Medical Microbiology laboratory manual
This laboratory practical procedure manual was developed
For MBBS Students (foreign students)
Department of Microbiology, China Three Gorges University,
Yi Chang
Han Li
Contents
Page
1
Introduction
4
Generals rules of microbiology laboratory
4
Microscopy
Procedure to see the smear
5
Laboratory growth media
Types of media
6
Microbiological specimens
Specimens rejection criteria
Specimen labeling
6
Methods of isolation from clinical specimens
Streaking procedure
7
Candle jar technique
9
Method of culture of anaerobes
10
Bacterial colony morphology
10
Preparation of smear for staining
11
Procedure for identification of isolates
Broth culture
An agar slant culture
An agar deep
Terms to describe growth pattern
12
Motility test
13
Gram staining procedure
14
Sterilization
Autoclave
Hot air oven
16
Preparation of routine culture media
Blood agar
Chocolate agar
Mac conkey agar
Mannitol salt agar
Dnase agar
17
2
S-S agar
Tests for bacteria associated enzymes
Catalase
Coagulase
Citrate ultilization
Nitrate reduction
Oxidase
20
Identification media
TSI
SIM
MR-VP
26
Antibiotic sensitivity testing
30
Wet mount microscopy of intestinal parasites
Concentration method
Floatation
Sedimentation
31
Examination of blood parasites
Thick smear
Thin smear
33
Staining of mycobacteria
AFB staining
Mycobacteria culture
34
An introduction to mycology practical
Collection procedure
Wet mount microscopy
35
Serological tests
Technique of blood drawing
Transfer of blood
Factors causing of haemolysis
Precautionary measures
37
RPR testing
39
Widal test
40
Some illustrations of pictures of laboratory concerns
Microscope
41-52
3
Candle technique
Anerobic jar
Loop sterilization
Haemolysis pattern in blood agar
Blood agar and nutrient agar – staph. Aureus
Mac conkey agar : lactose fermenter / nonfermenter
Transfer of bacteria
TSI interpretation
SIM interpretation
Eggs of intestinal helminthes
Cestodes : egg and plroglottids
Protozoal parasites (E. hystolytica & G lamblia )
Malaria parasites : Blood smear, P vivax, P malariae & P falciparum
AFB staining procedure
Fungus (Lactophenol cotton blue) Mucor, Penicillin & Aspergillus
Fungus (Lactophenol cotton blue) Microsporum & Trichophyton
There shall be lecture followed by demonstration and then
the students will practice.
Introduction
The clinical microbiology laboratory of the hospital is more commonly known as
a Diagnostic Laboratory of Infectious Diseases. There will be several sections or
units such as Bacteriology, Parasitology, Mycology, Mycobacteriology, Serology /
Immunology etc. The several categories of clinical microbiological specimens
collected from the hospitalized patients as well as out patients services (OPD )
will be submitted in those different sections or units for laboratory diagnosis to
help the clinicians for treatment management of the patients. These clinical
materials always contain the pathogens; and therefore every one must follow :
General rules / Guidelines of Microbiology laboratory: as a part of contamination /
infection prevention
General rules of microbiology laboratory
1. No eating or drinking in the microbiology laboratory.
2. Always disinfect the tables before and after lab. works
3. Wash your hands with soap both before and after lab, and in between the
lab work. (alcoholic hand rub is also recommended)
4. Do not carry the nonessential materials (the only essential thing is the lab
handouts and a notebook to write in).
5. Always use the proper aseptic technique when transferring cultures
from one medium to another.
4
6. Heat the entire piece of wire loop in the flame: it should be red hot. Be
sure to cool your inoculation instrument before picking up the colony or
liquid broth.
7. Use an inoculating wire for agar deeps and an inoculating loop for the
agar plate and the broths.
8. Label all test tubes and petri plates with your name (initials), your table #,
date, exercise #, and name of organism. Do not use tongue to gum paper
for label.
9. After finishing the lab work, discard the left over microbial cultures, test
tubes in the rack, agar plate in autoclave bag on the autoclave cart for
autoclave.
10. Do not dump ANY microbial suspension down the drain—only on the
discard cart.
11. Place test tubes in racks when working at your table: never lay the tubes
down—they leak.
12. Keep cover on the test tubes and petridish to reduce contamination it
does not matter whether it is sterile media or already cultured
13. Always check agar plates carefully to make sure that there are no mold
or bacterial contaminants on the plate. Do the same with any tube
media that you pick up. if contaminated, discard the plate / tube on the
discard cart.
14. All agar plates are incubated upside down before inoculation because It
reduces bacterial contamination.
Microscopy
The microscope is absolutely essential to the microbiology laboratory. This is the
first equipment of the laboratory.
Therefore every student should know
- the parts of the microscope and their functions.
- how to use the microscope effectively, and particularly the oil immersion
objective. This will be demonstrated and explained in the practical exercises
Procedure to see the smear
Place the prepared smear (or wet mount) on the stage, and secure it inside of the
stage clips.
1. First use low power objective (10Xlens) Try to know where the specimen
is located on the slide, and place it in the center of the hole allowing light
through the stage; and then focus the smear
2. While looking through the ocular eyepiece, lower the stage slowly using
the coarse adjustment knob.
3. As soon as you see the specimen, Stop using the coarse adjustment, and
then use the fine adjustment knob
4. Locate good field for examination by oil immersion objective (100Xlens).
5
5. Now ready to move with oil immersion objective. Keep a small drop of
microscopic oil. And then rotate the 100X objective (oil immersion) into
place. The lens will touch to the oil drop.
6. Use fine adjustment to see the bacteria. Be sure that condenser is raised
and light is adjusted. High power (40Xlens) is usually used for wet mount
microscopy
Note.
a. Be sure to remove any oil by lens paper before covering the
microscope. Do not use other tissue paper
b. Once you have gone into oil immersion, do not go back to the 40X lens;
oil will contact to the high power objective while changing the objective.
The 10X can be returned if necessary, since the lens should not touch the
slide anyway.
c. Clean all lenses (oculars, objective lenses, and lens on condenser) with
lens paper. One should not forget to clean the oil lens with lens paper.
Laboratory growth media :
Types of media: The types of media are described in following pages
An agar medium: This contains agar (1.5-2%) as a solidifying agent; and this
helps for isolation and purification e.g. Blood agar, Chocolate agar. MacConkey
agar, Salmonella-Shigella agar etc.
A broth medium: This has no agar; it is used for fast, luxuriant growth of
bacteria e.g. Brain Heart Infusion broth (BHI broth)
A semi-solid medium: This has a reduced percentage of agar, and can be used
for motility e.g. SIM medium
Microbiological specimens
All the clinical specimens ( such as sputum, urine, pus swab, body fluid etc)
must be proceeded soon after its reception. Delay causes the either overgrowth
or pathogens may die because of drying of specimen. The specimen should be
kept in the refrigerater particularly csf specimens because cold sensitive
organisms die soon.
6
Specimen Rejection / Test Cancellation
All specimens must be collected, labeled, transported, and processed according
to standard laboratory procedure. Selecting the container type, volume, and
proper handling of clinical material is essential for correct microbial diagnosis.. If
the criteria for these processes are not met, the specimen may be rejected or the
test may be canceled. The following represent some reasons for specimen
rejection or test cancellation:
• Insufficient volume for analysis
• Improperly labeled specimen
• Inappropriate specimen container
• Improper specimen transport
• Specimen that has leaked in transit
• Specimen that has been sent in incorrect or expired transport media
• Incomplete or incorrect test request form (e.g., testing not marked)
• Test request form without a specimen
• Specimen without a test request form
• No specimen type, no source provided
• Inappropriate specimen type (e.g. saliva in stead of expectorated sputum
specimens).
Specimen Labeling
To assure positive identification and optimum integrity of patient specimens from
the time of collection until testing is completed and results reported, all
specimens submitted for testing must be adequately labeled.
The label must have the patient’s name, age, sex with a identifying number (e.g.,
medical record number) and specimen type or source. Clients will be notified of
inappropriately labeled specimens which will be returned to the client upon
request
Method of isolation from the clinical specimen :
Streaking an agar plate
1. Gather all the necessary materials ( growth media, bunsen burner, transfer
tools, etc).
2. Practice adjusting the flame of the bunsen burner.
5. Sterilize the loop or wire by holding in the flame of a gas burner. The metal
must glow red before sterilization is considered complete.
6. Specimen is taken using sterile wire loop and streaked as demonstrated for
the isolation of pathogen
Routinely three kinds of isolation media are used in the microbiology laboratory.
These are Blood agar, Chocolate agar and MacConkey agar. Until you become
7
well-acquainted with this procedure, you might want to draw the 3 sections that
you will streak inside of, on the back (bottom of plate containing agar medium)
with a sharpie pen.
STREAKING AN AGAR PLATE
Streaking Procedure:
1. Pick up a loopful of your inoculum from clinical specimen, a broth or an
agar culture.
2. Using a sterile agar medium plate (lift the lid just enough to insert the
loop), streak a vertical line straight down.
3. When streaking the agar, keep the loop horizontal and only streak the
surface of the agar: do not dig into the agar.
4. Move the loop in a zig-zag pattern across the agar until 1/3 of the plate is
covered, finishing the first section.
5. Sterilize the loop in the flame and let it cool before continuing to spread
the bacteria.
6. You can do this by i) sticking the hot loop in the agar at the edge of the
agar away from the bacteria, or ii) just holding the loop for a few seconds
while it cools.
7. Rotate the plate about 90 degrees and spread the bacteria from the first
streak into a second area using the same zig-zag spread technique.
8. Sterilize the loop again. Rotate the plate about 90 degrees and spread
the bacteria from the second streak into the 3rd area in the same pattern.
Sterilize the loop again.
9. Replace the lid and invert the plate. Incubate the plate.
8
All the plates, tubes or others if any are incubated aerobically in normal
incubator at 36o C for 24-48 hours. You can see bacterial cells growing
along streak lines and in isolated areas.
Beside use of normal conventional incubator for normal incubation there are also
necessary of other incubation environments such as reduced oxygen content and
increased levels of carbon dioxide, anaerobic environment.
Candle Jar technique : method to reduce oxygen content and elevate levels
of carbon dioxide
Purpose
Certain specimens such as csf, sputum contain the pathogens that require
reduced oxygen content and elevated levels of carbon dioxide during the
incubation. The candle jar method creates an atmosphere with reduced oxygen
and elevated levels of carbon dioxide. These conditions enhance the growth of
microaerophiles. The candle used for this purpose normal plain candle (not
colored candle).
Principle
The flame of the candle within a closed environment, will use up a certain
percentage of the oxygen. When the available oxygen is reduced and elevated
carbon dioxide created by the flame is increased, the flame will be extinguished.
The plated medium within this atmosphere, will show enhanced growth of certain
bacteria such as Haemophilus influenzae. The candle jar is incubated at 36o C
as usual incubator.
9
Method of culture for anaerobes
Anaerobic Jar (Brewer's Gas Pak or Mc Intosh Field Jar or BBL Jar)
Purpose
The anaerobic jar creates an artificial anaerobic environment (devoid of oxygen)
which permits the growth of anaerobic bacteria.
Principle
The anaerobic jar employs a chemical reaction to generate hydrogen gas. In the
presence of a palladium catalyst, the hydrogen gas will react with free oxygen in
the air to form water. This reaction removes the oxygen from the sealed
atmosphere. The jar is then incubated at the desired temperature. There will be
slight negative pressure inside.
Bacterial Colony Morphology
Purpose Identifying and categorizing different, isolated bacterial colonial
morphology (form and structure) will permit the selection and transfer of different
species from a mixed culture, and allow transfer of a single colony to a sterile
medium for cultivation of a pure culture. This exercise also shows how many
diverse bacteria and fungi are present in our environment, as exhibited by the
varied morphologies
10
Principles When a single bacterial cells is deposited on the surface of a nutritive
medium, it begins to divide exponentially. After thousands (up to billions) of cells
are formed, a visible mass appears. This mass of cells is called a colony.
Therefore is defined as a visible mass of microorganisms all originating from a
single mother cell, therefore a colony constitutes a clone of bacteria all
genetically alike. Each species of bacterial or fungal organism will exhibit
characteristic colonies
Colony shape and size
Margin (edge): Elevation: Color: Texture
Colony characteristics (colonial morphology colony shape and size) are normally
described as
round, irregular, punctiform (tiny), margin (edge): entire (smooth), convex,
umbonate, flat, raised, pigmented, opaque, translucent, shiny or dull:
moist, mucoid, dry (or rough).
Surface of colony:smooth, glistening, rough, dull (opposite of glistening),
rugose (wrinkled)
Consistency: butyrous (buttery), viscid (sticks to loop, hard to get off),
brittle/friable (dry, breaks apart), mucoid
Emulsifiability of colony: Is it easy or difficult to emulsify? Does it form a
uniform suspension, a granular suspension, or does not emulsify at all? To
determine emulsifiabiliti, the colony is suspended in a tube or water or saline
solution to see how it goes into solution. Pick the colony with a loop, take it into
the tube of liquid, and smear the inoculum against the glass wall right above the
line of fluid within the tube. This ensures that the inoculum suspension becomes
an even suspension rather than an undissolved mass of bacteria floating in the
water.
11
Preparation of smear for Staining
From an agar plate culture:
Pick a bit of a colony from the plate showing the growth by use of flamed
sterilized loop or straight wire. This is transferred to the slide containing a small
drop of sterile normal saline and emulsify. A uniform thin smear is necessary.
1. The smear is thoroughly dried in air
2. The slide is then heat-fixed using one of the following techniques:
a. by passing the slide through a flame to heat-fix the smear.
b. use of a slide warmer can be used to heat-fix the slide.
The heating adheres the smear to the glass surface and kills the cells
Heat-fixing a bacterial smear to the slide by the use of the technique of
passing the slide through a bunsen burner flame which can easily destroy
bacterial cells if the slide is held in the flame longer than necessary.
Inoculating the new medium for identification of isolates
Procedure of taking the inoculum
1. Get the new medium into which which you are transferring the inoculum. Be
sure that the new medium is labeled so you do not confuse the various cultures.
2. Heat the inoculating wire of the loop or needle until red-hot, and be sure that
the entire wire is sterilized. You are now ready to pick the inoculum from the
bacterial culture.
A broth culture:
The inoculums is transferred into the inner sides of the tube containing broth by
use of bacteriological loop aseptically .
An agar slant culture:
Place the loop with the bacteria into the slant tube, all the way down to the
bottom of the slant. There are 2 ways to inoculate the slant. If you want to identify
the type of growth pattern, then just bring the loop straight up the slant.
If you want to have a luxuriant culture, inoculate in a zig-zag pattern, starting at
the bottom of the slant. This increases the surface area of the culture.
An agar plate culture: as described on plate streak technique
An agar deep:
Use the needle to inoculate the deeps or semi-solid agars. Stab the inoculum
down to the bottom of the deep in a clean, straight stroke.
12
Terms to describe growth patterns
bacterial cultures in broth:
Study of growth patterns in broth can be used for identification of organisms.
These patterns can be characteristic for certain species. Remember, however,
that these growth patterns can be influenced by growth conditions such as type
of medium used or the temperature of incubation
1. uniform turbidity, or diffuse growth
2. flocculent (clumps)
3. sediment, and
4. ring or pellicle growth on surface
Motility test:
The motility of the organisms can be identified in different ways:
The hanging drop wet mount and
Semi-solid motility medium (SIM medium tube is also used for this purpose)
1. The hanging drop slide method :
Use a small drop of bacterial suspension, but do not let it dry out (use a young
culture of bacteria).
Materials needed:
i. fresh cultures (broth medium less that 24 hours old is optimal) of E. coli and
either Bacillus or Pseudomonas
ii. depression slides or coverslips and petroleum jelly
Procedure:
1. Place a drop of the bacterial culture (optimally from a young broth culture) in
the middle of a cover slip.
2. Place a thin line of petroleum jelly around the edge of the cover slide.
3. Turn the depression slide upside-down (depressed area facing down) and
gently touch the cover slide. The jelly holds the cover slip to the slide and also
keeps the suspension from drying out.
4. Now flip the entire microscope slide/cover slip combination over.
It should look as shown in this figure
13
Looking at living bacteria are not as easy as one would think. First of all, living
bacteria have no color, and they are small: therefore, they are really difficult to
see, even with the oil immersion lens. Second, all bacteria have some vibrational
movement, even nonmotile ones. This Brownian movement is caused by water
molecules bouncing around in the solution, knocking up against each other and
the microorganisms. Kinetic energy inherent to all molecules causes this kind of
movement. On the other hand, those bacteria with flagella will be very apparently
moving about the field of vision, although perhaps not all of the bacteria will be
moving. Some cells will "run" straight across the field, others will "tumble" across
the field in a slower motion.
The medium is inoculated with an inoculating needle. The semisolid medium is
stabbed with inoculum and then incubated. Next day we can see the result of the
growth without the special stain and microscope. The movement of bacteria is
detected along the line of inoculation
Principle The lower agar concentration in the medium allows limited movement
of motile bacteria from the area of the stab. Motility will be detectable as diffuse
growth radiating from the stab line. A special dye, a tetrazolium salt (TTC) may
be added to make the radiating growth more visible as the reddish diffuse area
NOTE : The stab is performed with an inoculating needle. Care must be taken to
insert the needle straight in and to pull it straight out. If the needle "slices" the
agar horizontally, it may be difficult to tell if the resulting bacterial growth is from
motile bacterial, or is just growth along a wide stab line.
Motile bacteria move about with structures called flagella. Nonmotile bacteria
without flagella are called atrichous.
Categories of flagellation:
monotrichous = single flagellum
peritrichous = flagella all around
lophotrichous = tuft of many flagella at one end or both ends
14
Gram staining procedure
This staining is named after the Danish physician, Christian Gram, who
developed this technique in 1884. It involves a series of simple steps.
Gram staining method separates bacteria into two types. Gram-positive cells
retain the crystal violet-iodine complex and thus appear purple Gram-negative
cells are decolourised by the alcohol or acetone treatment, and then stained with
safranin so they appear pink
Gram-positive bacteria have a relatively thick wall composed of many layers of
the polymer peptidoglycan (sometimes termed murein). The thickness of this
wall blocks the escape of the crystal violet-iodine complex when the cells are
washed with alcohol or acetone.
Gram-negative bacteria have only a thin layer of peptidoglycan, surrounded by a
thin outer membrane composed of lipopolysaccharide (LPS). The region
between the peptidoglycan and LPS layers is termed the periplasmic space; it
is a fluid or gel-like zone containing many enzymes and nutrient-carrier proteins.
The crystal violet-iodine complex is easily lost through the LPS and thin
peptidoglycan layer when the cells are treated with a solvent.
Procedure:
Make the bacterial smear from broth, slant, or plate as described earlier
1. If taken from a broth, use a loopful of the solution.
2. If taken from plate or slant, suspend the inoculum in a drop of normal
saline on the slide and mix it well.
3. Spread the suspension on the slide so that it covers the size of one cm.
circle
4. Place a piece of tape on the side of the smear so you know which way is
up.
5. Air-dry and heat-fix the slide.
6. Place slide into container of crystal violet, leaving in for 1 minute. Wash
well with water
7. Place slide into container of Gram's iodine, leaving in for 1 minute. Wash
well with water.
8. Flood acetone-alcohol quickly on the slide, and wash off within 5-10
seconds (from beginning of decolorizer added). Wash well with water.
9. Place slide into container of safranin, leaving in for 1 minute. Wash well
with water.
10. Blot dry with bibulous paper.
11. Focus on smear using low power lens, ending up on 100X oil immersion.
Interpretation of microscopic finding:
Gram positive bacteria will be blue/purple/violet.
Gram negative bacteria will be light pink.
Note: We have cultures of bacteria such as Staphylococcus, E. coli, or
15
Bacillus for doing Gram stain. We will give you gram + ve and Gram –ve
bacteria to check your procedure
Sterilization
This is very important in microbiology laboratory. We do sterilization to destroy
the bacteria and other microganisms. We do sterilization to growth media for
use. We do sterilization for all the specimen collecting tools or containers. There
are many methods of which autoclave and hot air oven are extensively used.
In some cases we use disinfectant to kill the microorganism such as
Glutaraldehyde, Hypochloride solution, Lysol disinfectant etc.
Autoclave:
This steam sterilizer is the most frequently used in the laboratory. You
must know the details of this equipment
Basically an autoclave is a huge steam cooker. Steam enters into a jacket
surrounding the chamber. When the pressure from the steam is at a certain
point in the jacket, a valve allows the steam to enter the chamber. The pressure
will go up over 15 pounds per square inch (psi): at this point the timer begins to
count down, usually for 15 minutes, depending on the type of media. The high
pressure in a closed container allows the temperature to go above the highest
temperature one could get by just boiling, around 121 degrees C. Therefore, the
parameters for sterilization with an autoclave are 121o C at 15 lbs for 15
minutes. Fifteen minutes is the thermal death time for most organisms (except
some really hardy sporeformers).
16
Hot Air oven
Beside autoclave there is another very important equipment that is Hot air oven.
This equipment Hot Air Oven is used for the sterilization of articles in dry
condition. Hot air trapped after closing the door of the oven is heated to the 160 o
C . This temperature is maintained for one hour (holding time). In this time period
of 160o C ( the holding time) the articles will be free from the viable
microorganisms including the spore formed organisms. The operation of Hot Air
Oven will be demonstrated.
Preparation of Routine Microbiology Media
Normally there are two type of media : nutrient broth and nutrient agar.
These two media; one is a liquid and the other is a solid. The solid media is
prepared by addition of agar agar (a sea weeds), an extract from the cell walls of
red algae.
These days all the laboratory growth media are easily available in the dehydrated
form in the market. We simply need to rehydrate the powder form of the medium
as described in the instruction procedure contained in the purchased dehydrated
media; and all these prepared media are autoclaved.
17
The laboratory staff will demonstrate making of NA (Nutrient agar) plates, NA
slants, and Nutrient broth
The procedure:
The flasks of melted nutrient agar are kept in the water bath which is set to
45oC. This is to prevent them from solidifying, since agar solidifies at around
42oC. The flask will be taken from the bath only when ready to pour the plates.
Set the petri dishes out (small side down), tops covering the dishes. When it will
be ready to pour the plates, the staff will take the flask of agar and pour the
plates normally 20 ml per petri dish. Cover the bottom of each plate, Allow all
plates to stand until they are completely solidified. Once they have solidified,
place them on the tray upside down (bottom dish with agar on top).
Why are agar plates incubated upside down? Two reasons:
There may be air contaminants in the incubator, it will be more difficult for
them to get into the plates.
Often we will see a bit of water condensation on the top of the agar plate,
particularly when they have been at 37 oC. The water molecules are cohesive
and tend to run together. If the plate was sitting right side up, the water droplets
could then fall onto the agar, creating a kind of little lake on the agar plate and
messing up your plates. Upside down plates prevents the condensation from
dropping on the agar surface.
Routine used media
Blood Agar
Blood agar is used both as an enriched medium for growing fastidious bacteria
and as a differential medium to study the pattern of haemolysis. The hemolysis
pattern is an especially useful tool for identification of many of the Gram positive
cocci. Hemolysins are grouped in three categories:
1. Beta hemolysins completely lyse the red blood cells and hemoglobin; this
results in complete clearing around colonies.
2. Alpha hemolysis refers to the partial lysis of RBC's and hemoglobin and
produces a greenish discoloration of the blood agar around the colonies.
3. Gamma hemolysis, sometimes called NO hemolysis results in no change
of the medium.
When reading plates for type of hemolysis, be sure to look for color changes in
the surrounding agar. A common mistake is to look at the color of the bacteria,
rather than the agar.
MacConkey Agar
18
MacConkey agar is a widely-used culture medium which is both selective and
differential. The medium is primarily used to differentiate between Gram negative
bacteria while inhibiting the growth of most Gram positive bacteria. The medium
also differentiates between lactose-fermenting coliforms and non-lactose
fermenters, which include potential pathogens
This medium contains bile salts and crystal violet that will inhibit the growth of
most Gram positive bacteria, making MacConkey agar selective. Lactose, a
fermentable carbohydrate, and neutral red, a pH indicator, are added to
differentiate the lactose fermentors from the pathogenic lactose nonfermenters.
When lactose is fermented, acid products lower the pH below 6.8, with the
resulting colonial growth turning pinkish-red. If an organism is unable to ferment
lactose, the colonies will be colorless.
MacConkey is selective: Gram negative organisms will grow, Gram positive
organisms will not. This is due to the addition of bile salts and crystal violet.
MacConkey is differential: Lactose fermenters will appear pink, non-lactose
fermenters will appear colorless. This is due to the addition of the indicator,
neutral red.
Mannitol Salt Agar
Staphylococcus is the most commonest isolates’; therefore this medium is used
in the laboratory. Mannitol salt agar is both a selective and differential growth
medium. This medium helps determine two characteristics of bacteria, whether
they are salt tolerant or not, and whether they are able to ferment mannitol or not
1. Salt Tolerance : This medium contains 7.5% salt higher than normal mediuim
( about 0.5% NaCl in the routine isolation medium) and therefore "selects" for
organisms that are able to tolerate the presence of high levels of salt. If the
organism grows, it is salt tolerant; if no growth it is not salt tolerant.
2. Fermentation of mannitol. This medium contains an indicator, phenol red.
The indicator is a pinkish-red at neutral pH, is really red at pH 7.4 or above and
is yellow below pH 6.8 (acidic). An organism which can ferment (metabolize)
mannitol will be indicated by a color change on the MSA plate(which makes this
a differential test; and if no fermentation the medium will remain red (no change).
If the organism does ferment mannitol, it will create metabolic by-products which
are acidic--and the surrounding medium will be yellow.
DNase Test
DNase test is used to identify bacteria capable of producing the exoenzyme
DNase This enzyme catalyzes the depolymerization of DNA into smaller
19
fragments is called a deoxyribonuclease, or DNase. The test agar contains an
emulsion of DNA, peptides as a nutrient source, and methyl green dye. The dye
and polymerized DNA form a complex that gives the agar a blue-green color at
pH 7.5 . Bacterial colonies that secrete DNase will hydrolyze the DNA in the
medium. This results in clearing around the bacterial growth
Salmonella Shigella agar (SS agar)
This is the selective media for isolation of Salmonella and Shigella organism. In
this media all coliform bacilli group of bacteria are inhibited. However certain
strain of coliform bacteria can grow
The stool culture findings are usually positive in approximately 85-90% of
patients with typhoid fever during the first week, declining to 20-30% later in the
course of the disease. The conventional culture techniques usually take 48-72
hours from acquisition until the organism is identified.
The bacterial dysentery caused by Shigella species are also isolated by this
medium
The stool specimen is collected in a wide mouth sterile (preferably) wide mouth
container. One loopful of mucus portion preferably blood containing material is
and inoculated into SS Agar (DCA Dexycholate agar is also used in parallel in
many countries. A single rectal swab culture at hospital admission can be
expected to detect S typhi in 30-40% of patients. The plates are incubated
overnight at 37 oC. Salmonella and Shigella bacteria are non lactose fermenter..
The isolates are further identified by motility test and biochemical tests The
biochemical tests includes TSI, SIM, Citrate, MR-VP
Serology of isolate is done for species identification.
Tests for bacteria associated enzymes
Catalase test :
catalase is an enzyme that splits hydrogen peroxide into water and
oxygen. Hydrogen peroxide is a by-product of respiration and is lethal if it
accumulates in the cell. Catalase is an enzyme that can degrade the hydrogen
20
peroxide in the cell before it can do any cell damage. It splits the H2O2 to free
oxygen (bubbles) and water.
Important point : Use growing bacterial colony (plate or tube)
You can drop 3% H2O2 directly on an agar plate / slant, but it may kill the culture.
The procedure: Slide method
Pick the inoculum from a plate culture or slant culture and place it on a slide.
Add one drop of 3% H2O2.
Result: You will see a reaction if the test is positive, most often lots of bubbles.
Slight bubbles also indicate a positive reaction.
CATALASE TEST
Coagulase test
This is one of the most common test
Why we do this test? To differentiate potentially pathogenic Staphylococcus
species from other Gram positive and catalase-positive cocci.
It is thought that an infective organism that produces the coagulase enzyme may
protect itself by inducing clotting in surrounding tissues, thereby inhibiting
destruction by normal body defense such as phagocytosis or antibodies
21
Purpose This test is used to detect the ability of certain Staphylococcus species
to clot plasma. Coagulase production is a characteristic of the potentially
pathogenic S. aureus.
There are 2 methods (slide test and tube test) : Both tests use the same
substrate; The tube test is more accurate, but the slide test is faster.
Material needed: 0.5 ml rabbit plasma per test
Slide agglutination test
Make a 2 cms diameter circle on a clean glass slide using a wax pencil. Place
two drops of thawed rabbit plasma into the circle, using a wooden pick or a clean
loop. Add a single colony and emulsify it in the plasma. Add a single colony and
emulsify it in the plasma. Fibrin threads form between the cells, causing them to
agglutinate, or clump. There will a visible clumping of cells within 10-15 seconds.
This test is for the bound coagulase enzyme.
Tube agglutination test
Inoculate a tube with a ½ ml of rabbit plasma with the bacterial inoculum. Place
at 37C and check at ½ hour and after (some strains will give a + reaction in a few
hours, other strains take longer) by tipping the slide at an angle. Any degree of
coagulation is considered a positive test for the free coagulase enzyme. Add a
single colony and emulsify it in the plasma. Fibrin threads form between the cells,
causing them to agglutinate, or clump. There will a visible clumping of cells within
10-15 seconds. This test is for the bound coagulase enzyme.
Rabbit plasma is inoculated with
the organism
22
Citrate Utilization Test
Purpose The citrate utilization test is used to determine the ability of an
organism, using the enzyme citrase, to use citrate as its sole carbon source. The
citrate test identifies the use of citrate as a sole carbon source, since there are no
other nutrients in this medium. The basic end products will cause the brom
thymol blue indicator in the medium to turn from forest green to royal blue.
Principle Simmon's citrate agar is a medium containing sodium citrate as the
sole carbon source and the ammonium ion as the sole nitrogen source. The pH
indicator, bromthymol blue, will turn from green at neutral pH (6.9) to blue when a
pH higher than 7.6 is reached (basic or alkaline). If the citrate is utilized, the
resulting growth will produce alkaline products (pH >7.6), changing the color of
the medium from green to blue.
In this medium, sodium citrate is the sole source of carbon and energy. This test
determines whether or not an organism is able to metabolize citrate for energy.
The uninoculated medium is green. An indicator, bromthymol blue, is added to
the medium, which changes color based on pH. The citrate will be yellow if the
metabolic products are acidic. A color change to royal blue indicates alkaline byproducts. Any color change (from green to either yellow or blue) represents a
positive test for citrate utilization.
Nitrate Reduction Test
Purpose This test detects the ability of an organism to reduce nitrate (NO 3) to
nitrite (NO2) or some other nitrogenous compound, such as molecular nitrogen
(N2), using the enzyme nitrate reductase
Principle Nitrate (NO3) may be reduced to several different compounds, either by
anaerobic respiration or by denitrification. This test is used to detect whether or
not the reduction has taken place. The nitrate medium contains potassium nitrate
23
as the substrate. If the organism reduces the nitrate to nitrite, the nitrite will react
with added reagents sulfanilic acid and a-naphthylamine to produce a red color.
If no color is produced, this can indicate either of two reactions:
(1) the nitrate was not reduced
(2) the nitrate was reduced even further to compounds other than nitrite.
To distinguish between the negative reaction, or the complete reduction, zinc
dust is added. If nitrate remains in the medium, zinc will reduce it to nitrate, and a
pink color is observed. This is a negative reaction. No color change after zinc is
added means that nitrate has been reduced to compounds other than nitrite. This
is interpreted as positive and is often call positive complete to distinguish it from
the first positive test discussed
Reagents :
Nitrate broth
nitrate reagents A (sulfanilic acid) and B (naphthylamine)
wooden sticks for zinc
zinc powder
Procedure:
Inoculate the nitrate broths with your bacterial unknown.
Incubate at the optimal temperature, 30 or 37C, for your organisms.
Result:
After incubation: Look for N2 gas first before adding reagents; and then
add 6-8 drops of nitrite reagent A. After this add the same number of drops of
nitrite reagent B. One can see a reaction within a minute or less.
If you have not seen either nitrite or N2 gas, you need to add a bit of powdered
zinc. A bit of zinc is about the amount that sticks to the end of a wood stick.
Result
Reaction
N2 gas
NO3 to NO2
NO3 to NO2
NO3 to ammonia
NO3 – no reaction
none
yes
None
None
Color after
adding reagent
red
no color
no color
no color
Color after
adding zinc
Not added
no color
no color
pink color
Nitrate Test Reagents
Reagent A: Sulfanilic Acid
Reagent B: alpha-naphthylamine
24
Zinc dust is added to tubes that appear to be negative (remain clear) after the
addition of
Nitrate Reagents A & B
Oxidase test
This test is used to identify bacteria containing the respiratory enzyme
cytochrome oxidase.The oxidase test is a key test to differentiate between the
families of Pseudomonadaceae (Oxidase positive) and Enterobacteriaceae
(Oxidase negative). The enzyme cytochrome oxidase is involved with the
reduction of oxygen at the end of the electron transport chain. The colorless
reagent used in the test will detect the presence of the enzyme oxidase and,
reacting with oxygen, turn a color.
Principle
The cytochrome oxidase enzyme catalyzes the transport of electrons from a
donor compound to the final electron acceptor, oxygen. In this test, an artificial
electron donor, tetramethyl-p-phenylenediamine, a redox dye in its reduced form,
is used to reduce the cytochrome oxidase. If the enzyme is present, the colorless
dye will turn a purple to blue color. No color change is a negative test.
A couple of key points when doing this test:
We keep the oxidase reagent either frozen or unopened in tubes until
needed. If old reagent is sitting out on the bench and is purple, ask for a new
tube from the instructor
Use a young culture, preferably less than 24 hrs old.
Use fresh reagent, less than a couple of hours old (it is taken out of the
freezer).
Pick your inoculum, not with a metal loop (reagent may react with the
metal), but with a wooden stick.
Read the reaction within 20 seconds (not after), usually it will change in
less than 15 seconds. The oxygen will change the reagent color as time passes,
so it must be read quickly.
25
Materials needed:
oxidase reagent (Tetramethyl-p-phenylenediamine)
wooden rods
Procedcure:
Pick a good-sized amount of inoculum from a plate culture or slant culture and
place it on a piece of filter paper.
Add one drop of the reagent (if it is dark blue, it is old and should not be used).
OR you can drop the reagent directly onto the slant or plate, but that might
damage your culture.
Result
A positive reaction, bluish-purple will occur within 20 seconds. color that
progressively becomes more purple Do not read the reaction after 30 seconds.
Decarboxylase Test
This test is used to detect the ability of an organism to decarboxylate an amino
acid. Decarboxylation is a reaction which removes the carboxyl group (COOH)
of an amino acid, producing an amine and carbon dioxide. The amino acid is
added to the test medium, along with the pH indicator, bromcresol purple, and
the medium is sealed with mineral oil after inoculation. This creates anaerobic
conditions which promote fermentation. Accumulation of acid end products from
fermentation is necessary because decarboxylase enzymes are inducible only in
the presence of substrate and acid environment. The decarboxylation of the
amino acid by the enzyme then results in alkaline end products. These in turn will
cause the pH indicator to turn purple (positive).
Lysine and ornithine are the amino acids which are tested for decarboxylation.
Arginine, another amino acid tested, is a different chemical reaction:
dihydroxylation. However, the test results are interpreted in the same way.
Identification media
After the isolation of pathogens it must be identified by use of idendtification
media as well as the biochemical tests
Triple sugar iron agar (TSI)
This is a stab and streak inoculation technique.
This is a good medium for culture of bacteria. This medium is used for Gram
negative rods usually .
TSI agar tests for knowing three things :
They are sugar fermentation (glucose/lactose/sucrose), CO2, and H2S. Carbon
dioxide is identified by cracks and bubbles inside of the medium, sometimes a
few bubbles and sometimes enough to push the slant up to the top.
26
TSI agar contains peptone, glucose, sucrose, lactose, and thiosulfate. Phenol red
is the pH indicator (yellow at pH less than 6.8 and red above 6.8). It is prepared
on an agar slant, with a deep butt to provide for anaerobic growth.
Procedure
Inoculate the medium using an inoculating wire. Stab the inoculum down through
the butt, then pull the needle out and streak up the slant ( Do not take another
inoculum to do the slant).
Incubate at 37 o C.
There will be demostration so that you can see some different reactions using
this medium.
There are 3 possibilities of sugar reactions and different reactions
in different areas (butt vs. slant) of the medium, so the
physiology behind it is pretty complex. The outcome of sugar use
is always acid, so the pH indicator phenol red will turn yellow--reported as A. No use of the sugar or alkaline by-products
(which is NO sugar use) from the other non-sugar nutrients in the
medium will cause the indicator to stay the same color red/orange
or maybe even change it to a red---reported as a K.
The reactions in TSIA are reported as slant (A or K), butt (A or K), a circle around
the butt for CO2, and + for H2S. For example K/A +H2S = red slant, yellow
butt, with both CO2, and H2S. The glucose (=dextrose) is 1/10 in concentration
as the other 2 sugars. The fermentation of the sugars causes the anaerobic butt
to turn yellow and stay yellow. However, if only glucose is used, even though the
slant turns yellow only after a few hours it will revert to red because the protein in
the medium is broken down to alkaline products when the small amount of
glucose is used up. If lactose and/or sucrose are used, the large amount of
fermentation products neutralizes the basic products and the slant stays yellow.
The following symbol is used to express the result
A/A = glucose and lactose and/or sucrose are used
K/A = glucose alone
K/K = no sugars used
NOTE: Purpose of TSI test is to differentiate bacteria based on their ability to
ferment glucose, lactose and/or sucrose, and to reduce sulfur to hydrogen
sulfide. It is used primarily to distinguish the morphologically similar bacteria of
Enterobacteriaceae, all of which ferment glucose to an acid end product.
There is no way to get a A/K reaction when using a Gram negative rods on this
medium. IF you do, it means that 1) you did not inoculate correctly with a stab
and streak or 2) you inoculated something other than a Gram negative rods.
27
SOME COMMON SUGAR REACTIONS IN TSI
Slant /butt
Symbol
Interpretation
Red / Yellow
Yellow / yellow
K/A
A/A
Red / Red
K/K
Red / no color change
K/NC
Yellow / yellow with gas
A/A, G
Red / yellow with Gas
And black precipitate
Yellow / Red
And black precipitate
Yellow / yellow
And black precipitate
No change / No change
K/A,G, H2S
Glucose fermentation,
Glucose & Lactose and or
Sucrose fermented
No fermentation
Peptone catabolised
No fermentation
Peptone used aerobically
Glucose & Lactose and or
Sucrose fermented with Gas
Glucose fermented and gas
H2S produced
Glucose fermented only
H2S produced
Glucose, Lactose and or
Sucrosed fermented, H2S
No fermentation
K/A, H2S
A/A, H2S
NC/NC
A = Acid production;K = Alkaline reaction
H2S = Sulphur reduction
G = Gas
Interpretation:
A/A = yellow throughout e.g. Esch.coli
K/A = red slant, yellow butt e.g. Salmonella, Shigella
K/K = red or red throughout e.g. Pseudomonas
carbon dioxide = bubbles or breaks in medium
black precipitate = hydrogen sulfide
SIM Medium
This medium is used for three tests: they are Sulphur Reduction test, Indole
Production test, Motility test. This is particularly important in differentiating
certain enteric organisms
Sulphur Reducton
Hydrogen sulfide, H2S, can be formed by putrefaction or anaerobic
respiration. The medium contains cysteine, an amino acid containing sulfur, and
sodium thiosulfate plus peptonized iron or ferrous sulfate. The H2S will react with
the iron or ferrous sulfate, forming a black precipitate. If the black precipitate is
present, the test is positive for H2S production. No precipitate is a negative test
28
Test for Indole Production
The indole test is used to identify bacteria capable of producing indole
using the enzyme tryptophanase.
The enzyme tryptophanase can convert the amino acid, tryptophan, to
indole, ammonia, and pyruvic acid. The by-product, indole, is the metabolite
identified by this test. When Kovac's reagent, which contains hydrochloric acid
and dimethylaminobenzaldehyde and amyl alcohol, a red layer will from when
indole is present. No color in this layer is a negative result.
Motility test from semisolid agar
This medium can be stab-inoculated with an inoculating needle to indicate
motility. The lower agar concentration in the medium allows limited movement of
motile bacteria from the area of the stab. Motility will be detectable as diffuse
growth radiating from the stab line.
Methyl red (MR) and Voges-Proskauer (VP) tests
The MR - VP tests are run together in the same broth and then split into 2 tubes
when ready to be tested for the end products.
The methyl red test determines the use of glucose, with the subsequent
production of acid, tested for by the pH indicator methyl red. The VogesProskauer test also determines glucose use, but for a different end product---not
acid but a neutral product called acetoin (or acetylmethylcarbinol). The VP is
really important for identification of many bacteria, and must be done carefully.
Determine the various reactions for these media: MRVP broth, .
1 MRVP broth per unknown


Barritt's reagents A (alpha-naphthol) and B (KOH)
methyl red reagent
The procedure:
Inoculate into the MRVP broth.
Incubate at 25 or 37 degrees C.
AFTER INCUBATION: Pour 1/3 of the suspension into a clean nonsterile
tube: run the MR test in the tube with 2/3, and the VP test in the open
tube with 1/3.

for methyl red: Add 6-8 drops of methyl red reagent.
29


for Voges-Proskauer: Add 12 drops of Barritt's A, mix, 4 drops of
Barritt's B, mix.
Let sit, undisturbed, for at least 20 minutes.
Interpretation:
Methyl red
Within just a few seconds after adding methyl red reagent, you can see
the red-pink color of acid presence from glucose use..
Voges-Proskauer tests
The reagents MUST be added in the correct order, in the correct amounts,
and the tube must sit undisturbed, and open to the air (no cap) for at least
30 minutes (45 minutes is even better) as the light pink color intensifies at
the top of the tube (the reagents react with acetoin). Do not shake the
tube after sitting it down for the waiting period.
Antibiotic Sensitivity Testing
Antibiotic sensitivity testing is done to determine the susceptibility of bacteria to
various antibiotics. This standardized test is used to measure the effectiveness of
a variety of antibiotics on a specific organism in order to prescribe the most
suitable antibiotic therapy
There are several methods of doing the antimicrobial susceptibility testing. Of
those different method Disc Diffusion Technique is one widely followed in all
clinical microbiology laboratories of the globe. This technique is now well
standardized and recommended by NCCLS (National Committee of Clinical
Laboratory Standard) U S A.
A series of antibiotic-impregnated paper disks are placed on a plate inoculated to
form a bacterial lawn (even, confluent growth). The plates are incubated to allow
growth of the bacteria and time for the antibiotics to diffuse into the agar. If an
organism is susceptible to an antibiotic, a clear zone will appear around the disk
where the growth has been inhibited. The size of this zone of inhibition depends
on the sensitivity of the bacteria to the specific antibiotic and the antibiotic's ability
to diffuse through the agar
The Kirby-Bauer test must be carefully standarized. This means that a special
agar, Mueller-Hinton agar, is used along with a prescribed inoculum of broth. The
antibiotic disks are also standardized to contain a specific amount of antibiotic.
After 18 hours of
incubation at 35oC, the clear zones are measured. These are compared with
tables giving the interpretation of measurement for each antibiotic.
In order to determine if an antibiotic will be effective in treating the bacterial
infection, the zone of inhibition must be measured and compared to a standard
chart provided by NCCLS, USA. Compare the zone of inhibition of test result with
30
the Standard Chart and issue result as Sensitive, Partial sensitive or Resistant .
An organism is not considered to be sensitive to an antibiotic unless the zone of
no growth is matched with NCCLS Chart
The sensitivity test result
Reading & reporting
Wet mount microscopy of Intestinal parasites
Wet mount microscopy:
Procedure
1. A drop of normal saline (0.9% Sodium Chloride) is kept on a new slide.
2. A small amount of stool specimen is taken by use of wooden applicator
3. This stool is emulsified with the normal saline.
4. This mixed suspension should not very thick
5. A cover glass is kept over this mixed suspension
6. It is examined by low power objective of the microscope (X100) and
checked if necessary by high power objective (x 400)
7. Examination must be completed before drying of the preparation.
31
Concentration method / technique: This method is used when the ova, cysts,
larvae are in scant numbers
Types of Concentration method
1. Flotation method uses the high specific gravity of a solution to float the
lighter ova and cysts
a. Make saturated solution of Sodium chloride
b. Keep a little quantity amount 0.5 – 1ml into the penicillin vial or the
tube that can hold coverglass.
c. Keep stool sample about 3-4 times of matchstick head and emulsify
with the saturated solution of Sodium chloride
d. After smooth emulfication keep the saturated NaCl and mix it;
continue mixing this way to the neck of the vial of tube. The tube
must be in vertical position.
e. Keep cover glass in touch with the solution and wait for 30 minutes.
2. Sedimentation method : Modified Ridley-Allen method for sedimentation
technique.This technique uses formalin as a preservative and ether or ethyl
acetate as an extractor of fat and debris from faeces. Note: Ether is flammable
and formalin is an irritant
Materials.
a. Formalin water (100ml formaldehyde and 900ml distilled water)
b. Ether or ethyl acetate.
c. Mesh (425µm) brass wire filter, 3 inches in diameter (Endcott Sieves Ltd.
Lombard Road, London SW19 3BR). If this is not available ordinary nylon tea
strainers provides an alternative substitute (This is of low cost)
d. Small 3inches porcelain or stainless steel dish.
Method.
a. Using swab sticks, select a quantity of faeces (approximately 1g or pea size)
to include external and internal portions.
b. Place the faeces in a centrifuge tube containing 7 ml of 10% formalin.
c. Emulsify the faeces in the formalin and filter through the brass/nylon filter into
the dish. This filtration process allow the parasites to go through the solution.
d. Wash the filter and discard any lumpy residue.
e. Transfer the filtrate to a tube and add 3 ml of ether/ethyl acetate. Mix well on a
vortex mixer for 15 seconds or by hand for 1 minute.
32
f. Transfer back to the centrifuge tube and centrifuge at 3,000 rpm. for 2 minute.
g. Loosen the fatty plug with an orange stick and pour the supernatant away by
quickly inverting the tube.
h. Allow the fluid on the side of the tube to drain on to the deposit and mix well.
Transfer a drop to a slide for examination under a coverslip.
i. Use the x10 and x40 objectives to examine the whole of the deposit for ova
and cysts.
Note:
The important points to be considered when performing a concentration
technique are:
1. In a specimen the whole of the sample (equivalent to 1 gram of faeces) should
be concentrated and the whole of the deposit examined. This corresponds to
good practice with clinical
samples.
2. It is important to vortex the sample for at least
15 seconds after the addition of ether or ethyl acetate, as failure to do so may
result in excess deposit, thus obscuring ova and
cysts.
3. Adequate centrifugal force must be used
because if this is below the required value, there may be insufficient gravitational
force to sediment the ova and cysts. The centrifugal time is also critical, since the
ova and cysts may remain in suspension if the sample is not centrifuged for the
minimum required time. It is recommended that the sample be centrifuged at
1000g for 2 minute.
Examination of Blood Parasites
Preparation of thick and thin blood films :Thick films:- place a drop of blood in the middle of a clean microscope slide and
with the corner of a second slide spread the drop until it is about the size of a five
cent coin. The thickness should be such that it is just possible to see news print
through it.
Thin film :- Thin film are made as demonstrated ( also shown in the figure).
Allow the films to dry, do not leave on the bench in a laboratory which is not fly
proofed otherwise the film will be eaten
When the films are dry, fix and stain the films by any method of Romanosky
stain such as Giemsa stain, Field’s stain, Wright’s stain, Leishman stain etc but
remember the pH of the stain should be a slightly alkaline (pH 7.2) as an acid
stain may fail to show the parasites.
33
The films are stained by Giemsa stain : Giemsa is diluted (1/20) and keep
in the staining jar so that the film is in an upright position, this will allow any
debris to fall to the bottom of the jar. Stain for about 30 minutes, wash gently with
clean water and allow to dry. If available use a positive control.
Field's stain is also used because it is very quick. Field's stain comprises
two solutions; a polychrome methylene blue (A) and eosin (B). The solutions are
kept in covered staining jars.
Dip the dry but unfixed film into solution A for 1 or 2 seconds.
Remove from solution A and immediately rinse in clean water ( a 250ml beaker
with water gently flowing into it is suitable)
Dip the film into solution B for 1 or 2 seconds.
Rinse in clean water for a few seconds.
Place in a vertical position to dry.
Under the microscope examine the stained slide using an oil immersion
Staining for Mycobacteria
(Demonstration of AFB and Practice of AFB staining)
AFB staining procedure ( Also known as Ziehl Nelson’ stain):
The term AFB 'acid fast bacilli' refers to an organism's ability to retain the
carbolfushcin stain despite subsequent treatment with an ethanol-hydrochloric
acid mixture. The high lipid content (approximately 60%) of their cell wall makes
mycobacteria acid-fast.
1. Flame slides to heat fix
2. Flood the entire slide with Carbol Fuchsin
3. Ensure enough stain is added to keep the slides covered throughout the
entire staining step.
4. Using a Bunsen burner, heat the slides slowly until they are
steaming. Maintain steaming for 5 minutes by using low or intermittent
heat (i.e. by occasionally passing the flame from the Bunsen burner over
the slides)
5. Rinse the slide thoroughly with water
6. Flood the slide with 3% acid-alcohol and allow to decolorize for 5
minutes.Throughout the 5 minutes, continue to flood the slides with 3%
acid-alcohol until the slides are clear of stain visible to the naked eye.
7. Rinse the slide thoroughly with water and then drain any excess from the
slides.
8. Flood the slide with the counterstain, Methylene Blue. Keep the
counterstain on the slides for 1 minute.
9. Rinse the slide thoroughly with water
.Result : AFB – Red color bacilli; and Pus cells and other materials - Blue
34
AFB Staining : China modified method
1. A smear is made, dried and fixed by heat (flaming)
2. Flood the entire smear with dye # 1 for 3-5 minutes.
3. Rinse the slide with water
4. Flood the slide with dye #2 for one minute
5. Rinse the slide with water
6. Dry it and look by microscope using oil immersion objective
Note:
Dye #1
Basic red
4g
Carbolic acid
8g
90% Ethanol
20 ml
Water
100ml
Dye#2
Methylene blue
1g
Conc. Sulphuric acid
20ml
Ethanol
30ml
Water
50ml
Mycobacteria culture Culture : a brief introduction
Lowenstein-Jensen medium (For culture of Mycobacterium tuberculosis)
After digestion and concentration by treatment with sodium hydrochloride
solution, the mucopurulent sputum material is cultured on this selective media,
and then incubated for up to 8 weeks at 36o C.
Result : The Positive growth appears pigmented (Yellow pigment) which is
rough and hard to remove from the growth medium
Culture of AFB is carried out in safety cabinet because the bacteria can remain
viable for several days or week; and easily transmitted to the staffs.
An introduction to Mycology practical
Dermatophytes are fungi that can cause infections of the skin, hair, and nails.
The organisms colonize the keratin tissues and utilize keratin.
These infections are also known as ringworm or tinea. Some time the organisms
do invade the subcutaneous tissues
There are three genera of dermatophytes,
Trichophyton
Microsporum
Epidermophyton
35
Specimen collection : Skin, nail or hair
scrapings or cutting collected in nonsticky glazed paper or sterile container
e.g. petriplate in case of skin and nail
Collection Procedure : Skin, nails and hair
Cleanse the area with 70% alcohol prior to specimen collection.
Skin should be taken from the active border of the lesion
Nail should be from a subsurface portion of the infected nail.
scrapings or cutting collected in nonsticky glazed paper or sterile container
e.g. petriplate in case of skin and nail.
Hair should be plucked, not cut, from the edge of lesion. Take 10-12 hairs.
Specify the source of the specimen and include any pertinent clinical
information.
Wet mount microscopy (Direct Examination)
• A small sample of the specimen is selected for direct microscopic
examination and investigated for the presence of fungal elements.
• The specimen is mounted in a small amount of potassium hydroxide or
calcofluor white.
• The KOH slides are gently heated and allowed to clear for 30 to 60
minutes before examining on a light or phase contrast microscope.
Note : 10% KOH is for skin scraped material
15% KOH is for nail pieces
20% KOH is for hair pieces
Result
When present in the direct examination dermatophytes appear as hyaline (nonpigmented), septated elements. Fungal hyphae appears as branched filaments
making up a mycelium
When hair is involved the arthroconidia may be found on the periphery of the hair
shaft (ectothrix) or within the shaft (endothrix)
Cultures are incubated up to 3 weeks before a final report is issued. The cultures
are incubated at 30°C and examined frequently for 4 weeks. If the dimorphic
fungi is suspected the there should be incubation at 37o C also.
36
Dermatophytosis
Skin KOH
Serological test
Introduction
Serological tests are done to demonstrate either antigens or antibodies in the
serum. The VDRL / RPR / TPHA / HIV testing are the serological tests. Usually
the specific antibody is detected in the serological testing. Enzyme linked
immunosorbent assay (ELISA) is one method extensively followed in many
laboratories. In ELISA technique the microplate in which the wells are coated
with the specific antigens is used.
Technique of drawing the blood:
1. The patient is seated and the fore arm is stretched on the table with
supporting cushion if necessary
2. Using the tourniquet on the upper arm the patient is told to clench the fist.
37
3. The median cubital vein is identified
4. Clean the area with 75% alcohol swab.(Ethanol / Isopropanol)
5. The vein is palpated if required
6. Insert the needle with bevel side up and parallel to the vein
7. Withdraw the 5ml. of blood at least
8. Tourniquet is released and the patient is told to open the hand
Alcohol swab is applied and pressed for some time or Adhesive tape is applied.
Transfer of blood:
1. Blood is transferred slowly into the well cleaned and dried tube which is
already labeled with patient identity.
2. The needle is destroyed with use of needle destroyer and then discard
into the puncture proof container.
3. The tube is centrifuged at slow speed (1500-2000rpm) for 5 minutes
4. The separated serum is transferred into a cleaned tube for RPR, TPHA
and HIV testing as per necessary; and the remaining serum is transferred
into another sterile plastic vials( e.g. cryotubes / eppendorf tubes) for
storage at -20oC labeled with patient’s identity number
The factors causing of haemolysis
(Haemolysed blood is not suitable for serological test)
Haemolysis often occurs in
1. blood taking by too small bore of a needle
2. expelling blood quickly from syringe
3. forced suction of blood in the syringe during blood collection.
4. vigorous shaking of blood
5. centrifuging blood sample at a high speed before clotting
6. freezing and thawing of blood
7. unclean tubes with residual detergents
water in the tube
Precautionary measures in practical of blood specimens
1. Avoid skin contact with blood.
2. Any blood that comes in contact with floor must be decontaminated with
hypo chlorite
3. All the other details of the laboratory procedures will be described in this
procedure manual in the following chapter.
4. Avoid recapping needles because there is high risk of infection through
needle stick injuries. If recapping is necessary, recap using only one hand
(i.e. by placing the cap on the table and inserting the needle inside the
cap)
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Rapid Plasma Reagin (RPR )Test
Introduction
The RPR test is the primary screening test for syphilis antibodies in serum.
This is non-treponemal flocculation test . Treponema pallidum, a causative
organism of Syphilis, produces antibody in response to tissue damage. This test
if positive the titration is required. The titration indicate the possibility of current
activity and post treatment. The RPR positive test is confirmed by TPHA.
In RPR technology the antibody is detected by cardiolipin antigen; and
therefore this antibody also known as cardiolipin antibody. In the earlier years
Wasserman reaction test was done for sorodiagnosis of Syphilis; later on VDRL
test came in the market; now RPR test is followed by many laboratories because
- it is very easy to perform
- it is not necessary to heat serum
- this test can be done even by plasma.
Before testing the entire reagent kept in refrigerator (2 –8o C) is taken and
allowed to stand outside to acquire room temperature.
Test Procedure
1. With use of a clean and dried Pasteur pipette with rubber teat or disposable
plastic dropper dispense one drop of serum on to a circle of the test card and
spread it using an applicator stick.
2. Keep one drop of antigen suspension using a dispensing dropper (provided in
the test kit) into the test card
3. Rotate this test card at 100 rpm for 8 minutes keeping on an rotator
4. As a part of quality control every test run should include a control serum
Interpretation of results :
Medium and large aggregates
Finely dispersed aggregates
No aggregate (i.e. smooth appearance)
Reactive
weakly reactive
No reactive
Note: False positive reaction may occur in diseases for example Tuberculosis,
Leprosy, Malaria, Infectious mononucleosis, Cancer etc. Because of this reason
a specific treponemal test should be performed i.e. TPHA for the detection of
specific Antitreponemal antibodies.
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Widal test
This is a serological test; method of doing varies depending on the manufactures.
Therefore manufacture’s testing kit procedure (leaflet) is to followed. The
principle of the test is similar to RPR testing.
The Widal test is the traditional serologic test used for the diagnosis of typhoid
fever. The test measures agglutinating antibodies against flagellar (H) and
somatic (O) antigens of S typhi.
In acute infection, O antibody appears first, rising progressively and later on falls
often disappearing within a few months. High O antibody titers generally indicate
acute infection. Rising or high O antibody titers generally indicate acute infection.
The H antibody appears slightly later and persists longer. The elevations of H
antibody help to identify the type of enteric fever.
Numerous studies have shown that the sensitivity, specificity, and predictive
values of Widal test vary rendering the test's value to the clinician questionable.
This wide variation is caused by differences in patients’ stages of infection and
antigens variations, The Widal reaction is indicative of typhoid fever in only 4060% of patients.
These are other serological method is available such as ELISA method (enzymelinked immunosorbent assay) for immunoglobulin M (IgM) and immunoglobulin G
antibodies to S typhi polysaccharide, the monoclonal antibodies against S typhi
flagellin and DNA probes and these are confirmatory test.
The other lab. tests (Blood culture)
Clinical diagnosis is supported by tests that identify Salmonella antibodies and is
then confirmed by isolation of the organism and this is done by blood culture. The
definitive diagnosis of typhoid fever requires isolation of the organism from blood
or bone marrow. With culture-positive rates of approximately 90%, the most
sensitive method of isolating S typhi is obtaining a bone marrow aspirate (BMA)
culture. BMA and blood are cultured in a selective medium, such as 10%
aqueous oxgall, or a nutritious medium, such as tryptic soy broth, and are
incubated at 37°C for at least 7 days before reporting negative. Subcultures are
made daily to one selective medium, such as MacConkey agar, and one
inhibitory medium, such as Salmonella-Shigella agar
40
Some illustrations of pictures of laboratory concerns
41
42
43
Blood agar plate
Different hemolytic reactions
Staphylococcus aureus
showing beta haemolysis in Blood agar and
colonial characteristic in Nutrient agar
44
Lactose fermenters will appear pink, non-lactose
fermenters will appear colorless
45
Triple Sugar Iron Agar (TSI)
TSI agar contains
peptone,
glucose,
sucrose,
lactose, and
thiosulfate
46
MOTILITY TEST
• OrganismTest Result
• 1. Ps aeruginosa
• 2. S aureus
• 3. B subtilis
Motile
Non-motile
Motile
Egg of Ascaris lumbricoides, Ankylostoma duodenale,
Enterobius vermacularis, Trichuris trichuria
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Egg of Taena and proglottid of T sagina & T solium
egg of Hymenolepis nana
Trophozoite and cyst
Giardia lamblia & Entamoeba hystolytica
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49
Microfilaria bancrofti and brugia
50
Ziehl Neelson’s staining
(AFB Staining)
Continued AFB Staining
51
Mucor / Penicillium / Aspergillus
Microsporum gypseum &
Trichophyton mentagrophytes
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54
55
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