1. Stadnitskaia, A. et al. Molecular and carbon isotopic variability of hydrocarbon gases from mud volcanoes in the Gulf of Cadiz, NE Atlantic. Mar. Pet. Geol. 23, 281–296 (2006).

2. Mastalerz, V., de Lange, G. J. & Dahlmann, A. Differential aerobic and anaerobic oxidation of hydrocarbon gases discharged at mud volcanoes in the Nile deep-sea fan. Geochim. Cosmochim. Acta 73, 3849–3863 (2009).

3. Sassen, R. et al. Free hydrocarbon gas, gas hydrate, and authigenic minerals in chemosynthetic communities of the northern Gulf of Mexico continental slope: relation to microbial processes. Chem. Geol. 205, 195–217 (2004).

4. Adams, M. M., Hoarfrost, A. L., Bose, A., Joye, S. B. & Girguis, P. R. Anaerobic oxidation of short-chain alkanes in hydrothermal sediments: potential influences on sulfur cycling and microbial diversity. Front. Microbiol. 4, 110 (2013).

5. Bose, A., Rogers, D. R., Adams, M. M., Joye, S. B. & Girguis, P. R. Geomicrobiological linkages between short-chain alkane consumption and sulfate reduction rates in seep sediments. Front. Microbiol. 4, 386 (2013).

6. Kniemeyer, O. et al. Anaerobic oxidation of short-chain hydrocarbons by marine sulphate-reducing bacteria. Nature 449, 898–901 (2007).

7. Suarez-Zuluaga, D. A., Weijma, J., Timmers, P. H. A. & Buisman, C. J. N. High rates of anaerobic oxidation of methane, ethane and propane coupled to thiosulphate reduction. Environ. Sci. Pollut. Res. Int. 22, 3697–3704 (2015).

8. Singh, R., Guzman, M. S. & Bose, A. Anaerobic oxidation of ethane, propane, and butane by marine microbes: a mini review. Front. Microbiol. 8, 2056 (2017).

9. Laso-Pérez, R. et al. Thermophilic archaea activate butane via alkyl-coenzyme M formation. Nature 539, 396–401 (2016).

10. Dombrowski, N., Seitz, K. W., Teske, A. P. & Baker, B. J. Genomic insights into potential interdependencies in microbial hydrocarbon and nutrient cycling in hydrothermal sediments. Microbiome 5, 106 (2017).

11. Evans, P. N. et al. Methane metabolism in the archaeal phylum Bathyarchaeota revealed by genome-centric metagenomics. Science 350, 434–438 (2015).

12. McKay, L. et al. Thermal and geochemical influences on microbial biogeography in the hydrothermal sediments of Guaymas Basin, Gulf of California. Environ. Microbiol. Rep. 8, 150–161 (2016).

13. Bowles, M. W., Samarkin, V. A., Bowles, K. M. & Joye, S. B. Weak coupling between sulfate reduction and the anaerobic oxidation of methane in methane-rich seafloor sediments during ex situ incubation. Geochim. Cosmochim. Acta 75, 500–519 (2011).

14. Boetius, A. et al. A marine microbial consortium apparently mediating anaerobic oxidation of methane. Nature 407, 623–626 (2000).

15. Knittel, K. & Boetius, A. Anaerobic oxidation of methane: progress with an unknown process. Annu. Rev. Microbiol. 63, 311–334 (2009).

16. Milucka, J. et al. Zero-valent sulphur is a key intermediate in marine methane oxidation. Nature 491, 541–546 (2012).

17. Jaekel, U. et al. Anaerobic degradation of propane and butane by sulfate-reducing bacteria enriched from marine hydrocarbon cold seeps. ISME J. 7, 885–895 (2013).

18. Savage, K. N. et al. Biodegradation of low-molecular-weight alkanes under mesophilic, sulfate-reducing conditions: metabolic intermediates and community patterns. FEMS Microbiol. Ecol. 72, 485–495 (2010).

19. McGlynn, S. E., Chadwick, G. L., Kempes, C. P. & Orphan, V. J. Single cell activity reveals direct electron transfer in methanotrophic consortia. Nature 526, 531–535 (2015).

20. Wegener, G., Krukenberg, V., Riedel, D., Tegetmeyer, H. E. & Boetius, A. Intercellular wiring enables electron transfer between methanotrophic archaea and bacteria. Nature 526, 587–590 (2015).

21. Scheller, S., Goenrich, M., Boecher, R., Thauer, R. K. & Jaun, B. The key nickel enzyme of methanogenesis catalyses the anaerobic oxidation of methane. Nature 465, 606–608 (2010).

22. Thauer, R. K. Anaerobic oxidation of methane with sulfate: on the reversibility of the reactions that are catalyzed by enzymes also involved in methanogenesis from CO 2 . Curr. Opin. Microbiol. 14, 292–299 (2011).

23. Callaghan, A. V. Metabolomic investigations of anaerobic hydrocarbon-impacted environments. Curr. Opin. Biotechnol. 24, 506–515 (2013).

24. Milkov, A. V. & Etiope, G. Revised genetic diagrams for natural gases based on a global dataset of >20,000 samples. Org. Geochem. 125, 109–120 (2018).

25. Nauhaus, K., Boetius, A., Krüger, M. & Widdel, F. In vitro demonstration of anaerobic oxidation of methane coupled to sulphate reduction in sediment from a marine gas hydrate area. Environ. Microbiol. 4, 296–305 (2002).

26. Makarova, K. S., Yutin, N., Bell, S. D. & Koonin, E. V. Evolution of diverse cell division and vesicle formation systems in Archaea. Nat. Rev. Microbiol. 8, 731–741 (2010).

27. Thauer, R. K. Biochemistry of methanogenesis: a tribute to Marjory Stephenson. 1998 Marjory Stephenson Prize Lecture. Microbiology 144, 2377–2406 (1998).

28. Scheller, S., Goenrich, M., Thauer, R. K. & Jaun, B. Methyl-coenzyme M reductase from methanogenic archaea: isotope effects on label exchange and ethane formation with the homologous substrate ethyl-coenzyme M. J. Am. Chem. Soc. 135, 14985–14995 (2013).

29. Ermler, U., Grabarse, W., Shima, S., Goubeaud, M. & Thauer, R. K. Crystal structure of methyl-coenzyme M reductase: the key enzyme of biological methane formation. Science 278, 1457–1462 (1997).

30. Adam, P. S., Borrel, G. & Gribaldo, S. Evolutionary history of carbon monoxide dehydrogenase/acetyl-CoA synthase, one of the oldest enzymatic complexes. Proc. Natl Acad. Sci. USA 115, E1166–E1173 (2018).

31. Widdel, F. in Handbook of Hydrocarbon and Lipid Microbiology Ch. 298 (ed. Timmis, K. N.) 3787–3798 (Springer, Berlin Heidelberg, 2010).

32. Laso-Pérez, R., Krukenberg, V., Musat, F. & Wegener, G. Establishing anaerobic hydrocarbon-degrading enrichment cultures of microorganisms under strictly anoxic conditions. Nat. Protoc. 13, 1310–1330 (2018).

33. Cord-Ruwisch, R. A quick method for the determination of dissolved and precipitated sulfides in cultures of sulfate-reducing bacteria. J. Microbiol. Methods 4, 33–36 (1985).

34. Jaekel, U., Vogt, C., Fischer, A., Richnow, H. H. & Musat, F. Carbon and hydrogen stable isotope fractionation associated with the anaerobic degradation of propane and butane by marine sulfate-reducing bacteria. Environ. Microbiol. 16, 130–140 (2014).

35. Pruesse, E., Peplies, J. & Glöckner, F. O. SINA: accurate high-throughput multiple sequence alignment of ribosomal RNA genes. Bioinformatics 28, 1823–1829 (2012).

36. Quast, C. et al. The SILVA ribosomal RNA gene database project: improved data processing and web-based tools. Nucleic Acids Res. 41, D590–D596 (2013).

37. Bolger, A. M., Lohse, M. & Usadel, B. Trimmomatic: a flexible trimmer for Illumina sequence data. Bioinformatics 30, 2114–2120 (2014).

38. Nurk, S., Meleshko, D., Korobeynikov, A. & Pevzner, P. A. metaSPAdes: a new versatile metagenomic assembler. Genome Res. 27, 824–834 (2017).

39. Mikheenko, A., Saveliev, V. & Gurevich, A. MetaQUAST: evaluation of metagenome assemblies. Bioinformatics 32, 1088–1090 (2016).

40. Edgar, R. C. MUSCLE: multiple sequence alignment with high accuracy and high throughput. Nucleic Acids Res. 32, 1792–1797 (2004).

41. Stamatakis, A. RAxML version 8: a tool for phylogenetic analysis and post-analysis of large phylogenies. Bioinformatics 30, 1312–1313 (2014).

42. Wu, Y. W., Simmons, B. A. & Singer, S. W. MaxBin 2.0: an automated binning algorithm to recover genomes from multiple metagenomic datasets. Bioinformatics 32, 605–607 (2016).

43. Lagesen, K. et al. RNAmmer: consistent and rapid annotation of ribosomal RNA genes. Nucleic Acids Res. 35, 3100–3108 (2007).

44. Westram, R. et al. in Handbook of Molecular Microbial Ecology I: Metagenomics and Complementary Approaches (ed. de Bruijn, F. J.) 399–406 (Wiley-Blackwell, New Jersey, 2011).

45. Cao, M. D. et al. Scaffolding and completing genome assemblies in real-time with nanopore sequencing. Nat. Commun. 8, 14515 (2017).

46. Walker, B. J. et al. Pilon: an integrated tool for comprehensive microbial variant detection and genome assembly improvement. PLoS ONE 9, e112963 (2014).

47. Parks, D. H., Imelfort, M., Skennerton, C. T., Hugenholtz, P. & Tyson, G. W. CheckM: assessing the quality of microbial genomes recovered from isolates, single cells, and metagenomes. Genome Res. 25, 1043–1055 (2015).

48. Wu, M. & Scott, A. J. Phylogenomic analysis of bacterial and archaeal sequences with AMPHORA2. Bioinformatics 28, 1033–1034 (2012).

49. Overbeek, R. et al. The SEED and the rapid annotation of microbial genomes using subsystems technology (RAST). Nucleic Acids Res. 42, D206–D214 (2014).

50. Kanehisa, M., Furumichi, M., Tanabe, M., Sato, Y. & Morishima, K. KEGG: new perspectives on genomes, pathways, diseases and drugs. Nucleic Acids Res. 45, D353–D361 (2017).

51. Finn, R. D. et al. The Pfam protein families database: towards a more sustainable future. Nucleic Acids Res. 44, D279–D285 (2016).

52. Huerta-Cepas, J. et al. Fast genome-wide functional annotation through orthology assignment by eggNOG-mapper. Mol. Biol. Evol. 34, 2115–2122 (2017).

53. Lowe, T. M. & Chan, P. P. tRNAscan-SE On-line: integrating search and context for analysis of transfer RNA genes. Nucleic Acids Res. 44, W54–W57 (2016).

54. Capella-Gutiérrez, S., Silla-Martínez, J. M. & Gabaldón, T. trimAl: a tool for automated alignment trimming in large-scale phylogenetic analyses. Bioinformatics 25, 1972–1973 (2009).

55. Ludwig, W. et al. ARB: a software environment for sequence data. Nucleic Acids Res. 32, 1363–1371 (2004).

56. Pruesse, E. et al. SILVA: a comprehensive online resource for quality checked and aligned ribosomal RNA sequence data compatible with ARB. Nucleic Acids Res. 35, 7188–7196 (2007).

57. Maidak, B. L. et al. The RDP (Ribosomal Database Project) continues. Nucleic Acids Res. 28, 173–174 (2000).

58. Pernthaler, A., Pernthaler, J. & Amann, R. Fluorescence in situ hybridization and catalyzed reporter deposition for the identification of marine bacteria. Appl. Environ. Microbiol. 68, 3094–3101 (2002).

59. Pernthaler, A. & Pernthaler, J. Fluorescence in situ hybridization for the identification of environmental microbes. Methods Mol. Biol. 353, 153–164 (2007).

60. Yang, C., Kublik, A., Weidauer, C., Seiwert, B. & Adrian, L. Reductive dehalogenation of oligocyclic phenolic bromoaromatics by Dehalococcoides mccartyi strain CBDB1. Environ. Sci. Technol. 49, 8497–8505 (2015).

61. Vizcaíno, J. A. et al. 2016 update of the PRIDE database and its related tools. Nucleic Acids Res. 44, D447–D456 (2016).

62. Jariwala, F. B., Wood, R. E., Nishshanka, U. & Attygalle, A. B. Formation of the bisulfite anion (HSO 3 −, m/z 81) upon collision-induced dissociation of anions derived from organic sulfonic acids. J. Mass Spectrom. 47, 529–538 (2012).

63. Waterhouse, A. et al. SWISS-MODEL: homology modelling of protein structures and complexes. Nucleic Acids Res. 46, W296–W303 (2018).

64. Benkert, P., Tosatto, S. C. E. & Schomburg, D. QMEAN: a comprehensive scoring function for model quality assessment. Proteins 71, 261–277 (2008).

65. Stryhanyuk, H. et al. Calculation of single cell assimilation rates from SIP-NanoSIMS-derived isotope ratios: a comprehensive approach. Front. Microbiol. 9, 2342 (2018).

66. Polerecky, L. et al. Look@NanoSIMS—a tool for the analysis of nanoSIMS data in environmental microbiology. Environ. Microbiol. 14, 1009–1023 (2012).

67. Musat, N. et al. The effect of FISH and CARD–FISH on the isotopic composition of 13C- and 15N-labeled Pseudomonas putida cells measured by nanoSIMS. Syst. Appl. Microbiol. 37, 267–276 (2014).

68. Dowell, F. et al. Microbial communities in methane- and short chain alkane-rich hydrothermal sediments of Guaymas Basin. Front. Microbiol. 7, 17 (2016).

69. Welhan, J. A. & Lupton, J. E. Light hydrocarbon gases in Guaymas Basin hydrothermal fluids: thermogenic versus abiogenic origin. Bull. Am. Assoc. Petrol. Geol. 71, 215–223 (1987).

70. Orcutt, B. N. et al. Impact of natural oil and higher hydrocarbons on microbial diversity, distribution, and activity in Gulf of Mexico cold-seep sediments. Deep Sea Res. II Top. Stud. Oceanogr. 57, 2008–2021 (2010).

71. Pachiadaki, M. G., Kallionaki, A., Dählmann, A., De Lange, G. J. & Kormas, K. A. Diversity and spatial distribution of prokaryotic communities along a sediment vertical profile of a deep-sea mud volcano. Microb. Ecol. 62, 655–668 (2011).

72. Pape, T. et al. Gas hydrates in shallow deposits of the Amsterdam mud volcano, Anaximander Mountains, Northeastern Mediterranean Sea. Geo-Mar. Lett. 30, 187–206 (2010).

73. Heijs, S. K., Laverman, A. M., Forney, L. J., Hardoim, P. R. & van Elsas, J. D. Comparison of deep-sea sediment microbial communities in the Eastern Mediterranean. FEMS Microbiol. Ecol. 64, 362–377 (2008).

74. Egorov, A. V. & Ivanov, M. K. Hydrocarbon gases in sediments and mud breccia from the central and eastern part of the Mediterranean Ridge. Geo-Mar. Lett. 18, 127–138 (1998).

75. Robertson, A. H. F. et al. Collision-related break-up of a carbonate platform (Eratosthenes Seamount) and mud volcanism on the Mediterranean Ridge: preliminary synthesis and implications of tectonic results of ODP Leg 160 in the Eastern Mediterranean Sea. Geol. Soc. Spec. Publ. 131, 243–271 (1998).

76. Lloyd, K. G., Lapham, L. & Teske, A. An anaerobic methane-oxidizing community of ANME-1b archaea in hypersaline Gulf of Mexico sediments. Appl. Environ. Microbiol. 72, 7218–7230 (2006).

77. Brooks, J. M. et al. Association of gas hydrates and oil seepage in the Gulf of Mexico. Org. Geochem. 10, 221–234 (1986).

78. Maignien, L. et al. Anaerobic oxidation of methane in hypersaline cold seep sediments. FEMS Microbiol. Ecol. 83, 214–231 (2013).

79. Nuzzo, M. et al. Origin of light volatile hydrocarbon gases in mud volcano fluids, Gulf of Cadiz — evidence for multiple sources and transport mechanisms in active sedimentary wedges. Chem. Geol. 266, 350–363 (2009).

80. Roalkvam, I. et al. New insight into stratification of anaerobic methanotrophs in cold seep sediments. FEMS Microbiol. Ecol. 78, 233–243 (2011).

81. Vaular, E. N., Barth, T. & Haflidason, H. The geochemical characteristics of the hydrate-bound gases from the Nyegga pockmark field, Norwegian Sea. Org. Geochem. 41, 437–444 (2010).

82. Cruaud, P. et al. Comparative study of Guaymas Basin microbiomes: cold seeps vs. hydrothermal vents sediments. Front. Mar. Sci. 4, 417 (2017).

83. Simoneit, B. R. T., Lonsdale, P. F., Edmond, J. M. & Shanks, W. C. Deep-water hydrocarbon seeps in Guaymas Basin, Gulf of California. Appl. Geochem. 5, 41–49 (1990).

84. Case, D. H. et al. Methane seep carbonates host distinct, diverse, and dynamic microbial assemblages. MBio 6, e01348-15 (2015).

85. Milkov, A. V. et al. Co-existence of gas hydrate, free gas, and brine within the regional gas hydrate stability zone at Hydrate Ridge (Oregon margin): evidence from prolonged degassing of a pressurized core. Earth Planet. Sci. Lett. 222, 829–843 (2004).

86. Pop Ristova, P., Wenzhöfer, F., Ramette, A., Felden, J. & Boetius, A. Spatial scales of bacterial community diversity at cold seeps (Eastern Mediterranean Sea). ISME J. 9, 1306–1318 (2015).