Pour Plate Method: Procedure, Uses,
(Dis) Advantages
Pour plate method is usually the method of choice for counting the number of
colony-forming bacteria present in a liquid specimen. Because the sample is
mixed with the molten agar medium, a larger volume can be used than with
the spread plate. In this method, a fixed amount of inoculum (generally 1 ml)
from a broth/sample is placed in the center of a sterile Petri dish using a sterile
pipette. Molten cooled agar (approx. 15mL) is then poured into the Petri dish
containing the inoculum and mixed well. After the solidification of the agar, the
plate is inverted and incubated at 37°C for 24-48 hours.
Microorganisms will grow both on the surface and within the medium. Colonies
that grow within the medium generally are small in size and maybe confluent;
the few that grow on the agar surface are of the same size and appear like those
on a streak plate. Each (both large and small) colony is carefully counted (using
magnifying colony counter if needed). Each colony represents a “colony-forming
unit” (CFU).
The number of microorganisms present in the particular test sample is
determined using the formula:
CFU/mL= CFU * dilution factor * 1/aliquot
For accurate counts, the optimum count should be within the range of 30-300
colonies/plate. To insure a countable plate a series of dilutions should be
plated.The pour plate method of counting bacteria is more precise than
the streak plate method, but, on average, it will give a lower count as heatsensitive microorganisms may die when they come contact with hot, molten agar
medium.
Uses:
The pour plate technique can be used to determine the number of microbes/mL
in a specimen. It has the advantage of not requiring previously prepared plates,
and is often used to assay bacterial contamination of food stuffs.
Materials and Equipments
1. Test sample
2. Plate count agar (PCA) or nutrient agar
3. Hot water bath 45°C
4. Sterile Petri dishes
5. Flame
6. Colony counter with magnifying glass
7. Sterile capped 16*150 mm test tubes
8. Pipettes of various sizes (e.g. 01, 1.0 and 2.0 mL)
Procedure of Pour plate technique
1. Prepare the dilution of the test sample expected to contain between 30300 CFU/mL. (Follow serial dilution technique)
2. Inoculate labeled empty petri dish with specified mL (0.1 or 1.0 mL) of
diluted specimen
Note: for the detailed description regarding use of pipette, inoculation of sample,
dilution technique etc, follow the reference 1.
Pouring the molten agar and incubation
1. Collect one bottle of sterile molten agar (containing 15 mL of melted Plate
Count Agar or any other standard culture media) from the water bath
(45°C).
2. Hold the bottle in the right hand; remove the cap with the little finger of
the left hand.
3. Flame the neck of the bottle.
4. Lift the lid of the Petri dish slightly with the left hand and pour the sterile
molten agar into the Petri dish and replace the lid.
5. Flame the neck of the bottle and replace the cap.
6. Gently swirl the plate on the benchtop to mix the culture and the medium
thoroughly. Ensure that the medium covers the plate evenly and do not
slip the agar over the edge of the Petri dish.
7. Allow the agar to completely gel without disturbing it, it will take
approximately 10 minutes.
8. Seal and incubate the plate in an inverted position at 37°C for 24-48 hours.
Results:
After 24-48 hours, count all the colonies (again: note that the embedded colonies
will be much smaller than those which happen to form on the surface). A
magnifying colony counter can aid in counting small embedded colonies.
Calculate CFU/mL using the formula: CFU/mL= CFU * dilution factor * 1/aliquot
(the volume of diluted specimen (aliquot) is either 0.1 or 1.0 mL)
Disadvantages of Pour plate method
1. Preparation for the pour plate method is time-consuming compared with
the streak plate/and or spread plate technique.
2. Loss of viability of heat-sensitive organisms coming into contact with hot
agar.
3. Embedded colonies are much smaller than those which happen to be on
the surface. Thus, one must be careful to count these so that none are
overlooked.
4. Reduced growth rate of obligate aerobes in the depth of the agar.
Blood Smear Test: Procedure, Staining & Interpretation
The Blood Smear Test
The blood smear test plays an important role in the speedy diagnosis of certain
infections or diseases. This test uses a drop of blood spread onto a glass
microscope slide that is then treated with a colored stain and examined using a
microscope. The blood smear test shows a sample of blood components
including platelets, leukocytes (white blood cells), and erythrocytes (red blood
cells) that are present in plasma, the fluid part of blood. A blood smear test is
typically used as a follow-up test after abnormal results were shown in a
complete blood count test (CBC). The blood smear is a vital diagnostic aid.
Procedure
A blood smear test is performed by first obtaining a 5 mL blood sample from the
patient. The patient should be educated about the procedure before taking a
sample from their vein, or, less often, from a capillary. The blood sample is stored
in a bottle containing an anticoagulant called ethylene diamine tetra-acetic acid
(EDTA). Anticoagulants prevent the blood from clotting. This sample is sent to
the laboratory for testing within two hours of collection.
Preparation of the microscope slide is performed by trained personnel such as a
pathologist, medical laboratory technologist, hematologist, or laboratory
assistant. This personnel uses a base slide, a blood spreader slide, and a pipette
or capillary tube. The wedge method is the most common way to prepare the
slide for testing. Using this method, a mixed drop of blood 1 to 2 mm in diameter
is placed in the center line about 1/4 inch from the edge of the microscope slide
using a pipette or capillary tube. The slide containing the blood drop is called the
base slide.
Another microscope slide called a spreader slide is used. This slide should have
chipped edges, along with another smooth end. The side of the spreader slide
with chipped edges is placed on the original slide (base slide) in front of the
blood and moved backwards to touch the blood. This makes the blood spread
along the base of the slide.
The smear is made with the spreader inclined at an angle of approximately 30° to
the blood. The smear should cover two-thirds of the base slide and should have a
feathered end. The smear should then be air dried. The frosted end of the slide
should be labeled with the patient's name, identification number, and date. The
dried smear is then fixed with methanol or ethyl alcohol and stained.
The smear is covered with stain for approximately ten minutes, then diluted with
water and allowed an additional ten minutes for the cells to properly stain.
Following the stain application, the slide is rinsed under running water. The slide
should be wiped underneath with cotton to remove excess stain. Finally, the slide
is placed on a rack to dry.
Tips for Perfect Tests
It is best to follow instructions for the procedure in obtaining, creating, and
staining a blood smear. It is also better to create a new smear if the procedure is
compromised than it is to interpret an inadequate smear. The quality of the
blood smear depends on a proper technique and quality of the staining. Some
additional guidelines should be followed to create the best blood smear.
At least two slides should be made during testing. A smear will be too thin if the
spreader slide is moved too quickly or if the angle of the spreader is less than
30°. Conversely, the smear will be too thick if the spreader is moved slowly or if
the angle is greater than 30°. Large blood drops may extend the smear over too
much of the base slide, while a small drop can be insufficient for the smear. The
stain needs adequate time with the sample to avoid over-staining or understaining. If a sample is over-stained, debris might show up in the sample. This can
also happen if the stain is not washed enough with running water
The Triple Sugar Iron (TSI) Test –
Principle, Procedure, Uses and
Interpretation
Most bacteria have the ability to ferment carbohydrates, particularly sugars.
Among them, each bacteria can ferment only some of the sugars, while it
cannot ferment the others. Thus, the sugars, which a bacteria can ferment and
the sugars, which it cannot is the characteristic of the bacteria and thus an
important criterion for its identification.
The Triple Sugar Iron (TSI) test is a microbiological test named for its ability to
test a microorganism’s ability to ferment sugars and to produce hydrogen
sulfide. An agar slant of a special medium with multiple sugars constituting a
pH-sensitive dye (phenol red), 1% lactose, 1% sucrose, 0.1% glucose, as well
as sodium thiosulfate and ferrous sulfate or ferrous ammonium sulfate is used
for carrying out the test. All of these ingredients when mixed together and
allowed solidification at an angle result in a agar test tube at a slanted angle.
The slanted shape of this medium provides an array of surfaces that are either
exposed to oxygen-containing air in varying degrees (an aerobic environment)
or not exposed to air (an anaerobic environment) under which fermentation
patterns of organisms are determined.
Objective
To determine the ability of an organism to ferment glucose, lactose, and
sucrose, and their ability to produce hydrogen sulfide.
Principle
The triple sugar- iron agar test employing Triple Sugar Iron Agar is designed
to differentiate among organisms based on the differences in carbohydrate
fermentation patterns and hydrogen sulfide production. Carbohydrate
fermentation is indicated by the production of gas and a change in the colour
of the pH indicator from red to yellow.
To facilitate the observation of carbohydrate utilization patterns, TSI Agar
contains three fermentative sugars, lactose and sucrose in 1% concentrations
and glucose in 0.1% concentration. Due to the building of acid during
fermentation, the pH falls. The acid base indicator Phenol red is incorporated
for detecting carbohydrate fermentation that is indicated by the change in
color of the carbohydrate medium from orange red to yellow in the presence
of acids. In case of oxidative decarboxylation of peptone, alkaline products
are built and the pH rises. This is indicated by the change in colour of the
medium from orange red to deep red. Sodium thiosulfate and ferrous
ammonium sulfate present in the medium detects the production of hydrogen
sulfide and is indicated by the black color in the butt of the tube.
To facilitate the detection of organisms that only ferment glucose, the glucose
concentration is one-tenth the concentration of lactose or sucrose. The
meagre amount of acid production in the slant of the tube during glucose
fermentation oxidizes rapidly, causing the medium to remain orange red or
revert to an alkaline pH. In contrast, the acid reaction (yellow) is maintained in
the butt of the tube since it is under lower oxygen tension.
After depletion of the limited glucose, organisms able to do so will begin to
utilize the lactose or sucrose. To enhance the alkaline condition of the slant,
free exchange of air must be permitted by closing the tube cap loosely.
Media:
TSI Agar
Enzymatic digest of casein (5 g), enzymatic digest of animal tissue (5 g), yeast
enriched peptone (10 g), dextrose (1 g), lactose (10 g) sucrose (10 g), ferric
ammonium citrate (0.2 g), NaCl (5 g), sodium thiosulfate (0.3 g), phenol red
(0.025 g), agar (13.5 g), per 1000 mL, pH 7.3.
Method
1. With a straight inoculation needle, touch the top of a well-isolated
colony.
2. Inoculate TSI by first stabbing through the center of the medium
to the bottom of the tube and then streaking the surface of the
agar slant.
3. Leave the cap on loosely and incubate the tube at 35°-37°C in
ambient air for 18 to 24 hours.
4. Examine the reaction of medium.
Expected Results





An alkaline/acid (red slant/yellow butt) reaction: It is indicative
of dextrose fermentation only.
An acid/acid (yellow slant/yellow butt) reaction: It indicates
the fermentation of dextrose, lactose and/or sucrose.
An alkaline/alkaline (red slant, red butt) reaction: Absence of
carbohydrate fermentation results.
Blackening of the medium: Occurs in the presence of H2
Gas production: Bubbles or cracks in the agar indicate the
production of gas ( formation of CO2and H2)
Triple sugar iron agar. A, Acid slant/acid butt with gas, no H2S
(A/A). B, Alkaline slant/acid butt, no gas, H2S-positive (K/A H2S+). C, Alkaline
slant/alkaline butt, no gas, no H2S (K/K). D, Uninoculated tube.
Uses


The test is used primarily to differentiate members of the
Enterobacteriaceae family from other gram-negative rods.
It is also used in the differentiation among Enterobacteriaceae on
the basis of their sugar fermentation patterns.
Limitations


It is recommended that biochemical, immunological, molecular, or
mass spectrometry testing be performed on colonies from pure
culture for complete identification.
It is important to stab the butt of the medium. Failure to stab the
butt invalidates this test. The integrity of the agar must be
maintained when stabbing. Caps must be loosened during this
test or erroneous results will occur.




TSI Agar must be read within the 18-24 hour stated incubation
period. A false-positive reaction may be observed if read too early.
A false-negative reaction may be observed if read later than 24
hours.
An organism that produces hydrogen sulfide may mask acid
production in the butt of the medium. However, hydrogen sulfide
production requires an acid environment, thus the butt portion
should be considered acid.
TSI is not as sensitive in detecting hydrogen sulfide in comparison
to other iron containing mediums, such as Sulfide Indole Motility
(SIM) Medium.
Certain species or strains may give delayed reactions or
completely fail to ferment the carbohydrate in the stated manner.
Oxidase test: Principle, Procedure,
Results
The oxidase test is used to identify bacteria that produce cytochrome c
oxidase, an enzyme of the bacterial electron transport chain. When present,
the cytochrome c oxidase oxidizes the reagent (tetramethyl-pphenylenediamine dihydrochloride) to indophenols, a purple or dark blue
color end product. When the enzyme is not present, the reagent remains
reduced and is colorless.
Mechanism of the Cytochrome Oxidase Reaction
All bacteria that are oxidase-positive are aerobic and can use oxygen as a
terminal electron acceptor in respiration. This does NOT mean that they are
strict aerobes. Bacteria that are oxidase-negative may be anaerobic, aerobic,
or facultative; the oxidase negative result just means that these organisms do
not have the cytochrome c oxidase that oxidizes the test reagent. They may
respire using other oxidases in electron transport.
Test requirements for Oxidase test

Moist filter paper with the substrate (1% tetramethyl-pphenylenediamine dihydrochloride), or commercially prepared paper
disk, wooden wire, or platinum wire.

Kovács oxidase reagent (1% tetra-methyl-p-phenylenediamine
dihydrochloride, in water). Store refrigerated in a dark bottle for no
longer than 1 week.
Procedure of Oxidase test
Oxidase test can be performed in various ways. These include, but are not limited to, the filter
paper test, filter paper spot test, direct plate method, and test tube method.
Filter Paper Test Method
1. Soak a small piece of filter paper in 1% Kovács oxidase reagent and let dry.
2. Use a loop and pick a well-isolated colony from a fresh (18- to 24- hour culture)
bacterial plate and rub onto treated filter paper
3. Observe for color changes.
Results
 Oxidase positive: color changes to dark purple within 5 to 10 seconds.
 Delayed oxidase-positive: color changes to purple within 60 to 90
seconds.
 Oxidase negative: color does not change or it takes longer than 2
minutes.
Filter Paper Spot Method
1. Use a loop and pick a well-isolated colony from a fresh bacterial plate
and rub it onto a small piece of filter paper.
2. Place 1 or 2 drops of 1% Kovács oxidase reagent on the organism
smear.
3. Observe for color changes.
Results
 Oxidase positive: color changes to dark purple within 5 to 10 seconds.
 Delayed oxidase-positive: color changes to purple within 60 to 90
seconds.
 Oxidase negative: color does not change or it takes longer than 2
minutes.
Direct Plate Method
1. Grow a fresh culture (18 to 24 hours) of bacteria on nutrient agar
or trypticase soy agar using the streak plate method so that wellisolated colonies are present.
2. Place 1 or 2 drops of 1% Kovács oxidase reagent on the organisms.
3. Do not invert or flood plate.
4. Observe for color changes.
Results
 Oxidase positive: color changes to dark purple within 5 to 10 seconds.
 Delayed oxidase-positive: color changes to purple within 60 to 90
seconds.
 Oxidase negative: color does not change or it takes longer than 2
minutes.
Test Tube Method
1. Grow a fresh culture (18 to 24 hours) of bacteria in 4.5 ml of nutrient
broth (or standard media that does not contain a high concentration of
sugar).
2. Add 0.2 ml of 1% α-naphthol, then add 0.3 ml of 1%
paminodimethylaniline oxalate (Gaby and Hadley reagents).
3. Observe for color changes.
Results
 Oxidase positive: color changes to blue within 15 to 30 seconds.
 Delayed oxidase-positive: color changes to purple within 2 to 3 minutes.
 Oxidase negative: no change in color
Uses of oxidase test
 Oxidase test is most helpful in screening colonies suspected of being a
member of the Enterobacteriaceae family; all the members of the
Enterobacteriaceae family including E. coli are oxidase negative.
 To avoid misidentification, perform an oxidase test on all Gram-negative
rods. Oxidase test is especially important in
separating Aeromonas from Enterobacteriaceae.
Note: If you see swarming colonies in a culture media, do not perform
oxidase test, as its unique characteristics of Proteus spp, which are oxidase
negative.
 Oxidase test is used as a major characteristic for the identification of
Gram-negative rods that are not in the Enterobacteriaceae family.
Colonies suspected of belonging to other genera Aeromonas,
Pseudomonas, Neisseria, Campylobacter, and Pasteurella are oxidase
positive.
Gram-negative diplococci give a positive reaction. All members of the
genus Neisseria are oxidase positive. Moraxella spp. that are either
Gram-negative diplococci or coccobacilli are also oxidase-positive.
Quality Control
Bacterial species showing positive and negative reactions should be run as
controls at frequent intervals. The following are suggested:
A. Oxidase positive: Pseudomonas aeruginosa
B. Oxidase negative: Escherichia coli
Precautions and Limitations:
 Timing is critical to accurate testing.
 Use fresh reagents, no older than 1 week, older reagents can
autooxidize thus giving erroneous results. Do not use if reagent or filter
paper is purple.
 Do not test organisms growing on media that contain glucose or dyes
(e.g., MacConkey agar or EMB agar).
 Do not use nickel-base alloy wires containing chromium and iron
(nichrome) to pick the colony and make smear as this may give falsepositive results.
 Bacteria grown on media containing dyes may give aberrant results.
 Older cultures are less metabolically active so may give false-negative
results within the mentioned observation time.
Try this class experiment to detect the presence of enzymes as they
catalyse the decomposition of hydrogen peroxide
Enzymes are biological catalysts which increase the speed of a chemical
reaction. They are large protein molecules and are very specific to certain
reactions. Hydrogen peroxide decomposes slowly in light to produce oxygen
and water. The enzyme catalase can speed up (catalyse) this reaction.
In this practical, students investigate the presence of enzymes in liver, potato
and celery by detecting the oxygen gas produced when hydrogen peroxide
decomposes. The experiment should take no more than 20–30 minutes.
Equipment
Apparatus
 Eye protection
3
 Conical flasks, 100 cm , x3
3
 Measuring cylinder, 25 cm
 Bunsen burner

Wooden splint
 A bucket or bin for disposal of waste materials
Chemicals
 Hydrogen peroxide solution, ‘5 volume’
 Small pieces of the following (see note 4):
o Liver
o Potato
o Celery
Health, safety and technical notes
1. Read our standard health and safety guidance.
2. Wear eye protection throughout. Students must be instructed NOT to
taste or eat any of the foods used in the experiment.
3. Hydrogen peroxide solution, H2O2(aq) – see CLEAPSS Hazcard
HC050 and CLEAPSS Recipe Book RB045. Hydrogen peroxide solution of
‘5 volume’ concentration is low hazard, but it will probably need to be
prepared by dilution of a more concentrated solution which may be
hazardous.
4. Only small samples of liver, potato and celery are required. These should
be prepared for the lesson ready to be used by students. A disposal bin
or bucket for used samples should be provided to avoid these being put
down the sink.
Procedure
1. Measure 25 cm3 of hydrogen peroxide solution into each of three
conical flasks.
2. At the same time, add a small piece of liver to the first flask, a small
piece of potato to the second flask, and a small piece of celery to the
third flask.
3. Hold a glowing splint in the neck of each flask.
4. Note the time taken before each glowing splint is relit by the evolved
oxygen.
5. Dispose of all mixtures into the bucket or bin provided.
Teaching notes
Some vegetarian students may wish to opt out of handling liver samples, and
this should be respected.
Before or after the experiment, the term enzyme will need to be introduced.
The term may have been met previously in biological topics, but the notion

that they act as catalysts and increase the rate of reactions may be new.
Similarly their nature as large protein molecules whose catalytic activity can be
very specific to certain chemical reactions may be unfamiliar. The name
catalase for the enzyme present in all these foodstuffs can be introduced.
To show the similarity between enzymes and chemical catalysts, the teacher
may wish to demonstrate (or ask the class to perform as part of the class
experiment) the catalytic decomposition of hydrogen peroxide solution by
manganese(IV) oxide (HARMFUL – see CLEAPSS Hazcard HC060).
If students have not performed the glowing splint test for oxygen for some
time, they may need reminding of how to do so by a quick demonstration by
the teacher.
Additional information
This is a resource from the Practical Chemistry project, developed by the
Nuffield Foundation and the Royal Society of Chemistry. This collection of over
200 practical activities demonstrates a wide range of chemical concepts and
processes. Each activity contains comprehensive information for teachers and
technicians, including full technical notes and step-by-step procedures.
Practical Chemistry activities accompany Practical Physics and Practical
Biology.
Gram Staining: Principle, Procedure, Interpretation, Examples and Animation
Last updated: June 12, 2018 by Sagar Aryal
Gram Staining is the common, important, and most used differential staining
techniques in microbiology, which was introduced by Danish Bacteriologist
Hans Christian Gram in 1884. This test differentiate the bacteria into Gram
Positive and Gram Negative Bacteria, which helps in the classification and
differentiations of microorganisms.
Principle of Gram Staining
When the bacteria is stained with primary stain Crystal Violet and fixed by the
mordant, some of the bacteria are able to retain the primary stain and some
are decolorized by alcohol. The cell walls of gram positive bacteria have a thick
layer of protein-sugar complexes called peptidoglycan and lipid content is
low. Decolorizing the cell causes this thick cell wall to dehydrate and shrink,
which closes the pores in the cell wall and prevents the stain from exiting the
cell. So the ethanol cannot remove the Crystal Violet-Iodine complex that is
bound to the thick layer of peptidoglycan of gram positive bacteria and
appears blue or purple in colour.
In case of gram negative bacteria, cell wall also takes up the CV-Iodine
complex but due to the thin layer of peptidoglycan and thick outer layer which
is formed of lipids, CV-Iodine complex gets washed off. When they are
exposed to alcohol, decolorizer dissolves the lipids in the cell walls, which
allows the crystal violet-iodine complex to leach out of the cells. Then when
again stained with safranin, they take the stain and appears red in color.
Reagents Used in Gram Staining
 Crystal Violet, the primary stain
 Iodine, the mordant
 A decolorizer made of acetone and alcohol (95%)
 Safranin, the counterstain
Procedure of Gram Staining
1. Take a clean, grease free slide.
2. Prepare the smear of suspension on the clean slide with a loopful of
sample.
3. Air dry and heat fix
4. Crystal Violet was poured and kept for about 30 seconds to 1 minutes
and rinse with water.
5. Flood the gram’s iodine for 1 minute and wash with water.
6. Then ,wash with 95% alcohol or acetone for about 10-20 seconds and
rinse with water.
7. Add safranin for about 1 minute and wash with water.
8. Air dry, Blot dry and Observe under Microscope.
Examples
Gram Positive Bacteria: Actinomyces, Bacillus, Clostridium, Corynebacterium,
Enterococcus, Gardnerella, Lactobacillus, Listeria, Mycoplasma, Nocardia,
Staphylococcus, Streptococcus, Streptomyces ,etc.
Gram Negative Bacteria: Escherichia coli (E. coli), Salmonella, Shigella, and
other
Enterobacteriaceae, Pseudomonas,Moraxella, Helicobacter, Stenotrophomonas,
Bdellovibrio, acetic acid bacteria, Legionella etc
Blood Agar- Composition, Preparation, Uses and Pictures
Last updated: October 26, 2018 by Sagar Aryal
Blood Agar (BA) are enriched medium used to culture those bacteria or
microbes that do not grow easily. Such bacteria are called “fastidious” as they
demand a special, enriched nutritional environment as compared to the
routine bacteria. Blood Agar is used to grow a wide range of pathogens
particularly those that are more difficult to grow such as Haemophilus
influenzae, Streptococcus pneumoniae and Neisseria species. It is also required
to detect and differentiate haemolytic bacteria,
especially Streptococcus species. It is also a differential media in allowing the
detection of hemolysis (destroying the RBC) by cytolytic toxins secreted by
some bacteria, such as certain strains of Bacillus, Streptococcus, Enterococcus,
Staphylococcus, and Aerococcus.
Blood agar can be made selective for certain pathogens by the addition
of antibiotics, chemicals or dyes. Examples includes crystal violet blood agar to
select Streptococcus pyogens from throat swabs, and kanamycin or neomycin
blood agar to select anaerobes from pus.
Composition of Blood Agar
 0.5% Peptone
 0.3% beef extract/yeast extract
 1.5% agar
 0.5% NaCl
 Distilled water
(Since Blood Agar is made from Nutrient Agar, above is the composition
of Nutrient Agar)
 5% Sheep Blood
 pH should be from 7.2 to 7.6 (7.4)
Preparation of Blood Agar
1. Suspend 28 g of nutrient agar powder in 1 litre of distilled water.
2. Heat this mixture while stirring to fully dissolve all components.
3. Autoclave the dissolved mixture at 121 degrees Celsius for 15 minutes.
4. Once the nutrient agar has been autoclaved, allow it to cool but not
solidify.
5. When the agar has cooled to 45-50 °C, Add 5% (vol/vol) sterile
defibrinated blood that has been warmed to room temperature and mix
gently but well.
6. Avoid Air bubbles.
7. Dispense into sterile plates while liquid.
Uses of Blood Agar
1. Blood Agar is a general purpose enriched medium often used to grow
fastidious organisms
2. To differentiate bacteria based on their hemolytic properties (βhemolysis, α-hemolysis and γ-hemolysis (or non-hemolytic)).
Read more about Haemolysis and its types.
Pictures of Blood Agar
Microbiology BIOL 275 Dr. Eby Bassiri ebassiri@sas.upenn.edu 1
PREPARATION OF MEDIA I. OBJECTIVES • To become familiar with the
necessary nutritional and environmental factors for culturing microorganisms
in the laboratory. • To understand the decontamination or sterilization process
using an autoclave. • To learn the procedures used in preparing media needed
for culturing microorganisms. II. INTRODUCTION Microorganisms depend on
a number of factors such as nutrients, oxygen, moisture and temperature to
grow and divide. In the laboratory, except for the above factors, the culture
medium should be sterile and contamination of a culture with other organisms
should be prevented. Let us briefly discuss a few of the more important factors
for the growth of microorganisms. Nutrients A microbiological culture
medium must contain available sources of hydrogen donors and acceptors,
carbon, nitrogen, sulfur, phosphorus, inorganic salts and, in certain cases,
vitamins or other growth-promoting substances. These were originally
supplied in the form of meat infusions that were, and still are in certain cases,
widely used in culture media. Beef or yeast extracts can replace meat infusions.
The addition of peptone provides a readily available source of nitrogen and
carbon. Peptone is used in culture media to mainly supply nitrogen. Most
organisms are capable of utilizing the amino acids and other simpler
nitrogenous compounds present in peptone. Thus, in many cases, the
complicated infusion media can be replaced by simpler media prepared by
using the proper peptones in place of the meat infusions. Certain bacteria
require the addition of other nutrients, such as serum, blood, etc. to the
culture medium upon which they are to be propagated. Carbohydrates may
also be desirable at times, and certain salts such as calcium, manganese,
magnesium, sodium, and potassium seem to be required. Dyes may be added
to media as indicators of metabolic activity or for their selective inhibitory
powers. Growth promoting substances of a vitamin-like nature are essential or
assist greatly in the development of certain types of bacteria. Many of these
substances are given for individual bacteria in Bergey's Manual of
Determinative Bacteriology (Incidentally, this is a reference text that you
should familiarize yourself with when working with microorganisms.)
Microbiology BIOL 275 Dr. Eby Bassiri ebassiri@sas.upenn.edu 2 Oxygen Most
bacteria are capable of growth under ordinary conditions of oxygen tension.
Certain types, however, are capable of deriving their oxygen from various
substrates. The aerobic organisms require the free admission of air, while the
anaerobes grow only in the absence of atmospheric oxygen. Between these
two groups are the microaerophiles that develop best under partial anaerobic
conditions and the facultative anaerobes that develop under aerobic as well as
anaerobic conditions. It is easy to provide oxygen to aerobic and facultative
anaerobic and even microaerophilic organisms; however, special gadgetry is
required to exclude the atmospheric oxygen and provide an anaerobic
condition. Such conditions are obtained by: • Addition of a reducing substance
to the medium • Displacement of the air by carbon dioxide • Absorption of the
oxygen by chemicals • Removal of oxygen by direct oxidation of readily
oxidizable substances such as burning a candle, heating of copper,
phosphorus or other readily oxidizable metals • Incubation in the presence of
germinating grain or pieces of potato • Inoculation into the deeper layers of
solid media, or under a layer of oil in liquid media or • A combination of the
above methods. Moisture Proper moisture conditions must prevail in the
culture media for the growth of microorganisms. A moist medium and
atmosphere are necessary for the continued luxuriant growth of cells. For
example, if a medium in a plate is inoculated with an organism and wet cotton
is placed in the plate and sealed, the organism will show profuse growth. The
same organism might fail to show growth if the medium plate is not sealed
and is too dry. pH The pH of the culture medium, expressed as hydrogen ion
concentration [H+], is extremely important for growth. The majority of
microorganisms prefer culture media that are approximately neutral, while
others may require a medium that is distinctly acidic. Temperature Every
organism shows a rather general curve of growth as affected by temperature.
Such a curve shows 1) a minimum temperature below which growth stops, 2)
an optimum temperature at which growth is luxuriant and 3) a maximum
temperature above which the organism dies. Microorganisms are divided into
three main groups (mesophilic, psychrophilic and thermophilic) as far as
optimum temperature requirements are concerned. The usual range of
temperature suitable for the growth of mesophilic microorganisms lies
between 15-43 °C. Psychrophilic microorganisms have, however, been known
to grow and multiply at 0 °C. Thermophilic organisms may grow at
temperatures even greater than 80 °C. In general, the Microbiology BIOL 275
Dr. Eby Bassiri ebassiri@sas.upenn.edu 3 pathogenic organisms have a
temperature requirement of around 37 °C (body temperature) while
saprophytes have a much broader latitude. Medium Support The consistency
of a liquid medium may be modified by the addition of agar, gelatin or
albumin in order to change it into a solid or semisolid state. In the early 19th
century, infusions of plant and animal tissues, solutions of organic
compounds, and gelatin (as a solidifying agent) were employed as media for
the growth of microorganisms. However, gelatin had two main disadvantages;
being liquid at 37 °C (incubation temperature), and being liquefied or digested
by many bacteria. Bacteriology as a science began with the development of
methods for the cultivation of bacteria, and the introduction of agar by Hesse
in the 1890's was a step of greatest importance. Agar is actually credited to
Fanny Hesse, wife, technician and assistant of the German physician Walter
Hesse. Agar-agar, long used as an agent in preparing fruit jellies was
suggested by Mrs. Hesse as a replacement and became the standard
solidifying agent in microbiology. The properties of agar which make it ideal in
bacteriology are 1) solid agar melts (dissolves) at 100 °C, 2) remains solid at all
incubation temperatures, 3) is transparent, 4) is not heat-labile and therefore
easily sterilized, and 5) is unaffected by almost all bacteria. Liquid agar
solidifies at 42-44 °C which is useful because sterile, heat-labile components
such as antibiotics, blood, serum, carbohydrates and even bacterial cultures
may be added before allowing the medium to solidify. Solid media generally
contain agar at a concentration of 1.5%. Semi-solid media contain 0.05-0.3%
agar and are useful in culturing anaerobic and microaerophilic organisms
because such media form an oxygen gradient in test tubes, allowing all
degrees of oxygen tension to exist in the culture vessels. Sterile Conditions &
Autoclaving The media upon which microorganisms are grown must be sterile
or free from all other forms of microbes. The usual method for sterilization of
culture media is by means of the autoclave in which steam under pressure is
the sterilizing agent. Autoclave sterilization for 15 minutes at 15 pounds of
pressure and at 121 °C is recommended for quantities of liquid media up to
one liter (1 L). These settings are called the standard autoclaving conditions. If
larger volumes are to be sterilized in one container, and if the medium is not
hot when placed in the autoclave, a longer period should be employed. The
medium is prepared according to formula, distributed in tubes or flasks which
are then plugged with nonabsorbent cotton or loosely capped before being
placed in the autoclave. Plugs should fit neither too loosely nor too tightly.
Screw cap tops or metal covers may also be used to close the tubes or flasks.
Tubes should be placed in racks or packed loosely in baskets. Flasks should
never be more than two-thirds full. After the sterilization period has been
completed, the source of steam is cut off and the autoclave is allowed to
return to atmospheric pressure. Pressure should not drop too rapidly or the
media will boil over, blowing the plugs from the tubes or flasks. Pressure
should, however, Microbiology BIOL 275 Dr. Eby Bassiri
ebassiri@sas.upenn.edu 4 drop rapidly enough to prevent excessive exposure
of the media to heat after the sterilization period. The usual procedure for
using the autoclave is as follows: 1. Open door, and place items to be sterilized
into the autoclave chamber. Be sure that anything containing fluid is plugged
with styrofoam, cheesecloth, cotton, a Morton cap or else screw caps are
slightly loose. 2. Close door. Push down door lock lever until door studs are
completely in place. 3. Turn hatch wheel clockwise until it is secured tightly. 4.
The temperature of the autoclave is set at 121°C. If not, set the temperature by
sliding the upper (yellow) arrow to the desired temperature. Do not touch the
bottom indicator arrow. If you adjust to any temperature other than 121 °C,
return it to 121 °C at the end of the run. 5. Set timer by turning the large knob
just below the hands to the desired setting. DO NOT TOUCH the hands, they
break very easily! 6. Set cycle selection knob to desired setting. Remember, all
liquids MUST be done at SLOW EXHAUST. Dry materials can be done at Fast
Exhaust or Fast Exhaust and Dry. 7. Crank operating handle around to the
Sterilize position till the red steam light goes on. 8. If you are the first person
to use the autoclave that day, it is a good idea to wait and be sure the
chamber reaches the proper temperature and pressure. 9. The Slow Exhaust
and Fast Exhaust & Dry cycles both take 12-15 minutes longer than the time
set to finish. 10. At the end of the run the white STERILE light will go on, and a
loud, obnoxious buzzer will come on: a. Turn the cycle knob to MANUAL. b.
Rotate the operating handle all the way to OFF. Check that the chamber
pressure is zero, and the temperature is below 100 °C. d. Turn hatch wheel
counterclockwise, push up door lock lever and slowly open door. Watch out
for steam! e. Use heatproof gloves to remove materials. 11. Allow liquid
materials to cool before tightening caps. Microbiology BIOL 275 Dr. Eby Bassiri
ebassiri@sas.upenn.edu 5 A maximum of 15 minutes is recommended for the
sterilization of carbohydrates media in tubes to be used for fermentation
studies. Oversterilization or prolonged heating will change the composition of
the medium. For example, oversterilization results in the breakdown of lactose
in lactose-containing media. Agar media on prolonged sterilization, heating or
repeated melting are apt to show a precipitate. Media containing agar may
also form a flocculent precipitate if the liquid medium is held in the water bath
at 43-45 °C for longer than 30 minutes. Reheating the medium, however, may
disperse this flocculent agar precipitate. Excessive heating of media may also
result in an increase in acidity. The reaction of the media will become more
acidic as heating is prolonged. Culture media that may be harmed by
autoclaving are sometimes sterilized by the discontinuous or intermittent
method. This procedure consists of heating the medium in a chamber of
flowing steam for a period of 20 or 30 minutes on several successive days.
Liquid media may be sterilized by filtration through membranes, molecular
filters or unglazed porcelain. Storage of Media Media should always be stored
in a cool moist atmosphere to prevent evaporation, preferably in screwcapped tubes or bottles. Prolonged storage of sterile media cannot, however,
be recommended unless stability is established. If tubes of media have been
kept for any length of time, they should be reheated just before use. Liquid
media should be heated in a boiling water bath or in flowing steam for a few
minutes, to drive off dissolved gases, and then cooled quickly in cold water
without agitation just prior to inoculation. Agar tubes should be melted and
allowed to solidify in order to secure a moist surface that is desired by most
microorganisms. These precautions for both liquid and solid media are
extremely important for the initiation of growth of highly parasitic organisms
such as those encountered in blood culture work. Types of Media Culture
media may be divided into two main categories, complex (undefined) and
synthetic (defined). In a defined medium, all components are known to the
investigator such as a synthetic medium containing glucose as the sole carbon
source, inorganic salts as sources of sodium, phosphate and many other
required minerals such as Fe++ or Mg++, and an ammonium salt as a source
of nitrogen. Some bacteria are able to grow on media like that described
above, while others require growth factors that they cannot synthesize for
themselves (i.e., they are fastidious). A complex medium contains animal or
plant tissue extracts such as beef extract or yeast autolysate. These extracts
provide a large variety of nutrients in the form of lipids, hydrolyzed proteins (a
source of nitrogen as amino acids), carbon sources and vitamins and other
cofactors. The exact components of these extracts are unknown; therefore any
medium containing them is called undefined. Other sources of these necessary
growth factors are brain or heart tissue infusions, whole blood, serum, etc.
Microbiology BIOL 275 Dr. Eby Bassiri ebassiri@sas.upenn.edu 6 Throughout
the semester you will be using several types of general complex growth
media, both in broth and solid agar form including: Brain-Heart Infusion (BHI),
Nutrient Agar (NA), Trypticase Soy Agar (TSA), Luria or Luria-Bertani agar (L or
LB) and Sheep's Blood Agar (SBA) which is actually TSA + 5% sheep's blood. In
today's experiment, you will be making Nutrient Broth (NB) and Nutrient Agar
(NA), the most common standard complex media for culturing many
microorganisms. III. LABORATORY SUPPLIES Flask, 1 L 1/table Graduated
cylinder, 1 L 1/table Glass stirring rod 1/table Spatula 3/lab Weigh boats as
needed Beef extract, bottle 1/table Bacto peptone, bottle 1/table Agar,
powder 1/table pH paper, 6.5-10, box 1/table HCl, 1N 25 ml/dropper bottle
1/table NaOH, 1N 25 ml/dropper bottle 1/table Test tube rack 2/table Test
tubes, 18 x 150 mm 20/table Morton closure, 16 mm clear 10/table Morton
closure, 16 mm metal 10/table Slant racks 2/ table Labeling tape, roll 1/room
Heatproof gloves 1/room Autoclave RESERVE Water baths, 48 °C 1/room Petri
plates 10/table Cheesecloth square 1/table Cotton or Styrofoam plug 1/table
IV. PROCEDURE Note
1: The students at each table will work together as a group. Note 2: Check the
BlackBoard site for a flow chart of this lab procedure. 1. Wipe down lab bench
carefully with Disinfectant to help prevent contamination of your media.
2. Measure approximately 250 ml of distilled water (located in 60°C water
bath) in a 1 L graduated cylinder and pour into a 1 L flask. Microbiology BIOL
275 Dr. Eby Bassiri ebassiri@sas.upenn.edu 7
3. Weigh out 1.5 g beef extract and 2.5 g peptone and add into the flask.
Wash your spatula between bottles and wipe dry. DO NOT return excess
material that is weighed out to the container - discard. Use approximately 100
ml of the water to rinse any powder stuck to the side of the flask down into
the mixture.
4. Stir over gentle heat from a bunsen burner to dissolve completely.
5. Pour the mixture into the 1 L graduated cylinder and add warm water to the
500 ml mark. Pour back into the flask.
6. Check the pH of the medium and adjust to pH 7.0, if necessary, using the
HCl and/or NaOH. Adding the agar in the next step will not appreciably
change the pH.
7. Using a 10 ml pipette, dispense 10 ml of the mixture into each test tube.
Make 10 tubes and place in a test tube rack.
8. Add 6.0 g of agar to the flask and label it NA. Heat to just boiling for 1-2
minutes while stirring constantly. The agar will not dissolve unless it is boiled;
the solution will become completely clear when it has dissolved. Allow agar to
cool until there is no danger of you being burned and then dispense into the
tubes using a 10 ml pipette. Make ten 10 ml tubes.
9. Close the flask with a Styrofoam plug covered with cheesecloth and tape it
on top of the flask. Use another piece of tape to go around the neck of the
flask and pass over the first tape. Cap all the tubes with Morton closures.
These should be pushed down completely or else they will be forced off
during the autoclave's exhaust cycle. They are still selfventing when pushed
down all the way.
10. Keep one tube of each type in a drawer until next period to demonstrate
the need for sterilization. Continue to observe growth for one more period.
11. Autoclave the flask and the tubes for 15 minutes at 121 °C and 15 lb/in2
pressure at the slow exhaust mode. Watch your instructor for the use of the
autoclave.
12. After removing the media from the autoclave, allow the broth tubes to
cool, and store for later use. Place the flask in the 48°C water bath. Quickly lay
the tubes of NA on the slant racks on the center table so that the medium
forms a long slant and a short butt, and allow them to cool and solidify. Do
not allow the agar to reach the top of the tube. Allow them to cool completely
before returning to the rack. Store for later use. Label rack.
13. Lay your petri dishes on the bench. The cover should be on top. Light your
bunsen burner, then remove the NA flask from the water bath. Carefully wipe
the bottom dry to prevent the dripping water from contaminating the plates.
14. Remove the tapes and cotton plug from the flask. Carefully flame the neck
of the flask, open the plate cover about half way and fill the plate about 1/2
full. The plates have a full Microbiology BIOL 275 Dr. Eby Bassiri
ebassiri@sas.upenn.edu 8 line on the side; fill to that or slightly above. Put in a
little too much rather than too little. If there is not enough medium in the
plate, it will dry up in the incubator.
15. Flame the neck of the flask between each plate. Each student must pour at
least two plates. IMMEDIATELY rinse the excess agar out of the flask with hot
tap water and place on the discard cart. Allow plates to solidify completely,
which will take 15 minutes. Then invert, label and incubate at 37 °C overnight
to dry off excess moisture and check for contamination.
16. Clean all glassware and leave on paper towels beside sink.
Southeast Asian Fisheries Development Center Aquaculture Department
SEAFDEC/AQD Institutional Repository http://repository.seafdec.org.ph
SEAFDEC/AQD-Government of Japan-Trust Fund (GOJ-TF) Laboratory Manuals
2004 Disk diffusion method Tendencia, Eleonor A. Aquaculture Department,
Southeast Asian Fisheries Development Center Tendencia, E. A. (2004). Disk
diffusion method. In Laboratory manual of standardized methods for
antimicrobial sensitivity tests for bacteria isolated from aquatic animals and
environment (pp. 13-29). Tigbauan, Iloilo, Philippines: Aquaculture
Department, Southeast Asian Fisheries Development Center.
http://hdl.handle.net/10862/1635 Downloaded from
http://repository.seafdec.org.ph, SEAFDEC/AQD's Institutional Repository 13
Laboratory Manual of Standardized Methods for Antimicrobial Sensitivity Tests
for Bacteria Isolated from Aquatic Animals and Environment Eleonor A.
Tendencia Aquaculture Department Southeast Asian Fisheries Development
Center Philippines CHAPTER 2 Disk Diffusion Method 14 CHAPTER 2. Disk
Diffusion Method Laboratory Manual of Standardized Methods for
Antimicrobial Sensitivity Tests for Bacteria Isolated from Aquatic Animals and
Environment 15 Laboratory Manual of Standardized Methods for Antimicrobial
Sensitivity Tests for Bacteria Isolated from Aquatic Animals and Environment
PRINCIPLE This method is based on the principle that antibiotic-impregnated
disk, placed on agar previously inoculated with the test bacterium, pick-up
moisture and the antibiotic diffuse radially outward through the agar medium
producing an antibiotic concentration gradient. The concentration of the
antibiotic at the edge of the disk is high and gradually diminishes as the
distance from the disk increases to a point where it is no longer inhibitory for
the organism, which then grows freely. A clear zone or ring is formed around
an antibiotic disk after incubation if the agent inhibits bacterial growth. MEDIA
The disk diffusion method is performed using Mueller-Hinton Agar (MHA),
which is the best medium for routine susceptibility tests because it has good
reproducibility, low in sulfonamide, trimethoprim, and tetracycline inhibitors,
and gives satisfactory growth of most bacterial pathogens. The inoculum for
the disk diffusion method is prepared using a suitable broth such as tryptic
soy broth. This medium is prepared according to manufacturer’s instructions,
dispensed in tubes at 4-5 ml and sterilized. Sterile 0.9% salt solution may also
be used. Media are supplemented with 1-2% sodium chloride (NaCl) if
intended for marine organisms. Preparation of agar medium 1 Prepare MHA
from the dehydrated medium according to the manufacturer’s instructions.
Media should be prepared using distilled water or deionized water. 2 Heat
with frequent agitation and boil to dissolve the medium completely. Sterilize
by autoclaving at 121°C for 15 min. 14 CHAPTER 2. Disk Diffusion Method
Laboratory Manual of Standardized Methods for Antimicrobial Sensitivity Tests
for Bacteria Isolated from Aquatic Animals and Environment 15 Laboratory
Manual of Standardized Methods for Antimicrobial Sensitivity Tests for
Bacteria Isolated from Aquatic Animals and Environment 3 Check the pH of
each preparation after it is sterilized, which should be between 7.2 and 7.4 at
room temperature. This is done by macerating a small amount of medium in a
little distilled water or by allowing a little amount of medium to gel around a
pH meter electrode. 4 Cool the agar medium to 40-50°C. Pour the agar into
sterile glass or plastic petri dish on a flat surface to a uniform depth of 4 mm.
5 Allow to solidify. 16 CHAPTER 2. Disk Diffusion Method Laboratory Manual
of Standardized Methods for Antimicrobial Sensitivity Tests for Bacteria
Isolated from Aquatic Animals and Environment 17 Laboratory Manual of
Standardized Methods for Antimicrobial Sensitivity Tests for Bacteria Isolated
from Aquatic Animals and Environment 6 Prior to use, dry plates at 30-37°C in
an incubator, with lids partly ajar, for not more than 30 minutes or until excess
surface moisture has evaporated. Media must be moist but free of water
droplets on the surface. Presence of water droplets may result to swarming
bacterial growth, which could give inaccurate results. They are also easily
contaminated. 1 If plates are not to be immediately used, they may be stored
in the refrigerator inside airtight plastic bags at 2-8°C for up to 4 weeks.
Storage 2 Unpoured media may be stored in airtight screw-capped bottles
under the conditions specified by the manufacturer. 16 CHAPTER 2. Disk
Diffusion Method Laboratory Manual of Standardized Methods for
Antimicrobial Sensitivity Tests for Bacteria Isolated from Aquatic Animals and
Environment 17 Laboratory Manual of Standardized Methods for Antimicrobial
Sensitivity Tests for Bacteria Isolated from Aquatic Animals and Environment
Control Before use, check the ability of the agar to support the growth of
control strains (listed in the Introduction) by streaking bacterial cultures on the
agar medium. It is also advisable to check the ability of each batch of media to
support the growth of a representative member of the species to be tested.
INOCULUM Preparation 2 Transfer colonies to 5 ml of Trypticase soy broth or
0.9% saline. 1 From a pure bacterial culture (not more than 48 hours, old
except for slow growing organisms), take four or five colonies with a wire loop.
18 CHAPTER 2. Disk Diffusion Method Laboratory Manual of Standardized
Methods for Antimicrobial Sensitivity Tests for Bacteria Isolated from Aquatic
Animals and Environment 19 Laboratory Manual of Standardized Methods for
Antimicrobial Sensitivity Tests for Bacteria Isolated from Aquatic Animals and
Environment 3 Incubate the broth at 30°C or at an optimum growth
temperature until it achieves or exceeds the turbidity of 0.5 MacFarland
standard (prepared by adding 0.5 ml of 0.048 M BaCl2 to 99.5 ml of 0.36 NH2
SO4 ; commercially available). 4 Compare the turbidity of the test bacterial
suspension with that of 0.5 MacFarland (vigorously shaken before use) against
a white background with contrasting black line under adequate light. Arrow
points to tube with correct turbidity. 5 Reduce turbidity by adding sterile
saline or broth. NOTE: Standardized inoculum has a concentration of 1-2 ×
108 cfu/ml. 18 CHAPTER 2. Disk Diffusion Method Laboratory Manual of
Standardized Methods for Antimicrobial Sensitivity Tests for Bacteria Isolated
from Aquatic Animals and Environment 19 Laboratory Manual of Standardized
Methods for Antimicrobial Sensitivity Tests for Bacteria Isolated from Aquatic
Animals and Environment Inoculation of plates 1 Dip a sterile cotton swab into
the standardized bacterial suspension. 2 Remove excess inoculum by lightly
pressing the swab against the tube wall at a level above that of the liquid. 3
Inoculate the agar by streaking with the swab containing the inoculum. 20
CHAPTER 2. Disk Diffusion Method Laboratory Manual of Standardized
Methods for Antimicrobial Sensitivity Tests for Bacteria Isolated from Aquatic
Animals and Environment 21 Laboratory Manual of Standardized Methods for
Antimicrobial Sensitivity Tests for Bacteria Isolated from Aquatic Animals and
Environment 4 Rotate the plate by 60° and repeat the rubbing procedure.
Repeat two times. This will ensure an even distribution of the inoculum. 5
Allow the surface of the medium to dry for 3-5 minutes but not longer than 15
minutes to allow for absorption of excess moisture. ANTIMICROBIAL DISKS
Selection The number of antimicrobial agents to be tested should be limited.
To make the test practical and relevant, include only one representative of
each group of related drugs; those indicated for veterinary use to control or
prevent disease, and those that can be useful for epidemiological or research
purposes. Use antibiotic disks purchased from a reputable manufacturer. The
disk diameter is approximately 6 mm. Disks should be properly stored in a
tightly sealed container with desiccant at 2-8°C. Expired disks should not be
used. Application 1 Using sterile forceps or disk dispenser, place antibiotic disk
on the surface of the inoculated and dried plate. 20 CHAPTER 2. Disk Diffusion
Method Laboratory Manual of Standardized Methods for Antimicrobial
Sensitivity Tests for Bacteria Isolated from Aquatic Animals and Environment
21 Laboratory Manual of Standardized Methods for Antimicrobial Sensitivity
Tests for Bacteria Isolated from Aquatic Animals and Environment 2
Immediately press it down lightly with the instrument to ensure complete
contact between the disk and the agar surface. Do not move a disk once it has
come into contact with the agar surface since some diffusion of the drug
occurs instantaneously. 3 Position disks such that the minimum center - center
distance is 24 mm and no closer than 10 to 15 mm from the edge of the petri
dish. A maximum of six disks may be placed in a 9-cm petri dish and 12 disks
on a 150 mm plate. Reduce the number of disks applied per plate if
overlapping zones of inhibition are encountered. CONTROL PLATE Include one
plate inoculated with a control strain (Appendix 2.1) for every set of plates and
incubate together. INCUBATION 1 Incubate plates in an inverted position at
30°C or at an optimum growth temperature. 22 CHAPTER 2. Disk Diffusion
Method Laboratory Manual of Standardized Methods for Antimicrobial
Sensitivity Tests for Bacteria Isolated from Aquatic Animals and Environment
23 Laboratory Manual of Standardized Methods for Antimicrobial Sensitivity
Tests for Bacteria Isolated from Aquatic Animals and Environment 2 Observe
for the zone of inhibition after 16 to 18 hours. Slow growing organisms may
require longer incubation period. READING AND MEASUREMENT OF ZONES
OF INHIBITION Description 1 The zone of inhibition (arrow) is the point at
which no growth is visible to the unaided eye. 2 Record the presence of
individual colonies (arrow) within zones of inhibition. 22 CHAPTER 2. Disk
Diffusion Method Laboratory Manual of Standardized Methods for
Antimicrobial Sensitivity Tests for Bacteria Isolated from Aquatic Animals and
Environment 23 Laboratory Manual of Standardized Methods for Antimicrobial
Sensitivity Tests for Bacteria Isolated from Aquatic Animals and Environment 3
Record occurrence of fuzzy zones (arrow). In measuring the zone diameter, the
fuzzy portion of the zone should be ignored as much as possible. The zone
limit is the inner limit of the zone of normal growth. Reading 1 Read and
record the diameter of the zones of inhibition using a ruler graduated to 0.5
mm. 2 Round up the zone measurement to the nearest millimeter. 24
CHAPTER 2. Disk Diffusion Method Laboratory Manual of Standardized
Methods for Antimicrobial Sensitivity Tests for Bacteria Isolated from Aquatic
Animals and Environment 25 Laboratory Manual of Standardized Methods for
Antimicrobial Sensitivity Tests for Bacteria Isolated from Aquatic Animals and
Environment INTERPRETATION OF RESULTS 1 Compare the diameter of the
zone of inhibition of the test isolates with those in the chart of interpretative
standard for veterinary pathogens (Appendix 2.2). 2 Report result as Resistant
(R), Intermediate (I) or Susceptible (S). Example Disk used: Chloramphenicol, 30
µg (C-30) Zone of inhibition: 16 mm Result/ interpretation: Intermediate à
based on the zone diameter interpretative chart (Appendix 2.2) 3 Susceptibility
test results using agents other than those listed in the chart are interpreted on
the basis of the presence or absence of a definite zone of inhibition and is
considered only as qualitative until such time as interpretative zones have
been established. REJECTION CRITERIA 1 Do not read plates on which growth
of test bacteria have isolated colonies or less than semi-confluent growth
(arrow). 24 CHAPTER 2. Disk Diffusion Method Laboratory Manual of
Standardized Methods for Antimicrobial Sensitivity Tests for Bacteria Isolated
from Aquatic Animals and Environment 25 Laboratory Manual of Standardized
Methods for Antimicrobial Sensitivity Tests for Bacteria Isolated from Aquatic
Animals and Environment 2 Do not read zones of inhibition of two adjacent
disks that overlap (arrow) to the extent that measurement of the zone
diameter cannot be made. 3 Do not read zones showing distortion from
circular (arrow). 4 Reject all data collected in a particular set if the zones of
inhibition produced on plate inoculated with a control strain are not within the
tolerance limits set. 26 CHAPTER 2. Disk Diffusion Method Laboratory Manual
of Standardized Methods for Antimicrobial Sensitivity Tests for Bacteria
Isolated from Aquatic Animals and Environment 27 Laboratory Manual of
Standardized Methods for Antimicrobial Sensitivity Tests for Bacteria Isolated
from Aquatic Animals and Environment REFERENCES Alderman, D.J. and P.
Smith. 2001. Development of draft protocols of standard reference methods
for antimicrobial agent susceptibility testing of bacteria associated with fish
diseases. Aquaculture, 196: 211- 243. Anonymous. 1986. Antimicrobial
Susceptibility Testing: A System for Standardisation. Becton Dickinson and
Company, Hong Kong, 13 pp. Bailey, W.R. and E.G. Scott. 1966. Diagnostic
Microbiology, Second Edition. Toppan Company Ltd., Japan, pp. 257- 270.
Finegold, S.M. and W.J. Martin. 1982. Bailey and Scott’s Diagnostic
Microbiology, Sixth Edition. The CV Mosby Company, London, pp. 532- 557.
NCCLS. 2002. Performance Standards for Antimicrobial Disk and Dilution
Susceptibility Tests for Bacteria Isolated from Animals; Approved
StandardSecond Edition. NCCLS document M31-A2 (ISBN 1-56238-461-9).
NCCLS, 940 West Valley Road, Suite 1400, Wayne, Pennsylvania 19087-1898,
USA Prescott, L.M., J.P. Harley and D.A. Klein. 1993. Microbiology, Second
Edition. Wm C Brown Publishers, England, pp. 325-343. 26 CHAPTER 2. Disk
Diffusion Method Laboratory Manual of Standardized Methods for
Antimicrobial Sensitivity Tests for Bacteria Isolated from Aquatic Animals and
Environment 27 Laboratory Manual of Standardized Methods for Antimicrobial
Sensitivity Tests for Bacteria Isolated from Aquatic Animals and Environment
APPENDIX.