Medical Microbiology

Pure Culture Techniques

In this first lab, you will be learning some very fundamental and important techniques. As is the case with most things, shorts cuts usually get you in trouble. This is especially true in Microbiology. The techniques you will be learning tonight, if mastered correctly, will make your life and learning experience in Microbiology much easier, if you don’t pay attention and practice these techniques incorrectly, well then……? Today you will be learning the following techniques:
1. Streak plate method for colony isolation
2. Aseptic transfer techniques
First the streak plate methods. In Microbiology it is often necessary to isolate pure colonies of one bacterium from mixed cultures, or to check the purity of cultures. You can do this by “spreading” the bacteria on a nutrient growth media in a specific manner as demonstrated in the lab tonight. In this lab we will be learning two methods to accomplish this task, the quadrant streak and the T streak.

1. Quadrant streak

2. T streak
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Lab exercise:
Following the demonstration of both of these isolation techniques, it is your job to see how well you can repeat the procedure, and get isolated colonies of bacteria. A colony of bacteria is nothing more than a visible mass of bacteria that has grown on a plate as a “descendent” of one bacterium that was deposited at that spot by the streaking technique. Isolated colonies, as the name implies, are masses of bacteria that are clearly separated from one another on the plate. Bacterial colonies that have a different shape, size, or color are generally different bacteria. In this exercise you are streaking only one bacterium, so all the colonies should look pretty much the same. You will need the following: 2 tryptic soy agar (TSA) plates Broth culture of Escherichia coli

After you have streaked your plates, they will be incubated at 37°C. The plates MUST be labeled and incubated in an inverted position. Next lab we will examine your technique!

Aseptic Transfer Technique

The term aseptic transfer refers to a set of procedures that are used to minimize the chances that a culture you are working with will become contaminated by organisms from the environment. In the microbiology lab this is a technique that you must learn, practice, and perform with the greatest of care. EVERY time that you open a culture of bacteria to perform any manipulation you must carry out this set of procedures. The procedure is not hard, but does follow a specific routine of manipulation of tubes and inoculation devices, loops or needles. One thing that will really help when you do transfers is that your work area is well organized, and not cluttered with other materials. In other words, get things set up in advance before you do transfers!

In the course of this semester in this lab you will be doing hundreds of transfers from a variety of media, both liquid and solid. The technique that will be demonstrated is exactly the same for ANY media. At first these techniques may seem a little awkward, but with practice these techniques will become second nature to you. Please remember to always follow the steps in the procedure EXACTLY, taking short cuts, or sloppy technique can really get you into trouble in this class. Here is summary of the technique:
1. Sterilize your loop until the entire thin length of the loop, not just the end, is red hot in the burner flame.
2. Allow the loop to cool for a few seconds.
3. Pick up the tube from which you want to transfer bacteria. Remove the cap, but KEEP THE CAP IN YOUR FINGERS, DO NOT PLACE THE CAP ON THE BENCH! Briefly flame the mouth of the tube, and then put your sterile loop into the culture tube to obtain a sample of bacteria. Remove the loop, re-flame the mouth of the tube, and replace the cap. Put the tube back in the rack.
4. Pick up the tube into which you want to transfer (inoculate) the sample of bacteria. Remove the cap as before, flame the mouth of the tube, inoculate the media, remove the loop, re-flame the mouth of the tube, and replace the cap. Put the tube into the rack.
5. Sterilize your loop by flaming the entire loop, not just the tip, to red hot in the burner flame.

That’s the procedure. It is the same for any type of media or culture. It is important to remember that this procedure is always followed when ever you transfer bacteria out of a tube. If you are making a slide for staining these procedures are followed! NO EXCEPTIONS! In microbiology one thing that you definitely don’t want are contaminated cultures. If you follow the procedures demonstrated then you should have few problems with unwanted bacteria from the environment.

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LAB EXERCISE

For your transfers you will be working with the bacteria E. coli growing in a liquid media (broth), and on solid media (slant). Each table will have a can with this organism; you will have to share with your tablemates. You will need the following: 2 tubes of tryptic soy broth (TSB) 2 tubes of tryptic soy agar slants (TSA)

You will do the following transfers:
1. E. coli on a slant to another slant
2. E. coli on a slant to a broth
3. E. coli in broth to a slant
4. E. coli in broth to another broth

Incubate all tubes in a can at 37°C. Please remember to make a tape label for all your tubes. The label should have your name, the name of the bacteria, the date and the type of media that you are using. Also put your name on the can with tape so you will be able to find it easily when we examine the results during the next lab.

Simple Stains

As we discussed in lecture, because bacteria are so exceedingly small, and at the very limit of the resolution of light microscopes, it is necessary to stain (color) the bacteria to in order to visualize them. The first of the staining procedures you will be learning is the simple stain. Such staining procedures use only ONE dye, and allow you to determine such things as the shape, size, and arrangement of bacteria. The stains used in this procedure are CATIONIC dyes, that is, these dye carry a positive charge. Bacterial cell walls tend to carry a slight negative charge, so the positive charge of the cationic dye will bind well to the cell and stain the cell so that it can be seen. Dyes such as CRYSTAL VIOLET, METHYLENE BLUE, and CARBOL FUCHSIN are all goods ones to use for the simple stain.

Simple staining procedure:
1. Prepare a smear of the culture. If the smear sample is from solid growth media, i.e., a slant or plate, put a drop of water of the slide to make the smear.
2. Allow the smear to air dry.
3. Heat fix
4. Cover the smear with the dye. Allow to stain for at least one minute.
5. Wash off the dye with a gentle stream of water.
6. Blot the slide dry with bibulous paper.
7. Observe under oil immersion on the microscope.

With a simple stain you want to note the shape of the cells, either rod shaped (bacillus) or round (coccus), and any type of arrangement for the cells.

LAB EXERCISE

Make simple stains of the following:

E. coli (TSA slant) – the one that you grew from the last lab.
Serratia marcescens (TSB)
Staphylococcus epidermidis (TSA slant)
Cheek cells – this will provide you with a good comparison between the size of the cells in your mouth and the bacteria associated with these cells

RESULTS:
You should record in your lab book the shape and arrangement of the organisms that you have just observed. These drawings are for your records and as a reference for future identifications.

E. coli

Serratia marcescens

Staphylococcus epidermidis

Cheek cells and bacteria

Gram Stain

The Gram staining procedure is THE most important differential staining procedure that you will learn in Microbiology. The identification of unknown bacteria, which you will be doing this semester, begins with the Gram stain. You need to be able to do the Gram stain with absolute reliability, guessing is not an option!
The Gram stain is a differential stain, this means two things, first, this procedure uses more than one dye, and secondly, this procedure allows you to differentiate between different groups of bacteria.

As you may remember from lecture, the basis of the Gram stain procedure lies with the differences in the composition of the cell wall between gram positive and gram negative bacteria. The major structural component of the cell wall is peptidoglycan. Gram positive cells have large amounts of this material, while gram negative cells have very little. In the Gram staining procedure, the primary dye, crystal violet, binds strongly to the peptidoglycan matrix. Cells with large amounts of this matrix, gram positive cells, retain the dye during the staining procedure and as a result strain blue. Gram negative cells with small amounts of the matrix do not retain the dye during the staining procedure and as a result stain red, the color of counter-stain, safranin.

Gram staining procedure:
1. Make a smear, air dry, heat fix.
2. Flood the smear with the primary dye, CRYSTAL VIOLET. Allow to stain for 1 minute.
3. Wash off the dye with water.
4. Flood the smear with GRAM’S IODINE. Leave the iodine on the slide for 1 minute.
5. Wash with water.
6. Decolorize the smear with 95% ethyl alcohol.
7. Rinse with water.
8. Counter-stain with SAFRANIN for 45 seconds.
9. Rinse with water.
10. Blot dry with bibulous paper, and observe under oil immersion.

In this procedure the Gram’s iodine is a MORDANT, or “fixative”. The iodine reacts with the crystal violet to form an insoluble complex within the peptidogylcan matrix. The 95% ethyl alcohol is the decolorizing agent that removes this complex from the thin peptidoglycan matrix of gram negative cells. Safranin is the counterstain that stains the gram negative cells red, the gram positive cells are already stained blue with the crystal violet and will not take up the counterstain. The procedure is actually quite simple and with some practice is easy to master. However, there are some ways to introduce error into the procedure, here are a few:
1. Not using fresh cultures. Cultures older than 48hrs can give variable Gram stain results.
2. Smears too heavy – heavy, thick smears can result in incomplete decolorizing with the ethyl alcohol resulting in mixed results.
3. Using too much alcohol to decolorize – too heavy a rinse with the alcohol can cause the crystal violet to leave the gram positive cells resulting in the cells appearing gram negative.
4. Forgetting to heat fix before staining.

LAB EXERCISE

Do Gram stains on the following cultures:
E. coli (TSB)
Staphylococcus epidermidis (TSB)
Bacillus cereus (TSA)
Moraxella catarrhalis (TSA)
Proteus vulgaris (TSA)

Remember some of these cultures will need a drop of water on the slide to make the smear. Which ones?

Have me check your gram stains for accuracy so you will know if you have the correct results.

|Bacteria |Gram reaction |Shape |
|E. coli | | |
|S. epidermidis | | |
|Bacillus cereus | | |
|M. catarrhalis | | |
|Proteus vulgaris | | |

Gram positive = blue
Gram negative = red

Bacterial Motility

Perhaps half of all bacteria are motile. They can move in their surroundings with speed and purpose by means of flagella. Sense staining the flagella can be somewhat difficult, how then can you determine if a bacterium shows true motility or random motility, Brownian motion? Brownian motion results from the cells being bombarded by water molecules which are in motion thus imparting a shaking “in place” motion to the cells. These cells aren’t “going anywhere”, they are just moving in place. Bacteria which show true motility are seen to tumble and roll, and move with speed through the media. So without doing the flagella stain how can you determine motility? They are basically two methods, direct observation, and indirect determination using growth media.

Direct observation
Bacteria are hard to see if they are not stained, but if you stain them they are dead, so motility can not be determined. But, in fact, you can observe bacteria alive and unstained; it’s called a wet mount and is a direct method to determine motility. The tricky part about this method is locating extremely small organisms that are virtually transparent to light and are not colored. The secret is to reduce the light coming through the slide by closing down the iris on the condenser of your microscope. By doing this you should be able to see the live bacteria and determine motility. You prepare a wet mount as follows:
1. Using the proper aseptic technique, place a drop of bacteria on a microscope slide. If you want to use a second loop of bacteria remember to repeat the aseptic technique. (Broth cultures are best to use for this technique).

2. Place a cover slip over the bacteria, the liquid will spread out to a thin film. Put a drop of immersion oil on the cover slip and observe with the oil lens. You will have to focus very carefully and reduce the amount of light coming through the slide.
3. When you have finished with the slide put the slide and cover slip into your disinfectant jar.

Indirect observation
The other method to determine motility is indirectly using a growth media that can illustrate growth and motility. The media we use for this purpose is called SIM media. This media is solid but with a low agar content. When inoculated in a special way it is possible to determine if an organism shows motility. The inoculation is called a stab inoculation and requires the use of your straight wire “needle”, not the usual loop. To inoculate this media you remove a sample of the bacteria to be tested from its growth media, either liquid or solid, and stab the SIM media straight down the center and back out. The stab should be as straight as possible. After incubation, bacteria that are non-motile will grow just along the stab line, motile bacteria with show spreading growth away from the stab line. The key to this procedure is to be as straight in and out as possible. If you “wiggle” the needle in the media as you inoculate it, then non-motile bacteria will appear to be spreading because of this inoculation error and will incorrectly judged as motile.

LAB EXERCISE

Do a wet mount and SIM inoculation on the following bacteria:

Staphylococcus epidermidis
Enterobacter aerogenes
Streptococcus lactis
Pseudomonas aeruginosa

RESULTS:

|Bacteria |Wet mount |SIM |
|S. epidermidis | | |
|E. aerogenes | | |
|St. lactis | | |
|P. aeruginosa | | |

Negative Stain

The negative stain is a simple stain that is used primarily to determine the size, shape, and arrangement of bacterial cells. When you do this staining procedure NO heat fixing step is done so the cells are not distorted by the heating process. Due to the absence of heat fixing you can often get a better idea of the shape of the cells, their size, and arrangement. This is an easy procedure than can yield a lot of information about the organism. The dye used in this procedure is called NIGROSIN, it is an anionic dye, meaning that the dye carries a negative charge. Remember, bacterial cell walls tend to also carry a negative charge so the like charges will repel each other; the net result is that the cell does not stain, the background is stained. The cells will appear a small white “ghosts” against a black background. You may see a small amount of staining of the cells, but in general only the background surrounding the cells is stained. If you have done a simple stain or Gram stain and still can’t tell the shape or arrangement of your bacteria do a negative stain, it can really help.

Negative staining procedure:
1. Place a small drop of nigrosin at one end of a slide
2. Aseptically transfer a sample of bacteria into the drop of nigrosin. Broth cultures work best but solid cultures will also work.
3. Holding another slide at about a 45° angle, place this slide into the nigrosin/bacteria mix and pull the mixture the length of the slide to make a smear.
4. Allow to air dry.
5. Put a drop of immersion oil on the smear and observe the results.

Negative staining of a bacillus. Note the outline of the cells against the black background.

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LAB EXERCISE
Do negative stains of the following bacteria
E. coli
Enterococcus faecalis
Bacillus cereus

RESULTS:

|Bacteria |Shape |Arrangement |
|E. coli | | |
|E. faecalis | | |
|B. cereus | | |

Acid-fast Stain

The acid-fast stain is another type of differential stain that has important medical applications. The acid-fast staining procedure is based on the presence of a waxy lipid in the cell wall of some bacteria called MYCOLIC ACID. If this substance is present the primary dye in the acid-fast procedure, carbol fuchsin, will bind very tightly to the mycolic acid and stain the cells a very bright purple-pink (fuchsia) color. If there is no mycolic acid present the cells will not retain the dye and will stain the color of the counter stain which is blue (see the detailed staining procedure below for the counter stain). So what about the medical importance? Bacteria which are acid-fast and therefore have mycolic acid in their cell walls are all members one genus, Mycobacterium. Two well known and important human pathogens that are members of this genus are Mycobacterium tuberculosis and Mycobacterium leprae, the causative agents of tuberculosis and leprosy. Tuberculosis kills well over one million people world-wide every year so the detection and diagnosis of this bacteria is very important. If you were the micro tech in a hospital lab and got a sample that was suspected of tuberculosis you would do the acid-fast procedure to determine if the characteristic fuchsia rods are present in the sample. If the fuchsia rods are present then tuberculosis is confirmed. This is an important differential stain, separating the microbial world into those with mycolic acid and those without. What about Gram staining these bacteria? Generally the Gram procedure is not done on these bacteria, but if you did gram stain members of this genus the result would be gram positive. In the lab today we will be doing the acid-fast procedure not on Mycobacterium tuberculosis but on some non-pathogen members of genus Mycobacterium.

Acid-fast procedure:
1. Make a smear of the sample and allow to air dry.
2. Heat fix.
3. Flood the smear with the primary dye, CARBOL FUCHSIN
4. Heat the slide as demonstrated for 3-4 minutes. Replenish the dye as it evaporates.
5. Wash off the dye with a gentle stream of water.
6. Decolorize the smear with a dropper full of ACID ALCOHOL.
7. Gently was off the excess acid alcohol with water.
8. Counterstain for 1 minute with METHYLENE BLUE.
9. Rinse off the methylene blue with water.
10. Blot the slide dry and observe under oil immersion.

Bacteria which are acid fast retain the primary dye, carbol fuchsin even after decolorizing with acid alcohol. These cells will be a bright purple-pink color and also probably somewhat clumpy because of the mycolic acid making the cells “sticky”. Cells which are not acid fast, i.e., lack mycolic acid, do not retain the primary dye after the acid alcohol rinse and are stained by the counter stain methylene blue and will appear blue on the slide. To summarize, acid-fast bacteria will stain fuchsia, non-acid fast bacteria will stain blue. Once again it’s all to do with the cell wall, but this time it’s not peptidoglycan amounts (Gram stain) but the presence or absence of mycolic acid.

LAB EXERCISE

Do acid fast stains on the following:
Mycobacterium smegmatis
Mycobacterium phlei
Staphylococcus epidermidis

Make the following smears:
1. One slide of either species of Mycobacterium by itself. The slant cultures of these bacteria are thick and sticky so remember to use a drop or two of water on the slide to make the smear.
2. One slide of either species of Mycobacterium mixed with the broth culture of Staphylococcus epidermidis. This slide will help you see the difference between acid fast and non-acid fast bacteria on the same slide.

From the second slide, what’s the answer to this question? What color are the Mycobacterium cells, and what color are the cells of the Staphylococcus?

Spore Stain

This is the last of the differential stains that you will have to learn. This stain differentiates bacteria into cells that have to ability to form spores from those cells that do not. Spores are extremely resistant structures that some species of bacteria can form when the cells become stressed by lack of nutrients or changes in their environment. Spores DO NOT represent a form of reproduction, but rather a mode of survival. Bacterial spores can lie dormant for hundreds even thousands of years but when placed in nutrient media will quickly germinate into active, live cells. There are only two genera of bacteria than have the ability to form spores. These are genus Bacillus and genus Clostridium. Anytime you see the genus name of a bacteria that is either one of these two you know that all the bacteria of these genera form spores. Medically there are some very important diseases caused by spore forming bacteria, these include diseases such as anthrax, tetanus, botulism, and gangrene to mentions just a few.

Spores are very resistant structures, and as such are resistant to staining as well. As in the acid fast procedure you must use heat to drive to primary stain into the spores. The spore stain procedure is as follows:
1. Make a smear.
2. Air dry, heat fix.
3. Flood the smear with the primary dye, MALACHITE GREEN. Heat by the same method used in the acid fast procedure for 3-4 minutes.
4. Wash off the primary dye with a stream of water.
5. Counterstain for 1 minute with SAFRANIN.
6. Wash with water
7. Blot dry and observe under oil immersion.

The primary dye in the spore stain, malachite green, binds to cells rather weakly, that is why water is used as a decolorizer. The dye however binds very strongly to the spores when heated, so water will not wash the dye away. When the spore stain is done properly you should see the following:
1. The spores, which are oval to round in shape, will stain greenish blue. The spores may be observed in several locations, either inside the cells near the center of the cell or near the end of the cell. You will also see many free spores, spores that are not associated with any cell because the “parent” cell ruptured and disintegrated.
2. Vegetative cells, cells that have not formed spores or are a non-spores forming species of bacteria, will appear red, the color of the counterstain safranin.

LAB EXERCISE

Cultures:
Bacullus laterosporus(TSA)
Stahhylococcus epidermidis(TSA)

Make the following spore stains:
1. One slide of the Bacillus laterosporus by itself.
2. One slide of a mixture of the Bacillus laterosporus and Staphylococcus epidermidis.

From these slides you should be able to distinguish between spores, vegetative cells and non-spore forming cells. These differences will show up as differences in colors.

Summary of Staining Procedures

Simple stains: Cationic dyes Crystal violet Methylene blue Carbol fuchsin Safranin Malachite green Anionic dyes Nigrosin – negative stain

Differential stains: Gram stain Crystal violet – primary dye 95% ethanol – decolorizer Safranin – counterstain Acid-fast stain Carbol fuchsin – primary dye Acid alcohol – decolorizer Methylene blue – counterstain Spore stain Malachite green – primary dye Water – decolorizer Safranin – counterstain

Determination of Bacterial Cell Density

If you were given a broth culture of bacteria, and asked to determine how many cells were in that sample, how would you do this? No idea probably, but after this lab you’ll know how it’s done. There are several ways to count bacteria. One method would be to use expensive instruments called electronic particle counters to do the work for you. That’s not an option in this lab. Another way to count bacteria is do to it directly, the direct count method. You make a dilution of the culture, put some of the cells in a special counting chamber than just start counting the cells, 1,2,3,4 etc. As you might imagine this can be tough on the eyes, and produce some errors as well. The other way to count bacteria, and the way we are going to do it in this lab, is called the viable count method. In this method you make a series of careful dilutions of the original sample, and at certain dilutions take a small sample of the dilution and spread it on a TSA plate. After the plate has incubated all you do is count the number of colonies on the plate, and sense you know the dilution of that plate you just multiply the number of colonies times the dilution to arrive at an estimate of the viable (live) number of bacteria in the original sample. It’s not hard, but you do need to make accurate dilutions and remove accurate samples to get good numbers. We’ll get into the actual details of the procedure next.

Dilutions are simply a method to make a concentrated solution less concentrated, more dilute, in a controlled and precise manner. In this exercise you will be making a series of sequential dilutions, or serial dilutions of a concentrated sample of bacteria making it much more dilute as you proceed with the dilutions. You will be making a series 10 fold dilutions, that is, every dilution you make will be 10 times more dilute than the one before it.

Our dilution scheme will proceed as follows:

Each table will have a dilution of E. coli in water, all the tables will have the same starting dilution which is 1:100 or 10-2. I made this dilution from the original broth culture simply by pipeting 1ml of the culture into 99 mls of sterile water, that’s a 1 to 100 dilution (1:100). Sense each table has the same starting dilution all the groups should get the same results for number of bacteria, we’ll see! From this initial 1:100 dilution each group will need to make a series of serial dilutions out to a final dilution of 10-7 or a dilution of 1:10,000,000. To make these dilutions and grow the bacteria each group will need the following: 5 tubes containing 9 ml of sterile water 9 TSA plates

To make the dilutions accurately your groups will using Pipetman pipetters. The use of these devices will be demonstrated to the class. You have two Pipetman, the large one is set to measure 1ml and uses the large BLUE tips, the smaller Pipetman is set to measure .1ml and uses the smaller YELLOW tips. Before you start the dilutions have all tubes and plates labeled so you don’t make pipeting or plating mistakes. Here’s the dilution scheme:

1. From the 10-2 flask transfer 1ml to 9ml of sterile water. This dilution is 10-3. Change tips.
2. From the 10-3 tube transfer 1ml to another 9ml water tube. This dilution is 10-4. Change tips.
3. From the 10-4 tube transfer 1ml to another 9ml water tube. This dilution is 10-5. Change tips.
4. From the 10-5 tube transfer 1ml to another 9ml water tube. This dilution is 10-6. Change tips.
5. From the 10-6 tube transfer 1ml to your last 9ml water tube. This dilution is the final 10-7 dilution. Remove the tip.

The serial dilutions are now done. Which ones to use?
Of the dilutions that you have just done, which one or ones are dilute enough such that when a sample of the dilution is spread on a TSA plate you will get colonies that can be counted? It’s not always easy to pick the right dilution factor so to be safe you always pick several dilutions to sample. We will take samples from the following dilution tubes: 10-5 10-6 10-7
Samples from these tubes will be placed on TSA plates and incubated at 37°C until the next lab. Here is procedure for sampling:
1. From the tube labeled 10-5 remove a sample of 0.1ml. You will use the smaller Pipetman with the yellow tips for this. Pipet the sample directly onto a TSA plate. Repeat the process two more times on the two other plates. You will have THREE TSA plates that have samples on them from the 10-5 dilution. You DO NOT have to change tips between samples in this part of the procedure.
2. Repeat this procedure for the other two dilutions, 10-6 and 10-7.
3. Spread the samples on the plate as demonstrated.
3. You will have NINE TSA plates to incubate.

Incubate all plates at 37°C in an inverted position. We will examine the results and do the counts the next lab period.

Determination of Bacterial Density
Part II

Let’s examine the plates from the last lab. If all the dilutions were done properly then the plates should reflect the dilutions, that is, the 10-5 plates should show the most colonies, the 10-7 the fewest, and the 10-6 plates somewhere in between the other two dilutions. If your plates don’t reflect this pattern then it is likely a dilution error has occurred during the procedure. Now, which plates do you count? We want to choose the plates which have from 30 to 300 colonies on them to count. Counting a plate with 10 colonies on it may be easy, but the results are statistically inaccurate. Counting the colonies on a plate with over 300 colonies is probably impossible to do with any accuracy so is also not done. There should be one series of plates, probably the 10-6 dilution, that will have the right range of colonies. Counting is just that, use a marking pen and mark out and count the colonies on the plate. Do all three plates and get an average number. If one plate in the series has an obviously different number, then throw that number out. For example, if the counts are 86, 77, and 20, then don’t average in the 20, just average the two closest plates. When you count colonies on a plate you are actually counting colony forming units (CFU), instead of individual cells, so our final calculation will reflect this as colony forming units per ml of original broth rather than cells per ml of original broth. Here’s the formula for the calculation:

Number of colonies
Original cell density(CFU) = Volume plated X dilution factor
For example: 120
Original cell density = .1ml X 10-6 = 1.2 X 108 CFU/ml (10-6 = .000001 in decimal format)
Now you do the counts and calculations and let’s see what we get!

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Antibiotic Sensitivity Testing

In this lab we will be assessing the effects of various antibiotics against a variety of bacteria. When a person has a bacterial infection it is important to pick the right antibiotic to treat the infection. Given the current crisis that exists with bacterial resistance to antibiotics, it is important to treat the infective agent with an antibiotic that will have the best action. Guessing which antibiotic to use is really not an option when the patient is sick. Using the wrong antibiotic may have no effect, and can lead to increased resistance. The way to determine the effectiveness of a given antibiotic is the subject of this lab. The procedure you will be learning is called the Kirby-Bauer antibiotic sensitivity test, also known as the disc diffusion test. You will be determining the resistance or sensitivity of a set of 12 antibiotics against various bacteria.

The procedure for this lab is very simple. You will be using extra large Petri dishes that contains a media that is only used for this procedure. The media is called Mueller-Hinton agar. It is a media that assures rapid growth of aerobic and facultative anaerobic bacteria. Rapid growth, overnight growth, would be critical if you have a sick patient and doctors waiting to find out which is the best antibiotic to use.

LAB EXERCISE:
Here is the procedure that you will use for this lab:
Get one of the large plates and a tube containing a sterile cotton swab. You will be assigned a bacterium to inoculate. Check the list and get your organism. It will be a broth culture. Use the cotton swab to inoculate the entire plate surface for a “lawn” of bacterial growth. After the plate is inoculated come up to the front desk and use the antibiotic disc dispenser to put the 12 discs on the plate. Wait for 10 minutes before you put the plate into the incubator. This will allow the discs a chance to adhere well to the surface of the plate. Incubate the plates, inverted, at 37°C. Next lab you will determine the sensitivity or resistance of the various antibiotics to your bacteria.

Bacteria:
Alcaligenes faecalis
Enterobacter aerogenes
E. coli
E. coli (ampicillin resistant)
Klebsiella pneumoniae
Pseudomonas aeruginosa
Enterococcus faecalis
Staphylococcus aureus
Staphylococcus epidermidis
Citrobacter diversus

Antibiotic Sensitivity Testing
Part II

Now that the plates have grown you can determine the effectiveness of the various antibiotics against your test bacteria. You will notice that around some of the discs there is a clear zone of inhibition where no bacteria have grown, and around others there is no zone of inhibition. If there is NO zone of inhibition around the disc then the answer to the sensitivity/resistance is easy, the bacteria is resistant (R) to this antibiotic. If there is a visible zone around the disc this does not automatically signify sensitivity. You must measure the diameter of the zone and then compare your measurement with the values in the lab book table that will tell you whether the zone indicates sensitivity (S), resistance (R), or intermediate (I).

Using a ruler, as shown below, measure the diameter of all the zones and write them down for interpretation from the table.

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Some of the zones may be faint or hard to measure. If this is the case take a marking pen and outline the zone of the bottom of the plate to make the measurement easier to do.

Antibiotic Sensitivity Test Results

|Antibiotic |Concentration |Zone Diameter |Interpretation |
| | | |(R, S or I) |
|Trimethoprim(TMP) |5mg | | |
|Rifampin(RA) |5mg | | |
|Penicillin(P) |10 units | | |
|Kanamycin(K) |30mg | | |
|Vancomycin(VA) |30mg | | |
|Ampicillin(AM) |10mg | | |
|Streptomycin(S) |10mg | | |
|Tetracycline(TE) |30mg | | |
|Erythromycin(E) |15mg | | |
|Chloramphenicol(C) |30mg | | |
|Norfloxacin(NOR) |10mg | | |
|Gentamycin(GA) |10mg | | |

This procedure is called disc diffusion because once the disc is placed on the plate the antibiotic in the disc begins to diffuse, or spread out, into the surrounding media. The amount of diffusion is related to the size of the antibiotic molecule, and the thickness of the agar. The larger the size of the antibiotic molecule the shorter the distance it will diffuse.

Table of Zones of Inhibition

|Antibiotic |Disc Potency(mg) |Resistance zones(mm) |Intermediate zones(mm) |Sensitive zones(mm) |
|Ampicillin |10 | | | |
|G- enterics | |18 |
|Staphylococci | |29 |
|Enterococci | |17 |
|Streptococci | |30 |
|Cephalothin |30 |18 |
|Chloramphenicol |30 |18 |
|Erythromycin |15 |23 |
|Gentamycin |10 |15 |
|Kanamycin |30 |18 |
|Nalidixic acid |30 |19 |
|Norfloxacin |10 |17 |
|Penicillin G |10 units | | | |
|Staphylococci | |29 |
|Enterococci | |15 |
|Streptococci | |28 |
|Rifampin |5 |20 |
|Streptomycin |10 |15 |
|Tetracycline |30 |19 |
|Tobramycin |10 |15 |
|Trimethoprim |5 |16 |
|Vancomycin |30 |17 |

Some of these antibiotics are called broad spectrum antibiotics, other narrow spectrum antibiotics. It is best, when possible, to use the narrow spectrum antibiotics to limit the spread of resistance. From the results of this exercise can you identify some bacteria that you would NOT like to meet in the infective sense?

Effects of pH on Bacterial Growth

As we discussed in lecture, pH is the measure of whether a solution is acidic or basic. Solutions that range from 0-7 are acidic, while those above 7 are considered basic (alkaline). Exactly 7 is neutral, this is the pH of pure water. The pH of human blood is slightly basic, about 7.2. The range of pH and some examples is shown in the figure below.

[pic]

Bacteria also have a characteristic range of pH tolerance, and that is the subject of this lab. Bacteria can be divided into three categories of pH tolerance: Acidophiles – pH range of 1-5.4 Neutrophiles – pH range of 5.4-8.0 Alkalipiles – pH range of 7.0-11.5
All bacteria have an optimum pH, the pH at which they grow best, but also a range, as shown above, at which they will grow. Sense the pH of humans, the host of most bacteria, has a pH of approximately 7, it is not surprising to find that most bacteria, and especially the pathogenic bacteria, have a pH range of a neutrophile. You will be testing the pH tolerance of several bacteria using media that has been adjusted to pH 3, 7, and10.

LAB EXERCISE:
Each person will have three TSA plates that have been adjusted to the test pH. You will have one plate at pH 3, another plate at pH 7, and the third plate is at pH 10. With a marking pen draw a quadrant on the bottom of each plate and inoculate each quadrant with a loop of the following bacteria: E. coli Staphyococcus aureus Staphylococcus epidermidis Alcaligenes faecalis
Incubate all plates at 37°C.

RESULTS:
From the growth on the plates what is the pH classification of the test bacteria?
E. coli
Saphylococcus aureus
Staphylococcus epidermidis
Alcaligenes faecalis

Effect of Osmotic Pressure on Bacterial Growth

One of the things that bacteria require for growth is water. In fact, bacterial cells are mostly water, 80-90% of the cell is water. If the cell is placed in an environment where there is a higher concentration of solutes than inside the cells the cells will lose water to the outside environment. This osmotic loss of water is called plasmolysis. The result is inhibition of growth as the plasma membrane pulls away from the cell wall. Figure 6.4 from your text illustrates this process.

[pic]

This inhibition of growth by high osmotic pressure can prevent bacterial growth in some foods, especially sweet and salty foods.
These foods present a hypertonic environment to the bacteria and prevent bacterial growth, a natural preservative in a way. The key word here, however, is to prevent growth. The cells are not killed, such an environment is bacteriostatic, inhibition of bacterial cell growth. If you were transfer the cells to an isotonic environment were the cell membrane is allowed to return to its normal position against the cell wall, the cells will once again resume growth. In this lab you will be inoculating nutrient agar plates that have been modified to exert varied osmotic pressures by the addition of NaCl. The plates have had salt added such that the concentration of salt in the media is .5, 5, 10, and 15%. Which bacteria will grow, and how much salt can they tolerate?

LAB EXERCISE:
Each person gets 4 NA plates, one for each salt concentration. Divide the plates by drawing this pattern on the bottom of the plate:

[pic]
A small loop inoculation of each area of the plate is all that is required. All plates will be incubated at 37°C until the next lab.

Effects of Osmotic Pressure on Bacterial Growth
Part II

Now that the plates have grown, what do we see for the relationship between salt concentration (osmotic pressure) and amount of growth? It is not possible to accurately measure the growth in this experiment, so we will make a qualitative estimation of growth. We’ll set up a scale of 0 to 3 for growth, 0 = no growth, 1= little growth, 2 = good growth, and 3 = excellent growth. It’s far from exact but will give us an estimate of growth.

Results
|Bacteria |.5% NaCl |5% NaCl |10% NaCl |15% NaCl |
|Streptococcus lactis | | | | |
|Staphylococcus aureus | | | | |
|Staphylococcus epidermidis | | | | |

In general, the members of genus Staphylococcus are more salt tolerant than genus Streptococcus. In particular, Staphylococcus aureus is very salt tolerant. Ever had a sore throat, and were told to gargle salt water? The reason that this home remedy might work is because the bacteria of genus Streptococcus that can be causing the sore throat are killed by the salt. Try this next time you have a sore throat, about a teaspoon of table salt in a cup of warm water. Just remember not to swallow the water!

Effects of Temperature on Bacterial Growth

As is the case for pH and osmotic pressure, bacteria also have specific temperature requirements. Bacteria will grow over a wide range of temperatures, but there is always an optimum temperature for growth, the temperature at which the cells grow the best. Shown below is a figure from your text that illustrates the range of bacterial growth.

[pic]

Temperature is related to growth rate by enzyme activity. Enzymes, as you will recall from metabolism, control all aspects of cell function. Enzymes also have an optimal temperature for their activity, which translates into the growth rate of the cells. If the temperature is low, enzyme activity slows and so does growth, if the temperature is too high the enzymes can denature due to heat, and growth slows or stops altogether. In this lab you will be investigating the growth of several different bacteria over a range of temperatures. From the above figure, the bacteria called mesophiles are the most important from a medical point of view. Why? Because the optimal growth temperature for humans corresponds with the optimal growth temperature for the mesophiles, 37°C. All the important human pathogens are mesophiles. There are several other temperature groups that also need mention. These are the psychrophiles, bacteria that grow at colder temperatures, as low as 10-12°C, and the more moderate psychrotrophs, bacteria that have an optimal growth temperature of around 20°C. At the other extreme we have the thermophiles, bacteria that grow at temperatures above 60°C. Because this temperature would kill humans, there are no thermophiles that are serious human pathogens. At the absolute upper end are the extreme thermophiles that grow at temperatures above 80°C. These bacteria have evolved very special proteins that are not denatured at such extreme temperatures.

In this lab exercise you will be inoculating several test bacteria onto TSA plates and incubating the plates at 5 different temperatures and assessing growth. The incubation temperatures will be 5° (refrigerator), 25° (room temperature), 37°, 45°, and 60°C. You will be inoculating each plate with four bacteria, you will need 5 TSA plates, one for each temperature.

LAB EXERCISE:
The bacteria for this exercise are broth cultures of the following:
1. Bacillus stearothermophilis
2. Pseudomonas aeruginosa
3. E. coli
4. Staphylococcus aureus
5. Bacillus subtilis
6. Klebsiella pneumoniae
7. Staphylococcus epidermidis
8. Serratia marcescens

If you sit at an ODD number seat use organisms 1-4 for inoculation. Even number seats use 5-8 for inoculation.

Divide the plate into quadrants and label each one with the bacteria that has been placed on that quadrant. Again, a loop inoculation spread to almost fill the quadrant is all that is needed.

[pic]

Incubate all plates at 37°C for evaluation during the next lab period.

Effect of Temperature on Bacterial Growth
Part II

Now that the plates have grown, let’s have a look at the growth in relation to temperature. As you have done before, you will use a qualitative scale for growth of 0 to 3, 0 being no growth, 1 is little growth, 2 is fair to good growth, and 3 is excellent growth.

RESULTS:
|Bacteria |5°C |25°C |37°C |45°C |60°C |
|B. stearothermophilus | | | | | |
|Pseudomonas aeruginosa | | | | | |
|E. coli | | | | | |
|Staphylococcus aureus | | | | | |
|Bacillus subtilis | | | | | |
|Staphylococcus epidermidis | | | | | |
|Klebsiella pneumoniae | | | | | |
|Serratia marcescens | | | | | |

Based on these results, classify the test bacteria as either psychrophiles, mesophiles, or thermophiles.

Bacillus stearothermophilis –
Pseudomonas aeruginosa –
E. coli –
Staphylococcus aureus –
Bacillus subtilis –
Staphylococcus epidermidis –
Klebsiella pneumoniae -
Serratia marcescens -

Growth Characteristics of Anaerobic Bacteria

So far this semester you have been working with bacteria that grow in the presence of oxygen. These bacteria have been either aerobes and facultative anaerobes. As you know from your work with these bacteria they are easy to grow, and have few special requirements for growth. There are, however, bacteria that do not grow in the presence of oxygen, it is poisonous to their metabolism. These bacteria are called strict anaerobes. As you might expect such bacteria have some special needs related to their growth, that is, an environment or media that is free from high levels of oxygen. There are some medically important bacteria in the anaerobe category; the bacteria that cause tetanus, botulism, and gas gangrene are all anaerobic bacteria. Because of the unique requirements for the growth of these bacteria, this will be about the only time this semester that you will have a chance to work with these bacteria.

One type of media that is used in the cultivation of anaerobes, but will support the growth of all types of bacteria, is called thioglycolate media. This media contains various growth factors, but also contains a compound called sodium thioglycolate which reduces the oxidation-reduction potential of the media. This provides a more suitable growth environment for anaerobic bacteria. The media also contains a dye called resazurin, which is an indicator for the presence of oxygen. You will notice that tubes of thioglycolate media have a light pink layer at the top of the broth indicating oxygen. DO NOT mix the tubes before inoculation. The oxygen levels in the rest of the tube are low, you want to keep it that way. When this tube is inoculated the aerobic bacteria grow near the surface of the tube, facultative anaerobes grow through out the tube, and the anaerobic bacteria grow away from the oxygen lower in the tube.

A second type of media that you will use is called an agar deep. This media is just TSA in a bigger tube that is inoculated while still liquid then allowed to harden. The media is kept molten by keeping it in a 50°C water bath until you are ready to use it. All you do is inoculate the media, gently mix the tube, let the tube harden, and then incubate as usual. An oxygen gradient sets up in the tube, highest at the top, lowest near the bottom, with growth patterns similar to those observed in the thioglycolate media. Gas production may also occur during growth indicated by cracks in the solid media. The figure below from your book illustrates the type of growth generally seen in a deep tube.

[pic]

Another way to grow anaerobes that you will use is shown above. This is called a GasPak. Using this apparatus you can inoculate ordinary media, plates or broth, put them in the chamber, open a packet of oxygen absorbing material, then close the lid. In a few minutes the oxygen will be removed from the chamber creating an anaerobic environment. An indicator strip is added to chamber before closing that will change color to indicate when the chamber is anaerobic. The whole chamber would then be incubated at 37°C. This method does have several drawbacks however, the small size of the chamber limits the numbers of samples that can be put into the chamber, and once the chamber is sealed it cannot be opened until the incubation is finished. By comparing patterns of growth on media incubated inside the chamber with the identical media incubated outside the chamber you can determine the oxygen requirement of the bacteria in question.

LAB EXERCISE:
You will need the following media: 3 tubes of thioglycolate media 3 melted agar deep tubes 2 TSA plates

Bacteria: Odd number seats Even number seats Staphylococcus aureus Pseudomonas aeruginosa Acaligenes faecalis Enterobacter aerogenes Clostridium perfringens Clostridium sporogenes
All the cultures are broth cultures. The two bacteria from genus Clostridium are in screw cap tubes of thioglycolate media because these bacteria, as are all members this genus, are anaerobes.

PROCEDURE:
1. Thioglycolate tubes – inoculate your assigned bacteria into these tubes are you would inoculate any broth. The thioglycolate media is in screw caps tubes. After inoculation tighten the caps to prevent oxygen from entering the tubes. DO NOT shake the tubes. Incubate at 37°C.
2. Agar deep tubes – get three of these tubes out of the water bath and inoculate with your assigned bacteria. Don’t get these tubes out of the water bath until you are ready to use them. If you let them sit at room temperature for more than a few minutes the tubes will solidify and cannot be inoculated. These tubes have metal slip caps, and are inoculated in the usual manner for a liquid media. After inoculation gently mix the tubes, and incubate the tubes at 37°C.

3. TSA plates

[pic]

Inoculate your two TSA plates with your assigned bacteria as shown above. One plate is incubated at 37°C as usual. The other plate will be incubated in the GasPak so oxygen requirements can be determined.

Next lab we will determine the oxygen requirements for our test bacteria.

Oxygen Requirements
Part II

From the results of your inoculations, which bacteria are aerobes, facultative anaerobes, and strict anaerobes?
RESULTS:
|Bacteria |Growth in |Growth in |GasPak plate |Control plate |
| |Agar Deep |Thio media |growth |growth |
|Staphylococcus | | | | |
|aureus | | | | |
|Alcaligenes | | | | |
|faecalis | | | | |
|Clostridium | | | | |
|perfringenes | | | | |
|Pseudomonas | | | | |
|aeruginosa | | | | |
|Enterobacter | | | | |
|aerogenes | | | | |
|Clostridium | | | | |
|sporogenes | | | | |

Based on the results from above classify the bacteria as an aerobe, anaerobe, or facultative anaerobe.
|BACTERIA |OXYGEN REQUIREMENT |
|Staphylococcus aureus | |
|Alcaligenes faecalis | |
|Clostridium perfringenes | |
|Pseudomonas aeruginosa | |
|Enterobacter aerogenes | |
|Clostridium sporogenes | |

Bacterial Transformation

This experiment will involve the use of DNA to change, or TRANSFORM, a bacterium. You will give new genes to the bacteria E. coli. These genes when expressed will produce a protein called the pGLO protein that will cause colonies of the bacteria to glow a bright green color when exposed to UV light. The phenomenon of transformation was discovered in 1928 by Fredrick Griffith. Experimenting with different strains of Streptococcus pneumoniae, Griffith discovered that one strain of harmless bacteria could be changed, or transformed, into a different strain that now was pathogenic. The actual mechanism of this change was unknown to Griffith, it was not until 1944 that Avery and co-workers discovered that the agent of transformation was in fact DNA.

For your experiment you will be making a suspension of E. coli in calcium chloride and adding a small of DNA to the suspension. The calcium chloride causes the cells to become competent, that is, the cells become changed in some unknown way and are more receptive to picking up DNA from the surrounding media. If the experiment is successful the colonies which result should produce a protein that glows green under UV light.

EXPERIMENTAL PROCEDURE
1. You will be working in groups for this experiment. Regardless of the group size each table will get one plate of E. coli. Each group will also be supplied with the following: 2 microfuge tubes labeled +DNA and –DNA each containing 250ul of CaCl2 , and another microfuge tube containing 1ml of LB broth. Each group will also need the following plates: 1 plate labeled LB, 2 plates labeled LB/amp, 1 plate labeled LB/amp/ara.

2. From the plate of E. coli remove a colony and place it in the tube labeled +DNA. Repeat this process and put a colony into the tube labeled –DNA. Mix both tubes well to disperse the bacteria in the CaCl2 . Place both tubes on ice for 10 minutes.

3. After 10minutes on ice it is time to add the pGLO gene to the cells. You have a small tube that contains 30ul of DNA. This tube is labeled pGLO. You need to add 20ul of the DNA to the tube labeled +DNA. The other tube labeled –DNA receives nothing. This is the control tube for the experiment. Keep both tubes on ice for another 10 minutes.

4. HEAT SHOCK! This is an important step in this procedure. After the cells have been on ice for 10 minutes, both tubes, +DNA and –DNA, are place in a water bath that is at 42°C for 50 seconds. This rapid and short change in temperature facilitates the uptake of the pGLO plasmid by the cells. After this short heat shock place both tubes back on ice for 10 minutes.

5. After 10 minutes on ice place the tubes at room temperature and add 250ul of LB media to both tubes. Allow the tubes to sit at room temperature for 10 minutes. This is called the recovery phase.

6. After recovery phase it is time to put some cells on the plates. Plate as follows:
1. –DNA tube: pipet 100ul of sample onto the plate labeled LB.
2. –DNA tube: pipet 100ul of sample onto the plate labeled LB/amp.
3. +DNA tube: pipet 100ul of sample onto the plate labeled LB/amp.
4. +DNA tube: pipet 100ul of sample onto the plate labeled LB/amp/ara.
Sterilize your loop and use the sterile loop to spread the sample over the surface of the plate. Incubate plates at 37°C.

Transformation
Part II
[pic]
This is a diagram of the pGLO plasmid. The abbreviations on this figure mean the following: araC – gene sequence for the breakdown of the sugar arabinose.
GFP – gene sequence for green fluorescent protein. bla – gene sequence for beta lactamase, resistance to ampicillin is transferred by this gene.

What growth is seen on your plates? If the experiment worked these are results that you should see:
1. Plate labeled LB, sample from the –DNA tube. This is the control plate, there should be good growth on this plate.
2. Plate labeled LB/amp, sample from the –DNA tube. No growth should be present on this plate. Why? These cells received NO plasmid, and the plate contains the antibiotic ampicillin. Sense these cells have no resistance to the antibiotic there will be no growth. No plasmid = no growth!
3. Plate labeled LB/amp, sample from the +DNA tube. There should be growth on this plate because you added DNA to this tube. If the cells took up the plasmid they acquired the bla gene which will allows these cells to grow on a plate containing ampicillin. If there is no growth on this plate, then the success of this experiment is doubtful.
4. Plate labeled LB/amp/ara, sample from the +DNA tube. If there is growth on this plate then the experiment has worked and these colonies should glow green when exposed to UV light. But what does ara mean. Ara stands for the sugar arabinose which is in this plate along with the ampicillin. The arabinose is the carbon source in the media, and the plasmid that these cells have acquired has a gene sequence that allows the cell to use this sugar as a carbon source. The pGLO gene is linked to the arabinose gene. When the arabinose gene is turned on so is the pGLO gene and the green glowing protein is produced. Your have TRANSFORMED the bacteria. The E. coli picked up the plasmid from the media and uses the DNA to make a green protein, and to be resistant to ampicillin.

DNA Fingerprinting

You have all heard of DNA fingerprinting. It’s in the news, and on all the popular crime scene investigation programs. But what is this really all about, how does it work, how can a molecule leave a print? Using a fine dust to look for hand finger prints is a time honored method in detective work, but in these days of the molecular detective a new, and highly specific method of evidence collecting is available. DNA fingerprinting is predicated on the fact that the DNA sequence of humans is unique to that individual. Unless you have an identical twin, your DNA pattern, your molecular fingerprint is unique just to you. DNA can be extracted from blood or other body fluids, analyzed, and used in court to identify potential suspects. If your blood is found at a crime scene and you are a suspect, then you had better have a good attorney! DNA evidence is admissible in court and has not only put people in jail, but also freed others, in some cases these individual were close to an execution date! DNA fingerprinting is however not just used to criminal investigation, this technique is also used in paternity testing, identification of human remains, and determining the relatedness of various human ancestors.

How is the DNA actually analyzed? DNA is a very large and complicated molecule, how could you tell my DNA sequence from yours? The secret lies in the unique base sequence of DNA which is different for each individual, and the use of a special type of enzyme called a restriction enzyme. Restriction enzymes were discovered in 1968. These are enzymes that cut DNA at specific base sequences in the DNA molecule. There are over 900 different restriction enzymes that have been isolated from various species of bacteria. The “cut sites” for these enzymes are all well characterized, and different for the different enzymes. When a DNA molecule is cut with restriction enzymes, fragments of DNA of various sizes are generated, it is these fragments that are analyzed. Because everyone has a different DNA base sequence, the fragments generated by restriction digestion are of different molecular sizes. The sizes of the fragments is analyzed by a technique call agarose gel electrophoresis. Different people have different fragment patterns. Shown below is an example of a restriction digest gel, note the different pattern of bands. The bands are the dark “bars” standing out from the light blue background.

If these samples were from human DNA, the patterns seen would represent three different individuals because the banding patterns in the gel are clearly different. In our lab today you will be performing restriction digests on DNA samples from a crime scene. The pattern of DNA bands in the gels that you will run from the restriction digests should identify the guilty suspect.

EXPERIMENTAL PROCEDURE:
You will be working in teams for this experiment. Each team will need the following:
1. 1 vial containing 75ul of restriction enzyme mix. The vial is labeled EZ. The enzyme is a mixture if EcoR1 and Pst1. Keep this vial on ice!
2. Set of DNA samples. Each vial contains 10ul of DNA sample, There are 6 vials per set, and coded as follows: 1 green vial = crime scene DNA 1 blue vial = suspect 1 1 orange vial = suspect 2 1 violet vial = suspect 3 1 red vial = suspect 4 1 yellow vial = suspect 5

RESTRICTION DIGEST
1. Add 10ul of restriction enzyme mix to each DNA sample tube. Mix with the DNA samples as demonstrated. Remember to change tips between each tube!
2. Incubate the vials at 37°C for 45 minutes to digest the DNA samples.
3. After incubation all the vials will be collected and frozen until the next lab.

DNA Fingerprinting
Part II

As mentioned above, DNA fragments generated by restriction digestion are analyzed by a process called agarose gel electrophoresis. This is a very simple technique based on two facts, one that the DNA fragments are of different sizes and will “move” at different rates in the gel, and two, when subjected to an electrical current the DNA fragments will move toward the positive end of the gel. What is agarose? It is nothing more than a more pure form of the agar you have been using in the lab to grow bacteria. For this experiment a gel of agarose has been made at a concentration of .8% agarose. You will notice that the gel has small wells formed at one end. This is where you place your samples of DNA, and that’s the most difficult part of this whole procedure. After the DNA samples have been loaded into the wells a current is applied to the gel. As mentioned, DNA fragments will migrate toward the positive end of the gel, but the agarose acts as a molecular sieve which means the fragments are separated according to size. The bigger the fragment the shorter the distance of migration, the smaller the fragment the greater the distance of migration.

LOADING AND RUNNING THE GEL:
After your digests from the last lab have thawed, add 5ul of loading dye to each sample. The samples will now be blue in color. To make sure that all the sample is in the bottom of the tube you will have to briefly centrifuge your samples. Each gel has 16 wells, but the two wells at each edge of the gel are not used. Your group will add sample to 6 wells in the gel, and another group will use the rest of wells in the same gel. Make sure you remember which wells your samples are in. You need to add 10ul of digest sample to each well. The gel is covered with buffer and the wells are also full of buffer. To add the sample you position the pipet tip right over the top of the well and slowly add you sample. The loading dye contains sucrose which makes the sample heavier than the buffer in the well so the sample will displace the buffer out of the well as it is added. It takes a little practice but you’ll get it. After the gel has been loaded with your samples and those of the other group sharing the gel, you will put the lid on the box, place the correct electrodes on the correct ends, and turn on the power. We will run the gels at about 100 volts for about an hour to allow the fragments to separate into easily seen bands.

RESULTS:
After the gels have run you need to look at the bands. What bands? The gels probably only show some blue marker dye, no dark lines or bands of DNA. Here’s the answer. When the gels were made a special dye called ethidium bromide was mixed in with the agarose. This dye reacts with DNA to “stain” the bands, but can only be seen under UV light. You must be careful with these gels because of the ethidium bromide. This dye is quite toxic and the gels must be handled with gloves. We will put the gels on a UV box and take a picture of the glowing bands that have been stained with ethidium bromide. You must wear glasses when looking at UV light, it is very hard on the retinas! By examining the picture of the gel, which banding pattern of suspect DNA corresponds to the crime scene sample?

Microbial Physiology

One of the major projects that you will start soon is the identification of unknown bacteria. What techniques do you use to do this? The Gram stain is step one, and isolation of pure colonies is step two, but then what? The next few labs will introduce you to the steps involved in this procedure. By understanding the differences in metabolic characteristics between bacteria you can use this information to identify different organisms. You will be learning how to interpret bacterial growth on different test media. You will be assigned an organism to work with. It is your job to inoculate the media, accurately record the results, and present the data for the rest of the class to use for identification of unknown bacteria. Accuracy is the key, everyone, including you, is depending on the data! We will be generating data on both Gram positive and Gram negative bacteria. These are new tests for you. Here is the list of all the physiological tests you will be doing over the next few lab periods:
1. Phenol red sugar fermentation tubes – glucose, lactose, sucrose, and mannitol.
2. Kligler’s iron agar (KIA)
3. Sulfur indole motility media (SIM)
4. Lysine decarboxylase media (LDC)
5. Methyl red/Voges Proskauer (MRVP)
6. Urease
7. Citrate
8. Phenylalanine deaminase agar (PDA)
9. Nitrate
10. Bile esculin agar (BEA)
11. Catalase (rapid substrate test)
12. Coagulase test

Microbial Physiology
Explanation of results

1. Phenol red sugars:
After the tube has been inoculated and grown these are the typical results:
Yellow tube = acid production. (The pH indicator phenol red turns yellow if acid is produced.) NOTATION = A
The small tube in the media is called a Durham tube. This small tube detects the production of gas as a metabolic by-product.
Yellow tube + bubble in the Durham tube = acid +gas. NOTATION = A/G
Tubes which are orange, NOT yellow are negative for acid.
Tubes which are a fuchsia color (purple/pink) indicate aerobic metabolism, the NOTATION=K for these cells. Bacteria such as Pseudomonas aeruginosa and Alcaligenes faecalis produce this reaction on the sugar tubes.
Summary:
A = acid
A/G = acid and gas (bubble in Durham tube)
K = alkaline reaction (aerobic metabolism)

2. SULFUR INDOLE MOTILITY(SIM)
This is an important media that gives you three pieces of metabolic information. The media is inoculated with a stab inoculation.
RESULTS:
Sulfur: media turns black. It is + for H2S production, no black color is – for H2S production.
Indole: metabolic by-product of tyrptophan metabolism. Add 5 drops of Kovac’s reagent to the tube. Red color = positive for indole, no color is negative of indole production.
Motility: spreading from the stab line, is + for motility.

3. UREASE: this media is supplied as a slant, and is inoculated by a stab/streak. The compound urea is in this media. Some bacteria produce the enzyme urease, which breaks down urea and releases ammonia into the media. The release of ammonia causes the pH of the media to rise, and the pH indicator in the media, phenol red, changes color to produce a very bright pink color in the tube. So bright pink is positive for urease, no color change is negative for urease. Some tubes may not be completely pink, but any pink color is considered positive for urease. Member of genus Morganella, Proteus and Providencia are all urease positive.

4. LYSINE DECARBOXYLASE (LDC): This media is supplied as a broth, and is inoculated as you would inoculate any broth media. The test results can be a little tricky to interpret. Here is the biochemistry of the reaction: the media contains the amino acid lysine as well as glucose as a carbon source, and a pH indicator bromcresol blue. The pH indicator is purple at a pH above 6.8, and yellow at a pH of 5.2 or less. The tube is inoculated, the bacteria uses glucose and produces acid, the pH falls and the media turns yellow. Yellow color is negative for LDC. HOWEVER, some bacteria have an enzyme called lysine decarboxylase (LDC) which is INDUCED by the acid production of glucose catabolism. This enzyme allows the cells to breakdown lysine and this causes the pH to rise and the indicator turns purple. So PURPLE color is the positive reaction for the presence of the enzyme lysine decarboxylase! When the media is inoculated remember to add a small layer of mineral oil to the top of the media. This assures that the correct environment is present for the reaction to occur.
Summary:
Yellow = negative for LDC
Purple = positive for LDC
This is an important test for differentiation of member of genus
Enterobacteriaceae, especially for differentiation of Enterobacter aerogenes from Enterobacter cloacae.

5. METHYL RED/VOGES PROSKAUER(MRVP)
Inoculation of this media, which is a broth, gives two results from one tube. First the MR reaction. This media contains glucose and phosphate buffer. The MR and VP tests are especially important in the identification of the gram negative enteric bacteria that ferment glucose by a metabolic process called MIXED ACID FERMENTATION to produce a variety of stable acid end products. These acid end products lower the pH of the media and are tested for in this part of the procedure. To do the test remove 2ml of incubated broth from the tube to a clean test tube. Add 5 drops of methyl red reagent to the 2ml you placed in the clean test tube. A positive reaction is a red or red-orange color upon addition of the MR reagent. Stable acid end products have been formed. No color change is negative for MR. The methyl red reagent is red at a pH of less than 4.4 and yellow at a pH of greater than 6.0.

VP test: The VP test is also used to detect fermentation products but of a different kind. Some bacteria ferment the glucose in the MRVP tube and produce the end products ACETOIN and 2,3-BUTANEDIOL. To do the VP test use the remaining volume in the tube, approximately 3 ml. To test for the presence of these end products you need to add two reagents to the remaining sample. FIRST add 10 drops of Barritt’s reagent A to the tube, followed by 10 drops of Barritt’s reagent B. Mix the tube to oxygenate the reaction. Allow the tube to sit for at least 30 minutes. A red layer will develop at the top of the tube. This is a positive VP reaction. No color development is a negative VP reaction. The VP test is very important and one that is often misread by students because they don’t want to be patient and wait for the results. The MR part of the test is immediate, the VP takes some time to develop. Just wait and get it correct, it could be the difference between getting the unknown right or wrong!

6. KLIGLER’S IRON AGAR (KIA)
This is a solid slant media that is used to differentiate bacteria based on their ability to ferment the sugars, lactose and glucose, and also to produce hydrogen sulfide, H2S. Gas production is determined by bubbles or cracks in the media. The media is inoculated with a stab/streak. There are several possible results for this media shown in the table below. In this table, butt refers to the bottom of the tube, slant refers to the slanted surface of the tube.

|Result |Interpretation |Data Notation |
|Yellow slant/yellow butt |Glucose and lactose fermentation |A/A |
|Red slant/yellow butt |Glucose fermentation, peptone catabolism |K/A |
|Red slant/red butt |No sugar fermentation, peptone catabolism |K/K |
|Yellow slant/yellow butt with bubbles or cracks|Glucose and lactose fermentation with gas production |A/A/G |
|Red slant/yellow butt with bubbles or cracks |Glucose fermentation only with gas production |K/A/G |
|Yellow slant/yellow butt with black precipitate|Glucose and lactose fermentation with hydrogen sulfide |A/A/H2S |
| |production | |
|Red slant/yellow butt with black precipitate |Glucose fermentation only and hydrogen sulfide production |K/A/H2S |
|Red slant/yellow butt, bubbles and black |Glucose fermentation only, gas and hydrogen sulfide |K/A/G/H2S |
|precipitate |production | |
|No change in slant or butt |No fermentation |NC/NC |

It’s a little complicated to read the results of the KIA reaction, but this is another very important diagnostic tool for unknown identification.

7. CITRATE AGAR
This media is a slant that it is inoculated with a stab/streak. The media will be dark green when inoculated. This media is used to determine if a bacteria has an enzyme called CITRASE that will allow the organism to break down the only carbon source in the media which is citrate, hence the name of the media. There is a pH indicator in the media, bromthymol blue, which is green at a pH of 7.6 or less. If the pH of the media is above 7.6 the indicator turns a deep blue color. If an organism can utilize citrate, the enzyme citrase converts citrate to oxaloacetic acid which is then converted to pyruvic acid with the release of CO2. As citrate catabolism continues, more CO2 is released and combines with other ingredients in the media to form alkaline compounds that causes the pH of the media to rise and the media changes to a deep blue color. So, a color change from green to blue is positive for citrate. It is not necessary for the entire tube to change to blue. Any blue color is considered a positive reaction.

8. PHENYLALANINE DEAMINASE AGAR(PDA)
This is another enzyme test. Does a given organism have an enzyme called phenylalanine deaminase? The media is supplied as a slant, and is inoculated with just the slant portion of the media being inoculated. This is also an easy test to read. The media contains the amino acid phenylalanine. In the presence of the deaminase enzyme the amino group (NH2) of the amino acid phenylalanine is removed and phenylalanine is converted to another compound called phenlypyruvic acid. This is the compound that you test for. To test for the presence of phenlypyruvic acid you add a few drops of 10% ferric chloride solution to the slant. If phenlypyruvic acid is present in just a few seconds the slant will turn a green color. Green is positive, no color change is negative for the enzyme phenylalanine deaminase. Member of genus Proteus, Morganella and Providencia are positive for this enzyme.

9. NITRATE REDUCTION TEST
This is another enzyme dependent reaction that can be a little hard to interpret correctly. The media is a broth and contains beef extract, peptone, and potassium nitrate. The media also contains a Durham tube to detect the production of nitrogen gas. There are no pH indicators in the media. You are testing for the enzyme NITRATE REDUCTASE which very simply converts nitrate (NO3) to nitrite (NO2). This is actually a form of anaerobic respiration. If nitrate is produced, it is converted to nitrous acid (HNO2) which is tested for in this procedure. Another process that can occur in this reaction is called DENITRIFICATION, in which nitrate and its products are reduced all the way to nitrogen gas which is trapped in the Durham tube. After inoculation and incubation look for gas in the Durham tube, if a bubble is present then you are done with the test, the organism is nitrate positive. This generally does not happen, and further testing is required. You now test for the presence of nitrous acid by the addition of two reagents to the tube. First add 10 drops of sulfanilic acid, follow by 10 drops of α-naphthylamine. If a red color develops then the test is POSITIVE for nitrate reduction. This color will develop very quickly, 30 seconds or less. If no color change is observed this DOES NOT mean the reaction is negative, at least not yet. If no color change is observed you add a SMALL amount of zinc dust to the tube and mix gently and wait. If there is a color change now, probably red or orange, this means that there is still nitrate in the media, NO nitrate reduction has occurred and the test now is considered NEGATIVE for nitrate reduction. HOWEVER, if NO color change is observed after zinc addition this means that the nitrate WAS reduced to some other compound that the reagent additions cannot test for, and the test is POSITIVE for nitrate reduction. The nitrate test has three possible positive reactions and one negative reaction. Confusing enough?

10. BILE ESCULIN AGAR
This test procedure is generally used for differentiation and identification of Gram positive bacteria, especially the group D streptococci and enterococci. Gram negative bacteria can also be identified with this media as well. The media is supplied as a slant that is almost the same color as TSA, maybe a little darker. The inoculation is done as a stab/streak. The media contains a compound called esculin which some bacteria are able to catabolize to another compound called esculetin. If esculetin is produced it reacts with ferric citrate in the media to from a dark brown/black color in the media. After inoculation and incubation if 50% or more of the media is black-brown then this is a positive exculin reaction. If, after a maximum of 72 hours of incubation the tube is not at least 50% black-brown the reaction is considered negative.

11. CATALASE(rapid substrate test)
This is a very easy and quick test. All you need to do is put some bacteria from a solid culture (plate or slant) on a microscope slide. Add one drop of hydrogen peroxide, H2O2, to the bacteria. If you see bubbles then the reaction is catalase positive, no bubbles is catalase negative. The reaction is usually very obvious, but sometimes you must look closely to see the bubbles. This reaction is very important for differentiating genus Staphylococcus from genus Streptococcus, and Enterococcus. All the members of genus Staphylococcus are catalase positive, while the members of genus Streptococcus, and Enterococcus are all catalase negative.

12. COAGULASE REACTION
This is THE test to identify the bacteria Staphylococcus aureus. All that you do is inoculate a small tube of rabbit plasma with solid growth bacteria. Place the tube in the incubator overnight, if the plasma has clotted (solid mass) after incubation the bacteria is Staphylococcus aureus. Staphylococcus aureus is the only coagulase positive bacteria with which we work.

Incubation Times for Differential Tests

The incubation times for some the tests just described can be important. Here are some guide lines for how long certain tests can be incubated and still give reliable results.

1. Phenol red sugars – 18-24 hours
2. MRVP – 5 days
3. Nitrate – 24-48 hours
4. Citrate – 24-72 hours
5. Lysine decarboxylase – 24-48 hours
6. Phenylalanine deaminase – 24-48 hours
7. Bile esculin agar – up to 72 hours
8. Urease – 24-48 hours
9. SIM – 24-48 hours
10. Kligler’s iron agar(KIA) – 24-48 hours
11. Coagulase reaction – overnight
12. Catalase – immediate

It is a good idea to try and observe these reaction times to get reliable results. It is possible, however, in some reactions to see the result in less than the prescribed time. For example, if you look at a urease tube in 18hrs and it’s turning pink, this is positive, it will only get more pink over time. If you don’t see the positive reaction in the allotted time then further incubation is not needed the reaction is just negative no matter what you would like it to be!

Data sheets for metabolic results:

Metabolic results 1

|Bacteria |Glucose |Sucrose |Lactose |Mannitol |Urease |Catalase |
|Alcaligenes faecalis | | | | | | |
|Citrobacter diversus | | | | | | |
|Citrobacter freundii | | | | | | |
|Enterobacter aerogenes | | | | | | |
|Enterobacter cloacae | | | | | | |
|E. coli | | | | | | |
|Klebsiella pneumoniae | | | | | | |
|Morganella morganii | | | | | | |
|Proteus mirabilis | | | | | | |
|Proteus vulgaris | | | | | | |
|Pseudomonas aeruginosa | | | | | | |
|Salmonella typhimurium | | | | | | |
|Shigella flexneri | | | | | | |
|Shigella sonnei | | | | | | |
|Serratia marcescens | | | | | | |
|Providencia stuartii | | | | | | |
|Bacillus cereus | | | | | | |
|Bacillus megaterium | | | | | | |
|Bacillus subtilis | | | | | | |
|Enterococcus faecalis | | | | | | |
|Enterococcus faecium | | | | | | |
|Gaffkya tetragena | | | | | | |
|Lactobacillus plantarum | | | | | | |
|Micrococcus luteus | | | | | | |
|Micrococcus roseus | | | | | | |
|Staphylococcus aureus | | | | | | |
|Staphylococcus epidermidis | | | | | | |
|Staphylococcus saprophyticus | | | | | | |
|Streptococcus lactis | | | | | | |
|Streptococcus salivaricus | | | | | | |
|Streptococcus agalactiae | | | | | | |

Metabolic results 2

|Bacteria |Citrate |Nitrate |SIM test |MR test |VP test |Coagulase |
|Alcaligenes faecalis | | | | | | |
|Citrobacter diversus | | | | | | |
|Citrobacter freundii | | | | | | |
|Enterobacter aerogenes | | | | | | |
|Enterobacter cloacae | | | | | | |
|E. coli | | | | | | |
|Klebsiella pneumoniae | | | | | | |
|Morganella morganii | | | | | | |
|Proteus mirabilis | | | | | | |
|Proteus vulgaris | | | | | | |
|Pseudomonas aeruginosa | | | | | | |
|Salmonella typhimurium | | | | | | |
|Shigella flexneri | | | | | | |
|Shigella sonnei | | | | | | |
|Serratia marcescens | | | | | | |
|Providencia stuartii | | | | | | |
|Bacillus cereus | | | | | | |
|Bacillus megaterium | | | | | | |
|Bacillus subtilis | | | | | | |
|Enterococcus faecalis | | | | | | |
|Enterococcus faecium | | | | | | |
|Gaffkya tetragena | | | | | | |
|Lactobacillus plantarum | | | | | | |
|Micrococcus luteus | | | | | | |
|Micrococcus roseus | | | | | | |
|Staphylococcus aureus | | | | | | |
|Staphylococcus epidermidis | | | | | | |
|Staphylococcus saprophyticus | | | | | | |
|Streptococcus lactis | | | | | | |
|Streptococcus salivaricus | | | | | | |
|Streptococcus agalactiae | | | | | | |

Metabolic results 3

|Bacteria |LDC test |PDA test |BEA |Kligler’s iron agar |
|Alcaligenes faecalis | | | | |
|Citrobacter diversus | | | | |
|Citrobacter freundii | | | | |
|Enterobacter aerogenes | | | | |
|Enterobacter cloacae | | | | |
|E. coli | | | | |
|Klebsiella pneumoniae | | | | |
|Morganella morganii | | | | |
|Proteus mirabilis | | | | |
|Proteus vulgaris | | | | |
|Pseudomonas aeruginosa | | | | |
|Salmonella typhimurium | | | | |
|Shigella flexneri | | | | |
|Shigella sonnei | | | | |
|Serratia marcescens | | | | |
|Providencia stuartii | | | | |
|Bacillus cereus | | | | |
|Bacillus megaterium | | | | |
|Bacillus subtilis | | | | |
|Enterococcus faecalis | | | | |
|Enterococcus faecium | | | | |
|Gaffkya tetragena | | | | |
|Lactobacillus plantarum | | | | |
|Micrococcus luteus | | | | |
|Micrococcus roseus | | | | |
|Staphylococcus aureus | | | | |
|Staphylococcus epidermidis | | | | |
|Staphylococcus saprophyticus | | | | |
|Streptococcus lactis | | | | |
|Streptococcus salivaricus | | | | |
|Streptococcus agalactiae | | | | |

Gram Negative Enteric Bacteria

The enteric bacteria are the organisms that live in your intestines, especially the large intestine. Many of the species are familiar to you, such as Escherichia, Enterobacter, and Proteus. Generally, these species are not pathogenic. However, there are other enteric pathogens of medical importance. These are members of genus Salmonella and Shigella. How can the various species be differentiated, and the pathogenic bacteria identified? That is the subject of this lab, and will provide you with the information that will aid you in the identification of your last set of unknowns.

You will be using differential and selective media to identify these organisms largely based on the ability or inability of an organism to ferment lactose. Many of the non-pathogenic enteric bacteria ferment lactose, while the pathogens do not. You will be inoculating several types of media in this lab designed to show these differences. These media are plates of:
1. Hektoen enteric agar (HE)
2. MacConkey agar (MAC)
3. Xylose Lysine Desoxycholate agar (XLD)
All of these media have special ingredients that help to differentiate and select for certain bacteria. None of these media would be used with Gram positive bacteria, unless it was to get rid of the Gram positive organisms. Here is a brief description of each type of media.

1. Hektoen enteric agar
The purpose of this media is to differentiate Salmonella and Shigella from other gram negative enteric bacteria. The media has lactose in it differentiate lactose fermenters from non-fermenters, and also other components that will indicate the production of hydrogen sulfide (H2S) by some bacteria. Color is the key to interpretation of this media. Bacteria that ferment lactose and produce acid, such as Enterobacter and Escherichia along with several others will produce colonies on this media that are bright yellow to salmon-pink in color. Bacteria which do not ferment lactose produce blue-green colonies, and some produce black colonies from H2S production. Member of genus Proteus and Salmonella, again, along with other species exhibit this type of growth on this media.

2. MacConkey agar
This is another media that is used to differentiate Gram negative lactose fermenters from non-fermenters. It is also a selective media in that it inhibits the growth of Gram positive bacteria. This media contains a pH indicator, neutral red. The indicator is red at a pH of 6.8 or less. Above this pH the indicator in colorless. When a lactose fermenting bacteria grows on this media it produces acid, the pH goes down, and colonies of lactose fermenting bacteria will appear red. Bacteria which do not ferment lactose produce colonies that have no color. The enteric non-pathogens ferment lactose, the enteric pathogens do not.

3. Xylose Lysine Desoxycholate agar
This last media is once again differential and selective. It is selective because Gram positive bacteria will not grow on this media, and differential in that it shows fermentation, deamination, and H2S production by various bacteria. Phenol red is the pH indicator in this media. Fermenting bacteria produce acid and yellow colonies, lysine deaminating bacteria produce red colonies, and still other bacteria produce black colonies from H2S production. For example, red colonies on this media are characteristic of members of genus Shigella, while red colonies with a black center are characteristic of genus Salmonella.

The key to using these media effectively is good streaking technique, isolating colonies that are easy to characterize, and sample for further testing.

LAB EXERCISE:
Each table will get a set of plates to inoculate. You will inoculate HE, MAC, and XLD plates. You will use the following bacteria: Escherichia coli Shigella flexneri Salmonella typhimurium Proteus mirabilis

Divide the plate into 4 quadrants and loop inoculate each plate with all four bacteria. Incubate at 37°C. Next lab we will examine the results.

RESULTS:
1. HE Escherichia coli Shigella flexneri Salmonella typhimurium Proteus mirabilis

2. MAC Escherichia coli Shigella flexneri Salmonella typhimurium Proteus mirabilis

3. XLD Escherichia coli Shigella flexneri Salmonella typhimurium Proteus mirabilis

Enteric Unknowns

You will be given a mixture of two bacteria. These bacteria will be a mixture of a lactose fermenting bacteria and one that does not ferment lactose. Using the differential and selective media discussed in the last lab and other diagnostic media that you have been using for your general unknowns, it is your job to identify both of these organisms. Other media that will be useful for this identification are SIM, urease, KIA, LDC, MRVP and citrate. Here is a list of the possible bacteria:

E. coli
Enterobacter aerogenes
Citrobacter freundii
Shigella flexneri
Proteus mirabilis
Proteus vulgaris
Klebsiella pneumoniae
Salmonella typhimurium

The first thing to do is streak the mixtures on the differential media that we used in the last lab, HE, MAC, and XLD. Pick different colonies based on fermentation results or other properties, grow them on KIA slants for working cultures, then use the other diagnostic media mentioned above to identify both bacteria. Sense the list of possible bacteria is not long, this identification should not be difficult. It really depends on techniques of isolation and proper interpretation of the results. Good luck!

-----------------------
Inoculate one area of the plate with the following bacteria:
Streptococcus lactis
Staphylococcus aureus
Staphylococcus epidermidis

B. stearothermophilus
P. aeruginosa
E. coli
S. aureus

B. subtilis
S. epidermidis
K. pneumoniae
S. marcescens

Staphylococcus aureus
Alcaligenes faecalis
Clostridum perfringens

Pseudomonas aeruginosa
Enterobacter aerogenes
Clostridium sporogenes

[pic]

This figure shows the analysis of a sexual assault case. DNA has been analyzed on an agarose gel. The gel acts as a molecular sieve to separate the DNA fragments based on size. The DNA bands are the thick, black bars on the photograph. Examine the pattern of the bands and see which suspect has some explaining to do.