Hostname: page-component-78c5997874-xbtfd Total loading time: 0 Render date: 2024-11-10T16:11:51.016Z Has data issue: false hasContentIssue false
  • English
  • Français

A simple method for artificial infection of tsetse, Glossina Morsitans Morsitans larvae with the dna virus of G. Pallidipes

Published online by Cambridge University Press:  19 September 2011

Walter G. Z. O. Jura ,
Jan Zdarek  and
Leonard H. Otieno
Walter G. Z. O. Jura
Affiliation:
The International Centre of Insect Physiology and Ecology (ICIPE), P. O. Box 30772, Nairobi, Kenya
Jan Zdarek
Affiliation:
Institute of Organic Chemistry and Biochemistry, Czechoslovakia Academy of Sciences, Prague, Czechoslovakia
Leonard H. Otieno
Affiliation:
The International Centre of Insect Physiology and Ecology (ICIPE), P. O. Box 30772, Nairobi, Kenya
Get access

Abstract

Newly deposited Glossina morsitans morsitans larvae were chilled over ice and inoculated with 1 μ1 of either virus suspension derived from Glossina pallidipes salivary gland homogenate or sterile tsetse physiological saline. They were allowed to pupariate and then maintained at 25°C, 70% r.h. until soon after emergence when their salivary glands were examined for enlargement and presence of virus particles. Teneral G. m. morsitans which received the virus inoculum (n = 135) as larvae all became infected as revealed by gross hypertrophy of their salivary glands and ultrastructural manifestation of virus particles within the glandular epithelial cells and lumina. In the control group, which received the tsetse physiological saline (n = 91), only 1.1% of the flies showed the salivary gland enlargement, a level equivalent to the prevalence of virus infection normally detectable in the G. morsitans colony. This technique opens the way for testing the biocontrol potential of this virus. The DNA virus from G. pallidipes is clearly infective to G. morsitans morsitans, suggesting that the hypertrophied, chalky-white salivary glands, reported in various Glossina spp., are a manifestation of infection by one and the same virus.

Résumé

Les larves de Glossina morsitans morsitans nouvelement déposées étaient refroidies sur la glace et inoculées avec 1 μl soit d'une suspension de virus provenant d'homogenat de glandes salivaires de Glossina pallidipes ou d'une solution physiologique sterile de tsétsé. Les larves se sont transformées en pupes puis maintenues à une temperature de 25°C et une humidité relative de 70 % jusqu' à l'émergence lorsque leur glandes salivaires étaient examinées pour la présence des virus. Tous les 135 jeunes G. m. morsitans provenant des larves ayant été inoculées par des virus ont subit I'infection commerévelé par I'hypertrophie des glandes salivaires et la manifestation ultrastructurale des particules des virus au niveau des cellules épitheliales glandulaires et du lumen. Quant aux mouches tsétsé qui n'ont reçu que la solution physiologique (et utilisés comme contrôle), seulement 1,1% de 91 jeune G. m. morsitans ont montré I'agrandissement des glandes salivaires, un niveau équivalent à l'incidence de l'infection du virus normalement rencontrée dans la colonie de G. morsitans. Le succès de cette technique ouvre une voie pour tester, I'utilisation de ce virus comme agent potentiel dans la lutte biologique. Le virus ADN provenant de G. pallidipes sans doute infecte G. m. morsitans. Ceci suggère que les glandes salivaires hypertrophiées observées chez différentes espèces de Glossines sont des manifestations d'infection causées par le méme virus.

Keywords

Glossina morsitans morsitansGlossina pallidipesDNA viruslarvaeartificial infectionGlossina morsitans morsitansGlossina pallidipesvirus ADNlarvesinfection artificielle
Type
Research Artilces
Information
International Journal of Tropical Insect Science , Volume 14 , Issue 3 , June 1993 , pp. 383 - 387
Copyright
Copyright © ICIPE 1993

Access options

Get access to the full version of this content by using one of the access options below. (Log in options will check for institutional or personal access. Content may require purchase if you do not have access.)

References

REFERENCES

Brenner, S. and Home, R. W. (1959) A negative staining method for high resolution electron microscopy of viruses. Biochim. Biophys. Acta 34, 103–110.CrossRefGoogle ScholarPubMed
Burtt, E. (1945) Hypertrophied salivary glands in Glossina: evidence that G. pallidipes with this abnormality is particularly suited to trypanosome infection. Ann. Trop. Med. Parasitai. 39, 11–13.CrossRefGoogle Scholar
David, W. A. L. (1975) The status of viruses pathogenic for insects and mites. Annu. Rev. Entomol. 20, 97–117.CrossRefGoogle ScholarPubMed
Ellis, D. S. and Maudlin, I. (1987) Salivary gland hyperplasia in wild caught tsetse from Zimbabwe. Entomol. exp. appi. 45, 167–173.CrossRefGoogle Scholar
Engelmann, F. (1970) The Physiology of Insect Reproduction, 1st ed.Pergamon, Oxford.Google Scholar
Gee, J. D., (1976) Active transport of sodium by Malpighian tubules of the tsetse fly, Glossina morsitans. J. Exp. Biol. 64, 357–368.CrossRefGoogle Scholar
Glauert, A. M. and Glauert, R. H. (1958) Aralditeasan embedding medium for electron microscopy. J. Biophys. Biochem. Cytol. 4, 191–194.CrossRefGoogle ScholarPubMed
Gouteux, J. P. (1987) Prevalence of enlarged salivary glands in Glossina palpalis, G. pallicera, and G. nigrofusca (Diptera: Glossinidae) from the Vavoua area, Ivory Coast. J. Med. Entomol. 24, 268.CrossRefGoogle Scholar
Hagan, H. R. (1951) Embryology of the Viviparous Inscris. Ronald Press, New York.Google Scholar
Hayat, M. A. (1970) Principles and Techniques of Electron Microscopy: Biological Applications. Vol. 1, 1st ed.Van Nostrand Reinhold, New York, Cincinnati, Toronto, London, Melbourne.Google Scholar
Ignoffo, C. and Hink, W. F. (1971) Propagation of arthropod pathogens in living systems. In Microbial Control of Insects and Mites (Edited by Burgess, H. D. and Hussey, N. W.), pp. 541–580. Academic Press, New York.Google Scholar
Jaenson, T. G. T. (1978) Virus-like rods associated with sali vary gland hyperplasia in tsetse, Glossina pallidipes. Trans. R. Soc. Trop. Med. Hyg. 67, 234–238.CrossRefGoogle Scholar
Jaenson, T. G. T. (1986) Sex ratio distortion and reduced lifespan of Glossina pallidipes infected with the virus causing salivary gland hyperplasia. Entomol. exp. appi. 41, 265–271.CrossRefGoogle Scholar
Jura, W. G. Z. O. (1988) Morphological and functional changes associated with virus infection in male Glossina morsitans morsitans Westwood (Diptera: Glossinidae). In OAU/STRC, 1988, pp. 363–372.Google Scholar
Jura, W. G. Z. O., Odhiambo, T. R., Otieno, L. H. and Tabu, N. O. (1988) Gonadal lesions in virus-infected male and female tsetse, Glossina pallidipes (Diptera: Glossinidae). J. Invertebr. Pathol. 52, 1–8.CrossRefGoogle ScholarPubMed
Jura, W. G. Z. O., Otieno, L. H. and Chimtawi, M. M. B. (1989) Ultrastructural evidence fortrans-ovum transmission of the DNA virus of tsetse, Glossina pallidipes (Diptera: Glossinidae). Curr. Microbiol 18, 1–4.CrossRefGoogle Scholar
Knipling, E. F. (1955) Possibilities of insect control or eradication through the use of sexually sterile males. J. econ. Entomol. 48, 459–462.CrossRefGoogle Scholar
Nash, T. A. M., Jordan, A. M. and Trewern, M. A. (1971) Mass rearing of tsetse flies (Glossina spp.): Recent advances. In Sterility Principle for Insect Control or Eradication (Edited by Self, L. A.), pp. 99–110. Proceedings of the Symposium on the Sterility Principle for Insect Control or Eradication. IAEA, Vienna.Google Scholar
Odindo, M. O. (1988) Glossina pallidipes: Its potential for use in biological control of tsetse. Insect Sci. Applic. 9, 399–403.Google Scholar
Odindo, M. O., Sabwa, D. M., Amutalla, P. A. and Otieno, W. A. (1981) Preliminary tests on the transmission of virus-like particles to the tsetse Glossina pallidipes. Insect Sci. Applic. 2, 219–221.Google Scholar
Odindo, M. O., Payne, C. C., Crook, N. E. and Jarrett, P. (1986) Properties of a novel DNA virus from the tsetse fly, Glossina pallidipes. J. Gen. Virol. 67, 527–536.CrossRefGoogle ScholarPubMed
Otieno, L. H., Kokwaro, E. D., Chimtawi, M. and Onyango, P. ( 1980) Prevalence of enlarged salivary glands in wild populations of Glossina pallidipes in Kenya, with a note on the ultrastructure of the affected organ. J. lnvertebr. Pathol. 36, 113–118.CrossRefGoogle Scholar
Pal, R. and Whitten, M. J. (1974) The Use of Genetics in Insect Control 1st ed.Elsevier, North-Holland.Google Scholar
Reynolds, E. S. (1963) The use of lead citrate at high pH as an electron-opaque stain in electron microscopy. J. Cell Biol. 17, 208–212.CrossRefGoogle ScholarPubMed
Summers, M. D. (1977) Baculoviruses. In Atlas of Insect and Plant Viruses (Edited by Maramorosch, K.), pp. 3–28. Academic Press, New York.Google Scholar
Watson, M. L. (1958) Staining of tissue sections for electron microscopy with heavy metals. J. Biophys. Biochem. Cytol. 4, 475–478.CrossRefGoogle ScholarPubMed
Whitnall, A. B. M. (1934) The trypanosome infections of Glossina pallidipes in the Umfolosi Game Reserve, Zululand. Onderst. J. Vet. Anim. Ind. 11, 7–21.Google Scholar
Zdarek, J. and Denlinger, D. L. (1990) Wandering behaviour and pupariation in tsetse larvae. Physiol. Entomol. 16, 523–5.CrossRefGoogle Scholar