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    Amplifications of the two ribosomal DNA fragments from the Anopheles funestus and An. minimus groups. A, Internal transcribed spacer 2. B, Domain 3. bp = basepairs.

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    Polymerase chain reaction-restriction fragment length polymorphism patterns for the internal transcribed spacer 2 (ITS2) and Domain 3 (D3) fragments of Anopheles funestus or An. minimus group after digestion with either Msp I or Bsi ZI. A, ITS2/Msp I. B, ITS2/Bsi ZI. C, D3/Msp I. D, D3/Bsi ZI. bp = basepairs.

  • View in gallery

    Msp I digestion patterns for Domain 3 (D3) showing the intragenomic variation within the Anopheles funestus populations. The boxes represent the restriction endonuclease sites. The cloning and sequencing of the D3 fragment show the nucleotide mutation at position 205 (underlined). bp = basepairs.

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RESTRICTION FRAGMENT LENGTH POLYMORPHISM METHOD FOR THE IDENTIFICATION OF MAJOR AFRICAN AND ASIAN MALARIA VECTORS WITHIN THE ANOPHELES FUNESTUS AND AN. MINIMUS GROUPS

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  • 1 Institute of Research for Development, Centre of Biology and Management of Populations, Montpellier, France; Vector Control Reference Unit, National Institute for Communicable Diseases, Johannesburg, South Africa; Department of Clinical Microbiology and Infectious Diseases, School of Pathology of the National Health Laboratory Service and University of the Witwatersrand, Johannesburg, South Africa; Centre for Biotechnology Research and Development, Nairobi, Kenya; Department of Parasitology, Prince Leopold Institute of Tropical Medicine, Antwerp, Belgium

The African Anopheles funestus and the Asian An. minimus groups are closely related and are probably considered distinct only because of their geographic separation. This study aimed at improving two identification methods based on polymerase chain reaction-restriction fragment length polymorphism (PCR-RFLP) already developed for either group. Each PCR-RFLP, either on the internal transcribed spacer 2 (ITS2) for the An. minimus group, and domain 3 (D3) for the An. funestus group, was applied to the other group for the standardization of one identification method applicable on both continents. The ITS2 fragment digested by Bsi ZI showed the highest diagnostic power. This assay allowed the discrimination of at least 13 Anopheles species within the subgenus Cellia from two continents (Africa and Asia), among which are five major malaria vectors. Moreover, digestion of the D3 with Msp I showed intragenomic variations within An. funestus populations. Two types of D3 copies (M and W) occurred in specimens from southern Africa. The populations from West-Central Africa presented only type W and East-Malagasy populations exhibited type M. Since An. funestus shows a great capacity of adaptation, these molecular variations, along with behavioral and ecologic ones, reinforce the hypothesis of a species complex that will need to be further investigated.

INTRODUCTION

Any biological study is meaningful and rigorous only if the concerned organism is accurately identified; this is particularly important in vector-borne disease studies where correct and precise identification of the target species has medical and practical implications, such as in vector control. Identification is mostly achieved using morphologic characters. However, there are a variety of circumstances in which a molecular approach will greatly improve the accuracy of species identification. This not only applies to sibling species but also to members of closely related groups with overlapping morphologic characters. The differentiation among the anopheline species of the Anopheles funestus and the An. minimus groups is based mainly on characteristics of the larval and egg stages, but none of these species can be identified with absolute certainty using the adult morphology. These two groups, which belong to the Myzomyia series, subgenus Cellia,1 are closely related.2

On the Afrotropical continent, the An. funestus group consists of eight species,3,4 namely An. brucei, An. confusus, An. fuscivenosus, and An. rivulorum, as well as the Funestus subgroup composed of An. aruni, An. funestus, An. parensis, and An. vaneedeni. Beside the anthropophilic and endophilic An. funestus, which is a highly efficient malaria vector, the other species of the group are mainly zoophilic and play little or no role in malaria transmission.4 Anopheles funestus presents a wide geographic distribution covering all subtropical Africa and Madagascar,4,5 while the other species have more restricted distributions in Africa.3,4 Although An. leesoni was recently removed from the An. funestus group,1 it was included in the identification assays of the members of the funestus group because of their morphologic similarities and this of misidentification with An. funestus s.l..6–8

Similarly, in the Asian region, the An. minimus group is composed of nine species, namely An. aconitus, An. filipinae, An. flavirostris, An. fluviatilis, An. mangyanus, An. pampanai, An. varuna, An. minimus A, and An. minimus C. The latter two belong to the An. minimus complex.1 Anopheles minimus s.l. is most commonly associated with rivers and streams throughout Southeast Asia, where it is one of the primary vectors of human malaria.2 Six species, An. aconitus, An. flavirostris, An. minimus species A and C, An. pampanai, and An. varuna, occur on mainland Southeast Asia.9–13 Since these species play different roles in malaria transmission, vector and non-vector species can be easily confused on the basis of adult morphology.

Molecular methods have been developed for distinguishing different species either in the An. minimus or the An. funestus group. The polymerase chain reaction-restriction fragment length polymorphism (PCR-RFLP) method has been shown to be more appropriate for dealing with a greater number of species, and particularly in the case when the local fauna is not well known. However, identification assays are often developed for specific geographic area, but these can be extended to closely related groups on other continents. Previous studies used PCR-RFLP assays for differentiating members of either the An. funestus or the An. minimus group.6,14 The assay developed by Koekemoer and others6 differentiates two species of the An. funestus group by digesting the domain 3 (D3) of the 28S ribosomal DNA (rDNA) with the endonuclease Hpa II (an isoschozomere of Msp I). Digestion of the internal transcribed spacer 2 (ITS2) rDNA with endonucleases BsiZ I and Msp I proved to be useful in the identification of the five members of the An. minimus group and related species.14 The present report shows a single PCR-RFLP assay that allows the identification of species belonging to both groups. The purpose of this study was to improve existing molecular techniques by selecting an unique, reliable, and simple method for standardizing the identification assays to members of both the An. funestus and An. minimus anopheline groups. Therefore, we tested these two PCR-RFLP assays on 10 members of both groups (Table 1)6,14 and studied the interspecific and intraspecific variations at both loci.

MATERIALS AND METHODS

Mosquito populations and standard identification.

Anopheline mosquitoes originating from different localities in Africa and Southeast Asia are shown in Table 1. All mosquitoes from Africa were wild collected, except for 30 specimens of An. funestus from Mozambique, which were from a colony reared under laboratory conditions for 19 months and maintained at the National Health Laboratory Service (Johannesburg, South Africa). Wild samples of adult An. funestus originated from eight countries, including Burkina Faso (Boromo) and Senegal (Kedougou) from West Africa; Cameroon (Simbock) from Central Africa; Angola (Calueque), South Africa (Kwa-Zulu Natal), and Mozambique from Southern Africa; and Kenya (Magaoni-Kwale) and Madagascar (Ambohimena) from East Africa. The four other species of the group were collected either in South Africa or Kenya (Table 1). Specimens from Southeast Asia were collected during extensive surveys, particularly landing catches on humans and bovines, resting captures, and larval collections. These surveys were made during May, August, and November 1998 and April 1999 in two villages, one located in northern Vietnam (Khoi, Hoa Binh Province) for the specimens of An. minimus A and C and the other in central Vietnam (Lang Nhot, Khanh Hoa Province), for the specimens of An. minimus A, An. aconitus, An. pampanai, and An. varuna (Table 1). Adult specimens were either dry-preserved (material from Africa) or stored at −80°C (material from Madagascar and Vietnam). After capture, all adult mosquitoes were identified on the basis of their morphology. Mosquitoes originated from Africa were identified using the keys of Gillies and Coetzee3 and Gillies and de Meillon.4 Mosquitoes from Southeast Asia were identified by use of a standard key for medically important anophelines of this region.15 The members of the An. funestus group were further differentiated using a single-stand conformation polymorphism-PCR8 or a species-specific rDNA PCR (ITS2).16 The allele-specific method developed by Kengne and others11 was used for identifying the members of the An. minimus group. Anopheles gambiae s.s. from Côte d’Ivoire was used as the outgroup.

Extraction, amplification, restriction, and sequencing of DNA.

Genomic DNA was extracted from individual adult mosquitoes, whole specimens, or mosquito parts, such as legs and wings (material from Madagascar and Kenya), according to the procedure of Linton and others.17 Amplifications were performed in a 25-μL reaction volume containing 200 μM dNTPs, 10× buffer, 10 μM of each primers, 0.5 units of Taq polymerase, and 3 μL of template. Two regions of rDNA were amplified using the PCR: ITS2 and D3 of the 28S region with the primers reported by Van Bortel and others for ITS214 and those reported by Koekemoer and others for D3.6 The PCR conditions were a denaturation step at 94°C for three minutes, followed by 35 cycles at 94°C for 30 seconds, 45°C for 40 seconds (for ITS2) or 63°C for 40 seconds (for D3), and 72°C for 30 seconds, and a final elongation at 72°C for 10 minutes. Products were visualized after electrophoresis on an ethidium bromide-stained 1.5% agarose gel.

Based on previous studies,6,14 two endonucleases were used, Msp I (5′-CCGG-3′) and Bsi ZI (5′-GGNCC-3′) (Promega, Madison, WI). The reaction mixture contained 2 μL of sterile distilled water, 2 μL of buffer provided by the manufacturer, 1 μL of enzyme and 15 μL of PCR-amplified template. The mixture was incubated for two hours either at 37°C for Msp I or 60°C for Bsi ZI. Samples were then subjected to electrophoresis on a 3% small-fragment agarose gel.

In a first step, four PCR-RFLP assays were tested using either the rDNA fragments containing ITS2 or D3 and the endonucleases Msp I or Bsi ZI. Three individuals from each species of both groups (when available) were used for the PCR-RFLP. For An. funestus, nine specimens were tested: three from Madagascar and one each from Burkina Faso, Senegal, Cameroon, Kenya, Mozambique, and South Africa. In a second step, all 381 anophelines (Table 1) were screened for the validation of the most discriminating PCR-RFLP assay (ITS2 digested with Bsi ZI). The PCR products were gel-purified with the QIAquick gel extraction kit (Qiagen, Valencia, CA) following the manufacturer’s instructions. Double strand of the PCR products were directly sequenced by MWG Biotech, Inc. (High Point, NC) with either primers ITS2 A and B or D3 A and B. Two specimens (when available) from each species and populations were sequenced, one in both directions and the second one in forward direction.

The ITS2 and D3 sequences were deposited in GenBank with the following accession numbers: AY259142–AY259162 and AY255105–AY255111. The alignment for each fragment was done using BioEdit.18

RESULTS

ITS2 and D3 fragments.

The ITS2 fragment length ranged from 850 basepairs for An. funestus and An. vaneedeni to 520 basepairs for members of the An. minimus group and An. leesoni. Anopheles parensis showed a band of 600 basepairs and An. rivulorum one of 540 basepairs (Figure 1A). The D3 amplified product was 400 basepairs in length for species of the An. funestus group and 380 basepairs for those of the An. minimus group (Figure 1B).

PCR-RFLP assays.

Of the four PCR-RFLP tested, three were able to differentiate between seven and ten populations and species (Figure 2A, B, and C). The amplified D3 fragment digested with Bsi ZI exhibited a monomorph pattern for species of each group, except for An. rivulorum (Figure 2D).

Bsi ZI restriction patterns.

The digestion of PCR-amplified ITS2 with Bsi ZI did not exhibit any intraspecific variations among the An. funestus populations, and all showed a quadruple banding pattern (120, 180, 230, and 330 basepairs) (Figure 2B). However, this PCR-RFLP produced a clear and distinct banding pattern differentiating all 11 species, including the outgroup species An. gambiae s.s. with three bands of 150, 200, and 210 basepairs. Within the An. funestus group, An. parensis, An. leesoni, and An. vaneedeni showed two bands of 230 and 380 basepairs, 200 and 240 basepairs, and 250 and 600 basepairs, respectively. Within the An. minimus group, An. minimus C also showed two bands, a characteristic one of 300 basepairs and one of 220 basepairs, whereas those of An. minimus A were 200 and 220 basepairs. Anopheles aconitus presented a restriction pattern with one band of 400 basepairs and An. varuna showed two major bands of 200 and 250 basepairs. The Bsi ZI restriction enzyme did not cut the ITS2 fragment of An. pampanai and An. rivulorum, which showed single bands of 520 and 540 basepairs, respectively. The results obtained with the assay that associated the amplified and digested ITS2 fragment with the Bsi ZI endonuclease were tested on 381 samples, including 203 An. funestus and all 178 specimens available for each other populations and species (Table 1). All individuals gave the appropriate species-specific pattern.

Msp I restriction patterns.

The digestion patterns of the D3 and ITS2 regions with Msp I showed an intraspecific variability within the An. funestus populations (Figure 2A and C). The assay on ITS2 showed two clusters, one containing specimens from West (Burkina Faso and Senegal) and Central (Cameroon) Africa, and the other containing those from East (Kenya, Mozambique, and Madagascar) Africa (Figure 2A). The same PCR-RFLP of ITS2 with Msp I also allowed the differentiation of six species, five belonging to the An. funestus group, as well as An. gambiae s.s. (Figure 2A). This assay did not separate all the species of the An. minimus group, which is consistent with previous results.14 Only An. varuna showed a distinct banding pattern (Figure 2A). The assay on the D3 fragment using Msp I showed only five patterns among the nine species of both groups, allowing the specific identification of only three species, An. rivulorum, An. minimus A, and An. pampanai, which showed an undigested D3 fragment (Figure 2C). Anopheles leesoni showed the same restriction pattern as An. minimus C, An. aconitus, and An. varuna. In addition, a smaller band of approximately 100 basepairs was observed in all four species of the An. funestus group, but was absent in all species of the An. minimus group and An. leesoni. This band was smaller in An. rivulorum than in the three other species in the An. funestus group.

Intragenomic variation of D3 within An. funestus.

The Mozambique D3 sequence showed two Msp I sites at 55 and 205 basepairs, whereas the restriction pattern is characterized by four bands, i.e., three cutting sites (Figure 3). The inconsistency between the restriction pattern and the sequence for this An. funestus population was resolved by reading the D3 chromatogram. At position 205, there were two peaks corresponding to a G or A, which suggested superimposed sequences (Figure 3). The restriction sites on the sequences of the West-Central or east African populations were congruent with the respective restriction patterns. Comparison of the D3 sequences showed that Malagasy and Kenyan sequences had a D3 fragment (type M) that was different from the one found in the westcentral African populations (type W). The two D3 sequences differed by one basepair substitution (G→A) at position 205 (Figure 3), which leads to the deletion of an Msp I restriction site in type M. Eighty-one individuals (14 from Angola, 42 from Mozambique [colony and wild specimens], and 25 from South Africa) (Table 1) showed the same four bands, and it appeared that this pattern resulted from the presence of both types M and W. The Mozambique specimens appear to have both copies M and W.

Digestion of the D3 locus with Msp I differentiated three intraspecific An. funestus clusters that included the populations from west and central Africa, the ones from Madagascar and Kenya, and those and the colony from southern Africa (Angola, Mozambique, and South Africa). This could be due either to hybridization of the west and central African populations with east African populations or to differences among copies of the rDNA cistron that are present in multiple copies within one genome. To test the latter hypothesis, we cloned, digested, and sequenced the D3 fragment of two An. funestus specimens from Burkina Faso, Madagascar, and Mozambique populations. For each individual, 30 clones were digested with the Msp I. Cloning showed that specimens from the Mozambique colony had copies of both types M and W within their genome, but type W was found to be dominant (70%) and independent of sex. Copies of types W and M were found in fixed proportions respectively in West-Central and east African populations.

DISCUSSION

Harrison pointed out that the An. funestus group from the Afrotropical region and the An. minimus group from Asia have never been studied jointly, even though they may contain species that are “so closely related that they are probably considered distinct only because of their geographical separation.”2 This is the first time that 10 species of these two groups have been studied together. The DNA method used for identifying members of these two groups was a PCR-RFLP that has been used in the discrimination of numerous anopheline species complexes (reviewed by Collins and Paskewitz).19 The amplification and digestion of the ITS2 with Bsi ZI allowed the differentiation of five species in the An. funestus group, five in the An. minimus group, and An. gambiae s.s. (outgroup). This PCR-RFLP was first developed by Van Bortel and others14 for the same five species of the Asian An. minimus group, along with hybrids between An. minimus A and C, An. jeyporiensis, and An. culicifacies B, two sympatric and related species of the Myzomyia series.14 Thus, by adding five species of the An. funestus group the diagnostic power of this RFLP-based method included 12 species within the Myzomyia series along with the heterozygotes between An. minimus A and C, and 13 species with the out-group An. gambiae s.s. This technique can be further extended to species not yet included in this study.

The ITS2 region had already shown its potential in distinguishing other species of the An. minimus group, such as An. flavirostris and the non-vector An. filipinae, with the Msp I endonuclease.20 This assay should be tested on the species of the An. fluviatilis complex, some of which are important malaria vectors in western regions of Asia.21 It is important to verify that there is a non-overlapping pattern of the digested fragments among the different species that composed both groups. The technique is easy to carry out, needs only a minimum amount of material (legs only) for the isolation of DNA, and requires only basic molecular equipment. In addition to having a high diagnostic power that allows the discrimination of at least 13 Anopheles species within the subgenus Cellia, which includes five major malaria vectors, this method helps standardize the identification techniques applicable in both continents, Africa and Asia. Moreover, this generalistic method is potentially applicable to anopheline species from other groups that may show new specific banding patterns. As suggested by the results obtained with this technique, both anopheline groups are closely related genetically, and this similarity might be of primary interest in the investigation of potentially common insecticide resistance mechanisms.

Many malaria vectors have sibling species that may greatly differ in their vectorial capacities, biology, and behavior. The D3 fragment digested with Msp I showed intraspecific variations within An. funestus, defining up to three clusters in populations from West and Central Africa, East Africa and Madagascar, and Southern Africa. The third cluster is represented by the specimens from Angola, Mozambique, and South Africa that showed copies of both types M and W. Since the rDNA is composed of tandemly repeated transcriptional units, these results showed clear intragenomic differences between copies of the rDNA D3 cistron in the Southern African populations. This was tested in 51 wild caught An. funestus mosquitoes from these three countries. The presence of either the M or W type was not found as a single variant in the samples tested. We are now investigating in more detail this intragenomic heterogeneity by looking for other possible similar variations in different rDNA fragments of An. funestus populations. Such intragenomic variations in the rDNA sequences have been reported in An. nuneztovari, a malaria vector in South America22 and in Simulium damnosum s.l., a vector of the agent of human onchocerciasis in West Africa.23 For these two species, data on large natural field populations have confirmed cytogenetic and molecular results, leading to the existence of species complex. Since this species shows a great capacity of adaptation as demonstrated by its wide distribution and ability to occupy quite diverse ecologic regions,24 it is still not clear whether the behavioral, ecologic, and molecular variations exhibited by An. funestus populations are linked to the presence of a species complex, although this is strongly suspected.25,26

Table 1

Species, geographic origins, and sample size of the Anopheles populations of the An. funestus and An. minimus groups used in the study*

Species (name abbreviations)Country, (province name, village name), regionSample size
* Samples from established colony.
An. funestus
    (AF B)Burkina Faso, (Boromo village), West Africa15
    (AF S)Senegal, (Kedougou village), West Africa25
    (AF C)Cameroon, (Simbock village), Central Africa21
    (AF G)Angola, (Calueque village), Southern Africa14
    (AF Mz*)Mozambique (colony), Southern Africa30
    (AF Mz)Mozambique (wild material), Southern Africa12
    (AF A)South Africa, (Kwa Zulu Natal Province), Southern Africa25
    (AF K)Kenya, (Magaoni-Kwale village), East Africa5
    (AF Md)Madagascar, (Ambohimena village), East Africa56
An. funestusTotal203
An. leesoni (AL)South Africa, (Tzaneen Province), Southern Africa2
An. parensis (AP)Kenya, (Mwea village), East Africa32
An. rivulorum (AR)South Africa, (KwaZulu Natal or Mpumalanga Province), Southern Africa36
An. vaneedeni (AV)South Africa, (Tzaneen Province), Southern Africa8
An. Funestus groupTotal281
    An. aconitus (A)Vietnam (Khanh Hoa Province, Lang Nhot village), Southeast Asia20
    An. minimus sp. A (MA)Vietnam (Khanh Hoa, Hoa Binh Provinces; Khoi, Lang Nhot villages), Southeast Asia20
    An. minimus sp. C (MC)Vietnam, (Hoa Binh Province, Khoi village), Southeast Asia20
    An. pampanai (P)Vietnam, (Khanh Hoa Province, Lang Nhot village), Southeast Asia20
    An. varuna (V)Vietnam, (Khanh Hoa Province, Lang Nhot village), Southeast Asia20
An. Minimus groupTotal100
An. Funestus and
    An. Minimus groupsTotal381
Figure 1.
Figure 1.

Amplifications of the two ribosomal DNA fragments from the Anopheles funestus and An. minimus groups. A, Internal transcribed spacer 2. B, Domain 3. bp = basepairs.

Citation: The American Journal of Tropical Medicine and Hygiene Am J Trop Med Hyg 70, 3; 10.4269/ajtmh.2004.70.260

Figure 2.
Figure 2.

Polymerase chain reaction-restriction fragment length polymorphism patterns for the internal transcribed spacer 2 (ITS2) and Domain 3 (D3) fragments of Anopheles funestus or An. minimus group after digestion with either Msp I or Bsi ZI. A, ITS2/Msp I. B, ITS2/Bsi ZI. C, D3/Msp I. D, D3/Bsi ZI. bp = basepairs.

Citation: The American Journal of Tropical Medicine and Hygiene Am J Trop Med Hyg 70, 3; 10.4269/ajtmh.2004.70.260

Figure 3.
Figure 3.

Msp I digestion patterns for Domain 3 (D3) showing the intragenomic variation within the Anopheles funestus populations. The boxes represent the restriction endonuclease sites. The cloning and sequencing of the D3 fragment show the nucleotide mutation at position 205 (underlined). bp = basepairs.

Citation: The American Journal of Tropical Medicine and Hygiene Am J Trop Med Hyg 70, 3; 10.4269/ajtmh.2004.70.260

Authors’ addresses: C. Garros and S. Manguin, Centre of Biology and Management of Populations, Campus de Baillarguet, CS 30016, 34988 Montferrier sur Lez, France, Telephone: 33- 4-99-62-33-28/27, Fax: 33-4-99-62-33-45, E-mails: garros@mpl.ird.fr and manguin@mpl.ird.fr. L. L. Koekemoer and M. Coetzee, Medical Entomology, Department of Clinical Microbiology and Infectious Diseases, School of Pathology of the South African Institute for Medical Research and the University of the Witwatersand, PO Box 1038, Johannesburg, South Africa, Telephone: 27-11-489-9390, Fax: 27-11-489-9399. L. Kamau and T. S. Awolola, Centre for Biotechnology Research and Development, Kenya Medical Research Institute, PO Box 54840, Nairobi, Kenya. W. Van Bortel and M. Coosemans, Department of Parasitology, Prince Leopold Institute of Tropical Medicine, Nationalestraat 155, B-2000, Antwerp, Belgium.

Acknowledgments: This study was made possible because of the assistance of a number of people who provided mosquito samples from both continents (subtropical Africa and Southeast Asia). We are especially grateful to D. Fontenille, B. M. Ntomwa, and H. D. Trung for providing us with mosquitoes populations from West-Central Africa, Angola, and Vietnam, respectively.

Financial support: This work was partially supported by a European Community grant (research project ERBIC 18CT970211); a VIH-PAL (HIV/Malaria) 2000 grant from the French Ministry of Research to S. Manguin; the World Health Organization/AFRO for a multi-center project to M. Coosemans; a grant from the UNDP/World Bank/World Health Organization Special Program for Research and Training in Tropical Diseases to L. L. Koekemoer; and a grant from South African Medical Research Council and the National Health Laboratory Service to L. L. Koekemoer.

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Author Notes

Reprint requests: C. Garros or S. Manguin, Centre of Biology and Management of Populations, Campus de Baillarguet, CS 30016, 34988 Montferrier sur Lez, France.
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