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Laboratory tools for diagnosis of malaria

Laboratory tools for diagnosis of malaria
Literature review current through: Jan 2024.
This topic last updated: Oct 12, 2023.

INTRODUCTION — Prompt and accurate diagnosis of malaria is critical for implementation of appropriate treatment to reduce associated morbidity and mortality. Accurate detection of malaria is also important for epidemiologic screening and surveillance to inform malaria control strategies, for research purposes in testing efficacy of antimalarial drugs and vaccines, and for blood bank screening.

Characteristics of a useful malaria diagnostic tool include the ability to definitively establish presence or absence of infection, determine which species of malaria is/are present, quantify parasitemia (ie, parasites per microliter of blood or percent red blood cells infected), detect low-level parasitemia, and allow monitoring of response to antimalarial therapy (including detection of recrudescence or relapse). Thus far, there is no single malaria diagnostic tool that meets all of these criteria. Test characteristics that are important for diagnosis vary depending on the epidemiology of infection and goals for control in the region where the test is used.

Tools for diagnosis of malaria will be reviewed here. Issues related to the clinical approach to diagnosis of malaria, as well as issues related to the epidemiology, clinical manifestations, treatment, and prevention of malaria, are discussed separately:

(See "Malaria: Epidemiology, prevention, and control".)

(See "Malaria: Clinical manifestations and diagnosis in nonpregnant adults and children".)

(See "Treatment of uncomplicated falciparum malaria in nonpregnant adults and children".)

(See "Treatment of severe malaria".)

(See "Malaria in pregnancy: Prevention and treatment".)

(See "Non-falciparum malaria: P. vivax, P. ovale, and P. malariae".)

(See "Non-falciparum malaria: Plasmodium knowlesi".)

GENERAL PRINCIPLES — Tools for diagnosis of malaria include microscopy (visualization of parasites in stained blood smears) and rapid diagnostic tests (RDTs; which detect antigen or antibody). Smear examination via light microscopy is the standard tool for diagnosis of malaria; RDTs should be used if microscopy is not readily available. Molecular techniques for detection of genetic material are limited to research settings [1].

LIGHT MICROSCOPY — Detection of parasites on Giemsa-stained blood smears by light microscopy is the standard tool for diagnosis of malaria [2]. In expert hands, the sensitivity of microscopy can be excellent, with detection of malaria parasites at densities as low as 4 to 20 parasites/mcL of blood (approximately 0.0001 to 0.0005 percent parasitemia) [3-6]. Diagnostic errors occur more commonly in the setting of low-density parasitemia (10 to 100 parasites/mcL of blood), although errors can also occur with higher densities [7-9].

Microscopy allows identification of the Plasmodium species as well as quantification of parasitemia. It also enables diagnosis of hematologic abnormalities and other infectious diseases such as filariasis, trypanosomiasis, babesiosis, and others.

Drawbacks to microscopy include that it is labor intensive and requires substantial training and expertise [6,10,11]. The sensitivity and specificity of malaria microscopy in resource-limited settings is often below levels achievable in reference or research laboratories [8,12-15]. In well-equipped laboratories outside of endemic areas, variation in techniques used for preparation and interpretation can influence results [7,16-18], and errors may occur in laboratories with limited exposure to tropical infections [2,19].

Microscopy cannot reliably detect very low parasitemia (<5 to 10 parasites/mcL) or cases where the majority of the parasite biomass is sequestered (eg, in the case of placental sequestration during pregnancy) [10]. If malaria is suspected and the initial smear is negative, additional smears should be prepared and examined over the subsequent 48 to 72 hours. (See 'Blood smear interpretation' below.)

In general, there is no role for routine use of pre-emptive treatment if quality-assured malaria diagnostic testing is negative. Presumptive treatment may be appropriate for circumstances in which prompt laboratory diagnosis is not available and clinical suspicion for severe disease is high.

The sensitivity of light microscopy for detecting placental infection is low [20-22]; in one study including more than 590 pregnant women in Ghana, the sensitivity of light microscopy, rapid antigen test, and polymerase chain reaction were 42, 80, and 97 percent, respectively [20].

Blood smear preparation — Preparation of smears consists of applying a drop of blood to a glass microscopy slide, followed by drying and staining procedures. Capillary blood from a fingerprick (or ear lobe or infant heelstick) or anticoagulated venous blood may be used. Smears should be prepared as soon as possible after blood collection to avoid alteration in parasite morphology and staining properties.

Two types of blood smears are used in malaria microscopy: thin and thick smears. Thin smear preparation maintains the integrity and morphology of erythrocytes so that parasites are visible within red blood cells. Thin smears allow identification of the infecting parasite species and can be used to measure parasite density. Thick smear preparation involves mechanical lysis of red blood cells so that malaria parasites can be visualized independent of cell structures. Thick smears allow the microscopist to review a relatively large quantity of blood and are typically used to screen for presence or absence of parasites and to estimate parasite density.

The United States Centers for Disease Control and Prevention (CDC) has online guides for preparation and staining of blood smears.

Blood smear interpretation — Malaria parasites are best seen under 100x magnification using the oil immersion objective lens; smear evaluation should include examination of at least 200 to 500 fields or examination for 20 to 30 minutes.

If malaria is suspected and the initial smear is negative, additional smears should be prepared and examined over the subsequent 48 to 72 hours [23]; the CDC recommends repeating a thick and thin smear every 12 to 24 hours for a total of three sets before ruling out malaria [24,25].

Species identification — Thin smears allow identification of the malaria species. The malaria species morphology is variable depending on the stage of infection (figure 1 and figure 2 and table 1 and table 2).

The CDC has bench aids for identification of P. falciparum, P. vivax, P. ovale, and P. malariae.

Plasmodium knowlesi, which may cause severe disease, may be indistinguishable by microscopy from P. malariae (which generally causes milder illness). (See "Non-falciparum malaria: Plasmodium knowlesi".)

Administration of antimalarial therapy can alter the morphologic appearance of parasites and affect their identification in blood smears [6,26,27].

Quantification of parasitemia — Parasitemia density may be quantified using a thin blood smear or a thick blood smear:

For thin blood smears, the parasite density can be estimated by examining a monolayer of red blood cells (RBCs) using the oil immersion objective at 100x. The slide should be examined where the RBCs are more or less touching (approximately 400 RBCs per field). The parasite density can then be estimated from the percentage of infected RBCs; at least 500 RBCs should be counted [28].

For thick blood smears, a standard approach to estimating parasite density involves counting asexual parasite forms and white blood cells in each microscopy field until 200 white blood cells have been counted. If fewer than 10 asexual parasites are counted per 200 white blood cells, counting should continue to a total of 500 white blood cells. Subsequently, the measured white blood cell count (in cells per microL) is divided by the number of white blood cells counted (200 or 500), and the result is multiplied by the number of parasites counted, giving the parasite density (parasites per microL). Variation in white blood cell counts may confound estimates that use an average rather than a measured white blood count [29].

For example, a white blood cell count of 8000/microL divided by 200 white blood cells counted, times 500 parasites counted, gives a parasite density of 20,000 parasites/microL. An alternate expression of parasitemia is the percentage of erythrocytes that are parasitized. In this example, the percent parasitemia is 20,000 parasites/microL divided by 4,000,000 (the average number of erythrocytes per microL in human blood), or 0.5 percent.

As another example, a white blood cell count of 5000/microL, divided by 500 white blood cells counted, times 5 parasites counted, gives a parasite density of 50 parasites/microL. The percent parasitemia is 50 divided by 4,000,000, or 0.001 percent.

Only asexual parasite forms are counted in calculating the parasite density (figure 2). The presence of gametocytes alone indicates a recent infection (and potential for mosquitoes taking a blood meal to become infected) but not infection with actively replicating malaria parasites, which may cause symptoms. Because gametocytes are less susceptible than asexual parasites to many antimalarial drugs, persistence of gametocytes alone following treatment does not indicate drug resistance [25].

P. falciparum, P. malariae, and P. knowlesi infect mature erythrocytes and therefore can mount relatively high parasite densities; a parasitemia of ≥5 percent is very unlikely with any species except P. falciparum. P. vivax, and P. ovale infect only young erythrocytes, so parasite density for these species is typically lower.

Uncomplicated versus severe malaria — In general, the higher the parasite density, the higher the likelihood of severe malaria; this is particularly true for individuals who do not have partial immunity (such as travelers from non-malarious areas and residents of areas with low malaria transmission). However, this association is not observed in all cases, and the relationship between parasite density, total parasite biomass, patient immunity, and severity of presenting symptoms remains to be elucidated [30].

The World Health Organization definition of severe P. falciparum malaria includes hyperparasitemia (>2 percent or 100,000 parasites/microL in low-intensity transmission areas or >5 percent or 250,000 parasites/microL in areas of high stable malaria transmission intensity) [31]. The CDC definition of severe malaria includes parasitemia of ≥5 percent [24]. The peripheral blood smear may reflect a low parasite density relative to the parasite burden present, since parasites frequently sequester in capillaries in the setting of severe malaria.

RAPID DIAGNOSTIC TESTS

General principles — Rapid diagnostic tests (RDTs) for detection of malaria parasite antigens have become important diagnostic tools in resource-limited endemic settings over the past decade due to their accuracy and ease of use. They require no electricity or laboratory infrastructure, give results within 15 to 20 minutes, and can be performed successfully even by health workers with limited training. RDTs provide a qualitative result but cannot provide quantitative information regarding parasite density. RDTs that detect antibodies produced by an infected host are also available; however, these are less useful for diagnosing acute infection.

The approach to RDT selection depends on the epidemiology of infection and goals for control in the region where the test is used [32]. In regions where infection is primarily caused by P. falciparum, use of an assay that detects P. falciparum only may be sufficient and cost-effective. In regions where falciparum and non-falciparum parasites coexist or where P. vivax predominates, use of an RDT that can detect and distinguish between these species is warranted (table 3).

Assay types — RDTs based on antigen detection detect one or more of the following: histidine-rich protein 2 (HRP2; for detection of P. falciparum), Plasmodium lactate dehydrogenase (pLDH; for detection of all species or specific detection of P. falciparum or P. vivax), and aldolase (for detection of all species). Depending on the target antigen(s), an RDT may identify Plasmodium genus only or may distinguish P. falciparum and/or P. vivax infections.

In general, for diagnosis of P. falciparum, RDTs that detect HRP2 are somewhat more sensitive than those that detect pLDH. For diagnosis of non-falciparum species, RDTs that detect pLDH and aldolase appear to be comparable.

Most antigen-detecting RDTs are based on immunochromatographic lateral flow technology; they consist of a nitrocellulose wick packaged as a dipstick, plastic cassette, or paper card (figure 3) [6,10,11,33,34]. One end of the test strip contains labeled antibodies and an agent to lyse red blood cells; a blood sample (5 to 20 mcL) and buffer are placed there, and the liquid migrates along the strip via capillary action, together with the labeled antibodies. The strip also contains a test line (bound antibody that binds parasite antigen, if present) and a control line (bound antibody that binds the migrating-labeled antibody to confirm adequate flow). The development time is typically 15 to 20 minutes.

HRP2 (may detect P. falciparum)

Detection of P. falciparum – Histidine-rich protein 2 (HRP2) is part of a family of P. falciparum histidine-rich proteins [35]. It is only produced by P. falciparum; therefore, use of HRP2-based RDTs is appropriate in regions where P. falciparum is the predominant species (eg, much of sub-Saharan Africa). Combination tests may be useful in areas endemic for multiple Plasmodium species.

HRP2-based RDTs can detect lower levels of parasitemia than RDTs based on other target antigens [36-41]. However, detectable HRP2 antigen may persist in the bloodstream for anywhere from a few days to several weeks after parasitemia is no longer present. In the absence of good quality confirmatory microscopy, there is currently no way to distinguish HRP2 antigenemia resulting from a new infection or persistent infection (ie, resulting from treatment failure) from antigenemia persisting from a recently treated infection. Some endemic countries' national guidelines therefore advise that a positive HRP2 result should be considered to represent a new infection only if the specimen was drawn 7 to 14 days or longer following a previously treated infection; however, it is recognized that this provides general guidance only and the duration of persistent antigenemia varies considerably among individual cases. Therefore, the usefulness of HRP2-based assays may be limited in areas of intense malaria transmission where positive results may occur as a result of prior infection. For the same reason, HRP2-based assays are not useful for monitoring following treatment [42-46].

Interpreting negative HRP2 results – Variability in HRP2 gene sequence may affect test performance in some circumstances [47-49]. Deletions in the genes encoding for HRP2 (and the similar HRP3 protein), leading to false-negative RDT results, have been identified in P. falciparum parasites from the Peruvian Amazon [50,51] and Eritrea [52]. Therefore, HRP2-based tests are not reliable for detection of malaria infection contracted in these areas.

Reports of pfhrp2 and pfhrp3 mutations or deletions have emerged in regions of Africa, Asia, and the Middle East, leading to false-negative RDT results in some cases [53,54]. Strains with both pfhrp2 and pfhrp3 gene deletions are undetectable by HRP2-based RDTs; sometimes these assays can sometimes detect strains with only a pfhrp2 deletion, especially in high-density infections, due to cross reactivity with HRP3 epitopes [48].

The global distribution, frequency, and operational implications of these deletions are not yet fully understood [55]. The World Health Organization (WHO) has identified the Horn of Africa as a region with high levels of pfhrp2 deletions, and Eritrea moved its national policy away from use of HRP2-based RDTs in 2016. The WHO has issued epidemiologic and laboratory guidance for assessment of mutations, as well as a global action plan for surveillance and response [48,53,56,57].

Rarely, false-negative HRP2 results may occur at very high levels of antigenemia or parasitemia due to a prozone-like effect [49,58,59].

pLDH (may detect all species) — Plasmodium lactate dehydrogenase (pLDH) is the terminal enzyme in the malaria parasite glycolytic pathway and is produced by asexual and sexual forms of all Plasmodium species [60]. Plasmodium LDH can be distinguished from human LDH on the basis of unique epitopes and enzymatic characteristics [61,62].

Two types of pLDH-based RDTs are available: those that target a conserved pLDH element in all human malaria species and those that target species-specific regions that distinguish P. falciparum or P. vivax. pLDH antibodies specific for P. ovale and P. malariae have been described but are not yet commercially available [34].

Serum pLDH levels correlate with parasite density and become undetectable at the same time blood smears become negative following antimalarial therapy [62] so that pLDH-based assays may be used for monitoring following treatment and to diagnose treatment failure [46,63-66].

pLDH-based RDTs are less sensitive than HRP2-based tests for detection of P. falciparum infection, especially at relatively low parasite densities (<500 parasites/mcL) [36,39,67,68]. In highly endemic areas, this may have little clinical significance since patients with relatively low parasite densities are often asymptomatic.

Antigenic variation does not appear to significantly affect pLDH-based detection of most parasite species [69].

Aldolase (may detect all species) — Aldolase is an enzyme in the malaria parasite glycolytic pathway that is conserved across all human malaria species. Serum aldolase levels correlate with parasite density and become undetectable with clearance of parasitemia.

For detection of P. falciparum, the sensitivity of aldolase-based assays is generally somewhat lower than that of HRP2-based assays [34,67,70-72]. For detection of non-falciparum infections, the sensitivity of aldolase and pLDH assays is comparable.

Genetic diversity does not appear to be a factor in aldolase-based RDT accuracy [73,74].

Accuracy — There are many different commercial RDT products available. Between 2009 and 2017, the WHO-FIND global RDT testing program conducted standardized laboratory evaluations of RDTs, including testing against well-characterized panels of parasites and antigens [75]. Periodic reports from this program are available online and provided the most systematic and reliable information on accuracy of available malaria RDTs at any given time. As of 2018, the WHO switched to a "prequalification" program, and a list of prequalified malaria RDTs is maintained online [76].

The validity of individual reports on RDT accuracy must be considered in light of the parasite antigen(s) targeted, the comparison standard(s) used (routine microscopy, expert microscopy, polymerase chain reaction [PCR]), malaria epidemiology in the study area (including presence of Plasmodium species and level of transmission), the population in which the RDTs were evaluated, and the personnel performing the RDTs. Other factors influencing RDT accuracy include the type of antibody used (monoclonal versus polyclonal), the source of antigen used to induce the test antibodies, and the epitope targeted by test antibodies [10,34].

Given the volume and variety of data, any statement of RDT accuracy must include a number of caveats. A meta-analysis of RDT diagnosis for uncomplicated P. falciparum malaria in endemic countries (in comparison with microscopy) noted sensitivity and specificity of 93 to 98 percent for assays targeting HRP2 and pLDH [77]. A separate meta-analysis of RDT diagnosis for travelers returning from endemic areas noted that for P. falciparum, tests utilizing HRP2 were more accurate than those utilizing pLDH (negative likelihood ratios 0.08 and 0.13, respectively) [78]. Three-band HRP2 tests had similar negative likelihood ratios but higher positive likelihood ratios compared with two-band tests (34.7 versus 98.5; p = 0.003). Evidence was limited for species other than P. falciparum.

Use

In endemic areas — Use of RDTs in endemic areas has increased rapidly [79]. Globally, the proportion of suspected malaria cases in the public health care sector who receive a parasitologic test has increased in most endemic regions since 2010; the greatest estimated rise, in Africa, was from 36 to 87 percent between 2010 and 2016, mostly due to an increase in RDT use [79].

However, the safety and acceptability of withholding antimalarial treatment from patients with negative RDT results remains an important concern [31,80-82]; administration of presumptive antimalarial treatment for fever has long been a recommended standard of care in many endemic areas [83-85]. Favorable outcomes for management for uncomplicated febrile illness in children based on RDT results have been observed in a number of endemic settings, including Benin [86], Ghana [87], Papua New Guinea [88], Tanzania [89,90], and Zambia [91]. However, RDTs do not have adequate negative predictive value to justify withholding treatment in the setting of severe illness [39,92,93].

Use of RDTs for diagnosis of asymptomatic malaria in pregnancy is the subject of ongoing study [94-97].

Highly sensitive RDTs, also called ultrasensitive RDTs, detect parasite antigen at lower thresholds than conventional RDTs. These have been proposed for detection of asymptomatic parasitemia in malaria elimination programs. Highly sensitive RDTs detect more cases of low-level parasitemia than conventional RDTs but are less sensitive than molecular tests [98-101]. (See 'Molecular tests' below.)

Outside endemic areas — One RDT has been approved by the United States Food and Drug Administration (FDA): BinaxNOW Malaria. The test is a combination assay with antibodies for detection of HRP2 and aldolase.

Among 256 febrile returned travelers, the sensitivity of BinaxNOW for the detection of P. falciparum and non–P. falciparum infection was 94 and 84 percent, respectively [102]. Subsequent experience has confirmed excellent but not perfect performance. Both false-positive and false-negative test results have been described, the latter especially in cases with low parasite density and/or non-falciparum infections [10,103-106]. The CDC advises that RDTs should be used with a positive control (stored blood containing P. falciparum), and both negative and positive results should be confirmed with microscopy [107].

Interpretation of discordant results (between RDT and microscopy) depends on the clinical context and the assay used. For example:

A positive HRP2 RDT with negative microscopy may represent residual antigenemia after successful treatment; alternatively, such results may reflect incomplete treatment. These possibilities must be distinguished based clinical judgment.

A negative HRP2 RDT with positive microscopy may occur in the presence of P. falciparum parasites with pfhrp2/3 gene deletions.

A negative pLDH RDT with positive microscopy may represent a low-grade parasitemia below the RDT detection threshold.

RDTs should not be used for screening donated blood, since the thresholds of detection are not sufficiently sensitive to detect low parasite density [10,108].

Self use — Use of RDTs by travelers for self-diagnosis may be acceptable if expert diagnosis is not immediately available; however, the diagnosis should be confirmed as soon as possible [78,102,109-112]. In one study including more than 150 symptomatic British travelers, 9 percent failed to successfully perform a valid RDT for self-diagnosis; among those who did, the sensitivity and specificity were 97 and 95 percent, respectively (compared with microscopy) [113].

MOLECULAR TESTS — Use of molecular tests for malaria detection is generally limited to reference laboratories and is primarily for research and epidemiologic purposes [114]. The United States Centers for Disease Control and Prevention offers polymerase chain reaction (PCR) confirmation of species and identification of drug resistance mutations for malaria cases diagnosed in the United States [115].

In clinical practice, if P. knowlesi is suspected on epidemiologic grounds, following microscopy to detect parasitemia, PCR should be used to confirm the species and determine the appropriate treatment. (See "Non-falciparum malaria: Plasmodium knowlesi".)

Nucleic acid tests (eg, PCR) are typically used as a gold standard in efficacy studies for antimalarial drugs, vaccines, and evaluation of other diagnostic agents [32,116,117]. The theoretical limit of detection for PCR has been estimated at 0.02 to 1 parasite/microL [118]. Nested PCR is the most sensitive nucleic acid amplification technology; its sensitivity is 400 parasites/mL [119,120]. Parasite deoxyribonucleic acid (DNA) may be amplified after extraction from small volumes (typically 50 to 200 mcL) of whole blood from dried blood spots stored on filter paper following extraction [121,122]. Commonly used PCR assays target genus-specific and species-specific sequences of the 18S small-subunit ribosomal ribonucleic acid (RNA), circumsporozoite surface protein (a nuclear gene encoding a cysteine protease), and the cytochrome b gene [6,123-126].

Low-density malaria infections that are detectable by PCR but below the detection threshold of microscopy or rapid diagnostic tests can contribute to transmission [9,127-134]. Accurate detection of low-density malaria infection is of increasing importance as some malaria-endemic areas move toward elimination [135], with surveillance and screening playing larger roles in program management [136-138]. However, the infrastructure and training required for use of PCR limits its utility for these applications.

Loop-mediated isothermal amplification (LAMP) assays for detection of malaria parasite DNA are being developed to facilitate use of molecular technology in endemic areas [139-144]. Use of LAMP for amplification of DNA occurs at a single temperature so does not require the thermocycler instrumentation necessary for PCR. LAMP generates 109 to 1010 replicates in 15 to 60 minutes, and the resultant turbidity can be detected visually or by turbidimetry, without need for further manipulation. Use of LAMP assays with a variety of target sequences and processing methods has demonstrated variable sensitivity and specificity [145-153].

SOCIETY GUIDELINE LINKS — Links to society and government-sponsored guidelines from selected countries and regions around the world are provided separately. (See "Society guideline links: Malaria".)

SUMMARY

The approach to diagnosis of malaria consists of clinical diagnosis and parasite diagnosis. Parasite-based confirmation should be pursued whenever malaria is suspected clinically. Forms of parasite diagnosis include light microscopy (visualization of parasites in stained blood samples), rapid diagnostic tests (RDTs; detecting antigen or antibody), and molecular techniques detecting parasite genetic material. (See 'Introduction' above.)

Detection of parasites on Giemsa-stained blood smears by light microscopy is the standard tool for diagnosis of malaria; it allows identification of the Plasmodium species as well as quantification of parasitemia. However, microscopy cannot reliably detect very low parasitemia (<5 to 10 parasites/mcL); it is also labor intensive and requires substantial training and expertise. Issues related to smear preparation and interpretation are discussed in detail above. (See 'Light microscopy' above.)

Rapid diagnostic tests for detection of malaria parasite antigens are becoming increasingly important diagnostic tools in resource-limited endemic settings due to their accuracy and ease of use. They require no electricity or laboratory infrastructure, give results within 15 to 20 minutes, and can be performed successfully even by health workers with limited training. RDTs provide a qualitative result but cannot provide quantitative information regarding parasite density. (See 'Rapid diagnostic tests' above.)

The approach to RDT selection depends on the epidemiology of infection and goals for control in the region where the test is used. In regions where infection is primarily caused by P. falciparum, use of an assay that detects P. falciparum only may be sufficient and cost-effective. In regions where falciparum and non-falciparum parasites coexist or where P. vivax predominates, use of an RDT that can detect and distinguish between these species is warranted (table 3). (See 'Rapid diagnostic tests' above.)

RDTs detect one or more of the following antigens: histidine-rich protein 2 (HRP2), Plasmodium lactate dehydrogenase (pLDH), and aldolase. In general, for diagnosis of P. falciparum, RDTs that detect HRP2 are more sensitive than those that detect pLDH. Reliance on HRP2-based RDTs should be avoided in areas where pfhrp2/3 deletions are prevalent including the Peruvian Amazon and Eritrea. For diagnosis of non-falciparum species, RDTs that detect pLDH and aldolase appear to be comparable (see 'Assay types' above).

Use of molecular tests for malaria detection is generally limited to reference laboratories and is primarily for research and epidemiologic purposes. Nucleic acid tests (eg, polymerase chain reaction) are typically used as a gold standard in efficacy studies for antimalarial drugs, vaccines, and evaluation of other diagnostic agents. Loop-mediated isothermal amplification for detection of malaria parasite DNA is being developed as a field assay to facilitate use of molecular technology in endemic areas. (See 'Molecular tests' above.)

ACKNOWLEDGMENT — The UpToDate editorial staff acknowledges Dr. Clinton Murray, who contributed to an earlier version of this topic review.

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Topic 5707 Version 51.0

References

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