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    “Candidatus Dechloromonas phosphoritropha” and “Ca. D. phosphorivorans”, novel polyphosphate accumulating organisms abundant in wastewater treatment systems

    1.Nielsen PH, Mcilroy SJ, Albertsen M, Nierychlo M. Re-evaluating the microbiology of the enhanced biological phosphorus removal process. Curr Opin Biotechnol. 2019;57:111–8.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    2.Marques R, Santos J, Nguyen H, Carvalho G, Noronha JP, Nielsen PH, et al. Metabolism and ecological niche of Tetrasphaera and Ca. Accumulibacter in enhanced biological phosphorus removal. Water Res. 2017;122:159–71.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    3.Camejo PY, Oyserman BO, Mcmahon KD, Noguera DR. Integrated omic analyses provide evidence that a “Candidatus Accumulibacter phosphatis” strain performs denitrification under microaerobic conditions. mSystems. 2019;4:1–23.Article 

    Google Scholar 
    4.Oyserman BO, Noguera DR, Del Rio TG, Tringe SG, McMahon KD. Metatranscriptomic insights on gene expression and regulatory controls in Candidatus Accumulibacter phosphatis. ISME J. 2016;10:810–22.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    5.Tu Y, Schuler AJ. Low acetate concentrations favor polyphosphate-accumulating organisms over glycogen-accumulating organisms in enhanced biological phosphorus removal from wastewater. Environ Sci Technol. 2013;47:3816–24.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    6.Marques R, Ribera-guardia A, Santos J, Carvalho G, Reis MAM, Pijuan M, et al. Denitrifying capabilities of Tetrasphaera and their contribution towards nitrous oxide production in enhanced biological phosphorus removal processes. Water Res. 2018;137:262–72.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    7.Fernando EY, McIlroy SJ, Nierychlo M, Herbst F-A, Petriglieri F, Schmid MC, et al. Resolving the individual contribution of key microbial populations to enhanced biological phosphorus removal with Raman–FISH. ISME J. 2019;13:1933–46.8.Kawaharasaki M, Tanaka H, Kanagawa T, Nakamura K. In situ identification of polyphosphate-accumulating bacteria in activated sludge by dual staining with rRNA-targeted oligonucleotide probes and 4’,6-diaimidino-2-phenylindol (DAPI) at a polyphosphate-probing concentration. Water Res. 1999;33:257–65.CAS 
    Article 

    Google Scholar 
    9.Crocetti GR, Hugenholtz P, Bond PL, Schuler AJ, Keller J, Jenkins D, et al. Identification of polyphosphate-accumulating organisms and design of 16SrRNA-directed probes for their detection and quantitation. Appl Environ Microbiol. 2000;66:1175–82.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    10.Kong Y, Nielsen JL, Nielsen PH. Identity and ecophysiology of uncultured Actinobacterial polyphosphate-accumulating organisms in full-scale enhanced biological phosphorus removal plants. Appl Environ Microbiol. 2005;71:4076–85.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    11.Kong Y, Xia Y, Nielsen JL, Nielsen PH. Structure and function of the microbial community in a full-scale enhanced biological phosphorus removal plant. Microbiology. 2007;153:4061–73.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    12.Göel R, Sanhueza P, Noguera D. Evidence of Dechloromonas sp. participating in enhanced biological phosphorous removal (EBPR) in a bench-scale aerated-anoxic reactor. Proc Water Environ Fed. 2005;41:3864–71.Article 

    Google Scholar 
    13.Terashima M, Yama A, Sato M, Yumoto I, Kamagata Y, Kato S. Culture-dependent and -independent identification of polyphosphate-accumulating Dechloromonas spp. predominating in a full-scale oxidation ditch wastewater treatment plant. Microbes Environ Environ. 2016;31:449–55.Article 

    Google Scholar 
    14.Wang B, Jiao E, Guo Y, Zhang L, Meng Q, Zeng W, et al. Investigation of the polyphosphate-accumulating organism population in the full-scale simultaneous chemical phosphorus removal system. Environ Sci Pollut Res. 2020;27:37877–86.CAS 
    Article 

    Google Scholar 
    15.Stokholm-Bjerregaard M, McIlroy SJ, Nierychlo M, Karst SM, Albertsen M, Nielsen PH. A critical assessment of the microorganisms proposed to be important to enhanced biological phosphorus removal in full-scale wastewater treatment systems. Front Microbiol. 2017;8:718.PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    16.Achenbach LA, Michaelidou U, Bruce RA, Fryman J, Coates JD. Dechloromonas agitata gen. nov., sp. nov. and Dechlorosoma suillum gen. nov., sp. nov., two novel environmentally dominant (per)chlorate-reducing bacteria and their phylogenetic position. Int J Syst Evol Microbiol. 2001;51:527–33.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    17.Horn MA, Ihssen J, Matthies C, Schramm A, Acker G, Drake HL, et al. Dechloromonas denitrificans sp. nov., Flavobacterium denitrificans sp. nov., Paenibacillus anaericanus sp. nov. and Paenibacillus terrae strain MH72, N2O-producing bacteria isolated from the gut of the earthworm Aporrectodea caliginosa. Int J Syst Evol Microbiol. 2005;55:1255–65.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    18.Günther S, Trutnau M, Kleinsteuber S, Hause G, Bley T, Röske I, et al. Dynamics of polyphosphate-accumulating bacteria in wastewater treatment plant microbial communities detected via DAPI (4’,6’-Diamidino-2-Phenylindole) and Tetracycline Labeling. Appl Environ Microbiol. 2009;75:2111–21.PubMed 
    PubMed Central 
    Article 
    CAS 

    Google Scholar 
    19.Acevedo B, Murgui M, Borrás L, Barat R. New insights in the metabolic behaviour of PAO under negligible poly-P reserves. Chem Eng J. 2017;311:82–90.CAS 
    Article 

    Google Scholar 
    20.Yuan Y, Liu J, Ma B, Liu Y, Wang B, Peng Y. Improving municipal wastewater nitrogen and phosphorous removal by feeding sludge fermentation products to sequencing batch reactor (SBR). Bioresour Technol. 2016;222:326–34.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    21.Lv X, Shao M, Li C, Li J, Gao X, Sun F. A comparative study of the bacterial community in denitrifying and traditional enhanced biological phosphorus removal processes. Microbes Environ. 2014;29:261–8.PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    22.Salinero KK, Keller K, Feil WS, Feil H, Trong S, Bartolo Di G, et al. Metabolic analysis of the soil microbe Dechloromonas aromatica str. RCB anaerobic pathways for aromatic degradation. BMC Genom. 2009;23:1–23.
    Google Scholar 
    23.McIlroy SJ, Starnawska A, Starnawski P, Saunders AM, Nierychlo M, Nielsen PH, et al. Identification of active denitrifiers in full-scale nutrient removal wastewater treatment systems. Environ Microbiol. 2016;18:50–64.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    24.Hesselsoe M, Fu S, Schloter M, Bodrossy L, Iversen N, Roslev P, et al. Isotope array analysis of Rhodocyclales uncovers functional redundancy and versatility in an activated sludge. ISME J. 2009;3:1349–64.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    25.Ahn J, Schroeder S, Beer M, McIlroy S, Bayly RC, May JW, et al. Ecology of the microbial community removing phosphate from wastewater under continuously aerobic conditions in a sequencing batch reactor. Appl Environ Microbiol. 2007;73:2257–70.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    26.Dueholm MS, Andersen KS, McIlroy SJ, Kristensen JM, Yashiro E, Karst SM, et al. Generation of comprehensive ecosystems-specific reference databases with species-level resolution by high-throughput full-length 16S rRNA gene sequencing and automated taxonomy assignment (AutoTax). mBio. 2020;11:e01557–20.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    27.Singleton CM, Petriglieri F, Kristensen JM, Kirkegaard RH, Michaelsen TY, Andersen MH, et al. Connecting structure to function with the recovery of over 1000 high-quality activated sludge metagenome-assembled genomes encoding full-length rRNA genes using long-read sequencing. Nat Commun. 2021;12:2009.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    28.Albertsen M, Hugenholtz P, Skarshewski A, Nielsen KL, Tyson GW, Nielsen PH. Genome sequences of rare, uncultured bacteria obtained by differential coverage binning of multiple metagenomes. Nat Biotechnol. 2013;31:533–8.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    29.Parks DH, Rinke C, Chuvochina M, Chaumeil P, Woodcroft BJ, Evans PN, et al. Recovery of nearly 8,000 metagenome-assembled genomes substantially expands the tree of life. Nat Microbiol. 2017;2:1533–42.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    30.GJF Smolders, Meij Van Der J, Loosdrecht Van MCM, Heijnen JJ. Model of the anaerobic metabolism of the biological phosphorus removal process: stoichiometry and pH influence. Biotechnol Bioeng. 1994;43:461–70.Article 

    Google Scholar 
    31.Jørgensen MK, Nierychlo M, Nielsen AH, Larsen P, Christensen ML, Nielsen PH. Unified understanding of physico-chemical properties of activated sludge and fouling propensity. Water Res. 2017;120:117–32.PubMed 
    Article 
    CAS 
    PubMed Central 

    Google Scholar 
    32.Nielsen JL. Protocol for fluorescence in situ hybridization (FISH) with rRNA-targeted oligonucleotides. FISH handbook for biological wastewater treatment. 2009. pp 73–84.33.McIlroy SJ, Kirkegaard RH, McIlroy B, Nierychlo M, Kristensen JM, Karst SM, et al. MiDAS 2.0: an ecosystem-specific taxonomy and online database for the organisms of wastewater treatment systems expanded for anaerobic digester groups. Database. 2017;2017:1–9.Article 

    Google Scholar 
    34.Nierychlo M, Andersen KS, Xu Y, Green N, Jiang C, Albertsen M, et al. MiDAS 3: an ecosystem-specific reference database, taxonomy and knowledge platform for activated sludge and anaerobic digesters reveals species-level microbiome composition of activated sludge. Water Res. 2020;182:115955.35.R Core Team. R: A language and environment for statistical computing. 2020. R Foundation for Statistical Computing, Vienna, Austria.36.RStudio Team. RStudio: Integrated Development Environment for R. 2015. Boston, MA.37.Albertsen M, Karst SM, Ziegler AS, Kirkegaard RH, Nielsen PH. Back to basics—the influence of DNA extraction and primer choice on phylogenetic analysis of activated sludge communities. PLoS One. 2015;10:1–15.Article 
    CAS 

    Google Scholar 
    38.Wickham H. ggplot2—elegant graphics for data analysis. Springer. 2009. Springer Science & Business Media.39.Ludwig W, Strunk O, Westram R, Richter L, Meier H, Yadhukumar A, et al. ARB: a software environment for sequence data. Nucleic Acids Res. 2004;32:1363–71.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    40.Quast C, Pruesse E, Yilmaz P, Gerken J, Schweer T, Yarza P, et al. The SILVA ribosomal RNA gene database project: improved data processing and web-based tools. Nucleic Acids Res. 2013;41:590–6.Article 
    CAS 

    Google Scholar 
    41.Sayers EW, Agarwala R, Bolton EE, Brister JR, Canese K, Clark K, et al. Database resources of the National Center for Biotechnology Information. Nucleic Acids Res. 2019;47:D23–D28.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    42.Yilmaz LS, Parnerkar S, Noguera DR. MathFISH, a web tool that uses thermodynamics-based mathematical models for in silico evaluation of oligonucleotide probes for fluorescence in situ hybridization. Appl Environ Microbiol. 2011;77:1118–22.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    43.Daims H, Stoecker K, Wagner M. Fluorescence in situ hybridization for the detection of prokaryotes. In: Osborn AM, Smith CJ (eds). Molecular Microbial Ecology. 2005. Taylor & Francis, New York, pp 213–39.44.Daims H, Lücker S, Wagner M. Daime, a novel image analysis program for microbial ecology and biofilm research. Environ Microbiol. 2006;8:200–13.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    45.Chaumeil P, Mussig AJ, Parks DH, Hugenholtz P. Genome analysis GTDB-Tk: a toolkit to classify genomes with the Genome Taxonomy Database. Bioinformatics. 2019;36:1925–7.PubMed Central 

    Google Scholar 
    46.Olm MR, Brown CT, Brooks B, Banfield JF. dRep: a tool for fast and accurate genomic comparisons that enables improved genome recovery from metagenomes through de-replication. ISME J. 2017;11:2864–8.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    47.Pritchard L, Glover RH, Humphris S, Elphinstone JG, Toth IK. Genomics and taxonomy in diagnostics for food security: soft-rotting enterobacterial plant pathogens. Anal Methods. 2016;8:12–24.Article 

    Google Scholar 
    48.Kanehisa M, Sato Y, Kawashima M, Furumichi M, Tanabe M. KEGG as a reference resource for gene and protein annotation. Nucleic Acids Res. 2016;44:457–62.Article 
    CAS 

    Google Scholar 
    49.Seemann T. Prokka: rapid prokaryotic genome annotation. Bioinformatics. 2014;30:2068–9.CAS 
    Article 

    Google Scholar 
    50.Nawrocki EP, Kolbe DL, Eddy SR. Infernal 1.0: inference of RNA alignments. Bioinformatics. 2009; 25:1335–7.51.Nguyen L, Schmidt HA, Haeseler Von A, Minh BQ. IQ-TREE: a fast and effective stochastic algorithm for estimating maximum-likelihood phylogenies. 2014; 32:268–74.52.Wolterink A, Kim S, Muusse M, Kim IS, Roholl PJM, Ginkel Van GC. et al. Dechloromonas hortensis sp nov strain ASK-1, two Nov (per)chlorate-reducing Bact, taxonomic description strain GR-1. Int J Syst Evolut Microbiol. 2005;1:2063–8.Article 
    CAS 

    Google Scholar 
    53.Zilles JL, Peccia J, Noguera DR. Microbiology of enhanced biological phosphorus removal in aerated-anoxic orbal processes. Water Environ Res. 2002;74:428–36.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    54.Dueholm MS, Nierychlo M, Andersen KS, Rudkjøbing V, Knudsen S, the MiDAS Global Consortium, et al. MiDAS 4—a global WWTP ecosystem-specific full-length 16S rRNA gene catalogue and taxonomy for studies of bacterial communities. bioRxiv 2021.55.Oehmen A, Lemos PC, Carvalho G, Yuan Z, Blackall LL, Reis MAM. Advances in enhanced biological phosphorus removal: from micro to macro scale. Water Res. 2007;41:2271–2300.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    56.Qiu G, Zuniga-montanez R, Law Y, Swa S. Polyphosphate-accumulating organisms in full-scale tropical wastewater treatment plants use diverse carbon sources. Water Res. 2019;149:469–510.Article 
    CAS 

    Google Scholar 
    57.Petriglieri F, Petersen JF, Peces M, Nierychlo M, Hansen K, Baastrand CE, et al. Quantification of biologically and chemically bound phosphorus in activated sludge from full-scale plants with biological P-removal. biorxiv 2020. https://doi.org/10.1101/2021.01.04.425262.58.Hesselmann RPX, Von Rummel R, Resnick SM, Hany R, Zehnder AJB. Anaerobic metabolism of bacteria performing enhanced biological phosphate removal. Water Res. 2000;34:3487–94.CAS 
    Article 

    Google Scholar 
    59.Acevedo B, Oehmen A, Carvalho G, Seco A, Borrás L, Barat R. Metabolic shift of polyphosphate-accumulating organisms with different levels of polyphosphate storage. Water Res. 2012;6:1889–1900.Article 
    CAS 

    Google Scholar 
    60.Flowers JJ, He S, Malfatti S, Glavina T, Tringe SG, Hugenholtz P, et al. Comparative genomics of two ‘Candidatus Accumulibacter’ clades performing biological phosphorus removal. ISME J. 2013;7:2301–14.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    61.Qiu G, Liu X, Saw NMMT, Law Y, Zuniga-Montanez R, Thi SS, et al. Metabolic traits of Candidatus Accumulibacter clade IIF Strain SCELSE-1 using amino acids as carbon sources for enhanced biological phosphorus removal. Environ Sci Technol. 2019;54:2448–58.PubMed 
    Article 
    CAS 
    PubMed Central 

    Google Scholar 
    62.Kristiansen R, Thi H, Nguyen T, Saunders AM, Nielsen JL, Wimmer R, et al. A metabolic model for members of the genus Tetrasphaera involved in enhanced biological phosphorus removal. ISME J. 2013;7:543–54.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    63.Parks DH, Chuvochina M, Chaumeil P-A, Rinke C, Mussig AJ, Hugenholtz P. A complete domain-to-species taxonomy for Bacteria and Archaea. Nat Biotechnol. 2020;38:1079–86.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    64.McIlroy SJ, Albertsen M, Andresen EK, Saunders AM. ‘Candidatus Competibacter’-lineage genomes retrieved from metagenomes reveal functional metabolic diversity. ISME J. 2014;8:613–24.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    65.Saunders AM, Mabbett AN, Mcewan AG, Blackall LL. Proton motive force generation from stored polymers for the uptake of acetate under anaerobic conditions. FEMS Microbiol Lett. 2007;274:245–51.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    66.Erdal UG, Erdal ZK, Daigger GT, Randall CW. Is it PAO-GAO competition or metabolic shift in EBPR system? Evidence from an experimental study. Water Sci Technol. 2008;58:1329–34.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    67.Zhou Y, Pijuan M, Zeng RJ, Lu H, Could ÃZY. polyphosphate-accumulating organisms (PAOs) be glycogen-accumulating organisms (GAOs)? Water Res. 2008;42:2361–8.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    68.Weissbrodt DG, Lopez-vazquez CM, Welles L. “Candidatus Accumulibacter delftensis”: a clade IC novel polyphosphate-accumulating organism without denitrifying activity on nitrate. Water Res. 2019;161:136–51.PubMed 
    Article 
    CAS 
    PubMed Central 

    Google Scholar 
    69.Camejo PY, Owen BR, Martirano J, Ma J, Kapoor V, Santo J, et al. Candidatus Accumulibacter phosphatis clades enriched under cyclic anaerobic and microaerobic conditions simultaneously use different electron acceptors. Water Res. 2016;102:125–37.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    70.Skennerton CT, Barr JJ, Slater FR, Bond PL, Tyson GW. Expanding our view of genomic diversity in Candidatus Accumulibacter clades. Environ Microbiol. 2015;17:1574–85.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    71.Hendriks J, Oubrie A, Castresana J, Urbani A, Gemeinhardt S, Saraste M. Nitric oxide reductases in bacteria. Biochim Biophys Acta—Bioenerg. 2000;1459:266–73.CAS 
    Article 

    Google Scholar 
    72.Murray RGE, Stackebrandt E. Taxonomic note: implementation of the provisional status Candidatus for incompletely described procaryotes. Int J Syst Evol Microbiol. 1995;45:186–7.CAS 

    Google Scholar  More

  • in

    Emergent “core communities” of microbes, meiofauna and macrofauna at hydrothermal vents

    1.Wisz MS, Pottier J, Kissling WD, Pellissier L, Lenoir J, Damgaard CF, et al. The role of biotic interactions in shaping distributions and realised assemblages of species: implications for species distribution modelling. Biol Rev Camb Philos Soc. 2013;88:15–30.PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    2.Thompson LR, Sanders JG, McDonald D, Amir A, Ladau J, Locey KJ, et al. A communal catalogue reveals Earth’s multiscale microbial diversity. Nature. 2017;551:457–63.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    3.Cho BC, Azam F. Major role of bacteria in biogeochemical fluxes in the oceans interior. Nature. 1988;332:441–3.CAS 
    Article 

    Google Scholar 
    4.Rousk J, Bengtson P. Microbial regulation of global biogeochemical cycles. Front Microbiol. 2014;5:103.PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    5.Guilhon M, Montserrat F, Turra A. Recognition of ecosystem-based management principles in key documents of the seabed mining regime: implications and further recommendations. ICES J Marine Sci. 2020:fsaa229.6.Sherman K, Sissenwine M, Christensen V, Duda A, Hempel G, Ibe C, et al. A global movement toward an ecosystem approach to management of marine resources. Mar Ecol Prog Ser. 2005;300:275–9.Article 

    Google Scholar 
    7.Passarelli C, Olivier F, Paterson DM, Hubas C. Impacts of biogenic structures on benthic assemblages: microbes, meiofauna, macrofauna and related ecosystem functions. Mar Ecol Prog Ser. 2012;465:85–97.Article 

    Google Scholar 
    8.Baldrighi E, Aliani S, Conversi A, Lavaleye M, Borghini M, Manini E. From microbes to macrofauna: an integrated study of deep benthic communities and their response to environmental variables along the Malta Escarpment (Ionian Sea). Sci Mar. 2013;77:625–39.Article 

    Google Scholar 
    9.Foshtomi MY, Braeckman U, Derycke S, Sapp M, Van Gansbeke D, Sabbe K, et al. The link between microbial diversity and nitrogen cycling in marine sediments is modulated by macrofaunal bioturbation. PLoS ONE. 2015;10:e0130116.10.Hope JA, Paterson DM, Thrush SF. The role of microphytobenthos in soft-sediment ecological networks and their contribution to the delivery of multiple ecosystem services. J Ecology. 2020;108:815–30.Article 

    Google Scholar 
    11.Lima-Mendez G, Faust K, Henry N, Decelle J, Colin S, Carcillo F, et al. Ocean plankton. Determinants of community structure in the global plankton interactome. Science. 2015;348:1262073.PubMed 
    Article 
    CAS 
    PubMed Central 

    Google Scholar 
    12.Blanchet FG, Cazelles K, Gravel D. Co-occurrence is not evidence of ecological interactions. Ecol Lett. 2020;23:1050–63.PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    13.Pearson K. Mathematical contributions to the theory of evolution—on a form of spurious correlation which may arise when indices are used in the measurement of organs. Proc R Soc Lond. 1897;60:489–98.Article 

    Google Scholar 
    14.Jackson DA. Compositional data in community ecology: the paradigm or peril of proportions? Ecology. 1997;78:929–40.Article 

    Google Scholar 
    15.Gloor GB, Reid G. Compositional analysis: a valid approach to analyze microbiome high-throughput sequencing data. Can J Microbiol. 2016;62:692–703.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    16.Lovell D, Pawlowsky-Glahn V, Egozcue JJ, Marguerat S, Bahler J. Proportionality: a valid alternative to correlation for relative data. PLoS Comput Biol. 2015;11:e1004075.PubMed 
    PubMed Central 
    Article 
    CAS 

    Google Scholar 
    17.Sievert SM, Vetriani C. Chemoautotrophy at deep-sea vents: past, present, and future. Oceanography. 2012;25:218–33.Article 

    Google Scholar 
    18.Huber JA, Butterfield DA, Baross JA. Temporal changes in archaeal diversity and chemistry in a mid-ocean ridge subseafloor habitat. Appl Environ Microbiol. 2002;68:1585–94.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    19.Karl DM, Wirsen CO, Jannasch HW. Deep-sea primary production at the Galapagos hydrothermal vents. Science. 1980;207:1345–7.CAS 
    Article 

    Google Scholar 
    20.Meyer JL, Akerman NH, Proskurowski G, Huber JA Microbiological characterization of post-eruption “snowblower” vents at Axial Seamount Juan de Fuca Ridge. Front Microbiol. 2013;4:153.21.Orcutt BN, Sylvan JB, Knab NJ, Edwards KJ. Microbial ecology of the dark ocean above, at, and below the seafloor. Microbiol Mol Biol Rev. 2011;75:361–422.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    22.Dubilier N, Bergin C, Lott C. Symbiotic diversity in marine animals: the art of harnessing chemosynthesis. Nat Rev Microbiol. 2008;6:725–40.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    23.Yamanaka T et al. A Compilation of the Stable Isotopic Compositions of Carbon, Nitrogen, and Sulfur in Soft Body Parts of Animals Collected from Deep-Sea Hydrothermal Vent and Methane Seep Fields: Variations in Energy Source and Importance of Subsurface Microbial Processes in the Sediment-Hosted Systems. In: Ishibashi J, Okino K, Sunamura M, editors. Subseafloor Biosphere Linked to Hydrothermal Systems. Tokyo, Japan: Springer Open; 2015. p. 105–29.24.Bergquist D, Eckner J, Urcuyo I, Cordes E, Hourdez S, Macko S, Fisher C. Using stable isotopes and quantitative community characteristics to determine a local hydrothermal vent food web. Mar Ecol Prog Ser. 2007;330:49–65.Article 

    Google Scholar 
    25.Colaço A, Dehairs F, Desbruyères D. Nutritional relations of deep-sea hydrothermal fields at the Mid-Atlantic Ridge: a stable isotope approach. Deep-Sea Res Part I-Oceanogr Res Pap. 2002;49:395–412.Article 

    Google Scholar 
    26.Van Dover C, Fry B. Stable isotopic compositions of hydrothermal vent organisms. Mar Biol. 1989;102:257–63.Article 

    Google Scholar 
    27.Colaço A, Desbruyères D, Guezennec J. Polar lipid fatty acids as indicators of trophic associations in a deep-sea vent system community. Marine Ecology-an Evolut Perspect. 2007;28:15–24.Article 
    CAS 

    Google Scholar 
    28.Limen H, Stevens CJ, Bourass Z, Juniper SK. Trophic ecology of siphonostomatoid copepods at deep-sea hydrothermal vents in the northeast Pacific. Mar Ecol Prog Ser. 2008;359:161–70.Article 

    Google Scholar 
    29.Van Dover CL. Trophic relationships among invertebrates at the Kairei hydrothermal vent field (Central Indian Ridge). Mar Biol. 2002;141:761–72.Article 

    Google Scholar 
    30.Lamy T, Koenigs C, Holbrook SJ, Miller RJ, Stier AC, Reed DC. Foundation species promote community stability by increasing diversity in a giant kelp forest. Ecology. 2020;101:e02987.PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    31.Bruno JF, Bertness MD Habitat modification and facilitation in benthic marine communities. In: Bertness MD, Gaines SD, Hay ME, editors. Marine Community Ecology. Sunderland, MA: Sinauer Associates; 2001. p. 201–18.32.Dayton PK Toward an Understanding of Community Resilience and the Potential Effects of Enrichments to the Benthos at McMurdo Sound, Antarctica. Pages 81-95. In: Parker BC, editor. Proceedings of the Colloquium on Conservation Problems. Lawrence, Kansas, USA.: Allen Press; 1972.33.Tunnicliffe V, Cordes EE The tubeworm forests of hydrothermal vents and cold seeps. In: Rossi S, Bramanti L, editors. Perspectives on the Marine Animal Forests of the World Springer; 2020. p. 147–92.34.López-García P, Gaill F, Moreira D. Wide bacterial diversity associated with tubes of the vent worm Riftia pachyptila. Environ Microbiol. 2002;4:204–15.PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    35.Rincon-Tomas B, Francisco Javier González, Luis Somoza, Kathrin Sauter, Pedro Madureira, Teresa Medialdea et al. Siboglinidae Tubes as an Additional Niche for Microbial Communities in the Gulf of Cadiz-A Microscopical Appraisal. Microorganisms. 2020;8:367.36.Page A, Juniper SK, Olagnon M, Alain K, Desrosiers G, Querellou J, et al. Microbial diversity associated with a Paralvinella sulfincola tube and the adjacent substratum on an active deep-sea vent chimney. Geobiology. 2004;2:225–38.Article 

    Google Scholar 
    37.Govenar B Shaping Vent and Seep Communities: Habitat Provision and Modification by Foundation Species. In: Kiel S, editor. The vent and seep biota: aspects from microbes to ecosystems. Dordrecht: Springer; 2010. p. 403–32.38.Tunnicliffe V, Germain CS, Hilario A Phenotypic Variation and Fitness in a Metapopulation of Tubeworms (Ridgeia piscesae Jones) at Hydrothermal Vents. PLoS ONE. 2014;9:e110578.39.Sarrazin J, Juniper SK. Biological characteristics of a hydrothermal edifice mosaic community. Mar Ecol Prog Ser. 1999;185:1–19.Article 

    Google Scholar 
    40.Sarrazin J, Juniper SK, Massoth G, Legendre P. Physical and chemical factors influencing species distributions on hydrothermal sulfide edifices of the Juan de Fuca Ridge, northeast Pacific. Mar Ecol Prog Ser. 1999;190:89–112.CAS 
    Article 

    Google Scholar 
    41.Govenar BW, Bergquist DC, Urcuyo IA, Eckner JT, Fisher CR. Three Ridgeia piscesae assemblages from a single Juan de Fuca Ridge sulphide edifice: structurally different and functionally similar. Cah Biol Mar. 2002;43:247–52.
    Google Scholar 
    42.Forget NL, Juniper SK. Free-living bacterial communities associated with tubeworm (Ridgeia piscesae) aggregations in contrasting diffuse flow hydrothermal vent habitats at the Main Endeavour Field, Juan de Fuca Ridge. MicrobiologyOpen. 2013;2:259–75.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    43.Danovaro R, Gambi C, Dell’Anno A, Corinaldesi C, Fraschetti S, Vanreusel A, et al. Exponential decline of deep-sea ecosystem functioning linked to benthic biodiversity loss. Curr Biol. 2008;18:1–8.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    44.Dick GJ. The microbiomes of deep-sea hydrothermal vents: distributed globally, shaped locally. Nature Rev Microbiol. 2019;17:271–83.CAS 

    Google Scholar 
    45.Lee W-K, Juniper SK, Perez M, Ju S-J, Kim S-J Diversity and characterization of bacterial communities of five co-occurring species at a hydrothermal vent on the Tonga Arc. Ecol Evol. 2021;11:4481–93.46.Sogin ML, Morrison HG, Huber JA, Mark Welch D, Huse SM, Neal PR, et al. Microbial diversity in the deep sea and the underexplored “rare biosphere”. Proc Natl Acad Sci USA. 2006;103:12115–20.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    47.Eren AM, Vineis JH, Morrison HG, Sogin ML. A filtering method to generate high quality short reads using illumina paired-end technology. PLoS One. 2013;8:e66643.PubMed 
    PubMed Central 
    Article 
    CAS 

    Google Scholar 
    48.Schloss PD, Westcott SL, Ryabin T, Hall JR, Hartmann M, Hollister EB, et al. Introducing mothur: open-source, platform-independent, community-supported software for describing and comparing microbial communities. Appl Environ Microbiol. 2009;75:7537–41.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    49.Caron DA, Countway PD, Savai P, Gast RJ, Schnetzer A, Moorthi SD, et al. Defining DNA-based operational taxonomic units for microbial-eukaryote ecology. Appl Environ Microbiol. 2009;75:5797–808.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    50.Quast C, Pruesse E, Yilmaz P, Gerken J, Schweer T, Yarza P, et al. The SILVA ribosomal RNA gene database project: improved data processing and web-based tools. Nucleic Acids Res. 2013;41(Database issue):D590–6.CAS 
    PubMed 
    PubMed Central 

    Google Scholar 
    51.Guillou L, Bachar D, Audic S, Bass D, Berney C, Bittner L, et al. The Protist Ribosomal Reference database (PR2): a catalog of unicellular eukaryote Small Sub-Unit rRNA sequences with curated taxonomy. Nucleic Acids Res. 2013;41:D597–604.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    52.Quinn TP, Ionas Erb, Greg Gloor, Cedric Notredame, Mark F Richardson, Tamsyn M Crowley et al. A field guide for the compositional analysis of any-omics data. Gigascience. 2019;8:giz107.53.Martín-Fernández JA, Palarea-Albaladejo J, Olea RA Dealing with Zeros. In: Pawlowsky‐Glahn V, Buccianti A, editors. Compositional Data Analysis2011. p. 43-58.54.Palarea-Albaladejo J, Martin-Fernandez JA. zCompositions – R Package for multivariate imputation of left-censored data under a compositional approach. Chemometr Intell Lab. 2015;143:85–96.CAS 
    Article 

    Google Scholar 
    55.Aitchison J The statistical analysis of compositional data. London: Chapman & Hall; 1986. p. 416.56.Aitchison J, Barcelo-Vidal C, Martin-Fernandez JA, Pawlowsky-Glahn V. Logratio analysis and compositional distance. Math Geol. 2000;32:271–5.Article 

    Google Scholar 
    57.Comas-Cufí M coda.base: A Basic Set of Functions for Compositional Data Analysis. R package version 0.2.1 2019 [Available from: https://CRAN.R-project.org/package=coda.base.58.Oksanen J et al. vegan: Community Ecology Package. R package version 2.2-1. 2015 [Available from: http://CRAN.R-project.org/package=vegan.59.Fernandes AD, Macklaim JM, Linn TG, Reid G, Gloor GB. ANOVA-like differential expression (ALDEx) analysis for mixed population RNA-Seq. PLoS One. 2013;8:e67019.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    60.Quinn TP, Richardson MF, Lovell D, Crowley TM. propr: an R-package for identifying proportionally abundant features using compositional data analysis. Sci Rep. 2017;7:16252.PubMed 
    PubMed Central 
    Article 
    CAS 

    Google Scholar 
    61.Shannon P, Markiel A, Ozier O, Baliga NS, Wang JT, Ramage D, et al. Cytoscape: a software environment for integrated models of biomolecular interaction networks. Genome Research. 2003;13:2498–504.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    62.Huber JA, Butterfield DA, Baross JA. Bacterial diversity in a subseafloor habitat following a deep-sea volcanic eruption. FEMS Microbiol Ecol. 2003;43:393–409.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    63.Akerman NH, Butterfield DA, Huber JA Phylogenetic diversity and functional gene patterns of sulfur-oxidizing subseafloor Epsilonproteobacteria in diffuse hydrothermal vent fluids. Front Microbiol. 2013;4:185.64.Tsurumi M, Tunnicliffe V. Tubeworm-associated communities at hydrothermal vents on the Juan de Fuca Ridge, northeast Pacific. Deep-Sea Res Part I-Oceanogr Res Pap. 2003;50:611–29.Article 

    Google Scholar 
    65.Butterfield DA, Massoth GJ, McDuff RE, Lupton JE, Lilley MD. Geochemistry of hydrothermal fluids from Axial Seamount Hydrothermal Emissions Study vent field, Juan de Fuca Ridge: subseafloor boiling and subsequent fluid-rock interaction. J Geophys Res. 1990;95:12895–921.Article 

    Google Scholar 
    66.Johnson KS, Beehler CL, Sakamotoarnold CM, Childress JJ. insitu measurements of chemical-distributions in a deep-sea hydrothermal vent field. Science. 1986;231:1139–41.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    67.Du Preez C, Fisher CP Long-Term Stability of back-Arc basin hydrothermal vents. Front Mar Sci. 2018;5:54.68.Urcuyo IA, Bergquist DC, MacDonald IR, VanHorn M, Fisher CR. Growth and longevity of the tubeworm Ridgeia piscesae in the variable diffuse flow habitats of the Juan de Fuca Ridge. Mar Ecol Prog Ser. 2007;344:143–57.Article 

    Google Scholar 
    69.Perner M, Bach W, Hentscher M, Koschinsky A, Garbe-Schönberg D, Streit WR, et al. Short-term microbial and physico-chemical variability in low-temperature hydrothermal fluids near 5 degrees S on the Mid-Atlantic Ridge. Environ Microbiol. 2009;11:2526–41.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    70.Orcutt BN, Bradley JA, Brazelton WJ, Estes ER, Goordial JM, Huber JA, et al. Impacts of deep-sea mining on microbial ecosystem services. Limnology Oceanogr. 2020;65:1489–510.CAS 
    Article 

    Google Scholar 
    71.Gollner S, Ivanenko VN, Arbizu PM, Bright M. Advances in taxonomy, ecology, and biogeography of Dirivultidae (copepoda) associated with chemosynthetic environments in the deep sea. PLoS One. 2010;5:e9801.PubMed 
    PubMed Central 
    Article 
    CAS 

    Google Scholar 
    72.Kalanetra KM, Nelson DC. Vacuolate-attached filaments: highly productive Ridgeia piscesae epibionts at the Juan de Fuca hydrothermal vents. Mar Biol. 2010;157:791–800.PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    73.Girguis PR, Lee RW. Thermal preference and tolerance of alvinellids. Science. 2006;312:231.PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    74.Burgaud G, Le Calvez T, Arzur D, Vandenkoornhuyse P, Barbier G. Diversity of culturable marine filamentous fungi from deep-sea hydrothermal vents. Environ Microbiol. 2009;11:1588–600.PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    75.Murdock SA, Juniper SK. Hydrothermal vent protistan distribution along the Mariana arc suggests vent endemics may be rare and novel. Environ Microbiol. 2019;21:3796–815.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    76.Meier DV, Bach W, Girguis PR, Gruber-Vodicka HR, Reeves EP, Richter M, et al. Heterotrophic Proteobacteria in the vicinity of diffuse hydrothermal venting. Environ Microbiol. 2016;18:4348–68.PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    77.Stokke R, Dahle H, Roalkvam I, Wissuwa J, Daae FL, Tooming-Klunderud A, et al. Functional interactions among filamentous Epsilonproteobacteria and Bacteroidetes in a deep-sea hydrothermal vent biofilm. Environ Microbiol. 2015;17:4063–77.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    78.Adl SM, Bass D, Lane CE, Lukeš J, Schoch CL, Smirnov A, et al. Revisions to the classification, nomenclature, and diversity of eukaryotes. J Eukaryot Microbiol. 2019;66:4–119.PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    79.Brown MW, Sharpe SC, Silberman JD, Heiss AA, Lang BF, Simpson AG, et al. Phylogenomics demonstrates that breviate flagellates are related to opisthokonts and apusomonads. Proc Biol Sci. 2013;280:20131755.PubMed 
    PubMed Central 

    Google Scholar 
    80.Hamann E, Gruber-Vodicka H, Kleiner M, Tegetmeyer HE, Riedel D, Littmann S, et al. Environmental Breviatea harbour mutualistic Arcobacter epibionts. Nature. 2016;534:254–8.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    81.Gollner S, Riemer B, Arbizu PM, Le Bris N, Bright M. Diversity of meiofauna from the 9 degrees 50’ N East Pacific rise across a gradient of hydrothermal fluid emissions. PLoS ONE. 2010;5:e12321.82.Sarrazin J, Legendre P, de Busserolles F, Fabri MC, Guilini K, Ivanenko VN, et al. Biodiversity patterns, environmental drivers and indicator species on a high-temperature hydrothermal edifice, Mid-Atlantic Ridge. Deep-Sea Res Part Ii-Topical Stud Oceanogr. 2015;121:177–92.CAS 
    Article 

    Google Scholar 
    83.Bates AE, Harmer TL, Roeselers G, Cavanaugh CM. Phylogenetic characterization of episymbiotic bacteria hosted by a hydrothermal vent limpet (lepetodrilidae, vetigastropoda). Biol Bull-US. 2011;220:118–27.Article 

    Google Scholar 
    84.Schratzberger M, Ingels J. Meiofauna matters: the roles of meiofauna in benthic ecosystems. J Exp Mar Biol Ecol. 2018;502:12–25.Article 

    Google Scholar 
    85.Cronin-O’Reilly S, Joe D Taylor, Ian Jermyn, A Louise Allcock, Michael Cunliffe, Mark P Johnson et al. Limited congruence exhibited across microbial, meiofaunal and macrofaunal benthic assemblages in a heterogeneous coastal environment. Sci Rep-UK. 2018;8:15500.86.Reimann F, Schrage M. The mucus-trap hypothesis on feeding of aquatic nematodes and implications for biodegradation and sediment texture. Oecologia. 1978;34:75–88.Article 

    Google Scholar 
    87.Léveillé RJ, Levesque C, Juniper SK Biotic interactions and feedback processes in deep-sea hydrothermal vent ecosystems. In: Kristensen E, Haese RR, Kostka JE, editors. Interactions between macro- and microorganisms in marine sediments. Washington, DC: American Geophysical Union; 2005. p. 299–321.88.Ingels J, Ann Vanreusel, Ellen Pape, Francesca Pasotti, Lara Macheriotou, Pedro Martínez Arbizu et al. Ecological variables for deep-ocean monitoring must include microbiota and meiofauna for effective conservation. Nat Ecology Evolut. 2020: https://doi.org/10.1038/s41559-020-01335-6.89.Thompson KF, Miller KA, Currie D, Johnston P, Santillo D. Seabed mining and approaches to governance of the deep seabed. Front Mar Sci. 2018;5:480. More

  • in

    Escaping the choosiness trap

    1.Courtiol, A., Etienne, L., Feron, R., Godelle, B. & Rousset, F. Am. Nat. 188, 521–538 (2016).Article 

    Google Scholar 
    2.Jennions, M. D. & Petrie, M. Biol. Rev. 75, 21–64 (2000).CAS 
    Article 

    Google Scholar 
    3.Kokko, H. & Mappes, J. Evolution 59, 1876–1885 (2005).Article 

    Google Scholar 
    4.Hare, R. M. & Simmons, L. W. Biol. Rev. 94, 929–956 (2019).Article 

    Google Scholar 
    5.Kohlmeier, P., Zhang, Y., Gorter, J. A., Su, C.-Y. & Billeter, J.-C. Nat. Ecol. Evol. https://doi.org/10.1038/s41559-021-01482-4 (2021).Article 

    Google Scholar 
    6.Halliday, T. R. in Mate Choice (ed. Bateson, P.) 3–32 (Cambridge Univ. Press, 1983).7.Avila, F. W., Sirot, L. K., LaFlamme, B. A., Rubinstein, C. D. & Wolfner, M. F. Annu. Rev. Entomol. 56, 21–40 (2011).CAS 
    Article 

    Google Scholar 
    8.Perry, J. C. & Rowe, L. Cold Spring Harb. Perspect. Biol. 7, a017558 (2015).Article 

    Google Scholar 
    9.Hopkins, B. R., Avila, F. W. & Wolfner, M. F. in Encyclopedia of Reproduction (ed. Skinner, M. K.) 137–144 (Elsevier, 2018).10.de Boer, R. A., Vega-Trejo, R., Kotrschal, A. & Fitzpatrick, J. L. Nat. Ecol. Evol. https://doi.org/ggbb (2021). More

  • in

    Iron and sulfate reduction structure microbial communities in (sub-)Antarctic sediments

    1.D’Hondt S, Jørgensen BB, Miller DJ, Batzke A, Blake R, Cragg BA, et al. Distributions of microbial activities in deep subseafloor sediments. Science. 2004;306:2216–21.PubMed 
    Article 
    CAS 
    PubMed Central 

    Google Scholar 
    2.Froelich PN, Klinkhammer GP, Bender ML, Luedtke NA, Heath GR, Cullen D, et al. Early oxidation of organic matter in pelagic sediments of the eastern equatorial Atlantic: suboxic diagenesis. Geochim Cosmochim Acta. 1979;43:1075–90.CAS 
    Article 

    Google Scholar 
    3.Parkes RJ, Cragg B, Roussel E, Webster G, Weightman A, Sass H. A review of prokaryotic populations and processes in sub-seafloor sediments, including biosphere: geosphere interactions. Mar Geol. 2014;352:409–25.CAS 
    Article 

    Google Scholar 
    4.Arnosti C. Microbial extracellular enzymes and the marine carbon cycle. Annu Rev Mar Sci. 2011;3:401–25.Article 

    Google Scholar 
    5.Thamdrup B, Rosselló-Mora R, Amann R. Microbial manganese and sulfate reduction in Black Sea shelf sediments. Appl Environ Microbiol. 2000;66:2888–97.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    6.Thamdrup B. Bacterial manganese and iron reduction in aquatic sediments. In: Schink B, editor. Advances in microbial ecology. Boston, MA, US: Springer; 2000. p. 41–84.7.Jørgensen BB, Kasten S. Sulfur cycling and methane oxidation. In: Schulz HD, Zabel M, editors. Marine geochemistry. 2nd ed. Berlin, Heidelberg, Germany: Springer-Verlag; 2006. p. 271–309.8.Bowles MW, Mogollón JM, Kasten S, Zabel M, Hinrichs K-U. Global rates of marine sulfate reduction and implications for sub–sea-floor metabolic activities. Science. 2014;344:889–91.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    9.Jickells TD, An ZS, Andersen KK, Baker AR, Bergametti G, Brooks N, et al. Global iron connections between desert dust, ocean biogeochemistry, and climate. Science. 2005;308:67–71.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    10.Raiswell R, Hawkings JR, Benning LG, Baker AR, Death R, Albani S, et al. Potentially bioavailable iron delivery by iceberg-hosted sediments and atmospheric dust to the polar oceans. Biogeosciences. 2016;13:3887–900.CAS 
    Article 

    Google Scholar 
    11.Hawkings JR, Wadham JL, Tranter M, Raiswell R, Benning LG, Statham PJ, et al. Ice sheets as a significant source of highly reactive nanoparticulate iron to the oceans. Nat Commun. 2014;5:3929.CAS 
    PubMed 
    Article 

    Google Scholar 
    12.Death R, Wadham JL, Monteiro F, Le Brocq AM, Tranter M, Ridgwell A, et al. Antarctic ice sheet fertilises the Southern Ocean. Biogeosciences. 2014;11:2635–43.Article 

    Google Scholar 
    13.Monien D, Monien P, Brünjes R, Widmer T, Kappenberg A, Silva Busso AA, et al. Meltwater as a source of potentially bioavailable iron to Antarctica waters. Antarct Sci. 2017;29:277–91.Article 

    Google Scholar 
    14.Henkel S, Kasten S, Hartmann JF, Silva-Busso A, Staubwasser M. Iron cycling and stable Fe isotope fractionation in Antarctic shelf sediments, King George Island. Geochim Cosmochim Acta. 2018;237:320–38.CAS 
    Article 

    Google Scholar 
    15.Hodson A, Nowak A, Sabacka M, Jungblut A, Navarro F, Pearce D, et al. Climatically sensitive transfer of iron to maritime Antarctic ecosystems by surface runoff. Nat Commun. 2017;8:14499.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    16.Wang S, Bailey D, Lindsay K, Moore JK, Holland M. Impact of sea ice on the marine iron cycle and phytoplankton productivity. Biogeosciences. 2014;11:4713–31.CAS 
    Article 

    Google Scholar 
    17.Jørgensen BB, Findlay AJ, Pellerin A. The biogeochemical sulfur cycle of marine sediments. Front Microbiol. 2019;10:849.PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    18.Findlay AJ, Kamyshny A. Turnover rates of intermediate sulfur species (Sx2−, S0, S2O32−, S4O62−, SO32−) in anoxic freshwater and sediments. Front Microbiol. 2017;8:2551.19.Findlay AJ, Pellerin A, Laufer K, Jørgensen BB. Quantification of sulphide oxidation rates in marine sediment. Geochim Cosmochim Acta. 2020;280:441–52.CAS 
    Article 

    Google Scholar 
    20.Canfield DE, Jørgensen BB, Fossing H, Glud R, Gundersen J, Ramsing NB, et al. Pathways of organic carbon oxidation in three continental margin sediments. Mar Geol. 1993;113:27–40.CAS 
    PubMed 
    Article 

    Google Scholar 
    21.Michaud AB, Laufer K, Findlay A, Pellerin A, Antler G, Turchyn AV, et al. Glacial influence on the iron and sulfur cycles in Arctic fjord sediments (Svalbard). Geochim Cosmochim Acta. 2020;280:423–40.CAS 
    Article 

    Google Scholar 
    22.Jensen MM, Thamdrup B, Rysgaard S, Holmer M, Fossing H. Rates and regulation of microbial iron reduction in sediments of the Baltic-North Sea transition. Biogeochemistry. 2003;65:295–317.Article 

    Google Scholar 
    23.Beckler JS, Kiriazis N, Rabouille C, Stewart FJ, Taillefert M. Importance of microbial iron reduction in deep sediments of river-dominated continental-margins. Mar Chem. 2016;178:22–34.CAS 
    Article 

    Google Scholar 
    24.Riedinger N, Brunner B, Krastel S, Arnold GL, Wehrmann LM, Formolo MJ, et al. Sulfur cycling in an iron oxide-dominated, dynamic marine depositional system: the Argentine Continental Margin. Front Earth Sci. 2017;5:33.Article 

    Google Scholar 
    25.Thamdrup B, Fossing H, Jørgensen BB. Manganese, iron and sulfur cycling in a coastal marine sediment, Aarhus Bay, Denmark. Geochim Cosmochim Acta. 1994;58:5115–29.CAS 
    Article 

    Google Scholar 
    26.Arndt S, Jørgensen BB, LaRowe DE, Middelburg J, Pancost R, Regnier P. Quantifying the degradation of organic matter in marine sediments: a review and synthesis. Earth-Sci Rev. 2013;123:53–86.CAS 
    Article 

    Google Scholar 
    27.Algora C, Vasileiadis S, Wasmund K, Trevisan M, Krüger M, Puglisi E, et al. Manganese and iron as structuring parameters of microbial communities in Arctic marine sediments from the Baffin Bay. FEMS Microbiol Ecol. 2015;91:fiv056.PubMed 
    Article 
    CAS 

    Google Scholar 
    28.Franco M, De Mesel I, Diallo MD, Van Der Gucht K, Van Gansbeke D, Van, et al. Effect of phytoplankton bloom deposition on benthic bacterial communities in two contrasting sediments in the southern North Sea. Aquat Micro Ecol. 2007;48:241–54.Article 

    Google Scholar 
    29.Zonneveld KAF, Versteegh GJM, Kasten S, Eglinton TI, Emeis K-C, Huguet C, et al. Selective preservation of organic matter in marine environments; processes and impact on the sedimentary record. Biogeosciences. 2010;7:483–511.CAS 
    Article 

    Google Scholar 
    30.Jorgensen SL, Hannisdal B, Lanzén A, Baumberger T, Flesland K, Fonseca R, et al. Correlating microbial community profiles with geochemical data in highly stratified sediments from the Arctic Mid-Ocean Ridge. Proc Natl Acad Sci U S A. 2012;109:E2846–55.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    31.Zinke LA, Glombitza C, Bird JT, Røy H, Jørgensen BB, Lloyd KG, et al. Microbial organic matter degradation potential in Baltic Sea sediments is influenced by depositional conditions and in situ geochemistry. Appl Environ Microbiol. 2019;85:e02164-18.PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    32.Yang J, Jiang H, Wu G, Dong H. Salinity shapes microbial diversity and community structure in surface sediments of the Qinghai-Tibetan Lakes. Sci Rep. 2016;6:25078.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    33.Hicks N, Liu X, Gregory R, Kenny J, Lucaci A, Lenzi L, et al. Temperature driven changes in benthic bacterial diversity influences biogeochemical cycling in coastal sediments. Front Microbiol. 2018;9:1730.PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    34.Hamdan LJ, Coffin RB, Sikaroodi M, Greinert J, Treude T, Gillevet PM. Ocean currents shape the microbiome of Arctic marine sediments. ISME J. 2013;7:685–96.CAS 
    PubMed 
    Article 

    Google Scholar 
    35.Schulz HD, Zabel M, editors. Marine geochemistry. 2nd ed. Berlin, Heidelberg, Germany: Springer-Verlag; 2006.36.Geprägs P, Torres ME, Mau S, Kasten S, Römer M, Bohrmann G. Carbon cycling fed by methane seepage at the shallow Cumberland Bay, South Georgia, sub-Antarctic. Geochem, Geophys Geosystems. 2016;17:1401–18.Article 
    CAS 

    Google Scholar 
    37.Atkinson A, Whitehouse MJ, Priddle J, Cripps GC, Ward P, Brandon MA. South Georgia, Antarctica: a productive, cold water, pelagic ecosystem. Mar Ecol Prog Ser. 2001;216:279–308.CAS 
    Article 

    Google Scholar 
    38.Löffler B. Geochemische Prozesse und Stoffkreisläufe in Sedimenten innerhalb und außerhalb des Cumberland-Bay Fjordes, Süd Georgien. Bachelor Thesis. Bremen, Germany: University of Bremen; 2013.39.Köster M. (Bio-)geochemische Prozesse in den eisenreichen Seep-Sedimenten der Cumberland-Bucht Südgeorgiens, Subantarktis. Bachelor Thesis. Bremen, Germany: University of Bremen; 2014.40.Römer M, Torres M, Kasten S, Kuhn G, Graham AG, Mau S, et al. First evidence of widespread active methane seepage in the Southern Ocean, off the sub-Antarctic island of South Georgia. Earth Planet Sci Lett. 2014;403:166–77.Article 
    CAS 

    Google Scholar 
    41.Bohrmann G, Aromokeye AD, Bihler V, Dehning K, Dohrmann I, Gentz T, et al. R/V METEOR Cruise Report M134, emissions of free gas from cross-shelf troughs of South Georgia: distribution, quantification, and sources for methane ebullition sites in sub-Antarctic waters, Port Stanley (Falkland Islands)—Punta Arenas (Chile), 16 January–18 February 2017. 2017.42.Schnakenberg A, Aromokeye DA, Kulkarni A, Maier L, Wunder LC, Richter-Heitmann T, et al. Electron acceptor availability shapes Anaerobically Methane Oxidizing Archaea (ANME) communities in South Georgia sediments. Front Microbiol. 2021;12:726.Article 

    Google Scholar 
    43.Rückamp M, Braun M, Suckro S, Blindow N. Observed glacial changes on the King George Island ice cap, Antarctica, in the last decade. Global Planet Change. 2011;79:99–109.44.Seeberg-Elverfeldt J, Schlüter M, Feseker T, Kölling M. Rhizon sampling of porewaters near the sediment-water interface of aquatic systems. Limnol Oceanogr Methods. 2005;3:361–71.Article 

    Google Scholar 
    45.Oni OE, Miyatake T, Kasten S, Richter-Heitmann T, Fischer D, Wagenknecht L, et al. Distinct microbial populations are tightly linked to the profile of dissolved iron in the methanic sediments of the Helgoland mud area, North Sea. Front Microbiol. 2015;6:365.PubMed 
    PubMed Central 

    Google Scholar 
    46.Aromokeye DA, Richter-Heitmann T, Oni OE, Kulkarni A, Yin X, Kasten S, et al. Temperature controls crystalline iron oxide utilization by microbial communities in methanic ferruginous marine sediment incubations. Front Microbiol. 2018;9:2574.PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    47.Parada AE, Needham DM, Fuhrman JA. Every base matters: assessing small subunit rRNA primers for marine microbiomes with mock communities, time series and global field samples. Environ Microbiol. 2016;18:1403–14.CAS 
    PubMed 
    Article 

    Google Scholar 
    48.Herlemann DPR, Labrenz M, Jürgens K, Bertilsson S, Waniek JJ, Andersson AF. Transitions in bacterial communities along the 2000 km salinity gradient of the Baltic Sea. ISME J. 2011;5:1571–9.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    49.Ovreås L, Forney L, Daae FL, Torsvik V. Distribution of bacterioplankton in meromictic Lake Saelenvannet, as determined by denaturing gradient gel electrophoresis of PCR-amplified gene fragments coding for 16S rRNA. Appl Environ Microbiol. 1997;63:3367–73.PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    50.Takai K, Horikoshi K. Rapid detection and quantification of members of the archaeal community by quantitative PCR using fluorogenic probes. Appl Environ Microbiol. 2000;66:5066–72.51.Viollier E, Inglett P, Hunter K, Roychoudhury A, Van Cappellen P. The ferrozine method revisited: Fe(II)/Fe(III) determination in natural waters. Appl Geochem. 2000;15:785–90.CAS 
    Article 

    Google Scholar 
    52.Yin X, Kulkarni AC, Friedrich MW. DNA and RNA stable isotope probing of methylotrophic methanogenic Archaea. In: Dumont MG, Hernández García M, editors. Stable isotope probing: methods and protocols. New York, NY: Springer; 2019. p. 189–206.53.Aromokeye DA, Kulkarni AC, Elvert M, Wegener G, Henkel S, Coffinet S, et al. Rates and microbial players of iron-driven anaerobic oxidation of methane in methanic marine sediments. Front Microbiol. 2020;10:3041.PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    54.Eden PA, Schmidt TM, Blakemore RP, Pace NR. Phylogenetic analysis of Aquaspirillum magnetotacticum using polymerase chain reaction-amplified 16S rRNA-specific DNA. Int J Syst Evol Microbiol. 1991;41:324–5.CAS 

    Google Scholar 
    55.Yu Y, Lee C, Kim J, Hwang S. Group-specific primer and probe sets to detect methanogenic communities using quantitative real-time polymerase chain reaction. Biotechnol Bioeng. 2005;89:670–9.CAS 
    PubMed 
    Article 

    Google Scholar 
    56.Lueders T, Friedrich MW. Effects of amendment with ferrihydrite and gypsum on the structure and activity of methanogenic populations in rice field soil. Appl Environ Microbiol. 2002;68:2484–94.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    57.Lane DJ. 16S/23S rRNA sequencing. In: Stackebrandt E, Goodfellow M, editors. Nucleic acid techniques in bacterial systematics. New York: John Wiley and Sons; 1991. p. 115–75.58.Großkopf R, Janssen PH, Liesack W. Diversity and structure of the methanogenic community in anoxic rice paddy soil microcosms as examined by cultivation and direct 16S rRNA gene sequence retrieval. Appl Environ Microbiol. 1998;64:960–9.59.Reyes C, Schneider D, Thürmer A, Kulkarni A, Lipka M, Sztejrenszus SY, et al. Potentially active iron, sulfur, and sulfate reducing bacteria in Skagerrak and Bothnian Bay sediments. Geomicrobiol J. 2017;34:840–50.CAS 
    Article 

    Google Scholar 
    60.Kondo R, Nedwell DB, Purdy KJ, Silva SQ. Detection and enumeration of sulphate-reducing Bacteria in estuarine sediments by competitive PCR. Geomicrobiol J. 2004;21:145–57.CAS 
    Article 

    Google Scholar 
    61.Benjamini Y, Yekutieli D. The control of the false discovery rate in multiple testing under dependency. Ann Stat. 2001;29:1165–88.Article 

    Google Scholar 
    62.R Core Team. R: a language and environment for statistical computing, 3.6.1. Vienna, Austria: R Foundation for Statistical Computing; 2019. Available from: https://www.R-project.org.63.Oksanen J, Blanchet FG, Friendly M, Kindt R, Legendre P, McGlinn D, et al. vegan: Community Ecology Package, 2.5-6. 2019. Available from: https://CRAN.R-project.org/package=vegan.64.Quast C, Pruesse E, Yilmaz P, Gerken J, Schweer T, Yarza P, et al. The SILVA ribosomal RNA gene database project: improved data processing and web-based tools. Nucleic Acids Res. 2012;41:D590–6.PubMed 
    PubMed Central 
    Article 
    CAS 

    Google Scholar 
    65.Stamatakis A. RAxML version 8: a tool for phylogenetic analysis and post-analysis of large phylogenies. Bioinformatics. 2014;30:1312–3.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    66.Letunic I, Bork P. Interactive Tree Of Life (iTOL) v4: recent updates and new developments. Nucleic Acids Res. 2019;47:W256–9.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    67.Inkscape Team. Inkscape, 1.0.1. 2020. Available from: https://inkscape.org.68.Sun H, Spring S, Lapidus A, Davenport K, Glavina Del Rio T, Tice H, et al. Complete genome sequence of Desulfarculus baarsii type strain (2st14T). Stand Genom Sci. 2010;3:276–84.Article 

    Google Scholar 
    69.Kümmel S, Herbst F-A, Bahr A, Duarte M, Pieper DH, Jehmlich N, et al. Anaerobic naphthalene degradation by sulfate-reducing Desulfobacteraceae from various anoxic aquifers. FEMS Microbiol Ecol. 2015;91:fiv006.70.Belyakova EV, Rozanova EP, Borzenkov IA, Tourova TP, Pusheva MA, Lysenko AM, et al. The new facultatively chemolithoautotrophic, moderately halophilic, sulfate-reducing bacterium Desulfovermiculus halophilus gen. nov., sp. nov., isolated from an oil field. Microbiology. 2006;75:161–71.71.Rezadehbashi M, Baldwin SA. Core sulphate-reducing microorganisms in metal-removing semi-passive biochemical reactors and the co-occurrence of methanogens. Microorganisms. 2018;6:16.PubMed Central 
    Article 
    CAS 

    Google Scholar 
    72.Sheik CS, Jain S, Dick GJ. Metabolic flexibility of enigmatic SAR324 revealed through metagenomics and metatranscriptomics. Environ Microbiol. 2014;16:304–17.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    73.Sorokin DY, Chernyh NA. ‘Candidatus Desulfonatronobulbus propionicus’: a first haloalkaliphilic member of the order Syntrophobacterales from soda lakes. Extremophiles. 2016;20:895–901.CAS 
    PubMed 
    Article 

    Google Scholar 
    74.Lovley DR, Giovannoni SJ, White DC, Champine JE, Phillips E, Gorby YA, et al. Geobacter metallireducens gen. nov. sp. nov., a microorganism capable of coupling the complete oxidation of organic compounds to the reduction of iron and other metals. Arch Microbiol. 1993;159:336–44.75.Roden EE, Lovley DR. Dissimilatory Fe(III) reduction by the marine microorganism Desulfuromonas acetoxidans. Appl Environ Microbiol. 1993;59:734–42.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    76.Lovley DR, Coates JD, Saffarini DA, Lonergan DJ. Dissimilatory iron reduction. In: Winkelmann G, Carrano CJ, editors. Transition metals in microbial metabolism. Amsterdam: Harwood Academic Publishers; 1997. p. 187–215.77.Vandieken V, Finke N, Jørgensen BB. Pathways of carbon oxidation in an Arctic fjord sediment (Svalbard) and isolation of psychrophilic and psychrotolerant Fe(III)-reducing bacteria. Mar Ecol Prog Ser. 2006;322:29–41.CAS 
    Article 

    Google Scholar 
    78.Vandieken V, Thamdrup B. Identification of acetate-oxidizing bacteria in a coastal marine surface sediment by RNA-stable isotope probing in anoxic slurries and intact cores. FEMS Microbiol Ecol. 2013;84:373–86.CAS 
    PubMed 
    Article 

    Google Scholar 
    79.Hori T, Aoyagi T, Itoh H, Narihiro T, Oikawa A, Suzuki K, et al. Isolation of microorganisms involved in reduction of crystalline iron(III) oxides in natural environments. Front Microbiol. 2015;6:386.PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    80.Vandieken V, Mußmann M, Niemann H, Jørgensen BB. Desulfuromonas svalbardensis sp. nov. and Desulfuromusa ferrireducens sp. nov., psychrophilic, Fe(III)-reducing bacteria isolated from Arctic sediments, Svalbard. Int J Syst Evol Microbiol. 2006;56:1133–9.CAS 
    PubMed 
    Article 

    Google Scholar 
    81.Slobodkina GB, Reysenbach A-L, Panteleeva AN, Kostrikina NA, Wagner ID, Bonch-Osmolovskaya EA, et al. Deferrisoma camini gen. nov., sp. nov., a moderately thermophilic, dissimilatory iron(III)-reducing bacterium from a deep-sea hydrothermal vent that forms a distinct phylogenetic branch in the Deltaproteobacteria. Int J Syst Evol Microbiol. 2012;62:2463–8.CAS 
    PubMed 
    Article 

    Google Scholar 
    82.Tu T-H, Wu L-W, Lin Y-S, Imachi H, Lin L-H, Wang P-L. Microbial community composition and functional capacity in a terrestrial ferruginous, sulfate-depleted mud volcano. Front Microbiol. 2017;8:2137.PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    83.Lovley DR, Roden EE, Phillips EJP, Woodward JC. Enzymatic iron and uranium reduction by sulfate-reducing bacteria. Mar Geol. 1993;113:41–53.CAS 
    Article 

    Google Scholar 
    84.Bale SJ, Goodman K, Rochelle PA, Marchesi JR, Fry JC, Weightman AJ, et al. Desulfovibrio profundus sp. nov., a novel barophilic sulfate-reducing Bacterium from deep sediment layers in the Japan Sea. Int J Syst Evol Microbiol. 1997;47:515–21.85.Treude N, Rosencrantz D, Liesack W, Schnell S. Strain FAc12, a dissimilatory iron-reducing member of the Anaeromyxobacter subgroup of Myxococcales. FEMS Microbiol Ecol. 2003;44:261–9.CAS 
    PubMed 
    Article 

    Google Scholar 
    86.Hori T, Müller A, Igarashi Y, Conrad R, Friedrich MW. Identification of iron-reducing microorganisms in anoxic rice paddy soil by 13C-acetate probing. ISME J. 2010;4:267–78.87.Han Y, Perner M. The globally widespread genus Sulfurimonas: versatile energy metabolisms and adaptations to redox clines. Front Microbiol. 2015;6:989.PubMed 
    PubMed Central 

    Google Scholar 
    88.Roalkvam I, Drønen K, Stokke R, Daae FL, Dahle H, Steen IH. Physiological and genomic characterization of Arcobacter anaerophilus IR-1 reveals new metabolic features in Epsilonproteobacteria. Front Microbiol. 2015;6:987.PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    89.Schlosser C, Schmidt K, Aquilina A, Homoky WB, Castrillejo M, Mills RA, et al. Mechanisms of dissolved and labile particulate iron supply to shelf waters and phytoplankton blooms off South Georgia, Southern Ocean. Biogeosciences. 2018;15:4973–93.CAS 
    Article 

    Google Scholar 
    90.Sahade R, Lagger C, Torre L, Momo F, Monien P, Schloss I, et al. Climate change and glacier retreat drive shifts in an Antarctic benthic ecosystem. Sci Adv. 2015;1:e1500050.PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    91.Petro C, Starnawski P, Schramm A, Kjeldsen KU. Microbial community assembly in marine sediments. Aquat Micro Ecol. 2017;79:177–95.Article 

    Google Scholar 
    92.Petro C, Zäncker B, Starnawski P, Jochum LM, Ferdelman TG, Jørgensen BB, et al. Marine deep biosphere microbial communities assemble in near-surface sediments in Aarhus Bay. Front Microbiol. 2019;10:758.PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    93.Starnawski P, Bataillon T, Ettema TJ, Jochum LM, Schreiber L, Chen X, et al. Microbial community assembly and evolution in subseafloor sediment. Proc Natl Acad Sci USA. 2017;114:2940–5.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    94.Marshall IPG, Ren G, Jaussi M, Lomstein BA, Jørgensen BB, Røy H, et al. Environmental filtering determines family-level structure of sulfate-reducing microbial communities in subsurface marine sediments. ISME J. 2019;13:1920–32.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    95.Berner RA. Early diagenesis: a theoretical approach. Princeton, New Jersey: Princeton University Press; 1980.96.Cottrell MT, Kirchman DL. Natural assemblages of marine Proteobacteria and members of the Cytophaga-Flavobacter cluster consuming low- and high-molecular-weight dissolved organic matter. Appl Environ Microbiol. 2000;66:1692–7.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    97.Bissett A, Bowman JP, Burke CM. Flavobacterial response to organic pollution. Aquat Micro Ecol. 2008;51:31–43.Article 

    Google Scholar 
    98.Martinez-Garcia M, Brazel DM, Swan BK, Arnosti C, Chain PSG, Reitenga KG, et al. Capturing single cell genomes of active polysaccharide degraders: an unexpected contribution of Verrucomicrobia. PLoS ONE. 2012;7:e35314.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    99.Sabree ZL, Kambhampati S, Moran NA. Nitrogen recycling and nutritional provisioning by Blattabacterium, the cockroach endosymbiont. Proc Natl Acad Sci U S A. 2009;106:19521–6.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    100.Bowman JP, McCuaig RD. Biodiversity, community structural shifts, and biogeography of Prokaryotes within Antarctic continental shelf sediment. Appl Environ Microbiol. 2003;69:2463–83.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    101.Blazejak A, Schippers A. High abundance of JS-1- and Chloroflexi-related Bacteria in deeply buried marine sediments revealed by quantitative, real-time PCR. FEMS Microbiol Ecol. 2010;72:198–207.CAS 
    PubMed 
    Article 

    Google Scholar 
    102.Yamada T, Sekiguchi Y, Hanada S, Imachi H, Ohashi A, Harada H, et al. Anaerolinea thermolimosa sp. nov., Levilinea saccharolytica gen. nov., sp. nov. and Leptolinea tardivitalis gen. nov., sp. nov., novel filamentous anaerobes, and description of the new classes Anaerolineae classis nov. and Caldilineae classis nov. in the bacterial phylum Chloroflexi. Int J Syst Evol Microbiol. 2006;56:1331–40.CAS 
    PubMed 
    Article 

    Google Scholar 
    103.Storesund JE, Øvreås L. Diversity of Planctomycetes in iron-hydroxide deposits from the Arctic Mid Ocean Ridge (AMOR) and description of Bythopirellula goksoyri gen. nov., sp. nov., a novel Planctomycete from deep sea iron-hydroxide deposits. Antonie Van Leeuwenhoek. 2013;104:569–84.CAS 
    PubMed 
    Article 

    Google Scholar 
    104.Kovaleva OL, Merkel AY, Novikov AA, Baslerov RV, Toshchakov SV, Bonch-Osmolovskaya EA. Tepidisphaera mucosa gen. nov., sp. nov., a moderately thermophilic member of the class Phycisphaerae in the phylum Planctomycetes, and proposal of a new family, Tepidisphaeraceae fam. nov., and a new order, Tepidisphaerales ord. nov. Int J Syst Evol Microbiol. 2015;65:549–55.CAS 
    PubMed 
    Article 

    Google Scholar 
    105.Borrione I, Schlitzer R. Distribution and recurrence of phytoplankton blooms around South Georgia, Southern Ocean. Biogeosciences. 2013;10:217–31.Article 

    Google Scholar 
    106.Pfennig N, Biebl H. Desulfuromonas acetoxidans gen. nov. and sp. nov., a new anaerobic, sulfur-reducing, acetate-oxidizing bacterium. Arch Microbiol. 1976;110:3–12.CAS 
    PubMed 
    Article 

    Google Scholar 
    107.Finster K, Bak F, Pfennig N. Desulfuromonas acetexigens sp. nov., a dissimilatory sulfur-reducing eubacterium from anoxic freshwater sediments. Arch Microbiol. 1994;161:328–32.CAS 
    Article 

    Google Scholar 
    108.Lovley DR, Phillips EJP, Lonergan DJ, Widman PK. Fe(III) and S0 reduction by Pelobacter carbinolicus. Appl Environ Microbiol. 1995;61:2132–8.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    109.An TT, Picardal FW. Desulfuromonas carbonis sp. nov., an Fe(III)-, S0- and Mn(IV)-reducing bacterium isolated from an active coalbed methane gas well. Int J Syst Evol Microbiol. 2015;65:1686–93.CAS 
    PubMed 
    Article 

    Google Scholar 
    110.Pjevac P, Kamyshny A Jr, Dyksma S, Mußmann M. Microbial consumption of zero-valence sulfur in marine benthic habitats. Environ Microbiol. 2014;16:3416–30.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    111.Miao Z-Y, He H, Tan T, Zhang T, Tang J-L, Yang Y-C, et al. Biotreatment of Mn2+ and Pb2+ with sulfate-reducing bacterium Desulfuromonas alkenivorans S-7. J Environ Eng. 2018;144:04017116.Article 

    Google Scholar 
    112.Buongiorno J, Herbert L, Wehrmann L, Michaud A, Laufer K, Røy H, et al. Complex microbial communities drive iron and sulfur cycling in Arctic fjord sediments. Appl Environ Microbiol. 2019;85:e00949-19.PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    113.Zhang H, Liu F, Zheng S, Chen L, Zhang X, Gong J. The differentiation of iron-reducing bacterial community and iron-reduction activity between riverine and marine sediments in the Yellow River estuary. Mar Life Sci Technol. 2020;2:87–96.Article 

    Google Scholar 
    114.Ravenschlag K, Sahm K, Pernthaler J, Amann R. High bacterial diversity in permanently cold marine sediments. Appl Environ Microbiol. 1999;65:3982–9.115.Kashefi K, Holmes DE, Baross JA, Lovley DR. Thermophily in the Geobacteraceae: Geothermobacter ehrlichii gen. nov., sp. nov., a novel thermophilic member of the Geobacteraceae from the “Bag City” hydrothermal vent. Appl Environ Microbiol. 2003;69:2985–93.116.Holmes DE, Nicoll JS, Bond DR, Lovley DR. Potential role of a novel psychrotolerant member of the family Geobacteraceae, Geopsychrobacter electrodiphilus gen. nov., sp. nov., in electricity production by a marine sediment fuel cell. Appl Environ Microbiol. 2004;70:6023–30.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    117.Jørgensen BB. Mineralization of organic matter in the sea bed—the role of sulphate reduction. Nature. 1982;296:643–5.Article 

    Google Scholar 
    118.Bryant M, Campbell LL, 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. 1977;33:1162–9.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    119.Muyzer G, Stams AJ. The ecology and biotechnology of sulphate-reducing bacteria. Nat Rev Microbiol. 2008;6:441–54.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    120.Dalsgaard T, Bak F. Nitrate reduction in a sulfate-reducing bacterium, Desulfovibrio desulfuricans, isolated from rice paddy soil: sulfide inhibition, kinetics, and regulation. Appl Environ Microbiol. 1994;60:291–7.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    121.Holmes DE, Bond DR, Lovley DR. Electron transfer by Desulfobulbus propionicus to Fe (III) and graphite electrodes. Appl Environ Microbiol. 2004;70:1234–7.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    122.Lovley DR, Phillips EJP. Competitive mechanisms for inhibition of sulfate reduction and methane production in the zone of ferric iron reduction in sediments. Appl Environ Microbiol. 1987;53:2636–41.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    123.Finke N, Vandieken V, Jørgensen BB. Acetate, lactate, propionate, and isobutyrate as electron donors for iron and sulfate reduction in Arctic marine sediments, Svalbard. FEMS Microbiol Ecol. 2007;59:10–22.CAS 
    PubMed 
    Article 

    Google Scholar 
    124.Canfield DE, Thamdrup B, Hansen JW. The anaerobic degradation of organic matter in Danish coastal sediments: iron reduction, manganese reduction, and sulfate reduction. Geochim Cosmochim Acta. 1993;57:3867–83.CAS 
    PubMed 
    Article 

    Google Scholar 
    125.Jørgensen BB. The sulfur cycle of a coastal marine sediment (Limfjorden, Denmark). Limnol Oceanogr. 1977;22:814–32.Article 

    Google Scholar 
    126.Jørgensen BB, Laufer K, Michaud AB, Wehrmann LM. Biogeochemistry and microbiology of high Arctic marine sediment ecosystems—case study of Svalbard fjords. Limnol Oceanogr. 2021;66:S273–92.Article 
    CAS 

    Google Scholar 
    127.Laufer K, Michaud AB, Røy H, Jørgensen BB. Reactivity of iron minerals in the seabed toward microbial reduction—a comparison of different extraction techniques. Geomicrobiol J. 2020;37:170–89.Article 

    Google Scholar 
    128.Holmkvist L, Ferdelman TG, Jørgensen BB. A cryptic sulfur cycle driven by iron in the methane zone of marine sediment (Aarhus Bay, Denmark). Geochim Cosmochim Acta. 2011;75:3581–99.CAS 
    Article 

    Google Scholar 
    129.Riedinger N, Brunner B, Formolo MJ, Solomon E, Kasten S, Strasser M, et al. Oxidative sulfur cycling in the deep biosphere of the Nankai Trough, Japan. Geology. 2010;38:851–4.CAS 
    Article 

    Google Scholar  More

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    Novel metagenome-assembled genomes involved in the nitrogen cycle from a Pacific oxygen minimum zone

    Oxygen minimum zones (OMZs) are unique oceanic regions with strong redox gradients. Anoxic zones in OMZs are hotspots for fixed nitrogen loss and production of the greenhouse gas N2O [1, 2]. Microbes in OMZs make important contributions to biogeochemistry, which motivates us to reconstruct metagenome-assembled genomes (MAGs) from the Eastern Tropical South Pacific (ETSP) OMZ (Fig. 1a, b). Among 147 recovered MAGs, we present 39 high- and medium-quality MAGs with completeness >50% and contamination 100 nM d−1) at the same station [6], where MAGs were recovered. Consistently, Thaumarchaeota MAGs (AOAs) were nearly absent (only AOA-2 had a relative abundance higher than 0.01%) and NOB MAGs (NOB-1 and NOB-2) were much more abundant than AOA in the anoxic core (Fig. 1d). MAGs in this study will provide opportunities to discover novel processes and adaptation strategies.Most MAGs had their highest relative abundances in the anoxic zone (Fig. 1c). Many of them contribute to the loss of fixed nitrogen, which occurs by denitrification (the sequential reduction of nitrate to nitrite, NO, N2O, and finally N2) and anammox (anaerobic oxidation of ammonium to N2). Measured nitrate reduction rates at this [5, 8] and other [16, 17] nearby stations were much larger than rates of any subsequent denitrification steps (e.g., nitrite reduction to N2O or to N2). Consistently, preliminary prediction of metabolisms shows that more than half of the MAGs found here contained nar, which encodes nitrate reduction, and one-third of those contained only nar and none of the other denitrification genes (i.e., they are nitrate-reducing specialists) (Fig. 2). Consistently, a previous study found that nar dramatically outnumbered the other denitrification genes in contigs from the Eastern Tropical North Pacific (ETNP) OMZ [18]. Indeed, four of the five most abundant MAGs in the anoxic core were nitrate-reducing specialists (Fig. 2). The fifth was an anammox MAG, which was only assigned to the genus level (Candidatus Scalindua) in GTDB and was not represented at the species level in the Tara Oceans dataset (Table S1). However, this anammox MAG was highly related to 20 anammox single-cell amplified genomes (SAGs) from the ETNP OMZ [19]. The anammox MAG had at least 90% average nucleotide identity (ANI) to the SAGs, with the highest ANI (98.8%) to SAG K21. Consistent with the previous work [19], the anammox MAG also encoded cyanase, indicating its potential of using organic nitrogen substrates. The most abundant nitrate reducer MAG here is Marinimicrobia-1 (Fig. 1), which belongs to the newly proposed phylum Candidatus Marinimicrobia [20]. Notably, one nitrate reducer can only be assigned to phylum level (Candidatus Wallbacteria) and was not present in the Tara Oceans MAGs (Table S1).We also identified a novel archaeal MAG possessing multiple denitrification genes. MG-II MAG-2 encoded Nar alpha and beta subunits, nitrate/nitrite transporters, copper-containing nitrite reductase, and N2O reductase (Fig. 2). Two MAGs from the Tara Oceans metagenomes (Table S1) were identified as the same species as MG-II MAG-2. TOBG_NP-110 (ANI to MG-II MAG-2 = 99.8%) from the North Pacific encoded Nar and nitrate/nitrite transporters, and TOBG_SP-208 (ANI to MG-II MAG-2 = 99.6%) from the South Pacific also contained the same denitrification genes as MG-II MAG-2 (Table S2). In addition, two MG-II SAGs (AD-615-F09 and AD-613-O09) were found at a different station of the ETSP OMZ sampled on the same cruise as this study [21]. Partial 16S rRNA genes of both SAGs are 100% identical to that of MG-II MAG-2 (alignment length = 200 bp for AD-615-F09 and 183 bp for AD-613-O09), but only AD-615-F09 might be the same species as MG-II MAG-2 based on ANI analyses (MG-II MAG-2 had 99.5% ANI to AD-615-F09, and 80.9% to AD-613-O09). Both SAGs also encoded Nar and nitrate/nitrite transporters [21]. The absence of other denitrification genes may be due to the low completeness of the two SAGs (completeness = 5.61% for both SAGs) [21]. Nitrite reductase and N2O reductase genes were located on the same contig in both MG-II MAG-2 and TOBG_SP-208 (Table S2). MG-II MAG-2 and TOBG_SP-208 had low contamination (1.9% and 0.8%, respectively), and their contigs with nitrite reductase and N2O reductase genes contained single-copy marker genes present only once in each MAG (Supplementary Methods). Although these results suggest a nearly complete denitrification metabolism in MG-II archaea, especially N2O consumption metabolism, methods besides metagenomics (e.g. reconstructing SAGs with high completeness) are highly recommended to rule out possible artifacts introduced by metagenomic binning and confirm the presence of these genes and their denitrification activity. Nonetheless, MG-II MAG-2 was present (Fig. 1e) and transcriptionally active in both Pacific OMZs (Fig. S2), indicating its adaptation to low oxygen environments. The MG-III MAG did not have any denitrification genes but was abundant in the anoxic zone (Figs. 1e and 2). It had a GC value (43.2%) distinct from all other known MG-III MAGs [22] and is the most complete (86.0%) and the least contaminated (0%) (Table S1) among all reported MG-III MAGs, indicating that MG-III is a novel archaeon in this group. Bacterial and archaeal MAGs recovered here implied that nitrogen metabolisms were present in more microbial lineages than previously thought. Further analyses of these MAGs will shed light on adaptation strategies in the unique OMZ environment and novel functions related to important element cycles. More

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    Scenario simulation of land use and land cover change in mining area

    Data source and preprocessingConsidering factors such as amount of cloud and time intervals of image, four remote sensing images with a spatial resolution of 30 m, including Landsat 5 Thematic Mapper (TM) images for 08-21-2000, 09-04-2005 and 09-18-2010, and Landsat 8 Operational Land Imager (OLI) for 09-02-2016,were obtained from the Geospatial Data Cloud Platform (http://www.gscloud.cn). LULC information was extracted from these remote sensing images. In addition, the digital elevation model (DEM) with a spatial resolution of 30 m was obtained from the website. Elevation and slope information were derived from DEM data and used as terrain driving factors for scenario simulation. Other supporting data, such as Weishan County land use data, mine distribution data, general land use planing (2006–2020) and mineral resources planning (2008–2015), Jining City coal mining subsidence land rearrangement planning (2016–2030), were obtained from Weishan Natural Resources and Planning Bureau. These data were used for better data analysis.Considering severe ground subsidence and seeper in the study area, and referring to national standards: Current Land Use Classification (GB/T 21010-2017), remote sensing images were interpreted into six LULC types: farmland, other agricultural land, urban and rural construction land, subsided seeper area, water area, and tidal wetland.In the process of image interpretation, firstly, the remote sensing image was divided into two regions: one region were the lake and the surrounding tidal wetland, and the other region included farmland, other agricultural land, urban and rural construction land, subsided seeper area, etc.In region 1, decision tree classification, combined with the Modified Normalized Difference Water Index (MNDWI), was used to extract lakes. Then we masked them in region 1. The Normalized Difference Vegetation Index (NDVI) was calculated for the remaining image of region 1. Tidal wetland was mainly distributed along rivers and lakes, and NDVI value was higher than that of farmland and other vegetation. By analyzing its geographical distribution and NDVI value, and referring to Weishan County land use data, the appropriate threshold was selected to extract tidal wetland.The spectral signature of rivers, ditches and aquaculture ponds in other agricultural land in region 2 could be easily distinguished from other surface features. They could be extracted step by step by manual visual interpretation and empirical knowledge, referring to Weishan County land use data and water system data. Then we masked them separately in region 2. After extracting rivers, ditches, aquaculture ponds with high water content, the remaining LULC type with high water content in region 2 was subsided seeper area. According to the relationship of spectral signature of different LULC types, it was concluded that among the remaining LULC types in region 2, only TM3 band value of subsided seeper area was higher than TM5 band value. Using this characteristic, subsided seeper area could be distinguished from other LULC types. After extracting subsided seeper area, the remaining LULC types in region 2 were farmland and urban and rural construction land. The spectral characteristics of them were very different. Therefore, they could be distinguished using support vector machine (SVM) classification method, and their respective binary images were generated using decision tree method.The extracted six LULC types were shown in single layer and binary form respectively. Six LULC types were coded and synthesized into one image. We obtained 2000, 2005, 2010, 2016 LULC type maps (Fig. 2). Finally classification post-processing and accuracy evaluation were operated.Figure 2The LULC types maps of 2000, 2005, 2010 and 2016. Maps were generated using ArcGIS 10.1 for Desktop (http://www.esri.com/software/arcgis/arcgis-for-desktop).Full size imageThe accuracy of the interpretation results was verified by confusion matrix and kappa coefficient. The kappa coefficients of the four interpretation maps were 0.84, 0.85, 0.82 and 0.86, respectively (Table 1). The accuracy could meet the needs of further research.Table 1 Accuracy evaluation of the interpretation results (%).Full size tableBy reading previous research results37,38,39,40,41, based on the entropy theory, in the same study area, high spatial resolution data contains more entropy than low spatial resolution data, and reflecting more detailed information, but it will increase the uncertainty of prediction results and reduce the prediction accuracy. Although the prediction accuracy of low spatial resolution data increases, it will lose lots of detailed information. In order to ensure the accuracy of the simulation, considering the area of the study area and data requirement of the CLUE-S model, the interpreted LULC maps with a resolution of 30 m exceed the upper limit of the CLUE-S model data requirement, so the LULC maps were resampled to multiple scales including 60 m, 90 m, 120 m, and 150 m to facilitate logistic regression analysis of LULC types and driving factors.Selection and processing of driving factorsTo interpret the relationship between the LULC and its driving factors in the mining area, we not only need to identify the driving factors that have greater explanatory power for LULC change, but also need to quantitatively describe the relationship between driving factors and LULC types.Considering the accessibility, usability of the data and the actual conditions in the study area, seven driving factors were selected based on the land use map of Weishan County in 2005 and the DEM data5,10,11,13,26,28,29,30. The driving factors included: (1) terrain factors, including elevation and slope factors; (2) five accessibility factors, including the nearest distance between each grid pixel and the main roads, the major rivers, the residential area, the major mines, and the ditches. The 30 m grid data of each driving factor were resampled to 60 m, 90 m, 120 m and 150 m respectively.In this study, BLRM was used to explore the relationship between LULC change and the related 7 driving factors. BLRM is sensitive to multicollinearity. In order to eliminate the influence of collinearity on the regression results, the multicollinearity between independent variables was diagnosed before the regression model was established.The receiver operating characteristic (ROC) curve was used to evaluate the accuracy of regression analysis results at different scales. The results showed that using 60 m resolution provided more accurate regression analysis results and suffered less loss of LULC and driving factor information during resampling. Therefore, we used 60 m × 60 m grid cell data to driving forces analysis.Raster maps of each driving factor at a resolution scale of 60 m are shown in Fig. 3.Figure 3Raster maps of driving factors at a resolution scale of 60 m. Maps were generated using ArcGIS 10.1 for Desktop (http://www.esri.com/software/arcgis/arcgis-for-desktop).Full size imageLogistic regression analysis of LULC types and driving factorsBLRM is often used for regression analysis of explanatory binary variables. The presence and absence of a certain type of LULC in a specific area is set as 1 and 0, respectively, which is characteristic for binary variable. Therefore, we used BLRM to calculate the probability (P) of various LULC types in a specific spatial location, and its mathematical expression is:$$begin{aligned} ln left( frac{P}{1-P}right) = beta _0 + beta _1 X end{aligned}$$
    (1)
    where (frac{P}{1-P}) is the ’odds ratio’ of an event, abbreviated as ( Omega ), which represents the odds that an outcome will occur given a particular condition compared to the odds of the outcome occurring in the absence of that condition; (beta _0) is a constant; (beta _1) is the correlation coefficient of an explaining variable and an explained variable. Making mathematical transformation of the above expression, we get: (Omega = (frac{P}{1-P}) = e^{beta _0 + beta _1 X}).Regression analysis using BLRM, we divided the study area into many grid cells. Taking each LULC type as the explained variable, and the driving factor causing LULC change as the explanatory variable, we calculated the odds ratio of each LULC type in a specific spatial location, and analyzed the relationship between each LULC type and the driving factors. The calculating equation is:$$begin{aligned} mathrm{Logit} P = ln left( frac{P_i}{1-P_i}right) = beta _0 + beta _1 X_{1,i} + beta _2 X_{2,i} + cdots + beta _n X_{n,i} end{aligned}$$
    (2)
    Making mathematical transformation of the above equation, we get:$$begin{aligned} P_i = frac{e^{(beta _0 + beta _1 X_{1,i} + beta _2 X_{2,i} + cdots + beta _n X_{n,i})}}{1+e^{(beta _0 + beta _1 X_{1,i} + beta _2 X_{2,i} + cdots + beta _n X_{n,i})}} end{aligned}$$
    (3)
    where: (P_i) is the probability of a certain LULC type i in a grid cell, (X_{1,i}sim X_{n,i}) are the driving factors of LULC type i, (beta _0) is the constant, (beta _1sim beta _n) are the correlation coefficients of each driving factor and LULC type i.The receiver operating characteristic (ROC) was used to evaluate the accuracy of regression analysis results. The accuracy can be measured by calculating the area under the ROC curve. The area value is between 0.5 and 1. The closer the value is to 1, the higher the accuracy is. In general, the area under the ROC curve is greater than 0.7, which indicates that the selected factor has good explanatory power27,42.CLUE-S simulation and accuracy validationBefore using the CLUE-S model for futural LULC scenario simulation in mining area, the prediction accuracy needs to be verified. Based on the data of LULC in 2005, the spatial distribution pattern of LULC in 2016 was predicted firstly.The modeling accuracy was evaluated based on the Kappa index by comparing the actual LULC map in 2016 with the simulated in 201627,43,44. Equation (4) gives one of the most popular Kappa index equations: i.e.,$$begin{aligned} mathrm{Kappa}=frac{P_o-P_c}{P_p-P_c} end{aligned}$$
    (4)
    where (P_o) is the observed proportion correct, (P_c) is the expected proportion correct due to chance, (P_c) =1/n, n is the number of LULC types, and (P_p) is the proportion correct when classification is perfect.In order to further verify the accuracy of the model simulation, we also calculated kappa for quantity (Kquantity).Scenario setting of futural LULC simulationDue to the continuous population growth and mineral exploitation in the study area, the land resources, especially farmland resources, have become increasingly scarce and the environment has been deteriorating. Based on the simulation and validated results during 2005-2016, we defined three scenarios—namely natural development scenario, ecological protection scenario, and farmland protection scenario—to predict LULC spatial patterns for 2025.Natural development scenarioIn this scenario, the land use demand of the study area was basically not restricted by policies in near future. We assumed that the change rate of each LULC type in near future was consistent with the change trend from 2005 to 2016. So it is defined as natural development scenario. Using Markov model to obtain the area transition probability matrix of each year from 2017 to 2025, and taking the proportion of each LULC type area in the total study area in 2005 as the initial state matrix, the area of each LULC type in 2025 under the natural development scenario was predicted.Based on the characteristics and trend of the LULC change from 2005 to 2016, after appropriately adjusting the transition probability matrix of different LULC types, we predicted the demands of each LULC type in 2025 under ecological protection scenario and farmland protection scenario using Markov model45,46.Ecological protection scenarioThis scenario emphasizes protecting the ecological environment, restricting the conversion of the LULC types that have more regulatory effects on the ecosystem, such as tidal wetland and water area, to other land use types. Garden land, woodland, grassland, and aquaculture land, belong to other agricultural land, which have regulatory effects on the local ecosystem, so their conversion to other LULC types should be restricted as well.Farmland protection scenarioAccording to the guidelines of “the general land use planning in Weishan County (2006-2020)”, we should maximize the potential use of current construction land, implement intensive and economical utilization of construction land, and use less or not use farmland to economical construction. So in order to ensure the dynamic balance of total farmland amount and the regional food supply security, in the farmland protection scenario, the conversion from farmland to other land use types should be restricted. The projected land use demands for 2025 under the three different scenarios are shown in Table 2.Table 2 Areas of LULC types in 2025 under different scenarios (ha).Full size table More

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    Exploring physicochemical and cytogenomic diversity of African cowpea and common bean

    1.Lewis, G. P. Legumes of the World (Royal Botanic Gardens, 2005).
    Google Scholar 
    2.The Legume Phylogeny Working Group (LPWG). A new subfamily classification of the Leguminosae based on a taxonomically comprehensive phylogeny. Taxon 66, 44–77 (2017).Article 

    Google Scholar 
    3.Yahara, T. et al. Global legume diversity assessment: Concepts, key indicators, and strategies. Taxon 62, 249–266 (2013).Article 

    Google Scholar 
    4.Odendo, M., Bationo, A. & Kimani, S. Socio-economic contribution of legumes to livelihoods in Sub-Saharan Africa. In Fighting Poverty in Sub-Saharan Africa: The Multiple Roles of Legumes in Integrated Soil Fertility Management (eds Bationo, A. et al.) 27–46 (Springer, 2011).Chapter 

    Google Scholar 
    5.Dakora, F. D. & Keya, S. O. Contribution of legume nitrogen fixation to sustainable agriculture in Sub-Saharan Africa. Soil Biol. Biochem. 29, 809–817 (1997).CAS 
    Article 

    Google Scholar 
    6.Ajeigde, H. A., Singh, B. B. & Osenj, T. O. Cowpea-cereal intercrop productivity in the Sudan savanna zone of Nigeria as affected by planting pattern, crop variety and pest management. Afr. Crop Sci. J. 13, 269–279 (2005).
    Google Scholar 
    7.Rahmanian, M., Batello, C. & Calles, T. Pulse Crops for Sustainable Farms in Sub-Saharan Africa (FAO, 2018).
    Google Scholar 
    8.Rawal, V. & Navarro, D. K. The Global Economy of Pulses (FAO, 2017).
    Google Scholar 
    9.Plants of the World Online. http://powo.science.kew.org (2020).10.Broughton, W. J. et al. Beans (Phaseolus spp.)—Model food legumes. Plant Soil 252, 55–128 (2003).CAS 
    Article 

    Google Scholar 
    11.Delgado-Salinas, A., Bibler, R. & Lavin, M. Phylogeny of the genus Phaseolus (Leguminosae): A recent diversification in an ancient landscape. Syst. Bot. 31, 779–791 (2006).Article 

    Google Scholar 
    12.Greenway, P. J. Origins of some East African food plants: Part V. East Afr. Agric. J. 11, 56–63 (1945).
    Google Scholar 
    13.Wortmann, C. S. & Allen, D. J. African Bean Production Environments: Their Definition, Characteristics and Constraints. Occasional Publication Series 11 (CIAT, 1994).
    Google Scholar 
    14.Maxted, N. et al. African Vigna: Systematic and Ecogeographic Studies (International Plant Genetic Resource Institute, 2004).
    Google Scholar 
    15.Singh, B. B. Cowpea: The Food Legume of the 21st Century (Crop Science Society of America Inc., 2014).Book 

    Google Scholar 
    16.Catarino, S. et al. Conservation priorities for African Vigna species: Unveiling Angola’s diversity hotspots. Glob. Ecol. Conserv. 25, e01415. https://doi.org/10.1016/j.gecco.2020.e01415 (2021).Article 

    Google Scholar 
    17.Vidigal, P., Romeiras, M. M. & Monteiro, F. Crops diversification and the role of orphan legumes to improve the Sub-Saharan Africa farming systems. In Sustainable Crop Production (ed. Hasanuzzaman, M.) (IntechOpen, 2019).
    Google Scholar 
    18.Maréchal, R. Etude taxonomique d’un groupe complexe d’espèces des genres Phaseolus et Vigna (Papilionaceae) sur la base de données morphologiques et polliniques, traitées par l’analyse informatique. Boissiera 28, 1–273 (1978).
    Google Scholar 
    19.Peksen, E., Peksen, A. & Gulumser, A. Leaf and stomata characteristics and tolerance of cowpea cultivars to drought stress based on drought tolerance indices under rainfed and irrigated conditions. Int. J. Curr. Microbiol. Appl. Sci. 3, 626–634 (2014).CAS 

    Google Scholar 
    20.Iqbal, A., Khalil, I. A., Ateeq, N. & Khan, M. S. Nutritional quality of important food legumes. Food Chem. 97, 331–335 (2006).CAS 
    Article 

    Google Scholar 
    21.African Orphan Crops Consortium. http://africanorphancrops.org/meet-the-crops/ (2021)22.Boukar, O. et al. Cowpea. In Grain Legumes (ed. de Ron, A. M.) 219–250 (Springer, 2015).Chapter 

    Google Scholar 
    23.Animasaun, D. A., Oyedeji, S., Azeez, Y. K., Mustapha, O. T. & Azeez, M. A. Genetic variability study among ten cultivars of cowpea (Vigna unguiculata L. Walp) using morpho-agronomic traits and nutritional composition. J. Agric. Sci. 10, 119–130 (2015).
    Google Scholar 
    24.Timko, M. P. & Singh, B. B. Cowpea, a multifunctional legume. In Plant Genetics and Genomics: Crops and Models Vol. 1 (eds Moore, P. H. & Ming, R.) 227–258 (Springer, 2008).
    Google Scholar 
    25.Wortmann, S. C., Kirkby, A. R., Eledu, A. C. & Allen, J. D. Atlas of Common Bean (Phaseolus vulgaris L.) Production in Africa (International Centre for Tropical Agriculture, 2004).
    Google Scholar 
    26.Guignard, M. S. et al. Genome size and ploidy influence angiosperm species’ biomass under nitrogen and phosphorus limitation. New Phytol. 210, 1195–1206 (2016).PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    27.Sheidai, M. et al. Genetic diversity and genome size variability in Linum austriacum (Lineaceae) populations. Biochem. Syst. Ecol. 57, 20–26 (2014).CAS 
    Article 

    Google Scholar 
    28.Kron, P., Suda, J. & Husband, B. C. Applications of flow cytometry to evolutionary and population biology. Annu. Rev. Ecol. Evol. Syst. 38, 847–876 (2007).Article 

    Google Scholar 
    29.Wu, Y. Q. et al. Genetic analyses of Chinese Cynodon accessions by flow cytometry and AFLP markers. Crop Sci. 46, 917–926 (2016).Article 

    Google Scholar 
    30.Parida, A., Raina, S. N. & Narayan, R. K. J. Quantitative DNA variation between and within chromosome complements of Vigna species (Fabaceae). Genetica 82, 125–133 (1990).CAS 
    Article 

    Google Scholar 
    31.Nagl, W. & Treviranus, A. A flow cytometric analysis of the nuclear 2C DNA content in 17 Phaseolus species (53 genotypes). Bot. Acta 108, 403–406 (1995).CAS 
    Article 

    Google Scholar 
    32.Barow, M. & Meister, A. Endopolyploidy in seed plants is differently correlated to systematics, organ, life strategy and genome size. Plant Cell Environ. 26, 571–584 (2003).Article 

    Google Scholar 
    33.Lonardi, S. et al. The genome of cowpea (Vigna unguiculata [L.] Walp.). Plant J. 98, 767–782 (2019).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    34.The IUCN Red List of Threatened Species. Version 2020-2. https://www.iucnredlist.org/ (2020).35.Genesys. Plant Genetic Resources Accession. https://www.genesys-pgr.org/ (2021).36.Pope, G. V. & Polhill, R. M. Flora Zambesiaca, part 5 Vol. 3 (Royal Botanic Gardens, 2001).
    Google Scholar 
    37.Tomooka, N., Vaughan, D. A., Moss, H. & Maxted, N. The Asian Vigna: Genus Vigna Subgenus Ceratotropis Genetic Resources (Kluwer Academic Publishers, 2002).Book 

    Google Scholar 
    38.Debouck, D. G. Primary diversification of Phaseolus in the Americas: Three centers. Plant Genet. Resour. Newsl. 67, 2–8 (1986).
    Google Scholar 
    39.Plant Resources of Tropical Africa. https://www.prota4u.org/database/ (2021).40.Linder, H. P. The evolution of African plant diversity. Front. Ecol. Evol. 2, 38. https://doi.org/10.3389/fevo.2014.00038 (2014).Article 
    ADS 

    Google Scholar 
    41.Romeiras, M. M., Figueira, R., Duarte, M. C., Beja, P. & Darbyshire, I. Documenting biogeographical patterns of African timber species using herbarium records: A conservation perspective based on native trees from Angola. PLoS ONE 9, e103403. https://doi.org/10.1371/journal.pone.0103403 (2014).CAS 
    Article 
    PubMed 
    PubMed Central 
    ADS 

    Google Scholar 
    42.Catarino, S. et al. Spatial and temporal trends of burnt area in angola: Implications for natural vegetation and protected area management. Diversity 12, 307. https://doi.org/10.3390/d12080307 (2020).Article 

    Google Scholar 
    43.Catarino, S., Duarte, M. C., Costa, E., Carrero, P. G. & Romeiras, M. M. Conservation and sustainable use of the medicinal Leguminosae plants from Angola. PeerJ 7, e6736. https://doi.org/10.7717/peerj.6736 (2019).Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    44.Romeiras, M. M. et al. IUCN Red List assessment of the Cape Verde endemic flora: Towards a global strategy for plant conservation in Macaronesia. Bot. J. Linn. Soc. 180, 413–425 (2016).Article 

    Google Scholar 
    45.Gomes, A. M. et al. Drought response of cowpea (Vigna unguiculata (L.) Walp.) landraces at leaf physiological and metabolite profile levels. Environ. Exp. Bot. 175, 104060. https://doi.org/10.1016/j.envexpbot.2020.104060 (2020).CAS 
    Article 

    Google Scholar 
    46.The International Institute of Tropical Agriculture (IITA). https://www.iita.org/ (2021)47.Fatokun, C. et al. Genetic diversity and population structure of a mini-core subset from the world cowpea (Vigna unguiculata (L.) Walp.) germplasm collection. Sci. Rep. 8, 16035. https://doi.org/10.1038/s41598-018-34555-9 (2018).CAS 
    Article 
    PubMed 
    PubMed Central 
    ADS 

    Google Scholar 
    48.Rocha, V., Duarte, M. C., Catarino, S., Duarte, I. & Romeiras, M. M. Cabo Verde’s Poaceae flora: A reservoir of crop wild relatives diversity for crop improvement. Front. Plant Sci. 12, 630217. https://doi.org/10.3389/fpls.2021.630217 (2021).Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    49.Brilhante, M. et al. Tackling food insecurity in Cabo Verde Islands: The nutritional, agricultural and environmental values of the legume species. Foods 10, 206. https://doi.org/10.3390/foods10020206 (2021).Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    50.Pasquet, R. S. Wild cowpea (Vigna unguiculata) evolution. In Advances in Legume Systematics 8: Legumes of Economic Importance (eds Pickersgill, B. & Lock, J. M.) 95–100 (Royal Botanic Gardens, 1996).
    Google Scholar 
    51.Di Bella, G. et al. Mineral composition of some varieties of beans from Mediterranean and Tropical areas. Int. J. Food Sci. Nutr. 67, 239–248 (2016).PubMed 
    Article 
    CAS 
    PubMed Central 

    Google Scholar 
    52.Gelin, J. R., Forster, S., Grafton, K. F., McClean, P. E. & Rojas-Cifuentes, G. A. Analysis of seed zinc and other minerals in a recombinant inbred population of navy bean (Phaseolus vulgaris L.). Crop Sci. 47, 1361–1366 (2007).CAS 
    Article 

    Google Scholar 
    53.Dakora, F. D. & Belane, A. K. Evaluation of protein and micronutrient levels in edible cowpea (Vigna unguiculata L. Walp) leaves and seeds. Front. Sustain. Food Syst. 3, 70. https://doi.org/10.3389/fsufs.2019.00070 (2019).Article 

    Google Scholar 
    54.Yeken, M. Z., Akpolat, H., Karaköy, T. & Çiftçi, V. Assessment of mineral content variations for biofortification of the bean seed. Int. J. Agric. Sci. 4, 261–269 (2018).
    Google Scholar 
    55.Gondwe, T. M., Alamu, E. O., Mdziniso, P. & Maziya-Dixon, B. Cowpea (Vigna unguiculata (L.) Walp) for food security: An evaluation of end-user traits of improved varieties in Swaziland. Sci. Rep. 9, 15991. https://doi.org/10.1038/s41598-019-52360-w (2019).CAS 
    Article 
    PubMed 
    PubMed Central 
    ADS 

    Google Scholar 
    56.Sperotto, R. A., Ricachenevsky, F. K., Williams, L. E., Vasconcelos, M. W. & Menguer, P. K. From soil to seed: Micronutrient movement into and within the plant. Front. Plant Sci. 5, 438. https://doi.org/10.3389/fpls.2014.00438 (2014).Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    57.Maziya-Dixon, B., Kling, J. G., Menkir, A. & Dixon, A. Genetic variation in total carotene, iron, and zinc contents of maize and cassava genotypes. Food Nutr. Bull. 21, 419–422 (2000).Article 

    Google Scholar 
    58.Shewfelt, R. L. Sources of variation in the nutrient content of agricultural commodities from the farm to the consumer. J. Food Qual. 13, 37–54 (1990).Article 

    Google Scholar 
    59.World Health Organization. The World Health Report 2006: Working Together for Health. https://www.who.int/whr/2006/whr06_en.pdf?ua=1 (2006).60.Gödecke, T., Stein, A. J. & Qaim, M. The global burden of chronic and hidden hunger: Trends and determinants. Glob. Food Sec. 17, 21–29 (2018).Article 

    Google Scholar 
    61.Shankar, A. H. Mineral deficiencies. In Hunter’s Tropical Medicine and Emerging Infectious Diseases (eds Ryan, E. T. et al.) 1048–1054 (Elsevier, 2020).Chapter 

    Google Scholar 
    62.Muthayya, S. et al. The global hidden hunger indices and maps: An advocacy tool for action. PLoS ONE 8, e67860. https://doi.org/10.1371/journal.pone.0067860 (2013).CAS 
    Article 
    PubMed 
    PubMed Central 
    ADS 

    Google Scholar 
    63.Joy, E. J. et al. Dietary mineral supplies in Africa. Physiol. Plant. 151, 208–229 (2014).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    64.World Health Organization. World health statistics 2015. https://apps.who.int/iris/bitstream/handle/10665/170250/9789240694439_eng.pdf;jsessionid=9CFCB446F9217B60415DD216E70F6A49?sequence=1 (2015).65.Muriuki, J. M. et al. Estimating the burden of iron deficiency among African children. BMC Med. 18, 31. https://doi.org/10.1186/s12916-020-1502-7 (2020).CAS 
    Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    66.Official Journal of the European Union. Regulation (Eu) No 1169/2011 of the European Parliament and of the Council of 25 October 2011. https://eur-lex.europa.eu/legal-content/EN/TXT/PDF/?uri=CELEX:32011R1169&from=EN (2011).67.Nowicka, A. et al. Nuclear DNA content variation within the genus Daucus (Apiaceae) determined by flow cytometry. Sci. Hortic. 209, 132–138 (2016).CAS 
    Article 

    Google Scholar 
    68.Guilengue, N., Alves, S., Talhinhas, P. & Neves-Martins, J. Genetic and genomic diversity in a tarwi (Lupinus mutabilis Sweet) germplasm collection and adaptability to Mediterranean climate conditions. Agronomy 10, 21. https://doi.org/10.3390/agronomy10010021 (2020).Article 

    Google Scholar 
    69.Chable, V. et al. Embedding cultivated diversity in society for agro-ecological transition. Sustainability 12, 784. https://doi.org/10.3390/su12030784 (2020).Article 

    Google Scholar 
    70.Knight, C. A., Molinari, N. A. & Petrov, D. A. The large genome constraint hypothesis: Evolution, ecology and phenotype. Ann. Bot. 95, 177–190 (2005).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    71.Pati, K., Zhang, F. & Batley, J. First report of genome size and ploidy of the underutilized leguminous tuber crop Yam Bean (Pachyrhizus erosus and P. tuberosus) by flow cytometry. Plant Genet. Resour. 17, 456–459 (2019).CAS 
    Article 

    Google Scholar 
    72.Sliwinska, E. Flow cytometry—A modern method for exploring genome size and nuclear DNA synthesis in horticultural and medicinal plant species. Folia Hortic. 30, 103–128 (2018).Article 

    Google Scholar 
    73.Veselý, P., Bureš, P. & Šmarda, P. Nutrient reserves may allow for genome size increase: Evidence from comparison of geophytes and their sister non-geophytic relatives. Ann. Bot. 112, 1193–1200 (2013).PubMed 
    PubMed Central 
    Article 
    CAS 

    Google Scholar 
    74.African Plant Database. http://www.ville-ge.ch/musinfo/bd/cjb/africa/index. (2021).75.Hyde, M. A., Wursten, B. T., Ballings, P. & Coates Palgrave, M. Flora of Botswana. https://www.botswanaflora.com (2021).76.Hyde, M. A., Wursten, B. T., Ballings, P. & Coates Palgrave, M. Flora of Malawi. http://www.malawiflora.com (2021).77.Hyde, M. A., Wursten, B. T., Ballings, P. & Coates Palgrave, M. Flora of Mozambique. http://www.mozambiqueflora.com (2021)78.Bingham, M. G., Willemen, A., Wursten, B. T., Ballings, P. & Hyde, M. A. Flora of Zambia http://www.zambiaflora.com (2021).79.Hyde, M. A., Wursten, B. T., Ballings, P. & Coates Palgrave, M. Flora of Zimbabwe. http://www.zimbabweflora.co.zw (2021).80.International Legume Database & Information Service. https://ildis.org/LegumeWeb (2020).81.Exell, A.W. & Fernandes, A. Conspectus florae angolensis. Vol. 3, No. 2. Leguminosae (Papilionoideae: Hedysareae-Sophoreae) (Junta de Investigações do Ultramar, 1966)82.Pasquet, R. S. Notes on the genus Vigna (Leguminosae-Papilionoideae). Kew Bull 56, 223–227 (2001).Article 

    Google Scholar 
    83.van Zonneveld, M. et al. Mapping patterns of abiotic and biotic stress resilience uncovers conservation gaps and breeding potential of Vigna wild relatives. Sci. Rep. 10, 2111. https://doi.org/10.1038/s41598-020-58646-8 (2020).CAS 
    Article 
    PubMed 
    PubMed Central 
    ADS 

    Google Scholar 
    84.Global Biodiversity Information Facility. https://www.gbif.org/ (2021).85.GBIF Occurrence Download—Vigna. https://doi.org/10.15468/dl.bsjsk5 (2021).86.GBIF Occurrence Download—Phaseolus. https://doi.org/10.15468/dl.kjw72 (2021).87.QGIS Development Team. QGIS Geographic Information System. Open Source Geospatial Foundation Project. http://qgis.osgeo.org (2021).88.Doležel, J., Sgorbati, S. & Lucretti, S. Comparison of three DNA fluorochromes for flow cytometric estimation of nuclear DNA content in plants. Physiol. Plant. 85, 625–631 (1992).Article 

    Google Scholar 
    89.Loureiro, J., Rodriguez, E., Doležel, J. & Santos, C. Two new nuclear isolation buffers for plant DNA flow cytometry: A test with 37 species. Ann. Bot. 100, 875–888 (2007).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    90.Doležel, J. & Bartoš, J. Plant DNA flow cytometry and estimation of nuclear genome size. Ann. Bot. 95, 99–110 (2005).PubMed 
    PubMed Central 
    Article 
    CAS 

    Google Scholar 
    91.Doležel, J., Bartoš, J., Voglmayr, H. & Greilhuber, J. Nuclear DNA content and genome size of trout and human. Cytometry 51, 127–128 (2003).PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    92.Jelihovschi, E. G., Faria, J. C. & Allaman, I. B. ScottKnott: A package for performing the Scott-Knott clustering algorithm in R. TEMA 15, 3–17 (2014).MathSciNet 
    Article 

    Google Scholar 
    93.Wickham, H. ggplot2: Elegant Graphics for Data Analysis (Springer, 2016).MATH 
    Book 

    Google Scholar 
    94.R Core Team. R: A language and environment for statistical computing https://www.R-project.org/ (R Foundation for Statistical Computing, 2020). More

  • in

    Oil palm cultivation can be expanded while sparing biodiversity in India

    1.Vijay, V., Pimm, S. L., Jenkins, C. N. & Smith, S. J. The impacts of oil palm on recent deforestation and biodiversity loss. PLoS One 11, pe0159668 (2016).Article 

    Google Scholar 
    2.Rulli, M. C. et al. Interdependencies and telecoupling of oil palm expansion at the expense of Indonesian rainforest. Renew. Sustain. Energy Rev. 105, 499–512 (2019).Article 

    Google Scholar 
    3.Davis, K. F. et al. Tropical forest loss enhanced by large-scale land acquisitions. Nat. Geosci. 13, 482–488 (2020).ADS 
    CAS 
    Article 

    Google Scholar 
    4.Strona, G. et al. Small room for compromise between oil palm cultivation and primate conservation in Africa. Proc. Natl Acad. Sci. USA 115, 8811–8816 (2018).CAS 
    Article 

    Google Scholar 
    5.United States Department of Agriculture, Foreign Agricultural Service. Data retrieved from: https://apps.fas.usda.gov/psdonline/app/index.html#/app/advQuery (2020).6.Sagar, H. S. et al. India in the oil palm era: describing India’s dependence on palm oil, recommendations for sustainable production, and opportunities to become an influential consumer. Trop. Conserv. Sci. 12, 1940082919838918 (2019).Article 

    Google Scholar 
    7.Jadhav, R. Exclusive: India urges boycott of Malaysian palm oil after diplomatic row—sources. Reuters (13 January 2020).8.Srinivasan, U. Oil palm should not be expanded in Arunachal Pradesh. Arunachal Times (October 2016).9.Ministry of Agriculture and Farmers’ Welfare. National Mission on Oilseeds and Oil Palm; https://nmoop.gov.in (Government of India, 2020).10.Bose, P. Oil palm plantations vs shifting cultivation for indigenous peoples: analyzing Mizoram’s New Land Use Policy. Land Use Policy 81, 115–123 (2019).Article 

    Google Scholar 
    11.Dhar, A. Enter oil palm in northeast India: centre, Patanjali, Godrej bet big. The Citizen (16 September 2020).12.Raman, T. R. S. R. Is oil palm expansion good for Mizoram? The Frontier Despatch 3, 6–7 (2016).
    Google Scholar 
    13.Khandekar, N. Expanding oil palm plantations in the northeast could extract a long-term cost. The Wire (4 August 2020).14.Mandal, J. & Raman, T. R. S. R. Shifting agriculture supports more tropical forest birds than oil palm or teak plantations in Mizoram, northeast India. The Condor 118, 345–359 (2016).Article 

    Google Scholar 
    15.Nandi, J. Oil palm push on the northeast may impact biodiversity, water table, say experts. Hindustan Times 10, 51 (2020).
    Google Scholar 
    16.Myers, N., Mittermeier, R. A., Mittermeier, C. G., Da Fonseca, G. A. B. & Kent, J. Biodiversity hotspots for conservation priorities. Nature 403, 853–858 (2000).ADS 
    CAS 
    Article 

    Google Scholar 
    17.Global Agro-Ecological Zones, GAEZ v.3.0 (Food and Agriculture Organization, 2016); https://gaez.fao.org/pages/data-viewer18.Corley, R. H. V. How much palm oil do we need? Environ. Sci. Policy 12, 134–139 (2009).CAS 
    Article 

    Google Scholar 
    19.Meijaard, E. et al. The environmental impacts of palm oil in context. Nat. Plants 6, 1418–1426 (2020).Article 

    Google Scholar 
    20.West, P. C. et al. Leverage points for improving global food security and the environment. Science 18, 325–328 (2014).ADS 
    Article 

    Google Scholar 
    21.Fan, Y., Li, H. & Miguez-Macho, G. Global patterns of groundwater table depth. Science 339, 940–943 (2013).22.Shaktivadivel, R. The Agricultural Groundwater Revolution: Opportunities and Threats to Development (CAB International, 2007).
    Google Scholar 
    23.Lee, J. S. H., Miteva, D. A., Carlson, K. M., Heilmayr, R. & Saif, O. Does oil palm certification create trade-offs between environment and development in Indonesia? Env. Res. Lett. 15, 124064 (2020).Article 

    Google Scholar 
    24.Sankar, K. N. M. Oil palm finds favour with East Godavari farmers. The Hindu (25 January 2017).25.Curry, G. N. & Koczberski, G. Finding common ground: relational concepts of land tenure and economy in the oil palm frontier of Papua New Guinea. Geogr. J. 175, 98–111 (2009).Article 

    Google Scholar 
    26.DeVos, R., Kohne, M. & Roth, D. We’ll turn your water in Coca Cola: the atomising practices of oil palm development in Indonesia. J. Agrar. Change 1, 385–405 (2018).Article 

    Google Scholar 
    27.IPCC. Climate Change: Synthesis Report (eds Core Writing Team, Pachauri, R. K. & Meyer, L. A.) (IPCC, 2014).28.IPCC. IPCC Special Reports on Emissions Scenarios: Summary for Policymakers (IPCC, 2000).29.Schwalm, C. R., Glendon, S. & Duffy, P. B. RCP8. 5 tracks cumulative CO2 emissions. Proc. Natl Acad. Sci. USA 18, 19656–19657 (2020).ADS 
    Article 

    Google Scholar 
    30.Copernicus Land Monitoring Service (European Environment Agency, 2020).31.Hoffman, M., Koenig, K., Bunting, G., Cosntanza, J. & Willams, K. J. Biodiversity Hotspots v.2016.1 (2016); https://doi.org/10.5281/zenodo.326180632.IUCN World Database on Protected Areas, online April 2017 (UNEP-WCMC, 2016); www.protectedplanet.net33.QGIS Development Team. QGIS Geographic Information System (Open Source Geospatial Foundation, 2021); http://qgis.osgeo.org34.R Core Team. R: A Language and Environment for Statistical Computing (R Foundation for Statistical Computing, 2021). More