Laboratory 6
Enumeration
Introduction
One of the central questions a researcher asks when studying any bacterial system is
“how many bacteria are present in this sample?” This applies to environmental samples where
we want to learn about a native population as well as to lab cultures where we are measuring
physiological functions. There are many ways to count bacteria and each of them has advantages
and disadvantages.
Measuring the optical density (O.D.) of a sample is an indirect method of determining the
number of cells present. The amount of light of a specific wavelength that is absorbed by a
culture is related to the number of cells. This is a very fast and easy way to count cells when
possible. Unfortunately, in order to use this method, you first need to calculate a response factor
(the ratio of the number of cells to the change in absorbance) for the specific cells and media that
you are using. This requires use of an independent method for counting the cells.
Another common method for counting cells is to serially dilute the original sample and
spread-plate the resulting dilutions. This is known as the plate-count method. The colonies that
arise after incubation are counted to determine the original cell density. This method requires
more time and supplies than measuring the optical density but it does result in an absolute
number of cells rather than a relative value. The one important thing to remember when doing
plate-counts is that it only counts the viable cells (those cells that are able to grow under the
specific incubation conditions used).
If you want to count all of the cells present in a sample then a direct-counting method
should be chosen. This generally involves using a microscope to visualize the cells. Special
microscope slides that have surfaces calibrated into precise volumes can be used to determine
exactly how many cells there are in a milliliter of sample. Sometimes stains are used to make the
cells more visible. This method counts all cells whether they are alive or not. The decision as to
whether viable or total cell counts are more relevant will depend on the experimental question
being asked.
In this lab you will be counting the number of bacteria in an overnight culture of E. coli
using three different methods: optical density, direct counts and plate counts. You will then be
able to calculate a response factor for optical density and compare the direct count and viable
count methods.
Materials
Equipment
- spectrophotometer
- microscopes
- Petroff-Hauser chamber
- bunsen burners
- spreaders
Cultures
- Escherichia coli (broth)
Supplies
- Tryptic Soy Agar (TSA) plates
- Liquid TSB media
- Pasteur pipettes and bulbs
- Crystal violet stain
- 95% ethanol
Procedures
Plate-Counts (Counting by Dilution Series)
1. Make a dilution series of the E. coli culture in liquid TSB media.
a. Aseptically transfer 1 ml of the culture into a tube containing 9 ml of TSB
medium.
b. Transfer 1 ml of the resulting tube into another tube with 9 ml of medium.
c. Repeat this process until you have produced 9 tubes. (10-1 – 10-9).
2. Spread 0.1 ml of each dilution tube onto a TSA plate.
3. Incubate at 25C for two days.
Period 2
4. Count the number of colonies on plates that contain between 20 and 200 wellseparated colonies.
5. Calculate the number of cells in the starting culture by multiplying the number of
colonies times the inverse of the dilution.
Optical Density (Counting by Spectrophotometer)
1.
2.
3.
4.
5.
Turn on the spectrophotometer and set it to 620 nm.
Set the transmittance to 0 with no sample in the chamber.
Set the absorbance to 0 using un-inoculated TSB media as a blank.
Measure the absorbance of each of the dilution tubes used in the dilution series.
Plot the absorbance against the dilution factor.
Direct Counting (Petroff-Hauser Chamber)
1. Use a pipette to transfer 0.9 ml of the starting culture to a clean test-tube.
2. Add 0.1 ml of crystal violet stain to the culture.
3. Use a Pasteur pipette to transfer a drop of the stained culture into the Petroff-Hauser
slide.
4. Carefully place the cover slip onto the calibrated surface of the counting chamber.
5. Place the chamber on the microscope stage and focus under the 20X objective to
locate the large counting grids.
6. Switch to high power and count the number of bacteria in at least 20 of the small
squares.
(If there are too many bacteria to count, make a 1:10 dilution of the culture and start
over)
7. Average the number of cells and calculate the number per ml. (cubic centimeter).
Dilution Series
DATA
20 points
Name:
Date:
Plate counts
Count the number of colonies on plates that contain between 20 and 200 well-separated
colonies.
colonies
dilution
Calculate the number of cells in the starting culture by multiplying the number of colonies
times the inverse of the dilution.
Concentration of cells in starting culture =
cells/ml
Optical density
dilution tube
absorbance
dilution tube
culture
10-5
10-1
10-6
10-2
10-7
10-3
10-8
10-4
10-9
Plot the absorbance against the dilution factor. (Attach graph)
absorbance
Direct counts
Average the number of cells/square and calculate the number per ml. (cubic centimeter).
Each small square is 1/20th X 1/20th of one millimeter square and 1/50th of one millimeter
deep.
Average number of cells per square Cells per milliliter Cells per milliliter -
(corrected for dilution by dye)
Response factor
Use the absorbance graph, the plate-count data and the direct-count data to calculate two
separate response factors (# of cells / absorbance unit) for absorbance at 620nm for E. coli.
Plate count response factor
cells/absorbance unit
Direct count response factor
cells/absorbance unit
Question:
1). Which response factor is more accurate for correlating absorbance to cell numbers? Why?
2). Give an example of when you would want to use a direct count and an example of when you
would want to use a live count for quantifying bacteria.
Laboratory 7
Coliform Testing
Introduction
The determination of coliform bacteria in water is one of the oldest and most commonly
used techniques in environmental microbiology. Coliform bacteria are used as indicators of
potential health threats. The presence of bacteria in the coliform group in any sample is
considered to be evidence of fecal contamination and therefore of the possibility that pathogenic
organisms are present. The coliform group is functionally defined according to the way they are
detected as “all aerobic and facultative anaerobic, gram-negative, nonspore-forming, rod-shaped
bacteria that ferment lactose with gas and acid formation within 48hr at 35C”. This definition
was devised to detect E. coli but it also includes several other genera in the family
Enterobacteriaceae including Klebsiella, Enterobacter and Citrobacter. All of these organisms
are coliforms and will give a positive result in these tests but not all of them are necessarily the
result of fecal contamination. In order to help clarify the results, and understand the implications
of a coliform test, it is helpful to identify the number of “fecal coliforms” or to identify the
species of bacteria detected. Fecal coliforms are defined as those coliforms that are capable of
growth at 44C. The increased temperature is meant to select only those organisms adapted to
growth in the human body. Species typing is carried out by conducting several biochemical tests.
In this lab you will use two different methods two measure the number of coliforms
present in surface water samples. The first method (membrane-filtration) is a one-step process
where coliform colonies are counted directly from filters that have been incubated on EMB
media. This method is relatively quick and easy but it is more prone to errors than the alternative
“multiple tube fermentation” technique. Multiple tube fermentation is carried out in three
separate steps. The “presumptive” step involves testing for gas production from lactose
fermentation in selective media. The number of coliforms is determined by using the MPN
method during this step. This is followed by the “confirmed” step where positive samples from
the presumptive test are transferred to a more selective media and re-tested for gas production.
Finally, the test is “completed” by isolating single colonies with a positive reaction on EMB agar.
You will further characterize the isolates from the completed step by using a commercial
kit to determine the species.
Materials
Equipment
- incubator (35C)
- bunsen burners
- inoculating loops
- pipettes and bulbs
- sterile graduated cylinders
- 100 ml dilution bottles
- sterile filter holders and vacuum
flasks
Cultures
- environmental water samples
- Escherichia coli
- Enterobacter aerogenes
- Pseudomonas sp.
Media
- Durham tubes with lauryl tryptose
broth (LTB) (2X and 1X)
- Durham tubes with brilliant green
bile (BGB) broth
-
Eosine-Methylene Blue (EMB)
plates
TSA plates and TSB media in tubes
Tryptone broth
-
Enterotube II tests
sterile water
Kovac’s reagent
1% aqueous tetramethyl-pphenylenediamine HCl
Supplies and Reagents
- membrane filters
Procedures (Membrane Filtration Technique)
Period 1
1. Assemble a sterile filtration apparatus with a membrane filter
2. Aseptically transfer 25 ml of the sample to 225 ml of sterile distilled water.
3. Mixed diluted sample well.
4. Filter 10 ml of diluted water.
5. Aseptically transfer the membrane filter onto the surface of an EMB plate.
6. Replace the membrane filter and filter 50 ml of diluted water.
7. Aseptically transfer the membrane filter onto the surface of an EMB plate.
8. Replace the membrane filter and filter 100 ml of diluted water.
9. Aseptically transfer the membrane filter onto the surface of an EMB plate.
10. Incubate the plates upright at 35C for 24 hrs.
Period 2
11. Count the number of dark red colonies with a metallic sheen.
12. Calculate the number of coliforms per liter of the water sample.
Procedures (Multiple Fermentation Tube Technique)
Period 1
Most Probable Number (MPN) Determination (Presumptive Test)
1. Inoculate 5 tubes of 2X lauryl tryptose broth with 10 ml of your water sample and 5
tubes of 1X broth with 1 ml.
2. Dilute your water sample 1:10 by adding 10 ml to 90 ml of sterile water.
3. Inoculate 5 tubes of 1X broth with 1 ml of the diluted sample and 5 tubes of 1X broth
with 0.1 ml.
4. Incubate the tubes at 35C for two days.
Period 2
5. Examine each tube for gas production.
6. Record the results for each tube and use the MPN Table to calculate the number of
coliforms in the water samples.
Confirmed Test
1. Inoculate BGB tubes with a loop full of culture from each of the most dilute positive
tubes from the presumptive test.
2. Incubate the tubes at 35C for two days.
Period 3
3. Examine each tube for gas production.
Completed Test
1. Streak two EMB plates from each positive BGB tube.
2. Incubate one plate at 35C and the other at 44C for 24 hrs.
Period 4
3. Examine the plates for the presence of green, metallic colonies.
Species Typing
1. Select a positive E. coli colony from the EMB plates.
2. Streak a TSA plate with that colony from the EMB plate.
3. Incubate at 35C for 24 hrs.
Procedures (Enterotube II kit)
Period 5
Oxidase Test
1. Pick a colony from the TSA plate using a wooden applicator stick and smear it onto a
filter soaked in tetramethyl-p-phenylenediamine.
2. Development of a purple color indicates a positive reaction.
3. Record the result on the Data sheet.
Inoculating the Tube
1. Remove both caps from the Enterotube without touching the wire inside.
2. Use the tip of the inoculating wire to pick one large (2-3mm) colony from your TSA
plate (there should be a visible amount of cells on the wire).
3. Inoculate the tube by first twisting the wire then withdrawing it through all 12
compartments while slowly turning it.
4. Reinsert the wire back through all 12 compartments until the notch in the wire is even
with the opening of the tube.
5. Break off the handle end of the wire by bending it.
6. Use the handle of the wire to punch holes through the foil covering the air inlets of
the last eight compartments (adonitol, lactose, arabinose, sorbitol, Voges-Proskauer,
dulcitol/PA, urea and citrate).
7. Replace both caps.
8. Incubate at 37C for 24 hrs.
Period 6
Reading the Results
1. First read the color reactions for all of the tests except indole and Voges-Proskauer
and record the results on the ID form.
2. Use a syringe and needle to pierce the plastic film and add one or two drops of
Kovac’s reagent to the H2S / indole compartment.
3. Development of a red color in the compartment indicates a positive result.
4. Record the result on the ID form
5. Calculate the biotype number and determine the isolate’s identity from the code book.
6. If the code book indicates that it is necessary, perform the Voges-Proskauer test for
confirmation.
a.
Add two drops of 20% KOH and three drops of alpha naphthol to the
chamber.
b.
Development of a red color within 20 min. indicates a positive reaction.
DATA
20 points
Name:
Date:
Water sample name:
Membrane filter test
# of coliform colonies:
10 ml
50 ml
100 ml
# of coliform cells per liter
MPN test
Presumptive test
# of positive tubes
10 ml
1.0 ml
0.1 ml
0.01 ml
Most probable number
Confirmed test
How many of the BGB tubes are positive?
/
How does this change the estimate of how many coliforms were in the original sample?
Completed test
How many of the EMB plates showed typical coliform colonies at 35˚C?
How many of the EMB plates showed typical coliform colonies at 44˚C?
Species typing
Indole test
(Attach Enterotube ID form)
Species identity
Laboratory 8
Chlorination
Introduction
The final step in the treatment of sewage and other wastewaters often involves
disinfecting the treated water before discharging it into a river or lake. This is done to kill any
pathogenic organisms that may have survived through the initial stages in treatment.
Chlorination is the most common method used for the disinfecting stage of wastewater treatment.
Chlorine gas is bubbled through the water where it hydrolyses to form hypochlorous acid (HOCl)
and hypochlorite (OCl-).
Hydrolysis
CL2  HOCl + H+ + CL-
Ionization
HOCl  H+ + OCl-
Hypochlorous acid is a strong oxidizing agent that acts by reacting with cell components
to cause irreparable damage. Hypoclorite is less effective as an oxidizer. Together these two
compounds are referred to as the free residual chlorine. Hypoclorous acid can also be formed by
adding sodium hypochlorite (NaOCl) to the wastewater. Sodium hypochlorite is the active
ingredient in Clorox bleach.
NaOCl + H2O  HOCl + NaOH
In addition to killing cells, the free residual chlorine also oxidizes non-living organic
matter and several inorganic components of wastewater such as ammonia, iron and sulfides.
When hypochlorous acid reacts with ammonia, monochloramine and dichloramine are formed.
These compounds are also oxidizers and contribute to the disinfection process. They, along with
some chloro-organic compounds, are referred to as combined residual chlorine. When enough
chlorine has been added to completely oxidize the combined residual chlorine then free residual
chlorine will begin to accumulate. This process is known as breakpoint chlorination.
Although chlorination is performed in order to kill all of the remaining pathogens in
treated wastewater it is not always completely effective. Some organisms will only be injured by
the chlorine and some will escape treatment altogether. When this happens, there is potential for
re-growth of the pathogens after they have been discharged into clean water. This is why it is so
important to be sure that the breakpoint has been reached. In this lab we will be carrying out a
simple chlorination series with different concentrations of sodium hypochlorite and measuring
the free residual chlorine with a colorimetric assay. Then we will evaluate the survival of
coliforms in the treated wastewater.
Materials
Equipment
- spectrophotometer
- incubator
Cultures
- sewage samples
Supplies
- clean water
- Clorox bleach
- orthotolidine reagent
- EMB plates
- beakers and flasks
- 16 X 150mm test tubes
- pipettes and bulbs
Procedures
Standard Curve
1. Dilute a sample of Clorox bleach by adding 1ml of bleach to 499ml of distilled water.
2. Set up a standard curve by labeling five 16 X 150mm, glass test tubes as 0, 0.25, 0.5,
1.0 and 2.0 mg/L Cl.
3. Add 5ml of distilled water to each tube.
4. Add 0, 10, 20, 40 or 83l of the diluted bleach to the appropriate tubes.
5. Add 1ml of orthotolidine reagent to each tube and incubate for 4min. at room
temperature.
6. Read the absorbance at 460nm.
7. Plot the data to obtain a standard curve.
Chlorination
1. Dispense 60ml portions of “sewage” into seven 125 ml Erlenmeyer flasks.
2. Add 0, 0.5, 1, 2, 3, 5 or 8 ml of diluted bleach (1:500) to each of the flasks.
3. Mix each sample well and allow it to sit for 20min.
4. Remove 5 ml from each flask and transfer it to a 16 X 150mm, glass test tube.
5. Add 1 ml of orthotolidine reagent to each tube and incubate at room temperature for
4min.
6. Read the absorbance at 460nm.
7. Calculate the concentration of residual chlorine.
8. Select three flasks: one that has no residual chlorine; one that has a residual chlorine
conc. between 0 and 0.4 mg/l; and one with a higher residual chlorine concentration.
9. Test each of these flasks for coliforms as described below.
Counting Coliforms
Period 1
1.
Make a dilution series of each sewage sample into buffered water.
a.
Aseptically transfer 0.1 ml of the sample into a tube containing 9.9 ml of
buffered water.
b. Transfer 0.1 ml of the resulting tube into another tube with 9.9 ml of water.
c. Repeat this process until you have produced 5 tubes. (10-2 – 10-10).
2. Spread 0.1 ml of each dilution tube onto an EMB plate.
3. Incubate the plates upside-down at 37C for 24 hrs.
Period 2
4. Count the number of dark red colonies with a metallic sheen.
5. Calculate the number of coliforms per milliliter of the water sample.
Dilution Series
DATA
15 points
Name:
Date:
Standard curve:
Chlorine conc.
0
0.25
0.5
1.0
0.5
1
2
3
10-5
10-7
10-9
10-11
2.0
Absorbance
Chlorination:
Chlorine
0
5
8
Abs.
Conc.
Coliform counts:
10-3
Flask 1
Flask 2
Flask 3
Observations:
cells/ml
Laboratory 9
Rossi-Cholodny Slides
Introduction
The distribution of microorganisms in soil is heterogeneous. Microbes need nutrients and
water to survive and these resources are not evenly distributed in soil. The structure of soils is
composed of particles of inorganic and organic matter and the pores in between these particles.
The pore spaces may be filled with water or air. Bacteria are mostly found attached to particles
growing in small micro-colonies wherever nutrients can be found. Filamentous organisms such
as actinomycetes and fungi make up much of the biomass. When they grow around soil particles
they help to cement them into aggregates. These aggregates have their own internal pore spaces.
The filaments also stretch between aggregates.
Often when soils are examined microbiologically, the native structure is destroyed and it
is impossible to determine the original distribution of the microbes in their natural habitat. One
way to visualize how microbes are distributed in soil is to use the soil contact slide method
developed by Rossi and Cholodny (1938). In this technique, glass slides are buried in soil and
allowed to incubate. Bacteria and fungi attach to the glass as though it was a mineral particle
and grow on the surface. When the slide is carefully uncovered and stained, the microbes remain
in their original positions and their associations with particles and each other can be seen.
In this lab we will be staining and examining slides that have been buried in soils that
have received the following amendments: 1.6g of yeast extract, 0.2g of inorganic fertilizer, 1.6g
of yeast extract and 0.2g of inorganic fertilizer, or no addition. All of the slides were incubated
for ~2 weeks.
Materials
Supplies
- staining racks and trays
- Pasteur pipettes and bulbs
Reagents
- acetic acid
- phenolic rose bengal stain
Cultures
- Rossi-Cholodny slides
Equipment
- Phase contrast microscopes
- bunsen burners
Procedures
Soil Slide Stain
1. Start with a microscope slide that has had ¾ of its length buried in a beaker of garden
soil and incubated for over one week.
2. Remove the slide by tilting it backwards so that the top of the slide is not scraped off.
3. Shake off any large soil particles and clean the bottom of the slide with a moist paper
towel.
4. Allow the slide to thoroughly dry.
5. Immerse the slide in 40% acetic acid for 1-3 min. (Handle the Acid Carefully!).
6. Wash off the acid with water, being careful not to disturb the cells).
7. Flood the slide with phenolic rose bengal stain for 5-10 minutes.
8. Wash off the excess stain and allow the slide to dry.
9. View under the microscope.
10. Look for fungal filaments, actinomycetes and bacterial micro-colonies.
11. Record all observations.
Adapted from “Environmental Microbiology: A laboratory Manual” by Pepper, Gerba and
Brendecke. 1995. Academic Press, New York.
DATA
Name:
Observations:
10 points
Date:
Laboratory 10
Soil Enzyme Activity Part I
(Polymer Degradation)
Inroduction
Complex polymers such as starch, cellulose, proteins and nucleic acids make up an
important nutrient source for bacteria living in the soil. These polymers are components of the
cells of other organisms (plants, animals, fungi and bacteria) in the soil. They become available
after those cells die. Because of their large size, most polymers are insoluble in water. This
means that in their native state, they cannot be taken up into bacterial cells for metabolism.
Some bacteria can only use the polymers after they have been broken down into soluble
monomers such as glucose, amino acids or nucleotides. They depend on other organisms to
degrade the polymer structure. Other bacteria are able to cleave the polymer bonds by hydrolysis
(breaking by the addition of water). This process requires the bacteria to produce enzymes that
travel outside of the cell to attack the insoluble polymers. Although these enzymes allow the
organism to break down a plentiful nutrient source in the soil, the cells need to expend a lot of
energy to make the enzymes. In addition, many of the solubilized monomers will diffuse away
from the cell and be taken up by other bacteria.
In this lab we will be examining four pure cultures for their ability to hydrolyze starch
casein and gelatin. We will also plate soil dilutions on these same substrates to identify native
soil organisms that are effective polymer degraders.
Materials
Equipment and Supplies
- incubator (30C)
- bunsen burners
- spreaders and inoculating loops
- 99ml bottles with sterile water
- 1ml pipettes and pumps
Cultures
- soil samples
- Serratia marcesens
-
Escherichia coli
Pseudomonas fluorescens
Bacillus megaterium
Media and Reagents
- starch plates
- gelatin plates
- skim milk plates
- iodine solution
Procedures
Period 1
1.
2.
3.
4.
Divide one of each type of polymer plate into four sections.
Inoculate each plate with all four of the pure cultures (one in each section).
Set up a soil dilution series as shown in the diagram.
Use the dilutions to spread plate onto all three types of polymer plate (12 plates in all).
Period 2 (after 1 week)
Casein Hydrolysis
1. Examine each of the skim milk plates for colonies that show a zone of clearing around them.
Starch Hydrolysis
1. Flood each of the starch plates with an iodine solution.
2. Look for clearing zones around the colonies.
Gelatin Hydrolysis
1. Look for clearing zones around the colonies.
DATA
10 points
Name:
Date:
Starch
Hydrolysis
Casein
Hydrolysis
Gelatin
Hydrolysis
Ps. fluorescens
S. marcesens
B. megaterium
E. coli
Soil Dilution Observations:
How many colonies? What percent have clearing zones? How much diversity?
Casein Plates
Gelatin plates
Starch plates