Coliform in Drinking Water

Coliform in Drinking Water

Modified from: Standard Methods for the Examination of Water and Wastewater, 14th Ed, (1975). pages 928-935.

Students can test the drinking water from their homes, school, etc using this protocol.  The health standard for drinking water allows no more than 5 coliform per hundred mL.  For this reason, bacteria from 250 mL will be collected on a membrane filter, and grown on top of m Endo medium.  Coliform bacteria appear as red colonies on this medium.  More than 13 colonies with this assay does not meet health standards.
One student in 2001 tested her tap water before and after filtration through a water “purification” device. The image shows two plates , the one on the left is tap water, the plate on the right is water filtered through the filter.  Clearly the filter is contributing bacterial load to the water.  These devices must be regularly changed and maintained in order  to prevent such contamination.

Sterile 250 mL capped bottles (1/student)
sterile 47 mm petri dishes (1 per student)
sterile 47 mm memb. filters, 0.45 µm pores
sterile 47 mm millipore pads
vacuum pump
3 vacuum hoses joined with “T” joint
2 strong hose clamps
1000 mL side arm filter flask
m-Endo Broth MF powder
sterilized repipet in 250 mL bottle

millipore filtering apparatus:
sintered glass platform in #8 stopper
glass cylinder, 300 mL capacity
800 mL beaker with 400 mL EtOH
150 mL beaker with 100 mL EtOH
triangle-tipped Tongs
forceps with bent tipped blade
Bunsen burner
protective eye wear

(pay close attention to the location of the valves)

Vacuum Filtering Apparatus:
note the labeled valves:
vacuum pump
on/off power switch
main vacuum line
main vacuum line clamp
T joint
relief valve clamp
filter flask
#8 stopper
screen platform
membrane filter
glass cylinder, 300 mL
cylinder clamp

Plate Ready for Incubation:
(exploded view)
50 mm petri dish top
47 mm membrane filter
47 mm pad with 2 mL m-Endo MF
50 mm petri dish bottom

I. COLLECTION OF WATER (Collect the same day as performance of assay):


Determine the precise name of your water district, record it in your notebook.
Run tap water until it is cold (to clear out pipes, at least a minute or so).
Fill sterile 250 mL capped bottle with water, rinse several times, finally fill to neck, cap securely, maintaining sterility .

II. PREPARATION OF 50 mL OF MEDIUM (for 20 assays):


a: For up to 20 determinations, weight out 2.4 g m-Endo Broth MF
powder into 150 mL beaker.
b: Add 49 mL dH20 and 1 mL EtOH.
c: Bring to boil, remove from heat immediately.
Using sterile technique, pour into sterilized repipet vessel. Securely screw down repipet to container. Clamp to ring stand for stability.


Label 47 mm top of sterile petri dish with initials, seat number, date & source.
Flame off EtOH from bent blade-tipped forceps, pick up sterile pad, place in bottom of sterile petri dish , replace cover.
Repipet 2.0 mL m-Endo Broth sterilely onto pad , replace lid, keep bottom down.

IV. SET UP PLATE: Note: Wear safety goggles, tie back hair, keep flammable materials away from flame.

PRELIMINARY: Check for adequate head space in the filter flask to contain the water you are about to filter. If inadequate, empty out the flask into the sink, and replace the support platform. If necessary, re-sterilize surface of support platform with EtOH, turn on vacuum briefly to dry, then tightly clamp the main vacuum line.
Also, apply vacuum to the platform to remove any water or alcohol remaining from the previous filtrations by closing the relief valve, opening the main valve, then closing the main valve

Once the platform is dry:
SET UP MEMBRANE FILTER: Flame off EtOH from blade forceps

pick up sterile membrane filter , (discard the blue spacer discs)

center it on screen platform . Open main vacuum line to hold membrane in place (turn on pump if not on yet)


STERILIZE THE CYLINDER (Dangerous step): Ensure that all surfaces of the glass cylinder are immersed in 95% EtOH , pick up with fingers (touch outside of the cylinder only), invert.
Grasp cylinder upside down with triangle-tipped tongs (Ekco, for instance) and allow excess EtOH to drip into 800 mL beaker, touching off the last drops on a paper towel



CAREFULLY flame off the EtOH away from EtOH beakers and other students). It will flame up fairly high, but should burn off in a few seconds. Pass quickly through the flame once more to ensure that all of the EtOH has been removed.

Grasp the outside of the sterilized cylinder with your fingers. (It should not be too hot if you touched off the EtOH before flaming.)

SET UP FILTER APPARATUS: Place sterile cylinder centered over the membrane filter and support platform.
Clamp in place with spring clamp , vacuum still on.


Pour your 250 mL water sample into the cylinder , monitoring that it is not leaking at the clamped joint .

The vacuum will draw the water through, and all bacteria which may be in the 250 mL sample will be trapped on the surface of the membrane filter.

When all of the water has been drawn through, unclamp and remove the cylinder , place it carefully back in the EtOH, top down. (Do not let it drop into the beaker.)

Clamp the main vacuum line shut and open the relief valve to release the vacuum in the flask. With sterile, EtOH-free blade forceps, gently lift the edge of the membrane filter and remove from the screen platform . (Caution: the membrane filter is brittle.)

TRANSFER MEMBRANE FILTER TO PREPARED PAD , avoiding bubbles by lowering from one side first.
Rest it on far edge of petri dish, slowly pull it across the edge down toward you until . . .

. . . it drops down onto the pad.
If done properly, it will be
centered on the pad .

Ensure that the membrane is completely flat on the pad.



INCUBATE the plate without inverting (pad-side down) at 35C for 24 hours.
COUNT THE COLONIES: Record the total number of colonies, and the number of coliform (red colonies) . Divide by 2.5 to yield the number of bacteria per 100 mL. According to national health standards for drinking water, the number of coliform/100 mL should not exceed 5.
Most chlorinated tap water will have no bacteria in 250 mL, as seen on the left membrane in the first of the nest set of pictures.
Enter your data in this sequence and format into your notebook, and then into the spreadsheet for the class table in the computer
Desk No. Initials Source in detail (water district) Coliform/100 mL

TOP: tap water and “filtered” water from the same home. Note that filters must be changed regularly, or else they become “nesting” places for bacteria.
MIDDLE: The two water samples shown are both heavily contaminated.
BOTTOM: For years, Williamsburg had problems with their water, as can be seen in the 8/14/00 sample. We even had a village official tell a student it was against the law for her bring in a sample of her own water to test…

grasp with fingers, then tongs:






Preparation of Wet Mount Slide

Preparation of Wet Mount Slide

Wet mount preparations are especially valuable for demonstrating motility in microorganisms. Fresh cultures must be used for maximum motility. No stain is employed since most stains kill the organisms (except vital stains). Therefore focusing is more difficult (see suggestions for focusing). If your microscope has dark field capabilities, this is ideal illumination for this purpose (use the “D” position on the sub-stage condenser dial).   Alternatively, a small disc of dark paper may be placed in the center of the condenser to approximate dark field optics.

Motility needs to be distinguished from Brownian motion which is due to molecular bombardment.  Brownian motion occurs in all microscopic bodies suspended in water and appears as a random shimmying-shaking. Motility will be in the form of cork-screw spiraling, movement in a given direction, or tumbling in place.

Wet mount preparations are also useful for giving clear images of fresh specimens under the microscope. Features which may be particulate, such as spores of fungi and ferns, and pollen grains may be best observed using this technique.

FOR CULTURES: Place 15 – 20 uL of the culture in the middle of the slide. (Here is a larger view of this image.)
FOR COLONIES: Place a small drop of dH2 O in the center of a slide. [For greater volume of sample, or for hanging drop preparations, use a depression slide for this preparation.]

CULTURES: The sample of liquid culture is place on the slide.
COLONIES: Sterilely transfer a tiny portion of a single colony to the drop with a loop and suspend (be certain to allow the loop to cool before picking up specimen). For solid specimens or dry spores, transfer a small portion of the specimen with a scalpel.


Lower a clean cover slip over the drop as though it were hinged at one side.


Finished preparation. Note that it is essentially transparent, making focusing difficult. (Ignore the handle of a drawer which appears in the background. It plays no role here…)



First focus with the 4x objective on the edge of the coverslip. It is easier to find and focus on than the nearly transparent suspension.


Find a bubble in the liquid suspension, and adjust the fine focus on the edge of the bubble.


Switch to the 10x objective, repeat the careful focusing.


Switch to the 40x objective, repeat the careful focusing. You should be able to discern bacteria at this power (magnification = 400x).

Apply oil and examine with the 100x oil immersion lens, again using the edge of the bubble as a focusing point. At 1000x, maximize the depth of field by narrowing the iris diaphragm, and adjust the focus so that most bacteria are in focus. (Because of the depth of the water, not all bacteria will be in focus at a given point.) Illustrate the types of motility you observe.
Clean up the slide with alcohol first (because it had live bacteria on it), followed by soap and water. Discard the cover slip.

Graph Construction by Hand

Graph Construction by Hand

While computer programs can conveniently construct graphs from data, hand construction of a graph remains an important means of analyzing and appreciating the value and patterns present in data.  Follow the following rules in order to produce a properly sized and accurately plotted graph.


Examine the data set and note the minimum and maximum values:
X-axis:     ordinate (independent or known variable):       time, added concentration, etc.
Y-axis:     abscissa (dependent or unknown variable):     what was measured: weight, A660 etc

(If the zero value of X or Y is important for your graph, it should be included in the limits.)


Count the number of squares available for the X and Y axes, leaving at least 3 square at the bottom and sides, and 9 squares at the top.  Graph-lined composition notebooks with 5 X 5 quad ruling allow for a graph of no more than 35 squares wide and 40 squares tall.


Assign values to the coordinates which meet the following requirements:

a.  They include the limits determined in step 1.
b.  They make an adequately large graph as large as the available space will accommodate.
c.  They do not exceed the space available on the page.

Divide the value of the range by the number of squares available along the given axis.  Round up so that the first significant figure of the result equals 1, 2, 5 or 10 units per square. For example, if you have Y axis range from 0.000 to 1.212, divide the 1.212 by the 40 squares available, which equals 0.0303/square.  This would be rounded up to 0.05/square.  Memorize the 1, 2, 5 and 10 values per square.  Other values will make plotting the data difficult, and it will cost you points when graded.  The quantity zero should often be the space most to the left and/or bottom.

Here is a page from a notebook illustrating the parameters for sizing a graph to a page.


Draw lines for the X-axis below and the Y-axis to the left of the selected open area on the graph paper.  Label each axis.  Mark off the selected regular values (often every 5 or 10 squares) with a small line corresponding to units/square selected in step 2.  Label each small line with its corresponding value.  (Do not label every square.)   Be certain to maintain linearity: all spaces must have equal value.


For the first point, locate the appropriate value along the X axis and then follow that line up until the appropriate value of Y is reached.  Double check that you have not shifted from the desired location, and make a small dot at the point.  Draw a small circle around the point, making it easier to see, but preserving the integrity of the point.  Repeat until all data have been entered.  Did you include the zero point if it is a significant data point?  Use squares to indicate a second data set, triangles the third, etc.


If the function you are graphing is linear, carefully connect the circles by lining a ruler up with the points and drawing a line between them.  (Do not violate the interior of the circles so that the value of the point will remain clear.)  Alternatively, if the function is non linear, you may either connect the circles or approximate the curve plots with a “best fit” curve.


Create a title which is meaningful and explicitly reflects the value of the experimental data you have graphed.  Place it in CAPITAL LETTERS as the title of the page.  Below the title, indicate from where the original data came with a cross reference.  Be certain that the axes are correctly labeled.  Label any significant break points or phases in the curve, briefly indicate their meaning, if known.

Bacterial Contamination of Milk: Pour Plate Assay

Bacterial Contamination of Milk: Pour Plate Assay

Pour Plate Technique for Bacterial Enumeration

Fresh food will typically have very low bacterial content, but as it is handled and stored, the bacterial concentration may increase dramatically. Pasteurized Grade A milk is required to have less than 20,000 bacteria/mL by standard plate count. Ground beef may contain up to 50 million bacteria/gm. Since the number of bacteria may vary by several orders of magnitude, samples of these foodstuffs must be diluted and several dilutions plated out in order to achieve the desired range of colonies per plate (50-500). Typically,  for pour plate technique, 1.0 mL of dilutions ranging from undiluted to 103 (or higher) should be plated.   (Greater dilution is necessary for highly contaminated samples.)  The following procedure is for that purpose:



    1. Milk to be tested. Have date of origin, if possible. Calculate age of material.
    2. Standard Plate count agar*
    3. Sterile dH2O in 4 repipets
    4. Clean sterile petri dishes

15 mL melted Plate Count Agar in:
sterile capped 16x150mm test tubes
45o C bath (deep enough to = agar
depth. Hot Block, or water bath)
paper towel
stainless steel spatula in test tube with 95% EtOH
sterile 16x150mm test tubes
0.1, 1.0, 2.0 & 10 mL pipets, sterile
colony counter with magnifying glass

dilute milk

Label two empty plates with your initials, the date, specimen (milk), aliquot volume (0.1 or 1.0 mL) and dilution factor (10^2).Prepare dilution blank: Add 9.9 mL sterile dH2O to a sterile 16x150mm test tube.


Dilute the milk: pipet 0.1 mL milk into above dilution tube (10^2 dilution), vortex to mix

Diluted sample is added directly to empty plate

Add the aliquots: Using a 2.0 mL pipet, pipet 0.1 mL into first plate, 1.0 into the second.


Pick up a tube of 15 mL melted plate count agar, cooled to 45C, (here in a hot block, a 45C water bath can be used.)


Add melted agar (dry off if maintained in hot water) to each plate in turn, swirl to mix completely. Plunge the emptied tube immediately into warm water before agar solidifies to ease cleansing.


When the agar is solid, invert the plate and incubate 35C for 48 hr.


Count colonies on the plates and calculate CFU per mL:
CFU on plate x dil’n factor (102) x aliquot factor (either 1/1.0 ml or 1/0.1 mL) = CFU/ mL milk
Example plate: 40 colonies on plate x 10^2 x 1/1.0 mL = 4000 CFU/mL of milk
Enter your results into the class table ( your initials, the milk manufacturer, its expiration date, CFU/mL)
* Standard Plate Count Agar: 5 g tryptone, 2.5 g yeast ext..1 g dextrose, 15 g agar, 1 L water
Suggested stations (at least two each? If enough repipets):

Empty plates station, wax pencil
sterile bags of empty plates
wax pencils
16×150 mm sterile capped test tubes (5 per student)
test tube racks to hold 16×150 tubes (one per student)

Milk Dilution Station:
sterile capped 16×150 mm test tubes
repipet with sterile dH2O, set for 9.9 mL
sterile pipets:
0.1, one per student
2.0, one per student

Pour Plate Station:
hot block with styrofoam cage, 45 C
wash tub with warm water in it.
16x150mm test tubes with 15 mL 45 C melted Standard Plate Count Agar (5 per student)
paper towel
Be sure to have enough tubes of SPCA (at least 3 tubes/person if they work in pairs on both experiments).

Accredited labs
advanced testing laboratories
4700 Smith
Standard methods agar

Plate count agar:
5 g tryptone
2.5 g Yeast est
1 g dextrose
15 g agar
1 L water (pH 7

suggested limit total viable
basic food microbiology book, Banwart, OSU Microbiology
100,000-50,000,000/ gm


Nutra Lab

Pour Plate Technique for Bacterial Enumeration

Pour Plate Technique for Bacterial Enumeration

RELATED PROTOCOLS: Bacterial Contamination of Milk and Meat , Yeast Plate Count

The pour plate technique can be used to determine the number of microbes/mL or microbes/gram in a specimen. It has the advantage of not requiring previously prepared plates, and is often used to assay bacterial contamination of foodstuffs. The principle steps are to

1) prepare/dilute the sample
2) place an aliquot of the diluted sample in an empty sterile plate
3) pour in 15 mL of melted agar which has been cooled to 45C, swirl to mix well
4) let cool undisturbed to solidify on a flat table top
5) invert and incubate to develop colonies.

Each colony represents a “colony forming unit” (CFU). For optimum accuracy of a count, the preferred range for total CFU/plate is between 30 to 300 colonies/plate.

One disadvantage of pour plates is that embedded colonies will be much smaller than those which happen to be on the surface, and must be carefully scored so that none are overlooked. Also, obligate aerobes may grow poorly if deeply imbedded in the agar.

15 mL sterile Plate Count Agar (PCA)*, in capped 16 x 150 mm test tubes, melted and cooled to 45C
Hot Block, 45C (or water bath), 3″ deep to equal agar depth
sterile capped 16 x 150 mm test tubes
0.1, 1.0 and 2.0 mL pipets, sterile
petri dishes, empty and sterile
colony counter with magnifying glass

1. Write out details of preparing and plating your specimen(s):
Construct a table in your notebook with a line for each plate: • the identity/source of the specimen (notebook entries should be detailed).
• the dilution of the specimen expected to contain between 30-300 CFU/0.1-1.0 mL and how you will prepare it
• the volume of diluted specimen you will plate (usually 0.1 to 1.0 mL)
Label the bottom of the empty, sterile plates your initials, seat number, date and the above data.


2. Dilute specimen to yield approximately 30 to 300 CFU per aliquot to be plated (from 1).

Diluted sample is added directly to empty plate

3. Inoculate labeled empty petri dish with the aliquot of diluted specimen (from 1)

Melted plate count agar, 45 C is added to sample and mixed

4. Pour 15 mL of melted Plate Count Agar (45C) into the inoculated petri dish.

5. Cover and mix thoroughly by gentle tilting and swirling the dish. Do not slop the agar over the edge of the petri dish.


6. Place on a flat surface undistrubed for about 10 minutes to allow the agar to completely gel. In this illustration, the agar is completely gelled and the surface is “smooth as glass.”


7. Invert and incubate at 37 C for 24-48 hours.


8. Count, record, calculate:
Count all colonies (note that the embedded colonies will be much smaller than those which happen to form on the surface). A magnifying colony counter can aid in counting small embedded colonies. Record the data. Calculate CFU/mL or CFU/g. Enter results in your table.

CFU/ mL = CFU/plate x dilution factor x 1/aliquot

On the plate shown, milk was diluted 1 to 100 (10 2), 1.0 mL of the dilution was plated and 40 colonies formed. Therefore the count per mL in the milk was:
40 colonies x 10^2 x 1/1 = 4 x 10^3/mL

* For 600 mL of NA + 1% glu: 9 g agar, 4.8 g nutrient broth, 6 g dextrose. Dissolve ingredients at 95C, repipet into 16 x 150 mm tubes, cap, autoclave, 15 lb, 15 min. Cool to 45C before using. Plate Count Agar may also be used.

Bacterial Contamination of Meat: Pour Plate Assay

Bacterial Contamination of Meat: Pour Plate Assay

Pour Plate Technique for Bacterial Enumeration
Bacterial Contamination of Milk, Pour Plate Assay

Ground beef for human consumption may legally contain up to (hard to believe) 50 million bacteria/gm. The number of bacteria in a given sample may vary by several orders of magnitude.  Therefore, when samples of these foodstuffs are assayed for bacterial count, several dilutions must be prepared over several orders of magnitude in order to achieve the desired range of colonies per plate (30-300). Typically, 1.0 mL of several dilutions from each of the dilutios ( undiluted to 103, or higher) should be plated.  Greater dilution is necessary for more highly contaminated samples. The following procedure is for that purpose:

Ground beef to be tested. Have date of origin, if possible. Calculate age of material.
Standard Plate Count Agar, sterile (SPCA) 15 mL in capped 16 x 150mm test tubes, melted, 45 C , three per student
Sterile dH2O in 4 repipets capable of adjustment to deliver 1 to 10 mL aliquots.
Clean sterile petri dishes, 3 per student.

wax pencil
sterile 16x150mm test tubes  (four per student)
stainless steel spatula in test tube with 95% EtOH
45o C  Hot Block (or water bath deep enough to equal agar depth.)   One per15 students
2.0 mL pipets, sterile , 3 per student
1.0 mL pipet, 1 per student
colony counter with magnifying glass




Label the bottoms of three empty plates with:
aliquot volume (1.0 mL each)
dilution factor (101, 102 or 103)
Prepare meat dilution blanks by labeling three 16 x 150mm sterile tubes with your initials and the exponents 1, 2 or 3 (dil’n factors 101, 102 or 103.)


Prepare dilution blanks:
Repipet 9.0 mL sterile dH2O into each of the three tubes.


Prepare a 10% suspension of meat in sterile water:
Weigh out 1.00 g ground beef sterilely into a fourth sterile capped 16x150mm test tube. (Record actual amount weighted out.)
(If you only weigh out 0.8 g of meat, suspend it in 8 mL water in the next several steps. I.e., a 10% suspension.)

Suspend the weighed meat with a vortex and spatula:
Add 3.0 mL sterile dH2O 2, vortex with sterile spatula inserted to suspend well. Be sure to hold the tube near the top to prevent spraying…)
After thorough suspension, add an additional 7.0 mL more sterile dH2O. Vortex to mix well again.


In the following steps, you will prepare the serial dilutions:
Prepare serial dilutions of the specimen in steps of 101, vortex after each dilution to mix completely. [Greater dilution factors may be achieved by using 0.1 mL into 9.9 mL in one or more of the serial dilutions, or plating 0.1 mL into the final petri dish.]:

For the first dilution, because of chunks of meat in suspension, use an inverted 2 mL pipet as instructed:
1) Use an inverted 2.0 mL pipet to deliver 1.0 mL of sample into first dilution tube = 101. (Invert so that meat particles do not clog up the pipet. Draw fluid to the 1.0 mark, deliver to the 0.0 mark.)

Add the diluted sample to the plate. (Then pour in the melted 45 C agar.)

2) Use a 2.0 mL pipet to deliver first 1.0 mL of 10dilution into the appropriately marked empty plate, THEN deliver the rest (1.0 mL) into the 10dilution tube, vortex


3) Use a fresh 2.0 mL pipet to deliver first 1.0 ml of 10into the appropriately marked empty plate, then deliver the rest (1.0 mL) into 10dilution tube, vortex.

4) Use a fresh 1.0 mL pipet, deliver 1.0 mL of the 10dilution into the appropriately marked empty plate

Melted plate count agar, 45 C is added to sample and mixed

Add 15 mL 45 C melted agar to each plate in turn, swirl well to mix completely. To ease cleaning, plunge the emptied agar tube immediately into warm water before agar solidifies

After the plate has completely solidified (be patient), invert plate and incubate 35 C for 48 hr.
Meanwhile, pour the contents of the initial meat suspension tube into the quart jar provided (do not pour down the stink, er…, I mean sink…)
The contents of the other dilution tubes may be washed down with hot soapy water.


Count the colonies on the plates and calculate CFU per mL.
Remember that colonies imbedded within the agar are quite small, while the colonies on the surface are quite large (see the image to the left).
Each counts as a colony when enumerating the total colonies formed.

Using which of the three dilution plates has between 30 and 300 colonies, calculate the CFU/g meat. (See formula below.)
Enter your results into the class table (your initials, the meat source, its expiration date, CFU /g


CFU/plate x meat suspension factor (10 mL/g) x dil’n factor x aliquot factor (1) = CFU/g meat

1  On the plate shown, 1 gram of meat was suspended in 10 mL, diluted 1 to 10, 1.0 mL of the suspension was plated and 293 colonies formed.  Therefore the count per gram in the meat was 293 x 10mL/g x 10 D.F. = 2.93 x 103/gram

If testing for E coli H7:O157, substitute MacConkey Broth for sterile water in this step. After dilutions are completed, place suspension in 37oC.