REAGENTSBack to top
A number of classical and rapid tests are used for
the identification of medically important
bacteria. Below are brief descriptions of commonly
performed tests and reagents used in
clinical microbiology. See references 5,13,
27, and 38 for more detailed
descriptions of these
tests and the reagents they use.
Biochemical Tests
Acetamide hydrolysis test (Nessler reagent)
Nessler reagent is used in the determination of
acetamide hydrolysis. This test is useful in
differentiating some gram-negative bacteria.
Acetamide agar or broth is inoculated. After
incubation at 35 to 37°C until colonies or
turbidity develops, 1 drop of Nessler reagent is
added to 1 ml of broth or directly to the plate. A
positive reaction is indicated by the
formation of a red-brown sediment. Nessler reagent
is prepared by dissolving 1 g of mercuric
chloride in 6 ml of distilled water and then
adding 2 or 3 drops of concentrated hydrochloric
acid to dissolve the sediment. Separately, 2.5 g
of potassium iodide is dissolved in 6 ml of
distilled water and then added to the mercuric
chloride solution. Then 6 g of potassium
hydroxide is dissolved in 6 ml of distilled water
and added to the mercuric chloride-iodide
solution along with an additional 13 ml of
distilled water. The solution is filtered using a
sintered glass funnel (not a Nalgene filter). The
Nessler reagent is stored in the dark and
should be useful for several weeks. The solution
should be checked for decomposition prior
to use (any color change other than yellow
indicates decomposition, and a fresh solution
should be prepared). Nessler reagent is toxic if
swallowed, inhaled, or absorbed through the
skin. It presents a neurological hazard, may act
as a carcinogen, and may be a reproductive
hazard. It is corrosive and causes burns.
Alkaline phosphatase
Alkaline phosphatase is detected by the hydrolysis
of a colorless phosphate-containing
compound to a colored product, e.g., p-nitrophenol
phosphate to p-nitrophenol, which is
yellow; phenolphthalein phosphate to
phenolthalein, which is red under alkaline conditions;
or indolyl phosphate to indigo, which is blue.
This test is useful in the differentiation
of Staphylococcus species and
non-glucose-fermenting gram-negative rods, and it is
incorporated in several commercial identification
systems.
Arginine arylamidase
(L-arginine-4-methoxy-β-naphthylamide)
Arginine arylamidase (trypsin) is included as a
diagnostic test in some commercial systems,
e.g., the API test system of bioMerieux. The
substrate for this test is L-arginine-4-methoxy-
β-naphthylamide. A negative test is colorless, and
a positive test produces an orange color.
This test is useful in differentiating
staphylococcal species and various other bacteria.
Benzidine test (benzidine hydrochloride)
The benzidine test is useful for differentiating
coagulase-negative Staphylococcus species.
This test is included in several commercial
systems. It is based upon the presence of ironporphyrin
compounds. Addition of a solution of 1 g of
benzidine hydrochloride dissolved in 20
ml of glacial acetic acid, 30 ml of water, and 50
ml of 95% ethanol followed by addition of a
5% solution of hydrogen peroxide (H2O2) results in
the formation of a blue-green to deep
blue color for positive organisms.
Bile solubility test (deoxycholate)
The bile solubility test is used in the
presumptive identification of Streptococcus
pneumoniae. The key reagent in this test is sodium deoxycholate, which is a
surface-active
bile salt. The test may be run in a tube or on
agar plates. The test is performed on alphahemolytic
streptococcal colonies. A few drops of a 10%
solution of sodium deoxycholate can
be applied directly to the surface of a colony.
The plate is then incubated for 30 min at 35°C.
Pneumococcal colonies are lysed, whereas viridans
group streptococci are not lysed.
Alternatively, a heavy suspension of cells can be
added to physiological saline solution (pH
7.0) and divided into two tubes. The 10% sodium
deoxycholate solution is added to one
tube, and sterile physiological saline is added to
the other. The tubes are incubated at 35°C
and are visually compared. If the organism is bile
soluble, the tube containing the
deoxycholate will exhibit reduced turbidity within
15 min and show an increase in viscosity
along with clearing of the solution.
CAMP test (beta-lysin)
The CAMP factor test is used to identify group B
beta- hemolytic streptococci based on their
formation of a substance (CAMP factor) that
enlarges the area of hemolysis formed by betahemolysin.
Hardy Diagnostics CAMP Spot Test Reagent is used
as a rapid CAMP test method.
The reagent, containing staphylococcal beta-lysin
(also called beta-toxin, beta-hemolysin, or
beta-staphylolysin), acts directly with the CAMP
factor that is diffused into the medium
around the suspect colony. The beta-lysin has a
synergistic effect in the presence of CAMP
factor, producing enhanced hemolysis of sheep
erythrocytes. Enhanced hemolysis is visible
within 30 min to 1 h of placing a drop of CAMP
Spot Test Reagent next to an isolated betahemolytic
Streptococcuscolony.
Catalase test (H2O2)
H2O2 is used to detect bacterial production of
catalase. A concentration of 15% is used for
the differentiation of anaerobes, which do not
produce catalase. A 30% peroxide
concentration is used to test Neisseria species.
Cells from a colony are transferred to a clean
glass slide, and a drop of hydrogen peroxide is
added. Production of bubbles indicates a
positive reaction. Blood must be avoided, as
erythrocytes produce catalase and can cause a
false-positive reaction. It is also possible to
add a drop of hydrogen peroxide directly to a
colony or slant as long as the medium does not
contain blood. Immediate bubbling indicates
a positive reaction.
Coagulase test (rabbit plasma)
Dehydrated rabbit plasma with EDTA is used to
detect free or bound (clumping factor)
coagulase produced byStaphylococcus species.
Human plasma is preferred for the detection
of bound coagulase produced byStaphylococcus
lugdunensis and Staphylococcus
schleiferi but is not routinely used because it may contain antibodies
against staphylococci. A
heavy suspension of cells is added to a clean
glass slide and mixed with a drop of distilled
water. If agglutination does not occur
spontaneously, the procedure can be performed by
adding a drop of rabbit plasma to the suspension
and mixing with a circular motion. The
formation of visible white clumps indicates the
presence of bound coagulase. Positive and
negative controls should be run. The test can also
be run in a test tube which detects both
free and bound coagulase. For this test, 0.5 ml of
rabbit plasma is added to a sterile tube.
The tube is inoculated with a loopful of the test
organism and incubated at 35°C for 4 h.
Observations for clotting should be made within
the first 4 h since some staphylococci
produce fibrolysin, which can dissolve the clot.
If no clotting is observed, however, the tube
should be incubated overnight at room temperature
and again observed for delayed clotting.
Decarboxylase tests (Moeller broth, bromcresol
purple)
Moeller broth, which contains bromcresol purple
and cresol red, is used to detect the pH
change due to decarboxylation of either of the
amino acids lysine and ornithine.
Decarboxylase tests are useful for differentiating
the Enterobacteriaceae. The broth at
neutral or slightly acidic pH containing an
individual amino acid being tested is inoculated for
at least 24 h in most cases. The test may also be
run after growth on other broths by adding
a solution of bromcresol purple to a drop of the
medium to determine if the pH is alkaline.
Moeller broth can be used to this purpose. A rapid
test has been described omitting glucose
from the medium and using a starting pH of 5.5 (17).
In a positive result, the increased pH is
indicated by a change in color of bromcresol
purple from yellow to purple.
Esculin hydrolysis (ferric ammonium citrate)
The hydrolysis of esculin to esculetin is detected
using a 1% solution of ferric ammonium
citrate. After incubation in esculin-containing
medium for 1 to 2 days, a few drops of ferric
ammonium citrate is added. The immediate formation
of a brown-black color indicates a
positive reaction. Esculin hydrolysis can also be
determined using esculin agar without bile,
which contains iron; using this preferred medium,
esculin hydrolysis is indicated by
blackening after overnight incubation.
β-Galactosidase (o-nitrophenyl-β-D-galactopyranoside
[ONPG])
ONPG at a concentration of 4 mg/ml is used to
detect β-galactosidase activity. This enzyme
facilitates growth on a carbon source like lactose
by cleaving it into a molecule of glucose
and a molecule of galactose which the cells can
catabolize and on which the cells can grow.
The substrate ONPG is used in place of lactose.
When the β-galactosidase cleaves ONPG, onitrophenol
is released. This compound has a yellow color.
This test is especially useful for
identification of members of the family Enterobacteriaceae.
ONPG-impregnated tablets can
be used for this test. In API ZYM,
2-naphthyl-β-galactopyranoside is used as the substrate.
Gelatin hydrolysis (gelatin)
Gelatin hydrolysis, sometimes referred to as
gelatin liquefaction, is performed to determine
the presence of the proteolytic enzyme gelatinase,
which liquefies/hydrolyzes gelatin.
Following inoculation of semisolid nutrient
gelatin medium with the test organism, the
medium, which is c ommercially available, is
incubated at 35 to 37°C for up to 1 week,
depending on the organism being tested. The
culture is then placed at 4°C for a minimum of
15 min. A positive test is denoted by the
observation of a completely liquid medium
indicative of the hydrolysis (liquefaction) of the
gelatin; in a negative test the medium is
solid at 4°C.
Gelatin hydrolysis can also be assessed by using a
plate method, which tends to give
superior results for gram-negative nonfermenting
bacteria. A plate with nutrient agar plus
0.4% gelatin is inoculated with a spot or a streak
and incubated until luxuriant growth is
obtained. When the isolate is gelatin hydrolysis
positive, visual clearing of the agar is usually
obvious. If not, a HgCl2 solution (12 g of HgCl2,
16 ml of 35% HCl, and 80 ml of distilled
water) can be added to enhance the observation of
clearing. Because of the high toxicity of
HgCl2, it often is better to use the test tube
method; it also is possible to replace it with
sulfosalicylic acid to avoid exposure to mercuric
chloride.
β-Glucuronidase ( p-nitrophenyl-β-D-glucopyranoside,
4-
methylumbelliferyl-β-D-glucuronide [MUG])
Detection of β-glucuronidase activity can be
accomplished using either a colorimetric
substrate (p-nitrophenyl-β-D-glucopyranoside)
or a fluorometric substrate (MUG). This test is
useful for the rapid identification ofEscherichia
coli, members of the Streptococcus
anginosus group, and other bacteria. For the colorimetric test, a solution
of 0.1% (wt/vol) pnitrophenyl-
β-D-glucopyranoside (colorimetric substrate) in
0.067 M Sorensen phosphate
buffer (pH 8.0) is prepared. Tubes containing 0.5
ml of the substrate solution are inoculated
with a loopful of bacteria from an overnight
culture. The tubes are incubated at 35°C and
examined after 4 h for the appearance of a yellow
color (liberated p-nitrophenol). In the
fluorometric test the substrate MUG yields the
product 4-methylumbelliferyl, which
fluoresces blue under long-wave UV light. The MUG
test is normally used for the presumptive
identification of E. coli and more recently
for streptococcal strains. To prepare MUG for the
fluorescent test, dissolve 50 mg of MUG in 10 ml
of 0.05 M Sorensen phosphate buffer, pH
7.5. Dilute 1:16 of the stock MUG and add 1.25 ml
to a vial containing 50 sterile paper disks.
Allow the disks to be thoroughly saturated until
no liquid remains in the vial. Spread the
saturated disks out and allow to dry completely.
The disks can be stored in a dark bottle at
−20°C for 1 year or at 4°C for 1 month. Wet the
disk with 1 drop of sterile water. Apply the
organism to the disk using a wooden stick or loop
and then incubate the disk for up to 2 h at
35°C. Shine a long-wave UV light on the disk. A
positive reaction is indicated by blue
fluorescence.
Hippurate hydrolysis (ninhydrin reagent) (ferric
chloride)
Hippurate hydrolysis to benzoic acid and glycine
is useful in the identification of group B
streptococci (GBS), some Listeria spp., Gardnerella
vaginalis, Campylobacter
jejuni, and Legionella pneumophila. Ninhydrin reagent can be used
to detect the production
of glycine. Ninhydrin reagent (3.5%) is prepared
by adding 3.5 g of ninhydrin to 50.0 ml of
acetone and 50.0 ml of 1-butanol. The ninhydrin
reagent is stored in the dark at room
temperature. A 1% (wt/vol) solution of sodium
hippurate is prepared in 0.067 M Sorensen
phosphate buffer (pH 6.4). Tubes containing 0.5 ml
of this solution are inoculated and
incubated at 35°C for 2 h, after which 0.2 ml of
the ninhydrin reagent is added. Development
of a deep blue-purple color within 5 min indicates
a positive reaction. For L.
pneumophila, 0.5 ml of 1% sodium hippurate solution is inoculated with a
loopful of
organism and incubated at 35°C in ambient air for
18 to 20 h, after which 0.2 ml of
ninhydrin reagent is added. The cells and
ninhydrin are mixed and incubated for an
additional 10 min at 35°C. The mixture is observed
for 20 min for blue-purple color
development, which is indicative of a positive
reaction. Ferric chloride can also be used to
detect hippurate hydrolysis. Ferric chloride
reagent (12 g of FeCl3 6H2O in 100.0 ml of 2%
HCl) is added to inoculated broth (e.g., heart
infusion broth or Todd-Hewitt broth)
supplemented with hippurate. An insoluble brown
ferric benzoate precipitate indicates a
positive hydrolysis reaction.
Indole test (Ehrlich reagent, Kovacs reagent, pdimethylaminocinnamaldehyde
[DMACA])
The indole test is used for the determination of
production of indole from deamination of
tryptophan by tryptophanase. This reaction can be
detected using Ehrlich reagent, Kovacs
reagent, or dimethylaminocinnamaldehyde (DMACA).
Kovacs reagent is added directly to the
medium; an extraction phase using xylene is
required before adding Ehrlich reagent. To
prepare Ehrlich reagent, add 1 g of p-dimethylaminobenzaldehyde
to 95 ml of 95% ethyl
alcohol. Then slowly add 10 ml of concentrated
hydrochloric acid. Using Ehrlich reagent, first
extract the indole by adding 1 ml of xylene to a
48-h-old tryptone broth or other tryptophancontaining
broth medium. Shake the tube vigorously for 20 s
and let stand for 1 to 2 min to
allow the xylene extract to come to the top of the
broth. Gently add 0.5 ml of the Ehrlich
reagent down the side of the tube. Do not shake
the tube. A red ring at the interface of the
medium and the reagent phase within 5 min
represents a positive test. Ehrlich reagent is
preferred for organisms that produce small amounts
of indole, such as nonfermenters and
anaerobes. To prepare Kovacs indole reagent, add
10 g of p-dimethylaminobenzaldehyde to
150 ml of either amyl or isoamyl alcohol. Then add
50 ml of concentrated hydrochloric acid.
Add 5 drops of Kovacs reagent to either 48-h-old
2% tryptone broth or an 18- to 24-h-old
tryptophan broth culture. Do not shake the tube
after the addition of reagent. A red color at
the surface of the medium indicates a positive
test. For the spot indole test, add 2 ml of
concentrated HCl to 18 ml of distilled water.
Allow the mixture to cool. Then add 200 mg of
DMACA. Moisten a piece of Whatman no. 3 paper with
a couple of drops of the reagent.
Remove a well-isolated colony from an 18- to
24-h-old culture onto a blood agar plate with a
sterile inoculating loop or a wooden stick and
smear it onto the moistened filter paper.
Observe for a blue to blue-green color within 2
min, which indicates a positive reaction. No
color change or a pinkish tinge is considered
negative. This test should be used only on
colonies from media containing sufficient
tryptophan and no glucose (blood agar). Colonies
from media containing dyes (e.g., MacConkey or
eosin-methylene blue [EMB] agar) may
cause misleading results and should not be used.
Colonies from mixed cultures should not be
used, as indole-positive colonies can cause
indole-negative colonies to appear weakly
positive. The test can also be run using a heavy
bacterial suspension in 0.3% tryptophan
solution and revelation with Kovacs reagent after
4 h.
LAP test (leucine naphthylamide)
The LAP test detects the presence of leucine
aminopeptidase (LAP). The substrate leucine
naphthylamide is hydrolyzed by LAP to leucine and
free naphthylamine. The LAP test is
helpful in the presumptive characterization of
catalase-negative, gram-positive cocci
(streptococci, enterococci, and streptococcus-like
organisms). S. pneumoniae, Streptococcus
pyogenes, Pediococcus, Lactococcus, and Enterococcus species are all LAP
positive, while
other beta-hemolytic streptococci are LAP
negative. Disks are impregnated with leucine-β-
naphthylamide or leucine-α-naphthylamide, which is
hydrolyzed by the enzyme LAP,
produced by LAP-positive organisms. This test is
performed by inoculating several colonies
from overnight growth of the test organism to a
moistened LAP disk aseptically placed in a
sterile petri disk at room temperature. One drop
of DMACA reagent is added. After 1 min,
enzymatic activity results in the release of
β-naphthylamine, which couples with DMACA
reagent to form a highly visible red color
indicating a positive test.
Lysozyme test (lysozyme)
The lysozyme test measures the ability of
organisms, such as Nocardia, to grow in the
presence of lysozyme. A solution of 50 mg of
lysozyme in 50 ml of 0.01 N HCl is used for this
test. The solution is filter sterilized and can be
stored refrigerated for up to a week. For the
lysozyme test, add 5 ml of lysozyme solution to 95
ml of basal glycerol broth (peptone, 1 g;
beef extract, 0.6 g; glycerol, 14.0 ml; distilled
water, 200 ml). Dispense in 5-ml aliquots and
keep refrigerated. Growth of the test organism in
the lysozyme-supplemented glycerol broth
is compared with growth in the unsupplemented
glycerol broth.
Nitrate reduction test (N,N-dimethyl-naphthylamine
and sulfanilic acid)
The nitrate reduction test is used to determine
the ability to reduce nitrate to nitrite or free
nitrogen gas. This test involves the use of two
Griess reagents. Griess reagent A (0.8 g of
sulfanilic acid in 100 ml of 5 N [i.e., 30%]
acetic acid) reacts with nitrite to produce
diazonium salt, which, after addition of Griess
reagent B (0.5 g of α-naphthylamine or 0.6 g
of N,N-dimethyl-naphthylamine in 100 ml of
5 N acetic acid), will react to produceparasulfobenzene-
azo-naphthylamine (prontosil), the red end product
of this reaction. The
reagents may be stored in the dark under
refrigeration. To perform the test, add 0.05 ml of
reagent A to 10 drops of an overnight growth from
the nitrate broth culture and incubate for
5 to 10 min. Then add 0.05 ml of reagent B and
incubate for an additional 5 to 10 min.
Incubation should be in the dark. (Note: reagents
A and B may be mixed and added together
as indicated in the previous edition of this Manual,
but this lowers the sensitivity of the test
since Griess reagent B reacts with the product
formed by the reaction of Griess reagent A
with nitrite.) An organism may be reported as
nitrate positive if a red or purple-magenta
color develops in the medium within a few minutes
after nitrate reagents A and B are added
to the medium, indicating that the organism has
reduced nitrate to nitrite. The absence of a
red-purple color after the addition of both
reagents does not automatically mean that the
organism is unable to reduce nitrate. Strains may
have reduced the nitrate to nitrite and
then reduced the nitrite completely to nitrogenous
gases which are not detected when nitrate
reagents A and B are added to the medium. If the
medium does not change color after the
addition of sulfanilic acid and α-naphthylamine, a
small amount (“knife point”) of zinc dust is
added to the incubated medium. The zinc dust will
catalyze the reduction of nitrate to nitrite
chemically. Thus, if the nitrate has not been
reduced by the organisms, i.e., they are nitrate
negative, it will be reduced by the zinc dust and
a red color will develop in the incubated
medium within 15 min. If no color develops in the
incubated medium after the addition of
zinc dust, the organisms not only have reduced
nitrate to nitrite but also have reduced nitrite
to nitrogenous gases; these organisms are also
nitrate positive. See Table 1 for nitrate and
nitrite reduction reactions.
Oxidase test (TMPD/DMPD)
The oxidase test is a test used in microbiology to
determine if a bacterium produces certain
cytochrome oxidases (16).
It uses disks impregnated with a reagent such as N,N,N′,N′-
tetramethyl-p-phenylenediamine
dihydrochloride (TMPD) or N,N-dimethyl-pphenylenediamine
dihydrochloride (DMPD), which is also a redox
indicator. TMPD is more
sensitive than DMPD and therefore generally the
preferred reagent. The reagent is a dark
blue to maroon color when oxidized and colorless
when reduced. A modified oxidase test is
used for the differentiation of Micrococcus and
related organisms from most other aerobic
gram-positive cocci. Six percent TMPD (the same
chemical used in Kovacs oxidase reagent)
dissolved in dimethyl sulfoxide is used as the
reagent. Keep the reagent away from light
because light degrades it. Commercially available
strips (Merck) containing the dimethyl
compound are much more stable. A loopful of
colonies from blood agar plates is smeared
onto filter paper, and the reagent is dropped onto
the bacterial growth. Development of a
blue to purple-blue color in 2 min indicates a
positive reaction.
Phenylalanine deaminase test (ferric chloride)
Phenylalanine deaminase activity can be determined
on 1% of DL-phenylalanine agar media
or agar slants, which are flooded with a 12% FeCl3
solution in 2% HCl after 1 to 2 days of
incubation. The hydrochloric acid is prepared by
adding 5.4 ml of concentrated HCl (37%) to
94.6 ml of distilled water. To perform the
phenylalanine deaminase test, 4 or 5 drops of
ferric chloride reagent are added to a culture
grown overnight on phenylalanine agar or
broth. The development of a green to brown color,
due to the reaction of phenylpyruvic acid
with Fe in the medium or on the slant, indicates a
positive reaction.
Pyrrolidonyl aminopeptidase activity (PYR test)
Pyrrolidonyl aminopeptidase (pyrrolidonyl
arylamidase) or PYR is a rapid colorimetric method
for presumptive identification of certain groups
of bacteria based on the activity of the
enzyme pyrrolidonyl arylamidase. This test is used
in the identification of gram-positive cocci
and nonfermentative gram-negative bacteria. The
reaction involves addition of DMACA,
which can be suspended in a solution of 2.5 ml of
sodium dodecyl sulfate, 2.5 ml of glacial
acetic acid, 5.0 ml of 2-methoxyethanol, and 90 ml
of distilled water (stored at 4°C in a dark
container). There also is a commercial kit in
which L-pyroglutamic acid β-naphthylamide is
impregnated into the test disk and serves as the
substrate for the detection of pyrrolidonyl
arylamidase. Hydrolysis of the substrate yields
β-naphthylamide, which combines with the
PYR reagent (DMACA) to form a bright pink to
cherry red color. A positive PYR tests allows
for the presumptive identification of group A
streptococci (Streptococcus pyogenes) and
group D enterococci.
Tributyrin esterase (tributyrate glycerol,
bromo-chloro-indolyl
butyrate)
Tributyrin esterase activity is used in the
differentiation of nonfermenting gram-negative
bacteria and for the identifcation of Moraxella
catarrhalis. Tributyrin esterase activity can be
detected using disks containing tributyrate
glycerol and phenol red, which are available from
Rosco, or strips (TRIBU strips) that are available
from Sigma. Tributyrin esterase activity
frees butyric acid, resulting in the formation of
a yellow color due to acidification. Tributyrin
esterase can also be detected using disks
impregnated with bromo-chloro-indolyl butyrate
(CatScreen) from Hardy Diagnostics. Hydrolysis of
this substrate by the butyrate esterase
yields a chromogenic compound which appears blue
to blue-violet. For this test a heavy
inoculum from a 24- to 72-h-old culture is smeared
onto a disk that has been wet with sterile
distilled water and incubated for 5 min. Longer
incubation can yield false positives.
Tween 80 (polysorbitol) hydrolysis (Tween 80)
The formation of a precipitate around colonies
after growth on Trypticase soy agar containing
1% Tween 80 and 0.01% calcium chloride indicates
Tween 80 hydrolysis due to esterase
activity. This method is used for differentiation
of nonfermenting gram-negative bacteria and
identification of Moraxella catarrhalis.
Esterase activity can also be detected using a medium
containing Tween 80 and neutral red; the neutral
red binds to the Tween 80, producing an
amber color. When Tween 80 is hydrolyzed by
esterase activity, a red color develops due to
the release of oleic acid. This method has been
used for identifcation of mycobacteria.
Urease test (phenol red)
The urease test is used to determine the ability
of an organism to split urea through the
production of the enzyme urease. Ammonia is
produced, which causes a rise in pH that is
detected by a change in color of the indicator
phenol red to pink under alkaline conditions
(pH 8.4). Bacteria are cultured on a medium
containing urea, e.g., Christensen urea agar.
While many enteric bacteria can hydrolyze urea,
only a few “rapid urease-positive”
organisms, e.g., Proteus species, can
degrade urea quickly (less than 4 h). Urea broth is
formulated to test for rapid urease-positive
organisms. The restrictive amount of nutrients,
coupled with the use of pH buffers, prevents all
but rapid urease-positive organisms from
producing enough ammonia to turn phenol red to
pink. The rapid urease test also is used for
the diagnosis of Helicobacter pylori. To
detect H. pylori, this test is performed on stomach
lining cells collected by biopsy at the time of
endoscopy. A basic broth for performing the
urease test can be made by adding 10.4 ml of a 20%
(wt/vol) aqueous solution of urea to a
solution containing 0.1 g of KH2PO4, 0.1 g of
K2HPO4, and 0.5 ml of 1:500 phenol red,
adjusted to pH 6.8 in 100 ml. To make 1:500 phenol
red, dissolve 0.2 g of phenol red in
NaOH and add distilled water to 100 ml. This
solution not only is easier to prepare than
Christensen agar but also is more sensitive for
assessing urease activity by nonfermenters
when a dense inoculum is used. Red color
developing within 4 h after inoculation indicates
urease activity.
Voges-Proskauer (VP) test (α-naphthol/KOH)
The VP test is used to detect acetoin
(acetyl-methylcarbinol), which is produced by certain
bacteria during growth in a buffered
peptone-glucose broth (methyl red VP [MR-VP] broth).
The VP test is commonly used to aid in
differentiation between genera (such as E. coli from
the Klebsiella and Enterobacter species)
and among species of the Enterobacteriaceae family.
The test can be used as a differential test for
other organism groups (viridans group
streptococci). The test uses 5%α-naphthol, which
is prepared by dissolving 5 g of α-naphthol
in 100 ml of absolute ethanol, and 40% KOH, which
is prepared by dissolving 40 g of
potassium hydroxide in 100 ml of distilled water.
To perform the test, MR-VP broth is
inoculated and incubated until good growth is
obtained. Then 0.6 ml of the α-naphthol
solution and 0.2 ml of the 40% KOH are added to
2.5 ml of culture broth. A positive reaction
is indicated by the formation of a pink-red
product within 5 min. However, allow 15 min for
color development before considering the test
negative.
Buffers
Bovine albumin fraction V
A 0.2% solution of bovine albumin fraction V is
used to buffer mycobacterial specimens
following decontamination with N-acetyl-L-cysteine-sodium
hydroxide (NALC-NaOH). The
solution is prepared by mixing 40.0 ml of 5%
bovine albumin with 8.5 g of NaCl and 960.0
ml of distilled water. The pH is adjusted to 6.8
using 4% NaOH. The solution is filter
sterilized and stored refrigerated. Before
addition to the buffer, samples are decontaminated
with NALC-NaOH and concentrated by centrifugation.
The sedimented sample is suspended
in 1 to 2 ml of the sterile 0.2% bovine albumin
solution. The preserved cells can then be
examined microscopically or inoculated into a
culture medium.
Glycine-buffered saline
Glycine-buffered saline (0.043 M glycine, 0.15 M
NaCl [pH 9.0]) is used in some serological
procedures and is also used as a transport medium
for enteric organisms. It is prepared by
dissolving 3.22 g of glycine and 8.77 g of NaCl in
1 liter of distilled water.
Phosphate-buffered saline
Phosphate-buffered saline solutions are made by
mixing various amounts of 0.1 N monoand
dibasic phosphates, depending upon the pH desired,
with 0.85% NaCl. These are
prepared as 10× stock solutions. For 0.1 M NaH2PO4
(sodium phosphate, monobasic),
dissolve 13.9 g of NaH2PO4 in 1 liter of deionized
water; for 0.1 M Na2HPO4 (sodium
phosphate, dibasic), dissolve 26.8 g of Na2HPO4・7H2O in 1 liter of
deionized water; and for
8.5% NaCl (sodium chloride), dissolve 85.0 g of NaCl
in 1 liter of deionized water. Sterilize
by autoclaving for 20 min or by filtration. Store
refrigerated. For the working phosphatebuffered
saline, combine the appropriate amounts of the 10×
stock solutions of the monoand
dibasic phosphate solutions that are combined with
100 ml of 8.5% NaCl and bring the
volume to 1 liter with distilled or deionized
water. See the ninth edition of this Manual (8a)
for the appropriate amounts of the mono- and
dibasic phosphate solutions needed to achieve
specific pH values.
Sorensen pH buffers
Sorensen pH buffers are prepared by mixing
appropriate amounts of 0.067 M dibasic sodium
phosphate and 0.067 M monobasic potassium
phosphate. To prepare 0.067 M dibasic sodium
phosphate, dissolve 9.464 g of anhydrous Na2HPO4
in 1 liter of distilled water. To prepare
0.067 M monobasic potassium phosphate, dissolve
9.073 g of anhydrous KH2PO4 in 1 liter of
distilled water. See the ninth edition of this Manual
(8a) for appropriate amounts of dibasic
and monobasic phosphate solutions to achieve
specific pH values.
Decontamination Agents
NALC-NaOH
NALC (mucolytic agent)-NaOH (decontamination
agent) is used in the processing of
mycobacterial specimens. The reagent consists of
50.0 ml of sterile 4% NaOH, 50.0 ml of
2.9% sodium citrate, and 0.5 g of NALC. The sodium
citrate is included to stabilize the
acetylcysteine. This reagent should be used within
24 h of preparation.
Cetylpuridium chloride-sodium chloride (CPC-NaCl)
CPC-NaCl is used for decontamination of
transported sputum specimens for culturing
mycobacteria. It is prepared by dissolving 1 g of
CPC and 2 g of NaCl in 100 ml of distilled
water. It can be stored in a sealed brown bottle
at room temperature. If crystals form, the
solution should be gently heated before use. An
equal amount of sputum and CPC-NaCl is
mixed until the specimen is liquefied, and then
the specimen can be shipped to the testing
site. Specimens treated with CPC-NaCl must be
cultured on egg-based media or else residual
CPC will inhibit mycobacterial growth.
Oxalic acid
Oxalic acid is used as a decontamination agent for
specimens that contain Pseudomonas spp.
when culturing for mycobacteria. The reagent is
especially helpful when processing
respiratory specimens from cystic fibrosis
patients. To prepare the solution, 50 g of oxalic
acid is added to 1.0 liter of distilled water. The
solution is autoclaved at 121°C for 15 min. It
can be stored at room temperature for up to a year.
STAINSBack to top
Microscopic examination is useful in the
identification of clinically important specimens.
Smears can be made from relevant tissues and body
fluids. If there are sufficient quantities
of cells, the smear may be prepared by direct
contact with a tissue sample or by applying a
drop of body fluid, e.g., sputum, to a clean glass
slide. Cytocentrifugation may be used to
concentrate cells (9, 20,
31). Samples are fixed to the slides with either
heat or methanol.
Methanol fixation is preferred since heating may
produce artifacts, may create aerosols, and
may not adhere the specimen adequately to the
slide. A variety of stains can then be used to
help visualize and differentiate bacteria from the
specimen. The following are some of the
commonly used staining procedures.
Acid-fast stain
Acid-fast staining is useful for the identification
of Mycobacterium, Nocardia, Rhodococcus,
Tsukamurella, Gordonia, and Legionella micdadei. These bacteria
have long-chain fatty acids
(mycolic acids) that make them difficult to stain
with crystal violet and other basic dyes.
Mycobacteria often appear as slender, slightly
curved rods and may show darker granules
that give the impression of beading. Mycobacterium
tuberculosis can appear as beaded rods
arranged in parallel strands or “cords”; Mycobacterium
kansasii may form long, often broad
and banded cells; and M ycobacterium aviumcomplex
cells appear as short, uniformly
staining coccobacilli. Nocardia spp. often
branch and almost always show a speckled
appearance. A number of staining procedures have
been developed for acid-fast staining.
In the Ziehl-Neelsen (Z-N) procedure, the slide is
heat fixed for 2 h at 70°C. The slide is then
flooded with carbol fuchsin (0.3 g of basic
fuchsin is dissolved in 10 ml of 95% ethanol, 5 ml
of phenol, and 95 ml of water; the solution is
filtered before use). The slide is slowly heated
to steaming and maintained for 3 to 5 min at 60°C.
After cooling, the slide is washed with
water and decolorized with acid-alcohol (97 ml of
95% ethanol in 3 ml of HCl). The slide is
counterstained for 20 to 30 s with methylene blue
(0.3 g of dye in 100 ml of water). An acidfast
organism will stain red, and the background of
cellular elements and other bacteria will
be blue, the color of the counterstain.
In the Kinyoun modification of the Z-N staining
procedure, heating during staining with
carbol fuchsin is eliminated and a higher
concentration of phenol is used in the p rimary
stain. The primary stain consists of 4 g of basic
fuchsin in 20 ml of 95% alcohol, 8 g of
phenol, and 100 ml of distilled water. The Z-N and
Kinyoun stains have the same sensitivity
and specificity; however, the Kinyoun (cold)
staining procedure is less time-consuming and is
easier to perform.
Another modification of the acid-fast staining
procedure has been the use of a weaker
decolorizing agent (0.5 to 1.0% sulfuric acid) in
place of the 3% acid-alcohol. This particular
stain helps differentiate those organisms known to
be partially or weakly acid-fast,
particularly Nocardia, Rhodococcus,
Tsukamurella, Gordonia, andDietzia. These organisms do
not stain well with the Z-N or Kinyoun stain.
Factors such as age, exposure to drugs, and a
particular acid-fast organism itself may vary
the acid-fast presentation. For example, while M.
tuberculosis is consistently acid fast (with
the Z-N or Kinyoun stain), rapidly growing mycobacteria
and Nocardia are not. Therefore,
use of the modified Kinyoun stain may be necessary
for these organisms. Other
modifications used in tissue preparations, such as
the Fite-Faraco stain and Pottz stain, may
be preferred for unusual isolates such as Mycobacterium
leprae.
Detection of small numbers of acid-fast organisms
in clinical specimens is generally
significant. However, the use of acid-fast stains
for gastric aspirates in the interpretation of
pulmonary disease in adults or for stool specimens
from human immunodeficiency viruspositive
patients in diagnosing Mycobacterium
avium-Mycobacterium intracellulare infection
yields very poor specificity (false-positive
smears with saprophytic organisms) as well as
poor sensitivity. In addition, patients receiving
adequate therapy may still have positive
smears without positive cultures for a number of
weeks.
Acridine orange stain
Acridine orange is a fluorochrome that can be
intercalated into nucleic acid in both the native
and the denatured states. Acridine orange is
useful in a number of miscellaneous infections,
such as Acanthamoebainfections, infectious
keratitis, and Helicobacter pylori gastritis (26).
In the acridine orange staining procedure, the
slide is flooded with acridine orange solution
(stock solution, 1 g of dye in 100 ml of water;
working solution, 0.5 ml of stock added to 5
ml of 0.2 M acetate buffer [pH 4.0]). The slide is
then examined by UV fluorescence
microscopy.
Auramine-rhodamine stain
Auramine and rhodamine are nonspecific fluorochromes
that bind to mycolic acids and that
are resistant to decolorization with acid-alcohol.
Staining procedures with these
fluorochromes are thus equivalent to the
fuchsin-based acid-fast procedures (34). Acid-fast
organisms fluoresce orange-yellow in a black
background. If the secondary stain is not used,
the organisms will fluoresce a yellow-green color.
In this procedure, the slide is heat fixed at
65°C for at least 2 h. It is then stained for 15
min with auramine-rhodamine solution (1.5 g
of auramine O, 0.75 g of rhodamine B, 75 ml of
glycerol, 10 ml of phenol, and 50 ml of H2O)
and rinsed with water, followed by decolorization
for 2 to 3 min with 0.5% HCl in 70%
ethanol. After being rinsed, the slide is
counterstained with 0.5% potassium permanganate
for 2 to 4 min. The slide is rinsed, dried, and
examined under UV fluorescence microscopy.
Gimenez stain
The Gimenez stain is used for the visualization of
Rickettsia and Coxiella from cell cultures
and L. pneumophila. Carbol fuchsin is the
primary stain, and fast green and malachite green
are the counterstains, allowing greater contrast
with the organisms and background for
easier visualization of the organisms. The stain
must be heated 48 h prior to use and filtered.
Gram stain
Gram staining is the differential staining
procedure most commonly used for microscopic
examination of bacteria. The procedure was first
described by Hans Christian Joachim Gram.
Based upon the staining reaction, bacteria are
classified as gram-positive organisms, which
retain the primary crystal violet dye and appear
deep blue or purple, and gram-negative
organisms, which can be decolorized, thereby
losing the primary stain and subsequently
taking up the counterstain safranin and appearing
red or pink. The staining reaction reflects
underlying differences in cell wall structure
which are relevant for antibiotic susceptibility as
well as identification. The Gram stain reaction
works well with most bacteria but is not useful
for bacteria that are too small or lack a cell
wall, i.e., Treponema, Mycoplasma,
Chlamydia, and Rickettsia (18). Mycobacteria are
generally not seen by Gram staining;
however, in smears illustrating heavy infections,
the organisms may give a beaded
appearance that is somewhat similar to that of Nocardia
spp. or may exhibit organism
“ghosts” (18). Anaerobic bacteria,
older cultures, and organisms that are exhibiting the
effects of antibiotics may be especially difficult
to interpret.
In the conventional Gram stain procedure used in
most clinical laboratories, the slide is first
flooded with a primary stain of crystal violet (10
g of 90% dye in 500 ml of absolute
methanol). After at least 15 s, the slide is
washed with water and flooded with the mordant
Gram’s iodine (6 g of I2 and 12 g of KI in 1,800
ml of H2O), which increases the affinity of
the primary stain to the bacterial cell. The slide
is washed with water after 15 s with the
decolorizing agent acetone-alcohol (400 ml of
acetone in 1,200 ml of 95% ethanol). The
decolorizing agent will remove the primary stain
from a gram-negative cell. Gram-positive
bacterial cells retain the primary stain. The
slide is washed immediately and counterstained
for at least 15 s with safranin (10 g of dye in 1
liter of distilled or deionized water). This slide
is then washed, blotted dry, and examined by light
microscopy at ×1,000 magnification.
Gram stain confirmation
The Gram stain reaction, which can be difficult to
properly interpret for some gram-variable
bacteria, can be confirmed using APNA K915 disks
(L-alanine-p-nitroanilide in Tris buffer)
from Key Scientific products or by using Gram-Sure
(L-alanine 7-amido-4-methycoumarin)
from Remel as reagent-impregnated disks. These
reagents detect the presence of cell wall
aminopeptidase, which is present in the cell walls
of gram-negative bacteria. Each lot of
disks should be tested prior to use with organisms
whose Gram reactions are known. A pure
colony of overnight growth is inoculated into
demineralized water and then inoculated onto
the disk. The Gram-Sure disk is incubated at room
temperature for 5 to 10 min. The APNA
K915 disk is incubated at 37°C for 5 to 20 min.
The aminopeptidase in the cell walls of gramnegative
organisms will hydrolyze the
L-alanine-7-amido-4-methycoumarin in the Gram-Sure
disk from a nonfluorescent substrate to a blue
fluorescent compound that can be observed
under long-wave UV light. Blue fluorescence is
indicative of gram-negative bacteria, and the
absence of blue fluorescence is indicative of
gram-positive bacteria. The APNA K915 disk will
yield a yellow color for a positive test for
gram-negative bacteria; no color change from the
white/cream colored disk indicates that the
organism is gram positive. Obligate anaerobes
and some microaerophiles may fail to give expected
results (29).
Immunofluorescent antibody stain
Immunofluorescent staining consists of labeling
antibodies with a fluorescent dye, allowing
the labeled antibodies to react with their
specific antigens, and observing the stained
bacterial cells under a fluorescence microscope (14).
This method allows the identification of
specific bacterial species and subtypes based upon
the specificity of the antibody reaction,
e.g., for Legionella spp. They are used in
bacteriology primarily for culture confirmation, as
other methods for direct specimen testing, such as
enzyme immunoassays and nucleic acid
amplification tests, have supplanted them.
Methylene blue stain
Staining with methylene blue is used to show
bacterial cell shape. This is useful for revealing
the morphology of fusiform bacteria and
spirochetes from oral infections (Vincent’s angina).
It may also establish the intracellular location
of microorganisms such
as Neisseria. Methylene blue is the stain
of choice for identification of the metachromatic
granules of diphtheria; however, one should be
careful about overstaining, because this will
lessen the contrast between the bacteria and the
granules. Methylene blue stains organisms
or leukocytes a deep blue against a light gray
background. Corynebacterium
diphtheriae appears as a blue bacillus with prominent darker blue
metachromatic granules.
For methylene blue staining, a 0.5 to 1.0% aqueous
solution of methylene blue is applied for
30 to 60 s and up to 10 min for possible C.
diphtheriaegranules. The slide is rinsed with
water, blotted dry, and examined by light
microscopy at magnifications of ×100 to ×1,000.
M’Fadyean stain
The M’Fadyean stain is a modification of the
methylene blue stain developed for
detecting Bacillus anthracis in clinical
specimens. The stain is prepared by dissolving 0.05 mg
of methylene blue per ml in 20 mM potassium
phosphate adjusted to pH 7.3. Slides are
stained for 1 min and then washed. As a safety
precaution, washing of the slide is performed
using a 10% hypochlorite solution. The dried slide
is examined by light microscopy.
Virulent B. anthracis rods will be
surrounded by a clearly demarcated zone giving the
appearance of a reddish pink capsule (M’Fadyean
reaction).
Spore stain
The Wirtz-Conklin spore stain is a differential
stain for detection of spores. This is very useful
for the identification of Bacillus and Clostridium
species. Using this procedure, spores stain
green while the rest of the cell stains pink.
Non-spore-forming bacteria are pink. In this
procedure the slide is flooded with 5 to 10%
aqueous malachite green. The stain is left on
the slide for 45 min. Alternatively, the slide can
be heated gently to steaming for 3 to 6 min.
Heating to steaming enhances the uptake of the
stain into the spores. The slide is then
rinsed with water. Aqueous safranin (0.5%) is used
as a counterstain for 30 s. The slide is
then washed, blotted dry, and examined by light
microscopy at ×1,000 magnification.
Wayson stain
The Wayson stain can be used to demonstrate
bipolar staining characteristics of Yersinia
pestis but is not commonly used in clinical microbiology laboratories. It
is no longer used for
screening cerebrospinal fluid for bacteria due to
the rarity of Haemophilus
influenzae infections. The staining reagents are prepared by dissolving 0.2 g
of basic fuchsin
in 10 ml of 95% ethyl alcohol and 0.75 g of
methylene blue in 10 ml of 95% ethyl alcohol.
The two solutions are added together slowly into
200 ml of 5% phenol in distilled water. The
stain is then filtered and stored in an opaque
bottle at room temperature. The stain is
applied for 1 min. The slide is then washed,
blotted dry, and examined by light microscopy at
×1,000 magnification.
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