Russian Journal of Genetics

, Volume 52, Issue 1, pp 38–48 | Cite as

Distinctive features of the microbial diversity and the polyketide synthase genes spectrum in the community of the endemic Baikal sponge Swartschewskia papyracea

  • O. V. KaluzhnayaEmail author
  • V. B. Itskovich
Genetics of Microorganisms


The diversity of the symbiotic community of the endemic Baikal sponge Swartschewskia papyracea was studied, and an analysis of the polyketide synthases genes spectrum in sponge-associated microorganisms was carried out. Six bacterial phyla were detected in the S. papyracea microbiome: Verrucomicrobia, Cyanobacteria, Actinobacteria, Bacteroidetes, Proteobacteria, and Planctomycetes. Unlike the microbial associations of other freshwater sponges, the community under study was dominated by the phylaVerrucomicrobia (42.1%) and Cyanobacteria (17.5%), while the proportion of the Proteobacteria was unusually low (9.7%). In the S. papyracea community metagenome, there were identified 18 polyketide synthases genes fragments, the closest homologues of which included the polyketide synthases of the microorganisms belonging to the bacterial phyla Cyanobacteria, Proteobacteria (classes Betaproteobacteria, Deltaproteobacteria, and Gammaproteobacteria), and Acidobacteria as well as the eukaryotic algae of the phylum Heterokonta (class Eustigmatophyceae). Polyketide synthase sequences from S. papyracea formed three groups on the phylogenetic tree: a group of hybrid NRPS/PKS complexes, a group of cyanobacterial polyketide synthases, and a group of homologues of the eukaryotic alga Nannochloropsis gaditana. Notably, the identified polyketide synthase genes fragments showed only a 57–88% similarity to the sequences from the databases, which implies the presence of genes controlling the synthesis of the novel, still unstudied, polyketide compounds in the S. papyracea community. It was proposed that the habitat conditions of S. papyracea affect the taxonomic composition of the microorganisms associated with the sponge, including the diversity of the producers of secondary metabolites.


Lake Baikal freshwater sponges Swartschewskia papyracea microbial community 16S rRNA genes polyketide synthases genes 


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  1. 1.
    Webster, N.S. and Taylor, M.W., Marine sponges and their microbial symbionts: love and other relationships, Environ. Microbiol., 2012, vol. 14, no. 2, pp. 335–346.CrossRefPubMedGoogle Scholar
  2. 2.
    Thomas, T.R., Kavlekar, D.P., and LokaBharathi, P.A., Marine drugs from sponge microbe association—a review, Mar. Drugs, 2010, vol. 8, pp. 1417–1468.CrossRefPubMedPubMedCentralGoogle Scholar
  3. 3.
    Efremova, S.M., Sponges, in Annotirovannyi spisok fauny ozera Baikal i ego vodosbornogo basseina (An Annotated List of the Fauna of Lake Baikal and Its Catchment Area), Timoshkin, O.A., Ed., Novosibirsk: Nauka, 2001, vol. 1, pp. 177–190.Google Scholar
  4. 4.
    Kozhov, M.M., Biologiya ozera Baikal (Biology of Lake Baikal), Moscow: Akad. Nauk SSSR, 1962.Google Scholar
  5. 5.
    Kaluzhnaya, O.V., Krivich, A.A., and Itskovich, V.B., Diversity of 16S rRNA genes in metagenomic community of the freshwater sponge Lubomirskia baicalensis, Russ. J. Genet., 2012, vol. 48, no. 8, pp. 855–858.CrossRefGoogle Scholar
  6. 6.
    Kaluzhnaya, O.V. and Itskovich, V.B., Phylogenetic diversity of microorganisms associated with the deepwater sponge Baikalospongia intermedia, Russ. J. Genet., 2014, vol. 50, no. 7, pp. 667–676.CrossRefGoogle Scholar
  7. 7.
    Gladkikh, A.S., Kaluzhnaya, O.V., Belykh, O.I., et al., Analysis of bacterial communities of two Lake Baikal endemic sponge species, Microbiology (Moscow), 2014, vol. 83, no. 6, pp. 787–797.CrossRefGoogle Scholar
  8. 8.
    Kaluzhnaya, O.V., Kulakova, N.V., and Itskovich, V.B., Diversity of polyketide synthase (PKS) genes in metagenomic community of freshwater sponge Lubomirskia baicalensis, Mol. Biol. (Moscow), 2012, vol. 46, no. 6, pp. 790–795.CrossRefGoogle Scholar
  9. 9.
    Kaluzhnaya, O.V., Lipko, I.A., Itskovich, V.B., et al., PCR-screening of bacteria isolated from freshwater sponge Lubomirskia baicalensis for detection of genes of secondary metabolites, Voda: Khim. Ekol., 2013, no. 7, pp. 70–74.Google Scholar
  10. 10.
    Lipko (Terkina), I.A., Kaluzhnaya, O.V., Kravchenko, O.S., and Parfenova, V.V., Identification of polyketide synthase genes in genome of Pseudomonas fluorescens strain 28Bb-06 from freshwater sponge Baikalospongia bacillifera, Mol. Biol. (Moscow), 2012, vol. 46, no. 4. pp. 609–611.CrossRefGoogle Scholar
  11. 11.
    Schröder, H.C., Efremova, S.M., Itskovich, V.B., et al., Molecular phylogeny of the freshwater sponges in Lake Baikal, J. Zool. Syst. Evol. Res., 2003, vol. 41, pp. 80–86.CrossRefGoogle Scholar
  12. 12.
    Itskovich, V., Gontcharov, A., Masuda, Y., et al., Ribosomal ITS sequences allow resolution of freshwater sponge phylogeny with alignments guided by secondary structure prediction, J. Mol. Evol., 2008, vol. 67, pp. 608–620.CrossRefPubMedGoogle Scholar
  13. 13.
    Rezvoi, P.D., Prenovodnye gubki (Freshwater Sponges), vol. 2, no. 2 of Fauna SSSR: gubki (Fauna of the Soviet Union: Sponges), Zernov S.A., Ed., Moscow: Akad. Nauk SSSR, 1936.Google Scholar
  14. 14.
    Masuda, Y., Studies on the taxonomy and distribution of freshwater sponges in Lake Baikal, Prog. Mol. Subcell. Biol., 2009, vol. 47, pp. 81–110.CrossRefPubMedGoogle Scholar
  15. 15.
    Timoshkin, O.A., Atlas i opredelitel’ pelagobiontov Baikala (s kratkimi ocherkami po ikh ekologii) (Atlas and Identification Book of Pellagobionts of Lake Baikal (with Brief Essays on Their Ecology)), Timoshkin, O.A., Mazepova, G.F., Mel’nik, N.G., et al., Eds., Novosibirsk: Nauka, 1995.Google Scholar
  16. 16.
    Pile, A.J., Patterson, M.R., Savarese, M., et al., Trophic effects of sponge feeding within Lake Baikal’s littoral zone: 2. Sponge abundance, diet, feeding efficiency, and carbon flux, Limnol. Oceonogr., 1997, vol. 42, no. 1, pp. 178–184.CrossRefGoogle Scholar
  17. 17.
    Hochmuth, T. and Piel, J., Polyketide synthases of bacterial symbionts in sponges—evolution-based applications in natural products research, Phytochemistry, 2009, vol. 70, nos. 15–16, pp. 1841–1849.CrossRefPubMedGoogle Scholar
  18. 18.
    Esteves, A.I., Hardoim, C.C., Xavier, J.R., et al., Molecular richness and biotechnological potential of bacteria cultured from Irciniidae sponges in the northeast Atlantic, FEMS Microbiol. Ecol., 2013, vol. 85, no. 3, pp. 519–536.CrossRefPubMedGoogle Scholar
  19. 19.
    Della Sala, G., Hochmuth, T., Teta, R., et al., Polyketide synthases in the microbiome of the marine sponge Plakortis halichondrioides: a metagenomic update, Mar. Drugs, 2014, vol. 12, pp. 5425–5440.CrossRefPubMedPubMedCentralGoogle Scholar
  20. 20.
    Santos, O.C., Soares, A.R., Machado, F.L., et al., Investigation of biotechnological potential of spongeassociated bacteria collected in Brazilian coast, Lett. Appl. Microbiol., 2015, vol. 60, no. 2, pp. 140–147.CrossRefPubMedGoogle Scholar
  21. 21.
    Jenke-Kodama, H. and Dittmann, E., Evolution of metabolic diversity: insights from microbial polyketide synthases, Phytochem., 2009, vol. 70, pp. 1858–1866.CrossRefGoogle Scholar
  22. 22.
    Barrios-Llerena, M.E., Burja, A.M., and Wright, P.C., Genetic analysis of polyketide synthase and peptide synthetase genes in cyanobacteria as a mining tool for secondary metabolites, J. Ind. Microbiol. Biotechnol., 2007, vol. 34, pp. 443–456.CrossRefPubMedGoogle Scholar
  23. 23.
    Hall, T.A., BioEdit: a user-friendly biological sequence alignment editor and analysis program for Windows 95/98/NT, Nucleic Acids. Symp., 1999, vol. 41, pp. 95–98.Google Scholar
  24. 24.
    Ashelford, K.E., Chuzhanova, N.A., Fry, J.C., et al., New screening software shows that most recent large 16S rRNA gene clone libraries contain chimeras, Appl. Environ. Microbiol., 2006, vol. 72, no. 9, pp. 5734–5741.CrossRefPubMedPubMedCentralGoogle Scholar
  25. 25.
    Altschul, S.F., Warren, G., Miller, W., et al., Basic local alignment search tool, J. Mol. Biol., 1990, vol. 215, pp. 403–410.CrossRefPubMedGoogle Scholar
  26. 26.
    Tamura, K., Dudley, J., Nei, M., and Kumar, S., MEGA4: molecular genetics analysis (MEGA) software version 4.0, Mol. Biol. Evol., 2007, vol. 24, pp. 1596–1599.CrossRefPubMedGoogle Scholar
  27. 27.
    Newton, R.J., Jones, S.E., Eiler, A., et al., A guide to the natural history of freshwater lake bacteria, Microb. Mol. Biol. Rev., 2011, vol. 75, no. 1, pp. 14–49.CrossRefGoogle Scholar
  28. 28.
    Lee, K.-C., Webb, R.I., Janssen, P.H., et al., Phylum Verrucomicrobia representatives share a compartmentalized cell plan with members of bacterial phylum Planctomycetes, BMC Microbiol., 2009, vol. 9, no. 5. Scholar
  29. 29.
    Cardman, Z., Arnosti, C., Durbin, A., et al., Verrucomicrobia are candidates for polysaccharide-degrading bacterioplankton in an Arctic fjord of Svalbard, Appl. Environ. Microbiol., 2014, vol. 80, no. 12, pp. 3749–3756.CrossRefPubMedPubMedCentralGoogle Scholar
  30. 30.
    Zhang, J., Zhang, X., Liu, Y., et al., Bacterioplankton communities in a high-altitude freshwater wetland, Ann. Microbiol., 2014, vol. 64, no. 3, pp. 1405–1411.CrossRefGoogle Scholar
  31. 31.
    Parfenova, V.V., Gladkikh, A.S., and Belykh, O.I., Comparative analysis of biodiversity in the planktonic and biofilm bacterial communities in Lake Baikal, Microbiology (Moscow), 2013, vol. 82, no. 1, pp. 91–101.CrossRefGoogle Scholar
  32. 32.
    Costa, R., Keller-Costa, T., Gomes, N.C.M., et al., Evidence for selective bacterial community structuring in the freshwater sponge Ephydatia fluviatilis, Microb. Ecol., 2013, vol. 65, pp. 232–244.CrossRefPubMedGoogle Scholar
  33. 33.
    King, G.M., Smith, C.B., Tolar, B., and Hollibaugh, J.T., Analysis of composition and structure of coastal to mesopelagic bacterioplankton communities in the northern Gulf of Mexico, Front. Microbiol., 2013, vol. 3, no. 438. Scholar
  34. 34.
    Arnds, J., Knittel, K., Buck, U., et al., Development of a 16S rRNA-targeted probe set for Verrucomicrobia and its application for fluorescence in situ hybridization in a humic lake, Syst. Appl. Microbiol., 2010, vol. 33, pp. 139–148.CrossRefPubMedGoogle Scholar
  35. 35.
    Gernert, C., Glockner, F.O., Krohne, G., et al., Microbial diversity of the freshwater sponge Spongilla lacustris, Microb. Ecol., 2005, vol. 50, pp. 206–212.CrossRefPubMedGoogle Scholar
  36. 36.
    Belykh, O.I., Pomazkina, G.V., Tikhonova, I.V., and Tomberg, I.V., Characteristic of summer phytoplankton and autotrophic picoplankton of Lake Baikal in 2005, Algologiya, 2007, vol. 17, pp. 380–396.Google Scholar
  37. 37.
    Lemloh, M.-L., Fromont, J., Brümmer, F., and Usher, K.L., Diversity and abundance of photosynthetic sponges in temperate Western Australia, BMC Ecol., 2009, vol. 9, no. 4. Scholar
  38. 38.
    Erwin, P.M., López-Legentil, S., Gonzalez-Pech, R., and Turon, X., A specific mix of generalists: bacterial symbionts in mediterranean Ircinia spp., FEMS Microb. Ecol., 2012, vol. 79, no. 3, pp. 619–637.CrossRefGoogle Scholar
  39. 39.
    Ghai, R., Mizuno, C.M., Picazo, A., et al., Key roles for freshwater Actinobacteria revealed by deep metagenomic sequencing, Mol. Ecol., 2014, vol. 23, no. 24, pp. 6073–6090.CrossRefPubMedGoogle Scholar
  40. 40.
    Jezbera, J., Sharma, A.K., Brandt, U., et al., “Candidatus Planktophila limnetica,” an actinobacterium representing one of the most numerically important taxa in freshwater bacterioplankton, Int. J. Syst. Evol. Microbiol., 2009, vol. 59, pp. 2864–2869.CrossRefPubMedGoogle Scholar
  41. 41.
    Karlov, D.S., Mari, D., Chuvochina, M.S., et al., Microbial communities of water column of Lake Radok, East Antarctica, dominated by abundant actinobacterium “Candidatus Planktophila limnetica”, Microbiology (Moscow), 2011, vol. 80, no. 4, pp. 576–579.CrossRefGoogle Scholar
  42. 42.
    Zwart, G., Crump, B., Agterveld, M., et al., Typical freshwater bacteria: an analysis of available 16S rRNA gene sequences from plankton of lakes and rivers, Aquat. Microb. Ecol., 2002, vol. 28, pp. 141–155.CrossRefGoogle Scholar
  43. 43.
    Qu, J.-H. and Yuan, H.-L., Sediminibacterium salmoneum gen. nov., sp. nov., a member of the phylum Bacteroidetes isolated from sediment of a eutrophic reservoir, Int. J. Syst. Evol. Microbiol., 2008, vol. 58, pp. 2191–2194.CrossRefPubMedGoogle Scholar
  44. 44.
    Fernandez-Gomez, B., Richter, M., Schüler, M., et al., Ecology of marine Bacteroidetes: a comparative genomics approach, ISME J., 2013, vol. 7, pp. 1026–1037.CrossRefPubMedPubMedCentralGoogle Scholar
  45. 45.
    Wu, J., Gao, W., Johnson, R.H., et al., Integrated metagenomic and metatranscriptomic analyses of microbial communities in the mesoand bathypelagic realm of North Pacific Ocean, Mar. Drugs, 2013, vol. 11, pp. 3777–3801.CrossRefPubMedPubMedCentralGoogle Scholar
  46. 46.
    Villaescusa, J.A., Casamayora, E.O., Rochera, C., et al., Heterogeneous vertical structure of the bacterioplankton community in a non-stratified Antarctic lake, Antarct. Sci., 2013, vol. 25, no. 2, pp. 229–238.CrossRefGoogle Scholar
  47. 47.
    Taylor, M.W., Radax, R., Steger, D., and Wagner, M., Sponge-associated microorganisms: evolution, ecology, and biotechnological potential, Microbiol. Mol. Biol. Rev., 2007, vol. 71, no. 2, pp. 295–347.CrossRefPubMedPubMedCentralGoogle Scholar
  48. 48.
    Kasalicky V., Jezbera J., Hahn M.W., and Šimek K., The diversity of the Limnohabitans genus, an important group of freshwater bacterioplankton, by characterization of 35 isolated strains, PLoS One, 2013, vol. 8, no. 3. e58209. doi 10.1371/journal.pone.0058209CrossRefPubMedPubMedCentralGoogle Scholar
  49. 49.
    Jezbera, J., Jezberova, J., Kasalicky, V., et al., Patterns of Limnohabitans microdiversity across a large set of freshwater habitats as revealed by reverse line blot hybridization, PLoS One, 2013, vol. 8, no. 3. e58527. doi 10.1371/journal.pone.0058527Google Scholar
  50. 50.
    Paver, S.F., Hayek, K.R., Gano, K.A., et al., Interactions between specific phytoplankton and bacteria affect lake bacterial community succession, Environ. Microbiol., 2013, vol. 15, no. 9, pp. 2489–2504.CrossRefPubMedGoogle Scholar
  51. 51.
    Ehrenreich, I., Waterbury, J., and Webb, E., Distribution and diversity of natural product genes in marine and freshwater cyanobacterial cultures and genomes, Appl. Envir. Microb., 2005, vol. 71, pp. 7401–7413.CrossRefGoogle Scholar
  52. 52.
    Edwards, D.J., Marquez, B.L., Nogle, L.M., et al., Structure and biosynthesis of the jamaicamides, new mixed polyketide-peptide neurotoxins from the marine cyanobacterium Lyngbya majuscule, Chem. Biol., 2004, vol. 11, no. 6, pp. 817–833.CrossRefPubMedGoogle Scholar
  53. 53.
    Forli, S., Epothilones: from discovery to clinical trials, Curr. Top. Med. Chem., 2014, vol. 14, no. 20, pp. 2312–2321.CrossRefPubMedPubMedCentralGoogle Scholar
  54. 54.
    Khosla, C., Tang, Y., Chen, A.Y., et al., Structure and mechanism of the 6-deoxyerythronolide B synthase, Annu. Rev. Biochem., 2007, vol. 76, pp. 195–221.CrossRefPubMedGoogle Scholar
  55. 55.
    Wang, D., Ning, K., Li, J., et al., Nannochloropsis genomes reveal evolution of microalgal oleaginous traits, PLoS Genet., 2014, vol. 10, no. 1. e1004094. doi 10.1371/journal.pgen.1004094CrossRefPubMedPubMedCentralGoogle Scholar
  56. 56.
    Szaby M., Parker, K., Guruprasad, S., et al., Photosynthetic acclimation of Nannochloropsis oculata investigated by multi-wavelength chlorophyll fluorescence analysis, Bioresour. Technol., 2014, vol. 167, pp. 521–529.CrossRefGoogle Scholar
  57. 57.
    Fietz, S., Bleiß, W., Hepperle, D., et al., First record of Nannochloropsis limnetica (Eustigmatophyceae) in the autotrophic phytoplankton from Lake Baikal, J. Phycol., 2005, vol. 41, no. 4, pp. 780–790.CrossRefGoogle Scholar
  58. 58.
    Tamburic, B., Szaby M., Tran, N.-A.T., et al., Action spectra of oxygen production and chlorophyll a fluorescence in the green microalga Nannochloropsis oculata, Bioresour. Technol., 2014, vol. 169, pp. 320–327.CrossRefPubMedGoogle Scholar
  59. 59.
    Konga, W., Reamb, D.C., Priscuc, J.C., and MorganKissb, R.M., Diversity and expression of RubisCO genes in a perennially ice-covered Antarctic lake during the polar night transition, Appl. Environ. Microbiol., 2012, vol. 78, no. 12, pp. 4358–4366.CrossRefGoogle Scholar
  60. 60.
    Simionato, D., Block, M.A., La Rocca, N., et al., The response of Nannochloropsis gaditana to nitrogen starvation includes de novo biosynthesis of triacylglycerols, a decrease of chloroplast galactolipids, and reorganization of the photosynthetic apparatus, Eukaryot. Cell, 2013, vol. 12, no. 5, pp. 665–676.CrossRefPubMedPubMedCentralGoogle Scholar
  61. 61.
    Starkenburg, S., Kwon, K.J., Jha, R.K., et al., A pangenomic analysis of the Nannochloropsis organellar genomes reveals novel genetic variations in key metabolic genes, BMC Genomics, 2014, vol. 15, no. 212. Scholar
  62. 62.
    Krienitz, L., Hepperle, D., Stich, H.-B., and Weiler, W., Nannochloropsis limnetica (Eustigmatophyceae), a new species of picoplankton from freshwater, Phycologia, 2000, vol. 39, pp. 219–227.CrossRefGoogle Scholar
  63. 63.
    Fawley, K.P. and Fawley, M.W., Observations on the diversity and ecology of freshwater Nannochloropsis (Eustigmatophyceae), with descriptions of new taxa, Protist, 2007, vol. 158, no. 3, pp. 325–336.PubMedGoogle Scholar
  64. 64.
    Kehr, J.-C., Picchi, D.G., Dittmann, E., et al., Natural product biosyntheses in cyanobacteria: a treasure trove of unique enzymes, Beilstein. J. Org. Chem., 2011, vol. 7, pp. 1622–1635.CrossRefPubMedPubMedCentralGoogle Scholar
  65. 65.
    Bil, K., Titlyanov, E., Berner, T., et al., Some aspects of the physiology and biochemistry of Lubomirskia baikalensis, a sponge from Lake Baikal containing symbiotic algae, Symbiosis, 1999, vol. 26, pp. 179–191.Google Scholar
  66. 66.
    Sukenik, A., Beardall, J., Kromkamp, J.C., et al., Photosynthetic performance of outdoor Nannochloropsis mass cultures under a wide range of environmental conditions, Aquat. Microb. Ecol., 2009, vol. 56, pp. 297–308.CrossRefGoogle Scholar
  67. 67.
    Vinyard, D.J., Gimpel, J., Ananyev, G.M., et al., Natural variants of photosystem II subunit D1 tune photochemical fitness to solar intensity, J. Biol. Chem., 2013, vol. 288, no. 8, pp. 5451–5462.CrossRefPubMedPubMedCentralGoogle Scholar

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© Pleiades Publishing, Inc. 2016

Authors and Affiliations

  1. 1.Limnological Institute, Siberian Branch of the Russian Academy of SciencesIrkutskRussia

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