Over 200 RDT products from over 30 different manufacturers have been evaluated by the WHO and Foundation for Innovative New Diagnostics (FIND) and other malaria elimination stakeholders [56▪▪]. Operational detection thresholds have been set at a low density of 200 parasites/μl and a higher density of 2000–5000 parasites/μl [56▪▪]; the approximate RDT LOD is equivalent to 200 iRBCs/μl of blood (Table 2). RDTs detect P. falciparum histidine-rich protein II (PfHRP2), Plasmodium lactate dehydrogenase and aldolase [56▪▪]. Although they identify P. falciparum, specifically, RDTs have no species-specific capacity to identify all five malaria species and cannot provide information on developmental stages. As these methods have not been coupled with strategies for concentrating parasite protein from the blood prior to analysis, they are limited by the 5 μl sample volume applied to the RDT cartridge. Further complications linked to RDTs have arisen through reported false-positive and false-negative results systematically reviewed through the WHO–FIND collaboration [56▪▪]. False-positive results are associated with persistence of PfHRP2 in peripheral blood, cross reactivity against human rheumatoid factor, and other infectious diseases [56▪▪,57,58]. False-negative RDT results are associated with deletions of pfhrp2 and pfhrp3 genes . Cheng et al.[60▪▪] reported, however, that PfHRP2-detecting RDTs are effective for routine clinical case management in most malaria-endemic regions. When PfHRP2 deletion prevalence is greater that 10% in a region PfHRP2-detecting RDT usage is discouraged .
The LOD for NAA tests are at least two to four orders of magnitude lower than microscopy and RDTs (virtually all NAA strategies are able to detect 0.05–5 iRBCs/μl of blood; Table 2). Early NAA tests focused on detecting the 18S ribosomal RNA gene sequences (DNA template), present in 5–10 copies per Plasmodium genome. These sequences were either amplified using species-specific primers with products visualized following gel electrophoresis, or by genus-specific PCR amplification followed by post-PCR methods to distinguish species [67,68]. Plasmodium species NAA assays have expanded to target additional gene sequences [including the P. falciparum stevor multigene family , mitochondrial DNA (mtDNA) , and telomere-associated repetitive element 2 (TARE-2)  sequences and P. vivax Pvr64 sequences  and mtDNA . Assays targeting stevor, TARE-2 and Pvr64 are limited by their single species focus; 18S rRNA gene and mtDNA  assays are developed to identify all human malaria species]. Assay development focused on these sequences theoretically improves infection detection beyond assay formats that target 18S rRNA gene sequence due to increased copy number (stevor, 30–40 copies/parasite genome ; mt cytb, 30–100 copies/iRBC ; TARE-2,250 copies/parasite genome ). With a 100-fold increase in target sequence it is possible that the LOD could reach 0.005–0.05 iRBCs/μl or 1–10 iRBCs/200 μl of blood (Table 2)  as this very small number of iRBCs would be releasing hundreds of copies of the target DNA sequence into the final 200 μl volume of purified genomic DNA and just one template sequence would be needed to drive the NAA reaction.
Additional NAA strategies that have been shown to improve sensitivity of molecular diagnosis include amplification of expressed nucleic acid sequence (RNA template) and concentration of RBC away from serum and WBC of whole blood. Expressed nucleic acid compared with genomic DNA benefits from parasite amplification of target sequence 1000 to 3500-fold. This increase in nucleic acid template concentration would push the LOD to 0.0005–0.005 iRBCs/μl or 1–10 iRBCs/2 ml of blood (Table 2). To fully implement this increased capacity for detection requires RBC concentration strategies. Concerns that discourage RNA-based infection detection include the lability of RNA compared with the durability of DNA. RBC concentration strategies have been implemented in the context of malaria diagnosis in field-based malaria elimination [74▪] and in the context of clinical trials [75▪]. Nucleic acid extraction performed on the RBC fraction from 1 mL of whole blood (approximately, 5 × 109 RBCs) would enable surveillance 0.2% of the adult blood volume. Increasing the availability of nucleic acid template has facilitated successful pooling of patient samples to accelerate sample processing [30▪,71,76,77] that will be called for as malaria elimination requires increasing surveillance. Finally, loop-mediated isothermal amplification exhibits potential to release NAA from the laboratory and enable highly sensitive, NAA malaria diagnosis in remote healthcare settings [72,78]. Additionally, recent advances with lab-on-chip  and noninstrumented nucleic acid amplification  strategies advance promise for NAA point-of-care testing.
A number of recent studies have reported on strategies to exploit physical and/or electromagnetic features of hemozoin crystals to detect infection by malaria parasites [54,80–86]. Generally, the LOD of these methods is between 1 and 30 iRBCs/μl and, therefore, not as sensitive as NAA strategies. However, as these methods claim to rely on inexpensive and portable technologies, there is potential that they may contribute to more efficient and sensitive point-of-care malaria diagnostic strategies with further optimization. Finally, more recent work from Lukianova-Hleb et al.[87,88▪▪] describes a noninvasive method for detecting hemozoin. These authors indicate that hemozoin specificity for malaria and the susceptibility of this nanocrystal to optical excitation by laser pulse generates expansion and collapse of a vapor nanobubble. The resulting pressure pulse is easily detected through the skin with an ultrasound sensor. Preliminary studies in mice, one human infection, and mosquitoes have provided results that demonstrate a noninvasive strategy for malaria diagnosis.
Papers of particular interest, published within the annual period of review, have been highlighted as:
1. Oguike MC, Sutherland CJ. Dimorphism in genes encoding sexual-stage proteins of Plasmodium ovale curtisi and Plasmodium ovale wallikeri. Int J Parasitol 2015; 45:449–454.
2. Cox-Singh J. Zoonotic malaria
: Plasmodium knowlesi, an emerging pathogen. Curr Opin Infect Dis 2012; 25:530–536.
3. Sandeu MM, Moussiliou A, Moiroux N, et al. Optimized Pan-species and speciation duplex real-time PCR assays for Plasmodium parasites detection in malaria
vectors. PLoS ONE 2012; 7:e52719.
4. Wirtz RA, Zavala F, Charoenvit Y, et al. Comparative testing of monoclonal antibodies against Plasmodium falciparum sporozoites for ELISA development. Bull World Health Organ 1987; 65:39–45.
5. Ramasamy R. Zoonotic malaria
global overview and research and policy needs. Front Public Health 2014; 2:123.
6. Domingo GJ, Satyagraha AW, Anvikar A, et al. G6PD testing in support of treatment and elimination
: recommendations for evaluation of G6PD tests. Malar J 2013; 12:391.
7. Satyagraha AW, Sadhewa A, Baramuli V, et al. G6PD deficiency at Sumba in Eastern Indonesia is prevalent, diverse and severe: implications for primaquine therapy against relapsing Vivax malaria
. PLoS Negl Trop Dis 2015; 9:e0003602.
8. Roll Back Malaria
Partnership (RBMP). The Global Malaria
Action Plan for a Malaria
-Free World. Geneva: World Health Organization; 2008.
9. Smith DL, Cohen JM, Moonen B, et al. Infectious disease. Solving the Sisyphean problem of malaria
in Zanzibar. Science 2011; 332:1384–1385.
10▪▪. World Health Organization. World Malaria
Report 2014. Geneva: World Health Organization; 2014. p. 242.
This WHO document provides essential annual updates on malaria distribution around the world.
11. Alonso PL, Brown G, Arevalo-Herrera M, et al. A research agenda to underpin malaria
eradication. PLoS Med 2011; 8:e1000406.
12. Pampana EJ. A textbook of malaria
eradication. 1969; London:Oxford University Press, p. 360.
13▪▪. World Health Organization. From malaria
control to malaria elimination
: a manual for elimination
scenario planning: World Health Organization; 2014. p. 53.
This WHO document provides essential annual updates on all phases of malaria control and elimination.
14▪▪. Bousema T, Okell L, Felger I, et al. Asymptomatic malaria
infections: detectability, transmissibility and public health relevance. Nat Rev Miocrobiol 2014; 12:833–840.
This review article is the latest update on SMI and how they shape malaria epidemiology.
15. Okell LC, Bousema T, Griffin JT, et al. Factors determining the occurrence of submicroscopic malaria
infections and their relevance for control. Nat Commun 2012; 3:1237.
16. Tietje K, Hawkins K, Clerk C, et al. The essential role of infection-detection technologies for malaria elimination
and eradication. Trends Parasitol 2014; 30:259–266.
17▪▪. Hawkins K, Burton R, LaBarre P. Diagnostics to support malaria elimination
: choosing an appropriate biomarker to target the subclinical Plasmodium falciparum transmission reservoir. Proceedings of IEEE 2014 Global Humanitarian Technology Conference; 10–13 October 2014; San Jose, CA: IEEE; 2014.
This review article explains the relative importance of malaria parasite biomarkers for diagnosis of infection that has potential to continue transmission.
18▪▪. Cheng Q, Cunningham J, Gatton ML. Systematic review of sub-microscopic P. vivax infections: prevalence and determining factors. PLoS Negl Trop Dis 2015; 9:e3413.
As much of the malaria literature emphasizes the importance of P. falciparum, this review article shows specifically the evidence for submicroscopic infection involving P. vivax and the impact of this important parasite on global malaria epidemiology.
19. Cottrell G, Moussiliou A, Luty AJ, et al. Submicroscopic Plasmodium falciparum infections are associated with maternal anemia premature births, and low birth weight. Clin Infect Dis 2015; 60:1481–1488.
20. Isozumi R, Fukui M, Kaneko A, et al. Improved detection of malaria
cases in island settings of Vanuatu and Kenya by PCR that targets the Plasmodium mitochondrial cytochrome c oxidase III (cox3) gene. Parasitol Int 2015; 64:304–308.
21. Lo E, Zhou G, Oo W, et al. Low parasitemia in submicroscopic infections significantly impacts malaria
diagnostic sensitivity in the highlands of Western Kenya. PLoS ONE 2015; 10:e0121763.
22. Morris U, Xu W, Msellem MI, et al. Characterising temporal trends in asymptomatic Plasmodium infections and transporter polymorphisms during transition from high to low transmission in Zanzibar, 2005–2013. Infect Genet Evol 2015; 33:110–117.
23. Sema M, Alemu A, Bayih AG, et al. Evaluation of noninstrumented nucleic acid amplification by loop-mediated isothermal amplification (NINA-LAMP) for the diagnosis
in Northwest Ethiopia. Malar J 2015; 14:44.
24. Talha AA, Pirahmadi S, Mehrizi AA, et al. Molecular genetic analysis of Plasmodium vivax isolates from Eastern and Central Sudan using pvcsp and pvmsp-3alpha genes as molecular markers. Infect Genet Evol 2015; 32:12–22.
25. Ehtesham R, Heidari A, Raeisi A, et al. Detection of mixed-species infections of Plasmodium falciparum and Plasmodium vivax by nested PCR and rapid diagnostic tests in southeastern Iran. Am J Trop Med Hyg 2015.
26. Baum E, Sattabongkot J, Sirichaisinthop J, et al. Submicroscopic and asymptomatic Plasmodium falciparum and Plasmodium vivax infections are common in western Thailand: molecular and serological evidence. Malar J 2015; 14:95.
27. Chaturvedi N, Bhandari S, Bharti PK, et al. Sympatric distribution of Plasmodium ovale curtisi and P. ovale wallikeri in India: implication for the diagnosis
and its control. Trans R Soc Trop Med Hyg 2015; 109:352–354.
28. Waltmann A, Darcy AW, Harris I, et al. High rates of asymptomatic, submicroscopic Plasmodium vivax infection and disappearing Plasmodium falciparum malaria
in an area of low transmission in Solomon Islands. PLoS Negl Trop Dis 2015; 9:e0003758.
29▪. Thanh PV, Hong NV, Van Van N, et al. Epidemiology of forest malaria
in Central Vietnam: the hidden parasite reservoir. Malar J 2015; 14:86.
This manuscript illustrates the importance of remote, untreated P. vivax infections carrying gametocytes and hypnozoites that sustain blood stage infection. The authors conclude that until new approaches for diagnosing and treating this hidden reservoir, malaria elimination will be extremely challenging.
30▪. Cheng Z, Wang D, Tian X, et al. Capture and ligation probe-PCR (CLIP-PCR) for molecular screening, with application to active malaria
surveillance for elimination
. Clin Chem 2015; 61:821–828.
This manuscript presents a reverse-transcription PCR approach for amplification of 18S rRNA in pooled samples to detect malaria species.
31. Vallejo AF, Chaparro PE, Benavides Y, et al. High prevalence of sub-microscopic infections in Colombia. Malar J 2015; 14:201.
32. D’Acremont V, Kahama-Maro J, Swai N, et al. Reduction of antimalarial consumption after rapid diagnostic tests implementation in Dar es Salaam: a before-after and cluster randomized controlled study. Malar J 2011; 10:107.
33. Karl S, Gurarie D, Zimmerman PA, et al. A sub-microscopic gametocyte reservoir can sustain malaria
transmission. PLoS ONE 2011; 6:e20805.
34. Moore LR, Fujioka H, Williams PS, et al. Hemoglobin degradation in malaria
-infected erythrocytes determined from live cell magnetophoresis. Faseb J 2006; 20:747–749.
35. Gravenor MB, van Hensbroek MB, Kwiatkowski D. Estimating sequestered parasite population dynamics in cerebral malaria
. Proc Natl Acad Sci U S A 1998; 95:7620–7624.
36. Lee KS, Cox-Singh J, Singh B. Morphological features and differential counts of Plasmodium knowlesi parasites in naturally acquired human infections. Malar J 2009; 8:73.
37. Antinori S, Galimberti L, Milazzo L, et al. Biology of human malaria
plasmodia including Plasmodium knowlesi. Mediterr J Hematol Infect Dis 2012; 4:e2012013.
38. Kerlin DH, Gatton ML. Preferential invasion by Plasmodium merozoites and the self-regulation of parasite burden. PLoS ONE 2013; 8:e57434.
39. Gething PW, Elyazar IR, Moyes CL, et al. A long neglected world malaria
map: Plasmodium vivax endemicity in. PLoS Negl Trop Dis 2012; 6:e1814.
40. Gething PW, Patil AP, Smith DL, et al. A new world malaria
map: Plasmodium falciparum endemicity in. Malar J 2011; 10:378.
41. Moyes CL, Henry AJ, Golding N, et al. Defining the geographical range of the Plasmodium knowlesi reservoir. PLoS Negl Trop Dis 2014; 8:e2780.
42▪. Muller M, Schlagenhauf P. Plasmodium knowlesi in travellers, update. Int J Infect Dis 2014; 22:55–64.
This manuscript provides an update on the risk of P. knowlesi malaria for travelers to Malaysia and best approaches for malaria diagnosis and treatment.
43. Alonso PL, Bassat Q, Binka F, et al. A research agenda for malaria
eradication: drugs. PLoS Med 2011; 8:e1000402.
44. World Health Organization. Microscopy for the detection, identification and quantification of malaria
parasites on stained thick and thin blood films in research settings (version 1.0): procedure: methods manual.: World Health Organization; 2015. p. 32.
45. Dowling MA, Shute GT. A comparative study of thick and thin blood films in the diagnosis
of scanty malaria
parasitaemia. Bull World Health Organ 1966; 34:249–267.
46. Maguire JD, Lederman ER, Barcus MJ, et al. Production and validation of durable, high quality standardized malaria
microscopy slides for teaching, testing and quality assurance during an era of declining diagnostic proficiency. Malar J 2006; 5:92.
47. Zimmerman PA, Thomson JM, Fujioka H, et al. Diagnosis
by magnetic deposition microscopy. Am J Trop Med Hyg 2006; 74:568–572.
48. Karl S, David M, Moore L, et al. Enhanced detection of gametocytes by magnetic deposition microscopy predicts higher potential for Plasmodium falciparum transmission. Malar J 2008; 7:66.
49. Kremsner PG, Valim C, Missinou MA, et al. Prognostic value of circulating pigmented cells in African children with malaria
. J Infect Dis 2009; 199:142–150.
50. Metzger WG, Mordmuller BG, Kremsner PG. Malaria
pigment in leucocytes. Trans R Soc Trop Med Hyg 1995; 89:637–638.
51. Campo JJ, Aponte JJ, Nhabomba AJ, et al. Feasibility of flow cytometry for measurements of Plasmodium falciparum parasite burden in studies in areas of malaria
endemicity by use of bidimensional assessment of YOYO-1 and autofluorescence. J Clin Microbiol 2011; 49:968–974.
52. Jagannadh VK, Murthy RS, Srinivasan R, et al. Automated quantitative cytological analysis using portable microfluidic microscopy. J Biophotonics 2015; [Epub ahead of print].
53. Malleret B, Claser C, Ong AS, et al. A rapid and robust tri-color flow cytometry assay for monitoring malaria
parasite development. Sci Rep 2011; 1:118.
54. Rebelo M, Tempera C, Bispo C, et al. Light depolarization measurements in malaria
: a new job for an old friend. Cytometry A 2015; 87:437–445.
55. Shapiro HM, Apte SH, Chojnowski GM, et al.> Cytometry in malaria
: a practical replacement for microscopy? Current protocols in cytometry/editorial board, J Paul Robinson, managing editor [et al]. 2013; Chapter 11:Unit 11 20.
56▪▪. World Health Organization. Malaria
rapid diagnostic test performance: results of WHO product testing of malaria
RDTs: round 5 (2013). Geneva: World Health Organization; 2014. p. 130.
This publication reports on all of the latest efforts to test and evaluate malaria rapid diagnostic tests.
57. Gillet P, Mumba Ngoyi D, Lukuka A, et al. False positivity of nontargeted infections in malaria
rapid diagnostic tests: the case of human african trypanosomiasis. PLoS Negl Trop Dis 2013; 7:e2180.
58. Leshem E, Keller N, Guthman D, et al. False-positive Plasmodium falciparum histidine-rich protein 2 immunocapture assay results for acute schistosomiasis caused by Schistosoma mekongi. J Clin Microbiol 2011; 49:2331–2332.
59. Gamboa D, Ho MF, Bendezu J, et al. A large proportion of P. falciparum isolates in the Amazon region of Peru lack pfhrp2 and pfhrp3: implications for malaria
rapid diagnostic tests. PLoS ONE 2010; 5:e8091.
60▪▪. Cheng Q, Gatton ML, Barnwell J, et al. Plasmodium falciparum parasites lacking histidine-rich protein 2 and 3: a review and recommendations for accurate reporting. Malar J 2014; 13:283.
This manuscript reports on deletions involving the gene encoding P. falciparum histidine-rich protein II that underlie an important source of false-negative rapid diagnostic test results.
61. Udhayakumar V, Barnwell J. Molecular profile and survey of hrp2 and hrp3 genetic deletions in South America. Amazon Malaria
Initiative (AMI)/Amazon Network for the Surveillance of Antimalarial Drug Resistance (RAVREDA) Annual Meeting; Lima, Peru; 9 April 2013.
62. Erdman LK, Kain KC. Molecular diagnostic and surveillance tools for global malaria
control. Travel Med Infect Dis 2008; 6 (1–2):82–99.
63. Cordray MS, Richards-Kortum RR. Emerging nucleic acid-based tests for point-of-care detection of malaria
. Am J Trop Med Hyg 2012; 87:223–230.
64. World Health Organization. WHO Evidence Review Group on Malaria Diagnosis
in Low Transmission Settings. In: Meeting, M.P.A.C., editor. Geneva: World Health Organization; 2014. p. 33.
65. Okell LC, Ghani AC, Lyons E, et al. Submicroscopic infection in Plasmodium falciparum-endemic populations: a systematic review and meta-analysis. J Infect Dis 2009; 200:1509–1517.
66. Greenwood B. The molecular epidemiology of malaria
. Trop Med Int Health 2002; 7:1012–1021.
67. Snounou G, Viriyakosol S, Jarra W, et al. Identification of the four human malaria
parasite species in field samples by the polymerase chain reaction and detection of a high prevalence of mixed infections. Mol Biochem Parasitol 1993; 58:283–292.
68. McNamara DT, Kasehagen LJ, Grimberg BT, et al. Diagnosing infection levels of four human malaria
parasite species by a polymerase chain reaction/ligase detection reaction fluorescent microsphere-based assay. Am J Trop Med Hyg 2006; 74:413–421.
69. Cheng Q, Cloonan N, Fischer K, et al. stevor and rif are Plasmodium falciparum multicopy gene families which potentially encode variant antigens. Mol Biochem Parasitol 1998; 97 (1–2):161–176.
70. Steenkeste N, Incardona S, Chy S, et al. Towards high-throughput molecular detection of Plasmodium: new approaches and molecular markers. Malar J 2009; 8:86.
71. Hofmann N, Mwingira F, Shekalaghe S, et al. Ultra-sensitive detection of Plasmodium falciparum by amplification of multicopy subtelomeric targets. PLoS Med 2015; 12:e1001788.
72. Patel JC, Oberstaller J, Xayavong M, et al. Real-time loop-mediated isothermal amplification (RealAmp) for the species-specific identification of Plasmodium vivax. PLoS ONE 2013; 8:e54986.
73. Smits HL, Gussenhoven GC, Terpstra W, et al. Detection, identification and semi-quantification of malaria
parasites by NASBA amplification of small subunit ribosomal RNA sequences. J Microbiol Methods 1997; 28:65–75.
74▪. Imwong M, Hanchana S, Malleret B, et al. High-throughput ultrasensitive molecular techniques for quantifying low-density malaria
parasitemias. J Clin Microbiol 2014; 52:3303–3309.
This manuscript presents an approach for concentrating parasitized RBCs to increase the sensitivity of PCR detection of malaria parasite species in field settings.
75▪. Murphy SC, Hermsen CC, Douglas AD, et al. External quality assurance of malaria
nucleic acid testing for clinical trials and eradication surveillance. PLoS ONE 2014; 9:e97398.
This manuscript presents an approach for concentrating parasitized RBCs to increase the sensitivity of PCR detection of malaria parasite species in clinical trials.
76. Taylor SM, Juliano JJ, Trottman PA, et al. High-throughput pooling and real-time PCR-based strategy for malaria
detection. J Clin Microbiol 2010; 48:512–519.
77. Hsiang MS, Hwang J, Kunene S, et al. Surveillance for malaria elimination
in Swaziland: a national cross-sectional study using pooled PCR and serology. PLoS ONE 2012; 7:e29550.
78. Hopkins H, Gonzalez IJ, Polley SD, et al. Highly sensitive detection of malaria
parasitemia in a malaria
-endemic setting: performance of a new loop-mediated isothermal amplification kit in a remote clinic in Uganda. J Infect Dis 2013; 208:645–652.
79. Taylor BJ, Howell A, Martin KA, et al. A lab-on-chip for malaria diagnosis
and surveillance. Malar J 2014; 13:179.
80. Butykai A, Orban A, Kocsis V, et al. Malaria
pigment crystals as magnetic micro-rotors: key for high-sensitivity diagnosis
. Sci Rep 2013; 3:1431.
81. Khoshmanesh A, Dixon MW, Kenny S, et al. Detection and quantification of early-stage malaria
parasites in laboratory infected erythrocytes by attenuated total reflectance infrared spectroscopy and multivariate analysis. Anal Chem 2014; 86:4379–4386.
82. Mens PF, Matelon RJ, Nour BY, et al. Laboratory evaluation on the sensitivity and specificity of a novel and rapid detection method for malaria diagnosis
based on magneto-optical technology (MOT). Malar J 2010; 9:207.
83. Newman DM, Heptinstall J, Matelon RJ, et al. A magneto-optic route toward the in vivo diagnosis
: preliminary results and preclinical trial data. Biophys J 2008; 95:994–1000.
84. Orban A, Butykai A, Molnar A, et al. Evaluation of a novel magneto-optical method for the detection of malaria
parasites. PLoS ONE 2014; 9:e96981.
85. Peng WK, Kong TF, Ng CS, et al. Micromagnetic resonance relaxometry for rapid label-free malaria diagnosis
. Nat Med 2014; 20:1069–1073.
86. Yuen C, Liu Q. Magnetic field enriched surface enhanced resonance Raman spectroscopy for early malaria diagnosis
. J Biomed Opt 2012; 17:017005.
87. Lukianova-Hleb EY, Campbell KM, Constantinou PE, et al. Hemozoin-generated vapor nanobubbles for transdermal reagent- and needle-free detection of malaria
. Proc Natl Acad Sci U S A 2014; 111:900–905.
88▪▪. Lukianova-Hleb E, Bezek S, Szigeti R, et al. Transdermal diagnosis
using vapor nanobubbles. Emerg Infect Dis 2015; 21:1122–1127.
This manuscript presents a novel noninvasive method for detecting hemozoin resulting from a pressure pulse that is easily detected through the skin with an ultrasound sensor. Preliminary results have been shown for murine malaria, one human infection and mosquitoes.
89. Bustin SA, Benes V, Garson J, et al. The need for transparency and good practices in the qPCR literature. Nat Methods 2013; 10:1063–1067.
90▪. Dalrymple U, Mappin B, Gething PW. Malaria
mapping: understanding the global endemicity of falciparum and vivax malaria
. BMC Med 2015; 13:140.