*********** CONCENTRATION TECHNIQUES ***********
The concentration technique is effective in recovering cyst and
eggs of intestinal parasite. In the diagnosis of intestinal protozoan
infections, concentration techniques are useful in revealing light infections
in which cysts are present, but as yet no satisfactory method for concentrating
trophozoite exists.
The concentrates should be examined carefully in unstained wet
mount preparations. Iodine mounts can be prepared to stain the contents of the
cysts.
A.
Zinc Sulfate Flotation Technique
Flotation is a type of
concentration technique wherein the specific gravity of the parasite is lesser
than the suspending medium. The specific gravity of the medium should be 1.18 –
1.20.
The zinc sulfate flotation
technique is an effective concentration procedure for the recovery of both
protozoan cyst and helminth eggs and larvae from unpreserved specimens;
however, operculated eggs and those of the schistosomes are not recovered by
this method.
Procedure:
1.
Using 2 applicator stick, comminute a fecal
sample about the size of a small pea in a tube, half–filled with tap water.
Make certain that all obvious particles are broken up and that an even
suspension is formed.
2.
Add additional tap water until the tube is
2/3 full.
3.
Centrifuge for 1 minute at approximately
2,500 rpm.
4.
Pour off the supernatant fluid into a
container holding a disinfectant, for example, cresol.
5.
Repeat this washing only if the stool is
extremely oily.
6.
Add enough zinc sulfate to fill the tube half
full.
7.
Using an applicator, break up the packed
sediment very thoroughly
8.
Add additional zinc sulfate solution to fill
the tube within 1.3 cm of the top.
9.
Centrifuge this suspension for 1 minute at
2,500 rpm.
10. Without
shaking or spilling the solution, carefully place the tube in a rack.
11. Slowly fill
the tube brimful with zinc sulfate without allowing any runover.
12. Place a
clean, grease–free no.1 cover slip (22 x 22 mm) on top of the tube so that the
undersurface touches the meniscus. Leave undisturbed for about 10 minutes.
13. Deftly
remove the cover slip with a straight, upward motion. A drop containing egg and
cyst will adhere to the underside of the cover slip.
14. Lower this
onto a drop of iodine stain placed on a clean 2 x 3 inch slide. Seal the
preparation.
15. Examine
under a microscope for eggs and cysts.
B.
Formalin–Ether Sedimentation
Technique
Sedimentation is a concentration
technique wherein the specific gravity of the parasite is greater than the
suspending medium. This concentration procedure is efficient in recovering
protozoan cysts and helminth eggs and larvae, including operculated and schistosome
eggs. Less distortion of cysts occurs with this technique than with the zinc
sulfate method and is more effective in concentrating formalin treated
specimen.
Procedure with fresh specimen:
1.
Comminute a portion of the stool specimen in
sufficient saline so that upon centrifugation, 10 ml of emulsion will yield
about 2 ml of sediment. A portion about the size of walnut is usually enough.
The suspension can be prepared in the carton in which it is submitted or in a
beaker of flat–bottom paper cup.
2.
Using a small glass funnel, strain about 10
ml of the emulsion through one or two layers of wet gauze into a 15 ml pointed
centrifuge tube. With wide– mesh gauze, use two layers; with narrow meshed material,
use one layer.
3.
Centrifuge at 2,000 to 2,500 rpm for 1
minutes. Decant supernatant fluid.
4.
Resuspend the sediment in fresh saline,
centrifuge and decant as before. This step maybe repeated if cleaner sediment
is desired.
5.
Add about 10 ml of 10% formalin to the
sediment, mix thoroughly and allow standing for 5 minutes.
6.
Add 3 ml of ether then stopper the tube and
shake vigorously in an inverted position for a full 30 seconds. Remove the
stopper with care.
7.
Centrifuge at 1,500 rpm for about 1 minutes.
Four layers should result as follows:
a.
Ether at top
b.
Plug of debris
c.
Formalin solution
d.
Sediment
8.
Free the plug of debris from the sides of the
tube by ringing with an applicator stick and carefully decant the top three
layers. Use a cotton swab to remove any debris adhering to the sides of the
tube.
9.
Mix the remaining sediment with the small
amount of fluid that drains back from the sides of the tube (or if necessary,
add a small amount of formalin or saline) and prepare iodine and unstained
mounts in the usual manner for microscopic examination
Procedure with formalin–preserved
specimen
1.
Thoroughly stir the formalin–treated
specimen.
2.
Depending on the size and density of the
specimen, strain a sufficient quantity through gauze into a 15 ml pointed
centrifuge tube to give the desired amount of sediment indicated below.
3.
Add tap water, mix thoroughly and centrifuge
at 2,000 to 2,500 rpm for 1 minute. The resulting sediment should be about 1
ml.
4.
Decant supernatant fluid and, if desired,
wash again with tap water.
5.
Add about 10 ml of 10% formalin to the
sediment and mix thoroughly.
6.
Complete as fresh specimen, beginning with
step 6.
*********** STOLL’S EGG–COUNTING TECHNIQUE ***********
This is a method for determining the number of nematode eggs per
gram of feces in order to estimate the worm burden in humans. The advantage of this
technique is that it requires no specialized equipment. The disadvantage is the
counting takes a long time because of the amount of artifact materials on the
slide.
Procedure
1.
Fill a graduated 15 ml centrifuge tube to 14
ml mark with 0.1N sodium hydroxide.
2.
Using two applicators, add sufficient feces
to raise the fluid level to the 15 ml mark.
3.
If the feces are hard, allow the preparation
to stand for a while.
4.
Shake vigorously for 1 minute to secure a
homogenous suspension.
5.
The eggs and debris will begin to settle
immediately after the shaking is stopped. Quickly pipette 0.15 ml from the
middle area of the sample.
6.
Expel the entire contents onto a slide and
cover with coverslip
7.
Examine under LPO and count all hookworm eggs
present.
8.
Multiply the number obtained by 100. This
will yield the number of eggs per milliliter of formed feces.
Intensity grouping Clinical classification
(eggs/ml
of feces)
100 –
699 very
light
700 –
2,599 light
2,600
– 12,599 moderate
12,600
– 25,099 heavy
25,600
& over very
heavy
*********** KNOTTS CONCENTRATION TECHNIQUE ***********
This is used to detect the presence of microfilaria in peripheral
blood.
1.
To 1 ml venous blood, add 9 ml of 2%
formalin.
2.
Centrifuge at 1,500 rpm for 5 minutes.
3.
Pour off supernatant fluid and examine the
sediment
*********** GRADIENT CENTRIFUGATION TECHNIQUE ***********
This is used to detect the presence of microfilaria in peripheral
blood.
1.
Place 4 ml of Ficoll–Hypaque mixture in a 15
ml centrifuge tube and mix with 4ml of heparinized blood.
2.
Centrifuge at 150g for 40 minutes.
3.
Examine the middle Ficoll hypaque layer
separating the plasma and white cell layers.
*********** MEMBRANE FILTRATION TECHNIQUE ***********
This is a very sensitive method for detection of microfilaria in
peripheral blood.
1.
Around 50 ml of blood is filtered through nucleopore filter
of 5µm porosity.
2.
The membrane is dried and mounted on a slide
*********** TRIPLE CENTRIFUGATION PROCEDURE ***********
This is used for the diagnosis of malaria and trypanosoma
infections.
1.
9 ml blood is mixed with 1 ml of 6% sodium
citrate solution.
2.
Centrifuge at 100 g for 10 minutes.
3.
Collect the supernatant and centrifuge at 250
g for 10 minutes again.
4.
Centrifuge the supernatant at 700 g for 10
minutes.
5.
Examine the sediment as a wet film or as
stained smear.
*********** BENTONITE FLOCCULATION TEST ***********
Procedure
1.
Sera to be tested should be inactivated for
30 minutes at 56oC.
2.
Dilute serially each serum with 0.85% saline;
1:5, 1:10 and 1:20. Positive sera are further diluted until flocculation is
read negative.
3.
Pipette 0.1 ml of serum dilution into a well
of wax–ringed slide and add 1 drop of standardized test antigen.
4.
Rotate the slide in a horizontal plane on a
rotating for 15 minutes at 120 rotations per minute.
5.
Examine with dissecting microscope for the
presence of agglutination or flocculation.
Reading the test
Results are read
as follows:
4+ reaction – all
particles are agglutinated
3+ reaction –
75% of the particles are agglutinated
2+ reaction – 50%
of the particles are agglutinated
1+ reaction – 25%
of the particles are agglutinated
A 3+ or 4+
agglutination is considered positive. A 2+ or 1+ reaction is negative. In
each series of
tests, saline control and negative and positive serum controls
should be
included.
*********** PERIANAL SWAB
***********
Procedure
1.
Prepare a swab with a 9cm strip of scotch
tape, 1.9–2.5 cm in width and a standard 1 x 3 inch microscope slide. At one
end of the tape, 0.5cm is folded upon itself to provide a non–adhesive area for
handling. The remainder is applied to the slide with the gummed side down,
extending over the end and for about 1 cm on the undersurface of the slide.
2.
Employ the swab in the morning before bathing
or bowel movement.
3.
Hold the slide against a tongue depressor 2.5
cm below the end and lift the long portion of tape from the upper surface of
the slide.
4.
Loop the tape over the extended end of the
depressor to expose the gummed surface. Hold the tape and slide against the
depressor to provide tension and a firm support for the loop of scotch tape.
5.
Separate the buttocks and press the gummed
surfaces against several areas of the perianal region.
6.
Replace the tape on the side (to which it has
remained attached on the undersurface) and smooth the tape with cotton or
gauze.
7.
Examine the swab for eggs.
*********** CIRCUMOVAL PRECIPITIN TEST (COPT) ***********
The Circumoval Precipitin Test is a serological test used for
diagnosis of schistosomiasis japonica. Soluble egg antigens of Schistosoma
japonicum block the formation of the circumoval precipitin by serum from
infected humans.
1.
Use 1 drop of a suspension of eggs (about
100) in 1.75% saline and 1 drop or 0.05 ml of serum which has been inactivated
at 56oC for 30 minutes.
2.
Incubate at 37oC for 24 hours.
3.
Examine after 2 and 24 hours, recording
percentage of eggs showing precipitates, measuring length and noting fingerlike
shapes of the precipitate.
4.
Compare with negative and positive controls:
Weak reaction: precipitates 12.5 µm or less in length
Moderate reaction: precipitates 25 µm in length
Strong reaction: precipitates at 37.5 µm and over its
length
*********** McMASTER EGG COUNTING TECHNIQUE ***********
This method is for the
determination of the number of nematode eggs per gram of feces. The advantage
of this method is it is quick as the eggs are floated free of debris before
counting. The disadvantage is you must use a special counting chamber.
1.
Weigh out 2 grams of feces.
2.
Pass the feces through a sieve into a dish
containing 60 ml of ZnSO4 or saturated salt solution. Lift the sieve
and hold over the dish. Push out any remaining solution from the feces.
3.
While mixing vigorously (you may want to put
the solution into a flask to prevent spillage) take a sample of the mixture
with a pipette and transfer it to one of the chambers of the McMaster slide.
Repeat the procedure and fill the other chamber.
4.
Wait 30 seconds then count the total number
of eggs under both the etched areas on the slide. Focus first on the etched
lines of the grid, then go down a tiny bit, the eggs will be floating just
below the top of the chamber. Multiply the total number of eggs in the 2
chambers by 100, this is the eggs per gram (EPG).
5.
The volume under the etched area of each
chamber is 0.15 ml (the etched area is 1 cm x 1 cm and the chamber is 0.15cm
deep) so the volume examined is 0.3 ml. This is 1/200 of 60 ml. Since you
started with 2 grams of feces and then multiplied by 100, the final result is
eggs per gram of feces.
6.
Count both chambers.
(chamber 1 +
chamber 2) x 50 = eggs per gram (EPG)
*********** KATO–KATZ TECHNIQUE ***********
This technique is for the diagnosis of intestinal schistosomiasis
and intestinal helminths. This technique is not suitable for examining larvae,
cysts, or eggs from certain intestinal parasites.
1.
Soak the cellophane strips in a 50% glycerol–malachite
green (or methylene blue) solution for at least 24 hours before use.
2.
Transfer a small amount of feces on to a
piece of scrap paper (newspaper is ideal).
3.
Press the screen on top of the fecal sample.
4.
Using a flat sided applicator stick, scrape
across the upper surface of the screen to sieve the fecal sample.
5.
Place a template on a clean microscope slide.
6.
Transfer a small amount of sieved fecal
material into the hole of the template and carefully fill the hole. Level with
the applicator stick.
7.
Remove the template carefully so that all the
fecal materials is left on the slide and none is left sticking to the template
8.
Cover the faecal sample on the slide with a
glycerol–soaked cellophane strip.
9.
If an excess of glycerol is present on the
upper surface of the cellophane, wipe off the excess with a small piece of
toilet paper or absorbent tissue.
10. Invert the
microscope slide and press the faecal sample against the cellophane on a smooth
surface (a piece of tile or flat stone is ideal) to spread the sample evenly.
11. Do not lift
the slide straight up. The cellophane may separate. Gently slide the microscope
slide sideways holding the cellophane.
*********** BAERMAN TECHNIQUE ***********
The Baerman technique works on the principle that larvae will
migrate out of a fresh stool sample and will subsequently sink to the bottom of
their liquid environment. Larvae will therefore concentrate in the lowest point
and the contents of a large stool sample can be examined.
1.
Firmly attach a 6 inch glass funnel to a
retort stand using a ring adaptor. Attach rubber tubing with a secure clamp to
the stem of the funnel. Place a collection container under the end of the
tubing.
2.
On top of the funnel place a wire mesh with
two layers of gauze. Make sure that the gauze is trimmed to the size of the
funnel, so that none of the potentially infective solution will drip over the
slide of the funnel and contaminate the surrounding bench area. Fill the funnel
with water.
3.
Place the charcoal culture on top of the
gauze, making sure that it is in contact with the water.
4.
Allow the apparatus to stand for 2 hours or
longer before draining off a portion of the fluid directly above the clamp. Centrifuge
the fluid and examine for the presence of motile larvae.
*********** CITRATE ACID SAPONIN TECHNIQUE ***********
This procedure is based on the fact that most of the erythrocytes
will be destroyed as the citrated blood comes into contact with saponin, but
the microfilariae are unharmed to demonstrate motility.
1.
Mix 10 ml blood with 2 ml of freshly prepared
sodium citrate (two evacuated blood collection tubes with EDTA may be used).
The specimen should be mixed
immediately and inverted at intervals to avoid clotting.
2.
Centrifuge for 10 minutes at 1000 rpm and
carefully remove and discard plasma.
3.
Transfer all packed cells (approximately 8
ml) to 50 ml tubes containing freshly made saponin (0.5% w/v in saline).
4.
Mix gently at intervals and let stand for 15
mins.
5.
Centrifuge at 3500 rpm for 10 mins, decant
supernatant and discard.
6.
Examine the sediment under low power (x100)
magnification by spreading a few drops on a slide without coverslip. This
examination should be done quickly to observe the live, motile microfilariae
before allowing the slides to air dry for staining. Stained microfilariae will
appear coiled or curved.
7.
Mix the remainder of the sediment with two
drops of 1% acetic acid solution.
8.
Mix well with an applicator stick and spread
over the slide surface. Allow slides to air dry. (Microfilariae will be killed
and straightened)
9.
Dip the dried slides in methylene blue
phosphate solution and rinse in two changes of distilled water.
10. Place in
Giemsa stain for 8 to 10 minutes.
11. Rinse in
distilled water and air dry.
12. Search for
microfilaria under LPO.
*********** CASONI SKIN TEST ***********
Purpose
To determine skin test reactivity to Echinococcus
Principle
When hydatid cyst fluid is injected intradermally, patients with
hydatid cyst disease manifest hypersensitivity by developing redness and wheal
at the injection site. In strongly reactive cases, pseudopodia appear at the
periphery of the wheal. Although usually an immediate reaction, some are
delayed so that the injection site should be observed for 30 minutes and again
in 24 hours. A saline control should be injected intradermally below the test
site.
Reagent
The hydatid cyst fluid is obtained at the time of excision of a
cyst from an infected human. The contents of the excised cyst are aspirated
aseptically, centrifuged to remove hydatid sand (protoscolices), filtered and
then tested for sterility. If sterile, merthiolate is added to a final
concentration of 1:50,000 and the fluid is placed into a sterile vaccine
bottles. The shelf life at 25oC is 25 years and is longer at 5oC.
Procedure
1.
Obtain blood for serological testing prior to
injecting the antigen intradermally.
2.
Inject 0.05 ml of antigen intradermally into
the volar surface of one forearm.
3.
Inject 0.05 ml of saline (control) intradermally
into the volar surface of the forearm at a distance of not more than 10cm from
the site of injection of the antigen.
Interpretation
Positive test:
Appearance of redness and wheal
at the injection site of the antigen
Negative test:
No redness or wheal at the
injection site of antigen
*********** DEC (diethylcarbamazine) PROVOCATION TEST ***********
Suspected adults are given 100 mg of diethylcarbamazine orally in
daytime and peripheral blood smear examined for microfilariae in 30–45 minutes.
It is useful in situations where nocturnal periodicity occurs and night blood
collection is not feasible. The test has the disadvantage of provoking Mazzotti
reaction in areas with onchocerciasis.
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