Approach to Malaria Diagnosis

A clinician who faces a case of fever would need answers to the following questions:

  • Is it malaria?
  • If yes;
    • What is the species?
    • Is it severe?
    • Is it new/ recurrence?
    • Is it active?

At present, ONLY the peripheral smear can provide answers to ALL these questions on a single test. With trained technicians, blood smear remains the gold standard for detection, for speciation, for parasite count and for identification of different forms of the parasites. Therefore, rely ONLY on peripheral smear for speciation, parasite count and identification of parasite forms. The QBC test performed by trained personnel could match a thick blood smear for detection of malaria. The RDTs cannot be used as reliable tools for diagnosis of malaria.

Approach to Malaria Diagnosis

approach-diagnosisEvery case of fever, in an area of malaria transmission and among travelers returning from such areas, must be tested for malaria. In many remote and poor parts of India and elsewhere where malaria is endemic, facilities for malaria microscopy are either non existent or inaccessible. Microscopes, stains, power sources or technicians are unavailable either in toto or in parts. Diagnosing malaria in such areas is indeed a challenge.

The National Vector Borne Disease Programme in India recommends peripheral smear microscopy for all fever cases wherever facilities are available. In remote areas where microscopy is unavailable for 24 hours or more, bivalent RDT can be used to diagnose malaria. In some parts of India, QBC is being used instead of blood smear microscopy; however, this method is not approved by the NVBDCP. If a patient is positive with either microscopy or RDT, he/she must be started on relevant anti malaria drugs, based on the species. As RDT is neither very sensitive nor very specific for malaria, it is desirable to confirm every positive test with a blood smear at the earliest opportunity, the smear having been collected before initiating the treatment. All negative tests on microscopy may also need repeat tests at 12-24 hour intervals, on 2-3 occasions, to rule out malaria. Negative tests on RDT may also be candidates for repeat examination on blood smear, particularly when the patient is having features of severe illness wherein RDT may show a false negative test due to prozone effect. RDT may show false positive tests due to persistent antigenemia for 2 months after a primary malaria infection and this fact must be borne in mind while interpreting the test result in patients who return with fever within that period.

 © ©BS Kakkilaya | Last Updated: Mar 10, 2015

Other Tests

Non-Microscopic Tests

Several attempts have been made to take the malaria diagnosis out of the realm of the microscope and the microscopist. Important advances have been made in diagnostic testing, including fluorescence microscopy of parasite nuclei stained with acridine orange, rapid dipstick immunoassay, and Polymerase Chain Reaction assays.[1] These tests involve identification of the parasitic antigen or the antiplasmodial antibodies or the parasitic metabolic products. Nucleic acid probes and immunofluorescence for the detection of Plasmodia within the erythrocytes; gel diffusion, counter-immunoelectrophoresis, radio immunoassay, and enzyme immunoassay for malaria antigens in the body fluids; and hemagglutination test, indirect immunofluorescence, enzyme immunoassay, immunochromatography, and Western blotting for anti-plasmodial antibodies in the serum have all been developed. These tests have found some limited applications in research, retrograde confirmation of malaria, investigation of cryptic malaria, transfusion blood screening, and investigation of transfusion acquired infections.

Rapid Diagnostic Tests (RDTs) detect species-specific circulating parasite antigens targeting either the histidine-rich protein-2 of P. falciparum or a parasite-specific lactate dehydrogenase. Although the dipstick tests may enhance diagnostic speed, microscopic examination remains mandatory in patients with suspected malaria, because occasionally these dipstick tests are negative in patients with high parasitemia, and their sensitivity below 100 parasites/μl is low.

Tests based on polymerase chain reaction for species-specific Plasmodium genome are more sensitive and specific than are other tests, detecting as few as 10 parasites/μl blood. Antibody detection has no value in the diagnosis of acute malaria. It is mainly used for epidemiologic studies.[6-8]

Polymerase Chain Reaction (PCR): Using the non-isotopically labelled probe following PCR amplification, it is possible to detect malaria parasites. In travelers returning to developed countries, studies based on PCR have been found to be highly sensitive and specific for detecting all 4 species of malaria, particularly in cases of low level parasitemia and mixed infections. The PCR test is reportedly 10-fold more sensitive than microscopy, with one study reporting a sensitivity to detect 1.35 to 0.38 parasites/µL for P. falciparum and 0.12 parasites/µL for P. vivax. The PCR test has also been found useful in unraveling the diagnosis of malaria in cases of undiagnosed fever.

Detection Of Antimalarial Antibodies:

Antibodies to the asexual blood stages appear a few days after malarial infection, increase in titer over the next few weeks, and persist for months or years in semi-immune patients in endemic areas, where re-infection is frequent. In non-immune patients, antibodies fall more rapidly after treatment for a single infection and are undetectable in 3-6 months. Re-infection/relapse induces a secondary response with a rapidly increasing antibody titer.

Malarial antibodies can be detected by immunofluorescence or enzyme immuno assay. It is useful in epidemiological surveys, for screening potential blood donors and occasionally for providing evidence of recent infection in non-immunes. In future, detection of protective antibodies will be important in assessing the response to malaria vaccines.

Intraleucocytic malaria pigment: Intraleucocytic malaria pigment has been suggested as a measure of disease severity in malaria. In a study of 146 children aged 6 months to 14 years in 4 categories – cerebral malaria, mild malaria, asymptomatic malaria and ‘no malaria’- in Ibadan, Nigeria, an area of intense malaria transmission in Africa, the proportion of pigment-containing neutrophils showed a clear rise across the spectrum no malaria–asymptomatic malaria–mild malaria–cerebral malaria (median values 2.0%, 6.5%, 9.0% and 27.0%, respectively; P < 0.0001). The proportion of pigment-containing monocytes did not differ significantly between the mild malaria, asymptomatic malaria and no malaria groups but the cerebral malaria group had a higher median value than the other 3 groups. The ratio of pigment-containing neutrophils to pigment-containing monocytes showed the same trend across the groups of subjects as was observed with the number of pigment-containing neutrophils. The study concluded that the pigment-containing neutrophil count is a simple marker of disease severity in childhood malaria in addition to the parasite count. (Amodu OK, Adeyemo AA, Olumese PE, Gbadegesin RA. Intraleucocytic malaria pigment and clinical severity of malaria in children. Trans R Soc Trop Med Hyg. 1998 (Jan-Feb); 92(1):54-56)


Flowcytometry and automated hematology analyzers have been found to be useful in indicating a diagnosis of malaria during routine blood counts. In cases of malaria, abnormal cell clusters and small particles with DNA fluorescence, probably free malarial parasites, have been seen on automated hematology analyzers and it is suggested that malaria can be suspected based on the scatter plots produced on the analyzer. Automated detection of malaria pigment in white blood cells may also suggest a possibility of malaria with a sensitivity of 95% and specificity of 88%. On flow cytometric depolarized side scatter, the average relative frequency of pigment carrying monocytes was found to differ among semi-immune, non-immune and malaria negative patients.

Mass spectrometry:

A novel method for the in vitro detection of the malarial parasite at a sensitivity of 10 parasites/µL of blood has been recently reported. It comprises a protocol for cleanup of whole blood samples, followed by direct ultraviolet laser desorption time-of-flight mass spectrometry. Intense ion signals are observed from intact ferriprotoporphyrin IX (heme), sequestered by malaria parasites during their growth in human red blood cells. The laser desorption mass spectrum of the heme is structure-specific, and the signal intensities are correlated with the sample parasitemia. Many samples could be prepared in parallel and measurement per sample may not take longer than a second or so. However, the remote rural areas without electricity are not hospitable for existing high-tech mass spectrometers. Future improvements in the equipment and technique can make this method deployable and useful.

Other investigations: Total and differential count, hemoglobin, blood glucose, serum bilirubin, serum creatinine, BUN, AST, ALT, Prothrombin time, urine analysis etc. may be done as needed.

Widal test may be positive, even up to a dilution of 1:320 for ‘O’ and H’ and at lower titres for ‘AH’ and ‘BH’. Any or all the four may be positive, suggesting a non-specific response. A positive Widal test in a patient with confirmed malaria should not therefore be considered as suggestive of typhoid fever.

Further reading:

  1. Hanscheid T, Melo-Cristino J, Pinto BG. Automated detection of malaria pigment in white blood cells for the diagnosis of malaria in Portugal. Am J Trop Med Hyg.2001 May-Jun;64(5-6):290-2
  2. Ben-Ezra J, St. Louis M, Riley RS. Automated malarial detection with the Abbott Cell-Dyn 4000 hematology analyzer. Lab Hematol. 2001;7:61-64
  3. Kramer B, Grobusch MP, Suttorp N et al. Relative frequency of malaria pigment-carrying monocytes of nonimmune and semi-immune patients from flow cytometric depolarized side scatter. Cytometry. 2001 Oct 1;45(2):133-40.
  4. Demirev PA, Feldman AB, Kongkasuriyachai D et al. Detection of malaria parasites in blood by laser desorption mass spectrometry. Anal Chem 2002 Jul 15;74(14):3262-6
  5. Mann M. Mass tool for diagnosis. Nature 2002 Aug 15;418(6899):731-2

 © ©BS Kakkilaya | Last Updated: Mar 10, 2015

Rapid Diagnosis of Malaria

See Kakkilaya BS. Rapid Diagnosis of Malaria. Lab Medicine. 2003 Aug;8(34):602-608 [See]

Although the peripheral blood smear examination that provides the most comprehensive information on a single test format has been the “gold standard” for the diagnosis of malaria, the immunochromatographic tests for the detection of malaria antigens, developed in the past decade, have opened a new and exciting avenue in malaria diagnosis. However, their role in the management and control of malaria appears to be limited at present.

Immunochromatographic Tests for Malaria Antigens

Immunochromatographic tests are based on the capture of the parasite antigens from the peripheral blood using either monoclonal or polyclonal antibodies against the parasite antigen targets. Currently, immunochromatographic tests can target the histidine-rich protein 2 of P. falciparum, a pan-malarial Plasmodium aldolase, and the parasite specific lactate dehydrogenase. These RDTs do not require a laboratory, electricity, or any special equipment.

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 red cell membrane surface, and shown to remain in the blood for at least 28 days after the initiation of antimalarial therapy. Several RDTs targeting PfHRP2 have been developed.

Plasmodium aldolase is an enzyme of the parasite glycolytic pathway expressed by the blood stages of P. falciparum as well as the non-fa1ciparum malaria parasites. Monoclonal antibodies against Plasmodium aldolase are pan-specific in their reaction and have been used in a combined ‘P.f/P.v’ immunochromatographic test that targets the pan malarial antigen (PMA) along with PfHRP2.

Parasite lactate dehydrogenase (pLDH) is a soluble glycolytic enzyme produced by the asexual and sexual stages of the live parasites and it is present in and released from the parasite infected erythrocytes. It has been found in all 4 human malaria species, and different isomers of pLDH for each of the 4 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.

RDT Test Format

RDT Test Format

The Rapid Malaria Tests: The RDTs have been developed in different test formats like the dipstick, strip, card, pad, well, or cassette; and the latter has provided a more satisfactory device for safety and manipulation. The test procedure varies between the test kits. In general, the blood specimen (2 to 50µL) is either a finger-prick blood specimen, anticoagulated blood, or plasma, and it is mixed with a buffer solution that contains a hemolyzing compound and a specific antibody that is labeled with a visually detectable marker such as colloidal gold. In some kits, labeled antibody is pre-deposited during manufacture and only a lysing/washing buffer is added. If the target antigen is present in the blood, a labeled antigen/antibody complex is formed and it migrates up the test strip to be captured by the pre-deposited capture antibodies specific against the antigens and against the labeled antibody (as a procedural control). A washing buffer is then added to remove the hemoglobin and permit visualization of any colored lines formed by the immobilized antigen-antibody complexes. The pLDH test is formatted to detect a parasitemia of >100 to 200 parasites/µL and some of the PfHRP2 tests are said to detect asexual parasitemia of >40 parasites/µL.

rdtestThe PfHRP2 test strips have 2 lines, I for the control and the other for the PfHRP2 antigen. The PfHRP2/PMA test strips and the pLDH test strips have 3 lines, 1 for control, and the other 2 for P. falciparum (PfHRP2 or pLDH specific for P. falciparum) and non-falciparum antigens (PMA or pan specific pLDH), respectively. Change of color on the control line is necessary to validate the test and its non-appearance, with or without color changes on the test lines, invalidates the test. With color change only on the control line and without color change on the other lines, the test is interpreted as negative. With the PfHRP2 test, color change on both the lines is interpreted as a positive test for P. falciparum malaria. With the PfHRP2/PMA [The immuno chromatographic test (ICT Malaria P. f. /P.v.test)] and the pLDH tests, color change on the control line and the pan specific line indicates non-fa1ciparum infection and color change on all the 3 lines indicates the presence of P. falciparum infection, either as monoinfection or as a mixed infection with nonfa1ciparum species. Also, if the PfHRP2 line is visible when the PMA line is not, the test is interpreted as positive forP.falciparum infection. Mixed infections of P. falciparum with the non-falciparum species cannot be differentiated from pure P. falciparum infections. However, with regard to the pLDH test, it is claimed that in the presence of P. vivax infection, the genus specific line is much darker and more intense than the species specific line due to the presence of all the stages of the parasite in the blood.

It is also claimed that the rapid diagnostic tests can be performed by individuals with minimal training. With the different tests that are currently available, the procedure may involve 2 to 6 steps and take 5 to 30 minutes. The cost of the RDT also varies from test to test and from country to country, ranging from US $1.20 to $13.50 per test.

Problems with RDTs:

Cross-reactions with autoantibodies: Studies have reported cross reactivity of the various RDTs with autoantibodies such as rheumatoid factor, resulting in false positive tests for malaria. Studies in patients with positive rheumatoid factor have shown that the false positive reactions are higher with the PfHRP2 tests using IgG capture antibody (16.5% to 83% ) compared to the PfHRP2 tests using IgM antibodies (6.6%) and the pLDH test (3.3%). Cross reactivity of the PMA antibody with rheumatoid factor does not appear to occur.

Sensitivity: RDTs for the diagnosis of P. falciparum malaria generally achieve a sensitivity of >90% at densities above 100 parasites per µL blood and the sensitivity decreases markedly below that level of parasite density. Many studies have achieved >95% sensitivity at parasitemia of ~500 parasites/µL, but this high parasitemia is seen in only a minority of patients. For the diagnosis of P. vivax malaria, the PfHRP2/PMA test has a lower sensitivity compared to that for P. falciparum malaria; however, the pLDH test has an equal or better sensitivity for P. vivax malaria compared to P. falciparum malaria. For the diagnosis of P. malariae and P. ovale infections, the sensitivity is lower than that of P. falciparum malaria at all levels of parasitemia on both the PfHRP2/PMA and the pLDH tests. The specificity appears to be better with the pLDH test than the PfHRP2/PMA test for both P. falciparum and non-falciparum malaria.

The sensitivity of the RDTs at low levels of parasitemia and for non-immune populations remains a problem. Compared to microscopy, the PfHRP2/PMA tests were found to be less sensitive in detecting asymptomatic patients, particularly at low parasitemias. The sensitivity of the pLDH test in field studies was also found to be lower at low parasitemias in field studies. The comparisons between the PfHRP2/PMA test and the pLDH test in field studies have yielded variable results, but the pLDH tests were found to have a better specificity for P. vivax. In one study, PfHRP2, PfHRP2/PMA, and pLDH tests had a sensitivity of <75% at parasitemias of <1,000/m L. Of concern is the fact that in nonimmune individuals, symptomatic malaria can occur at parasite densities that are below the detection threshold of currently available RDTS. In a cross-sectional malaria survey, 84.1% patients with P. falciparum infection had a parasitemia of <500/µL and the sensitivity of the PfHRP2 test was only 23.3% at this level of parasitemia. The level of parasitemia encountered in P. vivax infection rarely exceeds 1% (50,000/µL) and usually is much lower. The level of parasitemia forP. malariae and P. ovale are lower than for P. vivax, and the affinity of the panspecific antibodies for these parasites is also lower. Lower levels of parasitemia are also common in nonimmune patients treated with antimalarial chemoprophylaxis. This would mean that P. falciparum infections with low levels of parasitemia and a significant proportion of symptomatic, non-immune patients with P. vivax (or other non-falciparum) malaria may be missed by the RDTs.

Further, the RDTs have been reported to give false negative results even at higher levels of parasitemia. Therefore, in cases of suspected severe malaria or complex health emergencies, a positive result may be confirmatory but a negative result may not rule out malaria. Further, a negative RDT result should always be confirmed by microscopy. It should be emphasized that P. falciparum malaria, a potentially lethal disease, must not be missed because of a false-negative dipstick test. It has been suggested that in such cases, 1 in 10 dilution of a negative sample with 0.9% sodium chloride may help to exclude the prozone phenomenon.

False Positivity: False positive tests can occur with RDTs for many reasons. Potential causes for PfHRP2 positivity, other than gametocytemia, include persistent viableasexual-stage parasitemia below the detection limit of microscopy (possibly due to drug resistance),persistence of antigens due to sequestration and incomplete treatment, delayed clearance of circulating antigen (free or in antigen-antibody complexes) and cross reaction with non-falciparum malaria or rheumatoid factor. Proportion of persistent positivity has been linked to the sensitivity of the test, type of test, degree of parasitemiaand possibly the type of capture antibody.

False negativity: On the other hand, false negative tests have been observed even in severe malaria with parasitemias >40000 parasites/µl. This has been attributed to possible genetic heterogeneity of PfHRP2 expression, deletion of HRP-2 gene, presence of blocking antibodies for PfHRP2 antigen or immune-complex formation, prozone phenomenon at high antigenemia or to unknown causes.

Cross reactions between Plasmodia species and problems in identifying non-falciparum species: Cross reaction of PfHRP2 with non-falciparum malaria could give false positive results for P. falciparum and mixed infections containing asexual stages of P. falciparum could be interpreted as negative in about one third of the patients.

Another major difficulty still encountered by the use of RDTs is the correct identification of Plasmodium species, particularly in areas where nonfalciparum malaria is prevalent. The PfHRP2 tests can detect only P. falciparum infection and would miss the more common non-falciparum malaria in areas where other Plasmodium species are co-endemic.

Multiple Influences: The performance of the RDTs is reported to be influenced by a multitude of factors like the type of the parasite and the level of parasitemia; the type of test; the target antigen and the capture antibody; the expression of the target antigens on the parasites and the presence of several isomers; the presence of gametocytemia; persistent antigenemia or sequestration of the parasites; cross-reactions with other malaria species and with autoantibodies; batch quality variations in test strips; prozone phenomenon; and prior treatment. The interpretation of the color changes to identify the malaria infection is influenced by the level of training, the type of instructions, and in case of self-use, by the state of the patient. The inability to quantify and differentiate between the sexual and asexual parasitemia could pose problems in the areas of high transmission and in cases of incomplete treatment.

The sensitivity and specificity of the RDTs, and hence the diagnosis and treatment of malaria based on the RDTs, are influenced by the positive results due to causes other than malaria antigenemia, and the negative results due to causes other than low parasitemia. Therefore, the identification of the color changes on the RDT strips may look simple but the interpretation of the result would require the knowledge of the malarial dynamics and of the possible errors with the RDTs. Otherwise, the RDTs may raise more questions than answers, and the insufficient accuracy of the RDTs could increase the number of incorrect malaria diagnoses.

Persistence of antigens: All the antigens targeted by the RDTs are expressed by the asexual as well as the sexual forms of the parasites and persistent antigenemia can cause positive tests on RDTs up to one month. With the schizonticidal drugs having no effect on the gametocytes of P.falciparum (except for the artemisinin compounds), RDTs may not be reliable tools to predict the therapeutic response.

Interpretation: Although the RDTs have been reported to be useful and easy tools for field surveys in remote forests and villages, some studies have found that the experience and the level of training of the field staff can influence the sensitivity and specificity of these tests and have reported questionable results or failure to interpret the results in 1.7% to 3.75% of the PfHRP2/PMA test strips.

Lower sensitivity in detecting asymptomatic patients, large numbers of positive tests due to persistent antigenemia following incomplete treatment, inability to differentiate the mixed infections and the non-falciparum species, and inability to differentiate between the various stages of the parasite limit the value of the RDTs in active surveillance in the field. The cost of the RDTs has also been considered as a major obstacle for their large scale use in field studies.

The RDTs have been evaluated for the diagnosis of malaria in travelers, as self-use kits and at the laboratories. Studies on self-use by travelers have raised doubts over the reliability of interpretation of the RDTs by travelers. In one study, only 68% of the European tourists to Kenya were able to perform the test correctly, and 10 out of 11 with malaria failed to diagnose themselves correctly. High number of false negative results have also been reported.

Comparison of Rapid Diagnostic Tests for Malaria Antigens
PfHRP2 tests PfHRP2 and PMA test pLDH test
Target antigen Histidine rich protein 2 of P. falciparum,water soluble protein expressed on RBC membrane Pan-specific Plasmodium aldolase. parasite glycolytic enzyme produced by all species and PfHRP2 Parasite lactate dehydrogenase. parasite glycolytic enzyme produced by all species
General test format 2 lines 3 lines 3 lines
Capability Detects P. falciparum only Can detect all 4 species Can detect all 4 species
Non-falciparum species Not detected Detected; differentiation between the 3 not possible Detected; differentiation between the 3 not possible
Mixed infections of P.falciparum with non-falciparum species Appear as P. falciparum; differentiation not possible Appear as P. falciparum; differentiation not possible Appear as P. falciparum; differentiation not possible
Detection limit >40-100 parasites/µL Higher for P. vivax and other non-falciparum species > 100-200 parasites/µL for P. falciparum andP. vivax; may be higher for P. malariae andP. ovale
Post-treatment persistence of antigens Reported up to 31 days Reported; longer for pan specific antigenemia than for PfHRP2 Reported up to 1 -3 weeks
Cross-reactivity between malarial species Reported Reported Reported
Cross-reactivity with auto antibodies Reported, high (up to 83% with rheumatoid factor) Not known Reported. low (3.3% with rheumatoid factor)
Indication of viability of parasites No No Positive test indicates presence of viable parasitemia
Comparison of Peripheral Blood Smear Examination and RDTs for Malaria
Peripheral Smear Rapid Diagnostic Tests
Format Slides with blood smear Test strip
Equipment Microscope Kit only
Training Trained microscopist ‘Anyone with a little training’
Test duration 20-60 minutes or more 5-30 minutes
Test result Direct visualization of the parasites Color changes on antibody coated lines
Capability Detects and differentiates all plasmodia at different stages Detects malaria antigens (PfHRP2/ PMA/pLDH) from asexual and/or sexual forms of the parasite
Detection threshold 5-10 parasites/µL of blood 1 00-500/µL for P. falciparum, higher for non-falciparum
Species differentiation Possible Cannot differentiate among non-falciparum species; mixed infections of P.falciparum and non-falciparum appear as P. falciparum
Quantification Possible Not possible
Differentiation between sexual and asexual stages Possible Not possible
Disadvantages Availability of equipment and skilled microscopists, particularly at remote areas and odd hours Unpredictable efficiency at low and very high parasitemia; cross reactions among plasmodial species and with auto-antibodies; persistence of antigens
Status Gold standard Not yet approved by the FDA
Cost per test US$ 0.12-0.40 US$ 1 .20-13.50

A potential problem with the dipstick test is that the circulating antigen will be detectable for many days even after the elimination of viable P. falciparum from the blood stream. A positive test therefore may not always indicate an active infection.

US FDA approves RDT: On June 13, 2007, the U.S. Food and Drug Administration (FDA) approved the first RDT for use in the United States. This RDT is approved for use by hospital and commercial laboratories, not by individual clinicians or by patients themselves. It is recommended that all RDTs are followed-up with microscopy to confirm the results and if positive, to quantify the proportion of red blood cells that are infected. [See] [Also See]

See this article cited in US FDA CBEA Presentation

Click for a RDT info from manufacturers and suppliers of RDTs

Further Reading:

  1. Anthony Moody. Rapid Diagnostic Tests for Malaria Parasites. Clinical Microbiology Reviews 2002;15(1):66-78 At
  2. Endeshaw T, Gebre T, Ngondi J et al. Evaluation of light microscopy and rapid diagnostic test for the detection of malaria under operational field conditions: a household survey in Ethiopia.  Malaria Journal. 2008;7:118 Full Text at
  3. Msellem MI, Mårtensson A, Rotllant G, Bhattarai A, Strömberg J, et al. Influence of Rapid Malaria Diagnostic Tests on Treatment and Health Outcome in Fever Patients, Zanzibar—A Crossover Validation Study PLoS Med2009;6(4): e1000070. Free Full Text
  4. Bisoffi Z, Gobbi F, Angheben A, Van den Ende J. The Role of Rapid Diagnostic Tests in Managing Malaria. PLoS Med. 2009;6(4): e1000063 Free Full Text
  5. Jacek Skarbinski et al. Effect of Malaria Rapid Diagnostic Tests on the Management of Uncomplicated Malaria with Artemether-Lumefantrine in Kenya: A Cluster Randomized Trial Am. J. Trop. Med. Hyg., 80(6), 2009, pp. 919-926 Available at
  6. Clinton K. Murray, Jason W. Bennett. Rapid Diagnosis of Malaria: Review Article. Interdisciplinary Perspectives on Infectious Diseases. Volume 2009, Article ID 415953, 7 pages doi:10.1155/2009/415953. Available at
  7. Laboratory demonstration of a prozone-like effect in HRP2-detecting malaria rapid diagnostic tests: implications for clinical management Jennifer Luchavez et al.Malaria Journal 2011;10:286 doi:10.1186/1475-2875-10-286 [Full Text] (Added Oct 9, 2011)

© ©BS Kakkilaya | Last Updated: Mar 27, 2015

Microscopic Tests

Diagnosis of malaria involves identification of malaria parasite or its antigens/products in the blood of the patient. Although this seems simple, the efficacy of the diagnosis is subject to many factors. The different forms of the four malaria species; the different stages of erythrocytic schizogony; the endemicity of different species; the population movements; the inter-relation between the levels of transmission, immunity, parasitemia, and the symptoms; the problems of recurrent malaria, drug resistance, persisting viable or non-viable parasitemia, and sequestration of the parasites in the deeper tissues; and the use of chemoprophylaxis or even presumptive treatment on the basis of clinical diagnosis can all have a bearing on the identification and interpretation of malaria parasitemia on a diagnostic test.

The diagnosis of malaria is confirmed by blood tests and can be divided into microscopic and non-microscopic tests.

Microscopic Tests

The microscopic tests involve staining and direct visualization of the parasite under the microscope. For more than hundred years, the direct microscopic visualization of the parasite on the thick and/or thin blood smears has been the accepted method for the diagnosis of malaria in most settings, from the clinical laboratory to the field surveys. The careful examination of a well-prepared and well-stained blood film currently remains the “gold standard” for malaria diagnosis. The most commonly used microscopic tests include the peripheral smear study and the Quantitative Buffy Coat (QBC) test.

The simplest and surest test is the time-honoured peripheral smear study for malarial parasites. None of the other newer tests have surpassed the ‘gold standard’ peripheral smear study.

Remember this:

  • Ask for MP test in all cases of fever and related symptoms and also whenever there is high level of suspicion
  • MP test can be done at any time. Do not wait for typical symptoms and signs or for chills
  • A negative test DOES NOT rule out malaria. Repeated tests may have to be done in all doubtful cases. Duration of the illness, level of parasitemia, expertise of the technician and the method of examination may all have a bearing on the result of the M.P. test

Peripheral smear study for malarial parasites – The MP test

Light microscopy of thick and thin stained blood smears remains the standard method for diagnosing malaria. It involves collection of a blood smear, its staining with Romanowsky stains and examination of the Red Blood Cells for intracellular malarial parasites. Thick smears are 20–40 times more sensitive than thin smears for screening of Plasmodium parasites, with a detection limit of 10–50 trophozoites/μl. Thin smears allow one to identify malaria species (including the diagnosis of mixed infections), quantify parasitemia, and assess for the presence of schizonts, gametocytes, and malarial pigment in neutrophils and monocytes.

The peripheral blood smear provides comprehensive information on the species, the stages, and the density of parasitemia. The efficiency of the test depends on the quality of the equipment and reagents, the type and quality of the smear, skill of the technician, the parasite density, and the time spent on reading the smear. The test takes about 20 to 60 minutes depending on the proximity of the laboratory and other factors mentioned above. It is estimated to cost about 12 to 40 US cents per slide in the endemic countries.

Before reporting a negative result, at least 200 oil immersion visual fields at a magnification of 1000× should be examined on both thick and thin smears, which has a sensitivity of 90%. The level of parasitemia may be expressed either as a percentage of parasitized erythrocytes or as the number of parasites per microliter of blood. In nonfalciparum malaria, parasitemia rarely exceeds 2%, whereas it can be considerably higher (>50%) in falciparum malaria. In nonimmune individuals, hyperparasitemia (>5% parasitemia or >250 000 parasites/μl) is generally associated with severe disease.

The smear can be prepared from blood collected by venipuncture, finger prick and ear lobe stab. In obstetric practice, cord blood and placental impression smears can be used. In fatal cases, post-mortem smears of cerebral grey matter obtained by needle necropsy through the foramen magnum, superior orbital fissure, ethmoid sinus via the nose or through fontanelle in young children can be used.

Preparation of the smear: Use universal precautions while preparing the smears for malarial parasites – use gloves; use only disposable needles/lancets; wash hands; handle and dispose the sharp instruments and other materials contaminated with blood carefully to avoid injury.

  • slides2Hold the third finger of the left hand and wipe its tip with spirit/Savlon swab; allow to dry
  • Prick the finger with disposable needle/lancet; allow the blood to ooze out
  • Take a clean glass slide. Take 3 drops of blood 1 cm from the edge of the slide, take another drop of blood one cm from the first drop of blood
  • Take another clean slide with smooth edges and use it as a spreader and make thick and thin smears. Allow it to dry
  • Slide number can be marked on the thin smear with a lead pencil

Thick smear: The thick smear of correct thickness is the one through which newsprint is barely visible. It is dried for 30 minutes and not fixed with methanol. This allows the red blood cells to be hemolyzed and leukocytes and any malaria parasites present will be the only detectable elements. However, due to the hemolysis and slow drying, the plasmodia morphology can get distorted, making differentiation of species difficult. Thick smears are therefore used to detect infection, and to estimate parasite concentration.

Thin smear: Air dry the thin smear for 10 minutes. After drying, the thin smear should be fixed in methanol. This can be done by either dipping the thin smear into methanol for 5 seconds or by dabbing the thin smear with a methanol-soaked cotton ball. While fixing the thin smear, all care should be taken to avoid exposure of the thick smear to methanol.

Staining: A number of Romanowsky stains like Field’s, Giemsa’s, Wright’s and Leishman’s are suitable for staining the smears. Thick films are ideally stained by the rapid Field’s technique or Giemsa’s stain for screening of parasites. The sensitivity of a thick blood film is 5-10 parasites/µl. Thin blood films stained by Giemsa’s or Leishman’s stain are useful for specification of parasites and for the stippling of infected red cells and have a sensitivity of 200 parasites/µl. The optimal pH of the stain is 7.2.

Slides should be clean and dry. It is better to use neutral distilled water.

Thick films: The thick film is first de-hemoglobinised in water and then stained with Giemsa.

Rapid Giemsa: Prepare a 10% Giemsa in buffered water at pH 7.1. Immerse the slide in the stain for 5 minutes. Rinse gently for 1 or 2 seconds in a jar of tap water. Drain, dry and examine.

Standard Giemsa: Prepare a 4% Giemsa in buffered solution at pH 7.1. Immerse the slide (at least 12 hours old) in stain for 30 minutes. Rinse with fresh water, drain, dry and examine.

Thin films: Thin film examination is the gold standard in diagnosis of malarial infection.

Giemsa stain: Fix with 1-2 drops of methanol. Cover the film with 10% Giemsa stain and leave for 30 minutes, wash with distilled water, drain, dry and examine.

Leishman’s stain: Add 7-8 drops of the stain and leave for 1-2 minutes. Then add 12-15 drops of buffered distilled water, mix thoroughly, leave for 4 – 8 minutes. Then wash off with clean water, drain, dry and examine.

microJaswant Singh Battacharya (JSB) Stain for thick and thin films: This is the standard method used by the laboratories under the National Malaria Eradication Programme in India.

Preparation of the stain:

JSB I stain: Medicinal methylene blue (0.5 g) is dissolved in 500 ml of distilled water and 3 ml of 1% sulphuric acid (H2SO4) is gradually added, followed by 0.5 g of potassium dichromate (K2Cr2O7) when a purple precipitate forms. 3.5 g of disodium hydrogen phosphate dihydrate (Na2HPO4.2H2O) is next added and when the precipitate has dissolved, the solution is boiled in a flask with a reflex condenser for 1 hour. The stain is ready for immediate use.

JSB II stain: 1 g Eosin is dissolved in 500 ml tap water.

Buffered water: 0.22 g of disodium hydrogen phosphate dihydrate (Na2HPO4.2H2O) and 0.74 g of potassium acid phosphate (KH2PO4) are added to 1000 ml of distilled water or filtered tap water.


After dehemoglobinisation, dip the thick smear in JSB II stain two to three times. Wash it by dipping in buffer water two to three times. Then keep the thick film dipped in JSB I stain for 40-60 seconds. Wash it with buffer water. Drain, dry and examine.

Differentiation of Malaria Parasites


Finding P. falciparum P. vivax P. malariae P. ovale
RBC Size Not enlarged Enlarged Not enlarged Enlarged
RBC Shape Round, sometimes crenated Round or oval, frequently bizarre Round Round or oval, often fimbriated
RBC Colour Normal, but may become darker; may have a purple rim Normal to pale Normal Normal
Stipling Maurer’s spots, appear as large red spots, loops and clefts; up to 20 or fewer. Schuffner’s dots, appear as small red dots, numerous. Ziemann’s dots, few tiny dots, rarely detected Schuffner’s dots (James’s dots). Numerous small red dots.
Pigment Black or dark brown; in asexual forms as one or two masses; in gametocytes as about 12 rods Seen as a haze of fine golden brown granules scattered through the cytoplasm Black or brown coarse granules; scattered Intermediate between P. vivax and P. malariae
Early trophozoite (ring) Smallest, delicate; sometimes two chromatin dots; multiple rings commonly found Relatively large; one chromatin dot, sometimes two; often two rings in one cell Compact; one chromatin dot; single Compact; one chromatin dot; single
Schizont Medium size; compact; numerous chromatin masses; coarse pigments; rarely seen in peripheral blood Large; amoeboid; numerous chromatin masses; fine pigments Small; compact; few chromatin masses; coarse pigments Medium size; compact; few chromatin masses; coarse pigments
Gametocyte Crescent shaped, larger and slender; central chromatin Spherical; compact Similar to P. vivax, but smaller and less numerous Like P. vivax, but smaller

Problems: The exacting needs of the blood smear examination are often not met in certain remote and poor parts of the world. Detection of low levels of parasitemia, sequestered parasites of P. falciparum and past infections in aspiring blood donors; ascertaining viability of the detected parasites; difficulties in maintaining the required technical skills and resultant misdiagnosis due to poor familiarity and problems in accessing and activating the facility in emergencies are some of the deficiencies with the blood smear examination.

In falciparum malaria, parasitized erythrocytes may be sequestered in tissue capillaries resulting in a falsely low parasite count in the peripheral blood (‘visible’ parasitemia). In such instances, the developmental stages of the parasite seen on blood smear may help to assess disease severity better than parasite count alone. The presence of more mature parasite forms (>20% of parasites as late trophozoites and schizonts) and of more than 5% of neutrophils containing malarial pigment indicates more advanced disease and a worse prognosis. One negative blood smear makes the diagnosis of malaria very unlikely (especially the severe form); however, smears should be repeated every 6–12 hours for 48 hours if malaria is still suspected.[1-5]

Sometimes no parasites can be found in peripheral blood smears from patients with malaria, even in severe infections. This may be explained by partial antimalarial treatment or by sequestration of parasitised cells in deep vascular beds. In these cases, parasites, or malarial pigment may be found in the bone marrow aspirates. Presence of malarial pigment in circulating neutrophils and monocytes may also suggest the possibility of malaria.

Alternative microscopic methods have been tried, including faster methods of preparation, dark-field microscopy, and stains like benzothiocarboxypurine, acridine orange and Rhodamine-123. Acridine orange has been tried as a direct staining technique, with concentration methods such as thick blood film or the centrifugal Quantitative Buffy Coat system and with excitation filter in the Kawamoto technique. Inability to easily differentiate the Plasmodium species, requirements of expensive equipment, supplies and special training as well as the high cost limit the use of these methods.

Parasitemia in blood films:

Thick Blood Film:

Infected erythrocytes are counted in relation to a predetermined number of WBCs and an average of 8000/µl is taken as standard. 200 leucocytes are counted in 100 fields (0.25 µl of blood). All parasite species and forms including both sexual and asexual forms are counted together.

If >10 parasites are counted, then the following formulae can be applied:

(No. of Parasites/ No. of WBCs counted) x 8000 = No. of parasites/µl

Or if 200 leukocytes are counted,

No. of parasites counted x 40 = No. of parasites/µl

If the parasites are <9, then 500 WBCs should be counted and the formula will be –

No. of parasites counted x 16 = No. of parasites/µl

In the Earle and Perez method, the number of asexual parasites per known volume of blood (usually 5µl) spread as a thick film are counted; this is used only in research studies.

Thin Blood Film:

Determining the percentage of parasitaemia will be essential for P. falciparum. The number of infected red cells (and not number of parasites) in 1000 RBCs is converted to percentage.

This method estimates the percentage of red blood cells infected with malarial parasites. The smear is scanned carefully, one ‘row’ at a time. The total number of red cells and the number of parasitised red cells are tabulated separately. If 1000 red cells are counted, then divide the number of parasitised red cells by 10 to get the percentage (i.e. if 30 out of 1000 cells are parasitised, then the parasitised red cell count is 3%). If lesser red cells are counted, then divide the number parasitized by the total number counted and multiply the result by 100 to obtain a percentage estimate of red blood cells parasitized. If occasional parasites are seen when scanning the smear, but none are identified during the process of counting 300-500 red blood cells, a percentage value of less than 1% of red blood cells parasitized is assigned. An estimate of less than 1% of red blood cells parasitized does not need to be refined, since no clinical predictive value is gained. It is values of 2-3% or above that are of clinical concern.

The “plus system” is less precise as variation in the thickness of the film results in false variation in parasite count.

+ = 1–10 per 100 thick fields.

++ = 11-100 per 100 thick fields.

+++ = 1–10 per thick field.

++++ = >10 per thick field.

Further Reading:

  1. Andrej Trampuz, Matjaz Jereb, Igor Muzlovic, Rajesh M Prabhu. Clinical review: Severe malaria. Critical Care 2003;7:315-323 Available at
  2. Moody AH, Chiodini PL. Methods for the detection of blood parasites. Clin Lab Haematol 2000;22:189-201.
  3. Torres JR. Malaria and babesiosis. Therapy of Infectious Diseases (Edited by: Baddour LM, Gorbach SL). Philadelphia, PA: Saunders 2003;597-613.
  4. Nguyen PH, Day N, Pram TD, Ferguson DJ, White NJ. Intraleucocytic malaria pigment and prognosis in severe malaria. Trans R Soc Trop Med Hyg 1995;89:200-204.
  5. White NJ. Malaria. Manson’s Tropical Diseases (Edited by: Cook GC, Zumla AI, Weir J). Philadelphia, PA: WB Saunders 2003;1205-1295
  6. Lee SH, Kara UA, Koay E, Lee MA, Lam S, Teo D. New strategies for the diagnosis and screening of malaria. Int J Hematol 2002;76(suppl 1):291-293.
  7. Moody A. Rapid diagnostic tests for malaria parasites. Clin Microbiol Rev 2002;15:66-78.
  8. Hanscheid T, Grobusch MP: How useful is PCR in the diagnosis of malaria? Trends Parasitol 2002;18:395-398
  9. New Perspectives – Malaria Diagnosis: Report Of A Joint Who/Usaid Informal Consultation 25–27 October 1999 p 11 At
  10. Castelli F, Carosi G. Diagnosis of malaria infection In Castelli F, Carosi G ed Handbook of malaria infection in the tropics. Organissazione per la cooperazione sanitaria internazionale 1997 pp 114
  13. Warhurst DC, William J Laboratory diagnosis of malaria ACP Broadsheet No 148 J Clin Path 1996;49:533-8
  14. Noppadon Tangpukdee, Chatnapa Duangdee, Polrat Wilairatana, Srivicha Krudsood. Malaria Diagnosis: A Brief Review Korean J Parasitol. 2009 Jun;47(2):93-102. Available at

 Quantitative Buffy Coat (QBC)

The QBC Test, developed by Becton and Dickenson Inc., is a new method for identifying the malarial parasite in the peripheral blood. It involves staining of the centrifuged and compressed red cell layer with acridine orange and its examination under UV light source. It is fast, easy and claimed to be more sensitive than the traditional thick smear examination.

qbcpicMethod: The QBC tube is a high-precision glass hematocrit tube, pre-coated internally with acridine orange stain and potassium oxalate. It is filled with 55-65 microliters of blood from a finger, ear or heel puncture. A clear plastic closure is then attached. A precisely made cylindrical float, designed to be suspended in the packed red blood cells, is inserted. The tube is centrifuged at 12,000 rpm for 5 minutes. The components of the buffy coat separate according to their densities, forming discrete bands. Because the float occupies 90% of the internal lumen of the tube, the leukocyte and the thrombocyte cell band widths and the top-most area of red cells are enlarged to 10 times normal. The QBC tube is placed on the tube holder and examined using a standard white light microscope equipped with the UV microscope adapter, an epi-illuminated microscope objective. Fluorescing parasites are then observed at the red blood cell/white blood cell interface.

The key feature of the method is centrifugation and thereby concentration of the red blood cells in a predictable area of the QBC tube, making detection easy and fast. Red cells containing Plasmodia are less dense than normal ones and concentrate just below the leukocytes, at the top of the erythrocyte column. The float forces all the surrounding red cells into the 40 micron space between its outside circumference and the inside of the tube. Since the parasites contain DNA which takes up the acridine orange stain, they appear as bright specks of light among the non-fluorescing red cells. Virtually all of the parasites found in the 60 microliter of blood can be visualized by rotating the tube under the microscope. A negative test can be reported within one minute and positive result within minutes.

Studies that have compared the QBC with the peripheral smear report that the test is as sensitive as the smear; however, identification of the species and quantification of parasitemia are difficult with the QBC technique. [See references below] Therefore, in spite of the speed and simplicity of QBC technique, it cannot be considered an acceptable alternative to GTF under routine clinical laboratory situation.

Comparison between peripheral smear and QBC test for detecting malaria>
Peripheral smear QBC
Method Cumbersome Easy
Time Longer, 60 – 120 minutes Faster, 15 – 30 minutes
Sensitivity 5 parasites/µl in thick film and 200 / µl in thin film Claimed to be more sensitive, at least as good as a thick film
Specificity Gold standard ? False positives, artifacts may be reported as positive by not-so-well-trained technicians
Species identification Accurate, gold standard Difficult to impossible
Cost Inexpensive Costly equipment and consumables
Acceptability 100% Not so
Availability Everywhere Limited

Accidentally can detect filarial worms

Therefore, whenever in doubt, ask for a peripheral smear study, particularly for species identification. There are instances of cases diagnosed as vivax malaria on the QBC, but soon after developed fatal complications of falciparum malaria.

See this article cited in US FDA CBEA Presentation

More from the Company Web site


  1. Adeoye GO, Nga IC. Comparison of Quantitative Buffy Coat technique (QBC) with Giemsa-stained thick film (GTF) for diagnosis of malaria. Parasitology International. 2007;56(4):308-312 At
  2. Cabezos J, Bada JL. The diagnosis of malaria by the thick film and the QBC: a comparative study of both technics Med Clin (Barc) 1993;101:91-4. At
  3. Pinto MJ, Rodrigues SR, Desouza R, Verenkar MP. Usefulness of quantitative buffy coat blood parasite detection system in diagnosis of malaria. Indian J Med Microbiol 2001;19:219-21. At
  4. Estacio RH, Edwin RE, Cresswell S, Coronel RF, Alora AT. The Quantitative Buffy Coat Technique (QBC) in Early Diagnosis of Malaria: The Santo Tomas University Hospital Experience At
  5. Wang X, Zhu S, Liu Q, Hu A, Zan Z, Yu Q, Yin Q. Field evaluation of the QBC technique for rapid diagnosis of vivax malaria. Bull World Health Organ. 1996;74(6):599-603. At
  6. Clendennen TE, Long GW, Baird JK. QBC® and Giemsa-stained thick blood films: diagnostic performance of laboratory technologists. Transactions of the Royal Society of Tropical Medicine and Hygiene 1995;89(2):183-184. At

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