Abstract
Human malaria, which remains a major public health problem, is transmitted by a subset of Anopheles mosquitoes belonging to only three out of eight subgenera: Anopheles, Cellia and Nyssorhynchus. Unlike almost every other insect species, it was shown that males of some Anopheles species produce and transfer steroid hormones to females during copulation and that this transfer mediates reproductive changes. Steroids are consequently seen as a potential target for malaria vector control. Here, we analysed the evolution of sexually-transferred steroids and their effects on female reproductive traits across Anopheles by using a set of 16 mosquito species (5 Anopheles, 8 Cellia, and 3 Nyssorhynchus), including malaria vector and non-vector species. We show that male steroid production and transfer are specific to the Cellia subgenus and that there is no correlation between mating-induced effects in females and sexually-transferred steroids. In the light of our results, male steroid production, transfer and post-mating effects in females do not correlate with their ability to transmit human malaria, which overturns the suggestion from previous studies and suggests that manipulation of steroid-response pathways in the field should be considered with caution in order to benefit malaria vector control strategies.
Introduction
Anopheles mosquitoes are mostly known for their ability to transmit to mammals malaria caused by Plasmodium parasites. Among 472 species named in this genus, 41 have been classified as dominant vector species (DVS) of human malaria [1–3]. The Anopheles genus is further subdivided into 8 subgenera of which 3 (Anopheles, Cellia, and Nyssorhynchus) contains all known DVS of human malaria [3]. Despite a significant decrease of malaria incidence in the last 15 years due to vector control and improvement of chemoprevention, diagnostic testing and treatment, human malaria remains a world-wide burden with more than 200 million cases reported and an estimated 445 000 deaths in 2016 [4]. With the increase of insecticide resistance in malaria mosquitoes [4, 5], new vector control strategies to limit and possibly eliminate malaria transmission are being developed such as replacement of wild malaria-susceptible mosquito populations by resistant ones and/or decrease of vector populations by manipulating mosquito reproduction [6–12]. Unlike vertebrates, it was considered that insect adult males do not produce significant amounts of steroid hormones until it was shown that males of Anopheles gambiae, the main vector of human malaria in Africa, produce and transfer high quantities of 20-hydroxyecdysone (20E) to females during copulation [13]. While ovarian steroid synthesis occurs in mosquito females upon blood meal triggering egg development [13–18], sexual transfer of steroids by males likely represents a nuptial gift that affects female reproduction in the malaria vector. Consistent with this, sexually-transferred steroids were shown to induce refractoriness to further copulation and to stimulate oviposition in An. gambiae females [19]. A study further reported that mating-induced phenotypes in Anopheles females would only occur in species whose males produce and transfer steroids to females during mating and that would be specific to dominant human malaria vectors. Since steroids contribute to promote oogenesis and possibly favouring Plasmodium survival, the authors concluded that sexual transfer of steroids is likely to have shape anopheline ability to transmit malaria to humans [20]. Steroids became as such a promising target to manipulate mosquito female reproduction with the aim to reduce malaria vector populations specifically. Consequently, field use of 20E receptor agonists has been recently proposed to reduce malaria transmission [21]. In the present work, we investigated the evolutionary history of male steroid production using a large set of mosquito species belonging to the three Anopheles subgenera (Anopheles, Cellia, Nyssorhynchus) that contain all known DVS of human malaria. We also investigated the post-mating effects on female reproductive traits whether the cognate males produce or not steroids. The main conclusions of our work are that i) production of steroids in male mosquitoes is restricted to the Cellia subgenus, and ii) the effect of mating on female reproductive potential vary across species of the 3 Anopheles subgenera and this is not correlated to male steroid production.
Results
Male steroid production is specific to the Cellia subgenus
We first measured steroid titers in sexually mature virgin males from 19 different mosquito species. These mosquito species were selected within two Culicidae subfamilies, Anophelinae and Culicinae. Within the Anophelinae subfamily, 16 species distributed all over the world were chosen to cover 3 different subgenera (i.e. Cellia, Nyssorhynchus and Anopheles) of the Anopheles genus (Figure 1A). In the Culicinae subfamily, Aedes aegypti, Aedes albopictus and Culex pipiens that are vectors of arboviruses were also analysed. As shown on Figure 1B (right panel), male 20E production occurs only within the Cellia subgenus (Anophelinae) and is absent in the Culicinae. Interestingly, Anopheles quadriannulatus whose males produce similar levels of steroids compared to Anopheles stephensi is not a malaria vector unlike An. stephensi and other Cellia species investigated in the present study. Indeed, its refractoriness or low susceptibility to the human malaria parasite has been experimentally determined [22, 23], confirming epidemiological data [24]. Conversely, production of 20E was undetected in male mosquitoes from the two other Anopheles subgenera, Anopheles and Nyssorhynchus of which all tested members are registered as DVS and/or experimentally shown to be highly susceptible to Plasmodium falciparum [25, 26].
To uncover the evolutionary history of male steroid production in mosquitoes, we further consolidated the phylogenetic relationships of the species we analysed. To this effect, we used DNA sequences of partial regions of the coding sequence of mitochondrial genes (COI, COII, ND5 and CYTB) and nuclear genes (g6pd and white as well as ribosomal subunits 18S and 28S) from the 19 mosquito species plus Chagasia bathana (Anophelinae subfamily, Chagasia genera) as outgroup for Anopheles mosquitoes [27]. Phylogenetic analysis of the data set was performed by Bayesian inference (Figure 1B left panel) and by maximum likelihood (Supplementary Fig. 1). Phylogenetic relationships inferred from these analyses resulted in different topologies mainly at the subgenus level and with varying node support values. In both analyses, the members of each subgenus formed monophyletic groups and the branching orders within the subgenus Cellia was identical. Our results are in agreement with ones obtained by Sallum et al. [28] with the Cellia clade being the outgroup of the subgenera Anopheles and Nyssorhynchus in the Bayesian approach while in the maximum likelihood approach, Nyssorhynchus is the outgroup of Anopheles and Cellia, also in agreement with recently published phylogenies [1, 29, 30]. The Anopheles subgenus has also been found by others to be a basal lineage to Cellia and Nyssorhynchus [31]. Thus, relationships between the different subgenera Cellia, Nyssorhynchus and Anopheles, are still not entirely resolved. The difficulty in determining the relationships among these subgenera might be due to limits and bias in gene and taxon sampling, but also to different methods used [3, 27]. This could also be due to the fact that the radiation of Anopheles and Cellia species happened roughly at the same time [29, 31].
From the Bayesian phylogenetic analysis and using fossil data, we obtained species divergence estimates, which largely conform to recent phylogenies [29, 31, 32] (Supplementary Table 1). The age of the last common ancestor of Anopheles genus is about 84.1 Ma (112.7-55.8, 95% confidence interval) and the most ancestral node within the Cellia subgenus is dated to 69.2 Ma (93.0-45.6, 95% confidence interval) (Figure 1B, Supplementary Table 1). As males from all Cellia species tested so far have the ability to produce 20E, according to the parsimony law steroid production by mosquito males probably originated once in the early Cellia lineage, at about 84.1-69.2 Ma. (112.7-45.6, 95% confidence interval), i.e. during the late Cretaceous. Thus, steroid production by mosquito males is most probably a shared derived character from the last common ancestor of Cellia mosquitoes and represents as such a synapomorphy of this subgenus. An. stephensi males transfer steroid upon mating to females (Supplementary Fig. 2) as do An. gambiae males [13] and males of other Cellia species such as Anopheles arabiensis and Anopheles dirus [20]. This strongly suggests that transfer of steroids to females during mating is part of the “male 20E production” synapomorphy of Cellia mosquitoes. Our geographical mapping of nearly all contemporary mosquito species belonging to the Cellia subgenus (224 species) against the ones of Anopheles subgenus (184 species) (Figure 2) suggests that the common ancestor of the Cellia subgenus diverged after separation of South America and Africa in agreement with previous observations [3, 33]. This biogeographic calibration is consistent with divergence times obtained from our phylogenetic analysis placing the origin of steroid production by males of the Cellia subgenus around 84.1-69.2 Ma (112.7-45.6, 95% highest posterior density interval), i.e. after the separation of South America and Africa, which started at least 100 Ma ago with no land bridge for about 80 to 50 Ma [34, 35].
Mating-induced effects do not correlate with sexually-transferred steroids
To get a deeper understanding of the effect(s) of sexually-transferred steroids in Cellia female mosquitoes, we investigated the influence of mating on two female reproductive traits which are known to be regulated by ovarian 20E. On the one hand, steroid hormones trigger ovarian follicle detachment from the germarium in An. stephensi and Ae. aegypti [14, 36], and ovarian steroids produced upon blood feeding stimulate vitellogenesis and egg development, on the other hand [14, 16–18, 37]. We therefore analysed these two phenotypes in virgin and mated females from 12 Anopheles species (8 Cellia, 3 Anopheles and 1 Nyssorhynchus). As expected, mating induces the separation of the secondary ovarian follicle from the germarium in An. stephensi (Figure 3, see also confocal pictures Supplementary Fig. 3), a Cellia species whose males produce and transfer steroids during mating. Mating induces as well the separation of the secondary follicle in Anopheles minimus and Anopheles merus, although to a lesser extent. However, no secondary follicle detachment was observed after mating in An. dirus nor detachment of the third follicle for the Cellia species whose secondary follicle is already fully detached in virgin females (Anopheles farauti, An. gambiae, An. arabiensis and An. quadriannulatus). Among the species whose males do not produce steroids, mating also induces secondary follicle detachment in some species (Anopheles atroparvus and Anopheles freeborni), but not in others, even in species showing partial detachment of the secondary follicle in virgin females (Anopheles quadrimaculatus and Anopheles albimanus). Similarly, mating increases the number of developed eggs in some mosquito species while not in others and this is not correlated to male steroid production (Figure 4 and Table 1). As an example, mating has an effect on egg development in An. albimanus (Nyssorhynchus), whose males do not produce steroids, but not in An. gambiae nor in An. arabiensis (Cellia). Intriguingly, those results are in discrepancy with previous studies reporting that mating increases egg development in An. gambiae and An. arabiensis [20, 38] but not in An. albimanus [20, 39]. These differences across studies are not due to the origin of the blood used to feed females (animal or human) as we obtained the same results with An. albimanus and An. gambiae fed on mouse or human blood (Supplementary Fig. 4). It is likely that some variations can be observed between strains of a single Anopheles species as already described for some Aedes species [40, 41]. It cannot be excluded that male steroids transferred upon mating benefit reproduction of some Cellia species but only under certain environmental conditions as different ecological pressures such as nutrition resources can favour or not the maintenance and importance of nuptial gifts, as shown for Ae. aegypti [42–44].
Importantly, across the species investigated, mating induces detachment of the secondary follicle from the germarium in some species of which a low proportion of virgin females has the secondary follicle already detached as well as in some species of which a high proportion of virgin females has a detached follicle. Similarly, the occurrence of mating-induced effects on egg development is not linked to the ability of virgin females to develop a low or high number of eggs after blood feeding. Moreover, among species within the Cellia subgenus, the occurrence or absence of mating-induced phenotypes in females are not linked to the different quantities of steroids produced by the cognate males (Figure 1 and Table 1) and likely transferred to females during mating as determined for An. gambiae, An. stephensi, An. arabiensis and An. dirus (Supplementary Fig. 2, [13, 20]). Indeed, as depicted in Table 1, mating triggers both secondary follicle detachment and a rise in the number of developed eggs in An. stephensi, or only an increase in egg development in An. dirus, while this does not hold for An. gambiae and An. arabiensis, two species whose males produce and transfer higher amounts of steroids than An. stephensi and An. dirus. Overall, our analysis demonstrates that post-mating responses increasing fecundity in females indeed exist in Cellia species, some of which are likely mediated by male steroids. Importantly, they also occur in mosquito species whose males do not produce and transfer steroids such as in species belonging to the Anopheles and Nyssorhynchus subgenera (Table 1). Recently, male 20E has been shown to induce refractoriness to further mating in An. gambiae mosquitoes [19]. Similarly to results obtained for the two post-mating responses analysed in this study, female monoandry (insemination by a single male) occurs in species whose males produce steroids but also in ones whose males do not [45–48]. While few data are available for mosquitoes, it is well known that post-mating responses are quite conserved among insects. However, the rapid evolution of insect reproductive systems often results in species-specific genes and signalling pathways that ultimately trigger similar post-mating changes in different insect species [49]. For instance, Ae. aegypti males transfer to females Juvenile Hormone (JH) and Drosophila flies transfer male accessory gland peptides such as Sex Peptide upon copulation to trigger physiological and behavioural changes in mated females [50, 51]. Likewise, males from Cellia species transfer steroid hormones while males from Anopheles and Nyssorhynchus species are likely to transfer other and not yet identified molecule(s) to achieve similar effects. As JH is transferred to female during mating in Ae. aegypti mosquitoes but also in the Lepidoptera Heliothis virescens [52], sexual transfer of steroid hormones in other mosquito subgenera, genera or even insect orders not yet tested cannot even be excluded.
Conclusions
A deep understanding of the selective forces driving reproductive strategy diversity and their functional consequences are critical for designing strategies for management of insect pests. It was previously suggested that post-mating responses in Anopheles mosquitoes only exist in species whose males produce and transfer steroids to females and that would be specific to dominant human malaria vectors [20]. Analysing the evolution of steroid production by male mosquitoes from a larger set of mosquito species reveals that this physiological trait is a synapomorphy of the Cellia subgenus. Importantly, there is no correlation between the evolution of sexually-transferred 20E and malaria transmission to humans (Table 1). Consistent with this, while we show that male steroid production and subsequent transfer to females is likely to have evolved only once in the common ancestor of Cellia species, phylogenetic analyses on malaria mosquitoes support a convergent evolution with independent acquisitions of vectorial capacities in Anopheles mosquitoes [53–55]. Furthermore, we demonstrate that mating-induced phenotypes are variable among species and possibly even among strains or under different environmental conditions. These differences are independent of males ability to produce and transfer steroids to females and are not correlated to malaria vectorial capacity. Apart from follicle detachment, increase in egg development and induction of refractoriness to mating, numerous other functions of steroids have been described in adult insects [19, 20, 56–61]. Thus, sexually-transferred steroids could mediate different functions with more or less direct benefit for the reproduction of Cellia females, due to the rapid evolution of reproductive systems between species. The recent interest in sexually-transferred steroids in anopheline mosquitoes has led to propose targeting 20E pathways with 20E receptor agonists to manipulate mosquito female reproduction as a mean to reduce vector populations and malaria transmission [21, 62]. In the light of our results, manipulating sexual-transfer of steroids in mosquitoes or using 20E receptor agonists in the field to manipulate Anopheles mosquito population and malaria transmission should be considered with caution in order to benefit malaria vector control strategies and also because it risks to affect other untargeted non-malaria vector species.
It remains open as what were the evolutionary forces that have initially promoted the acquisition and radiation of this presumably costly male steroid production and sexual gift to females in Cellia mosquitoes around 84.1-69.2 Ma. At this time, two main paleogeological events that had impact on environmental conditions may have led to pressures driving the evolution of steroid production and transfer by males in Cellia species: i) the Gondwana break up at 100 Ma with separation of South America and Africa [63]; ii) the Cretaceous-Paleogene extinction event at 65.5 Ma [64, 65]. Because such geographical isolation and environmental stresses are believed to drive traits contributing to animal species survival, it is likely that transfer of steroids by males to females have favoured Cellia species populations at this critical time.
Methods
Mosquito species and rearing
Ecdysteroid production by sexually mature males was analysed in 19 mosquito species. Sixteen (16) species belong to the subfamily Anophelinae (An. arabiensis, An. dirus, An. farauti, An. gambiae form M, An. merus, An. minimus, An. quadriannulatus, An. stephensi, An. albimanus, An. albitarsis, An. aquasalis, An. atroparvus, An. freeborni, An. plumbeus, An. pseudopunctipennis and An. quadrimaculatus) and 3 species belong to the subfamily Culicinae (Ae. aegypti, Ae. albopictus and Cx pipiens). An. gambiae form M, now called An. coluzzii (N’Gousso strain), An. stephensi (Sda 500) were permanently reared at Institut Pasteur (France). An. albimanus STECLA (MRA-126), An. arabiensis DONGOLA (males generously prepared by T. Bukhari and MRA-856), An. quadrimaculatus ORLANDO (MRA-139), An. minimus MINIMUS1 (MRA-729), An. dirus WRAIR2 (MRA-700), An. farauti FAR1 (MRA-489), An. freeborni F1 (MRA-130), An. atroparvus EBRO (MRA-493), An. quadriannulatus SANGWE (MRA-1155) and An. merus OPHANSI (MRA-803), were obtained through BEI Resources, NIAID, NIH, and contributed by M.Q Benedict (MRA-126,-856,-139,-729,-700,-489,-130,-493), C. Aranda (MRA-493), W. Takken (MRA-1155) and R. Mahraj (MRA-803). Ae. aegypti (Liverpool strain) and Ae. albopictus (Ho Chi Min Ville, Vietnam) were a generous gift from A-B. Failloux (Institut Pasteur, France). Eggs from Cx. pipiens (anautogenous strain) were kindly provided by M. Weill (ISEM, Montpellier, France). Mosquito larvae were reared at 27°C in deionized water supplemented with minerals and fed on TetraMin Baby-E fish food from the day of hatching to the fourth larval instar supplemented with pieces of cat food. Male and female adults were maintained at 27°C, under 68% relative humidity and a 12/12h light/dark cycle, and provided free access to a 10% wt/vol sucrose solution for the first 5 days post-emergence (PE). Female mosquitoes (first gonotrophic cycle) were allowed to feed for 30 min on the blood of an anesthetized mouse or rabbit depending on mosquito species preference (see Supplementary Table 2). An. plumbeus L4 larvae and pupae were harvested from a natural pond in Switzerland and reared up to the adult stage at the Institut Pasteur. Mature males of An. albitarsis and An. aquasalis were kindly prepared and provided by D. Valle and L. Moreira (Fiocruz, Brazil) and An. pseudopunctipennis by F. Lardeux (IRD, Bolivia).
Mosquito sampling, ecdysteroid extraction and quantification
To measure ecysteroid titers in virgin adult males and females, males and females were separated on the day of adult emergence, transferred individually 5 days later in methanol and stored at −20°C until ecdysteroid extraction. For the transfer experiments, An. stephensi males and females were separated on the day of adult emergence for 5 days to allow male sexual maturation. Mating experiments and sampling were performed as described in [13]. Total ecdysteroids from individual whole mosquitoes were extracted with methanol and re-dissolved in enzyme immuno assay (EIA) buffer. Empty tubes were treated similarly in parallel to be used as a negative control (referred as extraction blank). Ecdysteroids were quantified by EIA, with 20-hydroxyecdysone-2-succinate coupled to peroxydase as a tracer (dilution 1:100,000) and the L2 antiserum (gift from M. De Reggi (Marseille, France); dilution 1:100,000). Calibration curves were generated with ecdysone (E; 3,6 - 500 pg/tube) diluted in EIA buffer, and titers were expressed as E equivalents. Under these conditions, detection limit is 2 pg E equivalents. All measurements were performed in duplicate and the results are expressed as mean values ±SEM of several (n=20 at least) independent samples and have been repeated on two independent cohorts of mosquitoes. Samples at or above the highest value of the calibration curve were diluted and quantify again. The intra- and inter-assay variation coefficients were 3,9% and 5,6%, respectively. For steroid titers in whole males from different species, data were subjected to statistical analysis using Kruskall-Wallis test for nonparametric data followed by Dunn’s post-test (control group: extraction blank). The indicated p values are those obtained with Dunn’s test. For the transfer experiment in An. stephensi, results were subjected to statistical analysis using Mann-Whitney test.
Taxon sampling and DNA sequencing for phylogenetic analysis
We chose 20 Culicidae species for phylogenetic and comparative analysis. We selected 16 species of the genus Anopheles (Anophelinae subfamily) of which An. arabiensis, An. dirus, An. farauti, An. gambiae, An. merus, An. minimus, An. stephensi, An. quadriannulatus, An. atroparvus, An. freeborni, An. plumbeus, An. pseudopunctipennis, An. quadrimaculatus, An. albimanus, An. albitarsis, and An. aquasalis. As outgroups, we chose Chagasia bathana (Anophelinae subfamily, Chagasia genus), and 3 mosquito species belonging to the subfamily Culicinae with Ae. aegypti, Ae. albopictus (Aedes genus) and Cx. pipiens (Culex genus). Sequence data were generated for An. aquasalis, An. atroparvus, An. merus and An. plumbeus. Genomic DNA was obtained from single individuals using the DNeasy Blood and tissue kit (QIAGEN). A common set of molecular markers were chosen based on the availability of sequence data for Ch. bathana. Partial genomic regions of four nuclear genes (g6pd, white, 18S and 28S) and four mitochondrial genes (COI, COII, ND5 and CYTB) were amplified by PCR with gene-specific or degenerate primers (sequences in Supplementary Table 3). For PCR amplifications, we used 0.4 μM oligonucleotides, 1 U GoTaq® DNA Polymerase (Promega) per 35 μl reaction volume, 2 mM MgCl2, and 200 μM dNTP and reactions were carried out using standard thermocycle conditions. PCR products were purified and Sanger-sequenced with gene-specific primers or with T7, SP6 universal primers at Cogenics (www.cogenics.com, Beckman Coulter, GenBank Accession Numbers in Supplementary Table 4). Sequence data of the remaining species were obtained from GenBank and VectorBase (Supplementary Table 4). Sequences were examined and aligned with Geneious 6.1.3 (Biomatters). Our dataset was not complete, we did not find or generate sequence data for 10 / 160 (8 genes X 20 species) gene specific sequences and among the rest 15 / 150 of sequences were only partially covering the locus. The missing sequence data represent 5.6% of the total dataset and was annotated as a “?” (missing value) in the alignments for phylogenetic analysis. Introns were removed from g6pd and white sequences and the extremities of all protein coding sequences were trimmed to be in codon frame. Alignments for protein coding genes were re-aligned with the Geneious translation alignment program. In addition to gene specific alignments a concatenated dataset was generated in the following order: COI-COII-ND5-CYTB-18S-28S-g6pd-white. The number of informative sites was calculated using MEGA4 [66].
Phylogenetic analysis
We used DNA sequences of partial regions of the coding sequence of mitochondrial genes (COI, COII, ND5 and CYTB) and nuclear ones (18S, 28S, g6pd and white) from the 19 mosquito species plus Chagasia bathana. The combined dataset had 4398 positions, including 2602 constant positions, 1776 variable positions and 1356 parsimony informative positions. Phylogenetic analysis of the concatenated, five-partition data set was performed by maximum likelihood in PhyML [67] and by Bayesian inference in BEAST [68]. For all analyses, partition specific models of nucleotide substitution were selected using the Akaike Information Criterion as calculated in jModelTest 2.1.3 [69, 70]. Maximum likelihood inference was done on the concatenated dataset in PhyML [67] using a GTR+I+G model of nucleotide substitution. Node support was determined by performing 100 bootstrap replicates. Bayesian phylogenetic analysis was performed with BEAST v1.7.5 [68] on the concatenated data set using five partitions with unlinked models of nucleotide substitution (Supplementary Table 5). The partitions corresponded to a single partition for mitochondrial sequences (COI, COII, ND5 and CYTB) and four more partitions for the genes 18S, 28S, g6pd, and white. Mitochondrial genes were combined into one partition because they are closely linked in the mitochondrial genome and largely evolve as a single unit with little to no recombination [71]. We used a common strict clock model and a yule birth process as tree prior. While the Bayesian approach places the species of the subgenus Nyssorhynchus as sister group of the Anopheles subgenus species, they formed the outgroup of the Anopheles and Cellia lineages in the maximum likelihood approach. Overall, the Bayesian phylogeny revealed high posterior probabilities for each node (>0.9) while the maximum likelihood analysis lacked strong statistical support at the nodes that separate the lineages of the three Anopheles subgenera. For species divergence time estimates, we therefore used the Bayesian phylogenetic analysis. We assigned calibration fossil ages to set priors for most recent common ancestors (MRCA). We used Cx. winchesteri (33,9-55,8 Ma) [72] (see also http://mosquito-taxonomic-inventory.info/category/fossil-culicidae/fossil-culicidae) to approximate the age of the most recent node shared by Cx. pipiens – and Ae. aegypti/ Ae. albopictus; and An. dominicanus (33,9-40,4 Ma) [73, 74] to estimate the age of the most recent node shared by Anopheles species. MRCA priors that incorporate fossil calibration dates were assumed to follow an exponential distribution, in the two above mentioned cases with an offset of 33.9 Ma and a mean according to the mean ages of the fossils. Based on a recently published phylogeny of Kamali et al. [53], the root age of the Culicidae was set to 147 Ma and the prior was assumed to be normally distributed with a standard deviation of 20 Ma. The Markov-Chain Monte-Carlo (MCMC) run was performed with a chain length of 108 generations and was recorded every 1000 generations. Estimates were computed with Tracer version 1.5 (http://tree.bio.ed.ac.uk/software/tracer/) and MCMC output analysis was done using TreeAnnotator [68]. The first 2000 sampled trees were discarded as the burn-in. The phylogeny was visualized and annotated with Figtree version 1.4 (http://tree.bio.ed.ac.uk/software/figtree/).
Cellia and Anopheles species distribution mapping
Total numbers of mosquito species belonging to the Cellia or Anopheles subgenera per country were taken from the Walter Reed Biosystematics Unit (WRBU, http://www.wrbu.org/). Data were further represented on maps created with R [75] using the “sp” package [76].
Analysis of the secondary follicle detachment in virgin and mated non-blood fed females
Males and females were separated upon emergence to keep females virgin. A portion of females were put in a cage with males to obtain mated females. After 7 days, female ovaries were dissected, and the spermatheca of mated females checked for presence of spermatozoa. Ovaries from each female were then mounted on a slide in mounting media and observation of the secondary follicle detachment was performed using a transmitted light microscope. 30 to 60 females were analysed per species and per condition. Data were subjected to Chi-square test. For An. stephensi, ovaries from either virgin or mated females were fixed with 4% paraformaldehyde, washed 3 times with PBS-Tween 20 0.05% (PBS-T) and stained with DAPI. After 4 washes with PBS-T, ovaries were mounted on a slide in mounting media. Pictures were taken using a SP5 Leica confocal microscope.
Analysis of egg development in virgin and mated blood fed females
Virgin and mated females were prepared as described above. A portion of females were put in a cage with males to obtain mated females. On 7 days PE, females were allowed to blood feed on anesthetized mouse or rabbit (Supplementary Table 2). An. gambiae and An. albimanus females were also fed with fresh human blood (ICAReB Platform, Center for Translational Research, Institut Pasteur, Paris, France) for comparison with mouse-fed mosquitoes. Ovaries were dissected 48 hours after blood feeding, and the total number of eggs in virgin and mated females was counted. Mated status was verified by observing spermatozoa in the spermatheca and only females with a filled spermatheca were taken into account. 30 to 60 females were dissected per species and per condition. Data were subjected to Mann-Whitney non-parametric test.
Fundings
Support to E.P. was from an ANR-07-MIME-O25-01 award to C.B. and by the UK Medical Research Council to E.P. (MC_UU_12014/8), to C.B. from award no. ANR-10-LABX-62-IBEID. M.L. was hosted by V. Courtier-Orgogozo, financed by a FRM postdoctoral fellowship SPF20121226328 and by the CNRS.
Data availability
The sequence data reported in this paper are tabulated in the Supplementary information and archived at Genbank.
Contributions
E.P. and C.B. conceptualised and supervised the project as well as the experimental design; E.P., N.P., M.L., F.C., F.S., C.D-V. and E.B. performed the experiments; E.P., N.P., M.L., E.B. and C.B. contributed to data interpretation; E.P., N.P., M.L., F.S. and E.B. contribute to figure preparation; E.P. and C.B. wrote the manuscript; M.L., F.S., E.B. commented on the final manuscript.
Ethical compliance
This study complied with all relevant ethical regulations. Project (n° 2013-0132) approved by the Ministère de l’Enseignement Supérieur et de la Recherche – Direction Générale pour la Recherche et l’Innovation – Secrétariat « Autorisation de projet » - 1, rue Descartes, 75231 PARIS cedex 5.
Acknowledgments
We are grateful to L. Lambrechts, S. Blandin and J-P. Parvy for comments on the manuscript. We acknowledge D. Streicker, E. Hutchinson and M. Palmarini for discussions. We acknowledge M-T. Lecoq, S. Touron and C. Thouvenot for mosquito rearing, A-B. Failloux for providing Ae. aegypti and Ae. albopictus eggs, M. Weill (ISEM, Montpellier, France) for providing Cx. pipiens eggs, T. Bukhari for providing An. arabiensis samples, D. Valle and L. Moreira (FIOCRUZ, Brazil) for providing An. aquasalis and An. albitarsis samples, as well as F. Lardeux (IRD, Centre de Bolivie, La Paz, Bolivia) for providing An. pseudopunctipennis samples. We thank E. Perret (Dynamic Imaging Platform, Institut Pasteur) for her assistance with confocal microscopy. The following reagents were obtained through BEI resources, NIAID, NIH: eggs of An. albimanus STECLA (MRA-126), An. arabiensis DONGOLA (MRA-856), An. quadrimaculatus ORLANDO (MRA-139), An. quadriannulatus SANGWE (MRA-1155), An. merus OPHANSI (MRA-803), An. minimus MINIMUS1 (MRA-729), An. dirus WRAIR2 (MRA-700), An. farauti s.s. FAR1 (MRA-489), An. atroparvus EBRO (MRA-493) and An. freeborni F1 (MRA-130).
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