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 Staining Methods
 Direct saline, Iodine Mounts and BMB Mounts
 Materials and reagents
1. Coverslips
2. Dropping-bottles
containing: saline solution (isotonic), Lugol’s
iodine (1% solution), 1% BMB stain
3. Microscope
slides
4. Pens
or markers for labeling
5. Wire
loop (or applicator sticks, matchsticks, or toothpicks).
Technique
1. With a wax pencil write the patient’s name or
number and the date at the left-hand end of the slide.
2. Place a drop of saline in the centre of the
left half of the slide and place a drop of iodine solution in the right half
of the slide.
Note:
If the presence of amoebic trophozoites is suspected, warm saline (37° C) should be
used. In case of BMB stain, wait for five minutes to allow the stain to
penetrate the trophozoites. It will overstain the trophozoites in
30 minutes. Therefore the slide must be examined as early as possible.
3. With
an applicator stick (match or toothpick), pick up a small portion of the
specimen (size of a match head) and mix with the drop of saline.
Note
Formed stool: Take the portion of
stool from an area to include inside and outside parts of the specimen.
Stool with mucus: If mucus is
present, label a second slide with the patient’s name or number. Put a drop
of saline on the slide, pick up a small portion of mucus and mix with the
saline. Trophozoites, if present, are sometimes
more readily found in mucus than in the solid parts of the stool.
Loose watery stool: If mucus is
not present, pick up a small portion of the stool (any part) and mix with the
saline.
4. Similarly,
pick up a small amount of the stool and mix with the drop of iodine, to
prepare an iodine mount. If a wire loop is used, flame it after making the
mount. If applicator sticks are used, discard them.
5. Cover
the drop of saline and the drop of iodine with a coverslip.
Hold the coverslip at an angle, touch the edge of
the drop and lower gently on to the slide. This will reduce the chance of
including air bubbles in the mount.
Concentration
techniques
Formalin
ether concentration technique
If the number of organisms in
the stool specimen is low, examination of a direct wet mount may not detect
parasites. Thus, whenever possible, the stool should be concentrated. Worm
eggs, larvae and protozoan cysts may be recovered by concentration but
protozoan trophozoites will NOT be seen as they are
usually destroyed during the concentration procedure. This makes direct wet
mount examination obligatory as the initial phase of microscopic examination.
The concentration procedure is
indicated when the initial wet mount examination is negative despite the
clinical symptoms indicating parasitic infection of a patient, and for the
detection of Schistosoma and Taenia.
The concentration procedure
recommended is the formalin-ether (or formalin-ethyl acetate) method. All
types of worm eggs (roundworms, tapeworms, schistosomes,
and other fluke eggs), larvae, and protozoan cysts may be recovered by this
method.
Materials and reagents
1. Applicator sticks, wooden
2. Bottles,
dispensing or plastic "squeeze", 250 ml or
500 ml. These bottles are convenient for adding formalin to the centrifuge
tubes. However, any small bottles or flasks may be used.
3. Centrifuge,
with head and cups to hold 15-ml conical tubes. Sealed buckets must be used.
4. Centrifuge
tubes, 15 ml, conical (make a graduation at 7 ml and 10 ml with a grease
pencil).
5. Cotton
swabs
6. Coverslips
7. Funnel
8. Surgical
gauze
9. Microscope
slides
10. Pipettes,
Pasteur, with rubber bulbs
11. Rack
or support for tubes
12. 10%
Formalin. For everyday use, pour some of the solution into a
"squeeze" bottle. Label the bottle.
13. Ether
or ethyl acetate.
14. Lugol’s iodine, 1% solution - in a dispensing bottle with
a pipette
15. Saline
solution, isotonic
Caution
Ether
is a highly flammable compound and will ignite and explode quickly if there
is a flame or spark nearby. Store opened cans or bottles on an open shelf in
the coolest part of the laboratory. Be sure the cans or bottles are stoppered. DO NOT put an opened container of ether in a
refrigerator: fumes build up inside the refrigerator, even if the container
is closed and may explode when the door is opened. DO NOT put opened containers
in a cabinet. It is better to leave the container on an open shelf so that
the fumes can disperse readily.
Technique
1. Add 10 ml of 10% formalin to approximately 1 g
of faeces and stir an applicator stick, until you
get a slightly cloudy suspension. Fit a gauze filter into a funnel and place
the funnel on top of the centrifuge tube.
2. Pass
the faecal suspension through the filter into the
centrifuge tube until the 7 ml mark is reached.
3. Remove
the filter and discard the filter with the lumpy residue.
4. Add
3 ml of ether or ethyl acetate and mix well for one minute.
5. Transfer
back to the centrifuge tube and centrifuge for 1 minute.
6. Loosen
the fatty plug (debris) with an applicator stick, and pour away the
supernatant by quickly inverting the tube.
7. Replace
the tube in its rack and allow the fluid on the slides of the tube to drain
down to the sediment. Mix well and transfer a drop to a slide for examination
under a coverslip. Also make an iodine-stained
preparation.
8. Use
the X 10 and X 40 objectives to examine the whole area under the coverslip for ova, cysts, and larvae.
Concentration
technique for cryptospordium oocysts
Flotation
or sedimentation is particularly helpful in recovering oocysts
from non-liquid stool specimens. Oocysts float
easily in Sheather’s sucrose solution, in zinc sulphate (33% to saturated), and in sodium chloride (36%
to saturated). They pellet with formalin ethyl acetate or formalin ether.
Permanent
staining techniques
Permanently
stained slides are not made routinely in diagnostic practice and are not
required for the identification of worm, eggs or larvae. However, permanently
stained preparations are occasionally required for the following purposes:
Identification of oocysts
of Cryptosporidium;
Identification of protozoan trophozoites or cysts, if doubt exists;
Confirmation of the identified protozoan
cysts, where doubt exists;
Keeping a permanent record, and
Sending to a reference laboratory for an
expert opinion.
Staining
Technique for Oocysts of Cryptosporidium parvum
Occysts of Cryptosporidium passed in faeces
are spherical, measuring 4-6µ m in diameter. They may be concentrated using a
modified formalin-ether technique, but must be identified by staining
methods. The recommended method is the modified Ziel
- Neelsen technique.
Modified
Ziehl - Neelsen Technique
Materials
1. Applicator sticks, wooden
2. Coverslips
3. Forceps
4. Microscope
slides
5. Pen
or marker for labeling
6. Rod,
glass
7. Slide
holder, for finished slides
8. Small
bottle of mounting medium
9. Staining
dishes
10. Paper
towel or sponge
Reagents
1. Carbol – fuchsin, filter before use
2. Formalin (formaldehyde)
3. Hydrochloric
acid - ethanol solution
4. Glycerol
- malachite green (or methylene blue) solution
5. Hydrochloric
acid - methanol solution
6. Water
Preparation
1. Make a thin faecal
smear, leave it to air-dry and fix it with a flame for a few seconds or in
methanol for 2-3 minutes. Further fixation in formalin vapour
should be performed if possible, to reduce infectivity. (The sediment from
formalin - ether extraction cannot be used.)
2. Flood
the smear with cold carbol-fuchsin.
3. Heat
the slide until steaming but do not allow drying.
4. Allow
to stand for approximately 5 minutes.
5. Rinse
slide in tap water and drain.
6. Decolourize with 5% H2SO4 or 1% HCl–ethanol
until color ceases and flood out (1-2 minutes).
7. Rinse
slide in tap water and drain.
8. Counterstain with methylene
blue or 0.25% malachite green for 1-2 minutes.
9. Rinse
in tap water.
10. Blot
or drain dry.
11. Examine
using first x10 and then the high-power, dry objective x40 and confirm the
morphology using oil immersion. Measure the cysts. Cryptosporidium cysts
measure 4-6 µ m and appear as bright rose-pink spherules on a pale green
background.
Gram-chromotrope stain for Microsporidia
Materials and Reagents
Microscope slides
Coverslips
Applicators sticks, wooden and
glass
Forceps
Pen or marker for labeling
Slide holder, for finished
slides
Small bottle of mounting medium
Seven staining dishes
Paper towel or sponge (if not
available newspaper)
1% Crystal violet
Gram’s iodine solution
Ethyl ether acetone ( equal
concentrations of ethyl ether and acetone)
Chromotrope
stain
Chromotrope
2R 6.0 g
Fast green 0.15 g
Phosphotungstic
acid 0.7 g
Glacial acetic acid 3.0 g
Distilled water 100 ml
0.45% acid alcohol
Preparation
of chromotrope stain
Weigh
each dye powder separately. Put the dyes into a 100-ml flask. Weigh out phosphotungstic acid crystals and add to the flask with
the dyes. Measure the glacial acetic acid and pour into the flask. Swirl the
flask so that the acetic acid wets the dyes. Let it stand for 30 minutes.
Then mix with 100 ml of distilled water and adjust the pH to 2.5 with 1 M HCl. Pour the stain into a 100-ml, clean, glass-stoppered bottle. Label the bottle as Gram Chromotrope stain and write the date. Store on a shelf or
in cabinet away from the light. The stain will remain good for a year or
more.
Preparation
1. Make a faecal smear
and let it dry at room temperature.
2. Heat fix (Three times for l sec. each over a
low flame).
3. Flood
the slide with Crystal violet solution.
4. Rinse
with tap water.
5. Flood
slide with Gram’s iodine and allow to remain on the
slide for 1 min.
6. Rinse
gently with tap water (1-2 sec).
7. Flood
with tap water.
8. Place
slide in Chromotrope stain for at least 5-10 min.
9. Rinse
in 0.45% acid alcohol for 1-3 sec.
10. Rinse
in 95% ethanol for 1 min.
11. Rinse
in absolute ethanol for 30 sec twice.
12. Let
it dry at room temperature.
Examination
1. Put the slide with the mounts on the
microscope stage and focus on the mount with the 10x or low-power objective.
2. Regulate
the light in the microscope field with the substage
diaphragm. You should be able to see objects in the field distinctly. Too
much or too little light is not good.
3. Examine
the entire coverslip area with the X 10 objective:
focus the objective on the top left-hand corner and move the slide
systematically backwards and forwards, or up and down.
4. Use
the 40x objective to identify small parasites.
5. Use
the oil-immersion objectives to examine protozoan and coccidia.
This is
a systematic examination. If mounts are examined in this way, any parasites
present will usually be found. If the mount is not examined systematically,
parasites may be missed. Examine each microscopic field carefully, focusing
up and down, before moving to the next field.
Toluidine Stain for
Pneumocystis carinii
Staining
Procedure
1. Dry slide for 5 minutes.
2. Flood
slide with sulphonation reagent for 10 minutes.
3. Note:
Wear gloves and take care when handling this reagent.
4. Carefully
wash off reagent and flush down sink.
5. Place
slide in a container and flush with running water for 5 minutes.
6. Flood
slide with Toluidine blue for 3 minutes.
7. Wash
off and allow slide to air dry.
8. Examine
using 10x objective where clumps of cysts may be visible and 40x and under oil immersion for confirmation.
Results
Pneumocystis carinii cysts
of characteristic morphology and size 3-5 µm are violet or purple against a
bluish background.
Sulphonation
reagent
Fresh
reagent is made once a week. To 9 ml of glacial acetic acid in a bottle held
in a cool water bath (10-150C), slowly add 3 ml of concentrated sulphuric acid. Mix and leave in the water bath until the
solution cools.
Toluidine blue
Dissolve 0.3 g of Toluidine blue in 60 ml of distilled water. Add 2 ml of
concentrated hydrochloric acid followed by 140 ml of absolute ethyl alcohol.
Remember: Examine mounts
systematically 
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