Soil Microbial Ecology - Soil Molecular Ecology Laboratory

SOS 43
SOS4303C / SOS5305C
Soil Microbial Ecology
LABORATORY EXERCISES
ABID AL AGELY
ANDY OGRAM
2006

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INTRODUCTION .......................................................................................................................... 4
LABORATORY OPERATIONS ................................................................................................... 5
1. SAFETY ............................................................................................................................. 5
2. HANDLING BACTERIAL CULTURES .......................................................................... 6
3. AUTOCLAVING ............................................................................................................... 7
MICROSCOPY: ........................................................................................................................... 24
1. OBSERVE SOIL BACTERIA ......................................................................................... 25
2. OBSERVE SOIL FUNGI ................................................................................................. 30
SOIL MICROBIAL ENUMERATION:
1. DILUTION PLATE METHOD........................................................................................ 17
2. DIRECT MICROSCOPIC COUNT ................................................................................. 35
SOIL ALGAE ENUMERATION................................................................................................. 45
SOIL MICROBIAL ASSOCIATION
1. MYCORRHIZAE ............................................................................................................... 8
2. RHIZOBIA ....................................................................................................................... 41
SOIL METABOLIC ASSESMENT
1. ENZYME - PHOSPHATASE .......................................................................................... 47
2. RESPIRATION / BIOMASS............................................................................................ 51
3. DNA.................................................................................................................................. 57
SOIL DECOMPOSITION AND MICROBIAL COMMUNITY STRUCTURE ........................ 14
SOIL RHIZOSPHERE.................................................................................................................. 66

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LABORATORY SCHEDULE
08/29 Field Trip Sampling Forest and Garden Soils
09/12 Exercise 1 Mycorrhizal quantification experiment ...............................8 – 13
09/19 Exercise 2 Initiate soil decomposition experiment..............................14 - 16
09/26 Exercise 3 Dilution plating..................................................................17 - 23
00/03 Exercise 4 Microscopy ........................................................................24 - 37
Observe bacteria ..........................................................28 - 32
Start Riddell mounts ....................................................33
10/10 Exercise 5 Direct counts ......................................................................38 - 39
Observe fungi...............................................................33 - 37
10/17 Exercise 6 Initiate rhizobium experiment............................................40 - 43
10/24 Exercise 7 Initiate algae experiment....................................................44 - 49
Phosphatase assay ........................................................47 - 49
10/31 Exercise 8 Initiate respiration experiment...........................................50 - 56
11/07 Collect WK1 respiration data
11/14 Collect WK2 respiration data
11/21 Exercise 9 DNA analyses ...................................................................57 - 65
11/28 Collect mycorrhizal data
12/05 Exercise 10 Soil rhizosphere ................................................................66 - 67
12/07 End Classes
LABORATORY REPORTS
Report Grade Date Title Comment
1 15% 10/11 Enumeration of
bacteria & fungi
2 15% 11/21 Quantification of
microbial activity
Critical discussion of methods
employed Microbial observations
Soil Respiration/Biomass and
Phosphatase activity

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INTRODUCTION
"I hear and I forget, I see and I remember, I do and I understand"
Kung Fu-Tse
These laboratory exercises are designed to provide you with (i) exposure to selected
techniques in soil microbial ecology and (ii) experience in the quantification and
reporting of experimental data. We hope this "hands on" experience will give you greater
understanding of concepts in soil microbiology. Mere compliance with the instructions
for each exercise will not assure a good grade in the course. For each exercise, ask
yourself or the instructor: "Why is this done," "why does this happen," "what does it
mean"? Some questions are posed at the end of various exercises - but do not limit your
inquiry to these.
The laboratory is scheduled for Mondays, periods 8-9, in room 3096 McCarty Hall B.
Attendance in the laboratory is required. It is difficult, if not impossible, to make up
missed exercises since the materials will not be available after completion of an exercise.
On occasion, you may be required to come to the lab area briefly at times outside of the
scheduled laboratory period in order to carry out some manipulation that cannot wait
until the following week.
The laboratory portion of the course will count for approximately 30% of your grade.
Your grade will be based on two laboratory reports. The reports should be written in the
style of a scientific paper and should include:
Title: concise phrase describing the report, important for keyword searches
Introduction: providing pertinent background information
The introduction should end with a clear statement of objectives
Materials and Methods: only include changes from the procedure presented in the
lab manual
Results: best presented in tables and figures with supporting text
Discussion: what do the data mean in the greater context of soil microbial ecology;
what went wrong?
Literature cited: provide a few key references of supporting information.
In general, we will consider each student’s work as a replicate for each experiment.
Therefore, reports should present class means along with some statistical indication of
experimental error (for example, ANOVA or standard error of the mean). Obviously,
having several people involved in implementing an experiment will lead to greater
experimental error, but this approach will give you valuable experience in summarizing
experimental data. The reports should be approximately 10 double-spaced typed pages.
Laboratory notes and raw data should be kept in a research notebook (you may use the
back sides of pages in this manual for that purpose). Note that not all exercises will be
written up in a formal report; however, they should all be documented in your laboratory

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notebook. The instructor may ask to see these from time to time (without advance
notice) to check on your laboratory progress.
LABORATORY OPERATIONS
1. SAFETY
The prime rule in the microbiological laboratory is cleanliness! Your cooperation in
keeping the laboratory clean is requested, for reasons that should be obvious.
ORGANIZATION is closely allied to this rule. In fact, safe and aseptic operation in a
laboratory is nearly impossible unless individual activities are carefully ordered. The
following rules should be followed:
You may want to wear a laboratory coat or apron. This is not required, but it may
save an expensive item of clothing from stains, acids, or alkali.
Organize your laboratory work. The first step is to carefully read the instructions
before class. The instructor will outline the order of the day’s work. Pay close
attention during this portion of the period - it is, in many respects, the most
important.
Wipe the working area of your bench with disinfectant before and after the period’s
operations. The tops of the desks should be kept clean, neat, and free from wearing
apparel and non-essential books.
Wash your hands before leaving the laboratory.
Call the instructor immediately if you have an accident, especially if it involves
spillage of a culture, a cut or a burn.
Shoes must be worn in the laboratory at all times.
Long hair should be tied back to reduce the danger of ignition by the burner.
Do not eat or drink in the laboratory.
Wear safety glasses when working with hazardous materials such as acids and
bases.

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2. HANDLING BACTERIAL CULTURES
In this course, we will not be handling pathogenic bacteria, and with the bacteria of the
soil and air we can safely follow certain procedures that economize both time and effort,
but which are absolutely unacceptable when dealing with pathogens. When dealing with
nonpathogens, the primary aim of the technique is to avoid contamination by other
organisms.
The inoculating needle or loop must be heated to redness immediately before and
after use to destroy any bacteria on its surface. Never allowed the loop to touch
anything other than the place to which bacteria are being transferred until it again has
been heated to redness in the flame. Never lay the loop down without first flaming.
Large particles or drops of liquid should not be burnt off in the flame, but rather
should be gently shaken off in the culture tube before the wire is removed.
Cotton plugs should be handled only by the upper portion that projects out of the tube
and must never be placed on the table, but rather held between the fingers while the
tube or tubes are open, and then must be immediately replaced. Gentle rotation of the
plug while the tube is gripped firmly will allow easy removal of sticking plugs and
insertion of slightly oversized plugs. Plastic or metal caps are more readily removed
and replaced, but provide poorer protection against contamination.
After removing the plug, the mouth of the tube should be slowly rotated through the
flame to burn off wisps of cotton, to kill contaminating bacteria on the outer surface,
and to kill any bacteria from the culture that may have been carried by the wire to the
upper portion of the tube. This should be done before and after inoculating or
removing bacteria from a culture tube. Do not overheat the tube.
Any and all accidents, such as dropping a tube or plate or spilling a culture, must be
reported immediately to the instructor. Spilled cultures or broken tubes should be
immediately covered with germicidal solution and paper towels, and allowed to stand
for some time before any attempt is made to clean up the broken pieces or the spilled
material.
NO MOUTH PIPETTING!
All materials such as Petri plates, tubes, pipettes, etc. should be placed in the
receptacles provided for this purpose. Pipettes should never be placed on the desk.
Immerse them tip down in disinfectant solution. Discarded glass materials should be
returned to the appropriate place after all markings have been removed to facilitate
washing and recycling.

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3. AUTOCLAVING
Autoclaving (steam under pressure) is commonly used to sterilize media because it is
usually effective, cheap, and requires relatively short times. Its disadvantage is “heat
damage” which may or may not be tolerated depending on severity and satisfactory
alternative methods. Autoclaves equipped with a jacket shorten time in the autoclave
(they bring temperature up more quickly) and they reduce the amount of moisture left on
the material.
Complete air exhaustion is necessary to obtain sufficiently high temperatures for efficient
sterilization. Pressure does not indicate whether pure steam or a mixture of air and steam
is present. Therefore, consult the thermometer at the bottom exhaust line and not the
pressure gauge to determine if sterilization is occurring. Temperature and time required
for adequate sterilization of liquids are given below. With experience, and if heat
sensitivity of materials demand it, time and temperature may be reduced. The
recommendations given do not apply to materials that are hard to heat (oil, soil, seed, and
powders). These materials should be sterilized in small amounts and for longer periods.
Oil is often sterilized by dry heat and, if heat sensitive, by filter sterilization.
RECOMMENDED TIME IN AN AUTOCLAVE FOR STERILIZATION OF
VARIOUS VOLUMES
ITEMS
ERLENMEYERS
TEST TUBES
1L 20-25
500 ml 17-22
200 ml 12-14
125 ml 12-15
18 x 150 mm 12-14
38 x 200 mm 15-20
TEMPERATURE
In nutritional studies, possible adverse or beneficial effects of autoclaving should be
particularly recognized. For example, the pH is usually lowered 0.2-0.4 unit, and rarely
as much as 1 unit. Glucose may be inhibitory when autoclaved with amino acids.
Xylose forms toxic furfural. Agar is hydrolyzed when autoclaved too long, thus making
a mushy and sometimes inhibitory medium. Sucrose is partially hydrolyzed to readily
available glucose. Autoclaving of glucose with phosphate may be stimulatory. There
may be some stimulatory effects of autoclaving sucrose, glucose, and coconut milk in the
medium that is thought to be related to a chelating effect. The pH and the presence of
certain other substances influence the destruction of thiamine. An aqueous solution of
pH 3 to 5 is best. Separate autoclaving or filter-, gas- or liquid-sterilization of
carbohydrates, growth substances, and antibiotics is often advisable.

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E X E R C I S E
1
SOIL MICROBIAL ASSOCIATION
1. MYCORRHIZAE
OBJECTIVE:
To assess the mycorrhizal status of field area, plant species, or individual plants, to
estimate mycorrhizal inoculum potential, to isolate mycorrhizal spores, and to initiate
mycorrhizal pot culture
INTRODUCTION
The roots of most plant species form symbiotic associations with soil fungi. These
mycorrhizal (fungus/root) associations improve plant growth by effectively increasing
the absorptive surface of the root. Mycorrhizal plants generally have greater access to
poorly mobile nutrients (e.g. P, Cu, Zn) in soils with low fertility than do nonmycorrhizal
plants.
The arbuscular mycorrhizae (AM) or have the widest host range and distribution of all
mycorrhizal types. Hosts include many economically important plant families such as
the Rosaceae, Gramineae, and Leguminosae. The AM do not alter the gross morphology
of the host root and are, in fact, not recognizable without staining. Within the root, the
AM fungi form branch structures termed arbuscules and ovoid to globose thin-walled
structures termed vesicles. Nutrient exchange occurs at the interface of the arbuscule and
plant cell membrane. The vesicles are lipid-filled and are presumed to be for storage.
The fungi that form AM are classified in the order Glomales. There are six genera that
include more than 150 described species (see Fig). Spores produced by Gigaspora,
Scutellospora, Acaulospora, and Entrophospora are called azygospores, since they
resemble zygospores produced by Endogone. Spores produced by Glomus and
Sclerocystis are termed chlamydospores (thick-walled, asexual resting cells).
In this exercise, you will learn to identify and manipulate AM fungi.

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Glomaceae Acaulosporaceae Gigasporaceae
Glomus Entrophospora Acaulospora Gigaspora Scutellospora
Archaeosporaceae
Archaeospora
Paraglomaceae
● vesicles
auxiliary cells ●
● hyphae ●
● arbuscules ●
Al-Agely and Ogram © 2004
METHOD
1. SPORE ISOLATION - procedure will be demonstrated at takedown of experiment
described below. For 1st exercise, spores will be provided in Petri dishes and prepared
slides for observation at higher magnification.
Spores of VA mycorrhizal fungi are separated from soil by decanting and wet-sieving,
followed by sucrose centrifugation. Place 100 g of soil in a one-liter beaker and add tap
water. Stir vigorously, allowing the sand to settle for 10-15 sec, and pour supernatant
through a No. 40 (425 µm) and No. 325 (45 µm) sieve. Roots will be on the coarse sieve.
Spores will be on the fine sieve. Wash spores into 50 ml centrifuge tubes and fill to
within 2 to 3 cm of the top with tap water. The sieving should not be deeper than 1/2
inch thick at the bottom of the tube. If so, use two tubes for the sample. Remember to
balance the centrifuge. Spin at top speed for 3 min and allow stopping spinning by itself.
Gently remove tube and, in one smooth motion, pour off the water, taking care not to
pour off any sieving. You may need to remove floating organic matter that will adhere to
the upper walls of the tube with your finger.
Next, fill the tube up to within 2 to 3 cm of the top with cold 40% sucrose solution and
centrifuge at top speed for 1-1.5 minutes. Manually stop the centrifuge, or use the brake
to minimize the time the spores are in the sucrose solution due to potential damage by
osmosis. Pour the supernatant through the No. 325 sieve to collect the spores, rinse

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gently with water, and put spores in a Petri dish in water. Observe spores with the
dissecting microscope and identify to the level of genus. Also observe prepared slides of
spores under the compound microscopes set up for this purpose. There is also an
optional slide set that can be observed in my office if you have special interest in this
topic.
2. ROOT COLONIZATION: In the first lab period, you will be given cleared and
stained root material to work with. However, at takedown of experiment described below
you will prepare your own roots using the following procedure.
Put roots from the coarse sieve (see above) into tissue cassette and place in a beaker with
10% KOH. Heat to 90 o C for 10-15 min; do not boil roots. Rinse KOH from roots with 5
changes of water. Acidify with 0.1 N HCl for 5 min., and drain HCl from roots.
Cover the samples with Trypan blue-lactic acid (400 ml glycerol, 200 ml lactic acid, 400
ml H 2 O, and 600 mg Trypan blue) and store in a covered container. Heat to 90 o C for 10-
15 min and do not boil roots. Rinse the samples 1-2 times to completely remove all
excess stain, or until the water is clear. The samples can now be stored in water in a
refrigerator for 2-3 weeks or in double plastic bags.
To determine root colonization, prepare a 1/2 inch grid on the outside of the bottom half
of a plastic Petri dish. Remove stained roots from capsule and place in grid Petri dish
with a small amount of water. Pour in just enough water to cover the bottom of the dish
to facilitate uniplanar reading of the roots at 2X magnification of the dissecting
microscope. Carefully spread the roots randomly across the plate until you have
effectively covered the whole surface area evenly with roots.
With the microscope set at 2X magnification and the light source directed from the
bottom via the mirror, arrange the Petri dish in such a way that all of the vertical gridlines
can be followed one at a time by sliding the dish with your hand in an up and down
motion. With the two button counter operated by the other hand, record every root
segment that intersects a vertical gridline by pressing one button. If the segment at the
gridline is colonized by AM fungi, press both buttons. The final number multiplied by
two will give you centimeters of root. The % colonization is determined by dividing the
number of root intersections of the grid colonized by AM by the total number of root
intersections. Length is based on the following equation:
L = (pi * A * n) / 2 H; where
A = total area, H = total length of lines, n = # intersections.
Note: with 1/2 inch grid, 1 intersection is equal to 1 cm of root length. Determine the %
colonization of the root sample provided. From prepared slides learn to recognize
arbuscules and vesicles.

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3. POT CULTURE
AM mycorrhizal fungi are obligate symbionts and cannot yet be grown in pure culture.
Therefore, these fungi are maintained on plant roots in pot culture. Spores of a Glomus
sp. and Gigaspora sp. will be provided. Establish 2 greenhouse cultures with each
fungus and 2 controls (noninoculated) on a susceptible host such as maize. After 8 to 12
weeks document the results (% colonization, structures in root, sporulation, growth
compared to the control)
POT CULTURE METHOD:
Add soil to within 10 cm of the top of 6 growth tubes. Place 10 g of the appropriate soil
inoculum or sterilized inoculum on the soil surface and cover with additional soil. Seed
tubes with either bahiagrass or alfalfa. Water plants as needed and fertilize every other
week with a 20-0-20 soluble fertilizer.
QUESTIONS:
1. What benefits do AM mycorrhizal fungi provide plants? Are there any
disadvantages?
2. How does colonization by Glomus sp. differ from Gigaspora sp.?
3. What are some of the pit falls in the pot-culture method?
REFERENCES:
Smith, SE. and D.J. Read. 1997. Mycorrhizal symbiosis. 2nd ed. Academic Press
Schenck, N.C. (ed.). 1982. Methods and principles of mycorrhizal research. Amer.
Phytopath Soc., St. Paul, MN. (Chapters 1, 3, 4, 5 and 6).
Sylvia, D.M. 1994. Vesicular-arbuscular mycorrhizal fungi. p. 351-378. In R.W. Weaver
and et al. (ed.) Methods of soil analysis, Part 2. Microbiological and biochemical
properties. Soil Science Society of America, Madison, WI.

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COUNTING SOIL MICROORGANISMS
Determination of microorganism cells number or concentration in any soil plays
numerous and important roles in microbial characterization and ecological
experimentation.
METHODS OF CELL QUANTIFICATION:
A. DIRECT COUNT CELL NUMBERS:
The method yields a precise statistical measurement of cell quantification. The basis of
direct count is the actual counting of every organism (or every living organism) present
in a subsample of a population. Direct count can be a chafed by viable count or total
count.
1. VIABLE COUNT
It is the method of using dilution plates or most probable number (MPN) to count living
cells only.
a. DILUTION PLATES: Sold media in Petri dishes use to determine the count of
viable microorganisms. Dilution plate technique assumes that every colony is founded
by a single cell (Colony Forming Unit). That cell must have been alive in order to grow
and form a colony. Problems of this technique are lengthy incubation time, cell
clumping, large cell number often requires many serial dilutions, and few cells number
requires concentration by either centrifugation or filtration. Typically one can use a
minimum number which is 10-fold smaller than the maximum number, but greater than
30 and less than 300.
b. MOST PROBABLE NUMBER (MPN): Broth media in tubes use to
determine approximate viable count of culture in serial dilution. The culture has been
diluted to the point that the broth tubes were inoculated with on the order of only a single
microorganism (turbid) or fewer (not turbid). The concentration of the culture is then
taken to be equal to the amount of dilution necessary to have reached this point. MPN is
especially useful in situations where there is an advantage of using broth over solid
medium. For example, many organisms are not good at forming colonies, such as highly
motile organisms.
2. TOTAL COUNT
All cells are counted, whether dead or alive in a method. Generally, total count requires
the employment of microscopic Techniques. Direct microscopic count is a determination
of the number of microorganisms found within a demarcated region of a slide known to

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hold a certain volume of soil. This total count method of cell quantification is very rapid
but has the problem of requiring high cell concentrations (e.g. 10 7 / ml), and may include
dead and living cells with equal probability.
B. INDIRECT COUNT CELL NUMBERS (ESTIMATION):
Cell biological activity can be used to estimate cell numbers. Such method is often
preferable either for convenience or because direct counting is difficult or even
impossible in many situations (for example, when quantifying filamentous organisms).
Estimates of cell number include determinations of turbidity, metabolic activity, or dry
mass.
1. TURBIDITY
The cloudiness or turbidity of a culture is caused by the individual cells scattering light.
Degree of turbidity is a direct correlation of cell mass. The average cell mass of
individual cells in a culture must first be determined if the turbidity of a culture is to be
used to estimate cell count. This standardization works best for cultures in the
exponential growth phase.
2. METABOLIC ACTIVITY
The metabolic output or input of a culture may be used to estimate viable count. This
method has the same qualification as turbidity methods. The rate at which metabolic
products such as gases and/or acids are formed by culture reflects the mass of bacteria
present. The rate at which a substrate such as glucose or oxygen is used up also reflects
cell mass. Soil enzymes, soil DNA, and soil respiration are some of the widely used
techniques in estimating soil activities.
3. DRY MASS
Determinations of dry mass required to dry cultures before analyses. It is also required to
separate cells from medium by some physical means such as filtration or centrifugation.
The cells are then dried, and the resulting mass is weighed. For filamentous organisms
such as fungi, the method works sufficiently well compared to other methods listed that
dry mass determination is frequently employed.

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E X E R C I S E
2
SOIL DECOMPOSITION AND MICROBIAL COMMUNITY
STRUCTURE (WINOGRADSKY TECHNIQUE)
OBJECTIVES:
Study the impact of eutrophication and plant nutrients on soil decomposition and
microbial community construction in forest and agriculture environments using
Winogradsky technique.
INTRODUCTION
Soil decomposition is a necessary and important process in every ecosystem. It is the
process of recycling nutrients that all organism need for survival. Bacteria and fungi are
the main component of this process that involves the breaking down of detritus and dead
organic materials. Humus (decomposed material) is the final step in this process that
supply soil with nutrient such as calcium, phosphate, potassium, and other ions.
In any given condition, natural environment teems with microorganisms to provide a
specific combination of nutrients and oxygen that allows only certain microorganisms to
survive. We can use our own designed composed media and inoculum from Lake Alice
pond water to create and study a mini-ecosystem, called a Winogradsky column, in the
laboratory. This experiment will be a demonstration.

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METHOD
This will be a demonstration.
1. Obtain a clear container and label your initial, date, and soil type.
2. Place about 200 ml compost soils (Agriculture or Forest soil) to a mixing
container.
3. Add pond water and stir until it is about the consistency of applesauce.
4. Add 5-g grass cutting for the Agriculture soil and 5-g pine needle cutting for the
Forest soil as a carbon source (carbon source in the form of cellulose).
5. Add equal amount of calcium carbonate and calcium sulfate and mix until the
mixture become drier (source of carbon and sulfur).
6. Pour or spoon this mixture into the Decomposition Chamber to approximately 3-4
cm in depth.
7. Mix it well with a spoon or stirring rod to remove any air pockets.
8. Add plain Agriculture or Forest soil to the respective chamber until the depth of
the soil mixture reaches between 6 and 8 cm. DO NOT STIR!
9. Add pond water, leaving about 4 cm of headspace.
10. Insert a thermometer into the soil and record the starting temperature___ °C
11. The soil column in each Decomposition Chamber should be covered with
aluminum foil, seal with parafilm, and placed next to window.
12. Create data sheet for weekly observations and discussion of the developed soil
layers.

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Al-Agely and Ogram © 2004
QUESTIONS:
Why did grass and pine needle cuttings provide?
What did the calcium sulfate provide?
REFERENCES:
Procedure with modification from
http://www.sumanasinc.com/webcontent/anisamples/microbiology/microbiology.htm

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E X E R C I S E
3
SOIL MICROBIAL ENUMERATION:
1. DILUTION PLATE METHOD
OBJECTIVE
To analyze frequency and density and to compare similarity of microbial community for
soil comparative studies
INTRODUCTION
Soils generally contain an enormous and extremely diverse population of
microorganisms. Prokaryotes are the most abundant of these organisms. They consist of
only a single cell that lacks a distinct nuclear membrane and has a cell wall of a unique
composition. Prokaryotes are divided into two domains: Archaea (which include
methanogens and extremophiles; e.g. extreme halophiles, and extreme thermophiles); and
Bacteria, which include the vast majority of prokaryotes.

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Capsule
(Not always present)
Fimbriae
(Not always present)
Slime layer
(Not always present)
Ribosomes
Nuclear Region
Storage Granules
Wall
Membrane
Flagellum
Al-Agely and Ogram © 2004
Bacteria can be characterized based on their reaction with Gram stain or on the basis of
their shape and their metabolic requirements. The soil bacterial population is dominated
by species of Pseudomonas, Arthrobacter, Clostridium, Acinetobacter, Bacillus,
Micrococcus, Acidobacteria and Flavobacterium.
There are several methods for estimating the bacterial population in soils. These include
direct counts using the microscope with various special staining techniques, a statistical
approach of enumeration called the most probable number technique (see later exercises),
and any of several plating techniques employing a wide variety of culture media.
For enumerating the general soil heterotrophic bacterial population, the soil
microbiologist is usually content to use the dilution plating method, realizing full well
that it is not without limitations. The method measures only a small portion of the total
population. Nonetheless, it is quite useful for studying changes in the population that
grows on the chosen medium.
The same methods can be used to study actinomycete populations with appropriate
culture media for their growth requirements. The actinomycetes represent somewhat of a
transitional group between the bacteria and the fungi. They range from unicellular to
highly branched filamentous forms resembling fungi, but much smaller. The
actinomycetes share close affinities with the bacteria. Most notably, they are
prokaryotes. The actinomycetes are the source of many of our powerful antibiotics and
they have an earthy odor, similar to that of newly plowed soil, due to aromatic
compounds (geosmins) which they produce. The species most readily recognized on
dilution plates are members of the genus Streptomyces that form long chains of spores.

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Fungi constitute another extremely diverse group of soil microorganisms. The fungi are
classified into major groups primarily on the basis of their mechanism of sexual
reproduction. Thus the Zygomycetes reproduce sexually by the formation of zygospores,
the Ascomycetes (the sac fungi) through ascospores, the Basidiomycetes (mushrooms &
toadstools) by basidiospore formation, and the Fungi Imperfecti or Deuteromycetes
which lack a sexual stage or it has yet to be discovered. In addition to these sexual
spores, all of the groups produce at least one or more type(s) of asexual spore(s), and
these are generally the dominant mode for propagation. To make matters even more
confusing, many fungi can propagate simply through hyphal fragmentation.
The enumeration of soil fungi can give rise to quite misleading results if proper
constraints are not used in interpreting results. Much of our knowledge of soil fungi has
been derived through the use of dilution plating methods. These methods favor the
isolation of those organisms which grow most rapidly and sporulate profusely; thus, most
of the fungi observed are members of the Fungi Imperfecti. Notable examples in this
category are the genera Penicillium and Aspergillus. Slow growing fungi, which do not
sporulate readily, are often completely absent from dilution plates. Thus, rarely are
basidiomycetes observed by this method.
Estimations of fungal populations by dilution plate methods give much higher results
than estimation by direct methods. This is due primarily to the fact that each single spore
can give rise to a fungal colony on a dilution plate. So too, can fragments from
vegetative hyphae which may actually have been disrupted during the dilution procedure.
For these reasons, caution in interpretation is necessary. Despite its limitations, the
dilution plate method remains a convenient means for study of the distribution and
abundance of soil fungi.
Because bacteria grow much more rapidly, it is necessary to restrict their growth to allow
the fungi to grow. This can be accomplished in a number of ways. Two methods
frequently used are to (I) lower the pH of the culture medium since most bacteria do not
grow well at acid pH and (II) add antibacterial compounds such as Rose Bengal and
streptomycin or other antibiotics. Application of these procedures leads to the
formulation of media, which are selective for soil fungi.

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MATERIALS:
Note: Check expiration date on all media ingredients and submit sub samples for soil
moisture and chemistry analyses
Bacteria Media:
1. Tryptic-soy broth (3 g l -1 ) - note: this 1/10 of label concentration
2. Agar (12 g l -1 )
3. *Cycloheximide (25 mg l -1 ) - use water suspension.
4. *Antifoam agent
Fungi Media:
1. Potato dextrose agar (39 g l -1 )
2. Tergitol NP-10 (1 ml l -1 )
3. *Streptomycin (100 mg l -1 ) - use water suspension.
4. *Chlorotetracycline - HCl (50 mg l -1 ) Sterile Dilution Blanks (H 2 0), 90 and 9 ml.
* = Add after autoclaving
Sterile Petri plates and 1 ml pipettes
Funnels for placing soil in first blank
Vortex
Test tube racks
METHOD:
1. Half the students will process forest soil and the other half will process
agriculture soil.
2. Each student’s dilution series will serve as a replicate.
3. Before beginning the dilution series: (i) wipe down the workbench with Thymol
(a disinfectant); and (ii) label all tubes and dishes and line them up in order.
4. Prepare a dilution series of the soil sample from 10 -1 to 10 -8 (see figure on
following page). The 10 -1 dilution is prepared by weighing out 10 g of soil (*dry
weight basis) in a 90 ml H 2 0 dilution blank and shaking it vigorously for 1 min
(for a soil of average bulk density, this ratio of soil to solution dilutes the
microbial population by one-tenth).
5. From the 10 -1 dilution, transfer 1.0 ml of the solution to a 9.0 ml blank using a
sterile pipette and mix thoroughly (use the vortex), producing a 10 -2 dilution.

Page 21
6. Continue this procedure on out to the 10 -8 dilution using a new pipette for each
transfer; however, to conserve pipettes, when you reach the 10 -2 dilution use the
pipette that you transfer the sample with to also deliver 1 ml samples to each of
the labeled sterile Petri plates. Taking 1 ml from a dilution tube to a plate
represents an additional 10 -1 dilution, but this factor is canceled by the fact that 10
g of soil were added to the original dilution.
7. Each student will have 10 plates: 5 will receive fungi medium (1 each at 10 -2 , 10 -
3 , 10 -4 , 10 -5, and control from sterile water blank) and 5 will receive bacteria
medium (1 each at 10 -5 , 10 -6 10 -7 , 10 -8 and control).
8. When all samples have been distributed among the plates, carefully pour
approximately 10-15 ml of warm (50°C) melted medium into each plate. Pour
just enough to cover the plate - do not fill the plate! All media are kept in a hot
water bath until ready for use.
9. Gently swirl the plates clockwise and then counterclockwise to distribute the
inoculum. When the medium in each plate has solidified, invert the plates and
place them in the incubator (28°C).
10. Count the colonies after 3 days (either before or after class on Thursday) and
again after 1 wk.
11. Choose the dilutions, which yield between 30 and 300 colonies per plate and
make total counts on both media. The instructor will assist you in differentiating
bacteria cultures. Refer to the figure on next page for an example of how to arrive
at the final count of bacteria or fungi per gram of dry soil. Note the
characteristics of the colonies i.e. pigments, texture, and form. Compare your
results to the overall class data.
SOIL WATER CONTENT (PERCENTAGE MOISTURE) ESTIMATION
1. Weigh fresh subsample soil in aluminum containers
2. Oven-dry them (105°C for 72 hours)
3. Reweigh them
4. Water content calculation, use the formula:
Total wet weight - total dry weight
% water content = ⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯ X 100
Total wet weight - weight of container

Page 22
CALCULATE NUMBER OF COLONY FORMING UNITS (CFU) PER GRAM OF
DRY SOIL BY USE THE FORMULA:
A*D*100
N = -----------------
100 – X
WHERE:
N = number of organisms per gram of dry soil
A = average number of count on plates having between 30-300 colonies
D = Dilution factor
X = soil moisture percent, wet weight basis.
10 -1 10 -2 10 -5 10 -8 H 2 O
10
g
1 ml
90
ml
1 ml
10 -2 10 -8 9 ml
Fungi
H 2 O
10 -6
10 -5
Bacteria
Al-Agely and Ogram © 2004

Page 23
QUESTIONS:
1. Discuss the usefulness and limitations of the dilution plate method for
determining the abundance of bacteria, actinomycetes, and fungi in soil.
2. Why does one incubate culture plates in the inverted position?
3. In what respects are actinomycetes closely related to bacteria and fungi? How
can you distinguish bacteria colony from fungi?
4. How can you distinguish bacterial colonies from fungal colonies?
5. What factors make the culture media used in this exercise selective for fungi?
REFERENCES:
Casida, L.E. 1968. Methods for the isolation and estimation of activity of soil bacteria.
IN. Gray & Parkinson, ed. Ecology of Soil Bacteria.
Johnson, L.F. and E. A. Curl. 1972. Methods for research on the ecology of soil-borne
plant pathogens. Burgess Publ Company.
Zuberer, D.A. 1994. Recovery and enumeration of viable bacteria. Pp. 119-144 In:
Weaver, R.W. et al. (ed). 1994. Methods of Soil Analysis. Part 2. Microbiological and
biochemical properties. SSSA. Madison, WI.

Page 24
E X E R C I S E 4
MICROSCOPY:
EYE
Eye Piece
First Real
Objective piece
Objec
FINAL VIRTUAL
Al-Agely and Ogram © 2004

Page 25
OBJECTIVE:
Train to use safely the basic microscopes (dissecting and compound-oil immersion) and
to observe various soil microorganisms
INTRODUCTION
Microscopes are the most important tools of any microbiology studies. They are needed
to observe structures of microorganisms that are too small to be seen by naked eye.
Magnification and resolution are the two major steps to achieve quality observation.
Most are familiar with magnification, but not all are familiar with resolution. Optic
technology in the area of electron microscope allows the achievement of millions of
times magnification of small object, but these same principles limit magnification to
about 1000 of times light microscope. Why the two systems are so different?
Resolution is the answer to this question. The eye is the ultimate receptor of any image.
Eye resolution is determined by the distance between receptor cells in the retina. Pictures
of two objects that are received on the same receptor simultaneously cannot be
distinguished from each other. However, if these two pictures are received on adjacent
receptors, they may be resolved. The optics of the eye and the spacing of receptors on
the retina make the optimum focusing distance for any eye. Regular eye can distinguish
25-cm lines as separate lines if they are 60 to 100 µm apart. Lines less 60 µm apart are
unresolved and will appear as a single solid line. Eye resolution is an individual
phenomenon and it is different from one eye to another of the same person, and from one
person to another. For practical applications, 100 µm will be considered the resolving
power of human been eye.
Microscope magnification is required to observe an object that is smaller than 100 µm or
to distinguish between two structures that are closer together than 100 µm. Microscope
is also equipped to resolve mix light rays. Achieving this mainly depends on the ability
of glass to bend light, its refractive index, and the wavelength of light used to form the
image. These factors are quantified by this equation R = 0.61λ / NA (where: R = the
resolution in µm, λ = the wavelength of light in µm, NA = the numerical aperture of the
objective lens that includes the refractive index of the lens (n) and the sine of the lens
angle (U): NA = n * sine U).
Example:
Using blue light, λ = 45 µm
The 45X lens on your microscope (color coded yellow) NA = 0.66
R = (0.61 x 0.45) / 0.66 = 0.42 µm resolution
The smallest object that can be observed in this lens is 0.42 µm. Microscope
magnification is the product of the ocular lens (10x) and the objective lens (45x) or 450

Page 26
times together. In the above example the 0.42 µm object will be magnified 450 times and
will be appeared as 450 x 0.42 = 190 µm in size.
It is important to know that lens resolution is limited by the refractive index of air space
between the objective lens and the cover glass. The materials commonly found in this
region and their refractive indices (RI) are listed below:
SUBJECT
RI
Air 1.00
Water 1.33
Glass 1.50
Air space that is filled with liquid of higher refractive index will have a large effect on
NA. Some specific lens is designed to focus with a specific medium (water immersion or
oil immersion) in contact with the lens surface. The degree to which light is bent
depends on the ratio between the two materials. Compound microscopes in use for this
laboratory are equipped with an oil immersion lens.
Achieving best resolution depends mainly on proper microscope setting, proper
adjustment of stage condenser, and proper use of oil immersion with the right lens. Clean
lenses and thin specimens are also required to achieve good resolution.

Page 27
MICROSCOPE OPERATION
Microscope is an expensive and delicate instrument. Replacement cost is about $1800.
Treat it with care! Dropping or jarring it will cause considerable damage.
USE TWO HANDS TO CARRY the microscope as to grab the microscope arm with
one hand and support the microscope base with the other hand.
Gently remove the microscope from its cabinet. Plug the microscope cord into the
electrical outlet on your table. Adjust light setting to give reasonable light
illumination. Higher settings shorten the bulb life; bulb will burn out in 30 minutes.
Recent microscopes have a revolving head to change the direction of the oculars lens
unit. Find the proper interpupillary distance by adjusting the oculars with both hands
until a single image is seen.
Locate the coarse and fine focusing knobs and rotate them back and forth gently
while observing the movement of the objective lenses.
Rotate objective lens and notice that there is a positive click into position for each
lens. Look at the markings on each lens, colored rings, NA, and magnification.
Secure slide on stage with slide clips.
Adjust the condenser under the stage to achieve good resolution.
Put the 4x objective into position and adjust the condenser position to see the light
coverage in the field of view.
Put the 10X objective into position and do adjustments for better view
Move the iris diaphragm lever back and forth for the best amount of light passing
through.
Microscope oculars are focused independently of each other. Compound
microscopes in this laboratory have adjustable left ocular and fixed right ocular.
Close your left eye and bring the object on focus with the fine adjustments. Then
close the right eye and focus the left ocular by turning the focusing ring on the left
ocular tube. Recheck the right eye for proper focus. This should a regular practice
before any microscope use. The main cause of eyestrain and fatigues during
microscope work come from improper focus.
When using oil immersion lens use oil of 1.5 RI then gradually move from 4X, 10X,
45X, and finally 100X. Excessive oil will obscure large portions of the slide from
further examination at lower powers.
READING ASSIGNMENT:
Molling, F. K. 1981. Microscopy from the very beginning. Carl Zeiss, Oberkochen,
West Germany. (copies in 2170 McCarty).

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EXAMINING DILUTION PLATES
1. OBSERVE SOIL BACTERIA
Because of the transparency of unstained bacteria, it is very difficult to observe them
without either staining them or using special techniques of microscopy to view them. To
make the bacteria visible through the microscope, they must be stained with dyes that
have an affinity for the bacterial cytoplasm or other cell constituents such as the cell wall.
Many commonly used dyes are positively charged (cationic) molecules and hence
combine strongly with negatively charged cell constituents such as nucleic acids and
acidic polysaccharides. Cationic dyes include methylene blue, crystal violet and safranin.
Other dyes are negatively charged (anionic) molecules and combine with positively
charged cell constituents such as many proteins. Anionic dyes include eosin, acid
fuchsin, and Congo red. Still another group of dyes is called fat soluble. Dyes in this
group combine with fatty materials in the cell and are often utilized to reveal the location
of fat droplets or deposits. A common fat soluble dye is Sudan black.
Methylene blue is a good, simple stain that works well on many bacterial cells and does
not produce such an intense staining that cellular details are obscured. Another attribute
of methylene blue is that it does not combine well with most noncellular materials so that
it is especially useful in examining natural samples for the presence of bacteria.
PREPARATION OF A SMEAR MOUNT
MATERIALS
1. Soil isolates obtained in earlier exercises
2. Microscope slides
3. Inoculating loop
4. Staining solutions: Methylene blue
5. Bibulous paper or Kim wipes

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METHOD
1. Place a small drop of clear water on a clean microscope slide using the
inoculating loop.
2. Flame the inoculating loop, let it cool, and transfer a small portion of a bacterial
colony to the drop of water.
3. Using a circular motion, mix and spread the resulting cell suspension to cover an
area about the size of a dime. Flame the loop again immediately.
4. Allow this smear to air dry and then heat fixes it by passing it briefly through the
flame of the burner several times. Add a few drops of methylene blue to cover the
smear. Use sparingly to avoid making a mess with spillage.
5. After the dye has been on the smear for 4 min, gently rinse it in a stream of water
in the sink.
6. Blot the slide dry using a pad of bibulous paper. Examine the stained smear at the
different magnifications on the microscope. Be sure to use the oil immersion
objective to view the preparation.
PREPARATION OF WET MOUNTS:
It is not always desirable to view only stained preparations of bacteria since no
information is gained as to whether or not the organisms are motile i.e. if they have
flagella (the organelle of locomotion in bacteria: singular = flagellum). To determine
whether an organism is motile, it is usually observed in a wet mount.
METHOD
Place a small drop of water on a microscope slide using a flamed inoculating loop.
Flame the loop, let it cool and add some bacterial from a culture to the drop of water.
Mix but do not spread the drop out. Alternatively, if you have a broth culture, simply
place several loop-full onto the center of the slide to form a drop.
After you have placed the cell suspension on the slide, simply place a cover slip over the
drop to obtain as few air bubbles as possible. This is most easily done by bringing one
edge of the cover slip into contact with the edge of the drop and then laying the cover slip
down on an angle so the fluid flows evenly across the slide.
GRAM STAIN FOR BACTERIA
There are basically two types of bacteria, gram + (purple color) and gram - (no purple
color) when stained with Crystal Violet and Iodine. This is mainly due to the chemical

Page 30
and structural composition of the bacterial cell wall. In this part of the exercise you will
be given 2 different types of bacteria and will determine which types you have.
PROCEDURE
• Place a loop full of bacteria on a slide (mix very well).
• Add a drop of water, spread and allow them to dry.
• Hold the slide with a clothespin.
• Quickly pass the slide through a flame to fix most of the bacteria cells
• Once the slide cool down, flood with crystal violet and let sit for 30 seconds
• Rinse with water
• Flood the slide with iodine and let sit for 1 minute.
• Rinse the slide with 95% alcohol until no purple color drips off the slide.
• Rinse the slide with water.
• Flood the slide with safranin for 1 minute.
• Rinse the slide with water.
• Put cover slide, look at the slide under a compound microscope, and sketch what
you see.
BACTERIA FREQUENTLY ISOLATED FROM SOIL
GRAM-NEGATIVE CHEMOLITHOTROPHS
Nitrobacteraceae
Nitrobacter - short rods, reproduce by budding, yellow pigment, oxidize nitrite to nitrate
and fix CO 2 to fulfill energy and carbon needs, strict aerobes.
Nitrosomonas - ellipsoidal to short rods, obligately chemolithotrophic, oxidize ammonia
to nitrite and fix CO 2 to fulfill energy and carbon needs, strictly aerobic.
Sulfur metabolizing
Thiobacillus - small rod-shaped, energy derived from the oxidation of one or more
reduced sulfur compounds, mostly autotrophic.

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GRAM-NEGATIVE AEROBIC RODS AND COCCI
Pseudomonadaceae
Pseudomonas - straight or curved rods, motile by polar flagella, chemoorganotrophs,
most are strict aerobes (a few species can denitrify), abundant in rhizosphere.
Xanthomonas - straight rods, motile by polar flagellum, growth on agar yellow,
chemoorganotrophs, strict aerobes, mostly plant pathogens.
Azotobacteraceae
Azotobacter - large ovoid to coccoid cells, marked pleomorphism, form thick-walled
cysts and capsular slime, motile with peritrichous flagella or nonmotile, gram-neg. or
variable, fix atmospheric nitrogen.
Beijerinckia - straight to pear-shaped with rounded ends, up to 6 um with occasional
branching, cysts and capsules in some species, produces copious slime in culture,
fixes atmospheric nitrogen, acid tolerant.
Rhizobiaceae
Rhizobium - rods but pleomorphic under adverse conditions, motile by 2 to 6 peritrichous
flagella, nonsporing, copious extracellular slime in culture, chemoorganotrophs,
aerobic to microaerophilic, able to invade root hairs of leguminous plants.
Agrobacterium - rods, motile by 1 to 4 peritrichous flagella, nonsporing, slime
production, chemoorganotrophs, aerobic, most initiate plant hypertrophies.
Uncertain affiliation
Alcaligenes - rods to cocci, motile by up to 4 peritrichous flagella, chemoorganotrophs,
strict aerobes. Saprophytes in animals and soil.
Gram-Negative Facultative Anaerobic Rods
Flavobacterium - coccobacilli to slender rods, motile with peritrichous flagella or
nonmotile, pigmented in culture, chemoorganotrophs, fastidious as to requirement.
Gram-Negative Cocci and Coccobacilli
Acinetobacter - short rods to cocci, large irregular cells and filaments in culture, no
spores or flagella, chemoorganotrophic, strict aerobes, resistant to penicillin.
Gram-Positive Cocci
Micrococci - spherical, non-motile, chemoorganotrophs, aerobic.
Staphylococcus - spherical, chemoorganotrophs, metabolism respiratory or fermentative,
produce extracellular enzymes and toxins, facultative anaerobes.
Streptococcus - spherical to ovoid, chemoorganotrophs, metabolism fermentative,
facultative anaerobes.

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Sarcina - nearly spherical, non-motile, chemoorganotrophs, strictly fermentative
metabolism, strict anaerobes.
Endospore-Forming Rods
Bacillus - rod-shaped, motile, flagella lateral, heat-resistant endospore,
chemoorganotrophs, strict aerobes to facultative anaerobes, gram-positive.
Clostridium - rods, peritrichous flagella, form spherical to ovoid spores, gram-positive,
chemoorganotrophs, most strictly anaerobic.
Budding and/or Appendaged
Hyphomicrobium - rod-shaped with pointed ends, produce mono- or bipolar filamentous
outgrowths, gram stain unknown, multiply by budding at tip, growth in liquid culture
on surface, chemoorganotrophic, aerobic, temp 15-30 C.
Pedomicrobium - spherical to rod-shaped, multi. by budding at tip of cellular extension
producing uniflagellate swarmers, gram-neg., microaerophilic to aerobic,
chemoheterotrophic, mesophilic.
Caulobacter - rod-shaped to vibrioid, typically with stalk extending from one pole, cells
may adhere to each other in rosettes, cell division by asymmetrical fission, gram-neg.,
chemoorganotrophic, aerobic.
Metallogenium - coccoid, attached to surfaces, may form flexible filaments, multiply by
budding, manganese and iron oxides deposited on filaments, heterotrophic.
Coryneform Group
Corynebacterium - irregular shape with club-like swellings, non-motile, gram-positive,
chemoorganotrophs, aerobic and facultatively anaerobic.
Arthrobacter - old culture coccoid, on fresh media swellings from coccoid cells giving
rise to irregular rods, gram-positive, chemoorganotrophs, strict aerobes.
Cellulomonas - irregular rods, motile by one or more flagella, gram-positive,
chemoorganotrophs, decompose cellulose, aerobic.
Mycobacterizceae
Mycobacterium - curved to straight rods, filamentous growth may occur. Acid-fast
reaction, non-motile, lipid content in wall high, aerobic

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2. OBSERVE SOIL FUNGI
Examine selected colonies using the dissecting microscopes and low power objective of
the compound microscope. Look especially for fruiting structures (e.g., spores).
Aseptically remove small portions of fungal and an actinomycete culture from dilution
plates and make ‘squash’ mounts in various stains. What structures do you observe?
How can you distinguish fungi from actinomycetes? What are the benefits and limitations
of dilution plating?
Many soil fungi belong to the order "Hyphomycetes" of the Fungi Imperfecti - that is
they produce conidia (asexual spores) on conidiophores. These structures are very fragile
and require special techniques for observation. The Riddell mount is an excellent
technique to observe these fungi.
RIDDELL MOUNTS
Setup - Place two pieces of filter paper in a 9-cm glass Petri dish. Place a bent glass rod,
slide, and two cover slips in the dish. Note: carefully lean cover slips against glass rods
so they can be easily removed after autoclaving. Autoclave two units for each student
and distilled water to moisten filter paper.
One-cm squares are cut out of an agar medium (see fungus agar on page 20, but increase
to 2% agar) and placed on a sterilized microscope slide. A needlepoint inoculum is
placed on the corners of the agar and covered with a sterile cover slip. The slide is
incubated in a moist chamber on glass rods for 1-2 wk. Observe periodically and once
sporulation has occurred transfer cover slip to a drop of stain on a new slide and observe
conidiophores arrangement.
Try this procedure with at least 2 fungal cultures. Attempt to identify to genus using the
key in Barron.
CLASSIFICATION OF FUNGI
(Adapted from Cavalier-Smith, 1989 and Alexopoulus and Mims, 1983)
KINGDOM PROTISTA (PROTOZOANS)
Phagotrophic, organisms with somatic structures devoid of cell walls, we are including
them here because mycologists traditionally study them. These organisms are the
cellular slime molds and the true slime molds. The cellular slime molds have a
reproductive stalk that consist of walled cells and is simple. The most prevalent form of
the organisms is the myxamoeba that feeds by engulfing bacteria (Alexopoulus and
Mims, 1983). The true slime molds have the plasmodial somatic phase but produce
spores with definite walls from elaborate sporophores.

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KINGDOM STRAMENOPHILA
Eukaryotic organisms with either tubular ciliary mastigonemes or with chloroplast
bounded by an envelope of two membranes and surrounded by two chloroplast
endoplasmic reticulum membranes and a periplastidal space containing the periplastidal
reticulum; mitochondrial cristae are rounded tubules or flattened finger-like projections.
Pseudofungal organisms typically produce flagellate cells.
Phylum: Heterokonta
Anterior cilium with tubular retronemes; posterior cilium smooth or absent
Class Hyphochytridiomycetes
A very small group of aquatic fungi with motile anteriorly uniflagellate cells each with a
tinsel flagellum.
Class Oomycetes
Soma varied but usually filamentous, consisting of a coenocytic, walled mycelium;
hyphal wall containing glucans and cellulose, with chitin also present in one order
(Leptomitales); zoospores each bearing one whiplash and one tinsel flagellum; sexual
reproduction oogamous resulting in the formation of oospores. Root parasitic species,
Pythium and Phytophthora, belong to this class.
KINGDOM EUMYCOTA (FUNGI)
Eukaryotic organisms without chloroplasts or phagocytosis, but with saprobic or parasitic
nutrition and typically with chitinous walls and plate-like mitochondrial cristae; develop
from spores; cilium, when present, single posterior without rigid, tubular mastigonemes.
The kingdom is has four phyla: Chytridiomycota, Zygomycota, Ascomycota and
Basidiomycota.
PHYLUM CHYTRIDIOMYCOTA
Zoospores with single posterior whiplash cilium; perfect state spores are oospores or
zygospores; and have a coenocytic thallus of chitinous walls. These fungi are prevalent
in aquatic habitats but many inhabit the soil. Some parasitize and destroy algae and thus
form a link in the food chain.
PHYLUM ZYGOMYCOTA
Sexual reproductions are by the fusion of usually equal gametangia resulting in the
formation of a zygospore. Asexual reproduction is by the aplanospores, yeast cells,

Page 35
arthrospores or chlamydospores. Motile spores are absent. The phylum consists of two
classes, the Trichomycetes (arthropod parasites) and Zygomycetes.
CLASS ZYGOMYCETES:
Mainly terrestrial saprobes or parasites of plants or mammals, or predators of
microscopic animals; if parasitic, mycelium immersed in host tissue; asexual
reproduction by aplanospores borne singly or in groups within sporangial sacs; sexual
reproduction by fusion of usually equal gametangia resulting in the formation of a
zygosporangium containing a zygospore.
PHYLUM ASCOMYCOTA:
Unicellular or more generally with septate mycelium; sexual reproduction by formation
of meiospores (ascospores) in sac-like cells (asci) by free cell formation. Three Subphyla
based on ascus formation.
SUBPHYLUM EUASCOMYCOTINA:
Ascomata and ascogenous hyphae present; thallus mycelial. These classes include most
fungi imperfecti and lichen forming groups. The teleomorphs of Aspergillus,
Penicillium, Thielaviopsis and Fusarium belong to orders in this subphylum.
SUBPHYLUM LABOULBENIOMYCOTINA:
Thallus reduced; ascoma a perithecium. These are exoparasites of athropods and can
survive in the soil as resting structures.
SUBPHYLUM SACCHAROMYCOTINA:
These fungi lack ascogenous hyphae and have a yeast-like thallus or mycelial. They are
the budding yeast and their filamentous relatives.
PHYLUM BASIDIOMYCOTINA:
They are saprobic, symbiotic, or parasitic fungi. Morphologically are unicellular (yeastlike)
or more typically, with a septate mycelium with a vegetative heterokaryophase,
sexual reproduction by producing meiospores (basidiospores) on the surface of various
types of basidia.
CLASSES UREDINIOMYCETES AND USTOMYCETES:
Basidiocarps are lacking and resting spore germination results in formation of
basidiospores. These fungi cause rust and smuts of plants and their resting spores may
survive in the soil for decades.

Page 36
CLASS GELIMYCETES:
Basidia transversely or longitudinally septate (phragmobasidia) produced on various
types of sporophores or directly on the mycelium. These are mainly decomposing fungi
found on litter but Thanatephorus cucumeris (teleomorph of Rhizoctonia solani) also
belongs here.
CLASS HOLOBASIDIOMYCETES:
Basidia non-septate (holobasidia), produced on persistent hymenia on various types of
open sporophores or, rarely, directly on the mycelium; or inside closed sporophores
opening, if at all, after the spores are mature. These are the more commonly seen
mushrooms and wood rot fungi of which many species form ectomycorrhizal associations
with trees.
FORM PHYLUM DEUTEROMYCOTINA:
Generally are called the Imperfecti Fungi. Their main characteristic are: Teleomorph
absent, Saprobic, symbiotic, parasitic, or predatory fungi, unicellular or, more typically,
with a septate mycelium, usually producing conidia from various types of conidiogenous
cells. Sexual reproduction is unknown but a parasexual cycle may operate. A few
species produce no spores of any kind.
FORM CLASS BLASTOMYCETES:
Soma consisting of yeast cells with or without pseudomycelium; true mycelium, if
present, not well developed.
FORM CLASS COELOMYCETES:
True mycelium present; conidia produced in pycnidia or acervuli.
FORM CLASS HYPHOMYCETES:
True mycelium present; conidia produced on special conidiogenous hyphae
(conidiophores) arising in various ways other than in pycnidia or acervuli. A few species
do not produce spores of any kind.
SERIES ALEURIOSPORAE:
Spores develop terminally as blown-out ends of the sporogenus cells and usually thickwalled
and pigmented.

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SERIES ANNELLOSPORAE:
First spore produced terminally with each new spore blown out through the scar left by
the previous. A succession of proliferations is accompanied by increased length of
sporogeneous cells.
SERIES ARTHROSPORAE:
Conidia produced after separation and breaking up of sporogenous hyphae.
SERIES BLASTOSPORAE:
Develop in acropetal succession as blown out ends of conidiophore.
SERIES BOTRYOBLASTOSPORAE:
Conidia produced on well differentiated, swollen sporogenous cells.
SERIES MERISTEM BLASTOSPORAE:
Conidia borne singly at apex in irregular whorls which elongate from the base.
SERIES PHIALOSPORAE:
Sporogenous cells stay constant in length and conidia are abstricted successively in
basipetal succession from an opening.
REFERENCES
Barron, G.L. 1968. The genera of hyphomycetes from soil. Williams &Wilkins. 364
pp. 462.07 B277G
Domsch K.H., W. Gams, and T.H. Anderson. 1980. Compendium of Soil
Fungi.Academic Press, New York.
Hawksworth, D.C. 1983. Dictionary of Fungi. QK603 A5
Riddell, R.S. 1950. Permanent stained mycological preparations obtained by slide
culture. Mycologia 42:265-270
Cramer, West Germany. 3rd edition. 424 pp. QK603.2 A7X
Watanabe, T.1994. Pictorial atlas of soil and seed fungi : morphologies of cultured fungi
and key to species. Lewis Publishers, Boca Raton, FL QR111 .W26713 1994

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E X E R C I S E
5
SOIL MICROBIAL ENUMERATION:
2. DIRECT MICROSCOPIC COUNT
OBJECTIVE
Rapid quantification of microorganism numbers within a demarcated region of a slide
known to have a certain volume of soil by using Fluorescein Isothiocyanate (FITC) for
direct microscopic counting.
METHOD:
1. Prepare staining solution (FITC) as follows:
1.3 ml of 0.5 M Na 2 C0 3 - NaHC0 3 mixed 1:1, pH 9.6
6 ml 0.01 M phosphate buffer pH 7.2 (2.8 ml 0.2 M monobasic sodium
phosphate; 7.2 ml 0.2 M dibasic sodium phosphate; dilute to 20 ml with
distilled water)
5.7 ml 0.85% NaCl
5.3 mg crystalline fluorescein isothiocyanate – Note: this is not a vital stain.
2. Shake staining solution and use immediately or store in dark at 4°C (6 hr only).
3. Prepare smears from soil dilution as follows:
Mix 5 g of soil (agriculture or forest) with 45 ml of 0.1% agar solution and
shake vigorously
After 30 sec transfer 0.01 ml to a slide and spread evenly within a 1 cm 2 area
previously marked on the slide with a marking pen
Place slides on warmer to dry and then heat fix briefly. Note: dilution factor =
0.01/50.
4. Stain 3 min with FITC.
5. Wash slides thoroughly with 0.5 M Na 2 C0 3 - NaHC0 3 buffer.

Page 39
6. Mount smears immediately with glycerol (pH 9.6). Place cover slip over
specimen.
7. Observe with fluorescence microscopy (in Room 2170 McCarty A): Count the
number of bacteria within two microscope fields (normally at least 20 fields) at
400 X and record the mean.
Mean count X No of fields in 1 cm 2 (8264) 1
No bacteria / g = ⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯ X ⎯⎯⎯⎯⎯⎯⎯⎯
Soil dry weight
Dilution factor
QUESTION:
How and why do direct counts differ from dilution plating?
REFERENCES:
Babiuk LA and EA Paul 1970 Can J Microbiol 16:57-62

Page 40

Page 41
E X E R C I S E
6
SOIL MICROBIAL ASSOCIATION
2. RHIZOBIA
OBJECTIVE:
Examine root nodulation, isolate rhizobium bacteria, and inoculate new plants with and
without nitrogen source.
INTRODUCTION
Bacteria of the genera Rhizobium and Bradyrhizobium are capable of inducing the formation
of specialized structures called root nodules on the roots of many leguminous plants.
The legumes can be subdivided into what are known as cross inoculation groups. These
are groups of legumes which can be nodulated by the same rhizobial strain. There are 20
cross-inoculation groups; however, in practice only six of these receive much attention.
The genus Rhizobium is currently divided into species based on the legume crossinoculation
group which the particular strain is able to nodulate. The Rhizobium species
and the cross-inoculation groups which they nodulate are as follows: Rhizobium meliloti,
alfalfa group; R. trifolii, clover group; R. leguminosarum, pea group; R. phaseoli, bean
group, R. lupini, lupine group; B. japonicum, soybean group, and the “cowpea
miscellany,” the cowpea group.
The following exercise introduces several aspects of the rhizobia-legume symbiosis.

Page 42
MATERIALS:
Nodulated leguminous plants
Bean seeds (do not pregerminate seeds)
Rhizobium phaseoli (Nodulate the chosen legume)
Yeast-Extract-Mannitol-Congo Red-Agar (YEM-CRA) media - mannitol 10 g, yeast
extract 0.2 g; NaCl 0.l g; CaCO 3 3 g; Congo Red 2.5 ml of a 1% solution; and Agar
15 g in 1000 ml distilled water.
Modified Fahraeus solution of no nitrogen (-N) - CaCl 2 .H2O, 0.1 g; MgSO 4 .7H 2 O,
0.12 g; KH 2 PO 4, 0.1 g; Na 2 HPO 4 .12 H 2 O, 0.15 g; FeCl 3 .6H 2 O, 0.01 g;
Na 2 MoO4.2H 2 O, 0.002 g; trace elements (Hoagland’s minors), 1 ml. Add all and
bring the volume to one liter with distilled water. Adjust pH to 6.5.
Modified Fahraeus solution with nitrogen (+N) - same as above but with 2.5 g of
Ca(NO 3 ) 2 added. Adjust pH to 6.5
Plastic growth pouches
Ethanol, 95%
20%-Commercial bleach (5% sodium hypochlorite) for seed disinfestations
Sterile water in 13 x 50 mm tubes
Glass rods
Microscopes, slides, cover slips
Sterile Petri plates or 13 x 100 mm culture tubes
Scalpels or razor blades
Loeffler’s Alkaline Methylene Blue: dissolve 0.3 g Methylene Blue in 30 ml of 95%
Ethanol, and add 70 ml of a 0.01% KOH solution.

Page 43
METHOD:
EXAMINATION OF ROOT NODULE BACTERIA
Examine the root nodule bacteria from nodules of fresh plants preferably just removed
from soil. Remove a nodule, surface sterilize it in 20% Clorox for 5 minutes then wash
with several changes of sterile water. Crush the nodule in a small volume (1-2 ml) of
sterile water to achieve a cloudy suspension. Transfer some of this suspension to a small
drop of water on a clean slide and prepare a wet mount. Examine under oil immersion
with a cover slip.
Prepare a smear using the crushed nodule suspension, air dry, gently heat, fix and stain
with Loeffler's alkaline methylene blue (2-3 min). Wash, blot dry and examine using oil
immersion. Note the size, shape and staining characteristics of the bacteroids.
ISOLATION OF RHIZOBIUM FROM ROOT NODULES
Obtain 1 or 2 plates of yeast extract mannitol - Congo red medium. Using the nodule
suspension prepared above, streak the plates of YEM-CR medium to obtain isolated
colonies. Incubate the plates in an inverted position at 28ºC for 7-10 days. Observe the
colonies, which develop. Colonies, which take up much of the Congo red, are generally
not Rhizobium. Observe bacteria from the colonies, which are not highly colored
weekly. Note: let plates dry before sealing with parafilm.
NODULATION OF LEGUMES
Obtain 4 plastic growth pouches. Mark 2 of these "inoculated" and 2 "noninoculated."
Now mark 1 pouch of each "+NO 3 " and 1 pouch of each "-NO 3 ". Make sure you label
pouches with your name. Add 10 ml modified Fahraeus medium without added nitrogen
to each chamber marked "-NO 3 " and 10 ml of modified Fahraeus medium with added
nitrogen to each chamber marked "+NO 3 ".
To each of the pouches, transfer 2-4 surface sterilized bean seed or using sterile forceps.
Carefully place the seeds in the small "cradle" of folded paper at the top of each chamber.
When you have "seeded" all pouches, set aside the pouch marked "noninoculated" and
go on to inoculate the seeds in the pouch marked "inoculated" with the proper Rhizobium
supplied as a suspension of a pure culture. Be sure to inoculate only the pouch marked
"inoculated." It is important to maintain the noninoculated controls as such.
Maintain the growth pouches in the light in a growth chamber. Keep the plants well
supplied with the proper nutrient solution and water only with sterile water and nutrients.
After 1 week, thin to one plant per pouch.

Page 44
Note the rates of growth in all series and watch for differences between the inoculated
and noninoculated plants as well as between the "plus N" and "minus N" treatments.
After several weeks, record the degree of nodulation, vigor, color, and size of the plants.
QUESTIONS:
1.What is leghemoglobin and what is its significance?
2.Discuss the course of events in the formation of root nodules.
3.What effect does nitrate have on the nodulation of legumes?
REFERENCES:
Chapters 8 and 9 in Metting's Soil Microbial Ecology

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E X E R C I S E
7
SOIL ALGAE ENUMERATION
OBJECTIVE:
Quantify algal populations using the most probable number technique and enzymatically
quantify soil metabolic activity by using phosphatase assay.
ALGAE TEST
Since algae are photoautotrophs, they can be grown on selective culture media devoid of
organic carbon. This eliminates competing microbes, which would outgrow the algae
and possibly inhibit their development. One can make these media further selective for
cyanobacteria (blue-green algae) by eliminating all sources of fixed nitrogen (ammonium
or nitrate etc.). Enumeration of soil algae is most frequently accomplished by a dilution
technique involving inoculation of replicate tubes of an appropriate liquid medium with
subsamples from the dilution tubes. This technique is known as the most probable
number technique and represents a statistical probability approach to counting
microorganisms. It is employed when the group of organisms being investigated is not
readily cultured on solid media.
MATERIALS:
Soil samples - forest and agriculture
Modified Bristol’s solution (9 ml per test tube) - (NaNO 3 , 0.25 g; CaCl 2 , 0.025 g;
MgS0 4 .7H 2 O , 0.075 g; K 2 HPO 4 , 0.075 g; KH 2 PO 4 , 0.018 g; NaCl , 0.025 g; FeCl 3 ,
0.01 g; NaMoO 4 .2H 2 O , 0.002 g; and trace minerals* , 1ml combined and bring the
volume with distilled water to 1 L. *Trace element stock solution: MnSO 4 , 2.1 g;
H 3 BO 3 , 2.8 g; Cu (NO 3 ) 2 .3H 2 O , 0.4 g; ZnSO 4 . 7H 2 O , 0.24 g combined and bring
the volume with distilled water to 1 L. Use 1 ml stock solution per liter of medium
Sterile water blanks, 90 and 9 ml
Sterile pipettes, 1 ml
Test tube racks

Page 46
METHOD:
Prepare a dilution series of a soil sample from 10 -1 (1/10) to 10 -5 (1/100,000) as in
earlier exercises (remember that the 10 -1 dilution is prepared by adding 10 g
(approx. 5 cc) in a 95 ml dilution blank). Transfer 1 ml of this suspension to a 9 ml
sterile water blank to obtain a 10 -2 dilution and repeat until you have reached the 10 -5
dilution.
Inoculate 3 tubes of modified Bristol’s solution with 1 ml aliquots from each soil
dilution. Be sure to label tubes.
After inoculating all tubes, incubate them in diffuse light on a windowsill, in a
greenhouse, or in a growth chamber. Examine occasionally and make a final
observation after 30 days.
To determine the numbers of algae in your soil samples, determine the number of
tubes at each dilution, which exhibit noticeable growth. To facilitate this, hold the
tubes against a white background or up to the light and look for green coloration.
Record the number of positive tubes at each dilution and refer to Table. This table
indicates the most probable number (MPN) of algae per gram of soil based on the
number of positive tubes. Locate the series of numbers, which correlated to your
positive tubes at the highest dilutions showing growth. For example, consider the
following results:
Dilution 10 -1 10 -2 10 -3 10 -4 10 -5
Positive Tubes 3 3 2 1 0
The code (series) relating to these results is 3, 2, 1 (10 -2 , 10 -3 , 10 -4 ) or 2, 1, 0 (10 -3 ,
10 -4 , 10 -5 ).
Refer to the table and find these “codes” by reading across the columns. MPN
number for 3, 2, 1 is 1.50 and for 2, 1, 0 is 0.15.
The MPN number associated with the proper code is multiplied by the reciprocal of
the center dilution of the series (series B in this table) to obtain the most probable
number per gram of original soil.
Thus the 3, 2, 1 code yields 1.5 x 10 3 /g and 2, 1, 0 yields 0.15 x 10 -4 or 1.5 x 10 3 /g
also. It should be stressed here that the 3-tube MPN provides minimal accuracy and
is used here primarily to introduce the concept and the technique.
After you have calculated the most probable number of algae in your soil samples,
observe the contents of some tubes (low dilution & high dilution) microscopically
using simple wet mounts. Note the pigmentation and morphology of the organisms.
Try to classify one or more of these using Prescott’s book How to Know the Fresh
Water Algae available in the laboratory.

Page 47
The most probable number (MPN) technique is a statistical approach to quantification
and, as in the plate count technique, involves the preparation of a decimal dilution series.
The sample must be diluted to extinction. An aliquot of each dilution (usually 1.0 or 0.1
ml) is then added to a series of tubes containing a growth medium. The number of
replicate tubes employed dictates the degree of accuracy. Tables are available for 3, 5, 8,
and 10 tube determinations. The table below is a three-tube table. As an example,
consider the following: a sample is decimally diluted to extinction. Three 1-ml aliquots
from each dilution are dispensed to three tubes of nutrient broth. After incubation, growth
is determined by turbidity. Any turbidity is considered a positive test. At a dilution of 10 -
2 all three tubes are turbid, 10 -3 has two turbid tubes, 10 -4 has one turbid tube and 10 -5 has
none. The combination is 3, 2, 1, 0 or 2, 1, 0. Referring to the table, the combination 3, 2,
and 1 gives a MPN value of 1.50 for the center dilution and 2, 1, 0 gives a value of 0.15.
The MPN value is multiplied by the reciprocal of the dilution. In these examples, the
MPN is (1.5 x 10 3 ).
THREE-TUBES MOST PROBABLE NUMBER (MPN) TABLE
# OF POSITIVE TUBES # OF POSITIVE TUBES
SERIES SERIES SERIES MPN SERIES SERIES SERIES MPN
A B C
A B C
0 0 0 24.00
Al-Agely and Ogram © 2004

Page 48
SOIL METABOLIC ASSESMENT
1. ENZYME – PHOSPHATASE ACTIVITY
OBJECTIVE:
Estimate the in situ activity of phosphatase in two contrasting soils.
INTRODUCTION:
While enzymes in soil are sometimes associated with roots, many are of microbial origin.
Soil enzymes may retain their activity long after their release from microbial cells
primarily because the enzymes form complexes with humic and clay colloids.
Complexing with these colloids renders the enzymes highly resistant to denaturation and
degradation. Thus, soil can have significant enzymatic activity independent of a native
microbial community.
Phosphatase mediates the generalized reaction:
Phosphatas
C-PO 4 + H 2 O ⎯⎯⎯⎯⎯⎯⎯→ C-OH + HPO 4
In this experiment, phosphatase activity in soil will be estimated by using p-nitrophenylphosphate
as a substrate. As the phosphate is cleaved away, p-nitrophenol is left. This
hydrolyzed product is light sensitive so your samples should be kept covered with
aluminum foil to prevent photolysis. As before, half the class will assess agriculture soil
and the other half forest soil. Each student’s data will serve as a replicate.

Page 49
MATERIALS:
Soil samples (agriculture or forest)
Two 2.0-ml Eppendorf microcentrifuge tubes
P-nitrophenyl-phosphate solution (100 mM), SIGMA= LOT 054H6178, FW = 371.1
g/l = 1M, 0.7422 g / 20 ml = 100 mM
CaCl 2 solution (0.5M)
NaOH solution (0.5M)
P-nitrophenol standard stock solution (100 µM), PNP standard: PNP FW = 139.11 g,
0.013911 g / 100 ml = 1mM solution, to make 100 ml of 1 µM, take 10 ml of 1mM
solution + 90 ml water. Use the 1 µM solution to make five standard solutions
between 0 and 10 µM (10, 5, 2.5, 1.25, and 0).
Microplate reader (vertical path length photometer with 405 nm interference filter)
Two flat bottom 300-µl wells
96-well microplate reader rack
Aluminum foil
Micropipettes and micropipette tips
Spatula, marking Pen, and tape
METHOD:
Label the microcentrifuge tubes as enzyme assay treatment and control.
Wrap each microcentrifuge with aluminum foil
Weigh out 0.25 g soil (dry mass basis).
Add 1 ml dH 2 O to the soil and swirl gently
Initiate the reaction by adding 0.25 ml of 100 mM p-nitrophenyl-phosphate to the
treatment microcentrifuge tube but NOT to the control
Place microcentrifuge tubes in rack and shake at 175 rpm for 30 min.
NOTE: The algae MPN assay should be started during this incubation time.
After the incubation period, add 0.25 ml of the p-nitrophenyl-phosphate solution to
the control.
Terminate the reaction by adding 0.25 ml of each 0.5M CaCl 2 and 0.5M NaOH to all
tubes.

Page 50
Place the treatment and the control tubes of each soil opposite each other in the
microcentrifuge and centrifuge at 10,000 rpm for 10 min.
Remove tubes from the microcentrifuge and cover with aluminum foil.
Run your samples against of standard solutions of 0 to 10 µM of p-nitrophenol.
Your samples may require dilution to fit the standard curve. Use dH 2 O to make 1:3
dilutions (0.1-ml sample to 0.2-ml dH 2 O)
Place your sample wells in the microplate reader rack and mark their locations.
The Microplate Program Manager will calculate the concentration of p-nitrophenol in
each well based on the standard curve.
Subtract the control p-nitrophenol concentration from the treatment p-nitrophenol
concentration to get the net reaction result.
Because each mole of p-nitrophenol produces one mole of phosphate, the final data
can be directly expressed as µmol phosphate released. Multiply your net results by
the dilution factor to express the total as µmol phosphate released g -1 soil hr -1 .
REFERENCES
Dick, W.A. and Tabatabai, M.A. 1992. Significance and uses of soil enzymes. pp. 95-127
In: Metting, F.B. (ed). Soil Microbial Ecology. Marcel Dekker, Inc.
Tabatabi, M.A. 1994. Soil Enzymes. Pp. 775-833 In R.W. Weaver and et al. (ed.)
Methods of soil analysis, Part 2. Microbiological and biochemical properties. Soil
Science Society of America, Madison, WI.

Page 51
E X E R C I S E
8
SOIL METABOLIC ASSESMENT
2. RESPIRATION / BIOMASS TESTS
RESPIRATION
OBJECTIVE:
Quantify soil microbial activities by measuring carbon dioxide production in treatments
of different carbon sources.
INTRODUCTION:
Respiration of microorganisms in soil was one of the earliest, and still is one of the most
frequently, used indices of soil microbial activity. Measurements of respiration have
been found to be well correlated with other parameters of microbial activity such as
nitrogen or phosphorus transformations, metabolic intermediates, and average microbial
numbers
Carbon comprises about 45 to 50% of the dry matter of plant and animal tissues. When
these tissues or residues are metabolized by microorganisms, 0 2 is consumed and C0 2 is
evolved in accordance with the following generalized reaction:
(CH 2 0) x + 0 2 ⎯⎯⎯⎯⎯⎯→ C0 2 + H 2 0 + intermediates + cell material + energy
In this reaction, all of the organic carbon should eventually be released as C0 2 . In actual
practice, under normal aerobic conditions, only 60 to 80% of the carbon is evolved as
C0 2 because of incomplete oxidation and synthesis of cellular and intermediary
substances. The quantities of C0 2 evolved depend on the type of carbon substrate, the
environmental conditions, and the types and numbers of microorganisms involved.

Page 52
When a portion of the soil is fumigated (or otherwise sterilized - e.g. microwave
irradiation), the difference in CO 2 evolved between fumigated and nonfumigated soil can
be used to estimate microbial biomass.
The exercise, which follows, will introduce a simple titrimetric method for measuring
C0 2 evolution from soils, and demonstrates the effects that different carbonaceous
substrates have on the rate of C0 2 evolution (microbial respiration). In addition,
comparisons between the fumigated and nonfumigated samples will provide data for
estimating the microbial biomass of the original soil.
MATERIALS:
FIRST PERIOD
Soil samples sieved through a 2 mm sieve
Biometer vessel
Solution of NH 4 N0 3 where 2 ml = 12 mg
Glucose
Cellulose powder
NaOH (1N)
Balances
SUBSEQUENT PERIODS
HCl, 1.0 N (132.25 ml conc. HCl in 1 L distilled water), to standardize titrate with 1
ml of phenolphthalein as the indicator, against ca. 1.5 g THAM buffer in 25 ml
distilled water. One mole of THAM = one mole of HCl.
BaCl 2 , 50% solution
Phenolphthalein (1 g phenolphthalein in 100 ml of 95% ETOH)
Pasteur pipettes
Burettes

Page 53
METHOD:
RESPONSE TO CARBON ADDITION
Treatments are Agriculture Soil (alone, with glucose, or with cellulose additions) and
Forest Soil (alone, with glucose, or with cellulose additions).
Each student will do either agriculture soil or forest soil.
Weigh out 50 g (dry mass basis) of soil (soil should be at ca. 10% moisture) and place
in 1 pint Mason jar.
Each soil will receive 2 ml of NH 4 N0 3 solution, 250 mg glucose plus NH 4 N0 3 or 250
mg cellulose powder plus NH 4 N0 3 (note, add dry ingredients and mix before adding
N-solution).
Place a 50 ml beaker on the soil.
Add exactly 10 ml of 1 N NaOH to the 50 ml beaker and seal with the Mason jar lid.
C0 2 produced by the soil is absorbed by the alkali.
After 1 wk of incubation, remove NaOH with a syringe, place in beaker, and recharge
beaker with fresh NaOH for the second wk reading.
To determine the amount of C0 2 produced:
1. Add 1 ml of phenolphthalein and 1 ml of 50% BaCl 2 to the beaker of NaOH to
precipitate the carbonate as an insoluble barium carbonate
2. Titrate to neutralize the unused alkali with 1 N HCl and record the exact used acid
volume.
3. Calculate the amount (mg) of C or C0 2 = (B-V) NE
Note: (B-V) is a simplification of (T-V)-(T-B)
Where: T = Total volume of NaOH at the start of the experiment, V = Volume
(ml) of acid to titrate the alkali in the C0 2 collectors from treatments (i.e., with
glucose and cellulose) to the endpoint, B = Volume (ml) of acid to titrate the
alkali in the CO 2 collectors from controls (i.e., without C addition) to the
endpoint, N = Normality of the acid, and E = Equivalent weight.
If data are expressed in terms of carbon, E=6; if expressed as CO 2 , E=22
4. Express the results as mg carbon dioxide produced per g soil. Show the
calculation, tabulate the class results and discuss.

Page 54
BIOMASS (DEMO ONLY):
Treatments are: 2 soil samples (Agriculture and Forest) and 2 microwave irradiation (+ or
-)
METHOD
Weigh 50 g dry mass of well-mixed soil (soil should contain about 10% moisture) and
place in Erlenmeyer flask of a Biometer Vessel (see fig. below).
One vessel from each soil is microwaved for 30 seconds (high power, 700 W, 2450 MHz
microwave). Note, this is sufficient to kill approximately 90% of the cell. After
microwaving, close vessel with rubber stopper.
A similar vessel is assembled with non-microwaved soil.
Charge side tubes with exactly 10 ml of 1.0 N Na0H and seal.
After 10 days, titrate the NaOH as above to determine, the CO 2 evolved.
Calculate the mg C biomass by the formula:
(HCl used in control - HCl used in microwaved soil) / 0.5 1 = mg C biomass
(Assuming 50% conversion of biomass to C in 10 days)

Page 55
A BIOMETER VESSEL
A
G
F
B
C
H
D
E
I
A) rubber policeman B) 15-gauge needle C) 50-mL tube D) alkali solution E) polyethylene tubing F)
Ascarite filter G) stopcock H) 250-mL flask I) 50-g soil sample (From Bartha and Pramer (1965) Soil
Science 100:68-70 Used with permission)

Page 56
QUESTIONS:
1. Why is the barium chloride added to the absorption vessel prior to titration?
2. What is the source of the carbon dioxide recovered in the nonamended soils?
How did the addition of glucose and cellulose affect microbial respiration -
why?
3. What factors influence the reliability of these methods as an index of
microbial activity?
4. How would you expect CO 2 evolution from a soil amended with groundsoybean
plants to compare to that of a soil amended with straw? Support your
answer!
REFERENCES:
Bartha, R. and Pramer, D. 1965. Features of a flask and method for measuring the
persistence and biological effects of pesticides in soil. Soil Science 100:68-70.
Hendricks, C.W. and Pascoe, N. 1988. Soil microbial biomass estimates using 2450
MHz microwave irradiation. Plant and Soil 110:39-47.
Horwath, W.R. and E.A. Paul. 1994. Microbial biomass. p. 753-773. In R.W. Weaver and
et al. (ed.) Methods of soil analysis, Part 2, Microbiological and biochemical properties --
SSSA book series, no. 5. Soil Science Society of America, Madison, WI .

Page 57
Further example of calculations for CO 2 evolution from soil (Note: this example is not
related directly to lab exercise).
1. CO 2 evolved from the soils is absorbed into the basic solution
CO 2 + 2NaOH ⎯⎯⎯⎯⎯→ Na 2 CO 3 + H 2 O
Therefore the Eqwt of CO 2 is 2 of the Mwt. = 44/2 = 22
(Meqwt = 0.222)
2. Na 2 CO 3 + BaCl 2 ⎯⎯⎯⎯⎯⎯⎯⎯→ BaCO 3 + 2NaCl
3. HCl + unreacted NaOH ⎯⎯⎯⎯⎯→ NaCl + H 2 0
4. Milliliters of HCl used X (Normality) of HCl = Meq of HCl
5. Meq of HCl = Meq of NaOH left unreacted with CO 2
6. Milliliters of NaOH X N = Total meq of NaOH used in the Experiment.
7. Total meq NaOH - meq of NaOH left = Meq of NaOH consumed by CO 2
8. Meq of Anything = Meq of Anything else.
9. Therefore Meq of NaOH consumed by CO 2 = Meq of CO 2 evolved.
10. Meq CO 2 X Meqwt of CO 2 (0.022) = Total grams of CO 2 evolved.
11. Grams X 1,000 = mg CO 2 evolved.
12. This is mg of CO 2 evolved from 50 grams of soil.
13. Therefore mg/50 = mg/g of soil (mg of CO 2 that is)

Page 58
E X E R C I S E
9
SOIL METABOLIC ASSESMENT
3. DNA EXTRACTION AND QUANTIFICATION
OBJECTIVE
Use MO BIO protocol to extract DNA of two contrasting soils
INTRODUCTION
Use kit MO BIO Catalog # 12800-100 for isolating DNA from 0.25 - 1gm soil samples.
Please wear gloves and avoid all skin contact with protocol materials. In case of accident contact,
wash thoroughly with water and do not ingest. See Material Safety Data Sheets for emergency
procedures in case of accidental ingestion or contact. Reagents labeled flammable should be kept
away from open flames and sparks.
MATERIALS:
Micro centrifuge (10,000 x g)
Pipette (volumes required 50 µl - 500 µl), and
Vortex
Kit Contents: 2 ml Bead Solution tubes (contains 550µl solution) 100
Description
Amt.
Solution S1
6.6 ml
IRS solution
22 ml
Solution S2
27.5 ml
Solution S3
143 ml
Solution S4
30 ml
Solution S5
6 ml
Spin filters units in 2 ml tubes 100
Collection tubes (2 ml) 300

Page 59
PROTOCOL
• To the 2ml Bead Solution tubes provided, add 0.25 gm of soil sample.
• Gently vortex to mix
• Check Solution S1. If Solution S1 is precipitated, heat at 60°C until dissolved
• Add 60µl of Solution C1 and invert several times or vortex briefly.
• Secure bead tubes horizontally using the Mo Bio Vortex Adapter and vortex at
maximum speed for 10 minutes.
• Centrifuge at 10,000 x g for 30 seconds. Be sure not to exceed 10,000 x g.
• Transfer the supernatant to a clean microcentrifuge tube (provided).
• Note: With 0.25gm of soil and depending upon soil type, expect between 400 to
450µl of supernatant. Supernatant may still contain some soil particles.
• Add 250µl of Solution C2 and vortex for 5 sec. Incubate 4°C for 5 min.
• Centrifuge the tubes for 1 minute at 10,000-x g.
• Avoiding the pellet, transfer 450µl of supernatant to a clean microcentrifuge tube
(provided).
• (To transfer entire volume, follow alternative protocol steps 12 through 21.)
• Add 200µl of Solution C3 to the supernatant and vortex for 5 seconds.
• Load approximately 700µl onto a spin filter and centrifuge at 10,000-x g for 1
minute. Discard the flow through, add the remaining supernatant to the spin filter,
and centrifuge at 10,000-x g for 1 minute. Note: Two loads for each sample
processed are required.
• Add 1200µl of Solution C4 and centrifuge for 30 seconds at 10,000-x g.
• Discard the flow through.
• Centrifuge again for 1 minute.
• Carefully place spin filter in a new clean tube (provided). Avoid splashing any
Solution S4 onto the spin filter.
• Add 50µl of Solution S5 to the center of the white filter membrane.
• Centrifuge for 30 seconds.
• Discard the spin filter. DNA in the tube is now application ready. No further steps
are required.
• Storing DNA at frozen (-20°C). Solution S5 contains no EDTA.

Page 60
DETAILED PROTOCOL (DESCRIBES EACH STEP)
PLEASE WEAR GLOVES AT ALL TIMES
1. To the 2ml Bead Solution tubes provided, add 0.25 - 1gm of soil sample. (For larger
sample sizes up to 10 grams, try using our Mega Prep Kit, catalog number 12900-10.
For amounts of sample to process see Hints and Troubleshooting Guide).
What is happening: Your soil or fecal sample has now been loaded into the bead
tube. This is the first part of the lysis procedure. The Bead Solution is a buffer that
will disperse the soil particles and begin to dissolve humic acids.
2. Gently vortex to mix
What is happening: This step mixes the sample and Bead Solution.
3. Check Solution S1. If Solution S1 is precipitated, heat solution to 60°C until
dissolved before use.
What is happening: Solution S1 contains SDS. If it gets cold, it will precipitate.
Heating to 60°C will dissolve the SDS. The Solution S1 can be used while it is still
hot.
4. Add 60µl of Solution S1 and invert several times or vortex briefly.
What is happening: Solution S1 contains SDS. This is a detergent that aids in cell
lysis. The detergent breaks down fatty acids and lipids associated with the cell
membrane of several organisms.
5. Add 200µl of Solution IRS (Inhibitor Removal Solution). Only required if DNA is to
be used for PCR.
What is happening: IRS is a proprietary reagent designed to precipitate humic acids
and other PCR inhibitors. This precipitation step is required if the intended use of the
DNA is for PCR. Humic acids are generally brown in color. They belong to a large
group of organic compounds associated with most soils that are high in organic
content.
6. Secure bead tubes horizontally using the Mo Bio Vortex Adapter tube holder for the
vortex (cat.13000-V1. Call 1-800-606-6246 for information) or secure tubes
horizontally on a flat-bed vortex pad with tape. Vortex at maximum speed for 10
minutes, (See alternative lysis method for less DNA shearing).
What is happening: The method you use to secure tubes to the vortex is critical. We
have designed the vortex adapter as a simple tool that keeps tubes tightly attached to
the vortex. It should be noted that although you can attach tubes with tape, often the
tape becomes loose and not all tubes will shake evenly or efficiently. This may lead to
inconsistent results or lower yields. The use of the vortex adapter is highly
recommended for maximum DNA yields.

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Mechanical lysis is introduced at this step. The protocol uses a combination of
mechanical and chemical lysis. By randomly shaking the beads, they collide with one
another and with microbial cells causing them to break open.
7. Make sure the 2ml tubes rotate freely in your centrifuge without rubbing. Centrifuge
tubes at 10,000-x g for 30 seconds. CAUTION be sure not to exceed 10,000 x g or tubes
may break.
What is happening: Particulates including cell debris, soil, beads, and humic acids,
will form a pellet at this point. DNA is in the liquid supernatant at this stage.
8. Transfer the supernatant to a clean microcentrifuge tube (provided).
9. Note: With 0.25gm of soil and depending upon soil type, expect between 400 to
450µl of supernatant. Supernatant may still contain some soil particles.
10. Add 250µl of Solution S2 and vortex for 5 sec. Incubate 4°C for 5 min.
What is happening: Solution S2 contains a protein precipitation reagent. It is
important to remove contaminating proteins that may reduce DNA purity and inhibit
downstream applications for the DNA.
11. Centrifuge the tubes for 1 minute at 10,000 x g.
12. Avoiding the pellet, transfer entire volume of supernatant to a clean microcentrifuge
tube (provided).
What is happening: The pellet at this point contains residues of humic acid, cell
debris, and proteins. For the best DNA yields, and quality, avoid transferring any of
the pellets.
13. Add 1.3ml of Solution S3 to the supernatant (careful, volume touches rim of tube)
and vortex for 5 seconds.
What is happening: Solution S3 is a DNA binding salt solution. DNA binds to silica in
the presence of high salt concentrations.
14. Load approximately 700µl onto a spin filter and centrifuge at 10,000 x g for 1 minute.
Discard the flow through, add the remaining supernatant to the spin filter, and
centrifuge at 10,000 x g for 1 minute. Repeat until all supernatant has passed through
the spin filter. Note: A total of three loads for each sample processed is required.
What is happening: DNA is selectively bound to the silica membrane in the spin filter
device. Almost all contaminants pass through the filter membrane, leaving only the
desired DNA behind.
15. Add 300µl of Solution S4 and centrifuge for 30 seconds at 10,000-x g.
What is happening: Solution S4 is an ethanol based wash solution used to further
clean the DNA that is bound to the silica filter membrane in the spin filter. This wash
solution removes residues of salt, humic acid, and other contaminants while allowing

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the DNA to stay bound to the silica membrane. Note: You can wash more than one
time to further clean DNA if desired. In some cases where soils have very high humic
acid content, it will be beneficial to repeat this wash step. There is 10% extra
Solution S4 in the bottle for this purpose. Solution S4 is also sold separately.
16. Discard the flow through from the collection tube.
What is happening: This flow through is just waste containing ethanol wash solution
and contaminants that did not bind to the silica spin filter membrane.
17. Centrifuge again for 1 minute.
What is happening: This step removes residual Solution S4 (ethanol wash solution). It
is critical to remove all traces of wash solution because it can interfere with down
stream applications for the DNA.
18. Carefully place spin filter in a new clean tube (provided). Avoid splashing any
Solution S4 onto the spin filter.
What is happening: Once again, it is important to avoid any traces of the ethanol
based wash solution.
19. Add 50µl of Solution S5 to the center of the white filter membrane.
What is happening: Placing the Solution S5 (sterile elution buffer) in the center of the
small white membrane will make sure the entire membrane is wetted. This will
increase release the desired DNA.
20. Centrifuge for 30 seconds.
What is happening: As the Solution S5 (elution buffer) passes through the silica
membrane, DNA is released, and it flows through the membrane, and into the
collection tube. The DNA is released because it can only bind to the silica spin filter
membrane in the presence of salt. Solution S5 is 10mM Tris pH 8 and does not
contain salt.
21. Discard the spin filter. DNA in the tube is now application ready. No further steps are
required.
Storing DNA at frozen (-20°C). Solution S5 contains no EDTA.

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HINTS AND TROUBLESHOOTING GUIDE
AMOUNT OF SOIL TO PROCESS
Depending on soil type, usually 0.25gm -1gm works well. Typically, only 0.25 g of the more
absorbent soil types, such as potting soils, can be processed. For wet soils, see “Wet soil sample”
below.
WET SOIL SAMPLE
If soil sample is high in water content remove contents from bead tube (beads and
solution) and set aside. Add soil sample to bead tube and centrifuge for 30 seconds at
10,000 x g. Remove as much liquid as possible with a pipette tip. Add beads and bead
solution back to bead tube and follow protocol starting at step 2.
IF DNA DOES NOT AMPLIFY
This is due to high humic acid content in soil sample. If the humic acid content is sample
is high, you can do the following:
1. Diluting template DNA may also work because this will also dilute the inhibitors
of the reaction.
2. Perform two to three washes of Solution S4 in steps 15 through 18.
3. Dilute the elution three fold and add two volumes of Solution S3. Run through
spin filter, wash and elute.
4. Make sure to check DNA yields by gel electrophoresis or spectrophotometer
reading. An excess amount of DNA will also inhibit a PCR reaction.
5. If DNA will still not amplify after trying the steps above, then PCR optimization
may be needed.
ELUTION SAMPLE STILL BROWN
This is due to high humic acid content in soil sample. If the humic acid content is sample
is high, you can do two to three washes of Solution S4 in steps 15 through 18. If elution
solution is still brown, dilute the elution three fold and add two volumes of Solution S3.
Run through spin filter, wash and elute.
ALTERNATIVE LYSIS METHOD
After adding Solution S1, vortex 3-4 seconds
Add the IRS Solution, vortex 3-4 seconds then heat to 70°C for 5 min. Vortex 3-4 seconds AND
Heat another 5 minutes, Vortex 3-4 seconds. This alternative procedure will reduce
shearing but may reduce yield.

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CONCENTRATING THE DNA
Your final volume will be 50µl. If this is too dilute for your purposes, add 2µl of 5M NaCl and
mix. Add 100µl of 100% cold ethanol and mix. Centrifuge at 10,000-x g for 5 min
Decant all liquid.
Dry residual ethanol in speed vacuum desiccators, or air dry.
Resuspend precipitated DNA in desired volume.
DNA FLOATS OUT OF WELL WHEN LOADED ON A GEL
You may have inadvertently transferred some residual Solution S4 into the final sample. Prevent
this by being careful in step 17 not to transfer liquid onto the bottom of the spin filter basket.
Ethanol precipitation is the best way to remove Solution S4 residue. (See concentrating DNA
above)
STORING DNA
DNA is eluted in Solution S5 (10mM Tris) and must be stored at -20°C or it may degrade over
time. DNA can be eluted in TE but the EDTA may inhibit reactions such as PCR and automated
sequencing.
CELLS ARE DIFFICULT TO LYSE
If cells are difficult to lyse, 10-min incubation at 70°C, after adding Solution S1, can be
performed. Follow by continuing with protocol step 5.
REFERENCES:
Procedure with modification from MO BIO Laboratories Inc (http://www.mobio.com)

Al-Agely and Ogram © 2004
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Al-Agely and Ogram © 2004
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E X E R C I S E 10
SOIL RHIZOSPHERE:
OBJECTIVE:
Isolate and directly observe your desire plant rhizosphere microorganisms using dilution
techniques and compound microscopes
INTRODUCTION
The rhizosphere is defined as the soil volume in close contact with plant roots. Roots
excrete a wide range of organic materials into the soil and these greatly influence the
development of a rhizosphere microbial community. Microbial populations differ both
quantitatively and qualitatively from those in the bulk soil. Bacteria populations in the
rhizosphere are often 10-100x higher than in root-free portions of the same soil. The
rhizosphere effect may be expressed in terms of a R/S ratio (rhizosphere population/rootfree
soil population). The rhizosphere also contains a higher proportion of gram
negative, nonsporulating, rod-shaped bacteria, and a lower proportion of gram positive,
nonsporulating rods, cocci, and pleomorphic forms.
Microorganisms in the rhizosphere can have a marked influence on plant growth
(positive or negative). Indeed, the rhizosphere is a complex zone of interactions among
microbial populations and between microorganisms and plant roots. In this exercise you
will use the dilution-plating technique to isolate rhizosphere microorganisms and also
observe microorganisms the root surface by direct microscopy.
METHODS
1. DILUTION-PLATE TECHNIQUE
Obtain a block of soil (approximately 10 cm 3 ) that contains herbaceous plant roots.
Crush the block, with as little tearing of the roots as possible. Remove roots and gently
shake to remove superfluous soil. Place roots, along with adhering soil in weighed flasks
containing 100 ml sterile H 2 0 and glass beads. Shake flasks vigorously for 5 min. on a

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mechanical shaker. Prepare dilutions as in exercise #1 and Plate in bacteria (start at 10 -3
because less soil goes into the first dilution compared with soil dilution lab).
To determine weight of rhizosphere soil, the roots are removed from the original dilution
flask, washed, and the wash water is collected in the original flask. The water is
evaporated and the soil residue is dried to constant weight in an oven at 105-110 o C. The
flask containing dry soil is weighed and dilution factors are calculated, allowance being
made for the amount of soil removed in preparing the dilutions.
Count bacterial colonies at appropriate time intervals during the week and in the
following laboratory session. Describe the morphology of typical colonies. How do
these counts compare to the bulk soil counts in Exercise #1?
2. DIRECT MICROSCOPY.
Obtain 5 roots (< 2 mm diam.) and gently wash to remove excess soil. Place washed
roots in FITC solution (see page 27) for several minutes. Wash roots in sodium
carbonate buffer (5 changes). Cut roots into 1-2 cm lengths and mount in glycerol
adjusted to pH 9.6 with buffer. Observe roots with epifluorescent microscopy (2170
McC).
REFERENCE:
Chapter 2 in Metting's Soil Microbial Ecology