Letter | Published:

Anaerobic oxidation of ethane by archaea from a marine hydrocarbon seep


Ethane is the second most abundant component of natural gas in addition to methane, and—similar to methane—is chemically unreactive. The biological consumption of ethane under anoxic conditions was suggested by geochemical profiles at marine hydrocarbon seeps1,2,3, and through ethane-dependent sulfate reduction in slurries4,5,6,7. Nevertheless, the microorganisms and reactions that catalyse this process have to date remained unknown8. Here we describe ethane-oxidizing archaea that were obtained by specific enrichment over ten years, and analyse these archaea using phylogeny-based fluorescence analyses, proteogenomics and metabolite studies. The co-culture, which oxidized ethane completely while reducing sulfate to sulfide, was dominated by an archaeon that we name ‘Candidatus Argoarchaeum ethanivorans’; other members were sulfate-reducing Deltaproteobacteria. The genome of Ca. Argoarchaeum contains all of the genes that are necessary for a functional methyl-coenzyme M reductase, and all subunits were detected in protein extracts. Accordingly, ethyl-coenzyme M (ethyl-CoM) was identified as an intermediate by liquid chromatography–tandem mass spectrometry. This indicated that Ca. Argoarchaeum initiates ethane oxidation by ethyl-CoM formation, analogous to the recently described butane activation by ‘Candidatus Syntrophoarchaeum’9. Proteogenomics further suggests that oxidation of intermediary acetyl-CoA to CO2 occurs through the oxidative Wood–Ljungdahl pathway. The identification of an archaeon that uses ethane (C2H6) fills a gap in our knowledge of microorganisms that specifically oxidize members of the homologous alkane series (CnH2n+2) without oxygen. Detection of phylogenetic and functional gene markers related to those of Ca. Argoarchaeum at deep-sea gas seeps10,11,12 suggests that archaea that are able to oxidize ethane through ethyl-CoM are widespread members of the local communities fostered by venting gaseous alkanes around these seeps.

Access optionsAccess options

Rent or Buy article

Get time limited or full article access on ReadCube.


All prices are NET prices.

Data availability

Metagenome sequence data are archived in the NCBI database under BioProject number PRJNA495932, including the draft genomes of Ca. Argoarchaeum ethanivorans (SAMN10235260), Eth-SRB1 (SAMN10235261) and Eth-SRB2 (SAMN10235262). The 16S rRNA gene amplicon reads have been submitted to the NCBI Sequence Read Archive (SRA) database under the accession number SRR8089822. The proteomics dataset has been deposited with the ProteomeXchange Consortium identifier PXD011597. Source Data for the quantitative growth experiments (Fig. 1a), FT–ICR–MS (Fig. 3b, c) and LC–MS/MS measurements (Fig. 3d–f) are provided. All other data are available in the paper or the Supplementary Information.

Additional information

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


  1. 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. 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. 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. 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. 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. 6.

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

  7. 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. 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. 9.

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

  10. 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. 11.

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

  12. 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. 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. 14.

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

  15. 15.

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

  16. 16.

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

  17. 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. 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. 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. 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. 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. 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 CO2. Curr. Opin. Microbiol. 14, 292–299 (2011).

  23. 23.

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

  24. 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. 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. 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. 27.

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

  28. 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. 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. 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. 31.

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

  32. 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. 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. 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. 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. 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. 37.

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

  38. 38.

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

  39. 39.

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

  40. 40.

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

  41. 41.

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

  42. 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. 43.

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

  44. 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. 45.

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

  46. 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. 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. 48.

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

  49. 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. 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. 51.

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

  52. 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. 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. 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. 55.

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

  56. 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. 57.

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

  58. 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. 59.

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

  60. 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. 61.

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

  62. 62.

    Jariwala, F. B., Wood, R. E., Nishshanka, U. & Attygalle, A. B. Formation of the bisulfite anion (HSO3 , m/z 81) upon collision-induced dissociation of anions derived from organic sulfonic acids. J. Mass Spectrom. 47, 529–538 (2012).

  63. 63.

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

  64. 64.

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

  65. 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. 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. 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. 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. 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. 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. 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. 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. 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. 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. 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. 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. 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. 78.

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

  79. 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. 80.

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

  81. 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. 82.

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

  83. 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. 84.

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

  85. 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. 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).

  87. 87.

    Zaikova, E. et al. Microbial community dynamics in a seasonally anoxic fjord: Saanich Inlet, British Columbia. Environ. Microbiol. 12, 172–191 (2010).

Download references


We acknowledge R. Appel for assistance in cultivation and chemical analyses, K. Nerlich for CARD–FISH, J. M. Kaesler for FT–ICR–MS analyses, B. Scheer for proteomics analyses and K. T. Konstantinidis for support with metagenomics analyses. Research was funded by the Helmholtz Association and by the Max Planck Society. Further financial support was provided by the Helmholtz Association (grant ERC-RA-0020 to F.M.), the Strategic Priority Research Program of the Chinese Academy of Sciences (grants XDB15020302 and XDB15020402 to Y.-G.Z.) and the Chinese Scholarship Council (scholarship to S.-C.C.). We thank the shipboard science parties and submersible operation teams of the RV Seward Johnson II. Funding for the Gulf of Mexico cruise was provided by the US National Science Foundation (OCE-0085549) and the ACS Petroleum Research Fund (PRF-36834-AC2). The US Department of Energy and the US National Undersea Research Program provided funding for submersible operations. We acknowledge the Centre for Chemical Microscopy (ProVIS) at the Helmholtz Centre for Environmental Research for the use of their analytical facilities. ProVIS is supported by European Regional Development Funds (EFRE – Europe funds Saxony).

Reviewer information

Nature thanks Mike Jetten, Derek Lovley, Rudolf K. Thauer and the other anonymous reviewer(s) for their contribution to the peer review of this work.

Author information

S.-C.C., H.-H.R., F.W. and F.M. designed the research. S.B.J. retrieved the original sediment sample and contributed to establishment of the enrichment. U.J. and F.M. performed cultivation and physiology experiments. N.M. designed CARD–FISH probes and performed CARD–FISH and fluorescence microscopy. M.S., N.S. and N.M. designed and performed helium ion microscopy analyses. O.J.L. and H.P. performed FT–ICR–MS analyses, and LC–MS/MS method development and analyses. S.-C.C., D.P., Y.-G.Z. and F.M. performed metagenomics analyses. S.-C.C. performed phylogenetic and proteomics analyses. F.C., H.S. and N.M. designed and performed nanoSIMS analyses. S.-C.C., F.W. and F.M. wrote the manuscript with contributions from all co-authors.

Competing interests

The authors declare no competing interests.

Correspondence to Florin Musat.

Extended data figures and tables

  1. Extended Data Fig. 1 Helium ion microscopy images showing vesicular structures interpreted as budding of Ca. A. ethanivorans.

    Budding cells remained loosely attached to each other, leading to formation of small clusters or aggregates. To avoid false stacking of cells, centrifugation was avoided during sample preparation. The samples were fixed and filtered on gold–palladium-sputtered polycarbonate filters. All subsequent procedures (dehydration and critical point drying) were done with filter pieces. Images are representative of n = 20 recorded images of samples from n = 3 independent cultures.

  2. Extended Data Fig. 2 Proposed pathway for ethane oxidation by Ca. Argoarchaeum based on proteogenomics analyses.

    a, Candidatus Argoarchaeum uses an MCR-like enzyme (alkyl-CoM reductase (‘ACR’)) to activate ethane to ethyl-CoM. Reactions converting the alkyl thioether to the acyl thioester are currently unknown: we hypothesize an involvement of the detected methyl-transferase (mtr; see b) in these reactions. The proposed acetyl-CoA is oxidized to CO2 through the reverse (oxidative) Wood–Ljungdahl pathway. The physiological role of acetyl-CoA synthetase (Acs) and the acetate/cation symporter (ActP) is presently unclear. All enzymes depicted are encoded by the Ca. Argoarchaeum genome. Enzymes depicted in green were fully or partly detected in protein extracts (see b). Fpo, F420H2 dehydrogenase; Ftr, formylmethanofuran:tetrahydromethanopterin formyltransferase; Fwd, formylmethanofuran dehydrogenase; Hdr, heterodisulfide reductase; Mch, methenyltetrahydromethanopterin cyclohydrolase; Mer, 5,10-methylenetetrahydromethanopterin reductase; MF, methanofuran; Mnh, multicomponent Na+:H+ antiporter; MPT, methanopterin; Mtd, methylenetetrahydromethanopterin dehydrogenase; NAD, nicotinamide adenine dinucleotide; Ntp, vacuolar/archaea (V/A)-type H+/Na+-transporting ATPase; Nuo, NADH-quinone oxidoreductase. b, Organization of the genes that encode enzymes for the pathway proposed in a. Genes for which products were detected in protein extracts are depicted in green, all other genes are shown in yellow and genes unrelated to ethane oxidation are shown in grey.

  3. Extended Data Fig. 3 NanoSIMS analysis of the Ethane12 enrichment culture.

    af, Ion images of analysed areas (representative of n = 6 recorded fields of view). b, The cell types shown in b are Ca. Argoarchaeum (cocci; indicated by the arrows), Eth-SRB1 (curved rods; indicated by small arrowheads) and Eth-SRB2 (large, oval; indicated by large arrowheads). a, d, The 12C14N ion image was used as indicator of biomass (all cell types). b, c, e, f, Images of 32S (b, e) and 32S:12C14N ratios (c, f) show that cells of Ca. Argoarchaeum are enriched in sulfur, compared with bacterial cells—similar to AOM consortia16. Regions of interest used to define cells were drawn based on the single ion images (12C14N and 32S) and superimposed on the ratio images. g, Relative abundance of sulfur in Ca. Argoarchaeum compared to the sulfate-reducing bacteria at three different incubation times (95, 110 and 120 days). The average relative sulfur content of Ca. Argoarchaeum was more than twofold higher than in Eth-SRB1 and Eth-SRB2. Each dot represents the S:CN ion ratio of a single cell; in total, over 650 cells were analysed (517 of Ca. Argoarchaeum, 58 of Eth-SRB1 and 105 of Eth-SRB2). The box plots show the total ion ratio range (vertical line with whiskers), the clustering of 50% of all cells analysed (box) and the mean value for all cells of each strain. To calculate the ratio values in g, regions of interest were drawn in the LOOK@NanoSIMS software (not shown); these were smaller than those displayed in af, to avoid inclusion of filter material in the calculation. Scale bars, 2 μm (af).

  4. Extended Data Fig. 4 Homology models of MCR from Ca. A. ethanivorans and Ca. S butanivorans.

    a, Sequence alignment of the α subunit (McrA) and β subunit (McrB) of MCR from methanogens or ANME-1 methanotrophs (green), Ca. A. ethanivorans (AEth_00344, red) and Ca. S. butanivorans (SBU_000314, SBU_000718, SBU_001343 and SBU_001328; red); the functionally conserved residues in McrA and McrB are highlighted (yellow). The numbers below the alignment indicate the residue position in M. marburgensis MCR29M. barkeri, Methanosarcina barkeri; M. formicicus, Methanotorris formicicus; M. kandleri, Methanopyrus kandleri; M. thermauto, Methanothermobacter thermautotrophicus; M. thermolitho, Methanothermococcus thermolithotrophicus; M. wolfeii, Methanothermobacter wolfeii. b. Crystal structure of M. marburgensis MCR (Protein Data Bank (PDB) accession 1MRO) bound to coenzyme B (CoB) and coenzyme M (CoM). cg, Modelled active site regions in the MCR-like enzymes of Ca. A. ethanivorans (c) and Ca. S. butanivorans (dg). The predicted structure was superimposed on M. marburgensis MCR (blue wireframes in cg). Residues in McrA and McrB are indicated as green and cyan sticks, respectively; CoB and CoM are shown as yellow sticks, coenzyme F430 as pale yellow sticks and the arginine residue coordinating CoM as grey sticks (bg).

  5. Extended Data Table 1 Quantification of the anaerobic consumption of ethane, and sulfate reduction to sulfide in the Ethane12 enrichment culture
  6. Extended Data Table 2 Genes and proteins involved in ethane activation and complete oxidation by Ca. A. ethanivorans
  7. Extended Data Table 3 Genes and proteins involved in the energy metabolism of Ca. A. ethanivorans
  8. Extended Data Table 4 Genes that encode type IV pili in the genomes of Eth-SRB1 and Eth-SRB2
  9. Extended Data Table 5 Genes that encode cytochromes in the genomes of Eth-SRB1 and Eth-SRB2
  10. Extended Data Table 6 Environmental distribution of phylotypes related to Ca. A. ethanivorans

Supplementary information

  1. Supplementary Information

    This file contains Supplementary Tables 1–6 and Supplementary References.

  2. Reporting Summary

Source data

  1. Source Data Fig. 1a

  2. Source Data Fig. 3b–f

Rights and permissions

To obtain permission to re-use content from this article visit RightsLink.

About this article

Publication history

  • Received

  • Accepted

  • Published

  • Issue Date



Further reading

Fig. 1: Ethane oxidation with sulfate and major 16S rRNA gene phylotypes.
Fig. 2: Microscopic characterization of the Ethane12 culture.
Fig. 3: Phylogeny of McrA and identification of ethyl-coM.
Fig. 4: Basic reactions in archaeal oxidation of gaseous alkanes compared to methanogenesis.
Extended Data Fig. 1: Helium ion microscopy images showing vesicular structures interpreted as budding of Ca. A. ethanivorans.
Extended Data Fig. 2: Proposed pathway for ethane oxidation by Ca. Argoarchaeum based on proteogenomics analyses.
Extended Data Fig. 3: NanoSIMS analysis of the Ethane12 enrichment culture.
Extended Data Fig. 4: Homology models of MCR from Ca. A. ethanivorans and Ca. S butanivorans.


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.