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    Figure 1.

    Sequence alignment of the internal transcribed spacer region 2 of the unidentifiable Malawian (MalaF) samples and Anopheles funestus. Blocks highlight sequence variation. – indicates deletions.

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    Figure 2.

    Primer MalaFB included in the polymerase chain reaction (PCR) cocktail mixture for Anopheles funestus group identifications. Lane 1, 100-basepair DNA ladder; lanes 2–4, positive controls: An. vaneedeni, An. funestus, and An. rivulorum, respectively; lane 5, MalaF sample; lanes 6 and 7, positive controls: An. parensis and An. leesoni, respectively; lane 8, PCR negative control.

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    Figure 3.

    Sequence alignment of the partial D3 region of the unidentifiable Malawian (MalaF) samples (133 basepairs) and Anopheles funestus (135 basepairs). Blocks highlight sequence variation. – indicates deletions. The forward D3A primer binds approximately 90 basepairs upstream from the start of this sequence.

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    Figure 4.

    Asynapsis in hybrid polytene chromosomes indicated by arrows.

  • 1

    Gillies MT, de Meillon B, 1968. The Anophelinae of Africa South of the Sahara. Publications of the South African Institute for Medical Research, no. 54. Johannesburg, South Africa.

  • 2

    Gillies MT, Coetzee M, 1987. A Supplement to the Anophelinae of Africa South of the Sahara (Afrotropical Region). Publications of the South African Institute for Medical Research, no 55. Johannesburg, South Africa.

  • 3

    Harbach RE, 2004. The classification of genus Anopheles (Diptera: Culicidae): a working hypothesis of phylogenetic relationships. Bull Entomol Res 94 :537–553.

    • Search Google Scholar
    • Export Citation
  • 4

    Garros C, Harbach RE, Manguin S, 2005. Morphological assessment and molecular phylogenetics of the Funestus and Minimus Groups of Anopheles (Cellia). J Med Entomol 42 :522–536.

    • Search Google Scholar
    • Export Citation
  • 5

    Hackett BJ, Gimnig J, Guelbeogo W, Costantini C, Koekemoer LL, Coetzee M, Collins FH, Besansky NJ, 2000. Ribosomal DNA internal transcribed spacer (ITS2) sequences differentiate Anopheles funestus and An. rivulorum, and uncover a cryptic taxon. Insect Mol Biol 9 :369–374.

    • Search Google Scholar
    • Export Citation
  • 6

    Couhet A, Simard F, Toto J-C, Kengne P, Coetzee M, Fontenille D, 2003. Species identification within the Anopheles funestus group of malaria vectors in Cameroon and evidence for a new species. Am J Trop Med Hyg 69 :200–205.

    • Search Google Scholar
    • Export Citation
  • 7

    Green CA, 1982. Cladistic analysis of chromosome data (Anopheles (Cellia) Myzomyia). J Hered 73 :2–11.

  • 8

    Wilkes TJ, Matola YG, Charlwood JD, 1996. Anopheles rivulorum, a vector of human malaria in Africa. Med Vet Entomol 10 :108–110.

  • 9

    De Meillon B, Van Eeden GJ, Coetzee L, Coetzee M, Meiswinkel R, Du Toit CL, Hansford CF, 1977. Observations on a species of the Anopheles funestus subgroup, a sustpected exophilic vector of malaria parasites in North Eastern Transvaal, South Africa. Mosq News 37 :657–661.

    • Search Google Scholar
    • Export Citation
  • 10

    Evans AM, Symes CB, 1937. Anopheles funestus and its allies in Kenya. Ann Trop Med Parasitol 31 :105–111.

  • 11

    Evans AM, 1938. Mosquitoes of the Ethiopian Region. London: Bernard Quartich Ltd., Dulau and Co. Ltd., and The Oxford University Press.

  • 12

    De Meillon B, 1947. New records and species of biting insects from the Ethiopian Region II. J Entomol Soc S Afr 10 :110–124.

  • 13

    Green CA, Hunt RH, 1980. Interpretations of variation in ovarian polytene chromosomes of Anopheles funestus Giles, A. parensis Gillies and A. aruni? Genetica 51 :187–195.

    • Search Google Scholar
    • Export Citation
  • 14

    Lochouarn L, Dia I, Boccolini D, Coluzzi M, Fontenille D, 1998. Bionomical and cytogenetic heterogeneities of Anopheles funestus in Senegal. Trans R Soc Trop Med Hyg 92 :607–612.

    • Search Google Scholar
    • Export Citation
  • 15

    Costantini C, Sagnon, N’F, Ilboudo-Sanogo E, Coluzzi M, Boccolini D, 1999. Chromosomal and bionomic heterogeneities suggest incipient speciation in Anopheles funestus from Burkina Faso. Parassitologia 41 :595–611.

    • Search Google Scholar
    • Export Citation
  • 16

    Koekemoer LL, Kamau L, Hunt RH, Coetzee M, 2002. A cocktail polymerase chain reaction assay to identify members of the Anopheles funestus (Diptera: Culicidae) group. Am J Trop Med Hyg 6 :804–811.

    • Search Google Scholar
    • Export Citation
  • 17

    Collins FH, Mendez MA, Rasmussen MO, Meheffey PC, Besansky NJ, Finnerty V, 1987. A ribosomal RNA gene probes differentiates members of the Anopheles gambiae complex. Am J Trop Med Hyg 37 :37–41.

    • Search Google Scholar
    • Export Citation
  • 18

    Litvaitis MK, Nunn G, Thomas WK, Kocher TD, 1994. A molecular approach for the identification of Meiofaunal turbellarians (Platyhelminthes: Turbellaria). Mar Biol 158 :17–35.

    • Search Google Scholar
    • Export Citation
  • 19

    Hunt RH, 1973. A cytological technique for the study of Anopheles gambiae complex. Parassitologia 15 :137–139.

  • 20

    Davidson G, Paterson H, Coluzzi M, Mason G, Micks D, 1967. The Anopheles gambiae complex. Wright J, Pal R, eds. Genetics of Insect Vectors of Disease. London: Elsevier Publishing Company, 211–250.

  • 21

    Wirtz RA, Zavala F, Charoenvit Y, Cambell GH, Burkot TR, Schneider I, Esser KM, Beaudoin RL, Andre GR, 1987. Comparative testing of Plasmodium falciparum sporozoite monoclonal antibodies for ELISA development. Bull World Health Organ 65 :39–45.

    • Search Google Scholar
    • Export Citation
  • 22

    Singh OP, Chandra D, Nanda N, Raghavendra K, Sunil S, Sharma SK, Dua VK, Subbarao SK, 2004. Differentiation of members of the Anopheles fluviatilis species complex by an allele-specific polymerase chain reaction based on 28S ribosomal DNA sequences. Am J Trop Med Hyg 70 :27–32.

    • Search Google Scholar
    • Export Citation
  • 23

    Sharpe RG, Harbach RE, Butlin RK, 2000. Molecular variation and phylogeny of the members of the Minimus group of Anopheles subgenus Cellia (Diptera: Culicidae). Syst Entomol 25 :263–272.

    • Search Google Scholar
    • Export Citation
  • 24

    Paskewitz SM, Wesson DM, Collins FH, 1993. The internal transcribed spacers of ribosomal DNA in five members of the Anopheles gambiae species complex. Insect Mol Biol 2 :247–257.

    • Search Google Scholar
    • Export Citation
  • 25

    Frizzi G, 1947. Salivary gland chromosomes of Anopheles. Nature 160 :226–227.

  • 26

    Frizzi G, 1953. Étude cytog’enetique d’anopheles maculipennis en Italie. Extension des recherches ‘a d’autres esp’eces d’anoph’eles. Bull World Health Organ 9 :335–344.

    • Search Google Scholar
    • Export Citation
  • 27

    Coluzzi M, Sabatini A, 1967. Cytogenetic observations on species A and B of the Anopheles gambiae complex. Parassitologia 9 :73–88.

  • 28

    Coluzzi M, Sabatini A, 1968. Cytogenetic observations on species C of the Anopheles gambiae complex. Parassitologia 10 :155–166.

  • 29

    Coluzzi M, Sabatini A, 1969. Cytogenetic observations on the salt water species Anopheles merus and Anopheles melas, of the Gambiae complex. Parassitologia 11 :155–166.

    • Search Google Scholar
    • Export Citation
  • 30

    Thongwat D, Morgan K, O’Loughlin SM, Walton C, Choochote W, Somboon P, 2008. Crossing experiments supporting the specific status of Anopheles maculatus chromosomal form K. J Am Mosq Control Assoc 24 :194–202.

    • Search Google Scholar
    • Export Citation
  • 31

    Coetzee M, Estrada-Franco JG, Wunderlich CA, Hunt RH, 1999. Cytogenetic evidence for a species complex within Anopheles pseudopunctipennis Theobald (Diptera: Culicidae). Am J Trop Med Hyg 60 :649–653.

    • Search Google Scholar
    • Export Citation
  • 32

    Somboon P, Thongwat D, Choochote W, Walton C, Takagi M, 2005. Crossing experiments of Anopheles minimus species A and putative species E. J Am Mosq Control Assoc 21 :5–9.

    • Search Google Scholar
    • Export Citation
  • 33

    Kaiser PE, 1988. Cytotaxonomy as a tool for identification of siblings of the Anopheles quadrimaculatus complex. Fla Entomol 71 :311–323.

    • Search Google Scholar
    • Export Citation
  • 34

    Foley DH, Bryan JH, 1991. Anopheles annulipes Walker (Diptera: Culicidae) at Griffith New South Wales. 1. Two sibling species in sympatry. Aust J Entomol 30 :109–112.

    • Search Google Scholar
    • Export Citation
  • 35

    Hunt RH, Coetzee M, Fettene M, 1998. The Anopheles gambiae complex: a new species from Ethiopia. Trans R Soc Trop Med Hyg 92 :231–235.

    • Search Google Scholar
    • Export Citation
  • 36

    Garros C, Koekemoer LL, Kamau L, Awolola TS, Van Bortel W, Coetzee M, Coosemans M, Manguin S, 2004. Restriction fragment length polymorphism method for the identification of major African and Asian malaria vectors within the Anopheles funestus and An. minimus groups. Am J Trop Med Hyg 70 :260–265.

    • Search Google Scholar
    • Export Citation
  • 37

    Michel AP, Ingracsi MJ, Schemerhorn BJ, Kern M, Le Goff G, Coetzee M, Elissa N, Fontenille D, Vulule J, Lehmann T, Sagnon N’F, Costantini C, Besansky NJ, 2005. Rangewide population genetics structure of the African malaria vector Anopheles funestus. Mol Ecol 14 :4235–4248.

    • Search Google Scholar
    • Export Citation
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A New Species Concealed by Anopheles funestus Giles, a Major Malaria Vector in Africa

Belinda L. SpillingsVector Control Reference Unit, National Institute for Communicable Diseases, National Health Laboratory Service, Johannesburg, South Africa; Division of Virology and Communicable Disease Surveillance, School of Pathology of the University of the Witwatersrand and the National Health Laboratory Service, Johannesburg, South Africa; School of Animal, Plant and Environmental Sciences, University of the Witwatersrand, Johannesburg, South Africa; National Research Foundation Chair in Medical Entomology and Vector Control, School of Pathology, University of the Witwatersrand, Johannesburg, South Africa; National Malaria Control Programme, Lilongwe, Malawi

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Basil D. BrookeVector Control Reference Unit, National Institute for Communicable Diseases, National Health Laboratory Service, Johannesburg, South Africa; Division of Virology and Communicable Disease Surveillance, School of Pathology of the University of the Witwatersrand and the National Health Laboratory Service, Johannesburg, South Africa; School of Animal, Plant and Environmental Sciences, University of the Witwatersrand, Johannesburg, South Africa; National Research Foundation Chair in Medical Entomology and Vector Control, School of Pathology, University of the Witwatersrand, Johannesburg, South Africa; National Malaria Control Programme, Lilongwe, Malawi

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Lizette L. KoekemoerVector Control Reference Unit, National Institute for Communicable Diseases, National Health Laboratory Service, Johannesburg, South Africa; Division of Virology and Communicable Disease Surveillance, School of Pathology of the University of the Witwatersrand and the National Health Laboratory Service, Johannesburg, South Africa; School of Animal, Plant and Environmental Sciences, University of the Witwatersrand, Johannesburg, South Africa; National Research Foundation Chair in Medical Entomology and Vector Control, School of Pathology, University of the Witwatersrand, Johannesburg, South Africa; National Malaria Control Programme, Lilongwe, Malawi

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John ChiphwanyaVector Control Reference Unit, National Institute for Communicable Diseases, National Health Laboratory Service, Johannesburg, South Africa; Division of Virology and Communicable Disease Surveillance, School of Pathology of the University of the Witwatersrand and the National Health Laboratory Service, Johannesburg, South Africa; School of Animal, Plant and Environmental Sciences, University of the Witwatersrand, Johannesburg, South Africa; National Research Foundation Chair in Medical Entomology and Vector Control, School of Pathology, University of the Witwatersrand, Johannesburg, South Africa; National Malaria Control Programme, Lilongwe, Malawi

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Maureen CoetzeeVector Control Reference Unit, National Institute for Communicable Diseases, National Health Laboratory Service, Johannesburg, South Africa; Division of Virology and Communicable Disease Surveillance, School of Pathology of the University of the Witwatersrand and the National Health Laboratory Service, Johannesburg, South Africa; School of Animal, Plant and Environmental Sciences, University of the Witwatersrand, Johannesburg, South Africa; National Research Foundation Chair in Medical Entomology and Vector Control, School of Pathology, University of the Witwatersrand, Johannesburg, South Africa; National Malaria Control Programme, Lilongwe, Malawi

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Richard H. HuntVector Control Reference Unit, National Institute for Communicable Diseases, National Health Laboratory Service, Johannesburg, South Africa; Division of Virology and Communicable Disease Surveillance, School of Pathology of the University of the Witwatersrand and the National Health Laboratory Service, Johannesburg, South Africa; School of Animal, Plant and Environmental Sciences, University of the Witwatersrand, Johannesburg, South Africa; National Research Foundation Chair in Medical Entomology and Vector Control, School of Pathology, University of the Witwatersrand, Johannesburg, South Africa; National Malaria Control Programme, Lilongwe, Malawi

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The major malaria vector Anopheles funestus belongs to a group of morphologically similar species that are commonly distinguished from one another through the use of chromosomal and molecular techniques. Indoor resting collections of mosquitoes from Malawi were initially identified as An. funestus by morphology, but failed to have this confirmed by the species-specific polymerase chain reaction assay. Sequence analysis of the internal transcribed spacer region 2 identified variations within the An. funestus-specific primer binding site and showed a sequence variation of 4.5% compared with An. funestus. Domain 3 analysis showed sequence variation of 1.5% from An. funestus. Cytogenetic analysis of the polytene chromosome banding arrangements showed that the specimens were homosequential with An. funestus, with fixed inverted arrangements of the 3a, 3b, and 5a inversions commonly polymorphic in An. funestus. The chromosomes of hybrid females showed levels of asynapsis typical of inter-species crosses. These molecular and cytogenetic observations support the conclusion that this Malawi population is a new species and it has provisionally been named An. funestus-like.

INTRODUCTION

The Anopheles funestus Giles group consists of nine African species,13 of which five belong to the Funestus subgroup on the basis of phylogenetic analyses: An. funestus, An. vaneedeni Gillies and Coetzee, An. parensis Gillies, An. aruni Sobti, and An. confusus Evans and Leeson. 3,4 The remaining members of the group belong to the Rivulorum subgroup: An. brucei Service, An. fuscivenosus Leeson, An. rivulorum Leeson, and the An. rivulorum-like species from west Africa. 3,57 All the species are morphologically similar at the adult stage and species identification at the egg and larval stages is possible only for An. confusus.

Although the members of the An. funestus group may be similar in morphology, their efficiencies as malaria vectors vary greatly. Because of its highly anthropophilic and endophilic nature, An. funestus s.s. is one of the primary vectors of malaria in sub-Saharan Africa.1 Anopheles rivulorum has only once been implicated in malaria transmission in Tanzania8 but generally elects to blood feed on domestic animals rather than humans. The remaining members of the An. funestus group have never been shown to be malaria vectors in nature, 1,2 although An. vaneedeni was infected experimentally in the laboratory.9

Because of the different vectorial capacities, biting and resting behaviors and the close morphologic similarity of members of the An. funestus group, accurate identification of field-caught material is critical for vector control programs. Early identification methods relied solely on morphologic characters that detailed minor differences between the members of the group. 1,2,1012 This process of identification relies on the availability of multiple life stages and requires a high level of expertise because the characteristics used are frequently overlapping and only two of the species can be unequivocally identified using morphology.

The success of cytogenetics to elucidate the members of the An. gambiae complex led to similar studies on the An. funestus group. Analysis of the banding arrangements of giant polytene chromosomes distinguished An. parensis from An. funestus.13 However, An. vaneedeni (formerly An. aruni?) had homosequential chromosomal banding patterns with An. funestus.13 Cross-mating studies between An. vaneedeni and An. funestus produced sterile male hybrids and asynapsis of the hybrid polytene chromosomes, 13 thereby confirming the specific status of An. vaneedeni. More recent cytogenetic studies in west Africa have shown clear evidence of genetic differentiation in sympatric populations of An. funestus, indicating that this taxon may consist of a complex of cryptic species. 14,15 The need for simpler identification methods resulted in the development of a multiplex polymerase chain reaction (PCR) assay 16 targeting the variable internal transcribed spacer region 2 (ITS2) regions for the identification of the four most common species of the group: An. funestus, An. vaneedeni, An. parensis, and An. rivulorum.

In this paper, we report evidence of a new species of Anopheles mosquito from Malawi, which occurs in sympatry with An. funestus and shows morphologic overlap with this species and cannot be identified using the standard multiplex PCR assay.

MATERIALS AND METHODS

Study site and collection method.

Anopheline collections were carried out in December 2007 in rural villages around Karonga in northern Malawi (10°18.627′S, 34°07.901′E), close to the shoreline of Lake Malawi. The collections were predominantly indoor resting catches with the exception of samples that were collected from tyres stacked at the entrance to a reed hut. The adult specimens were identified as members of the An. funestus group, according to morphologic keys. 1,2 Male specimens were dry-preserved on silica and the females were transported alive to the insectaries of the Vector Control Reference Unit, National Institute for Communicable Diseases (Johannesburg, South Africa).

Laboratory rearing of wild-caught material.

A total of 63 An. funestus females were placed in glass vials for egg laying. They were maintained on 10% sucrose and offered a blood meal every alternate day. After egg laying, females that survived were offered a blood meal and kept until half gravid for chromosomal studies. The morphology of the egg batches confirmed that the samples belonged to the An. funestus subgroup.1 Each egg batch was treated as an individual family and the resulting F1 progeny were reared to adulthood and used for molecular identifications and cross-mating experiments.

Molecular identification of field material.

DNA was extracted 17 from an individual adult from each family. The multiplex PCR for the identification of An. funestus, An. rivulorum, An. vaneedeni, An. parensis, and An. leesoni16 was performed with all DNA samples. Each sample was tested 2–3 times to ensure accuracy of identification. A negative control containing no DNA and positive controls of An. funestus from laboratory colony material and An. leesoni from a previous wild collection were included in each PCR.

Sequencing and primer design for unidentified Malawi samples.

A segment of the ITS2 region of four unidentified Malawian (MalaF) samples and an An. funestus s.s. control were amplified using primers ITS2A (5′-TGT GAA CTG CAG GAC ACA T- 3′) and ITS2B (5′-TAT GCT TAA ATT CAG GGG GT-3′). 16 The 25-μL PCR mixture contained 50 pmol of each primer, 1.5 mM MgCl2, 200 μM of each dNTP, 1.25 units of Taq DNA polymerase, and 1 μL of DNA.5 The cycling conditions were initial denaturation at 94°C for 2 minutes; 40 cycles of denaturation at 94°C for 30 seconds, annealing at 50°C for 30 seconds, extension at 72°C for 40 seconds; and a final extension at 72°C for 10 minutes. The resulting amplicons were subjected to electrophoresis on a 1.8% low-melting temperature Tris-acetate-EDTA (TAE) agarose gel stained with ethidium bromide. Amplicons (approximately 850 basepairs) were excised and cleaned using the Qiaquick Gel Extraction kit (catalog no. 28704; Qiagen, Valencia, CA) and sequenced with the ITS2A and ITS2B primers. Direct sequencing of the PCR products was performed by Inqaba Biotechnical Industries (Pretoria, South Africa). Sequencing was carried out for both strands using the above primers. Sequence alignment and analysis was carried out using (Lasergene version 6; DNAStar; Madison, WI). A consensus sequence was created for the four MalaF samples and aligned to the An. funestus s.s. control sequence. Primer annealing sites specific to the MalaF samples were identified and new primers were designed to yield an amplicon with a different size to the amplicon yielded for An. funestus s.s. in the species-specific PCR. 16

Domain 3 (D3) of the 28s rDNA gene was amplified for the same MalaF samples and An. funestus s.s. control used for the ITS2 sequencing. The primers D3A (5′-GAC CCG TCT TGA AAC ACG GA-3′) and D3B (5′-TCG GAA GGA ACC AGC TAC TA-3′) were used for amplification. 18 Each reaction was carried out in a volume of 25 μL that contained 25 pmol of each primer, 1.5 mM MgCl2, 200 μM of each dNTP, 2 units of Taq DNA polymerase, and 1 μL of DNA. The cycling conditions were initial denaturation at 94°C for 3 minutes; 30 cycles of denaturation at 94°C for 30 seconds, annealing at 63°C for 40 seconds, extension at 72°C for 40 seconds; and a final extension at 72°C for 10 minutes. The resulting amplicons were subjected to electrophoresis and sequenced as for the ITS2 amplicons. Sequencing was carried out for both strands using the D3A and D3B primers. Sequence alignment and analysis was carried out using Lasergene version 6. A consensus sequence was created for the four MalaF samples and this was then aligned to the An. funestus s.s. control sequence.

Application of the MalaF-specific PCR primer.

Two potential primers (MalaFA: 5′-CCT GCG TCC CAA GGT T-3′; MalaFB: 5′-GTT TTC AAT TGA ATT CAC CAT T-3′) were individually tested for their efficiency in the species-specific PCR 16 for the identification of An. funestus group members. Each of the newly designed primers was included in the reaction mixture and the products were subjected to electrophoresis on a 3% TBE agarose gel stained with ethidium bromide. All 61 of the unidentifiable An. funestus-like samples were tested with the MalaFB primer. Any samples that failed to amplify with the new MalaFB primer were confirmed for the presence of nucleic acids by using a nanodrop spectrophotometer (NanoDrop Technologies Inc., Wilmington, DE). To confirm the presence of DNA in these samples, they were subjected to a PCR assay designed to detect, but not distinguish, members of the Funestus subgroup. The primers used were: UF (5′-TGT GAA CTG CAG GAC ACA T-3′) and LRev (5′-CCA AGC ACG TTG ATC CAG TAT TAC-3′). Each 25-μL PCR reaction mixture contained 6.6 pmol of each primer, 1.5 mM MgCl2, 200 μM of each dNTP, 1 unit of Taq DNA polymerase, and 1 μL of DNA. The cycling conditions were initial denaturation at 94°C for 2 minutes; 35 cycles of denaturation at 94°C for 30 seconds, annealing at 45°C for 30 seconds, extension at 72°C for 30 seconds; and a final extension at 72°C for 10 minutes. The resulting amplicons were subjected to electrophoresis on a 2.5% TAE gel stained with ethidium bromide. The presence of a product (approximately 440 basepairs) confirmed the presence of DNA for these samples.

Cytogenetics.

Half-gravid wild females and F1 progeny from the MalaF × An. funestus s.s. (Fumoz) crosses were dissected and their ovaries prepared for cytogenetic analysis. 13,19 The banding patterns of the polytene chromosomes were compared by chromosomal analyses with other members of the An. funestus group.7,13 Slides were viewed and photographed under phase contrast microscopy. The arm nomenclature follows that of Green,7 Green and Hunt, 13 Lochouarn and others, 14 and Costantini and others. 15

Cross-mating studies.

Virgin F1 females from the MalaF families were cross-mated with males from a laboratory colony of An. funestus s.s. (Fumoz). Their hybrid F1 female progeny were given a blood meal and ovaries were dissected at the half gravid stage. Polytene chromosomes were prepared as above and checked for asynapsis. 20

The testes of eight hybrid males from the cross-mating were dissected and viewed by phase contrast microscopy. The gross morphology of each pair of testes was examined, after which the testes were squashed to release the spermatozoa. The morphology of the spermatozoa was examined for signs of infertility. The reciprocal cross (MalaF males × Fumoz females) was also attempted.

Enzyme-linked immunosorbent assay for sporozoite detection.

The heads and thoraces of the entire group (n = 63) of wild-caught MalaF females were tested for the presence of Plasmodium falciparum circumsporozoite protein. The sandwich enzyme-linked immunosorbent assay (ELISA) technique 21 was used. The ELISA plates were read and analyzed using a Multiskan Ascent plate reader (Thermo Electron Corporation, Shanghai, China).

RESULTS

Species identification of field material.

Of the 63 wild females that were brought back to the laboratory for egg laying, only two specimens could be positively identified using the multiplex PCR assay 16 and these were both An. rivulorum, confirmed by morphology. The remaining 61 samples repeatedly failed to amplify any PCR product using the species-specific primers.

Sequence analysis, primer design, and application.

The ITS2 region of four of the unidentifiable MalaF samples (735 basepairs) and an An. funestus control (736 basepairs) were sequenced and aligned (Figure 1). The sequence data for the unidentifiable MalaF samples showed a three-basepair deletion and a T to C transition within the An. funestus specific primer binding site. The MalaF consensus sequence (GenBank accession no. FJ438963) showed a 4.5% (33 of 740 basepairs) difference in sequence to the An. funestus control, including insertions and deletions.

Primers were designed to anneal to the two most variable regions in the MalaF consensus sequence (Figure 1). Both of the MalaF specific primers were tested on MalaF samples prior to being combined into the species identification PCR cocktail. 16 The MalaFA primer was inconsistent and did not always result in an amplicon although the same template sample was used. The MalaFB primer consistently gave good amplicon yield on the same, as well as different, MalaF template samples. The MalaFB primer, when combined in the species cocktail PCR, resulted in good amplicon yield with a product of 390 basepairs (Figure 2). The MalaFB primer has been tested on the 61 An. funestus-like samples and amplicons were obtained for 54 (88.5%) of the samples. The remaining seven An. funestus–like samples failed to amplify. The presence of DNA in these samples was confirmed using a Funestus subgroup–specific PCR and spectrophotometry.

The D3 sequence data for the MalaF samples showed a five-basepair (GenBank accession no. FJ843022) change from the An. funestus control sequence (Figure 3). This change includes a two-basepair deletion, two base transversions and a single base transition, which translates into 1.5% (5 of 330 basepairs) sequence variation from the An. funestus control.

Cytogenetics.

Chromosomes from the wild MalaF females displayed homosequential banding arrangements with An. funestus. Autosomal inversions 3a, 3b, and 5a that are common as polymorphisms in An. funestus were each fixed in their inverted (theoretically derived) arrangements in the MalaF sample. A single, rare, polymorphic inversion was seen on autosome arm 2 from one female.

Cross-mating studies.

Cross-mating between F1 MalaF females and laboratory-reared An. funestus males was successful and resulted in hybrid progeny. The polytene chromosomes obtained from the hybrid females displayed asynapsis between homologous chromosomes (Figure 4). Examination of the testes of the hybrid males by microscopy showed fully developed testes that appeared normal in gross morphology. Although the head region of the spermatozoa appeared to be slightly narrower than expected, we could not carry out back-crosses to laboratory An. funestus to determine their viability and fertility because of insufficient material.

The reciprocal crosses of MalaF males to laboratory reared An. funestus females resulted in numerous egg batches (> 900 eggs). Only two of these eggs hatched, yielding a hatch rate of less than 0.2%. The adults that emerged were both females and died prematurely before taking a blood-meal. Thus, no further studies could be carried out.

ELISA for sporozoite detection.

All of the wild-caught females that were brought back to the laboratory for rearing were tested for the presence of P. falciparum and were negative.

DISCUSSION

The Malawian malaria vectors are predominantly An. gambiae, An. arabiensis, and An. funestus (Chiphwanya J, unpublished data). A previous collection of An. funestus carried out in Karonga in August 2007 resulted in 80% failure to identify the specimens using the multiplex PCR assay. 16 The identification of the specimens collected for this study again failed to give amplicons for 61 of 63 specimens collected although these samples were identified morphologically as the An. funestus group. 1,2 The failure of primer annealing in the PCR assay is similar to observations for An. rivulorum-like in Cameroon.6

Sequencing of the ITS2 region of four of the unidentified samples showed the presence of a three-basepair deletion and a T to C transition in the An. funestus-specific primer site. This change in sequence resulted in the species-specific primer not annealing and subsequent failure to amplify the region. Further analysis of this ITS2 region showed a high level of sequence variation (4.5%) when compared with the An. funestus control.

The D3 sequence analysis of the MalaF specimens showed a 1.5% variation compared with the An. funestus control, which for this gene region, is significant. Differences as small as 2–3 basepairs within the D3 region have been used to differentiate the members of the An. fluviatilis complex;22 An. minimus species A and C differ by five subsitutions. 23 Combined, the levels of variation seen in the MalaF specimens are significant. Levels of inter-specific sequence divergence can range from lows of 0.4% to 1.6% differences, as in the An. gambiae complex, 24 to 19% as observed between An. rivulorum in eastern and southern Africa and An. rivulorum-like in west and central Africa.5

A PCR primer (MalaFB) specific to the ITS2 region of the unidentified samples was designed and effectively used to amplify this region. However, this amplicon (approximately 390 basepairs) is too close in size to that of An. rivulorum (approximately 411 basepairs) to be incorporated into the multiplex PCR mixture, as seen by the species product ladder in Figure 2. The optimum difference in amplicon size for easy visualization on an agarose gel is approximately 50 basepairs, a criterion used in the design of the species-specific multiplex PCR assay. 16 However, the MalaFB primer does enable identification of our unidentified specimens where the initial multiplex PCR fails. We are currently testing the new primer on field-caught material from other regions that have failed to amplify using the An. funestus multiplex PCR. The seven MalaF samples that failed to amplify with the MalaFB primer are undergoing further molecular analysis. Because DNA extractions are routinely carried out with positive controls from insectary material, the quality of the DNA can be ensured. The DNA integrity of these samples was tested using primers known to amplify the members of the Funestus subgroup, and the presence of amplicons confirms that the failure to amplify using the MalaFB primer was not caused by DNA degradation.

Prior to the advent of DNA-based technologies, salivary gland and ovarian polytene chromosomes were used to distinguish members of sibling species complexes. Members of the European An. maculipennis complex were distinguished by the chromosomal banding patterns seen in salivary gland polytene chromosomes. 25,26 This success in cytotaxonomy was quickly followed by the cytogenetic description of the members of the An. gambiae complex.2729 The chromosomal banding patterns of the Malawian specimens displayed homosequential banding arrangements with An. funestus, but were fixed for the inverted arrangements 3a, 3b, and 5a, which are commonly polymorphic in An. funestus. Although An. vaneedeni also has homosequential chromosomes with An. funestus,13 the fixed inverted arrangements on arms 3 and 5 of MalaF distinguish it from An. vaneedeni. Even though cytogenetic studies on west African populations of An. funestus have provided evidence of species differentiation, 14,15 these investigators did not undertake cross-mating studies because of lack of colonized An. funestus at that time.

Species-crossing experiments have been widely used to prove the distinction of sibling species within anopheline complexes (An. maculatus form K, 30 An. pseudopunctipennis species C,31 An. minimus species E, 32 An. quadrimaculatus types A and B, 33 and An. annulipes species A and G 34) with hybrids being scored for asynapsis between homologous chromosomes and hybrid infertility. In the present study, the hybrid chromosomes resulting from the MalaF females × An. funestus males showed consistent asynapsis between homologous chromosomes, typical of inter-species crosses. 13,20,35 The male hybrids appeared to have normal testis morphology with the possible exception that the head region of the spermatozoa appeared narrower. Unfortunately, the effect of this narrower morphology in terms of male fertility is unknown because we were unable to carry out back crosses. Eggs were produced from the reciprocal crosses, but their viability was extremely low (< 0.2% hatch rate), suggesting a genetic discontinuity between the parental samples.

On the basis of the combined molecular, cytogenetic, and cross-mating evidence, we conclude that the Malawi population is a new member of the An. funestus subgroup. We provisionally designate it An. funestus-like until a formal description is published. Further molecular investigations are needed to determine how this new species impacts on the variation seen in restriction fragment length polymorphism 36 and mitochondrial DNA 37 analyses of An. funestus populations from the southern African region.

Further investigations into the biology of this new species are also required. Although none of the 61 specimens examined for malaria parasite infection during this study were positive for P. falciparum, the fact that these mosquitoes are common inside houses makes them potential vectors. Collections at different times of the year are needed to clarify their vector status and to provide data on the interactions between this new species and An. funestus s.s. in areas where they occur in sympatry.

Figure 1.
Figure 1.

Sequence alignment of the internal transcribed spacer region 2 of the unidentifiable Malawian (MalaF) samples and Anopheles funestus. Blocks highlight sequence variation. – indicates deletions.

Citation: The American Journal of Tropical Medicine and Hygiene Am J Trop Med Hyg 81, 3; 10.4269/ajtmh.2009.81.510

Figure 2.
Figure 2.

Primer MalaFB included in the polymerase chain reaction (PCR) cocktail mixture for Anopheles funestus group identifications. Lane 1, 100-basepair DNA ladder; lanes 2–4, positive controls: An. vaneedeni, An. funestus, and An. rivulorum, respectively; lane 5, MalaF sample; lanes 6 and 7, positive controls: An. parensis and An. leesoni, respectively; lane 8, PCR negative control.

Citation: The American Journal of Tropical Medicine and Hygiene Am J Trop Med Hyg 81, 3; 10.4269/ajtmh.2009.81.510

Figure 3.
Figure 3.

Sequence alignment of the partial D3 region of the unidentifiable Malawian (MalaF) samples (133 basepairs) and Anopheles funestus (135 basepairs). Blocks highlight sequence variation. – indicates deletions. The forward D3A primer binds approximately 90 basepairs upstream from the start of this sequence.

Citation: The American Journal of Tropical Medicine and Hygiene Am J Trop Med Hyg 81, 3; 10.4269/ajtmh.2009.81.510

Figure 4.
Figure 4.

Asynapsis in hybrid polytene chromosomes indicated by arrows.

Citation: The American Journal of Tropical Medicine and Hygiene Am J Trop Med Hyg 81, 3; 10.4269/ajtmh.2009.81.510

*

Address correspondence to Belinda L. Spillings, Vector Control Reference Unit, National Institute for Communicable Diseases, National Health Laboratory Service, Private Bag X4, Sandringham, Johannesburg, 2131, and Division of Virology and Communicable Disease Surveillance, School of Pathology of the University of the Witwatersrand, Johannesburg, South Africa. E-mail: belindas@nicd.ac.za

Authors’ addresses: Belinda L. Spillings, Basil D. Brooke, Lizette L. Koekemoer, and Maureen Coetzee, Vector Control Reference Unit, National Institute for Communicable Diseases, National Health Laboratory Service, Private Bag X4, Sandringham, Johannesburg, 2131, and Division of Virology and Communicable Disease Surveillance, School of Pathology of the University of the Witwatersrand, Johannesburg, South Africa. John Chiphwanya, National Malaria Control Programme, Lilongwe, Malawi. Richard H. Hunt, School of Animal, Plant and Environmental Sciences, University of the Witwatersrand, 1 Jan Smuts Avenue, Johannesburg, 2001, South Africa.

Acknowledgments: We thank Samuel Vezenegho for assistance with ELISA, Mike Lo for laboratory assistance, Christophe Kikankie for assistance with field collections, Paladin Resources Ltd., for facilitating the field collections, and R. A. Wirtz (Entomology Branch, Centers for Disease Control and Prevention, Atlanta, GA) for kindly supplying P. falciparum positive controls and the monoclonal antibody P. falciparum 2A10 for use in the indirect ELISA.

Financial support: This study was supported by the South African Medical Research Council, the South African Malaria Initiative and the South African Research Chair Initiative of the Department of Science and Technology, and the National Research Foundation.

REFERENCES

  • 1

    Gillies MT, de Meillon B, 1968. The Anophelinae of Africa South of the Sahara. Publications of the South African Institute for Medical Research, no. 54. Johannesburg, South Africa.

  • 2

    Gillies MT, Coetzee M, 1987. A Supplement to the Anophelinae of Africa South of the Sahara (Afrotropical Region). Publications of the South African Institute for Medical Research, no 55. Johannesburg, South Africa.

  • 3

    Harbach RE, 2004. The classification of genus Anopheles (Diptera: Culicidae): a working hypothesis of phylogenetic relationships. Bull Entomol Res 94 :537–553.

    • Search Google Scholar
    • Export Citation
  • 4

    Garros C, Harbach RE, Manguin S, 2005. Morphological assessment and molecular phylogenetics of the Funestus and Minimus Groups of Anopheles (Cellia). J Med Entomol 42 :522–536.

    • Search Google Scholar
    • Export Citation
  • 5

    Hackett BJ, Gimnig J, Guelbeogo W, Costantini C, Koekemoer LL, Coetzee M, Collins FH, Besansky NJ, 2000. Ribosomal DNA internal transcribed spacer (ITS2) sequences differentiate Anopheles funestus and An. rivulorum, and uncover a cryptic taxon. Insect Mol Biol 9 :369–374.

    • Search Google Scholar
    • Export Citation
  • 6

    Couhet A, Simard F, Toto J-C, Kengne P, Coetzee M, Fontenille D, 2003. Species identification within the Anopheles funestus group of malaria vectors in Cameroon and evidence for a new species. Am J Trop Med Hyg 69 :200–205.

    • Search Google Scholar
    • Export Citation
  • 7

    Green CA, 1982. Cladistic analysis of chromosome data (Anopheles (Cellia) Myzomyia). J Hered 73 :2–11.

  • 8

    Wilkes TJ, Matola YG, Charlwood JD, 1996. Anopheles rivulorum, a vector of human malaria in Africa. Med Vet Entomol 10 :108–110.

  • 9

    De Meillon B, Van Eeden GJ, Coetzee L, Coetzee M, Meiswinkel R, Du Toit CL, Hansford CF, 1977. Observations on a species of the Anopheles funestus subgroup, a sustpected exophilic vector of malaria parasites in North Eastern Transvaal, South Africa. Mosq News 37 :657–661.

    • Search Google Scholar
    • Export Citation
  • 10

    Evans AM, Symes CB, 1937. Anopheles funestus and its allies in Kenya. Ann Trop Med Parasitol 31 :105–111.

  • 11

    Evans AM, 1938. Mosquitoes of the Ethiopian Region. London: Bernard Quartich Ltd., Dulau and Co. Ltd., and The Oxford University Press.

  • 12

    De Meillon B, 1947. New records and species of biting insects from the Ethiopian Region II. J Entomol Soc S Afr 10 :110–124.

  • 13

    Green CA, Hunt RH, 1980. Interpretations of variation in ovarian polytene chromosomes of Anopheles funestus Giles, A. parensis Gillies and A. aruni? Genetica 51 :187–195.

    • Search Google Scholar
    • Export Citation
  • 14

    Lochouarn L, Dia I, Boccolini D, Coluzzi M, Fontenille D, 1998. Bionomical and cytogenetic heterogeneities of Anopheles funestus in Senegal. Trans R Soc Trop Med Hyg 92 :607–612.

    • Search Google Scholar
    • Export Citation
  • 15

    Costantini C, Sagnon, N’F, Ilboudo-Sanogo E, Coluzzi M, Boccolini D, 1999. Chromosomal and bionomic heterogeneities suggest incipient speciation in Anopheles funestus from Burkina Faso. Parassitologia 41 :595–611.

    • Search Google Scholar
    • Export Citation
  • 16

    Koekemoer LL, Kamau L, Hunt RH, Coetzee M, 2002. A cocktail polymerase chain reaction assay to identify members of the Anopheles funestus (Diptera: Culicidae) group. Am J Trop Med Hyg 6 :804–811.

    • Search Google Scholar
    • Export Citation
  • 17

    Collins FH, Mendez MA, Rasmussen MO, Meheffey PC, Besansky NJ, Finnerty V, 1987. A ribosomal RNA gene probes differentiates members of the Anopheles gambiae complex. Am J Trop Med Hyg 37 :37–41.

    • Search Google Scholar
    • Export Citation
  • 18

    Litvaitis MK, Nunn G, Thomas WK, Kocher TD, 1994. A molecular approach for the identification of Meiofaunal turbellarians (Platyhelminthes: Turbellaria). Mar Biol 158 :17–35.

    • Search Google Scholar
    • Export Citation
  • 19

    Hunt RH, 1973. A cytological technique for the study of Anopheles gambiae complex. Parassitologia 15 :137–139.

  • 20

    Davidson G, Paterson H, Coluzzi M, Mason G, Micks D, 1967. The Anopheles gambiae complex. Wright J, Pal R, eds. Genetics of Insect Vectors of Disease. London: Elsevier Publishing Company, 211–250.

  • 21

    Wirtz RA, Zavala F, Charoenvit Y, Cambell GH, Burkot TR, Schneider I, Esser KM, Beaudoin RL, Andre GR, 1987. Comparative testing of Plasmodium falciparum sporozoite monoclonal antibodies for ELISA development. Bull World Health Organ 65 :39–45.

    • Search Google Scholar
    • Export Citation
  • 22

    Singh OP, Chandra D, Nanda N, Raghavendra K, Sunil S, Sharma SK, Dua VK, Subbarao SK, 2004. Differentiation of members of the Anopheles fluviatilis species complex by an allele-specific polymerase chain reaction based on 28S ribosomal DNA sequences. Am J Trop Med Hyg 70 :27–32.

    • Search Google Scholar
    • Export Citation
  • 23

    Sharpe RG, Harbach RE, Butlin RK, 2000. Molecular variation and phylogeny of the members of the Minimus group of Anopheles subgenus Cellia (Diptera: Culicidae). Syst Entomol 25 :263–272.

    • Search Google Scholar
    • Export Citation
  • 24

    Paskewitz SM, Wesson DM, Collins FH, 1993. The internal transcribed spacers of ribosomal DNA in five members of the Anopheles gambiae species complex. Insect Mol Biol 2 :247–257.

    • Search Google Scholar
    • Export Citation
  • 25

    Frizzi G, 1947. Salivary gland chromosomes of Anopheles. Nature 160 :226–227.

  • 26

    Frizzi G, 1953. Étude cytog’enetique d’anopheles maculipennis en Italie. Extension des recherches ‘a d’autres esp’eces d’anoph’eles. Bull World Health Organ 9 :335–344.

    • Search Google Scholar
    • Export Citation
  • 27

    Coluzzi M, Sabatini A, 1967. Cytogenetic observations on species A and B of the Anopheles gambiae complex. Parassitologia 9 :73–88.

  • 28

    Coluzzi M, Sabatini A, 1968. Cytogenetic observations on species C of the Anopheles gambiae complex. Parassitologia 10 :155–166.

  • 29

    Coluzzi M, Sabatini A, 1969. Cytogenetic observations on the salt water species Anopheles merus and Anopheles melas, of the Gambiae complex. Parassitologia 11 :155–166.

    • Search Google Scholar
    • Export Citation
  • 30

    Thongwat D, Morgan K, O’Loughlin SM, Walton C, Choochote W, Somboon P, 2008. Crossing experiments supporting the specific status of Anopheles maculatus chromosomal form K. J Am Mosq Control Assoc 24 :194–202.

    • Search Google Scholar
    • Export Citation
  • 31

    Coetzee M, Estrada-Franco JG, Wunderlich CA, Hunt RH, 1999. Cytogenetic evidence for a species complex within Anopheles pseudopunctipennis Theobald (Diptera: Culicidae). Am J Trop Med Hyg 60 :649–653.

    • Search Google Scholar
    • Export Citation
  • 32

    Somboon P, Thongwat D, Choochote W, Walton C, Takagi M, 2005. Crossing experiments of Anopheles minimus species A and putative species E. J Am Mosq Control Assoc 21 :5–9.

    • Search Google Scholar
    • Export Citation
  • 33

    Kaiser PE, 1988. Cytotaxonomy as a tool for identification of siblings of the Anopheles quadrimaculatus complex. Fla Entomol 71 :311–323.

    • Search Google Scholar
    • Export Citation
  • 34

    Foley DH, Bryan JH, 1991. Anopheles annulipes Walker (Diptera: Culicidae) at Griffith New South Wales. 1. Two sibling species in sympatry. Aust J Entomol 30 :109–112.

    • Search Google Scholar
    • Export Citation
  • 35

    Hunt RH, Coetzee M, Fettene M, 1998. The Anopheles gambiae complex: a new species from Ethiopia. Trans R Soc Trop Med Hyg 92 :231–235.

    • Search Google Scholar
    • Export Citation
  • 36

    Garros C, Koekemoer LL, Kamau L, Awolola TS, Van Bortel W, Coetzee M, Coosemans M, Manguin S, 2004. Restriction fragment length polymorphism method for the identification of major African and Asian malaria vectors within the Anopheles funestus and An. minimus groups. Am J Trop Med Hyg 70 :260–265.

    • Search Google Scholar
    • Export Citation
  • 37

    Michel AP, Ingracsi MJ, Schemerhorn BJ, Kern M, Le Goff G, Coetzee M, Elissa N, Fontenille D, Vulule J, Lehmann T, Sagnon N’F, Costantini C, Besansky NJ, 2005. Rangewide population genetics structure of the African malaria vector Anopheles funestus. Mol Ecol 14 :4235–4248.

    • Search Google Scholar
    • Export Citation

Author Notes

Reprint requests: Maureen Coetzee, Vector Control Reference Unit, National Institute for Communicable Diseases, National Health Laboratory Service, Private Bag X4, Sandringham, Johannesburg, 2131, South Africa, E-mail: maureenc@nicd.ac.za.
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