A unique methodology for detecting the spread of chloroquine-resistant strains of Plasmodium falciparum, in previously unreported areas, by analyzing anophelines of malaria endemic zones of Orissa, India

Infection, Genetics and Evolution 9 (2009) 462–467

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A unique methodology for detecting the spread of chloroquine-resistant strains of Plasmodium falciparum, in previously unreported areas, by analyzing anophelines of malaria endemic zones of Orissa, India Amitav Mohanty a, Sunita Swain b, Durg V. Singh a, Namita Mahapatra b, Santanu K. Kar b, Rupenangshu K. Hazra b,* a b

Division of Vector Borne Diseases, Institute of Life Sciences, Bhubaneswar, India Regional Medical Research Centre, Bhubaneswar, India

A R T I C L E I N F O

A B S T R A C T

Article history: Received 24 October 2008 Received in revised form 5 January 2009 Accepted 9 January 2009 Available online 20 January 2009

Generally, clinical data is referred to study drug-resistance patterns of Plasmodium falciparum in an area. This is only possible after a clear manifestation of drug-resistance parasites inside the human host, and thereafter detection by healthcare persons. The detection of spread of drug-resistant P. falciparum in a population, before any pathological symptoms detected in humans is possible by analyzing the anopheline vectors, transmitting malaria. In the present study we implemented a new strategy to detect the spread of chloroquine-resistant (CQR) strains of P. falciparum by the major malaria vectors prevalent in selected endemic regions of Orissa, India. We screened P. falciparum positive vectors by using polymerase chain reaction (PCR)-based assay and thereafter detected K76T mutation in the Pfcrt gene, the chloroquine-resistance marker, of parasites present within the vectors. This study showed higher transmission rate of chloroquine-resistant P. falciparum parasites by Anopheles culicifacies and Anopheles fluviatilis. This study will help in assigning chloroquine-resistant P. falciparum sporozoite transmission potential of malaria vectors and suggest that by adopting the mentioned methodologies, we can detect the spreading of the drug-resistant P. falciparum in its transmission. This approach of studying the anophelines during regular vector collection and epidemiological analysis will give the knowledge of chloroquine-resistance pattern of P. falciparum of an area and help in devising effective malaria control strategy. ß 2009 Elsevier B.V. All rights reserved.

Keywords: Malaria Chloroquine resistance (CQR) Anopheles Plasmodium falciparum Pfcrt K76T

1. Introduction We are still in the timeline waiting for an effective malaria vaccine, and disease management with prompt and effective treatment with drug therapy remains the mainstay of malaria control. Chloroquine has been by far the most important and successful drug used to treat falciparum malaria (Wellems and Plowe, 2001). After its introduction to malaria therapy in the 1940s, chloroquine soon became the drug of choice for therapy and prevention. It was because of low toxicity, low production costs and high efficacy against malaria parasites. However, due to extensive use, resistance developed simultaneously in the late 1950s in Southeast Asia and South America and spread rapidly to other endemic areas (Jelinek et al., 2002; Keen et al., 2007).

* Corresponding author at: Regional Medical Research Centre, Nalco Square, Chandrasekharpur, Bhubaneswar, 751 023, Orissa. India. Tel.: +91 6742301416; fax: +91 6742301351. E-mail address: [email protected] (R.K. Hazra). 1567-1348/$ – see front matter ß 2009 Elsevier B.V. All rights reserved. doi:10.1016/j.meegid.2009.01.005

Resistant parasites manage to lessen the accumulation of chloroquine, thereby nullify the drug effect (Bray et al., 1998). Chloroquine-resistant (CQR) falciparum malaria has been associated with great increase in morbidity and mortality in malaria endemic regions (Greenberg et al., 1989; Trape et al., 1998). In spite of its diminishing efficacy due to emergence of resistant strains of parasite, chloroquine remains the primary antimalarial agent in many endemic areas because no affordable alternatives are available. This also further aid in spreading of the resistance. Chloroquine acts by interfering with heme metabolism in the digestive vacuole of Plasmodium falciparum. In resistant parasites, the accumulation of chloroquine inside the vacuole is diminished (Fitch, 1970; Bray et al., 1998). The P. falciparum multidrug resistance 1 (Pfmdr1) homolog that encodes the P-glycoprotein (Pgh1) was initially proposed as determinant of the CQR phenotype (Foote et al., 1989), but a direct association of mutations in this gene with resistance could not be established (Zakeri et al., 2008). The locus for CQR was mapped to a 36-kb segment on chromosome 7 of P. falciparum and specifically linked to the polymorphic gene Pfcrt, a gene with 13 exons, encodes PfCRT, a trans-membrane

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protein in the digestive vacuoles of malaria parasites. CQR in P. falciparum is conferred by mutations in the parasite PfCRT, a putative transporter localized to the digestive vacuole (Fidock et al., 2000b; Das and Dash, 2007; Tinto et al., 2008). A single mutation, Lysine to Threonine at codon 76 (K76T), led to in vitro resistance in all progeny of a genetic cross between chloroquinesensitive and chloroquine-resistant parental clones and among a set of geographically diverse parasite isolates (Fidock et al., 2000a,b; Djimde et al., 2001; Sidhu et al., 2002). Several field studies have since confirmed the specificity of the Pfcrt K76T to clinical CQR (Wellems and Plowe, 2001). In a recent study from India, the Pfcrt K76T mutation has been shown to be rampant (96– 100%) in blood samples collected from patients infected with P. falciparum, especially in the northern and northeastern parts of India (Vinayak et al., 2003). Due to the widespread occurrence of chloroquine-resistant malaria parasites, there is an urgent need to monitor their spread and adopt proper drug regimen in the control strategy. In vivo studies remain the gold standard for monitoring antimalarial drug efficacy and are the primary source of information used by policy makers to shape recommendations for malaria chemotherapy and prophylaxis (Plowe et al., 2001; Plowe, 2003). In vitro methods for measuring drug resistance (Nguyen-Dinh and Payne, 1980) have proven to be more limited in scope and suitability for surveillance. However, in vitro methods are essential for characterizing resistance and confirming the molecular mechanisms of resistance. Easy and efficient molecular assays has been developed and effectively adopted for determining drug resistance in malaria parasites (Wernsdorfer and Noedl, 2003; Schonfeld et al., 2007; Marfurt et al., 2008). Molecular markers are used in many studies focusing on malaria drug resistance (Wellems and Plowe, 2001; Kublin et al., 2002). The majority of experimenters used nested polymerase chain reaction (PCR) followed by restriction fragment length polymorphism (RFLP), a relatively more rapid method than DNA sequencing, though still labor-intensive. Elsewhere, nestedmutation-specific PCR assays were used under controlled conditions in order to get reproducibility (Dorsey et al., 2001). One of the main advantages of molecular assays is in processing a large number of samples at a low cost in a short time. Therefore there is need to adopt molecular methods to identify drug-resistant malaria parasite in vectors and monitor its spread in malaria endemic zones in order to modify therapeutics practice. It has been established that the extent of drug resistance in the parasites is slowly spreading throughout the malaria endemic zones (Millet, 1995; Hallett et al., 2006). In Indian context, Directorate of National Vector Borne Disease Control Programme (NVBDCP, 2007) has recommended the use of artemisinin combination therapy (ACT) as the first line of drug for the treatment of P. falciparum in the areas of chloroquine resistance. For this purpose, identification of such areas carries utmost importance in malaria control strategy. Any sign of spread of chloroquine-resistant parasites can be first detected by analyzing the anopheline vectors transmitting malaria parasites. Thus vector evaluation for the presence of chloroquine-resistant parasite will give first hand information of the spread of drug resistance even long before any clinical manifestations from human host point of view. Many species complexes of Anopheles mosquitoes in Orissa, India transmit malaria, principally An. culicifacies, An. fluviatilis, An. annularis, and An. minimus (Christophers, 1933). An. culicifacies s.l. is a major vector of malaria in India accounting for 60–70% of malaria cases in the country, comprised of five sibling species provisionally designated as species A, B, C, D, and E (Green and Miles, 1980; Subbarao et al., 1983; Suguna et al., 1988; Vasantha et al., 1991; Kar et al., 1999). Species A, C, D, and E are identified as vectors of malaria and species B is a very poor vector or a nonvector (Subbarao et al., 1980, 1988a,b, 1992). An. fluviatilis James is

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a primary vector in the hilly and foothill regions of India and ranks second contributing to the total malaria cases of the country. An. fluviatilis exists as three distinct reproductively isolated cryptic species viz. S, T and U. Species S is primarily anthropophagic (90%) and is active in transmitting malaria whereas species T and U are mainly zoophagic (99%) and normally considered as nonvector (Nanda et al., 1996, 2000). An. annularis is found extensively in Orissa state (Venkat Rao, 1961; Dash et al., 1983), and found to be a complex of two reproductively isolated species, A and B (Atrie et al., 1999). Species A is a vector for malaria and highly anthropophagic where as the other species is not a vector. An. minimus s.l. is also a species complex comprising species A, C and E (Nicolescu et al., 2004). Almost all the vector types of the anophelines cryptic species mentioned above are present in Orissa and are responsible for high rates of malaria transmission. In the present study, Anopheles vectors were collected from different malaria endemic regions of Orissa, India, tested for the presence of chloroquine-resistant P. falciparum and Pfcrt haplotype K76T mutation. The target was to ascertain the transmission of chloroquine-resistant strains of P. falciparum parasite in the malaria endemic zones of Orissa. And thereafter to associate specific malaria vectors those are transmitting drug-resistant P. falciparum strains in the selected malaria endemic areas. This approach for the detection of spread of chloroquine-resistant strains in malaria endemic zones by malaria vector study can be very useful in modifying the therapeutics strategy well before any clinical manifestations of drug resistance. 2. Materials and methods 2.1. Mosquito collection and sampling In order to detect the chloroquine-resistant P. falciparum strains active in transmission in malaria endemic zones of Orissa, India, we have made an attempt to screen the principal malaria vectors in the region for the presence of resistant strains of parasite. Indoor resting female Anopheles mosquitoes were collected from different malaria endemic areas from the state of Orissa, India with the help of mechanical aspirators and CDC light trap. The study areas were selected from high-endemicity malaria to sporadic occurrence without any previous clinical reports of chloroquine resistance. Attempts were made to collect the malaria vector specimens based on their behavioral biting habits, i.e. adult female mosquitoes were collected from indoor and outdoor resting habits in early morning from 6.00 a.m. to 9.00 a.m. and evening from 6.00 p.m. to 10.00 p.m. Also resting mosquitoes were collected from 6.00 p.m. to 6.00 a.m. on hourly intervals. Mosquito collection was performed throughout the year with particular emphasis during the peak transmission seasons. Effort was taken to collect and process only the malaria vectors reported in the region. Mainly An. fluviatilis, An. culicifacies, An. annularis and An. minimus mosquitoes were considered for further analysis. Preliminary morphological identification was carried out soon after the catch, as per the standard key of Christophers (1933) and Nagpal et al. (2005). 2.2. Mosquito processing and DNA isolation For the rapid screening of vectors from the pool of non-vectors, we adopted a unique mosquito processing method as described by Mohanty et al. (2007). Immediately after identification, each individual mosquito was homogenized in two separate parts in two different 500 ml microcentrifuge tubes containing 30 ml phosphate-buffered saline (PBS), with the help of a micropestle. In one of the microcentrifuge tube only the head-thoracic region and in the other the rest of the body was homogenized. The homogenates were then dispensed on to properly coded spots on

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FTA Classic cards (Whatman, USA) separately. Half of the headthoracic region homogenate was dispensed to an area on the card coded as spot A and the rest part to another area coded spot B for the same mosquito. Then half of the homogenate of the rest body parts was dispensed to spot B and the rest part to another spot C. The punched discs obtained from the spot A on FTA cards were subjected to DNA isolation using the FTA Classic card protocol, followed by PCR and PCR-RFLP assays required for drug-resistance screening. Both spot B and spot C were useful for the regular vectorial attributes studies viz. blood meal analysis and species identification during routine surveillance.

Table 1 Mosquito collection areas from Orissa (India), with the numbers of different types of Anopheles species, malaria vectors in particular, collected. Area

An. culicifacies

An. annularis

An. minimus

Angul Phulbani Keonjhar Kalahandi

117 72 81 69

An. fluviatilis 84 71 33 48

26 32 21 9

9 11 3 6

Total

339

236

88

29

2.3. PCR for determination of P. falciparum-positive mosquitoes Using the designed, specific primers for the identification of P. falciparum as described by Mohanty et al. (2007), determination of P. falciparum-positive mosquitoes was carried out. The PCR reaction mixture contains; 1 PCR buffer containing 1.5 mM MgCl2, 200 mM each dNTP, 1.8 mM PF1 primer (50 -AGC GTG ATG AGA TTG AAG TCA G-30 ) and 1.8 mM PF2 primer (50 -CCC TAA ACC CTC TAA TCA TTG TC30 ) and 1 unit of Taq DNA polymerase (JumpStartTM Taq, Sigma, USA) in 30-ml reaction volume. DNA isolated from punched disc from spot A was taken as template for the reaction. The amplification was performed in a thermocycler (MJ Research, USA) under the following condition; initial denaturation at 95 8C for 4 min, followed by 35 cycles of denaturation at 95 8C for 30 s, annealing at 55 8C for 30 s and extension at 72 8C for 1 min; and a final extension at 72 8C for 8 min. All the PCR reactions were strictly accompanied by proper negative and positive controls. PCR amplified products (10 ml) were subjected to gel electrophoresis on a 2% agarose gel, stained with ethidium bromide, and visualized in UltraLum gel documentation system (USA). 2.4. Detection of Pfcrt mutations by restriction digestion of PCR products DNA extracted from the spot A was amplified by PCR using the primer pair PFCRT1 (50 -GGCTCACGTTTAGGTGGA-30 ) and PFCRT2 (50 -TGAATTTCCCTTTTTATTTCCAAA-30 ) to give a 264-bp product corresponding to amino acid residues 32–119 of the Pfcrt gene product. Following amplification of the fragments, polymorphisms in the Pfcrt gene were assessed by the mutation-specific restriction endonuclease digestion to detect single nucleotide polymorphisms (SNPs) in Pfcrt at positions K76T. For Pfcrt SNP detection, the PCR products were digested with ApoI, to determine the polymorphisms at codon 76. Digestions were done in 20 ml reactions containing 10 ml of PCR products according to the manufacturer’s instructions (New England Biolab, USA). All the assays were carried out with proper negative and positive controls. PCR product was incubated with 0.5 U of the restriction enzyme ApoI at 50 8C for 6 h. Digested products were electrophoresed on 1.5% agarose gels stained with ethidium bromide, and visualized under UV transillumination.

Fig. 1. Ethidium bromide-stained gel electrophoresis of PCR products of the Anopheles species, those are predominately transmit P. falciparum sporozoite. Lane 15 is the negative control without any DNA template. Lanes 1–3: An. fluviatilis species; lanes 4–6: An. annularis species; lanes 7–9: An. minimus species; lanes 10– 13: An. culicifacies species; lanes 3, 11 and 12 indicates the presence of P. falciparum sporozoite (205 bp) in the field collected specimens. Lane 14 is 100-bp DNA ladder (NEB).

PCR assay. The PCR amplified product of P. falciparum-specific band of 205 bp was shown in Fig. 1. 3.3. Pfcrt mutation detection by restriction digestion of PCR products Restriction enzyme mediated digestion of this PCR product with ApoI results in two fragments of 128 and 136 bp if the CQ-sensitive haplotype (CVMNK) is present. The K76T mutation renders the fragment resistant to digestion with ApoI (Fig. 2). Table 2 shows the details of the total number of P. falciparum-positive mosquitoes and K76T mutants. Of the total collected 339 specimens of An. culicifacies, 117 were from Angul district, of which 9 showed Pf positive including 3 individuals with K76T mutation. 72 An. culicifacies individuals were collected from Phulbani district, of which 6 showed Pf positive including 2 individuals with K76T mutation. 81 individuals were collected from Keonjhar district, of which 4 showed Pf positive including one specimen with K76T mutation. From Kalahandi district 69 An. culicifacies specimens were collected, of which 7 showed Pf positive including 1 individual with K76T mutation. Accordingly, of the total collected

3. Results 3.1. Mosquito collection and sampling A total of 339 An. culicifacies specimens, 236 An. fluviatilis specimens, 88 An. annularis specimens and 29 An. minimus specimens collected from four malaria endemic regions of Orissa. The details of the collected specimen are given in Table 1. 3.2. Determination of P. falciparum-positive mosquitoes by PCR To detect the P. falciparum DNA within the Anopheles mosquitoes, the collected samples were tested by PF-specific

Fig. 2. Ethidium bromide-stained gel electrophoresis of PCR products of the Pfcrt gene of P. falciparum present within the salivary glands of Anopheles species, those are predominately transmitting sporozoite. Lane 1 is 100 bp DNA ladder (NEB). Lane 2 is the positive control for chloroquine-sensitive parasite strain and lane 3 is the positive control for chloroquine-resistance parasite strain. Lanes 4–11 are the Pfcrt gene product of DNA taken from anophelines which, showed P. falciparum positive during the screening steps. All the sporozoite-positive samples yielded a 264 bp product. All the samples were subjected to ApoI digestion. Lanes 5, 7, 9 10, and 11 are showing digested product of 136 bp and 128 bp suggesting chloroquinesensitive strains. These two smaller products appeared together due to less resolution. While rest lanes (4, 6, and 8) are chloroquine-resistant strains.

A. Mohanty et al. / Infection, Genetics and Evolution 9 (2009) 462–467 Table 2 Number of anophelines collected from different areas from Orissa (India), showing the number of Plasmodium falciparum-positive individuals and K76T mutants. Area

An. culicifacies

An. fluviatilis

Angul Phulbani Keonjhar Kalahandi

117 [9pf, 3cq] 72 [6pf, 2cq] 81 [4pf, 1cq] 69 [7pf, 1cq]

84 71 33 48

Total

339 [26pf, 7cq]

236 [12pf, 3cq]

[6pf, [2pf, [1pf, [3pf,

2cq] 0cq] 0cq] 1cq]

An. annularis

An. minimus

26 [2pf, 0cq] 32 [2pf, 0cq] 21 [1pf, 0cq] 9 [0pf, 0cq]

9 [0pf, 0cq] 11 [1pf, 0cq] 3 [0pf, 0cq] 6 [1pf, 0cq]

88 [5pf, 0cq]

29 [2pf, 0cq]

pf

: the total number of P. falciparum-positive mosquitoes; cq: the total number of K76T mutant P. falciparum-positive mosquitoes out of the total P. falciparumpositive mosquitoes.

236 specimens of An. fluviatilis, 84 were from Angul district, of which 6 showed Pf positive including 2 individuals with K76T mutation. 71 An. fluviatilis individuals were collected from Phulbani district, of which 2 showed Pf positive without any K76T mutation. 33 individuals were collected from Keonjhar district, of which one showed Pf positive without K76T mutation. From Kalahandi district 48 An. fluviatilis specimens were collected, of which 3 showed Pf positive including 1 individual with K76T mutation. Among the An. annularis, of the total collected 88 specimens, 26 were from Angul district, of which 2 showed Pf positive without K76T mutation. 32 An. annularis individuals were collected from Phulbani district, of which 2 showed Pf positive without any K76T mutation. 21 individuals were collected from Keonjhar district, of which 1 showed Pf positive without K76T mutation. From Kalahandi district 9 An. annularis specimens were collected and all were negative for Pf. Of the total 29 An. minimus, 9 were collected from Angul district, 11 from Phulbani district, 3 from Keonjhar district, and 6 from Kalahandi district. Of which one each from Phulbani and Kalahandi district showed Pf positive without K76T mutation. 4. Discussion PCR-based assays were widely used to detect the presence of P. falciparum in mosquito, for convenience and in diagnostics where multiple parameters were evaluated simultaneously (Singh et al., 2004; Goswami et al., 2006; Alam et al., 2007). Moreover, PCR assays to detect K76T type P. falciparum parasites were very fast and effective for the evaluation. In the present study, we adopted this method to study anophelines as a means to evaluate CQR status of P. falciparum parasites in different malaria endemic zones of Orissa. As anopheline vectors transmit malaria parasites, any sign of spread of CQR parasites can be first detected by analyzing the vectors, that will give first hand information of the spread of drug resistance even long before onset of clinical manifestations in human host. The implementation of this approach will help in getting early warning of spreading of chloroquine-resistance strains of P. falciparum in an area without prior report of resistance in human host. In the present study, we selected four anophelines viz. An. culicifacies, An. fluviatilis, An. annularis and An. Minimus, prevalent vector species in Orissa. From the results it was evident that, of the total collection, 7.6% of An. culicifacies from Angul district and 8.3% from Phulbani district were Pf positive, among which 33.3% had K76T mutation, indicating high malaria sporozoite rates in comparison to previous reports (Parida et al., 1991). However, among 4.9% of Keonjhar district, and 10.1% of Kalahandi district Pf positive individuals, 25% and 14.3% had K76T mutation, respectively. From the results it is clear that, sporozoite, possessing K76T mutation, carrying rate among An. culicifacies is very high in all studied regions of Orissa. Similarly of the 7.1% An. fluviatilis from Angul district, and 6.2% from Kalahandi Pf positive, 33.3% had K76T; indicating An. fluviatilis, like An. culicifacies, also had high

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sporozoite carrying capacity in these regions of Orissa. In contrast, An. annularis and An. minimus collected from different regions, although were Pf positive but none had K76T mutation. Moreover, the results showed that, the sporozoite infectivity rates of An. annularis and An. minimus are not high as that of An. culicifacies and An. fluviatilis in studied regions of Orissa. We found that there was a little difference of malaria sporozoite positive rate and chloroquine resistance compared to earlier studies in Orissa, India (Satpathy et al., 1997; Sahu et al., 2008). The difference may be due to methodology adopted and nature of samples selected in this study. A major reason of variation of the previous work was due to focus on malaria patient either admitted to hospitals or in the zones of high transmission rate having chloroquine-resistance areas due to decade long drug administration. Further, the limited numbers of patients admitted to hospital, as reported by Sahu et al. (2008), might have received treatment or from the regions of already established chloroquine resistance due to greater malaria control measures by the local authorities using chloroquine as a first line treatment, resulting in high rates of CQR. Similar results reported by Pati et al. (2007) may be due to related facts. However, the present study intended to provide an early idea of CQR parasites transmission in a given area and can add-on the information gained from the conventional analysis of CQR status in an area by patient blood analysis. Moreover, the study can supplement the CQR epidemiological data and help malariacontrolling authorities in preparing proper strategy for combination drug therapy. The present work was carried out in the areas previously not reported to be CQR by clinical data and situated adjacent to active malaria transmission zones. Thus due to lack of chloroquine selection pressure the percentage of CQR parasite may be low in the total pool of parasites. In addition, as the study focuses on the methodology depending on the collection of malaria vectors and analyzing the parasites presents within, the results will also vary depending on the vector behavioral attributes, which is certainly a great factor influencing the malaria transmission and needs attention while studying the malaria transmission epidemiology of a region. The method used in this study can be employed along with routine vector survey of a region, the large number of specimens collected will not be considered as a waste, as the other parameters like human blood feeding and species type can be analyzed from the same catch. This need to be further elaborated, whether this high coincidence of Pf transmission with chloroquine-resistant strains by An. culicifacies and An. fluviatilis is due to vector abundance or biting habits along with aggressive drug control measures or due to some other reasons. More widespread study in the region is essential to map the CQR pattern in order to ascertain the distribution of the parasite resistance. In many malaria endemic regions, the therapeutic efficacy of CQ has decreased considerably. This, therefore, has led to the change in the first line drug for the treatment of malaria to artemisininbased combination, although, CQ is still widely used in the country for the treatment of non-complicated cases (NVBDCP, 2007). Constant observation of the existing parasite population concerning their genetic makeup determining the resistance to CQ became even more important since it was shown that after CQ withdrawal for therapy CQ-sensitive parasite re-occurred (Laufer et al., 2006). There are widespread cases and reports of mixed infections of P. falciparum parasites in malaria endemic regions (Anderson et al., 2000). However, the case of mixed infection of chloroquine-sensitive and -resistant parasites could not be discriminated from the pure infections by the present method of detection of CQR parasites transmitted within the malaria vectors. It is very much crucial to determine the fraction of each population in the mixed infection, as a factor affecting the drug test results, during the evaluation of parasite responses to anti-malarial drugs in the in vitro studies (Liu

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et al., 2008). In the context of the present study, the screening of potential anopheline vectors, which transmits the drug-resistant parasite in the malaria endemic zones, is informative in delineating the vectorial capacity as well as the sign of transmission of drugresistant strains. Once the transmission of resistant strains in an area is confirmed using the present method, further approaches for determining the accurate degree of mixed infection can be made to characterize the nature of drug resistance by adopting various novel tools, viz. PCR methods, microsatellite markers, pyrosequencing, and real-time qPCR (Su et al., 1999; Ferdig and Su, 2000). In long-term studies with wider coverage, the results of this type of study may provide a rationale for considering the withdrawal of chloroquine as a first line treatment of uncomplicated malaria. The rapid and inexpensive genomic assay that is now available to detect the chloroquine-resistant Pfcrt K76T mutation along with the approach adopted in the present study could expand the possibilities for monitoring resistance and use of drugs at hand with more pragmatic manner by modifying the therapeutics strategy well before any clinical manifestations of drug resistance come to notice. Acknowledgments The authors thank Dr. B. Ravindran, Director of the Institute of Life Sciences, Bhubaneswar, India, for his support. We are grateful to Mr. H.K. Tripathy for technical help and staffs at Regional Medical Research Centre, Bhubaneswar, India. The authors duly acknowledge the financial support provided by Department Biotechnology, and Indian Council of Medical Research, Government of India. Senior Research Fellowship provided by Council of Scientific and Industrial Research, Government of India, New Delhi to Amitav Mohanty is gratefully acknowledged. References Alam, M.T., Das, M.K., Dev, V., Ansari, M.A., Sharma, Y.D., 2007. Identification of two cryptic species in the Anopheles (Cellia) annularis complex using ribosomal DNA PCR-RFLP. Parasitol. Res. 100, 943–948. Anderson, T.J., Haubold, B., Williams, J.T., Estrada-Franco, J.G., Richardson, L., Mollinedo, R., Bockarie, M., Mokili, J., Mharakurwa, S., French, N., Whitworth, J., Velez, I.D., Brockman, A.H., Nosten, F., Ferreira, M.U., Day, K.P., 2000. Microsatellite markers reveal a spectrum of population structures in the malaria parasite Plasmodium falciparum. Mol. Biol. Evol. 17, 1467–1482. Atrie, B., Subbarao, S.K., Pillai, M.K.K., Rao, S.R.V., Sharma, V.P., 1999. Population cytogenetic evidence for sibling species within the taxon Anopheles annularis (Diptera: Culicidae). Ann. Entomol. Soc. Am. 92, 243–249. Bray, P.G., Mungthin, M., Ridley, R.G., Ward, S.A., 1998. Access to hematin: the basis of chloroquine resistance. Mol. Pharmacol. 54, 170–179. Christophers, S.R., 1933. The Fauna of British India. Dipter, Vol-IV. Family-Culicidae. Tribe-Anophelini. Today and Tomorrow’s Printers and Publishers, New Delhi. Das, A., Dash, A.P., 2007. Evolutionary paradigm of chloroquine-resistant malaria in India. Trends Parasitol. 23, 132–135. Dash, A.P., Bendley, M.S., Das, A.K., Das, M., Dwivedi, S.R., 1983. Roll of An. annularis as a vector of malaria in land of Orissa. J. Commun. Dis. 14, 224. Djimde, A., Doumbo, O.K., Cortese, J.F., Kayentao, K., Doumbo, S., Diourte, Y., Dicko, A., Su, X.Z., Nomura, T., Fidock, D.A., Wellems, T.E., Plowe, C.V., Coulibaly, D., 2001. A molecular marker for chloroquine-resistant falciparum malaria. N. Engl. J. Med. 344, 257–263. Dorsey, G., Kamya, M.R., Singh, A., Rosenthal, P.J., 2001. Polymorphisms in the Plasmodium falciparum pfcrt and pfmdr-1 genes and clinical response to chloroquine in Kampala, Uganda. J. Infect. Dis. 183, 1417–1420. Ferdig, M.T., Su, X.Z., 2000. Microsatellite markers and genetic mapping in Plasmodium falciparum. Parasitol. Today 16, 307–312. Fidock, D.A., Nomura, T., Cooper, R.A., Su, X., Talley, A.K., Wellems, T.E., 2000a. Allelic modifications of the cg2 and cg1 genes do not alter the chloroquine response of drug-resistant Plasmodium falciparum. Mol. Biochem. Parasitol. 110, 1–10. Fidock, D.A., Nomura, T., Talley, A.K., Cooper, R.A., Dzekunov, S.M., Ferdig, M.T., Ursos, L.M., Sidhu, A.B., Naude, B., Deitsch, K.W., Su, X.Z., Wootton, J.C., Roepe, P.D., Wellems, T.E., 2000b. Mutations in the P. falciparum digestive vacuole transmembrane protein PfCRT and evidence for their role in chloroquine resistance. Mol. Cell 6, 861–871. Fitch, C.D., 1970. Plasmodium falciparum in owl monkeys: drug resistance and chloroquine binding capacity. Science 169, 289–290. Foote, S.J., Thompson, J.K., Cowman, A.F., Kemp, D.J., 1989. Amplification of the multidrug resistance gene in some chloroquine-resistant isolates of P. falciparum. Cell 57, 921–930.

Goswami, G., Singh, O.P., Nanda, N., Raghavendra, K., Gakhar, S.K., Subbarao, S.K., 2006. Identification of all members of the anopheles culicifacies complex using allele-specific polymerase chain reaction assays. Am. J. Trop. Med. Hyg. 75, 454– 460. Green, C.A., Miles, S.J., 1980. Chromosomal evidence for sibling species of the malaria vector Anopheles (Cellia) culicifacies Giles. J. Trop. Med. Hyg. 83, 75–78. Greenberg, A.E., Ntumbanzondo, M., Ntula, N., Mawa, L., Howell, J., Davachi, F., 1989. Hospital-based surveillance of malaria-related paediatric morbidity and mortality in Kinshasa, Zaire. Bull. World Health Organ. 67, 189–196. Hallett, R.L., Dunyo, S., Ord, R., Jawara, M., Pinder, M., Randall, A., Alloueche, A., Walraven, G., Targett, G.A., Alexander, N., Sutherland, C.J., 2006. Chloroquine/ sulphadoxine-pyrimethamine for Gambian children with malaria: transmission to mosquitoes of multidrug-resistant Plasmodium falciparum. PLoS Clin. Trials 1, e15. Jelinek, T., Aida, A.O., Peyerl-Hoffmann, G., Jordan, S., Mayor, A., Heuschkel, C., el Valy, A.O., von Sonnenburg, F., Christophel, E.M., 2002. Diagnostic value of molecular markers in chloroquine-resistant falciparum malaria in Southern Mauritania. Am. J. Trop. Med. Hyg. 67, 449–453. Kar, I., Subbarao, S.K., Eapen, A., Ravindran, J., Satyanarayana, T.S., Raghavendra, K., Nanda, N., Sharma, V.P., 1999. Evidence for a new malaria vector species, species E, within the Anopheles culicifacies complex (Diptera: Culicidae). J. Med. Entomol. 36, 595–600. Keen, J., Farcas, G.A., Zhong, K., Yohanna, S., Dunne, M.W., Kain, K.C., 2007. Real-time PCR assay for rapid detection and analysis of PfCRT haplotypes of chloroquineresistant Plasmodium falciparum isolates from India. J. Clin. Microbiol. 45, 2889– 2893. Kublin, J.G., Dzinjalamala, F.K., Kamwendo, D.D., Malkin, E.M., Cortese, J.F., Martino, L.M., Mukadam, R.A., Rogerson, S.J., Lescano, A.G., Molyneux, M.E., Winstanley, P.A., Chimpeni, P., Taylor, T.E., Plowe, C.V., 2002. Molecular markers for failure of sulfadoxine–pyrimethamine and chlorproguanil–dapsone treatment of Plasmodium falciparum malaria. J. Infect. Dis. 185, 380–388. Laufer, M.K., Thesing, P.C., Eddington, N.D., Masonga, R., Dzinjalamala, F.K., Takala, S.L., Taylor, T.E., Plowe, C.V., 2006. Return of chloroquine antimalarial efficacy in Malawi. N. Engl. J. Med. 355, 1959–1966. Liu, S., Mu, J., Jiang, H., Su, X.Z., 2008. Effects of Plasmodium falciparum mixed infections on in vitro antimalarial drug tests and genotyping. Am. J. Trop. Med. Hyg. 79, 178–184. Marfurt, J., Muller, I., Sie, A., Oa, O., Reeder, J.C., Smith, T.A., Beck, H.P., Genton, B., 2008. The usefulness of twenty-four molecular markers in predicting treatment outcome with combination therapy of amodiaquine plus sulphadoxine–pyrimethamine against falciparum malaria in Papua New Guinea. Malar. J. 7, 61. Millet, P., 1995. Current status and prospects of malaria vaccines. J. Travel. Med. 2, 96–98. Mohanty, A., Kar, P., Mishra, K., Singh, D.V., Mohapatra, N., Kar, S.K., Dash, A.P., Hazra, R.K., 2007. Multiplex PCR assay for the detection of Anopheles fluviatilis species complex, human host preference, and Plasmodium falciparum sporozoite presence, using a unique mosquito processing method. Am. J. Trop. Med. Hyg. 76, 837–843. Nagpal, B.N., Srivastava, A., Saxena, R., Ansari, M.A., Dash, A.P., Das, S.C., 2005. Pictorial Identification Key For Indian Anophelines. National Institute of Malaria Research (Indian Council of Medical Research), New Delhi. Nanda, N., Joshi, H., Subbarao, S.K., Yadav, R.S., Shukla, R.P., Dua, V.K., Sharma, V.P., 1996. Anopheles fluviatilis complex: host feeding patterns of species S, T, and U. J. Am. Mosq. Control Assoc. 12, 147–149. Nanda, N., Yadav, R.S., Subbarao, S.K., Joshi, H., Sharma, V.P., 2000. Studies on Anopheles fluviatilis and Anopheles culicifacies sibling species in relation to malaria in forested hilly and deforested riverine ecosystems in northern Orissa, India. J. Am. Mosq. Control Assoc. 16, 199–205. Nguyen-Dinh, P., Payne, D., 1980. Pyrimethamine sensitivity in Plasmodium falciparum: determination in vitro by a modified 48-hour test. Bull. World Health Organ. 58, 909–912. Nicolescu, G., Linton, Y.M., Vladimirescu, A., Howard, T.M., Harbach, R.E., 2004. Mosquitoes of the Anopheles maculipennis group (Diptera: Culicidae) in Romania, with the discovery and formal recognition of a new species based on molecular and morphological evidence. Bull. Entomol. Res. 94, 525–535. NVBDCP, 2007. National Drug Policy on Malaria. In, Directorate of National Vector Borne Disease Control Programme (Directorate General of Health Services), Ministry of Health and Family Welfare, Government of India, Delhi. Parida, S.K., Gunasekaran, K., Sadanandane, C., Patra, K.P., Sahu, S.S., Jambulingam, P., 1991. Infection rate and vectorial capacity of malaria vectors in Jeypore hill tract. Indian J. Malariol. 28, 207–213. Pati, S.S., Mishra, S., Mohanty, S., Mohapatra, D.N., Sahu, P.K., Priyadarshi, N., Kumar, S., Sharma, S.K., Tyagi, P.K., Chitnis, C.E., Das, B.S., 2007. Pfcrt haplotypes and invivo chloroquine response in Sundergarh district, Orissa, India. Trans. R. Soc. Trop. Med. Hyg. 101, 650–654. Plowe, C.V., Doumbo, O.K., Djimde, A., Kayentao, K., Diourte, Y., Doumbo, S.N., Coulibaly, D., Thera, M., Wellems, T.E., Diallo, D.A., 2001. Chloroquine treatment of uncomplicated Plasmodium falciparum malaria in Mali: parasitologic resistance versus therapeutic efficacy. Am. J. Trop. Med. Hyg. 64, 242–246. Plowe, C.V., 2003. Monitoring antimalarial drug resistance: making the most of the tools at hand. J. Exp. Biol. 206, 3745–3752. Sahu, P.K., Pati, S.S., Satpathy, R., 2008. Association of msp-1, msp-2 and pfcrt genes with the severe complications of Plasmodium falciparum malaria in children. Ann. Trop. Med. Parasitol. 102, 377–382. Satpathy, S.K., Jena, R.C., Sharma, R.S., Sharma, R.C., 1997. Status of Plasmodium falciparum resistance to chloroquine in Orissa. J. Commun. Dis. 29, 145–151.

A. Mohanty et al. / Infection, Genetics and Evolution 9 (2009) 462–467 Schonfeld, M., Barreto Miranda, I., Schunk, M., Maduhu, I., Maboko, L., Hoelscher, M., Berens-Riha, N., Kitua, A., Loscher, T., 2007. Molecular surveillance of drugresistance associated mutations of Plasmodium falciparum in south-west Tanzania. Malar. J. 6, 2. Sidhu, A.B., Verdier-Pinard, D., Fidock, D.A., 2002. Chloroquine resistance in Plasmodium falciparum malaria parasites conferred by pfcrt mutations. Science 298, 210–213. Singh, O.P., Chandra, D., Nanda, N., Raghavendra, K., Sunil, S., Sharma, S.K., Dua, V.K., Subbarao, S.K., 2004. Differentiation of members of the Anopheles fluviatilis species complex by an allele-specific polymerase chain reaction based on 28S ribosomal DNA sequences. Am. J. Trop. Med. Hyg. 70, 27–32. Su, X., Ferdig, M.T., Huang, Y., Huynh, C.Q., Liu, A., You, J., Wootton, J.C., Wellems, T.E., 1999. A genetic map and recombination parameters of the human malaria parasite Plasmodium falciparum. Science 286, 1351–1353. Subbarao, S.K., Adak, T., Sharma, V.P., 1980. Anopheles culicifacies: sibling species distribution and vector incrimination studies. J. Commun. Dis. 12, 102–104. Subbarao, S.K., Vasantha, K., Adak, T., Sharma, V.P., 1983. Anopheles culicifacies complex: evidence for a new sibling species, species C. Ann. Entomol. Soc. Am. 76, 985–986. Subbarao, S.K., Adak, T., Vasantha, K., Joshi, H., Raghvendra, K., Cochrane, A.H., Nussenzweig, R.S., Sharma, V.P., 1988a. Susceptibility of Anopheles culicifacies species A and B to Plasmodium vivax and Plasmodium falciparum as determined by immunoradiometric assay. Trans. R. Soc. Trop. Med. Hyg. 82, 394–397. Subbarao, S.K., Vasantha, K., Raghavendra, K., Sharma, V.P., Sharma, G.K., 1988b. Anopheles culicifacies: siblings species composition and its relationship to malaria incidence. J. Am. Mosq. Control Assoc. 4, 29–33. Subbarao, S.K., Vasantha, K., Joshi, H., Raghavendra, K., Usha Devi, C., Sathyanarayan, T.S., Cochrane, A.H., Nussenzweig, R.S., Sharma, V.P., 1992. Role of Anopheles

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culicifacies sibling species in malaria transmission in Madhya Pradesh state, India. Trans. R. Soc. Trop. Med. Hyg. 86, 613–614. Suguna, S.G., Tewari, S.C., Mani, T.R., Hiriyan, J., Reuben, R., 1988. A cytogenetic description of a new species of the Anopheles culicifacies complex. Genetica 78, 225–230. Tinto, H., Guekoun, L., Zongo, I., Guiguemde, R.T., D’Alessandro, U., Ouedraogo, J.B., 2008. Chloroquine-resistance molecular markers (Pfcrt T76 and Pfmdr-1 Y86) and amodiaquine resistance in Burkina Faso. Trop. Med. Int. Health 13, 238– 240. Trape, J.F., Pison, G., Preziosi, M.P., Enel, C., Desgrees du Lou, A., Delaunay, V., Samb, B., Lagarde, E., Molez, J.F., Simondon, F., 1998. Impact of chloroquine resistance on malaria mortality. C. R. Acad. Sci. III 321, 689–697. Vasantha, K., Subbarao, S.K., Sharma, V.P., 1991. Anopheles culicifacies complex: population cytogenetic evidence for species D (Diptera: Culicidae). Ann. Entomol. Soc. Am. 84, 531–536. Venkat Rao, V., 1961. Vectors of malaria in India. Nat. Soc. Ind. Mal. Mosq. Dis., Delhi. Vinayak, S., Biswas, S., Dev, V., Kumar, A., Ansari, M.A., Sharma, Y.D., 2003. Prevalence of the K76T mutation in the pfcrt gene of Plasmodium falciparum among chloroquine responders in India. Acta Trop. 87, 287–293. Wellems, T.E., Plowe, C.V., 2001. Chloroquine-resistant malaria. J. Infect. Dis. 184, 770–776. Wernsdorfer, W.H., Noedl, H., 2003. Molecular markers for drug resistance in malaria: use in treatment, diagnosis and epidemiology. Curr. Opin. Infect. Dis. 16, 553–558. Zakeri, S., Afsharpad, M., Kazemzadeh, T., Mehdizadeh, K., Shabani, A., Djadid, N.D., 2008. Association of pfcrt but not pfmdr1 alleles with chloroquine resistance in Iranian isolates of Plasmodium falciparum. Am. J. Trop. Med. Hyg. 78, 633–640.