Читать книгу Diagnostic Medical Parasitology - Lynne Shore Garcia - Страница 16
ОглавлениеDepending on the life cycle, a number of parasites may be recovered in a blood specimen, either whole blood, buffy coat preparations, or various types of concentrations. These parasites include Plasmodium, Babesia, and Trypanosoma species, Leishmania donovani, and microfilariae. Although some organisms are motile in fresh, whole blood, species identification is normally accomplished from the examination of both thick and thin permanent stained blood films. Blood films can be prepared from fresh, whole blood collected with no anticoagulants, from anticoagulated blood, or from sediment from the various concentration procedures. Although for many years Giemsa stain has been the stain of choice, the parasites can also be seen on blood films stained with Wright’s stain, a Wright/Giemsa combination stain, or one of the more rapid stains such as Diff-Quik (American Scientific Products, McGaw Park, IL), Wright’s Dip Stat stain (Medical Chemical Corp., Torrance, CA), or Field’s stain. Delafield’s hematoxylin stain is often used to stain the microfilarial sheath; in some cases, Giemsa stain does not provide sufficient stain quality to allow differentiation of the microfilariae. A complete discussion of the proper way to examine a blood film is presented later in this chapter. This information is important and particularly relevant when one is examining proficiency testing blood films in the absence of clinical information about patient history and possible etiologic agents.
It is important to remember that standard precautions should be used at all times when blood or body fluids are handled (1). Remember that all requests for malaria diagnosis are considered STAT requests, and specimens should be ordered, collected, processed, examined, and reported accordingly.
Preparation of Thick and Thin Blood Films
Some parasites (microfilariae and trypanosomes) can be detected in fresh blood by their characteristic shape and motility, but specific identification of the organisms requires a permanent stain. Two types of blood films are recommended. Thick films allow a larger amount of blood to be examined, which increases the possibility of detecting light infections (2). However, species identification by thick film, particularly for malaria parasites, can usually be made only by experienced workers. The morphologic characteristics of blood parasites are best seen in thin films, in which the red blood cell (RBC) morphology is preserved and the size relationship between infected and uninfected red cells can be determined after staining. This characteristic is often valuable in determining the species of Plasmodium present from the thin blood film.
The accurate examination of thick and thin blood films and identification of parasites depend on the use of absolutely clean, grease-free slides for preparation of all blood films. Old (unscratched) slides should be cleaned first with detergent and then 70% ethyl alcohol; new slides should also be cleaned with alcohol before use. When a new box of slides is opened, the slides are coated with a substance that allows them to be pulled apart; these slides should be cleaned before use for preparation of blood films. Do not use cotton; gauze is recommended with 70% alcohol. The advantages and disadvantages of the thin and thick blood films can be seen in Table 7.1.
Blood should be collected immediately on admission or when the patient is first seen in the emergency room and/or clinic; if the initial blood films are negative, collect daily specimens for 2 or 3 additional days (ideally between paroxysms if present; however, there is often no periodicity seen). After a finger stick, the blood should flow freely; blood that has to be “milked” from the finger will be diluted with tissue fluids, decreasing the number of parasites per field. If the specimen is sent directly to the laboratory, thus eliminating laboratory-patient contact, the following approach can be used. Unless you are positive that you will receive well-prepared slides, request a tube of fresh blood (EDTA anticoagulant is preferred/lavender top) and prepare the smears. In general, the use of finger stick blood has declined, particularly in areas of the world where automated hematology instruments have become much more widely used. For detection of stippling, the smears should be prepared within 1 h after the specimen is drawn. After that time, stippling may not be visible on stained films; however, the overall organism morphology will still be acceptable. Potential problems with anticoagulants can be seen in Table 7.2. Although blood films can be prepared from the small amount of blood left in the needle after the venipuncture collection using anticoagulant, it is not recommended for several reasons. The blood tends to clot fairly quickly in the needle, and there are safety recommendations that limit the handling of needles. Therefore, the finger stick and EDTA venipuncture are recommended for collection of specimens for blood film preparation.
The time when the specimen was drawn should be clearly indicated on the tube of blood and also on the result report. The physician will then be able to correlate the results with any fever pattern or other symptoms that the patient may have. However, with travelers who are immunologically naive (have never come in contact with malaria before), there may not be any fever periodicity at all, and the symptoms may be very general and nonspecific for a malaria infection. There should also be some indication on the report that is sent back to the physician that one negative specimen does not rule out the possibility of a parasitic infection.
Note Although most laboratories use commercially available blood collection tubes, the following approach can be used when necessary. EDTA (Sequestrene) can be prepared and tubed as follows. Dissolve 5 g of EDTA in 100 ml of distilled water. Aliquot 0.4 ml into tubes, and evaporate the water. This amount of anticoagulant is sufficient for 10 ml of blood. One can also use 20 mg of EDTA (dry) per tube (20 mg/10 ml of blood). The tube should be filled with blood to provide the proper blood/anticoagulant ratio.
Fresh Blood
To prepare the thick film, place 2 or 3 small drops of fresh blood (no anticoagulant) on an alcohol-cleaned slide. With the corner of another slide and using a circular motion, mix the drops and spread them over an area ∼2 cm in diameter. Continue stirring for 30 s to prevent the formation of fibrin strands that may obscure the parasites after staining.
Anticoagulant
If blood containing an anticoagulant is used, 2 or 3 drops may be spread over an area about 2 cm in diameter; it is not necessary to continue stirring for 30 s, since fibrin strands do not form. If the blood is too thick or any grease remains on the slide, the blood may flake off during staining. It is far better to make the preparation too thin, rather than too thick.
Allow the thick film to air dry (room temperature) in a dust-free area. Never apply heat to a thick film, since heat will fix the blood, causing the RBCs to remain intact during staining; the result is stain retention and inability to identify the parasites. After the thick films are thoroughly dry, they can be laked (lysed) to remove the hemoglobin. Rupture of the RBCs during laking removes the RBCs from the final stained blood film; the only structures remaining on the thick film are the white blood cells (WBCs), the platelets, and any parasites present. To lake the films, place them in buffer solution before staining or directly into Giemsa stain, which is an aqueous stain. If thick films are to be stained later, they should be laked before storage. Potential problems with the preparation and staining of thick blood films can be seen in Table 7.3.
The thin blood film is routinely used for specific parasite identification, although the number of organisms per field is much reduced compared with the thick film. The thin film is prepared exactly as one used for a differential count (Fig. 7.1). A well-prepared film is thick at one end and thin at the other (one layer of evenly distributed RBCs with no cell overlap). The thin, feathered end should be at least 2 cm long, and the film should occupy the central area of the slide, with free margins on both sides. The presence of long streamers of blood indicates that the slide used as a spreader was dirty or chipped. Streaks in the film are usually caused by dirt, and holes in the film indicate the presence of grease on the slide. After the film has air dried (do not apply heat), it may be stained. The necessity for fixation before staining will depend on the stain selected. Since Giemsa is an aqueous stain, laking of the RBCs occurs during the staining process. However, when Wright’s stain is used, a fixing agent is incorporated into the stain, so that laking of the blood films must occur prior to 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.
Figure 7.1 Method for preparation of thin blood film. (A) Position of spreader slide; (B) well-prepared thin film. Arrows indicate the area of the slide (feather edge) used to observe accurate cell morphology. (Illustration by Sharon Belkin.) doi:10.1128/9781555819002.ch7.f1
The instrument-prepared monolayer method or coverslip methods generally do not provide the best morphology for malarial parasites within the RBCs. However, the selection of a slide preparation method can be dictated by personal preference, since a malarial infection can be diagnosed from either type of slide. Potential problems with the preparation and staining of thin blood films can be seen in Table 7.4.
Combination Thick and Thin Blood Films (on the Same Slide)
In some instances (field surveys), it is helpful to prepare slides with both a thick and a thin film on the same slide. With this type of preparation, remember the following.
1. Sufficient time must be allowed for the thick portion of the smear to dry before staining.
2. If Giemsa stain is used, the thin film only must be fixed in absolute methanol before staining.
Combination Thick and Thin Blood Films (Can Be Stained as Either)
To prepare a slide containing blood that can be stained for either a thick or thin blood film, the following method was developed (Fig. 7.2). The specimen usually consists of fresh whole blood collected by finger stick or whole blood containing EDTA collected by venipuncture less than 1 h earlier.
Figure 7.2 Method of thick-thin combination blood film preparation. (a) Position of the drop of EDTA-containing blood. (b) Position of the applicator stick in contact with blood and glass slide. (c) Rotation of the applicator stick. (d) Completed thick-thin combination blood film prior to staining. (Illustration by Sharon Belkin.) Reprinted from reference 9. doi:10.1128/9781555819002.ch7.f2
Visually, the smear should consist of alternating thick and thin portions throughout the length of the glass slide. One should be able to barely read newsprint through the wet or dry film. Also, the film itself should not have any clear areas or smudges, indicating that grease or fingerprints were on the glass.
Detailed Procedure
1. Wear gloves when performing this procedure or preparing any blood films.
2. The procedure depends on the source of the specimen.
A. Blood from a finger puncture is not recommended, since the procedure does not lend itself to “stirring” to prevent fibrin strands.
B. For blood from venipuncture, place a clean 1- by 3-in. glass microscope slide on a horizontal surface. Place a drop (30 to 40 µl) of blood onto one end of the slide about 0.5 in. from the end. Using an applicator stick lying across the glass slide and keeping the applicator in contact with the blood and glass, rotate (do not “roll”) the stick in a circular motion while moving the stick down the glass slide to the opposite end. The appearance of the blood smear should be alternate thick and thin areas of blood that cover the entire slide. Immediately place the film over some small print and be sure that the print is just barely readable.
3. Allow the film 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.
4. This slide can be stained as either a thick or thin blood film.
5. Label the slide appropriately.
6. 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. Rapid stains are also acceptable.
Procedure Limitations
1. If the smears are prepared from anticoagulated blood that is more than 1 h old, the morphology of the parasites may not be typical and the film may wash off the slide during the staining procedure. If a tube of blood containing EDTA cools to room temperature and the cap has been removed, several parasite changes can occur. The parasites within the RBCs will respond as if they were now inside the mosquito after being taken in with a blood meal. The morphology of these changes in the life cycle and within the RBCs can cause confusion when examining blood films prepared from this blood (3–8).
A. Stippling (Schüffner’s dots) may not be visible.
B. The male gametocyte (if present) may exflagellate and may resemble spirochetes (Fig. 7.3).
Figure 7.3 (Left) Photograph of exflagellation of the malarial microgametocyte. This may occur when anticoagulated blood is left standing at room temperature for some time prior to smear preparation. The life cycle of the parasite continues in the tube of blood as it would if the parasite had been ingested by the mosquito during a blood meal. (Right) Borrelia in blood. Note that the Plasmodium microgametes can resemble these organisms, particularly if they appear free in the background of the smear. doi:10.1128/9781555819002.ch7.f3
C. The ookinetes of Plasmodium species other than P. falciparum may develop as if they were in the mosquito and may mimic the crescent-shaped gametocytes of P. falciparum.
2. Identification to species, particularly between P. ovale and P. vivax and between the ring forms of P. falciparum and Babesia spp., may be impossible without examining one of the slides stained as a thin blood film. Also, Trypanosoma cruzi trypomastigotes are frequently distorted in thick films.
3. Excess stain deposition on the film may be confusing and make the detection of organisms difficult.
4. If the EDTA blood is 4 to 6 h old prior to blood films preparation, parasites are actually lost from the specimen; the parasitemia may be too low or represent a false negative.
Fixation of Thin Blood Films: Acetone Dip for Thick Blood Films
Thin-Film Fixation. Thin blood films must be completely dry before being fixed in absolute methanol (Giemsa staining). Drying times will be extended if the humidity 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 (12) has recommended the addition of 1% (by volume) 2,6-di-t-butyl-p-cresol (butylated hydroxytoluene [Sigma Chemical Co. catalog no. B1253]) to the mounting medium. Without the addition of this antioxidant, mounted stained smears eventually become pink; stains protected with this compound remain unchanged in color for many years.
Note Any slide that is protected by a coverglass and is going to be examined with an oil immersion lens must be covered by a no. 1 coverglass. If a no. 2 coverglass is used, the extra thickness may prevent the oil immersion lens from focusing properly.
Giemsa stain is sold as a concentrated stock solution. Each new lot should be tested for optimal staining times before being used on patient specimens. If the blood cells appear to be adequately stained, the timing and stain dilution should be appropriate to demonstrate the presence of malarial and other parasites. Giemsa stain is also available as a powder for those who wish to make up their own stain. The use of prepared liquid stain or stain prepared from the powder depends on personal preference; there is apparently little difference between the two preparations. Directions for preparing the stain follow (4, 5, 9).
Note Quality control for blood film stains can be any negative or positive blood film; it is not necessary for the control slides to contain actual parasites. The actual patient slide being stained serves as its own control; if the WBCs look acceptable, any parasites present will also be acceptable in terms of morphology and staining colors. If the RBCs and WBCs stain correctly, any parasites present would also stain correctly with the same nuclear and cytoplasmic colors as the blood cells (see Fig. 7.4 to 7.8).
Reagents
Stock Giemsa Stain
1. Grind together small portions of stain and glycerin in a mortar, and collect mixtures in a 500- or 1,000-ml flask until all measured material is mixed.
2. Stopper the flask with a cotton plug, cover the plug with heavy paper, and place the flask in a 55 to 60°C water bath for 2 h. Make sure the water in the water bath is above the level of the stain. Shake gently at 30-min intervals.
3. After grinding the powder and glycerin in the mortar, use the 50 ml of methyl alcohol to wash the last bit of stain from the mortar; then pour the alcohol into a small, airtight bottle.
4. Remove the glycerin-stain powder mixture from the water bath, and allow it to cool to room temperature. Add alcohol washing from the mortar, and shake well.
5. Before use, filter through Whatman no. 1 paper into a brown bottle. Although the stain can be used immediately, it is better to let it stand for 2 to 3 weeks with intermittent shaking.
6. Label and store protected from light; the shelf life is 36 months, provided that results are within quality control guidelines.
Note The stock stain is stable for many years; however, it must be protected from moisture. The staining reaction is oxidative; any oxygen in water will initiate the staining reaction and destroy the stock stain. This is why the aqueous working solution of stock stain is good only for 1 day.
10% Stock Solution of Triton X-100
Mix thoroughly, and store at room temperature. This solution will keep indefinitely if kept tightly stoppered.
Stock Buffers
Disodium Phosphate (Dibasic)
Monosodium Phosphate (Monobasic)
Phosphate-Buffered Water (Table 7.6)
Triton-Phosphate Buffer
0.01% Triton-Buffered Water Stock
0.1% Triton-Buffered Water
For thin blood films or a combination of thin and thick blood films, use 0.01% Triton-buffered water; for thick blood films, use 0.1% Triton-buffered water.
Liquid
The commercial liquid stain or the stock solution prepared from powder should be diluted approximately the same amount to prepare the working stain solution. Stock Giemsa liquid stain is diluted 1:10 with buffer for thin blood films; for thick films, a dilution of up to 1:50 may be used. Some people prefer to stain both thick and thin smears for a longer period in a more dilute solution. Phosphate buffer used in dilution of the stock stain should be neutral or slightly alkaline. Phosphate buffer solution may be used to obtain the right pH (Table 7.6). In some laboratories, tap water has a satisfactory pH and may be used for the entire staining procedure and the final rinse. Some workers recommend using pH 6.8 to emphasize Schüffner’s dots.
Procedure for Staining Thin Films
1. Fix blood films in absolute methyl alcohol (acetone free) for 1 min.
2. Allow the slides to air dry.
3. Immerse the slides in a solution of 1 part Giemsa stock (commercial liquid stain or stock prepared from powder) to 10 to 50 parts phosphate buffer (pH 7.0 to 7.2). Stain for 10 to 60 min. Fresh working stain should be prepared from stock solution each day.
Note A good general rule for stain dilution versus staining time is as follows. If the dilution is 1:20, stain for 20 min; if the dilution is 1:30, stain for 30 min, etc. However, a series of stain dilutions and staining times should be tried to determine the best dilution and time for each batch of stock stain.
4. Dip the slides briefly in phosphate-buffered water, or rinse under gently running tap water.
Note Excessive washing will decolorize the film.
5. Drain thoroughly in a vertical position, and allow to air dry. The bottom of the slide should be wiped to remove excess stain before drying.
Procedure for Staining Thick Films
The procedure to be followed for thick films is the same as for thin films, except that the first two steps are omitted. If the slide has a thick film at one end and a thin film at the other, fix only the thin portion and then stain both parts of the film simultaneously. Normally, a stain/buffer dilution of 1:50 (vol/vol) for a staining time of 50 min is recommended for thick films. The longer staining time seems to give better results than do the shorter times that can be used for the thin film.
Results
Giemsa stain colors the components of blood as follows: RBCs, pale red; nuclei of WBCs, purple with pale purple cytoplasm; eosinophilic granules, bright purple-red; and neutrophilic granules, deep pink-purple. If malaria parasites are present, the cytoplasm stains blue and the nuclear material stains red to purple-red. Schüffner’s dots and other inclusions in the RBCs will stain red (Fig. 7.4). However, as seen in Fig. 7.4, there can be tremendous variation in color. Nuclear and cytoplasmic staining characteristics of the other blood parasites such as Babesia spp., trypanosomes, and leishmaniae are like those of the malaria parasites (Fig. 7.5 to 7.8). While the sheath of microfilariae may not always stain with Giemsa, the nuclei within the microfilaria itself stain blue to purple. The sheath of Brugia malayi will tend to stain pink, while the sheath of Wuchereria bancrofti may not stain with Giemsa (Fig. 7.9).
Figure 7.4 Plasmodium spp. seen in stained thin blood films. (Left column from top to bottom) Plasmodium vivax: (1) developing ring (note the enlarged RBC, lack of Schüffner’s dots [EDTA blood], and ameboid rings); (2) developing trophozoite (note the enlarged RBC and Schüffner’s dots); (3) mature schizont with approximately 16 merozoites; (4) thick blood film showing developing ring forms; (5) normal WBCs (including polymorphonuclear leukocytes [PMNs], eosinophil, basophil, monocyte, and lymphocyte). (Second column from top to bottom) Plasmodium ovale: (1) young ring form (note the nonameboid ring and Schüffner’s dots that appear earlier than in rings of P. vivax); (2) developing trophozoites (note Schüffner’s dots and fimbriated edges of the infected RBCs); (3) maturing schizont containing merozoites and malarial pigment; (4) thick blood film showing developing schizont; (5) normal PMNs. (Third column from top to bottom) Plasmodium malariae: (1) young ring (note the normal-sized RBC and no stippling); (2) developing trophozoite (note the normal-sized RBC and “band form” configuration of the trophozoite); (3) mature schizont (note the size of the RBC) containing approximately eight merozoites and brownish malarial pigment; (4) thick blood film with mature schizonts containing developing merozoites; (5) normal WBCs (including PMN, eosinophil, basophil, monocyte and lymphocyte). (Last column from top to bottom) Plasmodium falciparum: (1) typical ring forms (note appliqué form at the side of the RBC and multiple rings/cell); (2) typical ring forms (note the ring appearing to be partially outside the RBC—typical for P. falciparum); (3) crescent-shaped gametocyte; (4) thick blood film showing many ring forms; (5) typical PMNs. doi:10.1128/9781555819002.ch7.f4
Figure 7.5 Trypomastigotes. (Left) Trypanosoma brucei gambiense. (Right) Trypanosoma cruzi (note the undulating membrane on both trypomastigotes and the larger kinetoplast in T. cruzi). doi:10.1128/9781555819002.ch7.f5
Figure 7.6 Babesia sp. (Top) Note some of the ring forms are outside the RBCs (box, rare to see in Plasmodium infections). (Bottom) Note the numerous ring forms per RBC; also note the Maltese cross ring formation in the lowermost RBC (arrow). doi:10.1128/9781555819002.ch7.f6
Figure 7.7 Leishmania donovani in an impression smear. Note the numerous small amastigotes containing a nucleus and bar-shaped primitive flagellum. doi:10.1128/9781555819002.ch7.f7
Figure 7.8 Plasmodium falciparum. (Upper row) Note the ring forms (multiple rings per RBC and presence of the “headphone” ring configuration). The photograph at the far right is a good mimic of Babesia organisms, but the rings are not quite as pleomorphic as in Babesia spp. (Lower row) Two examples of P. falciparum gametocytes, two of which appear to be outside the RBC. However, although the RBC membrane is not visible, the gametocyte is intracellular. The image on the far right is an ookinete from the mosquito cycle; this can occur within the EDTA blood specimen if the blood cools and is left standing with the cap removed. In the cooled, aerated blood, the parasites begin the cycle that normally occurs within the mosquito. This artifact can easily be confused with the crescent-shaped gametocyte. doi:10.1128/9781555819002.ch7.f8
Figure 7.9 (Upper) Brugia malayi microfilaria on a blood film stained with Giemsa stain. Note the presence of the sheath (stains pink with Giemsa stain). (Lower) Brugia malayi microfilaria on a thick blood film stained with Giemsa stain (note some of the individual nuclei are visible). (Images courtesy of the CDC Public Health Image Library; lower, photograph by Dr. Mae Melvin.) doi:10.1128/9781555819002.ch7.f9
Wright’s stain is available commercially in liquid form, ready to use, and also as a powder which must be dissolved in anhydrous, acetone-free methyl alcohol before use. Directions for preparing the stain follow.
Reagent
Wright’s Stain
1. Grind 0.9 g of Wright’s stain powder with 10 to 15 ml of methanol (anhydrous, acetone free) in a clean mortar. Gradually add methanol while grinding. As the dye is dissolved in the methanol, pour that solution off and add more methanol to the mortar. Repeat this process until the entire 500 ml of methanol has been used.
2. Store the stain in a tightly stoppered glass bottle at room temperature. Shake the bottle several times daily for at least 5 days.
3. Allow the precipitate to settle, and pour off some of the supernatant fluid into dropping bottles for use. If the stock solution has been disturbed, the supernatant fluid can be filtered through Whatman no. 1 paper into a brown bottle.
4. Label and store protected from light; the shelf life is 36 months, provided that results are within quality control guidelines.
The staining procedure requires phosphate buffer solutions (see directions in the preceding discussion of Giemsa stain); the pH required for Wright’s stain is 6.6 to 6.8.
Procedure for Staining Thin Films
Since Wright’s stain contains alcohol, the slides do not require fixation before staining.
1. Place a slide on a rack in a horizontal, level position, and cover the surface with stain.
2. Count the number of drops of stain needed to cover the surface. Let stand for 1 to 3 min (the optimal staining time varies with each batch of stain).
3. Add an equal number of drops of phosphate-buffered water to the slide; mix the stain and buffer by blowing on the surface of the fluid.
4. After 4 to 8 min, flood the stain from the slide with phosphate buffer. Do not pour the stain off before washing. If you do, a precipitate will be deposited on the slide.
5. Wipe the bottom of the slide to remove excess stain.
6. Allow the slide to drain and air dry.
Procedure for Staining Thick Films
Thick films stained with Wright’s stain are usually inferior to those stained with Giemsa solution. Great care should also be taken to avoid excess stain precipitate on the slide during the final rinse. Before being stained, thick films must be laked in distilled water (to rupture and remove RBCs) and air dried. The staining procedure is the same as for thin films, but the staining time is usually somewhat longer and must be determined for each batch of stain.
Results
Wright’s stain colors blood components as follows: RBCs, light tan, reddish, or buff; nuclei of WBCs, bright blue with contrasting light cytoplasm; eosinophilic granules, bright red; and neutrophilic granules, pink or light purple.
If malaria parasites are present, the cytoplasm stains pale blue and the nuclear material stains red. Schüffner’s dots and other inclusions in the RBCs usually do not stain or stain very pale with Wright’s stain. Nuclear and cytoplasmic staining characteristics of the other blood parasites such as Babesia spp., trypanosomes, and leishmaniae are like those of the malaria parasites. While the sheath of microfilariae may not always stain with Wright’s stain, the nuclei within the microfilaria itself stain pale to dark blue.
General Notes on Staining Procedures
1. When large numbers of slides are being processed, remember that dry films should be stored in dust-free containers before staining, to protect fresh smears from insects.
2. If the slides cannot be stained within 48 h, thin films should be fixed in methyl alcohol and thick films should be laked in distilled water before storage (some users prefer buffered water at pH 6.8 to 7.2). Rapid stains can also be used. Parasites will stain like the WBCs.
3. Slides should be stored at reasonably low temperatures (below 80°F [ca. 27°C]) before being stained. If the slides are exposed to high temperatures, thick films that have not been laked will become heat fixed; hemoglobin will remain fixed in the RBCs, and heavy stain retention will then prevent parasite identification. Thin films also stain poorly after exposure to high temperatures.
4. Fresh working stain should be prepared just before use. If a large number of thick films are being laked during staining, the stain should be changed after 50 slides because of the accumulation of excess hemoglobin.
Proper Examination of Thin and Thick Blood Films
In any examination of thin blood films for parasitic organisms, the initial screen should be carried out with the low-power objective of a microscope. Microfilariae may be missed if the entire thin film is not examined. Microfilariae are rarely present in large numbers, and frequently only a few organisms occur in each thin-film preparation. Microfilariae are commonly found at the edges of the thin film or at the feathered end of the film, because they are carried to these sites during the process of spreading the blood. The feathered end of the film where the RBCs are drawn out into one single, distinctive layer of cells should be examined for the presence of malaria parasites and trypanosomes. In these areas, the morphology and size of the infected RBCs are most clearly seen.
Depending on the training and experience of the microscopist, examination of the thin film usually takes 5 to 10 min or less (200 to 300 oil immersion fields) at a magnification of ×1,000. Although some people use a 50× or 60× oil immersion objective to screen stained blood films, there is some concern that small parasites such as plasmodia, Babesia spp., or L. donovani may be missed at this smaller total magnification (×500 or ×600) compared with the ×1,000 total magnification obtained with the more traditional 100× oil immersion objective. Because people tend to scan blood films at different rates, it is important to examine a minimum number of fields, regardless of the time it takes to perform this procedure. If something suspicious has been seen in the thick film, the number of fields examined on the thin film is often considerably more than 200 to 300. The request for blood film examination should always be considered a STAT procedure, with all reports (negative as well as positive) being reported immediately to the physician as soon as possible. Appropriate governmental agencies (local, state, and federal) should be notified within a reasonable time frame in accordance with guidelines and laws.
Diagnostic problems with the use of automated differential instruments have been reported (13, 14). Both malaria and Babesia light infections (like those seen with travelers with no prior exposure to malaria) were missed with these instruments, and therapy was delayed. Although these instruments are not designed to detect intracellular blood parasites, the inability of the automated systems to discriminate between uninfected RBCs and those infected with parasites may pose serious diagnostic problems.
In the preparation of a thick blood film, the greatest concentration of blood cells will be in the center of the film. A search for parasitic organisms should be carried out initially at low magnification to detect microfilariae more readily. Examination of a thick film usually requires 3 to 5 min (approximately 100 oil immersion fields). Search for malarial organisms and trypanosomes is best done under oil immersion (total magnification, ×1,000). Intact RBCs are frequently seen at the very periphery of the thick film; such cells, if infected, may prove useful in malaria diagnosis, since they may demonstrate the characteristic morphology necessary to identify the organisms to the species level.
It is important to report the level of parasitemia when blood films are reviewed and found to be positive for malaria parasites. Because of the potential for drug resistance in some of the Plasmodium species, particularly P. falciparum and P. vivax, it is important that every positive smear be assessed and the parasitemia reported exactly the same way on follow-up specimens as on the initial specimen. This allows the parasitemia to be monitored after therapy has been initiated. In cases where the patient is hospitalized, monitoring should be performed at 24, 48, and 72 h after initiating therapy. Generally, the parasitemia drops very quickly within the first 2 h; however, in cases of drug resistance, the level may not decrease but actually may increase over time.
Malarial infections should be reported as the percentage of infected RBCs per 100 RBCs counted (0.5%, 1.0%, etc.) (3). Considering the low parasitemia frequently seen in patients within the United States, several hundred RBCs may have to be counted to arrive at an accurate count and determination of the percentage. The thin blood film must be used for this approach.
Another approach is to count the number of parasites per 100 WBCs on the smear. Either the thick or thin film can be used for this purpose. This figure can be converted to the number of parasites per microliter of blood; divide the number of parasites per 100 WBCs by 100, and multiply that figure by the number of WBCs per microliter of blood. Depending on the parasitemia, 200 or more WBCs may have to be counted, so the denominator may vary (it may be 200 or even more). In this case, blood for both the peripheral smears and cell counts must be collected at the same time.
It is critical that the same reporting method be used consistently for every subsequent set of blood films so that the parasitemia can be tracked for decrease or possible increase, indicating resistance. Also, remember that drug resistance may not become evident for a few days. The parasitemia may initially appear to decline but may then begin to increase after several days. Therefore, it is very important that patient parasitemia be monitored, particularly if an infection with P. falciparum has been diagnosed. Drug resistance has also been reported in P. vivax cases; mixed infections are also much more common than suspected.
Although therapy for Babesia infections is usually quite effective, calculation of parasitemia for positive Babesia infections is highly recommended. Babesiosis usually results in self-cure or cure after administration of azithromycin plus atovaquone or clindamycin plus quinine. Although certain individuals who are severely immunocompromised may not respond to these drug regimens, there has been no reported evidence that treatment failure is due to drug-resistant strains of Babesia microti. Preliminary studies indicate that reduction of pathogen loads and WBC inactivation within blood components using riboflavin and ultraviolet light have been used as an alternative for gamma-irradiation for B. microti, Babesia divergens, T. cruzi, HIV, and bacteria (15).
Diagnosis of Malaria: Review of Alternatives to Conventional Microscopy
It is well known that malaria causes significant morbidity and mortality worldwide, including in countries where imported cases are seen. In many developing countries where malaria is highly endemic, diagnostic testing is often inadequate or unavailable due to a lack of trained personnel or funds or both. Although microscopic examination of stained thick and thin blood films remains the standard of practice, this approach is time-consuming, is based on the need for a great deal of expertise in microscopic morphology, and requires the purchase and maintenance of expensive equipment. Rapid diagnostic tests (RDTs) offer great potential to improve the diagnosis of malaria, particularly in remote areas (World Health Organization. 2012. Malaria rapid diagnostic test performance. Summary results of WHO product testing of malaria RDTs: Round 1–4 [2008–2012]; http://www.who.int/malaria/publications/rdtmalaria_ [accessed 6/20/2013]). This provides very comprehensive lists of product testing to date. There are a number of new approaches to the diagnosis of malaria, including the use of fluorescent stains (QBC), dipstick antigen detection of histidine-rich protein 2 (HRP2), parasite lactate dehydrogenase (pLDH), and Flow anti-pLDH Plasmodium monoclonal antibodies), PCR, and automated blood cell analyzers. Parasitemia and its clinical correlates are given in Table 7.7, while some of the testing options for malaria can be found in Table 7.8. Testing options for some of the other blood parasites can be seen in Table 7.9.
Histidine-rich protein 2 of P. falciparum (PfHRP2) is a water-soluble protein that is produced by the asexual stages and gametocytes of P. falciparum, expressed on the RBC membrane surface, and shown to remain in the blood for at least 28 days after the initiation of antimalarial therapy. Many RDTs are based on the detection of PfHRP2, but reports from field tests have questioned their sensitivity and reliability. However, the variability in the results of PfHRP2-based RDTs may be related to the variability in the target antigen (16). This hypothesis was tested by examining the genetic diversity of PfHRP2, which includes numerous amino acid repeats, in 75 P. falciparum lines and isolates originating from 19 countries and testing a subset of parasites by use of two PfHRP2-based RDTs. There is extensive diversity in PfHRP2 sequences, both within and between countries. Logistic regression analysis indicated that two types of repeats were predictive of RDT detection sensitivity (87.5% accuracy), with predictions suggesting that only 84% of P. falciparum parasites in the Asia-Pacific region are likely to be detected at densities of ≤250 parasites/µl. Data also indicate that PfHRP3 may play a role in the performance of PfHRP2-based RDTs. These findings provide an alternative explanation for the variable sensitivity in field tests of malaria RDTs that is not due to the quality of the RDTs (16).
The persistence of parasite HRP2 in the circulation after parasite clearance has been considered a drawback for RDTs targeting HRP2 and a major cause of false-positive results. In one study when PCR was used as the gold standard rather than microscopy, the high rate of RDT false-positive parasitemia results in comparison with microscopy was shown to predominantly represent cases that had a parasite density below the threshold for detection by microscopy (17). Despite the generally low disease-endemic prevalence of malaria in the area, there was a high prevalence of chronic infections with low, fluctuating parasite densities that were better detected by RDT. In areas known to have low-density parasitemias, RDTs targeting HRP2 may increase the diagnostic sensitivity in comparison with microscopy. While microscopy remains the standard for comparison of the diagnostic accuracy for malaria, the limitations of microscopy, and the possibility that RDTs may have superior accuracy in some circumstances, should be taken into account when interpreting the results of diagnostic trials.
To determine the accuracy of RDTs for ruling out malaria in nonimmune travelers returning from areas where malaria is endemic, 21 studies and 5,747 individuals were surveyed from the published literature (18). Diagnostic accuracy studies of nonimmune individuals with suspected malaria were included if they compared rapid tests with expert microscopic examination or PCR tests. The authors concluded that RDTs for malaria may be a useful diagnostic adjunct to microscopy in centers without major expertise in tropical medicine. Initial decisions on treatment initiation and choice of antimalarial drugs can be based on travel history and posttest probabilities after rapid testing. However, expert microscopy is still required for species identification and confirmation.
Diagnostic accuracy of RDTs for self-diagnosis was variable, and requires improvements before widespread acceptance (19). RDTs have limitations that include the difficulty in detecting mixed infections, all species of Plasmodium, and infections at low concentrations of parasites, along with an inability to monitor response to therapy. Also, if the result is negative, microscopy is still mandatory. Therefore RDTs do not eliminate the need to examine thick and thin blood films. The maintenance of expert microscopy remains a diagnostic priority until a new gold standard is developed.
QBC Microhematocrit Centrifugation Method
Microhematocrit centrifugation with use of the QBC malaria tube (glass capillary tube and closely fitting plastic insert; CBC malaria blood tubes [Becton Dickinson, Tropical Disease Diagnostics, Sparks, MD]) has been used for the detection of blood parasites (20, 21). At the end of centrifugation of 50 to 60 µl of capillary or venous blood (5 min in a QBC centrifuge; 14,387 × g), parasites or erythrocytes containing parasites are concentrated into a small, 1- to 2-mm region near the top of the RBC column and are held close to the wall of the tube by the plastic float, thereby making them readily visible by microscopy. Tubes precoated with acridine orange provide a stain which induces fluorescence in the parasites. This method automatically prepares a concentrated smear, which represents the distance between the float and the walls of the tube. Once the tube is placed into the plastic holder (Paraviewer) and immersion oil is applied onto the top of the hematocrit tube (no coverslip is necessary), the tube is examined with a ×40 to ×60 oil immersion objective (there must be a working distance of 0.3 mm or greater) (Fig. 7.10).
Figure 7.10 The QBC Malaria Test components include the ParaLens UV microscope adapter (blue-violet light module providing fiber-optic illumination with AC outlet or bulb illumination) with rechargeable battery for mobile use (attachable to any conventional microscope), the ParaFuge battery-powered centrifuge, and the ParaViewer microscope tube holder, a specially designed QBC tube viewing block that accepts standard microscope oil. doi:10.1128/9781555819002.ch7.f10
Note Although a malaria infection could be detected by this method, appropriate thick and thin blood films must be examined to accurately identify the species of the organism causing the infection.
The range of sensitivities for the QBC test has been as low as 55% to >90% compared with microscopy. While most workers think the method is rapid, some think there are some disadvantages such as the high cost of capillary tubes and equipment, problems in species identification and quantification, technical problems, broken capillaries, and the fact that the capillaries cannot be stored for later reference. This may not be the most appropriate approach for a laboratory that receives a small number of specimens for malaria testing, but it could be helpful in other settings. In a recent study of 745 specimens from an area of low endemicity along the Colombian Pacific coast, the agreement between the QBC method and thick blood smear was reported to be 99.5% (22). However, another study concluded that in spite of the simplicity of the QBC method, it cannot be considered an acceptable alternative to stained thick blood films in routine laboratory situations (23). An excellent review of this and other methods is provided in reference 24.
This method has been reported to have a high degree of sensitivity in the detection of cases of human filariasis (25). In one study evaluating the technique in the detection of canine Dirofilaria immitis as a model for human filariasis, the QBC analysis was more sensitive (55%) than the thick blood smear (39%) and was more efficient (79% versus 72%). However, accurate identification to the species level was impossible using the QBC method.
Diagnosis of tick-borne relapsing fever has also been reported using the QBC system with a high level of sensitivity, more so than the thick blood smear. This method seems to be useful for this purpose and might be considered in the management of fever in travelers returning from tropical regions (26).
A P. falciparum antigen detection system, the ParaSight F test (Becton Dickinson), is very effective in field trials (27). This procedure is based on an antigen capture approach and has been incorporated in a dipstick format; the entire test takes approximately 10 min (Fig. 7.11 and 7.12) (27). The overall performance of the ParaSight F test was reviewed in 1995 at a World Health Organization meeting looking at 15 studies with a total of 7,926 assays for the detection of P. falciparum. Overall, the sensitivity ranged from 84 to 94%, with specificities ranging from 81 to 99% (28). However, the test detects only P. falciparum, and at low parasitemias (100/µl) the sensitivity drops to 11 to 40%. Also, since HRP2 is not present in mature gametocytes, cases where only gametocytes were present could be missed. There is also a possibility that some false-negative results, even at high parasitemias, are due to the lack of the HRP2 gene (29). After successful therapy, many patients continue to have circulating HRP2 antigen for 7 to 14 days after microscopic and clinical cure. Also, as many as 60% of patients positive for rheumatoid factor have a false-positive test (30). The ParaSight F test may be used in situations where no trained microscopists are available or where malaria is strongly suspected and the microscopy results are negative. This kit has proven to be very useful in many areas of the world.
Figure 7.11 Diagram of the ParaSight F test format. (Adapted from reference 24 with permission.) doi:10.1128/9781555819002.ch7.f11
Figure 7.12 ParaSight F test showing (from left to right) the positive test strip with a reagent control mark above the positive test result and a negative test strip with the reagent control mark. doi:10.1128/9781555819002.ch7.f12
In one systematic review and meta-analysis of controlled studies evaluating the diagnostic accuracy of the ParaSight F test in comparison with light microscopy, data sources included 15,359 subjects (4,119 with P. falciparum malaria) in 32 studies reported in 29 publications. Overall, the ParaSight F test demonstrated 90% sensitivity and 94% specificity. Both the sensitivity and specificity were significantly higher in the nonresident population than in the resident population. The posttest probability indicates that in settings of low malaria prevalence a negative test almost absolutely excludes infection, while in settings of high prevalence the same result still gives a substantial chance of infection being present. The authors conclude that the ParaSight F test is a simple and accurate test for the diagnosis of P. falciparum infection. The test could be of particular value in the diagnosis of malaria in travelers returning from areas of endemic infection (31).
The ICT Malaria P.f. test (ICT Diagnostics, Brookvale, New South Wales, Australia) was the first malaria rapid diagnostic device designed for convenience of use in a booklet format with the test strip mounted on cardboard. Like other nonmicroscopic malaria rapid tests, ICT Malaria P.f. was an immunochromatographic assay. The original ICT assay detected only P. falciparum HRP2. By 1999, AMRAD (French’s Forest, New South Wales, Australia) acquired ICT Diagnostics and continued manufacturing the product. At about the same time, the product was enhanced by adding the capability to detect non-falciparum malaria. The refined assay was renamed ICT Malaria P.f./P.v. This improvement was achieved by using monoclonal antibodies to capture Plasmodium aldolase, in addition to the HRP2 test. Plasmodium aldolase is an enzyme of the parasite glycolytic pathway expressed by the blood stages of P. falciparum as well as the non-falciparum malaria parasites. Monoclonal antibodies against Plasmodium aldolase are panspecific in their reaction and have been used in a combined P. falciparum-P. vivax immunochromatographic test that targets the panmalarial antigen along with PfHRP2.
In July 2000, AMRAD ceased the production of ICT and sold its ICT division to Binax, Inc. (Portland, ME), where further developmental work was done to refine the test. The new ICT test was released under the name NOW ICT Malaria P.f./P.v. for Whole Blood and is presently called the Binax NOW malaria test for whole blood (Binax, Inc.). The NOW malaria test uses an immunochromatographic format (Fig. 7.13). The test is now available from Alere (Alere, Inc., Waltham, MA) and is FDA approved for use within the United States.
Figure 7.13 Diagram of the NOW malaria test format. (Adapted from package insert, Binax, Portland, ME.) doi:10.1128/9781555819002.ch7.f13
Prior to the development of the improved product (Binax NOW malaria test), in a number of studies testing over 1,300 assays, the overall sensitivity ranged from 80 to 100% (32). Although some of the same issues apply to this test as to the ParaSight F test, the false-positive rate from the presence of rheumatoid factor appears to be less of an issue (33).
In many areas of the world where both P. falciparum and P. vivax occur, rapid diagnostic tests for malaria must be able to differentiate the two species. It is also interesting that when a mixed-species infection with both P. falciparum and P. vivax is misdiagnosed as a single-species P. vivax infection, treatment for P. vivax can lead to a surge in P. falciparum parasitemia. The combined P. falciparum-P. vivax immunochromatographic test (NOW ICT Malaria Pf/Pv) was used in Indonesia, where both species occur (34). Blinded microscopy was used as the gold standard, with all discordant and 20% of concordant results cross-checked blindly. Of those with a presumptive clinical diagnosis of malaria, only 50% were parasitemic. The NOW ICT Malaria Pf/Pv test was sensitive (95.5%) and specific (89.8%) for the diagnosis of P. falciparum malaria, with a positive predictive value of 88.1% and a negative predictive value of 96.2%. Although the specificity and negative predictive value for the diagnosis of P. vivax malaria were 94.8 and 98.2%, respectively, the overall sensitivity of 75% and positive predictive value of 50% were lower than desired. With parasitemias of >500/µl, the sensitivity for the diagnosis of P. vivax malaria was 96%; with parasitemias of <500/µl, it was only 29%. However, by using this test, undertreatment rates would be reduced from 14.7 to 3.6% with a modest increase in the rate of overtreatment of microscopy-negative patients from 7.1 to 15.4%. Unfortunately, cost remains an obstacle to widespread implementation.
In a study using the improved product (NOW malaria test), the sensitivity for P. falciparum was 100% and the specificity was 96%; the sensitivity for P. vivax was 89%, and the specificity was 98%. Testing was performed using P. falciparum- and P. vivax-positive specimens. These results suggest improved performance over NOW ICT predecessors (Fig. 7.13) (35). In another study, the NOW ICT test showed a sensitivity of 94% for the detection of P. falciparum malaria (96% for pure P. falciparum infection) and 84% for non-P. falciparum infections (87% for pure P. vivax infections and 62% for pure P. ovale and P. malariae infections) compared with PCR, with an overall specificity of 99% (36). The Binax NOW ICT may represent a useful adjunct for the diagnosis of P. falciparum and P. vivax malaria in febrile returned travelers. However, particularly in this patient group and in tests with a negative result, the rapid test should be performed in association with more traditional methods such as examination of thick and thin blood films (37, 38).
Flow Anti-pLDH Plasmodium Monoclonal Antibodies
pLDH is a soluble glycolytic enzyme produced by the asexual and sexual stages of the live parasites and is present in and released from the parasite-infected erythrocytes. It has been found in all five human malaria species, and different isomers of pLDH for each of the five species exist. With pLDH as the target, a quantitative immunocapture assay, a qualitative immunochromatographic dipstick assay using monoclonal antibodies, an immunodot assay, and a dipstick assay using polyclonal antibodies have been developed.
Flow anti-pLDH Plasmodium monoclonal antibodies detect a malaria pLDH antigen by using an antibody incorporated into the dipstick format (39) (Fig. 7.14). The test is positive only when viable parasites are present. Although the test has both a P. falciparum-specific and a panspecific antibody against all four species, the panspecific antibody detects only P. vivax with any degree of consistency (40, 41). The sensitivity and specificity of the Flow monoclonal antibodies are comparable to those of microscopy in detecting malaria infections at a parasitemia of >100/µl; however, the test failed to identify more than half of the patients with a parasitemia of <50/µl. In one study, the sensitivity of the Flow monoclonal antibodies was 97% at a high parasitemia (>100/µl) but fell to 39% at <50/µl (42). Therefore, the test is similar to the HRP2 assays and should not replace conventional microscopy in the diagnosis of malaria infection (43). However, a definite advantage is the ability to confirm a cure, since the test detects only viable organisms. Also, the number of false-positive results due to rheumatoid factor is much smaller than for some of the other tests. In a comparison of results with the ICT Malaria Pf and ParaSight F tests, low monoclonal antibodies correctly identified P. falciparum malaria in patient blood samples more often than the other two procedures did. In this study, the Flow monoclonal antibodies exhibited 94% sensitivity and 100% specificity for P. vivax and 88% sensitivity and 99% specificity for P. falciparum (41). In another study, compared with PCR, the sensitivity was 93% and the specificity was 99.5%. The authors felt that the test had sufficient sensitivity and specificity to detect P. vivax under laboratory conditions and could also be useful for malaria diagnosis in the field in Mexico (44). As with the other rapid malaria tests, cost is always an issue; however, Flow monoclonal antibodies provide a simple, rapid, and effective test in the diagnosis of malaria, especially where well-trained microscopists are not available or the work load is too high (45).
Figure 7.14 Flow anti-pLDH Plasmodium monoclonal antibodies rapid malaria test. (Left) Diagram of blood flow on the membrane through the timed test; note the positive control line and positive test line. (Right) Diagram of negative control, a positive P. vivax result, and a positive P. falciparum result. Note that the line for P. vivax is a panspecific antibody against all four species; however, the panspecific antibody has been shown to detect only P. vivax with any degree of consistency. (Photographs adapted from Flow’s website, with permission.) doi:10.1128/9781555819002.ch7.f14
Although the first PCR methods were developed to identify P. falciparum, many can now detect several or all five species. It has been well demonstrated that PCR can detect lower parasitemias than any of the traditional blood film or nontraditional dipstick methods, and it may be preferable to microscopy as the reference standard when evaluating new diagnostic tests (22, 24, 42, 46–48). One advantage of PCR is the ability to confirm false-negative results by microscopy as true positives. Another advantage is the enhanced ability to identify organisms to the species level, particularly in mixed infections, compared with microscopy. However, even PCR may miss some cases as a result of PCR inhibitors, DNA degradation, or genotypic variants. Unfortunately, the time required for PCR presents a problem for the clinical laboratory; this approach does not lend itself to routine use. With the development of automation for PCR, these tests may become much more user-friendly for the routine laboratory; however, currently PCR is reserved for reference centers and special circumstances. As PCR becomes more widely used, it will be important to remember that the mode of collection and storage of blood samples may influence the sensitivity of Plasmodium detection. This may be critical in studies of individuals with low parasitemia or mixed infections and in comparison of data from different settings, including field settings.
Other developments include the post-PCR/ligase detection reaction-fluorescent microsphere assay (LDR-FMA) (49). This assay, which uses Luminex FlexMAP microspheres, provides simultaneous, semiquantitative detection of infection by four human malaria parasite species at a sensitivity and specificity equal to those of other PCR-based assays. In blinded studies using P. falciparum-infected blood from in vitro cultures, the authors identified infected and uninfected samples with 100% concordance. Also, in analyses of P. falciparum in vitro cultures and P. vivax-infected monkeys, comparisons between parasitemia and LDR-FMA signal intensity showed very strong positive correlations (r > 0.95). Application of this multiplex Plasmodium LDR-FMA diagnostic assay will increase the speed, accuracy, and reliability of diagnosing human Plasmodium infections.
Certainly molecular testing offers many advantages over microscopy, including identification to species levels; this capability is becoming much more important with the shortage of experienced microscopists. A recent multiplex quantitative real-time PCR can rapidly and simultaneously identify all five Plasmodium species known to cause malaria in humans (P. vivax, P. ovale, P. malariae, P. falciparum, P. knowlesi) (50). In another recent study, malaria loop-mediated isothermal amplification (LAMP) used in a remote Ugandan clinic achieved sensitivity similar to that achieved by a single-well nested PCR in a United Kingdom reference laboratory (51).
Automated Blood Cell Analyzers
Although the use of automated blood cell analyzers is not yet clinically relevant for the diagnosis of blood parasites, improvements in the systems may offer future information that will supplement the routine microscopy procedures currently in use (52, 53). Unfortunately, when the parasitemia is light, automation tends to have some of the same problems seen with other alternative procedures. In many cases, the changes seen using analyzers are not specific to malaria infection and could occur in many other diseases (increases in the number of large, unstained cells and thrombocytopenia). The accuracy for malaria diagnosis appears to vary according to the Plasmodium species, parasitemia, immunity, and clinical context.
The Cell-Dyn 4000 automated hematology analyzer (Abbott, Chicago, IL) has the ability to detect 91.2% of malaria patients. In one study on day 3 of follow-up posttreatment, the sensitivity was 96.7% that of microscopy. The atypical polarizing events, which indicate the presence of malarial parasites in the analyzer, were highly correlated with the levels of parasitemia in serially diluted samples of the leukocyte-depleted blood; parasites were detected down to the level of 288 ± 17.7/µl). These data suggest that the atypical light depolarization could be influenced by parasitemia and could be used as a screening method for P. vivax malaria patients, as well as for therapeutic monitoring (52). Microscopy is still required for species determination and parasite quantitation.
The potential of automated depolarization analysis in detecting malaria infection as part of the routine full blood count performed by the Cell-Dyn 4000 analyzer has been described previously (52). In these cases, abnormal depolarizing patterns are due to the presence of leukocyte-associated malaria hemozoin, a pigment which depolarizes the laser light. Abnormal polarizing events have also been described for samples from three individual patients infected by the nematode Mansonella perstans. The observed depolarizing pattern consisted of a normal depolarizing eosinophil population plus an abnormal depolarizing population that showed a close “linear” relationship between “granularity” (90° depolarization) and “lobularity” (90° polarization). This atypical population was smaller than that of normal leukocytes and thus clearly different from the patterns associated with malaria infection. Abnormal depolarization patterns of M. perstans clearly do not reflect leukocyte-associated malaria hemozoin. It is possible, however, that the erythrocyte-lysing agent used to facilitate leukocyte analysis by the instrument may have caused microfilaria fragmentation and thus the distinctive straight-line features of the abnormal scatter plots (54).
Diagnosis of Leishmaniasis: Review of Alternatives to Conventional Microscopy
ICT for Detection of Anti-rK-39 Antibodies
In one study, the diagnostic utility of an immunochromatographic test for detection of anti-rK-39 antibodies for the diagnosis of kala azar and post-kala azar dermal leishmaniasis (PKDL) was evaluated (55). Of the 120 samples tested, 57 were positive by ICT; 51 of these were diagnosed as kala azar, and 6 were diagnosed as PKDL. The controls included individuals from areas of endemic (21) and nonendemic infection with malignancies, hemolytic disorders, chronic liver disease, hypersplenism, portal hypertension, metabolic disorders, or sarcoidosis. In addition, 47 sera from patients with confirmed cases of tuberculosis, malaria, typhoid, filariasis, leptospirosis, histoplasmosis, toxoplasmosis, invasive aspergillosis, amebic liver abscess, AIDS, leprosy, cryptococcosis, strongyloidiasis, and cyclosporosis, as well as from patients with collagen vascular diseases and patients with hypergammaglobulinemia, were tested to check the specificity of the test. Of the 51 patients with kala azar, all had fever lasting from <1 month to 1.5 years (median, 4.5 months). All six PKDL patients gave a history of having kala azar in the past, and their slit skin test smears were microscopically positive for Leishman-Donovan bodies. The strip test was positive in all the cases of kala azar and PKDL (estimated sensitivity, 100%), and all control sera were negative by the ICT (specificity, 100%). The rK-39 ICT is a highly sensitive and specific test and may be suitable for a rapid, cost-effective, and reliable field diagnosis of kala azar and PKDL (55).
This test has also been adapted to the use of urine (56) and sputum (57) for the detection of visceral leishmaniasis antibody. Both screening approaches appear to be very promising, with distinct advantages over the invasive use of blood specimens.
Cytocentrifugation (cytospin), which uses an apparatus for concentrating cells in suspension on a microscope slide, is commonly used in most histopathology laboratories. The use of a hemolyzing and isotonic saponin solution to lyse RBCs and platelets, which contains formalin as a fixative, has led to an improved technique for the detection of Plasmodium spp., L. donovani, microfilariae, and WBCs containing malaria pigment. The concentration of the parasites present in the sediment from 100 µl of blood spread on a 6-mm-diameter circle results in good morphology that is well stained using Giemsa, Wright’s, or rapid stains. This new method costs very little to perform and offers the possibility of isolating and identifying the main blood-stage parasites in the same sediment. The possible exception would be young trophozoites of P. falciparum, which do not concentrate well due to their small size (58).
The Knott concentration procedure is used primarily to detect the presence of microfilariae in the blood, especially when a light infection is suspected (4, 59) (Fig. 7.15). The disadvantage of the procedure is that the microfilariae are killed by the formalin and are therefore not seen as motile organisms.
Figure 7.15 Microfilariae at low power from a Knott concentration, unstained. Note that the sheath is not visible. doi:10.1128/9781555819002.ch7.f15
Figure 7.16 African trypanosomes obtained using the triple centrifugation method. doi:10.1128/9781555819002.ch7.f16
1. Place 1 ml of whole or citrated blood into a centrifuge tube containing 10 ml of 2% formalin. Thoroughly mix the tube contents.
2. Centrifuge for 2 min at 500 × g or 5 min at 300 × g.
3. Decant the supernatant fluid, examine some sediment as a wet mount at low (×100) and high dry (×400) power, prepare thick films from the remaining sediment, and allow the films to air dry.
4. Stain films with Giemsa, Wright’s, or rapid stains.
Note Use alcohol-cleaned slides for preparation of the films made from the sediment.
The membrane filtration technique as modified by Desowitz and others has proved highly efficient in demonstrating filarial infections when microfilaremias are of low density. It has also been successfully used in field surveys (3, 4, 60, 61). (See also Fig. 5.9, which shows a membrane filtration system that can also be used for blood filtration; see below.)
1. Draw 1 ml of fresh whole blood or anticoagulated blood into a 15-ml syringe containing 10 ml of distilled water.
2. Gently shake the mixture for 2 to 3 min to ensure that all blood cells are lysed.
3. Place a 25-mm Nuclepore filter (5-µm porosity) over a moist 25-mm filter paper pad. (This method is unsatisfactory for the isolation of M. perstans microfilariae because of their small size. A 3-µm-pore-size filter could be used for recovery of this organism. Other filters of similar pore size are not as satisfactory as the Nuclepore filter.) Place the filter in a Swinney filter adapter.
4. Attach the Swinney filter adapter to the syringe containing the lysed blood.
5. With gentle but steady pressure on the piston, push the lysed blood through the filter.
6. Without disturbing the filter, remove the Swinney adapter from the syringe and draw approximately 10 ml of distilled water into the syringe. Replace the adapter, and gently push the water through the filter to wash the debris from the filter.
7. Remove the adapter again, draw the piston of the syringe to about half the length of the barrel, replace the adapter, and push the air in the barrel through the filter to expel excess water.
8. To prepare the filter for staining, remove the adapter, draw the piston about half the length of the barrel, and then draw 3 ml of absolute methanol into the syringe. Holding the syringe vertically, replace the adapter and push the methanol followed by the air through the filter to fix the microfilariae and expel the excess methanol.
9. To stain, remove the filter from the adapter, place it on a slide, and allow it to air dry thoroughly. Stain with Giemsa stain as for a thick film (with 0.1% Triton X-100) or with Delafield’s hematoxylin.
10. To cover the stained filter, dip the slide in toluene before mounting the filter with neutral mounting medium and a coverslip. This will lessen the formation of bubbles in or under the filter.
Gradient Centrifugation Technique
The gradient centrifugation technique is another technique for the concentration of microfilariae (62).
1. Mix 30 ml of 50% Hypaque with 14 ml of distilled water; add 1 part of this mixture to 2.4 parts of 9% Ficoll.
2. Place 4 ml of the Ficoll-Hypaque mixture in a 15-ml plastic centrifuge tube; overlay this mixture with 4 ml of heparinized venous blood.
3. Centrifuge the tube at 400 × g for 40 min.
4. Microfilariae will be found in the middle Ficoll-Hypaque layer, which separates the overlying plasma and WBC layers from the underlying RBCs.
Triple-Centrifugation Method for Trypanosomes
The triple-centrifugation procedure may be valuable in demonstrating the presence of trypanosomes in the peripheral blood when the parasitemia is light (4, 9) (Fig. 7.16).
1. Centrifuge anticoagulated blood for 10 min at 300 × g.
2. Remove the supernatant fluid, and transfer it to another centrifuge tube.
3. Centrifuge this fluid for 10 min at 500 × g.
4. Again, remove the supernatant fluid to another centrifuge tube.
5. Centrifuge this fluid for 10 min at 900 × g.
6. Decant the supernatant fluid, and examine the sediment as a wet preparation.
7. The sediment may be used to prepare thin films that can then be stained with any of the blood stains.
Note Remember, you are saving the supernatant fluid in step 2 for subsequent centrifugation; do not accidentally pour this fluid off for disposal. The final sediment after the third centrifugation step is used to prepare stained films for examination.
Special Stain for Microfilarial Sheath
Some of the material that is obtained from the concentration procedures can be allowed to dry as thick and thin films and then stained with Delafield’s hematoxylin, which demonstrates greater nuclear detail as well as the microfilarial sheath, if present. In addition, fresh thick films of blood containing microfilariae can be stained by this hematoxylin technique (Fig. 7.17 and 7.18) (4, 9).
Figure 7.17 Wuchereria bancrofti microfilaria on a thick film stained with Delafield’s hematoxylin. Note the presence of the sheath. (Image courtesy of the CDC Public Health Image Library.) doi:10.1128/9781555819002.ch7.f17
Figure 7.18 Wuchereria bancrofti microfilaria. (Upper) Microfilaria stained with Giemsa stain. Note that the sheath is not visible. (Lower) Microfilaria stained with Delafield’s hematoxylin stain. Note that the sheath is visible. doi:10.1128/9781555819002.ch7.f18
Preparation of Stain
Dissolve 180 g of aluminum ammonium sulfate in 1 liter of distilled water (saturated solution). Heat until dissolved. The cooled supernatant fluid is saturated aluminum alum.
Hematoxylin Solution
Dissolve the hematoxylin in alcohol, and add it to the alum solution. Expose the solution to sunlight and air for ripening in a clear, cotton-plugged bottle for approximately 1 week; then filter and add the following:
Age for 1 month or longer in sunlight, and then run test smears to determine whether the solution is properly aged. Nuclei should stain blue, and cytoplasm should stain different shades of red.
Procedure
1. Prepare thick films from the concentration material or from fresh blood.
2. Allow the films to air dry.
3. Lake the films in 0.85% sodium chloride or distilled water for 15 min (not necessary for films prepared from the Knott sediment). Allow to air dry.
4. Fix all films in absolute methanol for 5 min. Allow to air dry.
5. Stain in undiluted Delafield’s hematoxylin for 10 to 15 min.
6. Wash off excess stain in tap water.
7. Intensify the blue color by placing the films into tap water containing several drops of ammonia (NH4OH) for several minutes.
8. Rinse again in running tap water for 5 min.
9. Air dry.
10. Mount in Permount or some other mounting medium; use a no. 1 coverglass.
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