LABORATORY MANUAL
MICR231: General Microbiology
2017
KELLY PESEK
Table of Contents
Lab #1: Care and Use of the Microscope
p2–8
Lab #2: Examination of Living Microorganisms
p 9 – 15
Lab #3: Microbes in the Environment
p 16 – 22
Lab #4: Gram Staining
p 23 – 27
Lab #5: Effectiveness of Hand Scrubbing
p 28 - 31
Lab #6: Epidemiology
p 32 – 36
Lab #7: Agglutination Reactions: Slide Agglutination
p 37 – 42
Lab #8: Isolation of Bacteria by Dilution Technique
p 43 – 48
Lab #9: Physical Methods of Control – Heat
p 49 – 54
Lab #10: Oxygen and the Growth of Bacteria
p 55 – 59
Lab #11: Chemical Methods of Control – Antimicrobial Drugs
p 60 – 65
Lab #12: Chemical Methods of Control – Disinfectants and Antiseptics
p 66 – 69
Lab #13: Bacteria of the Skin
p 70 – 76
Lab #14: Make-up Lab – Microbes in Food: Contamination
p 77 - 81
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Lab #1: Care and Use of the Microscope
Objectives:
After completing this experiment, students should be able to:
1. Demonstrate the correct use of a brightfield microscope.
2. Name the key components of a brightfield microscope and determine their uses.
3. Calculate total magnification.
Brightfield Compound Light Microscope
The light microscope is an important tool in the study of microorganisms. The compound light microscope
uses visible light to directly illuminate specimens in a two lens system, resulting in the illuminated specimen
appearing dark against a bright background. The two lenses present in a compound microscope are the ocular
lens in the eyepiece and the objective lens located in the revolving nosepiece.
Compound light microscopes typically have the following components:
Illuminator: the light source in the base of the microscope.
Condenser: a two lens system that collects and concentrates light from the illuminator and directs it to
the iris diaphragm.
Iris Diaphragm: regulates the amount of light entering the lens system.
Mechanical Stage: a platform used to place the slide on which has a hole in the center to let light from
the illuminator pass through. Often contains stage clips to hold the slide in place.
Body tube: Houses the lens system that magnifies the specimens
Upper end of body tube -- Oculars/Eye pieces: what you view through
Lower end of body tube -- Nose-piece: revolves and contains the objectives
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Principles of Microscopy
Basically, a light microscope magnifies small objects not normally visible to the naked eye and makes them
visible. The science of microscopy is based on the following concepts and principles:
Magnification is taking the specimen and making it larger. In a compound lens system, each lens further
enlarges or magnifies the specimen. The objective lens magnifies the specimen, producing an image that is
then further magnified by the ocular lens resulting in the final image that we see when we look in the
microscope. The total magnification can be calculated by multiplying the objective lens value by the ocular
lens value. (example: 40x objective x 10x oculars = total magnification of 400x
Resolving power is the ability of a lens to show definition between two objects right next to each other. In
general, the shorter the wavelength of light, the better the resolution, which is why a blue filter is usually
connected to the condenser to produce short light waves for optimum resolution.
Resolving power is also dependent on the refractive index or the bending power of light. Because air has a
lower refractive index than glass, light waves have a tendency to bend and scatter as they pass through the air
from the glass slide to the objective lens. Addition of immersion oil, which has the same refractive index as
glass, diminishes the loss of refracted light and improves resolution.
Contrast is the ability to distinguish an object on a slide from the background. Since most microbes are
relatively colorless when viewed under a standard light microscope, they can be difficult to identify. Using a
stain that will bind to the microorganism and not the slide will dramatically improve the contrast allowing
microorganisms to be observed more clearly and easily.
Depth-of-focus is the “thickness” of the sample that is seen clearly at a particular magnification. As the
magnification increases the depth-of-focus decreases, or the “slice” of the sample that we see in focus gets
thinner (think layers in a cake). Many newer compound microscopes are parfocal, meaning that if you are in
focus using one objective, you can switch to the next objective and still retain your primary focus. Only fine
focus is required to see a crisp and clear image. This is due to the fact that as you increase the magnification
and the “slice” of the sample you see in focus becomes thinner, the correct plane-of-focus will always be
within the depth of focus of the previous objective. After you get the sample into focus on low power using
the course adjustment knob, you only need to use the fine focus knob at the higher magnifications.
Field-of-View is the area of the slide that you are seeing when you look into the microscope. As you increase
the magnification, the area of the slide that you are looking at gets smaller, thus the field-of-view becomes
smaller. Since you are now closer to an object on the slide, you see a smaller area of the slide at a time.
(Think about using Google Maps – as you zoom in closer to a location on a map, you see less of the map
overall.)
Working distance is the distance between the bottom of the objective and the slide. As you increase
magnification, the working distance decreases (the bottom lens of the objective gets closer to the slide you
are examining. When you are using the oil-immersion objective (100X), the objective is almost touching the
slide, allowing the immersion oil to touch both the slide and objective. There are two primary reasons why
you should always be aware of the working distance. The first is so that you do not inadvertently push the
objective through the slide, causing damage to the slide and possibly the objective. The second is to estimate
whether you are in the correct plane-of-focus.
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Care of Microscopes
Microscopes are very expensive pieces of scientific equipment and must be treated with care.
Some basic rules of microscope care include the following:
1. Always carry a microscope with two hands, one on the base and one on the arm.
2. Never tilt the microscope, as the eyepieces may fall out.
3. Look at the slide with both eyes open, to avoid eye strain.
4. Always focus by moving the objective away from the slide.
5. Before using the oil immersion lens, have your slide in focus under high power. ALWAYS focus with
low power first.
6. Use the coarse focus knob on the lowest objective only. NEVER use coarse focus on high power or oil
immersion, or you may damage the objective lenses.
7. Always clean all lenses thoroughly with lens paper (and lens cleaner if needed) before putting away.
Immersion oil which is not removed immediately can dry on the lenses, making it difficult to view any
specimen. Dried immersion oil is also quite difficult to remove from the lenses, not mention can cause
permanent, irreparable damage to the microscope.
8. Always store the microscope with the low power objective (10x) in place. NEVER store the microscope
with the oil immersion objective in place, as it can damage the lens.
9. When you have a problem with the microscope you are using, ask the instructor for help.
Review of Focusing
Place the microscope in a position which allows you to look into the eyepiece comfortably.
1. Properly adjust illumination according to the light source used.
2. Rotate the nosepiece and move the low power (10X) objective into the lock directly above the slide.
3. Adjust the oculars for the correct distance between your eyes.
4. Place the slide (side with identification up) on the stage so that it is held firmly by the mechanical
stage.
5. Use the mechanical stage knob to position the slide so the area to be examined is directly over the
illuminated part of the stage.
Focusing on Low Power
6. Rotate the coarse adjustment knob and bring the stage up slowly to its positive stop or within a few
millimeters on the slide.
7. Look through the oculars and rotate the coarse adjustment knob slowly to focus away until the
specimen comes into view.
8. Use the fine adjustment knob to bring the specimen into sharp focus on low power.
Focusing on High Power
9. Rotate the microscope nosepiece and lock the high power 40X objective into the viewing position.
10. Rotate the fine adjustment knob slowly to bring the specimen into sharp focus.
11. Readjust the iris diaphragm as needed to avoid eye strain.
Focusing on Oil Immersion
12. Rotate the nosepiece slightly to provide an open space above the illuminated area of the slide. Place a
drop of immersion oil on the spot.
13. Rotate the nosepiece away from the high power objective and bring the oil immersion objective (100X)
into contact with the drop of oil.
14. Rotate the fine adjustment knob slightly and bring the specimen into sharp focus.
15. Readjust the iris diaphragm to avoid eye strain.
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Stage of the Microscope
Basic Shapes of Bacteria
Materials
Brightfield microscope
Immersion oil
Lens paper
Prepared slides of bacteria, protozoa, fungi and algae
PROCEDURE
1. Place the microscope on the lab table directly in front of you.
2. Obtain a slide of algae or fungi and secure it to the stage using the stage clips.
3. Go through the steps of proper focusing using low power, high power and oil immersion, drawing a
picture of the microorganisms seen on low, high and oil-immersion. Complete this part of the lab
report form.
4. When you have completed your observations with the first slide, swing the low power objective into
the viewing position (never switch back to high-power when there is oil on the slide as this will get on
the objective). Take the slide off the stage and clean it with lens paper. Return the slide to the slide
box. Make sure all oil is wiped off of the oil immersion objective.
5. Repeat the same steps with remaining slides. When observing bacteria, note the three different
morphologies, or shapes. (See picture above.)
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Lab #1: Care and Use of the Microscope
Laboratory Report
Name ______________________________
Lab Section _____________
Results
Microscope Number: _________
Monocular or binocular: ____________________
Draw a few cells from each slide, and show how they appeared at each magnification. Note the differences in
size at each magnification.
Algae
Slide of _______________________
Total
magnification ____x
_____x
_____x
_____x
_____x
Fungi
Slide of ___________________
Total
magnification ____x
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Protozoa
Slide of __________________
Total
magnification ____x
_____x
_____x
_____x
_____x
Bacteria
Slide of _________________________
Total
magnification ____x
Conclusions
1. What was the largest organism you observed?
2. What was the smallest organism you observed?
3. What three shapes of bacteria did you see?
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4. How does increased magnification affect the field of view?
Questions
1. Why do we want to use a microscope objective that is parfocol?
2. Which objective has the shortest working distance?
3. Name three controls on the microscope that can be used to change the amount of light that is used
when looking at a slide.
4. Let’s say that the diameter of the field of vision in your microscope is 2 mm under low power. If one
bacillus cell is 2 um, how many bacillus cells could fit end to end across the field? How many 10 um
yeast cells could fit across the field?
5. Tell me two different ways we can increase the resolving power of a microscope.
6. What is an advantage of using the low-power objective over the oil-immersion objective for viewing
fungi or algae?
7. What do you think would happen if we used water instead of immersion oil when using the oilimmersion objective?
8
Lab #2: Examination of Living Microorganisms
Objectives:
After completing this experiment, students should be able to:
1. Prepare and observe wet-mount slides and hanging-drop slides.
2. Distinguish between true motility and Brownian movement.
3. Describe the difference between phase-contrast microscopy and brightfield microscopy.
4. Use a phase-contrast microscope to observe living microbes.
INFORMATION
When using a brightfield microscope, objects appear dark against a light or “bright” background. A brightfield
microscope can be used to examine unstained microorganisms and specimens with little color. As both the background
and the specimen have little to no color, very little contrast exists between the two. This makes it difficult to see much
detail of the specimen being observed. A phase-contrast microscope, however, is useful in the examination of living
organisms. Phase-contrast microscopes are able to increase contrast between the microbes you are trying to see and
the background by using special condensers.
In phase-contrast microscopy, small differences in the refractive properties of the objects and the liquid
environment have variations of brightness. In phase-contrast microscopy, a ring of light passes through the object, and
light rays are diffracted and out of phase with the light rays not hitting the object. The phase-contrast microscope
enhances these phase differences so that the eye detects the difference as contrast between the organisms and
background and between structures within a cell. The organisms appear as degrees of brightness against a darker
background. Phase-contrast microscopes make it possible for us to see structural details within living cells.
Studying Living Microbes
Using a wet-mount technique, you will examine different environments to help you become aware of the numbers and
varieties of microorganisms found in nature. The microorganisms will demonstrate either Brownian movement or true
motility. Brownian movement is not true motility, but rather movement caused by the molecules in the liquid striking
an object and causing the object to shake or bounce. In Brownian movement, the particles and microorganisms all
vibrate at the same rate and maintain their positions. Motile microbes move from one position to another; we call this
“purposeful movement”. Their movement appears more directed than Brownian movement, and occasionally the cells
may spin or roll.
Many different microbes, such as protozoa, algae, fungi, and bacteria, can be found in pond water and in
infusions of organic matter. Van Leeuwenhoek made some of his discoveries using a peppercorn infusion similar to the
one you will see in our lab experiment. Direct examination of living microorganisms is useful in determining size, shape,
and movement. A wet mount is a fast way to observe bacteria. Larger microbes and true motility are easier to see in
the greater depth seen with the hanging-drop procedure. A petroleum jelly seal is used to reduce evaporation of the
suspended drop of fluid.
In this lab, you will examine living microbes using both a brightfield and phase-contrast microscope.
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MATERIALS
Disposable glass slides
Disposable coverslips
Depression slides (reusable)
Petroleum jelly
Toothpicks
Disposable pipettes
Alcohol preps
Phase-contrast microscope
Brightfield microscope
CULTURES
Hay infusion (hay and tap water), incubated 1 week in the light
Hay infusion (hay and tap water), incubated 1 week in the dark
Peppercorn infusion (peppercorn and tap water)
18 – 24 hour-old broth culture of Bacillus
PROCEDURE
Wet-Mount Technique
1. Using one of the disposable pipettes, suction up some of one hay infusion (try and get some of the solid material
from around the hay) and transfer a small drop to the center of a glass slide.
2. Carefully place a coverslip on top of the drop of fluid.
3. Place the slide on the stage of your microscope and observe with low power (10x). Adjust the iris diaphragm so
that only a small amount of light is admitted. Concentrate your observations on the larger, more rapidly moving
organisms. At this magnification, bacteria are barely discernible as tiny dots.
4. Next, examine the slide with the 40x objective. Increase the light using the iris diaphragm as needed. Some
microbes are motile, whereas others only exhibit Brownian movement.
5. After recording your observations, examine the slide with the oil immersion lens. Bacteria should now be
magnified sufficiently to be seen.
6. Record your observations, noting the relative size and shape of the organisms.
7. Make a wet mount from the other hay infusion, and observe it, using the low and high power objectives. Record
your observations.
8. As slides are made of glass, dispose of them, along with the coverslips, in the sharps container.
Hanging-Drop Procedure
1. Obtain a clean depression slide.
2. Pick up a small amount of petroleum jelly on a toothpick.
3. Using the technique demonstrated by the instructor, carefully touch the petroleum jelly to all four edges of a
coverslip to get a small rim of petroleum jelly around the entire coverslip. Be careful not to get petroleum jelly
on the other side of the coverslip or smeared on the middle of the coverslip.
4. Place the coverslip on a Kimwipe, with the petroleum jelly-side up.
5. Transfer a small drop of the peppercorn infusion to the coverslip.
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6. Place a depression slide over the drop, scoot the Kimwipe to the edge of the table, and carefully and quickly flip
the slide over so that the drop of fluid is suspended upside down from the coverslip into the space provided by
the depression slide.
7. Examine the drop under low power by locating the edge of the drop and moving the slide so the edge of the
drop crosses the center of the field.
8. Reduce the light with the iris diaphragm and focus. Observe the different sizes, shapes, and types of movement.
9. Switch to high-power and record your observations. Do not attempt to use the oil immersion objective!
10. When finished, clean your slide with an alcohol prep and Kimwipes and return to the appropriate area. Do NOT
throw the depression slides away! Coverslips may still be thrown away in the biohazard container.
11. Using a new coverslip, repeat the procedure with the culture of Bacillus. Record your observations.
12. When completely finished, make sure your microscope has been cleaned and return it to the proper cupboard.
Phase-Contrast Microscopy
1. As a lab group, go in the storeroom and examine the wet prep of the dark hay infusion with the phase-contrast
microscope. The slide has already been prepared and focused by the instructor.
2. Record your group’s observations on your lab report and compare your results for the hay infusion using the
brightfield microscope to the results seen with the phase-contrast microscope.
Algae – Diatoms
Algae – Spirogyra
Algae – Euglena
Algae – Chlamydomonas
Algae – Volvox
Algae – Scenedesmus
Protozoa – Paramecium
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Protozoa – Vorticella
Protozoa – Amoeba
Hanging-drop Procedure
Depression Slide
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Lab #2: Examination of Living Microorganisms
Laboratory Report
Name ___________________________
Lab Section _________
RESULTS
Wet-Mount Technique
Draw the types of protozoa, algae, fungi, and bacteria observed. Indicate their relative sizes and shapes. Record the
magnification.
Sample:
Hay infusion, light
Total magnification:
_____X
Hay infusion, dark
_____X
Compare the size and shape of the organisms observed in the “light” and “dark” hay infusions. Note your observations
here:
Hanging-Drop Procedure
Draw the types of microorganisms seen under high-power magnification. Indicate their relative sizes and shapes.
Sample:
Peppercorn infusion
Total magnification:
______X
Bacillus culture
______X
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Phase-Contrast Microscopy
Carefully draw the organisms and their internal structure.
Sample:
Hay infusion, dark
Total magnification:
_______X
Bacteria
In the following table, record the relative numbers of each bacterial shape observed. Record your data as 4+ (most
abundant), 3+, 2+, +, - (none seen).
Culture
Bacilli (rods)
Shape
Cocci
Spiral
Hay infusion, dark
Hay infusion, light
Peppercorn infusion
Bacillus culture
CONCLUSIONS
1. Did any of the bacteria you observed exhibit true motility?
from Brownian movement?
How would we distinguish true motility
2. Compare the appearance of the microbes observed in the dark hay infusion using the phase-contrast
microscope to those seen with the brightfield microscope.
3. Which infusion had the most overall bacteria?
Explain why you think this is:
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Questions
1. What, if any, practical value do these techniques have? (What are the benefits to utilizing these procedures)
2. Why don’t we use the oil immersion objective when examining a hanging-drop slide?
3. Why are microbes difficult to see in wet preps?
4. Where did the organisms seen in the light and dark hay and peppercorn infusions come from? (hint: I am
looking for more than just “hay” for an answer!)
5. What advantage (if any) does the hanging-drop procedure have over the wet mount procedure?
6. Why do we use petroleum jelly in the hanging-drop procedure?
7. Is it possible to distinguish eukaryotic organisms from prokaryotic organisms? Explain your answer.
8. Why do we use petroleum jelly in the hanging-drop procedure?
9. Is it possible to distinguish eukaryotic organisms from prokaryotic organisms? Explain your answer.
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Lab #3: Microbes in the Environment
Objectives:
After completing this experiment, students should be able to:
1. Explain the use of agar in culture media.
2. Discuss technique required for proper collection of specimens for microbial culture.
3. Compare microbial growth on solid and liquid culture media.
4. Describe colonial morphology using appropriate descriptions.
INTRODUCTION
Microbes are ubiquitous – all around us. They are found in the air we breathe, the water we drink, the dirt outside and
on every surface around us. Microbes even live in and on our bodies. Most of these organisms are harmless. In
microbiology, we have to be careful to avoid contamination of specimens, sterile media and other materials with
microbes.
In this experiment, we will attempt to culture and grow microorganisms from the environment. When we select media
for setting up cultures, we have to think about the nutrients (carbon, nitrogen, phosphorous and sulfur), source of
energy, and other necessary growth factors any microbes present may require. When we are working with a medium
where the exact chemical composition is known, we call this a chemically defined medium.
Glucose-Minimal Salts Broth
Ingredient
Amount/100mL
Glucose
0.5 g
Sodium chloride (NaCl)
0.5 g
Ammonium dihydrogen phosphate (NH4H2PO4)
0.1 g
Dipotassium phosphate (K2HPO4)
0.1 g
Magnesium sulfate (MgSO4)
0.02 g
Distilled water
100 mL
Culture Media
Many chemoheterotrophs can be grown on media for which the precise chemical composition varies slightly between
batches. Organic carbon, energy, and nitrogen sources are oftentimes supplied by protein in the form of meat extracts
and partially digested proteins called peptones. Nutrient broth is a commonly used liquid medium. When agar is added,
it becomes a solid media we call nutrient agar.
Nutrient Agar
Ingredient
Amount/100mL
Peptone
0.5 g
Beef extract
0.3 g
Sodium chloride (NaCl) 0.8 g
Agar
1.5 g
Distilled water
100 mL
Agar comes from marine red algae. It has unique, useful properties that make it perfect for culture media. Few
microbes are able to break-down agar, so it stays solid during microbial growth. It liquefies at 100℃ and remains liquid
until cooled to 40℃. Solidified agar can be placed in the lab incubator, where it will remain solid up to 100℃.
16
Media must be sterilized after being prepared. The most common method of sterilizing culture media that are
heat stable is steam sterilization, also known as autoclaving, which uses steam under pressure. During autoclaving,
materials are placed inside the autoclave and then heated to 121℃ at 15 pounds of pressure (15 psi) for 15 minutes.
Growing Bacteria
There are a number of forms culture media can be prepared in, depending on the needs of your lab. Petri plates contain
solid media and provide a large surface area for the examination of colonies. Microbes will be inoculated, or placed,
onto nutrient agar and into nutrient broth. Once bacteria are inoculated into culture media, they will increase in
number during the incubation period. After a suitable period of incubation, liquid media will become turbid, or cloudy,
which is a sign of bacterial growth. On solid media, colonies can be seen with the naked eye. A colony is a population
of cells that develops from a single bacterial cell. A colony can arise from a group of the same microbes attached to one
another, which is called a colony-forming-unit (CFU). Although many different species of bacteria have colonies that
appear similar, each colony that appears different is typically a different species.
MATERIALS
250 mL Erlenmeyer flask of sterilized melted nutrient agar
Sterile Petri dishes (2)
Tube of sterile nutrient broth
Sterile cotton swabs (2)
Tube of sterile water
Sharpie
PROCEDURE
1. Pouring your Nutrient Agar plates:
a. Set two sterile, unopened Petri dishes in front of you with the cover (the larger half) on top. The smaller
half (the deeper half) is the side the agar will go into, so it should be face-up.
b. Obtain a flask of melted nutrient agar from the instructor. You will need to work quickly so that the
melted agar does not cool. If your agar cools and solidifies, you will need to heat it to liquefy it again.
Make sure that you are ready to pour when you get the melted agar from the instructor.
c. The sterile nutrient agar will need to be poured aseptically – you do not want to introduce microbes
into the medium. Hold the flask at an angle and carefully remove the foil without touching the mouth of
the flask. Keep the flask at an angle the entire time as this will keep microbes in the air from entering
the flask while the foil is off.
d. Remove the cover from the first Petri dish – keep it in your hand; do not set it down on the lab table.
e. Quickly and carefully, pour melted nutrient agar into the dish until the bottom is just covered.
f. Place the lid back on the plate at an angle to vent and swirl gently to evenly distribute the agar over the
inside of the plate. You want a nice smooth surface. Since the agar is hot, condensation will form on
the lid. Venting the plate slightly will keep this from happening.
g. Repeat the same procedure with the second plate.
h. Return any unused melted agar to the instructor.
i. Wait at least 5 minutes before checking your agar to see if it has solidified. How do you know if it is
solid? Hold the bottom against the back of your hand – does it feel warm yet? Try jiggling the plate – if
it still seems liquid-like, it is not completely solid.
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j.
When you are sure your plates have solidified, place the lids on completely and tip your plates over so
that the agar side is facing up.
2. Culturing microbes from the environment:
a. Your lab group will be culturing one environmental surface and one body site. Decide as a group what
you will culture.
i. Some ideas: environmental – money; cell phone; laptop; backpack; car keys; water fountain;
restroom; body site – mouth; underarm; between fingers/toes; eyelashes; hair
b. Label your plates appropriately. Necessary information: date; group name; site cultured. Plates should
be labeled on the media side.
c. If you are culturing a site that is NOT wet already, you will need to dip your sterile swab in sterile water
before culturing the area. You do not want your swab dripping wet, just slightly damp to help microbes
adhere to the surface. Scrub the site thoroughly with all sides of the swab. Be careful not to
contaminate your swab with other extraneous areas (lab bench-top, your hand, etc.)
d. If you are culturing a site that is wet, you do not need to wet your swab with sterile water. Scrub the
site thoroughly with all sides of the swab.
e. You will then take your swab and inoculate the surface of the agar plate. To inoculate, gently roll the
swab across the agar surface. Be careful not to gouge into the agar. Don’t leave the lid off of the plate
any longer than necessary to prevent contamination. Repeat this procedure with the second plate.
f. After inoculating the agar plate for your environmental surface, place the swab into the nutrient broth
tube. If the handle on the swab is too long for the tube, it is acceptable to break off part of the handle
to fit the swab into the tube.
g. Inoculated agar plates from body sites will go into the 37℃ incubator. Inoculated agar plates and
nutrient broth tubes from environmental surfaces will be incubated at 20℃. Agar plates are always
incubated with the media side (the side that you labeled) facing up.
h. Incubate all inoculated media at least 48 hours.
PROCEDURE – RETURN TO CHECK
1. Observe and describe the growth seen on the plates. Notice each different-appearing colony, and describe the
colonial characteristics – size, pigment and morphology using the characteristics described in the pictures below.
Determine the approximate number of each type of colony. When many colonies are present, write TNTC (too
numerous to count) as the number of colonies seen.
2. Describe the appearance of the nutrient broth labeled “unsterilized”, a nutrient broth tube that is sterilized and
not inoculated, and the broth tube your group inoculated.
Look for clumps of microbial cells, called flocculent. Is there a membrane, or pellicle, across the surface of the
broth? See whether microbial cells have settled on the bottom of the tube, forming a sediment.
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Colony Descriptions
Example of Colonies on Nutrient Agar
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Lab #3: Microbes in the Environment
Laboratory Report
Name _____________________________
Lab Section __________________
RESULTS
Fill in the following table with the descriptions of the bacterial colonies on your plates. Use a different row for each
different-looking colony.
Colony Description
Area sampled:
___________
Diameter
Form
Margin
Elevation
Pigment
Number of
This Type
Pigment
Number of
This Type
___________
Incubated at
____℃ for
____ days
Colony Description
Area sampled:
___________
Diameter
Form
Margin
Elevation
___________
Incubated at
____℃ for
____ days
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Nutrient Broth
Site: _____________________________ Incubated at: ______℃ for: ______ days
Tube of
Tube of Sterilized Broth
Unsterilized Broth (not
(not inoculated)
inoculated)
Turbidity (yes/no)
Your Group’s Inoculated
Broth
Flocculent (y/n)
Sediment (y/n)
Pellicle (y/n)
Pigment (if any)
CONCLUSIONS
1. What is the minimum number of different bacteria types present on one of your culture plates?
How can you tell?
2. Compare the results of the sterilized and unsterilized broths that have not been inoculated. Explain what you
see and why they appear this way.
3. Which environment has the largest number of bacteria overall?
Explain why one area had more bacteria than the other.
4. Which environment has the most different types of bacteria?
Explain the differences in bacteria from these two areas.
5. Which culture technique gives us the most information, agar or broth?
Explain your answer
QUESTIONS
1. What are the bacteria using for nutrients in nutrient agar?
2. What purpose does agar serve in solid media?
3. Why are Petri plates useful in microbiology?
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4. Do you think all the microorganisms living in or on the environments we sampled grew on the nutrient agar
plates? Explain why you think this is.
5. Why do we use agar instead of gelatin to solidify culture media?
6. How could you figure out if the turbidity in your group’s nutrient broth tube is caused by a mixture of different
microbes or from the growth of only one kind of microbe?
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Lab #4: GRAM STAINING
OBJECTIVES
After completing this experiment, students should be able to:
1. Explain the rationale and procedure for the Gram stain.
2. Perform and interpret Gram stains.
The Gram stain is a differential stain which distinguishes bacteria based on properties of their cell wall. Bacterial cell
walls are composed primarily of peptidoglycan and bacteria can be classified into two main groups depending on the
amount of peptidoglycan in their cell wall. Gram-positive organisms have a thick layer of peptidoglycan, whereas Gramnegative organisms have a thin layer of peptidoglycan, plus an additional outer membrane that is absent in Grampositive organisms.
In the first step of the Gram stain procedure, crystal violet, also known as the primary stain, is added to the smear. All
cells take up the purple crystal violet stain. Following the primary stain, Gram’s Iodine is applied to the smear. The iodine
acts as a mordant by binding with crystal violet, creating CV-I, a much larger molecule within the cell wall. This CV-I
complex is now locked within the peptidoglycan. The decolorizing agent used in the gram staining procedure is 95%
ethanol, which is a lipid solvent that dissolves the Gram negative outer membrane. Without the outer membrane, the
CV-I washes out of the thin peptidoglycan of Gram-negative cells. In Gram-positive cells, the decolorizer dehydrates the
protein of the cell wall, trapping the CV-I within. Safranin, the counter stain, is then added. Safranin stains the
decolorized Gram-negative cells pink. At the end of the Gram staining procedure, Gram-positive cells are purple and
Gram-negative cells are pink/red.
It is important to note that Gram stains should only be performed on young cultures of bacteria (less than 24 hours old).
When bacteria die, their cell wall will degrade and have a difficult time holding onto the crystal violet, which can cause
inaccurate results. Older cultures may stain “Gram-variable”(a mix of purple and pink).
PREPARATION OF BACTERIAL SPECIMENS
When staining bacterial specimens using the Gram stain procedure, the bacteria must be placed on a glass slide. We call
this a bacterial smear. A smear is a small volume (oftentimes a loopful) of specimen either suspended in a broth or
taken directly from a plate that is spread (smeared) onto a microscope slide. When bacteria growing on a plate is used,
sterile water must be added to the slide to insure that the smear is not too thick. Practice and care are required when
preparing a smear that is the correct thickness. If smears are too thick, you will have trouble seeing individual cells and
23
there are oftentimes problems with staining. If smears are too thin, you may not find the organism. If you stir the drop
of medium too much as you spread it on the slide, you will disrupt cell arrangements such as chains or clusters.
Once the smear has been properly prepared, it must be allowed to air dry. When the smear is dry, it will need to be
fixed. There are two methods of fixing: heat fixing and chemical fixing. Heat fixation does three things: (1) it kills the
organisms; (2) it causes the organisms to adhere to the slide; and (3) it alters the organisms so that they more readily
accept stains (dyes). Timing with heat-fixing is important as well. If you heat-fix too little, the organism will wash off the
slide. Over heat-fixing of the smear can cause alterations to the bacterial cells and they won’t stain or look as they
should. Chemical fixing requires dipping an air-dried smear into methanol for a few seconds. Allow the methanol to
evaporate and proceed with staining.
MATERIALS
Gram-staining reagents:
Primary Stain
Mordant
Decolorizer
Counter Stain
Crystal Violet
Gram’s Iodine
95% Ethanol
Safranin
CULTURES NEEDED
Nutrient broth tubes or plates of the following:
Escherichia coli
Staphylococcus epidermidis
Bacillus subtilis
PROCEDURE:
1. Prepare your smear. Using a clean slide, make a large circle on the slide with a china marker/crayon. Label the
slide with the initials of your group’s assigned culture.
a. Using a broth culture:
i. Gently agitate your culture broth tube to disperse the bacteria.
ii. Sterilize your inoculating loop using an incinerator and let it cool for 20-30 seconds.
iii. Place loop in the bacterial broth and put the loopful of the broth onto the glass slide. Rub the
drop into a nickel-sized smear. Sterilize the loop again to kill any remaining bacteria. Let the
smear air dry completely. Do not use heat to dry your smear!
b. Using an agar plate:
i. Place a small drop of sterile water in the center of the slide. Do not use too much water or it
will take a long time for your smear to dry.
ii. Sterilize your inoculating loop using an incinerator and let cool for 20-30 seconds.
iii. Use the sterile loop to pick up a small amount of bacterial growth from the surface of the plate.
Do not dig into the agar. Put the loopful of bacteria into the drop of water on the glass slide and
swirl, spreading the liquid out. Stop swirling when the water becomes slightly cloudy. If your
smear is too thick, it will not stain correctly!
iv. Sterilize the loop again to kill any remaining bacteria. Let the smear air dry completely. Do not
use heat to dry your smear!
2. Fix the slide.
a. Using methanol:
i. Dip your air-dried smear into a beaker of methanol and immediately remove it. Allow to air dry
before staining.
b. Using heat-fixing:
i. Hold the back-side of the slide directly in front of the incinerator for 8 seconds. Be careful not to
touch the front of the incinerator. Place the slide against your hand. If the slide is warm, you
may proceed with staining. If the slide is not warm, repeat heat-fixing.
24
3. Stain your smear. Use a slide rack to hold your slide.
a. Cover the smear with crystal violet and let it sit for 1 minute.
b. Rinse the slide carefully with cold water. Do not turn the water on too high as it will wash the smear off
your slide.
c. Cover the smear with iodine and let it sit for 1 minute.
d. Repeat rinse step.
e. Pick up slide off of staining rack and tilt downward over the sink.
f. Squirt decolorizer onto the slide, directly ABOVE your smear. The decolorizer should run off of the slide.
You should see purple running off of your slide. Stop decolorizing when there is no longer any excess
purple running off. (5 – 10 seconds)
g. IMMEDIATELY rinse smear with water
h. Place smear back onto staining rack
i. Cover the smear with safranin and let it sit for 30 seconds.
j. Rinse the slide once last time with water
4. Shake excess water off of your slide and carefully blot between two layers of Kimwipes.
5. Examine the stained slide microscopically, using the low, high and oil immersion objectives. When you are done
with your slide, dispose of it in the sharps container.
Common Sources of Gram-Staining Errors
1. The loop is too hot.
2. Smears were over heat-fixed.
3. The decolorizer was left on the smear too long.
4. The culture was too old.
5. The smear was too thick.
25
Lab #4: Gram Staining
Laboratory Report
Name ___________________________
Lab Section _________
RESULTS
Draw a few bacteria seen using oil-immersion. Look at your classmate’s slides of the other two bacterium.
Bacteria:
Staphylococcus epidermidis
Escherichia coli
______X
______X
_____________________
____________________
___________________
_____________________
____________________
___________________
____________________
____________________
Total Magnification:
Morphology:
______X
Bacillus subtilis
(cocci, bacilli, spiral)
Arrangement:
(clusters, chains, pairs, tetrads, singles)
Color:
_____________________
CONCLUSIONS
1. Which bacteria were Gram-positive?
Which bacteria were Gram-negative?
2. Which bacteria is the largest?
Which bacteria is the smallest?
3. Do your results agree with the information found in your textbook/online?
If not, why?
QUESTIONS
1. What would happen if you tried to Gram stain human cells? (what would they stain and why)
26
2. You are looking at a Gram-stained field of purple cocci and red bacilli through the microscope. What do you
think is causing this?
3. Why do Gram-positive cells more than 24 hours old stain Gram-negative?
4. It is impossible to identify bacteria from a Gram stain alone. Why then, would a doctor request a Gram stain on
a culture before beginning antibiotics?
5. You are performing a Gram stain on a slide made from a pure culture of bacteria and see that there are both red
and purple cocci present. Adjacent cells are not always the same color. What is causing this?
6. Can you add iodine before crystal violet when preparing a Gram stain?
7. List the steps of the Gram-staining procedure in order (omitting washings), and fill in the color of Gram-positive
and Gram-negative cells after each step.
Appearance
Step
Chemical
Gram-Positive Cells
Gram-Negative Cells
1
2
3
4
Which step could be left out without affecting your ability to determine the Gram-stain reaction?
27
Lab #5: Effectiveness of Hand Scrubbing
Objectives:
After completing this experiment, students should be able to:
1. Evaluate the effectiveness of handwashing and a surgical scrub.
2. Explain the importance of aseptic technique in the hospital environment.
BACKGROUND
The skin is sterile during fetal development. After birth, a baby’s skin becomes colonized by many different bacteria for
the rest of its life. As a person ages and moves from one environment to another, the microbial population will change
to match the new environment’s conditions. Normal microbiota are microbes that are considered permanent.
Transient microbiota are microorganisms that are present only for days or weeks.
Importance of Handwashing
The discovery of the importance of handwashing in preventing disease is credited to Ignaz Semmelweis at Vienna
General Hospital in 1846. He recognized that the lack of aseptic technique was directly related to the incidence of
puerperal fever and other diseases. Medical students would go to the patient’s bedside directly from the autopsy room
to assist in child delivery without washing their hands. Less puerperal sepsis occurred in patients treated by midwives,
who did not touch cadavers. Semmelweis ordered the medical students to wash their hands with a chloride lime
solution, a policy that caused the death rate due to puerperal sepsis to drop from 12% to 1.2% in one year. Guidelines
from the Centers for Disease Control and Prevention (CDC) state that “handwashing is the single most important
procedure for preventing nosocomial infections,” yet recent studies in hospitals show handwashing rates as low as 31%.
Clinical Handwashing
The structure of the skin and a layer of oil prevent handwashing from removing all bacteria. Soap helps remove oil,
however, and scrubbing will help increase the removal of bacteria. Hospital personnel are required to wash their hands
before attending patients, and to perform a complete surgical scrub – removing the transient as well as many of the
resident microbiota – before surgery. Usually, a 10 to 15 minutes of scrubbing with soap will remove transient
microbiota. The surgeon’s skin is never sterilized. Only burning or scraping it off would achieve this.
In this experiment, we will check the effectiveness of washing skin with soap and water. Only organisms capable
of growing aerobically on nutrient agar will be observed. Because organisms with different nutritional and
environmental requirements will not grow, this procedure will involve only a small number of the skin microbiota.
MATERIALS
Petri plates containing nutrient agar (2)
Scrub brush
Bar soap or liquid soap
Waterless hand sanitizer
PROCEDURE – SET UP
1. You will need two nutrient agar plates.
28
2.
3.
4.
5.
6.
7.
8.
a. Divide one nutrient agar plate into four quadrants. Label the sections 1 through 4. Label the plate
“water”. (Don’t forget to put the date, your section, and your group’s name on your plate.)
b. Divide the other nutrient agar plate into five sections. Label the sections 1 through 5. Label the plate
“bar soap” or “liquid soap” depending on which your group is using in the experiment.
Use the “water” plate first. Using 1 finger (use the same finger throughout the experiment), touch section 1
with your finger (without wetting). Wash well WITHOUT soap, shake off the excess water, and, while your hands
are still wet, touch section 2. Do not dry your fingers with a paper towel. Wash again, and, while your hands are
still wet, touch section 3. Wash a final time, and touch section 4.
Use your same hand on the plate labeled “soap.” Wash well with soap, rinse, shake off the excess water, and
then touch section 1.
Wash again with soap, rinse, shake off the excess water, and then touch section 2.
Using a brush and soap, scrub your hand for 2 minutes, rinse, and shake off the excess water; then touch section
3.
Repeat the soap-and-brush scrub for 4 minutes, rinse, and shake off the excess water; then touch section 4.
Use the waterless hand sanitizer as you normally would, then touch section 5.
Incubate the plates, inverted, at 35 C for 24 hours.
PROCEDURE – RETURN TO CHECK
1. Record the growth as:
 “-“
- no growth
 “+”
- minimal growth
 “2+” - moderate growth
 “3+” - heavy growth
 “4+” - maximum growth
29
Lab #5: Effectiveness of Hand Scrubbing
Laboratory Report
Name ___________________________
Lab Section _________
Hypothesis
1. What do you expect to see when you check back on your plates.
a. Will you see a difference between washing with water and washing with soap? (Explain)
b. What do you think will happen as you repeatedly wash with soap? Will you get more or less bacteria
each time you wash? (Explain)
RESULTS
Water Alone Plate
Indicate the relative amounts of growth in each
quadrant.
Type your group tested: ___________________
Indicate the relative amounts of growth in each
quadrant.
Section
Section
1 – no washing
Amount of Growth
Amount of Growth
1 – wash with soap and
water
2 – first wash with water
2 – wash with soap and
water
3 – second wash with water
3 – wash with soap, water
and scrub brush (2 minutes)
4 – third wash with water
4 – wash with soap, water
and scrub brush (4 minutes)
5 – hand sanitizer
Soap Plate
CONCLUSIONS
1. Did your results differ from your expected results? Briefly explain why or why not.
30
2. Using your classmates’ data, compare the results seen with bar soap and liquid soap.
QUESTIONS
1. If most of the normal and transient microbes are not harmful to us, why do surgeons need to scrub their hands
before surgery?
2. What is the difference between transient microbiota and normal microbiota?
3. What results would a surgeon expect to see after scrubbing for 10 minutes with a brush and then using an
antiseptic?
4. The following data were collected from soaps after 1 week of use at a hospital nurses’ handwashing station.
Neither bacteria nor fungi were isolated from any of the products before use.
Aerobic bacteria were isolated from 25 soap products. Data are expressed as percentage of soap products
contaminated.
Liquid Soap: Type of Closure
Organisms
Bar Soap
Screw Top
Slit / Flip
Flip/Pump
Pump
Total bacteria
95%
71%
39%
10%
0%
Gram-positive cocci
95%
71%
39%
10%
0%
Gram-negative rods
12%
1%
1%
1%
0%
What conclusions can you draw from this data? (Explain the reason for the differences we see between the
types of soap used)
31
Lab #6: Epidemiology
Objectives:
1. Define the following terms: epidemiology, epidemic, reservoir, and carrier.
2. Describe three methods of transmission.
3. Define index case and case definition.
4. Attempt to determine the source of a simulated epidemic.
Background
In every infectious disease, the disease-producing microorganisms, the pathogen, must come in contact with the host,
the organism that harbors the pathogen. Communicable diseases can be spread either directly or indirectly from one
host to another. Some microorganisms cause disease only if the body is weakened or if a predisposing event such as a
wound allows them to enter the body. Such diseases are called noncommunicable diseases – that is, they cannot be
transmitted from one host to another. The science that deals with when and where diseases occur and how they are
transmitted in the human population is called epidemiology. Endemic diseases such as pneumonia are consistently
present in the population. When many people in a given area acquire the disease in a relatively short period of time, it
is referred to as an epidemic disease. The first case reported in a patient in a disease outbreak is the index case. One of
the first steps in analyzing a disease outbreak is to make a case definition, which should include the typical symptoms of
patients so that you know who should be included.
Disease Transmission
Diseases can be transmitted by direct contact between hosts. Droplet infection, which occurs when microorganisms are
carried on liquid drops from a cough or sneeze, is a method of direct contact. Diseases can also be transmitted by
contact with contaminated inanimate objects, or fomites. Drinking glasses, bedding, and towels are examples of
fomites that can be contaminated with pathogens from feces, sputum, or pus.
Some diseases are transmitted from one host to another by vectors. Vectors are insects and other arthropods that carry
pathogens. In mechanical transmission, insects carry a pathogen on their feet and may transfer the pathogen to a
person’s food. For example, houseflies may transmit typhoid bacteria from the feces of an infected person to food.
Transmission of a disease by an arthropod’s bite is called biological transmission. An arthropod ingests a pathogen
while biting an infected host. The pathogen can multiply or mature in the arthropod and then be transferred to a
healthy person in the arthropod’s feces or saliva.
The continual source of an infection is called the reservoir. Humans who harbor pathogens but who do not exhibit any
signs of disease are called carriers.
Tracking a Disease
An epidemiologist compiles data on the incidence of a disease and its method of transmission and tries to locate the
source of infection to decrease to decrease the incidence. The time course of an epidemic is shown by graphing the
number of cases and their date of onset. This epidemic curve gives a visual display of the outbreak’s magnitude and
time trend. An epidemic curve provides a great deal of information. First, you will usually be able to tell where you are
in the course of the epidemic and possibly be able to project its future course. Second, if you have identified the disease
and know its usual incubation period, you may be able to estimate a probably time period of exposure and can then
develop a questionnaire focusing on that time period.
32
By graphing the cases, an epidemiologist may be able to locate the index case. With more information from
patients, you may be able to determine whether the outbreak has resulted from a common-source exposure, from
person-to-person spread, or both. The latter occurs, for example, when someone gets an infection from a food and then
transmits the infection to family members who did not eat that food. The infected family members are called secondary
cases.
In this exercise, an epidemic will be simulated. Although you will be in the “epidemic”, you will be the
epidemiologist who, by deductive reasoning and with luck, determines the source of the epidemic.
Materials
Tube of clear liquid (instructor will assign)
Disposable pipette
Gloves
Pen
Indicator
Procedure
1. Obtain your assigned tube of liquid and a disposable pipette from the instructor.
2. Using the disposable pipette, suction up half of the liquid in your tube and trade with another student. (You will
put half of your tube in another student’s tube and they will put half of the liquid in their tube into yours). Make
sure that you only trade half of your liquid. You will need to keep track of all the people you trade with and in
what order you traded. Write down the number of the person you traded with on your glove. This trade is
simulating “exposure” to that person.
3. Find another student and trade half of your liquid with them. Make sure that you move around the room when
you trade liquid.
4. Complete the trade as many times as the instructor tells you to (as this depends on the class size).
5. Record all of your trades in the correct order on your lab report.
6. When everyone has completed all of their “exposures”, the instructor will add the indicator to each student’s
tube. Students whom are “infected” will have pink liquid, whereas students not “infected” will have clear or
white/milky looking liquid.
7. Using the group’s data, deductively try to determine who was the index case for the simulated epidemic.
33
Lab#6: Epidemiology
Lab Report
Name _______________________________
Lab Section _________
Results
1. Your tube # is ________. What was in the “infected” tubes? _____________________
2. What is the case definition for our classroom “epidemic”? ____________________________________________
___________________________________________________________________________________________
Student’s Name
Tube Number
Were they infected?
First Trade
Second Trade
Third Trade
Fourth Trade
Conclusions
1. Were you able to figure out who was the index case?
2. What number was the “infected” tube?
If so, who was it?
Explain how you figured this out.
3. Could you have been the “infected” person and not have the liquid in your tube turn pink?
Questions
1. In general, do all people who contact an infected person acquire the disease? Explain your answer.
2. How can an epidemic stop without medical intervention (e.g. quarantine, chemotherapy, vaccination, etc.)?
34
Assume that you work in the Infectious Disease Branch of the CDC. You are notified of the following two incidents. (1)
On June 20, cruise ship X reported that 84 of 2,318 passengers reported to the infirmary with norovirus gastroenteritis
during a 7-day vacation cruise. According to federal regulations, when the incidence of acute gastroenteritis among
passengers and crew exceeds 3%, an outbreak is defined and requires a formal investigation. Is this an outbreak?
_____________ (2) A nursing home reported 125 cases of norovirus gastroenteritis among residents and staff during
the week of June 23rd.
Data collected from the cruise ship and nursing home are shown in the following table. Use this data to answer
the questions.
Cruise Ship Data
Nursing Home Data
Date
Number of Cases Date Number of Cases
Cruise 1 6/9
2
6/23 1
6/10
4
6/24 8
6/11
5
6/25 12
6/12
3
6/26 12
6/13
3
6/27 50
6/14
2
6/28 32
6/15
1
6/29 10
6/30 8
Cruise 2 6/16
2
6/17
10
6/18
13
6/19
13
6/20
84
6/21
46
6/22
20
Cruise 3 6/23
10
6/24
23
6/25
39
6/26
41
6/27
18
6/28
17
6/29
9
1. How is norovirus transmitted (you will probably have to look this up).
2. What happened on cruse 2? Explain how you came to this conclusion.
3. What would you do before cruise 4?
4. Three nursing home residents were passengers on cruise 2. What can you conclude?
35
A health department received a report from hospital A that 15 patients had been admitted on October 12 with
unexplained pneumonia. On October 21, hospital B, located 15 miles from hospital A, reported a higher-than-normal
pneumonia census for the first 2 weeks of October. Legionella pneumophila was eventually identified in 23 patients; 21
were hospitalized, and 2 died. To identify potential exposures associated with L. pneumophila, a questionnaire was
developed, and a case-control study was initiated on November 2 to identify the source of infection. Three healthy
controls were selected for each confirmed case; controls were matched by age, gender, and underlying medical
conditions. Of the 15 case patients for whom a history was available, 14 had visited a large home improvement center 2
weeks before onset of illness. Results of the questionnaire are shown below.
Number of patients
Visited home improvement center
Average time at center (min)
Looked at whirlpool spa X
Looked at whirlpool spa Y
Visited greenhouse sprinkler system display
Visited decorative fish pond
Used drinking fountain
Used urinals
Used restroom hot water
Used restroom cold water
Case Patients
15
14
79
13
13
10
14
13
10
6
8
Healthy Controls
45
12
29
9
1
10
12
10
4
4
8
1. Where is Legionella pneumophila found in the environment (hint: what do all of these sites that were tested
have in common?)
2. What is the most likely source of this outbreak of legionellosis?
3. How would you prove this was the source?
4. How do you think this disease was probably transmitted (hint: Legionella cannot be transmitted from person to
person)?
5. Provide an explanation for the infected patient who did not go to the home improvement center. (How did they
become infected with Legionella if they didn’t go to the home improvement center?)
36
Lab #7: Agglutination Reactions: Slide Agglutination
Objectives:
1. Compare and contrast the terms agglutination and hemagglutination.
2. Use agglutination to identify a pathogenic bacterium.
3. Determine ABO and Rh blood types.
4. Determine possible compatible transfusions.
Background
The surfaces of bacterial cells contain antigens that can be used in agglutination reactions. Agglutination reactions
occur between particulate antigens – such as cell walls, flagella, or capsules bound to cells – and antibodies.
Agglutination, or the clumping of bacteria by antibodies, is a useful laboratory diagnostic technique. When the cells
involved are red blood cells, the reaction is called hemagglutination.
Agglutination of Bacteria
In a single agglutination test, an unknown bacterium is suspended in a saline solution on a slide and mixed with a drop of
known antiserum. This test is done with different antisera on separate slides. The bacteria will agglutinate when mixed
with antibodies produced against the same species and strain. A positive test can identify the bacterium. In some
diagnostic tests, the antiserum is coupled to latex particles to enhance the visibility of the positive agglutination test.
Hemagglutination
Hemagglutination reactions are used in the typing of blood. The presence or absence of two very similar carbohydrate
antigens (designated A and B) located on the surface of red blood cells is determined using specific antisera.
Hemagglutination occurs when anti-A antiserum is mixed with type A red blood cells. When anti-A antiserum is mixed
with type B red blood cells, no hemagglutination occurs. People with type AB blood possess both A and B antigens on
their red blood cells, and those with type O blood lack A and B antigens.
Many other blood antigen series exist on human red blood cells. Another surface antigen on red blood cells is
designated the Rh factor. The Rh factor is a complex of many antigens. The Rh factor that is used routinely in blood
typing is the Rh0 antigen, or D antigen. Individuals are Rh-positive when D antigen is present. The presence of the Rh
factor is determined by a hemagglutination reaction between anti-D anti-serum and red blood cells with D antigen on
their surfaces.
A person possesses antibodies to the alternate A-B antigen. Thus, people of blood type A will have antibodies to
the B antigen on their sera. Rh-negative individuals do not naturally have anti-D antibodies in their sera. Anti-D
antibodies are produced when red blood cells with D antigen are introduced into Rh-negative individuals.
The ABO and Rh systems place restrictions on how blood may be transfused from one person to another. An
incompatible transfusion results when antigens of the donor red blood cells react with the antibodies in the recipient’s
serum or induce the formation of antibodies.
A summary of the major characteristics of the ABO and Rh blood groups is presented in the table on the next
page. Blood group antigens are genetically determined, and thus they are inherited. Differences in percentages of
blood groups among groups of people are created by geographical isolation during the development of populations.
37
Characteristics
Antigen present on the red blood
cells
Antibody normally present in the
serum
Serum causes agglutination of red
blood cells of these types
THE ABO AND RH BLOOD GROUP SYSTEMS
Blood Group
A
B
AB
O
A
B
Both A and Neither A
B
nor B
Anti-B
Anti-A
Neither
Both anti-A
anti-A nor
and anti-B
anti-B
B, AB
A, AB
None
A, B, AB
Rh+
D
RhNo D
No anti-D
No anti-D
Neither
Rh+ nor
Rh85
Neither
Rh+ nor
Rh- **
15
Percent occurrence in a mixed
41
10
4
45
white population
Percent occurrence in a mixed
27
20
7
46
90
10
black population
Percent occurrence in a mixed
28
27
5
40
98
2
Asian population
**Anti-D antibodies are not naturally present in the serum of Rh- people. Anti-D antibodies can be produced upon
exposure to the D antigen through blood transfusions or pregnancy.
Source: Adapted from G.J. Tortora, B.R. Funker, and C.L. Case. Microbiology: An Introduction, 10th ed. San Francisco, CA:
Benjamin Cummings.
38
MATERIALS
BACTERIAL AGGLUTINATION
Staphaccol Coagulase test
Toothpicks
HEMAGGLUTINATION
Sterile gauze
Alcohol preps
Sterile lancet
Sharps and biohazard containers
Anti-A, Anti-B and Anti-D antisera
Glass slides (2)
Wax pencil
Toothpicks
CULTURES
Unknown bacterium A
Unknown bacterium B
PROCEDURE
**You can either perform your own fingerstick or you can have the instructor perform the fingerstick on you.
Hemagglutination
1. With a wax pencil, draw two circles on a clean glass slide, and label one “A” and the other “B”. Draw a circle on
the second slide, and label it “D”.
2. Disinfect a finger with the alcohol prep (use either your ring or middle finger. Wipe the alcohol if with sterile
gauze or allow it to dry. Perform the fingerstick using a sterile lancet. Dispose of the lancet immediately after
use into a sharps container.
3. Wipe off the first drop of blood with sterile gauze and proceed to add one drop of blood to each circle on your
slides (3 drops for 3 circles). Touch the slide to the drop of blood, try not to touch your finger to the slide. Apply
pressure to the site of the fingerstick with sterile gauze until it has stopped bleeding (but keep working, or the
blood will clot on your slides).
4. To the A circle, add one drop of anti-A antiserum. Add one drop of anti-B antiserum to the B circle and one drop
of anti-D antiserum to the D circle. Be careful not to touch the end of the dropper to the drop of blood (this will
cause contamination and ruin the antisera!)
5. Mix each suspension with toothpicks. Use a different toothpick/end of the toothpick for each antiserum to
prevent cross-contamination.
6. Observe for agglutination, and determine your blood type. Have the instructor check to make sure your
interpretation is correct.
**Discard the slides, and lancet in the sharps container. Toothpicks and anything else that has blood that is not sharp
can go in the regular biohazard garbage. Anything uncontaminated can go in the regular garbage.
39
Bacterial Agglutination
1. Cut two circles from the provided card on which to perform the test.
2. Using a toothpick, add a few colonies of bacteria from each plate to individual circles (each bacteria gets its own
circle.) Be careful not to pick up any agar as this can interfere with test results.
3. Add one drop of well-mixed Staphaccol latex reagent to each circle, next to the bacteria (don’t add it directly to
the bacteria).
4. Using a new toothpick, mix the reagent and bacteria together and watch for agglutination. Agglutination will
appear as red “chunks” or “flecks” against a blue or clear background. This should be interpreted as a positive
result. If the suspension remains smooth, the test is negative.
5. Discard toothpicks and card in biohazard container.
40
Lab #7: Agglutination Reactions – Slide Agglutination
Lab Report
Name ____________________________
Lab Section _________
Results
Bacterial Agglutination
Bacteria
Unknown A
Description of the Results
Agglutination?
Unknown B
Hemagglutination
Antiserum
Anti-A
Hemagglutination (+ or 0)
Anti-B
Anti-D
Conclusions
1. Which unknown bacterial sample was Staphylococcus aureus?
2. What is your blood type?
QUESTIONS
1. How do agglutination tests detecting microbes in a patient’s blood or fluids differ from techniques used to
detect antibodies to microbes in a patient’s serum?
2. How would agglutination reactions be used to locate the source of an epidemic?
3. What is hemolytic disease of the newborn? How does RhoGAM prevent it?
41
4. Individuals may have antibodies to the A and B blood antigens not found on their red blood cells. What is the
origin of these antibodies in someone who has never had a transfusion?
5. A woman with type A blood can have a healthy baby with type B blood. Why doesn’t the baby develop
hemolytic disease of the newborn?
6. A woman with type A, Rh-negative blood can have a healthy baby with type B, Rh-positive blood and usually will
not develop antibodies to the Rh antigen. Why?
42
Lab #8: Isolation of Bacteria by Dilution Technique
Objectives:
1. Isolate bacteria by using the streak plate and pour plate methods.
2. Prepare and maintain a pure culture.
BACKGROUND
In nature, most microorganisms are found growing in environments that contain many different organisms.
Unfortunately, mixed cultures are of little use in study microbes due to the difficulty they present in figuring out which
organism is responsible for any observed activities. To study growth characteristics, pathogenicity, metabolism,
antibiotic susceptibility, or other characteristics requires a pure culture, one that contains a single kind of
microorganism. Because bacteria are too small to separate directly without sophisticated micromanipulation
equipment, indirect methods of separation must be used.
In the 1870’s, Joseph Lister attempted to obtain pure cultures by performing serial dilutions until each of his
containers theoretically contained one bacterium. However, success was very limited, and contamination, the presence
of unwanted microorganisms, was common. In 1880, Robert Koch prepared solid media, after which microbiologists
could separate bacteria by dilution and trap them on the solid media. An isolated bacterium grows into a visible colony
that consists of one kind of bacterium.
Isolating Bacteria
Currently, three dilution methods are most often used for isolating bacteria: the streak plate, the spread plate, and the
pour plate. In the streak plate technique, a loop is used to streak the mixed sample many times over the surface of a
solid culture medium in a Petri plate. Theoretically, the process of streaking the loop repeatedly over the agar surface
causes the bacteria to fall off the loop one by one and ultimately to be distributed over the agar surface, where each cell
develops into a colony. The streak plate is the most common isolation technique used today.
Counting Bacteria
The spread plate and pour plate are quantitative techniques to determine the number of bacteria in a sample. The
number of bacteria in a sample can be determined by counting the number of colony forming units. The sample must
be diluted, because even 1 milliliter of milk or water could yield 20,000 colonies – too many to count. A series of
dilutions is made and cultured because you don’t know the number of bacteria in the sample. In the spread plate
technique, a small amount of previously diluted specimen is spread over the surface of a solid medium using a spreading
rod.
In the pour plate technique, a small amount of diluted sample is mixed with melted agar and poured in empty,
sterile Petri dishes. After incubation, bacterial growth is visible as colonies in and on the agar of a pour plate. To
determine the number of bacteria in the original sample, a plate with between 25 and 250 colonies is selected. Fewer
than 25 colonies is inaccurate because a single contaminant causes at least a 4% error. A plate with greater than 250
colonies is difficult to count.
The number of bacteria in the original sample is calculated using the following equation:
Colony-forming units =
Number of colonies
per mL
Dilution* x Amount plated
43
*In this exercise, 1 mL of sample is put into each plate. Dilution refers to the dilution of the sample. For example, if 37 colonies were present on the 1:8000 plate, the
calculation would be as follows:
Colony-forming units per mL =
37
= 37 x 8000 = 296,000 = 2.96 x 105
1:8000 x 1
MATERIALS
FOR CLASS
Petri plate containing nutrient agar
Tubes containing melted nutrient agar (3)
Sterile Petri dishes (3)
250-mL beaker
Sterile 1-mL disposable pipettes (3)
Incinerator
Inoculating loop
RETURN TO CHECK (1st time)
Petri plate containing nutrient agar
CULTURES
Mixed broth culture of bacteria
TECHNIQUES REQUIRED
Brightfield microscopy
Aseptic technique
Pipetting
Serial dilution technique
PROCEDURE – FOR CLASS
Streak Plate
1.
2.
3.
Label the bottom of one nutrient agar plate with your specimen type (“mixed culture”), the date, your group’s name and
lab section.
Sterilize the inoculating loop to redness, allow it to cool, and aseptically obtain a loopful of the broth culture.
With your plate laying lid down on the lab bench, pick up the agar side of the plate in your non-dominant hand (the loop
should be in your dominant hand).
a. Streak the first sector by gliding over the surface of the agar in a zig-zag pattern, making about 3 – 4 streaks. Each
“zig-zag” should not touch the one before, but try and get them as close together as possible. Be careful not to
gouge the agar. Hold the loop as you would hold a pencil or paintbrush, and gently touch the surface of the agar.
b. Set the plate back down in the lid (to keep it from becoming contaminated) and then sterilize your inoculating loop
and let it cool.
c. Turn the plate a quarter of a turn and then pick it back up out of the lid. Starting in the first streak, you
will streak out into the second sector of your plate. Streak two “zig-zags” while touching the first sector
and then streak a few without touching the first.
d. Set the plate back down in the lid and then sterilize your inoculating loop and let it cool.
e. Turn the plate another quarter of a turn and pick it back up out of the lid. Starting in the second streak,
you will streak out into the third sector of your plate. Streak two “zig-zags” while touching the second
sector and then streak a few without touching the second.
f. Set the plate back down in the lid and then sterilize your inoculating loop and let it cool.
g. Turn the plate another quarter of a turn and pick it back up out of the lid. Starting in the third streak,
you will streak out into the fourth sector of your plate. Streak two “zig-zags” while touching the third
44
sector and then create zig-zags across the rest of the empty space on the plate in a tornado pattern (see
pictures.) Be careful not to make contact with any streaks in the previous sections.
h. Set the plate back down in the lid and sterilize your loop one final time.
i. Incubate the plate, media side up, at 37 C until discrete, isolated colonies develop (usually 24 – 48
hours).
Pour Plate
1. Label the bottom of three empty, sterile Petri plates with your group’s name, lab section and the date. Label
one plate “1:20”, another “1:400”, and the third one “1:8000”. Place the labeled plates on your lab table lid-side
up.
2. Get a beaker with 3 tubes of melted nutrient agar from the instructor. Each tube contains 19 mL of nutrient
agar.
3. Obtain a mixed broth culture.
4. Using a sterile disposable pipette, suction up and aseptically transfer 1 mL of the broth to a tube of melted agar
and mix well. Using a second pipette, suction up and aseptically transfer 1 mL from this tube into a second tube.
Work quickly so the agar does not solidify in the pipette. Aseptically pour the contents of the first tube into the
1:20 Petri plate and place the lid on at an angle to vent. Discard the pipettes in the biohazard container.
5. Mix the second tube. With the third pipette, aseptically transfer 1 mL to the third tube. Pour the contents of
the second tube into the 1:400 plate and place the lid on. Mix the third tube and pour its contents into the
remaining 1:8000 plate.
6. Return the beaker of empty tubes to the instructor. Let the agar solidify in the plates. When the plates are
solid, incubate them, media side up, at 37 C until growth is seen (usually 24 – 48 hours). Suggestion: When
incubating multiple plates, use a rubber band or tape to keep the plates together.
PROCEDURE – RETURN TO CHECK (first time)
Streak Plate
1. Record the results of your streak plate. Use proper terms to describe the colonies (refer to Microbes in the
Environment for terminology).
2. Prepare a subculture of one colony. Sterilize your loop and let it cool. To subculture, touch the center of a small
isolated colony located on a streak line, and then aseptically streak a sterile nutrient agar plate using the same
technique used in your original streak plate. Be careful not to pick up or touch more than one colony as you are
attempting to produce a pure culture of only one colony type.
3. Incubate the plate at 37 C until good growth is observed.
Pour Plate
1. Count the number of colonies on the pour plates. Remember that more than 250 is too numerous to count and
fewer than 25 too few to count. Determine the number of bacteria per mL in the original culture.
PROCEDURE – RETURN TO CHECK (second time)
Streak Plate
1. Record the appearance of your subculture.
45
46
Lab #8: Isolation of Bacteria by Dilution Technique
Laboratory Report
Name _________________________
Lab Section ____________
RESULTS
Streak Plate
Draw the appearance of the streak plates.
Mixed Culture
Subculture
Fill in the following table using colonies from the most isolated streak areas.
Colony Description (Describe each different-appearing colony.)
Culture
Diameter
Appearance
Margin
Elevation
Color
Mixed culture
Pour Plate
Dilution
1:20
Number of Colonies
1:400
1:8000
Calculate the number of colony-forming units per milliliter in
the mixed culture. Which plate will you use for your
calculations? _________________
Show your calculations.
_______________ Colony-forming units per mL
47
CONCLUSIONS
1.
How many different bacteria were in the mixed culture?
How can you tell?
2.
When looking at a pour plate, how do the colonies on the surface differ from the colonies suspended in the agar?
3.
How could your group’s streak plate technique be improved?
4.
Did your group get a pure culture in your subculture?
How do you know?
QUESTIONS
1.
Will you always see isolated colonies in the fourth sector on the streak plate?
2.
What is a contaminant?
3.
How can you tell if a colony is a contaminant on a streak plate?
4.
What is a disadvantage of the streak plate technique?
5.
Could some bacteria grow on the streak plate and not be seen using the pour plate technique?
6.
What would happen if you incubated the plates for an additional week?
What about on a pour plate?
What about the pour plate technique?
Explain.
An additional month?
48
Lab #9: Physical Methods of Control – Heat
Objectives
1. Compare the effectiveness of dry heat and moist heat in killing bacteria.
2. Evaluate the heat tolerance of microorganisms.
3. Define and provide a use for each of the following: incineration, dry-heat oven, pasteurization, boiling, and
autoclaving.
BACKGROUND
Extreme temperature is widely used to control the growth of microorganisms. Generally, if heat is applied, microbes are
killed; if cold temperatures are used, microbial growth is inhibited.
Heat Sensitivity
Bacteria exhibit different tolerances to the application of heat. Heat sensitivity is genetically determined and is partially
reflected in the optimal growth ranges, which are psychrophilic (about 0 C to 20 C), psychrotrophic (20 C – 30 C),
mesophilic (25 C to 40 C), thermophilic (45 C – 65 C), hyperthermophilic (about 80 C or higher), and by the presence of
heat-resistant endospores. Overall, bacteria are more heat resistant than most other forms of life. Heat sensitivity of
organisms can be affected by container size, cell density, moisture content, pH, and medium composition.
Types of Heating
Heat can be applied as dry or moist heat. Dry heat, such as that in hot-air ovens is 170 C for 2 hours. The heat of hot air
is not readily transferred to a cooler body such as a microbial cell. Moisture transfers heat energy to the microbial cell
more efficiently than dry air, resulting in the denaturation of enzymes. Moist heat methods include pasteurization,
boiling, and autoclaving. In pasteurization, the temperature is maintained at 63 C for 30 minutes or 72 C for 15 seconds
to kill designated organisms that are pathogenic or cause spoilage. Boiling (100 C) for 10 minutes will kill vegetative
bacterial cells; however, boiling does not inactivate endospores. The most effective method of moist heat sterilization is
autoclaving, the use of steam under pressure. Increased pressure raises the boiling point of water and produces steam
with a higher temperature. Standard conditions for autoclaving are 15 psi, at 121 C, for 15 minutes. This is usually
sufficient to kill endospores and render materials sterile.
Measuring Effectiveness
The effectiveness of heat against a specific microbe can be expressed as the thermal death time. Thermal death time
(TDT) is the length of time required to kill all bacteria in a liquid culture at a given temperature. The less common
thermal death point (TDP) is the temperature required to kill all bacteria in a liquid culture in 10 minutes. Decimal
reduction time (DRT, or D value) is the time, in minutes, in which 90% of a population of bacteria at a given temperature
will be killed.
MATERIALS
IN CLASS SETUP
Petri plates containing nutrient agar (2 per group)
Water bath (63 C and 72 C)
Thermometer
Beaker
49
CULTURES
Group 1
Old (48 -72 hours) Bacillus subtilis
Young (24 hours) Bacillus subtilis
Group 2
Staphylococcus epidermidis
Escherichia coli
Group 3
Young (24 hours) Bacillus subtilis
Escherichia coli
DEMONSTRATION
Autoclaved and dry-heated soil
TECHNIQUES
Inoculating loop technique
Aseptic technique
Plate streaking
Graphing
PROCEDURE
IN CLASS
Carefully read the steps before beginning. Remember to use aseptic techniques even though you must work quickly. A
summary of the procedure is shown in the chart below:
Inoculate Plate
Total Time Microbes Are in Water
Bath (min)
Start
Inoculate 0 time section.
Place tube in water bath.
After 30 sec
Inoculate “30 sec) section.
After 1 min, 30 sec
Inoculate “2 min” section.
After 3 min
Inoculate 5 minute section.
After 10 more min
Inoculate: “15 min” section.
Each lab group is assigned two cultures and a temperature.
Group
Group
1: 63 C ______
1: 72 C ______
2: 63 C _______
2: 72 C ______
3: 63 C _______
3: 72 C ______
You can share water baths as long as the effect of the same temperature is being evaluated.
1. Divide two plates of nutrient agar into five sections each. Label the sections “0”, “30 sec“, “2 min”, “5 min” and
“15 min”. Make sure that you label your plates with the temp and organism!
2. Streak the assigned organisms on the “0” time section of the appropriate plate
50
3. Place the broth tubes of your organism into the bath when the temperature is at the desired point. After 30
seconds, remove the tubes, resuspend the culture, streak a loopful on the corresponding sections, and return
the tubes to the water bath. Repeat at 2, 5, and 15 minutes.
4. When you are done, return the materials to the instructor.
PROCEDURE – RETURN TO CHECK
1. Record your results and the results for the other organisms tested: (-) = no growth, (+) = minimum growth, (2+) =
moderate growth, (3+) = heavy growth, and (4+) = maximum growth.
2. Examine the demonstration plates and record your observations. Collect results from your classmates to
complete the data table.
http://web2.mendelu.cz/af_291_projekty2/vseo/stranka.php?kod=4639
MICROBIAL DEATH RATE:
AN EXAMPLE OF A DRT VALUE OF 1 MINUTE
Number of Survivors
Time Deaths per Minute
(min)
0
1
2
3
4
5
6
0
900,000
90,000
9,000
900
90
9
1,000,000
100,000
10,000
1000
100
10
1
Source: G. J. Tortora, B. R. Funke, and C. L. Case, Microbiology, An
Introduction, 10th ed.m San Francisco, Benjamin Cummings, 2010.
Demonstration Plates
Comparing the effects of dry heat and autoclaving. The control is a pour plate inoculated with soil left at room
temperature (no heat applied). The dry heat plate was inoculated with soil and exposed to dry heat (121 C for 1 hour).
The third plate was autoclaved (121 C for 15 min).
51
Lab #9: Physical Methods of Control – Heat
Laboratory Report
Name ______________________________
Lab Section ________
RESULTS
Record growth on a scale from (-) to (4+).
Temperature/Time
0
30 sec
63 C
2 min
5 min
15 min
0
30 sec
72 C
2 min
5 min
15 min
Organism
Old Bacillus subtilis
Young Bacillus subtilis
Staphylococcus aureus
Escherichia coli
Demonstration Plates
Control
Autoclaved
Dry-Heated
Number of Colonies
Number of Different Colonies
Graph your cultures at ______C.
Organism: ____________________________________
52
Time (min)
Organism: _____________________________________
Time (min)
Conclusions
1. What conclusions can you draw from your data?
2. In the exercise, were we attempting to determine the thermal death time or thermal death point?
Questions
1. Why were “old” and “new” Bacillus cultures used in this exercise?
2. The decimal reduction time (DRT) is the time it takes to kill 90% of cells present. Assume that a DRT value for
autoclaving a culture is 1.5 minutes. How long would it take to kill all the cells if 106 cells were present? What
would happen if you stopped the heating process at 9 minutes?
3. Explain the difference in the heat sensitivity of fungal spores and bacterial endospores.
53
4. Compare the effectiveness of autoclaving and dry heat.
5. Why do fungus and Bacilllus sometimes grow better after being heated?
6. Give an example of a use of thermal death time.
7. A biological indicator used in autoclaving is a vial containing 109 Geobacillus stearothermophilus cells that is
placed in the autoclave with the material to be sterilized. After autoclaving, the vial is incubated and examined
for growth. Why is this species used as opposed to E. coli or Bacillus subtilis?
8. Give an example of a nonlaboratory use of each of the following methods to control microbial growth:
a. Incineration:
b. Pasteurization:
c. Autoclaving:
9. Define pasteurization. What is the purpose of pasteurization?
10. Indicators are used in autoclaving to ensure that sterilization is complete. One type of chemical indicator turns
color when it has reached a specific temperature; the other type turns color when it has reached a specific
temperature and has been exposed to steam. Which type of indicator should be used?
54
Lab #10: Oxygen and the Growth of Bacteria
OBJECTIVES
1. Identify the oxygen needs for each of the following types of organisms: obligate aerobes, obligate anaerobes,
aerotolerant anaerobes, microaerophiles, and facultative anaerobes.
2. Name three methods of culturing anaerobes.
3. Culture anaerobic bacteria.
BACKGROUND
The presence or absence of molecular oxygen (O2) can be very important to the growth of bacteria. Some bacteria,
called obligate aerobic bacteria, require oxygen, whereas others, called anaerobic bacteria, do not use oxygen. One
reason obligate anaerobes cannot tolerate the presence of oxygen is that they lack catalase, and the resultant
accumulation of hydrogen peroxide is lethal. Aerotolerant anaerobes, cannot use oxygen but tolerate it fairly well,
although their growth may be enhanced by microaerophilic conditions. Most of these bacteria use fermentative
metabolism.
Some bacteria, the microaerophiles, grow best in an atmosphere with increased carbon dioxide (5% to 10%) and
lower concentrations of oxygen. Microaerophiles will grow in a solid nutrient medium at a depth to which small
amounts of oxygen have diffused into the medium. To culture microaerophiles on Petri plates and nonreducing media, a
CO2 jar is used. Inoculated plates and tubes are placed in a large jar with a lighted candle or CO2-generating packet.
The majority of bacteria are capable of living with or without oxygen; these bacteria are called facultative
anaerobes.
Five genera of bacteria lacking catalase are Streptococcus, Enterococcus, Leuconostoc, Lactobacillus, and
Clostridium. Species of Clostridium are obligate anaerobes, but members of the other four genera are aerotolerant
anaerobes. The four genera of aerotolerant anaerobes lack the cytochrome system to produce hydrogen peroxide and
therefore do not need catalase. Determining the presence or absence of catalase can help in identifying bacteria. When
a few drops of 3% hydrogen peroxide are added to a microbial colony and catalase is present, molecular oxygen is
released as bubbles.
2H2O2 → 2H2O + O2↑
In the laboratory, we can culture anaerobes either by excluding free oxygen from the environment or by using
reducing media. Many anaerobic culture methods involve both processes. Some of the anaerobic culturing methods
are as follows:
1. Reducing media contain reagents that chemically combine with free oxygen, reducing the concentration of
oxygen. In thioglycolate broth, sodium thioglycolate (HSCH2COONa) will combine with oxygen. A small
amount of agar is added to increase the viscosity, which reduces the diffusion of air into the medium.
Usually dye is added to indicate where oxygen is present in the medium. Resazurin, which is pink in the
presence of excess oxygen and colorless when reduced, or methylene blue are commonly used indicators.
2. Conventional, nonreducing media can be incubated in an anaerobic environment. Oxygen is excluded from
a Brewer anaerobic jar by adding a GasPak and a palladium catalyst to catalytically combine the hydrogen
with oxygen to form water. Carbon dioxide and hydrogen are given off when water is added to the GasPak
envelope of sodium bicarbonate and sodium borohydride. A methylene blue indicator strip is placed in the
jar; methylene blue is blue in h=the presence of oxygen and white when reduced. One of the disadvantages
of the Brewer jar is that the jar must be opened to observe or use one plate. An inexpensive modification of
the Brewer jar has been developed using disposable plastic bags for one or two culture plates. In this
55
technique, the bag is coated with antifogging chemicals, and a wet sodium bicarbonate tablet is added to
generate carbon dioxide. Iron (Fe0, e.g., steel wool) activated with water removes O2 as iron oxide (Fe2O3) is
produced. One plate can be observed without opening the bag.
3. Anaerobic incubators and glove boxes can also be used for incubation. Air is evacuated from the chamber
and can be replaced with a mixture of carbon dioxide and nitrogen.
All methods of anaerobic culturing are effective only if the specimen or culture of anaerobic organisms is
collected and transferred in a manner that minimizes exposure to oxygen. In this exercise, we will try two methods of
anaerobic culturing and will perform the catalase test.
MATERIALS
FIRST PERIOD
Petri plates containing nutrient agar (2)
Tubes containing thioglycolate broth (1)
BioBag Type A with resazurin indicator and palladium catalyst
SECOND PERIOD
3% hydrogen peroxide (H2O2)
CULTURES
Alcaligenes faecalis
Clostridium sporogenes
Enterococcus faecalis
Escherichia coli
TECHNIQUES REQUIRED
Inoculating loop technique
Aseptic technique
Plate streaking
Selective media
Catalase test
PROCEDURE – FIRST PERIOD
1. Don’t shake the thioglycolate.
2. Label one tube of thioglycolate broth (per group), and aseptically transfer one with a loopful of Alcaligenes, one
with Clostridium, one with Enterococcus, or one with Escherichia. Each group will set up a different organism.
3. Incubate the tube at 37 C for at least 24 hours.
4. Using a marker, divide two nutrient agar plates into four sections. Label one plate “aerobic” and the other
“anaerobic”. (Don’t forget to label your plates with the usual information as well!)
5. Streak a single line of Alcaligenes, Clostridium, Enterococcus and Escherichia in the appropriate sector on both
plates. Remember to sterilize your inoculating loop well between each organism!
6. Incubate the “aerobic” plate at 37 C for 24 hours. Place the “anaerobic” plate into one of the anaerobic BioBags
(see instructor for help) with a catalyst, generator and indicator. Seal the bag and break the generator and
indicator. The tablet in the CO2 generator should drop into the solution down below and begin to boil. The
indicator should turn pink.
7. After 5 minutes have passed, put the bag in the 37 C incubator for 24 hours.
56
PROCEDURE – SECOND PERIOD
1. Record the appearance of growth in each of the thioglycolate tubes. Record the growth on each plate.
2. Perform the catalase test by adding a few drops of 3% H2O2 to the different colonies on the nutrient agar plate.
A positive catalase test produces a bubbling white froth. A dissecting microscope can be used if more
magnification is required to detect bubbling. The catalase test may also be done by transferring bacteria to a
slide and adding the H2O2 to it.
57
Lab #10: Oxygen and the Growth of Bacteria
Laboratory Report
Name _______________________________
Lab Section __________
RESULTS
Thioglycolate
Draw the location of bacterial growth in each of the four tubes.
Alcaligenes
Clostridium
Enterococcus
Escherichia
Nutrient Agar Plates
Bacteria
Alcaligenes
Growth
Aerobic
Catalase Reaction
Bacteria
Anaerobic
Catalase Reaction
Clostridium
Enterococcus
Escherichia
Does the anaerobic plate have an odor?
CONCLUSIONS
1. Categorize the four organisms used in this exercise in terms of their oxygen requirements.
Alcaligenes: ______________________________
Clostridium: ______________________________
Enterococcus: _____________________________
Escherichia: ______________________________
58
QUESTIONS
1. Can aerobic bacteria grow without any O2 present? ________ What would you need to do to figure
out whether the bacteria growing on the anaerobic plate are truly obligate anaerobes?
2. Why will obligate anaerobes grow in thioglycolate?
3. When using a thio tube with indicator, what does the appearance of a pink color in the tube mean?
4. What is causing the odors of anaerobic decomposition?
5. Pseudomonas aeruginosa is an aerobe, but able to grow when there is no oxygen present. How is this
possible?
6. Bergey’s Manual describes Streptococcus and Escherichia as facultative anaerobes. How do the oxygen
requirements of these organisms differ? Which one could correctly be called an aerotolerant
anaerobe? Explain.
7. The catalase test is often used clinically to distinguish between two genera of Gram-positive cocci,
___________________________ and _________________________. It is also used to distinguish two
genera of Gram-positivie rods, ____________________________ and __________________________.
59
Lab #11: Chemical Methods of Control: Antimicrobial Drugs
Objectives
1. Define the following terms: antibiotic, antimicrobial drug, and MIC.
2. Perform an antibiotic sensitivity test.
3. Provide the rationale for the agar diffusion technique.
Background
The observation that some microbes inhibited the growth of others was made as early as 1874. Pasteur and others
observed that infecting an animal with Pseudomonas aeruginosa protected the animal against Bacillus anthracis. Later
investigators coined the word antibiosis (against life) for this inhibition and called the inhibiting substance an antibiotic.
In 1928, Alexander Fleming observed antibiosis around a Penicillium mold growth on a culture of staphylococci. He
found that culture filtrates of Penicillium inhibited the growth of many gram-positive cocci and Neisseria spp. In 1940,
Selman A. Waksman isolated the antibiotic streptomycin, produced by an actinomycete. This antibiotic was effective
against many bacteria that penicillin did not affect. Actinomycetes remain an important source of antibiotics. Today,
research investigators look for antibiotic-producing actinomycetes and fungi in soil and have synthesized many
antimicrobial substances in the laboratory. Antimicrobial chemicals absorbed or used internally, whether natural
(antibiotics) or synthetic, are called antimicrobial drugs.
To treat an infectious disease, a physician or dentist needs to select the correct antimicrobial agent intelligently
and administer the appropriate dose; then the practitioner must follow that treatment to be aware of resistant forms of
the organism that might occur. The clinical laboratory isolates the pathogen (disease-causing organism) from a clinical
sample and determines its sensitivity to antimicrobial agents.
Disk-Diffusion Method
In the disk-diffusion method, a Petri plate containing an agar growth medium is inoculated uniformly over its entire
surface. Paper disks impregnated with various antimicrobial agents are placed on the surface of the agar. During
incubation, the antimicrobial agent diffuses from the disk, from an area of high concentration to an area of lower
concentration. An effective agent will inhibit bacterial growth, and measurements can be made of the size of the zones
of inhibition around the disks. The zone size is affected by such factors as the diffusion rate of the antimicrobial agent
and the growth rate of the organism. To minimize the variance between laboratories, the standardized Kirby-Bauer test
for agar diffusion methods is performed in many clinical laboratories with strict quality controls. This test uses MuellerHinton agar. Mueller-Hinton agar allows the antimicrobial agent to diffuse freely.
Minimum Inhibitory Concentration
The minimum inhibitory concentration (MIC) of an antibiotic is determined by testing for bacterial growth in dilutions
of the antibiotic in nutrient broth. When the MIC is determined, inhibition zones can be correlated with MICs. Then,
inhibition zones can be compared to a standard table (Table 1) to use the disk-diffusion method to determine
susceptibility.
In this exercise, we will evaluate antimicrobial agents by the disk-diffusion method.
60
Materials
First Period
Mueller-Hinton agar plate (1 per group)
Sterile swab
Dispensers of antimicrobial disks
Forceps
Incinerator
Return to Check
Ruler
Cultures
Staphylococcus aureus (broth)
Escherichia coli (broth)
Pseudomonas aeruginosa (broth)
Procedure (First Period)
1. Mix broth tube of organism well. Carefully remove sterile swab from wrapper and saturate with broth.
2. Carefully swab the culture onto the plate, using a zig-zag pattern. Swab in three directions to make sure the
entire plate is covered (you want to achieve a solid lawn of bacteria).
3. Let the plate set for 5 minutes (make sure the lid is on the plate).
4. Using sterile forceps, remove an antimicrobial disk from its packaging and gently place it on the surface of your
Mueller-Hinton plate. Be careful not to gouge into the agar. Tap the disk with the forceps to ensure that the
disk adheres to the surface of the plate.
5. Using the same technique, aseptically transfer all required antimicrobial disks to the plate. Try to avoid placing
the disks too close to one another or the edges of the plate (star pattern).
6. Incubate the plate, inverted, at 37 C.
Return to Check
1. Using a ruler, measure the zones of inhibition (mm) on the underside of the plate.
a. If you are unable to measure the diameter, measure the radius and multiply by 2.
b. If you can see any visible colonies at all within a zone, do NOT measure that as part of your zone of
inhibition. To be included in your measurements, the area has to be completely clear of growth.
2. Record the zone sizes on your lab report and, based on the values in Table 1, determine if the organism is
susceptible, intermediate, or resistant.
3. Repeat steps 1 and 2 for the other 2 organisms used in this experiment.
61
quest.arc.nasa.gov
62
Table 1
Disk
Symbol
AM
INTERPRETING INHIBITION ZONES OF TEST CULTURES
Antimicrobial Agent
Disk Content
Diameter of Zones of Inhibiton (mm)
Resistant
Intermediate
Susceptible
<= 13
14 – 16
>=17
<=28
>=29
<=12
13 – 17
>=18
<=17
18 – 20
>=21
<=19
20 – 22
>=23
<=13
14 – 16
>=17
Ampicillin (gram-negative)
20 ug
Ampicillin (gram-positive)
10 ug
C
Chloramphenicol
30 ug
CAZ
Ceftazidime
30 ug
CB
Carbenicillin
100 ug
Carbenicillin when testing
100 ug
Pseudomonas
CF
Cephalothin
30 ug
<=14
15 – 17
>=18
CIP
Ciprofloxacin
5 ug
<=15
16 – 20
>=21
E
Erythromycin
15 ug
<=13
14 – 22
>=23
FOX
Cefoxitin
30 ug
<=14
15 – 17
>=18
G
Sulfisoxazole
25 ug
<=12
13 – 16
>=17
GM
Gentamicin
10 ug
<=12
13 – 14
>=15
IPM
Imipenem
10 ug
<=19
20 – 22
>=23
P
Penicillin (staphylococci)
10 units
<=28
>=29
Penicillin (other bacteria)
10 units
<=14
>=15
PB
Polymyxin
300 units
<=8
9 – 11
>=12
R
Rifampin
5 ug
<=16
17 – 19
>=20
S
Streptomycin
10 ug
<=11
12 – 14
>=15
SXT
Trimethoprim1.25 ug
<=10
11 – 15
>=16
Sulfamethoxazole
23.75 ug
Te
Tetracycline
30 ug
<=14
15 – 18
>=19
Va
Vancomycin
30 ug
>=15
(Staphylococcus spp.)
Vancomycin (enterococci)
30 ug
<=14
15 – 16
>=17
Clinical and Laboratory Standards Institute. Performance Standards for Antimicrobial Disk Susceptibility Tests, 2010.
63
Lab #11: Chemical Methods of Control: Antimicrobials
Laboratory Report
Name _______________________________
Lab Section __________
Results
Antimicrobial
Agent
1. Ampicillin
Disk
Staphylococcus aureus
Code
Zone Size
S, I, or R *
Escherichia coli
Zone
S, I, or R
Size
*
Pseudomonas aeruginosa
Zone Size
S, I, or R *
2. Erythromycin
3. Penicillin
4. SXT
5. Tetracycline
6. Gentamycin
7. Imipenem
8. Ciprofloxacin
9. Vancomycin
* S = susceptible; I = intermediate; R = resistant.
Questions
1.
Which antimicrobial agent was most effective against Staphylococcus aureus?
2. Which antimicrobial agent was most effective against Escherichia coli?
3. Which antimicrobial agent was most effective against Pseudomonas aeruginosa?
4. Why is it so important that the entire Mueller-Hinton agar plate be completely covered with a lawn of bacteria?
5. Why can’t we use the same antimicrobial agent for all microorganisms?
6. Using the Kirby-Bauer test, are we measuring bacteriostatic or bactericidal activity? Explain.
64
7. The following results were obtained from a disk-diffusion test against a bacterium:
Antibiotic Zone of Inhibition (mm)
A
19
B
12
C
19
D
14
Which drug should be used to treat an infection caused by this bacterium? Briefly explain.
8. Antibiotics are substances produced by ___________________, which inhibit the growth of other organisms.
a. Ben and Jerry
b. Fungi and viruses
c. Viruses and parasites
d. Bacteria and fungi
9. An antibiotic that has a lethal effect on a microorganism is said to be:
a. An analogue
b. A porin
c. Bacteriostatic
d. Bactericidal
10. The disk diffusion technique is being used to determine the antibiotic susceptibility of E. coli isolated from a
urinary tract infection. Forty minutes elapses between the time the bacterium is applied to the surface of the
Mueller-Hinton agar plate and the application of the antimicrobial disks. What effect might this have on the
interpretation of zone sizes?
a. No effect
b. Potential for false resistance
c. Potential for false susceptibility
11. Why is the disk-diffusion technique not an exact indication of how the drug will work in vivo? What other
factors do we need to think about before using an antimicrobial agent?
65
Lab #12: Chemical Methods of Control – Disinfectants and Antiseptics
Objectives:
1. Define the following terms: disinfectant and antiseptic.
2. Describe the use-dilution test.
3. Evaluate the relative effectiveness of various chemical substances as antimicrobial agents.
Wide varieties of chemicals called antimicrobial agents are available for controlling the growth of microbes.
Chemotherapeutic agents are used internally and will be evaluated in another exercise. Disinfectants are chemical
agents used on inanimate objects to lower the level of microbes on their surfaces; antiseptics are chemicals used on
living tissue to decrease the number of microbes. Disinfectants and antiseptics are able to affect bacteria in a number of
ways. Those that result in the death of bacteria are called bactericidal agents. Those that cause only temporary
inhibition of growth are called bacteriostatic agents.
There is no single chemical out there that is the best to use in every situation. Antimicrobial agents must be
matched to specific organisms and environmental conditions. Additional variables to consider in choosing an
antimicrobial agent include pH, solubility, toxicity, organic material present, and cost. When evaluating the
effectiveness of antimicrobial agents, the concentration, length of contact, and whether it is lethal (-cidal) or inhibiting (static) are most important. The standard method for measuring the effectiveness of a chemical agent is the American
Official Analytical Chemist’s use-dilution test. For most purposes, three strains of bacteria are used in this test:
Salmonella enterica choleraesuis, Staphylococcus aureus, and Pseudomonas aeruginosa. To perform a use-dilution test,
metal rings are dipped into standardized cultures of the test bacteria grown in liquid media, removed, and dried. The
rings are then placed into a solution of the disinfectant at the concentration recommended by the manufacturer for 10
minutes at 20 C. The rings are then transferred to a nutrient medium to permit the growth of any surviving bacteria.
The effectiveness of the disinfectant can then be determined by the amount of resulting growth. The use-dilution test is
limited to bactericidal compounds and can’t be used to evaluate bacteriostatic compounds.
In this exercise, we will perform a modified use-dilution test.
MATERIALS
Petri plate containing nutrient agar (1 per group)
5 mL test tube of test substance / lab disinfectant (1 per group)
Various Test Substances: chemical agents such as bathroom cleaner, floor cleaner, mouthwash, lens
cleaner and acne cream
Lab Disinfectant: 10% bleach, liquid from sani-cloth wipes, etc.
Sterile pipette (1)
Timer
CULTURES
Pseudomonas aeruginosa (broth)
Staphylococcus aureus (broth)
Techniques Required
66
Inoculating loop technique
Aseptic technique
Pipetting
PROCEDURE (in class)
1. Each group will be assigned one organism and one test substance/lab disinfectant by the instructor. The
disinfectants that we will be testing have already been diluted to the strength that they would normally be used
at. i.e. mouthwash (undiluted), bleach (diluted 1:10), etc.
2. Label your nutrient agar plate with the name of the organism and the test substance your group has been
assigned. Divide the plate up into five sections. Label the sections “0”, “2.5”, “5”, “10”, and “20”.
3. Inoculate the 0 sector with a loopful of your assigned bacteria.
4. Aseptically add 0.5 mL of the assigned culture to your tube of disinfectant.
5. Transfer one loopful from your tube to a corresponding sector of the plate at 2.5 minutes, 5 minutes, 10 minutes
and 20 minutes.
6. Incubate the plates, inverted, at 37 C. You may return to check as early as 24 hours.
7. Discard the chemical/bacteria mixtures in the rack indicated by the instructor.
PROCEDURE (return to check)
1.
Observe the plates for growth. Record the growth as (-) = no growth, (+) = minimum growth, (2+) = moderate
growth, (3+) = heavy growth, and (4+) = maximum growth. Observe the results of students using the other
organism and the same disinfectant that your group used as well as the two organisms tested with another
disinfectant. You will look at 4 plates total.
67
Lab #12: Chemical Methods of Control – Disinfectants and Antiseptics
Laboratory Report
Name ____________________________
Lab Section ________________
RESULTS
1.
What organism did your group use?
2.
What disinfectant did your group use?
Time of Exposure
(min)
0
Control
Amount of Growth
Staphylococcus aureus Pseudomonas aeruginosa
2.5
5
10
20
Disinfectant Tested:
Now look at another group’s data:
Time of Exposure
(min)
0
Control
Amount of Growth
Staphylococcus aureus Pseudomonas aeruginosa
2.5
5
10
20
Disinfectant Tested:
QUESTIOS
1. Was the disinfectant your group tested effective? Why/Why not?
2.
Was your disinfectant bactericidal or bacteriostatic?
3.
Was this a fair test? Is it representative of the effectiveness of the test substance?
68
4.
How could the procedure we just used in this lab be changed to measure bacteriostatic effects?
5.
In the use-dilution test, a chemical is evaluated by its ability to kill 106 to 108 dried Clostridium sporogenes or Bacillus subtilis
endospores. Why is this considered a stringent/strict test?
6.
The effectiveness of disinfectants can be measured in DRT values. DRT or decimal reduction time, is the length of time it
takes to kill 90% of a test population of bacteria. The DRT values for contact lens disinfectants against Serratia marcescens
are as follows:
Disinfectant
DRT Value (min) Disinfectant
DRT Value (min)
Chlorhexidine, 0.005%
2.8
Thimerosal, 0.002%
138.9
Hydrogen peroxide, 3%
3.1
Polyquaternium-1, 0.001%
383.3
a.
Which disinfectant is most effective?
b.
What is the minimum time that lenses with 10 2 bacteria should be soaked in chlorhexidine?

What about polyquaternium-1?
c.
What if the lenses are contaminated with Staphylococcus or Acanthamoeba?
d.
Why isn’t a higher concentration of disinfectant used?
69
Lab #13: Bacteria of the Skin
Objectives:
1.
2.
3.
4.
Isolate and identify bacteria from the human skin
Provide an example of normal skin microbiota.
List characteristics used to identify the staphylococci.
Explain why many bacteria are unable to grow on human skin.
The skin is generally an inhospitable environment for most microorganisms. The dry layers of keratin-containing cells
that make up the epidermis (the outermost layer of the skin) are not easily colonized by most microbes. Sebum,
secreted by oil glands, inhibits bacterial growth, and salts in perspiration create a hypertonic environment. Perspiration
and sebum are nutritive for certain microorganisms, however, which establishes them as part of the normal microbiota
of the skin.
Normal microbiota of the skin tend to be resistant to drying and to relatively high salt concentrations. More
bacteria are found in moist areas, such as the axilla (armpit) and the sides of the nose, than on the dry surfaces of arms
or legs. Transient microbiota are present on hands and arms in contact with the environment.
Skin Microbiota
Propionibacterium live in hair follicles on sebum from oil glands. The propionic acid they produce maintains the pH of
the skin between 3 and 5, which suppresses the growth of other bacteria. Most bacteria on the skin are gram-positive
and salt-tolerant. Mannitol salt agar is selective for salt-tolerant organisms and is differential in that mannitolfermenting organisms will produce acid, turning the indicator in the media yellow.
Staphylococcus aureus is part of the normal microbiota of the skin and is also considered a pathogen. S. aureus,
which produces coagulase, an enzyme that coagulates (clots) the fibrin in blood, is pathogenic. A test for the presence
of coagulase is used to distinguish S. aureus from other species of Staphylococcus.
Although many different bacterial genera live on human skin, in this exercise we will attempt to isolate and
identify a catalase-positive, gram-positive coccus.
Materials
In Class
Mannitol salt agar plate (1 per group)
Sterile swab (1 per group)
Sterile saline
Incinerator and inoculating loop
First Return to Check
Mannitol salt agar plate (MSA)
3% hydrogen peroxide
Gram-staining reagents
Toothpick
Incinerator and inoculating loop
70
Second Return to Check
Coagulase kit
Procedure – In class
1. Choose a site to culture. Possible areas: sides of the nose, axilla, elbow, between fingers, etc.
2. Label a MSA plate with your group’s information. Make sure that you write the source of the inoculum on the
plate.
3. Wet a sterile swab with sterile saline, push the swab against the wall of the test tube to express excess saline.
Swab the surface you wish to culture. Make sure that you scrub the swab over the area well.
4. Swab 1/3 of the MSA plate with the swab. Using a sterile loop, streak back and forth into the swabbed area a
few times and then streak away from the inoculum, covering about 1/3 of the agar. Sterilize your loop and
spread the bacteria over the rest of the agar. You should have three separate areas on your “streak plate”.
5. Incubate the plate, at 37 C for 24 – 48 hours.
Procedure – First Return to Check
1. Examine the colonies on your streak plate. Record the appearance of the colonies on your lab report. Look at
the colonies for pigment production. Remember, a pigment is a color produced by the cells, not a change in the
pH indicator. Record any mannitol fermentation (yellow halos).
2. If you do not have any colonies with a yellow halo, you will use one of Kelly’s plates for the remaining tests.
3. Perform a Gram stain of the colonies with yellow halos. Record the results on your lab report.
4. Test for catalase production. Record the results on your lab report.
5. Subculture a colony growing in an area with a yellow halo on another MSA plate. Make sure that you label your
plate “subculture” and include the lab section that you are normally in (i.e. Wpm).
6. Incubate your subculture, at 37 C for 24 – 48 hours.
Procedure – Second Return to Check
1. Examine your subculture for purity and test it for coagulase.
71
When you come back to check the first time:
1. Examine the colonies on your plate. Record the appearance of the colonies and any mannitol fermentation
(yellow halos) that you see.
****If your plate does not have any colonies with yellow colonies, please use one of Kelly’s plates on the counter in the
storeroom.***
2. Gram Stain (use one of the sinks in room 532 or 534)
-prepare smear using drop of sterile water (remember to make your smear thin / not too cloudy)
-don’t forget to allow your slide to air dry
-heat fix your slide before staining
1. Crystal Violet – 1 min
2. Rinse
3. Iodine – 1 min
4. Rinse
5. Decolorizer – 5 – 10 seconds (depending)
6. Rinse Immediately
7. Safranin – 1 min
8. Rinse
9. blot with Kimwipe and examine under oil-immersion
2. Catalase Test (second fridge from the left; brown bottle in the door)
-add drop of hydrogen peroxide to glass slide
-add colony from plate with yellow halo to hydrogen peroxide using sterilized inoculating loop or toothpick and
watch closely for the formation of bubbles.
3. Set-up subculture (using a catalase-positive, gram-positive cocci from your /Kelly’s plate)
-use another Mannitol Salt Agar plate and streak for isolation (attempt to get a pure culture)
-extra plates will be in the second fridge from the left
When you come back the second time:
1. Coagulase Test (Sure Vue Staph ID – white box with pink label on bottom shelf in second fridge from left)
1. Cut out one of the circles from a paper card with a scissors.
2. Add a drop of well-mixed latex reagent (pinkish/red liquid) to one side of the circle (make sure you don’t
accidentally grab the positive/negative controls). Make absolute sure that you have mixed the reagent
before adding a drop to the circle.
3. Add two colonies from your subculture to the other side of the circle using one of the plastic picks provided
in the kit.
4. Using the plastic pick, mix the bacterial colonies into the drop of latex.
5. Rotate the card for 25 seconds observing for red clumps. Positive reactions (agglutination/clumping) usually
take 5 – 20 seconds.
72
Streaking for Isolation
faculty.ccbcmd.edu
Catalase Test
www.cdc.gov
Coagulase Test
73
Lab #13: Bacteria of the Skin
Name _____________________________________
Laboratory Report
Lab Section _________________
Results
Source of inoculum: ________________________________________________________________
First Mannitol Salt Plate
1
Colony
2
3
Colony description
Pigment
Mannitol fermentation
Which colony will you use to make your subculture? _____________________________
Catalase reaction: __________________ (Positive or Negative)
Gram stain
Reaction: _________________ Morphology: ____________________ Arrangement: _______________
Second Mannitol Salt Plate
Coagulase Test
Clumping: ____________ Positive or Negative: ___________________
74
Conclusions
1. What organism did you identify? _________________________________________________ (Staphylococcus
aureus or not Staphylococcus aureus)
2. Is this organism a pathogen?
Explain.
3. After you have observed a Gram-positive cocci, what additional information do you need before performing a
coagulase test?
4. Why shouldn’t you perform a subculture on a colony that has not been tested for catalase?
Questions
1. What makes mannitol salt agar selective and differential for microorganisms of the skin?
2. List three ways our skin protects us from infection.
3. Mannitol salt agar is used for:
a. Differentiating fermenters from oxidizers.
b. Selective inhibition of salt-tolerant organisms.
c. Selective isolation of gram-positive organisms.
d. Selective isolation of pathogenic staphylococci.
4. Which of the following correctly describes Staphylococcus aureus?
a. Gram-negative cocci in clusters
b. Catalase positive
c. Coagulase negative
d. Salt intolerance
5. Growth surrounded by yellow halos on mannitol salt agar indicates:
a. The organism cannot ferment mannitol
b. The organism cannot tolerate high salt concentrations
c. The organism can sustain high salt concentrations and ferment mannitol
d. None of the above
6. What is coagulase? How is it related to pathogenicity?
75
7. An 8-year-old boy went to the pediatrician complaining of pain in his right hip. He had no previous injury to the
hip or leg. His temperature was 38 C. A bone scan revealed damage to the head of the femur. Bacterial
cultures of blood and fluid from the hip joint were positive for a Gram-positive, catalase-positive, coagulasepositive cocci that ferment mannitol. Identify the species of this bacterium.
76
Lab #14: MAKE-UP LAB – Microbes in Food: Contamination
Objectives:
1. Determine the approximate number of bacteria in a food sample using a standard plate count.
2. Provide reasons for monitoring the bacteriologic quality of foods.
3. Explain why the standard plate count is used in food quality control.
Illness and food spoilage can result from microbial growth in foods. The sanitary control of food quality is concerned
with testing foods for the presence of pathogens. During processing (grinding, washing and packaging), food may be
contaminated with soil microbes and microbiota from animals, food handlers, and machinery.
Foods are the primary vehicle responsible for transmitting diseases of the digestive system. For this reason,
they are examined for the presence of coliforms because the presence of coliforms usually indicates fecal
contamination.
Standard plate counts are routinely performed on food and milk by food-processing companies and public
health agencies. The standard plate count is used to determine the total number of viable bacteria in a food sample.
The presence of large numbers of bacteria is undesirable in most foods because it increases the likelihood that
pathogens will be present, and it increases the potential for food spoilage.
Calculating Numbers of Bacteria
In a standard plate count, the number of colony-forming units (cfu) is determined. Each colony may arise from a group
of cells rather than from one individual cell. The initial sample is diluted through serial dilutions to obtain a small number
of colonies on each plate. A known volume of the diluted sample is placed in a sterile Petri dish, and melted cooled
nutrient agar is poured over the inoculum. After incubation, the number of colonies is counted. Plates with between 25
and 250 colonies are suitable for counting. A plate with fewer than 25 colonies is unsuitable for counting because a
single contaminant could influence the results. A plate with more than 250 colonies is extremely difficult to count. The
microbial population in the original food sample can then be calculated using the following equation:
Colony forming units/gram or mL of sample
=
Number of colonies
Amount plated x dilution *
A limitation of the standard plate count is that only bacteria capable of growing in the culture medium and
environmental conditions provided will be counted. A medium that supports the growth of most heterotrophic bacteria
is used.
*Dilution refers to the tube prepared by serial dilutions. For example, if 250 colonies were present on the 1:10 6 plate, the
calculation would be as follows:
Colony-forming units/grams = 250 colonies
0.1 mL x 10-5
= 250 x 105 x 101
= 250,000,000
= 2.5 x 108
77
Materials
Melted standard plate count or nutrient agar, cooled to 45 C
Sterile 1 mL pipettes (8)
Sterile Petri dishes (10)
Sterile weighing dish of food samples/sterile beaker of milk
Sterile 99-mL dilution blanks (3)
Sterile 9-mL dilution blanks (2)
Vortex mixer
Techniques Required
Aseptic technique
Pour-plate technique
Pipetting
Serial dilution technique
Procedure (In class set-up)
Bacteriologic Examination of Milk
1. Obtain a sample of pasteurized milk.
2. Using a sterile 1-mL pipette, aseptically transfer 1 mL of the milk into a 9-mL dilution blank; label the tube
“1:10”, and discard the pipette. Mix the contents of the tube on a vortex mixer.
3. Using a sterile 1-mL pipette, aseptically transfer 1 mL of the 1:10 dilution into a 99-mL dilution blank; label the
bottle “1:103” and discard the pipette. Shake the bottle 20 times, with your elbow resting on the table and the
container hanging over the edge of the counter (be careful not to hit the lab table with the container).
4. Label the bottoms of four sterile Petri dishes with the dilutions: “1:10”, “1:102”, “1:103”, “1:104”. Make sure you
also label the top plate in the stack with your lab information: group name, section, date and the source of your
inoculum (milk).
5. Using a 1-mL pipette, aseptically transfer 0.1 mL of the 1:103 dilution into the bottom of the 1:104 dish. Note:
0.1 mL of the 1:103 dilution results in a 1:104 dilution of the original sample. Using the same pipette, transfer 1.0
mL of the 1:103 dilution into the dish labeled 1:103. Pipette 0.1 mL and 1.0 mL from the 1:10 dilution into the
1:102 and 1:10 dishes, respectively, with the same pipette.
6. Check how hot the melted nutrient agar is in your hand before you start. It should be warm, but not too hot for
your bare hand.
7. Pour the melted nutrient agar into one of the dishes (to about 1/3 full). Cover the plate with the lid slightly
askew to allow the steam to escape, and swirl it gently to distribute the milk sample evenly through the agar.
Continue until all the plates are poured.
8. When each plate has solidified, invert it, tape all of the plates together with the plate with the identification on
the top, and incubate all plates at 37C for 24 – 48 hours.
Bacteriologic Examination of Hamburger, Hot Dog, or Frozen Vegetables
1. Obtain 1 g of raw hamburger (or hot dog) or frozen, thawed vegetables on a covered weighing dish.
2. Transfer the 1 g of food into a 9-mL dilution blank; label the tube “1:10”. Mix the contents on a vortex mixer,
Mixing will shake bacteria off of the food and put them in suspension.
3. Using a sterile 1-mL pipette, aseptically transfer 1 mL of the 1:10 dilution into a 99-mL dilution blank; label the
bottle “1:103” and discard the pipette. Shake the bottle 20 times, with your elbow resting on the table. Make a
1:105 dilution using another 99-mL dilution blank. Shake the bottle as before.
78
4. Label the bottoms of six sterile Petri dishes with the dilutions: “1:10”, “1:102”, “1:103”, “1:104”, “1:105”, and
“1:106”. Make sure you also label the top plate in the stack with your lab information: group name, section,
date and the source of your inoculum.
5. Using a 1-mL pipette, aseptically transfer 0.1 mL of the 1:105 dilution into the 1x106 dish. Note: 0.1 mL of a 1:105
dilution results in a 1:106 dilution of the original sample. Using the same pipette, repeat this procedure with the
1:103 dilution and the 1:10 dilution until all the dishes have been inoculated.
6. Check how hot the melted nutrient agar is in your hand before you start. It should be warm, but not too hot for
your bare hand.
7. Pour the melted nutrient agar into one of the dishes (about 1/3 full). Cover the plate with the lid slightly askew
to allow the steam to escape, and swirl it gently to distribute the sample through the agar evenly. Continue until
all the plates are poured.
8. When each plate has solidified, invert it, tape all of the plates together with the plate with the identification on
the top, and incubate all plates at 37 for 24 – 48 hours.
Procedure (return to check)
1. Arrange each plate in order from lowest to highest dilution.
2. Select the plate with 25 to 250 colonies. Record data for plates with fewer than 25 colonies as too few to count
(TFTC) and those with more than 250 colonies, too numerous to count (TNTC).
3. Count the number of bacteria in the original food. For example, if 129 colonies were counted on a 1:103 dilution:
79
Make-up Lab: Microbes in Food – Contamination
Name ________________________
Laboratory Report
Lab Section ____________________
RESULTS
Milk sample
Dilution
1:
Food sample: ________________
Colonies per Plate
Dilution
1:
1:
1:
1:
1:
1:
1:
Colonies per Plate
1:
1:
Conclusions
1. What did your group test?
Number of colony-forming units per mL (or gram) of original sample:
Make your calculations here:
2. In a quality-control laboratory, each dilution is plated in duplicate or triplicate. Why would be increase the
accuracy of a standard plate count?
Questions
1. Why are plates with 25 to 250 colonies used for calculation?
2. There are other techniques for counting bacteria, such as a direct microscopic count and turbidity. Why is the
standard plate count preferred for food?
3. Why is ground beef a better bacterial growth medium than a steak or roast?
80
4. Why does repeated freezing and thawing increase bacterial growth in meat?
5. Assume that a standard plate count indicates that substantial numbers of microbes are present in the food
sample. What should be done next to determine whether the food sample is a danger to the consumer?
6. Pasteurized milk is allowed 20,000 cfu/mL. How many colonies would be present if 1 mL of a 10-3 dilution was
plated?
Is 20,000 cfu/mL a health hazard?
An outbreak of botulism associated with home-canned chili peppers killed 16 members of one family. Would a standard
plate count have detected the etiologic agent in the canned chili p
81