A novel real-time PCR assay for the detection of Plasmodium falciparum and Plasmodium vivax malaria in low parasitized individuals

Acta Tropica 120 (2011) 40–45

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A novel real-time PCR assay for the detection of Plasmodium falciparum and Plasmodium vivax malaria in low parasitized individuals Seung-Young Hwang a , So-Hee Kim a,b , Ga-Young Lee a , Vu Thi Thu Hang a,b , Chi-Sook Moon c , Jeong Hwan Shin d , Wan-Lim Koo e , Seong-Youl Kim e , Hae-Joon Park e , Han-Oh Park e , Weon-Gyu Kho a,b,f,∗ a

Department of Parasitology, Inje University College of Medicine, Busan 614-735, South Korea Department of Infecious diseases and Malaria, Paik Institute for Clinical Research, Inje University, Busan 614-735, South Korea c Department of Internal Medicine, Busan Paik Hospital, Inje University, Busan 614-735, South Korea d Department of Laboratory Medicine and Paik Institute for Clinical Research, Inje University College of Medicine, Busan 614-735, South Korea e Molecular Diagnostic Research Institute, Bioneer Corporation, Daejeon 306-220, South Korea f Mitochondrial Research Group, Frontier Inje Research for Science and Technology, Inje University, Busan 614-735, South Korea b

a r t i c l e

i n f o

Article history: Received 18 May 2010 Received in revised form 13 April 2011 Accepted 16 May 2011 Available online 6 June 2011 Keywords: Malaria Diagnosis Real-time PCR Low parasitemia

a b s t r a c t The rapid, accurate diagnosis of Plasmodium spp. is essential for the effective control of malaria, especially in asymptomatic infections. In this study, we developed a sensitive, genus-specific, real-time quantitative PCR assay. It was compared with the microscopic examination of Giemsa-stained blood smears and two different molecular diagnostic techniques: nested PCR and multiplex PCR. For the effective quantitative detection of malaria parasites, all reagents were designed with a lyophilized format in one tube. Plasmodium was detected successfully in all 112 clinically suspected malaria patients, including 32 individuals with low parasitemia (1–100 parasites/␮l). The sensitivity threshold was 0.2 parasites/␮l and no PCR-positive reaction occurred when malaria parasites were not present. This may be a useful method for detecting malaria parasites in endemic areas. © 2011 Elsevier B.V. All rights reserved.

1. Introduction Malaria is a highly prevalent disease in tropical and subtropical regions, affecting half of the world’s population in 108 countries. In 2008, there were an estimated 243 million cases, with an estimated 863,000 deaths (WHO, 2009). Therefore, a rapid and accurate diagnosis is essential for effective treatment and control of malaria (Vo et al., 2007). It is especially necessary to develop diagnostic techniques with a high degree of sensitivity for detecting malaria in individuals with asymptomatic malaria and relatively low parasite rates. The microscopic examination of Giemsa-stained blood films is current gold standard laboratory method for malaria diagnosis. However, this technique requires a long observation time and well-trained microscopist to exam samples with low parasitemia (<100 parasites/␮l) (Amexo et al., 2004). Therefore, alternative

diagnostic methods having sensitivity enough to detect a small number of parasites have been developed. Conventional PCR assays have higher sensitivity and specificity than microscopic examination, but they are labor-intensive, timeconsuming, and have amplicon contamination problems (Igbinosa et al., 2010). The recent real-time PCR methods allow the rapid amplification, simultaneous detection, and quantification of target DNA through the use of specific fluorophore-labeled probes (Elsayed et al., 2006). In this report, we described a novel pair of TaqMan probebased real-time PCR assays that have improved reaction time and threshold sensitivity, and that can simultaneously quantify human malaria parasites. And it was also compared with microscopic examination of Giemsa-stained blood smears and two different molecular diagnostic techniques, nested PCR (Snounou et al., 1993) and multiplex PCR (Kho et al., 2003). 2. Materials and methods

∗ Corresponding author at: Department of Parasitology, Inje University College of Medicine, 633-165, Gaegum-dong, Jin-Gu, Busan 614-735, South Korea. Tel.: +82 51 890 6731; fax: +82 51 894 6709. E-mail address: [email protected] (W.-G. Kho). 0001-706X/$ – see front matter © 2011 Elsevier B.V. All rights reserved. doi:10.1016/j.actatropica.2011.05.006

2.1. Blood sample collection and DNA extraction The study enrolled 112 clinically suspected malaria patients who attended Paik Hospital, South Korea between 2003 and 2008.

U10115

M19173, U03079 M54897, L48987

M19172 U03079

M19173 U03079

M19173, U03079 M54897, L48987

GenBank accession no.

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This study was approved by institutional review board for human research of Inje University Busan Paik Hospital, South Korea, and samples were taken only after obtaining written consent regarding participation. Blood samples (5 ml) were drawn into sterile tubes containing EDTA, and thin blood films were prepared at the time of sample collection. Aliquots (500 ␮l) of blood were stored at −20 ◦ C and used subsequently to extract DNA. As negative controls, blood samples from 80 healthy individuals with no history of malaria were collected and stored similarly. Genomic DNA was extracted from 20 ␮l of whole blood using a QIAamp DNA blood kit (QIAGEN, Valencia, CA) according to the manufacturer’s instructions. The DNA was eluted in 100 ␮l of TE buffer (10 mM Tris–HCl, pH 8.0, and 1 mM EDTA) and kept at −70 ◦ C until used.

Mus musculus

GGTATTGGCCTAACATGGCTATG CCTGCTGCCTTCCTTAGATGTG FAM-AGTTCGATTCCGGAGAGGGAGCCT-BHQ1 ACGCACCAGCTCTTCCTCACT CGCGGACAATGGCACTCAT TAMRA-CAGCTCAGTGCCTGGCGCCC-BHQ1 MAL-F MAL-R MAL-TMp IPC-F IPC-R IPC-TMp

Plasmodium sp.

TCA GCT TTT GAT GTT AGG GTA TT GCA TCA AAG ATA CAA ATA TAA GC TAA ACT CCG AAG AGA AAA TTC T UF FR VR

Plasmodium sp. P. falciparum P. vivax

TTA AAC TGG TTT GGG AAA ACC AAA TAT ATT ACA CAA TGA ACT CAA TCA TGA CTA CCC GTC CGC TTC TAG CTT AAA CCA CAT AAC TGA TAC ACT TCC AAG CCG AAG CAA AGA AAG TCC TTA rFAL1 rFAL2 rVIV1 rVIV2

P. falciparum P. vivax

TTA AAA TTG TTG CAG TTA AAA CG CCT GTT GTT GCC TTA AAC TTC rPLU6 rPLU5

Plasmodium sp.

Sequence (5 –3 ) Primer or probe name

For each patient, thin blood films were prepared and stained with Diff-Quick solution (International Reagents, Japan). In order to prevent the biased decision, two experienced technicians examined the smears independently using microscopy in blind manner. Parasites were quantified by counting the number of infected erythrocytes per 200 leukocytes in thin smear fields. Parasitemia was calculated assuming 8000 leukocytes per 1 ␮l of blood (Wilcox, 1960). Patients were considered negative when no parasites were detected in an area where 1000 leukocytes were observed. 2.3. Nested PCR and multiplex PCR Nested PCR was performed according to the protocol established by Snounou et al. (1993). The first PCR reagent mixture consisted of 250 nM each primer (Table 1), 124 ␮M dNTP, 10× PCR buffer (500 mM KCl, 20 mM MgCl2 , 100 mM Tris–HCl, pH 8.3), and 2 units of Taq DNA polymerase (TaKaRa Ex-Taq, TaKaRa, Kyoto, Japan). For each PCR tube, 95 ␮l of PCR mixture and 5 ␮l of sample DNA were used. The reaction was performed with a preliminary 5 min denaturation at 95 ◦ C, followed by 25 cycles of 1 min at 94 ◦ C, 2 min at 58 ◦ C, and 2 min at 72 ◦ C, with a final 5 min extension at 72 ◦ C. The second PCR was performed similarly, except for 30 cycles. Each sample was analyzed in duplicate. The amplification product was detected via ethidium bromide staining after 2% agarose gel electrophoresis. As expected from the sequences, the sizes of the fragments on a gel were 205 bp for Plasmodium falciparum (P. falciparum) and 120 bp for Plasmodium vivax (P. vivax). Multiplex PCR was performed using the three primers previously reported by our laboratory (Kho et al., 2003). The PCR was carried out in a total volume of 20 ␮l, which included 2 ␮l of extracted DNA, 20 mM dNTP, 0.5 pmol forward primer (UF, Table 1), 0.25 pmol each reverse primer (FR and VR, Table 1), 10× PCR buffer, and 0.25 units of Taq DNA polymerase. The reaction involved a 5 min denaturation at 94 ◦ C, followed by 35 cycles of 45 s at 94 ◦ C, 45 s at 57 ◦ C, and 45 s at 72 ◦ C, with a 5 min final extension at 72 ◦ C. Each sample was analyzed in duplicate on a 1% agarose gel. PCR amplification produced DNA fragments of 1451 and 833 bp for P. falciparum and P. vivax, respectively.

Real-time

Multiplex

Nested – second

Nested – first

2.4. Real-time PCR standardization

Types of PCR

Table 1 Primers and probes for PCR of 18S rRNA gene in malaria parasites.

Species

2.2. Microscopic examination

Using Primer Express software (Applied Biosystems, Foster City, CA), a series of primers and dual-labeled fluorescent probes were designed to amplify a species-conserved region of the Plasmodium 18S rRNA gene. The amplicon size was determined as 101 bp. The selected forward/reverse primers (MAL-F/MAL-R and IPC-F/IPC-R) and probes (MAL-TMp and IPC-TMp) of malaria and of internal positive control are shown in Table 1. All components required for the PCR reaction, including 5 units of DNA polymerase, 20 mM dNTPs,

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30 pmol species-conserved primer set, 15 pmol internal positive control primer set, and 30 pmol probe set, were aliquoted for one reaction per microtube and lyophilized using freeze dryer (Virtis, Gardiner, NY). The lyophilized PCR tubes were covered with silver foil to prevent exposure to sunlight and kept at −20 ◦ C until used. To each tube, we added 5 ␮l of sample DNA, 1 ␮l of internal positive control DNA, and 44 ␮l of distilled water. The 50 ␮l PCR mixtures were amplified in an ExicyclerTM 96 real-time PCR system (Bioneer, Daejeon, South Korea). The reaction was performed with an initial 5 min denaturation at 95 ◦ C, followed by 45 cycles of 20 s at 95 ◦ C and 30 s at 55 ◦ C. Each experiment included one reaction mixture without DNA as negative control, and each specimen was runned three times in duplicate for real-time PCR assay. 2.5. Determination of a parasite DNA standard curve As there is no consensus regarding the choice of a quantitative standard curve for real-time PCR, we investigated quantitative values from two different sources, a plasmid DNA containing a specific region of P. vivax 18S rRNA gene that synthesized de novo and parasite genomic DNA. The 500 bp of P. vivax 18S rRNA gene containing the region of PCR amplicon was cloned into pGEM T-easy vector (Promega). The concentration of the plasmid DNA was measured using ultraviolet spectrophotometry. The mass of single copy of plasmid DNA was determined as 3.85 × 10−6 pg. To construct a PCR standard curve, five serial diluted amplicon DNA samples, ranging from 1 × 106 to 1 × 102 copies/reaction, was analyzed three times in duplicate. Meanwhile, for the parasite genomic DNA, a P. vivax infected blood sample was diluted with uninfected erythrocytes from a healthy individual with a known baseline erythrocyte count, and a 10-fold serial dilution was made to obtain parasitemia ranging from 2 × 104 to 2 × 10−1 parasites/reaction. The genomic DNAs were extracted from the six diluted samples, and analyzed three times in duplicate to construct a PCR standard curve. 2.6. Analytical sensitivity and specificity of real-time PCR assay The minimum detection limit of the real-time PCR assay was evaluated using the same serial diluted blood samples that were used for determining the standard curve. We prepared genomic DNA from 6 samples with parasite concentrations of 2 × 104 to 2 × 10−1 parasites/reaction. The PCR sensitivity assay was performed three times in duplicate. To establish a threshold for real-time PCR, the variation and reproducibility of the threshold cycle (CT ) for each dilution were determined. To estimate the analytical specificity of the assay, DNA was obtained from healthy individuals who have never been infected with malaria (n = 10) and patients diagnosed with candidiasis (n = 10). 2.7. Sensitivity and specificity of clinical specimens The clinical sensitivity and specificity of nested PCR, multiplex PCR and the real-time PCR assay were calculated using blood from 112 malaria patients, including 32 patients with low parasitemia (1–100 parasites/␮l) and from 80 healthy controls. Microscopic examination was used as the reference gold standard for comparison with the results of the three PCR assays. 2.8. Stability tests of the lyophilized PCR tubes To study the effect of its temperature and time on diagnosis and quantitation of parasites, the lyophilized PCR tubes were stored at two different temperatures, and at six time points their activity were evaluated using the real-time PCR assay. For stability comparison of the lyophilized tubes, they were stored under accelerated degradation temperature (40 ◦ C) and control storage

temperature (−20 ◦ C) up to 15 weeks, and then were processed to detect malaria standard DNAs at selected time points. For 10 days, some tubes of accelerated temperature groups were daily withdrawn and amplified in duplicate using a standardized real-time PCR assay as described above. A standard curve was generated using five serial diluted samples with ranging from 1 × 106 to 1 × 102 copies/reaction. In the same ways, we also evaluated their stability under control temperature groups every week up to 15 weeks.

2.9. Statistical analysis The variation of the values of CT pairs obtained for each sample was assessed using Wilcoxon’s test. The significance of the correlation between parasitemia ascertained by microscopy and CT values of the real-time PCR was estimated using Pearson’s correlation coefficient.

3. Results 3.1. Detection of the genus Plasmodium Of 112 clinically suspected cases of malaria, only 80 (71.4%) were diagnosed with malaria based on microscopy, with 62.5% (70/112) determined to be P. vivax and 8.9% (10/112) determined to be P. falciparum. Microscopy failed to detect 32 malaria infections. By contrast, nested PCR and multiplex PCR diagnosed all of the 112 samples as malaria, resulting in 100 samples as P. vivax and 12 samples as P. falciparum.

3.2. Determining the DNA quantification standard curve Each of the two standard curves was linear: the synthetic amplicon, over five log ranges (1 × 106 to 1 × 102 copies/reaction, Fig. 1A), and the genomic DNA, over six log ranges (2 × 104 to 2 × 10−1 parasites/reaction, Fig. 1B). The mean slope and the coefficient of correlation (R2 ) for three duplicate experiments were −0.28 and 0.999 for the synthetic amplicon and −0.30 and 0.998 for the genomic DNA curve. A significant coefficient of correlation was found for the mean CT values and parasite concentration. Consequently, as all curves had slopes close to 100%, both of them were considered to be valid quantification standard curves.

3.3. Analytical sensitivity and specificity of the real-time PCR assay The sensitivity and intra- and inter-assay variations for three independent experiments are shown in Fig. 1C. Positive signals were found for all dilutions except for 0.02 parasite/reaction. Accordingly, the limit of detection was 0.2 parasite/␮l. The mean CT values (standard deviation) ranged from 20.8 (0.5) for 20,000 parasites per PCR volume to 37.3 (0.1) for 0.2 parasite per PCR volume. Of the low-parasitized blood samples, only one gave a CT coefficient of variation >0.2. No statistical differences were found between the CT pairs from each DNA sample (p > 0.05), and all CT values generated from each parasite DNA sample in different experiments were positively associated (p < 0.01). No DNA amplification was observed with real-time PCR of samples obtained from healthy individuals without history of malaria and patients infected with Candida albicans. Based on the signal of internal positive control, no amplification inhibition was observed in the positive sample.

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Fig. 1. The standard curve obtained from our real-time assay in two reproducibility validation experiments. (A) Synthetic amplicon DNA dilutions containing 107 , 106 , 105 , 104 , and 103 copies per PCR reaction, respectively and (B) Plasmodium vivax genomic DNA containing 20,000, 2,000, 200, 20, 2, and 0.2 parasites per PCR reaction were amplified in triplicate. The x-axis plots the threshold cycle value, CT , which indicates the quantity of the target gene at which the fluorescence exceeds a pre-set threshold. The y-axis plots the log of the (A) DNA copy number and (B) parasite count. The plots of the mean CT and DNA input fit linear functions with R2 values of 0.999 (A) and 0.998 (B), respectively. (C) The detection limits and intra- and inter-assay variation of parasite DNA quantification. Results obtained from three independent tests run in duplicate for the real-time PCR assay using TaqMan probe. (D) Correlation between parasite counts using microscopic measurement and the CT of patients who are suspicious malaria infections measured using the real-time PCR assay. There was a significant reciprocal correlation between the parasite count and CT for 80 malaria patients with parasitemia (r= –0.382, p = 0.000). Open circles indicate each patient. The significance was estimated as the Pearson correlation using SPSS (version 14) software.

3.4. Sensitivity, specificity, and quantification of clinical specimens

(r= –0.382, p = 0.000) by the real-time assay, but the correlation was relatively low.

Eighty out of 112 malaria patients were diagnosed as malaria using microscope. The mean parasitemia of them was 2870.82 parasites/␮l (standard deviation, SEM = 2825.74) with a range from 215.52 to 14024.55 parasites/␮l. The mean CT value ± SEM (range) evaluated using the real-time PCR were 27.74 ± 2.65 (23.09–39.50). Meanwhile, 32 patients identified with Gimesa-stained blood smear as negative were positive with the real-time PCR. They showed low parasitemia (1–100 parasites/␮l), and the mean CT ± SEM (range) was 34.8 ± 3.65 (29.16–41.49). Using microscopy as gold standard, the sensitivity and specificity of all three PCR assays were 100% (sensitivity, 80/80 + 0; specificity, 112/112 + 0), respectively. In contrast, using nested PCR assay as the gold standard, the sensitivity was 71.4% (80/80 + 32) with microscopy, and 100% (112/80 + 32) with the multiplex and real-time PCR assays; the specificity was 100% (80/80 + 0) for all techniques. Predictably, the specificity and sensitivity of multiplex and the real-time PCR assays increased when nested PCR assay was used as the gold standard, whereas the sensitivity and specificity of microscopy decreased. Furthermore, Fig. 1D indicates the real-time PCR quantification values in comparision to parasitemia levels of patients. There was a reciprocal correlation between parasitemia as determined by microscopy and the CT values for parasites

3.5. Time comparisons among the three PCR assays and microscopy Compared with all three molecular PCR techniques, microscopy had the shortest hands-on time (0.8 h) for a single sample. From DNA extraction to obtaining a result, nested PCR required the longest time, approximately 10 h. The multiplex PCR and real-time PCR required 4.5 h and 2.5 h, respectively. On the other hand, thermal cycling time of nested PCR, multiplex PCR and real-time PCR required 7 h, 2.5 h and 1.5 h, respectively. 3.6. Stability of the lyophilized PCR tubes We analyzed stability of the lyophilized PCR tubes over time in accelerated or real storage temperature. At 40 ◦ C storage condition, significant activity loss was not detected up to 8 days, but there had been some degradation since 10 days (Fig. 2A). There was little observed loss of activity at −20 ◦ C up to 12 weeks and the activity was slowly decreased since (Fig. 2B). Although there was degradation at higher temperatures, there was little observed degradation after 12 weeks at −20 ◦ C. However, this observed stability is in line

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Fig. 2. Stability of lyophilized materials according to storage times. They were stored under (A) accelerated temperature (40 ◦ C) and (B) real storage temperature (−20 ◦ C) and assayed using real-time PCR at selected time points. (A) At 40 ◦ C degeneration temperature, CT values of serial diluted standard DNA were similar to control group up to 8 days (). (B) At −20 ◦ C real storage temperature, there was little observed loss of activity up to 12 weeks ().

with that observed for the WHO International Standards for P. falciparum DNA nucleic acid amplification techniques (Padley et al., 2008; Fryer and Minor, 2009). Therefore, results from this temperature and period indicate that the lyophilized materials are very stable and reproducible devices for rapid and convenient diagnosis of malaria. 4. Discussion We developed a novel real-time PCR assay (MAL-1111, Bioneer, Korea) for the diagnosis of malaria, and found the assay to be rapid, sensitive, and specific for the detection of malaria parasites in febrile patients as well as asymptomatic individuals. Our realtime PCR assay presented the excellent threshold sensitivity value (0.2 parasite/␮l) (Fig. 1C). The threshold was 50 times more sensitive than thin blood smear examination (10–50 parasites/␮l) (Berry et al., 2005). The new real-time PCR-assay was more sensitive than other methods using 18S ribosomal RNA (rRNA), in conventional PCR assays (Tham et al., 1999) or real-time PCR assays (Perandin et al., 2004; Mangold et al., 2005; Gama et al., 2007). Gama et al. (2007) suggested that owing to the different proportion of schizonts, clinical sample of P. vivax infections with similar parasitemia on thin blood smear would vary in copy number of the rRNA gene and consequently in real-time PCR parasite quantification. In view of this, we evaluated parasite number corrected according to schizont proportion, and then our standard curve generated from P. vivax genomic DNA showed a significant correlation between the mean CT values and parasite density, like other curve generated from synthetic amplicon (Fig. 1A and

B). However, in clinical samples, a significant, but low correlation (r= −0.382, p = 0.000) between parasitemia, as determined by microscopy, and parasite number, as calculated from CT values of the standard curve, are somewhat confounded phenomenon (Fig. 1D). It is suggested that the parasitemia of the clinical samples evaluated by microscopy may not reflect correct proportion of schizonts. In general, PCR assays are more sensitive than microscopic examination for malaria diagnosis (Hänscheid and Grobusch, 2002; Kain et al., 1993). Using the real-time PCR assay, malaria was correctly diagnosed in 32 clinical samples that had been determined to be negative by microscopy. Accordingly, our real-time PCR assay may be useful for diagnosing low-parasitized individuals or asymptomatic malaria patients in endemic regions. The procedures of conventional PCR assays including nested PCR are relatively labor intensive, time-consuming, and the results of them are unquantifiable (Farcas et al., 2004). Our new real-time PCR assay requires approximately 1.5 h to generate quantitative results. Of the three PCR assays tested, the real-time PCR is the least time consuming method and gives the most sensitive detection limit of the three PCR assays. In addition to high sensitivity, and time-saving properties, realtime PCR assay has more merits. Real-time based assay can reduce post-amplification DNA contamination among samples and the results are not technician dependent (Gama et al., 2007). The assay des not need for hazardous, time-consuming gel electrophoresis (Boonma et al., 2007). Finally, considering sensitivity and time, we are tempted to conclude that the real-time PCR may be used instead of other PCR assays regardless of its higher cost.

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Real-time PCR assay format developed in this study is a ready to use lyophilized format with extended storage stability and enhanced reproducibility. All reagents required for PCR reaction including DNA polymerase, target-specific primers & probe, internal positive control-specific primers & probe, dNTPs, buffers, and stabilizers were mixed, aliquot into real-time PCR tubes and lyophilized. The assay format can reduce the possibility of humanerror handling because user add only DNA extracted from sample and PCR-grade water in each tube without any pre-preparation of master mix solution. Internal control DNA was applied to real-time PCR step for the confirmation of proper PCR reaction, and thus can give reliable interpretation. This assay format may be beneficial in endemic areas that do not established good medical environments. An ideal preparation should be stable for many years to sever as an international standard (WHO, 2004). The stability study on the lyophilized materials was performed to predict the long term stability at various temperatures. Their degradation was very little observed at lower temperatures (−20 ◦ C), whereas unstability was observed in the samples stored at 40 ◦ C since 8 days (Fig. 2). This may be due to natural degradation expected at this temperature, but may also be the result of increasing difficulty in resuspending samples stored at elevated temperature over time. These results suggest that this preparation has adequate stability for the long term use of the material. Moreover, it is suggested that elevated temperature for a relatively short time such as during shipment of the products should not cause unacceptable loss of potency. In summary, compared with nested and multiplex PCR assays and microscopy, our real-time PCR assay is an equally rapid, sensitive, specific, but a more rapid quantitative method for detecting malaria patients. This new real-time PCR assay may be useful for the primary identification of infected patients without microscopic analysis. In addition, it may be valuable for the identification of malaria cases with low parasitemia and for epidemiological studies that require high-throughput analyses. This assay may have potential applications in detecting malaria parasites in asymptomatic infections; evaluating candidate malaria vaccines; screening blood donors, especially in endemic areas, and monitoring malaria treatment. Acknowledgments This work was financially supported by Bioneer Corporation, Daejeon, South Korea. W.L.K., S.Y.K., H.J.P., and H.O.P. are employed by Bioneer Corporation. We thank Yee-Gyung Kwak and Dr. Jin-Ho Chun for their important contributions. References Amexo, M., Tolhurst, R., Barnish, G., Bates, I., 2004. Malaria misdiagnosis: effects on the poor and vulnerable. Lancet 364, 1896–1898.

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