Advertisement
Research Article

A Large Proportion of P. falciparum Isolates in the Amazon Region of Peru Lack pfhrp2 and pfhrp3: Implications for Malaria Rapid Diagnostic Tests

  • Dionicia Gamboa,

    Affiliations: Instituto de Medicina Tropical “Alexander von Humboldt” Universidad Peruana Cayetano Heredia, Lima, Peru, Departamento de Bioquimica, Biologia Molecular y Farmacologia, Facultad de Ciencias, Universidad Peruana Cayetano Heredia, Lima, Peru

    X
  • Mei-Fong Ho,

    Affiliations: Clinical Tropical Medicine, Queensland Institute of Medical Research, Brisbane, Queensland, Australia, Drug Resistance and Diagnostics, Australian Army Malaria Institute, Brisbane, Queensland, Australia

    X
  • Jorge Bendezu,

    Affiliation: Instituto de Medicina Tropical “Alexander von Humboldt” Universidad Peruana Cayetano Heredia, Lima, Peru

    X
  • Katherine Torres,

    Affiliation: Instituto de Medicina Tropical “Alexander von Humboldt” Universidad Peruana Cayetano Heredia, Lima, Peru

    X
  • Peter L. Chiodini,

    Affiliation: Hospital for Tropical Diseases, London, United Kingdom

    X
  • John W. Barnwell,

    Affiliation: Centers for Disease Control and Prevention, Atlanta, Georgia, United States of America

    X
  • Sandra Incardona,

    Affiliation: Foundation for Innovative New Diagnostics (FIND), Geneva, Switzerland

    X
  • Mark Perkins,

    Affiliation: Foundation for Innovative New Diagnostics (FIND), Geneva, Switzerland

    X
  • David Bell,

    Affiliations: Foundation for Innovative New Diagnostics (FIND), Geneva, Switzerland, World Health Organization - Regional Office for the Western Pacific, Manila, Philippines

    X
  • James McCarthy,

    Affiliations: Clinical Tropical Medicine, Queensland Institute of Medical Research, Brisbane, Queensland, Australia, School of Medicine, University of Queensland, Brisbane, Queensland, Australia

    X
  • Qin Cheng mail

    qin.cheng@defence.gov.au

    Affiliations: Drug Resistance and Diagnostics, Australian Army Malaria Institute, Brisbane, Queensland, Australia, Malaria Drug Resistance and Chemotherapy, Queensland Institute of Medical Research, Brisbane, Queensland, Australia

    X
  • Published: January 25, 2010
  • DOI: 10.1371/journal.pone.0008091

Abstract

Background

Malaria rapid diagnostic tests (RDTs) offer significant potential to improve the diagnosis of malaria, and are playing an increasing role in malaria case management, control and elimination. Peru, along with other South American countries, is moving to introduce malaria RDTs as components of malaria control programmes supported by the Global Fund for AIDS, TB and malaria. The selection of the most suitable malaria RDTs is critical to the success of the programmes.

Methods

Eight of nine microscopy positive P. falciparum samples collected in Iquitos, Peru tested negative or weak positive using HRP2-detecting RDTs. These samples were tested for the presence of pfhrp2 and pfhrp3 and their flanking genes by PCR, as well as the presence of HRP proteins by ELISA. To investigate for geographic extent of HRP-deleted parasites and their temporal occurrence a retrospective study was undertaken on 148 microscopy positive P. falciparum samples collected in different areas of the Amazon region of Peru.

Findings

Eight of the nine isolates lacked the pfhrp2 and/or pfhrp3 genes and one or both flanking genes, and the absence of HRP was confirmed by ELISA. The retrospective study showed that 61 (41%) and 103 (70%) of the 148 samples lacked the pfhrp2 or pfhrp3 genes respectively, with 32 (21.6%) samples lacking both hrp genes.

Conclusions

This is the first documentation of P. falciparum field isolates lacking pfhrp2 and/or pfhrp3. The high frequency and wide distribution of different parasites lacking pfhrp2 and/or pfhrp3 in widely dispersed areas in the Peruvian Amazon implies that malaria RDTs targeting HRP2 will fail to detect a high proportion of P. falciparum in malaria-endemic areas of Peru and should not be used. RDTs detecting parasite LDH or aldolase and quality microscopy should be use for malaria diagnosis in this region. There is an urgent need for investigation of the abundance and geographic distribution of these parasites in Peru and neighbouring countries.

Introduction

Despite several decades of control efforts malaria remains a major infectious disease, causing at least 250 million infections and nearly 1 million deaths per year [1]. The recent significant reductions in prevalence documented in some settings [1] through effective use of Artemisinin-based Combination Therapy (ACT), insecticide-treated bed nets and indoor residual spraying has given cause for optimism and placed malaria eradication back on the global health agenda. The ability to accurately and rapidly diagnose malaria infection in different settings is essential to the success of malaria control and elimination [2]. Accurate diagnosis facilitates appropriate and prompt treatment of febrile illness, reduces drug misuse, and minimises the risk of the development of drug resistance.

Microscopic examination of blood smears has been the traditional method for detecting malaria parasites. The accuracy of microscopy largely relies on the experience and training of the microscopists and the quality of smears. Unfortunately, the quality of microscopy varies significantly, and is often unreliable. Importantly, good quality microscopy is particularly hard to maintain in remote areas where malaria commonly occurs [3]. Rapid Diagnostic Tests (RDTs) for malaria have become increasingly recognized as essential components of efforts to improve diagnostic accuracy in areas where a large part of the population at risk of malaria.

Malaria RDTs are lateral flow devices that detect parasite proteins using antibodies. The tests are easy to perform and provide rapid results (in 15 to 20 min) without the need of electricity, expensive equipment or extensive training. RDTs have, therefore, great potential for rapid and accurate malaria diagnosis in most malaria-endemic areas. Today, over 50 brands of malaria RDTs are manufactured, with over 150 individual products being commercially available. Some detect P. falciparum only, while others detect P. falciparum and other Plasmodium species. Most P. falciparum-detecting RDTs target histidine rich protein 2 (HRP2). HRP2 is an abundant protein produced by all blood stages of P. falciparum [4], and is notable for a number of alanine and histidine rich repeats [5].

Peru is endemic for both P. falciparum and P. vivax, with most cases reported in the Amazon Region [6]. The rate of infection/person/season detected by active and passive case detections in Iquitos, the main city in the Peruvian Amazon region, was reported as 0.13 for P. falciparum and 0.39 for P. vivax in 2003[7]. Like other malaria endemic countries, rapid and accurate malaria diagnosis is critical for malaria control in remotes areas such as the Amazonian region. In areas where microscopy services are not available, diagnosis relies on blood films sent to distant centers for evaluation, with results taking up to 3 weeks to return [8]. Presumptive treatment base on clinical symptoms is common in these areas with a sensitivity as low as 39% [9].

Generally, HRP2-detecting RDTs show comparable sensitivity to quality microscopy, their performance, however, has been reported to be variable in different settings [10][18] including Peru. One study carried out in the Peruvian Amazon during 1998 and 1999 using ParaSight F™ commercial kit, which targets HRP2 antigen, showed 95% of sensitivity [19], [20]. However, other trials conducted in subsequent years using other RDTs that target HRP2, including ParaScreen™[21] and ICT MALARIA pf/pv (AMRAD™) [22], showed a sensitivity below 75%.

The performance of RDTs in the field is influenced by many factors including quality of manufacture, storage conditions [23], ability of the user to correctly interpret the result [24], [25] and parasite density [26]. For RDTs detecting HRP2, genetic diversity of the Pfhrp2 gene and the protein could also affect their performance. Indeed, the deduced amino acid sequences of HRP2 of different isolates are highly polymorphic [27], whereby the number and sequence of specific repeats present in the PfHRP2 proteins varies widely. Moreover the epitopes recognised by the Monoclonal antibodies (MAbs) specific for HRP that are available for and used in RDTs also vary significantly between parasites [28]. While available evidence indicates that HRP2 is the major target detected in HRP2 RDTs, HRP3 is also secreted and can be detected by MAbs raised against HRP2 [28]. The role of HRP3 in performance of HRP diagnostic tests is not well defined.

Another possible factor affecting the sensitivity of HRP2-detecting RDTs is failure of the parasite to express the antigen, due to deletion of the gene or frame shift mutations, or alterations in protein expression. However, to date parasite lines lacking either or both pfhrp2 or pfhrp3 genes have only been identified following laboratory adaptation to in vitro culture [29], [30], and in the progeny of a genetic cross [31], No reports exist in the literature of parasites collected from human patients lacking one or both of these genes.

During our recent characterization of pfhrp2 in parasite isolates collected from several countries as part of the World Health Organisation (WHO) – Foundation for Innovative New Diagnostics (FIND) malaria RDT evaluation program [32], we identified a number of P. falciparum samples collected in Iquitos, Peru, that were positive by microscopy but tested negative with HRP2-detecting RDTs. Further characterisation of these samples revealed that they lacked either pfhrp2 or pfhrp3 or both. Following these findings, we conducted a retrospective study to determine the prevalence of pfhrp2/3 gene deletion in P. falciparum samples collected from different malaria endemic areas in Peru. In this paper we report the genetic characterization and the geographic distribution of the parasites lacking pfhrp2 and/or pfhrp3 in the Peruvian Amazon region. The implications of these findings for various countries in South America that are moving to introduce malaria RDTs under programmes supported by the Global Fund to fight AIDS, Tuberculosis and Malaria (GFATM) are discussed.

Methods

Study Sites and Sample Collection

The samples used in this study came from two sources:

  1. An active case detection survey conducted in 2007 in communities around Iquitos, the main city in the Peruvian Amazon area (between Nanay river, Itaya river and the left margin of the Amazon river), Peru (Figure 1) as part of the WHO –FIND malaria RDT evaluation program. Finger prick blood was collected from 16 P. falciparum patients onto filter paper and dried. Of these 9 samples met the inclusion criteria set for the RDT QA program (HIV-hepatitis B/C free, with no history of antimalarial chemotherapy in the last month, and parasitemia >2000 parasites/µL). Blood spots were transported to the Australian Army Malaria Institute for characterisation. Aliquots of frozen blood samples were sent to the malaria specimen bank at the Centers for Disease Control and Prevention (CDC), Atlanta, USA for species confirmation by 18s rRNA nested PCR and archiving, and to the Hospital for Tropical Disease (HTD), London, UK, for ELISA. The collection, transport and storage of the blood samples were approved by the Human Ethics Committee from Universidad Peruana Cayetano Heredia (UPCH 52707), Peru, while the testing of blood samples for pfhrp2 diversity was approved by the Australian Defence Human Research Ethics Committee (ADHREC 377/05). The remaining 7 samples were characterised together with the retrospective samples.
  2. Samples collected during previous malaria studies from different areas of the Peruvian Amazon. A total of 159 P. falciparum samples, collected in different studies conducted between 2003 and 2007 by the Laboratory of Malaria at the Institute of Tropical Medicine “Alexander von Humboldt”-UPCH, were available for testing. 114/159 samples came from three different projects carried out at 11 Health post centers in peri-urban areas around Iquitos city (Table 1). The location and distance of these health centers to Iquitos city are: Cardozo (3.5 Km), Mazan (38 Km), Moronacocha (2 Km), San Juan de Miraflores (4.6 Km), 9 de octubre (2 Km) El progreso (2.5 Km), Santa Clara de Nanay (12 Km), Santo Tomas (16 Km), Moralillo (15 Km), Varillal (20 Km) and Zungarococha (15 Km) (Figure 1).
thumbnail

Figure 1. Map of Iquitos and the Peruvian Amazon.

The locations of health centers in Iquitos where the P. falciparum samples were collected are shown in solid circles with numbers: 1, Mazan; 2, Moronacocha; 3, Cardozo; 4, San Juan de Miraflores; 5, 9 de octubre; 6, El progreso; 7, Santa Clara de Nanay; 8, Santo Tomas; 9, Zungarococha; 10, Moralillo; 11, Varillal. The locations where samples were collected in other Peruvian Amazon areas are shown as red dots.

doi:10.1371/journal.pone.0008091.g001
thumbnail

Table 1. Origin of the P. falciparum samples used.

doi:10.1371/journal.pone.0008091.t001

The remaining 34 samples came from other areas in the Peruvian jungle as well as from different projects (Table 1). From this group, 12 samples were collected in Yurimaguas, located between the Marañon and Paranapura rivers (445 Km south of Iquitos); 15 samples were from San Lorenzo, located on the shores of the Huallaga river and nine hours from Yurimaguas by boat (110 Km south-west from Iquitos); 7 samples came from the area called High Jungle 2 from Jaen located 620 Km west of Iquitos and 5 samples from Condorcanqui located 524 Km west from Iquitos (Figure 1). The Universidad Peruana Cayetano Heredia Ethical Committee approved the use of parasite DNA for this study.

Other Parasite Isolates and Laboratory Lines

Genomic DNA from 10 P. falciparum field isolates collected as part of the same WHO program: Cambodia (n = 3), Myanmar (n = 2), Nigeria (n = 2), Tanzania (n = 2) and Madagascar (n = 1), as well as from 4 laboratory lines (3D7, 7G8, Dd2 and D10) was used for PCR amplification of genes immediately flanking pfhrp2 and pfhrp3.

Extraction of Parasite DNA

Genomic DNA was isolated from dried blood spots on filter paper, frozen blood or cryopreserved laboratory cultures using the QIAblood Mini™, Quelex™-100 or Qiagen™ kits (QIAGEN) in accordance with the manufacturer's instructions. Phenol-Chloroform method was used to extract parasite DNA from nitrocellulose strips of used RDTs [33].

Confirmation of P. falciparum Infection by PCR

P. falciparum infections were confirmed by PCR amplification of 4 P. falciparum specific genes:18sRNA gene, pfmsp1, pfmsp2 and pfglurp. The P. falciparum 18sRNA gene was amplified using methods described by Padley et al [34], modified by shortening the extension time to 50 sec and extending the number of cycles to 45, or as described by Rubio et al [35]. The amplification pfmsp1, pfmsp2 and pfglurp was performed using the nested primers described elsewhere [36] for 45 cycles.

Detection of the pfhrp2 and pfhrp3 Genes by PCR

The amplification of pfhrp2 and pfhrp3 exon 2 was performed using primers and conditions as described elsewhere [27] for 45 cycles. For the 9 samples collected in 2007, three further PCR protocols were undertaken: to amplify across exons 1 and 2 of both pfhrp2 and pfhrp3 and across exon 1 and the entire exon 2 of pfhrp3 using a second set of primers (Table S1).

Detection of Genes Flanking pfhrp2 and pfhrp3 by PCR

Primers (Table S1) were designed to amplify genes immediately flanking pfhrp2: MAL7P1.230 (5.535 kb upstream) and MAL7P1.228 (6.49 kb downstream), and those immediately flanking pfhrp3: MAL13P1.485 (4.404 kb upstream) and MAL13P1.475 (1.684 kb downstream). The PCR conditions and expected product sizes are described in Table S1.

DNA Sequencing

PCR products were purified by the use of spin columns (Roche), and used in a standard dye-terminator DNA sequencing reaction (ABI).

Detection of HRP2 Protein Using ELISA

The quantity of HRP2 protein in the 9 samples collected in 2007 was measured using the Malaria Ag CELISA kit (CELLABS Pty, Sydney, Australia) following manufacturer's instructions. The protein was quantified by plotting absorbance values against a standard curve of recombinant HRP2. Protein levels for parasite pLDH (Standard Diagnostics, Kyonggi-do, Korea) and aldolase (Centers for Disease Control, Atlanta, USA) were also measured by ELISA with recombinant protein standard curves [37].

Results

Eight of the 9 P. falciparum Samples Collected during the 2007 Survey in Iquitos Lack pfhrp2 and pfhrp3

As part of the WHO – FIND malaria RDT evaluation program [32] we undertook characterization of pfhrp2 in 9 P. falciparum patient samples collected in Iquitos, Peru. Eight of the nine samples did not yield the expected PCR product using our standard primers spanning the full length of pfhrp2 exon 2, or in subsequent experiments using primers spanning a shorter sequence of exon 1 and exon 2 (Table 2, Figure S1, S2), indicating that these parasites lacked the pfhrp2 gene. In further testing 4 of these 9 samples also tested negative for the pfhrp3 gene by PCR specific for exon 2 of this gene. Among the 5 samples that had yielded the expected PCR products for exon 2 of pfhrp3 two (PE01F07 and PE01F11) failed to yield a product with primers spanning exon 1 and exon 2, or exon 1 and full length exon 2 of pfhrp3 (Table 2, Figure S2) indicating a truncation of exon 1.

thumbnail

Table 2. Gene amplification and protein detection results for the 9 samples collected in 2007 (positive and negative results are represented by + and − respectively).

doi:10.1371/journal.pone.0008091.t002

Confirmation that the Parasite Isolates Were P. falciparum Infections and Investigation for Genetic Diversity of the Parasites

To verify the malaria parasite species and to investigate genetic diversity in the parasites, a multiplex PCR reaction that amplifies the 18s rRNA gene of 4 Plasmodium species commonly infecting humans, and three polymorphic P. falciparum genes pfmsp1, pfmsp2 and pfglurp was undertaken. Results of these experiments confirmed that all 9 samples were P. falciparum, with no evidence of contamination or co-infection with another species (Table 2, Figure S2). Of note, the sizes of the PCR products of the polymorphic genes pfmsp1, pfmsp2 and pfglurp were observed to vary, revealing 2, 2 and 4 allelic types at these loci respectively and establishing that the isolates were not clonal in nature. The combined three-loci-type separated the isolates into four different haplotypes: 1) PE01F04 and PE0F06, 2) PE01F07, 3) PE01F11 and 4) PE01F15 to PE01F19 (Table 2).

Investigation of the Genetic Basis of These Deletions

To investigate for larger chromosomal deletions encompassing the two hrp genes, PCR reactions were undertaken to amplify the two genes flanking each of pfhrp2 and pfhrp3. The specificity of these PCR reactions was first tested on well characterized laboratory lines of P. falciparum. PCR products of the expected size were amplified from two laboratory lines, 3D7 and 7G8, and the identity of the 3D7 PCR product confirmed by DNA sequence analysis. PCR amplification for pfhrp2 in Dd2, a laboratory line known to lack pfhrp2 failed to amplify pfhrp2 and both flanking genes; appropriate PCR products for pfhrp3 and both flanking genes were amplified from this line. PCR reactions undertaken with D10, another laboratory line known to lack pfhrp2, failed to amplify pfhrp2 and the upstream gene, but a PCR product of the expected size was present for the downstream gene, MAL7P1.228. Likewise, PCR amplification of pfhrp3 and its flanking genes was successful for D10. PCR reactions were then undertaken with 10 field isolates from different geographic areas that had yielded the expected pfhrp2 and pfhrp3 products, and also yielded products of the expected size for the four genes flanking both pfhrp2 and pfhrp3 (Table S2).

When the Peruvian isolate PE01F04, the only isolate positive for pfhrp2 in this subset, was subject to PCR for the genes flanking pfhrp2, it yielded products for both flanking genes as was observed with 3D7 and 7G8. PCR reactions undertaken with the remaining 8 Peruvian samples, which were negative for pfhrp2, resulted in amplification of the expected product for the downstream gene but failed to amplify the gene upstream of pfhrp2, in a pattern resembling that observed with D10. Isolates PE01F04, PE01F06 and PE01F15 that had yielded phfhp3 PCR products were also PCR-positive for both genes flanking pfhrp3. PE01F07 and PE01F11, isolates where evidence had indicated that may have truncated exon 1 were PCR positive for the upstream gene but negative for the downstream gene. The four samples that had failed to yield a PCR product for pfhrp3 were all PCR-negative for the upstream flanking gene, and 3 of the 4 samples were also negative for the downstream gene (Table 2, Figure S2).

Verification that These Parasites Do Not Produce HRP Protein

To investigate whether these parasites lacked histidine rich protein all 9 samples were screened for HRP2 protein levels by a HRP antigen capture ELISA. Only one of the 9 samples (PE01F04) had detectable histidine rich protein 2, giving an estimated 235 ng/mL protein (Table 2). In contrast, all 9 samples had measurable levels of pLDH and aldolase. When tested on 3 different commercially available HRP2-detecting RDTs, PE01F04 again was the only sample test positive with all 3 RDTs. Two samples lacking pfhrp2 but having intact pfhrp3 (PE01F06 and PE01F15) tested weakly positive on the ICT combo kit (Cape Town, South Africa). PE01F06 tested weakly positive but PE01F15 was negative with 2 other RDT brands (CareStart Malaria HRP2 and CareStart Malaria Combo. AccessBio, Monmouth, NJ, USA). The remaining 5 samples lacking both phfrp2 and pfhrp3 were negative on all 3 different RDTs.

A High Proportion of Retrospectively Collected Samples from the Amazon Region of Peru Lack pfhrp2 and pfhrp3 Genes

We then undertook study of the presence of hrp-deleted parasites from other areas of Peru. PCR amplifications of the 18SrDNA gene and pfglurp was performed for 159 samples that had been collected in previous studies, of which 148 were positive for both markers confirming P. falciparum infection and indicating a good quality of DNA. The size of the PCR product for pfglurp varied ranging from 550 to 900 bp, indicating a total of 6 genotypes (Table 3). These 148 samples were then tested for the presence of pfhrp2 and pfhrp3 by PCR.

thumbnail

Table 3. pfglurp allelic type detected in the retrospective samples.

doi:10.1371/journal.pone.0008091.t003

Of 114 samples collected from peri-urban areas of Iquitos, PCR for the pfhrp2 and the pfhrp3 genes failed in 36% and 67% of the samples respectively (Table 4), despite being positive for 18sRNA and pfglurp. The proportion of samples failing to amplify both pfhrp2 and pfhrp3 was 18.4% (21/114), while the proportion of samples positive for both genes was only 15.78% (18/114).

thumbnail

Table 4. Proportion of P. falciparum isolates positive or negative for pfhrp2/3 genes in different areas of the Peruvian Amazon.

doi:10.1371/journal.pone.0008091.t004

pfhrp2 and pfhrp3 Deleted Parasites are Widely Distributed in the Peruvian Amazon

Parasites lacking pfhrp2 and pfhrp3 were also identified in other areas in the Peruvian jungle. The proportion of pfhrp2 and pfhrp3 positive and negative in samples collected in Condorcanqui, San Lorenzo, Yurimaguas and Jaen is shown in table 4. The proportion varied between parasites collected in different areas. While the number of samples collected in these areas was small, overall a high proportion of parasites negative for pfhrp2 and pfhrp3 was observed in these areas and was comparable to levels observed around Iquitos. When all areas are combined, we observed that a total of 41% and 70% of the samples were negative to pfhrp2 and pfhrp3, respectively (Table 4). The percentage of samples that were negative for both pfhrp2 and pfhrp3 was 21.6% (32/148), while the percentage of samples that were positives for both pfhrp2 and pfhrp3 was 13.5% (20/148).

Amplification Stringency

In order to rule out the possibility that failure to PCR amplify the pfhrp2/3 genes was due to possible polymorphisms at the primer binding sites, an additional PCR reaction was undertaken for samples that were negative for either genes using the same primers with lower annealing temperatures (down to 42°C). All samples and the negative controls remained negative, while the positive control (NF54) was positive, suggesting that no small mismatches occurred at the primer sites (Figure S3).

Discussion

Laboratory lines of P. falciparum that lack pfhrp2 or pfhrp3 or both genes have been reported following adaptation to and long term growth under in vitro culture conditions [29][31]. However, this is the first report of clinical isolates of parasites taken directly from infected human subjects in endemic areas lacking these genes. As part of the WHO-FIND malaria RDT evaluation program we have examined pfhrp2 sequence variation of over 500 clinical parasite samples collected from many different malaria endemic areas in Africa, South East Asia, West Pacific and South America ([27] and Baker et al unpublished) and have not encountered P. falciparum samples lacking pfhrp2 and/or pfhrp3 elsewhere. In Peru, we identified, for the first time, not only parasites from clinical patients lacking these hrp genes but also a wide spread pattern of parasites lacking these genes in the Peruvian Amazon with an overall frequency of 41% and 70% of parasites without pfhrp2 and pfhrp3, respectively and 21.6% for double deletions.

pfhrp2 and/or pfhrp3 PCR Negative Samples Lack These Genes Due to Chromosomal Deletions

A plausible hypothesis for the negative PCR results for pfhrp2 and/or pfhrp3 is that these parasites lack these genes as a consequence of chromosomal deletion. A commonly used method for characterizing genetic rearrangements and deletions is a genomic hybridization where parasite genomic DNA or parasite chromosomes are hybridised with the target gene (generally by Southern Blot), in this case, pfhrp2 or pfhrp3. Since we only had blood spots on filter paper and small aliquots of frozen whole blood, there was insufficient parasite genomic DNA to use this method. Instead, we performed a number of independent PCR experiments to characterise the samples to obtain supporting evidence that the parasites actually lack these genes. These experiments also ensured that the negative PCR reactions for pfhrp2 and pfhrp3 were not due to poor quality or insufficient parasite DNA. To rule out the possibility of sequence polymorphism at primer binding sites, further PCR experiments were undertaken using alternative primers designed to anneal at highly conserved sequence stretches in both pfhrp2 and pfhrp3. All isolates that were PCR negative using the exon 2 pfhrp2 and pfhrp3 primers were also PCR negative using these latter primers.

Deletions of pfhrp2 and pfhrp3 previously reported in laboratory lines have all been shown to result from chromosomal deletions, where a large fragment of a chromosome usually containing several genes is deleted [29][31]. Further evidence that 8 of the 9 and 6 out 9 Iquitos parasites lack pfhrp2 or both pfhrp2 and pfhrp3, respectively, is provided by the observation that the negative PCR reactions for the flanking genes were consistent with those for pfhrp2 and pfhrp3. Taken together, the results suggest that a fragment on chromosome 7 that contains pfhrp2 and its immediate upstream gene (MAL7P1.230) encoding a hypothetical protein has been deleted in these parasites. In all 8 isolates negative for pfhrp2, the gene immediately downstream from pfhrp2 encoding HSP70 is preserved. On chromosome 13, PCR reactions for both pfhrp3 and the genes immediately upstream (MAL13P1.485), and in most cases for the downstream gene (MAL13P1.475) were negative for 4 of the 9 samples, indicating a chromosomal deletion. This finding is supported by independent data showing that chromosomal deletions in both the region on chromosome 7 where pfhrp2 is located, and the region on chromosome 13 where pfhrp3 is located, are possible and have been observed in some laboratory parasite lines [38].

Parasite Lack pfhrp2 and/or pfhrp3 Do Not Express HRP2 Protein and Are Not Detectable by HRP2 Detecting RDTs

Most importantly from the standpoint of diagnostic assay performance, the final evidence of gene deletion or inactivation, came from the assay for histidine-rich protein by ELISA where results show that with the exception of one parasite isolate none have detectable histidine rich protein 2, though all isolates have detectable pLDH. The single isolate in this subset of samples in which histidine-rich protein could be detected was positive for both pfhrp2 and pfhrp3. When tested using HRP2-detecting RDTs, parasites having double gene deletions all tested negative on different HRP2-detecting RDTs demonstrating the lack of antigens in these parasites. Parasites having deleted only phfrp2 but not pfhrp3 were detectable by some RDTs but not others resembling the ELISA results due, most likely, to different antibodies used in different RDTs or ELISA test which recognise different epitopes that may or may not be shared between pfhrp2 and pfhrp3 [28].

A Large Proportion of P. falciparum Parasites in Peruvian Amazon Lack pfhrp2 and pfhrp3

Evidence for a widespread distribution of HRP-deleted parasites across the Peruvian Amazon comes from our observation that a large proportion of parasites lacking pfhrp2 and or pfhrp3 were observed in other areas from the Peruvian jungle between 110 and 620 kilometres from Iquitos (Condorcanqui, San Lorenzo, Yurimaguas and Jaen) and collected over a period of 4 years. These findings indicate that the hrp-negative parasites are not restricted to a specific area, but are distributed in much of the malaria endemic areas in Peru, and raises the possibility that such parasites may be present in neigbouring countries with identical patterns of malaria and sharing the same river system for transport.

The Absence of HRP2 and HRP3 Likely Explains the Poor Performance of Some HRP2-Detecting RDTs Previously Reported in the Peruvian Amazon

The previously reported sensitivity for these tests ranged from 70% [21] to 65% (MALARIA pf/pv (AMRAD™)) [22], consistent with our findings, while pfLDH-detecting RDTs (OptiMAL) have shown higher sensitivity in the Peruvian Amazon [39][42]. ParaSight F (Becton-Dickensen, USA) was trialed in Iquitos for several years before 2000 showing good sensitivity in detecting P. falciparum infections [19], [20]. It is possible that at that time, parasites lacking pfhrp2 were not present in high proportions to affect the performance of this test. Therefore, the widespread of parasites lacking pfhrp2 gene may be a relatively recent event.

Parasite Evolution

The evolutionary process that produced these parasites is unknown but intriguing. It is notable that the proportion of parasites lacking pfhpr3 is higher than those lacking pfhrp2 in samples collected from almost all locations, while the proportion of parasites lacking both genes is 21.6%. This suggests that parasites lacking pfhrp3 may have been present in Peru longer than those lacking pfhrp2. It is possible that parasites lacking pfhrp3 had undergone a genetic cross with parasites lacking pfhrp2 and produced progeny lacking both genes. As four different pfmsp1/pfmsp2/pfglurp haplotypes were observed in the subset of 9 isolates lacing pfhrp2, chromosomal deletions on chromosome 7 may have occurred frequently in the area. A similar genetic cross event and outcome has been reported in a laboratory cross where Dd2 (pfhrp2 negative) and HB3 (pfhrp3 negative) were crossed. While most progeny inherited either a pfhrp2 or a pfhrp3 deletion, one progeny (3BD5) lacked both pfhrp2 and pfhrp3 according to genetic linkage analysis [31].

While the function(s) of pfhrp2 and pfhrp3 is unclear, they do not appear to be essential for in vitro growth because progeny lacking either gene or both pfhrp2 and pfhrp3 remain viable in vitro, with exhibition of a rapid growth phenotype. This observation of wild-type parasites lacking both pfhrp2 and pfhrp3 from patients in Peru indicate that these parasites are viable in vivo. Further study of parasite virulence and fitness in these deletion mutants would be of interest, but they are obviously capable of transmission and of causing disease as all cases were symptomatic. Certainly, the mechanism of selection that has driven these parasite genotypes to be common in this Peruvian population of P. falciparum remains to be determined.

HRP2-Detecting RDTs Are Not Reliable in Peru

The most important implication of this finding is that, in most malaria endemic areas of Peru, diagnosis of the malaria due to P. falciparum, the species responsible for almost all malaria mortality and much of the morbidity, would commonly fail if HRP2-detecting RDTs are used. This raises a serious question concerning on the value of the use of HRP2-detecting RDTs in Peru. Because P. falciparum is endemic in a great area of the Peruvian Amazon with 98% of the falciparum malaria cases in Peru occur in this area [6], any malaria RDT used here must detect P. falciparum with high sensitivity. The high proportion of samples in which pfhrp2 could not be amplified (41%) and high proportion of samples lacked both pfhrp2 and pfhrp3 (21.6%) in this study, which were collected from different areas and at different time points, clearly demonstrate that HRP2-detecting RDTs have limited reliability for detecting P. falciparum in Peru. RDTs targeting other antigens (pLDH and aldolase) and quality microscopy should be used for malaria diagnosis.

As Peru borders on several countries that share the Amazon River basin and where malaria transmission occurs without respect for national borders, it is unlikely that parasites lacking pfhrp2 and pfhrp3 are confined to Peru. It is therefore important that investigations be performed in other areas in South America urgently where P. falciparum is endemic to determine the presence and geographical spread of parasites lacking the pfhrp2 and pfhrp3 genes before the large scale implementation of malaria RDTs in this area. Investigations should also be carried out to monitor the presence and spread of parasites with gene deletions in areas outside of South America to ensure the best performance of malaria RDTs globally.

Supporting Information

Table S1.

Primer sequences, PCR conditions and expected product sizes

doi:10.1371/journal.pone.0008091.s001

(0.01 MB DOC)

Table S2.

Amplification of pfhrp2, pfhrp3 and their immediate flanking genes in laboratory lines and field isolates.

doi:10.1371/journal.pone.0008091.s002

(0.01 MB DOC)

Figure S1.

Schematic illustration of pfhrp2 and pfhrp3 gene structures and primer binding sites. Filled arrows represent primers that amplify the full length exon 2; open arrows representing primers amplify across exon 1 and exon 2.

doi:10.1371/journal.pone.0008091.s003

(0.05 MB TIF)

Figure S2.

PCR products visualised on agarose gels. Panels a) to l) shows the PCR results for: a) pfhrp2 exon 2, b) pfhrp2 exon 1-exon 2, c) pfhrp3 exon 2, d) pfhrp3 exon 1-exon 2, e) 18s rRNA, f) pfglurp, g) pfmsp1, h) pfmsp2, i) MAL7P1.228, j) MAL7P1.230, k) MAL13P1.475, l) MAL13P1.480. Numbers 1 to 12 represent lanes on each gel: 1: Marker; 2: PE01 F04; 3: PE01 F06; 4: PE01 F07; 5: PE01 F11; 6: PE01 F15; 7: PE01 F16; 8: PE01 F17; 9: PE01 F18; 10: PE01 F19; 11: 3D7; 12: No DNA

doi:10.1371/journal.pone.0008091.s004

(0.51 MB TIF)

Figure S3.

Temperature Gradient for PCR. A: pfhrp2 positive samples. Lane 1: 100 pb plus Molecular marker (MM) - RocheTM; Lane 2: 42.0°C; Lane 3: 43.0°C; Lane 4: 45.0°C; Lane 5: 47.0°C; Lane 6: 48.0°C; Lane 7: 50.0°C; Lane 8: NF54; Lane 9: Negative control (NC) to 50.0°C. B: pfhrp2 negative samples. Lane 1: MM; Lane 2: 50.0°C; Lane 3: 48°C; Lane 4: 47.0°C; Lane 5: 45.0°C; Lane 6: 44.0°C; Lane 7: 43.0°C; Lane 8: 42.0°C; Lane 9: NF54 (50.0°C); Lane 10: NC. C: pfhrp3 positive samples. Lane 1: MM; Lane 2: 50.0°C; Lane 3: 48.0°C; Lane 4: 47.0°C; Lane 5: 45.0°C; Lane 6: 44.0°C; Lane 7: 43.0°C; Lane 8: 42.0°C; Lane 9: NC. D: pfhrp3 negative samples. Lane 1: MM; Lane 2: 50.0°C; Lane 3: 48.0°C; Lane 4: 47.0°C; Lane 5: 45.0°C; Lane 6: 44.0°C; Lane 7: 43.0°C; Lane 8: 42.0°C; Lane 9: 41.0°C; Lane 10: 40.0°C; Lane 11: NC.

doi:10.1371/journal.pone.0008091.s005

(0.29 MB TIF)

Acknowledgments

The authors would like to thank colleagues in Institut Pasteur du Cambodge, Cambodia; Ifakara Health Research and Development Centre, Tanzania; Department of Medical Research, Myanmar; University of Lagos, Nigeria and Institut Pasteur de Madagascar of the WHO Malaria RDT Quality Assurance Network for providing help in sample collection and shipment. The opinions expressed herein are those of the authors and do not necessarily reflect those of the WHO or Defence Health Service or any extant policy of Department of Defence, Australia.

Author Contributions

Conceived and designed the experiments: DG PC JWB MDP DB JM QC. Performed the experiments: MFH JB KJT. Analyzed the data: DG MFH JB KJT PC JWB SI DB JM QC. Wrote the paper: DG MFH JWB DB JM QC. Coordinated sample collections: SI MDP DB.

References

  1. 1. World Health Organization (2008) World Malaria Report 2008. Geneva, Switzerland. http://www.who.int/malaria/publications/​atoz/9789241563697/en/index.html.
  2. 2. Perkins MD, Bell DR (2008) Working without a blindfold: the critical role of diagnostics in malaria control. Malar J 7: Suppl 1S5.
  3. 3. Wongsrichanalai C, Barcus MJ, Muth S, Sutamihardja A, Wernsdorfer WH (2007) A review of malaria diagnostic tools: microscopy and rapid diagnostic test (RDT). Am J Trop Med Hyg 77: 119–127.
  4. 4. Howard RJ, Uni S, Aikawa M, Aley SB, Leech JH, et al. (1986) Secretion of a malarial histidine-rich protein (Pf HRP II) from Plasmodium falciparum-infected erythrocytes. J Cell Biol 103: 1269–1277.
  5. 5. Wellems TE, Howard RJ (1986) Homologous genes encode two distinct histidine-rich proteins in a cloned isolate of Plasmodium falciparum. Proc Natl Acad Sci U S A 83: 6065–6069.
  6. 6. Minsa (2009) Boletin Epidemiologico 2009. (Lima) 18 (07). www.dge.gob.pe.
  7. 7. Branch O, Casapia WM, Gamboa DV, Hernandez JN, Alava FF, et al. (2005) Clustered local transmission and asymptomatic Plasmodium falciparum and Plasmodium vivax malaria infections in a recently emerged, hypoendemic Peruvian Amazon community. Malar J 4: 27.
  8. 8. Durand S, Cesar R, Maria H, Cabezas C (2005) Oportunidad en el diagnostico y tratamiento de la Malaria en comunidades periurbanas de la Amazonia Peruana. Rev Peru Med Exp Salud Publica 22, (1): 47–53.
  9. 9. Arróspide N (2005) Tratamiento presuntivo de Malaria por promotores de salud en comunidades periurbanas de Iquitos. Rev Peru Med Exp Salud Publica 22: 241–242.
  10. 10. Wongsrichanalai C, Chuanak N, Tulyayon S, Thanoosingha N, Laoboonchai A, et al. (1999) Comparison of a rapid field immunochromatographic test to expert microscopy for the detection of Plasmodium falciparum asexual parasitemia in Thailand. Acta Trop 73: 263–273.
  11. 11. Bechem NN, Leke RF, Tietche F, Taylor DW (1999) Evaluation of a rapid test for histidine rich protein 2 for diagnosis of Plasmodium falciparum infection in Cameroonian children. Trans R Soc Trop Med Hyg 93: 46.
  12. 12. Stow NW, Torrens JK, Walker J (1999) An assessment of the accuracy of clinical diagnosis, local microscopy and a rapid immunochromatographic card test in comparison with expert microscopy in the diagnosis of malaria in rural Kenya. Trans R Soc Trop Med Hyg 93: 519–520.
  13. 13. Gaye O, Diouf M, Dansokho EF, McLaughlin G, Diallo S (1998) Diagnosis of Plasmodium falciparum malaria using ParaSight F, ICT malaria PF and malaria IgG CELISA assays. Parasite 5: 189–192.
  14. 14. Beadle C, Long GW, Weiss WR, McElroy PD, Maret SM, et al. (1994) Diagnosis of malaria by detection of Plasmodium falciparum HRP-2 antigen with a rapid dipstick antigen-capture assay. Lancet 343: 564–568.
  15. 15. Rubio JM, Buhigas I, Subirats M, Baquero M, Puente S, et al. (2001) Limited level of accuracy provided by available rapid diagnosis tests for malaria enhances the need for PCR-based reference laboratories. J Clin Microbiol 39: 2736–2737.
  16. 16. Iqbal J, Khalid N, Hira PR (2002) Comparison of two commercial assays with expert microscopy for confirmation of symptomatically diagnosed malaria. J Clin Microbiol 40: 4675–4678.
  17. 17. Singh N, Valecha N (2000) Evaluation of a rapid diagnostic test, ‘Determine malaria pf’, in epidemic-prone, forest villages of central India (Madhya Pradesh). Ann Trop Med Parasitol 94: 421–427.
  18. 18. Huong NM, Davis TM, Hewitt S, Huong NV, Uyen TT, et al. (2002) Comparison of three antigen detection methods for diagnosis and therapeutic monitoring of malaria: a field study from southern Vietnam. Trop Med Int Health 7: 304–308.
  19. 19. Forney JR, Wongsrichanalai C, Magill AJ, Craig LG, Sirichaisinthop J, et al. (2003) Devices for rapid diagnosis of Malaria: evaluation of prototype assays that detect Plasmodium falciparum histidine-rich protein 2 and a Plasmodium vivax-specific antigen. J Clin Microbiol 41: 2358–2366.
  20. 20. Forney JR, Magill AJ, Wongsrichanalai C, Sirichaisinthop J, Bautista CT, et al. (2001) Malaria rapid diagnostic devices: performance characteristics of the ParaSight F device determined in a multisite field study. J Clin Microbiol 39: 2884–2890.
  21. 21. Bendezu J (2008) Evaluación en campo de la prueba rápida ParaScreen® comparado con Semi-nested Multiplex PCR (SnM-PCR) y Microscopía para el diagnóstico de Malaria. Universidad Nacional Federico Villareal, Lima 110.
  22. 22. Llanos-Zavalaga F, Villacorta J, Reyes R, Lecca L, Mendoza D, et al. (2002) Evaluación de la prueba ICT MALARIA P. f/P.v (AMRAD®) para la deteccion de P. falciparum y P. vivax en una zona endémica de la Amazonía Peruana. Rev Peru Med Exp Salud Publica 19: 39–42.
  23. 23. Chiodini PL, Bowers K, Jorgensen P, Barnwell JW, Grady KK, et al. (2007) The heat stability of Plasmodium lactate dehydrogenase-based and histidine-rich protein 2-based malaria rapid diagnostic tests. Trans R Soc Trop Med Hyg 101: 331–337.
  24. 24. Harvey SA, Jennings L, Chinyama M, Masaninga F, Mulholland K, et al. (2008) Improving community health worker use of malaria rapid diagnostic tests in Zambia: package instructions, job aid and job aid-plus-training. Malar J 7: 160.
  25. 25. Rennie W, Phetsouvanh R, Lupisan S, Vanisaveth V, Hongvanthong B, et al. (2007) Minimising human error in malaria rapid diagnosis: clarity of written instructions and health worker performance. Trans R Soc Trop Med Hyg 101: 9–18.
  26. 26. Bell DR, Wilson DW, Martin LB (2005) False-positive results of a Plasmodium falciparum histidine-rich protein 2-detecting malaria rapid diagnostic test due to high sensitivity in a community with fluctuating low parasite density. Am J Trop Med Hyg 73: 199–203.
  27. 27. Baker J, McCarthy J, Gatton M, Kyle DE, Belizario V, et al. (2005) Genetic diversity of Plasmodium falciparum histidine-rich protein 2 (PfHRP2) and its effect on the performance of PfHRP2-based rapid diagnostic tests. J Infect Dis 192: 870–877.
  28. 28. Lee N, Baker J, Andrews KT, Gatton ML, Bell D, et al. (2006) Effect of sequence variation in Plasmodium falciparum histidine- rich protein 2 on binding of specific monoclonal antibodies: Implications for rapid diagnostic tests for malaria. J Clin Microbiol 44: 2773–2778.
  29. 29. Kemp DJ, Thompson JK, Walliker D, Corcoran LM (1987) Molecular karyotype of Plasmodium falciparum: conserved linkage groups and expendable histidine-rich protein genes. Proc Natl Acad Sci U S A 84: 7672–7676.
  30. 30. Wellems TE, Oduola AMJ, Phenton B, Des Jardins R, Panton LJ, et al. (1988) Chromosome size variation occurs in cloned Plasmodium falciparum on in vitro cultivation. Braz J Genet 11: 813–825.
  31. 31. Su X, Ferdig MT, Huang Y, Huynh CQ, Liu A, et al. (1999) A genetic map and recombination parameters of the human malaria parasite Plasmodium falciparum. Science 286: 1351–1353.
  32. 32. Bell D, Wongsrichanalai C, Barnwell JW (2006) Ensuring quality and access for malaria diagnosis: how can it be achieved? Nat Rev Microbiol 4: S7–20.
  33. 33. Veron V, Carme B (2006) Recovery and use of Plasmodium DNA from malaria rapid diagnostic tests. Am J Trop Med Hyg 74: 941–943.
  34. 34. Padley D, Moody AH, Chiodini PL, Saldanha J (2003) Use of a rapid, single-round, multiplex PCR to detect malarial parasites and identify the species present. Ann Trop Med Parasitol 97: 131–137.
  35. 35. Rubio JM, Benito A, Roche J, Berzosa PJ, Garcia ML, et al. (1999) Semi-nested, multiplex polymerase chain reaction for detection of human malaria parasites and evidence of Plasmodium vivax infection in Equatorial Guinea. Am J Trop Med Hyg 60: 183–187.
  36. 36. Ranford CL, Taylor J, Umasunthar T, Taylor LH, Babiker HA, et al. (1997) Molecular analysis of recrudescent parasites in a Plasmodium falciparum drug efficacy trial in Gabon. Trans R Soc Trop Med Hyg 91: 719–724.
  37. 37. WHO Regional Office for the Western Pacific, UNDP/World Bank/WHO/TDR, Foundation for Innovative New Diagnostics (FIND) (2008) Methods manual for laboratory quality control testing of malaria rapid diagnostic tests. http://www.wpro.who.int/sites/rdt/who_rd​t_evaluation.
  38. 38. Jiang H, Yi M, Mu J, Zhang L, Ivens A, et al. (2008) Detection of genome-wide polymorphisms in the AT-rich Plasmodium falciparum genome using a high-density microarray. BMC Genomics 9: 398.
  39. 39. Arróspide N, Puray M, Guzmán E, Verano M, Medina S, et al. (2001) Estudio multicentrico de DIAMED OptiMAL para el diagnostico de P. falciparum y P.vivax en áreas endémicas del Perú Rev Peru Med Exp Salud Publica 19: 76–81.
  40. 40. Tarazona AS, Zerpa LS, Requena DM, Llanos-Cuentas A, Magill A (2004) Evaluation of the Rapid Diagnostic Test OptiMAL for Diagnosis of Malaria due to Plasmodium vivax. The Brazilian Journal of Infectious Diseases 8: 151–155.
  41. 41. Arróspide N, Flores R, Ruíz J (2006) Evaluación de una prueba rápida para el diagnóstico de Malaria en áreas endémicas del Perú. Rev Peru Med Exp Salud Publica 23: 81–86.
  42. 42. Arróspide N, Puray M, Guzman E, Verano M, Gutierrez S, et al. (2004) Uso de pruebas rápidas inmunocromatográficas para la detección de Plasmodium falciparum en donantes de sangre en Perú. Rev Peru Med Exp Salud Publica 21: 76–81.