Microscopic Techniques
Most parasitic agents occur within the gastrointestinal region, body tissues and blood system. Most helminth parasites (nematodes, trematodes, cestodes) and some protozoa inhabit the intestine as well as the bile and pancreatic ducts that empty into the intestinal lumen. The helminths produce characteristic eggs, which pass out in faeces that are used for identification. Either the protozoa exist as active trophozoites or as inactive cysts and their presence in a stool sample is an indication of an infection.
Some important stains for blood films
Method for cleaning microscope slides
Slides used in laboratories for identification purposes should be clean from dust and grease. Films do not stick well if the slides are dirty and greasy and stained artefacts may give the wrong results. There are several ways of cleaning blood slides but the most practical method is the one that uses Decon
Make up a 2 – 3 % solution of Decon in hot water.
Place slides into the solution individually.
Allow slides to stand in solution for at least ½ hour (1 – 2 hours is better).
Rinse thoroughly in running tap water for an hour.
Rinse briefly in distilled water.
Store slides in ethanol – dry as required or dry and store in clean boxes.
Giemsa’s Stain
This stain can be used in different dilutions. A one in ten dilution is convenient, as it gives rapid staining, although the time varies according to the quality of the stain.
The stain can be used for thin films and thick films.
Method for the thin films
Pour 2 ml of Giemsa into a measuring cylinder and make up to 20 ml with distilled water. Empty the dilute stain into a staining jar.
Fix the blood film by immersing it in methyl alcohol for 2 – 3 minutes.
Wash the slide containing the film in distilled water.
Drain off the water and transfer the slide into a staining jar containing Giemsa.
Stain for 30 – 60 minutes.
Remove the slide and rapidly wash it in distilled water.
Wipe off stain from the back of the slide and leave it in an upright position to dry.
Examine in immersion oil at 100 % magnification.
Method for thick films
Prepare the final stain as for the thin film.
Without any fixation, place the slide in the stain.
Leave for 15 – 30 minutes in the stain (by agitating the staining jar gently from time to time, haemoglobin is removed from the area of the film).
Remove the slide from the stain and wash in distilled water.
Let it dry
Leishman’s Stain
Method for thin films
Place the slide on a staining rack over sink or dish, with the film facing uppermost.
Cover the film with 12 drops of the stain and leave for 1½ minutes (the methyl alcohol in the stain fixes the film).
Add double the quantity of freshly distilled water.
Mix the solution thoroughly with a pipette or by rocking very gently the slide, taking care not to spill the stain.
Allow to stain for 15 to 20 min, the actual time depending on the quality of the stain. The film should be quite a strong pink colour, when sufficiently stained. The colour is then reduced or the film differentiated by washing with distilled water.
Flood the slide with distilled water, then tip and allow the water to run off. Continue to wash with distilled water until the film is a rather pale pink.
Stand the slide in an upright position to drain and dry. Heat must not be used to dry films but the film can be gently dabbed with blotting paper and waved in the air to expedite drying.
Examine in immersion oil at 100% magnification.
Method for thick films
Thick films must be treated very gently at all stages, as they are not firmly attached to the slides.
Place the slide film side downwards in a dish (or stand it upright in ajar) containing tap or distilled water.
Leave until all the haemoglobin has been removed (2 to 3 min) and the film has become completely white.
Stand the slide in an upright position to drain and dry.
Proceed as for staining thin films.
Field’s stain for thick films
Field’s stain is used for rapid diagnostic and identification purposes. For thin films, fix in methyl alcohol for 2 to 3 minutes prior to staining.
Preparation
A. Methylene blue (medicinal) 0.4 g
Azur 0.25 g
Buffered water (B), pH, and 6.8 – 7.0. 250 ml
B. Disodium hydrogen phosphate 10.0g
Potassium dihydrogen phosphate 12.5 g
Distilled water 1000 ml
C. Eosin 0.5 g
Buffered water (B) 250 ml
Method
Dip slide in A 1- 2 sec
Rinse in B 2- 3 sec
Dip in C 1- 3 sec
Rinse gently in tap water 2- 3 sec
Place slide upwards to drain and dry
Examination of Stool for Intestinal Protozoa
Direct Method
Place a fresh sample of stool, about the size of a bean seed, on a slide, add a drop or two of physiological saline, cover with a cover slip, and examine at a magnification 400 - 500X. This technique should reveal the motile forms of intestinal flagellates.
To identify the cysts and the nuclei of Amoeba, one or two drops of a 4% Lugo
Examination of Stool for Worm Eggs
a) Direct Examination
A sample of faeces, about the size of a bean seed, is placed on a slide, mixed with tap water or physiological saline, covered with a cover slip, and examined at low microscope magnification (100 to 200X).
The smear should not have lumps of faeces to obscure the eggs. A drop of Lugo
b) Thick Smear Technique (Kato and Miura)
Preparation of the Kato stain
Distilled water 500 ml
Glycerine 500 ml
Malachite green, 3% in water 5 ml
Place about 100 mg of faeces on a slide and cover it with a cellophane strip (size 26 x 28 mm), which had been soaked in the Koto Stain for at least 24 hr. Press over the strip with a spatula to have a thin uniform spread of the faeces sample. Allow the preparation to stay at room temperature for ½ to 1 hr. Examine the stool. The glycerine clears the faeces and helminth eggs can be seen at a magnification of 100x.
c) Concentration Method
Place 1 g of stool in a test tube and thoroughly mix it with a concentrated salt solution. Remove any plant particles floating on the surface and allow the mixture to remain undisturbed for 20 to 30 min.
Helminth eggs float on the surface because they are lighter than the salt solution, Transfer the eggs to a microscope slide using a wire loop. Examine at a magnification of 100-200x.
d) Universal Concentration Method
Place 1 g of faeces in a small beaker, add 7 ml of 50% hydrochloric acid and the same amount of ether. Mix the mixture to form a homogeneous solution. Pass the mixture into a centrifuge tube over two layers of muslin placed in a funnel. Centrifuge the solution for a minute or two.
Four layers are formed: the uppermost layer is a yellowish zone of ether below which is a zone consisting of food particles and below this is a zone of hydrochloric acid. The bottommost zone consists of small particles and worm eggs. Carefully insert a pipette into the bottom layer, suck out liquid, and examine it for eggs. Be careful, ether is very volatile and can explode!
Sedimentation Concentration Stool Examination Technique
Add an estimated 5 g of stool to 100 ml of formol glycerol solution (5 ml formaldehyde, 10 ml glycerol, 985 ml of water) in a flask and mix with a glass rod.
Sieve through a mesh into another flask.
Add more formol glycerol up to 3 cm below the top and allow to sediment for a minimum of 20 min at room temperature.
Pour off the supernatant to leave approximately 25 to 30 ml of fluid containing sedimented deposit
Resuspend in formol glycerol and sediment for 20 minutes.
Gently pour off supernatant to leave 15 to 20 ml of suspended deposit.
Use straw to transfer about 0.1 ml of the deposit to each of 3 microscope slides and cover each slide with a cover slip.
Examine
No comments:
Post a Comment