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Journal of Medical Entomology logoLink to Journal of Medical Entomology
. 2020 Jul 2;57(6):1782–1792. doi: 10.1093/jme/tjaa132

Genetic Diversity of Anopheles coustani (Diptera: Culicidae) in Malaria Transmission Foci in Southern and Central Africa

Ilinca I Ciubotariu 1, Christine M Jones 2, Tamaki Kobayashi 3, Thierry Bobanga 4,5, Mbanga Muleba 6, Julia C Pringle 2, Jennifer C Stevenson 2,7, Giovanna Carpi 1,#, Douglas E Norris 2,#,✉, for the Southern and Central Africa International Centers of Excellence for Malaria Research
Editor: David Severson
PMCID: PMC7899271  PMID: 32614047

Abstract

Despite ongoing malaria control efforts implemented throughout sub-Saharan Africa, malaria remains an enormous public health concern. Current interventions such as indoor residual spraying with insecticides and use of insecticide-treated bed nets are aimed at targeting the key malaria vectors that are primarily endophagic and endophilic. Anopheles coustani s.l., an understudied vector of malaria, is a species previously thought to exhibit mostly zoophilic behavior. Like many of these understudied species, An. coustani has greater anthropophilic tendencies than previously appreciated, is often both endophagic and exophagic, and carries Plasmodium falciparum sporozoites. The aim of this study was to explore genetic variation of An. coustani mosquitoes and the potential of this species to contribute to malaria parasite transmission in high transmission settings in Zambia and the Democratic Republic of the Congo (DRC). Morphologically identified An. coustani specimens that were trapped outdoors in these study sites were analyzed by PCR and sequencing for species identification and bloodmeal sources, and malaria parasite infection was determined by ELISA and qPCR. Fifty An. coustani s.s. specimens were confirmed by analysis of mitochondrial DNA cytochrome c oxidase subunit I (COI) and ribosomal internal transcribed spacer region 2 (ITS2). Maximum likelihood phylogenetic analysis of COI and ITS2 sequences revealed two distinct phylogenetic groups within this relatively small regional collection. Our findings indicate that both An. coustani groups have anthropophilic and exophagic habits and come into frequent contact with P. falciparum, suggesting that this potential alternative malaria vector might elude current vector control measures in northern Zambia and southern DRC.

Keywords: malaria, Anopheles coustani, mosquito, transmission

Malaria is transmitted to humans by the infectious bite of female mosquitoes of the Anopheles genus, and approximately 70 Anopheles species are potential vectors of malaria that transmit the disease to humans effectively worldwide (Sinka et al. 2012). Species are often characterized as primary or secondary vectors: primary vectors are those mosquitoes that are abundant, most commonly feed on humans, have measurable sporozoite rates, and contribute most to malaria transmission in an area, while secondary or alternative vectors can be uncommon, have low sporozoite rates, but may still play a role in malaria transmission (Institute of Medicine, Committee for the Study on Malaria Prevention and Control 1991).

Zambia, in sub-Saharan Africa, is a malaria-endemic country that has experienced high mortality and morbidity from this disease for decades (Mukonka et al. 2014, National Malaria Elimination Centre [NMEC] 2018). Significant strides here and across Africa have been made to reduce malaria transmission, largely due to the implementation of vector control interventions (Bhatt et al. 2015). These interventions include vector control through the distribution of long-lasting insecticide-treated nets (LLINs) and indoor residual spraying (IRS), in addition to treatment through intermittent preventive treatment in pregnancy (IPTp), and case management through the use of rapid diagnostic tests (RDTs) and artemisinin-combination therapy (ACT) (Chizema-Kawesha et al. 2010, Sutcliffe et al. 2012, NMEC 2018, President’s Malaria Initiative [PMI] 2019a). Despite successful reductions in morbidity and mortality, malaria remains endemic in Zambia with over 5 million reported cases in 2018 (NMEC 2018, World Health Organization [WHO] 2019a). While the scaling-up of malaria interventions such as widespread coverage by LLINs and IRS reduced transmission and parasitemia throughout many parts of Zambia, the disease continues to be a significant public health concern, especially in the northern region where Nchelenge District, Luapula Province, is recognized as a high transmission focus (Chanda et al. 2013, Mukonka et al. 2014, Nambozi et al. 2014, Hast et al. 2019). This region of Zambia reports over 350 confirmed cases per 1,000 population (Moss et al. 2012, PMI 2019a, WHO 2019a). This has raised doubts about whether the progress made across Zambia could be maintained and called for more enhanced and targeted interventions, especially in northern Zambia (Kamuliwo et al. 2013).

Like Zambia, the Democratic Republic of the Congo (DRC) is a malaria-endemic country in the central region of sub-Saharan Africa, located northwest of Zambia, and in which malaria is a leading cause of mortality and morbidity. It is the country with the second highest burden of malaria globally, accounting for approximately 12% of malaria cases and 11% of deaths worldwide (Messina et al. 2011, Stone et al. 2015, WHO 2019a). In 2018, the DRC had an estimated 26 million cases of malaria (WHO 2019a). Moreover, in this country, malaria accounts for 19% of deaths among children under the age of 5 (Ferrari et al. 2016, PMI 2019b). Despite actions taken to scale-up interventions in the DRC, such as the distribution of LLINs with over 50% household coverage on average, malaria transmission still remains high and progress appeared to have stalled according to the World Malaria Reports of 2017 and 2018 (WHO 2017, 2019a). This country was part of the launch of the WHO and RBM Partnership to End Malaria in 2018 through a high burden-to-high impact country-led approach in the hopes of continuing progress and reaching the 2030 goals of the Global Technical Strategy for Malaria (WHO 2015, 2019b).

Zambia and the DRC exhibit seasonal transmission that follows rainfall patterns, in which malaria peaks after the rains when mosquito populations increase (Masaninga et al. 2013). However, in Nchelenge District, malaria transmission is intense with limited seasonal fluctuations (Mharakurwa et al. 2012). In Nchelenge, year-round malaria prevalence by RDT ranges from 30% to more than 50% across all age groups and the primary vectors of malaria in both the dry and wet seasons have been found to be An. funestus s.s. Giles and Anopheles gambiae s.s. Giles (Das et al. 2016, Stevenson et al. 2016b, Jones et al. 2018, Hast et al. 2019). Malaria is also holoendemic in Haut-Katanga Province in the DRC with prevalence rates by microscopy in children in excess of 25%, but much less is known about malaria vectors and their phenology, other than that An. funestus s.s., An. gambiae s.s., and An. coluzzii Coetzee & Wilkerson are major vector species throughout the region, with the first two species exhibiting high biting rates (Bobanga et al. 2016, Nardini et al. 2017, Wat’senga et al. 2018, PMI 2019b). Vector control methods such as IRS and LLINs, which have been implemented throughout all of Zambia, and to a much less extent in the DRC, are aimed at targeting these indoor vectors preferentially. In addition to a potential suboptimal coverage of vector control in northern Zambia and southern DRC, malaria may remain intractable due to the presence of alternative vector species with behaviors that allow them to escape indoor control and that have largely remained unrecognized.

The focus on control and elimination methods for the well-recognized endophagic vector species highlights the fact that alternative vectors are rarely considered in existing malaria control programs, and are thought of as negligible because of their often zoophilic behavior and outdoor behaviors which result in lower human-vector contact (Fornadel et al. 2011). However, it has been observed that after primary vectors are reduced in a population, alternative vectors have the potential to sustain malaria transmission (Antonio-Nkondjio et al. 2006). Previous studies have indicated the presence of Plasmodium falciparum Welch parasites in these alternative vectors in Kenya, Ethiopia, Zambia, and other regions in Africa (Stevenson et al. 2012, Degefa et al. 2015, Lobo et al. 2015, Nepomichene et al. 2015, St. Laurent et al. 2016, Stevenson et al. 2016a). One of these alternative vectors, An. coustani s.l. Laveran, is a complex of at least two species (An. coustani A and B) that can only be differentiated by fixed differences on the X chromosome (Coetzee 1983). Morphologically similar species comprise the An. coustani species group and include An. tenebrosus Dönitz, An. symesi Edwards, An. ziemanni Grünberg, An. namibiensis Coetzee, and An. paludis Theobald (Gillies and De Meillon 1968, Gillies and Coetzee 1987). Like An. coustani, species within this group are reported to exhibit mostly zoophilic behavior (Gillies and De Meillon 1968). There is a significant paucity of entomological data on most species within this group and very little molecular data are available for comparison. However, recent studies from countries in southern Africa are bringing to light the potential contribution of An. coustani s.l. to malaria transmission. In Zambia, An. coustani s.l. displays an unexpectedly high degree of anthropophilic tendencies (Fornadel et al. 2011). In Kenya, this vector is both endophagic and exophagic and is thought to play a major role in outdoor malaria transmission (Mwangangi et al. 2013). In Madagascar, An. coustani mosquitoes have been shown to carry P. falciparum infections in both indoors and outdoors collections, and more recently, to act as a major local vector even though it is was previously a suspected alternative vector (Nepomichene et al. 2015, Goupeyou-Youmsi et al. 2019). These findings emphasize that mosquito species such as An. coustani may contribute more significantly to malaria transmission than previously recognized. These findings warrant further study of these alternative vectors, including foraging behaviors, ecology, genetics, and potential roles in the transmission of malaria.

As anophelines are commonly part of species complexes, integrating molecular and phylogenetic analysis to collections of An. coustani enable the confirmation of species identification and allow for further assessment of genetic diversity and relatedness within and between species complexes. In some species complexes, sibling species exhibit very different behaviors and therefore transmission potential. This study integrated an analysis of An. coustani phylogenetic structure and bloodmeal preference, and further assessed the potential role An. coustani may play in P. falciparum transmission in northern Zambia and southern DRC.

Methods

Mosquito Collection and Handling

As part of the International Centers of Excellence for Malaria Research (ICEMR) in Southern and Central Africa, mosquito specimens were collected solely outdoors from Nchelenge District, northern Zambia, using standard Centers for Disease Control and Prevention (CDC) light traps, and from two villages (Kilwa and Kashobwe) in Haut-Katanga Province, southern DRC, using CDC light traps and pyrethrum spray catches (PSC) (Fig. 1A and B). Households were geolocated and identified from satellite imagery and were randomly selected from those already enrolled for malaria epidemiological study for the ICEMR program (Stevenson et al. 2016b).

Fig. 1.

Fig. 1.

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Maps displaying the collection sites for the 50 Anopheles coustani samples from Zambia and the Democratic Republic of the Congo. This map was created using qGIS (v. 3.10.5) with Google Hybrid 2020 images. (A) The first map illustrates the general area of collection in context of southern and central Africa, and (B) the second map is a zoomed-in version displaying the study sites. The green-yellow circles represent the collection sites in Kilwa and Kashobwe from the DRC, and the purple circles indicate sites in Nchelenge from Zambia.

In Nchelenge District, Zambia, CDC light traps were placed overnight in the three following scenarios: outdoors where humans congregate, outdoors next to animal pens, and outdoors with a commercial human analogue bait (BG Lure, BioGents, Regensburg, Germany). Households were selected using a similar sampling frame to that of other studies under the ICEMR program (Pinchoff et al. 2015, Stevenson et al. 2016b). In brief, the ICEMR sampling frame facilitated identification of households that represented two ecological zones and had outdoor animal pens. Collections were performed over a 2-wk period in August 2016 at eight households (four inland along a stream and more than 3 km from Lake Mweru and four lakeside, close to Lake Mweru) for a total of 74 trap nights. Using a Latin Square design, trap scenarios were rotated through each household, such that each treatment occurred in each household at least once. Traps were activated at 6 p.m. and tied shut at 6 a.m. the following morning and retrieved.

In the DRC, the Kilwa Health Zone is located near Lake Mweru, across the lake from Nchelenge, Zambia, and the Kashobwe Health Zone is located near the Luapula River as it enters the south end of Lake Mweru, providing abundant vector breeding sites throughout the year (Fig. 1). In the Kilwa and Kashobwe Health Zones, 60 study households (Kilwa = 30 households, Kashobwe = 30 households) were randomly selected from village household census. Mosquito collections were performed in July 2016 by one of three collection scenarios: hanging CDC light traps indoors overnight next to a home occupant sleeping under a LLIN, hanging CDC light traps outdoors overnight by a window, or PSC early in the morning.

Mosquito Processing, Morphological Identification, and Isolation of DNA

Anophelines collected in Nchelenge were killed by freezing, while those from the DRC were left at room temperature before being packaged. Anophelines were identified by sex and morphology with the aid of a dissecting microscope and dichotomous key at all field sites (Gillies and Coetzee 1987). Although existing morphological keys do not identify males, suspected male An. coustani co-collected with females were retained for molecular confirmation. All mosquitoes were placed individually in 0.5 ml microcentrifuge tubes that contained silica gel desiccant and a cotton wool or paper plug. They were transported and stored at room temperature until processed in the laboratory at Johns Hopkins Bloomberg School of Public Health in Baltimore, Maryland.

In the laboratory, the abdomen of each collected anopheline was separated from the head and thorax using sterile forceps, and then stored in separate tubes at −20°C. Genomic DNA was extracted from the frozen mosquito abdomens with a salt extraction method as previously described (Post et al. 1993, Norris et al. 2001, Das et al. 2016).

Mosquito Species Identification by PCR

A fragment of the mitochondrial cytochrome c oxidase subunit I (COI) gene used for the Barcode of Life Database (BOLD), a molecular target that has been previously used for phylogeny construction of anophelines, was amplified and sequenced from 50 specimens (35 females and 15 males) that were morphologically/molecularly identified as An. coustani (Beebe 2018). The 698 bp BOLD fragment of the COI gene was amplified using LCO1490 and HCO2198 primers as previously published (Lobo et al. 2015). The 25 μl PCR mixture consisted of 2.5 μl of 10× buffer, 2.5 mM dNTP mixture, 30 pmol each of the forward and reverse primers, 2.0 U of Taq DNA polymerase (Invitrogen, Carlsbad, CA), and 1 μl of mosquito DNA template. The thermocycler (MultiGene OptiMax Thermal Cycler, Labnet International, Inc., Edison, NJ) conditions were identical to the ones described by Lobo et al. (2015).

A 750 bp fragment of the ribosomal DNA internal transcribed spacer region 2 (ITS2), a nuclear gene located between the 5.8S and 28S large subunit RNA genes that is often used for species verification and identification in anophelines, was also targeted for sequencing (Mohanty et al. 2009, Norris and Norris 2015). The ITS2 region was amplified from genomic DNA using ITS2A and ITS2B primers, and the 25 μl PCR mixture was identical to that for the BOLD fragment amplification PCR (Lobo et al. 2015). The thermocycler conditions were identical to the ones described by Lobo et al. (2015).

The PCR-amplified products of all specimens were visualized by electrophoresis on a 2% agarose gel. Resulting PCR products were then purified using the QIAquick PCR Purification Kit (Qiagen, Hilden, Germany) before Sanger sequencing by the Sequencing Facility at the Johns Hopkins School of Medicine.

Bloodmeal Analysis

A multiplexed PCR was performed on DNA extracted from mosquito abdomens to detect multiple bloodmeals by a protocol that differentiates between possible mammalian host blood in female mosquitoes as animal-specific products (human, cow, dog, pig, goat) amplified from the cytochrome b mitochondrial gene as described in detail elsewhere (Kent and Norris 2005).

Species Assignment and Phylogenetic Analyses

For both COI and ITS2 targets, forward and reverse sequences for each sample were trimmed to remove ends with low Phred quality and then high-quality trimmed forward and reverse sequences were aligned to generate a single consensus sequence for each individual sample using Geneious (Biomatters, Auckland, New Zealand) version 11.1.5 (https://www.geneious.com). Each COI and ITS2 consensus sequence was then queried against the NCBI database using BLASTn for molecular species identification/assessment (Altschul et al. 1997). Samples were confirmed as a particular species when the COI BLAST results indicated that there was a minimum nucleotide identity of greater than 95% to previously reported An. coustani sequences and a significant E-value < 1 × 10−5 as reported in previous papers with phylogenetic analyses (Lobo et al. 2015). Full consensus sequences for each sample were submitted to GenBank and assigned accession numbers (Supp Table 1 [online only]).

COI sequences that were confirmed as An. coustani after comparison with NCBI BLASTn were combined to generate a multiple sequence alignment using the MUSCLE algorithm and default parameters in the Geneious version 11.1.5 aligner. The multiple sequence alignment of the 50 An. coustani sequences was then trimmed to a final length of 552 bp for COI and 626 bp for ITS2. Furthermore, a multiple alignment was created with the confirmed An. coustani samples and other Anopheles species from the Hyrcanus and Coustani groups from the NCBI database for the COI target, and with the confirmed An. coustani samples and other An. coustaniITS2 sequences for the ITS2 target, as this latter gene region is not highly conserved between species.

Phylogenetic analysis was conducted using maximum likelihood (ML) inference as implemented in MEGA X (Kumar et al. 2018, Stecher et al. 2020). The evolutionary relationship was inferred by the Maximum Likelihood method and General Time Reversible model (Nei and Kumar 2000). One tree with the highest log likelihood for each target was included in this study. Branch support was included by bootstrap with 1,000 replications. The same steps were performed for analysis of the ITS2 sequence.

Detection of P. falciparum

Head and thoraces of female An. coustani mosquitoes were homogenized in a phosphate-buffered saline based solution in 1.5 ml microcentrifuge tubes for enzyme-linked immunosorbent assay (ELISA) and genomic DNA was extracted from half of the homogenate using the DNeasy Blood and Tissue Kit protocol (Qiagen) (Beier 2002, Gomes et al. 2017). As a quality control, genomic DNA concentration of each extract was quantified using High Sensitivity double-stranded DNA (dsDNA HS) assay on a Qubit 2.0 Fluorometer (Life Technologies, Grand Island, NY). All female An. coustani head and thoraces were subjected to the ELISA assay which uses monoclonal antibodies (CDC, Atlanta, GA) targeting the circumsporozoite protein (CSP) of P. falciparum sporozoites (Burkot et al. 1984). In addition, the genomic DNA extracts from the same individual mosquitoes were screened for P. falciparum DNA using a SYBR Green qPCR assay that targets an 85 bp fragment of the P. falciparum lactate dehydrogenase (Pfldh) gene using primers described elsewhere (Parr et al. 2016). The 25 μl PCR mixture consisted of 12.5 μl SYBR Green PCR Master Mix (Life Technologies, Warrington, United Kingdom), 1 μM each of the forward and reverse primers, and 4 μl of template DNA. Each reaction was performed in a StepOne Real-Time PCR System (Applied Biosystems, Foster City, CA) and the cycling conditions were the following: 50°C for 2 min, 95°C for 10 min, followed by 45 cycles of denaturation at 95°C for 15 s, and annealing at 60°C for 1 min. All samples were replicated in each reaction plate, no template controls (NTCs) were included and run alongside standard dilutions of gDNA of P. falciparum NF54 strain (1–105P. falciparum genome equivalent/μl). The qPCR is a more sensitive assay than ELISA, but the pitfall of this approach is that although it can specifically detect P. falciparum DNA, it cannot determine if it was from infectious sporozoites, early developmental stages, or even intact parasites.

Genetic Diversity

DNAsp (version 5.10.1) was used to assess diversity and polymorphism statistics on the sequences obtained from the mosquito samples collected in Zambia and the DRC (Librado and Rozas 2009). The number of polymorphic sites (S), the number of haplotypes, average nucleotide diversity (nucleotide differences per site based on pairwise comparisons among DNA sequences) (π), mean number of nucleotide differences (k), genetic diversity (θ), and haplotype diversity (Hd) (probability that two samples randomly sampled are unique) were all calculated for both COI and ITS2 sequences (Nei 1987).

Results

Morphological Identifications and Mosquito Processing

A total of 42 female (31 from northern Zambia and 11 from southern DRC) and 15 male (northern Zambia) morphological An. coustani were caught outdoors as part of this study, representing less than 5% of all mosquitoes captured from these study sites as part of a larger collection. Following extractions in the laboratory, both a 698 bp (BOLD) COI fragment and a 750 bp ITS2 fragment were amplified from the abdomen of all samples. Hundred percent of the males and 83.3% of the females successfully amplified with these PCRs; 7 females were excluded from the remainder of the study after multiple failed attempts of PCR amplification, and 50 total samples (15 males and 35 females) were included in the remaining assays and analyses.

Molecular Species Confirmation

Following Sanger sequencing and comparison to NCBI databases, the 50 mosquitoes that were morphologically identified as An. coustani were molecularly confirmed. All 50 An. coustaniCOI sequences matched to previously reported An. coustani sequences deposited in NCBI (as of January 18, 2020). For the COI fragment, 29 (58%) of these samples did not yield any close matches (closest match of ~80% was An. yatsushiroensis Miyazaki, a member of the Hyrcanus Group that is widely distributed in Oriental and Palaearctic areas) when their consensus sequences were blasted on the NCBI databases, which is presumably due to the relative lack of data for this locus and species in the database (Fang et al. 2017). There are over 20 An. coustaniCOI sequences publicly available (NCBI BLASTn, January 2020) and this gene is highly conserved, with little interspecies variation (Margoliash 1963). In contrast, there are currently only three previously submitted sequences in NCBI GenBank for the less conserved ITS2 gene for An. coustani (Coleman 2007).

Bloodmeal and P. falciparum Parasite Detection

Of the 35 female An. coustani samples, 4 (10.8%) (all from Zambia) had bloodmeals identified solely as goat, 3 (8.6%) (2 from Zambia and 1 from the DRC) had bloodmeals identified solely as human, and 3 (8.6%) (all from Zambia) had mixed bloodmeals identified as goat and human.

None of the 35 female anopheline samples were CSP ELISA positive. All 35 female An. coustani samples were also analyzed by qPCR for Pfldh. Six samples, all from Zambia (17.1%, 95% CI 6.5–33.6%), yielded positive signals for P. falciparum (parasite load was in the range of 1 parasite/μl – 5 parasite/μl, with Ct value of <37.88), of which two were identified as having a human bloodmeal.

Phylogenetic Analyses

The Maximum Likelihood analysis was performed for both the COI and ITS2 loci. The resulting phylogenetic tree that was constructed for the COI fragment (Fig. 2) revealed that all specimens that were morphologically identified as An. coustani clustered together. The COI clustering revealed two well-supported molecular groups (hereafter “An. coustani group 1” and “An. coustani group 2”) with 74 and 99% bootstrap support, respectively, among the An. coustani samples. Anopheles coustani group 1 encompasses 21 samples from the field and previously described An. coustani sequences from Zambia (KR014841 and KR014843), Mali (MK585958), and the Republic of Guinea-Bissau (KM097027). Within group 1, there is substructure that does not reflect geography, with samples from both Zambia and the DRC grouping closely with the aforementioned samples published from other parts of Africa. Anopheles coustani group 2, which has 100% bootstrap support, contains the remaining 29 newly reported sequences from this collection, and has two well-supported groups.

Fig. 2.

Fig. 2.

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COI maximum likelihood tree. This evolutionary analysis was performed using the Maximum Likelihood method and General Time Reversible model, with 1,000 bootstraps for branch support (Nei and Kumar 2000). The tree with the highest log likelihood (−1603.26) is shown. The tree is drawn to scale, with branch lengths shown in the number of substitutions per site. This analysis contains 61 samples, including 50 samples from this collection, and 11 sequences obtained from NCBI BLASTn with accession numbers. Evolutionary analyses were conducted in MEGA X (Kumar et al. 2018, Stecher et al. 2020).

Similarly, the phylogenetic tree that was created for the ITS2 fragment (Fig. 3) showed that all of the specimens morphologically identified as An. coustani clustered together. Moreover, the structuring revealed two groups (hereafter “An. coustani group 1” and “An. coustani group 2”) with 94 and 99% bootstrap support, respectively. Anopheles coustani group 1 from the ITS2 analysis contained the same 21 sequences from field mosquitoes as An. coustani group 1 from the COI clustering, and the same was observed for the samples in the group 2 clusters for both genetic targets, providing a topological concordance between mtDNA COI and nuclear ITS2 phylogenetic trees. As observed for the COI target, previously published sequences of An. coustani clustered into An. coustani group 1, even though they were from different geographical areas (KR014826 and KR014824 from Zambia, MK129245 from Madagascar, and KJ522815 from Kenya).

Fig. 3.

Fig. 3.

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Ribosomal ITS2 maximum likelihood tree. This evolutionary analysis was performed using the Maximum Likelihood method and General Time Reversible model, with 1,000 bootstraps for branch support (Nei and Kumar 2000). The tree with the highest log likelihood (−4723.20) is shown. The tree is drawn to scale, with branch lengths shown in the number of substitutions per site. This analysis contains 61 samples, including 50 samples from this collection, and 11 sequences obtained from NCBI BLASTn with accession numbers. Evolutionary analyses were conducted in MEGA X (Kumar et al. 2018, Stecher et al. 2020).

Genetic Diversity Analysis

Genetic diversity parameters were calculated for each gene and An. coustani group (Table 1). The mean haplotype diversity was higher for the An. coustani group 1 of both targets (0.943 ± 0.033 for COI and 0.762 ±0.050 for ITS2) when compared to the An. coustani group 2 (0.860 ± 0.038 for COI and 0.655 ± 0.041 for ITS2). Haplotype diversity is driven by an abundance of singleton haplotypes. Of the 31 haplotypes in the total set of 50 COI sequences, 78.5% (11/21) were singleton haplotypes (observed only once) in group 1 and 58.8% (10/17) in group 2. Nucleotide diversity was also higher for the An. coustani group 1 according to both genetic targets (0.011 for COI and 0.052 for ITS2) when compared to the An. coustani group 2 (0.007 for COI and 0.002 for ITS2) (Table 1). The lower number of nucleotide differences in the An. coustani group 2 is supported visually by both phylogenetic trees which generally show less clustering when compared to the group 1. The nucleotide diversity values are comparable to that of other within-species comparisons. A quick analysis of An. gambiaeCOI sequences from BLASTn revealed a value of 0.01, while that between sequences from species An. coluzzii and An. funestus was around 0.11 and between An. coluzzii and An. gambiae was much higher, around 0.40 (data not shown). Moreover, a previous study found a mean nucleotide diversity of 0.0065 across 114 individually sequenced An. gambiae samples for the nad5 gene, another mtDNA gene, and from a similar spatial range of the current study (Lukindu et al. 2018). Overall, haplotype and nucleotide diversity did not differ greatly between the groups 1 and 2 identified by the Maximum Likelihood phylogenies.

Table 1.

Genetic diversity by genetic target and phylogenetic An. coustani group

COI (552 bp) ITS2 (626 bp)
An. coustani group 1 An. coustani group 2 An. coustani group 1 An. coustani group 2
Number of sequences 21 29 21 29
S 23 22 58 11
h 14 17 7 7
Hd ± SD 0.943 ± 0.033 0.860 ± 0.038 0.762 ± 0.050 0.655 ± 0.041
π 0.011 0.007 0.052 0.002
k 6.200 3.774 28.859 1.333
θ 0.012 0.009 0.024 0.004
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DNAsp (v. 5.10.1) was used to calculate the number of polymorphic sites (S), the number of DNA haplotypes (h), the haplotype diversity (Hd), the nucleotide diversity (π), the average number of nucleotide differences (k), and theta per site from Eta (θ).

Discussion

All of the female An. coustani mosquitoes included in this study were caught outdoors, and some were found to be blooded either with human, goat, or a mixed bloodmeal of these two mammals. Although the sample size is small and there were no indoor collections conducted in northern Zambia during this study, these findings support reports that An. coustani is primarily exophagic (Nepomichene et al. 2015, Degefa et al. 2017). While none of the female An. coustani mosquitoes in this study had a positive signal from CSP ELISA assays, six samples from Zambia were qPCR-positive for P. falciparum. In this study, head and thoraces of the mosquitoes were used, which should restrict the presence of other parasite stages such as oocysts, but the presence of gametocytes present in a bloodmeal cannot be excluded (Carpi, unpublished). While this does not characterize the mosquitoes as infectious, finding human bloodmeals in 6 of 35 (3 solely human and 3 in combination with goat bloodmeal—5 from Zambia and 1 from the DRC), and detection of P. falciparum DNA in 6 (from Zambia) of 35 female An. coustani suggest that this mosquito feeds on human hosts frequently. Two of the samples that were shown to have fed on humans by PCR also tested qPCR-positive for P. falciparum confirming that An. coustani encounters P. falciparum parasite present in infected humans.

Phylogenetic analyses of both COI and ITS2 loci reveal that An. coustani from northern Zambia and southern DRC partition into two strongly supported groups, herein defined as An. coustani group 1 and An. coustani group 2 (Figs. 2 and 3). These groups have 100% bootstrap support in the COI analysis and 99% bootstrap support in the ITS2 analysis. Additionally, given that the nucleotide diversity found within groups 1 and 2 here is similar to that of other within-species for primary Anopheles malaria vectors within a species complex, it is plausible that these two An. coustani groups are two species. However, more samples must be collected and further phylogenetic and biological analyses should be conducted to elucidate this possibility. It is known that An. coustani comprises at least two species that are differentiated by fixed inversions on the X chromosome (Coetzee 1983). As the mosquitoes in this study were not processed for polytene chromosomes, it is unknown how our group 1 and 2 correspond to Coetzee species A and B. Until these species and/or molecular groups are better studied, the biological significance of this phylogenetic structure will remain unknown. Further studies are required to determine whether behaviors or vectorial capacity may vary between groups. Importantly, the tree topology for both genetic targets, which incorporates identical sample sets, strengthens the observed clustering of these specimens. Moreover, female mosquitoes in both An. coustani groups 1 and 2 (according to both targets) had taken human bloodmeals. It is of particular interest that two samples from northern Zambia that were found to have fed on human blood and found to be qPCR-positive for P. falciparum fell in the An. coustani group 2, illustrating the convergence of human host, parasite, and vector, although no causality can be proven here. Moreover, the presence of the identical haplotypes for both genetic targets at low frequency from northern Zambia and southern DRC (Figs. 2 and 3) may suggest that inter-breeding and migrations might be occurring between the An. coustani subpopulations. However, larger sample sizes and further genetic investigation are required to confirm this scenario.

Given that An. coustani has been linked to malaria transmission in other regions of Africa and that specimens within this limited sample set are associated both with biting humans and acquiring P. falciparum, further studies in Zambia and the DRC are warranted (St. Laurent et al. 2016, Stevenson et al. 2016a). More extensive investigations are necessary to truly understand the foraging behavior and malaria transmission potential of An. coustani and other understudied vectors. Such vectors are easily overlooked and may be behaviorally resistant to the indoor-based vector interventions deployed across sub-Saharan Africa (Afrane et al. 2016). Behavioral resistance may result from genetic adaptations or phenotypic plasticity in primary and alternative vectors (Govella et al. 2013, Killeen and Chitnis 2014). For instance, in the Solomon Islands, the scale-up of insecticide treated bed nets (ITNs) and indoor residual spraying (IRS) has successfully eliminated the primary vector An. koliensis Owen and has left An. punctulatus Donitz with a fragmented distribution, allowing an exophagic species (An. farauti Laveran) to emerge as the new sole primary vector as it changed its behavior to avoid insecticides (Bugoro et al. 2011, Russell et al. 2013). These findings highlight the urgency to expand entomological surveillance to include other anopheline species, especially in transmission settings where primary endophagic and endophilic vector populations have been mostly successfully controlled by elimination strategies, but transmission still remains. In these regions, residual transmission may be maintained by mosquito populations that preferentially bite outdoors and stay clear of indoor-based control measures. Recent studies such as that conducted in Madagascar, in which An. coustani was the main vector in one village and had a high exophagic rate, further emphasize the potential role of such vectors as major local vectors in these regions (Goupeyou-Youmsi et al. 2019). In sub-Saharan Africa, exophagic species such as An. coustani are ready to play a more significant role in transmission.

Supplementary Material

tjaa132_suppl_Supplementary_Table
Click here for additional data file. (13.8KB, xlsx)

Acknowledgments

The authors kindly acknowledge the Southern and Central Africa ICEMR field teams in Nchelenge District in Zambia and Kashobwe and Kilwa Health Zones in the Democratic Republic of the Congo (DRC) for their logistical assistance and participation in field collections. The authors are very grateful to the communities in Zambia and the DRC from which household collections were drawn and the National Malaria Control/Elimination Programs under the Ministries of Health in both DRC and Zambia. This work was supported in part by funding from the National Institutes of Health International Centers of Excellence for Malaria Research (U19AI089680), Bloomberg Philanthropies, the Johns Hopkins Malaria Research Institute, and NIH T32 Training Grant (T32AI0074717) support to CMJ. DEN and IIC conceived and developed the study. JCS, MM, GC, CMJ, and TK carried out the field collections and compiled collection data. Samples were processed by IIC, CMJ, and TK and phylogenetic trees were constructed and analyzed by IIC, GC, DEN, and CMJ. IIC, GC, and DEN drafted the manuscript. All authors read and reviewed the final manuscript. The authors affirm that there are no competing interests to be declared.

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