analis laboratorium kesehatan

PRINCIPLE

Preparation of thick and thin blood smears, appropriate staining procedure and detection and identification of hemo-parasites are crucial to clinical diagnosis of many parasitic diseases. These include species of malaria, trypanosomes, babesias and microfilariae of filarial nematodes. On properly prepared smears hemo-parasites can be identified on the basis of their morphologic features.

CLINICAL SIGNIFICANCE

Malaria infection is the most widespread, fatal human parasitic disease. All of the parasites infecting human blood can cause significant disease and prompt identification and treatment are necessary.

SPECIMEN

1. SLIDE PREPARATION: For making blood smears, use 3” x 1” slides with a frosted end to be used for patient identification. Slides must be free of scratches, grease, dust, and acid or alkali contamination. The recommended procedure is to use new slides, wash them individually in a solution of warm water and detergent with a soft brush or cloth, rinse thoroughly in distilled water, let drain briefly on a clean towel, dip in 100% Reagent Alcohol and dry with lint-free cloth or lens paper. Store cleaned slides in a closed container to protect from dust. Handle slides by the edges to avoid fingerprints.

PRECAUTION: Gloves should be worn throughout procedure. Patient specimens and all materials coming into contact with them should be handled as if capable of transmitting infections and disposed of with proper precautions.

2. FINGER PRICK PROCEDURE: Capillary blood is preferred to venous blood. Use the tip of the middle or “ring” finger. The skin area to be punctured should be warm so that blood flow will be adequate. Depending on the physical setting and the patient’s condition, warming the hand in warm water, covering the hand with a hot, wet towel, or briskly rubbing the hand may be used to warm the hands prior to the finger stick. Dry skin thoroughly and prior to performing the finger stick use an alcohol wipe to cleanse the finger; rub dry with a piece of gauze. Perform a finger stick using a sterile lancet. Use the gauze to wipe away the first drop of blood. Allow the blood to well up in a large drop. Gentle pressure may be used on the finger if needed.

3. Prepare at least two thin and two thick smears on separate slides. (Combination thick/thin smears are not recommended for diagnostic use although may be used for field screening use or to prepare slides for educational purposes.)

4. LABEL ALL SLIDES WITH THE PATIENT NAME, DATE AND TIME OF COLLECTION.

5. FOR THICK FILMS:
Touch the slide to the drop of blood. Use the corner of another clean slide to spread the drop of blood evenly, with a circular motion, until the drop is about the size of a dime (about 18mm in diameter) and of such a density that fine print can just be read through it. Two drops may be put on one slide.

6. FOR THIN FILMS:
Place one drop (approximately 0.50 ml) of blood near one end of a clean microscope slide. Hold a second slide at a 30° angle and immediately draw into the drop of blood. Allow the blood to spread almost to the width of the slide; rapidly and smoothly push the spreader slide to the opposite end of the slide, pulling the blood behind it.

Note: A properly prepared thin film is thick at one end and thin at the other. The thin, feathered end of the film should be centrally located on the slide with free margins on both sides; the feathered edge should be only one layer thick. Streaks in the blood film indicate dirt on the slides; holes indicate the presence of grease on the slide.

7. FOR COMBINATION THICK/THIN FILMS:
Combination thick/thin smears are not recommended for diagnostic use because of the amount of time that must be allowed for drying the thick smear before the smears can be stained. The combination film may be used for field screening use or to prepare slides for educational purposes.

Make a thin smear first on about 2/3 of the slide and then make a thick smear. Leave a space between the two smears so that the thin smear can be fixed.

8. After the collection of the specimen, pressure should be applied to the puncture site with sterile cotton or gauze until bleeding stops. Apply a bandage strip to the site.

9. DRYING OF BLOOD FILMS: Films should be allowed to dry in a horizontal position at room temperature. Protect smears from dust, insects or other contaminants. Thin smears will dry and be ready to fix and stain in about 15 minutes. Thick smears should dry several hours or overnight. Thick smears must be thoroughly dry to ensure that the smear does not wash off during staining. Combination thick/thin smears should be treated the same as a thick smear.

REAGENTS

Wright (Wright-Giemsa) Stain
Used in hematology, this stain is not optimal for blood parasites. It can be used if rapid results are needed, but should be followed up when possible with a confirmatory Giemsa stain, so that Schüffner’s dots can be demonstrated.

Giemsa Stain

1. Methanol

2. Giemsa Stain- Azure B (Certified) or Wright-Giemsa Stain

3. Stock Buffers:

A. Alkaline Buffer:
Disodium hydrogen phosphate-anhydrous (Na2HPO4) - 9.5gm
Distilled Water - 1000.0ml
Store at room temperature in a tightly capped container.

B. Acid Buffer
Sodium dihydrogen phosphate-monohydrate (NaH2PO4· H2O) - .9.2gm
Distilled Water - 1000.0ml
Store at room temperature in a tightly capped container.

4. STOCK SOLUTION OF TRITON X-100
10% aqueous solution. Store tightly stoppered at room temperature.

5. WORKING BUFFERS
A. Working Buffer for Thin Smears or combination thick/thin smears
Stock acid buffer . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 39.0ml
Stock alkaline buffer . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .61.0ml
Distilled Water . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 900ml
Add 1 ml of the stock TRITON X-100 10% aqueous dilution per 1000mls
Buffer solution. (Final concentration = 0.01% Triton X-100.)
Adjust pH of Buffer to 6.8 -- 7.0.

B. Working Buffer for Thick Smears
Stock acid buffer . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 39.0ml
Stock alkaline buffer . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .61.0ml
Distilled Water . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 900ml
Add 10 mls of the stock TRITON X-100 10% aqueous dilution per 1000 mls buffer solution. (Final concentration = 0.1% Triton X-100.)
Adjust pH of Buffer to 6.8 -- 7.0.

CALIBRATION

1. pH Meter--Calibration of the pH meter is done before each use (if available).

2. Ocular Micrometer--An ocular micrometer is recommended for measuring parasite size. The ocular micrometer is calibrated with a stage micrometer and the adjustment factors are posted on each microscope.

QUALITY CONTROL

Smears, which are positive for hemo-parasites may not always be available. A blood smear, thick and/or thin, is run once each month or with each staining procedure if done less often than once each month. The blood smear does not have to be positive for blood parasites. Good color differentiation of red and white cells is an indication of a good quality stain.

Results of Q.C. smears for blood parasites are recorded on the QC Record Sheet for Giemsa Staining. This record is maintained in the QC Log Book.

STAINING PROCEDURE

A. Thin Blood Smears
1. Fix smears in absolute methanol. Slides may be briefly dipped in a coplin jar or while holding slide at an angle, methanol can be dropped on the smear. Allow to air dry completely.

2. Prepare the working stain solution. Add 1 ml of stock Giemsa or Wright-Giemsa to 19 mls of Working Buffer for Thin Smears. Place the smear side down in a petri dish or other tray with a low edge. Rest the end of the slide on an applicator stick or the edge of the pan. Flood the stain underneath the slide and stain for 20 minutes. This method minimizes stain precipitate and debris on the stained smear. (Time and dilution can be adjusted as necessary for proper staining reactions.)

3. Wash briefly by gently rinsing or dipping in and out of a jar of working buffer for thin smears.

4. Allow slides to air-dry in a vertical position.

B. Thick Blood Smears
1. Blood films must be thoroughly dry. Dry for several hours or overnight. Do not dry in an incubator or by exposure to heat as this will fix the blood cells and interfere with lysing the red blood cells prior to staining. DO NOT FIX SLIDES IN METHANOL.

2. Place slides in a coplin jar of tap water for 1-2 minutes. This will lyse the red cells on the smear.

3. Prepare the working stain solution. Add 1 ml of stock Giemsa or Wright-Giemsa to 19 mls of Working Buffer for Thick Smears. Place the smear side down in a petri dish or other tray with a low edge. Rest the end of the slide on an applicator stick or the edge of the pan. Flood the stain underneath the slide and stain for 20 minutes. This method minimizes stain precipitate and debris on the stained smear. (Time and dilution can be adjusted as necessary for proper staining reactions.)

4. To rinse the smear, immerse in a coplin jar of working buffer for thick smears for 3 to 5 minutes.

5. Air dry in a vertical position.

C. For Combination Thick/Thin Smears

1. Blood films must be thoroughly dry. Dry for several hours or overnight. Do not dry in an incubator or by exposure to heat as this will fix the blood cells and interfere with lysing the red blood cells prior to staining.

2. Dip the thick portion of the smear in tap water for 1-2 minutes. Do not allow any water to get on the thin smear portion. Air-dry the smear.

3. Hold the slide with the thin portion down and drop methanol onto the smear. Allow alcohol to drain off and air dry. Smear must be completely dry before staining.

4. Prepare the working stain solution. Add 1 ml of stock Giemsa or Wright-Giemsa to 19 mls of Working Buffer for Thin smears. Place the smear side down in a petri dish or other tray with a low edge. Rest the end of the slide on an applicator stick or the edge of the pan. Flood the stain underneath the slide and stain for 20 minutes. This method minimizes stain precipitate and debris on the stained smear. (Time and dilution can be adjusted as necessary for proper staining reactions.)

5. To rinse the smear, immerse in a coplin jar of Working Buffer for Thin Smears for 3 to 5 minutes.

6. Allow slide to air dry in a vertical position with the Thick film down.

MICROSCOPIC READING OF SMEARS FOR HEMO-PARASITES

If microfilariae are suspected, scan both thick and thin smears entirely on the low power objective. Use higher power objectives to identify any parasites found.

Smears for Plasmodium species, Babesia species, Trypanosomes are read with the oil immersion (100X) objective. The entire Thick Smear should be examined; a minimum of 300 fields are read before a thin smear may be reported as negative.

Use reference materials available in the lab to identify any parasites found. If parasites are present, carefully study the morphology to determine:
· Stages found
· Size and shape of infected RBC’s
· Presence of schuffner’s dots
· Appearance of parasite cytoplasm
· Color of pigment (malaria)
· Number of merozoites in schizonts (malaria)
· Indicate the relative number of parasites seen as rare, few, moderate or many
Determine the species is possible.

QUANTIFICATION OF PARASITES

In some cases (especially malaria) quantification of parasites yields clinically useful information. If this information is needed by the physician, malaria parasites can be quantified against blood elements such as RBCs or WBCs.
To quantify malaria parasites against RBCs, count the parasitized RBCs among 500-2,000 RBCs on the thin smear and express the results as % parasitemia.
% parasitemia = (parasitized RBCs/total RBCs) × 100
If the parasitemia is high (e.g., > 10%) examine 500 RBCs; if it is low (e.g., <1%) blood="(parasites/WBCs)">