PCR-Based Pooling of Dried Blood Spots for Detection of Malaria

By Craig Cole,2014-05-28 17:28
8 views 0
PCR-Based Pooling of Dried Blood Spots for Detection of Malaria

     JOURNAL OF CLINICAL MICROBIOLOGY, Oct. 2010, p. 35393543 Vol. 48, No. 10 0095-1137/10/$12.00 doi:10.1128/JCM.00522-10 Copyright ? 2010, American Society for Microbiology. All Rights Reserved.

    PCR-Based Pooling of Dried Blood Spots for Detection of Malaria

    Parasites: Optimization and Application to a Cohort

    of Ugandan Children

    1,2345Michelle S. Hsiang,* Michael Lin,Christian Dokomajilar,Jordan Kemere, 444Christopher D. Pilcher,Grant Dorsey,and Bryan Greenhouse

    123Global Health Group, Department of Global Health Sciences,Department of Pediatrics,School of Medicine,and Department of 4Medicine,University of California at San Francisco, San Francisco, California 94110, and School of Medicine, 5University of North Carolina at Chapel Hill, Chapel Hill, North Carolina 27599

    Received 11 March 2010/Returned for modication 27 April 2010/Accepted 28 July 2010

Sensitive, high-throughput methods to detect malaria parasites in low-transmission settings are needed.

    PCR-based pooling strategies may offer a solution. We ;rst used laboratory-prepared samples to compare 2

    DNA extraction and 4 PCR detection methods across a range of pool sizes and parasite densities. Pooled

    Chelex extraction of DNA, followed by nested PCR of cytochrome b, was the optimal strategy, allowing reliable detection of a single low-parasitemic sample (100 parasites/ l) in pool sizes up to 50. This PCR-based pooling

    strategy was then compared with microscopy using 891 dried blood spots from a cohort of 77 Ugandan children followed for 2 years in an urban setting of low endemicity. Among 419 febrile episodes, 35 cases of malaria were detected using the PCR-based pooling strategy and 40 cases using microscopy. All ;ve cases of malaria not

    detected by PCR were from samples stored for >2 years with parasitemia of <6,000/ l, highlighting the issue

    of possible DNA degradation with long-term storage of samples. Among 472 samples collected from asymp- tomatic children as part of routine surveillance, 15 (3.2%) were positive by PCR-based pooling compared to 4

    (0.8%) by microscopy (P 0.01). Thus, this PCR-based pooling strategy for detection of malaria parasites

    using dried blood spots offers a sensitive and ef;cient approach for malaria surveillance in low-transmission settings, enabling improved detection of asymptomatic submicroscopic infections and dramatic savings in

    labor and costs.

     Due to large-scale implementation of effective control mea- care of individual patients. One exception would be real-time sures, many countries where malaria is endemic are experienc- PCR, which has a short turnaround time but carries with it cost ing dramatic declines in disease burden. With this success has and capacity constraints that make it unavailable for rapid come a shift in the end goal from control to elimination (9, 10). diagnosis in most settings. However, there is a potential role However, when the goal is elimination, accurate detection of for PCR in situations where large numbers of samples are persons infected with malaria parasites becomes essential (11). being screened: a high-throughput system could allow accu- Standard surveillance systems depend on diagnosis by micros- rate, rapid, and cost-effective assessments to quantify preva- copy, a method that is technically challenging, labor-intensive, lence and identify species. and often inaccurate in operational settings. More recently The need for a high-throughput method to detect malaria available rapid diagnostic tests (RDTs) provide convenience parasites is especially important for surveillance in low-trans- and ease of use, but they have limitations in specicity, sensi- mission and elimination settings, where low prevalence renders tivity, species identication, and cost (22). diagnostic testing at the individual sample level impractical and PCR-based methods for malaria parasite detection are rel- inefcient. Detection methods based on pooled nucleic acid atively simple and provide improved sensitivity compared to could provide a simple solution. The pooling of sera or plasma microscopy and RDTs, especially in settings where infections specimens has long been used to improve the accuracy, cost- have low parasitemia or contain mixed species (22). PCR can effectiveness, and throughput of screening for many infections also be performed on dried blood spots, which are convenient (e.g., HIV-1, hepatitis B virus, hepatitis C virus, and West Nile for collection, storage, and transport. Nonetheless, PCR has a virus) in low-prevalence settings, such as blood banks (4, 26). long turnaround time, making it an impractical tool for clinical We recently reported methods for pooled analysis of dried blood spots for HIV-1 nucleic acids (1, 18). The goal of this study was to develop and apply a novel * Corresponding author. Mailing address: Global Health Group, PCR-based pooling strategy for the detection of malaria par- Department of Global Health Sciences, Division of Pediatric Infec- asites using dried blood spots. We rst established reference tious Diseases, Department of Pediatrics, University of California, San standards by using laboratory strains to create dried blood Francisco, 50 Beale Street, 12th oor, San Francisco, CA 94105. Phone: (415) 597-8180. Fax: (415) 597-8299. E-mail: hsiangm@peds spots with known Plasmodium falciparum parasite densities. After extracting DNA from pools of dried blood spots, we Supplemental material for this article may be found at http://jcm compared the sensitivities and specicities of different nucleic acid amplication tests. The most robust pooling strategy was Published ahead of print on 4 August 2010.



    TABLE 1. Sensitivity and specicity of pooled Chelex DNA identied and then applied to 891 eld samples collected using extraction followed by cytochrome b nested PCR passive and active surveillance from a cohort of 77 children living in Kampala, Ugandaa city with relatively low malaria % a % Sensitivity (n 30)Specicity endemicitywho were followed for up to 2 years. The results aPool size (n 15) were compared to those obtained by microscopy. 0 p/ l 100 p/ l 10 p/ l

    MATERIALS AND METHODS 50 100 70 100

    25 100 65 100 Preparation of control dried blood spots with known parasite densities. P. 10 100 93 100 falciparum strain W2 parasites were cultured and synchronized at the ring stage 5 100 80 100 using standard methods (14). Parasitemia was determined using ow cytometry. Positive controls were prepared by mixing infected red blood cells with unin- arefers to independent pooled extractions followed by PCR; p/ l, para- nfected whole blood to create parasite densities of 10 and 100 parasites/ l. Neg- sites/ l. ative controls were prepared using blood from uninfected persons. To simulate collection of dried blood spots in the eld, blood was spotted in 20- l aliquots onto Whatman 3MM lter paper, air dried overnight, and stored at room tem- perature in plastic bags sealed with desiccant. Plasmodium vivax dried blood tion based on AluI restriction digestion (31). Species were identied by compar- spots were obtained from the Centers for Disease Control and Prevention of the ing digestion patterns to those of known controls. (See the supplemental material Republic of Korea. Punches 3 mm in diameter were removed from the dried for a detailed protocol of the pooling strategy.) blood spots using handheld hole punches. Statistical analysis. Data were analyzed using Stata 10.0 (StataCorp LP, TX). Optimizing the pooling methodology. We used the laboratory-prepared P. P values were generated using a chi-square or Fishers exact test as appropriate. falciparum blood samples to compare several DNA extraction and amplication methods across a range of pool sizes (10, 50, and 100) and parasite densities (0, 10, and 100 parasites/ l). Once the optimal strategy was identied, additional RESULTS tests were performed in pool sizes of 5, 10, 25, and 50 to generate estimates of sensitivity. Pool sizes of 100 did not easily t into the 1.5-ml tubes used for Optimizing the pooling strategy. Pooled Chelex extraction extraction and were therefore not pursued further. The optimal strategy was also used to test the P. vivax clinical samples in pools of 25. of DNA followed by cytochrome b nested PCR was identied Test pools contained one punch from a positive dried blood spot, the remain- as the most robust strategy. In pools containing one very low-P. der being punches from malaria-negative dried blood spots. A single DNA falciparum-parasitemic sample (10 parasites/ l), the sensitivity extraction was performed on each pool of punches, and the extraction product ranged from 40% with pools of 100 to 93% with pools of 10. was used as a template for amplication. For all pool sizes that included one low-P. falciparum-para- Two different DNA extraction methods were evaluated: the saponin/Chelex method (24) and the QIAamp DNA minikit (Qiagen, CA). The sensitivities of sitemic sample (100 parasites/ l), 100% sensitivity was four detection methods were compared: two nested techniques, 18S ribosomal achieved. The specicity was 100% throughout (Table 1). Sen- DNA (rDNA) nested PCR (29) and cytochrome b nested PCR (30), and two sitivity results from other DNA extraction/PCR combinations single-round techniques, 18S rDNA PCR (21) and 18S rDNA probe-based real- are shown in the supplemental material. time PCR (28). Real-time amplication was performed using a 7500 Real-Time We also evaluated eld samples from three patients diag- PCR System (Applied Biosystems, CA). Other PCR amplications were per- formed on a Bio-Rad Thermocycler C1000 or S1000 (Bio-Rad Laboratories, nosed with P. vivax infection in a similar manner. The sensi- CA). Amplication products were detected by agarose gel electrophoresis or via tivities for the detection of one positive sample in pools of 25 real-time PCR, as appropriate, by persons blinded to the status of the specimens. were 100% (10/10) when the sample had 640 parasites/ l, 90% Field samples from the study cohort. Once Chelex extraction followed by (9/10) at 2,000 parasites/ l, and 100% at 8,118 parasites/ l cytochrome b nested PCR was identied as the optimal strategy, it was used to test dried blood spots collected using passive and active surveillance of a pedi- (10/10). atric cohort randomly selected from the Mulago III parish of Kampala, Uganda Cohort characteristics. A total of 419 samples were col- (T. D. Clark, D. Njama-Meya, D. B. Nzarubara, C. Maiteki-Sebuguzi, B. Green- lected through passive surveillance in patients who presented house, S. G. Staedke, M. R. Kamya, G. Dorsey, and P. J. Rosenthal, unpublished with fever, and 472 samples were collected through active data). Children were enrolled from February to May 2007 and followed until surveillance in asymptomatic subjects. The cohort included 77 December 2008 (7). As part of passive surveillance, blood smears and dried blood spots were collected whenever the children presented to the clinic with a children; the mean age at the start of the study period was 3 new episode of fever (subjective or based on a temperature of 38?C in the years (range, 1 to 10 years), and the median time of follow-up previous 24 h). As part of active surveillance, blood smears and dried blood spots was 663 days (range, 616 to 730 days). were collected from asymptomatic children presenting for routine visits approx- Application of pooling strategy to dried blood spots col- imately every 90 days. Final microscopy results were based on a rigorous quality control system that included rereading of all blood smears by a second micros- lected from the cohort. We used a three-stage pooling strategy, copist and resolution of any discrepancies between the rst and second readings excluding samples that tested negative at each stage (Fig. 1). by a third microscopist. To prepare dried blood spots, blood was blotted on Ultimately, 50 of 891 dried blood spot samples were identied Whatman 3MM lter paper, air dried, and stored at room temperature in plastic as positive, resulting in an overall prevalence of 5.6%. The bags sealed with desiccant. Pooling strategy applied to ;eld samples. We used a three-stage hierarchical efciency of pooling, calculated as the number of tests per- pooling strategy (23). Samples were rst divided into master pools with the formed per total number of individual samples evaluated, was optimal size dependent on the estimated prevalence, as well as the sensitivity and 0.33. In other words, our pooling strategy, compared to per- specicity of the pooling assay (33, 34). Positive master pools were divided into forming individual PCRs on all samples, reduced labor and subpools, with the optimal subpool size being the square root of the master pool supply costs by about two-thirds. size. Positive subpools were tested as individual specimens. Based on the known prevalence by microscopy in this study ( 5%) and the expected high sensitivity Passive surveillance. Of the 50 samples that were positive by and specicity of the assay ( 100% for parasite densities of 100 parasites/ l), the pooling, 35 were among the 419 samples collected from febrile most efcient master and subpool sizes were determined to be nine and three, patients. All 35 of these samples were also positive by micros- respectively (34). copy. Five samples that were positive by microscopy were neg- In the nal step, we performed a restriction digestion, as the cytochrome b ative by pooling (Table 2). The incidence of symptomatic in- PCR assay begins with genus-specic detection, followed by species determina-


     FIG. 2. AluI digestion for species determination. P. ovale digestion patterns may vary depending on the strain type. Our samples showed a pattern that correlates with a published sequence (GenBank acces- sion number AB182497). However, another sequence (GenBank ac- cession number AB182496) would produce a different AluI restriction pattern with expected band sizes of 205 and 610 bp. Pf, P. falciparum; Pm, P. malariae; Po, P. ovale; Pv, P. vivax.

    years) with those that did not. The sensitivity of pooling for

    detection of parasites was 29% (2/7) among samples with both

    characteristics versus 100% (33/33) among samples without

    both characteristics (P 0.001).

    Active surveillance. Of the 50 samples positive by pooling, 15

    were among the 472 samples collected from asymptomatic

    patients. Four were detected by microscopy, and 11 were not. FIG. 1. Schematic of pooling strategy applied to cohort samples. Pooling successfully identied all the microscopy-positive sam- The white circles represent Plasmodium-negative dried blood spots ples (Table 2). The prevalence of asymptomatic infections was (DBS); the solid black circles represent Plasmodium-positive dried blood spots. At each stage, the dried blood spots were extracted to- 0.8% by microscopy versus 3.2% by pooling (P 0.01). gether. The DNA extraction product was then used as a template for For the 11 samples that were negative by microscopy but cytochrome b nested PCR. positive by pooling, reviews of the medical histories suggested that many represented asymptomatic parasitemia. Two sam- ples were from patients who presented symptomatic with pos- fections was 0.32 case per person-year by microscopy versus itive blood smears 5 to 11 days later. Five samples represented 0.28 case per person-year by our pooling strategy. consecutive collections from two patients. One sample was Of the 5 microscopy-positive samples that were missed by obtained from a patient who had gametocytes on smear 3 days pooling, 3 were excluded at the master pool level and 2 at the later. Two samples were obtained at enrollment, when the subpool level. These samples had two characteristics: they had prevalence of asymptomatic parasitemia was high in the cohort low parasite densities (64, 400, 400, 880, and 5,680 parasites/ l) (8). The last sample was from a patient who did not have any and were tested more than 2 years after collection. Among all follow-up smears; we were therefore unable to assess the pos- 40 microscopy-positive samples from symptomatic subjects, we sibility of persistent infection. compared those that had both characteristics (parasite density Species determination. Species determination was based on of less than 6,000 parasites/ l and storage for more than 2 AluI restriction digestion of the amplied cytochrome b gene (Fig. 2). Among the 35 samples that were positive by both microscopy and pooling, there was agreement in the detection TABLE 2. Microscopy versus PCR-based pooling in passive of 31 P. falciparum infections and 1 Plasmodium ovale infec- surveillance of symptomatic subjects and active tion. Three samples identied as P. falciparum by microscopy surveillance of asymptomatic subjects were identied by pooling as 2 P. ovale infections and 1 mixed aMicroscopy P. falciparum-P. vivax infection. Of the 11 asymptomatic cases aSurveillance Pooling Total positive by PCR but not microscopy, there were 7 P. falciparum

    infections, 1 Plasmodium malariae infection, 1 P. ovale infec- Passive 35 0 35 tion, and 2 mixed P. falciparum-P. ovale infections. 5 379 384 Total 40 379 419

    DISCUSSION 4 11 15 Active

    0 457 457 We developed and applied a novel strategy for PCR-based Total 4 468 472 pooled detection of malaria parasites using dried blood spots. a , positive; , negative. After evaluating numerous methods on laboratory-prepared


samples, we found the optimal strategy to be Chelex extraction much of the labor associated with DNA extraction. The nd-

    ings that sensitivity was maintained and that experiments were of pooled 3-mm punches from dried blood spots, followed by

    easy to perform at pool sizes up to 50 are particularly impor- cytochrome b nested PCR for Plasmodium detection and AluI

    tant, since the ability to pool many samples dramatically in- restriction digestion for species determination. Because of the

    sensitive detection associated with this method, we were able creases efciency, especially in low-prevalence settings. For

    example, if our pooling strategy were used in a setting where to reliably identify a single 100-parasite/ l sample in pool sizes

    the prevalence of parasitemia was 0.01%, using pool sizes of 49 up to 50. We then applied the optimized strategy to 891 eld

    samples collected through both passive and active surveillance would result in an efciency of 0.05, saving approximately 95% in a well-characterized pediatric cohort. Compared to micros- of the labor and supply costs (34). Also, of particular relevance copy, we detected fewer cases among symptomatic patients to low-prevalence settings, our proposed strategy can poten- (35/419 versus 40/419), missing 5 microscopy-positive samples tially provide improved specicity, given that an individual

    that were all stored for longer than 2 years and that tended to sample must be amplied by PCR three times to be identied

    have lower parasitemia. Among asymptomatic patients, we as positive. Specicity was not a problem in our study, but false detected 3 times as many infections as by microscopy (15/472 positives could occur in situations where there is cross-contam- versus 4/472). The pooling strategy also allowed improved de- ination. The fact that our strategy involves pooled DNA ex- tection of non-falciparum and mixed infections. tractions with resampling from the original dried blood spot at Monitoring, evaluation, and appropriate targeting of inter- each stage is a further advance over methods that require ventions depend on accurate surveillance, particularly in low- resampling from a potentially contaminated DNA extraction transmission and elimination settings. Compared to micros- product.

    copy and RDTs, PCR is well known to provide improved When applied to eld samples, we found that the sensitivity

    sensitivity and specicity (22). However, the long turnaround of low-parasitemic samples stored for at least 2 years was time for PCR makes it impractical for rapid diagnosis of indi- compromised, highlighting the possibility of DNA degradation vidual patients, and in surveillance settings, it is labor and cost with long-term storage of samples. No microscopy-positive prohibitive to perform individual PCRs on large numbers of samples stored for less than 2 years were missed by pooling, samples. Pooling is a simple strategy that has recently been suggesting that pooled testing of samples within 2 years of applied to the detection of malaria parasites in surveillance collection is reliable. In addition, extremely low parasite den- settings. In an exploratory study using 18S rDNA nested PCR sities, around 10 parasites/ l, may not be detected reliably, and pool sizes of 10, there was successful detection of 6 posi- likely due to the diluting effect of pooling. However, increased tives among 200 serum samples (2). In a larger study that sensitivity was demonstrated with smaller pool sizes (see Table utilized dried blood spot samples collected from a cohort of S1 in the supplemental material). Also, this strategy enabled pregnant women, individual DNA extractions were performed, the detection of 11 submicroscopic infections, 6 of which were and then the extraction products from 4 samples were pooled detected in samples that had been stored for more than 2 in a single 18S rDNA real-time PCR. Among 1,092 microsco- years.

    py-negative samples that were tested by pooling, 35 positive There are several potential applications for our pooling samples were identied. Among 176 microscopy-positive sam- strategy. First, it is a sensitive and high-throughput method ples that were tested in individual PCRs, 74 were positive (32). that can improve the detection of malaria infections in com- Our study offers advances over prior studies. Importantly, we munity surveys of parasite prevalence (12). Second, there is used easily collected dried blood spots, optimized strategies potential for our pooling strategy to be used for quality assur- after comparison of multiple related methods, validated the ance of standard diagnostics in research or clinical settings sensitivity and specicity of our strategy by comparison with (17). Third, it allows the identication of asymptomatic sub-

    laboratory controls, and were able to pool large numbers of microscopic infections. Similar to the results of a recently per- samples. formed systematic review of submicroscopic infections, we de- The inexpensive Chelex method performed better than the tected about three times as many parasitemic samples as with commonly used spin column system, and at lower parasite microscopy (20). Submicroscopic asymptomatic infections are densities and higher pool sizes, cytochrome b nested PCR known to be a major driver of malaria transmission (3, 6, 29), provided the highest sensitivity. Others have also found the but how and to what extent remains unclear. Using samples cytochrome b method to be more sensitive than 18S rDNA from easily collected dried blood spots, our pooling strategy nested PCR (13, 30), likely because there are more copies of can facilitate further studies on this topic. the gene in each parasite (15, 25). It also performed better than Lastly, there are several possible applications of our pooling two other single-round PCR methods, including one real-time strategy for malaria elimination programs. It can be applied in method. The cytochrome b PCR method is also advantageous mass screening and treatment of asymptomatic populations, in that amplication targets a DNA sequence common to all which is emerging as a potentially important strategy to facilitate species (30). For this reason, our pooling strategy requires tests malaria elimination (19). For example, mass screening and treat- of speciation only at the nal stage of pooling, further simpli- ment are being applied as part of recent efforts to eliminate fying the assay. artemisinin-resistant parasites that have emerged at the Thai- Unlike published strategies in which pooling occurred at the Cambodian border (World Health Organization, presented at the time of PCR detection (32, 34), we pooled at the time of DNA Strategy for the Containment of Artemisinin Tolerant Malaria extraction. By using this approach and by using optimal pool Parasites in South-East Asia Project Meeting, Pailin, Cambodia, sizes based on the anticipated prevalence, our strategy allowed 2009). Areas that have already achieved elimination have insti- pooled detection of large numbers of samples and reduced tuted high-volume border-screening programs in order to prevent


and R. W. Snow. 2008. The limits and intensity of Plasmodium falciparum reintroduction of malaria. Mauritius (5), Oman (27), and Singa- transmission: implications for malaria control and elimination worldwide. pore (16) are examples of countries that perform individual PLoS Med. 5:e38. screening of asymptomatic visitors and/or immigrants using mi- 13. Hasan, A. U., S. Suguri, J. Sattabongkot, C. Fujimoto, M. Amakawa, M. Harada, and H. Ohmae. 2009. Implementation of a novel PCR based croscopy or RDTs. While such mass screening strategies are fea- method for detecting malaria parasites from naturally infected mosquitoes in sible using existing techniques in resource-rich countries, use of Papua New Guinea. Malar. J. 8:182. the proposed pooling strategy may put mass screening within the 14. Jensen, J. B. 2002. In vitro culture of Plasmodium parasites. Methods Mol. Med. 72:477488. reach of resource-poor sites (5, 35). 15. Langsley, G., J. E. Hyde, M. Goman, and J. G. Scaife. 1983. Cloning and In summary, we have developed and applied a robust and novel characterisation of the rRNA genes from the human malaria parasite Plas- strategy for high-throughput PCR-based detection of malaria par- modium falciparum. Nucleic Acids Res. 11:87038717. 16. Lee, Y. C., C. S. Tang, L. W. Ang, H. K. Han, L. James, and K. T. Goh. 2009. asites using dried blood spots. Compared to microscopy, our Epidemiological characteristics of imported and locally-acquired malaria in PCR-based pooling strategy allows improved detection of species Singapore. Ann. Acad. Med. Singapore 38:840849. and, most importantly, asymptomatic infections. There is a need 17. McMorrow, M. L., M. I. Masanja, E. Kahigwa, S. M. Abdulla, and S. P. Kachur. 2010. Quality assurance of rapid diagnostic tests for malaria in for improved surveillance in low-transmission settings, and our routine patient care in rural Tanzania. Am. J. Trop. Med. Hyg. 82:151155. proposed strategy holds promise as a simple new tool to facilitate 18. Nugent, C. T., J. Dockter, F. Bernardin, R. Hecht, D. Smith, E. Delwart, C. Pilcher, D. Richman, M. Busch, and C. Giachetti. 2009. Detection of HIV-1 progress toward malaria elimination. in alternative specimen types using the APTIMA HIV-1 RNA Qualitative Assay. J. Virol. Methods 159:1014. ACKNOWLEDGMENTS 19. Ogutu, B., A. B. Tiono, M. Makanga, Z. Premji, A. D. Gbadoe, D. Ubben, A. C. Marrast, and O. Gaye. 2010. Treatment of asymptomatic carriers with This work was supported by grants from the Pediatric Infectious artemether-lumefantrine: an opportunity to reduce the burden of malaria? Diseases Society, National Institutes of Health (grants U01 AI52142, Malar. J. 9:30. R01 MH068686, and P01 AI071713), the Bill and Melinda Gates 20. Okell, L. C., A. C. Ghani, E. Lyons, and C. J. Drakeley. 2009. Submicroscopic Foundation (grant 48820), and the University of California, San Fran- infection in Plasmodium falciparum-endemic populations: a systematic re- cisco, Deans Research Fellowship Program. view and meta-analysis. J. Infect. Dis. 200:15091517. 21. Padley, D., A. H. Moody, P. L. Chiodini, and J. Saldanha. 2003. Use of a We thank Nicolas Steenkeste for sharing details of the cytochrome rapid, single-round, multiplex PCR to detect malarial parasites and identify b nested PCR and AluI digestion for species detection, Ric Price for the species present. Ann. Trop. Med. Parasitol. 97:131137. sharing standard operating procedures of other tested PCR methods, 22. Perkins, M. D., and D. Bell. 2008. Working without a blindfold: the critical and Jung-Yeon Kim for sharing of clinical P. vivax samples. We thank role of diagnostics in malaria control. Malar. J. 7(Suppl. 1):S5. Phil Rosenthal and Jay Tureen for their guidance and helpful review of 23. Pilcher, C. D., J. J. Eron, Jr., S. Galvin, C. Gay, and M. S. Cohen. 2004. the manuscript and Richard Feachem for his support of the project. Acute HIV revisited: new opportunities for treatment and prevention. We declare that we have no conicts of interest for this article. J. Clin. Invest. 113:937945. 24. Plowe, C. V., A. Djimde, M. Bouare, O. Doumbo, and T. E. Wellems. 1995. REFERENCES Pyrimethamine and proguanil resistance-conferring mutations in Plasmo- dium falciparum dihydrofolate reductase: polymerase chain reaction meth- 1. Bebell, L. M., C. D. Pilcher, G. Dorsey, D. Havlir, M. R. Kamya, M. P. Busch, ods for surveillance in Africa. Am. J. Trop. Med. Hyg. 52:565568. J. D. Williams, C. T. Nugent, C. Bentsen, P. J. Rosenthal, and E. D. Char- 25. Preiser, P. R., R. J. Wilson, P. W. Moore, S. McCready, M. A. Hajibagheri, K. J. lebois. 2010. Acute HIV-1 infection is highly prevalent in Ugandan adults Blight, M. Strath, and D. H. Williamson. 1996. Recombination associated with with suspected malaria. AIDS 24:19451952. replication of malarial mitochondrial DNA. EMBO J. 15:684693. 2. Bharti, A. R., S. L. Letendre, K. P. Patra, J. M. Vinetz, and D. M. Smith. 26. Quinn, T. C., R. Brookmeyer, R. Kline, M. Shepherd, R. Paranjape, S. 2009. Malaria diagnosis by a polymerase chain reaction-based assay using a Mehendale, D. A. Gadkari, and R. Bollinger. 2000. Feasibility of pooling sera pooling strategy. Am. J. Trop. Med. Hyg. 81:754757. for HIV-1 viral RNA to diagnose acute primary HIV-1 infection and esti- 3. Boudin, C., M. Olivier, J. F. Molez, J. P. Chiron, and P. Ambroise-Thomas. mate HIV incidence. AIDS 14:27512757. 1993. High human malarial infectivity to laboratory-bred Anopheles gambiae 27. Roll Back Malaria Partnership. 2008. Elimination and eradication: achiev- in a village in Burkina Faso. Am. J. Trop. Med. Hyg. 48:700706. ing zero transmission, Global malaria action plan, part II.3. Roll Back Ma- 4. Busch, M. P., D. J. Wright, B. Custer, L. H. Tobler, S. L. Stramer, S. H. laria, Geneva, Switzerland. Kleinman, H. E. Prince, C. Bianco, G. Foster, L. R. Petersen, G. Nemo, and 28. Shokoples, S. E., M. Ndao, K. Kowalewska-Grochowska, and S. K. Yanow. S. A. Glynn. 2006. West Nile virus infections projected from blood donor 2009. Multiplexed real-time PCR assay for discrimination of Plasmodium screening data, United States, 2003. Emerg. Infect. Dis. 12:395402. species with improved sensitivity for mixed infections. J. Clin. Microbiol. 4a.Clark, T. D., D. Njama-Meya, B. Nzarubara, C. Maiteki-Sebuguzi, B. Green- 47:975980. house, S. G. Staedke, M. R. Kamya, G. Dorsey, and P. J. Rosenthal. 2010. 29. Snounou, G., S. Viriyakosol, X. P. Zhu, W. Jarra, L. Pinheiro, V. E. do Incidence of malaria and efcacy of combination antimalarial therapies over Rosario, S. Thaithong, and K. N. Brown. 1993. High sensitivity of detection 4 years in an urban cohort of Ugandan children. PLoS One 5:e11759. of human malaria parasites by the use of nested polymerase chain reaction. 5. Cohen, J. M., D. L. Smith, A. Vallely, G. Taleo, G. Malefoasi, and O. Sabot. Mol. Biochem. Parasitol. 61:315320. 2009. Holding the line, p. 4060. In R. G. A. Feachem, A. Phillips, and G. A. 30. Steenkeste, N., S. Incardona, S. Chy, L. Duval, M. T. Ekala, P. Lim, S. Targett (ed.), A prospectus on malaria elimination. Global Health Group, Hewitt, T. Sochantha, D. Socheat, C. Rogier, O. Mercereau-Puijalon, T. San Francisco, CA. Fandeur, and F. Ariey. 2009. Towards high-throughput molecular detection 6. Coleman, R. E., C. Kumpitak, A. Ponlawat, N. Maneechai, V. Phunkitchar, of Plasmodium: new approaches and molecular markers. Malar. J. 8:86. N. Rachapaew, G. Zollner, and J. Sattabongkot. 2004. Infectivity of asymp- 31. Steenkeste, N., W. O. Rogers, L. Okell, I. Jeanne, S. Incardona, L. Duval, S. tomatic Plasmodium-infected human populations to Anopheles dirus mos- Chy, S. Hewitt, M. Chou, D. Socheat, F. X. Babin, F. Ariey, and C. Rogier. quitoes in western Thailand. J. Med. Entomol. 41:201208. 2010. Sub-microscopic malaria cases and mixed malaria infection in a remote 7. Davis, J. C., T. D. Clark, S. K. Kemble, N. Talemwa, D. Njama-Meya, S. G. Staedke, and G. Dorsey. 2006. Longitudinal study of urban malaria in a area of high malaria endemicity in Rattanakiri province, Cambodia: impli- cohort of Ugandan children: description of study site, census and recruit- cation for malaria elimination. Malar. J. 9:108. 32. Taylor, S. M., J. J. Juliano, P. A. Trottman, J. B. Grif;n, S. H. Landis, P. ment. Malar. J. 5:18. 8. Dorsey, G., S. Staedke, T. D. Clark, D. Njama-Meya, B. Nzarubara, C. Kitsa, A. K. Tshefu, and S. R. Meshnick. 2010. High-throughput pooling and Maiteki-Sebuguzi, C. Dokomajilar, M. R. Kamya, and P. J. Rosenthal. 2007. real-time PCR-based strategy for malaria detection. J. Clin. Microbiol. 48: 512519. Combination therapy for uncomplicated falciparum malaria in Ugandan children: a randomized trial. JAMA 297:22102219. 33. Westreich, D. J., M. G. Hudgens, S. A. Fiscus, and C. D. Pilcher. 1 March 9. Feachem, R., and O. Sabot. 2008. A new global malaria eradication strategy. 2010, posting date. Optimal pooling strategies for acute HIVweb calcula- tor v1.0. mhudgens/optimal.pooling.htm. Lancet 371:16331635. 10. Feachem, R. G. A., and the Malaria Elimination Group. 2009. Shrinking the 34. Westreich, D. J., M. G. Hudgens, S. A. Fiscus, and C. D. Pilcher. 2008. malaria map, a guide on malaria elimination for policy makers. Global Optimizing screening for acute human immunodeciency virus infection Health Group, San Francisco, CA. with pooled nucleic acid amplication tests. J. Clin. Microbiol. 46:17851792. 11. Greenwood, B. M. 2008. Control to elimination: implications for malaria 35. Zanzibar Malaria Control Program. 2009. Malaria elimination in Zanzibar: research. Trends Parasitol. 24:449454. a feasibility assessment. Ministry of Health and Social Welfare, Zanzibar 12. Guerra, C. A., P. W. Gikandi, A. J. Tatem, A. M. Noor, D. L. Smith, S. I. Hay, City, Zanzibar.

Report this document

For any questions or suggestions please email