Open access peer-reviewed chapter

Challenges in the Control and Elimination of Plasmodium vivax Malaria

Written By

Puji BS Asih, Din Syafruddin and John Kevin Baird

Submitted: 05 July 2017 Reviewed: 09 April 2018 Published: 18 July 2018

DOI: 10.5772/intechopen.77082

From the Edited Volume

Towards Malaria Elimination - A Leap Forward

Edited by Sylvie Manguin and Vas Dev

Chapter metrics overview

1,943 Chapter Downloads

View Full Metrics

Abstract

The human malaria parasite Plasmodium vivax imposes unique challenges to its control and elimination. Primary among those is the hypnozoite reservoir of infection in endemic communities. It is the dominant source of incident malaria and exceedingly difficult to attack due to both inability to diagnose latent carriers and the potentially life-threatening toxicity of primaquine in patients with an inborn deficiency of G6PD, the only therapeutic option against hypnozoites. Large segments of endemic populations are not eligible for primaquine, and alternative strategies for managing the threat of relapse in any group have not been optimized or validated. Association of risk of primaquine failure against latent P. vivax with impaired alleles of P450 2D6 exacerbates the substantial pool of primaquine ineligibles. Resistance to chloroquine against acute P. vivax malaria commonly occurs; alternative therapies like ACTs are effective but seldom evaluated as a partner drug to primaquine in the essential radical cure. Many of the Anopheles mosquito vector of P. vivax in South and Southeast Asia, where >90% of infections occur, thrive in a diversity of habitats and exhibit wide ranges of feeding and breeding behavior. This chapter explores many of these challenges and possible approaches in controlling and eliminating endemic vivax malaria.

Keywords

  • Plasmodium vivax
  • malaria
  • latent malaria
  • hypnozoites
  • glucose-6-phosphtate dehydrogenase (G6PD)
  • Anopheles mosquitoes
  • species-sanitation
  • control

1. Introduction

Human malaria caused by Plasmodium vivax currently has the widest geographical distribution among all malaria parasites with about 35% of the world population living at risk of this physically debilitating and sometime lethal infection [1, 2, 3]. Figure 1 illustrates this global distribution most heavily weighing upon South and Southeast Asia (SEA) [1]. In most endemic countries, chloroquine (CQ) remains the first-line therapy for acute vivax malaria after more than 70 years of continuous use. CQ-resistant P. vivax, documented nearly 30 years ago, now commonly occurs across much of SEA [4, 5]. Unlike the other dominant species causing human malaria, P. falciparum, some sporozoites (called bradysporozoites) of P. vivax develop into dormant forms in the liver called hypnozoites. This single feature—latency—defines and distinguishes the prevention, treatment, and control of vivax malaria. Other sporozoites (called tachysporozoites) immediately develop into actively dividing hepatic schizonts over the 7-day to 18-day incubation period and cause the primary parasitemia and acute attack of patent vivax malaria. Hypnozoites activate weeks, months, or even years later, causing a renewed clinical attack called relapses [6].

Figure 1.

Distribution of Plasmodium vivax malaria in the world.

In natural endemic settings, it may not be known if any given patient presenting with patent acute vivax malaria is experiencing a tachysporozoite-borne primary attack or a bradysporozoite-borne relapse. This uncertainty poses a fundamental problem of interpretation of parasitemia that may follow therapy [5, 6, 7, 8]. The origin of the parasitemia may be a consequence of new primary attack (reinfection), therapeutic failure against blood stages (recrudescence), or renewed latent malaria (relapse). These ambiguities may not be addressed by molecular genotyping techniques because relapses may be either homologous or heterologous to the primary infection event [9, 10]. Estimating the efficacy of blood schizonticidal therapy may thus be complex and difficult [11].

Another drug, a hypnozoitocide, is necessary to treat latent vivax malaria and prevent future attacks. Primaquine (PQ) has been the only available therapeutic option to kill hypnozoites since 1952. A single dose of 30 mg PQ within 48 hours of infection appears sufficient to kill stages of P. vivax or P. falciparum attempting to develop into hepatic schizonts or hypnozoites [12]. Beyond that period, presumably after formation of dormant hypnozoites, relatively large doses totaling 210 to 420 mg of PQ (delivered over 7 to 56 days) are required to prevent relapse [13]. Pharmacokinetic or pharmacodynamic interaction of blood schizonticidal and hypnozoitocidal therapies combined for the radical cure of vivax malaria has been observed and requires consideration in assessing the safety and efficacy of either or both in clinical use [14].

This chapter reviews the challenges vivax malaria poses in efforts to control and eliminate malaria in accordance with the Global Technical Strategy of the WHO [15] as they occur in many parts of the world. Experts advising the WHO formulated “Plasmodium vivax Control and Elimination: A Technical Brief” [16] highlighted the distinct character of this species in the context of control and elimination strategy. Conventional control aimed at diagnosing and treating of the acute attack and minimizing exposure to biting Anopheles mosquitoes will not suffice, largely due to the scale and importance of the latent hypnozoite reservoir in endemic communities. Decades of scientific, clinical, and public health neglect of this specific feature of vivax malaria leaves us poorly equipped to attack it safely and effectively. The biological basis of this problem is detailed with the aim of guiding discovery and development of sustainable solutions.

Advertisement

2. Biology of Plasmodium vivax

The broad and prolonged neglect of research on P. vivax has been highlighted by many researchers [17, 18, 19, 20]. Although this lack of research certainly derives from complex and multiple factors, the misperception of this species as intrinsically benign perhaps dominates among them [21]. Today, we accept that a diagnosis of vivax malaria is sometimes associated with severe disease syndromes associated with fatal outcomes [16, 22]. The manner in which P. vivax threatens life with such typically low-grade parasitemias (usually tenfold lower than P. falciparum) is an important and relatively new question. Nonetheless, some researchers suggest that vivax malaria may be primarily an infection of hematopoietic tissues rather than of the vascular sinuses per se [21, 23, 24]. If most P. vivax biomass in the human host resides within tissues of the bone marrow and spleen, it would have far-reaching scientific, clinical, and public health implications with respect to measuring and combatting the threats imposed.

We already know the likely importance of the latent hypnozoite reservoir and sub-patent/asymptomatic parasitemias [25, 26, 27]. Adding an as-yet unacknowledged sequestered trophozoite reservoir—very few or no asexual parasites in vascular sinuses but many in the extravascular spaces of erythropoietic tissues—would greatly amplify concerns regarding the effectiveness of diagnosis and treatment in control and elimination. It is possible that most P. vivax parasites—certainly hypnozoites but perhaps also trophozoites—occur beyond the vascular sinuses in both asymptomatic and acutely ill patients and, therefore, also beyond the reach of standard diagnostics.

Although the Duffy antigen on the surface of the red blood cell has long been considered essential to P. vivax invasion—and its absence in many African populations thought to explain the relative rarity of P. vivax on that continent—recent evidence from a variety of African locales has shown patent P. vivax parasitemia in patients who are negative for that molecule [28]. Moreover, P. vivax has been shown to be present in parts of Africa where it is not prevalent [29] and is indeed prevalent in other areas of that continent like Madagascar, the Horn, and across the northern Sahel [30].

Advertisement

3. Chloroquine-resistant acute P. vivax

Resistance to CQ by the asexual stages of P. vivax has been documented in most endemic regions [4, 5]. Resistant strains dominate the malarious Western Pacific and Indonesian archipelago and nations there have adopted highly efficacious ACTs [11] as first-line therapy. With the possible exception of artesunate combined with sulfadoxine-pyrimethamine, all ACTs have shown superb efficacy in killing asexual blood stages of P. vivax [31]. The safety and efficacy of PQ against relapse when combined with partner blood schizonticides other than CQ, quinine, or dihydroartemisinin-piperaquine [32, 33] require validation in clinical trials [14]. Elsewhere, for now, resistance appears sporadically and at relatively low frequencies. Despite substantial efforts to identify molecular markers of P. vivax resistance to antimalarial drugs, none have yet been validated. In vivo testing in patients or relatively difficult ex vivo drug testing procedures remain necessary [34]. The monitoring of antimalarial efficacy offers possible relief from risk of failure due to parasite resistance to specific therapies, but this is carried out relatively infrequently.

Advertisement

4. Latent and sub-patent P. vivax

The latent and sub-patent parasitemia caused by P. vivax is difficult or impractical to detect using available technologies. These unnoticed or invisible infections probably represent a dominant majority in most endemic settings. Thus, the primary blow to therapeutic effectiveness (the proportion of patients needing a particular therapy and receiving high-quality drug in a full and adequately absorbed dose) is simply the inability to identify those in need of therapy.

The human host also imposes important barriers to the effectiveness of antimalarial therapies in the real world. Clinical contraindications, patient adherence, provider prescribing practices, provider and patient access to the drug, and its quality and availability; all further chip away the realizable effectiveness of any given antimalarial agent. The contraindications are particularly important in the case of P. vivax and the crucial therapy against relapses with PQ, the only current therapeutic option for that clinical indication. Primaquine (and all other 8-aminoquinoline compounds evaluated) invariably provokes an acute hemolytic anemia in patients receiving therapeutic doses against relapse and having an inherited X chromosome-linked deficiency in glucose-6-phosphate dehydrogenase (G6PD) enzymatic activity [35]. This abnormality affects approximately 400 million people or 8% of people residing in malaria endemic countries [36]. Safe access to PQ for radical cure of vivax malaria may require access to point-of-care diagnostics for G6PD deficiency [37]. Even with such testing, however, there remains the problem of treating those diagnosed as G6PD-deficient, pregnant or lactating women, and infants below the age of 6 months [38]. There are no optimized or validated means of preventing relapse without 8-aminoquinoline drugs, e.g., by chemopreventive or presumptive periodic preventive therapeutic strategies [39, 40]. The 8-aminoquinoline drug, called tafenoquine, is in late clinical development and will likely soon offer a single-dose option to PQ, virtually eliminating the important adherence problem with that therapy [41, 42].

Another potential problem in the human host may be the inability to metabolize PQ to its active hypnozoite-killing metabolite by cytochrome P450 2D6 (CYP2D6) [43]. Natural polymorphism in the gene expressing CYP2D6 leads to a range of metabolic activities ranging anywhere between far above normal and null. Patients in need of PQ anti-relapse therapy and having significantly impaired or null CYP2D6 activity may relapse even with full compliance to good quality drug. We do not yet know the extent of this problem with regard to the frequencies of CYP2D6 alleles associated with PQ therapeutic failure, but the significantly impaired CYP2D6 *10 allele (a particular genetic variant of CYP2D6 gene) is relatively common among Southeast Asians, at about 35% frequency [44]. It may be that many Asians will be unable to adequately metabolize PQ and achieve successful radical cure [45].

The ambiguity of geographically variable frequency and timing of relapse—along with reinfection and recrudescence in recurrent P. vivax malaria after PQ therapy—makes estimating PQ efficacy in endemic settings very difficult. This is true even with directly observed therapy using high-quality drug. After decades of recommending a 5-day regimen of PQ against relapse, on the basis of observed low rates of relapse following therapy, investigators in India ultimately included a relapse control group (placebo) and discovered that efficacy to be nil [46]—the low rate of relapse was naturally occurring. John et al. [47] systematically reviewed recurrence rates after standard 0.25 mg/kg daily for 14-day regimen with rates of recurrence averaging about 8% at 1 month, 10% at 2–3 months, 14% at 4–6 months, and 20% at 7–12 months. In two randomized controlled trials of PQ given at high dose (0.5 mg/kg) to 257 Indonesian soldiers infected by P. vivax in eastern Indonesia and followed for a year where reinfection was not possible, 35 (14%) experienced at least one relapse [32, 33]. Among the 21 subjects whose CYP2D6 genotype and phenotype were examined, 20 showed evidence of significant functional impairment of CYP2D6 [48].

Evidence supports the notion of providing presumptive anti-relapse therapy to all patients diagnosed with any species of malaria agents, especially P. falciparum. In a retrospective analysis of over 10,000 research subjects naturally infected by P. falciparum in Thailand or Myanmar, 912 were treated with rapidly excreted blood schizonticides, and within 2 months, just over 50% experienced a P. vivax attack [49]. The people infected by one species in any given community must be considered at high risk of harboring latent and perhaps sub-patent infections of the other co-endemic species. Species-specific therapies, especially in an age of dominant CQ resistance among the plasmodia, may not be sensible in an elimination context.

Effective diagnosis and treatment represents the cornerstone of current control and elimination strategies, and the obstacles described here require consideration in realizing gains against this tenacious endemic problem. Indeed, such gains have been achieved both historically and recently. At the turn of the twentieth century, endemic vivax malaria occurred across much of southeastern North America, northern and southern Europe, the Middle East, and northern Australia—areas where it no longer appears. Much of this success was achieved applying environmental modifications against local Anopheles vectors, but more recent elimination successes using principally diagnosis and treatment strategy have occurred in nations like Turkey, Azerbaijan, and Sri Lanka, as examples [50]. The same had been achieved on the Korean Peninsula during the 1970s, but endemic vivax malaria transmission reappeared during the 1990s and persists today [51]. Post-elimination vigilance that includes not only diagnosis and treatment services but also vector control may be essential to protecting and sustaining the elimination of endemic vivax malaria [50].

Advertisement

5. Vector control in vivax malaria

Vector control of endemic vivax malaria may not have immediate impacts due to the hypnozoite reservoir contributing >80% of acute attacks of vivax malaria in low or high endemic settings [52, 53]. Success in reducing malaria incidence and local transmission to zero in a malaria endemic area, particularly where sympatric P. falciparum and P. vivax occur, may require greater sustainability of vector control measures. Vivax malaria transmission will outlast falciparum malaria, and reestablishment of local transmission may occur without imported cases, i.e., by local hypnozoites. Prevention of the seeding of new hypnozoites in liver cells by biting Anopheles mosquitoes obviously may contribute positively to the control and elimination of vivax malaria in the long term, but no randomized controlled trials yet affirm this. In one large cluster-randomized trial in Myanmar, insecticide-treated bed netting (ITN) had no impact whatsoever on the risk of malaria [54], an outcome attributed to the dominant Anopheles vector, A. dirus s.s., feeding predominantly outdoors and early in evening or morning [55]. Relatively modest effects were reported from a similarly cluster-randomized trial in Vietnam, again attributed to mosquito behaviors unfavorable to control by this means [56]. The main Asian vector species tend to feed early in the evening and outdoors where they also rest [57], minimizing their exposure to household insecticides. In other studies of strategies for minimizing exposure to Anopheles, much greater impacts against falciparum malaria were demonstrated relative to those against vivax malaria [58, 59, 60].

Over the last decade, attempts of using spatial repellents (SRs) to minimize exposure to biting insects have shown some success in diverse settings [61]. Repellency is distinct from the killing action of insecticides in more than one way, i.e., no direct contact is required, and lacking lethality does not select for resistance. SRs are effective irrespective of indoor or late-night feeding and resting behavior like conventional netting or indoor spraying. SRs should be evaluated for added benefit in areas where traditional long-lasting insecticidal net (LLIN) or indoor residual spraying (IRS) of insecticides interventions may not offer full protection or have reached their efficacy limits—especially in areas with residual transmission or in areas where elimination may be considered feasible. Control of disease in these areas will require new approaches, and possibly spatial repellency would be practical and effective [62, 63]. SRs may be useful as stand-alone tools of personal or household protection where other interventions may not reach. Also, they may be combined with conventional interventions to augment their impacts.

Another vector control strategy for eliminating P. vivax in the Asian-Pacific region may be the method of environmental modification called “species sanitation.” This approach offers prevention independently of the myriad problems and challenges of diagnosis and treatment or the limitations of insecticidal strategies. Species sanitation is simply sanitizing the environment against specific incriminated vector species by exploiting detailed knowledge of their bionomics (behavior and ecology) [64]. Malcolm Watson in British Malaya, along with Nicholas Swellengrebel and Raden Soesilo in the East Indies, invented, optimized, and validated species sanitation in malaria control [65]. A systematic analysis of 16 such interventions (most conducted before 1945) showed an average 88% reduction of malaria burden [66]. As new cases occur by relapse, reinfection, or importation, making the subsequent infection of mosquitoes improbable (by simply reducing their numbers) eventually suffocates transmission.

Although the implementation of LLIN, IRS, and species sanitation in different environmental settings rendered significant success rates [67], it is evident that the key determining factors for the success of any vector intervention selected is a thorough knowledge of the vector bionomics, local malaria transmission dynamics, and residual efficacy of choice insecticide. Knowledge of vector bionomics includes ascertaining breeding and resting preferences and feeding behavior of incriminated vector species. Transmission dynamics include information related to entomological inoculation rates, sibling species composition of vectors (based on reliable PCR identification assays), seasonality of malaria prevalence, and risk factors that may support the human-mosquito contact, while suitable insecticide means any available insecticide that renders knockdown effect and/or mortality to the incriminated vector population.

Another important issue to be considered is the ability of the Anopheles mosquitoes to adapt to the ongoing vector interventions by changing host-seeking behavior, such as from indoor to outdoor or vice versa, and selection of insecticide-resistant strain [68]. With current trends in globalization and population migration, deforestation, and resettlement of populations, reintroduction of malaria into areas that have been declared free from transmission is a clear and present risk. Therefore, no single intervention method may guarantee long-term efficacy; thus, regular monitoring of vector density and behavior should be a routine operation wherever this risk occurs. Most malaria control programs no longer have the entomological expertise needed to carry out these important tasks—addressing this problem may be the greatest and most important challenge within the context of a malaria elimination agenda.

Advertisement

6. Vaccination

A vaccine that prevents the seeding of human livers by both active schizonts and dormant hypnozoites of P. vivax would provide a conspicuously useful tool in eliminating this species. Mass or routine vaccination now seems impractical with non-sterilizing vaccines of short-lived immunity needing 3 or 4 doses. These may improve in the future, but even now a malaria vaccine could be applied in geographically or demographically narrowed settings to potentially great impacts. For example, high-risk and hard-to-reach populations like migrant workers or soldiers having sterile immunity to malaria (even if for just a season or two) may not only protect those people from harm but also greatly slow importation of malaria into receptive areas where transmission has been interrupted. Likewise, people living in areas prone to reintroduction of endemic malaria by high volumes of immigration from high-risk areas may be immunized and protected against very dangerous outbreaks and epidemics [69].

Today, there is no vaccine available that can prevent infection by P. vivax with high levels of sterilizing immune protection. That is also true for all other plasmodial species. The half-century-long efforts to develop a vaccine against P. falciparum—greatly aided by the ability to cultivate this species in continuous laboratory cultures since the late 1970s—culminated in the molecular subunit vaccine called RTS,S ASO1 (mimicking a protein-coating infectious sporozoites) [70] with the registered trademark name Mosquirix™ (GlaxoSmithKline). The vaccine did not prevent infection in the African infants and young children vaccinated but had the modest effects against higher parasitemias and signs of illness [67]. The modest efficacy combined with worrying and puzzling signals like increased risk of pneumococcal meningitis and significantly higher all-cause mortality among vaccinated females apparently explains the WHO position to withhold a favorable opinion on the vaccine until further studies involving targeted and limited rollout in several African nations are completed [71]. Molecular subunit vaccines targeting P. vivax molecules have not progressed beyond Phase 2a and show similar inability to achieve high levels of sterilizing protection [72].

Over the past decade, investigators applying live-attenuated sporozoites of P. falciparum have shown high levels of durable (~12 months) sterilizing protection in malaria-naïve adult volunteers in controlled human malaria infection (CHMI) experiments using a challenge strain homologous to the vaccine strain [73]. This approach relies on laboratory harvest of infectious sporozoites from laboratory-reared aseptic anopheline mosquitoes infected by P. falciparum maintained in the laboratory. Deriving live-attenuated sporozoites of P. vivax is possible [74] but exceedingly difficult, not strain-specific, and not sustainable as a source of vaccine. Nonetheless, immunization by irradiated sporozoites of the murine species Plasmodium berghei cross-protected against the murine species Plasmodium yoelii and vice versa in murine challenge models [75]. The possibility of sporozoites of P. falciparum cross-protecting against P. vivax challenge has not been examined directly, but proteomic analyses showed that these two human plasmodia species shared substantially more common probable T-cell epitopes than that between P. berghei and P. yoelii. A vaccine derived from laboratory-kept P. falciparumsystems offering protection against P. vivax would represent a quantum leap forward for vaccination against this species by effectively sidestepping the requirement for continuous laboratory cultivation for a live vaccine.

Advertisement

7. Challenges and recommendations

The greatest challenge in eliminating vivax malaria—the hypnozoite reservoir—may also be the greatest opportunity to accomplish the task. If >80% of incident malaria cases indeed derive from hypnozoites, then surely attacking and shrinking that reservoir would deliver substantial reductions in the burden of morbidity and mortality. Despite the availability of PQ for over 65 years, sustained and systematic assault on that reservoir has not been accomplished in the endemic tropics—largely due to the unsolved clinical problem of its hemolytic toxicity in G6PD-deficient patients.

Eliminating P. vivax malaria will require accepting the inadequacy of conventional falciparum malaria-focused control strategy, tactics, and tools and committing to the optimizing and validating of interventions suited to this stubborn parasite. This effectively means striving to solve the wrenchingly difficult problem of the hemolytic toxicity of PQ in G6PD-deficient patients by almost any means. The obstacles presented in managing populations and individual patients carrying this infection emphasize the great advantage of preventing it in the first place with an effective vector control strategy. In this context, species sanitation has proven highly effective against endemic Asian malarias a century ago [54] and would probably do so again.

Taking all these factors into considerations, we recommend the following measures for eliminating endemic vivax malaria:

  1. Active case detection and early treatment are essential steps, fundamental to eliminating any endemic malaria; however, this measure alone will not lead to elimination—too many infections are latent, sub-patent, sequestered, and asymptomatic.

  2. Adoption of safe and universal access to radical cure for cases of vivax malaria along with universal access to alternative means of relapse prevention for people ineligible for therapy with 8-aminoquinolines would accelerate progress to elimination. Achieving that will likely also require better diagnostics for both the parasite and G6PD deficiency than are currently available.

  3. Adoption of radical cure with an 8-aminoquinoline and ACT with diagnosis of any species of malaria where P. vivax also occurs as a means of targeting likely carriers of hypnozoites.

  4. Reduce new vivax infections/seeding of the liver with hypnozoites by substantially reducing human contact with malaria vectors, effectively stranding extant parasites in all stages of human infection—latent, sub-patent, patent, and eventually vanishing without Anopheles contact and onward transmission. Interrupting transmission by species sanitation measures may be the most durable and effective means of achieving this goal.

  5. Examine the possibility of sterilizing immune protection against P. vivax provided by attenuated P. falciparum sporozoite vaccines providing an immediately highly relevant tool for eliminating endemic P. vivax.

Advertisement

8. Conclusion

Plasmodium vivax passes substantial challenges that may hinder achievement of global malaria elimination by 2030. The most challenging evidence is the lack of technology to detect the latent infection caused by hypnozoite. Therefore, the only tool to prevent P. vivax transmission originated from reactivation of hypnozoites is by vector control.

Advertisement

Acknowledgments

We gratefully acknowledge Prof. Amin Soebandrio MD, PhD, Clin. Microbiol, Director of the Eijkman Institute for Molecular Biology for his encouragement and advice, and Prof. A. Asadul I. Tahir, MD, PhD, FICS, Dean of the Faculty of Medicine, Hasanuddin University for the support to DS. JKB is supported by the Vietnam Major Overseas Programme grant from the Wellcome Trust.

Advertisement

List of acronyms

ACTartemisinin-based combination therapy
CHMIcontrolled human malaria infection
CQchloroquine
CYP2D6cytochrome P450 2D6
G6PDglucose-6-phosphate dehydrogenase
IRSindoor residual spraying
ITNinsecticide-treated bed nets
LLINlong-lasting insecticide treated nets
PQprimaquine
SEASoutheast Asia
SRspatial repellent
WHOWorld Health Organization

References

  1. 1. Gething PW, Elyazar IR, Moyes CL, Smith DL, Battle KE, Guerra CA, Patil AP, Tatem AJ, Howes RE, Myers MF, George DB, Horby P, Wertheim HF, Price RN, Müeller I, Baird JK, Hay SI. A long-neglected world malaria map: Plasmodium vivax endemicity in 2010. PLoS Neglected Tropical Diseases. 2012;6(9):1814. DOI: 10.1371/journal.pntd.0001814
  2. 2. Battle KE, Gething PW, Elyazar IR, Moyes CL, Sinka ME, Howes RE, Guerra CA, Price RN, Baird KJ, Hay SI. The global public health significance of Plasmodium vivax. Advances in Parasitology. 2012;80:1-111. DOI: 10.1016/B978-0-12-397900-1.00001-3
  3. 3. Rosalind EH, Katherine EB, Kamini NM, David LS, Richard EC, Kevin Baird J, Hay SI. Global epidemiology of Plasmodium vivax. The American Journal of Tropical Medicine and Hygiene. 2016;95(6):15-34. DOI: 10.4269/ajtmh.16-0141
  4. 4. Price RN, von Seidlein L, Valecha N, Nosten F, Baird JK, White NJ. Global extent of chloroquine-resistant Plasmodium vivax: A systematic review and meta-analysis. The Lancet Infectious Diseases. 2014;14(10):982-991. DOI: 10.1016/S1473-3099(14)70855-2
  5. 5. Baird JK. Resistance to therapies to infection by Plasmodium vivax. Clinical Microbiology Reviews. 2009;22:508-534. DOI: 10.1128/CMR.00008-09
  6. 6. White NJ, Imwong M. Relapse. Advances in Parasitology. 2012;80:113-150. DOI: 10.1016/B978-0-12-397900-1.00002-5
  7. 7. Baird JK, Leksana B, Masbar S, Fryauff DJ, Sutanihardja MA, Suradi, Wignall FS, Hoffman SL. Diagnosis of resistance to chloroquine by Plasmodium vivax: Timing of recurrence and whole blood chloroquine levels. The American Journal of Tropical Medicine and Hygiene. 1997;56:618-620. PMID: 9230792
  8. 8. Asih PB, Syafruddin D, Leake J, Sorontou Y, Sadikin M, Sauerwein RW, Vinetz J, Baird JK. Phenotyping clinical resistance to chloroquine in Plasmodium vivax in northeastern Papua, Indonesia. International Journal for Parasitology: Drugs and Drug Resistance. 2011;1(1):28-32. DOI: 10.1016/j.ijpddr.2011.08.001
  9. 9. Chen N, Auliff A, Rieckmann K, Gatton M, Cheng Q. Relapses of Plasmodium vivax result from clonal hypnozoites activated at predetermined intervals. The Journal of Infectious Diseases. 2007;195:934-941. PMID: 17330782
  10. 10. Imwong M, Snounou G, Pukrittayakamee S, Tanomsing N, Kim JR, Nandy A, Guthmann JP, Nosten F, Carlton J, Looareesuwan S, Nair S, Sudimack D, Day NP, Anderson TJ, White NJ. Relapses of Plasmodium vivax infection usually result from activation of heterologous hypnozoites. The Journal of Infectious Diseases. 2007;195:927-933. PMID: 17330781
  11. 11. Baird JK, Valecha N, Duparc S, White NJ, Price RN. Diagnosis and treatment of Plasmodium vivax malaria. The American Journal of Tropical Medicine and Hygiene. 2016;95(6):35-51. DOI: 10.4269/ajtmh.16-0171
  12. 12. Baird JK, Fryauff DJ, Hoffman SL. Primaquine for prevention of malaria in travellers. Clinical Infectious Diseases. 2003;37(12):1659-1667. DOI: https://doi.org/10.1086/379714
  13. 13. Baird JK, Hoffman SL. Primaquine therapy for malaria. Clinical Infectious Diseases. 2004;39(9):1336-1345. DOI: https://doi.org/10.1086/424663
  14. 14. Baird JK. Resistance to chloroquine unhinges vivax malaria therapeutics. Antimicrobial Agents and Chemotherapy. 2011;55(5):1827-1830. DOI: 10.1128/AAC.01296-10
  15. 15. WHO. Global Technical Strategy for Malaria 2016-2030. Geneva, World Health Organization (WHO); 2015. http://apps.who.int/iris/bitstream/10665/176712/1/9789241564991_eng.pdf?ua=1&ua=1 [Accessed: 31 January 2018]
  16. 16. WHO. Control and Elimination of Plasmodium vivax: A Technical Brief. Geneva, Switzerland: World Health Organization (WHO); 2015. http://www.searo.who.int/entity/malaria/control-and-elimination-plasmodium.pdf?ua=1 [Accessed: 31 January 2018]
  17. 17. Mendis K, Sina BJ, Marchesini P, Carter R. The neglected burden of Plasmodium vivax malaria. The American Journal of Tropical Medicine and Hygiene 2001;64(1):97-106. PMID: 11425182
  18. 18. Baird JK. Neglect of Plasmodium vivax. Trends in Parasitology 2007;23:533-539. PMID: 17933585
  19. 19. Price RN, Tjitra E, Guerra CA, Yeung S, White NJ, Anstey NM. Vivax malaria: Neglected and not benign. The American Journal of Tropical Medicine and Hygiene 2007;77(6):79-87. PMID:18165478
  20. 20. Mueller I, Galinski MR, Baird JK, Carlton JM, Kochar DK, Alonso PL, del Portillo HA. Key gaps in the knowledge of Plasmodium vivax, a neglected human malaria parasite. The Lancet Infectious Diseases. 2009;9(9):555-566. DOI: 10.1016/S1473-3099(09)70177-X
  21. 21. Baird JK. Evidence and implications of mortality associated with acute Plasmodium vivax malaria. Clinical Microbiology Reviews. 2013;26(1):36-57. DOI: 10.1128/CMR.00074-12
  22. 22. Anstey NM, Douglas NM, Poespoprodjo JR, Price RN. Plasmodium vivax clinical spectrum, risk factors and pathogenesis. Advances in Parasitology. 2012;80:151-201. DOI: 10.1016/B978-0-12-397900-1.00003-7
  23. 23. Barber BE, William T, Grigg MJ, Parameswaran U, Piera KA, Price RN, Yeo TW, Anstey NM. Parasite biomass-related inflammation, endothelial activation, microvascular dysfunction and disease severity in vivax malaria. PLoS Pathogens. 2015;11. DOI: https://doi.org/10.1371/journal.ppat.1004558
  24. 24. Malleret B, Li A, Zhang R, Tan KS, Suwanarusk R, Claser C, Cho JS, Koh EG, Chu CS, Pukrittayakamee S, Ng ML, Ginhoux F, Ng LG, Lim CT, Nosten F, Snounou G, Rénia L, Russell B. Plasmodium vivax: Restricted tropism and rapid remodeling of CD71-positive reticulocytes. Blood. 2015;125(8):1314-1324. DOI: 10.1182/blood-2014-08-596015
  25. 25. Cheng Q, Cunningham J, Gatton ML. Systematic review of sub-microscopic P. vivax infections: Prevalence and determining factors. PLoS Neglected Tropical Diseases. 2015;9(1):3413. DOI: 10.1371/journal.pntd.0003413
  26. 26. Moreira CM, Abo-Shehada M, Price RN, Drakeley CJ. A systematic review of sub-microscopic Plasmodium vivax infection. Malaria Journal. 2015;14:360. DOI: 10.1186/s12936-015-0884-z
  27. 27. Okell LC, Bousema T, Griffin JT, Ouedraogo AL, Ghani AC, Drakeley CJ. Factors determining the occurrence of submicroscopic malaria infections and their relevance for control. Nature Communications. 2012;3:1237. DOI: 10.1038/ncomms2241
  28. 28. Menard D, Barnadas C, Bouchier C, Henry-Halldin C, Gray LR, Ratsimoasoa A, Thonier V, Carod JF, Domarle O, Colin Y, Bertrand O, Picot J, King CL, Grimberg BT, Mercereau-Puijalon O, Zimmerman PA. Plasmodium vivax clinical malaria is commonly observed in Duffy-negative Malagasy people. Proceedings of the National Academy of Sciences of the United States of America. 2010;107:5967-5971. DOI: 10.1073/pnas.0912496107
  29. 29. Culleton R, Ndounga M, Zeyrek FY, Coban C, Casimiro PN, Takeo S, Tsuboi T, Yadava A, Carter R, Tanabe K. Evidence for the transmission of Plasmodium vivax in the Republic of the Congo, West Central Africa. The Journal of Infectious Diseases. 2009;200:1465-1469. DOI: 10.1086/644510
  30. 30. Howes RE, Reiner RC Jr, Battle KE, Longbottom J, Mappin B, Ordanovich D, Tatem AJ, Drakeley C, Gething PW, Zimmerman PA, Smith DL, Hay SI. Plasmodium vivax transmission in Africa. PLoS Neglected Tropical Diseases. 2015;9:0004222. DOI: 10.1186/s12936-1118-8
  31. 31. Gogtay N, Kannan S, Thatte UM, Olliaro PL, Sinclair D. Artemisinin-based combination therapy for treating uncomplicated Plasmodium vivax malaria. Cochrane Database of Systematic Reviews. 2013;10:CD008492. DOI: 10.1002/14651858.XCD008492.pub3
  32. 32. Sutanto I, Tjahjono B, Basri H, Taylor WR, Putri FA, Meilia RA, Setiabudy R, Nurleila S, Ekawati LL, Elyazar I, Farrar J, Sudoyo H, Baird JK. Randomized, open-label trial of primaquine against vivax relapse in Indonesia. Antimicrob Agents Chemother. 2013;57:1128-1135. DOI: 1128/AAX.01879-12
  33. 33. Nelwan EJ, Ekawati LL, Tjahjono B, Setiabudy R, Sutanto I, Chand K, Ekasari T, Djoko D, Basri H, Taylor WR, Duparc S, Subekti D, Elyazar I, Noviyanti R, Sudoyo H, Baird JK. Randomized trial of primaquine hypnozoitocidal efficacy when administered with artemisinin-combined blood schizontocides for radical cure of Plasmodium vivax in Indonesia. BMC Medicine. 2015;13:294. DOI: 10.1186/s12916-015-0535-9
  34. 34. Commons RJ, Thriemer K, Humphreys S, Suay I, Sibley CH, Guerin PJ, Price RN. The vivax surveyor: Online mapping database for Plasmodium vivax clinical trials. International Journal for Parasitology: Drugs and Drug Resistance. 2017;7:181-190. DOI: 10.1016/ijpddr.2017.03.003
  35. 35. Luzzatto L, Nannelli C, Notaro R. Glucose-6-phosphate dehydrogenase deficiency. Hematology/Oncology Clinics of North America. 2016;30(2):373-393. DOI: 10.1016/j.hoc.2015.11.006
  36. 36. Howes RE, Piel FB, Patil AP, Nyangiri OA, Gething PW, Dewi M, Hogg MM, Battle KE, Padilla CD, Baird JK, Hay SI. G6PD deficiency prevalence and estimates of affected populations in malaria endemic countries: A geostatistical model-based map. PLoS Medicine. 2012;9:1001339. DOI: https://doi.org/10.1371/journal.pmed.1001339
  37. 37. Baird JK. Point-of-care G6PD diagnostics for Plasmodium vivax malaria is a clinical and public health urgency. BMC Medicine. 2015;13:296. DOI: 10.1186/s12916-015-0531-0
  38. 38. WHO. Guidelines for the treatment of malaria. 3rd ed. Geneva, World Health Organization (WHO); 2015. http://apps.who.int/iris/bitstream/10665/162441/1/9789241549127_eng.pdf [Accessed: 31 January 2018]
  39. 39. Peters AL, Van Noorden CJ. Glucose-6-phosphate dehydrogenase deficiency and malaria: Cytochemical detection of heterozygous G6PD deficiency in women. The Journal of Histochemistry and Cytochemistry. 2009;57(11):1003-1011. DOI: 10.1369/jhc.2009.953828
  40. 40. Padilla CD, Therrell BL. Working group of the Asia Pacific Society for human genetics on consolidating newborn screening efforts in the Asia Pacific R. Consolidating newborn screening efforts in the Asia Pacific region: networking and shared education. Journal of Community Genetics. 2012;3:35-45. DOI: 10.1007/s12687-011-0076-7
  41. 41. Douglas NM, Poespoprodjo JR, Patriani D, Malloy MJ, Kenangalem E, Sugiarto P, Simpson JA, Soenarto Y, Anstey NM, Price RN. Unsupervised primaquine for the treatment of Plasmodium vivax malaria relapses in Southern Papua: A hospital-based cohort study. PLoS Medicine. 2017;14:1002379. DOI: 10.1371/journal.pmed.1002379
  42. 42. Rjapaske S, Rodrigo C, Fernando SD. Tafenoquine for preventing relapse in people with Plasmodium vivax malaria. Cochrane Database of Systematic Reviews. 2015;29. DOI: CD010458, 10.1002/14651858.CD010458.pub2
  43. 43. Marcsisin SR, Reichard G, Pybus BS. Primaquine pharmacology in the context of CYP 2D6 pharmacogenomics: Current state of the art. Pharmacology & Therapeutics. 2016;161:1-10. DOI: 10.1016/j.pharmthera.2016.03.011
  44. 44. Sistonen J, Fuselli S, Palo JU, Chauhan N, Padh H, Sajantila A. Pharmacogenetic variation at CYP2C9, CYP2C19, and CYP2D6 at global and microgeographic scales. Pharmacogenetics and Genomics. 2009;19(2):170-179. DOI: 10.1097/FPC.0b013e32831ebb30
  45. 45. 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. Malaria Journal. 2018;17:42. DOI: 10.1186/s12936-018-2190-z
  46. 46. Gogtay NJ, Desai S, Kamtekar KD, Kadam VS, Dalvi SS, Kshirsagar NA. Efficacies of 5- and 14-day primaquine regimens in the prevention of relapses in Plasmodium vivax infections. Annals of Tropical Medicine and Parasitology. 1999;93:809-812. PMID: 9861412
  47. 47. John GK, Douglas NM, von Seidlein L, Nosten F, Baird JK, White NJ, Price RN. Primaquine radical cure of Plasmodium vivax: A critical review of the literature. Malaria Journal. 2012;11:280. DOI: 10.1186/1475-2875-11-1280
  48. 48. Baird JK, Louisa M, Noviyanti R, Ekawati L, Elyazar I, Subekti D, Chand K, Gayatri A, Instiaty, Soebianto S, Crenna-Darusallam C, Joki D, Silistyanto BP, Meriyenes D, Wesche D, Nelwan EJ, Sutanto I, Sudoyo H, Setiabudy R. Impaired cytochrom P-450 2D6 activity genotype and phenotype associated with primaquine treatment failure against latent Plasmodium vivax malaria: A nested case control study. JAMA Open Network, in press
  49. 49. Douglas NM, Nosten F, Ashley EA, Phaiphun L, van Vugt M, Singhasivanon P, White NJ, Price RN. Plasmodium vivax recurrence following falciparum and mixed species malaria: Risk factors and effect of elimination kinetics. Clinical Infectious Diseases. 2011;52:612-620. DOI: 10.1093/cid/ciq249
  50. 50. WHO. Eliminating Malaria. Global Malaria Program, World Health Organization, Geneva; 2016. WHO/HTM/GMP/2016.3
  51. 51. Park JW, Jun G, Yeom JS. Plasmodium vivax malaria: Status in the Republic of Korea following re-emergence. The Korean Journal of Parasitology. 2009;47:S39-S50. DOI: 10.3347/kjp.2009.47.S.S39
  52. 52. Adekunle AI, Pinkevych M, McGready R, Luxemburger C, White LJ, Nosten F, Cromer D, Davenport MP. Modeling the dynamics of Plasmodium vivax infection and hypnozoite reactivation in vivo. PLOS Neglected Tropical Diseases. 2015;9:0003595. DOI: https://doi.org/10.1371/journal.pntd.0003595
  53. 53. Robinson LJ, Wampfler R, Betuela I, Karl S, White MT, CSN LWS, Hofmann NE, Kinboro B, Waltmann A, Brewster J, Lorry L, Tarongka N, Samol L, Mueller I. 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 Medicine. 2015:1001891. DOI: https://doi.org/10.1371/journal.pmed.1001891
  54. 54. Smithuis FM, Kyaw MK, Phe UO, van der Broek I, Katterman N, Rogers C, Almeida P, Kager PA, Stepniewska K, Lubell Y, Simpson JA, White NJ. 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. Malaria Journal. 2013;12:363. DOI: https://doi.org/10.1186/1475-2875-12-363
  55. 55. Smithuis FM, Kyaw MK, Phe UO, van der Broek I, Katterman N, Rogers C, Almeida P, Kager PA, Stepniewska K, Lubell Y, Simpson JA, White NJ. Entomological determinants of insecticide-treated bed net effectiveness in Western Myanmar . Malaria Journal. 2013;12:364 DOI: https://doi.org/10.1186/1475-2875-12-364
  56. 56. Thang ND, Erhart A, Speybroeck N, Xa NX, Thanh NN, Ky PV, Hung LH, Thuan LK, Coosemans M, D'Alessandro U. Long-lasting insecticidal hammocks for controlling forest malaria: A community-based trial in a rural area of Central Vietnam. PLoS One. 2009;4(10):7369. DOI: http://doi.org/10.1371/journal.pone.0007369
  57. 57. Sinka ME, Bangs MJ, Manguin S, Chareonviriyaphap T, Patil AP, Temperley WH, Gething PW, Elyazar IR, Kabaria CW, Harbach RE, Hay SI. The dominant vectors of human malaria in the Asia-Pacific region: Occurrence data, distribution maps and bionomics precis. Parasites & Vectors. 2011;4:89. DOI: 10.1186/1756-3305-4-89
  58. 58. Ferreira MU, Castro MC. Challenges for malaria elimination in Brazil. Malaria Journal. 2016;15:284. DOI: 10.1186/s12936-016-1335-1
  59. 59. Avikar AR, Shah N, Dhariwal AC, Sonal GS, Prdhan MM, Ghosh SK, Valecha N. Epidemiology of Plasmodium vivax in India. The American Journal of Tropical Medicine and Hygiene. 2016;95(6Suppl):108-120. DOI: 10.4269/ajtmh.16—163
  60. 60. Sharma VP, Dev V, Phookan S. Neglected Plasmodium vivax malaria in northeastern states of India. Indian Journal of Medical Research. 2015;141:546-555. DOI: 10.4103/0971-5916.159511
  61. 61. Syafruddin D, Bangs MJ, Sidik D, Elyazar I, Asih PB, Chan K, Nurleila S, Nixon C, Hendarto J, Wahid I, Ishak H, Bøgh C, Grieco JP, Achee NL, Baird JK. Impact of a spatial repellent on malaria incidence in two villages in Sumba, Indonesia. The American Journal of Tropical Medicine and Hygiene. 2014;91(6):1079-1087. DOI: 10.4269/ajtmh.13-0735
  62. 62. Ogoma SB, Lorenz LM, Ngonyani H, Sangusangu R, Kitumbukile M, Kilalangongono M, Simfukwe ET, Mbeyela E, Roman D, Moore J, Kreppel K, Maia MF, Moore SJ. An experimental hut study to quantify the effect of DDT and airborne pyrethroids on entomological parameters of malaria transmission. Malaria Journal. 2014;13:131. DOI: https://doi.org/10.1186/1475-2875-13-131
  63. 63. Achee NL, Bangs MJ, Farlow R, Killeen GF, Lindsay S, Logan JG, Moore SJ, Rowland M, Sweeney K, Torr SJ, Zwiebel LJ, Grieco JP. Spatial repellents: From discovery and development to evidence-based validation. Malaria Journal. 2012;11:164. DOI: http://doi.org/10.1186/1475-2875-11-164
  64. 64. Baird JK. Malaria control by commodities without practical malariology. BMC Public Health. 2017;17:590. DOI: https://doi.org/10.1186/s12889-017-4454-x
  65. 65. Keiser J, Singer BH, Utzinger J. Reducing the burden of malaria in different eco-epidemiological settings with environmental management: A systematic review. The Lancet Infectious Diseases. 2005;5(11):695-708. DOI: 10.1016/S1473-3099(05)70268-1
  66. 66. Takken W, Snellen WB, Verhave JP, Knols BGJ, Atmosoedjono S. Environmental measures for malaria control in Indonesia -an historical review on species sanitation. Wageningen Agricultural University Papers. 1990;90-7:1-16. http://edepot.wur.nl/282643
  67. 67. Lengeler C. Insecticide-treated bed nets and curtains for preventing malaria. Cochrane Database of Systematic Reviews. 2004;2:CD000363. DOI: 10.1002/14651858.CD000363.pub2
  68. 68. Sokhna C, Ndiath MO, Rogier C. The changes in mosquito vector behaviour and the emerging resistance to insecticides will challenge the decline of malaria. Clinical Microbiology and Infection. 2013;19(10):902-907. DOI: 10.1111/1469-0691.12314
  69. 69. Gosling R, von Seidlein L. The future of the RTS,S/AS01 malaria vaccine: An alternative development plan. PLoS Medicine. 2016;13:1001994. DOI: 10.1371/journal.pmed.1001994
  70. 70. Fowkes FJI, Simpson JA, Beeson JG. Implications of the licensure of a partially efficacious malaria vaccine on evaluating second-generation vaccines. BMC Medicine. 2013;11:232. DOI: 10.1186/1741-7015-11-232
  71. 71. WHO. Malaria vaccine: World Health Organization position paper – January 2016. Weekly Epidemiological Record. 2016;91:33-52. http://www.who.int/wer/2016/wer9104.pdf
  72. 72. Sandoval-Reyes A, Bachmann MF. Plasmodium vivax vaccines: Why are we where we are? Human Vaccines & Immunotherapeutics. 2013;9:2558-2565. DOI: 10.4161/hv.26157
  73. 73. Hoffman SL, Vekemans J, Richie TL, Duffy PE. The march toward malaria vaccines. Vaccine. 2015;33(Suppl 4):D13-D23. DOI: 10.1016/j.vaccine.2015.07.091
  74. 74. Chattopadhyay R, Velmurugan S, Chakiath C, Donkor LA, Milhous W, Barnwell JW, Collins WE, Hoffman SL. Establishment of an in vitro assay for assessing the effects of drugs on the liver stages of Plasmodium vivax malaria. PLoS One. 2010;5:14275. DOI: 10.1371/journal.pone.0014275
  75. 75. Sedegah M, Weiss WW, Hoffman SL. Cross-protection between attenuated Plasmodium berghei and P. yoelii sporozoites. Parasite Immunology. 2007;29:559-565. DOI: 10.1111/j.1365-3024.2007.00976.x

Written By

Puji BS Asih, Din Syafruddin and John Kevin Baird

Submitted: 05 July 2017 Reviewed: 09 April 2018 Published: 18 July 2018