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Small World Initiative Protocols
SMALL WORLD INITIATIVE PILOT TRAINING WORKSHOP
PROTOCOLS
TABLE OF CONTENTS
BLAST ANALYSIS OF SEQUENCE .......................................................................................................... 104
FREEZING DOWN GLYCEROL STOCKS ..................................................................................................... 105
SERIAL DILUTIONS ............................................................................................................................ 106
SCREEN FOR ISOLATE ANTIBIOTIC PRODUCTION #1 – PATCH/PATCH ............................................................ 110
SCREEN FOR ISOLATE ANTIBIOTIC PRODUCTION #2 – SPREAD/PATCH .......................................................... 112
SCREEN FOR ISOLATE ANTIBIOTIC PRODUCTION #3 – SOFT AGAR ................................................................ 113
GRAM STAIN ................................................................................................................................... 114
PLATING SOIL SAMPLE ....................................................................................................................... 118
CATALASE REACTION......................................................................................................................... 119
COLONY MORPHOLOGY PROTOCOL ..................................................................................................... 122
COLONY PCR .................................................................................................................................. 125
MACCONKEY AGAR TEST ................................................................................................................... 127
PICKING AND PATCHING COLONIES ....................................................................................................... 129
CHEMOTAXIS ................................................................................................................................... 130
GEL ELECTROPHORESIS ...................................................................................................................... 132
SPREAD PLATE ................................................................................................................................. 133
STREAK PLATE ................................................................................................................................. 135
ANALYZING ORGANIC EXTRACTS FOR ANTIBIOTIC PRODUCTION ................................................................... 139
ANTIBIOTIC RESISTANCE TESTS ............................................................................................................ 141
APPENDIX A: HISTORY OF GROWTH MEDIA, AGAR, AND PETRI DISHES ......................................................... 142
APPENDIX B: PREPARING AND POURING PLATES..................................................................................... 142
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BLAST Analysis of Sequence
Genomes contain large amounts of information – the human genome is over 3 billion base pairs long and bacterial
genomes like Escherichia coli contain nearly 5 million base pairs. Making sense of all this information not only
requires biological knowledge of how genes work and what they encode, but also great computational tools to sort
through information and solve problems. Sequenced genomes are submitted into large databases like GenBank
that are impossible to navigate without search tools, just as Google Search helps us find specific information in the
whole of the World Wide Web.
The Basic Local Alignment Search Tool or BLAST (Altschul, Gish, Miller, Myers, & Lipman, 1990) is a bioinformatics
tool that allows us to navigate through huge databases and compare an amino acid or nucleotide sequence to a
library of published or submitted sequences. Using BLAST to compare DNA sequences allows us to find closely
related genes or regions of DNA in the database. These closely related genes give us information about the
function of the protein product or the identity of the organism they belongs to.
MATERIALS


16S ribosomal RNA gene sequence and chromatogram
BLAST website
PROTOCOL
1.
2.
3.
4.
5.
6.
7.
Go to the BLAST website: http://www.ncbi.nlm.nih.gov/BLAST/
Choose BLAST program to run  “nucleotide blast”
Enter your sequence* into the “Enter query sequence” field and enter appropriate nucleotide range*
Under “Choose search set”, select appropriate database to make search: click “Others” and select
“Nucleotide collection (nr/nt)”
Once you have submitted your sequence and set the parameters, click “BLAST” at the bottom of the page
BLAST will take a couple of seconds to run your sequence against other sequences in the database
Once the search is done, analyze you BLAST data: the “Descriptions” column lists identifiers for similar
sequences in the database. These identifiers are ranked by “max ident”, which is the percentage of
matching nucleotides. Normally, a “max ident” of 97% or higher means that your sequences matches the
specific description, which corresponds to a strain or species. The expectation value (“E value”) tells you
how statistically significant your match is, hence, the lower the “e value”, the more reliable the match.
*Before entering your sequence into the “Enter query field”, be sure to assess the quality of your sequence. This
can be done by looking at your sequence’s trace chromatogram, which shows the fluorescence peaks given off by
each of the four nucleotides during Sanger sequencing. Peaks are considered low quality when they are hard to
distinguish; this usually happens at the ends of the sequence. Based on this information, pick a “clean” nucleotide
range in your sequence to BLAST. After pasting your full sequence into the appropriate box, enter the appropriate
region into the “Query subrange” fields.
REFERENCE
Altschul, S. F., Gish, W., Miller, W., Myers, E. W., & Lipman, D. J. (1990). Basic local alignment search tool. J Mol
Biol, 215(3), 403-410. doi: 10.1016/S0022-2836(05)80360-2
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Freezing down glycerol stocks
Adapted from:
http://openwetware.org/wiki/
Once you have bacterial cells of interest, it is especially important to preserve the cells for years for future use. For
long-term storage, a glycerol stock of those cells should be made as soon as possible to avoid accidental loss of the
strain. E. coli can survive at -80oC for years if the cells are prepared properly. What kills cells upon freezing is the
formation of ice crystals rupturing the plasma membrane. Therefore, we add glycerol to prevent ice from forming.
Generally, glycerol stocks are 20% glycerol final at the final concentration.
MATERIALS
▪
▪
▪
80% glycerol solution
Day/overnight culture or fresh streak-out plates
Cryogenic vials/1.5mL microfuge tube
PROTOCOLS
1.
2.
3.
4.
5.
6.
Pick a single colony of the clone off a plate and grow an overnight in the appropriate selectable liquid
medium (3-5ml).
Add 0.25 ml of 80% glycerol in H2O to a cryogenic vial.
Add 0.75 ml sample from the culture of bacteria to be stored. OR 0.75 ml media with loopful of a fresh
streak-out of the isolate of interest
Gently vortex the cryogenic vial to ensure the culture and glycerol is well-mixed.
▪ Alternatively, pipet to mix.
On the side of the vial list all relevant information - strain, date, researcher, etc.
Freeze glycerol stock in liquid nitrogen and store in a -80C freezer.
▪ This will also be a good time to record the strain information and record the location in a external
database
Notes
▪
While it is possible to make a long term stock from cells in stationary phase, ideally your culture should be
in logarithmic growth phase.
Certain antibiotics in the medium should be removed first as they are supposedly toxic over time, ex)Tetracycline.
To do this, spin the culture down and resuspend in same volume of straight LB medium.
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Serial Dilutions
Adapted from:
Jackie Reynolds – Microbe Library
Determining microbial counts for liquid and solid samples is a common practice in the lab; whether it is to quantify
the biomass of a soil sample, calculate an antibiotic’s minimal inhibitory concentration (MIC) or the population
density in a liquid culture. In most environmental samples, bacteria are highly numerous, ranging from the tens of
thousands to the millions in as little as 1 ml of seawater or 1 g of soil (citation needed). Given the magnitude of
these numbers and rate of cell turnover, it would be nearly impossible to get an exact count of all the cells in a
sample. Instead of counting cells one by one, microbiologists calculate cell density through colony forming units or
CFUs, which give us an approximation of the number of viable cells per milliliter or gram of a sample. To calculate
CFUs, a sample must be diluted in a saline solution that keeps the cells in suspension alive. Diluting 1 g of soil with
saline solution to a final volume of 10 ml would create a 10-fold or 1:10 dilution of our soil sample; therefore, if the
cells are properly suspended in the solution, all the cells contained in 1 g of sample will be evenly distributed in 10
ml of solution. Making serial dilutions of a sample in 10-fold increments allow us to further reduce the number of
cell per volume to a proportion that is easier to work with and easier to count. Once we have reached a desired
dilution of our sample, we can add the dilution to a solid medium that will support the growth of the bacteria.
Once the bacteria grow to colonies (1 bacterium giving rise to 1 colony of clones) we can determine how many
bacteria were plated and calculate the cell density in the original sample, measured in CFUs. For example, if we
serially dilute 1 g of soil sample by a factor of 103, spread and incubate the dilution on a solid medium, and then
observe 130 colonies, we would obtain 130 x 103 or 1.3 x 105 CFUs / g of soil. This number represents the number
of viable cells, i.e. cells in an environmental sample that can survive lab conditions and grow in culture. While the
number of bacteria we can successfully grow in the lab remains scant (citation needed), plating various dilutions on
different media formulations and under different conditions (e.g., lighting and temperature) can increase our
access to diverse bacteria.
Standard formula for culture density:
Culture density (cells/mL) =
colony count (CFUs) on an agar plate total dilution X volume plated (mL)
To work the problem, you need 3 values:
 a colony count from the pour or spread plates
 a dilution factor for the dilution tube from which the countable agar plate comes
 the volume of the dilution that was plated on the agar plate.
Value obtained is cells/ml or colony forming units (CFU)/mL. If want to convert CFU/g:
CFU/g = CFU/mL X volume of suspended soil sample (mL)
Calculate dilution factor:
Determine the dilution factor of each tube in the set.
dilution factor for a tube =
amount of sample
volume of specimen transferred + volume of diluent in tube
(total volume)
Total dilution factor = multiply the individual dilution factor for the tube and all previous tubes. 106
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Serial dilution and plating schematic. A series of 10-fold dilution is made from the original inoculum, which
contains cells in suspension from environmental sample. Each subsequent dilution and plate will have 10-fold
fewer bacteria than the previous one, making it easier for us to count colonies and calculate CFUs.
(Image acquired from <http://faculty.irsc.edu/FACULTY/TFischer/micro/serial%20dilution.jpg>)
MATERIALS




Choice of media plates
Conical tube
Environmental [soil] sample
Phosphate buffered saline (PBS)
PROTOCOL
1.
2.
3.
4.
5.
6.
7.
Obtain and label appropriate number of plates and 1.5ml microcentrifuge tubes, one for each subsequent
dilution. Dilutions should be made in increments of 10 (10-1, 10-2, 10-3, etc.)
Take soil sample and weigh out 1g
Transfer to 15ml conical tube.
Add 9ml PBS to 1g soil
Place tube in sonicator bath and sonicate for 30 sec.
Determine the dilution series and calculate appropriate volumes for each. Again, dilutions should be
made in increments of 10, thus add 900ul of diluent (PBS) into each dilution tube.
900ul diluent + 100ul specimen transferred = 1000ul
Plate 100ul of each dilution to appropriate plates.* Note volume and dilutions plated.
*Check with your instructor about spread plating technique.
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SAFETY
Tubes and agar plates should be discarded properly in a biohazard container for proper sterilization. The pipettes
will also be sterilized (washed first if using reusable glass pipettes).
Do not pipette by mouth.
Use sterile
technique in the transfer of microorganisms from tube to tube, as well as in the production of the pour
plates.
The ASM advocates that students must successfully demonstrate the ability to explain and practice safe
laboratory techniques. For more information, read the laboratory safety section of the ASM Curriculum
Recommendations: Introductory Course in Microbiology and the Guidelines for Biosafety in Teaching
Laboratories.
COMMENTS AND TIPS
Greater than 300 colonies on the agar plate and less than 30 leads to a high degree of error. Air contaminants can
contribute significantly to a really low count. A high count can be confounded by error in counting too many small
colonies, or difficulty in counting overlapping colonies. Use sterile pipettes for the dilutions, and use different
ones in between the different dilutions. To do otherwise will increase the chances of inaccuracy because of carryover of cells.
Accuracy in quantitation is determined by accurate pipette use and adequate agitation of dilution
tubes.
PRACTICE EXERCISES
1.
You are given a test tube containing 10 mL of a solution with 8.4 x 107 cells/mL. You are to produce a solution
that contains less than 100 cells/mL. What dilutions must you perform in order to arrive at the desired result?
ANSWER: You should perform a series of three 1:100 dilutions to yield 84 cells/mL.
1 mL of original solution to 99 mL of water = 8.4 x 10 5 cells/mL.
1 mL of second solution to 99 mL of water = 8.4 x 103 cells/mL.
1 mL of third solution to 99 mL of water = 8.4 x 101 or 84 cells/mL.
2.
You have a microtube containing 1 mL of a solution with 4.3 x 104 cells/mL and you are to produce a solution
that contains 43 cells/mL. What dilutions must you perform?
ANSWER: You could perform the following dilutions:
10 µL of original solution to 990 µL of water = 4.3 x 102 cells/mL.
100 µL of second solution to 900 µL of water = 4.3 x 10 1 or 43 cells/mL.
3.
You are given a container with 5 mL of a solution containing 5.1 x 103 cells/mL. You are to produce a solution
that contains approximately 100 cells/mL.
ANSWER: You would perform the following dilutions:
0.5 mL of original solution to 4.5 mL of water = 5.1 x 102 cells/mL
1 mL of second solution to 4 mL of water = 1.02 x 102 cells/mL or 102 cells/mL
4.
You are given a container of yeast cells for which Klett units have been determined on a Klett Summerson
Colorimeter. The container contains a population whose concentration is 2.6 x10 6 cells/mL. You are to
prepare a suspension which, when you spread 1 mL of the suspension on appropriate media, will result in
about 100 cells.
ANSWER:
10 µl of original solution to 990 µl (or 1.0 mL) of sterile water = 2.6 x 104 cells/mL
10 µl of second solution to 990 µl (or 1.0 mL) of sterile water = 2.6 x 102 cells/mL
0.5 mL of third solution to 0.5 mL of sterile water = 1.3 x 10 2 or 130 cells/mL
0.77 mL of fourth solution to 0.23 mL of sterile water = 100 cells/mL
Note: Corrected 9 March 2005 108
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Screen for isolate antibiotic production #1 – patch/patch
Alexander Fleming is credited for having discovered the first naturally produced and clinically successful antibiotic
that humans came across with, penicillin. What he did on a whim back in 1928 is what many microbiologists do
systematically today to find antibiotic producers. *He noticed that a mold (Penicillum notatum) had contaminated
one of his staph cultures and inhibited the growth of the cells bacterial cells around it, creating a pronounced zone
of inhibition. These microbial producers secrete their powerful chemical weapons into their surroundings, diffusing
into the broth or agar they are growing in. Susceptible microorganisms that come in contact with this chemical are
inhibited in their ability to survive or reproduce, which can ultimately result in death. Areas on a plate that would
normally be lush with colonies or a spread/lawn of bacteria become clear, creating visible zones of inhibition.
Microbiologists apply these basic principles when conducting activity assays, tests that determine the presence of
antimicrobial compounds in a culture. Many researchers are concerned with finding compounds that specifically
target bacteria, especially those that pose a great threat to our health. Therefore, human pathogens or related
bacteria (due to the risk to the research of working with the pathogens themselves) are used as test subjects in
activity assays to find compounds that are “active” against them.
THEORY
The patch-patch protocol assays for bioactivity in bacteria that are in close proximity to but not necessarily in
physical contact with a particular tester strain. This approach assumes that bacteria will synthesize and secrete
active compounds independently of their neighboring microorganisms. Many prolific antibiotic producers, such as
streptomycetes, rely on other biochemical and physiological cues to onset their secondary metabolism, which
involves the production of accessory compounds such as pigments and antibiotics. For example, in Streptomyces
coelicolor, antibiotic production occurs in a growth-phase dependent-manner or in response to nutrient limitation
(Bibb, 1996). Therefore, even if susceptible microorganisms are separated by a small gap from S. coelicolor, if the
secreted antibiotic diffuses towards the tester strain, we should expect to observe inhibition.
Patch-patch schematic. Isolates are patched around the edge of plate and tester strain is patched in the middle,
separated by small gap.
MATERIALS




2 media plates of choice
Master plates (made in previous experiment) with isolates
Plate of safe ESKAPE relative
Sterile toothpicks
PROTOCOL
1.
2.
3.
Obtain your master plate and the culture containing your safe ESKAPE relative of choice
Choose a fresh medium plate for safe ESKAPE relative to grow
Pick isolates from your master plate and patch them around the perimeter of the plate (see diagram
below)
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4.
5.
Label each patch with the appropriate name on the back of the plate.
Patch your tester strain (safe ESKAPE pathogen) in the center of the plate without touching the soil
isolates’ patches.
REFERENCES
Bibb, M. (1996). The regulation of antibiotic production in Streptomyces coelicolor A3(2). Microbiology-Uk, 142,
1335-1344.
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Screen for isolate antibiotic production #2 – spread/patch
THEORY
The spread-patch protocol assays for bioactivity in bacteria that are in close physical contact with the tester strain.
In this protocol, the testers train is spread on a plate and then the isolates are patched onto the spread. If the
isolate is active against the tester strain, it should theoretically have no trouble growing on the spread. Yet, this is
not always the case as an antibiotic producer may need time to establish itself on the medium or may not be
successful at invading a growing culture of susceptible bacteria (Wiener, 1996). While in this experiment we can
ignore establishment times, this protocol cannot ignore that some antibiotic producers may require physical or
biochemical contact with other microbes to onset antibiotic production. Therefore, this protocol assumes that
microbe-microbe interactions may induce antibiotic production and will increase our chances of identifying a
producer.
While this approach is simple, its implications are grand. We know that in their natural habitats, microbes are part
of intricate networks and constantly interact with other microbes. Extracellular signals coming from different
organisms or the same species (quorum sensing) can unleash a signaling cascade within a cell that ultimately
affects its regulation of genes. For many years, researchers have been attempting to find what specific microbemicrobe interactions or other biochemical or environmental cues would trigger the expression of antibiotic genes.
Spread-patch schematic. Isolates are patched (in the same grid arrangement as master plate) onto safe ESKAPE
relative spread.
MATERIALS






2 appropriate media plates (same as master plate)
Liquid culture of safe ESKAPE relative
Master plates with isolates
Spread beads
Square grid
Sterile toothpicks
PROTOCOL
1.
2.
3.
4.
5.
6.
7.
Obtain appropriate media plate for your safe ESKAPE relative to growth.
Label plate with respective medium, safe ESKAPE relative, and master plate used. Add vertical line as
point of orientation, aligned with master plate.
Obtain a liquid culture of your safe ESKAPE relative from TA.
Add 150 μL of the safe ESKAPE relative liquid culture onto the medium plate. This is the inoculation step.
Add 5-10 spread beads to the plate, being careful not to splash the inoculum, cover plate with lid and
shake side by side to spread the inoculum. Carefully shake the beads off into appropriate container. The
liquid should be absorbed into medium within minutes.
Place the new plate face-up on top of the grid and align line of orientation with grid.
Pick isolates from the master plate and patch onto safe ESKAPE relative spread arranged as master plate.
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Screen for isolate antibiotic production #3 – soft agar
THEORY
Another method of screening for antibiotic production that we will be using is called “soft agar.” In this technique,
liquefied agar is mixed with the tester strain and poured over patches of the soil isolates. Since the tester strain is
uniformly distributed on the overlying agar (which soon after solidifies), it could almost allow for the threedimensional visualization of zones of inhibition.
MATERIALS







2 of each appropriate media plates
Glass test tube
Liquid culture of safe ESKAPE relative
Master plates with isolates
Soft agar
Square grid
Sterile toothpicks
PROTOCOL
1.
2.
3.
4.
5.
Patch isolates of interest onto appropriate solid medium for safe ESKAPE relative to grow. Do not use
excessive inoculum and patch lightly.
In a test tube, add ___ mL of overnight liquid culture of your safe ESKAPE relative with ___ ml of soft agar.
Mix contents of test tube by swirling and tapping gently.
Gently pour soft agar with safe ESKAPE relative over the patched plate. Pour onto one side of the plate
and tip over to flood entire plate.
Cover plate and let sit until soft agar solidifies. Incubate plate under appropriate conditions.
REFERENCES
Wiener, Pamela. (1996). Experimental studies on the ecological role of antibiotic production in bacteria.
Evolutionary Ecology, 10(4), 405-421. doi: 10.1007/BF01237726
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Gram Stain
Adapted from:
Ann C. Smith and Marise A. Hussey – Microbe Library
HISTORY
The Gram stain was first used in 1884 by Hans Christian Gram (Gram, 1884). Gram was searching for a method
that would allow visualization of cocci in tissue sections of lungs of those who had died of pneumonia. Already
available was a staining method designed by Robert Koch for visualizing turbercle bacilli. Gram devised his method
that used Crystal Violet (Gentian Violet) as the primary stain, an iodine solution as a mordant (sets the dye)
followed by treatment with ethanol as a decolorizer. This staining procedure left the nuclei of eukaryotic cells in
tissue samples unstained while the cocci found in the lungs of those who had succumbed to pneumonia were
stained blue/violet. Gram found that his stain worked for visualizing a series of bacteria associated with disease
such as the “cocci of suppurative arthritis following scarlet fever”. He found however that Typhoid bacilli were
easily decolorized after the treatment with crystal violet and iodine, when ethanol was added. We now know that
those organisms that stained blue/violet with Gram’s stain are gram-positive bacteria and include Streptococcus
pneumoniae (found in the lungs of those with pneumonia) and Streptococcus pyogenes (from patients with Scarlet
fever) while those that were decolorized are gram-negative bacteria such as the Salmonella typhi that is associated
with Typhoid fever.
You may read the original publication of the staining procedure in the translated article "The Differential Staining
of Schizomycetes in tissue sections and in dried preparations".
PURPOSE
The Gram stain is fundamental to the phenotypic characterization of bacteria. The staining procedure
differentiates organisms of the domain Bacteria according to cell wall structure. Gram-positive cells have a thick
peptidoglycan layer and stain blue to purple. Gram-negative cells have a thin peptidoglycan layer and stain red to
pink.
THEORY
The Gram stain, the most widely used staining procedure in bacteriology, is a complex and differential staining
procedure. Through a series of staining and decolorization steps, organisms in the Domain Bacteria are
differentiated according to cell wall composition. Gram-positive bacteria have cell walls that contain thick layers of
peptidoglycan (90% of cell wall). These stain purple. Gram-negative bacteria have walls with thin layers of
peptidoglycan (10% of wall) and high lipid content. These stain pink. This staining procedure is not used for
Archeae or Eukaryotes as both lack peptidoglycan. The performance of the Gram Stain on any sample requires
four basic steps that include applying a primary stain (crystal violet) to a heat-fixed smear, followed by the addition
of a mordant (Gram’s Iodine), rapid decolorization with alcohol, acetone, or a mixture of alcohol and acetone and
lastly, counterstaining with safranin.
Details of the chemical mechanism of the Gram stain were determined in 1983 (Davies et al.,1983 and Beveridge
and Davies, 1983). In aqueous solutions crystal violet dissociates into CV+ and Cl – ions that penetrate through the
wall and membrane of both gram-positive and gram-negative cells. The CV+ interacts with negatively charged
components of bacterial cells, staining the cells purple. When added, iodine (I- or I3-) interacts with CV+ to form
large CV-I complexes within the cytoplasm and outer layers of the cell. The decolorizing agent, (ethanol or an
ethanol and acetone solution), interacts with the lipids of the membranes of both gram-positive and gramnegative Bacteria. The outer membrane of the gram-negative cell is lost from the cell, leaving the peptidoglycan
layer exposed. Gram-negative cells have thin layers of peptidoglycan, one to three layers deep with a slightly
different structure than the peptidoglycan of gram-positive cells (Dmitriev, 2004). With ethanol treatment, gramnegative cell walls become leaky and allow the large CV-I complexes to be washed from the cell. The highly crosslinked and multi-layered peptidoglycan of the gram-positive cell is dehydrated by the addition of ethanol. The
multi-layered nature of the peptidoglycan along with the dehydration from the ethanol treatment traps the large
CV-I complexes within the cell. After decolorization, the gram-positive cell remains purple in color, whereas the
gram-negative cell loses the purple color and is only revealed when the counterstain, the positively charged dye
safranin, is added. At the completion of the Gram stain the gram-positive cell is purple and the gram-negative cell
is pink to red.
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Some bacteria, after staining with the Gram Stain yield a pattern called gram-variable where a mix of pink and
purple cells are observed. The genera Actinomyces, Arthrobacter, Corynebacterium, Mycobacterium, and
Propionibacterium have cell walls particularly sensitive to breakage during cell division, resulting in gram-negative
staining of these gram-positive cells. In cultures of Bacillus, Butyrivibrio, and Clostridium a decrease in
peptidoglycan thickness during growth coincides with an in increasing number cells that stain gram-negative
(Beveridge, 1990). In addition, in all bacteria stained using the Gram stain, the age of the culture may influence
the results of the stain.
Some bacteria do not stain as expected with the Gram stain. For example, members of the genus Acinetobacter
are gram-negative cocci that are resistant to the decolorization step of the Gram stain. Acinetobacter spp. often
appear gram-positive after a well prepared Gram stain (Visca et al. 2001). For Mycobacterium spp., the waxy
nature of the coat renders the bacteria not readily stainable with dyes used in the Gram stain, though the bacteria
are considered to be gram positive (Saviola and Bishai, 2000). Gardnella has an unusual gram-positive cell wall
structure that causes bacteria of this genus to stain gram-negative or gram-variable (Sadhu et al 1989).
Misinterpretation of the Gram stain has led to misdiagnosis or delayed diagnosis of infectious disease (Visca et al.,
2001, Noviello et al., 2004 )
RECIPE (Gephardt et al., 1981)
This is Hucker’s modification of the Gram Stain method. Gram originally used Gentian Violet as the primary stain in
the Gram stain. Crystal violet is generally used today. In Hucker’s method ammonium oxalate is added to prevent
precipitation of the dye (McClelland, 2001) and uses an alcoholic solution of the counterstain. Burke’s
modification of the Gram Stain adds sodium bicarbonate to the crystal violet solution. Sodium bicarbonate
prevents the acidification of the solution as iodine oxidizes (McClelland, 2001) and uses an aqueous solution of
Safranin for the counterstain (Gephardt et al., 1981).
The reagents listed below can be made or purchased commercially from biological supply houses
1. Primary Stain: Crystal Violet Staining Reagent.
Solution A for crystal violet staining reagent
Crystal violet (certified 90% dye content) 2g
Ethanol, 95% (vol/vol)
20 ml
Solution B for crystal violet staining reagent
Ammonium oxalate
0.8 g
Distilled water
80 ml
Mix A and B to obtain crystal violet staining reagent. Store for 24 h and filter through paper prior to use.
2. Mordant: Gram's Iodine
Iodine
Potassium iodide
Distilled water
1.0 g
2.0 g
300 ml
Grind the iodine and potassium iodide in a mortar and add water slowly with continuous grinding until the iodine is
dissolved. Store in amber bottles.
3. Decolorizing Agent
Ethanol, 95% (vol/vol)
*Alternate Decolorizing Agent
Some professionals prefer an acetone decolorizer while others use a 1:1 acetone and ethanol mixture.
Commercially, a variety of mixtures are available, most using 25 – 50% acetone with the ethanol. A few include a
small quantity of isopropyl alcohol and/or methanol in the formulation.
Acetone
50 ml
Ethanol (95%)
50 ml
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4. Counterstain: Safranin
Stock solution:
Safranin O
95% Ethanol
Working Solution:
Stock Solution
Distilled water
2.5g
100 ml
10 ml
90 ml
PROTOCOL (Gephardt et al, 1981, Feedback from ASMCUE participants, ASMCUE , 2005)
1.
2.
3.
4.
Place 25ul water onto a slide and spread as much as possible.
Using a sterile stick, toothpick or loop, obtain a very small sample of a bacterial colony.
Gently mix the bacteria into the water on the slide to make a smear.
Let the bacterial smear air-dry. Then, using forceps, pass the dried slide through the flame of a Bunsen
burner 3 or 4 times, smear side facing up. Once the slide is heat fixed, it can then be stained.
5. Flood smear of cells for 1 minute with crystal violet staining reagent. Please note that the quality of the
smear (too heavy or too light cell concentration) will affect the Gram Stain results.
6. Wash slide in a gentle and indirect stream of tap water for 2 seconds.
7. Flood slide with the mordant: Gram's iodine. Wait 1 minute.
8. Wash slide in a gentle and indirect stream of tap water for 2 seconds.
9. Flood slide with decolorizing agent. Wait 15 seconds or add drop by drop to slide until decolorizing agent
running from the slide runs clear (see Comments and Tips section).
10. Flood slide with counterstain, safranin. Wait 30 seconds to 1 minute.
11. Wash slide in a gentile and indirect stream of tap water until no color appears in the effluent and then
blot dry with absorbent paper.
12. Observe the results of the staining procedure under oil immersion using a Brightfield microscope. At the
completion of the Gram Stain, gram-negative bacteria will stain pink/red and gram-positive bacteria will
stain blue/purple.
In a smear that has been stained using the Gram Stain
protocol, the shape, arrangement and gram reaction of a
bacterial culture will be revealed.FIG. 1. shows grampositive (blue/purple) rods and FIG. 2. shows gramnegative (pink/red) rods.
SAFETY
Dispose of glass slides in the Biohazard Sharps container
after visualization.
COMMENTS AND TIPS:
Comments and tips come from
discussions at ASM Conference for Undergraduate Educators
2005.
The thickness of the smear used in the Gram stain will affect
the result of the stain. The step that is most crucial in
effecting the outcome of the stain is the decolorizing step.
Over-decolorizing will lead to an erroneous result where
gram-positive cells may stain pink to red indicating a gramnegative result, and under-decolorizing will lead to an
erroneous result where gram-negative cells may appear blue
to purple indicating a gram-positive result. The degree of
decolorizing required is determined by the thickness of the
smear (number of cells on the slide). The Gram stain was
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FIG. 1
FIG. 2
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discussed in detail at the American Society for Microbiology Conference for Undergraduate Educators in 2005
when this protocol was reviewed. The group recommends that cells be prepared with a thin smear with no areas
of clumping or inconsistency. When staining the thin smear a short decolorizing time should be used. Some
individuals will flood the slide for 15 seconds or less with decolorzing agent, while others recommend adding
decolorizing agent drop wise for 5- 15 seconds or until the color of the decolorizing agent running from the slide no
longer shows any color.
It is recommended that young, actively growing cultures be used for gram staining. An intact cell wall is required
for an accurate gram stain. Older cultures may have breaks in the cell wall and often give gram-variable results
where a mixture of pink/red cells are seen among blue/purple cells.
Using a gram stain control is recommended. On the same slide as the test culture, include a sample of cells with a
known gram stain reaction to serve as a control for success in the gram stain technique.
Gram-stained bacteria should be viewed with a brightfield microscope at 1000X magnification with oil immersion.
If the smear of cells is crowded it will be difficult to note cell shape and arrangement.
When viewing slides use brightfield microscopy and adjust the brightness sufficiently to reveal the color of the
specimen.
Freshly made staining reagents are recommended. With older staining reagents, filter stains before use.
In the Gram Stain technique, two positively charged dyes are used: crystal violet and safranin. The use of the
designation “gram-positive” should not be confused with the concept of staining cells with a simple stain that has a
positive charge.
KOH string test may be used as a confirmatory test for the Gram Stain (Powers, 1995, Arthi et al., 2003): The
formation of a string (DNA) in 3% KOH indicates that the isolate is a gram-negative organism. Procedure:
Place a drop of 3% KOH onto a glass slide.
Emulsify in KOH a loopful of a 18-24 hour culture.
Continue
to mix the suspension for 60 sec and by slowly lifting the loop, observe for the formation of a string.
Interpretation: Gram-negative cells form a string within 60 seconds. Gram-positive cells are not affected.
Various formulations of decolorizing agents may be used (acetone, acetone/ethanol, ethanol). Acetone is the
most rapid decolorizer followed by acetone/ethanol and then ethanol. Ethanol is recommended for student use to
prevent over-decolorization of samples (McClelland, 2001)
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Plating soil sample
We will be using a culture-dependent approach to study bacteria, which relies on our ability to effectively grow
bacteria in the lab. Microbiologists have used this approach for over a century and while the premise has not
changed, our increased understanding of bacterial nutrition and growth has allowed us to develop media and lab
conditions to more closely meet their requirements for survival. This has also increased the reproducibility of
natural phenomena from lab to lab, allowing microbiologists to agree on observations and make more
advancement in the field. While we are far from attaining perfection in culturing bacteria and accurately
replicating their natural habitats, we have the tools of observation, inquiry and ingenuity to come a step closer. In
addition, the more we learn about environments like the soil, the more we realize that this is truly an inexhaustible
source of bacteria that we still have much to learn from.
After collecting a soil sample, the first step is to suspend the bacteria in liquid so that we can easily transfer them
to another medium (e.g. a nutrient plate), leaving behind debris and inorganic matter in the soil. It is important to
take consideration of the type of soil they were isolated from (may range from moist to dry, organic to sandy,
sunlit to shady) and using these observations to select appropriate media and growth conditions. The soil contains
a wide range of bacteria so it is likely to observe growth in almost any standard bacterial medium.
MATERIALS





3 LB plates/group
Conical tube
Soil sample from TA
Spread beads
Sterile water
PROTOCOL
1.
2.
3.
Take soil sample, weigh 1 g, and place into conical tube.
Determine 3 different ways to plate this sample to visualize microbes.
Record all three procedures in lab notebook.
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Catalase Reaction
Adapted from:
Karen Reiner – Microbe Library
HISTORY
In order to survive, organisms must rely on defense mechanisms that allow them to repair or escape the oxidative
damage of hydrogen peroxide (H2O2). Some bacteria produce the enzyme catalase, which facilitates cellular
detoxification. Catalase neutralizes the bactericidal effects of hydrogen peroxide (13) and its concentration in
bacteria has been correlated with pathogenicity (8).
Enzyme-based tests play a crucial part in the identification of
bacteria. In 1893, a publication by Gottstein brought attention to bacterial catalase, making it one of the first
bacterial enzymes to be described (6, 9). Some 30 years later, McLeod and Gordon (9) developed and published
what is thought to be the first bacterial classification scheme based on catalase production and reactions (6).
PURPOSE
The catalase test facilitates the detection of the enzyme catalase in bacteria. It is essential for differentiating
catalase-positive Micrococcaceae from catalase-negative Streptococcaceae. While it is primarily useful in
differentiating between genera, it is also valuable in speciation of certain gram positives such as Aerococcus urinae
(positive) from Aerococcus viridians (negative) and gram-negative organisms such as Campylobacter fetus,
Campylobacter jejuni, and Campylobacter coli (all positive) from other Campylobacter species (7, 8). Some have
reported its value in the presumptive differentiation between certain Enterobacteriaceae (11). The catalase test is
also valuable in differentiating aerobic and obligate anaerobic bacteria, as anaerobes are generally known to lack
the enzyme (8, 9). In this context, the catalase test is valuable in differentiating aerotolerant strains of Clostridium,
which are catalase negative, from Bacillus, which are catalase positive (8).
THEORY
The catalase enzyme serves to neutralize the bactericidal effects of hydrogen peroxide (13). Catalase expedites
the breakdown of hydrogen peroxide (H2O2) into water and oxygen (2H2O2 + Catalase → 2H2O + O2). This reaction
is evident by the rapid formation of bubbles (2, 7).
RECIPE
For routine testing of aerobes, use commercially available 3% hydrogen peroxide (2, 7). Store the
hydrogen peroxide refrigerated in a dark bottle. For the identification of anaerobic bacteria, a 15% H2O2 solution is
necessary (1). In this context, the catalase test is used to differentiate aerotolerant strains of Clostridium, which
are catalase negative, from Bacillus species, which are positive (8).
The superoxol catalase test used for the
presumptive speciation of certain Neisseria organisms requires a different concentration of H2O2. Refer to the
“Additional Recommendations” section for details.
PROTOCOL
There are many applications and method variations of the catalase test. These include the slide or drop catalase
test, the tube method, the semi-quantitative catalase for the identification of Mycobacterium tuberculosis, the
heat-stable catalase used for the differentiation of Mycobacterium species, and the capillary tube and cover slip
method (7). We will focus on the slide (drop) and tube method here. One of the most popular methods in clinical
bacteriology is the slide or drop catalase method, because it requires a small amount of organism and relies on a
relatively uncomplicated technique. This protocol delineates the procedure for the qualitative slide and tube
catalase methods, which are primarily used for the differentiation of staphylococci and streptococci.
Slide (drop) method
1. Place a microscope slide inside a petri dish. Keep the petri dish cover available. The use of a petri dish is
optional as the slide catalase can be properly performed without it. However, to limit catalase aerosols,
which have been shown to carry viable bacterial cells (4), the use of a petri dish is strongly recommended.
2. Using a sterile inoculating loop or wooden applicator stick, collect a small amount of organism from a
well-isolated 18- to 24-hour colony and place it onto the microscope slide. Be careful not to pick up any
agar. This is particularly important if the colony isolate was grown on agar containing red blood cells.
Carryover of red blood cells into the test may result in a false-positive reaction (5, 7).
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3.
4.
5.
6.
Using a dropper or Pasteur pipette, place 1 drop of 3% H 2O2 onto the organism on the microscope slide.
Do not mix. Immediately cover the petri dish with a lid to limit aerosols and observe for immediate
bubble formation (O2 + water = bubbles).
Observing for the formation of bubbles against a dark background enhances readability. Positive
reactions are evident by immediate effervescence (bubble formation) (Fig. 1).
Place microscope slide over a dark background and use a magnifying glass or microscope to observe weak
positive reactions. No bubble formation (no catalase enzyme to hydrolyze the hydrogen peroxide)
represents a catalase-negative reaction (Fig. 1).
Quality control is performed by using organisms known to be positive and negative for catalase.
Note: If a platinum inoculating loop is used, do not add 3% H 2O2 to the slide before the organism, as the platinum
wire in the loop may produce a false-positive result. FIG. 1. Slide catalase test results. (Top) The positive reaction was produced by Staphylococcus aureus; (bottom)
the negative reaction was produced by Streptococcus pyogenes.
Tube method (10)
1. Add 4 to 5 drops of 3% H2O2 to a 12 x 75-mm test tube (10).
2. Using a wooden applicator stick, collect a small amount of organism from a well-isolated 18- to 24-hour
colony and place into the test tube. Be careful not to pick up any agar. This is particularly important if the
colony isolate was grown on agar containing red blood cells. Carryover of red blood cells into the test
may result in a false-positive reaction (5, 7).
3. Place the tube against a dark background and observe for immediate bubble formation (O 2 + water =
bubbles) at the end of the wooden applicator stick.
4. Positive reactions are evident by immediate effervescence (bubble formation) (Fig. 2A). Use a magnifying
glass or microscope to observe weak positive reactions.
5. No bubble formation (no catalase enzyme to hydrolyze the hydrogen peroxide) represents a catalasenegative reaction (Fig. 2B).
6. Quality control is performed by using organisms known to be positive and negative for catalase.
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A B
FIG. 2. Tube catalase test results. (A) The positive reaction was produced by Staphylococcus aureus; (B) the
negative reaction was produced by Streptococcus pyogenes.
SAFETY
COMMENTS AND TIPS
Always use strict aseptic techniques. If possible, perform the test under a biosafety hood.
Do not use this procedure for Mycobacteria testing. In addition to potential self-contamination by aerosol
exposure, Mycobacteria speciation by the catalase test is achieved by different techniques not discussed in this
protocol (3, 5, 7).
Positive superoxol organisms such as Neisseria gonorrhoeae produce immediate, vigorous
bubbling (8). Other Neisseria species produce delayed bubbling or none at all (8). The superoxol test is performed
the same way as the 3% catalase test.
REFERENCES
1. Bartelt, M. 2000. Diagnostic bacteriology, a study guide. F. A. Davis Co., Philadelphia, PA.
2. Clarke, H., and S. T. Cowan. 1952. Biochemical methods for bacteriology. J. Gen. Microbiol. 6:187–197.
3. Cowan, S. T., and K. J. Steel. 1965. Identification of medical bacteria. University Press, Cambridge, MA.
4. Duke, P. B., and J. D. Jarvis. 1972. The catalase test—a cautionary tale. J. Med. Lab Technol. 29(2):203–
204.
5. Forbes, B. A., D. F. Sahm, and A. S. Weissfeld. 2007. Bailey and Scott’s diagnostic microbiology, 12th ed.
Mosby Company, St. Louis, MO.
6. Gagnon, M., W. Hunting, and W. B. Esselen. 1959. A new method for catalase determination. Anal. Chem.
31:144.
7. MacFaddin, J. F. 2000. Biochemical tests for identification of medical bacteria, 3rd ed. Lippincott Williams &
Wilkins, Philadelphia, PA.
8. Mahon, C. R., D. C. Lehman, and G. Manuselis. 2011. Textbook of diagnostic microbiology, 4th ed. W. B
Saunders Co., Philadelphia, PA.
9. McLeod, J. W., and J. Gordon. 1923. Catalase production and sensitiveness to hydrogen peroxide amongst
bacteria: with a scheme for classification based on these properties. J. Pathol. Bacteriol. 26:326–331.
10. South Bend Medical Foundation. 2010. Catalase test protocol. South Bend Medical Foundation, South Bend,
IN.
11. Taylor, W. I., and D. Achanzar. 1972. Catalase test as an aid to the identification of Enterobacteriaceae. J.
Appl. Microbiol. 24:58–61.
12. Thomas, M. 1963. A blue peroxide slide catalase test. Mon. Bull. Min. Health 22:124–125.
13. Wheelis, M. 2008. Principles of modern microbiology. Jones & Bartlett Publishers, Inc., Sudbury, MA.
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Colony Morphology Protocol
Adapted from:
Donald Breakwell, Christopher Woolverton, Bryan MacDonald, Kyle Smith, Richard Robison – Microbe Library
HISTORY
Since bacteria were first cultured on solid media, describing the appearance of bacterial colonies has been an
important tool for microbiologists. Observing colony morphology is a tool used by clinical microbiologists, in
particular, and descriptions of colonies are often found in the primary literature. Distinguishing colony
morphology is one of the first skills taught to microbiology students.
In an early text, Principles of Microbiology, Moore defined the practice as “The examination of plate
cultures…determining the character of the different colonies, their action upon the medium, the rapidity of their
development, and in case of quantitative analysis, the number and variety of colonies” (2). This practice remains
consistent. Many different terms have been used to classify colonies themselves, however, and systems differ
from simple to complex. Another early text suggested that colonies be described as conglomerate, rhizoid, curled,
or myceloid (4). Later, additional morphological terms such as granular, arborescent, wavy interlaced, and
filamentous” (1) and elevation, edge, color, and texture were used. In Methods for General and Molecular
Bacteriology the terms were categorized by descriptions of color, form, elevation, margin, opacity, and texture (5).
Regardless of terminology, the practice of making observations of colony morphology remains a common exercise
in introductory microbiology laboratory courses. Current laboratory manuals seem to be fairly universal in their
use of the system proposed in Methods for General and Molecular Bacteriology (5). While different morphological
characteristics are never comprehensively described or exemplified using photographic images of bacterial
colonies, simple drawings are used to demonstrate just a few of the morphological characteristics instead (Fig. 1).
PURPOSE
Determining the morphology of a single colony growing on the surface of a plate culture can be an important tool
in the description and identification of microorganisms.
THEORY
On solid media, a colony is theoretically derived from a single cell. If well separated from other colonies, a colony
will have a characteristic shape (both in elevation and margin), size, color, and consistency. Observation is often
made with the naked eye, but dissecting microscopes are also used. The characteristics defined by a colony’s
morphology may be used at a superficial level to distinguish between types of microorganisms. For example, there
are differences in morphologies when rough and smooth colonies of Streptococcus pneumonieae are examined.
Another comparison can be made when describing pigmented colonies.
RECIPES
Cultures of microorganisms can be grown on any medium that is appropriate for their isolation and cultivation.
Since morphology is influenced by medium type and growth conditions, care should be taken to record these
parameters. Good determination of colony morphology is predicated on good streak technique because it
requires good separation of colonies.
PROTOCOL
Smibert and Krieg (5)
1. Measure the colony diameter in millimeters.
2. Describe the pigmentation (distinguishing between pigmented colonies and those secreting diffusible
pigments).
3. Describe the form, elevation, and margin as indicated in Fig. 1. Also indicate whether the colonies are
smooth (shiny glistening surface), rough (dull, bumpy, granular, or matte surface), or mucoid (slimy or
gummy appearance).
4. Record the opacity of the colonies (transparent, translucent, or opaque) and their texture when tested
with a needle: butyrous (buttery texture), viscous (gummy), or dry (brittle or powdery).
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FIG. 1. Diagram
illustrating the various
forms, elevations, and
margins of bacterial
colonies (3).
FIG. 2. Colonies of Sinorhizobium meliloti grown on trypticase
soy agar are approximately 1 to 2 mm in diameter. They lack
pigmentation and are translucent. Colonies are smooth and
are circular in form with an entire margin. They have a convex
elevation. When manipulated with a needle, the colonies are
viscous.
SAFETY
Students working with live cultures of microbes must be able to explain and practice safe laboratory techniques.
Good laboratory practice for microbiology students includes appropriate aseptic technique for protecting
themselves and others. Physical barriers and cleanliness are simple measures against contamination of students
and laboratory equipment. Working with gloves is a good idea when streaking with pathogens or suspected
pathogens. Use of biological safety cabinets, splatter shields, and protective eye equipment is also recommended.
TIPS AND COMMENTS
Current availability of inexpensive digital cameras can enhance the teaching environment. In addition to having
students make representative drawings of what they observe, they can provide a digital image for the instructor to
compare. For many of the images originally submitted, a Leica EZ4 D 3.0 MPixel stereomicroscope with camera
connected to a personal computer with Las EZ software was used for image collection. Images were cropped using
Adobe Photoshop with the only manipulation being image and file sizes. Images were taken with culture dishes
flat in the field of view or placed at varying angles up to about 80 o. Beyond 80o, the edge of the dish obstructed
the view. It is difficult when using a dissecting scope to determine the exact magnification of colonies, hence
this parameter is missing from legends in this Atlas collection. However, most colonies were between 1 and 3 mm
in diameter. The purpose of this Atlas collection is to provide examples and descriptions of commonly seen colony
morphologies, any of which might be similar to those found in the literature.
Colonies should be well isolated.
This is particularly true when several colonies have coalesced and may appear to have an irregular margin. Please
examine Figure 9 in the Atlas as an example of this.
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REFERENCES
1. Lamanna, C., and F. Mallette. 1953. Basic bacteriology: and its biological and chemical background. Williams
and Wilkins Company, Baltimore, MD.
2. Moore, V. 1912. Principles of microbiology. Carpenter and Company, Ithaca, NY.
3. Pelczar, M. J., Jr. 1957. Manual of microbiological methods. McGraw-Hill Book Co., New York, NY. 4. Reed, H. 1914. A manual of bacteriology: for agricultural and general science students. Ginn and Company,
Boston, MA. 5. Smibert, R. M., and N. R. Krieg. 1994. Phenotypic characterization, p. 615. In P. Gerhardt, R. Murray, W.
Wood, and N. Krieg (ed.), Methods for general and molecular bacteriology. ASM Press, Washington, DC. 124
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Colony PCR
Polymerase chain reaction (PCR) is one of the most commonly used techniques in biology labs, used to amplify
DNA for a variety of purposes. Its applications range from DNA sequencing to functional analysis of genes to
finding genetic fingerprints in a paternity test. PCR takes away the limitations of having too little material, and
often full of impurities, for nucleotide sequence analysis, which is fundamental to fields like molecular biology and
medicine. This has also paved the way for the development of functional genomics and projects like unraveling the
human genome.
Developed in 1983 by biochemist Kary Mullis, this elegant technique uses the same mechanism cells use to make
copies of their genetic material during reproduction, taking advantage of the basic molecular properties of DNA
and the DNA replication machinery. During the reaction, the template DNA strand must be denatured (at 95 °C),
allowed to anneal with the appropriate primers (at about 55 °C), and elongated by DNA polymerase at the
enzyme’s optimal temperature (72 °C) all in the same reaction mix, and this must be repeated up to 30 times to
make millions of copies! Therefore, finding a heat-resistant DNA polymerase that would not be denatured and lost
in every cycle was key to increasing the efficiency and reliability of the reaction.
An extreme bacterium held the answer to this problem: a thermophilic or “heat loving” bacterium isolated from
the hot spring of Yellowstone National Park, Themus aquaticus. Its DNA polymerase could withstand near-boiling
temperatures and replicate DNA with high fidelity. Taq DNA polymerase became the standard enzyme used in PCR
(Saiki et al., 1988) revolutionizing our ability to study DNA and transforming the fields of biology and medicine.
MATERIALS




1492R primer
23F primer
PCR beads
Sterile nuclease-free water
PROTOCOL*
1.
Acquire tubes containing PCR bead, which contains Taq polymerase (heat resistant enzyme) and
other necessary reagents. Label tubes appropriately.
Dispense 23 μL of ultra-pure sterile water.
Add 1 μL of forward primer (27F, 25 μM).
Add 1 μL of reverse primer (1492R, 25 μM).
Carefully scrape off colony from plate with micropipette tip. This contains your template DNA.
Mix with tube contents by gently swishing up and down with pipette.
Cap tubes and gently flick to mix contents. If necessary, vortex and centrifuge for a few seconds.
Transfer tubes to thermal cycler (PCR machine).
Select appropriate program* to start cycling (about 2 hours).
Once cycling is complete, remove tubes and keep in ice until the next step.
2.
3.
4.
5.
6.
7.
8.
9.
10.
*(Protocol adapted from “puRe Taq Ready-To-Go PCR Beads” guide)
Thermal cycler program: On Main Menu, select 16amp, Standard, and customize stages in the protocol as follows:
1. 95 °C for 10 mins (1 cycle)
2. 2: 95 °C for 30 secs (1 cycle)
3. 58 °C for 45 s (1 cycle)
4. 72 °C for 1:40 mins* (30 cycles)
5. 20 °C for ∞ (until retrieved)
*1 min per kb of DNA template
REFERENCES
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Saiki, R. K., Gelfand, D. H., Stoffel, S., Scharf, S. J., Higuchi, R., Horn, G. T., . . . Erlich, H. A. (1988). Primer-directed
enzymatic amplification of DNA with a thermostable DNA polymerase. Science, 239(4839), 487-491.
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MacConkey Agar Test
Adapted from:
Mary E. Allen – Microbe Library
HISTORY
MacConkey agar was the first solid differential media to be formulated. It was developed at the turn of the 20th
century by Alfred Theodore MacConkey, M.D, then Assistant Bacteriologist to the Royal Commission on Sewage
Disposal, in the Thompson-Yates Laboratories of Liverpool University, England. The goal was to formulate a
medium that would select for the growth of gram-negative microorganisms and inhibit the growth of grampositive microorganisms. Dr. MacConkey first developed a bile salt medium containing glycocholate, lactose and
litmus, to be incubated at 22°C (MacConkey, 1900). This formula was soon altered by the replacement of
glycocholate with taurocholate and the incubation temperature was raised to 42°C (MacConkey, 1901).
MacConkey later changed the recipe again by substituting neutral red for litmus (MacConkey, 1905), following the
suggestion that neutral red be used as an indicator in bile salt lactose medium (Grunbaum and Hume, 1902). The
final media formulation was designed to support growth of Shigella and is the one that is most commonly used
today.
PURPOSE
MacConkey agar is used for the isolation of gram-negative enteric bacteria and the differentiation of lactose
fermenting from lactose non-fermenting gram-negative bacteria. It has also become common to use the media to
differentiate bacteria by their abilities to ferment sugars other than lactose. In these cases lactose is replaced in
the medium by another sugar. These modified media are used to differentiate gram-negative bacteria or to
distinguish between phenotypes with mutations that confer varying abilities to ferment particular sugars.
THEORY
MacConkey agar is a selective and differential media used for the isolation and differentiation of non-fastidious
gram-negative rods, particularly members of the family Enterobacteriaceae and the genus Pseudomonas. The
inclusion of crystal violet and bile salts in the media prevent the growth of gram-positive bacteria and fastidious
gram-negative bacteria, such as Neisseria and Pasteurella. The tolerance of gram-negative enteric bacteria to bile
is partly a result of the relatively bile-resistant outer membrane, which hides the bile-sensitive cytoplasmic
membrane (Nikaido, 1996). Other species-specific bile-resistance mechanisms have also been identified
(Provenzano, et al. 2000; Thanassi et al. 1997).
Gram-negative bacteria growing on the media are differentiated by their ability to ferment the sugar lactose.
Bacteria that ferment lactose cause the pH of the media to drop and the resultant change in pH is detected by
neutral red, which is red in color at pH's below 6.8. As the pH drops, neutral red is absorbed by the bacteria, which
appear as bright pink to red colonies on the agar.
The color of the medium surrounding gram-negative bacteria may also change. Strongly lactose-fermenting
bacteria produce sufficient acid to cause precipitation of the bile salts, resulting in a pink halo in the medium
surrounding individual colonies or areas of confluent growth. Bacteria with weaker lactose fermentation growing
on MacConkey agar will still appear pink to red but will not be surrounded by a pink halo in the surrounding
medium.
Gram-negative bacteria that grow on MacConkey agar but do not ferment lactose appear colorless on the medium
and the agar surrounding the bacteria remains relatively transparent.
Lactose can be replaced in the medium by other sugars and the abilities of gram-negative bacteria to ferment
these replacement sugars are detectable in the same way as is lactose fermentation (for example Farmer and
Davis, 1985).
RECIPE
Peptone (Difco) or Gelysate (BBL)
Proteose peptone (Difco) or Polypeptone (BBL)
Lactose
17.0 g
3.0 g
10.0 g
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NaCl
5.0 g
Crystal Violet
1.0 mg
Neutral Red
30.0 mg
Bile Salts
1.5 g
Agar
13.5 g
Distilled Water
Add to make 1 Liter
Adjust pH to 7.1 +/-0.2. Boil to dissolve agar. Sterilize at 121° C for 15 minutes. (Holt and Krieg, 1994, Remel
2005)
PROTOCOL
Streak a plate of MacConkey's agar with the desired pure culture or mixed culture. If using a mixed culture use a
streak plate or spread plate to achieve colony isolation. Good colony separation will ensure the best
differentiation of lactose fermenting and non-fermenting colonies of bacteria.
Streak plate of Escherichia coli and Serratia marcescens on MacConkey agar as controls. Both microorganisms
grow on this selective media because they are gram-negative non-fastidious rods. Growth of E. coli, which
ferments lactose, appears red/pink on the agar. Growth of S. marcescsens, which does not ferment lactose,
appears colorless and translucent.
REFERENCES
1. Difco Manual, Tenth Edition. 1984. Difco Laboratories, Inc. Detroit, MI., U.S.
Grunbaum, A.S. and Hume, E. H. 1902. "Note on media for distinguishing B. coli, B. typhosus and related
species." British Medical Journal, i: 1473-1474.
2. Holt, J.G. and Krieg, N.R. 1994. "Chapter 8. Enrichment and Isolation." In [Eds.] Gerhardt, P., R.G.E. Murray,
W.A. Wood and N.R. Krieg. Methods for General and Molecular Bacteriology. ASM Press, Washington, D.C.
pg.205
3. Collard, Patrick. 1976. "The Development of Microbiology". Cambridge University Press, pp.31-32.
4. Farmer JJ 3rd and Davis BR. 1985. "H7 antiserum-sorbitol fermentation medium: a single tube screening
medium for detecting Escherichia coli O157:H7 associated with hemorrhagic colitis." J Clin Microbiol. (4):620-5.
5. Gerhardt, P., R.G.E. Murray, W.A. Wood and N.R. Krieg. Methods for General and Molecular Bacteriology.
ASM Press, Washington, D.C. pg.205
6. MacConkey, A. 1900. "A note on a new medium for the growth and differentiation of the bacillus Coli
communis and the bacillus Typhi abdominalis." Lancet, ii:20.
7. MacConkey, A. 1901. "Corrigendum et addendum." Zentralblatt fur Bakteriologie, 29: 740.
8. MacConkey, A. 1905. Lactose-fermenting bacteria in feces. J. Hyg.. 5:333-378.
9. Nikaido, H. 1996. Outer membrane, p. 29-47. In F. C. Neidhardt, R. Curtiss III, J. L. Ingraham, E. C. C. Lin, K.
B. Low, Jr., B. Magasanik, W. S. Reznikoff, M. Riley, M. Schaechter, and H. E. Umbarger (ed.), Escherichia coli
and Salmonella: cellular and molecular biology, 2nd ed. ASM Press, Washington, D.C.
10. Provenzano D, Schuhmacher DA, Barker JL, Klose KE. 2000 The virulence regulatory protein ToxR mediates
enhanced bile resistance in Vibrio cholerae and other pathogenic Vibrio species. Infect Immun. 68(3):1491-7
11. Remel Microbiology Products. Instructions for Use of MacConkey Agar. Accessed June 2005,
http://www.remelinc.com/IFUs/IFU1550.pdf
12. Ryan, K.J. and C.G. Ray (Ed.). 2004. Sherris Medical Microbiology. An Introduction to Infectious Disease. 4th
Edition. McGraw-Hill, New York City, U.S.
13. Thanassi, D. G., L. W. Cheng, and H. Nikaido. 1997. Active efflux of bile salts by Escherichia coli. J.
Bacteriol. 179:2512-2518
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Picking and patching colonies
Isolating single species of bacteria from a mixed culture containing tens or hundreds of species is a routine
procedure that allows microbiologists to carefully examine an organism and its unique characteristics. Dilution
plating allows us to spread individual bacteria on a plate enough for them to grow into distinct colonies. Yet
colonies overgrow with time and mix with one another, cells migrate, and cultures get contaminated, so keeping
timing, sterile condition in your work environment and proper techniques in mind is very important to isolating a
bacterium and starting a pure culture.
The isolation approach we will use is aptly called “pick and patch” and involves precisely those things: “picking”
bacteria from a mixed culture (e.g., usually a dilution plate) and “patching” them onto a fresh plate, which may be
a pure culture or a “master plate” containing all your unique bacteria of interest for your study. The slight touch of
a colony with a sterile toothpick or a metal rod picks up thousands of bacteria that can be transferred and smeared
or patched onto a fresh plate. At the end of this procedure you will have a “master plate” that will serve as a
bacterial catalog for your experiments.
MATERIALS


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2 media plates of choice
Square grid
Sterile toothpicks
PROTOCOL
1.
2.
3.
4.
5.
Obtain 2 plates of media of choice. Label appropriately and draw small vertical line at edge of plate as
point of orientation.
Tape the plate face-up on a square grid plate. Align grid with vertical line on plate.
Using a sterile toothpick, pick a unique colony from your dilution plate (10 2 or 103). Patch (gently zigzag)
colony smear on toothpick onto fresh media plate within the boundaries of one square on square grid.
Continue to pick and patch colonies onto media plate, each time occupying a new square. Make sure
patches do not overlap or touch as this will contaminate the patch, assuming that you picked a single
colony. Pick 24 morphologically unique colonies (i.e., different textures, colony margins, pigments), yet do
not delve to much into morphology as this is part of a separate experiment.
This collection of colony patches be your master plate.
Master plate schematic. This shows how your plate should look after incubation and allowing the isolates to grow.
Notice how zigzagging patches are separated by a grid (not shown) and are not touching; this is critical.
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Chemotaxis
Bacteria are constantly monitoring their surroundings for molecules that tell them about the presence of other
bacteria or their proximity to a nutrient or the threat of a toxic chemical, among many other extracellular cues.
Many have coupled receptors on the cell surface with flagellar motors that allow them to move towards or away
from an attractant or a repellent, respectively. Known as chemotaxis, this mechanism directs the movement of
motile bacteria in response to organic chemicals in their environment (Grimm & Harwood, 1997). This is mediated
by the binding of specific compounds to chemoreceptors, which in turn transduce the signal intracellularly to
motor proteins in the cytoplasm through a series of subsequent phosphorylations. Chemotaxis gives bacteria the
ability to respond to changes in the environment by moving towards a nutrient (e.g., an amino acid or a sugar) or
away from chemicals that could damage the cell. Alternatively, it could allow certain bacteria to make a bold move
to breaking down toxic organic chemicals, such as Pseudomonas putida, which is chemotactically attracted to the
toxin naphthalene and degrades it (Grimm & Harwood, 1997). Other pseudomonads are attracted to nutrients,
giving them a competitive advantage over other bacteria. In an experiment by de Weert et al., P. fluorescens was
found to out-compete mutants incapable of flagella-driven chemotaxis at colonizing tomato roots in the soil, even
when the mutants retained their ability to move (de Weert et al., 2002). P. fluorescens was attracted to certain
amino acids and organic acids present in root exudates, giving it a great advantage at procuring essential nutrients.
***This example of chemotaxis makes competition an important theme when studying bacteria living in soil
environments, as is the case of antibiotic producing bacteria.
In this experiment, we will perform a “drop assay” to test bacteria for motility and chemotaxis towards nutrients.
In this protocol, which is a modification of the “drop assay” as described by Grimm and Hardwood (Grimm &
Harwood, 1997), bacteria will be suspended in soft agar, poured into a small petri dish and spotted with a drop of a
chemo-attractant, such as malic acid, citric acid or an amino acid (de Weert et al., 2002). Within hours, the bacteria
will aggregate around the chemo-attractant, if they express this particular chemotaxis phenotype. In the process,
the bacteria will oxidize TTC present in the medium and leave behind a visible red trail. This will allow us to assess
motility, which is an important means of differentiating and classifying bacteria, and chemotaxis.
PROTOCOL*
Drop Assay
1. Make overnight culture, dilute 100-fold in 1% succinic acid solution (with minimal medium)
2. Resuspend in 12 ml chemotaxis buffer (100 uM potassium phosphate [ph 7.0], 20 uM EDTA)
3. Add 3 ml of hydroxypropylmethylcellulose (an animal gelatin alternative) in aqueous solution
Making motility medium:
1. Make 10% TSA (0.4 % agar) and add 5 ml of 1% TTC
2. Pour mixture (with cell suspension) into 60 mm petri dish to depth of about 3 mm
3. Add 10 ul drop of appropriate attractant to the center
4. Incubate for 0.5 to 2 h
*I have modified this protocol by combining Jess’ motility assay with Fahrner et al.’s drop assay (Fahrner, Block,
Krishnaswamy, Parkinson, & Berg, 1994).
New protocol (7/17/13)
Grew overnight cultures of S. cohnii (50% TSB) and P. putida (LB)
Diluted 1:10 in respective liquid broths (S. cohnii was switched to 50% TSB)
Mixed with respective cooled down (warm to the touch, but not solidified) 0.5% [soft] agar 10% TSA and LB to a
1:10 concentration. Poured 20 mL into small (60 mm) petri dish.
Allowed agar to solidify.
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Added (to the center of the medium) 10 uL drops of ~ 0.1 M L-isoleucine (dissolved in MeOH), malic acid, and
sucrose.
Incubated at 28 °C.
REFERENCES
de Weert, S., Vermeiren, H., Mulders, I. H., Kuiper, I., Hendrickx, N., Bloemberg, G. V., . . . Lugtenberg, B. J. (2002).
Flagella-driven chemotaxis towards exudate components is an important trait for tomato root
colonization by Pseudomonas fluorescens. Mol Plant Microbe Interact, 15(11), 1173-1180. doi:
10.1094/MPMI.2002.15.11.1173
Fahrner, K. A., Block, S. M., Krishnaswamy, S., Parkinson, J. S., & Berg, H. C. (1994). A mutant hook-associated
protein (HAP3) facilitates torsionally induced transformations of the flagellar filament of Escherichia coli. J
Mol Biol, 238(2), 173-186. doi: 10.1006/jmbi.1994.1279
Grimm, A. C., & Harwood, C. S. (1997). Chemotaxis of Pseudomonas spp. to the polyaromatic hydrocarbon
naphthalene. Appl Environ Microbiol, 63(10), 4111-4115.
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Gel electrophoresis
A simple and highly common technique is agarose gel electrophoresis, which allows us to confirm that we have
acquired a desired product through PCR (polymerase chain reaction). Agarose is a polysaccharide polymer related
to the solidifying agent that is used to make solid cultures, agar, both of which are extracted from seaweed. When
mixed with a specific buffer solution, it solidifies into a gelatinous, porous matrix that allows DNA molecules to
migrate through it when placed in an electric field. This is due to the partial negative charge of DNA molecules,
which attracts them towards the positively charged terminal on the gel electrophoresis apparatus. Differences in
charge, size and shape will cause individual molecules to migrate at different speeds as they force themselves
through the matrix. For example, a small DNA molecule of just a couple hundred base pairs (bp) will migrate much
farther through the gel than a thousand bp DNA strand over time. This property allows us to separate DNA by size,
which is a useful method to determine what genes are in a given sample based on their size, for example, whether
we have amplified a specific piece of DNA of a known size (e.g., the 16S ribosomal RNA gene, which is about 1500
bp long) versus an undesired product of a different size due to a PCR error, contamination in our sample, or a
degrading DNA molecules. The distance traveled by these “bands” of DNA molecules is compared to a marker or
“ladder” of fragments of known sizes that run side-by-side. This allows us to make estimations of the sizes of the
molecules in a PCR product.
MATERIALS
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Agarose
TBE buffer
Ethidium bromide
Erlenmeyer flask
Microwave oven
Loading dye
Gel tray, comb, and gel electrophoresis apparatus
UV chamber
PROTOCOL for making 1% agarose gel:
1.
2.
3.
4.
5.
6.
7.
8.
9.
Weigh 1 g of agarose and mix with 100 mL TBS buffer.
Microwave until boiling (1-2 mins) or until mixture becomes transparent.
Allow mixture to cool down (10-15 mins)
Carefully add 10 μL of ethidum bromide to mixture
(Warning: ethidium bromide is a DNA intercalating agent and mutagen. Handle this compound only in
designated areas. Wear gloves and avoid contact with skin).
Pour mixture into gel tray with appropriate comb and allow mixture to solidify (about 15 mins).
Carefully pull comb off gel. Place gel tray into chamber with TBS buffer and start loading wells:
a. Load 1 kb latter into first well.
b. Mix 5 μL of PCR product with 1 μL loading dye and dispense into wells.
Plug electrodes on appropriate terminals of the chamber (keep in mind that DNA has a partial
negative charge).
Allow gel to run for 40-60 minutes (at 100 milliamps/volt).
Finally, carefully remove gel from tray and place in UV chamber to get a photograph (expect to see
continuous band at about 1.7 kb).
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Spread Plate
Adapted from:
By: Kathryn Wise – Microbe Library
PURPOSE
One method of distributing bacteria evenly over the surface of an agar plate medium is commonly referred to as
the spread plate method. Classically a small volume of a bacterial suspension is spread evenly over the agar
surface using a sterile bent glass rod or glass beads as the spreading device. The goal in evenly distributing the
bacterial suspension is typically to permit the growth of colonies that can subsequently be enumerated (see Serial
Dilution Protocols) and/or sampled following incubation. Each plate is spread with a single inoculum of the
bacterial suspension. An alternative approach to spreading a single inoculum volume with a smooth device is to
apply a smaller volume and tip the plate, allowing gravity to distribute the inoculum in a band or track (track
method) or to allow the inoculum to dry in place (drop method). With this alternative approach, several sample
dilutions can be distributed on a single agar plate.
HISTORY
Since the development of the agar plate in Robert Koch's laboratory, several methods have been used to achieve
an even distribution of bacterial growth on or in the agar. The most common methods used to achieve this type of
distribution are: spread, pour, thin-layer, layered, and membrane filter (2).
PRINCIPLES
Using the spread method a small volume of a bacterial suspension is distributed evenly over the surface of an agar
plate using a smooth sterilized spreader (2). In the case of track plates, gravity is used to spread the inoculum
down the agar in a column forming a track (1).
PROTOCOL
Agar plates:
Select and prepare an agar medium based upon the type of bacteria to be enumerated or selected.
Freshly prepared plates do not work as well as dry plates as it takes longer for the inoculum to absorb into the
agar. Plates may be dried by keeping them at room temperature for roughly 24 hours. Plates will dry faster in
lower humidity so placing them in a laminar flow hood will speed the drying process. Once dried, plates may be
used or refrigerated in closed bags or containers until required. Refrigerated plates should be warmed to room
temperature prior to use. Inoculations:
When enumerating colony-forming units (CFUs), plates with 20 to 200 (or 25 to 250) CFUs can be used to calculate
the number of CFUs/ml of the original sample. Typically a dilution series is prepared, often a ten-fold dilution
series, using a suitable diluent such as phosphate-buffered saline.
Serial Dilution Protocols:
A convenient inoculum volume, in terms of spreading, absorption, and calculations, is 0.1 ml (100 μL). Since some
bacteria rapidly attach to the agar surface, the inoculum should be spread soon after it is applied. Working from
the most dilute suspension to the most concentrated is advised. Proceeding from most dilute to most
concentrated makes it unnecessary to change pipette tips between the dilutions.
Spreading:
A reusable glass or metal spreader should be flame sterilized by dipping in alcohol (such as 70% isopropyl or
ethanol), shaking off the excess alcohol, and igniting the residue. The spreader is then allowed to cool. The
spreader is placed in contact with the inoculum on the surface of the plate and positioned to allow the inoculum to
run evenly along the length of the spreader. Apply even, gentle pressure as the plate or spreader is spun or
rotated. If glass beads are used, ensure they have been properly autoclaved and stored under sterile conditions.
Pour 5-10 beads onto a plate either prior to the addition of inoculum and shake to evenly distribute until the liquid
is absorbed into the plate.
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The goal is to evenly distribute the inoculum and to allow it to be absorbed into the agar. The plate, or spreader,
should be rotated long enough to avoid pooling along the spreader once the rotation is stopped. Incubation:
After the spread plates have been permitted to absorb the inocula for 10 to 20 minutes they may be inverted and
incubated as desired. Observe the plates before the colonies have had time to fully develop. Closely positioned colonies may be difficult
to resolve as separate colonies later. Continue the incubation as necessary. Incubation in closed humidified
containers will help avoid problems with plates drying out when working with slow-growing colonies. Counting and Selection: After appropriate incubation, plates are inspected. When plating a dilution series, the growth on the plates should
reflect the predictable drop in CFUs/plate as illustrated in this of a 10-fold dilution series prepared from an
overnight broth culture of Escherichia coli.
FIG. 1. Picture of spread plates showing
bacterial growth (E. coli, 40 hours, room
temperature) on five plates prepared from a
ten-fold dilution series. Care was taken to
avoid spreading to the edges of the plates, as it
is more difficult to count colonies along the
edge of the agar.
Duplicate or triplicate plates with 30 to 300
CFUs/plate are used to calculate CFUs/ml.
Plates with well isolated colonies may be
inspected and, if desired, colonies "picked" to
establish new cultures.
COMMENTS AND TIPS
 Make sure plates are sufficiently dry prior to use.  Plates should be prepared in duplicate or triplicate.  Do not delay in spreading the inoculum once it has been applied to the plate since some cells will rapidly
attach to the agar, especially if the plate agar is nice and dry.  Avoid spreading the inoculum to the edge of the agar as it is more difficult to inspect and count colonies
along the agar's edge.  Once the dilution series has been made, inoculate plates within 30 minutes to minimize changes in the
number of cells in each dilution due to cell division or death.  Make sure even pressure is applied to the spreader so that fluid is evenly distributed along its length as
the plate or spreader is rotated.  Once the dilutions are made, work backwards spreading the most dilute samples first.  When making your own spreaders do not make the spreading edge too long, it should conveniently fit
into the alcohol container as well as the plate. Fire-polish the end of the spreader the student will hold.
Bending the glass to form a triangle rather than an "L" will help assure only smooth even surfaces touch
the agar and minimize pooling.  Distributing the organisms by rotating the spreader rather than the plate tends to cause more pooling of
the inoculum.  Sterile glass or plastic beads are sometimes used to spread the inoculum over the surface of an agar plate
when an even distribution of colonies is not an important outcome.
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Streak Plate
Adapted from:
Microbe Library
HISTORY
The modern streak plate procedure has evolved from attempts by Robert Koch and other early microbiologists to
obtain pure bacterial cultures in order to study them, as detailed in an 1881 paper authored by Koch (5).
The earliest appearance of the three-sector streak pattern (called the T streak) commonly used today may be the
1961 photos published in Finegold and Sweeney (4). An illustration detailing how to perform this streak is in the
1968 edition of the Manual of BBL Products and Laboratory Procedures (1). In addition to the T streak, the BBL
Manual illustrates two other streak patterns, neither of which is the simple mono-directional streak pattern used
earlier in the century.
FIG. 1. A three sector T streak of Serratia
marcescens grown on trypticase soy agar.
This illustrates a streak plate which has
many isolated colonies.
FIG. 2. This plate illustrates a streak plate which did not
achieve isolation, and which would not be considered a
good streak plate example. This photograph is by Dr.
Min-Ken Liao, Furman University.
PURPOSE
The purpose of the streak plate is to obtain isolated colonies from an inoculum by creating areas of increasing
dilution on a single plate. Isolated colonies represent a clone of cells, being derived from a single precursor cell.
When culture media is inoculated using a single isolated colony, the resulting culture grows from that single clone.
Historically, most microbiology research and microbial characterization has been done with pure cultures.
THEORY
One bacterial cell will create a colony as it multiplies. The streak process is intended to create a region where the
bacteria are so dilute that when each bacterium touches the surface of the agar, it is far enough away from other
cells so that an isolated colony can develop. In this manner, spreading an inoculum with multiple organisms will
result in isolation of the different organisms.
PROTOCOL
Mesophilic bacteria are generally streaked onto media solidified with 1.5% agar or agarose. Gelatin can be used if
a high enough concentration of gelatin protein or a low enough incubation temperature is used. Thermophiles and
hyperthermophiles can also be streaked onto growth media solidified with agar substitutes, such as Gelrite and
guar gum.
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One-hundred-mm-diameter petri dishes are the most commonly used size of plate for streaking. The agar surface
of the plate should be dry without visible moisture such as condensation drops. Traditionally, inoculated petri
dishes are incubated with the agar side up to prevent condensed moisture from falling onto the agar surface,
which would ruin the isolation by allowing bacteria to move across the moist surface creating areas of confluent
growth instead.
The inoculum for a streak plate could come from any type of source, for example clinical specimen, sedimented
urine, environmental swab, broth, or solid culture. The two most common streak patterns are the three sector T
streak and the four sector quadrant streak.
In a streak plate, dilution is achieved by first spreading the specimen over the agar surface of one sector. If a
cotton swab or disposable loop or needle was used to inoculate the first sector, it is now discarded into an
appropriate container, while reusable loops, usually with nichrome or platinum wire (24 gauge), are flamed to
incinerate any organisms on the loop. When cooled, the sterile loop is streaked through the initial sector and
organisms are carried into the second sector where they are spread using a zig-zag movement. In a similar
manner, the organisms present on the loop are incinerated after the second sector is streaked, and the third
sector is streaked. For a four quadrant plate, the process is carried an extra step.
Detailed procedure for a Three Sector Streak, the T Streak:
Reference J. Lammert, Techniques in Microbiology, A Student Handbook (6)
MATERIALS
▪ Specimen to be streaked; this protocol is written for a test tube culture
▪ Wooden stick, plastic, disposable transfer loop, or reusable metal transfer loop (usually nichrome, a
nickel-chromium alloy, or platinum; the single-use disposable plastic loop can be discarded between
sectors rather than resterilized)
▪ Bunsen burner
▪ Sterile petri dish with appropriate bacterial media
▪ Labeling pen
▪ Sterile cotton swabs (if necessary to remove condensation from the agar surface and from around the
inner rim of the petri dish)
PROTOCOL
1. Label a petri dish: Petri dishes are labeled on the bottom rather than on the lid. In order to preserve
area to observe the plate after it has incubated, write close to the edge of the bottom of the plate. Labels
usually include the organism name, type of agar, date, and the plater's name or initials.
2.
Obtain sterile loop or wooden stick or sterilize a metal transfer loop by flaming in Bunsen burner so that
the entire wires is red-hot: When manipulating bacteria, transfer loops, sticks, etc are usually held like a
pencil. If plastic disposable loops are being utilized, they are removed from the packaging to avoid
contamination and after being used, are discarded into an appropriate container. A new loop is
recommended for each sector of an isolation streak plate.
3.
Open the culture and collect a sample of specimen using the sterile loop or the sterile wooden stick:
Isolation can be obtained from any of a variety of specimens. This protocol describes the use of a mixed
broth culture, where the culture contains several different bacterial species or strains. The specimen
streaked on a plate could come in a variety of forms, such as solid samples, liquid samples, and cotton or
foam swabs. Material containing possibly infectious agents should be handled appropriately in the lab
(see biosafety references below), only by students with appropriate levels of skill and expertise. 4.
Remove the test tube cap: It is recommended that the cap be kept in your right hand (the hand holding
the sterile loop). Curl the little finger of your right hand around the cap to hold it or hold it between the
little finger and third finger from the back. See the illustration. Modern test tube caps extend over the
top of the test tube, keeping the rim of the test tube sterile while the rim of the cap has not been exposed
to the bacteria. The cap can also be placed on the disinfected table, if the test tube is held at an angle so
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that air contamination does not fall down into the tube and the test tube cap is set with the sterile rim on
the table.
Insert the loop or wooden stick into the culture tube and remove the loop or stick. Replace
the cap of the test tube and put it back into the test tube rack. 5.
Streak the plate: Inoculating the agar means that the lid will have to be opened. Minimize the amount of
agar and the length of time the agar is exposed to the environment during the streak process.
a. Streak the first sector: Raise the petri dish lid to insert the loop or wooden stick. Touch the
loop/stick to the agar area on the opposite side of the dish, the first sector. Bacteria on the
loop/stick will be transferred to the agar. Spread the bacteria in the first sector of the petri dish
by moving the loop/stick in a back and forth manner across the dish, a zig-zag motion. Make the
loop movements close together and cover the entire first region. The loop should glide over the
surface of the agar; take care not to dig into the agar.
b. Between sectors: Remove the loop/stick from the petri dish and obtain another sterile loop/stick
before continuing to the second sector. Either incinerate the material on the loop or obtain a
sterile loop if using plastic disposable loops. The loop must be cool before streaking can
continue. Metal loops can be touched to an uninoculated area of agar to test whether they are
adequately cooled. If the loop is cool, there will be no sizzling or hissing and the agar will not be
melted to form a brand. If a brand is formed, avoid that area when continuing with the streaking
process.
c. Streak the second sector: Open the petri dish and insert the loop/stick. Touch the loop/stick to
the first sector once, invisibly drawing a few of the bacteria from the first sector into the second
sector. The second sector is streaked less heavily than the first sector, again using a zig-zag
motion.
d. Obtain a sterile loop/stick for the third sector (see 2, above).
e. Streak the third sector: Open the petri dish and insert the loop/stick again. Touch the loop/stick
once into the second sector and draw bacteria from the second sector into the third sector.
Streak the third sector with a zig-zag motion. This last sector has the widest gap between the
rows of streaking, placing the bacteria a little further apart than in the previous two sectors.
Watch closely to avoid touching the first sector as the streak is completed.
SAFETY Always dispose of used loops, sticks, or any inoculating tools appropriately. Flame the metal loop one last time to
sterilize for proper storage.
COMMENTS AND TIPS
 Alternate streak patterns and different culture media: A variety of alternate streak patterns exist. Some
are used for specific inocula, such as a urine specimen. The patterns also differ in the number of sectors
as well as in the number of times the loop is sterilized. The four quadrant streak pattern would be
recommended for use when large amounts of bacteria are expected in the inoculum. The extra sector will
provide additional dilution and increase the probability of isolated colonies on the plate. The four
quadrant streak plate is described in a variety of references, e.g., in Cappuccino and Sherman's
Microbiology, A Laboratory Manual, 8th ed. (3).

Sometimes, cultures will be streaked on enrichment media or various selective and differential media.
For instance, a culture which is expected to have a gram-negative pathogen will be streaked on a
MacConkey agar plate, which inhibits the growth of gram-positive organisms.

Incinerating material on transfer loops—flaming: Reusable microbiological loops and needles are
sterilized by flaming. A Bunsen burner is traditionally used for this process. Most microbiology manuals
show the microbiologist positioned with his/her hand above the burner, with the loop placed into the
flame. To avoid possible contact with the flame, the microbiologist might consider placing his/her hand
below the flame with the loop/needle above the hand in the flame. The flame of the Bunsen burner
should be adjusted to blue, with the darker blue cone of cooler air visible in the center of the flame. The
loop or needle should be placed into the hotter part of the flame and kept there until it glows red. When
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
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







flaming, the wire loop is held in the light blue area of a bunsen burner just above the tip of inner flame of
the flame until it is red-hot. There is a possible aerosolization hazard if the loop or needle contains liquid
or a bacterial clump. These loops and needles should be placed into the heat slowly so that the moisture
evaporates rather than sputters. Once sterile, the loop is allowed to cool by holding it still. Do not wave
it around to cool it or blow on it. .
If an incinerator such as a Bacti-Cinerator is used to sterilize the loop, the loop is to remain inside the
incinerator for 5 to 7 seconds. When warmed up (which will take 5 minutes), the temperature inside the
incinerator is 815°C. The incinerator will take 5 to 10 minutes to warm up to working temperature.
Several techniques decontaminate transfer loops between sectors of a streak plate: flame, dig into agar,
flame once and rotate loop
A variety of methods exist for removing organisms from the loop between sectors. Beginning students
are generally taught to sterilize the loop between each sector by incinerating and then cooling the loop.
Clinical microbiologists practice a variety of methods. Some flame once after the initial sector and then
rotate the loop so that the next sectors can be streaked with an unused side of the loop. Other
laboratorians (as clinical microbiologists name themselves) stab the loop several times into the agar to
clear the loop between sectors.
Isolated colony appearances: Isolated colonies can be described using the traditional colony descriptions.
The Colony Morphology Atlas-Protocol project provides information about bacterial colony appearance
and characteristic photographs. The appearance of an organism can vary. For instance, a colony of an
organism growing in a crowded sector of the plate will not grow as large as the identical organism
growing in isolation. The media composition, pH, and moistness, as well as the length of time and
temperature can all affect the organism's appearance. Colonies selected for subculturing should be
colonies that are isolated, i.e., there is no other colony visibly touching the colony.
Agar with a surface layer of water is not suitable for obtaining isolated colonies. Obvious water drops
should be removed from the surface of the plate and from the rim of the plate by using sterile cotton
swabs. Plates should be incubated agar side up, to avoid condensation that would drop onto the growing
colonies on the agar surface.
Flaming tube mouths: Many protocols suggest flaming the tube mouth before and after removing
organisms from a tube. Flaming was important when test tubes were capped with a cotton plug. Flaming
would still be appropriate if a foam plug were being used. If a screw cap, KimKap, or similar test tube cap
is used, the open end of the tube remains sterile since the cap normally covers that area.
Rehearsing the streak procedure: Some instructors have students practice the streaking procedure on a
piece of paper. The process helps the student visualize the completed product and practice the fine
muscle movements that are required in successful streaking for isolation.
Students may also find that they can visualize the pattern better if they mark the back of the petri dish
(for instance, a T streak divide the plate into three sectors).
Before learning to streak, students should have had the opportunity to work with 1.5% agar media.
Ideally they will have also previously had the opportunity to practice using a loop on a plate to determine
the best angle of approach and the amount of force required to glide the loop over the surface of the agar
without gouging the surface.
Holding the plate while streaking: If possible, adequate lighting should be available to help the
microbiologist follow the tracings of the loop on the agar. For most labs, this means that the petri dish
should be held in one's hand while being streaked in order to reflect the light properly. Additionally, the
length of time the petri dish lid is removed should be minimized in order to limit contamination. There
are several ways to hold the petri dish. Beginning students may find that they obtain the best results
leaving the plate on the lab bench and lifting the lid to work. Other students may find that they can place
the plate upside down on the workbench and lift the agar-containing bottom, hold it to streak and then
quickly replace it into the lid. Yet other students may have the manual dexterity to manipulate the entire
dish in their hand, raising the lid with thumb and two fingers while balancing the plate in the rest of their
hand.
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Analyzing organic extracts for antibiotic production
DAY 1 – PREPARING PLATES
MATERIALS
 2 of each appropriate plate
 Plastic loop
 Streak out plate of isolate
PROTOCOL
1. Obtain 2 plates of the appropriate media used to grow isolates. Find streak-out plates of isolates.
2. Using the large end of the plastic loop, fill half of the loop with single colonies of an isolate.
3. Gently inoculate the entire surface of the fresh media plate by rubbing the loop back and forth on the
plate until the whole plate is covered.
4. Ensure the plate is evenly coated with bacteria. If needed, turn the plate 90o and continue to rub the
loop back and forth.
5. Incubate plates at appropriate temperatures and conditions.
DAY 2 – CHOPPING UP PLATE
MATERIALS
 100ml bottles – enough for all isolates
 Reagent spatula
PROTOCOL
1. Obtain your plates from the previous period.
2. Ensure your isolates have adequately grown on each plate to high concentration.
3. Using a reagent spatula, cut the agar into 5cm pieces.
4. Scoop all the agar pieces of each isolate into a 100ml bottle – label with appropriate isolate name. Push
all the pieces to the bottom of bottle.
5. Place the bottle in a -20oC freezer for two days.
DAY 3 – ORGANIC EXTRACTION
MATERIALS
 50ml conical tube
 Ethyl acetate
 Glass Pasteur pipettes
 Scintillation vials
 Steriflip® Filter Units
 Water
PROTOCOL
1. The TAs will remove all the bottles from the freezer ahead of class to thaw each sample.
2. Ethyl acetate (20 mL) and place on a shaker for ~ 1 hr.
3. Pre-weigh scintillation vials. Note weights in table.
4. Carefully remove the ethyl acetate from the bottle with a glass Pasteur pipette and transfer to clean preweighed scintillation vial.
5. Label vial.
6. Hand vials and table of vial weights to instructor.
DAY 4 – ANALYZE EXTRACTION FOR ANTIBIOTIC ACTIVITY
MATERIALS
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Beaker with ethanol
Empty petri dish
Forceps
Liquid culture of Safe ESKAPE relative
Methanol
Micropipettes
Sterile filter discs
PROTOCOL
1. Your supernatants are now dried to a white powder.
2. Re-suspend the dry powder in methanol to achieve a final concentration of 1mg/ml.
3. Place sterile filter discs onto bottom of empty petri dish. Label plate lid with isolate names.
4. Load 20μl onto filter disc. Cover dish and let dry.
5. Load another 20μl and let dry. Repeat until 80μl total volume is loaded onto the filter disc.
6. While the filter discs are drying, spread 150μl of your safe ESKAPE relative on appropriate media.
7. With sterile forceps, pick up the filter disc and transfer to plate with safe ESKAPE relative spread over.
Use plate lid with the isolates names in the appropriate positions.
8. Add 10μl of water to help diffuse contents of filter disc.
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Antibiotic Resistance Tests
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Appendix A: History of growth media, agar, and petri dishes
Slices of sterilized potatoes became the first solid media employed on which to grow bacteria. This procedure only
worked for a few organisms and only until the bacteria decomposed the potato surface. A search for other
materials led to experimentation with the suitability of gelatin and agar-agar as solidifying agents. Gelatin was
difficult to prepare and difficult to use at room temperature, let alone at the higher temperature of an incubator,
and many bacteria digest the protein. Agar, because of its characteristics of melting only when boiled, rarely being
digested by bacteria, and providing a substance in which other nutrients could be dissolved, proved to be a
suitable material on which to grow bacteria. Agar was originally called agar-agar and is derived from seaweed.
The agar that we use today is the same substance as agar-agar, but it has been processed by the manufacturer.
Agar, as purchased 100 years ago, required filtering before it was clear enough to use in media (12). In the early
eras of microbiology, making media was an extensive process of preparing the extracts of meat or other nutrient
sources, as well as purifying and filtering the gelatin or agar. Before the invention of the autoclave, sterilizing the
media properly was also time consuming. The 1939 edition of An introduction to Laboratory Technique in
Bacteriology, an early microbiology lab manual, contains extensive instruction for students to prepare their own
media from "scratch" (7) for use in the lab. Before R. J. Petri invented the petri dish, flat plates of glass covered
by glass lids were most commonly used to culture organisms in gelatin.
Appendix B: Preparing and pouring plates
After autoclaving, cool the agar to between 45°C and 50°C prior to pouring the plates to minimize the amount of
condensation that forms. The thickness of the agar should be roughly 0.3 cm, which can be achieved by
pouring 15 to 20 ml of media per 100 x 15 mm plate. REFERENCES
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