Welcome page

Timetable

Background

• Local info
• Water issues
• Projects
• Characterization
• Sample article (pdf)
• How to use Bergey's Manuals

Methods

• Rules
• Media/culture
• Aseptic technique
• Staining/interpretation
• Isolation strategies
• Assays/characterization
• Microscopy
• Cultural characteristics

Media preparation
and training

Projects and Rationale

Our projects will focus upon microbiological analysis of water samples from the proposed Lakes Region Recreational Area. In Project 1 we will isolate and characterize selected bacterial species that are commonly found in the pond systems. Project 1will be initiated as a team project and then completed by individuals who take responsibility for characterizing some number of bacterial isolates. Each individual will write a monograph on one selected species, according to the guidelines listed in the Characterization resource. Monographs will incorporated into a Compendium of bacterial species inhabiting the lakes in the Eggstain region. In Project 2 we will conduct a microbiological analysis of water samples from each of the area's individual ponds, including estimates of total bacterial density, concentration of coliform bacteria, and concentration of thermotolerant coliforms, which are indicative of fecal contamination. The work in project 2 will be divided among individual teams, the findings pooled, and each team willl present findings, interpretation, and recommendations in a poster.

Project 1:  Qualitative analysis of bacterial populations in the Lakes Recreational Area

Rationale

Project 1 is part of a (hypothetical) larger initiative to census the entire biota of the area. We will attempt to isolate pure cultures from two kinds of agar plates. Tryptic soy agar (TSA) is a rich medium that supports the growth of fastidious organisms that may not grow at at all on less nutrient-rich agars such as nutrient agar. It also promotes growth of very aggressive species that may crowd out slower growing bacteria. Reasoner's 2A (R2A) agar is formulated to prevent fast-growing bacteria from taking over the agar surface, allowing slower-growing bacteria to compete for nutrients. Between these two types of agar, we hope to obtain a diverse collection of bacterial isolates that is at least somewhat representative of the flora in our lake system.

We will not be able to isolate every bacterial species that inhabits these waters. In fact, it has been estimated that less that 10% of resident bacteria in the environment can be cultured in a laboratory. Each team should nevertheless attempt to isolate as many unique species as possible and preserve the cultures in agar slant tubes for analysis. Each team is to record information for each isolate collected (see below). Ideally we would attempt to characterize each isolate to the species level, but that ambitious goal is not practical given the time frame of the study. Instead, isolates are to be divided among team members and each team member is to characterize at least one unique isolate. A general strategy for identifying isolates is outlined in chapters 1 and 2 of Bergey's Manual of Determinative Bacteriology. Identification should proceed by working from broader to narrower categories, using the least number of necessary tests.

Processing samples and obtaining isolates

Project 1 will begin the first week with lab tour and training sessions in which you will learn your way around the laboratory, how to prepare media, use the autoclave, learn aseptic technique, etc. You will also learn how to prepare dilutions and spread agar plates as outlined in Appendix C: Spread plate procedures. Each team will also prepare media for culturing bacteria. At the start of the second week you will process samples and incubate your spread plates. At the second lab session you will

• select one R2A plate from each sample from which to obtain total bacterial density (heterotrophic plate count) for the lake the sample represents (see Total bacterial density, below, for why we obtain our information from R2A plates
• examine your R2A and TSA plates and identify unique species based upon colony characteristics
• obtain a culture from each unique colony type by transferring colony material to a fresh TSA plate

At the time you process your samples (which takes relatively little time), you will learn and practice a dilution streak method for obtaining separate colonies and ensuring that you have pure isolates. As soon as possible after obtaining sufficient growth (usually 24-48h at 30 degrees C) you will need to compare cultures and eliminate redundant isolates. Once you have a pure, unique isolate you will transfer to culture to a TSA agar slant tube.

Total bacterial density

In order to complete your analysis and poster presentation on pollution patterns in the lake system you will need an estimate of total bacterial density for each pond in our study. Each team is to report the following information to the instructor for each pond water sample.

• colony count for the selected plate
• dilution used for the selected plate
• CFU/ml calculation
• e.g., for pond A 0.1 ml sample resulted in 76 colonies on 1/100 dilution plate; final concentration is 76,000 CFU/ml

Rationale
For bacterial density we usse procedures outlined in the Standard Methods for the Examination of Water and Wastewater, 20th Edition, Baltimore: American Public Health Association, American Water Works Association, Water Environment Federation, 1998. Page 9-1 explains the rationale. In brief, the heterotrophic plate count provides an estimate of total number of viable bacteria per unit sample volume. High bacterial counts can be indicative of contamination with human or animal waste. We will use the spread plate method, described on pages 9-3, 9-35-36, and 9-38 to 9-40t 1. As recommended in the Standard Methods, we will obtain heterotrophic plate counts from the R2A plates and not from the TSA plates. Total bacterial density for each pond is to be determined from a single R2A plate that gives the best distribution of colonies (numbering between 30 and 300).

Information on isolates to be recorded by each team

Note that much of this information requires microscopic observation of bacteria. We will conduct a short workshop on microscopy, including the oil immersion technique, at the time that you begin to obtain isolates. The workshop will include observations of Gram stains and preparation/observation of wet mounts.

• Source of isolate, including pond ID, agar type, dilution, and date processed
• Colony description
• Gram stain result (+), (-) or variable
• Cell type:  straight rods, curved rods, spiral rods, irregular rods, cocci, other
  
• Cell size:  rod or cocci diameter
  
• Growth patterns:  (rods) solitary cells, chains, or (cocci) solitary, irregular clusters, chains, pairs, tetrads
• Is there apparent motility (observation of wet mounts)
• Relationship to oxygen:  obligate aerobe, facultative anaerobe, microaerophilic; we will not find obligately anerobic bacteria, which cannot grow in the presence of oxygen

Characterizing selected isolates

The online manual describes the available assays for characterizing isolates, instructions will be posted in the laboratory, and instructors will be available to assist with the methodology.

Your choice of assays to conduct will depend on the characteristics of the isolate. For example, any rod shaped cell type should be checked for motility. Any Gram (-) isolate should be subject to an oxidase test. Gram (+) rods should be examined for the presence/absence of spores. Characterization to the species level will likely require metabolic tests. For example, Gram (-) facultatively anaerobic oxidase negative rods should be tested with a combination of indole, methyl red, Voges-Proskauer, phenol red, and/or decarboxylation assays. These assays may or may not be useful for characterizing other types of isolates. Materials for assays not described in the manual may be available and/or purchased if given sufficient notice.

One principle that applies to microbiology as much as to any subject is that "absence of evidence is not evidence of absence." For example, many cells quickly lose their Gram stain properties as they age. If you see clear evidence of Gram (+) cells in a pure culture that has been carefully and properly stained, you have a Gram (+) species. However, a Gram (-) finding should be accepted tentatively unless experience suggests it is clearly not Gram (+). Similarly, many rods lose their flagella on solid media or even in broth under relatively uncrowded conditions. The uninitiated should assume that rod-shaped cells isolated from the environment are indeed motile, unless further testing produces inconsistent results that suggest the assumption of motility must be incorrect.There is no substitute for experience when characterizing an isolate, therefore your instructors are prepared to assist you in making decisions.

Project 1 culminates in an individually written monograph, described separately (see Characterization).

Project 2: Microbiological examination of water samples and analysis of coliform contamination patterns in the Lakes Recreational Area

Reference: Standard Methods for the Examination of Water and Wastewater (22nd ed.), Washington, D.C.: American Public Health Association, 2012

Project 2 addresses a potential pollution problem in the (hypothetical) Lakes Region, namely high bacterial counts and presence of fecal coliform bacteria, suggesting contamination with human or animal waste. This problem jeopardizes the entire revitalization project, therefore it is essential that the extent, patterns, and potential source(s) of contamination be identified. To complete the project each team will analyze several samples, each taken from single lake in the system. For each sample, representing an individual pond, we will estimate (1) total coliform concentration, and (2) concentration of thermotolerant coliforms (coliforms of probable fecal origin).

All procedures will be conducted as described in the Standard Methods. Each team will receive a packet of copies of selected pages, including the rationale behind microbiological examination of water and the individual tests, details of the individual methodologies, and the rationale behind data analysis and interpretation. A summary of the methodology that we will employ is provided below. Supplies and media needed for this project are described in "Media needed" under Media preparation and training, side menu.

Rationale

It is not practical to directly analyze water for the presence of pathogenic bacteria, parasitic protozoa, and related microorganisms. Giardia, Cryptosporidium, Shigella, Salmonella, Escherichia coli 0157:H7, and the viruses Hepatitis A, Coxsackie A and B, and Norwalk viruses all cause gasteroenteritis, and often produce further serious complications particularly in the infirm and in individuals with compromised immune systems. Leptospirosis, caused by bacteria of genus Leptospira, is not an enteric infection but it does produce high fevers, jaundice, and possible kidney damage or meningitis. Occurrence of these pathogenic organisms is nearly always traced to contamination with fecal wastes.

It is imperative that you read and understand the concept of coliform bacteria as indicator organisms. The EPA offers a very good introduction to the use of indicator organisms to test for fecal contamination of waters. Bacteria that we call coliform are members of Family Enterobacteriaceae that inhabit the lower intestines of warm-blooded animals. They are named for the principal coliform species found in the intestines of mammals, namely Escherichia coli. They are usually not harmful themselves, but they indicate the possible presence of pathogenic (disease-causing) bacteria, viruses, and protists that also live in human and animal digestive systems.

The Environmental Protection Agency (EPA) has identified members of the coliform group (especially Escherichia coli), fecal streptococci and a subset of coliform bacteria called thermotolerant coliforms (formerly called fecal coliforms) as indicators of the suitability of water for "domestic, industrial, or other uses." Fecal streptococci are more closely correlated with disease risk than are thermotolerant coliforms, however many communities, including ours, continue to estimate densities of thermotolerant coliforms to assess water quality. To assess the quality of waters in the Lakes region we will employ the multiple-tube fermentation technique for total coliforms and for thermotolerant coliforms.

Estimating bacterial densities using the MTF technique

The full rationale behind the Multiple-Tube Fermentation (MTF) technique is given in the introduction on page 9-1 of the Standard Methods. The introduction to the multiple-tube fermentation technique for members of the coliform group starts on page 9-65 of the Standard Methods, followed by a description of the methodology in sections 9221A and B. To conduct the assay for total coliforms we will follow the flow chart in figure 9221:1, page 9-68.

The MTF method employs broth tubes with Durham inserts. These tubes are filled, capped, sterilized, and cooled to room temperature in advance of the day the samples are processed. A Durham insert is simply a small inverted culture tube that is placed into the larger broth tube prior to capping it. Air in the inserts will be expelled during steam sterilization, therefore Durham tubes are self-filling. Media must be sterilized within an hour or so after mixing them, otherwise they will grow bacteria and be ruined.

To conduct an MTF assay on a sample we will inoculate 5 tubes of broth with 10 ml sample, 5 tubes with 1 ml sample, and 5 tubes with 0.1 ml sample. We will conduct both assays on each sample. For the total coliform assay we will use lauryl tryptose broth and for thermotolerant bacteria we will use EC medium. To estimate bacterial density we will use table 9221.IV (MPN Index for five tubes per dilution, volumes 10 ml, 1.0 ml, and 0.1 ml, respectively).

We will use 18 mm tubes and closures for the 10 ml water samples and 13 mm tubes and closures for the 1 ml and 0.1 ml samples. To cover the Durham inserts and hold the necessary volume of sample the starting volume of broth in each 18 mm tube should be 10 ml, so that the final volume will be 20 ml after adding a 10 ml water sample. Each 13 mm tube should contain 5 ml broth before adding a water sample.

The tubes to receive 10 ml samples and 1 ml samples will require concentrated broth so that after a water sample is added, diluting the broth in the tube, the final concentration will be 1x (normal full strength). An easy way to plan the preparation is to base the amount of powdered media you weigh out on the final volume in the tubes, then mix that amount into the initial volume. For example, to prepare 20 18x150 mm MTF tubes, each to start off with 10 ml medium and receive a 10 ml water sample, multiply the 20 ml final volume by 20 tubes, add 10% to account for waste, and weigh out the corresponding amount of powder (or in this case simply double the amount of prepared medium per liter). Mix the powder into a volume 20 x 10 ml plus 10% and then distribute 10 ml into each tube.

On the day samples are obtained, each team will receive four water samples in 500 ml bottles. Each sample will be from a different pond in the area, labeled with the pond system number (see map) and a letter from A to D. For example, the sample from system 2 pond B to be used for MTF incubations will be labeled "2B"). Recall that in each system the ponds are labeled A-D so that pond A (upstream) drains into B, which drains into C, which in turn drains into D (downstream). Teams will aseptically add the pre-determined volume of sample to each tube and then incubate at 35 ± 0.5°C (total coliforms) or 44.5 ± 0.2°C (thermotolerant bacteria). A tube is considered to be positive if gas appears in the Durham insert in the form of a bubble.

For thermotolerant coliforms, if there is no gas after 24 h incubation, a tube is recorded as negative for coliform presence, and the assay is completed. For total coliforms we check again after 48 h incubation, after which a tube without gas is considered to be negative. We conduct a confirmed test on all lauryl tryptose tubes showing positive for the presumptive test.

Confirmed tests for total coliform bacteria. For total coliforms we will follow each positive presumptive test with a confirmed test (page 9-67). Inoculate a brilliant green lactose bile broth tube with a loopful of material from each tube showing positive for gas production. Of course you won't know in advance how many positives you will have, so please plan to prepare the bile tubes as needed. Bacteria in the original tubes will remain viable for at least a week. Sterilize 5 ml broth in each 13 x 100 mm culture tube with Durham insert and stainless steel closure. We will not conduct completed tests because we do not need quality control data and because we are conducting a simultaneous assay for thermotolerant coliforms. Please report to the instructor all data for all of the tests, but for your final report (in poster format) base your estimates of total coliform densities on the results of the confirmed tests.

Appendix A: Elevations of selected lakes in the Lakes Region

We will examine samples from lakes throughout the region, including lakes that are not part of the redevelopment project. An objective is to obtain a big picture that reveals pollution patterns, if such patterns exist. The CFFCA harvested all remaining fish and treated each lake with a piscicide prior to abandoning the lakes at least six months before the start of this study.

Table 1. Elevations of lakes to be examined. Elevation refers to height in feet above sea level of the lake surface. Average depth of these lakes is less than fifty feet.

Appendix B: Spread plate procedures

Dilution procedure

You can choose to do each 1:10 dilution either by diluting 0.1 ml into 0.9 ml of sterile deionized water, or by diluting 1.0 ml into 9.0 ml. The former is somewhat faster, the latter makes accurate pipetting somewhat easier.

You will be given a rack of sterile 20 X 150 mm culture tubes and a bottle of sterile water. If you are going to use 9 ml dilution tubes, use a 10 ml pipette to put 9 ml into each of the needed dilution tubes. As long as you maintain sterility, you can use the same 10 ml pipette to fill all of your dilution tubes. If you choose to use 0.9 ml dilution tubes, you can use a 1.0 ml pipette to add 0.9 ml to each tube. For more efficient distribution, you can use a 5 ml pipette; fill it and then run 0.9 ml from that pipette into each of 5 tubes. You can reuse the same pipette for the next 5 tubes, etc, assuming that you have maintained sterility.

Note: This repeated use of the same pipette applies only to situations in which you are pipetting the same solution into tubes that are empty, clean, and sterile. Never use the same pipette to pipette 2 different solutions or 2 different dilutions of the same sample.

Sterility precautions: There are lots of bacteria floating in the air around you. To minimize the possibility that those bacteria might contaminate your samples, each vessel is opened for the minimum time necessary to pipette in or out, and pipettes are kept out in the air for the minimum time necessary. Handle the pipettes only by their back end, so you don't contaminate the portion of the pipette that goes into the tubes. For each pipetting:

1. Take the pipette out of the can and fit it snugly into the seal of the pipette pump, in a position so you can see the numbers on the pipette. (The following instructions assume that you are right-handed. If you are left-handed, reverse the instructions.)
2. Holding the pipette pump (with pipette firmly attached) in your right hand, with your thumb on the control wheel, and holding the tube that you are pipetting out of in your left hand, lift the cap off of that tube, using the little finger of the right hand to hold the tube cap
3. With the tip of the pipette resting on the bottom of the tube, fill the pipette to the desired line, by rolling the thumb wheel downward. It's easiest to fill it to a little above the desired line, and then move the meniscus down to the desired line by gently rolling the thumb wheel upward.
4. Immediately put the cap back on that tube and the tube back in the rack.
5. In the same way, lift the tube that you are pipetting into with your left hand, and remove its cap with your right little finger.
6. Place the pipette into that tube so its tip is just above the surface of the water in that tube.
7. Expel the liquid from the pipette by rolling the thumb wheel upward, so the liquid in the pipette goes directly into the liquid in the tube.
8. Lift the pipette out and immediately replace the cap on the tube.
9. Thoroughly mix the contents of that tube. The best way to do that is to use a Vortex Genie. We'll demonstrate how to do that. If no Vortex is available, you can mix by swirling the tube (for 1 ml volumes) or flicking it with a finger (for 10 ml volumes). The latter is less reliable, so if you plan to use 10 ml volumes, be sure that a Vortex is available
10. Detach the pipette from the pipette pump and put the pipette gently, tip downwards, into a dirty pipette bucket that is half full of water. You are now ready to make the next dilution.

Organizing tubes

The tubes should be labeled with marking pen, so you can keep track of which is which. For instance, if your samples are labeled A, B, C, and D, your dilution tubes for sample A would be labeled A -1, A -2, A -3, and A -4, where -1 refers to the 10-1 dilution, etc. To keep track of where you are, it's best to set up the tubes so there is an empty space in the rack next to the tubes that you are going to dilute into, so you can move each tube over one space after it has been diluted into.

Using pipette pumps

You must never pipette by mouth. The pipette pumps that we use are color coded: Green for 5 or 10 ml pipettes, Blue for 1 ml pipettes, Yellow for 0.2 ml pipettes. The pumps are a little tricky to use, so you should practice using them by pipetting water several times, and checking that you can control how much you pipette out. It's very easy to suck too hard and pull the liquid all the way up into the pump, particularly with the smaller volume (yellow) pumps. Turn the thumb wheel very gently at first, until you have a feel for how much pressure is needed to suck the liquid in. When filling a pipette, always let the tip rest on the bottom of the vessel that you are pipetting out of. That prevents you from raising the tip enough to get above the surface and start sucking air. When that happens, any liquid that was already in the pipette will fly up into the pump. If you do get liquid into a pump, don't try to use that pump any more that day, and report it to an instructor so we can clean the pump. For pipetting 0.1 ml aliquots, you can either use 0.2 ml pipettes, filled to the 0.1 ml line, or 1 ml pipettes, filled to the 0.9 ml line. The 0.2 ml pipettes are more precise, but the yellow pumps are more sensitive and harder to control.

All of the pipettes that we will be using are "blowout pipettes". That is, you fill the pipette to the desired line and then blow all of the contents out of the pipette to deliver the correct volume. (This is distinguished from "difference pipettes", where you fill it to one line and then eject a measured amount by moving the meniscus down to another line. Our pipettes can also be used for difference pipetting, as discussed for the 5 ml pipettes above.) In order to be able to blow out the entire contents, before starting to fill the pipette, you have to raise the piston of the pump upward a few millimeters, by rolling the thumb wheel downward. That gives the piston a few more millimeters to travel when blowing out the contents, so instead of just moving the meniscus to the tip of the pipette, where a drop might stick in the pipette, it drives even the last drop out of the pipette.

Just before you start diluting a sample, shake the bottle containing the sample, to mix it so the bacteria are uniformly distributed in the sample. Pipette from that sample into the first dilution tube, mix that, and then, using a fresh pipette, dilute from that tube into the next dilution tube, and continue through 4 dilution tubes for each sample.

Plating

0.1 ml aliquots of each undiluted sample, and each of the dilution tubes, are plated onto each of two types of plates, R2A and TSA. For maximum efficiency, you can use a single pipette to deliver both aliquots from a particular tube. For instance, fill a 0.2 ml pipette to the top line. Deliver 0.1 ml of that to the R2A plate by difference pipetting, touch the tip of the pipette to the plate, to be sure that there isn't a partly extruded drop still clinging to the pipette. Then deliver the remaining 0.1 ml to the TSA plate, again touching the tip to the plate to be sure that the last drop is plated. Flame the spreader before spreading each plate.