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Dramatic developmental changes in larval knockdown response enhance genetic sexing based on DDT resistance in Anopheles stephensi (Diptera: Culicidae)

Published online by Cambridge University Press:  09 March 2007

C.A. Malcolm  and
A.S. Robinson
C.A. Malcolm*
Affiliation:
School of Biological Sciences, Queen Mary, University of London, Mile End Road, London E1 4NS, UK
A.S. Robinson
Affiliation:
Entomology Unit, FAO/IAEA Agriculture and Biotechnology Laboratory, A-2444 Seibersdorf, Austria
*
*Fax: 44 (0)20 8983 0973 E-mail: c.a.malcolm@qmw.ac.uk
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Abstract

Genetic sexing systems based on a conditional lethal require good discrimination between the different phenotypes. DDT resistance in the early instars of Anopheles stephensi Liston is not a good candidate when based on mortality, but this study shows that the knockdown response gives exceptional discrimination between heterozygous resistant and homozygous susceptible individuals. One- and two-day-old larvae of the DlDDT strain showed high (417-fold) resistance to knockdown by DDT, but very low resistance to mortality (3.3-fold). This changes with the onset of the third instar, so that by the fourth instar, mortality resistance is high (108-fold) and knockdown resistance is low (6.5-fold). Susceptibility to DDT decreases from first to fourth instar in the susceptible strain by 443-fold for knockdown and 15-fold for mortality and in the resistant strain by 8.5-fold for knockdown and 491-fold for mortality. The DDT knockdown response in young larvae was successfully used to identify two Y-autosome translocations linked to the resistance gene, DDT. T(Y-3)69 and T(Y-3)72 gave recombination values between the translocation breakpoint and the DDT locus of 4.1 and 10.1 crossover units, respectively. T(Y-3)69 proved to be an adequate genetic sexing system for laboratory studies.

Type
Research Article
Information
Bulletin of Entomological Research , Volume 91 , Issue 6 , December 2001 , pp. 471 - 476
Copyright
Copyright © Cambridge University Press 2001

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References

Baker, R.H., Reisen, W.K., Sakai, R.K., Rathor, H.R., Raana, K., Azra, K. and Niaz, S. (1980) Anopheles culicifacies: mating behavior and competitiveness in nature of males carrying a complex chromosomal aberration. Annals of the Entomological Society of America 73, 581–588.CrossRefGoogle Scholar
Busvine, J.R. (1971) A critical review of the techniques for testing insecticides. 2nd edn. 344 pp. Commonwealth Agricultural Bureaux, Farnham Royal, UK.Google Scholar
Catteruccia, F., Nolan, T., Loukeris, T.G., Blass, C., Savakis, C., Kafatos, F.C. and Crisanti, A. (2000) Stable germline transformation of the malaria mosquito Anopheles stephensi. Nature 405, 959–962.CrossRefGoogle ScholarPubMed
Curtis, C.F. (1978) Genetic sex separation in Anopheles arabiensis and the production of sterile hybrids. Bulletin of the World Health Organization 56, 453–454.Google ScholarPubMed
Curtis, C.F. and Lines, J.D. (2000) Should DDT be banned by international treaty? Parasitology Today 16, 119–121.CrossRefGoogle ScholarPubMed
Curtis, C.F., Akiyama, J. and Davidson, G. (1976) A genetic sexing system in Anopheles gambiae species a. Mosquito News 36, 492–498.Google Scholar
Finney, D.J. (1971) Probit analysis. 3rd edn. 333 pp. Cambridge University Press.Google Scholar
Galvin, T.J. and Wyss, J.H. (1996) Screwworm eradication program in Central America. Annals of the New York Academy of Sciences 791, 233–240.CrossRefGoogle ScholarPubMed
Garms von, R. (1959) Die empfindlichkeit von Anopheles atroparvus und Anopheles stephensi (larven und imagines) gegenuber DDT in abhangigkeit von alter, ernahrung und populationsdichte. Zeitschrift für Tropenmedizin und Parasitologie 10, 426–441.Google Scholar
Glick, J.I. (1992) Illustrated key to the female Anopheles of southwestern Asia and Egypt (Diptera: Culicidae). Mosquito Systematics 24, 125–153.Google Scholar
Hati, A.K. (1997) Urban malaria vector biology. Indian Journal of Medical Research 106, 149–163.Google ScholarPubMed
IAEA (2000) Servicing immediate human needs. Resolution GC(44)/RES/24. 44th International Atomic Energy Agency General Conference 18–22 September 2000, Vienna, Austria.Google Scholar
Jayaraman, K.S. (1997) Consortium aims to revive sterile-mosquito project. Nature 389, 6.Google Scholar
Kaiser, P.E., Seawright, J.A., Dame, D.A. and Joslyn, D.J. (1978) Development of a genetic sexing for Anopheles albimanus. Journal of Economic Entomology 71, 766–771.CrossRefGoogle Scholar
Laven, H. (1967) Eradication of Culex pipiens fatigans through cytoplasmic incompatibility. Nature 216, 383–384.CrossRefGoogle ScholarPubMed
Laven, H., Cousserans, J. and Guille, G. (1972) Eradicating mosquitoes using translocations: a first field experiment. Nature 236, 456–457.CrossRefGoogle ScholarPubMed
Mahmood, F. and Sakai, R.K. (1984) Inversion polymorphisms in natural populations of Anopheles stephensi. Canadian Journal of Genetics and Cytology 26, 538–546.CrossRefGoogle ScholarPubMed
Malcolm, C.A. (1988) Genetic analysis of reduced susceptibility to permethrin and its relationship to DDT resistance in larvae of Anopheles stephensi. Medical and Veterinary Entomology 2, 37–46.CrossRefGoogle ScholarPubMed
Malcolm, C.A. (1990) Location of a gene conferring DDT resistance but no pyrethroid cross-resistance in larvae of Anopheles stephensi. Genetica 82, 51–55.CrossRefGoogle ScholarPubMed
Malcolm, C.A. and Mali, P. (1986) Genetic sexing of Anopheles stephensi with the larval morphological mutant Bl. Genetica 70, 37–42.CrossRefGoogle Scholar
Patterson, R.S., Weidhaas, D.E., Ford, H.R. and Lofgren, C.S. (1970) Suppression and elimination of an island population of Culex pipiens quinquefasciatus with sterile males. Science 168, 1368–1370.CrossRefGoogle ScholarPubMed
Rao, T.R. (1984) The anophelines of India. Malaria Research Centre (ICMR), Delhi.Google Scholar
Reichard, R. (1999) Case studies of emergency management of screwworm. Reviews in Science and Technology 18, 145–163.CrossRefGoogle ScholarPubMed
Robinson, A.S. (1986) Genetic sexing in Anopheles stephensi using dieldrin resistance. Journal of the American Mosquito Control Association 2, 93–95.Google ScholarPubMed
Robinson, A.S. and Pham, V.L. (1987) Cytological, linkage and insecticide studies on a genetic sexing line in Anopheles stephensi Liston. Heredity 58, 95–101.CrossRefGoogle ScholarPubMed
Robinson, A.S., Malcolm, C.A., Groenstijn, P. and Schelling, G. (1986) Breakpoint distribution of male linked translocations in Anopheles stephensi. Journal of Heredity 77, 394–398.CrossRefGoogle ScholarPubMed
Subbarao, S.K., Vasantha, K., Adak, T., Sharma, V.P. and Curtis, C.F. (1987) Egg-float ridge number in Anopheles stephensi: ecological variation and genetic analysis. Medical and Veterinary Entomology 1, 265–271.CrossRefGoogle ScholarPubMed
Tan, K.H. (2000) Area-wide control of insect pests integrating the sterile insects technique and related and other techniques. Proceedings of the FAO/IAEA International Conference Penang Malaysia May 29-June 2, 1998.Google Scholar
Vreysen, M.J.B., Saleh, K.M., Ali, M.Y., Abdulla, A.M., Zhu, Z.R., Juma, K.J., Dyck, V.A., Msangi, A.R., Mkonyi, P.A. and Feldmann, H.U. (2000) Glossina austeni (Diptera: Glossinidae) eradicated on the Island of Unguga, Zanzibar, using the sterile insect technique. Journal of Economic Entomology 93, 123–135.CrossRefGoogle ScholarPubMed
Weidhaas, D.E., Breeland, S.G., Lofgren, C.S., Dame, D.A. and Kaiser, R. (1974) Release of chemosterilized males for the control of Anopheles albimanus in El Salvador. IV. Dynamics of the test population. American Journal of Tropical Medicine and Hygiene 23, 298–308.CrossRefGoogle ScholarPubMed