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Laboratory Manual in General Microbiology
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- v
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
Front Matter
Preface
© The McGraw-H
Companies, 2001
Pref,
This eighth edition of Microbiological Applications
differs from the previous edition in that it has acquired
four new exercises and dropped three experiments. It
retains essentially the same format throughout, how-
ever. In response to requests for more emphasis on lab-
oratory safety, three new features have been incorpo-
rated into the text. In addition, several experiments
have been altered to improve simplicity and reliability.
The three exercises that were dropped pertain to fla-
gellar staining, bacterial conjugation, and nitrification in
soil. All of these exercises were either difficult to per-
form, unreliable, or of minimal pedagogical value.
To provide greater safety awareness in the labora-
tory, the following three features were added: (1) an
introductory laboratory protocol, (2) many cautionary
boxes dispersed throughout the text, and (3) a new ex-
ercise pertaining to aseptic technique.
The three-page laboratory protocol, which fol-
lows this preface, replaces the former introduction. It
provides terminology, safety measures, an introduc-
tion to aseptic technique, and other rules that apply to
laboratory safety.
To alert students to potential hazards in performing
certain experiments, caution boxes have been incorpo-
rated wherever they are needed. Although most of these
cautionary statements existed in previous editions, they
were not emphasized as much as they are in this edition.
Exercise 8 (Aseptic Technique) has been struc-
tured to provide further emphasis on culture tube han-
dling. In previous editions it was assumed that students
would learn these important skills as experiments were
performed. With the risk of being redundant, six pages
have been devoted to the proper handling of culture
tubes when making inoculation transfers.
Although most experiments remain unchanged,
there are a few that have been considerably altered.
Exercise 27 (Isolation of Anaerobic Phototrophic
Bacteria), in particular, is completely new. By using
the Winogradsky column for isolating and identifying
the phototrophic sulfur bacteria, it has been possible
to greatly enrich the scope of this experiment. Another
exercise that has been altered somewhat is Exercise
48, which pertains to oxidation and fermentation tests
that are used for identifying unknown bacteria.
The section that has undergone the greatest reor-
ganization is Part 10 (Microbiology of Soil). In the
previous edition it consisted of five exercises. In this
edition it has been expanded to seven exercises. A
more complete presentation of the nitrogen cycle is of-
fered in Exercise 58, and two new exercises (Exercises
61 and 62) are included that pertain to the isolation of
denitrifiers.
In addition to the above changes there has been
considerable upgrading of graphics throughout the
book. Approximately thirty-five illustrations have been
replaced. Several critical color photographs pertaining
to molds and physiological tests were also replaced to
bring about more faithful color representation.
I am greatly indebted to my editors, Jean Fornango
and Jim Smith, who made the necessary contacts for
critical reviews. As a result of their efforts the following
individuals have provided me with excellent sugges-
tions for improvement of this manual: Barbara Collins
at California Lutheran University, Thousand Oaks, CA
Alfred Brown of Auburn University, Auburn, AL
Lester A. Scharlin at El Camino College, Torrance, CA
and Hershell Hanks at Collin County Community
College, Piano, TX.
VII
Benson: Microbiological
Front Matter
Laboratory Protocol
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Companies, 2001
Laboratory Protocol
Welcome to the exciting field of microbiology ! The
intent of this laboratory manual is to provide you with
basic skills and tools that will enable you to explore a
vast microbial world. Its scope is incredibly broad in
that it includes a multitude of viruses, bacteria, proto-
zoans, yeasts, and molds. Both beneficial and harmful
ones will be studied. Although an in-depth study of
any single one of these groups could constitute a full
course by itself, we will be able to barely get ac-
quainted with them.
To embark on this study it will be necessary for
you to learn how to handle cultures in such a way that
they are not contaminated or inadvertently dispersed
throughout the classroom. This involves learning
aseptic techniques and practicing preventive safety
measures. The procedures outlined here address these
two aspects. It is of paramount importance that you
know all the regulations that are laid down here as
Laboratory Protocol.
Scheduling During the first week of this course
your instructor will provide you with a schedule of
laboratory exercises arranged in the order of their per-
formance. Before attending laboratory each day,
check the schedule to see what experiment or experi-
ments will be performed and prepare yourself so that
you understand what will be done.
Each laboratory session will begin with a short
discussion to brief you on the availability of materials
and procedures. Since the preliminary instructions
start promptly at the beginning of the period, it is ex-
tremely important that you are not late to class.
Personal Items When you first enter the lab, place
all personal items such as jackets, bags, and books in
some out of the way place for storage. Don't stack
them on your desktop. Desk space is minimal and
must be reserved for essential equipment and your
laboratory manual. The storage place may be a
drawer, locker, coatrack, or perimeter counter. Your
instructor will indicate where they should be placed.
Attire A lab coat or apron must be worn at all times
in the laboratory. It will protect your clothing from ac-
cidental contamination and stains in the lab. When
leaving the laboratory, remove the coat or apron. In
addition, long hair must be secured in a ponytail to
prevent injury from Bunsen burners and contamina-
tion of culture material.
Terminology
Various terms such as sterilization, disinfection, ger-
micides, sepsis, and aseptic techniques will be used
here. To be sure that you understand exactly what they
mean, the following definitions are provided.
Sterilization is a process in which all living mi-
croorganisms, including viruses, are destroyed. The
organisms may be killed with steam, dry heat, or in-
cineration. If we say an article is sterile, we understand
that it is completely free of all living microorganisms.
Generally speaking, when we refer to sterilization as it
pertains here to laboratory safety, we think, primarily,
in terms of steam sterilization with the autoclave. The
ultimate method of sterilization is to burn up the in-
fectious agents or incinerate them. All biological
wastes must ultimately be incinerated for disposal.
Disinfection is a process in which vegetative,
nonsporing microorganisms are destroyed. Agents
that cause disinfection are called disinfectants or
germicides. Such agents are used only on inanimate
objects because they are toxic to human and animal
tissues.
Sepsis is defined as the growth (multiplication) of
microorganisms in tissues of the body. The term asep-
sis refers to any procedure that prevents the entrance
of infectious agents into sterile tissues, thus prevent-
ing infection. Aseptic techniques refer to those prac-
tices that are used by microbiologists to exclude all
organisms from contaminating media or contacting
living tissues. Antiseptics are chemical agents (often
dilute disinfectants) that can be safely applied exter-
nally to human tissues to destroy or inhibit vegetative
bacteria.
Aseptic Techniques
When you start handling bacterial cultures as in
Exercises 9 and 10, you will learn the specifics of
aseptic techniques. Some of the basic things you will
do are as follows:
IX
Benson: Microbiological
Front Matter
Laboratory Protocol
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Companies, 2001
Laboratory Protocol
Hand Washing Before you start working in the lab,
wash your hands with a liquid detergent and dry them
with paper toweling. At the end of the period, before
leaving the laboratory, wash them again.
5.
glass. Don't try to pick up the glass fragments
with your fingers.
Contaminated material must never be placed in a
wastebasket.
Tabletop Disinfection. The first chore of the day
will be to sponge down your desktop with a disinfec-
tant. This process removes any dust that may be pre-
sent and minimizes the chances of bacterial contami-
nation of cultures that you are about to handle.
Your instructor will indicate where the bottles of
disinfectant and sponges are located. At the end of the
period before leaving the laboratory, perform the same
procedure to protect students that may occupy your desk
in the next class.
Bunsen Burner Usage When using a Bunsen burner
to flame loops, needles, and test tubes, follow the pro-
cedures outlined in Exercise 8. Inoculating loops and
needles should be heated until they are red-hot. Before
they are introduced into cultures, they must be allowed
to cool down sufficiently to prevent killing organisms
that are to be transferred.
If your burner has a pilot on it and you plan to use
the burner only intermittently, use it. If your burner
lacks a pilot, turn off the burner when it is not being
used. Excessive unnecessary use of Bunsen burners in
a small laboratory can actually raise the temperature
of the room. More important is the fact that unat-
tended burner flames are a constant hazard to hair,
clothing, and skin.
The proper handling of test tubes, while transfer-
ring bacteria from one tube to another, requires a cer-
tain amount of skill. Test-tube caps must never be
placed down on the desktop while you are making in-
oculations. Techniques that enable you to make trans-
fers properly must be mastered. Exercise 8 pertains to
these skills.
Pipetting Transferring solutions or cultures by
pipette must always be performed with a mechanical
suction device. Under no circumstances is pipetting
by mouth allowed in this laboratory.
Disposal of Cultures and Broken Glass The fol-
lowing rules apply to culture and broken glass disposal:
1 . Petri dishes must be placed in a plastic bag to be
autoclaved.
2. Unneeded test-tube cultures must be placed in a
wire basket to be autoclaved.
3. Used pipettes must be placed in a plastic bag for
autoclaving.
4. Broken glass should be swept up into a dustpan
and placed in a container reserved for broken
Accidental Spills
All accidental spills, whether chemical or biological,
must be reported immediately to your instructor.
Although the majority of microorganisms used in
this laboratory are nonpathogens, some pathogens
will be encountered. It is for this reason that we must
treat all accidental biological spills as if pathogens
were involved.
Chemical spills are just as important to report be-
cause some agents used in this laboratory may be car-
cinogenic; others are poisonous; and some can cause
dermal damage such as blistering and depigmentation.
Decontamination Procedure Once your instructor
is notified of an accidental spill, the following steps
will take place:
1. Any clothing that is contaminated should be
placed in an autoclavable plastic bag and auto-
claved.
2. Paper towels, soaked in a suitable germicide, such
as 5% bleach, are placed over the spill.
3. Additional germicide should be poured around
the edges of the spill to prevent further
aerosolization.
4. After approximately 20 minutes, the paper tow-
els should be scraped up off the floor with an
autoclavable squeegee into an autoclavable
dust pan.
5. The contents of the dust pan are transferred to an
autoclavable plastic bag, which may itself be
placed in a stainless steel bucket or pan for trans-
port to an autoclave.
6. All materials, including the squeegee and dust-
pan, are autoclaved.
Additional Important
Regulations
Here are a few additional laboratory rules:
1. Don't remove cultures, reagents, or other materi-
als from the laboratory unless you have been
granted specific permission.
2. Don't smoke or eat food in the laboratory.
3. Make it a habit to keep your hands away from your
mouth. Obviously, labels are never moistened
with the tongue; use tap water or self-adhesive la-
bels instead.
Benson: Microbiological
Front Matter
Laboratory Protocol
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Companies, 2001
4. Always clean up after yourself. Gram-stained
slides that have no further use to you should be
washed and dried and returned to a slide box.
Coverslips should be cleaned, dried, and returned.
Staining trays should be rinsed out and returned to
their storage place.
5. Return all bulk reagent bottles to places of storage.
6. Return inoculating loops and needles to your stor-
age container. Be sure that they are not upside
down.
Laboratory Protocol
7. If you have borrowed something from someone,
return it.
8 . Do not leave any items on your desk at the end of
the period.
9. Do not disturb another class at any time. Wait un-
til the class is dismissed.
10. Treat all instruments, especially microscopes,
with extreme care. If you don't understand how a
piece of equipment functions, ask your instructor.
1 1 . Work cooperatively with other students in group-
assigned experiments, but do your own analyses
of experimental results.
XI
Benson: Microbiological
1. Microscopy
Introduction
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Companies, 2001
Part
Microscopy
Although there are many kinds of microscopes available to the mi-
crobiologist today, only four types will be described here for our
use: the brightfield, darkfield, phase-contrast, and fluorescence
microscopes. If you have had extensive exposure to microscopy in
previous courses, this unit may not be of great value to you; how-
ever, if the study of microorganisms is a new field of study for you,
there is a great deal of information that you need to acquire about
the proper use of these instruments.
Microscopes in a college laboratory represent a considerable
investment and require special care to prevent damage to the
lenses and mechanicals. The fact that a laboratory microscope
may be used by several different individuals during the day and
moved around from one place to another results in a much greater
chance for damage and wear to occur than if the instrument were
used by only one individual.
The complexity of some of the more expensive microscopes
also requires that certain adjustments be made periodically.
Knowing how to make these adjustments to get the equipment to
perform properly is very important. An attempt is made in the five
exercises of this unit to provide the necessary assistance in getting
the most out of the equipment.
Microscopy should be as fascinating to the beginner as it is to
the professional of long standing; however, only with intelligent un-
derstanding can the beginner approach the achievement that oc-
curs with years of experience.
1
Benson: Microbiological
1. Microscopy
I.Brightfield Microscopy
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Companies, 2001
Brightfield Microscopy
A microscope that allows light rays to pass directly
through to the eye without being deflected by an in-
tervening opaque plate in the condenser is called a
brightfield microscope. This is the conventional type
of instrument encountered by students in beginning
courses in biology; it is also the first type to be used
in this laboratory.
All brightfield microscopes have certain things in
common, yet they differ somewhat in mechanical op-
eration. An attempt will be made in this exercise to
point out the similarities and differences of various
makes so that you will know how to use the instru-
ment that is available to you. Before attending the first
laboratory session in which the microscope will be
used, read over this exercise and answer all the ques-
tions on the Laboratory Report. Your instructor may
require that the Laboratory Report be handed in prior
to doing any laboratory work.
Care of the Instrument
Microscopes represent considerable investment and
can be damaged rather easily if certain precautions are
not observed. The following suggestions cover most
hazards.
Transport When carrying your microscope from
one part of the room to another, use both hands when
holding the instrument, as illustrated in figure 1.1. If
it is carried with only one hand and allowed to dangle
at your side, there is always the danger of collision
with furniture or some other object. And, incidentally,
under no circumstances should one attempt to carry
two microscopes at one time.
Clutter Keep your workstation uncluttered while
doing microscopy. Keep unnecessary books, lunches,
and other unneeded objects away from your work
area. A clear work area promotes efficiency and re-
sults in fewer accidents.
Electric Cord Microscopes have been known to
tumble off of tabletops when students have entangled
a foot in a dangling electric cord. Don't let the light
cord on your microscope dangle in such a way as to
hazard foot entanglement.
Lens Care At the beginning of each laboratory pe-
riod check the lenses to make sure they are clean. At
the end of each lab session be sure to wipe any im-
mersion oil off the immersion lens if it has been used.
More specifics about lens care are provided on page 5.
Dust Protection In most laboratories dustcovers
are used to protect the instruments during storage. If
one is available, place it over the microscope at the
end of the period.
Components
Before we discuss the procedures for using a micro-
scope, let's identify the principal parts of the instru-
ment as illustrated in figure 1.2.
Framework All microscopes have a basic frame
structure, which includes the arm and base. To this
framework all other parts are attached. On many of
the older microscopes the base is not rigidly attached
to the arm as is the case in figure 1.2; instead, a pivot
point is present that enables one to tilt the arm back-
ward to adjust the eyepoint height.
Stage The horizontal platform that supports the mi-
croscope slide is called the stage. Note that it has a
clamping device, the mechanical stage, which is
used for holding and moving the slide around on the
Figure 1.1 The microscope should be held firmly with
both hands while carrying it.
2
Benson: Microbiological
1. Microscopy
I.Brightfield Microscopy
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Companies, 2001
stage. Note, also, the location of the mechanical
stage control in figure 1.2.
Light Source In the base of most microscopes is po-
sitioned some kind of light source. Ideally, the lamp
should have a voltage control to vary the intensity of
light. The microscope in figure 1.2 has a knurled wheel
on the right side of its base to regulate the voltage sup-
plied to the light bulb. The microscope base in figure
1 .4 has a knob (the left one) that controls voltage.
Brightfield Microscopy • Exercise 1
Most microscopes have some provision for reduc-
ing light intensity with a neutral density filter. Such a
filter is often needed to reduce the intensity of light be-
low the lower limit allowed by the voltage control. On
microscopes such as the Olympus CH-2, one can simply
place a neutral density filter over the light source in the
base. On some microscopes a filter is built into the base.
Lens Systems All microscopes have three lens sys-
tems: the oculars, the objectives, and the condenser.
Oculars (Eyepieces)
Diopter Adjustment Ring
Rotatable Head
Nosepiece
Objective
Stage
ON/OFF Switch
Lock Screw
Arm
Mechanical Stage
Condenser
Coarse Adjustment Knob
Fine Adjustment Knob
Illuminator
Mechanical Stage Control
Voltage Regulator
Figure 1.2 The compound microscope
Courtesy of the Olympus Corporation, Lake Success, N.Y.
3
Benson: Microbiological
1. Microscopy
I.Brightfield Microscopy
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Companies, 2001
Exercise 1 • Brightfield Microscopy
Figure 1.3 illustrates the light path through these three
systems.
The ocular, or eyepiece, is a complex piece, lo-
cated at the top of the instrument, that consists of two
or more internal lenses and usually has a magnification
of 10 X . Although the microscope in figure 1 .2 has two
oculars (binocular), a microscope often has only one.
Three or more objectives are usually present.
Note that they are attached to a rotatable nosepiece,
which makes it possible to move them into position
over a slide. Objectives on most laboratory micro-
scopes have magnifications of 10 X, 45 X, and 100X,
designated as low power, high -dry, and oil immer-
sion, respectively. Some microscopes will have a
fourth objective for rapid scanning of microscopic
fields that is only 4 X .
The third lens system is the condenser, which is
located under the stage. It collects and directs the light
from the lamp to the slide being studied. The con-
denser can be moved up and down by a knob under
the stage. A diaphragm within the condenser regu-
lates the amount of light that reaches the slide.
Microscopes that lack a voltage control on the light
source rely entirely on the diaphragm for controlling
light intensity. On the Olympus microscope in figure
1.2 the diaphragm is controlled by turning a knurled
ring. On some microscopes a diaphragm lever is pres-
ent. Figure 1.3 illustrates the location of the condenser
and diaphragm.
Focusing Knobs The concentrically arranged
coarse adjustment and fine adjustment knobs on
the side of the microscope are used for bringing ob-
jects into focus when studying an object on a slide. On
some microscopes these knobs are not positioned con-
centrically as shown here.
Ocular Adjustments On binocular microscopes
one must be able to change the distance between the
oculars and to make diopter changes for eye differ-
ences. On most microscopes the interocular distance
is changed by simply pulling apart or pushing to-
gether the oculars.
To make diopter adjustments, one focuses first
with the right eye only. Without touching the focusing
knobs, diopter adjustments are then made on the left
eye by turning the knurled diopter adjustment ring
(figure 1.2) on the left ocular until a sharp image is
seen. One should now be able to see sharp images
with both eyes.
Figure 1.3 The light pathway of a microscope.
Resolution
The resolution limit, or resolving power, of a micro-
scope lens system is a function of its numerical aper-
ture, the wavelength of light, and the design of the
condenser. The optimum resolution of the best micro-
scopes with oil immersion lenses is around 0.2 |xm.
This means that two small objects that are 0.2 |jim
apart will be seen as separate entities; objects closer
than that will be seen as a single object.
To get the maximum amount of resolution from a
lens system, the following factors must be taken into
consideration:
• A blue filter should be in place over the light
source because the short wavelength of blue light
provides maximum resolution.
• The condenser should be kept at its highest posi-
tion where it allows a maximum amount of light
to enter the objective.
• The diaphragm should not be stopped down too
much. Although stopping down improves con-
trast, it reduces the numerical aperture.
• Immersion oil should be used between the slide
and the 100X objective.
Of significance is the fact that, as magnification is in-
creased, the resolution must also increase. Simply in-
creasing magnification by using a 20 X ocular won't
increase the resolution.
4
Benson: Microbiological
1. Microscopy
I.Brightfield Microscopy
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Companies, 2001
Lens Care
Keeping the lenses of your microscope clean is a con-
stant concern. Unless all lenses are kept free of dust,
oil, and other contaminants, they are unable to
achieve the degree of resolution that is intended.
Consider the following suggestions for cleaning the
various lens components:
Cleaning Tissues Only lint-free, optically safe tis-
sues should be used to clean lenses. Tissues free of
abrasive grit fall in this category. Booklets of lens
tissue are most widely used for this purpose.
Although several types of boxed tissues are also
safe, use only the type of tissue that is recommended
by your instructor.
Solvents Various liquids can be used for cleaning
microscope lenses. Green soap with warm water
works very well. Xylene is universally acceptable.
Alcohol and acetone are also recommended, but often
with some reservations. Acetone is a powerful solvent
that could possibly dissolve the lens mounting cement
in some objective lenses if it were used too liberally.
When it is used it should be used sparingly. Your in-
structor will inform you as to what solvents can be
used on the lenses of your microscope.
Oculars The best way to determine if your eyepiece
is clean is to rotate it between the thumb and forefin-
ger as you look through the microscope. A rotating
pattern will be evidence of dirt.
If cleaning the top lens of the ocular with lens
tissue fails to remove the debris, one should try
cleaning the lower lens with lens tissue and blowing
off any excess lint with an air syringe or gas cannis-
Brightfield Microscopy • Exercise 1
ter. Whenever the ocular is removed from the micro-
scope, it is imperative that a piece of lens tissue be
placed over the open end of the microscope as illus-
trated in figure 1.5.
Objectives Objective lenses often become soiled
by materials from slides or fingers. A piece of lens tis-
sue moistened with green soap and water, or one of
the acceptable solvents mentioned above, will usually
remove whatever is on the lens. Sometimes a cotton
swab with a solvent will work better than lens tissue.
At any time that the image on the slide is unclear or
cloudy, assume at once that the objective you are us-
ing is soiled.
Condenser Dust often accumulates on the top sur-
face of the condenser; thus, wiping it off occasionally
with lens tissue is desirable.
Procedures
If your microscope has three objectives you have three
magnification options: (1) low-power, or 100X total
magnification, (2) high-dry magnification, which is
450X total with a 45 X objective, and (3) 1000X total
magnification with a 100X oil immersion objective.
Note that the total magnification seen through an ob-
jective is calculated by simply multiplying the power
of the ocular by the power of the objective.
Whether you use the low-power objective or the oil
immersion objective will depend on how much magni-
fication is necessary. Generally speaking, however, it is
best to start with the low-power objective and progress
to the higher magnifications as your study progresses.
Consider the following suggestions for setting up your
microscope and making microscopic observations.
Figure 1.4 On this microscope, the left knob controls
voltage. The other knob is used for moving a neutral den-
sity filter into position.
Figure 1.5 When oculars are removed for cleaning,
cover the ocular opening with lens tissue. A blast from an
air syringe or gas cannister removes dust and lint.
5
Benson: Microbiological
1. Microscopy
I.Brightfield Microscopy
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Companies, 2001
Exercise 1 • Brightfield Microscopy
Viewing Setup If your microscope has a rotatable
head, such as the ones being used by the two students
in figure 1 .6, there are two ways that you can use the
instrument. Note that the student on the left has the
arm of the microscope near him, and the other student
has the arm away from her. With this type of micro-
scope, the student on the right has the advantage in
that the stage is easier to observe. Note, also that when
focusing the instrument she is able to rest her arm on
the table. The manufacturer of this type of microscope
intended that the instrument be used in the way
demonstrated by the young lady. If the microscope
head is not rotatable, it will be necessary to use the
other position.
Low-Power Examination The main reason for
starting with the low-power objective is to enable you
to explore the slide to look for the object you are plan-
ning to study. Once you have found what you are
looking for, you can proceed to higher magnifica-
tions. Use the following steps when exploring a slide
with the low-power objective:
1 . Position the slide on the stage with the material to
be studied on the upper surface of the slide.
Figure 1 .7 illustrates how the slide must be held
in place by the mechanical stage retainer lever.
2. Turn on the light source, using a minimum amount
of voltage. If necessary, reposition the slide so
that the stained material on the slide is in the ex-
act center of the light source.
3. Check the condenser to see that it has been raised
to its highest point.
4. If the low-power objective is not directly over the
center of the stage, rotate it into position. Be sure
that as you rotate the objective into position it
clicks into its locked position.
5. Turn the coarse adjustment knob to lower the ob-
jective until it stops. A built-in stop will prevent
the objective from touching the slide.
6. While looking down through the ocular (or ocu-
lars), bring the object into focus by turning the
fine adjustment focusing knob. Don't readjust the
coarse adjustment knob. If you are using a binoc-
ular microscope it will also be necessary to adjust
the interocular distance and diopter adjustment to
match your eyes.
7. Manipulate the diaphragm lever to reduce or in-
crease the light intensity to produce the clear-
est, sharpest image. Note that as you close
down the diaphragm to reduce the light inten-
sity, the contrast improves and the depth of
field increases. Stopping down the diaphragm
when using the low-power objective does not
decrease resolution.
8. Once an image is visible, move the slide about to
search out what you are looking for. The slide is
moved by turning the knobs that move the me-
chanical stage.
9. Check the cleanliness of the ocular, using the pro-
cedure outlined earlier.
10. Once you have identified the structures to be
studied and wish to increase the magnification,
you may proceed to either high-dry or oil immer-
sion magnification. However, before changing
objectives, be sure to center the object you wish
to observe.
High -Dry Examination To proceed from low-
power to high-dry magnification, all that is necessary
is to rotate the high-dry objective into position and
open up the diaphragm somewhat. It may be neces-
sary to make a minor adjustment with the fine adjust-
ment knob to sharpen up the image, but the coarse ad-
justment knob should not be touched.
If a microscope is of good quality, only minor
focusing adjustments are needed when changing
from low power to high- dry because all the objec-
tives will be parfocalized. Nonparfocalized micro-
Figure 1.6 The microscope position on the right has
the advantage of stage accessibility.
Figure 1 .7 The slide must be properly positioned as the
retainer lever is moved to the right.
6
Benson: Microbiological
1. Microscopy
I.Brightfield Microscopy
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Companies, 2001
scopes do require considerable refocusing when
changing objectives.
High-dry objectives should be used only on slides that
have cover glasses; without them, images are usually
unclear. When increasing the lighting, be sure to open
up the diaphragm first instead of increasing the volt-
age on your lamp; reason: lamp life is greatly ex-
tended when used at low voltage. If the field is not
bright enough after opening the diaphragm, feel free
to increase the voltage. A final point: Keep the con-
denser at its highest point.
Oil Immersion Techniques The oil immersion lens
derives its name from the fact that a special mineral oil
is interposed between the lens and the microscope
slide. The oil is used because it has the same refractive
index as glass, which prevents the loss of light due to
the bending of light rays as they pass through air. The
use of oil in this way enhances the resolving power of
the microscope. Figure 1.8 reveals this phenomenon.
Saved
Light Rays
/v^ri
Lost Light Rays
Due to Diffraction
Figure 1 Immersion oil, having the same refractive in
dex as glass, prevents light loss due to diffraction.
With parfocalized objectives one can go to oil
immersion from either low power or high-dry. On
some microscopes, however, going from low power
to high power and then to oil immersion is better.
Once the microscope has been brought into focus at
one magnification, the oil immersion lens can be ro-
tated into position without fear of striking the slide.
Before rotating the oil immersion lens into posi-
tion, however, a drop of immersion oil must be placed
on the slide. An oil immersion lens should never be
used without oil. Incidentally, if the oil appears
cloudy it should be discarded.
When using the oil immersion lens it is best to
open the diaphragm as much as possible. Stopping
Brightfield Microscopy • Exercise 1
down the diaphragm tends to limit the resolving power
of the optics. In addition, the condenser must be kept
at its highest point. If different colored filters are avail-
able for the lamp housing, it is best to use blue or
greenish filters to enhance the resolving power.
Since the oil immersion lens will be used exten-
sively in all bacteriological studies, it is of paramount
importance that you learn how to use this lens prop-
erly. Using this lens takes a little practice due to the
difficulties usually encountered in manipulating the
lighting. A final comment of importance: At the end of
the laboratory period remove all immersion oil from
the lens tip with lens tissue.
Putting It Away
When you take a microscope from the cabinet at the
beginning of the period, you expect it to be clean and
in proper working condition. The next person to use
the instrument after you have used it will expect the
same consideration. A few moments of care at the end
of the period will ensure these conditions. Check over
this list of items at the end of each period before you
return the microscope to the cabinet.
1. Remove the slide from the stage.
2. If immersion oil has been used, wipe it off the lens
and stage with lens tissue. (Do not wipe oil off
slides you wish to keep. Simply put them into a
slide box and let the oil drain off.)
3. Rotate the low-power objective into position.
4. If the microscope has been inclined, return it to an
erect position.
5. If the microscope has a built-in movable lamp,
raise the lamp to its highest position.
6. If the microscope has a long attached electric
cord, wrap it around the base.
7. Adjust the mechanical stage so that it does not
project too far on either side.
8. Replace the dustcover.
9. If the microscope has a separate transformer, re-
turn it to its designated place.
10. Return the microscope to its correct place in the
cabinet.
Laboratory Report
Before the microscope is to be used in the laboratory,
answer all the questions on Laboratory Report 1,2 that
pertain to brightfield microscopy. Preparation on your
part prior to going to the laboratory will greatly facil-
itate your understanding. Your instructor may wish to
collect this report at the beginning of the period on the
first day that the microscope is to be used in class.
7
Benson: Microbiological
1. Microscopy
2. Darkfield Microscopy
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Companies, 2001
Darkfield Microscopy
Delicate transparent living organisms can be more
easily observed with darkfield microscopy than with
conventional brightfield microscopy. This method is
particularly useful when one is attempting to identify
spirochaetes in the exudate from a syphilitic lesion.
Figure 2.1 illustrates the appearance of these organ-
isms under such illumination. This effect may be pro-
duced by placing a darkfield stop below the regular
condenser or by replacing the condenser with a spe-
cially constructed one.
Another application of darkfield microscopy is in
the fluorescence microscope (Exercise 4). Although
fluorescence may be seen without a dark field, it is
greatly enhanced with this application.
To achieve the darkfield effect it is necessary to
alter the light rays that approach the objective in such
a way that only oblique rays strike the objects being
viewed. The obliquity of the rays must be so extreme
that if no objects are in the field, the background is
completely light-free. Objects in the field become
brightly illuminated, however, by the rays that are re-
flected up through the lens system of the microscope.
Although there are several different methods for
producing a dark field, only two devices will be de-
scribed here: the star diaphragm and the cardioid con-
denser. The availability of equipment will determine
the method to be used in this laboratory.
The Star Diaphragm
One of the simplest ways to produce the darkfield
effect is to insert a star diaphragm into the filter slot
of the condenser housing as shown in figure 2.2.
This device has an opaque disk in the center that
blocks the central rays of light. Figure 2.3 reveals
the effect of this stop on the light rays passing
through the condenser. If such a device is not avail-
able, one can be made by cutting round disks of
opaque paper of different sizes that are cemented to
transparent celluloid disks that will fit into the slot.
If the microscope normally has a diffusion disk in
this slot, it is best to replace it with rigid clear cel-
luloid or glass.
An interesting modification of this technique is to
use colored celluloid stops instead of opaque paper.
Backgrounds of blue, red, or any color can be pro-
duced in this way.
In setting up this type of darkfield illumination it
is necessary to keep these points in mind:
1
2
Limit this technique to the study of large organ-
isms that can be seen easily with low-power mag-
nification. Good resolution with higher powered
objectives is difficult with this method.
Keep the diaphragm wide open and use as much
light as possible. If the microscope has a voltage
Figure 2.1 Transparent living microorganisms, such as
the syphilis spirochaete, can be seen much more easily
when observed in a dark field.
Figure 2.2 The insertion of a star diaphragm into the fil-
ter slot of the condenser will produce a dark field suitable
for low magnifications.
■
9
Benson: Microbiological
1. Microscopy
2. Darkfield Microscopy
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Companies, 2001
Exercise 2 • Darkfield Microscopy
regulator, you will find that the higher voltages
will produce better results.
3. Be sure to center the stop as precisely as possible.
4. Move the condenser up and down to produce the
best effects.
The Cardioid Condenser
The difficulty that results from using the star di-
aphragm or opaque paper disks with high-dry and oil
immersion objectives is that the oblique rays are not
as carefully metered as is necessary for the higher
magnifications. Special condensers such as the car-
dioid or paraboloid types must be used. Since the car-
dioid type is the most frequently used type, its use will
be described here.
Figure 2.4 illustrates the light path through such a
condenser. Note that the light rays entering the lower
element of the condenser are reflected first off a con-
vex mirrored surface and then off a second concave
surface to produce the desired oblique rays of light.
Once the condenser has been installed in the micro-
scope, the following steps should be followed to pro-
duce ideal illumination.
Materials:
slides and cover glasses of excellent quality
(slides of 1.15-1.25 mm thickness and
No. 1 cover glasses)
1. Adjust the upper surface of the condenser to a
height just below stage level.
2. Place a clear glass slide in position over the
condenser.
3. Focus the 10X objective on the top of the con-
denser until a bright ring comes into focus.
4. Center the bright ring so that it is concentric with
the field edge by adjusting the centering screws
on the darkfield condenser. If the condenser has a
light source built into it, it will also be necessary
to center it as well to achieve even illumination.
5. Remove the clear glass slide.
6. If a funnel stop is available for the oil immersion
objective, remove this object and insert this unit.
(This stop serves to reduce the numerical aperture
of the oil immersion objective to a value that is
less than the condenser.)
7. Place a drop of immersion oil on the upper surface
of the condenser and place the slide on top of the
oil. The following preconditions in slide usage
must be adhered to:
• Slides and cover glasses should be optically
perfect. Scratches and imperfections will cause
annoying diffractions of light rays.
• Slides and cover glasses must be free of dirt or
grease of any kind.
• A cover glass should always be used.
8. If the oil immersion lens is to be used, place a
drop of oil on the cover glass.
9. If the field does not appear dark and lacks con-
trast, return to the 10 X objective and check the
ring concentricity and light source centration. If
contrast is still lacking after these adjustments,
the specimen is probably too thick.
10. If sharp focus is difficult to achieve under oil im-
mersion, try using a thinner cover glass and
adding more oil to the top of the cover glass and
bottom of the slide.
Laboratory Report
This exercise may be used in conjunction with Part 2
when studying the various types of organisms. After
reading over this exercise and doing any special as-
signments made by your instructor, answer the ques-
tions on the last portion of Laboratory Report 1,2 that
pertain to darkfield microscopy.
Oblique Rays
Condenser
Reflected Rays
ii
Light Stop
Figure 2.3 The star diaphragm allows only peripheral
light rays to pass up through the condenser. This method
requires maximum illumination.
Glass Slide -
Cover Glass
^ ^ _ •m -, *•
■s'.lW
.1. 1
f r . ^ s ■' . m-
Immersion Oil
;■';>' J. -.1
Figure 2.4 A cardioid condenser provides greater
light concentration for oblique illumination than the
star diaphragm.
10
Benson: Microbiological
1. Microscopy
3. Phase-Contrast
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Microscopy
Companies, 2001
Phase- Contrast Microscopy
The difficulty that one encounters in trying to exam-
ine cellular organelles is that most protoplasmic ma-
terial is completely transparent and defies differentia-
tion. It is for this reason that stained slides are usually
used in brightfield cytological studies. Since the stain-
ing of slides results in cellular death, it is obvious that
when we study stained microorganisms on a slide, we
are observing artifacts rather than living cells.
A microscope that is able to differentiate trans-
parent protoplasmic structures without staining and
killing them is the phase-contrast microscope. The
first phase- contrast microscope was developed in
1933 by Frederick Zernike and was originally referred
to as the Zernike microscope. It is the instrument of
choice for studying living protozoans and other types
of transparent cells. Figure 3.1 illustrates the differ-
ences between brightfield and phase-contrast images.
Note the greater degree of differentiation that can be
seen inside cells when they are observed with phase-
contrast optics. In this exercise we will study the prin-
ciples that govern this type of microscope; we will
also see how different manufacturers have met the de-
sign challenges of these principles.
Image Contrast
Objects in a microscopic field may be categorized as
being either amplitude or phase objects. Amplitude
objects (illustration 1, figure 3.2) show up as dark ob-
jects under the microscope because the amplitude (in-
tensity) of light rays is reduced as the rays pass
through the objects. Phase objects (illustration 2, fig-
ure 3.2), on the other hand, are completely transparent
since light rays pass through them unchanged with re-
spect to amplitude. As some of the light rays pass
through phase objects, however, they are retarded by
l A wavelength.
This retardation, known as phase shift, occurs with
no amplitude diminution; thus, the objects appear
transparent rather than opaque. Since most biological
specimens are phase objects, lacking in contrast, it be-
comes necessary to apply dyes of various kinds to cells
that are to be studied with a brightfield microscope. To
understand how Zernike took advantage of the % wave-
length phase shift in developing his microscope we
must understand the difference between direct and dif-
fracted light rays.
BRIGHTFIELD
PHASE CONTRAST
Figure 3.1 Comparison of brightfield and phase-contrast images
11
Benson: Microbiological
1. Microscopy
3. Phase-Contrast
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Microscopy
Companies, 2001
Exercise 3 • Phase-Contrast Microscopy
Two Types of Light Rays
Light rays passing through a transparent object emerge
as either direct or diffracted rays. Those rays that pass
straight through unaffected by the medium are called
direct rays. They are unaltered in amplitude and
phase. The balance of the rays that are bent by their
slowing through the medium (due to density differ-
ences) emerge from the object as diffracted rays. It is
these rays that are retarded l A wavelength. Illustration
3, figure 3.2, illustrates these two types of light rays.
An important characteristic of these light rays is
that if the direct and diffracted rays of an object can be
brought into exact phase, or coincidence, with each
other, the resultant amplitude of the converged rays is
the sum of the two waves. This increase in amplitude
will produce increased brightness of the object in the
field. On the other hand, if two rays of equal ampli-
tude are in reverse phase QA wavelength off), their am-
plitudes cancel each other to produce a dark object.
This phenomenon is called interference. Illustration 4,
figure 3.2, shows these two conditions.
The Zernike Microscope
In constructing his first phase-contrast microscope,
Zernike experimented with various configurations of
diaphragms and various materials that could be used
to retard or advance the direct light rays. Figure 3.3 il-
lustrates the optical system of a typical modern phase-
contrast microscope. It differs from a conventional
brightfield microscope by having (1) a different type
of diaphragm and (2) a phase plate.
The diaphragm consists of an annular stop that
allows only a hollow cone of light rays to pass up
through the condenser to the object on the slide. The
phase plate is a special optical disk located at the rear
focal plane of the objective. It has a phase ring on it
that advances or retards the direct light rays % wave-
length.
Note in figure 3.3 that the direct rays converge on
the phase ring to be advanced or retarded l A wave-
length. These rays emerge as solid lines from the ob-
ject on the slide. This ring on the phase plate is coated
with a material that will produce the desired phase
shift. The diffracted rays, on the other hand, which
have already been retarded 1/4 wavelength by the
phase object on the slide, completely miss the phase
ring and are not affected by the phase plate. It should
be clear, then, that depending on the type of phase-
contrast microscope, the convergence of diffracted
and direct rays on the image plane will result in either
a brighter image (amplitude summation) or a darker
/T\ /'
Amplitude
i \
A f\
I
\
\
- v •
\
\
' i n i i
' v ^
A A \f\ A A A .
\j v \J v v v
/\ rt II
■>/X/
■J
J
\_y
Image
Wavelength
AMPLITUDE OBJECTS
h i- Va. Wavelength
^ A
A /\ '
V
A A
V
\,
.^
/
/
jL.
\
WW
ft/wv
■ -/
/
\
■-r- — ■-
mage
:S
/
PHASE OBJECTS
is
The extent to which the amplitude of light rays L
diminished determines the darkness of an object in
microscopic field.
Note that the retardation of light rays without amplitude
diminution results in transparent phase objects.
^
Illuminating
Beam
3rd Order
2nd Order
1 st Order
In Phase
^"~
" fc "*H.
/'
s.
/—
f
\
/
\
t
S
t
H
t
■ A
>*
^
Direct Ray
■s
v.
Reverse Phase
DIRECT AND DIFFRACTED RAYS
COINCIDENCE AND INTERFERENCE
A light ray passing through a slit or transparent object
emerges as a direct ray with several orders of
diffracted rays. The diffracted rays are 1 A wavelength
out of phase with the direct ray.
Note that when two light rays are in phase they will
unite to produce amplitude summation. Light rays in
reverse phase, however, cancel each other
(interference) to produce dark objects.
Figure 3.2 The utilization of light rays in phase-contrast microscopy
12
Benson: Microbiological
1. Microscopy
3. Phase-Contrast
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Microscopy
Companies, 2001
image (amplitude interference or reverse phase). The
former is referred to as bright phase microscopy; the
latter as dark phase microscopy. The apparent bright-
ness or darkness, incidentally, is proportional to the
square of the amplitude; thus, the image will be four
Phase-Contrast Microscopy • Exercise 3
times as bright or dark as seen through a brightfield
microscope.
It should be added here, parenthetically, that the
phase plates of some microscopes have coatings to
change the phase of the diffracted rays. In any event
Bright image with dark background results from light rays in exact
phase. Dark image with bright background results from light rays
in reverse phase.
— Image Plane
Direct light rays are retarded or
advanced V4 wavelength as they
pass through the phase ring.
Condenser
Amplitude contrast is achieved by these light
rays that are in phase or in reverse phase.
Phase Ring
Phase Plate
Most diffracted rays of light pass through
phase plate unchanged by missing phase
ring.
Diffracted rays (retarded Vi wavelength
after passing through phase objects).
Annular Stop
Figure 3.3 The optical system of a phase-contrast microscope
13
Benson: Microbiological
1. Microscopy
3. Phase-Contrast
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Microscopy
Companies, 2001
Exercise 3 • Phase-Contrast Microscopy
the end result will be the same: to achieve coincidence
or interference of direct and diffracted rays.
Microscope Adjustments
If the annular stop under the condenser of a phase-
contrast microscope can be moved out of position,
this instrument can also be used for brightfield stud-
ies. Although a phase-contrast objective has a phase
ring attached to the top surface of one of its lenses, the
presence of that ring does not seem to impair the res-
olution of the objective when it is used in the bright-
field mode. It is for this reason that manufacturers
have designed phase-contrast microscopes in such a
way that they can be quickly converted to brightfield
operation.
To make a microscope function efficiently in both
phase-contrast and brightfield situations one must
master the following procedures:
• lining up the annular ring and phase rings so that
they are perfectly concentric,
• adjusting the light source so that maximum illu-
mination is achieved for both phase-contrast and
brightfield usage, and
• being able to shift back and forth easily from
phase-contrast to brightfield modes. The follow-
ing suggestions should be helpful in coping with
these problems.
Alignment of Annulus and Phase Ring
Unless the annular ring below the condenser is
aligned perfectly with the phase ring in the objective,
good phase-contrast imagery cannot be achieved.
Figure 3.4 illustrates the difference between non-
alignment and alignment. If a microscope has only
one phase-contrast objective, there will be only one
annular stop that has to be aligned. If a microscope
has two or more phase objectives, there must be a
substage unit with separate annular stops for each
phase objective, and alignment procedure must be
performed separately for each objective and its annu-
lar stop.
Since the objective cannot be moved once it is
locked in position, all adjustments are made to the an-
nular stop. On some microscopes the adjustment may
be made with tools, as illustrated in figure 3.5. On
other microscopes, such as the Zeiss in figure 3.6
which has five phase-contrast objectives, the annular
rings are moved into position with special knobs on
the substage unit. Since the method of adjustment
varies from one brand of microscope to another, one
has to follow the instructions provided by the manu-
facturer. Once the adjustments have been made, they
Figure 3.4 The image on the right illustrates the ap-
pearance of the rings when perfect alignment of phase
ring and annulus diaphragm has been achieved.
Figure 3.5 Alignment of the annulus diaphragm and
phase ring is accomplished with a pair of Allen-type
screwdrivers on this American Optical microscope.
Figure 3.6 Alignment of the annulus and phase ring on
this Zeiss microscope is achieved by adjusting the two
knobs as shown.
14
Benson: Microbiological
1. Microscopy
3. Phase-Contrast
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Microscopy
Companies, 2001
Figure 3.7 If the ocular of a phase-contrast microscope
is replaced with a centering telescope, the orientation of
the phase ring and annular ring can be viewed.
Figure 3.8 Some microscopes have an aperture view-
ing unit that can be used instead of a centering telescope
for observing the orientation of the phase ring and annu-
lar ring.
Phase-Contrast Microscopy • Exercise 3
are rigidly set and needn't be changed unless someone
inadvertently disturbs them.
To observe ring alignment, one can replace the
eyepiece with a centering telescope as shown in fig-
ure 3.7. With this unit in place, the two rings can be
brought into sharp focus by rotating the focusing ring
on the telescope. Refocusing is necessary for each ob-
jective and its matching annular stop. Some manufac-
turers, such as American Optical, provide an aperture
viewing unit (figure 3.8), which enables one to ob-
serve the rings without using a centering telescope.
Zeiss microscopes have a unit called the Optovar,
which is located in a position similar to the American
Optical unit that serves the same purpose.
Light Source Adjustment
For both brightfield and phase-contrast modes it is
essential that optimum lighting be achieved. This is
no great problem for a simple setup such as the
American Optical instrument shown in figure 3.9.
For multiple phase objective microscopes, however,
(such as the Zeiss in figure 3.6) there are many more
adjustments that need to be made. A few suggestions
that highlight some of the problems and solutions
follow:
1
2
3
4
Figure 3.9 The annular stop on this American Optical
microscope has the annular stop located on a slideway.
When pushed in, the annular stop is in position.
Since blue light provides better images for both
phase-contrast and brightfield modes, make cer-
tain that a blue filter is placed in the filter holder
that is positioned in the light path. If the micro-
scope has no filter holder, placing the filter over
the light source on the base will help.
Brightness of field under phase-contrast is con-
trolled by adjusting the voltage or the iris di-
aphragm on the base. Considerably more light is
required for phase-contrast than for brightfield
since so much light is blocked out by the annu-
lar stop.
The evenness of illumination on some micro-
scopes, such as the Zeiss seen on these pages,
can be adjusted by removing the lamp housing
from the microscope and focusing the light spot
on a piece of translucent white paper. For the de-
tailed steps in this procedure, one should consult
the instruction manual that comes with the mi-
croscope. Light source adjustments of this na-
ture are not necessary for the simpler types of
microscopes.
Since each phase-contrast objective must be used
with a matching annular stop, make certain that
the proper annular stop is being used with the ob-
jective that is over the microscope slide. If image
quality is lacking, check first to see if the match-
ing annular stop is in position.
15
Benson: Microbiological
1. Microscopy
3. Phase-Contrast
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Microscopy
Companies, 2001
Exercise 3 • Phase-Contrast Microscopy
Working Procedures
Once the light source is correct and the phase ele-
ments are centered you are finally ready to examine
slide preparations. Keep in mind that from now on
most of the adjustments described earlier should
not be altered; however, if misalignment has oc-
curred due to mishandling, it will be necessary to
refer back to alignment procedures. The following
guidelines should be adhered to in all phase-con-
trast studies:
• Use only optically perfect slides and cover
glasses (no bubbles or striae in the glass).
• Be sure that slides and cover glasses are com-
pletely free of grease or chemicals.
• Use wet mount slides instead of hanging drop
preparations. The latter leave much to be desired.
Culture broths containing bacteria or protozoan
suspensions are ideal for wet mounts.
• In general, limit observations to living cells. In
most instances stained slides are not satisfactory.
The first time you use phase-contrast optics to ex-
amine a wet mount, follow these suggestions:
1 . Place the wet mount slide on the stage and bring
the material into focus, using brightfield optics at
low-power magnification.
2
3
4
5
6
Once the image is in focus, switch to phase op-
tics at the same magnification. Remember, it is
necessary to place in position the matching an-
nular stop.
Adjust the light intensity, first with the base di-
aphragm and then with the voltage regulator. In
most instances you will need to increase the
amount of light for phase-contrast.
Switch to higher magnifications, much in the
same way you do for brightfield optics, except
that you have to rotate a matching annular stop
into position.
If an oil immersion phase objective is used, add
immersion oil to the top of the condenser as well
as to the top of the cover glass.
Don't be disturbed by the "halo effect" that you
observe with phase optics. Halos are normal.
Laboratory Report
This exercise may be used in conjunction with Part 2
in studying various types of organisms. Organelles in
protozoans and algae will show up more distinctly
than with brightfield optics. After reading this exer-
cise and doing any special assignments made by your
instructor, answer the questions on combined
Laboratory Report 3-5 that pertain to this exercise.
16
Benson: Microbiological
1. Microscopy
4. Fluorescence
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Microscopy
Companies, 2001
Fluorescence Microscopy
The fluorescence microscope is a unique instrument
that is indispensible in certain diagnostic and research
endeavors. Differential dyes and immunofluores-
cence techniques have made laboratory diagnosis of
many diseases much simpler with this type of micro-
scope than with the other types described in Exercises
1, 2, and 3. If you are going to prepare and study any
differential fluorescence slides that are described in
certain exercises in this manual, you should have a ba-
sic understanding of the microscope's structure, its
capabilities, and its limitations. In addition, it is im-
portant that one be aware of the potential of experi-
encing eye injury if one of these instruments is not
used in a safe manner.
A fluorescence microscope differs from an ordi-
nary brightfield microscope in several respects. First
of all, it utilizes a powerful mercury vapor arc lamp
for its light source. Secondly, a darkfield condenser is
usually used in place of the conventional Abbe bright-
field condenser. The third difference is that it employs
three sets of filters to alter the light that passes up
through the instrument to the eye. Some general prin-
ciples related to its operation will follow an explana-
tion of the principle of fluorescence.
The Principle of Fluorescence
It was pointed out in the last exercise that light exists
as a form of energy propagated in wave form. An in-
teresting characteristic of such an electromagnetic
wave is that it can influence the electrons of mole-
cules that it encounters, causing significant interac-
tion. Those electrons within a molecule that are not
held too securely may be set in motion by the oscilla-
tions of the light beam. Not only are these electrons
interrupted from their normal pathways, but they are
also forced to oscillate in resonance with the passing
light wave. This excitation, caused by such oscilla-
tion, requires energy that is supplied by the light
beam. When we say that a molecule absorbs light, this
is essentially what is taking place.
Whenever a physical body absorbs energy, as in
the case of the activated molecule, the energy doesn't
just disappear; it must reappear again in some other
form. This new manifestation of the energy may be in
the form of a chemical reaction, heat, or light. If light
is emitted by the energized molecules, the phenome-
non is referred to as photoluminescence. In photolu-
minescence there is always a certain time lapse be-
tween the absorption and emission of light. If the time
lag is greater than 1/10,000 of a second it is generally
called phosphorescence. On the other hand, if the
time lapse is less than 1/10,000 of a second, it is
known as fluorescence.
Thus, we see that fluorescence is initiated when a
molecule absorbs energy from a passing wave of light.
The excited molecule, after a brief period of time, will
return to its fundamental energy state after emitting
fluorescent light. It is significant that the wavelength
of fluorescence is always longer than the exciting
light. This follows Stokes' law, which applies to liq-
uids but not to gases. This phenomenon is due to the
fact that energy loss occurs in the process so that the
emitting light has to be of a longer wavelength. This
energy loss, incidentally, occurs as a result of the mo-
bilization of the comparatively heavy atomic nuclei of
the molecules rather than the displacement of the
lighter electrons.
Microbiological material that is to be studied with
a fluorescence microscope must be coated with special
compounds that possess this quality of fluorescence.
Such compounds are called fluorochromes. Auramine
O, acridine orange, and fluorescein are well-known
fluorochromes. Whether a compound will fluoresce
will depend on its molecular structure, the tempera-
ture, and the pH of the medium. The proper prepara-
tion and use of fluorescent materials for microbiologi-
cal work must take all these factors into consideration.
Microscope Components
Figure 4.2 illustrates, diagrammatically, the light
pathway of a fluorescence microscope. The essential
components are the light source, heat filter, exciter fil-
ter, condenser, and barrier filter. The characteristics
and functions of each item follow.
17
Benson: Microbiological
1. Microscopy
4. Fluorescence
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Microscopy
Companies, 2001
Exercise 4 • Fluorescence Microscopy
Light Source The first essential component of a
fluorescence microscope is its bright mercury vapor
arc lamp. Such a bulb is preferred over an incandes-
cent one because it produces an ample supply of
shorter wavelengths of light (ultraviolet, violet, and
blue) that are needed for good fluorescence. To pro-
duce the arc in one of these lamps, voltages as high as
18,000 volts are required; thus, a power supply trans-
former is always used.
The wavelengths produced by these lamps in-
clude the ultraviolet range of 200-400 nm, the visible
range of 400-780 nm, and the long infrared rays that
are above 780 nm.
Mercury vapor arc lamps are expensive and po-
tentially dangerous. Certain precautions must be
taken, not only to promote long bulb life, but to pro-
tect the user as well. One of the hazards of these
bulbs is that they are pressurized and can explode.
Another hazard exists in direct exposure of the eyes
to harmful rays. Knowledge of these hazards is es-
sential to safe operation. If one follows certain pre-
cautionary measures, there is little need for anxiety.
However, one should not attempt to use one of these
instruments without a complete understanding of its
operation.
- i
^ 1
■B ^Cm ft- ^^^3
■ 1 M ■
^^^^^B
r 1
■ 1
^^^1
i
- 1
I i i
^VH
™
^3i m ' ■
^^Bj
tm
^^^^^^H
^^^^H
^^
^^^^ ^II ^^r *^^L 1
I
^^^^^
^^^■s
^^■^^^^fl
Figure 4.1 An early model American Optical fluores-
cence illuminator (Fluorolume) that could be adapted to
an ordinary darkfield microscope.
the majority of the ultraviolet light rays are deflected
by the condenser, protecting the observer's eyes. To
achieve this, the numerical aperture of the objective is
always 0.05 less than that of the condenser.
Heat Filter The infrared rays generated by the
mercury vapor arc lamp produce a considerable
amount of heat. These rays serve no useful purpose
in fluorescence and place considerable stress on
the filters within the system. To remove these rays,
a heat-absorbing filter is the first element in front
of the condensers. Ultraviolet rays, as well as most
of the visible spectrum, pass through this filter
unimpeded.
Exciter Filter After the light has been cooled down
by the heat filter it passes through the exciter filter,
which absorbs all the wavelengths except the short
ones needed to excite the fluorochrome on the slide.
These filters are very dark and are designed to let
through only the green, blue, violet, or ultraviolet
rays. If the exciter filter is intended for visible light
(blue, green, or violet) transmission, it will also allow
ultraviolet transmittance.
Condenser To achieve the best contrast of a fluo-
rescent object in the microscopic field, a darkfield
condenser is used. It must be kept in mind that weak
fluorescence of an object in a brightfield would be dif-
ficult to see. The dark background produced by the
darkfield condenser, thus, provides the desired con-
trast. Another bonus of this type of condenser is that
Barrier Filter This filter is situated between the ob-
jective and the eyepiece to remove all remnants of the
exciting light so that only the fluorescence is seen.
When ultraviolet excitation is employed with its very
dark, almost black- appearing exciter filters, the corre-
sponding barrier filters appear almost colorless. On
the other hand, when blue exciter filters are used, the
matching barrier filters have a yellow to deep orange
color. In both instances, the significant fact is that the
barrier filter should cut off precisely the shorter ex-
citer wavelengths without affecting the longer fluo-
rescence wavelengths.
Use of the Microscope
As in the case of most sophisticated equipment of this
type, it is best to consult the manufacturer's instruc-
tion manual before using it. Although different makes
of fluorescence microscopes are essentially alike in
principle, they may differ considerably in the fine
points of operation. Since it is not possible to be ex-
plicit about the operation of all makes, all that will be
attempted here is to generalize.
Some Precautions To protect yourself and others it
is well to outline the hazards first. Keep the following
points in mind:
18
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
I. Microscopy
4. Fluorescence
Microscopy
© The McGraw-H
Companies, 2001
Fluorescence Microscopy • Exercise 4
L
_J
fcn
=3
BARRIER FILTER
Removes any exciter wavelengths that
get past condenser without absorbing
longer wavelenghts of fluorescing objects.
■ ■■■!■■
FLUOROCHROME
Emits fluorescence due to activation
by exciting wavelength of light.
=a
DARKFIELD CONDENSER
Provides high contrast for
fluorescence.
MERCURY VAPOR
ARC LAMP
*
HEAT FILTER
Removes infrared rays
EXCITER FILTER
Allows only high-energy short
wavelengths through.
Figure 4.2 The light pathway of a fluorescence microscope
19
Benson: Microbiological
1. Microscopy
4. Fluorescence
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Microscopy
Companies, 2001
Exercise 4 • Fluorescence Microscopy
1
2
3
Remember that the pressurized mercury arc lamp
is literally a potential bomb. Design of the equip-
ment is such, however, that with good judgment,
no injury should result. When these lamps are
cold they are relatively safe, but when hot, the in-
side pressure increases to eight atmospheres, or
112 pounds per square inch.
The point to keep in mind is this — never at-
tempt to inspect the lamp while it is hot. Let it
cool completely before opening up the lamp
housing. Usually, 15 to 20 minutes cooling time
is sufficient.
Never expose your eyes to the direct rays of the
mercury arc lamp. Equipment design is such that
the bulb is always shielded against the scattering
of its rays. Remember that the unfiltered light
from one of these lamps is rich in both ultraviolet
and infrared rays — both of which are damaging to
the eyes. Severe retinal burns can result from ex-
posure to the mercury arc rays.
Be sure that the barrier filter is always in place
when looking down through the microscope.
Removal of the barrier filter or exciter filter or
both filters while looking through the microscope
could cause eye injury. It is possible to make mis-
takes of this nature if one is not completely famil-
iar with the instrument. Remember, the function
of the barrier filter is to prevent traces of ultravi-
olet light from reaching the eyes without blocking
wavelengths of fluorescence.
Warm-up Period The lamps in fluorescence mi-
croscopes require a warm-up period. When they are
first turned on the illumination is very low, but it in-
creases to maximum in about 2 minutes. Optimum il-
lumination occurs when the equipment has been op-
erating for 30 minutes or more. Most manufacturers
recommend leaving the instruments turned on for an
hour or more when using them. It is not considered
good economy to turn the instrument on and off sev-
eral times within a 2- or 3-hour period.
Keeping a Log The life expectancy of a mercury
arc lamp is around 400 hours. A log should be kept of
the number of hours that the instrument is used so that
inspection can be made of the bulb at approximately
200 hours. A card or piece of paper should be kept
conveniently near the instrument so that the individ-
ual using the instrument is reminded to record the
time that the instrument is turned on and off.
Filter Selection The most frequently used filter
combination is the bluish Schott BG12 (AO #702) ex-
citer and the yellowish Schott OG1 barrier filters.
CO
CO
E
CO
c
CO
OG1
Barrier
Filter
350
400 450 500
Wavelength (NM)
550
Figure 4.3
filters
Spectral transmissions of BG12 and OG1
Figure 4.3 shows the wavelength transmission of each
of these filters. Note that the exciter filter gives peak
emission of light in the 400 nm area of the spectrum.
These rays are violet. It allows practically no green or
yellow wavelengths through. The shortest wave-
lengths that this barrier filter lets through are green to
greenish-yellow.
If a darker background is desired than is being
achieved with the above filters, one may add a pale
blue Schott BG38 to the system. It may be placed on
either side of the heat filter, depending on the type of
equipment being used. If it is placed between the
lamp and heat filter, it will also function as another
heat filter.
Examination When looking for material on the
slide, it is best to use low- or high-power objectives.
If the illuminator is a separate unit, as in figure 4.1, it
may be desirable to move the illuminator out of posi-
tion and use incandescent lighting for this phase of the
work. Once the desirable field has been located, the
mercury vapor arc illuminator can be moved into po-
sition. One problem with fluorescence microscopes is
that most darkfield condensers do not illuminate well
through the low-power objectives (exception: the
Reichert-Toric setup used on some American Optical
instruments).
Keep in mind that there is no diaphragm control
on darkfield condensers. Some instruments are sup-
plied with neutral density filters to reduce light inten-
sity. The best system of illumination control, how-
ever, is achieved with objectives that have a built-in
iris control. These objectives have a knurled ring that
can be rotated to control the contrast.
20
Benson: Microbiological
1. Microscopy
4. Fluorescence
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Microscopy
Companies, 2001
For optimum results it is essential that oil be used
between the condenser and the slide. And, of course,
if the oil immersion lens is used, the oil must also be
interposed between the slide and the objective. It is
also important that special low-fluorescing immer-
sion oil be used. Ordinary immersion oil should be
avoided.
Although the ocular of a fluorescence microscope
is usually 10X, one should not hesitate to try other
size oculars if they are available. With bright-field mi-
croscopes it is generally accepted that nothing is
Fluorescence Microscopy • Exercise 4
gained by going beyond 1000X magnification. In a
fluorescence microscope, however, the image is
formed in a manner quite different from its brightfield
counterpart, obviating the need for following the
1000X rule. The only loss by using the higher magni-
fication is some brightness.
Laboratory Report
Complete all the answers to the questions on
Laboratory Report 3-5 that pertain to this exercise.
Lamp Condenser
Focus
Neutral Density
Filters
Field Diaphragm
Centering
Exciter Filters
Field Diaphragm Lever
Figure 4.4 An American Optical fluorescence microscope
21
Benson: Microbiological
1. Microscopy
5. Microscopic
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Measurements
Companies, 2001
Microscopic Measurements
With an ocular micrometer properly installed in the
eyepiece of your microscope, it is a simple matter to
measure the size of microorganisms that are seen in
the microscopic field. An ocular micrometer con-
sists of a circular disk of glass that has graduations
engraved on its upper surface. These graduations ap-
pear as shown in illustration B, figure 5.4. On some
microscopes one has to disassemble the ocular so that
the disk can be placed on a shelf in the ocular tube be-
tween the two lenses. On most microscopes, how-
ever, the ocular micrometer is simply inserted into
the bottom of the ocular, as shown in figure 5.1.
Before one can use the micrometer it is necessary to
calibrate it for each of the objectives by using a stage
micrometer.
The principal purpose of this exercise is to show
you how to calibrate an ocular micrometer for the
various objectives on your microscope. Proceed as
follows:
Calibration Procedure
The distance between the lines of an ocular microm-
eter is an arbitrary value that has meaning only if the
ocular micrometer is calibrated for the objective that
is being used. A stage micrometer (figure 5.2), also
known as an objective micrometer, has lines scribed
on it that are exactly 0.01 mm (10 jxm) apart.
Illustration C, figure 5.4 reveals the appearance of
these graduations.
To calibrate the ocular micrometer for a given
objective, it is necessary to superimpose the two
scales and determine how many of the ocular grad-
uations coincide with one graduation on the scale of
the stage micrometer. Illustration A in figure 5.4
shows how the two scales appear when they are
properly aligned in the microscopic field. In this
case, seven ocular divisions match up with one
stage micrometer division of 0.01 mm to give an oc-
ular value of 0.01/7, or 0.00143 mm. Since there are
1000 micrometers in 1 millimeter, these divisions
are 1 .43 |xm apart.
With this information known, the stage microme-
ter is replaced with a slide of organisms to be mea-
sured. Illustration D, figure 5.4, shows how a field of
microorganisms might appear with the ocular mi-
Figure 5.1 Ocular micrometer with retaining ring is in
serted into base of eyepiece.
yj^ta r -^^b
V IF "™JI n
■^D
_^HIHI_ ^ ^^™ ^^fc^P
^^r .
-^|^^H
Ptn
L.
^^^^^^^ "^^^H
pr
^H
r Li
■■ i^H
^^n~
^^^^^^^H^^^^^^^^^^^^^H
Ml^^^
^^^^^^ ,
^^^^^_ _
^^^
»^""
BF jM
Figure 5.2 Stage micrometer is positioned by centering
small glass disk over the light source.
1
■i
lyS" t
I^^^^^^IB
Figure 5.3 After calibration is completed, stage mi-
crometer is replaced with slide for measurements.
22
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
I. Microscopy
5. Microscopic
Measurements
© The McGraw-H
Companies, 2001
Microscopic Measurements • Exercise 5
View showing appearance of ocular
micrometer graduations. Spacing is
arbitrary.
View showing the alignment of stage
micrometer graduations (X) with ocular
micrometer graduations (Y). Since one
space of X (0.01 mm) is occupied by 7
spaces of Y, one space of Y =
.01
= .0014 mm, or 1.4 micrometers.
"• -£; ' 4 sS
." 5Sr£
Appearance of stage micrometer
graduations. Lines are exactly 0.01
mm (10 micrometers) apart.
mmmmzi
XJW
.V.-*.*
'* <
« r^
,l^^C/^£&!^^ *Y<
V.
' *r
^
lKi-££r*
'-r*';j
#
«*
L-V
-^
at"*;---:
»v
D
.O ■>- *5 n ->
s»*
*'#*»
On the basis of the calibration
calculations in view A above, what is
the total length of the yeast cell and
bud in this view?
Figure 5.4 Calibration of ocular micrometer
23
Benson: Microbiological
1. Microscopy
5. Microscopic
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Measurements
Companies, 2001
Exercise 5 • Microscopic Measurements
crometer in the eyepiece. To determine the size of an
organism, then, it is a simple matter to count the grad-
uations and multiply this number by the known dis-
tance between the graduations. When calibrating the
objectives of a microscope, proceed as follows.
Materials:
ocular micrometer or eyepiece that contains a
micrometer disk
stage micrometer
1 . If eyepieces are available that contain ocular mi-
crometers, replace the eyepiece in your micro-
scope with one of them. If it is necessary to in-
sert an ocular micrometer in your eyepiece, find
out from your instructor whether it is to be in-
serted below the bottom lens or placed between
the two lenses within the eyepiece. In either case,
great care must be taken to avoid dropping the
eyepiece or reassembling the lenses incorrectly.
Only with your instructor's prior approval shall
eyepieces be disassembled. Be sure that the grad-
uations are on the upper surface of the glass disk.
2. Place the stage micrometer on the stage and cen-
ter it exactly over the light source.
3 . With the low-power ( 1 X ) obj ecti ve in position,
bring the graduations of the stage micrometer
into focus, using the coarse adjustment knob.
Reduce the lighting. Note: If the microscope has
an automatic stop, do not use it as you normally
would for regular microscope slides. The stage
micrometer slide is too thick to allow it to func-
tion properly.
4. Rotate the eyepiece until the graduations of the
ocular micrometer lie parallel to the lines of the
stage micrometer.
5. If the low-power objective is the objective to be
calibrated, proceed to step 8 .
6. If the high-dry objective is to be calibrated,
swing it into position and proceed to step 8.
7. If the oil immersion lens is to be calibrated, place
a drop of immersion oil on the stage micrometer,
swing the oil immersion lens into position, and
bring the lines into focus; then, proceed to the
next step.
8
9
Move the stage micrometer laterally until the
lines at one end coincide. Then look for another
line on the ocular micrometer that coincides ex-
actly with one on the stage micrometer.
Occasionally one stage micrometer division will
include an even number of ocular divisions, as
shown in illustration A. In most instances, how-
ever, several stage graduations will be involved.
In this case, divide the number of stage microme-
ter divisions by the number of ocular divisions
that coincide. The figure you get will be that part
of a stage micrometer division that is seen in an
ocular division. This value must then be multi-
plied by 0.01 mm to get the amount of each ocu-
lar division.
Example: 3 divisions of the stage micrometer line
up with 20 divisions of the ocular micro-meter.
Each ocular division =
3
X 0.01
20
= 0.0015 mm
= 1.5 ^m
Replace the stage micrometer with slides of or
ganisms to be measured.
Measuring Assignments
Organisms such as protozoans, algae, fungi, and bac-
teria in the next few exercises may need to be mea-
sured. If your instructor requires that measurements
be made, you will be referred to this exercise.
Later on you will be working with unknowns. In
some cases measurements of the unknown organisms
will be pertinent to identification.
If trial measurements are to be made at this time,
your instructor will make appropriate assignments.
Important: Remove the ocular micrometer from
your microscope at the end of the laboratory period.
Laboratory Report
Answer the questions on combined Laboratory
Report 3-5 that pertain to this exercise.
24
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
II. Survey of
Microorganisms
Introduction
© The McGraw-H
Companies, 2001
Part
Survey of Microorganisms
Too often, in our serious concern with the direct applications of mi-
crobiology to human welfare, we neglect the large number of inter-
esting free-living microorganisms that abound in the water, soil,
and air. It is these free-spirited forms that we will study in the four
exercises of this unit. To observe these organisms we will examine
samples of pond water and Petri plates with special media that
have been exposed to the air and various items in our environment.
The principal organisms that we will encounter are protozoans, al-
gae, molds, yeasts, cyanobacteria, and bacteria.
The phylogenetic tree on this page illustrates where these or-
ganisms fit in the evolutionary scheme of organisms. The organ-
isms that you are likely to encounter are underlined on the diagram.
A few comments about each domain are presented here.
Domain Archaea Since the principal habitats of these organisms
are extreme environments such as volcanic waters, hot springs, or
waters of high salt conditions, you will not encounter any of these or-
ganisms in this study. These ancient organisms that exist in such
hostile environments have often been referred to as "extremophiles."
Domain Eukarya The protozoans, algae, and fungi fall in this do-
main. All members of this domain have distinct nuclei with nuclear
membranes and mitochondria. Some eukaryotes, such as the al-
gae, have chloroplasts, which puts them in the plant kingdom. The
eukaryotes appear to be more closely related to the Archaea than
to the Bacteria.
Domain Bacteria Members of this domain are also called
"prokaryotes." They are smaller than eukaryotes, lack distinct nu-
clei (no nuclear membrane), and are enclosed in a rigid cell wall with
a distinct cell membrane. In this study you will encounter various
species of cyanobacteria and bacteria.
ANftWti
PIANT5
'.iN^HSA ^OISWH
«CHHKHDA
tPPlCMWIADS
From: Extremophiles. Michael T. Madigan and Barry L. Marrs in Scientific American Vol. 276, Number 4, pages 82-87, April 1997.
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
II. Survey of
Microorganisms
6. Protozoa, Algae, and
Cyanobacteria
© The McGraw-H
Companies, 2001
Protozoa, Algae, and Cyanobacteria
In this exercise a study will be made of protozoans,
algae, and cyanobacteria that are found in pond wa-
ter. Bottles that contain water and bottom debris
from various ponds will be available for study.
Illustrations and text provided in this exercise will
be used to assist you in an attempt to identify the
various types that are encountered. Unpigmented,
moving microorganisms will probably be proto-
zoans. Greenish or golden-brown organisms are
usually algae. Organisms that appear blue-green
will be cyanobacteria. Supplementary books on the
laboratory bookshelf will also be available for as-
sistance in identifying organisms that are not de-
scribed in the short text of this exercise. If you en-
counter invertebrates and are curious as to their
identification, you may refer to Exercise 7; how-
ever, keep in mind that our prime concern here is
only with protozoans, algae, and cyanobacteria.
The purpose of this exercise is, simply, to provide
you with an opportunity to become familiar with the
differences between the three groups by comparing
their characteristics. The extent to which you will be
held accountable for the names of various organisms
will be determined by your instructor. The amount of
time available for this laboratory exercise will deter-
mine the depth of scope to be pursued.
To study the microorganisms of pond water, it
will be necessary to make wet mount slides. The
procedure for making such slides is relatively sim-
ple. All that is necessary is to place a drop of sus-
pended organisms on a microscope slide and cover
it with a cover glass. If several different cultures are
available, the number of the bottle should be
recorded on the slide with a china marking pencil.
As you prepare and study your slides, observe the
following guidelines:
Materials:
bottles of pond-water samples
microscope slides and cover glasses
rubber-bulbed pipettes and forceps
china marking pencil
reference books
1 . Clean the slide and cover glass with soap and wa-
ter, rinse thoroughly, and dry. Do not attempt to
study a slide that lacks a cover glass.
2
3
4
5
6
7
When using a pipette, insert it into the bottom of
the bottle to get a maximum number of organ-
isms. Very few organisms will be found swim-
ming around in middepth of the bottle.
To remove filamentous algae from a specimen
bottle, use forceps. Avoid putting too much mate-
rial on the slides.
Explore the slide first with the low-power objec-
tive. Reduce the lighting with the iris diaphragm.
Keep the condenser at its highest point.
When you find an organism of interest, swing the
high-dry objective into position and adjust the
lighting to get optimum contrast. If your micro-
scope has phase-contrast elements, use them.
Refer to Figures 6.1 through 6.6 and the text on
these pages to identify the various organisms that
you encounter.
Record your observations on the Laboratory
Reports.
The Protists
Single-celled eukaryons that lack tissue specialization
are called protists. Protozoologists group all protists
in Kingdom Protista. Those protists that are animal-
like are put in Subkingdom Protozoa and the protists
that are plantlike fall into Subkingdom Algae. This
system of classification includes all colonial species
as well as the single-celled types.
Subkingdom Protozoa
Externally, protozoan cells are covered with a cell
membrane, or pellicle; cell walls are absent; and dis-
tinct nuclei with nuclear membranes are present.
Specialized organelles, such as contractile vacuoles,
cytostomes, mitochondria, ribosomes, flagella, and
cilia, may also be present.
All protozoa produce cysts, which are resistant dor-
mant stages that enable them to survive drought, heat,
and freezing. They reproduce asexually by cell division
and exhibit various degrees of sexual reproduction.
The Subkingdom Protozoa is divided into
three phyla: Sarcomastigophora, Ciliophora, and
Apicomplexa. Type of locomotion plays an impor-
tant role in classification here. A brief description
of each phylum follows:
26
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
II. Survey of
Microorganisms
6. Protozoa, Algae, and
Cyanobacteria
© The McGraw-H
Companies, 2001
Protozoa, Algae, and Cyanobacteria • Exercise 6
~50/im
-0
-0
r
40JJI*
8
-0
|-20y m
-0
10
-25>jm
-0
-JOOv*)
-0
-400pm
L
13
14
15
16
-55pm
-0
-30pm
19
20
21
Lo
-1 5^m
-0
"|C
h20pm
-0
11
17
-20pm
-0
-lOOjjm
6
H4,0pm
-0
12
18
-J0pm
22
23
24
1 . Heteronema
2. Cercomonas
3. Co do si ga
4. Protospongia
5. T rich amoeba
Figure 6.1 Protozoans
6. Amoeba
7. May ore i la
8. Diffugia
9. Paramecium
10. Lacryrnaria
1 1 . Lionotus
12. Loxodes
13. Blepharisma
14. Coieps
15. Condyiostoma
16. Stentor
17, Vorticelta
13. Carchesium
19. Zoothamnium
20. Sty (onychia
21. Orsychodromos
11. Hypotrichidium
23. Eupiotes
24. Didinium
27
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
II. Survey of
Microorganisms
6. Protozoa, Algae, and
Cyanobacteria
© The McGraw-H
Companies, 2001
Exercise 6 • Protozoa, Algae, and Cyanobacteria
Phylum Sarcomastigophora
Members of this phylum have been subdivided into
two subphyla: Sarcodina and Mastigophora.
Sarcodina (Amoebae) Members of this subphylum
move about by the formation of flowing protoplasmic
projections called pseudopodia. The formation of
pseudopodia is commonly referred to as amoeboid
movement. Illustrations 5 through 8 in figure 6.1 are
representative amoebae.
Mastigophora (Zooflagellates) These protozoans
possess whiplike structures called flagella. There is
considerable diversity among the members of this
group. Only a few representatives (illustrations 1
through 4) are seen in figure 6.1.
Phylum Ciliophora
These microorganisms are undoubtedly the most ad-
vanced and structurally complex of all protozoans.
Evidence seems to indicate that they have evolved
from the zooflagellates. Movement and food-getting
is accomplished with short hairlike structures called
cilia. Illustrations 9 through 24 are typical ciliates.
Phylum Apicomplexa
This phylum has only one class, the Sporozoa.
Members of this phylum lack locomotor organelles
and all are internal parasites. As indicated by their
class name, their life cycles include spore-forming
stages. Plasmodium, the malarial parasite, is a signif-
icant pathogenic sporozoan of humans.
SUBKINGDOM ALGAE
The Subkingdom Algae includes all the photosyn-
thetic eukaryotic organisms in Kingdom Protista.
Being true protists, they differ from the plants
(Plantae) in that tissue differentiation is lacking.
The algae may be unicellular, as those shown in the
top row of figure 6.2; colonial, like the four in the lower
right-hand corner of figure 6.2; or filamentous, as those
in figure 6.3. The undifferentiated algal structure is of-
ten referred to as a thallus. It lacks the stem, root, and
leaf structures that result from tissue specialization.
These microorganisms are universally present
where ample moisture, favorable temperature, and suf-
ficient sunlight exist. Although a great majority of them
live submerged in water, some grow on soil. Others
grow on the bark of trees or on the surfaces of rocks.
Algae have distinct, visible nuclei and chloro-
plasts. Chloroplasts are organelles that contain
chlorophyll a and other pigments. Photosynthesis
takes place within these bodies. The size, shape, dis-
tribution, and number of chloroplasts vary consider-
ably from species to species. In some instances a sin-
gle chloroplast may occupy most of the cell space.
Although there are seven divisions of algae,
only five will be listed here. Since two groups, the
cryptomonads and red algae, are not usually en-
countered in freshwater ponds, they have not been
included here.
Division 1 Euglenophycophyta
(Euglenoids)
Illustrations 1 through 6 in figure 6.2 are typical eu-
glenoids, representing four different genera within
this relatively small group. All of them are flagellated
and appear to be intermediate between the algae and
protozoa. Protozoanlike characteristics seen in the eu-
glenoids are (1) the absence of a cell wall, (2) the pres-
ence of a gullet, (3) the ability to ingest food but not
through the gullet, (4) the ability to assimilate organic
substances, and (5) the absence of chloroplasts in
some species. In view of these facts, it becomes read-
ily apparent why many zoologists often group the eu-
glenoids with the zooflagellates.
The absence of a cell wall makes these protists
very flexible in movement. Instead of a cell wall they
possess a semirigid outer pellicle, which gives the or-
ganism a definite form. Photosynthetic types contain
chlorophylls a and b, and they always have a red
stigma (eyespot), which is light sensitive. Their char-
acteristic food- storage compound is a lipopoly sac-
charide, paramylum. The photosynthetic eugle-
noids can be bleached experimentally by various
means in the laboratory. The colorless forms that de-
velop, however, cannot be induced to revert back to
phototrophy.
Division 2 Chlorophycophyta
(Green Algae)
The majority of algae observed in ponds belong to this
group. They are grass-green in color, resembling the
euglenoids in having chlorophylls a and b. They dif-
fer from euglenoids in that they sythesize starch in-
stead of paramylum for food storage.
The diversity of this group is too great to explore
its subdivisions in this preliminary study; however,
the small flagellated Chlamy domonas (illustration 8,
figure 6.2) appears to be the archetype of the entire
group and has been extensively studied. Many colo-
nial forms, such as Pandorina, Eudorina, Gonium,
and Volvox (illustrations 14, 15, 19, and 20, figure
6.2), consist of organisms similar to Chlamy domonas.
It is the general consensus that from this flagellated
form all the filamentous algae have evolved.
Except for Vaucheria and Tribonema, all of the fil-
amentous forms in figure 6.3 are Chlorophycophyta.
All of the nonfilamentous, nonflagellated algae in fig-
ure 6.4 also are green algae.
28
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
II. Survey of
Microorganisms
6. Protozoa, Algae, and
Cyanobacteria
© The McGraw-H
Companies, 2001
Protozoa, Algae, and Cyanobacteria • Exercise 6
Courtesy of the U.S. Environmental Protection Agency, Office of Research & Development, Cincinnati, Ohio 45268.
1 . Euglena (700X)
2. Euglena (700X)
3. Pftacus (1 000X)
4. Phacus (350X)
5. Lepocinclis (350X)
Figure 6.2 Flagellated algae
6. Trachelomonas (1 000X)
7. Phacofivs (1 500X)
8. Chlamydomonas (1 000X)
9. Carter/a (1500X)
1 0. Chlorogonium (1 000X)
1 1 . Pyrobotrys (1 000X)
12. Chrysococcus (3000X)
13. Synura (3 50X)
1 4. Pandorina (350X)
1 5. Eudorina (1 75X)
1 6. Dinobyron (1 000X)
17. Peridinium {350X)
1 8. Ceratium (1 75X)
1 9. Gonium (350X)
20. \/o/\/ox (1 00X)
29
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
II. Survey of
Microorganisms
6. Protozoa, Algae, and
Cyanobacteria
© The McGraw-H
Companies, 2001
Exercise 6 • Protozoa, Algae, and Cyanobacteria
A unique group of green algae is the desmids (il-
lustrations 16 through 20, figure 6.4). With the excep-
tions of a few species, the cells of desmids consist of
two similar halves, or semicells. The two halves usu-
ally are separated by a constriction, the isthmus.
Division 3 Chrysophycophyta
(Golden Brown Algae)
This large diversified division consists of over 6,000
species. They differ from the euglenoids and green al-
gae in that (1) food storage is in the form of oils and
leucosin, a polysaccharide; (2) chlorophylls a and c
are present; and (3) fucoxanthin, a brownish pig-
ment, is present. It is the combination of fucoxanthin,
other yellow pigments, and the chlorophylls that
causes most of these algae to appear golden brown.
Representatives of this division are seen in fig-
ures 6.2, 6.3, and 6.5. In figure 6.2, Chrysococcus,
Synura, and Dinobyron are typical flagellated chryso-
phycophytes. Vaucheria and Tribonema are the only
filamentous chrysophycophytes shown in figure 6.3.
All of the organisms in figure 6.5 are chrysophy-
cophytes and fall into a special category of algae
called the diatoms. The diatoms are unique in that
they have hard cell walls of pectin, cellulose, or sili-
con oxide that are constructed in two halves. The two
halves fit together like lid and box.
Skeletons of dead diatoms accumulate on the
ocean bottom to form diatomite, or "diatomaceous
earth," which is commercially available as an excel-
lent polishing compound. It is postulated by some that
much of our petroleum reserves may have been for-
mulated by the accumulation of oil from dead diatoms
over millions of years.
Division 4 Phaeophycophyta
(Brown Algae)
With the exception of three freshwater species, all al-
gal protists of this division exist in salt water (ma-
rine); thus, it is unlikely that you will encounter any
phaeophycophytes in this laboratory experience.
These algae have essentially the same pigments seen
in the chrysophycophytes, but they appear brown be-
cause of the masking effect of the greater amount of
fucoxanthin. Food storage in the brown algae is in the
form of laminarin, a polysaccharide, and mannitol,
a sugar alcohol. All species of brown algae are multi-
cellular and sessile. Most seaweeds are brown algae.
Division 5 Pyrrophycophyta
(Fire Algae)
The principal members of this division are the di-
noflagellates. Since the majority of these protists are
marine, only two freshwater forms are shown in fig-
ure 6.2: Peridinium and Ceratium (illustrations 17 and
18). Most of these protists possess cellulose walls of
interlocking armor plates, as in Ceratium. Two fla-
gella are present: one is directed backward when
swimming and the other moves within a transverse
groove. Many marine dinoflagellates are biolumines-
cent. Some species of marine Gymnodinium, when
present in large numbers, produce the red tides that
cause water discoloration and unpleasant odors along
our coastal shores.
These algae have chlorophylls a and c and sev-
eral xanthophylls. Foods are variously stored in the
form of starch, fats, and oils.
The Prokaryotes
As indicated on the first page of this unit, the prokary-
otes differ from the protists in that they are consider-
ably smaller, lack distinct nuclei with nuclear mem-
branes, and are enclosed in rigid cell walls. Since all
members of this group are bacteria, the three-domain
system of classification puts them in Domain
Bacteria.
Division Cyanobacteria
Division Cyanobacteria in Domain Bacteria includes
a large number of microorganisms that were at one
time referred to as the blue-green algae. All these
prokaryotes are photosynthetic, utilizing chlorophyll
a for photosynthesis. They differ from the green sul-
fur and green nonsulfur photosynthetic bacteria in that
the latter use bacteriochlorophyll instead of chloro-
phyll a for photosynthesis.
Over 1,000 species of cyanobacteria have been
reported. They are present in almost all moist envi-
ronments from the tropics to the poles, including both
freshwater and marine. Figure 6.6 illustrates only a
random few that are frequently seen.
The designation of these bacteria as "blue-green"
is somewhat misleading in that many cyanobacteria
are actually black, purple, red, and various shades of
green instead of blue- green. These different colors are
produced by the varying proportions of the numerous
pigments present. These pigments are chlorophyll a,
carotene, xanthophylls, blue c-phycocyanin, and
red c-phycoerythrin. The last two pigments are
unique to the cyanobacteria and red algae.
Cellular structure is considerably different from
the eukaryotic algae. Although cells lack visible nu-
clei, nuclear material is present in the form of DNA
granules in a colorless area in the center of the cell.
Unlike the algae, the pigments of the cyanobacte-
ria are not contained in chloroplasts; instead, they are
located in granules (phycobilisomes) that are at-
tached to membranes (thylakoids) that permeate the
cytoplasm.
30
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
II. Survey of
Microorganisms
6. Protozoa, Algae, and
Cyanobacteria
© The McGraw-H
Companies, 2001
Protozoa, Algae, and Cyanobacteria • Exercise 6
Courtesy of the U.S. Environmental Protection Agency, Office of Research & Development, Cincinnati, Ohio 45268.
1 . Rhizoclonium (1 75X)
2. Cladophora {WOX)
3. Bulbochaete (100X)
4. Oedogonium (350X)
5. Vaucheria (1 00X)
6. Tribonema (300X)
7. Chara (3X)
8. Batrachospermum (2X)
9. Microspora (175X)
10. Ulothrix {M5X)
1 1 . Ulothrix (1 75X)
12. Desmidium (175X)
Figure 6.3 Filamentous algae
13. Mougeof/a (1 75X)
1 4. Spirogyra (1 75X)
1 5. Zygnema (1 75X)
1 6. Stigeoclonium (300X)
17. Draparnaldia (100X)
31
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
II. Survey of
Microorganisms
6. Protozoa, Algae, and
Cyanobacteria
© The McGraw-H
Companies, 2001
Exercise 6 • Protozoa, Algae, and Cyanobacteria
K
V
^■■BVJ ■■■■
■
■T*\
■i ■■
JT
^^^t^_j^
■■i
fc F-mT
^^^F^m—
■■■liiiiiiiiiiiiHBBBB
^^kmn«
^""■■■■■■■■■W ™t 1
L
^^■■1 ■■
■a
\ 6
^^m^^^^H
MB
T
r
1 8
15
■
■ H
ft.
■Br ■1^'
^BB^l JB
^^Uj
:ff - §!»^_ „
1
i
■Hh
r
■ i
U-b
17 I
"l "
■■"■■1" ■■! 1 i
L^^Jb
%™r^» . b^^ ^b j
^iW* 2
Courtesy of the U.S. Environmental Protection Agency, Office of Research & Development, Cincinnati, Ohio 45268.
1 . Chlorococcum (700X)
2. Oocyst is (700X)
3. Coelastrum (350X)
4. Chlorella (350X)
5. Sphaerocystis (350X)
6. Micractinium (700X)
7. Scendesmus (700X)
8. Actinastrum (700X)
9. Phytoconis (700X)
1 0. Ankistrodesmus (700X)
1 1 . Pame//a (70 OX)
12. BofAyococcus (700X)
13. Tefraec/ron (1 000X)
1 4. Pediastrum (1 00X)
1 5. Tetraspora (1 00X)
16. Staurastrum (7 00X)
17. Staurast rum (3 50X)
18. Closterium {M5X)
1 9. Euastrum (3 5 OX)
20. Micrasterias (1 75X)
Figure 6.4 Nonfilamentous and nonflagellated algae
32
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
II. Survey of
Microorganisms
6. Protozoa, Algae, and
Cyanobacteria
© The McGraw-H
Companies, 2001
Protozoa, Algae, and Cyanobacteria • Exercise 6
Courtesy of the U.S. Environmental Protection Agency, Office of Research & Development, Cincinnati, Ohio 45268.
1 . Diatoma (1 000X)
2. Gomphonema (175X)
3. Cymbella (M5X)
4. Cymbella (1 000X)
5. Gomphonema (2000X)
6. Cocconeis (750X)
Figure 6.5 Diatoms
7. Nitschia (1 500X)
8. P/'nnu/ar/a (1 75X)
9. Cyclotella (1 000X)
1 0. Tabellaria (1 75X)
1 1 . Tabellaria (1 000X)
12. Synedra (350X)
13. Synec/ra (1 75X)
1 4. Melosira (750X)
1 5. SL7/77-e//a (350X)
16. SteL7/-one/'s (350X)
17. Frag/'//ar/a (750X)
1 8. Fragillaria (750X)
19./\ster/'0A7e//a(175X)
20. Asterionella (750X)
21./Vaw'cu/a(750X)
22. Stephanodiscus (750X)
23. Meridion (750X)
33
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
II. Survey of
Microorganisms
6. Protozoa, Algae, and
Cyanobacteria
© The McGraw-H
Companies, 2001
Exercise 6 • Protozoa, Algae, and Cyanobacteria
Courtesy of the U.S. Environmental Protection Agency, Office of Research & Development, Cincinnati, Ohio 45268.
1 . Anabaena (350X)
2. Anabaena (350X)
3. Anabaena (175X)
4. Nodularia (350X)
5. Cylindrospermum (1 75X)
6. Arthrospira (700X)
Figure 6.6 Cyanobacteria
7. Microcoleus (350X)
8. Phormidium (350X)
9. Oscillatoria (1 75X)
1 0. Aphanizomenon (1 75X)
1 1 . Lyngbya (700X)
12. Tolypothrix (350X)
13. Entophysalis (1000X)
1 4. Gomphosphaeria (1 000X)
1 5. Gomphosphaeria (350X)
1 6. Agmenellum (700X)
1 7. Agmenellum (1 75X)
18. Calot h rix {350X)
1 9. Rivularia (1 75X)
20. /\nacysf/'s (700X)
21./\A?acysf/s(175X)
22. /\A?acysf/'s (700X)
34
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
II. Survey of
Microorganisms
7. Microscopic
Invertebrates
© The McGraw-H
Companies, 2001
Microscopic Invertebrates
While looking for protozoa, algae, and cyanobacteria
in pond water, one invariably encounters large, trans-
parent, complex microorganisms that, to the inexperi-
enced, appear to be protozoans. In most instances
these moving "monsters" are rotifers (illustrations 13
through 17, figure 7.1); in some cases they are cope-
pods, daphnia, or any one of the other forms illus-
trated in figure 7.1.
All of the organisms illustrated in figure 7.1 are
multicellular with organ systems. If organ systems are
present, then the organisms cannot be protists, be-
cause organs indicate the presence of tissue differen-
tiation. Collectively, these microscopic forms are des-
ignated as "invertebrates." It is to prevent you from
misinterpreting some of these invertebrates as proto-
zoans that they are described here.
In using figure 7.1 to identify what you consider
might be an invertebrate, keep in mind that there are
considerable size differences. A few invertebrates,
such as Dugesia and Hydra, are macroscopic in adult
form but microscopic when immature. Be sure to
judge size differences by reading the scale beside
each organism. The following phyla are listed ac-
cording to the degree of complexity, the simplest
first.
Phylum Coelenterata
(Illustration 1)
Members of this phylum are almost exclusively ma-
rine. The only common freshwater form shown in
figure 7.1 is Hydra. In addition, there are a few less-
common freshwater genera similar to the marine
hydroids.
The hydras are quite common in ponds and lakes.
They are usually attached to rocks, twigs, or other
substrata. Around the mouth at the free end are five
tentacles of variable length, depending on the
species. Smaller organisms, such as Daphnia, are
grasped by the tentacles and conveyed to the mouth.
These animals have a digestive cavity that makes up
the bulk of the interior. Since no anus is present, undi-
gested remains of food are expelled through the
mouth.
Phylum Platyhelminthes
(Illustrations 2, 3, 4, 5)
The invertebrates of this phylum are commonly re-
ferred to as flatworms. The phylum contains two
parasitic classes and one class of free-living organ-
isms, the Turbellaria. It is the organisms in this
class that are encountered in fresh water. The four
genera of this class shown in figure 7.1 are Dugesia,
Planaria, Macrostomum, and Provortex. The char-
acteristics common to all these organisms are dorso-
ventral flatness, a ciliated epidermis, a ventral
mouth, and eyespots on the dorsal surface near the
anterior end. As in the coelenterates, undigested
food must be ejected through the mouth since no
anus is present. Reproduction may be asexual by fis-
sion or fragmentation; generally, however, repro-
duction is sexual, each organism having both male
and female reproductive organs. Species identifica-
tion of the turbellarians is exceedingly difficult and
is based to a great extent on the details of the repro-
ductive system.
Phylum Nematoda
(Illustration 6)
The members of this phylum are the roundworms.
They are commonly referred to as nemas or nema-
todes. They are characteristically round in cross sec-
tion, have an external cuticle without cilia, lack eyes,
and have a tubular digestive system complete with
mouth, intestine, and anus. The males are generally
much smaller than the females and have a hooked
posterior end. The number of named species is only
a fraction of the total nematodes in existence.
Species identification of these invertebrates requires
very detailed study of many minute anatomical fea-
tures, which requires complete knowledge of
anatomy.
Phylum Aschelminthes
This phylum includes classes Gastrotricha and
Rotifera. Most of the members of this phylum are
microscopic. Their proximity to the nematodes in
35
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
II. Survey of
Microorganisms
7. Microscopic
Invertebrates
© The McGraw-H
Companies, 2001
Exercise 7 • Microscopic Invertebrates
classification is due to the type of body cavity
(pseudocoel) that is present in both phyla.
The gastrotrichs (illustrations 7, 8, 9, 10) range
from 10 to 540 |xm in size. They are very similar to
the ciliated protozoans in size and habits. The typical
gastrotrich is elongate, flexible, forked at the posterior
end, and covered with bristles. The digestive system
consists of an anterior mouth surrounded by bristles,
a pharynx, intestine, and posterior anus. Species iden-
tification is based partially on the shape of the head,
tail structure and size, and distribution of spines.
Overall length is also an important identification char-
acteristic. They feed primarily on unicellular algae.
The rotifers (illustrations 13, 14, 15, 16, and 17)
are most easily differentiated by the wheellike
arrangement of cilia at the anterior end and the pres-
ence of a chewing pharynx within the body. They are
considerably diversified in food habits: some feed on
algae and protozoa, others on juices of plant cells, and
some are parasitic. They play an important role in
keeping waters clean. They also serve as food for
small worms and crustaceans, being an important link
in the food chain of fresh waters.
Phylum Annelida
(Illustration 18)
This phylum includes three classes: Oligochaeta,
Polychaeta, and Hirudinea. Since polychaetes are pri-
marily marine and the leeches (Hirudinea) are mostly
macroscopic and parasitic, only the oligochaete is
represented in figure 7.1. Some oligochaetes are ma-
rine, but the majority are found in fresh water and soil.
These worms are characterized by body segmenta-
tion, bristles {setae) on each segment, an anterior
mouth, and a roundish protrusion — the prostomium —
anterior to the mouth. Although most oligochaetes
breathe through the skin, some aquatic forms possess
gills at the posterior end or along the sides of the seg-
ments. Most oligochaetes feed on vegetation; some
feed on the muck of the bottoms of polluted waters,
aiding in purifying such places.
Phylum Tardigrada
(Illustrations 11 and 12)
These invertebrates are of uncertain taxonomic posi-
tion. They appear to be closely related to both the
Annelida and Arthropoda. They are commonly re-
ferred to as the water bears. They are generally no
more than 1 mm long, with a head, four trunk seg-
ments, and four pairs of legs. The ends of the legs may
have claws, fingers, or disklike structures. The ante-
rior end has a retractable snout with teeth. Eyes are of-
ten present. Sexes are separate, and females are
oviparous. They are primarily herbivorous. Loco-
motion is by crawling, not swimming. During desic-
cation of their habitat they contract to form barrel-
shaped tuns and are able to survive years of dryness,
even in extremes of heat and cold. Widespread distri-
bution is due to dispersal of the tuns by the wind.
Phylum Arthropoda
(Illustrations 19, 20, 21, 22, 23)
This phylum contains most of the known Animalia,
almost a million species. Representatives of three
groups of the Class Crustacea are shown in figure
7.1: Cladocera, Ostracoda, and Copepoda. The char-
acteristics these three have in common are jointed ap-
pendages, an exo skeleton, and gills.
The cladocera are represented by Daphnia and
Latonopsis in figure 7.1. They are commonly known
as water fleas. All cladocera have a distinct head.
The body is covered by a bivalvelike carapace. There
is often a distinct cervical notch between the head
and body. A compound eye may be present; when
present, it is movable. They have many appendages:
antennules, antennae, mouth parts, and four to six
pairs of legs.
The ostracods are bivalved crustaceans that are
distinguished from minute clams by the absence of
lines of growth on the shell. Their bodies are not dis-
tinctly segmented. They have seven pairs of ap-
pendages. The end of the body terminates with a pair
of caudal f urea.
The copepods represented here are Cyclops and
Canthocamptus. They lack the shell-like covering of
the ostracods and cladocera; instead, they exhibit dis-
tinct body segmentation. They may have three simple
eyes or a single median eye. Eggs are often seen at-
tached to the abdomen on females.
Laboratory Report
There is no Laboratory Report for this exercise
36
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
II. Survey of
Microorganisms
7. Microscopic
Invertebrates
© The McGraw-H
Companies, 2001
Microscopic Invertebrates • Exercise 7
■*rtiM*P^
1
-WJjrfrn
-o
20
**G
8
-itojut)
-o
14
■40/**,
-«
US
\*1JJ
V V'
V
^?
•~.2fHf»t.
-o
10
-Jfl^hT
-Mjrfm
-O
-O
16
17
1 I —
4>Q
21
22
-«»>**
■GINS
3ft- i ft
J"? v : £*■
-o
^/4ty*w
12
-Jfmm
-o
AtaMitapUfWWVnPiP
1 . Wyrfra
2. Dugesia
3. Planaria
4. Macrostomum
5. Pro vortex
6. Nematodes
7. L&pidermeJIa
8. 9 f 10. Chaetonotus
11, 12, Hypsibius
13, 14. Philodina
15, 16, Rotaria
17. Euchlanis
18. Oligochaete
19. Daphnia
20. iatonopsis
21. Ostracod
22. Cyclops
23. Canthocamptus
Figure 7.1 Microscopic invertebrates
37
Benson: Microbiological
II. Survey of
8. Aseptic Invertebrates
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Microorganisms
Companies, 2001
Aseptic Technique
Exercises 9 and 1 of this survey involve the use of
sterile media in culture tubes and Petri plates. The
proper handling of these materials requires special
skills that you must master. It is the purpose of this ex-
ercise to provide you with procedures that will be-
come routine as you progress through this course. It's
all about aseptic technique.
The procedures put forth in the section of this man-
ual entitled Laboratory Protocol outline some of the
specifics to be observed to ensure that you understand
what is required in maintaining an aseptic environment
when handling cultures of microorganisms. In this ex-
ercise you will have an opportunity to actually work
with cultures and different kinds of media to develop
those skills that are required to maintain asepsis.
Aseptic transfer of a culture from one culture ves-
sel to another is successful only if no contaminating
microorganisms are introduced in the process. A
transfer may involve the transport of organisms from
an isolated colony on a plate of solid medium to a
broth tube, or inoculating various media (solid or liq-
uid) from a broth culture for various types of tests.
The general procedure is as follows:
Work Area Disinfection. The work area is first
treated with a disinfectant to kill any microorganisms
that may be present. This step destroys vegetative
cells and viruses; endospores, however, are not de-
stroyed in this brief application of disinfectant.
Loops and Needles. The transport of organisms
will be performed with an inoculating loop or needle.
To sterilize the loop or needle prior to picking up the
organisms, heat must be applied with a Bunsen burner
flame, rendering them glowing red-hot.
Culture Tube Flaming. Before inserting the cooled
loop or needle into a tube of culture, the tube cap is re-
moved and the mouth of the culture tube flamed.
Once the organisms have been removed from the
tube, the tube mouth must be flamed again before re-
turning the cap to the tube.
Liquid Medium Inoculation. If a tube of liquid
medium is to be inoculated, the tube mouth must be
flamed before inserting the loop into the tube. To dis-
perse the organisms on the loop, the loop should be
twisted back and forth in the medium.
If an inoculating needle is used for stabbing a solid
medium, the needle is inserted deep into the medium.
Final Flaming. Once the inoculation is completed,
the loop or needle is removed from the tube, flamed as
before, and returned to a receptacle. These tools
should never be placed on the tabletop. The inoculated
tube is also flamed before placing the cap on the tube.
Petri Plate Inoculation. To inoculate a Petri plate,
no heat is applied to the plate and a loop is used for the
transfer. When streaking the surface of the medium,
the cover should be held diagonally over the plate bot-
tom to prevent air contamination of the medium.
Final Disinfection. When all work is finished, the
work area is treated with disinfectant to ensure that
any microorganisms deposited during any of the pro-
cedures are eliminated.
To gain some practice in aseptic transfer of bacte-
rial cultures, three simple transfers will be performed
here in this exercise: (1) broth culture to broth,
(2) agar slant culture to agar slant, and (3) agar plate
to agar slant. Proceed as follows:
Transfer from Broth Culture
to Another Broth
Do a broth tube to broth tube inoculation, using the
following technique. Figure 8.1 illustrates the proce-
dure for removing organisms from a culture, and fig-
ure 8.2 shows how to inoculate a tube of sterile broth.
Materials:
broth culture of Escherichia coli
tubes of sterile nutrient broth
inoculating loop
Bunsen burner
disinfectant for desktop and sponge
china marking pencil
1 . Prepare your desktop by swabbing down its sur-
face with a disinfectant. Use a sponge.
2. With a china marking pencil, label a tube of ster-
ile nutrient broth with your initials and E. coli.
3. Sterilize your inoculating loop by holding it over
the flame of a Bunsen burner until it becomes
bright red. The entire wire must be heated. See il-
lustration 1 , figure 8.1.
4. Using your free hand, gently shake the tube to dis-
perse the culture (illustration 2, figure 8.1).
39
Benson: Microbiological
II. Survey of
8. Aseptic Invertebrates
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Microorganisms
Companies, 2001
Exercise 8 • Aseptic Technique
5. Grasp the tube cap with the little finger of your
hand holding the inoculating loop and remove it
from the tube. Flame the mouth of the tube as
shown in illustration 3, figure 8.1.
6. Insert the inoculating loop into the culture (illus-
tration 4, figure 8.1).
7. Remove the loop containing the culture, flame the
mouth of the tube again (illustration 5, figure 8.1),
and recap the tube (illustration 6). Place the cul-
ture tube back on the test-tube rack.
8. Grasp a tube of sterile nutrient broth with your
free hand, carefully remove the cap with your lit-
tle finger, and flame the mouth of this tube (illus-
tration 1, figure 8.2).
9. Without flaming the loop, insert it into the ster-
ile broth, inoculating it (illustration 2, figure
8.2). To disperse the organisms into the
medium, move the loop back and forth in the
tube.
Inoculating loop is heated until
it is red -hot.
10. Remove the loop from the tube and flame the
mouth (illustration 3, figure 8.2). Replace the cap
on the tube (illustration 4, figure 8.2).
11. Sterilize the loop by flaming it (illustration 5, fig-
ure 8.2). Return the loop to its container.
12. Incubate the culture you just inoculated at 37° C
for 24-48 hours.
Transfer of Bacteria from
Slant to Slant
To inoculate a sterile nutrient agar slant from an agar
slant culture, use the following procedure. Figure 8.4
illustrates the entire process.
Materials:
agar slant culture of E. coll
sterile nutrient agar slant
inoculating loop
Bunsen burner
china marking pencil
***
Organisms in culture are dis-
persed by shaking tube.
Tube enclosure is removed and
mouth of tube is flamed.
^^ ^ ^ ta
A loopfulof organisms is removed
from tube.
Loop is removed from culture
and tube mouth is flamed.
Tube enclosure is returned to
tube.
Figure 8.1 Procedure for removing organisms from a broth culture with inoculating loop
Benson: Microbiological
II. Survey of
8. Aseptic Invertebrates
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Microorganisms
Companies, 2001
1
2
3
4.
5
If you have not already done so, prepare your
desktop by swabbing down its surface with a dis-
infectant.
With a china marking pencil label a tube of nutri-
ent agar slant with your initials and E. coll.
Sterilize your inoculating loop by holding it over the
flame of a Bunsen burner until it becomes bright
red (illustration 1, figure 8.4). The entire wire must
be heated. Allow the loop to cool completely.
Using your free hand, pick up the slant culture of
E. coli and remove the cap using the little finger
of the hand that is holding the loop (illustration 2,
figure 8.4).
Flame the mouth of the tube and insert the cooled
loop into the tube. Pick up some of the culture on
the loop (illustration 3, figure 8.4) and remove the
loop from the tube.
6
7
8
9
Aseptic Technique • Exercise 8
Flame the mouth of the tube (illustrations 4 and 5,
figure 8.4) and replace the cap, being careful not
to burn your hand. Return tube to rack.
Pick up a sterile nutrient agar slant with your free
hand, remove the cap with your little finger as be-
fore, and flame the mouth of the tube (illustration
6, figure 8.4).
Without flaming the loop containing the culture,
insert the loop into the tube and gently inoculate
the surface of the slant by moving the loop back
and forth over the agar surface, while moving
up the surface of the slant (illustration 7, figure
8.4). This should involve a type of serpentine
motion.
Remove the loop, flame the mouth of the tube,
and recap the tube (illustration 8, figure 8.4).
Replace the tube in the rack.
i iM
Cap is removed from sterile broth
and tube mouth is flamed.
rtta^i
Unheated loop is inserted into
tube of sterile broth.
i fl M'*'
Loop is removed from broth and
tube mouth is flamed.
Tube enclosure is returned to tube.
n«PWN^iWH«
#PW^
■taB"^r
Loop is flamed and returned to receptacle.
wm^^
Figure 8.2 Procedure for inoculating a nutrient broth
41
Benson: Microbiological
II. Survey of
8. Aseptic Invertebrates
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Microorganisms
Companies, 2001
Exercise 8 • Aseptic Technique
10. Flame the loop, heating the entire wire to red-hot
(illustration 9, figure 8.4), allow to cool, and place
the loop in its container.
11. Incubate the inoculated agar slant at 30° C for
24-48 hours.
Working with Agar Plates
(Inoculating a slant from a Petri plate)
The transfer of organisms from colonies on agar plates
to slants or broth tubes is very similar to the procedures
used in the last two transfers (broth to broth and slant
to slant). The following rules should be observed.
Loops vs. Needles In some cases a loop is used. In
other situations a needle is preferred. When a large in-
oculum is needed in the transfer, a loop will be used.
Needles are preferred, however, when making transfers
in pure culture isolations and making stab cultures. In
pure culture isolations, a needle is inserted into the cen-
ter of a colony for the transfer. This technique is used,
primarily, when working with mixed cultures.
Plate Handling Media in plates must always be
protected against contamination. To prevent exposure
to air contamination, covers should always be left
closed. When organisms are removed from a plate
culture, the cover should be only partially opened as
shown in illustration 2, figure 8.3.
Flaming Procedures Inoculating loops or needles
must be flamed in the same manner that you used when
working with previous tubes. One difference when
working with plates is that plates are never flamed !
Plate Labeling Petri plates with media in them are
always labeled on the bottom. Inoculated plates are
preferably stored upside down.
Pta^n
fVMPMtartWk
1 Inoculating loop is heated until it is
red-hot.
With free hand, raise the lid of the Petri plate just enough to
access a colony to pick up a loopful of organisms.
After flaming the mouth of a ster-
ile slant, streak its surface.
IN ■ *!■
Fiame the mouth of the tube
and re-cap the tube.
Flame the inoculating loop
and return it to receptacle.
^^^■^^^^■^■^^^^•^^•■••^
Figure 8.3 Procedure for inoculating a nutrient agar slant from an agar plate
Benson: Microbiological
II. Survey of
8. Aseptic Invertebrates
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Microorganisms
Companies, 2001
Inoculating loop is heated until
it is red-hot.
Mouth of tube is flamed. Inocula-
ting loop is not flamed.
*+-^wmrr*m
Cap is removed from slant cul
ture and tube mouth is heated.
Slant culture is re-capped and
retruned to test tube rack.
Organism is picked up from
slant with inoculating loop.
Tube of sterile agar slant is un
capped and mouth is flamed.
Slant surface is streaked with un-
earned loop in serpentine manner.
Tube mouth is flamed, recap-
ped and incubated.
mm
Loop is flamed red-hot and re
turned to receptacle,
Figure 8.4 Procedure for inoculating a nutrient agar slant from a slant culture
43
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
II. Survey of
Microorganisms
8. Aseptic Invertebrates
© The McGraw-H
Companies, 2001
Benson: Microbiological
II. Survey of
8. Aseptic Invertebrates
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Microorganisms
Companies, 2001
To transfer organisms from a Petri plate to an agar
slant, use the following procedure:
Materials:
nutrient agar plate with bacterial colonies
sterile nutrient agar slant
inoculating loop
china marking pencil
1 . If you have not done so, swab your work area with
disinfectant. Allow area to dry.
2. Label a sterile nutrient agar slant with your name
and organism to be transferred.
3. Flame an inoculating loop until it is red-hot (il-
lustration 1, figure 8.3). Allow the loop to cool.
4. As shown in illustration 2, figure 8.3, raise the lid
of a Petri plate sufficiently to access a colony with
your sterile loop.
Do not gouge into the agar with your loop as
you pick up organisms, and do not completely re-
move the lid, exposing the surface to the air. Close
the lid once you have picked up the organisms.
5
6
7
8
9
Aseptic Technique • Exercise 8
With your free hand, pick up the sterile nutrient
agar slant tube. Remove the cap by grasping the
cap with the little finger of the hand that is hold-
ing the loop.
Flame the mouth of the tube and insert the loop
into the tube to inoculate the surface of the
slant, using a serpentine motion (illustration 3,
figure 8.3). Avoid disrupting the agar surface
with the loop.
Remove the loop from the tube and flame the
mouth of the tube. Replace the cap on the tube (il-
lustration 4, figure 8.3).
Flame the loop (illustration 5, figure 8.3) and
place it in its container.
Incubate the nutrient agar slant at 37° C for 24-48
hours.
Second Period
Examine all three tubes and record your results on
Laboratory Report 8,9.
45
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
II. Survey of
Microorganisms
9. The Bacteria
© The McGraw-H
Companies, 2001
The Bacteria
Of all the microorganisms studied so far, the bacteria
are the most widely distributed, the simplest in mor-
phology, the smallest in size, the most difficult to clas-
sify, and the hardest to identify. It is even difficult to
provide a descriptive definition of what a bacterial or-
ganism is because of considerable diversity in the
group. About the only generalization that can be made
for the entire group is that they are prokaryotic and are
seldom photosynthetic. The few that are photosyn-
thetic utilize a pigment that is chemically different
from chlorophyll a. It is called b act erio chlorophyll.
Probably the simplest definition that one can con-
struct from these facts is: Bacteria are prokaryons
without chlorophyll a.
Although the bacteria are generally smaller than
the cyanobacteria, some cyanobacteria are in the size
range of bacteria. Most bacteria are only 0.5 to 2.0 mi-
crometers in diameter.
Figure 9.1 illustrates most of the shapes of bacte-
ria that one would encounter. Note that they can be
grouped into three types: rod, spherical, and helical or
curved. Rod- shaped bacteria may vary considerably
in length; have square, round, or pointed ends; and be
motile or nonmotile. The spherical, or coccus-shaped,
bacteria may occur singly, in pairs, in tetrads, in
chains, and in irregular masses. The helical and
curved bacteria exist as slender spirochaetes, spiril-
lum, and bent rods (vibrios).
In this exercise an attempt will be made to demon-
strate the ubiquitousness of these organisms. No at-
tempt will be made to study detailed bacterial
anatomy or physiology. Many exercises related to
staining, microscopy, and physiology in subsequent
laboratory periods will provide a clear understanding
of these microorganisms.
Our concern here relates primarily to the wide-
spread distribution of bacteria in our environment.
Being thoroughly aware of their existence all around
us is of prime importance if we are to develop those
laboratory skills that we recognize as aseptic tech-
nique. The awareness that bacteria are everywhere
must be constantly in our minds when handling bac-
terial cultures. In the next laboratory period you will
be handling tube cultures of bacteria, much in the
Coccobacillus
Fusiform
ft
Diplococci
and Tetrads
ROD (BACILLUS) -
/,
•
3-;.
Streptococci
Staphylococci
SPHERICAL (COCCI)
./
i
Comma
Spirillum
HELICAL AND CURVED
Spirochaetes
Figure 9.1 Bacterial morphology
same manner that you did in Exercise 8 (Aseptic
Technique). A continuing constant effort will always
be made to develop the proper routine. Remember:
without pure cultures the study of bacteriology be-
comes a hopeless endeavor.
During this laboratory period you will be pro-
vided with three kinds of sterile bacteriological me-
dia that you will expose to the environment in vari-
ous ways. To ensure that these exposures cover as
wide a spectrum as possible, specific assignments
will be made for each student. In some instances a
moistened swab will be used to remove bacteria from
some object; in other instances a Petri plate of
medium will be exposed to the air or a cough. You
will be issued a number that will enable you to deter-
mine your specific assignment from the chart on the
next page.
46
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
II. Survey of
Microorganisms
9. The Bacteria
© The McGraw-H
Companies, 2001
Materials:
per student:
1 tube of nutrient broth
1 Petri plate of trypticase soy agar (TS A)
1 sterile cotton swab
china marking pencil
per two or more students:
1 Petri plate of blood agar
1
2
3
4.
5
Scrub down your desktop with a disinfectant in
the same manner you used in Exercise 8 (Aseptic
Technique) .
Expose your TS A plate according to your assign-
ment in the table below. Label the bottom of your
plate with your initials, your assignment number,
and the date.
Moisten a sterile swab by immersing it into a tube
of nutrient broth and expressing most of the broth
out of it by pressing the swab against the inside
wall of the tube.
Rub the moistened swab over a part of your body
such as a finger or ear, or some object such as a
doorknob or telephone mouthpiece, and return the
swab to the tube of broth. It may be necessary to
break off the stick end of the swab so that you can
replace the cap on the tube.
Label the tube with your initials and the source of
the bacteria.
6
7
The Bacteria • Exercise 9
Expose the blood agar plate by coughing onto it.
Label the bottom of the plate with the initials of
the individuals that cough onto it. Be sure to date
the plate also.
Incubate the plates and tube at 37° C for 48 hours.
Evaluation
After 48 hours incubation, examine the tube of nutri-
ent broth and two plates. Shake the tube vigorously
without wetting the cap. Is it cloudy or clear?
Compare it with an uncontaminated tube of broth.
What is the significance of cloudiness? Do you see
any colonies growing on the blood agar plate? Are the
colonies all the same size and color? If not, what does
this indicate? Group together a set of TSA plates rep-
resenting all nine types of exposure. Record your re-
sults on the Laboratory Report.
Your instructor will indicate whether these tubes
and plates are to be used for making slides in Exercise
13 (Simple Staining). If the plates and tubes are to be
saved, containers will be provided for their storage in
the refrigerator. Place the plates and tubes in the des-
ignated containers.
Laboratory Report
Record your results on the last portion of Laboratory
Report 8,9.
Exposure Method for TSA Plate
Student Number
1 . To the air in laboratory for 30 minutes
1, 10, 19,28
2. To the air in room other than laboratory for 30 minutes
2, 11,20,29
3. To the air outside of building for 30 minutes
3, 12,21,30
4. Blow dust onto exposed medium
4, 13,22,31
5. Moist lips pressed against medium
5, 14,23,32
6. Fingertips pressed lightly on medium
6, 15,24,33
7. Several coins pressed temporarily on medium
7, 16,25,34
8. Hair is combed over exposed medium (10 strokes)
8, 17,26,35
9. Optional: Any method not listed above
9, 18,27,36
47
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
II. Survey of
Microorganisms
10. The Fungi: Yeasts and
Molds
© The McGraw-H
Companies, 2001
10
The Fungi:
Yeasts and Molds
The fungi comprise a large group of eukaryotic non-
photo synthetic organisms that include such diverse
forms as slime molds, water molds, mushrooms, puff-
balls, bracket fungi, yeasts, and molds. Fungi belong
to Kingdom Myceteae. The study of fungi is called
mycology.
Myceteae consist of three divisions:
Gymnomycota (slime molds), Mastigomycota (water
molds and others), and Amastigomycota (yeasts,
molds, bracket fungi, and others). It is the last division
that we will study in this exercise.
Fungi may be saprophytic or parasitic and unicel-
lular or filamentous. Some organisms, such as the
slime molds (Exercise 25), are borderline between
fungi and protozoa in that amoeboid characteristics
are present and fungi like spores are produced.
The distinguishing characteristics of the group
as a whole are that they (1) are eukaryotic, (2) are
nonphotosynthetic, (3) lack tissue differentiation,
(4) have cell walls of chitin or other polysaccha-
rides, and (5) propagate by spores (sexual and/or
asexual).
In this study we will examine prepared stained
slides and slides made from living cultures of yeasts
and molds. Molds that are normally present in the air
will be cultured and studied macro scopically and mi-
croscopically. In addition, an attempt will be made to
identify the various types that are cultured.
Before attempting to identify the various molds,
familiarize yourself with the basic differences be-
tween molds and yeasts. Note in figure 10.1 that
yeasts are essentially unicellular and molds are
multicellular.
Mold and Yeast Differences
Species within the Amastigomycota may have cot-
tony (moldlike) appearance or moist (yeasty) charac-
teristics that set them apart. As pronounced as these
differences are, we do not classify the various fungi in
this group on the basis of their being mold or yeast.
The reason that this type of division doesn't work is
that some species exist as molds under certain condi-
tions and as yeasts under other conditions. Such
species are said to be dimorphic, or triphasic.
The principal differences between molds and
yeasts are as follows:
Molds
Hyphae Molds have microscopic filaments called hy-
phae (hypha, singular). As shown in figure 10.1, if the
filament has crosswalls, it is referred to as having sep-
tate hyphae. If no crosswalls are present, the coeno-
cytic filament is said to be nonseptate, or aseptate.
Actually, most of the fungi that are classified as being
septate are incompletely septate since the septae have
central openings that allow the streaming of cytoplasm
from one compartment to the next. A mass of inter-
meshed hyphae, as seen macroscopically, is a mycelium.
Asexual Spores Two kinds of asexual spores are
seen in molds: sporangiospores and conidia. Spor-
Nonseptate
Septate
HYPHAE
Blastospore
PSEUDOHYPHA
MOLDS
YEASTS
V^i^PPV
m
Figure 10.1 Structural differences between molds and yeasts
48
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
II. Survey of
Microorganisms
10. The Fungi: Yeasts and
Molds
© The McGraw-H
Companies, 2001
angiospores are spores that form within a sac called a
sporangium. The sporangia are attached to stalks
called sporangiophores. See illustration 1, figure 10.2.
Conidia are asexual spores that form on special-
ized hyphae called conidiophores. If the conidia are
small they are called microconidia; large multicellu-
lar conidia are known as macro conidia. The following
four types of conidia are shown in figure 10.2:
• Phialospores: Conidia of this type are produced
by vase-shaped cells called phialides. Note in fig-
ure 10.2 that Penicillium and Gliocadium produce
this type.
• Blastoconidia: Conidia of this type are produced
by budding from cells of preexisting conidia, as in
Cladosporium, which typically has lemon-shaped
spores.
• Arthrospores: This type of conidia forms by sep-
aration from preexisting hyphal cells. Example:
Oospora.
• Chlamydospores: These spores are large, thick-
walled, round, or irregular structures formed
within or on the ends of a hypha. Common to
most fungi, they generally form on old cultures.
Example: Candida albicans.
Sexual Spores Three kinds of sexual spores are
seen in molds: zygospores, ascospores, and ba-
sidiospores. Figure 10.3 illustrates the three types.
The Fungi: Yeasts and Molds • Exercise 1
Zygospores are formed by the union of nuclear
material from the hyphae of two different strains.
Ascospores, on the other hand, are sexual spores pro-
duced in enclosures, which may be oval sacs or elon-
gated tubes. Basidiospores are sexually produced on
club-shaped bodies called basidia. A basidium is con-
sidered by some to be a modified type of ascus.
Yeasts
Hyphae Unlike molds, yeasts do not have true hy-
phae. Instead they form multicellular structures called
pseudohyphae. See figure 10.1.
Asexual Spores The only asexual spore produced
by yeasts is called a blastospore, or bud. These
spores form as an outpouching of a cell by a budding
process. It is easily differentiated from the parent cell
by its small size. It may separate from the original cell
or remain attached. If successive buds remain at-
tached in the budding process, the result is the forma-
tion of a pseudohypha.
Subdivisions of the
Amastigomycota
Division Amastigomycota consists of four subdivi-
sions: Zygomycotina, Ascomycotina, Basidiomycotina,
and Deuteromycotina. They are separated on the basis
of the type of sexual reproductive spores as follows :
Sporangia
Columella
Phialide
MUCOR
SYNCEPHALASTRUM
PENICILLIUM
GLIOCADIUM
1. SPORANGIQSPGRES
(Within Sporangia)
2. PHIALOSPORES
(Conidia on Phialides)
Conidiophore
CLADOSPORIUM
3. BLASTOCONIDIA
(Formed by Budding)
OOSPORA
CANDIDA ALBICANS
FUSARIUM
4. ARTHROSPORES 5. CHLAMYDOSPORES
(By Separation) (Large, Round)
ALTERNARIA
6. MACROCONIDIA
(Multicelled Conidia)
MICROSPORUM
CANIS
Figure 10.2 Types of asexual spores seen in fungi
49
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
II. Survey of
Microorganisms
10. The Fungi: Yeasts and
Molds
© The McGraw-H
Companies, 2001
a
Exercise 10 • The Fungi: Yeasts and Molds
Zygomycotina
These fungi have nonseptate hyphae and produce zy-
gospores. They also produce sporangiospores.
Rhizopus, Mucor, and Syncephalastrum are represen-
tative genera of this subdivision.
Ascomycotina
Since all the fungi in this subdivision produce as-
cospores, they are grouped into one class, the
Ascomycetes. They are commonly referred to as the
ascomycetes" and are also called "sac fungi." All of
them have septate hyphae and most of them have
chitinous walls.
Fungi in this group that produce a single ascus are
called ascomycetes yeasts. Other ascomycetes pro-
duce numerous asci in complex flask- shaped fruiting
bodies called perithecia or pseudothecia, in cup-
shaped structures, or in hollow spherical bodies, as in
powdery mildews, Eupenicillium or Talaromyces (the
sexual stages for Penicillium) .
Basidiomycotina
All fungi in this subdivision belong to one class, the
Basidiomycetes. Puffballs, mushrooms, smuts, rust, and
shelf fungi on tree branches are also basidiomycetes.
The sexual spores of this class are basidiospores.
Deuteromycotina
This fourth division of the Amastigomycota is an artifi-
cial group that was created to place any fungus that has
not been shown to have some means of sexual repro-
duction. Often, species that are relegated to this division
remain here for only a short period of time: as soon as
the right conditions have been provided for sexual
spores to form, they are reclassified into one of the first
three subdivisions. Sometimes, however, the asexual
and sexual stages of a fungus are discovered and named
separately by different mycologists, with the result that
a single species acquires two different names. Although
generally there is a switch over to the sexual-stage
name, not all mycologists conform to this practice.
Members of this group are commonly referred to
as the fungi imperfecti or deuteromycetes. It is a large
group, containing over 15,000 species.
Laboratory Procedures
Several options are provided here for the study of
molds and yeasts. The procedures to be followed will
be outlined by your instructor.
Yeast Study
The organism Saccharomyces cerevisiae, which is
used in bread making and alcohol fermentation, will
be used for this study. Either prepared slides or living
organisms may be used.
Materials:
prepared slides of Saccharomyces cerevisiae
broth cultures of Saccharomyces cerevisiae
methylene blue stain
microscope slides and cover glasses
Prepared Slides If prepared slides are used, they
may be examined under high-dry or oil immersion.
One should look for typical blastospores and as-
cospores. Space is provided on the Laboratory Report
for drawing the organisms.
Living Material If broth cultures of Saccharomyces
cerevisiae are available they should be examined on a
wet mount slide with phase-contrast or brightfield op-
tics. Two or three loopfuls of the organisms should be
placed on the slide with a drop of methylene blue stain.
Oil immersion will reveal the greatest amount of de-
tail. Look for the nucleus and vacuole. The nucleus is
the smaller body. Draw a few cells on the Laboratory
Report.
Mold Study
Examine a Petri plate of Sabouraud's agar that has
been exposed to the air for about an hour and incu-
bated at room temperature for 3-5 days. This medium
Asci
Zygospore
Basidium
ZYGOSPORE
(Zygomycotina)
ASCOSPORES
(Ascomycotina)
BASIDIOSPORES
(Basidiomycotina)
Figure 10.3 Types of sexual spores seen in the Amastigomycota
50
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
II. Survey of
Microorganisms
10. The Fungi: Yeasts and
Molds
© The McGraw-H
Companies, 2001
The Fungi: Yeasts and Molds • Exercise 1
Top
Reverse
1. ALTERNARIA
Top
Reverse
3. CUNNINGHAMELLA
Top
Reverse
5. HELMINTHOSPORIUM
Top
Reverse
2. ASPERGILLUS
Top
Reverse
4. FUSARIUM
Top
Reverse
6. PENICILLIUM
Top
Reverse
7. PAECILOMYCES
Top
Reverse
8. SYNCEPHALASTRUM
Figure 10.4 Colony characteristics of some of the more common molds
51
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
II. Survey of
Microorganisms
10. The Fungi: Yeasts and
Molds
© The McGraw-H
Companies, 2001
Exercise 10 • The Fungi: Yeasts and Molds
has a low pH, which makes it selective for molds. A
good plate will have many different-colored colonies.
Note the characteristic "cottony" nature of the colonies.
Also, look at the bottom of the plate and observe how
the colonies differ in color here. The identification of
molds is based on surface color, backside color, hyphal
structure, and types of spores.
Figure 10.4 reveals how some of the more com-
mon molds appear when grown on Sabouraud's agar.
Keep in mind when using figure 10.4 that the appear-
ance of a mold colony can change appreciably as it
gets older. The photographs in figure 10.4 are of
colonies that are 10 to 21 days old.
Conclusive identification cannot be made unless
a microscope slide is made to determine the type of
hyphae and spores that are present. Figure 10.5 re-
veals, diagrammatically, the microscopic differences
that one looks for when identifying mold genera.
Two Options In making slides from mold colonies,
one can make either wet mounts directly from the
colonies by the procedure outlined here or make cul-
tured slides as outlined in Exercise 26. The following
steps should be used for making stained slides directly
from the colonies. Your instructor will indicate the
number of identifications that are to be made.
Materials:
mold cultures on Sabouraud's agar
microscope slides and cover glasses
lactophenol cotton blue stain
sharp-pointed scalpels or dissecting needles
1 . Place the uncovered plate on the stage of your mi-
croscope and examine the edge of a colored
colony with the low-power objective. Look for
hyphal structure and spore arrangement. Ignore
the white colonies since they generally lack
spores and are difficult to identify.
CAUTION
Avoid leaving the cover off the mold culture plates
or disturbing the colonies very much. Dispersal of
mold spores to the air must be kept to a minimum.
2
3
4
Consult figures 10.4 and 10.5 to make a preliminary
identification based on colony characteristics and
low-power magnification of hyphae and spores.
Make a wet mount slide by transferring a small
amount of the culture with a sharp scalpel or dis-
secting needle to a drop of lactophenol cotton
blue stain on a slide. Cover with a cover glass and
examine under low-power and high-dry objec-
tives. Refer again to figure 10.5 to confirm any
conclusions drawn from your previous examina-
tion of the edge of the colony.
Repeat the above procedure for each different
colony.
Laboratory Report
After recording your results on the Laboratory
Report, answer all the questions.
Figure 10.5 Legend
1.
2.
3.
4.
5.
Penicillium — bluish-green; brush arrangement of
phialospores.
Aspergillus — bluish-green with sulfur-yellow areas
on the surface. Aspergillus niger is black.
Vertici Ilium — pinkish-brown, elliptical microconidia.
Trichoderma — green, resemble Penicillium macro-
scopically.
Gliocadium — dark green; conidia (phialospores)
borne on phialides, similar to Penicillium; grows
faster than Penicillium.
6. Cladosporium (Hormodendrum) — light green to gray-
ish surface; gray to black back surface; blastoconidia.
7. Pleospora — tan to green surface with brown to black
back; ascospores shown are produced in sacs
borne within brown, flask-shaped fruiting bodies
called pseudothecia.
8 Scopulariopsis — light brown; rough-walled microconidia.
9. Paecilomyces — yellowish-brown; elliptical microconidia.
10. Alternaria — dark greenish-black surface with gray
periphery; black on reverse side; chains of macroconidia.
11. Bipolaris — black surface with grayish periphery;
macroconidia shown.
12. Pullularia — black, shiny, leathery surface; thick-
walled; budding spores.
13. Diplosporium — buff-colored wooly surface; reverse
side has red center surrounded by brown.
14. Oospora (Geotrichum) — buff-colored surface;
hyphae break up into thin-walled rectangular
arthrospores.
15. Fusarium — variants of yellow, orange, red, and pur-
ple colonies; sickle-shaped macroconidia.
16. Trichothecium — white to pink surface; two-celled
conidia.
1 7. Mucor — a zygomycete; sporangia with a slimy tex-
ture; spores with dark pigment.
1 8. Rhizopus — a zygomycete; spores with dark pigment.
1 9. Syncephalastrum — a zygomycete; sporangiophores
bear rod-shaped sporangioles, each containing a
row of spherical spores.
20. Nigrospora — conidia black, globose, one-celled,
borne on a flattened, colorless vesicle at the end of
a conidiophore.
21. Montospora — dark gray center with light gray
periphery; yellow-brown conidia.
52
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
II. Survey of
Microorganisms
10. The Fungi: Yeasts and
Molds
© The McGraw-H
Companies, 2001
The Fungi: Yeasts and Molds • Exercise 1
Figure 10.5 Microscopic appearance of some of the more common molds (legend on opposite page)
53
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
. Microscope Slide
Techniques
Introduction
© The McGraw-H
Companies, 2001
Part
Microscope Slide Techniques
(Bacterial Morphology)
The nine exercises in this unit include the procedures for ten slide
techniques that one might employ in morphological studies of bac-
teria. A culture method in Exercise 18 also is included as a substi-
tute for slide techniques when pathogens are encountered.
These exercises are intended to serve two equally important
functions: (1) to help you to develop the necessary skills in making
slides and (2) to introduce you to the morphology of bacteria.
Although the title of each exercise pertains to a specific technique,
the organisms chosen for each method have been carefully se-
lected so that you can learn to recognize certain morphological fea-
tures. For example, in the exercise on simple staining (Exercise 1 3),
the organisms selected exhibit metachromatic granules, pleomor-
phism, and palisade arrangement of cells. In Exercise 15, (Gram
Staining) you will observe the differences between cocci and bacilli,
as well as learn how to execute the staining routine.
The importance of the mastery of these techniques cannot be
overemphasized. Although one is seldom able to make species
identification on the basis of morphological characteristics alone, it
is a very significant starting point. This fact will become increasingly
clear with subsequent experiments.
Although the steps in the various staining procedures may
seem relatively simple, student success is often quite unpre-
dictable. Unless your instructor suggests a variation in the proce-
dure, try to follow the procedures exactly as stated, without impro-
visation. Photomicrographs in color have been provided for many
of the techniques; use them as a guide to evaluate the slides you
have prepared. Once you have mastered a specific technique, feel
free to experiment.
55
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
. Microscope Slide
Techniques
11. Negative Staining
© The McGraw-H
Companies, 2001
li
Negative Staining
The simplest way to make a slide of bacteria is to pre-
pare a wet mount, much in the same manner that was
used for studying protozoa and algae. Although this
method will quickly produce a slide, finding the bac-
teria on the slide may be difficult, especially for a be-
ginner. The problem one encounters is that bacteria are
quite colorless and transparent. Unless the diaphragm
is carefully adjusted, the beginner usually has consid-
erable difficulty bringing the organisms into focus.
A better way to observe bacteria for the first time is
to prepare a slide by a process called negative, or back-
ground, staining. This method consists of mixing the
microorganisms in a small amount of nigrosine or india
ink and spreading the mixture over the surface of a slide.
(Incidentally, nigrosine is far superior to india ink.)
Since these two pigments are not really bacterial
stains, they do not penetrate the microorganisms.
Instead they obliterate the background, leaving the or-
ganisms transparent and visible in a darkened field.
Although this technique has limitations, it can be
useful for determining cell morphology and size. Since
no heat is applied to the slide, there is no shrinkage of
the cells, and, consequently, more accurate cell-size de-
terminations result than with some other methods. This
method is also useful for studying spirochaetes that
don't stain readily with ordinary dyes.
Three Methods
Negative staining can be done by one of three differ-
ent methods. Figure 11.1 illustrates the more com-
monly used method in which the organisms are mixed
in a drop of nigrosine and spread over the slide with
another slide. The goal is to produce a smear that is
thick at one end and feather-thin at the other end.
Somewhere between the too thick and too thin areas
will be an ideal spot to study the organisms.
Figure 11.2 illustrates a second method, in which or-
ganisms are mixed in only a loopful of nigrosine instead
Organisms are dispersed into a small drop of nigro-
sine or india ink. Drop should not exceed 1/8" diam-
eter and should be near one end of the slide.
Spreader slide is moved toward drop of suspension
until it contacts the drop causing the liquid to be spread
along its spreading edge.
Once the spreader slide contacts the drop on the
bottom slide, the suspension will spread out along the
spreading edge as shown.
Spreader slide is pushed to the left, dragging the sus-
pension over the bottom slide. After the slide has air-
dried, it may be examined under oil immersion.
Figure 11.1 Negative staining technique, using a spreader slide
56
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
. Microscope Slide
Techniques
11. Negative Staining
© The McGraw-H
Companies, 2001
of a full drop. In this method the organisms are spread
over a smaller area in the center of the slide with an inoc-
ulating needle. No spreader slide is used in this method.
The third procedure (Woeste-Demchick's method),
which is not illustrated here, involves applying ink to a
conventional smear with a black felt marking pen. If this
method is used, it should be done on a smear prepared
in the manner described in the next exercise. Simply
put, the technique involves applying a single coat of
felt-pen ink over a smear.
Note in the procedure below that slides may be
made from organisms between your teeth or from spe-
cific bacterial cultures. Your instructor will indicate
which method or methods you should use and demon-
strate some basic aseptic techniques. Various options
are provided here to ensure success.
Materials:
microscope slides (with polished edges)
nigrosine solution or india ink
slant cultures of S. aureus and B. megaterium
inoculating straight wire and loop
sterile toothpicks
Bunsen burner
china marking pencil
felt marking pen (see Instructor's Handbook)
1. Swab down your tabletop with disinfectant in
preparation for making slides.
2
3
4.
Negative Staining • Exercise 1 1
Clean two or three microscope slides with Bon
Ami to rid them of all dirt and grease.
By referring to figure 11.1 or 11.2, place the
proper amount of stain on the slide.
Oral Organisms: Remove a small amount of ma-
terial from between your teeth with a sterile straight
toothpick or inoculating needle and mix it into the
stain on the slide. Be sure to break up any clumps of
organisms with the wire or toothpick. When using a
wire, be sure to flame it first to make it sterile.
CAUTION
If you use a toothpick, discard it into a beaker of
disinfectant.
5. From Cultures: With a sterile straight wire,
transfer a very small amount of bacteria from the
slant to the center of the stain on the slide.
6. Spread the mixture over the slide according to the
procedure used in figure 11.1 or 11.2.
7. Allow the slide to air-dry and examine with an oil
immersion objective.
Laboratory Report
Draw a few representative types of organisms on
Laboratory Report 11-14. If slide is of oral organ-
isms, look for yeasts and hyphae as well as bacteria.
Spirochaetes may also be present.
A loopful of nigrosine or india ink is placed in the center
of a clean microscope slide.
A sterile inoculating wire is used to transfer the organ-
isms to the liquid and mix the organisms into the stain
Suspension of bacteria is spread evenly over an area
of one to two centimeters with the straight wire.
Once the preparation has completely air-dried, it can
be examined under oil immersion. No heat should be
used to hasten drying.
Figure 1 1.2 A second method for negative staining
57
Benson: Microbiological
III. Microscope Slide
12. Smear Preparation
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Techniques
Companies, 2001
12
Smear Preparation
While negative staining is a simple enough process to
make bacteria more visible with a brightfield micro-
scope, it is of little help when one attempts to observe
anatomical micro structures such as flagella, granules,
and endospores. Only by applying specific bacterio-
logical stains to organisms can such organelles be
seen. However, success at bacterial staining depends
first of all on the preparation of a suitable smear of the
organisms. A properly prepared bacterial smear is one
that withstands one or more washings during staining
without loss of organisms, is not too thick, and does
not result in excessive distortion due to cell shrinkage.
The procedure for making such a smear is illustrated
in figure 12.1.
The first step in preparing a bacteriological smear
differs according to the source of the organisms. If the
bacteria are growing in a liquid medium (broths, milk,
saliva, urine, etc.), one starts by placing one or two
loopfuls of the liquid medium directly on the slide.
From solid media such as nutrient agar, blood
agar, or some part of the body, one starts by placing
one or two loopfuls of water on the slide and then uses
a straight inoculating wire to disperse the organisms
in the water. Bacteria growing on solid media tend to
cling to each other and must be dispersed sufficiently
by dilution in water; unless this is done, the smear will
be too thick. The most difficult concept for students to
understand about making slides from solid media is
that it takes only a very small amount of material to
make a good smear. When your instructor demon-
strates this step, pay very careful attention to the
amount of material that is placed on the slide.
The organisms to be used for your first slides may
be from several different sources. If the plates from
Exercise 9 were saved, some slides may be made from
them. If they were discarded, the first slides may be
made for Exercise 13, which pertains to simple stain-
ing. Your instructor will indicate which cultures to use.
From Liquid Media
(Broths, saliva, milk, etc.)
If you are preparing a bacterial smear from liquid me-
dia, follow this routine, which is depicted on the left
side of figure 12.1.
Materials:
microscope slides
Bunsen burner
wire loop
china marking pencil
slide holder (clothespin), optional
1 . Wash a slide with soap or Bon Ami and hot water,
removing all dirt and grease. Handle the clean
slide by its edges.
2. Write the initials of the organism or organisms on
the left-hand side of the slide with a china mark-
ing pencil.
3. To provide a target on which to place the or-
ganisms, make a V" circle on the bottom side of
the slide, centrally located, with a marking
pencil. Later on, when you become more
skilled, you may wish to omit the use of this
"target circle."
4. Shake the culture vigorously and transfer two
loopfuls of organisms to the center of the slide
over the target circle. Follow the routine for inoc-
ulations shown in figure 12.2. Be sure to flame the
loop after it has touched the slide.
CAUTION
Be sure to cool the loop completely before insert-
ing it into a medium. A loop that is too hot will
spatter the medium and move bacteria into the air.
5. Spread the organisms over the area of the target
circle.
6. Allow the slide to dry by normal evaporation of
the water. Don't apply heat.
7. After the smear has become completely dry, pass
the slide over a Bunsen burner flame to heat- kill
the organisms and fix them to the slide.
Note that in this step one has the option of using
or not using a clothespin to hold the slide. Use the op-
tion preferred by your instructor.
58
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
. Microscope Slide
Techniques
12. Smear Preparation
© The McGraw-H
Companies, 2001
FROM LIQUID MEDIA
FROM SOLID MEDIA
"Target circle" on bottom of slide
Two loopfuls of water are placed in
center of "target circle."
Two loopfuls of liquid containing
organisms are placed in the center of
the "target circle."
Organisms are dispersed over entire
area of the "target circle."
A very small amount of organisms is
dispersed with inoculating needle in
water over entire area of "target
circle."
The smear is allowed to dry at room
temperature.
Slide is passed through flame several times to
heat-kill and fix organisms to slide. Use of
clothespin is optional.
Figure 12.1 Procedure for making a bacterial smear
59
Benson: Microbiological
III. Microscope Slide
12. Smear Preparation
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Techniques
Companies, 2001
Exercise 12 • Smear Preparation
From Solid Media
When preparing a bacterial smear from solid media,
such as nutrient agar or a part of the body, follow
this routine, which is depicted on the right side of
figure 12.1.
Materials:
microscope slides
inoculating needle and loop
china marking pencil
slide holder (clothespin), optional
Bunsen burner
1
2
3
4.
Wash a slide with soap or Bon Ami and hot water,
removing all dirt and grease. Handle the clean
slide by its edges.
Write the initials of the organism or organisms on
the left-hand side of the slide with a china mark-
ing pencil.
Mark a "target circle" on the bottom side of the
slide with a china marking pencil. (See comments
in step 3 on page 58.)
Flame an inoculating loop, let it cool, and transfer
two loopfuls of water to the center of the target
circle.
5. Flame an inoculating needle then let it cool. Pick
up a very small amount of the organisms, and mix
it into the water on the slide. Disperse the mixture
over the area of the target circle. Be certain that
the organisms have been well emulsified in the
liquid. Be sure to flame the inoculating needle be-
fore placing it aside.
6. Allow the slide to dry by normal evaporation of
the water. Don't apply heat.
7. Once the smear is completely dry, pass the slide
over the flame of a Bunsen burner to heat-kill the
organisms and fix them to the slide. Use a
clothespin to hold the slide if it is preferred by
your instructor. Some workers prefer to hold the
slide with their fingers so that they can monitor
the temperature of the slide (to prevent over-
heating).
Laboratory Report
Answer the questions on Laboratory Report 11-14
that relate to this exercise.
60
Benson: Microbiological
III. Microscope Slide
12. Smear Preparation
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Techniques
Companies, 2001
Smear Preparation • Exercise 1 2
Shake the culture tube from side to
side to suspend organisms. Do not
moisten cap on tube.
Heat the loop and wire to red-hot
Flame the handle slightly also.
After allowing the loop to cool for at
least 5 seconds, remove a loopfu
of organisms. Avoid touching the
sides of the tube.
Flame the mouth of the culture tube
again.
Remove the cap and flame the
neck of the tube. Do not place the
cap down on the table.
Return the cap to the tube and
place the tube in a test-tube rack,
Place the loopful of organisms in the center
of the target circle on the slide.
Flame the loop again before removing
another loopful from the culture or setting the
inoculating loop aside.
Figure 12.2 Aseptic procedure for organism removal
61
Benson: Microbiological
III. Microscope Slide
13. Simple Staining
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Techniques
Companies, 2001
13
Simple Staining
The use of a single stain to color a bacterial organism
is commonly referred to as simple staining. Some of
the most commonly used dyes for simple staining are
methylene blue, basic fuchsin, and crystal violet. All
of these dyes work well on bacteria because they have
color-bearing ions (chromophores) that are positively
charged (cationic).
The fact that bacteria are slightly negatively
charged produces a pronounced attraction between
these cationic chromophores and the organism. Such
dyes are classified as basic dyes. The basic dye meth-
ylene blue (methylene + chloride - ) will be used in this
exercise. Those dyes that have anionic chromophores
are called acidic dyes. Eosin (sodium -1- eosinate - ) is
such a dye. The anionic chromophore, eosinate - , will
not stain bacteria because of the electrostatic repelling
forces that are involved.
The staining times for most simple stains are rel-
atively short, usually from 30 seconds to 2 minutes,
depending on the affinity of the dye. After a smear has
been stained for the required time, it is washed off
gently, blotted dry, and examined directly under oil
immersion. Such a slide is useful in determining basic
morphology and the presence or absence of certain
kinds of granules.
An avirulent strain of Coryne bacterium diphtheriae
will be used here for simple staining. In its pathogenic
form, this organism is the cause of diphtheria, a very se-
rious disease. One of the steps in identifying this
pathogen is to do a simple stain of it to demonstrate the
following unique characteristics: pleomorphism, meta-
chromatic granules, and palisade arrangement of cells.
Pleomorphism pertains to irregularity of form:
i.e., demonstrating several different shapes. While C.
diphtheriae is basically rod-shaped, it also appears
club-shaped, spermlike, or needle-shaped. Bergey's
Manual uses the terms "pleomorphic" and "irregular"
interchangeably.
Metachromatic granules are distinct reddish-
purple granules within cells that show up when the or-
ganisms are stained with methylene blue. These gran-
ules are considered to be masses of volutin, a
polymetaphosphate.
Palisade arrangement pertains to parallel
arrangement of rod-shaped cells. This characteristic,
also called "picket fence" arrangement, is common to
many corynebacteria.
Procedure
Prepare a slide of C. diphtheriae, using the proce-
dure outlined in figure 13.1. It will be necessary to
refer back to Exercise 12 for the smear preparation
procedure.
Materials:
slant culture of avirulent strain of
Coryne bacterium diphtheriae
methylene blue (Loeffler's)
wash bottle
bibulous paper
After examining the slide, compare it with the pho-
tomicrograph in illustration 1, figure 15.3 (page 66).
Record your observations on Laboratory Report 11-14.
A bacterial smear is stained with
methylene blue for one minute.
Stain is briefly washed off slide
with water.
Water drops are carefully blotted
off slide with bibulous paper.
Figure 13.1 Procedure for simple staining
62
Benson: Microbiological
III. Microscope Slide
14. Capsular Staining
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Techniques
Companies, 2001
Capsular Staining
14
Some bacterial cells are surrounded by a pronounced
gelatinous or slimy layer called a capsule. There is con-
siderable evidence to support the view that all bacteria
have some amount of slime material surrounding their
cells. In most instances, however, the layer is not of suf-
ficient magnitude to be readily discernible. Although
some capsules appear to be made of glycoprotein, oth-
ers contain polypeptides. All appear to be water-soluble.
Staining the bacterial capsule cannot be accom-
plished by ordinary simple staining procedures. The
problem with trying to stain capsules is that if you pre-
pare a heat-fixed smear of the organism by ordinary
methods, you will destroy the capsule; and, if you do
not heat-fix the slide, the organism will slide off the
slide during washing. In most of our bacteriological
studies our principal concern is simply to demonstrate
the presence or absence of a pronounced capsule. This
can be easily achieved by combining negative and
simple staining techniques, as in figure 14.1. To learn
about this technique prepare a capsule "stained" slide
of Klebsiella pneumoniae, using the procedure out-
lined in figure 14.1.
Materials:
36-48 hour milk culture of Klebsiella
pneumoniae
india ink
crystal violet
Observation: Examine the slide under oil im-
mersion and compare your slide with illustration 2,
figure 15.3 on page 66. Record your results on
Laboratory Report 11-14.
Two loopfuls of the organism are
mixed in a small drop of india ink.
The ink suspension of bacteria is
spread over slide and air-dried.
Smear is stained with crystal violet
for one minute.
Crystal violet is gently washed off
with water.
The slide is gently heat-dried to fix
the organisms to the slide.
Slide is blotted dry with bibulous
paper, and examined with oil
immersion objective.
Figure 14.1 Procedure for demonstration of capsule presence
63
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
. Microscope Slide
Techniques
15. Gram Staining
© The McGraw-H
Companies, 2001
15
Gram Staining
In 1884 the Danish bacteriologist Christian Gram de-
veloped a staining technique that separates bacteria
into two groups: those that are gram-positive and
those that are gram-negative. The procedure is based
on the ability of microorganisms to retain the purple
color of crystal violet during decolorization with al-
cohol. Gram-negative bacteria are decolorized by the
alcohol, losing the purple color of crystal violet.
Gram-positive bacteria are not decolorized and re-
main purple. After decolorization, safranin, a red
counterstain, is used to impart a pink color to the de-
colorized gram-negative organisms.
Figure 15.1 illustrates the effects of the various
reagents on bacterial cells at each stage in the
process. Note that crystal violet, the primary stain,
causes both gram-positive and gram-negative organ-
isms to become purple after 20 seconds of staining.
When Gram's iodine, the mordant, is applied to the
cells for one minute, the color of gram-positive and
gram-negative bacteria remains the same: purple.
The function of the mordant here is to combine with
crystal violet to form a relatively insoluble com-
pound in the gram-positive bacteria. When the de-
colorizing agent, 95% ethanol, is added to the cells
for 10-20 seconds, the gram-negative bacteria are
leached colorless, but the gram-positive bacteria re-
main purple. In the final step a counterstain,
safranin, adds a pink color to the decolorized gram-
negative bacteria without affecting the color of the
purple gram-positive bacteria.
Of all the staining techniques you will use in the
identification of unknown bacteria, Gram staining is,
undoubtedly, the most important tool you will use.
Although this technique seems quite simple, perform-
ing it with a high degree of reliability is a goal that re-
quires some practice and experience. Here are two
suggestions that can be helpful: first, don't make your
smears too thick, and second, pay particular attention
to the comments in step 4 on the next page that pertain
to decolorization.
When working with unknowns keep in mind that
old cultures of gram-positive bacteria tend to decol-
orize more rapidly than young ones, causing them to
appear gram-negative instead of gram-positive. For
reliable results one should use cultures that are ap-
proximately 16 hours old. Another point to remem-
ber is that some species of Bacillus tend to be gram-
REAGENT
NONE
(Heat-fixed Cells)
CRYSTAL VIOLET
(20 seconds)
GRAM'S IODINE
(1 minute)
ETHYL ALCOHOL
(1 0-20 seconds)
SAFRANIN
(20 seconds)
GRAM-POS.
O
o
o
o
o
o
o
o
o
o
o
o
o
o
o
GRAM-NEG.
o
o
o
o
o
o
o
o
o
o
o
o
o
o
o
Figure 15.1 Color changes that occur at each step in
the gram-staining process
variable; i.e., sometimes positive and sometimes
negative.
During this laboratory period you will be pro-
vided an opportunity to stain several different kinds of
bacteria to see if you can achieve the degree of suc-
cess that is required. Remember, if you don't master
this technique now, you will have difficulty with your
unknowns later.
Staining Procedure
Materials:
slides with heat-fixed smears
gram-staining kit and wash bottle
bibulous paper
64
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
. Microscope Slide
Techniques
15. Gram Staining
© The McGraw-H
Companies, 2001
1
2
3
4
Cover the smear with crystal violet and let stand
for 20 seconds.
Briefly wash off the stain, using a wash bottle of
distilled water. Drain off excess water.
Cover the smear with Gram's iodine solution and
let it stand for one minute. (Your instructor may
prefer only 30 seconds for this step.)
Pour off the Gram's iodine and flood the smear
with 95% ethyl alcohol for 10 to 20 seconds.
This step is critical. Thick smears will require
more time than thin ones. Decolorization has oc-
curred when the solvent flows colorlessly from
the slide.
Gram Staining • Exercise 15
5 . Stop action of the alcohol by rinsing the slide with
water from wash bottle for a few seconds.
6. Cover the smear with safranin for 20 seconds.
(Some technicians prefer more time here.)
7. Wash gently for a few seconds, blot dry with
bibulous paper, and air-dry.
8. Examine the slide under oil immersion.
Assignments
The organisms that will be used here for Gram stain-
ing represent a diversity of form and staining charac-
teristics. Some of the rods and cocci are gram-positive;
CRYSTAL
VIOLET
20 seconds
wash 2 seconds
GRAM'S
IODINE
1 minute
DECOLORIZE 1 0-20 seconds or
WITH until solvent flows
ALCOHOL colorlessly
WASH
2 seconds
safranin 20 seconds
WASH
2 seconds
BLOT DRY
Figure 15.2 The gram-staining procedure
65
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
. Microscope Slide
Techniques
15. Gram Staining
© The McGraw-H
Companies, 2001
Exercise 15 • Gram Staining
others are gram-negative. One rod-shaped organism is
a spore-former and another is acid-fast. The challenge
here is to make gram-stained slides of various combi-
nations that reveal their differences.
Materials:
broth cultures of Staphylococcus aureus,
Pseudomonas aeruginosa, and Moraxella
(Branhamella) catarrhalis
nutrient agar slant cultures of Bacillus
megaterium and Mycobacterium smegmatis
Mixed Organisms 1 (Triple Smear Practice Slides)
Prepare three slides with three smears on each slide.
On the left portion of each slide make a smear of
Staphylococcus aureus. On the right portion of each
slide make a smear of Pseudomonas aeruginosa. In
the middle of the slide make a smear that is a mixture
of both organisms, using two loopfuls of each organ-
ism. Be sure to flame the loop sufficiently to avoid
contaminating cultures.
Gram stain one slide first, saving the other two for
later. Examine the center smear. If done properly, you
should see purple cocci and pink rods as shown in il-
lustration 3, figure 15.3.
Call your instructor over to evaluate your slide. If
the slide is improperly stained, the instructor will be
able to tell what went wrong by examining all three
smears. He or she will inform you how to correct
your technique when you stain the next triple smear
reserve slide.
Record your results on Laboratory Report 15-18
by drawing a few cells in the appropriate circle.
Mixed Organisms II Make a gram-stained slide of
a mixture of Bacillus megaterium and Moraxella
(Branhamella) catarrhalis.
This mixture differs from the previous slide in
that the rods (B. megaterium) will be purple and the
cocci (M.B. catarrhalis) will be large pink diplococci.
See illustration 4, figure 15.3.
As you examine this slide look for clear areas on
the rods, which represent endospores. Since en-
dospores are refractile and impermeable to crystal vi-
olet they will appear as transparent holes in the cells.
Draw a few cells in the appropriate circle on your
Laboratory Report sheet.
Acid-Fast Bacteria To see how acid-fast mycobac-
teria react to Gram's stain, make a gram-stained slide
of Mycobacterium smegmatis. If your staining tech-
nique is correct, the organisms should appear gram-
positive.
Draw a few cells in the appropriate circle on your
Laboratory Report sheet.
i
i
■fc
4
1. SIMPLE STAIN
Corynebacterium diphtheriae
I
2. CAPSULE STAIN
Klebsiella pneumoniae
**¥
•
3. GRAM STAIN
P. aeruginosa and S. aureus
4. GRAM STAIN
B. megaterium and M. B. catarrhalis
%
5. SPORE STAIN (Schaeffer-Fulton)
Bacillus megaterium
6. ACID-FAST STAIN (Ziehl-Neelsen)
M. smegmatis and S. aureus
Figure 15.3 Photomicrographs of representative staining techniques (8000x)
66
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
. Microscope Slide
Techniques
16. Spore Staining: Two
Methods
© The McGraw-H
Companies, 2001
Spore Staining:
Two Methods
16
Species of bacteria, belonging principally to the gen-
era Bacillus and Clostridium, produce extremely
heat-resistant structures called endospores. In addi-
tion to being heat-resistant, they are very resistant to
many chemicals that destroy non- spore- forming bac-
teria. This resistance to heat and chemicals is due pri-
marily to a thick, tough spore coat.
It was observed in Exercise 15 that Gram staining
will not stain endospores. Only if considerable heat is ap-
plied to a suitable stain can the stain penetrate the spore
coat. Once the stain has entered the spore, however, it is
not easily removed with decolorizing agents or water.
Several methods are available that employ heat to
provide stain penetration. However, since the
Schaeffer-Fulton and Dorner methods are the princi-
pal ones used by most bacteriologists, both have been
included in this exercise. Your instructor will indicate
which procedure is preferred in this laboratory.
SCHAEFFER-FULTON METHOD
This method, which is depicted in figure 16.1, utilizes
malachite green to stain the endospore and safranin to
stain the vegetative portion of the cell. Utilizing this
technique, a properly stained spore- former will have a
green endospore contained in a pink sporangium.
Illustration 5, figure 15.3 on page 66 reveals what
such a slide looks like under oil immersion.
After preparing a smear of Bacillus megaterium,
follow the steps outlined in figure 16.1 to stain the
spores.
Materials:
24-36 hour nutrient agar slant culture of
Bacillus megaterium
electric hot plate and small beaker (25 ml size)
spore- staining kit consisting of a bottle each of
5% malachite green and safranin
Cover smear with small piece of paper toweling and
saturate it with malachite green. Steam over boiling
water for 5 minutes. Add additional stain if stain boils off.
After the slide has cooled sufficiently, remove the
paper toweling and rinse with water for 30 seconds.
Counterstain with safranin for
about 20 seconds.
Rinse briefly with water to remove
safranin.
Blot dry with bibulous paper and
examine slide under oil immersion.
Figure 16.1 The Schaeffer-Fulton spore stain method
67
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
. Microscope Slide
Techniques
16. Spore Staining: Two
Methods
© The McGraw-H
Companies, 2001
Exercise 16 • Spore Staining: Two Methods
Dorner Method
The Dorner method for staining endospores produces
a red spore within a colorless sporangium. Nigrosine
is used to provide a dark background for contrast. The
six steps involved in this technique are shown in fig-
ure 16.2. Although both the sporangium and en-
dospore are stained during boiling in step 3, the spo-
rangium is decolorized by the diffusion of safranin
molecules into the nigrosine.
Prepare a slide of Bacillus megaterium that utilizes
the Dorner method. Follow the steps in figure 16.2.
Materials:
nigrosine
electric hot plate and small beaker (25 ml size)
small test tube (10 X 75 mm size)
test-tube holder
24-36 hour nutrient agar slant culture of
Bacillus megaterium
Laboratory Report
After examining the organisms under oil immersion,
draw a few cells in the appropriate circles on
Laboratory Report 15-18.
Make a heavy suspension of bacteria by dispersing
several loopfuls of bacteria in 5 drops of sterile water.
U
Add 5 drops of carbolfuchsin to the bacterial
suspension.
Heat the carbolfuchsin suspension of bacteria in
beaker of boiling water for 10 minutes.
Spread the nigrosine-bacteria mixture on the slide
in the same manner as in Exercise 11 (Negative
Staining).
Mix several loopfuls of bacteria in a drop of nigrosine
on the slide.
Allow the smear to air-dry. Examine the slide under oi
immersion.
Figure 16.2 The Dorner spore stain method
68
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
. Microscope Slide
Techniques
17. Acid-Fast Staining:
Ziehl-Neelsen Method
© The McGraw-H
Companies, 2001
Acid-Fast Staining:
Ziekl-Neelsen Method
17
Most bacteria in the genus Mycobacterium contain
considerable amounts of waxlike lipoidal material,
which affects their staining properties. Unlike most
other bacteria, once they are properly stained with
carbolfuchsin, they resist decolorization with acid-
alcohol. Since they are not easily decolorized they are
said to be acid-fast. This property sets them apart
from many other bacteria.
This stain is used primarily in the identification of
the tuberculosis bacillus, Mycobacterium tuberculo-
sis, and the leprosy organism, Mycobacterium leprae.
After decolorization, methylene blue is added to the
organisms to counterstain any material that is not
acid-fast; thus, a properly stained slide of a mixture of
acid-fast organisms, tissue cells, and non-acid-fast
bacteria will reveal red acid-fast rods with bluish tis-
sue cells and bacteria. An example of acid-fast stain-
ing is shown in illustration 6 of figure 15.3.
The two organisms used in this staining exercise
are Mycobacterium smegmatis, a nonpathogenic acid-
fast rod found in soil and on external genitalia, and
Staphylococcus aureus, a non-acid-fast coccus.
Materials:
nutrient agar slant culture of Mycobacterium
smegmatis (48-hour culture)
nutrient broth culture of S. aureus
electric hot plate and small beaker
acid- fast staining kit (carbolfuchsin, acid
alcohol, and methylene blue)
Smear Preparation Prepare a mixed culture smear
by placing two loopfuls of S. aureus on a slide and
transferring a small amount of M. smegmatis to the
broth on the slide with an inoculating needle. Since
the smegma bacilli are waxy and tend to cling to each
other in clumps, break up the masses of organisms
with the inoculating needle. After air-drying the
smear, heat- fix it.
Staining Follow the staining procedure outlined in
figure 17.1.
Examination Examine under oil immersion and
compare your slide with illustration 6, figure 15.3.
Laboratory Report Record your results on
Laboratory Report 15-18.
nu ll um ■■> I' m
Cover smear with carbolfuchsin
Steam over boiling water for 5
minutes. Add additional stain if
stain boils off.
After slide has cooled, decolorize
with acid-alcohol for 15-20
seconds.
Stop decolorization action of acid
alcohol by rinsing briefly with
water.
Counterstain with methylene blue
for 30 seconds.
Rinse briefly with water to remove
excess methylene blue.
Blot dry with bibulous paper.
Examine directly under oil
immersion.
Figure 17.1 Ziehl-Neelsen acid-fast staining procedure
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
. Microscope Slide
Techniques
18. Acid-Fast Staining
Fluorescence Method
© The McGraw-H
Companies, 2001
18
Acid-Fast Staining:
Fluorescence Method
In laboratories where large numbers of sputum, gas-
tric washings, urine, and other body fluid samples are
tested for pathogenic mycobacteria, fluorochrome
acid-fast staining is used in conjunction with the
Ziehl-Neelsen technique. The advantage of using a
fluorescence method is that fluorochrome- stained
slides can be scanned under lower magnification.
While a Ziehl-Neelsen prepared slide must be exam-
ined under oil immersion (1000X magnification),
fluorochrome-stained slides can be examined with
60 X or 100 X magnification. In only a few minutes an
entire fluorochrome prepared slide can be scanned.
Because of this fact, many laboratories use this faster
technique as a screening tool. When they encounter a
positive slide with this method, they use a Ziehl-
Neelsen prepared slide as a means of confirmation.
The fact that dead or noncultivatable mycobacteria
may fluoresce makes it necessary to use a confirma-
tory technique.
The Truant method of fluorochrome staining
(figure 18.1) consists of staining smears with au-
ramine-rhodamine for 20 minutes, decolorizing with
acid-alcohol, and "counterstaining" with potassium
permanganate. As soon as the slides are dry, they are
examined with a fluorescence microscope. Bacteria
that are acid- fast will fluoresce as yellow-orange rods
in a dark field. Areas of fluorescence that show up
during scanning can be examined more critically un-
der high-dry or oil immersion.
In this exercise you will stain a mixture of
Staphylococcus aureus and Mycobacterium phlei by the
Truant method. It will be examined with a fluorescence
microscope. If the slide is prepared properly, only the
acid-fast rod-shaped mycobacteria will fluoresce.
Materials:
broth culture of S. aureus
slant culture of M. phlei (Lowenstein- Jensen
medium)
auramine-rhodamine stain
acid-alcohol (for fluorochrome staining)
potassium permanganate (0.5% solution)
microscope slides, inoculating loop, Bunsen
burner, wash bottle
1
2
3
4
5
6
7
8
9
Prepare a mixed smear of S. aureus and M. phlei
by adding a small amount of M. phlei to two loop-
fuls of S. aureus on a clean slide. The organisms
should be well dispersed on the slide by vigorous
manipulation of the inoculating loop on the
clumps of organisms.
Allow the smear to air-dry completely.
Flame- fix the slide over a Bunsen burner. Avoid
overheating.
In diagnostic work where pathogens are be-
ing stained, the smear is usually heat-fixed on a
slide warmer (65° C) for 2 hours.
Cover the smear with auramine-rhodamine
stain and let stand for 20 minutes at room
temperature.
Rinse off the stain with wash bottle.
Decolorize with acid-alcohol (2.5% HC1 in 70%
ethanol) for 7-10 seconds.
Rinse thoroughly with wash bottle.
Cover the smear with potassium permanganate
and let stand for 3 minutes. This solution elimi-
nates background fluorescence ("quenching").
Although this step is often referred to as "coun-
terstaining," in actuality it is not. Excessive coun-
terstaining must be avoided because fluorescence
will be completely eliminated.
Rinse with water and air-dry.
Observation
The fluorescence microscope should be equipped
with a BG12 exciter filter and an OG1 barrier filter.
Scan the slide with the lowest magnification that is
possible on the microscope. On some instruments this
may be high-dry, not low power. If high-dry is used,
it is necessary to place a cover glass on the smear with
immersion oil between the slide and cover glass. Be
sure to use only oil that is specific for fluorescence
viewing.
Once you have located areas of fluorescence
(yellow-orange spots), add oil and swing the oil im-
mersion lens into position for more critical observa-
tion. Record your results on Laboratory Report 15-18.
■
70
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
. Microscope Slide
Techniques
18. Acid-Fast Staining:
Fluorescence Method
© The McGraw-H
Companies, 2001
Acid- Fast Staining: Fluorescence Method • Exercise 1 8
Cover a conventionally prepared smear with
auramine-rhodamine. Stain for 20 minutes.
Remove all stain by washing with water.
Decolorize the stained smear with acid-alcohol for
7-10 seconds.
Cover the smear with
potassium permanganate for
3 minutes.
Stop the decolorization process by rinsing off the
acid-alcohol with water.
Rinse off the potassium
permanganate with water.
Shake off water and allow slide to
air-dry. Do not use bibulous paper.
Figure 18.1 Fluorochrome acid-fast staining routine
71
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
. Microscope Slide
Techniques
19. Motility Determination
© The McGraw-H
Companies, 2001
19
Motility Determination
When attempting to identify an unknown bacterium it
is usually necessary to determine whether the mi-
croorganism is motile. Although one might think that
this determination would be easily arrived at, such is
not always the case. For the beginner there are many
opportunities to err.
Four Methods
For nonpathogens, there are two slide techniques that
one might use. For pathogens, one tube and one Petri
plate method can be used. Each method has its advan-
tages and limitations. The method you use will de-
pend on which one is most suitable for the situation at
hand. A discussion of each procedure follows.
The Wet Mount Slide
When working with nonpathogens, the simplest way to
determine motility is to place a few loopfuls of the or-
ganism on a clean slide and cover it with a cover glass.
In addition to being able to determine the presence or
absence of motility, this method is useful in determin-
ing cellular shape (rod, coccus, or spiral) and arrange-
ment (irregular clusters, packets, pairs, or long chains).
A wet mount is especially useful if phase optics are
used. Unlike stained slides that are heat-fixed for stain-
ing, there is no distortion of cells on a wet mount.
One problem for beginners is the difficulty of be-
ing able to see the organisms on the slide. Since bac-
teria are generally colorless and very transparent, the
novice has to learn how to bring them into focus.
The Hanging Drop Slide
If it is necessary to study viable organisms on a mi-
croscope slide for a longer period of time than is pos-
sible with a wet mount, one can resort to a hanging
drop slide. As shown in illustration 4 of figure 19.1,
organisms are observed in a drop that is suspended
under a cover glass in a concave depression slide.
Since the drop lies within an enclosed glass chamber,
drying out occurs very slowly.
Tube Method
When working with pathogenic microorganisms such as
the typhoid bacillus, it is too dangerous to attempt to de-
termine motility with slide techniques. A much safer
method is to culture the organisms in a special medium
that can demonstrate the presence of motility. The pro-
cedure is to inoculate a tube of semisolid or SIM medium
that can demonstrate the presence of motility. Both me-
dia have a very soft consistency that allows motile bac-
teria to migrate readily through them causing cloudiness.
Figure 19.2 illustrates the inoculation procedure.
Soft Agar Plate Method
Although the tube method is the generally accepted
procedure for determining motility of pathogens, it is
often very difficult for beginners to interpret. Richard
Roller at the University of Iowa suggests that incu-
bating a Petri plate of soft agar that has been stab in-
oculated with a motile organism will show up motil-
ity more clearly than an inoculated tube. This method
will also be tried here in this laboratory period.
First Period
During the first period you will make wet mount and
hanging drop slides of two organisms: Proteus vul-
garis and Micrococcus luteus. Tube media (semisolid
medium or SIM medium) and a soft agar plate will
also be inoculated. The media inoculations will have
to be incubated to be studied in the next period.
Proceed as follows:
Materials:
microscope slides and cover glasses
depression slide
2 tubes of semisolid or SIM medium
1 Petri plate of soft nutrient agar (20-25 ml of
soft agar per plate)
nutrient broth cultures of Micrococcus luteus
and Proteus vulgaris (young cultures)
inoculating loop and needle
Bunsen burner
Wet Mounts Prepare wet mount slides of each of
the organisms, using several loopfuls of the organism
on the slides. Examine under an oil immersion objec-
tive. Observe the following guidelines:
• Use only scratch-free, clean slides and cover
glasses. This is particularly important when using
phase-contrast optics.
72
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
. Microscope Slide
Techniques
19. Motility Determination
© The McGraw-H
Companies, 2001
..-
%
'
■
&
©
A small amount of Vaseline is placed near
each corner of the cover glass with a
toothpick.
Two loopfuls of organisms are placed in
center of cover glass.
Depression slide is pressed against Vaseline
on cover glass and quickly inverted.
Cover Glass
k\W\\WWW^
Vaseline
,W\W\WVXJ
Organisms
The completed preparation can be examined
under oil immersion.
Figure 19.1 The hanging drop slide
Motility Determination • Exercise 1 9
Label each slide with the name of the organism.
By manipulating the diaphragm and voltage con-
trol, reduce the lighting sufficiently to make the
organisms visible. Unstained bacteria are very
transparent and difficult to see.
For proof of true motility, look for directional
movement that is several times the long dimen-
sion of the bacterium. The movement will also oc-
cur in different directions in the same field.
Ignore Brownian movement. Brownian move-
ment is vibrational movement caused by invisible
molecules bombarding bacterial cells. If the only
movement you see is vibrational and not direc-
tional, the organism is nonmotile.
If you see only a few cells exhibiting motility, con-
sider the organism to be motile. Characteristically,
only a few of the cells will be motile at a given
moment.
Don't confuse water current movements with
true motility. Water currents are due to capillary
action caused by temperature changes and dry-
ing out. All objects move in a straight line in one
direction.
And, finally, always examine a wet mount imme-
diately, once it has been prepared, because motil-
ity decreases with time after preparation.
Hanging Drop Slides By referring to figure 19.1
prepare hanging drop slides of each organism. Be sure
to use clean cover glasses and label each slide with a
china marking pencil. When placing loopfuls of or-
ganisms on the cover glass, be sure to flame the loop
between applications. Once the slide is placed on the
microscope stage, do as follows:
1
2
3
4
Examine the slide first with the low-power objec-
tive. If your microscope is equipped with an auto-
matic stop, avoid using the stop; instead, use the
coarse adjustment knob for bringing the image
into focus. The greater thickness of the depression
slide prevents one from being able to focus at the
stop point.
Once the image is visible under low power, swing
the high-dry objective into position and readjust
the lighting. Since most bacteria are drawn to the
edge of the drop by surface tension, focus near
the edge of the drop.
If your microscope has phase-contrast optics,
switch to high- dry phase. Although a hanging
drop does not provide the shallow field desired
for phase-contrast, you may find that it works
fairly well.
If you wish to use oil immersion, simply rotate the
high-dry objective out of position, add immersion
oil to the cover glass, and swing the oil immersion
lens into position.
73
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
. Microscope Slide
Techniques
19. Motility Determination
© The McGraw-H
Companies, 2001
Exercise 19 • Motility Determination
5. Avoid delay in using this setup. Water of conden-
sation may develop to decrease clarity and the or-
ganisms become less motile with time.
6. Review all the characteristics of bacterial motility
that are stated on pages 72 and 73 under wet
mounts.
Tube Method Inoculate tubes of semisolid or SIM
media with each organism according to the following
instructions:
1
2.
3
4
5
Label the tubes of semisolid (or SIM) media with
the names of the organisms. Place your initials on
the tubes, also.
Flame and cool the inoculating needle, and insert it
into the culture after flaming the neck of the tube.
Remove the cap from the tube of medium, flame
the neck, and stab it % of the way down to the bot-
tom, as shown in figure 19.2. Flame the neck of the
tube again before returning the cap to the tube.
Repeat steps 2 and 3 for the other culture.
Incubate the tubes at room temperature for 24 to
48 hours.
Plate Method Mark the bottom of a plate of soft
agar with two one-half inch circles about one inch
apart. Label one circle ML and the other PV. These
circles will be targets for your culture stabs. Put your
initials on the plate also.
Using proper aseptic techniques, stab the medium in
the center of the ML circle with M. luteus and the cen-
ter of the other circle with P. vulgaris. Incubate the
plate for 24 to 48 hours at room temperature.
Second Period
Assemble the following materials that were inocu-
lated during the last period and incubated.
Materials:
culture tubes of motility medium that have been
incubated
inoculated Petri plate that has been incubated
Compare the two tubes that were inoculated with
M. luteus and P. vulgaris. Look for cloudiness as evi-
dence of motility. Proteus should exhibit motility.
Does it? Record your results on the Laboratory
Report.
Compare the appearance of the two stabs in the
soft agar. Describe the differences that exist in the two
stabs.
Does the plate method provide any better differ-
entiation of results than the tube method?
Laboratory Report
Complete the Laboratory Report for this exercise
Wire with organisms is brought into
tube without touching walls of tube.
Wire penetrates medium to two-thirds
of its depth.
Wire is withdrawn from medium and
tube. Neck of tube is flamed and
plugged.
Figure 19.2 Stab technique for motility test
74
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
IV. Culture Methods
Introduction
© The McGraw-H
Companies, 2001
Part
Cult
lire
Meth
oas
All nutritional types are represented among the protists. This diver-
sity requires a multiplicity of culture methods. An attempt has been
made in this unit to present those techniques that have proven
most successful for the culture of autotrophic and heterotrophic
bacteria, molds, and slime molds.
The first four exercises (20, 21, 22, and 23) pertain to basic
techniques applicable to both autotrophs and heterotrophs. The
other four exercises are concerned with the culture of specific
types. In performing the last four experiments, you should be just
as concerned with understanding the growth conditions as with the
successful growth of a particular isolate. For example, the use of
an enrichment medium, such as in Exercise 27 (Anaerobic
Phototrophic Bacteria), has direct application in the culture of many
other autotrophic bacteria as well.
This unit culminates the basic techniques phase of this course.
A thorough understanding of microscopy, slide techniques, and
culture methods provides a substantial foundation for the remain-
der of the exercises in this manual. If independent study projects
are to be pursued as a part of this course, the completion of this
unit will round out the background knowledge and skills for such
work.
75
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
IV. Culture Methods
20. Culture Media
Preparation
© The McGraw-H
Companies, 2001
20
Culture Media Preparation
From now on, most of the laboratory experiments in
this manual will utilize bacteriological media. In
most instances it will be provided for you. However,
circumstances may arise when you will need some
special medium that is not already prepared, and it
will be up to you to put it together. It is for situations
like this that the information in this exercise will be
useful.
The first portion of this exercise pertains to the
different types of media and how they relate to the
needs of microorganisms. The last part of the exercise
pertains to the actual mechanics of making up a batch
of medium. Whether you will be provided an oppor-
tunity to prepare some media during a designated lab-
oratory period will depend on the availability of time
and classroom needs. Your instructor will indicate
how this exercise is to be used.
Media Consistency
A microbiological medium (media, plural) is the food
that we use for culturing bacteria, molds, and other
microorganisms. It can exist in three consistencies:
liquid, solid, and semisolid. If you have performed all
of the exercises in Part 3, you are already familiar
with all of them.
Liquid media include nutrient broth, citrate
broth, glucose broth, litmus milk, etc. These media
are used for the propagation of large numbers of or-
ganisms, fermentation studies, and various other
tests.
Solid media are made by adding a solidifying
agent, such as agar, gelatin, or silica gel, to a liquid
medium. A good solidifying agent is one that is not
utilized by microorganisms, does not inhibit bacterial
growth, and does not liquefy at room temperature.
Agar and silica gel do not liquefy at room temperature
and are utilized by very few organisms. Gelatin, on
the other hand, is hydrolyzed by quite a few organ-
isms and liquefies at room temperature.
Nutrient agar, blood agar, and Sabouraud's agar
are examples of solid media that are used for devel-
oping surface colony growth of bacteria and molds.
As we will see in the next exercise, the develop-
ment of colonies on the surface of a medium is es-
sential when trying to isolate organisms from mixed
cultures.
Semisolid media fall in between liquid and solid
media. Although they are similar to solid media in that
they contain solidifying agents such as agar and
gelatin, they are more jelly like due to lower percent-
ages of these solidifiers. These media are particularly
useful in determining whether certain bacteria are
motile (Exercise 19).
Nutritional Needs of Bacteria
Before one can construct a medium that will
achieve a desired result in the growth of organisms,
one must understand their basic needs. Any medium
that is to be suitable for a specific group of organ-
isms must take into account the following seven
factors: water, carbon, energy, nitrogen, minerals,
growth factors, and pH. The role of each one of
these factors follows.
Water Protoplasm consists of 70% to 85% water.
The water in a single-celled organism is continuous
with the water of its environment, and the molecules
pass freely in and out of the cell, providing a vehicle
for nutrients inward and secretions or excretions out-
ward. All the enzymatically controlled chemical reac-
tions that occur within the cell occur only in the pres-
ence of an adequate amount of water.
The quality of water used in preparing media is
important. Hard tap water, high in calcium and mag-
nesium ions, should not be used. Insoluble phosphates
of calcium and magnesium may precipitate in the
presence of peptones and beef extract. The best policy
is to always use distilled water.
Carbon Organisms are divided into two groups
with respect to their sources of carbon. Those that can
utilize the carbon in carbon dioxide for synthesis of
all cell materials are called autotrophs. If they must
have one or more organic compounds for their carbon
source, they are called heterotrophs. In addition to or-
ganic sources of carbon, the heterotrophs are also de-
pendent on carbon dioxide. If this gas is completely
excluded from their environment, their growth is
greatly retarded, particularly in the early stages of
starting a culture.
Specific organic carbon needs are as diverse as
the organisms themselves. Where one organism may
76
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
IV. Culture Methods
20. Culture Media
Preparation
© The McGraw-H
Companies, 2001
require only a single simple compound such as acetic
acid, another may require a dozen or more organic nu-
trients of various degrees of complexity.
Energy Organisms that have pigments that enable
them to utilize solar energy are called photoautotrophs
(photosynthetic autotrophs). Media for such organisms
will not include components to provide energy.
Autotrophs that cannot utilize solar energy but are
able to oxidize simple inorganic substances for energy
are called chemoautotrophs (chemo synthetic au-
totrophs). The essential energy-yielding substance for
these organisms may be as elemental as nitrite, nitrate,
or sulfide.
Most bacteria fall into the category of chemo-
heterotrophs (chemosynthetic heterotrophs) that re-
quire an organic source of energy, such as glucose or
amino acids. The amounts of energy-yielding ingredi-
ents in media for both chemosynthetic types is on the
order of 0.5%.
A small number of bacteria are classified as pho-
toheterotrophs (photosynthetic heterotrophs). These
organisms have photosynthetic pigments that enable
them to utilize sunlight for energy. Their carbon
source must be an organic compound such as alcohol.
Nitrogen Although autotrophic organisms can uti-
lize inorganic sources of nitrogen, the heterotrophs
get their nitrogen from amino acids and intermediate
protein compounds such as peptides, proteoses, and
peptones. Beef extract and peptone, as used in nutri-
ent broth, provide the nitrogen needs for the het-
erotrophs grown in this medium.
Minerals All organisms require several metallic el-
ements such as sodium, potassium, calcium, magne-
sium, manganese, iron, zinc, copper, phosphorus, and
Culture Media Preparation • Exercise 20
cobalt for normal growth. Bacteria are no exception.
The amounts required are very small.
Growth Factors Any essential component of cell
material that an organism is unable to synthesize
from its basic carbon and nitrogen sources is classi-
fied as being a growth factor. This may include cer-
tain amino acids or vitamins. Many heterotrophs are
satisfied by the growth factors present in beef extract
of nutrient broth. Most fastidious pathogens require
enriched media such as blood agar for ample growth
factors.
Hydrogen Ion Concentration The growth of or-
ganisms in a particular medium may be completely
inhibited if the pH of the medium is not within cer-
tain limits. The enzymes of microorganisms are
greatly affected by this factor. Since most bacteria
grow best around pH 7 or slightly lower, the pH of
nutrient broth should be adjusted to pH 6.8.
Pathogens, on the other hand, usually prefer a more
alkaline pH. Trypticase soy broth, a suitable
medium for the more fastidious pathogens, should
be adjusted to pH 7.3.
Exact Composition Media
Media can be prepared to exact specifications so that
the exact composition is known. These media are gen-
erally made from chemical compounds that are highly
purified and precisely defined. Such media are read-
ily reproducible. They are known as synthetic media.
Media such as nutrient broth that contain ingredients
of imprecise composition are called nonsynthetic
media. Both the beef extract and peptone in nutrient
broth are inexact in composition.
Figure 20.1 Basic supplies and equipment needed for
making up a batch of medium.
Figure 20.2 Correct amount of dehydrated medium is
weighed on balance.
77
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
IV. Culture Methods
20. Culture Media
Preparation
© The McGraw-H
Companies, 2001
Exercise 20 • Culture Media Preparation
Special Media
Two kinds of special media that will be widely used in
this manual are selective and differential media.
Selective media are media that allow only certain
types of organisms to grow in or on them because of
(1) the absence of certain critical nutrients that make
it unfavorable for most, but not all, organisms, or
(2) the presence of inhibitory substances that prevent
certain types of organisms to grow on them. The in-
hibitory substance may be salt (NaCl), acid, a toxic
chemical (crystal violet), an antibiotic (streptomycin),
or some other substance.
Differential media are media that contain sub-
stances that cause some bacteria to take on a different
appearance from other species, allowing one to dif-
ferentiate one species from another.
In some cases media have been formulated that
are both selective and differential. A good example is
Levine EMB agar, which is used to determine the
presence of coliforms in water analysis (Exercise 63).
Dehydrated Media
Until around 1930, the laboratory worker had to spend
a good deal of time preparing laboratory media from
various raw materials. If a medium contained five or
six ingredients, it was not only necessary to measure
the various materials, but, also, in many instances, to
fabricate some of the components such as beef extract
or veal infusion by long tedious cooking methods.
Today, dehydrated media have revolutionized media
preparation techniques in much the same way that
commercial cake mixes have taken over in the
kitchen. For most routine bacteriological work, media
preparation has been simplified to the extent that all
that is necessary is to dissolve a measured amount of
dehydrated medium in water, adjust the pH, dispense
into tubes, and sterilize. In many cases pH adjustment
is not even necessary.
Media Preparation Assignment
In this laboratory period you will work with your lab-
oratory partner to prepare tubes of media that will be
used in future laboratory experiments. Your instructor
will indicate which media you are to prepare. Record
in the space below the number of tubes of specific me-
dia that have been assigned to you and your partner.
nutrient broth
nutrient agar pours
nutrient agar slants
other
Several different sizes of test tubes are used for
media, but the two sizes most generally used are either
16 mm or 20 mm diameter by 15 cm long. Select the
correct size tubes first, according to these guidelines:
Large tubes (20 mm dia): Use these test tubes
for all pours: i.e., nutrient agar, Sabouraud's
agar, EMB agar, etc. Pours are used for
filling Petri plates.
Small tubes (16 mm dia): Use these tubes for all
broths, deeps, and slants.
If the tubes are clean and have been protected
from dust or other contamination, they can be used
without cleaning. If they need cleaning, scrub out the
insides with warm water and detergent, using a test-
tube brush. Rinse twice, first with tap water, and fi-
nally with distilled water to rid them of all traces of
detergent. Place them in a wire basket or rack, in-
verted, so that they can drain. Do not dry with a towel.
Figure 20.3 Dehydrated medium is dissolved in a mea-
sured amount of distilled water.
Figure 20.4 If medium contains agar, it must be brought
to a boil to bring agar into solution.
78
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
IV. Culture Methods
20. Culture Media
Preparation
© The McGraw-H
Companies, 2001
Measurement and Mixing
The amount of medium you make for a batch should
be determined as precisely as possible to avoid short-
age or excess.
Materials:
graduate, beaker, glass stirring rod
bottles of dehydrated media
Bunsen burner and tripod, or hot plate
1. Measure the correct amount of water needed to
make up your batch. The following volumes re-
quired per tube must be taken into consideration:
pours 12 ml
deeps 6 ml
slants 4 ml
broths 5 ml
broths with fermentation tubes 5-7 ml
2
3
Consult the label on the bottle to determine how
much powder is needed for 1 ,000 ml and then de-
termine by proportionate methods how much you
need for the amount of water you are using.
Weigh this amount on a balance and add it to the
beaker of water. If the medium does not contain
agar, the mixture usually goes into solution with-
out heating.
If the medium contains agar, heat the mixture
over a Bunsen burner (figure 20.4) or on an elec-
tric hot plate until it comes to a boil. To safe-
guard against water loss, before heating, mark
the level of the top of the medium on the side of
the beaker with a china marking pencil. As soon
as it "froths up," turn off the heat. If an electric
hot plate is used, the medium must be removed
from the hot plate or it will boil over the sides of
the container.
Culture Media Preparation • Exercise 20
Caution: Be sure to keep stirring from the bottom
with a glass stirring rod so that the medium does
not char on the bottom of the beaker.
4. Check the level of the medium with the mark on the
beaker to note if any water has been lost. Add suffi-
cient distilled water as indicated. Keep the temper-
ature of the medium at about 60° C to avoid solidi-
fication. The medium will solidify at around 40° C.
Adjusting the pH
Although dehydrated media contain buffering agents
to keep the pH of the medium in a desired range, the
pH of a batch of medium may differ from that stated
on the label of the bottle. Before the medium is tubed,
therefore, one should check the pH and make any nec-
essary adjustments.
If a pH meter (figure 20.5) is available and al-
ready standardized, use it to check the pH of your
medium. If the medium needs adjustment use the bot-
tles of HC1 and NaOH to correct the pH. If no meter
is available pH papers will work about as well. Make
pH adjustment as follows:
Materials:
beaker of medium
acid and base kits (dropping bottles of 1 N and
0.1NHC1 and NaOH)
glass stirring rod
pH papers
pH meter (optional)
1 . Dip a piece of pH test paper into the medium to
determine the pH of the medium.
2. If the pH is too high, add a drop or two of HC1 to
lower the pH. For large batches use IN HC1. If the
pH difference is slight, use the 0.1N HC1. Use a
glass stirring rod to mix the solution as the drops
are added.
Figure 20.5 The hydrogen ion concentration of a
medium must be adjusted to its recommended pH.
Figure 20.6 An automatic pipetting machine will deliver
precise amounts of media at a controlled rate.
79
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
IV. Culture Methods
20. Culture Media
Preparation
© The McGraw-H
Companies, 2001
Exercise 20 • Culture Media Preparation
3. If the pH is too low, add NaOH, one drop at a
time, to raise the pH. For slight pH differences,
use 0.1N NaOH; for large differences use IN
NaOH. Use a glass stirring rod to mix the solution
as the drops are added.
Filling the Test Tubes
Once the pH of the medium is adjusted it must be dis-
pensed into test tubes. If an automatic pipetting ma-
chine is to be used, as shown in figure 20.6, it will have
to be set up for you by your instructor. These machines
can be adjusted to deliver any amount of medium at
any desired speed. When large numbers of tubes are to
be filled, the automatic pipetting machine should be
used. For smaller batches, the funnel method shown in
figure 20.7 is adequate. Use the following procedure
when filling tubes with a funnel assembly.
Materials:
ring stand assembly
funnel assembly (glass funnel, rubber tubing,
hose clamp, and glass tip)
graduate (small size)
1
2
3
4
Fill one test tube with a measured amount of
medium. This tube will be your guide for filling
the other tubes.
Fill the funnel and proceed to fill the test tubes to
the proper level, holding the guide tube alongside
of each empty tube to help you to determine the
amount to allow into each tube.
Keep the beaker of medium over heat if it con-
tains agar.
If fermentation tubes are to be used, add one to
each tube at this time with the open end down.
Capping the Tubes
The last step before sterilization is to provide a clo-
sure for each tube. Plastic (polypropylene) caps are
suitable in most cases. All caps that slip over the tube
end have inside ridges that grip the side of the tube
and provide an air gap to allow steam to escape dur-
ing sterilization. If you are using tubes with plastic
screw-caps, the caps should not be screwed tightly be-
fore sterilization; instead, each one must be left partly
unscrewed.
If no slip-on caps of the correct size are available,
it may be necessary to make up some cotton plugs. A
properly made cotton plug should hold firmly in the
tube so that it is not easily dislodged.
Sterilization
As soon as the tubes of media have been stoppered
they must be sterilized. Organisms on the walls of the
Figure 20.7 A glass funnel assembly and hose clamp
are adequate for filling small batches of tubes.
Figure 20.8 Once the medium has been dispensed to
all the tubes, they are capped prior to sterilization.
Figure 20.9 Tubes of media are sterilized in an auto
clave for 20 to 30 minutes at 1 5 psi steam pressure.
80
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
IV. Culture Methods
20. Culture Media
Preparation
© The McGraw-H
Companies, 2001
tubes, in the distilled water, and in the dehydrated
medium will begin to grow within a short period of
time at room temperature, destroying the medium.
Prior to sterilization, the tubes of media should be
placed in a wire basket with a label taped on the out-
side of the basket. The label should indicate the type
of medium, the date, and your name.
Sterilization must be done in an autoclave. The
following considerations are important in using an au-
toclave:
• Check with your instructor on the procedure to be
used with your particular type of autoclave.
Complete sterilization occurs at 250° F (121.6°
C). To achieve this temperature the autoclave has
to develop 15 pounds per square inch (psi) of
steam pressure. To reach the correct temperature
there must be some provision in the chamber for
the escape of air. On some of the older units it is
necessary to allow the steam to force air out
through the door before closing it.
• Don 't overload the chamber. One should not at-
tempt to see how much media can be packed into
it. Provide ample space between baskets of media
to allow for circulation of steam.
• Adjust the time of sterilization to the size of load.
Small loads may take only 10 to 15 minutes. An
Culture Media Preparation • Exercise 20
autoclave full of media may require 30 minutes
for complete sterilization.
After Sterilization
Slants If you have a basket of tubes that are to be
converted to slants, it is necessary to lay the tubes
down in a near-horizontal manner as soon as they are
removed from the autoclave. The easiest way to do this
is to use a piece of rubber tubing (1/2" dia) to support
the capped end of the tube as it rests on the countertop.
Solidification should occur in about 30-60 minutes.
Other Media Tubes of broth, agar deeps, nutrient
gelatin, etc., should be allowed to cool to room tem-
perature after removal from the autoclave. Once they
have cooled down, place them in a refrigerator or
cold-storage room.
Storage If tubes of media are not to be used imme-
diately, they should be stored in a cool place. When
stored for long periods of time at room temperature
media tend to lose moisture. At refrigerated tempera-
tures media will keep for months.
Laboratory Report
Complete the Laboratory Report for this exercise
81
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
IV. Culture Methods
21. Pure Culture
Techniques
© The McGraw-H
Companies, 2001
21
Pure Culture Techniques
When we try to study the bacterial flora of the body,
soil, water, food, or any other part of our environ-
ment, we soon discover that bacteria exist in mixed
populations. It is only in very rare situations that
they occur as a single species. To be able to study
the cultural, morphological, and physiological char-
acteristics of an individual species, it is essential,
first of all, that the organism be separated from the
other species that are normally found in its habitat;
in other words, we must have a pure culture of the
microorganism.
Several different methods of getting a pure cul-
ture from a mixed culture are available to us. The two
most frequently used methods involve making a
streak plate or a pour plate. Both plate techniques in-
volve thinning the organisms so that the individual
species can be selected from the others.
In this exercise you will have an opportunity to
use both methods in an attempt to separate three dis-
tinct species from a tube that contains a mixture. The
principal difference between the three organisms will
be their colors: Serratia marcescens is red,
Micrococcus luteus is yellow, and Escherichia coli is
white. If Chromobacterium violaceum is used in place
of M. luteus ; the three colors will be red, white, and
purple.
Streak Plate Method
For economy of materials and time, this method is
best. It requires a certain amount of skill, however,
which is forthcoming with experience. A properly ex-
ecuted streak plate will give as good an isolation as is
desired for most work. Figure 21.1 illustrates how
colonies of a mixed culture should be spread out on a
properly made streak plate. The important thing is to
produce good spacing between colonies.
Materials:
electric hot plate (or tripod and wire gauze)
Bunsen burner and beaker of water
wire loop, thermometer, and china marking
pencil
1 nutrient agar pour and 1 sterile Petri plate
1 mixed culture of Serratia marcescens,
Escherichia coli, and Micrococcus luteus
(or Chromobacterium violaceum)
Figure 21.1 If your streak plate reveals well-isolated
colonies of three colors (red, white, and yellow), you will
have a plate suitable for subculturing.
1
2
3
Prepare your tabletop by disinfecting its surface
with the disinfectant that is available in the labo-
ratory (Roccal, Zephiran, Betadine, etc.). Use a
sponge to scrub it clean.
Label the bottom surface of a sterile Petri plate
with your name and date. Use a china marking
pencil.
Liquefy a tube of nutrient agar, cool to 50° C, and
pour the medium into the bottom of the plate, fol-
lowing the procedure illustrated in figure 21.2. Be
sure to flame the neck of the tube prior to pouring
to destroy any bacteria around the end of the tube.
After pouring the medium into the plate, gen-
tly rotate the plate so that it becomes evenly dis-
tributed, but do not splash any medium up over
the sides.
Agar-agar, the solidifying agent in this
medium becomes liquid when boiled and resolid-
ifies at around 42° C. Failure to cool it prior to
pouring into the plate will result in condensation
of moisture on the cover. Any moisture on the
cover is undesirable because if it drops down on
the colonies, the organisms of one colony can
spread to other colonies, defeating the entire iso-
lation technique.
82
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
IV. Culture Methods
21. Pure Culture
Techniques
© The McGraw-H
Companies, 2001
4. Streak the plate by one of the methods shown in
figure 21.4. Your instructor will indicate which
technique you should use.
Caution: Be sure to follow the routine in figure 21.3
Pure Culture Techniques • Exercise 21
for getting the organism out of culture.
5. Incubate the plate in an inverted position at 25° C
for 24-48 hours. By incubating plates upside down,
the problem of moisture on the cover is minimized.
Liquefy a nutrient agar pour by boiling for 5
minutes.
Cool down the nutrient agar pour to 50° C by pouring
off some of the hot water and adding cold water to the
beaker. Hold at 50° C for 5 minutes.
Remove the cap from the tube and flame the
open end of the tube.
Pour the contents of the tube into the bottom of
the Petri plate and allow it to solidify.
Figure 21.2 Procedure for pouring an agar plate for streaking
83
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
IV. Culture Methods
21. Pure Culture
Techniques
© The McGraw-H
Companies, 2001
Exercise 21 • Pure Culture Techniques
Shake the culture tube from side to
side to suspend organisms. Do not
moisten cap on tube.
Heat the loop and wire to red-hot
Flame the handle slightly also.
Remove the cap and flame the neck
of the tube. Do not place the cap
down on the table.
After allowing the loop to cool for
at least 5 seconds, remove a loop-
ful of organisms. Avoid touching the
sides of the tube.
Flame the mouth of the culture tube
again.
Return the cap to the tube and
place the tube in a test-tube rack.
Streak the plate, holding it as shown. Do not
gouge into the medium with the loop.
Flame the loop before placing it down
Figure 21.3 Routine for inoculating a Petri plate
84
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
IV. Culture Methods
21. Pure Culture
Techniques
© The McGraw-H
Companies, 2001
Pure Culture Techniques • Exercise 21
QUADRANT STREAK
(Method A)
1 . Streak one loopful of organisms over Area 1 near edge of
the plate. Apply the loop lightly. Don't gouge into the medium.
2. Flame the loop, cool 5 seconds, and make 5 of 6 streaks
from Area 1 through Area 2. Momentarily touching the loop to
a sterile area of the medium before streaking insures a cool
loop.
3. Flame the loop again, cool it, and make 6 or 7 streaks
from Area 2 through Area 3.
4. Flame the loop again and make as many streaks as
possible from Area 3 into Area 4, using up the remainder
of the plate surface.
5. Flame the loop before putting it aside.
QUADRANT STREAK
(Method B)
1 . Streak one loopful of organisms back and forth over Area 1 ,
starting at point designated by "s". Apply loop lightly. Don't
gouge into the medium.
2. Flame the loop, cool 5 seconds and touch the medium in
sterile area momentarily to insure coolness.
3. Rotate the dish 90 degrees while keeping the dish closed.
Streak Area 2 with several back and forth strokes, hitting the
original streak a few times.
4. Flame the loop again. Rotate the dish and streak Area 3
several times, hitting last area several times.
5. Flame the loop, cool it, and rotate the dish 90 degrees again
Streak Area 4, contacting Area 3 several times and drag out the
culture as illustrated.
6. Flame the loop before putting it aside.
RADIANT STREAK
1 . Spread a loopful of organisms in small area near the edge
of the plate in Area 1 . Don't gouge medium.
2. Flame the loop and allow it to cool for 5 seconds. Touching
a sterile area of the medium will insure coolness.
3. From the edge of Area 1 make 7 or 8 straight streaks to the
opposite side of the plate.
4. Flame the loop again, cool it sufficiently, and cross
streak over the last streaks, starting near Area 1.
5. Flame the loop again before putting it aside.
CONTINUOUS STREAK
1 . Starting at the edge of the plate (Area A) with a loopful of
organisms, spread the organisms in a single continuous
movement to the center of the plate. Use light pressure and
avoid gouging the medium.
2. Rotate the plate 1 80 degrees so that the uninoculated
portion of the plate is away from you.
3. Without flaming loop, and using the same face of the loop,
continue streaking the other half of the plate by starting at Area B
and working toward the center.
4. Flame your loop before putting it aside.
Figure 21 .4 Four different streak techniques
85
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
IV. Culture Methods
21. Pure Culture
Techniques
© The McGraw-H
Companies, 2001
Exercise 21 • Pure Culture Techniques
Pour Plate Method
(Loop Dilution)
This method of separating one species of bacteria
from another consists of diluting out one loopful of
organisms with three tubes of liquefied nutrient agar
in such a manner that one of the plates poured will
have an optimum number of organisms to provide
good isolation. Figure 21 .5 illustrates the general pro-
cedure. One advantage of this method is that it re-
quires somewhat less skill than that required for a
good streak plate; a disadvantage, however, is that it
requires more media, tubes, and plates. Proceed as
follows to make three dilution pour plates, using the
same mixed culture you used for your streak plate.
Materials:
mixed culture of bacteria
3 nutrient agar pours
3 sterile Petri plates
electric hot plate
beaker of water
thermometer
inoculating loop and china marking pencil
1 . Label the three nutrient agar pours I, II, and III
with a marking pencil and place them in a beaker
2
3
4.
5
6
7
8
9
of water on an electric hot plate to be liquefied. To
save time, start with hot tap water if it is available.
While the tubes of media are being heated, label
the bottoms of the three Petri plates I, II, and III.
Cool down the tubes of media to 50° C, using
the same method that was used for the streak
plate.
Following the routine in figure 21.5, inoculate
tube I with one loopful of organisms from the
mixed culture. Note the sequence and manner of
handling the tubes in figure 21.6.
Inoculate tube II with one loopful from tube I af-
ter thoroughly mixing the organisms in tube I by
shaking the tube from side to side or by rolling the
tube vigorously between the palms of both hands.
Do not splash any of the medium up onto the
tube closure. Return tube I to the water bath.
Agitate tube II to completely disperse the organ-
isms and inoculate tube III with one loopful from
tube II. Return tube II to the water bath.
Agitate tube III, flame its neck, and pour its con-
tents into plate III.
Flame the necks of tubes I and II and pour their
contents into their respective plates.
After the medium has completely solidified, incu-
bate the inverted plates at 25° C for 24-48 hours.
«
•-»
*»».»'
,» »
»*". t
V\ ' '-
"i
f*-
\f ' "» f»
o &:
• v.
I
one hop fu/
MIXED
CULTURE
Figure 21.5 Three steps in the loop dilution technique for separating out organisms
86
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
IV. Culture Methods
21. Pure Culture
Techniques
© The McGraw-H
Companies, 2001
Pure Culture Techniques • Exercise 21
Liquefy three nutrient agar
pours, cool to 50° C, and let
stand for 10 minutes.
After shaking the culture to dis-
perse the organisms, flame the
loop and necks of the tubes.
Flame the loop and the necks of
both tubes.
Replace the caps on the tubes and
return the culture to the test-tube
rack.
Transfer one loopful from tube I to
tube II. Return tube I to the water
bath.
After shaking tube II and
transferring one loopful to tube
flame the necks of each tube
Transfer one loopful of the culture
to tube
Disperse the organisms in tube
by shaking the tube or rolling it
between the palms.
Pour the inoculated pours into
their respective Petri plates.
Figure 21.6 Tube-handling procedure in making inoculations for pour plates
87
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
IV. Culture Methods
21. Pure Culture
Techniques
© The McGraw-H
Companies, 2001
Exercise 21 • Pure Culture Techniques
Evaluation of the Two
Methods
After 24 to 48 hours of incubation examine all four
Petri plates. Look for colonies that are well isolated
from the others. Note how crowded the colonies ap-
pear on plate I as compared with plates II and III. Plate
I will be unusable. Either plate II or III will have the
most favorable isolation of colonies. Can you pick out
three well-isolated colonies on your best pour plate
that are white, yellow, and red?
Draw the appearance of your streak plate and
pour plates on the Laboratory Report.
SUBCULTURING TECHNIQUES
The next step in the development of a pure culture is
to transfer the organisms from the Petri plate to a tube
of nutrient broth or a slant of nutrient agar. After this
subculture has been incubated for 24 hours, a stained
slide of the culture can be made to determine if a pure
culture has been achieved. When transferring the or-
ganisms from the plate, an inoculating needle
(straight wire) is used instead of the wire loop. The
needle is inserted into the center of the colony where
there is a greater probability of getting only one
species of organism. Use the following routine in sub-
culturing out the three different organisms.
Materials:
3 nutrient agar slants
inoculating needle
Bunsen burner
1 . Label one tube S. marc esc ens, another E. coli, and
the third M. luteus or C. violaceum.
2. Select a well-isolated red colony on either the streak
plate or pour plate for your first transfer. Insert the
inoculating needle into the center of the colony.
3
4.
5
In the tube labeled S. marcescens, streak the slant
by placing the needle near the bottom of the slant
and drawing it up over its surface. One streak is
sufficient.
Repeat this inoculating procedure on the other
two slants for a white colony and a yellow (or pur-
ple) colony.
Incubate for 24 to 48 hours at 25° C.
Evaluation of Slants
After incubation, examine the slants. Is S. marcescens
red? Is E. coli white? Is your third slant yellow or pur-
ple? If the incubation temperature has been too high,
S. marcescens may appear white due to the fact that
the red pigment forms only at a moderate temperature,
such as 25° C. Draw the appearance of the slants with
colored pencils on the Laboratory Report.
Although the colors of the growths on the slants
may lead you to think that you have pure cultures, you
cannot be absolutely certain until you have made a
microscopic examination of each culture. For exam-
ple, it is entirely possible that the yellow slant (M. lu-
teus) may have some E. coli present that are masked
by the yellow pigment.
To find out if you have a pure culture on each
slant, make a gram-stained slide from each slant.
Knowing that S. marcescens, E. coli, and C. vio-
laceum are gram-negative rods, and that M. luteus is a
gram-positive coccus, you should be able to evaluate
your slants more precisely microscopically. Draw the
organisms on the Laboratory Report.
Laboratory Report
Complete the Laboratory Report for this exercise
88
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
IV. Culture Methods
22. Cultivation of
Anaerobes
© The McGraw-H
Companies, 2001
Cultivation of Anaerobes
The procedures for culturing bacteria that were used
in the last exercise work well only if the organisms
will grow in the presence of oxygen. Unfortunately,
there are many bacteria that find oxygen toxic or at
least inhibitory to their existence. For these organisms
we need to create an anaerobic environment by using
special media deficient or lacking in oxygen and con-
tainers that are oxygen-free. In this laboratory period
we will learn how to find out what the oxygen re-
quirements are for specific organisms and how to
grow them in liquid and solid media. In doing so we
will be inoculating special media with several organ-
isms of different cultural requirements to evaluate
their oxygen needs.
The oxygen requirements of bacteria range from
strict (obligate) aerobes that cannot exist without this
gas to the strict (obligate) anaerobes that die in its
presence. In between these extremes are the faculta-
tives, indifferents, and microaerophilics. The faculta-
tives are bacteria that have enzyme systems enabling
them to utilize free oxygen or some alternative oxy-
gen source such as nitrate. If oxygen is present, they
tend to utilize it in preference to the alternative. The
indifferents, however, show no preference for either
condition, growing equally well in aerobic and anaer-
obic conditions. Microaerophiles, on the other hand,
are organisms that require free oxygen, but only in
limited amounts. Figure 22.1 illustrates where these
various types tend to grow with respect to the degree
of oxygen tension in a medium.
In this experiment we will inoculate one liquid
medium and two solid media with several organ-
isms that have different oxygen requirements. The
media are fluid thiogly collate medium (FTM), tryp-
tone glucose yeast agar (TGYA), and Brewer's
anaerobic agar. Each medium will serve a different
purpose. A discussion of the function of each
medium follows:
TGYA Shake This solid medium will be used in
what is called a "shake tube." The medium is not pri-
marily an anaerobic medium; instead it is a rich gen-
eral purpose medium that favors the growth of a
broad spectrum of organisms. It will be inoculated in
the liquefied state, shaken to mix the organisms
throughout the medium, and allowed to solidify.
After incubation one determines the oxygen require-
HIGH
Aerobes
Microaerophiles
Oxygen
Tension
LOW
Facultatives
and
Indifferents
Strict Anaerobes
Figure 22.1 Oxygen needs of microorganisms
ments on the basis of where the growth occurs in the
tube: top, middle, or bottom.
FTM Fluid thioglycollate medium is a rich liquid
medium that supports the growth of both aerobic and
anaerobic bacteria. It contains glucose, cystine, and
sodium thioglycollate to reduce its oxidation-reduc-
tion (O/R) potential. It also contains the dye resazurin
that is an indicator for the presence of oxygen. In the
presence of oxygen the dye becomes pink. Since the
oxygen tension is always higher near the surface of
the medium, the medium will be pink at the top and
colorless in the middle and bottom. The medium also
contains a small amount of agar that helps to localize
the organisms and favors anaerobiasis in the bottom
of the tube.
Brewer's Anaerobic Agar This solid medium is an
excellent medium for culturing anaerobic bacteria in
Petri dishes. It contains thioglycollate as a reducing
agent and resazurin as an O/R indicator. For strict
anaerobic growth it is essential that plates be incu-
bated in an oxygen-free environment.
To provide an oxygen-free incubation environ-
ment for the Petri plates of anaerobic agar we will
■
89
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
IV. Culture Methods
22. Cultivation of
Anaerobes
© The McGraw-H
Companies, 2001
Exercise 22 • Cultivation of Anaerobes
use the GasPak anaerobic jar. Note in figure 22.2
that hydrogen is generated in the jar, which removes
the oxygen by forming water. Palladium pellets cat-
alyze the reaction at room temperature. The genera-
tion of hydrogen is achieved by adding water to a
plastic envelope of chemicals. Note also that C0 2 is
produced, which is a requirement for the growth of
many fastidious bacteria. To make certain that
anaerobic conditions actually exist in the jar, an in-
dicator strip of methylene blue becomes colorless in
the total absence of oxygen. If the strip is not re-
duced (decolorized) within 2 hours, the jar has not
been sealed properly, or the chemical reaction has
failed to occur.
In addition to doing a study of the oxygen re-
quirements of six organisms in this experiment, an op-
portunity will be provided during the second period to
do a microscopic study of the types of endospores
formed by three spore-formers used in the inocula-
tions. Proceed as follows:
First Period
(Inoculations and Incubation)
Since six microorganisms and three kinds of media
are involved in this experiment, it will be necessary
for economy of time and materials to have each stu-
dent work with only three organisms. The materials
list for this period indicates how the organisms will be
distributed.
During this period each student will inoculate
three tubes of medium and only one Petri plate of
Brewer's anaerobic agar. The tubes and all of the
plates will be placed in a GasPak jar to be incubated
in a 37° C incubator. Students will share results.
Materials:
per student:
3 tubes of fluid thioglycollate medium
3 TGYA shake tubes (liquefied)
1 Petri plate of Brewer's anaerobic agar
broth cultures for odd-numbered students:
Staphylococcus aureus, Streptococcus
faecalis, and Clostridium sporogenes
broth cultures for even-numbered students:
Bacillus subtilis, Escherichia coli, and
Clostridium rubrum
GasPak anaerobic j ar, 3 GasPak generator
envelopes, 1 GasPak anaerobic generator
strip, scissors, and one 10 ml pipette
water baths at student stations (electric hot plate,
beaker of water, and thermometer)
1 . Set up a 45° C water bath at your station in which
you can keep your tubes of TGYA shakes from so-
lidifying. One water bath for you and your labo-
ratory partner will suffice. (Note in the materials
Catalyst Chamber
Contains palladium pellets
Inner Lid
Gas Generator Envelope
10 ml of water is added to
chemicals in envelope to generate
H 2 and C0 2 . Carbon dioxide
promotes more rapid growth of
organisms.
Lock Screw
Outer Lid
Rubber Gasket
Provides air-tight seal
Reaction
Oxygen is removed from chamber
by combining with hydrogen on
surface of palladium pellets.
Anaerobic Indicator Strip
Methylene blue becomes colorless
in absence of 2 .
Figure 22.2 The GasPak anaerobic jar
90
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
IV. Culture Methods
22. Cultivation of
Anaerobes
© The McGraw-H
Companies, 2001
Cultivation of Anaerobes • Exercise 22
2
3
4.
5
6
7
list that the agar shakes have been liquefied for
you prior to lab time.)
Label the six tubes with the organisms assigned to
you (one organism per tube), your initials, and as-
signment number.
Note: Handle the tubes gently to avoid taking on
any unwanted oxygen into the media. If the tubes
of FTM are pink in the upper 30%, they must be
boiled a few minutes to drive off the oxygen, then
cooled to inoculate.
Heavily inoculate each of the TGYA shake tubes
with several loopfuls of the appropriate organism
for that tube. To get good dispersion of the organ-
isms in the medium, roll each tube gently between
the palms as shown in figure 22.3. To prevent
oxygen uptake do not overly agitate the medium.
Allow these tubes to solidify at room temperature.
Inoculate each of the FTM tubes with the appro-
priate organisms.
Streak your three organisms on the plate of anaer-
obic agar in the manner shown in figure 22.4.
Note that only three straight-line streaks, well
separated, are made. Place the Petri plate (in-
verted) in a cannister with the plates of other stu-
dents that is to go into the GasPak jar.
Once all the students' plates are in cannisters,
place the cannisters and tubes into the jar.
To activate and seal the GasPak jar, proceed as
follows:
a. Peel apart the foil at one end of a GasPak indi-
cator strip and pull it halfway down. The indi-
cator will turn blue on exposure to the air.
Place the indicator strip in the jar so that the
wick is visible.
b. Cut off the corner of each of three GasPak gas
generator envelopes with a pair of scissors.
Place them in the jar in an upright position.
8
9
c. Pipette 10 ml of tap or distilled water into the
open corner of each envelope. Avoid forcing
the pipette into the envelope.
d. Place the inner section of the lid on the jar,
making certain it is centered on top of the jar.
Do not use grease or other sealant on the rim
of the jar since the O-ring gasket provides an
effective seal when pressed down on a clean
surface.
e. Unscrew the thumbscrew of the outer lid until
the exposed end is completely withdrawn into
the threaded hole. Unless this is done, it will be
impossible to engage the lugs of the jar with
the outer lid.
f. Place the outer lid on the jar directly over the
inner lid and rotate the lid slightly to allow it to
drop in place. Now rotate the lid firmly to en-
gage the lugs. The lid may be rotated in either
direction.
g. Tighten the thumbscrew by turning clockwise.
If the outer lid raises up, the lugs are not prop-
erly engaged.
Place the jar in a 37° C incubator. After 2 or 3
hours check the jar to note if the indicator strip
has lost its blue color. If decolorization has not oc-
curred, replace the palladium pellets and repeat
the entire process.
Incubate the tubes and plates for 24 to 48 hours.
Second Period
(Culture Evaluations and Spore Staining)
Remove the lid from the GasPak jar. If vacuum holds
the inner lid firmly in place, break the vacuum by slid-
ing the lid to the edge. When transporting the plates
fc".". ; \y wyr%
Figure 22.3 Organisms are dispersed in medium by
rolling tube gently between palms.
Figure 22.4 Three organisms are streaked on agar plate
as straight-line streaks.
91
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
IV. Culture Methods
22. Cultivation of
Anaerobes
© The McGraw-H
Companies, 2001
Exercise 22 • Cultivation of Anaerobes
and tubes to your desk take care not to agitate the
FTM tubes. The position of growth in the medium
can be easily changed if handled carelessly.
Materials:
tubes of FTM
shake tubes of TGYA
2 Brewer's anaerobic agar plates
spore- staining kits and slides
1. Compare the six FTM and TGYA shake tubes
that you and your laboratory partner share with
figure 22.5 to evaluate the oxygen needs of the
six organisms.
2. Compare the growths (or lack of growth) on your
Petri plate and the plate of your laboratory partner.
3. Record your results on the Laboratory Report.
4. If time permits, make a combined slide with three
separate smears of the three spore-formers, using
either one of the two spore- staining methods in
Exercise 16. Draw the organisms in the circles
provided on the Laboratory Report.
Laboratory Report
Complete the Laboratory Report for this exercise
:/>
• •*
0*
V
\J
Aerobic
Microaerophilic
i tf i """"
HHn
t ■
y
:■>■!!
Facultative
Anaerobic
Figure 22.5 Growth patterns for different types of bacteria
92
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
IV. Culture Methods
23. Bacterial Population
Counts
© The McGraw-H
Companies, 2001
Bacterial Population Counts
23
Many bacteriological studies require that we be able
to determine the number of organisms that are present
in a given unit of volume. Several different methods
are available to us for such population counts. The
method one uses is determined by the purpose of the
study.
To get by with a minimum of equipment, it is pos-
sible to do a population count by diluting out the or-
ganisms and counting the organisms in a number of
microscopic fields on a slide. Direct examination of
milk samples with this technique can be performed
very quickly, and the results obtained are quite reli-
able. A technique similar to this can be performed on
a Petrof-Hauser counting chamber.
Bacterial counts of gas-forming bacteria can be
made by inoculating a series of tubes of lactose broth
and using statistical probability tables to estimate
bacterial numbers. This method, which we will use
in Exercise 63 to estimate numbers of coliform bac-
teria in water samples, is easy to use, works well in
water testing, but is limited to water, milk, and food
testing.
In this exercise we will use quantitative plating
(Standard Plate Count, or SPC) and turbidity mea-
surements to determine the number of bacteria in a
culture sample. Although the two methods are some-
what parallel in the results they yield, there are dis-
tinct differences. For one thing, the SPC reveals in-
formation only as related to viable organisms; that is,
colonies that are seen on the plates after incubation
represent only living organisms, not dead ones.
Turbidimetry results, on the other hand, reflect the
presence of all organisms in a culture, dead and living.
Quantitative Plating Method
(Standard Plate Count)
In determining the number of organisms present in
water, milk, and food, the standard plate count
(SPC) is universally used. It is relatively easy to per-
form and gives excellent results. We can also use this
basic technique to calculate the number of organisms
in a bacterial culture. It is in this respect that this as-
signment is set up.
The procedure consists of diluting the organisms
with a series of sterile water blanks as illustrated in
*^£s
C.....5
1 ml
-«4ta
CULTURE
-\
■■:■■■ /
$
1 ml
^
^
B
1 ml
1:100
v
%
a
1:1,000,000
1:100,000
1:10,000
1.0 ml
1:10,000
1:1,000,000
Figure 23.1 Quantitative plating procedure
figure 23.1. Generally, only three bottles are needed,
but more could be used if necessary. By using the di-
lution procedure indicated here, a final dilution of
1:1,000,000 occurs in blank C. From blanks B and C,
measured amounts of the diluted organisms are trans-
ferred into empty Petri plates. Nutrient agar, cooled to
50° C, is then poured into each plate. After the nutri-
ent agar has solidified, the plates are incubated for 24
to 48 hours and examined. A plate that has between 30
and 300 colonies is selected for counting. From the
count it is a simple matter to calculate the number of
organisms per milliliter of the original culture. It
should be pointed out that greater accuracy can be
achieved by pouring two plates for each dilution and
averaging the counts. Duplicate plating, however, has
been avoided for obvious economic reasons.
Pipette Handling
Success in this experiment depends considerably on
proper pipetting techniques. Pipettes may be available
to you in metal cannisters or in individual envelopes;
they may be disposable or reusable. In the distant past
pipetting by mouth was routine practice. However,
the hazards are obvious, and today it must be avoided.
Your instructor will indicate the techniques that will
■
93
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
IV. Culture Methods
23. Bacterial Population
Counts
© The McGraw-H
Companies, 2001
Exercise 23 • Bacterial Population Counts
prevail in this laboratory. If this is the first time that
you have used sterile pipettes, consult figure 23.2,
keeping the following points in mind:
• When removing a sterile pipette from a cannister, do
so without contaminating the ends of the other
pipettes with your fingers. This can be accomplished
by gently moving the cannister from side to side in
an attempt to isolate one pipette from the rest.
• After removing your pipette, replace the cover on
the cannister to maintain sterility of the remaining
pipettes.
• Don't touch the body of the pipette with your fin-
gers or lay the pipette down on the table before or
after you use it. Keep that pipette sterile until
you have used it, and don't contaminate the table
or yourself with it after you have used it.
Always use a mechanical pipetting device such as
the one in illustration 3, figure 23.2. For safety
reasons, deliveries by mouth are not acceptable in
this laboratory.
Remove and use only one pipette at a time; if you
need 3 pipettes for the whole experiment and re-
move all 3 of them at once, there is no way that
you will be able to keep 2 of them sterile while
you are using the first one.
When finished with a pipette, place it in the dis-
card cannister. The discard cannister will have a
Reusable pipettes may be available in disposable envelopes
or metal cannisters. When using pipettes from cannisters
be sure to cap them after removing a pipette.
Never touch the tip or barrel of a pipette with your
fingers. Contaminating the pipette will contaminate
your work.
^SCrttfft
Use a mechanical pipetter for all pipetting in this
laboratory. Pipetting by mouth is too hazardous.
After using a pipette place it in the discard cannister.
Even "disposable" pipettes must be placed here.
Figure 23.2 Pipette-handling techniques
94
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
IV. Culture Methods
23. Bacterial Population
Counts
© The McGraw-H
Companies, 2001
disinfectant in it. At the end of the period reusable
pipettes will be washed and sterilized by the lab-
oratory assistant. Disposable pipettes will be dis-
carded. Students have been known to absent-
mindedly return used pipettes to the original
sterile cannister, and, occasionally, even toss
them into the wastebasket. We are certain that no
one in this laboratory would ever do that!
Diluting and Plating Procedure
Proceed as follows to dilute out a culture of E. coli and
pour four plates, as illustrated in figure 23.1.
Materials:
per 4 students:
1 bottle (40 ml) broth culture of E. coli per
student:
1 bottle (80 ml) nutrient agar
4 Petri plates
1 . 1 ml pipettes
3 sterile 99 ml water blanks
cannister for discarded pipettes
1 . Liquefy a bottle of nutrient agar. While it is being
heated, label three 99 ml sterile water blanks A, B,
and C. Also, label the four Petri plates 1:10,000,
1:100,000, 1:1,000,000, and 1:10,000,000. In ad-
dition, indicate with labels the amount to be pipet-
ted into each plate (0.1 ml or 1.0 ml).
2. Shake the culture of E. coli and transfer 1 ml of
the organisms to blank A, using a sterile 1 . 1 ml
pipette. After using the pipette, place it in the dis-
card cannister.
3. Shake blank A 25 times in an arc of 1 foot for 7
seconds with your elbow on the table as shown in
Bacterial Population Counts • Exercise 23
figure 23.3. Forceful shaking not only brings
about good distribution, but it also breaks up
clumps of bacteria.
4. With a different 1.1 ml pipette, transfer 1 ml from
blank A to blank B .
5. Shake water blank B 25 times in same manner.
6. With another sterile pipette, transfer 0.1 ml from
blank B to the 1:100,000 plate and 1.0 ml to the
1 : 10,000 plate. With the same pipette, transfer 1 .0
ml to blank C.
7. Shake blank C 25 times.
8. With another sterile pipette, transfer from blank C
0.1 ml to the 1:10,000,000 plate and 1.0 ml to the
1:1,000,000 plate.
9. After the bottle of nutrient agar has boiled for 8
minutes, cool it down in a water bath at 50° C for
at least 10 minutes.
10. Pour one-fourth of the nutrient agar (20 ml) into
each of 4 plates. Rotate the plates gently to get ad-
equate mixing of medium and organisms. This
step is critical! Too little action will result in poor
dispersion and too much action may slop inocu-
lated medium over the edge.
11. After the medium has cooled completely, incu-
bate at 35° C for 48 hours, inverted.
Counting and Calculations
Materials:
4 culture plates
Quebec colony counter
mechanical hand counter
felt pen (optional)
Figure 23.3 Standard procedure for shaking water
blanks requires elbow to remain fixed on table
Figure 23.4 Colony counts are made on a Quebec
counter, using a mechanical hand tally
95
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
IV. Culture Methods
23. Bacterial Population
Counts
© The McGraw-H
Companies, 2001
Exercise 23 • Bacterial Population Counts
1 . Lay out the plates on the table in order of dilution
and compare them. Select the plates that have no
fewer than 30 nor more than 300 colonies for
your count. Plates with less than 30 or more than
300 colonies are statistically unreliable.
2. Place the plate on the Quebec colony counter
with the lid removed. See figure 23.4. Start
counting at the top of the plate, using the grid
lines to prevent counting the same colony twice.
Use a mechanical hand counter. Count every
colony, regardless of how small or insignificant.
Record counts on the table in section A of the
Laboratory Report.
Alternative Counting Method: Another
way to do the count is to remove the lid and place
the plate upside down on the colony counter.
Instead of using the grid to keep track, use a felt
pen to mark off each colony as you do the count.
3. Calculate the number of bacteria per ml of undi-
luted culture using the data recorded in section A
of the Laboratory Report. Multiply the number of
colonies counted by the dilution factor (the recip-
rocal of the dilution).
Example: If you counted 220 colonies on the
plate that received 1.0 ml of the 1:1,000,000 di-
lution: 220 X 1,000,000 (or 2.2 X 10 8 ) bacteria
per ml. If 220 colonies were counted on the plate
that received 0.1 ml of the 1:1,000,000 dilution,
then the above results would be multiplied by 1
to convert from number of bacteria per 0.1 ml to
number of bacteria per 1 .0 ml (2,200,000,000, or
2.2 X 10 9 ).
Use only two significant figures. If the num-
ber of bacteria per ml was calculated to be
227,000,000, it should be recorded as
230,000,000, or 2.3 X 10 8 .
TURBIDIMETRY DETERMINATIONS
When it is necessary to make bacteriological counts
on large numbers of cultures, the quantitative plate
count method becomes a rather cumbersome tool. It
not only takes a considerable amount of glassware
and media, but it is also time-consuming. A much
faster method is to measure the turbidity of the culture
with a spectrophotometer and translate this into the
number of organisms. To accomplish this, however,
the plate count must be used to establish the count for
one culture of known turbidity.
To understand how a spectrophotometer works, it
is necessary, first, to recognize the fact that a culture
of bacteria acts as a colloidal suspension, which will
intercept the light as it passes through. Within certain
limits the amount of light that is absorbed is directly
proportional to the concentration of cells.
Figure 23.5 illustrates the path of light through a
spectrophotometer. Note that a beam of white light
passes through two lenses and an entrance slit into a
diffraction grating that disperses the light into hori-
zontal beams of all colors of the spectrum. Short
wavelengths (violet and ultraviolet) are at one end
and long wavelengths (red and infrared) are at the
other end. The spectrum of light falls on a dark
screen with a slit (exit slit) cut in it. Only that por-
tmmrmmmm
Lamp
Galvanometer
Percent Transmittance
4-0 5© fc
!■ ll'll
UU1U *
/
5*
Phototube
Sample
Figure 23.5 Schematic of a spectrophotometer
Entrance
Slit
Diffraction
Grating
Light
Control
^iMMMMMMI HUM 1441— HWWWWWP
96
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
IV. Culture Methods
23. Bacterial Population
Counts
© The McGraw-H
Companies, 2001
Bacterial Population Counts • Exercise 23
tion of the spectrum that happens to fall on the slit
goes through into the sample. It will be a monochro-
matic beam of light. By turning a wavelength control
knob on the instrument, the diffraction grating can
be reoriented to allow different wavelengths to pass
through the slit. The light that passes through the
culture activates a phototube, which, in turn, regis-
ters percent transmittance ( % T) on a galvanome-
ter. The higher the percent transmittance, the fewer
are the cells in suspension.
There should be a direct proportional relationship
between the concentration of bacterial cells and the
absorbance (optical density, O.D.) of the culture. To
demonstrate this principle, you will measure the %T
of various dilutions of the culture provided to you.
These values will be converted to O.D. and plotted on
a graph as a function of culture dilution. You may find
that there is a linear relationship between concentra-
tion of cells and O.D. only up to a certain O.D. At
higher O.D. values the relationship may not be linear.
That is, for a doubling in cell concentration, there may
be less than a doubling in O.D.
Materials:
broth culture of E. coli (same one as used for
plate count)
spectrophotometer cuvettes
(2 per student)
4 small test tubes and test-tube rack
5 ml pipettes
bottle of sterile nutrient broth
(20 ml per student)
Calibrate the spectrophotometer, using the proce-
dure described in figure 23.7. These instructions
are specifically for the Bausch and Lomb
Spectronic 20. In handling the cuvettes, keep the
following points in mind:
a. Rinse the cuvette several times with distilled
water to get it clean before using.
b. Keep the lower part of the cuvette spotlessly
clean by keeping it free of liquids, smudges,
and fingerprints. Wipe it clean with Kimwipes
or some other lint- free tissue. Don't wipe the
cuvettes with towels or handkerchiefs.
c. Insert the cuvette into the sample holder with
its index line registered with the index line on
the holder.
d. After the cuvette is seated, line up the index
lines exactly.
e. Handle these tubes with great care. They are
expensive.
2. Label a cuvette 1 : 1 (near top of tube) and four test
tubes 1:2, 1:4, 1:8, and 1:16. These tubes will be
used for the serial dilutions shown in figure 23.6.
1
\J
BACTERIAL CULTURE
4 ml
1:1
(undiluted)
1:2
1:4
C
1:16
4 ml of sterile nutrient broth in each of these tubes
Figure 23.6 Dilution procedure for cuvettes
3
4
5
6
7
8
With a 5 ml pipette, dispense 4 ml of sterile nutri-
ent broth into tubes 1:2, 1:4, 1:8, and 1:16.
Shake the culture of E. coli vigorously to suspend
the organisms, and with the same 5 ml pipette,
transfer 4 ml to the 1 : 1 cuvette and 4 ml to the 1 : 2
test tube.
Mix the contents in the 1 : 2 tube by drawing the
mixture up into the pipette and discharging it into
the tube three times.
Transfer 4 ml from the 1:2 tube to the 1:4 tube,
mix three times, and go on to the other tubes in a
similar manner. Tube 1:16 will have 8 ml of di-
luted organisms.
Measure the percent transmittance of each of the
five tubes, starting with the 1:16 tube first. The
contents of each of the test tubes must be trans-
ferred to a cuvette for measurement. Be sure to
close the lid on the sample holder when making
measurements. A single cuvette can be used for
all the measurements.
Convert the percent transmittance values to opti-
cal density (O.D.) using the following formula:
O.D. = 2 — log of percent transmittance
Example: If the percent transmittance of one of
your dilutions is 53.5, you would solve the prob-
lem in this way:
97
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
IV. Culture Methods
23. Bacterial Population
Counts
© The McGraw-H
Companies, 2001
Exercise 23 • Bacterial Population Counts
O.D.
= 2 — log of 53.5
= 2- 1.7284
= 0.272
This figure (the characteristic) is always one number
less than the number of digits of the figure you are
looking up.
Examples:
Table II of Appendix A is a log table. Of course, if
you have a calculator, all this is much simpler.
Logarithm Refresher In case you have forgotten
how to use logarithms, recall these facts:
Mantissa: The value you find in the log table
(0.7284 in the above example) is the mantissa.
Characteristic: The number to the left of the dec-
imal (1 in the example) is the characteristic.
9.
10.
number
characteristic
mantissa
5.31
.7251
531
2
.7251
Although the galvanometer may show absorbance
(O.D.) values, greater accuracy will result from
calculating them from percent transmittance.
Record the O.D. values in the table of the
Laboratory Report.
Plot the O.D. values on the graph of the
Laboratory Report.
ITurn on instrument by rotating
zero control knob clockwise. Do
this 20 minutes before measurements
are to be made. Also, set wave-
length knob (top of instrument) at 686
nanometers wavelength. Adjust the
meter needle to zero by rotating zero
control knob.
2 Insert a cuvette containing 3 m
of sterile nutrient broth into sample
holder. The cover must be closed.
Keep the index line of cuvette in line
with index line on the sample holder.
Refer to instructions 1a through 1e on
page 97 concerning care of cuvette.
3 Adjust the meter to read 100%
transmittance by rotating light-
control knob. Remove cuvette of
nutrient broth and close lid. If needle
does not return to zero, readjust
accordingly. Reinsert nutrient broth
again to see if 100% transmittance
still registers. If it has changed, re-
adjust with light-control knob. Once
meter is adjusted for and 100%,
transmittance, turbidity measurements
can be made. Recheck calibration
from time to time to make certain
instrument is set properly.
Figure 23.7 Calibration procedure for the B & L Spectronic 2 on page 97
98
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
IV. Culture Methods
24. Slide Culture:
Autotrophs
© The McGraw-H
Companies, 2001
■
Slide Culture: Autotrophs
2^i
There is probably no single medium or method that
one can use to do a comprehensive population count of
all living microorganisms in a specific biosphere.
Those media that we categorize as being "general pur-
pose" will, for various reasons, inhibit the growth of
many organisms. To make comparative studies of free-
living organisms in freshwater lakes, A. T. Henrici, in
1932, devised an immersed slide technique that re-
vealed the presence of many organisms that did not
show up by other methods. Although his original con-
cern was with algal populations, the technique worked
as well for bacteria and other microorganisms.
His method consists of suspending glass micro-
scope slides in the body of water for a specified period
of time. Microorganisms in the water adhere to the
glass and multiply to form small colonies that are ob-
servable under the microscope. Although there is no
guarantee that the organisms growing on the glass are
autotrophs, many of them are.
Materials:
adhesive tape QA" width)
2 microscope slides
copper wire
gummed labels
acid- alcohol
1
2
3
Clean 2 microscope slides as follows:
a. Scrub with green soap or Bon Ami.
b. Dip them in acid- alcohol for 1 minute and dry
with tissue.
c. Place them in a beaker of distilled water for
5 minutes to allow any residual solvent to
dissipate.
Tape a piece of copper wire to one edge as illus-
trated in figure 24.1. Hold the slides back to back
by their edges. Do not touch the flat surfaces with
your fingers. Wrap all four edges with tape. For
identification, attach a gummed label with your
name to the wire.
Suspend the slide in an aquarium or container of
water that is known to have a stabilized natural
flora of bacteria.
Copper Wire
Adhesive Tape
Figure 24.1 Preparation of slide for immersion
4. After 1 week remove the binding from the slides.
Prepare one slide with Gram's stain and place a
drop of water and cover glass on the other one.
5. Examine both slides under oil immersion and
record your observations on the first portion of
Laboratory Report 24, 25.
Reference: Henrici, A. T. 1933. Studies of fresh wa-
ter bacteria. /. Bact. 25 (3): 277-286.
99
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
IV. Culture Methods
25. Slime Mold Culture
© The McGraw-H
Companies, 2001
Slime Mold Culture
The classification system proposed by Alexopoulos
and Minis places the slime molds in Division
Gymnomycota of the Kingdom Myceteae. These het-
erotrophic microorganisms exist in cool, shady, moist
places in the woods — on decaying logs, dead leaves,
and other organic matter. Unlike the holophytic bac-
teria and other Myceteae, they ingest their food in a
manner similar to the amoebas; that is, they are
phagotrophic. In the vegetative stages, these microor-
ganisms are unlike the other Myceteae in that the cells
lack cell walls; when fruiting bodies are formed, how-
ever, cell walls are present.
The categorization of slime molds as protozoans
or as fungi has always been problematical. Certainly,
they are intermediate in that they have characteristics
of both groups.
Figure 25.1 illustrates the life cycle of one type of
slime mold, the plasmodial type. The genus Physarum
is the one to be studied in this experiment. The assim-
ilative stage of this organism is the Plasmodium. This
multinucleate structure is slimy in appearance and
moves slowly by flowing its cytoplasm in amoeboid
fashion over surfaces on which it feeds. Most species
feed on bacteria and possibly on other small organ-
isms that they encounter.
Plasmodial growth continues as long as ade-
quate food supply and moisture are available.
Eventually, however, environmental changes may
result in the formation of sclerotia or sporangia. A
sclerotium is a hardened mass of irregular shape
that forms from the Plasmodium when moisture and
temperature conditions become less than ideal.
When conditions improve, the sclerotium reverts
back to a Plasmodium. Figure 25.2 is a photograph
of two sclerotia that formed on a laboratory culture.
Sporangia are fructifications that form under con-
ditions similar to those required for sclerotia.
Exactly why sporangia form instead of sclerotia is
still not clearly understood. Sporangia form by the
separation of the Plasmodium into many rounded
mounds of protoplasm that extend upward on stalks.
The nuclei within the sporangia undergo meiosis to
become haploid spores with tough cell walls. The
sclerotia and sporangia of figures 25.2 and 25.3
were photographed on the same culture of labora-
tory-grown Physarum.
mmnmp^npvtwwiHWtata
muniwm
Encystment
Swarm Cells
Spore
Sporangium
(Meiosis)
¥
■Mr-*
Fructification
Figure 25.1 Life cycle of Physarum polycephalum
Isogametes
Zygote
(Amoeboid)
Sclerotium
^
\
.y'
y
■ t
\s
'S: :•
Plasmodium
100
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
IV. Culture Methods
25. Slime Mold Culture
© The McGraw-H
Companies, 2001
Slime Mold Culture • Exercise 25
Both sclerotia and spores may survive adverse en-
vironmental conditions for long periods of time. Once
environmental conditions improve, the spores germi-
nate to produce flagellated pear-shaped swarm cells.
These swarm cells may do one of three things:
(1) they may encyst if conditions suddenly become
adverse, (2) they may divide one or more times to
form isogametes, or (3) they may act as isogametes
and unite directly to form a zygote. Once a zygote is
formed, it takes on an amoeboid form and undergoes
a series of mitotic divisions to produce a Plasmodium.
This completes the life cycle.
Three procedures will be described here for the
study of Phys arum poly cephalum: (1) moist chamber
culture, (2) agar culture method, and (3) spore germi-
nation technique. The techniques used will be deter-
mined by the availability of time and materials.
2
Moist Chamber Culture
To grow large numbers of plasmodia, sclerotia, and
sporangia that can be used for an entire class, one
needs to create a rather large moisture chamber. Any
covered glass or plastic container that is 10 to 12
inches square or round is suitable.
Materials:
sclerotia of Physarum poly cephalum
container for culture (10M" dia Pyrex casserole
dish with cover or 10-12" square plastic box
with cover)
glass Petri dish cover
sharp scalpel
rolled oat flakes (long-cooking type)
10" dia filter paper or paper toweling
1 . In the center of the container place a Petri dish
cover, open end down. Lay a large piece of filter
paper or paper toweling over the Petri dish and
saturate with distilled water. The Petri dish pro-
3
4
vides a raised area above any excess water that
may make the paper too wet.
With a sharp scalpel transfer a small fragment of
sclerotium from the Physarum culture to the filter
paper. A sclerotium may vary from dark orange to
brown in color. See figure 25.2. Moisten the scle-
rotium with a drop of distilled water.
After a few hours the organism will be awakened
to activity and begin to seek food. At this point,
place a flake of rolled oats near the edge of the
spreading growth for it to feed on.
Incubate the moist chamber in a dark place at
room temperature. Add moisture (distilled wa-
ter) and oat flakes periodically as needed. It is
better to add a few fresh flakes daily than to
overfeed by applying all flakes at once. Such a
culture should keep for several weeks. To pro-
mote the formation of sclerotia, allow some of
the water to evaporate away by leaving the lid
partially open for a while. To bring about spo-
rangia formation, withhold food while keeping
the culture moist.
Agar Culture Method
(Plasmodial Study)
An actively metabolizing Plasmodium is dark yellow
and streaked with vessels. The streaming of proto-
plasm in these vessels is best observed under the mi-
croscope. To be able to study this unique structure, it
is best to culture the organism on non-nutrient agar.
Make such a culture as follows:
Materials:
rolled oat flakes
scalpel
Petri plate with 1 5 ml of nonsterile,
non-nutrient agar
"*
tf/.JN
V ,
I
V
w *c
- .
\
:
-i
■
s
II
Figure 25.2 Sclerotia of Physarum polycephalum (3X)
Figure 25.3 Sporangia of Physarum polycephalum (20X)
101
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
IV. Culture Methods
25. Slime Mold Culture
© The McGraw-H
Companies, 2001
Exercise 25 • Slime Mold Culture
1
2
3
4
5
Lift some occupied oat flakes from the filter pa-
per in the moist chamber and transfer to a plate of
nonsterile, non-nutrient 1.5% agar. Maintain this
culture by adding fresh oat flakes periodically, but
don't add water.
After a well-developed Plasmodium has formed,
study the streaming protoplasm under low
power of the microscope. Observation is made
by transmitted light through the agar on the mi-
croscope stage. Look for periodical reversal of
direction of flow.
Cut one of the vessels through in which the flow
is active and observe the effect.
Transfer a piece of Plasmodium to another part of
the medium and watch it reconstitute itself.
Leave the cover slightly open on the Petri dish for
several days and note any changes that might oc-
cur as time goes by.
Spore Germination
The observation of spore germination can be
achieved with a hanging drop slide. Once sporan-
gia are in abundance, one can make such a slide as
follows:
Materials:
depression slides (sterile)
plain microscope slides (sterile)
cover glasses (sterile)
1
2
3
4
5
6
7
Vaseline
toothpicks
sporangia of Physarum polycephalum
Bunsen burner
70% alcohol
With a toothpick, place a small amount of
Vaseline near each corner of the cover glass. (See
figure 19.1, page 73.)
Saturate a sporangium with a drop of 70% alcohol
on the center of a sterile plain microscope slide.
As soon as the alcohol has evaporated, add a drop
of distilled water and place another sterile slide
over the wet sporangium.
Crush the sporangium with thumb pressure on the
upper slide. Separate the two slides to expose the
crushed sporangium.
Transfer a few loopfuls of crushed sporangial ma-
terial to a drop of distilled water on a sterile cover
glass.
Place the depression slide over the cover glass,
make contact, and quickly invert to produce a
completed hanging drop slide.
Examine under low and high power.
Laboratory Report
Complete all the answers on Laboratory Report 24, 25
that pertain to this exercise.
102
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
IV. Culture Methods
26. Slide Culture: Molds
© The McGraw-H
Companies, 2001
Slide Culture
Molds
The isolation, culture, and microscopic examination
of molds require the use of suitable selective media
and special microscopic slide techniques. If simple
wet mount slides of molds were attempted in Exercise
10, it became apparent that wet mount slides made
from mold colonies usually don't reveal the arrange-
ment of spores that is so necessary in identification.
The process of merely transferring hyphae to a slide
breaks up the hyphae and sporangiophores in such a
way that identification becomes very difficult. In this
exercise a slide culture method will be used to prepare
stained slides of molds. The method is superior to wet
mounts in that the hyphae, sporangiophores, and
spores remain more or less intact when stained.
When molds are collected from the environment,
as in Exercise 10, Sabouraud's agar is most frequently
used. It is a simple medium consisting of 1 % peptone,
4% glucose, and 2% agar- agar. The pH of the medium
is adjusted to 5.6 to inhibit bacterial growth.
Unfortunately, for some molds the pH of
Sabouraud's agar is too low and the glucose content is
too high. A better medium for these organisms is one
suggested by C. W. Emmons that contains only 2%
glucose, with 1% neopeptone, and an adjusted pH of
6.8-7.0. To inhibit bacterial growth, 40 mg of chlo-
ramphenicol is added to one liter of the medium.
In addition to the above two media, cornmeal
agar, Czapek solution agar, and others are available
for special applications in culturing molds.
Figure 26.2 illustrates the procedure that will be
used to produce a mold culture on a slide that can be
stained directly on the slide. Note that a sterile cube of
Sabouraud's agar is inoculated on two sides with
spores from a mold colony. Figure 26.1 illustrates
how the cube is held with a scalpel blade as inocula-
tion takes place. The cube is placed in the center of a
microscope slide with one of the inoculated surfaces
placed against the slide. On the other inoculated sur-
face of the cube is placed a cover glass. The assem-
bled slide is incubated at room temperature for 48
hours in a moist chamber (Petri dish with a small
amount of water). After incubation the cube of
medium is carefully separated from the slide and dis-
carded.
During incubation the mold will grow over the
glass surfaces of the slide and cover glass. By adding
a little stain to the slide a semipermanent slide can be
made by placing a cover glass over it. The cover glass
Figure 26.1 Inoculation technique
103
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
IV. Culture Methods
26. Slide Culture: Molds
© The McGraw-H
Companies, 2001
Exercise 26 • Slide Culture: Molds
can also be used to make another slide by placing it on
another clean slide with a drop of stain on it. Before
the stain (lactophenol cotton blue) is used, it is desir-
able to add to the hyphae a drop of alcohol, which acts
as a wetting agent.
First Period
(Slide Culture Preparation)
Proceed as follows to make slide cultures of one or
more mold colonies.
Materials:
Petri dishes, glass, sterile
filter paper (9 cm dia, sterile)
glass U-shaped rods
mold culture plate (mixture)
1 Petri plate of Sabouraud's agar or Emmons'
medium per 4 students
scalpels
inoculating loop
sterile water
microscope slides and cover glasses (sterile)
forceps
1
2
3
4.
5
6
7
8
Aseptically, with a pair of forceps, place a sheet
of sterile filter paper in a Petri dish.
Place a sterile U-shaped glass rod on the filter pa-
per. (Rod can be sterilized by flaming, if held by
forceps.)
Pour enough sterile water (about 4 ml) on filter
paper to completely moisten it.
With forceps, place a sterile slide on the U-shaped
rod.
Gently flame a scalpel to sterilize, and cut a 5 mm
square block of the medium from the plate of
Sabouraud's agar or Emmons' medium.
Pick up the block of agar by inserting the scalpel
into one side as illustrated in figure 26.1.
Inoculate both top and bottom surfaces of the
cube with spores from the mold colony. Be sure to
flame and cool the loop prior to picking up spores.
Place the inoculated block of agar in the center of
a microscope slide. Be sure to place one of the in-
oculated surfaces down.
Aseptically, place a sterile cover glass on the up-
per inoculated surface of the agar cube.
9. Place the cover on the Petri dish and incubate at
room temperature for 48 hours.
10. After 48 hours examine the slide under low
power. If growth has occurred you should see hy-
phae and spores. If growth is inadequate and
spores are not evident, allow the mold to grow an-
other 24-48 hours before making the stained
slides.
Second Period
(Application of Stain)
As soon as there is evidence of spores on the slide,
prepare two stained slides from the slide culture, us-
ing the following procedure:
Materials:
microscope slides and cover glasses
95% ethanol
lactophenol cotton blue stain
forceps
1 . Place a drop of lactophenol cotton blue stain on a
clean microscope slide.
2. Remove the cover glass from the slide culture and
discard the block of agar.
3. Add a drop of 95% ethanol to the hyphae on the
cover glass. As soon as most of the alcohol has
evaporated place the cover glass, mold side
down, on the drop of lactophenol cotton blue
stain on the slide. This slide is ready for exami-
nation.
4. Remove the slide from the Petri dish, add a drop
of 95% ethanol to the hyphae and follow this up
with a drop of lactophenol cotton blue stain.
Cover the entire preparation with a clean cover
glass.
5. Compare both stained slides under the micro-
scope; one slide may be better than the other
one.
Laboratory Report
There is no Laboratory Report for this exercise
104
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
IV. Culture Methods
26. Slide Culture: Molds
© The McGraw-H
Companies, 2001
Slide Culture: Molds • Exercise 26
™*#
Hyphae on cover glass and slide are
first moistened with 95% ethanol and
then stained with lactophenol cotton
blue.
Figure 26.2 Procedure for making two stained slides from slide culture
105
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
IV. Culture Methods
27. Isolation of Anaerobic
Phototrophic Bacteria:
using the Winogradsky
Column
© The McGraw-H
Companies, 2001
27
Isolation of Anaerobic Phototrophic Bacteria:
Using the Winogradsky Column
The culture of photosynthetic bacteria requires spe-
cial culture methods to promote their growth. These
prokaryotes contain photopigments, such as chloro-
phyll and carotenoids, which convert solar energy
into cellular constituents. There are two groups of
phototrophic bacteria: (1) the aerobic phototrophic
cyanobacteria, which we studied in Exercise 6, and
(2) the anaerobic phototrophic bacteria, which in-
clude the purple and green bacteria. It is this latter
group that will be studied in this exercise.
As pointed out in Exercise 6, the cyanobacteria
contain chlorophyll a, carotenoids, and phycobili-
somes. The nonchlorophyll pigments in this group are
accessory pigments for capturing light. They resem-
ble higher plants in that they split water for a source
of reducing power and evolve oxygen in the process.
The anaerobic phototrophic bacteria, on the other
hand, differ in that they contain bacteriochlorophyll,
which is chemically distinct from chlorophyll.
Instead of utilizing water as a source of reducing
power, the purple and green bacteria use sulfide or
organic acids for the reduction of carbon dioxide. The
purple bacteria that utilize organic acids instead of
sulfide are essentially photoheterotrophic since they
derive their carbon from organic acids rather than
carbon dioxide.
These bacteria are ubiquitous in the ooze sedi-
ment of ditches, ponds, and lakes: i.e., mostly every-
where that freshwater lies relatively stagnant for long
periods of time and subject to sunlight. In this envi-
ronment, fermentation processes produce the sulfides
and organic acids that are essential to their existence.
Characterization
According to Bergey's Manual (Section 18, Vol. 3),
there are approximately 30 genera of anaerobic pho-
totrophic bacteria. The purple bacteria belong to the
family Chromatiaceae. The green ones are in the fam-
ily Chlorobiaceae. The morphological, cultural, and
physiological differences between the purple and
green sulfur bacteria are as follows:
Purple Sulfur Bacteria Members of this group are
all gram-negative, straight or slightly curved rods that
are motile with polar flagellation. Colors of the vari-
ous genera vary considerably — from orange-brown to
Aerobic zone-
Microaerophilic
zone
Anaerobic zone-
Paper
fragments
V
X.
Light brown zone
Beggiatoa
Thhbaciilus
Rust colored zone
Rhodospiriflum
Red/purple zone
Chromatium
Green zone
Chlorobium
Black zone
Clostridium
Desuifovibrio
Figure 27.1 Winogradsky's Column
brown, brownish-red to pink, and purple-red to pur-
ple-violet. Color variability is due to the blend of bac-
teriochlorophyll with the type of carotenoid present.
All species contain elemental -sulfur internally in the
form of globules. Some species are able to fix nitro-
gen. Sulfides are required as electron donors; bicar-
bonate, acetate, and pyrovate are also required. They
cannot utilize thiosulfate, sugars, alcohols, amino
acids, or benzoates.
Green Sulfur Bacteria All of these bacteria are
gram-negative, spherical to straight, or curved rods.
Arrangement of cells may be in chains like strepto-
cocci. Some are motile by gliding, others are non-
motile. Color may be grass-green or brown. Sulfur by-
product is excreted, not retained in cells. Some are
106
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
IV. Culture Methods
27. Isolation of Anaerobic
Phototrophic Bacteria:
using the Winogradsky
Column
© The McGraw-H
Companies, 2001
Isolation of Anaerobic Phototrophic Bacteria: Using the Winogradsky Column • Exercise 27
able to utilize thiosulfate. Many are mixotrophic in
that they can photoassimilate simple organic com-
pounds in the presence of sulfide and bicarbonate.
Winogradsky's Column
To create a small ecosystem that is suitable for the
growth of these bacteria, one can set up a
Winogradsky column as illustrated in figure 27.1.
Sergii Winogradsky, a Russian microbiologist, devel-
oped this culture technique to study the bacteria that
are involved in the sulfur cycle. From his studies he
defined the chemoautotrophic bacteria.
This setup consists of a large test tube or gradu-
ated cylinder that is packed with pond ooze, sulfate,
carbonate, and some source of cellulose (shredded pa-
per or cellulose powder). It is incubated for a period
of time (up to 8 weeks) while being exposed to incan-
descent light. Note that different layers of microor-
ganisms develop, much in the same manner that is
found in nature.
Observe that in the bottom of the column the cel-
lulose is degraded to fermentation products by
Clostridium. The fermentation products and sulfate
are then acted upon by other bacteria (Desulfovibrio)
to produce hydrogen sulfide, which diffuses upward
toward the oxygenated zone, creating a stable hydro-
gen sulfide gradient. Note, also, that the Chlorobium
species produce an olive-green zone deep in the col-
umn. A red to purple zone is produced by Chromatium
a little farther up. Ascending the column farther where
the oxygen gradient increases, other phototrophic
bacteria such as Rhodo spirillum, Beggiatoa, and
Thiobacillus will flourish.
Once the column has matured, one can make sub-
cultures from the different layers, using an enrich-
ment medium. The subcultures can be used for mak-
ing slides to study the morphological characteristics
of the various types of organisms. Figure 27.2 illus-
trates the overall procedure to be used for subcultur-
ing. Proceed as follows:
First Period
You will set up your Winogradsky column in a 1 00 ml
glass graduate. It will be filled with mud, sulfate, wa-
ter, phosphate, carbonate, and a source of fermentable
cellulose. The cellulose, in this case, will be in the
form of a shredded paper slurry.
The column will be covered completely at first
with aluminum foil to prevent the overgrowth of
amoeba and then later uncovered and illuminated
with incandescent light to promote the growth of
phototrophic bacteria. The column will be examined
at 2-week intervals to look for the development of
different-colored layers. Once distinct colored lay-
ers develop, subcultures will be made to tubes of en-
richment medium with a pipette. The subcultures
will be incubated at room temperature with exposure
to incandescent light and examined periodically for
color changes. Figure 27.2 illustrates the subcultur-
ing steps.
Materials:
graduated cylinder (100 ml size)
cellulose source (cellulose powder, newspaper,
or filter paper)
calcium sulfate, calcium carbonate, dipotassium
phosphate
mud from various sources (freshly collected)
water from ponds (freshly collected)
beaker (100 ml size)
glass stirring rod
aluminum foil
rubber bands
incandescent lamp (60-75 watt)
1 . Using cellulose powder or some form of paper,
prepare a thick slurry with water in a beaker. If
you are using paper, tear the paper up into small
pieces and macerate it in a small volume of water
with a glass rod. If you are using cellulose pow-
der, start with 1-2 g of powder in a small amount
of water. The slurry should be thick but not a
paste.
2. Fill the cylinder with the slurry until it is one-third
full.
3. To 200 g of mud, add 1.64 g of calcium sulfate
and 1.3 g each of calcium carbonate and dipotas-
sium phosphate. Keep a record of the source of
the mud you are using.
4. Add some "self water" (pond water collected with
the mud) to the mud and chemical mixture and
mix the ingredients well.
5. Pour the mud mixture into the cylinder on top of
the cellulose slurry.
6. With a glass rod, gently mix and pack the contents
of the cylinder. As packing occurs, you may find
that you need to add more "self water" to bring
the level up to two-thirds or three-fourths of the
graduate. Make sure all trapped air bubbles are re-
leased.
7. Top off the cylinder by adding pond water until
the graduate is 90% full.
8. Cap the cylinder with foil, using a rubber band to
secure the cover.
9. Record on the Laboratory Report the initial ap-
pearance of the cylinder.
10. Wrap the sides of the cylinder completely with
aluminum foil to exclude light.
1 1 . Incubate the cylinder at room temperature for one
and a half to two weeks.
107
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
IV. Culture Methods
27. Isolation of Anaerobic
Phototrophic Bacteria:
using the Winogradsky
Column
© The McGraw-H
Companies, 2001
Exercise 27 • Isolation of Anaerobic Phototrophic Bacteria: Using the Winogradsky Column
Two Weeks Later
Remove the aluminum foil from the sides of the
cylinder. Note the color of the mud, particularly in
the bottom. Its black appearance will indicate sulfur
respiration with the formation of sulfides by
Desulfovibrio and other related bacteria. Record the
color differences of different layers and the overall
appearance of the entire cylinder on the Laboratory
Report.
Place a lamp with a 75 watt bulb within a few
inches of the cylinder and continue to incubate the
cylinder at room temperature.
Subsequent Examinations
Examine the cylinder periodically at each laboratory
period, looking for the color changes that might occur.
The presence of green, purple, red, or brown areas on
the surface of the mud should indicate the presence of
blooms of anaerobic phototrophic bacterial growth.
Record your results on the Laboratory Report.
SUBCULTURING
After 6 to 8 weeks, make several subcultures from
your Winogradsky column following the procedure
shown in figure 27.2.
data
'■*■-*,«.*'
- -*
''- aal ^ J ^
Kj
Winogradsky Column
With a wide mouth pipette deliver ap-
proximately 1 gram of mud from each
colored layer to a tube containing Rho
dosplriilaceae enrichment medium.
Make wet mount slides from each tube
and examine with a phase-contrast
microscope.
J
Incubate the inoculated tubes at room
temperature while exposed to a 75 watt
lamp for 3 to 7 days.
^l^ri^^HWU^^dlillUltal^rilWMkMdUiHtUiUiil*
Figure 27.2 Procedure for subculturing and microscopic examination
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
IV. Culture Methods
27. Isolation of Anaerobic
Phototrophic Bacteria:
using the Winogradsky
Column
© The McGraw-H
Companies, 2001
Isolation of Anaerobic Phototrophic Bacteria: Using the Winogradsky Column • Exercise 27
Materials
5 screw-cap test tubes (13 X 200 mm size)
1 prescription bottle containing 200 ml of
Rhodospirillaceae enrichment medium.
5 wide-mouth 1 ml pipettes
1
2
3
Label the screw-cap test tubes with the colors of
the areas to be subcultured from your
Winogradsky column. They may be brown, red,
reddish-purple, or green. If such areas are not ob-
vious, collect mud from areas that are black to
grey.
With a pipette, deliver Rhodospirillaceae enrich-
ment medium from the prescription bottle to each
of the test tubes. Fill each tube about two- thirds
full with the medium.
With a pipette, collect about 1 g of mud from each
colored area of the column and deliver the mud to
the properly labeled tube. Use a fresh pipette for
each delivery.
4
5
6
7
After inoculating each tube, completely fill the
tubes with additional enrichment medium.
Place screw caps on each tube and tighten each
cap securely. Invert each tube several times to mix
the mud and enrichment medium.
Place all the tubes in front of a 75 watt incandes-
cent lamp and incubate at room temperature for
several days to a week.
Observe the cultures at several intervals. When
the cultures have developed a green, red-brown,
or red-purple coloration, make wet mount slides
and examine with a phase-contrast microscope. If
phase-contrast microscopy is unavailable, make
gram- stained slides. Record your results on the
Laboratory Report.
Laboratory Report
Complete the Laboratory Report for this exercise
109
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
V. Bacterial Viruses
Introduction
© The McGraw-H
Companies, 2001
Part
Bacterial Viruses: Isolation
and Propagation
The viruses differ from bacteria in being much smaller, noncellular
and intracellular parasites. In addition, they cannot be grown on or-
dinary media. Despite these seemingly difficult obstacles to labo-
ratory study, we are readily able to detect their presence by ob-
serving their effects upon the cells they parasitize.
Specific viruses are associated with all types of cells, eukary-
otic and prokaryotic. Their dependence on other cells is due to
their inability to synthesize enzymes needed for their own metab-
olism. By existing within cells, however, they are able to utilize the
Phage capsids, tails, and DNA begin
to appear within 1 2 minutes as phage
reorients cell metabolism to its own
fabrication processes.
•■•j *- . „
ilk ; -y-'J
Phage DNA enters cell to initiate
Eclipse Stage. Bacterial DNA begins
to disintegrate within minutes.
Components of phage are assembled
into mature infective virions. The
eclipse period ends with first
appearance of infective phage in cell.
Cell wall opens up due to enzymatic
action to release mature virions.
Burst size is the number of units
released by cell. Total time: 40
minutes.
Adsorption: Phage virion is adsorbed
to specific receptor site on bacterial
cell wall. This is Time Zero.
Figure V.1 The lytic cycle of a virulent bacteriophage
111
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
V. Bacterial Viruses
Introduction
© The McGraw-H
Companies, 2001
Part 5 • Bacterial Viruses: Isolation and Propagation
enzymes of their host. They may contain DNA or RNA, but never
both of these nucleic acids.
The study of viruses that parasitize plant and animal cells is
time-consuming and requires special tissue culture techniques.
Viruses that parasitize bacteria, however, are relatively easy to
study, utilizing ordinary bacteriological techniques. It is for this
reason that bacterial viruses will be studied here. Principles
learned from studying the viruses of bacteria apply to viruses of
eukaryotic cells.
Viruses that parasitize bacteria are called bacteriophage, or
phage. These viruses exist in many shapes and sizes. Some of the
simplest ones exist as a single-stranded DNA virion. Most of them
are tadpole-like, with "heads" and "tails" as seen in figure V.1 on the
previous page. The head, or capsid, may be round, oval, or polyhe-
dral and is composed of protein. It forms a protective envelope for
the DNA of the organism. The tail structure is hollow and provides
an exit for the DNA from the capsid into the cytoplasm of the bac-
terial cell. The extreme end of the tail has the ability to become at-
tached to specific receptor sites on the surface of phage-sensitive
bacteria. Once the tail of the virus attaches itself to a cell, it literally
digests its way through the wall of the host cell.
With the invasion of a bacterial cell by the DNA, one of two
things will occur: lysis or lysogeny. In the event that lysis occurs,
as illustrated on the previous page, the metabolism of the bacterial
cell becomes reoriented to the synthesis of new viral DNA and pro-
tein to produce mature phage particles. Once all the cellular mate-
rial is used up, the cell bursts to release phage virions that, in turn,
are prepared to invade other cells.
Phage that cause lysis are said to be virulent If the phage does
not cause lysis, however, it is termed temperate and establishes a
relationship with the bacterial cell known as lysogeny. In these
cells, the DNA of the phage becomes an integral part of the bacte-
rial chromosome. Lysogenic bacteria grow normally, but their cul-
tures always contain some phage. Periodically, however, phage
virions are released by lysogenized cells in lytic bursts similar to
that seen in the lytic cycle.
Visual evidence of lysis is demonstrated by mixing a culture of
bacteria with phage and growing the mixture on nutrient agar.
Areas where the phage are active will show up as clear spots called
plaques.
The most thoroughly studied bacterial viruses are those that
parasitize Escherichia coli. They are collectively referred to as the
coliphages. They are readily isolated from raw sewage and co-
prophagous (dung-eating) insects. Exercises 28 and 29 pertain to
these techniques. Exercise 30 provides a method for determining
the burst size of a phage. Before attempting any of these experi-
ments, be certain that you thoroughly understand the various
stages in the phage lytic cycle as depicted here.
112
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
V. Bacterial Viruses
28. Isolation of Phage from
Sewer
© The McGraw-H
Companies, 2001
■
Isolation of Phage from Sewage
28
Establishing the presence of phage virions in sewage
involves three steps. First, it is necessary to increase
the phage numbers by enrichment with special media.
Second, it is necessary to separate the phage from the
bacteria by filtration. The final step is to produce
plaque evidence by seeding a "lawn" of bacteria with
phage in the filtrate. Figures 28.1 and 28.2 illustrate
the three steps we will go through. Before beginning
this experiment, however, here are a few comments
about safety procedures that must be followed in han-
dling sewage:
• When collecting sewage samples always wear la-
tex gloves. Raw sewage is a potent source of bac-
terial, viral, and fungal pathogens.
• Raw sewage rich in bacteriophage is best collected
at municipal sewage treatment plants. Usually,
collection is made through manhole access.
• As emphasized in previous pages of this manual,
no mouth pipetting permitted!
Enrichment
To increase the number of phage virions in a raw
sewage sample, it is necessary to add 5 ml of deca-
strength phage broth (DSPB) and 5 ml of E. coli to 45
ml of raw sewage as in illustration 1 of figure 28.1.
The DSPB medium is 10 times as strong as ordinary
broth to accommodate dilution with 45 ml of sewage.
This mixture is incubated at 37° C for 24 hours.
5 ml DSPB
5 ml E. coli
37* C
24 hours
After adding 5 ml of E. coli and 5 ml of double-strength phage broth
(DSPB) to 45 ml. of raw sewage, mixture is incubated at 37° C. for 24
hours.
Sterile membrane filter is asep-
tically placed on filter base.
in mail in
Vacuum
E. coli-sewage culture is triple
centrifuged at 2,500 r.p.m.
Willi!
Supernatant from centrifuge
tubes is filtered.
^m^^rmmn
Filtrate is decanted into a small
sterile Erlenmeyer flask.
Figure 28.1 Enrichment and separation of phage from sewage
113
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
V. Bacterial Viruses
28. Isolation of Phage from
Sewer
© The McGraw-H
Companies, 2001
Exercise 28 • Isolation of Phage from Sewage
Materials:
flask of raw sewage
1 Erlenmeyer flask (125 ml size)
5 ml of DSPB medium
nutrient broth culture of E. coli (strain B)
graduate and 5 ml pipettes
1
2
3
With a graduate, measure out 45 ml of raw
sewage and decant into an Erlenmeyer flask.
Pour 5 ml of DSPB medium and 5 ml of E. coli
into the flask of sewage. If these constituents are
not premeasured, use a 5 ml pipette. If the
medium is pipetted first, the same pipette can be
used for pipetting the E. coli.
Place the flask in the 37° C incubator for 24 hours.
Filtration
Rapid filtration to separate the phage from E. coli in the
enrichment mixture requires adequate centrifugation
first. If centrifugation is inadequate, the membrane fil-
ter will clog quickly and impair the rate of filtration. To
minimize filter clogging, a triple centrifugation pro-
cedure will be used. To save time in the event that filter
clogging does occur, an extra filter assembly and an ad-
equate supply of membrane filters should be available.
These membrane filters have a maximum pore size of
0.45 (xm, which holds back all bacteria, allowing only
the phage virions to pass through.
Materials:
centrifuge and centrifuge tubes (6-12)
2 sterile membrane filter assemblies (funnel,
glass base, clamp, and vacuum flask)
package of sterile membrane filters
sterile Erlenmeyer flask with cotton plug (125
ml size)
forceps and Bunsen burner
vacuum pump and rubber hose
1 . Into 6 or 8 centrifuge tubes, dispense the sewage-£.
coli mixture, filling each tube to within V" of the
top. Place the tubes in the centrifuge so that the load
is balanced. Centrifuge the tubes at 2,500 rpm for
10 minutes.
2. Without disturbing the material in the bottom of
the tubes, decant all material from the tubes to
within 1" of the bottom into another set of tubes.
3. Centrifuge this second set of tubes at 2,500 rpm
for another 10 minutes. While centrifugation is
taking place, rinse out the first set of tubes.
4. When the second centrifugation is complete, pour
off the top two-thirds of each tube into the clean set
of tubes and centrifuge again in the same manner.
5. While the third centrifugation is taking place,
aseptically place a membrane filter on the glass
base of a sterile filter assembly (illustration 3, fig-
ure 28.1). Use flamed forceps. Note that the filter
is a thin sheet with grid lines on it.
114
6. Place the glass funnel over the filter and fix the
clamp in place.
7. Hook up a rubber hose between the vacuum flask
and pump .
8. Carefully decant the top three- fourths of each
tube into the filter funnel. Do not disturb the ma-
terial in the bottom of the tube.
9. Turn on the vacuum pump. If centrifugation has
removed all bacteria, filtration will occur almost
instantly. If the filter becomes clogged and you
have enough filtrate to complete the experiment,
go on to step 10. (If this filtrate is to be used by
the entire class, you will need 25-50 ml.)
If the filter clogs before you have enough fil-
trate, pour the unfiltered material from the funnel
back into another set of centrifuge tubes and re-
centrifuge for 10 minutes at 2,500 rpm.
While centrifugation is taking place, set up
the other filter assembly and pour whatever fil-
trate you have from the first flask into the funnel
of the new setup. After centrifugation, decant the
top three-fourths of material from each tube into
the funnel and turn on the vacuum pump.
Filtration should take place rapidly now.
10. Aseptically transfer the final filtrate from the vac-
uum flask to a sterile 125 ml Erlenmeyer flask
that has a sterile cotton plug. Putting the filtrate in
a small flask is necessary to facilitate pipetting.
Be sure to flame the necks of both flasks while
pouring from one to the other.
Seeding
Evidence of phage in the filtrate is produced by pro-
viding a "lawn" of E. coli and phage. The medium
used is soft nutrient agar. Its jelly like consistency al-
lows for better development of plaques. The soft agar
is poured over the top of prewarmed hard nutrient
agar. Prewarmed plates result in a smoother top agar
surface. Figure 28.2 illustrates the general procedure.
Materials:
nutrient broth culture of E. coli (strain B)
flask of enriched sewage filtrate
4 metal-capped tubes of soft nutrient agar (5 ml
per tube)
4 Petri plates of nutrient agar ( 1 5 ml per plate,
preferably prewarmed at 37° C)
1 ml serological pipettes
1
2
3
Liquefy 4 tubes of soft nutrient agar and cool to
50° C. Keep the tubes in a 50° C water bath to pre-
vent solidification.
Label the tubes 1, 2, 3, and 4. Label the plates 1,
2, 3, and control.
With a 1 ml pipette, transfer 1 drop of filtrate to
tube 1, 3 drops to tube 2, and 6 drops to tube 3.
Don't put any filtrate into tube 4.
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
V. Bacterial Viruses
28. Isolation of Phage from
Sewer
© The McGraw-H
Companies, 2001
4.
5
6
With a fresh 1 ml pipette, transfer 0.3 ml of E. coli
to each of the four tubes of soft agar.
After flaming the necks of each of the soft agar
tubes, pour the contents of each tube over the hard
agar of similarly numbered agar plates. Note that
tube 4 is poured over the control plate.
Once the agar is cooled completely, put the plates,
inverted, into a 37° C incubator. If possible, ex-
Isolation of Phage from Sewage • Exercise 28
amine the plates 3 hours later to look for plaque
formation. If some plaques are visible, measure
them and record their diameters on the Laboratory
Report. Plaque size should be checked every hour
for changes.
Laboratory Report
Record all results on Laboratory Report 28, 29.
Si-:-
V"
V&i
1*> •
Sewage Filtrate
E. coli
.3 m
3 Drops
.3 m
6 Drops
1
Four tubes of liquefied soft
nutrient agar are kept in water
bath at 50° C during inoculation
\^>
1
CONTROL
Tubes of seeded soft agar are poured over prewarmed nutrient agar in plates. The plates are
incubated at 37° C and examined 3 hours later to look for plaque formation.
Figure 28.2 Overlay method of seeding Escherichia coli cultures with phage
115
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
V. Bacterial Viruses
29. Isolation of Phage from
Flies
© The McGraw-H
Companies, 2001
29
Isolation of Phage from Flies
As stated earlier, coprophagous insects, as well as raw
sewage, contain various kinds of bacterial viruses.
Houseflies fall into the coprophagous category be-
cause they deposit their eggs in fecal material where
the young larvae feed, grow, pupate, and emerge as
adult flies. This type of environment is heavily popu-
lated by E. coll and its inseparable parasitic phages.
In this experiment we will follow a procedure that
is quite similar to the one used in working with raw
sewage. An enrichment medium, utilizing cyanide,
will be substituted for the DSPB, however. Figures
29.1 and 29.2 illustrate the procedure.
1
Fly Collection
To increase the probability of success in isolating
phage, it is desirable that one use 20 to 24 houseflies.
A smaller number might be sufficient; the larger num-
ber, however, increases the probability of initial suc-
cess. Houseflies should not be confused with the
smaller blackfly or the larger blowfly. An ideal spot
for collecting these insects is a barnyard or riding sta-
ble. One should not use a cyanide killing bottle or any
other chemical means. Flies should be kept alive until
just prior to crushing and placing them in the growth
medium. There are many ways that one might use to
capture them — use your ingenuity !
Enrichment
Within the flies' digestive tracts are several different
strains of E. coli and bacteriophage. Our first concern
is to enhance the growth of both organisms to ensure
an adequate supply of phage. To accomplish this the
flies must be ground up with a mortar and pestle and
then incubated in a special growth medium for a total
of 48 hours. During the last 6 hours of incubation, a
lysing agent, sodium cyanide, is included in the
growth medium to augment the lysing properties of
the phage.
Materials:
bottle of phage growth medium* (50 ml)
bottle of phage lysing medium* (50 ml)
Erlenmeyer flask (125 ml capacity) with cotton
plug
mortar and pestle (glass)
*see Appendix C for composition
2
3
4.
5
Into a clean nonsterile mortar place 24 freshly
killed houseflies. Pour half of the growth medium
into the mortar and grind the flies to a fine pulp
with the pestle.
Transfer this fly-broth mixture to an empty flask.
Use the remainder of the growth medium to rinse
out the mortar and pestle, pouring all the medium
into the flask.
Wash the mortar and pestle with soap and hot wa-
ter before returning them to the cabinet.
Incubate the fly-broth mixture for 42 hours at 37° C.
At the end of the 42-hour incubation period add
50 ml of lysing medium to the fly-broth mixture.
Incubate this mixture for another 6 hours.
Centrifugation
Before attempting filtration, you will find it necessary
to separate the fly fragments and miscellaneous bac-
teria from the culture medium. If centrifugation is in-
complete, the membrane filter will clog quickly and
filtration will progress slowly. To minimize filter
clogging, a triple centrifugation procedure will be
used. To save time in the event filter clogging does oc-
cur, an extra filter assembly and an adequate supply of
membrane filters should be available. These filters
have a maximum pore size of 0.45 jim, which holds
back all bacteria, allowing only the phage virions to
pass through.
Materials:
centrifuge
6-12 centrifuge tubes
2 sterile membrane filter assemblies (funnel,
glass base, clamp, and vacuum flask)
package of sterile membrane filters
sterile Erlenmeyer flask with cotton plug
(125 ml size)
vacuum pump and rubber hose
1
2
Into 6 or 8 centrifuge tubes, dispense the enrich-
ment mixture, filling each tube to within in l A" of
the top. Place the tubes in the centrifuge so that
the load is balanced. Centrifuge the tubes at 2,500
rpm for 10 minutes.
Without disturbing the material in the bottom of
the tubes, decant all material from the tubes to
within 1" of the bottom into another set of tubes.
116
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
V. Bacterial Viruses
29. Isolation of Phage from
Flies
© The McGraw-H
Companies, 2001
Isolation of Phage from Flies • Exercise 29
Twenty to twenty-four flies are ground up in phage
growth medium with a mortar and pestle.
Crushed flies are incubated in growth medium for 42
hours at 37° C. After adding lysing medium it is
incubated for another 6 hours.
Fly-broth culture is triple-centrifuged at 2,500 rpm
Membrane filter assembly is set up for filtration. This
step must be done aseptically.
Centrifuged supernatant is filtered to produce
bacteria-free phage filtrate.
Phage filtrate is dispensed to a sterile Erlenmeyer flask
from which layered plates will be made (Fig. 29.2).
Figure 29.1 Procedure for preparation of bacteriophage filtrate from houseflies
117
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
V. Bacterial Viruses
29. Isolation of Phage from
Flies
© The McGraw-H
Companies, 2001
Exercise 29 • Isolation of Phage from Flies
3
4,
Centrifuge this second set of tubes at 2,500 rpm
for another 10 minutes. While centrifugation is
taking place, rinse out the first set of tubes.
When the second centrifugation is complete, pour
off the top two-thirds of each tube into the clean set
of tubes and centrifuge again in the same manner.
Filtration
While the third centrifugation is taking place, asepti-
cally place a membrane filter on the glass base of a
sterile filter assembly (illustration 4, figure 29.1). Use
flamed forceps. Note that the filter is a thin sheet with
grid lines on it. Place the glass funnel over the filter
and fix the clamp in place. Hook up a rubber hose be-
tween the vacuum flask and pump.
Now, carefully decant the top three-fourths of
each tube into the filter funnel. Take care not to dis-
turb the material in the bottom of the tube. Turn on the
vacuum pump. If centrifugation and decanting have
been performed properly, filtration will occur almost
instantly. If the filter clogs before you have enough
filtrate, recentrifuge all material and pass it through
the spare filter assembly.
Aseptically, transfer the final filtrate from the
vacuum flask to a sterile 1 25 ml Erlenmeyer flask that
has a sterile cotton plug. Putting the filtrate in a small
flask is necessary to facilitate pipetting. Be sure to
flame the necks of both flasks while pouring from one
to the other.
Inoculation and Incubation
To demonstrate the presence of bacteriophage in the
fly-broth filtrate, a strain of phage-susceptible E. coli
will be used. To achieve an ideal proportion of phage
to bacteria, a proportional dilution method will be
used. The phage and bacteria will be added to tubes of
soft nutrient agar that will be layered over plates of
hard nutrient agar. Soft nutrient agar contains only
half as much agar as ordinary nutrient agar. This
medium and E. coli provide an ideal "lawn" for phage
growth. Its jelly-like consistency allows for better dif-
fusion of phage particles; thus, more even develop-
ment of plaques occurs.
Figure 29.2 illustrates the overall procedure. It is
best to perform this inoculation procedure in the
morning so that the plates can be examined in late af-
ternoon. As plaques develop, one can watch them in-
crease in size with the multiplication of phage and si-
multaneous destruction of E. coli.
Materials:
nutrient broth cultures of Escherchia coli (ATCC
#8677 phage host)
flask of fly-broth filtrate
1 tubes of soft nutrient agar (5 ml per tube)
with metal caps
10 plates of nutrient agar (15 ml per plate, and
pre warmed at 37° C)
1 ml serological pipettes, sterile
1 . Liquefy 1 tubes of soft nutrient agar and cool to
50° C. Keep tubes in water bath to prevent solid-
ification.
2. With a china marking pencil, number the tubes of
soft nutrient agar 1 through 10. Keep the tubes se-
quentially arranged in the test-tube rack.
3. Label 10 plates of prewarmed nutrient agar 1
through 10. Also, label plate 10 negative control.
Prewarming these plates will allow the soft agar
to solidify more evenly.
4. With a 1 ml serological pipette, deliver 0.1 ml of
fly-broth filtrate to tube 1, 0.2 ml to tube 2, etc.,
until 0.9 ml has been delivered to tube 9. Refer to
figure 29.2 for sequence. Note that no fly-broth
filtrate is added to tube 10. This tube will be
your negative control.
5. With a fresh 1 ml pipette, deliver 0.9 ml of E. coli
to tube 1, 0.8 ml to tube 2, etc., as shown in figure
29.2. Note that tube 10 receives 1.0 ml of E. coli.
6. After flaming the necks of each of the tubes, pour
them into similarly numbered plates.
7. When the agar has cooled completely, put the
plates, inverted, into a 37° C incubator.
8. After about 3 hours incubation, examine the
plates, looking for plaques. If some are visible,
measure them and record their diameters on the
Laboratory Report.
9. If no plaques are visible, check the plates again in
another 2 hours.
10. Check the plaque size again at 12 hours, if possi-
ble, recording your results. Incubate a total of 24
hours.
11. Complete Laboratory Report 28, 29.
118
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
V. Bacterial Viruses
29. Isolation of Phage from
Flies
© The McGraw-H
Companies, 2001
Isolation of Phage from Flies • Exercise 29
Fly-Broth Filtrate
\J
E. coil culture
.6
.4
.5
.5
A
.6
.3
J
3
.1
.9
1.0 mi
ml
4
<J
U 1
VJ'
l <J
8
10
\J>
1 Tubes of Soft Agar in 50 C Water-bath
~<^^_. kiibiiii |
J,
Tubes of inoculated soft nutrient agar are
poured over plates of hard nutrient agar and
incubated at 37°C, inverted.
After 3 hours incubation plates are exam-
ined for plaque formation. Periodic exami-
nation after first 3 hours should be made to
observe plaque size changes.
Figure 29.2 Inculation of Escherichia coli with bacteriophage from fly-broth filtrate
119
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
V. Bacterial Viruses
30. Burst Size
Determination: A One-Step
Growth Curve
© The McGraw-H
Companies, 2001
30
Burst Size Determination:
A One- Step Growth Curve
The average number of mature phage virions released
by the lysis of a single bacterial cell is between 20 and
200. This number is called the burst size. It can be de-
termined by adding a small amount of phage to a
known quantity of bacteria and then lysing the cells at
5-minute intervals with chloroform. The chloroform-
lysed cells, in turn, are mixed with bacteria, plated
out, and incubated. By counting the plaques, it is pos-
sible to determine the burst size. In this experiment
we will determine the burst size of coliphage T4 on
host cells E. coli, strain B .
Adsorption
Figure 30.1 illustrates the procedure of this experi-
ment. The first step is to add the phage to the suscep-
tible bacteria. As soon as the two are mixed, adsorp-
tion begins. The phage collide in random fashion with
the bacterial cells and attach their tails to specific re-
ceptor sites on the surfaces of host cells. The adsorp-
tion process can be stopped at any time by dilution.
Time zero of adsorption is the time of mixture of
phage and bacteria.
Note in figure 30.1 that 0.1 ml of coliphage T4 (2
X 10 8 /ml) and 2 ml of E. coli (5 X 10 8 /ml) are mixed
in the first tube, which is labeled ADS. The ratio of
phage to bacteria in this case is 0.02, which calculates
out in this manner:
0.1 X 2 X 10 8 0.2 X 10 8
2 X 5 X 10
8
10 X 10 8
= 0.02, or 1/50
This ratio is called the multiplicity of infection, or
m.o.i.
By referring back to figure V.l on page 111, we
can see what is occurring in this experiment. Note that
during the adsorption stage, DNA in the capsid passes
down through the tail into the host through a hole pro-
duced in the cell wall by enzymatic action at the tip of
the phage tail.
Eclipse Stage
As soon as the phage DNA gets inside the bacterial
cell, the phage enters the eclipse stage. During this
stage, which lasts approximately 12 minutes, the en-
tire physiology of the host cell is reoriented toward
the production of phage components: capsids, tails,
and DNA. If the cell is experimentally lysed with
chloroform during this period of time, it will be seen
that the incomplete components of phage are unable
to infect new cells (no plaques are formed).
Maturation Stage
As phage components begin to assemble late in the
eclipse stage to form mature infective virions, the
phage enters the maturation stage. The lysing of
cultures with chloroform beyond 1 2 minutes of time
zero will reveal the presence of these mature units
by producing plaques on poured plates. Lysis of a
population of infected cells does not occur instanta-
neously, but instead follows a normal distribution
curve, or rise period. The rise period, which lasts
for several minutes, represents the growth in num-
bers of mature phage present. The peak of the curve
is the burst size. It is this value that will be deter-
mined here.
Two Methods
To accommodate the availability of time and mate-
rials, there are two options for performing this ex-
periment. The first option is for students to work in
pairs to perform the entire experiment. Figure 30.1
illustrates the procedure for this method. The other
option, which requires much less media and time,
utilizes a team approach in which students, working
in pairs, do just a portion of the experiment; in this
case, data are pooled to complete the experiment.
Figure 30.3 illustrates the procedure for this
method. Your instructor will indicate which method
will be used.
The Entire Experiment
To perform the experiment in its entirety, follow the
procedures that are shown in figure 30. 1 .
Materials:
1 sterile serological tube (for ADS tube)
15 tubes tryptone broth (9.9 ml in each one)
8 tubes of nutrient soft agar (3 ml per tube)
8 Petri plates of tryptone agar
16 pipettes (1 ml size)
120
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
V. Bacterial Viruses
30. Burst Size
Determination: A One-Step
Growth Curve
© The McGraw-H
Companies, 2001
Burst Size Determination: A One-Step Growth Curve • Exercise 30
2
5 min. later
0.1 ml.
1
(0 + 5 min.)
WATER BATH
37° C
ADS
TIME ZERO
2 ml. E. co// and
0.1 ml. phage
added to ADS tube
0.3 ml. E. coli
0.1 ml.
1:1,000,000
Soft Agar
50° C
1:10,000,000
0.3 ml. E. coli
0.1 ml.
1:10,000
1:10,000,000
Soft Agar
50° C
1:10,000,000
At proper time intervals (every 5 minutes) 0.1 ml. is pipetted from ADS-2 tube through two tubes of tryptone broth
and into soft agar (as in steps 3 and 4) to overlay five more plates. All plates are incubated at 37° C.
50
1:10,000,000
1:10,000,000
1:10,000,000
1:10,000,000
1:10,000,000
Figure 30.1 Procedure for entire experiment
121
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
V. Bacterial Viruses
30. Burst Size
Determination: A One-Step
Growth Curve
© The McGraw-H
Companies, 2001
Exercise 30 • Burst Size Determination: A One-Step Growth Curve
1 dropping bottle of chloroform
1 small wire basket to hold 7 tubes of soft agar
in water bath
1 wire test-tube rack
2 water baths (37° C and 50° C)
1 culture of E. coli, strain B (5 ml) with
concentration of 5 X 1 8 per ml
1 tube of T4 phage (2 X 10 8 per ml)
Preli
1
2
3.
4.
5
lm inane s
Liquefy 8 tubes of soft nutrient agar by boiling in
a beaker of water. Cool to 50° C and place in a
wire basket or rack in 50° C water bath.
Label a sterile serological tube "ADS" to signify
the adsorption tube.
Label 1 tube of tryptone broth "ADS-2."
Label 8 tryptone agar plates: control, 15, 25, 30,
35, 40, 45, and 50.
Arrange the ADS, ADS-2, and the 14 tryptone
broth tubes in a rack as shown in figure 30.2.
Place the rack in a 37° C water bath.
ADS
OOOOOO O
ooooooo
ADS-2
Figure 30.2 Tube arrangement
6. Dispense 3 to 4 drops of chloroform in each of the
7 tubes of tryptone broth that are in the front row.
Inoculations and Dilutions
1
2
3
4
Pipette 0.1 ml of E. coli, strain B, into a tube of
liquefied soft nutrient agar and pour into the con-
trol plate. Swirl the plate gently to spread evenly.
This plate will indicate whether any phage was in
the original bacterial culture. Set this plate aside
to harden.
With the same pipette as above, transfer 0.3 ml of
E. coli into each of the tubes of soft nutrient agar.
Keep the tubes in the 50° C water bath.
Still using the same pipette, transfer 2.0 ml of E.
coli into the ADS tube.
With a fresh pipette, deliver 0.1 ml of T4 phage
into the ADS tube and immediately record the
time {time zero) of this mixing with E. coli in the
following table. Mix gently and allow to remain
in the 37° C water bath for 5 minutes.
TIME
Time zero
Step 6 time (5 min later)
Step 7 time (10 min later)
Step 8 time (10 min later)
Five minutes later
Five minutes later
Five minutes later
Five minutes later
Five minutes later
PLATE
none
none
15
25
30
35
40
45
50
5. While the mixture is incubating, fill in the table,
recording all the projected times so that you will
know when each step is to begin.
6. After the 5-minute incubation time, transfer 0.1
ml of the mixture to the ADS-2 tube, gently mix,
and incubate at 37° C for another 10 minutes.
7. After 10 minutes, transfer 0.1 ml from the ADS-2
tube to the first front row tube of tryptone broth.
Keep the ADS-2 tube in the water bath. Mix this
dilution tube gently and transfer 0.1 ml to the ad-
jacent tryptone broth tube in the second row. Mix
this tube gently, also.
8. Transfer 0.1 ml from the second tube of tryptone
broth to a tube of soft nutrient agar, mix gently,
flame the tube neck, and pour the soft agar over
the tryptone agar plate that is labeled "15."
Swirl the plate carefully to disperse the soft
agar mixture evenly.
9. Follow the above procedure 10 minutes later to
produce a soft agar overlay plate on the plate la-
beled "25."
10. Repeat at the allotted times for 30-, 35-, 40-, 45-,
and 50-minute plates.
11. Invert and incubate all plates for 24-48 hours at
37° C.
Examination of the Plates
Once the plates have been incubated, count the
plaques on all the plates, using a Quebec colony
counter and hand tally counter. Record all counts on
the Laboratory Report and determine burst size.
Abbreviated Procedure
(Team Method)
Performance of this experiment in teams will require
a minimum of seven pairs of students. Each pair of
students (team) will follow the procedure shown in
122
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
V. Bacterial Viruses
30. Burst Size
Determination: A One-Step
Growth Curve
© The McGraw-H
Companies, 2001
Burst Size Determination: A One-Step Growth Curve • Exercise 30
figure 30.3 to produce one soft agar overlay plate for
a designated time.
Materials:
per team:
1 sterile serological tube
3 tubes of tryptone broth (9.9 ml per tube)
2 tubes of soft nutrient agar (3 ml per tube)
2 Petri plates of tryptone agar
4 1 ml pipettes
1 dropping bottle of chloroform
1 wire test-tube rack (small size)
1 small beaker (150 ml size)
1 tube of T4 phage (2 X 10 8 per ml)
1 culture of E. coli, strain B (5 X 1 8 per ml)
water bath at 37° C (a small pan that will hold a
test-tube rack)
Preli
lm inane s
1.
Liquefy two tubes of soft nutrient agar in boiling
water. Use a small beaker. Cool the water to 50°
C and keep the tubes of media at this temperature.
2
3
4
5
6
7
Label a sterile serological tube "ADS" to signify
the adsorption tube.
Label one tube of tryptone broth "ADS-2."
Label the other tryptone tubes "I" and "II."
Label one tryptone agar plate "control" and the
other your designated time (15, 25, 30, 35, 40, 45,
or 50). Your instructor will assign you a specific
time. Put your names on both plates.
Arrange the ADS, ADS-2, and two tryptone tubes
in a small test-tube rack in same order as shown
in figure 30.3.
Place the rack of tubes in a pan of 37° C water.
Although it is only necessary to incubate the ADS
and ADS-2 tubes, it will be more convenient if
they are all together.
Inoculations and Dilutions
1. Pipette 0.1 ml of E. coli, strain B, into a tube of
liquefied soft nutrient agar and pour it into the
control plate. Swirl the plate gently to spread
evenly.
.1 ml.
4 Drops Chloroform
1 ml.
(TIME ZERO)
2.0 ml. f. cofi
. 1 ml. Phage
4DS
ADS2
37" C.
9.9
^
v_
1:100
9.9
9,9
1:10,000
1 : 1 ,000,000
1:10,000,000
Figure 30.3 Abbreviated procedure (team) method
123
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
V. Bacterial Viruses
30. Burst Size
Determination: A One-Step
Growth Curve
© The McGraw-H
Companies, 2001
Exercise 30 • Burst Size Determination: A One-Step Growth Curve
2
3
4
This plate will indicate whether any phage
was in the original bacterial culture. Set this plate
aside to harden.
With the same pipette, transfer 0.3 ml of E. coli to
the other tube of soft nutrient agar. Keep this tube
in the beaker of water at 50° C.
Still using the same pipette, transfer 2.0 ml of E.
coli into the ADS tube.
With a fresh pipette, deliver 0. 1 ml of T4 phage
into the ADS tube.
Record this time (time zero):
5
6
7
After 5 minutes, transfer 0.1 ml of the E.
<%>//-phage mixture from the ADS tube to ADS -2
tube. Mix the ADS-2 tube gently.
After the designated time (time zero plus desig-
nated time), transfer 0.1 ml from ADS-2 tube to
tryptone broth tube I. Mix gently.
Add 3 or 4 drops of chloroform to tube I.
8. With a fresh pipette, transfer 0. 1 ml from tube I to
tube II. Mix tube II gently.
9. With another fresh pipette, transfer 0.1 ml from
tube II to the tube of soft agar.
10. After mixing the soft agar tube, pour it over the
tryptone agar plate. Swirl the plate carefully to
disperse the soft agar. Set aside to cool for a few
minutes.
11. Incubate both plates at 37° C for 24-48 hours.
Examination of Plates
Once the plates have been incubated, examine both of
them on a Quebec colony counter. The control plate
should be free of plaques. Count the plaques on the
other plate, using a hand tally counter if the number is
great. Record your count on the Laboratory Report
and on the chalkboard.
124
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
VI. Microbial
Interrelationships
Introduction
© The McGraw-H
Companies, 2001
Part
Microbial Interrelationships
Populations within the microbial world relate to each other in vari-
ous ways. Although many of them will be neutralistic toward each
other by not interacting in any way, others will establish relation-
ships that are quite different.
The three exercises in this unit reveal how certain organisms
have developed relationships that are commensalistic, synergistic,
and antagonistic. While most of these relationships are between
bacteria, some are between bacteria and molds.
125
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
VI. Microbial
Interrelationships
31. Bacterial
Commensalism
© The McGraw-H
Companies, 2001
■
Bacterial Commensalism
There are many commensalistic relationships that exist
between organisms in a mixed microbial population.
The excretory products of one organism often become
the nutrients of another. The oxygen usage of one
species may produce the desired oxidation-reduction
potential for another organism. In all cases of com-
mensalism, the beneficiary contributes nothing in the
way of benefit or injury to the other.
In this exercise we will culture two organisms sep-
arately and together to observe an example of commen-
salism. One of the organisms is Staphylococcus aureus
and the other is Clostridium sporogenes. From your ob-
servations of the results, you are to determine which or-
ganism profits from the association and what control-
ling factor is changed when the two are grown together.
Materials:
3 tubes of nutrient broth
1 ml pipette
nutrient broth culture of S. aureus
fluid thioglycollate medium culture of C.
sporogenes
1
2
3
4
First Period
Label one tube of nutrient broth S. aureus, a sec
ond tube C. sporogenes, and the third tube S. au
reus and C. sporogenes.
Inoculate the first and third tubes with one loop
ful each of S. aureus.
With a 1 ml pipette, transfer 0.1 ml of C. sporo
genes to tubes 2 and 3.
Incubate the three tubes at 37° C for 48 hours.
1
2
Second Period
Compare the turbidity in the three tubes, noting
which ones are most turbid. Record these results
on the Laboratory Report.
After shaking the tubes for good dispersion, make
a gram-stained slide of the organisms in each tube
and record your observations on combined
Laboratory Report 31-33.
126
Benson: Microbiological
VI. Microbial
32. Bacterial Synergism
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Interrelationships
Companies, 2001
Bacterial Synergism
32
Two or more organisms acting together to produce a
substance that none can produce separately is a syner-
gistic relationship. Such relationships are not uncom-
mon among microorganisms. This phenomenon is
readily demonstrated in the ability of some bacteria
acting, synergistically, to produce gas by fermenting
certain disaccharides.
In this exercise we will observe the fermentation
capabilities of three organisms on two disaccharides.
The two sugars, lactose and sucrose, will be inocu-
lated with the individual organisms as well as with
various combinations of the organisms to detect
which organisms can act synergistically on which
sugars. To conserve on media, the class will be di-
vided into three groups (A, B, and C). Results of in-
oculations will be shared.
Materials:
per pair of students:
3 Durham* tubes of lactose broth with
bromthymol blue indicator
3 Durham* tubes of sucrose broth with
bromthymol blue indicator
1 nutrient broth culture of S. aureus
1 nutrient broth culture of P. vulgaris
1 nutrient broth culture of E. coli
*A Durham tube is a fermentation tube of sugar
broth that has a small inverted vial in it. See
figure 48.3, page 164.
First Period
Group A
1 . Label one tube of each kind of broth E. coli.
2. Label one tube of each kind of broth P. vulgaris,
3. Label one tube of each kind of broth E. coli and P.
vulgaris.
4. Inoculate each tube with one loopful of the ap-
propriate organisms.
5. Incubate the six tubes at 37° C for 48 hours.
Group B
1 . Label one tube of each kind of broth E. coli.
2. Label one tube of each kind of broth S. aureus.
3. Label one tube of each kind of broth E. coli and
S. aureus.
4. Inoculate each tube with one loopful of the ap-
propriate organisms.
5. Incubate the six tubes at 37° C for 48 hours.
Group C
1.
2.
3.
4.
5.
1.
2.
3.
Label one tube of each kind of broth S. aureus.
Label one tube of each kind of broth P. vulgaris.
Label one tube of each kind of broth S. aureus and
P. vulgaris.
Inoculate each tube with one loopful of the ap-
propriate organisms.
Incubate the six tubes at 37° C for 48 hours.
Second Period
Look for acid and gas production in each tube,
recording your results on the Laboratory Report.
Determine which organisms acted synergistically
on which disaccharides.
Answer the questions for this exercise on com-
bined Laboratory Report 31-33.
127
Benson: Microbiological
VI. Microbial
33. Microbial Anatagonism
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Interrelationships
Companies, 2001
■
33
Microbial Antagonism
Microbial antagonisms, in which one organism is in-
hibited and the other is unaffected, are easily demon-
strated. Usually, the inhibitor produces a substance
that inhibits or kills one or more organisms. The sub-
stance may be specific in its action, affecting only a
few species, or it may be nonspecific, affecting a large
number of organisms.
In this exercise we will attempt to evaluate the an-
tagonistic capabilities of three organisms on two test
organisms. The antagonists are Bacillus cereus var. my-
coides, Pseudomonas fluoresceins, and Penicillium no-
tatum. The test organisms are Escherichia coli (gram-
negative) and Staphylococcus aureus (gram-positive).
Materials:
6 nutrient agar pours
6 sterile Petri plates
nutrient broth cultures of E. coli, S. aureus, B.
cereus var. mycoides, and P. fluorescens
flask culture of Penicillium notatum (8-12 day
old culture)
2
First Period
1 . Liquefy six nutrient agar pours and cool to 50° C
Hold in 50° C water bath.
3
4
5
6
1
2
While the pours are being liquefied, label six
plates as follows:
Test Organism
I S. aureus
II S. aureus
III S. aureus
IV E. coli
V E. coli
VI E. coli
Antagonist
B. cereus var. mycoides
P. fluorescens
Penicillium notatum
B. mycoides
P. fluorescens
Penicillium notatum
Label three liquefied pours S. aureus, and label
the other three E. coli.
Inoculate each of the pours with a loopful of the
appropriate organisms, flame their necks, and
pour into their respective plates.
After the nutrient agar in the plates has hardened,
streak each plate with the appropriate antagonist.
Use a good isolation technique.
Invert and incubate the plates for 24 hours at 37° C.
Second Period
Examine each plate carefully, looking for evi-
dence of inhibition.
Record your results on combined Laboratory
Report 31-33 and answer all the questions.
128
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
VII. Environmental
Influences and Control of
Microbial Growth
Introduction
© The McGraw-H
Companies, 2001
Part
Environmental Influences and
Control of Microbial Growth
The 1 1 exercises of this unit are concerned with two aspects of mi-
crobial growth: promotion and control. On the one hand, the mi-
crobiologist is concerned with providing optimum growth condi-
tions to favor maximization of growth. The physician, nurse, and
other members of the medical arts profession, on the other hand,
are concerned with the limitation of microbial populations in dis-
ease prevention and treatment. An understanding of one of these
facets of microbial existence enhances the other.
n Part 4 we were primarily concerned with providing media for
microbial growth that contain all the essential nutritional needs.
Very little emphasis was placed on other limiting factors such as
temperature, oxygen, or hydrogen ion concentration. An organism
provided with all its nutritional needs may fail to grow if one or more
of these essentials are not provided. The total environment must be
sustained to achieve the desired growth of microorganisms.
Microbial control by chemical and physical means involves
the use of antiseptics, disinfectants, antibiotics, ultraviolet light,
and many other agents. The exercises of this unit that are related
to these aspects are intended, primarily, to demonstrate methods
of measurement; no attempt has been made to make in-depth
evaluation.
129
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
VII. Environmental
Influences and Control of
Microbial Growth
34. Temperature: Effects on
Growth
© The McGraw-H
Companies, 2001
34
Temp er ature :
Effects
on
Gr
ow
th
Temperature is one of the most important factors influ-
encing the activity of bacterial enzymes. Unlike warm-
blooded animals, the bacteria lack mechanisms that
conserve or dissipate heat generated by metabolism,
and consequently their enzyme systems are directly af-
fected by ambient temperatures. Enzymes have mini-
mal, optimal, and maximal temperatures. At the opti-
mum temperature the enzymatic reactions progress at
maximum speed. Below the minimum and above the
maximum temperatures the enzymes become inactive.
At some point above the maximum temperature, de-
struction of a specific enzyme will occur. Low temper-
atures are less deleterious in most cases.
Microorganisms grow in a broad temperature range
that extends from approximately 0° C to above 90° C.
They are divided into three groups: mesophiles that
grow between 10° C and 47° C, psychrophiles that are
able to grow between 0° C and 5° C, and thermophiles
that grow at high temperatures (above 50° C).
The psychrophiles and thermophiles are further
subdivided into obligate and facultative groups.
Obligate psychrophiles seldom grow above 22° C and
facultative psychrophiles (psychrotrophs) grow very
well above 25° C. Thermophiles that thrive only at
high temperatures (above 50° C and not below 40° C)
are considered to be obligate thermophiles; those that
will grow below 40° C are considered to be faculta-
tive thermophiles.
In this experiment we will attempt to measure the
effects of various temperatures on two physiological
reactions: pigment production and growth rate.
Nutrient broth and nutrient agar slants will be inocu-
lated with three different organisms that have differ-
ent optimum growth temperatures. One organism,
Serratia marcescens, produces a red pigment called
prodigiosin that is produced only in a certain temper-
ature range. It is our goal here to determine the opti-
mum temperature for prodigiosin production and the
approximate optimum growth temperatures for all
three microorganisms. To determine optimum growth
temperatures we will be incubating cultures at five
different temperatures. A spectrophotometer will be
used to measure turbidity densities in the broth cul-
tures after incubation.
First Period
(Inoculations)
To economize on time and media it will be necessary
for each student to work with only two organisms and
seven tubes of media. Refer to table 34.1 to deter-
mine your assignment. Figure 34.1 illustrates the
procedure.
Materials:
nutrient broth cultures of Serratia marcescens,
Bacillus stearothermophilus, and
Escherichia coli
per student:
2 nutrient agar slants
5 tubes of nutrient broth
1
2
3
Label the tubes as follows:
Slants: Label both of them S. marcescens; label
one tube 25° C and the other tube 38° C.
Broths: Label each tube of nutrient broth with
your other organism and one of the following five
temperatures: 5° C, 25° C, 38° C, 42° C, or 55° C.
Inoculate each of the tubes with the appropriate
organisms. Use a wire loop.
Place each tube in one of the five baskets that is
labeled according to incubation temperature.
Table 34.1 Inoculation Assignments
Student Number
S. marcescens
B. stearothermophilus
E. coli
1,4,7, 10,13,16,19,22,25
2 slants and 5 broths
2,5,8, 11,14,17,20,23,26
2 slants
5 broths
3,6,9, 12,15,18,21,24,27
2 slants
5 broths
130
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
VII. Environmental
Influences and Control of
Microbial Growth
34. Temperature: Effects on
Growth
© The McGraw-H
Companies, 2001
Temperature: Effects on Growth • Exercise 34
Note: The instructor will see that the 5° C basket
is placed in the refrigerator and the other four are
placed in incubators that are set at the proper
temperatures.
Second Period
(Tabulation of Results)
Materials:
slants and broth cultures that have been
incubated at various temperatures
spectrophotometer and cuvettes
tube of sterile nutrient broth
1 . Compare the nutrient agar slants of S. marcescens.
Using colored pencils, draw the appearance of the
growths on the Laboratory Report.
2. Shake the broth cultures and compare them, not-
ing the differences in turbidity. Those tubes that
3
4
5
appear to have no growth should be compared
with a tube of sterile nutrient broth.
If a spectrophotometer is available, determine the
turbidity of each tube following the instructions
on the Laboratory Report.
If no spectrophotometer is available, record tur-
bidity by visual observation. The Laboratory
Report indicates how to do this.
Exchange results with other students to complete
data collection for experiment.
Laboratory Report
After recording all data, answer the questions on the
Laboratory Report for this exercise.
S. marcescens
s
S. marcescens, B. stearothermophilus, E. coli
\
*»-"f
25
o
-y.
' '"-;'■* ■•
.f. »
* '
' .* *■'/*»
-.$ •:
■ T.V ' .
38
o
fFT
"\
- » .
■4'
*
V
O
* r ". ' . .
i * ■. ■ .
- • * ' • C** i
' ■,
". v i*v'
38
o
■ • .* -
.•• •
J,";:/ • '<Jy£ ■
■«»•• . ..;:«:
• \ * "I
..\
•.• ,r
55
o
■v
■v
Two nutrient agar slants are
streaked with S. marcescens and
incubated at different temperatures
for pigment production.
Five nutrient broths are inoculated with one of three organisms
and incubated at five different temperatures to determine
optimum growth temperatures for each organism.
Figure 34 Inoculation procedure
131
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
VII. Environmental
Influences and Control of
Microbial Growth
35. Temperature: Lethal
Effects
© The McGraw-H
Companies, 2001
35
Temp er atur e :
Lethal Effects
In attempting to compare the susceptibility of differ-
ent organisms to elevated temperatures, it is necessary
to use some yardstick of measure. Two methods of
comparison are used: the thermal death point and the
thermal death time. The thermal death point (TDP)
is the temperature at which an organism is killed in 10
minutes. The thermal death time (TDT) is the time
required to kill a suspension of cells or spores at a
given temperature. Since various factors such as pH,
moisture, composition of medium, and age of cells
will greatly influence results, these variables must be
clearly stated.
In this exercise we will subject cultures of three
different organisms to temperatures of 60°, 70°, 80°,
90°, and 100° C. At intervals of 10 minutes organisms
will be removed and plated out to test their viability.
The spore-former Bacillus megaterium will be com-
pared with the non- spore-formers Staphylococcus au-
reus and Escherichia coli. The overall procedure is il-
lustrated in figure 35.1.
Note in figure 35.1 that before the culture is
heated a control plate is inoculated with 0. 1 ml of the
organism. When the culture is placed in the water
bath, a tube of nutrient broth with a thermometer in-
serted into it is placed in the bath at the same time.
Timing of the experiment starts when the thermome-
ter reaches the test temperature.
Due to the large number of plates that have to be
inoculated to perform the entire experiment, it will be
necessary for each member of the class to be assigned
a specific temperature and organism to work with.
Table 35.1 provides assignments by student number.
After the plates have been incubated, each student's
results will be tabulated on a Laboratory Report chart
at the demonstration table. The instructor will have
copies made of it to give each student so that every-
one will have all the pertinent data needed to draw the
essential conclusions.
Although this experiment is not difficult, it often
fails to turn out the way it should because of student
error. Common errors are (1) omission of the control
plate inoculation, (2) putting the thermometer in the
culture tube instead of in a tube of sterile broth, and
(3) not using fresh sterile pipettes when instructed to
do so.
Materials:
per student:
5 Petri plates
5 pipettes ( 1 ml size)
1 tube of nutrient broth
1 bottle of nutrient agar (60 ml)
1 culture of organisms
class equipment:
water baths set up at 60
100° C
o
70°, 80°, 90°, and
broth cultures:
Staphylococcus aureus, Escherichia coli, and
Bacillus megaterium (minimum of 5
cultures of each species per lab section)
1. Consult table 35.1 to determine what organism
and temperature has been assigned to you. If
several thermostatically controlled water baths
have been provided in the lab, locate the one that
you will use. If a bath is not available for your
temperature, set up a bath on an electric hot plate
or over a tripod and Bunsen burner.
If your temperature is 100° C, a hot plate and
beaker of water are the only way to go. When set-
Table 35.1 Inoculation Assignments
Organism
Student Number
60° C
70° C
80° C
90° C
100° c
Staphylococcus aureus
1, 16
4,19
7,22
10,25
13,28
Escherichia coli
2,17
5,20
8,23
11,26
14,29
Bacillus megaterium
3,18
6,21
9,24
12,27
1 5, 30f
132
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
VII. Environmental
Influences and Control of
Microbial Growth
35. Temperature: Lethal
Effects
© The McGraw-H
Companies, 2001
2
3
4,
5
6
ting up a water bath use hot tap water to start with
to save heating time.
Liquefy a bottle of 60 ml of nutrient agar and cool
to 50° C. This can be done while the rest of the ex-
periment is in progress.
Label five Petri plates: control, 10 min, 20 min,
30 min, and 40 min.
Shake the culture of organisms and transfer 0.1 ml
of organisms with a 1 ml pipette to the control plate.
Place the culture and a tube of sterile nutrient
broth into the water bath. Remove the cap from the
tube of nutrient broth and insert a thermometer
into the tube. Don't make the mistake of inserting
the thermometer into the culture of organisms!
As soon as the temperature of the nutrient broth
reaches the desired temperature, record the time
here: .
Watch the temperature carefully to make sure it
does not vary appreciably.
7
8
9
Temperature: Lethal Effects • Exercise 35
After 10 minutes have elapsed, transfer 0.1 ml
from the culture to the 10-minute plate with a
fresh 1 ml pipette. Repeat this operation at 10-
minute intervals until all the plates have been in-
oculated. Use fresh pipettes each time and be sure
to shake the culture before each delivery.
Pour liquefied nutrient agar (50° C) into each
plate, rotate, and cool.
Incubate at 37° C for 24 to 48 hours. After evalu-
ating your plates, record your results on the chart
on the Laboratory Report and on the chart on the
demonstration table.
Laboratory Report
Complete the Laboratory Report once you have a
copy of the class results.
Every ten minutes 0.1 ml
of culture is pipetted into
one of four plates.
Control
20 Minutes
40 Minutes
Figure 35.1 Procedure for determining thermal endurance
133
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
VII. Environmental
Influences and Control of
Microbial Growth
36. pH and Microbial
Growth
© The McGraw-H
Companies, 2001
36
pH and Microbial Growth
Aside from temperature, the hydrogen ion concentra-
tion of an organism's environment exerts the greatest
influence on its growth. The concentration of hydro-
gen ions, which is customarily designated by the term
pH (—log 1/H + ), limits the activity of enzymes with
which an organism is able to synthesize new proto-
plasm. As in the case of temperature, there exists for
each organism an optimum concentration of hydrogen
ions in which it grows best. The pH values above and
below which an organism fails to grow are, respec-
tively, referred to as the minimum and maximum hy-
drogen ion concentrations. These values hold only
when other environmental factors remain constant. If
the composition of the medium, incubation tempera-
ture, or osmotic pressure is varied, the hydrogen ion
requirements become different.
In this exercise we will test the degree of inhibi-
tion of microorganisms that results from media con-
taining different pH concentrations. Note in the mate-
rials list that tubes of six different hydrogen
concentrations are listed. Your instructor will indicate
which ones, if not all, will be tested.
1.
First Period
Materials:
per student:
1 tube of nutrient broth of pH 3.0
1 tube of nutrient broth of pH 5.0
1 tube of nutrient broth of pH 7.0
1 tube of nutrient broth of pH 8.0
1 tube of nutrient broth of pH 9.0
1 tube of nutrient broth of pH 10.0
class materials:
broth cultures of Escherichia coli
broth cultures of Staphylococcus aureus
broth cultures of Alcaligenes faecalis*
broth cultures of Saccharomyces cerevisiae**
Inoculate a tube of each of these broths with one
organism. Use the organism following your as-
signed number from the table below:
Student Number Organism
1,5,9, 13, 17,21,25
Escherichia coli
2,6, 10, 14, 18,22,26
Staphylcoccus aureus
3,7, 11, 15, 19,23,27
Alcaligenes faecalis*
4,8, 12, 16,20,24,28
Saccharomyces cerevisiae**
2. Incubate the tubes of E. coli, S. aureus, and A. fae-
calis at 37° C for 48 hours. Incubate the tubes of
S. ureae, C. glabrata, and S. cervisiae at 20° C for
48 to 72 hours.
Second Period
Materials:
spectrophotometer
1 tube of sterile nutrient broth
1
2
tubes of incubated cultures at various pHs
Use the tube of sterile broth to calibrate the spec-
trophotometer and measure the %T of each cul-
ture (page 98, Exercise 23). Record your results in
the tables on the Laboratory Report.
Plot the O.D. values in the graph on the
Laboratory Report and answer all the questions.
*Sporosarcina ureae can be used as a substitute for Alcaligenes
faecalis.
**Candida glabrata is a good substitute for Saccharomyces cere-
visiae.
134
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
VII. Environmental
Influences and Control of
Microbial Growth
37. Osmotic Pressure and
Bacterial Growth
© The McGraw-H
Companies, 2001
Osmotic Pressure and Bacterial Growth
37
Growth of bacteria can be profoundly affected by the
amount of water entering or leaving the cell. When
the medium surrounding an organism is hypotonic
(low solute content), a resultant higher osmotic pres-
sure occurs in the cell. Except for some marine
forms, this situation is not harmful to most bacteria.
The cell wall structure of most bacteria is so strong
and rigid that even slight cellular swelling is gener-
ally inapparent.
In the reverse situation, however, when bacteria
are placed in a hypertonic solution (high solute
content), their growth may be considerably inhib-
ited. The degree of inhibition will depend on the
type of solute and the nature of the organism. In me-
dia of growth-inhibiting osmotic pressure, the cyto-
plasm becomes dehydrated and shrinks away from
the cell wall. Such plasmolyzed cells are often sim-
ply inhibited in the absence of sufficient cellular
water and return to normal when placed in an iso-
tonic solution. In other instances, the organisms are
irreversibly affected due to permanent inactivation
of enzyme systems.
■
-■
■
■
■
J
f
! 1 1 "■
■ ■ - ■ - L\ ■ r
■
■
■
r m 11"
■ r V '
■
*
■
1
■
P . L I . ., I . 1 . . \ I .. ¥ I ■ ,P . ■ I .
,*-
Hypotonic Isotonic
Figure 37.1 Osmotic variabilities
Hypertonic
used differ in their tolerance of salt concentrations.
The salt concentrations will be 0.5, 5, 10, and 15%.
After incubation for 48 hours and several more days,
comparisons will be made of growth differences to de-
termine their degrees of salt tolerances.
Materials:
per student:
1 Petri plate of nutrient agar (0.5% NaCl)
1 Petri plate of nutrient agar (5% NaCl)
1 Petri plate of nutrient agar ( 1 0% NaCl)
1 Petri plate of milk salt agar (15% NaCl)
cultures:
Escherichia coli (nutrient broth)
Staphylococcus aureus (nutrient broth)
Halobacterium salinarium (slant culture)
1
2
3
4
Mark the bottoms of the four Petri plates as indi-
cated in figure 37.2.
Streak each organism in a straight line on the agar,
using a wire loop.
Incubate all the plates for 48 hours at room tem-
perature with exposure to light (the pigmentation
of H. salinarium requires light to develop).
Record your results on the Laboratory Report.
Continue the incubation of the milk salt agar plate
for several more days in the same manner, and
record your results again on the first portion of
Laboratory Report 37, 38.
Organisms that thrive in hypertonic solutions are
designated as halophiles or osmophiles. If they re-
quire minimum concentrations of salt (NaCl and other
cations and anions) they are called halophiles.
Obligate halophiles require a minimum of 13%
sodium chloride. Osmophiles, on the other hand, re-
quire high concentrations of an organic solute, such as
sugar.
In this exercise we will test the degree of inhibi-
tion of organisms that results with media containing
different concentrations of sodium chloride. To ac-
complish this, you will streak three different organ-
isms on four plates of media. The specific organisms
S. aureus
E. coti
H. salinarium
I ■■ ^Vdb
Figure 37.2 Streak pattern
135
Benson: Microbiological
VII. Environmental
38. Oligodynamic Action
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Influences and Control of
Microbial Growth
Companies, 2001
38
Oligodynamic Action
The ability of small amounts of heavy metals to exert a
lethal effect on bacteria is designated as oligodynamic
action (Greek: oligos, small; dynamis, power). The ef-
fectiveness of these small amounts of metal is probably
due to the high affinity of cellular proteins for the
metallic ions. Although the concentration of ions in so-
lution may be miniscule (a few parts per million), cells
die due to the cumulative effects of ions within the cell.
The success of silver amalgam fillings to prevent
secondary dental decay in teeth over long periods of
time is due to the small amounts of silver and mercury
ions that diffuse into adjacent tooth dentin. Its success in
this respect has led to much debated concern that its tox-
icity may cause long-term injury to patients. In addition
to its value (or harm) as a dental restoration material,
oligodynamic action of certain other heavy metals has
been applied to water purification, ointment manufac-
ture, and the treatment of bandages and fabrics.
In this exercise we will compare the oligody-
namic action of three metals (copper, silver, and alu-
minum) to note the differences.
Materials:
1 Petri plate
1 nutrient agar pour
forceps and Bunsen burner
acid- alcohol
1
2
3
4
broth culture of E. coli and S. aureus
3 metallic disks (copper, silver, aluminum)
water bath at student station (beaker of water
and electric hot plate)
Liquefy a tube of nutrient agar, cool to 50° C, and
inoculate with either E. coli or S. aureus (odd: E.
coli; even: S. aureus).
Pour half of the medium from each tube into a
sterile Petri plate and leave the other half in a wa-
ter bath (50° C). Allow agar to solidify in the
plate.
Clean three metallic disks, one at a time, and
place them on the agar, evenly spaced, as soon as
they are cleaned. Use this routine:
• Wash first with soap and water; then rinse with
water.
• With flamed forceps dip in acid- alcohol and
rinse with distilled water.
Pour the remaining seeded agar from the tube over
the metal disks. Incubate for 48 hours at 37° C.
Laboratory Report
After incubation compare the zones of inhibition and
record your results on the last portion of Laboratory
Report 37, 38.
136
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
VII. Environmental
Influences and Control of
Microbial Growth
39. Ultraviolet Light: Lethal
Effects
© The McGraw-H
Companies, 2001
Ultraviolet Light:
Lethal Effects
39
Except for the photo synthetic bacteria, most bacteria
are harmed by ultraviolet radiation. Those that con-
tain photosynthetic pigments require exposure to sun-
light in order to synthesize substances needed in their
metabolism. Although sunlight contains the complete
spectrum of short to long wavelengths of light, it is
only the short, invisible ultraviolet wavelengths that
are injurious to the no npho to synthetic bacteria.
Wavelengths of light may be expressed in nano-
meters (nm) or angstrom units (A). The angstrom unit
is equal to 10 -8 cm. In terms of nanometers, 10A equal
-8
one nanometer. Thus, a wavelength of 4500 X 10~ cm
would be expressed as 4500A, 450 nm, or 0.45 urn.
Figure 39.1 illustrates the relationship of ultravi-
olet to other types of radiations. By definition, ultra-
violet light includes electromagnetic radiations that
fall in the wavelength band between 40 and 4000A. It
bridges the gap between the X rays and the shortest
wavelengths of light visible to the human eye. The
visible range is approximately between 4000 and
7 800 A. Actually, the practical range of ultraviolet, as
far as we are concerned, lies between 2000 and
4000A. The "extreme" range (40-2000 A) includes ra-
diations that are absorbed by air and consequently
function only in a vacuum. This region is also referred
to as vacuum ultraviolet.
Ultraviolet is not a single entity, but is a very wide
band of wavelengths. This fact is often not realized.
Extending from 40 to 4000A, it encompasses a span
of 1:100; visible wavelengths (4000-7800A), on the
other hand, represent only a twofold spread.
The germicidal effects of the ultraviolet are lim-
ited to only a specific region of the ultraviolet spec-
trum. As indicated in figure 39.1, the most effective
wavelength is 2650A. Low-pressure mercury vapor
lamps, which have a high output (90%) of 2437A,
make very effective bactericidal lamps.
In this exercise organisms that have been spread
on nutrient agar will be exposed to ultraviolet radia-
tion for various lengths of time to determine the min-
imum amount of exposure required to effect a 100%
kill. One-half of each plate will be shielded from the
radiation to provide a control comparison. Bacillus
megaterium, a spore-former, and Staphylococcus au-
reus, a non- spore- former, will be used to provide a
comparison of the relative resistance of vegetative and
spore types.
Exposure to ultraviolet light may be accom-
plished with a lamp as shown in figure 39.2 or with a
UV box that has built-in ultraviolet lamps. The UV
exposure effectiveness varies with the type of setup
used. The exposure times given in table 39.1 work
well for a specific type of mercury arc lamp. Note in
the table that space is provided under the times for
adding in different timing. Your instructor will inform
you as to whether you should write in new times that
will be more suited to the equipment in your lab.
Proceed as follows to do this experiment.
100-
ULTRAVIOLET
VISIBLE
INFRARED
Q
LU
*:
LU
<
LU
O
LT
LU
Q_
0-
40A
2000A
4000A
6000A
8000A
10,000A
2650A
Figure 39.1 Lethal effectiveness of ultraviolet light
■
137
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
VII. Environmental
Influences and Control of
Microbial Growth
39. Ultraviolet Light: Lethal
Effects
© The McGraw-H
Companies, 2001
Exercise 39 • Ultraviolet Light: Lethal Effects
Materials:
Petri plates of nutrient agar (one or more per
student)
ultraviolet lamp or UV exposure box
timers (bell type)
cards (3" X 5")
nutrient broth cultures of S. aureus with swabs
1
2
3
saline suspensions of B. megaterium with swabs
Refer to table 39.1 to determine which organism
you will work with. You may be assigned more
than one plate to inoculate. If different times are
to be used, your instructor will inform you what
times to write in. Since there are only 16 assign-
ment numbers in the table, more student assign-
ment numbers can be written in as designated by
your instructor.
Label the bottoms of the plates with your assign-
ment number and your initials.
Using a cotton-tipped swab that is in the culture
tube, swab the entire surface of the agar in each
plate. Before swabbing, express the excess culture
from the swab against the inner wall of the tube.
4. Place the plates under the ultraviolet lamp with the
lids removed. Cover half of each plate with a 3" X
5" card as shown in figure 39.2. Note that if your
number is 8 or 16, you will not remove the lid from
your plate. The purpose of this exposure is to see to
what extent, if any, UV light can penetrate plastic.
CAUTION
Avoid looking directly into the ultraviolet lamp. These rays
can cause cataracts and other eye injury.
5. After exposing the plates for the correct time du-
rations, re-cover them with their lids, and incu-
bate them inverted at 37° C for 48 hours.
Laboratory Report
Record your observations on the Laboratory Report
and answer all the questions.
Table 39.1 Student Inoculation Assignments
Exposure Times
(Student Assignments)
S. aureus
1
2
3
4
5
6
7
8
10 sec
20 sec
40 sec
80 sec
2.5 min
5 min
10 min
20 min*
B. megaterium
9
10
11
12
13
14
15
16
1 min
2 min
4 min
8 min
15 min
30 min
60 min
60 min*
These Petri plates will be covered with dish covers during exposure.
Figure 39.2 Plates are exposed to UV light with 50% coverage
138
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
VII. Environmental
Influences and Control of
Microbial Growth
40. Evaluation of
Disinfectants: The
Use-Dilution Method
© The McGraw-H
Companies, 2001
Evaluation of Disinfectants
Tke Use-Dilution Method
When considering the relative effectiveness of differ-
ent chemical agents against bacteria, some yardstick of
comparison is necessary. Many different methods
have been developed over the years since Robert
Koch, in 1881, worked out the first scientific proce-
dure by measuring the killing power of various germi-
cides on silk threads that were impregnated with
spores of Bacillus anthracis. Koch's method and many
that followed proved unreliable for various reasons.
Finally, in 1931, the United States Food and Drug
Administration adopted a method that was a modifi-
cation of a test developed in England in 1903 by
Rideal and Walker. In 1950 the Association of
Official Agricultural Chemists adopted it as the offi-
cial method of testing disinfectants. This method
compares the effectiveness of various agents with
phenol. A value called the phenol coefficient is ar-
rived at that has significant meaning with certain lim-
itations. The restrictions are that the test should be
used only for phenol-like compounds that do not ex-
ert bacteriostatic effects and are not neutralized by the
subculture media used. Many excellent disinfectants
cannot be evaluated with this test. Disinfectants such
as bichloride of mercury, iodine, metaphen, and qua-
ternary detergents are unlike phenol in their germici-
dal properties and should not be evaluated in terms of
phenol coefficients. Notwithstanding, however, many
pharmaceutical companies have applied this test to
such disinfectants with misleading results. A more
suitable test for these nonphenolic disinfectants is the
use-dilution method.
The use-dilution method makes use of small
glass rods on which test organisms are dried for 30
minutes. The seeded rods are then exposed to the test
solutions at 20° C for 1, 5, 10, and 30 minutes, rinsed
with water or neutralizing solution, and transferred to
the tubes of media. After incubation at 37° C for 48
hours, the tubes are examined for growth. When the
results of this test are applied to practical conditions
of use, they are found to be completely reliable.
MMMMMk
+****^^^^*mm#**mr*****m
Common Pins
Bacterial Culture
Pins Dried on
Filter Paper
J*^ - "f
■**.'* "TW.
r" *
!■ ' - ^"" T"L
K *■ k " '
' ■ * t ■ . ■m J *Sm
-fl^_ *
• ■■■ u u u ■ ■ ~ " ^h3
*:•"■ -.StH
■ 1^^ a
:- vJ
" -^M
■ .m juyl
iJfli" ■
" '■ "■ ?C j\X
:/■- ViSI
"■■■': -^1
P - -"..
•■■*- ^Jf
ITii ■
: *J
::::::■
-i,
Disinfection
(Timed)
Neutralization
ncubation
(Broth)
II M II II M II II III I I II II II II II
******
Figure 40.1 Procedure for use-dilution evaluation of a disinfectant
139
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
VII. Environmental
Influences and Control of
Microbial Growth
40. Evaluation of
Disinfectants: The
Use-Dilution Method
© The McGraw-H
Companies, 2001
Exercise 40 • Evalation of Disinfectants: The Use-Dilution Method
In this experiment you will follow a modified pro-
cedure of the use- dilution method to compare the rel-
ative merits of three different disinfectants on two
kinds of bacteria: a spore-former, Bacillus mega-
terium, and a non-spore-former, Staphylococcus au-
reus. Instead of glass rods, we will use rustproof com-
mon pins.
Since each student will be performing only a
small portion of the entire experiment it will be nec-
essary to make student assignments with respect to
agents used and timing. Table 40.1 indicates, accord-
ing to student numbers, which agent each student will
be working with, and the length of time to apply the
agent to the pin. Note that a blank column is provided
for write-in substitutions. Proceed as follows:
Materials:
per student:
2 tubes of one of the following agents:
1:750 Zephiran for students 1, 4, 7, 10, 13,
16, 19, 22, 25, 28
5% phenol for students 2, 5, 8, 11, 14, 17,
20, 23, 26, 29
8% formaldehyde for students 3, 6, 9, 12,
15, 18,21,24,27,30
2 tubes of sterile water (about 7 ml each)
2 tubes of nutrient broth (about 7 ml each)
forceps
on demonstration table:
1 nutrient broth culture of S. aureus
1 physiological saline suspension of a 48 hour
nutrient agar slant culture of B. megaterium
2 sterile Petri plates with filter paper in bottom
several forceps and Bunsen burner
2 test tubes containing 36 sterile common pins in
each one (pins must be plated brass, which
are rustproof)
1. Consult table 40.1 or the materials list to deter-
mine which disinfectant you are to use.
2. Get two tubes of the disinfectant, two tubes of
sterile water, and two tubes of nutrient broth from
3
4
5
6
7
the table. Label one of each pair B. megaterium
and the other S. aureus.
Instructor: While the students are getting their
supplies together, you can start the experiment by
pouring the broth culture of S. aureus into one of
the tubes of pins and the saline suspension of B.
megaterium into the other tube of pins. After de-
canting the organisms into a beaker of disinfec-
tant, the pins are deposited onto filter paper in
separate Petri plates to dry. Plates should be
clearly labeled as to contents. Allow a few min-
utes for the pins to dry before allowing students to
take them. Make certain, also, that a Bunsen
burner and forceps are set up near the two dishes
of pins.
Gently flame a pair of forceps, let cool, and trans-
fer one pin from each Petri plate to the separate
tubes of disinfectant. Be sure to put them into the
right tubes.
Leave the pins in the disinfectant for the length of
time indicated in table 40.1. Find your number
under the time indicated for your disinfectant.
At the end of the assigned time, flame the mouths
of the tubes of disinfectant and carefully pour the
disinfectant into the sink without discarding the
pins. Then, transfer the pins into separate tubes of
sterile water. Avoid transferring any of the disin-
fectant to the water tubes with the pins.
After 1 minute in the tubes of water, flame the
mouths of the water and broth tubes, pour off the
water, and shake the pins out of the emptied tubes
into separate, labeled tubes of nutrient broth.
Instructor: At this point the instructor, or a des-
ignated class member, should put one pin from
each of the Petri plates into separate labeled tubes
of nutrient broth to be used as positive controls
for each organism.
Incubate all nutrient broth tubes with pins for 48
hours at 37° C. Examine them and record your re-
sults on the Laboratory Report.
Table 40.1 Student Assignments for Agents and Timing
Disinfectant
Time in Minutes
Substitution
1
5
10
30
60
1:750 Zephiran
1,16
4, 19
7,22
10,25
13,28
5% Phenol
2,17
5,20
8,23
11,26
14,29
8% Formaldehyde
3,18
6,21
9,24
12, 27
15,30
140
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
VII. Environmental
Influences and Control of
Microbial Growth
41. Evaluation of Alcohol:
Its Effectiveness as a Skin
Degerming Agent
© The McGraw-H
Companies, 2001
Evaluation of Alcohol:
Its Effectiveness as a Skin Degerming Agent
41
As a skin disinfectant, 70% alcohol is undoubtedly the
most widely used agent. The ubiquitous prepackaged
alcohol swabs used by nurses and technicians are evi-
dence that these items are indispensible. The question
that often arises is: How really effective is alcohol in
routine use? When the skin is swabbed prior to pene-
tration, are all, or mostly all, of the surface bacteria
killed? To determine alcohol effectiveness, as it might
be used in routine skin disinfection, we are going to
perform a very simple experiment here that utilizes
four thumbprints and a plate of enriched agar. Class re-
sults will be pooled to arrive at a statistical analysis.
Figure 41.1 illustrates the various steps in this
test. Note that the Petri plate is divided into four parts.
On the left side of the plate an unwashed left thumb is
first pressed down on the agar in the lower quadrant
of the plate. Next the left thumb is pressed down on
the upper left quadrant. With the left thumb we are try-
ing to establish the percentage of bacteria that are re-
moved by simple contact with the agar.
On the right side of the plate an unwashed right
thumb is pressed down on the lower right quadrant of
the plate. The next step is to either dip the right thumb
into alcohol or to scrub it with an alcohol swab and
dry it. Half of the class will use the dipping method
and the other half will use alcohol swabs. Your in-
structor will indicate what your assignment will be.
The last step is to press the dried right thumb on the
upper right quadrant of the plate.
After inoculating the plate it is incubated at 37° C
for 24-48 hours. Colony counts will establish the ef-
fectiveness of the alcohol.
Without touching any other surface the
left thumb is pressed against the agar
in quadrant B.
B
The pad of the unwashed left thumb is
momentarily pressed against the agar
in quadrant A.
The pad of the treated right thumb is
pressed against the agar in the D
quadrant.
The alcohol-treated right thumb is
allowed to completely air-dry.
D
The pad of the right thumb is
immersed in 70% alcohol or scrubbed
with an alcohol swab for 10 seconds.
The pad of the unwashed right thumb
is momentarily pressed against the
agar in quadrant C.
Figure 41 .1 Procedure for testing the effectiveness of alcohol on the skin
141
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
VII. Environmental
Influences and Control of
Microbial Growth
41. Evaluation of Alcohol:
Its Effectiveness as a Skin
Degerming Agent
© The McGraw-H
Companies, 2001
ExerCJS6 41 • Evaluation of Alcohol: Its Effectiveness as a Skin Degerming Agent
Materials:
1 Petri plate of veal infusion agar
small beaker
70% ethanol
alcohol swab
1
2
3
Perform this experiment with unwashed hands.
With a china marking pencil, mark the bottom of
the Petri plate with two perpendicular lines that
divide it into four quadrants. Label the left quad-
rants A and B and the right quadrants C and D as
shown in figure 41.1. {Keep in mind that when
you turn the plates over to label them, the A and
B quadrants will be on the right and C and D will
be on the left.)
Press the pad of your left thumb against the agar
surface in the A quadrant.
4. Without touching any other surface, press the left
thumb into the B quadrant.
5. Press the pad of your right thumb against the agar
surface of the C quadrant.
6. Disinfect the right thumb by one of the two fol-
lowing methods:
• dip the thumb into a beaker of 70% ethanol for
5 seconds, or
• scrub the entire pad surface of the right thumb
with an alcohol swab.
7. Allow the alcohol to completely evaporate from
the skin.
8. Press the right thumb against the agar in the D
quadrant.
9. Incubate the plate at 37° C for 24-48 hours.
10. Follow the instructions on the Laboratory Report
for evaluating the plate and answer all of the
questions.
142
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
VII. Environmental
Influences and Control of
Microbial Growth
42. Evaluation of
Antiseptics: The Filter
Paper Disk Method
© The McGraw-H
Companies, 2001
Evaluation of Antiseptics:
Tke Filter Paper Disk Method
42
The term antiseptic has, unfortunately, been some-
what ill-defined. Originally, the term was applied to
any agent that prevents sepsis, or putrefaction. Since
sepsis is caused by growing microorganisms, it fol-
lows that an antiseptic inhibits microbial multiplica-
tion without necessarily killing them. By this defini-
tion, we can assume that antiseptics are essentially
bacteriostatic agents. Part of the confusion that has re-
sulted in its definition is that the United States Food
and Drug Administration rates antiseptics essentially
the same as disinfectants. Only when an agent is to be
used in contact with the body for a long period of time
do they rate its bacteriostatic properties instead of its
bactericidal properties.
If we are to compare antiseptics on the basis of their
bacteriostatic properties, the filter paper disk method
(figure 42.1) is a simple, satisfactory method to use. In
this method a disk of filter paper QA" diameter) is im-
pregnated with the chemical agent and placed on a
seeded nutrient agar plate. The plate is incubated for 48
hours. If the substance is inhibitory, a clear zone of in-
hibition will surround the disk. The size of this zone is
an expression of the agent's effectiveness and can be
compared quantitatively against other substances.
In this exercise we will measure the relative ef-
fectiveness of three agents (phenol, formaldehyde,
and iodine) against two organisms: Staphylococcus
aureus (gram-positive) and Pseudomonas aeruginosa
(gram-negative). Table 42.1 will be used to assign
each student one chemical agent to be tested against
one organism. Note that space has been provided in
the table for different agents to be written in as substi-
tutes for the three agents listed. Your instructor may
wish to make substitutions. Proceed as follows:
Liquefied nutrient agar is inoculated
with one Joopful of organisms.
Seeded nutrient agar is poured into
plate and allowed to solidify.
Sterile disk is dipped halfway into
agent. If completely submerged it
will be too wet.
r -ui M 'i»t^.- .•»' rf w 'hi» »i n '^ i i ii r. !!■■
■ ^ hi ^ ^ liW
mpregnated disk is placed in cen-
ter of nutrient agar and pressed
down lightly to secure it.
After 24-48 hours incubation the zone of inhibition is
measured on bottom of plate. Note that measurement
is between disk edge and growth.
Figure 42.1 Filter paper disk method of evaluating an antiseptic
143
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
VII. Environmental
Influences and Control of
Microbial Growth
42. Evaluation of
Antiseptics: The Filter
Paper Disk Method
© The McGraw-H
Companies, 2001
Exercise 42 • Evaluation of Antiseptics: The Filter Paper Disk Method
First Period
(Disk Application)
Materials:
per student:
1 nutrient agar pour and 1 Petri plate
broth culture of S. aureus or P. aeruginosa on
demonstration table:
Petri dish containing sterile disks of filter paper
QA" dia)
forceps and Bunsen burner
chemical agents in small beakers (5% phenol,
5% formaldehyde, 5% aqueous iodine)
1 . Consult table 42.1 to determine your assignment.
2. Liquefy a nutrient agar pour in a water bath and
cool to 50° C.
3. Label the bottom of a Petri plate with the names
of the organism and chemical agent.
4. Inoculate the agar pour with one loopful of the or-
ganism and pour into the plate.
5
6
1
2
After the medium has solidified in the plate, pick
up a sterile disk with lightly flamed forceps, dip
the disk halfway into a beaker of the chemical
agent, and place the disk in the center of the
medium.
To secure the disk to the medium, press
lightly on it with the forceps.
Incubate the plate at 37° C for 48 hours.
Second Period
(Evaluation)
Measure the zone of inhibition from the edge of
the disk to the edge of the growth (see illustration
5, figure 42.1).
Exchange plates with other members of the class
so that you will have an opportunity to complete
the table on the Laboratory Report.
Table 42.1 Student Assignments
Chemical Agent
Student Number
Substitution
S. aureus
P. aeruginosa
5% Phenol
1,7, 13,19,25
2, 8, 1 4, 20, 26
5% Formaldehyde
3,9, 15,21,27
4,10, 16,22,28
5% Iodine
5, 11,17,23,29
6,12, 18,24,30
144
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
VII. Environmental
Influences and Control of
Microbial Growth
43. Antimicrobic Sensitivity
Testing: The Kirby-Bauer
Method
© The McGraw-H
Companies, 2001
Antimicrobic Sensitivity Testing:
Tke Kirby-Bauer Method
43
Once the causative organism of a specific disease in a
patient has been isolated, it is up to the attending
physician to administer a chemotherapeutic agent that
will inhibit or kill the pathogen without causing seri-
ous harm to the individual. The method must be rela-
tively simple to use, be very reliable, and yield results
in as short a time as possible. The Kirby-Bauer method
of sensitivity testing is such a method. It is used for
testing both antibiotics and drugs. Antibiotics are
chemotherapeutic agents of low molecular weight pro-
duced by microorganisms that inhibit or kill other mi-
croorganisms. Drugs, on the other hand, are antimi-
crobic agents that are man-made. Both types of agents
will be tested in this laboratory session according to
the procedure shown in figure 43.1.
The effectiveness of an antimicrobic in sensitivity
testing is based on the size of the zone of inhibition
that surrounds a disk that has been impregnated with
a specific concentration of the agent. The zone of in-
hibition, however, varies with the diffusibility of the
agent, the size of the inoculum, the type of medium,
and many other factors. Only by taking all these vari-
ables into consideration can a reliable method be
worked out.
The Kirby-Bauer method is a standardized sys-
tem that takes all variables into consideration. It is
sanctioned by the U.S. FDA and the Subcommittee on
Antimicrobial Susceptibility Testing of the National
Committee for Clinical Laboratory Standards.
Although time is insufficient here to consider all
facets of this test, its basic procedure will be followed.
The recommended medium in this test is Mueller
Hinton II agar. Its pH should be between 7.2 and 7.4,
and it should be poured to a uniform thickness of 4
mm in the Petri plate. This requires 60 ml in a 150 mm
plate and 25 ml in a 1 00 mm plate. For certain fastid-
ious microorganisms, 5% defibrinated sheep blood is
added to the medium.
Inoculation of the surface of the medium is made
with a cotton swab from a broth culture. In clinical ap-
plications, the broth turbidity has to match a defined
standard. Care must also be taken to express excess
broth from the swab prior to inoculation.
High-potency disks are used that may be placed
on the agar with a mechanical dispenser or sterile for-
ceps. To secure the disks to the medium, it is neces-
sary to press them down onto the agar.
After 1 6 to 18 hours incubation the plates are ex-
amined and the diameters of the zones are measured
to the nearest millimeter. To determine the signifi-
cance of the zone diameters, one must consult a table
(Appendix A) .
In this exercise we will work with four microor-
ganisms: Staphylococcus aureus, Escherichia coli,
Proteus vulgaris, and Pseudomonas aeruginosa. Each
student will inoculate one plate with one of the four
organisms and place the disks on the medium by
whichever method is available. Since each student
will be doing only a portion of the total experiment,
student assignments will be made. Proceed as follows:
First Period
(Plate Preparation)
Materials:
1 Petri plate of Mueller-Hinton II agar
nutrient broth cultures (with swabs) of
S. aureus, E. coli, P. vulgaris, and
P. aeruginosa
disk dispenser (BBL or Difco)
cartridges of disks (BBL or Difco)
forceps and Bunsen burner
zone interpretation charts (Difco or BBL)
1. Select the organisms you are going to work with
from the following table.
Organism Student Number
S. aureus
1,5,9, 13, 17,21,25
E. coli
2,6, 10, 14, 18,22,26
P. vulgaris
3,7, 11, 15, 19,23,27
P. aeruginosa
4,8, 12, 16,20,24,28
2. Label your plate with the name of your organism.
3. Inoculate the surface of the medium with the
swab after expressing excess fluid from the swab
by pressing and rotating the swab against the in-
side walls of the tube above the fluid level. Cover
the surface of the agar evenly by swabbing in
three directions. A final sweep should be made of
the agar rim with the swab.
4. Allow 3 to 5 minutes for the agar surface to dry
before applying disks.
145
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
VII. Environmental
Influences and Control of
Microbial Growth
43. Antimicrobic Sensitivity
Testing: The Kirby-Bauer
Method
© The McGraw-H
Companies, 2001
Exercise 43 • Antimicrobic Sensitivity Testing: The Kirby-Bauer Method
5. Dispense disks as follows:
a. If an automatic dispenser is used, remove the
lid from the plate, place the dispenser over the
plate, and push down firmly on the plunger.
With the sterile tip of forceps, tap each disk
lightly to secure it to medium.
b. If forceps are used, sterilize them first by flaming
before picking up the disks. Keep each disk at
least 15 mm from the edge of the plate. Place no
more than 13 on a 150 mm plate, nor more than
5 on a 100 mm plate. Apply light pressure to each
disk on the agar with the tip of a sterile forceps
or inoculating loop to secure it to medium.
6. Invert and incubate the plate for 16 to 1 8 hours at
37° C.
Second Period
(Interpretation)
After incubation, measure the zone diameters with a
metric ruler to the nearest whole millimeter. The zone
of complete inhibition is determined without magnifi-
cation. Ignore faint growth or tiny colonies that can be
detected by very close scrutiny. Large colonies grow-
ing within the clear zone might represent resistant
variants or a mixed inoculum and may require reiden-
tification and retesting in clinical situations. Ignore
the "swarming" characteristics of Proteus, measuring
only to the margin of heavy growth.
Record the zone measurements on the table of the
Laboratory Report and on the chart on the demonstra-
tion table, which has been provided by the instructor.
Use table 43.1 or 43.2 for identifying the various
disks. Although BBL and Difco use essentially the
same code numbers, there are slight differences in the
two charts. Careful comparison of the charts will re-
veal that each company has certain antibiotics that are
not listed by the other company.
To determine which antibiotics your organism is
sensitive to (S), or resistant to (R), or intermediate (I),
consult Table VII in Appendix A. It is important to
note that the significance of a zone of inhibition varies
with the type of organism. If you cannot find your an-
tibiotic on the chart, consult a chart that is supplied by
BBL or Difco that is on the demonstration table or
bulletin board. Table VII is incomplete.
Table 43.1
Code for BBL Disks
AMD-10
Amdinocillin
E-15
Erythromycin
AN-30
Amikacin
GM-120
Gentamicin
AmC-30
Amoxicillin/
IPM-10
Imipenem
Clavulanic Acid
K-30
Kanamycin
AM-10
Ampicillin
LOM-10
Lomefloxacin
SAM-20
Ampicillin/
LOR-30
Loracarbef
Sulbactam
DP-5
Methicillin
AZM-15
Azithromycin
MZ-75
Meziocillin
AZ-75
Azlocillin
Ml -30
Minocycline
ATM -30
Aztreonam
MOX-30
Moxalactam
B-10
Bacitracin
NF-1
Nafcillin
CB-100
Carbenicillin
NA-30
Nalidixic Acid
CEC-30
Cefactor
N-30
Neomycin
MA-30
Cefamandole
NET-30
Netilmicin
CZ-30
Cefazolin
NOR-10
Norfloxacin
CFM-5
Cefixime
NB-30
Novobiocin
CMZ-30
Cefmetrazole
OFX-5
Ofloxacin
CID-30
Cefonicid
OX-1
Oxacillin
CFP-75
Cefoperazone
OA-2
Oxolinic Acid
CTX-30
Cefotaxime
P-10
Penicillin
CTT-30
Cefotetan
PIP-100
Piperacillin
FOX-30
Cefoxitin
PB-300
Polymyxin B
CPD-10
Cefpodoxime
RA-5
Rifampin
CPR-30
Cefprozil
SPT-1 00
Spectinomycin
CAZ-30
Ceftazidime
S-300
Streptomycin
ZOX-30
Ceftizoxime
G-25
Sulfisoxazole
CRO-30
Ceftriaxone
Te-30
Tetracycline
CXM-30
Cefuroxime
TIC-75
Ticarcillin
CF-30
Cephalothin
TIM-85
Ticarcillin/
C-30
Chloramphenicol
Clavulanic Acid
CIN-100
Cinoxacin
NN-10
Tobramycin
CIP-5
Ciprofloxacin
TMP-5
Trimethoprim
CLR-15
Clarithromycin
SXT
Trimethoprim/
CC-2
Clindamycin
Sulfamethoxazole
CL-10
Colistin
Va-30
Vancomycin
Table 43.2
Code for Difco Disks
AN 30
Amikacin
E15
Erythromycin
AMC30
Amoxicillin/
FLX5
Fleroxacin
Clavulanic Acid
GM 10
Gentamycin
AM 10
Ampicillin
IPM10
Imipenem
SAM 20
Ampicillin/
K30
Kanamycin
Sulbactam
LOM 10
Lomefloxacin
AZM 15
Azithromycin
LOR 30
Loracarbef
AZ75
Azlocillin
MZ75
Meziocillin
ATM 30
Aztreonam
Mi 30
Minocycline
CB100
Carbenicillin
MOX30
Moxalactam
CEC30
Cefactor
NF1
Nafcillin
MA 30
Cefamandole
NA30
Nalidixic Acid
CZ30
Cefazolin
NET 30
Netilmicin
FEP30
Cefepime
FD300
Nitrofurantoin
CAT 10
Cefetamet
NOR 10
Norfloxacin
CFM 5
Cefixime
OFX5
Ofloxacin
CMZ30
Cefmetrazole
P-10
Penicillin G
CID30
Cefonicid
PTZ110
Piperacillin/
CFP75
Cefoperazone
Tazobactam
CTX30
Cefotaxime
RA5
Rifampin
CTT30
Cefotetan
S10
Streptomycin
FOX 30
Cefoxitin
G300
Sulfisoxazole
CPD10
Cefpodoxime
TEC 30
Telcoplanin
CPR30
Cefprozil
TE30
Tetracycline
CAZ30
Ceftazidime
TIC 75
Ticarcillin
OX 30
Ceftizoxime
TIM 85
Ticarcillin/
CRO30
Ceftriaxone
Clavulanic Acid
CXM30
Cefuroxime
TN 10
Tobramycin
CF30
Cephalothin
TMP5
Trimethoprim
C30
Chloramphenicol
SxT
Trimethorprim/
CIN 100
Cinoxacin
Sulfamethoxazole
CLR15
Clarithromycin
VA30
Vancomycin
CC2
Clindamycin
D30
Doxycycline
ENX10
Enoxacin
146
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
VII. Environmental
Influences and Control of
Microbial Growth
43. Antimicrobic Sensitivity
Testing: The Kirby-Bauer
Method
© The McGraw-H
Companies, 2001
Antimicrobic Sensitivity Testing: The Kirby-Bauer Method • Exercise 43
The entire surface of a plate of nutrient
medium is swabbed with organism to
be tested.
Handle of dispenser is pushed down to place 12
disks on the medium. In addition to dispensing
disks, this dispenser also tamps disks onto
medium.
Cartridges (Difco) can be used to dispense
individual disks. Only 4 or 5 disks should be
placed on small (1 00 mm.) plates.
After 18 hours incubation, the zones of inhibition
(diameters) are measured in millimeters.
Significance of zones is determined from Kirby-
Bauer chart.
Figure 43.1 Antimicrobic sensitivity testing
147
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
VII. Environmental
Influences and Control of
Microbial Growth
44. Effectiveness of Hand
Scrubbing
© The McGraw-H
Companies, 2001
44
Effectiveness of Hand Scrubbing
The importance of hand disinfection in preventing
the spread of disease is accredited to the observa-
tions of Semmelweis at the Lying-in Hospital in
Vienna in 1846 and 1847. He noted that the number
of cases of puerperal fever was closely related to the
practice of sanitary methods. Until he took over his
assignment in this hospital, it was customary for
medical students to go directly from the autopsy
room to a patient's bedside and assist in deliveries
without scrubbing and disinfecting their hands.
When the medical students were on vacation, only
the nurses, who were not permitted in the autopsy
room, attended the patients. Semmelweis noted that
during this time, deaths due to puerperal fever fell
off markedly.
As a result of his observations, he established a
policy that no medical students would be allowed to
examine obstetric patients or assist in deliveries until
they had cleansed their hands with a solution of chlo-
ride of lime. This ruling caused the death rate from
puerperal infections to drop from 12% to 1.27% in
one year.
Today it is routine practice to wash hands prior
to the examination of any patient and to do a com-
plete surgical scrub prior to surgery. Scrubbing the
hands involves the removal of transient (contami-
nant) and resident microorganisms. Depending on
the condition of the skin and the numbers of bacteria
present, it takes from 7 to 8 minutes of washing with
soap and water to remove all transients, and they can
be killed with relative ease using suitable antisep-
tics. Residents, on the other hand, are firmly en-
trenched and are removed slowly by washing. These
organisms, which consist primarily of staphylococci
of low pathogenicity, are less susceptible than the
transients to the action of antiseptics.
In this exercise, an attempt will be made to eval-
uate the effectiveness of the length of time in removal
of organisms from the hands using a surgical scrub
technique. One member of the class will be selected
to perform the scrub. Another student will assist by
supplying the soap, brushes, and basins, as needed.
During the scrub, at 2-minute intervals, the hands will
be scrubbed into a basin of sterile water. Bacterial
counts will be made of these basins to determine the
effectiveness of the previous 2-minute scrub in reduc-
ing the bacterial flora of the hands. Members of the
class not involved in the scrub procedure will make
the inoculations from the basins for the plate counts.
Scrub Procedure
The two members of the class who are chosen to
perform the surgical scrub will set up their materi-
als near a sink for convenience. As one student per-
forms the scrub, the other will assist in reading the
instructions and providing materials as needed. The
basic steps, which are illustrated in figure 44.1, are
also described in detail below. Before beginning
the scrub, both students should read all the steps
carefully.
Materials:
5 sterile surgical scrub brushes, individually
wrapped
5 basins (or 2000 ml beakers), containing 1000
ml each of sterile water. These basins should
be covered to prevent contamination
1 dispenser of green soap
1 tube of hand lotion
Step 1 To get some idea of the number of transient
organisms on the hands, the scrubber will scrub all
surfaces of each hand with a sterile surgical scrub
brush for 30 seconds into Basin A. No green soap will
be used for this step. The successful performance of
this step will depend on
• spending the same amount of time on each hand
(30 seconds),
• maintaining the same amount of activity on each
hand, and
• scrubbing under the fingernails, as well as work-
ing over their surfaces.
After completion of this 60-second scrub, notify
Group A that their basin is ready for the inoculations.
Step 2 Using the same brush as above, begin scrub-
bing with green soap for 2 minutes, using cool tap wa-
ter to moisten and rinse the hands. One minute is de-
voted to each hand.
The assistant will make one application of green
soap to each hand as it is being scrubbed.
148
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
VII. Environmental
Influences and Control of
Microbial Growth
44. Effectiveness of Hand
Scrubbing
© The McGraw-H
Companies, 2001
Effectiveness of Hand Scrubbing • Exercise 44
Sixty-second hand scrub into Basin
A. No soap.
Two-minute soap scrub with running
water.
Same as 2.
Sixty-second hand scrub into Basin
C. No soap.
Sixty-second hand scrub into
Basin D. No soap.
Same as 2.
Sixty-second hand scrub into
Basin B. No soap.
Same as 2
Sixty-second hand scrub into
Basin E. No soap.
Figure 44.1 Hand scrubbing routine
149
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
VII. Environmental
Influences and Control of
Microbial Growth
44. Effectiveness of Hand
Scrubbing
© The McGraw-H
Companies, 2001
Exercise 44 • Effectiveness of Hand Scrubbing
Rinse both hands for 5 seconds under tap water at
the completion of the scrub.
Discard the brush.
Note: This same procedure will be followed exactly
in steps 4, 6, and 8 of figure 44.1.
Step 3 With afresh sterile brush, scrub the hands
into Basin B in a manner that is identical to step 1.
Don't use soap. Notify Group B when this basin is
ready.
Note: Exactly the same procedure is used in steps 5,
7, and 9 of figure 44.1, using Basins C, D, and E.
Remember: It is important to use a fresh sterile brush
for the preparation of each of these basins.
After Scrubbing After all scrubbing has been com-
pleted, the scrubber should dry his or her hands and
apply hand lotion.
Making the Pour Plates
While the scrub is being performed, the rest of the
class will be divided into five groups (A, B, C, D, and
E) by the instructor. Each group will make six plate
inoculations from one of the five basins (A, B, C, D,
or E). It is the function of these groups to determine
the bacterial count per milliliter in each basin. In this
way we hope to determine, in a relative way, the ef-
fectiveness of scrubbing in bringing down the total
bacterial count of the skin.
Materials:
30 veal infusion agar pours — 6 per group
1 ml pipettes
30 sterile Petri plates — 6 per group
70% alcohol
1
2
3
4
L- shaped glass stirring rod (optional)
Liquefy six pours of veal infusion agar and cool
to 50° C. While the medium is being liquefied, la-
bel two plates each: 0.1 ml, 0.2 ml, and 0.4 ml.
Also, indicate your group designation on the
plate.
As soon as the scrubber has prepared your basin,
take it to your table and make your inoculations as
follows:
a. Stir the water in the basin with a pipette or an
L-shaped stirring rod for 15 seconds. If the
stirring rod is used (figure 44.2), sterilize it be-
fore using by immersing it in 70% alcohol and
flaming. For consistency of results all groups
should use the same method of stirring.
b. Deliver the proper amounts of water from the
basin to the six Petri plates with a sterile sero-
logical pipette. Refer to figure 44.3. If a pipette
was used for stirring, it may be used for the de-
liveries.
c. Pour a tube of veal infusion agar, cooled to
50° C, into each plate, rotate to get good dis-
tribution of organisms, and allow to cool.
d. Incubate the plates at 37° C for 24 hours.
After the plates have been incubated, select the
pair that has the best colony distribution with no
fewer than 30 or more than 300 colonies. Count
the colonies on the two plates and record your
counts on the chart on the chalkboard.
After all data are on the chalkboard, complete the
table and graph on the Laboratory Report.
Figure 44.2 An alternative method of stirring utilizes an
L-shaped glass stirring rod.
Figure 44.3 Scrub water for count is distributed to six
Petri plates in amounts as shown.
150
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
VIM. Identification of
Unknown Bacteria
Introduction
© The McGraw-H
Companies, 2001
Part
Identification of Unknown
Bacteria
One of the most interesting experiences in introductory microbiol-
ogy is to attempt to identify an unknown microorganism that has
been assigned to you as a laboratory problem. The next seven ex-
ercises pertain to this phase of microbiological work. You will be
given one or more cultures of bacteria to identify. The only infor-
mation that might be given to you about your unknowns will pertain
to their sources and habitats. All the information needed for identi-
fication will have to be acquired by you through independent study.
Although you will be engrossed in trying to identify an unknown
organism, there is a more fundamental underlying objective of this
series of exercises that goes far beyond simply identifying an un-
known. That objective is to gain an understanding of the cultural
and physiological characteristics of bacteria. Physiological charac-
teristics will be determined with a series of biochemical tests that
you will perform on the organisms. Although correctly identifying
the unknowns that are given to you is very important, it is just as
important that you thoroughly understand the chemistry of the
tests that you perform on the organisms.
The first step in the identification procedure is to accumulate in-
formation that pertains to the organisms' morphological, cultural,
and physiological (biochemical) characteristics. This involves mak-
ing different kinds of slides for cellular studies and the inoculation of
various types of media to note the growth characteristics and types
of enzymes produced. As this information is accumulated, it is
recorded in an orderly manner on Descriptive Charts, which are lo-
cated toward the back of the manual with the Laboratory Reports.
After sufficient information has been recorded, the next step is to
consult a taxonomic key, which enables one to identify the organism.
For this final step, Bergey's Manual of Systematic Bacteriology will be
used. Copies of volumes 1 and 2 of this book will be available in the
laboratory, library, or both. In addition, a CD-ROM computer simula-
tion program called Identibacter interactus may be available, which
can be used for identifying and reporting your unknown. Exercise 51
pertains to the use of Bergey's Manual and Identibacter interactus.
Success in this endeavor will require meticulous techniques, in-
telligent interpretation, and careful recordkeeping. Your mastery of
aseptic methods in the handling of cultures and the performance of
inoculations will show up clearly in your results. Contamination of
your cultures with unwanted organisms will yield false results, mak-
ing identification hazardous speculation. If you have reason to
doubt the validity of the results of a specific test, repeat it; don't rely
on chance! As soon as you have made an observation or com-
pleted a test, record the information on the Descriptive Chart. Do
not trust your memory — record data immediately.
151
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
VIM. Identification of
Unknown Bacteria
45. Preparation and Care of
Stock Cultures
© The McGraw-H
Companies, 2001
45
Preparation and Care of Stock Cultures
Your unknown cultures will be used for making many
different kinds of slides and inoculations. Despite
meticulous aseptic practice on your part, the chance of
contamination of these cultures increases with fre-
quency of use. If you were to attempt to make all your
inoculations from the single tube given to you, it is
very likely that somewhere along the way contamina-
tion would result.
Another problem that will arise is aging of the cul-
ture. Two or three weeks may be necessary for the per-
formance of all tests. In this period of time, the organ-
isms in the broth culture may die, particularly if the
culture is kept very long at room temperature. To ensure
against the hazards of contamination or death of your
organisms, it is essential that you prepare stock cultures
before any slides or routine inoculations are made.
Different types of organisms require different
kinds of stock media, but for those used in this unit,
nutrient agar slants will suffice. For each unknown,
you will inoculate two slants. One of these will be
your reserve stock and the other one will be your
working stock.
The reserve stock culture will not be used for
making slides or routine inoculations; instead, it will
be stored in the refrigerator after incubation until
some time later when a transfer may be made from it
to another reserve stock or working stock culture.
The working stock culture will be used for mak-
ing slides and routine inoculations. When it becomes
too old to use or has been damaged in some way, re-
place it with a fresh culture that is made from the re-
serve stock.
7^
T
20° C
Inoculate two nu-
trient agar slants from
the unknown broth cul-
ture. To inoculate make a
straight streak from the
bottom to the top of
the slant.
24 hours
Select the tube with best
growth for your reserve
stock, and designate its
temperature as the presumed
optimum growth temperature.
in - t>
7r=n
37° C
24 hours
Use the other tube for your
working stock culture. Pro-
vide additional incubation
necessary to get good growth
of this culture.
\J>
Figure 45.1 Stock culture procedure
152
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
VIM. Identification of
Unknown Bacteria
45. Preparation and Care of
Stock Cultures
© The McGraw-H
Companies, 2001
Preparation and Care of Stock Cultures • Exercise 45
Note in figure 45.1 that one slant will be incu-
bated at 20° C and the other at 37° C. This will enable
you to learn something about the optimum growth
temperature of your unknown, which will be pertinent
in Exercise 47. Proceed as follows:
First Period
Inoculate two nutrient agar slants from each of your
unknowns as follows:
Materials:
for each unknown:
2 nutrient agar slants (screw-cap type)
gummed labels
1
2
3
Label two slants with the code number of the un-
known and your initials. Use gummed labels.
Also, mark one tube 20° C and the other 37° C.
With a loop, inoculate each slant with a straight
streak from the bottom to the top. Since these
slants will be used for your cultural study in
Exercise 47, a straight streak is more useful than
one that is spread over the entire surface.
Place the two slants in separate baskets on the
demonstration table that are designated with la-
bels for the two temperatures (20° C and 37° C).
Although the 20° C temperature is thought of
as "room temperature," it should be incubated in
a biological incubator instead of leaving it out at
laboratory room temperature. Laboratory temper-
atures are often quite variable in a 24-hour period.
Second Period
After 24 hours incubation, evaluate the slants made
from each of your unknowns, as follows:
1 . Examine the slants to note the extent of growth.
Some organisms require close examination to see
the growth, especially if the growth is thin and
translucent.
2
3
4
5
6
7
8
Determine which temperature seems to promote
the best growth.
Record on the Descriptive Chart the presumed op-
timum temperature. (Obviously, this may not be
the actual optimum growth temperature, but for all
practical purposes, it will suffice for this exercise.)
If there is no growth visible on either slant, there
are several possible explanations:
• It may be that the culture you were issued was
not viable.
• Another possibility might be that the organism
grows too slowly to be visible at this time.
• Or, possibly, neither temperature was suitable!
Think through these possibilities and decide what
you should do to circumvent the problem.
Label the tube with the best growth reserve
stock. Label the other tube working stock.
If both tubes have good growth, place them in the
refrigerator until needed.
If one tube has very scanty growth, refrigerate the
good one (reserve stock) and incubate the other
one at the more desirable temperature for another
24 hours, then refrigerate.
Remember these points concerning your stock
cultures:
• Most stock cultures will keep for 4 weeks in
the refrigerator. Some fastidious pathogens
will survive for only a few days. Although
none of the organisms issued in this unit are of
the extremely delicate type, don't wait 4 weeks
to make a new reserve stock culture; instead,
make fresh transfers every 10 days.
• Don't use your reserve stock culture for mak-
ing slides or routine inoculations.
• Don't store either of your stock cultures in
your desk drawer or a cupboard. After the ini-
tial incubation period cultures must be refrig-
erated. After 2 or 3 days at room temperature,
cultures begin to deteriorate. Some die out
completely.
Answer the questions on the Laboratory Report.
153
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
VIM. Identification of
Unknown Bacteria
46. Morphological Study of
Unknown
© The McGraw-H
Companies, 2001
46
Morphological Study of Unknown
The first step in the identification of an unknown bac-
terial organism is to learn as much as possible about
its morphological characteristics. One needs to know
whether the organism is rod-, coccus-, or spiral-
shaped; whether or not it is pleomorphic; its reaction
to gram staining; and the presence or absence of en-
dospores, capsules, or granules. All this morphologi-
cal information provides a starting point in the cate-
gorization of an unknown.
Figure 46.1 illustrates the steps that will be fol-
lowed in determining morphological characteristics
of your unknown. Note that fresh broth and slant cul-
tures will be needed to make the various slides and
perform motility tests. Since most of the slide tech-
niques were covered in Part 3, you will find it neces-
sary to refer to that section from time to time. Note
that gram staining, motility testing, and measure-
ments will be made from the broth culture; gram
staining and other stained slides will also be made
from the agar slant. The rationale as to the choice of
broth or agar slants will be explained as each tech-
nique is performed.
As soon as morphological information is ac-
quired be sure to record your observations on the
Descriptive Chart at the back of the manual. Proceed
as follows:
Materials:
gram- staining kit
spore- staining kit
acid-fast staining kit
Loeffler's methylene blue stain
nigrosine or india ink
tubes of nutrient broth and nutrient agar
gummed labels for test tubes
Grams Stain
Since a good gram-stained slide will provide you with
more valuable information than any other slide, this is
the place to start. Make gram- stained slides from both
the broth and agar slants, and compare them under oil
immersion.
Two questions must be answered at this time: (1) Is
the organism gram-positive, or is it gram-negative? and
(2) Is the organism rod- or coccus-shaped? If your
staining technique is correct, you should have no prob-
lem with the Gram reaction. If the organism is a long
rod, the morphology question is easily settled; how-
ever, if your organism is a very short rod, you may in-
correctly decide it is coccus-shaped.
Keep in mind that short rods with round ends
(coccobacilli) look like cocci. If you have what
seems to be a coccobacillus, examine many cells be-
fore you make a final decision. Also, keep in mind
that while rod-shaped organisms frequently appear
as cocci under certain growth conditions, cocci
rarely appear as rods. {Streptococcus mutans is
unique in forming rods under certain conditions.)
Thus, it is generally safe to assume that if you have a
slide on which you see both coccuslike cells and
short rods, the organism is probably rod-shaped. This
assumption is valid, however, only if you are not
working with a contaminated culture !
Record the shape of the organism and its reaction
to the stain on the Descriptive Chart.
Cell Size
Once you have a good gram-stained slide, determine
the size of the organism with an ocular micrometer.
Refer to Exercise 5. If the size is variable, determine
the size range. Record this information on the
Descriptive Chart.
New Inoculations
For all of these staining techniques you will need
24-48 hour cultures of your unknown. If your work-
ing stock slant is a fresh culture, use it. If you don't
have a fresh broth culture of your unknown inoculate
a tube of nutrient broth and incubate it at its estimated
optimum temperature for 24 hours.
Motility and Cellular Arrangement
If your organism is a nonpathogen make a wet mount
or hanging drop slide from the broth culture. Refer to
Exercise 19. This will enable you to determine
whether the organism is motile, and it will allow you
to confirm the cellular arrangement. By making this
154
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
VIM. Identification of
Unknown Bacteria
46. Morphological Study of
Unknown
© The McGraw-H
Companies, 2001
Morphological Study of Unknown • Exercise 46
WORKING STOCK
CULTURE
Inoculate a nutrient broth and
a nutrient agar slant from your
working stock culture.
Incubate both tubes at the optimum
temperature for 24 hours.
K
* i
t.
I:
rar.;
NUTRIENT BROTH
Make a gram-stained slide and
perform the proper motility tests
from this broth culture.
NUTRIENT AGAR SLANT
Use organisms from this young
culture to make specialized
stained slides that might be
needed.
T
MOTILITY TESTS
GRAM-STAINED SLIDE
y
/
WET MOUNT SLIDE
If organism is a non-
pathogen make a wet
mount or hanging drop
slide.
MICROSCOPIC MEASUREMENTS
(see Exercise 5)
SEMISOLID MEDIUM
(for pathogens)
STAINED SLIDES
GRAM STAIN: Make a gram-
stained slide from the slant and
compare it with slide made from
nutrient broth.
SIMPLE STAIN: Use Loeffler's
methylene blue if metachro-matic
granules are suspected.
SPORE STAIN: If the organism is
a gram-positive rod, do a spore
stain.
ACID-FAST STAIN: If the or-
ganism is a gram-positive rod,
make an acid-fast stained slide.
Figure 46.1 Procedure for morphological study
155
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
VIM. Identification of
Unknown Bacteria
46. Morphological Study of
Unknown
© The McGraw-H
Companies, 2001
Exercise 46 • Morphological Study of Unknown
slide from broth instead of the agar slant, the cells will
be well dispersed in natural clumps. Note whether the
cells occur singly, in pairs, masses, or chains.
Remember to place the slide preparation in a beaker
of disinfectant when finished with it.
If your organism happens to be a pathogen do not
make a slide preparation of the organisms; instead,
stab the organism into a tube of semisolid or SIM
medium to determine motility (Exercise 19). Incubate
for 48 hours.
Be sure to record your observations on the
Descriptive Chart.
Endospores
If your unknown is a gram-positive rod, check for en-
dospores. Only rarely is a coccus or gram-negative rod
a spore-former. Examination of your gram-stained
slide made from the agar slant should provide a clue,
since endospores show up as transparent holes in
gram-stained spore-formers. Endospores can also be
seen on unstained organisms if studied with phase-
contrast optics.
If there seems to be evidence that the organism is
a spore- former, make a slide using one of the spore-
staining techniques you used in Exercise 16. Since
some spore-formers require at least a week's time of
incubation before forming spores, it is prudent to
double -check for spores in older cultures.
Record on the Descriptive Chart whether the
spore is terminal, subterminal, or in the middle of
the rod.
Acid- Fast Staining
If your organism is a gram-positive, non- spore- forming
rod, you should determine whether or not it is acid-fast.
Although some bacteria require 4 or 5 days growth to
exhibit acid-fastness, most species become acid-fast
within 2 days. For best results, therefore, do not use cul-
tures that are too old.
Another point to keep in mind is that most acid-
fast bacteria do not produce cells that are 1 00% acid-
fast. An organism is considered acid- fast if only por-
tions of the cells exhibit this characteristic. Refer to
Exercise 17 for this staining technique.
A final bit of advice: If you feel insecure about
your adeptness at Gram staining and think that you
might possibly have a gram-positive organism, even
though your organism seems to be gram- negative,
make an acid-fast stained slide. Many students find
(much to their chagrin later) that they didn't do acid-
fast staining because their organism seemed to be
gram-negative. An improperly gram-stained slide can
be very misleading when it comes to unknown identi-
fication.
Other Structures
If the protoplast in gram- stained slides stains un-
evenly, you might wish to do a simple stain with
Loeffler's methylene blue (Exercise 13) for evidence
of metachromatic granules.
Although a capsule stain (Exercise 14) may be
performed at this time, it might be better to wait until
a later date when you have the organism growing on
blood agar. Capsules usually are more apparent when
the organisms are grown on this medium.
Laboratory Report
There is no Laboratory Report to fill out for this exer-
cise. All information is recorded on the Descriptive
Chart.
156
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
VIM. Identification of
Unknown Bacteria
47. Cultural Characteristics
© The McGraw-H
Companies, 2001
Cultural Characteristics
The cultural characteristics of an organism pertain
to its macroscopic appearance on different kinds of
media. Descriptive terms, which are familiar to all
bacteriologists, and are used in Bergey's Manual,
must be used in recording cultural characteristics.
The most frequently used media for a cultural study
are nutrient agar, nutrient broth, and nutrient
gelatin. For certain types of unknowns it is also de-
sirable to inoculate a blood agar plate; if necessary,
this plate can be inoculated later. In addition to these
media, you will be inoculating a fluid thioglycollate
medium to determine the oxygen requirements of
your unknown.
First Period
(Inoculations)
During this period one nutrient agar plate, one nutri-
ent gelatin deep, two nutrient broths, and one tube of
fluid thioglycollate medium will be inoculated.
Inoculations will be made with the original broth cul-
ture of your unknown. The reason for inoculating two
tubes of nutrient broth here is to recheck the optimum
growth temperature of your unknown. In Exercise 45
you incubated your nutrient agar slants at 20° C and
37° C. It may well be that the optimum growth tem-
perature is closer to 30° C. It is to check out this in-
termediate temperature that an extra nutrient broth is
being inoculated. Proceed as follows:
Materials:
for each unknown:
1 nutrient agar pour
1 nutrient gelatin deep
2 nutrient broths
1 fluid thioglycollate medium (FTM)
1 Petri plate
1
2
3
4
Pour a Petri plate of nutrient agar for each un-
known and streak it with a method that will give
good isolation of colonies. Use the original broth
culture for streaking.
Inoculate the tubes of nutrient broth with a loop.
Make a stab inoculation into the gelatin deep by
stabbing the inoculating needle (straight wire) di-
rectly down into the medium to the bottom of the
tube and pulling it straight out. The medium must
not be disturbed laterally.
Inoculate the tube of FTM with a loopful of your
unknown. Mix the organisms throughout the tube
by rolling the tube between your palms.
Filiform
Echinulate
Beaded
Effuse
Arborescent
Rhizoid
Figure 47.1 Types of bacterial growth on nutrient agar slants
157
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
VIM. Identification of
Unknown Bacteria
47. Cultural Characteristics
© The McGraw-H
Companies, 2001
Exercise 47 • Cultural Characteristics
5. Place all tubes except one nutrient broth into a
basket and incubate for 24 hours at the tempera-
ture that seemed best in Exercise 45. Incubate the
remaining tube of nutrient broth separately at 30°
C. Incubate the agar plate, inverted, at the pre-
sumed best temperature.
Second Period
(Evaluation)
After the cultures have been properly incubated, carry
them to your desk in a careful manner to avoid dis-
turbing the growth pattern in the nutrient broths and
FTM. Before studying any of the tubes or plates, place
the tube of nutrient gelatin in an ice water bath. It will
be studied later. Proceed as follows to study each type
of medium and record the proper descriptive termi-
nology on the Descriptive Chart.
Materials:
reserve stock agar slant of unknown
spectrophotometer and cuvettes
hand lens
ice water bath near sink
Nutrient Agar Slant (Reserve Stock)
Examine your reserve stock agar slant of your un-
known that has been stored in the refrigerator since
the last laboratory period. Evaluate it in terms of the
following criteria:
Amount of Growth The abundance of growth may
be described as none, slight, moderate, and abundant.
Color Pigmentation should be looked for on the or-
ganisms and within the medium. Most organisms will
lack chromogenesis, exhibiting a white growth; oth-
ers are various shades of different colors. Some bac-
teria produce soluble pigments that diffuse into the
medium. Hold the slant up to a strong light to exam-
ine it for diffused pigmentation.
Opacity Organisms that grow prolifically on the
surface of a medium will appear more opaque than
those that exhibit a small amount of growth. Degrees
of opacity may be expressed in terms of opaque,
transparent, and translucent (partially transparent).
Form The gross appearance of different types of
growth are illustrated in figure 47.1. The following
descriptions of each type will help in differentiation:
Filiform: characterized by uniform growth along
the line of inoculation
Echinulate: margins of growth exhibit toothed
appearance
Beaded: separate or semiconfluent colonies
along the line of inoculation
Effuse: growth is thin, veil-like, unusually
spreading
Arborescent: branched, treelike growth
Rhizoid: rootlike appearance
Nutrient Broth
The nature of growth on the surface, subsurface, and
bottom of the tube is significant in nutrient broth cul-
tures. Describe your cultures as thoroughly as possi-
ble on the Descriptive Chart with respect to these
characteristics:
Surface Figure 47.2 illustrates different types of
surface growth. A pellicle type of surface differs from
the membranous type in that the latter is much thinner.
Aflocculent surface is made up of floating adherent
masses of bacteria.
ddoooc
Ring
Pellicle
Flocculent
Membranous
Figure 47.2 Types of surface growth in nutrient broth
Subsurface Below the surface, the broth may be
described as turbid if it is cloudy; granular if specific
small particles can be seen; flocculent if small masses
are floating around; and flaky if large particles are in
suspension.
Sediment The amount of sediment in the bottom of
the tube may vary from none to a great deal. To de-
scribe the type of sediment, agitate the tube, putting
the material in suspension. The type of sediment can
be described as granular, flocculent, flaky, and viscid.
Test for viscosity by probing the bottom of the tube
with a sterile inoculating loop.
Amount of Growth To determine the amount of
growth, it is necessary to shake the tube to disperse
the organisms. Terms such as slight (scanty), moder-
ate, and abundant adequately describe the amount.
158
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
VIM. Identification of
Unknown Bacteria
47. Cultural Characteristics
© The McGraw-H
Companies, 2001
Cultural Characteristics • Exercise 47
Optimum Temperature To determine which tem-
perature produced the best growth, pour the contents
from each tube of nutrient broth into separate cuvettes
and measure their percent transmittances on the spec-
trophotometer. If the percent transmittance is less at
30° C than at the other presumed optimum tempera-
ture, revise the optimum temperature on your
Descriptive Chart.
Fluid Thioglycollate Medium
Since the primary purpose of inoculating a tube of
fluid thioglycollate medium is to determine oxygen
requirements of your unknown, examine the tube to
note the position of growth in the tube. Compare your
tube with figure 22.5 on page 92 to make your analy-
sis. Designate your organism as being aerobic, mi-
cro aerophilic, facultative, or anaerobic on the
Descriptive Chart.
Gelatin Stab Culture
Remove your tube of nutrient gelatin from the ice wa-
ter bath and examine it. Check first to see if liquefac-
tion has occurred. Organisms that are able to liquefy
gelatin produce the enzyme gelatinase.
Liquefaction Tilt the tube from side to side to see if
a portion of the medium is still liquid. If liquefaction
has occurred, check the configuration with figure 47.3
to see if any of the illustrations match your tube. A de-
scription of each type follows:
Crateriform: saucer- shaped liquefaction
Napiform: turniplike
Infundibular: funnel-like or inverted cone
Saccate: elongate sac, tubular, cylindrical
Stratiform: liquefied to the walls of the tube in
the upper region
Note: The configuration of liquefaction is not as
significant as the mere fact that liquefaction takes
place. If your organism liquefies gelatin, but you
are unable to determine the exact configuration,
don't worry about it. However, be sure to record
on the Descriptive Chart the presence or absence
of gelatinase production.
Another important point: Some organisms
produce gelatinase at a very slow rate. Tubes that
are negative should be incubated for another 4 or
5 days to see if gelatinase is produced slowly.
Type of Growth (No Liquefaction) If no liquefac-
tion has occurred, check the tube to see if the organ-
ism grows in nutrient gelatin (some do, some don't).
If growth has occurred compare the growth with the
left-hand illustration in figure 47.3. It should be
pointed out, however, that, from a categorization
standpoint, the nature of growth in gelatin is not very
important.
Nutrient Agar Plate
Colonies grown on plates of nutrient agar should be
studied with respect to size, color, opacity, form, ele-
vation, and margin. With a dissecting microscope or
hand lens study individual colonies carefully. Refer to
figure 47.4 for descriptive terminology. Record your
observations on the Descriptive Chart.
Laboratory Report
There is no Laboratory Report for this exercise
Record all information on the Descriptive Chart.
!
Filiform
Beaded
Papillate
Villous
Arborescent
GROWTH WITHOUT LIQUEFACTION
n
L
^
\y
Crateriform Napiform Infundibular Saccate Stratiform
LIQUEFACTION CONFIGURATIONS
Figure 47.3 Growth in gelatin stabs
159
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
VIM. Identification of
Unknown Bacteria
47. Cultural Characteristics
© The McGraw-H
Companies, 2001
Exercise 47 • Cultural Characteristics
1 . Round
5. Concentric
9. Round with
Radiating Margin
■ ■.•/• : : : "a 4
• . .-.-lifts .vs
.•.■TSriX •!
2. Round with
Scalloped Margin
V-fc; .■-,::■ -Wi
3. Round with
Raised Margin
6. Irregular and
Spreading
,■ - ".-• > ^>V--'~. > /— >
iMaBBi^'
■ ' fta K^ '
1 0. Filiform
7. Filamentous
1 1 . Rhizoid
CONFIGURATIONS
4. Wrinkled
8. L-Form
1 2. Complex
1 . Smooth
(Entire)
2. Wavy
(Undulate)
3. Lobate
4. Irregular
(Erose)
5. Ciliate
6. Branching
V*.'
7. Wooly
8. Thread-Like
9. "Hair-Lock"-Like
MARGINS
1 . Flat
2. Raised
3. Convex
4. Drop-Like
5. Umbonate
6. Hilly
7. Ingrowing
Into Medium
8. Crateriform
ELEVATIONS
Figure 47.4 Colony characteristics
160
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
VIM. Identification of
Unknown Bacteria
48. Physiological
Characteristics: Oxidation
and Fermentation Tests
© The McGraw-H
Companies, 2001
Physiological Characteristics:
Oxidation and Fermentation Tests
48
The assemblage of morphological and cultural charac-
teristics on your Descriptive Chart during the past few
laboratory periods may be leading you to believe that
you already know the name of your unknown.
Students at this stage often begin to draw premature
conclusions. To provide you with a clearer perspective
of where you are in the categorization process, refer to
the separation outlines in figures 51.1 and 51.2, pages
178 and 179. Note that morphological and cultural
characteristics can lead you to 11 separate groups of
genera. It is very likely that one of these groups con-
tains the genus that includes your unknown.
Although morphological and cultural characteris-
tics are essential in getting to the genus, species de-
termination requires a good deal more information.
The physiological information that will be accumu-
lated here and in the next two exercises will make
species identification possible.
Before we get into the details of the various inoc-
ulations and tests, let's review some of the essentials
of microbial metabolism.
Metabolic Reactions
The chemical reactions that occur within the cells of
all living organisms are referred to as metabolism.
These reactions are catalyzed by protein molecules
called enzymes. The majority of enzymes function
within the cell and are called endoenzymes. Many
bacteria also produce exoenzymes, which are released
by the cell to catalyze reactions outside of the cell.
Figure 48.1 illustrates how these enzymes function.
In deriving energy from food, bacteria may be
either oxidative or fermentative. Oxidative bacteria
utilize oxygen to yield carbon dioxide and water.
These bacteria have a cytochrome enzyme system.
By utilizing organic compounds as electron donors,
with oxygen as the ultimate electron (and hydro-
gen) acceptor, they produce C0 2 and water as end-
products. Fermentative bacteria, on the other hand,
also utilize organic compounds for energy, but they
lack a cytochrome system. Instead of producing
only C0 2 and water, they produce complex end-
products, such as acids, aldehydes, and alcohols.
Various gases, such as carbon dioxide, hydrogen,
and methane, are also produced. In fermentative
bacteria, the organic compounds act both as elec-
tron donors and electron acceptors.
Sugars, particularly glucose, are the compounds
most widely used by fermenting organisms. Other sub-
stances such as organic acids, amino acids, purines,
and pyrimidines also can be fermented by some bacte-
ria. The end-products of a particular fermentation are
determined by the nature of the organism, the charac-
teristics of the substrate, and environmental conditions
such as temperature and pH.
Although fermentation and oxidation represent
two different types of energy-yielding reactions, they
can both be present in the same organism, as is true of
facultative anaerobes. It was pointed out in Exercise
H ?
r
Exoenzymes
Energy
Endoenzymes
<
Cellular Material
ritaH*
tfrvV-WU*
Waste
Products
BACTERIUM
Figure 48.1 Note that the hydrolytic exoenzymes split larger molecules into smaller ones, utilizing water in the process
The smaller molecules are then assimilated by the cell to be acted upon by endoenzymes to produce energy and cellu-
lar material
■
161
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
VIM. Identification of
Unknown Bacteria
48. Physiological
Characteristics: Oxidation
and Fermentation Tests
© The McGraw-H
Companies, 2001
Exercise 48 • Physiological Characteristics: Oxidation and Fermentation Tests
22 that in the presence of molecular oxygen these or-
ganisms shift from fermentation to oxidation. An ex-
ception, however, is seen in the lactic acid bacteria
where fermentation occurs in the presence of air (0 2 ).
Tests to Be Performed
Six types of reactions will be studied in this exercise:
(1) Durham- tube sugar fermentations, (2) mixed acid
fermentation, (3) butanediol fermentation, (4) cata-
lase production, (5) oxidase production, and (6) ni-
trate reduction. The performance of all these tests on
your unknown will involve a considerable number of
inoculations because a set of positive test controls will
also be needed. Although photographs of positive test
results are provided in this exercise, seeing the actual
test results in test tubes will make it more meaningful.
As you perform these various tests, attempt to
keep in mind what groups of bacteria relate to each
test. Although some tests are not very specific in
pointing the way to unknown identification, others are
very narrow in application.
One last comment of importance: It is not routine
practice to perform all these tests in identifying every
unknown. Remember that although it might appear
that our prime concern here is to identify an organism,
our most important goal is to learn about the various
types of tests for enzymes that are available. The use
of unknown bacteria to learn about them simply
makes it more of a challenge. In actual practice, phys-
iological tests are used very selectively. The "shotgun
approach" employed here is used to expose you to the
multitude of tests that are available.
First Period
(Inoculations)
The following two sets of inoculations (unknown and
test controls) may be done separately or combined
into one operation. The media for each set of inocula-
tions are listed separately under each heading.
Unknown Inoculations
Figure 48.2 illustrates the procedure for inoculating
seven test tubes and one Petri plate with your un-
known. Since your instructor may want you to inoc-
ulate some different sugar broths, blanks have been
provided in the materials list for write-ins. If differ-
ent media are distinguished from each other with
different-colored tube caps, write down the colors
after each medium below.
Materials: (for each unknown)
Durham tubes with phenol red indicator
1 glucose broth
1 lactose broth
1 mannitol broth
1
2
3
2 MR- VP medium
1 nitrate broth
1 nutrient agar slant
1 Petri plate of trypticase soy agar (TS A)
Label each tube with the number of your un-
known and an identifying letter as designated in
figure 48.2.
Label one half of the Petri plate UNKNOWN and
the other half P. AERUGINOSA.
Inoculate all broths and the slant with a loop.
Inoculate one half of the TS A plate with your un-
known, using an isolation technique.
Test Control Inoculations
Figure 48.4 on page 165 illustrates the procedure that
will be used for inoculating five test tubes to be used
for positive test controls. The Petri plate shown on the
right side is the same one that is shown in figure 48.2;
thus, it will not be listed in the materials below.
Materials:
1 glucose broth (Durham tube)
2 MR- VP medium
1 nitrate broth
1 nutrient agar slant
nutrient broth cultures of Escherichia coli,
Enterobacter aerogenes, Staphylococcus
aureus, and Pseudomonas aeruginosa
1
Label each tube with the code letter assigned to it
as listed:
glucose broth
MR- VP medium
MR- VP medium
nitrate broth
nutrient agar slant
A 1
D 1
E 1
F 1
G 1
2
Inoculate each of these tubes with a loopful of
the appropriate test organism according to fig-
ure 48.4.
3. Inoculate the other half of the TSA plate with P.
aeruginosa.
Incubation
Except for tube E (MR-VP), all the unknown inocula-
tions should be incubated for 24-48 hours at the un-
known's optimum temperature. Tube E should be in-
cubated for 3-5 days at the optimum temperature.
Except for Tube E 1 of the test controls, incubate
all the test-control tubes and the TSA plate at 37° C for
24-48 hours. Tube E 1 should be incubated at 37° C for
3-5 days.
162
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
VIM. Identification of
Unknown Bacteria
48. Physiological
Characteristics: Oxidation
and Fermentation Tests
© The McGraw-H
Companies, 2001
Physiological Characteristics: Oxidation and Fermentation Tests • Exercise 48
INOCULATIONS:
A minimum of seven tubes and one plate of tryp-
ticase soy agar are inoculated with the unknown.
Additional tubes of various sugars may also be
required.
TSA PLATE:
Inoculate one half of plate with
unknown and use other half for
positive test control (R aeruginosa).
This plate is the same one used in
figure 48.4.
TRYPTICASE SOY AGAR
■ k
X:
m
A
GLUCOSE
BROTH
w •
1
B
LACTOSE
BROTH
m
c
MANNITOL
BROTH
D
MR-VP
MEDIUM
TJ7
•I;
• r
- 1
E
MR-VP
MEDIUM
it.
n
'Si
•it*:
!-■■.
!!§
•»'v\Vr
■'■>».
K.'
F
NITRATE
BROTH
24
37° C 24 to 48 hours
G
NUTRIENT
AGAR SLANT
\J
INCUBATION
EVALUATION AND TESTS
POSITIVE TEST RESULTS
Except for tube E, incubate all tubes
for 24-48 hours at the optimum
temperature for the unknown.
Tube E should be incubated for 3-5
days at the optimum temperature.
Tubes A, B, and C: If broth turns
yellow, acid has been produced.
If a bubble is present in the inver-
ted vial, gas has been produced.
Tube D: Do methyl red test.
Tube E: Do Voges-Proskauertest.
Tube F: Do nitrite test.
Tube G: Do catalase test.
TSA plate: Do oxidase test.
Tube D: Red color is positive for
mixed acids production.
Tube E: Pink or red color is posi-
tive for butanediol production.
Tube F: Red color is positive for
nitrate reduction to nitrite.
Tube G: Bubbles effervescing
from streak indicate that cata-
lase is produced.
TSA plate: Black colonies indicate
that oxidase is produced.
Figure 48.2 Procedure for performing oxidation and fermentation tests
163
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
VIM. Identification of
Unknown Bacteria
48. Physiological
Characteristics: Oxidation
and Fermentation Tests
© The McGraw-H
Companies, 2001
Exercise 48 • Physiological Characteristics: Oxidation and Fermentation Tests
Second Period
(Test Evaluations)
After 24 to 48 hours incubation, arrange all your tubes
(except tubes E and E 1 ) in a test-tube rack in alphabet-
ical order, with the unknown tubes in one row and the
test controls in another row. As you interpret the re-
sults, record the information on the Descriptive Chart
immediately. Don't trust your memory. Any result that
is not properly recorded will have to be repeated.
Durham Tube Sugar Fermentations
When we use a bank of Durham tubes containing
various sugars, we are able to determine what sugars
an organism is able to ferment. If an organism is able
to ferment a particular sugar, acid will be produced
and gas may be produced. The presence of acid is de-
tectable with the color change of a pH indicator in
the medium. Gas production is revealed by the for-
mation of a void in the inverted vial of the Durham
tube. If it were important to know the composition of
the gas, we would have to use a Smith tube as shown
in figure 48.3. For our purposes here, the Durham
tube is preferable.
G&!i Enirbp-e™
IrwirrlFfJ Val
9nl)jftT|Af
nuifiBm TUb*
Interpretation Examine the glucose test control
tube (tube A 1 ), which you inoculated with E. coli.
Note that the phenol red has turned yellow, indicating
acid production. Also, note that the inverted vial has a
gas bubble in it. These observations tell us that E. coli
ferments glucose to produce acid and gas. The left-
hand illustration in figure 48.5, page 167, illustrates
how this positive tube compares with a negative tube
and an uninoculated one.
Now examine the three sugar broths (tubes A, B,
and C) that were inoculated with your unknown and
record your observations on the Descriptive Chart. If
there is no color change, record NONE after the spe-
cific sugar. If the tube is yellow with no gas, record
ACID. If the inverted vial contains gas and the tube is
yellow, record ACID AND GAS .
An important point to keep in mind at this time
is that a negative result on an unknown is as impor-
tant as a positive result. Don't feel that you have
failed in your technique if many of your tubes are
negative !
Figure 48.3 Two types of fermentation tubes
Mixed Acid Fermentation
(Methyl-Red Test)
A considerable number of gram-negative intestinal
bacteria can be differentiated on the basis of the
end-products produced when they ferment the glu-
cose in MR-VP medium. Genera of bacteria such as
Escherichia, Salmonella, Proteus, and Aeromonas
ferment glucose to produce large amounts of lactic,
acetic, succinic, and formic acids, plus C0 2 , H 2 , and
ethanol. The accumulation of these acids lowers the
pH of the medium to 5.0 and less.
If methyl red is added to such a culture, the indi-
cator turns red, an indication that the organism is a
mixed acid fermenter. These organisms are generally
great gas producers, too, because they produce the en-
zyme formic hydrogenylase, which splits formic acid
into equal parts of C0 2 and H 2 .
Media The sugar broths used here contain 0.5% of
the specific carbohydrate plus sufficient amounts of
beef extract and peptone to satisfy the nitrogen and
mineral needs of most bacteria. The pH indicator phe-
nol red is included for acid detection. This indicator is
red when the pH is above 7 and yellow below this
point.
Although there are many sugars that one might
use, glucose, lactose, and mannitol are logical ones to
begin with. Your instructor may have had you include
one or more additional kinds, and it is very likely that
you may wish to use some others later.
HCOOH f ° rmiC Mrogenylasc > C( v + H
Medium MR-VP medium is essentially a glucose
broth with some buffered peptone and dipotassium
phosphate.
Test Procedure Perform the methyl-red test first on
your test-control tube (D 1 ) and then on your unknown
(tube D). Proceed as follows:
164
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
VIM. Identification of
Unknown Bacteria
48. Physiological
Characteristics: Oxidation
and Fermentation Tests
© The McGraw-H
Companies, 2001
Physiological Characteristics: Oxidation and Fermentation Tests • Exercise 48
P.
aeruginosa
" ;■ * "• .« • r.
I ".' ■** •
:V-
:0 :&■
1 . «.• .
■ », '"'"-,
.;' •#*./•
1 r. *>.*
1 v ?%*•
| ?. . ■■!•'..
:: :>■■
■:.»-■
•.v' ■:.**•:.
A1
."■*■ "*•*'
.V •#'
:v -:>
•' .-*{ ,
-' ■••■>•
D1
r-
•I
*(.
■'. V" i
.'t-'k.'..'
:£*''
• u
F1
■■#'■
Unknown
37° C 24 to 48 hours
GLUCOSE
BROTH
MR-VP
MEDIUM
NITRATE
BROTH
E1
MR-VP
MEDIUM
G1
NUTRIENT
AGAR SLANT
\j
<j
OXIDASE TEST
\
NCUBATE AT 37° C FOR 24 TO 48 HOURS
■»■ 1
vT.
A
• i * '. ' '
A1
/"\
^J :
F1
VJ*
G1
GLUCOSE
FERMENTATION
MIXED ACID
FERMENTATION
NITRATE
REDUCTION
BUTANEDIOL
PRODUCTION
CATALASE
PRODUCTION
Look for acid
(yellow) and gas
(bubble in tube).
Add methyl red
to culture.
Add nitrite test re-
agents to culture.
Confirm negatives
with zinc dust.
Add Barritt's re-
agents to culture
Add hydrogen per-
oxide to slant. Look
for bubbles.
Figure 48.4 Procedure for doing positive test controls
165
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
VIM. Identification of
Unknown Bacteria
48. Physiological
Characteristics: Oxidation
and Fermentation Tests
© The McGraw-H
Companies, 2001
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
VIM. Identification of
Unknown Bacteria
48. Physiological
Characteristics: Oxidation
and Fermentation Tests
© The McGraw-H
Companies, 2001
Physiological Characteristics: Oxidation and Fermentation Tests • Exercise 48
Materials:
dropping bottle of methyl-red indicator
1 . Add 3 or 4 drops of methyl red to test-control tube
D 1 , which was inoculated with E. coll The tube
should become red immediately. A reddish color,
as shown in the left-hand tube of the middle illus-
tration of figure 48.5 is a positive methyl-red test.
2. Repeat the same procedure with your unknown
culture (tube D) of MR-VP medium. If your un-
known culture becomes yellow like the right-
hand tube in figure 47.5, your unknown is nega-
tive for this test.
3. Record your results on the Descriptive Chart.
Butanediol Fermentation
(Voges-Proskauer Test)
A negative methyl-red test may indicate that the or-
ganism being tested produced a lot of 2,3 butanediol
and ethanol instead of acids. All species of
Enterobacter and Serratia, as well as some species of
Erwinia, Bacillus, and Aeromonas, do just that. The
production of these nonacid end-products results in
less lowering of the pH in MR-VP medium, causing
the methyl-red test to be negative.
Unfortunately, there is no satisfactory test for 2,3
butanediol; however, acetoin (acetylmethylcarbinol),
a precursor of 2,3 butanediol, is easily detected with
Barritt's reagent.
Barritt's reagent consists of alpha naphthol and
KOH. When added to a 3 to 5 day culture of MR-VP
medium, and allowed to stand for some time, the
medium changes to pink or red in the presence of ace-
toin. Since acetoin and 2,3 butanediol are always si-
multaneously present, the test is valid. This indirect
method of testing for 2,3 butanediol is called the
Voges-Proskauer test.
Test Procedure Perform the Voges-Proskauer test
on your unknown and test-control tubes of MR-VP
medium (tubes E and E 1 ). Note that the test-control
tube was inoculated with E. aero genes. Follow this
procedure:
Materials:
Barritt's reagents
2 pipettes (1 ml size)
2 empty test tubes
1 . Label one empty test tube E (for unknown) and
the other E 1 (for control).
2. Pipette 1 ml from culture tube E to the empty tube
E and 1 ml from culture tube E 1 to the empty tube
E 1 . Use separate pipettes for each tube.
3. Add 18 drops (about 0.5 ml) of Barritt's solution
A (alpha naphthol) to each of the tubes that con-
tain 1 ml of culture.
4. Add an equal amount of Barritt's solution B
(KOH) to the same tubes.
5. Shake the tubes vigorously every 20 seconds un-
til the control tube (E 1 ) turns pink or red. Let the
tubes stand for 1 or 2 hours to see if the unknown
turns red. Vigorous shaking is very important to
achieve complete aeration.
A positive Voges-Proskauer reaction is pink
or red. The left-hand tube in the right-hand illus-
tration of figure 48.5 shows what a positive result
looks like.
6. Record your results on the Descriptive Chart.
DURHAM TUBES
From left to right: uninoculated
positive, and negative.
METHYL RED TEST
Tube on left is positive (E. coli);
tube on right is negative.
VOGES-PROSKAUER TEST
Tube on left is positive (E. aerogenes);
tube on right is negative.
Figure 48.5 Durham tubes, mixed acid, and butanediol fermentation tests
167
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
VIM. Identification of
Unknown Bacteria
48. Physiological
Characteristics: Oxidation
and Fermentation Tests
© The McGraw-H
Companies, 2001
Exercise 48 • Physiological Characteristics: Oxidation and Fermentation Tests
Catalase Production
Most aerobes and facultatives that utilize oxygen pro-
duce hydrogen peroxide, which is toxic to their own
enzyme systems. Their survival in the presence of this
antimetabolite is possible because they produce an en-
zyme called catalase, which converts the hydrogen
peroxide to water and oxygen:
2H 2 2
catalase
* 2H 2 + 2
It has been postulated that the death of strict
anaerobes in the presence of oxygen may be due to the
suicidal act of H 2 2 production in the absence of cata-
lase production. The presence or absence of catalase
production is an important means of differentiation
between certain groups of bacteria.
Test Procedure To determine whether or not cata-
lase is produced, all that is necessary is to place a few
drops of 3% hydrogen peroxide on the organisms of a
slant culture. If the hydrogen peroxide effervesces,
the organism is catalase-positive.
Materials:
3% hydrogen peroxide
test-control tube G 1 with S. aureus growth and
unknown tube G
1
2
While holding test-control tube G 1 at an angle, al-
low a few drops of H 2 2 to flow slowly down
over the S. aureus growth on the slant. Note how
bubbles emerge from the organisms.
Repeat the test on your unknown (tube G) and
record your results on the Descriptive Chart.
Oxidase Production
The production of oxidase is one of the most signifi-
cant tests we have for differentiating certain groups of
bacteria. For example, all the Enterobacteriaceae are
oxidase-negative and most species of Pseudomonas
are oxidase-positive. Another important group, the
Neisseria, are oxidase producers.
Two methods are described here for performing
this test. The first method utilizes the entire TSA
plate; the second method is less demanding in that
only a loopful of organisms from the plate is used.
Both methods are equally reliable.
Materials:
TSA plate streaked with unknown and P.
aeruginosa
oxidase test reagents (1% solution of dimethyl-
p-phenylenediamine hydrochloride)
Whatman no. 2 filter paper
Petri dish
Entire Plate Method Onto the TSA plate where
you streaked your unknown and P. aeruginosa, pour
some of the oxidase test reagent, covering the
colonies of both organisms.
Observe that the Pseudomonas colonies first be-
come pink, then change to maroon, dark red, and fi-
nally black. Refer to figure 48.6. If your unknown fol-
lows the same color sequence, it, too, is oxidase-positive.
Record your results on the Descriptive Chart.
Filter Paper Method On a piece of Whatman no. 2
filter paper in a Petri dish, place several drops of oxi-
dase test reagent. Remove a loopful of the organisms
from one of the colonies and smear the organisms over
a small area of the paper. The positive color reaction
described above will show up within 10-15 seconds.
Record your results on the Descriptive Chart.
Nitrate Reduction
Many facultative bacteria are able to use the oxygen in
nitrate as a hydrogen acceptor in anaerobic respiration,
thus converting nitrate to nitrite. This enzymatic reaction
is controlled by an inducible enzyme called nitratase.
Figure 48.6 Oxidase Test: The colonies on the left are
positive; the ones on the right are negative.
Figure 48.7 Nitrate Reduction Test Tube on left is pos
itive (E. coli); tube on right is negative.
168
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
VIM. Identification of
Unknown Bacteria
48. Physiological
Characteristics: Oxidation
and Fermentation Tests
© The McGraw-H
Companies, 2001
Physiological Characteristics: Oxidation and Fermentation Tests • Exercise 48
The chemical reaction for this enzymatic reaction
is as follows:
N0 3 " + 2e~ + 2H + mtratase > NQ 2 ~ 4- H 2
Since the presence of free oxygen prevents nitrate
reduction, actively multiplying organisms will use up
the oxygen first and then utilize the nitrate. In cultur-
ing some organisms, it is desirable to use anaerobic
methods to ensure nitrate reduction.
Test Procedure The nitrate broth used in this test
consists of beef extract, peptone, and potassium ni-
trate. To test for nitrite after incubation, we use two
reagents designated as A and B .
Reagent A contains sulfanilic acid and reagent B
contains dimethyl-alpha-naphthylamine. In the pres-
ence of nitrite, these reagents cause the culture to turn
red. Negative results must be confirmed as negative
with zinc dust.
Materials:
nitrate broth cultures of unknown (tube F) and
test control E. coli (tube F 1 )
nitrite test reagents (solutions A and B)
zinc dust
1 . Add 2 or 3 drops of nitrite test solution A (sulfanilic
acid) and an equal amount of solution B (dimethyl-
2
3
alpha- naphthylamine) to the nitrate broth culture of
E. coli (tube F 1 ). A red color should appear almost
immediately (see figure 48.7), indicating that ni-
trate reduction has occurred.
CAUTION
Avoid skin contact with solution B. Dimethyl-alpha-
naphthylamine is carcinogenic.
Repeat this procedure with your unknown (tube
F). If the red color does not develop, your un-
known is negative for nitrate reduction. All nega-
tive results should be confirmed as being negative
as follows:
Negative Confirmation: Add a pinch of zinc dust
to the tube and shake it vigorously. If the tube be-
comes red, the test is confirmed as being negative.
Zinc causes this reaction by reducing nitrate to ni-
trite; the newly formed nitrite reacts with the
reagents to produce the red color.
Record your results on the Descriptive Chart.
Laboratory Report
Answer the questions on Laboratory Report 48-50
that pertain to this exercise.
169
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
VIM. Identification of
Unknown Bacteria
49. Physiological
Characteristics: Hydrolytic
Reactions
© The McGraw-H
Companies, 2001
49
Physiological Characteristics:
Hydrolytic Reactions
As indicated in the last exercise, many bacteria pro-
duce hydrolases, which split complex organic com-
pounds into smaller units. These exoenzymes accom-
plish molecular splitting in the presence of water. We
have already observed one example of protein hy-
drolysis in Exercise 47: gelatin hydrolysis by gelati-
nase. In this exercise we will observe the hydrolysis
of starch, casein, fat, tryptophan, and urea. Each test
plays an important role in the identification of certain
types of bacteria. This exercise will be performed in
the same manner as the previous one with test controls
being made for comparisons.
Figure 49.1 illustrates the general procedure to
be used. Three agar plates and four test tubes will be
inoculated. After incubation, some of the plates and
tubes will have test reagents added to them; others
will reveal the presence of hydrolysis by changes
that have occurred during incubation. Proceed as
follows:
2
3
4
Label a tube of urea broth P. VULGARIS and a
tube of tryptone broth E. COLL These will be
your test controls for urea and tryptophan hydrol-
ysis. Inoculate each tube accordingly.
For each unknown, label one tube of urea broth
and one tube of tryptone broth with the code num-
ber of your unknown. Inoculate each tube with
the appropriate unknown.
Incubate the plates and two test-control tubes at
37° C. Incubate the unknown tubes of urea broth
and tryptone broth at the optimum temperatures
for the unknowns.
Second Period
(Evaluation of Tests)
After 24 to 48 hours incubation of unknowns and test
controls, compare your unknowns with the test con-
trols, recording all data on the Descriptive Chart.
First Period
(Inoculations)
If each student is working with only one unknown,
students can work in pairs to share Petri plates. Note
in figure 49.1 how each plate can serve for two un-
knowns with the test-control organism streaked down
the middle. If each student is working with two un-
knowns, the plates will not be shared. Whether or not
the two tubes for test controls will be shared depends
on the availability of materials.
Materials:
per pair of students with one unknown each, or
for one student with two unknowns:
1 starch agar plate
1 skim milk agar plate
1 spirit blue agar plate
3 urea broths
3 tryptone broths
nutrient broth cultures of B. subtilis, E. coli,
S. aureus, and P. vulgaris.
1 . Label and streak the three different agar plates in
the manner shown in figure 49 . 1 . Note that straight
line streaks are made on each plate. Indicate, also,
the type of medium in each plate.
Starch Hydrolysis
Since many bacteria are capable of hydrolyzing
starch, this test has fairly wide application. The starch
molecule is a large one consisting of two constituents:
amy lose, a straight chain polymer of 200 to 300 glu-
cose units, and amylopectin, a larger branched poly-
mer with phosphate groups. Bacteria that hydrolyze
starch produce amylases that yield molecules of mal-
tose, glucose, and dextrins.
Materials:
Gram's iodine
starch agar culture plate
Iodine solution (Gram's) is an indicator of starch.
When iodine comes in contact with a medium con-
taining starch, it turns blue. If starch is hydrolyzed
and starch is no longer present, the medium will have
a clear zone next to the growth.
By pouring Gram's iodine over the growth on the
medium, one can see clearly where starch has been
hydrolyzed. If the area immediately adjacent to the
growth is clear, amylase is produced.
Pour enough iodine over each streak to com-
pletely wet the entire surface of the plate. Rotate and
tilt the plate gently to spread the iodine. Compare
170
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
VIM. Identification of
Unknown Bacteria
49. Physiological
Characteristics: Hydrolytic
Reactions
© The McGraw-H
Companies, 2001
INOCULATIONS:
Three agar plates, a urea broth, and a tryptone
broth are inoculated with the unknown. Note
that straight-line streaks are used on the
plates and that test control organisms are also
used on the plates.
INCUBATION:
The three plates and the two tubes should be
incubated at the optimum temperature of the
unknown for 24-48 hours.
TEST CONTROL TUBE INOCULATIONS:
A tryptone broth is inoculated with E. coli, and a
urea broth is inoculated with P. vulgaris. Incu-
bate at 37° C for 24-48 hours.
UNKNOWN
W
.1
• «.
**u.
STARCH AGAR
SKIM MILK AGAR
SPIRIT BLUE AGAR
J7.
11
O'
UREA
BROTH
TRYPTONE
BROTH
UREA
BROTH
TEST CONTROL PLATE INOCULATION
STARCH HYDROLYSIS
CASEIN HYDROLYSIS
FAT HYDROLYSIS
UREA HYDROLYSIS
TRYPTOPHAN
HYDROLYSIS
Add several drops
of Gram's iodine to
growth on plate.
Clear areas next to
growth indicate
that starch is
hydrolyzed.
If clear zone is seen
adjacent to growth
on medium, casein
is hydrolyzed.
If growth streak
exhibits a dark blue
precipitate, the
organism is posi-
tive for fat hydro-
lysis.
If broth exhibits a
red or cerise color,
the organism can
hydrolyze urea.
If after adding
Kovacs' reagent, a
red ring appears
on the surface of
the broth, the
organism is able to
hydrolyze
tryptophan (indole-
positive).
Figure 49.1 Procedure for doing hydrolysis tests on unknowns
171
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
VIM. Identification of
Unknown Bacteria
49. Physiological
Characteristics: Hydrolytic
Reactions
© The McGraw-H
Companies, 2001
Exercise 49 • Physiological Characteristics: Hydrolytic Reactions
your unknowns with the positive result seen along the
growth of B. subtilis. The left-hand illustration of fig-
ure 49.2 illustrates what it looks like.
Casein Hydrolysis
Casein is the predominant protein in milk. Its pres-
ence causes milk to have its characteristic white ap-
pearance. Many bacteria produce the exoenzyme ca-
seinase, which hydrolyzes casein to produce more
soluble, transparent derivatives. Protein hydrolysis
also is referred to as proteolysis, or peptonization.
Examine the streaks on the skim milk agar
plates. Note that a clear zone exists adjacent to the
growth of B. subtilis. This is evidence of casein hy-
drolysis. The middle illustration in figure 49.2
shows what it looks like. Compare your unknown
with this positive result and record the results on the
Descriptive Chart.
Fat Hydrolysis
The ability of organisms to hydrolyze fat is accom-
plished with the enzyme lipase. In this reaction the fat
molecule is split to form one molecule of glycerol and
CH, — o
CH — O
CH
-C — R
/
- C — R' •
O— C — R"
CH 2 OH RCOOH
-I- 3 H 2
lipase
CHOH + R'COOH
TRIGLYCERIDE
CHjOH R'COOH
GLYCEROL FATTY ACIDS
three fatty acid molecules. The glycerol and fatty
acids can be used by the organism to synthesize bac-
terial fats and other cell components. In many in-
stances they are even oxidized to yield energy under
aerobic conditions. This ability of bacteria to decom-
pose fats plays a role in the rancidity of certain foods,
such as margarine.
Spirit blue agar contains a vegetable oil that,
when hydrolyzed by most organisms, results in the
lowering of the pH sufficiently to produce a dark
blue precipitate. Unfortunately, the hydrolytic ac-
tion of some organisms on this medium does not pro-
duce a blue precipitate because the pH is not lowered
sufficiently.
Examine the S. aureus growth carefully. You
should be able to see this dark blue reaction. The
right-hand illustration in figure 49.2 exhibits what it
should look like.
Compare the positive reaction of S. aureus with
the reaction of your unknown. If your unknown ap-
pears to be negative, hold the plate up toward the light
and look for a region near growth where oil droplets
are depleted. If you see depletion of oil drops, con-
sider your organism to be positive for this test. Record
the results on the Descriptive Chart.
Tryptophan Hydrolysis
Certain bacteria, such as E. coli, have the ability to
split the amino acid tryptophan into indole and pyru-
vic acid. The enzyme that causes this hydrolysis is
tryptophanase. Indole can be easily detected with
Ko vacs' reagent. This test is particularly useful in dif-
ferentiating E. coli from some closely related enteric
bacteria.
STARCH
Dtiftf tow almig leh wreak indicates
starch hydrtfyaia
CASEIN
Oaor tart aldpg teh sireek inck^Hs-
casc-i nydrtityals
QB'k tJ'ua pl^Tflnl^tign on ■ inn
organ^m indicates fl hydrciyiw t?\
Figure 49.2 Hydrolysis test plates: Starch, casein, fat
172
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
VIM. Identification of
Unknown Bacteria
49. Physiological
Characteristics: Hydrolytic
Reactions
© The McGraw-H
Companies, 2001
Physiological Characteristics: Hydrolytic Reactions • Exercise 49
CH - COOH
I
NH : tryptophanase
* H 2 0™ ■
/\
*
I
H
/
TRYPTOPHAN
INDOLE
CH
+ C -- O + NHj
COOH
PYRUVIC
ACID
NH
c = o
NH 2
UREA
+ H 2
urease
2NH 3 + C0 2
AMMONIA
Tryptone broth (1%) is used for this test because
it contains a great deal of tryptophan. Tryptone is a
peptone derived from casein by pancreatic digestion.
Materials:
Ko vacs' reagent
tryptone broth cultures of unknown
and E. coli
To test for indole, add 10 or 12 drops of Ko vacs'
reagent to the E. coli culture in tryptone broth. A red
layer should form at the top of the culture, as shown
in figure 49.3. Repeat the test on your unknown and
record the results on the Descriptive Chart.
Urea Hydrolysis
The differentiation of gram-negative enteric bacteria
is greatly helped if one can demonstrate that the un-
known can produce urease. This enzyme splits off
ammonia from the urea molecule, as shown nearby.
Note in the separation outline in figure 51.3 that three
genera (Proteus, Providencia, and Morganella) are
positive for the production of this hydrolytic enzyme.
Urea broth is a buffered solution of yeast extract
and urea. It also contains phenol red as a pH indicator.
Since urea is unstable and breaks down in the auto-
clave at 15 psi steam pressure, it is usually sterilized
by filtration. It is tubed in small amounts to hasten the
visibility of the reaction.
When urease is produced by an organism in this
medium, the ammonia that is released raises the pH.
As the pH becomes higher, the phenol red changes
from a yellow color (pH 6.8) to a red or cerise color
(pH 8.1 or more).
Examine your tube of urea broth that was inocu-
lated with Proteus vulgaris. Compare your unknown
with this standard. Figure 49.4 reveals how positive
and negative results of the test should appear. If your
unknown is negative, incubate the tube for a total of 7
days to check for a slow urease producer. Record your
result on the Descriptive Chart.
Figure 49.3 Indole Test: Tube on the left is positive (E.
coli); tube on the right is negative.
Figure 49.4 Urease Test: From left to right — uninocu
lated, positive (Proteus) and negative
173
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
VIM. Identification of
Unknown Bacteria
50. Physiological
Characteristics:
Miscellaneous Tests
© The McGraw-H
Companies, 2001
50
Physiological Characteristics:
Miscellaneous Tests
There are several additional physiological tests used
in unknown identification that are best grouped sep-
arately as "miscellaneous tests." They include tests
for hydrogen sulfide production, citrate utilization,
phenylalanine deaminization, and litmus milk reac-
tions. During the first period, inoculations of four
kinds of media will be made for these tests. An ex-
planation of the value of the IMViC tests will also be
included.
First Period
(Inoculations)
Since test controls are included in this exercise, two
sets of inoculations will be made. For economy of ma-
terials, one set of test controls will be made by stu-
dents working in pairs.
Materials:
for test controls, per pair of students:
1 Kligler's iron agar deep or SIM medium
1 Simmons citrate agar slant
1 phenylalanine agar slant
nutrient broth cultures of Proteus vulgaris.
Staphylococcus aureus, and Enterobacter
aerogenes
per unknown, per student:
1 Kligler's iron agar deep or SIM medium
1 Simmons citrate agar slant
1 phenylalanine agar slant
1 litmus milk
1
2
3
4
Label one tube of Kligler's iron agar (or SIM
medium) P. VULGARIS and additional tubes with
your unknown numbers. Inoculate each tube by
stabbing with a straight wire.
Label one tube of Simmons citrate agar E. AERO-
GENES and additional tubes with your unknown
numbers. Use a straight wire to streak- stab each
slant; i.e., streak the slant first, and then stab into
the middle of the slant.
Label one tube of phenylalanine agar slant P.
VULGARIS and the other with your unknown
code number. Streak each slant with the appropri-
ate organisms.
With a loop, inoculate one tube of litmus milk
with your unknown. (Note: A test control for this
medium will not be made. Figure 50.2 will take
its place.)
5. Incubate the unknowns at their optimum temper-
atures. Incubate the test controls at 37° C for
24-48 hours.
Second Period
(Evaluation of Tests)
After 24 to 48 hours incubation, examine the tubes to
evaluate according to the following discussion.
Record all results on the Descriptive Chart.
Hydrogen Sulfide Production
Certain bacteria, such as Proteus vulgaris, produce hy-
drogen sulfide from the amino acid cysteine. These or-
ganisms produce the enzyme cysteine desulfurase,
which works in conjunction with the coenzyme pyri-
doxyl phosphate. The production of H 2 S is the initial
step in the deamination of cysteine as indicated below:
COOH
cysteine
I
*-H«S -r C NH:
COOH
a amino
acrylic acid
I
C - NH
I
COOH
tmmo
acid
CH
H,0
O
C '
I
COOH
pyruvic
acid
NH:<
Kligler's iron agar or SIM medium is used here to
detect hydrogen sulfide production. Both of these me-
dia contain iron salts that react with H 2 S to form a
dark precipitate of iron sulfide.
Kligler's iron agar also contains glucose, lactose,
and phenol red. When this medium is used in slants it
is an excellent medium for detecting glucose and lac-
tose fermentation. SIM medium, on the other hand,
can also be used for determining motility and testing
for indole production.
Examine the tube of one of these media that was
inoculated with P. vulgaris. If it is Kligler's iron agar
it will look like the left-hand tube in figure 50.1. A
positive reaction in SIM medium will look like the
small tube on the right.
Compare your unknown with this control tube
and record your results on the Descriptive Chart.
174
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
VIM. Identification of
Unknown Bacteria
50. Physiological
Characteristics:
Miscellaneous Tests
© The McGraw-H
Companies, 2001
Physiological Characteristics: Miscellaneous Tests • Exercise 50
Citrate Utilization
The ability of some organisms, such as E. aerogenes
and Salmonella typhimurium, to utilize citrate as a
sole source of carbon can be a very useful differenti-
ation characteristic in working with intestinal bacte-
ria. Koser's citrate medium and Simmons citrate agar
are two media that are used to detect this ability in
bacteria. In both of these synthetic media sodium cit-
rate is the sole carbon source; nitrogen is supplied by
ammonium salts instead of amino acids.
Examine the test control slant of this medium that
was inoculated with E. aerogenes. Note the distinct
Prussian blue color change that has occurred. Refer
to the right-hand illustration in figure 50.1. Record
your results on the Descriptive Chart.
Phenylalanine Deamination
A few bacteria, such as Proteus, Morganella, and
Providencia, produce the deaminase phenylalanase,
that deaminizes the amino acid phenylalanine to pro-
duce phenylpyruvic acid (PPA). This characteristic is
used to help differentiate these three genera from
other genera of the Enterobacteriaceae. The reaction
is as follows:
NH2
, L J "-
f •.
<( y)-CH 2 CHCOOH
phenylalanase
\ VcH 2 COCOOH + NHa
PHENYLALANINE
PHENYLPYRUVIC ACID
Proceed as follows to test for the production of
phenylpyruvic acid, which is evidence that the en-
zyme phenylalanase has been produced:
Materials:
dropping bottle of 10% ferric chloride
Allow 5-10 drops of 10% ferric chloride to flow down
over the slants of the test control (P. vulgaris) and your
unknowns. To hasten the reaction, use a loop to emul-
sify the organisms into solution. A deep green color
should appear on the test control slant in 1-5 minutes.
Refer to the middle illustration in figure 50.1.
Compare your unknown with the control and record
your results on the Laboratory Report.
The IMViC Tests
In the differentiation of E. aerogenes and E. coli, as
well as some other related species, four physiological
tests have been grouped together into what are called
the IMViC tests. The / stands for indole; the M and V
stand for methyl red and Voges-Proskauer tests; i sim-
ply facilitates pronunciation; and the C signifies cit-
rate utilization. In the differentiation of the two col-
iforms E. coli and E. aerogenes, the test results appear
as charted below, revealing completely opposite reac-
tions for the two organisms on all tests.
1
M
V
c
E. coli
+
+
—
E. aerogenes
+
+
The significance of these tests is that when test-
ing drinking water for the presence of the sewage in-
dicator E. coli, one must be able to rule out E. aero-
genes, which has many of the morphological and
physiological characteristics of E. coli. Since E.
aerogenes is not always associated with sewage, its
HYDROGEN SULFIDE TEST
Positive tubes have black precipitate.
Large tubes: Kligler; Small tube is SIM
PPA TEST
Left-hand tube exhibits a positive re-
action (green). Other tube is negative
CITRATE UTILIZATION
Left to right: uninoculated, positive
(E. aerogenes), and negative.
Figure 50.1 Hydrogen sulfide, PPA, and citrate utilization tests
175
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
VIM. Identification of
Unknown Bacteria
50. Physiological
Characteristics:
Miscellaneous Tests
© The McGraw-H
Companies, 2001
Exercise 50 • Physiological Characteristics: Miscellaneous Tests
presence in water would not necessarily indicate
sewage contamination.
If you are attempting to identify a gram- negative,
facultative, rod-shaped bacterial organism, group
these series of tests together in this manner to see how
your unknown fits this combination of tests.
Litmus Milk Reactions
Litmus milk contains 1 0% powdered skim milk and a
small amount of litmus as a pH indicator. When the
medium is made up, its pH is adjusted to 6.8. It is an
excellent growth medium for many organisms and
can be very helpful in unknown characterization. In
addition to revealing the presence or absence of fer-
mentation, it can detect certain proteolytic character-
istics in bacteria. A number of facultative bacteria
with strong reducing powers are able to utilize litmus
as an alternative electron acceptor to render it color-
less. Figure 50.2 reveals the color changes that cover
the spectrum of litmus milk changes. Since some of
the reactions take 4 to 5 days to occur, the cultures
should be incubated for at least this period of time;
they should be examined every 24 hours, however.
Look for the following reactions:
Acid Reaction Litmus becomes pink. Typical of
fermentative bacteria.
Alkaline Reaction Litmus turns blue or purple.
Many proteolytic bacteria cause this reaction in the
first 24 hours.
Litmus Reduction Culture becomes white; ac-
tively reproducing bacteria reduce the O/R potential
of medium.
Coagulation Curd formation. Solidification is due
to protein coagulation. Tilting tube at 45° will indicate
whether or not this has occurred.
Peptonization Medium becomes translucent. It of-
ten turns brown at this stage. Caused by proteolytic
bacteria.
Ropiness Thick, slimy residue in bottom of tube.
Ropiness can be demonstrated with sterile loop.
Record the litmus milk reactions of your unknown on
the Descriptive Chart.
Laboratory Report
Complete Laboratory Report 48-50, which reviews
all physiological tests performed in the last three
exercises.
Figure 50.2 Litmus milk reactions: (A) Alkaline. (B) Acid. (C) Upper transparent portion is peptonization; solid white por
tion in bottom is coagulation and litmus reduction; overall redness is interpreted as acid. (D) Coagulation and litmus re-
duction in lower half; some peptonization (transparency) and acid in top portion. (E) Litmus indicator is masked by pro-
duction of soluble pigment (Pseudomonas); some peptonization is present but difficult to see in photo.
176
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
VIM. Identification of
Unknown Bacteria
51. Use of Bergey's Manual
and Indentibacter
interactus
© The McGraw-H
Companies, 2001
Use of Bergey'd Manual and
Identibacter Interactus
51
Once you have recorded all the data on your
Descriptive Chart pertaining to morphological, cul-
tural, and physiological characteristics of your un-
known, you are ready to determine its genus and
species. Determination of the genus should be rela-
tively easy; species differentiation, however, is con-
siderably more difficult.
The most important single source of information
we have for the identification of bacteria is Bergey's
Manual of Systematic Bacteriology. This monumental
achievement, which consists of four volumes, re-
placed a single- volume eighth edition of Bergey's
Manual of Determinative Bacteriology. Although the
more recent publication consists of four volumes,
only volumes 1 and 2 will be used for the identifica-
tion of the unknowns in this course.
In addition to using Bergey's Manual, you may
have an opportunity to use a computer simulation pro-
gram called Identibacter interactus, which is avail-
able on a CD-ROM disc. Details of the application of
this computer program are discussed on page 181 and
in Appendix F.
Bergey's Manual is a worldwide collaborative ef-
fort that has an editorial board of 13 trustees. Over
200 specialists from 19 countries are listed as contrib-
utors to the first two volumes. One of the purposes of
this exercise is to help you glean the information from
these two volumes that is needed to identify your un-
known. Before we get into the mechanics of using
Bergey's Manual, a few comments are in order per-
taining to the problems of bacterial classification.
Classification Problems
Compared with the classification of bacteria, the clas-
sification of plants and animals has been relatively
easy. In these higher forms, a hierarchy of orders,
families, and genera is based, primarily, on evolution-
ary evidence revealed by fossils laid down in sedi-
mentary layers of Earth's crust. Some of the earlier
editions of Bergey's Manual attempted to use the
same hierarchial system, but the attempt had to be
abandoned when the eighth edition was published;
without paleontological information to support the
system it literally fell apart.
The present system of classification in Bergey's
Manual uses a list of "Sections" that separate the var-
ious groups. Each section is described in common
terms so that it is easily understood (even for begin-
ners). For example, Section 1 is entitled The
Spirochaetes. Section 4 pertains to Gram -Negative
Aerobic Rods and Cocci. If one scans the Table of
Contents in each volume after having completed all
tests, it is possible, usually, to find a section that con-
tains the unknown being studied.
A perusal of these sections will reveal that some
sections have a semblance of hierarchy in the form of
orders, families, and genera. Other sections list only
genera.
Thus, we see that the classification system of bac-
teria, as developed in Bergey 's Manual, is not the tidy
system we see in higher forms of life. The important
thing is that it works.
Our dependency over the years on Bergey's
Manual has led many to think of its classification sys-
tem as the "Official Classification." Staley and Krieg
in their Overview in Volume 1 emphasize that no of-
ficial classification of bacteria exists; in other words,
the system offered in Bergey's Manual is simply a
workable system, but in no sense of the word should
it be designated as the official classification system.
Presumptive Identification
The place to start in identifying your unknown is to
determine what genus it fits into. If Bergey's Manual
is available, scan the Tables of Contents in Volumes 1
and 2 to find the section that seems to describe your
unknown. If these books are not immediately avail-
able you can determine the genus by referring to the
separation outlines in figures 51.1 and 51.2. Note that
seven groups of gram-positive bacteria are winnowed
out in figure 51.1 and four groups of gram-negative
bacteria in figure 51.2.
To determine which genus in the group best fits
the description of your unknown, compare the genera
descriptions provided below. Note that each group has
a section designation to identify its position in
Bergey 's Manual.
Group I (Section 13, Vol. 2) Although there are only
three genera listed in this group, Section 13 in
Bergey's Manual lists three additional genera, one of
which is Sporosarcina, a coccus-shaped organism
177
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
VIM. Identification of
Unknown Bacteria
51. Use of Bergey's Manual
and Indentibacter
interactus
© The McGraw-H
Companies, 2001
Exercise 51 • Use of Bergey'j Manual and Identibacter Interactus
(see Group V). Most members of Group I are motile
and differentiation is based primarily on oxygen
needs.
Bacillus Although most of these organisms are aer-
obic, some are facultative anaerobes. Catalase is
usually produced. For comparative characteris-
tics of the 34 species in this genus refer to Table
13.4 on pages 1122 and 1123.
Clostridium While most of members of this genus
are strict anaerobes, some may grow in the pres-
ence of oxygen. Catalase is not usually pro-
duced. An excellent key for presumptive species
identification is provided on pages 1143-1148.
Species characterization tables are also provided
on pages 1149-1154.
Sporolactobacillus Microaerophilic and catalase-
negative. Nitrates are not reduced and indole is
not formed. Spore formation occurs very infre-
quently (1% of cells).
Since there is only one species in this genus,
one needs only to be certain that the unknown is
definitely of this genus. Table 13.11 on page
1140 can be used to compare other genera that
are similar to this one.
Group II (Section 16, Vol. 2) This group consists of
Family Mycobacteriaceae, with only one genus:
Mycobacterium. Fifty-four species are listed in
Section 16. Differentiation of species within this
group depends to some extent on whether the organ-
ism is classified as a slow or a fast grower. Tables on
pages 1439-1442 can be used for comparing the char-
acteristics of the various species.
Group III (Section 14, Vol. 2) Of the seven diverse
genera listed in Section 14, only three have been in-
cluded here in this group.
Lactobacillus Non- spore-forming rods, varying
from long and slender to coryneform (club-
shaped) coccobacilli. Chain formation is com-
mon. Only rarely motile. Facultative anaerobic
or microaerophilic. Catalase-negative. Nitrate
usually not reduced. Gelatin not liquefied.
Indole and H 2 S not produced.
Listeria Regular, short rods with rounded ends; oc-
cur singly and in short chains. Aerobic and fac-
ultative anaerobic. Motile when grown at 20-25°
C. Catalase-positive and oxidase-negative.
Methyl red positive. Voges-Proskauer positive.
Negative for citrate utilization, indole produc-
tion, urea hydrolysis, gelatinase production, and
casein hydrolysis. Table 14.12 on page 1241 pro-
vides information pertaining to species differen-
tiation in this genus.
Gram-positive
Rods and Cocci
Rods
Spore-former
Non-spore-former
Acid-Fast
Not Acid -Fast
Regular
Pleomorphic
Group I
Bacillus
Clostridium
Sporolactobacillus
Group II
Mycobacterium
Group
Lactobacillus
Listeria
Kurthia
Group IV
Corynebacterium
Propionibacterium
Arthrobacter
Cocci
Spore-former
Non-spore-former
Catalase + Catalase -
rregular & Tetrad Pairs or Chain
Arrangement Arrangement
Group V
Sporosarcina
Group VI
Micrococcus
Planococcus
Staphylococcus
Group VII
Streptococcus
Figure 51.1 Separation outline for gram-positive rods and cocci
178
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
VIM. Identification of
Unknown Bacteria
51. Use of Bergey's Manual
and Indentibacter
interactus
© The McGraw-H
Companies, 2001
Use of Bergeyj Manual and Identibacter Interactus • Exercise 51
Kurthia Regular rods, 2-A micrometers long with
rounded ends; in chains in young cultures; coc-
coidal in older cultures. Strictly aerobic.
Catalase-positive, oxidase-negative. Also neg-
ative for gelatinase production and nitrate re-
duction. Only two species in this genus.
Group IV (Section 15, Vol. 2) Although there are 21
genera listed in this section of Bergey's Manual, only
three genera are described here.
Cory neb acterium Straight to slightly curved rods
with tapered ends. Sometimes club-shaped.
Palisade arrangements common due to snapping
division of cells. Metachromatic granules
formed. Facultative anaerobic. Catalase-positive.
Most species produce acid from glucose and
some other sugars. Often produce pellicle in
broth. Table 15.3 on page 1269 provides informa-
tion for species characterization.
Proprionibacterium Pleomorphic rods, often
diphtheroid or club-shaped with one end
rounded and the other tapered or pointed. Cells
may be coccoid, bifid (forked, divided), or even
branched. Nonmotile. Some produce clumps of
cells with "Chinese character" arrangements.
Anaerobic to aerotolerant. Generally catalase-
positive. Produce large amounts of proprionic
and acetic acids. All produce acid from glucose.
Arthrobacter Gram-positive rod and coccoid
forms. Pleomorphic. Growth often starts out as
rods, followed by shortening as growth contin-
ues, and finally becoming coccoidal. Some V
and angular forms; branching by some. Rods
usually nonmotile; some motile. Oxidative,
never fermentative. Catalase-positive. Little or
no gas produced from glucose or other sugars.
Type species is Arthrobacter globiformis. For
species differentiation see tables on pages 1 294
and 1 295 .
Group V (Section 13, Vol. 2) This group, which has
only one genus in it, is closely related to genus
Bacillus.
Sporosarcina Cells are spherical or oval when sin-
gle. Cells may adhere to each other when divid-
ing to produce tetrads or packets of eight or more.
Endospores formed (see photomicrographs on
page 1203). Strictly aerobic. Generally motile.
Only two species: S. ureae and S. halophila.
Group VI (Section 12, Vol. 2) This section contains
two families and 15 genera. Our concern here is with
only three genera in this group. Oxygen requirements
and cellular arrangement are the principal factors in
differentiating the genera. Most of these genera are
not closely related.
Micrococcus Spheres, occurring as singles, pairs, ir-
regular clusters, tetrads, or cubical packets.
Usually nonmotile. Strict aerobes (one species is
facultative anaerobic). Catalase- and oxidase-pos-
itive. Most species produce carotenoid pigments.
All species will grow in media containing 5%
NaCl. For species differentiation see Table 12.4 on
page 1007.
Gram-negative
Rods and Cocci
Rods
Cocci
Aerobic
Facultative Anaerobic
Motile
Nonmotile
Group VIM
Pseudomonas
Alcaligenes
Halobactehum
Flavobactehum
Group IX
Escherichia Proteus
Enterobacter Providencia
Citrobacter Morganella
Erwinia Salmonella
Group X
Shigella
Klbesiella
Group XI
Neisseria
Veillonella
Figure 51.2 Separation outline for gram-negative rods and cocci
179
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
VIM. Identification of
Unknown Bacteria
51. Use of Bergey's Manual
and Indentibacter
interactus
© The McGraw-H
Companies, 2001
Exercise 51 • Use of Bergey^ Manual and Identibacter Interactus
Planococcus Spheres, occurring singly, in pairs,
in groups of three cells, occasionally in
tetrads. Although cells are generally gram-
positive, they may be gram-variable. Motility
is present. Catalase- and gelatinase-positive.
Carbohydrates not attacked. Do not hydrolyze
starch or reduce nitrate. Refer to Table 12.9 on
page 1013 for species differentiation.
Staphylococcus Spheres, occurring as singles,
pairs, and irregular clusters. Nonmotile.
Facultative anaerobes. Usually catalase-posi-
tive. Most strains grow in media with 10%
NaCl. Susceptible to lysis by lysostaphin.
Glucose fermentation: acid, no gas. Coagulase
production by some. Refer to Exercise 78 for
species differentiation, or to Table 12.10 on
pages 1016 and 1017.
Group VII (Section 1 2, Vol. 2) Note that the single
genus of this group is included in the same section of
Bergey's Manual as the three genera in group VI.
Members of the genus Streptococcus have spherical to
ovoid cells that occur in pairs or chains when grown in
liquid media. Some species, notably, S. mutans, will de-
velop short rods when grown under certain circum-
stances. Facultative anaerobes. Catalase- negative.
Carbohydrates are fermented to produce lactic acid
without gas production. Many species are commen-
sals or parasites of humans or animals. Refer to
Exercise 79 for species differentiation of pathogens.
Several tables in Bergey's Manual provide differenti-
ation characteristics of all the streptococci.
Group VIII (Section 4, Vol. 1) Although there are
many genera of gram-negative aerobic rod- shaped
bacteria, only four genera are likely to be encoun-
tered here.
Pseudomonas Generally motile. Strict aerobes.
Catalase-positive. Some species produce solu-
ble fluorescent pigments that diffuse into the
agar of a slant. Many tables are available in
Bergey's Manual for species differentiation.
Alcaligenes Rods, coccal rods, or cocci. Motile.
Obligate aerobes with some strains capable of
anaerobic respiration in presence of nitrate or
nitrite.
Halobacterium Cells may be rod- or disk-shaped.
Cells divide by constriction. Most are strict aer-
obes; a few are facultative anaerobes. Catalase-
and oxidase-positive. Colonies are pink, red, or
red to orange. Gelatinase not produced. Most
species require high NaCl concentrations in me-
dia. Cell lysis occurs in hypotonic solutions.
Flavobacterium Gram-negative rods with parallel
sides and rounded ends. Nonmotile. Oxidative.
Catalase-, oxidase-, and phosphatase-positive.
Growth on solid media is typically pigmented
yellow or orange. Nonpigmented strains do ex-
ist. For differentiation see tables on pages 356
and 357.
Groups IX and X (Section 5, Vol. 1) Section 5 in
Bergey 's Manual lists three families and 34 genera; of
these 34 only 10 genera of Family Enterobacteriaceae
MOTILITY
Motile
Nonmotile
C it rate +
Citrate
C it rate +
Citrate
Lactose +
Lactose
Lactose+
Urease-
Klebsiella
Shigella
Lactose -
Urease +
MR+
MR
MR+
MR
Escherichia Proteus
Providencia
Morganella
Citrobacter Enterobacter
Salmonella
Erwinia
Figure 51.3 Separation outline for groups IX and X
180
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
VIM. Identification of
Unknown Bacteria
51. Use of Bergey's Manual
and Indentibacter
interactus
© The McGraw-H
Companies, 2001
Use of Bergey'** Manual and Identibacter Interactus • Exercise 51
have been included in these two groups. If your un-
known appears to fall into one of these groups, use the
separation outline in figure 51.3 to determine the
genus. Another useful separation outline is provided in
figure 80.1 on page 270. Keep in mind, when using
these separation outlines, that there are some minor
exceptions in the applications of these tests. The di-
versity of species within a particular genus often pre-
sents some problematical exceptions to the rule. Your
final decision can be made only after checking the
species characteristics tables for each genus in
Bergey 's Manual.
Group XI These genera are morphologically quite
similar, yet physiologically quite different.
Neisseria (Section 4, Vol. 1) Cocci, occurring
singly, but more often in pairs (diplococci); ad-
jacent sides are flattened. One species (N. elon-
gata) consists of short rods. Nonmotile. Except
for N. elongata, all species are oxidase- and
catalase-positive. Aerobic.
Veillonella (Section 8, Vol. 1) Cocci, appearing as
diplococci, masses, and short chains. Diplococci
have flattening at adjacent surfaces. Nonmotile.
All are oxidase- and catalase-negative. Nitrate is
reduced to nitrite. Anaerobic.
Problem Analysis
If you have identified your unknown by following the
above procedures, congratulations! Not everyone suc-
ceeds at first attempt. If you are having difficulty, con-
sider the following possibilities:
• You may have been given the wrong unknown!
Although this is a remote possibility, it does hap-
pen at times. Occasionally, clerical errors are
made when unknowns are put together.
• Your organism may be giving you a ' 'false-negative' '
result on a test. This may be due to an incorrectly
prepared medium, faulty test reagents, or improper
testing technique.
• Your unknown organisms may not match the de-
scription exactly as stated in Bergey 's Manual. By
now you are aware that the words generally, usu-
ally, and sometimes are frequently used in the
book. It is entirely possible for one of these words
to be inadvertently left out in Bergey's assign-
ment of certain test results to a species. In other
words, test results, as stated in the manual, may
not always apply!
Your culture may be contaminated. If you are not
working with a pure culture, all tests are unreliable.
You may not have performed enough tests. Check
the various tables in Bergey's Manual to see if
there is some other test that will be helpful. In ad-
dition, double check the tables to make sure that
you have read them correctly.
Confirmation of Results
There are several ways to confirm your presumptive
identification. One method is to apply serological
techniques, if your organism is one for which typing
serum is available. Another alternative is to use one of
the miniature multitest systems that are described in
the next section of this manual. Your instructor will
indicate which of these alternatives, if any, will be
available.
Identibacter Interactus
Identibacter interactus is a computer simulation pro-
gram on a CD-ROM disc that was developed by Allan
Konopka, Paul Furbacher, and Clark Gedney at
Purdue University. The program is copyrighted by the
Purdue Research Foundation, and McGraw-Hill Co.
has exclusive distribution rights.
If your laboratory is set up with a computer that
has this program installed in it, use the program to
confirm the identification of your unknown. A group
of 50 tests can be pulled down from menus. The tests
are shown in color exactly as you would see them
when performed in the laboratory. It is your responsi-
bility to interpret each test as applied to your un-
known. Before you attempt to use this program, read
the comments pertaining to it in Appendix F.
Laboratory Report
There is no Laboratory Report for this exercise
181
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
IX. Miniaturized Multitest
Systems
Introduction
© The McGraw-H
Companies, 2001
Part
Miniaturized Multitest Systems
Having run a multitude of tests in Exercises 45 through 50 in an at-
tempt to identify an unknown, you undoubtedly have become aware
of the tremendous amount of media, glassware, and preparation
time that is involved just to set up the tests. And then, after perform-
ing all of the tests and meticulously following all the instructions, you
discover that finding the specific organism in "Encyclopedia Bergey"
is not exactly the simplest task you have accomplished in this
course. The question must arise occasionally: "There's got to be an
easier way!" Fortunately, there is: miniaturized multitest systems.
Miniaturized systems have the following advantages over the
macromethods you have used to study the physiological charac-
teristics of your unknown: (1) minimum media preparation, (2) sim-
plicity of performance, (3) reliability, (4) rapid results, and (5) uniform
results. These advantages have resulted in widespread acceptance
of these systems by microbiologists.
Since it is not possible to describe all of the systems that are
available, only four have been selected here: two by Analytab
Products and two by Becton-Dickinson. All four of these products
are designed specifically to provide rapid identification of medically
important organisms, often within 5 hours. Each method consists
of a plastic tube or strip that contains many different media to be
inoculated and incubated. To facilitate rapid identification, these
systems utilize numerical coding systems that can be applied to
charts or computer programs.
The four multitest systems described in this unit have been se-
lected to provide several options. Exercises 52 and 53 pertain to the
identification of gram-negative oxidase-negative bacteria (Entero-
bacteriaceae). Exercise 54 (Oxi/Ferm Tube) is used for identifying
gram-negative oxidase-positive bacteria. Exercise 55 (Staph-ldent)
is a rapid system for the differentiation of the staphylococci.
As convenient as these systems are, one must not assume that
the conventional macromethods of Part 8 are becoming obsolete.
Macromethods must still be used for culture studies and confirma-
tory tests; confirmatory tests by macromethods are often neces-
sary when a particular test on a miniaturized system is in question.
Another point to keep in mind is that all of the miniaturized multi-
test systems have been developed for the identification of med-
ically important microorganisms. If one is trying to identify a sapro-
phytic organism of the soil, water, or some other habitat, there is no
substitute for the conventional methods.
If these systems are available to you in this laboratory, they may
be used to confirm your conclusions that were drawn in Part 8 or
they may be used in conjunction with some of the exercises in Part
14. Your instructor will indicate what applications will be made.
183
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
IX. Miniaturized Multitest
Systems
52. Enterobacteriaceae
Identification: The API 20E
System
© The McGraw-H
Companies, 2001
Enterobacteriaceae Identification
Tke API 20E System
52
The API 20E System is a miniaturized version of
conventional tests that is used for the identification of
members of the family Enterobacteriaceae and other
gram-negative bacteria. It was developed by
Analytab Products, of Plainview, New York. This
system utilizes a plastic strip (figure 52.1) with 20
separate compartments. Each compartment consists
of a depression, or cupule, and a small tube that con-
tains a specific dehydrated medium (see illustration
4, figure 52.2). The system has a capacity of 23 bio-
chemical tests.
To inoculate each compartment it is necessary to
first make up a saline suspension of the unknown or-
ganism; then, with the aid of a Pasteur pipette, each
compartment is filled with the bacterial suspension.
The cupule receives the suspension and allows it to
flow into the tube of medium. The dehydrated
medium is reconstituted by the saline. To provide
anaerobic conditions for some of the compartments it
is necessary to add sterile mineral oil to them.
After incubation for 18-24 hours, the reactions
are recorded, test reagents are added to some com-
partments, and test results are tabulated. Once the test
results are tabulated, & profile number (7 or 9 digits) is
computed. By finding the profile number in a code
book, the Analytical Profile Index, one is able to de-
termine the name of the organism. If no Analytical
Profile Index is available, characterization can be
done by using Chart ffl in Appendix D.
Although this system is intended for the identifica-
tion of nonenterics, as well as the Enterobacteriaceae,
only the identification of the latter will be pursued in
this experiment. Proceed as follows to use the API 20E
System to identify your unknown enteric.
First Period
Two things will be accomplished during this period:
(1) the oxidase test will be performed if it has not been
previously performed, and (2) the API 20E test strip
will be inoculated. All steps are illustrated in figure
52.2. Proceed as follows to use this system:
Materials:
agar slant or plate culture of unknown
test tube of 5 ml 0.85% sterile saline
Figure 52.1 Positive and negative test results on API 20E test strips
185
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
IX. Miniaturized Multitest
Systems
52. Enterobacteriaceae
Identification: The API 20E
System
© The McGraw-H
Companies, 2001
Exercise 52 • Enterobacteriaceae Identification: The API 20E System
1
2
3
4
5
6
7
8
9
API 20E test strip
API incubation tray and cover
squeeze bottle of tap water
test tube of 5 ml sterile mineral oil
Pasteur pipettes (5 ml size)
oxidase test reagent
Whatman no. 2 filter paper
empty Petri dish
Vortex mixer
If you haven't already done the oxidase test on
your unknown, do so at this time. It must be es-
tablished that your unknown is definitely oxidase-
negative before using this system. Use the filter
paper method that is described on page 168.
Prepare a saline suspension of your unknown by
transferring organisms from the center of a well-
established colony on an agar plate (or from a
slant culture) to a tube of 0.85% saline solution.
Disperse the organisms well throughout the
saline.
Label the end strip of the API 20E tray with your
name and unknown number. See illustration 2,
figure 52.2.
Dispense about 5 ml of tap water into the tray
with a squeeze bottle. Note that the bottom of
the tray has numerous depressions to accept the
water.
Remove an API 20E test strip from the sealed
pouch and place it into the tray (see illustration 3).
Be sure to reseal the pouch to protect the remain-
ing strips.
Vortex mix the saline suspension to get uniform
dispersal, and fill a sterile Pasteur pipette with the
suspension. Take care not to spill any of the or-
ganisms on the table or yourself. You may have a
pathogen!
Inoculate all the tubes on the test strip with the
pipette by depositing the suspension into the
cupules as you tilt the API tray (see illustration 4,
figure 52.2).
Important: Slightly underfill ADR LDC ODC
H 2 S and URE. (Note that the labels for these
compartments are underlined on the strip.)
Underfilling these compartments leaves room for
oil to be added and facilitates interpretation of the
results.
Since the media in ICITL IVPL and I GEL I com-
partments require oxygen, completely fill both the
cupule and tube of these compartments. Note that
the labels on these three compartments are brack-
eted as shown here.
To provide anaerobic conditions for the ADH,
LDC, ODC H^L and URE compartments, dis-
pense sterile mineral oil to the cupules of these
compartments. Use another sterile Pasteur pipette
for this step.
10. Place the lid on the incubation tray and incubate
at 37° C for 1 8 to 24 hours. Refrigeration after in-
cubation is not recommended.
Second Period
(Evaluation of Tests)
During this period all reactions will be recorded on
the Laboratory Report, test reagents will be added to
four compartments, and the seven-digit profile num-
ber will be determined so that the unknown can be
looked up in the API 20E Analytical Profile Index.
Proceed as follows:
Materials:
incubation tray with API 20E test strip
10% ferric chloride
Barritt's reagents A and B
Ko vacs' reagent
nitrite test reagents A and B
zinc dust or 20-mesh granular zinc
hydrogen peroxide (1.5%)
API 20E Analytical Profile Index
Pasteur pipettes
1
2
3
4
5
6
7
Before any test reagents are added to any of the
compartments, consult Chart I, Appendix D, to
determine the nature of positive reactions of each
test, except TDA, VP, and IND.
Refer to Chart II, Appendix D, for an explanation
of the 20 symbols that are used on the plastic test
strip .
Record the results of these tests on the Laboratory
Report.
If GLU test is negative (blue or blue-green), and
there are fewer than three positive reactions
before adding reagents, do not progress any fur-
ther with this test as outlined here in this experi-
ment. Organisms that are GLU-negative are
nonenterics.
For nonenterics, additional incubation time is
required. If you wish to follow through on an or-
ganism of this type, consult your instructor for
more information.
If GLU test is positive (yellow), or there are
more than three positive reactions, proceed to
add reagents as indicated in the following steps.
Add one drop of 10% ferric chloride to the TDA
tube. A positive reaction (brown-red), if it occurs,
will occur immediately A negative reaction color
is yellow.
Add one drop each of Barritt's A and B solutions
to the VP tube. Read the VP tube within 10 min-
186
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
IX. Miniaturized Multitest
Systems
52. Enterobacteriaceae
Identification: The API 20E
System
© The McGraw-H
Companies, 2001
Enterobacteriaceae Identification: The API 20E System • Exercise 52
0.85% Saline
Select one well-isolated colony to make a saline
suspension of the unknown organism. Suspension
should be well dispersed with a Vortex mixer.
Place an API 20E test strip into the bottom of the
moistened tray. Be sure to seal the pouch from which
the test strip was removed to prevent contamination
of remaining strips.
After labeling the end tab of a tray with your name
and unknown number, dispense approximately 5 ml
of tap water into bottom of tray.
Tube
Cupule
OIL
<:
^J
\J
\_/
^>
\J
^J
888
\*J
ffi5A£H UPC OpC gr H g § UW| Ttft INT? Jg_
r
If
To provide anaerobic conditions for chambers ADH,
LDC, ODC, H 2 S, and URE, completely fill cupules
of these chambers with sterile mineral oil. Use a fresh
sterile Pasteur pipette.
Dispense saline suspension of organisms into cupules
of all twenty compartments. Slightly underfill ADH,
LDC, ODC, H 2 S, and URE. Completely ////cupules
of CIT, VP, and GEL.
0NP6
\
ADH
Z
LDC
4
4-
-h
ooc
+
CIT
z
H,S
URE
I
TDA
_2__
IND
4
+
VP
GEL
GUU
4
+
MAN
+
After incubation and after adding test reagents to
four compartments, record all results and total
numbers to arrive at 7-digit code. Consult the
Analytical Profile Index to find the unknown.
Figure 52.2 The API 20E procedure
187
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
IX. Miniaturized Multitest
Systems
52. Enterobacteriaceae
Identification: The API 20E
System
© The McGraw-H
Companies, 2001
Exercise 52 • Enterobacteriaceae Identification: The API 20E System
utes. The pale pink color that occurs immediately
has no significance. A positive reaction is dark
pink or red and may take 1 minutes before it ap-
pears.
8. Add one drop of Ko vacs' reagent to the IND
tube. Look for a positive (red ring) reaction
within 2 minutes.
After several minutes the acid in the reagent
reacts with the plastic cupule to produce a color
change from yellow to brownish-red, which is
considered negative.
9. Examine the GLU tube closely for evidence of
bubbles. Bubbles indicate the reduction of nitrate
and the formation of N 2 gas. Note on the
Laboratory Report that there is a place to record
the presence of this gas.
10. Add two drops of each nitrite test reagent to the
GLU tube. A positive (red) reaction should show
up within 2 to 3 minutes if nitrates are reduced.
If this test is negative, confirm negativity
with zinc dust or 20-mesh granular zinc. A pink-
orange color after 10 minutes confirms that ni-
trate reduction did not occur. A yellow color re-
sults if N 2 was produced.
1 1 . Add one drop of hydrogen peroxide to each of
the MAN, INO, and SOR cupules. If catalase is
produced, gas bubbles will appear within 2 min-
utes. Best results will be obtained in tubes that
have no gas from fermentation.
Final Confirmation
After all test results have been recorded and the
seven-digit profile number has been determined, ac-
cording to the procedures outlined on the Laboratory
Report, identify your unknown by looking up the pro-
file number in the API 20E Analytical Profile Index.
Cleanup
When finished with the test strip be sure to place it in
a container of disinfectant that has been designated
for test strip disposal.
188
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
IX. Miniaturized Multitest
Systems
53. Enterobacteriaceae
Identification: The
Enterotube II System
© The McGraw-H
Companies, 2001
Enterobacteriaceae Identification
Tke Enterotube II System
53
The Enterotube II miniaturized multitest system
was developed by Becton-Dickinson of Cock-
eysville, Maryland, for rapid identification of
Enterobacteriaceae. It incorporates 12 different
conventional media and 15 biochemical tests into a
single ready-to-use tube that can be simultaneously
inoculated in a moment's time with a minimum of
equipment.
If you have an unknown gram-negative rod or
coccobacillus that appears to be one of the Entero-
bacteriaceae, you may wish to try this system on it.
Before applying this test, however, make certain that
your unknown is oxidase -negative, since with only a
few exceptions, all Enterobacteriaceae are oxidase-
negative. If you have a gram-negative rod that is oxi-
dase-positive you might try the Oxi/Ferm Tube II in-
stead, which is featured in the next exercise.
Figure 53.1 illustrates an uninoculated tube (up-
per) and a tube with all positive reactions (lower).
Figure 53.2 outlines the entire procedure for utilizing
this system.
Each of the 1 2 compartments of an Enterotube II
contains a different agar-based medium. Compartments
that require aerobic conditions have openings for access
to air. Those compartments that require anaerobic con-
ditions have layers of paraffin wax over the media.
Extending through all compartments of the entire tube is
an inoculating wire. To inoculate the media, one simply
picks up some organisms on the end of the wire and
pulls the wire through each of the chambers in a single,
rotating action.
After incubation, the reactions in all the compart-
ments are noted and the indole test is performed. The
Voges-Proskauer test may also be performed as a
confirmation test. Positive reactions are given nu-
merical values, which are totaled to arrive at a five-
digit code. Identification of the unknown is achieved
by consulting a coding manual, the Enterotube II
Interpretation Guide, which lists these numerical
codes for the Enterobacteriaceae. Proceed as follows
to use an Enterotube II in the identification of your
unknown.
UNINOCULATED
COLORS
REACTED
COLORS
#
#
a-
*
#
&
#
$
s-
!3'-«&-
<^<^
4?
#
^
#
^ £
^
<r<r <r<r ^
-?
#
^
/ /
^ A J?
&
## ^
S
a*
^
^
r<?
*
^
GAS PRODUCTION
INDOLE
VOGES-
PROSKAUER
PA
Figure 53.1 Enterotube II color differences between uninoculated and positive tests
Courtesy of Becton-Dickinson, Cockeysville, Maryland.
189
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
IX. Miniaturized Multitest
Systems
53. Enterobacteriaceae
Identification: The
Enterotube II System
© The McGraw-H
Companies, 2001
Exercise 53 • Enterobacteriaceae Identification: The Enterotube II System
First Period
Inoculation and Incubation
The Enterotube II can be used to identify
Enterobacteriaceae from colonies on agar that have
been inoculated from urine, blood, sputum, etc. The
culture may be taken from media such as MacConkey,
EMB, SS, Hektoen enteric, or trypticase soy agar.
Materials:
culture plate of unknown
1 Enterotube II
1
2
3
4
5
6
7
8
9
Write your initials or unknown number on the
white paper label on the side of the tube.
Unscrew both caps from the Enterotube II. The tip
of the inoculating end is under the white cap.
Without heat-sterilizing the exposed inoculating
wire, insert it into a well-isolated colony
Inoculate each chamber by first twisting the wire
and then withdrawing it through all 12 compart-
ments. Rotate the wire as you pull it through. See
illustration 2, figure 53.2.
Again, without sterilizing, reinsert the wire, and
with a turning motion, force it through all 1 2 com-
partments until the notch on the wire is aligned
with the opening of the tube. (The notch is about
VA from handle end of wire.) The tip of the wire
should be visible in the citrate compartment. See
illustration 3, figure 53.2.
Break the wire at the notch by bending, as shown
in step 4, figure 53.2. The portion of the wire re-
maining in the tube maintains anaerobic condi-
tions essential for fermentation of glucose, pro-
duction of gas, and decarboxylation of lysine and
ornithine.
With the retained portion of the needle, punch
holes through the thin plastic coverings over the
small depressions on the sides of the last eight
compartments (adonitol, lactose, arabinose, sor-
bitol, Voges-Proskauer, dulcitol/PA, urea, and cit-
rate). These holes will enable aerobic growth in
these eight compartments.
Replace the caps at both ends.
Incubate at 35° to 37° C for 18 to 24 hours with
the Enterotube II lying on its flat surface. When
incubating several tubes together, allow space be-
tween them to allow for air circulation.
Second Period
Reading Results
Reading the results on the Enterotube may be done in
one of two ways: (1) by simply comparing the results
with information on Chart IV, Appendix D, or (2) by
finding the five-digit code number you compute for
your unknown in the Enterotube II Interpretation
Guide. Of the two methods, the latter is much pre-
ferred. The chart in the appendix should be used only
if the Interpretation Guide is not available.
Whether or not the Interpretation Guide is
available, these three steps will be performed during
this period to complete this experiment: (1) positive
test results must first be recorded on the Laboratory
Report, (2) the indole test, a presumptive test, is per-
formed on compartment 4, and (3) confirmatory
tests, if needed, are performed. The Voges-
Proskauer test falls in the latter category. Proceed as
follows:
Materials:
Enterotube II, inoculated and incubated
Ko vacs' reagent
10% KOH with 0.3% creatine solution
5% alpha- naphthol in absolute ethyl alcohol
syringes with needles, or disposable Pasteur
pipettes
test-tube rack
Enterotube II Results Pad (optional)
coding manual: Enterotube II Interpretation
Guide
1
2
3
4
5
6
Compare the colors of each compartment of your
Enterotube II with the lower tube illustrated in
figure 53.1.
With a pencil, mark a small plus ( + ) or minus ( — )
near each compartment symbol on the white label
on the side of the tube.
Consult table 53.1 for information as to the sig-
nificance of each compartment label.
Record the results of the tests on the Laboratory
Report. All results must be recorded before doing
the indole test.
Record results on the Laboratory Report.
Important: If at this point you discover that
your unknown is GLU-negative, proceed no fur-
ther with the Enterotube II because your un-
known is not one of the Enterobacteriaceae.
Your unknown may be Acinetobacter sp. or
Pseudomonas maltophilia. If an Oxi/Ferm Tube
is available, try it, using the procedure outlined
in the next exercise.
Indole Test: Perform the indole test as follows:
a. Place the Enterotube II into a test-tube rack
with the GLU-GAS compartment pointing up-
ward.
b. Inject one or two drops of Ko vacs' reagent
onto the surface of the medium in the H 2 S/
indole compartment. This may be done with a
syringe and needle through the thin Mylar
plastic film that covers the flat surface, or with
a disposable Pasteur pipette through a small
hole made in the Mylar film with a hot inocu-
lating needle.
190
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
IX. Miniaturized Multitest
Systems
53. Enterobacteriaceae
Identification: The
Enterotube II System
© The McGraw-H
Companies, 2001
Remove organisms from a well-isolated colony. Avoid
touching the agar with the wire. To prevent damaging
Enterotube II media, do not heat-sterilize the inoculat-
ing wire.
noculate each compartment by first twisting the wire
and then withdrawing it all the way out through the 1 2
compartments, using a turning movement.
Reinsert the wire (without sterilizing), using a turning
motion through all 12 compartments until the notch on
the wire is aligned with the opening of the tube.
Break the wire at the notch by bending. The portion of
the wire remaining in the tube maintains anaerobic con
ditions essential for true fermentation.
Punch holes with broken off part of wire through the thin
plastic covering over depressions on sides of the last
eight compartments (adonitol through citrate). Replace
caps and incubate at 35° C for 18-24 hours.
After interpreting and recording positive results on the
sides of the tube, perform the indole test by injecting 1
or 2 drops of Kovacs' reagent into the H 2 S/lndole
compartment.
N
H ? S
I
N
3
A
D
N
L
■
C
I A
R
A
a
S
R
B
D
U
L
P
A
U
R
i E
A
©+ J
®+ z + (D J
i 4 +©+(Dj
[<s + © +■
* * ^*
♦
<^
>
< )
>
Perform the Voges-Proskauer test, if needed for con-
firmation, by injecting the reagents into the H 2 S/indole
compartment.
After encircling the numbers of the positive tests
on the Laboratory Report, total up the numbers of each
bracketed series to determine the 5-digit code number.
Refer to the Enterotube II Interpretation Guide for iden-
tification of the unknown by using the code number.
Figure 53.2 The Enterotube II procedure
191
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
IX. Miniaturized Multitest
Systems
53. Enterobacteriaceae
Identification: The
Enterotube II System
© The McGraw-H
Companies, 2001
Exercise 53 • Enterobacteriaceae Identification: The Enterotube II System
c. A positive test is indicated by the development
of a red color on the surface of the medium or
Mylar film within 10 seconds.
7. Voges-Proskauer Test: Since this test is used as
a confirmatory test, it should be performed only
when called for in the Enterotube II Interpretation
Guide. If it is called for, perform the test in the fol-
lowing manner:
a. Use a syringe or Pasteur pipette to inject two
drops of potassium hydroxide containing crea-
tine into the V-P section.
8.
b. Inject three drops of 5% alpha-naphthol.
c. A positive test is indicated by a red color
within 10 minutes.
Record the indole and V-P results on the
Laboratory Report.
Laboratory Report
Determine the name of your unknown by following the
instructions on the Laboratory Report. Note that two
methods of making the final determination are given.
Table 53.1 Biochemical Reactions of Enterotube II
SYMBOL
UNINOCULATED REACTED
COLOR
COLOR
TYPE OF REACTION
GLU-GAS
Glucose (GLU) The end products of bacterial fermentation of
glucose are either acid or acid and gas. The shift in pH due to
the production of acid is indicated by a color change from red
(alkaline) to yellow (acidic). Any degree of yellow should be
interpreted as a positive reaction; orange should be considered
negative.
Gas Production (GAS) Complete separation of the wax overlay
from the surface of the glucose medium occurs when gas is
produced. The amount of separation between the medium and
overlay will vary with the strain of bacteria.
LYS
ORN
H 2 S/IND
Lysine Decarboxylase Bacterial decarboxylation of lysine,
which results in the formation of the alkaline end product
cadaverine, is indicated by a change in the color of the indi-
cator from pale yellow {acidic) to purple (alkaline). Any degree of
purple should be interpreted as a positive reaction. The medium
remains yellow if decarboxylation of lysine does not occur.
Ornithine Decarboxylase Bacterial decarboxylation of ornithine
causes the alkaline end product putrescine to be produced. The
acidic (yellow) nature of the medium is converted to purple as
alkalinity occurs. Any degree of purple should be interpreted as
a positive reaction. The medium remains yellow if decarboxyla-
tion of ornithine does not occur.
H2S Production Hydrogen sulfide, liberated by bacteria that
reduce sulfur-containing compounds such as peptones and
sodium thiosulfate. reacts with the iron salts in the medium to
form a black precipitate of ferric sulfide usually along the line
of inoculation. Some Proteus and Providencia strains may pro-
duce a diffuse brown coloration in this medium, which should
not be confused with true H2S production.
Indole Formation The production of indole from the metab-
olism of tryptophan by the bacterial enzyme tryptophanase is
detected by the development of a pink to red color after the
addition of Kovac's reagent.
192
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
IX. Miniaturized Multitest
Systems
53. Enterobacteriaceae
Identification: The
Enterotube II System
© The McGraw-H
Companies, 2001
Enterobacteriaceae Identification: The Enterotube II System • Exercise 53
Table 53.1 Biochemical Reactions of Enterotube II (continued)
UNINOCULATED REACTED
SYMBOL COLOR COLOR TYPE 0F REACTION
ADON
Adonitol Bacterial fermentation of adonitol, which results in
the formation of acidic end products, is indicated by a change
in color of the indicator present in the medium from red
{alkaline) to yellow (acidic). Any sign of yellow should be inter-
preted as a positive reaction; orange should be considered
neqative.
LAC
Lactose Bacterial fermentation of lactose, which results in the
formation of acidic end products, is indicated by a change in
color of the Indicator present in the medium from red (alkaline)
to yellow (acidic). Any sign of yellow should be interpreted as a
positive reaction; orange should be considered negative,
ARAB
Arabinose Bacterial fermentation of arabinose, which results in
the formation of acidic end products, is indicated by a change
in color from red (alkaline) to yellow (acidic). Any sign of yellow
should be interpreted as a positive reaction; orange should be
considered negative.
SORB
Sorbitol Bacterial fermentation of sorbitol, which results in the
formation of acidic end products, is indicated by a change in
color from red (alkaline) to yellow (acidic). Any sign of yellow
should be interpreted as a positive reaction; orange should be
considered negative.
V.P.
\
Voges-Proskauer Acetyl methyl carbi no 1 (acetoin) is an inter-
mediate in the production of butylene glycol from glucose fer-
mentation. The presence of acetoin is indicated by the develop-
ment of a red color within 20 minutes. Most positive reactions i
are evident within 10 minutes.
Dulcitol Bacterial fermentation of dulcitol, which results in the
formation of acidic end products, is indicated by a change in
color of the indicator present in the medium from green
(alkaline) to yellow or pale yellow (acidic).
DUL-PA
Phenylalanine Deaminase This test detects the formation of
pyruvic acid from the deamination of phenylalanine. The pyruvic
acid formed reacts with a ferric salt in the medium to produce a
characteristic black to smoky gray color.
UREA
Urea The production of urease by some bacteria hydrolyzes
urea in this medium to produce ammonia, which causes a shift
in pH from yellow (acidic) to reddish-purple (alkaline). This test
is strongly positive for Proteus in 6 hours and weakly positive
for Klebsiella and some Enterobacter species in 24 hours.
CIT
Citrate Organisms that are able to utilize the citrate in this
medium as their sole source of carbon produce alkaline metabo-
lites that change the color of the indicator from green (acidic)
to deep blue (alkaline). Any degree of blue should be considered
positive.
Courtesy of Becton-Dickinson, Cockeysville, Maryland
193
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
IX. Miniaturized Multitest
Systems
54. O/F Gram-Negative
Rods Identification: The
Oxi/Ferm Tube II System
© The McGraw-H
Companies, 2001
54
O/F Gram-Negative Rods Identification:
The Oxi/Ferm Tube II System
The Oxi/Ferm Tube II, produced by Becton-Dickinson,
takes care of the identification of the oxidase-positive,
gram-negative bacteria that cannot be identified by us-
ing the Enterotube II system. The two multitest systems
were developed to work together. If an unknown gram-
negative rod is oxidase-negative, the Enterotube II is
used. If the organism is oxidase-positive, the Oxi/Ferm
Tube II must be used. Whenever an oxidase-negative
gram-negative rod turns out to be glucose-negative on
the Enterotube II test, one must move on to use the
Oxi/Ferm Tube II.
The Oxi/Ferm Tube II system is intended for
the identification of non-fastidious species of
oxidative-fermentative gram-negative rods from
clinical specimens. This includes the following gen-
era: Aeromonas, Plesiomonas, Vibrio, Achromobacter,
Alcaligenes, Bordetella, Moraxella, and Pasteurella.
Some other gram-negative bacteria can also be identi-
fied with additional biochemical tests. The system in-
corporates 1 2 different conventional media that can be
inoculated simultaneously in a moment's time with a
minimum of equipment. A total of 14 physiological
tests are performed.
Like the Enterotube II system, the Oxi/Ferm Tube
II has an inoculating wire that extends through all 1 2
compartments of the entire tube. To inoculate the me-
dia, one simply picks up some organisms on the end
of the wire and pulls the wire through each of the
chambers in a rotating action.
After incubation, the results are recorded and
Kovacs' reagent is injected into one of the compart-
ments to perform the indole test. Positive reactions
are given numerical values that are totaled to arrive at
a five-digit code. By looking up the code in an
Oxi/Ferm Biocode Manual, one can quickly deter-
mine the name of the unknown and any tests that
might be needed to confirm the identification.
Figure 54.1 illustrates an uninoculated tube and a
tube with all positive reactions. Figure 54.2 illustrates
the entire procedure for utilizing this system. A mini-
mum of two periods are required to use this system.
Proceed as follows:
UNINOCULATED
COLORS
REACTED
COLORS
GAS INDOLE
Figure 54.1 Oxi/Ferm Tube II color differences between uninoculated and positive tests
194
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
IX. Miniaturized Multitest
Systems
54. O/F Gram-Negative
Rods Identification: The
Oxi/Ferm Tube II System
© The McGraw-H
Companies, 2001
Remove organisms from a well-isolated colony. Avoid
touching the agar with the wire. To prevent damaging
Enterotube II media, do not heat-sterilize the inoculat-
ing wire.
noculate each compartment by first twisting the wire
and then withdrawing it all the way out through the 12
compartments, using a turning movement.
Reinsert the wire (without sterilizing), using a turning
motion through all 12 compartments until the notch on
the wire is aligned with the opening of the tube.
Break the wire at the notch by bending. The portion of
the wire remaining in the tube maintains anaerobic con
ditions essential for true fermentation.
Punch holes with broken-off part of wire through the thin
plastic covering over depressions on sides of the last
eight compartments (sucrose/indole through citrate).
Replace caps and incubate at 35° C for 18-24 hours.
After interpreting and recording positive results on the
sides of the tube, perform the indole test by injecting 1
or 2 drops of Kovacs' reagent into the sucrose/indole
compartment.
After encircling the numbers of the positive tests on
the Laboratory Report, total up the numbers of each
bracketed series to determine the 5-digit code number.
Refer to the Biocode Manualtor identification of the
unknown by using the code number.
Figure 54.2 The Oxi/Ferm Tube II procedure
195
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
IX. Miniaturized Multitest
Systems
54. O/F Gram-Negative
Rods Identification: The
Oxi/Ferm Tube II System
© The McGraw-H
Companies, 2001
Exercise 54 • O/F Gram-Negative Rods Identification: The Oxi/Ferm Tube II System
First Period
Inoculation and Incubation
The Oxi/Ferm Tube II must be inoculated with a large
inoculum from a well-isolated colony. Culture purity,
of course, is of paramount importance. If there is any
doubt of purity, a TS A plate should be inoculated and
incubated at 35° C for 24 hours, followed by 24 hours
incubation at room temperature. If no growth occurs
on TS A, but growth does occur on blood agar, the or-
ganism has special growth requirements. Such organ-
isms are too fastidious and cannot be identified with
the Oxi/Ferm Tube II.
Materials:
culture plate of unknown
1 Oxi/Ferm Tube II
1 plate of trypticase soy agar (TS A) (for purity
check, if needed)
1
2
3
4
5
6
7
8
9
10
Write your initials or unknown number on the
side of the tube.
Unscrew both caps from the Oxi/Ferm Tube II. The
tip of the inoculating end is under the white cap.
Without heat-sterilizing the exposed inoculating
wire, insert it into a well-isolated colony. Do not
puncture the agar.
Inoculate each chamber by first twisting the wire
and then withdrawing it through all 12 compart-
ments. Rotate the wire as you pull it through. See
illustration 2, figure 54.2.
If a purity check of the culture is necessary,
streak a Petri plate of TSA with the inoculating
wire that has just been pulled through the tube.
Do not flame.
Again, without sterilizing, reinsert the wire, and
with a turning motion, force it through all 1 2 com-
partments until the notch on the wire is aligned
with the opening of the tube. (The notch is about
V/" from the handle end of the wire.) The tip of
the wire should be visible in the citrate compart-
ment. See illustration 3, figure 54.2.
Break the wire at the notch by bending, as noted
in step 4, figure 54.2. The portion of the wire re-
maining in the tube maintains anaerobic condi-
tions essential for true fermentation.
With the retained portion of the needle, punch
holes through the thin plastic coverings over the
small depressions on the sides of the last eight
compartments (sucrose/indole, xylose, aerobic
glucose, maltose, mannitol, phenylalanine, urea,
and citrate). These holes will enable aerobic
growth in these eight compartments.
Replace both caps on the tube.
Incubate at 35° to 37° C for 24 hours, with the
tube lying on its flat surface or upright. At the end
of 24 hours inspect the tube to check results and
continue incubation for another 24 hours. The 24-
hour check may be needed for doing confirmatory
tests as required in the Biocode Manual.
Occasionally, an Oxi/Ferm Tube II should be in-
cubated longer than 48 hours.
Second Period
Evaluation of Tests
During this period you will record the results of the
various tests on your Oxi/Ferm Tube II, do an indole
test, tabulate your results, use the Biocode Manual,
and perform any confirmatory tests called for.
Proceed as follows:
Materials:
Oxi/Ferm Tube II, inoculated and incubated
Ko vacs' reagent
syringes with needles, or disposable Pasteur
pipettes
Becton-Dickinson Biocode Manual (a booklet)
1
2
3
4
5
6
Compare the colors of each compartment of your
Oxi/Ferm Tube II with the lower tube illustrated
in figure 54.1.
With a pencil, mark a small plus ( + ) or minus ( — )
near each compartment symbol on the white label
on the side of the tube.
Consult Table 54.1 for information as to the sig-
nificance of each compartment label.
Record the results of all the tests on the Laboratory
Report. All results must be recorded before doing
the indole test.
Indole Test (illustration 6, figure 54.2): Do an in-
dole test by injecting two or three drops of
Ko vacs' reagent through the flat, plastic surface
into the sucrose/indole compartment. Release the
reagent onto the inside flat surface and allow it to
drop down onto the agar.
If a Pasteur pipette is used instead of a syringe
needle, it will be necessary to form a small hole in
the Mylar film with a hot inoculating needle to
admit the tip of the Pasteur pipette.
A positive test is indicated by the develop-
ment of a red color on the surface of the medium
or Mylar film within 10 seconds.
Record the results of the indole test on the
Laboratory Report.
Laboratory Report
Follow the instructions on the Laboratory Report for
determining the five-digit code. Use the Biocode
Manual booklet for identifying your unknown.
196
Benson: Microbiological
Applications Lab Manual, Systems
Eighth Edition
IX. Miniaturized Multitest
54. O/F Gram-Negative
Rods Identification: The
Oxi/Ferm Tube II System
© The McGraw-H
Companies, 2001
O/F Gram-Negative Rods Identification: The Oxi/Ferm Tube II System • Exercise 54
Table 54.1 Biochemical Reactions of the Oxi/Ferm Tube II
Reaction
Negative
Positive
Special Remarks
Anaerobic
Glucose
Positive fermentation is shown by change in color from green (neutral) to
yellow (acid). Most oxidative-fermentative, gram-negative rods are negative.
Arginine
Dihydrolase
Decarboxylation of arginine results in the formation of alkaline end
products that changes bromcresol purple from yellow (acid) to purple
(alkaline). Grey is negative.
Lysine
Decarboxylation of lysine results in the formation of alkaline end products
that changes bromcresol purple from yellow (acid) to purple (alkaline). Grey
is negative.
Lactose
Fermentation of lactose changes the color of the medium from red (neutral)
to yellow (acid). Most O/F gram-negative rods are negative.
ISh Gas-
production
^
<&
Gas production causes separation of wax overlay from medium.
Occasionally, the gas will also cause separation of the agar from the
compartment wall.
Sucrose
Bacterial oxidation of sucrose causes a change in color from green
(neutral) to yellow (acid).
Indole
The bacterial enzyme tryptophanase metabolizes tryptophan to produce
indole. Detection is by adding Kovacs' reagent to the compartment 48
hours after incubation.
Xylose
Bacterial oxidation of xylose causes a color change of green (neutral) to
yellow (acid).
Aerobic
Glucose
Bacterial oxidation of glucose causes a color change of green (neutral) to
yellow (acid).
Maltose
Bacterial oxidation of maltose causes a color change of green (neutral) to
yellow (acid).
Mannitol
Bacterial oxidation of this carbohydrate is evidenced by a change in color
from green (neutral) to yellow (acid).
Phenylalanine
Pyruvic acid is formed by deamination of phenylalanine. The pyruvic acid
reacts with a ferric salt to produce a brownish tinge.
Urea
The production of ammonia by the action of urease on urea increases the
alkalinity of the medium. The phenol red in this medium changes from
beige (acid) to pink or purple. Pale pink should be considered negative.
Citrate
Organisms that grow on this medium are able to utilize citrate as their sole
source of carbon. Utilization of citrate raises the alkalinity of the medium.
The color changes from green (neutral) to blue (alkaline).
197
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
IX. Miniaturized Multitest
Systems
55. Staphylococcus
Identification: The API
Staph-ldentification
System
© The McGraw-H
Companies, 2001
55
Staphylococcus Identification:
The API Staph-Ident System
The API Staph-Ident System, produced by Analytab
Products of Plainview, New York, was developed to
provide a rapid (5-hour) method for identifying 13 of
the most clinically important species of staphylo-
cocci. This system consists of 10 microcupules that
contain dehydrated substrates and/or nutrient media.
Except for the coagulase test, all the tests that are
needed for the identification of staphylococci are in-
cluded on the strip.
Figure 55.1 illustrates two inoculated strips: the
lower one just after inoculation and the upper one
with all positive reactions. Note that the appearance of
each microcupule undergoes a pronounced color
change when a positive reaction occurs.
Figure 55.2 illustrates the overall procedure. The
first step is to make a saline suspension of the organ-
ism from an isolated colony. A Staph-Ident strip is
then placed in a tray that has a small amount of water
added to it to provide humidity during incubation.
Next, a sterile Pasteur pipette is used to dispense two
to three drops of the bacterial suspension to each mi-
crocupule. The inoculated tray is then covered and in-
cubated aerobically at 35° to 37° C for 5 hours. After
incubation, a few drops of Staph-Ident reagent are
added to the tenth microcupule and the results are read
immediately. Finally, a four-digit profile is computed
that is used to determine the species from a chart in
Appendix D.
As simple as this system might seem, there are a
few limitations that one must keep in mind. Final
species determination by a competent microbiologist
must take into consideration other factors such as the
source of the specimen, the catalase reaction, colony
characteristics, and antimicrobial susceptibility pat-
tern. Very often there are confirmatory tests that must
also be made.
If you have been working with an unknown that
appears to be one of the staphylococci, use this system
to confirm your conclusions. If you have already done
the coagulase test and have learned that your organ-
ism is coagulase-negative, this system will enable you
to identify one of the numerous coagulase-negative
species that are not identifiable by the procedures in
Exercise 78.
ALL TESTS: POSITIVE
STAPH-IDENT
JUST INOCULATED: ALL NEGATIVE
Courtesy of Analytab Products, Plainview, N.Y.
Figure 55.1 Positive and negative results on API Staph-Ident test strips
198
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
IX. Miniaturized Multitest
Systems
55. Staphylococcus
Identification: The API
Staph-ldentification
System
© The McGraw-H
Companies, 2001
nvmmnMiwitwiiiBiitk
0.85% Saline
Use several loopfuls of organisms to make saline
suspension of unknown. Turbidity of suspension
should match McFarland No. 3 barium sulfate
standard.
Place a STAPH-IDENT test strip into the bottom of
the moistened tray. Take care not to contaminate
the microcupules with fingers when handling test
strip.
I'OOOOOOOOO
j'."AW ■![>£» r 1
J
ij
After incubation, record results of first 9 micro-
cupules and add 1-2 drops of STAPH-IDENT reagent
to tenth microcupule as shown. A plum-purple color
is positive. Record result.
After labeling the end tab of a tray with your name
and unknown number, dispense approximately 5 ml
of tap water into bottom of tray.
V
w
lloioodooobo
i& © ^} & © {: £) *b' © ® ^
:':f j h
i-
With a Pasteur pipette dispense 2 to 3 drops of the
bacteria) suspension into each of the 10 microcu-
pules. Cover the tray with the lid and incubate at
35°-37° C for 5 hours.
F'HS UNf Glf, MNE MAN 'RE SAL GLC A-.G MGP
I 2 « ? 4 1 2 4
Once all results are recorded on Laboratory Report,
total up positive values in each group to determine
4-digit profile. Consult chart VII, appendix D, to find
unknown.
Figure 55.2 The API Staph-ldent procedure
199
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
IX. Miniaturized Multitest
Systems
55. Staphylococcus
Identification: The API
Staph-ldentification
System
© The McGraw-H
Companies, 2001
Exercise 55 • Staphylococcus Identification: The API Staph-Ident System
First Period
(Inoculations and Coagulase Test)
Before setting up this experiment, take into consider-
ation that it must be completed at the end of 5 hours
Holding the test strips overnight is not recommended
Materials:
API Staph-Ident test strip
API incubation tray and cover
blood agar plate culture of unknown (must not
have been incubated over 30 hours)
blood agar plate (if needed for purity check)
serological tube of 2 ml sterile saline
test-tube rack
sterile swabs (optional in step 2 below)
squeeze bottle of tap water
tubes containing McFarland No. 3 (BaS0 4 )
standard (see Appendix B)
sterile Pasteur pipette (5 ml size)
1
2
3
4
5
6
7
8
If the coagulase test has not been performed, re-
fer to Exercise 78, page 260, for the procedure
and perform it on your unknown.
Prepare a saline suspension of your unknown by
transferring organisms to a tube of sterile saline
from one or more colonies with a loop or sterile
swab. Turbidity of the suspension should match a
tube of No. 3 McFarland barium sulfate standard.
Important: Do not allow the bacterial suspen-
sion to go unused for any great length of time.
Suspensions older than 15 minutes become less
effective.
Label the end strip of the tray with your name and
unknown number. See illustration 2, figure 55.2.
Dispense about 5 ml of tap water into the bottom
of the tray with a squeeze bottle. Note that the bot-
tom of the tray has numerous depressions to ac-
cept the water.
Remove the API test strip from its sealed enve-
lope and place the strip in the bottom of the tray.
After shaking the saline suspension to disperse
the organisms, fill a sterile Pasteur pipette with
the bacterial suspension.
Inoculate each of the microcupules with two or
three drops of the suspension. If a purity check is
necessary, use the excess suspension to inoculate
another blood agar plate.
Place the plastic lid on the tray and incubate the
strip aerobically for 5 hours at 35° to 37° C.
Figure 55.3 Test results of a strip inoculated with S.
aureus.
(Courtesy of Analytab Products)
Second Period
(Five Hours Later)
During this period the results will be recorded on the
Laboratory Report, the profile number will be deter-
mined, and the unknown will be identified by looking
up the number on the Staph-Ident Profile Register (or
Chart VII, Appendix D).
Materials:
API Staph-Ident test strip (incubated 5 hours)
1 bottle of Staph-Ident reagent (room
temperature)
Staph-Ident Profile Register
1. After 5 hours incubation, refer to Chart V,
Appendix D, to interpret and record the results of
the first nine microcupules (PHS through ARG) .
2. Record the results on the Profile Determination
Table on the Laboratory Report. Chart VI,
Appendix D, reveals the biochemistry involved in
these tests.
3. Add one or two drops of Staph-Ident reagent to
the NGP microcupule. Allow 30 seconds for the
color change to occur.
A positive test results in a change of color to
plum-purple. Record the results of this test.
4. Construct the profile number according to the in-
structions on the Laboratory Report and deter-
mine the name of your unknown.
If no recent Profile Register is available, use
Chart VII, Appendix D. Since the API Register is
constantly being updated, the one in the appendix
may be out of date.
Disposal
Once all the information has been recorded be sure to
place the entire incubation unit in a receptacle that is
to be autoclaved.
200
Benson: Microbiological
X. Microbiology of Soil
Introduction
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Companies, 2001
Part
Microbiology of Soil
With ideal temperature and moisture conditions, soils provide ex-
cellent culture media for many kinds of microorganisms. This is es-
pecially true of cultivated and improved soils. In many different
ways these organisms contribute to the fertility of the very medium
they inhabit. The action of certain autotrophic protists on minerals
produces substances, organic and inorganic, that are available to
plants. Maintaining a proper balance of available nitrogen to pho-
tosynthetic plants is one of the most important activities of some
forms of bacteria. Several experiments in this unit pertain to the cy-
cling of nitrogen between the soil, water, and atmosphere. The de-
composition of lifeless plant and animal tissues returns materials to
soils in a form that is reusable by plants.
n addition to studying phases of the nitrogen cycle, an attempt
will be made in Exercise 57 to isolate antibiotic producers from soi
samples. This study will concentrate primarily on Actinomyces-Wke
colonies.
201
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
X. Microbiology of Soil
56. Microbial Population
Counts of Soil
© The McGraw-H
Companies, 2001
56
Microbial Population Counts of Soil
Soils contain enormous numbers and kinds of mi-
croorganisms. In addition to the multitudes of bacte-
ria, there are protozoans, yeasts, molds, algae, and
microscopic worms in unbelievable numbers. Types
that predominate will depend on the composition of
the soil, moisture, pH, and other related environmen-
tal factors. No one technique can be used for count-
ing all organisms since such great variability in types
exists.
In this exercise we will use the plate count pro-
cedure that was used in Exercise 23 to determine the
numbers of bacteria, actinomycetes, and molds. It
will be necessary to use different kinds of media for
each group of organisms. For economy of time and
materials, the class will be divided into three
groups.
Materials:
1 bottle (50 ml) of nutrient agar ( l A of class)
1 bottle (50 ml) of glucose peptone acid agar ( l A
of class)
1 bottle (50 ml) of glycerol yeast extract agar ( l A
of class)
3 sterile water blanks (99 ml) (per pair of
students)
4 sterile Petri plates per student
1 . 1 ml dilution pipettes
soil sample
1
2
3
4
5
6
7
8
Liquefy and cool to 50° C a bottle of medium to
be used for the organisms that you will attempt to
count. The chart below indicates your assignment:
Label four Petri plates according to type of or-
ganisms and dilutions. Since the numbers of each
type will vary, different dilutions are necessary.
B acteria
Actinomycetes
Molds
1:10,000
1:1000
1:100
1:100,000
1:10,000
1:1000
1 : 1 ,000,000
1:100,000
1:10,000
1:10,000,000
1:1,000,000
1:100,000
Label three 99 ml sterile water blanks as you did
in Exercise 23.
Add 1 gram of soil to blank A, shake vigorously
for 5 minutes, and carry out the dilution of blanks
B and C.
With 1.1 ml pipettes, distribute the proper
amounts of water from the blanks to the plates for
your final dilutions. See figure 23.1. For the
1 : 1000 and 1 : 100 plates, you will need 0. 1 ml and
1 .0 ml, respectively, from blank A.
Pour the appropriate medium into each plate and
allow them to cool.
Incubate the plates in your locker for 3 to 7 days.
Count the colonies, using the procedures outlined
in Exercise 23. Record the results on the first por-
tion of Laboratory Report 56, 57.
Student Number
Organisms
Medium
1,4,7, 10, 13, 16, 19,22,25,28
Bacteria
Nutrient agar
2,5,8, 11, 14, 17,20,23,26,29
Actinomycetes
Glycerol yeast extract agar
3,6,9, 12, 15, 18,21,24,27,30
Molds
Glucose peptone acid agar
202
Benson: Microbiological
X. Microbiology of Soil
57. Isolation of an
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Antibiotic Producer from
Soil
Companies, 2001
■
Isolation of an Antibiotic Producer from Soil
The constant search of soils throughout the world has
yielded an abundance of antibiotics of great value for
the treatment of many infectious diseases. Pharma-
ceutical companies are in constant search for new
strains of bacteria, molds, and Actinomyces that can
be used for antibiotic production. Although many or-
ganisms in soil produce antibiotics, only a small por-
tion of new antibiotics are suitable for medical use. In
this experiment an attempt will be made to isolate an
antibiotic-producing Actinomyces from soil. Students
will work in pairs.
First Period
(Primary Isolation)
Unless the organisms in a soil sample are thinned out
sufficiently, the isolation of potential antibiotic pro-
ducers is nearly impossible. As indicated in figure
57.1, it will be necessary to use a series of six dilution
tubes to produce a final soil dilution of 10~ 6 . Proceed
as follows:
Materials:
per pair of students:
6 large test tubes
1 bottle of physiological saline solution
3 Petri plates of glycerol yeast extract agar
L- shaped glass rod
beaker of alcohol
6 1 ml pipettes
1 1 ml pipette
1 . Label six test tubes 1 through 6, and with a 10 ml
pipette, dispense 9 ml of saline into each tube.
2. Weigh out 1 g of soil and deposit it into tube 1 .
3. Vortex mix tube 1 until all soil is well dispersed
throughout the tube.
A tenfold serial dilution of the soil is made by transferring 1 .0
ml of solution from each tube to the next one to achieve a final
dilution of 1:1,000,000 in tube 6.
1 ml 1 ml 1m
1 m
1 m
One gram of soil is added
to tube 1 , containing 9 ml
of saline solution.
Soil in tube 1 is thoroughly
vortex-mixed.
1
\J>
v_^
^J
6
Each tube contains 9 ml of saline solution.
1 .0 ml is transferred from
tubes 4, 5, and 6 to Petri
plates of glycerol yeast ex-
tract agar.
An alcohol-flamed glass rod is used
to spread the 1 .0 ml of soil suspen-
sion on the surface of each of the
agar plates.
The three primary isolation plates of glycerol yeast
extract agar plates are incubated at 30° C for 7 days
Figure 57.1 Primary isolation of antibiotic-producing Actinomyces
203
Benson: Microbiological
X. Microbiology of Soil
57. Isolation of an
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Antibiotic Producer from
Soil
Companies, 2001
Exercise 57 • Isolation of an Antibiotic Producer from Soil
4
5
6
7
Make a tenfold dilution from tube 1 through tube
6 by transferring 1 ml from tube to tube. Use a
fresh pipette for each transfer and be sure to
pipette-mix thoroughly before each transfer.
Label three Petri plates with your initials and the
dilutions to be deposited into them.
From each of the last three tubes transfer 1 ml to
a plate of glycerol yeast extract agar.
Spread the organisms over the agar surfaces on
each plate with an L- shaped glass rod that has
been sterilized each time in alcohol and open
flame. Be sure to cool rod before using.
CAUTION
Keep Bunsen burner flame away from beaker of al-
cohol. Alcohol fumes are ignitable. Be sure to flame
the glass rod when finished.
8. Incubate the plates at 30° C for 7 days.
Second Period
(Colony Selection and Inoculation)
The objective in this laboratory period will be to se-
lect Actinomyces -like colonies that may be antibiotic
producers. The organisms will be streaked on nutri-
ent agar plates that have been seeded with
Staphylococcus epidermidis. After incubation we
will look for evidence of antibiosis. Students will
continue to work in pairs. Figure 57.2 illustrates the
procedure.
Materials:
per pair of students:
4 trypticase soy agar pours (liquefied)
4 sterile Petri plates
TSB culture of Staphylococcus epidermidis
1 ml pipette
3 primary isolate plates from previous period
water bath at student station (50° C)
1. Place four liquefied agar pours in water bath (50°
C) to prevent solidification, and then inoculate
each one with 1 ml of S. epidermidis.
2. Label the Petri plates with your initials and
date.
3. Pour the contents of each inoculated tube into
Petri plates. Allow agar to cool and solidify.
4. Examine the three primary isolation plates for the
presence of Actinomyces -like colonies. They have
Spores from primary isolate are
streaked on TSA plate that was
seeded with S. epidermidis
I
SECOND
PERIOD
PRIMARY ISOLATION PLATE
30° C 48 Hours
TO FOURTH
PERIOD
30° C 2-7 days
THIRD
PERIOD
Antibiotic producer is cross-streaked with
Staphylococcus epidermidis on TSA plate
]!
Figure 57.2 Second and third period inoculations
Benson: Microbiological
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57. Isolation of an
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Antibiotic Producer from
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Isolation of an Antibiotic Producer from Soil • Exercise 57
a dusty appearance due to the presence of spores.
They may be white or colored. Your instructor
will assist in the selection of colonies.
5. Using a sterile inoculating needle, scrape spores
from Actinomyces-likQ colonies on the primary
isolation plates to inoculate the seeded TSA
plates. Use inoculum from a different colony for
each of the four plates.
6. Incubate the plates at 30° C until the next labora-
tory period.
Materials:
1 Petri plate of trypticase soy agar
TSB culture of S. epidermidis
If antibiosis is present, make two streaks on the TSA
plate as shown in figure 57.2. Make a straight line
streak first with spores from the Actinomyces colony,
using a sterile inoculating needle. Cross-streak with
organisms from a culture of S. epidermidis. Incubate
at 30° C until the next period.
Third and Fourth Periods
(Evidence of Antibiosis and Confirmation)
Examine the four plates you streaked during the last
laboratory period. If you see evidence of antibiosis
(inhibition of S. epidermidis growth), proceed as fol-
lows to confirm results.
Laboratory Report
After examining the cross- streaked plate during the
fourth period, record your results on the Laboratory
Report and answer all the questions.
205
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58. The Nitrogen Cycle
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58
The Nitrogen Cycle
The next three exercises have one thing in common:
they all pertain to the nitrogen cycle. This exercise is
presented here to unify these experiments as you
study the different phases of the nitrogen cycle. There
is no laboratory report for this exercise.
As pointed out in Exercise 20, nitrogen is one of
the essential elements needed by all living organ-
isms. Although nearly 80% of the atmosphere con-
sists of molecular nitrogen, very few life-forms are
able to utilize it in its free state. Instead, most or-
ganisms can utilize it only if it is combined
("fixed") with another element such as oxygen or
hydrogen. Nitrates (NO J), nitrites (NO J), ammo-
nium (NH4 ), or organic nitrogenous compounds
(proteins and nucleic acids) are the principal forms
of fixed nitrogen.
Most plants are able to utilize nitrates and am-
monia. Animals, on the other hand, derive their ni-
trogen from plants and other animals in the form of
organic compounds. Microorganisms, however, vary
considerably in their nitrogen uptake in that they
may get it from all of the sources listed above, plus
free nitrogen.
Figure 58.1 illustrates the four phases of the ni-
trogen cycle: ammonification, nitrification, nitrogen
fixation, and denitrification. A discussion of each
phase follows.
Ammonification
Most of the nitrogen in soil exists in the form of or-
ganic molecules, mostly proteins and nucleic acids
that are derived from the decomposition of dead plant
and animal tissue. When an organism dies, its proteins
are attacked by proteases of soil bacteria to produce
polypeptides and amino acids. The amino groups on
the amino acids are then removed by a process called
deamination and converted into ammonia (NH 3 ).
This production of ammonia is called ammonifica-
tion. In addition to the ammonification of protein and
nucleic acids of dead animals and plants, other wastes
Amino Acids
Many Bacteria
Ammonia
such as urea and uric acid from animal wastes go
through the ammonification process. Bacteria and
plants that are able to assimilate ammonia convert it
into amino acids needed for their own enzyme and
protoplasm construction.
Nitrification
The next sequence of reactions in the nitrogen cycle in-
volves the oxidation of the nitrogen in the ammonium
ion to produce nitrite. This step is followed by the oxi-
dation of nitrites to produce nitrates. This two-step
process is called nitrification. Note in the reaction be-
low that the first stage is controlled by autotrophs of the
genera Nitrosomonas and Nitrosococcus. The second
Nitrosomonas
+ Nitrosococcus
NH 4
NO,
Nitrobacter
Nitrococcus
Ammonium ion
a
Nitrite ion
a
— N0 3
Nitrate ion
stage is performed by members of the genera
Nitrobacter and Nitrococcus. Although there are other
nitrifying bacteria in soil that can perform these con-
versions, they are insignificant contributors to this
process.
Nitrogen Fixation
The conversion of atmospheric nitrogen to ammonia
is called nitrogen fixation. This process, which is il-
lustrated on the right side of figure 58.1, is performed
by three groups of microorganisms: (1) free-living
bacteria, (2) cyanobacteria, and (3) symbiotic bacteria
in root nodules of leguminous plants.
Of the various free-living bacteria, the most ben-
eficial ones belong to genus Azotobacter. Since these
organisms are strict aerobes, they function effi-
ciently in well- aerated garden soils. Due to the fact
that soils usually lack abundant sources of carbohy-
drates, most of the other free-living bacteria, such as
Clostridium and Klebsiella, fail to contribute as
much fixed nitrogen.
206
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X. Microbiology of Soil
58. The Nitrogen Cycle
©The McGraw-Hill
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Cyanobacteria, such as Nostoc and Anabaena,
which do not require a carbohydrate source for en-
ergy, are excellent fixers of atmospheric nitrogen.
These chlorophyll-packed organisms usually carry
their nitrogen-fixing enzymes in specialized struc-
tures called heterocysts. The fact that these organisms
are so productive in nitrogen fixation explains why
they often contribute to organic pollution of freshwa-
ter ponds and lakes.
The symbiotic nitrogen-fixing bacteria are the
most important contributors to soil enrichment. They
develop in root nodules of leguminous plants, such as
peas, beans, peanuts, clover, and alfalfa. The principal
genera are Rhizobium and Brady rhizobium. They are
symbiotic in that they produce nourishment for the
host plant and the host provides anaerobic conditions
and nutrients for the bacteria. Farmers utilize this bac-
terial relationship through crop rotation to pump liter-
ally millions of tons of fixed nitrogen into their soils
annually.
The Nitrogen Cycle • Exercise 58
Denitrification
Under anaerobic conditions, some microbes can uti-
lize nitrates as electron acceptors to metabolize other
organic substances. This conversion of nitrates to free
nitrogen is called denitrification. The denitrification
process takes place as follows:
N0 3
Nitrate
on
N0 2
Nitrite
on
NO
Nitric
Oxide
- N 2
Nitrous
Oxide
N
Dinitrogen
Pseudomonas aeruginosa and Paracoccus deni-
trificans are examples of two species that can bring
about the denitrification of nitrates and nitrites. Since
the denitrification process occurs in waterlogged soils
where there is a deficiency of oxygen, farmers mini-
mize nutrient loss from soil by constant cultivation to
promote aeration.
Figure 58.1 The nitrogen cycle
207
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X. Microbiology of Soil
59. Nitrogen-Fixing
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59
Nitrogen- Fixing Bacteria
Among the most beneficial microorganisms of the
soil are those that are able to convert gaseous nitro-
gen of the air to "fixed forms" of nitrogen that can be
utilized by other bacteria and plants. Without these
nitrogen-fixers, life on this planet would probably
disappear within a relatively short period of time.
The utilization of free nitrogen gas by fixation
can be accomplished by organisms that are able to
produce the essential enzyme nitrogenase. This en-
zyme, in the presence of traces of molybdenum, en-
ables the organisms to combine atmospheric nitro-
gen with other elements to form organic compounds
in living cells. In organic combinations nitrogen is
more reduced than when it is free. From these or-
ganic compounds, upon their decomposition, the ni-
trogen is liberated in a fixed form, available to
plants either directly or through further microbial
action.
The most important nitrogen-fixers belong to two
families: Azotobacteraceae and Rhizobiaceae.
Other organisms of less importance that have this
ability are a few strains of Klebsiella, some species of
Clostridium, the cyanobacteria, and photosynthetic
bacteria.
In this exercise we will concern ourselves with
two activities: the isolation of Azotobacter from gar-
den soil and the demonstration of Rhizobium in root
nodules of legumes.
Azotobacteraceae
Bergey's Manual of Systematic Bacteriology, vol-
ume 1 , section 4, lists two genera of bacteria in fam-
ily Azotobacteraceae that fix nitrogen as free-living
organisms under aerobic conditions: Azotobacter
and Azomonas. The basic difference between these
two genera is that Azotobacter produces drought-
resistant cysts and Azomonas does not. Aside from
the presence or absence of cysts, these two genera
are very similar. Both are large gram-negative
motile rods that may be ovoid or coccoidal in shape
(pleomorphic). Catalase is produced by both gen-
era. There are six species of Azotobacter and three
species of Azomonas.
Figure 59.1 illustrates the overall procedure that
we will use for isolating Azotobacteraceae from gar-
den soil. Note that a small amount of rich garden soil
is added to a bottle of nitrogen-free medium that con-
tains glucose as a carbon source. The bottle of
medium is incubated in a horizontal position for 4 to
7 days at 30° C.
After incubation, a wet mount slide is made
from surface growth to see if typical azotobacter-
like organisms are present. If organisms are present,
an agar plate of the same medium, less iron, is used
to streak out for isolated colonies. After another 4 to
7 days incubation, colonies on the plate are studied
and more slides are made in an attempt to identify
the isolates.
The N 2 -free medium used here contains glucose
for a carbon source and is completely lacking in ni-
trogen. It is selective in that only organisms that can
use nitrogen from the air and use the carbon in glu-
cose will grow on it. All species of Azotobacter and
Azomonas are able to grow on it. The metallic ion
molybdenum is included to activate the enzyme nitro-
genase, which is involved in this process.
First Period (Enrichment)
Proceed as follows to inoculate a bottle of the nitrogen
free glucose medium with a sample of garden soil.
Materials:
1 bottle (50 ml) N 2 -free glucose medium
(Thompson-Skerman)
rich garden soil (neutral or alkaline)
spatula
1
2
With a small spatula, put about 1 gm of soil into
the bottle of medium. Cap the bottle and shake it
sufficiently to mix the soil and medium.
Loosen the cap slightly and incubate the bottle at
30° C for 4 to 7 days. Since the organisms are strict
aerobes, it is best to incubate the bottle horizontally
to provide maximum surface exposure to air.
Second Period (Plating Out)
During this period a slide will be made to make cer-
tain that organisms have grown on the medium. If
the culture has been successful, a streak plate will be
208
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X. Microbiology of Soil
59. Nitrogen-Fixing
©The McGraw-Hill
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Bacteria
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made on nitrogen-free, iron-free agar. Proceed as
follows:
1
Materials:
microscope slides and cover glasses
microscope with phase-contrast optics
1 agar plate of nitrogen-free, iron-free glucose
medium
2
Nitrogen- Fixing Bacteria • Exercise 59
After 4 to 7 days incubation, carefully move the
bottle of medium to your desktop without agitat-
ing the culture.
Make a wet mount slide with a few loopfuls from
the surface of the medium and examine under oil
immersion, preferably with phase-contrast optics.
Look for large ovoid to rod-shaped organisms,
singly and in pairs.
One gram of rich garden soil is
added to 50 ml of selected enrich
ment medium.
Thompson
Skerman
Medium
noculated medium is incubated
at about 30° C for 4-7 days in
horizontal position.
After incubation and before
making streak plate, a wet
mount slide is made to deter-
mine if organisms are present,
If organisms are present,
an agar plate of iron-free
medium is streaked out.
Isolated colonies are used for
making gram-stained slides, doing
motility studies, and looking for
fluorescent water-soluble pigmen-
tation of the medium. Further sub-
culturing may also be done for
other tests.
Figure 59.1 Enrichment and isolation procedure for Azotobacter and Azomonas
209
Benson: Microbiological
X. Microbiology of Soil
59. Nitrogen-Fixing
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Bacteria
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Exercise 59 • Nitrogen-Fixing Bacteria
3. If azotobacter-like organisms are seen, note
whether or not they are motile and if cysts are
present. Cysts look much like endospores in that
they are refractile. Since cysts often take 2 weeks
to form, they may not be seen.
4. If the presence of azotobacter-like organisms is
confirmed, streak an agar plate of nitrogen-free,
iron-free medium, using a good isolation streak
pattern. Ferrous sulfate has been left out of this
medium to facilitate the detection of water-soluble
pigments .
5. Incubate the plate at 30° C for 4 or 5 days. A
longer period of incubation is desirable for cyst
formation.
Third Period (Identification)
Azotobacter chroococcum is the type species of genus
Azotobacter. The cells are 1.5-2.0 micrometers in di-
ameter and pleomorphic, ranging from rods to coc-
coidal in shape. They occur singly, in pairs, and in ir-
regular clumps. Motility exists with peritrichous
flagella. Drought-resistant cysts are produced. They
are strict aerobes. Catalase is produced and starch is
hydrolyzed. Morphologically, the other five species
of this genus look very much like this organism.
Azomonas agilis is the type species of genus
Azomonas. Except for the absence of cysts, this
species and the other two species in this genus are
morphologically very similar to Azotobacter
chroococcum. Practically all of them produce water-
soluble fluorescent pigments.
Differentiation of the six species of the genus
Azotobacter and three species of Azomonas is based
primarily on the presence or absence of motility, the
type of water-soluble pigment produced, and carbon
source utilization. Table 59.1 reveals how the organ-
isms can be differentiated. For presumptive identifi-
cation, use the following character information to
identify your isolate.
Materials:
agar plate from previous period
ultraviolet lamp
Motility Note in table 59.1 that four species of
Azotobacter and all three species of Azomonas are
motile.
Pigmentation Although these organisms produce
both water-soluble and water-insoluble pigments,
only the water-soluble ones (those capable of diffus-
Table 59.1 Differential characteristics of the Azotobacteraceae
Water-Soluble Pigments
/ / / / / \fr / / / JS /
/ / / / / jt / / / ^ /
A A A A /<?/ A / //^
A A Af/f/f/*/ A«* A/ A
/ e> / a$P / jH / & / & / NT / <^ / <& / y? /
Azotobacter
A. chroococcum
+
+
—
—
—
—
—
—
—
A. vinelandii
+
+
—
—
—
d
d
+
—
A. beijerinckii
+
—
—
—
—
—
—
—
—
A. nigricans
+
—
—
d
+
d
—
—
—
A. armeniacus
+
+
—
—
+
+
—
—
—
A. paspali
+
+
+
—
—
+
—
+
—
Azomonas
A. agilis
—
+
—
—
—
—
—
+
+
A. insignis
—
+
—
d 1
—
d
—
d
—
A. macrocytogenes
—
+
—
—
—
—
—
d
d
d = 11%-89% positive
d 1 = 11%-89% positive on benzoate
From Bergey's Manual of Systematic Bacteriology, volume 1 , section 4
210
Benson: Microbiological
X. Microbiology of Soil
59. Nitrogen-Fixing
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ing into an agar medium) are important from the
standpoint of species differentiation.
Note in table 59.1 that two of the water-soluble pig-
ments are fluorescent: one is yellow-green and the
other is blue- white. To observe fluorescence the cul-
tures must be exposed to ultraviolet light (wavelength
364 nm) in a darkened room. The characteristics of
pigment production in each species may be limited by
certain factors, as indicated below:
Brown-black: If the colonies produce this hue
of diffusible pigment without becoming red-
violet, the organism is A. nigricans. Although
the table indicates that A. insignis can produce
the brown-black pigment, it can do so only if
the medium contains benzoate.
Brown-black to red-violet: As indicated in the
table, A. nigricans and A. armeniacus are the
only genera that produce this type of pigment.
Motility is a good way to differentiate these
two species.
Red-violet: Although table 59.1 reveals that five
species can produce this color of diffusible
pigment, one (A. insignis) cannot produce it
on the medium we used. A red- violet isolate
is unlikely to be A. paspali because this or-
ganism has been isolated from the rhizo-
sphere of only one species of grass (Paspalum
notatum). Thus, isolates that produce this pig-
ment are probably one of the other three in the
table.
Green: Note that only A. vinelandii can produce
this water-soluble pigment; however, only
1 1 %-89% of them produce it.
Yellow-green fluorescent: A. vinelandii, A. pas-
pali, and all species of Azomonas are able to
produce this pigment on the medium we used.
Check for fluorescence with an ultraviolet
lamp in a darkened room.
Blue-white fluorescent: Note in table 59.1 that
two species of Azomonas can produce this
type of diffusible pigment; no Azotobacter
are able to produce it. Check for fluorescence
with an ultraviolet lamp in a darkened room.
Carbon Source The medium we used in this exper-
iment contains 1% glucose, which can be utilized by
all Azotobacter and Azomonas. Selectivity can be
achieved by replacing the glucose with rhamnose,
caproate, caprylate, myoinositol, mannitol, mal-
onate, or several other carbon sources. If more precise
differentiation is desirable, the student is referred to
Tables 4.48 and 4.49 on pages 231 and 232 in
Bergey's Manual, volume 1.
Nitrogen- Fixing Bacteria • Exercise 59
Laboratory Report
Record your observations and conclusions for the
Azotobacter aceae on the Laboratory Report.
Rhizobiaceae
Although the free-living Azotobacteraceae are benefi-
cial nitrogen- fixers, their contribution to nitrogen en-
richment of the soil is limited due to the fact that they
would rather utilize NH 3 in soil than fix nitrogen. In
other words, if ammonia is present in the soil, nitro-
gen fixation by these organisms is suppressed. By
contrast, the symbiotic nitrogen-fixers of genus
Rhizobium, family Rhizobiaceae, are the principal ni-
trogen enrichers of soil.
Bergey's Manual lists three genera in family
Rhizobiaceae: Rhizobium, Brady rhizobium, and
Agrobacterium. Although the three genera are related,
only genus Rhizobium fixes nitrogen. This genus of
symbiotic nitrogen- fixers contains only three species.
Differentiation of these species relies primarily on
plant inoculation tests. A partial list of the host plants
for each species is as follows:
R. leguminosarum: peas, vetch, lentils, beans,
scarlet runner, and clover
R. meliloti: sweet clover, alfalfa, and fenugreek
R. loti: trefoil, lupines, kidney vetch, chickpea,
mimosa, and a few others
All three of these species are gram-negative pleomor-
phic rods (bacteroids), often X-, Y-, star-, and club-
shaped; some exhibit branching. Refractile granules
are usually observed with phase-contrast optics. All
are aerobic and motile. Our study of Rhizobium will
be of crushed root nodules from whatever legume is
available.
Materials:
washed nodules from the root of a legume
methylene blue stain
microscope slides
1
2
3
Place a nodule on a clean microscope slide and
crush it by pressing another slide over it. Produce
a thin smear by sliding the top slide over the lower
one.
After air-drying and fixing with heat, stain the
smear with methylene blue for 30 seconds.
Examine under oil immersion and draw some of
the organisms on the Laboratory Report. Look for
typical bacteroids of various configurations.
Laboratory Report
Complete the Laboratory Report for this exercise
211
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X. Microbiology of Soil
60. Ammonification in Soil
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Ammonification in Soil
As indicated in our discussion of the nitrogen cycle in
Exercise 58, the nitrogen in most plants and animals
exists in the form of protein. When these organisms
die, the protein is broken down to amino acids, which,
in turn, are deaminated to liberate ammonia. This
process of the production of ammonia from organic
compounds is called ammonification. Since most
bacteria and plants can assimilate ammonia, this is a
very important step in the nitrogen cycle. The majority
of bacteria in soil are able to take part in this process.
To demonstrate the existence of this process we
will inoculate peptone broth with a sample of soil, in-
cubate it for a few days, and test for ammonia produc-
tion. After a total of 7 days' incubation it will be tested
again to see if the amount of ammonia has increased.
First Period
(Inoculation)
Materials:
2 tubes of peptone broth
rich garden soil
1 . Inoculate one tube of peptone broth with a loop-
ful of soil. Save the other tube for a control.
2. Incubate the tube at room temperature for 3-4
days and 7 days.
Second and Third Periods
(Ammonia Detection)
After 3 or 4 days, test the medium for ammonia with
the following procedure. Repeat these tests again af-
ter a total of 7 days of incubation.
Materials:
Nessler's reagent
bromthymol blue and indicator chart spot plate
1
2
3
4
Deposit a drop of Nessler's reagent into two sep-
arate depressions of a spot plate.
Add a loopful of the inoculated peptone broth to
one depression and a loopful from the sterile
uninoculated tube in the other. Interpretation of
ammonia presence is as follows:
Faint yellow color — small amount of ammonia
Deep yellow — more ammonia
Brown precipitate — large amount of ammonia
Check the pH of the two tubes by placing several
loopfuls of each in separate depressions on the
spot plate and adding 1 drop of bromthymol blue
to each one. Compare the color with a color chart
or set of indicator tubes to determine the pH.
Record results on the Laboratory Report.
212
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61. Isolation of a Denitrifier
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Isolation of a Denitrifier from Soil
Using Nitrate Agar
61
Denitrification is defined as the reduction of nitrate
(N0 3 ) to gaseous dinitrogen (N 2 ). The consequence of
this process is the loss of fixed nitrogen from the soil
and water. As far as we know today, the only organisms
that are able to denitrify fixed nitrogen are the prokary-
otes. Although the percentage of prokaryotes that can
perform this phenomenon is not very high, that which
they are able to accomplish is truly extensive. The four
steps in the denitrification process are as follows:
1
N0 3 ---
Nitrate
Ion
N0 2
Nitrite
Ion
NO ---■► NoO
N
Nitric
Oxide
Nitrous
Oxide
Dinitrogen
Gas
That denitrification is extremely important in
ecological and geochemical terms is undeniable. A
summary of the effects of denitrification on ecology is
as follows:
• Without the existence of denitrification, the nitro-
gen in our atmosphere would become completely
depleted within a very short period of time.
Prokaryotic denitrification is essentially the only
source of nitrogen in our atmosphere.
• Denitrification is responsible for the extensive
depletion of fixed nitrogen in fertilizers that are
put into the soil by farmers. (It has been estimated
that somewhere between 5% and 80% of fixed ni-
trogen is removed from soils by this process.)
• Denitrification plays a major role in the return of
N 2 to the atmosphere from fixed nitrogen that ex-
ists in runoff water of rivers into the ocean.
• Denitrification is the most practical means of re-
ducing fixed nitrogen from sewage in sewage-
treatment plants.
• Nitrous oxide generated in the lower atmosphere
diffuses upward to the stratosphere where it is con-
verted to nitric oxide by a photochemical reaction.
Result: nitric oxide reacts with ozone to bring
about ozone depletion, which threatens our princi-
pal barrier against ultraviolet damage to all living
organisms. (It should be noted here that the signif-
icance of the cumulative effects of industrial, au-
tomotive, and other factors on nitrous oxide de-
pletion of the ozone layer is highly controversial.)
The essential function of denitrification to or-
ganisms is the generation of ATP. Although some or-
ganisms can carry the reaction completely from ni-
trate to dinitrogen, there are many organisms that are
able to act only at stages 1 , 2, or 3. If an organism can
work from one stage to another without final pro-
duction of dinitrogen, it is not considered to be a
denitrifier.
As far as we know at this time, organisms that can
convert nitrate to ammonia do not generate ATP from
the production of ammonia; rather, ATP is produced
only when the nitrate is first converted to nitrite. This
process has been referred to as nitrate respiration.
This reaction, as seen in the metabolism of E. coli, ap-
pears to be a means of detoxification of nitrite by con-
version to ammonia.
Habitats of Denitrifiers
Although most denitrifiers grow only in an anaerobic
environment, they are not all restricted to such places.
Those that grow elsewhere have alternative mecha-
nisms such as aerobic respiration, photosynthesis, or
fermentation to satisfy their ATP needs.
The most favorable environments for these or-
ganisms are heavily fertilized agricultural soils and
sewage where nitrogenous compounds abound in
considerable quantity. However, denitrifying
prokaryotes have been isolated from soils in the
Arctic and Antarctic, as well as from sediments in
freshwater, brackish water, and salt water. A ther-
mophilic denitrifier has even been isolated from a hot
spring. It is obvious, thus, that these organisms are
ubiquitous.
Organisms
Lists that have been compiled of denitrifiers reveal
that almost all groups of bacteria contain denitrifiers.
Typical groups are the phototrophic bacteria, gliding
bacteria, spiral and curved bacteria, gram-negative
aerobic bacteria, gram-negative cocci, chemolithic
sulfur bacteria, gram-positive spore formers, and
gram-positive non-spore formers.
Of all the groups listed, the gram-negative aerobic
group appears to have the largest number of denitri-
fiers, with genus Pseudomonas predominating.
213
Benson: Microbiological
X. Microbiology of Soil
61. Isolation of a Denitrifier
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
from Soil: Using Nitrate
Agar
Companies, 2001
Exercise 61 • Isolation of a Denitrifier from Soil: Using Nitrate Agar
Procedure
To isolate denitrifiers from a soil sample, the follow-
ing conditions must be met in the growth medium:
• Some nitrate must be available, which will pro-
vide the only terminal electron acceptor for the
generation of ATP.
• A carbon source must be present that cannot be
fermented by denitrifiers that have a fermentative
metabolism. Being unable to ferment the carbon
they are forced to use nitrate or nitrogenous oxide
for ATP generation.
• Some peptone must be present to provide essen-
tial amino acids needed by some denitrifiers.
Once we get an organism that grows on a medium
with these characteristics, the next step is to demon-
strate the ability of the organism to generate visible
nitrogen gas. An isolate that grows on nitrate media
and generates gas can be presumed to be a denitrifier.
It is these principles that govern the procedure that we
will follow here. Figure 61.1 illustrates the procedure
that involves a minimum of three laboratory periods.
First Period
Note that the water used in the blender contains 0.1%
Tween 80. Tween 80 is a surface active agent that low-
ers the surface tension around bacteria to improve dis-
persion of the organisms. The nitrate agar used in the
Petri plate is essentially nutrient agar to which 0.5%
KN0 3 is added.
Materials:
blenders
fresh soil sample
90 ml distilled water with 0.1% Tween 80
graduate
1 ml pipette
1 Petri plate of nitrate agar
GasPak anaerobic jar, generator envelopes, and
generator strips
1. Add 10 grams of soil to 90 ml of water that con-
tains Tween 80.
2. Blend for 2 minutes.
3. Label the bottom of a nitrate agar plate with your
name and date of inoculation.
4. Pipette 1 .0 ml of the blended mix onto the surface
of a plate of nitrate agar.
5. Spread the inoculum over the surface of the agar
with a bent glass rod.
6. Incubate the plate, inverted, at 30° C for 3 to 5
days in a GasPak anaerobic jar.
Second Period
During this period, nitrate agar plates will be examined
to select colonies that have developed during the incu-
bation period. Since the presence of growth doesn't nec-
essarily mean that the organism is a denitrifier, it will be
necessary to see if any of the isolates are nitrogen gas
producers; thus, Durham tube nitrate broths must be in-
oculated and incubated anaerobically. Nitrate broth con-
sists of nutrient broth plus 0.5% KN0 3 .
Materials:
nitrate agar plates with colonies
3 Durham tubes of nitrate broth
GasPak anaerobic j ar, generator envelopes, and
generator strips
1
2
3
Examine the nitrate agar plate. Look for colonies
that might be Pseudomonas aeruginosa, which
produces a soluble pigment into the medium. P.
fluorescens is also a denitrifier.
Select three different colonies to inoculate sepa-
rate tubes.
Note: Keep a record of the appearance of the
colonies transferred to the tubes.
Incubate the tubes at 30° C for 3 to 5 days in a
GasPak anaerobic j ar.
Third Period
Although most denitrifiers grow only in an anaerobic
environment, they are not all restricted to such places.
Those that grow elsewhere have alternative mecha-
nisms such as aerobic respiration, photosynthesis, or
fermentation to satisfy their ATP needs.
Materials:
3 Durham tubes from last period
1 Petri plate of sterile nitrate agar
GasPak anaerobic j ar, generator envelopes, and
generator strips
1. Record on the Laboratory Report whether you
have positive or negative results on the three
Durham tubes. The presence of gas is presump-
tive evidence that a denitrifier has been isolated.
2. If you plan to carry this experiment on further to
make more specific identification of an isolate,
make a streak plate from the positive tube and
proceed as indicated in figure 61.1.
Fourth Period
This period of inoculations is in preparation of trying to
do a definitive identification of a denitrifier. Note in fig-
ure 61.1 that from an isolated colony a nutrient broth is
inoculated and a gram-stained slide is made. After incu-
bation, the broth culture can be used as a stock culture
for doing further tests to identify your isolate. The slide
will reveal the morphological nature of your organism.
Laboratory Report
Complete the Laboratory Report for this exercise.
214
Benson: Microbiological
X. Microbiology of Soil
61. Isolation of a Denitrifier
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
from Soil: Using Nitrate
Agar
Companies, 2001
I , |i i ,!■■ ■qiMli
Ten grams of soil are added to
90 ml ol water with. Tween 80.
Mix in blender lor 2 minutes.
Third Period: Tubes positive for
gas formation indicate the presence
of denitrifiers.
30° C - 2 to 3 days,
in CasPak jnr
One ml of mix is pipetted to plate of
nitrate agar and spread-plated with
bent glass rod.
h
30° C - 2 to 3 days.
in CasPak jar
30° C - 2 to 3 days.
in CasPak jar
Second Period: From selected
colonies three Durham tubes of
nitrate broth are inoculated.
Fourth Period: If species identifi-
cation is to be performed, a streak
plate is made on nitrate agar from
Durham tube.
Fifth Period: From streak plate a
nutrient broth is inoculated to be used
as a stock culture for further tests and a
gram stained slide is made for study.
***W^»*W*P^^^P^P^^"^^^*^^"¥^**#^
p**H
Figure 61.1 Procedure for isolating a denitrifier
215
Benson: Microbiological
X. Microbiology of Soil
62. Isolation of a Denitrifier
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
from Soil: Using an
Enrichment Medium
Companies, 2001
Isolation of a Denitrifier from Soil
Using an Enrichment Medium
62
If you have already performed the experiment in
Exercise 61, much of the introductory information
that follows has already been discussed. For students
who have not done Exercise 61, however, this infor-
mation is critical to understanding this experiment.
Denitrification is defined as the reduction of ni-
trate (N0 3 ) to gaseous dinitrogen (N 2 ). The conse-
quence of this process is the loss of fixed nitrogen
from soil and water. As far as we know today, the only
organisms that are able to denitrify fixed nitrogen are
the prokaryotes. Although the percentage of prokary-
otes that can perform this phenomenon is not very
high, that which they are able to accomplish is truly
extensive. The four steps in the denitrification process
are as follows:
1
N0 3
Nitrate
Ion
N0 2
Nitrite
Ion
NO
Nitric
Oxide
N 2 ■
Nitrous
Oxide
N
Dinitrogen
Gas
That denitrification is extremely important in
ecological and geochemical terms is undeniable. A
summary of the effects of denitrification on ecology is
as follows:
• Without the existence of denitrification, the nitro-
gen in our atmosphere would become completely
depleted within a very short period of time.
Prokaryotic denitrification is essentially the only
source of nitrogen in our atmosphere.
• Denitrification is responsible for the extensive
depletion of fixed nitrogen in fertilizers that are
put into the soil by farmers. (It has been estimated
that somewhere between 5% and 80% of fixed ni-
trogen is removed from soils by this process.)
• Denitrification plays a major role in the return of
N 2 to the atmosphere from fixed nitrogen that ex-
ists in runoff water of rivers in the ocean.
• Denitrification is the most practical means of re-
ducing fixed nitrogen from sewage in sewage-
treatment plants.
• Nitrous oxide generated in the lower atmosphere
diffuses upward to the stratosphere where it is con-
verted to nitric oxide by a photochemical reaction.
Result: nitric oxide reacts with ozone to bring
about ozone depletion, which threatens our princi-
pal barrier against ultraviolet damage to all living
organisms. (It should be noted here that the signif-
icance of the cumulative effects of industrial, au-
tomotive, and other factors on nitrous oxide de-
pletion of the ozone layer is highly controversial.)
The essential function of denitrification to organ-
isms is the generation of ATP. Although some organ-
isms can carry the reaction completely from nitrate to
dinitrogen, there are many organisms that are able to
act only at stages 1, 2, or 3. If an organism can work
from one stage to another without final production of
dinitrogen, it is not considered to be a denitrifier.
As far as we know at this time, organisms that can
convert nitrate to ammonia do not generate ATP from
the production of ammonia; rather, ATP is produced
only when the nitrate is first converted to nitrite. This
process has been referred to as nitrate respiration. This
reaction, as seen in the metabolism of E. coli, appears
to be a means of detoxification of nitrite by conversion
to ammonia.
Habitats of Denitrifiers
Although most denitrifiers grow only in an anaerobic
environment, they are not all restricted to such places.
Those that grow elsewhere have alternative mecha-
nisms such as aerobic respiration, photosynthesis, or
fermentation to satisfy their ATP needs.
The most favorable environments for these or-
ganisms are heavily fertilized agricultural soils and
sewage where nitrogenous compounds abound in con-
siderable quantity. However, denitrifying prokaryotes
have been isolated from soils in the Arctic and
Antarctic, as well as from sediments in freshwater,
brackish water, and salt water. A thermophilic denitri-
fier has even been isolated from a hot spring. It is ob-
vious, thus, that these organisms are ubiquitous.
Organisms
Lists that have been compiled of denitrifiers reveal
that almost all groups of bacteria contain denitrifiers.
Typical groups are the phototrophic bacteria, gliding
bacteria, spiral and curved bacteria, gram-negative
217
Benson: Microbiological
X. Microbiology of Soil
62. Isolation of a Denitrifier
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
from Soil: Using an
Enrichment Medium
Companies, 2001
Exercise 62 • Isolation of a Denitrifier from Soil: Using an Enrichment Medium
aerobic bacteria, gram-negative cocci, chemolithic
sulfur bacteria, gram-positive spore formers, and
gram-positive non-spore formers.
Of all the groups listed, the gram-negative aero-
bic group appears to have the largest number of deni-
trifiers, with genus Pseudomonas predominating.
Paracoccus denitrificans In our experiment here we
will focus on isolating Paracoccus denitrificans, a
member of this gram-negative aerobic group.
According to Bergey's Manual, the characteristics of
this denitrifier are as follows:
Cells may be spherical or short rods, gram-negative,
and nonmotile. They are aerobic, having a strictly respi-
ratory type of metabolism. Anaerobic growth does oc-
cur, however, if nitrate, nitrite, or nitrous oxide are avail-
able as terminal electron acceptors. Under anaerobic
conditions, nitrate is reduced to nitrous oxide and dini-
trogen gas. Colonies on nutrient agar are 2 to 3 mm in
diameter, usually circular, entire, smooth, glistening
white, and opaque.
To isolate P. denitrificans we will use an enrich-
ment culture technique, which employs a nitrate
succinate-mineral salts medium. This medium
contains sodium succinate, potassium nitrate, and
basic mineral salts. Being able to oxidize the succi-
nate and use nitrate as a terminal electron acceptor
to produce nitrogen gas, P. denitrificans will grow
very well.
Procedure
Figure 62. 1 illustrates the procedure for the first two
laboratory sessions. This phase of the experiment will
yield a mixed culture of P. denitrificans and other soil
bacteria. To get a pure culture of P. denitrificans we
will follow the procedure in figure 62.2. Proceed as
follows:
First Period
Materials:
fresh soil sample
flask containing 200 ml nitrate
succinate-mineral salts broth
sterile glass-stoppered bottle (60 ml size)
Petri dish top or bottom
1. Add 1 g of soil to the nitrate succinate-mineral
salts broth. Shake the flask vigorously and allow
the soil contents to settle.
2. Carefully decant some of the supernatant into a
glass-stoppered bottle, filling it to total capacity.
3. Insert the stopper into the bottle in such a way that
the medium in the neck of the bottle is expelled.
4. Place the bottle in the top or bottom of a Petri dish to
collect any liquid that is expelled during incubation.
5. Incubate the bottle at 30° C until the next labora-
tory session.
Second Period
Materials:
flask of sterile nitrate succinate-mineral salts
broth
culture in glass- stoppered bottle (from previous
lab period)
sterile glass- stoppered bottle (60 ml size)
1 ml pipette
microscope slides and cover glasses
gram- staining kit
1
2
3
4
5
Examine the bottled culture for the presence of
gas. A stream of nitrogen-gas bubbles should be
visible extending up from the bottom of the bottle
and collecting at the top of the culture. The glass
stopper is often displaced by the force of the gas.
Prepare a second enrichment by aseptically trans-
ferring 1 ml of the initial culture to a sterile sec-
ond glass- stoppered bottle. Fill the bottle com-
pletely and stopper it. Set this bottle aside in a
Petri dish cover to incubate at 30° C until the next
laboratory period.
Prepare a gram- stained slide from the initial cul-
ture and examine it under oil immersion.
Make a wet mount slide from the initial culture
and examine with phase-contrast optics.
Record all your observations on the Laboratory
Report.
Third Period
Materials:
new culture in glass- stoppered bottle (from
previous lab period)
Petri plate with nitrate succinate-mineral salts
agar
microscope slides and cover glasses
gram staining kit
GasPak anaerobic j ar, generator envelopes, and
generator strips
1
2
3
Examine the second enrichment bottle, looking
for bubbles. Check its clarity, also.
Streak a nitrate succinate-mineral salts agar plate
from this second enrichment bottle. Set the plate
aside to incubate in a GasPak jar at 30° C until the
next laboratory period.
Prepare a gram-stained slide from the second
enrichment culture and examine it under oil
immersion.
218
Benson: Microbiological
X. Microbiology of Soil
62. Isolation of a Denitrifier
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
from Soil: Using an
Enrichment Medium
Companies, 2001
^^MMH
^-^,,-^
One gram of fresh soil is added to a
flask of enrichment broth, which is
shaken vigorously to mix.
Supernatant is carefully decanted into a
glass stoppered bottle that is placed on
a Petri dish cover to contain overflow
during incubation.
Nitrate Succinate-Mineral Salts Broth
Incubated at 30° C for 3 to 5 days.
Culture is examined for the presence of
gas. A stream of bubbles should be visi-
ble rising from the bottom to the top
near the stopper.
Incubated at 30° C for 3 to 5 days.
£U/
/
M
Wet mount slides and gram-stained
slides are made for microscopic exam
ination. Phase-contrast microscopy
should be used for wet mounts.
A second enrichment culture is made by trans-
ferring 1 ml of initial enrichment to a second
stoppered bottle and filling the bottle with
fresh enrichment medium.
Figure 62.1 Procedure for culturing Paracoccus denitrificans from a soil sample
219
Benson: Microbiological
X. Microbiology of Soil
62. Isolation of a Denitrifier
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
from Soil: Using an
Enrichment Medium
Companies, 2001
Exercise 62 • Isolation of a Denitrifier from Soil: Using an Enrichment Medium
4. Make a wet mount slide from the same cuture and
examine with phase-contrast optics.
5. Record all your observations on the Laboratory
Report.
Fourth Period
Materials:
Petri plate culture from last period
microscope slides and gram- staining kit
1 . Examine the colonies on the streak plate you in
cubated anaerobically from the last period.
Compare the characteristics of the colonies with
the characteristics of Paracoccus denitrificans
that are given on page 218. Do the colonies look
like P. denitrificans?
2. Make a gram-stained slide from one of the
colonies and examine the slide under the micro-
scope. How do the characteristics of the organism
match Be r gey 's Manual description? Record your
results on the Laboratory Report.
Laboratory Report
Complete the Laboratory Report for this exercise.
Wet mount and gram-stained slides are
made from the second enrichment cul-
ture. Microscopic characteristics are
noted.
After incubation, the second enrichment
bottle is examined (or the presence of
gas bubbles.
A nitrate succinate- mineral salts agar
plate is inoculated from the second
enrichment culture, using a good iso-
lation technique.
Incubate in GasPak jar 30° C - 2 to 3 days
Colony ^characteri sties are noted on the
nitrate succinate-mineral salts agar plate.
A gram-stained slide is made from a
typical colony to confirm characteristics.
^^■^■'■"■ M ' MW h MW ■ 1 1 Pi'i iJllhH iilJPMIh^fr^^— ^^
Figure 62.2 Procedure for getting a pure culture out of second enrichment culture
220
Benson: Microbiological
XI. Microbiology of Water
Introduction
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Companies, 2001
Part
Microbiology of Water
The microorganisms of natural waters are extremely diverse. The
numbers and types of bacteria present will depend on the amounts
of organic matter present, the presence of toxic substances, the wa-
ter's saline content, and environmental factors such as pH, temper-
ature, and aeration. The largest numbers of heterotrophic forms will
exist on the bottoms and banks of rivers and lakes where organic
matter predominates. Open water in the center of large bodies of wa-
ter, free of floating debris, will have small numbers of bacteria. Many
species of autotrophic types are present, however, that require only
the dissolved inorganic salts and minerals that are present.
The threat to human welfare by contamination of water supplies
with sewage is a prime concern of everyone. The enteric diseases
such as cholera, typhoid fever, and bacillary dysentery often result
in epidemics when water supplies are not properly protected or
treated. Thus, our prime concern in this unit is the sanitary phase
of water microbiology. The American Public Health Association in
its Standard Methods for the Examination of Water and Wastewater
has outlined acceptable procedures for testing water for sewage
contamination. The exercises of this unit are based on the proce-
dures in that book.
221
Benson: Microbiological
XI. Microbiology of Water
63. Bacteriological
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Examination of Water:
Qualitative Tests
Companies, 2001
63
Bacteriological Examination of Water:
Qualitative Tests
Water that contains large numbers of bacteria may be
perfectly safe to drink. The important consideration,
from a microbiological standpoint, is the kinds of mi-
croorganisms that are present. Water from streams
and lakes that contain multitudes of autotrophs and
saprophytic heterotrophs is potable as long as
pathogens for humans are lacking. The intestinal
pathogens such as those that cause typhoid fever,
cholera, and bacillary dysentery are of prime concern.
The fact that human fecal material is carried away by
water in sewage systems that often empty into rivers
and lakes presents a colossal sanitary problem; thus,
constant testing of municipal water supplies for the
presence of fecal microorganisms is essential for the
maintenance of water purity.
Routine examination of water for the presence of
intestinal pathogens would be a tedious and difficult,
if not impossible, task. It is much easier to demon-
strate the presence of some nonpathogenic intestinal
types such as Escherichia coli or Streptococcus fae-
calis. Since these organisms are always found in the
intestines, and normally are not present in soil or wa-
ter, it can be assumed that their presence in water in-
dicates that fecal material has contaminated the water
supply.
E. coli and S. faecalis are classified as good
sewage indicators. The characteristics that make
them good indicators of fecal contamination are (1)
they are normally not present in water or soil, (2) they
are relatively easy to identify, and (3) they survive a
little longer in water than enteric pathogens. If they
were hardy organisms, surviving a long time in water,
they would make any water purity test too sensitive.
Since both organisms are non-spore-formers, their
survival in water is not extensive.
E. coli and S. faecalis are completely different
organisms. E. coli is a gram-negative non-spore-
forming rod; S. faecalis is a gram-positive coccus.
The former is classified as a coliform; the latter is an
enterococcus. Physiologically, they are also com-
pletely different.
The series of tests depicted in figure 63.1 is based
on tests that will demonstrate the presence of a coliform
in water. By definition, a coliform is a facultative
anaerobe that ferments lactose to produce gas and is a
gram-negative, non-spore-forming rod. Escherichia
coli and Enterobacter aerogenes fit this description.
Since S. faecalis is not a coliform, a completely differ-
ent set of tests must be used for it.
Note that three different tests are shown in figure
63.1: presumptive, confirmed, and completed. Each
test exploits one or more of the characteristics of a co-
liform. A description of each test follows.
Presumptive Test In the presumptive test a series
of 9 or 12 tubes of lactose broth are inoculated with
measured amounts of water to see if the water con-
tains any lactose-fermenting bacteria that produce
gas. If, after incubation, gas is seen in any of the lac-
tose broths, it is presumed that coliforms are present
in the water sample. This test is also used to determine
the most probable number (MPN) of coliforms pres-
ent per 100 ml of water.
Confirmed Test In this test, plates of Levine EMB
agar or Endo agar are inoculated from positive (gas-
producing) tubes to see if the organisms that are
producing the gas are gram-negative (another co-
liform characteristic). Both of these media inhibit
the growth of gram-positive bacteria and cause
colonies of coliforms to be distinguishable from
nonconforms. On EMB agar coliforms produce
small colonies with dark centers (nucleated
colonies). On Endo agar coliforms produce reddish
colonies. The presence of coliform-like colonies
confirms the presence of a lactose-fermenting
gram-negative bacterium.
Completed Test In the completed test our concern
is to determine if the isolate from the agar plates truly
matches our definition of a coliform. Our media for
this test include a nutrient agar slant and a Durham
tube of lactose broth. If gas is produced in the lactose
tube and a slide from the agar slant reveals that we
have a gram-negative non-spore-forming rod, we can
be certain that we have a coliform.
The completion of these three tests with posi-
tive results establishes that coliforms are present;
however, there is no certainty that E. coli is the co-
liform present. The organism might be E. aero-
genes. Of the two, E. coli is the better sewage indi-
cator since E. aerogenes can be of nonsewage
origin. To differentiate these two species, one must
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XI. Microbiology of Water
63. Bacteriological
Examination of Water:
Qualitative Tests
© The McGraw-H
Companies, 2001
Bacteriological Examination of Water: Qualitative Tests • Exercise 63
I M M^II
^^W*¥4*fa
WATER
SAMPLE
T
•r
1
*.'■ +■ 1
1
■s;
! Vii
*•
>
■ i.
^
■ ■
ml
■ **■•
-■
-J:
1/
*.
."■■
y
, J
.■
mm'
■J
■ ■ .V'
-■
"*
If
"■ /■-*
.
n
r>
V.
D
10 10
,S. Lactose Broth
1.0
n
1.0
^
4
.5:
n
0.1
n
0.1
Single Strength Lactose Broth
35° C
NEGATIVE
PRESUMPTIVE
The absence of gas in
all lactose broth tubes
indicates col if or ms
are absent and water
is safe to drink. Test
stops here.
DOUBTFUL
POSITIVE
If gas is present only
after 48 hours, the gas
is probably not due to
coliforrns. Further
testing is necessary.
NEGATIVE
n
POSITIVE RESULT
If 1 0% or more gas is
present in one or more
tubes in 24 hours, water
is presumed to be unsafe
to drink.
P
R
E
S
u
M
P
T
I
V
E
DOUBTFUL RESULT
After 24 and 48 hours
incubation the tubes
of lactose broth are
examined for gas pro-
duction. MPN deter-
mination is made
from Table VI,
Appendix A.
POSITIVE RESULT
J
Plates of Levine EMB
agar are streaked from pos
itive and doubtful tubes of
lactose broth. Endo agar
may also be used. Plates
are incubated at 35° C for
24 hours.
C
o
N
F
I
R
M
E
D
NEGATIVE CONFIRMED
The absence of typical con-
form colonies on Levine
EMB agar indicates that gas
in presumptive test was not
due to coliforms. Test
stops here.
I
Typical col i form colo-
nies are selected for in-
oculation of nutrient
agar slant and lactose
broth.
LACTOSE BROTH
POSITIVE CONFIRMED
The presence of typical
colifornn colonies indicates
that gas in presumptive
tubes was due to conforms.
These colonies have dark
centers and may have a
greenish metallic sheen.
W
NUTRIENT AGAR
SLANT
After 24 hours incubation at 35° C a gram-
stained slide is made from the slant. If the
organisms present are gram-negative non-
spore-forming rods and produce gas from
lactose, the completed test is positive.
C
O
M
P
L
E
T
E
D
11
*p^p
JdilMIII^
Figure 63, Bacteriological analysis of water
223
Benson: Microbiological
XI. Microbiology of Water
63. Bacteriological
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Examination of Water:
Qualitative Tests
Companies, 2001
Exercise 63 • Bacteriological Examination of Water: Qualitative Tests
perform the IMViC tests, which are described on
page 175 in Exercise 50.
In this exercise, water will be tested from local
ponds, streams, swimming pools, and other sources
supplied by students and instructor. Enough known
positive samples will be evenly distributed through-
out the laboratory so that all students will be able to
see positive test results. All three tests in figure 63.1
will be performed. If time permits, the IMViC tests
may also be performed.
The Presumptive Test
As stated earlier, the presumptive test is used to de-
termine if gas-producing lactose fermenters are pres-
ent in a water sample. If clear surface water is being
tested, nine tubes of lactose broth will be used as
shown in figure 63.1. For turbid surface water an ad-
ditional three tubes of single strength lactose broth
will be inoculated.
In addition to determining the presence or ab-
sence of coliforms, we can also use this series of lac-
tose broth tubes to determine the most probable
number (MPN) of coliforms present in 100 ml of
water. A table for determining this value from the
number of positive lactose tubes is provided in
Appendix A.
Before setting up your test, determine whether
your water sample is clear or turbid. Note that a sep-
arate set of instructions is provided for each type of
water.
Clear Surface Water
If the water sample is relatively clear, proceed as
follows:
Materials:
3 Durham tubes of DSLB
6 Durham tubes of SSLB
1 1 ml pipette
1 1 ml pipette
Note: DSLB designates double strength lactose
broth. It contains twice as much lactose as
SSLB (single strength lactose broth).
1 . Set up 3 DSLB and 6 SSLB tubes as illustrated in
figure 63.1. Label each tube according to the
amount of water that is to be dispensed to it: 10
ml, 1.0 ml, and 0.1 ml, respectively.
2. Mix the bottle of water to be tested by shaking 25
times .
3. With a 10 ml pipette, transfer 10 ml of water to
each of the DSLB tubes.
4. With a 1.0 ml pipette, transfer 1 ml of water to
each of the middle set of tubes, and 0. 1 ml to each
of the last three SSLB tubes.
5
6
7
8
Incubate the tubes at 35° C for 24 hours.
Examine the tubes and record the number of tubes
in each set that have 10% gas or more.
Determine the MPN by referring to table VI,
Appendix A. Consider the following:
Example: If you had gas in the first three tubes
and gas only in one tube of the second series, but
none in the last three tubes, your test would be
read as 3-1-0. Table VI indicates that the MPN
for this reading would be 43. This means that this
particular sample of water would have approxi-
mately 43 organisms per 100 ml with 95% prob-
ability of there being between 7 and 210 organ-
isms. Keep in mind that the MPN figure of 43 is
only a statistical probability figure.
Record the data on the Laboratory Report.
Turbid Surface Water
If your water sample appears to have considerable
pollution, do as follows:
Materials:
3 Durham tubes of DSLB
9 Durham tubes of SSLB
1 1 ml pipette
2 1 ml pipettes
1 water blank (99 ml of sterile water)
Note: See comment in previous materials list
concerning DSLB and SSLB.
1. Set up three DSLB and nine SSLB tubes in a test-
tube rack, with the DSLB tubes on the left.
2. Label the three DSLB tubes 10 ml, the next three
SSLB tubes 1.0 ml, the next three SSLB tubes
0.1 ml, and the last three tubes 0.01 ml.
3. Mix the bottle of water to be tested by shaking
25 times.
4. With a 10 ml pipette, transfer 10 ml of water to
each of the DSLB tubes.
5. With a 1.0 ml pipette, transfer 1 ml to each of the
next three tubes, and 0. 1 ml to each of the third set
of tubes.
6. With the same 1 ml pipette, transfer 1 ml of wa-
ter to the 99 ml blank of sterile water and shake
25 times.
7. With afresh 1 ml pipette, transfer 1.0 ml of water
from the blank to the remaining tubes of SSLB.
This is equivalent to adding 0.01 ml of full-
strength water sample.
8. Incubate the tubes at 35° C for 24 hours.
9. Examine the tubes and record the number of tubes
in each set that have 10% gas or more.
10. Determine the MPN by referring to table VI,
Appendix A. This table is set up for only 9 tubes.
To apply a 12-tube reading to it, do as follows:
224
Benson: Microbiological
XI. Microbiology of Water
63. Bacteriological
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Examination of Water:
Qualitative Tests
Companies, 2001
Bacteriological Examination of Water: Qualitative Tests • Exercise 63
a. Select the three consecutive sets of tubes that
have at least one tube with no gas.
b. If the first set of tubes (10 ml tubes) are not
used, multiply the MPN by 10.
Example: Your tube reading was 3-3-3-1. What
is the MPN?
The first set of tubes (10 ml) is ignored and
the figures 3-3-1 are applied to the table. The
MPN for this series is 460. Multiplying this by 10,
the MPN becomes 4600.
Example: Your tube reading was 3-1-2-0. What
is the MPN?
The first three numbers are (3-1-2) applied to
the table. The MPN is 210. Since the last set of
tubes is ignored, 210 is the MPN.
The Confirmed Test
Once it has been established that gas-producing lac-
tose fermenters are present in the water, it is presumed
to be unsafe. However, gas formation may be due to
noncoliform bacteria. Some of these organisms, such
as Clostridium perfringens, are gram-positive. To
confirm the presence of gram-negative lactose fer-
menters, the next step is to inoculate media such as
Levine eosin-methylene blue agar or Endo agar from
positive presumptive tubes.
Levine EMB agar contains methylene blue,
which inhibits gram-positive bacteria. Gram-negative
lactose fermenters (coliforms) that grow on this
medium will produce "nucleated colonies" (dark cen-
ters). Colonies of E. coli and E. aerogenes can be dif-
ferentiated on the basis of size and the presence of a
greenish metallic sheen. E. coli colonies on this
medium are small and have this metallic sheen,
whereas E. aerogenes colonies usually lack the sheen
and are larger. Differentiation in this manner is not
completely reliable, however. It should be remem-
bered that E. coli is the more reliable sewage indica-
tor since it is not normally present in soil, while E.
aerogenes has been isolated from soil and grains.
Endo agar contains a fuchsin sulfite indicator
that makes identification of lactose fermenters rela-
tively easy. Coliform colonies and the surrounding
medium appear red on Endo agar. Nonfermenters of
lactose, on the other hand, are colorless and do not af-
fect the color of the medium.
In addition to these two media, there are several
other media that can be used for the confirmed test.
Brilliant green bile lactose broth, Eijkman's medium,
and EC medium are just a few examples that can be
used.
To demonstrate the confirmation of a positive
presumptive in this exercise, the class will use Levine
EMB agar and Endo agar. One half of the class will
use one medium; the other half will use the other
medium. Plates will be exchanged for comparisons.
Materials:
1 Petri plate of Levine EMB agar (odd-
numbered students)
1 Petri plate of Endo agar (even-numbered
students)
1
2
3
Select one positive lactose broth tube from the
presumptive test and streak a plate of medium ac-
cording to your assignment. Use a streak method
that will produce good isolation of colonies. If all
your tubes were negative, borrow a positive tube
from another student.
Incubate the plate for 24 hours at 35° C.
Look for typical coliform colonies on both kinds
of media. Record your results on the Laboratory
Report. If no coliform colonies are present, the
water is considered bacteriologically safe to
drink.
Note: In actual practice, confirmation of all pre-
sumptive tubes would be necessary to ensure ac-
curacy of results.
The Completed Test
A final check of the colonies that appear on the con-
firmatory media is made by inoculating a nutrient
agar slant and a Durham tube of lactose broth. After
incubation for 24 hours at 35° C, the lactose broth is
examined for gas production. A gram-stained slide is
made from the slant, and the slide is examined under
oil immersion optics.
If the organism proves to be a gram- negative,
non- spore-forming rod that ferments lactose, we
know that coliforms were present in the tested water
sample. If time permits, complete these last tests and
record the results on the Laboratory Report.
The IMViC Tests
Review the discussion of the IMViC tests on page
175. The significance of these tests should be much
more apparent at this time. Your instructor will indi-
cate whether these tests should also be performed if
you have a positive completed test.
Benson: Microbiological
XI. Microbiology of Water
64. The Membrane Filter
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Method
Companies, 2001
The Membrane Filter Method
In addition to the multiple tube test, a method utilizing
the membrane filter has been recognized by the United
States Public Health Service as a reliable method for
the detection of coliforms in water. These filter disks
are 150 micrometers thick, have pores of 0.45 microm-
eter diameter, and have 80% area perforation. The pre-
cision of manufacture is such that bacteria larger than
0.47 micrometer cannot pass through. Eighty percent
area perforation facilitates rapid filtration.
To test a sample of water, the water is passed
through one of these filters. All bacteria present in the
sample will be retained directly on the filter's surface.
The membrane filter is then placed on an absorbent
pad saturated with liquid nutrient medium and incu-
bated for 22 to 24 hours. The organisms on the filter
disk will form colonies that can be counted under the
microscope. If a differential medium such as m Endo
MF broth is used, coliforms will exhibit a characteris-
tic golden metallic sheen.
The advantages of this method over the multiple
tube test are (1) higher degree of reproducibility of re-
sults; (2) greater sensitivity since larger volumes of
water can be used; and (3) shorter time (one-fourth)
for getting results.
Figure 64.1 illustrates the procedure we will use
in this experiment.
Materials:
vacuum pump or water faucet aspirators
membrane filter assemblies (sterile)
side-arm flask, 1000 ml size, and rubber hose
sterile graduates (100 ml or 250 ml size)
sterile, plastic Petri dishes, 50 mm dia
(Millipore #PD 1 047 00)
sterile membrane filter disks (Millipore
#HAWG 047 AO)
sterile absorbent disks (packed with filters)
sterile water
5 ml pipettes
bottles of m Endo MF broth (50 ml)*
water samples
1
2
3
4
5
6
7
8
Prepare a small plastic Petri dish as follows:
a. With a flamed forceps, transfer a sterile ab-
sorbent pad to a sterile plastic Petri dish.
b. Using a 5 ml pipette, transfer 2.0 ml of m Endo
MF broth to the absorbent pad.
Assemble a membrane filtering unit as follows:
a. Aseptically insert the filter holder base into the
neck of a 1 -liter side-arm flask.
b. With a flamed forceps, place a sterile mem-
brane filter disk, grid side up, on the filter
holder base.
c. Place the filter funnel on top of the membrane
filter disk and secure it to the base with the
clamp.
Attach the rubber hose to a vacuum source (pump
or water aspirator) and pour the appropriate
amount of water into the funnel.
The amount of water used will depend on wa-
ter quality. No less than 50 ml should be used.
Waters with few bacteria and low turbidity permit
samples of 200 ml or more. Your instructor will
advise you as to the amount of water that you
should use. Use a sterile graduate for measuring
the water.
Rinse the inner sides of the funnel with 20 ml of
sterile water.
Disconnect the vacuum source, remove the fun-
nel, and carefully transfer the filter disk with ster-
ile forceps to the Petri dish of m Endo MF broth.
Keep grid side up.
Incubate at 35° C for 22 to 24 hours. Don't
invert.
After incubation, remove the filter from the dish
and dry for 1 hour on absorbent paper.
Count the colonies on the disk with low-power
magnification, using reflected light. Ignore all
colonies that lack the golden metallic sheen. If
desired, the disk may be held flat by mounting
between two 2" X 3" microscope slides after dry-
ing. Record your count on the first portion of
Laboratory Report 64, 65.
See Appendix C for special preparation method.
226
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XI. Microbiology of Water
64. The Membrane Filter
Method
© The McGraw-H
Companies, 2001
The Membrane Filter Method • Exercise 64
Sterile absorbent pad is aseptically placed in the bottom
of a sterile plastic Petri dish.
A sterile membrane filter disk is placed on filter holder
base with grid side up.
Absorbent pad is saturated with 20 ml of m Endo MF
broth.
u i i ■> : Lii iij- m ,
Water sample isvptftired into assembled funnel, utilizing
vacuum. A rinse of 20 ml of sterile water follows.
Filter disk is carefully removed with sterile forceps after
disassembling the funnel.
Membrane filter disk is placed on medium-soaked absor
bent pad with grid side up. Incubate at 35" C 24 hours.
Figure 64.1 Membrane filter routine
227
Benson: Microbiological
XI. Microbiology of Water
65. Standard Plate Count: A
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Quantitative Test
Companies, 2001
■
Standard Plate Count:
A Quantitative Test
In determining the total numbers of bacteria in wa-
ter, we are faced with the same problems that are en-
countered with soil. Water organisms have great
variability in physiological needs, and no single
medium, pH, or temperature is ideal for all types.
Despite the fact that only small numbers of organ-
isms in water will grow on nutrient media, the stan-
dard plate count can perform an important function
in water testing. Probably its most important use is
to give us a tool to reveal the effectiveness of vari-
ous stages in the purification of water. Plate counts
made of water before and after storage, for example,
can tell us how effective holding is in reducing bac-
terial numbers.
In this exercise, various samples of water will be
evaluated by routine standard plate count proce-
dures. Since different dilution procedures are re-
quired for different types of water, two methods are
given.
3
Tap Water Procedure
If the water is of low bacterial count, such as in the
case of tap water, use the following method.
Materials:
1 .0 ml pipettes
2 tryptone glucose extract agar pours (TGEA)
2 sterile Petri plates
Quebec colony counter and hand counters
water samples
1 . Liquefy two tubes of TGEA and cool to 45° C.
2. After shaking the sample of water 25 times trans-
fer 1 ml of water to each of the two sterile Petri
plates.
4
5
Pour the medium into the dishes, rotate suffi-
ciently to get good mixing of medium and water,
and let cool.
Incubate at 35° C for 24 hours.
Count the colonies of both plates on the Quebec
colony counter and record your average count of
the two plates on the Laboratory Report.
Surface Water Procedure
If the water is likely to have a high bacterial count, as
in the case of surface water, proceed as follows:
Materials:
1 bottle (75 ml) of tryptone glucose extract agar
(TGEA)
6 sterile Petri plates
2 water blanks (99 ml)
1 .0 ml pipettes
1
2
3
4
5
6
Liquefy a bottle of TGEA medium and cool to
45° C
After shaking your water sample 25 times, pro-
duce two water blanks with dilutions of 1 : 100 and
1:1000. See Exercise 23.
Distribute aliquots from these blanks to six Petri
dishes, which will provide you with two plates
each of 1:100, 1:1000, and 1:10,000 dilutions.
Pour one-sixth of the TGEA medium into each
plate and rotate sufficiently to get even mixing of
the water and medium.
Incubate at 35° C for 24 hours.
Select the pair of plates that has 30 to 300
colonies on each plate and count all the colonies
on both plates. Record the average count for the
two plates on the second portion of Laboratory
Report 64, 65.
228
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XII. Microbiology of Milk
and Food Products
Introduction
© The McGraw-H
Companies, 2001
Part
Microbiology of Milk and Food
Products
Milk and food provide excellent growth media for bacteria when
suitable temperatures exist. This is in direct contrast to natural wa-
ters, which lack the essential nutrients for pathogens. The intro-
duction of a few pathogens into food or milk products becomes a
much more serious problem because of the ability of these sub-
stances to support tremendous increases in bacterial numbers.
Many milk-borne epidemics of human diseases have been spread
by contamination of milk by soiled hands of dairy workers, unsan-
itary utensils, flies, and polluted water supplies. The same thing can
be said for improper handling of foods in the home, restaurants,
hospitals, and other institutions.
We learned in Part 1 1 that bacteriological testing of water is pri-
marily qualitative — emphasis being placed on the presence or ab-
sence of coliforms as indicators of sewage. Bacteriological testing
of milk and food may also be performed in this same manner, us-
ing similar media and procedures to detect the presence of coli-
forms. However, most testing by public health authorities is quan-
titative. Although the presence of small numbers of bacteria in
these substances does not necessarily mean that pathogens are
lacking, low counts do reflect better care in handling of food and
milk than is true when high counts are present.
Standardized testing procedures for milk products are outlined
by the American Public Health Association in Standard Methods for
the Examination of Dairy Products. The procedures in Exercises 66,
67, and 67 are excerpts from that publication. Copies of the book
may be available in the laboratory as well as in the library.
Exercises 69, 70, and 71 pertain to bacterial counts in dried fruit
and meats, as well as to spoilage of canned vegetables and meats.
Since bacterial counts in foods are performed with some of the
techniques you have learned in previous exercises, you will have an
opportunity to apply some of those skills here. Exercises 72 and 73
pertain to fermentation methods used in the production of wine and
yogurt.
229
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XII. Microbiology of Milk
and Food Products
66. Standard Plate Count of
Milk
© The McGraw-H
Companies, 2001
Standard Plate Count of Milk
The bacterial count in milk is the most reliable indi-
cation we have of its sanitary quality. It is for this rea-
son that the American Public Health Association rec-
ognizes the standard plate count as the official method
in its Milk Ordinance and Code. Although human
pathogens may not be present in a high count, it may
indicate a diseased udder, unsanitary handling of
milk, or unfavorable storage temperatures. In general,
therefore, a high count means that there is a greater
likelihood of disease transmission. On the other hand,
it is necessary to avoid the wrong interpretation of low
plate counts, since it is possible to have pathogens
such as the brucellosis and tuberculosis organisms
when counts are within acceptable numbers. Routine
examination and testing of animals act as safeguards
against the latter situation.
In this exercise, standard plate counts will be made
of two samples of milk: a supposedly good sample and
one of known poor quality. Odd-numbered students will
work with the high-quality milk and even-numbered stu-
dents will test the poor-quality sample. A modification
of the procedures in Exercise 23 will be used.
1
High-Ouality Milk
Materials:
milk sample
1 sterile water blank (99 ml)
4 sterile Petri plates
1 . 1 ml dilution pipettes
1 bottle of TGEA (40 ml)
Quebec colony counter
mechanical hand counter
2
3
Following the procedures used in Exercise 23,
pour four plates with dilutions of 1 : 1 , 1:10, 1 : 1 00,
and 1:1000. Before starting the dilution proce-
dures, shake the milk sample 25 times in the cus-
tomary manner.
Incubate the plates at 35° C for 24 hours and
count the colonies on the plate that has between
30 and 300 colonies.
Record your results on the first portion of
Laboratory Report 66, 67.
Poor-Quality Milk
Materials:
milk sample
3 sterile water blanks (99 ml)
4 sterile Petri plates
1 . 1 ml dilution pipettes
1 bottle TGEA (50 ml)
Quebec colony counter
mechanical hand counter
1
2
3
Following the procedures used in Exercise 23,
pour four plates with dilutions of 1:10,000,
1:100,000, 1:1,000,000, and 1:10,000,000. Before
starting the dilutions, shake the milk sample 25
times in the customary manner.
Incubate the plates at 35° C for 24 hours and
count the colonies on the plate that has between
30 and 300 colonies.
Record your results on the first portion of
Laboratory Report 66, 67.
230
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XII. Microbiology of Milk
and Food Products
67. Direct Microscopic
Count of Organisms in
Milk: The Breed Count
© The McGraw-H
Companies, 2001
Direct Microscopic Count of Organisms
in Milk:
The Breed Count
67
When it is necessary to determine milk quality in a
much shorter time than is possible with a standard
plate count, one can make a direct microscopic
count on a slide. This is accomplished by staining a
measured amount of milk that has been spread over an
area one square centimeter on a slide. The slide is ex-
amined under oil and all of the organisms in an entire
microscopic field are counted. To increase accuracy,
several fields are counted to get average field counts.
Before the field counts can be translated into organ-
isms per milliliter, however, it is necessary to calcu-
late the field area.
High-quality milk will have very few organisms
per field, necessitating the examination of many
fields. A slide made of poor-quality milk, on the other
hand, will reveal large numbers of bacteria per field,
thus requiring the examination of fewer fields. An ex-
perienced technician can determine, usually within
15 minutes, whether or not the milk is of acceptable
quality.
In addition to being much faster than the SPC, the
direct microscopic count has two other distinct ad-
vantages. First of all, it will reveal the presence of
bacteria that do not form colonies on an agar plate at
35° C; thermophiles, psychrophiles, and dead bacteria
would fall in this category. Secondly, the presence of
excessive numbers of leukocytes and pus-forming
streptococci on a slide will be evidence that the ani-
mal that produced the milk has an udder infection
(mastitis).
In view of all these advantages, it is apparent that
the direct microscopic count has real value in milk
testing. It is widely used for testing raw milk in
creamery receiving stations and for diagnosing the
types of contamination and growth in pasteurized
milk products.
In this exercise, samples of raw whole milk will
be examined. Milk that has been separated, blended,
homogenized, and pasteurized will lack leukocytes
and normal flora.
Slide Preparation
There are several acceptable ways of spreading the
milk onto the slide. Figure 67.1 illustrates a method
using a guide card. The Breed slide used in figure 67.2
Figure 67.1 Using a guide card to spread milk sample
over one square centimeter on a slide
has five one- centimeter areas that are surrounded by
ground glass, obviating the need for a card. Proceed as
follows:
Materials:
Breed slide or guide card
Breed pipettes (0.01 ml)
methylene blue, xylol, 95% alcohol
beaker of water and electric hot plate
samples of raw milk (poor and high quality)
1
2
3
4
5
6
Shake the milk sample 25 times to completely dis-
perse the organisms and break up large clumps of
bacteria.
Transfer 0.01 ml of milk to one square on the
slide. The pipette may be filled by capillary ac-
tion or by suction, depending on the type of
pipette. The instructor will indicate which
method to use. Be sure to wipe off the outside tip
of the pipette with tissue before touching the slide
to avoid getting more than 0.01 ml on the slide.
Allow the slide to air-dry and then place it over
a beaker of boiling water for 5 minutes to steam-
fix it.
Flood the slide with xylol to remove fat globules.
Remove the xylol from the slide by flooding the
slide with 95 % ethyl alcohol.
Gently immerse the slide into a beaker of distilled
water to remove the alcohol. Do not hold it under
running water; the milk film will wash off.
231
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XII. Microbiology of Milk
and Food Products
67. Direct Microscopic
Count of Organisms in
Milk: The Breed Count
© The McGraw-H
Companies, 2001
Exercise 67 • Direct Microscopic Count of Organisms in Milk: The Breed Count
7
8
9
Stain the smear with methylene blue for 15 sec-
onds and dip the slide again in water to remove
the excess stain.
Decolorize the smear to pale blue with 95% alco-
hol and dip in distilled water to stop decolorization.
Allow the slide to completely air-dry before ex-
amination.
Calibration of Microscope
(Microscope Factor [MF])
Before counting the organisms in each field it is nec-
essary to know what part of a milliliter of milk is rep-
resented in that field. The relationship of the field to a
milliliter is the microscope factor (MF). To calculate
the MF, it is necessary to use a stage micrometer to
measure the diameter of the oil immersion field. By
applying the formula irr 2 to this measurement, the
area is easily determined. With the amount of milk
(0.01 ml) and the area of the slide (1 cm 2 ), it is a sim-
ple matter to calculate the MF.
Materials:
stage micrometer
1
2
3
4
5
Place a stage micrometer on the microscope stage
and bring it into focus under oil. Measure the di-
ameter of the field, keeping in mind that each
space is equivalent to 0.01 mm.
Calculate the area of the field in square millime-
ters, using the formula ttt 2 (it = 3.14).
Convert the area of the field from square mil-
limeters to square centimeters by dividing by 100.
Calculate the number of fields in one square cen-
timeter by dividing one square centimeter by the
area of the field in square centimeters.
To get the part of a milliliter that is represented in
a single field (microscope factor), multiply the
number of fields by 100. The value should be
around 500,000. Therefore, a single field repre-
sents 1/500,000 of a ml of milk. Record your
computations on the Laboratory Report.
Examination of Slide
Two methods of counting the bacteria can be used: in-
dividual cells may be tallied or only clumps of bacte-
ria may be counted. In both cases, the number per mil-
liliter will be higher than a standard plate count, but a
A measured amount of milk (.01 ml)
is spread over one sq. cm. area of
Breed slide.
Slide is flooded gently with xylol to
remove fat. Removal of xylol is ac-
complished with alcohol.
Smears are air-dried. Four or five
minutes may be required for com-
plete drying.
tttfcrita^Mfe
lta— —ta— ^
* ■"- H f *
Smears are steamed over boiling
water to fix organisms to the slide,
After immersing slide in distilled water
to remove alcohol, smears are stained
with methylene blue for 1 5 seconds.
Smears are decolorized to a robin's
egg blue with alcohol. Immersion in
distilled water stops decolorization.
Figure 67.2 Procedure for making a stained slide of a raw milk sample
232
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XII. Microbiology of Milk
and Food Products
67. Direct Microscopic
Count of Organisms in
Milk: The Breed Count
© The McGraw-H
Companies, 2001
Direct Microscopic Count of Organisms in Milk: The Breed Count • Exercise 67
clump count will be closer to the SPC. Both methods
will be used.
3
1
2.
After the microscope has been calibrated, replace
the stage micrometer with the stained slide.
Examine it under oil immersion optics.
Count the individual cells in five fields and record
your results on the Laboratory Report. A field is the
entire area encompassed by the oil immersion lens.
As you see leukocytes, record their numbers, also.
4.
Count only clumps of bacteria in five fields,
recording the numbers of leukocytes as well.
Record the totals on the Laboratory Report.
Calculate the number of organisms, clumps, and
body cells per milliliter using the microscope factor.
Laboratory Report
Complete the last portion of Laboratory Report 66, 67
Clean high-grade milk will have very few, if any, bacteria.
Milk that is placed in improperly cleaned utensils wi
exhibit masses of miscellaneous bacteria.
Milk from a cow with mastitis. Long chain streptococci
and numerous leukocytes are visible.
High-grade milk that is allowed to stand without cooling
will reveal numerous streptococci as short chains and
diplococci.
Figure 67.3 Microscopic fields of milk samples
233
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XII. Microbiology of Milk
and Food Products
68. The Reductase Test
© The McGraw-H
Companies, 2001
Reductase Test
Milk that contains large numbers of actively growing
bacteria will have a lowered oxidation-reduction po-
tential due to the exhaustion of dissolved oxygen by
microorganisms. The fact that methylene blue loses
its color (becomes reduced) in such an environment
is the basis for the reductase test. In this test, 1 ml of
methylene blue (1:25,000) is added to 10 ml of milk.
The tube is sealed with a rubber stopper and slowly
inverted three times to mix. It is placed in a water
bath at 35° C and examined at intervals up to 6 hours.
The time it takes for the methylene blue to become
colorless is the methylene blue reduction time
(MBRT). The shorter the MBRT, the lower the qual-
ity of milk. An MBRT of 6 hours is very good. Milk
with an MBRT of 30 minutes is of very poor quality.
The validity of this test is based on the assump-
tion that all bacteria in milk lower the oxidation-
reduction potential at 35° C. Large numbers of psy-
chrophiles, thermophiles, and thermodurics, which do
not grow at this temperature, would not produce a
positive test. Raw milk, however, will contain pri-
marily Streptococcus lactis and Escherichia coli,
which are strong reducers; thus, this test is suitable for
screening raw milk at receiving stations. Its principal
value is that less technical training of personnel is re-
quired for its performance.
Methylene
Blue
.r. h 'ii
Rubber
Stopper
35° C
Water Bath
GOOD QUALITY MILK
Methylene blue is not
reduced within 6 hours.
One ml methylene blue is
added to 10 ml milk. I
Tube is inverted three
times after plugging with
stopper.
POOR QUALITY MILK
Methylene blue is
reduced within 2 hours
Figure 68.1 Procedure for testing raw milk with reductase test
234
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XII. Microbiology of Milk
and Food Products
68. The Reductase Test
© The McGraw-H
Companies, 2001
In this exercise, samples of low- and high-quality
raw milk will be tested.
Materials:
2 sterile test tubes with rubber stoppers for each
student
raw milk samples of low- and high-quality
(samples A and B)
water bath set at 35° C
methylene blue (1:25,000)
1 ml pipettes
1 ml pipettes
gummed labels
1 . Attach gummed labels with your name and type
of milk to two test tubes. Each student will test a
good-quality as well as a poor-quality milk.
2. Using separate 10 ml pipettes for each type of
milk, transfer 10 ml to each test tube. To the milk
in the tubes add 1 ml of methylene blue with a 1
ml pipette. Insert rubber stoppers and gently in-
3
4
The Reductase Test • Exercise 68
vert three times to mix. Record your name and the
time on the labels and place the tubes in the water
bath, which is set at 35° C.
After 5 minutes incubation, remove the tubes
from the bath and invert once to mix. This is the
last time they should be mixed.
Carefully remove the tubes from the water bath
30 minutes later and every half hour until the end
of the laboratory period. When at least four- fifths
of the tube has turned white, the end point of re-
duction has taken place. Record this time on the
Laboratory Report. The classification of milk
quality is as follows:
Class 1: Excellent, not decolorized in 8 hours.
Class 2: Good, decolorized in less than 8 hours,
but not less than 6 hours.
Class 3: Fair, decolorized in less than 6 hours,
but not less than 2 hours.
Class 4: Poor, decolorized in less than 2 hours.
235
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XII. Microbiology of Milk
and Food Products
69. Bacterial Counts of
Foods
© The McGraw-H
Companies, 2001
Bacterial Counts of Foods
The standard plate count, as well as the multiple tube
test, can be used on foods much in the same manner that
they are used on milk and water to determine total counts
and the presence of coliforms. To get the organisms in
suspension, however, a food blender is necessary.
In this exercise, samples of ground meat, dried
fruit, and frozen food will be tested for total numbers
of bacteria. This will not be a coliform count. The in-
structor will indicate the specific kinds of foods to be
tested and make individual assignments. Figure 69.1
illustrates the general procedure.
Materials:
per student:
3 Petri plates
1 bottle (45 ml) of Plate Count agar or
Standard Methods agar
1 99 ml sterile water blank
2 1.1 ml dilution pipettes
per class:
food blender
sterile blender jars (one for each type of food)
sterile weighing paper
1
2
3
4
5
6
7
180 ml sterile water blanks (one for each type
of food)
samples of ground meat, dried fruit, and frozen
vegetables, thawed 2 hours
Using aseptic techniques, weigh out on sterile
weighing paper 20 grams of food to be tested.
Add the food and 1 80 ml of sterile water to a ster-
ile mechanical blender jar. Blend the mixture for
5 minutes. This suspension will provide a 1:10
dilution.
With a 1.1 ml dilution pipette dispense from the
blender 0. 1 ml to plate I and 1 .0 ml to the water
blank. See figure 69.1.
Shake the water blank 25 times in an arc for 7 sec-
onds with your elbow on the table as done in
Exercise 23 (Bacterial Population Counts).
Using a fresh pipette, dispense 0. 1 ml to plate III
and 1 .0 ml to plate II.
Pour agar (50° C) into the three plates and incu-
bate them at 35° C for 24 hours.
Count the colonies on the best plate and record
the results on the Laboratory Report.
20 grams of food is blended in
180 ml of sterile water for 5
minutes.
1:10,000
1:100
1:1000
Figure 69.1 Dilution procedure for bacterial counts of food
236
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XII. Microbiology of Milk
and Food Products
70. Microbial Spoilage of
Canned Foods
© The McGraw-H
Companies, 2001
Microbial Spoilage of Canned Food
70
Spoilage of heat-processed, commercially canned
foods is confined almost entirely to the action of bac-
teria that produce heat-resistant endospores. Canning
of foods normally involves heat exposure for long pe-
riods of time at temperatures that are adequate to kill
spores of most bacteria. Particular concern is given to
the processing of low- acid foods in which
Clostridium botulinum can thrive to produce botulism
food poisoning.
Spoilage occurs when the heat processing fails to
meet accepted standards. This can occur for several
reasons: (1) lack of knowledge on the part of the
processor (usually the case in home canning); (2)
carelessness in handling the raw materials before can-
ning, resulting in an unacceptably high level of con-
tamination that ordinary heat processing may be inad-
equate to control; (3) equipment malfunction that
results in undetected underprocessing; and (4) defec-
tive containers that permit the entrance of organisms
after the heat process.
Our concern here will be with the most common
types of food spoilage caused by heat-resistant spore-
forming bacteria. There are three types: "flat sour,"
"T.A. spoilage," and "stinker spoilage."
Flat sour pertains to spoilage in which acids are
formed with no gas production; result: sour food in
cans that have flat ends. T.A. spoilage is caused by
thermophilic anaerobes that produce acid and gases
(C0 2 and H 2 , but not H 2 S) in low-acid foods. Cans
swell to various degrees, sometimes bursting. Stinker
spoilage is due to spore-formers that produce hydro-
gen sulfide and blackening of the can and contents.
Blackening is due to the reaction of H 2 S with the iron
in the can to form iron sulfide.
In this experiment you will have an opportunity to
become familiar with some of the morphological and
_
h>
1
Each can of corn or peas is
perforated with an awl or ice pick.
To create an air space under
the cover, some liquid is
poured off.
Contents of each can is
inoculated with one of five
different organisms.
SECOND PERIOD
1 . Type of spoilage caused by each orga-
nism is noted.
2. Gram- and spore-stained slides are made
from contents of cans.
24-48 Hours
Incubation
For Temperature
See text
4
Hole in each can is sealed by
soldering over it.
Figure 70.1 Canned food inoculation procedure
237
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XII. Microbiology of Milk
and Food Products
70. Microbial Spoilage of
Canned Foods
© The McGraw-H
Companies, 2001
Exercise 70 • Microbial Spoilage of Canned Food
physiological characteristics of organisms that cause
canned food spoilage, including both aerobic and anaer-
obic endospore formers of Bacillus and Clostridium, as
well as a non-spore-forming bacterium.
Working as a single group, the entire class will in-
oculate 10 cans of vegetables (corn and peas) with
five different organisms. Figure 70.1 illustrates the
procedure. Note that the cans will be sealed with sol-
der after inoculation and incubated at different tem-
peratures. After incubation the cans will be opened so
that stained microscope slides can be made to deter-
mine Gram reaction and presence of endospores. Your
instructor will assign individual students or groups of
students to inoculate one or more of the 10 cans. One
can of corn and one can of peas will be inoculated
with each of the organisms. Proceed as follows:
First Period
(Inoculations)
Materials:
5 small cans of corn
5 small cans of peas
cultures of B. stearothermophilus,
B. coagulans, C. sporogenes,
C. thermosaccharolyticum, and E.
ice picks or awls
hammer
solder and soldering iron
plastic bags
gummed labels and rubber bands
1
2
3
4
coli
5
Label the can or cans with the name of the organ-
ism that has been assigned to you. Use white
gummed labels. In addition, place a similar label
on one of the plastic bags to be used after sealing
of the cans.
With an ice pick or awl, punch a small hole
through a flat area in the top of each can. This can
be done easily with the heel of your hand or a
hammer, if available.
Pour off a small amount of the liquid from the can
to leave an air space under the lid.
Use an inoculating needle to inoculate each can of
corn or peas with the organism indicated on the
label.
Take the cans up to the demonstration table where
the instructor will seal the hole with solder.
6
7
After sealing, place each can in two plastic bags.
Each bag must be closed separately with rubber
bands, and the outer bag must have a label on it.
Incubation will be as follows till the next period:
• 55° C — C. thermosaccharolyticum and
B. stearothermophilus
• 37° C — C. sporogenes and B. coagulans
• 30° C — E. coli
Note: If cans begin to swell during incubation,
they should be placed in refrigerator.
Second Period
(Interpretation)
After incubation, place the cans under a hood to open
them. The odors of some of the cans will be very
strong due to H 2 S production.
Materials:
can opener, punch type
small plastic beakers
Parafilm
gram- staining kit
spore- staining kit
1. Open each can carefully with a punch-type can
opener. If the can is swollen, hold an inverted
plastic funnel over the can during perforation to
minimize the effects of any explosive release of
contents.
2. Remove about 10 ml of the liquid through the
opening, pouring it into a small plastic beaker.
Cover with Parafilm. This fluid will be used for
making stained slides.
3 . Return the cans of food to the plastic bags, reclose
them, and dispose in a proper trash bin.
4. Prepare gram-stained and endospore-stained
slides from your canned food extract as well as
from the extracts of all the other cans. Examine
under brightfield oil immersion.
5. Record your observations on the report sheet on
the demonstration table. It will be duplicated and
a copy will be made available to each student.
Laboratory Report
Complete the first portion of Laboratory Report 70, 7 1
238
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XII. Microbiology of Milk
and Food Products
71. Microbial Spoilage of
Refrigerated Meats
© The McGraw-H
Companies, 2001
Microbial Spoilage of Refrigerated Meat
71
Contamination of meats by microbes occurs during
and after slaughter. Many contaminants come from
the animal itself, others from utensils and equip-
ment. The conditions for rapid microbial growth in
freshly cut meats are very favorable, and spoilage
can be expected to occur rather quickly unless steps
are taken to prevent it. Although immediate refriger-
ation is essential after slaughter, it will not prevent
spoilage indefinitely, or even for a long period of
time under certain conditions. In time, cold-tolerant
microbes will destroy the meat, even at low refriger-
ator temperatures.
Microorganisms that grow at temperatures be-
tween 5° and 0° C are classified as being either psy-
chrophilic or psychro trophic. The difference be-
tween the two groups is that psychrophiles seldom
grow at temperatures above 22° C and psy-
chrotrophs (psychrotolerants or low-temperature
mesophiles) grow well above 25° C. While the opti-
mum growth temperature range for psychrophiles is
15°-18° C, psychrotrophs have an optimum growth
temperature range of 25°-30° C. It is the psy-
chrotrophic microorganisms that cause most meat
spoilage during refrigeration.
The majority of psychrophiles are gram-negative
and include species of Aeromonas, Alcaligenes,
Cytophagia, Flavobacterium, Pseudomonas, Serratia,
and Vibrio. Gram-positive psychrophiles include
species of Arthrobacter, Bacillus, Clostridium, and
Micrococcus.
Psychrotrophs include a much broader spectrum
of gram-positive and gram-negative rods, cocci,
vibrios, spore-formers, and non-spore-formers. Typi-
cal genera are Acinetobacter, Chromobacterium, Cit-
robacter, Cory neb acterium, Enter ob act er, Escheri-
chia, Klebsiella, Lactobacillus, Moraxella, Staphy-
lococcus, and Streptococcus.
The widespread use of vacuum or modified at-
mospheric packaging of raw and processed meat has
resulted in food spoilage due to facultative and obli-
gate anaerobes, such as Lactobacillus, Leuconostoc,
Pediococcus, and certain Enterobacteriaceae.
Although most of the previously mentioned psy-
chrotrophic representatives are nonpathogens, there
are significant pathogenic psychrotrophs such as
Aeromonas hydrophila, Clostridium botulinum, Lis-
teria monocytogenes, Vibrio cholera, Yersinia enter-
colitica, and some strains of E. coli.
In addition to bacterial spoilage of meat there are
many yeasts and molds that are psychrophilic and psy-
chrotrophic. Examples of psychrophilic yeasts are
Cryptococcus, Leucosporidium, and Torulopsis.
Psychrotrophic fungi include Candida, Cryptococcus,
Saccharomyces, Alternaria, Aspergillus, Cladosporium,
Fusarium, Mucor, Penicillium, and many more.
Our concern in this experiment will be to test
one or more meat samples for the prevalence of
psychrophilic-psychrotrophic organisms. To accom-
plish this, we will liquefy and dilute out a sample of
ground meat so that it can be plated out and then in-
cubated in a refrigerator for 2 weeks. After incuba-
tion, colony counts will be made to determine the
number of organisms of this type that exist in a gram
of the sample.
Figure 71.1 illustrates the overall procedure.
Work in pairs to perform the experiment.
First Period
Materials:
at demonstration table:
ground meat and balance
sterile foil- wrapped scoopula
1 blank of phosphate buffered water (90 ml)
blender with sterile blender jar
sterile Petri dish or sterile filter paper
per pair of students:
4 large test tubes of sterile phosphate buffered
water (9 ml each)
4 TS A plates
9 sterile 1 ml pipettes
L- shaped glass spreading rod
beaker of 95% ethyl alcohol
At Demonstration Table
1 . With a sterile scoopula, weigh 1 Og of ground meat
into a sterile Petri plate or onto a sterile piece of
filter paper.
239
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XII. Microbiology of Milk
and Food Products
71. Microbial Spoilage of
Refrigerated Meats
© The McGraw-H
Companies, 2001
Exercise 71 • Microbial Spoilage of Refrigerated Meat
2. Pour 90 ml of sterile buffered water from water
blank into a sterile blender jar and add the meat.
3. Blend the meat and water at moderate speed for 1
minute.
Student Pair
1
2
3
4
5
6
7
Label the four water blanks 1 through 4.
Label the four Petri plates with their dilutions, as
indicated in figure 71.1. Add your initials and
date also.
Once blender suspension is ready, pipette 1 ml
from jar to tube 1.
Using a fresh 1 ml pipette, mix the contents in
tube 1 and transfer 1 ml to tube 2.
Repeat step 4 for tubes 3 and 4, using fresh
pipettes for each tube.
Dispense 0. 1 ml from each tube to their respective
plates of TSA. Note that by using only 0.1 ml per
plate you are increasing the dilution factor by 1
times in each plate.
Using a sterile L- shaped glass rod, spread the or-
ganisms on the agar surfaces. Sterilize the rod
8.
each time by dipping in alcohol and flaming gen-
tly. Be sure to let rod cool completely each time.
Incubate the plates for 2 weeks in the back of the
refrigerator (away from door-opening) where the
temperature will remain between 0° and 5° C.
Second Period
Materials:
Quebec colony counters
hand tally counters
gram- staining kit
1
2
After incubation, count the colonies on all the
plates and calculate the number of psychrophiles
and psychrotrophs per gram of meat.
Select a colony from one of the plates and prepare
a gram-stained slide. Examine under oil immer-
sion and record your observations on the
Laboratory Report.
Laboratory Report
Complete the last portion of Laboratory Report 70, 7 1
1
Ten grams of ground meat is
added to 90 ml of water and
blended for 1 minute.
A tenfold serial dilution is made by
transferring 1 ml from each tube to the
next one.
9 ml water
per tube
1
v..
^
<J
\J>
I J I
0.1 ml is dispensed from each tube to a TSA plate
1:1,000
1:10,000
1:100,000 1:1,000,000
An alcohol-flamed glass rod is used to spread
organisms on the surfaces of each of the four
agar plates.
4
After spreading out of organisms on the agar
surfaces, the plates are incubated at 0°-5° C for
2 weeks.
Figure 71.1 Dilution and inoculation procedure
240
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XII. Microbiology of Milk
and Food Products
72. Microbiology of
Alcohol Fermentation
© The McGraw-H
Companies, 2001
Microbiology of Alchohol Fermentation
72
Fermented food and beverages are as old as civiliza-
tion. Historical evidence indicates that beer and wine
making were well established as long ago as 2000 B.C.
An Assyrian tablet states that Noah took beer aboard
the ark.
Beer, wine, vinegar, buttermilk, cottage cheese,
sauerkraut, pickles, and yogurt are some of the more
commonly known products of fermentation. Most of
these foods and beverages are produced by different
strains of yeasts (Saccharomyces) or bacteria
(Lactobacillus, Acetobacter, etc.).
Fermentation is actually a means of food preser-
vation because the acids formed and the reduced en-
vironment (anaerobiasis) hold back the growth of
many spoilage microbes.
Wine is essentially fermented fruit juice in which
alcoholic fermentation is carried out by Saccharomyces
cerevisiae var. ellipsoideus. Although we usually asso-
ciate wine with fermented grape juice, it may also be
made from various berries, dandelions, rhubarb, etc.
Three conditions are necessary: simple sugar, yeast,
and anaerobic conditions. The reaction is as follows:
QH 12 o 6
yeast
> 2CoH.OH + 2CO
2 AA 5
2
Commercially, wine is produced in two forms: red
and white. To produce red wines, the distillers use red
grapes with the skins left on during the initial stage of
the fermentation process. For white wines either red
or white grapes can be used, but the skins are dis-
carded. White and red wines are fermented at 13° C
(55° F) and 24° C (75° F), respectively.
In this exercise we will set up a grape juice fer-
mentation experiment to learn about some of the char-
acteristics of sugar fermentation to alcohol. Note in
figure 72.1 that a balloon will be attached over the
mouth of the fermentation flask to exclude oxygen up-
take and to trap gases that might be produced. To de-
tect the presence of hydrogen sulfide production we
will tape a lead acetate test strip inside the neck of the
flask. The pH of the substrate will also be monitored
before and after the reaction to note any changes that
occur.
Mouth of flask is sealed with
rubber balloon before incubation
1 00 ml of grape juice is
inoculated with 3 ml of yeast
culture.
Lead acetate test strip is taped
to inside of flask neck.
15°-17° C
2-5 Days
Balloon is removed after incu-
bation. Odor of gas and test strip
change are noted.
pH of juice-yeast mixture is de-
termined before incubation.
pH of fermented juice is checked
after incubation.
Figure 72.1 Alcohol fermentation setup
241
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XII. Microbiology of Milk
and Food Products
72. Microbiology of
Alcohol Fermentation
© The McGraw-H
Companies, 2001
Exercise 72 • Microbiology of Alchohol Fermentation
First Period
Materials:
100 ml grape juice (no preservative)
bottle of juice culture of wine yeast
1 25 ml Erlenmeyer flask
1 1 ml pipette
balloon
hydrogen sulfide (lead acetate) test paper
tape
pH meter
1
2
3
4
5
Label an Erlenmeyer flask with your initials and
date.
Add about 100 ml of grape juice to the flask (fer-
menter) .
Determine the pH of the juice with a pH meter
and record the pH on the Laboratory Report.
Agitate the container of yeast juice culture to sus-
pend the culture, remove 5 ml with a pipette, and
add it to the flask.
Attach a short strip of tape to a piece of lead- acetate
test paper (3 cm long), and attach it to the inside
surface of the neck of the flask. Make certain that
neither the tape nor the test strip protrudes from the
flask.
6. Cover the flask opening with a balloon.
7. Incubate at 1 5 °-l 7° C for 2-5 days.
Second Period
Materials:
pH meter
1
2
3
4
Remove the balloon and note the aroma of the
flask contents. Describe the odor on the
Laboratory Report.
Determine the pH and record it on the Laboratory
Report.
Record any change in color of the lead- acetate- test
strip on the Laboratory Report. If any H 2 S is pro-
duced, the paper will darken due to the formation
of lead sulfide as hydrogen sulfide reacts with the
lead acetate.
Wash out the flask and return it to the drain rack.
Laboratory Report
Complete the first portion of Laboratory Report 72,
73 by answering all the questions.
242
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XII. Microbiology of Milk
and Food Products
73. Microbiology of Yogurt
Production
© The McGraw-H
Companies, 2001
Microbiology of Yogurt Production
73
For centuries, people throughout the world have
been producing fermented milk products using
yeasts and lactic acid-producing bacteria. The yo-
gurt of eastern central Europe, the kefir of the
Cossacks, the koumiss of central Asia, and the leben
of Egypt are just a few examples. In all of these fer-
mented milks, lactobacilli act together with some
other microorganisms to curdle and thicken milk,
producing a distinctive flavor desired by the pro-
ducer. Kefir of the Cossacks is made by charging
milk with small cauliflower-like grains that contain
Streptococcus lactis, Saccharomyces delbrueckii,
and Lactobacillus brevis. As the grains swell in the
milk they release the growing microorganisms to fer-
ment the milk. The usual method for producing yo-
gurt in large-scale production is to add pure cultures
of Streptococcus thermophilus and Lactobacillus
bulgaricus to pasteurized milk.
In this exercise you will produce a batch of yogurt
from milk by using an inoculum from commercial yo-
gurt. Gram-stained slides will be made from the fin-
ished product to determine the types of organisms that
control the reaction. If proper safety measures are fol-
lowed, the sample can be tasted.
Two slightly different ways of performing this ex-
periment are provided here. Your instructor will indi-
cate which method will be followed.
Method A
(First Period)
Figure 73.1 illustrates the procedure for this method.
Note that 4 g of powdered milk are added to 1 00 ml of
whole milk. This mixture is then heated to boiling and
cooled to 45°C. After cooling, the milk is inoculated with
yogurt and incubated at 45° C for 24 hours. Proceed:
Dried Milk Powder
1
Four grams of dried milk powder is
dissolved in 100 ml of whole milk.
2
Milk is brought to boiling point while
stirring constantly.
noculum
1.
2.
SECOND PERIOD
Product is evaluated with respect
to texture, color, aroma, and taste
Slides, stained with methylene
blue, are studied to determine
morphology of organisms.
45° C
24 Hours
3
Once heated milk has cooled to 45° C, one
teaspoonful of yogurt is stirred into it. Beaker is
then covered with plastic wrap and incubated.
Figure 73.1 Yogurt production by Method A
243
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XII. Microbiology of Milk
and Food Products
73. Microbiology of Yogurt
Production
© The McGraw-H
Companies, 2001
Exercise 73 • Microbiology of Yogurt Production
Materials:
dried powdered milk
whole milk
commercial yogurt (with viable organisms)
small beaker, graduate, teaspoon, stirring rod
plastic wrap
filter paper (for weighing)
1
2
3
4.
5
On a piece of filter paper weigh 4 grams of dried
powdered milk.
To a beaker of 1 00 ml of whole milk add the pow-
dered milk and stir thoroughly with sterile glass
rod to dissolve.
Heat to boiling, while stirring constantly.
Cool to 45 ° C and inoculate with 1 teaspoon of the
commercial yogurt. Stir. Be sure to check the la-
bel to make certain that product contains a live
culture. Cover with plastic wrap.
Incubate at 45° C for 24 hours.
Method B
(First Period)
Figure 73.2 illustrates a slightly different method of
culturing yogurt, which, due to its simplicity, may be
preferred. Note that no whole milk is used and provi-
sions are made for producing a sample for tasting.
Materials:
small beaker, graduate, teaspoon, stirring rod
dried powdered milk
commercial yogurt (with viable organisms)
plastic wrap
filter paper for weighing
1
2
3
4
5
1
paper Dixie cup (5 oz size) and cover
electric hot plate or Bunsen burner and tripod
On a piece of filter paper weigh 25 grams of dried
powdered milk.
Heat 100 ml of water in a beaker to boiling and
cool to 45° C.
Add the 25 grams of powdered milk and 1 tea-
spoon of yogurt to the beaker of water. Mix the in-
gredients with a sterile glass rod.
Pour some of the mixture into a sterile Dixie cup
and cover loosely. Cover the remainder in the
beaker with plastic wrap.
Incubate at 45° C for 24 hours.
Second Period
(Both Methods)
Examine the product and record on the Laboratory
Report the color, aroma, texture, and, if desired,
the taste.
CAUTION
Refrain from working with other bacteria or doing
other exercises while tasting the yogurt.
2. Make slide preparations of the yogurt culture. Fix
and stain with methylene blue. Examine under oil
immersion and record your results on Laboratory
Report 72, 73.
Laboratory Report
Complete the last portion of Laboratory Report 72, 73
by answering all the questions.
1
2
100 ml of water is boiled in
a clean small beaker.
Twenty-five grams of dried powdered milk
and a teaspoonful of commercial yogurt
are stirred into the 100 ml of water at 45° C
Water is cooled down
to 45° C.
3
SECOND PERIOD
1 . Product is evaluated with respect to texture,
color, aroma, and taste. Sample in Dixie cup can be
used for tasting.
2. Slides, stained with methylene blue, are studied
to determine morphology of organisms.
ncubated at 45° C
for 24 hours.
Sample for tasting
Figure 73.2 Yogurt production by Method B
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XIII. Bacterial Genetic
Variations
Introduction
© The McGraw-H
Companies, 2001
Part
Bacterial Genetic Variations
Variations in bacteria that are due to environmental factors and that
do not involve restructuring DNA are designated as temporary
variations. Such variations may be morphological or physiological
and disappear as soon as the environmental changes that brought
them about disappear. For example, as a culture of E. coli becomes
old and the nutrients within the tube become depleted, the new
cells that form become so short that they appear coccoidal.
Reinoculation of the organism into fresh media, however, results in
the reappearance of distinct bacilli of characteristic length.
Variations in bacteria that involve alteration of the DNA macro-
molecule are designated as permanent variations. It is because
they survive a large number of transfers that they are so named.
Such variations are due to mutations. Variations of this type occur
spontaneously. They also might be induced by physical and chem-
ical methods. Some permanent variations also are caused by the
transfer of DNA from one organism to another, either directly by
conjugation or indirectly by phage. It is these permanent genetic
variations that the three exercises of this unit represent.
Exercises 74 and 75 of this unit demonstrate how spontaneous
mutations are constantly occurring in bacterial populations. The
genetic change that occurs in these two exercises pertains to the
development of bacterial resistance to streptomycin. In Exercise 76
we will study how chemically induced mutagenicity that causes
back mutations is used in the Ames test to determine possible car-
cinogenicity of chemical compounds.
245
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XIII. Bacterial Genetic
Variations
74. Mutant Isolation by
Gradient Plate Method
© The McGraw-H
Companies, 2001
7A
Mutant Isolation by Gradient Plate Method
An excellent way to determine the ability of organ-
isms to produce mutants that are resistant to antibi-
otics is to grow them on a gradient plate of a par-
ticular antibiotic. Such a plate consists of two
different wedgelike layers of media: a bottom layer
of plain nutrient agar and a top layer of nutrient agar
with the antibiotic. Since the antibiotic is only in the
top layer, it tends to diffuse into the lower layer, pro-
ducing a gradient of antibiotic concentration from
low to high.
In this exercise we will make a gradient plate us-
ing streptomycin in the medium. E. coli, which is nor-
mally sensitive to this antibiotic, will be spread over
the surface of the plate and incubated for 4 to 7 days.
Any colonies that develop in the high concentration
area will be streptomycin-resistant mutants.
Plate Preparation
The gradient plate used in this experiment will have a
high concentration of 100 meg of streptomycin per
milliliter of medium. This concentration is 10 times
the strength used in sensitivity disks in the Kirby-
Bauer test method. Prepare a gradient plate as follows:
Materials:
1 sterile Petri plate
2 nutrient agar pours (10 ml per tube)
1 tube of streptomycin solution (1%)
1 wood spacer (%" X M" X 2")
1 ml pipette
1 . Liquefy two pours of nutrient agar and cool to
50° C.
2. With wood spacer under one edge of Petri plate
(see figure 74.1), pour contents of one agar pour
into plate. Let stand until solidification has
occurred.
3. Remove the wood spacer from under the plate.
4. Pipette 0.1 ml of streptomycin into second agar
pour, mix tube between palms, and pour contents
over medium of plate that is now resting level on
the table.
5. Label the low and high concentration areas on the
bottom of the plate.
Inoculation
The inoculation procedure is illustrated in figure 74.2.
The technique involves spreading a measured amount
Plain nutrient agar is poured into Petri dish with plate in
slant position.
Streptomycin agar is poured over plain agar with plate in
normal position.
Figure 74.1 Procedure for pouring a gradient plate
246
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XIII. Bacterial Genetic
Variations
74. Mutant Isolation by
Gradient Plate Method
© The McGraw-H
Companies, 2001
Mutant Isolation by Gradient Plate Method • Exercise 74
of the culture on the surface of the medium with a
glass bent rod to provide optimum distribution.
Materials:
1 beaker of 95% ethanol
1 glass rod spreader
nutrient broth culture of E.
1
2
3
4.
coli
1 ml pipette
Pipette 0.1 ml of E. coli suspension onto surface
of medium in Petri plate.
Sterilize glass spreading rod by dipping it in alco-
hol first and then passing it quickly through the
flame of a Bunsen burner. Cool the rod by placing
against sterile medium in plate before contacting
organisms.
Spread the culture evenly over the surface with
the glass rod.
Invert and incubate the plate at 37° C for 4 to 7
days in a closed cannister or plastic bag. Unless
incubated in this manner, excessive dehydration
might occur.
First Evaluation
After 4 to 7 days, look for colonies of E. coli in the
area of high streptomycin concentration. Count the
colonies that appear to be resistant mutants and record
your count on the Laboratory Report.
Select a well-isolated colony in the high concen-
tration area and, with a sterile loop, smear the colony
over the surface of the medium toward the higher con-
centration portion of the plate. Do this with two or
three colonies. Return the plate to the incubator for
another 2 or 3 days.
Final Evaluation
Examine the plate again to note what effect the
spreading of the colonies had on their growth. Record
your observations on Laboratory Report 74, 75.
Spreading rod is dipped in ethanol for cleaning.
Rod is sterilized in Bunsen burner
flame.
Organisms are spread evenly over surface of agar.
Figure 74.2 Procedure for spreading organisms on gradient plate
247
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XIII. Bacterial Genetic
Variations
75. Mutant Isolation by
Replica Plating
© The McGraw-H
Companies, 2001
75
Mutant Isolation by Replica Plating
In the last exercise it was observed that E. coll could
develop mutant strains that are streptomycin-resistant.
If we had performed this experiment with other organ-
isms and with other antibiotics, the results would have
been quite similar. The question that logically devel-
ops in one's mind from this experiment is: What mech-
anism is involved here? Is a mutation of this sort in-
duced by the antibiotic? Or does the mutation occur
spontaneously and independently of the presence of
the drug? If we could demonstrate the presence of a
streptomycin-resistant mutant occurring on a medium
that lacks streptomycin, then we could assume that the
mutation occurs spontaneously.
To determine whether or not such a colony exists
on a plain agar plate having 500 to 1,000 colonies
could be a laborious task. One would have to transfer
organisms from each colony to a medium containing
streptomycin. This is somewhat self-defeating, too, in
light of the low incidence of mutations that occur.
Many thousands of the transfers might have to be
made to find the first mutant. Fortunately, we can re-
sort to replica plating to make all the transfers in one
step. Figure 75.1 illustrates the procedure. In this
technique a velveteen-covered colony transfer device
is used to make the transfers.
Note in figure 75.1 that organisms are first dis-
persed on nutrient agar with a glass spreading rod.
After incubation, all colonies are transferred from the
nutrient agar plate to two other plates: first to a nutri-
ent agar plate and second to a streptomycin agar plate.
After incubation, streptomycin-resistant strains are
looked for on the streptomycin agar.
1
Organisms are spread over nutrient
agar with a steril bent glass rod.
v.
:•;.
4
Streptomycin agar is inoculated with
same carrier in same manner.
2
After incubation, colonies are picked
up with velveteen colony carrier.
3
Nutrient agar is inoculated by lightly
pressing the carrier onto it.
Figure 75.1 Replica plating technique
248
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XIII. Bacterial Genetic
Variations
75. Mutant Isolation by
Replica Plating
© The McGraw-H
Companies, 2001
First Period
Materials:
1 Petri plate of nutrient agar
1 bent glass spreading rod
1 ml serological pipette
beaker of 95% ethanol
Bunsen burner
broth culture of E. coli
1
2
Pipette 0. 1 ml of E. coli from broth culture to sur-
face of medium in Petri dish.
With a sterile bent glass rod, spread the organisms
over the plate following the routine shown in fig-
ure 75.1.
3. Incubate this plate at 37° C for 24 hours.
Second Period
Materials:
1 Petri plate culture of E. coli from previous
period
1 Petri plate of nutrient agar per student
Mutant Isolation by Replica Plating • Exercise 75
1 Petri plate of streptomycin agar
( 1 00 micrograms of streptomycin per ml
of medium)
1 sterile colony carrier per student
1 . Carefully lower the sterile colony carrier onto the
colonies of E. coli on the plate from the previous
period.
2. Inoculate the plate of nutrient agar by lightly
pressing the carrier onto the medium.
3. Now without returning the carrier to the original
culture plate, inoculate the streptomycin agar in
the same manner.
4. Incubate both plates at 37° C for 2 to 4 days in an
enclosed cannister.
Third Period
Materials:
Quebec colony counter and hand counter
1. Examine both plates and record the information
called for on Laboratory Report 74, 75 .
2. Tabulate the results of other members of the class.
249
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XIII. Bacterial Genetic
Variations
76. Bacterial Mutagenicity
and Carcinogenesis: The
Ames Test
© The McGraw-H
Companies, 2001
76
Bacterial Mutagenicity and Carcinogenesis:
The Ames Test
The fact that carcinogenic compounds induce in-
creased rates of mutation in bacteria has led to the use
of bacteria for screening chemical compounds for
possible carcinogenesis. The Ames test, developed
by Bruce Ames at the University of California-
Berkeley, has been widely used for this purpose.
The conventional way to determine whether a
chemical substance is carcinogenic is to inject the
material into animals and look for the development of
tumors. If tumors develop, it is presumed that the
substance can cause cancer. Although this method
works well, it is costly, time-consuming, and cum-
bersome, especially if it is applied to all the industrial
chemicals that have found their way into food and
water supplies.
The Ames test serves as a screening test for the
detection of carcinogenic compounds by testing the
ability of chemical agents to induce bacterial muta-
tions. Although most mutagenic agents are carcino-
genic, some are not; however, the correlation between
carcinogenesis and mutagenicity is high — around
83%. Once it has been determined that a specific
agent is mutagenic, it can be used in animal tests to
confirm its carcinogenic capability.
The standard way to test chemicals for mutagene-
sis has been to measure the rate of back mutations in
strains of auxotrophic bacteria. In the Ames test a
strain of Salmonella typhimurium, which is aux-
otrophic for histidine (unable to grow in the absence of
histidine), is exposed to a chemical agent. After chem-
ical exposure and incubation on histidine-deficient
medium, the rate of reversion (back mutation) to pro-
totrophy is determined by counting the number of
colonies that are seen on the histidine-deficient
medium.
Although testing of chemicals for mutagenesis in
bacteria has been performed for a long time, two new
features are included in the Ames test that make it a
powerful tool. The first is that the strain of S. ty-
phimurium used here lacks DNA repair enzymes,
which prevents the correction of DNA injury. The sec-
ond feature of the test is the incorporation of mam-
malian liver enzyme preparations with the chemical
agent. The latter is significant because there is evi-
dence that liver enzymes convert many noncarcino-
genic chemical agents to carcinogenic ones.
There are two ways to perform the Ames test. The
method illustrated in figure 76.1 is a spot test that is
widely used for screening purposes. The other method
is the plate incorporation test, which is used for
quantitative analysis of the mutagenic effectiveness
of compounds. Our concern here will be with the spot
test; however, since the concentration of the liver ex-
tract is very critical, we will omit using it in our test.
The test, as performed here, will work well without it.
Success in performing the spot test requires con-
siderable attention to careful measurements and tim-
ing. It is for this reason that students will work in pairs
to perform the test.
Note in figure 76.1 that 0.1 ml of S. typhimurium
is first added to a small tube that contains 2 ml of top
agar that is held at 45° C. This top agar contains
0.6% agar, 0.5% NaCl, and a trace of histidine and
biotin. The histidine allows the bacteria to go
through several rounds of cell division, which is es-
sential for mutagenesis to occur. Since the histidine
deletion extends through the biotin gene, biotin is
also needed. This early growth of cells produces a
faint background lawn that is barely visible to the
naked eye.
Before pouring the top agar over the glucose-
minimal salts agar, the tube must be vortexed at slow
speed for 3 seconds and poured quickly to get even
distribution. The addition of the bacteria, vortexing,
and pouring must be accomplished in 20 seconds.
Failure to move quickly enough will cause stippling
of the top agar.
There are two ways that one can use to apply the
chemical agent to the top agar: a filter paper disk may
be used, or the chemical can be applied directly to the
center of the plate without a disk. The procedure
shown in figure 76.1 involves using a disk.
Note the unusual way in which a filter paper disk
is impregnated in figure 76.1. To get it to stand on
edge it must be put in position with sterile forceps and
pressed in slightly to hold it upright. Just the right
amount of the chemical agent is then added with a
Pasteur pipette to the upper edge of the disk to com-
pletely saturate it without making it dripping wet;
then the disk is lowered onto the top agar.
Once the test reagent is deposited on the top agar,
the plate is incubated at 37° C for 48 hours. If the
250
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XIII. Bacterial Genetic
Variations
76. Bacterial Mutagenicity
and Carcinogenesis: The
Ames Test
© The McGraw-H
Companies, 2001
Bacterial Mutagenicity and Carcinogenesis: The Ames Test • Exercise 76
0.1 ml of S. typhimurium, Ames
strain TA98 is pipetted into
liquefied top agar and mixed for 3
seconds on a Vortex mixer.
FOUR TEST PLATES ARE PREPARED FROM
FOUR TUBES OF TOP AGAR USING THIS
PROCEDURE. THE DISKS ON EACH PLATE
WILL RECEIVE DIFFERENT SOLUTIONS.
Top agar, containing traces
of histidine and biotin, is
liquefied and kept at 45° C.
Within 20 seconds of pipetting in
step 1 , top agar is poured over
glucose-minimal salts agar in plate
Filter paper disk is positioned on its edge
with sterile forceps. A Pasteur pipette is
used to saturate it with test compound.
37° C
48 hours
Disk is lowered with pipette tip
so that it lies horizontally
in contact with medium.
After 48 hours incubation, a positive test plate will have a
halo of high density revertants growing around the disk. The
large colonies beyond the halo are spontaneous back
mutations.
Figure 76.1 Procedure for performing a modified Ames test
251
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XIII. Bacterial Genetic
Variations
76. Bacterial Mutagenicity
and Carcinogenesis: The
Ames Test
© The McGraw-H
Companies, 2001
Exercise 76 • Bacterial Mutagenicity and Carcinogenesis: The Ames Test
agent is mutagenic, a halo of densely packed revertant
colonies will be seen around the disk. Scattered larger
colonies will show up beyond the halo that represent
spontaneous back mutations, not related to the test
reagent.
You will be issued an unknown chemical agent to
test and you will have an opportunity to test some
other substance you have brought to the laboratory. In
addition to these two tests you will be inoculating pos-
itive and negative test controls: thus, each pair of stu-
dents will be responsible for four plates.
Keep in mind as you perform this experiment that
there is a lot more to the Ames test than revealed here.
While we are using only one tester strain of S. ty-
phimurium, there are several others that are used in
routine testing. The additional strains are needed to
accommodate different kinds of chemical com-
pounds. While one chemical agent may be mutagenic
on one tester strain, it may produce a negative result
on another strain. Also, keep in mind that we are not
taking advantage of using the liver extract.
First Period
(Inoculations)
Materials:
per pair of students:
4 plates of glucose-minimal salts agar
(30 ml per plate)
4 tubes of top agar (2 ml per tube)
tube of sterile water
Vortex mixer
sterile Pasteur pipettes, forceps
serological pipettes ( 1 ml size)
filter paper disks, sterile in Petri dish
test reagents:
4-NOPD (10 |xg/ml) solution*
tube of unknown possible carcinogen
substance from home for testing
culture of S. typhimurium, Ames strain, TA98 in
trypticase soy broth
*4-nitro-o-phenylenediamine
1
2
3
4
Working with your laboratory partner, label the
bottoms of four glucose-minimal salts agar plates
as follows: POSITIVE CONTROL, NEGATIVE
CONTROL, UNKNOWN, and OPTIONAL.
Liquefy four tubes of top agar and cool to 45° C.
With a 1 ml serological pipette, inoculate a tube
of top agar with 0.1 ml of S. typhimurium.
Thoroughly mix the organisms into the top agar
by vortexing (slow speed) for 3 seconds, or
rolling the tube between the palms of both hands.
Pour the contents onto the positive control plate
of glucose-minimal salts agar. The agar plate
must be at room temperature. Work rapidly to
achieve pipetting, mixing, and spreading in 20
seconds.
5. Repeat steps 3 and 4 for each of the other three
tubes of top agar.
6. With sterile forceps place a disk on its edge near
the center of the positive control plate. Sterilize
the forceps by dipping in alcohol and flaming.
7. With a sterile Pasteur pipette, deposit just enough
4-NOPD on the upper edge of the disk to saturate
it; then, push over the disk with the pipette tip
onto the agar so that it lies flat.
8. Insert a sterile disk on the negative control plate
in the same manner as above. Moisten this disk
with sterile water, and reposition it flat on the agar
surface. Be sure to use a fresh Pasteur pipette.
9. Place a disk on the unknown plate, and, using the
same procedures, infiltrate it with your unknown,
and position it flat on the agar.
10. On the fourth plate (optional) deposit a drop of
your unknown from home. If the test substance
from home is crystalline, place a few crystals di-
rectly on the top agar of the optional plate in its
center. Liquid substances should be handled in
same manner as above.
11. Incubate all four plates for 48 hours at 37° C.
Second Period
(Evaluation)
Examine all four plates. You should have a pro-
nounced halo of revertant colonies around the disk on
the positive control plate and no, or very few, rever-
tants on the negative control plate. The presence of a
few scattered revertants on the negative control plate
is due to spontaneous back mutations, which always
occur. Examine the areas beyond the halo to see if you
can detect a faint lawn of bacterial growth.
CAUTION
Since much of the glassware in this experiment con-
tains carcinogens, do not dispose of any of it in the
usual manner. Your instructor will indicate how this
glassware is to be handled.
Record your results on the Laboratory Report and
answer all the questions.
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XIV. Medical Microbiology
and Immunology
Introduction
© The McGraw-H
Companies, 2001
Part
Medical Microbiology and
Immunology
Although many of the exercises up to this point in this manual per-
tain in some way to medical microbiology, they also have applica-
tions that are nonmedical. The exercises of this unit, however, are
primarily medical or dental in nature.
Medical (clinical) microbiology is primarily concerned with the
isolation and identification of pathogenic organisms. Naturally, the
techniques for studying each type of organism are different. A com-
plete coverage of this field of microbiology is very extensive, en-
compassing the Mycobacteriaceae, Brucellaceae, Enterobacte-
riaceae, Corynebacteriaceae, Micrococcaceae, ad infinitum. It is
not possible to explore all of these groups in such a short period of
time; however, this course would be incomplete if it did not include
some of the routine procedures that are used in the identification of
some of the more common pathogens.
Exercise 77 in this unit differs from the other 1 3 exercises in that
it pertains to the spread of disease (epidemiology) rather than to
specific microorganisms. Its primary function is to provide an un-
derstanding of some of the tools used by public health epidemiol-
ogists to determine the sources of infection in the disease trans-
mission cycle.
Since the most frequently encountered pathogenic bacteria are
the gram-positive pyogenic cocci and the intestinal organisms,
Exercises 78, 79, and 80 have been devoted to the study of those
bacteria. The exercise that provides the greatest amount of depth
is Exercise 79 (The Streptococci). To provide assistance in the iden-
tification of streptococci, it has been necessary to provide supple-
mentary information in Appendix E.
Four exercises (82, 83, 84, and 85) are related to various appli-
cations of the agglutination reaction to serological testing. Two of
these exercises pertain to slide tests and two of them are tube
tests. It is anticipated that the instructor will select those tests from
this group that fit time and budget limitations.
Exercises 87, 88, and 89 cover some of the basic hematological
tests that might be included in a microbiology laboratory. The last
exercise (90) pertains to an old test that has been revived pertain-
ing to caries susceptibility.
253
Benson: Microbiological
XIV. Medical Microbiology
77. A Synthetic Epidemic
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
and Immunology
Companies, 2001
77
A Synthetic Epidemic
A disease caused by microorganisms that enter the
body and multiply in the tissues at the expense of the
host is said to be an infectious disease. Infectious dis-
eases that are transmissible to other persons are con-
sidered to be communicable. The transfer of commu-
nicable infectious agents between individuals can be
accomplished by direct contact, such as in handshak-
ing, kissing, and sexual intercourse, or they can be
spread indirectly through food, water, objects, ani-
mals, and so on.
Epidemiology is the study of how, when, where,
what, and who are involved in the spread and distrib-
ution of diseases in human populations. An epidemi-
ologist is, in a sense, a medical detective who searches
out the sources of infection so that the transmission
cycle can be broken.
Whether an epidemic actually exists is deter-
mined by the epidemiologist by comparing the num-
ber of new cases with previous records. If the number
of newly reported cases in a given period of time in a
specific area is excessive, an epidemic is considered
to be in progress. If the disease spreads to one or more
continents, a pandemic is occurring.
In this experiment we will have an opportunity to
approximate, in several ways, the work of the epi-
demiologist. Each member of the class will take part
in the spread of a "synthetic infection." The mode of
transmission will be handshaking. For obvious safety
reasons, the agent of transmission will not be a
pathogen.
Two different approaches to this experiment are
given: procedures A and B. In procedure A a white
powder is used. In Procedure B two non-pathogens
(Micrococcus luteus and Serratia marcescens) will
be used. The advantage of procedure A is that it can
be completed in one laboratory session. Procedure B,
on the other hand, is more realistic in that viable or-
ganisms are used; however, it involves two periods.
Your instructor will indicate which procedure is to be
followed.
ered the infectious agent. The other members will be
issued a transmissible agent that is considered nonin-
fectious. After each student has spread the powder on
his or her hands, all members of the class will engage
in two rounds of handshaking, directed by the in-
structor. A record of the handshaking contacts will be
recorded on a chart similar to the one on the
Laboratory Report. After each round of handshaking,
the hands will be rubbed on blotting paper so that a
chemical test can be applied to it to determine the
presence or absence of the infectious agent.
Once all the data are compiled, an attempt will be
made to determine two things: (1) the original source
of the infection, and (2) who the carriers are. The type
of data analysis used in this experiment is similar to
the procedure that an epidemiologist would employ.
Proceed as follows:
Materials:
1 numbered container of white powder*
1 piece of white blotting paper
spray bottles of "developer solution"*
Preli
1
2
3
lminaries
After assembling your materials, write your name
and unknown number at the top of your sheet of
blotting paper. In addition, draw a line down the
middle, top to bottom, and label the left side
ROUND 1 and the right side ROUND 2.
Wash and dry your hands thoroughly.
Moisten the right hand with water and prepare it
with the agent by thoroughly coating it with the
white powder, especially on the palm surface.
This step is similar to the contamination that
would occur to one's hand if it were sneezed into
during a cold.
IMPORTANT: Once the hand has been prepared
do not rest it on the tabletop or allow it to touch
any other object.
Procedure A
In this experiment each student will be given a num-
bered container of white powder. Only one member in
the class will be given a powder that is to be consid-
*lnstructor: To prevent students from preguessing the outcome of
this experiment, the compositions of the powders and developer
solution are known only to the instructor. The Instructor's
Handbook provides this information.
254
Benson: Microbiological
XIV. Medical Microbiology
77. A Synthetic Epidemic
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
and Immunology
Companies, 2001
Round 1
1
2
3
On the cue of the instructor, you will begin the
first round of handshaking. Your instructor will
inform you when it is your turn to shake hands
with someone. You may shake with anyone, but it
is best not to shake your neighbor's hand. Be sure
to use only your treated hand, and avoid ex-
tracurricular glad-handing.
In each round of handshaking you will be selected
by the instructor only once for handshaking; how-
ever, due to the randomness of selection by the
handshakers, it is possible that you may be se-
lected as the "shakee" several times.
After every member of the class has shaken some-
one's hand, you need to assess just who might
have picked up the "microbe." To accomplish
this, wipe your fingers and palm of the contami-
nated hand on the left side of your blotting paper.
Press fairly hard, but don't tear the surface.
IMPORTANT: Don't allow your hand to touch
any other object. A second round of handshaking
follows.
Round 2
1
2
On the cue of your instructor, shake hands with
another person. Avoid contact with any other ob-
jects.
Once the second handshaking episode is finished,
rub the fingers and palm of the contaminated hand
on the right side of the blotting paper.
CAUTION: Keep your contaminated hand off
the left side of the blotting paper.
Chemical Identification
1
2
To determine who has been "infected" we will
now spray the developer solution on the hand-
prints of both rounds. One at a time, each student,
with the help of the instructor, will spray his or
her blotting paper with developer solution.
Color interpretation is as follows:
Blue: — positive for infectious agent
Brown or yellow: — negative
Tabulation of Results
1
2
3
Tabulate the results on the chalkboard, using a
table similar to the one on the Laboratory Report.
Once all results have been recorded, proceed to
determine the originator of the epidemic. The eas-
iest way to determine this is to put together a
flowchart of shaking.
Identify those persons that test positive. You will
be working backward with the kind of informa-
4.
A Synthetic Epidemic • Exercise 77
tion an epidemiologist has to work with (contacts
and infections). Eventually, a pattern will emerge
that shows which person started the epidemic.
Complete the Laboratory Report.
Procedure B
In this experiment each student will be given a piece
of hard candy that has had a drop of Micrococcus lu-
teus or Serratia marcescens applied to it. Only one
person in the class will receive candy with S.
marcescens, the presumed pathogen. All others will
receive M. luteus.
After each student has handled the piece of candy
with a glove-covered right hand, he or she will shake
hands (glove to glove) with another student as di-
rected by the instructor. A record will be kept of who
takes part in each contact. Two rounds of handshaking
will take place. After each round, a plate of trypticase
soy agar will be streaked.
After incubating the plates, a tabulation will be
made for the presence or absence of S. marcescens on
the plates. From the data collected, an attempt will be
made to determine two things: (1) the original source
of the infection and (2) who the carriers are. The type
of data analysis used in this experiment is similar to
the procedure that an epidemiologist would employ.
Proceed as follows:
CAUTION
Although the pathogenicity of S. marcescens is con-
sidered to be relatively low, avoid allowing any skin
contact during this experiment.
Materials:
sterile rubber surgical gloves ( 1 per student)
hard candy contaminated with M. luteus
hard candy contaminated with S. marcescens
sterile swabs (2 per student)
TS A plates (1 per student)
Preli
1
2
3
lm inane s
Draw a line down the middle of the bottom of a
TSA plate, dividing it into two halves. Label one
half ROUND 1 and the other ROUND 2.
Put a sterile rubber glove on your right hand.
Avoid contaminating the palm surface.
Grasp the piece of candy in your gloved hand,
rolling it around the surface of your palm. Discard
the candy into a beaker of disinfectant set aside
for disposal. You are now ready to do the first-
round handshake.
Benson: Microbiological
XIV. Medical Microbiology
77. A Synthetic Epidemic
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
and Immunology
Companies, 2001
Exercise 77 • A Synthetic Epidemic
Round 1
1
2
3
On the cue of your instructor, select someone to
shake hands with. You may shake with anyone,
but it is best not to shake hands with your
neighbor.
In each round of handshaking you will be selected
by the instructor only once for handshaking; how-
ever, due to the randomness of selection by the
handshakers, it is possible that you may be se-
lected as the "shakee" several times. The instruc-
tor or a recorder will record the initials of the
shaker and shakee each time.
After you have shaken someone's hand, swab the
surface of your palm and transfer the organisms to
the side of your plate designated as ROUND 1 .
Discard this swab into the appropriate container
for disposal.
Round 2
1
2
Again, on the cue of your instructor, select some-
one at random to shake hands with. Be sure not to
contaminate your gloved hand by touching some-
thing else.
With a fresh swab, swab the palm of your hand
and transfer the organisms to the side of your
3.
plate designated as ROUND 2. Make sure that
your initials and the initials of the shakee are
recorded by the instructor or recorder.
Incubate the TSA plate at room temperature for
48 hours.
Tabulation and Analysis
1
2
3
After 48 hours' incubation look for typical red S.
marcescens colonies on your Petri plate. If such
colonies are present, record them as positive on
your Laboratory Report chart and on the chart on
the chalkboard.
Fill out the chart on your Laboratory Report
with all the information from the chart on the
chalkboard.
Identify those persons that test positive. You
will be working backwards with the kind of in-
formation an epidemiologist has to work with
(contacts and infections). Eventually a pattern
will emerge that shows which person started the
epidemic.
Laboratory Report
Complete the Laboratory Report for this exercise
256
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XIV. Medical Microbiology
and Immunology
78. The Staphylococci:
Isolation and Identification
© The McGraw-H
Companies, 2001
The Staphylococci:
Isolation and Identification
78
Often in conjunction with streptococci, the staphylo-
cocci cause abscesses, boils, carbuncles, osteomyelitis,
and fatal septicemias. Collectively, the staphylococci
and streptococci are referred to as the pyogenic (pus-
forming) gram-positive cocci. Originally isolated from
pus in wounds, the staphylococci were subsequently
demonstrated to be normal inhabitants of the nasal
membranes, the hair follicles, the skin, and the per-
ineum of healthy individuals. The fact that 90% of hos-
pital personnel are carriers of staphylococci portends
serious epidemiological problems, especially since
most strains are penicillin-resistant.
The staphylococci are gram-positive spherical
bacteria that divide in more than one plane to form ir-
regular clusters of cells. They are listed in section 12,
volume 2, of Bergey's Manual of Systematic
Bacteriology. The genus Staphylococcus is grouped
with three other genera in family Micrococcaceae:
SECTION 12 GRAM-POSITIVE COCCI
Family I Micrococcaceae
Genus I Micrococcus
Genus II Stomatococcus
Genus III Planococcus
Genus IV Staphylococcus
Family II Deinococcaceae
Genus I Deinococcus
Genus II Streptococcus
Although the staphylococci make up a coherent
phylogenetic group, they have very little in common
with the streptococci except for their basic similar-
ities of being gram-positive, non- spore-forming
cocci. Note that Bergey's Manual puts these two
genera into separate families due to their inherent
differences.
Of the 19 species of staphylococci listed in
Bergey's Manual, the most important ones are S. au-
reus, S. epidermidis, and S. saprophyticus. The single
most significant characteristic that separates these
species is the ability or inability of these organisms to
coagulate plasma: only S. aureus has this ability; the
other two are coagulase-negative.
Although S. aureus has, historically, been con-
sidered to be the only significant pathogen of the
three, the others do cause infections. Some cere-
brospinal fluid infections (2), prosthetic joint infec-
Figure 78.1 Staphylococci
tions (3), and vascular graft infections (1) have been
shown to be due to coagulase-negative staphylo-
cocci. Numbers in parentheses designate references
at the end of this exercise.
Our concern in this exercise will pertain exclu-
sively to the differentiation of only three species of
staphylococci. If other species are encountered, the
student may wish to use the API Staph-Ident minia-
turized test strip system (Exercise 55).
In this experiment we will attempt to isolate
staphylococci from (1) the nose, (2) a fomite, and (3)
an "unknown-control." The unknown-control will be
a mixture containing staphylococci, streptococci, and
some other contaminants. If the nasal membranes and
fomite prove to be negative, the unknown-control will
yield positive results, providing all inoculations and
tests are performed correctly
Since S. aureus is by far the most significant
pathogen in this group, most of our concern here will
be with this organism. It is for this reason that the
characteristics of only this pathogen will be outlined
next.
Staphylococcus aureus cells are 0.8 to 1.0 |xm in
diameter and may occur singly, in pairs, or as clusters.
Colonies of S. aureus on trypticase soy agar or blood
agar are opaque, 1 to 3 mm in diameter, and yellow,
orange, or white. They are salt- tolerant, growing well
257
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XIV. Medical Microbiology
and Immunology
78. The Staphylococci:
Isolation and Identification
© The McGraw-H
Companies, 2001
Exercise 78 • The Staphylococci: Isolation and Identification
on media containing 10% sodium chloride. Virtually
all strains are coagulase-positive. Mannitol is fer-
mented aerobically to produce acid. Alpha toxin is
produced that causes a wide zone of clear (beta-type)
hemolysis on blood agar; in rabbits it causes local
necrosis and death.
The other two species lack alpha toxin (do not ex-
hibit hemolysis) and are coagulase-negative. Mannitol
is fermented to produce acid (no gas) by all strains of
S. aureus and most strains of S. saprophyticus. Table
78.1 lists the principal characteristics that differentiate
these three species of staphylococcus.
Table 78.1 Differentiation of three species of staphylococci
s.
aureus
s.
epider-
S. sapro-
midis
phyticus
Alpha toxin
+
—
—
Mannitol
+
—
(+)
(acid only)
Coagulase
+
—
—
Biotin for growth
—
+
NS
Novobiocin
S
S
R
Note: NS = not significant; S = sensitive; R = resistant;
(+) = mostly positive
To determine the incidence of carriers in our
classroom, as well as the incidence of the organism on
common fomites, we will follow the procedure illus-
trated in figure 78.2. Results of class findings will be
tabulated on the chalkboard so that all members of the
class can record data required on the Laboratory
Report. The characteristics we will look for in our iso-
lates will be (1) beta- type hemolysis (alpha toxin), (2)
mannitol fermentation, and (3) coagulase production.
Organisms found to be positive for these three char-
acteristics will be presumed to be S. aureus. Final con-
firmation will be made with additional tests. Proceed
as follows:
First Period
(Specimen Collection)
Note in figure 78.2 that swabs that have been applied
to the nasal membranes and fomites will be placed in
tubes of enrichment medium containing 10% NaCl
(m- staphylococcus broth). Since your unknown-
control will lack a swab, initial inoculations from
this culture will have to be done with a loop.
Materials:
1 tube containing numbered unknown-control
3 tubes of m- staphylococcus broth
2 sterile cotton swabs
1
2
3
4
5
Label the three tubes of ra-staphylococcus
broth NOSE, FOMITE, and the number of your
unknown-control .
Inoculate the appropriate tube of m-staphylo-
coccus broth with one or two loopfuls of your
unknown-control.
After moistening one of the swabs by immersing
partially into the "nose" tube of broth, swab the
nasal membrane just inside your nostril. A small
amount of moisture on the swab will enhance the
pickup of organisms. Place this swab into the
"nose" tube.
Swab the surface of a fomite with the other swab
that has been similarly moistened and deposit this
swab in the "fomite" tube.
The fomite you select may be a coin, drinking
glass, telephone mouthpiece, or any other item
that you might think of.
Incubate these tubes of broth for 4 to 24 hours at
37° C.
Second Period
(Primary Isolation Procedure)
Two kinds of media will be streaked for primary
isolation: mannitol salt agar and staphylococcus
medium 110.
Mannitol salt agar (MSA) contains mannitol,
7.5% sodium chloride, and phenol red indicator.
The NaCl inhibits organisms other than staphylo-
cocci. If the mannitol is fermented to produce acid,
the phenol red in the medium changes color from
red to yellow.
Staphylococcus medium 110 (SMI 10) also con-
tains NaCl and mannitol, but it lacks phenol red. Its
advantage over MSA is that it favors colony pigmen-
tation by different strains of S. aureus. Since this
medium lacks phenol red, no color change takes place
as mannitol is fermented.
Materials:
3 culture tubes from last period
2 Petri plates of MSA
2 Petri plates of SMI 10
1
2
3
Label the bottoms of the MSA and SMI 10 plates
as shown in figure 78.2. Note that to minimize the
number of plates required, it will be necessary to
make half-plate inoculations for the nose and
fomite. The unknown-control will be inoculated
on separate plates.
Quadrant streak the MSA and SMI 10 plates with
the unknown control.
Inoculate a portion of the nose side of each plate
with the swab from the nose tube; then, with a
258
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XIV. Medical Microbiology
and Immunology
78. The Staphylococci:
Isolation and Identification
© The McGraw-H
Companies, 2001
The Staphylococci: Isolation and Identification • Exercise 78
UNKNOWN-CONTROL
FROM NOSE
FROM FOMITE
"j*k
■iwi
Tubes of selective media (m-staph-
ylococcus broth) are inoculated and
incubated at 37° C for 24 hours.
\L
NOSE
Plates of mannitol salt agar (MSA)
and staphylococcus medium 110
(SM1 10) are streaked from broth.
FOMITE
37° C, 24-36 hr
Blood Agar
Typical well-isolated colonies are
selected from either medium for
inoculating blood agar and plasma,
as well as for making gram-stained slides.
S. aureus colonies cause
MSA medium to turn yellow.
37° C, 24-36 hr
GRAM STAIN
Blood Agar
S. aureus colonies tend
to be pigmented on
SM110 medium.
GRAM STAIN
Blood Agar
COAGULASE TESTS
Tubes are checked
every 30 min for up to
4 hr for evidence of
coagulation of plasma.
37° C, 18-24 hr
v^
COAGULASE TEST
¥
Negative tubes
are checked again
after 24 hours.
37° C, 18-24 hr
Second Check
Presence or absence of beta -type hemolysis is looked for
on blood agar plates to confirm existence of S. aureus.
I
I
R
S
T
L
A
B
ir
T
H
I
R
D
L
A
B
if
F
O
U
R
T
H
L
A
B
}[
Figure 78.2 Procedure for presumptive identification of staphylococci
259
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XIV. Medical Microbiology
and Immunology
78. The Staphylococci:
Isolation and Identification
© The McGraw-H
Companies, 2001
Exercise 78 • The Staphylococci: Isolation and Identification
4.
5.
sterile loop, streak out the organisms on the re-
mainder of the agar on that half of each plate.
The swabbed areas will provide massive growth;
the streaked-out areas should yield good colony
isolation.
Repeat step 3 to inoculate the other half of each
agar plate with the swab from the fomite tube.
Incubate the plates aerobically at 37° C for 24 to
36 hours.
2
3
Select staphylococcus-like colonies from the
MSA and SMI 10 plates from the nose and fomites
for streaking out on another blood agar plate. Use
half-plate streaking methods, if necessary.
Incubate the blood agar plates at 37° C for 18 to
24 hours. Don 't leave plates in incubator longer
than 24 hours. Overincubation will cause blood
degeneration.
Third Period
(Plate Evaluations and Coagulase/DNase Tests)
During this period we will perform the following
tasks: (1) evaluate the plates from the previous pe-
riod, (2) inoculate blood agar plates, (3) make gram-
stained slides, and (4) perform coagulase and/or
DNase tests on organisms from selected colonies.
Proceed as follows:
Materials:
MSA and SMI 10 plates from previous period
2 blood agar plates
serological tubes containing 0.5 ml of 1 :4 saline
dilution of rabbit or human plasma (one tube
for each isolate)
Petri plates of DNase agar
gram- staining kit
Evaluation of Plates
1
2
3
Examine the mannitol salt agar plates. Has the
phenol red in the medium surrounding any of the
colonies turned yellow?
If this color change exists, it can be pre-
sumed that you have isolated a strain of S. au-
reus. Record your results on the Laboratory
Report and chalkboard. (Your instructor may
wish to substitute a copy of the chart from the
Laboratory Report to be filled out at the demon-
stration table.)
Examine the plates of SMI 10. The presence of
growth here indicates that the organisms are salt-
tolerant. Note color of the colonies (white, yel-
low, or orange) .
Record your observations of these plates on the
Laboratory Report and chalkboard.
Blood Agar Inoculations
1. Label the bottom of one blood agar plate with
your unknown-control number, and streak out the
organisms from a staph-like colony.
Coagulase Tests
The fact that 97% of the strains of S. aureus have
proven to be coagulase-positive and that the other two
species are always coagulase-negative makes the co-
agulase test an excellent definitive test for confirming
identification of S. aureus.
The procedure is simple. It involves inoculating a
small tube of plasma with several loopfuls of the or-
ganism and incubating it in a 37° C water bath for sev-
eral hours. If the plasma coagulates, the organism is
coagulase-positive. Coagulation may occur in 30
minutes or several hours later. Any degree of coagula-
tion, from a loose clot suspended in plasma to a solid
immovable clot, is considered to be a positive result,
even if it takes 24 hours to occur.
It should be emphasized that this test is valid
only for gram -positive, staphylococcus-like bacte-
ria, because some gram-negative rods, such as
Pseudomonas, can cause a false-positive reaction.
The mechanism of clotting in such organisms is not
due to coagulase. Proceed as follows:
1
2
3
4
5
Label the plasma tubes NOSE, FOMITE, or UN-
KNOWN, depending on which of your plates
have staph-like colonies.
With a wire loop, inoculate the appropriate tube
of plasma with organisms from one or more
colonies on SMI 10 or MSA. Use several loop-
fuls. Success is more rapid with a heavy inocula-
tion. If positive colonies are present on both nose
and fomite sides, be sure to inoculate a separate
tube for each side.
Place the tubes in a 37° C water bath.
Check for solidification of the plasma every 30
minutes for the remainder of the period. Note in
figure 78.3 that solidification may be complete, as
in the lower tube, or show up as a semisolid ball,
as seen in the middle tube.
Any cultures that are negative at the end of
the period will be left in the water bath. At 24
hours your instructor will remove them from the
water bath and place them in the refrigerator, so
that you can evaluate them in the next laboratory
period.
Record your results on the Laboratory Report.
260
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XIV. Medical Microbiology
and Immunology
78. The Staphylococci:
Isolation and Identification
© The McGraw-H
Companies, 2001
The Staphylococci: Isolation and Identification • Exercise 78
COAGULASE-NEGATIVE
COAGULASE-POSITIVE
COAGULASE-POSITIVE
Figure 78.3 Coagulase test results: one negative and
two positive tests
Materials:
coagulase tubes from previous tests
blood agar plates from previous period
DNase test agar plates from previous period
0.1NHC1
1
2
3
4
Examine any coagulase tubes that were carried
over from the last laboratory period that were
negative at the end of that period. Record your re-
sults on the Laboratory Report.
Examine the colonies on your blood agar plates.
Look for clear (beta-type) hemolysis around the
colonies. The presence of alpha toxin is a defini-
tive characteristic of S. aureus. Record your re-
sults on the Laboratory Report.
Look for zones of clearing near the streaks on the
DNase agar plate. If none is seen, develop by
flooding the plate with 0.1N HC1. The acid will
render the hydrolyzed areas somewhat opaque.
Record your results on the chart on the chalk-
board or chart on demonstration table. If an
instructor- supplied tabulation chart is used, the
instructor will have copies made of it to be sup-
plied to each student.
DNase Test
The fact that coagulase-positive bacteria are also able
to hydrolyze DNA makes the DNase test a reliable
means of confirming S. aureus identification. The fol-
lowing procedure can be used to determine if a staph-
like organism can hydrolyze DNA.
1
2
Heavily streak the organism on a plate of DNase
test agar. One plate can be used for several test
cultures by making short streaks about 1 inch
long.
Incubate for 18-24 hours at 35° C.
Gram-Stained Slides
While your tubes of plasma are incubating in the wa-
ter bath, prepare gram-stained slides from the same
colonies that were used for the blood agar plates and
coagulase tests.
Examine the slides under oil immersion lens and
draw the organisms in the appropriate areas on the
Laboratory Report.
Fourth Period
(Confirmation)
During this period we will make final assessment of
all tests and perform any other confirmatory tests that
might be available to us.
Further Testing
In addition to using the API Staph-Ident miniaturized
test strip system (Exercise 55) to confirm your identi-
fication of staphylococci, you may wish to use the la-
tex agglutination slide test described in Exercise 83.
Your instructor will inform you as to the availability
of these materials and the desirability of proceeding
further.
Laboratory Report
After recording your results on the chalkboard (or on
chart on demonstration table), complete the chart on
the Laboratory Report and answer all the questions.
1
2
3
Literature Cited
Liekweg, W. G., Jr., and L. T. Greenfield. 1977.
Vascular prosthetic infection: Collected experi-
ence and results of treatment. Surgery 81:
355-400.
Schoenbaum, S. C, P. Gardner, and J. Shillito.
1975. Infections in cerebrospinal shunts:
Epidemiology, clinical manifestations, and ther-
apy. /. Infect Dis. 131:543-52.
Wilson, P. D., Jr., E. A. Salvati, P. Aglietti, and
L. J. Kutner. 1973. The problem of infection in
endoprosthetic surgery of the hip joint. Clin.
Orthop. Relat. Res. 96:213-21.
261
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XIV. Medical Microbiology
and Immunology
79. The Streptococci:
Isolation and Identification
© The McGraw-H
Companies, 2001
79
The Streptococci:
Isolation and Identification
The streptococci differ from the staphylococci in that
they are arranged primarily in chains rather than in
clusters. In addition to causing many mixed infections
with staphylococci, the streptococci can also, sepa-
rately, cause diseases such as pneumonia, meningitis,
endocarditis, pharyngitis, erysipelas, and glomeru-
lonephritis.
Several species of streptococci are normal inhab-
itants of the pharynx. They can also be isolated from
surfaces of the teeth, the saliva, skin, colon, rectum,
and vagina.
The streptococci of greatest medical significance
are S. pyogenes, S. agalactiae, and S. pneumoniae. Of
lesser importance are S. faecalis, S. faecium, and S.
bovis. Appendix E describes in greater detail the char-
acteristics and significance of these and other strepto-
coccal species.
The purpose of this exercise is twofold: (1) to
learn about standard procedures for isolating strepto-
cocci from the pharynx and (2) to learn how to differ-
entiate between the most significant medically impor-
tant streptococci.
Figure 79.2 illustrates the overall procedure to be
followed in the pursuit of the above two goals. Note
that blood agar is used to separate the streptococci
into two groups on the basis of the type of hemolysis
they produce on blood agar. Those organisms that
produce alpha hemolysis on blood agar can be differ-
entiated by four tests. Those that produce beta- type
hemolysis can be differentiated with the CAMP test
and three other tests. The procedure outlined here is,
primarily, designed to achieve presumptive identifica-
tion of seven groups of streptococci. A few extra tests
are usually required to confirm identification.
To broaden the application of these tests you may
be given two or three unknown cultures of strepto-
cocci to be identified along with the pharyngeal iso-
lates. If unknowns are to be used, they will not be is-
sued until physiological media are to be inoculated.
First Period
(Making a Streak-Stab Agar Plate)
During this period a plate of blood agar is swabbed
and streaked in a special way to determine the type of
hemolytic bacteria that are present in the pharynx.
Figure 79.1 Streptococci
Before making such a streak plate, however, clini-
cians prefer to use a tube of enrichment broth (TSB)
or a selective medium of TSB with a little crystal vi-
olet added to it (TSBCV). Media of this type are usu-
ally incubated at 37° C for 24 hours. This is particu-
larly useful if the number of organisms might be low
or if the swab cannot be applied to blood agar imme-
diately. Although this enrichment/ selective step has
been omitted here, it should be understood that the
procedure is routine.
Since swabbing one's own throat properly can be
difficult, it will be necessary for you to work with
your laboratory partner to swab each other's throats.
Once your throat has been swabbed, you will proceed
to use the swab to streak and stab your own agar plate
according to a special procedure shown in figure 79.3.
Materials:
1 tongue depressor
1 sterile cotton swab
inoculating loop
1 blood agar plate
1 . With the subject's head tilted back and the tongue
held down with the tongue depressor, rub the back
262
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XIV. Medical Microbiology
and Immunology
79. The Streptococci:
Isolation and Identification
© The McGraw-H
Companies, 2001
T
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A blood agar plate is streaked and
stabbed directly from the pharynx or
from enrichment/selective media.
Incubate at 37° C for 24 hours
'3*a
\^
Colony with alpha hemolysis is sub-
cultured by inoculating a tube of tryp-
ticase soy broth.
Tubes of TSB are incubated at 37° C for 24 hours,
Colony with beta hemolysis is subcul-
tured by inoculating a tube of trypticase
soy broth.
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6.5% NaCI
HIPPURATE
HYDROLYSIS
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All media incubated at 37° C for 24 hours
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OPTOCHIN SENSITIVITY: Pneumococci are sen-
sitive and viridans organisms are resistant to these
disks.
BILE SOLUBILITY: TSB culture is used for this test.
Pneumococci are always soluble in bile.
BILE ESCULIN HYDROLYSIS: All group D strep-
tococci are positive for this test.
SALT TOLERANCE: Group D enterococci are salt
tolerant. Other group D organisms are not.
CAMP TEST: If positive, the organism is very likely
S. agalactiae.
BACITRACIN SENSITIVITY: If sensitivity is present,
organism is probably S. pyogenes.
SXT SENSITIVITY: This test, together with baci-
tracin sensitivity test, is used for identification of
group C streptococci.
HIPPURATE HYDROLYSIS: If sodium hippurate is
hydrolyzed, organism is S. agalactiae.
Figure 79.2 Media inoculations for the presumptive identification of streptococci
263
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XIV. Medical Microbiology
and Immunology
79. The Streptococci:
Isolation and Identification
© The McGraw-H
Companies, 2001
Exercise 79 • The Streptococci: Isolation and Identification
surface of the pharynx up and down with the ster-
ile swab.
Also, look for white patches in the tonsillar
area. Avoid touching the cheeks and tongue.
2. Since streptococcal hemolysis is most accu-
rately analyzed when the colonies develop
anaerobically beneath the surface of the agar, it
will be necessary to use a streak-stab technique
as shown in figure 79.3. The essential steps are
as follows:
• Roll the swab over an area approximating one-
fifth of the surface. The entire surface of the
swab should contact the agar.
• With a wire loop, streak out three areas as
shown to thin out the organisms.
• Stab the loop into the agar to the bottom of
the plate at an angle perpendicular to the sur-
face to make a clean cut without ragged
edges.
• Be sure to make one set of stabs in an un-
streaked area so that streptococcal hemolysis
will be easier to interpret with a microscope.
CAUTION
Dispose of swabs and tongue depressors in beaker of
disinfectant.
Second Period
(Analysis and Subculturing)
During this period, two things must be accomplished:
first, the type of hemolysis must be correctly determined
and, second, well-isolated colonies must be selected for
making subcultures. The importance of proper subcul-
turing cannot be overemphasized: without a pure cul-
ture, future tests are certain to fail. Proceed as follows:
Materials:
blood agar plate from previous period
tubes of TSB (one for each different type of
colony)
dissecting microscope
1
2
3
3. Incubate the plate aerobically at 37° C for 24
hours. Do not incubate longer than 24 hours.
Look for isolated colonies that have alpha or beta
hemolysis surrounding them. Streptococcal
colonies are characteristically very small.
Do any of the stabs appear to exhibit hemoly-
sis? Examine these hemolytic zones near the
stabs under 60 X magnification with a dissect-
ing microscope.
Consult figure 79.4 to analyze the type of hemol-
ysis. Note that the illustrations on the left side in-
dicate what the colonies would look like if they
were submerged under a layer of blood agar (two-
layer pour plate). The illustrations on the right in-
dicate the nature of hemolysis around stabs on
streak-stab plates. Although this illustration is
very diagrammatic, it reveals the microscopic dif-
ferences between three kinds of hemolysis: alpha,
alpha-prime, and beta.
Swab is rolled over approx-
imately 1/5 area of plate.
Organisms are thinned out
by streaking from swabbed
area.
Thinning out of organisms
is completed with inocu-
lating loop.
Loop is stabbed several
times perpendicular to sur
face to bottom of plate.
Inoculating loop is used
to further thin out the
organisms.
Figure 79.3 Streak-stab procedure for blood agar inoculations
264
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XIV. Medical Microbiology
and Immunology
79. The Streptococci:
Isolation and Identification
© The McGraw-H
Companies, 2001
The Streptococci: Isolation and Identification • Exercise 79
4.
5
6
7
Only those stabs that are completely free of
red blood cells in the hemolytic area are consid-
ered to be beta hemolytic. The chance of isolat-
ing a colony of this type from your own throat is
very slim, for the beta hemolytic streptococci are
the most serious pathogens.
If some red blood cells are seen dispersed
throughout the hemolytic zone, the organism is
classified as alpha-prime hemolytic. Viridans
streptococci often fall in this category.
Record your observations on the Laboratory Report.
Select well-isolated colonies that exhibit hemoly-
sis (alpha, beta, or both) for inoculating tubes of
TSB. Be sure to label the tubes ALPHA or BETA.
Whether or not the organism is alpha or beta is
crucial in identification.
Since the chances of isolating beta hemolytic
streptococci from the pharynx are usually quite
slim, notify your instructor if you think you have
isolated one.
Incubate the tubes at 37° C for 24 hours.
Important: At some time prior to the next labo-
ratory session, review the material in Appendix E
that pertains to this exercise.
Third Period
(Inoculations for Physiological Tests)
Presumptive identification of the various groups of
streptococci is based on seven or eight physiological
tests. Table 79.1 on page 267 reveals how they perform
on these tests. Note that groups A, B, and C are all beta
hemolytic; a few enterococci are also beta hemolytic.
The remainder are all alpha hemolytic or nonhemolytic.
Since each of the physiological tests is specific
for differentiating only two or three groups, it is not
desirable to do all the tests on all unknowns. For econ-
omy and preciseness, only four tests that are men-
tioned for the third period in figure 79.2 should be
performed on an isolate or unknown.
Before any inoculations are made, however, it is de-
sirable to do a purity check on each TSB culture from
the previous period. To accomplish this it will be neces-
sary to make a gram-stained slide of each of the cultures.
If unknowns are to be issued, they will be given
to you at this time. They will be tested along with your
pharyngeal isolates. The only information that will be
given to you about each unknown is its hemolytic type
so that you will be able to determine what physiologi-
cal tests to perform on each one. Proceed as follows:
Gram-Stained Slides (Purity Check)
Materials:
TSB cultures from previous period
gram- staining kit
ALP
ME
POUR PLATE
STREAK-STAB
Figure 79.4 Comparison of hemolysis types as seen on
pour plates and streak-stab plates
1
2
Make a gram- stained slide from each of the pha-
ryngeal isolates and examine them under oil im-
mersion lens. Do they appear to be pure cultures?
Draw the organisms in the appropriate circles on
the Laboratory Report.
Beta-Type Inoculations
Use the following procedure to perform tests on each
isolate that has beta- type hemolysis:
Materials:
for each isolate:
1 blood agar plate
1 tube of sodium hippurate broth
1 bacitracin differential disk
1 SXT sensitivity disk
1 broth culture of S. aureus
dispenser or forceps for transferring disks
1 . Label a blood agar plate and a tube of sodium hip-
purate broth with proper identification informa-
tion of each isolate and unknown to be tested.
2. Follow the procedure outlined in figure 79.5 to in-
oculate each blood agar plate with the isolate (or
unknown) and S. aureus.
265
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XIV. Medical Microbiology
and Immunology
79. The Streptococci:
Isolation and Identification
© The McGraw-H
Companies, 2001
Exercise 79 • The Streptococci: Isolation and Identification
3
4
5
Note that a streak of the unknown is brought
down perpendicular to the S. aureus streak, keep-
ing the two organisms about 1 cm apart.
With forceps or dispenser, place one bacitracin
differential disk and one SXT disk on the heavily
streaked area at points shown in figure 79.5. Press
down on each disk slightly.
Inoculate one tube of sodium hippurate broth for
each isolate or unknown.
Incubate the blood agar plates at 37° C, aerobi-
cally, for 24 hours, and the hippurate broth tubes
at 35° C, aerobically, for 24 hours. If the hippurate
broths prove to be negative or weakly positive at
24 hours, they should be given more time to see if
they change.
Alpha-Type Inoculations
As shown in figure 79.2, four inoculations will be made
for each isolate or unknown that is alpha hemolytic.
Materials:
1 blood agar plate (for up to 4 unknowns)
1 6.5% sodium chloride broth
1 trypticase soy broth (TSB)
1 bile esculin (BE) slant
1 optochin (Taxo P) disk
candle jar setup or C0 2 incubator
1 . Mark the bottom of a blood agar plate to divide it
into halves, thirds, or quarters, depending on the
number of alpha hemolytic organisms to be
tested. Label each space with the code number of
each test organism.
2. Completely streak over each area of the blood
agar plate with the appropriate test organism, and
place one optochin (Taxo P) disk in the center of
each area. Press down slightly on each disk to se-
cure it to the medium.
3. Inoculate one tube each of TSB, BE, and 6.5%
NaCl broth with each test organism.
4. Incubate all media at 35°-37° C as follows:
Blood agar plates: 24 hours in a candle jar
6.5% NaCl broths: 24, 48, and 72 hours
Bile esculin slants: 48 hours
Trypticase soy broths: 24 hours
Note: While the blood agar plates should be in-
cubated in a candle jar or C0 2 incubator, the re-
maining cultures can be incubated aerobically.
Fourth Period
(Evaluation of Physiological Tests)
Once all of the inoculated media have been incubated
for 24 hours, you are ready to examine the plates and
tubes and add test reagents to some of the cultures.
Some of the tests will also have to be checked at 48
and 72 hours.
After you have assembled all the plates and tubes
from the last period, examine the blood agar plates
first that were double- streaked with the unknowns and
S. aureus. Note that the second, third, and fourth tests
listed in table 79.1 can be read from these plates.
Proceed as follows:
CAMP Reaction
If you have an unknown that produces an enlarged
arrowhead-shaped hemolytic zone at the juncture
where the unknown meets the S. aureus streak, as seen
Bacitracin and SXT differential
disks are placed as shown in
area streaked by the unknown.
Unknown is heavily streaked out
over 40% of the area and brought
straight downward in a single
line.
A loopful of S, aureus is streaked
perpendicular to unknown streak.
A gap of one centimeter should
separate the two streaks .
Figure 79 Blood agar inoculation technique for the CAMP, bacitracin, and SXT tests
266
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XIV. Medical Microbiology
and Immunology
79. The Streptococci:
Isolation and Identification
© The McGraw-H
Companies, 2001
The Streptococci: Isolation and Identification • Exercise 79
in figure 79.6, the organism is S. agalactiae. This phe-
nomenon is due to what is called the CAMP factor. The
only problem that can arise from this test is that if the
plate is incubated anaerobically, a positive CAMP re-
action can occur on S. pyogenes inoculated plates.
Record the CAMP reactions for each of your iso-
lates or unknowns on the Laboratory Report.
Bacitracin Susceptibility
Any size zone of inhibition seen around the bacitracin
disks should be considered to be a positive test result.
Note in table 79.1 that S. pyogenes is positive for this
characteristic.
This test has two limitations: (1) the disks must be
of the differential type, not sensitivity type, and (2) the
test should not be applied to alpha hemolytic strepto-
cocci. Reasons: Sensitivity disks have too high a con-
centration of the antibiotic, and many alpha hemolytic
streptococci are sensitive to these disks.
Record the results of this test on table under D of
your Laboratory Report.
SXT Sensitivity Test
The disks used in this test contain 1.25 mg of
trimethoprim and 27.75 mg of sulfamethoxazole
(SXT). The purpose of this test is to distinguish
Table 79.1 Physiological Tests for Streptococcal Differentiation
GROUP / <&/ &/&*? /& / *<$/ * <b- / cf / & /
Group A
S. pyogenes
beta
+
1 '
R
^"^^
^"^^
*^~ ^~
i^_^_
Group B
S. agalactiae
beta
_*
+
R
—
+
—
—
Group C
S. equi
S. equisimilis
S. zooepidemicus
beta
_*
S
•ff-ff
Group D
(enterococci)
S. faecalis
S. faecium
etc.
alpha
beta
none
R
+
+
Group D**
(nonenterococci)
S. bovis
etc.
alpha
none
R/S
+
Viridans
S. mitis
S. salivarius
S. mutans
etc.
alpha
none
*
*
S
Pneumococci
S. pneumoniae
alpha
+
—
—
—
+
+
Note: R = resistant; S = sensitive; blank = not significant.
"Exceptions occur occasionally.
**See comments on pp. 457 and 458 concerning correct genus.
267
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XIV. Medical Microbiology
and Immunology
79. The Streptococci:
Isolation and Identification
© The McGraw-H
Companies, 2001
Exercise 79 • The Streptococci: Isolation and Identification
Figure 79.6 Note positive SXT disk on right, negative
bacitracin disk on left, and positive CAMP reaction (ar-
rowhead). Organism: S. agalactiae.
groups A and B from other beta hemolytic strepto-
cocci. Note in table 79.1 that both groups A and B are
uniformly resistant to SXT.
If a beta hemolytic streptococcus proves to be
bacitracin-resistant and SXT-susceptible, it is classi-
fied as being a non-group-A or -B beta hemolytic
streptococcus. This means that the organism is prob-
ably a species within group C. Keep in mind that an
occasional group A streptococcal strain is susceptible
to both bacitracin and SXT disks. One must always re-
member that exceptions to most tests do occur; that is
why this identification procedure leads us only to pre-
sumptive conclusions.
Record any zone of inhibition (resistance) as pos-
itive for this test.
Hippurate Hydrolysis
Note in table 79.1 that hippurate hydrolysis and the
CAMP test are grouped together as positive tests for
S. agalactiae. If an organism is positive for both tests,
or either one, one can assume with almost 100% cer-
tainty that the organism is S. agalactiae.
Proceed as follows to determine which of your
isolates are able to hydrolyze sodium hippurate:
Materials:
serological test tubes
serological pipettes (1 ml size)
ferric chloride reagent
centrifuge
1. Centrifuge the culture for 3 to 5 minutes.
2. Pipette 0.2 ml of the supernatant and 0. 8 ml of fer-
ric chloride reagent into an empty serological test
tube. Mix well.
3
4
Look for a heavy precipitate to form. If the pre-
cipitate forms and persists for 10 minutes or
longer, the test is positive. If the culture proves to
be weakly positive, incubate the culture for an-
other 24 hours and repeat the test.
Record your results on the Laboratory Report.
Bile Esculin Hydrolysis
This is the best physiological test that we have for the
identification of group D streptococci. Both entero-
coccal and nonenterococcal species of group D are
able to hydrolyze esculin in the agar slant, causing the
slant to blacken.
A positive BE test tells us that we have a group D
streptococcus; differentiation of the two types of group
D streptococci depends on the salt-tolerance test.
Examine the BE agar slants, looking for black-
ening of the slant, as illustrated in figure 79.7. If
less than half of the slant is blackened, or if no
blackening occurs within 24 to 48 hours, the test is
negative.
Figure 79.7 Positive bile esculin hydrolysis on left;
negative on right
Salt Tolerance (6.5% NaCl)
All enterococci of group D produce heavy growth in
6.5% NaCl broth. As indicated in table 79.1, none of
the nonenterococci, group D, grow in this medium.
This test, then, provides us with a good method for
differentiating the two types of group D streptococci.
A positive result shows up as turbidity within 72
hours. A color change of purple to yellow may also
be present. If the tube is negative at 24 hours, incubate
it and check it again at 48 and 72 hours. If the organ-
268
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XIV. Medical Microbiology
and Immunology
79. The Streptococci:
Isolation and Identification
© The McGraw-H
Companies, 2001
The Streptococci: Isolation and Identification • Exercise 79
ism is salt-tolerant and BE-positive, it is considered to
be an enterococcus. Parenthetically, it should be
added here that approximately 80% of group B strep-
tococci will grow in this medium.
Optochin Susceptibility
Optochin susceptibility is used for differentiation of
the alpha hemolytic viridans streptococci from the
pneumococci. The pneumococci are sensitive to these
disks; the viridans organisms are resistant.
Materials:
blood agar plates with optochin disks
plastic metric ruler
1
2
Measure the diameters of zones of inhibition that
surround each disk, evaluating whether the zones
are large enough to be considered positive. The
standards are as follows:
• For 6 mm diameter disks, the zone must be at
least 14 mm diameter to be considered positive.
• For 10 mm diameter disks, the zone must be at
least 16 mm diameter to be considered positive.
Record your results on the Laboratory Report.
Bile Solubility
If an alpha hemolytic streptococcal organism is solu-
ble in bile and positive on the optochin test, presump-
tive evidence indicates that the isolate is S. pneumo-
niae. Perform the bile solubility test on each of your
alpha hemolytic isolates as follows:
Materials:
2 empty serological tubes (per test)
dropping bottle of phenol red indicator
1
2
3
4
5
dropping bottle of 0.05N NaOH
TSB culture of unknown
2% bile solution (sodium desoxycholate)
bottle of normal saline solution
2 serological pipettes (1 ml size)
water bath (37° C)
Mark one empty serological tube BILE and the
other SALINE. Into their respective tubes, pipette
0.5 ml of 2% bile and 0.5 ml of saline.
Shake the TSB unknown culture to suspend the
organisms and pipette 0.5 ml of the culture into
each tube.
Add one or two drops of phenol red indicator to
each tube and adjust the pH to 7.0 by adding drops
of 0.05N NaOH.
Place both tubes in a 37° C water bath and exam-
ine periodically for 2 hours. If the turbidity clears
in the bile tube, it indicates that the cells have dis-
integrated and the organism is S. pneumoniae.
Compare the tubes side by side.
Record your results on the Laboratory Report.
Final Confirmation
All the laboratory procedures performed so far lead us
to presumptive identification. To confirm these con-
clusions it is necessary to perform serological tests on
each of the unknowns. If commercial kits are avail-
able for such tests, they should be used to complete
the identification procedures.
Laboratory Report
Complete the Laboratory Report for this exercise
269
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XIV. Medical Microbiology
and Immunology
80. Gram-Negative
Intestinal Pathogens
© The McGraw-H
Companies, 2001
80
Gram-Negative Intestinal Pathogens
The enteric pathogens of prime medical concern are
the salmonella and shigella. They cause enteric
fevers, food poisoning, and bacillary dysentery.
Salmonella typhi, which causes typhoid fever, is by
far the most significant pathogen of the salmonella
group. In addition to the typhoid organism, there are
10 other distinct salmonella species and over 2,200
serotypes. The shigella, which are the prime causes of
human dysentery, comprise four species and many
serotypes. Serotypes within genera are organisms of
similar biochemical characteristics that can most eas-
ily be differentiated by serological typing.
Routine testing for the presence of these
pathogens is a function of public health laboratories at
various governmental levels. The isolation of these
pathogenic enterics from feces is complicated by the
fact that the colon contains a diverse population of
bacteria. Species of such genera as Escherichia,
Proteus, Enterobacter, Pseudomonas, and Clos-
tridium exist in large numbers: hence it is necessary to
use media that are differential and selective to favor
the growth of the pathogens.
Figure 80.1 is a separation outline that is the basis
for the series of tests that are used to demonstrate the
presence of salmonella or shigella in a patient's blood,
urine, or feces. Note that lactose fermentation sepa-
rates the salmonella and shigella from most of the
other Enterobacteriaceae. Final differentiation of the
two enteric pathogens from Proteus relies on motility,
hydrogen sulfide production, and urea hydrolysis.
The differentiation information of the positive lactose
fermenters on the left side of the separation outline is
provided here mainly for comparative references that
can be used for the identification of other unknown
enterics.
The procedural diagram in figure 80.2 on the op-
posite page reveals how we will apply these facts in
the identification of an unknown salmonella or
shigella. The entire process will involve four labora-
tory periods.
In this experiment you will be given a mixed cul-
ture containing a coliform, Proteus, and a salmonella
or shigella. The pathogens will be of the less danger-
ous types, but their presence will, naturally, demand
utmost caution in handling. Your problem will be to
isolate the pathogen from the mixed culture and make
a genus identification. There are five steps that are
used to prove the presence of these pathogens in a
Lactose
Lactose +
Lactose
ndole+
Indole
Glucose+
Glucose
C it rate +
Citrate
Urea +
Urea
Motile
Citrobacter
Escherichia
Klebsiella
Enterobacter
Nonmotile
H 2 S-
Pseudomonas
Alcaligenes
Shigella
Urea +
I
Proteus
Providencia
Morganella
Urea
Salmonella
Figure 80.1 Separation outline of Enterobacteriaceae
270
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XIV. Medical Microbiology
and Immunology
80. Gram-Negative
Intestinal Pathogens
© The McGraw-H
Companies, 2001
Gram -Negative Intestinal Pathogens • Exercise 80
37° C
24-48 hr
SELECTIVE MEDIUM
(such as MacConkey, HE,
or XLD agar)
ENRICHMENT
Tkv-
Slants that show glucose fermenta-
tion are selected for subculturing.
These tubes have yellow butts with
red slants. If Kligler iron agar slants
have a black precipitate, H 2 S is
produced.
RF..<
7,t '■ •' ' •
-t.
i<>i:-/ ■■■!
>•.:•
37° C
1 8-24 hr
NO
FERMENTATION
GLUCOSE
ONLY
GLUCOSE
AND LACTOSE
Slants of RDS or Kligler iron agar
are streaked and stabbed from typical
Salmonella and Shigella colonies
Tubes of urea broth and SIM medium
are inoculated from each tube that
exhibits glucose fermentation.
SIM medium is stabbed to 2/3 of
depth of medium. Both media are
incubated at 37° C for 1 8 to 24 hours.
Vj
UREA
BROTH
\^
SIM
MEDIUM
GRAM - STAINED SLIDE is made to
see if culture is pure. Serotyping is
generally necessary.
Figure 80.2 Isolation and presumptive identification of Salmonella and Shigella
271
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XIV. Medical Microbiology
and Immunology
80. Gram-Negative
Intestinal Pathogens
© The McGraw-H
Companies, 2001
Exercise 80 • Gram-Negative Intestinal Pathogens
stool sample: (1) enrichment, (2) isolation, (3) fer-
mentation tests, (4) final physiological tests, and
(5) serotyping.
Enrichment
There are two enrichment media that are most fre-
quently used to inhibit the nonpathogens and favor the
growth of pathogenic enterics. They are selenite F and
gram- negative (GN) broths. While most salmonella
grow unrestricted in these two media, some of the
shigella are inhibited to some extent in selenite F
broth; thus, for shigella isolation, GN broth is pre-
ferred. In many cases, stool samples are plated di-
rectly on isolation media.
In actual practice, 1 to 5 grams of feces are placed
in 10 ml of enrichment broth. In addition, plates of var-
ious kinds of selective media are inoculated directly.
The broths are usually incubated for 4 to 6 hours.
Since we are not using stool samples in this exer-
cise, the enrichment procedure is omitted. Instead,
you will streak the isolation media directly from the
unknown broth.
First Period
(Isolation)
There are several excellent selective differential me-
dia that have been developed for the isolation of these
pathogens. Various inhibiting agents such as brilliant
green, bismuth sulfite, sodium desoxycholate, and
sodium citrate are included in them. For Salmonella
typhi, bismuth sulfite agar appears to be the best
medium. Colonies of S. typhi on this medium appear
black due to the reduction of sulfite to sulfide.
Other widely used media are MacConkey agar,
Hektoen Enteric agar (HE), and Xylose Lysine
Desoxycholate (XLD) agar. These media may contain
bile salts and/or sodium desoxycholate to inhibit
gram-positive bacteria. To inhibit coliforms and other
nonenterics, they may contain a citrate. All of them
contain lactose and a dye so that if an organism is a
lactose fermenter, its colony will take on a color char-
acteristic of the dye present.
Since the enrichment procedure is being omitted
here, you will be issued an unknown broth culture
with a pathogenic enteric. Your instructor will indi-
cate which selective media will be used. Proceed as
follows to inoculate the selective media with your un-
known mixture:
Materials:
unknown culture (mixture of a coliform,
Proteus, and a salmonella or shigella)
1 or more Petri plates of different selective
media: MacConkey, Hektoen Enteric (HE),
or Xylose Lysine Desoxycholate (XLD) agar
1. Label each plate with your name and unknown
number.
2. With a loop, streak each plate with your unknown
in a manner that will produce good isolation.
3. Incubate the plates at 37° C for 24 to 48 hours.
Second Period
(Fermentation Tests)
As stated above, the fermentation characteristic that
separates the SS pathogens from the coliforms is their
inability to ferment lactose. Once we have isolated
colonies on differential media that look like salmo-
nella or shigella, the next step is to determine whether
the isolates can ferment lactose. All media for this
purpose contain at least two sugars, glucose and lac-
tose. Some contain a third sugar, sucrose. They also
contain phenol red to indicate when fermentation oc-
curs. Russell Double Sugar (RDS) agar is one of the
simpler media that works well. Kligler iron agar may
also be used. It is similar to RDS with the addition of
iron salts for detection of H 2 S. Your instructor will in-
dicate which one will be used.
Proceed as follows to inoculate three slants from
colonies on the selective media that look like either
salmonella or shigella. The reason for using three
slants is that the you may have difficulty distinguish-
ing Proteus from the SS pathogens. By inoculating
three tubes from different colonies, you will be in-
creasing your chances of success.
Materials:
3 agar slants (RDS or Kligler iron)
streak plates from first period
1. Label the three slants with your name and the
number of your unknown.
2. Look for isolated colonies that look like salmo-
nella or shigella organisms. The characteristics to
look for on each medium are as follows:
• MacConkey agar — Salmonella, Shigella and
other non-lactose-fermenting species produce
smooth, colorless colonies. Coliforms that fer-
ment lactose produce reddish, mucoid, or
dark-centered colonies.
• Hektoen Enteric (HE) agar — Salmonella
and Shigella colonies are greenish-blue. Some
species of Salmonella will have greenish-blue
colonies with black centers due to H 2 S pro-
duction. Coliform colonies are salmon to or-
ange and may have a bile precipitate.
• Xylose Lysine Desoxycholate (XLD) agar —
although most Salmonella produce red
colonies with black centers, a few may pro-
duce red colonies that lack black centers.
Shigella colonies are red. Coliform colonies
are yellow.
272
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XIV. Medical Microbiology
and Immunology
80. Gram-Negative
Intestinal Pathogens
© The McGraw-H
Companies, 2001
Gram -Negative Intestinal Pathogens • Exercise 80
3
4
Some Pseudomonas produce false-positive red
colonies.
With a straight wire, inoculate the three agar
slants from separate SS-appearing colonies. Use
the streak- stab technique. When streaking the sur-
face of the slant before stabbing, move the wire
over the entire surface for good coverage.
Incubate the slants at 37° C for 18 to 24 hours.
Longer incubation time may cause alkaline rever-
sion. Even refrigeration beyond this time may
cause reversion.
Alkaline reversion is a condition in which
the medium turns yellow during the first part of
the incubation period and then changes to red
later due to increased alkalinity.
Third Period
(Slant Evaluations and Final Inoculations)
During this period you will inoculate tubes of SIM
medium and urea broth with organisms from the
slants of the previous period. Examination of the sep-
aration outline in figure 80.1 reveals that the final step
in the differentiation of the SS pathogens is to deter-
mine whether a non-lactose-fermenter can do three
things: (1) exhibit motility, (2) produce hydrogen sul-
fide, and (3) produce urease. You will also be making
a gram-stained slide to perform a purity check. If
miniaturized multitest media are available, they can
also be inoculated at this time.
Materials:
RDS or Kligler iron agar slants from previous
period
1 tube of SIM medium for each positive slant
1 tube of urea broth for each positive slant
miniaturized multitest media such as API 20E or
Enterotube II (optional)
1 . Examine the slants from the previous period and
select those tubes that have a yellow butt with
a red slant. These tubes contain organisms that
ferment only glucose (non-lactose- fermenters). If
you used Kligler 's iron agar, a black precipitate in
the medium will indicate that the organism is a
producer of H 2 S.
Note in figure 80.1 that slants that are com-
pletely yellow are able to ferment lactose as well
as glucose. Tubes that are completely red are ei-
ther nonfermenters or examples of alkaline rever-
sion. Ignore those tubes.
2. With a loop, inoculate one tube of urea broth from
each slant that has a yellow butt and red slant
(non-lactose-fermenter) .
3
4
5
6
7
With a straight wire, stab one tube of SIM
medium from each of the same agar slants. Stab
in the center to two-thirds of depth of medium.
Incubate these tubes at 37° C for 18 to 24 hours.
Make gram-stained slides from the same slants
and confirm the presence of gram-negative rods.
If miniaturized multitest media are available,
such as API 20E or Enterotube II, inoculate and
incubate for evaluation in the next period. Consult
Exercises 52 and 53 for instructions.
Refrigerate the positive RDS and Kligler iron
slants for future use, if needed.
Fourth Period
(Final Evaluation)
During this last period the tubes of SIM medium, urea
broth, and any miniaturized multitest media from the
last period will be evaluated. Serotyping can also be
performed, if desired.
Materials:
tubes of urea broth and SIM medium from
previous period
Ko vacs' reagent and chloroform
5 ml pipettes
miniaturized multitest media from previous
period
serological testing materials (optional)
1
2
3
4
5
Examine the tubes of SIM medium, checking for
motility and H 2 S production. If you see cloudi-
ness spreading from the point of inoculation, the
organism is motile. A black precipitate will be
evidence of H 2 S production.
Test for indole production by pipetting 2 ml of chlo-
roform into each SIM tube and then adding 2 ml of
Ko vacs' reagent. A pink to deep red color will
form in the chloroform layer if indole is produced.
Salmonella are negative. Some Shigella may
be positive. Citrobacter and Escherichia are pos-
itive.
Examine the urea broth tubes. If the medium has
changed from yellow to red or cerise color, the
organism is urease-positive.
If a miniaturized multitest media was inoculated
in the last period, complete them now.
If time and materials are available, confirm the
identification of your unknown with serological
typing. Refer to Exercise 80.
Laboratory Report
Record the identity of your unknown on the
Laboratory Report and answer all the questions.
273
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XIV. Medical Microbiology
and Immunology
81. Urinary Tract
Pathogens
© The McGraw-H
Companies, 2001
81
Urinary Tract Pathogens
Chronic or acute infections of the urinary tract may
involve the kidneys, ureters, bladder, or urethra. Such
infections may cause high blood pressure, kidney
damage, uremia, or death. In some instances the in-
fections are inapparent and may go unnoticed for
some time. Most infections of this tract enter by way
of the urethra; very few originate in the blood.
A multitude of organisms can cause urinary in-
fections. The most common cause of such infections
in women of childbearing age is Escherichia coli. In
order of frequency after E. coli are other members of
the Enterobacteriaceae, Pseudomonas aeruginosa
and Staphylococcus species.
The importance of performing microbial analyses
of urine on patients with urinary infections cannot be
overemphasized. Some physicians tend to treat pa-
tients with antimicrobics and watch for symptomatic
improvement without performing follow-up urinary
tests, but this practice is not reliable. Clinical testing
of urine 48 to 78 hours after the start of chemotherapy
should be performed to evaluate the effectiveness of
the therapy. If the antimicrobics are effective, the
urine will be free of bacteria at this time.
A thorough microbial analysis of urine from a pa-
tient with urinary distress should be both quantitative
and qualitative. The steps are as follows:
• Collect a urine sample as aseptically as possible.
• Do a plate count to determine the presence or ab-
sence of infection.
• Isolate the pathogen, if an infection is known to
be present.
• Make a presumptive identification of the pathogen.
• Do an antimicrobic sensitivity test.
Except for antimicrobic testing, all of the above
steps will be addressed in this laboratory exercise.
Note that there are two parts to this experiment. The
first portion pertains to doing a plate count. Note,
also, that in figures 81.1 and 81.2 two different meth-
ods are available for making the inoculations. Your in-
structor will indicate which method will be used.
The second portion of this exercise is concerned
with the protocol that one can follow to identify the
genus of a pathogen that might be causing a urinary
infection. Figure 81.3 depicts the routine that we will
use to make this determination. Completion of this
second portion of the exercise will yield only a pre-
sumptive identification of a pathogen. Further phys-
iological tests, which are not included in this exercise,
would be necessary to make species identification.
Aseptic Collection of Urine
Since the urethra in all individuals contains some bac-
teria, especially near its external orifice, the mere
presence of bacteria in urine does not necessarily in-
dicate that a urinary infection exists. One might as-
sume that aseptic collection can be achieved with a
catheter. However, collection by catheterization is,
generally, not desirable because bacteria may be dis-
lodged in the urethra, and there is the danger of caus-
ing an infection with this procedure.
Meaningful results, however, can be obtained
with midstream voided specimens collected in sterile
containers. For best results, the external genitalia
should be cleansed with liquid soap containing
chlorhexidine. Even with midstream samples, how-
ever, one can expect to find low counts of the follow-
ing contaminants in normal urine: coagulase-negative
staphylococci, diphtheroid bacilli, enterococci,
Proteus, hemolytic streptococci, yeasts, and aerobic
gram-positive spore-forming rods.
Specimens are most reliable when plated out im-
mediately after collection. If bacterial tests cannot be
performed immediately, refrigeration is mandatory. It
must be kept in mind that urine is an ideal growth
medium for many bacteria. Specimens not properly re-
frigerated should be considered unsatisfactory for study.
The Plate Count
Before attempting to isolate pathogens from a urine
sample, one should determine, first of all, that an in-
fection actually exists. Since normal urine always
contains some bacteria, yeasts, and other organisms,
we need to know if there is an excess number of or-
ganisms present due to an infection somewhere in the
urinary tract. The best way to make this evaluation is
to do a plate count to determine the number of organ-
isms per ml that are present.
Generally speaking, if a urine sample contains
100,000 or more organisms per ml, one may assume
that significant bacteriuria exists. In some instances,
however, urine from completely normal individuals
■
274
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XIV. Medical Microbiology
and Immunology
81. Urinary Tract
Pathogens
© The McGraw-H
Companies, 2001
may exceed these numbers. It is also possible for
counts between 1,000 and 100,000 to be significant.
Thus, it is apparent that precise evaluation of plate
counts must take into consideration other factors. The
clinician, aware of the effects of certain variables, will
subjectively evaluate the results. Our purpose here in
this experiment is not to interpret, but simply to be-
come familiar with the basic procedures.
Proceed to inoculate two plates of trypticase soy
agar with inocula from a urine sample. After 24 hours
incubation, the colonies will be counted on the best
plate. Your instructor will indicate whether Method A
or Method B will be used.
Method A: Using Calibrated Inoculating
Loops
Note in figure 81.1 that calibrated loops are used to in-
oculate tubes of TSA. After the poured plates have
been incubated for 24 hours, counts will be made.
Proceed as follows:
Materials:
First period:
urine sample
1 sterile empty shake bottle
2 sterile Petri plates
2 trypticase soy agar pours
calibrated wire loops (0.01 |xm and 0.001 (xm)
Second period:
plates from previous period
Quebec colony counter
mechanical hand counter
1. Liquefy two TSA pours and cool to 50° C.
2
3
4
5
6
7
8
Urinary Tract Pathogens • Exercise 81
Label one Petri plate 1:100 and the other 1:1000.
Pour the urine into an empty sterile shake bottle,
cap it tightly, and shake 25 times, as in figure
23.3, page 95.
With a 0.01 calibrated sterile loop, transfer 1
loopful to one of the pours, mix by rolling the tube
between both palms, and pour into the 1:100
plate. Be sure to flame the neck of the tube before
pouring into the plate.
Repeat step 4 using the 0.001 calibrated loop and
the other TSA pour. Pour into the 1 : 1000 plate.
Incubate the plates at 37° C for 24 hours.
After incubation, select the plate that contains be-
tween 30 and 300 colonies. Count all colonies on
a Quebec colony counter, using a mechanical
hand counter to tally.
Record your count and number of organisms per
ml on the Laboratory Report.
Method B: Using Pipettes
This method differs from Method A in that pipettes
are used instead of calibrated loops for making the di-
lutions. Proceed as follows:
Materials:
First period:
urine sample
1 99 ml sterile water blank
1 sterile empty shake bottle
2 sterile Petri plates
2 trypticase soy agar pours
2 1.1 ml dilution pipettes
mechanical pipetting device
Urine Sample
Urine sample is transferred to
sterile shake bottle and shaken
25 times by standard shake
technique.
Using calibrated inoculating loops
0.01 and 0.001 ml of urine are
dispensed to liquefied TSA pours.
Shake
Bottle
SECOND PERIOD
After incubation, colonies on the two
plates are counted. Only counts be-
tween 30 and 300 are considered
significant.
After completely mixing TSA
pours between palms, contents
are poured into Petri plates
and cooled.
ncubate
37° C 24 hr
1:100
Figure 81.1 Method A procedure for doing a plate count
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XIV. Medical Microbiology
and Immunology
81. Urinary Tract
Pathogens
© The McGraw-H
Companies, 2001
Exercise 81 • Urinary Tract Pathogens
Second period:
plates from previous period
Quebec colony counter
mechanical hand counter
1
2
3
4.
5
6
7
8
9
Liquefy two TS A pours and cool to 50° C.
Label one Petri plate 1:100 and the other 1:1000.
Pour the urine into an empty sterile shake bottle,
cap it tightly, and shake 25 times, as in figure
23.3, page 95.
Transfer 1 ml of the mixed urine to a 99 ml ster-
ile water blank. Use a mechanical delivery device
with the pipette.
Mix the water blank with 25 shakes and, with a
fresh pipette, transfer 0.1 ml to the 1:1000 plate
and 1 .0 ml to the 1 : 1 00 plate.
Empty the tubes of TSA into the plates, swirl
them, and let stand to cool.
Incubate the plates at 37° C for 24 hours.
After incubation, select the plate that contains be-
tween 30 and 300 colonies. Count all colonies on
a Quebec colony counter, using a mechanical
hand counter to tally.
Record your count and number of organisms per
ml on the Laboratory Report.
Presumptive Identification
Once it is established that an infection exists, the next
step is to isolate the pathogen and identify it. Figure 81.3
illustrates the overall procedure for making a presump-
tive identification of the genus of a urinary pathogen.
The minimum number of laboratory periods re-
quired to arrive at a presumptive identification is two;
however, if one wishes to be more explicit in identi-
fying the unknown, a total of three or four periods will
be required.
First Period
Note in figure 81.3 that two things will be done dur-
ing this period with a concentrated sample of urine:
(1) two microscope slides will be made for direct ex-
amination, and (2) four kinds of media will be inocu-
lated. Proceed as follows:
Materials:
1 sterile centrifuge tube (with screw cap)
1 tube of thioglycollate medium (BBL135C)
1 plate of blood agar (TSA base)
1 plate of desoxycholate lactose agar (DLA)
1 plate of phenylethyl alcohol medium with
blood (PEA-B)
1 . Shake the sample to resuspend the organisms and
decant 10 ml into a centrifuge tube. Keep the tube
capped.
CAUTION
Be sure to balance centrifuge by placing a capped tube
with 10 ml of water opposite your urine sample tube.
2. Centrifuge for 10 minutes at 2,000 rpm.
3. Decant all but 0.5 ml from the tube and resuspend
the sediment with a sterile wire loop.
Urine Sample
~J>
1.0 m
99 ml
Shake
Bottle
1.100
1 : 1 000
Figure 81.2 Method B procedure for doing a plate count
276
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XIV. Medical Microbiology
and Immunology
81. Urinary Tract
Pathogens
© The McGraw-H
Companies, 2001
4.
5
6
7
8
Inoculate a tube of thioglycollate medium with a
loopful of the sediment.
Streak out a loopful of the sediment on each of the
three agar plates (blood agar, DLA, and PEA-B).
Use a good isolation technique.
Incubate the thioglycollate tube and three plates
at 37° C for 18 to 24 hours.
Make a wet mount slide from material in the bot-
tom of the centrifuge tube and examine under
high-dry, preferably with phase optics. Look for
casts, pus cells, and other elements.
Refer to figure 81.4 for help in identifying ob-
jects that are present. Normal urine will contain
an occasional leukocyte, some epithelial cells,
mucus, bacteria, and crystals of various kinds.
Make a gram- stained slide and examine it under oil
immersion. Determine the morphology and stain-
ing reaction of the predominant organism. Record
your observations on the Laboratory Report.
Urinary Tract Pathogens • Exercise 81
Second Period
After the plates have been incubated for 18 to 24
hours, lay them out and evaluate them according to
the characteristics of each medium.
Thioglycollate Medium This medium is inocu-
lated to promote the growth of organisms that are not
present in large numbers or are too fastidious to grow
readily in nutrient broth. In the event that none of the
plates produce colonies from the urine of a patient
known to have a urinary infection, this tube can be
used for reinoculation or to provide information per-
taining to growth characteristics.
Blood Agar Practically all pathogens of the urinary
tract will grow on this medium. This includes the co-
nforms, Proteus, Pseudomonas, Candida, staphy-
locci, streptococci, and others.
10 ml of urine is
centrifuged in
copper tube.
All but 0.5 ml of
urine is decanted
Gram-stained and
wet mount slides are
made and examined.
Four kinds of media
are inoculated and
incubated at 37°C
for 24 hours.
Thioglycollate
Medium
Blood Agar
Look for:
S. aureus
Hemolytic Strepto-
cocci
Yeasts
etc.
DLA
Look for:
Coliforms
Proteus
Pseudomonas
Salmonella
Shigella
PEA-B
Look for:
Staphylococci
Enterococci
Figure 81.3 Procedure for presumptive identification of urinary pathogens
277
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XIV. Medical Microbiology
and Immunology
81. Urinary Tract
Pathogens
© The McGraw-H
Companies, 2001
Exercise 81 • Urinary Tract Pathogens
Subculturing from this plate to trypticase soy broth
can provide pure cultures of the pathogen for physio-
logical testing or antimicrobic sensitivity testing.
The presence or absence of hemolytic activity can
also be determined at this time. If the pathogen ap-
pears to be a hemolytic, gram-positive coccus, one
should follow the procedures outlined in Exercises 78
and 79 for identification.
Desoxycholate Lactose Agar The presence of
sodium desoxycholate and sodium citrate in this medium
is inhibitory to gram-positive bacteria. Conforms and
other gram-negative bacteria grow well on it.
If the predominant organism is gram- negative,
some differentiation may be made at this point. If the
colonies on this medium are flat and rose-red in color,
the organism is E. coll. Pseudomonas and Proteus,
which do not ferment the lactose in the medium, pro-
duce white colonies. Proteus can be confirmed with
the urease test, being positive for urease production.
Pseudomonas species give a positive reaction
with Taxo N disks. Fermentation and additional phys-
iological testing may be necessary for species identi-
fication. Exercises 48, 49, and 80 should be consulted
for further testing.
Phenylethyl Alcohol Medium This medium, to
which blood has been added, is highly inhibitory to
gram-negative organisms. Proteus, in particular, is
prevented from growing on it. If considerable
growth occurs on the DLA plate, and very little or no
growth here, then one can assume that the disease is
due to a gram-negative organism. This, of course,
would be confirmed by the findings on the gram-
stained slide.
The findings on this plate should be correlated
with those on blood agar. If enterococci (S. faecalis)
are suspected, a plate of Mead agar should be streaked
and incubated at 37° C. Enterococci produce pink
colonies on this medium.
Laboratory Report
Record all findings on the Laboratory Report
Figure 81.4 Microscopic elements in urine
278
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XIV. Medical Microbiology
and Immunology
82. Slide Agglutination
Test: Serological Typing
© The McGraw-H
Companies, 2001
■
Slide Agglutination Test:
Serological Typing
82
Organisms of different species not only differ mor-
phologically and physiologically, but they also differ
in protein makeup. The different proteins of bacterial
cells that are able to stimulate antibody production
when injected into an animal are antigens. The anti-
genic structure of each species of bacteria is unique to
that species and, like the fingerprint of an individual,
can be used to identify the organism. Many closely re-
lated microorganisms that are identical physiologi-
cally can be differentiated only by determining their
antigenic nature.
The method of determining the presence of spe-
cific antigens in a microorganism is called serological
typing (sero typing). It consists of adding a suspen-
sion of the organisms to antiserum, which contains
antibodies that are specific for the known antigens. If
the antigens are present, the antibodies in the anti-
serum will combine with the antigens, causing agglu-
tination, or clumping, of the bacterial cells.
Serotyping is particularly useful in the identification
of various organisms that cause salmonella and
shigella infections. In the identification of the various
serotypes of these two genera, the use of antisera is
generally performed after basic biochemical tests
have been utilized as in Exercise 80.
In this exercise you will be issued two unknown
organisms, one of which is a salmonella. By follow-
ing the procedure shown in figure 82.1, you will de-
termine which one of the unknowns is salmonella.
Note that you will use two test controls. A negative
test control will be set up in depression A on the slide
to see what the absence of agglutination looks like.
The negative control is a mixture of antigen and
saline (antibody is lacking). A positive test control
will be performed in depression C with standardized
antigen and antiserum to give you a typical reaction
of agglutination.
Materials:
2 numbered unknowns per student (slant
cultures of a salmonella and a coliform)
salmonella O antigen, group B
(Difco #2840-56)
salmonella O antiserum, poly A-I
(Difco #2264-47)
depression slides or spot plates
Unknown Organisms
Antiserum
Phenolized Saline
Suspension
Phenolized
Saline
Antigen
Negative Positive Positive
Control Test Result Control
Figure 82.1 Slide agglutination technique
dropping bottle of phenolized saline solution
(0.85% sodium chloride, 0.5% phenol)
2 serological tubes per student
1 -ml pipettes
CAUTION
Keep in mind that Salmonella typhimurium is a
pathogen and can cause gastroenteritis. Be careful!
1 . Label three depressions on a spot plate or depres-
sion slide A, B, and C, as shown in figure 82.1.
2. Make a phenolized saline suspension of each un-
known in separate serological tubes by suspend-
ing one or more loopfuls of organisms in 1 ml of
279
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XIV. Medical Microbiology
and Immunology
82. Slide Agglutination
Test: Serological Typing
© The McGraw-H
Companies, 2001
Exercise 82 • Slide Agglutination Test: Serological Typing
phenolized saline. Mix the organisms sufficiently
to ensure complete dispersion of clumps of bacte-
ria. The mixture should be very turbid.
Transfer 1 loopful (0.05 ml) from the phenolized
saline suspension of one tube to depressions A
andB.
3
4
5
6
To depressions B and C add 1 drop of salmonella
O polyvalent antiserum. To depression A, add 1
drop of phenolized saline, and to depression C,
add 1 drop of salmonella O antigen, group B .
Mix the organisms in each depression with a
clean wire loop. Do not go from one depression to
the other without washing the loop first.
Compare the three mixtures. Agglutination
should occur in depression C (positive control),
7.
but not in depression A (negative control). If ag-
glutination occurs in depression B, the organism
is salmonella.
Repeat this process on another slide for the other
organism.
CAUTION
Deposit all slides and serological tubes in container
of disinfectant provided by the instructor.
Laboratory Report
Record your results on the first portion of Laboratory
Report 82, 83.
280
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XIV. Medical Microbiology
and Immunology
83. Slide Agglutination
(Latex) Test: For S. aureus
Identification
© The McGraw-H
Companies, 2001
Slide Agglutination (Latex) Test:
For S. aureus Identification
83
Many manufacturers of reagents for slide agglutina-
tion tests utilize polystyrene latex particles as carriers
for the antibody particles. By adsorbing reactive anti-
body units to these particles, an agglutination reaction
results that occurs rapidly and is much easier to see
than ordinary precipitin type reactions that might be
used to demonstrate the presence of a soluble antigen.
In this exercise we will use reagents manufac-
tured by Difco Laboratories to determine if a sus-
pected staphylococcus organism produces coagulase
and/or protein A. The test reagent {Difco Staph
Reagent) is a suspension of yellow latex particles sen-
sitized with antibodies for coagulase and protein A.
Reagents are also included to provide positive and
negative controls in the test. Instead of using depres-
sion slides or spot plates, Difco provides disposable
cards with eight black circles printed on them for per-
forming the test. As indicated in figure 83.1, only
three circles are used when performing the test on one
unknown. The additional circles are provided for test-
ing five additional unknowns at the same time. The
black background of the cards facilitates rapid inter-
pretation by providing good contrast for the yellow
clumps that form.
There are two versions of this test: direct and in-
direct. The procedure for the direct method is illus-
trated in figure 83.1. The indirect method differs in
that saline is used to suspend the organism being
tested.
It should be pointed out that the reliability corre-
lation between this test for coagulase and the tube test
(page 260) is very high. Studies reveal that a reliabil-
ity correlation of over 97% exists. Proceed as follows
to perform this test.
+
One drop of positive control
reagent is added to circle #1
One drop of latex reagent is added to
each of three circles as shown.
Disposable test slide (Difco)
TEST CULTURE
The slide is rocked by hand for 45 seconds and
placed on a slide rotator for another 45 seconds.
One drop of negative control
reagent is added to circle #2.
Two complete colonies are quickly
and completely emulsified into re-
agent in circle #3.
Figure 83.1 Slide agglutination test (direct method) for the presence of coagulase and/or protein A
281
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XIV. Medical Microbiology
and Immunology
83. Slide Agglutination
(Latex) Test: For S. aureus
Identification
© The McGraw-H
Companies, 2001
Exercise 83 • Slide Agglutination (Latex) Test: For S. aureus Identification
Materials:
plate culture of staphylococcus-like organism
(trypticase soy agar plus blood)
Difco Staph Latex Test kit #3850-32-7, which
consists of:
bottle of Bacto Staph Latex Reagent
bottle of Bacto Staph Positive Control
bottle of Bacto Staph Negative Control
bottle of Bacto Normal Saline Reagent
disposable test slides (black circle cards)
mixing sticks (minimum of 3)
slide rotator
Direct Method
If the direct method is to be used, as illustrated in fig-
ure 83.1, follow this procedure:
1
2
3
4.
5
6
7
Place 1 drop of Bacto Staph Positive Control
reagent onto circle # 1 .
Place 1 drop of Bacto Staph Negative Control
reagent on circle #2.
Place 1 drop of Bacto Staph Latex Reagent onto
circles #1, #2, and #3.
Using a sterile inoculating needle or loop, quickly
and completely emulsify two isolated colonies
from the culture to be tested into the drop of Staph
Latex Reagent in circle #3 .
Also, emulsify the Staph Latex Reagent in the
positive and negative controls in circles #1 and #2
using separate mixing sticks supplied in the kit.
All mixing in these three circles should be
done quickly to minimize drying of the latex on the
slide and to avoid extended reaction times for the
first cultures emulsified.
Rock the slide by hand for 45 seconds.
Place the slide on a slide rotator capable of pro-
viding 1 1 to 1 20 rpm and rotate it for another 45
seconds.
Read the results immediately, according to the de-
scriptions provided in the table at right. If agglu-
tination occurs before 45 seconds, the results may
be read at that time. The slide should be read at
normal reading distance under ambient light.
Indirect Method
The only differences between the direct and indirect
methods pertain to the amount of inoculum and the
use of saline to emulsify the unknown being tested.
Proceed as follows:
1
2
3
4
5
6
7
8
9
Place 1 drop of Bacto Staph Positive Control
reagent onto test circle #1.
Place 1 drop of Bacto Staph Negative Control
onto circle #2.
Place 1 drop of Bacto Normal Saline Reagent
onto circle #3.
Using a sterile inoculating needle or loop, com-
pletely emulsify four isolated colonies from the
culture to be tested into the circle containing the
drop of saline (circle #3).
Add 1 drop of Bacto Staph Latex Reagent to each
of the three circles.
Quickly mix the contents of each circle, using in-
dividual mixing sticks.
Rock the slide by hand for 45 seconds.
Place the slide on a slide rotator capable of pro-
viding 1 1 to 1 20 rpm and rotate it for another 45
seconds.
Read the results immediately according to the de-
scriptions provided in the table below. If aggluti-
nation occurs before 45 seconds, the results may
be read at that time. The slide should be read at
normal reading distance under ambient light.
POSITIVE REACTIONS
4 + Large to small clumps of aggregated yellow latex
beads; clear background
3 + Large to small clumps of aggregated yellow latex
beads; slightly cloudy background
2 + Medium to small but clearly visible clumps of aggre
gated yellow latex beads; moderately cloudy back-
ground
1 + Fine clumps of aggregated yellow latex beads;
cloudy background
NEGATIVE REACTIONS
+
Smooth cloudy suspension; particulate grainy
appearance that cannot be identified as agglutina
tion
Smooth, cloudy suspension; free of agglutination or
particles
Laboratory Report
Record your results on the last portion of Laboratory
Report 82, 83.
282
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XIV. Medical Microbiology
and Immunology
84. Tube Agglutination
Test: The Heterophile
Antibody Test
© The McGraw-H
Companies, 2001
Tube Agglutination Test:
The Heterophile Antibody Test
84
Infectious mononucleosis (IM) is a benign disease,
occurring principally in individuals in the 13 to 25
year age group. It is caused by the Epstein-Barr virus
(EB V), a herpesvirus, that is one of the most ubiqui-
tous viruses in humans. Studies have shown that the
virus can be isolated from saliva of patients with IM,
as well as from some healthy, asymptomatic individ-
uals. Between 80% and 90% of all adults possess an-
tibodies for EBV.
The disease is characterized by a sudden onset of
fever, sore throat, and pronounced enlargement of the
cervical lymph nodes. There is also moderate leuko-
cytosis with a marked increase in the number of lym-
phocytes (50% to 90%).
The serological test for IM takes advantage of an
unusual property: the antibodies produced against the
EBV coincidentally agglutinate sheep red blood cells.
This is an example of a heterophile antigen — a sub-
stance isolated from a living form that stimulates the
production of antibodies capable of reacting with tis-
sues of other organisms. The antibodies are referred to
as heterophile antibodies.
This test is performed by adding a suspension of
sheep red blood cells to dilutions of inactivated pa-
tient's serum and incubating the tubes overnight in the
refrigerator. Figure 84. 1 illustrates the overall proce-
dure. Agglutination titers of 320 or higher are consid-
ered significant. Titers of 40,960 have been obtained.
Proceed as follows to perform this test on a sam-
ple of test serum:
First Period
Materials:
test-tube rack (Wasserman type) with 10 clean
serological tubes
0.5 ML TRANSFERRED FROM TUBE TO TUBE
1
1:5
KJ,
%J
%J
7
8
%JJ
10
DISCARD
\JJ
1:10 1:20 1:40 1:80 1:160 1:320 1:640 1:1280 CONTROL
0.2 ml inactivated patient's
serum and 0.8 ml saline
0.5 ML SALINE PER TUBE
Figure 84.1 Procedure for setting up heterophile antibody test
283
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XIV. Medical Microbiology
and Immunology
84. Tube Agglutination
Test: The Heterophile
Antibody Test
© The McGraw-H
Companies, 2001
Exercise 84 • Tube Agglutination Test: The Heterophile Antibody Test
1
2
3
4
5
6
bottle of saline solution (0.85% NaCl), clear or
filtered
1 ml pipettes
5 ml pipettes
2% suspension of sheep red blood cells
patient's serum (known to be positive)
Place the test serum in a 56° C water bath for 30
minutes to inactivate the complement.
Set up a row of 10 serological tubes in the front
row of a test-tube rack and number them from 1
to 10 (left to right) with a marking pencil.
Into tube 1, pipette 0.8 ml of physiological saline.
Dispense 0.5 ml of physiological saline to tubes 2
through 10. Use a 5 ml pipette.
With a 1 ml pipette add 0.2 ml of the inactivated
serum to tube 1 . Mix the contents of this tube by
drawing into the pipette and expelling about five
times .
Transfer 0.5 ml from tube 1 to tube 2, mix five
times, and transfer 0.5 ml from tube 2 to tube 3,
etc., through the ninth tube. Discard 0.5 ml from
the ninth tube after mixing. Tube 10 is the control.
7
8
Add 0.2 ml of 2% sheep red blood cells to all
tubes (1 through 10) and shake the tubes. Final di-
lutions of the serum are shown in figure 84.1.
Allow the rack of tubes to stand at room temper-
ature for 1 hour, then transfer the tubes to a small
wire basket, and place in a refrigerator to remain
overnight.
Second Period
Set up the tubes in a tube rack in order of dilution and
compare each tube with the control by holding the
tubes overhead and looking up at the bottoms of the
tubes. Nonagglutinated cells will tumble to the bot-
tom of the tube and form a small button (as in control
tube). Agglutinated cells will form a more- amorphous
"blanket."
The titer should be recorded as the reciprocal of the
last tube in the series that shows positive agglutination.
Laboratory Report
Complete the first portion of Laboratory Report 84, 85
284
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XIV. Medical Microbiology
and Immunology
85. Tube Agglutination: The
WidalTest
© The McGraw-H
Companies, 2001
Tube Agglutination Test:
The Widal Test
85
A tube test for determining the quantity of agglutinat-
ing antibodies, or agglutinins, in the serum of a pa-
tient with typhoid fever was described by Grunbaum
and Widal in 1896. This technique is still in use today
and has been adapted to many other diseases as well.
The procedure involves adding a suspension of
dead typhoid bacterial cells to a series of tubes con-
taining the patient's serum, which has been diluted out
to various concentrations. After the tubes have been
incubated for 30 minutes at 37° C, they are cen-
trifuged and examined to note the amount of aggluti-
nation that has occurred.
The reciprocal of the highest dilution at which ag-
glutination is seen is designated as the antibody titer
of the patient's serum. For example, if the highest di-
lution at which agglutination occurs is 1 :320, the titer
is 320 antibody units per milliliter of serum.
Naturally, the higher the titer, the greater is the anti-
body response of the individual to the disease.
This technique can be used clinically to deter-
mine whether a patient with typhoidlike symptoms
actually has the disease. If successive daily tests on a
patient's serum reveal no antibody titer, or a low titer
that does not increase from day to day, it can be as-
sumed that some other disease is present. On the
other hand, if one sees a daily increase in the titer, it
can be assumed that a typhoid infection does exist.
Since the treatment of typhoid fever requires power-
ful antibiotics that are not widely used on other sim-
ilar diseases, it is very important to diagnose this dis-
ease early to begin the proper form of chemotherapy
as soon as possible.
In this exercise you will be given a sample of
blood serum that is known to contain antibodies for
the typhoid organism. By using the Widal tube agglu-
tination method, you will determine the antibody titer.
Materials:
test-tube rack (Wassermann type) with 1 clean
serological tubes
bottle of saline solution (0.85%), clear or
filtered
0.5 ML TRANSFERRED FROM TUBE TO TUBE
1
u
7
8
\\
10
ONE ML OF PATIENT'S
SERUM (1:10)
x>
s>
<§>
\
%
%
DISCARD
0.5 ML SALINE PER TUBE
Figure 85.1 Procedure for dilution of serum
285
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XIV. Medical Microbiology
and Immunology
85. Tube Agglutination: The
WidalTest
© The McGraw-H
Companies, 2001
Exercise 85 • Tube Agglutination Test: The Widal Test
1 ml pipettes
5 ml pipettes
water bath at 37° C
centrifuges
antigen (1:10 dilution) Salmonella typhi "O"
patient's serum (1:10 dilution), known 2
positive "O"
1 . Dilute the patient's serum as shown in figure 85. 1 .
Follow this routine:
a. Set up 1 clean serological test tubes in the front
row of a test-tube rack and number them from 1
to 10 (left to right) with a marking pencil.
b. Into tube 1 pipette 1 ml of the patient's serum
(1:10 dilution). For convenience, the instructor
may wish to dispense this material to each stu-
dent.
c. With a 5 ml pipette, dispense 0.5 ml of saline
to each of the remaining nine tubes.
d. With a 1 ml pipette, transfer 0.5 ml of the
serum from tube 1 to tube 2. Mix the serum
and saline in tube 2 by carefully drawing the
liquid up into the pipette and discharging it
slowly back down into the tube a minimum of
three times.
e. Repeat this process by transferring 0.5 ml from
tube 2 to 3, tube 3 to 4, 4 to 5, etc. When you
3
4
5
6
get to tube 9, discard the 0.5 ml drawn from it
instead of adding it to tube 10; thus, tube 10
will contain only saline and can be used as a
negative test control for comparing with the
other tubes.
With a fresh 5 ml pipette, transfer 0.5 ml of anti-
gen to each tube. Shake the rack to completely
mix the antigen and diluted serum.
Place the rack in a water bath at 37° C for 30 min-
utes.
Centrifuge all tubes for 3 minutes at 2,000 rpm.
(If time permits, 7 minutes centrifugation is
preferable.)
Examine each tube for agglutination and record
the titer as the reciprocal of the highest dilution in
which agglutination is seen.
When examining each tube, jar it first by rap-
ping the side of the tube with a snap of the finger
to suspend the clumps of agglutinated cells. Hold
it up against the light of a desk lamp in the man-
ner shown in figure 85.2. Do not look directly into
the light. The reflection of the light off the parti-
cles is best seen against a dark background.
Compare each tube with tube 10, which is your
negative test control.
Record your results on the last portion of
Laboratory Report 84, 85.
EYEPOINT
BLACK SURFACE
Figure 85.2 Agglutination is more readily seen when the tube is examined against a black surface
286
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Applications Lab Manual,
Eighth Edition
XIV. Medical Microbiology
and Immunology
86. Phage Testing
© The McGraw-H
Companies, 2001
■
Phage Typing
86
The host specificity of bacteriophage is such that it is
possible to delineate different strains of individual
species of bacteria on the basis of their susceptibility
to various kinds of bacteriophage. In epidemiological
studies, where it is important to discover the source of
a specific infection, determining the phage type of the
causative organism can be an important tool in solv-
ing the riddle. For example, if it can be shown that the
phage type of S. typhi in a patient with typhoid fever
is the same as the phage type of an isolate from a sus-
pected carrier, chances are excellent that the two cases
are epidemiologically related. Since all bacteria are
probably parasitized by bacteriophages, it is theoreti-
cally possible, through research, to classify each
species into strains or groups according to their phage
type susceptibility. This has been done for
Staphylococcus aureus, Salmonella typhi, and several
other pathogens. The following table illustrates the
lytic groups of S. aureus as proposed by M. T. Parker.
Lytic Group
Phages in Group
I
29 52 52A 79 80
II
3A 3B 3C 55 71
III
6 7 42E 47 53 54 75 77 83A
IV
42D
not allotted
81 187
In bacteriophage typing, a suspension of the or-
ganism to be typed is swabbed over an agar surface.
The bottom of the plate is marked off in squares and
labeled to indicate which phage types are going to be
used. To the organisms on the surface, a small drop of
each phage type is added to their respective squares.
After incubation, the plate is examined to see which
Agar is swabbed
with organism to
be typed.
/
37° C 24 hr
Different phage
types are added
to swabbed sur-
face of medium.
Bacteriophages that
cause plaque formation
determine the phage
type of the unknown.
Figure 86.1 Bacteriophage typing
phages were able to lyse the organisms. This is the
procedure to be used in this exercise. See figure 86.1.
Materials:
1 Petri plate of tryptone yeast extract agar
bacteriophage cultures (available types)
nutrient broth cultures of S. aureus with swabs
1
2
3
4
5
Mark the bottom of a plate of tryptone yeast ex-
tract agar with as many squares as there are phage
types to be used. Label each square with the
phage type numbers.
Swab the entire surface of the agar with the or-
ganisms.
Deposit 1 drop of each phage in its respective
square.
Incubate the plate at 37° C for 24 hours and record
the lytic group and phage type of the culture.
Record your results on the Laboratory Report.
287
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XIV. Medical Microbiology
and Immunology
87. White Blood Cell Study:
The Differential WBC
Count
© The McGraw-H
Companies, 2001
87
White Blood Cell Study:
The Differential WBC Count
In 1883, at the Pasteur Institute in Paris, Metchnikoff
published a paper proposing the phagocytic theory of
immunity. On the basis of his studies performed on
transparent starfish larvae, he postulated that amoe-
boid cells in the tissue fluid and blood of all animals
are the major guardians of health against bacterial in-
fection. He designated the large phagocytic cells of
the blood as macrophages and the smaller ones as mi-
crophages. Today, Metchnikoff s macrophages are
known as monocytes and his microphages as neu-
trophils or polymorphonuclear leukocytes.
Figure 87.1 illustrates the five types of leukocytes
that are normally seen in the blood. Blood platelets
and erythrocytes also are shown to present a complete
picture of all formed elements in the blood. When ob-
served as living cells under the microscope, they ap-
pear as refractile, colorless structures. As shown here,
however, they reflect the dyes that are imparted by
Wright's stain.
In this exercise we will do a study of the white
blood cells in human blood. This study may be made
from a prepared stained microscope slide or from a
slide made from your own blood. By scanning an en-
tire slide and counting the various types, you will
have an opportunity to encounter most, if not all,
types. The erythrocytes and blood platelets will be
ignored.
Figures 87.1 and 87.2 will be used to identify the
various types of cells. Figure 87.3 illustrates the pro-
cedure for preparing a slide stained with Wright's
stain. The relative percentages of each type will be de-
termined after a total of 100 white blood cells have
been identified. This method of white blood cell enu-
meration is called a differential WBC count.
■
VN^H^n
-
I
EaSfJCJPHLEi
BASCFHLb
*
X
-
r
MONOCYTES
^
IUTELETB
K.P. Talaro
Figure 87.1 Formed elements of blood
288
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XIV. Medical Microbiology
and Immunology
87. White Blood Cell Study:
The Differential WBC
Count
© The McGraw-H
Companies, 2001
White Blood Cell Study: The Differential WBC Count • Exercise 87
As you proceed with this count, it will become
obvious that the neutrophils are most abundant
(50%-70%). The next most prominent cells are the
lymphocytes (20%-30%). Monocytes comprise about
2%-6%; eosinophils, l%-5%; and basophils, less
than 1%.
A normal white blood cell count is between 5,000
and 10,000 white cells per cubic millimeter. Elevated
white blood cell counts are referred to as leukocytosis;
counts of 30,000 or 40,000 represent marked leuko-
cytosis. When counts fall considerably below 5,000,
leukopenia is said to exist. Both conditions can have
grave implications.
The value of a differential count is immeasurable
in the diagnosis of infectious diseases. High neu-
trophil counts, or neutrophilia, often signal localized
infections, such as appendicitis or abscesses in some
other part of the body. Neutropenia, a condition in
which there is a marked decrease in the numbers of
neutrophils, occurs in typhoid fever, undulant fever,
and influenza. Eosinophilia may indicate allergic con-
ditions or invasions by parasitic roundworms such as
Trichinella spiralis, the "pork worm." Counts of
eosinophils may rise to as high as 50% in cases of
trichinosis. High lymphocyte counts, or lymphocyto-
sis, are present in whooping cough and some viral in-
fections. Increased numbers of monocytes, or mono-
cytosis, may indicate the presence of the Epstein-Barr
virus, which causes infectious mononucleosis.
Note in the materials list that items needed for
making a slide (option B) are listed separately. If a
prepared slide (option A) is to be used, ignore the in-
structions under the heading "Preparation of Slide,"
and proceed to the heading "Performing the Cell
Count." Your instructor will indicate which option
will be used. Proceed as follows:
NEUTROPHIL
EOSINOPHIL
GRANULOCYTES
BASOPHIL
BLOOD PLATELETS
LYMPHOCYTES
MONOCYTE
AGRANULOCYTES
PLASMA CELL
(abnormal)
Figure 87.2 Photomicrographs of formed elements in blood
289
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XIV. Medical Microbiology
and Immunology
87. White Blood Cell Study:
The Differential WBC
Count
© The McGraw-H
Companies, 2001
Exercise 87 • White Blood Cell Study: The Differential WBC Count
PRECAUTIONS
When working with blood observe the following
precautions:
1 . Always disinfect the finger with alcohol prior to
piercing it.
2. Use sterile disposable lancets only one time.
3. Dispose of used lancets by placing them into a
beaker of disinfectant.
4. Avoid skin contact with blood of other students.
Wear disposable latex gloves.
5. Disinfect finger with alcohol after blood has
been taken.
Materials:
prepared blood slide (option A):
stained with Wright's or Giemsa's stains
for staining a blood smear (option B):
2 or 3 clean microscope slides (should have
polished edges)
sterile disposable lancets
disposable latex gloves
sterile absorbent cotton, 70% alcohol
Wright's stain, wax pencil, bibulous paper
distilled water in dropping bottle
Preparation of Slide
Figure 87.3 illustrates the procedure that will be used
to make a stained slide of a blood smear. The most dif-
ficult step in making such a slide is getting a good
spread of the blood, which is thick at one end and thin
at the other end. If done properly, the smear will have
a gradient of cellular density that will make it possi-
ble to choose an area that is ideal for study. The angle
at which the spreading slide is held in making the
smear will determine the thickness of the smear. It
may be necessary for you to make more than one slide
to get an ideal one.
1
2
Clean three or four slides with soap and water.
Handle them with care to avoid getting their flat
surfaces soiled by your fingers. Although only
two slides may be used, it is often necessary to
repeat the spreading process, thus the extra
slides.
Scrub the middle finger with 70% alcohol and
stick it with a lancet. Put a drop of blood on the
slide V" from one end and spread with another
slide in the manner illustrated in figure 87.3.
Note that the blood is dragged over the slide,
not pushed. Do not pull the slide over the smear a
second time. If you don't get an even smear the
A small drop of blood is placed about 3/4 inch
away from one end of slide. The drop should
not exceed 1/8" diameter.
Trie spreader slide is moved in direction of
arrow, allowing drop of blood to spread along
slide's back edge.
wax tines
The spreader slide is pushed along the slide,
dragging the blood over the surface of the
slide.
A china marking pencil is used to mark off both
ends of the smear to retain the staining
solution on the slide.
Figure 87.3 Smear preparation technique for making a stained blood slide
290
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XIV. Medical Microbiology
and Immunology
87. White Blood Cell Study:
The Differential WBC
Count
© The McGraw-H
Companies, 2001
White Blood Cell Study: The Differential WBC Count • Exercise 87
3
4.
5
first time, repeat the process on a fresh clean slide.
To get a smear that will be the proper thickness,
hold the spreading slide at an angle somewhat
greater than 45 ° .
Draw a line on each side of the smear with a wax
pencil to confine the stain that is to be added.
(Note: This step is helpful for beginners, and usu-
ally omitted by professionals.)
Cover the film with Wright's stain, counting the
drops as you add them. Stain for 4 minutes and
then add the same number of drops of distilled
water to the stain and let stand for another 10
minutes. Blow gently on the mixture every few
minutes to keep the solutions mixed.
Gently wash off the slide under running water for
30 seconds and shake off the excess. Blot dry with
bibulous paper.
Performing the Cell Count
Whether you are using a prepared slide or one that you
have just stained, the procedure is essentially the
same. Although the high-dry objective can be used for
the count, the oil immersion lens is much better.
Differentiation of some cells is difficult with high-dry
optics. Proceed as follows:
1
2
Scan the slide with the low-power objective to
find an area where cell distribution is best. A good
area is one in which the cells are not jammed to-
gether or scattered too far apart.
Systematically scan the slide, following the path-
way indicated below in figure 87.4. As each
leukocyte is encountered, identify it, using fig-
ures 87.1 and 87.2 for reference.
«MM
Figure 87.4 Path to follow when seeking cells
3. Tabulate your count on the Laboratory Report
sheet according to the instructions there. It is best
to remove the Lab Report sheet from the back of
the manual for this identification and tabulation
procedure.
Laboratory Report
Complete the first portion of Laboratory Report
87-89.
291
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XIV. Medical Microbiology
and Immunology
88. Total WBC Count
© The McGraw-H
Companies, 2001
Total WBC Count
Although the differential white blood cell count provides
us with the relative percentages of leukocytes, it alone
cannot reveal the true picture of the extent of an infec-
tion. For a more complete picture, one must also know
the total number of WBCs per cubic millimeter of blood.
Although the number of leukocytes may vary
with the time of day, exercise, and other factors, a
range of 5,000 to 9,000 WBCs per cubic millimeter is
considered normal. If an individual were to have an
abnormally high neutrophil percentage, and a total
count of, say, 17,000 WBCs, the presence of an infec-
tion of some sort would be highly probable.
To determine the number of leukocytes in a cubic
millimeter of blood, one must dilute the blood and
count the WBCs on a specialized slide called a hema-
cytometer. Figure 88.2 shows the sequence of steps in
performing for this count.
Note in illustration 1 of figure 88.2 that blood is
drawn up into a special pipette and then diluted in the
pipette with a weak acid solution (illustration 2). After
shaking the pipette (illustration 3) to mix the acid and
blood, a small amount of diluted blood is allowed to
flow under the cover glass of the hemacytometer (il-
lustration 4). The count of white blood cells is then
made with the low-power microscope objective.
Preparation of
Hemacytometer
Working with your laboratory partner, assist each other
to prepare a "charged" hemacytometer as follows:
PRECAUTIONS
Review the precautions that are stated in the previous
exercise and use a mechanical suction device as
shown in figure 88.1.
Materials:
hemacytometer and cover glass
WBC diluting pipette
WBC diluting fluid
mechanical hand counter
mechanical suction device
pipette cleaning solutions
cotton, alcohol, lancets, clean cloth
Figure 88.1 Using a mechanical suction device to draw
up fluids into pipette
1
2
3
Wash the hemacytometer and cover glass with
soap and water, rinse well, and dry with a clean
cloth or Kim wipes.
Produce a free flow of blood, wipe away the first
drop and draw the blood up into the diluting
pipette to the 0.5 mark. See illustration 1 of figure
88.2. If the blood happens to go a little above the
mark, the volume can be reduced to the mark by
placing the pipette tip on a piece of blotting paper.
If the blood goes substantially past the 0.5
mark, discharge the blood into a beaker of disin-
fectant, wash the pipette in the four cleansing so-
lutions (illustration 6, figure 88.2), and start over.
The ideal way is to draw up the blood exactly to
the mark on the first attempt. To clean the pipette,
rinse it first with acid, then water, alcohol, and fi-
nally with acetone.
As shown in illustration 2, figure 88.2, draw the
WBC diluting fluid up into the pipette until it
reaches the 11.0 mark.
292
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Eighth Edition
XIV. Medical Microbiology
and Immunology
88. Total WBC Count
© The McGraw-H
Companies, 2001
Total WBC Count • Exercise 88
4.
5
6
7
Place your thumb over the tip of the pipette, and
place your third finger over the other end (illus-
tration 3, figure 88.2).
Mix the blood and diluting fluid in the pipette for
2-3 minutes by holding it as shown in illustration
3, figure 88.2. The pipette should be held parallel
to the tabletop and moved through a 90° arc, with
the wrist held rigidly.
Discharge one-third of the bulb fluid from the
pipette by allowing it to drop onto a piece of pa-
per toweling.
While holding the pipette as shown in illustration
1, figure 88.2, deposit a tiny drop on the polished
surface of the counting chamber next to the edge
of the cover glass. Do not let the tip of the
pipette touch the polished surface for more
than an instant. If it is left there too long, the
chamber will overfill.
A properly filled chamber will have diluted
blood filling only the space between the cover
glass and counting chamber. No fluid should run
down into the moat.
8 . Charge the other side if the first side was overfilled.
Performing the Count
Place the hemacytometer on the microscope stage and
bring the grid lines into focus under the low-power
(10X) objective. Use the coarse adjustment knob and
reduce the lighting somewhat to make both the cells
and lines visible.
Locate one of the large "W" (white) areas shown
in illustration 5, figure 88.2. One of these areas should
fill up the entire field. Since the diluting fluid contains
acid, all red blood cells have been destroyed; only the
leukocytes will show up as very small dots.
Blood is drawn up into pipette
to the 0.5 mark.
WBC dilution fluid is drawn up
to 11.0 mark.
Blood and dilution fluid is mixed
for 2-3 minutes by shaking.
Hemacytometer chamber is charged
with diluted blood.
All leukocytes are counted in the
four large "W" areas.
After using, pipette must be
rinsed with four solutions.
Figure 88.2 Procedure for charging a hemacytometer and doing a total white blood cell count
293
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XIV. Medical Microbiology
and Immunology
88. Total WBC Count
© The McGraw-H
Companies, 2001
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XIV. Medical Microbiology
and Immunology
88. Total WBC Count
© The McGraw-H
Companies, 2001
Total WBC Count • Exercise 88
Do the cells seem to be evenly distributed? If not,
charge the other half of the counting chamber after
further mixing. If the other chamber had been previ-
ously charged unsuccessfully by overflooding, wash
off the hemacytometer and cover glass, shake the
pipette for 2-3 minutes, and recharge it.
Count all the cells in the four "W" areas, using a
mechanical hand counter. To avoid overcounting of
cells at the boundaries, count the cells that touch the
lines on the left and top sides only. Cells that touch
the boundary lines on the right and bottom sides
should be ignored. This applies to the boundaries of
each entire "W" area.
Discharge the contents of the pipette and rinse it
out by sequentially flushing with the following fluids:
acid, water, alcohol, and acetone. See illustration 6,
figure 88.2.
Calculations
To determine the number of leukocytes per cubic mil-
limeter, multiply the total number of cells counted in the
four "W" areas by 50. The factor of 50 is the product of
the volume correction factor and dilution factor, or
2.5 X 20 = 50
Laboratory Report
Answer the questions that pertain to this experiment
on Laboratory Report 87-89.
295
Benson: Microbiological
XIV. Medical Microbiology
89. Blood Grouping
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
and Immunology
Companies, 2001
89
Blood Grouping
Exercises 82 through 85 illustrate three uses of agglu-
tination tests as related to (1) the identification of
serological types, (2) species identification (S. au-
reus), and (3) disease identification (infectious
mononucleosis and typhoid fever). The typing of
blood is another example of a medical procedure that
relies on this useful phenomenon.
The procedure for blood typing was developed
by Karl Landsteiner around 1900. He is credited
with having discovered that human blood types can
be separated into four groups on the basis of two
antigens that are present on the surface of red blood
cells. These antigens are designated as A and B. The
four groups (types) are A, B, AB, and O. The last
group type O, which is characterized by the absence
of A or B antigens, is the most common type in the
United States (45% of the population). Type A is
next in frequency, found in 39% of the population.
The incidences of types B and AB are 12% and 4%,
respectively.
Blood typing is performed with antisera con-
taining high titers of anti-A and anti-B antibodies.
The test may be performed by either slide or tube
methods. In both instances, a drop of each kind of
antiserum is added to separate samples of saline
suspension of red blood cells. Figure 89.1 illustrates
the slide technique. If agglutination occurs only in
the suspension to which the anti-A serum was
added, the blood is type A. If agglutination occurs
only in the anti-B mixture, the blood is type B.
Agglutination in both samples indicates that the
blood is type AB. The absence of agglutination in-
dicates that the blood is type O.
Between 1900 and 1940 a great deal of research
was done to uncover the presence of other antigens
in human red blood cells. Finally, in 1940,
Landsteiner and Wiener reported that rabbit sera
containing antibodies against the red blood cells of
the rhesus monkey would agglutinate the red blood
cells of 5% of white humans. This antigen in hu-
mans, which was first designated as the Rh factor
(in due respect to the rhesus monkey), was later
found to exist as six antigens: C, c, D, d, E, and e. Of
these six antigens, the D factor is responsible for the
Rh-positive condition and is found in 85% of Cau-
casians, 94% of blacks, and 99% of orientals.
Typing blood for the Rh factor can also be per-
formed by both tube and slide methods, but there are
certain differences in the two techniques. First of all,
the antibodies in the typing sera are of the incomplete
albumin variety, which will not agglutinate human
red cells when they are diluted with saline. Therefore,
it is necessary to use whole blood or dilute the cells
with plasma. Another difference is that the test must
be performed at higher temperatures: 37° C for tube
test, 45° C for the slide test.
In this exercise, two separate slide methods are
presented for typing blood. If only the Landsteiner
ABO groups are to be determined, the first method
may be preferable. If Rh typing is to be included, the
second method, which utilizes a slide warmer, will be
followed. The availability of materials will determine
which method is to be used.
PRECAUTIONS
Review the precautionary comments highlighted
on page 290.
ABO Blood Typing
Materials:
small vial (10 mm dia X 50 mm long)
disposable lancets (B-D Microlance,
Serasharp, etc.)
70% alcohol and cotton
china marking pencil
microscope slides
typing sera (anti-A and anti-B)
applicators or toothpicks
saline solution (0.85% NaCl)
1 ml pipettes
disposable latex gloves
1
2
3
Mark a slide down the middle with a marking
pencil, dividing the slide into two halves as
shown in figure 89.1. Write "anti-A" on the left
side and "anti-B" on the right side.
Pipette 1 ml of saline solution into a small vial or
test tube.
Scrub the middle finger with a piece of cotton
saturated with 70% alcohol and pierce it with a
296
Benson: Microbiological
XIV. Medical Microbiology
89. Blood Grouping
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
and Immunology
Companies, 2001
4.
5.
sterile disposable lancet. Allow 2 or 3 drops of
blood to mix with the saline by holding the finger
over the end of the vial and washing it with the
saline by inverting the tube several times.
Place a drop of this red cell suspension on each
side of the slide.
Add a drop of anti-A serum to the left side of the
slide and a drop of anti-B serum to the right side.
Blood Grouping • Exercise 89
Do not contaminate the tips of the serum
pipettes with the material on the slide.
6. After mixing each side of the slide with separate
applicators or toothpicks, look for agglutination.
The slide should be held about 6" above an illu-
minated white background and rocked gently for
2 or 3 minutes. Record your results on the
Laboratory Report as of 3 minutes.
ANTI-A '.
SERUM V
I
t
Saline Suspension of
Red Blood Cells
••
ANTI-B
SERUM
Type O
(No Agglutination)
Type A
Type B
Type A B
Figure 89.1 Typing of ABO blood groups
297
Benson: Microbiological
XIV. Medical Microbiology
89. Blood Grouping
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
and Immunology
Companies, 2001
Exercise 89 • Blood Grouping
Combined ABO and Rh Typing
As stated, Rh typing must be performed with heat on
blood that has not been diluted with saline. A warm-
ing box such as the one in figure 89.2 is essential in
this procedure. In performing this test, two factors
are of considerable importance: first, only a small
amount of blood must be used (a drop of about 3 mm
diameter on the slide) and, second, proper agitation
must be executed. The agglutination that occurs in this
antibody-antigen reaction results in finer clumps;
therefore, closer examination is essential. If the agi-
tation is not properly performed, agglutination may
not be as apparent as it should be.
In this combined method we will use whole blood
for the ABO typing as well as for the Rh typing.
Although this method works satisfactorily as a class-
room demonstration for the ABO groups, it is not as
reliable as the previous method in which saline and
room temperature are used. This method is not recom-
mended for clinical situations.
Materials:
slide warming box with a special marked slide
anti-A, anti-B, and anti-D typing sera
applicators or toothpicks
1
2
3
4
5
70% alcohol and cotton
disposable sterile lancets
Scrub the middle finger with a piece of cotton sat-
urated with 70% alcohol and pierce it with a ster-
ile disposable lancet. Place a small drop of blood
in each of three squares on the marked slides on
the warming box.
To get the proper proportion of serum to
blood, do not use a drop larger than 3 mm diame-
ter on the slide.
Add a drop of anti-D serum to the blood in the
anti-D square, mix with a toothpick, and note the
time. Only 2 minutes should be allowed for ag-
glutination.
Add a drop of anti-B serum to the anti-B square
and a drop of anti-A serum to the anti-A square.
Mix the sera and blood in both squares with sep-
arate fresh toothpicks.
Agitate the mixtures on the slide by slowly rock-
ing the box back and forth on its pivot. At the end
of 2 minutes, examine the anti-D square carefully
for agglutination. If no agglutination is apparent,
consider the blood to be Rh-negative. By this time
the ABO type can also be determined.
Record your results on Laboratory Report 87-89.
One drop of each antiserum is sufficient,
'anti-D:
beru
anti-B;
eru
.»'
&nti-Aii
rum
<.
V"
■ • :*tf
&8
: T -. r -'"\ r ■ l j ■ ''•' ...\ r-.. '.x •■:'. ■ ■■>,-' ^-.>-'->^:- - A<;
Whole blood or plasma-diluted
blood must be used for Rh
typing. Saline-diluted blood is
preferred for the ABO typing.
^
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c.
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■ +
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Agitation is acheived by slowly
rocking box back and forth for
2 minutes.
Figure 89 Blood typing with warming box
298
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XIV. Medical Microbiology
and Immunology
90. The Snyder Caries
Susceptibility Test
© The McGraw-H
Companies, 2001
The Snyder Caries Susceptibility Test
90
The degradation of enamel and dentin in the forma-
tion of tooth decay (dental caries) occurs as a result of
the production of lactic acid by bacteria
(Streptococcus mutans and others) in the presence of
high levels of sucrose. Of the various methods that
have been devised to determine one's susceptibility to
tooth decay, M. L. Snyder's caries susceptibility test
is a relatively simple test that has been shown to have
a fairly high reliability correlation.
This method relies on the rapidity of organisms in
saliva to lower the pH in a medium that contains 2% dex-
trose (Snyder test agar). Since decalcification of enamel
begins at a pH of 5.5, and progresses rapidly as the pH is
lowered to 4.4 and less, the demonstration of pH lower-
ing becomes evidence of susceptibility to caries.
To indicate the presence of acid production in the
medium, the indicator bromcresol green is incorpo-
rated in it. This indicator is green at pH 4.8 and be-
comes yellow at pH 4.4, remaining yellow below 4.4.
Figure 90. 1 illustrates the procedure that is used
in the Snyder caries susceptibility test. Note that 0.2
ml of saliva is added to a tube of liquefied Snyder test
agar (50° C) and mixed well by rotating the tube be-
tween the palms of both hands. After the medium has
solidified, the tube is incubated at 37° C for a period
of 24-72 hours. If the medium turns yellow in 24-48
hours, the individual is said to be susceptible to caries.
Although we will be performing this test only
once, it should be noted that test reliability is en-
hanced by performing the test on three consecutive
days at the same time each day. If the test is performed
right after toothbrushing, it is not as reliable as if 2 or
3 hours have elapsed after brushing. Proceed as fol-
lows to perform this test:
Materials:
1 tube of Snyder test agar (5 ml in 15 mm dia
tube)
1 30 ml sterile beaker
1 piece of paraffin (1/4" X 1/4" X 1/8")
1 ml pipette
1 gummed label
Liquefy a tube of Snyder test agar and cool it to
50° C.
After allowing a piece of paraffin to soften under
the tongue for a few minutes, start chewing it. Chew
it for 3 minutes, moving it from one side of the
1.
2.
0.2 ml
SALIVA
GREEN
(NEGATIVE)
37° C
24-72
HOURS
SALIVA AND MEDIUM
ARE MIXED
LIQUIFIED SNYDER
TEST AGAR— 50° C
YELLOW
(POSITIVE)
Figure 90.1 Snyder caries susceptibility test
299
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XIV. Medical Microbiology
and Immunology
90. The Snyder Caries
Susceptibility Test
© The McGraw-H
Companies, 2001
Exercise 90 • The Snyder Caries Susceptibility Test
3
4.
5
mouth to the other. Do not swallow the saliva. As it
accumulates, deposit it in the small sterile beaker.
Vigorously shake the sample in the beaker from side
to side for 30 seconds to disperse the organisms.
With a 1 ml pipette transfer 0.2 ml of saliva to the
tube of agar. Do not allow the pipette to touch the
side of the tube or agar.
Before the medium solidifies, mix the contents of
the tube by rotating the tube vigorously between
the palms of the hands.
6. Write your name on a gummed label and attach it
to the tube.
7. Incubate the tube at 37° C. Examine the tube
every 24 hours to see if the bromcresol green in-
dicator has changed to yellow. If it has, the test is
positive. The degree of caries susceptibility is de-
termined from the table below.
8. Record your results on the Laboratory Report.
Caries Susceptibility
Medium turns yellow in:
24 hours
48 hours
72 hours
Marked
Positive
Moderate
Negative
Positive
Slight
Negative
Negative
Positive
Negative
Negative
Negative
Negative
300
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Laboratory Reports
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Laboratory Report
Student:
1.2
Desk No.:
Ex. 1 Brightfield Microscopy
Section
A Completion Questions
Record the answers to the following questions in the column at the right.
1. List three fluids that may be used for cleaning lenses.
2. How can one greatly increase the bulb life on a microscope lamp if
voltage is variable?
3. What characteristic of a microscope enables one to switch from one
objective to another without altering the focus?
4. What effect (increase or decrease) does closing the diaphragm have
on the following?
a. Image brightness
b. Image contrast
c. Resolution
5. In general, at what position should the condenser be kept?
6. Express the maximum resolution of the compound microscope in
terms of micrometers (jim).
7. If you are getting 225 X magnification with a 45 X high-dry objec-
tive, what is the power of the eyepiece?
8. What is the magnification of objects observed through a 100X oil
immersion objective with a 7.5 X eyepiece?
9. Immersion oil must have the same refractive index as
of any value.
to be
10. Substage filters should be of a
color to get the
maximum resolution of the optical system.
B. True-False
Record these statements as true or false in the answer column.
1 . Eyepieces are of such simple construction that almost anyone can
safely disassemble them for cleaning.
2. Lenses can be safely cleaned with almost any kind of tissue or cloth.
3. When swinging the oil immersion objective into position after us-
ing high-dry, one should always increase the distance between the
lens and slide to prevent damaging the oil immersion lens.
4. Instead of starting first with the oil immersion lens, it is best to use
one of the lower magnifications first, and then swing the oil im-
mersion into position.
5. The 45 X and 100X objectives have shorter working distances than
the 10 X objective.
Answers
Completion
La_
b._
c._
2. .
3. .
4,a. _
b. _
c. _
6, .
7. .
8.
9, .
10. .
True- False
1. .
2. ,
3. .
4. ,
5. .
Benson: Microbiological
Back Matter
Laboratory Reports
©The McGraw-Hill
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Eighth Edition
Companies, 2001
Brightfield Microscopy
C. Multiple Choice
Select the best answer for the following statements
1 . The resolution of a microscope is increased by
1 . using blue light.
2. stopping down the diaphragm.
3. lowering the condenser.
4. raising the condenser to its highest point.
5 . B oth 1 and 4 are correct.
2.
3.
4.
5.
The magnification of an object seen through the 10X objective
with a 10 X ocular is
1. ten times.
2. twenty times.
3. 1000 times.
4. None of the above are correct.
The most commonly used ocular is
1. 5X.
2. 10X.
3. 15X.
4. 20X.
Microscope lenses may be cleaned with
1 . lens tissue.
a soft linen handkerchief.
2.
3.
4.
5.
an air syringe.
Both 1 and 3 are correct.
1,2, and 3 are correct.
Answers
Multiple Choice
1.
2.
3.
5.
■■'——-i i- ■■ ■■' — . - ■'■■ ■
When changing from low power to high power, it is generally necessary to
1 . lower the condenser.
2. open the diaphragm.
3. close the diaphragm.
4. Both 1 and 2 are correct.
5. Both 1 and 3 are correct.
Ex. 2 Darkfield Microscopy
A Questions
1 . What characteristic of living bacteria makes them easier to see with a darkfield condenser than with a
regular brightfield condenser?
2. If a darkfield condenser causes all light rays to bypass the objective, where does the light come from that
makes an object visible in a dark field?
3. What advantage does a cardioid condenser have over a star diaphragm?
302
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Laboratory Reports
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Laboratory Report student:
Desk No.: Section
Ex. 3 Phase- Contrast Microscopy
A Questions
1 . Which rays (direct or diffracted) are altered by the phase ring on the phase plate?
2. How much phase shift occurs in the light rays that emerge from a transparent object?
3. Differentiate:
Bright phase microscope:
Dark phase microscope:
4. List two items that can be used for observing the concentricity of the annulus and phase ring
a.
b.
Ex. 4 Fluorescence Microscopy
A Questions
1. Differentiate:
Phosphorescence
Fluorescence:
2. List three fluorochromes that are used in staining bacteria
a. b.
3. What are the two most serious hazards when using mercury vapor arc lamps?
a.
b.
4. What relationship exists between the wavelength of light and its energy?
Ex. 5 Microscopic Measurements
A. Questions
1 . What is the distance between each of the graduations on the stage micrometer? mm
2. Why must the entire calibration procedure be performed for each objective?
(See reverse page for more questions pertaining to Exercises 3 and 4.)
303
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Laboratory Reports
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Phase-Contrast and Fluroscence Microscopy
B. Multiple Choice Questions for Exercises 3 and 4
Select the answer that best completes the following statements.
1. If direct rays passing through an object are advanced % wavelength by the
phase ring, the diffracted rays are
1 . in phase with the direct rays.
Yi wavelength out of phase with the direct rays.
wavelength out of phase with direct rays,
in reverse phase with the direct rays.
B oth 2 and 4 are correct.
2.
3.
4.
5.
2. The best barrier filter to use with the Schott BG12 exciter filter is the
1. blueA0702. 3. Wratten G-2.
2. Schott OG1. 4. None of these are correct.
3. The examination of ordinary stained slides is not enhanced with a
1. brightfield microscope. 3. fluorescence microscope.
2. phase-contrast microscope. 4. Both 2 and 3 are correct.
4.
5.
6
7
8
9.
10.
Special immersion oil is required for
1 . brightfield microscopy.
2. phase-contrast microscopy.
3. fluorescence microscopy.
4. Both 2 and 3 are correct.
Amplitude summation occurs in phase-contrast optics when both direct and
diffracted rays are
1. in phase.
2. in reverse phase.
3. off X A wavelength.
4. None of these are correct.
The phase-contrast microscope is best suited for observing
1 . living organisms in an uncovered drop on a slide.
2. stained slides with cover glasses.
3. living organisms in hanging drop slide preparations.
4. living organisms on a slide with a cover glass.
The wavelength of fluorescent light rays is
1 . always longer than the exciting wavelength.
2. always shorter than the exciting wavelength.
3. about the same length as the exciting wavelength.
4. sometimes shorter and at other times longer in wavelength.
The barrier filter in the fluorescence microscope is kept in position to
1 . block all light rays from getting through.
2. allow fluorescence light rays to pass through.
3. screen out exciting light rays.
4. Both 2 and 3 are correct.
A phase centering telescope is used to
1 . improve the resolution of the ocular.
2. increase magnification with the oil immersion objective.
3. observe the relationship of the annular diaphragm to the phase ring.
4. None of the above are correct.
The visible spectrum of light is between
1. 200 and 800 nanometers.
2. 400 and 780 nanometers.
3. 300 and 800 nanometers.
4. None of these are correct.
Answers
Multiple Choice
2.
3.
4.
5-
6.
7.
8,
9.
10.
304
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Back Matter
Laboratory Reports
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Companies, 2001
Laboratory Report
Student:
Desk No.:
Section
Protozoa, Algae, and Cyanobacteria
A Tabulation of Observations
In this study of freshwater microorganisms, record your observations in the following tables. The number of
organisms to be identified will depend on the availability of time and materials. Your instructor will indicate
the number of each type that should be recorded.
Record the genus of each identifiable type. Also, indicate the phylum or division to which the organism
belongs. Microorganisms that you are unable to identify should be sketched in the space provided. It is not
necessary to draw those that are identified.
PROTOZOA
GENUS
PHYLUM
BOTTLE
NO.
SKETCHES OF UNIDENTIFIED
ALGAE
GENUS
DIVISION
BOTTLE
NO.
SKETCHES OF UNIDENTIFIED
305
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Laboratory Reports
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Companies, 2001
Protozoa, Algae, and Cyanobacteria
CYANOBACTERIA
GENUS
BOTTLE
NO.
SKETCHES OF UNIDENTIFIED
B. General Questions
Record the answers to the following questions in the answer column.
It may be necessary to consult your text or library references for one
or two of the answers.
1. Give the kingdom in which each of the following groups of or-
ganisms is found:
a. protozoans
b. algae
c. cyanobacteria
d. bacteria
e. fungi
f . microscopic invertebrates
2. Four kingdoms are represented by the organisms in the above
question. Name the fifth kingdom.
3. What is the most significant characteristic seen in eukaryotes that
is lacking in prokaryotes?
4. What characteristic in the microscopic invertebrates distinguishes
them from protozoans?
5. Which protozoan phylum was not found in pond samples because
phylum members are all parasitic?
6. Indicate whether the following are present or absent in the algae:
a. cilia
b. flagella
c. chloroplasts
7. Indicate whether the following are present or absent in the proto
zoans:
a. cilia
b. chloroplasts
c. mitochondria
d. mitosis
8. Which photosynthetic pigment is common to all algae and cyano-
bacteria?
9. Name two photosynthetic pigments that are found in the cyanobac-
teria but not in the algae.
10. What photosynthetic pigment is found in bacteria but is lacking in
all other photosynthetic organisms?
11. What type of movement is exhibited by the diatoms?
1.a.
Answers
tL
r„
d.
e.
f.
2.
3.
4.
5.
6_a,
h.
c.
7 .a.
b.
r..
d.
8.
9.a.
h.
10.
11.
306
Benson: Microbiological
Back Matter
Laboratory Reports
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Companies, 2001
Protozoa, Algae, and Cyanobacteria
C. Protozoan Characterization
Select the protozoan groups in the right-hand column that have the following
characteristics:
1 . move with flagella
2. move with cilia
3. move with pseudopodia
4. have nuclear membranes
5 . lack nuclear membranes
6. all species are parasitic
7. produce resistant cysts
1. Sarcodina
2. Mastigophora
3. Ciliophora
4. Sporozoa
5. all of above
6. none of above
D. Characterization of Algae and Cyanobacteria
Select the groups in the right-hand column that have the following characteristics
Pigments
1 . chlorophyll a
2. chlorophyll b
3. chlorophyll c
4. fucoxanthin
5. c-phycocyanin
6. c-phycoerythrin
Food Storage
7. fats
8. oils
9. starches
10. laminarin
11. leucosin
12. paramylum
13. mannitol
Other Structures
14. pellicle, no cell wall
15. cell walls, box and lid
16. chloroplasts
17. phycobilisomes
18. thylakoids
1. Euglenophycophyta
2. Chlorophycophyta
3. Chrysophycophyta
4. Phaeophycophyta
5. Pyrrophycophyta
6. Cyanobacteria
7. all of above
8. none of above
1.
2.
5.
7.
1.
4.
6.
7.
8.
9.
10.
11.
12.
13.
14.
15.
16.
17.
18.
Answers
Protozoa
3.„
l¥¥*¥TPrt-»«l-Wl
Algae
1-%-i ■■ <■ l , W • — "■-
307
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
Back Matter
Laboratory Reports
© The McGraw-H
Companies, 2001
Benson: Microbiological
Back Matter
Laboratory Reports
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Companies, 2001
Laboratory Report student:
y
Desk No.: Section
Ex. 8 Aseptic Technique
A. Results
1. Were all your transfers successful?
2. What evidence do you have that they were successful?
3. What evidence do you have that a transfer is unsuccessful?
B. Questions:
1 . What kinds of organisms are destroyed when your desktop is scrubbed down with a disinfectant?
2. Are bacterial endospores destroyed?
3. How hot should inoculating loops and needles be heated?
4. Why is it necessary to flame the mouth of the tube before and after performing an inoculation?
Ex. 9 The Bacteria
A Questions: After you have tabulated your results on the back of this sheet, answer the following
questions:
1 . Using the number of colonies as an indicator, which habitat sampled by the class appears to be the most
contaminated one?
2. Why do you suppose this habitat contains such a high microbial count?
3. a. Were any plates completely lacking in colonies?
b. Do you think that the habitat sampled was really sterile?
c. If your answer to b is no, then how can you account for the lack of growth on the plate?
d. If your answer to b is yes, defend it:
4. In a few words describe some differences in the macroscopic appearance of bacteria and mold colonies
309
Benson: Microbiological
Back Matter
Laboratory Reports
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Companies, 2001
Bacteria
B. Tabulation
After examining your TS A and blood agar plates, record your results in the following table and on a similar
table that your instructor has drawn on the chalkboard. With respect to the plates, we are concerned with a
quantitative evaluation of the degree of contamination and differentiation as to whether the organisms are
bacteria or molds. Quantify your recording as follows:
no growth
+ 1 to 1 colonies
+ + 11 to 50 colonies
+ + + 51 to 100 colonies
+ + + + over 100 colonies
After shaking the tube of broth to disperse the organisms, look for cloudiness (turbidity). If the broth is clear,
no bacterial growth occurred. Record no growth as 0. If tube is turbid, record + in last column.
^■^^■^^"^^
STUDENT
INITIALS
i m iiiaiiiii Jii n i a
PLATE EXPOSURE METHOD
TSA
-
w
i "i h i q ■ 1 1 !■ i ■
^^Fm^^i^^h^
■ ■■■■I III
Blood Agar
COLONY COUNTS
Bacteria
Mold
■■■■■■■■■■
BROTH
Source
I ■ I III!
1 1 I I I M-T^d 1 ^ M h WW
^^mi^
Result
WB>»>>»*ta-B>«-B>»>B
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310
Benson: Microbiological
Back Matter
Laboratory Reports
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Companies, 2001
Laboratory Report
Student:
10
Desk No.:
The Fungi:
Yeasts and Molds
Section
A Yeast Study
Draw a few representative cells of Saccharomyces cerevisiae in the appropriate circles below. Blastospores
(buds) and ascospores, if seen, should be shown and labeled.
Prepared Slide
Living Cells
B. Mold Study
In the following table, list the genera of molds identified in this exercise. Under colony description, give the
approximate diameter of the colony, its topside color and backside (bottom) color. For microscopic appear-
ance, make a sketch of the organism as it appears on slide preparation.
GENUS
1 1 1 1^^^ " ■ i ^^^^^
^*i* w
COLONY DESCRIPTION
MICROSCOPIC APPEARANCE
(DRAWING)
■ I' i ■ i ■ .■ j^i-i ■
i ' m
*A«ltHHM<WWfP^<^
*AMA*riMt«t«
alBH^^riri
^^^^^*P
311
Benson: Microbiological
Back Matter
Laboratory Reports
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Companies, 2001
Fungi: Yeasts and Molds
C. Questions
Record the answers for the following questions in the answer column.
1. The science that is concerned with the study of fungi is called
2. The kingdom to which the fungi belong is
3. Microscopic filaments of molds are called
4. A filamentlike structure formed by a yeast from a chain of blas-
tospores is called a .
5. A mass of mold filaments, as observed by the naked eye, is called
a
6. Most molds have
septate).
hyphae (septate or non-
7. List three kinds of sexual spores that are the basis for classifying
the molds.
8. What is the name of the rootlike structure that is seen in Rhizopus ?
9. What type of hypha is seen in Mucor and Rhizopus?
10. What kind of asexual spores are seen in Mucor and Rhizopus?
11. What kind of asexual spores are seen in Penicillium?
12. What kind of asexual spores are seen in Alternaria?
13. Which subdivision of the Amastigomycota contains individuals
that lack sexual spores?
14. What division of Myceteae consists of slime molds?
15. Fungi that exist both as yeasts and molds are said to be .
1.
10.
11.
12.
13.
14.
15.
Answers
2.
3.
4-
5.
6.
7_a.
b.
c.
8.
9.
— 1^1--
rww ■ ■ ■ ■ m ■ t i
312
Benson: Microbiological
Back Matter
Laboratory Reports
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Companies, 2001
Laboratory Report
Student:
11
u
Desk No.:
Section
Negative Staining, Smear Preparation,
Simple and Capsular Staining
A Negative Staining (Exercise 1 1)
1
2
Drawing: Make a drawing in the circle at the right of some of the
organisms as seen under oil immersion.
In addition to nigrosine, what other agent is often used for making
negative- stained slides?
3. Other than bacteria, what kinds of microorganisms might one en-
counter in the mouth?
B. Smear Preparation (Exercise 12)
1 . Give two reasons for heating the slide after the smear is air-dried
a.
b.
Oral organisms
(nigrosine)
2. Why is an inoculating needle preferred to a wire loop when making smears from solid media?
C. Simple Staining (Exercise 13)
1
Drawing: Draw a few cells of C. diphtheriae from the portion of
the slide that exhibits metachromatic granules and palisade
arrangement.
2. Why are basic dyes more successful on bacteria than acidic dyes?
3. List three basic dyes that are used to stain bacteria: a
Corynebacterium diphtheriae
b.
,, c.
313
Benson: Microbiological
Back Matter
Laboratory Reports
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Negative Staining, Smear Preparation, Simple and Capsular Staining
D. Capsular Staining (Exercise 14)
1 . List some of the chemical substances that have been identified in
bacterial capsules.
2. What relationship is there between capsules and bacterial virulence?
Klebsiella pneumoniae
(capsular stain)
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Desk No.:
Section
Differential Staining:
Gram, Spore, and Acid- Fast
A Drawings
With colored pencils, draw the various organisms as seen under oil immersion. Extra circles are provided for
additional assignments, if needed.
P. aeruginosa & S. aureus
(Gram stain)
B. megaterium & M. B. catarrhalis
(Gram stain)
M. smegmatis
(Gram stain)
B. megaterium
(Schaeffer-Fulton method)
B. megaterium
(Dorner method)
M. smegmatis & S. aureus
(Ziehl-Neelsen method)
M. phlei & S. aureus
(Truant method)
OPTIONAL STAINING
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B. Completion Questions
1. What color would you expect S. aureus to be if the iodine step were omitted in the Gram staining
procedure?
Explanation
2. What part of the bacterial cell (cell wall or protoplast) appears to play the most important role in deter-
mining whether an organism is gram-positive?
3. Why would methylene blue not work just as well as safranin for counterstaining in the Gram staining
procedure?
4. Why are endo spores so difficult to stain?
5. How do the following two genera of spore- formers differ physiologically?
Bacillus:
Clostridium:
6. How do you differentiate S. aureus and M. B. catarrhalis from each other on the basis of morphological
characteristics?
7. Are the acid-fast mycobacteria gram-positive or gram-negative?
8. For what two diseases is acid- fast staining of paramount importance?
a.
b.
9. What advantage does the Ziehl-Neelsen technique have over the Truant method?
10. What advantage does the Truant method have over the Ziehl-Neelsen technique?
11. Why is it desirable to combine S. aureus with acid-fast organisms such as M. smegmatis when applying
an acid-fast staining technique?
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Desk No.:
Motility Determination
Section
A. Test Results
1. Which of the two organisms exhibited true motility on the slides?
2. Did the semisolid medium inoculations confirm the results obtained from the slides?
3. Sketch in the appearance of the two tube inoculations:
Micrococcus luteus
Proteus vulgaris
B. Questions
1 . How does Brownian movement differ from true motility?
2. How do you differentiate water current movement from true motility?.
3. Make sketches that illustrate each of the following flagellar arrangements:
Monotrichic
Lophotrichic
Amphitrichic
Peritrichic
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Desk No.: Section
Culture Media Preparation
1. How do the following types of organisms differ in their carbon needs?
Photoautotrophs :
Photoheterotrophs :
2. Where do the above two types of organisms get their energy?
3. Where do chemoheterotrophs get their energy?
4. What is a growth factor?
5. Give two reasons why agar is such a good ingredient for converting liquid media to solid media
a.
b.
6. Differentiate between the following two types of media
Synthetic medium :
Nonsynthetic medium:
7. Differentiate between the following two types of media
Selective medium:
Differential medium:
8. Briefly, list the steps that you would go through to make up a batch of nutrient agar slants
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Desk No.:
Section
Pure Culture Techniques
A Evaluation of Streak Plate
Show within the circle the distribution of the colonies on your streak plate. To identify the colonies, use red
for Serratia marcescens, yellow for Micrococcus luteus, and purple for Chromobacterium violaceum. If time
permits, your instructor may inspect your plate and enter a grade where indicated.
Grade
B. Evaluation of Pour Plates
Show the distribution of colonies on plates II and III, using only the quadrant section for plate II. If plate III
has too many colonies, follow the same procedure. Use colors.
plate II
plate III
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Pure Culture Techniques
C. Subculture Evaluation
With colored pencils, sketch the appearance of the growth on the slant diagrams below. Also, draw a few cells
of each organism as revealed by Gram staining in the adjacent circle.
Serratia marcescens
D. Questions
Micrococcus luteus
or
Chromobacterium violaceum
Escherichia coli
1 . Which method of separating organisms seems to achieve the best separation?
2. Which method requires the greatest skill?
3. Do you think you have pure cultures of each organism on the slants?
Can you be absolutely sure by studying its microscopic appearance?
Explain:
4. Give two reasons why the nutrient agar must be cooled to 50° C before inoculating and pouring
5. Why should a Petri plate be discarded if media is splashed up the side to the top?
6. Give two reasons why it is important to invert plates during incubation
7. Why is it important not to dig into the agar with the loop?
8. Why must the loop be flamed before entering a culture?
Why must it be flamed after making an inoculation?
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Desk No.:
Cultivation of Anaerobes
Section
A Tube Inoculations
After carefully comparing the appearance of the six cultures belonging to you and your laboratory partner,
select the best tube for each organism and sketch its appearance in the tubes below. Indicate under each name
the type of medium (FTM or TGYA).
E. coli
( )
S. faecalis
S. aureus
( )
B. subtilis
( >
C. sporogenes
( )
C. rubrum
( )
B. Plate Inoculations
After comparing the growths on the two plates of Brewer's anaerobic agar with the growths in the six tubes,
classify each organism as to its oxygen requirements:
Escherichia coli:
Bacillus subtilis:
Streptococcus faecalis:
Staphylococcus aureus:
Clostridium sporogenes
Clostridium rubrum:
C. Questions
1 . What is the function of oxygen at the cellular level?
2. Why are facultative organisms able to grow with or without oxygen while aerobes grow only in its
presence?
3. How do "indifferents" differ from "facultatives"?
4. What is the function of the following agents in the media used in this experiment?
Sodium thiogly collate:
Resazurin:
Agar in FTM:
Agar in TGYA shake
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5. How is oxygen removed from the air in a GasPak anaerobic jar?
D. Spore Study
If a spore-stained slide is made of the three spore-formers, draw a few cells of each organism in the spaces
provided below:
B. subtilis
C. rubrum
C. sp or o genes
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Desk No.:
Section
Bacterial Population Counts
A. Quantitative Plating Method
1 . Record your plate counts in this table
DILUTION PLATED
ML PLATED
NUMBER OF COLONIES
1:10,000
1.0
1:100,000
0.1
1:1,000,000
1.0
1:10,000,000
0.1
2. How many cells per ml were there in the undiluted culture?
3. How would you inoculate a plate to get 1 : 100 dilution?
4. How would you inoculate a plate to get 1:10 dilution?
5. Give two reasons why it is necessary to shake the water blanks as recommended
a.
b.
B. Turbidimetric Determinations
1 . Record the percent transmittance and optical density values for your dilutions in the following table
DILUTION
PERCENT TRANSMITTANCE
OPTICAL DENSITY
1:1
1:2
1:4
1:8
1:16
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Bacterial Population Counts
2. Plot the optical densities versus the concentration of organisms. Complete the graph by drawing a line
between plot points.
OPTICAL DENSITY
>
II
II
II
II
u
1:16 1:8 1:4 1:2 1:1
DILUTION
C. Questions
1. What is the maximum O.D. that is within the linear portion of the curve?
2
What is the corrected or true O.D. of the undiluted culture? (Hint: If the O.D. for the 1 :2 dilution but not
the 1:1 dilution is within the linear portion of the curve, then the O.D. of the 1:1 dilution should not be
considered correct. The correct or true O.D. of the undiluted culture in this example could be estimated
by multiplying the O.D. of the 1:2 dilution by 2.)
3. What is the correlation between corrected O.D. and cell number for your culture?
4. Why is it necessary to perform a plate count in conjunction with the turbidimetry procedure?
5. If your medium were pale blue instead of amber-colored, as is the case of nutrient broth, would you set
the wavelength control knob higher or lower than 686 nanometers?
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25
Desk No.:
Ex. 24 Slide Culture
Autotrophs
Section
A Microscopic Examination
While examining the two slides, move them around to different areas to note the various types of organisms
that are present. Draw representative types.
GRAM'S STAIN
LIVING
B. Questions
1. With respect to Gram's stain, which type (gram-positive or gram-negative) seems to predominate?
2. List as many different kinds of autotrophic protists as you can that can be cultured on this type of slide
3. Some organisms that grow on this type of slide are chemo synthetic heterotrophs. What would be the
source of their food?
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Slime Mold Culture
Ex. 25 Slime Mold Culture
A Observations
1 . What happened when the flow of protoplasm on a Plasmodium was interrupted by severance with a
scalpel?
2. Describe your observations of the crushed spores on the hanging drop slide.
B. Questions
1 . List two functions served by fructification (sporangia formation) in Physarum.
a.
b.
2. What is the principal function of the plasmodial stage of Physarum?
3. List two characteristics that the Myxobacterales and Gymnomycota have in common,
a.
b.
4. Postulate as to the evolutionary relationship between the myxobacteria and slime molds
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Desk No.: Section
Anaerobic Phototrophic Bacteria:
Isolation and Culture
A Observations:
1 . Column Appearance: Describe in a few words the appearance of the Winogradsky column during the
First Period:
One Week Later:
Two Weeks Later:
Subsequent Weeks
2. Subculture Appearance: Describe the appearance of your subculture tubes at the end of two weeks:
3 . Microscopic Appearance : Describe the characteristics of the cells you observed on wet mount slides and
gram- stained slides:
B. Questions:
1 . What roles do the following organisms perform in the Winogradsky column?
Clostridium:
Desulfovibrio :
2. How do Chromatium and Chlorobium differ as to where they put the sulfur that they produce?
3. Give the equations for photosynthesis in the following
Algae:
Chromatium and Chlorobium:
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Anaerobic Phototrophic Bacteria: Isolation and Culture
C. Tabulation of Results
If different mud samples were used by class members, record your results on the chart below and on a simi
lar chart constructed by your instructor on the chalkboard. Record the presence (+) or absence (— ) of the var
ious species identified. In the "Other" column list any other species that you might have encountered.
^>p^^w«p«Hfwin
Student
Initials
Source of
Pond Mud
— lfcfc ^ — ^^^^ ml ^^^^
Chromatium
^™^^^^^"^W^^^*^^"»*»^WWWta*l
Chlorobium
^^^H*
WWHtali
«*M
Other
v-i
■'^^
WWWWPnVtta^ta^^H^^
■■■ '!■■■■ *»'!■■
^i^Mia
^i^VWhMtaW^i
"■
■
MMUMU^
ri*^M^^
wn-n
^^^+m
^^^m^m
^^^^^V^^^ABMaBafe^
V^nMrHaaBBBBBBi
UH^i^
PWUrftfi^ta
*m
h^hhu
WW
■■■■■■p
iU^AWhU
^i^P¥M^^*fi IBP
■■■['^■■■hM
^^^WWI
D. Conclusions from above table,
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29
Desk No.:
Section
Isolation of Phage from Sewage and Flies
A Plaque Size Increase (for both Exercises 28 and 29)
With a china marking pencil, circle and label three plaques on one of the plates and record their sizes in mil
limeters at 1-hour intervals.
1
TIME
PLAQUE SIZE
(millimeters)
Plaque No. 1
Plaque No. 2
Plaque No. 3
When first seen
1 hour later
i
2 hours later
i
3 hours later
B. Questions (for Exercise 28 only)
1 . Were any plaques seen on the negative control plate?
2. Do plates 1 , 2, and 3 show a progressive increase in number of plaques with increased amount of sewage
filtrate?
3. Did the phage completely "wipe out" all bacterial growth on any of the plates?
If so, which plates?
C. Observations (for Exercise 29 only)
Count all the plaques on each plate and record the counts in the following table. If the plaques are very nu-
merous, use a Quebec colony counter and hand counting device. If this exercise was performed as a class
project with individual students doing only one or two plates from a common fly-broth filtrate, record all
counts on the chalkboard on a table similar to the one below.
Plate Number
1
2
3
4
5
6
7
8
9
10
E, coii (ml)
0.9
0.3
QJ
0.6
0.5
0.4
0.3
0.2
0.1
1.0
Filtrate (ml)
0.1
0.2
0.3
0.4
0.5
0.6
0.7
0.8
0.9
Number of plaques
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Isolation of Phage from Sewage and Flies
D. Questions (for Exercise 29 only)
1 . Which plate was used as the negative control?
2. Were there any plaques on the negative control plate?
3. What would be the explanation for the presence of plaques on the negative control plate?
4. Were any plates completely "wiped out" by phage action?
If so, which ones?
E. Terminology (for both Exercises 28 and 29)
1. Differentiate between the following:
Lysis:
Lysogeny:
2. Differentiate between the following
Virulent phage :
Temperate phage:
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Desk No.:
Section
Burst Size Determination:
A One- Step Growth Curve
A Plaque Counts
Record the counts of plaques on each of the plates in the following table. Record the peak number of plaques
as the burst size. The drop in plaque numbers after a peak results from adsorption of mature phage virions on
other bacterial cells and cell debris.
15
25
30
35
40
45
50
Burst size:
B. Dilution Interpretation
Answer the following questions to clarify your understanding of the dilutions that occur in this experiment.
1 . How many cells were present in each milliliter of the original bacterial culture?
2. How many bacterial cells (total) were dispensed into the ADS tube?
3 . If the bacterial dilution per plate is 1 : 1 0,000,000, how many bacterial cells were distributed to each plate?
4. How many phage virions were present in 1 ml of the original phage suspension?
5. How many phage virions were present in the 0. 1 ml of phage suspension that was added to the ADS tube?
6. What was the numerical ratio of phage virions to bacterial cells in the ADS tube?.
What is this ratio called?
7. How many bacterial cells were placed in the ADS-2 tube?
8. What effect does dilution have on adsorption?
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Desk No.:
Section
Microbial Interrelationships
Ex. 31 Bacterial Commensalism
A. Results
Indicate the degree of turbidity (none, +, + +, + + +)in the following table. With colored pencils, draw
the appearance of the gram-stained slides where indicated.
ORGANISMS
TURBIDITY
GRAM STAIN
Staphylococcus aureus
Clostridium sporogenes
S. aureus and C. sporogenes
B. Questions
1 . Does C. sporogenes grow well in nutrient broth?
2. How does S. aureus assist C. sporogenes in growth?
Ex. 32 Bacterial Synergism
A. Results
Examine the six tubes of media, looking for acid and gas. In the presence of acid, bromthymol blue turns
yellow. Record your results in the table below. Consult other students for their results and complete the
table.
INDIVIDUAL
ORGANISMS
LACTOSE
SUCROSE
COMBINATIONS
LACTOSE
SUCROSE
Acid
Gas
Acid
Gas
Acid
Gas
Acid
Gas
E. coli
E. coli and P. vulgaris
P. vulgaris
E. coil and S. aureus
i
S, aureus
S. aureus and P. vulgaris
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Bacterial Synergism and Microbial Antagonism
B. Questions
1 . Did any of the three organisms produce gas in either lactose or sucrose broth when alone in the medium?
2. Which organisms act synergistically to produce gas in
lactose?
sucrose?
Ex. 33 Microbial Antagonism
A Questions
1 . Which organisms are antagonistic to E. coli ?
2. Which organisms are antagonistic to S. aureus?
3. Name an antibiotic substance that is derived from each of the following types of organisms. Also, indi
cate whether the substance is effective against gram-positive or gram-negative organisms.
A bacterium:
An actinomyces:
A fungus:
4. What role does microbial antagonism play in nature?
5. In what physiological way does penicillin affect penicillin-sensitive bacteria?
6. How do sulfonamides inhibit bacterial growth?
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Desk No
Temperature:
Effects on Growth
Section:
A Pigment Formation and Temperature
1
2
3
Draw the appearance of the growth of Serratia marcescens on the
nutrient agar slants using colored pencils.
Which temperature seems to be closest to the optimum temperature
for pigment formation?
What are the cellular substances that control pigment formation and
are regulated by temperature?
25° C
38° C
B. Growth Rate and Temperature
If a spectrophotometer is available, dispense the cultures into labeled cuvettes and determine the percent
transmittance of each culture. Calculate the O.D. values from the percent transmittance, using the formula
given in Exercise 23.
If no spectrophotometer is available, record only the visual reading as +, + +,+ + + , and none.
r ' 1
Temp.
SERRATIA MARCESCENS
ESCHERICHIA COU
■
Visual
Reading
Spectrophotometer
Visual
Reading
Spectrophotometer
%T
O.D.
%T
O.D.
5
i
25
38
i
42
i
i
55
i
i
i
i
i
1
1
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Temperature: Effects on Growth
Growth curves of Serratia marcescens and Escherichia coli as related to temperature
c
o
Q
o
Q.
o
5
o
25
o
38
o
42
o
55
o
Temperature (Centigrade)
1 . On the basis of the above graph, estimate the optimum growth temperature of the two organisms
Serratia marcescens:
Escherichia coli:
2. To get more precise results for the above graph, what would you do?
3. Differentiate between the following
Thermophile:
Mesophile:
Psychrophile:
4. What is the optimum growth temperature range for most psychrophiles?
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Desk No.:
Section
Temperature:
Lethal Effects
A. Tabulation of Results
Examine your five Petri plates, looking for evidence of growth. Record on the chalkboard, using a chart sim-
ilar to the one below, the presence or absence of growth as (+) or (— ). When all members of the class have
recorded their results, complete this chart.
ORGANISM
60° C
70° C
80° C
90° C
100° c
C*
10
20
30
40
C*
10
20
30
40
C*
10
20
30
40
C*
10
20
30
40
c*
10
20
30
40
S. aureus
E. coli
B. megaterium
C = control tube
1 . If they can be determined from the above information, record the thermal death point for each of the
organisms.
S. aureus:
E. coli:
B. megaterium:
2. From the following table, determine the thermal death time for each organism at the tabulated temperatures
ORGANISM
THERMAL DEATH TIME
60° C
70° C
80° C
90° C
100°C
S. aureus
E. coli
B. megaterium
B. Questions
1 . Give three reasons why endospores are much more resistant to heat than are vegetative cells
a.
b.
c.
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2. Differentiate between the following
Thermoduric:
Thermophilic:
3. List four diseases caused by spore-forming bacteria,
a. b.
c. d.
4. Since boiling water is unreliable in destroying endospores, how should one use heat in medical applica
tions to ensure spore destruction? (three ways)
a.
b.
c.
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Desk No.:
Section
pH and Microbial Growth
A. Tabulation of Results
If a spectrophotometer is available, dispense the cultures into labeled cuvettes and determine the percent
transmittance of each culture. Calculate the O.D. values from the percent transmittance, using the formula
given in Exercise 23. To complete the tables, get the results of the other three organisms from other members
of the class, and delete the substitution organisms in the tables that were not used.
If no spectrophotometer is available, record only the visual reading as +, + +,+ + +, and none.
PH
Escherichia coli
Staphylococcus aureus
Visual
Reading
Spectrophotometer
Visual
Reading
Spectrophotometer
%T
O.D.
%T
O.D
3
5
7
8
9
10
PH
Alcaligenes faecalis or
Sporosarcina ureae
Saccharomyces cervisiae or
Candida glabrata
Visual
Reading
Spectrophotometer
Visual
Reading
Spectrophotometer
%T
O.D
%T
O.D
3
5
7
8
9
10
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pH and Microbial Growth
B. Growth Curves
Once you have computed all the O.D. values on the two tables, plot them on the following graph. Use dif-
ferent colored lines for each species.
Q
ft
O
pH
3
5 7
Hydrogen Ion Concentration
8
9
10
C. Questions
1 . Which organism seems to grow best in acid media?
2. Which organism seems to grow best in alkaline media?
3. Which organism seems to tolerate the broadest pH range?.
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Desk No.:
Section:
Ex. 37 Osmotic Pressure and Bacterial Growth
A. Results
Record the amount of growth of each organism at the different salt concentrations, using, + , + + , + + + , and
none to indicate degree of growth.
ORGANISM
SODIUM CHLORIDE CONCENTRATION
0.5%
5%
10%
15%
48 hr
■
96 hr
48 hr
! 96 hr
48 hr
96 hr
48 hr
96 hr
Escherichia colt
Staphylococcus aureus
.
Haiobacterium salinarium
■
B. Questions
1 . Evaluate the salt tolerance of the above organisms.
Tolerates very little salt:
Tolerates a broad range of salt concentration:
Grows only in the presence of high salt concentration:
2. How would you classify Haiobacterium salinarium as to salt needs? Check one
Obligate halophile
Facultative halophile
3. Differentiate between the following:
Halophile:
Osmophile:
4. Supply the following information concerning mannitol salt agar (Refer to the Difco Manual)
Composition:
For what organism is this medium selective?
What ingredient makes it selective?
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Oligodynamic Action
Ex. 38 Oligodynamic Action
A. Tabulation of Results
Measure the zone of inhibition from the edge of each piece of metal with a millimeter ruler. Record the mea
surements in the table. Spaces are provided for write-in of additional metals.
METAL
MILLIMETERS OF INHIBITION
E. cofi
S. aureus
Copper
Silver
Aluminum
B. Questions
1 . Which metal seems to exhibit the greatest amount of oligodynamic action?
2. Which metal or metals seem to be ineffective?
3. Do these two organisms seem to differ in their susceptibility to oligodynamic action?
Explain:
4. What specific chemical substances in bacterial cells are inactivated by heavy metals, affecting growth?
344
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39
Desk No.:
Section
Ultraviolet Light:
Lethal Effects
A. Tabulation of Results
Your instructor will construct a table similar to the one below on the chalkboard for you to record your re-
sults. If substantial growth is present in the exposed area, record your results as + + +. If three or fewer
colonies survived, record + . Moderate survival should be indicated as + + . No growth should be recorded
as — . Record all information in the table.
ORGANISMS
EXPOSURE TIMES
S. aureus
10 sec
20 sec
40 sec
80 sec
2.5 min
5 min
10 min
• 20 min
Survival
._
■
B. megaterium
1 min
2 min
4 min
8 mtn
15 min
30 min
60 min
*6 min
Survival
■
.....
,
*Plates covered during exposure.
B. Questions
1 . What length of time is required for the destruction of non-spore-forming bacteria such as Staphylococcus
aureus?
2. Can you express, quantitatively, how much more resistant B. megaterium spores are to ultraviolet light
than S. aureus vegetative cells (i.e., how many times more resistant are they)?
3. Why is it desirable to remove the cover from the Petri dish when making exposures?
4. In what specific way does ultraviolet light destroy microorganisms?
5. What adverse effect can result from overexposure of human tissues to ultraviolet light?
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Ultraviolet Light: Lethal Effects
6. What wavelength of ultraviolet is most germicidal?
7. List several practical applications of ultraviolet light to microbial control
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40
Desk No.:
Section
Evaluation of Disinfectants
The Use-Dilution Method
A. Tabulation of Results
The instructor will draw a table on the chalkboard similar to the one below. Examine your tubes of nutrient
broth and pins by shaking them and looking for growth (turbidity). If you are doubtful as to whether growth
is present, compare the tubes with a tube of sterile nutrient broth. Record on the chalkboard a plus (+) sign
if growth is present and a minus (— ) sign if no growth is visible. After all students have recorded their results,
complete the following chart.
DISINFECTANT
MINUTES
Staphylococcus aureus
Bacillus megaterium
1 :750 Zephiran
Substitution
, ... t
1
5
10
30
60
C*
1
5
10
30
60
■
■
i
5% phenol
■
i
8% formaldehyde
■
■
■
C = control tube
B. Questions
1 . What conclusions can be drawn from this experiment?
2. Distinguish between the following
Disinfectant :
Antiseptic:
347
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Evaluation of Disinfectants: The Use-Dilution Method
3. What factors other than time influence the action of a chemical agent on bacteria?
4. Fill in the equation that explains how the phenol coefficient is determined
RC. =
5. What are some drawbacks that one encounters when attempting to apply the phenol coefficient to all dis
infectants?
348
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Student:
41
Desk No.:
Section
Evaluation of Alcohol:
Its Effectiveness as a Skin Degerming Agent
A. Tabulation of Results
Count the number of colonies that appear on each of the thumbprints and record them in the following table
If the number of colonies has increased in the second press, record a in percent reduction. Calculate the per-
centages of reduction and record these data in the appropriate column. Use this formula:
Percent Reduction =
(Colony Count 1st press) — (Colony Count 2nd press)
(Colony count 1st press)
X 100
LEFT THUMB (Control)
RIGHT THUMB (Dipped)
RIGHT THUMB (Swabbed)
Colony
Count
1st Press
Colony
Count
2nd Press
Percent
Reduction
Colony
Count
1st Press
Colony
Count
2nd Press
Percent
Reduction
I
Colony
Count
1st Press
Colony
Count
2nd Press
Percent
Reduction
i
i
i
i
■
I — — ■ ■"* *""!
I r-rr^ . -' ' -^
■■
Av. % Reduction, Left (C)
Av. % Reduction, Right (D)
Av. % Reduction, Right (S)
:
349
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Evaluation of Alcohol: Its Effectiveness as a Skin Degerming Agent
B. Questions
1. In general, what effect does alcohol have on the level of skin contaminants?
2. Is there any difference between the effects of dipping versus swabbing?
Which method appears to be more effective?
3. There is definitely survival of some microorganisms even after alcohol treatment. Without staining or mi-
croscopic scrutiny, predict what types of microbes are growing on the medium where you made the right
thumb impression after treatment.
350
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Laboratory Report
Student:
42
Desk No.:
Section
Evaluation of Antiseptics:
The Filter Paper Disk Method
A. Tabulation of Results
With a millimeter scale, measure the zones of inhibition between the edge of the filter paper disk and the or-
ganisms. Record this information. Exchange your plates with other students' plates to complete the mea
surements for all chemical agents.
DISINFECTANT
MILLIMETERS OF INHIBITION
Staphylococcus aureus
Pseudomonas aeruginosa
5% phenol
5% formaldehyde
5% iodine
B. Questions
1 . What conclusions can be derived from these results?
2. What factors influence the size of the zone of inhibition?
351
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Benson: Microbiological
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Laboratory Report
Student:
43
Desk No.:
Section
Antimicrobic Sensitivity Testing:
The Kirby- Bauer Method
A. Tabulation
List the antimicrobics that were used for each organism. Consult tables 43.1 and 43.2 to identify the various
disks. After measuring and recording the zone diameters, consult table VII in Appendix A for interpretation.
Record the degrees of sensitivity (R, I, or S) in the sensitivity column. Exchange data with other class mem-
bers to complete the entire chart.
ANTIMICROBIC
ZONE
DIA.
RATING
(R, 1, S)
ANTIMICROBIC
ZONE
DIA.
RATING
(R, I, S)
S. aureus
P. aeruginosa
:
Proteus vulgaris
i
;
1
i
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Antimicrobic Sensitivity Testing: The Kirby-Bauer Method
B. Questions
1. Which antimicrobics would be suitable for the control of the following organisms?
S. aureus:
E. coli:
P. vulgaris:
P. aeruginosa:
2. Differentiate between the following
Narrow spectrum antibiotic:
Broad spectrum antibiotic:
3 . Which antimicrobics used in this experiment would qualify as being excellent broad spectrum antimicrobics?
4. Differentiate between the following
Antibiotic:
Antimicrobic:
354
5. How can drug resistance in microorganisms be circumvented?
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Laboratory Report
Student:
44
Desk No.:
Section
Effectiveness of Hand Scrubbing
A Tabulation of Results
The instructor will draw a table on the chalkboard similar to the one below. Examine the six plates that your
group inoculated from the basin of water. Select the two plates of a specific dilution that have approximately
30 to 300 colonies and count all of the colonies of each plate with a Quebec colony counter. Record the counts
for each plate and their averages on the chalkboard. Once all the groups have recorded their counts, record
the dilution factors for each group in the proper column. To calculate the organisms per milliliter multiply the
average count by the dilution factor.
GROUP
0.1 ml COUNT
0.2 ml COUNT
0.4 ml COUNT
DILUTION
FACTOR*
ORGANISMS
PER
MILLILITER
Per
Plate
Average
Per
Plate
Average
Per
Plate
Average
A
B
C
■
D
i
E
•Dilution factors: 0.1 ml = 10; 0.2 ml = 5; 0.4 ml = 2.5
355
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Effectiveness of Hand Scrubbing
B. Graph
After you have completed this tabulation, plot the number of organisms per milliliter that were present in each
basin.
MINUTES:
BASIN:
1
A
3
B
6
C
9
D
12
E
C. Questions
1 . What conclusions can be derived from this exercise?
2. What might be an explanation of a higher count in Basin D than in B, ruling out contamination or faulty
techniques?
3. Why is it so important that surgeons scrub their hands prior to surgery even though they wear rubber
gloves?
356
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Laboratory Report student:
45
Desk No.: Section
Preparation and Care of Stock Cultures
1 . Why shouldn't cultures be stored at room temperature or in the incubator for any length of time?
2. Why should stock cultures be reinoculated to new media ("rotated") even if they are stored in the
refrigerator?
3. For what types of inoculations do you use your
reserve stock culture?
working stock culture?
4. What is lyophilization?
What advantage does this procedure have over the method we are using for maintaining stock cultures?
357
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Benson: Microbiological
Back Matter
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Eighth Edition
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Laboratory Report
Student:
48^50
Desk No.:
Section
Physiological Characteristics of Bacteria
A. Media
List the media that are used for the following tests:
1. Butanediol production
2. Hydrogen sulfide production
3. Indole production
4. Starch hydrolysis
5. Urease production
6. Citrate utilization
7. Fat hydrolysis
8. Casein hydrolysis
9. Catalase production
10. Mixed acid fermentation
1 1 . Glucose fermentation
12. Nitrate reduction
B. Reagents
Select the reagents that are used for the following tests:
C.
1 . Indole test
2. Voges-Proskauer test
3 . Catalase test
4. Starch hydrolysis
Barritt's reagent — 1
Gram's iodine — 2
Hydrogen peroxide-
Ko vacs' reagent — 4
None of these — 5
■3
Ingredients
Select the ingredients of the reagents for the following tests. Consult
Appendix B . More than one ingredient may be present in a particular reagent.
1.
Oxidase test
a-naphthol — 1
2.
Voges-Proskauer test
Dimethyl- a- naphthylamine — 7
3.
Indole test
Dimethyl- p-pheny lenedi amine
4.
Nitrite test
hydrochloride — 3
p-dimethylamine benzaldehyde
Potassium hydroxide — 5
Sulfanilic acid — 6
4
D. Enzymes
What enzymes are involved in the following reactions?
1. Urea hydrolysis
2. Hydrogen gas production from formic acid
3. Casein hydrolysis
4. Indole production
5 . Nitrate reduction
6. Starch hydrolysis
7. Fat hydrolysis
8. Gelatin hydrolysis (Ex. 47)
9. Hydrogen sulfide production
Answers
Media
1.
o
3.
4.
Ui
6
7.
fl
9.
m_
12.
Reagents
Ingredients
1.
2.
3.
4
i
1-
2.
3.
4.
Enzymes
i ■ ,, _,-_ .,.., _.. _
2.
3.
4.
5.
6.
■ ■
7
r -
8.
9.
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Physiological Characteristics of Bacteria
E. Test Results
Indicate the appearance of the following positive test results
1 . Glucose fermentation, no gas
2. Citrate utilization
3. Urease production
4. Indole production
5. Acetoin production
6. Hydrogen sulfide production
7. Coagulation of milk
8. Peptonization in milk
9. Litmus reduction in milk
10. Nitrate reduction
11. Catalase production
12. Casein hydrolysis
13. Fat hydrolysis
Answers
1.
2.
"•
4.
5.
6.
7.
a.
9.
10.
11.
12.
13.
F. General Questions
1 . Differentiate between the following
Respiration:
Fermentation:
Oxidation:
Reduction:
Catalase:
Peroxidase:
2. List two or three difficulties one encounters in trying to differentiate bacteria on the basis of physiologi
cal characteristics.
3. Now that you have determined the morphological, cultural, and physiological characteristics of your un-
known, what other kinds of tests might you perform on the organism to assist in identification?
360
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Laboratory Report
Student:
52
Desk No.:
Section
Enterobacteriaceae Identification
The API 20E System
A. Tabulation of Results
By referring to charts I and II, Appendix D, determine the results of each test and record these results as pos-
itive ( + ) or negative ( — ) in the table below. Note that the results of the oxidase test must be recorded in the
last column on the right side of the table.
ONPGADH LDC
ODC CIT H2S
URE TDA IND
VP GEL GLU
■
MAN INO SOR
RHA SAC MEL
AMY ARA OXI
1
2
4
1
2
4
1
2
4
1
2
4
1
2
4
1
2
4
1
2
4
N0 2 G N A 2 S MOT
MACOF-OOF-F
1
2
4
1
2
4
Additional Digits
B. Construction of Seven-Digit Profile
Note in the above table that each test has a value of 1, 2, or 4. To compute the seven-digit profile for your un-
known, total up the positive values for each group.
Example:
5 1 44 572 = E. coti
ONPG ADH LDC ODC CIT H^S URE TDA IND VP GEL GLU
+
1
+
1
1
+
+
MAN INO SOR RHA SAC MEL AMY ARA OX
+
+
+
+
1
1
361
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Enterobacteriaceae Identification: The API 20E System
C. Using the API 20E Analytical Index or the API Characterization Chart
If the API 20E Analytical Index is available on the demonstration table, use it to identify your unknown, us-
ing the seven-digit profile number that has been computed. If no Analytical Index is available, use
Characterization Chart III in Appendix D.
Name of Unknown:
D. Additional Tabulation Blank
If you need another form, use the one below
^^Uita
I-20E
Reference Number
Patient
Date
Source /Site
Physician
Dept. /Service
1
ONPG
1
ADH
2
LDC:
4
ODC
1
CIT
2
4
URE
1
TOA
2
IND
4
VP
1
■ " i
■ 5 h
I
— i— i-i i— H i
24 h
48 h
Profile
Number
GEL
2
GLU
4
MAN
1
INO
2
SOR 1
4 |
iRHA
1
SAC
2
MEL
4
AMY
1
ARA
2
OXJ
4
■4-H .
NO,
1
N a
GAS
2
MOT
4
MAC
1
OFO
2
OFF
4
5 h
24 h
48 h
Additional
Digits
Additional Information
Identification
00-42^012
(7/80)
bf+nt^^
E. Questions
1 . What is the intended function of the API 20E system?
2. In the "real world" who would use this system?
3. What might be an explanation for the failure of this system to work with some of the bacterial cultures
we use?
362
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Laboratory Report
Student:
53
Desk No.:
Section
Enterobacteriaceae Identification
The Enterotube II System
A. Tabulation of Results
Record the results of each test in the following table with a plus ( + ) or minus ( — )
B. Identification by Chart Method
If no Interpretation Guide is available, apply the above results to chart IV, Appendix D, to find the name of
your unknown. Note that the spacing of the above table matches the size of the spaces on chart IV. If this page
is removed from the manual, folded, and placed on chart IV, the results on the above table can be moved down
the chart to make a quick comparison of your results with the expected results for each organism.
C. Using the Enterotube II Interpretation Guide
If the Interpretation Guide is available, determine the five-digit code number by circling the numbers (4, 2,
or 1) under each test that is positive, and then totaling these numbers within each group to form a digit for
that group. Note that there are two tally charts on the next page of this Laboratory Report for your use.
ID Value
L
Y
S.
o
Ft
: N.
H2S
^®
+ 2
+ V
a>
1
A
L
N
D
A
D.
C.
N.
4
+ @
+©)
<H>
<5>
p
U
R
c
1
A.
LLI <
T.
4
+@
♦©,
<3>
The "ID Value" 34363 can be found by thumbing the pages of the Interpretation Guide. The listing is as
follows:
ID Value
34363
Organism
Klebsiella pneumoniae
Atypical Test Results
None
Conclusion: Organism was correctly identified as Klebsiella pneumoniae. In this case, the identification was
made independent of the V-P test.
363
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Enterobacteriaceae Identification: The Enterotube II System
D. Tally Charts
ENTEROTUBE® II*
G
L
U.
G
A
S
2 + 1
L
Y
S.
*4
o
R
N
H*S
I
N
A
D
O
ISI.
L
A
C
v 4+2+1
4 + 2 + 1
W
A
R
A
B
S
o
R
B.
D
U
L.
▼
V
4 + 2 + 1
p
A
U
R
E
A
C
I
JD Value
▼
A.
4 + 2+1
▼
Confirmatory Test
Result
i n ji
Culture Number, Case Number or Patient Name
■ VP utilized as confirmatory lest only
Date
Organism Idenlified
ENTEROTUBE® If*
G
L
U
G
A
S
2 + 1
L
Y
S
^■■ i > W H
R
N
ii ii
I
H 2 S N i
D
A
D
O
N
A.
L
A
C
V UnMM
JD Value
4 + 2+1 4+2+1
,' V
TT
A
R
A
B
S
o
R
B
D
U
L
4 + 2 + 1
p
A
U
R
E
A
C
I
T.
>V
4 + 2+1
▼
...y
Confirmatory Test
Result
Culture Number, Case Number or Patient Name
*VP utilized as confirmatory lest only,
Date
Organism Identified
E. Questions
1 . What is the intended function of the Enterotube II System?
2. In the "real world" who would use this system?
3. What might be an explanation for the failure of this system to work with some of the bacterial cultures
we use?
364
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Laboratory Report
Student:
54
Desk No.:
Section
O/F Gram-Negative Rods Identification:
The Oxi/Ferm Tube II System
A Tabulation of Results and Code Determination
Once you have marked the positive reactions on the side of the tube and circled the numbers that are assigned
to each of the positive chambers, as indicated in the example below, add the numbers in each bracketed group
to get the five-digit code.
The final step is to look up the code number in the Oxi/Ferm Tube II Biocode Manual to determine the
genus and species. If confirmatory tests are necessary, the manual will tell you which ones to perform.
In the example below, the code number is 32303. If you look up this number in the Biocode Manual you
will find on page 25 that the organism is Pseudomonas aeruginosa.
Use this procedure to identify your unknown by applying your results to the blank diagrams provided.
B. Results Pads
o
9
03
C
<
CD
c
E?
<
\/
CD
CO
4 + 2 + 1
v
CD
CO
O
O
03
CM
CD
CO
O
o
c/d
_CD
o
CD
(/)
O
X
o
O
i
CD
<
4+2+1 4 + 2+1
CD
CO
O
05
03
2
4 + 2 + 1
X
X
o
O
i
03
C
<
CD
C
c
E?
<
CD
C
C/)
CD
CO
O
03 [
CM
CD
CO
O
O
CO
_CD
O
"D
CD
CO
O
X
o
O
i
<
4 + 2 + 1 4+2+1 4 + 2+1
CD
CO
_o
03
03
£
4 + 2 + 1
y \
365
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O/F Gram-Negative Rods Identification: The Oxi/Ferm Tube II System
C. Questions
1 . What is the intended function of the Oxi/Ferm Tube II System?
2. In the "real world" who would use this system?
3. What might be an explanation for the failure of this system to work with some of the bacterial cultures
we use?
366
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Laboratory Report
Student:
55
Desk No.:
Section
Staphylococcus Identification:
The API Staph- Ident System
A. Tabulation of Results
By referring to charts V and VI, Appendix D, determine the results of each test, and record these results as
positive ( + ) or negative ( — ) in the Profile Determination Table below. Note that two more of these tables
have been printed on the next page for tabulation of additional organisms.
PHS
1
URE
2
GLS
4
MNE
1
MAN
2
TRE
4
SAL
1
GLC
2
ARG
4
NGP
1
RESULTS
4_ ^ l _ m
1 1 — l
PROFILE NUMBER
Additio
nal Info
rrnation
Identification
GRAM STAIN
COAGULASE
MORPHOLOGY
C ATA LAS E
:
B. Construction of Four-Digit Profile
Note in the above table that each test has a value of 1, 2, or 4. To compute the four-digit profile for your un
known, total up the positive values for each group.
PHS URE GLS MNE MAN TRE SAL GLC ARG NGP
4-
1
+
+
+
1
+
+
C. Final Determination
Refer to the Staph-Ident Profile Register (chart VII, Appendix D) to find the organism that matches your pro-
file number. Write the name of your unknown in the space below and list any additional tests that are needed
for final confirmation. If the materials are available for these tests, perform them.
Name of Unknown:
Additional Tests:
367
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Styphylococcus Identification: The API Staph-ldent System
PHS
1
URE
2
GLS
4
MNE
1
MAN
2
TRE
4
SAL
1
GLC
2
ARG
4
NGP
1
RESULTS
PROFILE NUMBER
r— *■-— " ..-™_™
GRAM STAIN
MORPHOLOGY
COAGULASE
CATALASE
Additional Information
Identification
GRAM STAIN
MORPHOLOGY
COAGULASE
CATALASE
Additional Information
PHS
1
URE
2
GLS
4
MNE
1
MAN
2
TRE
4
SAL
1
GLC
2
ARG
4
NGP
1
RESULTS
"H
PROFILE NUMBER
Identification
D. Questions
1 . What is the intended function of the API Staph-ldent System?
2. In the "real world" who would use this system?
3. What might be an explanation for the failure of this system to work with some of the bacterial cultures
we use?
368
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Laboratory Report
Student:
56,57
Desk No.:
Section:
Ex. 56 Microbial Population Counts of Soil
A. Tabulation of Results
Select the best plate and count the organisms on a Quebec colony counter. Tabulate your results and the re
suits of other students near you who cultured the other types of soil organisms. Be sure to count only repre
sentative types.
ORGANISMS
COUNT PER PLATE
DILUTION
ORGANISMS PER GRAM
OF SOIL
Bacteria
Actinomycetes
Molds
B. Conclusions
What generalizations can you make from this exercise?
C. Questions
1 . Why would the number of bacteria present in the soil actually be higher than the number determined by
your plate count?
2. What other types of microbes might be present in your soil sample?
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Isolation of an Antibiotic Producer from Soil
Ex. 57 Isolation of an Antibiotic Producer from Soil
A. Results
Describe in detail how the experiment turned out.
B. Questions
1 . What four genera of microbes produce the vast majority of antibiotics?
2. What is known about the importance of these antibiotics in the natural habitat of these microbes?
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Desk No.:
Section
Nitrogen Fixing Bacteria
A Questions
1 . What enzyme is responsible for nitrogen fixation? _
2. What metal is essential for this enzyme to function?
3. Why is nitrogen fixation so important?
B. Azotobacteraceae
Identify the characteristics of your isolate that led to your decision as to its identification
1 . Were the colonies pigmented?
If pigmented, what color?
Was fluorescence present?
If fluorescent, what color?
2. Were you able to see cysts?
3. Is the organism motile?
4. Gram reaction?
5. Any other pertinent characteristic?
Azotobacteraceae
Name of Organism:
Draw a few cells of organisms in circle at right.
C. Rhizobiaceae
Draw a few representative cells of Rhizobium in the circle to the right.
1. What is the most important criterion for species identification in the
family of nitrogen-fixers?
2. From the standpoint of amount of nitrogen fixation, is this group of
nitrogen-fixers more or less important than the Azotobacteraceae?
Rhizobiaceae
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Desk No.:
Ammonification in Soil
Section
A. Tabulation of Results
Record the presence or absence of ammonia in the tubes of media
+
+ +
+ + +
slight ammonia (faint yellow)
moderate ammonia (deep yellow)
much ammonia (brown precipitate)
no ammonification
INCUBATION
TIME
AMOUNT OF AMMONIA
pH
Control
Peptone
with Soil
Control
Peptone
with Soil
4 days
7 days
B. Questions
1 . In the natural situation, what compounds serve as the source of the ammonia released in ammonification?
2. In simple terms, what occurs during decay?
3. Differentiate between the following
Peptonization:
Ammonification :
4. What happens to ammonia in soil when denitrification of ammonia takes place?
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Desk No.: Section
Ex. 61 Isolation of a Denitrifier from Soil
Using Nitrate Agar
A Observations:
1 . Second Period. Describe the types of colonies observed growing on the nitrate agar plate
2. Third Period. What evidence do you have of the presence of a denitrifier in the soil sample you are work-
ing with?
3. Microscopic Examination. Describe the appearance of organisms that were observed on a gram-stained
slide.
4. Further Testing. If you performed any further tests for species identification, state any conclusions that
were made.
B. Questions:
1 . Why are denitrifying bacteria costly to farmers?
2. Why are denitrifying bacteria essential to the existence of life on plant earth?
3. Of what value is denitrification to the organism?
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Isolation of a Denitrifier from Soil: Using an Enrichment Medium
Ex. 62 Isolation of a Denitrifier from Soil:
Using an Enrichment Medium
A. Observations:
1 . First Enrichment Culture Appearance. Describe the appearance of the first enrichment culture in the
second laboratory period:
2. Microscopic Examination: Describe the appearance of the cells as seen on
Wet Mount:
Gram- stained Slide:
3. Second Enrichment Culture Appearance. Describe the appearance of the second enrichment culture
after five days incubation:
4. Agar Plate Colonies. Describe the characteristics of the colonies on the nitrate succinate-mineral salts
agar:
Do these characteristics match those of Paracoccus denitrificans?
5. Final Gram-stained Slide. How do the microscopic characteristics of your organism match the de
scription of Paracoccus denitrificans?
B. Questions: Answer the questions that are provided on the front page of this Laboratory Report.
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Desk No.:
Section
Bacteriological Examination of Water:
Qualitative Tests
A. Results of Presumptive Test (MPN Determination)
Record the number of positive tubes on the chalkboard and on the following table. When all students have
recorded their results with the various water samples, complete this tabulation. Determine the MPN accord-
ing to the instructions on page 224.
WATER SAMPLE
(SOURCE)
NUMBER OF POSITIVE TUBES
MPN
3 Tubes DSLB
10 ml
3 Tubes SSLB
1.0 ml
3 Tubes SSLB
0.1 ml
3 Tubes SSLB
0.01 ml
.
B. Results of Confirmed Test
Record the results of the confirmed tests for each water sample that was positive on the presumptive test
WATER SAMPLE
(SOURCE)
POSITIVE
NEGATIVE
■
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Bacteriological Examination of Water: Qualitative Tests
C. Results of Completed Test
Record the results of completed tests for each water sample that was positive on the confirmed test
WATER SAMPLE
(SOURCE)
LACTOSE
FERMENTATION
RESULTS
MORPHOLOGY
EVALUATION
■
D. Questions
1 . Does a positive presumptive test mean that the water is absolutely unsafe to drink?
Explain:
2. What might cause a false positive presumptive test?
3. List three characteristics required of a good sewage indicator: a
b.
c.
4. What enteric bacterial diseases are transmitted in polluted water?
5. Name one or more protozoan diseases transmitted by polluted water.
6. Why don't health departments routinely test for pathogens instead of using a sewage indicator?
7. Give the functions of the various media used in these tests
Lactose broth:
Levine EMB agar
Nutrient agar slant:
8. What media, other than the ones used here, can be used for confirmatory tests?
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>
65
Desk No.:
Section
Ex. 64 Membrane Filter Method
A. Tabulation
A table similar to the one below will be provided for you, either on the chalkboard or as a photocopy. Record
your coliform count on it. Once all data are available, complete this table.
SAMPLE
SOURCE
COLIFORM COUNT
AMOUNT OF WATER
FILTERED
MPN*
A
B
C
i
D
E
F
G
H
MPW
Coiiform Count x 100
Amount of Water Filtered
B. Questions
1 . Give two limitations of the membrane filter technique
2. Even if the membrane filter removed all bacteria from water being tested, is the water that passes through
sterile?
Explain:
3. List some other applications of membrane filter technology in microbiology.
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Standard Plate Count: A Quantitative Test
Ex. 65 Standard Plate Count
A Quantitative Test
A. Tabulation of Results
After you have made your plate counts, record your results on the chalkboard and on the following table. After
all students have recorded their results, complete this table.
SAMPLE
B
■ > ■ ■■> ■
SOURCE
PLATE COUNT
(AVERAGE)
DILUTION
ORGANISMS PER ML
D
^—^J—lm
,
^^^^^•^^^^^^^^^^
H
ii 1 1 i ii ■ ■ — *•
ivfvp^mwm
What generalizations, if any, can be drawn from these results?
B. Questions
1 . What kinds of organisms in water will not produce colonies on TGEA?
2. Would a high plate count indicate that the water is unsafe to drink?
Explain:
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Desk No.:
Section
Ex. 66 Standard Plate Count of Milk
A. Tabulation of Results
After you have made your plate count, record your results on the following table. Get the results of the other
milk sample from some other member of the class.
TYPE OF MILK
PLATE COUNT
DILUTION
ORGANISMS PER ML
High-quality
Poor-quality
B. Questions
1 . Do plate count figures represent numbers of organisms or numbers of clumps of bacteria?
2. What are some factors that will produce errors in the SPC technique?
3. What might be the explanation of a very high count in raw milk that has been properly refrigerated from
the time of collection?
4. What is the most common source of bacteria in milk?
5. Why is milk a more suitable vector of disease than water?
6. What infectious diseases of cows can be transmitted to humans via milk?
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Direct Microscopic Count of Organisms in Milk: The Breed Count
Ex. 67 Direct Microscopic Count of Organisms in Milk
The Breed Count
A Microscope Factor
Show your computations in determining the MF of your microscope
1 . Field diameter:
2
2. Field area (in* ):
millimeters
square millimeters
square centimeters
3. Number of fields in one square centimeter
4. Microscope factor (100 X no. of fields): _.
B. Cell Counts
Record the counts of each field, total them, and determine their average counts. Get the results of the other
type of milk from a class member.
GOOD MILK
POOR MILK
Bacterial
Cells
Bacterial
Clumps
Leukocytes
Bacterial
Cells
Bacterial
Clumps
Leukocytes
1
2
3
4
5
Total
Av.
Determine the number of cells, clumps, and leukocytes per ml for each type of milk.
Good Milk
Poor Milk
Bacterial Cells:
Bacterial Cells:
Bacterial Clumps:
Leukocytes:
Bacterial Clumps
Leukocytes:
C. Questions
1 . Should milk from a healthy cow be completely free of bacteria?
2. Are leukocytes in milk always an indication of infection (mastitis)?
3. Why are direct counts higher than standard plate counts?
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Desk No.: Section
Reductase Test
1. How would you grade the two samples of milk that you tested? Give the MBRT for each one
Sample A:
Sample B:
2. Is milk with a short reduction time necessarily unsafe to drink?
Explain:
3. What other dye can be substituted for methylene blue in this test?
4. What advantage do you see in this method over the direct count method?
5. What kinds of organisms may be plentiful in a milk sample, yet give a negative reductase test?
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Desk No.:
Section
Bacterial Counts of Foods
A. Tabulation of Results
Record your count and the bacterial counts of various other foods made by other students
TYPE OF FOOD
PLATE COUNT
DILUTION
ORGANISMS PER ML
i
i
B. Questions
1 . Why is there such great variability in organisms per ml between different kinds of food?
2. What dangers and undesirable results may occur from ground meats of high bacterial counts?
3. What bacterial pathogens might be present in frozen foods?
4. What harm can result from repeated thawing and freezing of foods?
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Bacterial Counts of Foods
5. What precautions are taken to prevent the spoilage of foods?
6. Which methods in question 5 are most effective?
Least effective?
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)
71
Desk No.:
Section:
Ex. 70 Microbial Spoilage of Canned Food
A. Results
Record your observations of the effects of each organism on the cans of vegetables. Share results with other
students.
ORGANISM
PEAS
CORN
Gas Production Odor
+ or -
Gas Production Odor
+ or -
£ coii
B. coagulans
B. stearothermophilus
C. sporogenes
C. thermosaccharolyticum
B. Microscopy
After making gram-stained and spore-stained slides of all organisms from the canned food extracts, sketch in
representatives of each species:
^^^^P^WWWWWWWWWWPPH***
^^W^4n^^ff**P^^^^^ff*P^^ff *^4mn^B^^^M*fcUBiMtatata^^Hl
E. cofi
B. coagulans
B. stearothermophilus
C. Questions
1. Which organisms, if any, caused "flat sour spoilage"?
^^^^^^^^^^^^w^^
C. sporogenes
C, thermosac-
charolyticum
2. Which organisms, if any, caused "T.A. spoilage"?
3. Which organisms, if any, caused "stinker spoilage"?
4. Does "flat sour" cause a health problem?
5. Describe how typical spoilage resulting in botulism occurs
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Microbial Spoilage of Refrigerated Meat
Ex. 71 Microbial Spoilage of Refrigerated Meat
A. Results
1 . After performing the colony count on your best plates, how many psychrophilic-psychrotrophic bacteria
do you find were present in each gram of the meat sample?
2. Describe the appearance of the organism that you stained with Gram's stain.
B. Questions
1 . Differentiate between the following
Psy chrophile :
Psy chrotroph :
2. Why was it necessary to incubate the plates for 2 weeks?
3. List some genera of bacteria that might be psychrophilic or psychrotrophic in the meat sample
Gram -negative types:
Gram -positive types:
4. List three or four pathogenic psychrotrophs that might be found in refrigerated meat
5. What are the probable sources of psychrophilic microbes found growing in meat?
6. What types of measures can be taken to prevent spoilage of meats by psychrophilic-psychrotrophic
bacteria?
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Desk No.: Section:
Ex. 72 Microbiology of Fermented Beverages
A. Results
Record here your observations of the fermented product
Aroma:
pH:
H 2 S production
B. Questions
1 . Why must the fermentor be sealed?
Why with a balloon?
2. What compound in the grape juice is being fermented?
3. Why would production of hydrogen sulfide by the yeast be of importance?
4. Why are we concerned about the pH of the fruit juice and the wine?
5. What happens to wine if Acetobacter takes over?
6. What process can be used to prevent the action of Acetobacter in the production of wine and beer?
Ex. 73 Microbiology of Fermented Milk Products
A Results
1 . Record here your observations of the fermented product:
Color:
Aroma:
Texture:
Taste:
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Microbiology of Fermented Milk Products
2. In the space provided below sketch in the microscopic appearance of the organisms as seen on the slide
stained with methylene blue.
B. Questions
1 . What type of fermentation is involved in the production of yogurt?
2. What is the nutritional value of yogurt?
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Desk No.:
Section:
Ex. 74 Mutant Isolation by Gradient Plate Method
A. Results
1 . How many colonies of E. coll did you count in the high concentration area of the plate?
2. When the streptomycin-resistant colonies were smeared with a loop toward the high concentration side
and reincubated, did they continue to grow in the new area?
What does this result indicate?
Ex. 75 Mutant Isolation by Replica Plating
A. Tabulation of Results
Count the colonies that occur on both plates and record the information on the chalkboard on a table similar
to the one below. After all students have recorded their counts, complete this table.
STUDENT
INITIALS
■
NUMBER OF COLONIES
STUDENT
INITIALS
NUMBER OF COLONIES
Nutrient
Agar
Streptomycin
Agar
Nutrient
Agar
Streptomycin
Agar
■
i
TOTALS
TOTALS j
AVERAGE
PER PLATE
AVERAGE
PER PLATE
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Mutant Isolation by Replica Plating
B. Questions
1 . After determining the average number of streptomycin-resistant colonies per plate, calculate the muta
bility rate of the organisms, assuming that there were 100,000,000 organisms per milliliter in the origi-
nal culture. Show your mathematical computations.
Mutability rate:
2. What does replica plating of mutants attempt to prove that is not established by the gradient plate method?
3. How can the frequency of mutation in bacteria be increased?
4. In what way does the presence of an antibiotic increase the numbers of resistant forms of bacteria?
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Desk No.:
Section
Bacterial Mutagenicity and Carcinogenesis
The Ames Test
A. Tabulation of Results
Record the results of your tests in the following table and on a similar table on the chalkboard. Also record
the results of substances tested by other students. A positive result will exhibit a zone of colonies similar to
the zone shown on the plate in illustration 5, figure 76.1.
TEST SUBSTANCE
RESULT
(+ or -)
TEST SUBSTANCE
RESULT
(+ or -)
TEST SUBSTANCE
i
■
•
RESULT
(+ or -)
i
i
•
•
•
•
i
B. Questions
1.
Did you observe a zone of inhibition between the growing colonies and the impregnated disk on your
positive plates?
What is the cause of such a zone?
2. Differentiate between the following
Prototroph:
Auxotroph :
3 . Define back mutation
4. List two characteristics of the Ames test that made this test so much superior to previous mutagenesis
tests :
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Bacterial Mutagenicity and Carcinogenesis: The Ames Test
5. Does the fact that a chemical substance is carcinogenic in animals necessarily mean that it is also
carcinogenic in humans?
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Desk No.:
A Synthetic Epidemic
Section
A. Tabulation of Results
Record in the table below the information that has been tabulated on the chalkboard. The SHAKER is the
person designated by the instructor to shake hands with another class member. The SHAKEE is the individ-
ual chosen by the shaker. For Procedure A a blue color is positive; yellow or brown is negative. For
Procedure B red colonies (S. marcescens) is positive; no red colonies is negative.
SHAKER
Round 1
RESULT
+ or -
SHAKEE
Round 1
RESULT
+ or -
SHAKER
Round 2
RESULT
+ or -
SHAKEE
Round 2
RESULT
+ or -
! 1.
1.
2.
2.
3.
3.
4.
4.
5.
5.
6.
6.
7.
7.
8.
8.
9.
9.
10.
10.
11.
11.
12.
r
12.
13.
13.
14.
14.
15.
15.
16.
16. ;
17.
17.
18.
18.
19.
19.
20. '
20.
21.
21.
22.
22.
23.
23.
24.
24.
■
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A Synthetic Epidemic
B. Questions
1. Who in the group was "patient zero," the starter of the epidemic?
2. How many carriers resulted after
Round 1?
Round 2?
3. If this were a real infectious agent, such as a cold virus or influenza, list some other factors in transmis
sion besides the ones we tested:
4. How would it have been possible to stop this infection cycle?
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Desk No.:
Section
The Staphylococci:
Isolation and Identification
A. Tabulation
At the beginning of the third laboratory period, the instructor will construct a chart similar to this one on the
chalkboard. After examining your mannitol salt agar and staphylococcus medium 110 plates, record the pres-
ence (+) or absence (— ) of staphylococcus growth in the appropriate columns. After performing coagulase
tests on the various isolates record the results also as (+) or (— ) in the appropriate columns.
STUDENT
INITIALS
NOSE
FOMITES
Staph Colonies
Coagulase
Item
Staph Colonies
Coagulase
MSA
SM110
MSA
SM110
i
1
" " ""]
■
■
■
i
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The Staphylococci: Isolation and Identification
8. Microscopy
Provide drawings here of the various isolates as seen under oil immersion (gram staining)
UNKNOWN-CONTROL
i
i
NOSE
FOMITE
C. Percentages
From the data in the table on the previous page, determine the incidence (percentage) of individuals and
fomites that harbor coagulase-positive and coagulase-negative staphylococci in this experiment.
SOURCE
TOTAL
TESTED
TOTAL
POSITIVE
PERCENTAGE
POSITIVE
TOTAL
NEGATIVE
PERCENTAGE
NEGATIVE
Humans (Nose)
Fomites
i
D. Record of Test Results
Record here the results of each test performed in this experiment. Under GRAM STAIN indicate cellular
arrangement as well as Gram reaction.
ISOLATE
GRAM STAIN
ALPHA TOXIN
MANNITOL
(ACID)
COAGULASE
Unknown-Control No.
Nose Isolate No. 1
Nose Isolate No. 2
Fornite Isolate
E. Final Determination
Record here the name of your unknown-control. If API Staph-Ident miniaturized multitest strips are avail
able, confirm your conclusions by testing each isolate. See Exercise 55.
Name of unknown-control:
Staph-Ident results:
E Questions
1. What are nosocomial infections?
2. What bacterial organism causes most nosocomial infections?
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Desk No.:
Section
The Streptococci:
Isolation and Identification
A Tabulation of Pharynx Isolates
The instructor will construct a chart similar to this one on the chalkboard. After examining the blood agar
plates that were inoculated with pharynx organisms, record the types and size range of colonies that are pres-
ent on your plates. Record these data first on this table, then on the chalkboard. After all students have
recorded their results on the board, complete the tabulation of their results here, also. The names of the or-
ganisms will not be recorded until all tests are completed.
STUDENT
INITIALS
TYPE OF HEMOLYSIS
(ALPHA, ALPHA-PRIME,
BETA)
SIZE RANGE
OF COLONIES
(MM)
NAMES OF ORGANISMS
i
_
■
■
I
i
■
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The Streptococci: Isolation and Identification
8. Microscopy
Provide drawings here of the various pharyngeal isolates as seen under oil immersion (gram staining)
C. Percentages
From the data in the table on the previous page, calculate the percentages for each type of streptococci that
were isolated from classmates.
S. pyogenes:
S. agalactiae:
Group C streptococci:
Group D enterococci:
S. pneumoniae:
Group D nonenterococci
Viridans streptococci:
D. Record of Test Results
Record here all information pertaining to the identification of pharyngeal isolates and unknowns
SOURCE OF UNKNOWN \ \ \ \ \ \ \ \ \ \
E. Final Determination
Record here the identities of your various isolates and unknowns
Pharyngeal isolates:
Unknowns:
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The Streptococci: Isolation and Identification
F. Questions
Record the answers for the following questions in the answer column.
1 . What two physiological tests are significant in the identification of
S. agalactiae?
2. If an alpha hemolytic streptococcus is able to hydrolyze bile esculin,
what test can be used to tell whether the organism is an enterococ-
cus?
3. What test is used for differentiating group A from group C strepto-
cocci if both organisms are bacitracin- susceptible?
4. What two tests are used to differentiate pneumococci from the virid-
ians group?
5. What test is used for differentiating S. pyogenes from other beta he-
molytic streptococci?
6. Hemolysis in streptococci can only be evaluated when the colonies
develop (aerobically or anaerobic ally) in blood agar.
7. Which streptococcal species is frequently present in the vagina of
third- trimester pregnant women?
8. Only one beta hemolytic streptococcus is primarily of human origin.
Which one is it?
9. Who developed the system of classifying streptococci into groups A,
B, C, etc.?
10. Who is credited with grouping streptococci according to the type of
hemolysis?
1 1 . Which streptococcal species is seen primarily as paired cells (diplo-
cocci)?
1 2. Name two species of streptococci that are implicated in dental caries.
13. Where in the body can S. bovis be found?
14. After performing all physiological tests, what type of tests must be
performed to confirm identification?
15. Which hemolysin produced by S. pyogenes is responsible for the
beta-type hemolysis that is characteristic of this organism?
b.
2.
3.
4,a.
b.
5.„
8.
10.
11.
12.a.
13,
14.
15.
Answers
I iQl . ■ +m . .
I I ' FI .Ifat bll I I I I !■ I
■-■ — *■■ 'I I I
401
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Applications Lab Manual,
Eighth Edition
Back Matter
Laboratory Reports
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Benson: Microbiological
Back Matter
Laboratory Reports
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Companies, 2001
Laboratory Report student:
80
Desk No.: Section
Gram-Negative Intestinal Pathogens
A. Unknown Identification
1 . What was the genus of your unknown?
Genus No.
2. What problems, if any, did you encounter?
3. Now that you know the genus of your unknown, what steps would you follow to determine the species?
B. General Questions
1 . Why are bile salts and sodium desoxycholate used in certain selective media in this exercise?
2. How can one identify coliforms on MacConkey agar?
3. How does one differentiate Salmonella from Shigella colonies on XLD agar?
4. What characteristics do the salmonella and shigella have in common?
403
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Laboratory Reports
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Gram- Negative Intestinal Pathogens
5. How do the salmonella and shigella differ?
6. What restrictions might be placed on a person who is a typhoid carrier?
404
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Laboratory Reports
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Eighth Edition
Companies, 2001
Laboratory Report
Student:
81
Desk No.:
Section
Urinary Tract Pathogens
A Quantitative Evaiuation
After counting the colonies on the TS A plate, record the count as follows
Number of colonies:
Dilution:
No. of organisms/ml of urine:
Gram-Stained Slide. If organisms are seen on a gram- stained slide of an uncentrifuged sample, sketch in
color in the circle below.
Conclusion: Do the plate count and gram- stained slide of the
uncentrifuged sample provide presumptive evidence of a uri-
nary infection?
B. Microscopic Study (Centrifuged Sample)
Illustrate in the circle below the microscopic appearance of a centrifuged sample.
Conclusion: Describe here the morphological appearance of
the predominant organism seen:
C. Culture Analyses
After studying the organisms on the three plates and thioglycollate medium, what organism do you believe
is causing the infection?
Organism:
What further testing should be performed for confirmation?
405
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Applications Lab Manual,
Eighth Edition
Back Matter
Laboratory Reports
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Benson: Microbiological
Back Matter
Laboratory Reports
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Companies, 2001
Laboratory Report student:
82,83
Desk No.: Section
Ex. 82 Slide Agglutination Test:
Serological Typing
1 . Record the unknown number that proved to be a salmonella.
2. Why was phenolized saline used instead of plain physiological saline?
3. If your results were negative for both cultures, what might be the explanation?
Ex. 83 Slide Agglutination (Latex) Test:
For S. aureus Identification
1 . If your test turned out to be positive for S. aureus, record the degree of positivity here:
2. What other test for S. aureus is highly correlated with this test?
3. What two kinds of antibodies are attached to the latex particles in the Difco latex reagent?
a.
b.
4. What role, if any, do the staphylococci cells play in this reaction?
407
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Eighth Edition
Back Matter
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Benson: Microbiological
Back Matter
Laboratory Reports
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Companies, 2001
Laboratory Report student:
84,85
Desk No.: Section
Ex. 84 Tube Agglutination Test:
The Heterophils Antibody Test
1 . What was the titer of the serum that you tested?
2. For what disease is this diagnostic test used?
3. Below what titer would this test be considered to be negative?
4. What is the name of the virus that causes this disease?
5. What is unusual about a heterophile antibody?
6. If you are not going to perform the Widal test (Ex. 85), answer all the questions below except for ques
tion 1. It may be necessary for you to read the introduction to Ex. 85 for some of the answers.
Ex. 85 Tube Agglutination Test
The Widal Test
1 . What was the titer of the serum that you tested?
2. What is the exact meaning of the "titer" of a serum?
3. Differentiate between the following
Serum:
Antiserum:
Antitoxin :
4. How would you prepare antiserum for an organism such as E. coll?
409
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Laboratory Reports
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Tube Agglutination Test: The Widal Test
5.
Indicate the type of antigen (soluble protein, red blood cells, or bacteria) that is used for each of the fol
lowing serological tests:
Agglutination :
Precipitation:
Hemolysis:
6. For which one of the above tests is complement necessary?
410
Benson: Microbiological
Back Matter
Laboratory Reports
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Companies, 2001
Laboratory Report student:
86
Desk No.: Section
Phage Typing
1 . To which phage types was this strain of S. aureus susceptible?
2. To what lytic group does this strain of staphylococcus belong?
3. In what way can bacteriophage alter the genetic structure of a bacterium?
411
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
Back Matter
Laboratory Reports
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Benson: Microbiological
Back Matter
Laboratory Reports
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Companies, 2001
Laboratory Report
Student:
87-89
Desk No.:
Section
Ex. 87 White Blood Cell Study:
The Differential WBC Count
A. Tabulation of Results
As you move the slide in the pattern indicated in figure 87.4, record all the different types of cells in the fol-
lowing table. Refer to figures 87.1 and 87.2 for cell identification. Use this method of tabulation: 1^-rl 1-l-rf
1 1 . Identify and tabulate 100 leukocytes. Divide the total of each kind of cell by 100 to determine percentages.
NEUTROPHILS
LYM PHOCYTES
MONOCYTES
EOSINOPHILS
BASOPHILS
Total
Percent
B. Questions
1 . Were your percentages for each type within the normal ranges?
2. What errors might one be likely to make when doing this count for the first time?
3. Differentiate between the following
Cellular immunity:
Humoral immunity:
4. Do cellular and humoral immunity work independently?
Explain
413
Benson: Microbiological
Back Matter
Laboratory Reports
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Companies, 2001
Total WBC Count and Blood Grouping
Ex. 88 Total WBC Count
A Calculations
Using the formula provided on page 295, calculate the number of leukocytes per cubic millimeter.
Total Count:
B. Questions
1 . What is the normal WBC count range?
2. List several causes of a high WBC count
Ex. 89 Blood Grouping
1 . On the basis of this test, what is your blood type?
2. What antibodies are present in each type of blood?
Type A:
Type B :
Type AB :
Type O:
3. Why does a person of type A blood go into a transfusion reaction when given type B blood?
4. Why can Rh-positive blood be given only once to a person who is Rh-negative?
414
Benson: Microbiological
Back Matter
Laboratory Reports
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Applications Lab Manual,
Eighth Edition
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Laboratory Report student:
90
Desk No.: Section
The Snyder Caries Susceptibility Test
1 . What degree of caries susceptibility was indicated for you as a result of this test?
2. Is this substantiated by the amount of dental work you have had or should have had on your teeth?
3. What factors could affect the reliability of this test?
4. To increase the validity of this test how many times should it be performed and at what time of the day?
415
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
XIV. Medical Microbiology
and Immunology
Laboratory Report 90
© The McGraw-H
Companies, 2002
Benson: Microbiological
Back Matter
Descriptive Charts
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Companies, 2001
Descriptive Chart
STUDENT
' I M !■*
LAB SECTION:
Habitat:
Source:
Culture No.:
!■ I I Ul
Organism: —
**-
MORPHOLOGICAL CHARACTERISTICS
PHYSIOLOGICAL CHARACTERISTICS
Cell Shape:
Arrangement:
Size:
Spores:
Gram's Stain:
Motility:
Capsules:
Special Stains:
CULTURAL CHARACTERISTICS
Colonies:
Nutrient Agar:
Blood Agar:
Agar Slant:
Nutrient Broth:
Gelatin Stab:
Oxygen Requirements:
Optimum Temp.:
TESTS
o
s
s
RESULTS
Glucose
Lactose
Sucrose
i-i-i — ■ — «-
Mannitol
'So
u
>
•p^ ■ ^^r"
Gelatin Liquefaction
Sta rch
Casein
Fat
Indole
Methyl Red
V-P (acetylmethylcarbinol)
Citrate Utilization
Nitrate Reduction
HzS Production
Urease
Catalase
Oxidase
DNase
Phenylalanase
hjj— p^^b— i"i-
ii ■ ■ i ■ 'i
■ !■ M* !■
^
3
S
2
REACTION
Acid
Alkaline
Coagulation
Reduction
Peptonization
No Change
i ■ ■! i ■■«
417
Benson: Microbiological
Back Matter
Descriptive Charts
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Companies, 2001
Descriptive Chart
STUDENT
' I M !■*
LAB SECTION:
Habitat:
Source:
Culture No.:
!■ I I Ul
Organism: —
**-
MORPHOLOGICAL CHARACTERISTICS
PHYSIOLOGICAL CHARACTERISTICS
Cell Shape:
Arrangement:
Size:
Spores:
Gram's Stain:
Motility:
Capsules:
Special Stains:
CULTURAL CHARACTERISTICS
Colonies:
Nutrient Agar:
Blood Agar:
Agar Slant:
Nutrient Broth:
Gelatin Stab:
Oxygen Requirements:
Optimum Temp.:
TESTS
o
s
s
RESULTS
Glucose
Lactose
Sucrose
i-i-i — ■ — «-
Mannitol
'So
u
>
•p^ ■ ^^r"
Gelatin Liquefaction
Sta rch
Casein
Fat
Indole
Methyl Red
V-P (acetylmethylcarbinol)
Citrate Utilization
Nitrate Reduction
HzS Production
Urease
Catalase
Oxidase
DNase
Phenylalanase
hjj— p^^b— i"i-
ii ■ ■ i ■ 'i
■ !■ M* !■
^
3
S
2
REACTION
Acid
Alkaline
Coagulation
Reduction
Peptonization
No Change
i ■ ■! i ■■«
41
Benson: Microbiological
Back Matter
Descriptive Charts
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Companies, 2001
^ 4
Descriptive Chart
STUDENT:
' I M !■*
LAB SECTION:
Habitat:
Source:
Culture No.:
!■ I I Ul
Organism: —
MORPHOLOGICAL CHARACTERISTICS
PHYSIOLOGICAL CHARACTERISTICS
Cell Shape:
Arrangement:
Size:
Spores:
Gram's Stain:
Motility:
Capsules
Special Stains:
CULTURAL CHARACTERISTICS
Colonies:
Nutrient Agar:
Blood Agar:
Agar Slant:
Nutrient Broth:
Gelatin Stab:
Oxygen Requirements:
Optimum Temp.:
TESTS
o
v ■
c
g
U
RESULTS
Glucose
Lactose
Sucrose
Mannitol
"So
'o
u
Gelatin Liquefaction
Starch
Casein
Fat
._ ^ "._.j a *_j
Indole
Methyl Red
V-P (acetylmethylcarbinol)
Citrate Utilization
Nitrate Reduction
H 2 S Production
Urease
Catalase
Oxidase
DNase
Phenylalanase
rjj— p^^b— i"i-
ii ■ ■ i ■ 'i
■ !■ \W !■
^
3
S
2
REACTION
Acid
Alkaline
Coagulation
Reduction
Peptonization
No Change
i ■ ■! i ■■«
419
Benson: Microbiological
Back Matter
Descriptive Charts
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Companies, 2001
Descriptive Chart
STUDENT:
' I M !■*
LAB SECTION:
Habitat:
Source:
Culture No.:
!■ I I III
Organism: —
MORPHOLOGICAL CHARACTERISTICS
PHYSIOLOGICAL CHARACTERISTICS
Cell Shape:
Arrangement:
Size:
Spores:
Gram's Stain:
Motility:
Capsules
Special Stains:
CULTURAL CHARACTERISTICS
Colonies:
Nutrient Agar:
Blood Agar:
Agar Slant:
Nutrient Broth:
Gelatin Stab:
Oxygen Requirements:
Optimum Temp.:
TESTS
o
v ■
g
RESULTS
Glucose
Lactose
Sucrose
Mannitol
"So
'o
u
Gelatin Liquefaction
Starch
Casein
Fat
■ _ S %_■! a '_J
Indole
Methyl Red
V-P (acetylmethylcarbinol)
Citrate Utilization
Nitrate Reduction
H 2 S Production
Urease
Catalase
Oxidase
DNase
Phenylalanase
iun^^^^
ii ■ ■ i ■ 'i
■ !■ im !■
^
S
2
REACTION
TIME
Acid
Alkaline
Coagulation
Reduction
Peptonization
No Change
i ■ ■! i ■■«
421
Benson: Microbiological
Back Matter
Descriptive Charts
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Companies, 2001
Descriptive Chart
STUDENT
n-»¥»i-ta"^
LAB SECTION:
Habitat:
Source:
Culture No.
Organism: —
■ ■.*
MORPHOLOGICAL CHARACTERISTICS
PHYSIOLOGICAL CHARACTERISTICS
Cell Shape:
Arrangement:
Size:
Spores:
Gram's Stain:
Motility:
Capsules:
Special Stains:
CULTURAL CHARACTERISTICS
Colonies:
Nutrient Agar:
Blood Agar:
Agar Slant:
Nutrient Broth:
Gelatin Stab:
Oxygen Requirements:
Optimum Temp.:
TESTS
c
s
'53
U
RESULTS
Glucose
Lactose
Sucrose
Mannitol
■hi* ^^p"
Gelatin Liquefaction
Starc h
Casein
Fat
Indole
Methyl Red
1 1 ■ _ - 1*— ■ ■■
V-P (acetylmethylcarbinol)
Citrate Utilization
Nitrate Reduction
H2S Production
Urease
Catalase
Oxidase
DNase
Phenylalanase
^^■^*^^-
i 'm !■ pi p«
^
V}
3
S
3
REACTION
Acid
Alkaline
Coagulation
Reduction
Peptonization
No Change
i ■ ii i ■■«
423
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
Back Matter
Appendix A: Tables
© The McGraw-H
Companies, 2001
A
Appendix
Tabl
Table I International Atomic Weights
Element Symbol
Aluminum Al
Antimony Sb
Arsenic As
Barium Ba
Beryllium He
Bismuth Hi
Boron B
Bromine Br
Cadmium Cd
Calcium Ca
Chlorine CI
Chromium Cr
Cobalt Co
Copper Cu
Fluorine F
Cold Au
Hvdrogen „ H
Iodine I
Iron Fe
Lead Pb
Magnesium Mg
Manganese Mil
Mercury Hg
Nickel ' Ni
Nitrogen N
Oxygen O
Palladium Pd
Phosphorus P
Platinum Pt
Potassium K
Radium Ra
Selenium Se
Silicon Si
Silver Ag
Sodium Na
Strontium Sr
Sulfur S
Titanium Ti
Tungsten W
Uranium U
Vanadium V
Zinc Zn
Zirconium Zr
Atomic Atomic
Number Weight
13
26.97
51
121.76
33
74.91
56
137.36
4
9.013
83
209.00
5
10.82
35
79.916
48
1.12.41
20
40.08
6
12.010
17
35.457
24
52.01
27
58.94
29
63.54
9
19.00
79
197.2
1
1,0080
53
126.92
26
55.85
82
207.21
12
2432
25
51.93
80
200.61
28
58.69
i
14.008
8
16.0000
46
106.7
15
30.98
78
195.23
19
39.096
88
226.05
34
78.96
14
28.06
47
107.880
11
22.997
38
87.63
16
32.066
50
118.70
22
47.90
74
183.92
92
238.07
23
50.95
30
65.38
40
91.22
425
Benson: Microbiological
Back Matter
Appendix A: Tables
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Companies, 2001
Appendix A
Tabl
es
Table II Four-Place Logarithms
N
O
1
2
3
4
5
6
7
8
9
10
0000
0043
0086
0128
0170
0212
0253
0294
0334
0374
11
0414
0453
0492
0531
0569
0607
0645
0682
0719
0755
12
0792
0828
0864
0899
0934
0969
1004
1038
1072
1106
13
1139
1173
1206
1239
1271
1303
1335
1367
1399
1430
14
1461
1492
1523
1553
1584
1614
1644
1673
1703
1732
15
1761
1790
1818
1847
1875
1903
1931
1959
1987
2014
16 !
2041
2068
2095
2122
2148
2175
2201
2227
2253
2279
17
2304
2330
2355
2380
2405
2430
2455
2480
2504
2529
18
2553
2577
2601
2625
2648
2672
2695
2718
2742
2765
19
2788
2810
2833
2856
2878
2900
2923
2945
2967
2989
20
3010
3032
3054
3075
3096
3118
3139
3160
3181
3201
21
3222
3243
3263
C284
3304
3324
3345
3365
3385
3404
22
3424
3444
3464
3483
3502
3522
354 1
3560
3579
3598
23
3617
3636
3655
3674
3692
3711
3729
3747
3766
3784
24
3802
3820
3838
3856
3874
3892
3909
3927
3945
3962
25
3979
3997
4014
4031
4048
4065
4082
4099
4116
4133
26
4150
4166
4183
4200
4216
4232
4249
4265
4281
4298
27
4314
4330
4346
4362
4378
4393
4409
4425
4440
4456
28
4472
4487
4502
4518
4533
4548
4564
4579
4594
4609
29
4624
4639
4654
4669
4683
4698
4713
4728
4742
4757
30
4771
4786
4800
4814
4829 |
4843
4857
4871
4886
4900
31
4914
4928
4942
4955
4969
4983
4997
50 1 1
5024
5038
32
5051
5065
5079
5092
5105
5119
5132
5145
5159
5172
33
5185
5198
5211
5224
5237 ■
5250
5263
5276
5289
5302
34
5315
5328
5340
5353
5366
5378
5391
5403
5416
5428
35
5441
5453
5465
5478
5490
5502
55 1 4
5527
5539
5551
36
5563
5575
5587
5599
5611
5623
5635
5647
5658
5670
37
5682
5694
5705
5717
5729
5740
5752
5763
5775
5786
38
5798
5809
5821
5832
5843
5855
5866
5877
5888
5899
39
5911
5922
5933
5944
5955
5966
5977
5988
5999
6010
40
6021
6031
6042
6053
6064
6075
6085
6096
6107
6117
41
6128
6138
6149
6160
6170
6180
6191
6201
6212
6222
42
6232
6243
6253
6263
6274
6284
6294
6304
6314
6325
43
6335
6345
6355
6365
6375
6385
6395
6405
6415
6425
44
6435
6444
6454
6464
6474
6484
6493
6503
6513
6522
45
6532
6542
6551
6561
6571
6580
6590
6599
6609
6618
46
6628
6637
6646
6656
6665 ;
6675
6684
6693
6702
6712
47
6721
6730
6739
6749
6758
6767
6776
6785
6794
6803
48
6812
6821
6830
6839
6848
6857
6866
6875
6884
6893
49
6902
6911
6920
6928
6937
6946
6955
6964
6972
6981
50
6990
6998
7007
7016
7024
7033
7042
7050
7059
7067
51
7076
7084
7093
7101
7110
7118
7126
7135
7143
7152
52
7160
7168
7177
7185
7193
7202
7210
7218
7226
7235
53
7243
7251
7259
7267
7275
7284
7292
7300
7308
7316
54
7324
7332
7340
7348
7356
7364
7372
7380
7388
7396
N
1
2
3
4
5
6
7
8
9
426
Benson: Microbiological
Back Matter
Appendix A: Tables
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Companies, 2001
Tables • Appendix A
rable II
(continued)
N
1
2
3
4
5
6
7
8
9
55
7404
7412
7419
7427
7435
7443
7451
7459
7466
7474
56
7482
7490
7497
7505
7513
7520
7528
7536
7543
7551
57
7559
7566
7574
7582
7589
7597
7604
7612
7619
7627
58
7634
7642
7649
7657
7664
7672
7679
7686
7694
7701
59
7709
7716
7723
7731
7738
7745
7752
7760
7767
7774
60
7782
7789
7796
7803
7810
7818
7825
7832
7839
7846
61
7853
7860
7868
7875
7882
7889
7896
7903
7910
7917
62
7924
7931
7938
7945
7952
7959
7966
7973
7980
7987
63
7993
8000
8007
8014
802 1
8028
8035
8041
8048
8055
64
8062
8069
8075
8082
8089
8096
8102
8109
8116
8122
65
8129
8136
8142
8149
8156
8162
8169
8176
8182
8189
66
8195
8202
8209
8215
8222
8228
8235
8241
8248
8254
67
8261
8267
8274
8280
8287
8293
8299
8306
8312
8319
68
8325
8331
8338
8344
8351
8357
8363
8370
8376
8382
69
8388
8395
8401
8407
8414
8420
8426
8432
8439
8445
70
8451
8457
8463
8470
8476
8482
8488
8494
8500
8506
71
8513
8519
8525
8531
8537
8543
8549
8555
8561
8567
72
8573
8579
8585
8591
8597
8603
8609
8615
8621
8627
73
8633
8639
8645
8651
8657
8663
86S9
8675
8681
8686
74 I
8692
8698
8704
8710
8716
8722
8727
8733
8739
8745
75
8751
8756
8762
8768
8774
8779
8785
8791
8797
8802
76
8808
8814
8820
8825
8831
8837
8842
8848
8854
8859
77
8865
8871
8876
8882
8887
8893
8899
8904
8910
8915
78
8921
8927
8932
8938
8943
8949
8954
8960
8965
8971
79
8976
8982
8987
8993
8998
9004
9009
9015
9020
9025
SO
9031
9036
9042
9047
9053
9058
9063
9069
9074
9079
81
9085
9090
9096
9101
9106
9112
9117
9122
9128
9133
82
9138
9143
9149
9154
9159
9165
9170
9175
9180
9186
83
9191
9196
9201
9206
9212
9217
9222
9227
9232
9238
: 84
9243
9248
9253
9258
9263
9269
9274
9279
9284
9289
85
9294
9299
9304
9309
9315
9320
9325
9330
9335
9340
86
9345
9350
9355
9360
9365
9370
9375
9380
9385
9390
i 87
9395
9400
9405
9410
9415 ,
9420
9425
9430
9435
9440
88
9445
9450
9455
9460
9465
9469
9474
9479
9484
9489
89
9494
9499
9504
9509
9513
9518
9523
9528
9533
9538
90
9542
9547
9552
95,57
9562
9566
9571
9576
9581
9586
91
9590
9595
9600
9605
9609
9614
9619
9624
9628
9633
92
9638
9643
9647
9652
9657
9661
9666
9671
9675
9680
93
9685
9689
9694
9699
9703
9708
9713
9717
9722
9727
94
9731
9736
9741
9745
9750
9754
9759
9763
9768
9773
95
9777
9782
9786
9791
9795
9800
9805
9809
9814
9818
96
9823
9827
9832
9836
9841
9845
9850
9854
9859
9863
97
9868
9872
9877
9881
9886
9890
9894
9899
9903
9908
98 J
9912
9917
9921
9926
9930
9934
9939
9943
9948
9952
99
9956
9961
9965
9969
9974
9978
9983
9987
9991
9996
100
0000
0004
0009
0013
0017
0022
0026
0030
0035
0039
N
1
2
3
4
5
6
7
8
9
427
Benson: Microbiological
Back Matter
Appendix A: Tables
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Companies, 2001
Appendix A
Tabl
es
Table
Temperature Conversion Table Centigrade to Fahrenheit
c
1
2
3
4
5
6
7
i
8
9
-50
-58.0
-59.8
-61.6
-63.4
-65.2
-67,0
-68.8
-70.6
-72.4
-74.2
-40
40.0
-41.8
-43.6
-45.4
-47.2
-49.0
-50.8
-52.6
-54.4
-56.2
-30
-22.0
-23.8
-25.6
-27.4
-29.2
-31.0
-32.8
^34.6
-36.4
-38.2
-20
- 4.0
- 5.8
- 7.6
- 9.4
-11.2.
-13.0
-14.8
-16.6
-18.4
-20.2
-10
+ 14.0
+ 12.2
+ 10.4
+ 8.6
+ 6.8
+ 5.0
+ 3.2
+ 1.4
- 0.4
— 2,2
-
+32.0
+30.2
+28.4
+26.6
+24.8
+23.0
+21.2
+ 19.4
+ 17.6
+ 15.8
32.0
33.8
35.6
37.4
39.2
41.0
42.8
44.6
46.4
48.2
10
50.0
51.8
53.6
55.4
57.2
59.0
60.8
62.6
64.4
66.2
20
68.0
69.8
71.6
73.4
75.2
77.0
78.8 ;
80.6
82.4
84.2
30
86.0
87.8
89.6
91.4
93.2
95.0
96.8
98.6
100.4
102.2
40
104.0
105,8
107.6
109.4
111.2
113.0
114.8
116.6
118.4
120.2
50
122.0
123.8
125.6
127.4
129.2
131.0
132.8
134.6
136.4
138.2
60
140,0
141.8
143.6
145.4
147.2
149.0
150.8
152.6
154.4
156.2
70
158.0
159.8
161,6
163.4
165.2
167.0
168.8
170.6
172,4
174.2
80
176.0
177.8
179.6
181.4
183.2
185.0
186.8
188.6
190,4
192.2
90
194.0
195.8
197.6
199.4
201.2
203.0
204.8
206.6 :
208.4
210.2
100
; 212.0
213.8
215.6
217.4
219.2
221.0
222.8
224.6
226,4
228.2
110
230.0
231.8
233.6
235.4
237.2
239.0
240.8
242,6
244.4
246.2
120
248.0
249.8
251.6
253.4
255.2
257.0
258.8
260.6
262.4
264.2
130
266.0
267.8
269.6
271.4
273.2
275.0
276.8
278.6
280.4
282.2
140
284.0
285.8
287.6
289.4
291.2
2930
294.8
i 295.6
298.4
300.2
150
302.0
303.8
305.6
307.4
309.2
311.0
312.8
314.6
316.4
318.2
160
320.0
321.8
323.6
325.4
327.2
329,0
330.8
332.6
334.4
336.2
170
338.0
339.8
341.6
343.4
345.2
347.0
348.8
350.6
352.4
354.2
180
356.0
357.8
359.6
361.4
363.2
365.0
! 366.8 :
368.6
370.4
372,2
190
374.0
375.8
377.6
379.4
381.2
383.0
384.8
386.6
388.4
390.2
200
392.0
393.8
395.6
397.4
399.2
401,0
402.8
404.6
406.4
408.2
210
410.0
411.8
413.6
415.4
417.2
419.0
420.8
422.6
424.4
426.2
220
428.0
429.8
43L6
433.4
435.2
437.0
438.8
440.6
442.4
444.2
230
446,0
447.8
449.6
451.4
453.2
455.0
456.8
458.6
460.4
462.2
j 240
464.0
465.8
467.6
469.4
471.2
473-0
474.8
476.6
478,4
480.2
250
482.0
483.8
485.6
487.4
489.2
491.0
492.8
494.6
496.4
498.2
F
C X 9/5 + 32
°C
F
32 X 5/9
428
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
Back Matter
Appendix A: Tables
© The McGraw-H
Companies, 2001
Tabl
es
Appendix A
Table IV Autoclave Steam Pressures and Corresponding Temperatures
Steam
j
Steam
Steam
Pressure
lb/sq
Temperature
Pressure
lb/sq
Temperature
Pressure
lb/sq
Temperature
in
°C
°F
in
T
F
in
5 C
*F
100,0
212,0
1
101.9
215.4
11
116.4
241.5
21
126.9
260.4
2
103.6
218.5
12
117.6
243.7
127.8
262.0
3
105.3
221.5
13
118.8
245.8
23
128.7
263.7
4
106.9
224.4
14
119.9
247,8
24
129,6
265,3
108.4
227.1
15
121.0
249.8
25
130.4
266.7
6
109.S
229.6
16
122.0
251.6
26
131.3
268.3
7
111.3
232.3
17
123.0
253.4
27
132,1
269.8
8
112.6
234.7
18
1241
255.4
28
132.9
271.2
9
113.9
237.0
19
125.0
257.0 i
133.7
272.7
10
115.2
239.4
20
126.0
258.8
30
134.5
274.1
Figures are for steam pressure only and the presence of any air in the autoclave invalidates temperature read
ings from the above table.
429
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
Back Matter
Appendix A: Tables
© The McGraw-H
Companies, 2001
Appendix A
Tabl
es
Table V Autoclave Temperatures as Related to the Presence of Air
Gauge
Pressure,
lb
Pure steam,
complete air
discharge
Two-thirds
air discharge,
20-in. vacuum
One-half
air discharge,
15-in. vacuum
One-third
air discharge,
10-in« vacuum
No air
discharge
J C
°F
a C
°F
«C
*F
°C
°F
°C
'F
109
228
100
212
94
202
90
193
72
162
10
115
240
109
228
105
220
100
212
90
193
15
121
250
115
240
112
234
109
228
100
212
20
126
259
121
250
118
245
115
240
109
228
25
130
267
126
259
124
254
121
250
115
240
30
135
275
130
i
267
i
128
263
126
259
121 ;
250
430
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
Back Matter
Appendix A: Tables
© The McGraw-H
Companies, 2001
Tabl
es
Appendix A
Table VI MPN Determination from Multiple Tube Test
"
NUMBER
OF TUBES GIVING
% m rin.T
95 PERCENT
POSITIVE
REACTION OUT OF
MPN
Index
per
CONFIDENCE LIMITS
3 of 10
3 of 1
3 of 0.1
ml each
ml each
ml each
100 ml
Lower
Upper
1
3
<0.5
9
1
3
<0.5
13
1
4
<0.5
20
1
1
i
1
21
I
I
1
23
I
1
1
11
3
36
1
()
U
3
36
2
9
1
36
2
1
14
3
37
1
15
3
44
2
1
1
20
*
1
89
2
2
21
4
47
2
2
1
28
10
150
3
23
4
120
3
1
39 :
f
130
3
2
64
15
380
3
1
43
7
210
3
1
t
75
14
230
3
1
2
120
30
380
3
2
93
15
380
3
2
1
150
30
440
3
2
2
210
35
470
3
3
240
36
1,300
3
3
1
460
71
2,400
3
3
2
1,100
1 50
4,800
From Standard Methods for the Examination of Water and Wastewater, Twelfth edition. (New York: The American Public
Health Association, Inc.), p. 608.
431
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
Back Matter
Appendix A: Tables
© The McGraw-H
Companies, 2001
Appendix A
Tabl
es
Table VII Significance of zones of inhibition in Kirby-Bauer Method of antimicrobic sensitivity testing (1 995)
Antimicrobial
Disk
R
I
s
Agent
Potency
Resistant
Intermediate
Sensitive
mm
mm
mm
Amikacin
30 meg
<14
15-16
>17
Amoxicillin/Clavulinic Acid
30 meg
Staphylococci
<19
14-17
>20
Other gram-positive organisms
<13
14-17
>18
Ampiclllln
75 meg
Gram-negative enterics
<13
14-16
>17
Staphylococci
<28
>29
Enterococci
<16
>17
Streptococci (not S. pneumoniae)
<21
22-29
>30
Haemophilus spp.
i
<18
19-21
>22
Listeria monocytogenes
<19
>20
AzIocIlHn {Pseudomonas aeruginosa)
75 meg
<17
>18
CarbenlclJIln (P. aeruginosa)
100 meg
<13
14-16
>17
Other gram-negative organisms
<19
20-22
>23
Cefaclor
30 meg
<14
15-17
>18
Cephalothfn
30 meg
<14
15-17
>18
Chloramphenicol
30 meg
<12
13-17
I
>18
S. pneumoniae
<20
>21
Clarithromycin
15 meg
<13 |
14-17
>18
S, pneumoniae
<16
17-20
>21
Clindamycin
2 meg
<14
15-20
>21
S, pneumoniae
<20
>21
Erythromycin
15 meg
<13
14-22
>23
S. pneumoniae
<15
16-20
>21
Gentamlcin
10 meg
<12
13-14
>15
lmpenem
10 meg
<13
14-15
>16
Haemophilus spp.
>16
Kanamycin
30 meg
<13
14-17
>18
Lomelloxacln
10 meg
<18
19-21
>22
Loracarbef
30 meg
<14
15-17
>18
Mezlocillin {P. aeruginosa)
75 meg
<15 |
>16
Other gram-negative organisms
<17
18-20
>19
Minocycline
30 meg
<14
15-18
>19
Moxalactam
30 meg
<14
15-22
>23
Nafcillln
1 meg
<10
11-12
>13
Nalidixic Acid
30 meg
<13
14-18
>19
Netilmicin
30 meg
<12
13-14
>15
Norfloxacin
10 meg
<12
13-16
>17
Ofloxacin
5 meg
<12
13-15
>16
Penicillin G (Staphylococci)
10 units
<28 |
>29
Enterococci
<14
>15
Streptococci (not S, pneumoniae)
<19
20-27
>28
Neisseria gonorrhoeae
<26
27-46
>47
L monocytogenes
<19
>20
Piperaclllln/Tazobactam
100/10 meg
Staphylococci
<17
>18
P. aeruginosa
<17
>18
Other gram-negative organisms
<14
15-19
>20
Rifampin
5 meg
<16
17-19
>20
Haemophilus spp.
<16
17-19
>20
S. pneumoniae
I
<16
17-18
>19
Streptomycin
10 meg
<11
12-14
>15
Sulfisoxazole
300 meg
<12
13-16
>17
Tetracycline
30 meg
<14
15-18
>19
S. pneumoniae
<17
18-21
>22
Tobramycin
10mcg
<12
13-14
>15
Trimethoprim/Sulfamethoxazole
1.25/23.75
*10
11-15
>16
| Vancomycin
30 meg
<14
15-16
>17
432
Benson: Microbiological
Back Matter
Appendix A: Tables
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Companies, 2001
Tabl
es
Appendix A
Table VIII Indicators of Hydrogen Ion Concentration
Many of the following indicators are used in the media of certain exercises in this manual. This table indicates
the pH range of each indicator and the color changes that occur. To determine the exact pH within a particular
range one should use a set of standard colorimetric tubes that are available from the prep room. Consult your lab
instructor.
Indicator
Full Acid
Color
Full Alkaline
Color
pH Range
Cresol Red
red
yellow
0.2-1.8
Metacresol Purple (acid range)
red
yellow
1.2-2.8
Thymol Blue
red
yellow
1.2-2.8
i
Bromphenol Blue
yellow
blue
3.0-4.6
Bromcresol Green
yellow
blue
3.8-5.4
Chlorcresol Green
yellow
blue
4.0 - 5.6
Methyl Red
red
yellow
4.4 ~ 6.4
Chlorphenol Red
yellow
red
4.8 - 6.4
Bromcresol Purple
yellow
purple
5.2-6.8
Bromthymol Blue
yellow
blue
6.0-7.6
Neutral Red
red
amber
6.8 - 8.0
Phenol Red
yellow
red
6.8 - 8.4
Cresol Red
yellow
red
7.2 - 8.8
Metacresol Purple (alkaline range)
yellow
purple
7.4-9.0 :
Thymol Blue (alkaline range)
yellow
blue
8.0-9.6
Cresolphthalein
colorless
red
8.2 - 9.8
Phenolphthalein
colorless
red
8.3-10.0
433
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
Back Matter
Appendix B: Indicators,
Stains, Reagents
© The McGraw-H
Companies, 2001
Appendix
Indicators, Stains, Reagents
Indicators
All the indicators used in this manual can be made by (1) dissolving a measured amount of the indicator in 95%
ethanol, (2) adding a measured amount of water, and (3) filtering with filter paper. The following chart provides
the correct amounts of indicator, alcohol, and water for various indicator solutions.
Indicator Solution
Indicator
(gm)
95% Ethanol
(ml)
Distilled H 2
(ml)
-- — -• - —
Brorncresol green
0.4
500
500
Bromcresol purple
0.4
500
500
Bromthymol blue
0.4
500
500
Cresol red
02
500
500
Methyl red
0.2
500
500
Phenolphthalein
1.0
50
50
Phenol red
0.2
500
500
Thymol blue
0.4
500
500
Stains and Reagents
Acid-Alcohol (for Ziehl-Neelsen stain)
3 ml concentrated hydrochloric acid in 1 00 ml of
95% ethyl alcohol.
Acid-Alcohol (for fluorochrome staining)
HC1 2.5 ml
Ethyl alcohol, 70% 500.0 ml
NaCl 2.5 gm
Alcohol, 70% (from 95%)
Alcohol, 95% 368.0 ml
Distilled water 132.0 ml
Auramine-Rhodamine Stain
(for mycobacteria)
Auramine 3.0 gm
Rhodamine B 1.5 gm
Glycerol 150.0 ml
Phenol crystals (liquefied at 50° C) ... .20.0 ml
Distilled water 100.0 ml
Clarify by filtration with glass wool or
Whatman #2 filter paper. Do not heat. Store at
room temperature.
Barritt s Reagent (Voges-Proskauer test)
Solution A: 6 gm alpha- naphthol in 100 ml 95%
ethyl alcohol.
Solution B: 16 gm potassium hydroxide in 100 ml
water.
Note that no creatine is used in these reagents as
is used in O'Meara's reagent for the V-P test.
Carbolfuchsin Stain (Ziehl's)
Solution A: Dissolve 0.3 gm of basic fuchsin
(90% dye content) in 10 ml 95% ethyl
alcohol.
Solution B: Dissolve 5 gm of phenol in 95 ml of
water.
Mix solutions A and B .
Crystal Violet Stain (Hucker modification)
Solution A: Dissolve 2.0 gm of crystal violet
(85% dye content) in 20 ml of 95% ethyl
alcohol.
Solution B: Dissolve 0.8 gm ammonium oxalate
in 80.0 ml distilled water.
Mix solutions A and B .
435
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
Back Matter
Appendix B: Indicators,
Stains, Reagents
© The McGraw-H
Companies, 2001
Appendix B
Indicators, Stains, Reagents
Diphenylamine Reagent (nitrate test)
Dissolve 0.7 gm diphenylamine in a mixture of 60
ml of concentrated sulfuric acid and 28.8 ml of
distilled water.
Cool and add slowly 11.3 ml of concentrated
hydrochloric acid. After the solution has stood for
12 hours some of the base separates, showing that
the reagent is saturated.
Ferric Chloride Reagent (Ex. 77)
FeQ r 6H 2 12 gm
2% Aqueous HC1 100 ml
Make up the 2% aq. HC1 by adding 5.4 ml of con-
centrated HC1 (37%) to 94.6 ml H 2 0. Inoculate
with two or three colonies of beta hemolytic
streptococci, incubate at 35° C for 20 or more
hours. Centrifuge the medium to pack the cells,
and pipette 0.8 ml of the clear supernate into a
Kahn tube. Add 0.2 ml of the ferric chloride
reagent to the Kahn tube and mix well. If a heavy
precipitate remains longer than 10 minutes, the
test is positive.
Gram's Iodine (Lugol's)
Dissolve 2.0 gm of potassium iodide in 300 ml of
distilled water and then add 1 .0 gm iodine crystals.
Iodine, 5% Aqueous Solution (Ex. 41)
Dissolve 4 gm of potassium iodide in 300 ml of
distilled water and then add 2.0 gm iodine crystals.
ride in 100 ml of 95% ethyl alcohol. Let stand
overnight at room temperature to complete
dissolution.
Solution B: Dissolve 3 gm of tannic acid in 100
ml of distilled water.
Solution C: Dissolve 1.5 gm of sodium chloride
in 100 ml of distilled water.
Mix equal volumes of solutions A, B , and C and let
stand for 2 hours. Store in a stoppered bottle in a re-
frigerator (up to 2 months). Disregard precipitate
that forms in bottom of bottle. Do not filter. Will
store indefinitely, if frozen. Frozen stain solution
must be thoroughly mixed after thawing since the
water separates from the alcohol. After mixing, the
precipitate should be allowed to settle to the bottom.
Note: Pararosaniline compounds should be certi-
fied for flagellar staining.
Malachite Green Solution (spore stain)
Dissolve 5.0 gm malachite green oxalate in 100
ml distilled water.
JVlcFarland Nephelometer Barium
Sulfate Standards (Ex. 55)
Prepare 1% aqueous barium chloride and 1%
aqueous sulfuric acid solutions.
Add the amounts indicated in table 1 to clean,
dry ampoules. Ampoules should have the same
diameter as the test tube to be used in subsequent
density determinations.
Seal the ampoules and label them.
Kovacs' Reagent (indole test)
n-amyl alcohol 75.0 ml
Hydrochloric acid (cone.) 25.0 ml
p-dimethylamine-benzaldehyde 5.0 gm
Lactophenol Cotton Blue Stain
Phenol crystals 20 gm
Lactic acid 20 ml
Glycerol 40 ml
Cotton blue 0.05 gm
Dissolve the phenol crystals in the other ingredi-
ents by heating the mixture gently under a hot wa-
ter tap.
Leifson s Flagellar Stain
Prepare three separate solutions as follows:
Solution A: Dissolve 0.9 gm of pararosaniline ac
etate and 0.3 gm of pararosaniline hydrochlo
Table 1
Amounts for Standards
Tube
Barium
Chloride
1 % (ml)
Sulfuric
Acid
1 % (ml)
Corresponding
Approx.
Density of
Bacteria
(million/ml)
1
0.1
9.9
300
2
0.2
9.8
600
3
0.3
9.7
900
4
0.4
9.6
1200
5
0.5
9.5
1500
6
0.6
9.4
1800
7
0.7
9.3
2100
8
0.8
9.2
2400
9
0.9
9.1
2700
10
1.0
9.0
3000
436
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
Back Matter
Appendix B: Indicators,
Stains, Reagents
© The McGraw-H
Companies, 2001
Indicators, Stains, Reagents
Appendix B
Methylene Blue (Loeffler's)
Solution A: Dissolve 0.3 gm of methylene blue
(90% dye content) in 30.0 ml ethyl alcohol
(95%).
Solution B: Dissolve 0.01 gm potassium hydrox-
ide in 100.0 ml distilled water. Mix solutions
AandB.
Naphthol, alpha
5% alpha-naphthol in 95% ethyl alcohol
Caution: Avoid all contact with human tissues
Alpha-naphthol is considered to be carcino-
genic.
Nessler s Reagent (ammonia test)
Dissolve about 50 gm of potassium iodide in 35
ml of cold ammonia-free distilled water. Add a
saturated solution of mercuric chloride until a
slight precipitate persists. Add 400 ml of a 50%
solution of potassium hydroxide. Dilute to 1 liter,
allow to settle, and decant the supernatant for use.
Nigrosine Solution (Dorner's)
Nigrosine, water soluble 10 gm
Distilled water 100 ml
Boil for 30 minutes. Add as a preservative 0.5 ml
formaldehyde (40%). Filter twice through double
filter paper and store under aseptic conditions.
Nitrate Test Reagent
(see Diphenylamine)
Nitrite Test Reagents
Solution A: Dissolve 8 gm sulfanilic acid in 1000
ml 5N acetic acid (1 part glacial acetic acid to
2.5 parts water).
Solution B: Dissolve 5 gm dimethyl- alpha-naph-
thylamine in 1000 ml 5N acetic acid. Do not
mix solutions.
Caution: Although at this time it is not known for
sure, there is a possibility that dimethyl-a-
naphthylamine in solution B may be carcino-
genic. For reasons of safety, avoid all contact
with tissues.
Oxidase Test Reagent
Mix 1.0 gm of dimethyl- p-phenylenedi amine hy
drochloride in 100 ml of distilled water.
Preferably, the reagent should be made up
fresh, daily. It should not be stored longer than
one week in the refrigerator. Tetramethyl-p-
phenylenediamine dihy drochloride (1%) is even
more sensitive, but is considerably more expen-
sive and more difficult to obtain.
Phenolized Saline
Dissolve 8.5 gm sodium chloride and 5.0 gm phe
nol in 1 liter distilled water.
Physiological Saline
Dissolve 8.5 gm sodium chloride in 1 liter dis
tilled water.
Potassium permanganate
(for fluorochrome staining)
KMn0 4 2.5 gm
Distilled water 500.0 ml
Safranin (for gram staining)
Safranin O (2.5% sol'n in 95% ethyl
alcohol) 10.0 ml
Distilled water 100.0 ml
Trommsdorf s Reagent (nitrite test)
Add slowly, with constant stirring, 100 ml of a
20% aqueous zinc chloride solution to a mixture
of 4.0 gm of starch in water. Continue heating un-
til the starch is dissolved as much as possible, and
the solution is nearly clear. Dilute with water and
add 2 gm of potassium iodide. Dilute to 1 liter, fil-
ter, and store in amber bottle.
Vaspar
Melt together 1 pound of Vaseline and 1 pound of
paraffin. Store in small bottles for student use.
Voges-Proskauer Test Reagent
(see Barritt's)
White Blood Cell (WBC) Diluting Fluid
Hydrochloric acid 5 ml
Distilled water 495 ml
Add 2 small crystals of thymol as a preservative.
437
Benson: Microbiological
Back Matter
Appendix C: Media
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Companies, 2001
Appendix
Medi
la
Conventional Media The following media are used in the experiments of this manual. All of these media are
available in dehydrated form from either Difco Laboratories, Detroit, Michigan, or Baltimore Biological
Laboratory (BBL), a division of Becton, Dickinson & Co., Cockeysville, Maryland. Compositions, methods of
preparation, and usage will be found in their manuals, which are supplied upon request at no cost. The source of
each medium is designated as (B) for BBL and (D) for Difco.
Bile esculin (D)
Brewer's anaerobic agar (D)
Desoxycholate citrate agar (B,D)
Desoxycholate lactose agar (B,D)
DNase test agar (B,D)
Endo agar (B,D)
Eugonagar (B,D)
Fluid thioglycollate medium (B,D)
Heart infusion agar (D)
Hektoen Enteric Agar (B,D)
Kligler iron agar (B,D)
Lead acetate agar (D)
Levine EMB agar (B,D)
Lipase reagent (D)
Litmus milk (B,D)
Lowenstein- Jensen medium (B,D)
MacConkey Agar (B,D)
Mannitol salt agar (B,D)
MR- VP medium (D)
Mueller- Hinton medium (B,D)
Nitrate broth (D)
Nutrient agar (B,D)
Nutrient broth (B,D)
Nutrient gelatin (B,D)
Phenol red sucrose broth (B,D)
Phenylalanine agar (D)
Phenylethyl alcohol medium (B)
Russell double sugar agar (B,D)
Sabouraud's glucose (dextrose) agar (D)
Semisolid medium (B)
Simmons citrate agar (B,D)
Snyder test agar (D)
Sodium hippurate (D)
Spirit blue agar (D)
SS agar (B,D)
m-Staphylococcus broth (D)
Staphylococcus medium 110 (D)
Starch agar (D)
Trypticase soy agar (B)
Trypticase soy broth (B)
Tryptone glucose extract agar (B,D)
Urea (urease test) broth (B,D)
Veal infusion agar (B,D)
Xylose Lysine Desoxycholate Agar (B,D)
Special Media The following media are not included in the manuals that are supplied by Difco and BBL; there-
fore, methods of preparation are presented here.
Ammonium Medium (for N it rod o monad)
(NH 4 ) 2 S0 4 2.0 gm
MgS0 4 • 7H 2 0.5 gm
FeS0 4 • 7H 2 0.03 gm
NaCl 0.3 gm
MgC0 3 10.0 gm
K 2 HP0 4 1.0 gm
Water 1000.0 ml
For the isolation of Nitrosomonas from soil, steriliza-
tion is not necessary if the inoculations are made as
soon as the medium is made up. The pH should be ad-
justed to 7.3. If sterilization is desirable for storage or
other reasons, adjust the pH aseptically after steriliza-
tion with sterile IN HC1.
Bile Esculin Slants (Ex. 79)
Heart infusion agar 40.0 gm
Esculin 1.0 gm
Ferric chloride 0.5 gm
Distilled water 1000.0 ml
Dispense into sterile 15 X 1 25 mm screw-capped
tubes, sterilize in autoclave at 121° C for 15 min-
utes, and slant during cooling.
Blood Agar
Trypticase soy agar powder 40 gm
Distilled water 1000 ml
Final pH of 7.3
Defibrinated sheep or rabbit blood 50 ml
439
Benson: Microbiological
Back Matter
Appendix C: Media
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Companies, 2001
Appendix C
Media
Liquefy and sterilize 1000 ml of trypticase soy
agar in a large Erlenmeyer flask. While the TSA
is being sterilized, warm up 50 ml of defibrinated
blood to 50° C. After cooling the TSA to 50° C,
aseptically transfer the blood to the flask and mix
by gently rotating the flask (cold blood may cause
lumpiness).
Pour 10-12 ml of the mixture into sterile Petri
plates. If bubbles form on the surface of the
medium, flame the surface gently with a Bunsen
burner before the medium solidifies. It is best to
have an assistant to lift off the Petri plate lids
while pouring the medium into the plates. A full
flask of blood agar is somewhat cumbersome to
handle with one hand.
Bromthymol Blue Carbohydrate Broths
Make up stock indicator solution:
Bromthymol blue 8 gm
95% ethyl alcohol 250 ml
Distilled water 250 ml
Indicator is dissolved first in alcohol and then wa-
ter is added.
Make up broth:
Sugar base (lactose, sucrose, glucose, etc.) .5 gm
Tryptone 10 gm
Yeast extract 5 gm
Indicator solution 2 ml
Distilled water 1000 ml
Final pH 7.0
Deca-Strength Phage Broth (Ex. 28)
Peptone 100 gm
Yeast extract 50 gm
NaCl 25 gm
K 2 HP0 4 80 gm
Distilled water 1000 ml
Final pH 7.6
Emmons' Culture Medium for Fungi
C. W. Emmons developed the following recipe
as an improvement over Sabouraud's glucose
agar for the cultivation of fungi. Its principal ad-
vantage is that a neutral pH does not inhibit cer-
tain molds that have difficulty growing on
Sabouraud's agar (pH 5.6). Instead of relying on
a low pH to inhibit bacteria, it contains chlor-
amphenicol, which does not adversely affect the
fungi.
Glucose 20 gm
Neopeptone 10 gm
Agar 20 gm
Chloramphenicol 40 mg
Distilled water 1000 ml
After the glucose, peptone, and agar are dis-
solved, heat to boiling, add the chloramphenicol
which has been suspended in 10 ml of 95% alco-
hol and remove quickly from the heat. Autoclave
for only 10 minutes.
Glucose— Minimal Salts Agar
(Ex. 76, Ames test)
This medium is made from glucose, agar, and
Vogel-Bonner medium E (50 X).
Vogel-Bonner Medium E (50 X )
Distilled water (45° C) 670 ml
MgS0 4 • 7H 2 10 gm
Citric acid monohydrate 1 00 gm
K 2 HP0 4 (anhydrous) 500 gm
Sodium ammonium phosphate
(NaHNH 4 P0 4 • 4H 2 0) 175 gm
Add salts in the order indicated to warm water
(45° C) in a 2-liter beaker or flask placed on a
magnetic stirring hot plate. Allow each salt to dis-
solve completely before adding the next. Adjust
the volume to 1 liter. Distribute into two 1 -liter
glass bottles. Autoclave, loosely capped, for 20
minutes at 121° C.
Plates of Glucose-Minimal Salts Agar
Agar 15 gm
Distilled water 930 ml
50X V-B salts 20 ml
40% glucose 50 ml
Add 15 gm of agar to 930 ml of distilled water in
a 2- liter flask. Autoclave for 20 minutes using
slow exhaust. When the solution has cooled
slightly, add 20 ml of sterile 50 X V-B salts and 50
ml of sterile 40% glucose. For mixing, a large
magnetic stir bar can be added to the flask before
autoclaving. After all the ingredients have been
added, the solution should be stirred thoroughly.
Pour 30 ml into each Petri plate. Important: The
50 X V-B salts and 40% glucose should be auto-
claved separately.
440
Benson: Microbiological
Back Matter
Appendix C: Media
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Companies, 2001
Media
Appendix C
Glucose Peptone Acid Agar
Glucose 10 gm
Peptone 5 gm
Monopotassium phosphate 1 gm
Magnesium sulfate (MgS0 4 • 7H 2 0) 0.5 gm
Agar 15 gm
Water 1000 ml
While still liquid after sterilization, add sufficient
sulfuric acid to bring the pH down to 4.0.
Glycerol Yeast Extract Agar
Glycerol 5 ml
Yeast extract 2 gm
Dipotassium phosphate 1 gm
Agar 15 gm
Water 1000 ml
m Endo MF Broth (Ex. 64)
This medium is extremely hygroscopic in the de-
hydrated form and oxidizes quickly to cause dete-
rioration of the medium after the bottle has been
opened. Once a bottle has been opened it should
be dated and discarded after one year. If the
medium becomes hardened within that time it
should be discarded. Storage of the bottle inside a
larger bottle that contains silica gel will extend
shelf life.
Failure of Exercise 64 can often be attributed
to faulty preparation of the medium. It is best to
make up the medium the day it is to be used. It
should not be stored over 96 hours prior to use.
The Millipore Corporation recommends the fol-
lowing method for preparing this medium. (These
steps are not exactly as stated in the Millipore
Application Manual AM302.)
1 . Into a 250 ml screw-cap Erlenmeyer flask place
the following:
Distilled water 50 ml
95% ethyl alcohol 2 ml
Dehydrated medium (m Endo MF
broth) 4.8 gm
Shake the above mixture by swirling the flask un-
til the medium is dissolved and then add another
50 ml of distilled water.
2. Cap the flask loosely and immerse it into a pan of
boiling water. As soon as the medium begins to
simmer, remove the flask from the water bath. Do
not boil the medium any further.
3. Cool the medium to 45° C, and adjust the pH to
between 7.1 and 7.3.
4. If the medium must be stored for a few days, place
it in the refrigerator at 2°-10° C, with screw-cap
tightened securely.
Milk Salt Agar (15% NaCl)
Prepare three separate beakers of the following
ingredients:
1. Beaker containing 200 grams of sodium chloride.
2. Large beaker (2000 ml size) containing 50 grams
of skim milk powder in 500 ml of distilled water.
3. Glycerol-peptone agar medium:
MgS0 4 • 7H 2 5.0 gm
MgN0 3 • 6H 2 1.0 gm
FeCl 3 • 7H 2 0.025 gm
Difco proteose-peptone #3 5.0 gm
Glycerol 10.0 gm
Agar 30.0 gm
Distilled water 500.0 ml
Sterilize the above three beakers separately. The
milk solution should be sterilized at 113°- 115°
C (8 lb pressure) in autoclave for 20 minutes.
The salt and glycerol-peptone agar can be steril-
ized at conventional pressure and temperature.
After the milk solution has cooled to 55° C, add
the sterile salt, which should also be cooled
down to a moderate temperature. If the salt is too
hot, coagulation may occur. Combine the milk-
salt and glycerol-peptone agar solutions by gen-
tly swirling with a glass rod. Dispense asepti-
cally into Petri plates.
Nitrate Broth
Beef extract 3 gm
Peptone 5 gm
Potassium nitrate 1 gm
Distilled water 1000 ml
Final pH 7.0 at 25° C
Nitrate Agar
Beef extract 3 gm
Peptone 5 gm
Potassium nitrate 1 gm
Agar 12 gm
Distilled water 1000 ml
Final pH 6.8 at 25° C
441
Benson: Microbiological
Back Matter
Appendix C: Media
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Companies, 2001
Appendix C
Media
Nitrate Succinate— Mineral Salts Broth
(Ex. 62)
This medium is used as an enrichment medium
for isolating denitrifying bacteria from soil. Note
that two stock solutions (A and B) should be made
up before attempting to put together the complete
medium.
Solution A (Trace Mineral Salts)
FeS0 4 • 7H 2 300 mg
MnCl 2 • 4H 2 180 mg
Co(N0 3 )2 • 7H 2 130 mg
ZnS0 4 • 7H 2 40 mg
H 2 Mo0 4 20 mg
CuS0 4 • 5H 2 1 mg
CaCl 2 1000 mg
This solution should be stored at 4° C until used.
Solution B
NH 4 C1 1 gm
Na 2 HP0 4 2.14 gm
KH 2 P0 4 1.09 gm
MgS0 4 • 7H 2 0.2 gm
Trace mineral salts (Sol A) 10 ml
Water to 1000 ml
Complete Medium
Solution B 1000 ml
Sodium succinate 2 gm
Potassium nitrate 3 gm
Adjust the pH to 6.8, dispense into bottles, and
autoclave at standard conditions.
Nitrate Succinate— Mineral Salts Agar
(Ex. 62)
Add 15 g agar to 1000 ml of the above complete
medium. Dispense into Petri plates and sterilize in the
autoclave.
Nitrite Medium (for Nltrobacter)
NaN0 2 1.0 gm
MgS0 4 • 7H 2 0.5 gm
FeS0 4 • 7H 2 0.03 gm
NaCl 0.3 gm
Na 2 C0 3 1.0 gm
K 2 HP0 4 1.0 gm
Water 1000.0 ml
For the isolation of Nitrobacter from soil, steril-
ization is not necessary if the inoculations are
made as soon as the medium is made up. The pH
should be adjusted to 7.3. If sterilization is desir-
able for storage or other reasons, adjust the pH
aseptically after sterilization with sterile IN HC1.
Nitrogen-Free Glucose Agar (Ex. 59)
Add 1 5 grams of agar to the basal salts portion of
the above recipe, bring to a boil, and sterilize in
the autoclave at 121° C for 15 minutes. The glu-
cose is dissolved in 100 ml of water and sterilized
separately in similar manner. Mix the two solu-
tions aseptically, and dispense into sterile Petri
plates.
Nitrogen-Free Medium
(Ex. 59, Azotobacter)
K 2 HP0 4 1.0 gm
MgS0 4 • 7H 2 0.2 gm
FeS0 4 • 7H 2 0.05 gm
CaCl 2 • 2H 2 0.1 gm
Na 2 Mo0 4 • 2H 2 0.001 gm
*Glucose 10.0 gm
Distilled water 1000 ml
* Sterilize separately.
Adjust pH to 7.2.
If this medium is to be used immediately to iso-
late Azotobacter from soil, sterilization is not nec-
essary. When it must be stored for any length of
time, it should be sterilized.
If it is to be sterilized, the glucose should be
dissolved separately in 100 ml of water and ster-
ilized at 121° C for 15 minutes. The remainder of
the medium is sterilized in a similar manner.
After sterilization, the two solutions are
mixed aseptically and dispensed into sterile 8-oz
prescription bottles (50 ml per bottle).
442
Benson: Microbiological
Back Matter
Appendix C: Media
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Companies, 2001
Media
Appendix C
Phage Growth Medium (Ex. 29)
KH 2 P0 4 1.5 gm
Na 2 HP0 4 3.0 gm
NH 4 C1 1.0 gm
MgS0 4 • 7H 2 0.2 gm
Glycerol 10.0 gm
Acid-hydrolyzed casein 5.0 gm
dl- Tryptophan 0.01 gm
Gelatin 0.02 gm
Tween-80 0.2 gm
Distilled water 1000.0 ml
Sterilize in autoclave at 121° C for 20 minutes.
Phage Lysing Medium (Ex. 29)
Add sufficient sodium cyanide (NaCN) to the
above growth medium to bring the concentration
up to 0.02M. For 1 liter of lysing medium this will
amount to about 1 gram (actually 0.98 gm) of
NaCN. When an equal amount of this lysing
medium is added to the growth medium during
the last 6 hours of incubation, the concentration of
NaCN in the combined medium is 0.01 M.
Rhodospirillaceae Medium (Ex. 27)
This culture medium is used for the enrichment
and culture of anaerobic phototrophic bacteria. To
make up this medium you need to first prepare
three stock solutions (A, B, and C) before putting
together the entire batch.
A. Iron Citrate Solution
Ammonium ferrous sulfate 748 mg
Sodium citrate 1180 mg
Water to 500 ml
Store this stock solution at 4° C until needed.
Complete Enrichment Medium
The final batch of this medium has the following
ingredients. The succinate provides the organic
carbon and the yeast extract provides essential vi-
tamins for certain strains.
KH 2 P0 4 0.5 gm
MgS0 4 • 7H 2 0.2 gm
NaCl 0.4 gm
NH 4 C1 0.4 gm
CaCl 2 • 2H 2 0.05 gm
Sodium succinate 1.0 gm
Yeast extract 0.2 gm
Iron citrate solution (A) 5 ml
Vitamin B 12 solution (B) 0.1 ml
Trace elements solution (C) 1 ml
Water to 1000 ml
Adjust the pH to 6.8, dispense into bottles and au-
toclave at standard conditions.
Russell Double Sugar Agar (Ex. 80)
Beef extract 1 gm
Proteose Peptone No. 3 (Difco) 12 gm
Lactose 10 gm
Dextrose 1 gm
Sodium chloride 5 gm
Agar 15 gm
Phenol red (Difco) 0.025 gm
Distilled water 1000 ml
Final pH 7.5 at 25° C
Dissolve ingredients in water, and bring to boil-
ing. Cool to 50°-60° C, and dispense about 8 ml
per tube (16 mm dia tubes). Slant tubes to cool.
Butt depth should be about V".
B. Vitamin B 12 Solution
Certain strains require this vitamin. To make up
100 ml of this solution add 1 mg to 100 ml of wa-
ter. Store at 4° C until needed.
C. Trace Metals Solution
H 3 B0 4 2.86 gm
MnCl 2 • H 2 1.81 gm
ZnS0 4 • 7H 2 0.222 gm
Na 2 Mo0 4 • 2H 2 0.390 gm
CuS0 4 • 5H 2 0.079 gm
Co(N0 3 ) 2 • 6H 2 0.0494 gm
Water to 1000 ml
Store at 4° C until needed.
Skim Milk Agar
Skim milk powder 100 gm
Agar 15 gm
Distilled water 1000 ml
Dissolve the 15 gm of agar into 700 ml of distilled
water by boiling. Pour into a large flask and ster-
ilize at 121° C, 15 lb pressure.
In a separate container, dissolve the 100 gm
of skim milk powder into 300 ml of water heated
to 50° C. Sterilize this milk solution at 113°-115°
C (8 lb pressure) for 20 minutes.
After the two solutions have been sterilized,
cool to 55° C and combine in one flask, swirling
gently to avoid bubbles. Dispense into sterile
Petri plates.
443
Benson: Microbiological
Back Matter
Appendix C: Media
©The McGraw-Hill
Applications Lab Manual,
Eighth Edition
Companies, 2001
Appendix C
Media
Sodium Chloride (6.5%) Tolerance
Broth (Ex. 79)
Heart infusion broth 25 gm
NaCl 60 gm
Indicator (1.6 gm bromcresol purple in 100 ml
95% ethanol) 1 ml
Dextrose 1 gm
Distilled water 1000 ml
Add all reagents together up to 1 000 ml (final vol-
ume). Dispense in 15X125 mm screw-capped
tubes and sterilize in an autoclave 15 minutes at
121° C.
A positive reaction is recorded when the indi-
cator changes from purple to yellow or when
growth is obvious even though the indicator does
not change.
Sodium Hippurate Broth (Ex. 79)
Heart infusion broth 25 gm
Sodium hippurate 10 gm
Distilled water 1000 ml
Sterilize in autoclave at 121° C for 15 minutes af-
ter dispensing in 15 X 125 mm screw-capped
tubes. Tighten caps to prevent evaporation.
Soft Nutrient Agar (for bacteriophage)
Dehydrated nutrient broth 8 gm
Agar 7 gm
Distilled water 1000 ml
Sterilize in autoclave at 121° C for 20 minutes.
Spirit Blue Agar (Ex. 49)
This medium is used to detect lipase production
by bacteria. Lipolytic bacteria cause the medium
to change from pale lavender to deep blue.
Spirit blue agar (Difco) 35 gm
Lipase reagent (Difco) 35 ml
Distilled water 1000 ml
Dissolve the spirit blue agar in 1000 ml of water
by boiling. Sterilize in autoclave for 15 minutes at
15 psi (121° C). Cool to 55° C and slowly add the
35 ml of lipase reagent, agitating to obtain even
distribution. Dispense into sterile Petri plates.
Streptomycin Agar (Ex. 74)
To 1000 ml sterile liquid nutrient agar (50° C),
aseptically add 100 mg of streptomycin sulfate.
Pour directly into sterile Petri plates.
Top Agar (Ex. 76, Ames test)
Tubes containing 2 ml of top agar are made up
just prior to using from bottles of top agar base
and his/bio stock solution.
His/Bio Stock Solution
D-Biotin (F.W. 247.3) 30.9 mg
L-Histidine • HC1 (F.W. 191.7) 24.0 mg
Distilled water 250 ml
Dissolve by heating the water to the boiling point.
This can be done in a microwave oven. Sterilize
by filtration through 0.22 (xm membrane filter, or
autoclave for 20 minutes at 121° C. Store in a
glass bottle at 4° C.
Top Agar Base
Agar 6 gm
Sodium chloride (NaCl) 5 gm
Distilled water 1000 ml
The agar may be dissolved in a steam bath or mi-
crowave oven, or by autoclaving briefly. Mix
thoroughly and transfer 100-ml aliquots to 250-
ml glass bottles with screw caps. Autoclave for 20
minutes with loosened caps. Slow exhaust. Cool
the agar and tighten caps.
Just before using, add 10 ml of the his/bio
stock solution to bottle of 100 ml of liquefied top
agar base (45° C). After thoroughly mixing, dis-
tribute, aseptically, 2 ml of this mixture to sterile
tubes (13 mm X 100 mm). Hold tubes at 45° C
until used.
Tryptone Agar
Tryptone 10 gm
Agar 15 gm
Distilled water 1000 ml
Tryptone Broth
Tryptone 10 gm
Distilled water 1000 ml
Tryptone Yeast Extract Agar
Tryptone 10 gm
Yeast extract 5 gm
Dipotassium phosphate 3 gm
Sucrose 50 gm
Agar 15 gm
Water 1000 ml
pH7.4
444
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
Back Matter
Appendix D: Identification
Charts
© The McGraw-H
Companies, 2001
D
Appendix
Identification Charts
445
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
Back Matter
Appendix D: Identification
Charts
© The McGraw-H
Companies, 2001
Appendix D
Identification Charts
Chart I Interpretation of Test Results of API 20E System
Interpretation of Reactions
Tube
Positive Negative
Comments
ONPG
Yellow Colorless
(1) Any shade of yellow is a positive reaction.
(2) VP tube, before the addition of reagents, can be used as a
negative control.
ADH
Incubation
1 8-24 h Red or Orange Yellow
1 36-48 h Red Yellow or Orange
Orange reactions occurring at 36^48 hours should be interpreted as
negative.
LDC
18-24 h Red or Orange Yellow
36-48 h Red Yellow or Orange
Any shade of orange wrthin 18-24 hours is a positive reaction. At 36—
48 hours, orange decarboxylase reactions should be interpreted as
negative.
ODC
18-24 h Red or Orange Yellow
36-46 h Red Yellow or Orange
Orange reactions occurring at 36-48 hours should be interpreted as
negative.
CIT
Turquoise or Dark Light Green or Yellow
Blue
(1) Both the tube and cupufe should be filled.
{2> Reaction is read in the aerobic (cupule) area-
H 2 S
Black Deposit No Black Deposit
(1) HaS production may range from a heavy black deposit to a very
thin black line around the tube bottom. Carefully examine the bottom
of the tube before considering the reaction negative. (2) A
'browning" of the medium is a negative reaction unless a black
deposit ia present. li Browning M occurs with TDA-posrtive organisms.
URE
1 8-24 h Red or Orange Yellow
36-48 h Red Yellow or Orange
A method of lower sensitivity has been chosen. Klebsiella, Proteus*
and Y&rsinia routinely give positivs reactions.
TDA
Add 1 drop 10% ferric chloride
(1) Immediate reaction. (2) Indole-positrve organisms may produce a
golden orange color due to indole production. This is a negative
reaction.
Brown -Red Yeltow
IND
Add 1 drop Kovacs' reagent
(1) The reaction should be read within 2 minutes after the addition of
the Kovacs' reagent and the results recorded, (2) After several
minutes, the HCI present rn Kovacs reagent may react with the
plastic of the cupule resulting in a change from a negative {yeltow}
color to a brownish-red. This is a negative reaction.
Red Ring Yellow
VP
Add 1 drop of 40% potassium hydroxide, then 1 drop of 6% alpha—
naphthol.
{1 1 Wait 10 minutes before considering the reaction negative.
(2) A pale pink color {after 10 min ) should be interpreted as negative.
A pale pink color appears immediately after the addition of
reagents but turns dark pink or red after 10 min should be
interpreted as positive.
Motility may be ob&erveci by hanging drop or wet mount preparation.
Red Colorless
GEL
Diffusion ol the No diffusion
pigment
(1) The solid gelatin particles may spread throughout the tube after
inoculation. Unless diffusion occurs, the reaction is negative. (2) Any
degree of diffusion is a positive reaction.
GLU
MAN
INO
SOR
RHA
SAC
MEL
AMY
ARA
Yellow or Gray Blue or
Blue-Green
Yellow Blue or
Blue-Green
Comments for all
Carbohydrates
Fermentation (Enterobacteriaceae, Aeromonas, Vibrio)
(1) Fermentation of the carbohydrates begins in the most
anaerobic portion (bottom) of the tube. Therefore, these
reactions should be read from the bottom of the tube to
the top, (2) A yellow cok>r at the bottom of the tube only
indicates a weak or delayed positive reaction.
Oxidation (Other Gram-negatives)
(1) Oxidative utilization of the carbohydrates begins #n the
most aerobic portion (top) of the tube. Therefore, these
reactions should be read from the top to the bottom of the
tube. (2) A yellow color in the upper portion of the tube
and a blue in the bottom of the tube indicates oxidative
utilization of the sugar. This reaction should be considered
positive only for non-Enterobacteriaceae gram-negative
rods, This is a negative reaction for fermentative
organisms such as Enterobacteriaceae.
GLU
Nitrate
Reduction
After reading GLU reaction, add 2 drops 0.8% sulfanilic acid and 2
drops 0.5% N, N-dJmethylalpha-naphthylamine
NO^ Red Yellow
N2 gas Bubbles; Yellow after Orange after
reagents and zinc reagents and zinc
(1) Before addition of reagents, observe GLU tube (positive or
negative) for bubbles. Bubbles are indicative of reduction of nitrate to
the nitrogenous {N2) state. (2) A positive reaction may take 2-3
minutes for the red color to appear. (3) Confirm a negative test by
adding zinc dust or 20-mesh granular zinc. A pink-orange color after
10 minutes confirms a negative reaction. A yellow color indicates
reduction of nitrates to nitrogenous (Na) state.
MAN
INO
SOR
Catalase
After reading carbohydrate reactk>n, add 1 drop 1 .5% H2O2
(1) Bubbles may take 1-2 minutes to appear (2) Best results wilt be
1.4 ■ h _* _ B _ _■ •_■ _ 1 J _ _
Bubbles No bubbles
obtained 1
fermentat
r trie test *s run m tubes which have no gas from
on.
Courtesy of Analytab Products, Plainview, N.Y,
446
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
Back Matter
Appendix D: Identification
Charts
© The McGraw-H
Companies, 2001
Identification Charts
Appendix D
Chart II Symbol Interpretation of API 20E System
Tube
Chemical/Physical Principles
Components
Reactive Ingredients
Quantity
Ref.
ONPG
Hydrolysis of ONPG by bet^gatactasidase releases yellow
orthonitro phenol from the colorless ONPG; ITPG
(isopropylthiogalactopyranoside) is used as inducer
ONPG
ITPG
0.2 mg
8.0 }ig
12
13
14
ADH
Arginine dihydrolase transforms arginine into ornithine, ammonia,
and carbon dioxide. This causes a pH rise in the acid-buffered
system and a change in the indicator from yeJlow to red
Arginine
2,0 mg
15
LDC
Lysine decarboxylase transforms lysine into a basic primary : Lysine
amine, cadaverine. This amine causes a pH rise in the acid- !
buffered system and a change in the indicator from yellow to red. j
2.0 mg
15
ODC
Ornithine decarboxylase transforms ornithine into a basic
primary amine, putrescine This amine causes a pH rise in the
acid-buffered system and a change fn the indicator from yellow to
red.
Ornithine
2.0 mg
15
CIT
Citrate is the sole carbon source Citrate utilization results \n a
pH rise and a change in the indicator from green to blue.
Sodium Citrate
0.3 mg
21
H 2 S
Hydrogen sulfide is produced from thiosulfate. The hydrogen
sulfide reacts with iron salts to produce a black precipitate.
Sodium Thiosjlfate
30.0 ^g
6
LIRE
Urease releases ammonia from urea; ammonia causes the pH tc
rise and changes the indicator from yellow to red.
Urea
0.8 mg
7
TDA
Tryptophane deaminase forms indolepyruvic acid from
tryptophane. Indolepyruvic acid produces a brownish-red color in
the presence of ferric chloride.
Tryptophane
Q.4 mg
22
IND
Metabolism of tryptophane results in the formation of indole.
Kovacs 1 reagent forms a colored complex [pink to red) with
Indole.
Tryptophane
0.2 mg
10
k u ■ 1
VP
Acetoin, an intermediary glucose metabolite, is produced from
sodium pyruvate and indicated by the formation ol a colored
complex. Conventional VP tests may take up tc 4 days, but by
using sodium pyruvate, API has shortened the required test time.
Creatine intensifies the color when tests are positive.
Sodium Pyruvate
Creatine
2.0 mg
0.9 mg
3
GEL
Liquefaction of gelatin by proteolytic enzymes releases a black
pigment which diffuses throughout the tube.
Kohn Charcoal Gelatin 1 0,6 mg
1
9
GLU
MAN
INO
SOR
RHA
SAC
MEL
AMY
ARA
Utilization of the carbohydrate resutts in acid formation and a
consequent pH drop. The indicator changes from biue to yellow.
Glucose
MannttQl
Inositol
Sorbitol
Rhamnose
Sucrose
Meiibiose
Amygdalfn
(1 +) Arabinose
2.0 mg
2,0 mg
1 2.0 mg
2.0 mg
2.0 mg
2.0 mg
2.0 mg
2.0 mg
2.0 mg
5
6
12
| GLU
Nitrate
Reduction
Nitrites form a red complex with sulfanilic acid and N, N-
dimethylalpha-naphthylamine. In case of negative reaction,
addition of zinc confirms the presence of unreduced nitrates by
reducing them to nitrites (pink-orange color). If there is no color
change after the addition of zinc, this is indicative of the
complete reduction of nitrates through nrtrites to nitrogen gas or
to an anaerogenic amine,
Potassium Nitrate
60.0 ug
6
MAN
INO
SOR
Catalase
Catalase releases oxygen gas from hydrogen peroxide.
24
Courtesy of Analytab Products, Plainview, N.Y.
447
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
Back Matter
Appendix D: Identification
Charts
© The McGraw-H
Companies, 2001
Appendix D
Identification Charts
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448
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
Back Matter
Appendix D: Identification
Charts
© The McGraw-H
Companies, 2001
Identification Charts
Appendix D
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03
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(J)
CD
O
O
449
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
Back Matter
Appendix D: Identification
Charts
© The McGraw-H
Companies, 2001
Appendix D
Identification Charts
Chart IV Characterization of Enterobacteriaceae — The Enterotube II System
Groups
ESCHERlCMiEAE
EDWARDSIELLEAE
UJ
<
UJ
UJ
z
O
—I
<
<jO
UJ
<
LU
111
I-
O
X
ClTROBACTER
PROTEUS
MORGANELLA
\
PROVfDENOA
Escherichia
Shigella
Ecfwardstefte
Safrrtonetta
Arizona
freundft
am&fon&tt'Girs
diversus
vulgaris
mirabilis
morg&nii
aicatitac/ens
stoartn
rsttgeri
cloacae
\ $$kazakir
LU
<
UJ
_l
UJ
53
CD
UJ
UJ
<
2
ce
UJ
>
ENTEROBACTER
\gsrgQvi3&
HA FM A
SERRAVA
KLEBSIELLA
YERSINIA
aerogenes
&ggtom$rans
alv&i
marcescerts
tiquefa&ens
rubidaez
pneun?oof&$
oxyioca
ozaenaQ
+ J
100.0 92,0
100
10Q.D
'- A
2.1
■I
99.4
100,0 !
1000
+ c
91.9
+
100.0
+■
100.0
rhfnoschferoma trs
&nt&rocoi?tica
pseutfo tuberculosis
I
100.0
+■
100,0
100.0
+■
100,0
^
10Q.0
100.0
+
10D.O
1
100.0
4-
100.0
+
100.0
\
100.0
+
100.0
•i
100.0
+
100.0
+
100.0
+
100
+
100,0
ido.g
10O.O
+
997
4-
91.4
97.0
+
97.3
±G
86.0
100.0
+ H
946
934
0,0
+ B
200
99.0
+ 1
92.7
1D0.0
d
17.2
00
0,0
00
\ G
96.0
66.0
85,2
0.0
-G
12.2
99.3
97.0
93.0
t
95.9
24.1
&S.9
+ G
52.6
d
72.5
d G
350
960
96.0
d
55.0
■t
1D0.D i 0.0
4-
100.0
+™.
100.0
0.0
0,0
0.0
0.0
0.0
0.0
t-
97.0
00
4
99.6
4E
91.5
9B.7
81.6
0.0
i ^ vhii pr t-i
4
996
0.0
4
99.0
4
97.0
1.2
0.0
0.0
0.0
0.0
±
64.0
+
97,5
0.0
+
99.6
4
996
64.2
61.0
4
97.2
4-
97.2
35. a
0.0
0.0
0.0
+
93.7
+
97,0
4
100.0
+
95,9
0.0
+
98.6
4
99.6
+
100
0.0
-.-..
0.0
0.0
1.0
0.0
907
0.0
00
0.0
95,0
94.5
0.0
0.0
00
0.0
0.0
0.0
0.0
0.0
00
0.0
o.o
o.o
0.0
00
+
37.B
+
990
11
2.0
6.7
4
99.0
4
100.0
4
91.4
0.0
00
O.D
0.0
00
o.o
+
100.0
0.0
32
99.5
+
994
•I
9B.6
4
959
00
160
0.0
08
19 7
0.0
w
D.1
— w
1.8
-w
2.0
00
0.0
00
0.0
DO
00
+
1000
0.0
0.0
26.7
0.0
O.O
0.0
+
94.3
12,4
4
990
26,0
0,0
0,0
4
97 5
7.5
56,0
83
88.0
B9.D
890
91.B
4
98-0
0.0
00
B
0.3
0.0
0.B
d
69.8
d
39.3
70.0
d
40-3
0.0
2.0
0.0
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3.6
d
10
t-
94.0
+
100
42,0
+
925
d
52.9
d
28
67.6
10.7
+
39.2
+
99.1
I
100.0
4
99.0
I
98.0
0.0
00
0.0
07
4.0
00
4
99.4
4-
100.0
100.0
100
97.5
993
29.1
0.2
4
94.1
+
97.1
+
96.2
+
97.0
I
98.2
0,0
0.0
00
D.6
3-4
1.0
+
1D0.0
0.0
0.0
0.0
o.o
0.0
0-0
0.0
0.0
d
5.4
0.0
dD
B65
0,0
d
59.8
110
0.0
00
15.0
00
0.0
00
DO
100.0
97.0
100.0
98.3
d
26.3
O.O
21
00
+
99.1
d
15.6
100.0
98.7
98.7
d
26.2
d
6.0
0.0
0.0
4
97.3
4
100.0
I
99.9
4
100. D
+
100.0
4
1000
+
98.7
±
55.0
4
97.3
80
4
99,4
4
98.D
78,0
I
98.D
+
98.7
0.0
100.0
64.6
+
65.0
4-
98.7
49.5
4
92.0
4
93.7
52.2
0.0
0.0
0.0
0.0
0.0
d
15.2
6.0
0,0
4.1
0.0
0.0
0.0
0.0
0.0
0.0
0.0
100.0
4-
99.6
+
950
4-
97,4
94.5
98.0
0.0
00
0.0
0,0
0,0
dw
89.4
81.0
dw
85.8
95.0
69.3
97 1
0.0
+
20.0
4-
100.0
74.6
00
0.0
d
12,9
2.4
00
0.0
0.0
330
I
93.7
0.0
0.0
0.1
0,0
33.0
0.0
0.0
0.0
0.0
4
27.6
O.O
0.D
0.9
0.0
0.0
0.0
00
0.0
O.O
00
0.0
4
100.0
00
d
34.1
3.0
d w
39.7
d w
3.7
d w
4.0
4
95,4
954
d
148
0.0
f
90.7
100.0
Courtesy of Roche Diagnostics, Nutley, N.J
E. S enter itid/s bioserotype Paratyphi A and some rare biotypes may be Ha S negative.
F. S. typhi. S. enteritidis bio serotype Paratyphi A and some rare biotypes are citrate- negative and S. cholerae-suis is usually delayed positive.
G. The amount of gas produced by Serratia, Proteus and Providencia aicaMaCiens is slight; therefore, gas production may not be evident in the ENTEROTUBE II.
H S, Qnteritidis bio serotype Paratyphi A Is negative lor lysine decarboxylase.
I. S. typhi and $ gaUinarum are ornithine decarboxylese-negaiive.
J. The Alkalescens-Oispar (A-D) group i3 included as a biotype of E, pott. Members of the AD group are generally anaerogenic, non-motile and do not ferment lactose.
K. An occasional strain may produce hydrogen sulfide,
L. An occasional strain may appear to utilize citrate.
0.0
0.0
dF
80.1
4
96.6
4
90,4
4
94.0
+
99.7
d
10.5
58.7
L
0.0
4
97 9
t
93.7
+
960
4
96.9
4
94.0
4
96.0
92.6
d
84.2
d
5.6
97.6
936
68
4
96.6
96.6
d
28.1
0.0
0.0
0.0
450
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
Back Matter
Appendix D: Identification
Charts
© The McGraw-H
Companies, 2001
Identification Charts
Appendix D
Chart V Reaction Interpretations for API Staph-ldent
MICROCUPULE
INTERPRETATION OF REACTIONS
NO
1
SUBSTRATE
POSITIVE
NEGATIVE
PHS
URE
GLS
4
MNE
5
MAN
6
TRE
7
SAL
a
10
GLC
Yellow
Clear or straw
colored
w^^^mam
COMMENTS AND REFERENCES
A positive result should be recorded only if significant color
development has occurred. (3)
Purple to Red-
Orange
Yellow or Yellow- j Phenol red has been added to the urea formulation to allow
Orange detection of alkaline end products resulting from urea utilization. (1)
^ i
Yellow
Clear or straw-
colored
Yellow or
Yellow-Orange
Red or Orange
Yellow
Clear or straw
colored
ARG
\ Purple to Red-
Orange
NGP
Yellow or Yellow
Orange
A positive result should be recorded only if significant color
development has occurred.
Cresol red has been added to each carbohydrate to allow
detection of acid production if the respective carbohydrates are
utilized. (1,7)
A positive result should be recorded only if significant color
development has occurred.
Phenol red has been added to the arginine formulation to allow
detection of alkaline end products resulting from arginine
utilization. (1)
Add 1-2 drops of STAPH-IDENT
REAGENT
Plum-Purple
(Mauve)
Yellow or
colorless
Color development will begin within 30 seconds of reagent
addition. (1,5)
Courtesy of Analytab Products, Plainview, N.Y.
Abbreviation
PHS
URE
GLS
MNE
MAN
TRE
SAL
GLC
ARG
NGP
Test
Phosphatase
Urea utilization
p-Glucosidase
Mannose utilization
Mannitol utilization
Trehalose utilization
Salicin utilization
(^-Glucuronidase
Arginine utilization
p-Galactosidase
451
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
Back Matter
Appendix D: Identification
Charts
© The McGraw-H
Companies, 2001
Appendix D
Identification Charts
Chart VI Biochemistry of API Staph-ldent Tests
M1CROCUPULE
NO
SUBSTRATE
CHEMICAL/PHYSICAL PRINCIPLES
REACTIVE INGREDIENTS
QUANTITY
1
2
PHS
URE
Hydrolysis of p-nitrophenyl-phosphate, disodium salt, by
alkaline phosphatase releases yellow paranitrophenol
from the colorless substrate.
p-nitrophenyl-phosphate,
disodium salt
0.2%
Urease releases ammonia from urea; ammonia causes
the pH to rise and changes the indicator from yellow to
red.
Urea
1.6%
GLS
Hydrolysis of p-nitrophenyl-^-D-glucopyranoside by 8-
glucosidase releases yellow para-nitrophenol from the
colorless substrate.
p-nitrophenyl-;?-D-
glucopyranoside
0.2%
4
MIME
5
MAN
6
TRE
7
SAL
8
GLC
ARG
Utilization of carbohydrate results in acid formation and a
consequent pH drop. The indicator changes from red to
yellow.
Mannose
Mannitol
Trehalose
Salicin
1.0%
1.0%
1.0%
1.0%
^^AUI
Hydrolysis of p-nitrophenyl-tf-D-glucuronide by /3-
glucuronidase releases yellow para-nitrophenol from the
colorless substrate.
p-nitrophenyl-,tf-D-glucuronide
0.2%
Utilization of arginine produces alkaline end products
which change the indicator from yellow to red.
Arginine
1 .6%
10
NGP
Hydrolysis of 2-naphthol-^-D-galactopyranoside by 8-
galactosidase releases free ,#-naphthol which complexes
with STAPH-IDENT REAGENT to produce a plum-purple
(mauve) color.
Courtesy of Analytab Products, Plainview, N.Y.
2-naphthol-£-D-
galactopyranoside
0.3%
452
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
Back Matter
Appendix D: Identification
Charts
© The McGraw-H
Companies, 2001
Identification Charts
Appendix D
Chart VII ,
API Staph-ldent Profile Register*
r
Profile
Identification
Profile
Identification
040
STAPH CAPITIS
4 700
STAPH AUREUS
COAG +
060
STAPH HAEMOLYTICUS
STAPH SCIURI
COAG -
100
STAPH CAPITIS
4 710
STAPH SCIURI
D 140
i
STAPH CAPITIS
5 040
STAPH EPIDERMIDIS
200
STAPH COHNII
5 200
STAPH SCIURI
240
STAPH CAPITIS
5 210
STAPH SCIURI
300
STAPH CAPITIS
5 300
STAPH AUREUS
COAG +
340
STAPH CAPITIS
STAPH SCIURI
COAG -
440
STAPH HAEMOLYTICUS
5 310
STAPH SCIURI
460
STAPH HAEMOLYTICUS
5 600
STAPH SCIURI
600
STAPH COHNII
5610
STAPH SCIURI
620
STAPH HAEMOLYTICUS
5 700
STAPH AUREUS
COAG +
640
STAPH HAEMOLYTICUS j
STAPH SCIURI
COAG -
660
STAPH HAEMOLYTICUS
5710
STAPH SCIURI
1 000
STAPH EPIDERMIDIS
5 740
STAPH AUREUS
1 040
STAPH EPIDERMIDIS
6 001
STAPH XYLOSUS
XYL + ARA +
1 300
STAPH AUREUS
STAPH SAPROPHYTICUS
XYL - ARA -
1 540
STAPH HYICUS (An)
6011
STAPH XYLOSUS
1 560
STAPH HYICUS (An)
i 5 021
STAPH XYLOSUS
2 000
STAPH SAPROPHYTICUS NOVO R
6 101
STAPH XYLOSUS
STAPH HOMINIS NOVO S
6 121
STAPH XYLOSUS
2 001
STAPH SAPROPHYTICUS
6 221
STAPH XYLOSUS
2 040
STAPH SAPROPHYTICUS NOVO R
6 300
STAPH AUREUS
STAPH HOMINIS NOVO S
6 301
STAPH XYLOSUS
2 041
STAPH SIMULANS
6311
STAPH XYLOSUS
2 061
STAPH SIMULANS
6 321
STAPH XYLOSUS
2 141
STAPH SIMULANS
6 340
STAPH AUREUS
COAG +
2 161
STAPH SIMULANS
STAPH WARNERI
COAG -
2 201
STAPH SAPROPHYTICUS
6 400
STAPH WARNERI
2 241
STAPH SIMULANS
6 401
STAPH XYLOSUS
XYL + ARA ■+■
2 261
STAPH SIMULANS
STAPH SAPROPHYTICUS
XYL - ARA -
2 341
STAPH SIMULANS
6 421
STAPH XYLOSUS
2 361
STAPH SIMULANS
6 460
STAPH WARNERI
2 400
STAPH HOMINIS NOVO S
6 501
STAPH XYLOSUS
STAPH SAPROPHYTICUS NOVO R
i 6 521
STAPH XYLOSUS
2 401
STAPH SAPROPHYTICUS
6 600
STAPH WARNERI
2 421
STAPH SIMULANS
6 601
STAPH SAPROPHYTICUS
XYL - ARA -
2 441
STAPH SIMULANS
STAPH XYLOSUS
XYL + ARA +
2 461
STAPH SIMULANS
6 611
STAPH XYLOSUS
2 541
STAPH SIMULANS
6 621
STAPH XYLOSUS
2 561
STAPH SIMULANS
6 700
STAPH AUREUS
2 601
STAPH SAPROPHYTICUS
6 701
STAPH XYLOSUS
2 611
STAPH SAPROPHYTICUS
6 721
STAPH XYLOSUS
2 661
STAPH SIMULANS
6 731
STAPH XYLOSUS
2 721
STAPH COHNII (SSP1)
7 000
STAPH EPIDERMIDIS
2 741
STAPH SIMULANS
7 021
STAPH XYLOSUS
2 761
STAPH SIMULANS
7 040
STAPH EPIDERMIDIS
3 000
STAPH EPIDERMIDIS
7 141
STAPH INTERMEDIUS (An)
3 040
, STAPH EPIDERMIDIS
7 300
STAPH AUREUS
3 140
i STAPH EPIDERMIDIS
7 321
STAPH XYLOSUS
3 540
STAPH HYICUS (An)
7 340
STAPH AUREUS
3 541
STAPH INTERMEDIUS (An)
7 401
STAPH XYLOSUS
3 560
STAPH HYICUS (An)
7 421
STAPH XYLOSUS
3 601
STAPH SIMULANS NOVO S
7 501
STAPH INTERMEDIUS (An)
COAG +
STAPH SAPROPHYTICUS NOVO R
STAPH XYLOSUS
COAG -
4 060
STAPH HAEMOLYTICUS
7 521
STAPH XYLOSUS
4 210
STAPH SCIURI
7 541
STAPH INTERMEDIUS (An)
4 310
STAPH SCIURI
7 560
STAPH HYICUS (An)
4 420
STAPH HAEMOLYTICUS
7 601
STAPH XYLOSUS
i
4 440
STAPH HAEMOLYTICUS
7 621
STAPH XYLOSUS
4 460
STAPH HAEMOLYTICUS
7 631
STAPH XYLOSUS
4 610
STAPH SCIURI
7 700
STAPH AUREUS
4 620
i
STAPH HAEMOLYTICUS \
7 701
STAPH XYLOSUS
4 660
STAPH HAEMOLYTICUS
7 721
STAPH XYLOSUS
7 740
STAPH AUREUS
*Date of Publication: March, 1984
Courtesy of Analytab Products, Plainview, N.Y.
453
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
Back Matter
Appendix E:The
Streptococci
© The McGraw-H
Companies, 2001
E
Appendix
The Streptococci: Classification, Habitat,
Pathology, and Biochemical Characteristics
To fully understand the characteristics of the various
species of medically important streptococci, this appen-
dix has been included as an adjunct to Exercise 79. The
table of streptococcal characteristics on this page is the
same one that is shown on page 267 of Exercise 79. It is
also the basis for much of the discussion that follows.
The first system that was used for grouping the
streptococci was based on the type of hemolysis and
was proposed by J. H. Brown in 1919. In 1933, R. C.
Lancefield proposed that these bacteria be separated
into groups A, B, C, etc., on the basis of precipitation-
type serological testing. Both hemolysis and serolog-
ical typing still play predominant roles today in our
classification system. Note below that the Lancefield
groups are categorized with respect to the type of he-
molysis that is produced on blood agar.
Beta Hemolytic Groups
Using a streak-stab technique, a blood agar plate is in-
cubated aerobically at 37° C for 24 hours. Isolates that
have colonies surrounded by clear zones completely free of
red blood cells are characterized as being beta hemolytic.
Three serological groups of streptococci fall in this cate-
gory: groups A, B, and C; a few species in group D are also
beta hemolytic.
Table 1 Physiological Tests for Streptococcal Differentiation
GROUP / * / «Sr /<? / ** / **& / K *>' / <# / *$ /
Group A
S. pyogenes
beta
+
■ i ■■
R
—
^—
I -
Group B
S. agafactiae
beta
*
+
R
—
±
"
Group C
S. equi
S. equisimifis
S. zooepidemicus
beta
_ *
S
i
!
Group D**
(enterococci)
S. faecatis
S. faecium
etc.
alpha
beta
none
R
4-
i
+
Group D**
(nonenterococci)
S. bo vis
etc.
alpha
none
R/S
+ ;
Viridans
S. mitis
S. salivarius
S. mutans
etc.
alpha
none
+
*
s
Pneumococci
S. pneumoniae
alpha
±
^—
^—
^—
+
+
*Exceptions occur occasionally **See comments on pp. 457 and 458 concerning correct genus
Note: R = resistant; S = sensitive; blank = not significant
455
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
Back Matter
Appendix E:The
Streptococci
© The McGraw-H
Companies, 2001
Appendix E
The Streptococci: Classification, Habitat, Pathology, and Biochemical Characteristics
Group A Streptococci
This group is represented by only one species:
Streptococcus pyogenes. Approximately 25% of all
upper respiratory infections (URIs) are caused by this
species; another 10% of URIs are caused by other
streptococci; most of the remainder (65%) are caused
by viruses. Since no unique clinical symptoms can be
used to differentiate viral from streptococcal URIs,
and since successful treatment relies on proper identi-
fication, it becomes mandatory that throat cultures be
taken in an attempt to prove the presence or absence
of streptococci. It should be added that if streptococ-
cal URIs are improperly treated, serious sequelae
such as pneumonia, acute endocarditis, rheumatic
fever, and glomerularnephritis can result.
S. pyogenes is the only beta hemolytic strepto-
coccus that is primarily of human origin. Although the
pharynx is the most likely place to find this species, it
may be isolated from the skin and rectum.
Asymptomatic pharyngeal and anal carriers are not
uncommon. Outbreaks of postoperative streptococcal
infections have been traced to both pharyngeal and
anal carriers among hospital personnel.
These coccoidal bacteria (0.6-1.0 (xm diameter)
occur as pairs and as short to moderate-length chains
in clinical specimens; in broth cultures, the chains are
often longer.
When grown on blood agar, the colonies are small
(0.5 mm dia.), transparent to opaque, and domed; they
have a smooth or semimatte surface and an entire
edge; complete hemolysis (beta- type) occurs around
each colony, usually two to four times the diameter of
the colony.
S. pyogenes produces two hemolysins: strep-
tolysin S and streptolysin O. The beta- type hemolysis
on blood agar is due to the complete destruction of red
blood cells by the streptolysin S .
There is no group of physiological tests that can
be used with absolute certainty to differentiate S. pyo-
genes from other streptococci; however, if an isolate
is beta hemolytic and sensitive to bacitracin, one can
be 95% certain that the isolate is S. pyogenes. The
characteristics of this organism are the first ones tab-
ulated in table I on the previous page.
Group B Streptococci
The only recognized species of this group is 5.
agalactiae. Although this organism is frequently
found in milk and associated with mastitis in cattle,
the list of human infections caused by it is as long as
the one for S. pyogenes: abscesses, acute endocarditis,
impetigo, meningitis, neonatal sepsis, and pneumonia
are just a few. Like S. pyogenes, this pathogen may
also be found in the pharynx, skin, and rectum; how-
ever, it is more likely to be found in the genital and in-
testinal tracts of healthy adults and infants. It is not
unusual to find the organism in vaginal cultures of
third-trimester pregnant women.
Cells are spherical to ovoid (0.6-1.2 |xm dia) and
occur in chains of seldom less than four cells; long
chains are frequently present. Characteristically, the
chains appear to be composed of paired cocci.
Colonies of S. agalactiae on blood agar often pro-
duce double zone hemolysis. After 24 hours incuba-
tion colonies exhibit zones of beta hemolysis. After
cooling, a second ring of hemolysis forms which is
separated from the first by a ring of red blood cells.
Reference to table I emphasizes the significant
characteristics of S. agalactiae. Note that this organ-
ism gives a positive CAMP reaction, hydrolyzes hip-
purate, and is not (usually) sensitive to bacitracin. It is
also resistant to SXT. Presumptive identification of
this species relies heavily on a positive CAMP test or
hippurate hydrolysis, even if beta hemolysis is not
clearly demonstrated.
Group C Streptococci
Three species fall in this group: S. equisimilis, S.
equi, and S. zooepidemicus. Although all of these
species may cause human infections, the diseases are
not usually as grave as those caused by groups A and
B. Some group C species have been isolated from im-
petiginous lesions, abscesses, sputum, and the phar-
ynx. There is no evidence that they are associated
with acute glomerularnephritis, rheumatic fever, or
even pharyngitis.
Presumptive differentiation of this group from S.
pyogenes and S. agalactiae is based primarily on (1)
resistance to bacitracin, (2) inability to hydrolyze hip-
purate or bile esculin, and (3) a negative CAMP test.
There are other groups that have some of these same
characteristics, but they will not be studied here.
Tables 12.16 and 12.17 on page 1049 of Bergey's
Manual, vol. 2, provide information about these other
groups.
Alpha Hemolytic Groups
Streptococcal isolates that have colonies with zones of
incomplete lysis around them are said to be alpha he-
molytic. These zones are often greenish; sometimes
they are confused with beta hemolysis. The only way
to be certain that such zones are not beta hemolytic is
to examine the zones under 60X microscopic magnifi-
cation. Figure 79.4, page 265, illustrates the differ-
ences between alpha and beta hemolysis. If some red
blood cells are seen in the zone, the isolate is classified
as being alpha hemolytic.
456
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
Back Matter
Appendix E:The
Streptococci
© The McGraw-H
Companies, 2001
The Streptococci: Classification, Habitat, Pathology, and Biochemical Characteristics
Appendix E
The grouping of streptococci on the basis of alpha
hemolysis is not as clear-cut as it is for beta hemolytic
groups. Note in table I that the bottom four groups that
have alpha hemolytic types may also have beta he-
molytic or nonhemolytic strains. Thus, we see that he-
molysis in these four groups can be a misleading char-
acteristic in identification.
Alpha hemolytic isolates from the pharynx are
usually S. pneumoniae, viridans streptococci, or
group D. Our primary concern here in this experiment
is to identify isolates of S. pneumoniae. To accom-
plish this goal, it will be necessary to differentiate any
alpha hemolytic isolate from group D and viridans
streptococci.
Streptococcus pneumoniae
(Pneumococcus)
This organism is the most frequent cause of bacterial
pneumonia, a disease that has a high mortality rate
among the aged and debilitated. It is also frequently
implicated in conjunctivitis, otitis media, pericarditis,
subacute endocarditis, meningitis, septicemia,
empyema, and peritonitis. Thirty to 70% of normal in-
dividuals carry this organism in the pharynx.
Spherical or ovoid, these cells (0.5-1.25 |xm dia)
occur typically as pairs, sometimes singly, often in
short chains. Distal ends of the cells are pointed or
lancet-shaped and are heavily encapsulated with poly-
saccharide on primary isolation.
Colonies on blood agar are small, mucoidal,
opalescent, and flattened with entire edges sur-
rounded by a zone of greenish discoloration (alpha
hemolysis). In contrast, the viridans streptococcal
colonies are smaller, gray to whitish gray, and opaque
with entire edges.
Presumptive identification of S. pneumoniae can
be made with the optochin and bile solubility tests. On
the optochin test, the pneumococci exhibit sensitivity
to ethylhydrocupreine (optochin). With the bile solu-
bility test, pneumococci are dissolved in bile (2%
sodium desoxycholate). Table I reveals that except for
bacitracin susceptibility (±), S. pneumoniae is nega-
tive on all other tests used for differentiation of strep-
tococci.
Viridans Group
Streptococci that fall in this group are primarily alpha
hemolytic; some are nonhemolytic. Approximately 10
species are included in this group. All of them are
highly adapted parasites of the upper respiratory tract.
Although usually regarded as having low pathogenic-
ity, they are opportunistic and sometimes cause seri-
ous infections. Two species (S. mutans and S. sanguis)
are thought to be the primary cause of dental caries,
since they have the ability to form dental plaque.
Viridans streptococci are implicated more often than
any other bacteria in subacute bacterial endocarditis.
When it comes to differentiation of bacteria of this
group from the pneumococci and enterococci, we will
use the optochin, bile solubility, and salt-tolerance
tests. See table I.
Group D Streptococci (Enterococci)
Members of this group are, currently, considered by
most taxonomists to belong to the genus
Enterococcus. During the preparation of volume 2 of
Bergey's Manual Schleifer and Kilper-Balz presented
conclusive evidence that S.faecalis, S. faecium, and S.
bovis were so distantly related to the other groups of
streptococci that they should be transferred to another
genus. Since the term Enterococcus had been previ-
ously suggested by others, Schleifer and Kilper-Balz
recommended that this be the name of a new genus to
include all of the Group D streptococci, nonentero-
cocci included. The fact that these papers came too
late for Bergey's Manual to include this new genus
caused the genus Streptococcus to be retained. To
avoid confusion in our use of Bergey's Manual, we
have retained the same terminology used in Bergey 's
Manual.
The enterococci of serological group D may be al-
pha hemolytic, beta hemolytic, or nonhemolytic. The
principal species of this enterococcal group are S. fae-
calis, S. faecium, S. durans, and S. avium.
Subacute endocarditis, pyelonephritis, urinary
tract infections, meningitis, and biliary infections are
caused by these organisms. All five of these species
have been isolated from the intestinal tract.
Approximately 20% of subacute bacterial endocardi-
tis and 10% of urinary tract infections are caused by
members of this group. Differentiation of this group
from other streptococci in systemic infections is
mandatory because S. faecalis, S. faecium, and S. du-
rans are resistant to penicillin and require combined
antibiotic therapy.
Since S. faecalis can be isolated from many food
products (not connected with fecal contamination), it
can be a transient in the pharynx and show up as an
isolate in throat cultures. Morphologically, the cells
are ovoid (0.5-1.0 |xm dia) occurring as pairs in short
chains. Hemolytic reactions of S. faecalis on blood
agar will vary with the type of blood used in the
medium. Some strains produce beta hemolysis on
agar with horse, human, and rabbit blood; on sheep
blood agar the colonies will always exhibit alpha he-
molysis. Other streptococci are consistently either
beta, alpha, or nonhemolytic.
Cells of S. faecium are morphologically similar to
S. faecalis except that motile strains are often encoun-
457
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
Back Matter
Appendix E:The
Streptococci
© The McGraw-H
Companies, 2001
Appendix E
The Streptococci: Classification, Habitat, Pathology, and Biochemical Characteristics
tered. A strong alpha-type hemolysis is usually seen
around colonies of S. faecium on blood agar.
Although presumptive differentiation of group D
enterococcal streptococci from groups A, B, and C is
not too difficult with physiological tests, it is more la-
borious to differentiate the individual species within
group D. As indicated in table I, the enterococci (1)
hydrolyze bile esculin, (2) are CAMP negative, and
(3) grow well in 6.5% NaCl broth.
Differentiation of the five species within this
group involves nine or ten physiological tests.
Group D Streptococci (Nonenterococci)
The only medically significant nonenterococcal
species of group D is S. bovis. This organism is found
in the intestinal tract of humans as well as in cows,
sheep, and other ruminants. It can cause meningitis,
subacute endocarditis, and urinary tract infections. On
blood agar, the organism is usually alpha hemolytic;
occasionally, it is nonhemolytic. The best way to dif-
ferentiate it from the group D enterococci is to test its
tolerance to 6.5% NaCl. Note in table I that S. bovis
will not grow in this medium, but all enterococci will.
458
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
Back Matter
Appendix F: Identibacter
interactus
© The McGraw-H
Companies, 2001
Appendix
Identibacter Interactus
As stated in Exercise 51, Identibacter interactus is a
computer program designed to assist students in iden-
tifying unknown bacterial cultures. This CD-ROM
program, which is distributed by WCB/McGraw-Hill
Co. in Dubuque, is a powerful program that includes
more than 50 tests to run on assigned bacterial un-
knowns. The organism data base includes about 60
species of chemoheterotrophic bacteria.
To run this program, you will select each test from
pull-down menus. A color image of each test result
will be displayed on the computer screen, and you
must be able to correctly interpret the test result that
is shown. Once you have tabulated sufficient infor-
mation, you can identify your unknown by typing in
the name of the organism. An audit trail of your
choices can be saved to disk which can be evaluated
by your instructor.
Before you attempt to use this program, read over
the following pages of this Appendix. These twelve
pages are the first portion of a 59 page instructional
manual that can be accessed from the CD-ROM. This
information will explain more in detail how the pro-
gram functions. A full copy of the manual should be
available to you in the laboratory.
459
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
Back Matter
Index
©TheMcGraw-H
Companies, 2001
Acetobacter, 241
acid-fast staining, 69
Actinomycetes, isolation of, 203-5
agglutination tests
Epstein-Barr virus, 283
heterophile antibody, 283
S. aureus, 281
Widal, 285
Alcaligenes, characteristics of, 1 80, 270
alcohol fermentation, 241
Algae, Subkingdom, 26
alpha hemolysis, 264
alpha toxin, 258
Amastigomycota, 49
ammonification, 212
amoeboid movement, 28
Anabaena, 34
anaerobe culture, 89
anaerobic phototrophic bacteria, 106
Annelida, 36
antagonism, microbial, 128
antibiotic production, in soil, 203
antibiotic testing, 145
antigens, heterophile, 283
antiseptics, evaluation of, 143
Apicomplexa, 28
API 20E system, 185
API Staph-Ident system, 198
Archaea, Domain, 25
Arthrobacter, characteristics of, 179
arthrospores, 49
Ascomycetes, 50
ascospores, 49
Aschelminthes, 35
aseptic technique, 39-45
Aspergillus, 52, 53
atomic weights, 425
autoclave steam pressure table, 429
autotrophs, 76
Azotobacter, 210
Bacillus, characteristics, 178
bacitracin susceptibility, 267
bacteria, definition, 46
Bacteria, Domain, 25
bacteriochlorophyll, 30, 46
bacteriophage, 111-24
Barritt's reagents, usage, 167
Basidiomycotina, 50
basidiospores, 4
basophils, 288, 289
B er gey ' s Manual, usage of, 177-81
beta hemolysis, 264
bile esculin hydrolysis, 268
bile solubility test, 269
blastoconidia, 49
blastospore, 49
blood agar usage, 260
blood cells
differential WBC count, 288
total WBC count, 292
typing, 295
blood types, 296
Brady rhizobium, 207, 211
Breed count, 231
burst size, phage, 120
butanediol fermentation, 1 67
CAMP test, 262, 266
capsid, 112
capsular staining, 63
cardioid condenser, 10
caries susceptibility test, 299
carotene, 30
casein hydrolysis test, 172
catalase test, 168
Ceratium, 30, 31
chemoautotrophs, 77
Chlamydomonas, 28, 29
chlamydospores, 49
Chlorobiaceae, 106
Chlorobium, 107
Chlorogonium, 29
chlorophyll, 28
chloroplasts, 28
Chromatiaceae, 106
Chromatium, 107
Chrysophycophyta, 30
Ciliophora, 28
citrate utilization test, 175
Citrobacter, 270
cladocera, 36, 37
Cladosporium, 52, 53
Clostridium, characteristics, 178
coagulase test, 258, 260
coelenterates, 35
commensalism, microbial, 126
conidia, 49
copepods, 36, 37
Corynebacterium, characteristics of, 179
cultural characteristics, bacteria, 157-60
Cyanobacteria, 30, 34, 207
Deinococcaceae, 257
denitrification, 207, 213, 217
Deuteromycotina, 50
diatomite, 30
diatoms, 30, 33
differential WBC count, 288
disinfectants
alcohol effectiveness, 141
evaluation, 139
DNase test, 261
475
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
Back Matter
Index
©TheMcGraw-H
Companies, 2001
Index
domains, classification, 25
Dorner method staining, 68
Durham tube usage, 164
endoenzymes, 161
endospores, 67, 156
Enterobacter, 270
Enterobacteriaceae identification, 185, 189
eosinophilia, 289
eosinophils, 288, 289
epidemic, 254
erythrocytes, 288
Escherichia, 270
Eudorina, 28, 29
Euglena, 29
euglenoids, 28
Euglenophycophyta, 28
Eukarya, Domain, 25
exoenzymes, 161
fat hydrolysis test, 172
fermentation, 161
flagellum, 28
flatworms, 35
Flavobacterium, characteristics of, 180
fluorescence method staining, 70
food spoilage, 237
formic hydrogenylase, 164
fungi, 48-53
fungi imperfecta 50
GasPak anaerobic jar, usage of, 90
gastrotrichs, 35
Gonium, 28, 29
gram staining, 64
green sulfur bacteria, 106
Gymnodinium, 30
Halobacterium, characteristics of, 180
halophile, 135
hand scrubbing evaluation, 148
hemacytometer usage, 293
hemolysis types, streptococci, 264
Henrici slide culture technique, 99
heterocysts, 207
heterotrophs, 76
hippurate hydrolysis, 268
Hirodinea, 36
hydrogen ion needs, bacterial, 77
hydrogen sulfide test, 174
hydrolysis tests for bacteria, 170
hyphae, 48
Identibacter Inter actus, 1 8 1 , 45 9-72
IMViC tests, 175
indole test, 190, 196
Kirby-Bauer table, 432
Kirby-Bauer test for antibiotics, 145
Klebsiella, 270
Kovacs' reagent, usage, 173, 273
Kurthia, characteristics of, 179
Lactobacillus brevis, 243
characteristics of, 178
leucosin, 30
leukocytosis, 289
leukopenia, 289
Listeria, characteristics of, 178
litmus milk reactions, 176
logarithm tables, 426
lymphocytes, 288, 289
lysis, phage, 112
lysogeny, 112
Mastigophora, 28
media preparation, 76-8 1
media usage
blood agar, 263, 276
Brewer's anaerobic agar, 89
desoxycholate lactose agar, 276
fluid thyogly collate medium, 89, 159
glucose broth, 162
Hektoen enteric agar, 272
Kligler's iron agar, 174
litmus milk, 176
MacConkey agar, 272
MR- VP medium, 162
nitrate broth, 162
nutrient agar, 152
nutrient broth, 158
nutrient gelatin, 151
Russell double sugar agar, 273
semisolid medium, 155
SIM medium, 273
Simmons citrate agar, 174
skim milk agar, 170
Snyder test agar, 299
spirit blue agar, 170
starch agar, 170
trypticase soy agar, 1 62
xylose lysine desoxycholate agar, 272
mesophiles, 130
metabolism, 161
metachromatic granules, 62
methyl red test, 1 64
microaerophiles, 89
Micrococcus, characteristics of, 179
microphages, 288
microscopy
brightfield, 2-8
darkfield, 9-11
fluorescence, 17-21
measurements, 22-24
phase contrast, 11-16
mixed acid fermentation, 164
molds, 48, 5 1
monocytes, 288, 289
monocytosis, 289
M organella, 210
MPN calculation, 225
MPN table, 43 1
Mucor, 50, 52, 53
mycelium, 48
Myceteae, Kingdom, 48
mycology, 48
negative staining, 56
Neisseria, characteristics of, 181
476
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
Back Matter
Index
©TheMcGraw-H
Companies, 2001
Index
Nematoda, 35
neutropenia, 289
neutrophilia, 289
neutrophils, 288
nitrate reduction test, 168
Nitrobacter, 206
Nitrococcus, 206
nitrogen cycle, 206
nitrogen fixation, 208
Nitrosococcus, 206
Nitrosomonas, 206
pseudohypha, 48
Pseudomonas, characteristics of, 1 80, 270
pseudopod, 28
psychrophiles, 130
purple sulfur bacteria, 106
Pyrrophycophyta, 30
oil immersion techniques, 7
oligodynamic action, 136
Oospora, 52, 53
optochin susceptibility test, 269
Oscillatoria, 34
osmophile, 135
osmotic pressure and growth, 135
ostracods, 36, 37
oxidase test, 168
Oxi/Ferm tube II system, 1 94
palisade arrangement, 62
pandemic, 254
Pandorina, 28, 29
Paracoccus denitrificans, 218
paramylum, 28
parfocal lenses, 7
Penicillium, 50, 51
Peridinium, 30, 31
perithecia, 50
P h
adjustment methods, 79
effect on bacterial growth, 134
indicators, table of, 433
Phaeophycophyta, 30
phage typing, 287
phagocytic theory of immunity, 288
phenylalanine deamination test, 175
phialospores, 49
photoautotrophs, 77
phycobilisomes, 30
phycocyanin, 30
phycoerythrin, 30
pipette handling technique, 93
Planococcus, characteristics of, 180
Plantae, 28
Plasmodium, 28
Platyhelminthes, 35
pleomorphism, 62
polychaetes, 36
population counts, bacterial
food, 236
meat, 239
milk, 230
soil, 202
population count technique, 93
pour plate techniques, 86
prokaryotes, 25, 30
Proprionibacterium, characteristics of, 179
Proteus, 270
Protista, Kingdom, 26
Protozoa, Subkingdom, 26
Providencia, 270
reductase test, 234
resolution, microscope, 4
Rh blood typing, 298
Rhizobium, 207 ', 211
Rhizopus, 50, 53
rotifers, 35, 37
roundworms, 35
Saccharomyces cerevisiae, 241
Saccharomyces delbrueckii, 243
Salmonella, 270
salt tolerance, streptococci, 268
Sarcodina, 28
Schaeffer-Fulton method, 67
serological typing, 279
serotypes, 270
Shigella, 270
simple staining, 62
slide culture of molds, 103
slime mold culture, 100
smear preparation, 58
Smith fermentation tube, 164
soil microbiology, 201-20
solutions
hypertonic, 135
hypotonic, 135
isotonic, 135
SPC, milk, 230
spectrophotometer usage, 97
spore staining, 67
Sporolactobacillus, characteristics of, 178
Sporosarcina, characteristics of, 179
Sporozoa, 28
staining
acid-fast, 62, 69, 70
capsular, 63
fluorescent staining, 70
gram, 64
negative, 56
simple, 62
spore, 67
Staphylococcus, characteristics of, 180
Staphylococcus aureus, 257, 258
Staphylococcus epidermidis, 258
Staphylococcus saprophyticus, 198, 257, 258
starch hydrolysis test, 170
stigma, 28
stock cultures, 152
streak plate techniques, 82
Streptococcus, 257, 267
Streptococcus agalactiae, 262, 267
Streptococcus bovis, 262, 267
Streptococcus faecalis, 262, 267
Streptococcus pneumoniae, 262, 267
Streptococcus pyogenes, 262, 267
Submastigophora, 28
SXT sensitivity test, 267
synergism, microbial, 127
477
Benson: Microbiological
Applications Lab Manual,
Eighth Edition
Back Matter
Index
©TheMcGraw-H
Companies, 2001
Index
Talaromycetes, 50
Tardigrada, 36
temperature
effect on growth, 130
lethal effects, 132
temperature conversion table, 488
thermal death point (TDP), 132
thermal death time (TDT), 132
thermophiles, 130
thylakoids, 30
titer, 285
Tribonema, 28, 31
trichinosis, 289
tryptophan hydrolysis test, 173
turbidimetry, 96
Vaucheria, 28, 31
Veillonella, characteristics of, 181
Voges-Proskauer test, 192
water bears, 36, 37
water fleas, 36, 37
Winogradsky's column, 107
Wright's stain, 290
xanthophylls, 30
yeasts, 48
yogurt production, 243
ultraviolet light, lethal effects, 137
urea hydrolysis test, 173
urease, 173
urinary tract pathogens, 274
use dilution method, 139
Zernike microscope, 12
Ziehl-Neelsen staining method, 69
zooflagellates, 28
Zygomycotina, 50
zygospores, 49
478
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