Microbiology
Applied Microbiology
Laboratory Manual Bio-300
By Dr. Ahmed Gharib adapted for the Queens Campus with revisions by
Vilma Greene, PhD & Elizabeth Gray, M.S.
With contributions from
SAINT PAUL'S SCHOOL OF NURSING
Safety
Microscopy
Preparation of Agar Plates
Aseptic Technique
Smears
Staining
Culturing and Isolation Techniques
Selective and Differential Media
Identification of Gram-positive Cocci
Throat Culture
Skin Flora
MIC
Identification of Gram-negative Bacilli
Motility
Hydrolysis of Polysaccharides
Deamination of Amino Acids
IMViC
Disc Diffusion Susceptibility Methods
Urinalysis
LAB ATTENDANCE AND MAKE-UP POLICY
Attendance is checked and recorded for each lab. Students are expected to attend all scheduled
laboratory classes. Absences may place a student in jeopardy of failing to learn or demonstrate
essential content, thus failing the course.
If a student occasionally cannot attend a lab, arrangements should be made with a fellow
classmate to get notes from the missed lab. Students are responsible for all announcements,
handouts, and other material presented in lab and for meeting all course requirements. A faculty
member has the right to deny entrance to the classroom if students arrive after the start of lab if it
is considered to be disruptive to the learning environment. Frequently arriving late or leaving
class early may negatively impact a student's success in the course. Please be aware that it may
not be possible to make-up class exercises or assignments.
With valid justifications, make-ups must be done within the same week of the missed lab in
order to be able to cover the same material. Otherwise there will be absolutely no make-ups.
There are four Micro lab sessions running on Thursdays only (9:00AM, 12:30PM, 4:30PM &
8:00PM). Previous coordination and approval form the instructor and the registrar (Claudia
Menjivar) is needed. It is the student’s responsibility to make an appointment with the course
coordinator before or right after of the missed lab to schedule a makeup. If this is not done, a
grade of zero (0) will be assigned.
Safety
Introduction
In the laboratory individuals are exposed to hazards not found in a regular classroom. It is essential that
students follow all rules established by the lab instructor, lab manager, or lab assistant to ensure the
safety of all individuals in the class. Failure to follow established rules may result in dismissal of the
individual from the class. Laboratories have certain standard safety equipment. These typically include a
general-purpose fire extinguisher, eyewash, safety shower and cut off switches for electrical and gas
outlets. It is the responsibility of the student to locate and know how to use the general safety
equipment in the lab. Additionally, students should be aware of exits from the room in case of
emergency, the location of the nearest fire call box, how to contact Campus Security, and how to obtain
emergency medical assistance.
The microbiology lab has some additional safety considerations. Since individuals work with potentially
pathogenic organisms, care must be taken to prevent possible infection or transmission of the
organisms from the lab. Students must wear protective clothing (lab coats) while working in the lab. Lab
coats must not be worn outside the lab. Intact skin is an adequate barrier against microorganisms so
gloves are not necessary in the lab. Gloves will be provided and students may wear gloves when
handling cultures if they so desire. Tabletops must be disinfected before and after lab using the
disinfectant provided. Instruction in aseptic technique will be demonstrated and must be followed while
working with microorganisms. Hand washing is a simple and effective way to prevent the transmission
of disease. While antibacterial soap may provide some additional protection, the major effect of hand
washing is the mechanical removal of microbes from the skin. Friction when washing hands is important
to mechanically remove organisms from the surface of the skin. Using a paper towel to turn off the tap
prevents recontamination of the hands with microorganisms. Hands must be washed whenever the
student leaves the lab.
Two copies of the Laboratory Safety Rules are included. One must be signed and returned to the lab
instructor at the end of class. The additional copy is for your reference.
Signed _____________________________
Date ____________
Name (please print) ____________________________________
Microbiology Laboratory Safety Rules
1. All materials and clothes other than those needed for the laboratory are to be kept away from the
work area.
2. A lab coat or other protective clothing must be worn during lab. The lab clothing is not to be worn
outside of the laboratory.
3. Clean the lab table before and after lab with the disinfectant solution provided.
4. Wash hands before leaving the lab.
5. Any item contaminated with bacteria or body fluids must be disposed of properly. Disposable items
are to be placed in the BIOHAZARD container. Reusable items are to be in the designated area for
autoclaving prior to cleaning. Sharps are to be disposed of in the appropriate container.
6. Reusable items should have all tape and marks removed by the student before being autoclaved.
7. Because organisms used in this class are potentially pathogenic, aseptic technique must be observed
at all times. NO eating, drinking, application of cosmetics or smoking is allowed. Mouth pipetting is not
allowed.
8. Cuts and scratches must be covered with bandages. Disposal gloves will be provided on request.
9. Long hair should be tied back while in the lab.
10. All accidents, cuts, and any damaged glassware or equipment should be reported to the lab
instructor immediately.
11. Sterilization techniques will involve the use of Bacticinerators that are fire and bum hazards.
Bacticinerators reach an internal temperature of 850°C or 1500°F. Keep all combustibles away from the
Bacticinerators. Do not leave inoculating loops or needles propped in the Bacticinerator.
12. Microscopes and other instruments are to be cared for as directed by the instructor.
13. It is the responsibility of the student to know the location and use of all safety equipment in the lab.
14. Cultures may not be removed from the lab. Visitors are not allowed in the lab.
15. Doors and windows are to be kept closed at all times.
16. For the best lab experience, read labs before coming to class. Make notes as necessary. Wait for a
laboratory introduction by the instructor before starting work.
I have read and understand the above rules and agree to follow them.
Signed _____________________________
Date ____________
Name (please print) ____________________________________
Safety Review Questions
1. List all emergency exits from the laboratory.
2. Describe how you would obtain emergency medical assistance.
3. What protective clothing must be worn during the lab?
4. What is a simple and effective way to prevent disease transmission?
5. What general safety equipment is found in the lab?
6. What do you need to bring to the lab table for each class?
7. How do you dispose of materials that may be contaminated with bacteria?
8. How do you dispose of a broken slide?
9. You have just disinfected your lab table. Where do you dispose of the paper towels you used?
10. After washing your hands, where do you dispose of your paper towels?
11. When discarding reusable contaminated material where do you put it?
12. It is the end of lab. What must you do before leaving? List the tasks in order of performance.
Microscopy
Introduction:
Microorganisms are too small to be seen with the naked eye so a microscope must be used to visualize
these organisms. While a microscope is not difficult to use, it does require some practice to develop the
skills necessary to use the microscope to its maximum capabilities. Bacteria and other cellular
microorganisms are measured in micrometers (µm) or 1 x 10-6 meters.
Viruses are even smaller and are measured in nanometers (nm) or 1 x 10-9 meters. When carrying a
microscope, always use both hands. One should be on the arm of the microscope and one should be
under the base of the microscope.
Discussion:
There are several types of microscopes but the only one used in this laboratory is the compound light or
bright-field microscope. Individual microscopes will vary depending on the manufacturer, but all
microscopes have the same basic features. These microscopes are known as compound microscopes
because there are two magnifying lenses in the microscope. One magnifying lens is in the ocular and one
is in the objective. Initial magnification occurs in the objective lens. Each objective has the magnification
power written on the lens. The magnification of the ocular lens, in which final magnification occurs, is
also inscribed on the lens. Low magnifications are used for quickly examining the slide to find an
appropriate area to examine. Higher magnifications allow the examination of a particular object on the
slide. Each contributes to the magnification of the object on the stage. The total magnification of any
set of lenses is determined by multiplying the magnification of the objective by the magnification of the
ocular. The nosepiece rotates, allowing the objectives to change and thus change the magnification of
the microscope. The stage is where the slide is placed. The stage adjustment knobs allow the slide to be
moved easily. Light provides the illumination for the specimen.
The amount of light reaching the eye is controlled through the iris diaphragm lever that opens and
closes the condenser diaphragm. On low magnifications less light is needed than on higher
magnifications. Too much light on low magnification may mask the specimen, particularly something as
small as a bacterial cell.
The coarse and fine adjustment knobs are used to focus on the specimen. When a slide is on the stage
there is a space between the objective and the slide. This space is known as the working distance. The
coarse adjustment knob will cause the working distance to visibly change while the fine adjustment
knob is used ONLY FOR FINAL, fine focusing. The ability to see things using a microscope is limited by the
resolving power of the microscope.
Resolution: the resolving power of a microscope is the distance two objects must be apart and still be
seen as separate and distinct. For the light microscope, this is 0.2 µm. Objects closer together than 0.2
µm will not be seen distinctly. Increasing the magnification will not make the objects more distinct, just
bigger. The limit of resolution of a lens is the minimum distance between two closely spaced points in an
object that can be seen as distinct points in the image.
Please note and identify the following parts of the microscope:
OCULAR
OBJECTIVES
NOSEPIECE
STAGE
ARM
BASE
IRIS DIAPHRAGM
CONDENSER
STAGE ADJUSTMENT KNOB
COARSE ADJUSTMENT KNOB
FINE ADJUSTMENT KNOB
LIGHT SOURCE
Examine your microscope and fill in the appropriate values in the table below below:
POWER OF OCULAR
POWER OF OBJECTIVE
Scanning =
Low power =
High dry =
Oil immersion =
TOTAL MAGNIFICATION
Now turn on your microscope and look through the ocular where you will see a lighted circle. This is
known as the field of view or the field. While looking through the microscope, move the iris diaphragm
lever. Notice how the brightness of the light changes. As you move the objectives to provide increased
magnification, you will look at a smaller section of the slide. Be sure you move the object you want to
view into the center of the field before moving to the next objective.
These microscopes are parfocal. Once you have focused on an object using one objective the object will
be approximately in focus on the next objective. Use of the fine focus knob will sharpen the focus.
PROCEDURE FOR FOCUSING
1. Obtain a slide.
2. Use the coarse adjustment knob to obtain maximum working distance.
3. Place the slide on the stage. The slide should fit into the slide holders but is not placed under the slide
holder. Use the stage adjustment knob to move the slide over the hole in the stage.
4. Rotate the scanning (4X) objective in place.
5. Use the coarse adjustment knob to obtain the minimum working distance. Develop the habit of
watching this process to be sure the objective does not hit the slide, thus breaking it.
6. Look through the ocular. Adjust the light with the iris diaphragm lever if necessary. Slowly turn the
coarse adjustment knob until something comes into focus. Use the fine adjustment to sharpen the
focus.
7. Using the stage adjustment knob, move the slide until the object you wish to examine is in the center
of the field.
8. Rotate the low power (10X) objective into place. Use the fine adjustment knob to sharpen the focus.
Do not use the coarse adjustment knob. Adjust the light using the iris diaphragm lever if necessary.
9. Rotate the high power (40X) into place. Again use the fine adjustment knob to sharpen the focus.
10. Rotate the high power halfway to the next position. Place a drop of immersion oil on the slide, and
then rotate the oil immersion (I OOX) objective into place. The objective should be immersed in the oil
on the slide. Use the fine adjustment knob to sharpen the focus. Adjust the light using the iris diaphragm
lever if necessary.
11. When finished viewing the slide, use the coarse adjustment knob to maximize the working
distance and remove the slide from the stage. If you want to look at another slide, begin the process
over. If you are finished with the microscope, clean the microscope and return it to storage.
Procedure for Preparing Microscopic Specimens
1.
2.
3.
4.
With a toothpick, obtain skin cells by scraping the inside of your cheek.
Transfer the scrapings on to a brand new microscope slide
Using a transfer pipette put one small drop of methylene blue stain directly on top of the smear
To place the coverslip over the specimen, put one edge on the slide and slowly let it drop over the
specimen
5. If there is any excess stain around the edges of the coverslip, you may use a paper towel to absorb it
6. Visualize cells under the microscope
Procedure for Cleaning a Microscope
1. Turn off the light and unplug the cord. Store the cord appropriately.
2. Using the coarse adjustment knob to obtain maximum working distance, remove the slide from the
stage.
3. Using lens paper, clean all the lenses starting with the ocular, scanning, low power, high power and
oil immersion. Use lens cleaner if necessary.
4. Clean any oil off the stage using paper towels.
5. Rotate the scanning objective into place. Use the coarse adjustment knob to obtain minimum
working distance.
6. Return the microscope to the appropriate storage area.
Review Questions
1. Define:
a. Resolving power
2.
3.
4.
5.
6.
b. Parfocal
c. Field
d. Working distance
What are the functions of each of the following:
a. Coarse adjustment knob
b. Fine adjustment knob.
c. Iris diaphragm.
d. Stage adjustment knob
What unit of measurement is used for measuring bacteria?
How do you determine the total magnification of a set of lenses?
Describe the process for focusing on a slide.
Describe how to properly clean a microscope.
Preparation of Microbiological Culture Media
Introduction
Media is typically prepared from commercial dehydrated products through a simple procedure. Liquid
medias are referred to as broths and do not contain the solidifying agent, agar. Semi-solid and solid
media do contain agar and therefore solidify at room temperature. The bottles of dehydrated media
will have instructions on the label specific for preparation. For example, to prepare the general media
known as tryptic soy agar, you would suspend 40g of the medium in one liter of purified water. Heat it
with frequent agitation and boil to completely dissolve the medium. If the medium lacks agar, it will
most likely dissolve without any heat. After dissolving the medium, it must next be sterilized in the
autoclave. The autoclave will sterilize the medium through exposure to steam at 121oC and 15 lbs of
pressure per square inch for 15 minutes. These conditions will kill all forms of life, including endospores.
As solid media types are typically used for the surface growth of microorganisms to observe colony
appearance, for pure culture isolations, storage of cultures or to observe specific biochemical reactions
it is important that the media is not contaminated. After being removed from the autoclave, the agar
will cool down and then is poured into a sterile petri dish while holding the top lid above the bottom
portion, to avoid contamination. In ten minutes or so, the petri dishes will cool and the agar will harden.
To store or incubate the petri dishes, place them in to an inverted position.
Instructions
1.
2.
3.
4.
5.
6.
Turn on the scale, and place a weighing tray on to it then hit tare
Weigh out 10g of the medium and place it in to a flask containing 250ml of distilled water
Drop a stir bar into the flask and place it on to the heater
Boil for 1 minute allowing all of the medium to dissolve completely
Cover top of flask with foil, place in autoclave and run autoclave cycle
Allow agar and temperature of flask to cool to the touch and then pour agar into sterile petri
dishes
Procedure for Autoclaving
1. Pour distilled water in to the water reservoir on top of the autoclave
2. Turn control knob to the FILL position; let water enter the chamber until it reaches the indicator
groove
3. Turn the control knob to the STE position (sterilization mode)
4. Place the flasks in to the autoclave
5. Close autoclave door
6. Move temperature knob to 121oC (250oF)
7. Set timer to 15 minutes
8. Move switch to START position
9. At the end of the cycle, turn control knob to EXH+DRY, make sure the pressure has dropped to
0, then open door and unload flasks using an autoclave glove
Aseptic Technique
Introduction
When working with microorganisms, it is desirable to work with a pure culture. A pure culture is
composed of only one kind of microorganism. Occasionally a mixed culture is used. In a mixed culture,
there are two or more organisms that have distinct characteristics and can be separated easily. In either
situation, the organisms can be identified. When unwanted organisms are introduced into the culture,
they are known as contaminants.
Aseptic technique is a method that prevents the introduction of unwanted organisms into an
environment. When changing wound dressings, aseptic techniques are used to prevent possible
infection. When working with microbial cultures, aseptic techniques are used to prevent introducing
additional organisms into the culture. Microorganisms are everywhere in the environment. When
dealing with microbial cultures it is necessary to handle them in such a way that environmental
organisms do not get introduced into the culture. Microorganisms may be found on surfaces and
floating in air currents. They may fall from objects suspended over a culture or swim in fluids. Aseptic
techniques prevent environmental organisms from entering a culture.
Doors and windows are kept closed in the laboratory to prevent air currents which may cause
microorganisms from surfaces to become airborne. Once these microbes are airborne they are more
likely to get into cultures. Transfer loops and needles are sterilized before and after use in the
Bacticinerator to prevent introduction of unwanted organisms. Agar plates are held in a manner that
minimizes the exposure of the surface to the environment. When removing lids from tubes, lids are held
in the hand and not placed on the countertop during the transfer of materials from one tube to another.
These techniques are the basis of laboratory aseptic techniques.
In this laboratory exercise the location of environmental organisms will be explored and how
microorganisms can be transmitted through contact with contaminated surfaces.
Materials
1 nutrient agar plates per student plus one plate per group
Markers
Instructions
1. Label one plate "Open". Write on the agar containing side of the plate, not on the lid.
Remove the lid from the plate and place it on the lab table, agar side up, until the end of lab.
2. Obtain one nutrient agar plate per student and draw a line on the agar containing side of
the plate to divide the plate in half. Label one side "dirty" and one side "clean."
3. Remove the lid and gently touch your finger tips to the agar on the "dirty" side. Replace the lid.
4. Wash your hands or apply hand sanitizer and gently touch your fingertips to the agar on
the "clean" side of the plate.
5. At the end of the lab period, replace the lid on the 'open' plate. Place all plates, agar side up, in the
incubator.
6. At the next lab period, examine plates for growth and record results in your notebook.
7. Discard all plates in the biohazard container.
Conclusions
1. Describe the growth on the plate labeled "open."
2. Are organisms found in the air? What results support your conclusions?
3. Record your results from the plate you inoculated with your hands. Differentiate between the dirty
side and the clean side.
4. What effect does hand washing have on microorganisms?
Smears
Introduction
The microscopic examination of microorganisms is a valuable identification technique. In order to view
microbes, it is necessary to prepare slides of the organisms. Microscopic preparations may be
either wet mounts or smears. Wet mounts involve placing cells in a drop of water, adding a coverslip
and viewing the material under the microscope. In microbiology most of the organisms viewed are
bacteria which are small and difficult to see without staining. Wet mounts are temporary preparations
and the ability to stain is limited. A smear is a thin preparation of cells allowed to dry on a slide. This
material is then fixed to the slide using heat or a chemical. A smear is a more permanent preparation
and may be stained using a variety of techniques.
Smears are made using plain tap water. While tap water is not sterile, it has too few organisms in it to
interfere with a bacterial smear. At least 500,000 cells per mm must be present in order to see one cell
per oil immersion field. Bacteria are mixed in water and allowed to dry on the slide to make a bacterial
smear. This is then fixed to the slide using heat. Heat fixing helps attach the cells to the slide so they are
not washed off during the staining process, kills the cells so the slide is not hazardous to handle, and
alters the cell wall for staining. The number of cells placed on the slide is important for viewing the cells.
Too few microbes and it is hard to find them on the slide while too many make it difficult to see
individual cells to determine their morphology.
Laboratory Procedure
General Instructions
1. Students work individually.
2. To sterilize an inoculating loop or needle, insert the loop or needle into the Bacticinerator and
observe it. It must glow red for 3 seconds to be sterilized. Loops and needles should never be propped in
the Bacticinerator. The handles are aluminum and will melt. Also they conduct heat readily and can
cause bums if the handles heat up. A hot loop or needle must cool slightly before touching a bacterial
colony to prevent killing the cells.
3. To aseptically remove a lid from a bottle or tube, grasp the lid with the little finger of the dominant
hand. Twist the bottle or tube to loosen and remove the lid. Do not put the lid on the table but keep it in
your hand while removing material from the bottle or tube. Return the lid to the bottle or tube by
turning the bottle or tube to tighten the lid.
Materials/Equipment
Clean glass slides
Prepared mixed of E. coli and M luteus
Inoculating loop
Bacticinerator
Laboratory marker
Instructions
1. Glass slides should be relatively clean and grease free. Slides that do not appear clean may be
washed in soap and water and dried with a paper towel.
2. Use transfer pipette to add a drop of water on your slide if using bacteria from agar plate or slant.
3. Sterilize an inoculating loop.
4. Touch the surface of the loop unto the agar plate containing bacteria #1; gently draw the loop
through to pick up colonies. A little will be enough.
5. Mix the bacteria on the loop with the water on the slide and spread thinly.
6. Sterilize the loop again and remove some bacteria from plate #2.
7. Mix the bacteria with the other already on the slide. Spread thinly once again.
8. Allow to air dry.
9. Heat-fix the slide by passing it 10 times over the top of the Bacticinerator.
10. The slide is now ready for staining or can be stored for later use.
Smear Review Questions
I. Describe 2 preparations that may be used to observe microorganisms.
2. What is the purpose of heat fixing?
3. Outline the procedure for making a smear?
4. Why must slides used in a smear preparation be grease-free?
5. Is it necessary to use sterile water when making a stain? Why or why not?
6. List two reasons for not propping inoculating loops and needles in the Bacticinerator during
sterilization.
7. How long does it take to sterilize an inoculating loop or needle?
8. When removing a lid from a tube or bottle using aseptic technique, what do you do with the lid?
Staining
Introduction
Bacteria have almost the same refractive index as water. This means when you try to view them using a
microscope they appear as faint, gray shapes and are difficult to see. Staining cells makes them easier to
see.
Simple stains use only one dye that stains the cell wall of bacteria much like dying eggs at Easter.
Differential stains use 2 or more stains and categorize cells into groups. Both staining techniques allow
the detection of cell morphology, or shape, but the differential stain provides additional information
concerning the cell. The most common differential stain used in microbiology is the gram stain.
Bacteria have 3 basic shapes. Round cells are known as cocci, rod-shaped cells are bacilli, and spiralshaped cells are spirilla.
Negative Stain: The negative stain is typically used when the bacterium does not stain well or when
observations about bacterial shape need to be made without any heat fixing technique, that may result
in changing the shape of the cells. Negative staining is also known as background staining as the stain is
not absorbed by the cells. The negatively charged bacterial cell wall repels the negatively charged stain,
which therefore produces a dark background so that the unstained cells appear clear and visible. Acidic
dyes such as nigrosin, india ink, or eosin may be used.
Simple Stain: The simple stain consists of one dye. The dye adheres to the cell wall and colors the cell
making it easier to see. Basic dyes such as methylene blue, crystal violet, or carbolfuchsin are typically
used as they bind to the cell by ionic interactions.
Gram Stain: The gram stain is a differential stain. Four different reagents are used and the results are
based on differences in the cell wall of bacteria. Some bacteria have relatively thick cell walls composed
primarily of a carbohydrate known as peptidoglycan. Other bacterial cells have thinner cell walls
composed of peptidoglycan and lipopolysaccharides. Peptidoglycan is not soluble in non polar or organic
solvents such as alcohol or acetone, but lipopolysaccharides are nonpolar and will dissolve in nonpolar
organic solvents.
Crystal violet acts as the primary stain. This stain can also be used as a simple stain because it colors the
cell wall of any bacteria. Gram's iodine acts as a mordant. A mordant is a substance used to set dyes by
forming a complex with the dye. In cells the mordant intensify stains. Gram's iodine also forms a
complex with crystal violet to make a large crystal that is not easily washed out of the cell. At this point
in the staining process, all cells will be the same color. The difference in the cell wall structure is
displayed by the use of the decolorizer (95% ethanol). The decolorizer does not affect those cell walls
composed primarily of peptidoglycan (gram positive) but those with the lipid component will have large
holes develop in the cell wall where the lipid is dissolved away by the acetone. These large holes will
allow the crystal-violet-iodine complex to be washed out of the cell leaving the cell colorless. A counter
stain, safranin, is applied to the cells which will now dye the colorless cells.
The cells that retain the primary stain will appear blue or purple (gram positive) and the cells stained
with the counter-stain will appear pink or red (gram negative). The lipopolysaccharide of the gram
negative cell not only accounts for the staining reaction of the cell but also acts as an endotoxin. This
endotoxin is released when the cell dies and is responsible for the fever and general feeling of malaise
that accompanies a gram negative infection.
The spiral-shaped bacteria of medical importance do not stain well with Gram's iodine and are usually
seen using a dark-field microscope. There are no standard abbreviations of gram stain reactions for the
spirilla.
On your slide you have both gram positive and gram negative bacteria. After staining with the simple
stain, view cells under microscope and write down your observations. Stain the same slide with the
gram stain and observe again. Enter all information in your lab notebook.
Acid-Fast Stain: Microorganisms with containing high lipid content (mycolic acid) in the cell wall are
known as acid-fast. They include species of bacteria in the genera Mycobacterium and Nocardia. They
do not stain well by simple stain methods, however do stain when they are heated and stained with
carbolfuchsin. The heat applied helps to drive the stain in to the cells, where it is not easily removed
from. Acid-fast organisms therefore appear red. Microorganisms that are not acid-fast appear blue.
The primary stain applied to all cells stained by this method is carbolfuchsin and the counterstain
methylene blue. Non-acid-fast microorganisms do not retain the primary stain after being decolorized
with acid-alcohol. The difference in the retention of carbolfuchsin differentiates acid-fast from nonacid-fast microorganisms.
Endospore Stain: Bacteria within the Bacillus and Clostridium genera produce highly resistant structures
capable of surviving in unfavorable conditions, and give rise to new bacterial cells known as endospores.
Endospores do not easily stain, however once stained they do not easily decolorize. Malachite green is
the stain applied with heat that penetrates and stains the endospores. After the use of malachite green,
the cell is decolorized and counterstained with safranin. In the end endospores appear green, while the
cell and other structures appear red.
Procedure – Negative Stain
1. Apply a small amount of bacteria from the broth culture to one end of a microscope slide using
an inoculating loop
2. Add 1 drop of nigrosin to the bacteria and mix
3. Placing a second slide at a 45o angle, spread the bacteria-nigrosin solution across the slide
forming a thin smear
4. Allow the smear to air dry, NO HEAT FIXING REQUIRED
5. Observe under low power objective
Procedure – Simple Stain
1. Place the heat fixed smear over the staining rack and stain with methylene blue for 1 minute
and 30 seconds
2. Wash the stain off the slide with water for a few seconds
3. Blot dry the slide being careful not to rub and remove the cells
4. Examine under the microscope
Procedure – Gram Stain
1. Place the slide on the staining rack and flood the slide with crystal violet for 1 minute.
2. Rinse with tap water, tilting the slide slightly to rinse all the stain from the slide. Tap gently to
remove excess water.
3. Place slide on staining tray and flood with gram stain for 1 minute.
4. Rinse with tap water.
5. Decolorize with 95% ethanol.
6. Rinse with tap water.
7. Stain with safranin for 1 minute.
8. Rinse again with tap water and blot slide gently with bibulous paper
9. Observe under microscope and record observations in your note book.
Procedure – Acid-Fast Stain
1. Place prepared smear above boiling water on the hot plate and cover with a piece of paper
towel cut to the same size as the microscope slide
2. Immediately saturate the paper with carbolfuchsin, adding more stain as the paper towel dries
over a period of 5 minutes, be careful and avoid over flooding
3. Remove the slide using forceps, let it cool and rinse with water for 30 seconds
4. Decolorize by adding acid-alcohol drop by drop until the slide appears only slightly pink (do not
exceed 30 seconds)
5. Rinse with water for 15 seconds
6. Counterstain with methylene blue for 2 minutes
7. Rinse with water for 30 seconds
8. Blot dry using bibulous paper
9. Examine under microscope
Procedure – Endospore Stain
1. Place prepared slide over boiling water on the hot plate, cover the slide with paper towel cut to
the same size and soak the paper with malachite green stain for 5 minutes
2. Apply more malachite green only as needed when it evaporates so the paper remains saturated
during the heating
3. Remove the slide, allow it to cool and rinse with water for 30 seconds
4. Counterstain with safranin for 60-90 seconds
5. Rinse with water for 30 seconds
6. Blot dry with bibulous paper
Review Questions
1. What is the purpose of staining bacteria?
2. List and describe the three basic bacterial shapes.
3. What are the differences between a simple stain and a differential stain?
4. What is the most common differential stain used in microbiology?
5. What is the basis for gram stain results between different bacteria?
6. List the reagents used in the gram stain and describe the function of each.
7. Which dyes or stains listed above are acidic? What is the importance of that?
8. For what diseases would you use an acid-fast stain?
9. What is an endospore?
10. Why is heat necessary to stain the endospores?
Culturing and Isolation Techniques
Introduction
Microorganisms must have a constant nutrient supply if they are to survive. Free-living organisms
acquire nutrients from the environment and parasitic organisms acquire nutrients from their host. When
trying to grow microbes in the lab adequate nutrition must be provided using artificial media. Media
may be liquid (broth) or solid (agar). Any desired nutrients may be incorporated into the broth or agar to
grow bacteria
Agar is the solidifying material used in solid media. It is an extract of seaweed and melts at 100°C and
solidifies at about 42°C. Most pathogenic bacteria prefer to grow at 37oc so agar allows for a solid
medium at incubator temperatures. Since agar remains solid until reaching 100°C, thermophiles that
prefer temperatures above 50°C for growth can still be grown on solid media.
Organisms grown in broth cultures, cause turbidity (cloudiness) in the broth. On agar, masses of cells,
known as colonies, appear after a period of incubation. Certain techniques will allow bacterial cells to be
widely separated on agar so that as the cell divides and produces a visible mass (colony), the colony will
be isolated from other colonies. Since the colony came from a single bacterial cell, all cells in the colony
are clones and are considered pure cultures.
Principle
A mixed culture contains two or more bacterial species that are known and can be easily separated
based on cultural or biochemical characteristics. Culturing techniques provide a means for maintaining
adequate nutrition for the organisms so they can continue to survive. As organisms grow in a
culture, they consume the available nutrients and periodically need to be transferred to fresh media to
continue to grow. Certain culturing techniques not only provide the organisms with a fresh supply of
nutrients but also allow for the separation of bacterial cells to obtain isolated colonies. These culturing
procedures are known as isolation techniques.
Streak plates allow for the growth of isolated colonies on the surface of the agar. An isolated colony is a
colony that is not touching any other colonies and is assumed to be a pure culture. These colonies are
easily accessible for performing staining and identification procedures. They also show colonial
morphology that may be useful in identifying the organism. Part of the identification of any organism
includes a description of colonial morphology. Since organisms may grow differently on different media,
the type of media used must be included as part of any colonial morphology. Other elements of a
colonial description include colony color, hemolysis (if grown on blood agar), form, elevation and
margin. (See Text: Chapter 4, page 92)
Form refers to the overall appearance of the colony. Elevation is the height that the colony achieves on
the surface of the agar. The appearance of the edge of the colony is referred to as the margin.
The pour plate is used for counting organisms in a solution. A standard volume of solution is mixed in
the liquefied agar. Each organism in the solution is separated from all others. When the agar solidifies,
the cells are trapped in the agar and develop into colonies. Each colony can be counted and represents a
single cell in the original solution. If a milliliter of solution is mixed in the agar, then the number of
colonies represents the number of organisms per ml of solution. Usually a portion of a ml is mixed in the
agar so the number of colonies counted must be multiplied by the dilution factor to determine the
number of organisms in a ml of solution.
When counting the colonies in agar it is difficult to accurately count more than 300 colonies on a plate.
Less than 30 colonies on a plate are considered statistically insignificant. When evaluating a solution for
bacteria, a series of dilutions is usually made and cultured. The plate with 30-300 colonies is counted
and the number multiplied by the dilution factor for that plate to determine the number of bacteria/ml
in the original solution. This method is used to evaluate the number of organisms in milk, drinking water,
and even the water at the beach. While the cells grow and are isolated from each other in a pour plate,
they will not develop typical colonial morphology and are not easily accessible for further testing.
Procedure - Inoculation of a Broth Culture
Materials
Mixed Culture in broth
Inoculating loop
Bacticinerator
Incubator
Sterile nutrient broth
Students work individually.
1. Label the sterile nutrient broth with the source of the culture and your initials.
2. Sterilize the loop.
3. Using the appropriate aseptic technique, remove a loopful of broth from the mixed culture
tube.
4. Insert the loop into the sterile broth tube and swirl gently. Sterilize the loop.
5. Incubate the broth at 37°C for 24-48 hours.
6. Observe broth for turbidity. Record results in your lab notebook.
Inoculating an Agar Slant
Materials
Mixed Culture in broth
Inoculating loop
Bacticinerator
Incubator
Sterile nutrient agar slant
Students work individually.
1. Label the sterile nutrient agar slant with the source of the culture and your initials.
2. Sterilize the loop.
3. Using appropriate aseptic technique, remove a loopful of broth from the mixed culture
tube.
4. Insert the loop into the sterile agar slant tube and starting at the base of the slant, draw the loop up
the slant. Do not penetrate the agar. Sterilize the loop.
5. Incubate the slant at 37°C for 24 - 48 hours.
6. Observe the slant for growth. Record results in your lab notebook.
Streak Plate
Materials
Mixed Culture in broth
Inoculating loop
Bacticinerator
Incubator
Sterile nutrient agar plate
Students work individually.
1. Label the sterile nutrient agar plate with the source of the culture and your initials.
2. Sterilize the loop.
3. Using appropriate aseptic technique, remove a loopful of broth from the mixed culture tube.
4. Lift the agar plate from the lid and streak about half of the plate. The loop should be parallel to the
agar surface to prevent digging into or gouging the agar.
5. Return the pate to the lid. Sterilize the loop. Lift the agar plate and make one streak into the
inoculated portion of the plate. Finish by streaking about one-fourth of the uninoculated plate.
6. Return the plate to the lid. Sterilize the loop. Lift the agar plate and make one streak into the second
inoculated portion of the plate. Finish by streaking the remaining one-fourth of the uninoculated plate.
Sterilize the loop.
7. Place the plate in a 37°C incubator for 24 - 48 hours. Observe growth and record your results in your
lab notebook.
Pour Plate
Materials
Mixed Culture in broth
Inoculating loop
Bacticinerator
Incubator
Nutrient agar deep, liquefied
Sterile pipette
2 Sterile petri dish per pair of students
1. Dilute the original sample from the mixed broth culture by transferring 100µl in to Tube 1 which
contains 900µl of saline. The new dilution factor for Tube 1 = 10-1.
2. Take 10µl of culture from Tube 1 and transfer it to Tube 2 which also contains 990µl of saline.
Dilution factor of Tube 2 is now 10-3.
3. Take 10µl of culture from Tube 2 and transfer it to 990 µl of saline in Tube 3. The dilution factor
of Tube 3 is now 10-5.
4. Transfer 1ml of the 10-1 dilution in to a petri dish labeled as number 1, and then add tryptic soy
agar from the water bath; gently mix while the dish is on the table top.
5. Transfer 1ml of the 10-5 dilution in to a petri dish labeled as number 2, and then add tryptic soy
agar from the water bath; gently mix while the dish is on the table top.
6. Allow the plates to cool and solidify.
7. Incubate the plates in an inverted position.
8. Examine results next week.
Review Questions
1. How can you tell growth has occurred in a broth culture?
2. What is the purpose of agar?
3. At what temperature does agar liquefy?
4. At what temperature does agar solidify?
5. Why is liquefied agar cooled to 60o C before adding organisms?
6. List two methods for obtaining isolated colonies.
7. What is the primary purpose of the streak plate?
8. How many colony types did you observe on your streak plate?
9. Describe each colony type observed using standard terms.
10. What is the primary purpose of the pour plate?
11. Was there a difference in the number of colonies from each pour plated dish?
Selective & Differential Media
Organisms can be selected based on specially prepared media called selective or differential media.
These different types of media contain at least one ingredient that favors multiplication of only one type
of bacteria while inhibiting other types. The ingredients are called selective agents and can range from
antibiotics to chemicals or dyes. Bacteria will grow on the media with different characteristics that can
distinguish more than one type of bacterial colony.
Principle:
Mannitol salt agar can differentiate between Staphylococcus aureus and Staphylococcus epidermidis.
Eosin Methylene blue (EMB) agar can be used to differentiate between gram negative
Escherichia coli and Salmonella enteritidis as well as lactose fermenters and nonfermenters.
Mannitol-salts agar contains 7.5% sodium chloride, 0.5% mannitol and phenol red - a pH indicator.
EMB contains 0.065 g/L Methylene blue and 0.4 g/L eosin - a dye that would inhibit gram-positive
bacteria. EMB can also differentiate between gram-negative bacteria that can ferment lactose and
those that cannot. Lactose fermenters will have deep purple colonies, or pinkish colonies with a dark
center. Bacteria that can precipitate eosin will have a metallic green sheen over the purple colony. If the
colonies are pinkish with dark centers, those bacteria can only precipitate a small amount of eosin.
Vogel-Johnson agar also contains the pH indicator phenol-red and mannitol which when fermented
results in a color change to yellow. It is a modified tellurite-glycine agar that is differential for mannitol
fermentation. The tellurite, lithium chloride and high glycine content inhibit the growth of many gram
negative and some gram positive organisms. S. aureus is one organism able to grow on the medium that
produces gray to black colonies from the production of free tellurium, that are surrounded by a yellow
halo due to mannitol fermentation. S. epidermidis will form smaller black colonies with no yellow zone
around due to the inability to ferment mannitol. Brilliant Green Agar is selective for the isolation of
Salmonella species. When grown on the BGA plate, Salmonella colonies appear reddish or pink in color,
whereas E. coli, another gram negative appears green to yellow.
Protocol:
1. Plates of MSA.
a. Cultures of S. aureus and S. epidermidis.
b. Label half of the plate with S. aureus and the other half with S. epidermidis and streak each organism
on to the correct side of the plate, never allowing both to touch.
c. Incubate plates in an inverted position.
d. Next period: record results and discard plates as per instructions.
2. Plates of EMB.
a.
b.
c.
d.
Cultures of E. coli and P. vulgaris.
Streak plate each organism on to half of each plate as before.
Incubate plates in an inverted position.
Next period: record results and discard plates as per instructions.
3. Plates of VJ.
a.
b.
c.
d.
Cultures of S. aureus and M. smeg
Streak plate each organism on to half of each plate
Incubate plates in an inverted position
Next period: record results and discard plates as per instructions
4.Plates of BGA.
a.
b.
c.
d.
Cultures of E. coli and Salmonella
Streak plate each organism on to half of each plate
Incubate plates in an inverted position
Next period: record results and discard plates as per instructions
Questions:
1. How and why are the media selective and/or differential?
Identification of Gram Positive Cocci
Staphylococcus
Introduction
The genus Staphylococcus contains both pathogenic and non-pathogenic organisms. They do not
produce endospores but are highly resistant to drying, especially when associated with organic matter
such as blood, pus, and other tissue fluids. Most staphylococci are found routinely on the surface of the
skin. Breaks in skin and mucous membranes allow entrance of these organisms into the body where
they may cause disease.
The three major species include Staphylococcus aureus, Staphylococcus epidermidis, and Staphylococcus
saprophyticus. The latter two are rarely implicated in disease, but have been isolated in cases of
endocarditis and UTIs under certain circumstances. S. Aureus is considered the pathogenic strain,
causing abscesses, boils, carbuncles, acne and impetigo. Less commonly, are pneumonia, osteomyelitis,
cystitis, pyelonephritis, and food poisoning. These three strains of Staphylococci can be distinguished
from each other by a number of biochemical tests.
Principle
The identification of organisms is based on cellular, cultural and biochemical characteristics.
All species of Staphylococcus are gram positive. On nutrient agar, they tend to be white, circular, convex
colonies. On blood agar, S. Aureus may show hemolysis of the agar in the area around the colony.
Additional biochemical tests that are useful in separating the species include catalase, coagulase, growth
and fermentation of mannitol salt, and resistance or susceptibility to the antibiotic novobiocin.
The catalase test determines if the organism produces the enzyme catalase that breaks down hydrogen
peroxide to water and oxygen. The reaction is:
2H202 à 2H20 + 02
↑
Catalase
This enzyme allows organisms to breakdown harmful metabolites of aerobic respiration and may be
seen in aerobic and facultatively anaerobic organisms. There are other enzymes that some organisms
produce to handle toxic end products of metabolism so not all aerobes or facultative anaerobes
produce catalase.
Pathogenic organisms require mechanisms to help them overcome host defense mechanisms. One
mechanism involves coating the bacterial cells in a body substance, such as fibrin, to fool the immune
system. The coating of a natural body substance will not trigger an immune response. The enzyme
coagulase causes fibrin to be deposited on bacterial cells.
Some organisms cannot tolerate a high osmotic pressure. Media containing higher than normal salt
concentrations may inhibit the growth of these non-tolerant organisms. Mannitol salt agar contains a
high salt concentration so only salt tolerant organisms will grow on it. Additionally, mannitol salt
agar contains the sugar mannitol. Some organisms can utilize mannitol as a food source and will
produce acid end products from this metabolic process. Since this process is invisible, an indicator is
added to the media to detect changes in pH. Phenol red is the indicator used in mannitol salt agar. It is
red at neutral pH but turns yellow if conditions in the media become acidic. Antibiotic susceptibility is
another test that can be used to identify organisms. A filter paper disc is impregnated with an antibiotic,
novobiocin. When the disc is placed on agar, the antibiotic diffuses through the agar. An organism
susceptible to the antibiotic will be unable to grow on the media containing the antibiotic. A zone of
inhibition (no growth) will be seen around the disc. The size of the zone indicates the resistance or
susceptibility of the organism to the antibiotic.
Procedure
Catalase
1. Place a drop of 3% H2 02 on a glass slide.
2. Touch a sterile loop to a culture of the organism to be tested and pick up a visible mass of
cells.
3. Mix the organism in the drop ofH202.
4. Observe for immediate and vigorous bubbling.
5. Dispose of the slide in the contaminated slide container.
Interpretation: Bubbling indicates a positive test and scant or no bubbling indicates a negative test.
Coagulase
1. Dispense 1 drop of Test Latex onto one of the circles on the reaction card and 1 drop of
Control Latex onto another circle.
2. Touch a sterile loop to a culture of the organism to be tested and pick up a visible mass of cells. Mix
the cells in the drop of Test Latex.
3. Repeat step 2 for the Control Latex.
4. Pick up and hand rock the card for up to 20 seconds and observe for agglutination or clumping of
the latex particles.
5. Dispose of the reaction card in the biohazard container.
Interpretation: Agglutination of the Test Latex with no agglutination of the Control Latex is considered a
positive test for coagulase. No agglutination in either the Test Latex or Control Latex is considered
negative for coagulase. All reactions occurring after 20 seconds should be ignored. If agglutination
occurs in the Control Latex the agglutination is due to some factor other than the enzyme coagulase and
the test results are invalid.
Mannitol Salt Agar
1. Label a tube of mannitol salt agar with the organism to be tested and your initials.
2. Using a sterile loop transfer the organism to be tested to the surface of the mannitol salt agar slant.
3. Incubate the tube at 35°C for a minimum of 18 hours.
4. Examine the tube for evidence of growth on the slant and for a color change from red to yellow.
5. Remove the markings from the tube using EtOH on a paper towel and place the tube in
the designated area for disposal.
Interpretation: Two different characteristics of the organism are determined with this agar. The first is
the organism's ability to tolerate a high salt environment. Organisms that can ferment the sugar
mannitol produce an acid end product that changes the red pH indicator in the media to yellow. Any
yellow in the media is considered a positive test for mannitol fermentation. It is possible for organisms
to grow on the media and not ferment mannitol.
Novobiocin Susceptibility
I. Divide. a nutrient agar plate into three sections.
2. Label a section with the name of the organism to be tested.
3. Using a sterile loop transfer the test organism to the plate and streak the section for
confluent growth.
4. Aseptically transfer a novobiocin antibiotic disc to the center of each streaked area.
Gently press the disc to the surface of the agar.
5. Invert the plate and place in the incubator for a minimum of 18 hours.
6. Examine the plate for a zone of inhibition of growth around the antibiotic disc.
7. Using a metric ruler, measure the diameter of the zone of inhibition and record the measurement in
mm.
8. Discard the plate in the biohazard container.
Interpretation: A zone of growth inhibition of 17 mm or less in diameter indicates resistance to
novobiocin. If the zone is greater than 17 mm, the organism is susceptible to novobiocin.
Instructions
Cultures provided: Staphylococcus aureus
Staphylococcus epidermidis
Staphylococcus saprophyticus
Students work individually unless otherwise noted.
I. Make a smear of one of the organisms provided. Complete the remainder of the laboratory work
before heat fixing, staining and examining the smear.
2. Perform a catalase test on all organisms.
3. Select one of the organisms and perform a coagulase test. Allow the other members of
your group to observe your results. Observe the results of the other 2 organisms.
4. Select one of the organisms and inoculate a mannitol salt agar slant. As in step 3, observe the results
of all three organisms.
5. Test each organism for novobiocin susceptibility. Each person should test all three organisms.
6. Record all results in your notebook.
7. Organisms can be gram stained if time permits.
Review questions
1. Which test differentiates S. aureus from the other species of Staphylococcus?
2. How can you differentiate S. epidermidis from S. saprophyticus?
Identification of Gram Positive Cocci
Streptococcus
Introduction
Members of the genus Streptococcus are responsible for disease as well as being part of the normal flora
of humans. Among the diseases caused are bacterial pneumonia, meningitis, tonsillitis, endocarditis,
scarlet fever, erysipelas, and UTIs. Streptococcus species are also found normally in the mouth and on
the surface of the skin. Streptococci are classified by two major methods: hemolytic activity and
serologic classification of Lancefield.
Classification Based on Hemolytic Activity
When grown on sheep blood agar, streptococci display one of three types of hemolysis of the red blood
cells in the agar:
Alpha hemolysis (α-hemolysis) - the red blood cells in the media are partially digested producing
a greenish color in the agar.
Beta hemolysis (β-hemolysis) - the red blood cells in the media are completely digested producing
a clear ring around the colony in the agar.
Gamma hemolysis (λ-hemolysis) -no change is noted in the agar. The red blood cells are not affected by
the organism.
Expected Hemolysis
Organism
S. pyogenes
S. agalactiae
S. bovis
S. pneumoniae
S. faecalis
α
Never
Never
Sometimes
Always
Sometimes
β
Always
Usually
Sometimes
Never
Sometimes
λ
Never
Sometimes
Usually
Never
Usually
Classification Based on Lancefield Proteins
Rebecca Lancefield, working with various streptococcal species, discovered proteins in the cell wall that
were unique to certain organisms. These proteins were labeled Group A, Group B, Group C, and so
on through Group M. Currently three Lancefield Groups are of medical importance: Groups A, B
and D. Of the organisms used in this lab the following correlations apply:
Group A Strep--Streptococcus pyogenes
Group B Strep--Streptococcus agalactiae
Group D Strep--Streptococcus bovis, Enterococcus (Streptococcus) faecalis.
Streptococcus pneumonia does not possess Lancefield proteins and is not classified in one of the
Lancefield groups. Viridans streptococci is the term applied to α -hemolytic Streptococcus species
that lack Lancefield proteins. (Viridans are from a large group most abundant in mouth: S. mutans)
Principle
All Streptococcus species are gram positive cocci. Some will only grow on an enriched agar such as 5%
sheep blood agar. On sheep blood agar, the colonies are usually gray, punctiform, convex, and entire.
Various species display alpha, beta or gamma hemolysis. Important biochemical tests include catalase,
bacitracin susceptibility, optochin susceptibility, growth in high salt broth, hemolysis patterns seen with
the CAMP test, and the ability to hydrolyze esculin.
The bacitracin and optochin susceptibility are similar to the novobiocin susceptibility test used for the
identification of Staphylococcus species. Filter paper discs impregnated with the appropriate chemical
are placed on an agar surface. The chemical diffuses through the agar. Organisms that are susceptible to
the chemical will not grow on the agar containing the chemical. The size of the zone of growth inhibition
determines the organism's susceptibility to the chemical.
CAMP factor is a diffusible protein produced by cetiain species of Streptococcus. This factor will react
with the beta toxin produced by S. aureus to rapidly lyse sheep red blood cells. When a CAMP producing
Streptococcus is grown near a toxin producing strain of S. aureus, a definite hemolytic pattern is
produced.
Only a few organisms can tolerate a salt concentration of 6.5% NaCl. Those that can will grow in high salt
broth.
Bile esculin agar contains bile that inhibits the growth of many organisms. Some organisms can
hydrolyze esculin to esculetin and dextrose. Esculetin will react with ferric citrate in the media to
produce a black-brown product.
Procedures
Bacitracin Susceptibility
1. Divide a sheep blood agar plate into four quadrants.
2. Label a quadrant with the name of the organism to be tested.
3. Using a sterile loop, aseptically transfer the test organism to the plate and streak the quadrant for
confluent growth.
4. Aseptically transfer a bacitracin disc to the center of the quadrant. Forceps may be used
to position the disc. Gently press the disc to the surface of the agar but to not embed the disc in the
agar.
5. Invert the plate and place in the incubator for a minimum of 18 hours.
6. Examine the plate for a zone of inhibition of growth around the disc. When finished, discard the plate
in the biohazard container.
Interpretation: Any zone of inhibition of growth is considered positive for this test. If a red ring can be
seen around the disc, then this is considered a negative test. This test should be done only or organisms
that display beta-hemolysis.
Optochin Susceptibility
1. Divide a sheep blood agar plate into four quadrants.
2. Label a quadrant with the name of the organism to be tested.
3. Using a sterile loop aseptically transfer the test organism to the plate and streak the quadrant for
confluent growth.
4. Aseptically transfer an optochin disc to the center of the quadrant. Forceps may be used to position
the disc. Gently press the disc to the surface of the agar but do not embed the disc in the agar.
5. Invert the plate and place in the incubator for a minimum of 18 hours.
6. Examine the plate for a zone of inhibition of growth around the disc. Using a metric ruler, measure
the diameter of the zone of inhibition and record the measurement in mm. When finished, discard the
plate in the biohazard container.
Interpretation: a growth inhibition zone of 15- 30 mm is considered a positive test. Zone sizes of
less than 15 mm are considered negative for this test. This test should be done only on organisms
that display alpha-hemolysis.
CAMP Test
l. Obtain a sheep blood agar plate that has been prepared for a CAMP test by having Staph aureus
streaked in a single line down the center of the plate.
2. Lines have been drawn on the plate perpendicular to the Staph streak. These will act as guidelines
for inoculating the plate. Label one of the lines on the CAMP plate with the organism to be tested.
3. Using a sterile loop obtain a sample of the test organism. Using a single streak and moving from
the outer edge of the CAMP plate toward the Staph streak, inoculate the plate with the test organism.
Do not allow the test organism to directly touch the Staph streak, or streak across the Staph streak. The
test organism should be streaked using one of the perpendicular lines as a guide.
4. Invert the plate and place it in the incubator for a minimum of 18 hours.
5. Observe the plate for the development of a distinct arrowhead pattern of hemolysis where the test
organism and the Staph almost touch.
6. Discard the plate in the biohazard container.
Bile Esculin
1. Label a bile esculin slant with the organism to be tested and your initials.
2. Using a sterile loop, transfer the organism to be tested to the surface of the bile esculin slant.
3. Incubate the tube for a minimum of 18 hours.
4. Examine the tube for a definite blackening of the agar.
5. Remove the markings from the tube using EtOH on a paper towel and place the tube in the
designated area.
Interpretation: Blackening of the agar is considered positive for this test. No change in the color of the
agar is considered negative. This test should be done on all suspected Streptococci.
High Salt
1. Label a high salt broth tube with the organism to be tested and your initials.
2. Using a sterile loop transfer the organism to be tested to the broth.
3. Incubate the tube for a minimum of 18 hours.
4. Examine the tube for evidence of growth (turbidity). It may be helpful to compare the tube to an
uninoculated tube. Do not agitate the tubes before you examine them.
5. Remove the markings from the tube using EtOH on a paper towel and place the tube in the
designated area.
Interpretation: Organisms that can tolerate a high salt environment (6.5%) NaCl) will grow in this broth
causing the broth to become cloudy or turbid. Turbidity is considered positive for this test. Organisms
that cannot tolerate the high salt environment will not grow and the broth will remain clear. Clear broth
is considered negative. This test should be done on all suspected streptococci.
Instructions
Cultures provided: Streptococcus pyogenes
Streptococcus agalactiae
Streptococcus pneumoniae
Enterococcus (Streptococcus) faecalis
Streptococcus bovis
Students work individually unless otherwise noted.
I. Make a smear of one of the organisms provided. Complete the remainder of the lab work before heat
fixing, staining and examining the smear.
2. Perform a catalase test on all organisms and record your results in your notebook.
3. Examine all cultures for hemolysis and record your observations in your notebook.
4. Refer to your Jab notebook and on all beta-hemolytic organisms, set up a Bacitracin
Susceptibility test.
5. Refer to your notebook and set up an Optochin Susceptibility test on all alpha-hemolytic orgamsms.
6. Refer to your lab notebook and set up a CAMP test on all beta and gamma organisms tested. Be sure
each member of the group sets up one test. Organisms may be used more than once, if necessary.
7. Working in groups, set up a bile esculin slant on all organisms. Each member of the group must set
up at least one test.
8. Working in groups, set up a high salt broth on all organisms. Each member of the group must set up
at least one test.
9. After appropriate incubation, examine all tests and record results.
10. If time permits, gram stain the smear prepared in Step 1.
Identification of Streptococcus
Organism
Gram
Catalase
Bacitracin Optochin
CAMP
Bile esculin High Salt
S. pyogenes
S. agalactiae
S. pneumoniae
E. faecalis
S. bovis
If a test is not done on an organism because it is an inappropriate test for that organism, mark the
results box with an X.
Review Questions
1. What characteristic do Staphylococcus and Streptococcus share?
2. What test would distinguish Staphylococcus from Streptococcus?
3. An organism is GPC, catalase negative, and alpha hemolytic. List all appropriate tests for
identification of this organism.
4. An organism is GPC, catalase negative, and beta hemolytic. List all appropriate tests for identification
of this organism.
5. An organism is GPC, catalase negative, and gamma hemolytic. List all appropriate tests for
identification of this organism.
6. Once you know an organism is GPC, what test should you do next?
Throat Culture
Introduction
The human mouth has numerous and varied organisms as part of its normal flora. Both aerobes and
anaerobes flourish in this warm, moist enviromnent. Virtually every type of microorganism can be
found in the mouth. The most prevalent are the viridians Streptococci. These α -hemolytic organisms
account for most of the organisms that grow aerobically in a throat culture. In addition to these gram
positive cocci, numerous species of Staphylococcus may also be found. Neisseria, Branhamella and the
anaerobic Veillonella, comprise the majority of gram negative cocci found in the mouth. Various gram
negative bacilli, such as Haemophilus species and Klebsiella pneumoniae, are also present. The
nonpathogenic Corynebacterium, or dipththeroids, are also α -hemolytic. Dipththeroids are
pleomorphic gram positive bacilli. Spirochetes, a few yeasts and occasional protozoa round out the
normal mouth flora. These organisms are commensals that probably protect us from other organisms
that may enter our mouths. The presence of our normal flora prevents other organisms from finding
space or nutrients to support their growth.
While our normal flora potentially protects us from certain diseases, they do contribute to one. The
organisms of the mouth contribute to the development of dental caries. Certain organisms adhere to
the teeth forming a network for others to adhere. These organisms produce the plaque found on your
teeth (biofilm). Some of the organisms involved in plaque metabolize sugars found in the mouth to acids
that etch the tooth enamel and weaken it. If the tooth enamel is damaged, organisms can penetrate to
the pulp of the tooth damaging it. Regular removal of these organisms and plaque helps prevent tooth
decay.
Principle
The one organism responsible for disease in the throat is Streptococcus pyogenes or Group A Strep. This
organism is beta hemolytic and not part of the normal throat flora. Sheep blood agar provides the
enrichment necessary for growing many of the Streptococcus species and also acts as a differential
media. The hemolysis produced on sheep blood agar helps separate the normal α -hemolytic organisms
from the pathogenic, β -hemolytic Streptococcus pyogenes.
Organisms that grow in the throat also need special atmospheric conditions to grow. These organisms
are exposed to the higher carbon dioxide content in exhaled breath. To successfully grow these
organisms this C02 rich atmosphere must be reproduced. Organisms that require less oxygen are known
as microaerophiles. In the lab this atmosphere may be produced by placing the plates in a large jar,
lighting a candle in the jar and replacing the lid. As the candle burns, some of the oxygen in the jar is
converted to C02.
Typically, pharyngitis would cause redness and possibly pockets of pus on the back of the throat. When
culturing a throat, these areas indicating inflammation should be swabbed to provide the specimen.
Usually a swab in a protective plastic sleeve is used to take a throat culture. Once the specimen has
been taken, the swab is returned to its protective sleeve and an ampule of transport media is broken in
the bottom of the sleeve. Transport media is a special purpose media that contains balanced salts to
protect the specimen from pH changes and keep the swab moist while in transit to the lab for culturing.
Nutrients are not provided so growth does not occur but the organisms can survive for several hours in
the transport media, particularly if refrigerated.
Procedure
1. Obtain a sheep blood agar plate, sterile swab and tongue depressor.
2. Label the agar plate with your "patient's" name.
3. Using the tongue depressor, flatten the patient's tongue. Having the patient say "Ahhh" helps
flatten the tongue. Being careful not to touch any other parts of the mouth, use the sterile swab to
firmly swab the back of the patient's throat. Use care. Some people have a very strong gag reflex and
this may induce vomiting.
4. Gently roll the swab across the surface of the blood agar plate. Using a sterile loop, streak
the plate for isolation. First streak through the area where you rolled the swab and cover approximately
half of the plate. Sterilize the loop and streak one quarter of the plate streaking into the original only
once. Repeat the procedure for the remaining quarter of the plate, streaking into the second streak
only once. Discard the swab and tongue depressor in the biohazard container.
5. Place the plate in a candle jar. The jar will be incubated at 35-37°C for a minimum of 18
hours.
6. Following incubation, examine the plate for the presence of β-hemolytic colonies. A predominance of
β -hemolytic colonies would indicate a possible throat infection with Streptococcus pyogenes.
7. When finished with the plate, discard in the biohazard container.
Review Questions
1. List 3 organisms that are considered normal throat flora.
2. Why is sheep blood agar used for throat cultures?
3. What organism is pathogenic in the throat?
4. What incubation conditions are required for throat cultures?
5. What is the purpose of the candle jar?
6. Define microaerophile.
7. What are viridans streptococci?
8. What are the most predominant aerobic organisms in the throat?
Skin Flora Experiment
With the possible exceptions of those areas on the human skin that contains numerous hair follicles and
sebaceous and sweat glands, the skin is a relatively dry environment that does not provide favorable
conditions for growth of most microbes. Nutrients are limited, the concentration of salt may slow
growth and the pH of skin secretions are between 4 - 6 which also discourages bacterial growth. Sebum
and sweat can provide growth of normal flora due to lipid and amino acid content that provide
nutrients. Normal flora of human skin is primarily gram positive and includes micrococci and
staphylococci. Corynebacteria and propioni bacteria as well as yeasts are also found on skin.
Materials
Blood agar
Mannitol salt agar with phenol red indicator
Sterile diluent
Procedure
1. Moisten a sterile swab in the diluents provided. Squeeze out the excess fluid by pressing the swab
against the inner surface of the tube.
2. Rub the swab on a portion of skin (between the fingers; in the area between the nose and cheek).
3. Streak the swab over a quadrant of a blood agar plate, carefully rolling the swab so that the entire
cotton surface comes in contact with the medium. Discard the swab.
4. Using a sterile inoculating loop, streak for isolated colonies.
5. Repeat with the mannitol salt agar.
6. Incubate for 24 hrs.
7. Examine the blood agar plates for hemolytic activity (clear zones around colonies).
8. Examine the mannitol salt agar for staphylococci (yellow zones around colonies).
9. Perform gram stains on the predominant colonies
Questions:
1.
Why is the skin a poor enviromnent for the growth of most microbes?
2. What are the predominant bacterial genera found on the skin?
3. Where on the skin are these organisms most commonly found?
4. What are some skin diseases caused by the normal flora?
Minimum Inhibitory Concentration
Introduction
The minimum inhibitory concentration is the lowest concentration of an antibiotic that inhibits the
visible growth of a microorganism after 18-24 hours of incubation. It is important in labs to analyze the
sensitivity of a bacterial strain to an antibiotic to confirm resistance or compare the effectiveness
against other antimicrobial agents to lower the chance of microbial resistance. In clinical application,
the minimum inhibitory concentration may also be used to determine the amount of antibiotic that a
patient will receive to effectively treat the infection.
In the experiment we will determine the sensitivity of E. coli and S. aureus to the antibiotic kanamycin.
The cells are prepared and grown overnight to the log phase of approximately 1 x 108 cells/mL, as a
barely cloudy sample from overnight incubation will show about one hundred million bacteria per
milliliter. The cells are further diluted in Mueller Hinton medium by transferring 100 µl to 9.9 ml of
Mueller Hinton broth which equals to 105 cells in each tube. For samples 3-9 in the chart the
concentration of the stock antibiotic will be 1g/L. Sample 1 is a control to make sure the Mueller Hinton
broth prepared is not contaminated, and sample 2 a control showing the growth of cells in the media.
Procedure
1. Label tubes then using a micropipette transfer appropriate amounts of MH broth, kanamycin,
and cells as listed in the chart below.
Tube
MH (µl)
Kanamycin (µl)
Cells (µl)
1
1000
0
0
2
900
0
100
3
890
10
100
4
880
20
100
5
870
30
100
6
860
40
100
7
850
50
100
8
840
60
100
9
830
70
100
2. After all additions mix and incubate for 16-20 hours at 37 degrees Celsius.
3. Determine which tube containing the antibiotic has no growth and record your results.
Identification of Gram Negative Bacilli
Introduction - Oxidase
The oxidase test can be used in the identification of GNB to distinguish non-fermenters (oxidase
positive) from fermenters (oxidase negative).
The oxidase test checks for the presence of the enzyme indophenols oxidase. Tetramethylpara-phenylenediamine (oxidase reagent) will be oxidized in the presence of atmospheric oxygen by
indophenols oxidase causing the formation of a dark-purple compound known as indophenols.
Procedure & Organisms Used:
Pseudomonas aeruginosa
E. coli
Proteus vulgaris
Students work in groups.
1. Obtain a sterile swab. Touch the swab to the organism being tested.
2. Place one drop of oxidase reagent on the organism on the swab. Using more than one drop of
reagent may dilute the color reaction and result in a false negative.
3. Observe the swab for 10-30 seconds for the development of a dark-purple color around the edge of
the organism. This is interpreted as a positive test. No color change or a color change after 30 seconds in
interpreted as a negative test.
4. Share your results with the other members of your group and record them in your lab
notebook.
5. Dispose of the swabs in the biohazard container. The reagent droppers may be discarded in the
regular trash.
Review Questions
1. Based on your results, which organism(s) could be classified as a nonfermenter?
2. E. coli and Proteus vulgaris are members of the family Enterobacteriaceae so their reactions
are representative of the entire family. Klebsiella pneumonia is also a member of the family
Enterobacteriaceae. What would its oxidase test result be? Is it a fermenter or non-fermenter?
Introduction - Urea
The urea test can be used in the identification of GNB, particularly those in the family
Enterobacteriaceae.
If an organism produces the enzyme urease, it will break down urea to ammonia and CO2.
CO(NH2)2 à NH3 + C02
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Urease
Ammonia will increase the pH of the media to 8.0 or higher. The media contains phenol red as a pH
indicator. At pH 8.0 or higher, the indicator is a bright pink color. If urea is split to NH3 and C02, the pH
change will cause the media to turn a bright pink and the test will be considered positive for urease.
Procedure & Organisms Used:
Pseudomonas aeruginosa
E. coli
Proteus vulgaris
Work in groups to complete this lab.
1. Urea media may either be broth or slant. Obtain media and inoculate tubes with the three organisms
listed above. Use appropriate aseptic technique when inoculating the tubes.
2. Incubate the tubes for a minimum of 18 hours at 35-37°C.
3. Examine the tubes for a color change. Tubes that are bright pink are considered positive for the test.
Any other color change is considered negative. Remove labels from the tubes and discard in the
designated area.
4. Record results in your lab notebook
Review questions
1. What enzyme is produced by organisms that can split urea?
2. What is the indicator used in urea media?
3. Why does the media tum pink when the test is positive?
Introduction - TSI
The TSI (triple sugar iron) agar provides information concerning glucose fermentation, utilization of the
sugars lactose and sucrose, and the anaerobic respiratory process that uses sulfur as the final
electron acceptor to produce hydrogen sulfide (H2S). This information is useful in the identification
of gram negative bacilli.
TSI agar contains three sugars: glucose (0.1%), lactose (1.0%), and sucrose (1.0%). It
also contains phenol red to indicate a change in pH and ferrous sulfate to demonstrate H2S
production.
Sugar Fermentation:
Fermentation is an anaerobic process. When sugar is fermented, an acid end product is produced and
sometimes gas. Phenol red turns yellow under acidic conditions and red under alkaline conditions.
Yellow agar ---> acid production ---> sugar fermentation.
The enzymes for glucose fermentation are constitutive enzymes so glucose is the first choice of an
organism for fermentation. The acid produced will turn the agar in the tube yellow. Some organisms also
produce gas from glucose fermentation. This gas may be trapped in the agar pushing the agar up in the
tube or causing cracks or bubbles in the tube. It is possible for the gas to escape around the agar and not
be detected.
If only glucose can be used, the organism quickly uses the available glucose in the tube. In order to
survive, the organism will use the protein in the agar as a carbon source. The first step of protein
utilization is deamination which will cause NH3 to form. Deamination is an aerobic process and will only
occur on the slant. Those organisms that can only ferment glucose will deaminate proteins to obtain
nutrients and survive. The slant will revert to red due to the alkalinity of the media.
Organisms that can use either sucrose or lactose (or both) will begin to ferment these sugars once the
glucose has been consumed. The enzymes required for utilization of these sugars are inducible so the
presence of sucrose and lactose in the media will activate the necessary operon. Acid will continue to
be produced as a result of the metabolism of the lactose and/or sucrose and the tube will remain
yellow. There is a sufficient quantity of either sugars in the media to support the organism for at least 23 days.
Hydrogen Sulfide (H,S) Production
Anaerobic respiration does not require oxygen since an inorganic salt acts as the final electron acceptor
instead of oxygen. Sulfur is one of the anaerobic electron acceptors used by some facultative and
obligate anaerobes. Sulfur is readily available in the environment and in media in both organic (amino
acids) and inorganic (sulfates) molecules.
H2 S is a colorless, volatile liquid. In order to detect its presence in the media, ferrous sulfate is used as
an indicator H2S + FeS04 --+ FeS (black precipitate)
H2S production is an anaerobic process so the black precipitate will appear only in the butt of the TSI
tube.
Procedure & Organisms Used:
Pseudomonas aeruginosa
E. coli
Proteus vulgaris
Students work in groups to complete this exercise
1. Label a TSI slant with one of the organisms to be tested.
2. All tests in this media rely on anaerobic conditions. To provide this, the organism must be introduced
into the media, not on the surface. Using a sterile inoculating needle, tough the organism to be tested
and stab the TSI media penetrating to the bottom of the tube. When removing the needle, streak the
slant.
3. Incubate tube for at least 18 hours at 35-37°C.
4. After incubation, examine the tubes for color changes. Record all results m your notebook.
5. When finished, remove all labels and markings and place in the designated area.
Review Questions
1. Why is the media stabbed when inoculating it?
2. Why does the slant tum red if only glucose is fermented?
3. Which organism(s) produced gas? How could you tell?
4. Which organism(s) produced H,S? How could you tell?
5. What is the interpretation of the results for Pseudomonas aeruginosa?
6. What is the interpretation of the results for E. coli?
7. What is the interpretation of the results for Proteus vulgaris?
8. Do the TSI fermentation results match the oxidase results?
Motility
Introduction
Bacteria have a single strand protein for a flagellum. The flagellum is the only organelle for motility in
prokaryotic cells. Eukaryotic cells may move by using flagella, cilia, or pseudo pods. Motility in bacteria
indicates that the organism has flagella.
Bacterial flagella may be stained using a special flagellar stain to demonstrate their presence. This
procedure is somewhat tedious. An alternate way to show bacteria have flagella is to demonstrate their
ability to move. Since the only organelle for motility in bacteria is the flagellum, movement indicates the
presence of flagella. Media that contains half the agar content is semisolid. It does not pour but is too
soft to produce a slant. This consistency will allow bacteria to swim through the agar from the initial
inoculation point if they possess flagella. Cells will distribute along the migration route and will cause
the media to become cloudy so the trail will be visible. If a tetrazolium salt (triphenyltetrazolium
chloride or TTC) is added to the medium, that bacterial presence in the medium will be much easier to
see. TTC is colorless and soluble in the oxidized form but becomes insoluble and turns red when
reduced. Sin any metabolic process involves the oxidation of molecules to produce energy, the dye is
readily reduced by microbial growth and other microbial activities, such as motility. When TTC is
present in the media, the cloudy trail left by bacteria swimming through the media is red. The semi-solid
agar technique is the most common test for motility in the clinical lab.
Motility may also be demonstrated using the hanging drop method. A drop containing bacteria is
suspended from a cover slip using a depression slide. The slide is then examined to see if the organisms
are moving directionally for a distance of2-3 cell lengths. This is considered true motility. Due to the size
of the bacterial cell it is possible to see Brownian movement if the cells are nonmotile. This movement
is the result of molecular bombardment of the cells and is not due to the presence of flagella.
Procedure & Organisms Used
E. coli
Klebsiella pneumoniae
Enterobacter cloacae
Students work in groups to complete this exercise.
1. Obtain a tube of motility media. Using a sterile inoculating needle, inoculate the motility media by
stabbing halfway into the agar.
2. Incubate for a minimum of 18 hours at 35°C.
3. After incubation, examine the stab line. If the line is sharp the organism did not move and there is no
motility. If the line is fuzzy, shows a cloud of growth around it, or is indistinct in any way, the organism
was able to move away from the initial stab line and is motile.
4. Record all results. Remove all marks from the tube and discard in the designated area.
Review Questions
1. What is Brownian movement? Is it motility?
2. What organelle(s) for motility do bacteria posses?
3. List tluee methods that may be used to demonstrate flagella in bacteria.
Hydrolysis of Polysaccharides
The polysaccharide starch is composed of the monosaccharide glucose joined together in chains. Starch
is a compound found plants and serves as an environment for the colonization of many microorganisms.
Hydrolysis of starch is catalyzed by the enzyme amylase to .form maltose and glucose. Maltose is a
disaccharide of two glucose molecules which would be further broken down to glucose by the enzyme
maltase. Glucose is used by cells as both carbon and energy sources.
Starch reacts with iodine to give a blue-black color. However if starch has been broken down into
maltose or glucose, iodine can no longer react and the solution will remain an orange color (the color of
iodine). If bacteria produce amylase, they will hydrolyze starch if grown on starch agar. To test for
amylase production, a few drops of an iodine solution are added to the agar plate. If starch has been
hydrolyzed the area of bacterial growth will be colorless. If no starch has been hydrolyzed the area will
be blue-black in color.
Materials:
E. coli and B. subtilis
Starch agar plates.
Spot inoculate the starch agar plate with both cultures in a small enough area so the two bacteria do not
mix.
Incubate.
Next period: Use only the amount of iodine solution needed to barely cover the surface of the agar. Add
iodine to the area immediately surrounding bacterial growth. Measure the zone of hydrolysis or non
hydrolysis and record results.
Questions:
1. What determines the size of the zone of hydrolysis?
2. Name some foods that would be spoiled by starch hydrolyzing bacteria.
Deamination of amino acids
Amino acids are recycled by organisms for use in other processes if necessary. Before synthesis, amino
acids must be deaminated (removal of the amine group). The removal of the amine group is catalyzed by
deaminases. The phenylalanine deaminase test can be used to differentiate among different bacteria.
Deamination of phenylalanine produces phenylpyruvic acid, which reacts with ferric chloride (FeCI3) to
form a green-colored compound. If bacteria is grown on agar containing phenylalanine, a green
color will be develop if a few drops of FeCl3 is added. This indicates that phenylalanine has been
deaminated.
Materials
E. coli and P. vulgaris
Phenylalanine agar slants.
10% aqueous solution of ferric chloride.
Label 2 tubes of phenylalanine agar with the orgamsms. Inoculate the slants with the
corresponding organism.
Next period: Add 5- 10 drops of 10% FeCI3 to each of the 2 cultures and tap the sides of the tube to mix.
Observe for the development of a green color and record results.
Questions:
I. Cite three ways in which phenylalanine can be used by E. coli.
2. Name three ways in which amino acids can be used by E. coli.
Identification of Gram Negative Bacilli-IMViC
Introduction
IMViC is a mnemonic to remember the four biochemical tests being used. Indole, Methyl red, VogesProskauer and Citrate. These four tests help divide the Entero-bacteriaceae into two major groups- the
E. coli group and the Enterobacter-Klebsiella group.
Indole
Organisms that possess the enzyme tryptohanase can break down the amino acid tryptophan to
indole. When indole reacts with para-dimethylarninobenzaldehyde (Kovac's reagent) a pink-colored
complex is produced. Tryptophan is plentiful in most media, but growth on blood agar or chocolate agar
produces the best effects.
Methyl Red
Some organisms produce acid from the metabolism of glucose in a sufficient enough quantity to
produce a pH of 4.4 in the media. These acids are not further metabolized and are said to be stable
acids. At a pH of 4.4 or less, the pH indicator, methyl red, is a bright cherry red.
Voges-Proskauer
Some organisms initially produce acid from glucose metabolism but further metabolize the acid
produced to neutral end products such as acetoin and 2,3-butanediol. Initially the pH may drop to 4.4
but the neutral end products raise the pH so the methyl red test will be negative. Acetoin and 2,3butanediol, under alkaline conditions, will react with alpha-napththol (!- naphthol) to produce a
mahogany red color.
Citrate
Citrate contains carbon. If an organism can use citrate as its only source of carbon, the citrate in the
media will be metabolized. Bromothymol blue is incorporated into the media as an indicator. Under
alkaline conditions, this indicator turns from green to blue. The utilization of citrate in the media
releases alkaline bicarbonate ions that cause the media pH to increase above 7.4
Procedure & Organisms Used:
E. coli
Klebsiella pneumoniae
Enterobacter cloacae
Students work in groups to complete this exercise.
1. Obtain a Dry Slide indole test card. Using a sterile loop, transfer cells from an agar plate or slant to
the test area on the card. Observe for the development of a pink color within 30 seconds. Record your
results in your notebook and discard the test card in the biohazard container.
2. Obtain an MRVP broth and using aseptic technique, inoculate the tube. It is important to inoculate
this test heavily. Incubate for at least 24 hours at 35°C. (COMPLETED AND PREPARED FOR YOU ALREADY)
3. Take the incubated MRVP broth, obtain a spot plate and a sterile dropper. Observe the MRVP for
turbidity. If turbidity is not noted the test results are not reliable. The tube may be re-incubated until
growth is evident. Place 3 drops of turbid broth into two of the wells on the spot plate.
Methyl Red Test-To one well add 1-2 drops of methyl red reagent. Observe for an immediate cherry red
color that indicates a positive test. Orange or yellow is considered negative. Record your results in your
notebook.
Voges-Proskauer Test-To the remaining well add 2 drops of alphanaphthol and one drop of potassium
hydroxide (KOH). Observe the development of a mahogany red color. The color development takes 20
minutes or longer. Be extremely careful with the KOH. It is caustic and may cause bums if it gets on your
skin. The mahogany red color is considered positive. Record your results in your notebook. Remove
all marks from the MRVP tube and discard in the designated area. Clean the spot plate by
covering the surface with disinfectant. Allow the disinfectant to sit for a few minutes, then rinse with
water, wash and dry.
4. Obtain a citrate slant. Aseptically inoculate the slant and incubate for at least 24 hours at 35°C.
5. After incubation, observe the citrate for a change from green to blue. Blue is considered positive.
Record your results in your notebook.
Review Questions
1. What is the lMViC pattern for the E. coli group?
2. What is the IMViC pattern for the Enterobacter-Klebsiella group?
3. What is the difference between a reagent and an indicator?
4. Is it possible for an organism to be Methyl red and Voges-Proskauer positive?
Explain your answer.
5. What might happen if the MRVP tests are read too soon?
Disc Diffusion Susceptibility Methods
Introduction
When a filter paper disc impregnated with a chemical is placed on agar, the chemical will diffuse from
the disc into the agar. This diffusion will place the chemical in the agar only around the disc. The
solubility of the chemical and its molecular size will determine the size of the area of chemical
infiltration around the disc. If an organism is placed on the agar it will not grow in the area around the
disc if it is susceptible to the chemical. This area of no growth around the disc is known as a "zone of
inhibition."
Antiseptics, disinfectants and antibiotics are used in different ways to combat microbial growth.
Antiseptics are used on living tissue to remove pathogens. Disinfectants are similar in use but are used
on inanimate objects. Antibiotics are substances produced by living organisms such as Penicillium or
Bacillus which kill or inhibit the growth of other organisms, primarily bacteria. Many antibiotics are
chemically altered to reduce toxicity, increase solubility, or give them some other desirable
characteristic that they Jack in their natural form. Other substances have been developed from plants or
dyes and are used like antibiotics. A better term for these substances is antimicrobials, but the term
antibiotic is widely used to mean all types of antimicrobial chemotherapy. Many conditions can affect a
disc diffusion susceptibility test. When performing these tests, certain things are held constant so only
the size of the zone of inhibition is variable. Conditions that must be constant from test to test include
the agar used, the amount of organism used, the concentration of chemical used, and incubation
conditions (time, temperature, and atmosphere). The amount of organism used is standardized using a
turbidity standard. This may be a visual approximation using a McFarland standard 0.5 or turbidity may
be determined by using a spectrophotometer (optical density of 1.0 at 600 nm). For antibiotic
susceptibility testing the antibiotic concentrations are predetermined and commercially available. Each
test method has a prescribed media to be used and incubation is to be at 35-37°C in ambient air for 1824 hours.
The disc diffusion method for antibiotic susceptibility testing is the Kirby-Bauer method. The agar used is
Meuller-Hinton agar that is rigorously tested for composition and pH. Further the depth of the agar in
the plate is a factor to be considered. This method is well documented and standard zones of inhibition
have been determined for susceptible and resistant values. There is also a zone of intermediate
resistance indicating that some inhibition occurs using this antimicrobial but it may not be sufficient
inhibition to eradicate the organism from the body.
The standardized methods for antiseptic and disinfectant testing are more rigorous and more difficult to
reproduce in a student lab. Two common tests are the Phenol Coefficient Test (a comparison of the
effect of the chemical and phenol on several organisms) and the Use Dilution Test (testing the chemical
under actual conditions of use). A disc diffusion test can be used to approximate the Use Dilution Test.
The chemical under consideration is used to saturate a filter paper disc. This disc is then used to
introduce the chemical to the agar for testing. The actual zone sizes have not been standardized as in
the Kirby-Bauer method, but a comparison of zone sizes for the same chemical among organisms will
provide an approximate effectiveness of the chemical.
Procedure - Kirby-Bauer Antimicrobial Susceptibility Test
Organisms to be tested:
Staphylococcus aureus
E. coli
Students will work independently in the laboratory exercise.
1. Obtain a plate culture of one of the organisms to be tested.
2. Using a sterile loop, emulsify a colony from the plate in the sterile saline solution. Mix thoroughly
making sure that no solid material from the colony is visible.
3. Repeat this procedure until the turbidity of the saline solution matches that of the standard
available for your class.
4. Pipette 0.5 ml of the solution and dispense into the center of a Mueller-Hinton or nutrient agar
plate. Using a spreader, spread the solution over the entire surface of the plate.
5. Allow the plate to dry for about 5 minutes.
6. Antibiotic discs can be placed on the surface of the agar using a dispenser or by obtaining individual
discs and placing them on the surface of the agar using sterile forceps. Gently press the discs onto the
surface of the agar, taking care not to press the discs into the agar.
7. Invert the plates and incubate for 24 hours at 37°C.
8. Using a metric ruler, measure the diameter of the zone of inhibition, if present, for each antibiotic
used.
9. Compare the measurement obtained from the individual antibiotics to the table of
standards to determine if the bacterial species tested is resistant or sensitive to the antibiotic.
10. Enter date in your lab notebook and discard plates in the biohazard container.
Procedure - Antiseptic/Disinfectant Susceptibility Test
Organisms Used:
Staphylococcus aureus
E. coli
Bacillus cereus
Pseudomonas aeruginosa
1. Students work individually on this laboratory exercise.
2. Obtain one of the organisms (in liquid culture to be tested) and 5 nutrient agar plates.
3. Dispense 0.5 ml of liquid onto the agar plates.
4. Using a spreader, spread liquid culture over entire surface of the plate and let dry 5- 10 minutes.
5. Label plate with name or organism and antiseptic/disinfectant used. Place a disc soaked in an
antiseptic or disinfectant in the center of each plate.
6. Incubate the plates at 37°C for 18 hours.
7. Measure the diameter of the zone of inhibition for each chemical. The class will share the data for all
chemicals used.
8. Discard plates in the biohazard container.
Review Questions
1. What conditions must be held constant when doing disc diffusion procedures?
2. Define
a. Antiseptic
b. Disinfectant
c. Antibiotic
d. Zone of inhibition
3. According to your results, which chemical is the most effective? On what do you base this
conclusion?
4. What are the standard tests used for determining the effectiveness of antiseptics and disinfectants?
5. What is the standard method used for antimicrobial susceptibility testing?
Urinalysis
Introduction
Urinalysis is the most frequently performed test.
The examination of urine consists of three major areas:
1. Physical
2. Chemical
3. Microscopic
The specimen is collected in a plastic disposable container. Note on the container the patient’s name,
date and time collected as well as the type of test to be done. Perform the test within 30 minutes of
collection or refrigerate specimen.
Timed Specimens
This method is used when the doctor wants samples taken at specific intervals during the day. Time
intervals can be:
24 hour
1. Requires that all urine voided within a 24 hour period be collected
2. Give patient verbal and printed instructions
3. Preservative is usually added to the container
Two hour postprandial
The specimen is collected two hours after eating (used in diabetic testing)
Clean-catch midstream
1. Specimen clears urethra of sloughed off cells, bacteria, mucus, and other
debris that could interfere with the test results
2. Have patient wipe external genitalia from front to back with several
towelettes
3. Instruct patient to void a little and then finish catching about 3 ounces of
urine in the container
Catheterization
This is the introduction of a sterile catheter through the urethra into the bladder to obtain urine. The
medical assistant can perform this procedure if physician or other authorized medical person has given
the release. Physicians usually perform the procedure on males because it is more complicated and
painful to the patient. Sterile techniques must be followed and maintained throughout the procedure
to avoid infection.
There are three reasons for catheterizing patients (not a routine procedure)
1. To obtain a sterile urine specimen for analysis
2. To relieve urinary retention
3. To instill medication into the bladder, after the bladder is emptied
Types of Catheters
1. French catheters are used to perform simple catheterizations
2. Foley catheters are used when the catheter is to remain in the urinary bladder
Pediatric Urine Collection
1. Collection bag fits over infant’s genital area
2. Instruct parent to do this at home as needed
3. Advise parent to transfer urine into a specimen cup with a lid (refrigerate as
needed) for transport to the office
Routine Urinalysis
1. Physical Properties
Appearance: standard color description includes – light straw, straw, dark straw, light amber,
amber and dark amber. Color can be affected by medications, food and disease. Hematuria, for
instance, gives urine a red or rusty color (beets also give urine a red color). Vitamin B gives urine a
bright yellow-green color.
Turbidity and Clarity: Urine can be clear, slightly cloudy, cloudy and very cloudy.
Specific Gravity (SG): Indicates the concentration of urine (weight of substances in urine). The
test is usually done with a urinometer, refractometer, or reagent strips and assessed by comparing the
SG of urine with that of distilled water which is 1.000. Normal specific gravity of urine is 1.010 – 1.030.
A lighter colored urine usually has a lower SG and darker urine has a higher SG.
Odor: Described as normal, strong ammonia (indicates a high concentration of bacteria), fruity
(glycosuria). Any abnormal odor is documented.
2.Chemical Urinalysis: refrigerated urine must be brought to room temperature before testing. Analysis
will be done using reagent strips. Reagent strips will reveal the presence of abnormal levels of
substances and provide both qualitative (what is present) and quantitative (how much is present)
assessment. Reagent strips have the following indicators:
1. Sugar (glucose)
2. Protein (albumin)
3. Ketone (acetones)
4. Bilirubin
5. Urobilinogen
6. Blood
7. Nitrite
8. pH
9. Leukocytes (WBCs)
10. Erythrocytes (RBCs)
11. Specific gravity
The reagents may change color in reaction to substances in the urine. Compare colors with the color
chart on the bottle. Fresh urine is best. Exact timing of reagent strip color blocks is vital and adequate
lighting is also important to view results. No reaction is a negative result. (Check expiration date on the
bottle) Protect reagent strips from heat, light and moisture. Keep bottle capped, and close the bottle
immediately after obtaining a strip.
Procedure
1. Mix the urine to be tested by inverting the sample several times
2. Completely immerse all reagent areas of the strip briefly but completely in the urine
3. Remove excess urine by tapping the edge of the strip against the side of the container and
drawing the strip across the top of the container. You can also press the edge of the strip
against absorbent paper
4. Time according to the manufacturer’s directions using a timing device with a second hand
5. Compare test areas closely with corresponding color charts on the bottle label at the times
specified. Hold strip horizontally and close to the color blocks. Read at the times listed on
the product you are using
Normal Readings
pH: 5.0-8.0
Urine containing bacteria is alkaline (above 8.0). Bacteria destroy casts formed in the kidneys which are
important in diagnosis of patient’s conditions.
Protein: Is not normally present in urine. Diseases of the kidney and UTI produce protein. Patients with
a trace of 1+ reading should repeat the test with the first morning specimen to see if protein is in the
concentrated urine.
Ketone (acetone bodies): appear as a result of fat metabolism and are not normally present in urine.
Presence may indicate severe diabetes mellitus, starvation, a high-fat diet, or body wasting. It can also
be high in the urine of patients fasting for tests.
Bilirubin (bile): an orange or yellowish pigment that is a product of degenerated RBCs that release
hemoglobin in the liver. This normally is excreted through the GI. This is often seen in the urine of
patients who have liver damage or disease before it appears as jaundice of the skin.
Nitrite: indicates microorganisms present in urine. Microorganisms will grow in urine unrefrigerated
more than 2 hours.
Phenylketonuria: phenistix is a reagent strip that detects its presence in urine of infants. Place a strip in
fresh urine, time reaction and read results.