Overnight Biochemical Tests

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The overnight biochemical tests are a group of tests that require inoculating one or more culture media containing specific substrates and chemical indicators that detect pH change or specific microbial by-product. Similar to rapid tests, the choice of overnight tests is based on Gram-stain morphology and the results of preliminary testing with rapid enzyme tests. These tests are also inexpensive and easy to perform and may be used in three different ways. They may be used to obtain important initial information with respect to the identity of an unknown organism, such as the MILS test, which is used to screen for the presence of enteric pathogens. They may be used to verify the results of a preliminary positive/negative test or they may be used to assess an indeterminate finding. For example, Taxo P is an overnight test that will demonstrate if an isolate with an equivocal bile solubility result is S. pneumoniae. Similarly, a tube coagulase test will substantiate if a suspicious isolate, that is slide coagulase negative, is truly a coagulase-negative staphylococci. Finally, these tests may be used as the sole identification system (classical biochemical identification) to identity an unknown organism. This is generally labor intensive and requires the technologist to inoculate, incubate, read, interpret, and chart a number of biochemical reactions over several days. This is then followed by using various identification schemes or flow charts to generate a final identification. As a rule, the classical biochemical identification system is used to identify fastidious or slow-growing organisms in the reference laboratories. These isolates are by and large rare biotypes that are not part of the commercial identification system's database. Table 6.1 depicts the list of biochemical tests that are commonly used to identify Gram-negative bacilli (Weyant et al., 1996).

Tube Coagulase Test

Tube coagulase test detects free coagulase (liberated by the cell) that acts on prothrombin to produce a thrombin-like product that then acts as fibrinogen to form a fibrin clot.

Prepare a heavy suspension of the Staphylococcus colonies in 0.5 mL of water. Place the suspension into a tube containing rabbit plasma and incubate at 35°C for 4 h. Examine for the presence of a clot. If negative for a clot, reincubate the tube and reexamine at 24 h. Any degree of clot formation at 4 h or 24 h is considered a positive reaction. No clot formation at 24 h is considered negative coagulase reaction (Koneman et al., 1997; Forbes et al., 2002; Murray et al., 2003).

DNA Hydrolysis

The DNA hydrolysis test detects the presence of enzyme DNase in an organism. Using this media, DNase-positive coagulase-positive staphylococci are differentiated from other Staphylococcus spp. The media contains either toluidine blue or methyl green, which upon hydrolysis of the incorporated DNA turns colorless.

The media is inoculated with the organism and incubated overnight at 35°C. The plate is examined for evidence of growth and loss of color (positive reaction). No color change indicates a negative reaction (Murray et al., 2003).

Vancomycin Disk Test

The vancomycin disk test is performed as a susceptibility procedure to help differentiate the Gram-positive, catalase-negative cocci. Aerococcus, Gemella, Lac-tococcus, Streptococcus, and some enterococci are susceptible to vancomycin. Leuconostoc, Pediococcus, Lactobacillus, and some enterococci are resistant to vancomycin.

A 0.5 McFarland suspension of the organism is prepared in sterile saline. Using a sterile swab, the bacterial suspension is inoculated onto a BAP. A vancomycin disk is placed in the center of the inoculated plate and incubated at 35°C in a CO2 incubator for 18-24 h. The plate is observed for the presence of a zone of inhibition around the vancomycin disk. Leuconostoc spp., Pediococcus spp., Lactobacillus spp., and some Enterococcus spp. are resistant to vancomycin with growth to the edge of the disk <9 mm. Aerococcus spp, Gemella spp., Lactococcus spp., Streptococcus spp., and some Enterococcus spp. are susceptible to vancomycin and produce a zone of inhibition >12 mm (Koneman et al., 1997; Murray et al., 2003).

Table 6.1. Commonly used biochemical tests for identification of a Gram-negative organism.

Date

Date

Table 6.1. Commonly used biochemical tests for identification of a Gram-negative organism.

Biochemicals

1

2

3

4

5

6

7

Motility

OF glucose (oxid)

OF glucose (Ferm)

Xylose

Mannitol

Lactose

Sucrose

Maltose

Catalase

Oxidase

MacConkey

Citrate

Sodium acetate

Urea

Nitrate

Nitrate to gas

Indole

TSI slant

TSI butt

H2S (TSI butt)

H2S (Pb ac paper)

Gelatin

Pigment

Arginine

Lysine

Growth at 42°C

Bacitracin Inhibition Test (Taxo A Disk)

The bacitracin inhibition test presumptively differentiates group A streptococci (GAS) from other beta-hemolytic streptococci. The bacitracin at concentration of 0.04 units will selectively inhibit growth of GAS. Although there are rare strains of GAS that are bacitracin resistant, approximately 5% to 10% of strains of non-group A beta hemolytic streptococci (b, C, and G) are bacitracin susceptible.

Using a pure culture of the test organism, inoculate a BAP with the bacterial suspension. Place a bacitracin disk in the center of the inoculated BAP and incubate at 35°C for 18-24 h. Any zone of inhibition around the bacitracin disk is considered a positive test. Uniform lawn of growth right up to the rim of the disk indicates a negative bacitracin inhibition test (Isenberg, 1992; Koneman et al., 1997; Murray et al., 2003).

Taxo P Disks (Optochin)

Hydrocupreine hydrochloride (optochin) at the concentration 5.0 |xg inhibits the growth of S. pneumoniae, but not of other streptococci. S. pneumoniae may, therefore, be differentiated from other alpha-hemolytic streptococci by the formation of a zone of inhibition around a disk impregnated with this compound.

Inoculate a BAP with apure culture of the alpha-hemolytic Streptococcus isolate. Place a Taxo P disk (optochin) onto the inoculated plate and incubate the plate aerobically at 35°C for 24 h or as needed to obtain good growth. Incubation in a CO2 enriched atmosphere will enhance growth but reduce zone size. Zones of inhibition of 14 mm or more are formed with pure cultures of S. pneumoniae. Other organisms may show zone sizes less than 14 mm in diameter. A diameter between 6 and 14 mm is questionable for S. pneumoniae and the strain should be tested for bile solubility (Murray et al., 2003).

CAMP Test

The CAMP test is based on the fact that group B streptococci produce a protein-like compound known as the CAMP factor that acts synergistically with a staphylococ-cal beta-hemolysin (p -lysin) on sheep erythrocytes to produce an enhanced zone of hemolysis.

Streak a loopful of p toxin-producing S. aureus in a straight line across the center of a BAP. Streak a loopful of group B streptococci perpendicular to and nearly touching the streak line of the staphylococci (positive control). Streak a loopful of group A streptococci perpendicular to and nearly touching the streak line of the staphylococci (negative control). Streak a loopful of unknown isolate perpendicular to and nearly touching the streak line of the staphylococci and incubate the plate at 35°C for 24 h in the aerobic non-CO2 incubator. Following the incubation, if the patient isolate demonstrates an arrowhead zone of enhanced hemolysis, the isolate is identified as group B streptococci. If the patient isolate does not demonstrate an arrowhead of enhanced hemolysis, the isolate is not group B streptococci.

Do not incubate the CAMP test plate in the presence of 5-10% CO2 incubator. This may result in an incorrect interpretation (Wilkinson, 1977; Isenberg, 1992).

Reverse CAMP Test

The reverse CAMP test is based on the fact that some organisms such as Arcanobacterium haemolyticum completely inhibit the effect of Staphylococcus aureus B-hemolysin on sheep erythrocytes. The (3-hemolysin inhibition zone in the form of a triangle is formed.

A loopful of ( toxin-producing Staphylococcus aureus is streaked in a straight line across the center of a BAP. Group B streptococci and group A streptococci are streaked perpendicular to and nearly touching the streak line of the staphylococci. Similarly, A. haemolyticum and the test isolate are streaked perpendicular to and nearly touching the line of the staphylococci. The plate is incubated at 35° C for 24 h in the aerobic non-CO2 incubator. Following the incubation, if the test isolate demonstrates a triangular-shaped inhibition of ( -hemolysis, it is a reverse camp test positive. If the test isolate does not demonstrate a triangle-shaped inhibition of ( -hemolysis, it is a reverse camp test negative.

Do not incubate the reverse CAMP test plate in the 5-10% CO2 incubator. This may result in an incorrect interpretation (Wilkinson, 1977; Isenberg, 1992).

Bile Esculin Agar Slant

Group D streptococci (including Enterococcus spp.) and a few other bacteria, such as Listeria spp., can grow in the presence of 40% bile and also hydrolyze esculin to esculetin. Esculetin reacts with ferric ions, supplied by ferric citrate in the agar medium, to form a diffusible black complex. Most strains of viridans streptococci that are capable of hydrolyzing esculin will not grow in the presence of 40% bile.

Streak the surface of the bile esculin agar slant with several colonies of the organism to be tested. Incubate at 35°C in non-CO2 for 24 to 48 h. A diffuse blackening of more than half of the slant within 24 to 48 h is considered positive. No growth or growth without blackening of the medium after 48 h is considered negative test.

If the inoculum is too heavy, viridans streptococci may give a false-positive test result. Approximately 3% of viridans streptococci are able to hydrolyze esculin in the presence of bile. Growth in the presence of 6.5% salt is used to differentiate en-terococci from non-enterococcal group D streptococci (Isenberg, 1992; Koneman et al., 1997).

6.5% Salt Broth

Trypticase soy broth is a general-purpose medium for the cultivation of both fastidious and nonfastidious organisms. With the addition of 6.5% sodium chloride, the medium can be used to differentiate between salt-tolerant and salt-intolerant organisms. It is especially useful for distinguishing Enterococcus spp., which are salt-tolerant, from non-enterococcal group D streptococci, such as S. bovis and S. equinus.

Inoculate the tube containing 6.5% sodium chloride with the organism and incubate at 35°C in non-CO2 for 24-48 h. A visible growth (turbidity) is considered positive and no growth is considered negative.

If the medium is inoculated too heavily, the inoculum may be interpreted as growth, resulting in a false-positive reaction. Aerococcus, Pediococcus, Staphylococcus, and up to 80% of group B Streptococcus can grow in 6.5% salt broth. In addition, Aerococcus may also be bile esculin positive (Isenberg, 1992; Koneman et al., 1997).

Indole Test

Indole, a benzyl pyrrole, is one of the metabolic degradation products of the amino acid tryptophan. Bacteria that possess the enzyme tryptophanase are capable of hydrolyzing and deaminating tryptphan with the production of indole, pyruvic acid, and ammonia. The indole test is based on the formation of a red color complex when indole reacts with the aldehyde group of p-dimethylaminobenzaldehyde, the active chemical in Kovac's reagent. In order to perform this test, the organism must be grown on a medium rich in tryptophan such as indole nitrite broth.

Inoculate the indole nitrite broth medium with 2-3 colonies of the organism to be tested. Incubate the tubes at 35°C in a non-CO2 incubator for 24-48 h. Examine the tubes for growth. When the broth is visibly turbid, use a sterile pipette to transfer 3 mL into a sterile tube. Add 1 mL of xylene to the contents of the tube, which extracts the indole, if present, from the broth into the xylene. Wait 1-2 min, and add 0.5 mL Kovac's reagent and observe for the production of a pink to red color in the xylene layer. A pink to red color at the interface the of the reagent and the broth within seconds after the addition of Kovac's reagent indicates a positive reaction. No color change indicates a negative reaction (Koneman et al., 1997).

Nitrite Test

Organisms that reduce nitrate have the ability to extract oxygen from nitrates to form nitrites and other reduction products. The presence of nitrites in the medium are detected by the formation of a red diazonium dye, p-sulfobenzeneazo-a-naphthylamine, following the addition of a-naphthylamine and sulfanilic acid. If no color develops after adding the reagents, this indicates that nitrates have not been reduced (a true negative reaction) or that they have been reduced beyond the oxidation level of nitrite to products such as ammonia, nitrogen gas (denitrifica-tion), nitric oxide (NO), or nitrous oxide (N2O) and hydroxylamine. Because the test reagents detect only nitrites, the latter process would lead to a false-negative result. Therefore, it is necessary to add a small amount of zinc dust to all negative reactions. Because zinc ions reduce nitrates to nitrites, the development of a red color after adding zinc dust indicates the presence of nitrates and confirms a true negative reaction.

Using a sterile inoculating loop, an indole nitrite broth medium is inoculated with 2-3 colonies of the organism to be tested and incubated at 35°C in a non-CO2 incubator for 24-48 h. When the broth is visibly turbid, 3 mL of the broth culture is transferred into a sterile tube and 5 drops of N,N-dimethyl-a-naphthylamine (nitrate reagent A) is added to the broth. Five drops of sulfanilic acid (nitrate reagent B) is then added to the broth and observed for the production of a pink to red color within 30 s. If no color change occurs within 30 s, a small amount of zinc dust is added and the production of a pink to red color within 10 min is looked for (Koneman et al., 1997).

ALA (Haemophilus influenzae Porphyrin Test)

The porphyrin test is used in the rapid speciation of Haemophilus by separating those species that require an exogenous source of X factor from those that do not. Haemophilus species (H. parainfluenzae and H. parahemolyticus) that produce the enzyme porphobilinogen synthase have the ability to synthesize heme (factor X) and therefore do not require an exogenous source of factor X for growth. Porphobilinogen and porphyrin, precursors in heme synthesis, can be detected in an enzyme substrate inoculated with a porphobilinogen synthase producing Haemophilus spp. by the addition of modified Ehrlich's (Kovac's) reagent or by examination with a Wood's lamp.

Suspend a loopful of organism in 0.5 mL of the enzyme substrate. Incubate at 35°C for 4 h if the suspension is heavy or 18-24 h if the suspension is light. After incubation add an equal volume of modified Ehrlich's (Kovac's) reagent and vortex the mixture. Allow the substrate and reagent to separate. After the addition of Kovac's reagent, a red (pink) color will form in the aqueous phase, indicating the presence of porphobilinogen, and therefore a positive test for Haemophilus spp. not requiring factor X. Alternatively, a Wood's lamp can be used to detect fluorescence in the reagent phase, indicating the presence of porphyrins, also a positive test. No coloration or fluorescence indicates a factor X dependent Haemophilus spp. and a negative test (Killian, 1974).

Motility Indole Lysine (MILS)

MILS medium is a semisolid medium useful in the identification of members of the Enterobacteriaceae, specifically for screening suspicious colonies from stool cultures for potential pathogens.

It is used to demonstrate motility, indole production, lysine decarboxylase and deaminase activity, and hydrogen sulfide production. A small amount of agar is added to the media for demonstration of motility along a stab line of inoculation. Growth of motile organisms extends out from the line of inoculation, whereas nonmotile organisms grow along the stab line.

The pH indicator bromcresol purple is used to facilitate detection of decarboxylase activity. When inoculated with an organism that ferments dextrose, acids are produced that lower the pH, causing the indicator in the medium to change from purple to yellow. The acidic pH also stimulates enzyme activity. Organisms that possess a specific decarboxylase degrade the amino acid provided in the medium, yielding a corresponding amine. Lysine decarboxylation yields cadaverine. The production of these amines elevates the pH and causes the medium in the bottom portion of the tube to return to a purple color. The medium in the upper portion of the tube remains acidic because of the higher oxygen tension. Lysine deami-nation produces a color change in the upper portion of MILS medium. Oxidative deamination of lysine yields a compound that reacts with ferric ammonium citrate, producing a burgundy-red color in the top of the medium. (The bottom portion of the medium remains acidic.) This reaction can only be detected if lysine decarboxylation is not produced, which is the case with Proteus, Morganella, and Providencia species.

Indole is produced in MILS medium by organisms that possess the enzyme tryptophanase. Tryptophanase degrades the tryptophan present in the casein peptone, yielding indole. Indole can be detected in the medium by adding Kovac's reagent to the agar surface. MILS medium is also used in the demonstration of hydrogen sulfide production. Hydrogen sulfide, which is produced by some enteric organisms from sulfur compounds contained in the medium, reacts with ferric ion, producing a characteristic black precipitate (BD Microbiology Systems, 1999).

ONPG (O-Nithrophenyl-beta-D-Galactopyranoside) Test

In order for an organism to ferment lactose, it must have the enzymes perme-ase to transport the lactose inside the cell and beta-galactosidase to cleave the transported sugar. Some organisms (delayed lactose fermenters) though possessing beta-galactosidase do not have the enzyme permease. These organisms can utilize the enzyme beta-galactosidase to hydrolyze ONPG. ONPG is a colorless compound similar to lactose. In the presence of beta-galactosidase, ONPG is hydrolyzed to galactose and a yellow compound o-nitrophenyl.

Inoculate an ONPG broth tube with the organism and add the ONPG disk and incubate at 35°C. Periodically examine the color change for up to 24 h. Yellow color indicates a positive reaction and no color change indicates a negative reaction (Murray et al., 2003).

Methyl Red (MR) Test

This assay determines if an organism metabolizing pyruvic acid utilizes mixed acid pathway and produces acid end products that are detected by the indicator methyl red.

A 5 mL MR-VP broth tube is inoculated with the organism and incubated at 35°C for 48 h, then 2.5 mL of the broth culture is transferred to a fresh tube and inoculated with 5 drops of methyl red indicator. Positive MR is indicated if the methyl red reagent remains red. Negative result is indicated if the reagent turns yellow-orange (Koneman et al., 1997; Murray et al., 2003).

Voges-Proskaure (VP) Test

Organisms such as Klebsiella, Enterobacter, and Serratia spp. that utilize the butylenes glycol fermentation pathway produce acetoin, an intermediate in the fermentation of butylenes glycol. The VP test detects the production of acetoin by these organisms. In the presence of air and potassium hydroxide, acetoin is oxidized to diacetyl, which produces a red-colored complex. The addition of alpha-naphtol increases the sensitivity of the test.

A 5 mL MR-VP broth tube is inoculated with the organism and incubated at 35°C for 18-24 h, then 2.5 mL of the broth culture is transferred to a fresh tube and inoculated with 6 drops of alpha naphtol followed by 3 drops of KOH. A positive result is indicated by the presence of a red color that develops within 15 min. No color change indicates negative VP (Koneman et al., 1997; Murray et al., 2003).

Pseudosel Agar Slant

Pseudosel agar is a medium used for the identification of Pseudomonas aerug-inosa. Magnesium chloride and potassium sulfate in the medium enhance the production of pyocyanin, a blue-green, water-soluble, nonfluorescent phenazine pigment. P. aeruginosa is the only Gram-negative rod known to excrete pyocyanin. In addition to the promotion of pyocyanin production, pseudosel agar also enables the detection of fluorescent products by some Pseudomonas species other than P. aeruginosa. Streak the surface of the pseudosel agar slant, and incubate at 35°C in non-CO2 for 18-24 h. A blue-green pigmentation surrounding the growth on the agar slant indicates a positive reaction. No pigmentation indicates a negative reaction.

Negative pseudosel slants should be examined under short wavelength (254 nm) ultraviolet light to check for fluorescent products produced by some Pseudomonas species. Pseudomonas aeruginosa typically produces fluorescein as well as pyocyanin (BD Microbiology Systems, 1992).

Urea Agar Slant

Microorganisms that possess the enzyme urease are capable of hydrolyzing urea, which releases ammonia. This reaction raises the pH of the medium and is detected by phenol red, which turns pink-red above pH 8.0. The color change first appears in the slant because the oxidative decarboxylation of amino acids in the air-exposed portion of the medium enhances the alkaline reaction. The color change eventually spreads deeper into the medium.

Streak the surface of the urea agar slant with a heavy inoculum of a pure culture. Incubate at 35°C in non-CO2 for 18 to 24 h. Production of intense pink-red color on the slant, which may penetrate into the butt, is considered a positive reaction. No color change indicates negative a reaction.

The medium is not specific for urease. The utilization of peptones or other proteins in the medium by some urease-negative organisms may raise the pH due to protein hydrolysis and release of amino acid residues, resulting in false-positive reactions (Koneman et al., 1997; BD Microbiology Systems, 1992).

Citrate Agar Slant

Some organisms have the ability to utilize citrate, an intermediate metabolite in the Krebs cycle, as the sole external source of carbon. These organisms also utilize inorganic ammonium salts in the medium as the sole source of nitrogen. The resulting production of ammonia creates an alkaline environment that turns the bromthymol blue indicator to an intense blue.

Using an inoculating loop, select a well-isolated colony with and streak the surface of the citrate slant (do not stab the agar) and incubate at 35°C in non-CO2incubator and examine daily for up to 4 days. Growth with an intense blue color on the agar slant indicates a positive reaction and no growth and no color change (green) indicates a negative reaction.

Luxuriant growth on the slant without an accompanying color change may indicate a positive test. This should be confirmed by incubating the tube for an additional 24 h. The biochemical reaction requires oxygen. Therefore, the medium should not be stabbed, and the cap must be kept loose during incubation. Carry-over of protein and carbohydrate substrates from previous media may provide additional sources of carbon and therefore cause false-positive reactions (BD Microbiology Systems, 1992).

Cetrimide Agar

Cetrimide agar is a selective differential medium used for the identification of P. aeruginosa. The principle of the test is to determine the ability of an organism to grow in the presence of cetrimide. Cetrimide acts as a detergent and inhibits the growth of most other organisms. The iron content of the medium stimulates the production of pyocanin and fluorescent yellow-green pigment by this organism.

Using an inoculating loop, select a well-isolated colony with and streak the surface of the cetrimide slant (do not stab the agar) and incubate at 35° C in non-CO2 incubator and examine daily for up to seven days. Growth on the agar slant indicates positive reaction and no growth indicates a negative reaction (BD Microbiology Systems, 1992; Forbes et al., 2002).

Gelatin

The gelatin test is used to identify bacteria that produce the proteolytic enzyme gelatinase. Organisms that produce gelatinase are capable of hydrolyzing gelatin and cause it to lose its gelling characteristics. A gelatin tube may be inoculated with the organism and incubated at 35° C in ambient air. The tubes are then removed daily and incubated at 4°C to check for liquefaction. Alternatively, strips of exposed but undeveloped x-ray film are placed in the bacterial suspension equivalent to at least 2 McFarland standard and incubated at 35°C in a non-CO2 incubator for 48 h.

The strip is examined after 24 h and 48 h for loss of gelatin coating that leave the radiograph clear (Murray et al., 2003).

Acetate Utilization

Some organisms have the ability to utilize acetate as a sole external source of carbon. Acetate slants contain a mixture of salts and sodium acetate in a medium without organic nitrogen. Organisms that can utilize acetate as a sole carbon source break down sodium acetate causing the pH of the medium to shift toward the alkaline range, turning the bromthymol blue indicator blue. Organisms that cannot utilize acetate as a sole carbon source do not grow on the medium. Acetate differential agar is useful in the differentiation of Neisseria and Moraxella spp.

Streak the surface of the acetate differential agar slant (do not stab the agar), with a colony and cap the tube loosely. Incubate at 35°C in non-CO2and examine daily for up to 7 days. Growth with an intense blue color on the agar slant indicates a positive test and no growth or no color change (green) indicates a negative test.

Luxuriant growth on the slant without an accompanying color change may indicate a positive test. This should be confirmed by incubating the tube for an additional 24 h. The biochemical reaction requires oxygen. Therefore, the medium should not be stabbed, and the cap must be kept loose during incubation. Carry-over of protein and carbohydrate substrates from previous media may provide additional sources of carbon and therefore cause false-positive reactions (BD Microbiology Systems, 1992).

Lead Acetate for Hydrogen Sulfide Detection

Some organisms are capable of enzymatically liberating sulfur from sulfur-containing amino acids or inorganic sulfur compounds. The released hydrogen sulfide reacts with lead acetate to yield lead sulfide, an insoluble black precipitate. Lead acetate is the most sensitive H2S indicator reagent and is useful with organisms that produce trace amounts of H2S, especially organisms that are not in the family Enterobacteriaceae. Inoculate a TSI medium with the isolate (stab once through the center of the butt of the tube to within 3 to 5 mm of the bottom, withdraw the inoculating needle, and streak the surface of the TSI agar slant). Place the lead acetate strip so that it hangs down approximately 1" inside the TSI tube. Incubate at 35°C in non-CO2 for 18 to 24 h. A brownish-black coloration of the paper strip indicates a positive reaction. No coloration of the strip indicates a negative reaction.

Lead acetate is toxic to bacterial growth. Do not allow the strip to touch the medium. The TSI medium must support the growth of the test organism for H2S production to occur (Koneman et al., 1997; Murray et al., 2003).

Lysine Iron Agar (LIA)

Lysine iron agar is a differential medium used for the identification of enteric bacilli based on their ability to decarboxylate or deaminate lysine and produce hydrogen sulfide. Dextrose serves as a source of fermentable carbohydrate. The pH indicator, bromcresol purple, is changed to a yellow color at or below pH 5.2 and is purple at or above pH 6.8. Ferric ammonium citrate and sodium thio sulfate are indicators of hydrogen sulfide formation. Lysine serves as the substrate for detecting the enzymes lysine decarboxylase and lysine deaminase. Lysine iron agar is designed for use with TSI (tripe sugar iron agar) for the identification of enteric pathogens.

Using a sterile inoculating needle, stab the butt of the LIA slant twice then streak back and forth along the surface of the agar with the organism. Incubate at 35°C ± 2°C in non-CO2 for 18 to 24 h.

Alkaline (purple) reaction in the butt indicates lysine decarboxylation; red slant indicates lysine deamination, and black precipitate indicates H2S production. H2S may not be detected in this medium by organisms that are negative for lysine decarboxylase activity because acid production in the butt may suppress H2S formation. For this reason, H2S producing Proteus species do not blacken this medium (BD Microbiology Systems, 1992).

Triple Sugar Iron (TSI) Agar Slant

TSI agar is a medium that differentiates Gram-negative bacilli on the basis of the ability to ferment carbohydrates and liberate hydrogen sulfide (H2S). The medium contains 1 part glucose to 10 parts each of lactose and sucrose. Phenol red serves as an indicator to detect pH change, and ferrous sulfate detects the formation of H2S. If the organism ferments glucose, the butt and slant of the agar will become acidic and turn yellow. If the organism ferments lactose and/or sucrose, the slant will remain acidic (yellow). If the organism is unable to ferment lactose or sucrose, the slant will revert to alkaline (red) when the glucose is used up and alkaline amines are produced in the oxidative decarboxylation of peptides (derived from protein in the medium) near the surface of the agar. Organisms unable to ferment glucose will not change the pH of the medium or will produce alkaline products, and the TSI tube will remain red. Blackening of the medium indicates H2S production. Gas production is indicated by splits or cracks in the butt of the agar. Gas may also push the agar up the tube.

Using a sterile inoculating needle, stab the butt of the LIA slant twice then streak back and forth along the surface of the agar with the organism. Incubate at 35°C ± 2°C in non-CO2 for 18 to 24 h. If acid slant-acid butt (yellow-yellow): glucose and sucrose and/or lactose fermented. If alkaline slant-acid butt (red-yellow): glucose fermented only. If alkaline slant-alkaline butt (red-red): glucose not fermented. The presence of black precipitate (butt) indicates hydrogen sulfide production, and presence of splits or cracks with air bubbles indicates gas production.

Early readings may result in false acid-acid results, and delayed readings may result in false alkaline-alkaline results. Copious amounts of H2S may mask the glucose reaction. If this occurs, glucose has been fermented even if it is not observable. The utilization of sucrose may suppress the enzyme mechanism that results in the production of H2S. Trace amounts of H2S may not be detectable with the ferrous sulfate indicator in the agar (BD Microbiology Systems, 1992; Koneman et al., 1997).

Phenylalanine Deaminase

This assay is used to detect the ability of an organism to oxidatively deaminate phenylalanine to phenylpyrovic acid. The phenylpyrovic acid is detected by adding a few drops of 10% ferric chloride.

Inoculate a phenylalanine agar slant with the organism and incubate at 35°C in non-CO2 incubator for 18-24 h. Following the incubation, add 4-5 drops of 10% ferric chloride solution to the slant. The development of green color on the surface of the slant indicates a positive reaction. No color change indicates a negative reaction (Isenberg, 1992; Murray et al., 2003).

Decarboxylase

Decarboxylases are a group of substrate-specific enzymes that are capable of reacting with the carboxyl (COOH) portion of amino acids, forming alkaline-reacting amines. Each decarboxylase enzyme is specific for an amino acid. Lysine, or-nithine, and arginine are the three amino acids used routinely in the identification of Enterobacteriaceae, Aeromonas, Plesiomonas, and Vibrio species. The decar-boxylation of lysine and ornithine yield cadaverine and putrescine, respectively. Arginine is converted to citrulline by a dihydrolase reaction. A control tube containing the base without an added amino acid to verify that the organism utilizes glucose must accompany all decarboxylase tests. Because decarboxylation is an anaerobic reaction, it must be overlaid with mineral oil prior to incubation. If the organism is viable, both the control and the test tube with amino acid should turn yellow because of fermentation of the small amount of glucose in the medium. If the amino acid is decarboxylated, the alkaline amines cause the indicator (brom-cresol purple) in the acid medium to revert back to its original purple color.

Inoculate a Moeller decarboxylase broth containing ornithine, lysine, and/or arginine. Overlay the contents of all tubes with 1 mL of sterile mineral oil and incubate in a non-CO2 incubator at 35°C for 18-24 h. Examine for a color change. Negative reactions are examined daily for no more than 4 days (BD Microbiology Systems, 1992).

OF Glucose Medium

Bacteria can utilize glucose and other carbohydrates by using various metabolic cascades. Some are fermentative routes; others are oxidative. Oxidationfermentation (OF) medium permits classification of organisms by a simple method that differentiates aerobic and anaerobic degradation of carbohydrates. The low protein to carbohydrate ratio in the medium prevents neutralization of acids by the alkaline products of protein metabolism, thus allowing small quantities of weak acids to be detected. Acid production results in a pH shift that changes the color of the bromthymol blue indicator from green to yellow.

Using an inoculating needle, 2 tubes of OF glucose medium are stab-inoculated halfway to the bottom of the tubes. The content of one tube is overlaid with 1 mL of sterile mineral oil. Both tubes are incubated at 35°C in non-CO2, and examine daily for 72 h or longer for slow-growing organisms. Yellow color indicates the production of acid. Acid production in the tube without oil overlay is considered oxidative reaction. Acid production in both tubes is considered fermentative. No acid production in either tube is considered nonsaccharolytic. Nonsaccharolytic organisms produce slight alkalinity (blue-green color) in the tube without oil overlay, but the tube with oil will not exhibit a color change and will remain green (BD Microbiology Systems, 1992).

OF Sugars

OF basal medium, when supplemented with an appropriate carbohydrate, is used to determine an organism's ability to utilize sugars such as lactose, xylose, sucrose, maltose, and mannitol. The low protein to carbohydrate ratio in OF basal medium prevents the neutralization of small quantities of weak acids by the alkaline products of protein metabolism, which makes this medium ideal for determining carbohydrate utilization. Acid production from carbohydrate metabolism results in a pH shift that changes the color of the bromthymol blue indicator from green to yellow. Yellow color indicates carbohydrate metabolism.

Using an inoculating needle, touch the center of one colony and stab-inoculate the OF medium with the appropriate carbohydrate once halfway to the bottom of the tube. Cap the tubes loosely and Incubate at 35°C in non-CO2, and examine daily for 72 h or longer for slow-growing organisms. A yellow color indicates carbohydrate utilization and no color change (green) or blue color indicates no carbohydrate utilization. The acid reaction produced by oxidative organisms is detected first at the surface and gradually extends throughout the medium. When oxidation is weak or slow, it is common to observe an initial alkaline reaction at the surface of the tube that may persist for several days. This must not be mistaken for a negative test. If the organism is unable to grow in the OF medium, add either 2% serum or 0.1% yeast extract prior to inoculation (BD Microbiology Systems, 1992).

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Atkins Low Carb Diet Recipes

Atkins Low Carb Diet Recipes

The Atkins Diet is here. Dr Atkins is known for his great low carb diets. Excluding, Dr Atkins carb counter and Dr Atkins New Diet Revolution.

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