- Open Access
Member species of the Anopheles gambiae complex can be misidentified as Anopheles leesoni
Malaria Journal volume 19, Article number: 89 (2020)
Accurate Anopheles species identification is key for effective malaria vector control. Identification primarily depends on morphological analysis of field samples as well as molecular species-specific identifications. During an intra-laboratory assessment (proficiency testing) of the Anopheles funestus group multiplex PCR assay, it was noted that Anopheles arabiensis can be misidentified as Anopheles leesoni, a zoophilic member of the An. funestus group. The aim of this project was, therefore, to ascertain whether other members of the Anopheles gambiae complex can also be misidentified as An. leesoni when using the standard An. funestus multiplex PCR.
The An. funestus multiplex PCR was used to amplify DNA from An. gambiae complex specimens. These included specimens from the laboratory colonies and field samples from the Democratic Republic of Congo. Amplified DNA from these specimens, using the universal (UV) and An. leesoni species-specific primers (LEES), were sequence analysed. Additionally, An. leesoni DNA was processed through the diagnostic An. gambiae multiplex PCR to determine if this species can be misidentified as a member of the An. gambiae complex.
Laboratory-colonized as well as field-collected samples of An. arabiensis, An. gambiae, Anopheles merus, Anopheles quadriannulatus, Anopheles coluzzii as well as Anopheles moucheti produced an amplicon of similar size to that of An. leesoni when using an An. funestus multiplex PCR. Sequence analysis confirmed that the UV and LEES primers amplify a segment of the ITS2 region of members of the An. gambiae complex and An. moucheti. The reverse was not true, i.e. the An. gambiae multiplex PCR does not amplify DNA from An. leesoni.
This investigation shows that An. arabiensis, An. gambiae, An. merus, An. quadriannulatus, An. coluzzii and An. moucheti can be misidentified as An. leesoni when using An. funestus multiplex PCR. This shows the importance of identifying specimens using standard morphological dichotomous keys as far as possible prior to the use of appropriate PCR-based identification methods. Should there be doubt concerning field-collected specimens molecularly identified as An. leesoni, the An. gambiae multiplex PCR and sequencing of the internal transcribed spacer 2 (ITS2) can be used to eliminate false identifications.
Malaria is a major vector borne disease that is most prevalent in sub-Saharan Africa. There were approximately 213 million cases and 380,000 malaria-related deaths in this region in 2018, accounting for 93% of cases and 94% of deaths from malaria reported globally .
A key component of malaria control is suppression of Anopheles mosquito vectors.
The primary methods used for malaria vector control are indoor residual spraying (IRS) of formulated insecticides, insecticide-treated nets (ITN) and larval source management (LSM) . These can be incorporated into broader, tailored strategies within an integrated vector management (IVM) framework . Other initiatives under development include attractive toxic sugar baits (ATSB), spatial repellents, housing improvements, endectocide use and genetic approaches [4,5,6,7,8].
The major malaria vector mosquito species in Africa are Anopheles gambiae, Anopheles arabiensis and Anopheles coluzzii of the An. gambiae species complex, and Anopheles funestus of the An. funestus species group [9,10,11,12]. In addition to these, other species within these taxa—including Anopheles merus of the An. gambiae complex, and Anopheles rivulorum, Anopheles parensis, Anopheles vaneedeni and Anopheles leesoni of the An. funestus group—have been implicated as secondary malaria vectors at various African localities [11, 13,14,15,16,17,18,19,20,21,22,23] to mention but a few. Importantly, primary and secondary vector species often occur in sympatry in varying combinations depending on locality , different species may display different behaviours, such as indoor or outdoor feeding and resting [21, 24], and may vary in their susceptibilities to insecticide [19, 25,26,27]. It is, therefore, necessary to identify the entomological drivers of localized malaria transmission by using tailored vector surveillance strategies. These include judicious use of sampling techniques followed by species identifications, vector incrimination (sporozoite detection) and insecticide susceptibility assessments of these populations. The information generated in this way provides the necessary baseline data needed to guide control interventions that target incriminated vector populations based on their specific traits, such as their resting and feeding preferences (indoor vs. outdoor), their preferred breeding sites (perennial vs. temporary) and their insecticide susceptibilities. The same surveillance techniques can also be used to assess the effectiveness of interventions post implementation.
The accurate identification of malaria vector species is, therefore, central to the application of successful vector control interventions, primarily by ensuring the efficient and effective use of limited resources available to vector control programmes. Misidentification of Anopheles species can lead to misapplication of vector control interventions [28,29,30]. An example comes from Zimbabwe in the early 1970s, when An. quadriannulatus, a non-vector member of the An. gambiae complex, could not easily be distinguished from the vector An. arabiensis. Insecticide susceptibility tests on mixed samples of An. quadriannulatus and An. arabiensis suggested susceptibility to the insecticide dieldrin [28, 29]. What was not however evident at the time was that the samples that succumbed to dieldrin exposure were An. quadriannulatus, while the few survivors were An. arabiensis, implying resistance in the vector population. The use of dieldrin for indoor residual spraying did not therefore achieve the desired effect on malaria transmission, and the insecticide regimen was subsequently changed once accurate species identifications were used to differentiate between resistance in the An. arabiensis vector population and susceptibility in the An. quadriannulatus non-vector population [28, 29].
Identification to species of field-collected mosquito specimens depends on the use of external morphological characters followed by molecular methods where indicated [9, 10, 31]. This is especially pertinent for members of the An. gambiae complex and An. funestus group whose member species vary significantly in their behavioural traits and vector competencies. The subsequent use of diagnostic molecular procedures to identify specimens to species is required because of morphological similarities between members within each taxon [32, 33].
Morphological identification of mosquitoes can be done at district level and is not reliant on expensive molecular equipment. Subsequent molecular analysis to identify indicated specimens to species (using multiplex PCR assays) is generally conducted at established laboratories at the national level or within research institutes with sufficient capacity [34,35,36]. These species-specific assays are an important diagnostic tool and are regularly used in laboratories for research and routine vector surveillance [34,35,36]. Molecular sequencing of target genes has been used for Anopheles species identifications [21, 37,38,39,40,41]. Laboratory infrastructure and cost, however, preclude this method from being routinely used in support of vector surveillance.
Regardless of the method used for molecular species identification, quality assurance (QA) of the data produced is critical. This is because the pertinence and relevance of all follow-on associative analyses (vector incrimination/sporozoite detection, insecticide susceptibility assessments, associated behaviours) is dependent on accurate species identification. An essential requirement of QA is regular proficiency testing of laboratory staff to monitor their competency in the application of diagnostic assays [42, 43]. A recent proficiency assessment exercise conducted at the Vector Control Reference Laboratory of the National Institute for Communicable Diseases (NICD) in Johannesburg was based on an intra-laboratory comparison using the An. funestus multiplex PCR method [35, 36]. Unexpectedly, An. arabiensis, which was used as a blind negative control, produced an amplicon of similar size to that of An. leesoni when using the An. funestus PCR.
It has recently been established that specimens not of the An. gambiae complex or An. funestus group can be misidentified as members of either of these taxa by using the corresponding multiplex PCR assays in the absence or misidentification of a priori morphological identification . Morphological identification on field samples can be problematic if samples are damaged due to mosquito handling (collection method, preservation processing) or due to age of the mosquito samples. Based on these data, the aim of this study was to ascertain whether An. gambiae complex specimens can easily be misidentified as An. leesoni when using the An. funestus multiplex PCR.
In silico sequence analysis of Anopheles funestus multiplex PCR primers and Anopheles gambiae complex species internal transcribed spacer 2 (ITS2) region
The sequences of primers used in the An. funestus multiplex PCR [35, 36] were compared with ITS2 sequences from the An. gambiae complex species to identify sequence similarities. Nucleotide Basic Local Alignment Search Tool (BLAST) (https://blast.ncbi.nlm.nih.gov/Blast.cgi) and Emboss Needle pairwise sequence alignment tool (https://www.ebi.ac.uk/Tools/psa/emboss_needle/nucleotide.html) were used.
Laboratory-reared Anopheles gambiae complex species samples
Specimens of An. funestus, An. arabiensis, An. gambiae, An. merus and An. quadriannulatus (FUMOZ, KGB, COGS, MAFUS and SANGWE colonies respectively) housed in the Botha De Meillon insectary at the National Institute for Communicable Diseases in Johannesburg were used. The An. leesoni positive control was obtained from a field sample from Limpopo Province, South Africa, in December 2016. This sample was verified as An. leesoni by morphological and PCR species identification as well as ITS2 sequence analysis.
DNA extraction: DNA was extracted from the An. funestus, An. leesoni, An. arabiensis, An. gambiae, An. merus and An. quadriannulatus specimens using prepGEM Insect DNA extraction kit (ZyGEM, PIN0020).
Anopheles funestus multiplex PCR: Each PCR reaction contained extracted DNA from An. funestus and An. leesoni positive controls; a “no DNA template” negative control (PCR master mix without DNA template); “extraction kit” negative controls (PCR master mix with extraction mix performed without mosquito sample), and extracted DNA from An. arabiensis, An. gambiae, An. merus and An. quadriannulatus specimens.
Several variations of the An. funestus multiplex PCR were performed during this investigation: (1) Standard An. funestus multiplex PCR with the annealing temperature set at 45 °C as per the protocol by Koekemoer et al.  and Cohuet et al.  or with the exception of the annealing temperature set at 50 °C; (2) Standard An. funestus multiplex PCR with the exception of the LEES primer being omitted from the PCR reaction, and with the annealing temperature set at 45 °C or 50 °C; (3). Standard An. funestus multiplex PCR with the exception of the PCR reaction only including the UV and LEES primers, and with the annealing temperature set at 45 °C or 50 °C. The different variations of the An. funestus multiplex PCR were used to test whether a non-specific PCR amplicon is produced while using the DNA of An. gambiae complex specimens in the PCR. Subsequently, the An. funestus multiplex PCRs with or without only the LEES reverse primer were used to establish whether this primer is responsible for amplification of DNA from An. gambiae complex specimens in the PCR. Different annealing temperatures were used in the PCRs to determine whether the annealing temperature reduces non-specific amplification of DNA from the An. gambiae complex when performing an An. funestus multiplex PCR.
Anopheles gambiae multiplex PCR: PCR was performed according to the protocol by Scott et al. . The PCR reaction contained extracted DNA from An. arabiensis, An. gambiae, An. merus and An. quadriannulatus positive controls; a “no DNA template” negative control (PCR master mix without DNA template); “extraction kit” negative controls (PCR master mix with extraction mix performed without mosquito sample) and extracted DNA from an An. leesoni positive control.
The PCR products from the An. funestus and An. gambiae amplifications were electrophoresed on a 2.5% agarose gel and viewed with a ChemiDoc XRS + Imaging system (Biorad).
The An. leesoni sized amplicons produced by the UV and LEES primers were purified and sequenced through Macrogen (http://www.macrogen.com). Subsequently, the chromatograms of the sequences were manually edited using BioEdit version 7.2.5  and analysed using the BLAST tool (https://blast.ncbi.nlm.nih.gov/Blast.cgi) to determine sequence identity between the PCR products and the ITS2 sequences of the An. gambiae complex.
Field sample investigations
Morphological identification was conducted on all field samples, which were (mis)identified as belonging to the An. funestus group. Species identification was performed on a subset of field samples (n = 28) molecularly identified as An. leesoni using the An. funestus multiplex PCR . The ITS2 PCR and mDNA cytochrome oxidase I (COI) loci [35, 37] PCR followed by sequencing of the PCR amplicons was used for these species identifications. The resulting sequences were analysed using nBLAST (https://blast.ncbi.nlm.nih.gov/Blast.cgi). In addition, these samples were also amplified using conventional PCR methods for the identification of mosquitoes in the An. gambiae complex [34, 45] and An. moucheti complex by multiplex PCR assays , to rule out the possibility of morphological misidentification at the start.
PCR using the UV and LEES primers of the field samples was performed. Anopheles gambiae complex specimens used as controls in the PCR were An. gambiae sensu stricto (s.s.) (KISUMU colony), An. coluzzii (AKRON colony), An. gambiae/coluzzii hybrid (ASEMBO colony), An. arabiensis (KGB colony) as well as An. funestus (s.s.) (FUMOZ colony). Sequencing analysis was performed on the resultant PCR amplicons of the field samples.
Intra-laboratory proficiency assessment of the An. funestus multiplex PCR assay revealed that An. arabiensis DNA amplifies a ~ 150 bp fragment and can therefore be incorrectly identified as An. leesoni, which amplifies a fragment of similar size . In silico analyses of the primer sequence similarity revealed a 100% sequence identity of UV to the 3′ region of the 5.8S region flanking the ITS2 region of members of the An. gambiae complex (Table 1) as can be expected from this highly conserved region . The species-specific reverse primers shared a variable degree of identity with the An. gambiae complex (Table 1). The LEES reverse primer had a 77% sequence identity with the ITS2 region of An. arabiensis. It was also the only primer which showed over 50% sequence identity with the ITS2 region of other members of the An. gambiae complex in the location 120 to 153 bp downstream of the UV primer binding site, therefore producing an amplicon size diagnostic for An. leesoni. Additionally, the LEES primer had the highest number of consecutive bases (7) at the 3′ end that directly bound with the ITS2 region of the An. gambiae complex member species (Table 1).
The An. funestus multiplex PCR assay was subsequently evaluated on other members of the An. gambiae complex, and all species tested produced An. leesoni diagnostic PCR product (~ 150 bp, Table 2). The exclusion of LEES primer resulted in no amplification (Table 2) regardless of An. gambiae complex species or annealing temperature analysed.
Amplification of DNA from members of the An. gambiae complex using only the UV and LEES primers and the An. funestus PCR protocol yielded a ~ 150 bp PCR product from all species (Fig. 1; Table 2). Sequence analysis of these PCR amplicons using the UV and LEES primers revealed that there was 99–100% sequence identity between the amplicons and the ITS2 region of An. gambiae complex species. Furthermore, sequencing of the PCR amplicons, using UV as the sequencing primer, revealed that the LEES primer sequence was incorporated into the PCR amplicon sequence. This confirms that the LEES and UV primers are responsible for the 150 bp fragment when An. funestus PCR is used to amplify the ITS2 of An. gambiae complex species, which leads to their misidentification as An. leesoni.
Field sample data
A large number of field-collected samples from the Democratic Republic of the Congo were morphologically identified as An. funestus group and subsequently molecularly identified as An. leesoni. The ITS2 and COI regions were amplified by PCR and sequenced, showing that a subset of these samples were An. gambiae s.s. (n = 13) and An. moucheti (n = 12). Those identified as An. gambiae s.s. through sequencing were further confirmed by An. gambiae complex PCR [34, 45]. The samples identified as An. moucheti through sequencing were further confirmed by An. moucheti multiplex PCR assay . PCR amplification of these samples using the UV and LEES primers produced an An. leesoni sized amplicon between 100 and 200 bp. Additionally, An. gambiae complex specimens that were used as controls in the PCR—An. gambiae s.s., An. coluzzii, An. gambiae/coluzzii hybrid and An. arabiensis—also produced similar-sized fragments (Fig. 2). Sequencing of the field samples using the UV and LEES primers in the PCR confirmed that the LEES primer fragment was incorporated in the sequences of the PCR amplicons.
Anopheles gambiae multiplex PCR does not amplify DNA from Anopheles leesoni
It has been demonstrated that An. gambiae complex member species can be misidentified as An. leesoni by PCR. In contrast, the An. gambiae multiplex PCR does not amplify DNA from An. leesoni and cannot therefore misidentify this species as a member of the An. gambiae complex.
The importance of correct identification of Anopheles species in malaria vector control programmes is critical in terms of choice of control intervention and insecticide product. Accurate species identification enables assessments of vector competence, insecticide susceptibilities and important behavioural characteristics (such as feeding and resting behaviours) by species, leading to the design of coherent insecticide-based control strategies that can be enhanced by additional methodologies for malaria elimination. These data indicate that if members of the An. gambiae complex (An. arabiensis, An. gambiae, An. coluzzii, An. merus and An. quadriannulatus) as well as An. moucheti are morphologically incorrectly identified as An. funestus group, they can be falsely identified as An. leesoni when using an An. funestus multiplex PCR.
This is due to high primer (specifically UV and LEES) sequence identity between the two species groups. The UV primer showed a 100% sequence identity to the ITS2 region of the An. gambiae complex. This is not surprising, since the UV primer is in the conserved region of the 5.8S ribosomal RNA gene . The LEES primer sequence identity with the An. gambiae complex ITS2 region ranged between 53% and 77%. The likely reason for the amplification of An. gambiae complex DNA using the LEES primer is due to the seven consecutive bases at its 3′end. These bases specifically bind to the ITS2 region of the An. gambiae complex. In a PCR reaction, this leads to the incorporation of the LEES primer 120 bp downstream of the UV primer binding region to produce an amplicon of the An. gambiae complex species ITS2, which had the LEES primer binding region in its sequence as was evidenced by the sequencing data. This is also true for An. moucheti. This scenario is expected to be the case in other species of the An. gambiae complex, such as An. coluzzii, Anopheles bwambae and Anopheles amharicus, because the same 7 bases of the LEES primer bind to the ITS2 regions of these species (GenBank Accession numbers: KT160244.1; GQ870320.1 and GQ870316.1). Indeed, an An. coluzzii sample that was used as a control in the PCR (using the UV and LEES primers), for the field samples analysis, produced a positive An. leesoni sized amplicon band. The fact that An. gambiae complex species can be misidentified as An. leesoni supports a recent publication by Erlank et al. , which demonstrated that Anopheles rufipes and Anopheles rhodesiensis can misleadingly be identified as An. leesoni with the use of An. funestus multiplex PCR.
Different Anopheles species vary in their malaria vectorial capacities as well as in their feeding and resting habits [11, 47]. They may also have different insecticide susceptibility profiles and, therefore, their correct identification to species is vital for the implementation of an efficient vector control strategy based on accurate vector incrimination and appropriate use of insecticides. Members of the An. gambiae complex and An. funestus group are often found in sympatry [11, 47, 48]. It is, therefore, likely that the collection of field samples could contain a mix of species, making accurate identification to species essential.
These data also raise concerns over previously published records of vector incrimination of species identified as An. leesoni by An. funestus multiplex PCR alone, which was common practice at the time . This stresses the importance to confirm species identity through ITS2 and/or COI sequencing to prevent mis-interpretation of data.
There are several steps necessary to minimize the misidentification of species from the An. gambiae complex as An. leesoni. The first step, which is also highlighted by Erlank et al. , is to accurately identify the samples morphologically. However, morphological species identification is largely dependent on the condition of the sample—field-collected samples may be damaged—as well as the skill of personnel involved, the equipment they have and their workload. In the event that a field sample is suspected to be An. leesoni via the An. funestus multiplex PCR, but the morphological identification is not certain, it is advisable to use An. gambiae multiplex PCR on the DNA of the sample. The results from this study indicate that DNA from a true An. leesoni sample does not amplify using the An. gambiae multiplex PCR, eliminating any uncertainty regarding the identity of the field sample. Additionally, should a suspected An. leesoni female test positive for P. falciparum sporozoites by ELISA  and/or PCR [50, 51], it is necessary to perform an ITS2 and/or COI sequence confirmation of the mosquito sample so as to eliminate any ambiguity regarding vector status [21, 35, 37].
Member species of the An. gambiae complex can be misidentified as An. leesoni when analysed using the An. funestus group multiplex PCR. This is best avoided by accurate morphological identification prior to PCR assessments and can also be resolved by further analysing samples using the An. gambiae multiplex PCR where sequencing technology is not available. Lastly, it is important for the reference laboratory performing species identifications to periodically conduct quality control assessments and proficiency testing of laboratory personnel. Sequence analysis should be performed to confirm the species identity in cases of conflicting results. This ensures that the correct species identifications are reported to malaria vector control programmes.
Availability of data and materials
The datasets used and/or analysed during the current study are available from the corresponding author on reasonable request.
Attractive toxic sugar baits
Basic Local Alignment Search Tool
Cytochrome oxidase I
Enzyme-linked immunosorbent assay
An. funestus species-specific reverse primer
Indoor residual spraying
Insecticide treated nets
Internal transcribed spacer 2
An. leesoni species-specific reverse primer
Larval source management
National Institute for Communicable Diseases
An. parensis species-specific reverse primer
Polymerase chain reaction
An. rivulorum species-specific reverse primer
An. rivulorum-like species specific reverse primer
Universal forward primer
An. vaneedeni species-specific reverse primer
World Health Organization
WHO. World malaria report 2019. Geneva: World Health Organization; 2019. https://www.who.int/publications-detail/world-malaria-report-2019. Accessed Dec 2019.
WHO. Global Vector Control Response. Geneva: World Health Organization; 2017. https://www.who.int/vector-control/publications/global-control-response/en/. Accessed Mar 2019.
WHO. Handbook for integrated vector management. Geneva: World Health Organization; 2012. https://www.who.int/neglected_diseases/vector_ecology/resources/9789241502801/en/. Accessed Mar 2019.
Norris EJ, Coats JR. Current and future repellent technologies: the potential of spatial repellents and their place in mosquito-borne disease control. Int J Environ Res Public Health. 2017;14:124.
Marshall JM, White MT, Ghani AC, Schlein Y, Muller GC, Beier JC. Quantifying the mosquito’s sweet tooth: modelling the effectiveness of attractive toxic sugar baits (ATSB) for malaria vector control. Malar J. 2013;12:291.
Tusting LS, Ippolito MM, Willey BA, Kleinschmidt I, Dorsey G, Gosling RD, et al. The evidence for improving housing to reduce malaria: a systematic review and meta-analysis. Malar J. 2015;14:209.
James S, Collins FH, Welkhoff PA, Emerson C, Godfray HCJ, Gottlieb M, et al. Pathway to deployment of gene drive mosquitoes as a potential biocontrol tool for elimination of malaria in sub-Saharan Africa: recommendations of a scientific working group. Am J Trop Med Hyg. 2018;98(Suppl 6):1–49.
Burrows J, Slater H, Macintyre F, Rees S, Thomas A, Okumu F, et al. A discovery and development roadmap for new endectocidal transmission-blocking agents in malaria. Malar J. 2018;17:462.
Gillies MT, De Meillon B. The Anophelinae of Africa south of the Sahara, vol. 54. Johannesburg: Publications of the South African Institute for Medical Research; 1968.
Gillies MT, Coetzee M. A supplement to the Anophelinae of Africa south of the Sahara, vol. 55. Johannesburg: Publications of the South African Institute for Medical Research; 1987.
Sinka ME, Bangs MJ, Manguin S, Rubio-Palis Y, Chareonviriyaphap T, Coetzee M, et al. A global map of dominant malaria vectors. Parasit Vectors. 2012;5:69.
Akogbéto MC, Salako AS, Dagnon F, Aïkpon R, Kouletio M, Sovi A, et al. Blood feeding behaviour comparison and contribution of Anopheles coluzzii and Anopheles gambiae, two sibling species living in sympatry, to malaria transmission in Alibori and Donga region, northern Benin, West Africa. Malar J. 2018;17:307.
Wilkes TJ, Matola YG, Charlwood JD. Anopheles rivulorum, a vector of human malaria in Africa. Med Vet Entomol. 1996;10:108–10.
Temu EA, Minjas JN, Tuno N, Kawada H, Takagi M. Identification of four members of the Anopheles funestus (Diptera: Culicidae) group and their role in Plasmodium falciparum transmission in Bagamoyo coastal Tanzania. Acta Trop. 2007;102:119–25.
Cuamba N, Mendis C. The role of Anopheles merus in malaria transmission in an area of southern Mozambique. J Vector Borne Dis. 2009;46:157–9.
Kipyab PC, Khaemba BM, Mwangangi JM, Mbogo CM. The bionomics of Anopheles merus (Diptera: Culicidae) along the Kenyan coast. Parasit Vectors. 2013;6:37.
Mulamba C, Irving H, Riveron JM, Mukwaya LG, Birungi J, Wondji CS. Contrasting Plasmodium infection rates and insecticide susceptibility profiles between the sympatric sibling species Anopheles parensis and Anopheles funestus s.s.: a potential challenge for malaria vector control in Uganda. Parasit Vectors. 2014;7:71.
Afrane YA, Bonizzoni M, Yan G. Secondary malaria vectors of sub-Saharan Africa: threat to malaria elimination on the continent? In: Rodriguez-Morales AJ, editor. Current topics in malaria. London: IntechOpen; 2016. p. 473–90.
Stevenson JC, Norris DE. Implicating cryptic and novel anophelines as malaria vectors in Africa. Insects. 2016;8:1.
St Laurent B, Cooke M, Krishnankutty SM, Asih P, Mueller JD, Kahindi S, et al. Molecular characterization reveals diverse and unknown malaria vectors in the western Kenyan highlands. Am J Trop Med Hyg. 2016;94:327–35.
Burke A, Dandalo L, Munhenga G, Dahan-Moss Y, Mbokazi F, Ngxongo S, et al. A new malaria vector mosquito in South Africa. Sci Rep. 2017;7:43779.
Kyalo D, Amratia P, Mundia CW, Mbogo CM, Coetzee M, Snow RW. A geo-coded inventory of anophelines in the Afrotropical Region south of the Sahara: 1898–2016. Wellcome Open Res. 2017;2:57.
Ogola EO, Fillinger U, Ondiba IM, Villinger J, Masiga DK, Torto B, et al. Insights into malaria transmission among Anopheles funestus mosquitoes, Kenya. Parasit Vectors. 2018;11:577.
Dandalo LC, Brooke BD, Munhenga G, Lobb LN, Zikhali J, Ngxongo SP, et al. Population dynamics and Plasmodium falciparum (Haemosporida: Plasmodiidae) infectivity rates for the malaria vector Anopheles arabiensis (Diptera: Culicidae) at Mamfene, KwaZulu-Natal, South Africa. J Med Entomol. 2017;54:1758–66.
Killeen GF, Marshall JM, Kiware SS, South AB, Tusting LS, Chaki PP, et al. Measuring, manipulating and exploiting behaviours of adult mosquitoes to optimise malaria vector control impact. BMJ Glob Health. 2017;2:e000212.
Killeen GF, Chaki PP, Reed TE, Moyes CL, Govella NJ. Entomological surveillance as a cornerstone of malaria elimination: a critical appraisal. In: Manguin S, Dev V, editors. Towards malaria elimination. London: InTech Open; 2018. p. 403–29.
Sherrard-Smith E, Skarp JE, Beale AD, Fornadel C, Norris LC, Moore SJ, et al. Mosquito feeding behavior and how it influences residual malaria transmission across Africa. Proc Natl Acad Sci USA. 2019;116:15086–95.
Green CA. Malaria epidemiology and Anopheline cytogenetics. In: Pal R, Kitzmiller JB, Kanda T, editors. Cytogenetics and genetics of vectors. Amsterdam: Elsevier Biomedical Press; 1981. p. 21–9.
Hunt RH, Mahon RJ. Collections of Anopheles quadriannulatus (Diptera:Culcidae) from human habitations in Southern Africa. J Entomol Soc South Afr. 1986;49:390–1.
Choochote W, Saeung A. Systematic techniques for the recognition of Anopheles species complexes. In: Manguin S, editor. Anopheles mosquitoes—new insights into malaria vectors. London: IntechOpen; 2013. p. 57–79.
WHO. Malaria surveillance, monitoring & evaluation: a reference manual. Geneva: World Health Organization; 2018. https://apps.who.int/iris/bitstream/handle/10665/272284/9789241565578-eng.pdf?ua=1. Accessed Dec 2019.
Coetzee M, Koekemoer LL. Molecular systematics and insecticide resistance in the major African malaria vector Anopheles funestus. Annu Rev Entomol. 2013;58:393–412.
Erlank E, Koekemoer LL, Coetzee M. The importance of morphological identification of African anopheline mosquitoes (Diptera: Culicidae) for malaria control programmes. Malar J. 2018;17:43.
Scott JA, Brogdon WG, Collins FH. Identification of single specimens of the Anopheles gambiae complex by the polymerase chain reaction. Am J Trop Med Hyg. 1993;49:520–9.
Koekemoer LL, Kamau L, Hunt RH, Coetzee M. A cocktail polymerase chain reaction assay to identify members of the Anopheles funestus (Diptera: Culicidae) group. Am J Trop Med Hyg. 2002;66:804–11.
Cohuet A, Simard F, Toto JC, Kengne P, Coetzee M, Fontenille D. Species identification within the Anopheles funestus group of malaria vectors in Cameroon and evidence for a new species. Am J Trop Med Hyg. 2003;69:200–5.
Lobo NF, St Laurent B, Sikaala CH, Hamainza B, Chanda J, Chinula D, et al. Unexpected diversity of Anopheles species in Eastern Zambia: implications for evaluating vector behavior and interventions using molecular tools. Sci Rep. 2015;5:17952.
Norris LC, Norris DE. Phylogeny of anopheline (Diptera: Culicidae) species in southern Africa, based on nuclear and mitochondrial genes. J Vector Ecol. 2015;40:16–27.
Mouatcho J, Cornel AJ, Dahan-Moss Y, Koekemoer LL, Coetzee M, Braack L. Detection of Anopheles rivulorum-like, a member of the Anopheles funestus group, in South Africa. Malar J. 2018;17:195.
Carter TE, Yared S, Hansel S, Lopez K, Janies D. Sequence-based identification of Anopheles species in eastern Ethiopia. Malar J. 2019;18:135.
Burke A, Dahan-Moss Y, Duncan F, Qwabe B, Coetzee M, Koekemoer L, et al. Anopheles parensis contributes to residual malaria transmission in South Africa. Malar J. 2019;18:257.
International Standard. ISO/IEC 17025:2017. 3rd edition. General requirements for the competence of testing and calibration laboratories. Published in Switzerland. ISO/IEC; 2017.
Clinical and Laboratory Standards Institute. GP27-A2 Vol.27 No.8. Using Proficiency Testing to Improve the Clinical Laboratory, Approved Guideline-second edition.
Hall TA. BioEdit: a user-friendly biological sequence alignment editor and analysis program for Windows 95/98/NT. Nucleic Acids Symp Ser. 1999;41:95–8.
Wilkins EE, Howell PI, Benedict MQ. IMP PCR primers detect single nucleotide polymorphisms for Anopheles gambiae species identification, Mopti and Savanna rDNA types, and resistance to dieldrin in Anopheles arabiensis. Malar J. 2006;5:125.
Kengne P, Antonio-Nkondjio C, Awono-Ambene HP, Simard F, Awolola TS, Fontenille D. Molecular differentiation of three closely related members of the mosquito species complex, Anopheles moucheti, by mitochondrial and ribosomal DNA polymorphism. Med Vet Entomol. 2007;21:177–82.
Das S, Henning TC, Simubali L, Hamapumbu H, Nzira L, Mamini E, et al. Underestimation of foraging behaviour by standard field methods in malaria vector mosquitoes in southern Africa. Malar J. 2015;14:12.
Zawada JW, Dahan-Moss YL, Muleba M, Dabire RK, Maïga H, Venter N, et al. Molecular and physiological analysis of Anopheles funestus swarms in Nchelenge, Zambia. Malar J. 2018;17:49.
Durnez L, Van Bortel W, Denis L, Roelants P, Veracx A, Trung HD, et al. False positive circumsporozoite protein ELISA: a challenge for the estimation of the entomological inoculation rate of malaria and for vector incrimination. Malar J. 2011;10:195.
Snounou G, Pinheiro L, Gonçalves A, Fonseca L, Dias F, Brown KN, et al. The importance of sensitive detection of malaria parasites in the human and insect hosts in epidemiological studies, as shown by the analysis of field samples from Guinea Bissau. Trans R Soc Trop Med Hyg. 1993;87:649–53.
Bass C, Nikou D, Blagborough AM, Vontas J, Sinden RE, Williamson MS, et al. PCR-based detection of Plasmodium in Anopheles mosquitoes: a comparison of a new high-throughput assay with existing methods. Malar J. 2008;7:177.
The U.S. President’s Malaria Initiative and the Institut National de Recherche Biomedical are thanked for their support in the collection of the DRC specimens. Elodie Ekoka is thanked for her participation in the proficiency testing. Prof. Maureen Coetzee is thanked for her advice regarding the manuscript.
This work was supported by the National Institute for Communicable Diseases (Grant number: 90932), DST/NRF South African Research Chairs Initiative (Grant number: 64763), and a Pilot Project Grant from the Eck Institute for Global Health, University of Notre Dame, IN.
Ethics approval and consent to participate
The research represented in this article did not require clearance from the Human research Ethics Committee. Waiver no: AREC-101210-002.
Consent for publication
The authors declare that they have no competing interests.
Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
About this article
Cite this article
Dahan-Moss, Y., Hendershot, A., Dhoogra, M. et al. Member species of the Anopheles gambiae complex can be misidentified as Anopheles leesoni. Malar J 19, 89 (2020). https://doi.org/10.1186/s12936-020-03168-x
- Species identification
- Anopheles leesoni
- Anopheles gambiae multiplex PCR
- An. funestus multiplex PCR
- Dichotomous keys