Open access peer-reviewed chapter

Increased Trends of P. vivax in Sub-Saharan Africa: What Does it Mean for Malaria Elimination?

Written By

Mary Aigbiremo Oboh, Mamadou Ndiath, Olumide Ajibola, Kolapo Oyebola and Alfred Amambua-Ngwa

Submitted: October 13th, 2020 Reviewed: March 12th, 2021 Published: April 14th, 2021

DOI: 10.5772/intechopen.97189

Chapter metrics overview

275 Chapter Downloads

View Full Metrics


Plasmodium vivax being the most geographically spread Plasmodium species is considered sparsely distributed in sub-Saharan Africa (sSA) while P. falciparum is the most prevalent species in this region. Thus, control strategies in sSA have been disproportionately targeted towards falciparum malaria. Nevertheless, with the use of more sensitive malaria diagnostic platforms, there are more reports of P. vivax and other non-falciparum malaria in sSA. In addition, P. vivax is presumed benign, however there are new findings of severe cases recorded from P. vivax single or mixed infection with other Plasmodium species. Besides, the extended dormant period (lasting for weeks or months) is a challenge for achieving effective cure for vivax infections. Although, chloroquine has been proscribed for treatment P. falciparum, it still remains the drug of choice for P. vivax in most Asian countries where it is predominant. In sSA, artemisinin combination-based therapies (ACTs) are used for treatment of falciparum malaria and, it is probable that the use of ACT could be enhancing adaptive selection for P. vivax in the face of its increasing prevalence in the population. Hence, understanding epidemiological and biological factors, and data that could be contributing to the observed steady increase in P. vivax prevalence in sSA is important. In this chapter, we discuss the mechanisms for invasion of red blood cells, trends in increasing prevalence of vivax malaria, diagnostic tools, and the public health implications of P. vivax and P. falciparum co-endemicity in Africa.


  • Plasmodium vivax
  • Plasmodium falciparum
  • sub-Saharan African
  • Malaria elimination
  • Duffy negative antigen

1. Introduction

In sub-Saharan Africa (sSA), Plasmodium falciparum is responsible for most of the human malaria infection and less frequently, malaria could be due to P. ovale, P. malariae and P. vivax [1]. Significant gains observed in massive reduction in global malaria cases and mortalities in the early 2000s have been largely due to strategic malaria interventions [1]. Nevertheless, signals of reversal in gains are beginning to emerge in most African countries [1]. This stall in progress has been attributed to increased prevalence of non-falciparum malaria parasites on one hand and the development of resistance by both the parasite and vector to drugs and insecticides respectively [2, 3, 4]. Thus, refocusing control strategies to include all circulating species of Plasmodium in sSA aimed at reducing the burden of these non-falciparum parasites will greatly contribute to malaria control progress.

P. vivax is particularly interesting as regards its mechanism of invasion, reproduction, virulence and development. Understanding the biology of vivax parasites is crucial for developing strategies aimed at controlling malaria. Regrettably, vivax malaria poses a huge challenge for in vitro analysis to adapt in cell culture, in order to shed more light on invasion patterns, virulence, growth, and reproduction. For example, P. vivax has high predisposition for immature red blood cells-RBCs (reticulocytes) which is not readily available [5]. Another drawback as regards vivax control is the use of primaquine (PQ) to achieve hypnozoites clearance, which causes haemolytic anaemia in glucose 6 phosphate dehydrogenase (G6PD) deficient individuals [6, 7]. In sSA with very limited resources for ascertaining the G6PD status of individuals, this presents a practical challenge as individuals are either treated without their G6PD status known or remain untreated. The implication of the latter is the prolonged carriage of hypnozoites capable of relapsing weeks or months later. It has also been found that individuals with specific cytochrome P450 gene polymorphic alleles such as the CYP2D6 do not efficiently metabolise PQ and eventually miss out in the clearance of dormant liver parasites [8].

Overcoming vivax resistance to chloroquine [9], which is also a major drawback in vivax control is looking promising, with a single dose of tafenoquine seemingly independent of CYP2D6-impairement is effective in the prevention of vivax relapse [10, 11].

Evolution is shaping the biological interaction between vivax parasites and the human host [12, 13]. For instance, there is new evidence counteracting the former notion of P. vivax emergence in Asia from vivax-like macaque parasites [14, 15]. Recent evolutionary genomic analyses have postulated that vivax most probably originated in Africa [16]. Selection for Duffy-negative mutations could have been due to the high circulation of P. vivax in ancient Africans [12] which led to the eventual disappearance of vivax malaria in most parts of Africa [17]. There is need to carry out a comprehensive characterisation of the Duffy-independent vivax strains in West [18, 19] and East Africa [20] in order to understand the genetic similarities and phylogeny between the strains and, relationship with emerging zoonotic non-human primate (NHP) malaria.

Although, non-human primates (NHPs) and human vivax species are considered non-interbreedable (allopatric) [12], it is possible that recombination events between vivax parasites of apes and human lineages have occurred to favour the adaptation of currently circulating vivax strains [21]. This hypothesis is being supported by binding studies of recombinant orthologues; reticulocyte binding protein 2- RBP2 and RBP3, to gorillas, humans and chimpanzees erythrocytes’ that displayed no evidence of host specificity to invasions of red cells [12]. Moreover, there has been detection of ape-like P. vivax in a European returnee from Africa [21], while human P. vivax has been identified in monkeys from the wild in South America [22]. To infer genetic epidemiology of P. vivax and relationship with other closely related species, population genetic studies is necessary. This has been employed in P. falciparum to demonstrate the origin, complexity, evolutionary history, changes in population size and relatedness between parasite populations.


2. Mechanism of invasion of P. vivax malaria to Duffy antigen receptor on red blood cells

Duffy antigen receptor for chemokines (DARC) gene is the fourth erythrocyte gene associated with resistance to Plasmodium species [23]. Polymorphisms in other genes associated with malaria are specifically accompanied by severity or protection against falciparum malaria. DARC confers protection against vivax malaria [24]. DARC has a 36–46 kDa glycoprotein major sub-unit and is made up of 338 amino acids (AAs) that is organised into seven transmembrane domain [25] with an extracellular epitope at the N-terminal that mediates P. vivax invasion of RBCs. DARC, a non-specific receptor is expressed on erythrocytes and employed by P. vivax merozoites in invasion of RBCs [26]. It is located on the long arm of chromosome 1 (1.q22–1.q23) and possesses two exons [27]. A mutation from a thymine to a cytosine at the GATA-box upstream of the promoter region (33rd position) results in the non-expression of DARC on RBCs, giving rise to the FYO* allele also known as the Duffy negative phenotype [28].

The FYO* allele is prevalent in sSA as well as in African-Americans but rare in other racial groups lacking black admixtures [27]. For instance, all 1000 individuals screened in a study in the Gambia were Duffy negative [29]; likewise more than 80% of the participants of a study carried out in northern Nigeria were Duffy-negative [30]. Similarly, another report among pregnant women in Nigeria confirmed a 9:1 Duffy negative to positive ratio [31]. Thus, the null expression of the Duffy gene and reduced prevalence of P. vivax in sSA suggests an interplay of adaptive features being employed by RBC overtime. Consequently, the rare observation of FYA and FYB alleles in this part of the world [32, 33] substantiates the argument of the selective adaptation of the Duffy gene on RBC and its exertion in sSA.

There are several reports of vivax malaria in hitherto low risk regions [34, 35, 36, 37] in sSA. The first Plasmodium invasion molecules identified was the Duffy binding proteins (DBP) otherwise referred to as, erythrocyte-binding-like (EBL) proteins. These proteins are characteristically homogenous in their cysteine-rich Duffy-binding-like (DBL) domains. The EBL protein of P. falciparum and P. knowlesi are able to use different receptors in invading the RBC due to variation in EBL gene numbers [38]. Therefore, it is likely that, P. vivax having phylogenetic similarity to P. knowlesi might have developed diverse EBL proteins also capable of binding to other receptors in the RBCs [2]. The high genome diversity in P. vivax [39, 40] provides a plausible explanation that supports and augments its capability of developing alternative invasion pathways through other receptors [41]. An alternate hypothesis is that, P. vivax is more endemic in north Africa as well as the Afro-Asiatic populations found in Sudan, Somalia [42, 43] and Ethiopia [20, 44] where there are higher populations of Duffy positive individuals. In this case, the Duffy positive carriers are reservoirs of infection to Duffy negative individuals. Despite the two assumptions, the precise route of transmission of P. vivax to/from Africa has not been fully established.


3. Increasing trends of P. vivax in sub-Saharan Africa and potential selection for Duffy-negative individuals

Previous reports on vivax epidemiology are now under scrutiny due to the use of more sensitive detection tools [37, 45, 46, 47, 48, 49, 50, 51]. P. vivax has been detected in south-eastern Senegal: a region sharing border with Mali, Guinea Conakry and The Gambia. In addition, the detection of Anopheles vector capable of transmitting vivax in Kenya [52], as well as identification of vivax in mosquito in a study carried out in Angola and Equatorial Guinea [35] points to active transmission of vivax malaria in this sub-region. In a case report, a pregnant Nigerian residing in Italy visited Nigeria briefly and was diagnosed with vivax infection [53], with no further investigation as to the source of infection. Whether these infections were transmitted from Duffy positive individuals serving as reservoir to the negative individuals was not determined.

Although DBP1 possess unique sequences and are highly polymorphic [53], as revealed from correlation analysis of DBP1 mutations, they do not explain the ability of P. vivax to infect Duffy-null Africans [54]. The high haplotype diversity of DBP1 and its flanking region in Ethiopia is predictive of multiple independent emergence of vivax variants that were not purged from the population [55]. Therefore, genome-wide comparative studies of Duffy-dependent and emerging Duffy-independent strains are critical for understanding the different forms of adaptive responses, fitness and selection operating in the parasites. The occurrence of DBP1 gene amplification in Ethiopia [55], Madagascar [56] and Cambodia [57] is implicative of adaptation to blood stage infection in the host. A substantial knowledge of the patterns of genetic variation in different geographic scales as well as genetic differences between populations permits inference of population relatedness or structural variation [58].


4. Strategies for P. vivax diagnosis in Africa in the era of malaria elimination

Diagnosis of P. vivax has relied primarily on two approaches; light microscopy and rapid diagnostic tests (non-molecular). However, modern tools (molecular based tests) such as PCR tests have been demonstrated to be useful, but has suffered widespread utilisation in African countries due to lack of expensive equipment and trained personnel to carry these complex assays.

4.1 Light microscopy (LM)

The World Health Organisation’s gold standard for P. vivax testing is the detection of parasites in Giemsa stained blood smears by light microscopy [59]. In detecting parasitemia from blood, thick smears are more sensitive, while thin smears allow for clearer visualisation of the morphology of the parasites for accurate speciation and calculation of parasite density. It is recommended that 3 sets of thick and thin smears should be collected for analysing P. vivax suspected blood smears. Light microscopy has a number of benefits in African health laboratories such as low cost of operation, and ease of deployment on the field. However, in Africa, the main constraint in accurate microscopy diagnosis is the lack of trained microscopists, and epileptic power supply in most parts of the continent. One of the main challenges in microscopic diagnosis of P. vivax is the presence of fewer number of parasites circulating in the blood compared to P. falciparum infections making it more likely to report false negatives. The preference of P. vivax for reticulocytes which accounts for less than 1% of the fraction of erythrocytes in circulation explains the reason for lower parasite densities compared to P. falciparum. In addition, clinical immunity to P. vivax infections occur earlier and the immune response is effective in the control of parasitemia [60].

4.2 Malaria rapid diagnostic tests (mRDTs)

The use of mRDTs has become very popular and critical to controlling malaria in Africa. Malaria RDTs provide results within 15 to 20 minutes, do not require special training to interpret results and special storage facilities. The use of mRDTs has contributed significantly to reducing the burden of malaria and is the most commonly used approach in rural laboratory settings with a sensitivity as high as 99% for P. falciparum. Malaria RDTs that test for P. vivax detect the P.vivax - pLDH antigen specific to P. vivax, or pan-pLDH or aldolase antigens common to all Plasmodium species. The sensitivity of mRDTs in detecting P. vivax, is lower [61, 62, 63] when compared to those for falciparum malaria. Moreover, cross reactions with other disease conditions, presence of certain immunological factors and gametocytaemia can result in false positives. They also have a limited shelf life (~24 months) and need to be kept dry and away from extreme temperatures (> 40°C). Malaria RDTs are very reliable at high parasitemia, while at lower parasitemia chances of false negatives increase [64]. A combination of RDTs with light microscopy would significantly improve P. vivax diagnosis in Africa.

4.3 Polymerase chain reaction (PCR)

LM has been the gold standard for malaria diagnosis, but with substantial progress in malaria control, interventions have begun to consider asymptomatic carriers as well. Hence, there is increase interests in the utilisation of molecular tools for detection of sub-microscopic infections. Molecular techniques allow investigation of 5–10 μL whole blood, which increases test sensitivity when compared to 0.025–0.0625 μL whole blood) used for LM [59].

PCR based tests such as conventional PCR and real time PCR have been used to detect P. vivax mostly in laboratory settings as part of research projects and not usually as routine clinical based tests. PCR based tests detect P. vivax with high sensitivities in diagnosing asymptomatic people particularly in samples with low parasitemia. Molecular based tests have a lower detection limit than both mRDTs and LM. The type of input material (DNA, RNA or whole blood) target gene, species detected, primer/probe composition and concentration, amplification technique (PCR or isothermal), read-out (gel electrophoresis, fluorescence detection, lateral flow) and whether it is qualitative or quantitative impact on the sensitivity of molecular detection tests. The standard molecular marker for Plasmodium species differentiation detection is the 18S rRNA genes [65]. In the reference genomes of P. vivax (Sal1 and P01), there are 3 distinct copies of 18S rRNA that are expressed at different stages [66]. Some of the Pv18S rRNA assays in use target only one of the three copies by qPCR [67]. Pv18S rRNA assays have also been used to target PV18S rRNA transcripts in addition to the genes [67, 68]. The detection of P. vivax in clinical samples can also be improved by targeting high multi-copy genes in PCR assays which provides improved sensitivity over low-copy Pv18S rRNA approaches. Targeting multicopy genes allows pooling samples in community surveys in areas with low parasite prevalence or in elimination settings. Pooling will also bring down costs of assays without compromising on the sensitivity. An example of a multi-copy gene that has been tested for P. vivax diagnosis is the non-coding subtelomeric repeat sequence Pvr 47 which has 14 copies in the P. vivax Sal1 reference genome. The Pvr47 single step PCR was almost as sensitive as the P. vivax 18S rRNA nested PCR [69]. Other targets such as vir/pir multigene family with high copy numbers similar to the var. gene family in P. falciparum have also been tested with limited success [70, 71]. Other molecular assays for P. vivax have also targeted the mitochondrial DNA in a one-step PCR reaction, loop mediated isothermal amplification or qPCR targeting cytochrome C oxidase I (cox1) [72, 73, 74], genus-specific PCR followed by targeting vivax specific non-coding regions between cytochrome B gene and cox1 [75].

Loop-mediated isothermal amplification (LAMP) assays amplify single or multi-copy molecular markers in an isothermal reaction. This technique is useful at point of care settings in the field because of ease of implementation and limited infrastructure requirement. LAMP is also useful in detecting submicroscopic infections with high sensitivity [76, 77].


5. Public health implications of P. vivax and P. falciparum co-endemicity

Co-infections of P. vivax and P. falciparum are often unrecognised in sSA, likely due to the predominance of P. falciparum as the major infecting species in most parts of the continent and the dogma that P. vivax was absent in sSA [78, 79]. With several lines of evidence, it is now being appreciated that P. vivax prevalence in sSA might actually be much higher than presumed [80, 81, 82]. Detection of mixed species infection with Plasmodium vivax, are also missed as most health care facilities use mRDTs specific for P. falciparum and usually do not have skilled man-power and resources to analyse all species that might be present by microscopy. In health facilities that use microscopy, misdiagnosis of co-infections of P. falciparum and P. vivax as P. vivax mono-infection, will lead to wrong treatment which could increase antimalarial resistance or P. falciparum parasitemia, leading to severe malaria. There are conflicting reports on the impact of co-infections of P. falciparum and P. vivax, some studies reported a decrease in disease severity through cross species immunity [83], while others have suggested an increased morbidity in children [84]. A recent meta-analysis also demonstrated a high prevalence of severe mixed malaria compared to P. falciparum mono-infection in the studies included in the meta-analysis [85]. This finding has serious implications on malaria public health control efforts and further reiterates the reason why malaria remains a leading cause of morbidity and mortality globally. Also, this emphasises the importance of improved diagnostic tools and the use of species-specific mRDTs in combination with routine microscopy or PCR where present for continuous surveillance. The possibility of mixed infections resulting in a significantly higher risk of developing severe malaria than patients with P. falciparum mono-infections is a grave public health risk, considering that mixed infections are underreported in sSA [86, 87]. This could be one of the reasons why treatment failure from ACT use and antimalarial drug resistance is on gradual increase on the continent. Taken together, it is important that clinical laboratories recognise the likelihood of other non-falciparum species especially vivax in their setting, considering the movement of persons (due to trade within the continent) that could carry parasites from one location to the other. Therefore, improved capacity for detection of suspected cases of mixed infections through a combination of species specific mRDTs together with microscopy is crucial for accurate treatment.


6. Conclusion

Duffy negativity is no longer a guaranteed barrier against P. vivax invasion, and the transmission of Duffy-independent adapted-vivax parasites will interrupt malaria control. National malaria elimination programmes should prioritise P vivax surveillance. Focus should also be on P. vivax biology especially as it concerns erythrocyte invasion in order to improve control strategies. Fine-scale epidemiological mapping is also required to elucidate the evolutionary dynamics of erythrocyte invasion by P. vivax.



The authors appreciate all their colleagues who provided valuable suggestions on this manuscript especially Dr. Eniyou C. Oriero.


Conflict of interest

All authors declare that they have no competing interest.


  1. 1. WHO. World Malaria Report. 2018.
  2. 2. Rosenberg R. Plasmodium vivax in Africa : hidden in plain sight ? Trends Parasitol. 2007;23:193-196.
  3. 3. Daniels RF, Deme AB, Gomis JF, Dieye B, Durfee K, Thwing JI, et al. Evidence of non-Plasmodium falciparum malaria infection in Kédougou, Sénégal. Malar J. 2017;16:9.
  4. 4. Diallo MA, Badiane AS, Diongue K, Deme A, Lucchi NW, Gaye M, et al. Non-falciparum malaria in Dakar: A confirmed case of Plasmodium ovale wallikeri infection. Malar J. 2016;15:429.
  5. 5. Malleret B, Li A, Zhang R, Tan KSW, Suwanarusk R, Claser C, et al. Plasmodium vivax : restricted tropism and rapid remodeling of CD71-positive reticulocytes. Blood. 2015;125:1314-1325.
  6. 6. Baird JK, Rieckmann KH. Can primaquine therapy for vivax malaria be improved? Trends Parasitol. 2003;19:115-120.
  7. 7. Baird JK, Hoffman SL. Primaquine Therapy for Malaria. Clin Infect Dis. 2004;39:1336-1345.
  8. 8. Bennett JW, Pybus BS, Yadava A, Tosh D, Sousa JC, McCarthy WF, et al. Primaquine Failure and Cytochrome P-450 2D6 in Plasmodium vivax Malaria. N Engl J Med. 2013;369:1381-1382.
  9. 9. Mekonnen SK, Aseffa A, Medhin G, Berhe N, Velavan TP. Re-evaluation of microscopy confirmed Plasmodium falciparum and Plasmodium vivax malaria by nested PCR detection in southern Ethiopia. Malar J. 2014;13:48. doi:10.1186/1475-2875-13-48.
  10. 10. Lacerda MVG, Llanos-Cuentas A, Krudsood S, Lon C, Saunders DL, Mohammed R, et al. Single-dose tafenoquine to prevent relapse of plasmodium vivax malaria. N Engl J Med. 2019;380:215-228.
  11. 11. St Jean PL, Xue Z, Carter N, Koh GCKW, Duparc S, Taylor M, et al. Tafenoquine treatment of Plasmodium vivax malaria: Suggestive evidence that CYP2D6 reduced metabolism is not associated with relapse in the Phase 2b DETECTIVE trial. Malar J. 2016;15:97.
  12. 12. Loy DE, Plenderleith LJ, Sundararaman SA, Liu W, Gruszczyk J, Chen Y-J, et al. Evolutionary history of human Plasmodium vivax revealed by genome-wide analyses of related ape parasites. Proc Natl Acad Sci. 2018;:1-10. doi:10.1073/pnas.1810053115.
  13. 13. Hamblin MT, Thompson EE, Di Rienzo A. Complex Signatures of Natural Selection at the Duffy Blood Group Locus. Am J Hum Genet. 2002;70:369-383. doi:10.1086/338628.
  14. 14. Mu J, Joy DA, Duan J, Huang Y, Carlton J, Walker J, et al. Host switch leads to emergence of Plasmodium vivax malaria in humans. Mol Biol Evol. 2005;22:1686-1693.
  15. 15. Cornejo OE, Escalante AA. The origin and age of Plasmodium vivax Omar. Trends Parasitol. 2006;22:558-563.
  16. 16. Liu W, Li Y, Shaw KS, Learn GH, Plenderleith LJ, Malenke JA, et al. African origin of the malaria parasite Plasmodium vivax. Nat Commun. 2014;5:e3346.
  17. 17. Niang M, Diop F, Niang O, Sadio BD, Sow A, Faye O, et al. Unexpected high circulation of Plasmodium vivax in asymptomatic children from Kédougou, southeastern Senegal. Malar J. 2017;16:497. doi:10.1186/s12936-017-2146-8.
  18. 18. Wurtz N, Mint Lekweiry K, Bogreau H, Pradines B, Rogier C, Ould Mohamed Salem Boukhary A, et al. Vivax malaria in Mauritania includes infection of a Duffy-negative individual. Malar J. 2011;10:336.
  19. 19. Fru-Cho J, Bumah V V., Safeukui I, Nkuo-Akenji T, Titanji VPK, Haldar K. Molecular typing reveals substantial Plasmodium vivax infection in asymptomatic adults in a rural area of Cameroon. Malar J. 2014;13:170.
  20. 20. Lo E, Yewhalaw D, Zhong D, Zemene E, Degefa T, Tushune K, et al. Molecular epidemiology of Plasmodium vivax and Plasmodium falciparum malaria among duffy-positive and Duffy-negative populations in Ethiopia. Malar J. 2015;14:84.
  21. 21. Prugnolle F, Rougeron V, Becquart P, Berry A, Makanga B, Rahola N, et al. Diversity, host switching and evolution of plasmodium vivax infecting african great apes. Proc Natl Acad Sci U S A. 2013;110:8123-8128.
  22. 22. Buery JC, Rodrigues PT, Natal L, Salla LC, Loss AC, Vicente CR, et al. Mitochondrial genome of Plasmodium vivax/simium detected in an endemic region for malaria in the Atlantic Forest of Espírito Santo state, Brazil: Do mosquitoes, simians and humans harbour the same parasite? Malar J. 2017;16:437.
  23. 23. Miller LH, Mason SJ, Clyde DF, McGinniss MH. The Resistance Factor to Plasmodium vivax in Blacks. N Engl J Med. 1976;295:302-304. doi:10.1056/NEJM197608052950602.
  24. 24. Livingstone FB. The Duffy blood groups, vivax malaria, and malaria selection in human populations: a review. Hum Biol. 1984;56:413-425.
  25. 25. Neote K, Mak JY, Kolakowski LF, Schall TJ. Functional and biochemical analysis of the cloned Duffy antigen: identity with the red blood cell chemokine receptor. Blood. 1994;84:44-52. doi:10.1016/S0887-7963(05)80035-5.
  26. 26. Carvalho GB de, Carvalho GB de. Duffy blood group system and the malaria adaptation process in humans. Rev Bras Hematol Hemoter. 2011;33:55-64. doi:10.5581/1516-8484.20110016.
  27. 27. Reid ME, Lomas-Francis C. The blood group antigen Factbook. Elsevier Academic Press; 2004. doi:10.1007/s13398-014-0173-7.2.
  28. 28. Tournamille C, Colin Y, Cartron JP, Kim CL Van. Disruption of a GATA motif in the Duffy gene promoter abolishes erythroid gene expression in Duffy-negative individuals. Nat Genet. 1995;10:196-201.
  29. 29. Welch SG, McGregor IA, Williams K. The duffy blood group and malaria prevalence in Gambian west africans. Trans R Soc Trop Med Hyg. 1977;71:295-296.
  30. 30. Isaac IZ, John RT, Udomah FP, Imoru M, Erhabor O, Femi A. Duffy Blood Group Distribution among Patients in a Malaria Endemic Region. Int J Trop Dis Heal. 2016;19:1-5.
  31. 31. Erhabor O, Shehu CE, Alhaji YB, Yakubu A. Duffy Red Cell Phenotypes among Pregnant Women in Sokoto, North Western Nigeria. J Blood Disord Transfus. 2014;5:7-11.
  32. 32. Howes RE, Patil AP, Piel FB, Nyangiri OA, Kabaria CW, Gething PW, et al. The global distribution of the Duffy blood group. Nat Commun. 2011;2:226-225.
  33. 33. King CL, Adams JH, Xianli J, Grimberg BT, McHenry AM, Greenberg LJ, et al. Fya/Fyb antigen polymorphism in human erythrocyte Duffy antigen affects susceptibility to Plasmodium vivax malaria. Proc Natl Acad Sci. 2011;108:20113-20118. doi:10.1073/pnas.1109621108.
  34. 34. Niang M, Thiam LG, Sow A, Loucoubar C, Bob NS, Diop F, et al. A molecular survey of acute febrile illnesses reveals Plasmodium vivax infections in Kedougou, southeastern Senegal. Malar J. 2015;14:281.
  35. 35. Mendes C, Dias F, Figueiredo J, Mora VG, Cano J, de Sousa B, et al. Duffy negative antigen is no longer a barrier to Plasmodium vivax - molecular evidences from the African West Coast (Angola and Equatorial Guinea). PLoS Negl Trop Dis. 2011;5:e1192.
  36. 36. Poirier P, Doderer-Lang C, Atchade PS, Lemoine J-P, de l’Isle M-LC, Abou-bacar A, et al. The hide and seek of Plasmodium vivax in West Africa: report from a large-scale study in Beninese asymptomatic subjects. Malar J. 2016;15:570. doi:10.1186/s12936-016-1620-z.
  37. 37. Mbenda HGN, Das A. Molecular evidence of plasmodium vivax mono and mixed malaria parasite infections in duffy-negative native cameroonians. PLoS One. 2014;9:e103262.
  38. 38. Michon P, Stevens JR, Kaneko O, Adams JH. Evolutionary relationships of conserved cysteine-rich motifs in adhesive molecules of malaria parasites. Mol Biol Evol. 2002;19:1128-1142.
  39. 39. Feng X, Carlton JM, Joy DA, Mu J, Furuya T, Suh BB, et al. Single-nucleotide polymorphisms and genome diversity in Plasmodium vivax. Proc Natl Acad Sci. 2003;100:8502-8507. doi:10.1073/pnas.1232502100.
  40. 40. Imwong M, Sudimack D, Pukrittayakamee S, Osorio L, Carlton JM, Day NPJ, et al. Microsatellite variation, repeat array length, and population history of Plasmodium vivax. Mol Biol Evol. 2006;23:1016-1018.
  41. 41. Gruszczyk J, Kanjee U, Chan LJ, Menant S, Malleret B, Lim NTY, et al. Transferrin receptor 1 is a reticulocyte-specific receptor for Plasmodium vivax. Science (80- ). 2018;359:48-55.
  42. 42. Abdelraheem MH, Albsheer MMA, Mohamed HS, Amin M, Hamid MMA. Transmission of Plasmodium vivax in Duffy-negative individuals in central Sudan. Trans R Soc Trop Med Hyg. 2016;110:258-260.
  43. 43. Dalgleish T, Williams JMG., Golden A-MJ, Perkins N, Barrett LF, Barnard PJ, et al. Dsitribution of blood groups in the east African Somali Population. J Exp Psychol Gen. 2007;136:23-42.
  44. 44. Mathews HM, Armstrong JC. Duffy blood types and vivax malaria in Ethiopia. Am J Trop Med Hyg. 1981;30:299-303.
  45. 45. Erhabor O, Babatunde S, Uko KE. Some haematological parameters in plasmodial parasitized HIV-infected Nigerians. Niger J Med. 2006;15:52-55.
  46. 46. Babamale O, Ugbomoiko U. Status of malaria infection in peri-urban community of north central region of Nigeria. J Bacteriol Parasitol. 2016;07:1-6. doi:10.4172/2155-9597.1000256.
  47. 47. Mbenda HGN, Gouado I, Das A. An additional observation of Plasmodium vivax malaria infection in Duffy-negative individuals from Cameroon. J Infect Dev Ctries. 2016;10:682-686.
  48. 48. Ménard D, Barnadas C, Bouchier C, Henry-halldin C, Gray LR. Plasmodium vivax clinical malaria is commonly observed in Duffy-negative Malagasy people. Proc Natl Acad Sci. 2010;107:5967-5971.
  49. 49. Rubio JM, Benito A, Roche J, Berzosa PJ, García ML, Micó M, et al. Semi-nested, multiplex polymerase chain reaction for detection of human malaria parasites and evidence of Plasmodium vivax infection in Equatorial Guinea. Am J Trop Med Hyg. 1999;60:183-187.
  50. 50. Russo G, Faggioni G, Paganotti GM, Djeunang Dongho GB, Pomponi A, De Santis R, et al. Molecular evidence of Plasmodium vivax infection in Duffy negative symptomatic individuals from Dschang, West Cameroon. Malar J. 2017;16:74.
  51. 51. Poirier P, Lang CD, Atchade PS, Lemoine JP, Louise M, De C, et al. The hide and seek of Plasmodium vivax in West Africa : report from a large - scale study in Beninese asymptomatic subjects. Malar J. 2016;15:570.
  52. 52. Ryan JR, Stoute JA, Amon J, Dunton RF, Mtalib R, Koros J, et al. Evidence for transmission of Plasmodium vivax among a Duffy antigen negative population in Western Kenya. Am J Trop Med Hyg. 2006;75:575-581.
  53. 53. Graffeo R, Masucci L, Bugli F, Damiano F, Pinnetti C, Federico G, et al. Multiple malaria infection in a pregnant woman from Nigeria: Detection by multiplex PCR. New Microbiol. 2008;31:565-567.
  54. 54. Gunalan K, Lo E, Hostetler JB, Yewhalaw D, Mu J, Neafsey DE, et al. Role of Plasmodium vivax Duffy-binding protein 1 in invasion of Duffy-null Africans. Proc Natl Acad Sci. 2016;113:6271-6276.
  55. 55. Auburn S, Getachew S, Pearson RD, Amato R, Miotto O, Trimarsanto H, et al. Genomic Analysis of Plasmodium vivax in Southern Ethiopia Reveals Selective Pressures in Multiple Parasite Mechanisms. J Infect Dis. 2019; Xx Xxxx.
  56. 56. Menard D, Chan ER, Benedet C, Ratsimbasoa A, Kim S, Chim P, et al. Whole Genome Sequencing of Field Isolates Reveals a Common Duplication of the Duffy Binding Protein Gene in Malagasy Plasmodium vivax Strains. PLoS Negl Trop Dis. 2013;7:e2489.
  57. 57. Chan ER, Menard D, David PH, Ratsimbasoa A, Kim S, Chim P, et al. Whole Genome Sequencing of Field Isolates Provides Robust Characterization of Genetic Diversity in Plasmodium vivax. PLoS Negl Trop Dis. 2012;6:e1811.
  58. 58. Neafsey DE, Kevin Galinsky, Jiang RHY, Young L, Sykes SM, Saif S, et al. The malaria parasite Plasmodium vivax exhibits greater genetic diversity than Plasmodium falciparum. Nat Genet. 2012;44:1046-1050.
  59. 59. World Health Organisation. Basic malaria microscopy. (2010).
  60. 60. Lin, E. et al. Differential Patterns of Infection and Disease with P. falciparum and P. vivax in Young Papua New Guinean Children. PLoS ONE 5, e9047 (2010).
  61. 61. Pattanasin, S. et al. Evaluation of a new Plasmodium lactate dehydrogenase assay (OptiMAL-IT) for the detection of malaria. Trans. R. Soc. Trop. Med. Hyg. 97, 672-674 (2003).
  62. 62. Tjitra, E., Suprianto, S., Dyer, M., Currie, B. J. & Anstey, N. M. Field evaluation of the ICT malaria P.f/P.v immunochromatographic test for detection of Plasmodium falciparum and Plasmodium vivax in patients with a presumptive clinical diagnosis of malaria in eastern Indonesia. J. Clin. Microbiol.37, 2412-2417 (1999).
  63. 63. Fransisca, L. et al. Comparison of rapid diagnostic test Plasmotec Malaria-3, microscopy, and quantitative real-time PCR for diagnoses of Plasmodium falciparum and Plasmodium vivax infections in Mimika Regency, Papua, Indonesia. Malar. J.14, 103 (2015).
  64. 64. Abba, K. et al. Rapid diagnostic tests for diagnosing uncomplicated non-falciparum or Plasmodium vivax malaria in endemic countries. Cochrane Database Syst. Rev. (2014) doi:10.1002/14651858.CD011431.
  65. 65. Gruenberg, M. et al. Plasmodium vivax molecular diagnostics in community surveys: pitfalls and solutions. Malar. J. 17, 55 (2018).
  66. 66. Li, J. et al. Regulation and trafficking of three distinct 18 S ribosomal RNAs during development of the malaria parasite. J. Mol. Biol. 269, 203-213 (1997).
  67. 67. Wampfler, R. et al. Strategies for Detection of Plasmodium species Gametocytes. PLoS ONE 8, e76316 (2013).
  68. 68. Pinzon, C. et al. Real-Time Quantitative Reverse Transcription PCR for Monitoring of Blood-Stage Plasmodium falciparum Infections in Malaria Human Challenge Trials. Am. J. Trop. Med. Hyg. 86, 383-394 (2012).
  69. 69. Demas, A. et al. Applied genomics: data mining reveals species-specific malaria diagnostic targets more sensitive than 18S rRNA. J. Clin. Microbiol. 49, 2411-2418 (2011).
  70. 70. Hester, J. et al. De Novo Assembly of a Field Isolate Genome Reveals Novel Plasmodium vivax Erythrocyte Invasion Genes. PLoS Negl. Trop. Dis. 7, e2569 (2013).
  71. 71. Chen, S.-B. et al. Whole-genome sequencing of a Plasmodium vivax clinical isolate exhibits geographical characteristics and high genetic variation in China-Myanmar border area. BMC Genomics 18, 131 (2017).
  72. 72. Cunha, M. G. et al. Development of a Polymerase Chain Reaction (PCR) method based on amplification of mitochondrial DNA to detect Plasmodium falciparum and Plasmodium vivax. Acta Trop. 111, 35-38 (2009).
  73. 73. Batista-dos-Santos, S., Raiol, M., Santos, S., Cunha, M. G. & Ribeiro-dos-Santos, Â. Real-time PCR diagnosis of Plasmodium vivax among blood donors. Malar. J. 11, 345 (2012).
  74. 74. Souza, C. R. T. et al. Prevalence of Plasmodium falciparum and P. vivax in an area of transmission located in Pará State, Brazil, determined by amplification of mtDNA using a real-time PCR assay. Genet. Mol. Res. GMR 11, 3409-3413 (2012).
  75. 75. Haanshuus, C. G. et al. A novel, single-amplification PCR targeting mitochondrial genome highly sensitive and specific in diagnosing malaria among returned travellers in Bergen, Norway. Malar. J.12, 26 (2013).
  76. 76. Katrak, S. et al. Performance of Loop-Mediated Isothermal Amplification for the Identification of Submicroscopic Plasmodium falciparum Infection in Uganda. Am. J. Trop. Med. Hyg. 97, 1777-1781 (2017).
  77. 77. Lau, Y.-L., Lai, M.-Y., Fong, M.-Y., Jelip, J. & Mahmud, R. Loop-Mediated Isothermal Amplification Assay for Identification of Five Human Plasmodium Species in Malaysia. Am. J. Trop. Med. Hyg. 94, 336-339 (2016).
  78. 78. Rosenberg, R. Plasmodium vivax in Africa: hidden in plain sight? Trends Parasitol. 23, 193-196 (2007).
  79. 79. Howes, R. E. et al. The global distribution of the Duffy blood group. Nat. Commun. 2, 266 (2011).
  80. 80. Guerra, C. A. et al. The international limits and population at risk of Plasmodium vivax transmission in 2009. PLoS Negl. Trop. Dis. 4, e774 (2010).
  81. 81. Price, R. N. et al. Vivax malaria: neglected and not benign. Am. J. Trop. Med. Hyg. 77, 79-87 (2007).
  82. 82. Mendis, K., Sina, B. J., Marchesini, P. & Carter, R. The neglected burden of Plasmodium vivax malaria. Am. J. Trop. Med. Hyg. 64, 97-106 (2001).
  83. 83. McKenzie, F. E. & Mason, D. P. Blood-stage dynamics and clinical implications of mixed Plasmodium vivax-Plasmodium falciparum infections. Am. J. Trop. Med. Hyg. 61, 367-374 (1999).
  84. 84. Bruce, M. C. et al. Effect of Transmission Setting and Mixed Species Infections on Clinical Measures of Malaria in Malawi. PLoS ONE 3, e2775 (2008).
  85. 85. Kotepui, M., Kotepui, K. U., De Jesus Milanez, G. & Masangkay, F. R. Plasmodium spp. mixed infection leading to severe malaria: a systematic review and meta-analysis. Sci. Rep.10, 11068 (2020).
  86. 86. Mbakilwa, H. et al. Quality of malaria microscopy in 12 district hospital laboratories in Tanzania. Pathog. Glob. Health 106, 330-334 (2012).
  87. 87. Frean, J. et al. External quality assessment of national public health laboratories in Africa, 2002-2009. Bull. World Health Organ. 90, 191-199 (2012).

Written By

Mary Aigbiremo Oboh, Mamadou Ndiath, Olumide Ajibola, Kolapo Oyebola and Alfred Amambua-Ngwa

Submitted: October 13th, 2020 Reviewed: March 12th, 2021 Published: April 14th, 2021