Advertisement
Browse Subject Areas
?

Click through the PLOS taxonomy to find articles in your field.

For more information about PLOS Subject Areas, click here.

  • Loading metrics

Updating the bionomy and geographical distribution of Anopheles (Nyssorhynchus) albitarsis F: A vector of malaria parasites in northern South America

  • Miguel A. Zúñiga,

    Roles Data curation, Formal analysis, Funding acquisition, Investigation, Methodology, Resources, Validation, Visualization, Writing – original draft, Writing – review & editing

    Affiliation Escuela de Microbiología, Facultad de Ciencias, Departamento Francisco Morazán, Universidad Nacional Autónoma de Honduras, Tegucigalpa, Honduras

  • Yasmin Rubio-Palis ,

    Roles Conceptualization, Data curation, Formal analysis, Investigation, Methodology, Project administration, Resources, Supervision, Validation, Visualization, Writing – original draft, Writing – review & editing

    rubiopalis@gmail.com

    Affiliations Departamento Clínico Integral, Facultad de Ciencias de la Salud, sede Aragua, Universidad de Carabobo, Maracay, Estado Aragua, Venezuela, Centro de Estudios de Enfermedades Endémicas y Salud Ambiental (CEEESA), Servicio Autónomo Instituto de Altos Estudios “Dr. Arnoldo Gabaldon”, Maracay, Estado Aragua, Venezuela

  • Helena Brochero

    Roles Conceptualization, Data curation, Formal analysis, Funding acquisition, Investigation, Methodology, Project administration, Resources, Supervision, Validation, Visualization, Writing – original draft, Writing – review & editing

    Affiliation Departamento de Agronomía, Facultad de Ciencias Agrarias, Bogotá, Universidad Nacional de Colombia, Bogotá, Distrito Capital, Colombia

Updating the bionomy and geographical distribution of Anopheles (Nyssorhynchus) albitarsis F: A vector of malaria parasites in northern South America

  • Miguel A. Zúñiga, 
  • Yasmin Rubio-Palis, 
  • Helena Brochero
PLOS
x

Abstract

Anopheles albitarsis F is a putative species belonging to the Albitarsis Complex, recognized by rDNA, mtDNA, partial white gene, and microsatellites sequences. It has been reported from the island of Trinidad, Venezuela and Colombia, and incriminated as a vector of malaria parasites in the latter. This study examined mitochondrially encoded cytochrome c oxidase I (MT-CO1) sequences of An. albitarsis F from malaria-endemic areas in Colombia and Venezuela to understand its relations with other members of the Complex, revised and update the geographical distribution and bionomics of An. albitarsis F and explore hypotheses to explain its phylogenetic relationships and geographical expansion. Forty-five MT-CO1 sequences obtained in this study were analyzed to estimate genetic diversity and possible evolutionary relationships. Sequences generated 37 haplotypes clustered in a group where the genetic divergence of Venezuelan populations did not exceed 1.6% with respect to Colombian samples. Anopheles albitarsis F (π = 0.013) represented the most recent cluster located closer to An. albitarsis I (π = 0.009). Barcode gap was detected according to Albitarsis Complex lineages previously reported (threshold 0.014–0.021). Anopheles albitarsis F has a wide distribution in northern South America and might play an important role in the transmission dynamics of malaria due to its high expansion capacity. Future studies are required to establish the southern distribution of An. albitarsis F in Venezuela, and its occurrence in Guyana and Ecuador.

Introduction

For over 40 years, the Anopheles (Nyssorhynchus) albitarsis complex has been considered a species complex based on chromosomal inversions [1, 2], allozymes [3, 4], random amplified polymorphic DNA-polymerase chain reaction (RAPD-PCR) [3, 5, 6] and, morphology and behavior [7, 8]. At present, the Albitarsis Complex includes five formally described species: An. albitarsis Lynch-Arribálzaga sensu stricto, An. deaneorum Rosa-Freitas, An. marajoara Galvão & Damasceno, An. oryzalimnetes Wilkerson & Motoki and An. janconnae Wilkerson & Sallum, and five putative species based on molecular studies: An. albitarsis F [9], An. albitarsis G [10, 11], An. albitarsis I [11, 12], An. albitarsis H [11] and An. albitarsis J [13]. Geographical distribution restriction has been found for some of these species [11, 14].

Anopheles albitarsis F was reported for the first time from Puerto Carreño, Vichada, Colombia [9] based on white gene, rDNA-ITS2 and consistently supported by microsatellite sequences [15, 16] and DNA mitochondrially encoded cytochrome c oxidase I (MT-CO1) sequences [11, 13], although morphologic diagnostic characters ranges for natural adult female populations have been reported [17, 18]. This species has been reported from coastal, piedmont, savannah, and interior lowland forest ecoregions [19, 20] in Colombia, the island of Trinidad and Venezuela [9, 11, 17, 21]. Natural populations of An. albitarsis F (as An. marajoara) from Colombia have been found naturally infected with Plasmodium falciparum and P. vivax [21, 22]. Anopheles albitarsis I, its sister species, occurs in northern Colombia and in sympatry with An. albitarsis F in Norte de Santander, Colombia and Zulia, north western Venezuela, near the border with Colombia [11, 12, 16, 23, 24]. It is considered that previous reports on geographic distribution, bionomy, and malaria vector incrimination of An. albitarsis sensu lato, An. marajoara and, An. allopha (nomen dubium) collected in Colombia, Trinidad and Tobago and Venezuela corresponded to either of these two lineages.

In Colombia, two genetic pools for these denominations were characterized based on microsatellite sequences [15, 16], one recognized now as An. albitarsis F, distributed east of the Eastern Andean Cordillera that corresponds to the cis-Andean genetic pool [9, 11, 21], and another, now recognized as An. albitarsis I [11, 24] corresponding to the trans-Andean genetic pool, distributed along the valleys between the Central and Eastern Andean Cordillera and lowlands of northern Colombia. Notably, Brochero et al. [16] found a higher number of hybrids between the cis and trans-gene pools as they were not completely isolated. It means that some foreign mosquitoes in a given population could be constituted of migrants with no genetic interchange with the original mosquitoes. The migration of individuals and the gene-flow levels detected were elevated, despite the fact that an important geographic barrier such as the Eastern Andean Cordillera separated the populations studied providing only partial differentiation of the two gene pools but addressing it. These genetic pools showed that their populations were remarkably similar, with divergence times of approximately 29 years when a mutation rate of 6.3x10-6 and twelve generations per year were estimated. These populations were characterized for having high historical effective numbers with a tendency of expansion in the absence of bottleneck events [15].

Rubio-Palis et al. [25], concluded that An. marajoara was the only species from the Albitarsis Complex present in Venezuela, based on morphometric analysis of 12 populations along the range of distribution of An. albitarsis s. l. in this country, and RAPD-PCR of one population from the center of the country. Based on the barcode region of mtDNA, sequences from Zulia and Cojedes States in western Venezuela and, a sequence from Portuguesa State, Venezuelan Llanos, assumed by Lehr et al. [26] to be An. janconnae (GenBank: DQ076234), were later confirmed as An. albitarsis F by Ruíz-López et al. [11]. Shortly after, Rubio-Palis et al. [17] reported An. albitarsis F for the first time from a malaria endemic area in Sucre Municipality, Bolívar State, south of the Orinoco River, which suggested a much wider range of distribution for this taxon in Venezuela. Additionally, An. albitarsis F has been confirmed from the island of Trinidad [11, 13] where previously Chadee and Wilkerson [27] had identified An. marajoara based on RAPD-PCR and ITS2-rDNA.

In the present study, we report an analysis of MT-CO1 sequences of An. albitarsis F samples collected in malaria-endemic areas of Colombia and Venezuela, a review of its bionomy and hypotheses of its origin and current geographical distribution.

Materials and methods

Study areas and mosquito collections

Specimens from Venezuela were collected in two localities: Calabozo, Guárico State (8°58’ N, 67°25’ W) in March 1999, preserved in 96% isopropanol, and San Rafael, Bolívar State (06° 47 N, 61 °34 W) in March 2014, preserved dry in silica gel. Collections were conducted by researchers and public health inspectors from the Ministry of Health involved in the Malaria Control Program who therefore did not need specific ethical clearance and no permits were required; permission for collecting in private properties was received from owners prior to collections. Calabozo is the capital of Francisco de Miranda Municipality; this area is characterized by large rice field plantations. Malaria was eradicated there in the early 1950’s and the incriminated vector was An. (Nys.) darlingi Root [28]. At present, malaria cases have been reported from the area, but no entomological information is available. San Rafael is a gold mining camp located in Sifontes Municipality, which in 2018 reported a total of 150,115 malaria cases [29]; incriminated vectors were An. darlingi, An. albitarsis s.l. (as An. marajoara) and An. (Anopheles) neomaculipalpus Curry based on their sporozoite and malaria entomological inoculation rates [30, 31]. Official records of malaria cases are usually sub-estimated due to self-medication, illegal migration from neighboring countries, and the limited diagnosis and treatment activities due to the economic crisis [32].

Specimens from Colombia were collected in three endemic malaria areas of the Departments of Meta, Guaviare and Vichada. Samples from Meta were collected in Puerto Gaitán (04°19′N, 72°05′W), where an urban P. falciparum malaria outbreak was reported during sampling [33]; specimens from Vichada were collected in peri urban areas of Puerto Carreño (06°11′16″N, 67°28′23″W), the capital of Vichada Department near the border with Venezuela; and, samples from Guaviare (02°34′43.8″N, 72°37′31.6″W) corresponded to peri urban localities where malaria transmission was enhanced by proliferation and intensification of the cultivation of illicit crops, internal armed conflict, intensive land use and immigration of people from hyperendemic malaria areas [34]. Detailed description of study sites and collection methods have been previously published [21, 34, 35]. Taxonomic identification was based on morphological characters [3638] and DNA MT-CO1 barcode sequences available in the GenBank and the Bold System public databases. Colombian samples were collected based on the protocols of the Malaria Vector Biology in Brazil project funded by The National Institute of Health (NIH, R01 AI5413), U.S., and the Universidad Nacional de Colombia, Quipu Code 201010012197. Collection of wild adult mosquitoes by human landing catches was conducted under an informed consent agreement using a protocol and collection procedures that were reviewed and approved by the Ethics Committee of the Faculty of Medicine of the Universidad Nacional de Colombia and by the Institutional Review Board of the New York State Department of Health, protocol No. 02–028. No additional permits were necessary.

DNA extraction and DNA MT-CO1 analysis

Total genomic DNA was extracted from three Venezuelan specimens, randomly chosen from each group of mosquitoes collected in each geographical locality: one from Calabozo and two from San Rafael. Additionally, gDNA of 17 mosquitos from Guaviare, 5 from Meta and 20 from Vichada, Colombia were obtained. DNA extractions were done by using the DNeasy blood and tissue kit (QIAGEN®, Hilden, Germany) but DNA was eluted in 30 μL of buffer AE. The DNA MT-CO1 gene sequence (corresponding to the DNA barcode region) was amplified using the universal primers [39] and PCR conditions corresponded to those described by Ruíz-López et al. [11]. All PCR reactions included negative controls. PCR products were subjected to bidirectional sequencing by Corporación Corpogen (Bogotá, Colombia, www.corpogen.org). Forward and reverse sequences were manually edited based on the chromatograms and consensus sequences were obtained by alignment using ClustalW [40] algorithm under Geneious v7.1.7 software (https://www.geneious.com). The MT-CO1 data sets amino acids translation did not show stop codons, meaning no pseudogenes were included in the analyses. Saturation levels between transition and transversion rates in relation to genetic distances were checked using the DAMBE v7.0 [41]. Sequence identification of species was confirmed by comparison to publicly available sequence data in the GenBank using the BLASTn algorithm [42] and using the BOLD Identification System (IDS) of the BOLD Systems database [43]. The intra and inter-specific genetic distances were calculated in MEGA v10.0 software [44] based on the Kimura-2-parameter (K2P) model [45].

Phylogenetic analyses based on barcode MT-CO1 sequences

Two analyses were done to estimate phylogenetic relationships. First, we analysed 45 sequences of An. albitarsis F obtained in this study and 123 publicly available sequences representative of the Albitarsis Complex members located in northern South America (N = 57 An. albitarsis F; N = 50 An. albitarsis I; N = 11 An. marajoara and N = 5 An. janconnae). Second, the analysis was done with 12 sequences from this study (N = 3 Venezuela; N = 9 Colombia) and 88 publicly available sequences representing all members of the Albitarsis Complex (N = 7 An. albitarsis s.s.; N = 9 An. oryzalimnetes; N = 11 An. marajoara; N = 5 An. deaneorum; N = 5 An. janconnae; N = 24 An. albitarsis F; N = 8 An. albitarsis G; N = 5 An. albitarsis H and, N = 14 for An. albitarsis I). The phylogenetic relationships were inferred using Neighbor Joining (NJ) and Bayesian inference (BI) methods. The NJ trees were constructed using the K2P distances model: branch support was assessed by bootstrapping with 1,000 replicates (Bootstrap values-BSV) [46, 47] using MEGA v10.0 software [44]. Bayesian inference (BI) analyses were performed in BEAST v1.10.4 [48] using the nucleotide substitution model HKY+I+G [49] previously selected with the Akaike Information Criterion (AIC) in the jModelTest2 program [50]. The Markov chain Monte Carlo (MCMC) algorithm was executed in duplicate runs of 10 million states each, sampling every 10,000 steps. The convergence of the MCMC chains was checked for effective sample size (ESS) values greater than 200 for each estimated parameter using Tracer v.1.7.1 [51]. The maximum credibility tree was determined using TreeAnnotator v1.10.4 after 25% burn-in and visualized with FigTree v1.4.3 [52]. Bayesian posterior probabilities (BPP) were used to assess nodal support. Anopheles (Nys.) braziliensis Chagas (GenBank: DQ076236) and An. darlingi (GenBank: DQ076238) were used as outgroup taxa.

Haplotypes were determined using DnaSP v6.12 [53]. Genetic diversity within populations was estimated by computing haplotype diversity (Hd) [47] and nucleotide diversity (π) [54]. Genealogical relationships were analysed through haplotype networks using PopART v1.7 [55] based on the TCS algorithm [56] to show relationships among the individuals sampled from different locations.

Analysis of published records of geographic distribution and bionomics of Anopheles albitarsis s.l., north of the Amazon River

Based on the revisions by Linthicum [36] and Sinka et al. [57] on the geographic distribution and bionomics of An. albitarsis s.l., a review protocol was established and agreed upon by all authors. The Guidelines from the Cochrane Handbook for Systematic Review was followed as standard methodology [58]. Online scientific bibliographic databases PubMed, LILACS, Web of Science and Scielo were searched using Anopheles albitarsis, An. marajoara and An. allopha as key words search terms followed by the Boolean operator “and” combined with one of each of the following ‘free text’ terms in succession: ‘entomological surveillance’, ‘larval collection’, ‘adult collection’, ‘resting collection’, ‘landing collection’, ‘vector density’, ‘geographical distribution’. The reference list of each of the included studies was also searched, and “grey literature” was sought by communication with authors for cited unpublished documents. The resulting citation library was then reviewed and refined, retaining all references that met one or more of the following criteria for inclusion: 1) The reported study was conducted in either of the following countries: Costa Rica, Panamá, Colombia, Venezuela, Trinidad and Tobago, Guyana, Suriname, French Guiana, and the Brazilian States of Amazonas, Roraima, Pará and Amapá, located north of the Amazon River; 2) the survey provided specific information on location to a precision of administrative unit level (municipality) or higher; 3) the surveys provided species-level information; 4) the surveys provided data on bionomics; 5) the surveys provided data on malaria vector incrimination. No limits were placed on year of publication and language. Globally, the literature search resulted in 1,837 publications or reports containing potential data to be reviewed. Of these publications, 621 were selected for full evaluation, of which 112 fulfilled the inclusion criteria.

The geographic distribution for Anopheles albitarsis s.l., An. albitarsis F, An. albitarsis I, An. marajoara and An. janconnae were plotted on a map using QGIS version 3.4 [59] based on images from Natural Earth raster (public domain: https://www.naturalearthdata.com) and data from published records extracted from the literature reviewed in this study.

Results

DNA extraction and DNA MT-CO1 analysis of Anopheles albitarsis F

Forty-five MT-CO1 sequences obtained from Venezuelan (n = 3) and Colombian (n = 42) specimens were identified as An. albitarsis F by high homology (99.20–100%) with sequences deposited in the GenBank (S1 Table) and the BOLD Identification System. The newly generated sequences were submitted to the GenBank database under the accession numbers [MW136004- MW136048].

The overall GC content was 32.0%; 43 variable polymorphic sites, and 20 parsimony informative sites were recorded. High MT-CO1 haplotype diversity (Hd = 0.990 ± 0.007 S.D.) and low nucleotide diversity (π = 0.012 ± 0.001 S.D.) were detected (Table 1). The overall mean genetic diversity (K2P) was 0.013 and the average genetic distance between populations from Colombia and Venezuela corresponded to 0.014. The highest genetic divergence (0.016) was found between localities from Venezuela (Guárico/Calabozo [GCA]; Bolívar/San Rafael/ [BSR]) and Colombia, Meta/Puerto Gaitán (MPG), while the lowest genetic divergence (0.012) occurred between the Colombian localities MPG and San José del Guaviare/Guaviare (SJG). From 37 haplotypes, closely related between Venezuelan localities and VPC-Colombia, 31 (83.78%) were singletons and 6 (16.22%) were shared, linked by single or few mutations (≤ 4) (S2 Table).

thumbnail
Table 1. Summary of MT-CO1 sequences datasets statistics.

https://doi.org/10.1371/journal.pone.0253230.t001

Phylogenetic analyses based on barcode MT-CO1 partial sequences

The dataset for the first analysis with An. albitarsis F, An. albitarsis I, An. marajoara and An. janconnae sequences from specimens collected in northern South America (N = 168) showed an overall GC of 32.2%, 89 variable polymorphic sites and 57 parsimony informative sites. The overall mean genetic distance (K2P) was 0.023, the average intra-specific genetic divergence was 0.009 (range: 0.006–0.013) and the average inter-specific divergence was 0.040 (range: 0.026–0.055). The most genetically divergent clusters were An. marajoara with An. janconnae (0.055), while lowest genetic divergence was found in An. albitarsis F with An. albitarsis I (0.026). For these taxa, a high MT-CO1 haplotype diversity (Hd = 0.985 ± 0.004 S.D.) and low nucleotide diversity (π = 0.020 ± 0.001 S.D.) were also detected (Table 1). A total of 106 haplotypes were calculated: 79 (74.53%) were singleton and 27 (25.47%) shared haplotypes linked by a single or a few mutations (≤ 6) (Fig 1, S3 Table). The largest haplotype of An. albitarsis I (H79) was represented with 17 sequences from the Caribbean region of Colombia while the haplotypes H1 to H73 corresponded to An. albitarsis F where H66 was the most widely distributed with sequences from Vichada/Colombia, Bolívar (Jabillal)/Venezuela and St. George/Trinidad. Colombia shared more haplotypes (H11, H13, H16, H29, H47, H52, H63 and H71) with Venezuela than with Trinidad (H3, H5). The highest number of mutational steps was found between An. albitarsis F with An. marajoara. The Neighbor Joining (NJ-K2P) and Bayesian inference (BI) trees revealed An. marajoara as the basal clade and An. albitarsis F as the most recent clade. However, NJ-K2P grouped a clade for three sister taxa (BSV = 80%): An. albitarsis F (BSV = 64%) closer to An. albitarsis I (BSV = 78%) and An. janconnae (BSV = 99%); meanwhile the BI tree showed sister taxa An. albitarsis F closer to An. albitarsis I but poorly supported (BPP = 0.56).

thumbnail
Fig 1. Genealogical analyses of Anopheles albitarsis complex species reported in the north of South America.

Haplotype network of 106 MT-CO1 haplotypes generated with TCS algorithm from 168 sequences dataset. Each taxa specimen and country are represented by a different color. The circle sizes indicate the frequency of individuals observed in each haplotype and, each notch on the links represents a mutated nucleotide position. The black solid circle indicates inferred missing intermediate steps between observed haplotypes (single nucleotide changes). mar: An. marajoara; jan: An. janconnae; albF: An. albitarsis F; albI: An. albitarsis I. CO: Colombia, VE: Venezuela, TT: Trinidad. *This analysis included all available sequences for An. albitarsis F and An. albitarsis I in the Genbank and the BOLD Systems database.

https://doi.org/10.1371/journal.pone.0253230.g001

The second analysis with sequences (N = 100) representing all members of the Albitarsis Complex generated a fragment length of 658bp, with a large region of overlap located at the positions 1462 to 2120 of the An. albitarsis s.s. mitochondrion complete genome (GenBank: HQ335344) excluding the primer regions. The average nucleotide composition percentages for this dataset were A = 29.3%, C = 15.8%, G = 16.3% and T = 38.6%; GC content = 31.2% with 92 variable polymorphic sites and 67 parsimony informative sites. The overall mean genetic distance (K2P) was 0.036, the average intra-specific genetic divergence was 0.010 (0.004–0.014) and the average inter-specific divergence corresponded to 0.040 (0.021–0.055). The most genetically divergent clusters were An. marajoara with An. janconnae (0.055) followed by An. marajoara with An. albitarsis F and An. albitarsis G with An. janconnae (identical values respectively 0.052). Most similar clusters were An. marajoara with An. albitarsis H (0.021). The Neighbor Joining (NJ-K2P) and Bayesian inference (BI) trees were represented with nine distinct clusters (Fig 2) and similar topologies. The NJ-K2P tree showed highly supported clades for all taxa in the Albitarsis Complex (BSV: 85–100%) and the BI tree showed robust probabilities (BPP: 0.95–1) for all terminal taxa. Two main clades were clearly identified; An. albitarsis F was located at the clade 1 with An. albitarsis I and An. janconnae. The MT-CO1 haplotype diversity (Hd = 0.991 ± 0.004 S.D.) and nucleotide diversity (π = 0.034 ± 0.001 S.D.) showed a similar pattern with the other analyses performed in this study (Table 1). The TCS network generated 75 haplotypes, 62 (82.67%) singletons and 13 (17.33%) were shared. Each of the clades identified in the phylogenetic trees was well-recognizable in the haplotypes network (Fig 3). Anopheles albitarsis F had the largest number of haplotypes (H1 –H34); An. albitarsis I (H35) and An. marajoara (H56) were the most shared haplotypes, including sequences from neighboring locations (S4 Table).

thumbnail
Fig 2. Phylogeny of Anopheles albitarsis complex in South America.

A. Bootstrapped NJ tree based on 1000 replicates of K2P data matrices from 100 sequences dataset. B. Bayesian inference tree based on posterior probabilities. Outgroup taxa include An. darlingi (GenBank: DQ076236) and An. braziliensis (GenBank: DQ076238). Color-coding represents the country of origin of MT-CO1 sequences taxa.

https://doi.org/10.1371/journal.pone.0253230.g002

thumbnail
Fig 3. Genealogical analyses for nine species of the Anopheles albitarsis complex in South America.

Haplotype network of 75 MT-CO1 haplotype generated with TCS algorithm from 100 sequences dataset. The circles represent individual haplotypes with size proportional to frequency and, each notch on the links represents a mutated nucleotide position. The black solid circle indicates inferred missing intermediate steps between observed haplotypes (single nucleotide changes).

https://doi.org/10.1371/journal.pone.0253230.g003

DNA barcodes showed not overlapping intra and interspecific divergences. This revealed a barcoding gap (threshold 0.014–0.021) between the largest intraspecific and smallest interspecific mean genetic distances (S1 Fig). The exhibiting interspecific divergence (≥2%) suggested distinct lineages within the Albitarsis Complex.

Published records of geographic distribution and bionomics of Anopheles albitarsis s.l., north of the Amazon River

Formal records of An. albitarsis F based on DNA MT-CO1 barcode, nDNA white gene, microsatellites and rDNA-ITS2 sequences showed a geographic distribution north of the Amazon river with reports from Colombia, the island of Trinidad, and Venezuela [9, 11, 13, 14, 16, 17, 21] (Fig 4).

thumbnail
Fig 4. Geographic distribution of Anopheles albitarsis complex species in the north of South America.

Note the distribution above the Amazon River of four species within the Anopheles albitarsis complex. (*) Species previously reported as An. albitarsis, An. marajoara or An. allopha were included as An. albitarsis s.l. The map was created in QGIS v 3.4 [59] using free vector and raster map data obtained from Natural Earth (public domain): naturalearthdata.com for illustrative purposes.

https://doi.org/10.1371/journal.pone.0253230.g004

In Colombia, An. albitarsis F (as An. marajoara, An. albitarsis s.l., An. allopha) corresponds to a gene pool associated with the Orinoquia Region (trans-Andean genetic pool) with reports from the Departments located east of the Andean Eastern Cordillera: Caquetá, Guaviare, Meta, Vichada [9, 15, 16, 21, 34], Putumayo [60], Casanare and Arauca [23] up to Norte de Santander near the border with Venezuela [16, 23]. Because local populations have been found naturally infected by P. falciparum [21, 22], and have been successfully infected by P. vivax in controlled conditions [61], this species has been considered a regional and opportunistic vector that maintains the transmission of malaria parasites throughout the year together with An. darlingi [15, 21]. Interestingly, Jimenez et al. [21] reported an Entomological Inoculation Rate (EIR) of 5.1 biting indoors and outdoors between 1800 and 1900 hrs in urban and peri urban neighborhoods of Puerto Carreño, Vichada. These authors also reported that this species is more active outdoors after sunset until 2100 hrs with a biting peak between 1800 and 1900 hrs, and a biting rate of 2.57 bites/person/hour [21]. The species has also been collected in urban areas of Villavicencio, Meta where it was more abundant outdoors with a biting rate of 11 bites/man/hour while indoors the biting rate was much lower (1.55); in rural areas the outdoor biting rate was 1.55 [62]. Anopheles albitarsis F (as An. marajoara) have been collected from a wide variety of larval habitats such as springs, streams, morichales (a flooded savannah area where the dominant tree is the palm Mauritia flexuosa or moriche) and lagoons; distinctly it has been also collected from man-made larval habitats exposed to sunlight such as fish ponds, wells and puddles [62].

Anopheles albitarsis I (as An. albitarsis s.l., An. marajoara) has been reported from the inter-Andean valleys of Colombia in the Departments of Antioquia and Huila [11, 15, 16, 23, 24, 63], and from the lowlands of the northern Departments of Bolívar, Córdoba and Magdalena [16, 23]; interestingly, in Norte de Santander Department, Colombia and Zulia State, Venezuela, this species occurs sympatrically with An. albitarsis F [11, 24]. This species has been collected at sunset in urban areas of El Banco, Magdalena and in Cáceres, Antioquia [15]. In general An. albitarsis I has been found in lowland dry and humid interior forests, strongly affected by human interventions as gold mining, cattle ranching, logging and small scale agriculture [16, 23, 6469]. Larvae have been collected in sunlit flooded pasture fields, fish ponds and ponds [63, 66]; adult abundance was low (< 1%) [67, 68] except in some localities of the Caucasia Municipality, Antioquia, where up to 38.5% of mosquitoes collected on human landing catches were identified as An. albitarsis s.l. [69], biting mainly outdoors before midnight [67, 68]. So far An. albitarsis I has not been found positive for Plasmodium spp.; in these regions of Colombia the incriminated vectors are An. darlingi, An. albimanus Wiedemann and An. nuneztovari Gabaldon [64, 6668].

In Venezuela, An. albitarsis F (as An. albitarsis s.l., An. marajoara) has a wide distribution; it has been reported from all the states at altitudes below 1000 m [17, 19, 25, 30, 31, 37, 7081]. Anopheles albitarsis F immature stages have been found in rice fields, streams, swamps, herbazales (flooded savannahs), river margins, lagoons in forests and in savannahs, and pools in riverbeds during the dry season; forest lagoons are mainly the result of abandoned gold mines [37, 72, 7476]. Larval habitats are usually located in sunlit open spaces, where the water color is amber and anopheline species are associated with grasses, aquatic vegetation such as green algae, Utricularia sp., Ludwigia sp., Eleocharis sp. and Mayaca sp. [74, 76]. This species was most abundant in lagoons and herbazales in August, three months after the peak of rains in gold mining areas of southern Venezuela [74]; adults showed no significant correlation with rainfall and were more abundant than An. darlingi all year round [73]. While in western Venezuela An. albitarsis F (as An. albitarsis s.l.) adults were significantly more abundant in August, one month after the peak of rains [79]. Longitudinal studies conducted in malaria endemic areas in western and southern Venezuela, as well as in the center of the country, the latter associated with rice fields, have shown a similar biting pattern along its range of distribution: it is active throughout the night indoors and outdoors and about 70% of bites occurred outdoors before midnight [37, 73, 79, 8284] with preference of feeding on humans relative to bovines [85]. Along its range of distribution in Venezuela, as well as other species of the subgenus Nyssorhynchus, this species does not rest inside houses, being collected early morning resting on vegetation around houses [79, 8385]. Anopheles albitarsis F (as An. albitarsis s.l.) has been found positive for malaria parasites in Venezuela in 1984 during an epidemic outbreak of P. vivax in Portuguesa State [37], where more recently Ruíz-López et al. [11] confirmed the presence of this lineage. This species is considered a secondary vector in western Venezuela where it was found positive for P. vivax-210 circumsporozoite protein [78], although during the dry season its vectorial capacity is similar to that of An. nuneztovari s.l., the principal malaria vector in the region [86]. In the malaria hotspot of southern Venezuela (Sifontes Municipality, Bolívar State), An. albitarsis F (as An. marajoara) has been found positive for circumsporozite protein of P. falciparum, P. vivax-247 and P. vivax-210 [30], with an overall sporozoite rate of 1.27 and entomological inoculation rate (EIR) of 1.25, while for An. darlingi the EIR was 2.21 [31]. Nevertheless, more recently [87] using nested PCR reported a marked increase of infection rates of anophelines in the hotspot of Sifontes Municipality where An. albitarsis F (as An. albitarsis s.l.) had higher infection rates than An. darlingi. Furthermore, the An. albitarsis F infection rate was 5.4%, followed by An. darlingi with 4.0 and An. nuneztovari s.l. with 0.5%. Plasmodium vivax accounted for 3.7% (61/1633) of infections in mosquitoes and 0.2% (4/1633) for P. falciparum. These authors concluded that the infection rate by Plasmodium spp. has increased more than three times in An. albitarsis F in ten years. Evaluations of susceptibility to insecticides conducted in 1994 and 2005 [88] have shown that populations from the Calabozo area, associated with rice crops, exhibit multiple resistance to the insecticides DDT, organophosphates and pyrethroids where the resistance mechanisms involved were increased level of nonspecific esterases, and evidence of insensitive acetylcholinesterase.

Anopheles albitarsis F (as An. marajoara) larvae have been collected in the island of Trinidad in rice fields, where they were more abundant during the rainy season [27]. Human landing catches between 1500 h and 2300 h showed a similar biting pattern indoors and outdoors with a peak of biting at 1900 hours [27, 89]. Exophilic resting behavior suggested that this species could play an important role in malaria transmission [89]. Nevertheless, the last outbreak of malaria in Trinidad and Tobago was reported in 1991 in villages located in the southwest tip of the island of Trinidad and the incriminated vector was An. (Nys.) aquasalis Curry [90]; until now An. albitarsis F has not been incriminated as a vector of malaria parasites in Trinidad and Tobago.

There are no recent published reports of An. albitarsis s.l. from Guyana. Giglioli [91] reported An. albitarsis from the savannah region of East Demerara and West Coast Berbice; Linthicum [36] examined specimens of An. marajoara collected by Aitken in 1962 in Georgetown, Berbice and Corentyne, and the most recent report is that of Rambajan [92] of An. albitarsis s.l. (as An. allopha) from coastal and interior parts of the country. The same seems to be the situation for Suriname; in fact, Hudson [93] reported An. albitarsis s.l. (as An. allopha) in New Nicherie associated with rice fields and Van der Kuyp [94] collected An. albitarsis in coastal areas. Apart from these reports An. albitarsis s.l. has not been collected in Suriname in over 30 years. However, in French Guiana, An. marajoara was confirmed by DNA MT-CO1 barcoding associated with an outbreak of P. vivax malaria in gold mining camps in forested areas [95, 96].

Linthicum [36] and Sinka et al. [57] revisions included reports of An. marajoara from Costa Rica and Panamá. There are no recent records from Costa Rica. The reports date from specimens collected by Komp in 1936, 1941 and 1942 [36]. Reports from Panamá are mainly from the Canal Zone by Baxter & Zetek [97], Arnett [98] and Blanton & Payton [99]. Loaiza et al. [100] provided an update on the distribution of anophelines from Panamá based on data collected between 1970 and 2005 by the personnel of the Ministry of Health, reporting An. albitarsis s.l. from the western coast of the Darien malaria endemic area; no DNA sequences are available to confirm the species within the Albitarsis Complex in Panamá.

The Albitarsis Complex has a wide distribution in Brazil and extends south to Bolivia, Paraguay and Argentina [11]; so far it has not been reported from Ecuador or Perú despite there being a record of An. albitarsis F from Putumayo Department in Colombia [60] on the border with Ecuador. In the states of northern Brazil, parts of which lie on the Guayana Shield, four species have been reported: An. marajoara, An. oryzalimnetes, An. janconnae and An. albitarsis G [11, 13]. Presently, the only species of the Albitarsis Complex identified in the State of Roraima, which borders with Venezuela, has been An. janconnae [11, 13, 101]. Collections of immature stages and adults around Boa Vista, have yield significantly large numbers of An. janconnae (as An. albitarsis E) compared to An. darlingi and they are incriminated in malaria transmission with lower infection rates than An. darlingi [102, 103]. McKeon et al. [101] studied larval habitats in the savannah ecoregion of Roraima and characterized An. janconnae as a specialist species in terms of the types of larval habitats and its abiotic and biotic characteristics required; this species showed a positive correlation with water flow and a negative relationship to sun exposure. Anopheles marajoara and An. oryzalimnetes were found in lowland forests in Pará States, north of the Amazon River; An. oryzalimnetes was a specialist species preferring saline larval habitats exposed to sun light, while An. marajoara was found in diverse larval habitats and could not be classified as specialist [101, 103105]. Although An. albitarsis G have been also collected in Pará State, south of the Amazon River [11], there is not available data on its bionomics and ecology. Also, in the State of Amapá, north of the Amazon River, An. janconnae and An. marajoara have been found sympatrically [11], but no data on their bionomics is available. Galardo et al. [106] reported that An. marajoara s.l. in Amapá was more abundant during the rainy season and positive for P. vivax; the species larval habitats were flooded forests and temporal or permanent open pastures. Nevertheless, it is not possible to determine to which species of the Complex this data is associated. So far, around Manaus, Amazonas State the only species confirmed is An. albitarsis G from forested areas [11, 107] but no information was provided in terms of abundance, biting rate, larval habitats and/or its medical importance.

Discussion

In the present study, we considered that previous reports on An. albitarsis s.l., An. allopha and An. marajoara from Colombia, Venezuela, and the island of Trinidad referred to either An. albitarsis F or An. albitarsis I based on the previous studies [9, 11, 13, 1517, 21, 23, 24]. Molecular analyses of DNA MT-CO1 barcodes revealed the presence of An. albitarsis F in endemic malaria regions of Colombia (MPG, SJG and VPC) and Venezuela (GCA, BSR). The genetic distance between Venezuelan and Colombian populations was 1.2% with Meta population (MPG) as the most divergent. In contrast, the haplotype analysis showed Venezuelan localities more closely related to VPC, Colombia, located in the Orinoquia Region, bordering Venezuela (S2 Table). In fact, Puerto Carreño (VPC) is located at the confluence of the Meta and Orinoco rivers on the border with Venezuela which suggests that previous reports of An. albitarsis s.l. (as An. marajoara) from Apure [25] and Amazonas States [77], with similar altitude, vegetation and larval habitats, are actually An. albitarsis F. The present results support the broader distribution and public health importance of this taxon in northern South America, as indicated in earlier studies [9, 11, 15, 16]. Furthermore, the confirmation in this study of An. albitarsis F from the center of the country in Venezuela and in the gold mining area of Bolívar State, together with previous reports from other localities [11, 17] suggest that An. albitarsis F is present throughout the country where An. albitarsis s.l./An. marajoara has been reported from all the states and all ecoregions [19, 25, 7088, 108]. Also, our findings and other reports [11, 15, 16, 21, 34, 60] suggest that An. albitarsis F is the only member of the Albitarsis Complex present in Colombia, east of the Eastern Andean Cordillera (trans-Andean genetic pool). Anopheles albitarsis F is the only species of the Albitarsis Complex present in the island of Trinidad [11] and it has not been reported from Ecuador, Perú and the island of Tobago; nevertheless, the confirmation of An. albitarsis F from Puerto Asís, Putumayo Department [60] on the Colombian border with Ecuador, suggests that this species might be also present in Ecuador since this is an active border with significant moving human population and similar ecology on both sides of the border. Also, this species shows rapid adaptation and expansion [14]. The absence of An. albitarsis F from the island of Tobago, located 35 km northeast of the island of Trinidad, might be due to its different geological origin, which resulted as in the rest of the Lesser Antilles, from the subduction of the South American Plate under the Caribbean Plate during the Mid-Eocene and Oligocene becoming a separate biogeographic region [109, 110]. As previously described in Results, this species is a generalist, exploiting different types of oviposition sites usually exposed to sunlight and associated with highly modify environments by human activities such as cattle ranching, agriculture and mining. It is important to point out that An. albitarsis F has not been collected in more pristine environments in Venezuela such as the Amerindian Yanomami territory of Amazonas State [111, 112] and the Amerindian Ye’kwana-Sanema territory along the Caura/Erebato rivers, Bolívar State [113], while it has been collected in Amazonas State around villages along the Orinoco river on the border with Colombia [77] and around criollo villages in the lower Caura river, Bolívar State [17, 76, 80, 82, 113]. Within its range of distribution, An. albitarsis F bites mainly outdoors before midnight [21, 62, 73, 79, 82, 83] and shows exophilic resting behavior [79, 85], characteristics that offer challenges for vector control since the traditional methods using long lasting treated nets (LLINS) and indoor residual spraying (IRS) are ineffective. Anopheles albitarsis F has been confirmed as a vector of malaria parasites in Colombia [21, 22] and Venezuela [30, 31, 78, 87]. Over the past 30 years the anarchic extensive deforestation and intensive human mobilization in Bolívar State, and specifically in the malaria hotspot of Sifontes municipality, southern Venezuela, has resulted in a change of species composition and abundance of anophelines. In fact, studies conducted in Sifontes between 1992–1997 showed that in general anopheline abundance was very low and the most frequent species collected was An. darlingi [114]; progressively it has been shown how species composition changed and abundance increased [7275, 84]. Furthermore, a dramatic increase in infection rates of anophelines has been reported, where An. albitarsis F showed a higher infection than that of An. darlingi, until then the principal vector in the region. In fact, An. albitarsis F (as An. albitarsis s.l.) has more than tripled its human Plasmodia infection rates in ten years [31, 87]. This phenomenon supports the prediction models proposed for the year 2070 by Laporta et al. [115] where climatic and landscape effects might lead to shifts in the importance of roles and distribution of species of the Albitarsis Complex over An. darlingi for transmission caused by global warming. Similar observations were reported from northern Brazil for An. janconnae (as An. albitarsis s.l.). This species was significantly more abundant than An. darlingi in urban areas of Boa Vista, Roraima State, and was an important vector with higher EIR despite An. darlingi showed a higher infection rate [102, 103]. Meanwhile in Amapá State, Galardo et al. [106] reported higher sporozoite rates for An. marajoara s.l. in relation to An. darlingi; nevertheless, in this State An. janconnae and An. marajoara have been found sympatrically [11], and therefore it is not possible to determine to which species of the Complex these data are associated. Anopheles marajoara have been confirmed from some malaria foci in southern French Guiana [95] and incriminated as an important vector during an outbreak with a P. vivax infection rate of 6.4% while An. darlingi showed an infection rate of 1.1% for P. falciparum [96]. Intensive entomological surveys in Suriname have not reported the presence of An. albitarsis s.l. for over 30 years; a similar situation is found in Guyana, although entomological surveillance has not been as intense as in Suriname. Due to the shared border between Guyana (Essequibo) and Venezuela (Bolívar State), similar epidemiological and ecological conditions, associated with gold mining and dynamic movements of populations across the border, it is likely that An. albitarsis F is present and might be involved in the transmission of malaria parasites in Guyana.

In this study, phylogenetic relationship and genetic variation of An. albitarsis F, An. albitarsis I, An. janconnae and An. marajoara confirmed these taxa as separated lineages; based on interspecific divergences greater than 2% and distinct haplotype clusters for each species represented in a TCS network linked by multiple mutational steps (Fig 1). In terms of species differentiation, the haplotype network demonstrated that An. albitarsis F, An. albitarsis I and An. janconnae had relatively little divergence; An. albitarsis I corresponded to sequences from the Caribbean region of Colombia (GenBank: GQ153597—GQ153610), with a single record from Venezuela (Río Socuavo/Zulia, GenBank: JQ615189) that was in a shared haplotype with two records from Colombia (Tibú/Norte de Santander, GenBank: JQ615190; JQ615197). The haplotype (H79) was the largest cluster of sequences (N = 17) for the An. albitarsis complex phylogenetic analysis (Fig 1, S3 Table). A similar pattern occurred in both localities for the sister taxon An. albitarsis F that has been detected in sympatry and with low level of divergence [11, 13]. The high haplotype diversity and broad distribution encountered among haplotypes for An. albitarsis F in the present study and previously [11, 13] and the most widely distributed shared haplotype with sequences from Colombia, the island of Trinidad, and Venezuela could be explained by the correspondence of geographical proximity for the species and its sister clusters, either sympatric or overlapping with little ecological divergence [14]. However, further sampling is required to clarify these relationships.

Anopheles albitarsis I have been reported only from northwestern Venezuela and northeastern Colombia and extending south along the inter-Andean valleys between the Central and Eastern Andean Cordilleras (trans-Andean genetic pool) [11, 23]. This suggested that Colombian Andean Central and Western mountain ranges might limit the geographic distribution of An. albitarsis I to the west and south of the country. Anopheles marajoara (the basal species from which An. albitarsis F originated) and other members of the Albitarsis Complex have not been reported from Perú [11, 14]. It is proposed that An. albitarsis I is a lineage derived from An. albitarsis F. Apparently, An. albitarsis I is found in Venezuela only in sympatry with An. albitarsis F in Zulia State, north-eastern Venezuela near the border with Colombia. A similar situation was found in Norte de Santander Department [11]. Since An. albitarsis F and An. albitarsis I are in sympatry in the northern zone of the border between Colombia and Venezuela, this might be a zone of hybridization for the two lineages or a recent hotspot of speciation for them [15, 16]. There are no reports of An. albitarsis I from Trinidad and Tobago or elsewhere. Although An. albitarsis s.l. has been reported from Panamá [100], there are no sequences available; taking into account the current geographical distribution proposed for the clade of An. albitarsis F, An. janconnae and An. albitarsis I, it is suggested that An. albitarsis s.l. from Panamá could correspond to An. albitarsis I, a lineage more adapted to high temperatures, high relative humidity, and with more plasticity to occupy a wide range of larval habitats [16, 23, 6466, 68, 69]. It is important to point out that so far, An. albitarsis I has not been found positive for Plasmodium spp. from either Colombia or Panamá, probably due to its low biting rate in relation to the incriminated vectors An. albimanus, An. darlingi and An. nuneztovari s.l. [64, 66].

Foley et al. [14] explored the possibility of finding a phylogenetic signal to explain ecological and environmental divergence for the proposed phylogenetic relationships of the Albitarsis Complex [11] which incorporated climatic and ecological data from the geographic locations of the sequencies included. Their results showed the Clade 1 comprised of An. albitarsis F, An. albitarsis I, and An. janconnae, as in the present study, and Clade 2 (all other species) were separated by the Amazon River. Events of dispersion and colonization of new habitats by these species might be determined by ecological requirements for each taxon, together with processes of landscape fragmentation, climatic changes and natural barriers that contributed to their recent speciation [14]. Furthermore, using MT-CO1 sequences from the present study and previous records for all the members of the Albitarsis Complex, the monophyly of the Albitarsis Complex was strongly supported by Bayesian analysis (BPP: 0.95–1) and nine distinct clusters of the Albitarsis Complex were recovered as in previous studies [11, 13]. We have found a well-supported clade of sister taxa with An. albitarsis F, An. janconnae, and An. albitarsis I, all geographically located north of the Amazon River. These species correspond to separated lineages with interspecific divergences greater than 2% and distinct haplotypes clusters [11, 13]. As previously described, these species have distinctive geographic distribution, ecology, bionomy and vector characteristics [14]. In this sense, geological studies based on radar image interpretation, sedimentology and radio carbon dating indicated that the Amazon River originated during the Plio-Pleistocene with significant landscape changes that resulted in speciation [116]. The Amazon River determines the separation of Clade 1 and Clade 2 of the Albitarsis Complex [11, 14]; it has been proposed that it represents a major barrier for gene flow [117, 118]. However, Santorelli et al. [119] have rejected the river barrier hypothesis based on species distribution of different taxonomic groups around the Madeira River, Brazil, suggesting that the river had functioned as a vicariance barrier only for a low percentage of the total species identified (<1%). These contrasting hypotheses suggest that novel analyses are necessary to explain species distribution around major rivers. In addition, further studies are required to elucidate the evolutionary and taxonomic status to understand speciation events and phylogenetic relationships between closely related vector and non-vector species, which would allow the targeting of important disease vector species groups and the development of novel genetic-based vector and pathogen control methods.

It has been estimated that the genus Anopheles originated in Western Gondwana [120] in what is now South America during the early Cretaceous period around 145 to 100 million years ago [121, 122]. For the Albitarsis Complex, a monophyletic group, it has been estimated that it diverged approximately 39 MYA from its ancestor An. darlingi [123], following a latitudinal migration and subsequent diversification [121, 124] and within the group the time divergence was estimated at 0.58–2.25 MYA [13]. In accordance with the current natural geographical distribution registered for the Albitarsis Complex, we hypothesize that the origin of An. albitarsis s.l. was in the Precambrian Guayana Shield in southern Venezuela as the result of vertical migrations triggered by glacial/interglacial alternations followed by dispersal [125]. Although Conn & Mirabello [126] considered that the ancestry of An. albitarsis s.l. was in Venezuela, they proposed that it was due to the interaction of Pleistocene refugia and Miocene-Pliocene marine incursion that determined the distribution pattern of An. albitarsis s.l. At present, based on paleoecological, palynological and molecular phylogenetic studies, reported evidence suggests that the large biodiversity in the Pantepui region was not the result of Pleistocene refuge [127131], but is the result of complex ecological and evolutionary trends initiated by Neogene tectonic events and paleogeographical reorganization, and maintained by the action of Pleistocene climatic changes [132]. The Guayana Shield, located between the Orinoco and Amazon basins, is a vast area which extends over half of Venezuela, Guyana, Suriname, French Guiana, north of Brazil and a small portion of south-east Colombia. This region, one of the riches in biodiversity and endemism, is characterized by the presence of steep table mountains or tepuis separated by large areas of savannahs and rain forests. Mayr and Phelps [133] used the term Pantepui to designate the region located between the summits of the tepuis of the Venezuelan Guayana Shield, including the “Gran Sabana” (Guayana Highlands) and adjacent regions of Brazil and Guyana. Steyermark [134] broadened this denomination to include the “Gran Sabana”, the savannahs and bornhardts of the Venezuelan Amazonas State and the northeast region of Bolívar State. This author defined all this region as the ‘Pantepui Refuge’. The ecological differences between the highlands and lowlands of the Guayana Shield were established and it was proposed that the term Pantepui should be used for ecosystems of middle and high altitudes (approximately above 1200 m) [135137]. Navarro et al. [138], based on Parsimony Analysis of Endemicity between immature Culicidae and their aquatic habitats in plants, concluded that the Venezuelan Guayana Shield was an ancestral center of speciation. Similar observations were made for other taxa. Furthermore, within the Pantepui 60% of the vascular plant species and 87% of the frog species are endemic [139141]. It seems that the Pantepui had emigrational pathways by physically connected valleys and slopes that contributed to radiation of species under the influence of cooling-warming oscillation patterns. Downward altitudinal migrations and dispersion of species followed by fragmentation and isolation of natural populations resulted in adaptive radiation and speciation [129].

The Orinoquia region in Colombia and the western savannahs of Venezuela have a unimodal rainfall pattern with a short dry period between December-April and wet season the rest of the year which produce flooding of lowlands providing available larval habitats for mosquitoes, particularly for species such as An. albitarsis F, that prefer oviposition sites totally exposed to sunlight. During the dry season, fragmentation of these habitats forces adults to find blood sources, natural refuges and to colonize new ecotopes far away from the initial oviposition sites. Since this condition is repeated every year, it can contribute to the dispersion of natural populations of mosquitoes. It is hypothesized that An. albitarsis F could be a lineage that separated from the of origin of basal members of the Albitarsis Complex such as An. marajoara, which could find a refuge in the Pantepui as samples have been collected from the Department of Caquetá in Colombia [15, 16] and in Jabillal, Bolívar State, Venezuela [17], both located on the borders of the Guayana Shield. Clear differences found in An. albitarsis F for the rDNA-ITS2, nDNA (partial loss of an intron in the white gene) [9] and microsatellite gDNA [16] and, mtDNA (MT-CO1 barcode region) [11, 13] could be interpreted as divergences associated with a recent speciation process. The Venezuelan Andes could be a barrier for the expansion of An. albitarsis F towards north western Venezuela and north-eastern Colombia, although the Yaracuy depression between the North-Central Cordillera and the Venezuelan Andes could have been a corridor facilitating the expansion of An. albitarsis F towards the depression of the Maracaibo Lake and into Colombia. The island of Trinidad was part of north-eastern Venezuela until the early Holocene period (about 7000/6000 BCE) when sea level increased by about 60 m [142]; this event clearly explains the presence of An. albitarsis F on this island where it found appropriate ecological niches and at present is associated to irrigated rice fields [27]. Following a similar pathway, the expansions of populations of An. albitarsis s.l. from the Pantepui as an ancestral niche could derive in An. janconnae, currently located in the States of Roraima and Pará, Brazil, in the southern border of the Guayana Shield [11, 101] about 230 km from the Venezuelan border, where it has been described as a specialist species in relation to the selection of larval habitats in moving and sunlight exposed waters [101].

Despite the criticism regarding the use of unique markers such as a DNA barcode for species identification [143, 144]; barcode region analysis has proved to be a more sensitive marker for discriminating between incipient or very recently separated species/lineages compared to rDNA (ITS2) and white gene for the Albitarsis Complex [11, 13, 23, 145]. Likewise, the findings of this study revealed that this DNA fragment was robust for separating all the species of the Albitarsis Complex and suggested that species had reciprocal monophyly according to the current taxonomy [11]. However standard limits between intra and inter-species divergence cannot be generalized to many groups of organisms, the reported “barcoding gap” in our analyses corresponded to an interspecific variation threshold of at least 2% suggested for delimit separated species [43, 146149]. Although sampling limitations in Venezuela (n = 3) could have led to bias in genetic differentiation among populations and species delimitation [150, 151], the present study generated 45 new sequences of An. albitarsis F, and the addition of publicly available MT-CO1 sequences (n = 123) belonging to the Albitarsis Complex from north of the Amazon River, resulted in robust analyses. Nevertheless, further studies on population genetics that include more samples from each of the analyzed localities as well as localities from the proposed geographic expansion of this species are required to elucidate the morphological and genetic variation of An. albitarsis F and sister species. This would thus contribute to strengthening the phylogenetic relationships, hypotheses of their origin, expansion, and geographic dispersion presented in this study, and address questions about its taxonomic status, which has not yet been formally described.

Conclusions

Phylogenetic analysis showed a well-supported clade with An. albitarsis F, An. janconnae and An. albitarsis I geographically restricted to northern South America. Biogeographic analysis based on the current geographical distribution of these sister taxa allowed the hypothesis that its ancestor came from An. marajoara and colonized the Pantepui region and then migrated to western Venezuela and Colombia suffering speciation events to generate An. albitarsis F and then An. albitarsis I. Migration south of Venezuela produced An. janconnae. Anopheles albitarsis F is recorded in Colombia, Venezuela and the island of Trinidad while An. albitarsis I is recorded in Venezuela and Colombia showing a hybridization zone in the border of both countries in Norte de Santander Department and Zulia State. Natural infections by P. vivax and P. falciparum in Colombia and Venezuela and epidemiological incrimination constitute strong evidence of the important role of An. albitarsis F in the transmission of malaria parasites. Entomological surveillance for these species in Colombia, Venezuela, Trinidad and Tobago, and also in Guyana, Ecuador, Perú, and Panamá, are mandatory to update the geographic distribution, ecological and biological aspects and its possible role as a regional vector of malaria parasites.

Supporting information

S1 Table. Comparison between DNA mitochondrially encoded cytochrome c oxidase I (MT-CO1) gene sequences obtained in this study and those available in the GenBank database.

: Numerical order of the sequences; Code: abbreviation of the sequences obtained; Percentage of Identify: Similarity of the sequences obtained in this study with the sequences available in the GenBank; Comparison in BLASTn, Author: References of the sequences compared; Species: Species identified; Accession N°: Accession number of the compared sequences.

https://doi.org/10.1371/journal.pone.0253230.s001

(DOCX)

S2 Table. Information of the 37 haplotypes generated with DNA mitochondrially encoded cytochrome c oxidase I (MT-CO1) gene sequences database from Colombia (n = 42) and Venezuela (n = 3).

H: Haplotypes; : Absolute frequency of individuals observed in each haplotype. Within parentheses are the numbers of individuals observed for each haplotype in each locality. CO: Colombia, VE: Venezuela.

https://doi.org/10.1371/journal.pone.0253230.s002

(DOCX)

S3 Table. Information of 106 haplotypes generated with the 168 DNA mitochondrially encoded cytochrome c oxidase I (MT-CO1) gene sequences database.

H: Haplotypes; : Absolute frequency of individuals observed in each haplotype. Within parentheses are the numbers of individuals observed for each haplotype in each locality. BR: Brazil, CO: Colombia, TT: Trinidad, VE: Venezuela.

https://doi.org/10.1371/journal.pone.0253230.s003

(DOCX)

S4 Table. Information of 75 haplotypes generated with the 100 DNA mitochondrially encoded cytochrome c oxidase I (MT-CO1) gene sequences database.

H: Haplotypes; : Absolute frequency of individuals observed in each haplotype. Within parentheses are the numbers of individuals observed for each haplotype in each locality. BR: Brazil, CO: Colombia, TT: Trinidad, VE: Venezuela.

https://doi.org/10.1371/journal.pone.0253230.s004

(DOCX)

S1 Fig. Barcode analysis plots of Anopheles albitarsis complex species reported in South America.

Barcode gap analysis of all species within the Anopheles albitarsis complex, plots are based in distance matrices of the clusters determined using NJ-K2P distances. Y-axis: genetic divergence and X-axis clusters. alb: An. albitarsis s.s.; ory: An. oryzalimnetes; mar: An. marajoara; dea: An. deaneorum; jan: An. janconnae; albF: An. albitarsis F; albG: An. albitarsis G; albH: An. albitarsis H; albI: An. albitarsis I.

https://doi.org/10.1371/journal.pone.0253230.s005

(TIF)

Acknowledgments

Special thanks are due to Jorge E. Moreno for providing specimens from San Rafael and sharing unpublished data. We thank MM Póvoa (Brazil); M Middelveen and R Allan (Canada); F Ruíz-López, M Correa and V Olano (Colombia); LF Chaves (Costa Rica); B Puno (Guyana); J Loaiza (Panamá); H. Hiwat (Suriname), and D. Strickman (USA) for valuable comments and references.

References

  1. 1. Kitzmiller JB. Chromosomal differences among species of Anopheles mosquitoes. Mosq Syst. 1977; 9(2): 112–22.
  2. 2. Kreutzer RD, Kitzmiller JB, Rabbani MG. Cytogenetically distinguishable sympatric and allopatric populations of the mosquito Anopheles albitarsis. Acta Amazon. 1976; 6: 473–81.
  3. 3. Narang SK, Klein TA, Perera OP, Lima JB, Tang AT. Genetic evidence for the existence of cryptic species in the Anopheles albitarsis complex in Brazil: allozymes and mitochondrial DNA restriction fragment length polymorphisms. Biochem Genet. 1993; 31(1–2): 97–112. pmid:8097085.
  4. 4. Steiner WWM, Narang SK, Kitzmiller JB, Swofford DL. Genetic divergence and evolution in neotropical Anopheles (subgenus Nyssorhynchus). In: Steiner WWM, Tabachnick WJK, Rai S, Narang SK, editors. Recent Developments Genetics Insects Disease Vectors: Stipes Publishing, Champaign; 1982. p. 523–50.
  5. 5. Wilkerson RC, Gaffigan TV, Bento Lima J. Identification of species related to Anopheles (Nyssorhynchus) albitarsis by random amplified polymorphic DNA-polymerase chain reaction (Diptera: Culicidae). Mem Inst Oswaldo Cruz. 1995; 90(6): 721–32. pmid:8731368.
  6. 6. Wilkerson RC, Parsons TJ, Klein TA, Gaffigan TV, Bergo E, Consolim J. Diagnosis by random amplified polymorphic DNA polymerase chain reaction of four cryptic species related to Anopheles (Nyssorhynchus) albitarsis (Diptera: Culicidae) from Paraguay, Argentina, and Brazil. J Med Entomol. 1995; 32(5): 697–704. pmid:7473625.
  7. 7. Motoki MT, Wilkerson RC, Sallum MAM. The Anopheles albitarsis complex with the recognition of Anopheles oryzalimnetes Wilkerson and Motoki, n. sp. and Anopheles janconnae Wilkerson and Sallum, n. sp. (Diptera: Culicidae). Mem Inst Oswaldo Cruz. 2009; 104: 823–50. pmid:19876554
  8. 8. Rosa-Freitas MG, Deane LM, Momen H. A morphological, isoenzymatic and behavioural study of ten populations of Anopheles (Nyssorhynchus) albitarsis Lynch-Arribálzaga, 1878 (Diptera: Culicidae) including from the type-locality—Baradero, Argentina. Mem Inst Oswaldo Cruz. 1990; 85: 275–89.
  9. 9. Brochero HL, Li C, Wilkerson RC. A newly recognized species in the Anopheles (Nyssorhynchus) albitarsis complex (Diptera: Culicidae) from Puerto Carreño, Colombia. Am J Trop Med Hyg. 2007; 76(6): 1113–7. pmid:17556620.
  10. 10. Krzywinski J, Li C, Morris M, Conn JE, Lima JB, Póvoa MM, et al. Analysis of the evolutionary forces shaping mitochondrial genomes of a Neotropical malaria vector complex. Mol Phylogenet Evol. 2011; 58(3): 469–77. pmid:21241811.
  11. 11. Ruíz-López F, Wilkerson RC, Conn JE, McKeon SN, Levin DM, Quiñones ML, et al. DNA barcoding reveals both known and novel taxa in the Albitarsis Group (Anopheles: Nyssorhynchus) of Neotropical malaria vectors. Parasit Vectors. 2012; 5: 44. pmid:22353437.
  12. 12. Gutiérrez LA, Orrego LM, Gómez GF, López A, Luckhart S, Conn JE, et al. A new mtDNA COI gene lineage closely related to Anopheles janconnae of the Albitarsis Complex in the Caribbean region of Colombia. Mem Inst Oswaldo Cruz. 2010; 105(8): 1019–25. pmid:21225199
  13. 13. Motoki MT, Linton YM, Conn JE, Ruíz-López F, Wilkerson RC. Phylogenetic Network of Mitochondrial COI Gene Sequences Distinguishes 10 Taxa Within the Neotropical Albitarsis Group (Diptera: Culicidae), Confirming the Separate Species Status of Anopheles albitarsis H (Diptera: Culicidae) and Revealing a Novel Lineage, Anopheles albitarsis J. J Med Entomol. 2020. pmid:33033825.
  14. 14. Foley DH, Linton YM, Ruíz-López JF, Conn JE, Sallum MA, Póvoa MM, et al. Geographic distribution, evolution, and disease importance of species within the Neotropical Anopheles albitarsis Group (Diptera, Culicidae). J Vector Ecol. 2014; 39(1): 168–81. pmid:24820570.
  15. 15. Brochero H. Estructura genética poblacional del mosquito Anopheles (Nyssorhynchus) marajoara Galvão & Damasceno 1942 (Diptera: Culicidae) de Colombia. [PhD Thesis]. Bogotá, Colombia: Pontificia Universidad Javeriana; 2006. https://biblos.javeriana.edu.co/uhtbin/cgisirsi/x/0/0/57/5/3?searchdata1=729923%7bCKEY%7d&searchfield1=GENERAL%5eSUBJECT%5eGENERAL%5e%5e&user_id=WEBSERVER
  16. 16. Brochero H, Li C, Wilkerson R, Conn JE, Ruiz-Garcia M. Genetic structure of Anopheles (Nyssorhynchus) marajoara (Diptera: Culicidae) in Colombia. Am J Trop Med Hyg. 2010; 83(3): 585–95. pmid:20810825.
  17. 17. Rubio-Palis Y, Ruíz-López F, Guzmán H, Sánchez V, Moreno J, Estrada Y, et al. Primer registro de Anopheles (Nyssorhynchus) oswaldoi B y Anopheles (Nys.) albitarsis F en la cuenca del río Caura, Estado Bolívar, Venezuela. Bol Mal Salud Amb. 2013; 53: 68–72.
  18. 18. Pacheco-Gómez MA, González-Obando R, Brochero H. Morphometric variations of two populations of Anopheles albitarsis F (Diptera: Culicidae) in the Orinoquia region, Colombia. Rev Fac Med Univ Nac Colomb. 2018; 66(2): 201–8.
  19. 19. Osborn FR, Rubio-Palis Y, Herrera M, Figuera A, Moreno JE. Caracterización ecoregional de los vectores de malaria en Venezuela. Bol Mal Salud Amb. 2004; 44: 77–92.
  20. 20. Rubio-Palis Y, Zimmerman RH. Ecoregional classification of malaria vectors in the neotropics. J Med Entomol. 1997; 34(5): 499–510. pmid:9379453.
  21. 21. Jiménez P, Conn JE, Wirtz R, Brochero H. Anopheles (Diptera: Culicidae) vectores de malaria en el municipio de Puerto Carreño, Vichada, Colombia. Biomédica. 2012; 32 Suppl 1: 13–21. pmid:23235809.
  22. 22. Herrera S, Suárez M, Sánchez G, Quiñones M, Herrera M. Uso de la técnica inmuno-radiométrica (IRMA) en Anopheles de Colombia para la identificación de esporozoitos de Plasmodium. Colomb Med. 1987; 18: 2–6.
  23. 23. Gómez G, Jaramillo L, Correa MM. Wing geometric morphometrics and molecular assessment of members in the Albitarsis Complex from Colombia. Mol Ecol Resour. 2013; 13(6): 1082–92. pmid:23702155.
  24. 24. Gómez GF, Correa MM. Discrimination of Neotropical Anopheles species based on molecular and wing geometric morphometric traits. Infect Genet Evol. 2017; 54: 379–86. pmid:28774799
  25. 25. Rubio-Palis Y, Wilkerson R, Guzmán H. Morphological characters of adult Anopheles (Nyssorhynchus) marajoara in Venezuela. J Am Mosq Control Assoc. 2003; 19(2): 107–14. pmid:12825659.
  26. 26. Lehr MA, Kilpatrick CW, Wilkerson RC, Conn JE. Cryptic Species in the Anopheles (Nyssorhynchus) albitarsis (Diptera: Culicidae) Complex: Incongruence Between Random Amplified Polymorphic DNA-Polymerase Chain Reaction Identification and Analysis of Mitochondrial DNA COI Gene Sequences. Ann Entomol Soc Am. 2005; 98(6): 908–17. pmid:17082822.
  27. 27. Chadee DD, Wilkerson RC. Ecology of the malaria vector, Anopheles (Nyssorhynchus) marajoara Galvão and Damasceno in Trinidad, West Indies. J Am Mosq Control Assoc. 2006; 22(1): 22–8. pmid:16646317.
  28. 28. Gabaldon A. Malaria eradication in Venezuela: doctrine, practice, and achievements after twenty years. Am J Trop Med Hyg. 1983; 32(2): 203–11. pmid:6340536
  29. 29. PAHO/WHO. Profiles: Malaria Champions in the Americas 2019. https://www.paho.org/en/topics/malaria/malaria-champions-2019/profiles-malaria-champions-americas-2019.
  30. 30. Moreno JE, Rubio-Palis Y, Páez E, Pérez E, Sánchez V, Vaccari E. Anopheles (Anopheles) neomaculipalpus: a new malaria vector in the Amazon basin?. Med Vet Entomol. 2005; 19(3): 329–32. pmid:16134983.
  31. 31. Moreno JE, Rubio-Palis Y, Páez E, Pérez E, Sánchez V, Vaccari E. Malaria entomological inoculation rates in gold mining areas of Southern Venezuela. Mem Inst Oswaldo Cruz. 2009; 104(5): 764–8. pmid:19820839.
  32. 32. Grillet ME, Moreno JE, Hernández JV, Vincenti-González MF, Noya O, Tami A, et al. Malaria in Southern Venezuela: The Hottest Hotspot in Latin America. 2020.
  33. 33. Buitrago LS, Brochero HL, McKeon SN, Lainhart W, Conn JE. First published record of urban malaria in Puerto Gaitán, Meta, Colombia. Mem Inst Oswaldo Cruz. 2013; 108(8): 1045–50. pmid:24402157.
  34. 34. Jiménez IP, Conn JE, Brochero H. Malaria vectors in San José del Guaviare, Orinoquia, Colombia. J Am Mosq Control Assoc. 2014; 30(2): 91–8. pmid:25102591.
  35. 35. Durán JS. Determinación del papel en la transmisión de malaria de las especies de Anopheles (Diptera: Culicidae) de dos localidades endémicas para malaria en Colombia. [M. Sc. Thesis]. Bogotá, Colombia: Universidad Nacional de Colombia; 2014. https://repositorio.unal.edu.co/handle/unal/52260
  36. 36. Linthicum KJ. A Revision of the Argyritarsis Section of the subgenus Nyssorhynchus of Anopheles (Diptera: Culicidae). Mosq Syst. 1988; 20(2): 98–271.
  37. 37. Rubio-Palis Y. Anopheles (Nyssorhynchus) de Venezuela: taxonomía, bionomía, ecología e importancia médica. Maracay, Venezuela: Escuela de Malariología y Saneamiento Ambiental “Dr. Arnoldo Gabaldon”; 2000. www.iaes.edu.ve/index.php/centro-de-descargas/viewcategory/3-libros-y-publicaciones
  38. 38. González R, Carrejo NS. Introducción al estudio taxonómico de Anopheles de Colombia. Claves y notas de distribución. 2nd ed. Santiago de Cali: Universidad del Valle; 2009.
  39. 39. Folmer O, Black M, Hoeh W, Lutz R, Vrijenhoek R. DNA primers for amplification of mitochondrial cytochrome c oxidase subunit I from diverse metazoan invertebrates. Mol Mar Biol Biotechnol. 1994; 3(5): 294–9. pmid:7881515.
  40. 40. Thompson JD, Higgins DG, Gibson TJ. CLUSTAL W: improving the sensitivity of progressive multiple sequence alignment through sequence weighting, position-specific gap penalties and weight matrix choice. Nucleic Acids Res. 1994; 22(22): 4673–80. pmid:7984417.
  41. 41. Xia X. Imputing missing distances in molecular phylogenetics. Peer J. 2018; 6: e5321. pmid:30065887.
  42. 42. Altschul SF, Gish W, Miller W, Myers EW, Lipman DJ. Basic local alignment search tool. J Mol Biol. 1990; 215(3): 403–10. pmid:2231712.
  43. 43. Ratnasingham S, Hebert PD. BOLD: The Barcode of Life Data System (http://www.barcodinglife.org). Mol Ecol Notes. 2007; 7(3): 355–64. pmid:18784790.
  44. 44. Kumar S, Stecher G, Li M, Knyaz C, Tamura K. MEGA X: Molecular Evolutionary Genetics Analysis across Computing Platforms. Mol Biol Evol. 2018; 35(6): 1547–9. pmid:29722887.
  45. 45. Kimura M. A simple method for estimating evolutionary rates of base substitutions through comparative studies of nucleotide sequences. J Mol Evol. 1980; 16(2): 111–20. pmid:7463489.
  46. 46. Felsenstein J. Confidence Limits on Phylogenies: An Approach Using the Bootstrap. Evolution. 1985; 39(4): 783–91. pmid:28561359.
  47. 47. Saitou N, Nei M. The neighbor-joining method: a new method for reconstructing phylogenetic trees. Mol Biol Evol. 1987; 4(4): 406–25. pmid:3447015.
  48. 48. Suchard MA, Lemey P, Baele G, Ayres DL, Drummond AJ, Rambaut A. Bayesian phylogenetic and phylodynamic data integration using BEAST 1.10. Virus Evol. 2018; 4(1): vey016. pmid:29942656.
  49. 49. Hasegawa M, Kishino H, Yano T. Dating of the human-ape splitting by a molecular clock of mitochondrial DNA. J Mol Evol. 1985; 22(2): 160–74. pmid:3934395.
  50. 50. Darriba D, Taboada GL, Doallo R, Posada D. jModelTest 2: more models, new heuristics and parallel computing. Nat Methods. 2012; 9(8): 772. pmid:22847109.
  51. 51. Rambaut A, Drummond AJ, Xie D, Baele G, Suchard MA. Posterior Summarization in Bayesian Phylogenetics Using Tracer 1.7. Syst Biol. 2018; 67(5): 901–4. pmid:29718447.
  52. 52. Rambaut A. FigTree, version 1.4.3 [internet]. Edinburgh, UK: University of Edinburgh; 2016. http://tree.bio.ed.ac.uk/software/figtree/.
  53. 53. Rozas J, Ferrer-Mata A, Sánchez-Del Barrio JC, Guirao-Rico S, Librado P, Ramos-Onsins SE, et al. DnaSP 6: DNA Sequence Polymorphism Analysis of Large Data Sets. Mol Biol Evol. 2017; 34(12): 3299–302. pmid:29029172.
  54. 54. Tajima F. Evolutionary relationship of DNA sequences in finite populations. Genetics. 1983; 105(2): 437–60. pmid:6628982.
  55. 55. Leigh JW, Bryant D. PopART: Full-feature software for haplotype network construction. Methods Ecol Evol 2015; 6(9): 1110–6.
  56. 56. Clement M, Snell Q, Walke P, Posada D, Crandall K. TCS: estimating gene genealogies. Proc 16th Int Parallel Distrib Process Symp. 2002; 2(184).
  57. 57. Sinka ME, Rubio-Palis Y, Manguin S, Patil AP, Temperley WH, Gething PW, et al. The dominant Anopheles vectors of human malaria in the Americas: occurrence data, distribution maps and bionomic precis. Parasit Vectors. 2010; 3: 72. pmid:20712879.
  58. 58. CRD (Centre for Reviews and Dissemination). Systematic reviews: CRD’s Guidance for undertaking reviews in health care. York, UK: York Publishing Services, Ltd.; 2009.
  59. 59. QGIS Geographic Information System. Open Source Geospatial Foundation Project; 2018. http://qgis.org/.
  60. 60. Orjuela LI, Herrera M, Erazo H, Quiñones ML. Especies de Anopheles presentes en el departamento del Putumayo y su infección natural con Plasmodium. Biomédica. 2013; 33(1):42–52.
  61. 61. Collins WE, Warren M, Skinner JC, Sutton BB. Infectivity of two strains of Plasmodium vivax to Anopheles albitarsis mosquitoes from Colombia. J Parasitol. 1985; 71(6): 771–3. pmid:3912483.
  62. 62. Brochero HL, Rey G, Buitrago LS, Olano VA. Biting activity and breeding sites of Anopheles species in the municipality Villavicencio, Meta, Colombia. J Am Mosq Control Assoc. 2005; 21(2): 182–6. pmid:16033120.
  63. 63. Posso CE, González R, Cárdenas H, Tascón R. Genetic structure of Anopheles darlingi Root, An. nuneztovari Gabaldon and An. marajoara Galvão & Damasceno from Colombia using RAPD-PCR. Rev Colomb Entomol. 2006; 32: 49–56.
  64. 64. Gutiérrez LA, Naranjo N, Jaramillo LM, Muskus C, Luckhart S, Conn JE, et al. Natural infectivity of Anopheles species from the Pacific and Atlantic Regions of Colombia. Acta Trop. 2008; 107(2): 99–105. pmid:18554564.
  65. 65. Gutiérrez LA, González JJ, Gómez GF, Castro MI, Rosero DA, Luckhart S, et al. Species composition and natural infectivity of anthropophilic Anopheles (Diptera: Culicidae) in the states of Córdoba and Antioquia, Northwestern Colombia. Mem Inst Oswaldo Cruz. 2009; 104(8): 1117–24. pmid:20140372.
  66. 66. Naranjo-Díaz N, Rosero DA, Rua-Uribe G, Luckhart S, Correa MM. Abundance, behavior and entomological inoculation rates of anthropophilic anophelines from a primary Colombian malaria endemic area. Parasit Vectors. 2013; 6: 61. pmid:23497535.
  67. 67. Montoya C, Bascunan P, Rodríguez-Zabala J, Correa MM. Abundance, composition and natural infection of Anopheles mosquitoes from two malaria-endemic regions of Colombia. Biomédica. 2017; 37: 98–105. pmid:29161482.
  68. 68. Naranjo-Díaz N, Altamiranda-Saavedra M, Correa MM. Anopheles species composition and entomological parameters in malaria endemic localities of North West Colombia. Acta Trop. 2019; 190: 13–21. pmid:30367837.
  69. 69. Hernández-Valencia JC, Rincón DS, Marín A, Naranjo-Díaz , Correa MM. Effect of land cover and landscape fragmentation on anopheline mosquito abundance and diversity in an important Colombian malaria endemic region. PLoS ONE. 2020; 15(10): e0240207. pmid:33057442
  70. 70. Berti J, Guzmán H, Liria J, González J, Estrada Y, Pérez E. Nuevos registros de mosquitos (Diptera: Culicidae) para el estado Bolívar, Venezuela: Dos de ellos nuevos para el país. Bol Mal Salud Amb. 2011; 51: 59–69.
  71. 71. Berti J, Ramírez R, Estrada Y, Guzmán H, Arias L. Registros de altitud de mosquitos anofelinos (Diptera: Culicidae: Anophelinae) del Municipio Gran Sabana, estado Bolívar, Venezuela, y nuevos datos altitudinales de importancia. Bol Mal Salud Amb. 2016; 56: 78–86.
  72. 72. Moreno J, Rubio-Palis Y, Acevedo P. Identificación de criaderos de anofelinos en un área endémica del estado Bolívar, Venezuela. Bol Mal Salud Amb. 2000; 40: 21–30.
  73. 73. Moreno JE, Rubio-Palis Y, Páez E, Pérez E, Sánchez V. Abundance, biting behaviour and parous rate of anopheline mosquito species in relation to malaria incidence in gold-mining areas of southern Venezuela. Med Vet Entomol. 2007; 21(4): 339–49. pmid:18092972.
  74. 74. Moreno JE, Rubio-Palis Y, Sánchez V, Martínez Á. Fluctuación poblacional y hábitat larval de anofelinos en el municipio Sifontes, estado Bolívar, Venezuela. Bol Mal Salud Amb. 2015; 55: 52–68.
  75. 75. Moreno JE, Rubio-Palis Y, Sánchez V, Martínez Á. Caracterización de hábitats larvales de anofelinos en el municipio Sifontes del estado Bolívar, Venezuela. Bol Mal Salud Amb. 2015; 55: 117–31.
  76. 76. Moreno JE, Rubio-Palis Y, Bevilacqua M, Sánchez V, Guzmán H. Caracterización de hábitats larvales de Anofelinos en el bajo Río Caura, región malárica del Estado Bolívar, Venezuela. Bol Mal Salud Amb. 2018; 58: 17–30.
  77. 77. Navarro E, Grillet ME, Menare C, González J, Frontado H. Inventario preliminar de anofelinos (Díptera: Culicidae) en áreas endémicas de malaria, municipios Atures y Autana, estado Amazonas, Venezuela. Bol Mal Salud Amb. 2015; 55: 194–8.
  78. 78. Rubio-Palis Y, Wirtz RA, Curtis CF. Malaria entomological inoculation rates in western Venezuela. Acta Trop. 1992; 52(2–3): 167–74. pmid:1363181.
  79. 79. Rubio-Palis Y, Curtis CF. Biting and resting behaviour of anophelines in western Venezuela and implications for control of malaria transmission. Med Vet Entomol. 1992; 6(4): 325–34. pmid:1463897
  80. 80. Rubio-Palis Y, Moreno JE, Sánchez V, Estrada Y, Anaya W, Bevilacqua M, et al. Can Mosquito Magnet® substitute for human-landing catches to sample anopheline populations?. Mem Inst Oswaldo Cruz. 2012; 107(4): 546–9. pmid:22666868.
  81. 81. Sutil E. Enumeración histórica y geográfica de las especies de Culicidae de Venezuela ordenadas según su taxonomía. Bol Dir Malariol San Amb. 1980; 20: 1–32.
  82. 82. Rubio-Palis Y, Bevilacqua M, Medina D, Moreno J, Cárdenas L, Sánchez V, et al. Malaria entomological risk factors in relation to land cover in the Lower Caura River Basin, Venezuela. Mem Inst Oswaldo Cruz. 2013; 108(2): 220–8. pmid:23579803
  83. 83. Rubio-Palis Y, Curtis CF. Evaluation of different methods of catching anopheline mosquitoes in western Venezuela. J Am Mosq Control Assoc. 1992; 8(3): 261–7. pmid:1402863.
  84. 84. Moreno J, Rubio-Palis Y, Pérez E, Sánchez V, Páez E. Evaluación de tres métodos de captura de anofelinos en un área endémica de malaria del estado Bolívar, Venezuela. Entomotropica. 2002; 17(2): 157–65.
  85. 85. Rubio-Palis Y, Curtis CF, Gonzáles C, Wirtz RA. Host choice of anopheline mosquitoes in a malaria endemic area of western Venezuela. Med Vet Entomol. 1994; 8(3): 275–80. pmid:7949319.
  86. 86. Rubio-Palis Y. Variation of the vectorial capacity of some anophelines in western Venezuela. Am J Trop Med Hyg. 1994; 50(4): 420–4. pmid:8166348.
  87. 87. Abou Orm S, Moreno JE, Carrozza M, Acevedo P, Herrera F. Tasas de infección de Plasmodium spp. para algunos Anopheles spp. del municipio Sifontes, Estado Bolívar, Venezuela. Bol Mal Salud Amb. 2017; 57: 17–25.
  88. 88. Molina de Fernández D, Figueroa LE, Pérez E. Resistencia múltiple a insecticidas en Anopheles marajoara Galvão & Damasceno, 1942 en zonas agrícolas. Salud & Desarrollo Social. 2007; 28: 108–16.
  89. 89. Chadee DD. Indoor and outdoor host-seeking rhythms of Anopheles albitarsis (Diptera: Culicidae) in Trinidad, West Indies. J Med Entomol. 1992; 29(3): 567–9. pmid:1625309.
  90. 90. Chadee DD, Le Maitre A, Tilluckdharry CC. An outbreak of Plasmodium vivax malaria in Trinidad, W.I. Ann Trop Med Parasitol. 1992; 86(6): 583–90. pmid:1304699.
  91. 91. Giglioli G. Fifteen years experience in malaria eradication in British Guiana. Maintenance policies and residual problems. Riv Parassitol. 1959; 20: 279–88.
  92. 92. Rambajan I. Reappearance of Anopheles darlingi Root and vivax malaria in a controlled area of Guyana, South America. Trop Geogr Med. 1984; 36(1): 61–6. pmid:6375050.
  93. 93. Hudson J. Anopheles darlingi Root (Diptera: Culicidae) in the Suriname rain forest. Bull Entomol Res. 1984; 74: 129–42.
  94. 94. Van Der Kuyp E. Mosquitoes in and around Wageningen, Nickerie District, Suriname. Sur Med Bull. 1985; 9: 108–15.
  95. 95. Dusfour I, Jarjaval F, Gaborit P, Mura M, Girod R, Pages F. Confirmation of the occurrence of Anopheles (Nyssorhynchus) marajoara in French Guiana. J Am Mosq Control Assoc. 2012; 28(4): 309–11. pmid:23393754.
  96. 96. de Santi VP, Girod R, Mura M, Dia A, Briolant S, Djossou F, et al. Epidemiological and entomological studies of a malaria outbreak among French armed forces deployed at illegal gold mining sites reveal new aspects of the disease’s transmission in French Guiana. Malar J. 2016; 15: 35. pmid:26801629.
  97. 97. Baxter CP, Zetek J. The Anopheles of Panama with special reference to hand lens identication and notes on collecting and care of specimens. J Trop Med Hyg. 1944; 21: 105–23.
  98. 98. Arnett JR. Notes on the distribution, habits and habitats of some Panamanian mosquitoes (Diptera: Culicidie). JNY Entomol Soc. 1947; 4: 185–200.
  99. 99. Blanton F, Peyton EL. Notes and distribution records of Anopheles and Chagasia mosquitoes in Panama based on a three-year light trap survey. Mosq News. 1956; 16: 22–7.
  100. 100. Loaiza JR, Bermingham E, Scott ME, Rovira JR, Conn JE. Species composition and distribution of adult Anopheles (Diptera: Culicidae) in Panama. J Med Entomol. 2008; 45(5): 841–51. pmid:18826025.
  101. 101. McKeon SN, Schlichting CD, Póvoa MM, Conn JE. Ecological suitability and spatial distribution of five Anopheles species in Amazonian Brazil. Am J Trop Med Hyg. 2013; 88(6): 1079–86. pmid:23546804.
  102. 102. da Silva-Vasconcelos A, Kato MY, Mourao EN, de Souza RT, Lacerda RN, Sibajev A, et al. Biting indices, host-seeking activity and natural infection rates of anopheline species in Boa Vista, Roraima, Brazil from 1996 to 1998. Mem Inst Oswaldo Cruz. 2002; 97(2): 151–61. pmid:12016435.
  103. 103. Póvoa MM, de Souza RT, Lacerda RN, Rosa ES, Galiza D, de Souza JR, et al. 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(2): 163–8. pmid:16830709.
  104. 104. dos Santos RLC, Padilha A, Costa MDP, Costa EM, Dantas-Filho HdC, Póvoa MM. Vectores de malária em duas reservas indígenas da Amazônia Brasileira. Rev Saude Publica. 2009; 43(5): 859–68. pmid:19851633.
  105. 105. McKeon SN, Lehr MA, Wilkerson RC, Ruiz JF, Sallum MA, Lima JB, et al. Lineage divergence detected in the malaria vector Anopheles marajoara (Diptera: Culicidae) in Amazonian Brazil. Malar J. 2010; 9: 271. pmid:20929572.
  106. 106. Galardo AK, Zimmerman RH, Lounibos LP, Young LJ, Galardo CD, Arruda M, et al. Seasonal abundance of anopheline mosquitoes and their association with rainfall and malaria along the Matapi River, Amapá, Brazil. Med Vet Entomol. 2009; 23(4): 335–49. pmid:19941599.
  107. 107. Rafael MS, dos Santos-Junior IP, Tadei WP, Sallum MAM, Forattini OP. Karyotype of Brazilian Anopheles albitarsis sensu lato (Diptera: Culicidae). Genet Mol Res. 2005; 4(4): 684–90. pmid:16475113.
  108. 108. Rubio-Palis Y, Ramírez Álvarez R, Guzmán H, Estrada Y. Evaluación de la trampa Mosquito Magnet® con y sin octenol para capturar mosquitos (Diptera: Culicidae). Bol Mal Salud Amb. 2014; 54(1): 100–2.
  109. 109. Burke K. Tectonic evolution of the Caribbean. Ann Rev Earth Planet Sci. 1988; 16: 201–230.
  110. 110. Hedges SB. Biogeography of the West Indies: An overview. In Woods CA, Sergile FE, editors. Biogeography of the West Indies: Patterns and perspectives. Boca Raton, FL: CRC Press; 2001. p. 15–33.
  111. 111. Rubio-Palis Y, Menare C, Quinto A, Magris M, Amarista M. Caracterización de criaderos de anofelinos (Diptera: Culicidae) vectores de malaria del Alto Orinoco, Amazonas, Venezuela. Entomotropica. 2005; 20(1): 29–38.
  112. 112. Rubio-Palis Y, Ramírez Álvarez R, Guzmán H, Suárez A, Navarro JC. Abundancia y diversidad de especies de Culicinae (Diptera: Culicidae) del Alto Orinoco, estado Amazonas, Venezuela. Bol Mal Salud Amb. 2014; 54(2): 186–98.
  113. 113. Rubio-Palis Y, Moreno JE, Guzmán H, Sánchez V, Bevilacqua MP, Cárdenas L. Mosquitos (Diptera: Culicidae) de la cuenca del río Caura, estado Bolívar, Venezuela. Nuevos registros para el país y el estado. Bol Mal Salud Amb. 2019; 59(2): 98–111.
  114. 114. Rubio-Palis Y, Manguin S, Ayesta C, Guzmán H, Arcia J, González J, et al. Revisión taxonómica de los anofelinos vectores de malaria en el sur de Venezuela. Bol Dir Malariol San Amb. 1997; 37: 35–48.
  115. 115. Laporta GZ, Linton YM, Wilkerson RC, Bergo ES, Nagaki SS, Sant’Ana DC, et al. Malaria vectors in South America: current and future scenarios. Parasit Vectors. 2015; 8: 426. pmid:26283539.
  116. 116. de Fátima Rossetti D, Mann de Toledo P, Góes AM. New geological framework for Western Amazonia (Brazil) and implications for biogeography and evolution. Quat Res. 2005; 63: 78.
  117. 117. Zeisset I, Beebee TJ. Amphibian phylogeography: a model for understanding historical aspects of species distributions. Heredity. 2008; 101(2): 109–19. pmid:18493262.
  118. 118. Pedro PM, Uezu A, Sallum MA. Concordant phylogeographies of 2 malaria vectors attest to common spatial and demographic histories. J Hered. 2010; 101(5): 618–27. pmid:20511380.
  119. 119. Santorelli S Jr, Magnusson WE, Deus CP. Most species are not limited by an Amazonian river postulated to be a border between endemism areas. Sci Rep. 2018; 8(1): 2294. pmid:29396491.
  120. 120. Marinotti O, Jasinskiene N, Fazekas A, Scaife S, Fu G, Mattingly ST, et al. Development of a population suppression strain of the human malaria vector mosquito, Anopheles stephensi. Malar J. 2013; 12: 142. pmid:23622561.
  121. 121. Krzywinski J, Grushko OG, Besansky NJ. Analysis of the complete mitochondrial DNA from Anopheles funestus: an improved dipteran mitochondrial genome annotation and a temporal dimension of mosquito evolution. Mol Phylogenet Evol. 2006; 39(2): 417–23. pmid:16473530.
  122. 122. Reidenbach KR, Cook S, Bertone MA, Harbach RE, Wiegmann BM, Besansky NJ. Phylogenetic analysis and temporal diversification of mosquitoes (Diptera: Culicidae) based on nuclear genes and morphology. BMC Evol Biol. 2009; 9: 298. pmid:20028549.
  123. 123. Martínez-Villegas L, Assis-Geraldo J, Koerich LB, Collier TC, Lee Y, Main BJ, et al. Characterization of the complete mitogenome of Anopheles aquasalis, and phylogenetic divergences among Anopheles from diverse geographic zones. PLoS ONE. 2019; 14(9): e0219523. pmid:31479460.
  124. 124. Krzywinski J, Besansky NJ. Molecular systematics of Anopheles: from subgenera to subpopulations. Ann Rev Entomol. 2003; 48: 111–39. pmid:12208816.
  125. 125. Rull V. Biotic diversification in the Guayana Highlands: a proposal. J Biogeogr. 2005; 32: 921–7.
  126. 126. Conn JE, Mirabello L. The biogeography and population genetics of neotropical vector species. Heredity. 2007; 99(3): 245–56. pmid:17534382.
  127. 127. Rull V. Biogeography of the ‘Lost World’: a palaeoecological perspective. Earth Sci Rev. 2004; 67(1–2): 125–37.
  128. 128. Rull V. An evaluation of the Lost World and Vertical Displacement hypotheses in the Chimantá massif, Venezuelan Guayana. Glob Ecol Biogeogr. 2004; 13: 141–8.
  129. 129. Rull V. Speciation timing and neotropical biodiversity: the Tertiary-Quaternary debate in the light of molecular phylogenetic evidence. Mol Ecol. 2008; 17(11): 2722–9. pmid:18494610.
  130. 130. Rull V, Montoya E, Nogué S, Safont E, Vegas-Vilarrúbia T. Climatic and ecological history of Pantepui and surrounding areas. In: Rull V, Vegas-Vilarrúbia T, Huber O, Señaris C, editors. Biodiversity of Pantepui: the pristine “Lost World” of the Neotropics. London: Elsevier-Academic Press; 2019. p. 33–54.
  131. 131. Schubert C, Fritz P, Aravena R. Late quaternary paleoenvironmental studies in the Gran Sabana (Venezuelan Guayana shield). Quat Int. 1994; 21: 81–90.
  132. 132. Rull V. Origins of biodiversity. Science. 2011; 331(6016): 398–9; author reply 9–400. pmid:21273468.
  133. 133. Mayr E, Phelps WH. The origin of the bird fauna of the south Venezuelan highlands. Bull Am Mus Nat Hist. 1967; 136: 273–327.
  134. 134. Steyermark J. Plant refuge and dispersal centres in Venezuela: their relict and endemic element. In: Larsen K, Holm-Nielsen L, editors. Tropical Botany. London: Academic Press; 1979. p. 185–221.
  135. 135. Huber O. Consideraciones sobre el concepto de Pantepui. Pantepui. 1987; 2:2–10.
  136. 136. Huber O. Guayana highlands versus Guayana lowlands, a reappraisal. Taxon. 1988; 37(3): 595–614.
  137. 137. Huber O, Zent S. Indigenous people and vegetation in the Venezuelan Guayana: Some ecological considerations. In: Heinen D, San José J, Caballero Arias H, editors. Nature and Human Ecology in the Neotropics. Scientia Guaianae 5. Caracas, Venezuela.1995. p. 37–64.
  138. 138. Navarro JC, Liria J, Piñango H, Barrera R. Biogeographic area relationships in Venezuela: A Parsimony analysis of Culicidae—Phytotelmata distribution in National Parks. Zootaxa. 2007; 1547(1): 19.
  139. 139. Duellman WE. Patterns of distribution of amphibians: a global perspective. Baltimore, MD: Johns Hopkins University Press; 1999.
  140. 140. Berry PE, Riina R. Insights into the diversity of the Pantepui flora and the biogeographic complexity of the Guayana Shield. Biologiske Skrifter. 2005; 55: 145–67.
  141. 141. McDiarmid RW, Donnelly MA. The herpetofauna of the Guayana Highlands: amphibians and reptiles of the Lost World. In: Donnelly MA, Crother BI, Guyer C, Wake MH, editors. Ecology and Evolution in the Tropics: A Herpetological Perspective. Chicago, Illinois: University of Chicago Press; 2005. p. 461–560.
  142. 142. Flinch JF, Rambaran V, Ali W, Lisa VD, Hernández G, Rodrigues K, et al. Chapter 17 Structure of the Gulf of Paria pull-apart basin (Eastern Venezuela-Trinidad). In: Mann P, editor. Sedimentary Basins of the World. 4: Elsevier; 1999. p. 477–94.
  143. 143. DeSalle R, Goldstein P. Review and Interpretation of Trends in DNA Barcoding. Front Ecol Evol. 2019; 7(302).
  144. 144. Dupuis JR, Roe AD, Sperling FA. Multi-locus species delimitation in closely related animals and fungi: one marker is not enough. Mol Ecol. 2012; 21(18): 4422–36. pmid:22891635.
  145. 145. Zink RM, Barrowclough GF. Mitochondrial DNA under siege in avian phylogeography. Mol Ecol. 2008; 17(9): 2107–21. pmid:18397219.
  146. 146. Hebert PD, Cywinska A, Ball SL, deWaard JR. Biological identifications through DNA barcodes. Proc Biol Sci. 2003; 270(1512): 313–21. pmid:12614582.
  147. 147. Meyer CP, Paulay G. DNA barcoding: error rates based on comprehensive sampling. PLoS Biol. 2005; 3(12): e422. pmid:16336051.
  148. 148. Cywinska A, Hunter FF, Hebert PD. Identifying Canadian mosquito species through DNA barcodes. Med Vet Entomol. 2006; 20(4): 413–24. pmid:17199753.
  149. 149. Teletchea F. After 7 years and 1000 citations: comparative assessment of the DNA barcoding and the DNA taxonomy proposals for taxonomists and non-taxonomists. Mitochondrial DNA. 2010; 21(6): 206–26. pmid:21171865.
  150. 150. Ahrens D, Fujisawa T, Krammer H-J, Eberle J, Fabrizi S, Vogler AP. Rarity and incomplete sampling in DNA-based species delimitation. Syst Biol. 2016; 65(3): 478–494. pmid:26797695
  151. 151. Phillips JD, Gillis DJ, Hanner RH. Incomplete estimates of genetic diversity within species: Implications for DNA barcoding. Ecol Evol. 2019; 9(5): 2996–3010. pmid:30891232.