SAN ANTONIO COLLEGE
BIOLOGY 2420
MICROBIOLOGY FOR
ALLIED HEALTH MAJORS
FRESHMAN LEVEL COURSE
Laboratory Manual
Xhavit Zogaj, Ph.D.
Office: Chance 345
Phone: 486-0840
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LABORATORY SUPPLEMENT
Please remember to budget your time and make the most of what little you have. Also, don't be timid. Roll up your sleeves
and dig in. When you come in to the lab get out all your lab supplies. Get out your Bunsen burners and any other equipment
you will need. Get out any experiments you have in progress from the previous lab. Do not sit at the bench studying for a
daily quiz instead of preparing to work. When you walk in the door, study time is over. Each week lab will begin with a
short quiz. Following the quiz, a short prelab lecture will be given. Because your labs are short, this lecture material is kept
to a minimum. However, if material is deemed worth mentioning by your instructor, you can insure it is information you will
want to record. Take notes during prelab!! Once prelab instructions are given, begin work quickly. If you make a mistake,
it will be a learning experience. Don't sweat it.
READ YOUR LAB MATERIAL BEFORE COMING TO CLASS!!!!!!!!!!!!!!!!!!
TAKE NOTES DURING PRELAB LECTURE!!!!!!! THIS IS ALL TESTABLE INFORMATION. This lab
supplement is provided because your lab book does not provide adequate explanation and background for most exercises.
However, enough material is provided between the book and prelab, that none of you should experience the confusion level
which was expressed prior to this supplement. The same procedural questions should not be asked repeatedly if the lab
material is read and notes are taken especially when the information has been given during prelab. Remember, the
microbiology laboratory is 25% of your final grade and cannot be ignored if you intend to perform well in this class.
DO NOT PUT PAPER TOWELS IN THE BIOHAZARD BUCKETS. THE ONLY THINGS IN THE BIOHAZARD
BUCKETS SHOULD BE PETRI PLATES CONTAINING LIVING ORGANISMS AND INFECTED SWABS.
PAPER TOWELS USED TO DISINFECT LAB BENCHES ARE NOT CONSIDERED BIOHAZARDOUS
MATERIAL.
REMEMBER BACTERIAL CULTURES ARE NOT REUSED UNLESS YOU ARE SPECIFICALLY INSTRUCTED
TO DO SO. ALL CULTURES IN GLASS TUBES SHOULD HAVE THE LABELS REMOVED AND THE TUBES
SHOULD BE PLACED IN ONE OF THE TEST TUBE RACKS ON THE CART AT THE BACK OF THE ROOM.
ANY CULTURES ON PETRI PLATES GO INTO THE BIOHAZARD BUCKETS.
REMEMBER EACH LAB SECTION HAS A SHELF DESIGNATED FOR LAB SUPPLIES AND SLIDE BOXES.
YOU SHOULD ALWAYS GET OUT ONE OF THOSE SLIDE BOXES PER TABLE AT THE BEGINNING OF
EACH LAB WHEN YOU NEED THEM, AND RETURN THOSE BOXES (CONTAINING ALL THE SLIDES YOU
HAVE USED AND THEN CLEANED) TO THE PROPER SHELF IN THE CABINET AT THE END OF EACH
LAB.
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SAFETY INFORMATION
All faculty, staff and students must follow the college safety policies and procedures while on campus. If the fire alarm
sounds during lecture or laboratory classes, everyone must evacuate and not return, until an authorized school representative
announces that it is safe to re-enter the building. Your laboratory instructor will conduct a general safety training session at
the beginning of your first laboratory class. Additional safety training and procedures for specific laboratory activities will be
discussed as needed by your laboratory instructor as you progress through the semester.
The following rules apply to all microbiology lab sessions:
1. Read your laboratory manual prior to the beginning of each laboratory. Wash your hands and disinfect your lab
bench at the beginning of each laboratory session.
2. Wear clothing which you can risk being ruined by the numerous biological stains present in the laboratory.
3. Absolutely no eating or drinking in the laboratory. Do not even bring food or beverage containers into the room. Those
materials must be discarded outside the laboratory. Liquid or solid food wastes in the bottom of trash receptacles serve as the
perfect medium for growing microbes.
4. Gas burners will be in use. Be extremely careful with hair and clothing. Individuals with long hair should pull the hair
back with a barrette or rubber band for safety. Make sure you know where the emergency fire blanket, fire extinguisher and
shower are located.
5. Treat every culture as if it is extremely pathogenic and use aseptic technique at all times. Dispose of all cultures and contaminated materials in the proper receptacles for sterilization. Never pour an unsterilized culture in the drain or in the
trash. Any student who is immunocompromised for any reason should inform the instructor as well as your personal
physician so that appropriate safety measures can be implemented to insure your health.
6. Use proper labeling procedures for any cultures you create. Disposable plastics should be labeled directly with a sharpie.
Glass items (test tubes, flasks, etc.) should have a tape label. Proper labeling includes your last name, course section (2420001, 2420-005, etc.), date, instructor's name and organism name or number. All labels must be removed before
discarding cultures.
7. At the beginning and end of each laboratory session, swab your work area with the disinfectant provided.
8. All purses, book bags, umbrellas and other personal belongings will be stowed in a cabinet at the front of the laboratory.
At your lab bench, you will only need lab supplies and your lab book. This rule is to maintain clear, safe walkways.
9. Each lab section will have a number and a specific area in which to incubate or refrigerate lab experiments. Ask your
professor where your assigned areas are.
10. Wash your hands at the end of each laboratory session and disinfect your lab bench.
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Aseptic Technique and Culture Handling
BACKGROUND

Subculturing is the process by which microbes are transferred from one medium to another using an inoculating
instrument of some kind. Aseptic techniques must be applied at all times when subculturing to insure the original
stock and the new culture do not become contaminated.

When pouring a Petri plate using molten agar in a tube, be sure to temper or cool the agar to ~50 degrees prior to
pouring. Use a paper towel to wipe off any water from the outside of the tube which you usually cool in the water bath.
This prevents contaminated water from dripping into your sterile Petri plate while pouring. Place a sterile Petri plate in
front of you with the lid up. Remove the cap from the tube of agar and flame the lip of the tube. As soon as you remove
the lid, discard the cap (This is the only time in Micro lab when it is OK to put test tube caps on your lab bench).
Raise the lid of the Petri plate, and pour the molten agar into the bottom of the dish while holding the Petri dish lid
above the tube and plate bottom. This prevents contaminants from falling into the freshly poured agar. Leave the Petri
plate stationary until solidified. If you stack the plates, it will take longer for them to cool. Once solidified, the agar
will have an opaque appearance.

When preparing a streak plate, label the bottom of the Petri plate with a sharpie. Place the Petri plate upside down in
front of you.

Sterilize an inoculating loop and remove a small sample of bacteria from the available culture using aseptic technique.
Remember to recap or close the culture once you have removed your sample and before you begin the streak plate. (You
will only go into the primary culture for the first quadrant streak. Do not add more culture for each quadrant!)
Lift the bottom of the Petri plate containing the agar in your hand, leaving the lid on the bench. Streak quadrant #1 so
that the bacteria are evenly spread. Then put the agar plate back onto the lid. (Never put the lid on the bench so that the
inside of the lid faces the bench. The inside of the lid will pick up bacteria from the bench) Flame the loop. When the
loop is cool, touch one corner of quadrant #1 pulling a sample into quadrant #2 and spread evenly. (You can test the
coolness of your loop by touching the agar around the edge of the Petri plate. If it sizzles, it is too hot. If it does not
sizzle, it is ready.) Once you have streaked quadrant #2, flame the loop. When cool, pull a small amount of material
from quadrant #2 into the third quadrant, and streak evenly. I recommend a fourth quadrant streak for beginning
students since most beginners use too much initial inoculum. The goal here is to dilute out the number of bacteria you are
moving with the loop with each additional streak. By the end, you should have only a few cells in the 3rd or 4th
quadrants. Each of these individual cells will begin dividing by binary fission, producing a population of clones that pile
on top of each other and form what is called a colony.

When you select a colony, you can be sure that you have generated a pure culture. This will be your most immediate
goal for your unknown. If you have two unknown organisms, the streak plate technique will be even more important
because you want an isolated colony of each organism to make your working stocks. When streaking a plate be careful
not to dig into the agar. Always incubate all bacterial plates in an inverted position. This prevents condensation
from gathering on the lid and dripping onto the agar surface. If you get a lot of moisture buildup on the agar
surface, you will not have colonies; you will generate a bacterial lawn. Please stack the Petri plates in the
incubators to conserve space and be sure to put your plates on the appropriate shelf for your lab section.

When inoculating a slant, select bacteria using a sterile loop and aseptic technique. Flame the lip of a sterile agar slant
keeping the cap in the crook of your pinkie finger. Do not put the cap on the bench because you will pick up
contaminants. (Remember to always check the agar and broth cultures you are inoculating to insure there are no
contaminating organisms already growing) Begin at the bottom of the slant. Touch the agar surface and make a
squiggle pattern across the agar surface moving up toward the opening of the tube. Be careful not to gouge the agar
surface. Likewise, when removing bacteria from an agar slant, do not dig into the agar with the loop. Slants should be
labeled with your name, lab section, date, instructor's name and organism name. When working with aerobic or
facultatively anaerobic organisms, keep the cap slightly loose during incubation.
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
When inoculating a deep, use an inoculating needle. (If you don't have a needle you can straighten out a loop). Flame
the needle and remove a small amount of bacterial growth. You will not be able to see the inoculum on the needle
unless you have taken too much. Stab the inoculum through the agar or gelatin. Go straight in and straight out. If your
inoculating needle is short, be careful not to stab the handle of the inoculating needle into the agar because you have
probably not sterilized the handle. After bacteria grow in an agar deep, you will see growth along the stab line when
you hold the tube up to the light. If the organism is also motile, you will see outward motility of the organism from the
initial stab line throughout the agar. The pattern is wider at the top of the tube (inverted Christmas tree pattern) since
most of our organisms are aerobes and fastest growth will occur toward the top of the tube. In gelatin, you will find that
some bacteria are capable of breaking down protein in the gelatin resulting in liquefaction. Stabs should be labeled with
your name, lab section, date, instructor's name and organism name. When working with aerobic or facultatively
anaerobic organisms, keep the cap slightly loose during incubation.

When inoculating liquid broth, remove an inoculum of bacteria from a culture using a sterile inoculating loop. Flame the
lip of the sterile broth culture and tilt the tube so that the liquid can be reached with the inoculating loop without having
to put the handle of the loop into the tube. Twirl the loop around so that bacteria are removed. You will not be able to
see the inoculum in the liquid unless you have added too much. Another technique is to transfer the inoculum from the
loop to the inside wall of the test tube you are inoculating. This will usually only work if the inside of the tube is not wet
which means you should not shake or tilt the tube prior to this procedure. Once you have an inoculum stuck to the inside
glass wall of a tube, you can tilt the tube so that the bacteria are washed from the wall and into the medium. Again with
this technique, the goal is to prevent introduction of the inoculating loop handle into the sterile tube. Heat the handle of
an inoculating loop to cut down on possible contamination. It is often impossible not to introduce the handle into a
culture if you are working with those large tubes. For this reason, try not to touch the handle of the loop against the
mouth of the tube. Never invert liquid cultures so that liquid contacts the cap. This procedure will lead to contamination
and spillage if the cap is not tight. Broth cultures should be labeled with your name, lab section and organism name.
When working with an aerobic organism, keep the cap slightly loose during incubation.
PURPOSE
•
To learn how to handle cultures in an aseptic manner. A pure culture is a specific microbe grown on
a medium. A medium is a collection of nutrients (solidified or liquid) used to support microbial
growth. You will be able to manipulate cultures without introducing contaminants into the culture
and without contaminating yourself or the environment. Also to learn how to inoculate various types
of media and to identify various types of growth.
CULTURES
•
Escherichia coli, Serratia marcescens, Micrococcus luteus
E. coli is an enteric organism [isolated from intestines of animals]. The colonies are beige and
mucoid. The cells are very short bacilli with peritrichous flagella. S. marcescens is also an enteric.
The colonies are beige/pink at 37oC [the organism’s optimum temperature] and red at 25oC. The
cells are very short bacilli with peritrichous flagella. E. coli and S. marcescens are opportunistic
pathogens. M. luteus is an organism found in soil and on skin of some animals. It is not a pathogen.
The colonies are citron (sunshine) yellow, have round margins and are raised (convex). The cells
are cocci which occur in the arrangement of tetrads. Most often, the cells will appear to be in the
staphylococcus arrangement when taken from agar. M. luteus is nonmotile, as are all cocci.
SUPPLIES
Bunsen burner, inoculating needle and loop, striker
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MEDIA
•
Tryptic soy broth (TSB), Tryptic soy agar (TSA) plates, TSA slants, Motility agar deeps
TSB, TSA slants and Motility agar are premade, sterilized and stored on the shelves at the front of the lab.
You must learn the proper method for pouring TSA plates using pre-sterilized TSA deeps. Melt TSA deeps
in the boiling water baths at the back of the lab. Agar melts at 100oC (boiling). Once melted the deeps are
transferred to the 50oC holding tank where they will be cooled. Agar will solidify at 45oC, so the media will
remain liquid at 50oC. When cooled, remove tubes from water bath and dry tubes with paper towels. This is
important to remove non-sterile water from the outside of the tube that can drip into your sterile Petri plates.
Remove the lid of the tube and flame the lip of the tube. Pour the agar into a sterile Petri plate holding the lid
of the Petri plate over the tube while pouring. Petri plates are stored on the shelves to the left of the water
baths. Always close the Petri plate bags after opening them.
CULTURE CONDITIONS

TSB is inoculated with an inoculating loop and incubated at 37oC for E. coli and M. luteus (25oC for
S. marcescens) with the caps loose. TSA plates are inoculated using the 4 quadrant streak plate
technique and incubated in an inverted position at temperatures as indicated above. Motility agar is
stabbed using an inoculating needle and incubated with loose caps as above. TSA slants are
inoculated by streaking the slant from bottom to top with bacteria using an inoculating loop.
Slants are incubated at 37oC with loose caps unless otherwise instructed.
STUDENT TASK
·
Pour two TSA (Tryptic Soy Agar) plates using aseptic technique
·
Inoculate 1 TSB (Tryptic Soy Broth) using M. luteus (37oC)
·
Stab inoculate one motility agar deep using E. coli (37oC)
·
Streak one TSA slant using any of the three organisms and incubate at the appropriate temperature.
·
Once solidified, streak the two TSA plates using the four quadrant method. One plate will be S.
marcescens (25oC) and the other can be either 37oC preference organism.
·
Leave all caps loose and store at appropriate temperatures. Freshly poured agar tears easily.
Use a very light touch with the inoculating loop.
REACTIONS

TSB is a general purpose liquid medium used to grow cultures. Growth will cause the broth to
become turbid (cloudy); however, you cannot tell by visual inspection if the culture is
contaminated. The streak plate technique is powerful for the generation of pure cultures. This
technique provides isolated colonies, each represents a single cell that was isolated away from the
population and grew by binary fission to create a visible clump on agar. Motility is the ability of
bacteria to move. In bacteria, this is most commonly associated with flagella. Motility agar is
relatively clear and allows you to visualize movement of bacteria through the agar (producing
cloudy agar) in all directions away from the stab line. In TSA slants look for smooth, complete
coverage of the slant without contamination. These slants are perfect for storing large numbers of
organisms on a relatively large surface that is sealable with a lid. Cultures may be preserved for
long periods of time in the refrigerator on slants.
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RESULTS

Look for turbidity in TSB cultures. S. marcescens may produce the red pigment giving the broth a
red appearance. On TSA plates, look for isolated colonies. Contaminants usually are of a
different color or texture and often do not lie on the streak line. Motility is often very subtle. It
must move in all directions from stab line. If you shake when inoculating the agar, there will be a
wide, ribbon-like pattern of growth that can be mistaken for motility, however it will only be wide
in one direction. Do not mistake motility for the inverted Christmas tree pattern that can be seen
when organisms grow faster (divide by binary fission at a faster rate) and therefore spread out across
the surface of the agar faster than deeper within the agar. M. luteus is nonmotile; E. coli and S.
marcescens are motile.
____________________________________________________________________________________
Microscopy
BACKGROUND
DO NOT PUT OIL ON THE 40x OBJECTIVES. The microscopes were recently cleaned and should be kept clean henceforth.
THESE ARE OTHER PROBLEMS OFTEN SEEN CONCERNING MICROSCOPE USE:
1. MICROSCOPES ARE TO BE STORED IN PROPER SLOTS. EACH MICROSCOPE HAS A NUMBER AND A
CORRESPONDING SPOT IN THE MICROSCOPE CABINET. EACH STUDENT WILL BE ASSIGNED A
MICROSCOPE AT THE BEGINNING OF THE SEMESTER. IT IS YOUR RESPONSIBILITY TO KEEP THIS
MICROSCOPE CLEAN AND REPORT ANY MISUSE BY STUDENTS IN OTHER LAB SECTIONS.
2. MICROSCOPES ARE NOT TO BE STORED WITH THE OIL IMMERSION LENS IN POSITION AND THE
STAGE UP. THE STAGE SHOULD BE AT THE LOWEST SETTING AND THE 4X OBJECTIVE SHOULD BE IN
POSITION. DO NOT WIND THE CORDS AROUND THE MICROSCOPES BECAUSE THIS MAY DISLODGE THE
ILLUMINATOR AND THE CONDENSER. THESE STRUCTURES ARE VERY WORN NOW BECAUSE OF THIS
PRACTICE.
3. DO NOT LEAVE SLIDES ON THE MICROSCOPES. CLEAN OFF ANY RESIDUAL OIL FROM THE STAGE
AND OBJECTIVES PRIOR TO STORAGE.
4. NEVER MOVE THE COARSE ADJUSTMENT WHEN THE OIL IMMERSION LENS IS IN POSITION. MANY
BEGINNING STUDENTS PLACE OIL ON THE SLIDES, THEN MOVE THE STAGE UP UNTIL THE LENS
TOUCHES THE OBJECTIVE. DO NOT USE THIS PRACTICE. YOU RISK BREAKING THE SLIDE AND THE
OBJECTIVE. YOU WILL ALSO EXPERIENCE DIFFICULTY IN FINDING MICROBES IN A TIMELY FASHION.
The binocular brightfield light microscope is shown in the textbook. Be familiar with all its features. First binocular means the
scope has two ocular lenses located at the top of the microscope head. These ocular or eyepiece lenses can be moved in and out
so that they will be positioned at the perfect width for your eyes. It is important that when you look through the microscope you
only see one circle of light. The eyepieces must be adjusted until the two circles become one. Most of the microscopes have a
pointer in the left ocular lens which can be moved by rotating the eyepiece. This will allow you to point out objects to your
instructor when you ask questions. The ocular lenses must be clean otherwise you will see large brown spots covering the
microbes. If you want to check for cleanliness, rotate each ocular while looking through the scope. If the brown or dark spots
rotate, you know they are on the lenses. Clean the ocular lenses by using lens paper and ethyl alcohol and rub firmly until the glass
squeaks. Be sure to get around the rim of the glass also. If finger print oils and mascara are on the lenses, you may have to clean
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several times. The magnification of the ocular lenses is 10X. This number may be multiplied by the magnification of the objective
in use to determine the total magnification.
The head of the microscope will pivot so that you can show images to your instructor without completely moving the whole scope.
This will insure that you have not accidentally moved the stage or slide. To rotate the head, loosen the silver screw on the side of
the head. Be sure to tighten the screw at the end of the day because if too loose, the entire headpiece will fall off when you tilt the
scope to return it to its cabinet. If this happens, not only will the head hit the floor, but the ocular lenses will also fly off and can
be damaged or broken.
Below the head, the microscope also has 4 objectives. The smallest one with the red ring at the top is the 4X objective. The best
use of this objective is to use it to scan the surface of a slide looking for color or objects on which to focus. Once you have located
an object with the 4X lens, higher magnification can be used. The next objective with the yellow ring is the 10X objective. This
can also be used for the purpose of scanning or observing large organisms like molds. The third objective is the 40X objective
also called the High Dry lens. This objective is the highest power objective on the scope which should never come in contact
with immersion oil. This objective is long and has a blue ring around it. Always check this objective for oil before using the
microscope. If any oil residue is on the lens, you will never be able to clearly focus on an object. The fourth objective is the oil
immersion objective or 100X. This objective will provide you with enough magnification power to see bacterial cells. This is
the only objective which should come into direct contact with immersion oil. Each objective should be cleaned thoroughly prior to
microscope use. This is best achieved by using ethyl alcohol on lens paper. Use a different piece of lens paper for each objective
to insure that oily residue is not picked up from one objective and transferred to another. Rub the lenses firmly with the lens paper
and alcohol and use a dry piece to dry the surface. The 100X objective may need to be cleaned several times.
The stage is the flat black platform directly below the objectives. There is a hole in the center that will allow light to pass through
your slides. If you look through the hole, you will see the substage condenser directly beneath the stage. The stage is mechanical
which means the slide clamps are controlled by two black knobs beneath the microscope stage on the right. The top knob moves
the slide front to back. The bottom knob moves the slide left and right. The mechanical stage will allow you to easily manipulate
the positioning of the slide. The position of the stage can be controlled by the two large knobs on the arm of the microscope.
These are the fine and coarse adjustment. The coarse adjustment moves the stage up and down in large increments and is to be
used only when the 4X or 10X objectives are in place. The fine adjustment can be used at any time but is most useful when using
the 40X and 100X objectives. The fine adjustment moves the stages in very fine increments allowing you to fine tune the focus on
an object.
Also beneath the stage on the left is a single black knob that controls the substage condenser. The substage condenser functions
by focusing light on a small area above the stage where the specimen is located. By moving the condenser down toward the
microscope base, the amount of light focused on the slide will decrease. By moving the condenser to the highest position, the
maximum light will be achieved. You may also control the amount of light entering the condenser by manipulating the iris
diaphragm. The iris diaphragm is located in the middle of the condenser and is controlled by a ring or lever on the front of the
condenser. With the microscope light on, look through the hole in the stage and watch the light while opening and closing the iris
diaphragm. You will see that the iris diaphragm works like the pupil of your eye by widening and narrowing to control the amount
of light passing through the condenser.
Below the condenser on the base of the microscope, you will find the illuminator. This is the light source. The source may be
turned on using the black wheel on the base to the left. Turn the knob until you hear a click then continue to turn the wheel until it
cannot be turned any more. You may notice that the black plastic covers over the light source are cracked and chipped. Many of
them are taped on. This damage is largely the result of microscope cords being wrapped around this area. When the cords are
unwound, the illuminator covers were damaged often ripped from the base and bounce on the floor. Please do not wrap the
microscope cords around the microscopes. This practice can also damage the substage condenser.
Below is the procedure for viewing slides using the binocular brightfield microscope.
1. Remove the microscope from the case carrying it with one hand on the arm, one hand beneath the base.
2. Clean the objectives and ocular lenses thoroughly.
3. Move the stage to the lowest point and place the slide specimen on the stage and secure with the silver clip.
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4. Turn on the illuminator.
5. Make sure the 4X objective is in place and move the stage up while looking through the ocular lenses until the specimen is in
focus.
6. Move to the 10X objective and refocus using the fine adjustment.
7. Move to the 40X objective making sure to only use the fine adjustment to focus.
8. Move to a position between the 40X and 100X objective and place a drop of immersion oil on the slide where the light is
coming through it.
9. Click the 100X objective lens into position. The 100X objective will touch the oil. Use the fine adjustment only to focus on
the specimen.
10. Adjust the condenser and substage diaphragm to insure you get the best possible image.
11. When through, move the 4X objective back in place. Lower the stage. Remove the slide.
12. Clean the microscope. Store in the cabinet with the 4X objective in place and the stage at the lowest setting.
PURPOSE

To demonstrate the use of the binocular brightfield light microscope and use of the oil
immersion lens. Students should also appreciate the size of bacteria (Prokaryotes) in relation to
molds, yeasts and protozoans (Eukaryotes).
CULTURES

None required. Use prepared slides. Observe all assigned slides. Proficiency in handling the
microscope at this stage is absolutely necessary for almost every exercise following this one.
SUPPLIES
·
Prepared microscope slides, paper towels, lens paper, immersion oil, ethyl alcohol.
Magnification
4X
4X
4X
10X
10X
10X
100X
100X
100X
100X
100X
100X
10X
10X
The following slides will be viewed by each student
Slide # Specimen
1
letter “e” - to appreciate that the image you see is inverted
2
silk fibers - to appreciate depth perception with the microscope
3
flea - further emphasized depth of this “large” organism
4
Mold sporangia
5
Mold zygospores
6
Mold conidia
13 or 50
Typical bacillus
14
Typical staphylococcus
15
Typical spiral
19
Peritrichous flagella
35
E. coli
45
Treponema
24
Amoeba proteus
26
Paramecium caudatum
TECHNIQUES
Use of oil immersion lens.
REACTIONS
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
Immersion oil has the same refractive index as glass, which means as the light leaves the prepared
specimen slide it does not bend if it passes through oil as it would if it passed through air on its way
to the objective lens. The absence of light scattering increases resolution. Use 100X (oil
immersion) for observing all bacterial slides. Use 4X and 10X for observing molds. The 40X may
be necessary for yeasts and protozoans.
RESULTS

The microscopes are parfocal, which means that once you have focused using one of the low power
objectives, you can switch to any other objective and still be pretty much in focus. You might have
to use the fine adjustment, but that is the only adjustment needed. Never use the coarse focus once
you have gone to the 40X or 100X lens.
____________________________________________________________________________________
Examination of Living Microbes
BACKGROUND

Motility is the ability of organisms to self propel. Motile bacteria include some bacilli and spiral organisms. Cocci are
never motile. Eucaryotic cells may also be motile. Do not confuse Brownian movement with true motility. Brownian
movement results from the bombardment of bacteria with randomly moving water molecules. If you are using the oil
immersion lens on a thick preparation, the objective depressing on the coverslip can cause fluid to rapidly flow past your
field of vision. Do not mistake these occurrences for motility.

Some bacteria possess flagella which are long hair-like structures extending from the bacterial cell wall. Bacteria with
peritrichous flagella have these projections all over the surface of the cells. Some bacteria have only one, two or a tuft
of flagella extending from one or both ends. Other bacteria such as the helical spirochetes possess axial filaments.
These are modified flagella that wrap around the cell giving it the spiral shape and causing it to corkscrew through
liquid. Lastly, although we will not see an example of these, some bacteria can glide over moist surfaces.

Eucaryotic cells may also possess flagella (Euglena) although eucaryotic flagella are structurally unique from bacterial
flagella. Eucaryotes may also possess cilia (Paramecium) which are short projections extending from the entire surface
of the cell which move in wave-like motions. Other eucaryotes exhibit amoeboid movement (Amoeba) which means
they ooze over a surface by extending a projection called a pseudopod (false foot). A process called cytoplasmic
streaming then occurs, and the body of the organism flows toward the direction of the pseudopod.

The purpose of the hanging drop slide is to confine protozoans and bacteria within the three dimensional area of a drop
of liquid. This technique is helpful for identifying motile bacteria, for observing motility of larger organisms over a
longer period of time and for providing a more realistic view of motility compared to a wet mount. The drawbacks to
using a wet mount are that the preparation dries up quickly, and the organisms are not confined to one area, so they
quickly move out of the field of vision.

A very tiny drop of liquid is best for preparing hanging drop slides. Large drops tend to move around easily and will
touch the bottom of the slide depression. When this happens, the slide is ruined and must be done again. Large drops
are best for wet mounts because they evaporate quickly.

Remember when observing unstained microbes, you need to lower the light coming through the specimen so that you
have enough contrast between organism and background. Depending on the size of the drop you have generated, you
may or may not be able to use oil immersion. Proceed with caution. If you use too much Vaseline, the slide will be too
thick to fit under the oil immersion. Be careful not to break the coverslips. Place the Vaseline on the coverslip rather
than on the depression slide. When discarding the hanging drop slide, keep in mind that you are working with a living
10
culture. Discard all coverslips in the red plastic biohazard container for sharps. Then clean the depression slides with
disinfectant.
PURPOSE

To demonstrate the difficulty in observing living unstained cells, especially if they are motile. Also
to appreciate the size difference between eucaryotic and procaryotic cells. To be able to identify
motility in procaryotes and eucaryotes.
CULTURES

Pseudomonas aeruginosa, Euglena, Amoeba proteus, Paramecium caudatum
P. aeruginosa is an extremely opportunistic pathogen which is ubiquitous in the environment.
This organism causes many nosocomial infections including infections in CF (cystic fibrosis)
patients, burn patients and individuals with indwelling instruments. The colonies are very mucoid,
produce fluorescent green pigments and have a very distinctive odor. Euglena is a small
protozoan having a single flagellum for motility. It is photosynthetic when growing in the
presence of sunlight and the chlorophyll is visible. Amoeba are large protozoans which move by
amoeboid movement through the formation of pseudopods and cytoplasmic streaming.
Paramecium is also a large protozoan which is ciliated.
SUPPLIES

Slides, Cover slips, Toothpicks, Depression slides, Inoculating loops, Transfer pipettes,
Vaseline, paper towels.
TECHNIQUES

Wet mounts and Hanging Drop Slides. (Practice correct culture handling techniques with
bacterial cultures)

A wet mount is generated by adding a generous drop of culture to a clean slide and adding a cover
slip on top of the drop. You don’t want so much fluid that the cover slip floats or fluid flows past
the edges of the cover slip. However, the wet mounts will dry out quickly so more fluid is better.
Beware of air bubbles. Find the organisms quickly. If the slide begins to dry out, make another.
Cover slips are discarded in sharps containers. Slides are washed with disinfectant followed
by soap and reused. A hanging drop slide is generated by adding Vaseline to 4 sides of a cover
slip. Then place the cover slip on the bench (Vaseline up) and add a loopful of culture to the
coverslip. (It is very important to use a loopful of culture. A drop will not work.) Then place the
depression slide (depression side down) on top of the cover slip. As soon as the Vaseline touches
the slide and the cover slip sticks, do not press down further. Flip the hanging drop slide over
quickly and observe the organisms moving in the suspended drop. Depression slides are very
expensive. Be careful with them, wash them well and return them to the box. Use oil immersion
lens for observing the bacterial cultures. You only need 10X for Paramecium and Amoeba. 40X
11
may be useful for Euglena. Be sure to reduce the light intensity of the microscope to observe
living unstained cells.
STUDENT TASK

Prepare a wet mount of the hay infusion culture (or individual cultures of protozoans) and look for
Amoeba on 4X or 10X. The Amoeba will be large and slow moving. Look for cytoplasmic
streaming. Go to 40X for Paramecium and Euglena. Paramecium will be a constant oval shape
and is smaller than Euglena. Euglena may have a green appearance and is long and tapered on one
end. Euglena will also pulsate by rounding and extending. Go to 100X and observe protozoans for
contractile vacuoles. Observe Paramecium for cilia and Euglena for a flagellum. The entire
flagellum will not be visible because it is so thin; however, you will see water displacement from
the beating motion. Be aware that eucaryotic flagella do not rotate like procaryotic flagella. Instead
eucaryotic flagella wave back and forth like a cat’s tail.

Prepare a hanging drop slide on Pseudomonas aeruginosa. Starting on 4X, look for the border of
the water drop. It will look like a crooked brown line. Move the slide so that the line is dead center
in the field of vision. Then move to the next objective and again find the border. Keep moving the
border line to the center of the view as you move up to 40X. If you cannot find the border, ask for
help. If you do not have the border in view when you go to 100X, you will never find your cells.
Once you make it to 100X, look for runs and tumbles. No flagella will be visible. Notice the size
difference of these procaryotes compared to the eucaryotic protozoans.
REACTIONS

Note that the protozoans will try to move out of your field of vision because they do not like the
light and heat. Using the protozoans, you will be able to observe the three types of eucaryotic
motility. Flagellar, amoeboid, and ciliary motility will be observed. Flagellar motility involves
flagella. It should be noted that bacterial and eucaryotic flagella are completely different
structurally. Amoeboid motility involves the formation of pseudopods and cytoplasmic streaming
to allow eucaryotes to ooze across surfaces. Ciliary motility involves cilia, which are short hair
like structures found only in eucaryotes. There are also three types of bacterial motility. You will
see flagellar motility by P. aeruginosa. The second type is corkscrew motility which is
demonstrated by the spirochetes which have axial filaments. The third type of bacterial motility is
gliding where an organism slides across a moist surface. These forms of motility are not to be
mistaken for Brownian movement which is not true motility.
RESULTS

Be sure to observe for contractile vacuoles in the protozoans. These organelles are involved in
pumping water out of the cells to counteract the build up of osmotic pressure. P. aeruginosa is a
motile bacillus.
____________________________________________________________________________________
Fungi: Yeasts and Molds
12
BACKGROUND

Molds are multicellular, filamentous fungi. The mold colony is called a thallus. The thalli are composed of bundles of
filaments called mycelia. These vegetative structures are composed of long filament-like structures called hyphae.
Vegetative hyphae grow on the agar surface; rhizoidal hyphae grow below the surface; reproductive hyphae extend into
the air. Hyphae are divided into individual cells by wall-like structures called septa. Molds that have these septa are
comprised of septate hyphae. Hyphae that lack septa are called coenocytic hyphae. The reproductive hyphae bear
sexual or asexual reproductive structures. The reproductive structures often appear as flower like structures of sacs.
These sacs are filled with spores. You will also be able to see spores that have broken free. When observing the mold
spores under oil immersion, you should be able to see new hyphae being formed from the fungal spores. Each
spore you see has the potential to produce a new organism. (Remember the production of spores by molds is a
method of sexual or asexual reproduction.)

When observing the yeast cells under 40X or oil immersion, you should be able to see the cells' nuclei as well as
budding. Over time, most of the buds detached themselves from the parent. Budding is a form of asexual
reproduction. Some yeast cells produce buds that did not detach themselves. These structures give the yeast cells the
appearance of producing hyphae, but they are not true hyphae. These structures are called pseudohyphae. Also
remember pathogenic yeasts exhibit dimorphism. Wet mounts will be performed using 1 drop each of water and Gram's
iodine. Fungi will then be added to the mixture and covered by a coverslip.

When you inoculate the mold spores onto the TSA plate, each spore produces a new mold. Even though you may not
see anything on the inoculating loop, you will probably add thousands of spores to the agar. Each of these produces a
new organism, which in turn produces more spores, you will see the mold spread out and cover the entire surface of the
plate.

Each individual cell streaked onto an agar surface will begin reproducing (by budding in the case of yeasts; by binary
fission in the case of bacteria) and will produce a visible clump of cells called a colony. Colonies should be separated
enough on a streak plate that you could remove one colony using your inoculating loop. The word colony is used only
to describe growth of microbes on an agar surface. Since each cell making up a colony is a clone of the original cell you
isolated, each cell is identical in its characteristics. The word colony is not used to describe cells growing in liquid
culture. After growth of yeast colonies on Sabouraud agar, notice that the colonies are very large. These colonies are
much larger than bacterial colonies you will see in upcoming labs because eucaryotic cells are larger than procaryotic
cells.

Fermentation by yeasts involves the conversion of carbohydrates to some other organic product such as alcohol.
Fermentation tests of yeasts are read as positive if the medium becomes turbid and carbon dioxide is produced. During
fermentation carbon dioxide is formed, and the media becomes acidified. As a result, you can visualize fermentation
indirectly by looking for the formation of a carbon dioxide bubble inside the Durham tube (the gas will not cause the
tube to rise). Additionally you can detect acidification by noticing the development of yellow color in the fermentation
tube. Acidification may occur if an organic acid byproduct is produced instead of alcohol. Alcohol production often
does not change the color of the medium. The medium contains phenol red (a pH indicator) which is red at neutral and
alkaline pH and yellow at acidic pH. In order to judge whether anything has happened in your fermentation
tubes, you must first be able to see growth. Growth in a liquid culture is noted as turbidity (cloudiness). If you do
not see turbidity, but you see a color change, something is wrong. You cannot judge any metabolic event in this class if
you cannot confirm growth. If you allow your fermentation tube to sit for 48 hours or longer prior to observation,
alcohol can be metabolized and converted to an alkaline product by microbes resulting in alkalinization of the medium.
In this case the color of the medium will change back to red.

Candida albicans is a yeast found in healthy individuals as normal flora. However, under conditions where this
organism overgrows or when an individual is immunocompromised, oral and genital candidiasis can result.
PURPOSE
To observe living fungal cultures and learn how to cultivate them.
13
CULTURES

Aspergillus niger, Saccharomyces, Baker’s yeast, Tongue culture
A. niger is a black bread mold which is fast growing and produces conidiospores arranged in chains
at the end of a conidiophore. Saccharomyces is a yeast which is commonly used in the bread
industry and ferments sugars. Baker’s yeast is a fermentative yeast as well. It is normal for some
people to have yeasts as part of the normal flora of their mouths and urogenital tracts. We will use
sterile swabs to take tongue cultures.
SUPPLIES

Clean slides, Cover slips, Inoculating instruments, Distilled water, Gram’s iodine
MEDIA
Sabouraud Dextrose Agar, Glucose and Sucrose fermentation tubes
Sabouraud agar is a low pH, high sugar medium which is selective for fungi.
CULTURE CONDITIONS

Fungal cultures will be cultivated at 25oC, which is the preferred temperature. Do not invert
the plates in the incubator. Leave caps loose since fungi are aerobic organisms. Inoculate
fermentation broths using an inoculating loop. Streak Saccharomyces on a plate using the
streak plate technique. Inoculate A. niger onto a slant by spot inoculation. Use a sterile swab
to swab the tongue and rub the swab over the surface of the plate.
TECHNIQUES

Prepare wet mounts of molds and yeasts using 2 loops of water and 1 loop of Gram’s iodine.
Mix a loopful of yeast into water mixture. Tease mold off of agar using an inoculating needle
then add to water mixture. Observe both at 40X. 100X may be used for yeasts and for mold
spores. Observe yeasts for reproduction in the form of budding. Some buds will fail to detach and
form pseudohyphae. Observe mold for sporangia and hyphae. Observe mold spores for several
minutes for the formation of new hyphae.
STUDENT TASKS
Per pair

Pour two Sabouraud agar plates.

Spot inoculate Sabouraud agar slant with Aspergillus niger and incubate at room temperature.

Inoculate one glucose and one sucrose fermentation tube with Saccharomyces cerevisiae and
incubate at body temp.

Once solidified, streak one plate with four quadrant technique using Saccharomyces cerevisiae and
incubate at room temperature. Streak plate two with a tongue swab and incubate at body
temperature.
Per Person
14


Prepare a wet mount of A. niger and observe on 10X for hyphae and sporangia. Go to 100X and
observe individual spores for germination and hyphae formation.
Prepare a wet mount of Baker’s yeast and observe on 100X for budding and pseudohyphae
formation.
REACTIONS

During fermentation, yeasts will convert organic molecules such as sugars to new organic products
such as alcohols, acids and CO2 with the production of a small amount of energy. In liquid
fermentation broth, acid production is seen when the red broth changes to a yellow color. Alcohol
or other alkaline products will produce a fuchsia color. Gas production is seen as the trapping of a
gas bubble in the Durham tube. When yeast colonies grow, they are larger than bacterial colonies,
they are white in color, raised and have the odor of fresh bread. Yeast cells divide by a process
called budding in which new cells bleb off the parent. Sometimes these buds do not detach and
form pseudohyphae. When molds grow on agar, they spread by producing and releasing spores
which land on new surfaces, and grow to become a new mold. A mold is composed of long branchlike strands called hyphae. One strand is called a hypha. Many hyphae bundled together are called
a mycelium that is visible to the naked eye and is the fuzzy appearance you see on bread molds.
Rhizoidal hyphae are those embedded in the agar like roots. Vegetative hyphae are those spread
out across the agar involved in gathering nutrients and oxygen. Aerial or reproductive hyphae are
those extending into the air, and are involved in reproduction (spore formation). The tips of the
reproductive hyphae may have sporangia, sacs filled with spores, or conidia, chains of spores
extending like fingers. Fungi may reproduce sexually or asexually. Pathogenic yeasts are
dimorphic, that is they grow as a yeast under certain conditions and as a mold under other
environmental conditions. Candida albicans is a common pathogenic yeast in humans. It is
responsible for urogenital infections (Candidiasis) as well as oral infections (Thrush). These
yeasts are part of the normal flora of some individuals and are opportunistically pathogenic in that
when the person is debilitated, taking antibiotics or during hormone and pH fluctuations, the yeasts
can overpopulate and cause an infection. In some cases the yeasts are part of the normal flora of a
sexual partner and can be transmitted between partners. If one person is more susceptible than the
other, the yeast infection will crop up repeatedly.
RESULTS

Observe budding, pseudohyphae, hyphae, spores, sporangia and germination of mold spores by
microscopy. Observe formation of yeast colonies on agar. Observe spread of molds by spread of
spores on agar. Observe growth of yeast colonies from some student’s tongues. Observe yeast
fermentation using liquid fermentation broth.
____________________________________________________________________________________
Negative and Simple Staining
BACKGROUND

The most important feature of negative staining is that no heat is applied, so there is no possibility of cell distortion as a
result of overheating; true morphology is seen. With this technique, acidic stains or dyes having negatively charged dye
15
molecules are repelled from the negatively charged surface of a bacterium, so the bacteria appear clear in a dark
background. Examples of stains or dyes used for negative stains include nigrosin, eosin and India ink.

A negative stain is prepared by adding a drop of nigrosin to one end of a clean slide. A loopful of bacteria is then mixed
with the drop of dye. Another slide is then used to smear the nigrosin/bacteria mixture across the surface of the slide.
The smear is allowed to air dry. Once dry, the slide is observed under oil immersion. In a negative stain, the bacteria
are seen as bright areas in a dark background.

All the staining techniques performed in this lab, with the exception of negative and capsule staining, involves the
preparation of a bacterial smear and heat fixing. In preparing a smear, the tendency of all beginning students is to add
too much inoculum. The general rule for preparing smears is that if the culture you are using is liquid, you do not need
to add water to the slide. If the culture is growing on agar, you must add water to the slide first. Remember to add only
a loopful of water to a slide followed by addition of a few organisms. Mix the organisms into the water using the
inoculating loop so an even emulsion is formed if possible. You may have problems with clumping of some organisms
but do your best. Do not get any clumps of agar on the slides. When completed, your bacterial smear should be the
consistency of very diluted milk. A bacterial smear need only be the diameter of a pencil eraser and in fact many
smears can be placed on a single slide.

For every staining procedure involving a smear, the bacterial smear must be completely dry prior to heat fixing. It is
not acceptable to speed up drying by blowing on the slide. It is not acceptable to speed drying by using the
Bunsen burner. It is not acceptable to speed drying by waving the slide around in the air. Even though we do
not work with overt pathogens, it is just not a healthy or wise practice to risk the generation of aerosols containing living
bacteria. The slide warmers located in the lab are designed to be used to speed up the drying of the slides. However,
most professors at SAC use the slide warmers for heat fixing instead of using Bunsen burners. For this reason, the slide
warmers are usually set at a very high temperature setting. I encourage beginning microbiology students to allow slides
to air dry rather than using the slide warmers. Keep in mind that if you use small loops of water to generate your
smears, they usually dry in less than one minute, and there is no need for a slide warmer anyway.

Heat fixing is a critical and necessary step for a good staining procedure. Heating fixing attaches the bacteria to the
glass so the slide is permanent and the cells cannot be washed off easily. In addition, heating kills cells and inactivates
enzymes within the bacterial cells which can cause cells to rupture or degrade. Too much heat fixing also causes
distortion and breakage of cells. Once cells are heat fixed, they may be stored prior to staining.

Once a bacterial smear is heat fixed, it is ready for staining. A variety of stains can be used in a simple stain as long as
they are basic or positively charge. Remember bacterial cells have negatively charged surfaces, so positively charged
dye molecules will be attracted to them. Positively charged stains include crystal violet, safranin, and methylene blue.
Carbolfuchsin is also a basic stain; however, it also contains phenol (carbolic acid) which makes it more suitable for
specialized stains such as the acid-fast technique. The time you expose a smear to a basic stain may differ with the stain.
Once the staining procedure is finished, residual dye is washed away, and the slide is blotted dry. The cells retain the
stain, and the background should be clear. No cover slip is required for these slides, and immersion oil is added directly
to the slide for observation. Stained slides may be stored and observed later.
PURPOSE

A negative stain is a technique in which a dye, which repels from bacterial cell walls, is used
as a background, and the bacteria that will not be stained will appear as bright shapes in the
darker background. A simple stain uses a dye that is attracted to bacterial cell walls to color
bacteria so that they appear as colored shapes in a clear background. These simple
techniques work the same for all bacteria regardless of their cell wall composition. The
negative stain is powerful because it does not utilize any heat fixing and therefore there can be
no cell distortion. The negative stain shows true cell morphology.
16
CULTURES
Bacillus subtilis, Micrococcus luteus, Corynebacterium pseudodiphtheriticum

B. subtilis is a large bacillus which produces endospores. Endospores are resistant, dormant
structures which are produced by members of the Bacillus and Clostridium genera as a means of
surviving harsh environments. Endospores are resistant to simple staining procedures. M. luteus is
a coccus which occurs in tetrads. C. pseudodiphtheriticum is a nonsporing bacillus which forms
club or cigar shapes.

Begin your “bug collection” today. Each table will save the cultures issued. These slants will
be labeled with each person’s last name, the table number and lab section and placed in a can
or rack at 25oC. You will use your bug collection when needed to set up experiments for the
rest of the semester and add new cultures to the collection as you go. Be careful to maintain
pure cultures. If the culture is contaminated, reisolate the organism on a TSA streak plate
and reestablish a working slant. If you run out of a culture, make a new slant from cells left
on the old one.
MEDIA
Will receive TSA slant cultures of each organism.
CULTURE CONDITIONS

Cultures were grown at 37oC with loose caps.
REACTIONS
Use aseptic techniques to transfer cultures to clean slides.
Simple Stain:
1. Generate smear (mix loopful of water with bacterial sample)
2. Air dry!
3. Heat fix! The purpose of heat fixing is to permanently attach bacterial cells to the
glass slide and allows storage of preserved cultures for long periods of time. Also the
heat inactivates enzymes within the bacterial cells which, if active, can degrade the
cell wall over time and destroy the sample.
4. Add methylene blue for 1 minute. Methylene blue is a basic dye (positively
charged) which is attracted to the negatively charged bacterial cell wall.
5. Wash with dH2O (distilled water).
6. Blot slide dry with paper towels. Do not rub.
7. Wash back of slide with alcohol. Observe the slide using immersion oil. No
cover slip is used. Oil is added directly on top of smear.
Negative Stain:
1. Add 1 drop of nigrosin to one end of a clean slide.
2. Mix in bacteria.
3. Use second slide at a 45o angle to smear dye and bacteria across slide.
4. Air dry slide completely. (Will look like patent leather with cracks)
17
5. Observe dried slide using immersion oil. Put oil directly on top of dried dye.
RESULTS

Using the simple stain, B. subtilis will be blue bacilli with endospores which will resist staining.
Endospores will look like clear grains of rice. M. luteus will be blue cocci arranged in tetrads,
diplococci, staphylococci and streptococci. C. pseudodiphtheriticum will be blue bacilli. Look for
unique shape of this organism. This organism is pleomorphic, which means it may have more than
one shape. Look for “P” or “9" shapes as well as cigar or club shapes. Using the negative stain, you
will see the true morphology of microbes. B. subtilis and C. pseudodiphtheriticum will appear as
bright bacilli in a purple background. M. luteus will be clusters of bright cocci in a dark purple
background.
____________________________________________________________________________________
Gram Staining
BACKGROUND

Know the Gram staining steps in detail including why each step is done and what color Gram positive and Gram
negative cells are at each step of the procedure.

It costs $100 to take apart and clean each objective which is contaminated with immersion oil. In addition many
contaminated objectives cannot be salvaged and must be replaced. Always check to be sure you are positioning
the correct objective (100X only) when using oil. The bulk of your lab fees go to the upkeep of the microscopes.

When preparing for the Gram stains, make sure the slides are clean. This will become very important in some of the
latter staining techniques also. Be sure to label your slides using a black sharpie or tape.

Remember, a small loopful of water and a tiny dot of organisms are sufficient for making a bacterial smear. It is not
necessary to cover the entire surface of a slide with a smear. But, of course the bigger the drop of water or liquid
culture, the more you will have to spread out the liquid to achieve fast drying.

Please note the amount and appearance of growth that you have on your table's agar slants or liquid cultures. These
tubes contain enough bacteria to perform hundreds of Gram stains. There is no reason for a tube to be scraped dry
for a few stains. Less is often better in microbiology.

When tubes are discarded, the labels should be removed from your table's cultures at the end of each lab, and the
tubes should be placed on the cart at the back of the room. Those tubes will be picked up later by the laboratory
technicians and autoclaved. Autoclaving is a method of sterilization using steam under pressure. If you have written on
a glass tube with a sharpie, remove the writing with alcohol before discarding the tube. If you label your tubes with
tape, remove the tape before discarding the culture. Once autoclaved, sharpie marks and tape become permanent on the
glass.

It is best to mark slides on the bottom. Otherwise the marker will easily erase with alcohol during your washing steps.

If you heat fix smears which are not dry, you are essentially boiling the bacteria in water. The cell walls will be broken,
and the cell shapes will be distorted. This can also occur if you apply too much heat during heat fixing.
18

Gram positive cultures which are over 24 hours old often will begin losing cell wall integrity and will destain easily.
This may result in a Gram negative appearance. You should record the correct Gram reactions of all the bacteria you
encounter in the lab as a reference. You can easily determine the Gram reaction of an organism by consulting Bergey's
manual or the text.

Too much destaining will also result in the decolorization of Gram positive cells and can lead to mistaken identity. For
future reference, members of the genus Micrococcus are Gram positive cocci which destain very easily with alcohol. In
fact, it is almost impossible to see completely purple Gram stains of this organism. Usually you will see what appears to
be both pink and purple cocci within the same cluster. When this happens with any organism, it is a clue that too much
decolorization has occurred. Do not mistake this phenomenon as representing a mixed culture. In addition the only
Gram negative coccus on the list of organisms you might encounter this term is an oxygen sensitive organism and
probably would not grow under normal conditions.

When using the microscope, don't forget to begin with lower power objectives first. You will not be able to distinguish
bacteria until you go to oil, but please sacrifice a few seconds of your time to work your way up through the objectives.
If you put oil onto a slide before placing it on the microscope and move the stage with the coarse adjustment until
the oil objective touches the oil, you are using incorrect procedure. You are risking breakage of the oil objective
lens and the slide.

Remember to close all the Gram staining reagent bottles before placing them back on the shelf. This prevents
evaporation especially of the alcohol and formation of crystals in the stains.

Now is the time to begin making observations of any bacterial cultures which you use in the lab. Remember from the
orientation session that many of the bacterial strains in the lab have distinguishing features such as color and texture
which can be seen with the naked eye. Many of these cultures will be potential unknowns.
PURPOSE

To learn this important staining technique and the difference between Gram positive and
Gram negative cells; the Gram stain is called a differential stain.
CULTURES

Staphylococcus aureus, Escherichia coli, Bacillus subtilis and mixture of all three
MEDIA
Cultures will be provided on TSA slants.
CULTURE CONDITIONS
REACTIONS
Cultures were cultivated at 37oC with loose caps.
1. Generate smear.
2. Air dry!
3. Heat fix!
4. Flood smear with crystal violet (a basic dye) and let stand for 30 seconds.
5. Wash with dH2O.
6. Flood smear with Gram’s iodine and let stand 1 minute. (A mordant which
intensifies the Gram stain by forming complexes with crystal violet. These
19
complexes will be trapped in the thick Gram positive cell walls but can still wash out
of Gram negative cell walls.)
7. Wash with dH2O.
8. Decolorize using acetone alcohol for ~2 sec. Be very careful during this step.
Wash only until color leaches out of smear.
9. Wash with dH2O immediately after decolorization to stop the destaining process.
10. Flood smear with safranin (a basic dye) and let stand for 2 minutes.
11. Wash with dH2O.
12. Blot slide dry with paper towels
13. Wash back of slide with alcohol. Observe using oil immersion.
RESULTS

E. coli is a Gram negative bacillus. S. aureus is a Gram positive staphylococcus. Gram positive
organisms are purple. Gram negative organisms are pink.
____________________________________________________________________________________
Acid fast and Endospore Staining
BACKGROUND

Acid fast staining is required to visualize organisms such as the Mycobacterium species. Because of the impenetrable,
waxy lipid cell wall of the mycobacteria, simple stains and Gram stains cannot be used. Remember the waxy lipid is
called mycolic acid, and the organism is named after this lipid. When the mycobacteria are heated in the presence of
carbolfuchsin (which contains the dye fuchsin and carbolic acid), the heat and the acid promote the penetration of the
acid-fast cell wall by the dye. When you wash with acid-alcohol afterward, the carbolfuchsin is soluble in lipid and not
in the alcohol so no amount of washing will decolorize the acid-fast organisms. Non-acid fast organisms such as E. coli
or any of the other microbes we used for a Gram stain would easily decolorize in acid alcohol. Therefore, when the
slide is counterstained with methylene blue, non-acid fast organisms will take up the blue dye.

One major problem with acid-fast stains is that if there is the slightest bit of dirt on the slide, the waxy organisms will
begin releasing from the slide during staining and washing. Therefore a clean slide, and proper heat fixing of the
bacterial smears are very important in this technique.

If you perform Gram stains on your unknown organism on the first day you receive it, please remember that an
attempted Gram stain of Mycobacterium smegmatis or Mycobacterium phlei will often give results which are
ambiguous and can be mistaken as other truly Gram sensitive organisms. Gram stains of mycobacteria may look
like Gram positive rods, Gram negative rods, Gram positive cocci, Gram negative cocci and often Gram negative
spirilla. Therefore, do not jump to a conclusion until you have observed the growth of your organism(s) on the agar
plates which you will generate. Remember the mycobacteria have a very distinctive, dry, waxy appearance on agar.

Remember endospores are highly resistant, dormant forms of bacteria which can survive extremely long periods of
time in harsh conditions. Members of the genus, Bacillus, are all Gram positive endospore producing cells. After
growth for a couple of days in a test tube, a Bacillus culture will exhaust its supply of nutrients and will actively produce
endospores in order to survive. Members of the genus, Clostridium, are anaerobic, Gram positive bacilli. In addition
to the depletion of nutrients, Clostridium species will respond to the presence of oxygen as a negative signal or hostile
environment and will begin to sporulate. The Clostridium used for the endospore stain will be grown in a candle jar.
The tubes are placed in a jar with a lighted candle, and the lid of the jar is screwed on tightly. When the supply of
oxygen is exhausted in the jar, the flame is extinguished. This is not an oxygen free environment, but the amount of
20
oxygen is reduced sufficiently for this anaerobe to grow. Therefore, the Clostridium will be grown in a sufficient
environment such that not many endospores will be formed when you observe them. However, by scanning the slide,
you should be able to find a few cells which contain endospores which have not broken free. In the Bacillus culture,
however, most of the spores will already be free and outside the remaining bacilli.

The boiling of the malachite green with the bacteria in an endospore stain facilitates the penetration of the green stain
into these resistant structures. Once the spores are stained, you can wash vigorously without worrying about removing
the stain. The bacilli are then counterstained with safranin. So you should see green endospores with pink bacilli.
PURPOSE

The Acid fast stain is a differential stain which allows students to differentiate between acid fast
and non-acid fast bacteria. Acid fast refers to the ability of certain cells to retain a lipid soluble dye
even when washed with acid. These bacteria include the Mycobacterium and Nocardia genera
which have a waxy lipid called mycolic acid as a component of their cell walls. Non-acid fast
bacteria lack this lipid and decolorize when washed with acid. The endospore stain is a special
stain used to identify bacteria capable of producing endospores. Endospores resist staining with
normal techniques and must be heated in the presence of dye in order to drive the dye into the
endospores.
CULTURES
Acid Fast Stain: Escherichia coli, Mycobacterium smegmatis or Mycobacterium phlei
Endospore Stain:
Bacillus subtilis and Clostridium butyricum

E. coli is a non-acid fast, Gram negative bacillus. Mycobacterium species are acid fast bacilli.
Bacillus and Clostridium species are endospore producers.
STUDENT TASK

Each student will prepare two acid fast stains. One of E. coli and another of Mycobacterium.

Each student will prepare two endospore stains. One on each of the organisms listed.
MEDIA
Cultures will be issued on TSA agar slants.
CULTURE CONDITIONS

Cultures were grown at 37oC with loose caps with the exception of Clostridium, which is an
anaerobe. The anaerobe was grown in an anaerobic chamber. The sporulating organisms
were grown at least 72 hours to insure they are running low on nutrients and have begun to
sporulate. The anaerobe was transferred out of the anaerobic chamber a couple of days in
advance to promote sporulation in the presence of oxygen.
REACTIONS
Acid Fast Stain:
1. Generate smear on extra clean slide.
2. Air dry!
3. Heat fix!
21
4. Place over steaming beaker of tap water. Add filter paper cut to size of smear.
Saturate smear with Carbolfuchsin (a basic, lipid soluble dye) and heat for 5
minutes, keeping paper moist so it doesn’t stick to smear.
5. Remove paper with forceps and wash with dH2O for 30 seconds.
6. Decolorize with acid-alcohol for 15 seconds.
7. Rinse with dH2O.
8. Flood smear with methylene blue for 2 minutes.
9. Rinse with dH2O.
10. Blot dry. Wash back of slide with an alcohol soaked Kimwipe.
11. Observe using immersion oil.
Endospore Stain:
1. Generate smear on extra clean slide.
2. Air dry!
3. Heat fix!
4. Place over steaming beaker of tap water. Add filter paper cut to size of smear.
Saturate smear with malachite green and heat for 5 minutes, keeping paper moist.
5. Remove paper with forceps and rinse with dH2O for 30 seconds.
6. Flood smear with safranin for 2-3 minutes.
7. Rinse with dH2O.
8. Blot dry. Wash back of slide with an alcohol soaked Kimwipe.
9. Observe using immersion oil.
RESULTS

E. coli is non-acid fast and will be blue following the acid fast procedure. Mycobacterium will be
fuchsia (pink) and is acid-fast. Endospores will take up the malachite green and will be pale teal
green in color. The endospore producing parent bacilli will be stained by the safranin and will be
pink.
____________________________________________________________________________________
Capsule Staining
BACKGROUND

CONTRARY TO WHAT LAB MANUALS MAY SAY, A NEGATIVE STAIN IS NOT A VIABLE
TECHNIQUE FOR IDENTIFYING CAPSULES.

Remember with capsule staining that the capsules are impenetrable, nonionic, carbohydrate structures, but they are not
indestructible. The smears should be prepared gently using a healthy loopful of culture.

The slides need to be very clean for this technique because residues left on the slide surface will cause the dried Congo
red/bacterium mixture to lift off the glass during washing. The solution labeled slide cleaner is an industrial detergent
called Alconox dissolved in water. If you run out, you can make more by adding approximately a teaspoon of white
powder detergent, always kept by the sink, to the bottle and add water.
22

The Congo red is a pH indicator. At neutral pH, Congo red is red. At acidic pH, Congo red is blue. You should
observe the color change from red to blue when you wash with acid alcohol. Therefore, the background color of a
capsule stain is blue.

Because no heat is applied during this technique and the background is stained, the capsule stain is most like a negative
stain.

Also because no heat is applied, the bacteria may still be alive following this procedure. Be sure to add disinfectant to
the slide when cleaning it.

After the capsule stain is treated with acid fuchsin, the bacterial cells themselves should appear pink because they have
taken up the fuchsin. The capsules will appear as clear halos, and the background will be blue.

Remember, you should expect a bacterium that produces a capsule to appear as sticky, slimy colonies on agar.
PURPOSE

To demonstrate the presence of capsules surrounding some species of bacteria.
CULTURES
Enterobacter aerogenes or Bacillus subtilis
SUPPLIES

Congo red, acid alcohol, acid fuchsin, clean slides, immersion oil, paper towels, lens paper,
ethyl alcohol
MEDIA
Cultures are provided as TSA slants.
CULTURE CONDITIONS
Cultures were grown at 37oC with loose caps.
REACTIONS

Congo red dye is a pH indicator that is red at neutral pH and blue at acid pH. This dye will serve
to create the blue background in which the bacilli will be embedded. Acid alcohol is required to
penetrate the capsule that will resist staining and is normally quite impenetrable. Acid fuchsin is
the secondary stain that will pass through the capsule and stain the bacilli.
Capsule Stain:
1. Add 1 drop Congo red to a clean slide.
2. Mix in bacteria and spread drop to the size of a quarter.
3. Allow mix to air dry completely.
4. Wash smear with acid alcohol. Note color change.
5. Rinse with dH2O.
6. Flood smear with acid fuchsin for 5 minutes.
7. Rinse with dH2O.
8. Observe using immersion oil.
RESULTS
23

For either B. subtilis or E. aerogenes, the bacilli will be stained pink by the acid fuchsin. The
capsules will appear as clear or white halos surrounding each bacillus like a line drawn around the
cell. The background will be blue. Move to an area where the background is a smooth light blue
color and the bacilli can be easily visualized. This technique is similar to the negative stain. E.
aerogenes bacilli are very short and can be mistaken for slightly oval cocci.
Hand Washing and Environmental Sampling
BACKGROUND

Direct contact of a disinfectant with bacterial cells is necessary in order for a chemical to penetrate and kill the cells.
Organic materials such as those encountered in wound cleansing can interfere with the effectiveness of a chemical.
Disinfectants are chemicals used to kill microbes on inanimate objects. Antiseptics are used to kill microbes on living
tissues.

The effectiveness of over the counter soaps will differ. Bar soaps are probably the least effective because these soaps
become contaminated with bacteria. Deodorant soaps do contain additional chemicals that inhibit Gram positives,
which are usually the microbes associated with production of body odor. Solutions such as Betadine contain iodine plus
a detergent, and are very effective because of the physical removal of microbes by the soap and killing by the
disinfectant (iodine).

Most normal flora are not harmful, however they must be removed during a surgical scrub because they may be harmful
if inoculated into the body where they become opportunistic pathogens. Surgery patients are often more susceptible to
these infections. Hospital scrubs often include both a detergent and an antiseptic to remove and kill as many bacteria as
possible.

Hand washing fails to remove all bacteria because microbes may remain in the pores of the skin. In addition, new
microbes are acquired from anything touched such as containers, sink handles, towels, soap itself and other fomites. For
this reason liquid soaps which can be dispensed with a foot pump are often used in hospitals.

Paper toweling in rest rooms minimize post-washing contamination since residual microbes from previous users can be
left on continuous-feed cloth towels. Bacteria such as Pseudomonas can also be transmitted person to person if towels,
washcloths, loofa sponges and back brushes are shared in family showers.
PURPOSE

To introduce students to their normal flora (organisms which normally live in or on them). All of
these organisms cannot be removed even with vigorous hand washing. Normal soaps do not kill
microbes. The purpose of normal soaps is to physically solubilize oils on the skin or surfaces and
cause organisms to be washed away with rinsing. Only soaps with a disinfectant or antiseptic have
a true antibacterial (meaning killing) effect. Transient flora or those organisms temporarily
contaminating hands and surfaces can be removed or decreased to a safe level by washing.
CULTURES
Normal flora or transient flora
MEDIA
Nutrient agar deeps (for Petri plates)
SUPPLIES
Sterile swabs, sterile Petri plates, soaps, scrub brushes
24
CULTURE CONDITIONS

After touching the plates pre- and post-washing, the plates will be inverted and placed in the 37oC
incubator. Environmental samples will be grown at room temperature (25oC). If any moisture is
on the plate surface, allow it to absorb before inverting the plates.
REACTIONS

Microbes that have been transferred to the medium will be allowed to grow. Nosocomial
infections are hospital acquired infections which are often the result of cross contamination of areas
which are supposed to be sterile with normal or transient flora. Even though the normal flora would
not normally make you sick, they are opportunistic pathogens, which can make you sick if given
the opportunity to do so. Transient organisms are those only present for a short time and are easily
removed by washing. Disinfectants are chemicals used to decrease the number of microbes on
inanimate surfaces. Antiseptics are chemicals used to decrease the number of microbes on skin and
mucous membranes. Notice the numbers are decreased. Microbes are not eliminated completely.
These chemicals do not sterilize under normal use. Divide 2 nutrient agar plates into 4 quadrants
each. Touch 1 quadrant with two unwashed fingers. Wash fingers with soap and scrub brush for 15
sec, 30 sec, 1 min, 2 min, 3 min, 4 min and 5 minutes (total washing time). Do not dry fingers and
touch remaining quadrants labeled to match washing times. Incubate plates at 37oC inverted. For
environmental samples, dip a sterile swab in sterile water and then swab a surface or object. Take
the plate with you if you leave the room to swab. Streak the swab over the surface of the plate in
several directions. Discard the swab in the biohazard trash. Incubate the plates at 25oC if the
sample came from an inanimate object. Incubate at 37oC if the sample is a body sample.
RESULTS

Only microbes capable of growth on nutrient agar will be seen. You might miss seeing many
microbes that need specialized growth conditions. Don’t be surprised to see an increase in the
amount of microbial growth on the quadrants of the plates that were touched after the fingers were
washed longer. This is due to the fact that longer washes solubilize more microbes so that they will
easily transfer to the agar. You will never remove all microbes. In the real world, hand washing is
most important in order to prevent the spread of transient flora such as cold viruses, which you may
have picked up from objects.
____________________________________________________________________________________
Carbohydrate Fermentation (Liquid Fermentation Broth)
BACKGROUND

The process of fermentation is a group of biochemical reactions in which an organic molecule such as a carbohydrate
(glucose, sucrose or lactose) serves as an electron donor, and the final electron acceptor is the new organic product.
Some ATP is produced from fermentation reactions.
25

During fermentation of a carbohydrate, bacteria may produce organic acids as byproducts. Just because a given
organism can ferment one sugar does not mean it can use all of them. In the case of E. coli, glucose and lactose should
be used with the production of an acid as a fermentation product resulting in the yellow color in the fermentation tubes
and in the positive methyl red test for the presence of acid. E. aerogenes ferments all three sugars with the production
of acid but no gas. The Durham tube inside the fermentation tube is present to trap any gas produced. Carbohydrate
fermentation that can be tested for in this lab include: glucose, sucrose, lactose, mannitol, sorbitol, dulcitol

Possible outcomes of fermentations tests can be: no fermentation, acid production without CO 2, acid and CO2
production, and CO2 production with no acid (Ethanol production).
PURPOSE

To identify bacterial cultures capable of fermenting various sugars with the production of acid or
alkaline products or gasses.
CULTURES
Escherichia coli, Alcaligenes faecalis, Staphylococcus aureus
MEDIA

3 each of Sucrose, Lactose and Glucose fermentation broths with Durham tubes
CULTURE
CONDITIONS
Inoculate broth with loop and incubate at 37oC with loose caps.
REACTIONS

Fermentation broth contains phenol red as a pH indicator. At neutral pH, the indicator is
orange/red. At alkaline pH, the medium becomes fuchsia (hot pink). At acid pH, the medium
becomes yellow. If gas is produced, it will be trapped in the Durham tube.
RESULTS

Yellow = acid; fuchsia = alkaline (perhaps an alcohol); gas in Durham tube could be CO2, H2S,
methane, nitrogen, Etc. See unknown charts for expected results for tested organisms.
Triple Sugar Iron (TSI) Agar Test
BACKGROUND
The TSI test is used in the identification of enteric bacteria, which metabolize the triple sugars and release hydrogen sulfide.
TSI agar contains lactose, sucrose and glucose. The pH indicator, phenol red, is used to monitor acid production from
carbohydrate fermentation. TSI slants are inoculated by streaking the slant, then stabbing the agar with an inoculating needle.
After incubation, several possible reactions can be observed. The agar is orange prior to inoculation. Glucose fermentation is seen
as a yellow butt and a red slant due to the fact that glucose is in low concentration and acid accumulates at the bottom. At the
surface, in the presence of oxygen, the slant remains alkaline. A yellow slant and butt indicates fermentation of the lactose and/or
sucrose, which are in high concentration and results in acidification of the entire medium. Gas formation is seen as splitting of the
agar. Hydrogen sulfide production is seen as blackening of the agar. This test should be read 18-24 hours post-inoculation
because acid products can be further metabolized resulting in alkalinization of the medium.
26
PURPOSE

To identify and differentiate among the enterics based on the sugar fermentation pattern and
the production of hydrogen sulfide gas.
CULTURES

Alcaligenes faecalis, Escherichia coli, Proteus vulgaris, Proteus mirabilis, Pseudomonas
aeruginosa

All Proteus species should produce H2S. Pseudomonas should synthesize alkaline products and
turn the slant fuchsia.
MEDIA
5 Triple Sugar Iron Agar (TSI) Slants
CULTURE CONDITIONS

Inoculate slants using the streak/stab method. Grow all cultures at 37oC with loose caps.
REACTIONS

TSI agar contains sucrose and lactose in high percentage. If organisms ferment these sugars
producing acid products, the slant and butt will turn yellow. If the organisms ferment glucose,
which is in low concentration, the butt will turn yellow; the slant will probably remain unchanged
with the original orange red color. If hydrogen sulfide gas is produced, the agar will turn black.
Hydrogen sulfide gas (H2S) is produced when sulfate containing compounds, such as ferrous
ammonium sulfate, are used as a sulfur source. The breakdown of ferrous ammonium sulfate
results in the release of ferrous iron compounds, which will precipitate forming a black sediment.
RESULTS
Check the unknown charts for the expected reactions for these organisms.
____________________________________________________________________________________
IMViC Series
BACKGROUND

The IMViC tests are important in the differentiation and identification of enteric (intestinal) bacteria, which often
contribute to waterborne and food-borne diseases.

SIM agar contains peptones and beef extract as a source of protein. The indole test identifies bacteria that hydrolyze
tryptophan (an amino acid which is found in most proteins) to produce indole, pyruvic acid and ammonia. Indole is
not further used in bacterial metabolism and accumulates in the medium. Indole can be detected in medium following
growth of bacteria by adding Kovacs' reagent, which produces a bright red color at the top of the medium if indole
is present.

The methyl red test is used to identify bacteria that convert glucose to an acidic end product such as lactic, acetic or
formic acids. Some bacteria are called mixed acid fermenters because they produce a mixture of fermentation acids
such as acetic, lactic and formic acid. Butanediol fermenters form butanediol, acetoin and a few organic acids that do
not acidify the medium detectably. Methyl red is a pH indicator, which will color the medium red at acidic pH and
27
will be yellow in neutral pH. In the MR-VP broth that is used to grow the bacteria, the yellow color cannot be seen
and a negative methyl red test is indicated.

The Voges-Proskauer test identifies bacteria that do not acidify medium but produce 2,3-butanediol (a neutral
product). The VP reagents are added to the medium to detect the presence of acetoin (a precursor of 2,3-butanediol). If
acetoin is present, a cherry-red color develops at the top of the medium.

If acid is present and remains present, the VP test for neutral products is negative. After 48 hrs, however, some
organisms are capable of converting acid to the neutral product, acetoin, and a positive VP test occurs. Be sure to
check MRVP tests no later than 2 days after inoculation.

The citrate utilization test is used to identify bacteria that can use citrate as a sole carbon source. These cells must
produce citrate permease, which is used to transport citrate into the bacterium. The Simmon's citrate agar is a bright
green color. It contains sodium citrate, ammonium as a nitrogen source, and a pH indicator. When citrate is utilized,
CO2 is released which combines with the sodium to produce sodium carbonate (alkaline). As the pH rises, the pH
indicator changes from green to royal blue. The blue color represents a positive citrate utilization test. This test is
performed on an agar slant because oxygen is required. Remember to leave the test tube cap loose.
PURPOSE

To differentiate among the enteric organisms. Enterics are intestinal bacteria.
Composed of four biochemical tests.

Indole - used to identify bacteria capable of tryptophan hydrolysis.

Methyl red - used to identify mixed acid fermenters.

Voges-Proskauer - used to identify butanediol fermenters

Citrate - used to identify bacteria capable of transporting and using citrate as a sole carbon source.
CULTURES

Indole: Escherichia coli, Proteus vulgaris, Proteus mirabilis, Enterobacter aerogenes

MR: E. coli, E. aerogenes

VP: E. coli, Bacillus mycoides

Citrate: E. coli, Proteus mirabilis
MEDIA

Indole: 4 SIM agar deeps (SIM also used for H2S detection, Indole and Motility)

MR: 2 MRVP broths

VP: 2 MRVP broths

Citrate: 2 Simmon’s citrate slants
CULTURE CONDITIONS

Stab inoculate SIM agar deeps; incubate at 37oC with loose caps, 48 hrs.

Inoculate MRVP broths and incubate with loose caps at 37oC, 48 hrs.

Streak/stab inoculate citrate slants and incubate at 37oC with loose caps, check after 48 hrs, but
may require 4 or more days.
REACTIONS
28

Indole: Tryptophan----------> Indole--------> Waste
Pyruvate-----> Nutrition
Ammonia----> Nutrition

MR: Glucose ---------> lactic, formic, or acetic acid (mixed acids)
pH drops to 4

VP: Glucose------->Acetoin--------->2,3-butanediol + little acid
pH drops to 6

Citrate:
Sodium citrate------->Pyruvic acid +CO2 + sodium
Requires O2 and citrate permease enzyme.
CO2 + sodium --------> sodium carbonate (a base)
pH goes up.
RESULTS

Indole: Add 5 drops of Kovac’s reagent to top of agar. Red ring developing after a few seconds is a
positive indole test. No red ring is a negative test. See unknown charts for expected results.

MR: Add 5 drops methyl red to MRVP broth culture. Do not shake. If red ring persists, positive
MR test. If red dye dissipates, negative test. See unknown charts for expected results.

VP: Add 15 drops VP-A reagent; 5 drops VP-B reagent to MRVP broth culture and shake
vigorously. Let stand with cap off for 15 minutes. Red ring at top is positive VP test. Positive VP
test indicates the presence of acetoin (butanediol precursor). No color or a cloudy beige color at top
is negative VP test. See unknown charts for expected results.

Citrate: pH indicator bromothymol blue changes from green (neutral) to royal blue (base) as pH
goes up. See unknown charts for expected results.
____________________________________________________________________________________
Oxygen Requirements of Bacteria
BACKGROUND

The GasPak system is useful for culturing anaerobic bacteria on standard microbiological media because the GasPak
generates carbon dioxide and hydrogen. The hydrogen will combine with oxygen present in an anaerobic jar to
produce water. This system can reproducibly attain oxygen levels in the parts per million range if used correctly. This
is the best method for determining the oxygen requirements of unknown organisms.

A candle jar is useful for culturing organisms that prefer reduced oxygen levels and increased carbon dioxide levels.
The candle jar is not an oxygen free system. This is the best method of culturing a microaerophile.

An obligate anaerobe is a microbe that cannot tolerate oxygen and will be killed in its presence. This type of organism
is killed because it lacks the enzymes necessary to eliminate toxic oxygen products.
29

An obligate aerobe is a bacterium that requires atmospheric levels of oxygen for normal growth.

A facultative anaerobe is a microorganism that can grow with or without oxygen but usually grows faster (produces
more ATP) in its presence.

An aerotolerant anaerobe is a microbe that prefers anaerobic conditions but can tolerate exposure to low levels of
oxygen.

A microaerophile is an organism that requires reduced levels of oxygen.

A sodium thioglycollate broth tubes contains substances that chemically combine with oxygen making it unavailable.
Anaerobes will grow in this medium. Aerobes can also grow but only in the upper layers of this medium.
PURPOSE
To identify bacteria based on growth in oxygen at differing levels.
CULTURES
Pseudomonas aeruginosa or Micrococcus luteus, Clostridium, Escherichia coli
(Aerobes)
(Anaerobe) (Facultative anaerobe)
MEDIA
3 TSA plates (for anaerobic chamber)
CULTURE CONDITIONS

Lightly streak 4 quadrants on TSA agar for anaerobic chamber. Make sure you can’t see bacteria
on agar surface prior to incubation because a heavy streak can be mistaken for growth. Place in
anaerobic chamber at 37oC with plates inverted.
REACTIONS

In an anaerobic chamber, a GasPak of chemicals is activated using 10ml of dH2O (distilled
water). The GasPak releases CO2 and H2 gases. Most anaerobes also prefer elevated CO2 levels.
The chamber has a mesh capsule in the lid that contains a catalyst called palladium. In the presence
of the catalyst, the H2 gas released from the GasPak and any O2 in the chamber chemically react
forming H2O. This will be seen as condensation inside the chamber. In addition, an oxygen
indicator strip is enclosed in the chamber. In the presence of O2, the paper pad is blue. In the
absence of O2, the pad is white. Always check for condensation and the color of the oxygen
indicator pad prior to opening the chamber.
RESULTS

In an anaerobic chamber, only the obligate anaerobes, aerotolerant anaerobes and facultative
anaerobes will grow. Obligate aerobes and microaerophiles will not grow. An organism is a
facultative anaerobe if it grows in the presence of oxygen as well as in the anaerobic chamber. A
microaerophile can be identified by growth in a candle jar in which a plate is sealed in a jar with a
lighted candle. As oxygen is consumed by the flame, the O2 level drops and the CO2 level
increases. This is the perfect environment for microaerophiles.
30
Catalase Activity
BACKGROUND

Any bacteria that grow in the presence of oxygen must deal with the production of toxic oxygen products. Toxic oxygen
products include superoxide anions and hydrogen peroxide. These products are strong oxidizing agents and cause
destruction of cellular components including DNA. Most aerobic bacteria have superoxide dismutase (SOD) as well
as catalase or peroxidase. SOD eliminates superoxide anions by combining them with hydrogen cations to produce
oxygen and H2O2. H2O2 is destroyed by catalase to produce water and oxygen. Peroxidase destroys hydrogen
peroxide to release water only. We tested several bacteria for the production of the enzyme catalase. A catalase test
is performed by dripping hydrogen peroxide onto isolated bacterial colonies. If the cells bubble (indicating the release
of oxygen), the bacterium is positive for the enzyme catalase. The catalase cannot be performed on colonies that are
growing on blood agar because the endogenous catalase activity of animal cells will give a false positive. If you
have cells growing on blood, scrape the cells off onto a slide and then drip on the hydrogen peroxide.

Strict anaerobes do not produce catalase therefore they cannot deal with toxic oxygen products. Aerobes,
facultative anaerobes, aerotolerant anaerobes and microaerophiles may express catalase. The majority of all bacteria are
facultative anaerobes (can grow with or without oxygen). The catalase test can be useful in differentiation between
bacteria such as the catalase negative Enterococcus and the catalase positive Staphylococcus.
PURPOSE

To identify bacteria capable of dealing which metabolism in the presence of oxygen.
CULTURES
Staphylococcus aureus, Enterococcus faecalis, Micrococcus luteus
MEDIA
3 TSA slants
CULTURE
CONDITIONS
Streak slants and incubate at 37oC with the caps loose.
REACTIONS

Normal growth of cells in O2 results in the production of hydrogen peroxide (H2O2) or superoxide
anions (O2-). H2O2 and O2- are toxic oxygen products that must be eliminated for survival.
Superoxide is also called free radical. These products destroy organic molecules such as DNA and
proteins in cells. Many bacteria as well as your cells have enzymes for the elimination of toxic O2
products.

O2--------------------------> O2 + H2O2
Superoxide dismutase

H2O2 ----------------> H2O +O2
Catalase

H2O2 ----------------> H2O
Peroxidase
RESULTS
31

Add H2O2 to a TSA slant culture. If catalase enzyme is present in the bacterial cells, it will be
broken down into water and oxygen. The release of oxygen results in vigorous bubbling. Bubbling
is a positive catalase test. Be aware that all cells in your body including blood cells produce
catalase. A catalase test cannot be performed directly on blood agar. Cells must be scraped off the
plate onto a slide or the lid, then the hydrogen peroxide is added. Also be aware that hydrogen
peroxide has a 6 month shelf life. Always check the date before using. Aerobes, microaerophiles,
aerotolerant anaerobes and facultative anaerobes are potentially catalase positive. Obligate
anaerobes are never catalase positive which is why they cannot survive in the presence of oxygen.
See unknown charts for expected results on tested organisms.
____________________________________________________________________________________
Litmus Milk
BACKGROUND

Milk is a good differential medium because it contains many macromolecules that can be metabolized by bacteria,
including proteins (casein), and carbohydrates (lactose). Several different outcomes can be used to gauge what has
occurred in the tube. The main caveat to using litmus milk cultures is that failure of a bacterium to grow can be
misinterpreted as a negative test because the liquid is opaque and turbidity cannot be used as a measure of growth.
Milk protein is called casein. The process by which milk protein is broken down is called peptonization.

The litmus milk test can also be used to detect fermentation of the carbohydrate lactose. When lactose is fermented
using the enzyme beta-galactosidase, you will see a pink color develop as a result of lactic acid production. Litmus
milk is purple prior to inoculation. You should expect that bacteria which ferment lactose and produce acid will also
produce positive reactions in lactose fermentation tubes.

The litmus indicator may also be reduced (used as an electron acceptor) during growth of some organisms in litmus
milk. When this occurs, a white precipitate is seen at the bottom of the tube. The top of the milk will usually remain
purple because the litmus is oxidized in the presence of oxygen.

If acid is produced (pH 4) and reduction occurs, some bacteria produce an enzyme that causes the milk proteins to
coagulate (curd). Lactose fermentation may result in production of CO2 and/or H2 which causes fissures or cracks in
the curd (stormy fermentation). Proteolysis (peptonization) of the curd may be seen as the appearance of brownish,
straw-colored fluid.

Once proteins and carbohydrates have been used up as the preferred energy source, amino acids may be catabolized.
When amino acids are used in a litmus milk culture or if the lactose cannot be used, proteins and amino acids may be
broken down for energy resulting in peptonization (hydrolysis of the milk protein, casein). This breakdown is seen as
settling or clearing of the suspension. You might also see alkalinization if the amino acids from the casein are further
broken down resulting in formation of a purple or deep blue color.

Reading litmus milk cultures can be confusing so consult with instructor if you are not sure of a result. Ropiness
means that when you pull an inoculating loop through a milk culture, strings or ropes of material will follow the loop as
you pull it up along the side of the test tube wall.
PURPOSE

The purpose of this exercise is to identify bacteria capable of breaking down the various nutrients in
milk and the extent to which they can do so. Many of the microbes that catabolize milk are
important in the food industry for making cheeses and yogurt.
32
CULTURES
Escherichia coli, Proteus vulgaris, Bacillus subtilis, Streptococcus lactis
MEDIA

Litmus Milk contains lactose, casein, and litmus. Lactose is milk sugar; casein is milk protein and
gives milk its white color, litmus is a pH and reduction/oxidation indicator that is purple when
oxidized.
CULTURE CONDITIONS

Inoculate 4 litmus broth cultures and incubate at 37oC with caps loose. Litmus milk is kept in the
“clean” media refrigerator until needed.
REACTIONS
Lactose----------------------> Acid
Beta-galactosidase
Casein ----------------------------> Amino acids
Proteases or peptidases
Casein ------------------> Curd
Rennin + acid
RESULTS
The following results may be seen alone or in combination

Lactose fermentation producing acid ---> Pink

Lactose fermentation producing alkaline products ---> Purple

Casein hydrolysis ---> Peptonization (straw colored, clear fluid)

Coagulation ---> curding of milk protein

Gas production ---> cracks in curd (stormy fermentation)

Reduction of litmus Redox indicator ---> off white color beginning at bottom

Ropiness ---> mucoid, stringy trails follow loop when passed through milk
See unknown charts for expected results on tested organisms.
____________________________________________________________________________________
H2S and Motility
BACKGROUND

Cysteine is a sulfur-containing amino acid found in some proteins. Following protein hydrolysis, cysteine may be
broken down by cysteine desulfurase. The sulfur is removed from the amino acid and joined to hydrogen to form
hydrogen sulfide gas (H2S). In addition, inorganic sulfur containing compounds may be reduced during anaerobic
respiration to produce hydrogen sulfide. In the hydrogen sulfide production test, SIM agar contains peptones that are
partially digested proteins some of which will contain cysteine. The ferrous ammonium sulfate is an indicator which
combines with hydrogen sulfide forming an insoluble, black ferrous sulfide precipitate. This black coloration will
33
appear along the stab line of SIM agar. If the organism is motile, the entire tube may turn black. Any semisolid agar
medium may be used as a matrix for determining motility. Remember that many motile organisms will produce a
growth pattern resembling an inverted Christmas tree. In addition to hydrogen sulfide production and motility, SIM
agar can also be used for the indole test (See the IMViC series).
PURPOSE

To demonstrate that some media have dual purposes. However, production of a product like H2S
often precludes your ability to visualize motility.
CULTURES
Proteus vulgaris, Escherichia coli
MEDIA
2 SIM agar deeps and 2 Motility agar deeps
CULTURE
CONDITIONS
Stab inoculate deeps and incubate at 37oC with caps loose.
REACTIONS

SIM agar contains ferrous ammonium sulfate. Some bacteria catabolize sulfur containing amino
acids and other compounds resulting in release of H2S gas and the liberation of iron. Iron is
insoluble in the presence of oxygen and quickly acquires a black appearance. H2S production
within an agar medium can make observing motility difficult. Motility agar is a clear, semisolid
medium, which allows visualization of motility. Motility is seen as cloudiness of the agar moving
outward in all directions away from the stab line. Cocci are never motile. Do not mistake motility
for a shaky inoculation. If your hand shakes while stabbing the agar, a slice will be made in the agar
particularly wider near the top. This will result in a wide growth pattern at the top that tapers off
toward the bottom. When you turn the tube it will be obvious because you only see growth in one
plane. Do not mistake motility for faster growth at the top for aerobes and facultative anaerobes.
The inverted Christmas tree appearance has to do with oxygen not motility. Even a nonmotile
organism can form this pattern. If you cannot confirm motility using this method, a hanging drop
slide may be helpful in some cases.
RESULTS

P. vulgaris and E. coli are both motile organisms. Both have peritrichous flagella. Proteus species
also produce H2S.
Lipid Hydrolysis
BACKGROUND
In bacteria, lipids are high energy molecules and play a vital role in biosynthesis of membrane components. Many fatty foods can
be spoiled by bacteria that produce lipases. This spoilage is termed rancidity. Pathogenic bacteria that produce lipases can attack
host membranes and break them down resulting in increased pathogenicity and spread of microbes. Some bacteria breakdown
triglycerides and phospholipids to release this energy. Triglycerides contain glycerol and 3 fatty acid molecules. A phospholipid
is a complex lipid containing glycerol, 2 fatty acids and a phosphate group. Triglycerides are broken down by lipases to produce
glycerol and free fatty acids that can be further catabolized during glycolysis, beta-oxidation, the Krebs cycle and other metabolic
pathways. When bacteria are grown on agar containing lipids, the release of fatty acids can be detected as acidification of the
34
medium. Spirit blue agar contains a lipase reagent that gives the plates a bright blue color. When the lipid is broken down, a
clearing area will be seen surrounding lipolytic bacterial colonies.
PURPOSE
To identify bacteria capable of breaking down fats for energy.
CULTURES
Proteus mirabilis, Staphylococcus epidermidis
MEDIA
1 Spirit blue agar (bright blue in color, stored in fridge prior to use)
CULTURE
CONDITIONS
Spot inoculate plate and incubate at 37oC in the inverted position.
REACTIONS
Triglycerides (fats) -------------------> Glycerol + 3 Fatty acids
Lipase
(Krebs or)
(Beta-oxidation)
(Glycolysis)
RESULTS

Following incubation, a clearing zone, lighter blue zone or yellowing zone is a positive test for fat
or lipid hydrolysis. Do not incubate plates too long (more than two days at 37oC) because they will
dry out. Thin, dried out plates are often mistaken for a positive result. Always check plates prior to
inoculation for contaminants. S. epidermidis is negative; P. mirabilis is positive.
____________________________________________________________________________________
DNase Activity
BACKGROUND
DNases (deoxyribonucleases) are enzymes that degrade DNA. DNase test agar contains DNA. When an organism breaks
down the DNA, the small nucleotides and other breakdown products will not precipitate with hydrochloric acid. Large, intact
DNA molecules will precipitate when HCl is added to the plate and will cause the agar to appear cloudy. A DNase positive culture
will form a clear area around the bacterial growth. Most pathogenic staphylococci produce DNase.
PURPOSE

To identify pathogenic Staphylococcus species. These organisms degrade foreign DNA to increase
their virulence when in an animal host. By degraded the host DNA, the host is debilitated and less
able to react. In addition, the DNA nucleotides can be used by the bacteria for energy and
production of new nucleic acids.
CULTURES
Staphylococcus aureus, Staphylococcus epidermidis
MEDIA
1 DNase Test Agar divided in half
CULTURE CONDITIONS
35

Spot inoculate Petri plate and incubate inverted at 37oC. Media is kept in the refrigerator prior to
use.
REACTIONS
Deoxyribonuclease
DNA -------------------------> Nucleotides
Intact DNA precipitates
Soluble nucleotides do not precipitate
(Cloudy Agar)
(Clear agar)

DNA is very large when intact. Intact DNA fragments will precipitate when hydrochloric acid
(HCl) is added to the agar. The precipitate DNA will give the agar a cloudy appearance. When
DNA is digested with the DNase enzyme, soluble nucleotides are released which do not precipitate
when HCl is added. S. epidermidis is negative; S. aureus is positive.
RESULTS

Add concentrated hydrochloric acid to the plate following growth. A clearing zone indicates
breakdown of DNA and is a positive test. Flood the entire surface of the plate with acid. Be
careful, it will burn the skin and eyes. Try not to directly inhale the acid fumes. Leave the lid on
the plate for a few minutes while the fumes dissipate. Use a black background to more easily
visualize clear zones. Discard the plate as soon as possible in the biohazard waste.
Starch Hydrolysis
BACKGROUND

A hydrolase is an enzyme that catalyzes the splitting of organic molecules by water. Starch is composed of amylose (a
glucose polymer) and amylopectin. These subunits may be broken down by alpha-amylases to yield dextrins, glucose
and maltose. These simple sugars can then be used for energy. A starch hydrolysis test is performed by dripping
Gram's iodine onto colonies grown on starch agar. When iodine contacts starch, it produces a dark purplish-brown
color. If a clear area appears around a colony, the starch has been broken down by amylase. Starch hydrolysis is a clear
cut yes or no answer. If you see starch hydrolysis by one of the organisms besides B. subtilis, it was most likely the
result of the spreading of an excreted exoenzyme that diffused throughout the agar giving one of the other organisms a
false positive. Starch agar also contains beef extract (protein and lipid source), so it is possible that a bacterium will
grow heavily on this agar without producing amylases.
PURPOSE
To identify bacteria capable of breaking down starch.
CULTURES
Bacillus subtilis, Escherichia coli, Proteus vulgaris
MEDIA
3 Starch agar plates (plates may have to be poured by student)
CULTURE
Spot inoculate starch agar plate and incubate inverted at 37oC.
36
CONDITIONS
REACTIONS
Amylases
Starch ----------------> Dextrins, glucose and maltose
Intact Starch
Simple sugars will not stain
Stains dark
Purple by iodine
RESULTS

Add Gram’s iodine to Starch agar culture. Flood entire surface. Iodine stains intact starch
molecules. Pour off residual iodine into a pan of disinfectant. Observe plate from bottom while
holding up to the light. Clear zones around bacterial growth represent areas of starch breakdown.
B. subtilis is positive, E. coli and Proteus are negative.
Urease Activity
BACKGROUND

Some bacteria break down urea using an enzyme called urease to produce ammonia, CO2 and water. A urease test is
performed using agar containing urea and a pH indicator. When urease is produced by bacteria growing on this
medium, ammonia accumulates in the medium, and an alkaline reaction is observed. We will perform this test if the
urea agar is available. You should expect bacteria that produce urease to be capable of living in environments
where urea might be encountered such as during urinary tract infections and during formation of ulcers.
PURPOSE

To detect bacteria which have the ability to break down nitrogen containing molecules such as urea.
CULTURES
Escherichia coli, Proteus vulgaris, Corynebacterium pseudodiphtheriticum
MEDIA
3 Urea Agar plates (kept in fridge prior to use)
CULTURE
CONDITIONS
Spot inoculate urea agar plate and incubate inverted at 37oC.
REACTIONS

Urea --------------> Ammonia, CO2 and H2O
Urease
pH increases because of release of ammonia (a base). pH indicator changes from pale yellow to
pale pink when pH increases.
RESULTS
37

Following growth of an organism on urea agar, a pale pink color indicates positive urease activity.
The color change is subtle and is best seen when the plate is placed on a white background such as
paper. See unknown charts for results of tested organisms.
____________________________________________________________________________________
Casein Hydrolysis
PURPOSE

To identify, by another means other than litmus milk, bacteria which have the ability to breakdown
milk protein, casein
CULTURES
Bacillus subtilis, Escherichia coli, Pseudomonas aeruginosa
MEDIA

3 Casein agar plates (made by mixing 1 tube of sterile skim milk with 1 tube of nutrient agar.
Swirl mixture to blend; but do not swirl so much that agar sloshes onto lid.)
CULTURE CONDITIONS

Spot inoculate one bacterium per plate. Expect copious amounts of growth on this agar. Only one
bacterium can be placed per plate because of growth rate.
REACTIONS

Casein is the large protein that gives milk the opaque white appearance. Note that autoclave
sterilized milk is not stark white like pasteurized. This is due to carmelization of lactose at high
temperature. When casein is broken down, proteases or peptidases must be released from the
bacterium and diffuse into the agar. These enzymes break down the casein into soluble amino acids
and small peptides that can be easily transported into the bacterium. Breakdown of the casein will
result in clearing of the agar.
RESULTS

Note that Bacillus subtilis will be positive for peptonization in litmus milk and on casein agar. E.
coli is negative. P. aeruginosa will be strongly positive and will also produce copious amounts of
the classic fluorescent green pigment characteristic for this genus.
Gelatin Hydrolysis
BACKGROUND

Know that proteins are made up of carbon, hydrogen, oxygen, nitrogen and sometimes sulfur. Subunits making up
proteins are amino acids that are joined by peptide bonds.
38

Many bacteria breakdown proteins that are too large to be transported into the cells by producing proteolytic
exoenzymes (enzymes which are secreted from the bacterial cells which break down proteins). These enzymes break
down proteins into small peptides and soluble amino acids.

Examples of protein catabolism you will see in this lab include gelatin hydrolysis and milk protein (casein) hydrolysis.
Gelatin is a soluble mixture of proteins that gels at temperatures below 37oC. Gelatin hydrolysis can be used to assess
the pathogenicity of some bacteria. Gelatinase (a proteolytic exoenzyme) production can be correlated in some
cases with the breakdown of tissue collagen and dissemination throughout the body.

To perform a gelatin hydrolysis test, bacteria are inoculated into nutrient gelatin by stab. The tube is incubated at the
optimum growth temperature for the organism. If the organism is capable of breaking down the protein in the gelatin,
the matrix will liquefy. Unfortunately, 37oC is warm enough to liquefy gelatin. Therefore following incubation, the
tube must be chilled on ice or in the refrigerator for 15-30 minutes. If the gelatin remains liquid after chilling, the
organism hydrolyzed the protein. If the gelatin solidifies (gels), there has been no protein hydrolysis. Gelatin hydrolysis
tubes must be incubated for a minimum of 7 days before the results are read. Short incubations can give a false
negative for hydrolysis.
PURPOSE

To identify bacteria which produce proteases or peptidases which breakdown proteins outside the
bacterium into amino acids and small peptides that can be transported into the bacteria for energy or
protein synthesis.
CULTURES
Enterobacter aerogenes, Escherichia coli, Proteus vulgaris, Bacillus subtilis
MEDIA
4 Nutrient gelatin deeps
CULTURE CONDITIONS

Stab inoculate deeps using a needle or loop and incubate at 37oC with caps loose for a minimum of
7 days.
REACTIONS
Gelatin ----------------------------------> soluble amino acids and peptides
(Solid)
(Liquid even after refrigeration)

Gelatin is a protein made by boiling tissue collagen causing polypeptides to solubilize.
RESULTS

Following incubation, the tubes are transferred to the refrigerator for 15-30 minutes. If no protein
breakdown has occurred, the gelatin will solidify. (Gelatin will liquefy at 37oC even if proteins are
not hydrolyzed) If the protein has been broken down, the gelatin will remain liquid following
refrigeration. B. subtilis should be positive. E. coli and E. aerogenes should be negative. P.
vulgaris may give variable results. Some very vigorous, fast growing species can hydrolyze gelatin
protein in two days; however, in general 7 days are required for most bacteria and insures there will
be no false negative results.
____________________________________________________________________________________
Nitrate Reduction
39
BACKGROUND

During aerobic and anaerobic respiration, the electron transport system requires electron acceptor molecules. In
aerobic respiration, the final electron acceptor is oxygen. In anaerobic respiration, no oxygen is present so other
molecules must be the acceptors. Most often in anaerobic respiration, the final electron acceptor is an inorganic
molecule such as sulfate or nitrate; however, carbonate may also be used. In fermentation, the final electron
acceptor is some organic molecule. When these molecules accept electrons, they are reduced. When sulfate is
reduced, hydrogen sulfide is produced (H2S) and this can be seen by growing an organism on a TSI (triple sugar iron)
slant. The use of carbonate results in the production of methane gas which we cannot measure in this lab. The
reduction of nitrate may result in the production of nitrite or some other byproduct that can be monitored by the
nitrate reduction test.

A nitrate reduction test is performed by inoculating nitrate containing broth with a bacterium and growing overnight.
Nitrate reagents are then added to the tube. If the medium turns red, then nitrate has been reduced to nitrite. If there
is no color change, either the nitrate was not converted at all or the nitrite was further reduced to ammonium ions,
nitrous oxide or nitrogen gas. To determine which is the case, elemental zinc is added to the tube. Zinc will convert
any nitrate present into nitrite providing a red color (a negative test). If no nitrate is present, there will still be no
color change in the medium telling you that nitrate has been reduced (a positive test). You should expect nitrate
reduction to occur in organisms that have the capability of growing anaerobically where they would use nitrate
as an alternative electron acceptor in the absence of oxygen. One of the primary functions of many soil bacteria is
the conversion of nitrogen containing compounds into a form useable by plants, and in recycling nitrogen and returning
nitrogen to the air (the nitrogen cycle). Therefore, you would expect many soil bacteria to be capable of nitrate
reduction. A control tube (uninoculated) should be used to demonstrate the color expected and behavior of a negative
test when nitrate remains present.
PURPOSE

To identify bacteria which are capable of anaerobic respiration using nitrate as a final electron
acceptor.
CULTURES
Escherichia coli, Streptococcus lactis, Staphylococcus epidermidis, soil
MEDIA
4 Nitrate broths
CULTURE CONDITIONS

Inoculate broth and incubate at 37oC with tight caps. E. coli and S. epidermidis are facultative
anaerobes; S. lactis is a microaerophile. Soil will contain anaerobes and facultative anaerobes.
REACTIONS
e- + NO3- ---------------------------------> NO2(Nitrate)
Nitrate reductase (Nitrite)
e- + NO2- ---------------------------------> N2 or
(Nitrite)
(Nitrogen gas)
Ammonia
RESULTS

Add 5 drops each of Nitrate reagents A and B. These chemicals detect the presence of nitrite and
form a red color if nitrite is present. A red color at this point is a positive nitrate reduction test. If
no red color develops, there are two possibilities. 1: Nitrate is present meaning there has been no
40
nitrate reduction. 2: Nitrite has been further reduced forming nitrogen gas or ammonia. To
determine which is the case, add elemental zinc. Zinc will reduce nitrate if present to nitrite.
When the nitrite is formed, the broth will turn red. A red color upon addition of zinc means nitrate
was not reduced by the bacterium and this is a negative nitrate reduction test. If no color develops
when zinc is added, this means ammonia or nitrogen gas are present and this is a positive
nitrate reduction test.

S. epidermidis should be positive for nitrate reduction. S. lactis and E. coli are negative. You
should expect soil microbes to be positive because one of their jobs in the ecosystem is to break
down nitrogenous waste products in the soil and return nitrogen to the atmosphere.
Oxidase Activity
BACKGROUND

In bacteria that have electron transport systems, cytochrome c is one of the electron acceptors. The oxidase test assays
the action of the enzyme cytochrome oxidase, which carries electrons from cytochrome c to oxygen in organisms
capable of aerobic respiration. If oxidase activity is present, this is an indirect test for the presence of cytochrome c.
Bacteria are grown on TSA and oxidase reagent is dripped onto a colony. If the colony turns purple within 20-30
seconds, this is a positive test for oxidase. In the presence of cytochrome oxidase and free oxygen, oxidase reagent
serves as an electron donor allowing the oxidase enzyme to transfer electrons to oxygen forming water. When electrons
are removed from the oxidase reagent (oxidized), the reagent becomes purple. Many Gram negative pathogenic species
(N. gonorrhoeae, P. aeruginosa, Vibrio species) are oxidase positive but the enterics are not. If an oxidase test is
positive, the bacterium being tested must be capable of aerobic respiration. If an organism is oxidase negative,
some other enzyme must be capable of transferring electrons from cytochrome to oxygen, or the bacterium must be
capable of anaerobic respiration. Anaerobes do not require oxidase because oxygen is not the final electron acceptor.
A drawback to oxidase tests is that color development due to auto-oxidation of the reagent after 20-30 seconds of
exposure to air can be misread as a positive result. Iron-containing inoculating loops and needles should not be used to
transfer colonies prior to an oxidase test because iron is a potent catalyst of redox reactions which can actively donate or
accept electrons and interfere with the test results.
PURPOSE

To identify bacteria capable of aerobic respiration in which electrons must be shuttled from
cytochrome c in the electron transport chain and donated to oxygen as the final electron acceptor.
Cytochrome oxidase is the enzyme required to reduce oxygen.
CULTURES

Alcaligenes faecalis, Escherichia coli, Pseudomonas aeruginosa
MEDIA
3 TSA agar plates
CULTURE CONDITIONS

Perform a 4 quadrant streak for isolated colonies and incubate inverted at 37oC.
REACTIONS
Cytochrome c -------------e- ----------------> Oxygen =========>H20
Cytochrome oxidase
41
RESULTS

Add oxidase reagent to an isolated colony. A positive oxidase test occurs when the colony
turns purple within 20-30 seconds. The reagent will oxidize in the presence of oxygen after a
few minutes so you will eventually see purple color even on a negative oxidase culture. The
microbe is only positive for oxidase enzyme if the purple color develops quickly. In addition,
once opened, the oxidase reagent tube has a very limited shelf life. Never use a reagent that is
already purple or dark in color. Often the purple color in a positive oxidase colony will be
seen as a purple ring around the colony first because the cells on the edge are the ones
exposed to oxygen and growing by aerobic respiration. Often the center of a colony is not
growing aerobically. See unknown charts for expected results.
____________________________________________________________________________________
Effect of UV Light on Microbes
BACKGROUND

Radiant energy of short wavelengths has more energy than longer wavelengths. Ionizing radiation such as
gamma rays and X rays kill bacteria and other cells by destroying DNA molecules and other macromolecules.

Ultraviolet radiation is a form of nonionizing radiation that is required for processes such as photosynthesis and
Vitamin D synthesis. However, DNA absorbs UV light at a wavelength of 260 nm and can be damaged.

Some bacterial cultures will tolerate higher doses of UV radiation because macromolecules in the culture, such as
protein, RNA and DNA, as well as dead cells within the culture, will absorb UV light.

UV light does have some application for the sterilization of materials such as surgical instruments and surfaces
that might be damaged by heat or moist sterilants.

Some of the limitations to the use of UV light for bactericidal purposes include the lack of penetration. UV light
is not useful to sterilize liquids, cloth or other material except on the surface. UV light will not penetrate glass
and most plastics. UV light is also hazardous to eyes and skin and may cause blindness or skin burns. In
addition UV light can facilitate mutations resulting in the formation of malignant tumors.

UV light kills bacteria by forming thymine dimers in bacterial DNA. A thymine dimer is formed when a bond is
formed between two thymine nucleotides that occur side-by-side in one strand of DNA rather than to the purine
nucleotide (dATP) on the opposite strand. These dimers interfere with normal transcription and replication of
the DNA. Bacteria will die unless they can repair the damage. Most bacteria have the capability of repair.
Repair of thymine dimers may occur in one of two ways. Light repair or photoreactivation occurs when UV
damaged cells are exposed to visible light. Visible light activates an enzyme called pyrimidine dimerase (breaks
thymine dimers) that restores the DNA without removal of the damaged thymine nucleotides that were bonded
together. Dark repair involves the use of DNA polymerase to clip out damaged thymine nucleotides and
replacement with normal thymine nucleotides. DNA strands are then joined by DNA ligase. Mutations as a
result of UV damage may occur during the repair phase. If a nucleotide other than dTTP (dATP, dCTP or
dGTP) is inserted in the place of the thymine nucleotide, a mutation has occurred. This mutation may be lethal
or advantageous to the bacterium.
PURPOSE
To demonstrate the mutagenic effect of UV light on bacteria.
CULTURES
Staphylococcus aureus (broth)
42
MEDIA
2 TSA plates
CULTURE
CONDITIONS
Swab plates with bacteria. Treat with UV light for various times. Incubate upside
down at 37oC.
REACTIONS

UV light has a direct effect on the DNA of cells causing the formation of thymine dimers.
Once the dimers have formed, the DNA is damaged and must be repaired prior to cell division
(DNA replication) and gene expression (transcription). DNA may be repaired by dark repair
or photoreactivation. In dark repair, the proofreading ability of DNA polymerase recognizes
the dimer, cuts it out and replaces the nucleotides with new thymine nucleotides. DNA ligase
then seals the gap in the DNA backbone. At a very low frequency, the DNA polymerase will
add some nucleotide other than TTP (such as GTP, ATP or CTP). When this occurs, a
mutation has taken place. A mutation is defined as any change in a DNA sequence. During
photoreactivation, an enzyme called thymine dimerase, is activated by visible light. This
enzyme cleaves the covalent bond between the thymine dimer and allows the proper hydrogen
bonds to reform between the thymines and the complementary adenines. During
photoreactivation there is no chance of mutation.
RESULTS

The longer the exposure of a bacterium or any cell to UV light, the higher the probability of
DNA damage. With the formation of more thymine damage, there is less of a chance that all
thymine dimers can be repaired and cells are at a greater risk to die. This is why after longer
periods of exposure, the bacteria will not regrow. During shorter periods of exposure,
thymine dimers may be repaired completely resulting in regrowth. In addition, some of the
colonies that regrow will be mutants. Look for visible changes in colony color or appearance.
Note that UV light of 260 nanometers is most effective in damaging DNA because that is the
most effectively absorbed wavelength. UV light is not particularly effective as an
antimicrobial agent because a long direct exposure is required and there is a chance of
developing adaptive mutants.
____________________________________________________________________________________
Antibiotic Sensitivity (Kirby-Bauer Test)
BACKGROUND

Antibiotics are naturally occurring (produced by microbes to kill other microbes) antibacterial compounds.
Antimicrobics are naturally occurring or man-made chemical agents which are effective against microorganisms.

In order to determine if the zone of inhibition obtained by the Kirby-Bauer test is due to death or growth
inhibition of a bacterium, you must subculture from the zone into fresh media without antibiotics to see if any
bacteria grow.

Many factors influence the effectiveness of an antibiotic using the Kirby-Bauer test. These factors include: size
of the bacterial inoculum, distribution of the inoculum, period of incubation, agar depth, diffusion rate of the
43
antibiotic, concentration of antibiotic in the disk, growth rate of the bacterium, growth phase of the bacterium,
as well as the presence or absence of antibiotic resistance genes in the bacterium.

Bacterial cultures in early exponential phase of growth are generally most sensitive to antibiotics. Remember
most antibiotics work on cell walls, cytoplasmic membranes or ribosomes so actively growing cultures will be
most susceptible.

Just because an antibiotic works against a specific bacterium in a clinical lab does not mean the antibiotic will
work in a patient. Errors can occur in handling patient samples and in culturing the microorganisms from the
patient. Contamination can occur. The dose required to kill a bacterium in a laboratory may not be achievable
in a patient. Some patients will be allergic to antibiotics. Antibiotics often respond differently in the presence of
organic material. Bacteria may acquire mutations or antibiotic resistance genes in response to antibiotic
therapy.

Some of the reasons bacteria are becoming more resistant to antibiotics is that the more they are used, the
greater the probability of selecting for resistant mutants. Most antibiotics are given orally allowing more
contact with bacteria of the gastrointestinal tract where the development of resistances often begins. The use of
antibiotics in animal feeds to increase size and growth rates of commercial animals for food can result in the
selection of resistant microbial strains and residual antibiotics in the tissues of animals. Uninformed patients and
unthinking physicians nondiscriminantly demand and prescribe antibiotics, respectively, when antibiotics are
not warranted. "Take two aspirins and call me next week" will often work just as well as an antibiotic.
Remember, antibiotics are of no use in the treatment of viral diseases.

The Kirby-Bauer method is not limited to the testing of antibiotics. Other antimicrobial agents such as garlic
can be tested. Garlic inhibits the growth of many bacteria.
PURPOSE
To determine sensitivity of various bacteria to a variety of antibiotics.
CULTURES
Escherichia coli, Staphylococcus aureus, Pseudomonas aeruginosa
MEDIA
3 Mueller Hinton Agar Plates
CULTURE CONDITIONS

Use sterile swab to completely cover the entire surface of the plate with an organism. Then
apply a set of antibiotic impregnated disks to the surface of each plate and incubate agar side
down at 37oC.
REACTIONS

The sensitivity of a bacterium to an antibiotic is influenced by many factors listed in your lab
supplement and the lab manual. Be familiar with all of these. In the Kirby Bauer test,
antibiotics will diffuse into the agar from the paper disks and inhibit (bacteriostatic) or kill
(bacteriocidal) the bacterial culture. Zones of inhibition (clearing zones) will be seen in the
bacterial lawn if the bacterium is affected.
RESULTS

Just because an antibiotic has an apparent effect on a bacterium on a Kirby Bauer test does
not mean the antibiotic will work in the human body. Just because there is a zone of
44
inhibition, does not mean the organism is susceptible to the antibiotic. The zone must be a
certain minimum size if the bacterium is to be considered sensitive. Following growth of the
plates, use a ruler to measure the diameters of the zones and plug those numbers (in
millimeters) into the chart. You cannot determine whether an antibiotic is bacteriostatic or
bacteriocidal based on the Kirby Bauer test. This can be determined only if a sample is taken
from a zone of inhibition and is subcultured in media without antibiotic. If sample grows, you
know the organism was only inhibited in the presence of the antibiotic. If the organism does
not grow, the cells were indeed killed.
Disk
Symbol
AM
Antibiotic
Concentration
Resistant
Intermediate
Susceptible
Ampicillin against G- and Enterococci
10 g
11 mm or less
12-13 mm
14 mm or more
AM
Ampicillin against Staphylococci
10 g
20 mm or less
21-28 mm
29 mm or more
CM
Chloramphenicol
30g
12 mm or less
13-17 mm
18 mm or more
E
Erythromycin
g
13 mm or less
14-17 mm
18 mm or more
N 30
Neomycin
g
12 mm or less
13-16 mm
17 mm or more
P 10
Penicillin G for staphylococci
P 10
Penicillin G or other microbes
g
11 mm or less
12-21 mm
22 mm or more
S10
Streptomycin
g
11 mm or less
12-14 mm
15 mm or more
SSS 300
Triple Sulfa
300 g
12 mm or less
13-16 mm
17 mm or more
TE 30
Tetracycline
30 g
14 mm or less
15-18 mm
19 mm or more
10 g
20 mm or less
21 mm or more
___________________________________________________________________________________
Microbiology of Water (MPN Test)
BACKGROUND

Coliforms are aerobic or facultatively anaerobic, Gram-negative, nonendospore-forming, rod-shaped
bacteria that ferment lactose with acid and gas formation within 48 hours at 35 oC. These organisms are not
usually pathogenic; however, their presence in water usually indicates fecal contamination.

Coliforms are selected as the indicator of water potability (fitness or suitability for drinking) because they
indicate the presence of lactose-fermenting Gram-negative rods. Coliforms include Enterobacter, Klebsiella,
Citrobacter, and Escherichia.

When any kind of plating test is used to determine the presence of bacteria, colony forming units (CFU) are
counted to determine the number of bacteria present in a given volume of sample. A colony forming unit
represents a single cell from the original sample which grew by binary fission to form a visible unit on agar
(a colony) that can be seen with the naked eye. A 1:10 dilution is the same as a 10 -1 dilution. A 1:100
dilution = 10-2. If 1 mL of a 1:100 dilution was plated on agar, the number of CFU that are present after
incubation represents the number of bacterial cells present in l mL of that dilution. To determine the
number of bacteria present in 1 gram of the original sample, the dilution factor must be multiplied by the
45
number of colonies counted. In this example, if 100 colonies were counted from 1 mL of a 1:100 dilution (l
gram of sample in 99 mL water), then 100 (colonies) is multiplied by 100 (the dilution factor) to give you the
number of bacterial cells in l gram of sample (10,000 or 10 4 in this case). If 100 colonies were counted from
0.1 mL of a 1:100 dilution, then the amount plated must be factored into the equation. 100 colonies X 100
(dilution factor) X 10 (the volume plated was 1/10 mL) to give 100,000 cells or 10 5 cells. Appendices of your
text book contain more information on Scientific Notation and Dilutions if you need review.

The MPN test is qualitative because it only estimates the probable number of microorganisms in 100 mL of
water without actually counting them.

In the MPN test, the fermentation of lactose is a presumptive indication of coliforms in water. A positive
presumptive test indicates the water may not be potable. A positive presumptive test is seen when water
contains bacteria that are capable of fermenting lactose. A false positive presumptive test may occur when
lactose is fermented but not by a coliform. The confirmed test utilizes brilliant green lactose bile broth that
is selective and differential for coliforms. Growth and gas production is a positive confirmed test. EMB is
used for the completed test indicating the presence of coliforms. Gram staining is also necessary to confirm
the presence of Gram negative rods.

On EMB agar coliforms grow as small dark centered colonies with a metallic green sheen or dark purple
color.

Many bacterial diseases can be transmitted in polluted water, some of which are not caused by coliforms but
occur at a statistically increased rate if coliform contamination is present. These include bacterial dysentery
(Shigella dysenteriae), typhoid fever (Salmonella typhi), cholera (Vibrio cholerae), ear infections
(Pseudomonas), Urinary tract infections (Proteus), Hemorrhagic fever (Leptospirosis), diarrhea (E. coli) and
atypical pneumonia (Klebsiella).
PURPOSE

To determine the degree of coliform contamination in water samples using the Most
Probable Number Test.
CULTURES
None required. Water samples from home (at least 60 mL)
SUPPLIES
Sterile 1 and 10 mL pipettes, Pipette pumps
MEDIA

Single and double strength lactose fermentation broth tubes, EMB agar plates, brilliant
green lactose bile broth fermentation tubes
CULTURE CONDITIONS

When testing for coliforms, grow all samples at 37oC with loose caps or inverted.
46
Most Probable Number (MPN) index for various combinations of Positive and Negative results when various amounts
of water are tested
Number of tubes giving Positive reaction out of:
Number of tubes giving Positive reaction out of:
5 tubes with
10 ml each
0
0
0
0
1
1
1
1
1
2
2
2
2
2
2
3
3
3
3
3
3
3
4
4
4
4
4
4
5 tubes with
1 ml each
0
0
1
2
0
0
1
1
2
0
0
1
1
2
3
0
0
1
1
2
2
3
0
0
1
1
1
2
5 tubes with
0.1 ml each
0
1
0
0
0
1
0
1
0
0
1
0
1
0
0
0
1
0
1
0
1
0
0
1
0
1
2
0
MPN Index
per 100 ml
<2
2
2
4
2
4
4
6
6
5
7
7
9
9
12
8
11
11
14
14
17
17
13
17
17
21
26
22
5 tubes with
10 ml each
4
4
4
4
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5 tubes with
1 ml each
2
3
3
4
0
0
0
1
1
1
2
2
2
3
3
3
3
4
4
4
4
4
5
5
5
5
5
5
5 tubes with
0.1 ml each
1
0
1
0
0
1
2
0
1
2
0
1
2
0
1
2
3
0
1
2
3
4
1
1
2
3
4
5
MPN Index
per 100 ml
26
27
33
34
23
31
43
22
46
63
49
70
94
79
110
140
180
130
170
220
280
350
240
350
540
920
1600
>1600
Most Probable Number Procedure
47
48
REACTIONS

The MPN test identifies coliforms based on three parts: presumptive, confirmed and
completed tests. Coliforms include the genera Enterobacter, Klebsiella, Citrobacter and
Escherichia. In the presumptive test, lactose fermentation broth cultures are cultivated 48
hours at 37oC and are positive if fermentation with gas production occurs. In the confirmed
test, the highest dilution from the presumptive test is subcultured into brilliant green lactose
bile broth (which is selective and differential for coliforms) and the cultures are positive if
growth (turbidity) and gas production is seen. In the completed test, any positive confirmed
cultures are subcultured onto EMB agar plates to determine which species is present.
Coliforms form colored colonies on EMB agar as described in the unknown section of your
lab manual. The number of positive presumptive, confirmed and completed tubes out of five
are plugged into the MPN chart to estimate the degree of coliform contamination per 100 mL
of water.
RESULTS

The degree of coliform contamination in water indicates the potability of water and should
imply the degree of disease causing potential of the water sample. The more coliforms
present, the more likely it is that the water will cause disease when ingested by humans. MPN
tests are routinely done on rivers, lakes and other water sources especially in times of drought
because bacterial numbers can greatly increase in stagnant or slow moving water. Keep in
mind that other pathogens such as Amoeba can also greatly increase in number in these
environmental conditions.
____________________________________________________________________________________
Microbial Contamination of Foods and Beverages
BACKGROUND

Pasteurization is a method of processing raw milk with sufficient heat to destroy pathogenic microorganism in
the milk without destroying the physical and nutrient properties of milk. Pasteurization usually involves heat at
62.9 oC for 30 minutes or "flash" pasteurization by heating to 71.6 oC for 15 seconds. When you purchase milk in
the grocery, it is not free of bacteria. The point of pasteurization is to reduce the total number of bacteria
present in milk, and in the process the total number of pathogenic bacteria is reduced. Milk can be sterilized but
if you have ever tried canned milk, you know that both taste and color is changed by heat sterilization.
Therefore, most people prefer the pasteurization method. The commercial sale of unpasteurized milk is
prohibited by law. But remember, that reducing the overall number of bacteria of milk decreases the probability
that pathogens will be present, but does not guarantee the absence of disease-causing bacteria.

The presence of coliforms in milk and milk products is a major indicator of the sanitary quality of milk. The
presence of many coliforms means there is an increased probability of pathogens present in the milk.

Milk that contains a large bacterial load will contain a lower oxygen concentration than milk without bacteria.
Methylene blue is white in the absence of oxygen and blue in the presence of oxygen. If methylene blue turns
white when added to a milk sample, the milk is of poor quality. This is known as a methylene blue reductase test.
The more contaminants which are present, the more quickly reduction will occur. Milk is placed into quality
classes based on the speed of reduction.

Milk sours at room temperature because the bacteria that are present grow better at 25-40oC (mesophiles) than
at 4oC (refrigerator temperature).
49

Milk can be contaminated by humans when skin and fecal microbes are transferred to equipment or animals
themselves, by coughing, sneezing and breathing (oral microorganisms) and by accidental contamination with
fomites (soil, dust, hair, etc.). Remember the Schwann Ice Cream contamination with Salmonella in the spring of
1995. The Schwann company also handles meat products which are a likely source of Salmonella.

As a microbiological growth medium, milk contains many types of nutrients for bacterial growth, but is often a
poor choice because of the opacity (growth cannot be visualized as easily).

Bacteria that normally grow in milk include Streptococcus species, Lactobacillus species and sometimes the
pathogenic species such as Salmonella, Brucella, Listeria, and Mycobacterium.

The standard heterotrophic plate count is performed on food products to determine the number of viable
microorganisms in 1 gram of food. If greater than 106 bacteria per gram is present, the food may be hazardous
to consume.

The number of bacteria recorded by the plate count method does not accurately reflect the total bacterial count
from a sample because many microorganisms present may not grow in the medium utilized.

Using the standard plate count, plates with 25 to 250 colonies are counted and used for calculations because this
range allows a reasonably accurate count without requiring excessive time consumption.

When performing plate counts of bacteria from beef, the sample is mixed with water to yield a 1/10 dilution so
that the sample will be of a consistency that will allow accurate pipetting and subsequent dilutions.

Food poisoning occurs when food contains microorganisms.

Food intoxication occurs when foods contain toxins produced by bacteria; however, the microbes may no longer
be alive.

Standard Methods Agar or nutrient agar is used to determine the number of bacteria in food because it supports
growth of many bacteria commonly found in food and is light in color, which facilitates colony counting.

Foods to be tested for bacterial contamination should not be repeatedly frozen and thawed because each time
thawing occurs, more bacteria grow in the food, and these microbes are not killed by freezing.

Indicator organisms such as E. coli and other coliforms are used to gauge the sanitary condition of food
products. High concentrations of indicator organisms suggest a food may be unsuitable for human consumption.
These indicators may not be pathogenic but a high level of contamination increases the chances that a pathogen
will be present. Likewise, the absence of these organisms does not mean that no pathogens are present. Brilliant
green bile lactose broth is highly selective and differential for coliforms because only they will tolerate the
presence of bile and ferment lactose with gas production. It is not advisable to thaw and refreeze chicken
because each thawing process allows the growth of bacteria at the higher temperature. When the meat is frozen
again, the bacteria are not killed just slowed. Each subsequent thawing results in higher and higher numbers of
bacteria present.

Chickens are often contaminated prior to leaving the processing plant. One probable source of contamination
comes from machinery used to disembowel the carcasses. These machines may result in tearing of the intestines
allowing fecal contaminants to contact the meat. Another source of contamination is through the washing vats
(Bug soup), which the carcasses are sent through prior to packaging. There is also risk of contamination from
countertops, instruments and personnel prior to shipment. You must also safeguard yourself against further
contamination once you get home by using sanitary procedures in the kitchen including proper disinfection of
cutting boards, knives and countertops.
50
PURPOSE
To determine the degree of bacterial contamination in foods.
CULTURES
Food and Beverage samples
MEDIA
Nutrient agar and Eosin Methylene Blue (EMB) agar
CULTURE CONDITIONS

Incubate plates inverted at 37oC. Nutrient agar plates will be formed using the pour plate
method. EMB plates will be inoculated using the spread plate technique.
REACTIONS

Samples will be diluted according to the below figure for determination of total bacterial
counts. Duplicate samples will also be plated on EMB agar for the determination of the
percentage of coliforms present. EMB samples will be dispersed by using alcohol to sterilize
glass hockey sticks and spread as below:
Procedure for Creating Serial Dilutions and Platings:
51
52
RESULTS

Coliforms are indicator organisms in foods and water which signal fecal contamination. The
number of the organisms present can affect the potability of water and the safety of foods.
Coliforms are Gram negative, facultatively anaerobic, nonsporing rods that ferment lactose
with the production of gas within 48 hours at 37oC. Examples are the enterics such as
Escherichia coli, Proteus, Serratia, Enterobacter, Klebsiella, etc. In order to determine the
number of bacteria present per gram of food, the sample must be diluted in sterile water or
saline and plated so that individual colonies can be counted. Clearly the more bacteria
present in the original sample, the more the sample will have to be diluted to find a plate with
a countable number of isolated colonies. Be familiar with scientific notation and serial
dilutions presented in the appendix of your lab manual and your text book. On nutrient agar,
all colonies will be the same color. Count all colonies present. Those embedded in the agar
will look like tiny footballs in shape. Those on top of the agar will be very large and often
spreading. Each is counted as a colony regardless of size. A colony counter with magnifying
lens will be provided for counting. On EMB agar, various coliform species will have different
colors. Consult your unknown section of the manual for this information. EMB is a selective
and differential medium which means it selects for or only allows Gram negative organisms to
grow and you will be able to differentiate among species based on color. In order for ground
beef to be safe, there must not be more than 106 bacteria per gram of meat.
********************************************************************************
Before your laboratory responsibilities are over for this term, you must clean out any areas in which
you may have cultures, slides or supplies. These areas include the green cabinet where you may have
personal materials. Any slides in that area are to be cleaned well and returned to the slide boxes.
Cultures in the refrigerators and incubators should have the labels removed and any glass tubes
should be discarded on the cart. Any Petri plates should be discarded in the biohazard buckets. Your
unknown reports can be turned in any time prior to the due date. Early submissions will be gratefully
accepted.
53
UNKNOWN
IDENTIFICATION
UNKNOWN IDENTIFICATION
This exercise requires that you apply knowledge gained from your laboratory exercises. It will be necessary for you to
complete each of the assignments yourself. It will be in your best interest during the course of this term to repeat any
exercises in which you are lacking before you begin your unknown identification. Prior to obtaining an unknown
organism, you will have learned enough of the techniques that will be required for the identification of the unknown.
We will also have completed most of the biochemical tests that you might find useful in your identification. Once your
unknown has been issued, you may not ask your instructor or other instructors for help in identifying your organism(s).
The proper identification of an unknown organism(s) is essential in order for you to score well in this laboratory. You
will be issued an overnight culture of unknown bacteria. The culture will be tagged with a number. It is your
responsibility to record that number for your reference and on every page of the report you hand to the instructor.
Make observations of the broth culture as soon as you receive it using scientifically accepted terminology. Once that
culture is transferred to your hands, it is your responsibility to maintain that culture in useable form and contaminant
free!
Using aseptic technique, transfer a sample of the broth culture to two TSA (Tryptic Soy Agar) plates immediately and
incubate the plates at 25 and 37oC to determine the appropriate growth temperature for the bacterium or bacteria.
Also inoculate two TSA slants and grow at 37 oC as a backup in case your plates do not grow for some unpredictable
reason. Remember to label every culture you create in the proper manner and discard all cultures appropriately
when you are through with them.
Use careful technique to streak the TSA plates so that you may analyze colony morphology and separate each
bacterium of a mixed culture. If you deem it necessary to streak additional slants or plates, it is your responsibility to
inform the instructor of your needs and justify those needs with sound logic. We cannot afford the unnecessary waste
of microbiological media.
Staining tests can be carried out on the broth culture on the day of issue prior to discarding the tube. It will be in
your best interest to set up at least two Gram stains and one each of the following: acid-fast stain, endospore stain
and negative stain. It is critical that you also perform the appropriate staining techniques on any colony or culture
that you generate from the original culture to insure it matches the original stains.
Any subculture you create must be monitored regularly and the appropriate observations concerning bacterial
growth should be recorded in good scientific terminology. If your cultures become contaminated, it is your responsibility to rescue the appropriate organism.
Once you are sure you have established two pure agar slant cultures, you will use one as your working culture and the
other as an emergency backup. Your lab section will be issued a test tube rack on the refrigerator shelf with your
instructor's name. It is your responsibility to keep both of your cultures there at all times when not in use. Use only
the working culture for biochemical tests or any other means of identification. Be sure to stain your organism after
every biochemical test you perform. You may make a fresh subculture from the working stock whenever necessary.
54
Before submitting your results, go back and insure by staining that your subcultures are identical to the emergency
stock that was not handled.
If necessary, you may request specialized media or additional material if you can justify their uses and if we have
them available in the laboratory. You may use any reference material you deem appropriate. Computer software in
the Biology Study Center may help you. You are not to take your cultures out of the laboratory at any time. You are
not to seek help from your instructor or other instructors in the identification of your organism(s). You may find it
necessary to put in time other than your scheduled laboratory hours. Do not ask another faculty member to assist
you in the laboratory.
55
AEROBES
FACULTATIVE ANAEROBES
Gram Neg. Bacilli
Pseudomonas
aeruginosa
Pseudomonas
fluorescens
Alcaligenes
faecalis
Enterobacter
aerogenes
Escherichia
coli
Proteus
vulgaris
Proteus
mirabilis
Temp Preference
37oC
25oC, no growth
37o
37oC
37oC
37oC
37oC
37oC
Colonies on TSA
tan, circular,
diffusable
fluorescent green
pigment
tan, circular,
diffusable
fluorescent green
pigment
Mucoid,
spreading,
beige/pink
Heavily mucoid,
round, beige,
convex
Mucoid, beige,
spreading
Swarming
at 25oC, tan
pigment
Swarming
at 25 and
37oC,
darker tan
Glucose
Alkaline
Negative
Neg/Alk
Acid
Acid/gas
Acid
Acid/gas
Sucrose
Alkaline
Negative
Neg/Alk
Acid
Variable
Acid
Variable
Lactose
Alkaline
Negative
Neg/Alk
Acid
Acid/gas
Negative
Variable
Indole
Negative
Negative
Negative
Negative
Positive
Positive
Negative
MRVP
MR-VP-
MR+VP+
MR-VP-
MR+VP-
MR+VP-
MR+VP-
MR-VP-
Citrate
Positive
Negative
Positive
Negative
Negative
Negative
Positive
Lipid
Positive
Negative
Positive
Positive
Negative
Positive
Positive
Hemolysis
Beta
Gamma
Alpha
Gamma
Gamma
Gamma
Gamma
Litmus Milk Acid
Negative
Negative
Negative
Positive
Negative
Negative
Negative
Alkaline
Positive
Positive
Negative
Negative
Negative
Negative
Negative
Peptonization
Positive
Positive
Negative
Negative
Negative
Negative
Negative
Ropiness
Positive
Negative
Negative
Positive
Negative
Negative
Negative
Coagulation
Negative
Negative
Negative
Negative
Negative
Negative
Negative
Reduction
Positive
Negative
Negative
Negative
Negative
Negative
Negative
Starch
Negative
Negative
Negative
Negative
Negative
Negative
Negative
DNase
Negative
Negative
Negative
Negative
Negative
Negative
Positive
TSI slant
Alkaline
Negative
Alkaline
Acid
Acid
Acid
Alkaline
TSI butt
Negative
Negative
Negative
Acid/gas
Acid/gas
Acid
Acid/gas
TSI H2S
Negative
Negative
Negative
Negative
Negative
Positive
Positive
Gelatin
Positive
Negative
Positive
Negative
Negative
Negative
Negative
Motility
Positive
Positive
Negative
Positive
Positive
Positive
Positive
Urease
Positive
Negative
Negative
Negative
Negative
Positive
Negative
Oxidase
Positive
Positive
Negative
Negative
Negative
Negative
Positive
EMB
Lavender
Lavender
Lavender
Fisheye
Metallic green
Purple
Purple
Nitrate Reduction
Positive
Negative
Negative
Negative
Negative
Positive
Positive
MSA
Not tested
Not tested
Negative
Not tested
Not tested
Alkaline
Positive
56
Aerobic
Facultative Anaerobes
Microaerophilic/Slightly Facultative
Gram Pos. Cocci
Micrococcus
luteus
Micrococcus
roseus
Staphylococcus
aureus
Staphylococcus
epidermidis
Streptococcus
lactis
Enterococcus
faecalis
Temp Prefered
30oC
30oC
37oC
37oC
37oC
37oC
37oC
Bile Growth
Not tested
Not tested
Not tested
Not tested
Positive
Negative
Negative
Glucose
Negative
Negative
Acid
Acid
Acid
Acid
Acid
Sucrose
Negative
Negative
Acid
Acid
Acid
Acid
Negative
Lactose
Negative
Negative
Acid
Acid
Acid
Acid
Acid
Colonies on TSA
convex,
citron yellow
convex,
peach color
circular,
beige/gold
circular, stark
white
small, slow,
transluscent
small, slow,
transluscent
small, slow,
transluscent
MRVP
MR-VP-
MR-VP-
MR+VP-
MR+VP-
MR+VP-
MR+VP-
MR+VP-
Cell morphology
tetrads, and
staph
tetrads and
staph
staphylococci
staphylococci
strep in liquid,
staph on agar
strep in liquid,
staph on agar
strep in liquid,
staph on agar
Lipid
Negative
Negative
Positive
Negative
Negative
Negative
Positive
Hemolysis
Gamma
Gamma
Alpha
Alpha
Gamma
Beta
Alpha
Litmus Milk Acid
Negative
Negative
Negative
Top positive
Top positive
Top positive
Alkaline
Negative
Negative
Negative
Negative
Negative
Negative
Negative
Peptonization
Negative
Negative
Negative
Negative
Top positive
Top positive
Top positive
Ropiness
Negative
Negative
Negative
Negative
Negative
Negative
Negative
Coagulation
Negative
Negative
Negative
Negative
Positive
Positive
Positive
Reduction
Negative
Negative
Negative
Negative
Positive
Positive
Positive
Growth at high
temp
Not tested
Not tested
Not tested
Not tested
no growth at
45oC
45 not 50oC
growth
Growth at 45
and 50oC
DNase
Negative
Negative
Positive
Negative
Negative
Negative
Negative
TSI slant
Negative
Negative
Acid
Acid
Acid
Variable
Acid
TSI butt
Negative
Negative
Acid
Variable
Acid
Acid
Acid
TSI H2S
Negative
Negative
Negative
Negative
Negative
Negative
Negative
Gelatin
Positive
Positive
Negative
Negative
Negative
Negative
Negative
Sorbitol
Not tested
Not tested
Not tested
Not tested
Positive
Negative
Negative
Urease
Positive
Negative
Negative
Negative
Negative
Negative
Negative
Oxidase
Positive
Negative
Negative
Negative
Negative
Negative
Negative
Catalase
Positive
Positive
Positive
Positive
Negative
Negative
Negative
Nitrate Reduction
Negative
Negative
Positive
Positive
Negative
Negative
Negative
MSA
Positive
Negative
Positive
Negative
Negative
Negative
Negative
Negative
Enterococcus
faecium
57
Gram Positive Sporulating Bacilli
Gram Positive Nonsporulating Bacilli
Gram Pos. Bacilli
Bacillus cereus Bacillus
licheniformis
Bacillus
megaterium
Bacillus
subtilis
Bacillus mycoides
Lactobacillus casei
Corynebacterium
pseudodiphtheriticum
Temp Preference
37oC
37oC
37oC
37oC
37oC
37oC
30oC
Oxygen
Facultative
Facultative
Aerobic
Aerobic
Facultative
Aerotolerant
Aerotolerant
Glucose
Acid
Acid
Acid
Acid
Negative
Acid
Peach
Sucrose
Acid
Acid
Acid
Acid
Negative
Acid
Acid
Lactose
Negative
Negative
Negative
Negative
Negative
Variable
Negative
Indole
Negative
Negative
Negative
Negative
Negative
Negative
Negative
MRVP
MR-VP-
MR+VP-
MR+VP-
MR-VP-
MR-VP+
MR+VP-
MR-VP-
Citrate
Positive
Positive
Negative
Positive
Negative
Negative
Negative
Lipid
Positive
Positive
Positive
Positive
Positive
Negative
Positive
Hemolysis
Beta
Beta
Beta
Beta
Beta
Gamma
Gamma
Litmus Milk Acid
Negative
Negative
Negative
Negative
Negative
Positive
Negative
Alkaline
Negative
Negative
Positive
Positive
Negative
Negative
Negative
Peptonization
Positive
Negative
Positive
Positive
Negative
Positive
Negative
Ropiness
Negative
Negative
Positive
Negative
Negative
Negative
Negative
Coagulation
Negative
Negative
Negative
Negative
Positive
Positive
Negative
Reduction
Positive
Positive
Positive
Positive
Negative
Positive
Negative
Starch
Positive
Positive
Positive
Positive
Positive
Negative
Negative
DNase
Positive
Negative
Positive
Negative
Negative
Negative
Positive
TSI slant
Alkaline
Alkaline
Alkaline
Alkaline
Acid
Acid
Acid
TSI butt
Acid
Acid/gas
Acid
Acid
Neg
Acid
Acid
TSI H2S
Negative
Negative
Negative
Negative
Negative
Negative
Negative
Gelatin
Positive
Positive
Positive
Positive
Positive
Negative
Negative
Motility
Negative
Positive
Positive
Negative
Negative
Negative
Negative
Urease
Negative
Positive
Negative
Negative
Positive
Negative
Negative
Oxidase
Negative
Variable
Variable
Variable
Negative
Negative
Negative
Catalase
Positive
Positive
Positive
Positive
Positive
Negative
Negative
Nitrate Reduction
Positive
Positive
Positive
Positive
Negative
Negative
Negative
MSA
Positive
Positive
Alkaline
Positive
Negative
Not tested
Negative
Unique feature
Large,
spreading,
waxy, flat
dull, rough,
strongly attached
to agar, slimy
dry, creamy,
undulate edges,
spreading
large,
creamy,
spreading
large, spreading,
lacy edges
small, translucent,
glossy
pleomorphic rods and
“0” shapes, partially
acid fast, transluscent,
small
58
Gram Pos., Acid Fast Bacilli
Mycobacterium smegmatis
Mycobacterium phlei
Temp Preference
37oC
37oC
Oxygen
Aerobic
Aerobic
Glucose
Negative
Negative
Sucrose
Negative
Negative
Lactose
Negative
Negative
Indole
Negative
Negative
MRVP
MR-VP-
MR-VP-
Citrate
Positive
Positive
Lipid
Positive
Positive
Hemolysis
Gamma
Gamma
Litmus Milk Acid
Negative
Negative
Alkaline
Negative
Negative
Peptonization
Negative
Negative
Ropiness
Negative
Negative
Coagulation
Negative
Negative
Reduction
Negative
Negative
Starch
Negative
Negative
DNase
Negative
Negative
TSI slant
Alkaline
Alkaline
TSI butt
Negative
Negative
TSI H2S
Negative
Negative
Gelatin
Negative
Negative
Motility
Negative
Negative
Urease
Positive
Positive
Oxidase
Negative
Negative
Catalase
Positive
Positive
Nitrate Reduction
Positive
Positive
MSA
Negative
Negative
Unique features
Waxy, clumpy colonies, Growth on LJ agar,
Growth at 45oC in TSB, but not at 52oC
Waxy, clumpy colonies, Growth on LJ agar,
Growth at both 45oC and 52oC in TSB
59
Growth Characteristics of Unknowns
Optimal Growth Temperature: Determine the optimum growth temperature by growing the unknown at 25 oC and
37oC and note where the organism grows best. Most of the possible unknown organisms will be mesophilic and will
grow well at either temperature. If an organism which is red at 25 oC but only slightly pink at 37oC, it is a mistake to
presume the organism prefers room temperature growth. 37 oC is the optimum temperature of growth for Serratia
marcescens. A characteristic phenotype such as a color is different from a growth rate. Preferred growth
temperature will be established on the first streak plates generated from the unknown broth.
Colony Appearance: Record the growth characteristics of the bacterial colonies. Include descriptions of colony
margins, colony diameter, elevation, color, consistency (Is it dry, mucoid, etc.?), degree of spreading. You will do
yourself a favor if you make notes throughout the term on each organism we use for lab experiments. Most of these
organisms will be possible unknowns. The following are terms used to describe colony characteristics on agar as you
would see them with the naked eye. These terms must only be applied to isolated colonies not growth on slants or as a
lawn. Colony form refers to the overall shape of the entire colony. Elevation refers to shape and distance a colony
comes up from the surface of the agar while looking at the side of the colony. The margin refers to the shape of the
colony border.
Colony Form
Colony
Elevation
Colony
Margin
Growth on
slant
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Staining Characteristics: Perform a Gram stain on the sample you obtain as your unknown. Be aware that if you
have an acid fast organism as your unknown or one of your unknowns, it will not consistently give you the correct
Gram stain if it Gram stains at all. Don't make the mistake of missing one of the unknown organisms just because you
don't have a clear-cut Gram stain. It is also critical that you perform the appropriate stain ( either a Gram stain or
acid fast stain is usually sufficient) following each biochemical test you perform. Every time you perform a test, you
will want to insure that there is only one organism in the tube and that it is the correct organism. One wrong result
can lead you on a wild goose chase and contamination will happen even to the most careful person when you least
expect it. When you report your Gram staining results, they should include the Gram reaction as well as the shape.
For instance you would report that you have a Gram Negative Bacillus or a Gram Positive Coccus. When you do the
acid fast test, you also report the reaction and shape. For instance an Acid Fast Bacillus or a Non-Acid Fast Coccus.
Growth Tests: You will also want to perform Gram Stains on cocci which have been grown in broth and compare
them to stains from agar cultures. This is because all the cocci will have a staphylococcus arrangement when grown
on agar. This is often an artificial result because you are scraping many groups off the agar and placing them on a
slide where they are all bunched together. In a broth, you will see the true arrangement because the groupings space
themselves out within the broth. You may also need to grow some of the Gram Positive Cocci at elevated
temperatures especially if you are trying to differentiate between streptococci (Streptococcus lactis, Enterococcus
faecium, and Enterococcus faecalis). We have a 45oC water bath kept at the front of the lab and the 50oC holding tank
can also be used for incubations. Put your organisms in TSB and check for ability to grow at elevated temperature by
looking for turbidity. Bile tolerance may also be determined by a simple growth test. Use brilliant green lactose bile
broth, inoculate and grow the organism at preferred temperature. If the organism grows, it is bile tolerant (positive).
This medium is also used in the MPN test where both bile tolerance and lactose fermentation is important. For bile
tolerance only, it is not important to see a gas bubble in the Durham tube.
Motility testing: Use a 24-hour culture of the unknown to determine motility. You may perform a hanging drop test
on the sample on the day you receive it, but be aware that if you have two organisms only one of them might be motile.
It is best to wait until you have a pure culture of each unknown organism prior to performing motility testing using
semisolid agar media. If you have a coccus, it will be nonmotile. It is a waste of your time and the lab media to
perform motility testing on a coccus.
Oxygen Requirements: If you suspect that you have an anaerobic or facultatively anaerobic organism, you may want
to grow the organism in the absence of oxygen. Growth of microaerophiles will be boosted if you use a candle jar;
however, microaerophiles will not grow in an anaerobic chamber. Microaerophiles will grow as tiny, pinpoint,
translucent colonies on agar in the presence of oxygen. Anaerobes, aerotolerant anaerobes and facultative anaerobes
will grow in a GasPak canister. Additionally, using these techniques, you will be able to exclude aerobes.
Biochemical Tests (This information will appear on Lab Exam II)
Sorbitol, Dulcitol, Mannitol, Glucose, Sucrose, and Lactose fermentation broths (see Sugar Fermentation Tests).
Gelatin Hydrolysis: The production of proteases may lead to gelatin hydrolysis. This test is performed in nutrient
gelatin by inoculating the gelatin by stabbing and allowing it to incubate at the organism’s preferred temperature for
7-10 days. A positive result is determined when the solid gel is liquefied and remains liquid after the tube has been
chilled to below 25oC. The tube is usually refrigerated 15-30 minutes (4oC) prior to reading results. Stain culture
before placing in the refrigerator to confirm only one organism is growing in the tube. It may be necessary to remelt
gelatin if it is chilled prior to staining. A positive gelatin hydrolysis test means the organism produces peptidases or
proteases which break down protein in the medium and allow the bacterium to transport amino acids and small
peptides into the cell for energy or protein synthesis. A positive test also may mean the organism is capable of
spreading through tissue during infection by producing enzymes to degrade protein.
Fat Hydrolysis: The production of lipases results in breakdown of triglyceride (simple fat) into glycerol and three
fatty acids. This test is performed by spot inoculating one unknown organism per plate in the center of a Spirit Blue
Agar plate. A positive reaction is indicated by a lightening or clearing of the blue color. The color change is the result
of the release of fatty acids which acidify the medium. The pH indicator in the agar responds to the drop of pH by
changing to a lighter color. The plate should be incubated at the organism's optimum temperature for 24-48 hours.
Stain culture following reading.
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Starch Hydrolysis: The production of amylase results in the breakdown of starch. Starch agar comes in the form of a
deep and will need to be melted and poured into a petri plate. Spot inoculate the unknown onto starch agar plate and
incubate at the optimum temperature for 24 hours. Prepare a smear for staining prior to adding the iodine. Flood the
plate with Gram's iodine and pour off residual iodine into a container of disinfectant. The iodine will stain intact
starch dark brown or purple. The areas where the starch has been broken down by amylase will be clear because
simple sugars do not stain.
Indole test: Some bacteria will hydrolyze tryptophan producing pyruvate, ammonia and indole. This test is
performed by stab inoculating a SIM deep and incubating 24-48 hours at the optimum temperature. Prepare a smear
from the growth at the top of the SIM agar prior to adding the Kovac’s reagent. 5 drops of Kovac's reagent
(dimethylaminobenzaldehyde) are added to the tube following growth. A positive test is read as a red ring at the top
of the tube.
Sugar Fermentation Tests: Fermentation reactions can be tested on a variety of sugars (glucose, sucrose, lactose,
sorbitol, mannitol, dulcitol, etc.). The test is performed using a broth with the appropriate sugar plus a phenol red
indicator. The tubes are inoculated and incubated 24-72 hours at the optimum temperature. Acid production is
indicated by a bright yellow color change. Gas production is seen as a gas bubble trapped inside the small inner tube
(Durham tube). Some bacteria may grow in the broth without producing acid and gas and are scored as negative.
Some bacteria will produce an alkaline pH which is seen as a pink or mauve color in the broth. In addition,
contamination can result in an incorrect result. Perform Gram or other appropriate stains following a fermentation
test to insure that there is only one organism in the tube.
MRVP (Methyl Red/ Voges-Proskauer Test): Some strains of bacteria will produce a mixture of acids during
fermentation which may be detected by the methyl red (MR) test. The test is performed by inoculating MRVP broth
with the unknown and growing 24 hours at the optimum temperature. Prepare a smear of the growth before going
on. Five drops of methyl red are added to the tube which is left undisturbed. A positive test results in a pink/red
color of the medium. Positive MR indicates the presence of a mixed acid fermenter which converts glucose in MRVP
broth to a mixture of acids and thus create a very low pH which is indicated by the red color of the Methyl Red pH
indicator. The Voges-Proskauer (VP) test allows identification of butanediol fermenters by reaction with the neutral
product acetoin. Butanediol fermenters produce acetoin as a neutral precursor produce of glucose fermentation
before converting it to 2,3-butanediol and some acids as well. Because of the production of this alcohol and less acid,
the pH of the medium is higher in this case. The unknown is added to MRVP broth and incubated at optimum
temperature for 24 hours. 15 Drops of V-P reagent I and 5 drops of V-P reagent II are added to the tube. The tube is
shaken to oxygenate the media and allowed to stand 15 minutes. A positive test is seen as a red to pink color at the
surface of the liquid. A negative VP test is seen as a cloudy beige layer or no change.
Catalase Production: Tests for the production of the enzyme, catalase. Grow organism on TSA agar at the optimum
temperature and prepare smear prior to testing. A drop of 3% hydrogen peroxide is placed on a bacterial colony.
Production of bubbles is a positive test for the breakdown of hydrogen peroxide producing water and oxygen. This
test cannot be performed on a blood agar surface; however, cells from a blood culture can be removed from the blood
then tested.
Triple Sugar Iron Agar (TSI): TSI medium contains three sugars (glucose, sucrose and lactose) and iron salts. The
test may be used to detect acid and gas production and also production of hydrogen sulfide. The slant is inoculated
with the unknown by streaking the slant and then using the loop to stab through the agar. The tube is incubated for
24-72 hours and observed for the following reactions: Acid production = yellow color; Alkalinity = red color; Gas =
bubbles or cracks in the agar; Hydrogen sulfide = blackening of the agar. Prepare a stain of growth seen.
Citrate Utilization: The unknown is streaked/stabbed into a Simmon's citrate agar slant and allowed to grow at the
optimum temperature. This test requires oxygen (loose cap), and the ensyme, citrate permease, must be present in the
bacterium. If citrate is used as a carbon source, the agar will change color from green to royal blue. Stain organisms
growing on agar. The color change is caused by the release of sodium carbonate (a base) which causes the
bromothymol blue to turn from green to blue.
Litmus Milk Reactions: Litmus milk (containing 10% skim milk and litmus pH indicator) is inoculated with the
unknown and allowed to incubate 2-7 days at the optimum temperature. There are several possible reactions. Pink
color = fermentation of sugars with acid production; Purple = alkaline reaction; Peptonization = clearing of liquid
indicates fat and protein hydrolysis; Reduction = a bone white color in the milk especially in the lower half;
Coagulation = curd formation due to denaturation of milk proteins; Ropiness = trailing of strings or ropes of
thickened material behind an inoculating loop when passed through the milk. Gas production will only be detected
62
when coagulation takes place and is seen as cracks in the curd. Don’t forget to stain growth; however, be aware that
a pink background color will develop when milk is Gram stained. The background may make it harder to verify
Gram negative cells.
Mannitol Salt Agar (MSA): Used to distinguish the Gram positive cocci from other organisms as well as other salt
tolerant species. The agar contains mannitol (a sugar/alcohol), phenol red pH indicator and 5% sodium chloride (the
Staphylococcus species are salt tolerant). The agar comes in the form of a deep. It may be stabbed and incubated. It
may also be melted, poured into a plate and streaked (4 quads) when solid. Temperature of incubation depends on the
organism’s optimum growth temperature. A positive MSA test is scored by growth and a bright yellow color change
in the agar. The yellow color comes from the production of acid from fermentation of mannitol, and the phenol red
pH indicator turns yellow in acid. Likewise, alkaline products cause phenol red to turn fuchsia. Some organisms will
be salt tolerant, but will produce alkaline produces. These are scored as negative in the MSA test. Don't be fooled if
you have a yellow colored organism growing on the agar. Streptococcus and Enterococcus species grow more slowly
on MSA with only slight fermentation. Micrococcus species grow slowest and are usually negative for the color
change. Stain organisms following growth.
Oxidase Production: Tests for the production of the enzyme, oxidase, which is produced by certain bacteria. Grow
bacteria on a TSA agar plate and prepare a smear prior to testing for oxidase. Temperature of incubation depends on
the organism’s optimum growth temperature. Following growth, prepare a Gram of AF stain. Do not add chemical
until staining result have been viewed and recorded. A drop of oxidase reagent (diphenylamine) is placed on a
bacterial colony. The colony will darken to purple after 20-30 seconds if the test is positive. Any purple color after 30
seconds is read as a negative test because of exposure of the chemical to oxygen. Make sure the chemical is fresh
before using.
Nitrate Reduction: Nitrate may be converted to nitrite by certain bacteria. The unknown is inoculated into nitrate
broth and incubated at optimum temperature for 24-48 hours and a smear is prepared prior to going on. Be sure tube
in incubated with tight cap since this test is used as in indicator of anaerobic respiration where the nitrate will be used
as a final electron acceptor (therefore reduced). Also be sure to check for turbidity before performing a test. You will
get false negative results if your organism does not grow. Equal volumes (5 drops) of reagent A (sulfanilic acid) and
reagent B (napthylamine) are added to the tube. A positive test results in bright pink or blood red color development.
Many Bacillus, Pseudomonas and Micrococcus species reduce nitrate. If no color develops, zinc is added to the tube.
If the reaction was truly negative, nitrate will be reduced to nitrite, and the broth will turn red in the presence of zinc
(negative test). If nitrite was reduced further to nitrogen or ammonia, zinc addition will still result in lack of color
(this is a positive test) for nitrate reduction. It may take about five minutes for zinc to form a red color.
Eosin Methylene Blue (EMB) Agar: This medium is selective and differential for Gram negative bacteria because the
dyes inhibit everything else. The dyes are also absorbed differently by each genus that tolerates them. The agar is
dark purple when solid and is in the form of a deep. The agar must be melted and poured into a petri plate. When
solid, perform a 4 quadrant streak. Grow 24-48 hours. Temperature of incubation depends on the organism’s
optimum growth temperature. The colony color is distinctive for several genuses: Escherichia = metallic green
colonies; Enterobacter = mucoid colonies with dark centers (Fisheye); Proteus = thin, purple, flat, spreading colonies;
Serratia = dark purple colonies, Pseudomonas and Alcaligenes = lavender colonies. Don’t forget to stain the organism
following growth.
MacConkey Agar: This medium is selective and differential for enteric bacteria. The agar is light sensitive and is
kept in a cabinet in the biology stock room. You must request it, then melt the agar and pour a petri plate. When
solid, perform a 4 quadrant streak and grow 24-48 hours. Temperature of incubation depends on the organism’s
optimum growth temperature. Bile salts in the medium inhibit growth of many bacteria especially Gram positives.
Lactose fermenters appear as pink colonies. Non-lactose fermenters are white. Gram stain following growth.
Blood Agar (Hemolysis): 5% sheep erythrocytes in an agar base are used to determine if bacteria are capable of
breaking down the red blood cells resulting in a clearing or other change in the agar. This is an important
differentiating test for streptococci and staphylococci; however, this medium is expensive and should not be used
unless truly warranted. For example, if you check the unknown charts, you should notice that all Gram negative rods
and Gram positive rods, respectively, yield the same results on blood agar. The hemolysis test does not provide any
relevant information toward the identification of organisms in the Gram negative or Gram positive rod categories and
should not be used. Please avoid needless waste of media in the laboratory at all times. Blood is kept in the clean
refrigerator at the back of the room. Obtain a plate, be sure to check the expiration date. Expired blood or even
plates close to the expiration date may give varying results. On thing sure to happen on older blood agar plates is
oxidation or “browning” of the agar when the plate is incubated at 37oC overnight. These brown plates cannot be
63
read reliably. If plate is not expired, warm to room temperature. Streak the blood agar plate using a 4 quadrant
streak and incubate for 24 hours. Check. If results are unclear, recheck in 24 hours. Do not incubate longer than 48
hours total. Temperature of incubation depends on the organism’s optimum growth temperature. Beta hemolysis is
indicated by complete clearing of the blood agar adjacent to the bacterial colony which means the RBCs were
completely destroyed. Alpha hemolysis is usually seen as a greenish opaque color around the colony when you hold
the plate up to the light indicating that the hemoglobin was modified into methemoglobin thus changing the color.
Gamma hemolysis is inapparent hemolysis. However, if you scrape a colony off the plate, you may be able to see a
clearing under the colony. Stain any growth seen to insure no contamination has occurred.
Lowenstein-Jensen (LJ) Agar: Used to identify the Mycobacterium species. The agar contains eggs and malachite
green dye which kills most other bacteria. If Mycobacterium is suspected, streak the slant and incubate at 37oC with
the cap loose. This genus requires oxygen so a loose cap is essential. This is one of the few tests used to identify this
genus. In addition growth at higher temperatures in TSB is the conclusive test. Mycobacterium phlei grows vigorously
on LJ agar and produces a tan pigment. Mycobacterium smegmatis grows more slowly on LJ agar and has a tan-white
color. Remove a sample for acid fast staining following growth.
64