Objectives

Standards Addressed Biology I Standards
Physical Science Standards

Introduction and Background

Methods Safety            Sample Collection Sample Preservation & Storage  Bacterial Analysis  Calculations

Classroom Activity
Geographic Sources
Effects of Stormwater
Effects of Temperature
Effects of Sunlight
Effects of Salinity
Effects of Soil             Effects of Time

Resources and Links

Teacher Guide

Glossary

Bacteria can be difficult to sample and analyze.  As a result the acceptable combined precision and accuracy of the standard methods is as much as 20%.  Some of this uncertainty is due to large natural variations.  Since bacteria are particles that can adsorb onto surfaces and each other, they tend to be unevenly distributed through water bodies.  Their numbers can also change greatly over short time periods especially following rain events.  Thus, if you can afford it, it is best to take at least two samples and average the results.  Three is even better.  Sampling techniques can also be a source of variability as water needs to be collected without adding any of the sampler's bacteria.  Sampling containers also need to be sterile.  The analytical procedure is not difficult but sterile conditions need to be maintained - coughing and sneezing are a problem.  It is best to do these procedures under a laminar flow hood or in a clean room.

The containers and surfaces that will come into contact with your samples must be sterile.  If sample containers are to be reused, they must be autoclaved in order to ensure sterility.  Plastic containers, either polyethylene or polypropylene are generally prefered to glass due to their ability to withstand breakage.  Plastic bottles must be able to withstand 15 minutes in an autoclave at a temperature of 121^oC without melting.  Once this is completed, the bottles should be marked or taped to indicate sterility. The size of the container is determined by the size of the sample you wish to test. Typically, we use 100-mL samples.  If bacterial numbers are high, use less sample so that you end up with about 20 colonies on your plates.  To measure out lower volumes of samples, purchase sterile pipettes.

It is helpful to use a sampling pole if you cannot get within arm's reach of the water at your sampling site.  See Equipment for instructions on how to obtain a low-cost sampling pole.

Safety

Since you and your students could be potentially exposed to pathogenic bacteria, we recommend the following standard precautions which are similar to those required for general laboratory work..

Use latex gloves and goggles when handling samples in the lab and field.  At the end of the sampling and laboratory sessions have students clean their hands with antimicrobial soap and water.  Do not permit any food or drink in the lab or field.  Students should wear closed shoes and preferably a lab coat in the lab.

Sample Collection

1. If factory sterilized, sealed bottles are used, no preparation is necessary. We recommend using pre-sterilized, 100-mL plastic containers as listed on the Resources page.  Otherwise, sterilize as described above.  The following instructions assume that you are using the 100-mL plastic containers.

2. Prior to leaving the laboratory, ensure that you have your sampling pole, cooler with ice, a permanent marker to label your sampling bottles, paper towels, and the field sample data sheet.  For safety, we also recommend using latex gloves and goggles during sampling and a hand sanitizer after students return from field work.

3. When you arrive at the collection site, label the bottle and attach to your sampling pole, or, if the sample site is within arm's reach, hold the container by its lid, submerge the bottle and allow it to fill to the top. Try to avoid incorporating large pieces of debris, such as leaves and twigs.   Take sample container off the sampling pole and carefully close the top.  Do not allow your fingers to touch the rim of the bottle, as this will contaminate your sample.

Sampling

4.  Place the container upright wedged in the ice in the cooler.  The ice will slow the bacteria life cycle and preserve your sample.  Fill out the field data sheet.

5.  Once all student groups have properly stored and  filled their sample bottles, return to the lab and continue with the analysis procedure.

Sample Preservation and Storage

Hold the temperature of all stream pollution, drinking, and wastewater samples below 10^oC during a maximum transport time of 6 hours. Start the microbiological analysis of your water samples as soon after collection as possible to avoid unpredictable changes.  Keep the samples on ice or in a refrigerator until you are ready to analyze them.    

For most high school science classes, the samples will be analyzed immediately after collection.  If necessary, students may process samples the next day.   Record the length of time and temperature of storage of all samples and consider this information when interpreting data.  As described under Research Themes, your students could elect to explore the effects of time on bacterial concentrations.

Bacterial Analysis

1. Sterilize the forceps by heating in a bunsen burner flame for 10 seconds.  The metal should turn orange when hot.  Let forceps cool before using.  

   Sterilizing forceps

   Breaking ampoule

2. Snap open an ampoule of media (m-Coli Blue for Total Coliforms and E. Coli) with the blue plastic ampoule breaker.  We recommend resting the bottom of  the ampoule on the top of your lab bench while you do this.  Be careful to keep fingers away from the ampoule's seal. 

   Media Ampoule

3. Take the cover off of a sterile petri dish containing a dry filter pad.  Put the cover onto the desk so that the top of the inside of the cover lies face up.  Pour the media onto the dry pad.

   Petri Dish

    

   Strive to spread the media as evenly as possible.  Put the cover back on and set aside to let the media soak into the pad.

4. Remove a sterile filter funnel from the plastic bag.  Discard red cap on bottom and insert filter funnel into manifold.

   Filter setup

    

5. Open sample container and hold in one hand.  Use the other hand to remove the top cap on the filter funnel.  Pour 100 mL of water into the filter funnel using the volume markings on the funnel as a guide.  Place the cap back on the funnel leaving a small air gap.

   Filter funnel w/ water

6. Connect the vacuum and completely filter the sample under a vacuum of no greater than 25 mm Hg.

7. Remove the cap and pour 10 to 20 mL of sterile dilution water onto the filter trying to wash down the sides of the filter funnel.  Filter this water through completely.  Allow the vacuum to remain engaged for at least one minute after all of the water appears to have passed through the filter.

8. Snap the upper cup off of filter funnel, leaving the filter behind on the filtration assembly.

   Snapping filter cup off

9. Remove the cover (and place on desk top as before) to the petri dish containing the media-soaked filter pad.  Pour off any excess media. 

10. Using sterile forceps, transfer the filter onto the media-soaked filter pad.  Be careful to make sure that the filter makes complete contact with the filter pad so that the bacteria on the filter come in contact with the media.  The best technique is to roll the filter onto the pad.

    Transfer filter to pad

    Roll the filter onto the pad

11. Put the cover back on the petri dish.

12. Using a permanent marker, label the petri dish with the sample name.

    Incubator

13. Place the petri dish in the incubator, upside down, for 18 to  24 hours at 35^oC.  Organisms from non-disinfected sources produce a red or blue metallic sheen at 16 to 18 hours.  The sheen may fade after 24 to 30 hours. (NOTE:  The dishes are placed upside down to prevent condensation from falling onto the filters causing the colonies to diffuse off the filter.  Excess liquids on the filter cause a smeared appearance.)

    Place dishes upside down in incubator

14. Count the red and blue colonies using magnification of 10 to 15X and a cool white fluorescent light source directed as nearly perpendicular as possible to the plane of the filter to bring out the sheen.  Optimally, use a binocular wide field dissecting microscope.  Do not use a microscope illuminator with an incandescent source.  Record the results on the field data sheet.

    Petri dish with blue and red colonies after incubation

The typical coliform colony has a pink to dark red color with a metallic surface sheen.  The E.coli colonies register as blue with a metallic sheen.  The sheen area may vary in size from a small pinhead to complete coverage of the colony surface. Count sheen colonies only. Colonies that lack sheen may be pink, red, white, or colorless and are considered to be noncoliforms.

Ideally, your 100-mL sample should yield no more than 20 to 50 colonies.  If your samples consistently have higher bacteria levels, try using a smaller sample volume, such as 50, 20, 10, 5 or even 1 mL.  You will have to transfer these smaller volumes using a sterile pipette. Disposable sterile pipettes are available through Fisher Scientific and other general scientific suppliers.

If you are faced with a plate with a lot of colonies or part of the filter did not produce colonies well, you can elect to count the number of colonies in each of ten randomly selected squares (the filters have a grid with 146 squares on it).  The field data sheet has a table for recording these results and details on converting the results into concentrations of CFU per 100 mL.