EDP Sciences logo
Submit your paper
Search
Menu
All issues Volume 31 (2024) Parasite, 31 (2024) 31 Full HTML
Open Access
Issue
Parasite
Volume 31, 2024
Article Number 31
Number of page(s) 7
DOI https://doi.org/10.1051/parasite/2024032
Published online 18 June 2024

© T.M. Andrianinarivomanana et al., published by EDP Sciences, 2024

Licence Creative CommonsThis is an Open Access article distributed under the terms of the Creative Commons Attribution License (https://creativecommons.org/licenses/by/4.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

Introduction

Malaria is still a public health concern in many countries, mainly in Africa [38]. Malaria is transmitted by mosquitoes belonging to the Anopheles genus. The Anopheles genus is further subdivided into eight subgenera of which three (Anopheles, Cellia, and Nyssorhynchus) contain all known dominant vector species (DVS) of human malaria [34]. For countries close to malaria elimination, residual malaria is of high concern. In this regard, the role of the so-called secondary malaria vectors, often outdoor biters, in maintaining malaria transmission has been considered [39]. However, for decades most laboratory and field studies focused on the primary or dominant malaria vectors [34].

With the stagnation in worldwide malaria incidence over the last decade, it is more and more important to better understand the bionomics of secondary malaria vectors for which limited data are available. This is especially crucial for secondary malaria vectors in Madagascar, a country that has recorded a worrying increase in malaria cases despite indoor residual spraying and distribution campaigns for insecticidal-treated nets (WHO 2022).

Anopheles coustani is an Afrotropical anopheline species belonging to the Anopheles subgenus. In several African countries, including Madagascar, An. coustani is now widely recognized as a secondary vector of human malaria parasites [15, 17, 20, 24, 26]. Although it was reported as being both anthropophilic and exophagic (feeding outdoors on humans) in South Africa [10], An. coustani is often referred to as a zoophilic mosquito [35]. Nevertheless, because of its abundance it can be a locally major malaria vector [15, 28]. Indeed, in Madagascar, An. coustani was shown to be the major vector of both Plasmodium vivax and Plasmodium falciparum in a village where it showed high anthropophilic behavior, associated with both endophilic and exophilic behavior [18]. Furthermore, An. coustani has been implicated as a potential vector of Rift Valley Fever virus in Madagascar [31] and of Chikungunya virus in Senegal [13].

Anopheles coustani Laveran was first described in 1900 by Alphonse Laveran from samples received from Madagascar [23]. Anopheles mauritianus described at about the same time from mosquitoes collected in the Mascarene Islands was later considered as identical to An. coustani Laveran [14, 32]; details are presented by M. Coetzee [11]. Using cytogenetics on larval polytene chromosomes, M. Coetzee reported the existence of two cryptic species within An. coustani collected in South Africa and Eswatini (former Swaziland) [10]. When crossed, these species named species A and species B produced infertile male F1. More recently, using An. coustani samples from Madagascar and morphology as well as cytogenetics, the same author confirmed that the Malagasy mosquitoes belong to the former species A, and conclude that species B was a different species named An. crypticus [11]. Of note, Malagasy specimens were then referred to as neotypes as the original type specimens were lost. In 2020, sampling the genetic diversity of An. coustani populations in Zambia and the Democratic Republic of Congo revealed two distinct phylogenetic groups [8].

To our knowledge, no colony of An. coustani has ever been established. As for other species, having at hand a colony will be instrumental for various types of studies: insecticide assays with the colony as a reference, behavioral study for refining vector control strategies, assessing whether the bacteria Asaia or Wolbachia could be considered as additional tools for controlling mosquito populations or their vectorial competence toward Plasmodium and viruses [6, 7, 33].

Establishing a novel mosquito colony from field collected samples can be very tricky. The main challenge is mating behavior. Indeed, many anopheline mosquitoes are eurygamous, mating in open space often in swarms, and anopheline females are also known for their refractoriness to multiple mating [9, 36]. Anopheles coustani is no exception, as evidenced in preliminary assays while working with large numbers of F1 progeny obtained by in-tube forced oviposition of gravid wild females [27]. Even though F1 females would readily feed on blood, none would lay eggs over more than a week post blood feeding. Observation of a subset of the spermatheca of F1 females revealed that none had been fertilized (Jean Jose Nepomichene and Bourgouin, unpublished).

To overcome the absence of mating in a constrained environment (rearing cages), a “forced mating” technology was developed [4], and successfully used for establishing anopheline colonies [1, 29], even though the technique is highly demanding in human resources and skill. However, forced mating does not equal female fertilization. Indeed, it was reported that males from major anopheline vectors from the Nyssorhynchus subgenus, including Anopheles darlingi, would readily copulate by the forced mating technique but would fail to inseminate the females ([22]; Puchot, personal observation). On the contrary, several colonies of anophelines from both the Anopheles and Cellia subgenus were created and maintained using the forced mating technique [1, 3, 29].

Based on successful published records on anopheline from the Anopheles subgenus and our preliminary observations, we initiated the establishment of the first An. coustani colony using the forced mating technique. Here, we provide details on the major initial steps until we obtained the eighth generation and discuss ways of improvement toward the establishment of a free-mating colony of An. coustani.

Material and methods

Producing the F1 generation from wild An. coustani

Anopheles coustani females were captured in Maroharona (17°36’44.33”S, 46°56’2.11”E), municipality of Andriba (Maevatanana district) in the Northwest fringe of the Central Highlands of Madagascar, in September 2019. The mosquitoes were collected in zebu parks using mouth aspirators, as previously described [27]. Engorged females were transferred into 30 cm3 cages, provided with a 10% sucrose solution on cotton bowls. The cages were covered with a wet black fabric and transported over the next day to the Medical Entomology Unit at the Institut Pasteur de Madagascar in Antananarivo, located a 6 h drive away from Andriba. Morphological identification on live mosquitoes was confirmed one by one using a dichotomous identification key adapted from Gjrebine [19], Fontenille [16], and Coetzee [12]. Females were transferred to a rearing cage (32.5 × 32.5 × 32.5 cm BugDorm-4® -Bioquip, Rancho Dominguez, CA, USA).

A beaker half-filled with dechlorinated water was placed inside the cage for egg collection and surveyed every day for 5 days. Each batch of eggs was treated for 10 min with a 0.01% formaldehyde solution and extensively rinsed with dechlorinated water for limiting potential contamination with micro-organisms such as microsporidia. Eggs were then transferred to larval rearing pans (26 × 26 × 11 cm (Gilac®), filled with 1L of dechlorinated tap water (water height ~ 1.5 cm). L1 and L2 larvae were fed on Tetramin Baby fish food (Tetra®), whereas L3 and L4 were fed on ground cat food. When pupae started appearing, they were counted every day and transferred to a small beaker containing dechlorinated water and placed in a novel rearing cage. A 10% sucrose solution-containing cotton bowl was placed on top of the rearing cage and changed every other day.

Insectary conditions

Larvae and adults were reared at 26.9 ± 0.7 °C, 77 ± 7.7% RH, with a 12:12 night and day light regimen.

Establishing the An. coustani colony

From the F1 adults obtained from wild F0 mosquitoes, the artificial mating (forced mating also called forced copulation) strategy of males and females was used. This followed the procedure initially described by Baker et al. [4], refined by Ow Yang et al. [29] and nicely documented by the MR4 staff [25]. Females were given a blood meal 24 h before forced copulation. Males and females were cold anesthetized just before the process as illustrated in Figure 1 panels B&C. Each male (3–5 days old) was used for fertilizing 2 females (5–10 days old). We found it unnecessary to decapitate males for copulation stimulation. After forced copulation, females were transferred to a novel rearing cage and provided a blood meal every other day to stimulate egg production. From egg production to adult emergence, the procedure described above was used and led to the successful production of eight generations between September 2019 and May 2020. Because of the general movement restriction imposed in Madagascar during the Covid-19 outbreak, the colony could not be maintained further than the F8.

thumbnail Figure 1

Illustrations of An. coustani forced copulation procedure and developmental stages. Female An. coustani (A and F). Forced copulation: the male is hooked on a dissecting needle and presented to a cold anesthetized female; decapitated male (B); non-decapitated male (C). Fourth instar larvae (D). Pupa (E).

Recorded rearing parameters

At each generation, the following parameters were recorded: number of females successfully fertilized and their egg production, L1 to pupae growing success, as well as larvae to adult developmental time.

Results and discussion

Establishing the first six generations of An. coustani

Starting with 31 gravid F0 wild females free to lay eggs in cage, we obtained 260 eggs, which is quite a low and challenging number (8.4 eggs/female). This number is actually very similar to previous results [27]. The 260 eggs were obtained from a unique gonotrophic cycle of the females as they refused to feed on blood provided through the Hemotek membrane feeding device. For the sake of the project, the females from each subsequent generation were fed on a rabbit multiple times.

As previous attempts revealed that An. coustani would not mate in cages, the production of each generation was performed by artificial copulation at the ratio of 1 male for 2 females. It was not necessary to decapitate the male to obtain efficient copulation and insemination of the females. Table 1 summarizes the number of males and females used for forced mating at each generation, as well as the number of blood meals that the mated females received, and the mean number of eggs laid per female at each generation. The mean number of eggs produced per female increased slightly between F1 and F5, despite a reduced number of blood meals. This is possibly due to the acquisition of better skills for performing the forced copulation technique. Accordingly, this led to an increase of total adults from F1 to F6 (Figure 2). The production of F7 and F8 was achieved; however, no data on egg and adult numbers were recorded due to the Covid-19 lockdown.

thumbnail Figure 2

Yield in eggs and adults from F1 to F6. The graph represents the number of eggs obtained at each generation (Purple line and left scale) and the subsequent number of adults (green line and right scale).

Table 1

Number of males and females used at each generation in the forced mating procedure and mean egg production by each mated female.

Bionomic features

Egg to adult production

As depicted in Table 2, each generation suffered a drastic reduction, with a mean survival rate from egg to adult of 36.3%, with a broad range from 24.2 to 62.2%. This latter value happened at generation F3, but no clue of this result could be identified. Nevertheless, the egg to adult survival rate increased slightly across the generation and was higher than previous results obtained by Jean Jose Nepomichene in 2017, on F1 only [21]. Except for the F3 generation, a high mortality rate occurred during the larval stages. Normalization of larval density and food amount might contribute to limit this rather high mortality rate. It is also striking that the mortality rate of pupae was quite high (18.7–44.7%). These mortality rates might be a consequence of poor larval development linked to restricted food availability and larval competition.

Table 2

Summary of the raw data obtained across the production of the first six generation of An. coustani starting with 31 wild gravid females producing 260 eggs.

Larval to adult development

Previous work at a field station (Andriba, Madagascar) revealed that the larval to adult development of An. coustani F1 was quite asynchronous and long. Indeed, larval development lengthened more than 25 days, while An. arabiensis F1 population produced side by side would reach the adult stage in 10–15 days, as usually observed in insectary with tightly controlled parameters (Bourgouin et al., unpublished, 2016). Under the rearing conditions reported here, the larval to adult stage of An. coustani decreased across the generations and exhibited a different pattern of adult emergence (Figure 3). Indeed, F1 adults emerged in two peaks. The first peak occured between day 16 and day 18 from the day of egg laying and the second peak at day 20. From F2 to F4, adult emergence followed a 2-peak pattern, but each peak occurred earlier, except for the second peak of F2 that was at day 20, as for F1. Interestingly, F5 exhibited a broad single peak of adult emergence between day 11 and day 17, while for F6, a sharp single peak of adult emergence centered at day 13. These changes in adult emergence patterns possibly reveal a trait of adaptation to colonization.

thumbnail Figure 3

Graphic representation of An. coustani adult emergence at each generation. (A) Individual graphs showing the pattern of emergence of F1 to F6. (B) Aggregated graphs highlighting the clear changing patterns of adult emergence from F1 to F6 both in mode and date of the peak of emergence. The scale of the abscissa axis corresponds to days till the first appearance of L1.

Concluding remarks

We report here the first successful estabishment of a colony of An. coustani. As this mosquito species would not mate in cages, each of the 8 generations produced resulted from forced mating. This is comparable to the Anopheles cracens colony recently established to the 6th generation by forced mating [1]. On the contrary, a novel Anopheles atroparvus colony was established to the 10th generation without forced mating [5]. Given that Anopheles coustani and An. atroparvus are from the Anopheles subgenus, while An. cracens belongs to the Cellia subgenus, no conclusions can be drawn as to whether species from the Anopheles subgenus would require forced copulation for colony establishment. Indeed, even a reference colony of Anopheles dirus (Cellia subgenus) is maintained by forced copulation at the MR4 repository (WRAIR2 strain, https://www.beiresources.org/Catalog/BEIVectors/MRA-700.aspx)

As maintaining a colony by forced copulation is highly demanding in human resources and skill, the biggest refinement would be to obtain a free-mating colony. Towards this goal, it would be of great interest to test whether the tools developed for establishing An. darlingi colonies would be beneficial, notably the use of light flashes to stimulate free copulation and shift in night temperature [2, 30, 37]. As discussed previously, additional refinements in the larval feeding regimen and larval density would possibly contribute to a decrease in the mortality rate during both larval and pupal stages. Overall, having at hand an An. coustani colony, whether free mating or maintained by forced copulation, as An. dirus, will provide an unvaluable tool to better understand the biology of this secondary malaria vector and assessing relevant malaria transmission control strategies.

Acknowledgments

We wish to thank the technicians of the Medical Entomology Unit from Institut Pasteur de Madagascar for their support in the field and the insectary work (Tata, Lala, Fidélis, Andrisoa, and Onja). Thanks are also due to Jean-Pierre Ratavilahy, our guide in Andriba; to Dadabe, the owner of the zebu park; and members of the CSBII and of the city hall in Andriba. We are grateful to Richard Paul (Ecology and Emergence of Arthropod-borne Pathogens, Institut Pasteur) for improving the quality of the manuscript. This work was supported by funds from Institut Pasteur de Madagascar to RG and a Girard PhD grant to TMA, who also received an award grant from L’Oréal-Unesco-FWIS Sub-Saharan Africa Young Talents Awards. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

Conflict of interest

The authors declare that there are no competing interests.

Author contribution

Conceived the project: TMA, CB and RG. Designed the experiments: TMA, NP, CB. Performed the experiments: TMA, FTR, MRA, MRR and HJVS. Analyzed the data: TMA and CB. Drafted the paper: TMA and CB. Reviewed and edited the manuscript: TMA, RG and CB. All authors contributed to the article and approved the submitted version.

References

  1. Amir A, Sum JS, Lau YL, Vythilingam I, Fong MY. 2013. Colonization of Anopheles cracens: A malaria vector of emerging importance. Parasites and Vectors, 6(1), 81. [CrossRef] [Google Scholar]
  2. Araujo MDS, Andrade AO, Santos N, Pereira DB, Costa GDS, Paulo PFM, Rios CT, Moreno M, Pereira-da-Silva LH, Medeiros JF. 2019. Brazil’s first free-mating laboratory colony of Nyssorhynchus darlingi. Revista da Sociedade Brasileira de Medicina Tropical, 52, e20190159. [CrossRef] [PubMed] [Google Scholar]
  3. Baker RH. 1964. Mating problems as related to the establishment and maintenance of laboratory colonies of mosquitos. Bulletin of the World Health Organization, 31, 467–468. [PubMed] [Google Scholar]
  4. Baker RH, Drench WL, Kitzmiller JB. 1962. Induced copulation in Anopheles mosquitoes. Mosquito News, 22(1), 16–17. [Google Scholar]
  5. Birnberg L, Aranda C, Talavera S, Núñez AI, Escosa R, Busquets N. 2020. Laboratory colonization and maintenance of Anopheles atroparvus from the Ebro Delta, Spain. Parasites and Vectors, 13(1), 394. [CrossRef] [Google Scholar]
  6. Capone A, Ricci I, Damiani C, Mosca M, Rossi P, Scuppa P, Crotti E, Epis S, Angeletti M, Valzano M, Sacchi L, Bandi C, Daffonchio D, Mandrioli M, Favia G. 2013. Interactions between Asaia, Plasmodium and Anopheles: New insights into mosquito symbiosis and implications in Malaria Symbiotic Control. Parasites and Vectors, 6(1), 182. [CrossRef] [Google Scholar]
  7. Caragata EP, Dutra HLC, Sucupira PHF, Ferreira AGA, Moreira LA. 2021. Wolbachia as translational science: Controlling mosquito-borne pathogens. Trends in Parasitology, 37(12), 1050–1067. [CrossRef] [PubMed] [Google Scholar]
  8. Ciubotariu II, Jones CM, Kobayashi T, Bobanga T, Muleba M, Pringle JC, Stevenson JC, Carpi G, Norris DE. 2020. Genetic diversity of Anopheles coustani (Diptera: Culicidae) in malaria transmission foci in Southern and Central Africa. Journal of Medical Entomology, 57(6), 1782–1792. [CrossRef] [PubMed] [Google Scholar]
  9. Clements A. 1999. The biology of mosquitoes. Vol. 2: Sensory reception and behaviour, vol 2, Cabi Publishing: Oxforshire. [CrossRef] [Google Scholar]
  10. Coetzee M. 1983. Chromosomal and cross-mating evidence for two species within Anopheles (A.) coustani (Diptera: Culicidae). Systematic Entomology, 8(2), 137–141. [CrossRef] [Google Scholar]
  11. Coetzee M. 1994. Anopheles crypticus, new species from South Africa is distinguished from Anopheles coustani (Diptera: Culicidae). Mosquito Systematics, 26, 125–131. [Google Scholar]
  12. Coetzee M. 2020. Key to the females of Afrotropical Anopheles mosquitoes (Diptera: Culicidae). Malaria Journal, 19(1), 70. [CrossRef] [PubMed] [Google Scholar]
  13. Diallo D, Sall AA, Buenemann M, Chen R, Faye O, Diagne CT, Faye O, Ba Y, Dia I, Watts D, Weaver SC, Hanley KA, Diallo M. 2012. Landscape ecology of sylvatic Chikungunya Virus and mosquito vectors in Southeastern Senegal. PLOS Neglected Tropical Diseases, 6(6), e1649. [CrossRef] [PubMed] [Google Scholar]
  14. Dyé L. 1902. Notes et observations sur les Culicides. Archives de Parasitologie, 6, 359–376. [Google Scholar]
  15. Finney M, McKenzie BA, Rabaovola B, Sutcliffe A, Dotson E, Zohdy S. 2021. Widespread zoophagy and detection of Plasmodium spp. in Anopheles mosquitoes in southeastern Madagascar. Malaria Journal, 20(1), 25. [CrossRef] [PubMed] [Google Scholar]
  16. Fontenille D. 1989. Clé de détermination des moustiques adultes de Madagascar, Antananarivo, Madagascar: Internal Document -Unité d’entomologie -Institut Pasteur de Madagascar. https://www.pasteur.mg/. [Google Scholar]
  17. Fornadel CM, Norris LC, Franco V, Norris DE. 2011. Unexpected anthropophily in the potential secondary malaria vectors Anopheles coustani s.l. and Anopheles squamosus in Macha, Zambia. Vector-Borne and Zoonotic Diseases, 11(8), 1173–1179. [CrossRef] [PubMed] [Google Scholar]
  18. Goupeyou-Youmsi J, Rakotondranaivo T, Puchot N, Peterson I, Girod R, Vigan-Womas I, Paul R, Ndiath MO, Bourgouin C. 2020. Differential contribution of Anopheles coustani and Anopheles arabiensis to the transmission of Plasmodium falciparum and Plasmodium vivax in two neighbouring villages of Madagascar. Parasites and Vectors, 13(1), 430. [CrossRef] [Google Scholar]
  19. Grjebine A. 1966. Insectes diptères Culicidae Anophelinae. Faune de Madagascar. Paris: ORSTOM, CNRS. p. 489. [Google Scholar]
  20. Hamon J, Mouchet J. 1961. Les vecteurs secondaires du paludisme humain en Afrique. Médecine Tropicale, 21(Special), 643–660. [Google Scholar]
  21. Jean Jose Nepomichene TN. 2017. Biologie d’Anopheles coustani (Diptera : Culicidae) et implications dans la transmission du Plasmodium et du Virus de la Fièvre de la Vallée du Rift à Madagascar. In École Doctorale Sciences de la Vie et de l’Environnement, Madagascar: Université d’Antananarivo. PhD Thesis. [Google Scholar]
  22. Lardeux F, Quispe V, Tejerina R, Rodriguez R, Torrez L, Bouchite B, Chavez T. 2007. Laboratory colonization of Anopheles pseudopunctipennis (Diptera: Culicidae) without forced mating. Comptes Rendus Biologie, 330(8), 571–575. [CrossRef] [Google Scholar]
  23. Laveran A. 1900. Sur un Anopheles provenant de Madagascar. Comptes Rendus de la Société de Biologie, 52, 109–110. [Google Scholar]
  24. Lobo NF, Laurent BS, Sikaala CH, Hamainza B, Chanda J, Chinula D, Krishnankutty SM, Mueller JD, Deason NA, Hoang QT, Boldt HL, Thumloup J, Stevenson J, Seyoum A, Collins FH. 2015. Unexpected diversity of Anopheles species in Eastern Zambia: Implications for evaluating vector behavior and interventions using molecular tools. Scientific Reports, 5(1), 17952. [CrossRef] [PubMed] [Google Scholar]
  25. MR4. Methods in Anopheles Research. 2015 [cited; Available from: https://www.beiresources.org/Portals/2/VectorResources/2016%20Methods%20in%20Anopheles%20Research%20full%20manual.pdf. [Google Scholar]
  26. Mwangangi JM, Muturi EJ, Muriu SM, Nzovu J, Midega JT, Mbogo C. 2013. The role of Anopheles arabiensis and Anopheles coustani in indoor and outdoor malaria transmission in Taveta District, Kenya. Parasites and Vectors, 6(1), 114. [CrossRef] [Google Scholar]
  27. Nepomichene TN, Andrianaivolambo L, Boyer S, Bourgouin C. 2017. Efficient method for establishing F1 progeny from wild populations of Anopheles mosquitoes. Malaria Journal, 16(1), 21. [CrossRef] [PubMed] [Google Scholar]
  28. Nepomichene TN, Tata E, Boyer S. 2015. Malaria case in Madagascar, probable implication of a new vector, Anopheles coustani. Malaria Journal, 14, 475. [CrossRef] [PubMed] [Google Scholar]
  29. Ow Yang CK, St Maria FL, Wharton RH. 1963. Maintenance of a laboratory colony of Anopheles maculatus (Theobald) by artificial mating. Mosquito News, 23, 34–35. [Google Scholar]
  30. Puchot N, Lecoq M-T, Carinci R, Duchemin JB, Gendrin M, Bourgouin C. 2022. Establishment of a colony of Anopheles darlingi from French Guiana for vector competence studies on malaria transmission. Frontiers in Tropical Diseases, 3. https://www.frontiersin.org/articles/10.3389/fitd.2022.949300. [CrossRef] [Google Scholar]
  31. Ratovonjato J, Olive M-M, Tantely LM, Andrianaivolambo L, Tata E, Razainirina J, Jeanmaire E, Reynes J-M, Elissa N. 2010. Detection, isolation, and genetic characterization of Rift Valley Fever Virus from Anopheles (Anopheles) coustani, Anopheles (Anopheles) squamosus, and Culex (Culex) antennatus of the Haute Matsiatra Region, Madagascar. Vector-Borne and Zoonotic Diseases, 11(6), 753–759. [Google Scholar]
  32. Senevet G. 1932. À propos d’Anopheles coustani Laveran. Archives de l’Institut Pasteur d’Algérie, 10, 478–484. [Google Scholar]
  33. Shane JL, Grogan CL, Cwalina C, Lampe DJ. 2018. Blood meal-induced inhibition of vector-borne disease by transgenic microbiota. Nature Communications, 9(1), 4127. [CrossRef] [PubMed] [Google Scholar]
  34. Sinka ME, Bangs MJ, Manguin S, Coetzee M, Mbogo CM, Hemingway J, Patil AP, Temperley WH, Gething PW, Kabaria CW, Okara RM, Van Boeckel T, Godfray HC, Harbach RE, Hay SI. 2010. The dominant Anopheles vectors of human malaria in Africa, Europe and the Middle East: Occurrence data, distribution maps and bionomic precis. Parasites and Vectors, 3, 117. [CrossRef] [Google Scholar]
  35. Tantely ML, Rakotoniaina JC, Tata E, Andrianaivolambo L, Razafindrasata F, Fontenille D, Elissa N. 2013. Biology of mosquitoes that are potential vectors of Rift Valley Fever virus in different biotopes of the central highlands of Madagascar. Journal of Medical Entomologie, 50(3), 603–610. [CrossRef] [PubMed] [Google Scholar]
  36. Tripet F, Touré YT, Dolo G, Lanzaro GC. 2003. Frequency of multiple inseminations in field-collected Anopheles gambiae females revealed by DNA analysis of transferred sperm. American Journal of Tropical Medicine and Hygiene, 68(1), 1–5. [CrossRef] [Google Scholar]
  37. Villarreal-Trevino C, Vasquez GM, Lopez-Sifuentes VM, Escobedo-Vargas K, Huayanay-Repetto A, Linton YM, Flores-Mendoza C, Lescano AG, Stell FM. 2015. Establishment of a free-mating, long-standing and highly productive laboratory colony of Anopheles darlingi from the Peruvian Amazon. Malaria Journal, 14, 227. [CrossRef] [PubMed] [Google Scholar]
  38. WHO. 2023. World Malaria Report 2023, ISBN 978-92-4-008617-3. [Google Scholar]
  39. Yaw Asare A, Mariangela B, Guiyun Y. 2016. Secondary malaria vectors of Sub-Saharan Africa: Threat to malaria elimination on the continent? in: Current Topics in Malaria, Chapter 20, Alfonso JR-M, Ed. Rijeka, Croatia: IntechOpen. [Google Scholar]

Cite this article as: Andrianinarivomanana TM, Randrianaivo FT, Andriamiarimanana MR, Razafimamonjy MR, Velonirina HJS, Puchot N, Girod R & Bourgouin C. 2024. Colonization of Anopheles coustani, a neglected malaria vector in Madagascar. Parasite 31, 31.

All Tables

Table 1

Number of males and females used at each generation in the forced mating procedure and mean egg production by each mated female.

In the text
Table 2

Summary of the raw data obtained across the production of the first six generation of An. coustani starting with 31 wild gravid females producing 260 eggs.

In the text

All Figures

thumbnail Figure 1

Illustrations of An. coustani forced copulation procedure and developmental stages. Female An. coustani (A and F). Forced copulation: the male is hooked on a dissecting needle and presented to a cold anesthetized female; decapitated male (B); non-decapitated male (C). Fourth instar larvae (D). Pupa (E).
In the text
thumbnail Figure 2

Yield in eggs and adults from F1 to F6. The graph represents the number of eggs obtained at each generation (Purple line and left scale) and the subsequent number of adults (green line and right scale).
In the text
thumbnail Figure 3

Graphic representation of An. coustani adult emergence at each generation. (A) Individual graphs showing the pattern of emergence of F1 to F6. (B) Aggregated graphs highlighting the clear changing patterns of adult emergence from F1 to F6 both in mode and date of the peak of emergence. The scale of the abscissa axis corresponds to days till the first appearance of L1.
In the text

Current usage metrics show cumulative count of Article Views (full-text article views including HTML views, PDF and ePub downloads, according to the available data) and Abstracts Views on Vision4Press platform.

Data correspond to usage on the plateform after 2015. The current usage metrics is available 48-96 hours after online publication and is updated daily on week days.

Initial download of the metrics may take a while.