Skip to content


Malaria Journal

Open Access

The importance of morphological identification of African anopheline mosquitoes (Diptera: Culicidae) for malaria control programmes

Malaria Journal201817:43

Received: 25 September 2017

Accepted: 15 January 2018

Published: 22 January 2018



The correct identification of disease vectors is the first step towards implementing an effective control programme. Traditionally, for malaria control, this was based on the morphological differences observed in the adults and larvae between different mosquito species. However, the discovery of species complexes meant that genetic tools were needed to separate the sibling species and today there are standard molecular techniques that are used to identify the two major malaria vector groups of mosquitoes. On the assumption that species-diagnostic DNA polymerase chain reaction (PCR) assays are highly species-specific, experiments were conducted to investigate what would happen if non-vector species were randomly included in the molecular assays.


Morphological keys for the Afrotropical Anophelinae were used to provide the a priori identifications. All mosquito specimens were then subjected to the standard PCR assays for members of the Anopheles gambiae complex and Anopheles funestus group.


One hundred and fifty mosquitoes belonging to 11 morphological species were processed. Three species (Anopheles pretoriensis, Anopheles rufipes and Anopheles rhodesiensis) amplified members of the An. funestus group and four species (An. pretoriensis, An. rufipes, Anopheles listeri and Anopheles squamosus) amplified members of the An. gambiae complex.


Morphological identification of mosquitoes prior to PCR assays not only saves time and money in the laboratory, but also ensures that data received by malaria vector control programmes are useful for targeting the major vectors.




Malaria continues to be an ongoing problem in African countries south of the Sahara and although a lot has been achieved in the past 15 years, millions of people still remain at risk of contracting the parasite [1]. Africa provides a stable and ecologically diverse ecosystem and is home to the most efficient malaria vectors in the world [24], and is likely to remain so in the face of global climate change [5]. The major anopheline malaria vectors across sub-Saharan Africa are Anopheles funestus s.s. and three members of the Anopheles gambiae complex: An. gambiae s.s., Anopheles coluzzii and Anopheles arabiensis [24, 69]. There are, however, many additional species outside of these that play a role in malaria transmission within their geographic distribution, for example the Anopheles moucheti and Anopheles nili groups [3], and a host of secondary or incidental vectors [1012]. Considering that the genus Anopheles contains over 500 species globally, of which only a few are considered important species for malaria transmission [2, 13], the morphological identification of species is crucial in order to target scarce resources for controlling the malaria vectors only.

Species groups and species complexes are common within the genus Anopheles [14] and this complicates vector control since not all species within a complex have similar behaviours or similar roles in malaria transmission [3, 4, 6]. In the An. gambiae complex, for example, species range from the non-vectors Anopheles quadriannulatus and Anopheles amharicus to minor vectors Anopheles melas, Anopheles merus and Anopheles bwambae, to the major vectors An. gambiae, An. coluzzii and An. arabiensis [4, 6, 9]. Genetic tools to identify the species have ranged from cross-mating wild mosquitoes with known laboratory colonies [15] to chromosomal banding patterns [16] and enzyme electrophoresis [17]. Today, the Scott et al. [18] molecular species-diagnostic PCR method is the gold standard for identifying members of the An. gambiae complex, and the five most common members of the An. funestus group are identified by the multiplex PCR assay of Koekemoer et al. [19]. However, prior to molecular analysis, accurate morphological identification is critical if correct interpretation of the results is to be used for vector control programme planning.

The purpose of the present study was to evaluate the impact of incorrect or no morphological identification of 150 specimens of 11 anopheline species from different African countries, by determining if these species could be misidentified using the standard vector-specific An. gambiae complex [18] and An. funestus group [19] PCR assays.


Mosquito collections

The species used in this study came from five African countries: Mali and Guinea in West Africa and Namibia, Botswana and South Africa in southern Africa. Collections were made between 2009 and 2017 by different entomological field teams on request and were made by sampling larvae, indoor resting catches and outdoor biting collections. Some were once off collections where field work was being conducted in other African countries, while other collections were made during routine surveillance around South African provinces. The inclusion of more specimens of the less abundant species in this study was limited due to methods of collection, species distribution and number of times that field sites were visited. Adults, larvae and pupae were brought back to Johannesburg, South Africa where further rearing of immatures to adults and processing of adult samples took place (Table 1). Adult mosquitoes were preserved in silica tubes for further morphological and molecular identification.
Table 1

Anopheline species collected from five African countries on which different species-diagnostic PCR analyses were conducted



GPS coordinates

Morphological ID

Molecular ID using An. gambiae complex PCR

Molecular ID using An. funestus group PCR

Molecular ID using An. funestus-like PCR

Molecular ID using An. rivulorum-like PCR





An. wellcomei group (n = 1)

Guinea Conakry




An. rufipes (n = 8)

An. leesoni (5)





An. rufipes (n = 10)

An. rivulorum (1)

An. leesoni (1)


Katima Mulilo



An. squamosus (n = 2)

An. gambiae (2)

South Africa




An. rufipes (n = 19)





An. crypticus (n = 17)


An. marshallii

(n = 1)





An. coustani (n = 6)


An. tenebrosus (n = 1)


An. pretoriensis (n = 20)


An. rufipes (n = 9)

An. gambiae/An. merus (1)


An. maculipalpis (n = 10)



23°55′ 12.00″S

31°15′ 36.00″E

An. maculipalpis (n = 1)


An. rhodesiensis (n = 1)

An. leesoni (1)


An. listeri (n = 12)

An. gambiae (3)

An. merus (2)


Kruger National Park



An. pretoriensis (n = 26)

An. arabiensis (1)

An. gambiae (3)

An. funestus (3)

An. vaneedeni (1)


An. rufipes (n = 6)

An. arabiensis (3)

An. gambiae (2)


The total sample size (n) is indicated per morphological species according to each location and the number of specimens that amplified during the PCR analyses is shown for each assay

Laboratory analyses

Adults were morphologically identified to species using the dichotomous key of Gillies and Coetzee [6]. DNA was extracted from a mosquito leg or wing using the prepGEM Insect DNA extraction kit (ZyGEM; E3BG6M; New Zealand). The amount of DNA used for each specimen was standard for the controls and specimens (1 µl). The DNA concentration that was present in the 1 µl of DNA extraction product was determined with the use of the NanoDrop One spectrophotometer (Thermo Scientific; Cat No. 13400518). A random sample (n = 42) produced an average of 39 ng/µl for the samples compared to the 44 ng/µl for the positive controls. Each specimen was then processed using PCR assays according to the standard protocols for the An. gambiae complex [18], An. funestus group [19], An. funestus-like [20] and Anopheles rivulorum-like [21]. Due to the fact that a number of non-specific amplification occurred during the latter two assays, only the first 111 samples were processed according to these protocols. Controls consisted of laboratory colony mosquitoes (An. funestus FUMOZ Mozambique; An. coluzzii NAG Nigeria; An. arabiensis AMAL South Africa; An. merus MAFUS South Africa and An. quadriannulatus SANGWE Zimbabwe) and two negative controls (one for DNA extraction and one for the PCR mix). DNA of a Malawian An. funestus-like specimen [20] was used for the An. funestus-like assay, and Kruger National Park specimens were used for the An. rivulorum-like control. The PCR product of each assay was electrophoresed on a 2.5% TAE agarose gel.

Gels were viewed with a Geldoc system and the amplicon sizes determined by comparing them with the base pair sizes of the DNA ladder. The base pair sizes were analysed with the QIAxcel Advanced system and DNA screening kit (2400) (Lot No. 154049015; Germany).


A total of 150 mosquito specimens belonging to 11 different anopheline species were analysed using four species-diagnostic PCR assays (Table 1). Five of the species amplified and produced diagnostic fragments for either An. funestus group or An. gambiae complex species or both (Table 2), but no amplification was observed for Anopheles crypticus, Anopheles coustani, Anopheles tenebrosus, Anopheles wellcomei, Anopheles marshallii or Anopheles maculipalpis. The assays for An. funestus-like and An. rivulorum-like produced non-specific amplifications where no distinct base pair size recognition could be made that was diagnostic for either of these two species.
Table 2

Morphological species that amplified to approximate base pair (bp) sizes of species within the An. gambiae complex and An. funestus group

Species complex

Expected band size (bp)

Morphological species that showed approximate amplification

An. gambiae complex

 An. merus


An. rufipes; An. listeri

 An. gambiae


An. squamosus; An. rufipes; An. listeri; An. pretoriensis

 An. arabiensis


An. pretoriensis; An. rufipes

 An. quadriannulatus



An. funestus group

 An. vaneedeni


An. pretoriensis

 An. funestus


An. pretoriensis

 An. rivulorum


An. rufipes

 An. parensis



 An. leesoni


An. rufipes; An. rhodesiensis

Using the diagnostic An. funestus multiplex PCR assay, 8% (12/150) of the samples amplified. Seven of 52 Anopheles rufipes specimens had amplicon sizes corresponding to either Anopheles leesoni (n = 6) or An. rivulorum (n = 1), while the single An. rhodesiensis showed an amplicon size similar to An. leesoni. Only four out of 46 Anopheles pretoriensis specimens showed amplification during this assay—three produced a fragment size similar to An. funestus s.s. and one had a base pair size corresponding to that of Anopheles vaneedeni.

For the An. gambiae species-diagnostic PCR assay, 11.3% (17/150) amplified. Two Anopheles squamosus specimens showed amplicons similar to An. gambiae s.s. (Table 2). Seven out of 52 An. rufipes had amplifications similar to An. gambiae s.s. (n = 3), An. arabiensis (n = 3) and An. merus (n = 1). Five of the 12 An. listeri specimens showed fragments diagnostic for either An. gambiae s.s. (n = 3) or An. merus (n = 2). Four out of 46 An. pretoriensis samples amplified products similar in size to An. arabiensis (n = 1) and An. gambiae s.s. (n = 3). Some specimens also revealed non-specific amplification, most likely due to the fact that these protocols are not optimized for other anopheline species. Both amplification and non-amplification was present for specimens of the same species, irrespective of the amount of DNA that was present. Examples are shown in Fig. 1.
Fig. 1

Anopheles gambiae species-diagnostic PCR gel electrophoresis showing the amplification by other anopheline species. Lanes 1 and 26: 100 bp DNA Ladder; lanes 2–5: positive controls for An. arabiensis; An. gambiae; An. merus and An. quadriannulatus; lanes 6, 7: negative controls for DNA extraction and PCR master mix; lane 10: An. rufipes; lanes 11–21: An. listeri; lanes 22, 23: An. squamosus; lanes 24, 25 contained no specimens

Interesting geographical variation was noted for An. rufipes in particular. Specimens from two West African countries, Guinea (Siguiri) and Mali (Yanfolila), had amplicons from the An. funestus species-diagnostic PCR assay but none for the An. gambiae species-diagnostic PCR assay. However, for South African sites it was the other way around with no amplification during the An. funestus species-diagnostic PCR assay and only amplicons produced by the An. gambiae assay in specimens from the Kruger National Park and surrounding areas in Mpumalanga.


As with other techniques used to identify mosquitoes, there are drawbacks to morphological species identification, such as when specimens have lost important external features of their anatomy (e.g. legs), a common occurrence when using collection methods such as CDC light traps where mosquitoes are damaged as they are sucked through the fan blades. In addition, the level of relevant skills to carry out the identifications may be inadequate or lacking.

A recent study carried out in eastern Zambia comparing morphological identifications with two molecular assays (COI mtDNA and ITS2 rDNA) [22] provides interesting insights. For example, of the 18 molecular species/forms they identified, 16 contained specimens that were morphologically identified as the An. funestus group and 12 as the An. gambiae complex. The molecular “Anopheles coustani” was being identified morphologically as both An. funestus and An. gambiae groups [22]. This variation either demonstrates a very low level of morphological skills, or there is a lot of variation in the molecular sequencing of individual mosquitoes. This latter is not unlikely given the amount of variation that was found in An. gambiae through the recently published 1000 Genome study [23]. Furthermore, sequence variability in the ITS2 region has been demonstrated previously between specimens of An. rivulorum from different countries [24] and this needs further investigation on a broad spectrum of species.

The present study showed that five of the 11 species amplified three members of the An. gambiae complex and four members of the An. funestus group. 30.8% of the An. pretoriensis gave amplicons of four different species, while 25% of the An. rufipes produced fragments of five species. But not every individual specimen of each species gave the same results, indicating considerable molecular variation. Some specimens also revealed non-specific amplification, most likely due to the fact that these protocols are not optimized for other anopheline species, thus possibly creating confusion in the routine analysis of the results.

It is well-known that almost every morphological taxon studied so far is a species complex. The best known are the An. gambiae complex and the An. funestus group based on various genetic techniques [6, 9, 25]. Others include Anopheles coustani/crypticus (chromosomes [26]), Anopheles nili/ovengensis/carnevalei (morphology, molecular [27, 28]) and Anopheles marshallii/letabensis/hughi (chromosomes [29, 30]). Complexes that still require formal morphological descriptions include An. pharoensis (chromosomes, 2 species [31]), Anopheles longipalpis (molecular, 2 species [32]) and An. squamosus (chromosomes, 5 species [Green and Hunt, unpublished data]). It is, therefore, not surprising when molecular studies show possible new species [22, 33], but in order to understand their importance in malaria transmission, these molecular forms need to be linked to iso-female lines that can be used to provide information on genetic variation within families and appropriate morphological descriptions. Such iso-female lines could also be used to produce reference sequence data for use in the development of accurate molecular species-diagnostic assays.

Morphological and molecular identification techniques should complement each other in order to intensify vector surveillance and our understanding of mosquito biodiversity [22, 34]. Morphological identification with the use of dichotomous keys forms the basis for the development of molecular techniques [35] and their subsequent use in surveillance or research [18, 19, 36, 37]. The above results indicate how important it is to carry out a priori morphological identification before using the standard molecular assays for the two major vector groups and if unusual species composition is reported, to initiate further investigations. Unfortunately, the loss of taxonomic expertise over the years has had a severe negative impact on describing mosquito biodiversity [14].


This study showed that poor morphological identification cannot necessarily be detected and corrected during molecular PCR identification and can negatively affect vector surveillance since control interventions can likely be based on wrong identifications. Furthermore, processing mosquitoes for molecular identification is expensive and scarce resources should be limited to those specimens that require them. Malaria control programmes should continue to invest in capacity building for entomology teams and morphological training should be prioritized for entomological surveillance, particularly as more reports appear on secondary or incidental vectors playing a role in residual malaria transmission [1012, 37, 38].


Authors’ contributions

MC carried out the morphological identifications using the keys of Gillies and Coetzee [6]. EE carried out the laboratory analysis and drafted the manuscript. LLK and MC conceived the project, analysed the data and drafted the manuscript. All authors read and approved the final manuscript.


The following are thanked for collection of field material: RH Hunt, Mali; N Venter, Guinea; P Tshikae, Limpopo; the SIT team, Kwazulu/Natal; and F Mbokazi, Mpumalanga Province. Prof L Braack is thanked for An. rivulorum-like DNA used as controls. Profs B Brooke and RH Hunt are thanked for comments on the manuscript.

Competing interests

The authors declare that they have no competing interests.

Availability of data and materials

All data generated or analysed during this study are included in this published article.

Consent for publication

Not applicable.

Ethics approval and consent to participate

Not applicable.


This work was supported by the International Atomic Energy Agency, the Industrial Development Corporation and the South African Nuclear Energy Corporation (Necsa) through its Nuclear Technologies in Medicine and the Biosciences Initiative (NTeMBI)—a national platform funded by the Department of Science and Technology; the Department of Science and Technology (DST)/National Research Foundation (NRF) competitive programme for rated researchers Grants (CPRR) and Incentive Funding [(UID) 85538] to LLK; DST/NRF Research Chair Initiative grant to MC; the Global Diseases Detection/CDC Grant (U19GH000622-01 MAL01), IAEA (Research Contract No. 17904; 19099 and SAF 5013/5014); and the MRC Collaborating Centre for Multidisciplinary Research on Malaria. The Grant-holders acknowledge that opinions, findings and conclusions or recommendations expressed in any publication generated by the NRF supported research are that of the author(s), and that the NRF accepts no liability whatsoever in this regard.

Publisher’s Note

Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Open AccessThis article is distributed under the terms of the Creative Commons Attribution 4.0 International License (, which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made. The Creative Commons Public Domain Dedication waiver ( applies to the data made available in this article, unless otherwise stated.

Authors’ Affiliations

Wits Research Institute for Malaria, MRC Collaborating Centre for Multidisciplinary Research on Malaria, School of Pathology, Faculty of Health Sciences, University of the Witwatersrand, Johannesburg, South Africa
Centre for Emerging Zoonotic and Parasitic Diseases, National Institute for Communicable Diseases, Johannesburg, South Africa


  1. Bhatt S, Weiss DJ, Cameron E, Bisanzio D, Mappin B, Dalrymple U, et al. The effect of malaria control on Plasmodium falciparum in Africa between 2000 and 2015. Nature. 2015;526:207–11.View ArticlePubMedPubMed CentralGoogle Scholar
  2. Sinka ME, Bangs MJ, Manguin S, Coetzee M, Mbogo CM, Hemingway J, et al. The dominant Anopheles vectors of human malaria in Africa, Europe and the Middle East: occurrence data, distribution maps and bionomic précis. Parasites Vectors. 2010;3:117.View ArticlePubMedPubMed CentralGoogle Scholar
  3. Wiebe A, Longbottom J, Gleave K, Shearer FM, Sinka ME, Massey NC, et al. Geographical distributions of African malaria vector sibling species and evidence for insecticide resistance. Malar J. 2017;16:85.View ArticlePubMedPubMed CentralGoogle Scholar
  4. 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. ArticlePubMedPubMed CentralGoogle Scholar
  5. Peterson A. Shifting suitability for malaria vectors across Africa with warming climates. BMC Infect Dis. 2009;9:59.View ArticlePubMedPubMed CentralGoogle Scholar
  6. Gillies MT, Coetzee M. Supplement to the Anophelinae of Africa south of the Sahara, vol. 55. Johannesburg: Publications of the South African Institute for Medical Research; 1987. p. 1–143.Google Scholar
  7. 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; 1987. p. 1–343.Google Scholar
  8. Coluzzi M. Heterogeneities of the malaria vectorial system in tropical Africa and their significance in malaria epidemiology and control. Bull World Health Org. 1984;62:107–13.PubMedPubMed CentralGoogle Scholar
  9. Coetzee M, Hunt RH, Wilkerson R, Della Torre A, Coulibaly MB, Besansky NJ. Anopheles coluzzii and Anopheles amharicus, new members of the Anopheles gambiae complex. Zootaxa. 2013;3619:246–74.View ArticlePubMedGoogle Scholar
  10. Antonio-Nkondjio C, Kerah CH, Simard F, Awono-Ambene P, Chouaibou M, Tchuinkam T, et al. Complexity of the malaria vectorial system in Cameroon: contribution of secondary vectors to malaria transmission. J Med Entomol. 2006;43:1215–21.View ArticlePubMedGoogle Scholar
  11. Nepomichene TNJJ, Tata E, Boyer S. Malaria case in Madagascar, probable implication of a new vector, Anopheles coustani. Malar J. 2015;14:475.View ArticlePubMedPubMed CentralGoogle Scholar
  12. Tabue RN, Awono-Ambene P, Etang J, Atangana J, Antonio-Nkondjio C, Toto JC, et al. Role of Anopheles (Cellia) rufipes (Gough, 1910) and other local anophelines in human malaria transmission in the northern savannah of Cameroon: a cross-sectional survey. Parasites Vectors. 2017;10:22.View ArticlePubMedPubMed CentralGoogle Scholar
  13. Garros C, Dujardin JP. Genetic and phenetic approaches to Anopheles systematics. In: Manguin S, editor. Anopheles mosquitoes: new insights into malaria vectors. Rijeka: InTech; 2013. p. 81–105.Google Scholar
  14. Harbach RE. The classification of genus Anopheles (Diptera: Culicidae): a working hypothesis of phylogenetic relationships. Bull Entomol Res. 2004;94:537–53.View ArticlePubMedGoogle Scholar
  15. Davidson G. The five mating-types in the Anopheles gambiae complex. Riv Malar. 1964;43:167–83.PubMedGoogle Scholar
  16. Coluzzi M. Morphological divergences in the Anopheles gambiae complex. Riv Malar. 1964;43:197–232.PubMedGoogle Scholar
  17. Mahon RJ, Green CA, Hunt RH. Diagnostic allozymes for routine identification of adults of the Anopheles gambiae complex (Diptera: Culicidae). Bull Entomol Res. 1976;66:25–31.View ArticleGoogle Scholar
  18. 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.View ArticlePubMedGoogle Scholar
  19. 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.View ArticlePubMedGoogle Scholar
  20. Choi KS, Coetzee M, Koekemoer LL. Simultaneous identification of the Anopheles funestus group and Anopheles longipalpis type C by PCR-RFLP. Malar J. 2010;9:316.View ArticlePubMedPubMed CentralGoogle Scholar
  21. 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.PubMedGoogle Scholar
  22. 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.View ArticlePubMedPubMed CentralGoogle Scholar
  23. The Anopheles gambiae 1000 Genomes Consortium. Genetic diversity of the African malaria vector Anopheles gambiae. Nature. 2017;552:96–100.Google Scholar
  24. Hackett BJ, Gimnig J, Guelbeogo W, Costantini C, Koekemoer LL, Coetzee M, et al. Ribosomal DNA internal transcribed spacer (ITS2) sequences differentiate Anopheles funestus and An. rivulorum, and uncover a cryptic taxon. Insect Mol Biol. 2000;9:369–74.View ArticlePubMedGoogle Scholar
  25. Costantini C, Sagnon NF, Ilboudo-Sanogo E, Coluzzi M, Boccolini D. Chromosomal and bionomic heterogeneities suggest incipient speciation in Anopheles funestus from Burkina Faso. Parassitologia. 1999;41:595–611.PubMedGoogle Scholar
  26. Coetzee M. Anopheles crypticus, new species from South Africa is distinguished from Anopheles coustani (Diptera: Culicidae). Mosq Syst. 1994;26:125–31.Google Scholar
  27. Brunhes J, Le Goff G, Geoffroy B. Afro-tropical anopheline mosquitoes. III. Description of three new species: Anopheles carnevalei sp.nov., An. hervyi sp.nov., and An. dualaensis sp.nov., and resurrection of An. rageaui Mattingly and Adam. J Am Mosq Control Assoc. 1999;15:552–8.PubMedGoogle Scholar
  28. Awono-Ambene P, Kengne P, Simard F, Antonio-Nkondjio C, Fontenille D. Description and bionomics of Anopheles (Cellia) ovengensis (Diptera: Culicidae), a new malaria vector species of the Anopheles nili group from south Cameroon. J Med Entomol. 2004;41:561–8.View ArticlePubMedGoogle Scholar
  29. Lambert DM. Anopheles marshallii (Theobald) is a complex of species. Mosq Syst. 1979;11:173–8.Google Scholar
  30. Lambert DM, Coetzee M. A dual genetic and taxonomic approach to the resolution of the mosquito taxon Anopheles (Cellia) marshallii (Culicidae). Syst Entomol. 1982;7:321–32.View ArticleGoogle Scholar
  31. Miles SJ, Green CA, Hunt RH. Genetic observations on the taxon Anopheles (Cellia) pharoensis Theobald (Diptera: Culicidae). J Trop Med Hyg. 1983;86:153–7.PubMedGoogle Scholar
  32. Koekemoer LL, Misiani EA, Hunt RH, Kent RJ, Norris DE, Coetzee M. Cryptic species within Anopheles longipalpis from southern Africa and phylogenetic comparison with members of the An. funestus group. Bull Entomol Res. 2009;99:41–9.View ArticlePubMedGoogle Scholar
  33. Stevenson J, St Laurent B, Lobo NF, Cooke MK, Kahindi SC, Oriango RM, et al. Novel vectors of malaria parasites in the western highlands of Kenya. Emerg Infect Dis. 2012;18:1547–9.View ArticlePubMedPubMed CentralGoogle Scholar
  34. Chan A, Chiang L, Hapuarachchi HC, Tan CH, Pang SC, Lee R, et al. DNA barcoding: complementing morphological identification of mosquito species in Singapore. Parasites Vectors. 2014;7:569.View ArticlePubMedPubMed CentralGoogle Scholar
  35. Kumar NP, Rajavel AR, Natarajan R, Jambulingam P. DNA barcodes can distinguish species of Indian mosquitoes (Diptera: Culicidae). J Med Entomol. 2007;44:1–7.View ArticlePubMedGoogle Scholar
  36. Collins FH, Paskewitz SM. A review of the use of ribosomal DNA (rDNA) to differentiate among cryptic Anopheles species. Insect Mol Biol. 1996;5:1–9.View ArticlePubMedGoogle Scholar
  37. Burke A, Dandalo L, Munhenga G, Dahan Y, Mbokazi F, Coetzee M, et al. A new malaria vector mosquito in South Africa. Sci Rep. 2017;7:43779.View ArticlePubMedPubMed CentralGoogle Scholar
  38. Stevenson JC, Simubali L, Mbambara S, Musonda M, Mweetwa S, Mudenda T, et al. Detection of Plasmodium falciparum infection in Anopheles squamosus (Diptera: Culicidae) in an area targeted for malaria elimination, Southern Zambia. J Med Entomol. 2016;53:1482–7.View ArticlePubMedPubMed CentralGoogle Scholar


© The Author(s) 2018


By submitting a comment you agree to abide by our Terms and Community Guidelines. If you find something abusive or that does not comply with our terms or guidelines please flag it as inappropriate.