Hostname: page-component-78c5997874-fbnjt Total loading time: 0 Render date: 2024-11-10T11:23:26.608Z Has data issue: false hasContentIssue false
  • English
  • Français

Fitness effects and transmission routes of a microsporidian parasite infecting Drosophila and its parasitoids

Published online by Cambridge University Press:  01 December 2005

P. H. FUTERMAN ,
S. J. LAYEN ,
M. L. KOTZEN ,
C. FRANZEN ,
A. R. KRAAIJEVELD  and
H. C. J. GODFRAY
P. H. FUTERMAN
Affiliation:
NERC Centre for Population Biology, Division of Biology, Imperial College London, Silwood Park Campus, Ascot, Berkshire SL5 7PY, UK
S. J. LAYEN
Affiliation:
NERC Centre for Population Biology, Division of Biology, Imperial College London, Silwood Park Campus, Ascot, Berkshire SL5 7PY, UK
M. L. KOTZEN
Affiliation:
NERC Centre for Population Biology, Division of Biology, Imperial College London, Silwood Park Campus, Ascot, Berkshire SL5 7PY, UK
C. FRANZEN
Affiliation:
Klinik und Poliklinik für Innere Medizin I, Universität Regensburg, Franz-Josef-Strauß Allee 11, D93042 Regensburg, Germany
A. R. KRAAIJEVELD
Affiliation:
NERC Centre for Population Biology, Division of Biology, Imperial College London, Silwood Park Campus, Ascot, Berkshire SL5 7PY, UK
H. C. J. GODFRAY
Affiliation:
NERC Centre for Population Biology, Division of Biology, Imperial College London, Silwood Park Campus, Ascot, Berkshire SL5 7PY, UK
Get access Rights & Permissions [Opens in a new window]

Abstract

A microsporidian infection was discovered in laboratory cultures of Drosophila species. Ultrastructural examination suggested it belonged to the poorly characterized species Tubulinosema kingi, and morphological and sequence data are presented. We explored how T. kingi affected the fitness of Drosophila melanogaster and D. subobscura, as well as the fitness of 2 of their parasitoids, Asobara tabida and Pachycrepoideus vindemiae. In Drosophila, infections caused changes in most of the traits we looked at that were associated with fitness, in particular causing a 34–55% reduction in early-life fecundity. Parasitoid fitness was affected more severely by infection than that of their hosts, with pupal mortality in particular increasing by 75–89%. We investigated the most important routes of transmission for T. kingi in a laboratory setting. Letting Drosophila larvae feed on medium contaminated with spores from infected dead flies resulted in 100% infection. Low levels of transmission (<10%) were found between larvae, and vertically between mothers and their offspring. Parasitoids developing in infected hosts all became infected, but infected adults were neither able to transmit the pathogen to their offspring nor to their offspring's Drosophila host, either directly, or via contamination of the ovipositor or other body parts. A field survey of Drosophila and their parasitoids in southern England revealed no natural infections. We discuss the potential importance of Microsporidia in parasitoid-host interactions, and for those working with Drosophila in the laboratory.

Keywords

Tubulinosema kingiDrosophilaAsobara tabidaPachycrepoideus vindemiaemicrosporidian parasitefitness coststransmission
Type
Research Article
Information
Parasitology , Volume 132 , Issue 4 , April 2006 , pp. 479 - 492
Copyright
2005 Cambridge University Press

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

Agnew, P., Bedhomme, S., Haussy, C. and Michalakis, Y. ( 1999). Age and size at maturity of the mosquito Culex pipiens infected by the microsporidian parasite Vavraia culicis. Proceedings of the Royal Society of London, B 266, 947–952.CrossRefGoogle Scholar
Van Alphen, J. J. M. and Thunnissen, I. ( 1983). Host selection and sex allocation by Pachycrepoideus vindemiae Rondani (Pteromalidae) as a facultative hyperparasitoid of Asobara tabida Nees (Braconidae; Alysiinae) and Leptopilina heterotoma (Thomson)(Cynipoidea; Eucoilidae). Netherlands Journal of Zoology 33, 497–514.Google Scholar
Armstrong, E. ( 1976). Transmission and infectivity studies on Nosema kingi in Drosophila willistoni and other Drosophilids. Zeitschrift für Parasitenkunde 50, 161–165.CrossRefGoogle Scholar
Armstrong, E. ( 1977). Transmission of Nosema kingi to offspring of Drosophila willistoni during copulation. Zeitschrift für Parasitenkunde 53, 311–315.CrossRefGoogle Scholar
Armstrong, E. and Bass, L. ( 1989 a). Effects of Nosema kingi on the development and weight of adult Drosophila melanogaster (OR-R-Strain). Journal of Invertebrate Pathology 53, 102–106.Google Scholar
Armstrong, E. and Bass, L. ( 1989 b). Nosema kingi: Effects on fecundity, fertility and longevity of Drosophila melanogaster. Journal of Experimental Zoology 250, 82–86.Google Scholar
Armstrong, E., Bass, L., Staker, K. and Harrell, L. ( 1986). A comparison of the biology of a Nosema in Drosophila melanogaster to Nosema kingi in Drosophila willistoni. Journal of Invertebrate Pathology 48, 124–126.CrossRefGoogle Scholar
Baker, M. D., Vossbrinck, C. R., Maddox, J. V. and Undeen, A. H. ( 1994). Phylogenetic relationships among Vairimorpha and Nosema species (Microspora) based on ribosomal RNA sequence data. Journal of Invertebrate Pathology 64, 100–106.CrossRefGoogle Scholar
Becnel, J. J. and Andreadis, T. G. ( 1999). Microsporidia in insects. In The Microsporidia and Microsporidiosis ( ed. Wittner, M. and Weiss, L. M.), pp. 447–501. ASM Press, Washington DC.CrossRef
Becnel, J. J., Garcia, J. J. and Johnson, M. A. ( 1995). Edhazardia aedis (Microspora: Culicosporidae) effects on the reproductive capacity of Aedes aegypti (Diptera: Culicidae). Journal of Medical Entomology 32, 549–553.CrossRefGoogle Scholar
Belshaw, R., Fitton, M., Herniou, E., Gimeno, C. and Quicke, D. L. J. ( 1998). A phylogenetic reconstruction of the Ichneumonoidea (Hymenoptera) based on the D2 variable region of 28S ribosomal RNA. Systematic Entomology 23, 109–123.CrossRefGoogle Scholar
Boheene, C. K., Geden, C. J. and Becnel, J. J. ( 2003). Development of microsporidia-infected Muscidifurax raptor (Hymenoptera: Pteromalidae) at different temperatures. Biological Control 26, 1–7.CrossRefGoogle Scholar
Brooks, W. M. ( 1993). Host-parasitoid-pathogen interactions. In Parasites and Pathogens of Insects: Pathogens ( ed. Beckage, N. E., Thompson, S. N. and Federici, B. A.), pp. 231–272. Academic Press, San Diego.
Burnett, R. G. and King, R. C. ( 1962). Observations on a microsporidian parasite of Drosophila willistoni Sturtevant. Journal of Insect Pathology 4, 104–112.Google Scholar
Canning, E. U., Refardt, D., Vossbrinck, C. R., Okamura, B. and Curry, A. ( 2002). New diplokaryotic microsporidia (Phylum Microsporidia) from freshwater bryozoans (Bryozoa, Phylactolaemata). European Journal of Protistology 38, 247–265.CrossRefGoogle Scholar
Clark, M. E., Anderson, C., Cande, J. and Karr, T. ( 2005). Widespread prevalence of Wolbachia in laboratory stocks and implications for Drosophila research. Genetics 170, 1667–1675.CrossRefGoogle Scholar
Dunn, A. M., Terry, R. S. and Smith, J. E. ( 2001). Transovarial transmission in the microsporidia. Advances in Parasitology 48, 57–100.CrossRefGoogle Scholar
Franzen, C. ( 2004). Microsporidia: how can they invade other cells? Trends in Parasitology 20, 275–279.Google Scholar
Franzen, C., Fischer, S., Schroeder, J., Schölmerich, J. and Schneuwly, S. ( 2005). Morphological and molecular investigations of Tubulinosema ratisbonensis gen. nov., sp. nov. (Microsporidia: Tubulinosematidae fam. nov.), a parasite infecting a laboratory colony of Drosophila melanogaster (Diptera: Drosophilidae). Journal of Eukaryotic Microbiology 52, 141–152.Google Scholar
Gatehouse, H. S. and Malone, L. A. ( 1998). The ribosomal RNA gene region of Nosema apis (Microspora): DNA sequence for small and large subunit rRNA genes and evidence for a tandem repeat unit size. Journal of Invertebrate Pathology 71, 97–105.CrossRefGoogle Scholar
Geden, C. J., Ferreira De Almeida, M. A. and Pires do Prado, A. ( 2003). Effects of Nosema disease on fitness of the parasitoid Tachinaephagus zealandicus (Hymenoptera: Encyrtidae). Environmental Entomology 32, 1139–1145.CrossRefGoogle Scholar
Geden, C. J., Long, S. J., Rutz, D. A. and Becnel, J. J. ( 1995). Nosema disease of the parasitoid Muscidifurax raptor (Hymenoptera: Pteromalidae): prevalence, patterns of transmission, management, and impact. Biological Control 5, 607–614.CrossRefGoogle Scholar
Hirt, R. P., Logsdon Jr, J. M., Healy, B., Dorey, M. W., Doolittle, W. F. and Embley, T. M. ( 1999). Microsporidia are related to fungi: evidence from the largest subunit of RNA polymerase II and other proteins. Proceedings of the National Academy of Sciences, USA 96, 580–585.CrossRefGoogle Scholar
Hoffmann, J. A. and Reichhart, J.-M. ( 2002). Drosophila innate immunity: an evolutionary perspective. Nature Immunology 3, 121–126.CrossRefGoogle Scholar
Jaenike, J. ( 1995). Interactions between mycophagous Drosophila and their nematode parasites: from physiological to community ecology. Oikos 72, 235–244.CrossRefGoogle Scholar
Kraaijeveld, A. R. and van Alphen, J. J. M. ( 1993). Successful invasion of North America by two Palearctic Drosophila species (Diptera: Drosophilidae): a matter of immunity to local parasitoids? Netherlands Journal of Zoology 43, 235–241.Google Scholar
Kraaijeveld, A. R., van Alphen, J. J. M. and Godfray, H. C. J. ( 1998). The coevolution of host resistance and parasitoid virulence. Parasitology 116, S29–S45.CrossRefGoogle Scholar
Kraaijeveld, A. R. and Godfray, H. C. J. ( 2003). Potential life history costs of parasitoid avoidance in Drosophila melanogaster. Evolutionary Ecology Research 5, 1251–1261.Google Scholar
Kraaijeveld, A. R. and Van Der Wel, N. N. ( 1994). Geographic variation in reproductive success of the parasitoid Asobara tabida in larvae of several Drosophila species. Ecological Entomology 19, 221–229.CrossRefGoogle Scholar
Kramer, J. P. ( 1964). Nosema kingi sp. n., a microsporidian from Drosophila willistoni Sturtevant, and its infectivity for other muscoids. Journal of Insect Pathology 6, 491–499.Google Scholar
Kramer, J. P. ( 1973). Susceptibility of sixteen species of muscoid flies to the microsporidean parasite Octosporea muscaedomesticae. Journal of the New York Entomological Society 81, 50–53.Google Scholar
Müller, A., Trammer, T., Chioralia, G., Seitz, H. M., Diehl, V. and Franzen, C. ( 2000). Ribosomal RNA of Nosema algerae and phylogenetic relationship to other microsporidia. Parasitology Research 86, 18–23.CrossRefGoogle Scholar
Müller, C. B. and Brodeur, J. ( 2002). Intraguild predation in biological control and conservation biology. Biological Control 25, 216–223.CrossRefGoogle Scholar
Nappi, A. J. ( 1981). Cellular immune response of Drosophila melanogaster against Asobara tabida. Parasitology 83, 319–324.CrossRefGoogle Scholar
Pell, J. and Canning, E. ( 1993). Light microscope and ultrastructural observations of a microsporidian parasite of Mesocyclops rarus (Copepoda: Cyclopoda) in Tanzania. Journal of Invertebrate Pathology 61, 275–280.CrossRefGoogle Scholar
Polis, G. A., Myers, C. A. and Holt, R. D. ( 1989). The ecology and evolution of intraguild predation: potential competitors that eat each other. Annual Review of Ecology and Systematics 20, 297–330.CrossRefGoogle Scholar
Rosenheim, J. A., Kaya, H. K., Ehler, L. E., Marois, J. J. and Jaffee, B. A. ( 1995). Intraguild predation among biological-control agents: theory and evidence. Biological Control 5, 303–335.CrossRefGoogle Scholar
Roxström-Lindquist, K., Terenius, O. and Faye, I. ( 2004). Parasite-specific immune response in adult Drosophila melanogaster: a genomic study. EMBO Reports 5, 207–212.CrossRefGoogle Scholar
Schuld, M., Madel, G. and Schmuck, R. ( 1999). Impact of Vairimorpha sp. (Microsporidia: Burnellidae) on Trichogramma chilonis (Hymenoptera, Trichogrammatidae), a hymenopteran parasitoid of the cabbage moth, Plutella xylostella (Lepidoptera, Yponomeutidae). Journal of Invertebrate Pathology 74, 120–126.Google Scholar
Street, D. A. and Henry, J. E. ( 1993). Ultrastructural study of Nosema acridophagus Henry (Microspora: Nosematidae) from grasshopper. Parasitology Research 79, 173–177.CrossRefGoogle Scholar
Terry, R. S., Smith, J. E., Sharpe, R. G., Rigaud, T., Littlewood, D. T. J., Ironside, J. E., Rollinson, D., Bouchin, D., Macneil, C., Dick, J. T. A. and Dunn, A. M. ( 2004). Widespread vertical transmission and associated host sex-ratio distortion within the eukaryotic phylum Microspora. Proceedings of the Royal Society of London, B 271, 1783–1789.CrossRefGoogle Scholar
Tzou, P., De Gregorio, E. and Lemaitre, B. ( 2002). How Drosophila combats microbial infection: a model to study innate immunity and host-pathogen interactions. Current Opinion in Microbiology 5, 102–110.CrossRefGoogle Scholar
Undeen, A. ( 1997). Microsporidia (Protozoa): A Handbook of Biology and Research Techniques. http://pearl.agcomm.okstate.edu/scsb387/content.htm
Vossbrinck, C. R., Baker, M. D., Didier, E. S., Debrunner-Vossbrinck, B. A. and Shadduck, J. A. ( 1993). Ribosomal DNA sequences of Encephalitozoon hellem and Encephalitozoon cuniculi: species identification and phylogenetic construction. Journal of Eukaryotic Microbiology 40, 354–362.CrossRefGoogle Scholar
Williams, B. A. P., Hirt, R. P., Lucocq, J. M. and Embley, T. M. ( 2002). A mitochondrial remnant in the microsporidian Trachipleistophora hominis. Nature, London 418, 865–869.CrossRefGoogle Scholar
Wilson, G. G. ( 1974). Effects of larval age at innoculation, and dosage of microsporidian (Nosema fumiferanae) spores, on mortality of spruce budworm (Choristoneura fumiferana). Canadian Journal of Zoology 52, 993–996.CrossRefGoogle Scholar
Wittner, M. and Weiss, L. M. ( 1999). The Microsporidia and Microsporidiosis. ASM Press, Washington DC.
Supplementary material: PDF

Futerman supplementary material

Supplementary figure S1

Download Futerman supplementary material(PDF)
PDF 453.5 KB