is high; drying films as quickly as possible produces the best results. However, they should never be heated to decrease drying times; heat tends to produce distorted parasite and blood cell morphology. Absolute methanol tends to absorb moisture from the air, so that absolute methanol for fixation should not be reused from day to day but needs to be fresh daily. Do not store this fixative in a Coplin jar. Thin blood films should be fixed and stained within 24 h; deterioration may occur if the slides are held too long before being processed. However, the slides can be frozen for long-term storage. All thin blood films must be fixed prior to shipment to a reference laboratory; both these fixed slides and the original tube of blood should be sent to the reference laboratory. This assumes that the original laboratory has examined the blood films on site, as well.
If the thin blood films are fixed for too long, some of the morphologic details may be reduced or lost. If they are fixed for too short a time, the RBCs may be distorted or partially lysed and the thin film will not be uniformly stained. If the thin films are fixed in methanol that contains water, the RBCs will be distorted and there will appear to be “holes” in the thicker areas of the films. As a reminder, do not reuse absolute methanol for fixation; discard after each use.
Thick-Film Acetone “Quick Dip.” Although thick blood films are not fixed with absolute methanol, after the thick films are thoroughly dry, they can be dipped twice in acetone and allowed to dry before being stained. This extra step does not interfere with RBC lysis that occurs either prior to or during staining. The acetone “quick dip” makes the thick film less likely to fall off during staining and provides a cleaner background for microscopic examination.
For patients with suspected malaria (negative thick and thin blood films), trypanosomiasis, filariasis, or leishmaniasis, concentration procedures increase the number of organisms recovered from blood specimens. The buffy coat containing WBCs and platelets, and the layer of RBCs just below the buffy coat layer, can be used to prepare thick and thin blood films. The sensitivity of this approach is greatly enhanced over that of the thick film. However, it is critical that the correct layer be sampled from the centrifuged blood.
L. donovani, trypanosomes, and Histoplasma capsulatum (a fungus with intracellular elements resembling those of L. donovani) are occasionally detected in the peripheral blood. The parasite or fungus is found in the large mononuclear cells in the buffy coat (a layer of white cells resulting from centrifugation of whole citrated blood). The nuclear material stains dark red-purple, and the cytoplasm is light blue (L donovani). H. capsulatum appears as a large dot of nuclear material (dark red-purple) surrounded by a clear halo area. Trypanosomes in the peripheral blood also concentrate with the buffy coat cells.
After centrifugation and aliquoting of the appropriate layers, some of the material can be examined as a wet mount; trypomastigotes and microfilariae may be seen as motile objects in the wet mount. After staining, L. donovani amastigotes may be found in the monocytes and Plasmodium parasites may be seen in the thick and thin films. Advantages and disadvantages of buffy coat films can be seen in Table 7.5.
Detailed Procedure
1. Wear gloves when performing this procedure or preparing any blood films.
2. Whole blood should be collected using EDTA anticoagulant. Although heparin can be used, if malaria films are to be prepared, EDTA is recommended.
3. Although capillary hematocrit tubes have been used in the past, the cutting and breaking of these tubes to remove the cells for film preparation is not considered a safe procedure and is not recommended. However, if you use a microhematocrit tube, the tube should be carefully scored and snapped at the buffy coat interface, and the white cells are prepared as a thin film. The tube can also be examined before removal of the buffy coat, at the low and high dry powers of the microscope. If trypanosomes are present, the motility may be observed in the buffy coat. Microfilarial motility would also be visible.
4. Using a capillary pipette, fill a Wintrobe tube with blood containing anticoagulant (EDTA is preferred), cap the tube, and centrifuge it for 30 min at 100 × g. Another option is to centrifuge the tube of anticoagulated blood at 100 × g for 15 min, transfer that buffy coat to another tube, and centrifuge the tube at 300 × g for 30 min. After centrifugation, the tube contains three layers: plasma on top, a layer of white cells (buffy coat), and the packed RBCs on the bottom.
5. Remove and discard most of the plasma above the buffy coat, leaving a small amount on top of the buffy coat layer. Then remove the remaining plasma, buffy coat, and the RBCs right below the buffy coat. Transfer this aliquot to a separate tube.
6. Examine the buffy coat directly for motile trypomastigotes and microfilariae by mixing 0.5 drop of saline with 1 drop of buffy coat sediment on a microscope slide. Add a coverslip, and examine at low power (×10 objective).
7. Mix the aliquot gently (avoid bubbles), and prepare thick and thin blood films on alcohol-cleaned slides.
8. Allow the films to air dry horizontally and protected from dust for at least 30 min to 1 h. Do not attempt to speed the drying process by applying any type of heat, because the heat fixes the RBCs and they subsequently will not lyse (lake) in the staining process for the thin films.
9. Label the slide appropriately.
10. If staining with Giemsa is delayed for more than 3 days or if the film is to be stained with Wright’s stain, lyse the RBCs in the thick film by placing the slide in buffered water (pH 7.0 to 7.2) for 10 min, remove it from the water, and place it in a vertical position to air dry.
For accurate identification of blood parasites, a laboratory should develop proficiency in the use of at least one good staining method (2, 9–11). It is better to select one method that will provide reproducible results than to use several on a hit-or-miss basis. Blood films should be stained as soon as possible, since prolonged storage may result in stain retention. Failure to stain positive malarial smears within a month may result in failure to demonstrate typical staining characteristics for individual species.
The most common stains are of two types. Wright’s stain has the fixative in combination with the staining solution, so that both fixation and staining occur at the same time; therefore, the thick film must be laked before staining. Giemsa stain has the fixative and stain separate; therefore, the thin film must be fixed with absolute methanol before staining. If using one of the rapid blood stains, read the package insert to confirm laking and/or fixation requirements for the thin and thick blood films.
When slides are removed from either type of staining solution, they should be dried in a vertical position. After being air dried, they may be examined under oil immersion by placing the oil directly on the uncovered blood film. Remember, do not wipe the film to remove oil; lay slide oil side down on a paper towel to blot off the excess oil before storing the slide. If films are to be kept for a permanent record, they should be protected with a coverglass after being mounted in a medium such as Permount.
Note Blood films stained with any of the Romanowsky stains that have been mounted with Permount or other resinous mounting media are susceptible to fading of the basophilic elements and generalized loss of stain intensity. Hollander
