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    Dimethyl sulfide mediates microbial predator–prey interactions between zooplankton and algae in the ocean

    1.Simó, R. Production of atmospheric sulfur by oceanic plankton: biogeochemical, ecological and evolutionary links. Trends Ecol. Evol. 16, 287–294 (2001).PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    2.Charlson, R. J., Lovelock, J. E., Andreae, M. O. & Warren, S. G. Oceanic phytoplankton, atmospheric sulphur, cloud albedo and climate. Nature 326, 655–661 (1987).CAS 
    Article 

    Google Scholar 
    3.Wang, S., Maltrud, M. E., Burrows, S. M., Elliott, S. M. & Cameron-Smith, P. Impacts of shifts in phytoplankton community on clouds and climate via the sulfur cycle. Glob. Biogeochem. Cycles 32, 1005–1026 (2018).Article 
    CAS 

    Google Scholar 
    4.Wolfe, G. V., Steinke, M. & Kirst, G. O. Grazing-activated chemical defence in a unicellular marine alga. Nature 387, 894–897 (1997).CAS 
    Article 

    Google Scholar 
    5.Seymour, J., Simó, R., Ahmed, T. & Stocker, R. Chemoattraction to dimethylsulfoniopropionate throughout the marine microbial food web. Science 329, 342–345 (2010).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    6.Alcolombri, U. et al. Identification of the algal dimethyl sulfide-releasing enzyme: a missing link in the marine sulfur cycle. Science 348, 1466–1469 (2015).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    7.Alcolombri, U., Lei, L., Meltzer, D., Vardi, A. & Tawfik, D. S. Assigning the algal source of dimethylsulfide using a selective lyase inhibitor. ACS Chem. Biol. 12, 41–46 (2017).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    8.Kettle, A. J. & Andreae, M. O. Flux of dimethylsulfide from the oceans: a comparison of updated data sets and flux models. J. Geophys. Res. Atmos. 105, 26793–26808 (2000).CAS 
    Article 

    Google Scholar 
    9.Carpenter, L. J., Archer, S. D. & Beale, R. Ocean–atmosphere trace gas exchange. Chem. Soc. Rev. 41, 6473–6506 (2012).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    10.Franklin, D. J., Steinke, M., Young, J., Probert, I. & Malin, G. Dimethylsulphoniopropionate (DMSP), DMSP-lyase activity (DLA) and dimethylsulphide (DMS) in 10 species of coccolithophore. Mar. Ecol. Prog. Ser. 410, 13–23 (2010).CAS 
    Article 

    Google Scholar 
    11.Keller, M. D. Dimethyl sulfide production and marine phytoplankton: the importance of species composition and cell size. Biol. Oceanogr. 6, 375–382 (1989).
    Google Scholar 
    12.Curson, A. R. J. et al. DSYB catalyses the key step of dimethylsulfoniopropionate biosynthesis in many phytoplankton. Nat. Microbiol. 3, 430–439 (2018).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    13.Sunda, W., Kieber, D. J., Kiene, R. P. & Huntsman, S. An antioxidant function for DMSP and DMS in marine algae. Nature 418, 317–320 (2002).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    14.Kirst, G. O. in Biological and Environmental Chemistry of DMSP and Related Sulfonium Compounds (eds Kiene, R. P. et al.) 121−129 (Springer, 1996).15.Darroch, L. et al. Effect of short-term light- and UV-stress on DMSP, DMS, and DMSP lyase activity in Emiliania huxleyi. Aquat. Microb. Ecol. 74, 173–185 (2015).16.Barak-Gavish, N. et al. Bacterial virulence against an oceanic bloom-forming phytoplankter is mediated by algal DMSP. Sci. Adv. 4, eaau5716 (2018).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    17.Amin, S. A. et al. Interaction and signalling between a cosmopolitan phytoplankton and associated bacteria. Nature 522, 98–101 (2015).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    18.Garcés, E., Alacid, E., Reñé, A., Petrou, K. & Simó, R. Host-released dimethylsulphide activates the dinoflagellate parasitoid Parvilucifera sinerae. ISME J. 7, 1065–1068 (2013).PubMed 
    PubMed Central 
    Article 
    CAS 

    Google Scholar 
    19.Steinke, M., Stefels, J. & Stamhuis, E. Dimethyl sulfide triggers search behavior in copepods. Limnol. Oceanogr. 51, 1925–1930 (2006).CAS 
    Article 

    Google Scholar 
    20.Breckels, M., Bode, N., Codling, E. & Steinke, M. Effect of grazing-mediated dimethyl sulfide (DMS) production on the swimming behavior of the copepod Calanus helgolandicus. Mar. Drugs 11, 2486 (2013).PubMed 
    PubMed Central 
    Article 
    CAS 

    Google Scholar 
    21.Procter, J., Hopkins, F. E., Fileman, E. S. & Lindeque, P. K. Smells good enough to eat: dimethyl sulfide (DMS) enhances copepod ingestion of microplastics. Mar. Pollut. Bull. 138, 1–6 (2019).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    22.Foretich, M. A., Paris, C. B., Grosell, M., Stieglitz, J. D. & Benetti, D. D. Dimethyl sulfide is a chemical attractant for reef fish larvae. Sci. Rep. 7, 2498 (2017).PubMed 
    PubMed Central 
    Article 
    CAS 

    Google Scholar 
    23.Savoca, M. S. & Nevitt, G. A. Evidence that dimethyl sulfide facilitates a tritrophic mutualism between marine primary producers and top predators. Proc. Natl Acad. Sci. USA 111, 4157–4161 (2014).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    24.Wright, K. L. B., Pichegru, L. & Ryan, P. G. Penguins are attracted to dimethyl sulphide at sea. J. Exp. Biol. 214, 2509–2511 (2011).PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    25.Owen, K. et al. Natural dimethyl sulfide gradients would lead marine predators to higher prey biomass. Commun. Biol. 4, 149 (2021).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    26.Wolfe, G. V. & Steinke, M. Grazing-activated production of dimethyl sulfide (DMS) by two clones of Emiliania huxleyi. Limnol. Oceanogr. 41, 1151–1160 (1996).CAS 
    Article 

    Google Scholar 
    27.Simó, R. et al. The quantitative role of microzooplankton grazing in dimethylsulfide (DMS) production in the NW Mediterranean. Biogeochemistry 141, 125–142 (2018).Article 

    Google Scholar 
    28.Evans, C., Kadner, S. V. & Darroch, L. J. The relative significance of viral lysis and microzooplankton grazing as pathways of dimethylsulfoniopropionate (DMSP) cleavage: an Emiliania huxleyi culture study. Limnol. Oceanogr. 52, 1036–1045 (2007).Article 

    Google Scholar 
    29.Kiene, R. P. Dimethyl sulfide production from dimethylsulfoniopropionate in coastal seawater samples and bacterial cultures. Appl. Environ. Microbiol. 56, 3292–3297 (1990).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    30.Bullock, H. A., Luo, H. & Whitman, W. B. Evolution of dimethylsulfoniopropionate metabolism in marine phytoplankton and bacteria. Front. Microbiol. https://doi.org/10.3389/fmicb.2017.00637 (2017).31.Strom, S. et al. Chemical defense in the microplankton I: feeding and growth rates of heterotrophic protists on the DMS-producing phytoplankter Emiliania huxleyi. Limnol. Oceanogr. 48, 217–229 (2003).CAS 
    Article 

    Google Scholar 
    32.Calbet, A. & Landry, M. R. Phytoplankton growth, microzooplankton grazing, and carbon cycling in marine systems. Limnol. Oceanogr. 49, 51–57 (2004).CAS 
    Article 

    Google Scholar 
    33.Schmoker, C., Hernández-León, S. & Calbet, A. Microzooplankton grazing in the oceans: impacts, data variability, knowledge gaps and future directions. J. Plankton Res. 35, 691–706 (2013).Article 

    Google Scholar 
    34.Steinke, M., Wolfe, G. V. & Kirst, G. O. Partial characterisation of dimethylsulfoniopropionate (DMSP) lyase isozymes in 6 strains of Emiliania huxleyi. Mar. Ecol. 175, 215–225 (1998).CAS 
    Article 

    Google Scholar 
    35.Breckels, M. N., Roberts, E. C., Archer, S. D., Malin, G. & Steinke, M. The role of dissolved infochemicals in mediating predator–prey interactions in the heterotrophic dinoflagellate Oxyrrhis marina. J. Plankton Res. 33, 629–639 (2011).Article 

    Google Scholar 
    36.Saló, V., Simó, R., Vila-Costa, M. & Calbet, A. Sulfur assimilation by Oxyrrhis marina feeding on a 35S-DMSP-labelled prey. Environ. Microbiol. 11, 3063–3072 (2009).PubMed 
    Article 
    CAS 
    PubMed Central 

    Google Scholar 
    37.Raina, J. B. et al. Subcellular tracking reveals the location of dimethylsulfoniopropionate in microalgae and visualises its uptake by marine bacteria. eLife 6, e23008 (2017).PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    38.Franklin, D. J. et al. Identification of senescence and death in Emiliania huxleyi and Thalassiosira pseudonana: cell staining, chlorophyll alterations, and dimethylsulfoniopropionate (DMSP) metabolism. Limnol. Oceanogr. 57, 305–317 (2012).CAS 
    Article 

    Google Scholar 
    39.Kettles, N. L., Kopriva, S. & Malin, G. Insights into the regulation of DMSP synthesis in the diatom Thalassiosira pseudonana through APR activity, proteomics and gene expression analyses on cells acclimating to changes in salinity, light and nitrogen. PLoS ONE 9, e94795 (2014).PubMed 
    PubMed Central 
    Article 
    CAS 

    Google Scholar 
    40.Poulsen, N., Chesley, P. M. & Kröger, N. Molecular genetic manipulation of the diatom Thalassiosira pseudonana (bacillariophyceae). J. Phycol. 42, 1059–1065 (2006).Article 

    Google Scholar 
    41.Armbrust, E. V. et al. The genome of the diatom Thalassiosira pseudonana: ecology, evolution, and metabolism. Science 306, 79–86 (2004).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    42.Malviya, S. et al. Insights into global diatom distribution and diversity in the world’s ocean. Proc. Natl Acad. Sci. USA 113, E1516–E1525 (2016).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    43.Apt, K. E. et al. In vivo characterization of diatom multipartite plastid targeting signals. J. Cell Sci. 115, 4061–4069 (2002).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    44.McParland, E. L., Wright, A., Art, K., He, M. & Levine, N. M. Evidence for contrasting roles of dimethylsulfoniopropionate production in Emiliania huxleyi and Thalassiosira oceanica. New Phytol. 226, 396–409 (2020).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    45.Keeling, P. J. et al. The Marine Microbial Eukaryote Transcriptome Sequencing Project (MMETSP): illuminating the functional diversity of eukaryotic life in the oceans through transcriptome sequencing. PLoS Biol. 12, e1001889 (2014).PubMed 
    PubMed Central 
    Article 
    CAS 

    Google Scholar 
    46.Olson, M. B. & Strom, S. L. Phytoplankton growth, microzooplankton herbivory and community structure in the southeast Bering Sea: insight into the formation and temporal persistence of an Emiliania huxleyi bloom. Deep-Sea Res. II 49, 5969–5990 (2002).CAS 
    Article 

    Google Scholar 
    47.Challenger, F. & Simpson, M. I. Studies on biological methylation; a precursor of the dimethyl sulphide evolved by Polysiphonia fastigiata; dimethyl-2-carboxyethylsulphonium hydroxide and its salts. J. Chem. Soc. 3, 1591–1597 (1948).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    48.Haas, P. The liberation of methyl sulphide by seaweed. Biochem. J. 29, 1297–1299 (1935).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    49.Stefels, J. & Dijkhuizen, L. Characteristics of DMSP-lyase in Phaeocystis sp. (Prymnesiophyceae). Mar. Ecol. 131, 307–313 (1996).CAS 
    Article 

    Google Scholar 
    50.Wolfe, G. V., Sherr, E. B. & Sherr, B. F. Release and consumption of DMSP from Emiliania huxleyi during grazing by Oxyrrhis marina. Mar. Ecol. 111, 111–119 (1994).CAS 
    Article 

    Google Scholar 
    51.Reisch, C. R., Moran, M. A. & Whitman, W. B. Bacterial catabolism of dimethylsulfoniopropionate (DMSP). Front. Microbiol. 2, 172 (2011).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    52.von Dassow, P. et al. Transcriptome analysis of functional differentiation between haploid and diploid cells of Emiliania huxleyi, a globally significant photosynthetic calcifying cell. Genome Biol. 10, R114 (2009).Article 
    CAS 

    Google Scholar 
    53.Strom, S., Wolfe, G., Slajer, A., Lambert, S. & Clough, J. Chemical defense in the microplankton II: inhibition of protist feeding by β-dimethylsulfoniopropionate (DMSP). Limnol. Oceanogr. 48, 230–237 (2003).CAS 
    Article 

    Google Scholar 
    54.Li, W. Eat-me signals: keys to molecular phagocyte biology and “appetite” control. J. Cell. Physiol. 227, 1291–1297 (2012).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    55.Tyssebotn, I. M. B. et al. Concentrations, biological uptake, and respiration of dissolved acrylate and dimethylsulfoxide in the northern Gulf of Mexico. Limnol. Oceanogr. 62, 1198–1218 (2017).Article 

    Google Scholar 
    56.Curson, A. R. J., Todd, J. D., Sullivan, M. J. & Johnston, A. W. B. Catabolism of dimethylsulphoniopropionate: microorganisms, enzymes and genes. Nat. Rev. Microbiol. 9, 849–859 (2011).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    57.Spiese, C. E., Le, T., Zimmer, R. L. & Kieber, D. J. Dimethylsulfide membrane permeability, cellular concentrations and implications for physiological functions in marine algae. J. Plankton Res. 38, 41–54 (2015).Article 
    CAS 

    Google Scholar 
    58.Hatton, A. D., Shenoy, D. M., Hart, M. C., Mogg, A. & Green, D. H. Metabolism of DMSP, DMS and DMSO by the cultivable bacterial community associated with the DMSP-producing dinoflagellate Scrippsiella trochoidea. Biogeochemistry 110, 131–146 (2012).CAS 
    Article 

    Google Scholar 
    59.Laber, C. P. et al. Coccolithovirus facilitation of carbon export in the North Atlantic. Nat. Microbiol. 3, 537–547 (2018).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    60.Endres, C. S. & Lohmann, K. J. Perception of dimethyl sulfide (DMS) by loggerhead sea turtles: a possible mechanism for locating high-productivity oceanic regions for foraging. J. Exp. Biol. 215, 3535–3538 (2012).PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    61.Savoca, M. S. Chemoattraction to dimethyl sulfide links the sulfur, iron, and carbon cycles in high-latitude oceans. Biogeochemistry 138, 1–21 (2018).CAS 
    Article 

    Google Scholar 
    62.Steinke, M., Malin, G. & Liss, P. Trophic interactions in the sea: an ecological role for climate relevant volatiles? J. Phycol. 38, 630–638 (2002).CAS 
    Article 

    Google Scholar 
    63.Pohnert, G., Steinke, M. & Tollrian, R. Chemical cues, defence metabolites and the shaping of pelagic interspecific interactions. Trends Ecol. Evol. 22, 198–204 (2007).PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    64.Lewis, N. et al. Grazing-induced production of DMS can stabilize food-web dynamics and promote the formation of phytoplankton blooms in a multitrophic plankton model. Biogeochemistry 110, 303–313 (2012).CAS 
    Article 

    Google Scholar 
    65.Lewis, N. D., Breckels, M. N., Steinke, M. & Codling, E. A. Role of infochemical mediated zooplankton grazing in a phytoplankton competition model. Ecol. Complex. 16, 41–50 (2013).Article 

    Google Scholar 
    66.Hansen, F. C., Reckermann, M., Breteler, W. C. M. K. & Riegman, R. Phaeocystis blooming enhanced by copepod predation on protozoa: evidence from incubation experiments. Mar. Ecol. Prog. Ser. 102, 51–57 (1993).Article 

    Google Scholar 
    67.Levasseur, M. et al. Production of DMSP and DMS during a mesocosm study of an Emiliania huxleyi bloom: influence of bacteria and Calanus finmarchicus grazing. Mar. Biol. 126, 609–618 (1996).CAS 
    Article 

    Google Scholar 
    68.Guillard, R. R. & Ryther, J. H. Studies of marine planktonic diatoms. I. Cyclotella nana Hustedt, and Detonula confervacea (cleve) Gran. Can. J. Microbiol. 8, 229–239 (1962).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    69.Parada, A. E., Needham, D. M. & Fuhrman, J. A. Every base matters: assessing small subunit rRNA primers for marine microbiomes with mock communities, time series and global field samples. Environ. Microbiol. 18, 1403–1414 (2016).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    70.Apprill, A., McNally, S., Parsons, R. & Weber, L. Minor revision to V4 region SSU rRNA 806R gene primer greatly increases detection of SAR11 bacterioplankton. Aquat. Microb. Ecol. https://doi.org/10.3354/ame01753 (2015).71.Callahan, B. J. et al. DADA2: high-resolution sample inference from Illumina amplicon data. Nat. Methods 13, 581–583 (2016).CAS 
    PubMed 
    PubMed Central 
    Article 

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

    Google Scholar 
    73.McMurdie, P. J. & Holmes, S. phyloseq: an R package for reproducible interactive analysis and graphics of microbiome census data. PLoS ONE 8, e61217 (2013).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    74.Frost, B. W. Effects of size and concentration of food particles on the feeding and behavior of the marine planktonic copepod Calanus pacificus. Limnol. Oceanogr. 17, 805–815 (1972).Article 

    Google Scholar 
    75.Johnson, M. D., Michelle, R. & Stoecker, D. K. Microzooplankton grazing on Prorocentrum minimum and Karlodinium micrum in Chesapeake Bay. Limnol. Oceanogr. 48, 238–248 (2003).Article 

    Google Scholar 
    76.Stoeck, T. et al. Multiple marker parallel tag environmental DNA sequencing reveals a highly complex eukaryotic community in marine anoxic water. Mol. Ecol. 19, 21–31 (2010).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    77.Piredda, R. et al. Diversity and temporal patterns of planktonic protist assemblages at a Mediterranean Long Term Ecological Research site. FEMS Microbiol. Ecol. https://doi.org/10.1093/femsec/fiw200 (2017).78.Guillou, 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. 41, D597–D604 (2013).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    79.Slamovits, C. H., Saldarriaga, J. F., Larocque, A. & Keeling, P. J. The highly reduced and fragmented mitochondrial genome of the early-branching dinoflagellate Oxyrrhis marina shares characteristics with both apicomplexan and dinoflagellate mitochondrial genomes. J. Mol. Biol. 372, 356–368 (2007).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    80.Untergasser, A. et al. Primer3Plus, an enhanced web interface to Primer3. Nucleic Acids Res. 35, W71–W74 (2007).PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    81.Dagg, M. J., Jackson, G. A. & Checkley, D. M. The distribution and vertical flux of fecal pellets from large zooplankton in Monterey Bay and coastal California. Deep-Sea Res. I 94, 72–86 (2014).Article 

    Google Scholar  More

  • in

    Heterotrophic bacterial diazotrophs are more abundant than their cyanobacterial counterparts in metagenomes covering most of the sunlit ocean

    1.Boyd PW. Toward quantifying the response of the oceans’ biological pump to climate change. Front Mar Sci. 2015. https://doi.org/10.3389/fmars.2015.00077.2.Charlson RJ, Lovelock JE, Andreae MO, Warren SG. Oceanic phytoplankton, atmospheric sulphur, cloud albedo and climate. Nature. 1987;326:655–61.CAS 
    Article 

    Google Scholar 
    3.Falkowski PG, Barber RT, Smetacek V. Biogeochemical controls and feedbacks on ocean primary production. Science (80-). 1998;281:200–6.CAS 
    Article 

    Google Scholar 
    4.Arrigo KR. Marine microorganisms and global nutrient cycles. Nature. 2005;437:349–55.CAS 
    PubMed 
    Article 

    Google Scholar 
    5.Sanders R, Henson SA, Koski M, De La Rocha CL, Painter SC, Poulton AJ, et al. The biological carbon pump in the North Atlantic. Prog Oceanogr e-pub print. 2014. https://doi.org/10.1016/j.pocean.2014.05.005.Article 

    Google Scholar 
    6.De Vargas C, Audic S, Henry N, Decelle J, Mahé F, Logares R, et al. Eukaryotic plankton diversity in the sunlit ocean. Science. 2015. https://doi.org/10.1126/science.1261605.7.Moore CM, Mills MM, Arrigo KR, Berman-Frank I, Bopp L, Boyd PW, et al. Processes and patterns of oceanic nutrient limitation. Nat Geosci. 2013;6:701–10.CAS 
    Article 

    Google Scholar 
    8.Tyrrell T. The relative influences of nitrogen and phosohorus on oceanic primary production. Nature. 1999;400:525–31.CAS 
    Article 

    Google Scholar 
    9.Dos Santos PC, Fang Z, Mason SW, Setubal JC, Dixon R. Distribution of nitrogen fixation and nitrogenase-like sequences amongst microbial genomes. BMC Genomics. 2012;13:162.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    10.Zehr JP, Capone DG. Changing perspectives in marine nitrogen fixation. Science. 2020. https://doi.org/10.1126/science.aay9514.11.Zehr JP, Jenkins BD, Short SM, Steward GF. Nitrogenase gene diversity and microbial community structure: a cross-system comparison. Env Microbiol. 2003;5:539–54.CAS 
    Article 

    Google Scholar 
    12.Galloway JN, Dentener FJ, Capone DG, Boyer EW, Howarth RW, Seitzinger SP, et al. Nitrogen cycles: Past, present, and future. Biogeochemistry. 2004. https://doi.org/10.1007/s10533-004-0370-0.13.Carpenter EJ, Capone DG, Rueter JG. Marine pelagic cyanobacteria: trichodesmium and other diazotrophs. Boston: Kluwer Academic Publishers; 1992.14.Carpenter EJ, Romans K. Major role of the cyanobacterium trichodesmium in nutrient cycling in the north atlantic ocean. Science. 1991;254:1356–8.CAS 
    PubMed 
    Article 

    Google Scholar 
    15.Karl D, Letelier R, Tupas L, Dore J, Christian J, Hebel D. The role of nitrogen fixation in biogeochemical cycling in the subtropical North Pacific Ocean. Nature. 1997;388:533–8.CAS 
    Article 

    Google Scholar 
    16.Capone DG. Trichodesmium, a globally significant marine cyanobacterium. Science (80-). 1997;276:1221–9.CAS 
    Article 

    Google Scholar 
    17.Dyhrman ST, Chappell PD, Haley ST, Moffett JW, Orchard ED, Waterbury JB, et al. Phosphonate utilization by the globally important marine diazotroph. Trichodesmium Nat. 2006;439:68–71.CAS 
    Article 

    Google Scholar 
    18.Pierella Karlusich JJ, Pelletier E, Lombard F, Carsique M, Dvorak E, Colin S, et al. Global distribution patterns of marine nitrogen-fixers by imaging and molecular methods. Nat Commun 2021 121. 2021;12:1–18.
    Google Scholar 
    19.Gómez F, Furuya K, Takeda S. Distribution of the cyanobacterium Richelia intracellularis as an epiphyte of the diatom Chaetoceros compressus in the western Pacific Ocean. J Plankton Res. 2005. https://doi.org/10.1093/plankt/fbi007.20.Hilton JA, Foster RA, James Tripp H, Carter BJ, Zehr JP, Villareal TA. Genomic deletions disrupt nitrogen metabolism pathways of a cyanobacterial diatom symbiont. Nat Commun. 2013. https://doi.org/10.1038/ncomms2748.21.Martínez-Pérez C, Mohr W, Löscher CR, Dekaezemacker J, Littmann S, Yilmaz P, et al. The small unicellular diazotrophic symbiont, UCYN-A, is a key player in the marine nitrogen cycle. Nat Microbiol 2016. https://doi.org/10.1038/nmicrobiol.2016.163.Article 
    PubMed 

    Google Scholar 
    22.Tripp HJ, Bench SR, Turk KA, Foster RA, Desany BA, Niazi F, et al. Metabolic streamlining in an open-ocean nitrogen-fixing cyanobacterium. Nature. 2010;464:90–94.CAS 
    PubMed 
    Article 

    Google Scholar 
    23.Moisander PH, Beinart RA, Hewson I, White AE, Johnson KS, Carlson CA, et al. (2010). Unicellular cyanobacterial distributions broaden the oceanic N2 fixation domain. Science (80-). https://doi.org/10.1126/science.1185468.24.Montoya JP, Holl CM, Zehr JP, Hansen A, Villareal TA, Capone DG (2004). High rates of N2 fixation by unicellular diazotrophs in the oligotrophic Pacific Ocean. Nature. https://doi.org/10.1038/nature02824.25.Church MJ, Short CM, Jenkins BD, Karl DM, Zehr JP. Temporal patterns of nitrogenase gene (nifH) expression in the oligotrophic North Pacific Ocean. Appl Environ Microbiol. 2005;71:5362–70.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    26.Church MJ, Björkman KM, Karl DM, Saito MA, Zehr JP. Regional distributions of nitrogen-fixing bacteria in the Pacific Ocean. Limnol Oceanogr. 2008;53:63–77.CAS 
    Article 

    Google Scholar 
    27.Zehr JP, Montoya JP, Jenkins BD, Hewson I, Mondragon E, Short CM, et al. Experiments linking nitrogenase gene expression to nitrogen fixation in the North Pacific subtropical gyre. Limnology and Oceanography. 2007;52:169–83.CAS 
    Article 

    Google Scholar 
    28.Fong AA, Karl DM, Lukas R, Letelier RM, Zehr JP, Church MJ. Nitrogen fixation in an anticyclonic eddy in the oligotrophic North Pacific Ocean. ISME J. 2008;2:663–76.CAS 
    PubMed 
    Article 

    Google Scholar 
    29.Moisander PH, Beinart RA, Voss M, Zehr JP. Diversity and abundance of diazotrophic microorganisms in the South China Sea during intermonsoon. ISME J. 2008;251:954–67.Article 
    CAS 

    Google Scholar 
    30.Man-Aharonovich D, Kress N, Zeev EB, Berman-Frank I, Béjà O. Molecular ecology of nifH genes and transcripts in the eastern Mediterranean Sea. Environ Microbiol. 2007;9:2354–63.CAS 
    PubMed 
    Article 

    Google Scholar 
    31.Benavides M, Moisander PH, Daley MC, Bode A, Arístegui J (2016). Longitudinal variability of diazotroph abundances in the subtropical North Atlantic Ocean. J Plankton Res. https://doi.org/10.1093/plankt/fbv121.32.Langlois RJ, LaRoche J, Raab PA (2005). Diazotrophic diversity and distribution in the tropical and subtropical Atlantic Ocean. Appl Environ Microbiol. https://doi.org/10.1128/AEM.71.12.7910-7919.2005.33.Bombar D, Paerl RW, Riemann L. Marine non-cyanobacterial diazotrophs: moving beyond molecular detection. Trends Microbiol. 2016;24:916–27.CAS 
    PubMed 
    Article 

    Google Scholar 
    34.Farnelid H, Andersson AF, Bertilsson S, Al-Soud WA, Hansen LH, Sørensen S, et al. Nitrogenase gene amplicons from global marine surface waters are dominated by genes of non-cyanobacteria. PLoS One 6. 2011. https://doi.org/10.1371/journal.pone.0019223.35.Riemann L, Farnelid H, Steward GF. Nitrogenase genes in non-cyanobacterial plankton: prevalence, diversity and regulation in marine waters. Aquat Micro Ecol. 2010;61:235–47.Article 

    Google Scholar 
    36.Moisander PH, Benavides M, Bonnet S, Berman-Frank I, White AE, Riemann L. Chasing after non-cyanobacterial nitrogen fixation in marine pelagic environments. Front Microbiol. 2017. https://doi.org/10.3389/fmicb.2017.01736.37.Moreira-Coello V, Mouriño-Carballido B, Marañón E, Fernández-Carrera A, Bode A, Sintes E, et al. Temporal variability of diazotroph community composition in the upwelling region off NW Iberia. Sci Rep. 2019. https://doi.org/10.1038/s41598-019-39586-4.38.Luo YW, Doney SC, Anderson LA, Benavides M, Berman-Frank I, Bode A, et al. Database of diazotrophs in global ocean: abundance, biomass and nitrogen fixation rates. Earth Syst Sci Data. 2012. https://doi.org/10.5194/essd-4-47-2012.39.Delmont TO, Quince C, Shaiber A, Esen ÖC, Lee ST, Rappé MS, et al. Nitrogen-fixing populations of Planctomycetes and Proteobacteria are abundant in surface ocean metagenomes. Nat Microbiol. 2018;3:804–13.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    40.Salazar G, Paoli L, Alberti A, Huerta-Cepas J, Ruscheweyh HJ, Cuenca M, et al. Gene expression changes and community turnover differentially shape the global ocean metatranscriptome. Cell. 2019. https://doi.org/10.1016/j.cell.2019.10.014.41.Sunagawa S, Acinas SG, Bork P, Bowler C, Eveillard D, Gorsky G, et al. Tara Oceans: towards global ocean ecosystems biology. Nat Rev Microbiol. 2020;18:428–45CAS 
    PubMed 
    Article 

    Google Scholar 
    42.Delmont TO, Gaia M, Hinsinger DD, Fremont P, Fernandez Guerra A, Murat et al. Functional repertoire convergence of distantly related eukaryotic plankton lineages revealed by genome-resolved metagenomics. bioRxiv. 2020. 2020.10.15.341214.43.Eren AM, Esen ÖC, Quince C, Vineis JH, Morrison HG, Sogin ML, et al. Anvi’o: an advanced analysis and visualization platform for ‘omics data. PeerJ. 2015;3:e1319.PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    44.Eren AM, Kiefl E, Shaiber A, Veseli I, Miller SE, Schechter MS, et al. Community-led, integrated, reproducible multi-omics with anvi’o. Nat Microbiol 2020;6:3–6.Article 
    CAS 

    Google Scholar 
    45.Gaby JC, Buckley DH (2012). A comprehensive evaluation of PCR primers to amplify the nifH gene of nitrogenase. PLoS One. https://doi.org/10.1371/journal.pone.0042149.46.Turk-Kubo KA, Karamchandani M, Capone DG, Zehr JP. The paradox of marine heterotrophic nitrogen fixation: abundances of heterotrophic diazotrophs do not account for nitrogen fixation rates in the Eastern Tropical South Pacific. Environ Microbiol. 2014;16:3095–114.CAS 
    PubMed 
    Article 

    Google Scholar 
    47.Zehr JP, Turner PJ. Nitrogen fixation: nitrogenase genes and gene expression. METHODS Microbiol. 2001;30:271–86.CAS 
    Article 

    Google Scholar 
    48.Galperin MY, Wolf YI, Makarova KS, Vera Alvarez R, Landsman D, Koonin EV. COG database update: focus on microbial diversity, model organisms, and widespread pathogens. Nucleic Acids Res. 2021. https://doi.org/10.1093/nar/gkaa1018.49.Aramaki T, Blanc-Mathieu R, Endo H, Ohkubo K, Kanehisa M, Goto S, et al. KofamKOALA: KEGG Ortholog assignment based on profile HMM and adaptive score threshold. Bioinformatics. 2020. https://doi.org/10.1093/bioinformatics/btz859.50.Kanehisa M, Furumichi M, Tanabe M, Sato Y, Morishima K. KEGG: New perspectives on genomes, pathways, diseases and drugs. Nucleic Acids Res. 2017. https://doi.org/10.1093/nar/gkw1092.51.Pesant S, Not F, Picheral M, Kandels-Lewis S, Le Bescot N, Gorsky G, et al. Open science resources for the discovery and analysis of Tara Oceans data. Sci Data 2015 21. 2015;2:1–16.
    Google Scholar 
    52.Farnelid H, Tarangkoon W, Hansen G, Hansen PJ, Riemann L. Putative N2-fixing heterotrophic bacteria associated with dinoflagellate-cyanobacteria consortia in the low-nitrogen Indian Ocean. Aquat Microb Ecol. 2010. https://doi.org/10.3354/ame01440.53.Farnelid H, Turk-Kubo K, Ploug H, Ossolinski JE, Collins JR, Van Mooy BAS, et al. Diverse diazotrophs are present on sinking particles in the North Pacific Subtropical Gyre. ISME J. 2019. https://doi.org/10.1038/s41396-018-0259-x.54.Foster RA, Carpenter EJ, Bergman B. Unicellular cyanobionts in open ocean dinoflagellates, radiolarians, and tintinnids: ultrastructural characterization and immuno-localization of phycoerythrin and nitrogenase. J Phycol. 2006. https://doi.org/10.1111/j.1529-8817.2006.00206.x.55.Scavotto RE, Dziallas C, Bentzon-Tilia M, Riemann L, Moisander PH. Nitrogen-fixing bacteria associated with copepods in coastal waters of the North Atlantic Ocean. Environ Microbiol. 2015. https://doi.org/10.1111/1462-2920.12777.56.Zani S, Mellon MT, Collier JL, Zehr JP. Expression of nifH genes in natural microbial assemblages in Lake George, New York, detected by reverse transcriptase PCR. Appl Environ Microbiol. 2000;66:3119–24.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    57.Geisler E, Bogler A, Rahav E, Bar-Zeev E. Direct Detection of Heterotrophic Diazotrophs Associated with Planktonic Aggregates. Sci Rep. 2019. https://doi.org/10.1038/s41598-019-45505-4.58.Martínez-Pérez C, Mohr W, Schwedt A, Dürschlag J, Callbeck CM, Schunck H, et al. Metabolic versatility of a novel N2-fixing Alphaproteobacterium isolated from a marine oxygen minimum zone. Environ Microbiol. 2018. https://doi.org/10.1111/1462-2920.14008.59.Rahav E, Bar-Zeev E, Ohayon S, Elifantz H, Belkin N, Herut B, et al. Dinitrogen fixation in aphotic oxygenated marine environments. Front Microbiol. 2013. https://doi.org/10.3389/fmicb.2013.00227.60.Bentzon-Tilia M, Severin I, Hansen LH, Riemann L. Genomics and ecophysiology of heterotrophic nitrogen-fixing bacteria isolated from estuarine surface water. MBio 6. 2015. https://doi.org/10.1128/mBio.00929-15.61.Cornejo-Castillo FM, Zehr JP. Intriguing size distribution of the uncultured and globally widespread marine non-cyanobacterial diazotroph Gamma-A. ISME J. 2021. https://doi.org/10.1038/s41396-020-00765-1.62.Carradec Q, Pelletier E, Da Silva C, Alberti A, Seeleuthner Y, Blanc-Mathieu R, et al. A global ocean atlas of eukaryotic genes. Nat Commun. 2018. https://doi.org/10.1038/s41467-017-02342-1.63.Güell M, Yus E, Lluch-Senar M, Serrano L. Bacterial transcriptomics: what is beyond the RNA horiz-ome? Nat Rev Microbiol. 2011. https://doi.org/10.1038/nrmicro2620.64.Cornejo-Castillo FM, Cabello AM, Salazar G, Sánchez-Baracaldo P, Lima-Mendez G, Hingamp P, et al. Cyanobacterial symbionts diverged in the late Cretaceous towards lineage-specific nitrogen fixation factories in single-celled phytoplankton. Nat Commun. 2016. https://doi.org/10.1038/ncomms11071.65.Needoba JA, Foster RA, Sakamoto C, Zehr JP, Johnson KS. Nitrogen fixation by unicellular diazotrophic cyanobacteria in the temperate oligotrophic North Pacific Ocean. Limnol Oceanogr. 2007. https://doi.org/10.4319/lo.2007.52.4.1317.66.Foster RA, Paytan A, Zehr JP. Seasonality of N2 fixation and nifH gene diversity in the Gulf of Aqaba (Red Sea). Limnol Oceanogr. 2009. https://doi.org/10.4319/lo.2009.54.1.0219.67.Thompson AW, Foster RA, Krupke A, Carter BJ, Musat N, Vaulot D, et al. Unicellular cyanobacterium symbiotic with a single-celled eukaryotic alga. Science. 2012;337:1546–50.CAS 
    PubMed 
    Article 

    Google Scholar 
    68.Zehr JP, Waterbury JB, Turner PJ, Montoya JP, Omoregie E, Steward GF, et al. Unicellular cyanobacteria fix N2 in the subtropical North Pacific Ocean. Nature. 2001;412:635–8.CAS 
    PubMed 
    Article 

    Google Scholar 
    69.Ohki K, Zehr JP, Fujita Y. Trichodesmium: establishment of culture and characteristics of N2- fixation. Mar pelagic cyanobacteria. 1992. https://doi.org/10.1007/978-94-015-7977-3_20.70.Li D, Liu CM, Luo R, Sadakane K, Lam TW. MEGAHIT: an ultra-fast single-node solution for large and complex metagenomics assembly via succinct de Bruijn graph. Bioinformatics. 2014;31:1674–6.Article 
    CAS 

    Google Scholar 
    71.Hyatt D, Chen G-L, Locascio PF, Land ML, Larimer FW, Hauser LJ. Prodigal: prokaryotic gene recognition and translation initiation site identification. BMC Bioinforma. 2010;11:119.Article 
    CAS 

    Google Scholar 
    72.Eddy SR. Accelerated profile HMM searches. PLoS Comput Biol. 2011;7:e1002195.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    73.Li H, Durbin R. Fast and accurate short read alignment with Burrows-Wheeler transform. Bioinformatics. 2009;25:1754–60.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    74.Li H, Handsaker B, Wysoker A, Fennell T, Ruan J, Homer N, et al. The sequence alignment/map format and SAMtools. Bioinformatics. 2009;25:2078–9.PubMed 
    PubMed Central 
    Article 
    CAS 

    Google Scholar 
    75.Alneberg J, Bjarnason BS, de Bruijn I, Schirmer M, Quick J, Ijaz UZ, et al. Binning metagenomic contigs by coverage and composition. Nat Methods. 2014;11:1144–6.CAS 
    PubMed 
    Article 

    Google Scholar 
    76.Delmont TO, Eren AM. Identifying contamination with advanced visualization and analysis practices: metagenomic approaches for eukaryotic genome assemblies. PeerJ. 2016;4:e1839.PubMed 
    PubMed Central 
    Article 
    CAS 

    Google Scholar 
    77.Delcher AL, Phillippy A, Carlton J, Salzberg SL. Fast algorithms for large-scale genome alignment and comparison. Nucleic Acids Res. 2002;30:2478–83.PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    78.Parks DH, Imelfort M, Skennerton CT, Hugenholtz P, Tyson GW. CheckM: assessing the quality of microbial genomes recovered from isolates, single cells, and metagenomes. Genome Res. 2015;25:1043–55.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    79.Chaumeil PA, Mussig AJ, Hugenholtz P, Parks DH. GTDB-Tk: A toolkit to classify genomes with the genome taxonomy database. Bioinformatics. 2020. https://doi.org/10.1093/bioinformatics/btz848.80.Bateman A, Birney E, Durbin R, Eddy SR, Howe KL, Sonnhammer ELL. The Pfam protein families database. Nucleic Acids Res. 2000;28:263–6.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    81.Zdobnov EM, Apweiler R. InterProScan – an integration platform for the signature-recognition methods in InterPro. Bioinformatics. 2001;17:847–8.CAS 
    PubMed 
    Article 

    Google Scholar 
    82.Haft DH, Selengut JD, White O. The TIGRFAMs database of protein families. Nucleic Acids Res. 2003;31:371–3.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    83.Aziz RK, Bartels D, Best AA, DeJongh M, Disz T, Edwards RA, et al. The RAST Server: rapid annotations using subsystems technology. BMC Genomics. 2008;9:75.PubMed 
    PubMed Central 
    Article 
    CAS 

    Google Scholar 
    84.Altschul SF, Gish W, Miller W, Myers EW, Lipman DJ. Basic local alignment search tool. J Mol Biol. 1990;215:403–10.CAS 
    Article 

    Google Scholar 
    85.Darling AE, Jospin G, Lowe E, Matsen FA, Bik HM, Eisen JA. PhyloSift: phylogenetic analysis of genomes and metagenomes. PeerJ. 2014;2:e243.PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    86.Price MN, Dehal PS, Arkin AP. FastTree 2 — Approximately maximum-likelihood trees for large alignments. PLoS One. 2010;5:e9490.PubMed 
    PubMed Central 
    Article 
    CAS 

    Google Scholar  More

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    Multivariate trait analysis reveals diatom plasticity constrained to a reduced set of biological axes

    Culture maintenance and growthTwelve strains of Thalassiosira spp. were obtained from the Provasoli-Guillard National Centre of Marine Phytoplankton (NCMA, https://ncma.bigelow.org/), and one strain from the Australian National Culture Collection, representing 7 species in total (Supplementary Table 1). Cultures were maintained in polystyrene tissue culture flasks in artificial seawater with f/2 media [37] at 20 °C, with 60 µmolm−2s−1 of light on a 12:12 light cycle.Three strains originally identified as Thalassiosira sp. in the NCMA collection were further classified to the species level using sequencing of the ITS2 gene region (Supplementary Table 1): CCMP1055 as T. auguste-lineata (84.64% similarity; [38]) and CCMP2929 as T. weisflogii (98.37% similarity to Strain 1587 used in our study; [39]). Strain CCMP1059 was tentatively identified as Cyclotella striata (94.17% identity match to clone ZX28-3-40; [40]) also from order Thalassiosirales, but this assignment requires further investigation.Experimental set upExperimental cultures (200 mL) were grown in 250 mL polystyrene tissue culture flasks in triplicate, at a starting concentration of 2500 cells ml−1. All 13 strains were grown in a “standard” environment (identical to maintenance conditions) with 9 phenotypic traits measured to describe the initial trait-scape. Five strains (1010, 1059, 2929, 3264, and 3367) were grown in two additional environments in triplicate: a high temperature and light treatment (HT: 30 °C, 200 µmol photons m−2s−1 of light, 12:12 light:dark), and a low nutrient treatment (LN: f/400 media with an adjusted N:P ratio of 10:1 achieved by reducing the nitrate concentration from 4.4 to 1.8 µM, 60 µmol photons m−2s−1 of light, 12:12 light:dark). Cultures for the two additional treatments were inoculated with 10,000 cells ml−1 (LN) and 5,000 cells ml−1 (HT) in anticipation of limited growth.Growth was tracked daily using in vivo fluorescence as a proxy for cell density [41]. One mL aliquots of experimental cultures were measured for chlorophyll-a fluorescence using a plate reader (TECAN Infinite M1000 Pro, Männedorf, Switzerland) using 455/680 nm excitation/emission spectra. Phenotypic traits were measured at mid-late exponential phase, assessed by visually examining in vivo fluorescence growth curves. In the case of the low nutrient treatment, where growth was limited to 3–5 days, cultures were harvested in early stationary phase. Duration of growth for each experiment is summarised in Supplementary Table 2.Phenotypic trait measurement methodsPhenotypic traits were selected to capture different commonly measured base physiological functions, and to include traits that are used in biogeochemical models. We also selected traits that demonstrated independence and orthogonality (i.e., not all co-varying), based on pilot studies, in order to successfully define the multivariate trait-scape [42].Growth rateGrowth rates for each time step were calculated from the daily in vivo fluorescence measurements according to the calculation:$$mu = frac{{{{{{{{{mathrm{ln}}}}}}}}left( {F_2} right)-{{{{{{{mathrm{ln}}}}}}}}left( {F_1} right)}}{{t_2 – t_1}}$$Maximum growth rates were determined by the average growth over 2–4 consecutive steps depending on the duration of exponential growth.Flow cytometry traitsFor flow cytometry trait measures (growth rate, size, chlorophyll a content, lipid content), 1 mL aliquots of experimental culture were fixed with EM grade paraformaldehyde (0.8% final concentration, Electron Microscopy Sciences, Ft Washington, PA) in 1.6 mL cryopreservation tubes (CryoPure, Sarstedt), frozen in liquid nitrogen, then stored at −80 °C prior to analysis. All measures were performed using a Cytoflex LX (Beckman Coulter, CA, USA).Cell counts and sizeCell counts were done by gating the diatom population using chlorophyll a (488 nm excitation, 690/50 nm detector) and forward scatter channel thresholds. Cell size was estimated using forward scatter values calibrated against spherical beads (2, 4, 6, 10, 15 µM diameters; Invitrogen, CA). This resulted in a conversion equation of equivalent spherical diameter (ESD) = (FSC + 194636)/75775, which was used to assess relative changes in cell size [43].Chlorophyll a contentChlorophyll a (Chl-a) fluorescence of the gated diatom population was quantified using 488 nm excitation, 690/50 nm detection. A standard bead (Cytoflex Daily QC Fluorospheres; Beckman Coulter) was used to calibrate the performance of the instrument and ensure comparable measures across samples. Chlorophyll values were divided by ESD to account for cell size differences.Side scatter/granularitySide scatter is an indicator of the internal complexity of a cell or “granularity”. This trait is measured in tandem with other flow cytometry measures and was included as a phenotypic trait. The interpretation of this trait is not straight forward, but is independent of other flow cytometry traits measured and has been used in other flow cytometry studies of microalgae [44]. This trait was divided by ESD to account for cell size differences.Neutral lipidsRelative neutral lipid content was determined using the fluorescent stain BODIPY™ 505/515 (Thermo Fisher, MA, USA) which is commonly used to assess neutral lipid content in phytoplankton [45,46,47]. Background fluorescence (488 nm excitation, 525/40 nm detector) of PFA-fixed cells was measured in tandem with the size, chlorophyll a, and side scatter. After this, 10 µL of BODIPY stain (2 mg mL−1 in DMSO) was added to each sample, resulting in a final BODIPY concentration of 2 μg mL−1. Samples were incubated for 10 min in the dark before being read again on the flow cytometer. Neutral lipid content was defined as the difference in median fluorescence per cell between the pre- and post-stained sample. This value was then divided by the ESD size to account for size-related effects.Photophysiological traitsPhotophysiological measures were taken by conducting a rapid light curve [48] with a water PAM (Water-PAM; Walz GmbH, Effeltrich, Germany) using 1 mL of experimental culture diluted in artificial seawater. The rapid light curve protocol exposes the culture to 8 steps of increasing irradiance for 10 seconds each, measuring the photophysiological response at each step. Maximum electron transport rate (ETRmax), Ik (half saturation irradience), and alpha (the photosynthetic rate during the light-limited linear region) were calculated using the regression fit function in the PAM WinControl software. Photophysiology measurements were taken between 4–5 h after the start of the photoperiod.Reactive oxygen speciesThe development of reactive oxygen species (ROS) was measured using the fluorescent probe 2’,7’-dichlorodihydrofluorescein diacetate (H2DCFDA; Thermo Fisher, MA, USA) which has been used in a number of phytoplankton studies [49,50,51]. Two 1 mL aliquots of experimental culture were transferred to a 48 well tissue culture plate; 2 µL of stain (2.5 mg mL−1 H2DCFDA was made in DMSO) was added to one aliquot, with the other acting as a blank. The plates were sealed (Breathe-Easy, Diversified Biotech) and incubated in the dark at growth temperature (20 or 30 °C) for 2 h. Incubation was done in the dark because of the effects of light on the dye itself, therefore the effects of the excess light treatment were not captured in this trait. Fluorescence of H2DCFDA was read using a plate reader with 488 nm excitation 525 nm emission (TECAN Infinite M1000 Pro, Männedorf, Switzerland). ROS concentration was estimated as the difference in fluorescence units per cell between the stained and unstained aliquots of each culture. This metric was also divided by ESD size to account for size effects.Taxonomic confirmation of strainsDNA from stock cultures (10 mL) was extracted using a DNeasy PowerSoil kit (QIAGEN Inc., CA, USA) and checked for quality with a NanopDrop™ 2000 (ThermoFIsher Scientific, MA, USA), before amplification and sequencing at the Australian Genome Research Facility (AGRF, Sydney, Australia). PCR conditions and primers used were those developed by Chappell et al. [52] for the ITS region: forward primer: 5ʹ-RCGAAYTGCAGAACCTCG-3ʹ, reverse primer: 5ʹ-TACTYAATCTGAGATYCA-3ʹ.Bioinformatics processing was conducted using Geneious Prime (Version 2020.0.5; Biomatters Ltd.). Strain sequences were compared to GenBank using the BLAST function to confirm species identity. Nucleotide sequences were aligned using the MUSCLE alignment [53], followed by Bayesian inference analysis using MrBayes [54] to generate a phylogenetic tree. The out-group for the tree was a strain of Chaetoceros atlanticus isolate TPV2 1146 obtained from GenBank. Percentage similarity between strains according to the alignment was used as a metric of genetic relatedness.Statistical analysisWe assessed the multivariate phenotypes for the Thalassiosira strains using principal component analysis (PCA). The input variables were the 9 independent trait measurements made on each replicate culture (n = 36, 3 biological replicates per strain). Trait data was standardized (mean = 0, SD = 1) for each trait prior to PCA analysis to account for differences in the units and scale of measurements. The resulting PCA plot was defined as the ‘trait-scape’.Hierarchical clustering analysis was performed on the 9-trait dataset used to assess similarity in multivariate phenotypes between each replicate for each strain (n = 3 per strain).To compare genetic vs. phenotypic similarity, percentage similarity between strains was correlated against the distance between strain centroids (multivariate means) within the trait-scape. Distances between multivariate means (centroids) were calculated using the equation:$${{{{{{{mathrm{distance}}}}}}}} = sqrt {left( {{{{{{{{mathrm{{Delta}}}}}}}PC}}1.{{{{{{{mathrm{a}}}}}}}}} right)^2,+,left( {{{{{{{{mathrm{{Delta}}}}}}}PC}}2.{{{{{{{mathrm{b}}}}}}}}} right)^2}$$ΔPC1 is the difference in PC1 co-ordinates between the two strains, a is the % variance explained by PC1, ΔPC2 is the difference in PC2 co-ordinates between the two strains, b is the % variance explained by PC2.To assess whether a trait-scape generated using fewer input traits (4 rather than 9) was representative of the full, 9-trait plot, we conducted PCA using 4 input traits, and then assessed whether the inter-strain distances (distances between centroids) within the plot were correlated using linear regression. This provided a quantitative assessment of whether the strains were in the same relative positions to each other within the trait-scape.Covariation of traitsTo compare the pairwise relationships between traits across the strains, correlation matrices were made using data collected in the standard environment, and for the HT and LN environments.Phenotypic plasticityThe change in phenotypes in the new environments were assessed firstly by conducting PCA on the full dataset, including trait data from the 13 strains grown in the standard environment, plus the 5 strains grown in the two additional environments. This generated an “expanded trait-scape”. In addition, correlation matrices were generated for the new environments’ trait dataset to assess differences in trait-trait relationships between the ‘standard’ and “expanded” datasets.Relative changes in trait values for each trait in the new environments were calculated as follows:$$ {{{{{{{mathrm{Relative}}}}}}}},{{{{{{{mathrm{change}}}}}}}} \ = frac{{{{{{{{{mathrm{trait}}}}}}}},{{{{{{{mathrm{value}}}}}}}},{{{{{{{mathrm{new}}}}}}}},{{{{{{{mathrm{environment}}}}}}}} – overline {{{{{{{mathrm{x}}}}}}}} ,,{{{{{{{mathrm{trait}}}}}}}},{{{{{{{mathrm{value}}}}}}}},{{{{{{{mathrm{standard}}}}}}}},{{{{{{{mathrm{environment}}}}}}}}}}{{overline {{{{{{{mathrm{x}}}}}}}} ,,{{{{{{{mathrm{trait}}}}}}}},{{{{{{{mathrm{value}}}}}}}},{{{{{{{mathrm{standard}}}}}}}},{{{{{{{mathrm{environment}}}}}}}}}}$$We used PCA to assess whether the relative changes in trait values were consistent between strains in the two different environments. i.e., was the relative change in whole phenotype consistent. If the changes were consistent across strains, we expected to see clustering in the PCA based on treatment.Statistical softwareStatistical analyses were performed in R [55], Matlab, and Microsoft Excel. Hierarchical clustering analysis with multiscale bootstrap resampling (1000 replicates) on trait values from biological replicates was done with the ‘pvclust’ package in R [56] using Euclidean distance and the average (UPGMA) method. Principal component analysis was used to generate the multivariate trait-scape was done using the “vegan package” in R [57]. The contributions of each trait to the PC axes (loadings) were extracted using the “factoextra” package in R [58]. Trait correlation matrices were generated using the “corrplot” package in R [59]. More

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    Acrylate protects a marine bacterium from grazing by a ciliate predator

    1.Yang, J. W. et al. Predator and prey biodiversity relationship and its consequences on marine ecosystem functioning-interplay between nanoflagellates and bacterioplankton. ISME J. 12, 1532–1542 (2018).PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    2.Zan, J. et al. A microbial factory for defensive kahalalides in a tripartite marine symbiosis. Science 364, eaaw6732 (2019).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    3.Yoch, D. C. Dimethylsulfoniopropionate: its sources, role in the marine food web, and biological degradation to dimethylsulfide. Appl. Environ. Microbiol. 68, 5804–5815 (2002).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    4.Bullock, H. A., Luo, H. & Whitman, W. B. Evolution of dimethylsulfoniopropionate metabolism in marine phytoplankton and bacteria. Front. Microbiol. 8, 637 (2017).PubMed 
    PubMed Central 

    Google Scholar 
    5.Curson, A. R. J. et al. DSYB catalyses the key step of dimethylsulfoniopropionate biosynthesis in many phytoplankton. Nat. Microbiol. 3, 430–439 (2018).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    6.Curson, A. et al. Dimethylsulfoniopropionate biosynthesis in marine bacteria and identification of the key gene in this process. Nat. Microbiol. 2, 17009 (2017).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    7.Williams, B. T. et al. Bacteria are important dimethylsulfoniopropionate producers in coastal sediments. Nat. Microbiol. 4, 1815–1825 (2019).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    8.Zhang, X. H. et al. Biogenic production of DMSP and its degradation to DMS—their roles in the global sulfur cycle. Sci. China Life. Sci. 62, 1296–1319 (2019).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    9.Alstyne, K. L. V., Wolfe, G. V., Freidenburg, T. L., Neill, A. & Hicken, C. Activated defense systems in marine macroalgae: evidence for an ecological role for DMSP cleavage. Mar. Ecol. Prog. Ser. 213, 53–65 (2001).Article 

    Google Scholar 
    10.Paul, V. J. & Van Alstyne, K. L. Activation of chemical defenses in the tropical green algae Halimeda spp. J. Exp. Mar. Biol. Ecol. 160, 191–203 (1992).CAS 
    Article 

    Google Scholar 
    11.Strom, S. et al. Chemical defense in the microplankton I: feeding and growth rates of heterotrophic protists on the DMS-producing phytoplankter Emiliania huxleyi. Limnol. Oceangr. 48, 217–229 (2003).CAS 
    Article 

    Google Scholar 
    12.Wolfe, G. V., Steinke, M. & Kirst, G. O. Grazing-activated chemical defence in a unicellular marine alga. Nature 387, 894–897 (1997).CAS 
    Article 

    Google Scholar 
    13.Liu, C. et al. Puniceibacterium antarcticum gen. nov., sp. nov., isolated from seawater. Int. J. Syst. Evol. Microbiol. 64, 1566–1572 (2014).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    14.Aronson, D. E., Costantini, L. M. & Snapp, E. L. Superfolder GFP is fluorescent in oxidizing environments when targeted via the Sec translocon. Traffic 12, 543–548 (2011).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    15.Coppellotti Krupa, O. & Vannucci, D. Citrate synthase from Antarctic ciliates: adaptation to low temperatures and comparison with temperate ciliates. Polar Biol. 26, 452–457 (2003).Article 

    Google Scholar 
    16.Asher, E. C., Dacey, J. W. H., Stukel, M., Long, M. C. & Tortell, P. D. Processes driving seasonal variability in DMS, DMSP, and DMSO concentrations and turnover in coastal Antarctic waters. Limnol. Oceanogr. 62, 104–124 (2017).Article 

    Google Scholar 
    17.Ahmed, M., Stal, L. J. & Hasnain, S. DTAF: an efficient probe to study cyanobacterial-plant interaction using confocal laser scanning microscopy (CLSM). J. Ind. Microbiol. Biotechnol. 38, 249–255 (2011).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    18.Hojo, F. et al. Ciliates expel environmental Legionella-laden pellets to stockpile food. Appl. Environ. Microbiol. 78, 5247–5257 (2012).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    19.Seymour, J. R., Simo, R., Ahmed, T. & Stocker, R. Chemoattraction to dimethylsulfoniopropionate throughout the marine microbial food web. Science 329, 342–345 (2010).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    20.Shemi, A. et al. Dimethyl sulfide acts as eat-me signal during microbial predator–prey interactions in the ocean. Research Square https://doi.org/10.21203/rs.3.rs-139243/v1 (2021).21.Chen, I. A. et al. IMG/M v.5.0: an integrated data management and comparative analysis system for microbial genomes and microbiomes. Nucleic Acids Res. 47, D666–D677 (2019).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    22.Wang, P. et al. Structural and molecular basis for the novel catalytic mechanism and evolution of DddP, an abundant peptidase-like bacterial dimethylsulfoniopropionate lyase: a new enzyme from an old fold. Mol. Microbiol. 98, 289–301 (2015).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    23.Li, C. Y. et al. Molecular insight into bacterial cleavage of oceanic dimethylsulfoniopropionate into dimethyl sulfide. Proc. Natl Acad. Sci. USA 111, 1026–1031 (2014).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    24.González, J. M., Whitman, W. B., Hodson, R. E. & Moran, M. A. Identifying numerically abundant culturable bacteria from complex communities: an example from a lignin enrichment culture. Appl. Environ. Microbiol. 62, 4433–4440 (1996).PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    25.Freier, D., Mothershed, C. P. & Wiegel, J. Characterization of Clostridium thermocellum JW20. Appl. Environ. Microbiol. 54, 204–JW211 (1988).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    26.Wang, P. et al. Development of an efficient conjugation-based genetic manipulation system for Pseudoalteromonas. Microb. Cell Fact. 14, 11 (2015).PubMed 
    PubMed Central 
    Article 
    CAS 

    Google Scholar 
    27.Obranic, S., Babic, F. & Maravic-Vlahovicek, G. Improvement of pBBR1MCS plasmids, a very useful series of broad-host-range cloning vectors. Plasmid 70, 263–267 (2013).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    28.Dinh, T. & Bernhardt, T. G. Using superfolder green fluorescent protein for periplasmic protein localization studies. J. Bacteriol. 193, 4984–4987 (2011).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    29.Yu, Z. C. et al. Development of a genetic system for the deep-sea psychrophilic bacterium Pseudoalteromonas sp. SM9913. Microb. Cell Fact. 13, 13 (2014).PubMed 
    PubMed Central 
    Article 
    CAS 

    Google Scholar 
    30.Walker, J. M. The bicinchoninic acid (BCA) assay for protein quantitation. Methods Mol. Biol. 32, 5–8 (1994).CAS 
    PubMed 
    PubMed Central 

    Google Scholar 
    31.Ansede, J. H., Pellechia, P. J. & Yoch, D. C. Metabolism of acrylate to beta-hydroxypropionate and its role in dimethylsulfoniopropionate lyase induction by a salt marsh sediment bacterium, Alcaligenes faecalis M3A. Appl. Environ. Microbiol. 65, 5075–5081 (1999).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    32.Liu, J. et al. Novel insights into bacterial dimethylsulfoniopropionate catabolism in the East China Sea. Front. Microbiol. 9, 3206–3206 (2018).PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    33.Shao, X. et al. Mechanistic insight into 3-methylmercaptopropionate metabolism and kinetical regulation of demethylation pathway in marine dimethylsulfoniopropionate-catabolizing bacteria. Mol. Microbiol. 111, 1057–1073 (2019).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    34.Dumon-Seignovert, L., Cariot, G. & Vuillard, L. The toxicity of recombinant proteins in Escherichia coli: a comparison of overexpression in BL21(DE3), C41(DE3), and C43(DE3). Protein Expr. Purif. 37, 203–206 (2004).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    35.Yamashita, A., Singh, S. K., Kawate, T., Jin, Y. & Gouaux, E. Crystal structure of a bacterial homologue of Na+/Cl−-dependent neurotransmitter transporters. Nature 437, 215–223 (2005).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    36.Barek, J., Pumera, M., Muck, A., Kadeřabkova, M. & Zima, J. Polarographic and voltammetric determination of selected nitrated polycyclic aromatic hydrocarbons. Anal. Chim. Acta 393, 141–146 (1999).CAS 
    Article 

    Google Scholar 
    37.Sherr, B. F., Sherr, E. B. & Fallon, R. D. Use of monodispersed, fluorescently labeled bacteria to estimate in situ protozoan bacterivory. Appl. Environ. Microbiol. 53, 958–965 (1987).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    38.Perez-Uz, B. Bacterial preferences and growth kinetic variation in Uronema marinum and Uronema nigricans (Ciliophora: Scuticociliatida). Microb. Ecol. 31, 189–198 (1996).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    39.Siegmund, L., Schweikert, M., Fischer, M. S. & Wostemeyer, J. Bacterial surface traits influence digestion by Tetrahymena pyriformis and alter opportunity to escape from food vacuoles. J. Eukaryot. Microbiol. 65, 600–611 (2018).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    40.Christaki, U. et al. Optimized routine flow cytometric enumeration of heterotrophic flagellates using SYBR Green I. Limnol. Oceanogr. Meth. 9, 329–339 (2011).Article 

    Google Scholar 
    41.Headland, S. E., Jones, H. R., D’Sa, A. S., Perretti, M. & Norling, L. V. Cutting-edge analysis of extracellular microparticles using ImageStream(X) imaging flow cytometry. Sci. Rep. 4, 5237 (2014).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    42.Hayduk, W. & Laudie, H. Prediction of diffusion coefficients for nonelectrolytes in dilute aqueous solutions. AIChE J. 20, 611–615 (1974).CAS 
    Article 

    Google Scholar 
    43.Schotte, W. Prediction of the molar volume at the normal boiling point. Chem. Eng. J. 48, 167–172 (1992).CAS 
    Article 

    Google Scholar 
    44.Carrión, O. et al. A novel pathway producing dimethylsulphide in bacteria is widespread in soil environments. Nat. Commun. 6, 6579 (2015).PubMed 
    Article 
    CAS 
    PubMed Central 

    Google Scholar 
    45.Zhang, W. et al. Marine biofilms constitute a bank of hidden microbial diversity and functional potential. Nat. Commun. 10, 517 (2019).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    46.Bailey, T. L. & Elkan, C. Fitting a mixture model by expectation maximization to discover motifs in biopolymers. Proc. Int. Conf. Intell. Syst. Mol. Biol. 2, 28–36 (1994).CAS 
    PubMed 
    PubMed Central 

    Google Scholar 
    47.Hoffman, K. & Stoffel, W. TMbase—a database of membrane spanning proteins segments. Biol. Chem. Hoppe-Seyler 374, 166 (1993).
    Google Scholar 
    48.Bansal, M. S., Alm, E. J. & Kellis, M. Efficient algorithms for the reconciliation problem with gene duplication, horizontal transfer and loss. Bioinformatics 28, i283–i291 (2012).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar  More

  • in

    Association of bacterial community types, functional microbial processes and lung disease in cystic fibrosis airways

    1.Filkins LM, Hampton TH, Gifford AH, Gross MJ, Hogan DA, Sogin ML, et al. Prevalence of Streptococci and increased polymicrobial diversity associated with cystic fibrosis patient stability. J Bacteriol. 2012;194:4709–17.CAS 
    Article 

    Google Scholar 
    2.Fodor AA, Klem ER, Gilpin DF, Elborn JS, Boucher RC, Tunney MM, et al. The adult cystic fibrosis airway microbiota is stable over time and infection type, and highly resilient to antibiotic treatment of exacerbations. PLoS One. 2012;7:e45001.CAS 
    Article 

    Google Scholar 
    3.Goddard AF, Staudinger BJ, Dowd SE, Joshi-Datar A, Wolcott RD, Aitken ML, et al. Direct sampling of cystic fibrosis lungs indicates that DNA-based analyses of upper-airway specimens can misrepresent lung microbiota. Proc Natl Acad Sci USA. 2012;109:13769–74.CAS 
    Article 

    Google Scholar 
    4.Guss AM, Roeselers G, Newton IL, Young CR, Klepac-Ceraj V, Lory S, et al. Phylogenetic and metabolic diversity of bacteria associated with cystic fibrosis. ISME J. 2011;5:20–9.Article 

    Google Scholar 
    5.Harris JK, De Groote MA, Sagel SD, Zemanick ET, Kapsner R, Penvari C, et al. Molecular identification of bacteria in bronchoalveolar lavage fluid from children with cystic fibrosis. Proc Natl Acad Sci USA. 2007;104:20529–33.CAS 
    Article 

    Google Scholar 
    6.Brown PS, Pope CE, Marsh RL, Qin X, McNamara S, Gibson R, et al. Directly sampling the lung of a young child with cystic fibrosis reveals diverse microbiota. Ann Am Thorac Soc. 2014;11:1049–55.Article 

    Google Scholar 
    7.Jorth P, Staudinger BJ, Wu X, Hisert KB, Hayden H, Garudathri J, et al. Regional isolation drives bacterial diversification within cystic fibrosis lungs. Cell Host Microbe. 2015;18:307–19.CAS 
    Article 

    Google Scholar 
    8.Sibley CD, Parkins MD, Rabin HR, Duan K, Norgaard JC, Surette MG. A polymicrobial perspective of pulmonary infections exposes an enigmatic pathogen in cystic fibrosis patients. Proc Natl Acad Sci USA. 2008;105:15070–5.CAS 
    Article 

    Google Scholar 
    9.van der Gast CJ, Walker AW, Stressmann FA, Rogers GB, Scott P, Daniels TW, et al. Partitioning core and satellite taxa from within cystic fibrosis lung bacterial communities. ISME J. 2011;5:780–91.Article 

    Google Scholar 
    10.Zhao J, Carmody LA, Kalikin LM, Li J, Petrosino JF, Schloss PD, et al. Impact of enhanced Staphylococcus DNA extraction on microbial community measures in cystic fibrosis sputum. PLoS One. 2012;7:e33127.CAS 
    Article 

    Google Scholar 
    11.Carmody LA, Zhao J, Schloss PD, Petrosino JF, Murray S, Young VB, et al. Changes in cystic fibrosis airway microbiota at pulmonary exacerbation. Ann Am Thorac Soc. 2013;10:179–87.Article 

    Google Scholar 
    12.Cox MJ, Allgaier M, Taylor B, Baek MS, Huang YJ, Daly RA, et al. Airway microbiota and pathogen abundance in age-stratified cystic fibrosis patients. PLoS One. 2010;5:e11044.Article 

    Google Scholar 
    13.Stressmann FA, Rogers GB, van der Gast CJ, Marsh P, Vermeer LS, Carroll MP, et al. Long-term cultivation-independent microbial diversity analysis demonstrates that bacterial communities infecting the adult cystic fibrosis lung show stability and resilience. Thorax. 2012;67:867–73.Article 

    Google Scholar 
    14.Zhao J, Schloss PD, Kalikin LM, Carmody LA, Foster BK, Petrosino JF, et al. Decade-long bacterial community dynamics in cystic fibrosis airways. Proc Natl Acad Sci USA. 2012;109:5809–14.CAS 
    Article 

    Google Scholar 
    15.Rogers GB, Bruce KD, Hoffman LR. How can the cystic fibrosis respiratory microbiome influence our clinical decision-making? Curr Opin Pulm Med. 2017;23:536–43.Article 

    Google Scholar 
    16.Widder S, Knapp S. Microbial metabolites in cystic fibrosis: a target for future therapy? Am J Respir Cell Mol Biol. 2019;61:132–3.17.Mahboubi MA, Carmody LA, Foster BK, Kalikin LM, VanDevanter DR, LiPuma JJ. Culture-based and culture-independent bacteriologic analysis of cystic fibrosis respiratory specimens. J Clin Microbiol. 2016;54:613–9.CAS 
    Article 

    Google Scholar 
    18.Carmody LA, Caverly LJ, Foster BK, Rogers MAM, Kalikin LM, Simon RH, et al. Fluctuations in airway bacterial communities associated with clinical states and disease stages in cystic fibrosis. PLoS One. 2018;13:e0194060.Article 

    Google Scholar 
    19.Zhao J, Li J, Schloss PD, Kalikin LM, Raymond TA, Petrosino JF, et al. Effect of sample storage conditions on cultureindependent bacterial community measures in cystic fibrosis sputum specimens. J Clin Microbiol 2011;49:3717–8.20.Hnizdo E, Yu L, Freyder L, Attfield M, Lefante J & Glindmeyer HW. The precision of longitudinal lung function measurements: Monitoring and interpretation. Occup Environ Med 2005;62:695–701.21.Konstan MW, Wagener JS, VanDevanter DR. Characterizing aggressiveness and predicting future progression of CF lung disease. J Cyst Fibros. 2009;8:S15–19.Article 

    Google Scholar 
    22.Schloss PD, Gevers D, Westcott SL. Reducing the effects of PCR amplification and sequencing artifacts on 16s rRNA-based studies. PLoS One. 2011;6:e27310.CAS 
    Article 

    Google Scholar 
    23.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 
    Article 

    Google Scholar 
    24.Cole JR, Wang Q, Cardenas E, Fish J, Chai B, Farris RJ, et al. The ribosomal database project: Improved alignments and new tools for rRNA analysis. Nucleic Acids Res. 2009;37:D141–145.CAS 
    Article 

    Google Scholar 
    25.Sung J, Kim S, Cabatbat JJT, Jang S, Jin YS, Jung GY, et al. Global metabolic interaction network of the human gut microbiota for context-specific community-scale analysis. Nat Commun 2017;8:15393.26.Friedman J, Alm EJ. Inferring correlation networks from genomic survey data. PLoS Comput Biol. 2012;8:e1002687.CAS 
    Article 

    Google Scholar 
    27.Holmes I, Harris K, Quince C. Dirichlet multinomial mixtures: generative models for microbial metagenomics. PLoS One. 2012;7:e30126.CAS 
    Article 

    Google Scholar 
    28.Ding T, Schloss PD. Dynamics and associations of microbial community types across the human body. Nature. 2014;509:357–60.CAS 
    Article 

    Google Scholar 
    29.Price KE, Hampton TH, Gifford AH, Dolben EL, Hogan DA, Morrison HG, et al. Unique microbial communities persist in individual cystic fibrosis patients throughout a clinical exacerbation. Microbiome 2013;1:27.30.Carmody LA, Zhao J, Kalikin LM, LeBar W, Simon RH, Venkataraman A, et al. The daily dynamics of cystic fibrosis airway microbiota during clinical stability and at exacerbation. Microbiome 2015;3:12.31.de Dios Caballero J, Vida R, Cobo M, Maiz L, Suarez L, Galeano J, et al. Individual patterns of complexity in cystic fibrosis lung microbiota, including predator bacteria, over a 1-year period. mBio 2017;8::e00959–17.32.Whelan, FJ, Heirali AA, Rossi L, Rabin HR, Parkins MD, & Surette MG. Longitudinal sampling of the lung microbiota in individuals with cystic fibrosis. PLoS One 2017:12:e0172811.33.Noecker C, Eng A, Srinivasan S, Theriot CM, Young VB, Jansson JK, et al. Metabolic model-based integration of microbiome taxonomic and metabolomic profiles elucidates mechanistic links between ecological and metabolic variation. mSystems 2016;1.34.Douglas, GM, Maffei, VJ, Zaneveld, JR, Yurgel, SN, Brown JR, Taylor CM, et al. PICRUSt2 for prediction of metagenome functions. Nat Biotechnol 2020;38:685–8.35.Caspi R, Billington R, Fulcher CA, Keseler IM, Kothari A, Krummenacker M, et al. The MetaCyc database of metabolic pathways and enzymes. Nucleic Acids Res 2018;46:D633–D639.36.Quinn RA, Comstock W, Zhang T, Morton JT, da Silva R, Tran A, et al. Niche partitioning of a pathogenic microbiome driven by chemical gradients. Sci Adv 2018;4:eaau1908.37.Quinn RA, Whiteson K, Lim YW, Zhao J, Conrad D, LiPuma JJ, et al. Ecological networking of cystic fibrosis lung infections. NPJ Biofilms Microbiomes. 2016;2:4.Article 

    Google Scholar 
    38.Pradeu T, Vivier E. The discontinuity theory of immunity. Sci Immunol. 2016;1:AAG0479.39.Flynn JM, Niccum D, Dunitz JM, Hunter RC. Evidence and role for bacterial mucin degradation in cystic fibrosis airway disease. PLoS Pathog. 2016;12:e1005846.Article 

    Google Scholar 
    40.Adamowicz EM, Flynn J, Hunter RC, Harcombe WR. Cross-feeding modulates antibiotic tolerance in bacterial communities. ISME J. 2018;12:2723–35.CAS 
    Article 

    Google Scholar 
    41.Rose MC & Voynow JA. Respiratory tract mucin genes and mucin glycoproteins in health and disease. Physiol Rev 2006;86:245–78.42.Tailford LE, Crost EH, Kavanaugh D. & Juge N. Mucin glycan foraging in the human gut microbiome. Front Genet 2015;6:81.43.Wheeler KM, Carcamo-Oyarce G, Turner BS, Dellos-Nolan S, Co JY, Lehoux S, et al. Mucin glycans attenuate the virulence of Pseudomonas aeruginosa in infection. Nat Microbiol 2019;4:2146–54.44.Twomey KB, O’Connell OJ, McCarthy Y, Dow JM, O’Toole GA, Plant BJ, et al. Bacterial cis-2-unsaturated fatty acids found in the cystic fibrosis airway modulate virulence and persistence of Pseudomonas aeruginosa. ISME J 2012;6:939–50.45.Zemanick ET, Wagner BD, Robertson CE, Ahrens RC, Chmiel JF, Clancy JP, et al. Airway microbiota across age and disease spectrum in cystic fibrosis. Eur Respir J 2017;50:1700832.46.Lu J, Carmody LA, Opron K, Simon RH, Kalikin LM, Caverly LJ, et al. Parallel analysis of cystic fibrosis sputum and saliva’reveals overlapping communities and an opportunity for sample decontamination. mSystems 2020;5.47.Jones KL, Hegab AH, Hillman BC, Simpson KL, Jinkins PA, Grisham MB, et al. Elevation of nitrotyrosine and nitrate concentrations in cystic fibrosis sputum. Pediatr Pulmonol 2000;30:79–85.48.Quinn RA, Lim YW, Maughan H, Conrad D, Rohwer F, Whiteson KL. Biogeochemical forces shape the composition and physiology of polymicrobial communities in the cystic fibrosis lung. mBio. 2014;5:e00956–00913.Article 

    Google Scholar 
    49.Mirkovic B, Murray MA, Lavelle GM, Molloy K, Azim AA, Gunaratnam C, et al. The role of short-chain fatty acids, produced by anaerobic bacteria, in the cystic fibrosis airway. Am J Respir Crit Care Med. 2015;192:1314–24.CAS 
    Article 

    Google Scholar 
    50.Trompette A, Gollwitzer ES, Pattaroni C, Lopez-Mejia IC, Riva E, Pernot J, et al. Dietary fiber confers protection against flu by shaping Ly6c(-) patrolling monocyte hematopoiesis and CD8(+) t cell metabolism. Immunity. 2018;48:992–1005.e1008.CAS 
    Article 

    Google Scholar 
    51.Flynn JM, Phan C, Hunter RC. Genome-wide survey of Pseudomonas aeruginosa PA14 reveals a role for the glyoxylate pathway and extracellular proteases in the utilization of mucin. Infect Immun. 2017;85:e00182–17.52.Jorth P, Ehsan Z, Rezayat A, Caldwell E, Pope C, Brewington JJ, et al. Direct lung sampling indicates that established pathogens dominate early infections in children with cystic fibrosis. Cell Rep. 2019;27:1190–204.e1193.CAS 
    Article 

    Google Scholar 
    53.Charalampous T, Kay GL, Richardson H, Aydin A, Baldan R, Jeanes C, et al. Nanopore metagenomics enables rapid clinical diagnosis of bacterial lower respiratory infection. Nat Biotechnol. 2019;37:783–92.CAS 
    Article 

    Google Scholar 
    54.Cowley ES, Kopf SH, LaRiviere A, Ziebis W, Newman DK. Pediatric cystic fibrosis sputum can be chemically dynamic, anoxic, and extremely reduced due to hydrogen sulfide formation. mBio. 2015;6:e00767.CAS 
    Article 

    Google Scholar 
    55.Cuthbertson L, Walker AW, Oliver AE, Rogers GB, Rivett DW, Hampton TH, et al. Lung function and microbiota diversity in cystic fibrosis. Microbiome 2020;8:45. More

  • in

    Sublethal effects of bifenazate on biological traits and enzymatic properties in the Panonychus citri (Acari: Tetranychidae)

    1.Zhang, Z. Y. et al. A shift pattern of bacterial communities across the life stages of the citrus red mite, Panonychus citri. Front. Microbiol. 11, 1620. https://doi.org/10.3389/fmicb.2020.01620 (2020).Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    2.Pan, D., Dou, W., Yuan, G. R., Zhou, Q. H. & Wang, J. J. Monitoring the resistance of the citrus red mite (Acari: Tetranychidae) to four acaricides in different citrus orchards in China. J. Econ. Entomol. 113, 918–923. https://doi.org/10.1093/jee/toz335 (2020).CAS 
    Article 
    PubMed 

    Google Scholar 
    3.Zhang, Y., Guo, L., Atlihan, R., Chi, H. & Chu, D. Demographic analysis of progeny fitness and timing of resurgence of Laodelphax striatellus after insecticides exposure. Entomol. Generalis 39, 221–230. https://doi.org/10.1127/entomologia/2019/0816 (2019).Article 

    Google Scholar 
    4.Quesada, C. R. & Sadof, C. S. Field evaluation of insecticides and application timing on natural enemies of selected armored and soft scales. Biol. Control 133, 81–90. https://doi.org/10.1016/j.biocontrol.2019.03.013 (2019).CAS 
    Article 

    Google Scholar 
    5.Ullah, F. et al. Fitness costs in chlorfenapyr-resistant populations of the chive maggot, Bradysia odoriphaga. Ecotoxicology 29, 407–416. https://doi.org/10.1007/s10646-020-02183-7 (2020).CAS 
    Article 
    PubMed 

    Google Scholar 
    6.Razik, M. A. R. A. M. A. Toxicity and side effects of some insecticides applied in cotton fields on Apis mellifera. Environ. Sci. Pollut. R. 26, 4987–4996. https://doi.org/10.1007/s11356-018-04061-6 (2019).CAS 
    Article 

    Google Scholar 
    7.Ochiai, N. et al. Toxicity of bifenazate and its principal active metabolite, diazene, to Tetranychus urticae and Panonychus citri and their relative toxicity to the predaceous mites, Phytoseiulus persimilis and Neoseiulus californicus. Exp. Appl. Acarol. 43, 181–197. https://doi.org/10.1007/s10493-007-9115-9 (2007).CAS 
    Article 
    PubMed 

    Google Scholar 
    8.Van Nieuwenhuyse, P. et al. On the mode of action of bifenazate: New evidence for a mitochondrial target site. Pestic. Biochem. Physiol. 104, 88–95. https://doi.org/10.1016/j.pestbp.2012.05.013 (2012).CAS 
    Article 

    Google Scholar 
    9.Wang, R. et al. Lethal and sublethal effects of a novel cis-nitromethylene neonicotinoid insecticide, cycloxaprid, on Bemisia tabaci. Crop Prot. 83, 15–19. https://doi.org/10.1016/j.cropro.2016.01.015 (2016).CAS 
    Article 

    Google Scholar 
    10.Ullah, F., Gul, H., Desneux, N., Gao, X. & Song, D. Imidacloprid-induced hormesis effects on demographic traits of the melon aphid, Aphis gossypii. Entomol. Generalis 39, 325–337 (2019).Article 

    Google Scholar 
    11.Dong, J., Wang, K., Li, Y. & Wang, S. Lethal and sublethal effects of cyantraniliprole on Helicoverpa assulta (Lepidoptera: Noctuidae). Pestic. Biochem. Physiol. 136, 58–63. https://doi.org/10.1016/j.pestbp.2016.08.003 (2017).ADS 
    CAS 
    Article 
    PubMed 

    Google Scholar 
    12.Elzen, G. W. Lethal and sublethal effects of insecticide residues on Orius insidiosus (Hemiptera: Anthocoridae) and Geocoris punctipes (Hemiptera: Lygaeidae). J. Econ. Entomol. 94, 55–59. https://doi.org/10.1603/0022-0493-94.1.55 (2001).CAS 
    Article 
    PubMed 

    Google Scholar 
    13.Desneux, N., Decourtye, A. & Delpuech, J. M. The sublethal effects of pesticides on beneficial arthropods. Annu. Rev. Entomol. 52, 81–106. https://doi.org/10.1146/annurev.ento.52.110405.091440 (2007).CAS 
    Article 
    PubMed 

    Google Scholar 
    14.Deng, D. et al. Assessment of the effects of lethal and sublethal exposure to dinotefuran on the wheat aphid Rhopalosiphum padi (Linnaeus). Ecotoxicology 28, 825–833. https://doi.org/10.1007/s10646-019-02080-8 (2019).CAS 
    Article 
    PubMed 

    Google Scholar 
    15.Guo, L. et al. Sublethal and transgenerational effects of chlorantraniliprole on biological traits of the diamondback moth, Plutella xylostella L.. Crop Prot. 48, 29–34. https://doi.org/10.1016/j.cropro.2013.02.009 (2013).CAS 
    Article 

    Google Scholar 
    16.Duke, S. O. et al. (eds) Pesticide Dose: Effects on the Environment and Target and Non-target Organisms 101–119 (American Chemical Society, 2017).
    Google Scholar 
    17.Guedes, N. M. P., Tolledo, J., Correa, A. S. & Guedes, R. N. C. Insecticide-induced hormesis in an insecticide-resistant strain of the maize weevil, Sitophilus zeamais. J. Appl. Entomol. 134, 142–148. https://doi.org/10.1111/j.1439-0418.2009.01462.x (2010).CAS 
    Article 

    Google Scholar 
    18.Haddi, K., Oliveira, E. E., Faroni, L. R., Guedes, D. C. & Miranda, N. N. Sublethal exposure to clove and cinnamon essential oils induces hormetic-like responses and disturbs behavioral and respiratory responses in Sitophilus zeamais (Coleoptera: Curculionidae). J. Econ. Entomol. 108, 2815–2822. https://doi.org/10.1093/jee/tov255 (2015).CAS 
    Article 
    PubMed 

    Google Scholar 
    19.Chen, X. D., Seo, M. & Stelinski, L. L. Behavioral and hormetic effects of the butenolide insecticide, flupyradifurone, on Asian citrus psyllid, Diaphorina citri. Crop Prot. 98, 102–107. https://doi.org/10.1016/j.cropro.2017.03.017 (2017).CAS 
    Article 

    Google Scholar 
    20.Wang, L., Zhang, Y., Xie, W., Wu, Q. & Wang, S. Sublethal effects of spinetoram on the two-spotted spider mite, Tetranychus urticae (Acari: Tetranychidae). Pestic Biochem. Physiol. 132, 102–107. https://doi.org/10.1016/j.pestbp.2016.02.002 (2016).CAS 
    Article 
    PubMed 

    Google Scholar 
    21.Xiao, L. F. et al. Genome-wide identification, phylogenetic analysis, and expression profiles of ATP-binding cassette transporter genes in the oriental fruit fly, Bactrocera dorsalis (Hendel) (Diptera: Tephritidae). Comp. Biochem. Physiol. D Genomics Proteomics 25, 1–8 (2018).ADS 
    CAS 
    Article 

    Google Scholar 
    22.Dubovskiy, I. M. et al. Effect of bacterial infection on antioxidant activity and lipid peroxidation in the midgut of Galleria mellonella L. larvae (Lepidoptera, Pyralidae). Comp. Biochem. Physiol. C Toxicol. Pharmacol. 148, 1–5 (2008).CAS 
    Article 

    Google Scholar 
    23.Jia, B. T., Hong, S. S., Zhang, Y. C. & Cao, Y. W. Effect of sublethal concentrations of abamectin on protective and detoxifying enzymes in Diagegma semclausum. J. Environ. Entomol. 38, 990 (2016).
    Google Scholar 
    24.Yang, Q., Wang, S., Zhang, W., Yang, T. & Liu, Y. Toxicity of commonly used insecticides and their influences on protective enzyme activity of multicolored Asian lady beetle Harmonia axyridis (Pallas). Acta Phytophylacica Sin. 42, 258–263 (2015).CAS 

    Google Scholar 
    25.Zhou, C., Yang, H., Wang, Z., Long, G. Y. & Jin, D. C. Protective and detoxifying enzyme activity and abcg subfamily gene expression in Sogatella furcifera under insecticide stress. Front. Physiol. 9, 1890. https://doi.org/10.3389/fphys.2018.01890 (2018).Article 
    PubMed 

    Google Scholar 
    26.Cui, L., Yuan, H., Wang, Q., Wang, Q. & Rui, C. Sublethal effects of the novel cis-nitromethylene neonicotinoid cycloxaprid on the cotton aphid Aphis gossypii Glover (Hemiptera: Aphididae). Sci. Rep. 8, 8915. https://doi.org/10.1038/s41598-018-27035-7 (2018).ADS 
    CAS 
    Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    27.Li, Y. Y. et al. Sublethal effects of bifenazate on life history and population parameters of Tetranychus urticae (Acari: Tetranychidae). Syst. Appl. Acarol. 22, 148–158. https://doi.org/10.11158/saa.22.1.15 (2017).CAS 
    Article 

    Google Scholar 
    28.Wang, Y., Huang, X., Chang, B. H. & Zhang, Z. The survival, growth, and detoxifying enzyme activities of grasshoppers Oedaleus asiaticus (Orthoptera: Acrididae) exposed to toxic rutin. Appl. Entomol. Zool. 55, 385–393. https://doi.org/10.1007/s13355-020-00694-7 (2020).CAS 
    Article 

    Google Scholar 
    29.Rasheed, M. A. et al. Lethal and sublethal effects of chlorpyrifos on biological traits and feeding of the aphidophagous predator Harmonia axyridis. Insects. https://doi.org/10.3390/insects11080491 (2020).Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    30.Yamamoto, A., Yoneda, H., Hatano, R. & Asada, M. Genetic analysis of hexythiazox resistance in the citrus red mite, Panonychus citri (MCGREGOR). J. Pestic. Sci. 20, 513–519 (1995).CAS 
    Article 

    Google Scholar 
    31.Chi, H. & Liu, H. Two new methods for the study of insect population ecology. Acad. Sin. 24(2), 225–240 (1985).MathSciNet 

    Google Scholar 
    32.Chi, H. et al. Age-Stage, two-sex life table: an introduction to theory, data analysis, and application. Entomol. Generalis 40, 103 (2019).Article 

    Google Scholar 
    33.Hsin, C. Letter to the editor. J. Econ. Entomol. 108, 1465 (2015).Article 

    Google Scholar 
    34.Akköprü, E. P., Atlıhan, R., Okut, H. & Chi, H. Demographic assessment of plant cultivar resistance to insect pests: A case study of the dusky-veined walnut Aphid (Hemiptera: Callaphididae) on five walnut cultivars. J. Econ. Entomol. 108, 378 (2015).Article 

    Google Scholar 
    35.Mousavi, M., Ghosta, Y. & Maroofpour, N. Insecticidal activity and sublethal effects of Beauveria bassiana (Bals.-Criv.) Vuill. isolates and essential oils against Aphis gossypii Glover, 1877 (Hemiptera: Aphididae). Acta Agric. Slovenica. https://doi.org/10.14720/aas.2020.115.2.1306 (2020).Article 

    Google Scholar 
    36.Rahmani, S. & Bandani, A. R. Sublethal concentrations of thiamethoxam adversely affect life table parameters of the aphid predator, Hippodamia variegata (Goeze) (Coleoptera: Coccinellidae). Crop Prot. 54, 168–175. https://doi.org/10.1016/j.cropro.2013.08.002 (2013).CAS 
    Article 

    Google Scholar 
    37.Papachristos, D. P. & Milonas, P. G. Adverse effects of soil applied insecticides on the predatory coccinellid Hippodamia undecimnotata (Coleoptera: Coccinellidae). Biol. Control 47, 77–81. https://doi.org/10.1016/j.biocontrol.2008.06.009 (2008).CAS 
    Article 

    Google Scholar 
    38.Ranjbar, F., Reitz, S., Jalali, M. A., Ziaaddini, M. & Izadi, H. Lethal and sublethal effects of two commercial insecticides on egg parasitoids (Hymenoptera: Scelionidae) of green stink bugs (Hem: Pentatomidae). J. Econ. Entomol. https://doi.org/10.1093/jee/toaa232 (2020).Article 

    Google Scholar 
    39.Zhao, Y. et al. Sublethal concentration of benzothiazole adversely affect development, reproduction and longevity of Bradysia odoriphaga (Diptera: Sciaridae). Phytoparasitica 44, 115–124. https://doi.org/10.1007/s12600-016-0506-5 (2016).CAS 
    Article 

    Google Scholar 
    40.Sani, B., Hamid, G. & Elham, R. Sublethal effects of chlorfenapyr on the life table parameters of two-spotted spider mite, Tetranychus urticae (Acari: Tetranychidae). Syst. Appl. Acarol. 23, 1342 (2018).
    Google Scholar 
    41.Leeuwen, T. V., Pottelberge, S. V. & Tirry, L. Biochemical analysis of a chlorfenapyr-selected resistant strain of Tetranychus urticae Koch. Pest Manage. Sci. 62, 425–433 (2010).Article 

    Google Scholar 
    42.Allen, R. G. & Balin, A. K. Oxidative influence on development and differentiation: An overview of a free radical theory of development. Free Radic. Biol. Med. 6, 631–661 (1989).CAS 
    Article 

    Google Scholar 
    43.Bolter, C. J. & Chefurka, W. Extramitochondrial release of hydrogen peroxide from insect and mouse liver mitochondria using the respiratory inhibitors phosphine, myxothiazol, and antimycin and spectral analysis of inhibited cytochromes. Arch. Biochem. Biophys. 278, 65–72 (1990).CAS 
    Article 

    Google Scholar 
    44.Liu, Y., Wang, C., Qi, S., He, J. & Bai, Y. The sublethal effects of ethiprole on the development, defense mechanisms, and immune pathways of honeybees (Apis mellifera L.). Environ. Geochem. Health. https://doi.org/10.1007/s10653-020-00736-7 (2020).Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    45.Zhang, S. et al. Sublethal effects of triflumezopyrim on biological traits and detoxification enzyme activities in the small brown Planthopper Laodelphax striatellus (Hemiptera: Delphacidae). Front. Physiol. 11, 261. https://doi.org/10.3389/fphys.2020.00261 (2020).Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    46.Ku, C. C., Chiang, F. M., Hsin, C. Y., Yao, Y. E. & Sun, C. N. Glutathione transferase isozymes involved in insecticide resistance of diamondback moth larvae. Pestic. Biochem. Physiol. 50, 191–197 (1994).CAS 
    Article 

    Google Scholar 
    47.Prapanthadara, L., Promtet, N., Koottathep, S., Somboon, P. & Ketterman, A. J. Isoenzymes of glutathione S-transferase from the mosquito Anopheles dirus species B: The purification, partial characterization and interaction with various insecticides. Insect Biochemi. Mol. Biol. 30, 395–403 (2000).CAS 
    Article 

    Google Scholar 
    48.Döker, İ, Kazak, C. & Ay, R. Resistance status and detoxification enzyme activity in ten populations of Panonychus citri (Acari: Tetranychidae) from Turkey. Crop Prot. https://doi.org/10.1016/j.cropro.2020.105488 (2021).Article 

    Google Scholar 
    49.Goel, A., Dani, V. & Dhawan, D. K. Protective effects of zinc on lipid peroxidation, antioxidant enzymes and hepatic histoarchitecture in chlorpyrifos-induced toxicity. Chem. Biol. Interact. 156, 131–140. https://doi.org/10.1016/j.cbi.2005.08.004 (2005).CAS 
    Article 
    PubMed 

    Google Scholar 
    50.Van Leeuwen, T., Van Pottelberge, S. & Tirry, L. Biochemical analysis of a chlorfenapyr-selected resistant strain of Tetranychus urticae Koch. Pest Manage. Sci. 62, 425–433. https://doi.org/10.1002/ps.1183 (2006).CAS 
    Article 

    Google Scholar  More

  • in

    The role of Medieval road operation on cultural landscape transformation

    1.Kaplan, J. O., Krumhardt, K. M. & Zimmermann, N. The prehistoric and preindustrial deforestation of Europe. Quatern. Sci. Rev. 28, 3016–3034. https://doi.org/10.1016/j.quascirev.2009.09.028 (2009).ADS 
    Article 

    Google Scholar 
    2.Słowiński, M. et al. Drought as a stress driver of ecological changes in peatland – A palaeoecological study of peatland development between 3500 BCE and 200 BCE in central Poland. Palaeogeogr. Palaeoclimatol. Palaeoecol. 461, 272–291. https://doi.org/10.1016/j.palaeo.2016.08.038 (2016).Article 

    Google Scholar 
    3.Dietze, E., Słowiński, M., Zawiska, I., Veh, G. & Brauer, A. Multiple drivers of Holocene lake level changes at a lowland lake in northeastern Germany. Boreas 45, 828–845. https://doi.org/10.1111/bor.12190 (2016).Article 

    Google Scholar 
    4.Woodward, C., Shulmeister, J., Larsen, J., Jacobsen, G. E. & Zawadzki, A. Landscape hydrology. The hydrological legacy of deforestation on global wetlands. Science 346, 844–847, doi:https://doi.org/10.1126/science.1260510 (2014).5.Dreibrodt, S., Lubos, C., Terhorst, B., Damm, B. & Bork, H. R. Historical soil erosion by water in Germany: scales and archives, chronology, research perspectives. Quatern. Int. 222, 80–95. https://doi.org/10.1016/j.quaint.2009.06.014 (2010).Article 

    Google Scholar 
    6.Trumbore, S., Brando, P. & Hartmann, H. Forest health and global change. Science 349, 814–818. https://doi.org/10.1126/science.aac6759 (2015).ADS 
    CAS 
    Article 
    PubMed 

    Google Scholar 
    7.Syvitski, J. P. & Kettner, A. Sediment flux and the Anthropocene. Philos. Trans. A Math. Phys. Eng. Sci. 369, 957–975. https://doi.org/10.1098/rsta.2010.0329 (2011).ADS 
    Article 
    PubMed 

    Google Scholar 
    8.McConnell, J. R. et al. Lead pollution recorded in Greenland ice indicates European emissions tracked plagues, wars, and imperial expansion during antiquity. Proc. Natl. Acad. Sci. USA https://doi.org/10.1073/pnas.1721818115 (2018).Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    9.Christian, D. Silk roads or steppe roads? The silk roads in world history. J. World Hist. 11, 1–26. https://doi.org/10.1353/jwh.2000.0004 (2000).Article 

    Google Scholar 
    10.Beck, C. W. The role of the scientist: the amber trade, the chemical analysis of amber, and the determination of Baltic provenience. J. Baltic Stud. 16, 191–199. https://doi.org/10.1080/01629778500000111 (1985).Article 

    Google Scholar 
    11.Gimbutas, M. East Baltic amber in the fourth and third millennia B.C. J. Baltic Stud. 16, 231–256, doi:https://doi.org/10.1080/01629778500000151 (1985).12.de Navarro, J. M. Prehistoric routes between Northern Europe and Italy defined by the amber trade. Geogr J 66, doi:https://doi.org/10.2307/1783003 (1925).13.Szilágyi, M. On the Road: The History and Archaeology of Medieval Communication Networks in East-Central Europe. (Archaeolingua Alapítvány, 2014).14.Schmid, B. V. et al. Climate-driven introduction of the Black Death and successive plague reintroductions into Europe. Proc. Natl. Acad. Sci. USA 112, 3020–3025. https://doi.org/10.1073/pnas.1412887112 (2015).ADS 
    CAS 
    Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    15.Rossignol, S., Kleingärtner, S., Newfield, T., Wehner, D. & Studies, P. I. o. M. Landscapes and societies in medieval Europe east of the Elbe: Interactions between environmental settings and cultural transformations. (Pontifical Institute of Mediaeval Studies, 2013).16.Drager, N. et al. Varve microfacies and varve preservation record of climate change and human impact for the last 6000 years at Lake Tiefer See (NE Germany). The Holocene 27, 450–464. https://doi.org/10.1177/0959683616660173 (2016).ADS 
    Article 

    Google Scholar 
    17.Theuerkauf, M., Drager, N., Kienel, U., Kuparinen, A. & Brauer, A. Effects of changes in land management practices on pollen productivity of open vegetation during the last century derived from varved lake sediments. Holocene 25, 733–744. https://doi.org/10.1177/0959683614567881 (2015).ADS 
    Article 

    Google Scholar 
    18.Izdebski, A., Koloch, G., Słoczyński, T. & Tycner, M. On the use of palynological data in economic history: New methods and an application to agricultural output in Central Europe, 0–2000AD. Explor. Econ. Hist. 59, 17–39. https://doi.org/10.1016/j.eeh.2015.10.003 (2016).Article 

    Google Scholar 
    19.Brauer, A. et al. The importance of independent chronology in integrating records of past climate change for the 60–8 ka INTIMATE time interval. Quatern. Sci. Rev. 106, 47–66. https://doi.org/10.1016/j.quascirev.2014.07.006 (2014).ADS 
    Article 

    Google Scholar 
    20.Ott, F. et al. Site-specific sediment responses to climate change during the last 140 years in three varved lakes in Northern Poland. The Holocene 28, 464–477. https://doi.org/10.1177/0959683617729448 (2017).ADS 
    Article 

    Google Scholar 
    21.Śląski, K. Lądowe szlaki handlowe Pomorza w XI-XIII wieku. Zapiski Historyczne 34 (1969).22.Zakrzewski, I. Vol. 1 (Poznań, 1877).23.Ślaski, K. Zasięg lasów Pomorza w ostatnim tysiącleciu [in English: Range of forests Pomerania in the last millennium]. Przegląd Zachodni 5(6), 207–263 (1951).
    Google Scholar 
    24.Górska-Gołaska, K. in Słownik historyczno-geograficzny ziem polskich w średniowieczu Vol. part 3 (ed A. Gąsiorowski) 89 (1993–1999).25.Szulist, W. Ważniejsze szlaki handlowo-komunikacyjne północno-zachodniego Pomorza Gdańskiego w XVI-XVII w. Zapiski Historyczne 35, 105–106 (1970).
    Google Scholar 
    26.Wilska, M. in Historical atlas of Poland in the 2nd half of the 16th century: voivodeships of Cracow, Sandomierz, Lublin, Sieradz, Łęczyca, Rawa, Płock and Mazovia Vol. 3 (ed M. Słoń) 637 (Peter Lang, 2014).27.Pluskowski, A. The archaeology of the military orders: The material culture of holy war. Mediev. Archaeol. 62, 105–134. https://doi.org/10.1080/00766097.2018.1451590 (2018).Article 

    Google Scholar 
    28.Związek, T. in Wielkopolska w drugiej połowie XVI w. Vol. 2 (eds K. Chłapowski & M. Słoń) 273 (Wydawnictwo Instytutu Historii PAN, 2017).29.Wielopolski, A. Polsko-pomorskie spory graniczne w latach 1536–1555. Przegląd Zachodni 10, 85 (1954).
    Google Scholar 
    30.Kowalkowski, K. Z dziejów gminy Kaliska oraz wsi do niej należących. (Wydawnictwo Region, 2010).31.Słowiński, M. et al. Paleoecological and historical data as an important tool in ecosystem management. J. Environ. Manage. 236, 755–768. https://doi.org/10.1016/j.jenvman.2019.02.002 (2019).Article 
    PubMed 

    Google Scholar 
    32.Błaszkiewicz, M. et al. Climatic and morphological controls on diachronous postglacial lake and river valley evolution in the area of Last Glaciation, northern Poland. Quatern. Sci. Rev. 109, 13–27. https://doi.org/10.1016/j.quascirev.2014.11.023 (2015).Article 

    Google Scholar 
    33.Woś, A. Climate of Poland (Wydawnictwo Naukowe PWN, (in Polish), Warszawa, 1999).34.Haldon, J. et al. History meets palaeoscience: Consilience and collaboration in studying past societal responses to environmental change. Proc. Natl. Acad. Sci. USA https://doi.org/10.1073/pnas.1716912115 (2018).Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    35.Stivrins, N. et al. Palaeoenvironmental evidence for the impact of the crusades on the local and regional environment of medieval (13th–16th century) northern Latvia, eastern Baltic. The Holocene 26, 61–69. https://doi.org/10.1177/0959683615596821 (2015).ADS 
    Article 

    Google Scholar 
    36.Grzegorz, M. Osady Pomorza Gdańskiego w latach 1309–1454. (Państwowe Wydawn. Nauk., 1990).37.Biskup, M. Prusy Królewskie w drugiej połowie XVI wieku. (Państwowe Wydawnictwo Naukowe, 1961).38.Chmielewski, S. Gospodarka rolna i hodowlana w Polsce w XIV i XV w. Technika i rozmiary produkcji, . (Państwowe Wydawnictwo Naukowe, 1962).39.Brown, A. et al. The ecological impact of conquest and colonization on a medieval frontier landscape: combined Palynological and geochemical analysis of lake sediments from Radzyń Chełminski Northern Poland. Geoarchaeology 30, 511–527. https://doi.org/10.1002/gea.21525 (2015).Article 

    Google Scholar 
    40.Brown, A. & Pluskowski, A. Detecting the environmental impact of the Baltic Crusades on a late-medieval (13th–15th century) frontier landscape: palynological analysis from Malbork Castle and hinterland, Northern Poland. J. Archaeol. Sci. 38, 1957–1966. https://doi.org/10.1016/j.jas.2011.04.010 (2011).Article 

    Google Scholar 
    41.Buntgen, U. et al. Filling the Eastern European gap in millennium-long temperature reconstructions. Proc. Natl. Acad. Sci. USA 110, 1773–1778. https://doi.org/10.1073/pnas.1211485110 (2013).ADS 
    CAS 
    Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    42.Luterbacher, J. et al. European summer temperatures since Roman times. Environ. Res. Lett. 11, 024001. https://doi.org/10.1088/1748-9326/11/2/024001 (2016).Article 

    Google Scholar 
    43.Czaja, R. & Radzimiński, A. (eds.), The Teutonic Order in Prussia and Livonia: The political and ecclesiastical Structures 13th–16th century. 407 (TNT, Bohlau Verlag, 2015).44.Bartlett, R. The making of Europe: conquest, colonization and cultural change, 950–1350. (Allen Lane, 1993).45.Broda, J. History of forestry in Poland [in Polish]. 368 (Wydawnictwo Akademii Rolniczej im. Augusta Cieszkowskiego w Poznaniu, 2000).46.Schreg, R. in Processing, storage, distribution of food in medieval rural environment (eds J. Klápště & P. Sommer) 301–320 (2011).47.Kaczmarczyk, Z. & Sczaniecki, M. Kolonizacja na prawie niemieckim w Polsce a rozwój renty feudalnej’. Czasopismo Prawno-Historyczne 3, 59–83 (1951).
    Google Scholar 
    48.Czerwiński, S. et al. Environmental implications of past socioeconomic events in Greater Poland during the last 1200 years. Synthesis of paleoecological and historical data. Quaternary Science Reviews 259, doi:https://doi.org/10.1016/j.quascirev.2021.106902 (2021).49.Schreg, R. in Settlement change across Medieval Europe. Old paradigms and new vistas (eds N. Brady & C. Theune) 61–70 (Sidestone Press, 2019).50.Mikulski, K. Osadnictwo wiejskie województwa pomorskiego od połowy XVI do końca XVII wieku. (1994).51.Biskup, M. & Labuda, G. Dzieje zakonu krzyżackiego w Prusach: gospodarka, społeczeństwo, państwo, ideologia. (Wydawn. Morskie, 1986).52.Kizik, E. in Dżuma, ospa, cholera. W trzechsetną rocznicę wielkiej epidemii na ziemiach Rzeczypospolitej w latach 1708–1711 (ed E. Kizik) 275 (2012).53.Brönnimann, S. Climatic Changes Since 1700. Vol. 55 360 (Springer, 2015).54.Dygdała, J. Lustracja województw Prus Królewskich. Vol. 1–2 (Towarzystwo Naukowe w Toruniu, 2003).55.Dietze, E. et al. Human-induced fire regime shifts during 19th century industrialization: A robust fire regime reconstruction using northern Polish lake sediments. PLoS ONE 14, e0222011. https://doi.org/10.1371/journal.pone.0222011 (2019).CAS 
    Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    56.Lamentowicz, M. et al. Always on the tipping point – A search for signals of past societies and related peatland ecosystem critical transitions during the last 6500 years in N Poland. Quat. Sci. Rev. 225, doi:https://doi.org/10.1016/j.quascirev.2019.105954 (2019).57.Theuerkauf, M., Couwenberg, J., Kuparinen, A. & Liebscher, V. A matter of dispersal: REVEALSinR introduces state-of-the-art dispersal models to quantitative vegetation reconstruction. Veg. Hist. Archaeobotany 25, 541–553. https://doi.org/10.1007/s00334-016-0572-0 (2016).Article 

    Google Scholar 
    58.Hoszowski, S. “Lustracja województwa pomorskiego 1565” (Gdańskie Towarzystwo Naukowe, 1961).59.Hoszowski, S. Lustracja województw Prus Królewskich, 1624, z fragmentami lustracji 1615 roku, Lustracje dóbr królewskich XVI-XVIII wieku (Gdańskie Towarzystwo Naukowe, 1967).60.Metryka Koronna (Crown’s Metric), Central Archives of Historical Records, Warsaw, Poland, vol. XVIII, f. 23261.Ott, F. et al. Constraining the time span between the Early Holocene Hasseldalen and Askja-S Tephras through varve counting in the Lake Czechowskie sediment record Poland. J. Quat. Sci. 31, 103–113. https://doi.org/10.1002/jqs.2844 (2016).Article 

    Google Scholar 
    62.Roeser, P. et al. Advances in understanding calcite varve formation: new insights from a dual lake monitoring approach in the southern Baltic lowlands. Boreas (in press).63.Wulf, S. et al. Holocene tephrostratigraphy of varved sediment records from Lakes Tiefer See (NE Germany) and Czechowskie (N Poland). Quatern. Sci. Rev. 132, 1–14. https://doi.org/10.1016/j.quascirev.2015.11.007 (2016).ADS 
    Article 

    Google Scholar 
    64.Sugita, S. Theory of quantitative reconstruction of vegetation I: pollen from large sites REVEALS regional vegetation composition. Holocene 17, 229–241. https://doi.org/10.1177/0959683607075837 (2007).ADS 
    Article 

    Google Scholar 
    65.Giętkowski, T. Temporal change of forest area in Tuchola Pinewoods region between 1938 and 2000 (in Polish: Zmiany lesistości Borów Tucholskich w latach 1938 – 2000). Promotio Geographica Bydgostiensia IV, 1–12 (2009). More

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    A robust multiple-objective decision-making paradigm based on the water–energy–food security nexus under changing climate uncertainties

    As stated, the primary goal of this study is to promote an objective decision support framework for water resource planning and management purposes within the context of the WEF security nexus, which takes into account the uncertainties imposed by the climate change phenomenon. Such a framework is “robust” since it takes the multi-dimensionality of water-related problems into account while addressing the uncertainties imposed by climate change projections. The basic components of this decision-making paradigm are depicted in Fig. 1. In principle, while this framework is sensitive to the uncertainties associated with the climate change projections, it can provide a dynamic water resources planning and management scheme promoted within the WEF security network. Thus, in addition to the status quo, a series of climate change projections (i.e., RCP 2.6, RCP 4.5, and RCP 8.5) are also integrated into the proposed decision support framework. In essence, the main components of the proposed framework are simulation and operation of the water resources system based on the standard operation policy (SOP), evaluating the system’s efficiency through a series of quantitative performance criteria, and finally, applying the MADM-based framework to opt for a robust system renovation setting.Figure 1Basic components of the robust decision-making paradigm for water resources planning and management.Full size imageSimulating the water resources systemSOP is a primitive, and perhaps the most-well-known real-time operation policy in water resources planning and management14. The core principle here is to minimize the prioritized water shortage at the current time step with no conservation policy (e.g., hedging rules) in place. SOP, as a standard rule curve (RC), determines how the operator should behave at any given state of a reservoir15,16. This rule curve is established as an attempt to balance various water demands including but not limited to flood control, hydropower, water supply, and recreation17. A SOP operating system attempts to release water to meet a water demand at the current time, with no regard to the future.In general, SOP can be mathematically expressed as18:$$R_{t} = left{ {begin{array}{*{20}c} {D_{t} } \ {AW_{t} } \ 0 \ end{array} – S_{min } } right.begin{array}{*{20}c} {} & {if} & {AW_{t} > S_{min } } \ {} & {if} & {AW_{t} > S_{min } } \ {} & {if} & {AW_{t} le S_{min } } \ end{array} begin{array}{*{20}c} {} & {and} & {AW_{t} – S_{min } ge D_{t} } \ {} & {and} & {AW_{t} – S_{min } < D_{t} } \ {} & {} & {} \ end{array} quad t = { 1},{ 2},{ 3}, , ... , ,T$$ (1a) where$$AW_{t} = S_{t} + Q_{t} - Loss_{t}$$ (1b) in which Rt = amount of water supplied during the tth time step; Dt = consumers’ water demand during the tth time step; AWt = amount of available water during the tth time step; St = amount of stored water during the tth time step; Smin = dead storage of the reservoir; Qt = inflow during the tth time step; Losst = net water loss (i.e., precipitation minus evaporation) of the reservoir during the tth time step; and T = total number of time steps in the operational horizon.In practice, however, a different type of water demand leads to a different interpretation of water shortage. There are cases in which the stakeholders’ needs are represented by a set of volumetric demand targets, and the decision-makers’ objective would be to minimize the water deficit based on a set of priorities for these demands. This is a typical case for agricultural, domestic, industrial, and environmental demands. For hydropower generation, however, a conventional interpretation of SOP would be to generate maximum electricity permitted by the power plant capacity (PPC) at each given time step19. For a hydropower system, the amount of water needed to reach a power plant capacity is given by19:$$R_{t} = frac{{86400 times PF times Countday_{t} times PPC}}{{gamma_{w} times g times eta times Delta H_{t} }}$$ (2) in which, γw = water specific weight; g = gravitational acceleration; η = efficiency of the hydropower system; ΔHt = height difference between the reservoir water level and the tailwater level at time step t; Countdayt = number of days within time step t; and PF = plant factor of the hydropower system.As stated earlier, applying an SOP-based plan requires a set of pre-defined priorities to advise decision-makers concerning the order, in which each of these demands is to be met. The major water demands include drinking, industry, environment, agriculture, and hydropower. Thus, according to the SOP’s principle, the decision-makers, first, allocate the available water to meet the demand of the stakeholder with the highest priority (i.e., the domestic and industrial demand). After this first water demand is fully satisfied, the available water can be used for the next demand. Such an allocation process continues until no water is available. It should be noted, however, that if the released water in each stage passes through the penstock equipped with the turbines, electricity can be generated. The amount of energy generated in previous stages must be accounted for before computing the amount of water released for hydropower purposes.Performance criteriaPerformance criteria are, in essence, quantitative measures that can provide a practical insight for the decision-makers regarding the status of a system. This definition covers a broad spectrum of mathematical representations, which can range from simple mathematical formulas such as the average of a specific output to more complex and probability-based entities20,21. The most fundamental and universal probability-based performance criteria are reliability, resiliency, and vulnerability22,23,24. In essence, reliability is the probability of successful function of a system; resiliency measures the probability of successful functioning following a system failure; and vulnerability quantifies the severity of failure during an operation horizon25. It should be noted that these three criteria assess different aspects of a water resources system, and as such, they complement one another26. For more information regarding these probabilistic performance criteria, the readers can refer to Sandoval-Solis et al.27 and Zolghadr-Asli et al.20.In this study, the concept of levelized cost of energy (LCOE) is utilized for economic evaluation. The LCOE of a given hydropower system is the ratio of lifetime costs to lifetime electricity generation, both of which are discounted back to a common year using a discount rate that reflects the average cost of capital28. The LCOE of renewable energy systems depends on the technology, geographic criteria, capital and operating costs, and the efficiency of the system. The LCOE can be mathematically expressed as follows29:$$LCOE = frac{{sumnolimits_{t = 1}^{n} {frac{{I_{t} + M_{t} + F_{t} }}{{left( {1 + r} right)^{t} }}} }}{{sumnolimits_{t = 1}^{n} {frac{{E_{t} }}{{left( {1 + r} right)^{t} }}} }}$$ (3) in which It = investment expenditures in year t; Mt = operation and maintenance expenditures in year t; Ft = fuel expenditures in year t; Et = electricity generation in year t; r = discount rate; and n = economic life expectancy of the system.MADMMADM is an umbrella term to describe a series of frameworks, which aim to help individuals or a group of individuals to prioritize a series of discretely defined alternatives with regard to a set of evaluation attributes30,31. MADM can provide the necessary means to conduct planning and management under changing circumstances such as those under climate change conditions10,32. According to one of the basic principles of MADM, the decision-maker can use the similarity of the feasible alternatives and the preferential result and/or incongruity of the undesirable alternatives. The notion mentioned above is, chiefly, the core principle of the reference-dependent theory33. Accordingly, the reference-based branch of the MADM methods can, itself, be classified into two major groups: screening methods and ranking methods. Screening methods eliminate alternatives that cannot satisfy the pre-determined conditions for the desirable solution, while ranking methods order all the alternatives from the best to the worst34.Pioneered by Hwang and Yoon35, the technique for order references by similarity to an ideal solution (TOPSIS) is a compensatory, objective MADM solving method rooted from the basic principles of the reference-dependent theory. The core idea is that the chosen alternative should have the shortest distance from the ideal solution and the farthest distance from the negative-ideal solution36. The basic computation algorithm of TOPSIS can be summarized as follows37,38:Step I: Construct the original decision matrix (X), where m feasible alternatives are to be evaluated based on n evaluation criteria:$$X = left[ {begin{array}{*{20}c} {x_{11} } & {x_{12} } & cdots & {x_{1n} } \ {x_{21} } & {x_{22} } & cdots & {x_{2n} } \ vdots & vdots & ddots & vdots \ {x_{m1} } & {x_{m2} } & cdots & {x_{mn} } \ end{array} } right]$$ (4) in which xij = the element of the ith alternative concerning the jth criterion.Step II: Defining the reference alternatives [i.e., the ideal solution (s+) and the negative-ideal solution (s−)]. To do so, first, the elements of the decision matrix that are associated with negative criteria must be redefined by using the following equation:$$x_{ij}^{ * } = frac{1}{{x_{ij} }}$$ (5a) The elements of the decision matrix that are associated with positive criteria would remain the same:$$x_{ij}^{ * } = x_{ij}$$ (5b) The ideal alternative is an arbitrarily defined vector, which describes the aspired solution to the given problem, while the inferior alternative is an arbitrarily defined solution that represents the most undesirable option for the given MADM problem. Here, the ideal and negative-ideal solutions would be represented with two separate vectors where each pair of the corresponding elements in these vectors is, respectively, the maximum and minimum values of (x_{ij}^{ * }) with regard to each of the evaluation criteria.Step III: Each element of the decision matrix should be normalized by using the following equation:$$p_{ij} = frac{{x_{ij}^{ * } }}{{sqrt {sumnolimits_{i = 1}^{m} {x_{ij}^{ * 2} } } }}$$ (6) in which pij = the normalized performance value for the ith alternative with respect to the jth criterion.Step IV: The weighted normalized preference value (zij) can be computed as follows:$$z_{ij} = p_{ij} times w_{j} quad forall i,j$$ (7) in which wj = the weight (i.e., the importance value) of the jth criterion. The weights assigned to the evolution criteria reflect their relative importance to the decision-makers. The higher the weights are, the more crucial their roles would be in the selection process. Chiefly, these weighting mechanisms are either subjective in nature or follow an objective procedure. In the subjective approaches, the weights of the attributes are assigned based on the performance information given by the decision-maker, whereas in the objective approaches, the weights of the evaluation attributes would be obtained by using the objective information extracted from the decision matrix39. Shannon’s Entropy method, used in this study as the weight assignment mechanism, is a well-known objective weighting technique40. This method tends to assign the highest weight to an evaluation attribute with the highest dispersity in its values. For more information on the computational framework of this method, the readers can refer to Lotfi and Fallahnejad41.Step V: In this step, every given alternative is compared to the reference points, namely, the ideal and inferior alternatives. The described procedure, which is known as the separation measurement in TOPSIS, can be mathematically expressed as follows35:$$D_{i}^{ + } = sqrt {sumlimits_{j = 1}^{n} {left( {z_{ij} - z_{j}^{ + } } right)^{2} } }$$ (8) And$$D_{i}^{ - } = sqrt {sumlimits_{j = 1}^{n} {left( {z_{ij} - z_{j}^{ - } } right)^{2} } }$$ (9) in which (D_{j}^{ + }) and (D_{j}^{ - }) = separation measurements of the jth criterion with respect to the ideal and inferior alternatives, respectively.Step VI: The relative closeness to the ideal solution (χi), which can be used to rank the desirability of the feasible alternative, can be computed as follows35:$$chi_{i} = frac{{D_{i}^{ - } }}{{D_{i}^{ + } + D_{i}^{ - } }}quad forall i$$ (10) The further this distance (i.e., larger values of χi), the more desirable the alternative would be.Robust multi-attribute frameworkAs stated, each climate change scenario depicts a unique future with regard to the changing climate, which in turn introduces an element of uncertainty to the projected performance of water resources systems during their operation horizon. Furthermore, downscaling methods, which link these projected changes in the global climatic pattern to a local or regional scale, can be another source of uncertainty. Naturally, for long-lasting water infrastructure such as a hydropower system, addressing these uncertainties in a proper and timely manner can be one of the key components of a robust project. Thus, this study aims to not only evaluate the system’s performance under the status quo but also assess the credibility of the system under the projected climate change conditions.The other characteristic one might expect from a robust project is its ability to take into account the multi-dimensionality nature of water-related infrastructure. Most notably, addressing the WEF security nexus must be a priority in water resources planning and management. Resultantly, any robust decision-making paradigm for water resources planning and management purposes should also account for the other pillars of the WEF nexus (i.e., energy and food sectors), as they would be consequentially affected by such decisions. It is also important to note that these sectors could be affected by the climate change phenomenon. The other crucial feature of a robust decision-making paradigm is that it should be able to account for the socio-economic, environmental, and technical factors that determine the overall quality of the project. Such a decision-making paradigm is depicted in Fig. 2. This notion in practice, however, can typically lead to a mega decision matrix composed of numerous criteria and alternatives that can be overwhelming if the subjective MADM methods are to be employed. This study, thus, employs an objective MADM framework (i.e., TOPSIS/Entropy) to help overcome the above-described problem. The basic idea is to promote a universal and practical decision support framework that enables the water resources planners and managers to account for the intricacies of the WEF security nexus while simultaneously taking the uncertainties of climate change projections into account. Figure 3 illustrates the flowchart of the proposed decision support framework.Figure 2Schematic diagram of the MADM problem.Full size imageFigure 3Flowchart of the proposed framework.Full size image More