Overview of Bacterial Identification Methods and Strategies

Differentiate microorganisms based on the ability to use acetamide as the sole source of carbon.

Bacteria capable of growth on this medium produce the enzyme acylamidase, which deaminates acetamide to release ammonia. The production of ammonia results in an alkaline pH, causing the medium to change color from green to royal blue.

Media: NaCl (5 g), NH4H2PO4 (1 g), K2HPO4 (1 g), agar (15 g), bromthymol blue indicator (0.8 g), per 1000 mL, acetamide (10 g), pH 6.8.

1. Inoculate acetamide slant with a needle using growth from an 18- to 24-hour culture. Do not inoculate from a broth culture, because the growth will be too heavy.

2. Incubate aerobically at 35°-37°C for up to 4 days. If equivocal, the slant may be reincubated for 2 additional days.

Positive: Deamination of the acetamide, resulting in a blue color (Figure 13-3, A).

Figure 13-3 Acetamide utilization. A, Positive. B, Negative.

Negative: No color change (Figure 13-3, B).

Growth without a color change may indicate a positive test result. If further incubation results in no color change, repeat test with less inoculum.

Positive: Pseudomonas aeruginosa (ATCC 27853)—growth; blue color

Negative: Escherichia coli (ATCC 25922)—no growth; green color

Procedure 13-2 Acetate Utilization

Differentiate organisms based on ability to use acetate as the sole source of carbon. Generally used to differentiate Shigella sp. from Escherichia coli.

This test is used to differentiate an organism capable of using acetate as the sole source of carbon. Organisms capable of using sodium acetate grow on the medium, resulting in an alkaline pH, turning the indicator from green to blue.

Media: NaC2H3O2 (2 g); MgSO4 (0.1 g); NaCl (5 g); NH4H2PO4 (1 g); agar (20 g); bromthymol blue indicator (0.8 g), per 1000 mL, pH 6.7.

1. With a straight inoculating needle, inoculate acetate slant lightly from an 18- to 24-hour culture. Do not inoculate from a broth culture, because the growth will be too heavy.

Positive: Medium becomes alkalinized (blue) as a result of the growth and use of acetate (Figure 13-4, A).

Figure 13-4 Acetate utilization. A, Positive. B, Negative.

Negative: No growth or growth with no indicator change to blue (Figure 13-4, B).

Some strains of E. coli may use acetate at a very slow rate or not at all, resulting in a false negative reaction in the identification process.

Positive: Escherichia coli (ATCC 25922)—growth; blue

Negative: Shigella sonnei (ATCC 25931)—small amount of growth; green

Procedure 13-3 Bacitracin Susceptibility

This test is used for presumptive identification and differentiation of beta-hemolytic group A streptococci (Streptococcus pyogenes–susceptible) from other beta-hemolytic streptococci. It is also used to distinguish staphylococci species (resistant) from micrococci (susceptible).

The antibiotic bacitracin inhibits the synthesis of bacterial cell walls. A disk (TaxoA) impregnated with a small amount of bacitracin (0.04 units) is placed on an agar plate, allowing the antibiotic to diffuse into the medium and inhibit the growth of susceptible organisms. After incubation, the inoculated plates are examined for zones of inhibition surrounding the disks.

1. Using an inoculating loop, streak two or three suspect colonies of a pure culture onto a blood agar plate.

2. Using heated forceps, place a bacitracin disk in the first quadrant (area of heaviest growth). Gently tap the disk to ensure adequate contact with the agar surface.

3. Incubate the plate for 18 to 24 hours at 35°-37°C in ambient air for staphylococci and in 5% to 10% carbon dioxide (CO₂) for streptococci differentiation.

4. Look for a zone of inhibition around the disk.

Positive: Any zone of inhibition greater than 10 mm; susceptible (Figure 13-5, A).

Figure 13-5 Bacitracin (A disk) Susceptibility. Any zone of inhibition is positive *(Streptococcus pyogenes);* growth up to the disk is negative *(Streptococcus agalactiae).*

Negative: No zone of inhibition; resistant (Figure 13-5, B).

Performance depends on the integrity of the disk. Proper storage and expiration dates should be maintained.

Positive: Streptococcus pyogenes (ATCC19615)—susceptible

Micrococcus luteus (ATCC10240)—susceptible

Negative: Streptococcus agalactiae (ATCC27956)—resistant

Staphylococcus aureus (ATCC25923)—resistant

Procedure 13-4 Bile Esculin Test

This test is used for the presumptive identification of enterococci and organisms in the Streptococcus bovis group. The test differentiates enterococci and group D streptococci from non–group D viridans streptococci.

Gram-positive bacteria other than some streptococci and enterococci are inhibited by the bile salts in this medium. Organisms capable of growth in the presence of 4% bile and able to hydrolyze esculin to esculetin. Esculetin reacts with Fe3+ and forms a dark brown to black precipitate.

Media: Beef extract (11 g), enzymatic digest of gelatin (34.5 g), esculin (1 g), ox bile (2 g), ferric ammonium citrate (0.5 g), agar (15 g), per 1000 mL, pH 6.6.

1. Inoculate one to two colonies from an 18- to 24-hour culture onto the surface of the slant.

2. Incubate at 35°-37°C in ambient air for 48 hours.

Positive: Growth and blackening of the agar slant (Figure 13-6, A)

Figure 13-6 Bile esculin agar. A, Positive. B, Negative.

Negative: Growth and no blackening of medium (Figure 13-6, B)

No growth (not shown)

As a result of nutritional requirements, some organisms may grow poorly or not at all on this medium.

Positive: Enterococcus faecalis (ATCC19433)—growth; black precipitate

Negative: Escherichia coli (ATCC25922)—growth; no color change

Streptococcus pyogenes (ATCC19615)—no growth; no color change

Procedure 13-5 Bile Solubility Test

This test differentiates Streptococcus pneumoniae (positive–soluble) from alpha-hemolytic streptococci (negative–insoluble).

Bile or a solution of a bile salt (e.g., sodium desoxycholate) rapidly lyses pneumococcal colonies. Lysis depends on the presence of an intracellular autolytic enzyme, amidase. Bile salts lower the surface tension between the bacterial cell membrane and the medium, thus accelerating the organism’s natural autolytic process.

1. After 12 to 24 hours of incubation on 5% sheep blood agar, place 1 to 2 drops of 10% sodium desoxycholate on a well-isolated colony.

Note: A tube test is performed with 2% sodium desoxycholate.

2. Gently wash liquid over the colony without dislodging the colony from the agar.

3. Incubate the plate at 35°-37°C in ambient air for 30 minutes.

4. Examine for lysis of colony.

Positive: Colony disintegrates; an imprint of the lysed colony may remain in the zone (Figure 13-7, A).

Figure 13-7 Bile solubility (desoxycholate) test. A, Colony lysed. B, Intact colony.

Negative: Intact colonies (Figure 13-7, B).

Enzyme activity may be reduced in old cultures. Therefore, negative results with colonies resembling S. pneumoniae should be further tested for identification with alternate methods.

Positive: Streptococcus pneumoniae (ATCC49619)—bile soluble

Negative: Enterococcus faecalis (ATCC29212)—bile insoluble

Procedure 13-6 Butyrate Disk

This is a rapid test to detect the enzyme butyrate esterase, to aid identification of Moraxella (Branhamella) catarrhalis.

Organisms capable of producing butyrate esterase hydrolyze bromochlorindolyl butyrate. Hydrolysis of the substrate in the presence of butyrate esterase releases indoxyl, which in the presence of oxygen spontaneously forms indigo, a blue to blue-violet color.

1. Remove a disk from the vial and place on a glass microscope slide.

2. Add 1 drop of reagent-grade water. This should leave a slight excess of water on the disk.

3. Using a wooden applicator stick, rub a small amount of several colonies from an 18- to 24-hour pure culture onto the disk.

4. Incubate at room temperature for up to 5 minutes.

Positive: Development of a blue color during the 5-minute incubation period (Figure 13-8, A).

Figure 13-8 Butyrate disk. A, Positive. B, Negative.

Negative: No color change (Figure 13-8, B).

Incubation longer than 5 minutes may result in a false-positive reaction.

False-negative reactions may occur if the inoculum is too small. If the organism is negative, repeat with a larger inoculum and follow-up with additional methods.

Positive: Moraxella catarrhalis (ATCC25240)—formation of blue color

Negative: Neisseria gonorrhoeae (ATCC43069)—no color change

Procedure 13-7 CAMP Test

The Christie, Atkins, and Munch-Peterson (CAMP) test is used to differentiate group B streptococci (Streptococcus agalactiae–positive) from other streptococcal species. Listeria monocytogenes also produces a positive CAMP reaction.

Certain organisms (including group B streptococci) produce a diffusible extracellular hemolytic protein (CAMP factor) that acts synergistically with the beta-lysin of Staphylococcus aureus to cause enhanced lysis of red blood cells. The group B streptococci are streaked perpendicular to a streak of S. aureus on sheep blood agar. A positive reaction appears as an arrowhead zone of hemolysis adjacent to the place where the two streak lines come into proximity.

1. Streak a beta-lysin–producing strain of S. aureus down the center of a sheep blood agar plate.

2. Streak test organisms across the plate perpendicular to the S. aureus streak within 2 mm. (Multiple organisms can be tested on a single plate).

3. Incubate overnight at 35°-37°C in ambient air.

Positive: Enhanced hemolysis is indicated by an arrowhead-shaped zone of beta-hemolysis at the juncture of the two organisms (Figure 13-9, A).

Figure 13-9 CAMP test. A, Positive; arrowhead zone of beta-hemolysis *(at arrow)*, typical of group B streptococci. B, Negative; no enhancement of hemolysis.

Negative: No enhancement of hemolysis (Figure 13-9, B).

A small percentage of group A streptococci may have a positive CAMP reaction. The test should be limited to colonies with the characteristic group B streptococci morphology and narrow zone beta-hemolysis on sheep blood agar.

Positive: Streptococcus agalactiae (ATCC13813)—enhanced arrowhead hemolysis

Negative: Streptococcus pyogenes (ATCC19615)—beta-hemolysis without enhanced arrowhead formation

Procedure 13-8 Catalase Test

This test differentiates catalase-positive micrococcal and staphylococcal species from catalase-negative streptococcal species.

Aerobic and facultative anaerobic organisms produce two toxins during normal metabolism, hydrogen peroxide (H₂O₂) and superoxide radical (O2−). These bacteria have two enzymes that detoxify the products of normal metabolism. One of these enzymes, catalase, is capable of converting hydrogen peroxide to water and oxygen. The presence of the enzyme in a bacterial isolate is evidenced when a small inoculum introduced into hydrogen peroxide (30% for the slide test) causes rapid elaboration of oxygen bubbles. The lack of catalase is evident by a lack of or weak bubble production.

1. Use a loop or sterile wooden stick to transfer a small amount of colony growth to the surface of a clean, dry glass slide.

2. Place a drop of 30% hydrogen peroxide (H₂O₂) onto the medium.

3. Observe for the evolution of oxygen bubbles (Figure 13-10).

Figure 13-10 Catalase test. A, Positive. B, Negative.

Positive: Copious bubbles are produced (Figure 13-10, A).

Negative: No or few bubbles are produced (Figure 13-10, B).

Some organisms (enterococci) produce a peroxidase that slowly catalyzes the breakdown of H₂O₂, and the test may appear weakly positive. This reaction is not a truly positive test.

False positives may occur if the sample is contaminated with blood agar.

Positive: Staphylococcus aureus (ATCC25923)

Negative: Streptococcus pyogenes (ATCC19615)

Procedure 13-9 Cetrimide Agar

This test is primarily used to isolate and purify Pseudomonas aeruginosa from contaminated specimens.

The test is used to determine the ability of an organism to grow in the presence of cetrimide, a toxic substance that inhibits the growth of many bacteria by causing the release of nitrogen and phosphorous, which slows or kills the organism. P. aeruginosa is resistant to cetrimide.

Media: Enzymatic digest of gelatin (20 g), MgCl2 (1.4 g), K2SO4 (10 g), cetrimide (cetyltrimethylammonium bromide) (0.3 g), agar (13.6), pH 7.2.

1. Inoculate a cetrimide agar slant with 1 drop of an 18- to 24-hour brain-heart infusion broth culture.

3. Examine the slant for bacterial growth.

Positive: Growth, variation in color of colonies (Figure 13-11, A).

Figure 13-11 Cetrimide agar. A, Positive. B, Negative.

Negative: No growth (Figure 13-11, B).

Some enteric organisms will grow and exhibit a weak yellow color in the media. This color change is distinguishable from the production of fluorescein.

Additional testing is necessary to confirm a diagnosis of P. aeruginosa.

Positive: Pseudomonas aeruginosa (ATCC27853)—growth and color change; yellow-green to blue-green colonies

Negative: Escherichia coli (ATCC25922)—no growth and no color change

Procedure 13-10 Citrate Utilization

The purpose of this test is to identify organisms capable of using sodium citrate as the sole carbon source and inorganic ammonium salts as the sole nitrogen source. The test is part of a series referred to as IMViC (indole, methyl red, Voges-Proskauer, and citrate), which is used to differentiate Enterobacteriaceae from other gram-negative rods.

Bacteria that can grow on this medium produce an enzyme, citrate-permease, capable of converting citrate to pyruvate. Pyruvate can then enter the organism’s metabolic cycle for the production of energy. Bacteria capable of growth in this medium use the citrate and convert ammonium phosphate to ammonia and ammonium hydroxide, creating an alkaline pH. The pH change turns the bromthymol blue indicator from green to blue.

Media: NH₄H₂PO₄ (1 g), K₂HPO₄ (1 g), NaCl (5 g), sodium citrate (2 g), MgSO₄ (0.2 g), agar (15 g), bromthymol blue (0.08 g), per 1000 mL, pH 6.9.

1. Inoculate Simmons citrate agar lightly on the slant by touching the tip of a needle to a colony that is 18 to 24 hours old. Do not inoculate from a broth culture, because the inoculum will be too heavy.

3. Observe for growth and the development of blue color, denoting alkalinization.

Positive: Growth on the medium, with or without a change in the color of the indicator. Growth typically results in the bromthymol blue indicator turning from green to blue (Figure 13-12, A).

Figure 13-12 Citrate utilization. A, Positive. B, Negative.

Negative: Absence of growth (Figure 13-12, B).

Some organisms are capable of growth on citrate and do not produce a color change. Growth is considered a positive citrate utilization test, even in the absence of a color change.

Positive: Enterobacter aerogenes (ATCC13048)—growth, blue color

Negative: Escherichia coli (ATCC25922)—little to no growth, no color change

Procedure 13-11 Coagulase Test

The test is used to differentiate Staphylococcus aureus (positive) from coagulase-negative staphylococci (negative).

S. aureus produces two forms of coagulase, bound and free. Bound coagulase, or “clumping factor,” is bound to the bacterial cell wall and reacts directly with fibrinogen. This results in precipitation of fibrinogen on the staphylococcal cell, causing the cells to clump when a bacterial suspension is mixed with plasma. The presence of bound coagulase correlates with free coagulase, an extracellular protein enzyme that causes the formation of a clot when S. aureus colonies are incubated with plasma. The clotting mechanism involves activation of a plasma coagulase-reacting factor (CRF), which is a modified or derived thrombin molecule, to form a coagulase-CRF complex. This complex in turn reacts with fibrinogen to produce the fibrin clot.

A Slide Test (Detection of ?)

1. Place a drop of coagulase plasma (preferably rabbit plasma with ethylenediaminetetraacetic acid [EDTA]) on a clean, dry, glass slide.

2. Place a drop of distilled water or saline next to the drop of plasma as a control.

3. With a loop, straight wire, or wooden stick, emulsify a portion of the isolated colony being tested in each drop, inoculating the water or saline first. Try to create a smooth suspension.

4. Mix well with a wooden applicator stick.

5. Rock the slide gently for 5 to 10 seconds.

Positive: Macroscopic clumping in 10 seconds or less in coagulated plasma drop and no clumping in saline or water drop (Figure 13-13, A, left side).

Figure 13-13 Coagulase test. A, Slide coagulase test for clumping factor. Left side is positive; right side is negative. B, Tube coagulase test for free coagulase. Tube on the left is positive, exhibiting clot. Tube on the right is negative.

Negative: No clumping in either drop.

Note: All negative slide tests must be confirmed using the tube test (Figure 13-13, B, right side).

B Tube Test

1. Emulsify several colonies in 0.5 mL of rabbit plasma (with EDTA) to give a milky suspension.

2. Incubate tube at 35°-37°C in ambient air for 4 hours.

3. Check for clot formation.

Positive: Clot of any size (Figure 13-13, A, left side).

Negative: No clot (Figure 13-13, B, right side).

Equivocal: Clumping in both drops indicates that the organism autoagglutinates and is unsuitable for the slide coagulase test.

Slide Test

Tube Test

1. Test results can be positive at 4 hours and then revert to negative after 24 hours.

2. If negative at 4 hours, incubate at room temperature overnight and check again for clot formation.

Positive: Staphylococcus aureus (ATCC25923)

Negative: Staphylococcus epidermidis (ATC12228)

Procedure 13-12 Decarboxylase Tests (Moeller’s Method)

This test is used to differentiate decarboxylase-producing Enterobacteriaceae from other gram-negative rods.

This test measures the enzymatic ability (decarboxylase) of an organism to decarboxylate (or hydrolyze) an amino acid to form an amine. Decarboxylation, or hydrolysis, of the amino acid results in an alkaline pH and a color change from orange to purple.

Media: Peptic digest of animal tissue (5 g), beef extract (5 g), bromcresol purple (0.1 g), cresol red (0.005 g), dextrose (0.5 g), pyridoxal (0.005 g), amino acid (10 g), pH 6.0.

A Glucose-Nonfermenting Organisms

1. Prepare a suspension (≥McFarland No. 5 turbidity standard) in brain-heart infusion broth from an overnight culture (18 to 24 hours old) growing on 5% sheep blood agar.

2. Inoculate each of the three decarboxylase broths (arginine, lysine, and ornithine) and the control broth (no amino acid) with 4 drops of broth.

3. Add a 4-mm layer of sterile mineral oil to each tube.

4. Incubate the cultures at 35°-37°C in ambient air. Examine the tubes at 24, 48, 72, and 96 hours.

B Glucose-Fermenting Organisms

1. Inoculate tubes with 1 drop of an 18- to 24-hour brain-heart infusion broth culture.

2. Add a 4-mm layer of sterile mineral oil to each tube.

3. Incubate the cultures for 4 days at 35°-37°C in ambient air. Examine the tubes at 24, 48, 72, and 96 hours.

Positive: Alkaline (purple) color change compared with the control tube (Figure 13-14, A).

Figure 13-14 Decarboxylase tests (Moeller’s method). A, Positive. B, Negative. C, Uninoculated tube.

Negative: No color change or acid (yellow) color in test and control tube. Growth in the control tube.

The fermentation of dextrose in the medium causes the acid color change. However, it would not mask the alkaline color change brought about by a positive decarboxylation reaction (Figure 13-14, B). An uninoculated tube is shown in Figure 13-14, C.

Lysine—Klebsiella pneumoniae (ATCC33495)

Positive:

Ornithine—Enterobacter aerogenes (ATCC13048)

Arginine—Enterobacter cloacae (ATCC13047)

Base—

Negative:

Lysine—Enterobacter cloacae (ATCC13047)

Ornithine—Klebsiella pneumoniae (ATCC13883)

Arginine—Klebsiella pneumoniae (ATCC33495)

Base—Klebsiella pneumoniae (ATC33495)

Procedure 13-13 DNA Hydrolysis (DNase Test Agar)

This test is used to differentiate organisms based on the production of deoxyribonuclease. It is used to distinguish Serratia sp. (positive) from Enterobacter sp., Staphylococcus aureus (positive) from other species, and Moraxella catarrhalis (positive) from Neisseria sp.

The test is used to determine the ability of an organism to hydrolyze DNA. The medium is pale green because of the DNA–methyl green complex. If the organism growing on the medium hydrolyses DNA, the green color fades and the colony is surrounded by a colorless zone.

Media: Pancreatic digest of casein (10 g), yeast extract (10 g), deoxyribonucleic acid (2 g), NaCl (5 g), agar (15 g), methyl green (0.5 g), pH 7.5.

1. Inoculate the DNase agar with the organism to be tested and streak for isolation.

2. Incubate aerobically at 35°-37°C for 13 to 24 hours.

Positive: When DNA is hydrolyzed, methyl green is released and combines with highly polymerized DNA at a pH of 7.5, turning the medium colorless around the test organism (Figure 13-15, A and B).

Figure 13-15 DNA hydrolysis. A, Positive, *Staphylococcus aureus.* B, Positive, *Serratia marcescens.* C, Negative.

Negative: If no degradation of DNA occurs, the medium remains green (Figure 13-15, C).

Agar must be inoculated with a suspension of a young broth culture (4 hours old) or an 18- to 24-hour colony in 1-2 mL of saline.

Positive: Staphylococcus aureus (ATCC25923)

Procedure 13-14 Esculin Hydrolysis

This test is used for the presumptive identification and differentiation of Enterobacteriaceae.

This test is used to determine whether an organism is able to hydrolyze the glycoside esculin. Esculin is hydrolyzed to esculetin, which reacts with Fe3+ and forms a dark brown to black precipitate.

Media: NaCl (8 g), K₂HPO₄ (0.4 g), KH₂PO₄ (0.1 g), esculin (5 g), ferric ammonium citrate (0.5 g), agar (15 g), per 1000 mL, pH 7.0.

1. Inoculate the medium with 1 drop of a 24-hour broth culture.

3. Examine the slants for blackening and, under the ultraviolet rays of a Wood’s lamp, for esculin hydrolysis.

Negative: No blackening and no loss of fluorescence under the Wood’s lamp, or slight blackening with no loss of fluorescence under the Wood’s lamp. An uninoculated tube is shown in Figure 13-16, B.

Positive: Blackened medium (Figure 13-16, A), which would also show a loss of fluorescence under the Wood’s lamp.

Figure 13-16 Esculin hydrolysis. A, Positive, blackening of slant. B, Uninoculated tube.

This medium is a nonselective agar. The bile esculin hydrolysis test presented in Procedure 13-4 is a selective differential method.

Positive: Enterococcus faecalis (ATCC29212)

Procedure 13-15 Fermentation Media

Fermentation media are used to differentiate organisms based on their ability to ferment carbohydrates incorporated into the basal medium. Andrade’s formula is used to differentiate enteric bacteria from coryneforms, and bromocresol purple is used to distinguish enterococci from streptococci.

Carbohydrate fermentation is the process microorganisms use to produce energy. Most microorganisms convert glucose to pyruvate during glycolysis; however, some organisms use alternate pathways. A fermentation medium consists of a basal medium containing a single carbohydrate (glucose, lactose, or sucrose) for fermentation. However, the medium may contain various color indicators, such as Andrade’s indicator, bromocresol, or others. In addition to a color indicator to detect the production of acid from fermentation, a Durham tube is placed in each tube to capture gas produced by metabolism.

Basal media: Pancreatic digest of casein (10 g), beef extract (3 g), NaCl (5 g), carbohydrate (10 g), specific indicator (Andrade’s indicator [10 mL, pH 7.4] or bromocresol purple [0.02 g, pH 6.8]).

A Peptone Medium with Andrade’s Indicator (for Enterics and Coryneforms)

1. Inoculate each tube with 1 drop of an 18- to 24-hour brain-heart infusion broth culture.

2. Incubate at 35°-37°C for up to 7 days in ambient air.

Note: Tubes are held only 4 days for organisms belonging to the Enterobacteriaceae family.

3. Examine the tubes for acid (indicated by a pink color) and gas production.

4. Tubes must show growth for the test to be valid. If no growth in the fermentation tubes or control is seen after 24 hours of incubation, add 1 to 2 drops of sterile rabbit serum per 5 mL of fermentation broth to each tube.

Negative: Growth, but no change in color. Medium remains clear to straw colored (Figure 13-17, A, right).

Positive: Indicator change to pink with or without gas formation in Durham tube (Figure 13-17, A, left and middle).

Figure 13-17 Fermentation media. A, Peptone medium with Andrade’s indicator. The tube on the left ferments glucose with the production of gas (visible as a bubble *[arrow]* in the inverted [Durham] tube); the tube in the middle ferments glucose with no gas production; and the tube on the right does not ferment glucose. B, Heart infusion broth with bromocresol purple indicator. The tube on the left is positive; the tube on the right is negative.

B Broth (Brain Heart Infusion Broth May Be Substituted) with Bromcresol Purple Indicator (for Streptococci and Enterococci)

1. Inoculate each tube with 2 drops of an 18- to 24-hour brain-heart infusion broth culture.

2. Incubate 4 days at 35°-37°C in ambient air.

3. Observe daily for a change of the bromcresol purple indicator from purple to yellow (acid).

Positive: Indicator change to yellow (Figure 13-17, B, left).

Negative: Growth, but no change in color. Medium remains purple (Figure 13-17, B, right).

Readings after 24 hours may not be reliable if no acid is produced. No color change or a result indicating alkalinity may occur if the organism deaminates the peptone, masking the evidence of carbohydrate fermentation.

Note: Appropriate organisms depend on which carbohydrate has been added to the basal medium. An example is given for each type of medium.

A Peptone Medium with Andrade’s Indicator

Dextrose:

Positive, with gas: Escherichia coli (ATCC25922)

Positive, no gas: Shigella flexneri (ATCC12022)

B Brain-Heart Infusion Broth with Bromocresol Purple Indicator

Dextrose:

Positive, with gas: Escherichia coli (ATCC25922)

Negative, no gas: Moraxella osloensis (ATCC10973)

Procedure 13-16 Flagella Stain (Wet Mount Technique)

This technique is used to visualize the presence and arrangement of flagella for the presumptive identification of motile bacterial species.

Flagella are too thin to be visualized using a bright field microscope with ordinary stains, such as the Gram stain, or a simple stain. A wet mount technique is used for staining bacterial flagella, and it is simple and useful when the number and arrangement of flagella are critical to the identification of species of motile bacteria. The staining procedures require the use of a mordant so that the stain adheres in layers to the flagella, allowing visualization.

1. Grow the organism to be stained at room temperature on blood agar for 16 to 24 hours.

2. Add a small drop of water to a microscope slide.

3. Dip a sterile inoculating loop into sterile water.

4. Touch the loopful of water to the colony margin briefly (this allows motile cells to swim into the droplet of water).

5. Touch the loopful of motile cells to the drop of water on the slide. Note: Agitating the loop in the droplet of water on the slide causes the flagella to shear off the cell.

6. Cover the faintly turbid drop of water on the slide with a cover slip. A proper wet mount has barely enough liquid to fill the space under a cover slip. Small air spaces around the edge are preferable.

7. Examine the slide immediately under 40× to 50× for motile cells. If motile cells are not seen, do not proceed with the stain.

8. If motile cells are seen, leave the slide at room temperature for 5 to 10 minutes. This allows the bacterial cells time to adhere either to the glass slide or to the cover slip.

9. Gently apply 2 drops of RYU flagella stain (Remel, Lenexa, Kansas) to the edge of the cover slip. The stain will flow by capillary action and mix with the cell suspension. Small air pockets around the edge of the wet mount are useful in aiding the capillary action.

10. After 5 to 10 minutes at room temperature, examine the cells for flagella.

11. Cells with flagella may be observed at 100× (oil) in the zone of optimum stain concentration, about halfway from the edge of the cover slip to the center of the mount.

12. Focusing the microscope on the cells attached to the cover slip rather than on the cells attached to the slide facilitates visualization of the flagella. The precipitate from the stain is primarily on the slide rather than the cover slip.

Observe the slide and note the following:

1. Presence or absence of flagella

2. Number of flagella per cell

3. Location of flagella per cell

a. Peritrichous (Figure 13-18, A)

Figure 13-18 Flagella stain (wet mount technique). A, *Alcaligenes* spp., peritrichous flagella *(arrows).* B, *Pseudomonas aeruginosa,* polar flagella *(arrows).*

b. Lophotrichous

c. Polar (Figure 13-18, B)

4. Amplitude of wavelength

a. Short

b. Long

5. Whether or not “tufted”

Even with a specific stain, visualization of flagella requires an experienced laboratory scientist and is not considered an entry-level technique.

Peritrichous: Escherichia coli

Polar: Pseudomonas aeruginosa

Negative: Klebsiella pneumoniae

Procedure 13-17 Gelatin Hydrolysis

The production of gelatinases capable of hydrolyzing gelatin is used as a presumptive test for the identification of various organisms, including Staphylococcus sp., Enterobacteriaceae, and some gram-positive bacilli.

This test is used to determine the ability of an organism to produce extracellular proteolytic enzymes (gelatinases) that liquefy gelatin, a component of vertebrate connective tissue.

Nutrient gelatin medium differs from traditional microbiology media in that the solidifying agent (agar) is replaced with gelatin. When an organism produces gelatinase, the enzyme liquefies the growth medium.

Media: Enzymatic digest of gelatin (5 g), beef extract (3 g), gelatin (120 g), per 1000 mL, pH 6.8.

1. Inoculate the gelatin deep with 4 to 5 drops of a 24-hour broth culture.

2. Incubate at 35°-37°C in ambient air for up to 14 days. Note: Incubate the medium at 25°C if the organism grows better at 25°C than at 35°C.

3. Alternatively, inoculate the gelatin deep from a 24-hour-old colony by stabbing four or five times, 0.5 inch into the medium.

4. Remove the gelatin tube daily from the incubator and place at 4°C to check for liquefaction. Do not invert or tip the tube, because sometimes the only discernible liquefaction occurs at the top of the deep where inoculation occurred.

5. Refrigerate an uninoculated control along with the inoculated tube. Liquefaction is determined only after the control has hardened (gelled).

Positive: Partial or total liquefaction of the inoculated tube (the control tube must be completely solidified) at 4°C within 14 days (Figure 13-19, A).

Figure 13-19 Gelatin hydrolysis. A, Positive; note liquefaction at top of tube. B, Uninoculated tube.

Negative: Complete solidification of the tube at 4°C (Figure 13-19, B).

Some organisms may grow poorly or not at all in this medium

Gelatin is liquid above 20°C; therefore determination of results must be completed following refrigeration.

Positive: Bacillus subtilis (ATCC9372)

Uninoculated control tube: medium becomes solid after refrigeration.

Procedure 13-18 Growth at 42°C

This test is used to differentiate a pyocyanogenic pseudomonads from other Pseudomonas sp.

The test is used to determine the ability of an organism to grow at 42°C. Several Pseudomonas species have been isolated in the clinical laboratory that are capable of growth at elevated temperatures.

1. Inoculate two tubes of trypticase soy agar (TSA) with a light inoculum by lightly touching a needle to the top of a single 13- to 24-hour-old colony and streaking the slant.

2. Immediately incubate one tube at 35°C and one at 42°C.

3. Record the presence of growth on each slant after 18 to 24 hours.

Positive: Good growth at both 35°and 42°C (Figure 13-20, A).

Figure 13-20 Growth at 42°C. A, Positive; good growth. B, Negative; no growth.

Negative: No growth at 42°C (Figure 13-20, B), but good growth at 35°C.

Positive: Pseudomonas aeruginosa (ATCC10145)

Negative: Pseudomonas fluorescens (ATCC13525)

Procedure 13-19 Hippurate Hydrolysis

Production of the enzyme hippuricase is used for the presumptive identification of a variety of microorganisms.

The end products of hydrolysis of hippuric acid by hippuricase include glycine and benzoic acid. Glycine is deaminated by the oxidizing agent ninhydrin, which is reduced during the process. The end products of the ninhydrin oxidation react to form a purple-colored product. The test medium must contain only hippurate, because ninhydrin might react with any free amino acids present in growth media or other broths.

1. Add 0.1 mL of sterile water to a 12 ×75 mm plastic test tube.

2. Make a heavy suspension of the organism to be tested.

3. Using heated forceps, place a rapid hippurate disk in the mixture.

4. Cap and incubate the tube for 2 hours at 35°C; use of a water bath is preferred.

5. Add 0.2 mL ninhydrin reagent and reincubate for an additional 15 to 30 minutes. Observe the solution for the development of a deep purple color.

Positive: Deep purple color (Figure 13-21, A).

Figure 13-21 Hippurate hydrolysis. A, Positive. B, Negative.

Negative: Colorless or slightly yellow pink color (Figure 13-21, B).

A false-positive result may occur if incubation with ninhydrin exceeds 30 minutes.

Positive: Streptococcus agalactiae (ATCC12386)

Negative: Streptococcus pyogenes (ATCC19615)

Procedure 13-20 Indole Production

This test is used to identify organisms that produce the enzyme tryptophanase.

The test is used to determine an organism’s ability to hydrolyze tryptophan to form the compound indole. Tryptophan is present in casein and animal protein. Bacteria with tryptophanase are capable of hydrolyzing tryptophan to pyruvate, ammonia, and indole. Kovac’s reagent (dimethylamine-benzaldehyde and hydrochloride), when added to the broth culture, reacts with the indole, producing a red color. An alternative method uses Ehrlich’s reagent. Ehrlich’s reagent has the same chemicals as the Kovac preparation, but it also contains absolute ethyl alcohol, making it flammable. Ehrlich’s reagent is more sensitive for detecting small amounts of indole. (The spot indole test is described in Procedure 13-39.)

Media: Casein peptone (10 g), NaCl (5 g), tryptophan (10 g), per 1000 mL.

1. Inoculate tryptophane broth with 1 drop from a 24-hour brain-heart infusion broth culture.

A Enterobacteriaceae

2. Incubate at 35°-37°C in ambient air for 48 hours.

3. Add 0.5 mL of Kovac’s reagent to the broth culture.

B Other Gram-Negative Bacilli

1. Inoculate tryptophane broth with 1 drop of a 24-hour broth culture.

2. Incubate at 35°-37°C in ambient air for 48 hours.

3. Add 1 mL of xylene to the culture.

4. Shake the mixture vigorously to extract the indole and allow it to stand until the xylene forms a layer on top of the aqueous phase.

5. Add 0.5 mL of Ehrlich’s reagent down the side of the tube.

Positive: Pink- to wine-colored ring after addition of appropriate reagent (Figure 13-22, A).

Figure 13-22 Indole production. A, Positive. B, Negative.

Negative: No color change after addition of the appropriate reagent (Figure 13-22, B).

Ehrlich’s method may also be used to differentiate organisms under anaerobic conditions.

A Kovac’s Method

Negative: Klebsiella pneumoniae (ATCC13883)

B Ehrlich’s Method

Positive: Haemophilus influenzae (ATCC49766)

Negative: Haemophilus parainfluenzae (ATCC76901)

C Ehrlich’s Method (Anaerobic)

Positive: Porphyromonas asaccharolytica (ATCC25260)

Negative: Bacteroides fragilis (ATCC25285)

Procedure 13-21 Leucine Aminopeptidase (LAP) Test

The LAP test is used for the presumptive identification of catalase-negative gram-positive cocci.

The LAP disk is a rapid test for the detection of the enzyme leucine aminopeptidase. Leucine-beta-naphthylamide–impregnated disks serve as a substrate for the detection of leucine aminopeptidase. After hydrolysis of the substrate by the enzyme, the resulting beta-naphthylamine produces a red color upon addition of cinnamaldehyde reagent.

1. Before incubation, slightly dampen the LAP disk with reagent-grade water. Do not supersaturate the disk.

2. Using a wooden applicator stick, rub a small amount of several colonies of an 18- to 24-hour pure culture onto a small area of the LAP disk.

3. Incubate at room temperature for 5 minutes.

4. After this incubation period, add 1 drop of cinnamaldehyde reagent.

Negative: No color change or development of a slight yellow color (Figure 13-23, B).

Positive: Development of a red color within 1 minute after adding cinnamaldehyde reagent (Figure 13-23, A; swab test is depicted)

Figure 13-23 LAP test. A, Positive. B, Negative.

The test result depends on the integrity of the substrate-impregnated disk.

Positive: Enterococcus faecalis (ATCC29212)—red color

Negative: Aerococcus viridans (ATCC11563)—no color change

Procedure 13-22 Litmus Milk Medium

This test differentiates microorganisms based on various metabolic reactions in litmus milk, including fermentation, reduction, clot formation, digestion, and the formation of gas. Litmus milk is also used to grow lactic acid bacteria.

This test is used to determine an organism’s ability to metabolize litmus milk. Fermentation of lactose is demonstrated when the litmus turns pink as a result of acid production. If sufficient acid is produced, casein in the milk is coagulated, solidifying the milk. With some organisms, the curd shrinks and whey is formed at the surface. Some bacteria hydrolyze casein, causing the milk to become straw colored and resemble turbid serum. Additionally, some organisms reduce litmus, in which case the medium becomes colorless in the bottom of the tube.

Media: Powdered skim milk (100 g), litmus (0.5 g), sodium sulphite (0.5 g), per 1000 mL, pH 6.8.

1. Inoculate with 4 drops of a 24-hour broth culture.

2. Incubate at 35°-37°C in ambient air.

3. Observe daily for 7 days for alkaline reaction (litmus turns blue), acid reaction (litmus turns pink), indicator reduction, acid clot, rennet clot, and peptonization. Multiple changes can occur over the observation period.

4. Record all changes.

Fermentation: Clostridium perfringens (ATCC13124)—gas production

Acid: Lactobacillus acidophilus (ATCC11506)—clot formation

Peptonization: Pseudomonas aeruginosa (ATCC27853)—clearing

Appearance of Indicator (Litmus Dye)

Litmus media reactions are not specific and should be followed up with additional tests for definitive identification of microorganisms.

Appearance of Indicator (Litmus Dye)

Color	pH Change to …	Record
Pink, mauve (Figure 13-24, A)	Acid	Acid (A)
Blue (Figure 13-24, B)	Alkaline	Alkaline (K)
Purple (identical to uninoculated control) (Figure 13-24, C)	No change	No change
White (Figure 13-24, D)	Independent of pH change; result of reduction of indicator	Decolorized

Figure 13-24 Litmus milk. A, Acid reaction. B, Alkaline reaction. C, No change. D, Reduction of indicator. E, Clot. (Note separation of clear fluid from clot at *arrow*.) F, Peptonization.

Appearance of Milk

Consistency of Milk	Occurs When pH Is …	Record
Coagulation or clot (Figure 13-24, E)	Acid or alkaline	Clot
Dissolution of clot with clear, grayish, watery fluid and a shrunken, insoluble pink clot (Figure 13-24, F)	Acid	Digestion
Dissolution of clot with grayish, watery fluid and a clear, shrunken, insoluble blue clot	Alkaline	Peptonization

Procedure 13-23 Lysine Iron Agar (LIA)

This test is used to differentiate gram-negative bacilli based on decarboxylation or deamination of lysine and the formation of hydrogen sulfide (H₂S).

Lysine iron agar contains lysine, peptones, a small amount of glucose, ferric ammonium citrate, and sodium thiosulfate. The medium has an aerobic slant and an anaerobic butt. When glucose is fermented, the butt of the medium becomes acidic (yellow). If the organism produces lysine decarboxylase, cadaverine is formed. Cadaverine neutralizes the organic acids formed by glucose fermentation, and the butt of the medium reverts to the alkaline state (purple). If the decarboxylase is not produced, the butt remains acidic (yellow). If oxidative deamination of lysine occurs, a compound is formed that, in the presence of ferric ammonium citrate and a coenzyme, flavin mononucleotide, forms a burgundy color on the slant. If deamination does not occur, the LIA slant remains purple. Bromocresol purple, the pH indicator, is yellow at or below pH 5.2 and purple at or above pH 6.8.

Media: Enzymatic digest of gelatin (5 g), yeast extract (3 g), dextrose (1 g), L-lysine (10 g), ferric ammonium citrate (0.5 g), sodium thiosulfate (0.04 g), bromocresol purple (0.02 g), agar (13.5 g), per 1000 mL, pH 6.7.

2. Cap the tube tightly and incubate at 35°-37°C in ambient air for 18 to 24 hours.

1. With a straight inoculating needle, inoculate LIA (Figure 13-25, E) by twice stabbing through the center of the medium to the bottom of the tube and then streaking the slant.

Figure 13-25 Lysine iron agar. A, Alkaline slant/alkaline butt (K/K). B, Alkaline slant/alkaline butt, H₂S positive (K/K H₂S+). C, Alkaline slant/acid butt (K/A). D, Red slant/acid butt (R/A). E, Uninoculated tube.

Alkaline slant/alkaline butt (K/K)—lysine decarboxylation and no fermentation of glucose (Figure 13-25, A)

Alkaline slant/acid butt (K/A)—glucose fermentation (Figure 13-25, C)

Note: Patterns shown in Figure 13-25, A and C, can be accompanied by a black precipitate of ferrous sulfide (FeS), which indicates production of H₂S (Figure 13-25, B).

Red slant/acid butt (R/A)—lysine deamination and glucose fermentation (Figure 13-25, D)

Proteus sp. that produce hydrogen sulfide will not blacken the medium. Additional testing, such as triple sugar iron agar, should be used as a follow-up identification method.

Alkaline slant and butt: H₂S positive: Citrobacter freundii (ATCC8090)

Alkaline slant and butt: Escherichia coli (ATCC25922)

Alkaline slant and butt: H₂S positive: Salmonella typhimurium (ATCC14028)

Red slant, acid butt: Proteus mirabilis (ATCC12453)

Procedure 13-24 Methyl Red/Voges-Proskauer (Mrvp) Tests

The combination test methyl red (MR) and Voges-Proskauer (VP) differentiates members of the Enterobacteriaceae family.

This test is used to determine the ability of an organism to produce and maintain stable acid end products from glucose fermentation, to overcome the buffering capacity of the system, and to determine the ability of some organisms to produce neutral end products (e.g., 2,3-butanediol or acetoin) from glucose fermentation. The methyl red detects mixed acid fermentation that lowers the pH of the broth. The MR indicator is added after incubation. Methyl red is red at pH 4.4 and yellow at pH 6.2. A clear red is a positive result; yellow is a negative result; and various shades of orange are negative or inconclusive. The VP detects the organism’s ability to convert the acid products to acetoin and 2,3-butanediol. Organisms capable of using the VP pathway produce a smaller amount of acid during glucose fermentation and therefore do not produce a color change when the methyl red indicator is added. A secondary reagent is added, alpha-naphthol, followed by potassium hydroxide (KOH); a positive test result is indicated by a red color complex.

Media: Peptic digest of animal tissue (3.5 g), pancreatic digest of casein (3.5 g), dextrose (5 g), KPO₄ (5 g), per 1000 mL, pH 6.9.

1. Inoculate MRVP broth with 1 drop from a 24-hour brain-heart infusion broth culture.

2. Incubate at 35°-37°C for a minimum of 48 hours in ambient air. Tests should not be made with cultures incubated less than 48 hours, because the end products build up to detectable levels over time. If results are equivocal at 48 hours, repeat the tests with cultures incubated at 35°-37°C for 4 to 5 days in ambient air; in such instances, duplicate tests should be incubated at 25°C.

3. Split broth into aliquots for MR test and VP test.

A MR (Methyl Red) Test

1. Add 5 or 6 drops of methyl red reagent per 5 mL of broth.

2. Read reaction immediately.

Positive: Bright red color, indicative of mixed acid fermentation (Figure 13-26, A).

Figure 13-26 Methyl red/Voges-Proskauer (MRVP) tests. A, Positive methyl red. B, Negative methyl red. C, Positive Voges-Proskauer. D, Negative Voges-Proskauer.

Weakly positive: Red-orange color.

Negative: Yellow color (Figure 13-26, B).

B VP (Voges-Proskauer) Test (Barritt’s Method) for Gram-Negative Rods

1. Add 0.6 mL (6 drops) of solution A (alpha-naphthol) and 0.2 mL (2 drops) of solution B (KOH) to 1 mL of MRVP broth.

2. Shake well after addition of each reagent.

3. Observe for 5 minutes.

Positive: Red color, indicative of acetoin production (Figure 13-26, C).

Negative: Yellow color (Figure 13-26, D).

C VP (Voges-Proskauer) Test (Coblentz Method) for Streptococci

1. Use 24-hour growth from blood agar plate to heavily inoculate 2 mL of MRVP broth.

2. After 6 hours of incubation at 35°C in ambient air, add 1.2 mL (12 drops) of solution A (alpha-naphthol) and 0.4 mL (4 drops) solution B (40% KOH with creatine).

3. Shake the tube and incubate at room temperature for 30 minutes.

The MR test should not be read before 48 hours, because some organisms will not have produced enough products from the fermentation of glucose.

MR-negative organisms may also not have had sufficient time to convert those products and will appear MR positive.

MR-VP testing should be used in conjunction with other confirmatory tests to differentiate organisms among the Enterobacteriaceae.

Methyl red—Voges-Proskauer

MR positive/VP negative: Escherichia coli (ATCC25922)

MR negative:/VP positive: Enterobacter aerogenes (ATCC13048)

Procedure 13-25 Microdase Test (Modified Oxidase)

This test is used to differentiate gram-positive, catalase-positive cocci (micrococci from staphylococci).

The microdase test is a rapid method to differentiate Staphylococcus from Micrococcus spp. by detection of the enzyme oxidase. In the presence of atmospheric oxygen, the oxidase enzyme reacts with the oxidase reagent and cytochrome C to form the colored compound, indophenol.

1. Using a wooden applicator stick, rub a small amount of several colonies of an 18- to 24-hour pure culture grown on blood agar onto a small area of the microdase disk. Note: Do not rehydrate the disk before use.

2. Incubate at room temperature for 2 minutes.

Positive: Development of blue to purple-blue color (Figure 13-27, A).

Figure 13-27 Microdase test. A, Positive. B, Negative.

Negative: No color change (Figure 13-27, B).

Staphylococci should yield a negative color change, except for S. sciuri, S. lentus, and S. vitulus.

Positive: Micrococcus luteus (ATCC10240)

Negative: Staphylococcus aureus (ATCC25923)

Procedure 13-26 Motility Testing

These tests are used to determine whether an enteric organism is motile. An organism must have flagella to be motile.

The inoculum is stabbed into the center of a semisolid agar deep. Bacterial motility is evident by a diffuse zone of growth extending out from the line of inoculation. Some organisms grow throughout the entire medium, whereas others show small areas or nodules that grow out from the line of inoculation.

Media: Enzymatic digest of gelatin (10 g), beef extract (3 g), NaCl (5 g), agar (4 g), per 1000 mL, pH 7.3.

1. Touch a straight needle to a colony of a young (18- to 24-hour) culture growing on agar medium.

2. Stab once to a depth of only to inch in the middle of the tube.

3. Incubate at 35°-37°C and examine daily for up to 7 days.

Positive: Motile organisms will spread out into the medium from the site of inoculation (Figure 13-28, A).

Figure 13-28 Motility test. A, Positive. B, Negative.

Negative: Nonmotile organisms remain at the site of inoculation (Figure 13-28, B).

Some organisms will not display sufficient growth in this medium to make an accurate determination, and additional follow-up testing is required.

Negative: Staphylococcus aureus (ATCC25923)

Procedure 13-27 MRS Broth

This test is used to determine whether an organism forms gas during glucose fermentation. Some Lactobacillus spp. and Leuconostoc sp. produce gas.

The MRS broth contains sources of carbon, nitrogen, and vitamins to support the growth of lactobacilli and other organisms. It is a selective medium that uses sodium acetate and ammonium citrate to prevent overgrowth by contaminating organisms. Growth is considered a positive result. A Durham tube may be added to differentiate Lactobacillus spp. from Leuconostoc sp.

Media: Enzymatic digest of animal tissue (10 g), beef extract (10 g), yeast extract (5 g), dextrose (20 g), NaC₂H₃O₂ (5 g), polysorbate 80 (1 g), KH₂PO₄ (2 g), ammonium citrate (2 g), MgSO₄ (0.1 g), MnSO₄ (0.05 g), per 1000 mL, pH 6.5.

1. Inoculate MRS broth with an 18- to 24-hour culture from agar or broth.

2. Incubate 24 to 48 hours at 35°-37°C in ambient air.

Positive: Leuconostoc sp.—Growth, gas production indicated by a bubble in the Durham tube, (Figure 13-29, A).

Figure 13-29 MRS broth. A, Positive; gas production by *Leuconostoc* sp. *(arrow).* B, Positive: growth, no gas production by *Lactobacillus* sp.

Positive: Lactobacillus spp.—Growth, no gas production (Figure 13-29, B).

Negative: No growth (not shown).

Positive: Lactobacillus lactis (ATCC19435)

Procedure 13-28 4-Methylumbelliferyl-β-D-Glucuronide (MUG) Test

This test is used to presumptively identify various genera of Enterobacteriaceae and verotoxin-producing Escherichia coli.

E. coli and other Enterobacteriaceae produce the enzyme β-d-glucuronidase, which hydrolyzes β-d-glucopyranosid-uronic derivatives to aglycons and d-glucuronic acid. The substrate 4-methylumbelliferyl-β-d-glucuronide is impregnated into the disk and is hydrolyzed by the enzyme to yield the 4-methylumbelliferyl moiety, which fluoresces blue under long wavelength ultraviolet light. However, verotoxin-producing strains of E. coli do not produce MUG, and a negative test result may indicate the presence of a clinically important strain.

1. Wet the disk with 1 drop of water.

2. Using a wooden applicator stick, rub a portion of a colony from an 18- to 24-hour-old pure culture onto the disk.

3. Incubate at 35°-37°C in a closed container for up to 2 hours.

4. Observe disk using a 366-nm ultraviolet light.

Positive: Electric blue fluorescence (Figure 13-30, A).

Figure 13-30 MUG test. A, Positive. B, Negative.

Negative: Lack of fluorescence (Figure 13-30, B).

Do not test colonies isolated from medias containing dyes (EMB, MAC), because it may make the interpretation difficult.

Only test on oxidase-positive organisms, because some oxidase-negative organisms naturally fluoresce.

Negative: Klebsiella pneumoniae (ATC13883)

Procedure 13-29 Nitrate Reduction

This test is used to determine the ability of an organism to reduce nitrate to nitrite. All members of the Enterobacteriaceae family reduce nitrate, but some members further metabolize nitrite to other compounds.

Anaerobic metabolism requires an electron acceptor other than atmospheric oxygen (O₂). Many gram-negative bacteria use nitrate as the final electron acceptor. The organisms produce nitrate reductase, which converts the nitrate (NO₃) to nitrite (NO₂). The reduction of nitrate to nitrite is determined by adding sulfanilic acid and alpha-naphthylamine. The sulfanilic acid and nitrite react to form a diazonium salt. The diazonium salt then couples with the alpha-naphthylamine to produce a red, water-soluble azo dye. If no color change occurs, the organism did not reduce nitrate or reduced it further to NH₃, NO, or N₂O₂. Zinc is added at this point; if nitrate remains, the zinc will reduce the compound to nitrite and the reaction will turn positive, indicating a negative test result for nitrate reduction by the organism. If no color change occurs after the addition of zinc, this indicates that the organism reduced nitrate to one of the other nitrogen compounds previously described. A Durham tube is placed in the broth for two reasons: (1) to detect deterioration of the broth before inoculation, as evidenced by gas formation in the tube; and (2) to identify denitrification by organisms that produce gas by alternate pathways; if gas is formed in the tube before the addition of the color indicator, the test result is negative for nitrate reduction by this method.

Media: Pancreatic digest of gelatin (20 g), KNO₃ (2 g), per 1000 mL.

2. Incubate for 48 hours at 35°-37°C in ambient air (some organisms may require longer incubation for adequate growth). Test these cultures 24 hours after obvious growth is detected or after a maximum of 7 days.

1. Inoculate nitrate broth (Figure 13-31, D) with 1 to 2 drops from a young broth culture of the test organism.

Figure 13-31 Nitrate reduction. A, Positive, no gas. B, Positive, gas *(arrow).* C, Positive, no color after addition of zinc *(arrow).*D, Uninoculated tube.

3. After a suitable incubation period, test the nitrate broth culture for the presence of gas, reduction of nitrate, and reduction of nitrite according to the following steps:

a. Observe the inverted Durham tube for the presence of gas, indicated by bubbles inside the tube.

b. Add 5 drops each of nitrate reagent solution A (sulfanilic acid) and B (alpha-naphthylamine). Observe for at least 3 minutes for a red color to develop.

c. If no color develops, test further with zinc powder. Dip a wooden applicator stick into zinc powder and transfer only the amount that adheres to the stick to the nitrate broth culture to which solutions A and B have been added. Observe for at least 3 minutes for a red color to develop. Breaking the stick into the tube after the addition of the zinc provides a useful marker for the stage of testing.

The nitrate reduction test is read for the presence or absence of three metabolic products: gas, nitrate (NO₃), and nitrite (NO₂). The expected results can be summarized as follows:

Reaction	Gas	Color after Addition of Solutions A and B	Color after Addition of Zinc	Interpretation
NO₃ → NO₂ (Figure 13-31, A)	None	Red	—	NO₃+, no gas
NO₃ → NO₂, gas partial nongaseous end products	None	Red	—	NO₃+, no
NO₃ → NO₂, gaseous end products (Figure 13-31, B)	Yes	Red	—	NO₃+, gas+
NO₃ → gaseous end product (Figure 13-31, C)	Yes	None	None	NO₃+, NO₂+, gas+ C)
NO₃ → nongaseous end products	None	None	None	NO₃+, NO₂+, no gas
NO₃ → no reaction	None	None	Red	Negative

Nitrate reduction is a supportive test for identification of Enterobacteriaceae to the genus level; however, additional follow-up, confirmatory testing is required for final identification.

Positive: NO₃+, no gas: Escherichia coli (ATCC25922)

Positive: NO₃+, gas: Pseudomonas aeruginosa (ATCC17588)

Negative: Acinetobacter baumannii (ATCC19606)

Procedure 13-30 Nitrite Reduction

This test is used to determine whether an organism can reduce nitrites to gaseous nitrogen or to other compounds containing nitrogen.

Media: Brain-heart infusion broth (2 g), pancreatic digest of casein (10 g), peptic digest of animal tissue (5 g), yeast extract (3 g), NaCl (5 g), NaNO₂ (0.1 g), per 1000 mL, pH 6.9.

Microorganisms capable of reducing nitrite to nitrogen do not turn color and do produce gas in the nitrate reduction test (see Procedure 13-29). The test does not require the addition of zinc dust.

1. Inoculate nitrite broth with 1 drop from a 24-hour broth culture.

2. Incubate for 48 hours at 35°-37°C.

3. Examine 48-hour nitrite broth cultures for nitrogen gas in the inverted Durham tube and add 5 drops each of the nitrate reagents A and B to determine whether nitrite is still present in the medium (reagents A and B are described under the nitrate reduction test in Procedure 13-29).

Positive: No color change to red 2 minutes after the addition of the reagents; gas production observed in the Durham tube (Figure 13-32, A).

Figure 13-32 Nitrite reduction. A, Positive, no color change after addition of zinc dust and gas in Durham tube *(arrow).* B, Negative.

Negative: The broth becomes red after the addition of the reagents. No gas production is observed (Figure 13-32, B).

If the broth does not become red and no gas production is observed, zinc dust is added to determine if the nitrite has not been oxidized to nitrate (thus invalidating the test). If oxidation has occurred, the mixture turns red after the addition of zinc.

Positive: Proteus mirabilis (ATCC12453)—colorless, gas production

Negative: Acinetobacter baumannii (ATCC19606)—red, no gas production

Procedure 13-31 o-Nitrophenyl-β-D-Galactopyranoside (ONPG) Test

This test is used to determine the ability of an organism to produce β-galactosidase, an enzyme that hydrolyzes the substrate ONPG to form a visible (yellow) product, orthonitrophenol. The test distinguishes late lactose fermenters from non–lactose fermenters of Enterobacteriaceae.

Lactose fermenters must be able to transport the carbohydrate (β-galactoside permease) and hydrolyze (β-galactosidase) the lactose to glucose and galactose. Organisms unable to produce β-galactosidase may become genetically altered through a variety of mechanisms and be identified as late-lactose fermenters. ONPG enters the cells of organisms that do not produce the permease but are capable of hydrolyzing the ONPG to galactose and a yellow compound, o-nitrophenol, indicating the presence of β-galactosidase.

Media (tube method): Na₂HPO₄ (9.46 g), phenylalanine (4 g), ONPG (2 g), KH₂PO₄ (0.907 g), per 1000 mL, pH 8.0.

1. Aseptically suspend a loop full of organism in 0.85% saline.

2. Place an ONPG disk in the tube.

3. Incubate for 4 hours at 37°C in ambient air.

4. Examine tubes for a color change.

Positive: Yellow (presence of β-galactosidase) (Figure 13-33, A).

Figure 13-33 OPNG test. A, Positive. B, Negative.

Negative: Colorless (absence of enzyme) (Figure 13-33, B).

Positive: Shigella sonnei (ATCC9290)

Negative: Salmonella typhimurium (ATCC14028)

Procedure 13-32 Optochin (P disk) Susceptibility Test

This test is used to determine the effect of Optochin (ethyl hydrocupreine hydrochloride) on an organism. Optochin lyses pneumococci (positive test), but alpha-streptococci are resistant (negative test).

Optochin is an antibiotic that interferes with the ATPase and production of adenosine triphosphate (ATP) in microorganisms. The Optochin-impregnated disk (TaxoP) is placed on a lawn of organism on a sheep blood agar plate, allowing the antibiotic to diffuse into the medium. The antibiotic inhibits the growth of a susceptible organism, creating a clearing, or zone of inhibition, around the disk. A zone of 14 to 16 mm is considered susceptible and presumptive identification for Streptococcus pneumoniae.

1. Using an inoculating loop, streak two or three suspect colonies of a pure culture onto half of a 5% sheep blood agar plate.

2. Using heated forceps, place an Optochin disk in the upper third of the streaked area. Gently tap the disk to ensure adequate contact with the agar surface.

3. Incubate the plate for 18 to 24 hours at 35°C in 5% CO₂. Note: Cultures do not grow as well in ambient air, and larger zones of inhibition occur.

4. Measure the zone of inhibition in millimeters, including the diameter of the disk.

Positive: Zone of inhibition ≥ 14 mm in diameter, with 6-mm disk (Figure 13-34, A).

Figure 13-34 Optochin (TaxoP disk) test. A, *Streptococcus pneumoniae* showing zone of inhibition greater than 14 mm. B, Alpha-hemolytic *Streptococcus* sp. growing up to the disk.

Negative: No zone of inhibition (Figure 13-34, B).

Equivocal: Any zone of inhibition less than 14 mm is questionable for pneumococci; the strain is identified as a pneumococcus with confirmation by a positive bile-solubility test.

Positive: Streptococcus pneumoniae (ATCC6305)

Negative: Streptococcus pyogenes (ATCC12384)

Procedure 13-33 Oxidase Test (Kovac’s Method)

This test determines the presence of cytochrome oxidase activity in microorganisms for the identification of oxidase-negative Enterobacteriaceae, differentiating them from other gram-negative bacilli.

To determine the presence of bacterial cytochrome oxidase using the oxidation of the substrate tetramethyl-p-phenylenediamine dihydrochloride to indophenol, a dark purple-colored end product. A positive test (presence of oxidase) is indicated by the development of a dark purple color. No color development indicates a negative test and the absence of the enzyme.

1. Moisten filter paper with the substrate (1% tetramethyl-p-phenylenediamine dihydrochloride) or select a commercially available paper disk that has been impregnated with the substrate.

2. Use a platinum wire or wooden stick to remove a small portion of a bacterial colony (preferably not more than 24 hours old) from the agar surface and rub the sample on the filter paper or commercial disk.

3. Observe the inoculated area of paper or disk for a color change to deep blue or purple (Figure 13-35) within 10 seconds (timing is critical).

Figure 13-35 Oxidase test. A, Positive. B, Negative.

Positive: Development of a dark purple color within 10 seconds (Figure 13-35, A).

Negative: Absence of color (Figure 13-35, B).

Using nickel-base alloy wires containing chromium and iron (nichrome) to rub the colony paste onto the filter paper may cause false-positive results.

Positive: Pseudomonas aeruginosa (ATCC27853)

Procedure 13-34 Oxidation/Fermentation (of) Medium (CDC Method)

This test is used to differentiate microorganisms based on the ability to oxidize or ferment specific carbohydrates.

This test is used to determine whether an organism uses carbohydrate substrates to produce acid byproducts. Nonfermentative bacteria are routinely tested for their ability to produce acid from six carbohydrates (glucose, xylose, mannitol, lactose, sucrose, and maltose). In addition to the six tubes containing carbohydrates, a control tube containing the OF base without carbohydrate is also inoculated. Triple sugar iron agar (TSI) (see Procedure 13-40) is also used to determine whether an organism can ferment glucose. OF glucose is used to determine whether an organism ferments (Figure 13-36, A) or oxidizes (Figure 13-36, B) glucose. If no reaction occurs in either the TSI or OF glucose, the organism is considered a non-glucose utilizer (Figure 13-36, C). Hugh and Leifson’s formula uses a low peptone-to-carbohydrate ratio and a limiting amount of carbohydrate. The reduced peptone limits the formation of alkaline amines that may mask acid production resulting from oxidative metabolism. Two tubes are required for interpretation of the OF test. Both are inoculated, and one tube is overlaid with mineral oil, producing an anaerobic environment. Production of acid in the overlaid tube results in a color change and is an indication of fermentation. Acid production in the open tube and color change is the result of oxidation. Media: Pancreatic digest of casein (2 g), glycerol (10.0 mL), phenol red (King method) (0.03 g), agar (3 g), per 1000 mL, pH 7.3.

1. To determine whether acid is produced from carbohydrates, inoculate agar deeps, each containing a single carbohydrate, with bacterial growth from an 18- to 24-hour culture by stabbing a needle 4 to 5 times into the medium to a depth of 1 cm. Note: Two tubes of OF dextrose are usually inoculated; one is overlaid with either sterile melted petrolatum or sterile paraffin oil to detect fermentation.

2. Incubate the tubes at 35°-37°C in ambient air for up to 7 days. Note: If screwcap tubes are used, loosen the caps during incubation to allow for air exchange. Otherwise, the control tube and tubes containing carbohydrates that are not oxidized might not become alkaline.

Positive: Acid production (A) is indicated by the color indicator changing to yellow in the carbohydrate-containing deep.

Weak-positive (Aw): Weak acid formation can be detected by comparing the tube containing the medium with carbohydrate with the inoculated tube containing medium with no carbohydrate. Most bacteria that can grow in the OF base produce an alkaline reaction in the control tube. If the color of the medium in a tube containing carbohydrate remains about the same as it was before the medium was inoculated and if the inoculated medium in the control tube becomes a deeper red (i.e., becomes alkaline), the culture being tested is considered weakly positive, assuming the amount of growth is about the same in both tubes.

Negative: Red or alkaline (K) color in the deep with carbohydrate equal to the color of the inoculated control tube.

No change (NC) or neutral (N): There is growth in the media, but neither the carbohydrate-containing medium nor the control base turns alkaline (red).

Note: If the organism does not grow at all in the OF medium, mark the reaction as no growth (NG).

Slow-growing organisms may not produce results for several days.

Note: Appropriate organisms depend on which carbohydrate has been added to the basal medium. Glucose is used as an example.

Fermenter: Escherichia coli (ATCC25922)

Oxidizer: Pseudomonas aeruginosa (ATCC27853)

Procedure 13-35 Phenylalanine Deaminase Agar

This test is used to determine the ability of an organism to oxidatively deaminate phenylalanine to phenylpyruvic acid. The genera Morganella, Proteus, and Providencia can be differentiated from other members of the Enterobacteriaceae family.

Microorganisms that produce phenylalanine deaminase remove the amine (NH₂) from phenylalanine. The reaction results in the production of ammonia (NH₃) and phenylpyruvic acid. The phenylpyruvic acid is detected by adding a few drops of 10% ferric chloride; a green-colored complex is formed between these two compounds.

Media: Phenylalanine (2 g), yeast extract (3 g), NaCl (5 g), Na₃PO₄ (1 g), agar (12 g), per 1000 mL, pH 7.3.

1. Inoculate phenylalanine slant with 1 drop of a 24-hour brain-heart infusion broth.

2. Incubate 18 to 24 hours (or until good growth is apparent) at 35°-37°C in ambient air with cap loose.

3. After incubation, add 4 to 5 drops of 10% aqueous ferric chloride to the slant.

Positive: Green color develops on slant after ferric chloride is added (Figure 13-37, A).

Figure 13-37 Phenylalanine deaminase. A, Positive. B, Negative.

Negative: Slant remains original color after the addition of ferric chloride (Figure 13-37, B).

Positive: Proteus mirabilis (ATCC12453)

Procedure 13-36 L-Pyrrolidonyl Arylamidase (PYR) Test

This test is used for the presumptive identification of group A streptococci (Streptococcus pyogenes) and enterococci by the presence of the enzyme L-pyrrolidonyl arylamidase.

The enzyme L-pyrrolidonyl arylamidase hydrolyzes the L-pyrrolidonyl- β-naphthylamide substrate to produce a β-naphthylamine. The β-naphthylamine can be detected in the presence of N,N-methylaminocinnamaldehyde reagent by the production of a bright red precipitate.

1. Before inoculation, moisten the disk slightly with reagent-grade water. Do not flood the disk.

2. Using a wooden applicator stick, rub a small amount of several colonies of an 18- to 24-hour pure culture onto a small area of the PYR disk.

3. Incubate at room temperature for 2 minutes.

4. Add a drop of detector reagent, N,N-dimethylaminocinnamaldehyde, and observe for a red color within 1 minute.

Positive: Bright red color within 5 minutes (Figure 13-38, A).

Figure 13-38 PYR test. A, Positive. B, Negative.

Negative: No color change or an orange color (Figure 13-38, B).

Positive: Enterococcus faecalis (ATCC29212)

Streptococcus pyogenes (ATCC19615)

Negative: Streptococcus agalactiae (ATCC10386)

Procedure 13-37 Pyruvate Broth

This test is used to determine the ability of an organism to utilize pyruvate. This aids in the differentiation between Enterococcus faecalis (positive) and Enterococcus faecium (negative).

Pyruvate broth is a carbohydrate-free, nutrient-limited medium. Pyruvic acid is added to the broth to determine whether the microorganism is able to use pyruvate, resulting in the formation of metabolic acids. Bromthymol blue indicator changes from blue to yellow in the presence of acid as a result of the decrease in pH.

Media: Pancreatic digest of casein (10 g), pyruvic acid, sodium (10 g), yeast extract (5 g), K₂HPO₄ (5 g), NaCl (5 g), bromthymol blue (40 g), per 1000 mL, pH 7.3.

1. Lightly inoculate the pyruvate broth with an 18- to 24-hour culture of the organism from 5% sheep blood agar.

2. Incubate at 35°-37°C in ambient air for 24 to 48 hours.

Positive: Indicator changes from green to yellow (Figure 13-39, A).

Figure 13-39 Pyruvate broth. A, Positive. B, Negative.

Negative: No color change; yellow-green indicates a weak reaction and should be regarded as negative (Figure 13-39, B).

Positive: Enterococcus faecalis (ATCC29212)

Negative: Streptococcus bovis (ATCC9809)

Procedure 13-38 Salt Tolerance Test

This test is used to determine the ability of an organism to grow in high concentrations of salt. It is used to differentiate enterococci (positive) from nonenterococci (negative).

The salt tolerance test is a selective and differential medium. Enterococci are resistant to high salt concentration. A heart infusion broth containing 6.5% NaCl is used as the test medium. This broth also contains a small amount of glucose and bromcresol purple as the indicator for acid production.

Media: Brain-heart infusion broth (BHI) may be used in place of the individual components with the addition of NaCl and indicator dye. Components: Heart digest (10 g), enzymatic digest of animal tissue (10 g), NaCl (65 g), dextrose (1 g), bromocresol purple (0.016 g), per 1000 mL.

1. Inoculate one or two colonies from an 18- to 24-hour culture into 6.5% NaCl broth.

2. Incubate the tube at 35°-37°C in ambient air for 48 hours.

3. Check daily for growth.

Positive: Visible turbidity in the broth, with or without a color change from purple to yellow (Figure 13-40, A).

Figure 13-40 Salt tolerance (6.5% NaCl) test. A, Positive. B, Negative.

Negative: No turbidity and no color change (Figure 13-40, B).

Positive: Enterococcus faecalis (ATCC29212)—growth, color change to yellow

Negative: Streptococcus bovis (ATCC9809)—inhibition, as demonstrated by little to no growth, no color change

Procedure 13-39 Spot Indole Test

This test is used to determine the presence of the enzyme tryptophanase. It is a rapid method that can be used in lieu of the tube test described in Procedure 13-20.

Tryptophanase breaks down tryptophan to release indole, which is detected by its ability to combine with certain aldehydes to form a colored compound. For indole-positive bacteria, the blue-green compound formed by the reaction of indole with cinnamaldehyde is easily visualized. The absence of enzyme results in no color production (indole negative).

1. Saturate a piece of filter paper with the 1% paradimethylaminocinnamaldehyde reagent.

2. Use a wooden stick or bacteriologic loop to remove a small portion of a bacterial colony from the agar surface and rub the sample on the filter paper. Rapid development of a blue color indicates a positive test result. Most indole-positive organisms turn blue within 30 seconds.

Positive: Development of a blue color within 20 seconds (Figure 13-41, A).

Figure 13-41 Spot indole test. A, Positive. B, Negative.

Negative: No color development or slightly pink color (Figure 13-41, B).

The bacterial inoculum should not be selected from MacConkey agar, because the color of lactose-fermenting colonies on this medium can interfere with test interpretation.

Negative: Klebsiella pneumoniae (ATCC13883)

Procedure 13-40 Triple Sugar Iron Agar (TSI)

TSI is used to determine whether a gram-negative rod ferments glucose and lactose or sucrose and forms hydrogen sulfide (H₂S). The test is used primarily to differentiate members of the Enterobacteriaceae family from other gram-negative rods.

The composition of TSI is 10 parts lactose:10 parts sucrose:1 part glucose and peptone. Phenol red and ferrous sulfate serve as indicators of acidification and H₂S formation, respectively. A glucose-fermenting organism turns the entire medium acidic (yellow) in 8 to 12 hours. The butt remains acidic after the recommended 18- to 24-hour incubation period because of the presence of organic acids resulting from the fermentation of glucose under anaerobic conditions in the butt of the tube. The slant, however, reverts to the alkaline (red) state because of oxidation of the fermentation products under aerobic conditions on the slant. This change is a result of the formation of CO₂ and H₂O and the oxidation of peptones in the medium to alkaline amines. When, in addition to glucose, lactose and/or sucrose are fermented, the large amount of fermentation products formed on the slant neutralizes the alkaline amines and renders the slant acidic (yellow), provided the reaction is read in 18 to 24 hours. Reactions in TSI should not be read beyond 24 hours of incubation, because aerobic oxidation of the fermentation products from lactose and/or sucrose proceeds, and the slant eventually reverts to the alkaline state. The formation of CO₂ and hydrogen gas (H₂) is indicated by the presence of bubbles or cracks in the agar or by separation of the agar from the sides or bottom of the tube. The production of H₂S (sodium thiosulfate reduced to H₂S) requires an acidic environment, and reaction with the ferric ammonium citrate produces a blackening of the agar butt in the tube.

Media: Enzymatic digest of casein (5 g), enzymatic digest of animal tissue (5 g), yeast-enriched peptone (10 g), dextrose (1 g), lactose (10 g) sucrose (10 g), ferric ammonium citrate (0.2 g), NaCl (5 g), sodium thiosulfate (0.3 g), phenol red (0.025 g), agar (13.5 g), per 1000 mL, pH 7.3.

1. With a straight inoculation needle, touch the top of a well-isolated colony.

2. Inoculate TSI (Figure 13-42, D) by first stabbing through the center of the medium to the bottom of the tube and then streaking the surface of the agar slant.

Figure 13-42 Triple sugar iron agar. A, Acid slant/acid butt with gas, no H₂S (A/A). B, Alkaline slant/acid butt, no gas, H₂S-positive (K/A H₂S+). C, Alkaline slant/alkaline butt, no gas, no H₂S (K/K). D, Uninoculated tube.

3. Leave the cap on loosely and incubate the tube at 35°-37°C in ambient air for 18 to 24 hours.