**Speciation in** *Anopheles gambiae* **— The Distribution of Genetic Polymorphism and Patterns of Reproductive Isolation Among Natural Populations**

Gregory C. Lanzaro and Yoosook Lee

Additional information is available at the end of the chapter

http://dx.doi.org/10.5772/56232

## **1. Introduction**

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The African malaria vector, *Anopheles gambiae*, is characterized by multiple polymorphic chromosomal inversions and has become widely studied as a system for exploring models of ecological speciation. An attempt to develop a molecular diagnostic for the chromosomal forms of *A. gambiae* s.s. led to the development of a PCR-based diagnostic to differentiate M and S molecular forms based on a marker located on the X chromosome. Near complete reproductive isolation between M and S molecular forms has led to the suggestion that *A. gambiae* is in early stages of speciation. Comparative genomic studies have been applied to gain an understanding of the evolutionary process resulting in these forms, but models based on these studies currently lack consensus. Furthermore, various studies suggest further subdivisions within each molecular form. These topics are discussed and suggestions for further research needed to elucidate the population structure of *A. gambiae* are presented.

## **2.** *Anopheles gambiae* **species complex**

Among the global vectors of human malaria arguably the most important species belong to the *Anopheles gambiae* complex, which include the most widespread and potent vectors of malaria in sub-Saharan Africa. The *Anopheles gambiae* species complex includes eight sibling species: *A. gambiae* s.s. Giles, *A. arabiensis* Patton, *A. bwambae* White, *A. melas* Theobald, *A. merus* Dönitz, *A. quadriannulatus* Theobald, *A. amharicus* Hunt, Coetzee and Fettene and *A. comorensis* Brunhes, le Goff and Geoffroy [1-4]. The status of these species was established via the demonstration of F1 hybrid sterility among crosses between populations [4-8], morpho‐

© 2013 Lanzaro and Lee; licensee InTech. This is an open access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited. © 2013 Lanzaro and Lee; licensee InTech. This is a paper distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

logical features [9] and fixed differences in chromosomal inversions [5, 10]. Although the species cannot be reliably distinguished morphologically they do differ in terms of their ecology and geographic distributions (Figure 1). Two species, *A. merus* and *A. melas*, are associated with saltwater larval habitats and so are restricted in distribution to brackish water breeding sites along the east and west coasts respectively. A third saltwater species, *A. bwambae*, is only known to occur in association with hot springs in Semliki Forest National Park in eastern Uganda. The species, *A. quadriannulatus* and *A. amharicus* are primarily zoophilic and are not considered to be involved in the transmission of malaria. *A. quadrian‐ nulatus* occurs in southeastern Africa and *A. amharicus* in Ethiopia [2, 4]. A population on the island of Grande Comore in the Indian Ocean was described as a distinct species, *A. comoren‐ sis*, on the basis of morphological characters [9]. Little is known about the biology of *A. comorensis*. The two remaining freshwater species, *A. gambiae sensu stricto (*hereafter referred to as *A. gambiae)* and *A. arabiensis*, have the broadest geographic distribution and are the most important vectors of human malaria (Figure 1) [11, 12]. *A. gambiae* has been the most studied with respect to molecular and population genetics, and its whole genome sequence was published in 2002 [13].

Natural populations of *A. gambiae* have an extremely complex genetic structure that has been the subject of a great deal of research, a summary of which will be the focus of this chapter. Populations of *A. gambiae* are thought to be undergoing speciation and have been the focus of numerous studies aimed at evaluating speciation models [14-16]. Discrete subpopulations of *A. gambiae* have been defined in two ways: *chromosomal form* and *molecular form*. Recently the M molecular form of *A. gambiae* was elevated to species status and designated *Anopheles coluzzii* Coetzee *et al.* [4]. We retain the designation M and S forms to facilitate discussion of the recent literature.

## **3. Chromosomal forms of** *Anopheles gambiae*

*Chromosomal forms*. The *A. gambiae* genome is organized on three chromosomes: two subme‐ tacentric autosomes and X/Y sex chromosomes, with males being the heterogametic sex. For descriptive purposes the autosomes are divided into two "arms" at the centromere. The longer arm is referred to as the right arm and the shorter the left arm. A high degree of chromosomal polymorphism, in the form of paracentric inversions, has been described in populations of *A. gambiae.* In a recent study Pombi *et al*. [18] describe 82 rare and 7 common inversions observed in natural populations. Inversions are not randomly distributed among chromosomes, but occur most often on the right arm of chromosome 2 (2R). Cytogenetic analysis is facilitated by the presence of giant polytene chromosomes in the cells of certain tissues. In early studies, the salivary glands of larvae were the source of material, but more recently ovarian nurse cells are used (the latter are easier to obtain and make better preparations for microscopic examination). Polytene chromosomes contain light and dark banding patterns that serve as critical landmarks for the determination of karyotypes (Figure 2). Protocols for the preparation of polytene chromosomes for karyotyping are available on-line at [19].

Speciation in *Anopheles gambiae* — The Distribution of Genetic Polymorphism and Patterns of Reproductive Isolation Among Natural Populations http://dx.doi.org/10.5772/56232 175

logical features [9] and fixed differences in chromosomal inversions [5, 10]. Although the species cannot be reliably distinguished morphologically they do differ in terms of their ecology and geographic distributions (Figure 1). Two species, *A. merus* and *A. melas*, are associated with saltwater larval habitats and so are restricted in distribution to brackish water breeding sites along the east and west coasts respectively. A third saltwater species, *A. bwambae*, is only known to occur in association with hot springs in Semliki Forest National Park in eastern Uganda. The species, *A. quadriannulatus* and *A. amharicus* are primarily zoophilic and are not considered to be involved in the transmission of malaria. *A. quadrian‐ nulatus* occurs in southeastern Africa and *A. amharicus* in Ethiopia [2, 4]. A population on the island of Grande Comore in the Indian Ocean was described as a distinct species, *A. comoren‐ sis*, on the basis of morphological characters [9]. Little is known about the biology of *A. comorensis*. The two remaining freshwater species, *A. gambiae sensu stricto (*hereafter referred to as *A. gambiae)* and *A. arabiensis*, have the broadest geographic distribution and are the most important vectors of human malaria (Figure 1) [11, 12]. *A. gambiae* has been the most studied with respect to molecular and population genetics, and its whole genome sequence was

Natural populations of *A. gambiae* have an extremely complex genetic structure that has been the subject of a great deal of research, a summary of which will be the focus of this chapter. Populations of *A. gambiae* are thought to be undergoing speciation and have been the focus of numerous studies aimed at evaluating speciation models [14-16]. Discrete subpopulations of *A. gambiae* have been defined in two ways: *chromosomal form* and *molecular form*. Recently the M molecular form of *A. gambiae* was elevated to species status and designated *Anopheles coluzzii* Coetzee *et al.* [4]. We retain the designation M and S forms to facilitate discussion of

*Chromosomal forms*. The *A. gambiae* genome is organized on three chromosomes: two subme‐ tacentric autosomes and X/Y sex chromosomes, with males being the heterogametic sex. For descriptive purposes the autosomes are divided into two "arms" at the centromere. The longer arm is referred to as the right arm and the shorter the left arm. A high degree of chromosomal polymorphism, in the form of paracentric inversions, has been described in populations of *A. gambiae.* In a recent study Pombi *et al*. [18] describe 82 rare and 7 common inversions observed in natural populations. Inversions are not randomly distributed among chromosomes, but occur most often on the right arm of chromosome 2 (2R). Cytogenetic analysis is facilitated by the presence of giant polytene chromosomes in the cells of certain tissues. In early studies, the salivary glands of larvae were the source of material, but more recently ovarian nurse cells are used (the latter are easier to obtain and make better preparations for microscopic examination). Polytene chromosomes contain light and dark banding patterns that serve as critical landmarks for the determination of karyotypes (Figure 2). Protocols for the preparation of polytene

published in 2002 [13].

174 *Anopheles* Anopheles mosquitoes - New insights into malaria vectors mosquitoes - New insights into malaria vectors

the recent literature.

**3. Chromosomal forms of** *Anopheles gambiae*

chromosomes for karyotyping are available on-line at [19].

**Figure 1.** Geographic distribution of members of the *A. gambiae* complex*.* A: *A. arabiensis* (red); B: *A. gambiae s.s.* (green); C: *A. melas* (Blue), *A. merus* (orange), and *A. bwambae* (cyan); D: *A. quadriannulatus* (former species A) (yel‐ low), *A. amharicus* (former *A. quadriannulatus* B) (magenta) and *A. comorensis* (cyan circle). Data and maps adapted from [17] and [14].

There is general agreement that inversions represent coadapted gene complexes that may enable individuals carrying them to occupy different ecological niches. The nonrandom distribution of inversion breakpoints along the chromosomes [18] and the distribution of inversion frequencies throughout the geographical ranges of the species strongly suggest that at least some of the inversions are maintained by selection that allows different species and, in the case of *A. gambiae*, populations, to survive and exploit a wide variety of habitats [21-23]. The best example is the strong association of inversions 2La and 2Rb with aridity, with the frequency of these inversions being highest in drier areas and even increasing in frequency during the dry season at single sites that experience distinct wet and dry seasons [21, 23, 24]. Specific inversion configurations are associated with specific habitats, leading to the term "ecophenotype" frequently applied to describe individuals carrying certain combinations of inversions [25]. Chromosomal forms have been defined based on the configuration of five paracentric chromosome inversions on the right arm of chromosome 2 (2Rj, b, c, d and u) and one on the left arm of chromosome 2 (2La). Based on this, five chromosomal forms of *A. gambiae* have been described and named *Mopti*, *Bamako*, *Bissau*, *Forest* and *Savanna* according to the geographic regions from which they were first collected and indicating an association of each with a particular type of habitat, as illustrated in Figure 3 [10]. Chromosomal forms are defined as follows: [1] the *Forest* form characterized by the typical non-inverted arrange‐ ment 2R+/+, 2L+/+, or by a single inversion polymorphism due to inversion 2Rb, 2Rd or 2La; [2] *Bissau* characterized by high frequencies of the 2Rd inversion and standard 2L+ arrange‐ ment; [3] *Savanna* exhibiting high frequencies of 2Rb and 2La inversions as well as polymor‐ phism involving the 2Rcu arrangements and polymorphism in the j, d and the rare k inversion; [4] *Bamako* characterized by the fixed 2Rjcu arrangement and polymorphism in the 2Rb inversion; [5] *Mopti* showing high frequencies of 2Rbc, 2Ru and nearly fixed for 2La (Figure 2). The *Savanna* form has the broadest distribution occurring throughout sub-Saharan Africa, the *Mopti* form predominates in drier habitats in West Africa, the *Forest* form occurs in wetter habitats in Africa, the *Bamako* form occurs in habitats along the Niger River in West Africa and the *Bissau* form is restricted to West Africa (Figure 3) [26, 27].

**Figure 2.** Photomap of polytene chromosomes of *A. gambiae* Forest-M form (collected from Tiko, Cameroon) depict‐ ing band positions. Six major inversions on the chromosome 2 used for identifying chromosomal forms are marked.

Speciation in *Anopheles gambiae* — The Distribution of Genetic Polymorphism and Patterns of Reproductive Isolation Among Natural Populations http://dx.doi.org/10.5772/56232 177

frequency of these inversions being highest in drier areas and even increasing in frequency during the dry season at single sites that experience distinct wet and dry seasons [21, 23, 24]. Specific inversion configurations are associated with specific habitats, leading to the term "ecophenotype" frequently applied to describe individuals carrying certain combinations of inversions [25]. Chromosomal forms have been defined based on the configuration of five paracentric chromosome inversions on the right arm of chromosome 2 (2Rj, b, c, d and u) and one on the left arm of chromosome 2 (2La). Based on this, five chromosomal forms of *A. gambiae* have been described and named *Mopti*, *Bamako*, *Bissau*, *Forest* and *Savanna* according to the geographic regions from which they were first collected and indicating an association of each with a particular type of habitat, as illustrated in Figure 3 [10]. Chromosomal forms are defined as follows: [1] the *Forest* form characterized by the typical non-inverted arrange‐ ment 2R+/+, 2L+/+, or by a single inversion polymorphism due to inversion 2Rb, 2Rd or 2La; [2] *Bissau* characterized by high frequencies of the 2Rd inversion and standard 2L+ arrange‐ ment; [3] *Savanna* exhibiting high frequencies of 2Rb and 2La inversions as well as polymor‐ phism involving the 2Rcu arrangements and polymorphism in the j, d and the rare k inversion; [4] *Bamako* characterized by the fixed 2Rjcu arrangement and polymorphism in the 2Rb inversion; [5] *Mopti* showing high frequencies of 2Rbc, 2Ru and nearly fixed for 2La (Figure 2). The *Savanna* form has the broadest distribution occurring throughout sub-Saharan Africa, the *Mopti* form predominates in drier habitats in West Africa, the *Forest* form occurs in wetter habitats in Africa, the *Bamako* form occurs in habitats along the Niger River in West Africa and

**Figure 2.** Photomap of polytene chromosomes of *A. gambiae* Forest-M form (collected from Tiko, Cameroon) depict‐ ing band positions. Six major inversions on the chromosome 2 used for identifying chromosomal forms are marked.

the *Bissau* form is restricted to West Africa (Figure 3) [26, 27].

176 *Anopheles* Anopheles mosquitoes - New insights into malaria vectors mosquitoes - New insights into malaria vectors

**Figure 3.** Distribution of chromosomal forms in West and Central Africa. Data from *PopI* [27]*.* BAM stands for *Bamako* chromosomal form, FOR for *Forest*, MOP for *Mopti*, SAV for *Savanna* and BIS for *Bissau*. OTHER refers to samples with karyotypes that do not fit any described chromosomal form designation.

The Xag inversions is fixed and used as a diagnostic marker to distinguish *A. gambiae* from other species in the com‐ plex. Chromosome photomap adapted from: [20]

It has furthermore been suggested that the chromosomal forms are to some extent reproduc‐ tively isolated and represent distinct species or incipient species that have evolved or are evolving via a process described as "ecotypic speciation" [15, 25]. Studies of karyotype frequencies at sites where the *Bamako*, *Mopti* and *Savanna* forms occur in sympatry have revealed significant departures from the Hardy-Weinberg equilibrium (H-W) [10, 28-30]. Specifically, heterokaryotypes representing hybrids between the *Savanna* form and the other two were under-represented and *Bamako*/*Mopti* hybrids were never encountered. This observation led to the suggestion that there is partial reproductive isolation between *Savan‐ na* and the other forms, nearly complete isolation between the *Bamako* and *Mopti* forms and that these forms represent incipient species. However, hybridization experiments involving crosses between the *Bamako* and *Mopti* forms resulted in viable offspring, demonstrating a lack of post-mating reproductive barriers between them [29, 31]. An estimate of genetic distance (based on allozyme frequencies) [32] between the *Bamako* and *Mopti* forms was reported as 0.015 [33], a value not higher than that typically found between local populations of a single mosquito species. We found that genotypic frequencies in a population composed of three chromosomal forms in Mali did not depart from Hardy-Weinberg expectations, suggesting that this population represents a single gene pool (Lanzaro, unpublished).

It should be emphasized that although these studies do not support reproductive isolation among chromosomal forms, they do not disprove it. Pre-mating isolating mechanisms may act as a barrier between subpopulations, even if post-mating mechanisms have not evolved, and isolation may be recent, so that not enough time has passed for the accumulation of substantial allozyme divergence between the forms. Lanzaro *et al.*[34] conducted a study based on 21 microsatellite loci distributed over the genome, examining genetic differentiation between the *Bamako* and *Mopti* forms in the villages of Banambani and Selinkenyi, Mali. This study revealed strong genetic differentiation between *A. gambiae* and *A. arabiensis*, used here as an outgroup. Within *A. gambiae*, different patterns of genetic differentiation, depending on the genomic location of the microsatellite loci, were observed. No genetic differentiation was found on the 3rd and X-chromosome whereas strong linkage disequilibrium and low levels of genetic differentiation were found for loci located on the 2nd chromosome in association with the inversions that occur there [34]. Similar results were obtained in a study also using microsatellites distributed on all three chromosomes for samples collected in the villages of Selinkenyi, Soulouba, and Kokouna, Mali [35].

Gene flow, like other forces, may be higher in some parts of the genome and lower in others. For example, favorable genes can still be exchanged successfully even when barriers to gene flow are strong. Such genes could be at loci that confer local adaptations and at any linked loci. The significance of this is that gene flow, even if estimated accurately, may still fail to account for variation among different parts of the genome. This effect may be particularly strong for genes contained within inversions, both because of potentially strong selection and because of linkage imposed by the reduced recombination associated with inversions. This effect was explored by Tripet et al. [36] in a study in which they examined divergence for microsatellite loci contained within the *j* and *b* inversions compared with loci outside of inversions. Indeed they did find elevated divergence estimated from loci contained within the inversions relative to those outside. This pattern of divergence, with a strongly non-random distribution over the genome, was later described as a 'mosaic genome architecture' in a paper by Wang-Sattler et al. [37]. As we shall see, this concept was later refined based on high resolution genome-wide analysis, ultimately leading to the recognition of 'islands of speciation' in the *A. gambiae* genome.

Using the chromosomal form concept to define genetically discrete populations is problematic because there is substantial overlap in inversions that define them, probably due to some level of contemporary gene flow. This creates ambiguities in assigning individuals to form, diminishing the utility of the chromosomal form concept for defining reproductive boundaries among populations. For example, in a recent survey of populations in Mali, we found that 26% of 2,459 individuals could not be assigned to a chromosomal form and in Cameroon 39% of 632 individuals could likewise not be assigned (Figure 3, data available at *PopI* [27]).

*The role of chromosome inversions in A. gambiae evolution: Ecotypic Speciation*. The chromosomal or ecotypic model of speciation was first described for anopheline mosquitoes by Coluzzi [38] and is the prevailing model applied to the chromosomal forms of *A. gambiae*[14, 15]. This model is founded on the observation that certain paracentric inversions that are polymorphic in *A. gambiae* are non-randomly distributed in nature. These are thought to contain multi-locus genotypes that are adaptive to specific aquatic habitats occupied by the immature stages of the mosquito. Under this model, populations carrying alternate gene arrangements would inhabit different, spatially isolated habitats. Genetic divergence, enhanced by reduced recombination associated with the inversions, would then evolve. Ultimately divergence would include genes resulting in reproductive isolation (reduced fitness in hybrids or behavioral differences preventing between form mating), explaining the observed deficiency of inversion heterozygotes. This model was initially adopted to describe the evolution of chromosomal forms of *A. gambiae* [15, 21, 28], but now has become the model for explaining the evolution of the molecular forms as described below [16, 39-42].

mosquito species. We found that genotypic frequencies in a population composed of three chromosomal forms in Mali did not depart from Hardy-Weinberg expectations, suggesting

It should be emphasized that although these studies do not support reproductive isolation among chromosomal forms, they do not disprove it. Pre-mating isolating mechanisms may act as a barrier between subpopulations, even if post-mating mechanisms have not evolved, and isolation may be recent, so that not enough time has passed for the accumulation of substantial allozyme divergence between the forms. Lanzaro *et al.*[34] conducted a study based on 21 microsatellite loci distributed over the genome, examining genetic differentiation between the *Bamako* and *Mopti* forms in the villages of Banambani and Selinkenyi, Mali. This study revealed strong genetic differentiation between *A. gambiae* and *A. arabiensis*, used here as an outgroup. Within *A. gambiae*, different patterns of genetic differentiation, depending on the genomic location of the microsatellite loci, were observed. No genetic differentiation was found on the 3rd and X-chromosome whereas strong linkage disequilibrium and low levels of genetic differentiation were found for loci located on the 2nd chromosome in association with the inversions that occur there [34]. Similar results were obtained in a study also using microsatellites distributed on all three chromosomes for samples collected in the villages of

Gene flow, like other forces, may be higher in some parts of the genome and lower in others. For example, favorable genes can still be exchanged successfully even when barriers to gene flow are strong. Such genes could be at loci that confer local adaptations and at any linked loci. The significance of this is that gene flow, even if estimated accurately, may still fail to account for variation among different parts of the genome. This effect may be particularly strong for genes contained within inversions, both because of potentially strong selection and because of linkage imposed by the reduced recombination associated with inversions. This effect was explored by Tripet et al. [36] in a study in which they examined divergence for microsatellite loci contained within the *j* and *b* inversions compared with loci outside of inversions. Indeed they did find elevated divergence estimated from loci contained within the inversions relative to those outside. This pattern of divergence, with a strongly non-random distribution over the genome, was later described as a 'mosaic genome architecture' in a paper by Wang-Sattler et al. [37]. As we shall see, this concept was later refined based on high resolution genome-wide analysis, ultimately leading to the recognition of 'islands of speciation' in the *A. gambiae*

Using the chromosomal form concept to define genetically discrete populations is problematic because there is substantial overlap in inversions that define them, probably due to some level of contemporary gene flow. This creates ambiguities in assigning individuals to form, diminishing the utility of the chromosomal form concept for defining reproductive boundaries among populations. For example, in a recent survey of populations in Mali, we found that 26% of 2,459 individuals could not be assigned to a chromosomal form and in Cameroon 39% of

*The role of chromosome inversions in A. gambiae evolution: Ecotypic Speciation*. The chromosomal or ecotypic model of speciation was first described for anopheline mosquitoes by Coluzzi [38]

632 individuals could likewise not be assigned (Figure 3, data available at *PopI* [27]).

that this population represents a single gene pool (Lanzaro, unpublished).

Selinkenyi, Soulouba, and Kokouna, Mali [35].

178 *Anopheles* Anopheles mosquitoes - New insights into malaria vectors mosquitoes - New insights into malaria vectors

genome.

The most thorough evaluation of the ecotypic speciation model has been its application to the *Bamako* and *Savanna* forms in Mali [15]. Central to this evaluation is the observation of niche partitioning with respect to larval habitat. This observation was based on a PCR identification method developed for detecting the 2R*j* inversion [43] among larval samples collected in rock pools vs. more typical larval sites (puddles/ponds) in the village of Banambani, Mali. We evaluated this PCR method on a set of 85 field-collected adults previously scored for the 2R*j* genotype cytogenetically. In total, we selected 25 2R*j* homozygotes (*j*/*j*), 40 2R*j* heterozygotes (+*j* /*j*) and 20 2R*j* standard (+*<sup>j</sup>* /+*j* ) from the villages of Banambani, Selinkenyi, Tinko and Seroume, Mali. The 2R*j* PCR was accurate in calling 2R*j* homozygotes (*j*/*j*) (100%) in all villages regardless of the presence of the 2R*c* and *u* inversions (Table 1). However, the PCR was much less accurate for the standard arrangement for 2R*j* (+*<sup>j</sup>* /+*j* ), resulting in consistent false identification as 2R*j* heterozygotes (+*<sup>j</sup>* /*j*) in 11 cases and 2Rj (*j*/*j*) homozygotes in 5 cases. Moreover, all true heter‐ ozytoes (+*<sup>j</sup>* /*j*) were misidentified as either *j*/*j* (N=13) or +*<sup>j</sup>* /+*j* (N=7). The low accuracy rate (=48.2%) of the 2R*j* diagnostic PCR casts doubt on this sole example of niche partitioning (rock pool vs. other) in larval habitat distinguishing the two forms.

The 2R*j* inversion polymorphism in Mali shows two mating patterns in different parts of the species range in this country. At sites along the Senegal River (e.g. villages of Sebetou, Seroume, Bantinngoungou, and Tinko), 2R*j* inversion heterozygotes are commonly found and 2R*j* karyotypes are in Hardy-Weinberg expectation (HWE). On the other hand, at sites along the Niger River and its tributaries (e.g. villages of Banambani, Doneguebougou, Senou, Kela, Selinkenyi, Soulouba, Yorobougoula, Kokouna), a severe deficiency of 2R*j* heterozygotes are observed and 2R*j* genotypes are not in HWE (Figure 4).

In the literature the *Bamako* form includes three genotypes, *jcu/jcu*, *jcu/jbcu*, and *jbcu/jbcu*, all homozygous for *j* [21]. Other individuals carrying 2R*j* inversion but not *c* and *u* inversions such as *jbd/jbd*, and *jb/b*, commonly found along the Senegal River, cannot be classified under the current definitions for chromosomal forms. 94% of the 2R*j* homozygotes along the Niger River are *Bamako* forms, while no *Bamako* forms are found along the Senegal River.

Overall these results weaken the argument that paracentric inversions play a role in the evolution of reproductive isolation via divergent selection (ecotypic speciation), both because

**Figure 4.** 2R*j* inversion distribution in Mali. For legend of the GlobCover 2009 land cover type used as background, see Figure 2.


**Table 1.** Evaluation of 2R*j* genotyping via a PCR identification method. Samples were karyotyped microscopically prior to being assayed using the PCR protocol of Coulibaly *et al*. [43] \* NA stands for 'no amplification'

they cast doubt on the association of inversions with distinct larval habitats and on evidence for reproductive isolation between individuals that differ with respect to the inversions they carry (e.g. a lack of *j* inversion heterozygotes). Genome-wide comparisons of individuals with and without inversions have been conducted and these cast doubt on the role of inversions as forming "coadapted gene complexes". These results are described in detail below.

Speciation in *Anopheles gambiae* — The Distribution of Genetic Polymorphism and Patterns of Reproductive Isolation Among Natural Populations http://dx.doi.org/10.5772/56232 181

*The role of chromosome inversions in A. gambiae evolution: Comparative Genomics*. Central to the "ecotypic speciation" model as applied to *A. gambiae*is the notion that inversions contain multilocus genotypes that are adaptive to different environments. These "coadapted gene com‐ plexes" arise and are maintained as the consequence of reduced recombination within and around the inversion. Ultimately these become, either directly or indirectly, associated with reproductive isolation. One expectation arising from this phenomenon, assuming that reproductive isolation is incomplete or has evolved recently, is higher levels of genetic divergence in regions of the genome contained within the inversion relative to elsewhere in the genome. Indeed, in a genome-wide scan comparing individuals with and without the 2L*a* inversion, significantly higher divergence was observed in a 3 Mb region of the genome within and proximal to the inversion [44]. However, in a subsequent study that included a comparison of inverted and uninverted genomes for the four common 2R inversions (*j*, *b*, *c* and *u*), a region of the genome spanning ~26 Mb, divergence was limited to just one small region (~100 kb) in the 2R*u* inversion [45]. In both studies the Affymetrix *Plasmodium*/*Anoph‐ eles* Genome Microarray (*P*/*A* array), which contains 142,065 25bp probes, representing roughly 13,000 predicted genes, was used. Lack of divergence associated with the inversions hypothe‐ sized to be driving the "ecotypic speciation" process was unexpected. Several explanations were provided including that divergence between the inversion arrangements escaped detection due to shared ancestral polymorphism, extensive recombination within the inver‐ sions (gene flux) and limits to the resolution of the microarray they used [45].

In a more recent study [46] the genomes of individuals homozygous for the *jbcu* arrangement (*Bamako* form) were compared with individuals homozygous for the standard arrangement, *+j +b+c+u* (*Savanna* form). In this case all individuals were of the S molecular form (unlike the comparisons made in the White et al. [45] study, which were a mixture of M and S form individuals). In addition, Lee et al. [46] utilized an *A. gambiae* whole genome tiling microarray (WGTM) which provides a far higher resolution of the genome than the *P*/*A* array (probe density = 1 probe per 100,000bp for the *P*/*A* array; 1 probe per 17bp for the WGTM). As in the White et al. [45] study, this new study revealed very little divergence associated with the chromosome 2R inversions. However, a 3Mb region of the genome on the X chromosome, proximal to the centromere was observed. This is the same region of the genome that contains the sequence divergence used to define the M and S molecular forms (discussed in detail in the following sections). X chromosome divergence is associated with reproductive isolation observed between both the M and S molecular forms and between the *Bamako* and *Savanna* chromosomal forms. These results suggest that the 2R inversions may not be involved in either the evolution or maintenance of reproductive isolation among *A. gambiae* populations.

## **4. Molecular forms of** *Anopheles gambiae*

they cast doubt on the association of inversions with distinct larval habitats and on evidence for reproductive isolation between individuals that differ with respect to the inversions they carry (e.g. a lack of *j* inversion heterozygotes). Genome-wide comparisons of individuals with and without inversions have been conducted and these cast doubt on the role of inversions as

**Figure 4.** 2R*j* inversion distribution in Mali. For legend of the GlobCover 2009 land cover type used as background, see

**2Rj N match mismatch NA\* match mismatch NA\*** j/j 25 24 0 1 24 0 1

/j 20 0 **20** 0 0 **19** 1

/+j 40 17 **14** 9 16 **16** 8

**Table 1.** Evaluation of 2R*j* genotyping via a PCR identification method. Samples were karyotyped microscopically prior

**Run 1 Run 2**

Figure 2.

180 *Anopheles* Anopheles mosquitoes - New insights into malaria vectors mosquitoes - New insights into malaria vectors

+j

+j

forming "coadapted gene complexes". These results are described in detail below.

to being assayed using the PCR protocol of Coulibaly *et al*. [43] \* NA stands for 'no amplification'

*Defining Molecular Forms*. An attempt to develop a molecular diagnostic for the chromosomal forms of *A. gambiae* identified 10 nucleotide residues that differ between the *Mopti* and the *Savanna* or *Bamako* chromosomal forms in a 2.3 kb fragment at the 5' end of the rDNA IGS region located on the X chromosome [47]. These findings led to the development of a PCR- based diagnostic to differentiate *Mopti* chromosomal forms from *Bamako* and *Savanna* forms based on a single base pair substitution at the 540th nucleotide position in a 28S rDNA amplimer sequence. *Mopti* form individuals carry a C/C genotype and both *Bamako* and *Savanna* individuals a T/T genotype (Genbank accession number AF470112-6) [48]. Individuals carrying C/C are referred to as M molecular form and those carrying the T/T genotype are known as S molecular form. There is good correspondence between the M molecular form and the *Mopti* chromosomal form in Burkina Faso and Mali, however, the *Bamako* and *Savanna* chromosomal forms cannot be distinguished (both are of the S molecular form). The association of M and S molecular forms and chromosomal forms breaks down at other locations in West Africa. For example, in western Senegal and Gambia the association between the *Savanna* chromosomal form and S molecular form does not hold [49] and the *Forest* form contains both M and S individuals. The M and S molecular forms, therefore, largely fail as a diagnostic for chromosomal form. However, the significance of the M and S forms of *A. gambiae* goes well beyond their utility as proxies for identifying chromosomal forms. The molecular form concept has now largely replaced chromosomal form for defining discrete sub-populations of *A. gambiae*, that are to some extent reproductively isolated.

M and S forms occur in sympatry at many sites in West and Central Africa, and typically there is a high degree of reproductive isolation between the two forms. M/S hybrids (C/T genotype) produced in the laboratory did yield clearly distinguishable hybrid patterns in females. Surprisingly, however, field collected individuals carrying "hybrid" karyotypes (putative hybrids between different chromosomal forms) did not produce results consistent with their being hybrid, but rather produced either M or S patterns [48]. This observation supports the notion that certain karyotypes, thought to be fixed in one chromosomal form or another, are in fact shared, occurring commonly in one form and rarely in another, due to ancestral polymorphism and/or ongoing gene flow [40, 50]. This diagnostic now forms the basis of recognizing two distinct subpopulations of *A. gambiae*, known as *molecular forms* (M and S).

*Alternate methods for distinguishing M and S forms.* The original PCR-based diagnostic used to distinguish the M and S forms [48] was further developed into a method using a restriction digestion of PCR amplimers that allowed distinguishing *A. gambiae* from one of its sibling species *A. arabiensis* while simultaneously distinguishing M from S [51]. This was useful in the field since *A. arabiensis* and both the M and S forms are morphologically indistinguishable and commonly occur in sympatry at study sites throughout West and Central Africa. In 2008, a new method for distinguishing the M and S forms was discovered which takes advantage of polymorphism in insertion sites for a group of retrotransposons known as short interspersed elements (SINEs). One of the SINE insertion sites, located on the X chromosome and referred to as SINE X6.1, was found to be fixed in the M form and absent in the S form. In subsequent studies, in which multiple M/S diagnostic methods were employed, some discrepancies in results were observed [52]. These were most common in populations where M/S hybridization is common, for example in Guinea-Bissau.

*Relationships between the M and S forms*. Understanding the relationship between the two molecular forms has been the focus of an intense and ongoing research effort. The S form has the broadest distribution occurring throughout sub-Saharan Africa, whereas the M form occurs throughout West and parts of Central Africa. With the exception of a single site in northern Zimbabwe [53], M is absent from eastern Africa (Figure 5) [49].

based diagnostic to differentiate *Mopti* chromosomal forms from *Bamako* and *Savanna* forms based on a single base pair substitution at the 540th nucleotide position in a 28S rDNA amplimer sequence. *Mopti* form individuals carry a C/C genotype and both *Bamako* and *Savanna* individuals a T/T genotype (Genbank accession number AF470112-6) [48]. Individuals carrying C/C are referred to as M molecular form and those carrying the T/T genotype are known as S molecular form. There is good correspondence between the M molecular form and the *Mopti* chromosomal form in Burkina Faso and Mali, however, the *Bamako* and *Savanna* chromosomal forms cannot be distinguished (both are of the S molecular form). The association of M and S molecular forms and chromosomal forms breaks down at other locations in West Africa. For example, in western Senegal and Gambia the association between the *Savanna* chromosomal form and S molecular form does not hold [49] and the *Forest* form contains both M and S individuals. The M and S molecular forms, therefore, largely fail as a diagnostic for chromosomal form. However, the significance of the M and S forms of *A. gambiae* goes well beyond their utility as proxies for identifying chromosomal forms. The molecular form concept has now largely replaced chromosomal form for defining discrete sub-populations of *A.*

M and S forms occur in sympatry at many sites in West and Central Africa, and typically there is a high degree of reproductive isolation between the two forms. M/S hybrids (C/T genotype) produced in the laboratory did yield clearly distinguishable hybrid patterns in females. Surprisingly, however, field collected individuals carrying "hybrid" karyotypes (putative hybrids between different chromosomal forms) did not produce results consistent with their being hybrid, but rather produced either M or S patterns [48]. This observation supports the notion that certain karyotypes, thought to be fixed in one chromosomal form or another, are in fact shared, occurring commonly in one form and rarely in another, due to ancestral polymorphism and/or ongoing gene flow [40, 50]. This diagnostic now forms the basis of recognizing two distinct subpopulations of *A. gambiae*, known as *molecular forms* (M and S).

*Alternate methods for distinguishing M and S forms.* The original PCR-based diagnostic used to distinguish the M and S forms [48] was further developed into a method using a restriction digestion of PCR amplimers that allowed distinguishing *A. gambiae* from one of its sibling species *A. arabiensis* while simultaneously distinguishing M from S [51]. This was useful in the field since *A. arabiensis* and both the M and S forms are morphologically indistinguishable and commonly occur in sympatry at study sites throughout West and Central Africa. In 2008, a new method for distinguishing the M and S forms was discovered which takes advantage of polymorphism in insertion sites for a group of retrotransposons known as short interspersed elements (SINEs). One of the SINE insertion sites, located on the X chromosome and referred to as SINE X6.1, was found to be fixed in the M form and absent in the S form. In subsequent studies, in which multiple M/S diagnostic methods were employed, some discrepancies in results were observed [52]. These were most common in populations where M/S hybridization

*Relationships between the M and S forms*. Understanding the relationship between the two molecular forms has been the focus of an intense and ongoing research effort. The S form has the broadest distribution occurring throughout sub-Saharan Africa, whereas the M form

*gambiae*, that are to some extent reproductively isolated.

182 *Anopheles* Anopheles mosquitoes - New insights into malaria vectors mosquitoes - New insights into malaria vectors

is common, for example in Guinea-Bissau.

**Figure 5.** Distribution of molecular forms in Sub-Saharan Africa. For legend of the GlobCover 2009 land cover type used as background, see Figure 2. Data from [27, 40, 49, 50, 54]

Although the M and S forms are largely reproductively isolated in most places where they occur together, this is not true everywhere. Hybridization between forms occurs rarely (~1%) in Mali [55] and reproductive isolation between M and S appears to be complete in Cameroon [56]. In The Gambia, M/S hybrids were identified from a number of sites at frequencies as high as 16.7% of the *A. gambiae* individuals sampled [57] and in Guinea-Bissau hybrids were recovered in over 20% of the individuals assayed [58, 59]. A cryptic subgroup of *A. gambiae* known as the "Goundry" population collected in Burkina Faso was recently found to be composed of 36% M/S hybrids [60]. The Goundry population discovered in the Sudan Savanna zone of Burkina Faso in larval collections but absent in indoor adult collection of the same locality, suggesting that adult stage of Goundry populations mostly rest outdoors [60, 61]. These results suggest that linkage between the M and S alleles and those genes that directly affect reproductive isolation has broken down in a much broader geographic area than previously suggested. Therefore, the notion of an M form and an S form that are largely reproductively isolated (incipient species) and that hybridization only occurs in the "Far-West" region of Africa [62] is an oversimplification.

In the laboratory, chromosomal and molecular forms, including the *Bamako* and *Savanna* forms, appear to display no post-zygotic isolation [31, 63, 64]. Analysis of sperm recovered from inseminated females [55] and the composition of mating swarms [65] support the existence of strong, but not complete, pre-mating reproductive isolation between the M and S molecular forms in nature.

The two molecular forms display phenotypic divergence in different locations within their geographic range [66]. Most notable among these phenotypic differences include differential insecticide resistance [67], desiccation resistance [68], larval habitat segregation [69], and wing morphological differentiation [70]. It has been proposed that the mechanism responsible for promoting divergence is pre-zygotic [63] and associated with mate selection either during swarm formation [71, 72] or within a swarm [65]. Diabate et al. found evidence of clustering of swarms composed of individuals of a single molecular form within the village of Donégué‐ bougou, Mali [71]. Mixed swarms of M and S forms were found elsewhere (Burkina Faso) but the occurrence of mixed swarms was lower than the frequency expected by chance. Manoukis *et al.* analyzed the shape of male swarms and suggested that a difference in swarm organization between M and S forms may enhance the behavioral isolation of the two forms [72].

### **5. Evolution of the M and S forms**

*Comparative genomics*. Early studies aimed at describing patterns of genetic divergence among chromosomal forms revealed what was termed a "mosaic genome architecture", with divergence distributed non-randomly over the genome (as described above, [29]). Comparisons of the M and S forms revealed a similar pattern. Initial work examined the distribution of microsatellite DNA polymorphism showing exceptionally high divergence in a region of the genome proximal to the centromere on the X chromosome, near the rDNA locus used to define the two forms [35, 73]. High levels of M/S form divergence on the X chromosome was substantiated through detailed examination of the centromeric re‐ gion using DNA sequencing [74, 75].

The first high density genome-wide comparison of M and S was conducted by Turner et al. [76] using samples collected in Cameroon. They utilized an Affymetrix *Plasmodium*/*Anopheles* Genome Microarray which contains 142,065 25bp probes representing roughly 13,000 predict‐ ed genes. Divergence between the M and S genomes was very low and restricted to three discrete regions, one on the X chromosome (corresponding with the location identified in the microsatellite studies) and two on chromosome 2, one on 2L and one very small (37kb) region on 2R. In total, these diverged regions cover less than 2.8Mb, roughly 1% of the genome. In a subsequent study, utilizing the same microarray, but with samples collected in Mali, the small 2R region of divergence was not observed, and so this small region was considered not to contribute to reproductive isolation between the two forms [77]. Later a third diverged region was observed on the left arm of chromosome 3L and this region, like the X and chromosome 2L regions, was proximal to the centromere [16]. Taken together these studies revealed that the M and S genomes are diverged over only about 3% of their genomes and that this diver‐ gence is organized into 3 small regions located near the centromere on the X, 2L and 3L chromosomes, with the remainder of their genomes essentially undifferentiated. These regions of divergence have been considered to represent *islands of speciation* because it is thought that they contain genes that are directly involved in reproductive isolation.

strong, but not complete, pre-mating reproductive isolation between the M and S molecular

The two molecular forms display phenotypic divergence in different locations within their geographic range [66]. Most notable among these phenotypic differences include differential insecticide resistance [67], desiccation resistance [68], larval habitat segregation [69], and wing morphological differentiation [70]. It has been proposed that the mechanism responsible for promoting divergence is pre-zygotic [63] and associated with mate selection either during swarm formation [71, 72] or within a swarm [65]. Diabate et al. found evidence of clustering of swarms composed of individuals of a single molecular form within the village of Donégué‐ bougou, Mali [71]. Mixed swarms of M and S forms were found elsewhere (Burkina Faso) but the occurrence of mixed swarms was lower than the frequency expected by chance. Manoukis *et al.* analyzed the shape of male swarms and suggested that a difference in swarm organization

between M and S forms may enhance the behavioral isolation of the two forms [72].

*Comparative genomics*. Early studies aimed at describing patterns of genetic divergence among chromosomal forms revealed what was termed a "mosaic genome architecture", with divergence distributed non-randomly over the genome (as described above, [29]). Comparisons of the M and S forms revealed a similar pattern. Initial work examined the distribution of microsatellite DNA polymorphism showing exceptionally high divergence in a region of the genome proximal to the centromere on the X chromosome, near the rDNA locus used to define the two forms [35, 73]. High levels of M/S form divergence on the X chromosome was substantiated through detailed examination of the centromeric re‐

The first high density genome-wide comparison of M and S was conducted by Turner et al. [76] using samples collected in Cameroon. They utilized an Affymetrix *Plasmodium*/*Anopheles* Genome Microarray which contains 142,065 25bp probes representing roughly 13,000 predict‐ ed genes. Divergence between the M and S genomes was very low and restricted to three discrete regions, one on the X chromosome (corresponding with the location identified in the microsatellite studies) and two on chromosome 2, one on 2L and one very small (37kb) region on 2R. In total, these diverged regions cover less than 2.8Mb, roughly 1% of the genome. In a subsequent study, utilizing the same microarray, but with samples collected in Mali, the small 2R region of divergence was not observed, and so this small region was considered not to contribute to reproductive isolation between the two forms [77]. Later a third diverged region was observed on the left arm of chromosome 3L and this region, like the X and chromosome 2L regions, was proximal to the centromere [16]. Taken together these studies revealed that the M and S genomes are diverged over only about 3% of their genomes and that this diver‐ gence is organized into 3 small regions located near the centromere on the X, 2L and 3L chromosomes, with the remainder of their genomes essentially undifferentiated. These regions

forms in nature.

**5. Evolution of the M and S forms**

184 *Anopheles* Anopheles mosquitoes - New insights into malaria vectors mosquitoes - New insights into malaria vectors

gion using DNA sequencing [74, 75].

*Islands of speciation model*. The widely held interpretation of this work is that *A. gambiae* forms represent incipient species, but with enough gene flow to prevent their genomes from diverging in all but a few, relatively small regions [34, 35, 37, 76, 77]. This interpretation is consistent with recent genic models of speciation that predict the existence of small regions of divergence between incipient species in the presence of some degree of gene flow (Figure 6) [78, 79]. The observation that putative "*islands of speciation*" in *A. gambiae* are located proximal to centromeres, where levels of recombination are known to be low, is likewise consistent with models that consider speciation to be driven by genes located in regions of the genome with reduced crossing-over [80, 81].

*Incidental islands model*. White et al. [16] developed PCR-RFLP assays to detect SNPs that occurred in each of the three islands of speciation and that were diagnostic for the M and S forms. They genotyped a total of 517 individuals including both M and S forms from Mali, Burkina Faso, Cameroon and Kenya. They found complete association among the three unlinked islands in 512 of the 517 individuals genotyped (275 M form and 237 S). Of the five exceptional genotypes, three were heterozygous at all three loci, suggesting these represented F1 hybrids. To account for the nearly complete linkage between the three diverged islands they suggest that gene flow between M and S must be nearly zero. The presence of F1 hybrids suggests that they have such low fitness that they contribute little to gene flow between the forms. As mentioned above F1 hybrids generated in the laboratory show no evidence of intrinsically low fitness, so it is assumed that these are maladapted to conditions in nature. Additional support for very low levels of between form gene flow come from comparisons of M and S based on high-density, genome-wide SNP genotyping [41] and whole genome sequences [42] which revealed widespread divergence between the M and S genomes. Collectively these studies propose an alternative model referred to as the "incidental islands" model [82, 83], which states that reproductive isolation between M and S is complete and that the observed islands of divergence may be incidental, meaning that the divergence observed in areas proximal to centromeres do not necessarily represent the location of genes underlying reproductive isolation but the divergence is due to segregating ancestral variation and not due to contemporary gene flow.

In summary, two opposing models exist that describe the relationship between the M and S forms. The "genomic islands of speciation" model suggests that divergence between the M and S genomes is restricted to small regions (~3% of the genome) that may contain the genes responsible for reproductive isolation between forms and that ongoing gene flow is respon‐ sible for very low levels of divergence over the remaining 97% of the genome. The second model, the "incidental islands of divergence" model, suggests that divergence between the two forms is far more extensive and widely distributed over the genome, that gene flow between the two forms is nearly zero and that the M and S forms therefore represent distinct species (Figure 6D).

**Figure 6.** A: Stage 1 - Population/races with differential adaptation; reproductive isolation (RI) not apparent. Green box represents diverged loci specific to Population 1 (Pop1) and Blue represents diverged loci specific to Pop2. Arrows indicate regions of gene flow. B: Stage 2 - Transition between races and species with some degree of RI; population may fuse or diverge. C: Stage 3 - Divergent populations beyond the point of fusion but still share a portion of their genome via gene flow; good species. D: Stage 4 - Species with complete RI. Adapted from [79].

## **6. Further sub-divisions within molecular forms**

Although most discussions consider M and S as the major and biologically relevant subdivi‐ sions of *A. gambiae* there is evidence that the two can be further subdivided into population groups that are significantly diverged.

*Subdivision within the S form.* In a continent-wide survey Lehmann et al. [73] found that S form populations fall into two well defined clades, based on analysis of microsatellite DNA. They refer to these clades as the Northwest (Nigeria, Gabon, Democratic Republic of Congo, NW Kenya) and Southeast (SW Kenya, Tanzania, Malawi) divisions. Wang-Sattler et al. [37] also conducted an analysis based on microsatellite DNA and likewise report that the S form in eastern Africa (Kenya) are distinct from S form populations in the west (Mali). In addition to the East vs. West division between allopatric S form populations is the division of sympatric S form populations in Mali. These are described in detail above (Section 2). In brief, the S form in Mali is divided into the *Bamako* and *Savanna* chromosomal forms which display strong asssortative mating where they occur in sympatry at sites along the Niger River ([21], also see Figure 4). These two populations can be distinguished by the *j* inversion, which is fixed in the *Bamako* form and absent in the *Savanna*. Interestingly, although the two share the X-linked allele that defines them as S molecular form, a detailed analysis revealed that they are strongly diverged at a 3Mb region of the X chromosome, proximal to the centromere [46].

*Subdivision within the M form.* A comparison of the M form in Mali and the M form in Cameroon has revealed that the two are very different genetically, in fact, divergence between these two is higher than the level of divergence between the M and S forms [23]. This observation has led to a recognition of two, distinct M form groups, the Mopti-M form, which is polymorphic with respect to the 2R *b*, *c*, and u chromosome inversions and the Forest-M form which lacks inversions on chromosome 2L and 2R [23, 84]. In addition to genetic divergence the Forest-M and Mopti-M forms differ in their ecology. The Mopti-M in Mali is most common in the dry northern part of the country whereas Forest-M is absent in the dry northern part of Cameroon and is restricted to the wet southern part of the country [23]. This observation lends support the notion that chromosome inversions are involved in adaptation to arid environments.

*The Goundry form.* Genetic analysis of *A. gambiae* larvae from roadside pools in Burkina Faso and adults collected from inside nearby houses revealed the occurrence of a genetically distinct population present in the larval sample, but absent from adult collections [60]. The larval population differed from the adult population with respect to the distribution of microsatellite alleles (FST=0.15), the presence of M/S hybrids (35% in the larval population, <1% in adults) and in the frequency of the 2L*a* inversion (2L*a* = 58% in larval population, 96% in adults). This distinct larval population is called the Goundry form, after one of the village collection sites. Based on these results it is supposed that the Goundry form is a unique form in which the adults rest nearly exclusively outdoors (exophilic) and which, although they carry the X-linked genetic markers that distinguish the M and S forms, the assortative mating associated with these markers is absent. Adults of the Goundry form have never been collected. Adults reared from larvae of the Goundry form were found to have increased susceptibility to infection with *P. falciparum* in laboratory experiments. [60]

## **7. Future directions**

**Figure 6.** A: Stage 1 - Population/races with differential adaptation; reproductive isolation (RI) not apparent. Green box represents diverged loci specific to Population 1 (Pop1) and Blue represents diverged loci specific to Pop2. Arrows indicate regions of gene flow. B: Stage 2 - Transition between races and species with some degree of RI; population may fuse or diverge. C: Stage 3 - Divergent populations beyond the point of fusion but still share a portion of their

Although most discussions consider M and S as the major and biologically relevant subdivi‐ sions of *A. gambiae* there is evidence that the two can be further subdivided into population

*Subdivision within the S form.* In a continent-wide survey Lehmann et al. [73] found that S form populations fall into two well defined clades, based on analysis of microsatellite DNA. They refer to these clades as the Northwest (Nigeria, Gabon, Democratic Republic of Congo, NW Kenya) and Southeast (SW Kenya, Tanzania, Malawi) divisions. Wang-Sattler et al. [37] also conducted an analysis based on microsatellite DNA and likewise report that the S form in eastern Africa (Kenya) are distinct from S form populations in the west (Mali). In addition to the East vs. West division between allopatric S form populations is the division of sympatric S form populations in Mali. These are described in detail above (Section 2). In brief, the S form in Mali is divided into the *Bamako* and *Savanna* chromosomal forms which display strong asssortative mating where they occur in sympatry at sites along the Niger River ([21], also see Figure 4). These two populations can be distinguished by the *j* inversion, which is fixed in the *Bamako* form and absent in the *Savanna*. Interestingly, although the two share the X-linked

genome via gene flow; good species. D: Stage 4 - Species with complete RI. Adapted from [79].

**6. Further sub-divisions within molecular forms**

groups that are significantly diverged.

186 *Anopheles* Anopheles mosquitoes - New insights into malaria vectors mosquitoes - New insights into malaria vectors

Reconciliation of the opposing speciation models and clarification of new "forms" await the resolution of a number of outstanding questions concerning interactions between the M and S forms. It is clear that the determination of the frequency of hybrid individuals requires that individuals be identified using multi-locus genotypes at unlinked loci, such as those employed by White et al. [16], as opposed to the widely used single locus X-linked markers. This would allow not only the recognition of F1 hybrids but backcross individuals as well. Determination of the frequencies of both F1 and backcross genotypes would provide information on the level of introgression. Moreover, multi-locus approach will allow identification of hybrid males. The application of this method to populations throughout the sympatric range of M and S would allow a description of spatial heterogeneity in levels of introgression that could be related to key environmental parameters that include mating cues that sustain assortative mating within forms as well as conditions that favor the survival of hybrid genotypes.

## **Author details**

Gregory C. Lanzaro\* and Yoosook Lee

\*Address all correspondence to: gclanzaro@ucdavis.edu

Department of Pathology, Microbiology and Immunology School of Veterinary Medicine University of California Davis, CA, USA

## **References**


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**Author details**

Gregory C. Lanzaro\*

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## **Advances and Perspectives in the Study of the Malaria Mosquito** *Anopheles funestus*

Ibrahima Dia, Moussa Wamdaogo Guelbeogo and Diego Ayala

Additional information is available at the end of the chapter

http://dx.doi.org/10.5772/55389

## **1. Taxonomy, biology and distribution of the species within the Funestus Group**

#### **1.1. Introduction**

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PMC3233708. Epub 2011/12/28. eng.

196 *Anopheles* Anopheles mosquitoes - New insights into malaria vectors mosquitoes - New insights into malaria vectors

*Anopheles funestus* Giles, 1900 is considered one of the most proficient malaria vectors world‐ wide [1]. It thrives in a wide range of habitats through the Afrotropical Region. Largely neglected with regard to its counterpart *Anopheles gambiae, An. funestus* cannot be ignored in any comprehensive control program aiming at the eradication of malaria from the African continent. Its transmission role goes beyond that of secondary vector, surpassing *An. gam‐ biae* in many parts of Africa [2]. One of the main reasons of this inattention is the difficulty of adapting this species to standard insectary conditions, despite noteworthy molecular and epidemiological advances over the past three decades. Currently, substantial evidence shows that a group of species belongs to the taxon "*An. funestus*", with different morphological, behavioural and epidemiological characteristics.

#### **1.2. The Funestus Group**

The term "Funestus Group" was first coined in its strictest sense by Gillies and De Meillon [3] to designate a group of species morphologically close to *An. funestus*. Seventy years after the first description of *An. funestussensu stricto* (hereafter *An. funestus*) by Giles in 1900, Mick Gillies and Botha De Meillon developed a new classification based on larva, pupa and adult stages. In fact, first suspicions of the existence of heterogeneity within *An. funestus* populations came from the early 1930's [4, 5]. They stated, based on larval studies, the presence of '*varieties'*, most of them were subsequently recognized as species within the group. These species showed

© 2013 Dia et al.; licensee InTech. This is an open access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited. © 2013 Dia et al.; licensee InTech. This is a paper distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

minor or no morphological differences at adult stage. They were then classified under the Funestus Group and their recognition was based on the identification of eggs, larvae or pharyngeal armature [3]. However, in Southern and Eastern Africa, several populations of outdoors resting mosquitoes were distinguishable from *An. funestus* by small morphological characters at the adult stage, while the larva were indistinguishable. These taxonomical observations were later confirmed by cytogenetic studies as different species of *An. funestus* [6-8].

Given the laborious nature of morphological and cytogenetic techniques, several studies were undertaken for the research of simple and useful molecular identification tools [9-12]. These techniques have the advantage to be applicable to all developmental stages. On the basis of morphological [13, 14] and molecular studies [15, 16], the status and position of each species within the Funestus Group was revisited. It is now accepted that *An. funestus* belongs to a group composed of five subgroups of which 3 groups containing 13 species are present in the Afrotropical region (Table 1) [17].


**Table 1.** Summary of ecological characteristics of Funestus Group in Africa.

#### **1.3. Geographical distribution**

Summary of ecological characteristics of Funestus group in Africa.

Among the species of the Funestus Group, *An. funestus*, *An. leesoni* and *An. rivulorum* exhibit the widest distribution. They are traditionally represented throughout the entire sub-Saharan Africa [1, 3]. Figure 1 presents the predicted distribution of these species [11, 12]. *Anopheles funestus* is found virtually all across the continent (Fig. 1A). Being predominantly a savannah mosquito [18], this malaria vector is present in many other areas, such as high altitude zones (900 m in Madagascar [19], 1400 m in Central Africa [20] and up to 2000 m in Kenya [21]) and forested areas of West and Central Africa [18, 22-25]. Moreover, it can inhabit extreme dry conditions in the Sahel, when suitable breeding place are available, such as human-made irrigation zones [26, 27]. On the other hand, *An. funestus* is scarce or completely absent along the coast [18]. *Anopheles funestus* disappeared from several parts of Africa after adverse climatic conditions (i.e recurrent droughts) and/or vector control programs [28]. Unfortunately, this mosquito gradually re-emerged once control measures stopped or suitable environmental conditions re-appeared [29-32], evidencing its extraordinary environmental plasticity and dispersion ability.

The other species of the group exhibit locally defined distribution (Fig. 1B, C). *Anopheles parensis*, *An. confusus* and *An. aruni* are localized in East Africa [33, 34]. In West and Central Africa, we find *An. rivulorum*-like and *An. brucei* [11, 12]. Finally, in Southern Africa, we find *An. vaneedeni, An. parensis* again, *An. fuscivenosus, An. funestus*-like and *An. longipalpis* types A (South Africa) and C (Zambia) [1, 35, 36]. Certainly, these records are based on sampling efforts, and we might expect changes in the number of species within the group as well in their distribution.

**Figure 1.** Distribution of the 13 species of the Funestus Group in Africa, A: *Anopheles funestus*, (modified from [37]); B: *An. leesoni*, *An. longipalpis* (type A and C), *An. aruni* and *An. parensis* (Courtesy of Dr. S. Manguin), C: *An. rivolorum, An. rivolorum-like, An. funestus-like, An. vaneedeni, An. fuscivenosus* and *An. brucei* (Courtesy of Dr. S. Manguin).

#### **1.4. Breeding place**

minor or no morphological differences at adult stage. They were then classified under the Funestus Group and their recognition was based on the identification of eggs, larvae or pharyngeal armature [3]. However, in Southern and Eastern Africa, several populations of outdoors resting mosquitoes were distinguishable from *An. funestus* by small morphological characters at the adult stage, while the larva were indistinguishable. These taxonomical observations were later confirmed by cytogenetic studies as different species of *An. funestus*

Given the laborious nature of morphological and cytogenetic techniques, several studies were undertaken for the research of simple and useful molecular identification tools [9-12]. These techniques have the advantage to be applicable to all developmental stages. On the basis of morphological [13, 14] and molecular studies [15, 16], the status and position of each species within the Funestus Group was revisited. It is now accepted that *An. funestus* belongs to a group composed of five subgroups of which 3 groups containing 13 species are present in the

**Subgroup Species Geographical distribution Host preference Vector role**

Among the species of the Funestus Group, *An. funestus*, *An. leesoni* and *An. rivulorum* exhibit the widest distribution. They are traditionally represented throughout the entire sub-Saharan Africa [1, 3]. Figure 1 presents the predicted distribution of these species [11, 12]. *Anopheles funestus* is found virtually all across the continent (Fig. 1A). Being predominantly a savannah mosquito [18], this malaria vector is present in many other areas, such as high altitude zones (900 m in Madagascar [19], 1400 m in Central Africa [20] and up to 2000 m in Kenya [21]) and forested areas of West and Central Africa [18, 22-25]. Moreover, it can inhabit extreme dry

*An. funestus* continental anthropophilic major *An. funestusͲlike* localƵŶŬŶŽǁŶƵŶŬŶŽǁŶ *An. aruni* localƵŶŬŶŽǁŶƵŶŬŶŽǁŶ *An. confusus* regional zoophilic unknowŶ *An. parensis* regionalƵŶŬŶŽǁŶ minor *An. vaneedeni* localƵŶŬŶŽǁŶ unknowŶ *An. longipalpis* type C local zoophilic unknown *An. leesoni* continental zoophilic minor *An. longipalpis* type A local zoophilic unknown *An. rivulorum* continental zoophilic minor *An. rivulorumͲlike* localƵŶŬŶŽǁŶƵŶŬŶŽǁŶ *An. brucei* local unknowŶƵŶŬŶŽǁŶ *An. fuscivenosus* localƵŶŬŶŽǁŶƵŶŬŶŽǁŶ

**African species of the Funestus Group**

[6-8].

Afrotropical region (Table 1) [17].

198 *Anopheles* Anopheles mosquitoes - New insights into malaria vectors mosquitoes - New insights into malaria vectors

Funestus

Minimus

Rivulorum

Summary of ecological characteristics of Funestus group in Africa.

**1.3. Geographical distribution**

**Table 1.** Summary of ecological characteristics of Funestus Group in Africa.

*Anopheles funestus* breeds in natural/artificial permanent and semi-permanent water bodies with floating or emerging vegetation. However, in areas with both vegetation types, this mosquito prefers the latter one [3]. Natural breeding occurs in edges of swamps, in weedy and grassy parts of rivers, streams, furrows, ditches and ponds. The presence of vegetation is crucial for mosquito breeding (Fig 2. A-C). Mainly because aquatic stages have a marked preference for shaded habitats and can barely survive in water bodies directly exposed to sunlight. Artificial breeding opportunities include rice fields, wells and domestic watercontainers [3]. The main limiting factors to their development include salinity, extreme like [1, 3, 36, 40].

in weedy and grassy parts of rivers, streams, furrows, ditches and ponds. The presence of vegetation is crucial for breeding (Fig 2. A-C), mainly because aquatic stages have a marked preference for shaded habitats and can barely survive in water bodies directly exposed to sunlight. Artificial breeding opportunities include rice fields, wells and domestic water-containers [3]. The main limiting factors to their development include salinity, extreme temperatures and sometimes, heavy rains. For the other species within the Funestus Group, the biology of aquatic stages is poorly understood. The larva of *An. leesoni*, *An. rivulorum* and *An. vaneedeni* are often found in association with those of *An. funestus*. In Kenya, *An. rivulorum* replaced *An. funestus* in rice fields after indoor residual [38]. The presence of vegetation appears to be essential too. These breeding sites are represented generally by slowmoving backwaters of grassy rivers and tide pools. In western Kenya, larva of *An. rivulorum* were recently found in hyacinth water protected by trees [39]. Similarly, *An. parensis* develops in permanent swamps and ponds between the reeds and the emergent vegetation. However, *An. parensis* is a species of stagnant water that has never been found in rivers. The larva were always collected in marshes, temporary and permanent ponds, among reeds and emerging vegetation [1, 3]. *Anopheles aruni* breeds in ponds, rice fields or ditches near human habitations. Larva of *An. brucei* were found in streams of forested river beds. A*nopheles confusus*, on the other hand, breeds in the vegetation of the edges of slow flowing rivers. *Anopheles longipalpis* prefers relatively calm water with abundant aquatic vegetation on the banks of fast-flowing rivers [3]. In many occasions, breeding places are very similar

Figure 2. **Breeding sites of** *Anopheles funestus* **(Photos D. Ayala, Cameroon).** *A:* Pitoa (Cameroon) is situated in the northern dry savannah, close to a permanent human-made lake, which provides a year-round breeding site for *An. funestus*. *B:* Tibati (Cameroon) is located in the central highlands of the country. *Anopheles funestus* breeds year-round in the lake, which provides shaded areas thanks to the lake vegetation. *C:* Mfou (Cameroon) is situated in the southern rainforest, in the surroundings of Yaoundé. The artificial water-body provides an excellent breeding site for *An. funestus*, making it the major vector of the village. **Figure 2.** Breeding sites of *Anopheles funestus* (Photos D. Ayala, Cameroon). *A:* Pitoa (Cameroon) is situated in the northern dry savannah, close to a permanent human-made lake, which provides a year-round breeding site for *An. funestus*. *B:* Tibati (Cameroon) is located in the central highlands of the country. *Anopheles funestus* breeds year-round in the lake, which provides shaded areas thanks to the lake vegetation. *C:* Mfou (Cameroon) is situated in the southern rainforest, in the surroundings of Yaoundé. The artificial water-body provides an excellent breeding site for An. funes‐ *tus*, making it the major vector of the village.

temperatures and sometimes, heavy rains. For the other species within the Funestus Group, the biology of aquatic stages is poorly understood. The larva of *An. leesoni*, *An. rivulorum* and *An. vaneedeni* are often found in association with those of *An. funestus*. In Kenya, *An. rivulo‐ rum* replaced *An. funestus* in rice fields after indoor residual spraying [38]. The presence of vegetation appears to be essential too. These breeding sites are represented generally by slowmoving backwaters of grassy rivers and tide pools. In western Kenya, larva of *An. rivulorum* were recently found in hyacinth water protected by trees [39]. Similarly, *An. parensis* develops in permanent swamps and ponds between the reeds and the emergent vegetation. However, *An. parensis* is a species of stagnant water that has never been found in rivers. The larva were always collected in marshes, temporary and permanent ponds, among reeds and emerging vegetation [1, 3]. *Anopheles aruni* breeds in ponds, rice fields or ditches near human habitations. Larva of *An. brucei* were found in streams of forested river beds. A*nopheles confusus*, on the other hand, breeds in the vegetation of the edges of slow flowing rivers. *Anopheles longipalpis* **1.5. Resting behaviour and host feeding preference: Their impact on vector capacity**  Despite the morphological similarities that exist between members of the group, these species show extreme behavioural differences that affect their vectorial capacities. To date, all malaria transmission studies have shown that *An. funestus* is the main malaria vector in the group, with infection rates up to 11% [41] and exceptionally 50% [42]. *Anopheles funestus* has late-night biting patterns, commonly between midnight and the early hours of the morning [22, 43, 44]. It is also the most endophilic and anthropophilic member of the Funestus group [45-47]. In savanna areas where its breeding sites are rain-dependant, *An. funestus* follows in peak abundance its counterpart *An. gambiae*, therefore extending malaria transmission from the beginning to the first part of the dry season [48, 49]. Overall, *An. funestus* shows fairly consistent host feeding preferences (human) and resting behaviour (indoor) throughout its entire range. However, behavioural differences linked to chromosomal polymorphisms have been documented. For instance, Lochouarn *et al.* [50] reported a west-east gradient of human to animal biting preference, corresponding  prefers relatively calm water with abundant aquatic vegetation on the banks of fast-flowing rivers [3]. In many occasions, breeding places are very similar to *An. funestus*. Unfortunately, no information exists about breeding places for *An. fuscivenosus*, *An. rivulorum*-like and *An. funestus*-like [1, 3, 36, 40].

#### **1.5. Resting behaviour and host feeding preference: Their impact on vector capacity**

Despite the morphological similarities that exist between members of the group, these species show extreme behavioural differences that affect their vectorial capacities. To date, all malaria transmission studies have shown that *An. funestus* is the main malaria vector in the group, with infection rates up to 11% [41] and exceptionally 50% [42]. *Anopheles funestus* has late-night biting patterns, commonly between midnight and the early hours of the morning [22, 43, 44]. It is also the most endophilic and anthropophilic member of the Funestus Group [45-47]. In savanna areas where its breeding sites are rain-dependant, *An. funestus* follows in peak abundance its counterpart *An. gambiae*, therefore extending malaria transmission from the beginning to the first part of the dry season [48, 49]. Overall, *An. funestus* shows fairly consistent host feeding preferences (human) and resting behaviour (indoor) throughout its entire range. However, behavioural differences linked to chromosomal polymorphisms have been docu‐ mented. For instance, Lochouarn *et al.* [50] reported a west-east gradient of human to animal biting preference, corresponding to chromosomal polymorphisms that also follow this cline. In Burkina Faso, different chromosomal inversion combinations (chromosomal forms, see below) were associated with different resting and biting activities [42]. These studies showed that carriers of inverted arrangements on the arm 2R and 3R feed predominantly on humans (anthropophilic) and rest inside dwellings, while the standard counterpart exhibit higher levels of zoophily and exophily (Guelbeogo, pers. Comm.). In Madagascar, the carriers of inverted arrangements 3Ra and 3Rb were less anthropophilic than carriers of standard arrangements [51]. In Senegal, the population of mosquitoes with inverted arrangements 3Ra and 3Rb was also more zoophilic. However, this heterogeneity in host preference might also be related to specific local conditions, such as host availability [52] or indoor microclimatic conditions (i.e. humidity).

The other species of the group are mainly zoophilic, but can occasionally feed on hu‐ mans [3]. *Anopheles rivulorum* has been incriminated as a malaria vector in Tanzania [53]. Indeed, this species was found naturally infected by *Plasmodium falciparum.* However, this species is mainly zoophilic (77% animal hosts) and shows a lower longevity compared to *An. funestus*. Positive infected specimens of *An. rivulorum* were also observed in coastal Tanzania by Temu et al. [54]. This study also found positive specimens of *An. leesoni* and *An. parensis* to *P. falciparum*, suggesting a secondary role of these mosquitoes in malaria transmission. *Plasmodium falciparum* infected *An. parensis* specimens were also observed during an entomological study in South Africa using an Enzyme-Linked Immunosorbent Assay (ELISA) [55]. *Anopheles vaneedeni* feeds rarely on humans outdoors (1.22%). Al‐ though experimentally infected with *P. falciparum* in the laboratory, it has never been found involved in transmission in natural conditions [56]. *Anopheles longipalpis* has never been involved in malaria transmission [1, 3, 57]. In East Africa (Tanzania and Ethiopia), different

temperatures and sometimes, heavy rains. For the other species within the Funestus Group, the biology of aquatic stages is poorly understood. The larva of *An. leesoni*, *An. rivulorum* and *An. vaneedeni* are often found in association with those of *An. funestus*. In Kenya, *An. rivulo‐ rum* replaced *An. funestus* in rice fields after indoor residual spraying [38]. The presence of vegetation appears to be essential too. These breeding sites are represented generally by slowmoving backwaters of grassy rivers and tide pools. In western Kenya, larva of *An. rivulorum* were recently found in hyacinth water protected by trees [39]. Similarly, *An. parensis* develops in permanent swamps and ponds between the reeds and the emergent vegetation. However, *An. parensis* is a species of stagnant water that has never been found in rivers. The larva were always collected in marshes, temporary and permanent ponds, among reeds and emerging vegetation [1, 3]. *Anopheles aruni* breeds in ponds, rice fields or ditches near human habitations. Larva of *An. brucei* were found in streams of forested river beds. A*nopheles confusus*, on the other hand, breeds in the vegetation of the edges of slow flowing rivers. *Anopheles longipalpis*

**Figure 2.** Breeding sites of *Anopheles funestus* (Photos D. Ayala, Cameroon). *A:* Pitoa (Cameroon) is situated in the northern dry savannah, close to a permanent human-made lake, which provides a year-round breeding site for *An. funestus*. *B:* Tibati (Cameroon) is located in the central highlands of the country. *Anopheles funestus* breeds year-round in the lake, which provides shaded areas thanks to the lake vegetation. *C:* Mfou (Cameroon) is situated in the southern rainforest, in the surroundings of Yaoundé. The artificial water-body provides an excellent breeding site for An. funes‐

**1.5. Resting behaviour and host feeding preference: Their impact on vector capacity** 

*An. funestus*, making it the major vector of the village.

*tus*, making it the major vector of the village.

Figure 2. **Breeding sites of** *Anopheles funestus* **(Photos D. Ayala, Cameroon).** *A:* Pitoa (Cameroon) is situated in the northern dry savannah, close to a permanent human-made lake, which provides a year-round breeding site for *An. funestus*. *B:* Tibati (Cameroon) is located in the central highlands of the country. *Anopheles funestus* breeds year-round in the lake, which provides shaded areas thanks to the lake vegetation. *C:* Mfou (Cameroon) is situated in the southern rainforest, in the surroundings of Yaoundé. The artificial water-body provides an excellent breeding site for

Despite the morphological similarities that exist between members of the group, these species show extreme behavioural differences that affect their vectorial capacities. To date, all malaria transmission studies have shown that *An. funestus* is the main malaria vector in the group, with infection rates up to 11% [41] and exceptionally 50% [42]. *Anopheles funestus* has late-night biting patterns, commonly between midnight and the early hours of the morning [22, 43, 44]. It is also the most endophilic and anthropophilic member of the Funestus group [45-47]. In savanna areas where its breeding sites are rain-dependant, *An. funestus* follows in peak abundance its counterpart *An. gambiae*, therefore extending malaria transmission from the beginning to the first part of the dry season [48, 49]. Overall, *An. funestus* shows fairly consistent host feeding preferences (human) and resting behaviour (indoor) throughout its entire range. However, behavioural differences linked to chromosomal polymorphisms have been documented. For instance, Lochouarn *et al.* [50] reported a west-east gradient of human to animal biting preference, corresponding

in weedy and grassy parts of rivers, streams, furrows, ditches and ponds. The presence of vegetation is crucial for breeding (Fig 2. A-C), mainly because aquatic stages have a marked preference for shaded habitats and can barely survive in water bodies directly exposed to sunlight. Artificial breeding opportunities include rice fields, wells and domestic water-containers [3]. The main limiting factors to their development include salinity, extreme temperatures and sometimes, heavy rains. For the other species within the Funestus Group, the biology of aquatic stages is poorly understood. The larva of *An. leesoni*, *An. rivulorum* and *An. vaneedeni* are often found in association with those of *An. funestus*. In Kenya, *An. rivulorum* replaced *An. funestus* in rice fields after indoor residual [38]. The presence of vegetation appears to be essential too. These breeding sites are represented generally by slowmoving backwaters of grassy rivers and tide pools. In western Kenya, larva of *An. rivulorum* were recently found in hyacinth water protected by trees [39]. Similarly, *An. parensis* develops in permanent swamps and ponds between the reeds and the emergent vegetation. However, *An. parensis* is a species of stagnant water that has never been found in rivers. The larva were always collected in marshes, temporary and permanent ponds, among reeds and emerging vegetation [1, 3]. *Anopheles aruni* breeds in ponds, rice fields or ditches near human habitations. Larva of *An. brucei* were found in streams of forested river beds. A*nopheles confusus*, on the other hand, breeds in the vegetation of the edges of slow flowing rivers. *Anopheles longipalpis* prefers relatively calm water with abundant aquatic vegetation on the banks of fast-flowing rivers [3]. In many occasions, breeding places are very similar to *An. funestus*. Unfortunately, no information exists about breeding places for *An. fuscivenosus*, *An. rivulorum*-like and *An. funestus*-

like [1, 3, 36, 40].

*B*

*A C*

200 *Anopheles* Anopheles mosquitoes - New insights into malaria vectors mosquitoes - New insights into malaria vectors

authors have reported human feeding behaviour of *An. longipalpis* from indoor and outdoor collections [58-60]. Recently, Kent et al., [57] reported that even when found in large numbers resting indoors together with *An. funestus* in Zambia, *An. longipalpis* remains predominantly zoophilic.

## **2. Insecticide susceptibility and vector control**

Because of its highly anthropophilic and endophilic behaviour, *An. funestus* has been an "easy" target in malaria control programs (i.e. insecticide treated materials or indoor residual spraying). *Anopheles funestus* has developed insecticide resistance in many parts of the African continent [61-64]. To date, *An. funestus* has been shown resistant to pyrethroids, carbamates and DDT. The first documented reports on insecticide resistance in this malaria mosquito (mainly to BHC, dieldrin, and malathion) were in West Africa (Mali, Ghana, Benin), Central Africa (Cameroon) and East Africa (Kenya), following vector control programs [65-68]. Recent studies have shown that dieldrin resistance is still high in *An. funestus* populations from Burkina Faso, despite the fact that this insecticide is no longer used in public health [47]. In agreement with Burkina Faso results, Wondji et al. [69] documented *An. funestus* resistant populations to dieldrin in Cameroon due to the remaining presence of RdlR target-site mutation. With regard to pyrethroids, resistant *An. funestus* populations were first detected in Southern Africa, being at the origin of the malaria outbreaks in the late 1990's [31, 62]. Pyrethroid resistant populations for this mosquito were also reported in Ghana, West Africa, combined with carbamate resistance [70]. Altogether, it is now clearly established that *An. funestus* populations in Africa show resistance to at least the 4 insecticide classes recommended for vector control by WHO.

During the last decade, efforts have been made in order to unravel the molecular mechanisms involved in insecticide resistance. The mechanisms discovered involve insecticide detoxifica‐ tion by one or multiple metabolic pathways mediated by glutathione S-transferases (GST), monooxygenases and/or esterases [61, 71-73]. No evidence for the presence of L1014F *kdr* mutation or G119S *Ace-1* mutation has been detected in *An. funestus* [63, 64, 71, 72]. However, a multiple insecticide resistance profile has been recently observed in Benin [74]. Insecticide resistance is an threat to effective malaria control. With the advent of malaria control program through the use of LLINs (Long Lasting Insecticidal Nets) and IRS (Indoor Residual Spraying), the presence of insecticide resistant populations should be carefully monitored. It would improve the implementation and management of current and future malaria vector control programs in Africa. In this context, a novel approach using the pyrrole insecticide chlorfenapyr against pyrethroid resistant *An. funestus* populations has led to valuable results [75]. An important challenge for the study of molecular mechanisms of insecticide resistance is the development and maintenance of laboratory colonies. To date, only two colonies are currently maintained at insectarium conditions, coming from southern Africa [76], although, some progress has been made and new strains have been established in Burkina Faso (Sagnon *et al.,* pers. comm.).

## **3. Molecular tools**

#### **3.1. Introduction**

authors have reported human feeding behaviour of *An. longipalpis* from indoor and outdoor collections [58-60]. Recently, Kent et al., [57] reported that even when found in large numbers resting indoors together with *An. funestus* in Zambia, *An. longipalpis* remains

Because of its highly anthropophilic and endophilic behaviour, *An. funestus* has been an "easy" target in malaria control programs (i.e. insecticide treated materials or indoor residual spraying). *Anopheles funestus* has developed insecticide resistance in many parts of the African continent [61-64]. To date, *An. funestus* has been shown resistant to pyrethroids, carbamates and DDT. The first documented reports on insecticide resistance in this malaria mosquito (mainly to BHC, dieldrin, and malathion) were in West Africa (Mali, Ghana, Benin), Central Africa (Cameroon) and East Africa (Kenya), following vector control programs [65-68]. Recent studies have shown that dieldrin resistance is still high in *An. funestus* populations from Burkina Faso, despite the fact that this insecticide is no longer used in public health [47]. In agreement with Burkina Faso results, Wondji et al. [69] documented *An. funestus* resistant populations to dieldrin in Cameroon due to the remaining presence of RdlR target-site mutation. With regard to pyrethroids, resistant *An. funestus* populations were first detected in Southern Africa, being at the origin of the malaria outbreaks in the late 1990's [31, 62]. Pyrethroid resistant populations for this mosquito were also reported in Ghana, West Africa, combined with carbamate resistance [70]. Altogether, it is now clearly established that *An. funestus* populations in Africa show resistance to at least the 4 insecticide classes recommended

During the last decade, efforts have been made in order to unravel the molecular mechanisms involved in insecticide resistance. The mechanisms discovered involve insecticide detoxifica‐ tion by one or multiple metabolic pathways mediated by glutathione S-transferases (GST), monooxygenases and/or esterases [61, 71-73]. No evidence for the presence of L1014F *kdr* mutation or G119S *Ace-1* mutation has been detected in *An. funestus* [63, 64, 71, 72]. However, a multiple insecticide resistance profile has been recently observed in Benin [74]. Insecticide resistance is an threat to effective malaria control. With the advent of malaria control program through the use of LLINs (Long Lasting Insecticidal Nets) and IRS (Indoor Residual Spraying), the presence of insecticide resistant populations should be carefully monitored. It would improve the implementation and management of current and future malaria vector control programs in Africa. In this context, a novel approach using the pyrrole insecticide chlorfenapyr against pyrethroid resistant *An. funestus* populations has led to valuable results [75]. An important challenge for the study of molecular mechanisms of insecticide resistance is the development and maintenance of laboratory colonies. To date, only two colonies are currently maintained at insectarium conditions, coming from southern Africa [76], although, some progress has been made and new strains have been established in Burkina Faso (Sagnon *et*

predominantly zoophilic.

202 *Anopheles* Anopheles mosquitoes - New insights into malaria vectors mosquitoes - New insights into malaria vectors

for vector control by WHO.

*al.,* pers. comm.).

**2. Insecticide susceptibility and vector control**

In 2002, the genome of *An. gambiae* s.s. was publicly released [77]. This event had a very large impact on the better understanding of the complexity of the malaria system. Furthermore, the publication of the *An. gambiae* genome brought with itself a rapid development of new genetic tools, from molecular markers (i.e. SNPs chips, microarrays, microsatellites, etc) to transgenic mosquitoes, for instance. To date, no other malaria mosquito genome has been released but progress has been made, and soon (2013), the release of several *Anopheles* genomes, including *An. funestus* [78], is expected.

Three inherent characteristics of *An. funestus*, have hampered the study of this mosquito at the molecular level. First, its "eternal" role as second important malaria vector. For decades, *An. funestus* has been neglected with regard to its well-studied congener *An. gambiae*. With virtually the same geographical distribution as *An. gambiae* across the African continent, *An. funestus* has been many times overruled because its mosaic-like presence (see previous section in this chapter). However, its major role in malaria transmission has been evidenced throughout the continent, surpassing in a number of locations *An. gambiae* and *An. arabiensis*[2] in many places. Second, the extreme difficulties to breed *An. funestus* in standard insectary conditions. To date, as mentioned earlier in this chapter, there exist only two colonies of *An. funestus* with published records: FANG and FUMOZ (and its pyrethroid resistance counterpart FUMOZ-R), originat‐ ing from Angola and Mozambique, respectively [76, 79]. Both colonies have been recurrently used in insecticide resistance studies of *An. funestus* [74, 79, 80]. Indeed, it is one of these colonies (FUMOZ), which has been elected as reference *An. funestus* genome for sequencing [78]. Unfortunately and besides the numerous efforts in many parts of Africa, only one new colony has been colonized (Sagnon *et al.,* pers. comm.). Third, polytene chromosomes of this species exhibit a poor quality in comparison with *An. gambiae* [7]. The assembly of the *An. gambiae* genome was primarily based on techniques, which required the identification of probes through polytene chromosomes [77]. Although polytene chromosomes are readable, as several studies assert, however, the effort involved is very high and the rate of success, significantly lower.

Despite these challenges, and the lack of a publicly available *An. funestus* genome, several noteworthy molecular and genetic advances have been reached in this malaria mosquito during the last decade. These advances have been inspired by those previously achieved in *An. gambiae*. Particularly, we can distinguish two fields: molecular markers and expression profiling analysis.

#### **3.2. Molecular markers**

In the late 70's and beginning of the 80's, several studies revealed the importance of chromo‐ somal inversions as genetic markers to differentiate species within the Funestus Group [6, 7]. These results mirrored those obtained in the *An. gambiae* complex [81, 82]. But, we had to wait until the end of the 90's and the past decade to settle the role of the chromosomal inversions in local adaptation and speciation within *An. funestus* populations [42, 52, 83-86]. Despite its evident interest, the technical demands of traditional karyotype analysis, the low rate of success in chromosome preparations, and the sex- and stage-specific limitations, have hampered the proliferation of this kind of studies. Nowadays, the new advances in molecular karyotyping in *An. gambiae* (based on quick, low-cost and convenient PCR reactions) have relaunched an interest in this field [87, 88]. Together with new high-throughput technology, the *An. funestus* genome will undoubtedly open new possibilities to develop molecular karyotyp‐ ing in this mosquito.


**Table 2.** Summary of microsatellite loci in *An. funestus* modified from Wondji et al. [89].

Table 2. Summary of microsatellite loci in *An. funestus* 

members, such as *An. gambiae*, *Ae. aegypti* or *Cx. pipiens*.

**3.3. Expression profiles** 

workers to explore the genetic basis of insecticide resistance in this malaria vectors [79]. Several genes including the P450 cytochrome (CYP6P9a and CYP6P9b) were associated to DDT resistance by Quantitative Trait Loci (QTL) analysis using both markers [72]. The role of microsatellites in population genetic studies is discussed in other sections of this chapter (see below). Despite these advances, we are still far from *An. gambiae molecular advances*. For instance, in *An. funestus* 75 microsatellite loci have been identified, compared to 300 in *An. gambiae*. With regards to SNPs, 509 have been reported in *An. funestus* [79, 89], compared to 400,000 in *An. gambiae* [80]. In *An. funestus*, several genes have been recurrently involved in genetic studies: three nuclear genes (ITS1, ITS2 and D3) and another three mitochondrial genes (COI, COII and ND5). Nuclear genes have been involved in species differentiation within the Funestus Group [15, 16], while mitochondrial genes revealed signatures of incipient speciation between popula‐ tions of Burkina Faso [85]. Another kind of molecular markers, Single Nucleotide Polymor‐ phisms (SNPs), have been recently developed in this malaria mosquito. Wondji et al. [79]

Considering the lack of *An. funestus* genome, transcriptome analysis appeared as a suitable alternative to whole genome sequencing. This technique is significantly cheaper and provides important information at the gene transcript level. Moreover, it provides valuable molecular tools for the analysis of gene expression evolution and comparative analysis among other *Culicidae*

In 2007, Calvo et al., [96] investigated salivary gland genes from 916 cDNA clones coming from adult females. This study debuted the analysis of transcripts in this mosquito, providing important clues about the evolution of salivary gland proteins in blood feeding insects and Culicidae. In particular, a 30 KDa allergen family and several *mucins* were exclusively found in *Culicidae* when compared to *Aedes albopictus*, *Aedes aegypti* and *Culex pipiens quinquefasciatus*. Moreover, ten proteins and peptide families were only found in *Anopheles* when included in the analysis *An. gambiae*, *Anopheles stephensi* and *Anopheles darlingi*. Later, two new studies emerged with the aim to analyze the transcriptome evolution and differences in expression profile between insecticide susceptible and resistant phenotypes of *An. funestus*, respectively [80, 97]. While, Serazin et al. [97] used SANGER sequencing technology for this purpose, Gregory et al. [80] employed *de novo* expression profiling by 454 pyrosequencing. In general, these two studies were largely complementary and boosted the available genetic information in *An. funestus*. However, 454 pyrosequencing allowed parallel DNA sequencing and increased sequencing depth and genome coverage. For instance, Gregory et al. [80] improved the number of ESTs (Exressed Sequence Tags) from 2,846 [97] to 18,103 contigs. Regarding comparative analysis with other mosquitoes, both studies agreed on the fact that the highest similarity pattern remains with *An. gambiae*. Interestingly, the mean percentage of similarity differs drastically between functional groups. Two groups of housekeeping functions show the highest amino acid sequence conservation: protein synthesis and degradation. On the other hand, three groups of interest patently showed very low similarity scores, suggesting accelerated rates of evolution. These three functional categories – salivary, immunity and extracellular structures – may be driven by environmental selection pressures. For instance, selective pressures imposed by parasites could explain both the highest genetic variability and the lowest conservation of immune genes between *An. funestus and An. gambiae*. Alternatively, *de novo* 454 sequencing offered the opportunity to identify new SNPs. In this sense, 31,000 potential SNPs were discovered over 4.579 Mb of sequence, meaning one SNP every 70 bp [80]. Thus, expression profile studies led to

These molecular markers have been key in numerous advances. For instance, SNPs and microsatellites allowed to Wondji and co-

reported a genome-wide set of SNP markers from 50 genes. A total of 494 SNPs were identified, which were added to 15 SNPs previously discovered by analyzing sequence traces of 11 physically mapped DNA fragments of cytochrome P450s of *An. funestus*. However, to date, microsatellites are the most frequently employed molecular markers in *An. funestus* [89-92]. Seventy-five microsatellites have been developed, although, only 32 were successfully revisited by Wondji et al [89] (Table 2). They are widely distributed across the *An. funestus* genome. They have allowed the analysis of population genetic structure, gene flow and demographic events across Africa [93], from Senegal [40], Cameroon [83, 86], Kenya [94] to Madagascar [95], revealing important signatures of local adaptation, dispersion or speciation.

These molecular markers have been key in numerous advances. For instance, SNPs and microsatellites allowed to Wondji and co-workers to explore the genetic basis of insecticide resistance in this malaria vectors [79]. Several genes including the P450 cytochrome (CYP6P9a and CYP6P9b) were associated to DDT resistance by Quantitative Trait Loci (QTL) analysis using both markers [72]. The role of microsatellites in population genetic studies is discussed in other sections of this chapter (see below). Despite, we are still far from the molecular advances carried out on *An. gambiae*. For instance, in *An. funestus* 75 microsatellite loci have been identified, compared to 300 in *An. gambiae*. With regards to SNPs, 509 have been reported in *An. funestus* [79, 89], compared to 400,000 in *An. gambiae* [80].

#### **3.3. Expression profiles**

in local adaptation and speciation within *An. funestus* populations [42, 52, 83-86]. Despite its evident interest, the technical demands of traditional karyotype analysis, the low rate of success in chromosome preparations, and the sex- and stage-specific limitations, have hampered the proliferation of this kind of studies. Nowadays, the new advances in molecular karyotyping in *An. gambiae* (based on quick, low-cost and convenient PCR reactions) have relaunched an interest in this field [87, 88]. Together with new high-throughput technology, the *An. funestus* genome will undoubtedly open new possibilities to develop molecular karyotyp‐

**Chromosome Locus Accession number Forward primer Reverse primer Allele size**

FUNE AY6009 GACCGGTTCTGGTATCGTC ATCGAGTCACCCAATTCTCC 136–154 FUNQ AY6021 GCAAACTGCTAGTAAATGTTTCC \*ACACAACGCCACCACTATGA 84–98 AFND6 AF171036 GCTTCTTCTCCCCTAATCTG TCCTGCTTTTTAGTTTGTCG 184–212 AFUB15 AY029722 GATGCCGGGAGTAATAGCAA AGACAGCCCGTAGAACGGTA 155–191 AFND2 AF171032 ATAAACCCGTCCATTCCCTT CCTATGATTCGCTCCTGACA 131–151 AFND32 AY291367 GAAGCATTTTGGGTTAGACTC GCAGTTGTTTACCTTTCACTG 103–121 AFUB14 AY029721 ATCAGTGCTCCTCCACATCC CGTGGTTGGCAATGTTACTG 152–188 AFND17 AF171047 AAAACGCCACAAAGAGCAC CGGGTCAAATTCTACCGTAAG 129–157 AFUB4 AY029711 CTATCAGCAGCCGCCACA GATGCCGATGAGGAATGTTG 183–192 AFUB25 AY029723 GTGGAAACGGTGGTACTGT CGCCATGTAGCTAGGGTTTG 212–224 AFUB10 AY029717 TGTCCATGTACAACCGCAAC TTCTCCAGCATCATCAGCAC 195–210 AFND37 AY291373 GATCGATACAATAAGTGTAGAAATAAT TCACGATGTGCAACCTATAA 161–189 AFUB30 AY029737 GCCAGTTTGCAGAACCAAAT CTGCTGCTGATGTTGCTGAT 154–163 AFUB7 AY029714 ATGGGACGATGGATTACCAA GCCAGTTTGCAGAACCAAAT 220–223 AFUB16 AY029723 CGTGGATGGCAATGTTACTG TGCGACTTATCAGTGCTCCT 179–209 AFND21 AF171051 CCGCACACCAACTTACACTC TGGCGTGGGATTAAATAGG 96–104 AFUB13 GACTTCCGCCACAGAACATC CTCAGGCTCGCAGTAGGAGT 207–210 AFND19 AF171049 CAGAACCACTTCGATTCAAC CCTGCACTCAGAAACACAC 172–205 FUND AY6008 GCTAACTACTCCGAAGCGCT GATCGCAAAACTTCCGGTT 145–177 FUNI AY6013 \*GCAACTAAGCTGGGACAGGA GCATCTAACCCTGCTGCTT 181–197 AFND3 AF171033 ACGACTGTAACCACAACACC TAGTAGCGAAGGCGAAAGAT 171–195 FUNF AY6010 CCTTCAGTTTCGATTGGCG AATAAGATGCGACCGTGGC 104–118 AFND10 AF171040 TTTTTTCTTCCCGTGTTGC TACCATTTGATTACAGCGCC 114–146 AFUB17 AY029724 GAAAACCGTACGAACGATGG TGCGACAGTAGCACAGGGTA 187–196 AFUB1 AY029708 CAGCAGCAGCAGCAACAG GACGTTAGCATCTCCACCAG 266–269 AFUB12 AY029719 TGGGGAACTGGTCGTTAGAG CTGGTGATGGGATTGAGGAT 152–158 FUNK AY6015 GCGCTTCCGCAAACATAC ACTCACACCCCATTCTTGTG 184–202 263B12 AGTGCGTCAGAGTTTGAA TCGATTGATGGCGATGATAA 230–242 261H03 CGCTCAAACTGAAAGCGATA GGATGCGGAGATGATGTTGT 208–220 263A06 CGTTCGGTTTCGCTAACTGT CGTTCTATTTCGGGGTGTGT 210–220 AFUB21 AY029728 \*AACGCAGCAGTGGAGAGAAT AACACCAACCCTTGTTGTGC 224–230 AFND30 AY291369 GCCAGTTTGCAGAACCAAAT CTGCTGCTGATGTTGCTGAT 81–107

These molecular markers have been key in numerous advances. For instance, SNPs and microsatellites allowed to Wondji and coworkers to explore the genetic basis of insecticide resistance in this malaria vectors [79]. Several genes including the P450 cytochrome (CYP6P9a and CYP6P9b) were associated to DDT resistance by Quantitative Trait Loci (QTL) analysis using both markers [72]. The role of microsatellites in population genetic studies is discussed in other sections of this chapter (see below). Despite these advances, we are still far from *An. gambiae molecular advances*. For instance, in *An. funestus* 75 microsatellite loci have been identified, compared to 300 in *An. gambiae*. With regards to SNPs, 509 have been reported in *An. funestus* [79, 89], compared to

In *An. funestus*, several genes have been recurrently involved in genetic studies: three nuclear genes (ITS1, ITS2 and D3) and another three mitochondrial genes (COI, COII and ND5). Nuclear genes have been involved in species differentiation within the Funestus Group [15, 16], while mitochondrial genes revealed signatures of incipient speciation between popula‐ tions of Burkina Faso [85]. Another kind of molecular markers, Single Nucleotide Polymor‐ phisms (SNPs), have been recently developed in this malaria mosquito. Wondji et al. [79]

**Table 2.** Summary of microsatellite loci in *An. funestus* modified from Wondji et al. [89].

Considering the lack of *An. funestus* genome, transcriptome analysis appeared as a suitable alternative to whole genome sequencing. This technique is significantly cheaper and provides important information at the gene transcript level. Moreover, it provides valuable molecular tools for the analysis of gene expression evolution and comparative analysis among other *Culicidae*

In 2007, Calvo et al., [96] investigated salivary gland genes from 916 cDNA clones coming from adult females. This study debuted the analysis of transcripts in this mosquito, providing important clues about the evolution of salivary gland proteins in blood feeding insects and Culicidae. In particular, a 30 KDa allergen family and several *mucins* were exclusively found in *Culicidae* when compared to *Aedes albopictus*, *Aedes aegypti* and *Culex pipiens quinquefasciatus*. Moreover, ten proteins and peptide families were only found in *Anopheles* when included in the analysis *An. gambiae*, *Anopheles stephensi* and *Anopheles darlingi*. Later, two new studies emerged with the aim to analyze the transcriptome evolution and differences in expression profile between insecticide susceptible and resistant phenotypes of *An. funestus*, respectively [80, 97]. While, Serazin et al. [97] used SANGER sequencing technology for this purpose, Gregory et al. [80] employed *de novo* expression profiling by 454 pyrosequencing. In general, these two studies were largely complementary and boosted the available genetic information in *An. funestus*. However, 454 pyrosequencing allowed parallel DNA sequencing and increased sequencing depth and genome coverage. For instance, Gregory et al. [80] improved the number of ESTs (Exressed Sequence Tags) from 2,846 [97] to 18,103 contigs. Regarding comparative analysis with other mosquitoes, both studies agreed on the fact that the highest similarity pattern remains with *An. gambiae*. Interestingly, the mean percentage of similarity differs drastically between functional groups. Two groups of housekeeping functions show the highest amino acid sequence conservation: protein synthesis and degradation. On the other hand, three groups of interest patently showed very low similarity scores, suggesting accelerated rates of evolution. These three functional categories – salivary, immunity and extracellular structures – may be driven by environmental selection pressures. For instance, selective pressures imposed by parasites could explain both the highest genetic variability and the lowest conservation of immune genes between *An. funestus and An. gambiae*. Alternatively, *de novo* 454 sequencing offered the opportunity to identify new SNPs. In this sense, 31,000 potential SNPs were discovered over 4.579 Mb of sequence, meaning one SNP every 70 bp [80]. Thus, expression profile studies led to

ing in this mosquito.

204 *Anopheles* Anopheles mosquitoes - New insights into malaria vectors mosquitoes - New insights into malaria vectors

Chr. X

Chr. 2

Chr. 3

Table 2. Summary of microsatellite loci in *An. funestus* 

Unknown

members, such as *An. gambiae*, *Ae. aegypti* or *Cx. pipiens*.

400,000 in *An. gambiae* [80].

**3.3. Expression profiles** 

Considering the lack of *An. funestus* genome, transcriptome analysis appeared as a suitable alternative to whole genome sequencing. This technique is significantly cheaper and provides important information at the gene transcript level. Moreover, it provides valuable molecular tools for the analysis of gene expression evolution and comparative analysis among other Culicidae members, such as *An. gambiae*, *Ae. aegypti* or *Cx. pipiens*.

In 2007, Calvo et al., [96] investigated salivary gland genes from 916 cDNA clones coming from adult females. This study debuted the analysis of transcripts in this mosquito, providing important clues about the evolution of salivary gland proteins in blood feeding insects and Culicidae. In particular, a 30 KDa allergen family and several *mucins* were exclusively found in *Culicidae* when compared to *Aedes albopictus*, *Aedes aegypti* and *Culex pipiens quinquefascia‐ tus*. Moreover, ten proteins and peptide families were only found in *Anopheles* when included in the analysis *An. gambiae*, *Anopheles stephensi* and *Anopheles darlingi*. Later, two new studies emerged with the aim to analyze the transcriptome evolution and differences in expression profile between insecticide susceptible and resistant phenotypes of *An. funestus*, respectively [80, 97]. While, Serazin et al. [97] used SANGER sequencing technology for this purpose, Gregory et al. [80] employed *de novo* expression profiling by 454 pyrosequencing. In general, these two studies were largely complementary and boosted the available genetic information in *An. funestus*. However, 454 pyrosequencing allowed parallel DNA sequencing and increased sequencing depth and genome coverage. For instance, Gregory et al. [80] improved the number of ESTs (Expressed Sequence Tags) from 2,846 [97] to 18,103 contigs. Regarding comparative analysis with other mosquitoes, both studies agreed on the fact that the highest similarity pattern remains with *An. gambiae*. Interestingly, the mean percentage of similarity differs drastically between functional groups. Two groups of housekeeping functions show the highest amino acid sequence conservation: protein synthesis and degradation. On the other hand, three groups of interest patently showed very low similarity scores, suggesting accel‐ erated rates of evolution. These three functional categories – salivary, immunity and extrac‐ ellular structures – may be driven by environmental selection pressures. For instance, selective pressures imposed by parasites could explain both the highest genetic variability and the lowest conservation of immune genes between *An. funestus and An. gambiae*. Alternatively, *de novo* 454 sequencing offered the opportunity to identify new SNPs. In this sense, 31,000 potential SNPs were discovered over 4.579 Mb of sequence, meaning one SNP every 70 bp [80]. Thus, expression profile studies led to identify genes under selective pressures (i.e. insecticide resistance, immunity genes) and might generate new functional genomic tools (i.e. microarrays or SNP platforms) while we wait for future genomic sequencing of *An. funestus*.

## **4. Population genetic structure across Africa**

#### **4.1. Introduction**

In malaria mosquitoes, population genetics have been revealed as an excellent tool for implementation of vector control programs. The study of gene flow among vector populations allows the analysis of mosquitoes' movement in natural populations, and therefore, how those populations are segregated. They can, for instance, assist to follow the expansion of genes of interest, such as those that confer insecticide resistance [98], or potentially help to introduce transgenic mosquitoes, refractory to parasite infection [99, 100]. On the other hand, these population genetic studies might be useful to investigate the genetic basis of speciation and/or local adaptation processes. They evidence a considerable importance in vector control measures [101].

The biology of *An. funestus* has supported several "*a priories*" about its population structure in natural conditions. As mentioned previously in this chapter, this malaria mosquito mainly breeds in permanent or semi-permanent water bodies, such as rice fields, swamps or artificial lakes, always linked to human presence (see above). Moreover, this mosquito has exhibited a very slow recolonization power of those areas treated with insecticide. Both characteristics have led to assume the population subdivision of *An. funestus*. In this section, we will discuss the population structure of this malaria vector across Africa as revealed by two types of markers: chromosomal inversions and molecular markers.

#### **4.2. Cytogenetic studies**

The study of chromosomal rearrangements – cytogenetics – of *An. funestus* debuted early in the 1980's [6, 7], preceded by the success of this kind of studies in its congener *An. gambiae* [81, 82, 102]. It allowed differentiating members of the Funestus Group, avoiding the challenging interpretation of taxo-morphological rules. Green & Hunt [7] and Green [6] showed differences in the chromosomal polymorphism within the species of the group. As in *An. gambiae*, several chromosomal inversions were species-specific, while other inversions were polymorphic in some species and fixed in others. Although, other cytogenetic studies appeared in the mean‐ time, we had to wait until 2001 when Sharakhov et al. [103] finally established the chromosome map of this species (Fig. 2), based on comparisons to the *An. gambiae* map [102].

**Figure 3.** Chromosome map of *An. funestus*

drastically between functional groups. Two groups of housekeeping functions show the highest amino acid sequence conservation: protein synthesis and degradation. On the other hand, three groups of interest patently showed very low similarity scores, suggesting accel‐ erated rates of evolution. These three functional categories – salivary, immunity and extrac‐ ellular structures – may be driven by environmental selection pressures. For instance, selective pressures imposed by parasites could explain both the highest genetic variability and the lowest conservation of immune genes between *An. funestus and An. gambiae*. Alternatively, *de novo* 454 sequencing offered the opportunity to identify new SNPs. In this sense, 31,000 potential SNPs were discovered over 4.579 Mb of sequence, meaning one SNP every 70 bp [80]. Thus, expression profile studies led to identify genes under selective pressures (i.e. insecticide resistance, immunity genes) and might generate new functional genomic tools (i.e. microarrays

or SNP platforms) while we wait for future genomic sequencing of *An. funestus*.

In malaria mosquitoes, population genetics have been revealed as an excellent tool for implementation of vector control programs. The study of gene flow among vector populations allows the analysis of mosquitoes' movement in natural populations, and therefore, how those populations are segregated. They can, for instance, assist to follow the expansion of genes of interest, such as those that confer insecticide resistance [98], or potentially help to introduce transgenic mosquitoes, refractory to parasite infection [99, 100]. On the other hand, these population genetic studies might be useful to investigate the genetic basis of speciation and/or local adaptation processes. They evidence a considerable importance in vector control

The biology of *An. funestus* has supported several "*a priories*" about its population structure in natural conditions. As mentioned previously in this chapter, this malaria mosquito mainly breeds in permanent or semi-permanent water bodies, such as rice fields, swamps or artificial lakes, always linked to human presence (see above). Moreover, this mosquito has exhibited a very slow recolonization power of those areas treated with insecticide. Both characteristics have led to assume the population subdivision of *An. funestus*. In this section, we will discuss the population structure of this malaria vector across Africa as revealed by two types of

The study of chromosomal rearrangements – cytogenetics – of *An. funestus* debuted early in the 1980's [6, 7], preceded by the success of this kind of studies in its congener *An. gambiae* [81, 82, 102]. It allowed differentiating members of the Funestus Group, avoiding the challenging interpretation of taxo-morphological rules. Green & Hunt [7] and Green [6] showed differences in the chromosomal polymorphism within the species of the group. As in *An. gambiae*, several chromosomal inversions were species-specific, while other inversions were polymorphic in

**4. Population genetic structure across Africa**

206 *Anopheles* Anopheles mosquitoes - New insights into malaria vectors mosquitoes - New insights into malaria vectors

markers: chromosomal inversions and molecular markers.

**4.1. Introduction**

measures [101].

**4.2. Cytogenetic studies**

For its predominant role as malaria vector and its wide geographical distribution across sub-Saharan Africa, *An. funestus* has been the most studied species of the group, although greatly exceeded by the studies in *An. gambiae* [82, 104, 105]. Seventeen chromosomal inversions have been recognized, with specific distribution through the African continent [6]; [52]; [84, 106-108]; (D. Ayala pers. comm.). Among them, four inversions are found all across the continent (2Ra, 3Ra, 3Rb, 3La), while others have a regional distribution (i.e. 2Rt in West Africa or 2Rh in South and Central Africa), or a very localized distribution (2Rd in the southern forested areas of Cameroon). These distributional patterns could be due to environmental selection, demographic effects or historical events [109].

Chromosomal inversions have been widely implicated in the process of speciation and local adaptation in a wide range of animals and plants [110, 111]. In recent years, studies on the chromosome composition of the populations of *An. funestus* were conducted in several African countries. These results showed a great complexity with different trends. In Burkina Faso, a deficit of heterozygotes and linkage disequilibrium among some rearrangements, led Costantini et al. [42] to identify two chromosomal forms: Kiribina and Folonzo, with a certain parallelism with the chromosomal forms of *An. gambiae* from Mali [104, 112]. These

two forms are also differentiated at the ecological level. While Kiribina appears better adapted to arid conditions, Folonzo inhabits more humid habitats [84, 113]. The presence of these two chromosomal forms was not observed in other countries such as Angola, Madagascar or Kenya [108, 114] (LeGoff, pers. comm.). Nevertheless, deficits of heterozy‐ gotes were also detected, particularly in inversions of the 3R and 3L arm, in some areas of Cameroon and Senegal [52, 83, 86, 115]. These studies did not show a clear division between the "chromosomal forms from Burkina Faso", rather a non-random distribution of chromo‐ somal inversions and their frequencies through different habitats and environments. This fact suggests that most inversions frequencies in *An. funestus* do not follow a neutral pattern. Ayala et al. [86] observed a sharp contrast between population structure measured at neutral microsatellite markers and at chromosomal inversions. Microsatellite data detected only a weak signal of population structure due to distance among geographical zones in Came‐ roon, as previously described by Cohuet et al. [83]. By contrast, strong differentiation among habitats was revealed by chromosomal inversions, strongly suggesting a role of environmen‐ tal selection in shaping their distribution. Moreover, in the same study, there was no apparent difference between microsatellite loci (*FST* estimates) lying within and outside polymorphic chromosomal inversions [86].

#### **4.3. Molecular markers**

The first assays to characterize wild populations of this mosquito were based on mitochondrial (Internal Transcribed Spacer 2, ITS2) and ribosomal DNA (cytochrome b gene, cyt-b) [116]. The results did not show any differentiation between chromosomal forms previously descri‐ bed by Costantini [42], rather one panmictic population. At the beginning of this century, new microsatellite markers were developed, which allowed more precise studies [89-92]. At the country scale, the results have evidenced a general trend to only one population, with a slight but significant isolation by distance. In Kenya, Braginets et al. [94] did not find any population genetic structure throughout the country, however, an important sub-division due to Rift Valley was found. A similar pattern was already observed in *An. gambiae*[117]. In Madagascar, Ayala et al. [95] did not find a population structure at the island level, rather a correlation between genetic and geographic distance across vector populations. In Senegal, Cohuet et al. [40] also showed genetic differentiation due to distance, without a clear relationship between "Burkina Faso chromosomal forms" and genetic data.

Similar results were obtained in Cameroon, where for the first time, a latitudinal cline across different environments was analyzed [83, 86]. As in previous studies, genetic differentiation among populations might be explained by isolation by distance. On the other hand, in Burkina Faso, Michel et al. [85] showed a genetic divergence between chromosomal forms on the basis of five microsatellite markers and sequence of a mitochondrial gene (ND-5). These results validated in some extend those precluded by Costantini et al. [42] and Guelbeogo et al. [84]. Unfortunately, they still remain restricted to Burkina Faso, similarly to chromosomal forms of *An. gambiae*in West Africa [118]. In recent years, several population genetic studies have been conducted at the sub-region and/or continental scale. Temu et al. [119], showed a similar pattern to the other studies at the country level for five countries in Eastern and Southern Africa: the genetic distance limited the gene flow among populations and promoted genetic differentiation among populations. A comprehensive study using samples across the continent provided important findings [93]. *Anopheles funestus* was subdivided into three large blocks: West Africa, East Africa and Central Africa [120, 121]. This subdivision was roughly similar than that observed in *An. gambiae* across Africa [122]. Despite these results and the unques‐ tionable accuracy of the analysis, the question about the incipient speciation of *An. funestus*, still remains to be elucidated.

The very rapid pace of development of genetic and molecular tools will allow characterizing *An. funestus* populations in a very detailed fashion. New molecular tools, such as SNP chip, RAD-tag or DNA microarrays, will certainly contribute to a better understanding of the biology of this mosquito. The expected *An. funestus* genome sequencing will undoubtedly boost new advances in order to elucidate a variety of biological processes involved in local adaptation, speciation, parasite transmission or the immunity system among others. It will also enable comparative studies with other anopheline species, particularly, *An. gambiae*.

## **5. Conclusion**

two forms are also differentiated at the ecological level. While Kiribina appears better adapted to arid conditions, Folonzo inhabits more humid habitats [84, 113]. The presence of these two chromosomal forms was not observed in other countries such as Angola, Madagascar or Kenya [108, 114] (LeGoff, pers. comm.). Nevertheless, deficits of heterozy‐ gotes were also detected, particularly in inversions of the 3R and 3L arm, in some areas of Cameroon and Senegal [52, 83, 86, 115]. These studies did not show a clear division between the "chromosomal forms from Burkina Faso", rather a non-random distribution of chromo‐ somal inversions and their frequencies through different habitats and environments. This fact suggests that most inversions frequencies in *An. funestus* do not follow a neutral pattern. Ayala et al. [86] observed a sharp contrast between population structure measured at neutral microsatellite markers and at chromosomal inversions. Microsatellite data detected only a weak signal of population structure due to distance among geographical zones in Came‐ roon, as previously described by Cohuet et al. [83]. By contrast, strong differentiation among habitats was revealed by chromosomal inversions, strongly suggesting a role of environmen‐ tal selection in shaping their distribution. Moreover, in the same study, there was no apparent difference between microsatellite loci (*FST* estimates) lying within and outside

The first assays to characterize wild populations of this mosquito were based on mitochondrial (Internal Transcribed Spacer 2, ITS2) and ribosomal DNA (cytochrome b gene, cyt-b) [116]. The results did not show any differentiation between chromosomal forms previously descri‐ bed by Costantini [42], rather one panmictic population. At the beginning of this century, new microsatellite markers were developed, which allowed more precise studies [89-92]. At the country scale, the results have evidenced a general trend to only one population, with a slight but significant isolation by distance. In Kenya, Braginets et al. [94] did not find any population genetic structure throughout the country, however, an important sub-division due to Rift Valley was found. A similar pattern was already observed in *An. gambiae*[117]. In Madagascar, Ayala et al. [95] did not find a population structure at the island level, rather a correlation between genetic and geographic distance across vector populations. In Senegal, Cohuet et al. [40] also showed genetic differentiation due to distance, without a clear relationship between

Similar results were obtained in Cameroon, where for the first time, a latitudinal cline across different environments was analyzed [83, 86]. As in previous studies, genetic differentiation among populations might be explained by isolation by distance. On the other hand, in Burkina Faso, Michel et al. [85] showed a genetic divergence between chromosomal forms on the basis of five microsatellite markers and sequence of a mitochondrial gene (ND-5). These results validated in some extend those precluded by Costantini et al. [42] and Guelbeogo et al. [84]. Unfortunately, they still remain restricted to Burkina Faso, similarly to chromosomal forms of *An. gambiae*in West Africa [118]. In recent years, several population genetic studies have been conducted at the sub-region and/or continental scale. Temu et al. [119], showed a similar pattern to the other studies at the country level for five countries in Eastern and Southern

polymorphic chromosomal inversions [86].

208 *Anopheles* Anopheles mosquitoes - New insights into malaria vectors mosquitoes - New insights into malaria vectors

"Burkina Faso chromosomal forms" and genetic data.

**4.3. Molecular markers**

During the last decade, we have seen how new molecular advances have elevated *An. gambiae* to the level of model species with regard to the number of data and tools available. *Anopheles funestus* is still far from this point. Undoubtedly, it is one of the major and more deadly malaria vectors worldwide. Its capacity to adapt to a wide range of ecological settings coupled with the appearance of insecticide resistance highlight the importance for studying this mosquito. However, the extreme difficulty to establish colonies in insectary conditions has hindered its study. Now, its upcoming genome sequencing and the availability of new molecular tools preclude a promising future for the study of this malaria mosquito.

The *An. funestus* geographical distribution mirrors *An. gambiae*'s across the whole African continent, with presumably similar environmental pressures. This mosquito exhibits a large number of chromosomal and genetic polymorphisms. Furthermore, it belongs to a group of morphologically undistinguishable species. This malaria mosquito is suspected to be at the heart of an ongoing speciation process, as its congener *An. gambiae*. Once the new techniques and vector control strategies have achieved their goals in *An. gambiae*, *An. funestus* will become the new target for succeeding malaria control programs. Moreover, the parallel study between both species will help to elucidate the ecological and genetics mechanisms involved in many biological processes from immunity system to local adaptation or speciation.

In this chapter, we revisited the *state-of-the-art* of this malaria mosquito as well as the other species of the Funestus Group. Detailed descriptions were provided on their biology, role in malaria transmission and insecticide resistance status. We examined the new genomic advances and how they can be useful for improving vector control strategies. To sum up, we strongly believe that a general knowledge about this mosquito is essential for the success of its control and the ultimate aim to reduce the malaria burden in Africa.

## **Author details**

Ibrahima Dia1 , Moussa Wamdaogo Guelbeogo2 and Diego Ayala3

\*Address all correspondence to: dia@pasteur.sn; guelbeogo.crnfp@fasonet.bf; diego.aya‐ la.g@gmail.com

1 Medical Entomology Unit, Institut Pasteur de Dakar, Dakar, Senegal

2 Centre National de Recherche et de Formation sur le Paludisme, Ouagadougou, Burkina Faso

3 UMR 224 MIVEGEC/BEES, Institut de Recherche pour le Développement, Montpellier, France

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**Author details**

210 *Anopheles* Anopheles mosquitoes - New insights into malaria vectors mosquitoes - New insights into malaria vectors

, Moussa Wamdaogo Guelbeogo2

1 Medical Entomology Unit, Institut Pasteur de Dakar, Dakar, Senegal

and Diego Ayala3

\*Address all correspondence to: dia@pasteur.sn; guelbeogo.crnfp@fasonet.bf; diego.aya‐

2 Centre National de Recherche et de Formation sur le Paludisme, Ouagadougou, Burkina

3 UMR 224 MIVEGEC/BEES, Institut de Recherche pour le Développement, Montpellier,

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