1887

Microbial Genomics logo

Volume 6, Issue 12

Diminutive, degraded but dissimilar: genomes from filarial nematodes do not conform to a single paradigm Open Access

Abstract

are alpha-proteobacteria symbionts infecting a large range of arthropod species and two different families of nematodes. Interestingly, these endosymbionts are able to induce diverse phenotypes in their hosts: they are reproductive parasites within many arthropods, nutritional mutualists within some insects and obligate mutualists within their filarial nematode hosts. Defining ‘species’ is controversial and so they are commonly classified into 17 different phylogenetic lineages, termed supergroups, named A–F, H–Q and S. However, available genomic data remain limited and not representative of the full diversity; indeed, of the 24 complete genomes and 55 draft genomes of available to date, 84 % belong to supergroups A and B, exclusively composed of from arthropods. For the current study, we took advantage of a recently developed DNA-enrichment method to produce four complete genomes and two draft genomes of from filarial nematodes. Two complete genomes, Ctub and Dcau, are the smallest genomes sequenced to date (863 988 bp and 863 427 bp, respectively), as well as the first genomes representing supergroup J. These genomes confirm the validity of this supergroup, a controversial clade due to weaknesses of the multilocus sequence typing approach. We also produced the first draft genome from a supergroup F filarial nematode representative (Mhie), two genomes from supergroup D (Lsig and Lbra) and the complete genome of Dimm from supergroup C. Our new data confirm the paradigm of smaller genomes from filarial nematodes containing low levels of transposable elements and the absence of intact bacteriophage sequences, unlike many from arthropods, where both are more abundant. However, we observe differences among the genomes from filarial nematodes: no global co-evolutionary pattern, strong synteny between supergroup C and supergroup J and more transposable elements observed in supergroup D compared to the other supergroups. Metabolic pathway analysis indicates several highly conserved pathways (haem and nucleotide biosynthesis, for example) as opposed to more variable pathways, such as vitamin B biosynthesis, which might be specific to certain host–symbiont associations. Overall, there appears to be no single –filarial nematode pattern of co-evolution or symbiotic relationship.

  • Received:
  • Accepted:
  • Published Online:
Keyword(s): filarial nematodes , genomics , symbiosis , target enrichment and Wolbachia
© 2020 The Authors
  • This is an open-access article distributed under the terms of the Creative Commons Attribution License.
Loading

Article metrics loading...

/content/journal/mgen/10.1099/mgen.0.000487
2020-12-09
2024-05-14
Loading full text...

Full text loading...

/deliver/fulltext/mgen/6/12/mgen000487.html?itemId=/content/journal/mgen/10.1099/mgen.0.000487&mimeType=html&fmt=ahah

References

  1. Zug R, Hammerstein P. Still a host of hosts for Wolbachia: analysis of recent data suggests that 40% of terrestrial arthropod species are infected. PLoS One 2012; 7:e38544 [View Article]
     [Google Scholar]
  2. Werren JH, Windsor DM. Wolbachia infection frequencies in insects: evidence of a global equilibrium?. Proc Biol Sci 2000; 267:1277–1285 [View Article]
     [Google Scholar]
  3. Brown AMV, Wasala SK, Howe DK, Peetz AB, Zasada IA et al. Genomic evidence for plant-parasitic nematodes as the earliest Wolbachia hosts. Sci Rep 2016; 6:34955 [View Article]
     [Google Scholar]
  4. Bandi C, Anderson TJC, Genchi C, Blaxter ML. Phylogeny of Wolbachia in filarial nematodes. Proc Biol Sci 1998; 265:2407–2413 [View Article]
     [Google Scholar]
  5. Werren JH, Baldo L, Clark ME. Wolbachia: master manipulators of invertebrate biology. Nat Rev Microbiol 2008; 6:741–751 [View Article]
     [Google Scholar]
  6. Kageyama D, Nishimura G, Hoshizaki S, Ishikawa Y. Feminizing Wolbachia in an insect, Ostrinia furnacalis (Lepidoptera: Crambidae). Heredity 2002; 88:444–449 [View Article]
     [Google Scholar]
  7. Bouchon D, Rigaud T, Juchault P. Evidence for widespread Wolbachia infection in isopod crustaceans: molecular identification and host feminization. Proc Biol Sci 1998; 265:1081–1090 [View Article]
     [Google Scholar]
  8. Duron O, Fort P, Weill M. Influence of aging on cytoplasmic incompatibility, sperm modification and Wolbachia density in Culex pipiens mosquitoes. Heredity 2007; 98:368–374 [View Article]
     [Google Scholar]
  9. Hosokawa T, Koga R, Kikuchi Y, Meng X-Y, Fukatsu T. Wolbachia as a bacteriocyte-associated nutritional mutualist. Proc Natl Acad Sci USA 2010; 107:769–774 [View Article]
     [Google Scholar]
  10. Hoerauf A, Mand S, Fischer K, Kruppa T, Marfo-Debrekyei Y et al. Doxycycline as a novel strategy against bancroftian filariasis–depletion of Wolbachia endosymbionts from Wuchereria bancrofti and stop of microfilaria production. Med Microbiol Immunol 2003; 192:211–216 [View Article]
     [Google Scholar]
  11. Badawi M, Moumen B, Giraud I, Grève P, Cordaux R. Investigating the molecular genetic basis of cytoplasmic sex determination caused by Wolbachia endosymbionts in terrestrial isopods. Genes 2018; 9:290 [View Article]
     [Google Scholar]
  12. Lindsey ARI, Werren JH, Richards S, Stouthamer R. Comparative genomics of a parthenogenesis-inducing Wolbachia symbiont. G3 2016; 6:2113–2123 [View Article]
     [Google Scholar]
  13. Bakowski MA, McNamara CW. Advances in antiwolbachial drug discovery for treatment of parasitic filarial worm infections. Trop Med Infect Dis 2019; 4:108 [View Article]
     [Google Scholar]
  14. Walker T, Klasson L, Sebaihia M, Sanders MJ, Thomson NR et al. Ankyrin repeat domain-encoding genes in the wPip strain of Wolbachia from the Culex pipiens group. BMC Biol 2007; 5:39 [View Article]
     [Google Scholar]
  15. Wu M, Sun LV, Vamathevan J, Riegler M, Deboy R et al. Phylogenomics of the reproductive parasite Wolbachia pipientis wMel: a streamlined genome overrun by mobile genetic elements. PLoS Biol 2004; 2:E69 [View Article]
     [Google Scholar]
  16. LePage DP, Metcalf JA, Bordenstein SR, On J, Perlmutter JI et al. Prophage WO genes recapitulate and enhance Wolbachia-induced cytoplasmic incompatibility. Nature 2017; 543:243–247 [View Article]
     [Google Scholar]
  17. Beckmann JF, Ronau JA, Hochstrasser M. A Wolbachia deubiquitylating enzyme induces cytoplasmic incompatibility. Nat Microbiol 2017; 2:17007 [View Article]
     [Google Scholar]
  18. Perlmutter JI, Bordenstein SR, Unckless RL, LePage DP, Metcalf JA et al. The phage gene wmk is a candidate for male killing by a bacterial endosymbiont. PLoS Pathog 2019; 15:e1007936 [View Article]
     [Google Scholar]
  19. Pichon S, Bouchon D, Liu C, Chen L, Garrett RA et al. The expression of one ankyrin pk2 allele of the WO prophage is correlated with the Wolbachia feminizing effect in isopods. BMC Microbiol 2012; 12:55 [View Article]
     [Google Scholar]
  20. Nikoh N, Hosokawa T, Moriyama M, Oshima K, Hattori M et al. Evolutionary origin of insect-Wolbachia nutritional mutualism. Proc Natl Acad Sci USA 2014; 111:10257–10262 [View Article]
     [Google Scholar]
  21. Ju J-F, Bing X-L, Zhao D-S, Guo Y, Xi Z et al. Wolbachia supplement biotin and riboflavin to enhance reproduction in planthoppers. ISME J 2020; 14:676–687
     [Google Scholar]
  22. Kamtchum-Tatuene J, Makepeace BL, Benjamin L, Baylis M, Solomon T. The potential role of Wolbachia in controlling the transmission of emerging human arboviral infections. Curr Opin Infect Dis 2017; 30:108–116 [View Article]
     [Google Scholar]
  23. Moreira LA, Iturbe-Ormaetxe I, Jeffery JA, Lu G, Pyke AT et al. A Wolbachia symbiont in Aedes aegypti limits infection with dengue, chikungunya, and Plasmodium . Cell 2009; 139:1268–1278 [View Article]
     [Google Scholar]
  24. Blagrove MSC, Arias-Goeta C, Failloux A-B, Sinkins SP. Wolbachia strain wMel induces cytoplasmic incompatibility and blocks dengue transmission in Aedes albopictus. Proc Natl Acad Sci USA 2012; 109:255–260 [View Article]
     [Google Scholar]
  25. Slatko BE, Luck AN, Dobson SL, Foster JM. Wolbachia endosymbionts and human disease control. Mol Biochem Parasitol 2014; 195:88–95 [View Article]
     [Google Scholar]
  26. Bourtzis K, Dobson SL, Xi Z, Rasgon JL, Calvitti M et al. Harnessing mosquito–Wolbachia symbiosis for vector and disease control. Acta Trop 2014; 132:S150–S163 [View Article]
     [Google Scholar]
  27. Ye YH, Woolfit M, Rances E, O'Neill SL, McGraw EA. Wolbachia-associated bacterial protection in the mosquito Aedes aegypti . PLoS Negl Trop Dis 2013; 7:e2362
     [Google Scholar]
  28. Bouchery T, Lefoulon E, Karadjian G, Nieguitsila A, Martin C. The symbiotic role of Wolbachia in Onchocercidae and its impact on filariasis. Clin Microbiol Infect 2013; 19:131–140 [View Article]
     [Google Scholar]
  29. Pfarr KM, Debrah AY, Specht S, Hoerauf A. Filariasis and lymphoedema. Parasite Immunol 2009; 31:664–672 [View Article]
     [Google Scholar]
  30. Taylor MJ, von Geldern TW, Ford L, Hübner MP, Marsh K et al. Preclinical development of an oral anti-Wolbachia macrolide drug for the treatment of lymphatic filariasis and onchocerciasis. Sci Transl Med 2019; 11:eaau2086 [View Article]
     [Google Scholar]
  31. Molyneux DH, Bradley M, Hoerauf A, Kyelem D, Taylor MJ. Mass drug treatment for lymphatic filariasis and onchocerciasis. Trends Parasitol 2003; 19:516–522 [View Article]
     [Google Scholar]
  32. Taylor MJ, Hoerauf A, Bockarie M. Lymphatic filariasis and onchocerciasis. Lancet 2010; 376:1175–1185 [View Article]
     [Google Scholar]
  33. Foster J, Ganatra M, Kamal I, Ware J, Makarova K et al. The Wolbachia genome of Brugia malayi: endosymbiont evolution within a human pathogenic nematode. PLoS Biol 2005; 3:e121 [View Article]
     [Google Scholar]
  34. Ghedin E, Hailemariam T, DePasse JV, Zhang X, Oksov Y et al. Brugia malayi gene expression in response to the targeting of the Wolbachia endosymbiont by tetracycline treatment. PLoS Negl Trop Dis 2009; 3:e525 [View Article]
     [Google Scholar]
  35. Voronin D, Bachu S, Shlossman M, Unnasch TR, Ghedin E et al. Glucose and glycogen metabolism in Brugia malayi is associated with Wolbachia symbiont fitness. PLoS One 2016; 11:e0153812 [View Article]
     [Google Scholar]
  36. Grote A, Voronin D, Ding T, Twaddle A, Unnasch TR et al. Defining Brugia malayi and Wolbachia symbiosis by stage-specific dual RNA-seq. PLoS Negl Trop Dis 2017; 11:e0005357 [View Article]
     [Google Scholar]
  37. Li Z, Carlow CKS. Characterization of transcription factors that regulate the type IV secretion system and riboflavin biosynthesis in Wolbachia of Brugia malayi. PLoS One 2012; 7:e51597 [View Article]
     [Google Scholar]
  38. Slatko BE, Taylor MJ, Foster JM. The Wolbachia endosymbiont as an anti-filarial nematode target. Symbiosis 2010; 51:55–65 [View Article]
     [Google Scholar]
  39. Clare RH, Cook DA, Johnston KL, Ford L, Ward SA et al. Development and validation of a high-throughput anti-Wolbachia whole-cell screen: a route to macrofilaricidal drugs against onchocerciasis and lymphatic filariasis. J Biomol Screen 2015; 20:64–69
     [Google Scholar]
  40. Xu Z, Fang S-M, Bakowski MA, Rateb ME, Yang D et al. Discovery of kirromycins with anti-Wolbachia activity from Streptomyces sp. CB00686. ACS Chem Biol 2019; 14:1174–1182 [View Article]
     [Google Scholar]
  41. Johnston KL, Cook DAN, Berry NG, David Hong W, Clare RH et al. Identification and prioritization of novel anti-Wolbachia chemotypes from screening a 10,000-compound diversity library. Sci Adv 2017; 3:eaao1551 [View Article]
     [Google Scholar]
  42. Hong WD, Benayoud F, Nixon GL, Ford L, Johnston KL et al. AWZ1066S, a highly specific anti-Wolbachia drug candidate for a short-course treatment of filariasis. Proc Natl Acad Sci USA 2019; 116:1414–1419 [View Article]
     [Google Scholar]
  43. Darby AC, Armstrong SD, Bah GS, Kaur G, Hughes MA et al. Analysis of gene expression from the Wolbachia genome of a filarial nematode supports both metabolic and defensive roles within the symbiosis. Genome Res 2012; 22:2467–2477
     [Google Scholar]
  44. Cotton JA, Bennuru S, Grote A, Harsha B, Tracey A et al. The genome of Onchocerca volvulus, agent of river blindness. Nat Microbiol 2017; 2:16216 [View Article]
     [Google Scholar]
  45. Lebov JF, Mattick J, Libro S, Sparklin BC, Chung M et al. Complete genome sequence of wBp, the Wolbachia endosymbiont of Brugia pahangi FR3. Microbiol Resour Announc 2020; 9:e00480-20 [View Article]
     [Google Scholar]
  46. Sullivan W. Wolbachia, bottled water, and the dark side of symbiosis. Mol Biol Cell 2017; 28:2343–2346 [View Article][PubMed]
     [Google Scholar]
  47. Comandatore F, Cordaux R, Bandi C, Blaxter M, Darby A et al. Supergroup C Wolbachia, mutualist symbionts of filarial nematodes, have a distinct genome structure. Open Biol 2015; 5:150099 [View Article]
     [Google Scholar]
  48. Ramírez-Puebla ST, Servín-Garcidueñas LE, Ormeño-Orrillo E, Vera-Ponce de León A, Rosenblueth M et al. Species in Wolbachia? Proposal for the designation of ‘Candidatus Wolbachia bourtzisii’, ‘Candidatus Wolbachia onchocercicola’, ‘Candidatus Wolbachia blaxteri’, ‘Candidatus Wolbachia brugii’, ‘Candidatus Wolbachia taylori’, ‘Candidatus Wolbachia collembolicola’ and ‘Candidatus Wolbachia multihospitum’ for the different species within Wolbachia supergroups. Syst Appl Microbiol 2015; 38:390–399 [View Article]
     [Google Scholar]
  49. Lindsey ARI, Bordenstein SR, Newton ILG, Rasgon JL. Wolbachia pipientis should not be split into multiple species: a response to Ramírez-Puebla et al., “Species in Wolbachia? Proposal for the designation of ‘Candidatus Wolbachia bourtzisii’, ‘Candidatus Wolbachia onchocercicola’, ‘Candidatus Wolbachia blaxteri’, ‘Candidatus Wolbachia brugii’, ‘Candidatus Wolbachia taylori’, ‘Candidatus Wolbachia collembolicola’ and ‘Candidatus Wolbachia multihospitum’ for the different species within Wolbachia supergroups”. Syst Appl Microbiol 2016; 39:220–222 [View Article]
     [Google Scholar]
  50. Chung M, Munro JB, Tettelin H, Dunning Hotopp JC. Using core genome alignments to assign bacterial species. mSystems 2018; 3:e00236-18 [View Article]
     [Google Scholar]
  51. Newton ILG, Slatko BE. Symbiosis comes of age at the 10th biennial meeting of Wolbachia researchers. Appl Environ Microbiol 2019; 85:e03071-18 [View Article]
     [Google Scholar]
  52. Gerth M. Classification of Wolbachia (Alphaproteobacteria, Rickettsiales): no evidence for a distinct supergroup in cave spiders. Infect Genet Evol 2016; 43:378–380 [View Article]
     [Google Scholar]
  53. Baldo L, Werren JH. Revisiting Wolbachia supergroup typing based on WSP: spurious lineages and discordance with MLST. Curr Microbiol 2007; 55:81–87 [View Article]
     [Google Scholar]
  54. Zhou W, Rousset F, O'Neill S. Phylogeny and PCR–based classification of Wolbachia strains using wsp gene sequences. Proc Biol Sci 1998; 265:509–515 [View Article]
     [Google Scholar]
  55. Lo N, Casiraghi M, Salati E, Bazzocchi C, Bandi C. How many Wolbachia supergroups exist?. Mol Biol Evol 2002; 19:341–346 [View Article]
     [Google Scholar]
  56. Ferri E, Bain O, Barbuto M, Martin C, Lo N et al. New insights into the evolution of Wolbachia infections in filarial nematodes inferred from a large range of screened species. PLoS One 2011; 6:e20843 [View Article]
     [Google Scholar]
  57. Glowska E, Dragun-Damian A, Dabert M, Gerth M. New Wolbachia supergroups detected in quill mites (Acari: Syringophilidae). Infect Genet Evol 2015; 30:140–146 [View Article]
     [Google Scholar]
  58. Lefoulon E, Bain O, Makepeace BL, d’Haese C, Uni S et al. Breakdown of coevolution between symbiotic bacteria Wolbachia and their filarial hosts. PeerJ 2016; 4:e1840 [View Article]
     [Google Scholar]
  59. Lo N, Paraskevopoulos C, Bourtzis K, O'Neill SL, Werren JH et al. Taxonomic status of the intracellular bacterium Wolbachia pipientis. Int J Syst Evol Microbiol 2007; 57:654–657 [View Article]
     [Google Scholar]
  60. Baldo L, Dunning Hotopp JC, Jolley KA, Bordenstein SR, Biber SA et al. Multilocus sequence typing system for the endosymbiont Wolbachia pipientis. Appl Environ Microbiol 2006; 72:7098–7110 [View Article]
     [Google Scholar]
  61. Casiraghi M, Bain O, Guerrero R, Martin C, Pocacqua V et al. Mapping the presence of Wolbachia pipientis on the phylogeny of filarial nematodes: evidence for symbiont loss during evolution. Int J Parasitol 2004; 34:191–203 [View Article]
     [Google Scholar]
  62. Bordenstein SR, Paraskevopoulos C, Dunning Hotopp JC, Sapountzis P, Lo N et al. Parasitism and mutualism in Wolbachia: what the phylogenomic trees can and cannot say. Mol Biol Evol 2009; 26:231–241 [View Article]
     [Google Scholar]
  63. Ma Y, Chen W-J, Li Z-H, Zhang F, Gao Y et al. Revisiting the phylogeny of Wolbachia in Collembola. Ecol Evol 2017; 7:2009–2017 [View Article]
     [Google Scholar]
  64. Konecka E, Olszanowski Z. A screen of maternally inherited microbial endosymbionts in oribatid mites (Acari: Oribatida). Microbiology 2015; 161:1561–1571 [View Article]
     [Google Scholar]
  65. Khoo JJ, Kurtti TJ, Husin NA, Beliavskaia A, Lim FS et al. Isolation and propagation of laboratory strains and a novel flea-derived field strain of Wolbachia in tick cell lines. Microorganisms 2020; 8:988 [View Article]
     [Google Scholar]
  66. Werren JH, Zhang W, Guo LR. Evolution and phylogeny of Wolbachia: reproductive parasites of arthropods. Proc Biol Sci 1995; 261:55–63
     [Google Scholar]
  67. Ros VID, Fleming VM, Feil EJ, Breeuwer JAJ. How diverse is the genus Wolbachia? Multiple-gene sequencing reveals a putatively new Wolbachia supergroup recovered from spider mites (Acari: Tetranychidae). Appl Environ Microbiol 2009; 75:1036–1043 [View Article]
     [Google Scholar]
  68. Bing X-L, Xia W-Q, Gui J-D, Yan G-H, Wang X-W et al. Diversity and evolution of the Wolbachia endosymbionts of Bemisia (Hemiptera: Aleyrodidae) whiteflies. Ecol Evol 2014; 4:2714–2737 [View Article][PubMed]
     [Google Scholar]
  69. Bordenstein S, Rosengaus RB. Discovery of a novel Wolbachia supergroup in Isoptera. Curr Microbiol 2005; 51:393–398 [View Article]
     [Google Scholar]
  70. Lefoulon E, Clark T, Borveto F, Perriat-Sanguinet M, Moulia C et al. Pseudoscorpion Wolbachia symbionts: diversity and evidence for a new supergroup S. BMC Microbiol 2020; 20:188 [View Article]
     [Google Scholar]
  71. Sironi M, Bandi C, Sacchi L, Di Sacco B, Damiani G et al. Molecular evidence for a close relative of the arthropod endosymbiont Wolbachia in a filarial worm. Mol Biochem Parasitol 1995; 74:223–227 [View Article]
     [Google Scholar]
  72. Haegeman A, Vanholme B, Jacob J, Vandekerckhove TTM, Claeys M et al. An endosymbiotic bacterium in a plant-parasitic nematode: member of a new Wolbachia supergroup. Int J Parasitol 2009; 39:1045–1054 [View Article]
     [Google Scholar]
  73. Lefoulon E, Gavotte L, Junker K, Barbuto M, Uni S et al. A new type F Wolbachia from Splendidofilariinae (Onchocercidae) supports the recent emergence of this supergroup. Int J Parasitol 2012; 42:1025–1036 [View Article]
     [Google Scholar]
  74. Baldo L, Lo N, Werren JH. Mosaic nature of the Wolbachia surface protein. J Bacteriol 2005; 187:5406–5418 [View Article]
     [Google Scholar]
  75. Bleidorn C, Gerth M. A critical re-evaluation of multilocus sequence typing (MLST) efforts in Wolbachia. FEMS Microbiol Ecol 2018; 94:fix163 [View Article]
     [Google Scholar]
  76. Lefoulon E, Vaisman N, Frydman HM, Sun L, Voland L et al. Large enriched fragment targeted sequencing (LEFT-SEQ) applied to capture of Wolbachia genomes. Sci Rep 2019; 9:5939 [View Article]
     [Google Scholar]
  77. Kent BN, Salichos L, Gibbons JG, Rokas A, Newton ILG et al. Complete bacteriophage transfer in a bacterial endosymbiont (Wolbachia) determined by targeted genome capture. Genome Biol Evol 2011; 3:209–218 [View Article]
     [Google Scholar]
  78. Geniez S, Foster JM, Kumar S, Moumen B, LeProust E et al. Targeted genome enrichment for efficient purification of endosymbiont DNA from host DNA. Symbiosis 2012; 58:201–207 [View Article]
     [Google Scholar]
  79. Lefoulon E, Bain O, Bourret J, Junker K, Guerrero R et al. Shaking the tree: multi-locus sequence typing usurps current Onchocercid (filarial nematode) phylogeny. PLoS Negl Trop Dis 2015; 9:e0004233 [View Article]
     [Google Scholar]
  80. Luck AN, Anderson KG, McClung CM, VerBerkmoes NC, Foster JM et al. Tissue-specific transcriptomics and proteomics of a filarial nematode and its Wolbachia endosymbiont. BMC Genomics 2015; 16:920 [View Article]
     [Google Scholar]
  81. Godel C, Kumar S, Koutsovoulos G, Ludin P, Nilsson D et al. The genome of the heartworm, Dirofilaria immitis, reveals drug and vaccine targets. FASEB J 2012; 26:4650–4661 [View Article]
     [Google Scholar]
  82. Comandatore F, Sassera D, Montagna M, Kumar S, Koutsovoulos G et al. Phylogenomics and analysis of shared genes suggest a single transition to mutualism in Wolbachia of nematodes. Genome Biol Evol 2013; 5:1668–1674 [View Article]
     [Google Scholar]
  83. Saha S, Hunter WB, Reese J, Morgan JK, Marutani-Hert M et al. Survey of endosymbionts in the Diaphorina citri metagenome and assembly of a Wolbachia wDi draft genome. PLoS One 2012; 7:e50067 [View Article]
     [Google Scholar]
  84. Koren S, Walenz BP, Berlin K, Miller JR, Bergman NH et al. Canu: scalable and accurate long-read assembly via adaptive k -mer weighting and repeat separation. Genome Res 2017; 27:722–736 [View Article]
     [Google Scholar]
  85. Nurk S, Bankevich A, Antipov D, Gurevich AA, Korobeynikov A et al. Assembling single-cell genomes and mini-metagenomes from chimeric MDA products. J Comput Biol 2013; 20:714–737 [View Article]
     [Google Scholar]
  86. Camacho C, Coulouris G, Avagyan V, Ma N, Papadopoulos J et al. BLAST+: architecture and applications. BMC Bioinformatics 2009; 10:421 [View Article]
     [Google Scholar]
  87. Zhang J, Kobert K, Flouri T, Stamatakis A. PEAR: a fast and accurate Illumina Paired-End reAd mergeR. Bioinformatics 2014; 30:614–620 [View Article]
     [Google Scholar]
  88. Langmead B, Salzberg SL. Fast gapped-read alignment with Bowtie 2. Nat Methods 2012; 9:357–359 [View Article]
     [Google Scholar]
  89. Sedlazeck FJ, Rescheneder P, Smolka M, Fang H, Nattestad M et al. Accurate detection of complex structural variations using single-molecule sequencing. Nat Methods 2018; 15:461–468 [View Article]
     [Google Scholar]
  90. Wick RR, Judd LM, Gorrie CL, Holt KE. Unicycler: resolving bacterial genome assemblies from short and long sequencing reads. PLoS Comput Biol 2017; 13:e1005595 [View Article]
     [Google Scholar]
  91. Gurevich A, Saveliev V, Vyahhi N, Tesler G. QUAST: quality assessment tool for genome assemblies. Bioinformatics 2013; 29:1072–1075 [View Article]
     [Google Scholar]
  92. Driscoll TP, Verhoeve VI, Gillespie JJ, Johnston JS, Guillotte ML et al. A chromosome-level assembly of the cat flea genome uncovers rampant gene duplication and genome size plasticity. BMC Biol 2020; 18:70 [View Article][PubMed]
     [Google Scholar]
  93. Yoon S-H, Ha S, Lim J, Kwon S, Chun J. A large-scale evaluation of algorithms to calculate average nucleotide identity. Antonie van Leeuwenhoek 2017; 110:1281–1286 [View Article]
     [Google Scholar]
  94. Meier-Kolthoff JP, Auch AF, Klenk H-P, Göker M. Genome sequence-based species delimitation with confidence intervals and improved distance functions. BMC Bioinformatics 2013; 14:60 [View Article]
     [Google Scholar]
  95. Auch AF, Klenk H-P, Göker M. Standard operating procedure for calculating genome-to-genome distances based on high-scoring segment pairs. Stand Genomic Sci 2010; 2:142–148 [View Article]
     [Google Scholar]
  96. Aziz RK, Bartels D, Best AA, DeJongh M, Disz T et al. The RAST server: rapid annotations using subsystems technology. BMC Genomics 2008; 9:75 [View Article]
     [Google Scholar]
  97. Varani AM, Siguier P, Gourbeyre E, Charneau V, Chandler M. ISsaga is an ensemble of web-based methods for high throughput identification and semi-automatic annotation of insertion sequences in prokaryotic genomes. Genome Biol 2011; 12:R30 [View Article]
     [Google Scholar]
  98. Arndt D, Grant JR, Marcu A, Sajed T, Pon A et al. PHASTER: a better, faster version of the PHAST phage search tool. Nucleic Acids Res 2016; 44:W16–W21 [View Article]
     [Google Scholar]
  99. Seemann T. Prokka: rapid prokaryotic genome annotation. Bioinformatics 2014; 30:2068–2069 [View Article]
     [Google Scholar]
  100. R Core Team R: a Language and Environment for Statistical Computing Vienna: R Foundation for Statistical Computing; 2017
  101. Moriya Y, Itoh M, Okuda S, Yoshizawa AC, Kanehisa M. KAAS: an automatic genome annotation and pathway reconstruction server. Nucleic Acids Res 2007; 35:W182–W185 [View Article]
     [Google Scholar]
  102. Emms DM, Kelly S. OrthoFinder: solving fundamental biases in whole genome comparisons dramatically improves orthogroup inference accuracy. Genome Biol 2015; 16:157 [View Article]
     [Google Scholar]
  103. Talavera G, Castresana J. Improvement of phylogenies after removing divergent and ambiguously aligned blocks from protein sequence alignments. Syst Biol 2007; 56:564–577 [View Article][PubMed]
     [Google Scholar]
  104. Kalyaanamoorthy S, Minh BQ, Wong TKF, von Haeseler A, Jermiin LS. ModelFinder: fast model selection for accurate phylogenetic estimates. Nat Methods 2017; 14:587–589 [View Article][PubMed]
     [Google Scholar]
  105. Ioannidis P, Dunning Hotopp JC, Sapountzis P, Siozios S, Tsiamis G et al. New criteria for selecting the origin of DNA replication in Wolbachia and closely related bacteria. BMC Genomics 2007; 8:182 [View Article][PubMed]
     [Google Scholar]
  106. Kurtz S, Phillippy A, Delcher AL, Smoot M, Shumway M et al. Versatile and open software for comparing large genomes. Genome Biol 2004; 5:R12 [View Article][PubMed]
     [Google Scholar]
  107. Balbuena JA, Míguez-Lozano R, Blasco-Costa I. PACo: a novel procrustes application to cophylogenetic analysis. PLoS One 2013; 8:e61048 [View Article][PubMed]
     [Google Scholar]
  108. Legendre P, Desdevises Y, Bazin E. A statistical test for host-parasite coevolution. Syst Biol 2002; 51:217–234 [View Article][PubMed]
     [Google Scholar]
  109. International Helminth Genomes Consortium Comparative genomics of the major parasitic worms. Nat Genet 2019; 51:163–174 [View Article][PubMed]
     [Google Scholar]
  110. Koutsovoulos G, Makepeace B, Tanya VN, Blaxter M. Palaeosymbiosis revealed by genomic fossils of Wolbachia in a strongyloidean nematode. PLoS Genet 2014; 10:e1004397 [View Article][PubMed]
     [Google Scholar]
  111. Casiraghi M, Bordenstein SR, Baldo L, Lo N, Beninati T et al. Phylogeny of Wolbachia pipientis based on gltA, groEL and ftsZ gene sequences: clustering of arthropod and nematode symbionts in the F supergroup, and evidence for further diversity in the Wolbachia tree. Microbiology 2005; 151:4015–4022 [View Article][PubMed]
     [Google Scholar]
  112. Kampfraath AA, Klasson L, Anvar SY, Vossen R, Roelofs D et al. Genome expansion of an obligate parthenogenesis-associated Wolbachia poses an exception to the symbiont reduction model. BMC Genomics 2019; 20:106 [View Article][PubMed]
     [Google Scholar]
  113. Martin C, Gavotte L. The bacteria Wolbachia in filariae, a biological Russian dolls' system: new trends in antifilarial treatments. Parasite 2010; 17:79–89 [View Article][PubMed]
     [Google Scholar]
  114. Siozios S, Ioannidis P, Klasson L, Andersson SGE, Braig HR et al. The diversity and evolution of Wolbachia ankyrin repeat domain genes. PLoS One 2013; 8:e55390 [View Article][PubMed]
     [Google Scholar]
  115. Fenn K, Blaxter M. Wolbachia genomes: revealing the biology of parasitism and mutualism. Trends Parasitol 2006; 22:60–65 [View Article][PubMed]
     [Google Scholar]
  116. Gerth M, Bleidorn C. Comparative genomics provides a timeframe for Wolbachia evolution and exposes a recent biotin synthesis operon transfer. Nat Microbiol 2016; 2:16241 [View Article][PubMed]
     [Google Scholar]
  117. McCutcheon JP, Moran NA. Extreme genome reduction in symbiotic bacteria. Nat Rev Microbiol 2011; 10:13–26 [View Article][PubMed]
     [Google Scholar]
  118. Fujii Y, Kubo T, Ishikawa H, Sasaki T. Isolation and characterization of the bacteriophage WO from Wolbachia, an arthropod endosymbiont. Biochem Biophys Res Commun 2004; 317:1183–1188 [View Article][PubMed]
     [Google Scholar]
  119. Gavotte L, Henri H, Stouthamer R, Charif D, Charlat S et al. A survey of the bacteriophage WO in the endosymbiotic bacteria Wolbachia. Mol Biol Evol 2007; 24:427–435 [View Article][PubMed]
     [Google Scholar]
  120. Masui S, Kuroiwa H, Sasaki T, Inui M, Kuroiwa T et al. Bacteriophage WO and virus-like particles in Wolbachia, an endosymbiont of arthropods. Biochem Biophys Res Commun 2001; 283:1099–1104 [View Article][PubMed]
     [Google Scholar]
  121. Wright JD, Sjöstrand FS, Portaro JK, Barr AR. The ultrastructure of the rickettsia-like microorganism Wolbachia pipientis and associated virus-like bodies in the mosquito Culex pipiens. J Ultrastruct Res 1978; 63:79–85 [View Article][PubMed]
     [Google Scholar]
  122. Bordenstein SR, Bordenstein SR. Eukaryotic association module in phage WO genomes from Wolbachia. Nat Commun 2016; 7:13155 [View Article][PubMed]
     [Google Scholar]
  123. Gerth M, Gansauge M-T, Weigert A, Bleidorn C. Phylogenomic analyses uncover origin and spread of the Wolbachia pandemic. Nat Commun 2014; 5:5117 [View Article][PubMed]
     [Google Scholar]
http://instance.metastore.ingenta.com/content/journal/mgen/10.1099/mgen.0.000487
Loading
Diminutive, degraded but dissimilar: Wolbachia genomes from filarial nematodes do not conform to a single paradigm
M Gen 6, e000487 (2020); https://doi.org/10.1099/mgen.0.000487
/content/journal/mgen/10.1099/mgen.0.000487
/content/journal/mgen/10.1099/mgen.0.000487
Loading

Data & Media loading...

Supplements

Supplementary material 1

PDF

Most cited Most Cited RSS feed

This is a required field
Please enter a valid email address
Approval was a Success
Invalid data
An Error Occurred
Approval was partially successful, following selected items could not be processed due to error