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Malaria is a significant public health problem and impediment to socioeconomic development in countries of the Greater Mekong Subregion (GMS), which comprises Cambodia, China’s Yunnan Province, Lao People’s Democratic Republic, Myanmar, Thailand, and Vietnam. Over the past decade, intensified malaria control has greatly reduced the regional malaria burden. Driven by increasing political commitment, motivated by recent achievements in malaria control, and urged by the imminent threat of emerging artemisinin resistance, the GMS countries have endorsed a regional malaria elimination plan with a goal of eliminating malaria by 2030. However, this ambitious, but laudable, goal faces a daunting array of challenges and requires integrated strategies tailored to the region, which should be based on a mechanistic understanding of the human, parasite, and vector factors sustaining continued malaria transmission along international borders. Malaria epidemiology in the GMS is complex and rapidly evolving. Spatial heterogeneity requires targeted use of the limited resources. Border malaria accounts for continued malaria transmission and represents sources of parasite introduction through porous borders by highly mobile human populations. Asymptomatic infections constitute huge parasite reservoir requiring interventions in time and place to pave the way for malaria elimination. Of the two most predominant malaria parasites, Plasmodium falciparum and P. vivax, the prevalence of the latter is increasing in most member GMS countries. This parasite requires the use of 8-aminoquinoline drugs to prevent relapses from liver hypnozoites, but high prevalence of glucose-6-phosphate dehydrogenase deficiency in the endemic human populations makes it difficult to adopt this treatment regimen. The recent emergence of resistance to artemisinins and partner drugs in P. falciparum has raised both regional and global concerns, and elimination efforts are invariably prioritized against this parasite to avert spread. Moreover, the effectiveness of the two core vector control interventions—insecticide-treated nets and indoor residual spraying—has been declining due to insecticide resistance and increased outdoor biting activity of mosquito vectors. These technical challenges, though varying from country to country, require integrated approaches and better understanding of the malaria epidemiology enabling targeted control of the parasites and vectors. Understanding the mechanism and distribution of drug-resistant parasites will allow effective drug treatment and prevent, or slow down, the spread of drug resistance. Coordination among the GMS countries is essential to prevent parasite reintroduction across the international borders to achieve regional malaria elimination.
Department of Entomology, Pennsylvania State University, USA
Yaming Cao
Department of Immunology, China Medical University, China
Jaranit Kaewkungwal
Faculty of Tropical Medicine, Mahidol University, Thailand
Amnat Khamsiriwatchara
Faculty of Tropical Medicine, Mahidol University, Thailand
Saranath Lawpoolsri
Faculty of Tropical Medicine, Mahidol University, Thailand
Than Naing Soe
Department of Public Health, Ministry of Health and Sports, Myanmar
Myat Phone Kyaw
Myanmar Medical Association, Myanmar
Jetsumon Sattabongkot
Faculty of Tropical Medicine, Mahidol University, Thailand
*Address all correspondence to: luc2@psu.edu
1. Introduction
With steady gains in the fight against malaria over the past decade, the international malaria community once again is embracing the global goal of malaria eradication. Meanwhile, the World Health Organization (WHO) has launched a new Global Technical Strategy for Malaria (http://www.who.int/malaria/areas/global_technical_strategy/en/) as the operational framework guiding malarious nations and regions in their pursuit of malaria elimination. In the Greater Mekong Subregion (GMS) of Southeast Asia (SEA), which comprises Cambodia, China’s Yunnan Province, Lao People’s Democratic Republic (Laos), Myanmar, Thailand, and Vietnam, malaria has been a significant public health problem and impediment to socioeconomic development [1, 2]. Intensified malaria control in recent years, fueled by increased international funding and local bustling economic development, has greatly reduced the regional malaria burden. Compared with confirmed malaria cases in 2010, the number of malaria cases in the GMS was reduced by ~50% in 2014. Driven by increasing political will and financial support and motivated by recent achievements in malaria control, the six GMS nations have endorsed a regional malaria elimination plan with an ultimate goal of eliminating Plasmodium falciparum malaria by 2025 and all malaria by 2030 [3]. Emerging artemisinin resistance in this region further escalated urgency for National Malaria Control Programmes (NMCPs) to make such a transition of their aims [4, 5]. However, this ambitious goal faces numerous technical challenges [6] and requires integrated strategies tailored to the whole region and individual countries. In the malaria elimination settings, control strategies need to align with the changing malaria epidemiology. Control measures such as long-lasting insecticide-treated bed nets (LLINs), indoor residual insecticide spraying (IRS), rapid diagnostic tests (RDTs), and artemisinin combined therapies (ACTs) used to effectively reduce malaria burden in hyperendemic regions may not be enough for the malaria elimination task. Additional tools such as mass drug administration (MDA) and innovative vector control programs may be needed. Here, we attempt to provide an updated view of the changing malaria epidemiology, the challenges, and prospect of malaria elimination in the GMS.
Malaria epidemiology in the GMS is complex and rapidly evolving. There is immense spatial heterogeneity in both regional and countrywide disease distribution (Figure 1 and Table 1). Within the GMS, Myanmar has the heaviest malaria burden and accounts for more than 53% of regionally confirmed malaria cases. Within each country, the pattern of malaria distribution remains similar, but transmission is still concentrated along international borders—the so-called border malaria. In border areas, there is poor accessibility to healthcare services, and surveillance for malaria is far less than optimal [8]. Given that these border regions represent probable malaria reservoirs and that importation and dispersal by migratory human populations are extremely difficult to monitor, border malaria constitutes one of the biggest obstacles for malaria elimination. Highly mobile populations crossing porous borders are a major contributor to parasite introduction and continued transmission [9]. Border areas also are home to ethnic minorities, hill tribes, temporary and seasonal migrants, refugees, and internally displaced people; many have poor educational level, limited access to healthcare services, and reduced legal rights. Geographical and cultural isolation leaves these groups at a high risk for infection and poor access to treatment [1, 2, 10, 11]. In Thailand, malaria makes up ~31% of communicable diseases diagnosed in migrants, as compared to 3% in Thai natives [12]. Heavy population flow along the extremely porous borders makes neighboring countries very vulnerable to malaria introduction and reintroduction [13, 14]. As a result, malaria prevalence on both sides of the border is often highly correlated [15]. In Yunnan Province of China, although autochthonous P. vivax malaria was still detected, P. falciparum infections were mostly associated with travel history to Myanmar [16]. There is also genetic evidence of asymmetric parasite flow from the more endemic to the less endemic side of the border [17, 18]. On a smaller geographical scale in a border village in Western Thailand, malaria incidence was clustered and significantly associated with citizen status indicating recent migration [19]. Moreover, there is a high probability that frequent border crossings by migrants will spread artemisinin-resistant P. falciparum [20, 21] beyond the “containment zone” [22, 23]. More sophisticated surveillance tools are needed to provide a clear picture of border malaria transmission so that targeted control measures are implemented to curb the spread of resistance and to prevent the reintroduction of parasites into populations where they have been eliminated. Thus, malaria elimination is a multinational, multipronged issue, with cross-border migration posing one of the largest threats to its success [24]. In recognition of this issue, the GMS countries have initiated bi- and multilateral coordination between the NMCPs. While the healthcare systems in the GMS countries are improving, further bolstering is needed to meet the malaria elimination challenge.
Figure 1.
The geographical proximity of countries and reported malaria cases for data based on 2016 in the Greater Mekong Subregion (GMS). Note: The majority of malaria cases in Yunnan Province of China were imported.
It has long been held as conventional wisdom that asymptomatic infections would be much less frequent in low-endemicity settings because the level of exposure-related immunity to malaria in human populations may be low [25]. However, asymptomatic infections represent the vast majority of infections in all endemic settings [26]. The use of molecular tools is essential for identifying submicroscopic infections. For both P. falciparum and P. vivax, microscopy detects only 1/3–1/2 of the infections detected by regular PCR [27, 28]. As the sensitivity of detection methods increases (e.g., with the use of a larger blood volume or reverse transcriptase-PCR targeting the parasite 18S rRNA), greater proportions of asymptomatic infections are discovered, revealing larger pools of infections [29, 30]. In Western Thailand and other GMS regions, qPCR and large-volume ultrasensitive qPCR could detect as much as 20% of the villagers harboring malaria infections as compared to ~5% detected by microscopy [31, 32]. Although we still do not have a clear picture about how much these asymptomatic infections actually contribute to malaria transmission in these areas [33], studies in Western Thailand have clearly demonstrated mosquito infectivity of submicroscopic P. falciparum and P. vivax [34], albeit the asymptomatic parasite carriers were found to be much less infective to mosquitoes than acute cases [35]. Since asymptomatic individuals are unlikely to seek treatment, they are missed by passive case detection, and submicroscopic infections also are missed by microscopy-based active case detection. It is highly possible that these asymptomatic infections act as important silent reservoirs of transmission. Even under such low-endemicity settings, it is estimated that submicroscopic carriers may be the source of 20–50% of all human-to-mosquito transmission [36], underlining the significance of managing this population in the malaria elimination phase. Therefore, information about the prevalence and seasonal dynamics of the asymptomatic infections in the border regions and their contribution to transmission is required to guide the efforts of NMCPs to achieve malaria elimination.
4. The burden of P. vivax malaria and G6PD deficiency
Another characteristic of the rapidly evolving malaria epidemiology in the GMS is that the prevalence of P. vivax is increasing proportionally to P. falciparum [37] (Table 1). The resilience of vivax malaria to control efforts may be attributed to some intrinsic biological features of this parasite. First, P. vivax only invades reticulocytes, and thus the resulting parasitemia is normally far lower than that of P. falciparum malaria. This makes microscopy-based diagnosis and RDTs not sufficiently sensitive in detecting P. vivax infections [38, 39, 40]. Second, during blood-stage infections with P. vivax, gametocytes are formed before the manifestation of clinical symptoms, which allows transmission of the parasite before treatment. Third, P. vivax develops dormant hypnozoites in the liver of the human host, which awaken in the weeks and months following a primary attack and cause relapses. Finally, vivax malaria is often transmitted by outdoor biting mosquitoes, making the current insecticide-based control measures (LLIN and IRS) less effective. Because of these unique features, traditional malaria control efforts often fail to control P. vivax transmission. In addition, containment of P. falciparum has been prioritized in the GMS, partially because of the emerging artemisinin resistance. As a result, P. falciparum prevalence has decreased, while the proportion of P. vivax has increased.
In the GMS, the first-line therapy for vivax malaria remains chloroquine (CQ) and primaquine (PQ) (Table 1) [41]. Reports of clinical CQ resistance in many regions of the world and falling efficacy of PQ are of great concern for vivax malaria control [42, 43, 44, 45]. Although some studies indicated that P. vivax in the GMS remained sensitive to CQ [46, 47, 48, 49, 50, 51], others clearly documented CQ-resistant P. vivax [52, 53, 54, 55]. In Myanmar, sporadic CQ-resistant P. vivax cases were first reported more than 20 years ago [52, 53]. A later report of 34% treatment failures in Dawei of Southern Myanmar suggests an increase of CQ resistance [55]. More recent studies identified both early and late treatment failures in Myawaddy of the Kayin State and Kawthaung of the Tanintharyi Region, Myanmar [56]. In northeastern Myanmar bordering China, a recent study showed 5.2% cumulative incidence of recurrent parasitemia during a 28-day follow-up of 587 P. vivax treated with CQ/PQ [57], suggesting sensitivity to CQ may also be deteriorating in this region. This reduced sensitivity of P. vivax to CQ requires close surveillance and potential implementation of more effective treatment measures such as ACTs [58].
Studies from Papua New Guinea suggest that 80% of the vivax infections may be attributed to relapses. A modeling approach predicts that as much as 96% of clinical attacks by P. vivax in Thailand are due to relapses [60]. For radical cure, WHO recommends a dose of 0.25–0.5 mg/kg of PQ daily for 14 days. However, the lower dose (total of 3.5 mg/kg) fails to prevent relapses in many different endemic sites [61]. Because of the potential risk of severe hemolysis that this drug could cause in patients with glucose-6-phosphate dehydrogenase (G6PD) deficiency, PQ is not widely prescribed [43, 62, 63]. In routine practice, G6PD status is not screened; the GMS nations still use the lower total dose of PQ in fear of the possible harm to those with G6PD deficiency. Because evaluation of PQ efficacy in preventing relapses requires longer-term follow-up, the clinical efficacy of the current PQ regimen for radical cure of vivax malaria in the GMS is unknown. Even with longer follow-ups, it is still not possible to reliably determine whether a recurrent infection after day 28 is due to relapse or reinfection given that a relapse infection may be from reactivation of a different hypnozoite clone [64, 65]. For PQ efficacy, host factors also need to be considered. Recently, failures of the PQ radical cure have been linked to reduced activity of the hepatic cytochrome P450 (CYP) 2D6 [66], which mediates activation of PQ to its active metabolite(s) [67, 68]. Different CYP2D6 activities have differential effects on the pharmacokinetics of PQ [69]. CYP2D6 is involved in the metabolism of as many as 25% of drugs in clinical use and is also a member of the CYP450 family with the greatest prevalence and genetic polymorphism [70, 71]. About 70 CYP2D6 allelic variants have been found and grouped into 4 phenotypic classes of ultra-rapid, extensive, intermediate, and abolished protein activity [72]. The frequency of alleles with reduced function is as high as 50% in most Asian populations [73]. Thus, it is important to determine the extent by which reduced CYP2D6 activity is responsible for PQ failures in radical cure of vivax malaria [74].
The G6PD gene is extraordinarily polymorphic with more than 400 variants discovered based on biochemical diagnosis [75], among which 186 mutations are associated with G6PD deficiency [76]. The prevalence of G6PD deficiency and distribution of G6PD variants vary geographically [77]. In the GMS, G6PD deficiency is often highly prevalent among ethnic groups. Along the Thailand-Myanmar border, the prevalence of G6PD deficiency was above 10% [78, 79, 80], whereas in the Kachin ethnicity along the China-Myanmar border, it almost reached 30% [81]. In Thailand and Myanmar, the Mahidol variant (487G>A) is the most predominant and often accounts for ~90% of all mutations [79, 81, 82, 83]. According to the WHO classification, the Mahidol variant is a Class III mutation or mild-deficient variant with 60% enzyme activity [76]. However, this classification may not be accurate since patients with the Mahidol variant often had <1% of the normal G6PD activity [79, 84, 85]. Patients having the G6PD Mahidol variant (487G>A) rarely had acute hemolytic anemia after taking the normal dose of PQ [84, 86]. In contrast to the belief that PQ only induces mild hemolysis in patients with the Mahidol variant, there have been case reports showing that the normal dosage of 15 mg/kg/day for 3 days in vivax patients with this G6PD variant could lead to acute hemolytic anemia that required blood transfusion or even cause renal failure [87, 88, 89]. It is noteworthy that G6PD activity can vary substantially between individuals with the same variant and even within the same individual over time. Therefore, with the prevalence of vivax malaria in this region and the goal of malaria elimination, the deployment of point-of-care G6PD deficiency diagnostics is urgent [90]. In addition, there is a need to test whether weekly PQ of 0.75 mg/kg for 8 weeks, a dosage considered safe for the G6PD African variant [91], could be prescribed in the GMS without prior testing for G6PD deficiency.
ACTs have played an indispensable role in reducing global malaria-associated mortality and morbidity. However, these achievements are threatened by the recent emergence of artemisinin resistance in P. falciparum in the GMS [92, 93, 94]. Artemisinin resistance is associated with a parasite clearance half-life of >5 h as compared to a normal value of ~2 h [94, 95, 96]. Clinical artemisinin resistance was first detected in western Cambodia [92, 93, 96, 97] but is now detected in other GMS regions including Thailand, Laos, Vietnam, Southern Myanmar, and the China-Myanmar border area [94, 95, 98, 99, 100, 101, 102, 103]. Out of fear of a catastrophic spread of artemisinin resistance to Africa, WHO deployed an artemisinin resistance containment plan in Cambodia [104]. Later, with the finding that artemisinin resistance has emerged independently in many areas of the GMS [105], the containment plan has been revised to a regional malaria elimination strategy [3, 4].
The principle of ACTs is that the fast-acting artemisinins rapidly reduce the parasite biomass, leaving the slow-eliminating partner drugs to clear the residual parasites. The emergence of artemisinin resistance means that a larger parasite mass is left for the partner drugs to clear after the usual 3-day ACT course, which increases the chance of resistance development to the partner drugs. Indeed, in the short period of time since the deployment of ACTs, clinical resistance to two ACTs, first artesunate/mefloquine [106] and more recently dihydroartemisinin/piperaquine (DHA/PPQ), has emerged in the GMS. These are the two most popular ACTs deployed in the GMS countries (Table 1). Since promising new antimalarials are still in the development pipeline, possible solutions to this problem include introduction of new ACTs, rotation of different ACTs, use of longer course of ACT treatment, and introduction of triple ACTs (artemisinin derivatives with two slow-eliminating partner drugs) [112]. To mitigate the threat of spread of artemisinin-resistant P. falciparum parasites, heightened surveillance is needed in sentinel sites of the GMS [113].
Tools for monitoring the epidemiology of antimalarial drug resistance include ex vivo or in vitro drug assays and molecular surveillance, which complement in vivo drug efficacy studies. It is noteworthy that the slow-clearance phenotype of clinical artemisinin resistance does not correspond to the 50% inhibitory concentrations of artemisinin drugs estimated from the conventional DNA replication-based in vitro assay but is better reflected in the newly developed ring-stage survival assay, which quantifies the number of early ring-stage parasites (0–3 h) that can survive the exposure to 700 nM of DHA for 6 h [114]. The discovery of mutations in the kelch domain protein K13 associated with artemisinin resistance provides a convenient molecular marker for a large-scale surveillance purpose [115]. To date, the correlations of K13 mutations with delayed parasite clearance have been established in several studies [95, 105, 115, 116, 117] but only a very limited number of K13 mutations were confirmed to confer in vitro artemisinin resistance through genetic manipulations [118, 119]. The K13 gene in the world P. falciparum populations harbors more than 108 nonsynonymous mutations, which showed marked geographic disparity in frequency and distribution [120]. Similarly, K13 mutations in the GMS also showed highly heterogeneous distribution [103, 121, 122, 123, 124, 125], possibly reflecting different drug histories and evolutionary origins of the parasite populations [126]. Clinical failures of DHA/PPQ have been associated with increased in vitro PPQ resistance and the molecular markers of PPQ resistance in western Cambodia include amplification of the aspartic protease genes plasmepsin 2–3 and point mutation E415G in an exonuclease gene (PF3D7_12362500) [127, 128]. Molecular surveillance of artemisinin resistance in western Cambodia, Thailand, and Laos has detected the spread of a parasite clone with a long K13 haplotype carrying the C580Y mutation (the artemisinin-resistant mutation reaching near fixation in western Cambodia) to northeastern Thailand and southern Laos, which indicates a transnational selective sweep [129]. Importantly, this parasite lineage also harbors the plasmepsin 2 amplification, which may preclude further use of DHA/PPQ in this region. In addition, this situation also necessitates implementation of stringent follow-ups of malaria cases after ACT treatment to ensure that recrudescent cases are treated with effective antimalarials. Thus, surveillance should be mandatory to delay the spread of the resistant parasites and to accelerate malaria elimination in the GMS.
LLINs and IRS are the key vector-based malaria interventions that have been found to be highly effective in sub-Saharan Africa. However, these measures are much less efficient in the GMS [130]. The GMS has a complex vector system; most of the malaria vectors belong to species complexes or groups such as Dirus, Minimus, Maculatus, and Sundaicus, which vary significantly in terms of geographic distribution, ecology, behavior, and vectorial competence [131, 132, 133]. At least 19 species are known malaria vectors, some of which comprise cryptic species complexes [132]. In order to apply the appropriate control approaches in relation to the biology of the vector species, we first need to identify the mosquitoes to their species level and to differentiate the vector from nonvector species, which requires molecular assays [134]. These vector species display significant variations in geographical distribution and seasonal dynamics, and accordingly their roles in malaria transmission also vary in space and time [135]. In many endemic areas of the GMS, perennial malaria transmission is maintained by Anopheles dirus during the rainy season and An. minimus during the drier periods of the year [132, 136]. Environmental changes such as deforestation have caused changes in the vector species composition [137, 138] and benefited the survivorship of major vectors [40]. Since many of these vector species exhibit early evening and outdoor biting preferences, LLINs alone are not sufficient for interrupting malaria transmission [140]. In addition, the emergence and spread of insecticide resistance further compromise the effectiveness of the mosquito control measures [141, 142, 143].
7. Technological innovation for malaria elimination
The technical challenges discussed here suggest that the currently used malaria control tools (RDT, ACT, LLIN, and IRS) that were instrumental for the gains against malaria may not be sufficient for malaria elimination [144]. Additional tools are needed to achieve the final goal of malaria elimination in the GMS. First, residual transmission requires MDA to eliminate asymptomatic and submicroscopic parasite reservoirs. For the success of MDA, better knowledge of malaria epidemiology is needed so that targeted MDA can be implemented. Successful MDA programs also require strong community engagement. MDA has proved successful in eliminating malaria in Asia-Pacific regions such as Vanuatu and central China [145, 146]. In an earlier study conducted in Cambodian villages, MDA of artemisinin-PPQ at 10-day intervals for 6 months drastically reduced P. falciparum rates [147]. A recent pilot MDA study conducted in villages of Kayin State, Myanmar, showed that a 3-day supervised course of DHA/PPQ was well tolerated and highly effective in reducing asymptomatic P. falciparum carriage, whereas the effect on reducing P. vivax was transient presumably due to relapse [148]. Thus, drugs targeting the P. vivax hypnozoite reservoir are required for MDA in the GMS, where P. vivax is becoming the predominant parasite species [149]. The high prevalence of G6PD deficiency in the target populations demands prescreening using a point-of-care diagnostic for G6PD deficiency. From a programmatic standpoint, such an operation requires substantial financial commitment. Second, effective management of malaria cases in the face of emergence and spread of drug resistance requires new therapies such as triple ACTs. Third, novel vector control approaches are desperately needed including larval control strategies [150], incorporation of ivermectin in the MDA program to reduce the life span of mosquitoes [151, 152], topical and spatial repellents against outdoor biting vectors [153, 154], genetically manipulated mosquitoes for population replacement [155], and next generation of LLINs and IRS [156]. It is imperative that new interventions are continuously developed and integrated into malaria elimination programs.
Malaria elimination in the GMS carries the urgency of eliminating artemisinin-resistant P. falciparum parasites before they become untreatable and spread to Africa. The changing malaria epidemiology with increasing proportion of P. vivax malaria requires an 8-aminoquinoline drug for radical cure, but it demands deployment of point-of-care diagnostics for G6PD deficiency due to its high prevalence in endemic human populations. In addition, the prevalent asymptomatic parasite reservoirs need to be targeted by a MDA approach. The diversity of Anopheles vectors in the GMS and decreasing effectiveness of indoor control measures, such as LLIN and IRS facing the outdoor malaria transmission, also require development and implementation of novel interventions for vector control. To meet the challenge of border malaria, coordinated efforts among the NMCPs targeting the mobile and migrant populations along international borders will prevent cross-border reintroduction of malaria. Altogether, a holistic attack on malaria using integrated approaches is necessary to achieve the goal of regional malaria elimination in the GMS.
We thank the National Institute of Allergy and Infectious Diseases, NIH, for financial support (U19AI089672).
References
1.Cui L, Yan G, Sattabongkot J, Cao Y, Chen B, Chen X, et al. Malaria in the greater Mekong subregion: Heterogeneity and complexity. Acta Tropica. 2012;121:227-239
2.Hewitt S, Delacollette C, Chavez I. Malaria situation in the greater Mekong subregion. The Southeast Asian Journal of Tropical Medicine and Public Health. 2013;44(Suppl 1):46-72; discussion 306-307
3.WHO. Strategy for malaria elimination in the greater Mekong subregion (2015-2030). http://iris.wpro.who.int/bitstream/handle/10665.1/10945/9789290617181_eng.pdf [Accessed: January 21, 2018]
4.Smith Gueye C, Newby G, Hwang J, Phillips AA, Whittaker M, MacArthur JR, et al. The challenge of artemisinin resistance can only be met by eliminating Plasmodium falciparum malaria across the greater Mekong subregion. Malaria Journal. 2014;13:286
5.Fidock DA. Microbiology. Eliminating malaria. Science. 2013;340:1531-1533
6.Hewitt S, Delacollette C, Poirot E. Malaria control in the greater Mekong subregion: An overview of the current response and its limitations. The Southeast Asian Journal of Tropical Medicine and Public Health. 2013;44(Suppl 1):249-305; discussion 306-247
7.WHO. World Malaria Report 2017. http://www.who.int/malaria/publications/world-malaria-report-2017/en/ [Accessed: January 21, 2018]
8.Liu H, Nie R-H, Li C-F, Sun Y-H, Li G-S. Active detection of malaria cases in Myanmar Wa ethnical villages of China-Myanmar border. Parasitoses and Infectious Diseases. 2009;7:6-9
9.Gueye CS, Teng A, Kinyua K, Wafula F, Gosling R, McCoy D. Parasites and vectors carry no passport: How to fund cross-border and regional efforts to achieve malaria elimination. Malaria Journal. 2012;11:344
10.Parker DM, Carrara VI, Pukrittayakamee S, McGready R, Nosten FH. Malaria ecology along the Thailand-Myanmar border. Malaria Journal. 2015;14:388
11.Xu J, Liu H. The challenges of malaria elimination in Yunnan Province, People's Republic of China. The Southeast Asian Journal of Tropical Medicine and Public Health. 2012;43:819-824
12.Huguet JW, Chamratrithirong A. Thailand Migration Report. Bangkok, Thailand: International Organization for Migration, Thailand Office; 2011 http://publications.iom.int/system/files/pdf/tmr_2011.pdf [Accessed: January 21, 2018]
13.Jitthai N. Migration and malaria. The Southeast Asian Journal of Tropical Medicine and Public Health. 2013;44(Suppl 1):166-200; discussion 306-167
14.Guyant P, Canavati SE, Chea N, Ly P, Whittaker MA, Roca-Feltrer A, et al. Malaria and the mobile and migrant population in Cambodia: A population movement framework to inform strategies for malaria control and elimination. Malaria Journal. 2015;14:252
15.Xu JW, Liu H. The relationship of malaria between Chinese side and Myanmar's five special regions along China-Myanmar border: A linear regression analysis. Malaria Journal. 2016;15:368
16.Zhou G, Sun L, Xia R, Duan Y, Xu J, Yang H, et al. Clinical malaria along the China-Myanmar border, Yunnan Province, China, January 2011-August 2012. Emerging Infectious Diseases. 2014;20:675-678
17.Lo E, Lam N, Hemming-Schroeder E, Nguyen J, Zhou G, Lee MC, et al. Frequent spread of Plasmodium vivax malaria maintains high genetic diversity at the Myanmar-China border without distance and landscape barriers. The Journal of Infectious Diseases. 2017;216(10):1254-1263
18.Lo E, Zhou G, Oo W, Lee MC, Baum E, Felgner PL, et al. Molecular inference of sources and spreading patterns of Plasmodium falciparum malaria parasites in internally displaced persons settlements in Myanmar-China border area. Infection, Genetics and Evolution. 2015;33:189-196
19.Parker DM, Matthews SA, Yan G, Zhou G, Lee MC, Sirichaisinthop J, et al. Microgeography and molecular epidemiology of malaria at the Thailand-Myanmar border in the malaria pre-elimination phase. Malaria Journal. 2015;14:198
20.Wangroongsarb P, Satimai W, Khamsiriwatchara A, Thwing J, Eliades JM, Kaewkungwal J, et al. Respondent-driven sampling on the Thailand-Cambodia border. II. Knowledge, perception, practice and treatment-seeking behaviour of migrants in malaria endemic zones. Malaria Journal. 2011;10:117
21.Wangroongsarb P, Sudathip P, Satimai W. Characteristics and malaria prevalence of migrant populations in malaria-endemic areas along the Thai-Cambodian border. The Southeast Asian Journal of Tropical Medicine and Public Health. 2012;43:261-269
22.Khamsiriwatchara A, Sudathip P, Sawang S, Vijakadge S, Potithavoranan T, Sangvichean A, et al. Artemisinin resistance containment project in Thailand. (I): Implementation of electronic-based malaria information system for early case detection and individual case management in provinces along the Thai-Cambodian border. Malaria Journal. 2012;11:247
23.Khamsiriwatchara A, Wangroongsarb P, Thwing J, Eliades J, Satimai W, Delacollette C, et al. Respondent-driven sampling on the Thailand-Cambodia border. I. Can malaria cases be contained in mobile migrant workers? Malaria Journal. 2011;10:120
24.Sturrock HJ, Roberts KW, Wegbreit J, Ohrt C, Gosling RD. Tackling imported malaria: An elimination endgame. The American Journal of Tropical Medicine and Hygiene. 2015;93:139-144
25.Ghani AC, Sutherland CJ, Riley EM, Drakeley CJ, Griffin JT, Gosling RD, et al. Loss of population levels of immunity to malaria as a result of exposure-reducing interventions: Consequences for interpretation of disease trends. PLoS One. 2009;4:e4383
26.Lin JT, Saunders DL, Meshnick SR. The role of submicroscopic parasitemia in malaria transmission: What is the evidence? Trends in Parasitology. 2014;30:183-190
27.Okell LC, Ghani AC, Lyons E, Drakeley CJ. Submicroscopic infection in Plasmodium falciparum-endemic populations: A systematic review and meta-analysis. The Journal of Infectious Diseases. 2009;200:1509-1517
28.Moreira CM, Abo-Shehada M, Price RN, Drakeley CJ. A systematic review of sub-microscopic Plasmodium vivax infection. Malaria Journal. 2015;14:360
29.Imwong M, Hanchana S, Malleret B, Renia L, Day NP, Dondorp A, et al. High-throughput ultrasensitive molecular techniques for quantifying low-density malaria parasitemias. Journal of Clinical Microbiology. 2014;52:3303-3309
30.Zhao Y, Lv Y, Liu F, Wang Q, Li P, Zhao Z, et al. Comparison of methods for detecting asymptomatic malaria infections in the China-Myanmar border area. Malaria Journal. 2017;16:159
31.Imwong M, Nguyen TN, Tripura R, Peto TJ, Lee SJ, Lwin KM, et al. The epidemiology of subclinical malaria infections in South-East Asia: Findings from cross-sectional surveys in Thailand-Myanmar border areas, Cambodia, and Vietnam. Malaria Journal. 2015;14:381
32.Baum E, Sattabongkot J, Sirichaisinthop J, Kiattibutr K, Davies DH, Jain A, et al. Submicroscopic and asymptomatic Plasmodium falciparum and Plasmodium vivax infections are common in western Thailand—Molecular and serological evidence. Malaria Journal. 2015;14:95
33.Pethleart A, Prajakwong S, Suwonkerd W, Corthong B, Webber R, Curtis C. Infectious reservoir of Plasmodium infection in Mae Hong Son Province, north-west Thailand. Malaria Journal. 2004;3:34
34.Coleman RE, Kumpitak C, Ponlawat A, Maneechai N, Phunkitchar V, Rachapaew N, et al. Infectivity of asymptomatic Plasmodium-infected human populations to Anophelesdirus mosquitoes in western Thailand. Journal of Medical Entomology. 2004;41:201-208
35.Kiattibutr K, Roobsoong W, Sriwichai P, Saeseu T, Rachaphaew N, Suansomjit C, et al. Infectivity of symptomatic and asymptomatic Plasmodium vivax infections to a southeast Asian vector, Anophelesdirus. International Journal for Parasitology. 2017;47:163-170
36.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
37.Cotter C, Sturrock HJ, Hsiang MS, Liu J, Phillips AA, Hwang J, et al. The changing epidemiology of malaria elimination: New strategies for new challenges. Lancet. 2013;382:900-911
38.Coleman RE, Maneechai N, Ponlawat A, Kumpitak C, Rachapaew N, Miller RS, et al. Short report: Failure of the OptiMAL rapid malaria test as a tool for the detection of asymptomatic malaria in an area of Thailand endemic for Plasmodium falciparum and P. vivax. The American Journal of Tropical Medicine and Hygiene. 2002;67:563-565
39.Yan J, Li N, Wei X, Li P, Zhao Z, Wang L, et al. Performance of two rapid diagnostic tests for malaria diagnosis at the China-Myanmar border area. Malaria Journal. 2013;12:73
40.Wongsrichanalai C, Barcus MJ, Muth S, Sutamihardja A, Wernsdorfer WH. A review of malaria diagnostic tools: Microscopy and rapid diagnostic test (RDT). The American Journal of Tropical Medicine and Hygiene. 2007;77:119-127
41.Baird KJ, Maguire JD, Price RN. Diagnosis and treatment of Plasmodium vivax malaria. Advances in Parasitology. 2012;80:203-270
42.Baird JK. Resistance to therapies for infection by Plasmodium vivax. Clinical Microbiology Reviews. 2009;22:508-534
43.Baird JK, Rieckmann KH. Can primaquine therapy for vivax malaria be improved? Trends in Parasitology. 2003;19:115-120
44.Goller JL, Jolley D, Ringwald P, Biggs BA. Regional differences in the response of Plasmodium vivax malaria to primaquine as anti-relapse therapy. The American Journal of Tropical Medicine and Hygiene. 2007;76:203-207
45.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;14:982-991
46.Luxemburger C, van Vugt M, Jonathan S, McGready R, Looareesuwan S, White NJ, et al. Treatment of vivax malaria on the western border of Thailand. Transactions of the Royal Society of Tropical Medicine and Hygiene. 1999;93:433-438
47.Pukrittayakamee S, Chantra A, Simpson JA, Vanijanonta S, Clemens R, Looareesuwan S, et al. Therapeutic responses to different antimalarial drugs in vivax malaria. Antimicrobial Agents and Chemotherapy. 2000;44:1680-1685
48.Tasanor O, Ruengweerayut R, Sirichaisinthop J, Congpuong K, Wernsdorfer WH, Na-Bangchang K. Clinical-parasitological response and in-vitro sensitivity of Plasmodium vivax to chloroquine and quinine on the western border of Thailand. Transactions of the Royal Society of Tropical Medicine and Hygiene. 2006;100:410-418
49.Muhamad P, Ruengweerayut R, Chacharoenkul W, Rungsihirunrat K, Na-Bangchang K. Monitoring of clinical efficacy and in vitro sensitivity of Plasmodium vivax to chloroquine in area along Thai Myanmar border during 2009-2010. Malaria Journal. 2011;10:44
50.Vijaykadga S, Rojanawatsirivej C, Congpoung K, Wilairatana P, Satimai W, Uaekowitchai C, et al. Assessment of therapeutic efficacy of chloroquine for vivax malaria in Thailand. The Southeast Asian Journal of Tropical Medicine and Public Health. 2004;35:566-569
51.Congpoung K, Satimai W, Sujariyakul A, Intanakom S, Harnpitakpong W, Pranuth Y, et al. In vivo sensitivity monitoring of chloroquine for the treatment of uncomplicated vivax malaria in four bordered provinces of Thailand during 2009-2010. Journal of Vector Borne Diseases. 2011;48:190-196
52.Myat-Phone-Kyaw, Myint-Oo, Myint-Lwin, Thaw-Zin, Kyin-Hla-Aye, Nwe-Nwe-Yin. Emergence of chloroquine-resistant Plasmodium vivax in Myanmar (Burma). Transactions of the Royal Society of Tropical Medicine and Hygiene. 1993;87:687
53.Marlar T, Myat Phone K, Aye Yu S, Khaing Khaing G, Ma S, Myint O. Development of resistance to chloroquine by Plasmodium vivax in Myanmar. Transactions of the Royal Society of Tropical Medicine and Hygiene. 1995;89:307-308
54.Rijken MJ, Boel ME, Russell B, Imwong M, Leimanis ML, Phyo AP, et al. Chloroquine resistant vivax malaria in a pregnant woman on the western border of Thailand. Malaria Journal. 2011;10:113
55.Guthmann JP, Pittet A, Lesage A, Imwong M, Lindegardh N, Min Lwin M, et al. Plasmodium vivax resistance to chloroquine in Dawei, southern Myanmar. Tropical Medicine & International Health. 2008;13:91-98
56.Htun MW, Mon NCN, Aye KM, Hlaing CM, Kyaw MP, Handayuni I, et al. Chloroquine efficacy for Plasmodium vivax in Myanmar in populations with high genetic diversity and moderate parasite gene flow. Malaria Journal. 2017;16:281
57.Yuan L, Wang Y, Parker DM, Gupta B, Yang Z, Liu H, et al. Therapeutic responses of Plasmodium vivax malaria to chloroquine and primaquine treatment in northeastern Myanmar. Antimicrobial Agents and Chemotherapy. 2015;59:1230-1235
58.Baird JK. Resistance to chloroquine unhinges vivax malaria therapeutics. Antimicrobial Agents and Chemotherapy. 2011;55:1827-1830
59.Robinson LJ, Wampfler R, Betuela I, Karl S, White MT, Li Wai Suen CS, 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 Medicine. 2015;12:e1001891
60.Adekunle AI, Pinkevych M, McGready R, Luxemburger C, White LJ, Nosten F, et al. Modeling the dynamics of Plasmodium vivax infection and hypnozoite reactivation in vivo. PLoS Neglected Tropical Diseases. 2015;9:e0003595
61.White NJ. Determinants of relapse periodicity in Plasmodium vivax malaria. Malaria Journal. 2011;10:297
62.Butcher GA. Antimalarial drugs and the mosquito transmission of Plasmodium. International Journal for Parasitology. 1997;27:975-987
63.White NJ. The role of anti-malarial drugs in eliminating malaria. Malaria Journal. 2008;7(Suppl 1):S8
64.Imwong M, Snounou G, Pukrittayakamee S, Tanomsing N, Kim JR, Nandy A, et al. Relapses of Plasmodium vivax infection usually result from activation of heterologous hypnozoites. The Journal of Infectious Diseases. 2007;195:927-933
65.Chen N, Auliff A, Rieckmann K, Gatton M, Cheng Q. Relapses of Plasmodium vivax infection result from clonal hypnozoites activated at predetermined intervals. The Journal of Infectious Diseases. 2007;195:934-941
66.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. The New England Journal of Medicine. 2013;369:1381-1382
67.Pybus BS, Marcsisin SR, Jin X, Deye G, Sousa JC, Li Q, et al. The metabolism of primaquine to its active metabolite is dependent on CYP 2D6. Malaria Journal. 2013;12:212
68.Pybus BS, Sousa JC, Jin X, Ferguson JA, Christian RE, Barnhart R, et al. CYP450 phenotyping and accurate mass identification of metabolites of the 8-aminoquinoline, anti-malarial drug primaquine. Malaria Journal. 2012;11:259
70.Zhou SF, Liu JP, Lai XS. Substrate specificity, inhibitors and regulation of human cytochrome P450 2D6 and implications in drug development. Current Medicinal Chemistry. 2009;16:2661-2805
71.Zhou SF. Polymorphism of human cytochrome P450 2D6 and its clinical significance: Part I. Clinical Pharmacokinetics. 2009;48:689-723
72.Ingelman-Sundberg M, Sim SC, Gomez A, Rodriguez-Antona C. Influence of cytochrome P450 polymorphisms on drug therapies: Pharmacogenetic, pharmacoepigenetic and clinical aspects. Pharmacology & Therapeutics. 2007;116:496-526
73.Bradford LD. CYP2D6 allele frequency in European Caucasians, Asians, Africans and their descendants. Pharmacogenomics. 2002;3:229-243
74.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
75.Nkhoma ET, Poole C, Vannappagari V, Hall SA, Beutler E. The global prevalence of glucose-6-phosphate dehydrogenase deficiency: A systematic review and meta-analysis. Blood Cells, Molecules & Diseases. 2009;42:267-278
76.Minucci A, Moradkhani K, Hwang MJ, Zuppi C, Giardina B, Capoluongo E. Glucose-6-phosphate dehydrogenase (G6PD) mutations database: Review of the “old” and update of the new mutations. Blood Cells, Molecules & Diseases. 2012;48:154-165
77.Howes RE, Dewi M, Piel FB, Monteiro WM, Battle KE, Messina JP, et al. Spatial distribution of G6PD deficiency variants across malaria-endemic regions. Malaria Journal. 2013;12:418
78.Charoenkwan P, Tantiprabha W, Sirichotiyakul S, Phusua A, Sanguansermsri T. Prevalence and molecular characterization of glucose-6-phosphate dehydrogenase deficiency in northern Thailand. The Southeast Asian Journal of Tropical Medicine and Public Health. 2014;45:187-193
79.Bancone G, Chu CS, Somsakchaicharoen R, Chowwiwat N, Parker DM, Charunwatthana P, et al. Characterization of G6PD genotypes and phenotypes on the northwestern Thailand-Myanmar border. PLoS One. 2014;9:e116063
80.Kotepui M, Uthaisar K, PhunPhuech B, Phiwklam N. Prevalence and hematological indicators of G6PD deficiency in malaria-infected patients. Infectious Diseases of Poverty. 2016;5:36
81.Li Q, Yang F, Liu R, Luo L, Yang Y, Zhang L, et al. Prevalence and molecular characterization of glucose-6-phosphate dehydrogenase deficiency at the China-Myanmar border. PLoS One. 2015;10:e0134593
82.Matsuoka H, Wang J, Hirai M, Arai M, Yoshida S, Kobayashi T, et al. Glucose-6-phosphate dehydrogenase (G6PD) mutations in Myanmar: G6PD Mahidol (487G>A) is the most common variant in the Myanmar population. Journal of Human Genetics. 2004;49:544-547
83.Nuchprayoon I, Sanpavat S, Nuchprayoon S. Glucose-6-phosphate dehydrogenase (G6PD) mutations in Thailand: G6PD Viangchan (871G>A) is the most common deficiency variant in the Thai population. Human Mutation. 2002;19:185
84.Charoenlarp P, Areekul S, Pholpothi T, Harinasuta T. The course of primaquine-induced haemolysis in G-6-PD-deficient Thais. Journal of the Medical Association of Thailand. 1973;56:392-397
85.Charoenlarp P, Areekul S, Harinasuta T, Sirivorasarn P. The haemolytic effect of a single dose of 45 mg of primaquine in G-6-PD deficient Thais. Journal of the Medical Association of Thailand. 1972;55:631-638
86.Buchachart K, Krudsood S, Singhasivanon P, Treeprasertsuk S, Phophak N, Srivilairit S, et al. Effect of primaquine standard dose (15 mg/day for 14 days) in the treatment of vivax malaria patients in Thailand. The Southeast Asian Journal of Tropical Medicine and Public Health. 2001;32:720-726
87.Karwacki JJ, Shanks GD, Kummalue T, Watanasook C. Primaquine induced hemolysis in a Thai soldier. The Southeast Asian Journal of Tropical Medicine and Public Health. 1989;20:555-556
88.Kheng S, Muth S, Taylor WR, Tops N, Kosal K, Sothea K, et al. Tolerability and safety of weekly primaquine against relapse of Plasmodium vivax in Cambodians with glucose-6-phosphate dehydrogenase deficiency. BMC Medicine. 2015;13:203
89.Chen X, He Y, Miao Y, Yang Z, Cui L. A young man with severe acute haemolytic anaemia. BMJ. 2017;359:j4263
90.Domingo GJ, Satyagraha AW, Anvikar A, Baird K, Bancone G, Bansil P, et al. G6PD testing in support of treatment and elimination of malaria: Recommendations for evaluation of G6PD tests. Malaria Journal. 2013;12:391
91.Alving AS, Johnson CF, Tarlov AR, Brewer GJ, Kellermeyer RW, Carson PE. Mitigation of the haemolytic effect of primaquine and enhancement of its action against exoerythrocytic forms of the Chesson strain of Plasmodium vivax by intermittent regimens of drug administration: A preliminary report. Bulletin of the World Health Organization. 1960;22:621-631
92.Noedl H, Se Y, Schaecher K, Smith BL, Socheat D, Fukuda MM. Evidence of artemisinin-resistant malaria in western Cambodia. The New England Journal of Medicine. 2008;359:2619-2620
93.Dondorp AM, Nosten F, Yi P, Das D, Phyo AP, Tarning J, et al. Artemisinin resistance in Plasmodium falciparum malaria. The New England Journal of Medicine. 2009;361:455-467
94.Phyo AP, Nkhoma S, Stepniewska K, Ashley EA, Nair S, McGready R, et al. Emergence of artemisinin-resistant malaria on the western border of Thailand: A longitudinal study. Lancet. 2012;379:1960-1966
95.Ashley EA, Dhorda M, Fairhurst RM, Amaratunga C, Lim P, Suon S, et al. Spread of artemisinin resistance in Plasmodium falciparum malaria. The New England Journal of Medicine. 2014;371:411-423
96.Amaratunga C, Sreng S, Suon S, Phelps ES, Stepniewska K, Lim P, et al. Artemisinin-resistant Plasmodium falciparum in Pursat province, western Cambodia: A parasite clearance rate study. The Lancet Infectious Diseases. 2012;12:851-858
97.Noedl H, Se Y, Sriwichai S, Schaecher K, Teja-Isavadharm P, Smith B, et al. Artemisinin resistance in Cambodia: A clinical trial designed to address an emerging problem in Southeast Asia. Clinical Infectious Diseases. 2010;51:e82-e89
98.Hien TT, Thuy-Nhien NT, Phu NH, Boni MF, Thanh NV, Nha-Ca NT, et al. In vivo susceptibility of Plasmodium falciparum to artesunate in Binh Phuoc Province, Vietnam. Malaria Journal. 2012;11:355
99.Bustos MD, Wongsrichanalai C, Delacollette C, Burkholder B. Monitoring antimalarial drug efficacy in the greater Mekong subregion: An overview of in vivo results from 2008 to 2010. The Southeast Asian Journal of Tropical Medicine and Public Health. 2013;44(Suppl 1):201-230; discussion 306-207
100.Kyaw MP, Nyunt MH, Chit K, Aye MM, Aye KH, Lindegardh N, et al. Reduced susceptibility of Plasmodium falciparum to artesunate in southern Myanmar. PLoS One. 2013;8:e57689
101.Carrara VI, Lwin KM, Phyo AP, Ashley E, Wiladphaingern J, Sriprawat K, et al. Malaria burden and artemisinin resistance in the mobile and migrant population on the Thai-Myanmar border, 1999-2011: An observational study. PLoS Medicine. 2013;10:e1001398
102.Thriemer K, Hong NV, Rosanas-Urgell A, Phuc BQ, Ha do M, Pockele E, et al. Delayed parasite clearance after treatment with dihydroartemisinin-piperaquine in Plasmodium falciparum malaria patients in Central Vietnam. Antimicrobial Agents and Chemotherapy. 2014;58:7049-7055
103.Huang F, Takala-Harrison S, Jacob CG, Liu H, Sun X, Yang H, et al. A single mutation in K13 predominates in southern China and is associated with delayed clearance of Plasmodium falciparum following artemisinin treatment. The Journal of Infectious Diseases. 2015;212:1629-1635
104.WHO. Global plan for artemisinin resistance containment (GPARC). 2011. p. 87
105.Takala-Harrison S, Jacob CG, Arze C, Cummings MP, Silva JC, Dondorp AM, et al. Independent emergence of artemisinin resistance mutations among Plasmodium falciparum in Southeast Asia. The Journal of Infectious Diseases. 2015;211:670-679
106.Wongsrichanalai C, Meshnick SR. Declining artesunate-mefloquine efficacy against falciparum malaria on the Cambodia-Thailand border. Emerging Infectious Diseases. 2008;14:716-719
107.Amaratunga C, Lim P, Suon S, Sreng S, Mao S, Sopha C, et al. Dihydroartemisinin-piperaquine resistance in Plasmodium falciparum malaria in Cambodia: A multisite prospective cohort study. The Lancet Infectious Diseases. 2016;16:357-365
108.Saunders DL, Vanachayangkul P, Lon C. Dihydroartemisinin-piperaquine failure in Cambodia. The New England Journal of Medicine. 2014;371:484-485
109.Leang R, Taylor WR, Bouth DM, Song L, Tarning J, Char MC, et al. Evidence of Plasmodium falciparum malaria multidrug resistance to Artemisinin and Piperaquine in western Cambodia: Dihydroartemisinin-Piperaquine open-label multicenter clinical assessment. Antimicrobial Agents and Chemotherapy. 2015;59:4719-4726
110.Spring MD, Lin JT, Manning JE, Vanachayangkul P, Somethy S, Bun R, et al. Dihydroartemisinin-piperaquine failure associated with a triple mutant including kelch13 C580Y in Cambodia: An observational cohort study. The Lancet Infectious Diseases. 2015;15:683-691
111.Phuc BQ, Rasmussen C, Duong TT, Dong LT, Loi MA, Menard D, et al. Treatment failure of dihydroartemisinin/piperaquine for Plasmodium falciparum malaria, Vietnam. Emerging Infectious Diseases. 2017;23:715-717
112.Menard D, Dondorp A. Antimalarial drug resistance: A threat to malaria elimination. Cold Spring Harbor Perspectives in Medicine. 2017;7:a025619
113.Cui L, Mharakurwa S, Ndiaye D, Rathod PK, Rosenthal PJ. Antimalarial drug resistance: Literature review and activities and findings of the ICEMR network. The American Journal of Tropical Medicine and Hygiene. 2015;93:57-68
114.Witkowski B, Amaratunga C, Khim N, Sreng S, Chim P, Kim S, et al. Novel phenotypic assays for the detection of artemisinin-resistant Plasmodium falciparum malaria in Cambodia: In-vitro and ex-vivo drug-response studies. The Lancet Infectious Diseases. 2013;13:1043-1049
115.Ariey F, Witkowski B, Amaratunga C, Beghain J, Langlois AC, Khim N, et al. A molecular marker of artemisinin-resistant Plasmodium falciparum malaria. Nature. 2014;505:50-55
116.Nyunt MH, Hlaing T, Oo HW, Tin-Oo LL, Phway HP, Wang B, et al. Molecular assessment of artemisinin resistance markers, polymorphisms in the k13 propeller, and a multidrug-resistance gene in the eastern and western border areas of Myanmar. Clinical Infectious Diseases. 2015;60:1208-1215
117.Miotto O, Amato R, Ashley EA, MacInnis B, Almagro-Garcia J, Amaratunga C, et al. Genetic architecture of artemisinin-resistant Plasmodium falciparum. Nature Genetics. 2015;47:226-234
118.Ghorbal M, Gorman M, Macpherson CR, Martins RM, Scherf A, Lopez-Rubio JJ. Genome editing in the human malaria parasite Plasmodium falciparum using the CRISPR-Cas9 system. Nature Biotechnology. 2014;32:819-821
119.Straimer J, Gnadig NF, Witkowski B, Amaratunga C, Duru V, Ramadani AP, et al. Drug resistance. K13-propeller mutations confer artemisinin resistance in Plasmodium falciparum clinical isolates. Science. 2015;347:428-431
120.Menard D, Khim N, Beghain J, Adegnika AA, Shafiul-Alam M, Amodu O, et al. A worldwide map of Plasmodium falciparum K13-propeller polymorphisms. The New England Journal of Medicine. 2016;374:2453-2464
121.Wang Z, Shrestha S, Li X, Miao J, Yuan L, Cabrera M, et al. Prevalence of K13-propeller polymorphisms in Plasmodium falciparum from China-Myanmar border in 2007-2012. Malaria Journal. 2015;14:168
122.Tun KM, Imwong M, Lwin KM, Win AA, Hlaing TM, Hlaing T, et al. Spread of artemisinin-resistant Plasmodium falciparum in Myanmar: A cross-sectional survey of the K13 molecular marker. The Lancet Infectious Diseases. 2015;15:415-421
123.Wang Z, Wang Y, Cabrera M, Zhang Y, Gupta B, Wu Y, et al. Artemisinin resistance at the China-Myanmar border and association with mutations in the K13 propeller gene. Antimicrobial Agents and Chemotherapy. 2015;59:6952-6959
124.Ye R, Hu D, Zhang Y, Huang Y, Sun X, Wang J, et al. Distinctive origin of artemisinin-resistant Plasmodium falciparum on the China-Myanmar border. Scientific Reports. 2016;6:20100
125.Putaporntip C, Kuamsab N, Kosuwin R, Tantiwattanasub W, Vejakama P, Sueblinvong T, et al. Natural selection of K13 mutants of Plasmodium falciparum in response to artemisinin combination therapies in Thailand. Clinical Microbiology and Infection. 2016;22:285 e281-285 e288
126.Talundzic E, Okoth SA, Congpuong K, Plucinski MM, Morton L, Goldman IF, et al. Selection and spread of artemisinin-resistant alleles in Thailand prior to the global artemisinin resistance containment campaign. PLoS Pathogens. 2015;11:e1004789
127.Witkowski B, Duru V, Khim N, Ross LS, Saintpierre B, Beghain J, et al. A surrogate marker of piperaquine-resistant Plasmodium falciparum malaria: A phenotype-genotype association study. The Lancet Infectious Diseases. 2017;17:174-183
128.Amato R, Lim P, Miotto O, Amaratunga C, Dek D, Pearson RD, et al. Genetic markers associated with dihydroartemisinin-piperaquine failure in Plasmodium falciparum malaria in Cambodia: A genotype-phenotype association study. The Lancet Infectious Diseases. 2017;17:164-173
129.Imwong M, Suwannasin K, Kunasol C, Sutawong K, Mayxay M, Rekol H, et al. The spread of artemisinin-resistant Plasmodium falciparum in the greater Mekong subregion: A molecular epidemiology observational study. The Lancet Infectious Diseases. 2017;17:491-497
130.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. Malaria Journal. 2013;12:364
131.Suwonkerd W, Ritthison W, Ngo CT, Tainchum K, Bangs MJ, Chareonviriphap T. Vector biology and malaria transmission in Southeast Asia. In: Manguin S, editor. Anopheles Mosquitoes—New Insights into Malaria Vectors. Rijeka: Intech; 2013. pp. 274-325
132.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 precis. Parasites & Vectors. 2011;4:89
133.Hii J, Rueda LM. Malaria vectors in the greater Mekong subregion: Overview of malaria vectors and remaining challenges. The Southeast Asian Journal of Tropical Medicine and Public Health. 2013;44(Suppl 1):73-165; discussion 306-167
134.Manguin S, Garros C, Dusfour I, Harbach RE, Coosemans M. Bionomics, taxonomy, and distribution of the major malaria vector taxa of Anopheles subgenus Cellia in Southeast Asia: An updated review. Infection, Genetics and Evolution. 2008;8:489-503
135.Morgan K, Somboon P, Walton C. Understanding Anopheles diversity in Southeast Asia and its applications for malaria control. In: Manguin S, editor. Anopheles Mosquitoes—New Insights into Malaria Vectors. Rijeka: Intech; 2013. pp. 327-355
136.Tainchum K, Ritthison W, Chuaycharoensuk T, Bangs MJ, Manguin S, Chareonviriyaphap T. Diversity of Anopheles species and trophic behavior of putative malaria vectors in two malaria endemic areas of northwestern Thailand. Journal of Vector Ecology. 2014;39:424-436
137.Sriwichai P, Karl S, Samung Y, Sumruayphol S, Kiattibutr K, Payakkapol A, et al. Evaluation of CDC light traps for mosquito surveillance in a malaria endemic area on the Thai-Myanmar border. Parasites & Vectors. 2015;8:636
138.Wang Y, Zhong D, Cui L, Lee MC, Yang Z, Yan G, et al. Population dynamics and community structure of Anopheles mosquitoes along the China-Myanmar border. Parasites & Vectors. 2015;8:445
139.Zhong D, Wang X, Xu T, Zhou G, Wang Y, Lee MC, et al. Effects of microclimate condition changes due to land use and land cover changes on the survivorship of malaria vectors in China-Myanmar border region. PLoS One. 2016;11:e0155301
140.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. Malaria Journal. 2013;12:363
141.Chareonviriyaphap T, Bangs MJ, Suwonkerd W, Kongmee M, Corbel V, Ngoen-Klan R. Review of insecticide resistance and behavioral avoidance of vectors of human diseases in Thailand. Parasites & Vectors. 2013;6:280
142.Van Bortel W, Trung HD, Thuan le K, Sochantha T, Socheat D, Sumrandee C, et al. The insecticide resistance status of malaria vectors in the Mekong region. Malaria Journal. 2008;7:102
143.Verhaeghen K, Van Bortel W, Trung HD, Sochantha T, Coosemans M. Absence of knockdown resistance suggests metabolic resistance in the main malaria vectors of the Mekong region. Malaria Journal. 2009;8:84
144.Baird JK. Asia-Pacific malaria is singular, pervasive, diverse and invisible. International Journal for Parasitology. 2017;47:371-377
145.Kaneko A, Taleo G, Kalkoa M, Yamar S, Kobayakawa T, Bjorkman A. Malaria eradication on islands. Lancet. 2000;356:1560-1564
146.Hsiang MS, Hwang J, Tao AR, Liu Y, Bennett A, Shanks GD, et al. Mass drug administration for the control and elimination of Plasmodium vivax malaria: An ecological study from Jiangsu province, China. Malaria Journal. 2013;12:383
147.Song J, Socheat D, Tan B, Dara P, Deng C, Sokunthea S, et al. Rapid and effective malaria control in Cambodia through mass administration of artemisinin-piperaquine. Malaria Journal. 2010;9:57
148.Landier J, Kajeechiwa L, Thwin MM, Parker DM, Chaumeau V, Wiladphaingern J, et al. Safety and effectiveness of mass drug administration to accelerate elimination of artemisinin-resistant falciparum malaria: A pilot trial in four villages of eastern Myanmar. Wellcome Open Research. 2017;2:81
149.Newby G, Hwang J, Koita K, Chen I, Greenwood B, von Seidlein L, et al. Review of mass drug administration for malaria and its operational challenges. The American Journal of Tropical Medicine and Hygiene. 2015;93:125-134
150.Geissbuhler Y, Kannady K, Chaki PP, Emidi B, Govella NJ, Mayagaya V, et al. Microbial larvicide application by a large-scale, community-based program reduces malaria infection prevalence in urban Dar es Salaam, Tanzania. PLoS One. 2009;4:e5107
151.Chaccour CJ, Kobylinski KC, Bassat Q, Bousema T, Drakeley C, Alonso P, et al. Ivermectin to reduce malaria transmission: A research agenda for a promising new tool for elimination. Malaria Journal. 2013;12:153
152.Kobylinski KC, Ubalee R, Ponlawat A, Nitatsukprasert C, Phasomkulsolsil S, Wattanakul T, et al. Ivermectin susceptibility and sporontocidal effect in greater Mekong subregion Anopheles. Malaria Journal. 2017;16:280
153.Sluydts V, Durnez L, Heng S, Gryseels C, Canier L, Kim S, et al. Efficacy of topical mosquito repellent (picaridin) plus long-lasting insecticidal nets versus long-lasting insecticidal nets alone for control of malaria: A cluster randomised controlled trial. The Lancet Infectious Diseases. 2016;16:1169-1177
154.Gryseels C, Uk S, Sluydts V, Durnez L, Phoeuk P, Suon S, et al. Factors influencing the use of topical repellents: Implications for the effectiveness of malaria elimination strategies. Scientific Reports. 2015;5:16847
155.Wang S, Jacobs-Lorena M. Genetic approaches to interfere with malaria transmission by vector mosquitoes. Trends in Biotechnology. 2013;31:185-193
156.Barreaux P, Barreaux AMG, Sternberg ED, Suh E, Waite JL, Whitehead SA, et al. Priorities for broadening the malaria vector control tool kit. Trends in Parasitology. 2017;33:763-774
Written By
Liwang Cui, Yaming Cao, Jaranit Kaewkungwal, Amnat
Khamsiriwatchara, Saranath Lawpoolsri, Than Naing Soe, Myat
Phone Kyaw and Jetsumon Sattabongkot
Submitted: 17 July 2017Reviewed: 09 March 2018Published: 18 July 2018
Open access peer-reviewed chapter
