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. Author manuscript; available in PMC: 2012 May 1.
Published in final edited form as: Acta Trop. 2011 Feb 21;118(2):128–135. doi: 10.1016/j.actatropica.2011.02.004

Evaluation of a PCR-RFLP- ITS2 assay for discrimination of Anopheles species in northern and western Colombia

Astrid V Cienfuegos a, Doris A Rosero a, Nelson Naranjo a, Shirley Luckhart b, Jan E Conn c,d, Margarita M Correa a
PMCID: PMC3085589  NIHMSID: NIHMS277456  PMID: 21345325

Abstract

Anopheles mosquitoes are routinely identified using morphological characters of the female that often lead to misidentification due to interspecies similarity and intraspecies variability. The aim of this work was to evaluate the applicability of a previously developed PCR-RFLP-ITS2 assay for accurate discrimination of anophelines in twelve localities spanning three Colombian malaria epidemiological regions: Atlantic Coast, Pacific Coast, and Uraba-Bajo Cauca-Alto Sinu Region. The evaluation of the stability of the PCR-RFLP patterns is required since variability of the ITS2 has been documented and may produce discrepancies in the patterns previously reported. The assay was used to evaluate species assignation of 939 mosquitoes identified by morphology. Strong agreement between the morphological and molecular identification was found for species An. albimanus, An. aquasalis, An. darlingi and An. triannulatus s.l. (p ≥ 0.05, kappa=1). However, disagreement was found for species An. nuneztovari s.l., An. neomaculipalpus, An. apicimacula and An. punctimacula (p ≤ 0.05; kappa ranging from 0.33–0.80). The ITS2-PCR-RFLP assay proved valuable for discriminating anopheline species of northern and western Colombia, especially those with overlapping morphology in the Oswaldoi Group.

Keywords: Anopheles, PCR-RFLP, ITS2, species identification, malaria

1. INTRODUCTION

Colombia is among the 31 countries with the highest malaria burden in the world, and it has the second highest number of malaria cases in Latin America, with an average of 145,000 cases per year, although, it is estimated that more than 400,000 cases occur annually in the country (WHO, 2008, 2009). The social and economical problems caused by malaria have turned the control and elimination of this parasitic disease into one of the priorities of public health programs in several endemic countries (WHO, 1993, 2005a). The Global Malaria Program has claimed that vector control is one of the most efficient strategies for reducing disease burden (WHO, 1993, 2005b, 2005c). Successful malaria control, however, is critically dependent on the accurate identification of Anopheles species involved in parasite transmission, since targeted vector control strategies should be designed according to species specific aspects, such as ecology and behavior. For example, in Binh Thuan province of Vietnam, Anopheles varuna Iyengar was mistargeted as the malaria vector after being erroneously identified as An. minimus Theobald, another species recognized as a primary vector in southeastern Asia (Van Bortel et al., 2001). Using a PCR-RFLP assay, An. dirus s.l. (Peyton & Ramalingam, 1988) was correctly identified as the primary vector species in Binh Thuan province and subsequently targeted for control (Van Bortel et al., 2001; Van Bortel et al., 2000). Similar reports of morphological misidentification of potential vectors are found for Colombian and Peruvian anopheline species. Quiñones et al. (2001) and Fernandez et al. (2004) found that specimens of An. benarrochi s.l. Gabaldon were erroneously identified as An. evansae (Brèthes) in the Department of Putumayo in southern Colombia and in the region of Ucayali in Peru. Anopheles benarrochi s.l. is considered a zoophilic species in Brazil (Faran, 1980), but recent studies have shown it could be implicated in parasite transmission in other areas (Fernandez et al., 2004; Flores-Mendoza et al., 2004). Specimens of An. benarrochi s.l. collected in malaria endemic areas in eastern Peru were found infected with Plasmodium falciparum and Plasmodium vivax VK210 and VK247, suggesting that this species played a role in transmitting and maintaining Plasmodium species at least locally and that misidentification could have adverse public health impacts (Flores-Mendoza et al., 2004).

Species of the Oswaldoi Group of Anopheles subgenus Nyssorhynchus, such as An. benarrochi s.l., An. evansae, An. nuneztovari s.l. Gabaldón, An. rangeli Gabaldón, Cova Garcia & Lopez, An. oswaldoi (Peryassú) and An. aquasalis Curry (Harbach, 2004), are difficult to distinguish because of the well-known intraspecific variability and the substantial morphological overlap in some characters used for their discrimination (Calle et al., 2002; Fajardo et al., 2008; Faran and Linthicum, 1981). The morphologically similar species of the Oswaldoi Group differ in their contribution to parasite transmission in several regions of South America. Anopheles nuneztovari s.l. is a primary malaria vector in Colombia (Brochero et al., 2006; Gutierrez et al., 2008; Gutierrez et al., 2009; Olano et al., 2001), where it was recently found naturally infected with Plasmodium vivax (Gutierrez et al., 2009); but it has little importance in parasite transmission in the Brazilian Amazon where it has zoophilic behavior and other vectors have a primary role in parasite transmission (da Rocha et al., 2008; Povoa et al., 2006; Tadei and Dutary Thatcher, 2000). Anopheles aquasalis is a vector in Venezuela, Trinidad and Tobago, Brazil and Guyana (Berti et al., 1993; Chadee and Kitron, 1999; Laubach et al., 2001; Povoa et al., 2003), but until now its involvement in parasite transmission in Colombia has not been demonstrated, except for an epidemiological implication in a malaria focus in the Department of Guajira in 2000 (INS, 2001). Other species such as An. rangeli and An. oswaldoi s.l. have only recently been implicated in transmission in southern Colombia (Quiñones et al., 2006). Such differences in the vectorial roles of morphologically similar species make it critical to develop alternative identification strategies to complement the traditional use of morphological keys (Collins et al., 2000).

In Colombia, several studies have contributed to the design and standardization of molecular tools for Anopheles species discrimination. An assay was developed for the molecular identification of An. oswaldoi s.l. and An. benarrochi s.l. specimens from Putumayo, southern Colombia (Ruiz et al., 2005). Later, a PCR-RFLP for discrimination of seven anopheline species collected in Antioquia, northwestern Colombia was designed by Zapata et al. (2007). In this study, spacers ITS1 and ITS2 of the ITS region were evaluated, however only the ITS2 region was used in the assay since it proved to be more reproducible and reliable as a marker for species differentiation. The ITS2 PCR-RFLP patterns were better defined and more clearly distinguishable among species than those for the ITS1. Also, various studies have found greater levels of sequence and length variation in the ITS1 compared with the ITS2 (Fairley et al., 2005; Bower et al., 2008; 2009). The Zapata et al. (2007) assay was subsequently optimized to include other species of the Oswaldoi Group reported in the country (Cienfuegos et al., 2008). Considering the importance of correct species discrimination in malaria endemic regions of Colombia, the aim of the present work was to evaluate a PCR-RFLP-ITS2 strategy (Cienfuegos et al., 2008; Zapata et al., 2007) in 12 localities of northern and western Colombia, spanning three malaria epidemiological regions: the Atlantic and Pacific Coasts, and the Uraba-Bajo Cauca-Alto Sinu region. The assessment of the ITS2 stability in specimens of localities from different areas is required since variability of this region has been documented and is due to the presence of insertions, deletions, transitions and transversions which could result in changes of the patterns previously reported. However, the PCR-RFLP-ITS2 band patterns for the specimens evaluated herein were in agreement with the band patterns reported for specimens from San Pedro de Uraba, Antioquia, and with the An. aquasalis pattern confirmed by Cienfuegos et al. (2008), suggesting that the assay can be applied to identify anopheline mosquitoes collected in the three malaria epidemiological regions. The implementation of this assay in mosquito surveillance would provide a vital tool for malaria control programs.

2. MATERIALS AND METHODS

2.1. Collection sites

Specimens were collected in 12 localities spanning three Colombian epidemiological regions: Atlantic Coast (annual parasite index, API: 0.4), Pacific Coast (API: 10.8) and Uraba- Bajo Cauca-Alto Sinu (API: 25.5) (INS, 2006). The latter two regions present the highest risk of parasite transmission in Colombia (API ≥ 10: high risk) (PAHO, 2001). The localities sampled are listed in Table 1 and depicted in Figure 1. The closest distances among collections sites were 12 km between VAL and TAL, followed by MLI and PLT (27 km); the rest have distances > 50 km. Adult mosquitoes were collected by human landing catches under an informed consent agreement and procedures approved by the Bioethics Committee for Human Research at University Research Center (SIU) of the University of Antioquia. In addition, isofemale lines and reared larvae with associated exuviae were obtained to support the identification of collected Anopheles species. Details on collection procedures in this study are found in Gutierrez et al. (2008).

Table 1.

Collection data for specimens analyzed in this study.

Region Department / Locality Abbreviations Coordinate Latitude, Longitude* No. specimens analyzed Collection date
Atlantic Magdalena
 Los Achiotes ACH 11.2578–73.6144 28 2005–2006
Bolivar
 Santa Rosa de Lima SRL 10.435–73.3558 78 2005–2006
Urabá-Bajo Cauca-Alto Sinú Córdoba
 Moñitos MOÑ 9.2561–16.1092 128 2005–2007
 Puerto Libertador PLT 7.9064–75.6733 200 2007
 Tierra Alta TAL 8.1942–76.0786 49 2007
 Montelibano MLT 7.9853–75.4214 223 2007
 Valencia VAL 8.2617–76.1528 17 2007
Antioquia
 Zaragoza ZAR 7.4944–74.8678 50 2008
Pacific Chocó
 Pizarro PIZ 2.0511–78.6247 44 2005
 Nuquí NUQ 5.7111–77.2639 39 2005–2006
Valle del Cauca
 Buenaventura BUE 3.4292–77.0531 51 2005–2006
Nariño
 Tumaco TUM 1.8067–78.7717 41 2005–2006
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*

Coordinates are expressed in decimal degrees

Figure 1.

Figure 1

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Localities sampled for Anopheles mosquitoes (1 to 12). San Pedro de Urabá (SP) was the source of specimens for standardizing the AluI PCR-RFLP-ITS2 assay of Zapata et al., 2007.

2.2. Morphological identification of anopheline mosquitoes

Identification of wild-caught females and larvae collected in the field and reared in the laboratory was performed using available morphological keys (Faran, 1980; Faran and Linthicum, 1981; González and Carrejo, 2007; Rubio-Palis, 2000; Wilkerson and Strickman, 1993). Mosquitoes were dissected, and the abdomen, legs and wings of each mosquito were placed in vials containing 95% ethanol. In addition, one wing and one hindleg were mounted on a glass slide with Euparal and conserved as part of the Anopheles species collection of the Molecular Microbiology Group, University of Antioquia.

2.3. PCR-RFLP-ITS2 analysis

Mosquito legs or wings were used as the source of DNA for PCR reactions. When amplification from this material was not accomplished, dissected abdomens were used for DNA extraction (Birungi and Munstermann, 2002). The ITS2 primers used were from Beebe and Saul (1995). PCR conditions were as previously described (Zapata et al., 2007).

Previous bioinformatic analyses (Cienfuegos et al., 2008) indicated that a PCR-RFLP-ITS2 assay performed with AluI, FspI or DraIII is useful for identification of species belonging to the Oswaldoi Group. As such, amplified ITS2 products (50 to 100 ng/μl) were selected for the enzyme restriction analysis and digested according to manufacturer’s instructions (Promega, Madison, WI and New England Biolabs, Ipswich, MA, USA). Each restriction digest contained 1X buffer, 2 μg acetylated BSA, 1 unit AluI (Promega, Madison, WI, USA), 1μg ITS2 amplimer and sterile deionized water in a 20μl final volume. In addition to single restriction digests, double restriction digests with AluI/FspI were used to discriminate An. nuneztovari s.l. from An. benarrochi s.l. (Cienfuegos et al., 2008; Matson et al., 2008). These double digests were performed in a 20 μl reaction containing 20 mM Tris-acetate, 50 mM potassium acetate, 10 mM magnesium acetate, 1 mM dithiothreitol, 1 unit FspI and 1 unit AluI (New England Biolabs, Ipswich, MA, USA), and incubated at 37°C for 4 hours or overnight. Digested products were visualized as previously described (Zapata et al., 2007). Positive controls corresponding to the ITS2 digest of each species being analyzed were included in every gel. These controls were prepared from sequenced clones containing complete ITS1 and ITS2 regions from individuals of isofemale lines or immature stages. Anopheles aquasalis controls were clones from voucher specimens collected in Suriname. In order to avoid bias during the PCR-RFLP pattern analyses, the morphological identification and the molecular confirmations were carried out by two different persons and species assignations were confirmed only at the end of the PCR-RFLP process; in addition, the patterns were interpreted by two observers, one of them not involved in the processing of the samples.

2.4. Statistical analysis

The percentage of concordance between the morphological and molecular identifications was calculated by comparison of the number of specimens given the same or different morphological and molecular identification for each species. To test the null hypothesis of no real disagreement between molecular and morphological identification, a χ2 was calculated using a McNemar test and the strength of agreement was evaluated using a kappa index.

3. RESULTS

Nine hundred and thirty-nine anopheline specimens were analyzed and their morphological identifications were evaluated by PCR-RFLP-ITS2 (Table 2, Figure 2). A 100% concordance was found between morphological and molecular identification for species An. albimanus, An. aquasalis, An. darlingi, An. nuneztovari s.l., An. triannulatus s.l., An neomaculipalpus and An. apicimacula, only 30% for An. punctimacula, and no concordance was found for An. rangeli, An. oswaldoi s.l., An. evansae and An. strodei s.l. (Table 3). Results of the McNemar test for species An. albimanus, An. aquasalis, An. darlingi and An. triannulatus s.l., did not show disagreement between the identification methods (p ≥ 0.05), and the kappa statistic showed a strong agreement (kappa=1), between both methods. Disagreement between molecular and morphological identification was found for specimens An. nuneztovari s.l., An. neomaculipalpus, An. apicimacula and An. punctimacula (p ≤0.05). The strength of agreement was higher in An. neomaculipalpus (kappa=0.8), but lower in An. nuneztovari (kappa=0.36), An. apicimacula (kappa=0.33) and An. punctimacula (kappa=0.45). The McNemar and kappa tests could not be calculated for An. rangeli, An. oswaldoi s.l., An. evansae, An. strodei s.l. since none of the specimens analyzed were identified as such by the PCR-RFLP method (Table 3).

Table 2.

Morphological and molecular identification of Anopheles mosquitoes from localities of three Colombian malaria endemic regions.

Region Locality No. individuals collected per species Morphological Identification No. specimens analyzed by PCR-RFLP Molecular Identification
Atlantic Coast ACH 66 An. albimanus 17 An. albimanus
6 An. punctimacula 6 An. punctimacula
4 An. aquasalis 4 An. aquasalis
1 An. darlingi 1 An. darlingi
SRL 1583 An. triannulatus s.l. 35 An. triannulatus s.l.
945 An. albimanus 37 An. albimanus
3 An. punctimacula 3 An. neomaculipalpus
4 An. neomaculipalpus 3 An. neomaculipalpus
Urabá-Bajo Cauca-Alto Sinu MON 2099 An. albimanus 50 An. albimanus
113 An. triannulatus s.l. 30 An. triannulatus s.l.
40 An. neomaculipalpus 34 An. neomaculipalpus
15 An. punctimacula 14 An. neomaculipalpus
PLT 144 An. nuneztovari s.l. 117 An. nuneztovari s.l.
52 An. rangeli 43 An. nuneztovari s.l.
31 An. darlingi 14 An. darlingi
17 An. triannulatus s.l. 10 An. triannulatus s.l.
6 An. oswaldoi s.l. 6 An. nuneztovari s.l.
5 An. evansae 5 An. nuneztovari s.l.
4 An. punctimacula 3 An. punctimacula
2 An. strodei s.l. 2 An. nuneztovari s.l.
MLI 145 An. nuneztovari s.l. 15 An. nuneztovari s.l.
134 An. oswaldoi s.l. 119 An. nuneztovari s.l.
78 An. rangeli 70 An. nuneztovari s.l.
8 An. evansae 8 An. nuneztovari s.l.
8 An. strodei s.l. 8 An. nuneztovari s.l.
5 An. darlingi 3 An. darlingi
TAL 127 An. nuneztovari s.l. 11 An. nuneztovari s.l.
22 An. rangeli 20 An. nuneztovari s.l.
8 An. oswaldoi s.l. 8 An. nuneztovari s.l.
1 An. triannulatus s.l. 1 An. triannulatus s.l.
VAL 37 An. nuneztovari s.l. 10 An. nuneztovari s.l.
11 An. oswaldoi s.l. 4 An. nuneztovari s.l.
3 An. darlingi 3 An. darlingi
ZAR 58 An. triannulatus s.l. 33 An. triannulatus s.l.
32 An. nuneztovari s.l. 10 An. nuneztovari s.l.
9 An. darlingi 3 An. darlingi
3 An. nuneztovari s.l./rangeli 3 An. nuneztovari s.l.
1 An. nuneztovari s.l./aquasalis 1 An. nuneztovari s.l.
Pacific Coast NUQ 3099 An. albimanus 34 An. albimanus
5 An. punctimacula 4 An. apicimacula
1 An. apicimacula 1 An. apicimacula
BUE 1630 An. albimanus 45 An. albimanus
6 An. rangeli 6 An. nuneztovari s.l.
PIZ 47 An. albimanus 44 An. albimanus
TUM 1687 An. albimanus 41 An. albimanus
TOTAL 939
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In bold: specimens without concordance between morphological and molecular identification

Figure 2.

Figure 2

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PCR-RFLP-ITS2 analysis of anopheline mosquitoes of northern and western Colombia. 2.5% agarose gel. Lanes: M: Molecular weight marker; 1 and 2: An. albimanus; 3 and 4: An. nuneztovari s.l.; 5 and 6: An. aquasalis; 7 and 8: An. triannulatus s.l.; 9 and 10: An. punctimacula; 11 and 12: An. apicimacula; 13 and 14: An. neomaculipalpus; 15: negative control.

Table 3.

Concordance between morphological vs. molecular identification.

Species Identified by Morphology No. Specimens No. Specimens confirmed as each species by PCR-RFLP Molecular Identification % Concordance pb Kappa
An. albimanus 268 268 An. albimanus 100 1.0 1.0
An. aquasalis 4 4 An. aquasalis 100 1.0 1.0
An. darlingi 24 24 An. darlingi 100 1.0 1.0
An. nuneztovari s.l. 167 167 An. nuneztovari s.l. 100 0.0 0.36
An. triannulatus s.l. 109 109 An. triannulatus s.l. 100 1.0 1.0
An. neomaculipalpusa 37 37 An. neomaculipalpusa 100 0.0 0.80
An. apicimaculaa 1 1 An. apicimaculaa 100 0.12 0.33
An. punctimacula An. punctimacula (n=9)
30 9 An. neomaculipalpusa (n=17) 30 0.0 0.45
An. apicimaculaa (n=4)
An. rangeli 139 0 An. nuneztovari s.l. 0 NC NC
An. oswaldoi s.l. 137 0 An. nuneztovari s.l. 0 NC NC
An. evansae 13 0 An. nuneztovari s.l. 0 NC NC
An. strodei s.l. 10 0 An. nuneztovari s.l. 0 NC NC
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a

A different PCR-RFLP pattern was found for specimens identified by morphology as An. neomaculipalpus and An. apicimacula. The species specificity of these patterns should be confirmed by PCR-RFLP-ITS2 analyses of progeny-reared samples

b

p: significance level; the null hypothesis of agreement between the morphological and molecular identification methods is rejected when p ≤ 0.05. The statistics were calculated comparing the morphological and molecular identifications of each species

NC: Value not calculated since molecular identification for these species was not obtained

The PCR-RFLP-ITS2 allowed the identification of An. aquasalis (n=4), a species that was not included in the previous study (Zapata et al., 2007). The restriction pattern for An. aquasalis agreed with the one predicted in silico using An. aquasalis ITS2 sequences retrieved from GenBank (Cienfuegos et al., 2008) and with that from voucher material (Figure 3). In An. albimanus specimens processed in this study, a slight variation of 15 bp was observed in the banding pattern in specimens of all localities (Figure 4). Sequence analyses confirmed that this banding alteration likely resulted from a single point mutation in the recognition site of the AluI enzyme.

Figure 3.

Figure 3

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PCR-RFLP of ITS2 region following AluI restriction of amplimers from An. aquasalis on 2.5% agarose gel. Lanes: M: Molecular weight marker; 1–3: An. aquasalis individuals collected in Los Achiotes; 4: positive control, clone of An. aquasalis ITS2 region.

Figure 4.

Figure 4

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PCR-RFLP-ITS2 analysis of An. albimanus. 2.5% agarose gel. Lanes: M: Molecular weight marker; 1: positive control, clone of An. albimanus ITS2 region; 2–9: An. albimanus specimens collected in Los Achiotes.

Only specimens of An. punctimacula from ACH (n=6) and PLT (n=3) showed a PCR-RFLP pattern similar to the one described in San Pedro de Uraba (317 and 77 bp) (Zapata et al., 2007). However, specimens from SRL (n=3) and MON (n=14) that were morphologically consistent with An. punctimacula yielded RFLP results consistent with specimens identified as An. neomaculipalpus (95, 155 and 264 bp) and specimens from NUQ (n=4) yielded RFLP results consistent with specimens identified as An. apicimacula (372, 61 and 47 bp, deduced by bioinformatic analysis of sequences of individuals identified by morphology as such). Specimens from SRL and MON identified morphologically as An. neomaculipalpus were confirmed as such by their RFLP pattern (Table 2). One specimen from NUQ was identified by both morphology and RFLP pattern as An. apicimacula.

Specimens of the subgenus Nyssorhynchus of the Oswaldoi Group that were identified morphologically as An. evansae, An. rangeli, An. benarrochi s.l., and An. oswaldoi s.l. were confirmed by PCR-RFLP of ITS2 as An. nuneztovari s.l. (Table 2). Amplimers from specimens identified morphologically as An. nuneztovari s.l. (n=94) were double digested with AluI/FspI for discrimination between An. nuneztovari s.l. and An. benarrochi s.l. (Cienfuegos et al., 2008; Matson et al., 2008). All specimens [MLT (n=21), VAL (n=5), PLT (n=67)] were confirmed as An. nuneztovari s.l. according to band pattern expected for this species (296, 83, 57, 45 and 16 bp), which differs from that for An. benarrochi s.l. (341, 83, 60 and 16 bp) (Figure 5).

Figure 5.

Figure 5

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Double digest (AluI/FspI) of ITS2 region amplified from individuals identified by the PCR-RFLP with AluI as An. nuneztovari s.l. Lanes: M: Molecular weight marker; 1: positive control, clone of An. nuneztovari s.l. ITS2 sequence; 2–12: specimens collected in Puerto Libertador.

4. DISCUSSION

Numerous investigators have reported conflicting morphological identification of female mosquitoes of species of the Oswaldoi Group (Calle et al., 2002; Fajardo et al., 2008; Quiñones et al., 2001; Zapata et al., 2007). In particular, An. nuneztovari s.l. can be confused with An. rangeli, with An. benarrochi s.l. or An. aquasalis, and to some extent, with An. trinkae Faran, An. strodei s.l. Root or An. evansae (Fajardo et al., 2008). Although alternative approaches have been proposed for the accurate discrimination of these problematic Anopheles species, e.g., traditional and geometric morphometric analyses (Calle et al., 2008; Calle et al., 2002), these strategies are not entirely effective, nor especially practical, and therefore molecular identification assays should be evaluated from across the known geographic distribution of each species. In response to this issue, our PCR-RFLP-ITS2 strategy showed that species-specific band patterns were conserved throughout malaria-endemic localities in Uraba-Bajo Cauca-Alto Sinu and the Atlantic and Pacific coasts, and agreed with those obtained in specimens from San Pedro de Uraba (Zapata et al., 2007).

Among the species in the Oswoldoi Group, previous bioinformatic analysis showed that the AluI restriction pattern obtained in silico for An. benarrochi s.l. was similar to the one for An. nuneztovari s.l. (Cienfuegos et al., 2008; Matson et al., 2008). This occurs because a deletion of a G at position 349 of the An. benarrochi s.l. sequence AF462384 of a specimen from Brazil creates a new AluI recognition site (AGGCT→AGCT). Of relevance is that differences in behavior, morphology of male genitalia and ITS2 sequences suggest the existence of different species within An. benarrochi s.l. in Brazil, Venezuela, Colombia and Peru (Flores-Mendoza, 2004; Quiñones, 2006; Deane et al. 1948, Klein et al. 1991; Sallum et al. 2008). The An. benarrochi B from southern Colombia (Ruiz et al., 2005) and Peru are anthropophilic, and in some areas of Peru have been found naturally infected by Plasmodium spp. (Flores-Mendoza, 2004; Quiñones, 2006). Conversely, An. benarrochi s.l. from Brazil showed zoophilic behavior and was not susceptible to infection by Plasmodium (Deane et al., 1948; Klein et al., 1991). In addition, preliminary descriptions of two morphological Forms (I and II) based on differences in male genitalia of specimens from Acrelandia, Brazil, showed identical ITS2 sequences (Sallum et al., 2008). Both forms are similar in ITS2 sequences (97%) to An. benarrochi B from Colombia (GenBank accession nos. AY684984-AY684976), while ITS2 sequences from Rondonia, Brazil only showed 86% similarity (GenBank accession nos. AF462383 and AF462384), to sequences of An. benarrochi B (Sallum et al., 2008). Specimens identified morphologically in the present study as An. nuneztovari s.l. and double digested with AluI/FspI for their discrimination from An. benarrochi s.l. were confirmed as An. nuneztovari s.l. In addition, bioinformatic predictions of the ITS2 digest of An. benarrochi B from Colombia reveal an AluI PCR-RFLP pattern that is distinct from An. benarrochi s.l. from Brazil and from An. nuneztovari s.l., suggesting that in Colombia these species should not be confused when a digest with AluI is performed.

The small variation in the band pattern observed for An. albimanus was likely the result of a single transversion (G→C) in the recognition site of the AluI enzyme (AGCT, mutation: ACCT), rendering this site unavailable for restriction. Accordingly, two An. albimanus PCR-RFLP patterns can be defined: a wild type or “A” pattern of three bands (381, 128, and 15 bp) similar to that identified by Zapata et al. (2007) and a transversion or “B” pattern of two bands (381 and 143 bp). These patterns were consistent with those predicted in silico for the wild type and transversion sequences (GenBank accession nos. GU477267- GU477274). In spite of this variation, patterns A and B were suitable for the discrimination of An. albimanus (Figure 4). Further analysis of the wild type and transversion sequences will be necessary to test the extent of concerted evolution in An. albimanus populations.

Within the Arribalzagia Series of subgenus Anopheles, An. apicimacula, An. punctimacula, An. neomaculipalpus, and An. calderoni have presented additional challenges. In particular, An. apicimacula, An. punctimacula, An. neomaculipalpus share a subset of adult female morphological characters (González and Carrejo, 2007; Wilkerson and Strickman, 1993). However, our analyses revealed that specimens of these species were differentiated by their PCR-RFLP-ITS2 patterns. The species specificity of these patterns will be confirmed with PCR-RFLP-ITS2 analyses of progeny-reared samples. Among the other species in this series, An. calderoni can be confused with An. punctimacula (Rubio-Palis and Moreno, 2003; Wilkerson, 1991). Anopheles calderoni was recently reported from two localities in Antioquia State, which is located in the Uraba-Bajo Cauca-Alto Sinu region (González and Carrejo, 2009). The fact that An. punctimacula was collected in the present study in this same region (PLT, SLU) affirms the importance of evaluating mosquitoes at different life stages to definitively identify An. calderoni.

Based on our data, we suggest that the PCR-RFLP-ITS2 assay evaluated in this study is, at this time, the most accurate method for discrimination of anopheline species of Colombia. A similar assay that discriminates An. benarrochi s.l. and An. oswaldoi s.l. in southern Colombia was proposed previously (Ruiz et al., 2005). However, bioinformatic predictions using ITS2 sequences retrieved from GenBank, selected based on the correctness of species assignation according to analyses conducted by Marrelli et al. (2006), showed that each of the patterns identified by Ruiz et al. (2005) can be confused with other species in the Oswaldoi Group. For example, An. benarrochi s.l. (GenBank accession nos. AF462383 and AF462384 from field-collected specimens in Brazil) and An. nuneztovari s.l. (GenBank accession nos. AY028081-AY028126) would yield the same pattern under the conditions proposed by Ruiz et al. (2005). Likewise, An. oswaldoi s.l. (GenBank accession nos. AF055068-AF055070), An. aquasalis (GenBank accession nos. DQ020123-DQ020137) and An. rangeli (GenBank accession nos. AF462381, AF462382,Y09239) would yield similar patterns (Cienfuegos et al., 2008).

In conclusion, the results suggest that the AluI PCR-RFLP assay previously developed in San Pedro de Uraba, Colombia (Zapata et al., 2007), has applicability in different localities of Colombian malaria endemic regions. The assay was useful for discriminating species of the Oswaldoi Group and species within the Arribalzagia Series. In general, the results suggest less species diversity in the localities than previously reported, which among others, may be attributable to misidentification of the adults of species of the Oswaldoi Group that are frequently confused with An. nuneztovari s.l. This molecular tool is easy to perform, affordable and highly accurate for discriminating anopheline species, and therefore it is recommended that this or a similar molecular methodology be implemented for high throughput analysis and correct identification of Anopheles species in malaria endemic regions of Colombia. In addition, the fact that the ITS2 region is generally conserved and the patterns are reproducible in specimens collected from different geographical regions, suggest that it has the potential to be used as a nuclear barcoding marker for Anopheles species identification.

Acknowledgments

The authors are grateful to G.F. Gomez, L.M. Jaramillo, J.J. Gonzalez and M.I. Castro for technical assistance. The specimens of An. aquasalis from Suriname were collected under NIH grant AI-31034 to L.P. Lounibos. We thank L.P. Lounibos (Florida Medical Entomology Laboratory, USA), C. Limon and L. Resida (Bureau of Public Health, Suriname) for logistical assistance in collections. This study was supported by USA NIH R03AI076710, and partly funded by Comite para el Desarrollo de la Investigación- CODI, Universidad de Antioquia, Grant numbers 8700-1416 and 8700-039 to MCO.

Footnotes

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Contributor Information

Astrid V. Cienfuegos, Email: acienfuegos@udea.edu.co.

Doris A. Rosero, Email: roserodoris@hotmail.com.

Nelson Naranjo, Email: jezzid@hotmail.com.

Shirley Luckhart, Email: sluckhart@ucdavis.edu.

Jan E. Conn, Email: jconn@wadsworth.org.

References

  1. Beebe NW, Saul A. Discrimination of all members of the Anopheles punctulatus complex by polymerase chain reaction--restriction fragment length polymorphism analysis. Am J Trop Med Hyg. 1995;53:478–481. doi: 10.4269/ajtmh.1995.53.478. [DOI] [PubMed] [Google Scholar]
  2. Berti J, Zimmerman R, Amarista J. Adult abundance, biting behavior and parity of Anopheles aquasalis, Curry 1932 in two malarious areas of Sucre State, Venezuela. Mem Inst Oswaldo Cruz. 1993;88:363–369. doi: 10.1590/s0074-02761993000300004. [DOI] [PubMed] [Google Scholar]
  3. Birungi J, Munstermann L. Genetic structure of Aedes albopictus (Diptera: Culicidae) populations based on mitochondrial ND5 sequences: evidence for an independent invasion into Brazil and United States. Ann Entomol Soc Am. 2002;95:125–132. [Google Scholar]
  4. Bower JE, Cooper RD, Beebe NW. Internal repetition and intraindividual variation in the rDNA ITS1 of the Anopheles punctulatus group (Diptera: Culicidae): multiple units and rates of turnover. J Mol Evol. 2009;68:66–79. doi: 10.1007/s00239-008-9188-z. [DOI] [PMC free article] [PubMed] [Google Scholar]
  5. Bower JE, Dowton M, Cooper RD, Beebe NW. Intraspecific concerted evolution of the rDNA ITS1 in Anopheles farauti sensu stricto (Diptera: Culicidae) reveals recent patterns of population structure. J Mol Evol. 2008;67:397–411. doi: 10.1007/s00239-008-9161-x. [DOI] [PMC free article] [PubMed] [Google Scholar]
  6. Brochero H, Pareja PX, Ortiz G, Olano VA. Breeding places and biting activity of Anopheles species in the municipality of Cimitarra, Santander, Colombia. Biomedica. 2006;26:269–277. [PubMed] [Google Scholar]
  7. Calle DA, Quinones ML, Erazo HF, Jaramillo N. Differentiation by geometric morphometrics among 11 Anopheles (Nyssorhynchus) in Colombia. Biomedica. 2008;28:371–385. [PubMed] [Google Scholar]
  8. Calle LD, Quinones ML, Erazo HF, Jaramillo ON. Morphometric discrimination of females of five species of Anopheles of the subgenus Nyssorhynchus from Southern and Northwest Colombia. Mem Inst Oswaldo Cruz. 2002;97:1191–1195. doi: 10.1590/s0074-02762002000800021. [DOI] [PubMed] [Google Scholar]
  9. Cienfuegos AV, Gómez GF, Córdoba LA, Luckhart S, Conn JE, Correa MM. Diseño y evaluación de metodologías basadas en PCR-RFLP de ITS2 para la identificación molecular de mosquitos Anopheles spp. (Diptera: Culicidae) de la Costa Pacífica de Colombia. Rev Biomed. 2008;19:35–44. [Google Scholar]
  10. Collins FH, Kamau L, Ranson HA, Vulule JM. Molecular entomology and prospects for malaria control. Bull World Health Organ. 2000;78:1412–1423. [PMC free article] [PubMed] [Google Scholar]
  11. Chadee DD, Kitron U. Spatial and temporal patterns of imported malaria cases and local transmission in Trinidad. Am J Trop Med Hyg. 1999;61:513–517. doi: 10.4269/ajtmh.1999.61.513. [DOI] [PubMed] [Google Scholar]
  12. da Rocha JA, de Oliveira SB, Povoa MM, Moreira LA, Krettli AU. Malaria vectors in areas of Plasmodium falciparum epidemic transmission in the Amazon region, Brazil. Am J Trop Med Hyg. 2008;78:872–877. [PubMed] [Google Scholar]
  13. Deane LM, Causey OR, Deane MP. Notas sôbre a distribuição e a biologia anofelinos das regiões nordestina e amazônica do Brasil. Revista do Serviço Especial de Saúde Pública. 1948;1:827–965. [Google Scholar]
  14. Fajardo M, González R, Suárez MF, López D, Wilkerson R, Mureb-Sallum MA. Morphological analysis of three populations of Anopheles (Nyssorhynchus) nuneztovari Gabaldón (Diptera: Culicidae) from Colombia. Mem Inst Oswaldo Cruz. 2008;103:85–92. doi: 10.1590/s0074-02762008000100013. [DOI] [PubMed] [Google Scholar]
  15. Faran M. A revision of the Albimanus section of the subgenus Nyssorhynchus of the Anopheles. Contrib Am Entomol Inst. 1980;15:214p. [Google Scholar]
  16. Faran M, Linthicum L. A handbook of the Amazonian species of Anopheles (Nyssorhynchus) (Diptera: Culicidae) Mosq Sys. 1981;13:1–81. [Google Scholar]
  17. Fairley TL, Kilpatrick CW, Conn JE. Intragenomic heterogeneity of internal transcribed spacer rDNA in neotropical malaria vector Anopheles aquasalis (Diptera: Culicidae) J Med Entomol. 2005;42:795–800. doi: 10.1093/jmedent/42.5.795. [DOI] [PubMed] [Google Scholar]
  18. Fernandez RL, Schoeler GB, Stancil J. Presencia de Anopheles (Nyssorhynchus) benarrochi en áreas de selva con transmisión malárica. Rev Peru Med Exp Salud Publica. 2004;21:217–222. [Google Scholar]
  19. Flores-Mendoza C, Fernandez R, Escobedo-Vargas KS, Vela-Perez Q, Schoeler GB. Natural Plasmodium infections in Anopheles darlingi and Anopheles benarrochi (Diptera: Culicidae) from eastern Peru. J Med Entomol. 2004;41:489–494. doi: 10.1603/0022-2585-41.3.489. [DOI] [PubMed] [Google Scholar]
  20. González R, Carrejo N. Primera Edición. Claves y notas de distribución. Universidad del Valle; 2007. Introducción al estudio taxonómico de Anopheles de Colombia. [Google Scholar]
  21. González R, Carrejo N. Segunda Edición. Claves y notas de distribución. Universidad del Valle; 2009. Introducción al estudio taxonómico de Anopheles de Colombia. [Google Scholar]
  22. Gutierrez LA, Naranjo N, Jaramillo LM, Muskus C, Luckhart S, Conn JE, Correa MM. Natural infectivity of Anopheles species from the Pacific and Atlantic Regions of Colombia. Acta Trop. 2008;107:99–105. doi: 10.1016/j.actatropica.2008.04.019. [DOI] [PubMed] [Google Scholar]
  23. Gutierrez LA, Gonzalez JJ, Gomez GF, Castro MI, Rosero DA, Luckhart S, Conn JE, Correa MM. Species composition and natural infectivity of anthropophilic Anopheles (Diptera: Culicidae) in the states of Cordoba and Antioquia, Northwestern Colombia. Mem Inst Oswaldo Cruz. 2009;104:1117–1124. doi: 10.1590/s0074-02762009000800008. [DOI] [PMC free article] [PubMed] [Google Scholar]
  24. Harbach RE. The classification of genus Anopheles (Diptera: Culicidae): a working hypothesis of phylogenetic relationships. Bull Entomol Res. 2004;94:537–553. doi: 10.1079/ber2004321. [DOI] [PubMed] [Google Scholar]
  25. INS. Sistema de Vigilancia en Salud Pública - SIVIGILA. 14. Vol. 6. Instituto Nacional de Salud; 2001. Informe Quincenal Epidemiológico Nacional; pp. 205–220. [Google Scholar]
  26. INS. Sistema de Vigilancia en Salud Pública - SIVIGILA. 4. Vol. 11. Instituto Nacional de Salud; 2006. Informe Quincenal Epidemiológico Nacional; pp. 49–64. [Google Scholar]
  27. Klein TA, Lima JB, Tada MS. Comparative susceptibility of anopheline mosquitoes to Plasmodium falciparum in Rondonia, Brazil. Am J Trop Med Hyg. 1991;44:598–603. doi: 10.4269/ajtmh.1991.44.598. [DOI] [PubMed] [Google Scholar]
  28. Laubach HE, Validum L, Bonilla JA, Agar A, Cummings R, Mitchell C, Cuadrado RR, Palmer CJ. Identification of Anopheles aquasalis as a possible vector of malaria in Guyana, South America. West Indian Med J. 2001;50:319–321. [PubMed] [Google Scholar]
  29. Marrelli MT, Sallum MA, Marinotti O. The second internal transcribed spacer of nuclear ribosomal DNA as a tool for Latin American anopheline taxonomy - a critical review. Mem Inst Oswaldo Cruz. 2006;101:817–832. doi: 10.1590/s0074-02762006000800002. [DOI] [PubMed] [Google Scholar]
  30. Matson R, Tong Rios C, Banda Chavez C, Gilman R, Florin D, Lopez Sifuentes V, Cardenas Greffa R, Fernandez R, Yori P, Velasquez Portocarrero D, Vinetz J, Kosek M. Improved molecular technique for the differentiation of neotropical anopheline species. Am J Trop Med Hyg. 2008;78:492–498. [PMC free article] [PubMed] [Google Scholar]
  31. Olano V, Brochero H, Saenz R, Quiñones M, Molina J. Mapas preliminares de la distribución de especies de Anopheles vectores de malaria en Colombia. Biomédica. 2001;21:402–408. [Google Scholar]
  32. PAHO. Informe de la situación de los Programas Regionales de Malaria en las Américas. Pan American Health Organization; 2001. [Google Scholar]
  33. Povoa MM, Conn JE, Schlichting CD, Amaral JC, Segura MN, Da Silva AN, Dos Santos CC, Lacerda RN, De Souza RT, Galiza D, Santa Rosa EP, Wirtz RA. Malaria vectors, epidemiology, and the re-emergence of Anopheles darlingi in Belem, Para, Brazil. J Med Entomol. 2003;40:379–386. doi: 10.1603/0022-2585-40.4.379. [DOI] [PubMed] [Google Scholar]
  34. Povoa MM, de Souza RT, Lacerda RN, Rosa ES, Galiza D, de Souza JR, Wirtz RA, Schlichting CD, Conn JE. The importance of Anopheles albitarsis E and An. darlingi in human malaria transmission in Boa Vista, state of Roraima, Brazil. Mem Inst Oswaldo Cruz. 2006;101:163–168. doi: 10.1590/s0074-02762006000200008. [DOI] [PubMed] [Google Scholar]
  35. Quiñones ML, Harbach RE, Calle DA, Ruiz F, Erazo HF, Linton YM. Variante morfológica de adultos hembras de Anopheles benarrochi (Diptera: Culicidae) en Putumayo, Colombia. Biomedica. 2001;21:351–359. [Google Scholar]
  36. Quiñones ML, Ruiz F, Calle DA, Harbach RE, Erazo HF, Linton YM. Incrimination of Anopheles (Nyssorhynchus) rangeli and An. (Nys.) oswaldoi as natural vectors of Plasmodium vivax in Southern Colombia. Mem Inst Oswaldo Cruz. 2006;101:617–623. doi: 10.1590/s0074-02762006000600007. [DOI] [PubMed] [Google Scholar]
  37. Rubio-Palis Y. Taxonomía, Bionomía, Ecología e Importancia Médica. Escuela de Malariologia y Saneamiento Ambiental “DrAmoldo Gabaldon” y el Proyecto Control de Enfermedades Endémicas; 2000. Anopheles (Nyssorhynchus) de Venezuela; p. 138. [Google Scholar]
  38. Rubio-Palis Y, Moreno JE. Primer registro de Anopheles (Anopheles) calderoni (Diptera: Culicidae) en Venezuela. Entomotropica. 2003;18:159–161. [Google Scholar]
  39. Ruiz F, Quinones ML, Erazo HF, Calle DA, Alzate JF, Linton YM. Molecular differentiation of Anopheles (Nyssorhynchus) benarrochi and An. (N.) oswaldoi from southern Colombia. Mem Inst Oswaldo Cruz. 2005;100:155–160. doi: 10.1590/s0074-02762005000200008. [DOI] [PubMed] [Google Scholar]
  40. Sallum MA, Marrelli MT, Nagaki SS, Laporta GZ, Dos Santos CL. Insight into Anopheles (Nyssorhynchus) (Diptera: Culicidae) species from Brazil. J Med Entomol. 2008;45:970–981. doi: 10.1603/0022-2585(2008)45[970:iiandc]2.0.co;2. [DOI] [PubMed] [Google Scholar]
  41. Tadei WP, Dutary Thatcher B. Malaria vectors in the Brazilian amazon: Anopheles of the subgenus Nyssorhynchus. Rev Inst Med Trop Sao Paulo. 2000;42:87–94. doi: 10.1590/s0036-46652000000200005. [DOI] [PubMed] [Google Scholar]
  42. Van Bortel W, Harbach RE, Trung HD, Roelants P, Backeljau T, Coosemans M. Confirmation of Anopheles varuna in Vietnam, previously misidentified and mistargeted as the malaria vector Anopheles minimus. Am J Trop Med Hyg. 2001;65:729–732. doi: 10.4269/ajtmh.2001.65.729. [DOI] [PubMed] [Google Scholar]
  43. Van Bortel W, Trung HD, Roelants P, Harbach RE, Backeljau T, Coosemans M. Molecular identification of Anopheles minimus s.l. beyond distinguishing the members of the species complex. Insect Mol Biol. 2000;9:335–340. doi: 10.1046/j.1365-2583.2000.00192.x. [DOI] [PubMed] [Google Scholar]
  44. WHO. A global strategy for malaria control. World Health Organization; 1993. p. 44. [Google Scholar]
  45. WHO. Global Strategic Plan 2005–2015. Roll Back Malaria Partnership; Géneva: 2005a. p. 43. [Google Scholar]
  46. WHO. Roll Back Malaria. World Health Organization; 2005b. Malaria Control Today; p. 74. [Google Scholar]
  47. WHO. Malaria Vector Control and Personal Protection, WHO Technical Report Series. World Health Organization; 2005c. p. 62. [PubMed] [Google Scholar]
  48. WHO. World Malaria Report, WHO Library Cataloguing-in-Publication Data. World Health Organization; Géneva: 2008. p. 191. [Google Scholar]
  49. WHO. World Malaria Report. WHO Library Cataloguing-in-Publication Data; Geneva: 2009. p. 163. [Google Scholar]
  50. Wilkerson R. Anopheles (Anopheles) calderoni n.sp., a malaria vector of the Arribalzagia Series from Peru (Diptera: Culicidae) Mosq Sys. 1991;23:25–38. [Google Scholar]
  51. Wilkerson R, Strickman D. Clave Ilustrada para la Identificación de las Hembras de Mosquitos de México y Centroamérica. Secretaria de Salud, Subsecretaria de Coordinación y Desarrollo; México: 1993. [Google Scholar]
  52. Zapata MA, Cienfuegos AV, Quiros OI, Quinones ML, Luckhart S, Correa MM. Discrimination of seven Anopheles species from San Pedro de Uraba, Antioquia, Colombia, by polymerase chain reaction-restriction fragment length polymorphism analysis of ITS sequences. Am J Trop Med Hyg. 2007;77:67–72. [PubMed] [Google Scholar]

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