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In most malaria-endemic countries, since the introduction of more effective but more expensive antimalarial drug combinations, such as artemisinin combination therapy as first-line treatment, parasitologic confirmation has become essential in routine malaria case management. This medical practice ensures that antimalarial drugs are administered to patients who need them. This is considered as a public health priority by the World Health Organization, in particular in limiting the unnecessary use of inappropriate treatments and thereby avoiding selection and spread of drug-resistant Plasmodium falciparum parasites.
Over the past two decades, malaria rapid diagnostic tests (RDTs) have been developed for use in any situation where the only realistic alternative was the clinical diagnosis of malaria. These diagnostic tests are fast and easy to perform, and do not require electricity or specific equipment.1–3 Currently, 86 malaria RDT products from 28 different manufacturers are available.4 They are all based on the same principle and use antibodies that detect only three groups of antigen. Most products are based on the detection of a P. falciparum-specific protein, either P. falciparum histidine-rich protein 2 (PfHRP2) or P. falciparum lactate dehydrogenase (pfLDH). Some tests detect P. falciparum-specific and pan-specific antigens (aldolase or pan-malaria pLDH) and distinguish a non-falciparum infection from P. falciparum or P. falciparum/ mixed infections.
The purpose of this study was to assess the performance of two new commercially available immunochromatographic assays: the SD Bioline Malaria Ag Pf® (ref. 05FK50) test and the SD Bioline Malaria Ag Pf/Pan® (ref. 05FK60) test (Standard Diagnostics Inc., Suwon City, South Korea). These tests both contain a membrane strip encased in a flat plastic housing. The strip is precoated with two antibodies: one that is specific for P. falciparum HRP2 (both kits) and one that is pan-specific for pLDH from Plasmodium species (SD Bioline Malaria Ag Pf/Pan®).
Our study was carried out between August and October during the season of low malaria transmission at the primary health center in Ampasimpotsy, a rural area in the western foothill of the central highlands in Madagascar. Malaria transmission in this area is low and predominantly seasonal. The main vector is Anopheles funestus and the number of infective bites associated with P. falciparum is estimated to be 1–2 per person each year.5,6 Patients with a fever, or who have had a fever within the past 24 hours, and with typical malaria symptoms were invited to participate in the study. Pregnant women and patients with signs of severe and complicated P. falciparum malaria, as defined by the World Health Organization (2001), were excluded.7 The study protocol was reviewed and approved by the expert committee of the National Malaria Control Program of the Ministry of Health of Madagascar. All subjects included provided informed consent.
Venous blood samples (3 mL) were collected in EDTA anti-coagulation tubes, and thick and thin blood films were prepared by a trained technician. Immunochromatographic tests were immediately carried out according to the manufacturers’ instructions. All tests were kept at room temperature and opened just before use to avoid humidity damage. An RDT result was considered positive when both the internal control and the test band were stained (irrespective of the intensity of the staining). A test result was considered negative if only the internal control was stained. The result of an RDT was considered invalid if the internal control was not stained. Patients with a positive result for the RDT were promptly treated, as specified by the National Malaria Policy, with a combination of artesunate and amodiaquine (Falcimon®; Cipla Ltd., Mumbai, India).8 Blood and blood films were sent to the malaria unit laboratory within 24 hours in a controlled cool box at 4°C. Thick and thin blood smears were stained with 10% Giemsa for 10 minutes. A microscopist who was blind to the results of clinical diagnosis and RDTs examined the blood films for parasites and identified the parasite species. A minimum of 200 consecutive fields were counted in the thick blood film before a slide was classified as negative. Depending on the parasite density, parasites in thick blood films were counted against 200 or 500 leukoytes. Parasite density was estimated assuming 8,000 leukocytes/µL of blood. As previously described,9 parasite DNA was extracted and Plasmodium species were detected using a real-time polymerase chain reaction (PCR) with a RotorGene 3000 thermocycler (Corbett Life Science, Sydney, New South Wales, Australia) carried out by technicians blind to the results of microscopy and RDT testing.
Data were entered, processed, and analyzed using Epi-Info version 3.3.2 software (Centers for Disease Control and Prevention, Atlanta, GA). The RDT results were compared with the reference method (combination of real-time PCR and Giemsa stain microscopy results) for sensitivity and specificity. Sensitivity was calculated as the proportion of samples with malaria parasites detected using the reference method that gave positive RDT results. Specificity was measured by the proportion of samples negative by the reference method that showed negative RDT results. Positive and negative predictive values were the proportion of all positive samples that were true positive samples and the proportion of all samples negative that were true negative samples.
A total of two hundred patients 0.5–63 years of age (mean ± SD age = 13.8 ± 14.9 years) were recruited; 40% were < 5 years of age, 26.5% were 5–15 years of age, and 33.5% were > 15 years of age. The male:female ratio was 1.2:1. The mean ± SD axillary temperature was 38.1 ± 1°C (range = 36.1–40.7°C) and the mean ± SD parasitemia was 16,757 ± 42,631 parasites/µL (range = 16–285,000 parasites/µL). Thirteen percent of these patients reported that they had received anti-malarial therapy before consultation (sulfamethoxazol-trimethoprim in 56%, chloroquine in 28%, sulfadoxine-pyrimethamine in 8%, and quinine in 8%).
Three results were discordant between microscopy and real-time PCR: among two isolates classified as P. falciparum malaria by real-time PCR, one was negative and one was classified as P. vivax malaria by microscopy; the third isolate was classified as mixed P. falciparum/P. vivax malaria by real-time PCR was classified as P. falciparum malaria by microscopy. The three isolates showed the same results as the real-time PCR results. The reference method was positive for 109 (54.5%) of the 200 malaria cases: P. falciparum was present in 78%, P. vivax in 17.4%, P. malariae in 2.8%, and mixed infections with P. falciparum/P. vivax or P. falciparum/P. malariae in 0.9% of the positive specimens (Table 1
). Three results were invalid: two (1%) with the SD Bioline Malaria Ag Pf® (ref. 05FK50) test and one (0.5%) with the SD Bioline Malaria Ag Pf/Pan® (ref. 05FK60) test. The performance of the two RDTs is shown in Table 2. The sensitivity of the RDTs for Plasmodium spp. at different levels of parasitemia is summarized in Table 3. Consistent with the World Health Organization recommendation for RDT performance, the two RDTs had sensitivities greater than 95% for samples with parasitemia levels 
100 parasites/µL (98.4%, 95% confidence interval = 92.5–99.9% for both tests).
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In conclusion, these two new RDTs performed to a similar level as other commercially available devices. These results are consistent with World Health Organization recommendations for RDT performance and offer a good alternative tool for the diagnosis of malaria in disease-endemic areas.
Received May 21, 2008. Accepted for publication July 17, 2008.
Acknowledgments: We thank the Ministry of Health of Madagascar for allowing us access to malaria patients and the patients for participating in the study.
Financial support: This study was supported by Kozone, representing Standard Diagnostics Inc. in Madagascar. Authors’ addresses: Arsène Ratsimbasoa, Laza Fanazava, Rogelin Radrianjafy, and Didier Ménard, Malaria Unit Research, Institut Pasteur de Madagascar, Antananarivo, Madagascar. Julien Ramilijaona and Hughes Rafanomezantsoa, Primary Health Centre of Ampasimpotsy, Ampasimpotsy, Madagascar.
* Address correspondence to Didier Ménard, Malaria Research Unit, Institut Pasteur de Madagascar, BP 1274, Antananarivo 101, Madagascar. E-mail: dmenard{at}pasteur.mg 
Authors’ addresses: Arsène Ratsimbasoa, Laza Fanazava, Rogelin Radrianjafy, and Didier Ménard, Malaria Research Unit, Institut Pasteur de Madagascar, BP 1274-Antananarivo 101, Madagascar. Julien Ramilijaona and Hughes Rafanomezantsoa, Primary Health Centre of Ampasimpotsy, NGO ASA, Ampasimpotsy, Madagascar.
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  M. T. Makler and R. C. Piper Rapid Malaria Tests: Where Do We Go After 20 Years? Am J Trop Med Hyg, December 1, 2009; 81(6): 921 - 926. [Abstract] [Full Text] [PDF]
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