Open access

Phylogeography, Vectors and Transmission in Latin America

Written By

Jan E. Conn, Martha L. Quiñones and Marinete M. Póvoa

Submitted: 10 February 2012 Published: 24 July 2013

DOI: 10.5772/55217

From the Edited Volume

Anopheles mosquitoes - New insights into malaria vectors

Edited by Sylvie Manguin

Chapter metrics overview

3,218 Chapter Downloads

View Full Metrics

1. Introduction

The overall focus of this chapter is the impact of phylogeographic studies on information pertinent to vector control, and an update on the relative importance and taxonomic status of five malaria vectors, some of which are species complexes, in the subgenus Nyssorhynchus: Anopheles albimanus Wiedmann, Albitarsis Complex, Anopheles aquasalis Curry, Anopheles darlingi Root, and Anopheles nuneztovari s.l. Gabaldón, considering literature predominantly since 2000. This cut-off date is to avoid repetition or overlap with some of the same subjects that have been covered in other places [1-4].

It is also of interest to vector control and elimination programs that, since 2000 and a more recent compilation that included a list of confirmed or potential Latin American malaria vectors [5], some vector species have been implicated in additional regions or countries by enzyme-linked immunosorbent assay [ELISA; 6], PCR techniques [7], VecTest [8] or more definitive biological and epidemiological evidence has been provided. Examples of these include An. rangeli Gabaldón, Cova Garcia and Lopez, initially implicated in Amapá state, Brazil [9] and subsequently in Putumayo, southern Colombia [10]; and An. triannulatus, (Neiva and Pinto) incriminated more broadly from Amazonian Brazil [11], then locally from Amapá, Brazil [12]. Furthermore, based on high frequency, biting behavior, seasonality, ELISA and nested-PCR, for the first time, An. rondoni (Neiva and Pinto) has been implicated in Matapá, Pará state, Brazil [13]. This is an understudied species, and its potential as a vector in other localities and regions in its distribution (Argentina, Bolivia, Brazil) is worth investigating.

A relatively early summary of information on the five most important malaria vectors in Latin America was published in 1986 [14]. This publication focused on four species in the Nyssorhynchus subgenus: Anopheles albimanus, Anopheles aquasalis, Anopheles darlingi, Anopheles nuneztovari and one in the Anopheles subgenus, Anopheles pseudopunctipennis Theobald. Naturally, more than 20 years later, this list of five is debatable, although most researchers would still consider An. darlingi to be the primary vector overall, and An. albimanus to be one of the most important. Nevertheless, some aspects of this publication are still relevant, and it serves as a useful historical introduction.

A review published in 2012 [15] summarized the overall findings of much of the available literature on genetic diversity of malaria vectors, including those in Latin America, and concluded that Pleistocene (0.01-2.6 mya; 16) environmental changes have been the primary drivers of divergence, at least at the species and population levels. These changes and earlier ones during the Miocene (2.6-5.3 mya)/Pliocene (5.3-23.0 mya) were hypothesized to have influenced the phylogeography of some co-distributed neotropical vector species, including Anopheles darlingi and selected Albitarsis Complex members [17].


2. Biology and vector status

2.1. An. albimanus

Throughout its broad, mostly coastal distribution (Figure 1), Anopheles albimanus is an important local vector and is considered to be ecologically adaptable (18). In general this species is crepuscular, zoophilic, exophagic, exophilic and seasonally abundant (19-20). Despite heteogeneity of several attributes, such as host-feeding behaviour, longevity, insecticide resistance and susceptibility to Plasmodium species, thoughout its distribution, it has maintained single species status (20).

Figure 1.

Distribution of Anopheles albimanus highlighted in green [21].

2.1.1. Colombia

An. albimanus is distributed in Colombia along the Atlantic and Pacific coasts (Figure 1). It is the main malaria vector on the Pacific coast, but its presence is considered a risk factor in other regions, even where malaria transmission is low [22]. Adult abundance of An. albimanus is associated with malaria transmission. The El Niño-Southern Oscillation Event (ENSO), that affects global climatic conditions every 2 to 7 years, has been strongly associated with increases in malaria cases, particularly in areas where An. albimanus is the main malaria vector, such as the Pacific coast [23]. An. albimanus breeding sites are very diverse, ranging from temporary small ponds to lagoons, and even include artificial containers. Its human biting rates can range between a few specimens per night night up to thousands, depending on the availablity of breeding sites in and around villages. Despite its considerable distribution on both coasts, An. albimanus has been found naturally infected with P. vivax only along the Pacific [24] (Table 1). In this region, its biting activity shows at least two peaks, one around midnight and a second one of less intensity before dawn, both indoors and outdoors [4]. Consistent use of insecticide-treated nets (ITNs) in this region could potentially reduce malaria transmission risk. In the Buenaventura peri-urban area, around 20% of the bites occur indoors and the main biting activity is outside houses between 18-21h. Then, at midnight, there is a second peak inside houses. Finally, between 05-06h, activity increases again outdoors [4].

Taxon Country Local transmission Regional transmission Evidence Reference
An. albimanus Colombia Pacific region ELISA, PCR 24
Albitarsis Complex* Brazil Marabá, Pará; Matapi River, Amapá; Amazon region; Macapá, Amapá; N. Amazon region, Roraima; Boa Vista, Roraima; Serra do Navio, Amapá Amazon region ELISA, PCR, VecTest 9; 11-13; 25-28; Póvoa 2010 (unpub. data)
Colombia Puerto Carreño, Vichada ELISA 29
Venezuela Sifontes, Bolivar ELISA 30-31
An. aquasalis Brazil São Luis, Maranhão; Belém, Pará ELISA 32-33
Guyana Mahdia VecTest 34
Suriname Brokobondo; Galibi; Paramaribo ELISA 35
Venezuela coastal areas 1
An. darlingi Brazil Marabá, Pará; Belém, Pará; Matapi River, Amapa; Macapá, Amapá; Boa Vista, Roraima; Serra do Navio, Amapá; Pará, N. Brazil; Anajás, Pará; Goianesia do Para, Pará Amazon region ELISA, PCR 9; 11-13; 25; 27-28; 33; 36-37; Póvoa 2012 (unpub. data)
Colombia Puerto Carreño, Vichada; Quibdó, Chocó; Dibulla, Guajira Córdoba; Villavicencio, Meta; Putumayo ELISA 4; 10; 29; 38-41
French Guiana Camopi; Saint Georges de l'Oyapock; Maroni River; Upper Maroni Amazonian forest; Village of Loca; Village of Twenke ELISA 42-45
Peru eastern region ELISA 46
Suriname Maroni River ELISA 43; 47
Venezuela Sifontes, Bolivar; Upper Orinoco River (southern) ELISA 30-31; 48
An. nuneztovari s.l. Brazil Matapi River, Amapa; Serra do Navio, Amapá; Anajás, Pará Amazon region ELISA 9; 11-12; 37
Colombia Tierralta, Córdoba ELISA 38
French Guiana Saint Georges de l'Oyapock ELISA 42
Venezuela Ocidente de Venezuela ELISA 49
*, Reference 27 refers to Anopheles albitarsis E (now An. janconnae) and Póvoa 2010 (unpublished data) refers to the first evidence of An. oryzalimnetes as a potential vector. References 12 and 25 refer to An. marajoara as a vector, while the remaining references refer to An. albitarsis s.l. as vectors.

Table 1.

Regional South American vectors subgenus Nyssorhynchus: evidence for malaria vector status.

2.1.2. Peru

An. albimanus in Peru is considered the main malaria vector along the Pacific coast, particularly in the north, where it is seasonal, and linked to agriculture [50]. Due to high insecticide application, mainly in rice fields, An. albimanus is resistant to all insecticides used in public health in this area [51]. Flooded rice fields provide ideal mosquito breeding habitat and An. albimanus density is associated with rice crops. Since 2005, the Peruvian Minister of Health, together with the Agricultural sector, implemented a modified irrigation system, so that the fields are dry for a week, and then intermittently irrigated, resulting in a decrease in mosquito larvae by 87% [52]. An important follow-up question would be whether this An. albimanus larval control has actually resulted in a decrease in malaria incidence rate (MIR), or in local An. albimanus adult female abundance, as measured by human biting rate (HBR) and entomological inoculation rate (EIR).

In several other South American countries (Ecuador, Panama, Venezuela) where An. albimanus is a malaria vector, as determined by sporozoite detection or other comparable information in earlier studies, data based on newer techniques are not available. However, some recent investigations have drawn attention to new distributions or larval habitat characterizations that pinpoint areas of fruitful potential research and possible targets for control measures [53-56].

2.2. Albitarsis Complex

Presently, there are eight recognized species (An. albitarsis s.s., An. albitarsis F, An. albitarsis G, An. albitarsis I, An. deaneorum, An. janconnae, An. marajoara, An. oryzalimnetes) and one lineage (An. albitarsis H) in the Albitarsis Complex [57]. The species described as near An. janconnae from Colombia [58] is now considered to be An. albitarsis I [57]. The overall distribution of members of this complex is wide-ranging, including both Central and South America, as well as some Caribbean Islands [59] (Figure 2).

Figure 2.

This map shows the predicted probability of occurrence of An. albitarsis in the Americas [2]

2.2.1. Brazil

Six species of the Albitarsis Complex are known from Brazil to date: An. albitarsis s.s., An. albitarsis G, An. deaneorum, An. janconnae, An. marajoara, and An. oryzalimnetes, [57,60] (Figure 3). The most broadly distributed member of this complex is An. marajoara Galvão and Damasceno [57,59]. It can be very abundant locally [12], and its breeding site types vary from swampy shores of lakes and ponds to small road puddles; it is generally associated with sunlight and often with aquatic or semi-aquatic vegetation [59]. It has been found infected by Plasmodium falciparum (Welsh), Plasmodium vivax (Grassi & Feletti) 210 and P. vivax 247, and Plasmodium malariae (Grassi & Feletti), and is a peri-urban as well as a rural vector, depending on locality, availability of breeding sites and hosts [12, 25]. It is also associated with deforested areas of the Vale do Ribeira in the southeastern Atlantic Forest of Brazil [61]. Although the EIR in Amapá state, Brazil was found to be lower than that of An. darlingi, it is an important local vector, at least in lowland rainforest in parts of the eastern Amazon [12, 62].

The distribution of An. janconnae Wilkerson and Sallum [classified previously as An. albitarsis E; [27, 60] appears to be limited to northern Amazonian Brazil, including along the Amazon River [57, 60]. Larval habitat types in several localities in Roraima state ranged from marsh to seepages to stream margins, and, based on analyses of several environmental variables, An. janconnae could be classified as a habitat specialist [McKeon, Conn & Povoa, unpublished data, 2012]. An. janconnae was incriminated as a local malaria vector around Boa Vista, the capital of Roraima state [27]. It is likely that the infected specimens identified as An. albitarsis s.l. from this region [26, 28] (Table 1), are An. janconnae, at least according to the geographic distribution [57].

An. oryzalimnetes (Wilkerson and Motoki), previously An. albitarsis B [60] has a broad distribution in Brazil that includes the Amazon region and southern Brazil [57]. It is frequently associated with rice fields, and is anthropophilic [60]. It was determined to be positive by ELISA for Plasmodium in Pará state [M.M. Povoa 2010, unpublished data], and may play a role in local transmission.

2.2.2. Colombia

At least three members of An. albitarsis s.l. are present in Colombia. Anopheles marajoara (some collections of which, according to the map [57], may be An. albitarsis I) is widely distributed [22], and its biology is similar to that described above under An. marajoara in Brazil. It is conisidered to be a regional vector in Colombia [22, 39, 63]. In the municipality of Puerto Carreno in eastern Colombia near the Venezuelan border, it was detected infected with P. falciparum [29] at a surprisingly high rate (1.92%; 3/152 specimens infected). Here, its peak biting time was 18-19h, with a minor peak from 20-21h, and it was collected both indoors and outdoors. It is suggested that together with An. darlingi, the dominant vector in the area, it is responsible for maintaining local malaria transmission in this municipality [29].

The second member of the complex is sympatric with An. darlingi in the east, and probably involved in malaria transmission. This species has been identified by various names, including An. allopha, An. marajoara, near An. janconnae [58], and most recently as An. albitarsis I [57]. Its known distribution thus far is restricted to Colombia. Relatively little is known about its biology, because of species identification issues, and there is no direct evidence yet for its involvement in malaria transmission. It appears that the specimens from Vichada, Colombia, identified as An. marajoara [64], are An. albitarsis I [57], so the distribution of An. marajoara in Colombia, and its involvement in malaria transmission, need re-evaluation.

The third species, An. albitarsis F, was first described from Puerto Carreño, near the Venezuelan border, [65]. In this locality it was found in sympatry with An. darlingi and a species in the Albitaris Complex now defined as An. albitarsis I. Its distribution is hypothesized to include Colombia, Venezuela and Trinidad [57]. Because it can easily be confused morphologically with An. marajoara, and it is found in regions of malaria endemnicity, this species is of some epidemiological importance.

2.2.3. Venezuela

It now appears that at least An. albitarsis I and possibly also An. albitarsis F are present is Venezuela [57]. Furthermore, An. albitarsis I could be sympatric in some regions with An. marajoara (the identification of which needs to be confirmed in Venezuela using molecular techniques), a local vector of P. vivax in western Venezuela [66]. An. marajoara also plays a significant role, together with An. darlingi, in malaria transmission in five localities in southern Venezuela [30-31]. In this gold mining region, where transmission is yearlong, the peak biting time, (19-21h, mostly before midnight), was comparable with most other reports of An. marajoara. Although An. marajoara was feeding both indoors and outdoors, it was significantly exophagic.

2.3. Anopheles aquasalis

This brackish-water breeder is found along the Pacific as far south as Ecuador and along the Atlantic to southern Brazil [1,14] (Figures 3,4). It is rarely found far from the ocean, but it can tolerate quite low salt ion concentrations, and has been detected in freshwater springs. It can be present in enormous numbers in marshy coastal areas, so that even if it is not extremely susceptible to Plasmodium, it can maintain malaria transmission when its abundance is high, especially during the rainy season [67]. It persists as an important local vector in Sucre state, eastern Venezuela, where a series of pioneering studies have identified hotspots of local transmission that are very useful for prevention and control efforts [68-69]. It has also been incriminated as a vector of P. falciparum and [or] P. vivax in Maranhão [32] and Pará states in Brazil [33, 70]. In Linden and Madia, Guyana [34], An. aquasalis was detected infected with P. vivax using VecTest, but the total sample size of anophelines collected was very small (n=45). In three towns in Suriname, Paramaribo, Brokopondo and Galibi, An. aquasalis was also detected infected with Plasmodium by ELISA [35], and is likely responsible for local, coastal transmission in this region.

2.4. Anopheles darlingi

The species considered to be the most important vector in the Amazon basin is Anopheles darlingi [2,3] (Figure 5). It is anthropophilic, adaptive and it has been incriminated in many localities in many countries, where it is often labeled a national vector (Table 1). As such, it has been the focus of a very wide range of research, monitoring and control efforts, and the publication for the first time of its complete genome is an exciting new development (GenBank accession number ADMH00000000).

2.4.1. Bolivia

An. darlingi is distributed in the northeastern Bolivian Amazon, in the departments of Pando, Beni and Santacruz de la Sierra, along the border with Brazil [71]. It shows a biting peak between 19-21h, with 83% of the bites occurring before 22h, when most local people go to bed. After this time, numbers decline, with little or no activity between 02:30-05h [72]. There have been relatively few studies on this species in Bolivia, and data are very scarce.

Figure 3.

South American localities where malaria vectors have been incriminated by various methods since the year 2000. Species codes: ALB, Anopheles albimanus; ALC, Albitarsis Complex; AQU, An. aquasalis; DAR, An. darlingi; NUN, An. nuneztovari. The darker grey area is Brazil.

Figure 4.

Distribution of Anopheles aquasalis highlighted in orange [21]

Figure 5.

Distribution of Anopheles darlingi highlighted in pink [21].

2.4.2. Brazil

Most of the newest incriminations of the continued involvement of An. darlingi in malaria transmission originate in Amazonian Brazil (Table 1, Figure 3). Rather than summarizing each new investigation, this section focuses on the findings on An. darlingi in a longitudinal study that investigated bloodmeal hosts, transmission, and seasonal abundance in three riverine villages along the Matapí River in Amapa state, northern Amazonian Brazil [12, 62, 73]. An ELISA analysis for IgG of common vertebrates found that the highest human blood indices (HBI) were in An. darlingi and An. marajoara. What was unexpected was that the HBIs of An. darlingi varied significantly among the three villages, which are only 1.5-7.0 km apart, likely because of host availability. It was found to be important to conduct a census of animals in each locality to be able to interpret the HBI results correctly. Even though An. darlingi was the most abundant species collected at human landing catches in each village, the HBI of An. darlingi resting collections, from under houses or in vegetation, ranged from 0.017-0.405, demonstrating how opportunistic this species can be, despite its anthropophily [12]. From the same study sites 113,117 mosquitoes collected from 2003-2005 were analyzed by ELISA. For this part of the study, An. darlingi and An. marajoara had the highest proportion of positives and also the highest EIRs, and thus the highest human-vector contact. Nevertheless, An. darlingi is still considered to be more important in this study area than An. marajoara because of its higher EIR [12]. Seasonal abundance was measured for 32 consecutive months of collection and showed that An. darlingi was most abundant during the wet-dry transition period between June and August, and that a strong positive correlation of An. darlingi abundance with rainfall lagged by several months. The latter finding may indicate that rainfall could be an important factor in predicting vector abundance, at least locally.

2.4.3. Colombia

The distribution of An. darlingi in Colombia is widespread but heterogeneous, and hypothesized to be interrupted by the Andes. It is found mainly south and east of the Andes, including the Amazon region, bordering Brazil and Peru, but also north and west of the Andes, along the main Colombian rivers (such as Magdalena, Cauca and Atrato) [22, 74]. An. darlingi is the main malaria vector throughout its distribution. Although associated with forest environments, it was also detected in the peri-urban area of the cities of Quibdó (Chocó) and Villavicencio (Meta), where malaria transmission occurs [40]. Breeding sites are the typical streams with slow water movement, but stationary water bodies such as natural and constructed fish ponds also provide good habitats. Similar to Peru (see below), the biting behaviour in Colombia is mostly before midnight, from 18-24h, with a smaller peak at sunrise (05-06h), but there is also persistant biting activity throughout the night [39].

2.4.4. French Guiana

In French Guiana, there has been renewed activity on malaria vectors, with most findings incriminating An. darlingi as the primary vector (Table 1). Between 2000-2002 in three Amerindian villages in the Upper-Maroni region of the Amazon forest, An. darlingi bit throughout the night, with peaks at 21:30-03:30h and again after 05:30h [44]. The biting rate was very high (255.5 bites/person/night) and specimens were infected with P. falciparum, P. vivax and P. malariae. Behavior was characterized as endo-exophagic and exophilic. The malaria transmission risk exists all year but probably it is greater during the rainy season when vectorial capacity was estimated to be higher [44]. A combination of ITNs and repellent is recommended; IRS is not efficient because of the housing materials and relative inaccessibility of this region. A second study in the Maroni area compared villages of Amerindian Wayanas and the Aloukous [45]. Significant findings include: the peak local malaria case reporting is the same timeframe (August to October) as the highest IMT (numbers of infected mosquitoes surviving long enough to transmit) of An. darlingi; the possibility that the persistent yearlong transmission is focused or perhaps limited to the Amerindian villages; and different bionomics of An. darlingi in the two villages which lead investigators to conclude that in this region there may exist two distinctive subspecies of An. darlingi. An analysis of collections from 2006-2011, in several regions of French Guiana, detected An. darlingi infected by P. vivax from Camopi and Saint Georges de l’Oyapock, both near the Oyapock River along the eastern border with Brazil [42].

2.4.5. Peru

An. darlingi is the main malaria vector species in eastern Amazonian Peru, the area with the highest malaria transmission in the country. It invaded this region in the 1990s [75], and its distribution now includes peri-urban settlements around the city of Iquitos, Loreto province. This change has been attributed to logging, agriculture and urban expansion, associated with deforestation [76-77]. To date, its greatest abundance is associated with areas of at least partial deforestation [78]. The main breeding sites in the Peruvian Amazon are streams and river margins in forested areas [77], however, the density of fish ponds has shown a positive association with malaria cases along roads in Loreto, suggesting that such ponds could be important local sources of this species [79]. An. darlingi was found naturally infected with P. vivax and P. falciparum in Loreto [46]. The human biting activity, which is similar indoors and outdoors, peaks two hours after sunset [77]. Because of the early evening biting peak, personal protection may be necessary to supplement bed-net use.

2.4.6. Panama

An. darlingi was recently detected in Panamá for the first time, in the eastern Darien region, near the border with Colombia, associated with the highest prevalence of drug-resistant P. falciparum [80]. This species was only collected by human landing catch, and not recorded in light traps, confirming its high anthropophily. Despite the extension and increased frequency of deforestation in Panama, An. darlingi has been detected only in the east. Its discovery suggests that unplanned deforestation should be avoided to prevent further expansion of this very anthropophilic species, and hence potential P. falciparum transmission, to other regions in the country.

2.4.7. Suriname

A timely and important new development in Suriname is the apparent collapse of populations of An. darlingi in the sparsely inhabited interior, in and around three study communities, correlated with two main factors: the introduction of ITNs and climatic events, i.e., unusual flooding which coincided with the beginning of the control activities in 2006 [47]. However, it should be noted that indoor residual spraying (IRS), active case detection (ACD), and a public awareness campaign also were implemented throughout the interior in 2006. As the authors point out, for Suriname, the next challenge is to try to find ways to use these methods to reduce or eliminate transmission among the gold-mining communities, where people are very mobile, and often active (not using ITNs) during potential biting times of An. darlingi. The latter are notoriously plastic, and vary locally and regionally. An important determination will be whether these results [47] can serve as a model for some communities where An. darlingi and malaria transmission are endemic in other countries.

2.4.8. Venezuela

Investigations along the Upper Orinoco River, southern Venezuela from 1994-1995 confirmed that An. darlingi was responsible for most, if not all of the local transmission of P. falciparum, P. vivax and P. malariae [48], that children under the age of 10 were at greatest risk, and that the EIR of An. darlingi was 129 positive bites/person/year. In a gold-mining region in southern Venezuela, studies from 1999-2000 [30-31] also determined that An. darlingi was one of two main vectors (the other was An. marajoara but see above under Albitarsis Complex). Surprisingly, many of the bionomic aspects of the two species in the five localities studied were quite similar (both more abundant during the rainy season, both biting indoors and outdoors with pronounced endophagic behavior), although An. marajoara was more abundant overall. The most striking bionomics difference between the two species was the peak biting time: An. darlingi bit throughout the night with two minor peaks (23-0h and 03-04h), whereas An. marajoara had a peak from 19-21h [30-31].

2.5. Anopheles nuneztovari s.l.

Anopheles nuneztovari s.l. is restricted to northern and Amazonian South America (Figure 6) and has been considered to be two genetically, ecologically and epidemiologically distinctive geographic populations, with the perception that the Colombian/western Venezuelan population was a regional vector (anthrophophilic and endo-exophagic) and the Amazonian population, mostly zoophilic and exophagic, was not [1, 5, 11]. Initially, evidence for malaria transmission by An. nuneztovari s.l. was found predominantly in Colombia and western Venezuela [22, 49]. However, a series of positive ELISA results and incriminations of malaria transmission involvement from localities in the Brazilian Amazon (Table 1; Figure 3) since 2000 soon undermined this relatively simple view. An. goeldii, which had been synonymized with An. nuneztovari, was resurrected as a valid species [81]. This work proposed different geographical distributions for each species, with An. goeldii in the Amazon region and An. nuneztovari more restricted to Colombia and Venezuela. The report of the discovery of An. nuneztovari infected with Plasmodium from Saint Georges de l'Oyapock, French Guiana, using results from a longitudinal study (2006-2011) is of at least local relevance, but it will be taxonomically important to determine whether this species is actually An. nuneztovari, or might possibly be An. goeldii, since susceptibility of An. goeldii to Plasmodium has not yet been tested [42].

An. nuneztovari s.l. is widely distributed in Colombia [22], particularly in the east, along the Venezuelan frontier, in the northwest region (Departments of Córdoba and Antioquia), where approximately 50% of the malaria cases occur, and in some areas along the Pacific Coast in the west, notably along the San Juan river (Chocó), and in the Buenaventura area (Valle). Specimens from Tierralta, Córdoba, a region of crop and livestock production, where An. nuneztovari s.l. was the most abundant species collected by human landing catches, were infected by P. vivax [38]. The breeding sites include small, permanent ponds, sunlit flooded pastures, and it has been determined that aquaculture ponds are one of this species’ most frequent breeding places. In the west, in Cimitarra (Santander), such ponds, characterized as permanent, completely exposed to sun and containing emerging vegetation, particularly grasses, represent approximately 81% of the breeding sites [63]. In Colombia, An. nuneztovari s.l. shows differing biting behavior by region. An exophagic tendency has been described in the northwest (Córdoba) [82], whereas in the east (Santander), a more endophagic behavior has been described [63]. The endophagic-exophilic variability makes control by residual insecticides very difficult.

Figure 6.

Distribution of Anopheles nuneztovari s.l. highlighted in blue [21].


3. Phylogeography

3.1. Anopheles albimanus

An exemplary study, based on large sample sizes using microsatellite markers and a mtDNA ND5 gene fragment, laid the groundwork for several Anopheles albimanus phylogeographic ideas [83]. These researchers detected restricted gene flow that they hypothesized to be the result of the physical barrier of the Central American Cordillera. Recent work, more geographically focused on one country or a region, with additional local sampling, provides additional insights into phylogeography in Central America [20, 85] and Colombia [84].

A mitochondrial DNA COI gene fragment and microsatellites were used to test for congruence with biogeographical provinces [86] in Colombia. In this case [84], one population, Turbo, was from Magdalena (Caribbean), three were from Maracaibo (Caribbean), and the four were from Choco (Pacific). The eight populations tested were clearly differentiated into two coastal regions, Caribbean and Pacific, with evidence for a late Pleistocene expansion (estimated to 21,994 years ago) or a selective sweep. Even though there was evidence for historical restrictions to gene flow (COI data), the microsatellites detected contemporary gene flow between the regions. Interestingly, a SAMOVA analysis found an unusual division. Only the three most easterly populations along the Caribbean coast grouped together. The fourth and most western Caribbean population, Turbo, was consistently more closely related to the four Pacific populations. Taken together, these data suggest possible semi-permeable boundaries among the three biogeographical provinces tested. Most relevant to malaria is the fact that the evidence for contemporary gene flow indicates that insecticide resistance genes, for example, could spread readily in these Colombian regions [84].

An. albimanus from Central America was examined using a fragment of the mtDNA COI gene to test the original hypothesis [83]. Physical barriers to gene flow were not detected (i.e., the Central America Cordillera was porous for An. albimanus) and contemporary isolation by distance was not supported [20]. Three divergent, co-occuring haplotype groups were detected using a statistical parsimony network, and these were not evenly distributed across Costa Rica and Panama. A new hypothesis suggested that they could be the result of multiple introductions into the region, probably caused by historical fragmentation and subsequent secondary contact. A more wide-ranging study incorporated the samples from Colombia [84], Ecuador, and Nicaragua with those from Costa Rica and Panama [20] and added two molecular fragments: the nuclear white gene and the ITS2. A SAMOVA analysis defined three large population demes, one from Nicaragua, Costa Rica and the Atlantic coast of western Panama; a second one incorporating the Pacific coast of western Panama, central-eastern Panama and the Caribbean Colombian coast; and a third one restricted to the Pacific coast of Colombia and Ecuador [85]. There were also four haplogroups, based on the COI fragment, which differed little from those found in the earlier Panamanian study [20] except for the addition of a fourth, restricted to the Pacific coast of Colombia and Ecuador, and separated by 18 mutation steps from its nearest haplogroup. Interestingly, because it tracks an earlier history, the white gene network showed much less divergence, supporting the overall conclusion that the primary time-frame for anopheline divergence at the species level is Pleistocene [15]. In summary, the combined An. albimanus data set strongly supported the presence of a single species in this region, which was expected, but also found very robust evidence for Pleistocene geographic fragmentation followed by range expansion across southern Central America [85].

3.2. The Albitarsis Complex

Following the newest revelations about the number of species (eight) plus a novel lineage (An. albitarsis H) in An. albitarsis s.l. [57], parts of an earlier study on the biogeography and population genetics of this complex [17] need to be reconsidered and modified. This is particularly the case for An. janconnae, which is more restricted than thought (under the taxonomic name of An. albitarsis E), the expanded distribution of An. albitarsis F (which now includes Venezuela and Trinidad as well as Colombia), the complexities of the distribution of An. marajoara, which really may have a very broad range, newly described An. albitarsis G, distributed along the Brazilian Amazon, and An. albitarsis I, restricted to northwestern Colombia [57]. Despite these problems, one recent study can be used to illustrate the phylogeography of at least An. albitarsis G [as far as it is known; 57] and part of the range of An. marajoara [87]. An. albitarsis G [lineage 2 in reference 87] may be restricted to localities near the Amazon River or its tributaries. It has little population structure and the small subdivisions that were detected in haplotype networks were unrelated to geographic locality. The evidence from the mtDNA COI fragment used in this study indicates that this lineage is older than An. marajoara [lineage 1 in reference 87]. On the other hand, the white gene and ITS2 data detected a single network between An. albitarsis G and An. marajoara, indicating that the divergence is recent. The most compelling result in this study concerning An. marajoara is that SAMOVA defined two population demes along the Amazon River, splitting this species into western and eastern entities with differing genetic characteristics. The boundary is located near Rio Jari in Amapá state, not far from one detected in An. darlingi [88] and an earlier one seen in a study of An. nuneztovari s.l. using restriction fragment length analysis of the mtDNA genome [89]. For An. marajoara, this boundary is permeable, since there were shared haplotypes on either side [87]. A denser sampling of all three species could more rigorously test whether this is the result of underlying geological boundaries or perhaps more recent climatic events.

3.3. An. darlingi

Several studies on the phylogeography of An. darlingi have been undertaken. The earliest one [90] used the mtDNA COI fragment and detected a significant genetic division between Central America/northwestern Colombia, and the rest of South America. According to the statistical parsimony network, the more widespread and ancestral haplotypes were in Amazonian and southern South America, suggesting that the Central American/Colombian haplotypes may have originated there. This division was also supported by sequences of the white gene, which found two genotypes, genotype I, restricted to the Amazon, and genotype II, in northwestern Colombia and Venezuela, and Central America [91]. A microsatellite analysis of 1,376 samples also strongly supported the initial COI genetic division, and found substantial structure within the Amazon Basin [91]. The conclusion was that there were two main drivers for this division: differences in effective population size among the divisions, and physical distances between the populations. A more sophisticated analysis of the mtDNA COI fragment included additional Brazilian samples and excluded the Central American samples [88]. These researchers detected six main population groups in South America, and found ancestral distribution to be central Amazonia. They proposed that populations became isolated by three barriers: the Amazon River, the Andes and the southeastern Brazilian coastal ranges. They also found that limited dispersal across some landscape types has promoted differentiation between other proximate populations. A local study of An. darlingi in Córdoba and Antioquia, Colombia, using mtDNA COI, microsatellites and the white gene [74] supported the earlier geographic hypothesis [90], discovering that the five populations tested were more closely related to the Central American populations of An. darlngi that they were to South American An. darlingi. Because of local high gene flow among the five populations, similar control strategies could be implemented in these two contiguous Colombian states. Similarly, newly detected An. darlingi from Panama were most closely related to Colombian and Central American An. darlingi [80]. Concordant phylogeographies were determined for the two neotropical vectors An. darlingi and An. triannulatus [92]. With the mtDNA COI fragment, SAMOVA detected four similar population subdivisions: one in southern coastal Brazil, two in central Brazil and one northeast of the Amazon. Both species originated south of the Amazon River and seem to have followed a similar expansion pathway to their present-day distributions. Other neotropical anophelines with similar distributions may share a common spatial and demographic history with these species, and remain to be evaluated.

3.4. An. aquasalis

The only study that attempted to analyse An. aquasalis within a phylogeographic framework was conducted using a fragment of the mtDNA COI gene with specimens from five localities on either side of the Amazon, in Amapá and Pará states, Brazil [93]. The most important findings from this study inferred that despite the width of the mouth of the Amazon, this freshwater delta was not a barrier for the salt-water tolerant An. aquasalis, likely because of so much tidal mixing, and the numerous islands and channels in the region. However, gene flow was restricted, based on isolation by distance that was detected using a Nested Clade Analysis [94]. The relative regional importance of An. aquasalis as a malaria vector has waned since the earlier publications [1, 14], so there may be fewer opportunities to pursue phylogeographic questions, especially because the distribution is relatively limited. However, no one has compared specimens from the Atlantic and Pacific coasts, and it is possible that population structure similar to that found for An. albimanus [85], could be detected in An. aquasalis, considering that both species share a relatively narrow coastal distribution in South America, and were subjected to the same kinds of Pleistocene environmental changes.

3.5. An. nuneztovari s.l.

The revision of the taxonomic status of An. nuneztovari s.l., that now includes An. nuneztovari s.s. and An. goeldii [81] has implications for the interpretation of the first study of An. nuneztovari phylogeography, which focused on the nuclear white gene [95]. Five lineages were detected [95], 2 and 3 in Colombia/Venezuela and 1, 4 and 5 in Amazonian Brazil. The earliest divergence, during the Pliocene (5.3-23.0 mya), is between Colombia and Venezuela west of the Andes (lineage 3) and Amazonian lineage 4. The most likely hypothesis to explain this divergence is an early uplift of the East Andean Cordillera [96]. Curiously, the levels of genetic divergences among the five lineages were high, although the minimum spanning network of the haplotypes connected all of them. There were five localities where two lineages were sympatric: in Brazil - Boa Vista, Roraima state; Altamira, Pará state and near Pôrto Velho, Rondonia state; Guayaramerín, Beni, Bolivia; and Rio Socuavó, Zulia, Venezuela. These localities are of special interest, since they may be admixture zones or hotspots of divergence. The simplest hypothesis to explain the five lineages taxonomically is that the two in Colombia/Venezuela are An. nuneztovari s.s. and the three in Brazil are An. goeldii. The sharing of haplotypes across the Andes, between eastern and western Venezuela [95], is congruent with and supports findings for An. albimanus [83], The Albitarsis Complex [57] and An. darlingi (Conn, unpublished data) that have hypothesized that the eastern Andean Cordillera is only a partial barrier for anopheline mosquitoes.

A second phylogeographic study was undertaken with some of the same samples, plus new ones from Amazonian Brazil, using a mtDNA COI fragment [97], which charts a more recent history of divergence, all within the Pleistocene, compared with the white gene fragment. In this work, there were two major monophyletic clades, I and II. Specimens from Bolivia/Colombia/Venezuela represent the most basal subclade, IIC; whereas the Amazonian specimens were found in clades I and II-A and II-B. There were also several localities of sympatry among the clades: five in Amapá, Amazonas and Pará states, Amazonian Brazil, and one in Suriname. None of these are the same as the ones detected by the white gene study, perhaps suggesting that these were later areas of sympatry. There was an intriguing connection detected between the specimens from Colombia/Venezuela and those from Amazonian Bolivia, which had previously been seen when sequences of the rDNA ITS2 were used [98]. This may be the signature of the marine incursion hypothesis [95]. One of clades I, II-A or II-B likely represents An. goeldii, but additional analyses are needed to determine which one, and also to test the hypothesis of multiple species in the Amazon.


4. Conclusions

There have been many changes in the incrimination, identification and several new insights into the phylogeography of the species discussed in this chapter. The most important taxonomic changes are those in the Albitarsis Complex, with the discovery of two new species, An. albitarsis G and I, and a new lineage, An. albitarsis H. Nothing is known about their involvement in malaria transmission, although their ranges all include malaria endemic areas, or their local contribution to diversity or to a better understanding of the complex patterns of Amazonian biogeography and phylogeography. The relative paucity of new work on An. aquasalis is a reminder that its relative importance appears to be lessening, although it is still likely to be important locally, particularly when in high abundance. Obviously, the importance of An. darlingi in still on the rise in several localities in many countries, attributable mainly to its remarkable adaptability and association with landscape changes. The resurrection of An. goeldii from synonymy is also a milestone, because it provides a first step toward resolving a longstanding discussion about the possible importance of An. nuneztovari s.l. in local transmission in Amazonian Brazil. It may also clarify some aspects of the recent phylogeographic inferences based on white and the mtDNA COI genes. Lastly, the detection of concordant phylogeographies, one of which is An. darlingi in Brazil, depict a clear path towards future research which will have important epidemiological consequences.



We thank Ricardo Guimarães (LabGeo/IEC/SVS) for creating the map in Figure 3. We thank Sara Bickersmith for tireless work on the table, editorial suggestions, and slight modifications to the map. Some of the unpublished work cited here was funded by a grant from the National Institutes of Health (USA) to JEC (AI R01 54139-02) and a grant from CNPq (Brazil) to MMP.


  1. 1. Lounibos LP, Conn JE. Malaria vector heterogeneity in South America. American Entomologist 2000;46 238-249.
  2. 2. Sinka ME, Rubio-Palis Y, Manguin S, Patil AP, Temperley WH, Gething PW, Van Boeckel T, Kabaria CW, Harbach RE, Hay SI. The dominant Anopheles vectors of human malaria in the Americas: occurrence data, distribution maps and bionomic précis. Parasites & Vectors 2010;3 72.
  3. 3. Hiwat H, Bretas G. Ecology of Anopheles darlingi Root with respect to vector importance: a review. Parasites & Vectors 2011;4 177.
  4. 4. Montoya-Lerma J, Solarte YA, Giraldo-Calderón GI, Quiñones ML, Ruiz-López F, Wilkerson RC, González R. Malaria vector species in Colombia: a review. Memórias do Instituto Oswaldo Cruz 2011;106: 223-238.
  5. 5. Marrelli MT, Sallum MAM, Marinotti O. The second internal transcribed spacer of nuclear ribosomal DNA as a tool for Latin American anopheline taxonomy – a critical review. Memórias do Instituto Oswaldo Cruz 2006;101(8) 817-832.
  6. 6. Wirtz RA, Burkot TR, Graves PM, Andre RG. Field evaluation of enzyme-linked immunosorbent assays for Plasmodium falciparum and Plasmodium vivax sporozoites in mosquitoes (Diptera: Culicidae) from Papua New Guinea. Journal of Medical Entomology 1987;24(4) 433-437.
  7. 7. Snounou G, Viriyakosol S, Jarra W, Thaithong S, Brown KN. Identification of the four human malaria parasite species in field samples by the polymerase chain reaction and detection of a high prevalence of mixed infections. Molecular and Biochemical Parasitology 1993;58(2) 283-292.
  8. 8. Ryan JR, Davé K, Collins KM, Hochberg L, Sattabongkot J, Coleman RE, Dunton RF, Bangs MJ, Mbogo CM, Cooper RD, Schoeler GB, Rubio-Palis Y, Magris M, Romer LI, Padilla N, Quakyi IA, Bigoga J, Leke RG, Akinpelu O, Evans B, Walsey M, Patterson P, Wirtz RA, Chan AS. Extensive multiple test center evaluation of the VecTest malaria antigen panel assay. Medical and Veterinary Entomology 2002;17 321-332.
  9. 9. Póvoa MM, Wirtz RA, Lacerda RN, Miles MA, Warhurst D. Malaria vectors in the municipality of Serra do Navio, State of Amapá, Amazon Region, Brazil. Memorias do Instituto Oswaldo Cruz 2001;96(2) 179-184
  10. 10. Quiñones ML, Ruiz F, Calle DA, Harbach RE, Erazo HF, Linton YM. 2006. Incrimination of Anopheles (Nyssorhynchus) rangeli and An. (Nys.) oswaldoi as natural vectors of Plasmodium vivax in Southern Colombia. Memórias do Instituto Oswaldo Cruz 2006;101 617-623.
  11. 11. Tadei WP, Dutary Thatcher B. Malaria vectors in the Brazilian amazon: Anopheles of the subgenus Nyssorhynchus. Revista do Instituto de Medicina Tropical de São Paulo 2000;42(2) 87-94.
  12. 12. Galardo AK, Arruda M, D’Almeida Couto AA, Wirtz R, Lounibos LP, Zimmerman RH. Malaria vector incrimination in three rural riverine villages in the Brazilian Amazon. American Journal of Tropical Medicine and Hygiene 2007;76(3) 461-469.
  13. 13. da Rocha JAM, de Oliveira SB, Póvoa MM, Moreira LA, Krettli AU. Malaria vectors in areas of Plasmodium falciparum epidemic transmission in the Amazon region, Brazil. American Journal of Tropical Medicine and Hygiene 2008;78(6) 872-877.
  14. 14. Fleming G. Biologia y ecologia de los vectores de la malaria en las Americas. PAHO/WHO, 1986; PNSP-86-72, Washington, 54 pp.
  15. 15. Loaiza JR, Bermingham E, Sanjur OI, Scott ME, Bickersmith SA, Conn JE. Review of genetic diversity in malaria vectors (Culicidae: Anophelinae). Infection, Genetics, and Evolution 2012;12(1) 1-12.
  16. 16. Gibbard PL, Head MJ. IUGS ratification of the Quaternary System/Period and the Pleistocene Series/Epoch with a base at 2.58 Ma. Quaternaire 2009; 20 271-272.
  17. 17. Conn JE, Mirabello LM. The biogeography and population genetics of neotropical vector species. Heredity 2007;99 245-256.
  18. 18. Faran ME. Mosquito studies (Diptera: Culicidae) XXXIV. A revision of the Albimanus section of the subgenus Nyssorhynchus of Anopheles. Contributions of the Annals of the Entomological Institute 1980;15 1-215.
  19. 19. Breeland SG. Studies on the ecology of Anopheles albimanus. American Journal of Tropical Medicine and Hygiene 1972;21 751-754.
  20. 20. Loaiza JR, Scott ME, Bermingham E, Rovira J, Conn JE. Evidence for Pleistocene population divergence and expansion of Anopheles albimanus in Southern Central America. American Journal of Tropical Medicine and Hygiene 2010;82(1) 156-164.
  21. 21. Manguin S, Carnevale P, Mouchet J, Coosemans M, Julvez J, Richard-Lenoble D, Sircoulon J. Biodiversity of malaria in the world. France: John Libbey Eurotext Ed.; 2008.
  22. 22. Olano VA, Brochero H, Saenz R, Quinones M. Molina J. Mapas preliminares de la distribucion de especies de Anopheles vectores de malaria en Colombia. Biomedica 2001; 21 402–408.
  23. 23. Poveda G, Rojas W, Quiñones ML, Velez IV, Mantilla RI, Ruiz D, Zuluaga JS, Rua GL. Coupling between Annual and ENSO Timescales in the Malaria - Climate Association in Colombia. Environmental Health Perspectives 2001;109(5) 489-493.
  24. 24. Gutiérrez LA, Naranjo N, Jaramillo LM, Muskus C, Luckart S, Conn JE, Correa MM. Natural infectivity of Anopheles species from the Pacific and Atlantic regions of Colombia. Acta Tropica 2008;107(2) 99-105.
  25. 25. Conn JE, Wilkerson RC, Segura MN, de Souza RT, Schlichting CD, Wirtz RA, Póvoa MM. Emergence of a new neotropical malaria vector facilitated by human migration and changes in land use. American Journal of Tropical Medicine and Hygiene 2002;66(1) 18-22.
  26. 26. de Barros FS, Honório NA, Arruda ME. Mosquito anthropophily: implications on malaria transmission in the Northern Brazilian Amazon. Neotropical Entomology 2010;39(6) 1039-1043.
  27. 27. Póvoa MM, de Souza RT, Lacerda RN, Rosa ES, Galiza D, de Souza JR, Wirtz RA, Schlichting CD, Conn JE. The importance of Anopheles albitarsis E and An. darlingi in human malaria transmission in Boa Vista, state of Roraima, Brazil. Memórias do Instituto Oswaldo Cruz 2006;101(2) 163-168.
  28. 28. da Silva-Vasconcelos A, Kató MY, Mourão EN, de Souza RT, Lacerda RN, Sibajev A, Tsouris P, Póvoa MM, Momen H, Rosa-Freitas MG. Biting indices, host-seeking activity and natural infection rates of anopheline species in Boa Vista, Roraima, Brazil from 1996 to 1998. Memórias do Instituto Oswaldo Cruz 2002;97(2) 151-161.
  29. 29. Jiménez P, Conn JE, Wirtz R, Brochero H. Anopheles (Diptera: Culicidae) vectores de malaria en el municipio de Puerto Carreño, Vichada, Colombia. Biomedica 2012;32 13-21.
  30. 30. Moreno JE, Rubio-Palis Y, Páez E, Pérez E, Sánchez V, Vaccari E. Malaria entomological inoculation rates in gold mining areas of Southern Venezuela. Memorias do Instituto Oswaldo Cruz 2009;104(5) 764-768.
  31. 31. Moreno JE, Rubio-Palis Y, Páez E, Pérez E, Sánchez V. Abundance, biting behavior and parous rate of anopheline mosquito species in relation to malaria incidence in gold-mining areas of southern Venezuela. Medical and Veterinary Entomology 2007;21(4) 339-349.
  32. 32. Ribeiro MCT, Goncalves EDD, Tauil PL, da Silva AR. Epidemiological aspects of a malaria focus in the districts of São Luis, MA. Revista da Sociedade Brasileira de Medicina Tropical 2005;38(3) 272-274.
  33. 33. Póvoa MM, Conn JE, Schlichting CD, Amaral JC, Segura MN, da Silva AN, dos Santos CC, Lacerda RN, de Souza RT, Galiza D, Santa Rosa EP, Wirtz RA. Malaria vectors, epidemiology, and the re-emergence of Anopheles darlingi in Belém, Pará, Brazil. Journal of Medical Entomology 2003;40(4) 379-386.
  34. 34. Laubach HE, Validum L, Bonilla JA, Agar A, Cummings R, Mitchell C, Cuadrado RR, Palmer CJ. Identification of Anopheles aquasalis as a possible vector of malaria in Guyana, South America. West Indian Medical Journal 2001;50(4) 319-21.
  35. 35. Póvoa MM. 2002. Report on Malaria transmission in Suriname. 5p
  36. 36. dos Santos RL, Padilha A, Costa MD, Costa EM, Dantas-Filho Hde C, Póvoa MM. Malaria vectors in two indigenous reserves of the Brazilian Amazon. Revista de Saúde Pública 2009;43(5) 859-868.
  37. 37. dos Santos RL, Sucupira IM, Lacerda RN, Fayal A da S, Póvoa MM. Entomological survey and infectivity during malaria outbreak in the Anajás municipality, Pará State. Revista da Sociedade Brasileira de Medicina Tropical 2005;38(2) 202-204.
  38. 38. Gutiérrez LA, González JJ, Gómez GF, Castro MI, Rosero DA, Luckart S, Conn JE, Correa MM. Species composition and natural infectivity of anthropophilic Anopheles (Diptera: Culicidae) in the states of Córdoba and Antioquia, Northwestern Colombia. Memórias do Instituto Oswaldo Cruz 2009;104(8) 1117-1124.
  39. 39. Brochero HL, Rey G, Buitrago LS, Olano VA. Biting activity and breeding sites of Anopheles species in the municipality Villavicencio, Meta, Colombia. Journal of the American Mosquito Control Association 2005;21(2) 182-186.
  40. 40. Ochoa J, Osorio L. Epidemiologia de malaria urbana en Quibdó, Chocó. Biomedica 2006;26 278-285.
  41. 41. Herrera M, Orjuela LI, Peñalver C, Quiñones ML. Characterization of the Anopheles species present in two ecologically different regions in La Guajira, northern Colombia. In Clark G & Rubio-Palis. Mosquito Vector Biology and Control in Latin America - a 19th Symposium. Journal of the American Mosquito Control Association 2009;25(4):486-499.
  42. 42. Dusfour I, Issaly J, Carinci R, Gaborit P, Girod R. Incrimination of Anopheles (Anopheles) intermedius Peryassú, An. (Nyssorhynchus) nuneztovari Gabaldón, An. (Nys.) oswaldoi Peryassú as natural vectors of Plasmodium falciparum in French Guiana. Memórias do Instituto Oswaldo Cruz 2012;107(3) 429-432.
  43. 43. Hiwat H, Issaly J, Gaborit P, Somai A, Samjhawan A, Sardjoe P, Soekhoe T, Girod R. Behavioral heterogeneity of Anopheles darlingi (Diptera: Culicidae) and malaria transmission dynamics along the Maroni River, Suriname, French Guiana. Transactions of the Royal Society of Tropical Medicine and Hygiene 2010;104(3) 207-213.
  44. 44. Girod R, Gaborit P, Carinci R, Issaly J, Fouque F. Anopheles darlingi bionomics and transmission of Plasmodium falciparum, Plasmodium vivax, and Plasmodium malariae in American villages of the Upper-Maroni Amazonian forest, French Guiana. Memórias do Instituto Oswaldo Cruz 2008;103(7) 702-710.
  45. 45. Fouque F, Gaborit P, Carinci R, Issaly J, Girod R. 2010. Annual variations in the number of malaria cases related to two different patterns of Anopheles darlingi transmission potential in the Maroni area of French Guiana. Malaria Journal 2010;9 80.
  46. 46. Flores-Mendoza C, Fernández R, Escobedo-Vargas KS, Vela-Perez Q, Schoeler GB. Natural Plasmodium infections in Anopheles darlingi and Anopheles benarrochi (Diptera: Culicidae) from Eastern Peru. Journal of Medical Entomology 2004; 41(3) 489-494.
  47. 47. Hiwat H, Mitro S, Samjhawan A, Sardjoe P, Soekhoe T, Takken W. Collapse of Anopheles darlingi populations in Suriname after introduction of insecticide-treated nets (ITNs); malaria down to near elimination level. American Journal of Tropical Medicine and Hygiene 2012;86(4) 649-655.
  48. 48. Magris M, Rubio-Palis Y, Menares C, Villegas L. Vector bionomics and malaria transmission in the Upper Orinoco River, Southern Venezuela. Mem Inst Oswaldo Cruz 2007;102(3) 303-311.
  49. 49. Rubio-Palis Y. 2000. Anopheles (Nyssorhynchus) de Venezuela: taxonomia, bionomía, ecología e importancia médica. Escuela de Malariología y Saneamiento Ambiental “Dr. Arnoldo Gabaldon” y Proyecto Control de Enfermedades Endémicas, Maracay, Venezuela.
  50. 50. Guthmann JP, Hall AJ, Jaffar S, Palacios A, Lines J, Llanos-Cuentas A. Environmental risk factors for clinical malaria: a case-control study in the Grau region of Peru. Transactions of the Royal Society of Tropical Medicine and Hygiene 2001;95(6) 577-583.
  51. 51. Vargas F, Córdova O, Alvardo A. Determinación de la Resistencia a insecticidas en Aedes aegypti, Anopheles albimanus y Lutzomyia peruensis procedentes del Norte Peruano. Revista Peruana de Medicina Experimental y Salud Publica 2006;23(4) 259-264.
  52. 52. DIGESA-Direccion General de Salud Ambiental, Ministerio de Salud de Perú. Plan de Implementación de la Estrategia de Riego con Secas Intermitentes en el Cultivo de Arroz para el Control Vectorial de la Malaria en Regiones Priorizadas del Perú. Lima, Perú, 2011. 33p.
  53. 53. Loaiza JR, Bermingham E, Scott ME, Rovira JR, Conn JE. Species composition and distribution of adult Anopheles (Diptera: Culicidae) in Panama. Journal of Medical Entomology 2008;45(5) 841-851.
  54. 54. Pinault LL, Hunter FF. New highland distribution records of multiple Anopheles species in the Ecuadorian Andes. Malaria Journal 2011;11(10) 236.
  55. 55. Pinault LL, Hunter FF. Larval habitat associations with human land uses, roads, rivers, and land cover for Anopheles albimanus, A. pseudopunctipennis, and A. punctimacula (Diptera: Culicidae) in coastal and highland Ecuador. Frontiers in Physiology 2012;3 59.
  56. 56. Pinault LL, Hunter FF. Characterization of larval habitats of Anopheles albimanus, Anopheles pseudopunctipennis, Anopheles punctimacula, and Anopheles oswaldoi s.l. populations in lowland and highland Ecuador. Journal of Vector Ecology 2012;37(1) 124-136.
  57. 57. Ruiz-Lopez F, Wilkerson RC, Conn JE, McKeon SN, Levin DM, Quiñones ML, Póvoa MM, Linton YM. DNA barcoding reveals both known and novel taxa in the Albitarsis Group (Anopheles: Nyssorhynchus) of Neotropical malaria vectors. Parasites & Vectors 2012;5:44.
  58. 58. Gutiérrez LA, Orrego LM, Gómez GF, López A, Luckart S, Conn JE, Correa MM. A new mtDNA COI gene lineage closely related to Anopheles janconnae of the Albitarsis complex in the Caribbean region of Colombia. Memórias do Instituto Oswaldo Cruz 2010;105 1019-1025.
  59. 59. Linthicum KJ. A revision of the Argyritarsis Section of the subgenus Nyssorhynchus of Anopheles (Diptera: Culicidae). Mosquito Systematics 1988;20(2) 98-271.
  60. 60. 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). Memórias do Instituto Oswaldo Cruz 2009;104(6) 823-850.
  61. 61. Laporta GZ, Ramos DG, Ribeiro MC, Sallum MA. Habitat suitability of Anopheles vector species and association with human malaria in the Atlantic Forest in south-eastern Brazil. Memórias do Instituto Oswaldo Cruz 2011;106 Suppl 1:239-45.
  62. 62. Galardo AK, Zimmerman RH, Lounibos LP, Young LJ, Galardo CD, Arruda M, D’Almeida Couto AA. Seasonal abundance of anopheline mosquitoes and their association with rainfall and malaria along the Matapí River, Amapí, Brazil. Medical and Veterinary Entomology 2009;23 335-349.
  63. 63. Brochero H, Pareja PX, Ortiz G, Olano VA. Sitios de cría y actividad de picadura de especies de Anopheles en el municipio de Cimitarra, Santander, Colombia. Biomedica 2006;26(2) 269-277.
  64. 64. Brochero H, Li C, Wilkerson RC, Conn JE, Ruiz-García. Genetic structure of Anopheles (Nyssorhynchus) marajoara (Diptera: Culicidae) in Colombia. American Journal of Tropical Medicine and Hygiene 2010;83(3) 585-595.
  65. 65. Brochero HL, Li C, Wilkerson RC. A newly recognized species in the Anopheles (Nyssorhynchus) albitarsis complex (Diptera: Culicidae) from Puerto Carreño, Colombia. American Journal of Tropical Medicine and Hygiene 2007;76(6) 1113-1117.
  66. 66. Rubio-Palis Y, Wilkerson R, Guzman H. Morphological characters of adult Anopheles (Nyssorhynchus) marajoara in Venezuela. Journal of the American Mosquito Control Association 2003;19(2)107-114.
  67. 67. Forattini OP. Culicidologia médica: identificação, biologia, epidemiologia, Vol. II., Edusp, São Paulo, 2002. 864 pp.
  68. 68. Grillet ME, Eudes Martínez JE, Barrera R. Malaria hot spot areas: implications for effective and targeted interventions in Venezuela. Boletin de Malariología y Salud Ambiental 2009;49(2) 193-208.
  69. 69. Grillet ME, Barrera R, Martínez JE, Berti J, Fortin MJ. Disentangling the effect of local and global spatial variation on a mosquito-borne infection in a neotropical heterogeneous environment. American Journal of Tropical Medicine and Hygiene 2010;82(2) 194-201.
  70. 70. da Silva Ade N, Fraiha-Neto H, dos Santos CC, Segura Mde N, Amaral JC, Gorayeb Ide S, Lacerda RN, Sucupira IM, Pimentel LN, Conn JE, Póvoa MM. Anophelines in Belém, Pará, Brazil: current and retrospective data. Cadernos de Saude Publica 2006;22(8) 1575-1585.
  71. 71. Lardeux F, Chávez T, Rodríguez R, Torrez L. Anopheles of Bolivia: new records with an updated and annotated checklist. Comptes Rendus Biologies 2009;332(5) 489-499.
  72. 72. Harris AF, Matias-Arnéz A, Hill N. Biting time of Anopheles darlingi in the Bolivian Amazon and implications for control of malaria. Transactions of the Royal Society of Tropical Medicine and Hygiene 2006;100(1) 45-47.
  73. 73. Zimmerman RH, Galardo AK, Lounibos LP, Arruda M, Wirtz R. Bloodmeal hosts of Anopheles species (Diptera: Culicidae) in a malaria-endemic area of the Brazilian Amazon. Journal of Medical Entomology 2006;43(5) 947-956.
  74. 74. Gutiérrez LA, Gómez GF, González JJ, Castro MI, Luckart S, Conn JE, Correa MM. Microgeographic genetic variation of the malaria vector Anopheles darlingi Root (Diptera: Culicidae) from Córdoba and Antioquia, Colombia. American Journal of Tropical Medicine and Hygiene 2010;83(1) 38-47.
  75. 75. Aramburu Guarda J, Ramal Asayag C, Witzig R. Malaria reemergence in the Peruvian Amazon region. Emerging Infectious Diseases 1999;5 209-215.
  76. 76. Turell MJ, Sardelis MR, Jones JW, Watts DM, Fernandez R, Carbajal F, Pecor JE, Klein TA. Seasonal distribution, biology, and human attraction patterns of mosquitoes (Diptera: Culicidae) in a rural village and adjacent forested site near Iquitos, Peru. Journal of Medical Entomology 2008;45(6) 1165-1172.
  77. 77. Reinbold-Wasson DD, Sardelis MR, Jones JW, Watts DM, Fernandez R, Carbajal F, Pecor JE, Calampa C, Klein TA, Turell MJ. Determinants of Anopheles seasonal distribution patterns across a forest to periurban gradient near Iquitos, Peru. American Journal of Tropical Medicine and Hygiene 2012;86(3) 459-463.
  78. 78. Vittor AY, Gilman RH, Tielsch J, Glass G, Shields T, Lozano WS, Pinedo-Cancino V, Patz JA. The effect of deforestation on the human-biting rate of Anopheles darlingi, the primary vector of Falciparum malaria in the Peruvian Amazon. American Journal of Tropical Medicine and Hygiene 2006:74(1) 3-11.
  79. 79. Maheu-Giroux M, Casapía M, Soto-Calle VE, Ford LB, Buckeridge DL, Coomes OT, Gyorkos TW. Risk of malaria transmission from fish ponds in the Peruvian Amazon. Acta Tropica. 2010 115(1-2);112-118.
  80. 80. Loaiza J, Scott M, Bermingham E, Rovira J, Sanjur O, Conn JE. Anopheles darlingi (Diptera: Culicidae) in Panama. American Journal of Tropical Medicine and Hygiene 2009;81(1) 23-26.
  81. 81. Calado DC, Foster PG, Bergo ES, dos Santos CLS, Galardo AKR, Sallum MAM. Resurrection of Anopheles goeldii from synonymy with Anopheles nuneztovari (Diptera, Culicidae) and a new record for Anopheles dunhami in the Brazilian Amazon. Memórias do Instituto Oswaldo Cruz 2008;103(8) 791-799.
  82. 82. Parra-Henao G, Alarcón-Pineda EP. Observaciones sobre la bionomía de Anopheles spp. (Diptera: Culicidae) en el municipio Valencia, departamento Córdoba, Colombia. Boletín de Malariologia y Saneamiento Ambiental 2008;48 95-98.
  83. 83. Molina-Cruz A, de Mérida AM, Mills K, Rodríguez F, Schoua C, Yurrita MM, Molina E, Palmieri M, Black WC 4th. Gene flow among Anopheles albimanus populations in Central America, South America, and the Caribbean assessed by microsatellites and mitochondrial DNA. American Journal of Tropical Medicine and Hygiene 2004;71(3) 350-359.
  84. 84. Gutiérrez LA, Naranjo NJ, Cienfuegos AV, Muskus CE, Luckhart S, Conn JE, Correa MM. Population structure analyses and demographic history of the malaria vector Anopheles albimanus from the Caribbean and the Pacific regions of Colombia. Malaria Journal 2009;8 259.
  85. 85. Loaiza JR, Scott ME, Bermingham E, Sajur OI, Wilkerson R, Rovira J, Gutiérrez LA, Correa MM, Gríjalva MJ, Birnberg L, Bickersmith S, Conn JE. Late Pleistocene environmental changes lead to unstable demography and population divergence of Anopheles albimanus in the northern Neotropics. Molecular Phylogenetics and Evolution 2010b;57(3) 1341-1346.
  86. 86. Morrone JJ. Biogeographic areas and transition zones of Latin America and the Caribbean Islands based on panbiogeographic and cladistic analyses of the entomofauna. Annual Review of Entomology 2006;51 467-494.
  87. 87. McKeon SN, Lehr MA, Wilkerson RC, Ruiz JF, Sallum MA, Lima JBP, Póvoa MM, Conn JE. Lineage divergence detected in the malaria vector Anopheles marajoara (Diptera: Culicidae) in Amazonian Brazil. Malaria Journal 2010;9 271.
  88. 88. Pedro PM, Sallum MAM. Spatial expansion and population structure of the neotropical malaria vector, Anopheles darlingi (Diptera: Culicidae). Biological Journal of the Linnean Society 2009;97 854-866.
  89. 89. Conn JE, Mitchell SE, Cockburn AF. Mitochondrial DNA analysis of the neotropical malaria vector Anopheles nuneztovari. Genome 1998;41(3) 313-327.
  90. 90. Mirabello LM, Conn JE. Molecular population genetics of the malaria vector Anopheles darlingi in Central and South America. Heredity 2006;96 311-321.
  91. 91. Mirabello L, Vineis JH, Yanoviak SP, Scarpassa VM, Póvoa MM, Padilla N, Achee NL, Conn JE. Microsatellite data suggest significant population structure and differentiation within the malaria vector Anopheles darlingi in Central and South America. BMC Ecology 2008;8 3.
  92. 92. Pedro PM, Uezu A, Sallum MAM. Concordant phylogeographies of 2 malaria vectors attest to common spatial and demographic histories. Journal of Heredity 2010;101(5) 618-627.
  93. 93. Fairley TL, Póvoa MM, Conn JE. Evaluation of the Amazon River delta as a barrier to gene flow for the regional malaria vector, Anopheles aquasalis (Diptera: Culicidae) in northeastern Brazil. Journal of Medical Entomology 2002;39(6) 861-869.
  94. 94. Templeton AR, Routman E, Phillips CA. Separating population structure from population history: a cladistics analysis of the geographical distribution of mitochondrial DNA haplotypes in the tiger salamander, Ambystoma tigrinum. Genetics 1995;140(2) 767-782.
  95. 95. Mirabello LM, Conn JE. Population analysis using the nuclear white gene detects Pliocene/Pleistocene lineage divergence within Anopheles nuneztovari in South America. Medical and Veterinary Entomology 2008;22 109-119.
  96. 96. de Fátima Rossetti D, Mann de Toledo P, Góes AM. New geological framework for Western Amazonia (Brazil) and implications for biogeography and evolution. Quaternary Research 2005; 63 78-89.
  97. 97. Scarpassa VM, Conn JE. Mitochondrial DNA detects a complex evolutionary history with Pleistocene epoch divergence for the neotropical malaria vector Anopheles nuneztovari s.l. American Journal of Tropical Medicine and Hygiene 2011;85(5) 857-867.
  98. 98. Fritz GN, Conn J, Cockburn AF, Seawright JA. Sequence analysis of the ribosomal DNA internal transcribed spacer 2 from populations of Anopheles nuneztovari (Diptera: Culicidae). Molecular Biology and Evolution 1994; 11 406-416.

Written By

Jan E. Conn, Martha L. Quiñones and Marinete M. Póvoa

Submitted: 10 February 2012 Published: 24 July 2013