Letter | Published:

Male-killing toxin in a bacterial symbiont of Drosophila

Naturevolume 557pages252255 (2018) | Download Citation


Several lineages of symbiotic bacteria in insects selfishly manipulate host reproduction to spread in a population1, often by distorting host sex ratios. Spiroplasma poulsonii2,3 is a helical and motile, Gram-positive symbiotic bacterium that resides in a wide range of Drosophila species4. A notable feature of S. poulsonii is male killing, whereby the sons of infected female hosts are selectively killed during development1,2. Although male killing caused by S. poulsonii has been studied since the 1950s, its underlying mechanism is unknown. Here we identify an S. poulsonii protein, designated Spaid, whose expression induces male killing. Overexpression of Spaid in D. melanogaster kills males but not females, and induces massive apoptosis and neural defects, recapitulating the pathology observed in S. poulsonii-infected male embryos5,6,7,8,9,10,11. Our data suggest that Spaid targets the dosage compensation machinery on the male X chromosome to mediate its effects. Spaid contains ankyrin repeats and a deubiquitinase domain, which are required for its subcellular localization and activity. Moreover, we found a laboratory mutant strain of S. poulsonii with reduced male-killing ability and a large deletion in the spaid locus. Our study has uncovered a bacterial protein that affects host cellular machinery in a sex-specific way, which is likely to be the long-searched-for factor responsible for S. poulsonii-induced male killing.

Access optionsAccess options

Rent or Buy article

Get time limited or full article access on ReadCube.


All prices are NET prices.

Additional information

Publisher’s note: Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.


  1. 1.

    Hurst, G. D. D. & Frost, C. L. Reproductive parasitism: maternally inherited symbionts in a biparental world. Cold Spring Harb. Perspect. Biol. 7, a017699 (2015).

  2. 2.

    Williamson, D. L. & Poulson, D. F. in The Mycoplasmas, Volume III: Plant and Insect Mycoplasmas (eds Whitcomb R.F. & Tully J.G.) Ch. 6, 175–208 (Academic, New York, 1979).

  3. 3.

    Williamson, D. L. et al. Spiroplasma poulsonii sp. nov., a new species associated with male-lethality in Drosophila willistoni, a neotropical species of fruit fly. Int. J. Syst. Bacteriol. 49, 611–618 (1999).

  4. 4.

    Haselkorn, T. S. The Spiroplasma heritable bacterial endosymbiont of Drosophila. Fly (Austin) 4, 80–87 (2010).

  5. 5.

    Tsuchiyama-Omura, S., Sakaguchi, B., Koga, K. & Poulson, D. F. Morphological features of embryogenesis in Drosophila melanogaster infected with a male-killing. Spiroplasma. Zool. Sci. 5, 375–383 (1988).

  6. 6.

    Veneti, Z., Bentley, J. K., Koana, T., Braig, H. R. & Hurst, G. D. D. A functional dosage compensation complex required for male killing in Drosophila. Science 307, 1461–1463 (2005).

  7. 7.

    Bentley, J. K., Veneti, Z., Heraty, J. & Hurst, G. D. D. The pathology of embryo death caused by the male-killing Spiroplasma bacterium in Drosophila nebulosa. BMC Biol. 5, 9 (2007).

  8. 8.

    Martin, J., Chong, T. & Ferree, P. M. Male killing Spiroplasma preferentially disrupts neural development in the Drosophila melanogaster embryo. PLoS One 8, e79368 (2013).

  9. 9.

    Harumoto, T., Anbutsu, H. & Fukatsu, T. Male-killing Spiroplasma induces sex-specific cell death via host apoptotic pathway. PLoS Pathog. 10, e1003956 (2014).

  10. 10.

    Cheng, B., Kuppanda, N., Aldrich, J. C., Akbari, O. S. & Ferree, P. M. Male-killing Spiroplasma alters behavior of the dosage compensation complex during Drosophila melanogaster embryogenesis. Curr. Biol. 26, 1339–1345 (2016).

  11. 11.

    Harumoto, T., Anbutsu, H., Lemaitre, B. & Fukatsu, T. Male-killing symbiont damages host’s dosage-compensated sex chromosome to induce embryonic apoptosis. Nat. Commun. 7, 12781 (2016).

  12. 12.

    Oishi, K. Spirochaete-mediated abnormal sex-ratio (SR) condition in Drosophila: a second virus associated with spirochaetes and its use in the study of the SR condition. Genet. Res. 18, 45–56 (1971).

  13. 13.

    Makarova, K. S., Aravind, L. & Koonin, E. V. A novel superfamily of predicted cysteine proteases from eukaryotes, viruses and Chlamydia pneumoniae. Trends Biochem. Sci. 25, 50–52 (2000).

  14. 14.

    Al-Khodor, S., Price, C. T., Kalia, A. & Abu Kwaik, Y. Functional diversity of ankyrin repeats in microbial proteins. Trends Microbiol. 18, 132–139 (2010).

  15. 15.

    Masson, F., Calderon Copete, S., Schüpfer, F., Garcia-Arraez, G. & Lemaitre, B. In vitro culture of the insect endosymbiont Spiroplasma poulsonii highlights bacterial genes involved in host-symbiont interaction. MBio 9, e00024-18 (2018).

  16. 16.

    Paredes, J. C. et al. Genome sequence of the Drosophila melanogaster male-killing Spiroplasma strain MSRO endosymbiont. MBio 6, e02437–14 (2015).

  17. 17.

    Campos, A. R., Rosen, D. R., Robinow, S. N. & White, K. Molecular analysis of the locus elav in Drosophila melanogaster: a gene whose embryonic expression is neural specific. EMBO J. 6, 425–431 (1987).

  18. 18.

    Salz, H. K. & Erickson, J. W. Sex determination in Drosophila: The view from the top. Fly (Austin) 4, 60–70 (2010).

  19. 19.

    Lucchesi, J. C. & Kuroda, M. I. Dosage compensation in Drosophila. Cold Spring Harb. Perspect. Biol. 7, a019398 (2015).

  20. 20.

    Madigan, J. P., Chotkowski, H. L. & Glaser, R. L. DNA double-strand break-induced phosphorylation of Drosophila histone variant H2Av helps prevent radiation-induced apoptosis. Nucleic Acids Res. 30, 3698–3705 (2002).

  21. 21.

    Siozios, S. et al. The diversity and evolution of Wolbachia ankyrin repeat domain genes. PLoS ONE 8, e55390 (2013).

  22. 22.

    Jernigan, K. K. & Bordenstein, S. R. Ankyrin domains across the Tree of Life. PeerJ 2, e264 (2014).

  23. 23.

    Wu, M. et al. Phylogenomics of the reproductive parasite Wolbachia pipientis wMel: a streamlined genome overrun by mobile genetic elements. PLoS Biol. 2, e69 (2004).

  24. 24.

    Yamada, R., Iturbe-Ormaetxe, I., Brownlie, J. C. & O’Neill, S. L. Functional test of the influence of Wolbachia genes on cytoplasmic incompatibility expression in Drosophila melanogaster. Insect Mol. Biol. 20, 75–85 (2011).

  25. 25.

    Beckmann, J. F. & Fallon, A. M. Detection of the Wolbachia protein WPIP0282 in mosquito spermathecae: implications for cytoplasmic incompatibility. Insect Biochem. Mol. Biol. 43, 867–878 (2013).

  26. 26.

    Beckmann, J. F., Ronau, J. A. & Hochstrasser, M. A. Wolbachia deubiquitylating enzyme induces cytoplasmic incompatibility. Nat. Microbiol. 2, 17007 (2017).

  27. 27.

    LePage, D. P. et al. Prophage WO genes recapitulate and enhance Wolbachia-induced cytoplasmic incompatibility. Nature 543, 243–247 (2017).

  28. 28.

    Zhou, W., Rousset, F. & O’Neil, S. Phylogeny and PCR-based classification of Wolbachia strains using wsp gene sequences. Proc. Biol. Sci. 265, 509–515 (1998).

  29. 29.

    Fukatsu, T. & Nikoh, N. Two intracellular symbiotic bacteria from the mulberry psyllid Anomoneura mori (Insecta, Homoptera). Appl. Environ. Microbiol. 64, 3599–3606 (1998).

  30. 30.

    Fukatsu, T. & Nikoh, N. Endosymbiotic microbiota of the bamboo pseudococcid Antonina crawii (Insecta, Homoptera). Appl. Environ. Microbiol. 66, 643–650 (2000).

  31. 31.

    Kelley, R. L. et al. Expression of msl-2 causes assembly of dosage compensation regulators on the X chromosomes and female lethality in Drosophila. Cell 81, 867–877 (1995).

  32. 32.

    Brand, A. H. & Perrimon, N. Targeted gene expression as a means of altering cell fates and generating dominant phenotypes. Development 118, 401–415 (1993).

  33. 33.

    Ashburner, M., Golic, K. G. & Hawley, R. S. Drosophila: a Laboratory Handbook. 2nd Edition, (Cold Spring Harbor, New York, 2005).

  34. 34.

    Rørth, P. Gal4 in the Drosophila female germline. Mech. Dev. 78, 113–118 (1998).

  35. 35.

    Pool, J. E., Wong, A. & Aquadro, C. F. Finding of male-killing Spiroplasma infecting Drosophila melanogaster in Africa implies transatlantic migration of this endosymbiont. Heredity (Edinb) 97, 27–32 (2006).

  36. 36.

    Ye, F., Melcher, U., Rascoe, J. E. & Fletcher, J. Extensive chromosome aberrations in Spiroplasma citri Strain BR3. Biochem. Genet. 34, 269–286 (1996).

  37. 37.

    Ku, C., Lo, W.-S., Chen, L.-L. & Kuo, C.-H. Complete genomes of two dipteran-associated spiroplasmas provided insights into the origin, dynamics, and impacts of viral invasion in Spiroplasma. Genome Biol. Evol. 5, 1151–1164 (2013).

  38. 38.

    Anbutsu, H. & Fukatsu, T. Population dynamics of male-killing and non-male-killing spiroplasmas in Drosophila melanogaster. Appl. Environ. Microbiol. 69, 1428–1434 (2003).

  39. 39.

    Osborne, S. E., Leong, Y. S., O’Neill, S. L. & Johnson, K. N. Variation in antiviral protection mediated by different Wolbachia strains in Drosophila simulans. PLoS Pathog. 5, e1000656 (2009).

  40. 40.

    Pfaffl, M. W. A new mathematical model for relative quantification in real-time RT–PCR. Nucleic Acids Res. 29, e45 (2001).

  41. 41.

    Musselman, L. P. Drosophila hemolymph collection procedure. YouTube https://www.youtube.com/watch?v=im78OIBKlPA (2013).

  42. 42.

    Field, D. et al. Open software for biologists: from famine to feast. Nat. Biotechnol. 24, 801–803 (2006).

  43. 43.

    Darling, A. C. E., Mau, B., Blattner, F. R. & Perna, N. T. Mauve: multiple alignment of conserved genomic sequence with rearrangements. Genome Res. 14, 1394–1403 (2004).

  44. 44.

    Darling, A. E., Mau, B. & Perna, N. T. progressiveMauve: multiple genome alignment with gene gain, loss and rearrangement. PLoS ONE 5, e11147 (2010).

  45. 45.

    Saillard, C. et al. The abundant extrachromosomal DNA content of the Spiroplasma citri GII3-3X genome. BMC Genomics 9, 195 (2008).

  46. 46.

    Seemann, T. Prokka: rapid prokaryotic genome annotation. Bioinformatics 30, 2068–2069 (2014).

  47. 47.

    Carle, P. et al. Partial chromosome sequence of Spiroplasma citri reveals extensive viral invasion and important gene decay. Appl. Environ. Microbiol. 76, 3420–3426 (2010).

  48. 48.

    Alexeev, D. et al. Application of Spiroplasma melliferum proteogenomic profiling for the discovery of virulence factors and pathogenicity mechanisms in host-associated spiroplasmas. J. Proteome Res. 11, 224–236 (2012).

  49. 49.

    Lo, W.-S., Chen, L.-L., Chung, W.-C., Gasparich, G. E. & Kuo, C.-H. Comparative genome analysis of Spiroplasma melliferum IPMB4A, a honeybee-associated bacterium. BMC Genomics 14, 22 (2013).

  50. 50.

    Davis, R. E. et al. Complete genome sequence of Spiroplasma kunkelii strain CR2-3X, causal agent of corn stunt disease in Zea mays L. Genome Announc. 3, e01216–15 (2015).

  51. 51.

    Marchler-Bauer, A. et al. CDD: a conserved domain database for the functional annotation of proteins. Nucleic Acids Res. 39, D225–D229 (2011).

  52. 52.

    Marchler-Bauer, A. et al. CDD: NCBI’s conserved domain database. Nucleic Acids Res. 43, D222–D226 (2015).

  53. 53.

    Finn, R. D. et al. InterPro in 2017-beyond protein family and domain annotations. Nucleic Acids Res 45, D190–D199 (2017).

  54. 54.

    Liu, W. et al. IBS: an illustrator for the presentation and visualization of biological sequences. Bioinformatics 31, 3359–3361 (2015).

  55. 55.

    Su, X. Z., Wu, Y., Sifri, C. D. & Wellems, T. E. Reduced extension temperatures required for PCR amplification of extremely A+T-rich DNA. Nucleic Acids Res. 24, 1574–1575 (1996).

  56. 56.

    Hartenstein, V. Atlas of Drosophila Development. (Cold Spring Harbor, New York, 1993).

  57. 57.

    Campos-Ortega, J. A. & Hartenstein, V. The Embryonic Development of Drosophila melanogaster. (Springer, Berlin, 1997).

  58. 58.

    Bopp, D., Bell, L. R., Cline, T. W. & Schedl, P. Developmental distribution of female-specific Sex-lethal proteins in Drosophila melanogaster. Genes Dev. 5, 403–415 (1991).

  59. 59.

    O’Neill, E. M., Rebay, I., Tjian, R. & Rubin, G. M. The activities of two Ets-related transcription factors required for Drosophila eye development are modulated by the Ras/MAPK pathway. Cell 78, 137–147 (1994).

  60. 60.

    Schindelin, J. et al. Fiji: an open-source platform for biological-image analysis. Nat. Methods 9, 676–682 (2012).

  61. 61.

    Pau, G., Fuchs, F., Sklyar, O., Boutros, M. & Huber, W. EBImage—an R package for image processing with applications to cellular phenotypes. Bioinformatics 26, 979–981 (2010).

  62. 62.

    Hollander, M., Wolfe, A. D. & Chicken, E. Nonparametric Statistical Methods, 3rd Edition, (John Wiley & Sons, New Jersey, 2013).

Download references


We thank the Bloomington Stock Center in the USA and the Department of Drosophila Genomics and Genetic Resources at Kyoto Institute of Technology in Japan (DGGR) for fly stocks; the Developmental Studies Hybridoma Bank at the University of Iowa for monoclonal antibodies; the Drosophila Genomics Resource Center in the USA for DNA vectors; J. Jaenike, T. Murata, and M. Kuroda for providing fly strains; T. Fukatsu for the S. poulsonii-infected fly stocks that were originally established in his laboratory by T.H; J. Lucchesi for providing antibodies; the BioImaging & Optics Platform (BIOP) in EPFL for confocal microscopy; K. Harshman and the Lausanne Genomic Technologies Facility (GTF) at the University of Lausanne (UNIL) for whole-genome sequencing and draft genome assemblies; and C. Vorburger and S. Perlman for comments and suggestions on the manuscript; and the members of the laboratory for discussion and support. This work was supported by the European Research Council (ERC) Advanced Grant 339970 and the Swiss National Science Foundation (SNSF) Sinergia Grant CRSII3_154396. The purchase of the PacBio RS II system in UNIL was supported in part by the Loterie Romande through the Fondation pour la Recherche en Médecine Génétique.

Reviewer information

Nature thanks P. Ferree and the other anonymous reviewer(s) for their contribution to the peer review of this work.

Author information


  1. Global Health Institute, School of Life Sciences, École Polytechnique Fédérale de Lausanne (EPFL), Lausanne, Switzerland

    • Toshiyuki Harumoto
    •  & Bruno Lemaitre


  1. Search for Toshiyuki Harumoto in:

  2. Search for Bruno Lemaitre in:


T.H. conceived of and designed the study, performed the experiments, analysed the data and wrote and edited the manuscript. B.L. conceived and administered the study and wrote and edited the manuscript.

Competing interests

The authors declare no competing interests.

Corresponding authors

Correspondence to Toshiyuki Harumoto or Bruno Lemaitre.

Extended data figures and tables

  1. Extended Data Fig. 1 Identification and characterization of the partial male-killing S. poulsonii strain.

    a, An illustration showing the origin of the S. poulsonii strains analysed in this study. Pictures show male-killing (MK) S. poulsonii of D. melanogaster. MSRO-Ug is the original male-killing strain maintained in the Oregon-R wild-type fly. Fly stocks (Df(3L)H99 and Sxl-eGFP) artificially infected with this original strain showed complete male killing (100% MK) for the first 20 generations. Afterwards, one strain (MSRO-SE) started to show the partial male-killing phenotype, whereas the other (MSRO-H99) kept the ability to induce complete male killing. See Methods section ‘Identification and characterization of the partial male-killing S. poulsonii strain’ for more detail. b, c, Sex-ratio analysis of the adult progeny obtained from Oregon-R flies infected with MSRO-Ug, MSRO-H99, and MSRO-SE. We repeated experiments three times on the fourth, fifth and sixth generations (G4–6) after the establishment of infection. In c, the relative number of male offspring (percentage of females) obtained from Oregon-R female flies infected with MSRO-SE are plotted. Data points were excluded if the total count of flies was below 10. d, Relative titre of S. poulsonii within individual female flies. Adult females infected with three MSRO strains were kept for 0, 7 and 14 days after eclosion and analysed by qPCR. Data were normalized with respect to females from day 0 that were infected with MSRO-Ug. Different letters indicate statistically significant different groups (P < 0.01; N.S., not significant, P > 0.05; Steel–Dwass test; see Supplementary Table 2). Please note that the titres of the three strains are comparable and even the higher titre in old females (see 14 days in d) fails to induce complete male killing in MSRO-SE (c). Box and dot plots are as in Fig. 1c and sample sizes (n, number of adult flies) are shown at the bottom. Source Data

  2. Extended Data Fig. 2 Whole-genome sequencing studies of the male-killing S. poulsonii variants.

    a, Comparison of the genomic features of three S. poulsonii strains. MSRO-H99 and MSRO-SE are newly obtained variants isolated in this study (Extended Data Fig. 1a). As a control, data from the previously reported male-killing S. poulsonii genome16 are also indicated. b, Whole-genome alignment of the three S. poulsonii strains. To start the alignment from the dnaA gene, contig 1 of MSRO-H99 was split into two fragments (contig 1.1 and 1.2). The locations of the contigs corresponding to extra chromosomes (putative plasmids; see Methods section ‘Genomic data analyses and protein domain searches’) are shown as ‘extra’. spaid (gene ID from GenBank BioProject PRJNA416288: SMH99_26490) and spaid ΔC (gene ID: SMSE_25110) are located on these extra chromosomes in MSRO-H99 (contig 4) and MSRO-SE (contig 2), respectively.

  3. Extended Data Fig. 3 Genetic alterations of the spaid locus in the partial male-killing S. poulsonii strain.

    The genome structures around the spaid loci in the male-killing (a, MSRO-Ug and MSRO-H99) and the partial male-killing (b, MSRO-SE) S. poulsonii strains. Genes encoded on opposite strands are shown in different colours (red and blue, respectively). An 828-bp deletion and nucleotide substitutions (coloured in red; corresponding amino acid sequences are presented in one-letter code) in the 3′ region of the spaid gene are indicated. These sequence alterations were confirmed using Sanger sequencing (see Methods).

  4. Extended Data Fig. 4 Neural defects of Spaid-expressing embryos.

    Representative images of stage 13–14 female (a, n = 14) and male (bn = 16) embryos maternally expressing Spaid and stained for TUNEL (green) and neural cells (Elav, magenta). Single-channel images of Elav are also shown. The outlined region in b is magnified in c with single-channel images of Elav and TUNEL. Embryos were co-stained for Elav, TUNEL, Sxl and DNA, and selected channels are shown in a–c and Fig. 2a, b, respectively.

  5. Extended Data Fig. 5 Spaid acts through the dosage compensation machinery.

    a–c, Representative images of stage 13–14 embryos ectopically expressing the MSL complex (H83M2 transgene), stained for TUNEL (green) and Sxl (magenta). GFP-expressing control female (a, n = 15), Spaid-expressing female (b, n = 20), and male (c, n = 19) embryos are shown. d, Quantification of TUNEL signals in H83M2 embryos at stages 13–14. Different characters indicate significantly different groups (P < 0.001; Steel–Dwass test; see Supplementary Table 2). The box and dot plot (females, red; males, blue) is as in Fig. 1c and sample sizes (n, number of embryos) are shown at the bottom. e, f, Representative images of epithelial cells of stage 8–10 male embryos expressing GFP (e, n = 25) and Spaid (f, n = 25), stained for DNA (green) and MSL1 (magenta) from the datasets analysed in Fig. 3. All UAS transgenes were expressed maternally. Source Data

  6. Extended Data Fig. 6 Expression of Spaid using a weak GAL4 driver.

    The number of adult progeny (females, red; males, blue) obtained from crosses between the armadillo-GAL4 driver line (weak and ubiquitous expression) and four UAS transgenic lines (GFP, Spaid, ΔANK, and ΔOTU; n = 6 independent crosses for GFP and Spaid, n = 8 independent crosses for ΔANK and ΔOTU). The UAS-GFP line was used as a negative control. With this weak GAL4 driver, Spaid still eliminated all male progeny, while both ΔANK and ΔOTU had no impact on male viability. An asterisk indicates the statistically significant difference (P < 0.01; N.S., not significant, P > 0.05; two-tailed Mann–Whitney U test; see Supplementary Table 2). Box and dot plots are as in Fig. 1c. The total numbers of adult counts for each genotype and sex are shown at the bottom. Source Data

  7. Extended Data Fig. 7 A proposed model for Spaid-induced male-killing phenotypes.

    Spaid utilizes the OTU domain and ankyrin repeats (ANK) to target the host nucleus and the male X chromosome. ‘MSL’ and ‘Ac’ indicate the dosage compensation complex and resultant histone acetylation, respectively. See text for other explanations.

Supplementary information

  1. Supplementary Information

    This file contains Supplementary Tables 1-2. Supplementary Table 1 contains a list of PCR primers (the names and sequences of PCR primers are indicated with the names of the target organisms) and Supplementary Table 2 contains exact P values (statistically significant pairs with an alpha level of 0.05 are indicated in red characters).

  2. Reporting Summary

  3. Supplementary Data

    Summary of conserved domain database search results for the proteins annotated in the Spiroplasma strains MSRO-H99 (Data 1), MSRO-SE (Data 2), and in the previously published genome (Data 3; MSRO, 2015). Spaid in MSRO-H99 (SMH99_26490) and Spaid ∆C in MSRO-SE (SMSE_25110) are shown in red characters.

Source data

About this article

Publication history




Issue Date




By submitting a comment you agree to abide by our Terms and Community Guidelines. If you find something abusive or that does not comply with our terms or guidelines please flag it as inappropriate.