Skip to main content

The biology of unconventional invasion of Duffy-negative reticulocytes by Plasmodium vivax and its implication in malaria epidemiology and public health


Plasmodium vivax has been largely neglected over the past century, despite a widespread recognition of its burden across region where it is endemic. The parasite invades reticulocytes, employing the interaction between Plasmodium vivax Duffy binding protein (PvDBP) and human Duffy antigen receptor for chemokines (DARC). However, P. vivax has now been observed in Duffy-negative individuals, presenting a potentially serious public health problem as the majority of African populations are Duffy-negative. Invasion of Duffy-negative reticulocytes is suggested to be through duplication of the PvDBP and a novel protein encoded by P. vivax erythrocyte binding protein (EBP) genes. The emergence and spread of specific P. vivax strains with ability to invade Duffy-negative reticulocytes has, therefore, drawn substantial attention and further complicated the epidemiology and public health implication of vivax malaria. Given the right environment and vectorial capacity for transmission coupled with the parasite’s ability to invade Duffy-negative individuals, P. vivax could increase its epidemiological significance in Africa. In this review, authors present accruing knowledge on the paradigm shift in P. vivax invasion of Duffy-negative reticulocytes against the established mechanism of invading only Duffy-positive individuals and offer a perspective on the epidemiological diagnostic and public health implication in Africa.


Malaria affects one-third of the world’s population and kills hundreds of thousands across the globe [1]. Plasmodium vivax is the most widespread species of human malaria and more than 3 billion people live in P. vivax-endemic regions [2]. Unlike Plasmodium falciparum, P. vivax was believed not to be a major health problem in Africa. However, until recently, P. vivax infections have been widely reported in several African countries and its emergence is not surprising given the presence of a suitable environment, competent vectors and susceptible human hosts [3]. Plasmodium vivax not only differs from P. falciparum in terms of its epidemiology, but also its biology and evolutionary trajectory. These unique features enable P. vivax to survive under a wide range of climatic and ecological settings with the abilities to adapt to both temperate and tropical climates. Other features, including P. vivax ability to form hypnozoites, a stage of which the parasite can remain quiescent from months to years [4], an invasion preference of reticulocytes that only make up a small proportion of circulating erythrocytes, the absence of in vitro cultures, and rapidly forming gametocytes all together make elimination of vivax malaria more difficult than P. falciparum [5].

The epidemiology of P. vivax in Africa remains poorly understood, although P. vivax transmission from almost all countries across the continent has been reported from various entomological, serological, community prevalence surveys, including clinical infection data from local residents and travellers returning to malaria-free countries [6]. These reports have provided substantive evidence that P. vivax transmission is ongoing throughout Africa [3, 7]. However, there is still a paucity of data on the contemporary distribution of P. vivax in the region. There are increasing reports of P. vivax infections in Duffy-negative populations of Africa [8] as determined by different diagnostic tools. Indeed, P. vivax has been observed to be less severe in Duffy-negative Africans compared to their Duffy-positive counterparts [9]. This may be underpinned by the fact that P. vivax is poorly adapted to Duffy-negative individuals and may change as the parasite evolves to optimize its ability to infect Duffy-negative populations.

Here, authors review the contemporary biology and epidemiology of P. vivax among the spectrum of Duffy phenotype populations across Africa and synthesize data accrued on the invasion mechanisms of Duffy-negative erythrocytes and novel parasite ligands and host receptors that enable the parasites to invade reticulocytes. To achieve this, published articles was searched in the following electronic databases: Google Scholar, ( and PubMed ( Search terms included Plasmodium vivax, Duffy OR CD234 OR DARC, parasite ligands, host receptors, molecular epidemiology, invasion, mechanism. No restriction on year of publication was imposed but only articles published in English were considered.

Studies were selected if they met the following criteria: (1) The study involved human participants; (2) infections analysed were either P. falciparum and P. vivax co-infections or pure P. vivax infections; and, (3) studies were conducted in African or South American populations. Selected studies were entered into Mendeley Reference Manager and the following information was recorded: (1) country where study was conducted; (2) the case prevalence of P. vivax reported; (3) Duffy phenotypes reported; (4) comments on disease pathogenesis, particularly for Duffy-negative infections; (5) proposed mechanisms of infections of Duffy-negative individuals; (6) novel receptor–ligand interactions reported; and, (7) study conclusions.

Diagnostic challenges associated with Plasmodium vivax infections

Plasmodium vivax infections persist with low levels of parasitaemia either as mono-infections or co-infections with P. falciparum in co-endemic settings, making it relatively difficult to diagnose with microscopy, which is the gold standard for malaria detection [10]. In most parts of Africa, particularly in Duffy-negative populations, malaria rapid diagnostic test kits (RDTs) are mainly aimed at detecting P. falciparum and sometimes minor species, such as Plasmodium ovale and Plasmodium malariae, but not specifically P. vivax, despite the increasing number of P. vivax reported across Africa and in Duffy-negative individuals. Therefore, P. vivax infections are often misdiagnosed either by mistake or because parasitaemia is too low given that its P. vivax invades reticulocytes that comprise only a small fraction of the circulating erythrocytes [5]. Owing to the increased ability to deploy molecular detection platforms, such as polymerase chain reaction (PCR), countries that had hitherto not reported any P. vivax infections are beginning to report increasing numbers of these infections in community surveys [11]. This is apparent that infections can be easily missed by the conventional microscopy and/or RDTs in such community-based surveillance [9, 12] and, therefore, hamper an accurate assessment of P. vivax prevalence in endemic areas across Africa. Moreover, with the advent of whole genome sequencing of Plasmodium spp, genome analysis of P. vivax from clinical sources will uncover the true magnitude of infections among Duffy-positive/negative populations [13].

Evolutionary and clinical significance of Plasmodium vivax in Duffy-negatives

For many years epidemiologist thought that Duffy-negative individuals were totally resistant to vivax malaria and the complete resistance of Duffy-negatives to vivax malaria was the most striking example of association of a genetic trait with an infectious disease [14]. On the contrary, recent studies have demonstrated the capacity of P. vivax to infect human erythrocytes with or without the Duffy antigen to cause blood-stage infections [12, 15, 16]. A longitudinal study conducted in Mali by Niangaly et al. [17], found 2% asymptomatic P. vivax infections in Duffy-negative anaemic children. In another study conducted in Cameroon by Russo et al. [18], 27 P. vivax-infected Duffy-negative patients presented with fever. In admixture populations, according to Menard et al. [15], Duffy-positive individuals may serve as a reservoir of P. vivax, providing an opportunity for this parasite to infect hepatocytes of Duffy-negative people with the selection of novel P. vivax strains with the ability to invade Duffy-negative erythrocytes. Taken together, these data suggest a paradigm shift in the understanding of P. vivax invasion biology and raises more questions than answers. Could the adaptation of vivax to infect Duffy-negative individuals be frequency dependent? If so, what ratio of Duffy phenotypes fuel vivax adaptation to invade Duffy-negatives? There is yet limited evidence which supports the inability of vivax to spread to populations such as those in West Africa where the Duffy-negative allele is fixed. At any rate, given that P. vivax can cause both clinical and sub-clinical infections in Duffy-negative patients, these evolutionary realities are of public health concern, particularly when major investments in interventions against P. falciparum are driving falciparum to elimination but creating a niche for other Plasmodium spp.

Epidemiology, disease severity and public health importance

While P. falciparum is considered the deadliest malaria parasite with most severe clinical outcomes, P. vivax is more widespread and often associated with high levels of morbidity. The overall burden, economic impact and severity associated with vivax malaria has been underestimated until some recent studies showing the association of P. vivax infections with severe malaria and death [19]. The clinical spectrum of P. vivax-associated malaria ranges from asymptomatic parasitaemia and uncomplicated febrile illness to severe and fatal malaria. Other severe clinical manifestations include multi-organ dysfunction with anaemia and thrombocytopaenia [20]. Another major public health concern of P. vivax is its association with spontaneous abortions, premature and low birth weight in pregnant women. These clinical features have mostly been described for Duffy-positive populations and may vary among Duffy-negative individuals in Africa. In areas where P. falciparum and P. vivax co-exist, while the total malaria burden decreases, the proportion of cases attributable to P. vivax increases [21].

In P. vivax, gametocytes appear simultaneously with the asexual schizonts [22] while in P. falciparum, gametocytes usually appear 10–14 days after infection. Such a difference in development not only increases the length of time the individual remains infectious, but also increases the likelihood of transmission before the infected individual seeks treatment [23]. Moreover, P. vivax ability to relapse from dormant liver-stage hypnozoites, from weeks to years after clearance of the primary blood-stage infection is a major obstacle to its control and elimination [24]. The increased population at risk of P. vivax infection and the growing clinical burden across regions with a spectrum of Duffy phenotypes puts vivax malaria on the spotlight as an important public health issue [25]. Definitively, Duffy-negative phenotype is not a barrier to P. vivax infection and the public health significance of vivax malaria should no longer be neglected [25].

Plasmodium vivax tropism to reticulocytes and Duffy antigens

The sparse distribution of P. vivax in Africa is viewed as a consequence of the lack of expression of the Duffy antigen on red blood cells (RBCs). Exhibiting distinct red cell tropism, malaria parasites use sophisticated invasion mechanisms to identify, penetrate and multiply in vector and human host cells. Plasmodium vivax blood-stage infection is restricted to human reticulocytes, whereas P. falciparum can infect all types of RBCs [26]. RBC invasion by Plasmodium species requires multiple interactions between parasite ligands and host receptors, some of which have overlapping and partially redundant roles [27]. With the evolutionary arms race between the malaria parasites and human hosts, variants of erythrocyte receptors and parasites ligands have evolved. In P. vivax, the connection between the PvDBP and DARC is a critical pathway, and the only pathway identified so far, for P. vivax invasion of human reticulocytes, in spite of the increasing reports of Pvivax infections in Duffy-negative individuals that rise questions on the specificity of Duffy antigens [28, 29]. With increasingly reports of P. vivax infection in Duffy-negative populations throughout Africa and South America, it is possible that these infections were historically not detected rather than a recent adaptation of certain P. vivax strains.

The invasion of human reticulocytes by Pvivax had been known to be completely dependent on the interaction between PvDBP and DARC [30], unlike Pfalciparum merozoites that use several erythrocyte receptors (e.g., glycophorin A (GPA) and glycophorin C (GPC) and ligands (e.g., EBA-175 and EBA-140) for erythrocyte invasion. Although DARC is present on the erythrocyte surface, P. vivax reticulocyte binding proteins (PvRBPs) bind receptors on reticulocytes that are subsequently lost during erythrocyte maturation [31]. This enhances the exposure of PvDBP binding pocket to young reticulocytes explaining P. vivax tropism [32]. Based on sequence homology, the PvRBP family comprises PvRBP1 (composed of PvRBP1a and PvRBP1b) and PvRBP2 (composed of PvRBP2a, PvRBP2b, PvRBP2c) [33]. PvRBP2d and PvRBP3 are pseudo-genes that share homology with PvRBPs but do not encode functional proteins. According to Miller et al., Duffy-negative individuals can be infected by P. vivax, but are protected from blood-stage P. vivax infection [34] indicating that such infections could persist as asymptomatic.

DARC polymorphisms and Plasmodium vivax invasion canon

DARC is a membrane protein present on the surface of erythrocytes and central to P. vivax merozoites invasion of RBCs. Due to its strong geographic differentiation and association with vivax malaria resistance, DARC is considered as best example to depict positive selection in human genome. It belongs to the G protein-coupled receptor family (a 35–50 kDa glycoprotein) expressed both on the surface of RBCs and endothelial tissue [35]. Duffy proteins are also expressed in erythroid precursor cells of the bone marrow where P. vivax invasion could be occurring to some degree [26]. The Duffy glycoprotein is encoded by the FY gene main alleles, FY*A (FY*01) and FY*B (FY*02) that are codominantly expressed (Table 1). Each allele can be inherited either from mother or father and both gene products, Duffy Fya and Fyb antigens, can be expressed on the RBCs. Fya and Fyb differ by a single amino acid at position 42 that encodes glycine in Fya and aspartic acid in Fyb. Antibodies against Fya and Fyb (anti-Fya and anti-Fyb) define four main phenotypes: Fy(a+b−), Fy(a+b+), Fy(a−b+) and Fy(a−b−). The Duffy-negative phenotype, Fy(a−b−), occurs in two-thirds of African–American Blacks [36] and is associated with a single point mutation c.1-67T>C (rs2814778) in the GATA-1 binding motif for the erythroid promoter of FY*B  [35] that prevents the expression of FY*B in RBCs (thus blocks the RBC invasion by P. vivax), but without affecting FY*B expression in other tissues [37, 38].

Table 1 Variants of the Duffy blood group (ISBY008)

The gene encoding the Duffy blood group, Fy, is characterized by a SNP in a GATA-1 transcription factor binding site associated with the erythrocyte silent (ES) phenotype that has been shown to protect against P. vivax infection and is at near fixation in sub-Saharan Africa and is virtually absent any where else. Indeed, the Duffy antigen exhibits extreme geographic differentiation in Africa, but nearly absent from Asia and Europe. While ES (Fy*BES allele; FY*02N.01) phenotype impairs promoter activity in erythroid cells by disrupting a binding site for the GATA-1 erythroid transcription factor [37] and confers complete protection from vivax malaria, low levels of P. vivax infections have been observed in Fy*01N.01 homozygotes [39], which indicates that P. vivax might be evolving escape-variants able to overcome the protective effect of Fy*01N.01/FY*01N.01 or has evolved a Fy-independent RBC invasion pathway or that the GATA-1 SNP does not abolish Fy expression. Different allelic forms of the Duffy blood group gene could also modify the antigen’s expression level, leading to weak phenotype. For example, two variants c.265C>T (rs34599082) and c.298G>A (rs13962) of glycoprotein Duffy have resulted in weak Fyb expression [40]. Another mutation c.145G>T generating the FY*02W.02 allele has also shown to weaken the expression of the Fyb antigen [41, 42]. The expression level of these different weak phenotypes needs verification through flow cytometry in future studies.

Plasmodium vivax DBP gene duplication and Duffy-negative reticulocyte invasion

Red blood cell invasion by Pvivax merozoites appears to rely heavily on the interaction between PvDBP and DARC [43]. Whole-genome sequencing of P. vivax clinical isolates collected from Duffy-negative individuals indicates the presence of two and higher copies of the PvDBP gene [44]. This observation suggests that the PvDBP gene is under strong positive selection [45]. Two different types of the duplication have been observed among P. vivax clinical isolates from Cambodia [46], Ethiopia [44] and Madagascar [47]. Moreover, analysis of Pvivax genomes from over 200 isolates originating from across the world indicated that copy number variation (CNV) at the PvDBP locus is one of the most common CNVs within the Pvivax genome irrespective of the geographical origin of the isolates [13]. For instance, in Cambodia, where Duffy-negativity is essentially absent, the frequency of PvDBP duplication had reached 40% [13]. PvDBP CNVs seem to be continuously arising de novo, occurring independently within defined geographic boundaries based on phylogeographic analysis. Apparently, PvDBP is also evolving rapidly among Duffy-negative population in most parts of the sub-Saharan Africa, presumed to be the origin of P. vivax [48]. In other geographic areas where almost all individuals are Duffy-positives, PvDBP duplications have also been observed with high frequency [13]. The outcome of this repeated gene duplications (or by population size expansion) from the ancestral lineage in P. vivax indicates that several sub-populations are present [49].

CNVs are common in both P. falciparum and Pvivax [13]. What initiates PvDBP expansion in P. vivax? For example, expansion of mdr1 has been shown to result in increased anti-malarial resistance, however, it is unlikely that anti-malarial drug pressure plays a role in PvDBP duplication unless the duplication confers an increased growth rate to P. vivax parasites [44]. Owing to the critical role that PvDBP-DARC interaction plays in Pvivax erythrocyte invasion, it is expected that the binding affinity of DARC with high-copies PvDBP could be higher than with single-copy PvDBP parasites. However, no association has been found between PvDBP duplications and the efficiency of parasite to invade reticulocytes with FY*A or FY*B genotypes [44]. Assuming a high adaptability of Plasmodium species, it is likely that the duplications emerged in the parasites in response to the variation in human Duffy blood group across malaria-endemic settings. Recent studies have shown that PvDBP gene amplification not only facilitates binding to an alternative lower affinity receptor in Duffy-negative reticulocytes [16, 44], but also allows P. vivax to evade host anti-PvDBP humoral immunity [50], reassuring PvDBPII region as a promising candidate for a blood-stage vaccine against P. vivax [34]. The polymorphic nature of PvDBP certainly allows P. vivax to colonize diverse ecological niches as well as to evade the host immune system.

A molecular epidemiology study conducted in Ethiopia observed that the proportion of parasites with PvDBP amplification was higher in individuals carrying the FY*A allele compared to individuals carrying the FY*B [51]. On the other hand, in Madagascar where both Duffy-negative and Duffy-positive individuals coexist, PvDBP amplification is not selected in response to the Duffy null homozygotes [52]. It is noteworthy to see a higher frequency (56%) of multiple PvEBP copies in Madagascar where Duffy-negative and Duffy-positive people coexist than in Cambodia (19%) where mostly Duffy-positive people are present [53]. Such a contrast may imply a functional role of PvEBP amplification towards Duffy-negative erythrocyte invasion.

Evidences of Plasmodium vivax in Duffy-negative individuals

The first indirect evidence of invasion of Duffy-negative reticulocytes by P. vivax came from travellers who presented with P. vivax infection after returning from countries in Africa where Duffy-negativity is at near fixation [54, 55]. Because P. vivax can remain as latent infections for months or even years before triggering a relapse infection, it is difficult to confidently identify the source of P. vivax responsible for the blood stage infection in those individuals. In addition, there is yet insufficient data demonstrating the presence of P. vivax in Anopheles mosquitoes collected in areas where the majority of people are Duffy-negative [39]. However, with the advent of sensitive molecular detection techniques, there has been a growing number of reports of P. vivax infections in Duffy-negative individuals.

The detection of P. vivax DNA by molecular and serological assays in a Duffy-negative individual’s blood sample could represent pre-erythrocytic stages of the parasites, independent of blood-stage infection [48]. Therefore, microscopic observation of P. vivax within Duffy-negative erythrocytes coupled by confirmation by genotyping has essentially proved Duffy-negative infection [50]. The first microscopic observation of P. vivax infection in a Duffy-negative erythrocyte was reported by Ryan et al., in Kenya in [39] followed by a definitive evidence of a P. vivax infection within Duffy-negative erythrocyte from patients in Madagascar [15]. Further experimental evidence on host susceptibility was obtained from a Nigerian who got infected with P. vivax through mosquito bite followed by the establishment of P. vivax blood-stage infection [55], though the Duffy status of the infected individual was not confirmed. Future investigations of expanded anti-P. vivax antibody testings in Duffy-negative populations would reveal the extent of exposure and possible adaption of P. vivax for continuous transmission in these populations. Furthermore, P. vivax infections of Duffy-negative people have also been reported from the American continent using serological, molecular, and microscopic detection tools [56, 57]. While Duffy negativity is clearly no longer a barrier against P. vivax infections, a recent study of P. vivax patients in the Brazilian Amazon region has shown that the null allele FY*02N.01 and weak allele FY*02W.01 were associated with low parasitaemia and low symptomatology [58]. The presence of the polymorphic alleles could lead to a possible reduction in P. vivax adhesion by the reduction of the Duffy glycoprotein, as supported by earlier studies showing erythrocytes, expressing the FY*/FY*02N.01, have a significant reduction in parasite adhesion when compared to erythrocytes expressing FY*02/FY*02 [59,60,61]. Therefore, the presence of FY*02N.01 and FY*02W.01 alleles may have an impact on the reduction of clinical manifestation in malaria, leading to the development of sub-clinical malaria.

Suggested mechanisms of Duffy-negative reticulocyte invasion by Plasmodium vivax

Understanding of the mechanisms of RBC invasion by P. vivax in the absence of Duffy-positive receptor is imperative with respect to parasite epidemiology, public health importance and elimination strategies against vivax malaria. To date, the question of whether P. vivax has recently evolved its strategies to infect Duffy-negative erythrocytes and utilized an alternative DARC-independent invasion pathway remains unclear. Characterizing this pathway is important in the development of innovative strategies to prevent spread of P. vivax infections in Africa, and vaccine development. Unfortunately, the inherent challenge of culturing P. vivax in vitro has made studies targeting vivax malaria very difficult [62]. The involvement of putative parasite ligands in the invasion of Duffy-negative individuals has been described in a number of studies, but it remains unclear what specific receptor on the human erythrocyte is aiding the entry of P. vivax in Duffy-negative reticulocytes [50].

One suggested alternate invasion pathway of P. vivax is through the interaction between the human Duffy antigen/receptor for chemokines (DARC, CD234) and the P. vivax EBP2, a new member of the DBP family [63]. Based on genome sequences of P. vivax field isolates and monkey-adapted strains, two possible invasion mechanisms have been proposed. The first mechanism is through duplication of the PvDBP gene observed in multiple P. vivax strains. The other is invasion through a novel PvEBP-driven pathway that contains conserved Duffy-binding-like and C-terminal cysteine-rich domains that can be recognized by the Duffy receptor (Fig. 1) [64]. It is unclear what other possible host receptor(s) are recognized by PvEBP during the invasion process in Duffy-negative individuals. However, panels of receptors on erythrocytes and reticulocytes as well as parasite ligands have been well described as the possible alternative pathways for Duffy-negative invasion [9].

Fig. 1
figure 1

Interaction between parasite ligands and reticulocyte receptors during invasion process of Duffy-positive and Duffy-negative cells by Plasmodium vivax (adapted from Popovici et al. [50]). A spectrum of ligands on the merozoite surface of the parasites that may bind to Duffy-negative and Duffy-positive reticulocytes are presented

One potential parasite ligands involved in Duffy-positive/Duffy-negative reticulocyte invasion is the family of P. vivax reticulocyte binding proteins (PvRBPs) [31]. Although PvRBPs ligands bind to erythrocytes, there are contradictory results regarding their specific binding to reticulocytes [9, 65]. Of the different PvRBPs previously examined, PvRBP1a and PvRBP2c bind to Duffy-negative erythrocytes [31]. However, PvRBP1a does not seem to be critical for P. vivax invasion of Duffy-positive cells [66]. The third member of the PvRBP family, PvRBP2b, was recently shown to be a key ligand involved in reticulocyte recognition and invasion through the transferrin receptor 1 (TfR1) [29]. However, the role of PvRBP1a and of PvRBP2b-TfR1 recognition of Duffy-negative red blood in P. vivax invasion of reticulocytes remain unclear. The membrane protein TfR1 is lost during red blood cell maturation and, therefore, absent in normocytes [67]. Theoretically, TfR1 should be present on the surface of both Duffy-positive and Duffy-negative reticulocytes. The critical interaction between PvRBP2b and TfR1 occurs upstream of the PvDBP-Duffy interaction, but is not independent of the latter [29]. Although the role of this interaction in Duffy-negative has not been assessed, it is proposed to play a key role in Duffy-independent invasion pathways [50]. PvRBP2b first binds to the TfR1 present on reticulocytes before PvDBP engages with the Duffy protein, allowing entry of the merozoite in the red blood cell during invasion in Duffy-positives [50]. It has been shown that DARC knockout and TfR mutants individually reduced P. vivax parasite invasion.

Plasmodium vivax merozoite surface protein-1 paralog (PvMSP1P) and P. vivax glycosylphosphatidylinositol-anchored micronemal antigen (PvGAMA) were also shown to bind to both Duffy-positive and Duffy-negative RBCs [68,69,70]. Taken together, this suggest that PvGAMA and PvMSP1P ligands could be involved in Duffy-independent reticulocyte invasion pathways even though their function is yet to be fully determined. Furthermore, P. vivax erythrocyte-binding protein (PvEBP, also called PvDBP2) has also been shown to bind moderately to Duffy-negative reticulocytes [64, 71].

It is as yet unclear whether a few Duffy molecules present on the surface of the erythrocyte enabling parasites with multiple PvDBP gene copies to invade the cell, or whether the invasion process of Duffy-negative reticulocytes could occur through alternate pathways by non-DBP ligands (PvEBP, PvMSP1P, or PvGAMA) [50]. Another plausible explanation for Duffy-negative invasion is that the Duffy-negative phenotype may not be entirely a Duffy null and that the Duffy receptor may be expressed at a subtle level sufficient enough for low level invasion. The c.-67T>C mutation in GATA-1 transcription factor has been confirmed in P. vivax-infected Duffy-negative individual [15]. This mutation is known to reduce the binding of the GATA-1 transcription factor and gene transcription. However, the reduced levels of RNA transcripts may still permit low expression and binding of the Duffy receptor protein with PvDBP. Thus, it is possible that Duffy-negative individuals could have been asymptomatic carriers of P. vivax for some time before symptoms emerged. These asymptomatic cases could have escaped the detection by current diagnostic tools and/or regimes in most African countries that primarily target P. falciparum. Plasmodium vivax is now confirmed not to be completely absent from western and central Africa, though the dominance of Duffy-negative individuals in these regions has kept the prevalence of P. vivax low. Moreover, the presence of very low proportion of Duffy-positive individuals could have minimized P. vivax transmission in these regions, though there is no evidence of direct transmission among Duffy-negative individuals and between Duffy-positive and Duffy-negative individuals. Lastly, the contribution of alternative hosts such as apes [72], the geographical range of P. vivax parasites [73], its red blood cell invasion protein repertoire [74], and the human and non-human primate hosts that are infected [75] cannot be neglected in facilitating transmission. Given that Duffy-negative people are not completely resistant to P. vivax blood stage malaria [11], new reservoirs of P. vivax within infected individuals must be considered [26].


Infection of Duffy-negative individuals by P. vivax remains an enigma for parasitology and evolution. Since most individuals of African origin are Duffy-negative and do not express the DARC antigen on the surface of their erythrocytes, it is unclear whether the high prevalence of Duffy-negative alleles in African populations results from P. vivax resistance selection [76] or there exists hidden transmission of P. vivax among Duffy-negative populations. The public health significance of P. vivax is increasingly apparent in Africa. Duffy-negative reticulocyte infection by P. vivax highlights the risk of emergence and spread of vivax malaria in Africa where it was non-existent before. Apart from the Horn of Africa, Madagascar and Mauritania, P. vivax is grossly overlooked in many parts of the African continent. Given the low parasitaemia associated with P. vivax infections in Duffy-negative individuals, microscopy and RDTs are not sensitive enough to detect. Therefore, the contemporary epidemiology of vivax malaria in Africa is vastly unknown [17]. This calls for sensitive molecular detection tools such as PCR for the diagnosis of low-density P. vivax infections and studies that further characterize genomic loci under selection in P. vivax isolates infecting Duffy-negative individuals. Future studies towards a better insight of erythrocyte invasion mechanism could be obtained by comparison of gene expression profiles of parasites infecting Duffy-negative and Duffy-positive people and identifying genes specifically up-regulated in Duffy-negativity infections. The evolution of P. vivax strains able to infect RBCs not expressing Duffy antigen could have important implications for vaccine development. Increasing efforts for continuous surveillance of P. vivax strains capable of invading red cell through Duffy-independent pathway is essential for designing P. vivax-specific vaccine candidates. A better understanding of the epidemiology of P. vivax enables the design of P. vivax-specific control and elimination strategies. Given the presence of an ideal temperature, highly competent vectors for its transmission, and the apparent increase in the host range, P. vivax seems to expand to areas where it was non-existent in the past in Africa. It is imperative to consider how the growing evidence that P. vivax is not restricted to a Duffy-dependent invasion pathway is affecting the African population at risk of vivax malaria.

Availability of data and materials

Not applicable.



Copy number variants


Glycophorin A


Glycophorin C


Duffy antigen receptor for chemokines


Malaria Atlas Project


Multidrug resistance 1


Single nucleotide polymorphism


Polymerase chain reaction


Plasmodium vivax Duffy binding protein


Plasmodium vivax erythrocyte binding protein


P. vivax glycosylphosphatidylinositol-anchored micronemal antigen


Plasmodium vivax merozoite surface protein-1 paralog


Plasmodium vivax reticulocyte binding proteins


Ribonucleic acid


Malaria Rapid diagnostics tests


Reticulocyte binding proteins


Red blood cells


Transferrin receptor


  1. Kumar H, Tolia NH. Getting in the structural biology of malaria invasion. PLoS Pathog. 2019;15:e1007943.

    CAS  PubMed  PubMed Central  Google Scholar 

  2. Battle KE, Lucas TCD, Nguyen M, Howes RE, Nandi AK, Twohig KA, et al. Mapping the global endemicity and clinical burden of Plasmodium vivax, 2000–17: a spatial and temporal modelling study. Lancet. 2019;394:332–43.

    PubMed  PubMed Central  Google Scholar 

  3. 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;2:266.

    PubMed  PubMed Central  Google Scholar 

  4. Adams JH, Mueller I. The biology of Plasmodium vivax. Cold Spring Harb Perspect Med. 2017;7:a0255585.

    Google Scholar 

  5. Mueller I, Galinski MR, Baird JK, Carlton JM, Kochar DK, Alonso PL, et al. Key gaps in the knowledge of Plasmodium vivax, a neglected human malaria parasite. Lancet Infect Dis. 2009;9:555–66.

    CAS  PubMed  Google Scholar 

  6. Guerra CA, Howes RE, Patil AP, Gething PW, van Boeckel TP, Temperley WH, et al. The international limits and population at risk of Plasmodium vivax transmission in 2009. PLoS Negl Trop Dis. 2010;4:e774.

    PubMed  PubMed Central  Google Scholar 

  7. 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.

    Google Scholar 

  8. Zimmerman PA, Ferreira MU, Howes RE, Mercereau-Puijalon O. Red blood cell polymorphism and susceptibility to Plasmodium vivax. Adv Parasitol. 2013;81:27–76.

    PubMed  PubMed Central  Google Scholar 

  9. Gunalan K, Niangaly A, Thera MA, Doumbo OK, Miller LH. Plasmodium vivax infections of Duffy-negative erythrocytes: historically undetected or a recent adaptation? Trends Parasitol. 2018;34:420–9.

    PubMed  PubMed Central  Google Scholar 

  10. Mendis K, Sina BJB, Marchesini P, Carter R. The neglected burden of Plasmodium vivax malaria. Am J Trop Med Hyg. 2001;64:97–106.

    CAS  PubMed  Google Scholar 

  11. Zimmerman PA. Plasmodium vivax infection in Duffy-negative people in Africa. Am J Trop Med Hyg. 2017;97:636–8.

    PubMed  PubMed Central  Google Scholar 

  12. 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.

    PubMed  PubMed Central  Google Scholar 

  13. Pearson RD, Amato R, Auburn S, Miotto O, Almagro-Garcia J, Amaratunga C, et al. Genomic analysis of local variation and recent evolution in Plasmodium vivax. Nat Genet. 2016;48:959–64.

    CAS  PubMed  PubMed Central  Google Scholar 

  14. Livingstone FB. The Duffy blood groups, vivax malaria, and malaria selection in human populations: a review. Hum Biol. 1984;56:413–25.

    CAS  PubMed  Google Scholar 

  15. Menard D, Barnadas C, Bouchier C, Henry-Halldin C, Gray LR, Ratsimbasoa A, et al. Plasmodium vivax clinical malaria is commonly observed in Duffy-negative Malagasy people. Proc Natl Acad Sci USA. 2010;107:5967–71.

    CAS  PubMed  Google Scholar 

  16. 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 USA. 2016;113:6271–6.

    CAS  PubMed  Google Scholar 

  17. Niangaly A, Gunalan K, Ouattara A, Coulibaly D, Sá JM, Adams M, et al. Plasmodium vivax infections over 3 years in Duffy blood group negative Malians in Bandiagara, Mali. Am J Trop Med Hyg. 2017;97:744–52.

    CAS  PubMed  PubMed Central  Google Scholar 

  18. 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.

    PubMed  PubMed Central  Google Scholar 

  19. Anstey NM, Russell B, Yeo TW, Price RN. The pathophysiology of vivax malaria. Trends Parasitol. 2009;25:220–7.

    CAS  PubMed  Google Scholar 

  20. Villamil-Gómez WE. Severe and complicated malaria due to Plasmodium vivax. In: Rodriguez-Morales A, editor. Current topics in malaria. Rijeka: InTech Open Publ; 2016.

    Google Scholar 

  21. WHO. World malaria report 2016. Geneva: World Health Organization; 2016.

    Google Scholar 

  22. Ngotho P, Soares AB, Hentzschel F, Achcar F, Bertuccini L, Marti M. Revisiting gametocyte biology in malaria parasites. FEMS Microbiol. 2019;43:401–14.

    CAS  Google Scholar 

  23. Olliaro PL, Barnwell JW, Barry A, Mendis K, Mueller I, Reeder JC, et al. Implications of Plasmodium vivax biology for control, elimination, and research. Am J Trop Med Hyg. 2016;95:4–14.

    CAS  PubMed  PubMed Central  Google Scholar 

  24. White NJ. Determinants of relapse periodicity in Plasmodium vivax malaria. Malar J. 2011;10:297.

    PubMed  PubMed Central  Google Scholar 

  25. Siqueira AM, Lacerda MVG, Magalhães BML, Mourão MPG, Melo GC, Alexandre MAA, et al. Characterization of Plasmodium vivax-associated admissions to reference hospitals in Brazil and India. BMC Med. 2015;13:57.

    PubMed  PubMed Central  Google Scholar 

  26. 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;125:1314–24.

    CAS  PubMed  PubMed Central  Google Scholar 

  27. Sim BKL, Chitnis CE, Wasniowska K, Hadley TJ, Miller LH. Receptor and ligand domains for invasion of erythrocytes by Plasmodium falciparum. Science. 1994;264:1941–4.

    CAS  PubMed  Google Scholar 

  28. Batchelor JJD, Malpede BMB, Omattage NNS, DeKoster GT, Henzler-Wildman KA, Tolia NH. Red blood cell invasion by Plasmodium vivax: structural basis for DBP engagement of DARC. PLoS Pathog. 2014;10:e1003869.

    PubMed  PubMed Central  Google Scholar 

  29. 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. 2018;359:48–55.

    CAS  PubMed  PubMed Central  Google Scholar 

  30. Salinas ND, Tolia NH. Red cell receptors as access points for malaria infection. Curr Opin Hematol. 2016;23:215–23.

    CAS  PubMed  PubMed Central  Google Scholar 

  31. Galinski MR, Medina CC, Ingravallo P, Barnwell JW. A reticulocyte-binding protein complex of Plasmodium vivax merozoites. Cell. 1992;69:1213–26.

    CAS  PubMed  Google Scholar 

  32. Ovchynnikova E, Aglialoro F, Bentlage AEH, Vidarsson G, Salinas ND, Von Lindern M, et al. DARC extracellular domain remodeling in maturating reticulocytes explains Plasmodium vivax tropism. Blood. 2017;130:1441–4.

    CAS  PubMed  PubMed Central  Google Scholar 

  33. Li J, Han ET. Dissection of the Plasmodium vivax reticulocyte binding-like proteins (PvRBPs). Biochem Biophys Res Commun. 2012;425:1–6.

    Google Scholar 

  34. Miller L, Mason S, Clyde D, McGinniss M. The resistance factor to Plasmodium vivax in blacks. N Engl J Med. 1976;295:302–4.

    CAS  PubMed  Google Scholar 

  35. Höher G, Fiegenbaum M, Almeida S. Molecular basis of the Duffy blood group system. Blood Transfus. 2018;16:93–100.

    PubMed  PubMed Central  Google Scholar 

  36. Parasol N, Reid M, Rios M, Castilho L, Harari I, Kosower NS. A novel mutation in the coding sequence of the FY*B allele of the Duffy chemokine receptor gene is associated with an altered erythrocyte phenotype. Blood. 1998;92:2237–43.

    CAS  PubMed  Google Scholar 

  37. Tournamille C, Colin Y, Cartron JP, Van Kim CL. Disruption of a GATA motif in the Duffy gene promoter abolishes erythroid gene expression in Duffy-negative individuals. Nat Genet. 1995;10:224–8.

    CAS  PubMed  Google Scholar 

  38. Peiper SC, Wang Z, Neote K, Martin AW, Showell HJ, Conklyn MJ, et al. The Duffy antigen/receptor for chemoklnes (DARC) is expressed in endothelial cells of duffy negative individuals who lack the erythrocyte receptor. J Exp Med. 1995;181:1311–7.

    CAS  PubMed  Google Scholar 

  39. 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;75:575–81.

    CAS  PubMed  Google Scholar 

  40. Lopez GH, Morrison J, Condon JA, Wilson B, Martin JR, Liew YW, et al. Duffy blood group phenotype-genotype correlations using high-resolution melting analysis PCR and microarray reveal complex cases including a new null FY*A allele: the role for sequencing in genotyping algorithms. Vox Sang. 2015;109:296–303.

    CAS  PubMed  Google Scholar 

  41. Lopez GH, Condon JA, Wilson B, Martin JR, Liew YW, Flower RL, et al. A novel FY*A allele with the 265T and 298A SNPs formerly associated exclusively with the FY*B allele and weak Fyb antigen expression: implication for genotyping interpretative algorithms. Vox Sang. 2015;108:52–7.

    CAS  PubMed  Google Scholar 

  42. Castilho L, Rios M, Pellegrino J, Saad STO, Costa FF, Reid ME. A novel FY allele in Brazilians. Vox Sang. 2004;87:190–5.

    CAS  PubMed  Google Scholar 

  43. Fang XX, Kaslow DDC, Adams JJH, Miller LHL. Cloning of the Plasmodium vivax Duffy receptor. Mol Biochem Parasitol. 1991;44:125–32.

    CAS  PubMed  Google Scholar 

  44. Hostetler JB, Lo E, Kanjee U, Amaratunga C, Suon S, Sreng S, et al. Independent origin and global distribution of distinct Plasmodium vivax Duffy binding protein gene duplications. PLoS Negl Trop Dis. 2016;10:e0005091.

    PubMed  PubMed Central  Google Scholar 

  45. Hupalo DN, Luo Z, Melnikov A, Sutton PL, Rogov P, Escalante A, et al. Population genomics studies identify signatures of global dispersal and drug resistance in Plasmodium vivax. Nat Genet. 2016;48:953–8.

    CAS  PubMed  PubMed Central  Google Scholar 

  46. Amaratunga C, Sreng S, Mao S, Tullo GS, Anderson JM, Chuor CM, et al. Chloroquine remains effective for treating Plasmodium vivax malaria in Pursat Province, western Cambodia. Antimicrob Agents Chemother. 2014;58:6270–2.

    PubMed  PubMed Central  Google Scholar 

  47. Menard D, Chan ER, Benedet C, Ratsimbasoa A, Kim S, Chim P, et al. Whole genome fequencing 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.

    PubMed  PubMed Central  Google Scholar 

  48. Abkallo HM, Liu W, Hokama S, Ferreira PE, Nakazawa S, Maeno Y, et al. DNA from pre-erythrocytic stage malaria parasites is detectable by PCR in the faeces and blood of hosts. Int J Parasitol. 2014;44:467–73.

    CAS  PubMed  Google Scholar 

  49. Parobek CM, Lin JT, Saunders DL, Barnett EJ, Lon C, Lanteri CA, et al. Selective sweep suggests transcriptional regulation may underlie Plasmodium vivax resilience to malaria control measures in Cambodia. Proc Natl Acad Sci USA. 2016;113:e8096–105.

    CAS  PubMed  Google Scholar 

  50. Popovici J, Roesch C, Carias LL, Khim N, Kim S, Vantaux A, et al. Amplification of Duffy binding protein-encoding gene allows Plasmodium vivax to evade host anti-DBP humoral immunity. Nat Commun. 2020;11:953.

    CAS  PubMed  PubMed Central  Google Scholar 

  51. Lo E, Hostetler JB, Yewhalaw D, Pearson RD, Hamid MMA, Gunalan K, et al. Frequent expansion of Plasmodium vivax Duffy binding protein in Ethiopia and its epidemiological significance. PLoS Negl Trop Dis. 2019;13:e0007222.

    CAS  PubMed  PubMed Central  Google Scholar 

  52. Roesch C, Popovici J, Bin S, Run V, Kim S, Ramboarina S, et al. Genetic diversity in two Plasmodium vivax protein ligands for reticulocyte invasion. PLoS Negl Trop Dis. 2018;12:e0006555.

    CAS  PubMed  PubMed Central  Google Scholar 

  53. Howes RE, Reiner RC, Battle KE, Longbottom J, Mappin B, Ordanovich D, et al. Plasmodium vivax transmission in Africa. PLoS Negl Trop Dis. 2015;9:e0004222.

    PubMed  PubMed Central  Google Scholar 

  54. Twohig KA, Pfeffer DA, Baird JK, Price RN, Zimmerman PA, Hay SI, et al. Growing evidence of Plasmodium vivax across malaria-endemic Africa. PLoS Negl Trop Dis. 2019;13:e0007140.

    PubMed  PubMed Central  Google Scholar 

  55. Bray RS. The susceptibility of Liberians to the Madagascar strain of Plasmodium vivax. J Parasitol. 1958;44:371–3.

    CAS  PubMed  Google Scholar 

  56. Kano FS, de Souza AM, de Menezes Torres L, Costa MA, Souza-Silva FA, Sanchez BAM, et al. Susceptibility to Plasmodium vivax malaria associated with DARC (Duffy antigen) polymorphisms is influenced by the time of exposure to malaria. Sci Rep. 2018;8:13851.

    PubMed  PubMed Central  Google Scholar 

  57. Carvalho TAA, Queiroz MG, Cardoso GL, Diniz IG, Silva ANLM, Pinto AYN, et al. Plasmodium vivax infection in Anajás, State of Pará: no differential resistance profile among Duffy-negative and Duffy-positive individuals. Malar J. 2012;11:430.

    PubMed  PubMed Central  Google Scholar 

  58. Abou-Ali RK, Dhyani A, Terço AL, Toro DM, Gomes KS, Tezza LC, et al. Impact of Duffy polymorphisms on parasite density in Brazilian Amazonian patients infected by Plasmodium vivax. Malar J. 2019;18:289.

    PubMed  PubMed Central  Google Scholar 

  59. King CL, Adams JH, Xianli J, Grimberg BT, McHenry AM, Greenberg LJ, et al. Fy a/Fy b antigen polymorphism in human erythrocyte duffy antigen affects susceptibility to Plasmodium vivax malaria. Proc Natl Acad Sci USA. 2011;108:20113–8.

    CAS  PubMed  Google Scholar 

  60. Woolley IJ, Wood EM, Sramkoski RM, Zimmerman PA, Miller PA, Kazura JW. Expression of Duffy antigen receptor for chemokines during reticulocyte maturation: using a CD71 flow cytometric technique to identify reticulocytes. Immunohematology. 2005;21:15–20.

    CAS  PubMed  Google Scholar 

  61. Tournamille C, Le C, Kim V, Gane P, Le Pennec PY, Roubinet F, et al. Arg89Cys substitution results in very low membrane expression of the Duffy antigen/receptor for chemokines in Fy(x) individuals. Blood. 1998;92:2147–56.

    CAS  PubMed  Google Scholar 

  62. Bermúdez M, Moreno-Pérez DA, Arévalo-Pinzón G, Curtidor H, Patarroyo MA. Plasmodium vivax in vitro continuous culture: the spoke in the wheel. Malar J. 2018;17:301.

    PubMed  PubMed Central  Google Scholar 

  63. Chootong P, Ntumngia FB, VanBuskirk KM, Xainli J, Cole-Tobian JLJ, Campbell CO, et al. Mapping epitopes of the Plasmodium vivax Duffy binding protein with naturally acquired inhibitory antibodies. Infect Immun. 2010;78:1089–95.

    CAS  PubMed  Google Scholar 

  64. Hester J, Chan ER, Menard D, Mercereau-Puijalon O, Barnwell J, Zimmerman PA, et al. De Novo assembly of a field isolate genome reveals novel Plasmodium vivax erythrocyte invasion genes. PLoS Negl Trop Dis. 2013;7:e2569.

    PubMed  PubMed Central  Google Scholar 

  65. França CT, White MT, He WQ, Hostetler JB, Brewster J, Frato G, et al. Identification of highly-protective combinations of Plasmodium vivax recombinant proteins for vaccine development. Elife. 2017;6:e28673.

    PubMed  PubMed Central  Google Scholar 

  66. Gupta ED, Anand G, Singh H, Chaddha K, Bharti PK, Singh N, et al. Naturally acquired human antibodies against reticulocyte-binding domains of Plasmodium vivax proteins, PvRBP2c and PvRBP1a, exhibit binding-inhibitory activity. J Infect Dis. 2017;215:1558–68.

    CAS  PubMed  PubMed Central  Google Scholar 

  67. Malleret B, Xu F, Mohandas N, Suwanarusk R, Chu C, Leite JA, et al. Significant biochemical, biophysical and metabolic diversity in circulating human cord blood reticulocytes. PLoS ONE. 2013;8:e76062.

    CAS  PubMed  PubMed Central  Google Scholar 

  68. Han JH, Cheng Y, Muh F, Ahmed MA, Cho JS, Nyunt MH, et al. Inhibition of parasite invasion by monoclonal antibody against epidermal growth factor-like domain of Plasmodium vivax merozoite surface protein 1 paralog. Sci Rep. 2019;9:3906.

    PubMed  PubMed Central  Google Scholar 

  69. Han JH, Cho JS, Cheng Y, Muh F, Yoo WG, Russell B, et al. Plasmodium vivax merozoite surface protein 1 paralog as a mediator of parasite adherence to reticulocytes. Infect Immun. 2018;86:e00239-18.

    PubMed  PubMed Central  Google Scholar 

  70. Cheng Y, Lu F, Wang B, Li J, Han JH, Ito D, et al. Plasmodium vivax GPI-anchored micronemal antigen (PvGAMA) binds human erythrocytes independent of Duffy antigen status. Sci Rep. 2016;6:35581.

    CAS  PubMed  PubMed Central  Google Scholar 

  71. Ntumngia FB, Thomson-Luque R, de Menezes Torres L, Gunalan K, Carvalho LH, Adams JH. A novel erythrocyte binding protein of Plasmodium vivax suggests an alternate invasion pathway into duffy-positive reticulocytes. MBio. 2016;7:e1261-16.

    Google Scholar 

  72. Culleton R, Carter R. African Plasmodium vivax: distribution and origins. Int J Parasitol. 2012;42:1091–7.

    PubMed  Google Scholar 

  73. Loy DE, Liu W, Li Y, Learn GH, Plenderleith LJ, Sundararaman SA, et al. Out of Africa: origins and evolution of the human malaria parasites Plasmodium falciparum and Plasmodium vivax. Int J Parasitol. 2017;47:87–97.

    PubMed  Google Scholar 

  74. Canier L, Khim N, Kim S, Sluydts V, Heng S, Dourng D, et al. An innovative tool for moving malaria PCR detection of parasite reservoir into the field. Malar J. 2013;12:405.

    PubMed  PubMed Central  Google Scholar 

  75. 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 USA. 2013;110:8123–8.

    CAS  PubMed  Google Scholar 

  76. Cao J, Sturrock HJW, Cotter C, Zhou S, Zhou H, Liu Y, et al. Communicating and monitoring surveillance and response activities for malaria elimination: china’s “1-3-7” strategy. PLoS Med. 2014;11:e1001642.

    PubMed  PubMed Central  Google Scholar 

Download references


Not applicable.


This work was supported through the DELTAS Africa Initiative [DELGEME grant 107740/Z/15/Z]. The DELTAS Africa Initiative is an independent funding scheme of the African Academy of Sciences (AAS)’s Alliance for Accelerating Excellence in Science in Africa (AESA) and supported by the New Partnership for Africa’s Development Planning and Coordinating Agency (NEPAD Agency) with funding from the Wellcome Trust [DELGEME grant 107740/Z/15/Z] and the UK government. The views expressed in this publication are those of the author(s) and not necessarily those of AAS, NEPAD Agency, Wellcome Trust or the UK government. This work was also funded by National Institutes of Health (R15 A1138002).The funders have no role in the design of the study and collection, analysis, and interpretation of data and in writing this manuscript.

Author information

Authors and Affiliations



LG did the initial draft. LME, EL and AAN, revised and made intellectual inputs into the manuscript. All authors read and approved the final manuscript.

Corresponding author

Correspondence to Lemu Golassa.

Ethics declarations

Ethics approval and consent to participate

Not applicable.

Consent for publication

Not applicable.

Competing interests

The authors declare that they have no competing interests.

Additional information

Publisher's Note

Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Rights and permissions

Open Access This article is licensed under a Creative Commons Attribution 4.0 International License, which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons licence, and indicate if changes were made. The images or other third party material in this article are included in the article's Creative Commons licence, unless indicated otherwise in a credit line to the material. If material is not included in the article's Creative Commons licence and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of this licence, visit The Creative Commons Public Domain Dedication waiver ( applies to the data made available in this article, unless otherwise stated in a credit line to the data.

Reprints and permissions

About this article

Check for updates. Verify currency and authenticity via CrossMark

Cite this article

Golassa, L., Amenga-Etego, L., Lo, E. et al. The biology of unconventional invasion of Duffy-negative reticulocytes by Plasmodium vivax and its implication in malaria epidemiology and public health. Malar J 19, 299 (2020).

Download citation

  • Received:

  • Accepted:

  • Published:

  • DOI: