Objective: To determine the organism's oxygen requirement.
1. Inoculate 5 ml of BHI broth with your unknown organism and incubate overnight. We have found that broth cultures provide much more accurate results than using inoculum from a plate. However, if you are inoculating from a plate, make sure you use a very light inoculum.
2. Obtain a thioglycollate tube and make sure that it does not have more than 20% of the medium in pink color. This may happen due to oxidation of the top layer of the medium. To restore anaerobic conditions, such a tube should be placed in boiling water for 10 minutes and then cooled to room temperature. If you do not see any pink color against a white background, the tube is good to use.
3. Use a sterile narrow thin needle (rather than a thick one), insert into your culture broth and slowly stab a thioglycollate tube to the bottom. Carefully remove the needle along the same stab line. Do not shake the tube or move the needle around, or you will introduce extra oxygen into the medium. The needle should reach all the way to the bottom of the tube.
4. Incubate the tube at 30°C (without any regard to the optimum temperature requirement of your species) for 24 hours before reading the tube.
-- Aerobe: band of growth on the top of the tube. Some species have a tendency to grow very rapidly in thioglycollate tube so that the growth covers a rather thick band from the top and extends to the line of stab where there is oxygen available (brought in by the needle). So it is best to look at the bottom 1-cm of the tube and if it is clear with no growth whatsoever, then you can be sure that you have an aerobe.
-- Microaerophile: band of no growth at the top, then a band of growth extending a short distance down proceeded by no growth to the bottom. The bottom 3-cm of the tube should be clear of any growth.
-- Facultative Anaerobe: growth can occur either throughout the tube or can begin at some point below the surface and extend all the way to the bottom, even in the 1-cm bottom of the tube.
-- Anaerobe: growth only at the bottom fifth of the tube.
Objective: To determine the presence of the oxidase enzymes (e.g. cytochrome c oxidase).
Test Procedure and Interpretation
1. Grow the culture on a BHI plate for 48 hours. Up to 7 day old cultures are fine.
2. Warm the plate to 20-37°C. Pick a good amount of the test organism with a sterile swab and rub onto the reaction area of a DrySlide card. If the organism is oxidase positive, a purple color will develop on the slide within 20 seconds. The slide is saturated with Kovacs' oxidase reagent (1% N, N, N', N' tetra-methyl-p-phenylene diamine dihydrochloride). Oxidase negative colonies do not change the color of the slide in 20 seconds, or if they do, it would be after 20 seconds and thus negative.
-- Most Gram-positive bacteria and all Enterobacteriaceae are oxidase negative.
-- Do not attempt to perform an oxidase test on any colonies growing on media containing glucose, as glucose fermentation will inhibit oxidase enzyme activity, and result in possible false negatives. Oxidase test on Gram-negative rods should be performed only on colonies from nonselective and/or non-differential media to ensure valid results.
-- The culture should not be older than a week, unless the species is a slowgrower. False results may be obtained if the culture is old.
-- The oxidase reagent quickly auto-oxidizes (by free oxygen in the air) and loses its sensitivity. The reagent should be discarded if any precipitate forms. Avoid undue exposure of the reagent to light. The reagent must be made up fresh each week.
-- Time period for color development must be adhered to since a purpleblack color may develop later due to auto-oxidation of reagent and/or a weak positive oxidase organism containing a small quantity of cytochrome c oxidase.
-- As an alternative to Kovacs' reagent, one may use a few drops of a 1:1 mixture of 1% α−naphthol in 95% ethanol and freshly prepared 1% aqueous dimethyl-p-phenylenediamine oxalate.
Objective: To test an organism's susceptibility to the chemical, optochin. Optochin susceptibility tests the fragility of the bacterial cell membrane. This test is mainly used to differentiate between Streptococcus pneumoniae (sensitive) and other Streptococcus species (resistant)
1. Pick a single pure colony with a sterile swab to inoculate a SBA plate. Streak the entire blood agar plate with the swab. Turn plate 90 degrees and re-streak with the same swab. Blood agar plate must be used for optochin testing since all species of Streptococcus are fastidious organisms and require extra enrichment for growth.
2. With alcohol flamed forceps, aseptically remove an optochin disc and apply to the center of the plate. Gently apply pressure to disc so that it adheres to the surface of the plate but do not press disc down into the medium.
3. Invert plate and incubate for 48 hours at your organism's optimum growth temperature.
-- Sensitive (S): A distinct zone of inhibition (5 to 30 mm) with a clear-cut margin around disc.
-- Resistant (R): Growth not inhibited around disc.
-- Occasionally, a few scattered optochin resistant colonies of S. pneumoniae may be observed in a wide zone of inhibition.
-- Occasionally an alpha-Streptococcus spp. may exhibit a very small zone (1 to 2 mm) of inhibition. S. pneumoniae exhibits a zone of inhibition at least 5 mm or greater in diameter.
Objective: To test an organism's susceptibility to the antibiotic novobiocin.
1. Streak a BHI plate using a sterile cotton swab. Turn the plate 90 degrees and restreak with the same swab
2. Using a pair of alcohol flamed forceps, aseptically place a novobiocin disc in the center of the plate. Apply gentle pressure to disc so it adheres to the surface of the agar but try not to press too much to embed the disc into the agar.
3. Incubate the inverted plate 48 hours at your organism's optimum growth temperature.
Sensitive (S): No growth around disc; clear zone around disc.
Resistant (R): Growth not inhibited; growth around disc.
4. Incubate at 37°C
5. Examine at intervals, e.g. after 6 h, and 1 and 2 days (depends on generation time of bacteria) . Freshly prepared medium containing 1% glucose can be used for motility tests on anaerobes.
Motile Bacteria typically give diffuse, hazy growths that spread throughout the medium rendering it slightly opaque.
Objective: To determine the ability of an organism to grow in 7.5% NaCl and ferment mannitol.
Any significant growth indicates the organism is a Staphylococcus species. The phenol red indicator changes to yellow at low (acid) pH, which is a product of fermentation. Therefore, fermentation of mannitol will change the color of agar to yellow. Orange is negative.
Positive: Growth, yellow color (mannitol "+").
Negative: Growth or no growth; red or orange color (mannitol "-").
Objective: Some pathogens are able to produce exoenzymes called hemolysins which lyse red blood cells and thus their action can be demonstrated on a blood agar plate.
1. Using a sterile loop, inoculate a blood plate (SBA) with the pure culture of the organism to be tested using the quadrant method. Also stab the medium in the second quadrant with your loop. (Some hemolysins show their effects better under lower oxygen concentrations.)
2. Incubate for 48 hours at optimum temperature for the organism.
Interpret by noting the reaction around isolated colonies as follows:
Alpha (α) hemolysis: formation of a green or brown zone around the colonies (due to loss of potassium from the red cells).
Beta (β) hemolysis: complete lysis of cells and reduction of released hemoglobin; a clear zone appears around isolated colonies.
Gamma (γ) hemolysis: no hemolytic reaction (no change of the medium surrounding isolated colonies).
-- The reaction should be checked only around isolated colonies. If you do not have isolated colonies on the blood agar, a lighter inoculation should be streaked and the test repeated.
Objective: DNase mediates the hydrolysis of DNA. Methyl Green indicator is stable at pHs above 7.5 but becomes colorless at lower pHs. The hydrolysis of DNA in the agar by bacterial DNase reduces the agar pH.
1. Using a sterile loop, inoculate a DNA+Methyl Green agar plate with the fresh bacterial culture. Use a heavy streak line for each bacterial strain to be tested. Be sure to label the plate bottom properly for each strain.
2. Incubate at 37°C for 48 hrs.
-- The test is positive if clearing develops around the areas of growth. If the color of the agar around the growth is unchanged, the test is negative (i.e., the organism is not able to produce DNase).
Objective: To determine the ability of an organism to ferment (degrade) a specific carbohydrate in a basal medium producing acid or acid with visible gas. The acid would change the color of the medium in a positive test. The following carbohydrate semi-solid media tubes are available at our lab:
1. Using a sterile needle, stab the tube within 1/4 inch of the bottom with medium inoculation.
2. Incubate for at least 48 hrs. Bacteria that are known to be slow growers should be given up to 96 hours.
Positive: Any yellow color (not orange). It does not necessarily have to be the whole tube. A positive result is referred to as ("+") or (A) or (Acid), as fermentation forms acidic products.
Negative: A red, pink or orange color - no yellow at all.
-- Gas production
Positive: Significant bubbling in semisolid medium (one small bubble is generally negative, caused by the stab). Gas may also cause the medium to get separated from tube. Record as (G) for positive gas production.
Negative: No gas bubbles except those produced by stabbing.
Objective: To determine if the organism is capable of breaking down starch into maltose through the activity of the extra-cellular α-amylase enzyme.
1. Use a sterile swab or a sterile loop to pick a few colonies from your pure culture plate. Streak a starch plate in the form of a line across the width of the plate. Several cultures can be tested on a single agar plate, each represented by a line or the plate may be divided into four quadrants (pie plate) for this purpose.
2. Incubate plate at 37 °C for 48 hours.
3. Add 2-3 drops of 10% iodine solution directly onto the edge of colonies. Wait 10-15 minutes and record the results.
-- Positive test ("+"): The medium will turn dark. However, areas surrounding isolated colonies where starch has been hydrolyzed by amylase will appear clear.
-- Negative test ("-"): The medium will be colored dark, right up to the edge of isolated colonies.