Protocols: Membrane Filtration Laboratory Analysis

Recommended methods for measuring fecal E. coli are:

1. Membrane filtration and plating on BBL MI agar, mColiBlue, BioRad Rapid E. coli 2 media, or Chromocult

● Works well for all sample types

● See the manufacturer’s instructions for use and correct identification of E. coli colonies

2. IDEXX Quantitray and Colilert media

● Not recommended for swabs

Important note on microbiological assays:

Be sure to use 100 mls of the sterile water or PBS used for washing items or making dilutions as a negative control in each assay. This is especially important if you are purchasing bottled water from a vendor, rather than using lab-prepared filtered or autoclaved water, as bottled water is not always sterile.

Membrane filtration protocol

A. Preparation

1. Wipe down bench or hood with 10% bleach followed by 70% ethanol.

2. Assemble the filter equipment, making sure all filter membrane holders have been autoclaved

3. Attach vacuum tubing to side arm of 1 liter flask and the vacuum source.

4. Attach tubing from output on side of manifold or vacuum flask to mouth of filter flask.

5. Insert filter base into mouth of manifold.

6. Prepare alcohol burner with lighter.

7. Prepare a small beaker with 100% ethanol for sterilizing the tips of the forceps. The ethanol should be 2-3 cm deep, just enough to cover the tips of the forceps when they are resting in the beaker. NOTE: The alcohol burner and ethanol should be on the same side as your dominant hand for easy forceps sterilization.

8. If you are using mColiBlue, prepare the plates then label the bottom of the plates with the date, dilution, sample ID, and initials. Be sure to include a plate for the negative control.

B. Sample Processing:

Materials and Equipment:

● Petri dishes, purchased or prepared

● 10-20 ml 100% ethanol in a small beaker (50-100 ml)

● Distilled or deionized water

● Sterile 1 x PBS

● 70% ethanol

● 10% bleach

● 10 ml serological pipets

● Pipet Aid or bulb

● Vacuum manifold or vacuum flask and tubing

● Vacuum pump

● Autoclaved membrane filter holder and vacuum funnel

● Sterile filters, mixed cellulose esters, 0.45um pore size, white gridded, 47mm diameter

● Flat blade forceps

● 1 liter side-arm flask

● Alcohol burner

● Lighter

1. Flame forceps for ~5 seconds to sterilize. Take care to hold the forceps horizontally to avoid burning your hand.

2. Remove a sterile filter from the packaging with sterile forceps.

3. Remove the filter holder and place the filter on the filter base, grid side up. Affix filter holder to the base.

4. Pour 10 mL sterile PBS on the filter.

5. Turn on vacuum, open the valve and close the manifold valves.

6. Pour another 10 mL of PBS on the filter and vacuum it through.

7. Carefully, remove membrane filter from filter base with sterile forceps, avoiding contact with the center of the membrane.

8. Place the filter, gridded side up, onto the plate labeled “Negative Control”. By rolling the filter onto the plate, you can avoid the formation of bubbles between the membrane and the agar surface, which can invalidate your results.

9. Replace the lid of the Petri plate.

10. Repeat steps 2 to 5

11. Add a minimum of 10 mL and up to 100 ml of liquid containing the highest dilution of the sample to the filter. Note: always start with highest dilution (lowest concentration) of sample to avoid introducing significant contamination from higher concentrations. If the test volume is 1ml, add 9mL PBS and spike the PBS with the 1mL sample aliquot (the total volume filtered is 10mL). This ensures that the solution is dispersed evenly around the filter surface.

12. Open the manifold valve and vacuum the liquid through the filter.

13. Use a 10 ml serological pipet to rinse the sides of the filter cup with 10 ml PBS.

14. Close valve on manifold and remove filter cup.

15. Flame forceps for ~5 seconds to sterilize.

16. Remove filter from base using sterile forceps, taking care not to disturb inner area of filter.

17. Place the filter onto a plate labeled with the Sample ID, date, dilution, and your initials. Take care to avoid the formation of bubbles between the filter and the agar.

18. Replace the plate lid.

19. Using the same filter holder, repeat steps 10 to 18 for the other dilutions of the sample being tested, going from most dilute to least dilute.

20. For each new sample you will need to re-sterilize gloves with alcohol and use a new filter holder.

21. Finish by processing the positive control (optional).

22. Invert the Petri plates, unless you are using mColiBlue, in which case plates should not be inverted to prevent the broth media from leaking and the plate drying.

23. Incubate the plates in a box at 37°C for 20 to 24 hours (according to media manufacturer’s guidelines). If using mColiBlue, incubate plates in box to avoid desiccation. Be sure not to close the lid tightly, it should just sit on top of the box so that oxygen can still circulate.

24. Record the date and time that the sample was placed in the incubator and your name (Lab Operator) on the laboratory form.

Recommended dilutions to be plated by Sample type:

C. Counting and recording colonies:

• • 1. Retrieve the laboratory form for your sample.

    2. Check the box of the concentration of sample tested.

    3. Retrieve the incubated samples and record the date and time the samples were removed from the incubator on the Laboratory.

    4. Refer to the manufacturer’s instructions for your media for proper identification of E. coli colonies.

a. Valid Reading: Any membrane filtration plate with colonies greater than 0 and less than or equal to 200

b. If there are > 200 colonies, record the results as 999 for “too numerous to count (TNTC)”.

c. If individual E. coli colonies cannot be clearly distinguished from background growth or dirt on the filter, record the result as 998 “too dirty to count” (TDTC).

d. If any E. coli colonies are found on the Negative Control plate, indicate the results next to “Negative Control”. Record 0 for no colonies.