Background

Africa has witnessed significant progress in the fight against malaria from 2000, thanks to extensive vector control using insecticide-treated nets and indoor spraying, alongside improved diagnosis and treatment [1]. However, this progress began to stagnate around 2015, and currently, malaria causes ≤ 249 million cases and 608,000 deaths annually, predominantly in sub-Saharan Africa [2].

Other than the weak health systems and socio-economic conditions [3, 4], the persistent malaria burden in the region is exacerbated by several biological challenges, notably insecticide resistance [5, 6], anti-malarial drug resistance [7,8,9], failing diagnostics [10,11,12,13] and an invasive vector species, Anopheles stephensi [14,15,16]. The problem is compounded by human behaviour and lifestyles leading to inadequate protection during peak transmission periods, and insufficient community and stakeholder engagement in malaria prevention efforts [17,18,19,20]. Evidence also suggests that the traditional methods used to disrupt malaria transmission, notably insecticide treated bed nets (ITNs) and indoor residual spraying (IRS) are insufficient against certain vector species with atypical behaviours, such as those that do not bite or rest primarily indoors [21,22,23].

In east and southern Africa, the major malaria vectors are Anopheles gambiae sensu stricto, Anopheles arabiensis and Anopheles funestus mosquitoes [24, 25]. In most settings, An. gambiae s.s. and An. funestus mosquitoes have been historically dominating the malaria transmission [26,27,28,29]. However, recently, in some localities, such as in parts of western Kenya and south-eastern Tanzania, the wide coverage of ITNs likely in concert with environmental changes, appears to have significantly suppressed An. gambiae s.s. leaving An. arabiensis and An. funestus s.s. as the main drivers of transmission [30,31,32,33]. In Tanzania, An. funestus is now the dominant malaria vector across the country [34]. More detailed studies have revealed that even when outnumbered by An. arabiensis, An. funestus s.s. mediates over 90% of the ongoing malaria transmission in south-eastern Tanzania [31]. With An. funestus being highly anthropophilic, and in some settings having stronger resistance to public health insecticides compared with the other major malaria vectors [35], this vector species poses a significant challenge to the existing vector control interventions. It is noteworthy, that most studies have so far focused on only one member of the An. funestus group, i.e. An. funestus s.s. despite this species being one member of a large species complex [36].

The An. funestus group is thought to comprise 13 sibling species: An. funestus s.s.[25], An. funestus-like, An. vaneedeni [37], An. parensis [38,39,40], An. rivulorum[41], An. rivulorum-like [42], An. leesoni, An. aruni, An. confusus, An. brucei, An. fuscivenosus and An. longipalpis types A and C [43]. Of these, An. funestus s.s. is the most competent malaria vector in the group, though other sibling species, such as An. rivulorum, An. leesoni and An. parensis have also been reported to carry Plasmodium falciparum to lesser extent [37,38,39, 41, 43]. Despite these important observations, the species composition, distribution, and role in malaria transmission of the An. funestus group remains understudied, and several members of this group are likely to be misidentified. For instance, Ogola et al. [44] reported an unidentified sibling species within the An. funestus group in Kenya, and existing polymerase chain reaction (PCR) assays commonly return unamplified samples, including those morphologically confirmed as belonging to the group [45, 46]. More importantly, our understanding of the ecological dynamics and potential roles of these in perpetuating persistent malaria transmission remains limited.

To bridge these knowledge gaps, our research team initiated and implemented a cross-country survey of malaria vectors aimed at determining the species composition, spatial distribution, and the relative contribution of different Anopheles spp. to malaria transmission in mainland Tanzania. This paper presents the results from the initial phase of these surveys, covering 14 districts in 12 regions across Tanzania.

Methods

Study area

Mosquitoes were collected from 14 districts in 12 administrative regions across Tanzania mainland (Fig. 1). Tanzania has a broadly tropical climate, with four primary climatic zones: the hot and humid coastal plain (i.e. Pwani, Tanga, Lindi and Mtwara), the semi-arid central plateau (i.e. Dodoma, Kigoma, Katavi and Rukwa), the high rainfall lake regions (i.e. Kagera and Mwanza) and the cooler highlands (i.e. Morogoro and Ruvuma). On the Tanzanian coast and offshore islands, temperatures typically fluctuate between 27 °C and 29 °C. In the central, northern, and western regions, temperatures vary between 20 °C and 30 °C. The extended rainy season spans from March to May, while the shorter rainy season extends from October to early December, with the dry season lasting from June to September. Overall, annual rainfall ranges from 550 mm in the central areas to 3690 mm in certain parts of the southwestern highlands [47]. In most of these districts, the majority of the rural households are subsistence farmers [48, 49]. Malaria prevalence in children under the age of 5 years differs significantly in the study area, with the highest in the north-western (i.e. Kagera and Kigoma) and south-eastern regions (i.e. Mtwara and Lindi) to less than 1% in the central region (i.e. Dodoma)[50].

Fig. 1
figure 1

Map of Tanzania showing the regions where Anopheles funestus mosquitoes were collected

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The data collection sites are shown in Fig. 1. Specific districts were: Misenyi in Kagera, Kakonko and Kibondo in Kigoma, Chamwino in Dodoma, Ulanga and Kilombero in Morogoro, Tunduru in Ruvuma, Bagamoyo in Pwani, Nkasi in Rukwa, Tanganyika in Katavi, Misungwi in Mwanza, Mtama in Lindi, Mahurunga in Mtwara and Muheza in Tanga. These collection sites represent diverse geographical regions, including the hot and humid coastal plain (i.e. Bagamoyo, Muheza, Mtama and Mahurunga), the semi-arid central plateau (i.e. Chamwino, Kakonko, Kibondo, Tanganyika and Nkasi), the high rainfall lake regions (i.e. Misenyi and Misungwi) and the cooler highlands (i.e. Kilombero, Ulanga and Tunduru).

Mosquito collection and processing

Mosquito collections, conducted as part of a larger project on the population genetics of An. funestus sensu lato, were sporadic and completed between December 2018 and December 2022. These collections spanned both dry and wet seasons. Whereas multiple mosquito species were collected, only An. funestus s.l. are used in this analysis. In each of the districts, at least two houses were selected upon consent from the household heads and used for the collection of adult mosquitoes. Centres for Disease Control and Prevention (CDC) light traps [51] and Prokopack aspirators [52] were used to sample indoor host-seeking and resting mosquitoes, respectively. The overall sampling approach had been specifically designed to collect An. funestus s.l for population genetics studies, and was specifically focused on indoor collections, with no outdoor trapping in this phase of the study. Thus, mosquitoes were morphologically sorted to the species level and females of An. funestus s.l individually packed in an Eppendorf tube with 80% ethanol. In addition, in some locations, such as Dodoma, Tanga and Morogoro regions, where sampling of adult An. funestus s.l was insufficient, larval collections were conducted using standard larval dippers [53]. The collected larvae were reared to adults as previously described [54], then also sorted by taxa as above.

Extraction of genomic DNA

Genomic DNA was extracted from the heads and thoraxes of collected mosquitoes using DNAzol method [55]. Bead Ruptor 96 well plate homogenizer (OMNI international, Kennesaw, GA, USA) was used for homogenization and the resultant DNA pellets were eluted in 50 µl of Tris–ethylenediaminetetraacetic acid (EDTA) buffer.

Identification of the sibling species in the An. funestus group and detection of Plasmodium spp. infections

A cocktail of species-specific primers for the identification of the sibling species in the An. funestus group was used, as previously described by Koekemoer et al. [56]; with a slight adaptation to include a primer for An. rivulorum-like (Table 1) in the cocktail [42]. A nested PCR assay was used for the detection of the Plasmodium spp., of which the first round of the PCR included universal forward and reverse primers for 18S rDNA Plasmodium spp. (Table 2) regardless of species; followed by a second round using the amplicon from the first round as DNA template. Species-specific primers for Plasmodium falciparum, Plasmodium ovale, Plasmodium vivax and Plasmodium malariae were used in the second round (Table 2).

Table 1 Primers for PCR detection of Anopheles funestus group sibling species
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Table 2 Primers for a nested PCR detection of Plasmodium infection and species
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Further analysis of the internal transcribed spacer 2 (ITS2) region in non-amplified An. funestus samples to investigate polymorphisms

A total of ten samples underwent cloning and sequencing, employing the following primers: ITS2A: 5′ TGT GAA CTG CAG GAC ACA T 3′ (forward) and ITS2B: 5′ TAT GCT TAA ATT CAG GGG GT 3′ (reverse). The PCR reaction mixture, conditions and procedures for the thermal cycling and electrophoresis were similar to those described earlier. The amplicons (approximately 840 base pairs) were excised from the gel and cleaned using Wizard® SV Gel and PCR Clean-Up System (Catalogue number: #A9281, Promega). The purified product was cloned using a plasmid vector pJET1.2/blunt (CloneJET PCR Cloning Kit, Catalogue number: #K1231, Thermo Scientific). The resulting recombinant plasmid DNA was isolated and purified (QIAprep Spin Miniprep Kit, Catalogue number #27106, Qiagen) and sent for sequencing. Sequencing of the recombinant plasmid DNA was carried out using the reverse PJET1.2 primer (5′-AAGAACATCGATTTTCCATGGCAG-3′). Plasmid primer regions trimming, sequence alignment and analysis were performed using SeaView software [57].

Data analysis

The data collected from the field included the number of traps used, the number of collection days, and the mosquitoes captured per trap, facilitating the calculation of trap nights (defined as the product of the number of traps and collection days). The annual entomological inoculation rate (EIR) was determined by multiplying human biting rates and Plasmodium sporozoite prevalence, then adjusted for 365 days [31]. A coefficient of 0.68 was used for conversion of the sampling efficiency of the CDC light trap relative to human landing catch (HLC) [31, 46]. Mosquitoes collected using Prokopack aspirators and from larval collections were excluded from the EIR calculation, as these methods do not accurately reflect host-seeking behaviour or the potential for infectivity. Basic Local Alignment Search Tool for nucleotides (BLASTn) analysis [58] was used to identify and characterize nucleotide sequences by finding homologous sequences in the National Center for Biotechnology Information nucleotide (NCBI nt) database. The top hits were retrieved and analysed to determine sequence similarity, alignment scores and query coverage. For the alignment and analysis of ITS2 sequences cloned from PCR-negative samples, SeaView software [57] was utilized.

Results

Species composition and distribution

A total of 1092 An. funestus s.l were analysed, of which 549 (50.3%) were An. funestus s.s., 124 were An. parensis (11.4%) and 12 were An. rivulorum (1.1%; Table 3). No An. vaneedeni or An. rivulorum-like were found during this study. While the other species, An. funestus s.s. and An. rivulorum were more widespread in the study sites, the An. parensis samples were found most abundantly in the central and northern regions (Table 3; Dodoma and Mwanza). There were 404 samples (37%) for which the DNA fragments did not amplify during the PCR (Table 3). Subsequent cloning and sequencing of the ITS2 region in these non-amplified samples revealed multiple polymorphisms within the reverse primer’s priming region specific to An. funestus s.s. (examples are shown in Additional file 1). The ITS2 sequences were found to be similar to those of An. funestus s.s., with the exception of polymorphisms within the priming site. BLASTn analysis revealed that the highest identity was 99.5%, considering sequences with 100% query coverage. The analysis confirmed that these sequences closely matched An. funestus s.s. sequences from the NCBI nt database.

Table 3 Composition and distribution of Anopheles funestus mosquitoes in mainland Tanzania
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Prevalence of Plasmodium sporozoite infections in the mosquitoes

Of 914 An. funestus s.l tested for Plasmodium spp. infection, 13 were found positive for P. falciparum (Table 4). The majority of the infections were in An. funestus s.s. (n = 11). In addition, there were two An. parensis mosquitoes infected with P. falciparum (n = 2). No other Plasmodium species were detected, nor were any sibling species of An. funestus (besides An. funestus s.s. and An. parensis) found to be infected with Plasmodium spp.

Table 4 Plasmodium spp. prevalence in Anopheles funestus sibling species collected from 14 districts across mainland Tanzania
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Transmission intensities mediated by An. funestus group

The annualized EIR estimates varied significantly across the regions. The highest EIR estimates were recorded in Kagera [108.3 infectious bites/person/year (ib/p/y)], Ruvuma (72.2 ib/p/y) and Morogoro regions (65.6 ib/p/y). Since no infected mosquitoes were collected in Dodoma, Rukwa, Lindi, Katavi and Mtwara, it was not possible to estimate EIRs from the An. funestus s.l collected in these regions (Table 5).

Table 5 Annual entomological inoculation rates owing to Anopheles funestus group by regions
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Discussion

Anopheles funestus mosquitoes are among the most widespread, and yet also among the least studied species of malaria vectors. However, in recent years, there has been an increasing awareness that populations of An. funestus s.s., known for their high degree of anthropophily and now marked by significant pesticide resistance [35, 46], are becoming predominant in many malaria transmission areas, particularly in East and Southern Africa [59, 60]. In areas, such as south-eastern Tanzania, this species now mediates 85–98% of new malaria infections, even in villages where it is outnumbered by other species, such as An. arabienesis [31, 46, 61]. Today, the species composition and distribution of the An. funestus group, particularly in Tanzania, are well described. However, despite field collections regularly capturing several other members of the complex in many locations, the ecology and vectorial importance of these potential secondary vectors are poorly understood. This current study was therefore aimed at expanding on the existing knowledge towards understanding the vectorial role of An. funestus species across Tanzania mainland.

We found four known and previously reported sibling species of the An. funestus group (i.e. An. funestus s.s., An. parensis, An. rivulorum and An. leesoni), with An. funestus s.s. dominating malaria transmission across all the 12 regions surveyed. Moreover, 37% of the collected mosquitoes were not amplified by the available species-specific PCR assay [56] designed for the An. funestus group, despite being morphologically identified as An. funestus s.l. While this is a significantly high failure rate of the recommended PCR assays, similar non-amplification problems have been reported in previous studies, albeit at lower rates, including in south-eastern Tanzania [35, 45, 46]. Nonetheless, upon cloning and sequencing, it was confirmed that the ITS2 sequences were similar to that of An. funestus s.s. with the exception of polymorphisms present within the priming site of the common and widely used species-specific reverse primer. This PCR mis-priming is hereby considered the main reason for the high rates of non-amplification observed in this study; and may also have affected the aforementioned past studies. It is noteworthy, that all technologies based on PCR amplification, including higher throughput species identification multilocus amplicon panel approaches [62], will at times face similar issues because of the highly polymorphic genomes of Anopheline vector species. This suggests the need to continue improving the methods for identifying members of such species groups and complexes.

The incrimination of An. parensis with transmission of P. falciparum in this study provides only the third such report in Tanzania in the past 15 years [39, 63]. The two previous reports [39, 63] utilized CDC light traps, pyrethrum spray catch and aspirators for indoor sampling of host-seeking An. parensis mosquitoes, with nested PCR and enzyme linked immunosorbent assay (ELISA) as methods of Plasmodium sporozoite detection. Furthermore, the first report which was based on four specimens reported a 25% sporozoite rate [39], whilst the one conducted within the similar geographical area surrounding Lake Victoria as our present study and based on hundreds of samples reported 1.1% rate [63]. Collectively, these repeated observations suggest that An. parensis may be playing a modest but considerable role as a secondary malaria vector in Tanzania and should be further investigated to optimize the control of malaria transmission.

In this study, the two Plasmodium-infected An. parensis mosquitoes were found in the village of Ngaya in the Misungwi district in north-western Tanzania where long-lasting insecticidal nets (LLINs) were already widely implemented [63]. A 2018 study [64] focusing on An. funestus group species composition in several villages of the same district reported over 90% An. funestus s.s. and 6.5% An. parensis. While the aquatic ecology of An. parensis was not within the scope of this paper, studies in rural south-eastern Tanzania noted that An. parensis generally shared aquatic habitats with An. funestus s.s. and An. rivulorum (Kahamba et al. Unpublished data). Further studies are required to understand how vector control interventions might have been associated with the apparently higher importance of An. parensis in this location.

On the basis of our present findings there is a possibility that An. parensis may be contributing to residual malaria transmission, particularly in localities where An. funestus s.s. and other major vector species have been significantly impacted by chemical control interventions. This has previously been observed in the north-eastern part of South Africa, where An. parensis was reported to minimally contribute to residual malaria transmission, following an almost complete suppression of An. funestus s.s. following large-scale IRS implementation [38].

In previous studies, various sibling species within the Anopheles funestus group have been implicated as malaria vectors [37,38,39, 41, 43], resulting in multiple questions regarding the factors influencing their prevalence and roles in disease transmission. For instance, a study conducted in central Kenya reported significant densities of An. parensis inside human dwellings, though with a low human blood index [40]. In our present study, we also collected a significant number of resting An. parensis inside houses in the northern region. Additionally, our current findings, coupled with a previous study [65] conducted in the Muheza district of north-eastern Tanzania, which reported that over 60% of An. parensis caught inside houses had fed on humans despite the availability of cattle, signify the potential role of An. parensis as a contributor to the residual malaria transmission. Consequently, it will be necessary to extend our control efforts beyond the current indoor vector control interventions, to address not just An. parensis, but also other important species, such as An. arabiensis, which is also widespread in Tanzania [34] and tends to bite outdoors [66]. Additionally, there is a need for a thorough understanding of the ecology of An. parensis and other sibling species within the An. funestus group; as well as their responsiveness to current vector control interventions.

Annual entomological inoculation rates (EIR) were computed for different regions and were found to be the highest in areas where An. funestus s.s. dominate as the member of the An. funestus group, such as north-western and southern regions of Kagera and Ruvuma. Notably, Kigoma exhibited the lowest measurable EIR at 18.2 infectious bites per person per year (ib/p/yr) among all regions where infected mosquitoes were found. Among infected mosquitoes, Plasmodium falciparum was the only malarial parasite detected. However, it is essential to note that other Plasmodium species, such as P. ovale and P. malariae, have been previously reported in other country-wide surveys [67,68,69]. One limitation of this study was that the mosquito sampling, primarily designed for species identification and genomic analysis, was insufficient to definitively rule out Plasmodium infections in regions where none of the tested mosquitoes were found to be infected. Consequently, areas reporting zero EIR estimates are simply categorized as having non-estimable EIRs, rather than being considered as having no risk of malaria transmission. It is expected that expanded surveys would reveal non-zero prevalence rates within either the An. funestus group or the An. gambiae complex. Additionally, another limitation of the present study was the inability of the available species-specific PCR assay [56] designed for the An. funestus group, to identify 37% of the collected mosquitoes that had otherwise been morphologically identified as An. funestus.

Conclusions

This study underscores the pivotal role of the An. funestus group in malaria transmission with a particular focus on the prominent An. funestus s.s. Additionally, the study sheds light on the lesser-studied sibling species, An. parensis, which is identified here, for the third time, as playing a role in the transmission of Plasmodium falciparum. Challenges in PCR amplification owing to ITS2 region polymorphisms highlight the limitations of current molecular tools for distinguishing species within the Funestus group. This study contributes to the body of knowledge on malaria vector composition and distribution in Tanzania and emphasizes the critical need for the adaptation of vector control interventions to regional specificities in malaria transmission dynamics. More importantly, the findings call for a deeper investigation into the ecology and vectorial capacity of secondary vectors to enhance malaria control strategies.