Nitrous oxide respiration in acidophilic methanotrophs.
Journal
Nature communications
ISSN: 2041-1723
Titre abrégé: Nat Commun
Pays: England
ID NLM: 101528555
Informations de publication
Date de publication:
18 May 2024
18 May 2024
Historique:
received:
03
01
2024
accepted:
22
04
2024
medline:
19
5
2024
pubmed:
19
5
2024
entrez:
18
5
2024
Statut:
epublish
Résumé
Aerobic methanotrophic bacteria are considered strict aerobes but are often highly abundant in hypoxic and even anoxic environments. Despite possessing denitrification genes, it remains to be verified whether denitrification contributes to their growth. Here, we show that acidophilic methanotrophs can respire nitrous oxide (N
Identifiants
pubmed: 38762502
doi: 10.1038/s41467-024-48161-z
pii: 10.1038/s41467-024-48161-z
doi:
Substances chimiques
nitrous oxide reductase
0
methane monooxygenase
0
Types de publication
Journal Article
Langues
eng
Sous-ensembles de citation
IM
Pagination
4226Informations de copyright
© 2024. The Author(s).
Références
IPCC. Summary for Policymakers. In: Climate Change 2021 – The Physical Science Basis: Working Group I Contribution to the Sixth Assessment Report of the Intergovernmental Panel on Climate Change (ed Intergovernmental Panel on Climate C). (Cambridge University Press, 2023).
Forster, P. et al. The Earth’s Energy Budget, Climate Feedbacks and Climate Sensitivity. In Climate Change 2021: The Physical Science Basis: Working Group I Contribution to the Sixth Assessment Report of the Intergovernmental Panel on Climate Change (eds Masson-Delmotte, V., P. Zhai, A. Pirani, S.L. Connors, C. Péan, S. Berger, N. Caud, Y. Chen, L. Goldfarb, M.I. Gomis, M. Huang, K. Leitzell, E. Lonnoy, J.B.R. Matthews, T.K. Maycock, T. Waterfield, O. Yelekçi, R. Yu, and B. Zhou) (Cambridge University Press United Kingdom and New York, NY, USA, 2023).
Prinn, R. G. et al. Evidence for variability of atmospheric hydroxyl radicals over the past quarter century. Geophys. Res. Lett. 32, L07809 (2005).
doi: 10.1029/2004GL022228
Myhre, G. et al. Anthropogenic and natural radiative forcing. in Climate change 2013: The physical science basis. Contribution of working group I to the fifth assessment report of the Intergovernmental Panel on Climate Change (eds Stocker T. F., et al.) (Cambridge University Press, 2013).
Szopa, S. et al. Short-Lived Climate Forcers. In: Climate Change 2021: The Physical Science Basis. Contribution of Working Group I to the Sixth Assessment Report of the Intergovernmental Panel on Climate Change (eds Masson-Delmotte V., et al.) (Cambridge University Press, 2021).
Prather, M. J. et al. Measuring and modeling the lifetime of nitrous oxide including its variability. J. Geophys. Res. Atmos. 120, 5693–5705 (2015).
pubmed: 26900537
pmcid: 4744722
doi: 10.1002/2015JD023267
Ravishankara, A. R., Daniel, J. S. & Portmann, R. W. Nitrous oxide (N
pubmed: 19713491
doi: 10.1126/science.1176985
Montzka, S. A., Dlugokencky, E. J. & Butler, J. H. Non-CO
pubmed: 21814274
doi: 10.1038/nature10322
Canadell, J. G. et al. Global Carbon and Other Biogeochemical Cycles and Feedbacks. In Climate Change 2021:The Physical Science Basis: Working Group I Contribution to the Sixth Assessment Report of the Intergovernmental Panel on Climate Change (eds Masson-Delmotte, V., P. Zhai, A. Pirani, S.L. Connors, C. Péan, S. Berger, N. Caud, Y. Chen, L. Goldfarb, M.I. Gomis, M. Huang, K. Leitzell, E. Lonnoy, J.B.R. Matthews, T.K. Maycock, T. Waterfield, O. Yelekçi, R. Yu, and B. Zhou) (Cambridge University Press, 2021).
Butterbach-Bahl, K., Baggs, E. M., Dannenmann, M., Kiese, R. & Zechmeister-Boltenstern, S. Nitrous oxide emissions from soils: how well do we understand the processes and their controls? Philos. Trans. R. Soc. Lond. B Biol. Sci. 368, 20130122 (2013).
pubmed: 23713120
pmcid: 3682742
doi: 10.1098/rstb.2013.0122
Ringeval, B. et al. Climate-CH
doi: 10.5194/bg-8-2137-2011
Beaulieu, J. J., DelSontro, T. & Downing, J. A. Eutrophication will increase methane emissions from lakes and impoundments during the 21st century. Nat. Commun. 10, 1375 (2019).
pubmed: 30914638
pmcid: 6435651
doi: 10.1038/s41467-019-09100-5
Murrell, J. C. & Jetten, M. S. The microbial methane cycle. Environ. Microbiol. Rep. 1, 279–284 (2009).
pubmed: 23765880
doi: 10.1111/j.1758-2229.2009.00089.x
Conrad, R. The global methane cycle: recent advances in understanding the microbial processes involved. Environ. Microbiol. Rep. 1, 285–292 (2009).
pubmed: 23765881
doi: 10.1111/j.1758-2229.2009.00038.x
Bürgmann, H. Methane oxidation (aerobic). in Encyclopedia of Geobiology, (eds Reitner, J., Thiel, V.) (Springer Netherlands, 2011).
Leu, A. O. et al. Anaerobic methane oxidation coupled to manganese reduction by members of the Methanoperedenaceae. ISME J. 14, 1030–1041 (2020).
pubmed: 31988473
pmcid: 7082337
doi: 10.1038/s41396-020-0590-x
Haroon, M. F. et al. Anaerobic oxidation of methane coupled to nitrate reduction in a novel archaeal lineage. Nature 500, 567 (2013).
pubmed: 23892779
doi: 10.1038/nature12375
Scheller, S., Yu, H., Chadwick, G. L., McGlynn, S. E. & Orphan, V. J. Artificial electron acceptors decouple archaeal methane oxidation from sulfate reduction. Science 351, 703–707 (2016).
pubmed: 26912857
doi: 10.1126/science.aad7154
Ettwig, K. F. et al. Archaea catalyze iron-dependent anaerobic oxidation of methane. Proc. Natl Acad. Sci. USA 113, 12792–12796 (2016).
pubmed: 27791118
pmcid: 5111651
doi: 10.1073/pnas.1609534113
Ettwig, K. F. et al. Nitrite-driven anaerobic methane oxidation by oxygenic bacteria. Nature 464, 543–548 (2010).
pubmed: 20336137
doi: 10.1038/nature08883
Kits, K. D., Campbell, D. J., Rosana, A. R. & Stein, L. Y. Diverse electron sources support denitrification under hypoxia in the obligate methanotroph Methylomicrobium album strain BG8. Front. Microbiol. 6, 1072–1072 (2015).
pubmed: 26500622
pmcid: 4594100
doi: 10.3389/fmicb.2015.01072
Kits, K. D., Klotz, M. G. & Stein, L. Y. Methane oxidation coupled to nitrate reduction under hypoxia by the Gammaproteobacterium Methylomonas denitrificans, sp. nov. type strain FJG1. Environ. Microbiol. 17, 3219–3232 (2015).
pubmed: 25580993
doi: 10.1111/1462-2920.12772
Dam, B., Kube, M., Dam, S., Reinhardt, R. & Liesack, W. Complete sequence analysis of two methanotroph-specific repABC-containing plasmids from Methylocystis sp. strain SC2. Appl. Environ. Microbiol. 78, 4373–4379 (2012).
pubmed: 22504811
pmcid: 3370566
doi: 10.1128/AEM.00628-12
Kox, M. A. R. et al. Complete genome sequence of the aerobic facultative methanotroph Methylocella tundrae strain T4. Microbiol. Resour. Announc. 8, e00286–00219 (2019).
pubmed: 31097502
pmcid: 6522787
doi: 10.1128/MRA.00286-19
Tian, H. et al. A comprehensive quantification of global nitrous oxide sources and sinks. Nature 586, 248–256 (2020).
pubmed: 33028999
doi: 10.1038/s41586-020-2780-0
Thomson, A. J., Giannopoulos, G., Pretty, J., Baggs, E. M. & Richardson, D. J. Biological sources and sinks of nitrous oxide and strategies to mitigate emissions. Philos. Trans. R. Soc. Lond. B Biol. Sci. 367, 1157–1168 (2012).
pubmed: 22451101
pmcid: 3306631
doi: 10.1098/rstb.2011.0415
Buessecker, S. et al. Coupled abiotic-biotic cycling of nitrous oxide in tropical peatlands. Nat. Ecol. Evol. 6, 1881–1890 (2022).
pubmed: 36202923
doi: 10.1038/s41559-022-01892-y
Su, Q., Domingo-Félez, C., Jensen, M. M. & Smets, B. F. Abiotic nitrous oxide (N
pubmed: 30816038
doi: 10.1021/acs.est.8b06193
Zumft, W. G. & Kroneck, P. M. Respiratory transformation of nitrous oxide (N
pubmed: 17027372
doi: 10.1016/S0065-2911(06)52003-X
Graf, D. R., Jones, C. M. & Hallin, S. Intergenomic comparisons highlight modularity of the denitrification pathway and underpin the importance of community structure for N
pubmed: 25436772
pmcid: 4250227
doi: 10.1371/journal.pone.0114118
Sanford, R. A. et al. Unexpected nondenitrifier nitrous oxide reductase gene diversity and abundance in soils. Proc. Natl Acad. Sci. USA 109, 19709–19714 (2012).
pubmed: 23150571
pmcid: 3511753
doi: 10.1073/pnas.1211238109
Hallin, S., Philippot, L., Löffler, F. E., Sanford, R. A. & Jones, C. M. Genomics and ecology of novel N
pubmed: 28803698
doi: 10.1016/j.tim.2017.07.003
Payne, W. J., Grant, M. A., Shapleigh, J. & Hoffman, P. Nitrogen oxide reduction in Wolinella succinogenes and Campylobacter species. J. Bacteriol. 152, 915–918 (1982).
pubmed: 7130133
pmcid: 221551
doi: 10.1128/jb.152.2.915-918.1982
Yoon, S., Nissen, S., Park, D., Sanford, R. A. & Löffler, F. E. Nitrous oxide reduction kinetics distinguish bacteria harboring clade I NosZ from those harboring clade II NosZ. Appl. Environ. Microbiol. 82, 3793–3800 (2016).
pubmed: 27084012
pmcid: 4907195
doi: 10.1128/AEM.00409-16
Park, D., Kim, H. & Yoon, S. Nitrous oxide reduction by an obligate aerobic bacterium, Gemmatimonas aurantiaca strain T-27. Appl. Environ. Microbiol. 83, e00502–00517 (2017).
pubmed: 28389533
pmcid: 5452805
doi: 10.1128/AEM.00502-17
Dam, B., Dam, S., Blom, J. & Liesack, W. Genome analysis coupled with physiological studies reveals a diverse nitrogen metabolism in Methylocystis sp. strain SC2. PLOS ONE 8, e74767 (2013).
pubmed: 24130670
pmcid: 3794950
doi: 10.1371/journal.pone.0074767
Knief, C. Diversity and habitat preferences of cultivated and uncultivated aerobic methanotrophic bacteria evaluated based on pmoA as molecular marker. Front. Microbiol. 6, 1346 (2015).
pubmed: 26696968
pmcid: 4678205
doi: 10.3389/fmicb.2015.01346
Reim, A., Lüke, C., Krause, S., Pratscher, J. & Frenzel, P. One millimetre makes the difference: high-resolution analysis of methane-oxidizing bacteria and their specific activity at the oxic–anoxic interface in a flooded paddy soil. ISME J. 6, 2128–2139 (2012).
pubmed: 22695859
pmcid: 3475382
doi: 10.1038/ismej.2012.57
Farhan Ul Haque, M., Crombie, A. T. & Murrell, J. C. Novel facultative Methylocella strains are active methane consumers at terrestrial natural gas seeps. Microbiome 7, 134 (2019).
pubmed: 31585550
pmcid: 6778391
doi: 10.1186/s40168-019-0741-3
Kantor, R. S., Miller, S. E. & Nelson, K. L. The water microbiome through a pilot scale advanced treatment facility for direct potable reuse. Front. Microbiol. 10, 993 (2019).
pubmed: 31139160
pmcid: 6517601
doi: 10.3389/fmicb.2019.00993
McGuirl, M. A., Bollinger, J. A., Cosper, N., Scott, R. A. & Dooley, D. M. Expression, purification, and characterization of NosL, a novel Cu(I) protein of the nitrous oxide reductase (nos) gene cluster. J. Biol. Inorg. Chem. 6, 189–195 (2001).
pubmed: 11293413
doi: 10.1007/s007750000190
Kang, C. S., Dunfield, P. F. & Semrau, J. D. The origin of aerobic methanotrophy within the Proteobacteria. FEMS Microbiol. Lett. 366, fnz096 (2019).
pubmed: 31054238
doi: 10.1093/femsle/fnz096
Berks, B. C., Palmer, T. & Sargent, F. Protein targeting by the bacterial twin-arginine translocation (Tat) pathway. Curr. Opin. Microbiol. 8, 174–181 (2005).
pubmed: 15802249
doi: 10.1016/j.mib.2005.02.010
Awala, S. I. et al. Verrucomicrobial methanotrophs grow on diverse C3 compounds and use a homolog of particulate methane monooxygenase to oxidize acetone. ISME J. 15, 3636–3647 (2021).
pubmed: 34158629
pmcid: 8630023
doi: 10.1038/s41396-021-01037-2
Zhang, L., Trncik, C., Andrade, S. L. A. & Einsle, O. The flavinyl transferase ApbE of Pseudomonas stutzeri matures the NosR protein required for nitrous oxide reduction. Biochim. Biophys. Acta Bioenerg. 1858, 95–102 (2017).
pubmed: 27864152
doi: 10.1016/j.bbabio.2016.11.008
Honisch, U. & Zumft, W. G. Operon structure and regulation of the nos gene region of Pseudomonas stutzeri, encoding an ABC-Type ATPase for maturation of nitrous oxide reductase. J. Bacteriol. 185, 1895–1902 (2003).
pubmed: 12618453
pmcid: 150149
doi: 10.1128/JB.185.6.1895-1902.2003
Simon, J., Einsle, O., Kroneck, P. M. & Zumft, W. G. The unprecedented nos gene cluster of Wolinella succinogenes encodes a novel respiratory electron transfer pathway to cytochrome c nitrous oxide reductase. FEBS Lett. 569, 7–12 (2004).
pubmed: 15225600
doi: 10.1016/j.febslet.2004.05.060
Suzuki, M., Cui, Z. J., Ishii, M. & Igarashi, Y. Nitrate respiratory metabolism in an obligately autotrophic hydrogen-oxidizing bacterium, Hydrogenobacter thermophilus TK-6. Arch. Microbiol. 175, 75–78 (2001).
pubmed: 11271424
doi: 10.1007/s002030000230
Sharp, C. E., den Camp, H.J.M.O., Tamas, I., Dunfield P. F. Unusual members of the PVC superphylum: the methanotrophic Verrucomicrobia genus “Methylacidiphilum”. in Planctomycetes: cell structure, origins and biology (ed Fuerst, J. A.) (Humana Press, 2013).
Bay, S. K. et al. Trace gas oxidizers are widespread and active members of soil microbial communities. Nat. Microbiol. 6, 246–256 (2021).
pubmed: 33398096
doi: 10.1038/s41564-020-00811-w
Dedysh, S. N., Knief, C. & Dunfield, P. F. Methylocella species are facultatively methanotrophic. J. Bacteriol. 187, 4665–4670 (2005).
pubmed: 15968078
pmcid: 1151763
doi: 10.1128/JB.187.13.4665-4670.2005
Awala, S. I. et al. Methylacidiphilum caldifontis gen. nov., sp. nov., a thermoacidophilic methane-oxidizing bacterium from an acidic geothermal environment, and descriptions of the family Methylacidiphilaceae fam. nov. and order Methylacidiphilales ord. nov. Int. J. Syst. Evol. Microbiol. 73, 006085 (2023).
doi: 10.1099/ijsem.0.006085
Thauer, R. K., Jungermann, K. & Decker, K. Energy conservation in chemotrophic anaerobic bacteria. Bacteriol. Rev. 41, 100–180 (1977).
pubmed: 860983
pmcid: 413997
doi: 10.1128/br.41.1.100-180.1977
Svensson-Ek, M. et al. The X-ray crystal structures of wild-type and EQ (I-286) mutant cytochrome c oxidases from Rhodobacter sphaeroides. J. Mol. Biol. 321, 329–339 (2002).
pubmed: 12144789
doi: 10.1016/S0022-2836(02)00619-8
Iwata, S., Ostermeier, C., Ludwig, B. & Michel, H. Structure at 2.8 Å resolution of cytochrome c oxidase from Paracoccus denitrificans. Nature 376, 660–669 (1995).
pubmed: 7651515
doi: 10.1038/376660a0
Chen, J. & Strous, M. Denitrification and aerobic respiration, hybrid electron transport chains and co-evolution. Biochim. Biophys. Acta Bioenerg. 1827, 136–144 (2013).
doi: 10.1016/j.bbabio.2012.10.002
Bergaust, L., Mao, Y., Bakken, L. R. & Frostegård, A. Denitrification response patterns during the transition to anoxic respiration and posttranscriptional effects of suboptimal pH on nitrous oxide reductase in Paracoccus denitrificans. Appl. Environ. Microbiol. 76, 6387–6396 (2010).
pubmed: 20709842
pmcid: 2950438
doi: 10.1128/AEM.00608-10
Van den Heuvel, R. N., Bakker, S. E., Jetten, M. S. & Hefting, M. M. Decreased N
pubmed: 21504539
doi: 10.1111/j.1472-4669.2011.00276.x
Palmer, K., Drake, H. L. & Horn, M. A. Association of novel and highly diverse acid-tolerant denitrifiers with N
pubmed: 20023077
doi: 10.1128/AEM.02256-09
Brenzinger, K., Dörsch, P. & Braker, G. pH-driven shifts in overall and transcriptionally active denitrifiers control gaseous product stoichiometry in growth experiments with extracted bacteria from soil. Front. Microbiol. 6, 961 (2015).
pubmed: 26441895
pmcid: 4585170
doi: 10.3389/fmicb.2015.00961
Lycus, P. et al. Phenotypic and genotypic richness of denitrifiers revealed by a novel isolation strategy. ISME J. 11, 2219–2232 (2017).
pubmed: 28696424
pmcid: 5607364
doi: 10.1038/ismej.2017.82
Almeida, J. S., Júlio, S. M., Reis, M. A. & Carrondo, M. J. Nitrite inhibition of denitrification by Pseudomonas fluorescens. Biotechnol. Bioeng. 46, 194–201 (1995).
pubmed: 18623304
doi: 10.1002/bit.260460303
Fang, F. C. Antimicrobial reactive oxygen and nitrogen species: concepts and controversies. Nat. Rev. Microbiol. 2, 820–832 (2004).
pubmed: 15378046
doi: 10.1038/nrmicro1004
Bueno, E., Sit, B., Waldor, M. K. & Cava, F. Anaerobic nitrate reduction divergently governs population expansion of the enteropathogen Vibrio cholerae. Nat. Microbiol. 3, 1346–1353 (2018).
pubmed: 30275512
pmcid: 6443258
doi: 10.1038/s41564-018-0253-0
Vadivelu, V. M., Keller, J. & Yuan, Z. Free ammonia and free nitrous acid inhibition on the anabolic and catabolic processes of Nitrosomonas and Nitrobacter. Water Sci Technol. 56, 89–97 (2007).
pubmed: 17951872
doi: 10.2166/wst.2007.612
Zhu-Barker, X., Cavazos, A. R., Ostrom, N. E., Horwath, W. R. & Glass, J. B. The importance of abiotic reactions for nitrous oxide production. Biogeochemistry 126, 251–267 (2015).
doi: 10.1007/s10533-015-0166-4
Cole, J. Anaerobic bacterial response to nitric oxide stress: Widespread misconceptions and physiologically relevant responses. Mol. Microbiol. 116, 29–40 (2021).
pubmed: 33706420
doi: 10.1111/mmi.14713
Auclair, J., Lépine, F., Parent, S. & Villemur, R. Dissimilatory reduction of nitrate in seawater by a Methylophaga strain containing two highly divergent narG sequences. ISME J. 4, 1302–1313 (2010).
pubmed: 20393572
doi: 10.1038/ismej.2010.47
Kamps, J. J., Hopkinson, R. J., Schofield, C. J. & Claridge, T. D. How formaldehyde reacts with amino acids. Commun. Chem. 2, 126 (2019).
doi: 10.1038/s42004-019-0224-2
Dedysh, S. N. & Dunfield, P. F. Facultative and obligate methanotrophs:how to identify and differentiate them.Methods Enzymol. 495, 31–44 (2011).
pubmed: 21419913
doi: 10.1016/B978-0-12-386905-0.00003-6
Purchase, M. L., Bending, G. D. & Mushinski, R. M. Spatiotemporal variations of soil reactive nitrogen oxide fluxes across the anthropogenic landscape. Environ. Sci. Technol. 57, 16348–16360 (2023).
pubmed: 37856795
pmcid: 10620987
doi: 10.1021/acs.est.3c05849
Pauleta, S. R., Dell’Acqua, S. & Moura, I. Nitrous oxide reductase. Coord. Chem. Rev. 257, 332–349 (2013).
doi: 10.1016/j.ccr.2012.05.026
Wang, Z., Vishwanathan, N., Kowaliczko, S. & Ishii, S. Clarifying microbial nitrous oxide reduction under aerobic conditions: tolerant, intolerant, and sensitive. Microbiol. Spectr. 11, e0470922 (2023).
pubmed: 36926990
doi: 10.1128/spectrum.04709-22
Suenaga, T., Riya, S., Hosomi, M. & Terada, A. Biokinetic characterization and activities of N
pubmed: 29692767
pmcid: 5902568
doi: 10.3389/fmicb.2018.00697
Hilgeri, H. & Humer, M. Biotic landfill cover treatments for mitigating methane emissions. Environ. Monit. Assess. 84, 71–84 (2003).
pubmed: 12733810
doi: 10.1023/A:1022878830252
Ross, M. O. & Rosenzweig, A. C. A tale of two methane monooxygenases. J. Biol. Inorg. Chem. 22, 307–319 (2017).
pubmed: 27878395
doi: 10.1007/s00775-016-1419-y
Hein, S., Witt, S. & Simon, J. Clade II nitrous oxide respiration of Wolinella succinogenes depends on the NosG, -C1, -C2, -H electron transport module, NosB and a Rieske/cytochrome bc complex. Environ. Microbiol. 19, 4913–4925 (2017).
pubmed: 28925551
doi: 10.1111/1462-2920.13935
Keltjens, J. T., Pol, A., Reimann, J. & Op den Camp, H. J. M. PQQ-dependent methanol dehydrogenases: rare-earth elements make a difference. Appl. Microbiol. Biotechnol. 98, 6163–6183 (2014).
pubmed: 24816778
doi: 10.1007/s00253-014-5766-8
Sirota, F. L., Maurer-Stroh, S., Li, Z., Eisenhaber, F. & Eisenhaber, B. Functional classification of super-large families of enzymes based on substrate binding pocket residues for biocatalysis and enzyme engineering applications. Front. Bioeng. Biotechnol. 9, 701120 (2021).
pubmed: 34409021
pmcid: 8366029
doi: 10.3389/fbioe.2021.701120
Le, T.-K., Lee, Y.-J., Han, G. H. & Yeom, S.-J. Methanol dehydrogenases as a key biocatalysts for synthetic methylotrophy. Front. Bioeng. Biotechnol. 9, 787791 (2021).
pubmed: 35004648
pmcid: 8741260
doi: 10.3389/fbioe.2021.787791
Bonnot, F., Iavarone, A. T. & Klinman, J. P. Multistep, eight-electron oxidation catalyzed by the cofactorless oxidase, PqqC: identification of chemical intermediates and their dependence on molecular oxygen. Biochemistry 52, 4667–4675 (2013).
pubmed: 23718207
doi: 10.1021/bi4003315
Matsushita, K. et al. Escherichia coli is unable to produce pyrroloquinoline quinone (PQQ). Microbiology 143, 3149–3156 (1997).
pubmed: 9353919
doi: 10.1099/00221287-143-10-3149
Zhang, W. et al. Guidance for engineering of synthetic methylotrophy based on methanol metabolism in methylotrophy. RSC Adv. 7, 4083–4091 (2017).
doi: 10.1039/C6RA27038G
Chu, F. & Lidstrom, M. E. XoxF Acts as the predominant methanol dehydrogenase in the type I methanotroph Methylomicrobium buryatense. J .Bacteriol. 198, 1317–1325 (2016).
pubmed: 26858104
pmcid: 4859581
doi: 10.1128/JB.00959-15
Kim, H. J. & Graham, D. W. Effect of oxygen level on simultaneous nitrogenase and sMMO expression and activity in Methylosinus trichosporium OB3b and its sMMO
pubmed: 11470351
doi: 10.1111/j.1574-6968.2001.tb10746.x
Kim, H. J. & Graham, D. W. Effects of oxygen and nitrogen conditions on the transformation kinetics of 1,2-dichloroethenes by Methylosinus trichosporium OB3b and its sMMO
pubmed: 14669871
doi: 10.1023/A:1027396619596
Smirnova, A. V. & Dunfield, P. F. Differential transcriptional activation of genes encoding soluble methane monooxygenase in a facultative versus an obligate methanotroph. Microorganisms 6, 20 (2018).
pubmed: 29509697
pmcid: 5874634
doi: 10.3390/microorganisms6010020
Theisen, A. R. et al. Regulation of methane oxidation in the facultative methanotroph Methylocella silvestris BL2. Mol. Microbiol. 58, 682–692 (2005).
pubmed: 16238619
doi: 10.1111/j.1365-2958.2005.04861.x
Reimer, L. C., Sarda Carbasse, J., Koblitz, J., Podstawka, A. & Overmann, J. Methylocella tundrae Dedysh et al. 2004. DSMZ. https://doi.org/10.13145/bacdive1659.20230509.8.1 (2023).
Reimer, L. C., Sarda Carbasse, J., Koblitz J., Podstawka, A., Overmann, J. Methylocystis echinoides (ex Gal’chenko et al. 1977) DSMZ. https://doi.org/10.13145/bacdive169085.20230509.8.1 (2023).
Qian, H. et al. Greenhouse gas emissions and mitigation in rice agriculture. Nat. Rev. Earth Environ. 4, 716–732 (2023).
doi: 10.1038/s43017-023-00482-1
Kolb, S. & Horn, M. A. Microbial CH
pubmed: 22403579
pmcid: 3291872
doi: 10.3389/fmicb.2012.00078
Ishii, S., Ohno, H., Tsuboi, M., Otsuka, S. & Senoo, K. Identification and isolation of active N
pubmed: 21677691
pmcid: 3223309
doi: 10.1038/ismej.2011.69
Taminskas, J. et al. Climate change and water table fluctuation: Implications for raised bog surface variability. Geomorphology 304, 40–49 (2018).
doi: 10.1016/j.geomorph.2017.12.026
Ratcliffe, J. L., Campbell, D. I., Clarkson, B. R., Wall, A. M. & Schipper, L. A. Water table fluctuations control CO
pubmed: 30577098
doi: 10.1016/j.scitotenv.2018.11.151
Evans, C. D. et al. Overriding water table control on managed peatland greenhouse gas emissions. Nature 593, 548–552 (2021).
pubmed: 33882562
doi: 10.1038/s41586-021-03523-1
Kotsyurbenko O. R., Glagolev M. V., Merkel A. Y., Sabrekov A. F., Terentieva I. E. Methanogenesis in Soils, Wetlands, and Peat. in Biogenesis of Hydrocarbons (eds Stams A. J. M., Sousa D. Z.) (Springer International Publishing, 2019).
Angle, J. C. et al. Methanogenesis in oxygenated soils is a substantial fraction of wetland methane emissions. Nat. Commun. 8, 1567 (2017).
pubmed: 29146959
pmcid: 5691036
doi: 10.1038/s41467-017-01753-4
Zhu, X., Burger, M., Doane, T. A. & Horwath, W. R. Ammonia oxidation pathways and nitrifier denitrification are significant sources of N
pubmed: 23576736
pmcid: 3631630
doi: 10.1073/pnas.1219993110
Hakobyan, A., Zhu, J., Glatter, T., Paczia, N. & Liesack, W. Hydrogen utilization by Methylocystis sp. strain SC2 expands the known metabolic versatility of type IIa methanotrophs. Metab. Eng. 61, 181–196 (2020).
pubmed: 32479801
doi: 10.1016/j.ymben.2020.05.003
Widdel, F., Bak, F. Gram-negative mesophilic sulfate-reducing bacteria. In The Prokaryotes: A Handbook on the Biology of Bacteria: Ecophysiology, Isolation, Identification, Applications (eds Balows, A., Trüper, H. G., Dworkin, M., Harder, W., Schleifer, K.-H.) (Springer International Publishing, 1992).
Bellosillo, L. A. Effects of environmental factors on methanotroph communities from a forest soil, lake sediment and a landfill soil. (Chungbuk National University, 2020).
Awala, S. I. et al. Methylococcus geothermalis sp. nov., a methanotroph isolated from a geothermal field in the Republic of Korea. Int. J. Syst. Evol. Microbiol. 70, 5520–5530 (2020).
pubmed: 32910751
doi: 10.1099/ijsem.0.004442
Weisburg, W. G., Barns, S. M., Pelletier, D. A. & Lane, D. J. 16S ribosomal DNA amplification for phylogenetic study. J. Bacteriol. 173, 697–703 (1991).
pubmed: 1987160
pmcid: 207061
doi: 10.1128/jb.173.2.697-703.1991
Hurt, R. A. et al. Simultaneous recovery of RNA and DNA from soils and sediments. Appl. Environ. Microbiol. 67, 4495–4503 (2001).
pubmed: 11571148
pmcid: 93195
doi: 10.1128/AEM.67.10.4495-4503.2001
Wick, R. R. et al. Trycycler: consensus long-read assemblies for bacterial genomes. Genome Biol. 22, 266 (2021).
pubmed: 34521459
pmcid: 8442456
doi: 10.1186/s13059-021-02483-z
Li, H. Minimap and miniasm: fast mapping and de novo assembly for noisy long sequences. Bioinformatics 32, 2103–2110 (2016).
pubmed: 27153593
pmcid: 4937194
doi: 10.1093/bioinformatics/btw152
Kolmogorov, M., Yuan, J., Lin, Y. & Pevzner, P. A. Assembly of long, error-prone reads using repeat graphs. Nat. Biotechnol. 37, 540–546 (2019).
pubmed: 30936562
doi: 10.1038/s41587-019-0072-8
Vaser, R. & Šikić, M. Time- and memory-efficient genome assembly with Raven. Nat. Comput. Sci. 1, 332–336 (2021).
pubmed: 38217213
doi: 10.1038/s43588-021-00073-4
Wick, R. R. & Holt, K. E. Polypolish: short-read polishing of long-read bacterial genome assemblies. PLoS Comput. Biol. 18, e1009802 (2022).
pubmed: 35073327
pmcid: 8812927
doi: 10.1371/journal.pcbi.1009802
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).
pubmed: 32589667
pmcid: 7347232
doi: 10.1371/journal.pcbi.1007981
Seemann, T. Prokka: rapid prokaryotic genome annotation. Bioinformatics 30, 2068–2069 (2014).
pubmed: 24642063
doi: 10.1093/bioinformatics/btu153
Tatusova, T. et al. NCBI prokaryotic genome annotation pipeline. Nucleic Acids Res. 44, 6614–6624 (2016).
pubmed: 27342282
pmcid: 5001611
doi: 10.1093/nar/gkw569
Hunter, S. et al. InterPro: the integrative protein signature database. Nucleic Acids Res. 37, D211–D215 (2009).
pubmed: 18940856
doi: 10.1093/nar/gkn785
Ashburner, M. et al. Gene ontology: tool for the unification of biology. Nat. Genet. 25, 25–29 (2000).
pubmed: 10802651
pmcid: 3037419
doi: 10.1038/75556
Consortium, T. G. O. The gene ontology resource: enriching a GOld mine. Nucleic Acids Res. 49, D325–d334 (2021).
doi: 10.1093/nar/gkaa1113
Finn, R. D. et al. Pfam: the protein families database. Nucleic Acids Res. 42, D222–D230 (2014).
pubmed: 24288371
doi: 10.1093/nar/gkt1223
Lu, S. et al. CDD/SPARCLE: the conserved domain database in 2020. Nucleic Acids Res. 48, D265–d268 (2020).
pubmed: 31777944
doi: 10.1093/nar/gkz991
Haft, D. H., Selengut, J. D. & White, O. The TIGRFAMs database of protein families. Nucleic Acids Res. 31, 371–373 (2003).
pubmed: 12520025
pmcid: 165575
doi: 10.1093/nar/gkg128
Huerta-Cepas, J. et al. eggNOG 5.0: a hierarchical, functionally and phylogenetically annotated orthology resource based on 5090 organisms and 2502 viruses. Nucleic Acids Res. 47, D309–D314 (2019).
pubmed: 30418610
doi: 10.1093/nar/gky1085
Almagro Armenteros, J. J. et al. SignalP 5.0 improves signal peptide predictions using deep neural networks. Nat. Biotechnol. 37, 420–423 (2019).
pubmed: 30778233
doi: 10.1038/s41587-019-0036-z
Krogh, A., Larsson, B., von Heijne, G. & Sonnhammer, E. L. Predicting transmembrane protein topology with a hidden Markov model: application to complete genomes. J. Mol. Biol. 305, 567–580 (2001).
pubmed: 11152613
doi: 10.1006/jmbi.2000.4315
Tu, Q., Lin, L., Cheng, L., Deng, Y. & He, Z. NCycDB: a curated integrative database for fast and accurate metagenomic profiling of nitrogen cycling genes. Bioinformatics 35, 1040–1048 (2018).
doi: 10.1093/bioinformatics/bty741
Olson, R. D. et al. Introducing the bacterial and viral Bioinformatics Resource Center (BV-BRC): a resource combining PATRIC, IRD and ViPR. Nucleic Acids Res. 51, D678–D689 (2022).
pmcid: 9825582
doi: 10.1093/nar/gkac1003
Katoh, K. & Standley, D. M. MAFFT multiple sequence alignment software version 7: improvements in performance and usability. Mol. Biol. Evol. 30, 772–780 (2013).
pubmed: 23329690
pmcid: 3603318
doi: 10.1093/molbev/mst010
Letunic, I. & Bork, P. Interactive Tree Of Life (iTOL) v5: an online tool for phylogenetic tree display and annotation. Nucleic Acids Res. 49, W293–W296 (2021).
pubmed: 33885785
pmcid: 8265157
doi: 10.1093/nar/gkab301
Bertelli, C. et al. IslandViewer 4: expanded prediction of genomic islands for larger-scale datasets. Nucleic Acids Res. 45, W30–w35 (2017).
pubmed: 28472413
pmcid: 5570257
doi: 10.1093/nar/gkx343
Onley, J. R., Ahsan, S., Sanford, R. A. & Löffler, F. E. Denitrification by Anaeromyxobacter dehalogenans, a common soil bacterium lacking the nitrite reductase genes nirS and nirK. Appl. Environ. Microbiol. 84, e01985–01917 (2018).
pubmed: 29196287
pmcid: 5795083
doi: 10.1128/AEM.01985-17
Miranda, K. M., Espey, M. G. & Wink, D. A. A rapid, simple spectrophotometric method for simultaneous detection of nitrate and nitrite. Nitric Oxide 5, 62–71 (2001).
pubmed: 11178938
doi: 10.1006/niox.2000.0319
Muyzer, G., De Waal, E. C. & Uitterlinden, A. G. Profiling of complex microbial populations by denaturing gradient gel electrophoresis analysis of polymerase chain reaction-amplified genes coding for 16S rRNA. Appl. Environ. Microbiol. 59, 695–700 (1993).
pubmed: 7683183
pmcid: 202176
doi: 10.1128/aem.59.3.695-700.1993
Andrews, S. FastQC: a quality control tool for high throughput sequence data http://www.bioinformatics.babraham.ac.uk/projects/fastqc (2010).
Bolger, A. M., Lohse, M. & Usadel, B. Trimmomatic: a flexible trimmer for Illumina sequence data. Bioinformatics 30, 2114–2120 (2014).
pubmed: 24695404
pmcid: 4103590
doi: 10.1093/bioinformatics/btu170
Kopylova, E., Noé, L. & Touzet, H. SortMeRNA: fast and accurate filtering of ribosomal RNAs in metatranscriptomic data. Bioinformatics 28, 3211–3217 (2012).
pubmed: 23071270
doi: 10.1093/bioinformatics/bts611
Langmead, B. & Salzberg, S. L. Fast gapped-read alignment with Bowtie 2. Nat. Methods 9, 357–359 (2012).
pubmed: 22388286
pmcid: 3322381
doi: 10.1038/nmeth.1923
Anders, S., Pyl, P. T. & Huber, W. HTSeq—a Python framework to work with high-throughput sequencing data. Bioinformatics 31, 166–169 (2014).
pubmed: 25260700
pmcid: 4287950
doi: 10.1093/bioinformatics/btu638
Hein, S. & Simon, J. Bacterial nitrous oxide respiration: electron transport chains and copper transfer reactions. Adv .Microb. Physiol. 75, 137–175 (2019).
pubmed: 31655736
doi: 10.1016/bs.ampbs.2019.07.001
Torres, M. J. et al. Nitrous oxide metabolism in nitrate-reducing bacteria: physiology and regulatory mechanisms. Adv. Microb. Physiol. 68, 353–432 (2016).
pubmed: 27134026
doi: 10.1016/bs.ampbs.2016.02.007