• View in gallery

    Plasmodium vivax and Plasmodium falciparum infected children from June 2009 to June 2010. Only six P. vivax infected children were identified on June 2011. This figure appears in color at www.ajtmh.org.

  • View in gallery

    The first time the children were infected with Plasmodium vivax from Figure 1. In total, there were 25 P. vivax infections in the 300 children. The date in the table shows the first-time P. vivax infection was observed in a child. The color coding marks each collection period and is used in the map in Figure 4. This figure appears in color at www.ajtmh.org.

  • View in gallery

    Plasmodium vivax infections in children from Bandiagara. The blood smears containing P. vivax parasites from two children in Bandiagara, Mali. Images of different stages of blood stage P. vivax parasites are shown from (A) child 44 and (B) child 32. Bars indicate 5 µm. This figure appears in color at www.ajtmh.org.

  • View in gallery

    Plasmodium vivax infected children’s locations in the Bandiagara Map. The figure shows the map of Bandiagara and the locations of each patient with P. vivax infections. The unique colors to the children are based on the first time a child had a P. vivax infection during the specific collection dates as shown in Figure 2. The river Yame passes through Bandiagara which is colored in blue as well as the water ponds. This figure appears in color at www.ajtmh.org.

  • View in gallery

    Plasmodium vivax and anemia in Malian children. The table shows the data of hemoglobin concentration in 25 P. vivax infected children. The data points highlighted in bold are the children only infected by P. vivax. The hemoglobin concentration was markedly reduced during P. vivax infection and returned to normal within 30–60 days. This figure appears in color at www.ajtmh.org.

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Plasmodium vivax Infections over 3 Years in Duffy Blood Group Negative Malians in Bandiagara, Mali

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  • 1 Malaria Research and Training Center, International Center for Excellence in Research, University of Sciences, Techniques and Technology of Bamako, Bamako, Mali;
  • | 2 Laboratory of Malaria and Vector Research, National Institutes of Allergy and Infectious Diseases, National Institutes of Health, Rockville, Maryland;
  • | 3 Division of Malaria Research, Institute for Global Health, University of Maryland School of Medicine, Baltimore, Maryland

Plasmodium vivax was thought to infect only the erythrocytes of Duffy blood group positive people. In the last decade, P. vivax has appeared throughout Africa, both in areas where Duffy positive and negative people live side by side as in Madagascar and Ethiopia and in areas where people are primarily Duffy negative, such as in western Kenya. We performed quantitative polymerase chain reaction on blood samples dried onto filter paper to determine the prevalence of P. vivax and Plasmodium falciparum in a cohort of 300 children (newborn to 6 years of age) in Bandiagara, a Sahelian area of Mali, west Africa, where the people are Duffy negative. We report 1–3 occurrences of P. vivax in each of 25 Duffy-negative children at six time points over two rainy seasons and the beginning of the third season. The prevalence of P. vivax infection was 2.0–2.5% at every time point (June 2009 to June 2010). All children with P. vivax infections were asymptomatic and afebrile, and parasite densities were extremely low. Anemia, however, was the main burden of infection. Plasmodium vivax could become a burden to sub-Saharan Africa, and the evidence of P. vivax existence needs to be taken into consideration in designing malaria control and elimination strategies in Africa.

INTRODUCTION

In the 1970s, Duffy blood group negative people were shown to be resistant to Plasmodium vivax infection in studies conducted on U.S. and Honduran volunteers and American soldiers.13 From these studies, it was assumed that all Duffy-negative people were resistant to P. vivax infection. In Madagascar, Ethiopia, and Sudan where Duffy-positive and Duffy-negative people live side by side, infections of P. vivax were found and assumed to be the result of passage back and forth between Duffy positive and negative individuals, with the Duffy-positive population possibly maintaining the infection.46 The P. vivax infections in Duffy-negative people showed lower parasite density and milder symptoms compared with P. vivax infections in Duffy-positive people, suggesting a less efficient infection.4,5 However, P. vivax infections were also found in Duffy blood group negative people in a predominantly Duffy-negative area of western Kenya.7 Menard and others4 in Madagascar and Gunalan and others8 in Ethiopia reported a large expansion of the gene-encoding Duffy-binding protein 1 (DBP1) in the P. vivax populations, especially in two patients in Ethiopia who were infected with P. vivax that had eight and three copies of the gene.8 A recent study in southeast Asia found two to three copies of DBP1 in parasites in Duffy-positive populations9; the reason for the increase in DBP1 gene copy number in southeast Asia is unknown. However, this finding suggests the possibility that the P. vivax parasites that are able to infect Duffy-negative erythrocytes may have been selected for their ability to use multiple copies of DBP1 to invade Duffy-negative erythrocytes through an alternative receptor.

Plasmodium vivax has now been reported from all over Africa, Angola,10 Benin,11 Cameroon,12,13 Ethiopia,5 Equatorial Guinea,10 Madagascar,4 Mauritania,14 Sudan,6 and Mali,15 raising the possibility that P. vivax was transmitted between Duffy-negative people as was first described in western Kenya.7 In Mali, P. vivax has been found in the northern part of the country. In 1991, Doumbo and others described a case of P. vivax in the region of Kidal,16 and subsequently a prevalence of 10–30% of P. vivax in febrile patients was reported in northern regions of Mali.17,18 It was possible that P. vivax was transmitted from Duffy-positive people in the north Sahelian region to Duffy-negative people in the Bandiagara region. However, it is also possible that P. vivax was transmitted between Duffy-negative individuals, as is the case in western Kenya.

We searched for the presence of P. vivax in a cohort of 300 children (newborn to 6 years of age) in Bandiagara, a Duffy-negative area over a 3-year period. We identified 25 P. vivax infections in children who were all Duffy blood group negative determined by molecular techniques. This suggests that P. vivax in Duffy-negative people is widespread in Mali and perhaps in all of Africa.

MATERIALS AND METHODS

Study site and ethical clearance.

The study was conducted in Bandiagara, a town located in the central region of Mali and approximately 700 km from the capital city, Bamako. The Yame River, a tributary of the Niger River, passes through the town and constitutes an important breeding site for the malaria vector mosquitoes Anopheles gambiae and Anopheles funestus during 5 months of transmission each year. The landscape is of the Sahelian type on a rocky plateau. The population of Bandiagara is around 13,364 and the economic activity is mainly agropastoral with some tourism.19 The climate is characterized by a rainy season from June to October, with an average rainfall of 600 mm/year and a longer dry season from November to May. Malaria transmission is seasonal with transmission from June or July to November or December. The peak incidence of clinical malaria is observed in August–September with more than 60 infected bites per person per month.19 The incidence of clinical malaria was 1.7 episodes per transmission season in children less than 10 years of age, and P. falciparum was the most frequent species with 97% of the malaria infection, Plasmodium malariae was 3% and Plasmodium ovale was rarely identified by microscopy, while P. vivax was not detected.19,20

The study was reviewed and approved by the institutional review board of the University of Maryland, Baltimore, MD, and the Institutional Ethical Review Committee of the Faculty of Medicine, Pharmacy and Odonto-Stomatology at the University of Sciences, Techniques and Technologies of Bamako, Bamako, Mali. The study protocol and risks and benefits of participation were explained in local languages to the population and community permission was obtained. Individual written informed consent was obtained from the parents or legal guardians of children before inclusion in the study. An identification code was assigned to each participant and personal identity was kept confidential.

Population and study design.

As part of a prospective cohort study of malaria incidence in Bandiagara, blood samples were collected during monthly scheduled visits from 300 children aged < 1 to 6 years. Blood was collected using Whatman 3MM chromatography filter paper (GE Healthcare, Buckinghamshire, UK). Dried blood spots were sealed in individual plastic bags with desiccant and kept in a dry area at room temperature. Plasmodium vivax sampling was done at time points in June, September, and December 2009, February and June 2010, and June 2011. Prevalence at each time point (except June 2011) was calculated based on the number of positive infections among the total samples analyzed.

Blood smear preparation.

Thick smears were performed on blood samples of each participant for active surveillance of malaria regardless of the fever or other malaria symptoms. The smears were air-dried and not fixed with methanol. The thick smears were stained using 5% Giemsa solution for 45 minutes. The number of parasites in fields containing 1,000 leukocytes was determined. The parasite count was determined assuming 8,000 leukocytes/µL.

Molecular detection of P. falciparum and P. vivax infection.

Molecular screening of P. falciparum and P. vivax was carried out using genomic DNA extracted from dried blood spot filter papers of 300 samples over five time points (June, September, December 2009, February, June 2010, and June 2011) from the beginning of one transmission season every 2–3 months to the beginning of the second season and for only P. vivax in the beginning of the third season (June 2011). DNA isolation was performed using QIAamp 96 DNA Blood Kit (Qiagen, Hilden, Germany) according to the manufacturer’s instructions. Plasmodium parasite DNA was qualitatively detected using a quantitative real-time polymerase chain reaction (qPCR) assay (LightCycler® 480, Roche, Basel, Switzerland) based on primers targeting the 18S small subunit ribosomal RNA (18S rRNA) gene as described.21 A two-plex P. falciparum and P. vivax PCR was performed to determine the presences of these two species in the children’s specimens. The assay consists of a combination of two different forward primers, two different TaqMan probes each labeled with a different fluorophore, and one reverse primer that amplifies and detects varied 18S rRNA regions specific to either P. falciparum or P. vivax. Detection of the appropriate fluorophore will indicate the presence of P. falciparum and P. vivax. The PCR amplification was performed using the following oligonucleotide primers and probes sequences: Forward P. falciparum primer: 5′-CCGACTAGGTGTTGGATGAAAGTGTTAA-3′; Forward P. vivax primer: 5′-CCGACTAGGCTTTGGATGAAAGATTTTA-3′; Reverse Plasmodium primer: 5′-AACCCAAAGACTTTGATTTCTCATAA-3′; TaqMan P. falciparum probe: 5′-(Cy5)-AGCAATCTAAAAGTCACCTCGAAAGATGACT-(BHQ-2)-3′; TaqMan P. vivax probe: 5′-(TAMRA)-AGCAATCTAAGAATAAACTCCGAAGAGAAAATTCT-(BHQ-2)-3′.21

The PCR amplification was performed using 5 µL of template DNA added to 20 µL of reaction master mix consisting of 0.5 µL of each individual primer at 45 µM, 0.5 µL of each probe at 0.25 µM, and 12.5 µL of 2× concentration of TaqMan master mix (Applied Biosystem, Foster City, CA). A final reaction volume of 25 µL was run with the following thermal cycling conditions: 50°C for 2 minutes followed by 95°C for 10 minutes and 40 cycles of 95°C for 15 seconds and 60°C for 1 minute. Plasmodium falciparum 3D7 DNA and P. vivax DNA were used as positive controls for each species and the PCR reaction master mix with no template DNA was used as a negative control.

Duffy blood group genotyping.

The absence of Duffy antigen receptor on the surface of the erythrocytes is due to the inability of the GATA1 transcription factor to bind to the upstream promotor region of Duffy blood group antigen, which is due to the mutation at 33rd nucleotide position upstream of the transcription initiation site. Hence, no Duffy blood group antigen is expressed on the surface of the erythrocyte.22 In this study, the Duffy blood group genotyping was performed using PCR amplification of the human Duffy antigen/chemokine receptor gene followed by sequencing. The primary PCR amplification (Nest 1) of a fragment of 997 basepair (bp) was performed using primer pairs (ExtForward 5′-GTGGGGTAAGGCTTCCTGAT-3′ and ExtReverse 5′ CAGAGCTGCGAGTGCTACCT-3′) in a reaction mixture of 26 µL containing 22.5 µL of Platinum PCR SuperMix (Invitrogen, Carlsbad, CA), 1 µL of each forward and reverse primer at 10 µM concentration, and 1.5 µL of DNA template as described.4 Nest 2 PCR amplification was performed using primer pairs (ExtForward 5′-GTGGGGTAAGGCTTCCTGAT-3′ and InterReverse 5′-CAAACAGCAGGGGAAATGAG-3) to determine a single-nucleotide polymorphism in the Duffy gene (223 bp fragment) which is the GATA-1 transcription factor–binding site in the promoter region that governs the erythrocyte Duffy negativity. Reaction mixture for PCR amplifications included 22.5 µL of Platinum PCR SuperMix (Invitrogen, Carlsbad, CA), 1 µL of each forward and reverse primer for each primer at 10 µM concentration, and 1.5 µL of primary PCR product in a final volume of 26 µL. Two known Duffy-positive controls were used.

PCR products were purified by filtration using MultiScreen PCR Plates for DNA cleanup (Merck Millipore, Billerica, MA). Sequencing reactions were performed for each strand using the ABI PRISM BigDye Terminator cycle sequencing ready-to-use reaction kit run on a 3730XL automatic sequencer (Applied Biosystems, Foster City, CA). Electrophoregrams were visualized and analyzed with Sequencher® version 5.4.1 sequence analysis software (Gene Codes Corporation, Ann Arbor, MI).

RESULTS

We searched for P. vivax infections in samples collected at six time points over 2 years among 300 children in Bandiagara. We identified a total of 25 P. vivax (Figures 1 and 2) and 109 P. falciparum (Figure 1) first-time infections in each individual in the cohort while the samples collected at the beginning of the third transmission season (June 2011) were not tested for P. falciparum. The prevalence of P. vivax was between 2.0% and 2.5% (Figure 1). The prevalence of P. falciparum was between 11% and 21% during the same periods (Figure 1). One child was infected with both P. falciparum and P. vivax (Figure 1). All cases of P. vivax were asymptomatic. None had fever defined by axillary temperatures equal or above 37.5°C. Of P. vivax cases (Figure 1), two had 37.0°C and one had 37.4°C; the rest had below 37.0°C. None of the children had a history of fever and chills, headache, or muscle ache, symptoms that may be associated with malaria. Three children had respiratory symptoms and two had rhinorrhea or nasal congestion. The children were not treated for P. vivax as they did not have fever and no parasites were seen on the smear.

Figure 1.
Figure 1.

Plasmodium vivax and Plasmodium falciparum infected children from June 2009 to June 2010. Only six P. vivax infected children were identified on June 2011. This figure appears in color at www.ajtmh.org.

Citation: The American Journal of Tropical Medicine and Hygiene 97, 3; 10.4269/ajtmh.17-0254

Figure 2.
Figure 2.

The first time the children were infected with Plasmodium vivax from Figure 1. In total, there were 25 P. vivax infections in the 300 children. The date in the table shows the first-time P. vivax infection was observed in a child. The color coding marks each collection period and is used in the map in Figure 4. This figure appears in color at www.ajtmh.org.

Citation: The American Journal of Tropical Medicine and Hygiene 97, 3; 10.4269/ajtmh.17-0254

The thick smears of all the P. vivax PCR-positive children were read as negative in the field. Slides from 24 of the 25 children who were PCR positive for P. vivax were re-read by two experts. One slide was PCR positive for both P. vivax and P. falciparum and was excluded because of the problem of identifying the difference between P. falciparum and P. vivax rings on a thick smear. Thirteen of the 23 slides were read as negative by two readers. Of the remaining 10 slides, the parasite density was between eight and 157 parasites/µL (88.7 ± 53.4 parasites/µL), calculated from the number of white blood cells assuming 8,000 white blood cells/µL. Plasmodium vivax parasites from the thick blood smears are shown in Figure 3. These results indicate an extremely low parasite density, indicating the importance of molecular tests for P. vivax infections. The location of the children infected with P. vivax was spread throughout the area of study (Figure 4).

Figure 3.
Figure 3.

Plasmodium vivax infections in children from Bandiagara. The blood smears containing P. vivax parasites from two children in Bandiagara, Mali. Images of different stages of blood stage P. vivax parasites are shown from (A) child 44 and (B) child 32. Bars indicate 5 µm. This figure appears in color at www.ajtmh.org.

Citation: The American Journal of Tropical Medicine and Hygiene 97, 3; 10.4269/ajtmh.17-0254

Figure 4.
Figure 4.

Plasmodium vivax infected children’s locations in the Bandiagara Map. The figure shows the map of Bandiagara and the locations of each patient with P. vivax infections. The unique colors to the children are based on the first time a child had a P. vivax infection during the specific collection dates as shown in Figure 2. The river Yame passes through Bandiagara which is colored in blue as well as the water ponds. This figure appears in color at www.ajtmh.org.

Citation: The American Journal of Tropical Medicine and Hygiene 97, 3; 10.4269/ajtmh.17-0254

The hemoglobin (Hb) concentration for all the children was measured during each visit. As anemia is associated with malaria, we looked for P. vivax-associated anemia in these children (Figure 5). Children (32, 33, 34, 38, 59, 66, and 94) highlighted in red and bold were infected only with P. vivax and observed to have decrease in Hb concentration to the anemic level. Child 32 had 10.7 g/dL Hb when infected with P. vivax and the concentration decreased to 9.6 g/dL a month later and started to increase to 11.9 g/dL in around 50 days and no P. vivax and P. falciparum was found in subsequent time points. Others such as child 33 had a marked reduction from 9.6 to 7.7 g/dL the month after P. vivax infection. We do not know the cause of the decrease of Hb to 7.7 g/dL, although untreated P. vivax could have resulted in the anemia. Another marked fall in Hb in child 94 was from 13.1 during P. vivax infection to 9.4 g/dL that decreased a month after P. vivax and returned to 13.2 a month later. Again, as the P. vivax was untreated, the anemia may have been caused by P. vivax. A similar effect was also observed in children 34, 38, and 66.

Figure 5.
Figure 5.

Plasmodium vivax and anemia in Malian children. The table shows the data of hemoglobin concentration in 25 P. vivax infected children. The data points highlighted in bold are the children only infected by P. vivax. The hemoglobin concentration was markedly reduced during P. vivax infection and returned to normal within 30–60 days. This figure appears in color at www.ajtmh.org.

Citation: The American Journal of Tropical Medicine and Hygiene 97, 3; 10.4269/ajtmh.17-0254

All P. vivax were found in Duffy blood group negative children with the mutation from T to C in the GATA1-binding area 5′ to the open reading frame.22 The location of the children in Bandiagara during each study period is shown in color related to time of their first infection (Figure 4). The 2–3 episodes of P. vivax infection occurrence in some of the Duffy-negative children suggest a possible reinfection (new infection) by mosquito bites with a different P. vivax strain or due to relapse from hypnozoites in the liver. The children 32/33 and 58/59 were siblings living in the same house and each pair was infected during the same study period (Figures 2 and 4), suggesting that the infection occurred within their respective houses. Infections occurred in most areas where the study children lived. In Bandiagara, the rainy season starts in May and continues through October. The six infections in February 2010 (in yellow, Figure 1) were unusual in that they occurred during the dry season, which could be due to relapse. However, during the dry season, the Yame River that runs through Bandiagara dries up, but the local people block the river for water to grow crops such as tomatoes and onions. We know that P. falciparum infection persists at low incidence during the dry season in Bandiagara.20 We speculate that P. vivax transmission may have continued through February by mosquitoes breeding in the dammed river.

DISCUSSION

We describe here a longitudinal study of P. vivax infection in a Duffy-negative population of 300 children spanning across three rainy seasons in Bandiagara. At every time point except June 2011, P. vivax was detected in about 2% of the children, and over the course of the study 8.3% of the children experienced P. vivax infection at least once. The prevalence data highlight that P. vivax infection occurs throughout the year irrespective of the rainy season, further suggesting a chronic low level infection or relapses. Approximately 36% of the children were infected with P. falciparum during the study period. It has been shown that there is an association between P. vivax relapse after P. falciparum infection and treatment.2325 Although none of the P. vivax infected children were symptomatic, however, 10 of 25 children were treated with the malaria drug at different time points during June 2009 to June 2012. Of the 10 children, only three children were treated with malaria drug before P. vivax infection (Supplemental Figure 1), the other seven were treated with malarial drugs after P. vivax infection. Hence, from this observation we cannot conclude whether P. falciparum infection and its treatment with malarial drugs would have led to triggering of P. vivax hypnozoites.

Ten of the 25 children who experienced P. vivax were infected more than once (range 2–3 infections), sometimes at consecutive time points, and sometimes separated by the absence of detectable P. vivax. The P. vivax-infected children were observed in different parts of the Bandiagara study area (Figure 4). Two siblings in two families were infected with P. vivax during the same study period, indicating that the infections were occurring in their home. All infections were seen in Duffy-negative children and not in Duffy positive, indicating that the P. vivax infections were similar to what was observed in western Kenya7 but different from those in Madagascar4,26 and Ethiopia5,8 where Duffy-positive P. vivax infections can serve as a source of infection for the Duffy blood group negative people.

What was the source of P. vivax in Bandiagara? It is possible that P. vivax could be introduced from the Sahara desert in the north where the Tuareg people may be Duffy positive,17,18 but the continuous infection for 3 years raises the possibility that P. vivax infection is maintained by infections from Duffy-negative children. The other possibility is tourists visiting Bandiagara who are Duffy positive could interact with Duffy negatives, but the area of the study is not near the tourist area.

The first evidence that P. vivax could exist in a Duffy blood group negative population was in western Kenya in 20067 where the evidence for Duffy negative was defined by the negativity with anti-Fy6 and anti-Fy3. In other areas such as in Cameroon, two studies found P. vivax in Duffy-negative people, presumably in different areas.12,13 These Duffy negative infections were identified by the mutation of T to C in the GATA1-binding domain 5′ to the open reading frame.22 Other areas of Africa had P. vivax, but the Duffy type was not determined.27,28

It is likely that P. vivax selected for the refractory Duffy-negative phenotype in Africa. The selection is accelerated by the protection of the heterozygote29 and may have occurred in 500 generations over 10,000 years, depending on the selective advantage of the homozygous null and partial dominance of the heterozygote. Anemia due to P. vivax has been well documented.3032 Our data in this study suggest that low-level P. vivax parasite density may have caused anemia in the children. Thus, the diagnosis and the treatment of P. vivax might benefit the children in Bandiagara.

Plasmodium vivax varies greatly in virulence with the Madagascar P. vivax isolate studied by James and others33 associated with marked virulence and mortality of 10–14% if left untreated. It has been presumed that P. vivax was responsible for the mortality in the Thames estuary over many centuries, preventing people from working in the zone.34 Today various reports describe the severity of P. vivax.30,35,36 Plasmodium vivax can cause severe disease and selected for the loss of the erythrocyte receptor, the Duffy blood group, to prevent the infection with P. vivax.

In recent years, with advanced molecular and cellular techniques, including conventional PCR, real-time PCR, and flow cytometry, we are able to detect P. vivax infection with little or no clinical symptoms, which could have gone unnoticed and undetected for an unknown period in Duffy-negative individuals. This further suggests that P. vivax is maintained in the Duffy-negative population in Bandiagara without any malarial disease (fever and chills). In addition, it is not clear whether the observed persistent infections in Bandiagara are due to new infections or relapses due to reactivation of dormant liver stage hypnozoites. It was shown that 70% and ∼96% of P. vivax infections in Papua New Guinea (PNG) and Thailand, respectively, are due to reactivation of hypnozoites, and this difference in PNG and Thailand is due to primaquine treatment failure in PNG.37

We are now faced with an enigma. In the 1970s, Duffy-negative people were shown to be refractory to P. vivax infection. The present data indicate that at some time in the past, P. vivax adapted to infect Duffy-negative people throughout Africa. The mutations in P. vivax that permitted infection may be the expansion of the DBP locus8 or other unknown mutations that may upregulate or change invasion ligands on merozoites that permit invasion.3840 Studies from Ethiopia and Madagascar show that the infection with P. vivax results in a mild disease. With continued mutations, will P. vivax become a more virulent infection in Duffy-negative populations as seen in Duffy-positive populations? For example, the Duffy blood group antigen in Saimiri and Aotus monkey erythrocytes are highly similar. However, the DBP1 of P. vivax Salvador I does not bind Saimiri monkey erythrocytes, although it binds to the erythrocytes of Aotus monkey. Yet the infection of Saimiri and Aotus monkeys is similar to the Salvador I P. vivax infection.8,41,42 As we learn more about P. vivax in Africa, the mutations in ligands that affect specificity or expression levels in the parasite that have allowed its spread throughout Africa today will be defined.

Supplementary Material

Acknowledgments:

We thank the BMP clinical research team, specifically, Karim Traore Ahmadou Dembele and Boureima Guindo for clinical support; Youssouf Tolo, Issa Diarra, Modibo Daou, Mody Sissoko, Bourema Kouriba, Agnès Guindo, and MRTC CLIA lab team for Clinical lab support; Ibrahima Kebe, Chebou Diallo, and Ismael Thera for data management; Bouréma Ouologuem, Abdoulaye Ouologuem, Tabanidjou Karembé, Boucari Djiguiba, Akouni Dougnon, Sadio Dolo, Domo Ouologuem, Issa Tapily, and Paul Dougnon for fieldwork; Danzélé Coulibaly, Moctar Traoré, Sékouba Mariko, Amadou Arama, Nicole Eddington, and Malathi Vadla for administrative and technical support; Aboubacar Sidiki Kouyaté, the chief medical officer of the Bandiagara District Hospital, for support; Karen Ball for regulatory support; and the community of Bandiagara, Mali for their dedication. The Malaria Incidence Study was conducted at the Bandiagara Malaria Project (BMP) Clinical Research Center, an ANDI Center of Excellence for Clinical Development of Malaria Products. We also thank Susan K. Pierce, NIH, for the constructive suggestions and critical reading of the manuscript, Roberto Moraes Barros and Joe Brzostowskij, NIH, for help in microscopy. Jianbing Mu, NIH, for the help in arranging the data table.

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Author Notes

Address correspondence to Louis H. Miller, Laboratory of Malaria and Vector Research, National Institutes of Allergy and Infectious Diseases, National Institutes of Health, 12735 Twinbrook Parkway Rockville, MD 20852, E-mail: lmiller@niaid.nih.gov or Ogobara K. Doumbo, Malaria Research and Training Center, International Center for Excellence in Research, University of Sciences, Techniques and Technology of Bamako, BP 1805, Bamako, Mali, E-mail: okd@icermali.org.

These authors contributed equally to this work.

Financial support: This work was supported by the cooperative agreement 5U01AI065683 from the National Institute of Allergy and Infectious Diseases; and the grant D43TW001589 from the Fogarty International Center, National Institutes of Health, Howard Hughes Medical Institute and the Intramural Research Program of the Division of Intramural Research, National Institute of Allergy and Infectious Diseases, National Institutes of Health.

Authors’ addresses: Amadou Niangaly, Amed Ouattara, Drissa Coulibaly, Abdoulaye K. Kone, Mahamadou A. Thera, and Ogobara K. Doumbo, Malaria Research and Training Center, International Center for Excellence in Research, University of Sciences, Techniques and Technology of Bamako, Bamako, Mali, E-mails: niangaly@icermali.org, amouattara@som.umaryland.edu, coulibalyd@icermali.org, fankone@icermali.org, mthera@icermali.org, and okd@icermali.org. Karthigayan Gunalan, Juliana M. Sá, and Louis H. Miller, Laboratory of Malaria and Vector Research and National Institutes of Allergy and Infectious Diseases, National Institutes of Health, Rockville, MD, E-mails: karthigayan.gunalan@nih.gov, jsa@niaid.nih.gov, and lmiller@niaid.nih.gov. Amed Ouattara, Matthew Adams, Mark A. Travassos, Jennifer Ferrero, Matthew B. Laurens, and Christopher V. Plowe, Division of Malaria Research, Institute for Global Health, University of Maryland School of Medicine, Baltimore, MD, E-mails: amouattara@som.umaryland.edu, madams@som.umaryland.edu, mtravass@som.umaryland.edu, jferrero@thellf.org, mlaurens@som.umaryland.edu, and cplowe@som.umaryland.edu.

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