CURRICULUM VITAE
Laboratoire des Productions, Valorisations Végétales et
Microbiennes (LP2VM), Département de biotechnologie, Faculté des sciences,
Université des Sciences et de la Technologie d’Oran Mohamed Boudiaf
Pr. KACEM Mourad was born in Sétif, Algeria, in 1964. He received the B.S. (in
1984), D.E.S. (in 1988), M.S. (in 1993), and Doctorat és Sciences (in 2003). He
received the Professor Grad (in 2010). His degrees are in molecular biology and
genetics from Oran University. He is currently with the Biotechnology
Department, Faculty of Sciences, University of Sciences and Technology, Oran,
Algeria, where he also occupies a position in several projects in the Laboratory of
Microbiology and Biotechnology. His research fields are on Lactic acid bacteria,
Rhizobia and Pseudomonas bacteria, metabolites, antibiotics, and bacteriocin
production. He has presented more than 40 papers in national and international
journals and conferences.
Microbiology
Chapter 1 Microorganisms in the Environment
o 1-1 Microbes in the environment
o 1-2 Hay infusion - you try it
o 1-3 Sampling various environments
Chapter 2 Basic Pure Culture Techniques
o 2-1 Aseptic technique
o 2-2 Flame sterilization and tube transfer
o 2-3 Making a medium - You try it
o 2-4 Streak plates
o 2-5 Spread plates and dilution plating
o 2-6 Pour Plates
Chapter 3 Microscopy and staining
o 3-1 Use of the microscope
o 3-2 Operating procedure
o 3-3 Common Problems
o 3-4 Proper care of the microscope
o 3-5 Staining microorganisms
o 3-6 Preparation of a Bacterial Smear for Staining
3-7 The Simple Stain
o 3-8 The Gram Stain
o 3-9 The Endospore Stain
o 3-10 The acid-fast stain
o 3-11 Practice staining
o 3-12 Summary of Microscopy and Staining
Chapter 4 Quantitative Microbiology
o 4-1 Why enumerate microbes
o 4-2 Viable plate counts
o 4-3 The mechanics of dilution plating
o 4-4 Calculating CFU from dilution plating results
o 4-5 A dilution plating protocol
o 4-6 Direct particle counts
o 4-7 Using a counting chamber
o 4-8 Indirect methods
Chapter 5 Selected aspects of bacterial growth and nutrition
o 5-1 Bacterial nutrition
o 5-2 Protocol for making media
o 5-3 Oxygen relationships
o 5-4 Example of a growth factor requirement and protocol
o 5-5 Growth of microbes in batch culture
o 5-6 Generating a growth curve
o 5-7 Examples of growth curve data and graphs
o 5-8 Summary of bacterial growth and nutrition
Chapter 6 Bacterial Motility
o 6-1 The different kinds of motility in microbes
o 6-2 Investigating bacterial motility by flagella
Chapter 7 Characterization and Identification of Bacteria
o 7-1 Putting a name to a microbe
o 7-2 Classic strain testing
o 7-3 Protocol for inoculation of medium
o 7-4 Typical results for biochemical tests
o 7-5 Determination of unknowns
o 7-6 Antibody tests
o 7-6 Identification by DNA probes and primers
o 7-7 Summary of identification of bacteria
Chapter 8 An Introduction to Bacterial Genetics
o 8-1 Definitions in bacterial genetics
o 8-2 Selection of mutants
o 8-3 DNA transfer by conjugation
o 8-4 Summary of bacterial genetics
Chapter 9 Bacteriophages, the Viruses of Bacteria
o 9-1 Defining and counting bacteriophage
o
9-2 Phage typing
o 9-4 Summary of bacteriophage
Chapter 10 Inhibition and Killing of Microorganisms
o 10-1 Controling the growth of microbes
o 10-2 The antibiotic disk sensitivity test
o 10-3 Streptomyces
Chapter 11 Isolation of Bacteria From Natural Sources
o 11-1 Introduction to isolation
o 11-2 Isolation of photosynthetics
o 11-3 Isolation of Bacillus
o 11-4 Nitrogen fixing bacteria
Chapter 12 Lactic Acid Bacteria and an Introduction to Food
Microbiology
o 12-1 An introduction to lactic acid bacteria
o 12-2 Lactic acid bacteria experiment
Chapter 13 Selected genera of medical importance
o 13-1 Microorganisms are friend and foe
o 13-2 Staphylococcus, Streptococcus and Neisseria
Chapter 14 The enteric bacteria (the family Enterobacteriaceae)
o 14-1 Introduction
o 14-2 Isolation and identification of an unknown enteric
o 14-3 Serological identification of a Salmonella culture
o 14-4 Outline of plating and preliminary screening media used in
experiment 14
Chapter 15 The Bacteriological Examination of Water
o 15-1 Introduction to microbial water analysis
o 15-2 Water analysis procedure
Chapter 16 Pathogenicity and virulence
o 16-1 Koch's Postulates
o 16-2 Soft rot of potatoes
Chapter 17 A final unknown identification
o 17-1 Instructions
o 17-2 Credit
Chapter 18 Appendices
o
Chapter 1 - Microorganisms in the Environment
1 - 1 Microbes in the environment
Microorganisms exist in almost every environment on earth and maybe beyond
this planet. They were most likely the first forms of life and will probably be the
last. It is not too bold a statement to say that if water is in the liquid state and a
source of carbon and energy is present, microorganisms can exist. Microbes can
grow at temperatures from < 0°C (the snow alga, Chlamydomas nivalis) to
113°C (Pyrolobus fumarii). Think about that, a microbe that can grow in water
over 100°C. Water boils at 100°C, how is that possible? The answer is that P.
fumarii grows under the sea at hydrothermal vents [1] where the water pressure
is very high. Figure 1-1 [2] shows a picture of a hydrothermal vent. Water
therefore does not boil above 100°C. P. fumarii has very special proteins and
membranes that help it deal with the intense heat and scientists are just
beginning to understand the adaptations this microbe must make. While
microbes that live at the extreme of temperatures are fascinating, most of the
organisms we will examine will grow at 20-50°C. Microbes are also present in
saturated salt lakes, in acid mine drainage that is below pH [3] 1, in
environments devoid of oxygen, in soil, and on you!
Figure 1-1 A hydrothermal vent
This particlular vent is located in the . Note the large amount of material being
precipitated out of the super-heated water (about 300C) as it comes in contact
with ocean water at the sea floor (4C).
Whether you measure them by population or total biomass [4], microbes are the
most common forms of life present in almost any environment. You contain
more microbes in your digestive track (about 100 trillion) than you have cells in
your body (about 10 trillion). The soil is also teeming with microbes. There is a
rich nutrient source in the form of leaf litter, dead animals and plants, and
organic waste.
Microbes have a large impact on the environments where they grow. In the most
general terms, this impact is in the form of their metabolism [5] where they
consume available nutrients [6], use them to create energy and cell [7] material,
and then discard waste products. They use this bounty to grow to large numbers.
In nature, bacteria [8] do not exist in isolation. Their biosphere [9] is crowded
with many different types of microorganisms competing for food and space.
1 - 2 Hay infusion - you try it
Producing a simulation of a pond environment in a cup is easy and fun to look
at. The method used at UW-Madison is called a hay infusion and despite the
concoctions being made for over 50 years, something novel pops up frequently.
The below directions describe how to make a hay infusion.
1. Go to your nearest body of water and collect a water sample. Any natural
water will work, but tap water will not. Tap water is chlorinated to
remove microbes and the water out of your tap will have enough chlorine
in it to kill or inhibit the growth of microbes.
2. Pour the water into a glass, or disposable cup, and add a handful of hay or
grass to the pond water. A glass you do not care about should be used, as
it is going to get pretty scummy.
3. Let the mixture incubate at room temperature or above for several days. If
desired, the addition of a light source (either the sun or a lamp) will
encourage the growth of photosynthetic microbes.
4. During the incubation, check the infusion and add more pond water as it
evaporates.
5. In 5 to 10 days the broth should turn dark and turbid. Examination under a
microscope will reveal a large number of microorganisms.
Figure 1-2 [10] shows the microbes present in pond water. Watch the video and
make a drawing of the various microbes that you see in the water
1 - 3 Sampling various environments
Although microorganisms are present in or on nearly everything, it is usually not
possible to demonstrate their presence by direct microscopic observation unless
their density is high. However, if sterile culture media are exposed to air or
inoculated with substances such as soil or lake water, a variety of
microorganisms will multiply in the media and can be examined subsequently.
To prove that microorganisms are in or on a substance, it is necessary that all
media and equipment used be sterile and that aseptic technique be employed in
performing inoculations and transfers.
The following procedures are meant to demonstrate colony formation by
microbial cells inoculated onto a petri dish medium [11]. Each cell [12] which
can utilize the medium as a source of nutrients [13] and can tolerate the physical
conditions present (temperature, pH [14], atmosphere, etc.) should multiply,
resulting, during incubation, in a visible colony of like cells. Different-appearing
colonies [15] imply different species of microorganisms; colony appearance is
often used in the characterization of bacterial species. When we observe
colonies, we cannot assume each arose from just one cell originally planted on
the medium, however. A pair, chain or cluster of cells or individual cells which
"land" on the medium in close proximity to each other can multiply and produce
a single colony. Thus, we use the term colony-forming unit when we consider
the common origin for the cells of any colony.
Another term we will often use is culture which is simply a large population of
living cells. Examples include a colony (above), a flask of organisms in a liquid
medium, and a tube of slanted agar medium on which organisms are growing. A
culture of cells, dividing every 20 minutes, can begin with one "new" cell and
result in 16,777,216 (i.e., 224) cells after just 8 hours! A pure culture is
composed of identical cells (except for occasional mutants), possibly having
arisen from one cell. A mixed culture contains two or more different kinds of
organisms.
We often refer to "young" and "old" cultures, depending on how long they have
been incubating since inoculation [16]. We do not, however, refer to "young"
and "old" individual cells in the same way, as the cells of most of the bacterial
species we work with undergo division every 15-30 minutes. Thus, an "old cell"
- just about to divide into two "brand new" cells - may be less than a half-hour in
age!
The three periods of this exercise are designed to coincide with Periods 1
through 3 of Experiment 2 during a regular semester when there are two or more
days between periods.
Period 1
Materials
5 plates of Plate Count Agar (PCA)
4 sterile cotton swabs
1 tube of sterile saline (3-4 ml)
1. Remove the lid from one of the plates. Expose the surface of the medium
to the air for 15-30 minutes and then replace the cover. Label the plate on
the bottom lid. (This is standard procedure for labeling petri plates [17].)
2. For the remaining plates, various sites can be sampled with sterile cottontipped swabs moistened with the sterile saline. Each swab is then streaked
across the entire surface of the medium in a petri plate and then discarded
into disinfectant. (Who knows for sure if we're picking up any pathogenic
organisms?) Examples of various items which can be sampled include
your skin, the lab bench, a doorknob, an appliance, some other object in
the vicinity, and one or more of the environmental samples provided for
microscopic observation in Experiment 2. Discard the tube into the
slanted tray on the discard cart.
3. Incubate the plates by placing them in an inverted (medium side up)
position in the 30°C incubator for 2-5 days. Note: As a rule, we will
always incubate our plates in an inverted position. Otherwise, moisture
collecting on the top lid may drop down on the developing colonies,
causing them to run together.
Period 2
Materials
Demonstrations of colonies of various species of bacteria [18] and molds
Tube containing 1 ml of a soil suspension (a 1/10,000 dilution)
Tube containing 1 ml of lake water (a 1/100 dilution)
2 tubes of melted Plate Count Agar (PCA; 15-20 ml/tube) - in 50°C water bath
2 empty, sterile petri plates
Figure 1-3 Exposure plates
The exposure plates prepared by students in Period 1. The colonies observed on
the plates is dependent upon the sample added to the plates.
Figure 1-4 Feather duster
Microbial contaminants of dust picked up by a feather duster.
Figure 1-5 Demonstration colonies
Three common, and colorful, species. Micrococcus luteus is a common
contaminant of dust.
1. Before observation of the plates prepared last period, another plating
method will be performed:
2. For each of the two samples, dump the entire tube contents into an empty,
sterile petri dish. Observe Figure 1-7 [19] to see the correct method.
3. Obtain two tubes of melted PCA from the water bath. Wipe off the excess
water with a paper towel, and pour the contents of each tube into a petri
dish sitting upright on the table, opening the lid just enough to pour out
the tube. Mix the sample and medium in each dish with a gentle, swirling
motion and let the medium solidify.
4. Incubate the plates inverted at 30°C for 2-5 days.
5. Note the demonstration of colonies of various species of bacteria and
molds. Keep the lids on the plates and observe the colonies through the
top lid.
6. As time permits (i.e., with everything else having been done in Exps. 1
and 2 for today), the following can be done. Space for recording results is
on page 3.
7. Observe the plates from Period 1, noting the various bacterial and mold
colonies. At this point, do not open the plates, especially if molds are
present. (Any fuzzy or hairy colonies are probably mold colonies. Their
spores are very easily dispersed into the air, causing subsequent
contamination problems and perhaps allergic reactions as well!) Note: As
a rule, we always observe colonies through the top lids of the plates. Very
little information about colony characteristics and differences can be
obtained by looking through the medium. Note the various shapes, sizes
and colors of the colonies.
8. From one or more of your plates which do not contain mold colonies,
choose two or more different colonies and record their visible
characteristics. Your observations can be recorded in the appropriate
pages of the observation manual.
9. OPTIONAL: For each of your chosen colonies, prepare a smear (with a
drop of water as described here [20]) and stain by the gram [21]-stain
procedure. What is the gram reaction and morphology of the cells?
Period 3
Figure 1-6 Results of pour plates
Typical colonies found after water and soil pour plates.
1. Observe the plates prepared last period. Note and count the surface and
sub-surface colonies. When counting the colonies, it is handy to draw a
few lines with the wax pencil on the back of the plate to mark off a grid.
The colonies can then be counted easily as you scan the sectors.
2. For the lake water sample, you can determine the density of colonyforming units (CFUs) that were in the original (undiluted) lake water if
you know three things: the dilution of the sample (see the previous page),
the amount inoculated into the plate (1 ml), and the colony count. For
example, if you count 40 colonies on the plate, it follows that 40 CFUs
had been in the 1 ml inoculum. As the inoculum was a 1/100 dilution of
the lake water (i.e., "diluted 100 times"), there would have been 40 X 100
(i.e., 4000) CFUs per ml of the undiluted lake water. This result is
expressed best in scientific notation: 4.0 X 103 CFUs/ml. The same
method applies to the soil sample, but the answer is expressed as CFUs
per gram of the soil.
3. Time permitting (i.e., with everything else having been done in Exps. 1
and 2 for today), macro- and microscopic observation of colonies can be
made as in the previous period.
As the instructor will explain, we treat milliliters and grams as equivalents, for
convenience. Bacterial quantitation will be dealt with more fully in Experiment
4 (with Appendix C), and you will note that we are always interested in the
concentration of CFUs (the number in one gram or ml) rather than the total
number in the entire sample.
Chapter 2 - Basic Pure Culture Techniques
2 - 1 Aseptic technique
Trying to study this mixed population is often difficult and in the tradition of the
scientific method; researchers dissect a system and study each piece in isolation.
For microorganisms this means separating the organisms and getting them into
pure culture. A pure culture is defined as a growth of microorganisms (a culture)
that contains one cell [1] type. It is essential in microbiology to be able to obtain
and preserve pure cultures. Over 100 years ago, Robert Koch [2] devised
methods to achieve this goal and the methods he developed are essentially still
used today. The protocols used to maintain pure cultures are a major part of
aseptic technique and are the subject of this chapter.
The goals of aseptic technique are two-fold. The first objective is to obtain pure
cultures and secondly to prevent cross-contamination. Microorganisms in
culture must not escape into the environment, and microbes in the environment
must not get into the cultures we are studying. It is essential that aseptic
technique be understood and practiced correctly. Contaminated cultures are
worthless for diagnosis or for doing research on, because it is unclear what
microbe is performing any action that is being observed.
Aseptic methods commonly used are flame sterilization [3], tube transfer, streak
plates, spread plates and pour plates. Flame sterilization is an easy method to
insure sterile transfer of a culture from a source to a growth medium [4]. Tube
transfer is useful for moving inocula from one tube to another. Mechanical
dilution by making streak plates is the preferred method for obtaining a pure
culture of a microorganism. Finally, spread plates and pour plates are common
methods for enumerating microorganisms and are sometimes useful for
obtaining isolated colonies.
2 - 2 Flame sterilization and tube transfer
Flame sterilization is a very quick simple method of killing microorganisms on
an inoculating loop or needle. The loop or needle is held inside a flame for a few
seconds to bring it to redness and then cooled. Once cool, the loop or needle can
be used for various culture manipulations. Make sure that the area that contacts
the culture is flamed to redness. Also, be patient and let the loop cool down, this
usually takes about 15-30 seconds. Learning this technique is essential to
everything else you do in microbiology.
Transfer of culture from agar plates to tubes, or from tube to tube, is a common,
simple procedure. It is important to perform these transfers in a consistent and
rapid manner. The following protocols have been found effective.
To transfer a culture from an agar plate to a broth or agar slant:
1. Place the Bunsen burner in front of you and assemble all necessary
equipment with in arms reach. Position everything so that you will not
burn yourself while trying to inoculate your tubes.
2. Label the tube of broth or agar to be inoculated with identifying marks.
The culture, the date, and your initials for example. Place it in a rack in
front of you.
3. Holding the inoculating loop handle, flame the entire wire to redness.
4. When the wire cools (about 15-30 seconds) remove the lid of the plate
with your other hand and obtain an inoculum by removing a small portion
of the surface growth on the agar plate. In most cases you will be picking
an isolated colony. Choose a well isolated one. Do not dig into the agar.
Replace the lid of the plate immediately.
5. Hold the tube to be inoculated with the free hand. Remove the cotton plug
or cap of the tube with the little finger of the hand holding the needle
holder. If a cotton-plugged tube is used, the mouth of the tube should be
passed briefly through the flame to singe off dust and lint particles. (Dust
or lint may fall into the tube and contaminate the medium.)
6. Introduce the inoculum into the tube. When inoculating a tube of broth,
rub the wire against the glass just above the fluid level and then tip the
tube slightly to wash the inoculum into the broth. The wire should not be
rattled against the sides of the tube to shake an inoculum into the broth;
this is unnecessary and may create a dangerous aerosol. If the transfer is
made to an agar slant, a single mid-line stroke over its surface is made
with the wire or loop.
7. Replace the cap or plug (the latter after reflaming the mouth of the tube).
8. Flame the inoculating wire again to redness, slowly to avoid spattering.
Put the loop holder down after the wire cools.
In tube to tube transfers by loop or straight wire, both the tube containing the
inoculum source and the tube to be inoculated are usually in the hand at the
same time. An easy procedure which prevents hand fatigue and the danger of
dropping the tubes is illustrated in Figure 2-1 [5].
Figure 2-1 Tube to tube transfer
The standard method for transferring microbes from one medium to another.
Each of the steps is described in the text.
1. The tubes are positioned in the hand as shown in Fig. 2-1 B-E. Plugs and
caps can be loosened by twisting them.
2. The needle holder is taken in the other hand and the wire flamed and
allowed to cool (Fig. 2-1 A).
3. The plugs or caps are removed with the last two fingers of the hand
holding the inoculating wire leaving the thumb and index finger free to
hold and to manipulate the loop holder with the second finger as a guide
and support. Flame the tops of the tubes.
4. Immerse the inoculating wire into the broth culture or scrape the wire
across a portion of surface growth on an agar slant to obtain inoculum.
Make the transfer from one tube to the other (Fig. 2-1 C).
5. Flame the tubes (Fig. 2-1 D).
6. Return the plugs or caps to the tubes (Fig. 2-1 E).
7. Flame the inoculating wire to sterilize it (Fig. 2-1 F).
At first, this procedure will be awkward, but after some practice it becomes
second nature. Figure 2-2 [6] contains a movie that demonstrates the technique
of flame sterilization and tube transfer.
2 - 3 Making a medium - You try it
Making medium is as simple as cooking and a crude medium can be made in
almost any kitchen with a few utensils and a source of heat. Below is described
the production of a chicken broth medium that will grow many common
microorganisms. The medium will be sterilized by tyndalization [7]. Simply
boiling a medium once, while it will kill most vegetative cells, does not kill
endospores. However, heating encourages the endospores to germinate, and a
second boiling kills these microbes. The boiling process is repeated a third time
to ensure that all spores have germinated and been killed.
1. Add 250 ml (about 1 cup) of water into a glass container or some other
vessel that can stand boiling water. The container should be something
you can cover. Glass bottles that can stand boiling or canning jars work
well.
2. To this add 15 grams, about 1 tablespoon of instant chicken broth crystals.
and stir until dissolved. Cover loosely so that steam can escape, but dust
and dirt cannot enter.
3. Microwave the broth on high for 3 minutes or until it just begins to boil.
4. Let the broth cool and sit for at least 2 hours up to overnight at room
temperature. Make sure that the medium vessel is covered to prevent
contamination from the air. At this point the vegetative cells are dead
and most endospores will germinate.
5. Repeat steps 3 and 4 a second time. This will kill the endospores that
germinated on day 1.
6. Repeat steps 3 and 4 a third time. This will eliminate the remaining
endospores, making the medium sterile.
7. Place the medium in a warm place overnight. If your tyndalization was
done correctly, your medium should remain clear and free of microbes.
8. While you are waiting for your medium, search for a sample you would
like to inoculate into it. Almost any sample should work, but natural
samples are a good idea for this experiment. Samples coming from food
or from your body may contain pathogens. Something you probably want
to avoid.
9. Check your medium. If it is still clear and sterile, add a pinch of your
collected sample to the medium. (The exact amount really does not
matter). Again place the medium in a warm place and check it
periodically. After a few days, you should start to see the medium become
more turbid.
10. If you have the equipment, examine a sample of the medium under a
microscope. What shapes of microbes are present? Is there more than one
species?
11. When finished, carefully dispose of the borth. It is basically a spoiled
food, but the properties of the spoilage microbe are unknown and should
be treated with care.
2 - 4 Streak plates
The streak plate method is a rapid and simple technique of mechanically diluting
a relatively large concentration of microorganisms to a small, scattered
population of cells. The goal is to obtain isolated colonies [8] on a large part of
the agar surface, so that desired species can then be brought into pure culture.
Proper streaking of plates is an indispensable tool in microbiology. In most
cases a closed inoculating loop is used for streaking plates. The wire loop should
not be badly oxidized or pitted or it will fail to dilute the inoculum and will
scratch the surface of the agar. Streak plates can be made from a broth culture,
an agar slant or from an agar plate. It is sometimes convenient to suspend a bit
of growth from a solid surface in sterile saline and use this as a source of
inoculum. Resuspension of colonies or cultures grown on solid surfaces dilutes
the culture and makes streak plating easier. A loopful of inoculum is transferred
from the source and put on the agar surface. When using a large inoculum (a
turbid culture or growth from a solid surface), a small spot is spread during the
initial transfer. If the inoculum is from a lightly turbid suspension, the first phase
of the streaking pattern is begun. Several basic patterns are illustrated in Figure
2-3 [9]. The three-phase streaking pattern is recommended for beginners
because it is most likely to give satisfactory results with suspensions having a
wide range of microbial density.
Figure 2-3 Streaking patterns
There are a number of different methods for mechanically diluting microbes on
a streak plate. The most common method is spreading microbes across a plate as
shown in the first four figures. As the concentration of microbes increases so do
the number of phases. Irrespective of the number of phases, loop is flamed
between each one. The fifth plate shows an alternative method, where the
streaks are not continuous, but are a series of parallel lines. foobar
Choosing a streaking pattern is a matter of individual preference and depends
upon the number of microorganisms in the sample. Figure 2-3 [10]
demonstrates the most common patterns, but they are not the only methods. The
object of any streaking pattern is the continuous dilution of the inoculum to give
many well isolated colonies. For multi-phase streaking it is crucial to flame the
loop before starting the next phase. Note the slight overlap into the previous
phase to pick up a small inoculum. To streak a plate...
1. Flame the loop to sterilize it and let cool.
2. Position the plate so that the spot of inoculum is nearest the hand not
holding the loop (the opposite hand).
3. Lift the plate lid with the opposite hand; just enough to get the loop inside
and touch the loop to the inoculum spot. It is often helpful to treat the
inoculating loop as if it were a pencil - steadying the loop by resting the
heel of the hand against the lab bench.
4. Move the loop back and forth across the spot and then gradually continue
toward the center of the plate as you sweep back and forth. Use a very
gentle and even pressure.
5. When creating each phase, do not worry about keeping each pass across
the plate separate from previous ones.
6. When about 30% of the plate has been covered by the first streaking
phase, remove the loop and flame sterilize it.
7. Repeat the above procedure for the second phase, but this time pick up
some inoculum by crossing into the first phase 2-3 times and then not
passing into it again (Figure 2-3).
8. Repeat as necessary for the third and fourth phases. After streaking the
plate, flame sterilize the loop before setting it down.
Figure 2-4 [11] demonstrates the technique of streak plates in a movie.
2 - 5 Spread plates and dilution plating
An absolute requirement for a microbiologist is to be able to determine the
concentration of microorganisms in a given sample. Various particle-counting
devices, spectrophotometric methods and microscopic techniques have been
used to count cells. However, one drawback to these methods is that they count
dead as well as living cells. The most common method of enumerating viable
cells is the plate-count method. Diluting microorganisms and placing them into
petri plates [12] (or plates) for incubation is another essential technique for
working with microorganisms.This method suffers from some problems. First,
only those organisms which can grow on the medium [13], and at the
temperature and atmospheric conditions of incubation, will divide and develop
into colonies [14]. Second, each colony may not represent the progeny from one
cell [15], as two or more cells (those in clusters, chains or otherwise close to one
another) can give rise to one colony. For these reasons the counts obtained from
the plate-count method are given as the number of colony-forming units (CFU's)
per ml (or gram [16]) rather than the number of cells per ml (or gram). Despite
these drawbacks, the plate-count method is a powerful means by which
concentrations of viable organisms may be estimated. Also, if it is desirable to
count a specific subgroup of microorganisms in a sample, selective media [17]
or special incubation conditions can often be used to encourage the growth of
only this class of organism. More information is available in the chapter on
quantitative microbiology [18].
As microbial quantitation involves the use of pipettes [19]"(or micropipettes
[20]) in preparing dilutions and inoculating plates, the beginning student in
microbiology must become familiar with their use.
Due to the possibility of ingesting pathogens and toxic liquids, mouth-pipetting
is forbidden in the laboratory! Pipettes are filled and subsequently emptied by
the use of propipettes or other pipette bulb. Pay close attention to the
demonstration of their use.
Important Safety Consideration: When fitting the pipette and pipette bulb
together, use very gentle pressure!! Do not jam these items together! (Force is
usually not the answer, a good general rule to live by in this lab and in life for
that matter.) The glass pipette will probably break and possibly cause severe
injury. Handle the pipette only at the top inch or so.
For volumes of 5 ml of less, micropipettes are often the tool of choice.
Instruments are available that are capable of dispensing 5 ml all the way to less
than 1 µl (one one-millionth of a liter). Micropipettes have made it possible to
miniaturize many experiments and greatly decrease the cost of running them.
They are also easy to use and can dispense volumes quickly, increasing the
number of experiments that can be performed in a set amount of time.
Micropipettes are the tool of choice for small volumes.
2 - 6 Pour Plates
A practical and common laboratory technique used in isolating pure cultures or
enumerating the living microorganisms in water, milk, foods, and other
materials is the pour plate technique.
To aseptically transfer liquid into a pour plate, raise one side of a Petri plate lid
only just enough to allow access of the sample (from a tube or pipette). Transfer
a known amount of the sample to the dish and cover immediately with the lid.
Then pour 15-20 ml of sterile agar culture medium which has been melted and
cooled to 45-50°C into the plate as shown in Figure 2-5 [21]. The inoculum and
medium are mixed by gentle rotation ten times in one direction and ten times in
the other direction. The agar must be allowed to solidify completely before the
plates are inverted for incubation. After incubation both surface and subsurface
colonies will be observed.
Figure 2-5 Pour plates
Pour plates allow the addition of larger amounts of liquid (1-5 ml) to an agar
dish. The sample is added to the bottom of a sterile Petri plate. Molten agar is
then added to the plate aseptically. It is important to only open the cover enough
to allow the pouring of the agar. This prevent contamination from the
environment.
Figure 2-6 [22] is a movie demonstrating the pour plate technique.
Chapter 3 - Microscopy and staining
3 - 1 Use of the microscope
The microscope, as shown in Figure 3-1 [1], is one of the most important
instruments utilized by the microbiologist. In order to study the morphological
and staining characteristics of microorganisms such as bacteria, yeasts, molds,
algae and protozoa, you must be able to use a microscope correctly.
Figure 3-1 The light microscope
A modern light microscope. This is an example of the kind used in the teaching
labs at the University of Wisconsin-Madison. The various parts of the
microscope are labeled. Please take the time to become familiar with their
names.
The compound microscope used in microbiology is a precision instrument; its
mechanical parts, such as the calibrated mechanical stage and the adjustment
knobs, are easily damaged, and all lenses, particularly the oil immersion
objective, are delicate and expensive. Handle the instrument with care and keep
it clean.
The microscope is basically an optical system (for magnification) and an
illumination system (to make the specimen visible). To help understand the
function of the various parts of the microscope, we will follow a ray of light as it
works its way through a microscope from the light source, through the lenses, up
to the eye. Figure 3-8 [2] traces the path of light through the parts of the
microscope
Figure 3-8 The path of light through a microscope
Modern microscopes are complex precision instruments. Light, originating in
the light source (1), is focused by the condensor (2) onto the specimin (3). The
light then enters the objective lens (4) and the image is magnified. Light then
passes through a series of glass prisms and mirrors, eventually entering the
eyepiece (5) where is it further magnified, finally reacing the eye.
First let us consider a primary feature of all microscopes, the light source.
Proper illumination is essential for effective use of a microscope. A tungsten
filament lamp usually serves as the source of illumination. If reflected
illumination is used, a separate lamp provides a focused beam of light which is
reflected upward through the condenser lenses by a mirror.
The light from the illuminating source is passed through the substage condenser.
The condenser serves two purposes; it regulates the amount of light reaching the
specimen and it focuses the light coming from the light source. As the
magnification of the objective lens increases, more light is needed. The iris
diaphragm (located in the condenser), regulates the amount of light reaching the
specimen. The condenser also collects the broad bundle of light produced by the
light source and focuses it on the small area of the specimen that is under
observation.
Light then passes up through the slide and into the objective lens where the first
magnification of the image takes place. Magnification increases the apparent
size of an object. In the compound light microscope two lenses, one near the
stage called the objective lens and another in the eyepiece, enlarge the sample.
The magnifying power of an objective lens is engraved in the lens mount.
Microscopes in most microbiology laboratories have three objective lenses: the
low power objective lens (10X), the high-dry objective lens (40X) and the oilimmersion objective lens (100X). The desired objective lens is rotated into
working position by means of a revolving nosepiece.
On both sides of the base of the microscope are the course and fine adjustment
knobs, used to bring the image into focus. Rotation of these knobs will either
move the specimen and the objectives closer or farther apart. The coarse
adjustment moves the nosepiece in large increments and brings the specimen
into approximate focus. The fine adjustment moves the nosepiece more slowly
for precise final focusing. In some microscopes, rotation of the fine and course
adjustment knobs will move the stage instead of the nosepiece.
Magnification alone is not the only aim of a microscope. A given picture may be
faithfully enlarged without showing any increase in detail. The true measure of a
microscope is its resolving power. The resolving power of the lens is its ability
to reveal fine detail and to make small objects clearly visible. It is measured in
terms of the smallest distance between two points or lines where they are visible
as separate entities instead of one blurred image. The resolving power of the
objective lens, engraved on the lens, allows us to predict which objective lens
should be used for observing a given specimen. However, having good
resolution in the microscope does not guarantee a visible image, the resolving
power of the human eye is quite limited. Often further magnification is needed
to obtain a good image.
When the oil-immersion objective lens is in use, the difference between the
light-bending ability (or refractive index of the medium holding the sample) and
the objective lens becomes important. Because the refractive index of air is less
than that of glass, light rays are bent or refracted as they pass from the
microscope slide into the air, as shown in Figure 3-9 [3]. Many of these light
rays are refracted at so great an angle that they completely miss the objective
lens. This loss of light is so severe that images are significantly degraded.
Placing a drop of immersion oil, which has a refractive index similar to glass,
between the slide and the objective lens decreases this refraction, and increases
the amount of light passing from the specimen into the objective lens. This
results in greater resolution and a clearer image.
Figure 3-9 Refraction of light at 100X
Light passing out of the slide, into the air, toward the objective lens is refracted,
due to the different in refractive index between air and glass. While the bending
cause by this difference is not important at 100X and 400X, at 1000X this
refraction is problematic, causing blurring of the image and significant loss of
light. Immersion oil has a refractive index very similar to that of glass.
Placement of a drop of oil between the objective lens and the slide prevents the
bending of light rays and clarifies the image. The blue dashed line represents a
potential light ray if immersion oil is not present. The red dashed line represents
a light ray if immersion oil is present.
The image of the specimen continues on through a series of mirrors and/or
prisms that bend it toward the eyepiece. A further magnification takes place at
the eyepiece producing what is called a virtual image. Total magnification is
equal to the product of the eyepiece magnification and the objective
magnification. Most often eyepiece lenses magnify 10-fold resulting in total
magnifications of 100, 400, or 1000X, depending upon which objective is in
place. Many modern microscopes will also have focusable eyepieces to
compensate for differences between individuals and even between individual's
eyes. The adjustment of these is important and is described below.
3 - 2 Operating procedure
Below we describe detailed directions for the use of a microscope. This will give
you an appreciation of their operation. These directions have been written as
generally as possible, but it may be necessary for your instructor to make
modifications for the exact microscopes you are using. Light microscopes used
in teaching laboratories are designed for ease of use and with some practice
should become automatic.
1. Raise the nosepiece using the course adjustment knob. This provides
greater access for positioning the slide on the stage.
2. Rotate the nosepiece so that the 10X objective lens is in operating
position.
3. Open the iris diaphragm approximately half way.
4. Turn on the in-base illuminator by depressing the push-type switch.
5. Place a stained specimen slide on the stage and with the naked eye
position the specimen directly above the center of the condenser.
6. Use the thumbwheel below the eyepieces to adjust the interpupillary
distance between the two eyepieces. This is important to be able to view
specimens with both eyes, maximizing the quality of the image and
preventing fatigue from prolonged use of one eye.
7. Move the microscope condenser by means of the condenser adjustment
knob until the top of the condenser is almost at the highest position. There
should be enough room to slide a piece of paper between the stage and the
condenser, but no more. This will focus the light onto the slide.
8. Rotate the coarse adjustment knob in a clockwise direction to bring the
10X objective closer to the slide. View through the eyepieces and, without
disturbing the coarse adjustment setting, slowly rotate the fine adjustment
knob until the specimen is in the sharpest possible focus.
9. The left eyepiece tube is focusable to compensate for refraction
differences of the eyes. The correct procedure is to bring the specimen
into sharp focus looking though the right eyepiece only. Then focus for
the left eye by turning the left eye tube collar fully counter-clockwise.
Next, while viewing the specimen with the left eye only turn the knurled
collar clockwise until the specimen is in sharp focus. Do not adjust the
fine adjustment knob during this procedure.
10. Remove an eyepiece to view the back aperture of the objective lens. Close
the condenser iris diaphragm, then re-open until the leaves of the
diaphragm just disappear from view. Replace the eyepiece and view the
specimen. The iris diaphragm may be closed slightly to enhance contrast,
especially when viewing unstained specimens.
Unstained specimens have only minimal contrast with their surrounding
environments. As a result they can usually be viewed more effectively by
setting the diaphragm at or near minimum opening. Reducing the
diaphragm setting increases definition, contrast, and depth of focus but
introduces diffraction problems and sacrifices resolution. Play with the
diaphragm setting and select the best compromise by trial and error.
11. Once the specimen is in sharp focus using the 10X objective lens, it is
then possible to rotate the nosepiece to the 40X objective lens without
changing the position of the coarse adjustment knob. Very little
refocusing with the fine adjustment knob is required since most light
microscope objective lenses are parfocal [4]. Remember that the iris
diaphragm setting must be changed to allow more light to pass though the
sample as the magnification increases.
12. If the specimen is to be viewed using the 100X oil immersion lens,
immersion oil must be applied to the slide.
13. Rotate the 40X objective lens slightly to the side so that a drop of
immersion oil may be placed on the specimen without getting it on the
40X lens.
14. Place a drop of immersion oil in the center of the circle of light formed on
the specimen slide.
15. Carefully turn the nosepiece until the 100X objective lens snaps into
place. The objective lens should be in the oil but must not touch the slide.
16. Increase the light intensity as required and rotate the fine adjustment knob
to obtain a sharp focus of the specimen. If necessary make further
adjustments to obtain optimal illumination.
If the microscope is not parfocal, it will be necessary to lower the objective lens
as close to the slide as possible without touching it. This is done only while
looking at the lens and slide from the side of the microscope. Bring the
specimen into view by slowly raising the objective lens with the coarse
adjustment knob. Next, focus with the fine adjustment knob and adjust the
illumination as necessary. If this is not successful the first time, repeat the entire
procedure.
In many cases, a preparation needs to be observed only under the oil immersion
lens. In this case, first locate the specimen and center it in the field with the low
power objective lens. Then add oil and rotate the oil immersion objective lens
into position.
3 - 3 Common Problems
Certain problems real or apparent, may be encountered while operating your
microscope. Here is a trouble shooting guide to help you if you are having
difficulty focusing a sample.
The sample can be focused at 10X, but it is difficult to find or blurry at 40X.
This is often caused by immersion oil on the 40X lens. Wipe the 40X lens with
lens paper to remove the oil and refocus. This can be prevented by never
viewing a specimen with the 40X objective after adding immersion oil to a slide.
The sample can be focused at 10X but when the 40X lens is rotated in place
it contacts the slide.
In most cases this is caused by the slide being place on the stage upside down
†with the smear facing the stage. Check your slide carefully to make sure
it is placed on the stage correctly.
The fine adjustment knob does not turn in the direction required for sharp
focusing.
This indicates that it has been turned to the limits of its threads, either upward or
downward, as the case may be. Screw it back to about one-half the thread
distance (About four turns), use the coarse adjustment to raise or lower the
objective lens sufficiently to bring the specimen into view; then refocus with the
fine adjustment.
What I am viewing does not look like bacteria.
Check to make sure you are in the correct focal plane that you are focusing on
the smear and not dust on the lenses. To verify this, move the slide while
looking at it. Anything in the smear should move in the field of view.
What I am viewing does not look like bacteria. Part II
If you are in the correct focal plane, there may be problems with smear
preparation. Did you heat fix too much? Was the amount of culture applied
sufficient? Did you stain the slide correctly? Many apparent microscope
problems can be attributed to poor slide preparation.
3 - 4 Proper care of the microscope
The following rules, cautions and maintenance hints will help keep your
microscope in good operating condition.
1. Use both hands when carrying the microscope: one firmly grasping the
arm of the microscope; the other beneath the base. Avoid jarring your
microscope.
To keep the microscope and lens systems clean:
2. Never touch the lenses. If the lenses become dirty, wipe them gently with
lens tissue.
3. If blurred specks appear in the field of view this may be due to lint or
smears on the eyepiece. If the specks move while rotating the eyepiece,
the dust is on the eyepiece and cleaning the outer lens of the eyepiece is in
order. If the quality of the image is improved by changing objective
lenses, clean the objective lens with lens paper.
4. Never leave a slide on the microscope when it is not in use.
5. Always remove oil from the oil-immersion objective lens after its use. If
by accident oil should get on either of the lower-power objective lenses,
wipe it off immediately with lens tissue.
6. Keep the stage of the microscope clean and dry. If any liquids are spilled,
dry the stage with a piece of cheesecloth. If oil should get on the stage
moisten a piece of cheesecloth with xylol and clean the stage, then wipe it
dry.
7. When not in use, store your microscope in its cabinet. Put the low power
objective lens into position at its lowest point above the stage. Be sure that
the mechanical stage does not extend beyond the edge of the microscope
stage. Wrap the electrical cord around the base.
To avoid breaking the microscope:
8. Never force the adjustments. All adjustments should work freely and
easily. If anything does not work correctly, do not attempt to fix it
yourself, immediately notify your instructor.
9. Never allow an objective lens to jam into or even to touch the slide or
cover-slip.
10. Never focus downward with the coarse adjustment while you are looking
through the microscope. Always incline your head to the side with eyes
parallel to the slide and watch the objective as you move it closer to the
slide. This will prevent you from smashing the objective into the slide.
11. Never exchange the objective or eyepiece lenses of different microscopes,
and never under any circumstances remove the front lenses from objective
lenses.
12. Never attempt to carry two microscopes at one time
If you follow these rules, you will never have trouble with your microscope.
3 - 5 Staining microorganisms
Preliminary identification of bacteria [5] is usually based upon their cell [6]
morphology and grouping and the manner in which they react to certain staining
procedures. The purpose of this section is to demonstrate some common staining
reactions used to categorize microorganisms.
Figure 3-2 An unstained bacterial smear
Unstained bacteria are mostly made of water and are nearly transparent when
viewed through a light microscope (pictured on the left). Note that most of the
microbes are not visible, but a dust spec in the center of the field of view is
visible. Stains cling to the positive and negative charges of bacteria, but do not
bind as readily to the background of a slide. They therefore differentiate
microbes from their surroundings. Stained bacteria are shown at 40X and 100X
in the center and right panels.
Unstained bacteria are practically transparent when viewed using the light
microscope and thus are difficult to see as shown in Figure 3-2 [7]. The
development of dyes to stain microorganisms was a significant advance in
microbiology. Stains serve several purposes:
Stains differentiate microorganisms from their surrounding environment
They allow detailed observation of microbial structures at high
magnification
Certain staining protocols can help to differentiate between different types
of microorganisms.
Most dyes consist of two functional chemical groups as shown in Figure 3-3
[8]. The chromophore group, which give dyes their characteristic color; and the
auxochrome group, containing an ionizable chemical structure, which helps to
solubilize the dye and facilitates binding to different parts of microorganisms.
Previously, dyes were classified as acidic or basic, depending upon whether the
pigment was negatively or positively charged at neutral pH [9]. More
accurately, dyes can be referred to as anionic (-) or cationic (+) and this is the
convention that will be used in this manual. Cationic dyes (crystal violet,
methylene blue) will react with groups on bacteria that have a negative charge.
Anionic dyes (eosin, nigrosine) will react with groups that have a positive
charge. Since most bacteria have many positive and negative groups in their cell
walls and other surfaces, they will react with both cationic and anionic dyes.
Figure 3-3 The structure of crystal violet
The auxochrome groups of crystal violet is the charged carbon in the center of
the molecule. This is typically neutralized by a Cl- ion. The chromophore group
consists of the three benzene rings and the central carbon. These structures
readily absorb light.
Staining protocols can be divided into 3 basic types, simple, differential, and
specialized. Simple stains react uniformly with all microorganisms and only
distinguish the organisms from their surroundings. Differential stains
discriminate between various bacteria, depending upon the chemical or physical
composition of the microorganism. The Gram stain is an example of a
differential stain. Specialized stains detect specific structures of cells such as
flagella [10] and endospores.
3 - 6 Preparation of a Bacterial Smear for Staining
Before staining and observing a microbe under a microscope, a smear must be
prepared. The goal of smear preparation is to place an appropriate concentration
of cells on a slide and then cement them there so that they do not wash off
during the subsequent staining procedure. Figure 3-4 [11] demonstrates smear
preparation.
The best smears are made from bacteria [12] that have grown on a solid surface
such as an agar slant or plate. A bit of growth from a culture is mixed with
distilled or tap water to form a slightly turbid solution and this is spread on a
clean grease free slide. When staining broth cultures, a drop of broth is
transferred directly to a slide, using no extra water. The procedure for making a
smear is as follows:
1. If more than one culture is to be examined using the same stain, it is
possible to prepare up to 6 smears on the same slide. Before preparing the
slide, divide it into the appropriate number of sections [13] and clearly
label each section on the underside of the slide.
2. If your culture has been grown on a agar slant or agar plate. Place a small
drop of water on a clean, grease-free slide. Next, using a sterile loop or
straight wire needle, transfer a bit of the growth to the drop of water and
rub the needle around until the material is evenly emulsified. Spread the
drop over a portion of the slide to make a thin film. The suspension
should be only slightly turbid.
3. If you are using a broth culture, the broth culture must have clearly visible
turbidity [14]. Transfer a loopful of culture from the broth onto a clean
grease free slide. Spread the drop over a portion of the slide to make a thin
film.
4. Allow the film to air-dry. To get a good stain, it is important to let the
smear dry completely. Excess water left on the slide will boil during the
fixing stage, causing most microbe present to rupture. Rushing this step
will result in a poor final stain.
5. Once dry, "fix" the smear to the slide by passing the bottom of the slide
through the tip of the burner flame several times for a one second. After
heat fixing, touch the heated portion of the slide to your hand. It should be
comfortably warm, but not burning hot.
6. Take care not to under-fix (the smear will wash off) or over-heat (the cells
will be ruptured or distorted) the slide. The correct amount of heat fixing
is learned by experience.
7. Allow the smear to cool and apply the stain.
3 - 7 The Simple Stain
In a simple stain, the smear is stained with a solution of a single dye which
stains all cells the same color. Differentiation of cell [15] types or structures is
not the objective of the simple stain. However, certain structures which are not
stained by this method may be easily seen, for example, endospores [16] and
lipid inclusions.
Simple stains are, well simple. One makes a smear and the applies a single stain
to the slide. Below is a procedure for a simple stain.
1. Prepare and heat-fix a smear of the organism to be studied.
2. Cover the smear with the staining solution. If crystal violet or safranin is
used, allow one minute for staining. The use of methylene blue requires 35 minutes to achieve good staining.
3. Carefully wash off the dye with tap water and blot the slide dry with
blotting paper, an absorbent paper pad or a paper towel.
Three steps, now wasn't that easy? Figure 3-5 [17] contains a movie
demonstrating the simple stain. Figure 3-10 [18] shows a light micrograph of
what a simple stain should look like.
Figure 3-10 The Simple Stain
A photomicrograph of a simple stain at 1000X magnification. Note that all cells,
regardless of species or cell wall construction, stain the same color.
3 - 8 The Gram Stain
The Gram stain, performed properly, differentiates nearly all bacteria [19] into
two major groups. For example, one group, the gram-positive bacteria, include
the causative agents of the diseases diphtheria, anthrax, tetanus, scarlet fever,
and certain forms of pneumonia and tonsillitis. A second group, the gramnegative bacteria, includes organisms which cause typhoid fever, dysentery,
gonorrhea and whooping cough. In Bacteria the reaction to Gram stain reagents
is explained by different cell [20] wall structures. Gram-positive microbes have
a much thicker cell wall, while that found in Gram-negative microbes is thinner.
Microbes from the Archaea domain contain different cell wall structures than
that seen in microbes commonly found in the lab (Bacteria domain). However,
they will still have a species specific Gram stain reaction, even though the
underlying macromolecular structures are different.
The Gram stain is one of the most useful differential stains in bacteriology,
including diagnostic medical bacteriology. The differential staining effect
correlates to differences in the cell wall structure of microorganisms (at least
Bacteria, but not Archaea as mentioned above). In order to obtain reliable results
it is important to take the following precautions:
The cultures to be stained should be young - incubated in broth or on a
solid medium [21] until growth is just visible (no more than 12 to 18
hours old if possible). Old cultures of some gram-positive bacteria will
appear Gram negative. This is especially true for endospore-forming
bacteria, such as species from the genus Bacillus. In this class, many of
the cultures will have grown for more than 2 days. For most bacteria this
is not a problem, but be aware that some cultures staining characteristics
may change!
When feasible, the cultures to be stained should be grown on a sugar-free
medium. Many organisms produce substantial amounts of capsular or
slime material in the presence of certain carbohydrates. This may interfere
with decolorization, and certain Gram-negative organisms such as
Klebsiella may appear as a mixture of pink and purple cells.
Gram stain procedure
Below is a procedure that works well in the teaching laboratories.
1. Cover the slide with crystal violet stain and wait one minute.
2. After one minute wash the stain off (gently!) with a minimum amount of
tap water. Drain off most of the water and proceed to the next step. It may
3.
4.
5.
6.
7.
help to hold the slide vertically and touch a bottom corner to paper
toweling or blotting paper.
Cover the slide with iodine solution for one minute. The iodine acts as a
mordant (fixer) and will form a complex with the crystal violet, fixing it
into the cell.
Rinse briefly with tap water.
Tilt the slide lengthwise over the sink and apply the alcohol-acetone
decolorizing solution (dropwise) such that the solution washes over the
entire slide from one end to the other. All smears on the slide are to be
treated thoroughly and equally in this procedure. Process the sample in
this manner for about 2-5 seconds and immediately rinse with tap water.
This procedure will decolorize cells with a Gram negative type of cell
wall but not those with a gram-positive type of cell wall, as a general rule.
Drain off most of the water and proceed.
As the decolorized gram-negative cells need to be stained in order to be
visible, cover the slide with the safranin counterstain for 30 seconds to
one minute.
Rinse briefly and blot the slide dry. Record each culture as Gram positive
(purple cells) or Gram negative (pink cells).
Figure in Figure 3-6 [22] demonstrates the Gram stain procedure, while Figure
3-11 [23] shows the results of a Gram stain for gram-positive and gram-negative
negative bacteria.
Figure 3-11 The Gram Stain
A photomicrograph of gram-positive and gram-negative bacteria. Note that
Gram reaction is dependent upon cell wall structure. A) E. coli a common gram-
negative rod found in the colon. B) Staphylococcus epidermidis a gram-positive
cocci found on the skin. C) Bacillus cereus a gram-positive rod found in the soil.
3 - 9 The Endospore Stain
Cells of Bacillus, Desulfotomaculum and Clostridium (and several other, lesserknown genera--see Bergey's Manual) may, as a response to nutrient limitations,
develop endospores [24] that possess remarkable resistance to heat, dryness,
irradiation and many chemical agents. Each cell [25] can produce only one
endospore. It is therefore not a reproductive spore [26] as seen for some
organisms such as Streptomyces and most molds. The endospore is essentially a
specialized cell, containing a full complement of DNA [27] and many proteins,
but little water. This dehydration contributes to the spores resistance and makes
it metabolically inert. The endospore develops in a characteristic position (for its
species) in the vegetative cell. Eventually the cell lyses, releasing a free
endospore. For more information on endospores, read the Figure 3-7 [28].
Endospore Stain Procedure
Endospore stains require heat to drive the stain into the cells. For a endospore
stain to be successful, the temperature of the stain must be near boiling and the
stain cannot dry out. Most failed endospore stains occur because the stain was
allowed to completely evaporate during the procedure.
1. Place the heat-fixed slide over a steaming water bath and place a piece of
blotting paper over the area of the smear. The blotting paper should
completely cover the smear, but should not stick out past the edges of the
slide. If it sticks out over the edges stain will flow over the edge of the
slide by capillary action and make a mess.
2. Saturate the blotting paper with the 5-6% solution of malachite green.
Allow the steam to heat the slide for five minutes, and replenish the stain
if it appears to be drying out.
3. Cool the slide to room temperature. Rinse thoroughly and carefully with
tap water.
4. Apply safranin for one minute. Rinse thoroughly but briefly with tap
water, blot dry and examine. Mature endospores stain green whether free
or in the vegetative cell. Vegetative cells stain pink to red.
Figure 3-12 [29] shows a photomicrograph of an endospore stain.
Figure 3-12 The Endospore Stain
A photomicrograph of an enodspore stain. Spores present in the picture stain
green, while the vegetative cells stain red. A) Staphylococcus epdiermidis which
does not form endospores. B) The endospore-forming rod, Bacillus cereus.
3 - 10 The acid-fast stain
Because of the waxy substance (mycolic acids) present on the cell [30] walls,
cells of species of Mycobacterium do not stain readily with ordinary dyes.
However, treatment with cold carbol fuchsin for several hours or at high
temperatures for five minutes will dye the cells. Once the cells have been
stained, subsequent treatment with a dilute hydrochloric acid solution or ethyl
alcohol containing 3% HCl (acid-alcohol) will not decolorize them. Such cells
are thus termed acid-fast in that the cell will hold the stain fast in the presence of
the acidic decolorizing agent. This property is possessed by few bacteria [31]
other than Mycobacterium.
Microscopic examination of tissues or of sputum stained by the acid-fast
staining procedure is an aid in the diagnosis of tuberculosis. If an individual has
pulmonary tuberculosis, and if the tubercles in the lungs are open, the bacteria
(Mycobacterium tuberculosis) will be present in the sputum. The bacteria which
cause leprosy (Hansen's disease; caused by M. leprae) can also be detected with
this staining procedure. The finding of acid-fast cells in milk, on the skin, or in
feces is of no great signifi-cance, because these bacteria may be commonlyfound saprophytic species of Mycobacterium.
1. After preparation of the heat-fixed smear, place the slide over a steaming
water bath.
2. Place a piece of paper towel or blotting paper over the smear. The paper
should be about as wide as the slide and cover an area just slightly greater
than the smear itself. Saturate the paper with carbol fuchsin and let the
slide remain above the steaming water bath for five minutes. Add more
carbol fuchsin to the paper if it appears the stain is drying out.
3. Allow the slide to cool to room temperature. Remove the paper and wash
off the excess stain with water.
4. Decolorize the smear with acid-alcohol for 10-15 seconds. Wash gently
with tap water.
5. Counterstain with methylene blue for 3 minutes. Rinse the slide gently
and dry.
6. Examine the smear first with the 10X and then the 100X (oil-immersion)
objective. Those cells which retained the primary stain (carbol fuchsin)
through the acid-alcohol treatment are stained red; these are the acid-fast
organisms. Mycobacterium cells characteristically appear as clusters of
long, red rods. All other cells are blue.
Below is pictured an example of an acid-fast stain
Figure 3-17 The acid fast stain
A photomicrograph of Mycobacterium smegmatis (pink) and Micrococcus luteus
(blue) at 1000x magnification. M. smegmatis is acid-fast, retaining the carbol
fuchsin dye, thus appearing pink. M. luteus is not acid-fast, loses the carbol
fuchsin during decolorizaiton, and is counter-stained with methylene blue.
Also, Figure 3-18 [32] is a movie showing the steps of the acid-fast stain.
3 - 11 Practice staining
In the study and identification of bacteria [33], the microscope is indispensable!
The series of micro-scopic observations in this exercise is designed to illustrate
how bacteria may be viewed individually in their basic form, the cell [34]. The
second and third periods herein coincide with those of Experiment 1 where
organisms isolated by the student are examined microscopically (and could be
found to be more interesting than those provided in this exercise!).
Period 1
Materials
Hay infusions and various other items from nature
Slide with smears of Bacillus cereus and Staphylococcus epidermidis
Simple stain.
1. You are provided with a microscope slide with two smears. Following the
directions for microscopy and staining, heat-fix the slide [35], making
sure the slide goes through the flame smear-side up.
2. Gloves are available for the staining procedure. Placing the slide on the
staining rack in the sink, cover the slide with crystal violet for one minute.
For a review, look at the directions for the simple stain [36].
3. Carefully rinse off the dye with tap water and blot the slide dry with paper
towel or blotting paper.
4. With both hands, obtain the light microscope from the cabinet
(corresponding to your desk number). This is the type of microscope
which we will always use to observe stained smears.
5. Unless the instructor has other directions more directly applicable to the
microscope you are using, use the simple procedure described in the
operating procedure [37]. Refer to this procedure as you study the cells in
the two smears in the following steps.
6. Place the slide on the stage such that it is oriented as illustrated above.
Make sure the clips on the stage hold the slide securely.
7. Begin your observations with the Bacillus cereus smear. (See figure
below) When observing this organism with the oil-immersion objective,
you will notice that the cells are relatively large and rod-shaped (bacilli)
and are usually in chains. Record your observations on the next page.
8. Repeat this procedure with the Staphylococcus epidermidis smear. Cells
of this organism are spheres (cocci) which are usually arranged in clusters
(staphylococci) and pairs.
9. When you are through, be sure the microscope is put away properly (i.e.,
all oil wiped off, 10X objective lens in place, stage centered). It is
recommended that you keep the slide. (To remove immersion oil from
smears, place a few pieces of lens paper on the slide to absorb the oil.
Then, add several drops of xylol to the lens paper. Peel the paper, now
soaked with xylol, off the slide. Xylol is flammable! Keep it away from
flames!)
Figure 3-13 Simple stain
A simple stain of S. epidermidis and B. cereus. S. epidermidis (A), B. cereus old
(B), B. cereus young (C)
Wet Mount
1. For observation of living microorganisms, various samples including a
hay infusion [38] are available. To study the microorganisms in the
aqueous materials available, it is necessary to make wet mounts. The
procedure is relatively simple:
2. Using a capillary pipette or inoculating loop, pick up some of the material
from around the surfaces of grass and leaves and from the bottom of the
sample. Place a drop of suspended material on a clean microscope slide.
3. With a toothpick, spread a very thin layer of vaseline over a small part of
the palm of your hand. Take a clean coverslip (always held by the edges)
and gently scrape all four edges along your palm, picking up a thin line of
vaseline along each edge.
4. Place the coverslip directly onto the drop on the slide in such a manner
that some air bubbles are trapped. Place a small, multilayered piece of
paper towel over the coverslip and press down. Discard the piece of paper
towel into the disinfectant.
5. Examine the wet mount with your light microscope or a phase
CONTRAST microscope set up by the instructor at a special station in the
back or side of the lab.
6. Without removing the coverslip, discard the slide into the disinfectant
container on the stage. (Refer to page viii for cleanup directions.)
If you haven't already, Figure 1-2 [39] presents a movie of the types of life
forms found in a hay infusion.
Period 2
Materials
Bacterial cultures growing either in a liquid medium [40] (Heart Infusion Broth)
or on a slant of an all-purpose medium followed by suspension in saline:
Escherichia coli - young culture, incubated 12-15 hours
Bacillus cereus - young culture, incubated 12-15 hours
Bacillus cereus - old culture, incubated 2-3 days
Figure 3-14 Gram stains
Gram stains of demonstration species. Below are shown typical Gram stain
reactions of two species. E. coli (A), B. cereus old (B), B. cereus young (C). The
images are slightly larger than what would be visible in a light microscope to
improve clarity.
1. On one clean glass slide, prepare smears of the three cultures. Go to
smear prepapration [41] if you need a refresher. Only when the smears
have dried completely should the slide be heat-fixed.
2. Perform the Gram [42] stain procedure as described.
3. As with any stained smear, definitive observations are made with the
100X, oil-immersion objective. Refer to the microscope directions
already given [43], remembering to focus the slide initially with the 10X
objective, moving then to the oil immersion objective.
4. Keep in mind that the young cultures of B. cereus and E. coli are your
positive and negative control cultures, respectively, for the observation of
probable gram-variability of the older B. cereus culture.
5. Using the figures below, record your observations in your notebook,
noting the Gram reaction (positive if purple, negative if red) and the
cellular shape. Is there any difference seen between the two cultures of
Bacillus cereus? Is gram-variability evident for the older culture? Recall
from the introduction to Experiment 1 that we can refer to old and young
cultures but should not do so for individual cells. (Remember to discard
the tubes and slides properly)
Period 3
Materials
This experiment will be done in class.
Young bacterial cultures growing on slants of Heart Infusion Agar:
Staphylococcus epidermidis
Pseudomonas fluorescens
An unknown
Record the number of your unknown!
Figure 3-15 Typical reactions of example strains for test
The classic Gram reactions for Staphylococcus epidermidis (A) and
Pseudomonas fluorescens (B). From this, determine whether they are Gram (+)
or Gram (-). Note we do not show an unknown as this must be done in class.
The images are slightly larger than what would be visible in a light microscope
to improve clarity.
1. On a clean glass slide, prepare heat-fixed smears of the three cultures,
noting that these cultures are growing on a solid medium. Therefore the
cells must be dispersed in a drop of water when preparing the smears, as a
smear is always a dried suspension of cells. Take care not to make the
smears too thick! S. epidermidis and P. fluorescens are your positive and
negative control cultures (respectively) for your unknown.
2. Perform the Gram stain procedure and note the Gram reaction and cellular
shape. Record your results. Fill out and turn in your description of your
unkonwn. Save your slide until your graded unknown is returned.
Period 4
Materials
For the capsule [44] stain:
36-48 hour culture of Klebsiella pneumoniae growing on a slant of EMB Agar
(a high-sugar medium)
Dropper bottle of filtered India ink
For the acid-fast stain:
3-day culture of Mycobacterium smegmatis growing on a slant of Trypticase
Soy Agar plus 1% glycerol
18-24 hour culture of Micrococcus luteus (the negative control culture) growing
in Nutrient Broth
Dropper bottles of carbol fuchsin (freshly-made), acid alcohol and methylene
blue
Capsule stain
Figure 3-16 The capsule stain
A capsule stain using India ink at 1000x magnification. The cells of Klebsiella
pneumoniaeare surrounded by a dark background. The capsule is the clear area
surrounding the cells. The photomicrographs is slightly enlarged for clarity.
capsule stain
1. Using the culture of Klebsiella pneumoniae, Place one loopful of water on
a slide and emulsify in it a bit of growth from the slant or plate culture of
the designated organism. Add a drop of filtered India ink to the cell
suspension. It often works out well to place the drop of India ink adjacent
to the cell suspension on the glass slide.
2. Obtain a clean coverslip (no fingerprints, smudges, dirt, etc.) and rim it
lightly with vaseline; the vaseline can be gently scraped from a thin layer
applied to the palm of the hand. Place a small, multi-layered piece (about
1-2 cm2) of paper towel over the coverslip and press down firmly; discard
the paper towel into the disinfectant.
3. Using the regular light microscope, focus initially with the 10X objective,
switching to the 45X objective and then - if needed - the 100X, oilimmersion objective. Adjust the light intensity as required with the iris
diaphragm. The outline of the cell can be seen within the area of the clear
capsule.
4. Alternatively, the phase microscope can be used. Heed the precautions
regarding use of this microscope. Excellent observations can be made
with just the 40X objective lens (which takes no immersion oil).
5. When finished, without removing the coverslip, discard the slide directly
into the disinfectant. Never discard capsule stains and other wet mounts
with the stained smears, as viable cells are still present and the slides must
be disinfected!. Record your observations below.
Acid-fast stain
Figure 3-17 The acid fast stain
A photomicrograph of Mycobacterium smegmatis (pink) and Micrococcus luteus
(blue) at 1000x magnification. M. smegmatis is acid-fast, retaining the carbol
fuchsin dye, thus appearing pink. M. luteus is not acid-fast, loses the carbol
fuchsin during decolorizaiton, and is counter-stained with methylene blue.
acid fast stain.
1. Prepare a mixed smear of two organisms as follows: Place a drop of the
Micrococcus luteus broth culture on a slide. Into this drop, add cells from
the Mycobacterium smegmatis culture. Disperse the cells as much as you
can (the Mycobacterium cells tend to clump), and prepare a smear about
the size of a nickel. Let it air-dry completely, and then heat-fix it well,
passing the slide through the flame an extra one or two times.
2. Perform the acid-fast procedure (page 148, observing the slide with the
regular light microscope) and record your observations below.
3. As with all stained smears, discard the slide in the appropriate container.
3 - 12 Summary of Microscopy and Staining
Staining and viewing microbes under the microscope is often necessary in for
their identification and classification. The identity of a microbe can help in
determining the cause of a disease or the source of food spoilage. Microscopes
also have important roles in genetics, cell [45] structure, biochemistry and many
other scientific disiplines. Hopefully, this short introduction has helped you to
understand the visualization of microorganisms.
Chapter 4 - Quantitative Microbiology
4 - 1 Why enumerate microbes
Many interesting chemical processes take place in the environment, in our
bodies an in the food we eat. In almost all cases microbes play an important roll
in these chemical conversions. Here are a few examples
When mines are abandoned they leave underground tunnels exposed to air
and water. This encourages the growth of sulfur-oxidizing bacteria [1]
that catabolize iron sulfide present in the rock into sulfuric acid. The high
concentration of acid produced will drop the pH [2] of the water to below
2.0, creating a very corrosive liquid that then exists the mine.
One result of urban pollution from cars and smokestacks is the releases
ammonia and other reduced nitrogen compounds into the air. This can be
used as a substrate [3] by ammonia-oxidizing bacteria, forming nitric
acid, which corrodes the surface where theses microbes live. Ammoniaoxidizing bacteria are often found on the surface, and deep in the
crevasses, of stone, and are thus found with statues and natural stone
buildings. The build up of nitric acid causes the decay of these statues and
structures. An example of this type of stone decay is found in the
Cölonge cathedral in Germany.
Microbes contribute to the global cycling of elements, converting carbon,
oxygen, nitrogen, sulfur and phosphorous into their various inorganic and
organic forms. In some cases only microbes are capable of certain
conversions. For example, only microbes can convert nitrogen gas into
ammonia using a process called nitrogen fixation [4].
The decay of dead animals and plants depends upon the action of
microorganisms, releasing the elements lock in the deceased so that they
can be reused by other life forms.
It is worthwhile to understand all of these processes and the microbes involved
in them. However, it is also just as important to know the population of
microbes carrying out these conversions. Just because a microbe has been found
that carries out a process does not mean that it is the major actor on any
chemical conversion. It may be at too low a concentration to contribute
significantly. Therefore, it is often vital to determine the number of microbes as
well as their identity.
There are also many other reasons one might need to known the population of
microorganisms in a given sample. For example, determining the rate at which a
microbe is killed by UV light or heat requires analysis of the number of viable
cells before, and at various times during treatment. In other experiments, it is
important to know that you have the right density for a procedure, such as
required for transforming a plasmid [5] constructed in vitro [6] into an E. coli
strain. Assessment of microbial populations in applied industries is also
important. Many food-processing plants will measure the level and type of
microorganisms present in their food by doing counts on selective medium [7].
In addition, viable cell [8] counts will be performed when optimizing heat
treatments for processing food. Sewage treatment plants will routinely sample
and count the microbes present in their treatment systems to insure the correct
type and numbers of bacteria are present. The microbial count can be
determined using a wide array of techniques. Note that these assays all require
somewhat different information and in different time frames, as explained
below. In this chapter we summarize some of the more common methods and
then do some virtual experiments performing them.
4 - 2 Viable plate counts
Viable plate counts
One of the most common methods of determining cell [9] number is the viable
plate count. A sample to be counted is diluted in a solution that will not harm the
microbe, yet does not support its growth (so they do not grow during the
analysis). In most cases a volume of liquid (or a portion of solid) from the
sample is first diluted 10-fold into buffer and mixed thoroughly. In most cases, a
0.1-1.0 ml portion of this first dilution is then diluted a further 10-fold, giving a
total dilution of 100-fold. This process is repeated until a concentration that is
estimated to be about 1000 cells per ml is reached. In the spread-plate technique
some of the highest dilutions (lowest bacterial density) are then taken and spread
with a sterile glass rod onto a solid medium [10] that will support the growth of
the microbe. It is important that the liquid spread onto the plate soaks into the
agar. This prevents left over liquid on the surface from causing colonies [11] to
run together and the need for dry plates restricts the volume to 0.1 ml or less. A
second method for counting viable bacteria [12] is the pour plate technique,
which consists of mixing a portion of the dilution with molten agar and pouring
the mixture into a petri plate. In either case, sample dilution is high enough that
individual cells are deposited on the agar and these give rise to colonies. By
counting each colony, the total number of colony forming units (CFUs) on the
plate is determined. By multiplying this count by the total dilution of the
solution, it is possible to find the total number of CFUs in the original sample.
Figure 4-1 Dilution plating and viable plate counts
(A) A demonstration of a decimal series of dilutions. The 100 sample is a
concentrated solution of methylene blue. A 0.2 ml portion of this was added to
1.8 ml (1:9 ratio) of 0.85% saline to create the 1:10 dilution. After mixing, 0.2
ml of the 10-1 dilution was added to a second tube containing 1.8 ml to create the
10-2 dilution. This was continued to generate the dilution series.
(B) A series of pour plates demonstrating the appearance of a viable plate count.
The 3 plates show a 10-7, 10-8, and 10-9 dilution of a natural sample. Note how
the number of colony forming units decreases 10 fold between the plates.
One major disadvantage of the viable plate count is the assumption that each
colony arises from one cell. In species where cells grow together in clusters, a
gross underestimation of the true population results. One example of this are
species of Staphylococcus, which is known to form clumps of microorganisms
in solution. Each clump is therefore counted as one colony. This problem is why
the term CFUs per ml is used instead ofâ bacteria per mlâ for the results of
such an analysis. It is a constant reminder that one colony does not equal one
cell. Great care must also be taking during dilution and plating to avoid errors.
Even one error in dilution can have large effects on the final numbers. The rate
at which bacteria give rise to an observable colony can also vary. If too short an
incubation time is used, some colonies may be missed. The temperature of
incubation and medium conditions must also be optimized to achieve the largest
colonies possible so that they are easily counted. Finally, this technique takes
time. Depending on the organism, one day to several weeks might be necessary
to determine the number of CFUs that were present when the experiment started.
Such information may no longer be useful for many experiments.
Despite its shortcomings, the viable plate count is a popular method for
determining cell number. The technique is sensitive and has the advantage of
only counting living bacteria, which is often the important issue. Any
concentration of microorganism can be easily counted, if the appropriate
dilution is plated. It is even possible to concentrate a solution before counting, as
is often done in water analysis, where bacterial populations are usually at low
density. The equipment necessary for performing viable plate counts is readily
available in any microbiology lab and is cheap in comparison to other methods.
Finally, by using a selective medium it is possible to determine the number of
bacteria of a certain class, even in mixed populations. These advantages have
made viable plate counts a favorite of food, medical, aquatic and research
laboratories for the routine determination of cell number.
4 - 3 The mechanics of dilution plating
Typical bacterial populations in a 1 ml or 1 g sample can be as high as 1011
CFU's. If most samples were spread directly onto a plate, there would be so
many organisms that individual colonies [13] would not form and a viable count
would be impossible. Therefore, samples almost always need to be diluted.
Using simple 1/10 or 1/100 dilutions is most convenient and makes later
calculations easier. For a 1/10 dilution 1 part sample is added to 9 parts diluent.
Note this is not 1 part to 10 parts (that would be a 1/11 dilution)! For a 1/100
dilution, 1 part is added to 99 parts.
When a known amount of a sample is added by pipette to a known amount of
diluent (called a dilution blank) the resulting suspension must be thoroughly
mixed. Rolling a tube between the hands is not as effective as giving the tube a
series of firm flicks with the index finger. (End-to-end mixing is not done with
the cotton plugged or capped tubes because they will leak.) A series of dilutions
may be made as desired, and from one or more of these dilutions, known
amounts are inoculated onto the surface of agar media in plates. Care must be
taken to open the plates just enough to admit the pipette, in order to decrease the
risk of contamination.
Following the placement of the inoculum on the surface of the plate, a sterile, Lshaped glass rod (often called a "hockey stick") is used to spread the inoculum
over the entire surface of the medium [14]. The part of the rod which enters the
plate is sterilized by immersion in 95% ethanol followed by passing the rod
through a flame such that the ethanol can burn off. (It should be noted that one
such flaming may not always kill all of the endospores [15] which may be
present on the rod!) After a short period of cooling, the rod is ready for use.
Once diluents are prepared and then added to a plate, the medium is incubated at
an appropriate temperture to allow growth of the microorganisms. Colonies
arising on the plate can then be enumerated, and this count used to calculate the
total number microbe present in the original sample.
4 - 4 Calculating CFU from dilution plating results
How des a count on a plates get converted to CFUs per gram [16] or ml of
sample? Let's illustrate the procedure with an example. Imagine that we perform
the following experiment:
Five ml of milk are added to 45 ml of sterile diluent. From this suspension, two
serial, 1/100 dilutions are made, and 0.1 ml is plated onto Plate Count Agar from
the last dilution. After incubation, 137 colonies [17] are counted on the plate.
This problem may be illustrated as follows:
Figure 4-2 A drawing of the dilution problem
It is normally a good idea to draw out dilution problems until you are
comfortable doing them.
Note that it is often a good idea to draw out dilution problems until you are
comfortable doing them. It will help you to develop a clear picture of what is
being done.
The first step in solving this problem is to work out the total dilution of the
sample. First 5 ml is added to 45 ml; This is a 1/10 dilution.
Figure 4-3 Initial dilution
The initial dilution is a 1 to 10 dilution.
Remember, there are many ways to make 1/10 and 1/100 dilutions. A 0.1 ml to
0.9 ml dilution is the same as a 1 ml to 9 ml dilution and a 13 ml to 117 ml
dilution. Next, 1 ml of the first dilution is added to 99 ml to make the second
dilution, that is a 1/100 dilution. This is repeated with third dilution giving
another 1/100 dilution. Then 0.1 ml of the third dilution is plated out on a plate
of PCA. The total dilution of the sample is cumulative and can be represented
mathematically as....
Figure 4-4 Calulating total dilution
The total dilution for the problem
Notice that the amount put on the plate is also a dilution. Normally CFUs are
reported per ml or per gram. In some cases less than 1 ml is put on the plate and
this must be taken into account. One way to solve this, is to factor it into the
total dilution. In this problem 0.1 ml was added to the plate, or 1/10th of a ml.
So multiply the total dilution by 1/10 for the amount added to the plate. This
leaves the total dilution as one-one millionth. The next step is to work out the
dilution factor. The dilution factor is the reciprocal of the total dilution. In this
case it would be......
Figure 4-5 Dilution factor
A mathematical representation of the diluction factor.
Finally, multiply the total dilution by the average number of colonies in the
plate(s) and report your answer in CFUs/ml or CFUs/gram depending upon
where the sample came from; in this case ml because we used milk as a sample.
Figure 4-6 Total colony forming units
A calculation of the total number of CFUs in the original milk sample.
With enough practice, dilution problems can be worked out quite easily and
rapidly. The method described above is just a suggested approach, if you find
another way to do these problems which is more intuitive for you, use it. When
doing dilution problems, remember the following:
Note that using this method, the answer in CFUs per one milliliter or per
one gram is derived. Answers may need to be adjusted if the number of
CFUs per sample (other than a milliliter or gram) is requested. Assume 1
gram = 1 ml. (1 ml of water does indeed weigh 1 gram. That is actually
how the ml is defined.)
Use only those plates with colony counts between 30 and 300. With
duplicate or triplicate plating from the same dilution, take the average of
the plate counts and then proceed.
Note that all individual dilutions and the amount plated are multiplied
together.
The initial dilution is often different from the subsequent dilutions. This is
generally due to the nature of the sample available for analysis.
Decimal (1:10) dilutions can be made by adding 1 ml to 9 ml.
Proportional amounts can be utilized such as by adding 0.1 ml to 0.9 ml
or 11 ml to 99 ml.
Centimal (1:100) dilutions can be made by adding 1 ml to 99 ml. This can
also be done by adding 0.1 ml to 9.9 ml.
Note that plating 0.1 ml of a 10-4 dilution results in the same dilution
factor (105) as plating 1 ml of a 10-5 dilution.
Here is another sample problem. Using any method you choose, solve the
problem.
One ml of a bacterial culture is pipetted into a 9 ml dilution blank. One-tenth ml
of this dilution is pipetted into a 9.9 ml dilution blank. From this dilution onetenth ml is plated using 25 ml of Plate Count Agar. 219 colonies arise after
incubation. How many colony-forming units were present per ml of the original
culture?
The correct answer: 2.19 X 106 CFUs / ml. (Note, the 25 ml of Plate Count Agar
plated is irrelevant. Why?)
4 - 5 A dilution plating protocol
Before beginning the dilution plating exercise, it is necessary for you to learn
how to use micropipettors. We will be using them in this experiment and
throughout the semester. Each micropipette costs $200 so please treat them with
extreme care. Always adhere to the following rules when using micropipettors:
Never set the pipettor to above the upper limit or below the lower limit.
Figure 4-19 [18] lists the limits for the micropipettes [19] that we use in
this course
Figure 4-19 Limits of micropipettes
Micropipette Volume Range (µl)
p20
0.5-20
p200
20-200
p1000
200-1000
The limits of the micropipettes. Never go above or below these ranges as
it may damage the micropipette.
Never point a pipettor up. This can cause liquid to run down into the
pipettor and destroy it's parts.
When withdrawing liquids with the pipettor, always release the plunger
slowly. This prevents liquid from rushing into the end of the pipette and
clogging it up and is especially important with large volume pipettors
(200-1000 µl).
Be sure you use the proper size tip for each pipettor.
Always use a new tip for each different liquid.
Use the correct pipettor for the volume that is to be dispensed. Never use
the 200-1000 µl pipette to dispense volumes below 200 µl.
Protocol for Experiment 2
Period 1
Materials
Test tube containing solution to pipette (practice solution)
4 1.5-ml microcentrifuge tubes
20-200 µl micropipette
200-1000 µl micropipette
Yellow pipette tips
Blue pipette tips
Microcentrifuge
1. Remove four 1.5-ml microcentrifuge tubes from the beaker. Label each
tube 1 through 4 on the frosted labeling spot on the side. Place each in a
microcentrifuge rack.
2. Add the volumes of water listed in Figure 4-20 [20] to tubes 1 through 4.
Figure 4-20 Volumes to add
Tube Measurement 1 Measurement 2 Measurement 3 Measurement 4
1
25
35
45
70
2
30
60
85
-
3
125
175
450
250
4
750
200
-
50
Add these volumed to tubes 1 through 4 as listed.
3. To add the first volume (Measurement 1), take the 20-200 µl micropipette
and set the dial to 20 µl. Follow the protocol below to dispense the
volume into the tube:
4. While holding the pipettor, open the box of yellow tips and firmly press
the end of the pipettor into a tip. Remove the pipettor from the box, the tip
should come along for the ride. Close the pipette tip box.
o Press the plunger down to the first stop and place the end of the
pipettor tip into the practice solution. The end of the pipette tip
should be below the surface of the liquid, but not touching the
bottom of the tube. Slowly release the plunger until it stops and
then remove the tip from the liquid dragging it along the side of the
test tube as you do. This will remove any excess liquid clinging to
the side of the pipette tip.
o Take the tube you labeled 1 and hold it at eye level. Place the
pipette tip against the side of the microcentrifuge tube and expel the
liquid by pushing the plunger down until the first stop. As you drag
the tip along the side, push the plunger to the second stop to blow
out any remaining liquid. Remove the micropipette from the tube.
o
o
o
o
o
To remove the pipette tip, hold the pipettor over the tip discard tray
and press the white tip eject button located near your thumb. This
will eject the tip.
Repeat step 3 for each measurement that you add to the tubes.
Continue until both tube 1 and tube 2 have been completed. Take
your tubes and place them opposite one another in the
microcentrifuge. It is very important that tubes are balanced in the
microcentrifuge, so make sure you place directly across from one
another. If you have questions, please ask your instructor.
Centrifuge the tubes for 2-3 seconds to force all liquid to the
bottom of the tubes. Remove the tube and return to your lab bench.
Since you added a total of 170 µl to tubes 1 and 2, set the 20-200 µl
micropipettor for 170 µl and withdraw the tubes contents. If the
tube volume exactly fills the micropipette tip, it is time to celebrate!
You did it right!.
Perform the additions for tubes 3 and 4 in a similar manner. Use the
200-1000 µl pipettor to dispense the larger volumes when
necessary. Next, mix each tube briefly on a vortex mixer, and pulse
2-3 seconds in a microcentrifuge. Since a total of 1000 µl (or 1 ml)
was added to tubes 3 and 4, set the 200-1000 µl pipettor to 1000 µl
and remove the contents of each tube to check the accuracy of your
pipetting.
Feel free to practice more until you are comfortable with the
micropipettes.
Figure 4-7 A drawing of the dilution
A schematic for the dilution plating to be performed. Measure pipetting volumes
carefully and mix tubes thoroughly. Eacg agar plate is inoculated with 0.1 ml.
Period 2
Materials
A 1/10 dilution of hamburger
4 saline or 0.85% saline dilution blanks (9 ml)
4 plates of Plate Count Agar (PCA)
4 plates of MacConkey Agar (MAC)
Pipettors and sterile tips
1. Label one plate of PCA and one plate of MAC for each of the following
plated dilutions (as we defined them on page 121, q.v.): 10-3, 10-4, 10-5 and
10-6.
2. 3. Label the four 9 ml dilution blanks with the dilutions to make of the
hamburger as follows: 10-2, 10-3, 10-4 and 10-5.
3. With the P1000 pipettor and a blue pipettor tip, aseptically transfer 1 ml
of the 1/10 hamburger dilution to the dilution blank labeled 10 -2; discard
the tip into the disinfectant. (Alternatively, if the hamburger dilution is
provided in a 1 ml amount, you can dump the contents of the dilu-tion
blank into the hamburger tube.)
4. Mix this dilution well by holding the tube on the Vortex mixer* for about
5-10 seconds as demonstrated by the instructor; an actual vortex must be
achieved for proper mixing. Note that this tube is a 1/100 (i.e., 10-2)
dilution of the original, undiluted hamburger.
5. Referring to the diagram on the next page, continue making serial
dilutions of the hamburger, using a new pipettor tip for each new dilution.
(Be sure you are discarding the tips into disinfec-tant!) Thus, by making 1
ml inoculations into the 9 ml dilution blanks, we will achieve the hamburger dilutions which you have labeled on the tubes.
6. With the P200 pipettor and a yellow tip, aseptically transfer 0.1 ml of the
10-2 dilution onto each of the plates marked 10-3. (Why are these plates
marked 10-3 and not 10-2?) With new tips, continue in like manner to
inoculate the remaining plates.
If such a device is not available, hold the tube between thumb and
forefinger in one hand and flick the bottom of the tube with the forefinger of the other hand in order to achieve a vortex. (Do not simply roll
the tube between the hands or lightly tap the tube. Don't be too gentle, yet
don't shake the tubes end-to-end!)
7. Perform the following with care! (See page 118 including the safety
precaution.)
o Sterilize a hockey stick according to the directions near the bottom
of page 118, making sure that the glass hockey stick has cooled
sufficiently before immersing it into the ethanol.
-6
o Spread the inoculum over the entire surface of the 10
plates.
Proceed to do the same with the remaining plates, moving from the
more dilute to the more concentrated inocula. During the course of
spreading the plates, the hockey stick need not be re-flamed, as
long as you proceed from the more dilute to the more concentrated
inocula. When all plates have been spread, resterilize the hockey
stick and let it cool before returning it to the drawer.
8. Incubate the plates (inverted!) and incubate at 30°C for 2 days. If the next
period is 3 or more days away, bring the plates to the tray on the stage for
2-day incubation (followed by refrigera-tion which will arrest growth and
development of the colonies [21], preventing overgrowth).
Figure 4-21 [22] depicts a movie showing the dilution plating technique.
Period 3
Figure 4-8 Dilution plating results
Colonies growing on PCA agar
Colonies growing on MAC agar
Example results from a dilution plating. One gram of fresh hamburger hours was
diluted as described in the procedure for this expeirment and then plated on
MAC agar and PCA Row 1, MacConkey Agar; Row 2 PCA. If you are doing
this virtually, you can count the colonies on the plate and determine the CFU per
gram of hamburger. Record your values in your notebook.
1. . Total Aerobic Plate Count: Choose one PCA plate which appears to
have between 30 and 300 colonies on it and count the colonies on the
plate. As in the example above, determine the number of colony-forming
units per gram [23] of the hamburger. Note that this method is technically
neither total nor aerobic with regard to microorganisms. Not all
microorganisms are able to grow on this medium [24] even though it is
termed an all-purpose medium. Aerobic refers to the incubation
conditions, not the oxygen relationship of the organisms. Figure 4-8 [25]
shows a picture of typical results. The PCA plate has been diluted to 10 -4.
This is the total dilution. This would be an appropriate plate to count.
2. . Total Gram-Negative Plate Count: (Note the selective and differential
aspects of this medium as discussed on page 126.) Choose one MAC plate
which appears to have between 30 and 300 colonies on it and count the
colonies on the plate. Determine the number of gram-negative colonyforming units per gram of the hamburger. If distinctly red or pink colonies
can be differentiated clearly, what appears to be the relative proportion of
lactose fermenters? (E. coli is among the gram-negative organisms which
can ferment lactose and thus produce red colonies on MacConkey Agar.)
The MAC plate has been diluted to 10-3. This is the total dilution.
4 - 6 Direct particle counts
Counting chambers
The most direct method of counting microorganism is by the use of a
microscope and a slide with special chambers of known volume. These slides
allow the counting of a small number of cells in a small volume and
extrapolating the result to determine the population. An example of such a
device is shown in Figure 4-10 [26]. A culture is placed on the slide marked
with precise grids. The number of cells present in each grid is counted and an
average determined. Conversion using a formula gives the number of cells per
milliliter in the culture. This method is rapid, a result can be known in just a few
minutes, and is easy to perform. However, it is impossible to distinguish living
cells from dead ones. If this distinction is important, direct microscopic counts
are not the solution. Finally, cultures containing less than 1 million cells per ml
are actually too dilute for direct counts since there will be too few cells in the
very small volume that is actually examined under the microscope for an
accurate count.
Figure 4-10 The Petroff-Hauser counting chamber
The center of the slide contains a precisely machined grid, with each square
(1/20 mm x 1/20 mm) having a known area. The coverslip also rests above the
slide a known distance (typcially 1/50 of a mm). Since each of these dimenstions
is known, it is possible to calculate the number of cells in each square and
pluggin it into a formula. On the left of the figure is a photograph of a PetroffHauser slide. On the right is a grid at 100 x magnification showing the size of
the squares.
a picture of the petroff-hauser cell [27] counter and then what it looks like on a
slide.
Electronic particle counters
Electronic particle counters are useful if the number of bacteria in a sample
needs to be counted on a routine basis. The method is based on the property that
nonconductive particles, such as bacteria, will cause a disruption in an electric
field as they pass through it. A Coulter counter is a type of electronic particle
counter in which there is a small opening between electrodes through which
suspended particles pass, see Figure 4-11 [28]. In this sensing zone, each
particle displaces its own volume of electrolyte, causing a current pulse. The
pulse is noted and recorded as one particle count. By precisely controlling the
rate at which solution passes through the opening, it is possible to get exact,
reproducible counts at a rate of up to several thousand bacteria per second.
Coulter counters are highly dependent upon particle size and those dependent
upon changes in current are near their detectable limits with microorganisms.
Particle counters that use light diffraction as a means of sizing and counting
particles are also manufactured and can detect particle less than 1 µm in
diameter.
Figure 4-11 The coulter counter
A picture of a Coulter Counter. The diagram at the right demonstrates the
opening that the microbes must pass through during counting. When microbes
pass through the aperture the electrical potential across the electrodes is
distrubes, which the dataprocessing system records as a count. In the lower right
is an actual picture of a Multisizerâ„¢ 3 by Beckmann instruments that uses the
Coulter counter priciple.
The advantage of this method is the simplicity of its operation and it
reproducibility. As in microscopic counts, the machine cannot distinguish
between living or dead cells or even between dust and bacteria. Any reasonably
sized particle in the solution will be counted. There is also the expense of buying
the counter, which can cost many thousands of dollars.
4 - 7 Using a counting chamber
The direct cell counts will be demonstrated using Petroff-Hauser cytometers.
These are glass slides with precisely machined chambers and coverslips, so that
cells in very small volumes can be counted using the microscope.
The chamber is formed by laying the coverslip on the elevated borders of the
central well area. See the diagram below:
Figure 4-12 Arrangement of the Petroff-Hauser counting chamber
The drawing shows the correct placement of the cover slip. Once the coverslip is
placed on the slide, a small amount of culture (about 10 µl) is placed in the
well on the left or right of the viewing area. Capillary action then brings the
liquid onto the viewing area with the grid.
1. Using a capillary pipette, place a drop of the broth culture at the edge of
the coverslip. Capillary action will draw the liquid under the coverslip.
Wait 1-2 min. for the movement to stop and the cells to settle. Then with
the low-power objective focus on the grid in the center of the slide; you
should see a crosshatched area containing 25 squares each containing 16
squares:
2. You will not be able to see many bacteria [29] at this magnification so
turn the high dry power lens (40X) into line. do not use the oil
immersion lens with these chamber, as it just goomers the chamber and
coverslip.
3. Count the number of bacteria in 10-15 of the 1/400 mm2 squares and
calculate an average cell [30] number.
4. To be statistically correct you should use a dilution such that there are no
more than three cells in each small (1/400 mm2) square and the total
number of cells counted is at least 100.
Figure 4-13 [31] shows two images of a Petroff-Hauser counting chamber. Use
this to see what it looks like and for performing a count of the microbes.
To determine the concentration of bacteria in the original culture use the
following formula
Figure 4-14 Formula for the counting chamber
Use this formula for calculating the number of cells per ml from the count
obtained using a counting chamber. Nc is the average number of cells counted
per square and D is the dilution of the samples placed on the slide.
For example:
The 103 is there as a conversion factor from mm3 as measured by the chamber to
cm3 (a.k.a. ml) as typically expressed for culture density. Here is a more detialed
explanation of that conversion factor:
1 ml = 1 cm3 = 1 cm x 1 cm x 1 cm
1 cm = 10 mm
so 1 ml = 10 mm x 10 mm x 10 mm
or 1 ml = 103mm3
If you use one drop (without dilution) from a broth culture: and find an average
of 2.31 squares per cell, your results would be:
Figure 4-15 Calculation of cell number from a counting chamber
If an average of 2.31 cells if found in a 10-1 dilution, the formula would appear
as shown here with a result of 4.62 x 108 cells per ml of culture.
4 - 8 Indirect methods
Change in the amount of a cell component
In situations where determining the number of microorganisms is difficult or
undesirable for other reasons, the use of indirect methods can be an excellent
alternative. These methods measure some quantifiable cell property that
increases as a direct result of microbial growth.
The simplest technique of this sort is to measure the weight of cells in a sample.
Portions of a culture can be taken at particular intervals and centrifuged at high
speed to sediment bacterial cells to the bottom of a vessel. The sedimented cells
(called a cell pellet) are then washed to remove contaminating salt, and dried in
an oven at 100-105 °C to remove all water, leaving only the mass of components
that make up the population of cells. An increase in the dry weight of the cells
correlates closely with cell growth. However, this method will count dead as
well as living cells. There might also be conditions where the dry weight per cell
changes over time or under different conditions. For example, some bacteria
[32] that excrete polysaccharides [33] will have a much higher dry weight per
cell when growing on high sugar levels (when polysaccharides are produced)
than on low. If the species under study forms large clumps of cells such as those
that grow filamentously, dry weight is a better measurement of the cell
population than is a viable plate count.
It is also possible to follow the change in the amount of a cellular component
instead of the entire mass of the cell. This method may be chosen because
determining dry weights is difficult or when the total weight of the cell is not
giving an accurate picture of the number of individuals in a population. In this
case, only one component of the cell is followed such as total protein [34] or
total DNA [35]. This has some of the same advantages and disadvantages listed
above for dry weight. Additionally, the measurement of a cellular component is
more labor-intensive than previously mentioned methods since the component
of interest has to be partially purified and then subjected to an analysis designed
to measure the desired molecule. The assumption in choosing a single
component such as DNA is that that component will be relatively constant per
cell. This assumption has a problem when growth rates are different because
cells growing at high rates actually have more DNA per cell because of multiple
initiations of replication.
Turbidity
A final widely used method for the determination of cell number is a
turbidometric measurement or light scattering. This technique depends on the
fact that as the number of cells in a solution increases, the solution becomes
increasingly turbid (cloudy). The solution looks turbid because light passing
through it is scattered by the microorganisms present and the turbidity is
proportional to the number of microorganisms in the solution. The turbidity of a
culture can be measured using a photometer or a spectrophotometer. The
difference between these instruments is the type of light they pass through the
sample. Photometers, such as the Klett-Summerson device, use a red, green or
blue filter providing a broad spectrum of light. Spectrophotometers use prisms
or diffraction gratings supplying a narrow band of wavelengths to the sample.
Both instruments measure the amount of transmitted light, the light that makes it
from the light source through the sample to the detector.
Figure 4-16 Measuring the turbidity of a culture
A spectrophotometer or photometer quantifies the amount of turbidity of a
culture. The amount of light scattered from a solution is proportional to cell
number. The instruments measures light that is not scattered by the sample.
When measuring light scattering it is important to consider the wavelength of
light used a bacterial culture. Microorganisms may contain numerous
macromolecules that will absorb light, including DNA (254 nm), proteins (280
nm), cytochromes (400-500 nm), and possible cell pigments. When measuring
bacteria by light scattering it is best to pick a wavelength where absorption is at
a minimum and for most bacterial cultures wavelengths around 600 nm are a
good choice. However, the exact wavelength chosen is species specific.
The amount of light transmitted through a sample is inversely proportional to
cell number and can be expressed in the following equation.
Figure 4-17 The transmittance equation
The ratio of light hitting the sample (I0) to light passing through a sample (I) is
the transmittance.
Where T is the light transmitted, I0 is the light entering the sample and I is the
light passing through to the detector.
Due to the nature of light scattering, transmittance decreases geometrically as
the cell numbers increase. It is more intuitive to think of the units increasing as
growth increases and for most bacterial analysis, transmittance is converted into
absorbance using the following equation.
Figure 4-18 Absorbance
The absorbance of a sample it the negative log base 10 of the transmittance.
Absorbance increases in a linear fashion as the cell number increases. When
measuring growth of a culture the term optical density [36] (OD) is normally
used to more correctly represent the light scattering that is occurring; under
optimal conditions, little light is actually absorbed by the culture so the term
absorbance is misleading. For most unicellular organisms changes in OD are
proportional to changes in cell number (within certain limits) and therefore can
be used as a method to follow cell growth. If a precise cell number for a given
OD is desired, a standard curve can be generated, where viable plate count or
cell mass is plotted as a function of OD. It also wise to develop a standard curve
to verify that the OD is actually an accurate portrayal of cell growth. After the
standard curve is made, it is then possible to simply measure the OD of the
culture and read the cell number from the curve.
The turbidity of a culture is dependent upon the shape and internal lightabsorbing components of the microorganism and therefore turbidity readings are
species-specific and cannot be compared between different microbes or even
between different strains of the same species. As above, there are microbes that
change cell size or shape at different stages of growth, which introduces some
inaccuracy to this method of cell counting. Also both living and dead cells
scatter light and are therefore counted. However, the method is very rapid and
simple to perform and provides reliable results when used with care, so it is an
extremely common method of real time analysis of prokaryotic populations. In
fact it is one of the methods we will use for measuring cell number in the
experiment on bacterial growth. Turbidometric measurements also do not
destroy the sample.
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Chapter 5 - Selected aspects of bacterial growth and nutrition
5 - 1 Bacterial nutrition
The growth of any microorganism, whether in its natural niche or in a
laboratory, is dependent upon the presence of certain essential compounds in its
environment. In the lab all these components must be provided for in the culture
medium [1] for the organisms to grow. The potential composition of media is as
diverse as the number of microorganisms under study. This section discusses the
basic requirements for all organisms and the components of culture media. If
you want to know more about microbial requirements for growth, read the
chapter on bacterial nutrition in your microbiology textbook
A fundamental and often ignored ingredient in all growth media is water. Water
is an amazing compound. It is the only natural substance found in all three
states; liquid, solid and gas at temperatures commonly found on earth. Water has
a high specific heat index and can hold a large amount of heat. Water is the
universal solvent, dissolving many more substances than any other liquid. All
biological reactions take place in water. Its unique properties allowed the
development of life on this planet. When humans discover life on other planets,
life forms will likely be water-based. Besides being the solvent of choice, water
can also donate H or O to certain reactions, but this contribution is minimal. In
the preparation of most culture media, the first ingredient added is water.
The second most important ingredient added to media is the carbon source. For
many of the media we will use in this course, an organic compound, such as
glucose, will serve as the carbon source. However, this is not always the case.
Autotrophs will utilize CO2 for cell [2] carbon and the medium used for the
growth of some autotrophs consists of just a few salts. In many cases the carbon
source will also serve as the source of energy. E. coli, growing in a minimal
glucose medium utilizes the glucose both as carbon and energy source. In other
cases, another source will be used to generate energy, including inorganic
compounds and light.
Carbon is not the only element needed in relatively large amounts by
microorganisms. Hydrogen, oxygen, nitrogen, phosphorous, sulfur, and
potassium are usually provided in a culture medium. Some microbes can
assimilate these elements in their most simple form (i.e. O2, H2,or N2), while in
other cases they have to be provided as a part of a larger molecule (KNO3,
amino acids [3], MgSO4). Other elements that are also essential, but are not
added in as large a quantity include, magnesium, iron, calcium, and potassium.
Whatever the source, all these elements must be provided to support growth.
Components of Media and its Classification
Since the list of growth requirements is quite extensive, providing all of them to
a microorganism in a culture medium would seem complicated. In practice,
most media is easy to make, but some formulations can be tremendously
difficult. The ingredients added to culture media can range from pure chemical
compounds to extracts and digests of plant or animal tissues. If all the
components of a medium are known both qualitatively and quantitatively, it is
referred to as chemically defined medium [4]. This type of medium is often
used to study the nutritional requirements of an organism or is necessary when
elucidating various metabolic processes. Complex medium [5] contains
components that are extracts or digests whose exact chemical composition is
impossible to determine and often varies from lot to lot. Therefore the exact
amount of ingredients in complex medium is generally unknown. Common
extracts and digests used in the preparation of microbial medium include; brain
heart infusion (boiled, concentrated cow brains and hearts), yeast extract (killed
and purified, dehydrated yeast) and various peptones (a digest of certain plant
and animal proteins). These complex materials can provide carbon and energy
sources, all necessary minerals, and growth factors [6] (known and unknown) to
an organism. Complex medium is often used in diagnostic tests, since they often
provide all necessary components for growth of many different microorganisms.
Media used in the cultivation of microorganisms can also be classified according
to the way in which it is used.
1. A medium that contains only the minimal components necessary for
growth of a microorganism is termed a minimal medium [7]. This type of
medium can be simple, containing a few salts ((NH)4SO4, KH2PO4,
MgSO4) and a carbon source (glucose). Minimal medium can also be
extremely complex. A medium formulated for the growth of Leuconostoc
mesenteroides, a fastidious organism, contains glucose, 7 salts, 19 amino
acids, 4 nucleotides [8], and 10 vitamins [9].
2. All purpose media are able to support the growth of a wide variety of
microorganisms. These media are usually complex. Some examples of all
purpose media include; Brain Heart Infusion Broth, All Purpose Tween
(APT), Penassay agar and Luria Broth.
3. Some media contain ingredients that inhibit the growth of a certain class
of microorganisms. For example, MacConkey's Agar contains bile salts
and crystal violet to inhibit most gram [10] positive microorganisms. This
type of medium is termed selective medium, since it selects for a certain
class of microorganisms. Other examples include Eosin Methylene Blue
(EMB) Agar and Lactose Lauryl Tryptose Broth.
4. Differential media help us distinguish between different groups of
microorganisms by some biochemical or physiological criteria. This type
of medium is useful in identifying the genera of microorganisms under
study. Examples of this type include MacConkey's agar sugar
fermentation [11] broths, and Kliger's Iron Agar.
If a solid medium is necessary, agar is usually added as the solidifying agent.
For plates or slants, a 1.5% concentration of agar is typical. For semi-solid
medium < 0.5% agar is employed. Agar is a complex, long chain polysaccharide
derived from certain marine algae and has several useful properties. When added
to a solution, it melts at 100 °C forming a slightly viscous liquid that solidifies at
~43 °C. After solidification, the agar will not melt unless the temperature is
again raised to 100 °C. This is a tremendously useful property as you will
discover later in the semester. Some other useful properties of agar include its
resistance to microbial degradation and its translucence for easy viewing of
colonies [12] embedded in the agar. One important disadvantage of agar is its
tendency to harbor impurities, which are virtually impossible to completely
extract. With certain organisms, these impurities can sometimes interfere with
nutritional studies, and even inhibit growth. Chemically defined medium that
contains agar must technically be considered complex. If agar presents a
problem in certain studies, silica based solidifying agents are usually used as a
substitute.
We owe the adoption of agar as a solidifying agent in microbiology to Fanny
Eilshemius Hesse. Robert Koch and her husband Walter Hesse were searching
for a method of culturing microorganisms on solid surfaces. Walter had tried
using potato slices and solidified gelatin, both with unsatisfactory results. In
1881, Fanny suggested he try agar (a thickener she often used in cooking). With
this new solid medium, Walter was able to develop pure culture techniques and
discover the causes of many diseases, including tuberculosis.
Sterilization of Media
In almost all cases, once a medium is made, it must be treated to eliminate any
microorganisms contaminating containers, media ingredients, weighing papers,
or other surfaces that come in contact with the medium. If this is not performed
correctly, contaminates arise during incubation, making microbiological
investigations impossible. Sterilization is defined as the inactivation (or
removal) of all life forms (including the pseudo-life forms, viruses [13]) in a
specific area. Culture media must be made sterile without inactivating nutrients
[14] necessary for growth of the microorganism. Equipment and media used in
the microbiology laboratory are most often sterilized using one of the methods
listed in Figure 5-14 [15].
Figure 5-14 Sterilization of media
Method
Mode of action
Materials
Autoclaving - moist
heat (121 �C) under
coagulates proteins
pressure
(15-17
lb/in2).
heat stable items such as
most culture media, glass
and metal, but not plastic.
Oven - dry heat (160
�C
for
several coagulates proteins
hours).
glass and metals but not
liquids nor plastics.
prevents organisms from
passing through the filter, but
Filtration - 0.22 to
does allow viruses to pass so
0.45 m pore size.
therefore not sterilization in
true sense.
Solutions of heat sensitive
compounds such as some
amino acids, vitamins,
some sugars, antibiotics.
Radiation
ultraviolet light or damages nucleic acids
gamma rays.
heat sensitive solids such
as
plastics,
however
effective on surfaces only.
heat sensitive solids such
as some plastics.
Gas-ethylene oxide. inactivates enzymes.
Some common methods for the sterilization of media and instruments. The
methods used depends upon the material and its intended use.
5 - 2 Protocol for making media
Period 1
Materials
13 tubes of Nutrient Broth
2 250 ml flasks
1 graduated cylinder (100 ml)
magnetic stirrer and stir bar
scales and weighing paper
8 empty sterile Petri plates [16]
1 tube of 50% glucose (2 ml)
1 bottle of 3% Agar + MgSO4 (50 ml, melted in 50 °C water bath)
Media ingredients (as listed below)
Minimal Medium - Part A g/L Nutrient Agar g/L
Ammonium Sulfate
7.0 Beef Extract
5.0
Dipotassium Phosphate
7.0 Peptone
3.0
Monopotassium Phosphate 3.0 Agar
Minimal Medium -Part B g/100 ml
Agar
3.0
15.0
Magnesium Sulfate
0.02
Minimal Medium Part C g/100 ml
Glucose
50
To become familiar with what goes into mixing up a batch of medium, 4 plates
of a minimal medium and 4 plates of a complex medium [17] will be prepared.
1. Go to one of the designated measuring areas and weigh out each
component required for the Minimal Medium - Part A. Use the recipe
listed above. Note: The above recipe is in grams/liter and you are making
100 ml not 1 liter of each medium.
2. Carry the measured amounts back to your lab bench. Use the graduated
cylinder to measure 50 ml of distilled water (dH2O) and pour this into one
250 ml flask.
3. Add the first ingredient on the list, gently swirl the flask, and let it
dissolve completely. Then add the next ingredient. Continue until all
ingredients have been added. When finished, place a foam plug in the
flask.
4. Fill the other 250 ml flask with 100 ml of dH2O. In this case we are using
a dehydrated ready-made medium. Weigh out an appropriate amount of
Nutrient Agar powder, add it to the flask, and swirl. That's it! Many types
of media come in dehydrated, premixed form. They are convenient and
cheap. Which brings up a rule to remember, scientists always try to make
necessary chores, like media making, as quick and convenient as possible,
thus freeing their time for more fruitful pursuits.
5. Place the finished medium in the trays provided. When everyone is
finished, attend the autoclaving demonstration. Autoclaving will take
about 45 minutes total. During the waiting period, perform the other
experiments scheduled for today.
6. When the medium is sterile, place it in the 50 °C water bath to cool down.
The minimal medium needs some additions (a carbon source and agar).
Once the minimal medium is cool, add 0.4 ml of 50% glucose (Minimal
Medium - Part C) to the medium and 50 ml of Minimal Medium - Part B
(This is pre-made and is located in the 50 °C water bath). Swirl the flask
and pour the now complete Minimal Medium into 4 plates. Also, swirl the
Nutrient Agar flask and pour 4 plates.
7. Let the plates cool and when completely solidified place them in your lab
bench drawer, upside down, until next period.
Period 3
Materials
2 plates of Minimal Medium (MM)*
2 plates of Nutrient Agar (NA)*
Cultures for streaking
Figure 5-1 A picture of plates prepared plates
The appearance of plates after they are made. The minimal medium is colorless
(left), while the nutrient agar is tan (right).
*Prepared last period by students
Media Making
1. Bring out your plates of MM and NA. Examine them for contamination. If
any colonies [18] are present, do not use that plate. If you're unsure, have
the instructors check them.
2. To test the microbial acceptance of your media, streak the supplied culture
for isolated colonies using a plate of minimal medium and rich medium.
3. Incubate all tubes at 30 °C.
5 - 3 Oxygen relationships
Oxygen has a tendency to form very reactive by-products (H2O2 and O2(superoxide)) inside a cell [19]. These by-products create havoc by reacting with
protein [20] and DNA [21], thus inactivating them. Cells that are able to live in
the presence of oxygen have evolved enzymes to cope with H2O2 and O2- and
thus are not inhibited by O2. Also many anaerobes have oxygen labile Fe-S
centers and no cellular machinery to protect them from the oxidizing power of
oxygen. Organisms that cannot deal with the problems presented by oxygen
cannot survive in air and are killed.
On the basis of oxygen tolerance, microorganisms can be placed into four
classes. Strict aerobes [22] cannot survive in the absence of oxygen and produce
energy only by oxidative phosphorylation. Strict anaerobes [23], in many cases,
generate energy by fermentation [24] or by anaerobic respiration [25] and are
killed in the presence of oxygen. Aerotolerant anaerobes [26] generate ATP
only by fermentation, but have mechanisms to protect themselves from oxygen.
Facultative anaerobes [27] prefer to grow in the presence of oxygen, using
oxidative phosphorylation, but can grow in an anaerobic environment using
fermentation.
Oxygen utilization is a primary diagnostic tool when identifying
microorganisms. Special media has been developed for the purposes of
determining the oxygen relationship and method of metabolism [28]
(fermentation vs. respiration) of microorganisms. One such medium [29],
Thioglycollate Agar is useful for determining the oxygen relationship of a
microorganism. The medium contains thioglycollic acid, cystine and 0.35%
agar, among other things. The thioglycollic acid and agar prevent oxygen from
entering the entire medium. A dye, resazurin, is used as an indicator of the
amount of oxygen in the medium. Resazurin is red in the presence of oxygen
and turns colorless under anaerobic conditions. The medium is steamed just
before use, which removes all oxygen from the tubes. After inoculation [30] and
incubation, oxygen is able to diffuse into the top part of the medium and support
growth aerobically, while the bottom half of the medium remains devoid of
oxygen.
A second medium used to investigate the general type of metabolism used by a
microorganism is glucose O/F medium. This is a rich medium that contains
glucose as primary carbon source. A pH indicator [31] dye, brom thymol blue,
is added and is green/blue under alkaline-Oxidative conditions or yellow under
acidic-Fermentative conditions. Each test organism is inoculated into two tubes
of glucose O/F medium. One tube is overlaid with mineral oil and the other is
not. The mineral oil serves as a barrier to oxygen, which helps to create an
anaerobic environment.
In this experiment you will first investigate the reactions of several known
microorganisms having different types of metabolism. You will determine the
characteristic reactions of thioglycollate medium and glucose O/F medium. You
will then use this information to determine the oxygen relationships and
catabolism [32] type of your two unknown isolates.
Protocol for testing oxygen relationships
Period 1
Materials
8 tubes of Glucose O/F Medium
4 tubes of sterile mineral oil
4 tubes of Thioglycollate Agar (melted, in 50 °C water bath)
Cultures of
Pseudomonas fluorescens
Clostridium sporogenes
Enterococcus faecalis
Escherichia coli
2 plates of Brain Heart Infusion Agar
Anaerobe jar
In this exercise we will be first testing the oxygen relationships of some known
organisms in Glucose O/F medium and Thioglycollate Agar. This will give you
a sense of inoculating test media and allow you to observe their characteristic
reactions.
1. Get four tubes of thioglycollate agar from the 50 °C water bath. The
thioglycollate agar has been steamed for several minutes to drive off any
oxygen. Keep the agar melted by incubating the tubes in a container
containing 50 °C water. Label the tubes with the culture names.
2. Inoculate each culture into one tube of Thioglycollate agar. Mix the tubes
by placing the palm of your hand over the top of the tube and moving the
bottom of the tube in a circular fashion.
3. Incubate the tubes at 30 °C for 2-5 days.
4. Inoculate each culture into two tubes of glucose O/F medium by stabbing
the medium the full length of the tube with the inoculating needle.
5. Overlay one tube of each culture of Glucose O/F medium with 2-3 cm of
mineral oil. What is the purpose of the mineral oil? Incubate at 30 °C for
2-5 days.
6. Divide each plate into four sectors. Label one plate aerobic and the other
anaerobic. Streak each culture onto one sector on each of the plates.
7. Incubate one plate aerobically at 30 °C. Place the other plate in the
anaerobe jar. The air will be evacuated and replaced with H2 + CO2
atmosphere. The jar will also be incubated at 30 °C.
Figure 5-2 Uninoculated thioglycollate agar and Glucose O/F medium
Glucose O/F medium (left) is green and clear. Thioglycollate agar (right) is
yellow and clear with no turbidity visible before inoculation with culture.
Period 2
1. Observe the thioglycollate tubes for the known cultures. Notice the
growth pattern of each organism. Is growth seen throughout the tube? Is
there more growth at the top of the tube? What does that mean? Did any
of the cultures exhibit growth only in the bottom of the tube?
2. Make drawings of each test tube culture in your lab notebook. From
looking at the tubes can you infer which organisms are strict aerobes,
facultative anaerobes, aerotolerant anaerobes or strict anaerobes? Record
this information in your lab notebook.
3. Observe the Glucose O/F medium. Look for growth in the tubes and
production of acid (yellow color). Bacteria [33] that grow only in the tube
without mineral oil have a respiratory form of metabolism. Bacteria that
grow in both tubes and produce acid under anaerobic conditions can also
use fermentation. What would be the reaction of a strict aerobe in this
medium? Record the results in your notebook. Do the results obtained
here agree with the results from the thioglycollate experiment?
4. Observe the BHI plates. Record anaerobic and aerobic growth as + or -.
Do the results here agree with that which was observed for the other
media in this experiment.
5. Catalase test. Add several drops of (H2O2) to each area of growth on the
plates incubated aerobically. Observe through the top lid of the closed
plates so you don't cause an aerosol of live cells to spread from a positive
reaction! A positive reaction is indicated by the constant evolution of
bubbles. Figure 5-15 [34] is a movie of the catalase test for these six
microbes. However, C. butyricum cannot be tested, why?
Figure 5-3 Reactions in thioglycollate agar
After preparation, Thioglycollate Agar will develop a stable oxygen gradient,
with high concentrations of oxygen near the surface of the agar and no oxygen
near the bottom. Microbes will display different growth patters depending upon
their oxygen relationship. Strict aerobes will only grow near the surface of the
agar (Af, Alcaligenes faecalis; Pf, Pseudomonas fluorescens). Aerotolerant
anaerobes grow at the same rate in presence or absence of oxygen (Lp.
Lactobacillus plantarum). Facultative anaerobes will grow throughout the tube,
but will display more growth near the top of the tube (Se, Staphylococcus
epidermidis; Ea, Enterobacter aerogenes). Strict aerobes will only grow in the
presence of oxygen, at the top of the tube (Cb, Clostridium butryricum). Strict
anaerobes will only grow in the bottom of the tube where oxygen is absent.
Figure 5-4 Reactions in Glucose O/F medium
Each organism is inoculated into two tubes of glucose O/F medium, one of
which is covered with mineral oil to exclude oxygen. There are three types of
reactions possible. Microbes that are incapable of utilizing glucose will have a
alkaline reaction (blue color) at the top of the aerobic tube (the one not covered
with mineral oil) and no reaction in the anaerobic tube (Af; Alcaligenes
faecalis). Oxidative microbes, those only capable of respiration, will only grow
significantly in the aerobic tube. Often there will be a small amount of acid
produced, turning the top of the aerobic tube yellow (Pf; Pseudomonas
fluorescens). Those capable of fermentative metabolism will grow in both tubes
and turn the medium yellow due to the production of acid while growing
anaerobically (Ea; Enterococcus aerogenes). The aerobic tube turns yellow for
fermentative organisms because they use up any available oxygen and begin to
ferment available glucose.
Figure 5-5 Growth of microbes on BHI plates
The left side plates were incubated anaerobically, while those on the right were
incubated aerobically. Strict aerobes cannot grow in the absence of oxygen,
while strict anaerobes cannot grow in the presence of oxygen. Note that Pf and
Af grew slightly under anaerobic conditions due to residual oxygen present in
the anaerobe jar. Cultures shown: Se, Staphylococcus epidermidis; Af,
Alcaligenes faecalis; Cb, Clostridium butyricum; Lp, Lactobacillus plantarum;
Ea, Enterococcus aerogenes; Pf, Pseudomonas fluorescens;
5 - 4 Example of a growth factor requirement and protocol
Iron is required by virtually all organisms. Many microorganisms, both
eukaryotes and prokaryotes, synthesize specific iron-chelating compounds called
siderophores [35] (also known as ironophores) which are extracellular (i.e.,
exported outside of the cell [36]) and are involved in the solubilization and
transport [37] of iron compounds into the cell. (Siderophores may perform
additional functions in the metabolism [38] of microorganisms.) Microbiallyproduced siderophores act as growth factors [39] (see Appendix D) for
organisms which cannot synthesize them. Thus, in the mixed microbial
populations of natural environments, siderophores essential to the growth of
certain organisms are supplied by the excessive secretions of these substances
by other organisms which can synthesize them. This experiment will do nothing
more than demonstrate the requirement of an exogenously-supplied siderophore
for growth of a siderophore auxotroph [40] and the ability of the auxotroph to
be supplied the siderophore artificially and by certain other microorganisms.
Period 1
Materials
Saline suspension of cells of the siderophore auxotroph, Arthrobacter flavescens
(strain
Broth cultures of Bacillus subtilis, Streptomyces griseus and Rhodotorula rosei
(a yeast)
3 plates of Brain Heart Infusion (BHI) Agar plates 3 sterile swabs
Sterile paper disc saturated with a solution of Deferrioxamine B (5µg/ml), a
commercially- available siderophore obtained from a species of Streptomyces
and normally used in the treatment of chronic iron storage disease.
Procedure
1. Inoculate each plate of BHI Agar with the Arthrobacter flavescens
suspension by means of the sterile swabs. Make sure each plate is
completely and evenly covered with the inoculum. Discard the swabs into
the disinfectant.
2. At the edge of one of the plates, aseptically (with flame-sterilized forceps)
place a Deferrioxamine-impregnated disc. At the opposite edge of the
same plate, spot-inoculate (just a dab with your loop - not a streak) the
Bacillus subtilis culture.
3. On the second plate, at opposite edges, spot-inoculate Streptomyces
griseus and Rhodotorula rosei.
4. Leave the third plate as is, with no further treatment. This is the control
plate which will show how poorly the A. flavescens grows (if it grows at
all) on a medium [41] not supplemented with the required growth factor
(i.e., the siderophore).
5. Incubate the plates at 30 °C for 3 or more days.
Period 2
Procedure
1. Observe the plates for the presence or absence of satellite growth of A.
flavescens around the inoculation [42] spots and the Deferrioxamine disc.
Which organisms can provide the siderophore? For the organism which
does not appear to provide a siderophore which the A. flavescens can
utilize, would you expect that organism to be producing one anyway?
Figure 5-6 Growth of A. flavescens near the cultures and deferrioxamine
Note the reaction of A. flavescens to the presence of the various cultures. (A,
Rhodotorula rosei a yeast; B, Streptomyces gresius; C, Deferrioxamine B; D,
Bacillus subtilis; E, control plate with nothing added.) Which ones are providing
the required siderophore? Does the deferrioxamine disk make up for the
deficiency? What can you conclude about the deferrioxamine? Why do you
think there is a zone of clearing around the Streptomyces gresius culture?
Example of phenotypic variation
Genotype [43] is defined as the entire array of genes possessed by a cell, i.e., the
sum of the genetic constitution of the organism, a blueprint in code. The
characteristics of an organism which are based on the genotype but expressed
within a given environment make up the phenotype [44] of the organism. Thus,
the genotype represents the potential of the organism, and the phenotype
describes what the organism actually is and does. For example, in some species
of bacteria [45], the production of a pigment is a phenotypic manifestation of
the genotype, and the degree of actual pigment production may be influenced by
temperature or nutritional variations. As another example, the ability to produce
a siderophore is a genetically-controlled characteristic, but it may or may not be
expressed depending on the amount of iron in the environment. When iron is in
low levels, the organism will produce much more of the siderophore in order to
scavenge the iron.
In this experiment, we will utilize an organism, Pseudomonas fluorescens,
which produces a pigmented siderophore (fluorescein) which is easy to detect
with the naked eye if it is produced in sufficient quantity. It also glows
(fluoresces) when illuminated with ultraviolet light. When inoculated onto two
media which differ in the concentration of iron compounds, we will see how the
same organism appears differently on each medium, an example of phenotypic
variation, a difference in fluorescein synthesis.
Since phenotypic characteristics are generally used to differentiate between
microorganisms, it is important that all tests be done in a standardized set of
conditions (medium, temperature, time, etc.).
SPECIAL SAFETY PRECAUTION: BE CAREFUL WHEN USING THE
ULTRAVIOLET LIGHT. USE THE SAFETY GLASSES PROVIDED.
Ultraviolet light damages the retina of the eye, and prolonged exposure to the
light reflecting off the plates can also cause damage.
Period 1
Materials
Broth culture of Pseudomonas fluorescens
1 plate each of Nutrient Agar and Pseudomonas Agar-F
Procedure
1. With a single streak with the loop, inoculate the culture onto each of the
plates.
2. Incubate the plates at 30 °C for 2 or more days.
Period 2
Materials
Ultraviolet lights set up in a dark room
Procedure
1. Observe the plates for production of fluorescein by the culture. This
usually appears as a yellowish-green color diffused into the medium.
2. To observe the fluorescence of the pigment (hence its name), take the
plates to the dark room.
Figure 5-7 Fluorescent pigment produced by Pseudomonas fluoresens
An example of a pigment produced in response to various nutrient conditions.
Note the intense fluorescence of the pigment(fluorescein) under UV light. (A.
Pseudomonas fluorescens growing on nutrient agar, ambient light; B,
Pseudomonas fluorescens growing on nutrient agar, ultraviolet light; C
Pseudomonas fluorescens growing on Pseudomonas Agar F, ambient light; D,
Pseudomonas fluorescens growing on Pseudomonas Agar F, ultraviolet light)
Remove the covers and expose the cultures to the ultraviolet lamp. SEE THE
SAFETY PRE-CAUTION ABOVE REGARDING ULTRAVIOLET LIGHT.
The glowing, yellow-green fluorescence of the pigment is not itself ultraviolet
light, but light within the normal, visible range. Which of the two media
apparently contains more iron? (Note that iron is a trace element in each
medium; this was also the case in the BHI Agar used in Experiment 2.
5 - 5 Growth of microbes in batch culture
The Phases of Growth
Growth in the bacterial context is normally described as an increase in cell [46]
number. Microorganisms, depending upon the specific species, increase their
numbers by binary fission [47], budding or by filamentous growth [48]. Binary
fission is the separation of the initial cell, the mother cell, into two daughter cells
of approximately equal size. This is a very common method of multiplication
and most of the organisms we will investigate divide in this manner.
Figure 5-8 The various forms of division
Cells can increase in number by binary fission, budding or filamentous growth.
This animation demonstrates each of these processes.
Budding division [49] involves the asymmetric creation of a growing bud, on
the mother cell. The bud increases in size and eventually is severed from the
parental cell. After division is complete, the mother cell reinitiates the process
by growing another bud. Yeast and some bacteria [50] (Caulobacter is one
example) use this form of division.
Figure 5-9 Budding cells
A picture of Histoplasma capsulatum var. duboisii cells, a yeast that causes
histoplasmosis, in the process of budding. Picture courtesy of Dr. Libero Ajello
and the Centers for Disease Control and Prevention.
Filamentous growth is characterized by the formation of long, branching, nondivided filaments, containing multiple chromosomes. Sometimes the filaments
will have cross walls separating chromosomes. As growth proceeds, the
filaments increase in length and number. Under nutrient limiting conditions,
some filamentous microorganisms will go through developmental changes, with
a fraction of the filaments differentiating to form spores. The structures and
mechanisms used to form these spores can be spectacular. Streptomyces species
and many molds grow in this manner.
Figure 5-10 Filamentous growth
The bacteria Nocardia asteroides. Note the long branching cells and the circular
spores in the picture.
The best characterized type of growth is binary fission and this is what we will
focus on in this experiment. When grown in liquid medium [51], bacterial
cultures progress through several distinguishable phases, which can be
characterized by plotting the log of the cell number vs. time. Figure 5-12 [52]
shows an example of a typical growth curve with the 4 phases of growth, lag
phase [53], exponential growth phase (also termed balanced growth), stationary
phase [54] and death phase [55].
Figure 5-11 Binary fission
As a cell divides by binary fission, a cross-wall is formed to separate the
daughter cells. In this EM of Staphylococcus aureus, the cross-wall is already
formed to divide the cells.
Figure 5-12 A bacterial growth curve
A bacterial growth curve generated from actual data obtained in the teaching
laboratories at the University of Wisconsin-Madison. The four phases of growth
(lag (1), exponential (2), stationary (3) and death phase(4)) are labeled.
When an organism is inoculated into a fresh medium, it needs to adapt to the
new nutrients [56] available, synthesize RNA, protein [57] and finally replicate
its DNA [58] before starting division. These processes take time and there is no
net increase in cell numbers, thus a lag phase is observed.
In the subsequent discussion, the numbers refer to the figure above. Once the
appropriate enzymes for growth in a particular medium have been expressed
cells begin to multiply. This period of maximal division can last for several
hours or days, depending upon the organism, and is called the log or
exponential growth phase (2).
Eventually the increase in cell number ceases, either because cells stop dividing
or the rate of division equals the rate of cell death, resulting in a stationary
phase (3). This is usually caused by limitation of a nutrient or the accumulation
of a toxic waste product. Depending on the bacterium, stationary phase can last
for several hours to many days.
The final chapter of a growth curve is the death phase (4). An exponential
decrease in the number of organisms due to cell death occurs during this phase.
Some microorganisms never experience a death phase or it is greatly delayed
due to their ability to survive for long periods without nutrients.
Measurement of Bacterial Growth
Growth of microorganisms can be measured by following the increase in cell
number or the increase in a cellular macromolecule such as DNA or protein. In
most cases, the increase in cell number is determined. Cell numbers can be
measured in a variety of ways. For this experiment we will use the viable plate
count, which you are familiar with, and turbidometric measurement, which is
was explained in the chapter on quantification of microorganisms [59].
So which wavelength of light do we use to measure cell numbers? Typically the
shorter the wavelength the greater the sensitivity of the measurement. Some
wavelengths cannot be used, however, because cellular constituents absorb light,
not scatter them. For example proteins absorb light at 280 nm. Likewise the
color of the medium affects absorbance. For this experiment we will be
measuring the growth of E. coli in either a yellow or a clear medium, 600 nm
works well. Beware, however, that the choice of wavelength must be considered
for each experimental condition.
The sample is placed in a sample chamber. The chamber will contain a holder,
an entrance for the selected wavelength, and an exit, leading to the detector. A
critical piece of equipment used in the sample chamber is the cuvette. A cuvette
is a special test tube that holds the sample. Cuvettes must be clean and free of
aberrations, both of which could scatter light, resulting in inaccurate readings.
Most good cuvettes are expensive and must be treated with care!
The actual operation of a spectrophotometer is much simpler than understanding
its parts. There are a few general, rules of thumb, when using any
spectrophotometer.
1. Before you begin, be sure the wavelength selector is set for the
wavelength of light you are going to use. This involves setting the correct
wavelength on the selector, making sure the correct lamp is on for the
wavelength you have selected and that you are using the correct cuvette.
2. Make sure to zero the spectrophotometer. When this is performed you are
adjusting the machine to 100% transmittance. This is done by using a
sample that contains all components of a mixture except the component to
3.
4.
5.
6.
be measured. For example, if you are measuring the growth of B. cereus
in nutrient broth, the machine would be blanked with sterile, uninoculated
nutrient broth. Realize that the machine is being blanked to read 100% T
(or 0 absorbance) with a specific solution in a specific cuvette, with the
cuvette in a specific orientation. This last consideration, nullifies any
aberrations in the cuvette.
With proper technique, nothing should be spilled into the
spectrophotometer. If it is, clean it up and immediately notify an
instructor. This type of spillage usually happens if cuvettes are filled over
a spectrophotometer, a very bad habit to form!
Clean the cuvette after each use by rinsing with distilled water and
allowing to dry upside down in a test tube rack. For more stubborn stains
use a dilute solution containing a mild soap (like Ivory). In rare cases,
difficult blemishes can be removed with dilute acid (1-5%) or with
ethanol. Never let a sample dry in a cuvette! Protein and nucleic acids
form a strong bond with glass and can be impossible to remove.
When measuring the turbidity [60] of a cell suspension, absorbance
values in the range of 0.1--0.8 are acceptable. Readings below 0.1 push
the limits of the spectrophotometer, since it is not sensitive enough in that
range. Absorbance values above 0.8 result in microorganisms casting
shadows onto one another and not being seen by the spectrophotometer. If
a reading is above this range, the culture must be diluted with sterile
medium. If a reading is below this range, concentrate your culture using a
centrifuge.
When graphing growth versus time, you must plot absorbance on a log
scale. Even though absorbance is a logarithmic unit, absorbance readings
are proportional to cell mass which is increasing exponentially during
growth.
5 - 6 Generating a growth curve
In this experiment, the classic bacterial growth curve will be demonstrated. A
culture of Escherichia coli will be sampled at hourly or half-hourly intervals
from the time of inoculation [61] of the culture (0-time) through a 7 to 9-hour
incubation period. The periodic samplings will be plated to determine viable
counts (as colony-forming units per ml of culture) over the incubation period
such that a growth curve may be plotted. From the graph, we may note the
stages in the growth of the culture as it grows into the stationary phase [62].
Additionally we will be able to determine the growth rate and generation time
of E. coli under our experimental conditions from two points in the exponential
phase [63] of the graph.
Figure 5-16 Bacterial Growth
Graphing of bacterial growth on a linear scale
By definition, bacterial growth is cell [64] replication - i.e., growth of the
culture. Most species of bacteria [65] replicate by binary fission [66], where
one cell divides into 2 cells, the 2 cells into 4, the 4 into 8, etc. If this cell
division occurs at a steady rate - such as when the cells have adequate nutrients
[67] and compatible growing conditions - we can plot numbers of cells vs. time
such as on the graph at right. Before too long, we will need to extend the paper
vertically as the population continues to double. For a culture where cells divide
every 20 minutes, one cell can result in 16,777,216 (i.e., 224) cells after just 8
hours - barring nutrient depletion or other growth-altering conditions.
Figure 5-17 Bacterial Growth
Graphing of bacterial growth with cell number on a log scale.
If we were to convert our vertical axis to a logarithmic scale - as on the graph at
right - we will not need as many sheets of graph paper, and we will find that a
steady rate of growth is reflected as a straight line. (On the vertical axis, the
same distance on the paper is covered with each doubling.) This type of graph
paper is called semilogarithmic graph paper on which we will be plotting our
class results. The numbers we plot will fall on the graph at the same place the
logarithms of these numbers would fall when plotted on conventional graph
paper.
The example below shows the type of graph we may obtain from our class data.
We can plot both colony-forming units (CFUs) per ml and absorbance on the
same graph, remembering that the absorbance units should also be on a
logarithmic scale. Rather than "connecting the dots," we draw the best straight
line among our CFU/ml plots to represent the phases of growth - lag,
exponential, and the start of the maximum stationary phase.
Figure 5-18 Two measurements of growth
Example data showing a plot of cell number by VPC and by turbidity.
For the growth rate formula we are about to use, we need to choose two points
on the straight line drawn through the exponential phase, also making note of the
time interval between them. As we will be converting our numbers to logarithms
for the formula, why not choose two points for which the logs are easy to
obtain? (For example, the log of 1X1010 is simply 10.)
Higher CFU/ml = Xt = 1X1010 (at 5.75 hours)
Lower CFU/ml = X0 = 1X108 (at 2.75 hours)
Time interval (in hours) between the 2 points = t = 3
Using the first formula, we find the growth rate which is the number of
generations (doublings) per hour:
Figure 5-19 Caluclating the growth rate
Use this formula to determine the growth rate k
With the second formula, we find the generation time which is the time it takes
for the population to double:
Figure 5-20 Generation time
The generation time is the reciprocal of the growth rate.
When we graph the CFUs/ml and absorbance on the same graph, we would hope
to see an upward trend for both. Sometimes the absorbance continues to rise
after the CFUs/ml level off into the maximum stationary phase. What would be
the cause of that?
With a clear graph, one should be able to determine the generation time without
the use of formulas. Just look for a doubling of the population and the time it
takes for that to happen. For example - in the above graph - the time it takes to
go from 3 X 109 to 6 X 109 appears to be approximately 30 minutes, which is
close to the generation time determined above.
In preparation for this exercise, be sure to read the relevant material in your
textbook, and look over the procedure below.
Precautions regarding observance of aseptic technique:
1. Remember the proper procedure for holding the tubes during inoculations.
Do not allow the plugs or caps to contact the desk top or anything else.
2. When using the pipettor, hold one tube at a time.
3. Do not pipette from tube to tube with the tubes sitting open and vertical in
the test tube rack!
4. Also, one person should not be holding the tubes while another makes the
transfers.
5. Remember to use a new tip each time you begin to use a more dilute
concentration of cells
Period 1
Materials
Samples (5-6 ml) which were taken at hourly or half-hourly intervals from a
culture of E. coli growing in Nutrient Broth+0.2% yeast extract, incubated at
37°C on a shaker. These samples have been kept on ice for use in this
experiment, and each pair will use one sample.
The following are provided for each pair of students:
7-9 nine ml dilution blanks
8 tubes of melted Plate Count Agar (PCA) in test tubes (15-20 ml/tube) - in
50°C water bath
8 empty, sterile petri dishes
Pipettors (P1000) and sterile tips
Spectrophotometer tube and spectrophotometer
1. Each pair will pick up one culture from the ice-water bath on the front
table. Record the number on the tube. It represents the age of the culture
at which time the sample was taken.
2. With the P1000 (blue) pipettor set at 1.0 ml, transfer 1 ml of the culture to
the first nine ml dilution blank (for the first 1/10 dilution to work with in
Step 5).
3. Aseptically dump the remainder of the culture into the small
spectrophotometer tube (to work with in step 4).
4. With the culture in the spectrophotometer tube, one person in the pair will
obtain and record the absorbance reading of the culture while the other
begins the next step.
5. With additional dilution blanks, make dilutions as specified below.
Inoculate 1 ml from each of the four specified dilutions into each of two
petri plates [68]; plate inoculations can be made concurrently with
preparation of the dilutions.
The dilutions to be plated are as follows:
For 0-hour through 2-hour sampling times:
10-4, 10-5, 10-6, 10-7
For 2.5-hour through 4.5-hour sampling times: 10-5, 10-6, 10-7, 10-8
For 5-hour through 9-hour sampling times:
10-6, 10-7, 10-8, 10-9
1. For each plate, obtain a tube of melted PCA from the water bath and pour
the contents into the plate. Mix the medium [69] and inoculum by
carefully swirling and allow the plates to solidify.
2. Incubate the plates inverted at 30°C until the next period.
Period 2
1. Each pair will determine the total plate count (no. of colony-forming
units/ml of culture). Be sure you are counting colonies [70] of all sizes.
(E. coli typically produces small, lens-shaped colonies when growing
below the surface.) Turn your result in to the instructor along with the
absorbance reading. Be sure you have indicated the sampling time!
Results will be compiled and presented next period.
For Your Assignment:
1. Plot the plate count data on semi-logarithmic graph paper. Rather than
generating a growth curve by connecting the dots, draw the best straight
lines through the lag and exponential phases. (Transitions between the
growth phases can be rounded out.)
2. Determine the growth rate and generation time for the particular strain
and cultural conditions in our experiment. Remember that the points you
need to calculate these values are to be taken from the best straight line
drawn through the exponential phase. Do not use individual data points
from the class data. Also, be sure to indicate the proper units (gen/hr or
hr/gen). Show your calculations!
3. Plot the absorbance readings on semi-logarithmic graph paper. Note any
similarities in the graph generated and that for the plate count data. Both
the absorbance and plate count plots can be made on the same graph. Do
not use the absorbance readings for any calculations of growth rate or
generation time.
5 - 7 Examples of growth curve data and graphs
The data shown here was generated by growing E. coli in four different media
and then comparing the results. Before you do anything, think about the various
media and hypothesize about which condition would have the fastest growth
rate. Then, use this data to prepare graphs of the four conditions. Once created,
choose appropriate points and determine the growth rate and generation time of
each culture.
Growth of E. coli in minimal medium [71] + lactose at 37 °C.
Time (hours) VPC1
VPC2
7.80 x 106
1.0
1.08 x 107
2.0
3.20 x 107
2.5
6.40 x 107
3.0
2.02 x 108
3.5
3.56 x 108
4.0
9.60 x 108
4.5
3.60 x 108
5.0
9.50 x 108
2.50 x 108
5.5
1.83 x 109
1.07 x 109
6.0
1.05 x 109
6.5
1.41 x 109
7.0
1.03 x 10100
7.5
1.00 x 109
24.5
4.20 x 108
25.5
1.00 x 108
26.5
7.00 x 108
4.1 x 108
E. coli grown in Minimal medium + glucose
Time VPC1
VPC2
0.0
1.20 x 107
0.5
1.13 x 107
1.0
1.03 x 107 1.40 x 107
1.5
2.48 x 107 2.10 x 107
2.0
6.00 x 107
2.5
8.40 x 107
3.0
1.66 x 108
3.5
3.30 x 108
4.0
2.90 x 109 1.80 x 109
4.5
6.20 x 108 9.40 x 108
5.0
7.50 x 108 1.09 x 109
5.5
2.00 x 109 1.60 x 109
6.0
1.70 x 109 2.42 x 109
6.5
2.00 x 109 1.48 x 109
7.5
3.48 x 109
8.0
4.00 x 109
12.0 4.10 x 109
16.0 3.70 x 109
20.0 8.00 x 108
24.5 1.25 x 108
26.5 1.23 x 108
E. coli grown in Minimal medium + glycerol
Time VPC1
VPC2
0.0
2.52 x 107
0.5
1.11 x 107
1.0
9.80 x 106
1.5
1.96 x 107
2.0
1.65 x 107
2.5
3.80 x 107
3.0
4.10 x 107
3.5
9.10 x 107
4.0
5.80 x 107 1.50 x 108
4.5
2.23 x 107
5.0
2.23 x 107 4.10 x 108
5.5
6.20 x 108 5.10 x 108
6.0
5.00 x 108 7.10 x 108
6.5
1.16 x 108 2.48 x 108
7.5
1.90 x 108
24.5 3.56 x 108
25.5 8.60 x 108 4.10 x 108
26.5 6.30 x 108 6.20 x 108
E. coli grown in Rich medium (Nutritent Broth + Glucose + Yeast Extract)
Time (hours) VPC1
VPC2
0.0
0.5
5.01 x 106
1.0
4.90 x 106
1.5
5.20 x 106
2.0
1.46 x 108
2.5
8.35 x 107
3.0
3.85 x 108
3.5
5.50 x 108
4.0
5.55 x 108
4.5
9.10 x 108
5.0
8.40 x 108
5.5
9.10 x 108
6.0
1.10 x 109
6.5
3.90 x 109 7.00 x 108
7.0
7.40 x 108
7.5
7.70 x 108
25.5
5.00 x 108
26.5
7.70 x 108
After looking at the data and determining your growth rates, did your actual
results match your hypothesis? In either case explain your reasoning.
5 - 8 Summary of bacterial growth and nutrition
Making media
Proper care and maintenance of microorganisms is essential for any successful
experiments, be they in academic research, industry or the health field. If you do
not understand what a microbe requires for growth it is very difficult to study its
properties. Even in the field of environmental microbiology [72], where
unculturable microbes are sometimes studied, the ultimate goal will be to bring
strains into the laboratory where they can be examined in detail. Microbes,
despite their great metabolic diversity, have the same basic requirements for
growth; a carbon source, an energy source and a source of reducing power.
Carbon can come from CO2 or organic molecules, energy will come from light
or chemicals and reducing power is obtained from organic or inorganic
chemicals.
A successful medium [73] for a target microbe must contain all the nutrients
[74] it needs for growth in a form that the microbe can utilize. These are mixed
in an aqueous solution and then sterilized, most often by autoclaving.
Ingredients that cannot stand the heat of the autoclave are often filter sterilized.
Once sterilized, all the components of a medium are mixed together and the
medium is then inoculated with a source of bacteria [75].
Oxygen relationships
One of the more important properties for a microbe is the relationship of the
microbe toward oxygen. Organisms capable of utilizing oxygen in their
metabolism [76] are aerobes, those that cannot are called anaerobes. Microbes
can be separated into 5 categories based upon their use of oxygen. Strict aerobes
[77] require oxygen for their growth, while strict anaerobes [78] cannot survive
in the presence of oxygen. Facultative anaerobes [79] can grow in the absence
of oxygen, but grow better in its presence. Aerotolerant anaerobes [80] have
mechanisms to protect themselves from oxygen, thus being able to grow in its
presence or absence, but do not use oxygen in their metabolism. Finally
microaerophiles [81] require oxygen for their metabolism, but cannot survive at
atmospheric levels of oxygen (21% O2). Microaerophiles are restricted to narrow
bands in between aerobic and anaerobic [82] habitats, where the oxygen
gradient is within an acceptable range. Some examples would be in a lake or
pond, or wet soil.
Measuring bacterial growth and growth curves
For unicellular orgnaisms, growth is normally thought of as an increase in
number. Microbes can growth very rapidly and the most heavily studied
microbes grow by binary fission [83]. A characteristic growth pattern is found
when microbes are growth in a closed vessel. Initially, the microbe adapts to a
fresh medium and no growth is observed. This is called lag phase [84]. The
microbe then begins dividing and the increase in population occurs by a
geometric progression, resulting in explosive growth until nutrients are used up
or toxic end products accumulate. This is termed exponential growth phase. At
this stage growth stops and stationary phase [85] begins. The population
remains constant, sometimes for long periods of time. Eventually, cells
accumulate damage and begin to die. The decrease in population, called death
phase [86], occurs exponentially.
The rate of growth can be calculated by determining the cell [87] number at
different times. The growth rate (k) is a measure of how rapidly the cells are
dividing, the faster cells are growing, the higher the growth rate will be.
Links
1. http://inst.bact.wisc.edu/inst/index.php?module=Book&func=displaygloss
ary#medium
2. http://inst.bact.wisc.edu/inst/index.php?module=Book&func=displaygloss
ary#cell
3. http://inst.bact.wisc.edu/inst/index.php?module=Book&func=displaygloss
ary#amino acids
4. http://inst.bact.wisc.edu/inst/index.php?module=Book&func=displaygloss
ary#chemically defined medium
5. http://inst.bact.wisc.edu/inst/index.php?module=Book&func=displaygloss
ary#complex medium
6. http://inst.bact.wisc.edu/inst/index.php?module=Book&func=displaygloss
ary#growth factors
7. http://inst.bact.wisc.edu/inst/index.php?module=Book&func=displaygloss
ary#minimal medium
8. http://inst.bact.wisc.edu/inst/index.php?module=Book&func=displaygloss
ary#nucleotides
9. http://inst.bact.wisc.edu/inst/index.php?module=Book&func=displaygloss
ary#vitamins
10. http://inst.bact.wisc.edu/inst/index.php?module=Book&func=displaygloss
ary#gram
11. http://inst.bact.wisc.edu/inst/index.php?module=Book&func=displaygloss
ary#fermentation
12. http://inst.bact.wisc.edu/inst/index.php?module=Book&func=displaygloss
ary#colonies
13. http://inst.bact.wisc.edu/inst/index.php?module=Book&func=displaygloss
ary#viruses
14. http://inst.bact.wisc.edu/inst/index.php?module=Book&func=displaygloss
ary#nutrients
15. http://inst.bact.wisc.edu/inst/index.php?module=Book&func=displayfigur
e&book_id=3&fig_number=14&chap_number=5
16. http://inst.bact.wisc.edu/inst/index.php?module=Book&func=displaygloss
ary#petri plates
17. http://inst.bact.wisc.edu/inst/index.php?module=Book&func=displaygloss
ary#protein
18. http://inst.bact.wisc.edu/inst/index.php?module=Book&func=displaygloss
ary#dna
19. http://inst.bact.wisc.edu/inst/index.php?module=Book&func=displaygloss
ary#strict aerobes
20. http://inst.bact.wisc.edu/inst/index.php?module=Book&func=displaygloss
ary#strict anaerobes
21. http://inst.bact.wisc.edu/inst/index.php?module=Book&func=displaygloss
ary#anaerobic respiration
22. http://inst.bact.wisc.edu/inst/index.php?module=Book&func=displaygloss
ary#aerotolerant anaerobes
23. http://inst.bact.wisc.edu/inst/index.php?module=Book&func=displaygloss
ary#facultative anaerobes
24. http://inst.bact.wisc.edu/inst/index.php?module=Book&func=displaygloss
ary#metabolism
25. http://inst.bact.wisc.edu/inst/index.php?module=Book&func=displaygloss
ary#inoculation
26. http://inst.bact.wisc.edu/inst/index.php?module=Book&func=displaygloss
ary#ph indicator
27. http://inst.bact.wisc.edu/inst/index.php?module=Book&func=displaygloss
ary#catabolism
28. http://inst.bact.wisc.edu/inst/index.php?module=Book&func=displaygloss
ary#bacteria
29. http://inst.bact.wisc.edu/inst/index.php?module=Book&func=displayfigur
e&book_id=3&fig_number=15&chap_number=5&width=320&height=
30. http://inst.bact.wisc.edu/inst/index.php?module=Book&func=displaygloss
ary#siderophores
31. http://inst.bact.wisc.edu/inst/index.php?module=Book&func=displaygloss
ary#transport
32. http://inst.bact.wisc.edu/inst/index.php?module=Book&func=displaygloss
ary#auxotroph
33. http://inst.bact.wisc.edu/inst/index.php?module=Book&func=displaygloss
ary#genotype
34. http://inst.bact.wisc.edu/inst/index.php?module=Book&func=displaygloss
ary#phenotype
35. http://inst.bact.wisc.edu/inst/index.php?module=Book&func=displaygloss
ary#binary fission
36. http://inst.bact.wisc.edu/inst/index.php?module=Book&func=displaygloss
ary#filamentous growth
37. http://inst.bact.wisc.edu/inst/index.php?module=Book&func=displaygloss
ary#budding division
38. http://inst.bact.wisc.edu/inst/index.php?module=Book&func=displayfigur
e&book_id=3&fig_number=12&chap_number=5
39. http://inst.bact.wisc.edu/inst/index.php?module=Book&func=displaygloss
ary#lag phase
40. http://inst.bact.wisc.edu/inst/index.php?module=Book&func=displaygloss
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41. http://inst.bact.wisc.edu/inst/index.php?module=Book&func=displaygloss
ary#death phase
42. http://inst.bact.wisc.edu/inst/index.php?module=Book&func=displayarticl
e&art_id=107
43. http://inst.bact.wisc.edu/inst/index.php?module=Book&func=displaygloss
ary#turbidity
44. http://inst.bact.wisc.edu/inst/index.php?module=Book&func=displaygloss
ary#exponential phase
45. http://inst.bact.wisc.edu/inst/index.php?module=Book&func=displaygloss
ary#environmental microbiology
46. http://inst.bact.wisc.edu/inst/index.php?module=Book&func=displaygloss
ary#microaerophiles
47. http://inst.bact.wisc.edu/inst/index.php?module=Book&func=displaygloss
ary#anaerobic
Kacem Mourad