Methods for the detection of blood parasites

The diagnosis and identification of blood parasites is becoming ever more important with the increase in imported parasitic disease in the temperate zone. Any clinical haematology laboratory can expect to be called upon to diagnose these parasites, especially malaria, and a high level of competence is required. This review considers the imported blood parasites that could be encountered and discusses the merits of the various diagnostic tests available. The accepted laboratory procedure for the diagnosis of malaria is the preparation and examination of Giemsa or Field's stained blood smears under the light microscope ( Warhurst & Williams 1996; General Haematology Workforce 1997). Smears may be prepared directly from a finger prick blood sample. Venipuncture specimens collected in EDTA coated tubes are also acceptable, but must be examined promptly to prevent alteration of parasite morphology due to the sequestrine effect. Both thick and thin smears should be made. The thick smear, which increases by a factor of 20–30 the number of red blood cells per given area on the slide compared to the thin film, is much more sensitive than the thin film for detection of a malarial infection. The thin smear is superior to the thick for speciation. This technique has undergone very little improvement over the last 100 years. Although considered the gold standard, diagnosis of malaria using blood films can be problematic; it requires up to 60 min of preparation time, is labour intensive and interpretation of the result requires considerable expertise, particularly at low levels of parasitaemia Warhurst & Williams (1996). In addition, in patients with Plasmodium falciparum malaria, the parasites are sequestered in tissue capillaries for part of the asexual cycle and, in a synchronous infection with this species, may not always be present in the peripheral blood. Thus a P. falciparum infection might be missed unless blood samples are repeated daily, or more frequently if the clinical picture dictates. Furthermore, detection of parasites by blood film on follow-up of treatment does not assess their viability ( Srinavasan et al. 2000) . Warhurst & Williams (1996) discuss several methods employed to quantify malaria parasites in thick blood smears. One technique is to count the total number of parasites per 200 white blood cells (WBC) and multiply this number by 40 to give the number of parasites/µl, assuming that there are 8000 WBC/µl blood. A second method involves making a thick smear with a known volume of blood (5 µl), and staining with Giemsa stain before counting all the parasites on the smear. The total parasite count is divided by 5 to obtain the number of parasites/µl. A similar technique again uses a defined amount of whole blood to make a thick smear and then the parasites present in 100 high-power fields (HPF) are counted. One parasite/100 HPF is assumed to be the equivalent to 50 parasites/µl. A thick smear is generally considered negative if no parasites are seen after 10 min searching. Parasite levels may also be quantified by examination of a thin blood smear. This is usually accomplished by counting the number of parasites/1000 red blood cells (RBC) and expressing as a percentage the number of erythrocytes infected, or again, by counting the number of parasites per 200 WBC as described for the thick film. A weakness in the estimation of parasite levels by counting parasites against a particular number of WBC is the assumption that all blood samples contain 8000 WBC/µl. If a suitable instrument is available, it is much better to count the WBC/µl of each blood sample under investigation, and so obtain an accurate conversion factor for each sample. For example, if there are 10 parasites/200 WBC and the total WBC is 10 000/µl then there are 500 parasites [i.e. (10 000/200) × 10] parasites/µl. The major disadvantage of the thick smear is that it is difficult to read and the necessary expertise may be unavailable. Using one method to evaluate the intensity of the infection, the examiner should, in theory, only count thick-smear fields containing 20 WBC/HPF. In reality, fields containing 20 WBC are often too thick to count accurately, whereas the fields that are easiest to read are those that contain only five or six WBC each. It has been suggested that the microscopist should be able to read print through a well-prepared, thick blood smear before it is stained. Warhurst & Williams (1996) report that examination of thin blood smears is only one tenth as sensitive as examination of thick blood smears for the quantification of malarial parasites, although identification of the Plasmodium species present is much easier using thin smears. Most laboratories involved in the identification and quantification of malaria parasites by microscopy therefore produce both thick and thin blood smears. In conclusion, even though examination of blood smears is accepted as the current, universal 'gold standard' for malaria diagnosis, no single method for estimation of parasitaemia is used by all investigators, which can lead to difficulties in comparing data between different studies ( Hanscheid & Valadas 1999). Three techniques using fluorescence of the parasite for the diagnosis of malaria have been described. The quantitative buffy coat or QBC11® method ( Baird et al. 1992 ; Benito et al. 1994 ; Clendeman, Long & Baird 1995) which is available as a commercial kit (Becton Dickinson, Franklin Lakes, NJ, USA); the Kawamoto acridine orange process ( Kawamoto 1991a; Kawamoto 1991b; Kong & Chung 1995; Bosch et al. 1996 ; Gay et al. 1996 ; Lowe et al. 1996 ) and the use of benzothiocarboxypurine (BCP) ( Makler et al. 1991 ; Cooke et al. 1992 ). These three techniques are rapid and relatively easy to perform (when there are > 100 parasites/µl) and demonstrate sensitivity and specificity equivalent to those achievable by examination of stained thick smears. Both the QBC and Kawamoto methods use acridine orange (AO) as the fluorochrome to stain the nucleic acids of any malaria parasites in the sample. BCP is another fluorochrome which stains nucleic acids. Although AO is a very intense fluorescent stain, it is nonspecific and stains nucleic acids from all cell types. Consequently, the microscopist using AO has to learn to distinguish fluorescent-stained parasites from other cells and cellular debris containing nucleic acids. The sensitivity of AO staining at parasite levels of < 100 parasites/µl has been reported to range from 41.1% to 93% ( Delacollett & VanderStuyft 1994). The specificity of AO staining for P. vivax infections was reported to be about 52%, whereas that for P. falciparum infections was around 93% ( Hakim et al. 1993 ). The BCP method has a reported sensitivity and specificity of > 90% ( Cooke et al. 1993 ). An important limitation of methods based on AO and BCP is their inability to differentiate easily between Plasmodium spp. In addition, AO is considered hazardous and has special disposal requirements. Comparing methodologies, both the QBC and the BCP fluorescent methods are more demanding technically than the Kawamoto AO method ( Hakim et al. 1993 ; Hind et al. 1994 ) and require special equipment and supplies. The QBC method requires a particular centrifuge and haematocrit tubes and was reported to increase costs to about US$ 1.70/sample ( Craig & Sharp 1997). The BCP method also requires a good fluorescent microscope, with a high intensity mercury vapour or quartz halogen lamp. The simpler Kawamoto method employs a pair of fluorescence filters and readily available AO and can make use of the sunlight as the exciting light source (if a shield to surround the observer's eyes is used). If sunlight is not used this technique also requires a high-intensity halogen lamp. In spite of its limitations, including the requirements for special training, expensive equipment and supplies, fluorescence microscopy for the rapid detection of malaria parasites in blood is a viable alternative to examination of Giemsa stained smears. Another approach to the laboratory diagnosis of malaria is based on the detection of nucleic acid sequences specific to Plasmodium. Several PCR assays have been developed for the diagnosis of malaria. The 18S rRNA gene has been used as a target for the differentiation of Plasmodium species by nested PCR and reverse transcription-PCR ( Barker et al. 1992 ; Snounou et al. 1993 ). Other DNA targets such as the circumsporozoite protein gene have also been investigated for species specific regions. The large-subunit rRNA gene is extensively conserved within Plasmodium species and is suitable as a genus-specific target region; the amplified target sequence is detected by internal probes or analysed by gel electrophoresis ( Kain et al. 1993 ). The major advantage of using a PCR-based technique is the ability to detect infection in patients with low parasitaemia; infection with five parasites/µl can be detected with 100% specificity ( Kawamoto et al. 1996 ). PCR-based techniques have been used to screen blood donors in Ho Chi Minh City, Vietnam, an area where malaria is endemic. The sensitivity of PCR-based methods is greater than microscopy and, following treatment, PCR yields positive results longer than microscopic examination ( Kain et al. 1994 ). Several workers have investigated the persistence of parasite DNA in peripheral blood following antimalarial treatment. In one report, PCR remained positive for a median of 144 h compared to 66 h for microscopy ( Sethabutr et al. 1992 ; Seesod et al. 1993 ). Such microscopically negative but PCR positive specimens probably had either subpatent parasitaemia or circulating P. falciparum DNA. Kain et al. (1994) also reported that if PCR yields positive results for 5–8 days after treatment, therapeutic failure, possibly due to drug resistance, might be predicted. The variable PCR results in different studies ( Table 1) reflect a number of variations in PCR technique from collection and storage of the specimens, DNA extraction, selection of primers, amplification conditions, and analysis of amplified product. Further advances in PCR technology may soon allow viable parasites to be distinguished from nonviable and facilitate use of PCR-based procedures in the field ( Wataya et al. 1993 ). Although PCR-based methods are arguably the most sensitive method for detection of patent malarial infections, they are also particularly useful for studies on strain differences, mutations, and genes involved in drug resistance in the parasites and to show the relationship between strains associated with different outbreaks of malaria. Use of PCR-based technology for malaria diagnosis may therefore become clinically relevant for acute diagnosis in both field and reference laboratory situations. Although alternative methods to stained blood films for diagnosis of malaria have been around for a considerable time ( Taylor & Voller 1993), the original fluorescent antibody detection tests for malaria were limited in sensitivity and in their ability to distinguish active from prior infections. The new generation antigen capture tests are capable of detecting fewer parasites and of producing a more rapid result (10–15 min). They are commercially available as kits, which include all the necessary reagents, and do not require extensive training or equipment to perform or to interpret their results. Two parasite antigens are currently used in the new, rapid immunochromatographic tests: Histidine-rich protein-2 (HRP-2) ( Rock et al. 1987 ), a water-soluble protein which is expressed by P. falciparum and parasite lactate-dehydrogenase (pLDH) antigen ( Makler et al. 1998 ), present as separate isomers for all four Plasmodium species infecting humans. Both of these antigens are also produced by gametocytes ( Odoula et al. 1987 ; Kilian et al. 1997 ). The latest antigen capture tests are rapid and simple to perform, and have detection limits comparable to those of high-quality microscopy (i.e. 100–200 parasites/µl). At parasitaemias of 60–100 parasites/µl, the HRP-2 based tests are > 90% sensitive and > 90% specific for P. falciparum, compared with thick-smear microscopy ( Dietz et al. 1995 ). Currently, two such tests are commercially available: the ParaSight® F-test (Becton Dickinson) and the ICT Malaria Pf Test® (Amrad ICT Diagnostics, Sydney, Australia). Both tests can be performed on finger-prick blood samples. They deploy labelled monoclonal antibodies to HRP-2 in a mobile phase, to capture HRP-2 antigen from P. falciparum and a further stripe of anti HRP-2 antibody immobilized onto nitro-cellulose strips, to capture labelled antigens in a dipstick format. In each test, a positive result is indicated by the appearance of a red line on the test strip at the site where the monoclonal antibodies are immobilized. Both tests also contain a built-in control line, which must appear for the test to be considered valid. A comparison of the ParaSight® F antigen-capture test with PCR-based diagnosis ( Peyron et al. 1994 ) gave a reported sensitivity and specificity of the ParaSight® F-test of 80% and 97%, respectively, compared with the results of PCR. Other studies on the sensitivity and specificity of the ParaSight® F-test for the detection of Plasmodium falciparum in blood samples. ( Makler & Hinrichs 1993; Schiff et al. 1993 ; Beadle et al. 1994 ; Miller et al. 1994 ; Peyron et al. 1994 ; Premji et al. 1994 ; Dietz et al. 1995 ; Verle et al. 1996 ; Kodisinghe et al. 1997 ; Singh et al. 1997 ) have shown an average sensitivity of 77–98% and specificity of 83–99%. Although HRP-2-based immunochromatographic tests permit rapid diagnosis of P. falciparum malaria, they do have limitations in their use. Firstly, since HRP-2 is only expressed by P. falciparum, tests based on the detection of HRP-2 give negative results for samples containing P. vivax, P. ovale or P. malariae. Thus, cases of nonfalciparum malaria may be misdiagnosed as malaria-negative if the only diagnostic test used is based upon HRP-2 detection. Another limitation is the persistence in the blood of HRP-2 after the clinical symptoms of malaria have disappeared and the parasites have apparently been cleared from the host. Results of investigations with the ParaSight® F-test on blood taken 0–14 days after treatment of falciparum malaria with artemether showed persistence of HRP-2 in 16 of 75 blood samples at day 14 after treatment had finished. Tests for HRP-2 may therefore be difficult to interpret when screening indigenous people of malaria-endemic areas, who may have persistent, low-level parasitaemia, yet not be ill ( Verle et al. 1996 ). The reason for the persistence of the HRP-2 antigens is not well understood; it may reflect the presence of latent, viable parasites (possibly the result of treatment failure) or of soluble, antigen/antibody complexes ( Humar et al. 1997 ). The persistence may also depend upon the type of antimalarial therapy instituted. Schiff et al. (1993) noted that 10% of patients treated with Fansidar® had detectable HRP-2 antigen on day 14. Using the ParaSight® F-test to investigate 19 patients given unspecified antimalarial chemotherapy, Humar et al. (1997) detected HRP-2 antigen in 68% of the patients on day 7 after initiation of treatment and in 27% on day 28. The HRP-2 signal has been shown to persist for 19 days after apparently effective quinine therapy. Karbwang et al. (1996) also detected persistent HRP-2 antigen during and after artemether therapy, acknowledging that the HRP-2 signal was of no value during the first week of treatment but appeared to be a precise indicator of treatment failure under 'field' conditions, when it was detected on day 14 post-treatment. More studies are required in this area. Other limitations are specifically related to technical aspects of the test system. For example, the monoclonal IgG antibody utilized in the ParaSight® F may cross-react with serum rheumatoid factor and be the cause of a false-positive response ( Laferi et al. 1997 ). Parasite lactate dehydrogenase (pLDH) ( Makler et al. 1993 ; Piper et al. 1996 ) is a glycolytic enzyme present in all species of malaria parasites of humans. Furthermore, pLDH from P. falciparum can be assayed in the presence of host LDH utilizing the coenzyme 3-acetylpyridine adenine dinucleotide (APAD), an analogue of nicotinamide adenine dinucleotide (NAD) ( Piper et al. 1999 ). Although the Michaelis–Menten constants (km) are similar, the keat (turnover number) of pLDH in the presence of APAD is much greater than that of the human enzyme with the same coenzyme ( Gomez et al. 1997 ). This characteristic is due in part to conformational changes that occur in the enzyme ( Dunn et al. 1996 ). pLDH-based assays are available in two formats: a semiquantitative, dry dipstick (OptiMAL®, Flow Inc., Portland, OR, USA) and a 96 well, quantitative, immunoenzymatic capture, plate assay ( Piper et al. 1999 ). The latter technique captures pLDH on a microtitre plate with a specific anti pLDH monoclonal antibody, then quantifies the amount of enzyme captured with an enzyme reaction utilizing APAD. The bound pLDH activity correlates well with viable parasite density on initial diagnosis. But the pLDH activity has also been shown to parallel the level of viable parasites during therapy ( Makler & Heinrichs 1993; Gomez et al. 1997 ; Piper et al. 1999 ). The latter observation is especially important, since this assay could thus be used to follow patient progress during antimalarial therapy, and alert the physician to the presence of a drug-resistant infection if pLDH levels fail to decline in sequential samples. Low levels of pLDH cannot easily be measured colourimetrically by this enzymatic method because of the presence of nonspecific reductases in blood lysates. Because of this, two assays have been formatted that utilize monoclonal antibodies (mAbs) which capture the pLDH. These mAbs were raised against pLDH purified from P. falciparum infected erythrocytes. They vary in respect of their reactivity with pLDH isoforms from the different human malarial species. Two of the mAbs are pan-specific, recognizing all four species of malaria. A third mAb recognizes only P. falciparum LDH isoforms. The monoclonal antibodies utilized in the OptiMAL® test have been exhaustively tested for cross-reactivity with LDH from Leishmania, Babesia and pathogenic bacteria or fungi and no evidence of such cross-reactivity has been found ( Makler & Heinrichs 1993). The immunochromatographic strip test (OptiMAL®) which is based on these mAbs uses a gold-conjugated pan-specific antibody (6C9) to capture pLDH from all species of Plasmodium from humans. The blood/antibody complex travels along a nitro-cellulose strip on which are two further monoclonal antibody capture stripes, one specific for P. falciparum (17E4) and a second pan-specific monoclonal antibody (19G7) for capturing all four Plasmodium species. A goat antimouse monoclonal antibody capture control line is present to indicate a successfully performed test. The gold conjugated antigen/antibody complex builds as a red line at the capture stripe. The presence of three lines (control + pan-specific + Pf specific) indicates P. falciparum antigen captured. Two lines (control + pan-specific but not Pf specific) indicates a species other than P. falciparum. Trials of the OptiMAL® test have been undertaken in many centres ( John et al. 1998 ; Palmer et al. 1998 ; Quitana et al. 1998 ; Hunt Cooke et al. 1999 ; Moody et al. 2000 ) and have reported sensitivities of 92–98%, with a specificity of 94%, for P. falciparum and a sensitivity of 96% for P. vivax. Studies to evaluate pLDH levels in sequential blood samples taken from patients undergoing antimalarial chemotherapy demonstrated that pLDH levels followed peripheral parasitaemia and fell to undetectable values 3–5 days after quinine therapy, which could provide a valuable tool in monitoring antimalarial therapy ( Srinavasan et al. 2000 ). Clearance of parasitaemia and clearance of pLDH appear to parallel each other. In the past few years, efforts to replace the and of blood smears have to techniques for the detection of malaria parasites with a sensitivity equivalent to or better than microscopy ( Taylor & Voller 1993). the of a more method for malaria diagnosis, on training and is by parasites of the more than 100 species have been only a few have been shown to cause human infection. Babesia and Babesia have been in most human cases ( but different have been seen in malaria-endemic where Babesia can easily be misdiagnosed as Plasmodium. 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