Cultivation and visualization of a methanogen of the phylum Thermoproteota (original) (raw)
Thauer, R. K., Kaster, A.-K., Seedorf, H., Buckel, W. & Hedderich, R. Methanogenic archaea: ecologically relevant differences in energy conservation. Nat. Rev. Microbiol.6, 579–591 (2008). CASPubMed Google Scholar
Garcia, P. S., Gribaldo, S. & Borrel, G. Diversity and evolution of methane-related pathways in archaea. Annu. Rev. Microbiol.76, 727–755 (2022). CASPubMed Google Scholar
Borrel, G. et al. Wide diversity of methane and short-chain alkane metabolisms in uncultured archaea. Nat Microbiol4, 603–613 (2019). CASPubMedPubMed Central Google Scholar
Wang, Y., Wegener, G., Hou, J., Wang, F. & Xiao, X. Expanding anaerobic alkane metabolism in the domain of Archaea. Nat. Microbiol.4, 595–602 (2019). CASPubMed Google Scholar
Evans, P. N. et al. Methane metabolism in the archaeal phylum Bathyarchaeota revealed by genome-centric metagenomics. Science350, 434–438 (2015). ADSCASPubMed Google Scholar
Vanwonterghem, I. et al. Methylotrophic methanogenesis discovered in the archaeal phylum Verstraetearchaeota. Nat. Microbiol.1, 1–9 (2016). Google Scholar
Saunois, M. et al. The Global Methane Budget 2000-2017. Earth Syst. Sci. Data12, 1561–1623 (2020). ADS Google Scholar
Conrad, R. The global methane cycle: recent advances in understanding the microbial processes involved. Environ. Microbiol. Rep.1, 285–292 (2009). CASPubMed Google Scholar
Thauer, R. K. Methyl (Alkyl)-Coenzyme M reductases: nickel F-430-containing enzymes involved in anaerobic methane formation and in anaerobic oxidation of methane or of short chain alkanes. Biochemistry58, 5198–5220 (2019). CASPubMed Google Scholar
Evans, P. N. et al. An evolving view of methane metabolism in the Archaea. Nat. Rev. Microbiol.17, 219–232 (2019). CASPubMed Google Scholar
Stephenson, M. & Stickland, L. H. Hydrogenase: the bacterial formation of methane by the reduction of one-carbon compounds by molecular hydrogen. Biochem. J.27, 1517–1527 (1933). CASPubMedPubMed Central Google Scholar
Rinke, C. et al. A standardized archaeal taxonomy for the Genome Taxonomy Database. Nat. Microbiol.6, 946–959 (2021). CASPubMed Google Scholar
McKay, L. J. et al. Co-occurring genomic capacity for anaerobic methane metabolism and dissimilatory sulfite reduction discovered in the Korarchaeota. Nat. Microbiol.4, 614–622 (2019). CASPubMed Google Scholar
McKay, L. J., Hatzenpichler, R., Inskeep, W. P. & Fields, M. W. Occurrence and expression of novel methyl-coenzyme M reductase gene (mcrA) variants in hot spring sediments. Sci. Rep.7, 7252 (2017). ADSPubMedPubMed Central Google Scholar
Hua, Z.-S. et al. Insights into the ecological roles and evolution of methyl-coenzyme M reductase-containing hot spring Archaea. Nat. Commun.10, 4574 (2019). ADSPubMedPubMed Central Google Scholar
Lynes, M. M. et al. Diversity and function of methyl-coenzyme M reductase-encoding archaea in Yellowstone hot springs revealed by metagenomics and mesocosm experiments. ISME Commun.3, 22 (2023). ADSPubMedPubMed Central Google Scholar
Buessecker, S. et al. Mcr-dependent methanogenesis in Archaeoglobaceae enriched from a terrestrial hot spring. ISME J.17, 1649–1659 (2023). CASPubMed Google Scholar
Wang, J. et al. Evidence for nontraditional mcr-containing archaea contributing to biological methanogenesis in geothermal springs. Sci. Adv.9, eadg6004 (2023). CASPubMedPubMed Central Google Scholar
Lynes, M. M., Jay, Z. J., Kohtz, A. J. & Hatzenpichler, R. Methylotrophic methanogenesis in the Archaeoglobi revealed by cultivation of Ca. Methanoglobus hypatiae from a Yellowstone hot spring. ISME J. 18, wrae026 (2024).
Liu, Y.-F. et al. Long-term cultivation and meta-omics reveal methylotrophic methanogenesis in hydrocarbon-impacted habitats. Engineering24, 264–275 (2023).
Oren, A., Garrity, G. M., Parker, C. T., Chuvochina, M. & Trujillo, M. E. Lists of names of prokaryotic Candidatus taxa. Int. J. Syst. Evol. Microbiol.70, 3956–4042 (2020). PubMed Google Scholar
Zeikus, J., Ben-Bassat, A. & Hegge, P. Microbiology of methanogenesis in thermal, volcanic environments. J. Bacteriol.143, 432–440 (1980). CASPubMedPubMed Central Google Scholar
McKay, L. J., Klingelsmith, K. B., Deutschbauer, A. M., Inskeep, W. P. & Fields, M. W. Draft genome sequence of Methanothermobacter thermautotrophicus WHS, a thermophilic hydrogenotrophic methanogen from Washburn Hot Springs in Yellowstone National Park, USA. Microbiol. Resour. Announc.10, e01157–01120 (2021). CASPubMedPubMed Central Google Scholar
Cheng, L., Dai, L., Li, X., Zhang, H. & Lu, Y. Isolation and characterization of Methanothermobacter crinale sp. nov., a novel hydrogenotrophic methanogen from the Shengli oil field. Appl. Environ. Microbiol.77, 5212–5219 (2011). ADSCASPubMedPubMed Central Google Scholar
Balk, M., Weijma, J. & Stams, A. J. Thermotoga lettingae sp. nov., a novel thermophilic, methanol-degrading bacterium isolated from a thermophilic anaerobic reactor. Int. J. Syst. Evol. Microbiol.52, 1361–1368 (2002). CASPubMed Google Scholar
Paulo, P. et al. Pathways of methanol conversion in a thermophilic anaerobic (55 C) sludge consortium. Appl. Microbiol. Biotechnol.63, 307–314 (2003). CASPubMed Google Scholar
Hatzenpichler, R., Krukenberg, V., Spietz, R. L. & Jay, Z. J. Next-generation physiology approaches to study microbiome function at single cell level. Nat. Rev. Microbiol., 18, 241–256 (2020).
Hatzenpichler, R. et al. Visualizing in situ translational activity for identifying and sorting slow-growing archaeal− bacterial consortia. Proc. Natl Acad. Sci. USA113, E4069–E4078 (2016). CASPubMedPubMed Central Google Scholar
Kohtz, A. J., Jay, Z. J., Lynes, M. M., Krukenberg, V. & Hatzenpichler, R. Culexarchaeia, a novel archaeal class of anaerobic generalists inhabiting geothermal environments. ISME Commun.2, 1–13 (2022). Google Scholar
Major, T. A., Liu, Y. & Whitman, W. B. Characterization of energy-conserving hydrogenase B in Methanococcus maripaludis. J. Bacteriol.192, 4022–4030 (2010). CASPubMedPubMed Central Google Scholar
Ma, K., Schicho, R. N., Kelly, R. M. & Adams, M. Hydrogenase of the hyperthermophile Pyrococcus furiosus is an elemental sulfur reductase or sulfhydrogenase: evidence for a sulfur-reducing hydrogenase ancestor. Proc. Natl Acad. Sci. USA90, 5341–5344 (1993). ADSCASPubMedPubMed Central Google Scholar
Lang, K. et al. New mode of energy metabolism in the seventh order of methanogens as revealed by comparative genome analysis of “Candidatus Methanoplasma termitum”. Appl. Environ. Microbiol.81, 1338–1352 (2015). ADSPubMedPubMed Central Google Scholar
Loh, H. Q., Hervé, V. & Brune, A. Metabolic potential for reductive acetogenesis and a novel energy-converting [NiFe] hydrogenase in Bathyarchaeia from termite guts–A genome-centric analysis. Front. Microbiol.11, 635786 (2021). PubMedPubMed Central Google Scholar
Kröninger, L., Berger, S., Welte, C. & Deppenmeier, U. Evidence for the involvement of two heterodisulfide reductases in the energy‐conserving system of Methanomassiliicoccus luminyensis. FEBS J.283, 472–483 (2016). PubMed Google Scholar
Kröninger, L. et al. Energy conservation in the gut microbe Methanomassiliicoccus luminyensis is based on membrane‐bound ferredoxin oxidation coupled to heterodisulfide reduction. FEBS J.286, 3831–3843 (2019). PubMed Google Scholar
Bryant, M., Campbell, L. L., Reddy, C. & Crabill, M. Growth of Desulfovibrio in lactate or ethanol media low in sulfate in association with H2-utilizing methanogenic bacteria. Appl. Environ. Microbiol.33, 1162–1169 (1977). ADSCASPubMedPubMed Central Google Scholar
McInerney, M. J. & Bryant, M. P. Anaerobic degradation of lactate by syntrophic associations of Methanosarcina barkeri and Desulfovibrio species and effect of H2 on acetate degradation. Appl. Environ. Microbiol.41, 346–354 (1981). ADSCASPubMedPubMed Central Google Scholar
Hwang, W. C. et al. LUD, a new protein domain associated with lactate utilization. BMC Bioinf.14, 1–9 (2013). Google Scholar
Young, L. N. & Villa, E. Bringing Structure to Cell Biology with Cryo-Electron Tomography. Annu. Rev. Biophys.52, 573–595 (2023). CASPubMedPubMed Central Google Scholar
Briegel, A. et al. Structural conservation of chemotaxis machinery across A rchaea and B acteria. Environ. Microbiol. Rep.7, 414–419 (2015). CASPubMed Google Scholar
Albers, S.-V. & Jarrell, K. F. The archaellum: an update on the unique archaeal motility structure. Trends Microbiol.26, 351–362 (2018). CASPubMed Google Scholar
Baidya, A. K., Bhattacharya, S., Dubey, G. P., Mamou, G. & Ben-Yehuda, S. Bacterial nanotubes: a conduit for intercellular molecular trade. Curr. Opin. Microbiol.42, 1–6 (2018). CASPubMed Google Scholar
Sivabalasarma, S. et al. Analysis of cell–cell bridges in Haloferax volcanii using electron cryo-tomography reveal a continuous cytoplasm and S-layer. Front. Microbiol.11, 612239 (2021). PubMedPubMed Central Google Scholar
Brandis, A. & Thauer, R. K. Growth of Desulfovibrio species on hydrogen and sulphate as sole energy source. Microbiology126, 249–252 (1981). CAS Google Scholar
Quast, C. et al. The SILVA ribosomal RNA gene database project: improved data processing and web-based tools. Nucleic Acids Res.41, D590–D596 (2012). PubMedPubMed Central Google Scholar
Stoecker, K., Dorninger, C., Daims, H. & Wagner, M. Double labeling of oligonucleotide probes for fluorescence in situ hybridization (DOPE-FISH) improves signal intensity and increases rRNA accessibility. Appl. Environ. Microbiol.76, 922–926 (2010). ADSCASPubMed Google Scholar
Stahl, D. A. in Nucleic Acid Techniques in Bacterial Systematics (eds Stackebrandt, E. & Goodfellow, M.) 205–248 (Wiley, 1991).
Wallner, G., Amann, R. & Beisker, W. Optimizing fluorescent in situ hybridization with rRNA‐targeted oligonucleotide probes for flow cytometric identification of microorganisms. Cytom.: J. Int. Soc. Anal. Cytol.14, 136–143 (1993). CAS Google Scholar
Daims, H. Use of fluorescence in situ hybridization and the daime image analysis program for the cultivation-independent quantification of microorganisms in environmental and medical samples. Cold Spring Harb. Protoc.2009, pdb. prot5253 (2009). PubMed Google Scholar
Daims, H., Lücker, S. & Wagner, M. Daime, a novel image analysis program for microbial ecology and biofilm research. Environ. Microbiol.8, 200–213 (2006). CASPubMed Google Scholar
Zhou, J., Bruns, M. A. & Tiedje, J. M. DNA recovery from soils of diverse composition. Appl. Environ. Microbiol.62, 316–322 (1996). ADSCASPubMedPubMed Central Google Scholar
Nurk, S., Meleshko, D., Korobeynikov, A. & Pevzner, P. A. metaSPAdes: a new versatile metagenomic assembler. Genome research27, 824–834 (2017). CASPubMedPubMed Central Google Scholar
Bushnell, B. BBMap: a fast, accurate, splice-aware aligner (Lawrence Berkeley National Lab., 2014).
Wu, Y.-W., Tang, Y.-H., Tringe, S. G., Simmons, B. A. & Singer, S. W. MaxBin: an automated binning method to recover individual genomes from metagenomes using an expectation-maximization algorithm. Microbiome2, 1–18 (2014). Google Scholar
Alneberg, J. et al. Binning metagenomic contigs by coverage and composition. Nat. Methods11, 1144–1146 (2014). CASPubMed Google Scholar
Kang, D. D. et al. MetaBAT 2: an adaptive binning algorithm for robust and efficient genome reconstruction from metagenome assemblies. PeerJ7, e7359 (2019). PubMedPubMed Central Google Scholar
Miller, I. J. et al. Autometa: automated extraction of microbial genomes from individual shotgun metagenomes. Nucleic Acids Res.47, e57–e57 (2019). CASPubMedPubMed Central Google Scholar
Sieber, C. M. et al. Recovery of genomes from metagenomes via a dereplication, aggregation and scoring strategy. Nat. Microbiol.3, 836–843 (2018). CASPubMedPubMed Central Google Scholar
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). CASPubMedPubMed Central Google Scholar
Kolmogorov, M. et al. metaFlye: scalable long-read metagenome assembly using repeat graphs. Nat. Methods17, 1103–1110 (2020). CASPubMedPubMed Central Google Scholar
Wick, R. R. & Holt, K. E. Polypolish: short-read polishing of long-read bacterial genome assemblies. PLoS Comput. Biol.18, e1009802 (2022). ADSCASPubMedPubMed Central Google Scholar
Zimin, A. V. & Salzberg, S. L. The genome polishing tool POLCA makes fast and accurate corrections in genome assemblies. PLoS Comput. Biol.16, e1007981 (2020). ADSCASPubMedPubMed Central Google Scholar
Apprill, A., McNally, S., Parsons, R. & Weber, L. Minor revision to V4 region SSU rRNA 806 R gene primer greatly increases detection of SAR11 bacterioplankton. Aquat. Microb. Ecol.75, 129–137 (2015). Google Scholar
Parada, A. E., Needham, D. M. & Fuhrman, J. A. Every base matters: assessing small subunit rRNA primers for marine microbiomes with mock communities, time series and global field samples. Environ. Microbiol.18, 1403–1414 (2016). CASPubMed Google Scholar
Reichart, N. J. et al. Activity-based cell sorting reveals responses of uncultured archaea and bacteria to substrate amendment. ISME J.14, 2851–2861 (2020). CASPubMedPubMed Central Google Scholar
Bolyen, E. et al. Reproducible, interactive, scalable and extensible microbiome data science using QIIME 2. Nat. Biotechnol.37, 852–857 (2019). CASPubMedPubMed Central Google Scholar
Callahan, B. J. et al. DADA2: high-resolution sample inference from Illumina amplicon data. Nat. Methods13, 581–583 (2016). CASPubMedPubMed Central Google Scholar
Nawrocki, E. P., Kolbe, D. L. & Eddy, S. R. Infernal 1.0: inference of RNA alignments. Bioinformatics25, 1335–1337 (2009). CASPubMedPubMed Central Google Scholar
Edgar, R. C. MUSCLE: multiple sequence alignment with high accuracy and high throughput. Nucleic Acids Res.32, 1792–1797 (2004). CASPubMedPubMed Central Google Scholar
Capella-Gutiérrez, S., Silla-Martínez, J. M. & Gabaldón, T. trimAl: a tool for automated alignment trimming in large-scale phylogenetic analyses. Bioinformatics25, 1972–1973 (2009). PubMedPubMed Central Google Scholar
Kalyaanamoorthy, S., Minh, B. Q., Wong, T. K., Von Haeseler, A. & Jermiin, L. S. ModelFinder: fast model selection for accurate phylogenetic estimates. Nat. Methods14, 587–589 (2017). CASPubMedPubMed Central Google Scholar
Guindon, S. et al. New algorithms and methods to estimate maximum-likelihood phylogenies: assessing the performance of PhyML 3.0. Syst. Biol.59, 307–321 (2010). CASPubMed Google Scholar
Katoh, K. & Standley, D. M. MAFFT multiple sequence alignment software version 7: improvements in performance and usability. Mol. Biol. Evol.30, 772–780 (2013). CASPubMedPubMed Central Google Scholar
Seemann, T. Prokka: rapid prokaryotic genome annotation. Bioinformatics30, 2068–2069 (2014). CASPubMed Google Scholar
Lu, S. et al. CDD/SPARCLE: the conserved domain database in 2020. Nucleic Acids Res.48, D265–D268 (2020). CASPubMed Google Scholar
Zimmermann, L. et al. A completely reimplemented MPI bioinformatics toolkit with a new HHpred server at its core. J. Mol. Biol.430, 2237–2243 (2018). CASPubMed Google Scholar
Chen, I.-M. A. et al. IMG/M v. 5.0: an integrated data management and comparative analysis system for microbial genomes and microbiomes. Nucleic Acids Res.47, D666–D677 (2019). ADSCASPubMed Google Scholar
Søndergaard, D., Pedersen, C. N. & Greening, C. HydDB: a web tool for hydrogenase classification and analysis. Sci. Rep.6, 1–8 (2016). Google Scholar
Patro, R., Duggal, G., Love, M. I., Irizarry, R. A. & Kingsford, C. Salmon provides fast and bias-aware quantification of transcript expression. Nat. Methods14, 417–419 (2017). CASPubMedPubMed Central Google Scholar
Ai, G., Zhu, J., Dong, X. & Sun, T. Simultaneous characterization of methane and carbon dioxide produced by cultured methanogens using gas chromatography/isotope ratio mass spectrometry and gas chromatography/mass spectrometry. Rapid Commun. Mass Spectrom.27, 1935–1944 (2013). ADSCASPubMed Google Scholar
Lagkouvardos, I. et al. IMNGS: a comprehensive open resource of processed 16S rRNA microbial profiles for ecology and diversity studies. Sci. Rep.6, 1–9 (2016). Google Scholar
Iancu, C. V. et al. Electron cryotomography sample preparation using the Vitrobot. Nat. Protoc.1, 2813–2819 (2006). CASPubMed Google Scholar
Mastronarde, D. N. Automated electron microscope tomography using robust prediction of specimen movements. J. Struct. Biol.152, 36–51 (2005). PubMed Google Scholar
Kremer, J. R., Mastronarde, D. N. & McIntosh, J. R. Computer visualization of three-dimensional image data using IMOD. J. Struct. Biol.116, 71–76 (1996). CASPubMed Google Scholar
Mastronarde, D. Correction for non‐perpendicularity of beam and tilt axis in tomographic reconstructions with the IMOD package. J. Microsc.230, 212–217 (2008). MathSciNetCASPubMed Google Scholar
Tegunov, D. & Cramer, P. Real-time cryo-electron microscopy data preprocessing with Warp. Nat. Methods16, 1146–1152 (2019). CASPubMedPubMed Central Google Scholar
Pettersen, E. F. et al. UCSF ChimeraX: Structure visualization for researchers, educators, and developers. Protein Sci.30, 70–82 (2021). CASPubMed Google Scholar
Schaible, G. A., Kohtz, A. J., Cliff, J. & Hatzenpichler, R. Correlative SIP-FISH-Raman-SEM-NanoSIMS links identity, morphology, biochemistry, and physiology of environmental microbes. ISME Commun.2, 52 (2022). PubMedPubMed Central Google Scholar
Fernando, E. Y. et al. Resolving the individual contribution of key microbial populations to enhanced biological phosphorus removal with Raman–FISH. ISME J.13, 1933–1946 (2019). CASPubMedPubMed Central Google Scholar
Majed, N. & Gu, A. Z. Application of Raman microscopy for simultaneous and quantitative evaluation of multiple intracellular polymers dynamics functionally relevant to enhanced biological phosphorus removal processes. Environ. Sci. Technol.44, 8601–8608 (2010). ADSCASPubMed Google Scholar