Efficient method for establishing F1 progeny from wild populations of Anopheles mosquitoes
© The Author(s) 2017
Received: 27 October 2016
Accepted: 4 January 2017
Published: 9 January 2017
The changing malaria situation in Madagascar requires additional knowledge on the physiology and behaviour of local mosquito vectors. However, the absence of established colonies for several anopheline species present in Madagascar constitutes a limiting factor. To avoid labour intensive work and uncertainty for success of establishing Anopheles colonies from Malagasy species, field collections of blood-fed females and in-tube forced oviposition were combined to reliably produce large numbers of F1 progeny.
Blood-fed females were captured in zebu stables or open zebu parks. Oviposition was induced by enclosing gravid females in eppendorf tubes as initially described for Anopheles funestus. The effect of cold anaesthesia on inducing in-tube forced oviposition and on egg yield was assessed for five Anopheles species, namely Anopheles coustani, An. funestus, Anopheles mascarensis, Anopheles arabiensis and Anopheles squamosus. The production of eggs from in-tube forced oviposition and standard egg laying in cages was compared.
For the five anopheline species studied, the in-tube forced oviposition method had different efficacy ranging from 35.6 to 71.1% females willing to lay eggs in tubes. Interestingly, prior anaesthesia increased significantly the proportion of ovipositing females for An. mascarensis. Prior anaesthesia has a marginal effect on the number of eggs produced. However, the overall yield in eggs collected using the in-tube forced oviposition method largely exceeds the number of eggs that can be produced by females free to oviposit in cages.
The efficiency of the method allowed the production of F1 progeny in numbers sufficiently large for developing detailed analyses of the five species tested, including behavioural studies, insecticide resistance assessment and molecular characterization, as well as vector competence studies. It should be applicable to other anopheline species difficult to colonize.
KeywordsWild anopheles Malaria Madagascar Forced-oviposition F1 production
Mosquitoes constitute a large group of arthropod vectors of pathogens. Among them the Anopheles genus includes all known vectors of malaria parasites infecting mammals. One feature of Anopheles mosquitoes as malaria vectors is that they exhibit a highly specific geographic distribution . As a consequence, each malaria endemic country harbours its own set of major and/or potential secondary anopheline vectors. This is in sharp contrast with the large geographic distribution of arbovirus culicine vectors, exemplified by Aedes aegypti, which can be found in most tropical areas, and Aedes albopictus, which expanded its distribution range from tropical to more continental areas in the last two decades .
Among the ~60 known human malaria vectors, very few have been easily colonized, limiting in-depth study of their biological characteristics . Over many years scientists have developed strategies for colonizing several anopheline species in the laboratory with successes and failures. A common feature of several anopheline mosquitoes is eurygamy, which hampers efficient mating in a confined environment. To counteract the absence of free mating in cages, the technique of forced mating was developed and shown to work for several species [4, 5], Anopheles dirus being among the best-known examples. This mating technique allowed the establishment of optimized production of some anopheline species at the cost of time. Alternatively, long-term efforts led to the selection of individuals that accept mating in a confined environment and to the establishment of so-called free-mating colonies . For other mosquito species introducing tricks such as a stroboscopic light has shown to be effective in inducing mating [7–9]. However, these tricks turn out not to be efficient for establishing colonies from every single anopheline species. Even in situations where rearing success has been reported, the reasons for success are often obscure and not repeatable outside the successful laboratory. Establishment of Anopheles funestus colonies was such an example [10, 11].
In Madagascar, three anopheline species (Anopheles gambiae, Anopheles arabiensis, An. funestus) are considered major vector species of malaria. Anopheles mascarensis, an endemic species, and Anopheles merus have been identified as secondary vectors of local importance [12–17]. Due to their high abundance, two other species, Anopheles squamosus and Anopheles coustani, are suspected to be involved in residual malaria transmission, in places where the major vectors are of low abundance. Indeed, An. coustani has recently been described as a potential secondary malaria vector in Madagascar . Whereas An. gambiae and An. arabiensis can be easily colonized, none of the other species have been successfully colonized yet, despite reports on existing current or past colonies of these species [10, 11, 19].
The malaria transmission pattern is currently changing in Madagascar with an epidemic situation for the past few years . Among several other causes, there is suspicion of increased transmission by An. mascarensis and An. coustani. The changing malaria situation in Madagascar advocates for the urgent need of gaining additional knowledge on these two species. To avoid labour intensive work and uncertainty for success of establishing Anopheles colonies from Malagasy species including An. mascarensis and An. coustani, field collection of blood-fed females was combined with in-tube forced oviposition to reliably produce F1 progenies. The efficiency of the method allowed the production of F1 in numbers sufficiently large to permit detailed analyses of those species, including behavioural studies, insecticide resistance assessment and molecular characterization, as well as vector competence studies. The in-tube forced oviposition method was first reported by Morgan et al. , successfully producing F1 from An. funestus and later included into the MR4 manual 2014 .
To the original method an additional step was introduced, which turns out to be highly efficient for some species for increasing the number of females willing to lay eggs and subsequently increasing the number of the F1 progeny. Herein is presented a detailed analysis of the benefit of this strategy for producing F1 from four Anopheles species encountered throughout Africa (An. arabiensis, An. funestus, An. coustani and An. squamosus) and one Malagasy species An. mascarensis. The demonstration that this method, initially developed for An. funestus, is easily applicable to additional anopheline species should facilitate a better characterization of malaria vectors from different countries, for which no sustainable colonies exist yet.
Mosquito collection methods
Eggs were transferred into a rearing pan containing dechlorinated tap water. L1 and L2 larvae were fed with Tetramin™ baby fish food and L3 and L4 larvae either with cat or mice finely ground food. Water from each larval pan was changed every other day. The F1 adults were mixed in cages for subsequent experiments.
Chi square test was used for comparing the proportion of female mosquitoes that laid eggs by in-tube forced oviposition with or without prior cold-anaesthesia. Wilcoxon test was used for comparing egg numbers produced by those females. These tests were performed using R Core Team (2013).
Results and discussion
Blood-fed Anopheles females (n = 1026) were captured over six sampling periods from April 2014 until April 2016 in three different locations. Most specimens were trapped using a large mosquito net set up on one side of a zebu park (see Fig. 2 and “Methods” section) from 7 pm till 2 am and collected using mouth aspirators. Resting mosquitoes were also collected within zebu stables in the morning. For each harvest, all captured mosquitoes were placed inside a single large cage before being morphologically identified and sorted on the next day. Fully fed females from each species were maintained in the insectary for an additional three days until fully gravid, with free access to only sugar solution. At that stage, females were individually placed into 1.5 ml Eppendorf ® tubes prepared as described in “Methods” section and observed over 5 days to determine their ability to lay eggs under these conditions. To control the gravid state of the females, a proportion of females were anaesthetized on ice and observed under a binocular microscope before being placed in tubes.
Estimation of egg production for 100 females by in-tube forced oviposition
Proportion of females laying eggs in tubes (%)
Egg number per female laying eggs
Yield for 100 females
The generation time from egg to adult was around 15–17 days for An. funestus, confirming the observation of Cuamba et al. . It was similar for An. arabiensis and An. mascarensis, but in the range of 21 days for both An. coustani and An. squamosus.
Egg collection increase by in-tube forced oviposition
Mean egg number per female
Increase % in egg yield
In-tube forced oviposition
In-cage free oviposition
This work demonstrates that the in-tube forced oviposition method initially developed for An. funestus applies efficiently to other anopheline species. For An. funestus the same efficiency as the one reported by Morgan et al. was observed . Furthermore, inclusion of a cold anaesthesia step to An. mascarensis increases significantly the number of females that oviposit in tubes. Although this does not significantly apply to the other mosquito species tested, this might be worth trying for other anopheline species. Overall the comparison of egg yield between in-tube forced oviposition and free oviposition in cages clearly shows that establishing large F1 populations can be easily achieved using the in-tube forced oviposition method and should be favoured for conducting detailed analyses on behavioural studies, insecticide resistance assessment and molecular characterization or vector competence studies.
CB conceived and designed the experiments. TNN, LA and CB performed the experiments. TNN, SB and CB analysed the data. SB contributed reagents/materials/logistics. TNN and CB wrote the paper. All authors read and approved the final manuscript.
We wish to thank Gael Millot (C3BI-Institut Pasteur) for his help with statistical analyses, Christophe Rogier for his constant support, the technical support of the Entomology Unit from the Institut Pasteur de Madagascar, and the farmers at the sites of mosquito collections. We are grateful to Richard Paul (GFMI-Institut Pasteur) for manuscript editing.
The authors declare that they have no competing interests.
Availability of data and materials
Most data generated or analysed during this study are included in this published article. Datasets for egg counts used and analysed during the current study are available from the corresponding author on reasonable request.
The study was supported by Institut Pasteur de Madagascar, and financial support to CB from the Institut Pasteur International Network-IPIN.
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- Sinka M, Bangs M, Manguin S, Rubio-Palis Y, Chareonviriyaphap T, Coetzee M, et al. A global map of dominant malaria vectors. Parasit Vectors. 2012;5:69.View ArticlePubMedPubMed CentralGoogle Scholar
- Kraemer MU, Sinka ME, Duda KA, Mylne AQ, Shearer FM, Barker CM, et al. The global distribution of the arbovirus vectors Aedes aegypti and Ae. albopictus. Elife. 2015;4:e08347.View ArticlePubMedPubMed CentralGoogle Scholar
- Benedict M, Knols B, Bossin H, Howell P, Mialhe E, Caceres C, Robinson A. Colonisation and mass rearing: learning from others. Malar J. 2009;8:S4.View ArticlePubMedPubMed CentralGoogle Scholar
- Baker RH. Mating problems as related to the establishment and maintenance of laboratory colonies of mosquitos. Bull World Health Organ. 1964;31:467–8.PubMedPubMed CentralGoogle Scholar
- Bryan JH, Southgate BA. Studies of forced mating techniques on anopheline mosquitos. Mosq News. 1978;38:338–42.Google Scholar
- Bates M. The laboratory colonization of Anopheles darlingi. J Natl Malar Soc. 1947;6:155–8.PubMedGoogle Scholar
- Lardeux F, Quispe V, Tejerina R, Rodriguez R, Torrez L, Bouchite B, et al. Laboratory colonization of Anopheles pseudopunctipennis (Diptera: Culicidae) without forced mating. C R Biol. 2007;330:571–5.View ArticlePubMedGoogle Scholar
- Moreno M, Tong C, Guzman M, Chuquiyauri R, Llanos-Cuentas A, Rodriguez H, et al. Infection of laboratory-colonized Anopheles darlingi mosquitoes by Plasmodium vivax. Am J Trop Med Hyg. 2014;90:612–6.View ArticlePubMedPubMed CentralGoogle Scholar
- Villarreal-Trevino C, Vasquez GM, Lopez-Sifuentes VM, Escobedo-Vargas K, Huayanay-Repetto A, Linton YM, et al. Establishment of a free-mating, long-standing and highly productive laboratory colony of Anopheles darlingi from the Peruvian Amazon. Malar J. 2015;14:227.View ArticlePubMedPubMed CentralGoogle Scholar
- Hunt RH, Brooke BD, Pillay C, Koekemoer LL, Coetzee M. Laboratory selection for and characteristics of pyrethroid resistance in the malaria vector Anopheles funestus. Med Vet Entomol. 2005;19:271–5.View ArticlePubMedGoogle Scholar
- Service MW, Oguamah D. Colonization of Anopheles funestus. Nature. 1958;181:1225.View ArticleGoogle Scholar
- Fontenille D, Campbell GH. Is Anopheles mascarensis a new malaria vector in Madagascar? Am J Trop Med Hyg. 1992;46:28–30.PubMedGoogle Scholar
- Marrama L, Laventure S, Rabarison P, Roux J. [Anopheles mascarensis (De Meillon, 1947): main vector of malaria in the region of Fort-Dauphin (south–east of Madagascar)](in French). Bull Soc Pathol Exot. 1999;92:136–8.PubMedGoogle Scholar
- Le Goff G, Randimby FM, Rajaonarivelo V, Laganier R, Duchemin JB, Robert V, et al. [Anopheles mascarensis of Meillon 1947, a malaria vector in the middle west of Madagascar?] (in French). Arch Inst Pasteur Madagascar. 2003;69:57–62.PubMedGoogle Scholar
- Pock Tsy JM, Duchemin JB, Marrama L, Rabarison P, Le Goff G, Rajaonarivelo V, et al. Distribution of the species of the Anopheles gambiae complex and first evidence of Anopheles merus as a malaria vector in Madagascar. Malar J. 2003;2:33.View ArticlePubMedPubMed CentralGoogle Scholar
- Marrama L, Jambou R, Rakotoarivony I, Leong Pock Tsi JM, Duchemin JB, Laventure S, et al. Malaria transmission in Southern Madagascar: influence of the environment and hydro-agricultural works in sub-arid and humid regions. Part 1. Entomological investigations. Acta Trop. 2004;89:193–203.View ArticlePubMedGoogle Scholar
- Andrianaivolambo L, Domarle O, Randrianarivelojosia M, Ratovonjato J, Le Goff G, Talman A, et al. Anthropophilic mosquitoes and malaria transmission in the eastern foothills of the central highlands of Madagascar. Acta Trop. 2010;116:240–5.View ArticlePubMedGoogle Scholar
- Nepomichene TN, Tata E, Boyer S. Malaria case in Madagascar, probable implication of a new vector Anopheles coustani. Malar J. 2015;14:475.View ArticlePubMedPubMed CentralGoogle Scholar
- Gerberg EJ. Manual for mosquito rearing and experimental techniques. Lake Charles: American Mosquito Control Association; 1994.Google Scholar
- Kesteman T, Rafalimanantsoa SA, Razafimandimby H, Rasamimanana HH, Raharimanga V, Ramarosandratana B, et al. Multiple causes of an unexpected malaria outbreak in a high-transmission area in Madagascar. Malar J. 2016;15:57.View ArticlePubMedPubMed CentralGoogle Scholar
- Morgan JC, Irving H, Okedi LM, Steven A, Wondji CS. Pyrethroid resistance in an Anopheles funestus population from Uganda. PLoS ONE. 2010;5:e11872.View ArticlePubMedPubMed CentralGoogle Scholar
- Methods in anopheles research. https://www.beiresources.org/portals/2/MR4/MR4_Publications/Methods%20in%20Anopheles%20Research%202014/2014MethodsinAnophelesResearchManualFullVersionv2tso.pdf Accessed Sept 2016.
- Raharimalala FN. Role des moustiques Culicidae, de leurs communautes microbiennes, et des reservoirs vertebres, dans la transmission d’arbovirus a Madagascar. Université de Lyon (France) et Université d’Antananarivo (Madagascar). 2011.
- Grjébine A. Insectes diptères Culicidae Anophelinae. Paris: ORSTOM, CNRS; 1966.Google Scholar
- Cuamba N, Morgan JC, Irving H, Steven A, Wondji CS. High level of pyrethroid resistance in an Anopheles funestus population of the Chokwe District in Mozambique. PLoS ONE. 2010;5:e11010.View ArticlePubMedPubMed CentralGoogle Scholar
- VassarStats: Website for statistical computation. http://vassarstats.net/index.html. Accessed Sept 2016.
- Newcombe RG. Two-sided confidence intervals for the single proportion: comparison of seven methods. Stat Med. 1998;17:857–72.View ArticlePubMedGoogle Scholar