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The global burden of Plasmodium vivax malaria is obscure and insidious

The global burden of Plasmodium vivax malaria is obscure and insidious

  • Katherine E. Battle, 
  • J. Kevin Baird
PLOS
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Summary points

  • Estimates of the global burdens of morbidity attributable to acute attacks of Plasmodium falciparum malaria typically dwarf those of Plasmodium vivax, i.e., hundreds of millions versus tens of millions of cases.
  • Global burden estimates take no account of latent and subpatent reservoirs of infections carrying more subtle burdens of illness and death in impoverished settings of malnutrition, coendemic infections, and limited access to quality healthcare. Impacts of chronic malaria on human health may be substantial and are excluded from estimates of burdens of acute malaria.
  • Compartments of human infection by P. vivax beyond vascular patency—vascular subpatency, extravascular subpatency, sexual latency, and hepatic latency—obscure endemic transmission and burdens of infection and illness.
  • Long thought to be absent from most of sub-Saharan Africa due to the high prevalence of the Duffy-negative phenotype among residents, recent investigations suggest that widespread reservoirs of transmission may occur across that region.
  • Human glucose-6-phosphate dehydrogenase (G6PD) deficiency may also affect susceptibility to infection and directly impact access to effective antirelapse therapy of P. vivax using 8-aminoquinolines that are dangerous to those patients. Natural polymorphisms of the human cytochrome P-450 2D6 gene impact parasite susceptibility to primaquine antirelapse therapy at population levels.
  • All these factors impose great complexity in considering estimates of burdens of P. vivax and access to effective mitigation of the harm caused. The conventional diagnostics underpinning epidemiological and clinical understanding of vivax malaria may be inadequate to the biology of this parasite.

Introduction

The global burden of malaria is often reported as a single value that combines the malarias caused by the 5 species of plasmodia that naturally infect humans [1]. The vast majority of this burden is attributable to Plasmodium falciparum and Plasmodium vivax malarias, which have only recently begun to be reported separately in the World Malaria Report [2]. The estimated annual burden of P. vivax malaria (14.3 million [13.7 to 15.0 million]) is an order of magnitude lower than that of P. falciparum (193.5 million [142.0 to 254.7 million]) [3,4]. Infections with Plasmodium ovale or Plasmodium malariae are geographically widespread but only rarely prevalent at detectable ranges, whereas the zoonosis caused by Plasmodium knowlesi of Southeast Asian macaques occurs only in forested areas of that region. The contributions of these 3 minority malarias to the global burden of acute malaria have not been credibly estimated but are likely to be <5%.

Authoritative estimates of burdens of P. falciparum and P. vivax refer specifically to events of clinical illness associated with patent parasitemia [3,4]. The incidence of infection per se is not accounted, despite asymptomatic carriers of infection representing an important and often dominant state of infection with regard to either species [5,6]. Semi-immune older children and adults through much of endemic sub-Saharan Africa, who greatly outnumber vulnerable young children and pregnant women, are very often infected at high rates of prevalence but only rarely suffer attacks of acute malaria [7]. Nonetheless, so-called asymptomatic infections may not be considered benign because chronic malaria carries substantial health consequences [810]. This naturally acquired immunity associated with chronic malaria has long been attributed to very high rates of exposure to repeated infections occurring almost exclusively in Africa [11]. However, recent work across the endemic globe demonstrates dominance of asymptomatic, microscopically subpatent infections even with relatively low levels of endemic transmission [1217]. Estimates of global burdens based on clinical attacks likely miss far broader and more subtle indicators of infection and the harm done in often impoverished communities facing myriad other challenges to good health.

A recent study from Indonesia offers important insights regarding that question applied to P. vivax. Dini and colleagues reported a retrospective analysis of 37,168 patients diagnosed with P. falciparum and 22,209 with P. vivax over a period of nearly 10 years [18]. Whereas a diagnosis of P. falciparum came with a higher risk of death within 14 days of diagnosis, over the longer term, greater numbers of repeated attacks and hospitalizations among P. vivax patients came with risk of death that was nearly twice that among patients with P. falciparum. As expressed by those authors in conclusion, “Whilst the acute management of malaria is paramount to prevent early death, our analysis highlights the importance of preventing recurrent malaria.” Repeated attacks of vivax malaria carried elevated risk of malaria morbidity and mortality by any cause. Renal, circulatory, and cognitive harm is done by chronic malaria infections [1924]. Harm is not limited to a single attack of acute malaria but logically extends to the varied insults to health repeatedly endured by many residents living under endemic malaria transmission. So-called silent reservoirs of infection may thus carry subtle but important burdens of illness and death not captured in global estimates of acute attacks of malaria.

The asymptomatic, microscopically subpatent reservoir of blood stage P. vivax has been described [2528], and the latent reservoir of hepatic hypnozoites has long been known [2932]. In both instances, however, understanding of their very significant contributions to both on-going transmission and acute attacks is only recent [3335]. Yet another reservoir may also prove relevant to this epidemiology: infection of the extravascular spaces of the marrow and spleen [3644]. This pathophysiology has only very recently been understood, and it transforms how P. vivax may be viewed biologically, clinically, and epidemiologically. Transferrin receptor CD71 occurs only on erythroblasts and the youngest (stage I of V) reticulocytes, and it is required for P. vivax merozoite invasion of those cells [45,46]. This points to an infection seated in deep hemopoietic tissues where CD71 naturally occurs in great abundance relative to its near absence among cells within vascular sinuses. Moreover, hematopoietic niches of plasmodial asexual schizogony (and gametocytogenesis) may be less sensitive to blood schizontocides like artemisinin [47]. Plasma-derived extracellular vesicles specific to P. vivax malaria cause up-regulation of P. vivax-adhesive molecules (ICAM-1) on spleen fibroblasts [48]. This fundamental biology of P. vivax—tropisms favoring the tissues of marrow and spleen rather than peripheral blood circulation—may often place the infection beyond the reach of conventional diagnosis, upon which global burden estimates are ultimately based. These biological intricacies of P. vivax malaria are thus important to consider in weighing estimates of its clinical burden and prevalence of infection ranges.

Important host factors further complicate those considerations of burden and countermeasures against them. Overwhelming predominance of Duffy-negative phenotype in most of sub-Saharan Africa has long been considered the basis of the relative paucity of endemic P. vivax in that region [49]. Nonetheless, travelers to that continent often acquire P. vivax infection, and recent work demonstrates widespread infection of Duffy-negative residents [5053]. Two other host factors—glucose-6-phosphate dehydrogenase (G6PDd) mutations and cytochrome P450 isozyme 2D6 (CYP2D6) polymorphisms—may also impact burdens of P. vivax. While only G6PDd may interfere with parasite development in the host [5457], both of these factors directly impact successful therapy of latent vivax malaria. The only therapeutic options against hepatic latency of malaria are the 8-aminoquinolines, and these compounds are both invariably toxic to G6PDd patients (causing a threatening acute hemolytic anemia) and appear dependent on CYP2D6 metabolic processing to generate the therapeutically active derivative [5860]. A study from Brazilian Amazonia described primaquine-induced acute hemolytic anemia as the dominant cause of blood transfusion [61], and some endemic nations have prohibited primaquine therapy for fear of such harm [62]. A survey of CYP2D6 genotypes in endemic Cambodia showed 29% of residents likely to have significantly impaired CYP2D6 activity phenotypes (predicted by genotype) at high risk of primaquine treatment failure [63].

This review considers the estimated global burdens of vivax malaria against this backdrop of great biological, clinical, and public health complexity. Aspiration for the elimination of endemic P. vivax transmission from much of Asia and the Americas within a few short years [6466] imposes the necessity of understanding burdens of infection, in addition to those of both direct and indirectly linked morbidity and mortality. Numbers of diagnosed clinical attacks as a measure of direct morbidity very likely represent a minority of infections and their health consequences.

Vector biology

Human infection by the plasmodia requires the presence of anopheline mosquitoes, and their distributions and abundance bear directly upon global burdens of malaria. Plasmodium vivax is known to be transmitted by over 70 Anopheles species with diverse bionomics. In a previous review, it was found that all Anopheles species that were incriminated to transmit P. falciparum could also transmit P. vivax [67]. However, the converse is not true—not all vectors that can transmit P. vivax can also naturally transmit P. falciparum. The minimal temperature at which sporogonic development occurs is lower for P. vivax than for P. falciparum. That biology, along with hepatic latency, largely explains the reach of endemic P. vivax transmission into temperate zones where P. falciparum only rarely occurs [68].

The bionomics of the dominant vector species in the parts of the world where P. vivax predominates are known to be more diverse than those that transmit P. falciparum in Africa [69]. The African region was identified to have 7 dominant vector species, most of them being within the Anopheles gambiae species complex. In the Asia-Pacific (where more than 80% of the global P. vivax burden is found), there are 19 described species in at least several species complexes [70,71]. Transmission of P. vivax by these diverse vectors defines varied endemic malaria ecologies: forest, coastal, plantation, paddy, hillside, and urban to name a few. Bionomic heterogeneity among the dominant vector species in P. vivax–endemic areas requires equally diverse vector control strategies extending well beyond mass net campaigns and emphasis on acute case management [7274]. Bed nets are less effective against Asian anophelines because, unlike indoor and late night feeding African vectors, those mosquitoes tend to favor feeding outdoors early in the evening [7476].

Parasite biology

Whereas in P. falciparum infectious gametocytes emerge only after several days of asexual parasitemia, in P. vivax, the asexual and sexual forms emerge together [77]. In a humanized liver rodent model, P. vivax gametocytes emerged directly from hepatic schizogony [78]. Transmission to mosquitoes may therefore occur before onset of symptoms, during early illness, and thus before treatment is obtained. Indeed, gametocytogenesis without prior blood schizogony would infer an ability to sustain wholly silent infection and transmission. Compared to P. falciparum, ordinary levels of parasitemia in uncomplicated acute attacks by P. vivax are very often an order of magnitude lower. That contrast has been attributed to the indiscriminate invasion of red blood cells of any age by P. falciparum versus the obligate preference of P. vivax for reticulocytes. This phenomenon also underpinned the spurrious idea that P. vivax is inherently unable to cause serious illness [79]. The bulk of harmful infectious biomass of P. vivax appears to lie beyond diagnostic reach in the extravascular spaces of deep hemopoietic tissues or splenic sinusoids. This biology imposes conspicuous diagnostic challenges, making both microscopic and immunochromatographic antigen detection (by rapid diagnostic tests (RDTs)) inherently less sensitive for infection by P. vivax relative to P. falciparum. Numerous studies bear this out [6,14,80]. It is a significant problem compounded by a presumably naturally acquired (or innate in the instance of Duffy negativity) immunity routinely suppressing asexual parasitemias below the limits of practical diagnosis by microscope or RDT [81, 82]. In any given survey of peripheral blood for the plasmodia by these means, the actual prevalence of P. vivax may be considerably higher than that detected, and perhaps higher still when including infections involving only the extravascular spaces of other tissues. There is an inverse correlation between prevalence of microscopically patent parasitemia as a proportion of all detectable (by PCR) parasitemias, i.e., vascular subpatency among the infected becomes more frequent as prevalence declines [6,83].

Latency in P. vivax involves dormant stages in hepatocytes called hypnozoites that awaken over the weeks, months, and few years following inoculation by a feeding mosquito. A single such event may seed the liver with any number of hypnozoites, but somewhere between 1 and 12 may be typical. The sporozoites that immediately develop to hepatic schizonts (tachysporozoites) simultaneously launch invasive merozoites into the blood to initiate symptomatic blood schizogony a week or two later. Sporozoites becoming latent hypnozoites (bradysporozoites) awaken to launch those attacks after variable (geographic location- and strain-dependent) intervals [32,84]. Latent infection may persist as long as several years and through a series of independent clinical attacks. This biology logically aligns with the estimated proportion of clinical attacks of vivax malaria originating from hypnozoites rather than recent mosquito bites being about 80% [85]. Untreated latent infections cause most patent attacks, and latency cannot now be detected by any diagnostic means.

Known features of the biology of P. vivax point to a relatively deep obscurity of presence. The actual prevalence of infection may substantially exceed that which is routinely detected by conventional means, with each of the following biological traits contributing to that character: (1) tropisms for extravascular spaces of deep organs like marrow and spleen; (2) sharply limited asexual multiplication in peripheral circulation; (3) naturally acquired or innate immune suppression of asexual blood stages; and (4) prolonged hepatic latency as a dominant state of infection.

Human genetic factors

People lacking expression of Duffy factor on the surface of red blood cells, as occurs in most of sub-Saharan Africa, has been linked to the relative scarcity of P. vivax infection in that region [86]. Several lines of evidence, however, point to endemic low-level transmission of P. vivax across much of Africa [8789]. Among Duffy-negative individuals, infections by P. vivax have been confirmed at African sites [9094]. Several surveys reported prevalent serological positivity to specific P. vivax antigens in areas where P. vivax malaria in patients or cross-sectional surveys of blood is exceedingly rare or unknown [9597]. Another study in peer review indicates expression of Duffy factor by erythroblasts in marrow despite Duffy-negative genotype and an absence of Duffy factor in circulating red blood cells [98], raising the possibility of endemic subpatent infections of hemopoietic tissues. If so, Duffy negativity may not protect from infection by P. vivax but would profoundly impact the character and detectability of that infection, i.e., limiting it to extravascular spaces of marrow (excepting transient gametocytemia). Endemic transmission of P. vivax may occur across Africa but is obscured by this host genetic factor and obligate parasite tropism for extravascular spaces of deep tissues.

G6PDd is the most common human inherited abnormality, affecting more than 400 million people and occurring at an average prevalence of 8% in malaria-endemic countries [99]. Many dozens of distinct single nucleotide polymorphisms (SNPs) are known, each being associated with variably impaired enzymatic function [100]. These variants tend to occur within distinct ethnic/geographic groups and are classified according to level of residual enzymatic activity compared to normal. The WHO classification is most often applied, where most variants are either Class II or III, representing phenotypes of <10% or >10% of normal activity, respectively [101]. One of the most conspicuous physiological distinctions between those classes may be G6PD activity phenotypes across red blood cells of increasing age. Whereas reticulocytes of the archetypical Class III A- variant of G6PDd of Africa exhibit nearly normal G6PD activity (which more sharply declines with red blood cell age compared to normal), those of archetypical Class II Mediterranean variant have almost none [102]. This biochemistry bears on global distributions of P. vivax and this inherited abnormality.

G6PDd is among the inherited blood disorders believed to have been selected by survival advantages against endemic malaria [5457,103,104]. As already explained, P. vivax exhibits profound tropisms of infection and anatomic location anchored upon CD71 as an essential molecule of invasion. Severely impaired G6PD activity in reticulocytes of Class II variants may thus have great impacts on the parasitism of P. vivax but almost none in the nearly G6PD-normal reticulocytes of Class III variants, e.g., Mahidol variant in Thailand and A- in Africa [105]. In South and Southeast Asia, where most endemic P. vivax transmission occurs, Class II variants predominate over those of Class III [106], suggesting a selection pressure at work. Populations residing within endemic zones most likely to benefit from 8-aminoquinoline therapies against latent P. vivax may also be the most likely to suffer harm caused by them. Rational fear of 8-aminoquinoline toxicity protects the hypnozoite reservoir from aggressive exposure to those drugs in clinical and public health practice [106].

Likewise, poor efficacy of 8-aminoquinolines due to natural polymorphisms of CYP2D6 may also bear upon global burdens of P. vivax. Initial demonstrations of CYP2D-dependent therapeutic activity of primaquine against hepatic plasmodia in murine malaria models has been similarly observed in humans infected by P. vivax [60,107]. CYP2D6 polymorphisms associated with null or relatively poor enzymatic activity occurred in patients experiencing failure of directly supervised high-dose primaquine therapy against relapse by P. vivax [59,107]. The most common allele of CYP2D6 in Asian populations is impaired *10. There may be relatively high risk of therapeutic failure of antirelapse therapies in as many as 40% of Asians exposed to risk of P. vivax infection [63,108].

Complex human genetic factors exert impacts on the global distribution of burdens of P. vivax infection, both directly (Duffy negativity and G6PDd) and indirectly through 8-aminoquinoline toxicity (G6PDd) or therapeutic activity (CYP2D6) problems. The direct effects of Duffy negativity may no longer be construed as wholly preventing infection by P. vivax but profoundly impacting the ability to detect the infection by conventional examinations of peripheral blood. Endemic infection and transmission may occur where Duffy negativity prevails and measures of prevalence of vascular patency seem vanishingly low, as across sub-Saharan Africa (Fig 1).

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Fig 1. The global prevalence of patent Plasmodium vivax in 2017.

The mean predicted parasite rate of P. vivax is in people 1 to 99 years of age based on the analysis from Battle and colleagues is shown on a scale of low prevalence (blue) to high (red) with very low prevalence in light grey and nonendemic regions in white [3]. Global national shapefile obtained from the Malaria Atlas Project (MAP; https://malariaatlas.org/) and available for download through the malariaAtlas R package [109].

https://doi.org/10.1371/journal.pmed.1003799.g001

Geographic distribution

The dormant liver stage and an ability to develop in its invertebrate host at lower temperatures allow P. vivax to be the most widely distributed cause of human malaria. Plasmodium vivax extends across the endemic tropics, subtropics, and well into temperate climates where the hypnozoite may lie in wait for the seasonal reappearance of feeding anophelines. Keeping in mind the challenges of detection described above, observed prevalence of patent parasitemia values vary greatly across endemic zones, very rarely exceeding 10% prevalence (Fig 1).

A century ago, the global distribution of endemic P. vivax transmission included most of North America, Europe, and northern Asia and Australia. All of these areas remain receptive to P. vivax transmission by numerous competent and seasonally abundant vectors. Outbreaks of P. vivax malaria sporadically occur in these regions, typically in association with migrant human populations [110113]. The latency of P. vivax—which is not now possible to diagnose—makes the prevention of imported malaria particularly difficult.

Populations at risk

Accepting prevalent P. vivax as evidence of endemic transmission, whether stable or unstable, well over one-third of world’s population (3.3 billion people) were estimated to be living at risk of P. vivax transmission in 2017 [3]. Of that population, 1.6 billion were in the Southeast Asian WHO region (SEARO), with 80% of the total population in that region being considered at risk. Following SEARO, the Western Pacific (WPRO) and African (AFRO) regions had 661 and 629 million people at risk, respectively. The Eastern Mediterranean Region (EMRO) has nearly half of the population at risk of these regions (311 million), but it disproportionately contributes to the global burden of clinical cases as shown in Fig 2.

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Fig 2.

Relative burdens of risk (A) and estimated illness (B) due to P. vivax malaria among the WHO regions: WPRO is Western Pacific; SEARO is Southeast Asia; EMRO is Eastern Mediterranean; AFRO is Sub-Saharan Africa; and PAHO is the Americas.

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The countries with the largest populations at risk are India (SEARO), Pakistan (EMRO), and Ethiopia (AFRO). In much of the P. vivax–endemic world, urban areas are not considered to be areas of transmission (though there may be incident infections due to relapses or travel). However, in and around the Indian subcontinent, urban areas were not excluded from the populations at risk because of the occurrence of Anopheles stephensi, a notoriously competent vector of malaria singularly able to thrive in urban settings [71]. Recent evidence demonstrates that An. stephensi also occurs on the Horn of Africa, including Ethiopia [113]. The range of this dangerously capable and adaptable vector thus includes a very substantial segment of the population living at risk with stable P. vivax transmission. Among the predicted 14.3 million cases in 2017, 15% were estimated to have been acquired in urban settings [3]. The near success but ultimate failure to eliminate malaria from India during the 1960s and 1970s has been attributed to the neglect of control in urban areas serving to reseed already cleared rural areas [114]. Plasmodium vivax malaria remains predominantly a disease of rural settings; of the 85% of cases occurring beyond urban centers, 64% (7.7 million) cases are estimated to have occurred in rural settings.

Burdens of infection and disease

Estimates of burden refer to the annual incidence of clinical (symptomatic) infections. The largest burden of these acute attacks occurs on the Indian subcontinent and the Horn of Africa [3], as shown in Fig 3. As discussed above, this metric may narrowly represent broader burdens of morbidity and mortality occurring in connection with chronic infection but more subtly than with a successfully diagnosed and reported event of clinical malaria. Chronic subpatency and latency are examples of this subtlety. While not every individual infected by P. vivax will experience multiple relapses, it very commonly occurs [30] with “remarkable periodicity,” which varies geographically [84]. The presence of the parasites in the liver is overdispersed—not every infection will lead to the formation of hypnozoites, but those individuals with hypnozoites are more likely to carry at least several more, and, absent specific therapy, will go on to experience multiple relapses within 2 or more years [35]. Infection with P. vivax in Indonesia, where P. vivax has a frequent and rapid relapse periodicity, has been shown to have chronic impacts on individuals leading to rehospitalizations with malaria and even early death [18]. In contrast, P. vivax in India tends to relapse infrequently and at long intervals after infection. Graphic representations of clinical burdens like that of Fig 3 may thus be insensitive to broader health impacts of endemic P. vivax malaria.

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Fig 3. National clinical burdens of P. vivax malaria.

The mean estimated clinical burdens of P. vivax malaria are shown on a scale of blue (very low burden) to red (high clinical burden) [3]. Global national shapefile obtained from the Malaria Atlas Project (MAP; https://malariaatlas.org/) and available for download through the malariaAtlas R package [109].

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What is true for the burden of clinical disease is also true for infection prevalence. Measured and modeled prevalence estimates like those in Fig 1 refer to infections patent by microscopy or RDT. Low-density subpatent infections are common with P. vivax, and, as already discussed, subpatency very likely includes infection of extravascular spaces of some deep organs. Standard diagnostic methods are unable to detect vascular or extravascular subpatency. That is also true of the vitally important hypnozoite reservoir. Prevalence of patent infection only narrowly represents broader and probably dominant states of infection, and conventional diagnostics-dependent estimates of burdens of infection are very likely to be minimally representative of true burdens of infection.

Credible vivax malaria diagnostics may require joining the list of human infections reliably diagnosed primarily or solely by serological means. Serological evidence of recent exposure to P. vivax could prompt therapy without regard to symptoms or conventional diagnostic outcomes. Serological surveys across sub-Saharan Africa, for example, suggested a true prevalence of P. vivax infection in the range of 11% to 60%, whereas by conventional diagnostics, it appears almost wholly absent [9496]. This contrast—highly prevalent versus absent—highlights the gravity of diagnostic approach in striving to assess the extent to which P. vivax infects human communities. Assessments of P. vivax global burdens of infection will likely require recalibrations based on validated serological diagnostic approaches. Work on those techniques is in progress [115,116].

Conclusions

Endemic P. vivax transmission occurs across the tropics and reaches into subtropical and temperate climates. Since 2000, the estimated number of patent P. vivax clinical cases as fallen from 24.5 (22.5 to 27.0 95% CI) million to 14.3 (13.7 to 15.0 95% CI) million in 2017. However, malaria control has seen stagnated progress since 2015, and this holds true for P. vivax specifically (Fig 4) [3,117]. There has been little change in the estimated burden of acute disease in recent years, and a few countries carry more than 80% of the global case load, i.e., India, Pakistan, and Ethiopia. Addressing case counts alone, however, overemphasizes densely populated endemic countries. Infection prevalence rates are highest in less populated parts of the world, likely driven by environmental suitability as well as high relapse rates, like those seen in Papua New Guinea and the Solomon Islands [83] as examples. Areas in South America, specifically Venezuela where prevalence rates have been increasing in recent years, also exhibit much higher infection rates than those seen in the countries with highest burden (Figs 1 and 3).

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Fig 4.

Global and region trends of P. vivax clinical incidence rate (A) and case counts (B) from 2000 to 2017 [3]. WPRO is Western Pacific; SEARO is Southeast Asia; EMRO is Eastern Mediterranean; AFRO is Sub-Saharan Africa; and PAHO is the Americas.

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Limited blood surveys for P. vivax malaria and the coarse resolution of routine surveillance data make estimating subnational patterns of prevalence and incidence challenging (Box 1). Existing data and model outputs indicate that P. vivax is primarily a rural disease affecting populations with limited access to effective treatment. Fine-scale estimates of the distribution of the burden of disease will continue to improve as routine data are made more readily available at the resolution at which they are collected, such as health facility or district. Finer resolution data also have the potential to disaggregate the populations most at risk of infection or clinical disease. Age-specific estimates for P. vivax, which are important for control planning and commodity forecasting, are currently derived from a model originally developed for P. falciparum [118] calibrated using data from Papua New Guinea and Indonesia [119]. Updated routine and clinical trial data would greatly increase the temporal and geographic coverage of age-specific data to inform regional P. vivax malaria age profiles. Robust models to standardize all-age metrics to age-specific results are imperative now that available treatment regimens may be tailored by age. Metrics indicating transmission or endemicity of P. vivax malaria are most commonly reported for all ages because of the challenges in detecting the parasite [120].

Box 1. Uncertainties in global malaria burden estimates

  1. Vascular patency is the basis of estimates of global burden of disease, but this is the minority state of infection by P. vivax in endemic zones where vascular subpatency, extravascular subpatency, sexual latency, and hepatic latency dominate.
  2. The parasitemia of P. vivax malaria is inherently lower than that in P. falciparum malaria, and it is therefore more often missed in peripheral blood examinations. Vivax malaria may often be considered a fever of unknown origin and treated presumptively.
  3. Innate and acquired immunity to P. vivax suppress or even prevent asexual parasitemia and acute illness, but the host remains infected and infectious. Geographic distributions of infection based on measurements of vascular patency and illness may disregard broad zones or even regions of endemic transmission. Duffy negativity in sub-Saharan Africa, for example, may deeply obscure endemic P. vivax transmission.
  4. The patent acute attack of malaria is the basis of global estimates of disease, but chronic malaria—be it repeated acute attacks borne of mosquitoes or hyponozoites, or persistant vascular or extravascular subpatency—causes more subtle morbidity and mortality that goes unaccounted.
  5. Estimates of the global burden of acute patent malaria certainly serve important surveillance goals, but those should not be misconstrued as representing all morbidity and mortality nor the number of infections or geographic distributions of endemic malaria transmission. The hard work of eliminating malaria will require maps of infection/infectiousness from new diagnostic technologies rather than simple vascular patency.
  6. Validated serological diagnosis of recent exposure to the plasmodia may prompt treatment and reporting as malaria without regard to vascular patency or illness. That surveillance may yield nearer-to-true prevalence and geographic distributions of infection by these insidious parasites.

The global burden of infection and disease imposed by endemic P. vivax transmission is obscured by its biology as both an active and latent infection seated in inaccessible tissues. Complex host genetic factors like Duffy factor negativity phenotype deepen that obscurity, and inherited G6PD deficiency may do likewise, i.e., by enhancing tropisms for those tissues. Likewise, due to the singular problem of 8-aminoquinoline toxicity and CYP2D6 dependency for activity against the latent hepatic infections, our ability to attack that important reservoir is deeply impaired. The insidious character of the harm done by P. vivax further sheltered this parasite from a determined assault upon it—for nearly 60 years, we neglected developing better diagnostics and more effective therapies [121124]. We thus inherited diagnostics, chemotherapies, and vector control strategies and tactics optimized and validated for an African P. falciparum problem that are conspicuously inadequate to endemic P. vivax anywhere. As endemic nations press the elimination agenda by plying those tools, P. falciparum wanes, but P. vivax has, unsurprisingly, proven more tenacious [124,125].

The conventional perspective of human malaria as an infection of peripheral blood obscures broader and more subtle burdens of P. vivax. The microscopic diagnostic standard for over a century (later joined by antigen capture immunochromatography) from finger stick blood specimens defined the presence or absence of infection. This standard remains firmly in place and underpins the global burden estimates described here, which, as a result, can only enumerate certain aspects of P. vivax burden (i.e., patent blood-stage infection and clinical/classically symptomatic cases). If infection beyond the vascular sinuses indeed dominates P. vivax biomass in any given human host, we may have to accept the inadequacy of conventional diagnostics to this biology and the epidemiology informed by it. Doing so opens promising avenues of alternative diagnostics like serology. Expanding the scope of burden estimation to consider (i) subpatent infection and (ii) indirect morbidity and mortality is important and firmly on the agenda as and when understanding improves and data allow for it.

Acknowledgments

The authors thank Peter Gething in Perth, Australia for his useful comments on the manuscript.

References

  1. 1. Institute for Health Metrics and Evaluation. Findings from the Global Burden of Disease Study 2017 [Internet]. Seattle, WA; 2018 [cited 2020 Oct 18]. Available from: www.healthdata.org.
  2. 2. World Health Organization. World Malaria Report 2015. Geneva; 2015. Available from: https://www.who.int/malaria/publications/world-malaria-report-2015/report/en/.
  3. 3. 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. pmid:31229233
  4. 4. Weiss DJ, Lucas TCD, Nguyen M, Nandi AK, Bisanzio D, Battle KE, et al. Mapping the global prevalence, incidence, and mortality of Plasmodium falciparum, 2000–17: a spatial and temporal modelling study. Lancet. 2019 Jul 27;394(10195):322–331. pmid:31229234
  5. 5. Bousema T, Okell L, Felger I, Drakeley C. Asymptomatic malaria infections: detectability, transmissibility and public health relevance. Nat Rev Microbiol. 2014;(October):1–8. pmid:25329408
  6. 6. Moreira CM, Abo-Shehada M, Price RN, Drakeley CJ. A systematic review of sub-microscopic Plasmodium vivax infection. Malar J. 2015 Sep 22;14:360. pmid:26390924
  7. 7. Doolan DL, Dobaño C, Baird JK. Acquired immunity to malaria. Clin Microbiol Rev. 2009;22:13–36. pmid:19136431
  8. 8. Chen I, Clarke SE, Gosling R, Hamainza B, Killeen G, Magill A, et al. “Asymptomatic” malaria: A chronic and debilitating infection that should be treated. PLoS Med. 2016;13(1):e1001942. pmid:26783752
  9. 9. de Mast Q, Brouwers J, Syafruddin D, Bousema T, Baidjoe AY, de Groot PG, et al. Is asymptomatic malaria really asymptomatic? Hematological, vascular and inflammatory effects of asymptomatic malaria parasitemia. J Infect. 2015 Nov 1;71(5):587–96. pmid:26304688
  10. 10. Peto TJ, Tripura R, Lee SJ, Althaus T, Dunachie S, Nguon C, et al. Association between subclinical malaria infection and inflammatory host response in a pre-elimination setting. PLoS ONE. 2016 Jul 1;11(7):e0158656. pmid:27386859
  11. 11. Snow RW, Marsh K. The epidemiology of clinical malaria among African children. Bull Inst Pasteur. 1998 Mar;96(1):15–23. pmid:23682192
  12. 12. McCreesh P, Mumbengegwi D, Roberts K, Tambo M, Smith J, Whittemore B, et al. Subpatent malaria in a low transmission African setting: A cross-sectional study using rapid diagnostic testing (RDT) and loop-mediated isothermal amplification (LAMP) from Zambezi region. Namibia Malar J. 2018 Dec 19;17(480).
  13. 13. Harris I, Sharrock WW, Bain LM, Gray KA, Bobogare A, Boaz L, et al. A large proportion of asymptomatic Plasmodium infections with low and sub-microscopic parasite densities in the low transmission setting of Temotu Province, Solomon Islands: Challenges for malaria diagnostics in an elimination setting. Malar J. 2010;9(1).
  14. 14. van Eijk AM, Sutton PL, Ramanathapuram L, Sullivan SA, Kanagaraj D, Priya GSL, et al. The burden of submicroscopic and asymptomatic malaria in India revealed from epidemiology studies at three varied transmission sites in India. Sci Rep. 2019 Dec 1;9(1). pmid:31745160
  15. 15. Hawash Y, Ismail K, Alsharif K, Alsanie W. Malaria prevalence in a low transmission area, Jazan district of southwestern Saudi Arabia. Korean J Parasitol. 2019 Jun 1;57(3):233–42. pmid:31284345
  16. 16. Baum E, Sattabongkot J, Sirichaisinthop J, Kiattibutr K, Jain A, Taghavian O, et al. Common asymptomatic and submicroscopic malaria infections in Western Thailand revealed in longitudinal molecular and serological studies: a challenge to malaria elimination. Malar J. 2016;15 (1):333. pmid:27333893
  17. 17. Rosas-Aguirre A, Gamboa D, Manrique P, Conn JE, Moreno M, Lescano AG, et al. Epidemiology of Plasmodium vivax malaria in Peru. Am J Trop Med Hyg 2016;95(6 Suppl):133–44. pmid:27799639
  18. 18. Dini S, Douglas NM, Poespoprodjo JR, Kenangalem E, Sugiarto P, Plumb ID, et al. The risk of morbidity and mortality following recurrent malaria in Papua, Indonesia: A retrospective cohort study. BMC Med. 2020 Feb 20;18(1):28. pmid:32075649
  19. 19. Da Silva Junior GB, Pinto JR, Barros EJG, Farias GMN, Daher EDF. Kidney involvement in malaria: An update. Rev Inst Med Trop Sao Paulo. 2017;59:e53. pmid:28793022
  20. 20. Langford S, Douglas NM, Lampah DA, Simpson JA, Kenangalem E, Sugiarto P, et al. Plasmodium malariae infection associated with a high burden of anemia: A hospital-based surveillance study. PLoS Negl Trop Dis. 2015 Dec 31;9(12):e0004195. pmid:26720002
  21. 21. Douglas NM, Lampah DA, Kenangalem E, Simpson JA, Poespoprodjo JR, Sugiarto P, et al. Major burden of severe anemia from non-falciparum malaria species in southern Papua: A hospital-based surveillance study. PLoS Med. 2013;10(12):1–17. pmid:24358031
  22. 22. Imtiaz S, Drohlia MF, Nasir K, Hussain M, Ahmad A. Morbidity and mortality associated with Plasmodium vivax and Plasmodium falciparum infection in a tertiary care kidney hospital. Saudi J Kidney Dis Transpl. 2015 Nov 1;26(6):1169–76. pmid:26586055
  23. 23. Brasil LMBF Vieira JLF, Araújo EC Piani PPF, Dias RM Ventura AMRS, et al. Cognitive performance of children living in endemic areas for Plasmodium vivax. Malar J. 2017 Sep 12;16(1). pmid:28899387
  24. 24. Fernando D, De Silva D, Carter R, Mendis KN, Wickremasinghe R. A randomized, double-blind, placebo-controlled, clinical trial of the impact of malaria prevention on the educational attainment of school children. Am J Trop Med Hyg. 2006;74(3):386–93. pmid:16525095
  25. 25. van den Eede P, Soto-Calle VE, Delgado C, Gamboa D, Grande T, Rodriguez H, et al. Plasmodium vivax sub-patent infections after radical treatment are common in Peruvian patients: Results of a 1-year prospective cohort study. PLoS ONE. 2011;6(1). pmid:21297986
  26. 26. Kho S, Marfurt J, Handayuni I, Pava Z, Noviyanti R, Kusuma A, et al. Characterization of blood dendritic and regulatory T cells in asymptomatic adults with sub-microscopic Plasmodium falciparum or Plasmodium vivax infection. Malar J. 2016 Jun;21:15(1). pmid:27328659
  27. 27. Van Nguyen H, Van Den Eede P, Van Overmeir C, Thang ND, Hung LX, D’Alessandro U, et al. Marked age-dependent prevalence of symptomatic and patent infections and complexity of distribution of human Plasmodium species in central Vietnam. Am J Trop Med Hyg. 2012 Dec;87(6):989–95. pmid:23128294
  28. 28. Sutanto I, Kosasih A, Elyazar IRF, Simanjuntak DR, Larasati TA, Dahlan MS, et al. Negligible impact of mass screening and treatment on mesoendemic malaria transmission at west Timor in eastern Indonesia: A cluster-randomized trial. Clin Infect Dis. 2018 Oct 15;67(9):1364–72. pmid:29579195
  29. 29. Tiburskaja NA, Sergiev PG, Vrublevskaja OS. Dates of onset of relapses and the duration of infection in induced tertian malaria with short and long incubation periods. Bull World Health Organ. 1968;38(3):447–57. pmid:4876427
  30. 30. White NJ. Determinants of relapse periodicity in Plasmodium vivax malaria. Malar J. 2011;10:297. pmid:21989376
  31. 31. Krotoski WA. The hypnozoite and malarial relapse. Prog Clin Parasitol. 1989. pmid:2491691
  32. 32. Ja Lysenko. A, Beljaev AE, Rybalka VM. Population studies of Plasmodium vivax. I. The theory of polymorphism of sporozoites and epidemiological phenomena of tertian malaria. Bull World Health Organ. 1977;55(5):541–9. pmid:338188
  33. 33. Robinson LJ, Wampfler R, Betuela I, Karl S, White MT, Li Wai Suen CSN, et al. Strategies for understanding and reducing the Plasmodium vivax and Plasmodium ovale hypnozoite reservoir in Papua New Guinean children: A randomised placebo-controlled trial and mathematical model. PLoS Med. 2015;12(10):e1001891. pmid:26505753
  34. 34. Adekunle AI, Pinkevych M, Mcgready R, Luxemburger C. Modeling the dynamics of Plasmodium vivax infection and hypnozoite reactivation in vivo. PLoS Negl Trop Dis. 2015;9(3):e0003595. pmid:25780913
  35. 35. White MT, Karl S, Battle KE, Hay SI, Mueller I, Ghani AC. Modelling the contribution of the hypnozoite reservoir to Plasmodium vivax transmission. Elife. 2014 Nov 18;3(November):1–19. pmid:25406065
  36. 36. Baird JK. Evidence and implications of mortality associated with acute Plasmodium vivax malaria. Clin Microbiol Rev. 2013 Jan;26(1):36–57. pmid:23297258
  37. 37. Barber BE, William T, Grigg MJ, Parameswaran U, Piera KA, Price RN, et al. Parasite biomass-related inflammation, endothelial activation, microvascular dysfunction and disease severity in vivax malaria. PLoS Pathog. 2015;11(1):1–13. pmid:25569250
  38. 38. Obaldia N, Meibalan E, Sa JM, Ma S, Clark MA, Mejia P, et al. Bone marrow is a major parasite reservoir in Plasmodium vivax infection. mBio. 2018 May 1;9(3):e00625–18. pmid:29739900
  39. 39. Brito MAM, Baro B, Raiol TC, Ayllon-Hermida A, Safe IP, Deroost K, et al. Morphological and transcriptional changes in human bone marrow during natural Plasmodium vivax malaria infections. J Infect Dis. 2020. pmid:32556188
  40. 40. Kho S, Qotrunnada L, Leonardo B, Wardani PAI, Fricot A, Henry B, et al. Hidden biomass of intact malaria parasites in the human spleen. N Engl J Med. 2021;384:2067–9. pmid:34042394
  41. 41. Kho S, Qotrunnada L, Leonardo L, Andries B, Wardani PAI, Fricot A, et al. Evaluation of splenic accumulation and colocalization of immature reticulocytes and Plasmodium vivax in asymptomatic malaria: a prospective human splenectomy study. PLoS Med. 2021;18:e1003632. pmid:34038413
  42. 42. Silva-Filho JL, Lacerda MVG, Recker M, Wassmer SC, Marti M, Costa FTM. Plasmodium vivax in hematopoietic niches: hidden and dangerous. Trends Parasitol. 2020;36:447–58. pmid:32298632
  43. 43. Siquiera AM, Magalhaes BML, Melo GC, Ferrer M, Castillo P, Martin-Jaular L, et al. Spleen rupture in a case of untreated Plasmodium vivax infection. PLoS Negl Trop Dis. 2012;6:e1934. pmid:23272256
  44. 44. Fernandez-Becerra C, Bernabeu M, Castellanos A, Correa BR, Ramirez M, Rui E, et al. Plasmodium vivax spleen-dependent genes encode antigens associated with cytoadhesion and clinical protection. Proc Natl Acad Sci U S A. 2020;117:13056–65. pmid:32439708
  45. 45. 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 Jan 5;359(6371):48–55. pmid:29302006
  46. 46. 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 Feb 19;125(8):1314–24. pmid:25414440
  47. 47. Lee RS, Waters AP, Brewer JM. A cryptic cycle in haemoatopoietic niches promotes initiation of malaria transmission and evasion of chemotherapy. Nat Commun. 2018;9:1689. pmid:29703959
  48. 48. Toda H, Diaz-Varela M, Segui-Barber J, Roobsoon W, Baro B, Garcia-Silva S, et al. Plasma-derived extracellular vesicles from Plasmodium vivax patients signal spleen fibroblasts via NF-kB facilitating parasite cytoadherence. Nat Commun. 2020;11:2761. pmid:32487994
  49. 49. Miller LH, Mason SJ, Dvorak JA, Mcginniss MH, Rothman IK. Erythrocyte receptors for (Plasmodium knowlesi) malaria: Duffy blood group determinants. Science. 1975 Aug 15;189(4202):561–3. pmid:1145213
  50. 50. Elliott JH, O’Brien D, Leder K, Kitchener S, Schwartz E, Weld L, et al. Imported Plasmodium vivax malaria: Demographic and clinical features in nonimmune travelers. J Travel Med. 2004;11(4):213–9. pmid:15541223
  51. 51. He X, Pan M, Zeng W, Zou C, Pi L, Qin Y, et al. Multiple relapses of Plasmodium vivax malaria acquired from West Africa and association with poor metabolizer CYP2D6 variant: A case report. BMC Infect Dis. 2019 Aug 9;19(1). pmid:31399061
  52. 52. Feng J, Xiao H, Zhang L, Yan H, Feng X, Fang W, et al. The Plasmodium vivax in China: Decreased in local cases but increased imported cases from Southeast Asia and Africa. Sci Rep. 2015 Mar 5;5. pmid:25739365
  53. 53. 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. pmid:29530446
  54. 54. Leslie T, Briceño M, Mayan I, Mohammed N, Klinkenberg E, Sibley CH, et al. The impact of phenotypic and genotypic G6PD deficiency on risk of Plasmodium vivax infection: A case-control study amongst Afghan refugees in Pakistan. PLoS Med. 2010 May;7(5):e1000283. pmid:20520804
  55. 55. Yi H, Li H, Liang L, Wu Y, Zhang L, Qiu W, et al. The glucose-6-phosphate dehydrogenase Mahidol variant protects against uncomplicated Plasmodium vivax infection and reduces disease severity in a Kachin population from northeast Myanmar. Infect Genet Evol. 2019 Nov 1;75:103980. pmid:31351234
  56. 56. Santana MS, de Lacerda MVG, Barbosa MDGV, Duarte Alecrim W, Costa Alecrim MDG. Glucose-6-phosphate dehydrogenase deficiency in an endemic area for malaria in Manaus: A cross-sectional survey in the Brazilian Amazon. PLoS ONE. 2009 Apr 16;4(4):e5259. pmid:19370159
  57. 57. Louicharoen C, Patin E, Paul R, Nuchprayoon I, Witoonpanich B, Peerapittayamongkol C, et al. Positively selected G6PD-mahidol mutation reduces Plasmodium vivax density in Southeast Asians. Science. 2009 Dec 11;326(5959):1546–9. pmid:20007901
  58. 58. Howes RE, Battle KE, Satyagraha AW, Baird JK, Hay SI. G6PD deficiency. Global distribution, genetic variants and primaquine therapy. Adv Parasitol. 2013;81:133–201. pmid:23384623
  59. 59. Baird JK, Louisa M, Noviyanti R, Ekawati L, Subekti D, Chand K, et al. Association of impaired cytochrome P450 2D6 activity henotype and phenotype with therapeutic efficacy of primaquine treatment for latent Plasmodium vivax malaria. JAMA Netw Open. 2018;1(4):181449.
  60. 60. Marcsisin SR, Reichard G, Pybus BS. Primaquine pharmacology in the context of CYP 2D6 pharmacogenomics: Current state of the art. Pharmacol Ther. 2016;161:1–10. pmid:27016470
  61. 61. Brito-Sousa JD, Santos TC, Avalos S, Fontecha G, Melo GC, Val F, et al. Clinical spectrum of primaquine-induced hemolysis in glucose-6-phosphate dehydrogenase deficiency: A 9-year hospitalization-based study from the Brazilian Amazon. Clin Infect Dis. 2019 Sep 27;69(8):1440–2. pmid:30753364
  62. 62. Recht J, Ashley EA, White NJ. Use of primaquine and glucose-6-phosphate dehydrogenase deficiency testing: Divergent policies and practices in malaria endemic countries. PLoS Negl Trop Dis. 2018 Apr 19;12(4): e0006230. pmid:29672516
  63. 63. Spring MD, Lon C, Sok S, Sea D, Wojnarski M, Chann S, et al. Prevalence of CYP2D6 genotypes and predicted phenotypes in a cohort of Cambodians at high risk for infections with Plasmodium vivax. Am J Trop Med Hyg. 2020 Aug 1;103(2):756–9. pmid:32394887
  64. 64. Newby G, Bennett A, Larson E, Cotter C, Shretta R, Phillips AA, et al. The path to eradication: A progress report on the malaria-eliminating countries. Lancet. 2016;387(10029):1775–84. pmid:27116283
  65. 65. Shretta R, Silal SP, Celhay OJ, Gran Mercado CE, Kyaw SS, Avancena A, et al. Malaria elimination transmission and costing in the Asia-Pacific: Developing an investment case. Wellcome Open Res. 2020;4.
  66. 66. Herrera S, Ochoa-Orozco SA, González IJ, Peinado L, Quiñones ML, Arévalo-Herrera M. Prospects for malaria elimination in Mesoamerica and Hispaniola. PLoS Negl Trop Dis. 2015 May 14;9(5):e0003700.
  67. 67. Battle KE, Gething PW, Elyazar IRF, Moyes CL, Sinka ME, Howes RE, et al. The global public health significance of Plasmodium vivax. Adv Parasitol. 2012;80:1–111. pmid:23199486
  68. 68. Coatney GR, Collins WE, Warren M, Contacos PG. The Primate Malarias. Bethesda, MD: National Institute of Health, National Institute of Allergy and Infectious Diseases; 1971. p. 366.
  69. 69. Sinka ME, Golding N, Massey NC, Wiebe A, Huang Z, Hay SI, et al. Modelling the relative abundance of the primary African vectors of malaria before and after the implementation of indoor, insecticide-based vector control. Malar J. 2016;15(1):142.
  70. 70. Sinka ME, Bangs MJ, Manguin S, Coetzee M, Mbogo CM, Hemingway J, et al. The dominant Anopheles vectors of human malaria in Africa, Europe and the Middle East: Occurrence data, distribution maps and bionomic précis. Parasit Vectors. 2010 Dec 3;3(1):1–34. pmid:20051120
  71. 71. Sinka ME, Bangs MJ, Manguin S, Chareonviriyaphap T, Patil AP, Temperley WH, et al. The dominant Anopheles vectors of human malaria in the Asia-Pacific region: Occurrence data, distribution maps and bionomic précis. Parasit Vectors. 2011 May 25;4:89. pmid:21612587
  72. 72. Ho DT, Van Bortel W, Sochantha T, Keokenchanh K, Briët OJT, Coosemans M. Behavioural heterogeneity of Anopheles species in ecologically different localities in Southeast Asia: A challenge for vector control. Trop Med Int Health. 2005 Mar 1;10(3):251–62. pmid:15730510
  73. 73. Baird KJ, Maguire JD, Price RN. Diagnosis and treatment of Plasmodium vivax malaria. Adv Parasitol. 2012; 80:203–70. pmid:23199489
  74. 74. Smithuis FM, Kyaw MK, Phe UO, van der Broek I, Katterman N, Rogers C, et al. The effect of insecticide-treated bed nets on the incidence and prevalence of malaria in children in an area of unstable seasonal transmission in western Myanmar. Malar J. 2013 Oct;12:363. pmid:24119916
  75. 75. Smithuis FM, Kyaw MK, Phe UO, Van Der Broek I, Katterman N, Rogers C, et al. Entomological determinants of insecticide-treated bed net effectiveness in Western Myanmar. Malar J. 2013;12(1). pmid:24119994
  76. 76. Hii J, Rueda LM. Malaria vectors in the Greater Mekong Subregion: overview of malaria vectors and remaining challenges. Southeast Asian J Trop Med Public Health. 2013;44(Suppl 1):73–165; discussion 306–7. 76. pmid:24159831
  77. 77. 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. pmid:19695492
  78. 78. Schafer C, Roobsoon W, Kangwanrangsan N, Bardelli M, Rawlinson TA, Dambrauskas N, et al. A humanized mouse model for Plasmodium vivax to test interventions that block liver stage to blood stage transition and blood stage infection. iScience. 2020;23:101381. pmid:32739836
  79. 79. Kitchen SF. Malariology: a comprehensive survey of all aspects of this group of diseases from a global standpoint. In: Boyd MF, editor. vol. II. Philadelphia, PA: W.B. Saunders Ltd; 1949. p. 966–1045.
  80. 80. Barber BE, William T, Grigg MJ, Yeo TW, Anstey NM. Limitations of microscopy to differentiate Plasmodium species in a region co-endemic for Plasmodium falciparum, Plasmodium vivax and Plasmodium knowlesi. Malar J. 2013 Jan 8;12:8 pmid:23294844
  81. 81. Steenkeste N, Incardona S, Chy S, Duval L, Ekala M-T, Lim P, et al. Towards high-throughput molecular detection of Plasmodium: new approaches and molecular markers. Malar J. 2009;8:86. pmid:19402894
  82. 82. Imwong M, Stepniewska K, Tripura R, Peto TJ, Lwin KM, Vihokhern B, et al. Numerical distributions of parasite densities during asymptomatic malaria. J Infect Dis. 2015;jiv596. pmid:26681777
  83. 83. Cheng Q, Cunningham J, Gatton ML. Systematic review of sub-microscopic P. vivax infections: Prevalence and determining factors. PLoS Negl Trop Dis. 2015;9(1). pmid:25569135
  84. 84. Battle KE, Karhunen MS, Bhatt S, Gething PW, Howes RE, Golding N, et al. Geographical variation in Plasmodium vivax relapse. Malar J. 2014 Apr 15;13:144. pmid:24731298
  85. 85. Commons RJ, Simpson JA, Watson J, White NJ, Price RN. Estimating the proportion of Plasmodium vivax recurrences caused by relapse: A systematic review and meta-analysis. Am J Trop Med Hyg. 2020 Sep 1;103(3):1094–9. pmid:32524950
  86. 86. 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(1). pmid:21468018
  87. 87. Twohig KA, Pfeffer DA, Baird JK, Price RN, Zimmerman PA, Hay SI, et al. Growing evidence of Plasmodium vivax across malaria-endemic Africa. Lacerda MV, editor. PLoS Negl Trop Dis. 2019 Jan 31;13(1):e0007140. pmid:30703083
  88. 88. Rosenberg R. Plasmodium vivax in Africa: hidden in plain sight? Trends Parasitol. 2007;23(5):193–6. pmid:17360237
  89. 89. Brazeau NF, Whitesell AN, Doctor SM, Keeler C, Mwandagalirwa MK, Tshefu AK, et al. Plasmodium vivax infections in Duffy-negative individuals in the Democratic Republic of the Congo. Am J Trop Med Hyg. 2018 Nov 1;99(5):1128–33. pmid:30203741
  90. 90. Zimmerman PA. Plasmodium vivax infection in Duffy-negative people in Africa. Am J Trop Med Hyg. 2017 Sep;97(3):636–638. pmid:28990906
  91. 91. 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 Apr 1;110(4):258–60. pmid:27076512
  92. 92. Woldearegai TG, Kremsner PG, Kun JFJ, Mordmü ller B. Plasmodium vivax malaria in Duffy-negative individuals from Ethiopia. Trans R Soc Trop Med Hyg. 2013 May;107(5):328–31. pmid:23584375
  93. 93. 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 Feb 14;16(1):1–9. pmid:28049519
  94. 94. Ménard 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 U S A. 2010 Mar 30;107(13):5967–71. pmid:20231434
  95. 95. Culleton R, Ndounga M, Zeyrek FY, Coban C, Casimiro PN, Takeo S, et al. Evidence for the transmission of Plasmodium vivax in the Republic of the Congo, West Central Africa J Infect Dis. 2009 Nov;200(9):1465–9. pmid:19803728
  96. 96. Poirier P, Doderer-Lang C, Atchade PS, Lemoine JP, De L’Isle MLC, 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 Nov 25;15(1). pmid:27887647
  97. 97. Niang M, Diop F, Niang O, Sadio BD, Sow A, Faye O, et al. Unexpected high circulation of Plasmodium vivax in asymptomatic children from Kedougou, southeastern Senegal. Malar J. 2017;16:497. pmid:29284488
  98. 98. Dechavanne C, Dechavanne S, Metral S, Roeper B, Krishnan S, Fong R, et al. Duffy antigen expression in erythroid bone marrow precursor cells of genotypically Duffy negative individuals. bioRxiv. 2018:508481.
  99. 99. Howes RE, Piel FB, Patil AP, Nyangiri O, Gething PW, Dewi M, et al. G6PD deficiency prevalence and estimates of affected populations in malaria endemic countries: A geostatistical model-based map. PLoS Med. 2012;9(11). pmid:23152723
  100. 100. Luzzatto L, Seneca E. G6PD deficiency: A classic example of pharmacogenetics with on-going clinical implications. Br J Haematol. 2014;164:469–80. pmid:24372186
  101. 101. WHO Working Group. Glucose-6-phosphate dehydrogenase deficiency. Bull World Health Organ. 1989;67(6):601–11. pmid:2633878
  102. 102. Piomelli S, Corash LM, Davenport DD, Miraglia J, Amorosi EL. In vivo lability of glucose-6-phosphate dehydrogenase in GdA- and GdMediterranean deficiency. J Clin Invest. 1968;47(4):940–8. pmid:5641629
  103. 103. Siniscalco M, Bernini L, Filippi G, Latte B, Meera Khan P, Piomelli S, et al. Population genetics of haemoglobin variants, thalassaemia and glucose-6-phosphate dehydrogenase deficiency, with particular reference to the malaria hypothesis. Bull World Health Organ. 1966;34(3):379–93. pmid:5296398
  104. 104. Awab GR, Aaram F, Jamornthanyawat N, Suwannasin K, Pagornat W, Watson JA, et al. Protective effect of Mediterranean-type glucose-6-phosphate dehydrogenase deficiency against Plasmodium vivax malaria. Elife. 2021;10:e62448. pmid:33543710
  105. 105. Bancone G, Malleret B, Suwanarusk R, Chowwiwat N, Chu CS, McGready R, et al. Asian G6PD-mahidol reticulocytes sustain normal Plasmodium vivax development. J Infect Dis. 2017 Jul 15;216(2):263–6. pmid:28591790
  106. 106. Howes RE, Dewi M, Piel FB, Monteiro WM, Battle KE, Messina JP, et al. Spatial distribution of G6PD deficiency variants across malaria-endemic regions. Malar J. 2013;12(1):418. pmid:24228846
  107. 107. 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 Oct 3;369(14):1381–2. pmid:24088113
  108. 108. Baird JK, Battle KE, Howes RE. Primaquine ineligibility in anti-relapse therapy of Plasmodium vivax malaria: The problem of G6PD deficiency and cytochrome P-450 2D6 polymorphisms. Malar J. 2018 Jan 22;17(1):42.
  109. 109. Pfeffer D, Lucas T, May D, Haris J, Rozier J, Twohig KA, et al. « malariaAtlas: an R interface to global malariometric data hosted by the Malaria Atlas Project. Malar J. 2018;17:352. pmid:30290815
  110. 110. Maldonado YA, Nahlen BL, Roberto RR, Ginsberg M, Orellana E, Mizrahi M, et al. Transmission of Plasmodium vivax malaria in San Diego County, California, 1986. Am J Trop Med Hyg. 1990;42(1):3–9. pmid:1967916
  111. 111. Robert LL, Santos-Ciminera PD, Andre RG, Schultz GW, Lawyer PG, Nigro J, et al. Plasmodium-infected Anopheles mosquitoes collected in Virginia and Maryland following local transmission of Plasmodium vivax malaria in Loudoun County, Virginia. J Am Mosq Control Assoc. 2005 Jun;21(2):187–93. pmid:16033121
  112. 112. Andriopoulos P, Economopoulou A, Spanakos G, Assimakopoulos G. A local outbreak of autochthonous Plasmodium vivax malaria in Laconia, Greece-a re-emerging infection in the southern borders of Europe? Int J Infect Dis. 2013 Feb;17(2). pmid:23098813
  113. 113. Sinka ME, Pironon S, Massey NC, Longbottom J, Hemingway J, Moyes CL, et al. A new malaria vector in Africa: Predicting the expansion range of Anopheles stephensi and identifying the urban populations at risk. Proc Natl Acad Sci. 2020 Sep 14;117(40):202003976. pmid:32929020
  114. 114. Baird JK. Essential guidance on malaria elimination in its history. J Vector Borne Dis. 2019;56:11–14. pmid:31070160
  115. 115. Longley RJ, White MT, Takashima E, Brewster J, Morita M, Harbers M, et al. Development and validation of serological markers for detecting recent Plasmodium vivax infection. Nat Med. 2020;26:741–9. pmid:32405064
  116. 116. Auburn S, Cheng Q, Marfurt J, Price RN. The changing epidemiology of Plasmodium vivax: insights from conventional and novel surveillance tools. 2021 Apr 23;18(4):e1003560. pmid:33891580
  117. 117. WHO. World Malaria Report 2020. Vol. WHO/HTM/GM, World Health. 2020. Available from: https://www.who.int/publications/i/item/9789240015791.
  118. 118. Smith DL, Guerra CA, Snow RW, Hay SI. Standardizing estimates of the Plasmodium falciparum parasite rate. Malar J. 2007 Sep 25;6:131.
  119. 119. Gething PW, Elyazar IRF, Moyes CL, Smith DL, Battle KE, Guerra CA, et al. A long neglected world malaria map: Plasmodium vivax endemicity in 2010. Carlton JM, editor. PLoS Negl Trop Dis. 2012 Sep 6;6(9):e1814. pmid:22970336
  120. 120. Howes RE, Battle KE, Mendis KN, Smith DL, Cibulskis RE, Baird JK, et al. Global epidemiology of Plasmodium vivax. Am J Trop Med Hyg. 2016 Dec 28;95(6 Suppl):15–34. pmid:27402513
  121. 121. Mendis K, Sina BJ, Marchesini P, Carter R. The neglected burden of Plasmodium vivax malaria. Am J Trop Med Hyg. 2001 Jan-Feb;64(1–2 Suppl):97–106. pmid:11425182
  122. 122. Price RN, Tjitra E, Guerra CA, Yeung S, White NJ, Anstey NM. Vivax malaria: Neglected and not benign. Am J Trop Med Hyg. 2007;77(SUPPL. 6):79–87. pmid:18165478
  123. 123. Baird K. Origins and implications of neglect of G6PD deficiency and primaquine toxicity in Plasmodium vivax malaria. Pathog Glob Health. 2015;109(3):93–106. pmid:25943156
  124. 124. Price RN, Commons RJ, Battle KE, Thriemer K, Mendis K. Plasmodium vivax in the era of the shrinking P. falciparum map. Trends Parasitol. 2020 Jun;36(6):560–570. pmid:32407682
  125. 125. Seidlein L, White NJ. Taking on Plasmodium vivax malaria: a timely and important challenge. PLoS Med. 2021;18:e1003593. pmid:33891584