More stories

  • in

    Energy efficiency and biological interactions define the core microbiome of deep oligotrophic groundwater

    Fennoscandian shield genomic database (FSGD)The Fennoscandian Shield bedrock contains an abundance of fracture zones with different groundwater characteristics that vary in water source, retention time, chemistry, and connectivity to surface-fed organic compounds (see Supplementary Data 1). The Äspö Hard Rock Laboratory (HRL) and Olkiluoto drillholes were sampled over time, covering a diversity of aquifers representing waters of differing ages and both planktonic and biofilm-associated communities. In order to provide a genome-resolved view of the Fennoscandian Shield bedrock Archaeal and bacterial communities, collected samples were used for an integrated analysis by combining metagenomes (n = 44), single-cell genomes (n = 564), and metatranscriptomes (n = 9) (see detailed statistics for the generated datasets in the Supplementary Data 1 and Supplementary Information). Assembly and binning of the 44 metagenomes (~1.3 TB sequenced data) resulted in the reconstruction of 1278 metagenome-assembled genomes (MAGs; ≥ 50% completeness and ≤ 5% contamination). By augmenting this dataset with 564 sequenced single-cell amplified genomes (SAGs; 114 of which were ≥ 50% complete with ≤ 5% contamination), we present a comprehensive genomic database for the archaeal and bacterial diversity of these oligotrophic deep groundwaters, hereafter referred to as the Fennoscandian Shield genomic database (FSGD; statistics in Fig. 1A & Supplementary Data 2). Phylogenomic reconstruction using reference genomes in the Genome Taxonomy Database (GTDB-TK; release 86) shows that the FSGD MAGs/SAGs span most branches on the prokaryotic tree of life (Fig. 2). Harboring representatives from 53 phyla (152 archaeal MAGs/SAGs in 7 phyla and 1240 bacterial MAGs/SAGs in 46 phyla), the FSGD highlights the remarkable diversity of these oligotrophic deep groundwaters. Apart from the exceptional case of a single-species ecosystem composed of ‘Candidatus Desulforudis audaxviator’ in the fracture fluids of an African gold mine17, other studies of deep groundwaters as well as aquifer sediments have also revealed a notable phylogenetic diversity of the Archaea and Bacteria10,11,18. For example, metagenomic and single-cell genomic analysis of the CO2-driven Crystal geyser (Colorado Plateau, Utah, USA) resulted in reconstructed genomes of 503 archaeal and bacterial species distributed across 104 different phylum-level lineages11.Fig. 1: Overview of the FSGD MAGs and SAGs.Statistics of the metagenome-assembled genomes (MAGs) and single-cell amplified genomes (SAGs) of the Fennoscandian Shield Genomic Database (a). The number of genome clusters present in borehole samples (centerline, median; hinge limits, 25 and 75% quartiles; whiskers, 1.5x interquartile range; points, outliers). Numbers on top of each box plot represent the number of metagenomes generated for borehole samples (b). NMDS plot of unweighted binary Jaccard beta-diversities of presence/absence of all FSGD reconstructed MAGs/SAGs (c) and MAG and SAG clusters belonging to the common core microbiome present in both Äspö HRL and Olkiluoto (d). Numbers in the parenthesis show the number of overlapping points. The data used to generate these plots are available in Supplementary Data 4 and the Source Data.Full size imageFig. 2: Phylogenetic diversity of reconstructed MAGs and SAGs of the fennoscandian shield genomic database (FSGD).Genomes present in genome taxonomy database (GTDB) release 86 were used as reference. Archaea and Bacteria phylogenies are represented separately in the top and bottom panels, respectively. MAGs and SAGs of the FSGD are highlighted in red. Legend in front of each number at the bottom of the figure shows the list of taxa in the tree that are marked with the same number.Full size imageClustering reconstructed FSGD MAGs/SAGs into operationally defined prokaryotic species (≥ 95% average nucleotide identity (ANI) and ≥ 70% coverage) produced 598 genome clusters. Based on the GTDB-TK affiliated taxonomy, a single FSGD cluster may represent a novel phylum, whereas at the lower taxonomic levels, the FSGD harbors genome clusters representing seven novel taxa at class, 58 at order, 123 at family, and 345 at the genus levels. In addition, more than 94% of the reconstructed MAGs/SAGs clusters (n = 568) represent novel species with no existing representative in public databases (Supplementary Data 2). Mapping metagenomic reads against genome clusters represented exclusively by SAGs (n = 38, Fig. 1A) revealed that 14 genome clusters (20 SAGs) were not detectable in the metagenomes, suggesting they might represent rare species in the microbial community of the investigated deep groundwaters (Supplementary Data 3).To explore the community composition of different groundwaters and their temporal dynamics, presence/absence patterns were computed by competitively mapping the metagenomics reads against all reconstructed MAGs/SAGs of the FSGD. Contigs were discarded from the mapping results if More

  • in

    The Welwitschia genome reveals a unique biology underpinning extreme longevity in deserts

    1.Jürgens, N., Oncken, I., Oldeland, J., Gunter, F. & Rudolph, B. Welwitschia: phylogeography of a living fossil, diversified within a desert refuge. Sci. Rep. 11, 2385 (2021).ADS 
    PubMed 
    PubMed Central 
    Article 
    CAS 

    Google Scholar 
    2.Herre, H. The age of Welwitschia bainesii (Hook. f) Cearr.: C14 research. S. Afr. J. Bot. 27, 139–140 (1961).
    Google Scholar 
    3.Bornman, C. H. Welwitschia mirabilis: structural and functional anomalies. Madoqua 10, 21–31 (1977).
    Google Scholar 
    4.Talalaj, S., Talalaj, D. & Talalaj, J. The strangest plants in the world. (Hill of Content, 1991).5.Hooker, J. I. On Welwitschia, a new genus of Gnetaceæ. Trans. Linn. Soc. Lond. 24, 1–48 (1862).Article 

    Google Scholar 
    6.Friedman, W. E. Development and evolution of the female gametophyte and fertilization process in Welwitschia mirabilis (Welwitschiaceae). Am. J. Bot. 102, 312–324 (2015).PubMed 
    Article 

    Google Scholar 
    7.Leebens-Mack, J. H. et al. One thousand plant transcriptomes and the phylogenomics of green plants. Nature 574, 679–685 (2019).Article 
    CAS 

    Google Scholar 
    8.Dilcher, D. L., Bernardes-De-Oliveira, M. E. & Pons, D. Welwitschiaceae from the lower Cretaceous of northeastern Brazil. Am. J. Bot. 92, 1294–1310 (2005).PubMed 
    Article 

    Google Scholar 
    9.Wickett, N. J. et al. Phylotranscriptomic analysis of the origin and early diversification of land plants. Proc. Natl Acad. Sci. USA 111, E4859 (2014).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    10.Li, Z. et al. Single-copy genes as molecular markers for phylogenomic studies in seed plants. Genome Biol. Evol. 9, 1130–1147 (2017).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    11.Doyle, J. A. Molecular and fossil evidence on the origin of angiosperms. Annu. Rev. Earth Planet. Sci. 40, 301–326 (2012).ADS 
    CAS 
    Article 

    Google Scholar 
    12.Bateman, R. Hunting the Snark: the flawed search for mythical Jurassic angiosperms. J. Exp. Bot. 71, 22–35 (2019).Article 
    CAS 

    Google Scholar 
    13.Wan, T. et al. A genome for gnetophytes and early evolution of seed plants. Nat. Plants 4, 82–89 (2018).CAS 
    PubMed 
    Article 

    Google Scholar 
    14.Leitch, I. J., Hanson, L., Winfield, M., Parker, J. & Bennett, M. D. Nuclear DNA C-values complete familial representation in gymnosperms. Ann. Bot. 88, 843–849 (2001).CAS 
    Article 

    Google Scholar 
    15.Khoshoo, T. N. & Ahuja, M. R. The chromosomes and relationships of Welwitschia mirabilis. Chromosoma 14, 522–533 (1963).Article 

    Google Scholar 
    16.Li, Z. et al. Early genome duplications in conifers and other seed plants. Sci. Adv. 1, e1501084 (2015).ADS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    17.Van de Peer, Y. Computational approaches to unveiling ancient genome duplications. Nat. Rev. Genet 5, 752–763 (2004).PubMed 
    Article 
    CAS 

    Google Scholar 
    18.Zhang, Q.-J. et al. The chromosome-level reference genome of tea tree unveils recent bursts of non-autonomous LTR retrotransposons to drive genome size evolution. Mol. Plant 13, 935–938 (2020).CAS 
    PubMed 
    Article 

    Google Scholar 
    19.Zhang, Q. J. & Gao, L. Z. Rapid and recent evolution of LTR retrotransposons drives rice genome evolution during the speciation of AA-genome Oryza species. G3 (Bethesda, Md.) 7, 1875–1885 (2017).CAS 
    Article 

    Google Scholar 
    20.Cossu, R. M. et al. LTR retrotransposons show low levels of unequal recombination and high rates of intraelement gene conversion in large plant genomes. Genome Biol. Evol. 9, 3449–3462 (2017).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    21.Roddy, A. et al. The scaling of genome size and cell size limits maximum rates of photosynthesis with implications for ecological strategies. Int. J. Plant. Sci. https://doi.org/10.1101/619585 (2019).22.Ausin, I. et al. DNA methylome of the 20-gigabase Norway spruce genome. Proc. Natl Acad. Sci. USA 113, E8106–e8113 (2016).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    23.Takuno, S., Ran, J.-H. & Gaut, B. S. Evolutionary patterns of genic DNA methylation vary across land plants. Nat. Plants 2, 15222 (2016).CAS 
    PubMed 
    Article 

    Google Scholar 
    24.Niederhuth, C. E. et al. Widespread natural variation of DNA methylation within angiosperms. Genome Biol. 17, 194 (2016).PubMed 
    PubMed Central 
    Article 
    CAS 

    Google Scholar 
    25.Zhang, X. et al. Genome-wide high-resolution mapping and functional analysis of DNA methylation in Arabidopsis. Cell 126, 1189–1201 (2006).CAS 
    PubMed 
    Article 

    Google Scholar 
    26.Matzke, M. A., Kanno, T. & Matzke, A. J. M. RNA-Directed DNA methylation: the evolution of a complex epigenetic pathway in flowering plants. Annu. Rev. Plant Biol. 66, 243–267 (2015).CAS 
    PubMed 
    Article 

    Google Scholar 
    27.Johnsen, Ø. et al. Climatic adaptation in Picea abies progenies is affected by the temperature during zygotic embryogenesis and seed maturation. Plant Cell Environ. 28, 1090–1102 (2005).CAS 
    Article 

    Google Scholar 
    28.Yakovlev, I. A., Carneros, E., Lee, Y., Olsen, J. E. & Fossdal, C. G. Transcriptional profiling of epigenetic regulators in somatic embryos during temperature induced formation of an epigenetic memory in Norway spruce. Planta 243, 1237–1249 (2016).CAS 
    PubMed 
    Article 

    Google Scholar 
    29.Trávníček, P. et al. Diversity in genome size and GC content shows adaptive potential in orchids and is closely linked to partial endoreplication, plant life-history traits and climatic conditions. N. Phytol. 224, 1642–1656 (2019).Article 
    CAS 

    Google Scholar 
    30.Cacciò, S. et al. Methylation patterns in the isochores of vertebrate genomes. Gene 205, 119–124 (1997).PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    31.Serres-Giardi, L., Belkhir, K., David, J. & Glémin, S. Patterns and evolution of nucleotide landscapes in seed plants. Plant Cell 24, 1379–1397 (2012).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    32.Ossowski, S. et al. The rate and molecular spectrum of spontaneous mutations in Arabidopsis thaliana. Science 327, 92–94 (2010).ADS 
    CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    33.Glémin, S. Surprising fitness consequences of GC-biased gene conversion: I. Mutation load and inbreeding depression. Genetics 185, 939–959 (2010).PubMed 
    PubMed Central 
    Article 
    CAS 

    Google Scholar 
    34.Vinogradov, A. E. DNA helix: the importance of being GC-rich. Nucleic Acids Res. 31, 1838–1844 (2003).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    35.Rocha, E. P. & Danchin, A. Base composition bias might result from competition for metabolic resources. Trends Genet. 18, 291–294 (2002).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    36.Shenhav, L. & Zeevi, D. Resource conservation manifests in the genetic code. Science 370, 683–687 (2020).ADS 
    CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    37.Kelly, S. The amount of nitrogen used for photosynthesis modulates molecular evolution in plants. Mol. Biol. Evol. 35, 1616–1625 (2018).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    38.Martens, P. Welwitschia mirabilis and neoteny. Am. J. Bot. 64, 916–920 (1977).Article 

    Google Scholar 
    39.Robert, J. R. Leaf anatomy of Welwitschia. i. Early development of the leaf. Am. J. Bot. 45, 90–95 (1958).Article 

    Google Scholar 
    40.Bornman, C. H. Welwitschia mirabilis: paradox of the Namib Desert. Endeavour 31, 95–99 (1972).
    Google Scholar 
    41.Pham, T. & Sinha, N. Role of KNOX genes in shoot development of Welwitschia mirabilis. Int. J. Plant Sci. 164, 333–343 (2003).CAS 
    Article 

    Google Scholar 
    42.Nishii, K. et al. A complex case of simple leaves: indeterminate leaves co-express ARP and KNOX1 genes. Dev. Genes Evol. 220, 25–40 (2010).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    43.Hacham, Y. et al. Brassinosteroid perception in the epidermis controls root meristem size. Dev. (Camb., Engl.) 138, 839–848 (2011).CAS 
    Article 

    Google Scholar 
    44.Sun, S. et al. Brassinosteroid signalling regulates leaf erectness in Oryza sativa via the control of a specific U-type cyclin and cell proliferation. Dev. Cell 34, 220–228 (2015).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    45.Wei, Z. & Li, J. Brassinosteroids regulate root growth, development, and symbiosis. Mol. Plant 9, 86–100 (2016).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    46.Jiang, C. K. & Rao, G. Y. Insights into the diversification and evolution of R2R3-MYB transcription factors in plants. Plant Physiol. 183, 637–655 (2020).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    47.Dubos, C. et al. MYB transcription factors in Arabidopsis. Trends Plant Sci. 15, 573–581 (2010).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    48.Pandey, A., Misra, P. & Trivedi, P. K. Constitutive expression of Arabidopsis MYB transcription factor, AtMYB11, in tobacco modulates flavonoid biosynthesis in favor of flavonol accumulation. Plant Cell Rep. 34, 1515–1528 (2015).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    49.Petroni, K. et al. The AtMYB11 gene from Arabidopsis is expressed in meristematic cells and modulates growth in planta and organogenesis in vitro. J. Exp. Bot. 59, 1201–1213 (2008).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    50.Gugger, P. F., Peñaloza-Ramírez, J. M., Wright, J. W. & Sork, V. L. Whole-transcriptome response to water stress in a California endemic oak, Quercus lobata. Tree Physiol. 37, 632–644 (2017).CAS 
    PubMed 
    PubMed Central 

    Google Scholar 
    51.Plomion, C. et al. Oak genome reveals facets of long lifespan. Nat. Plants 4, 440–452 (2018).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    52.Jaiwal, S. K. C. A., Mahajan, S., Kumar, S. & Sharma, V. K. The genome sequence of Aloe vera reveals adaptive evolution of drought tolerance mechanisms. iScience 24, 102078 (2021).ADS 
    Article 

    Google Scholar 
    53.Henschel, J. R. & Seely, M. K. Long-term growth patterns of Welwitschia mirabilis, a long-lived plant of the Namib desert (including a bibliography). Plant Ecol. 150, 7–26 (2000).Article 

    Google Scholar 
    54.Stortenbeker, N. & Bemer, M. The SAUR gene family: the plant’s toolbox for adaptation of growth and development. J. Exp. Bot. 70, 17–27 (2019).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    55.Wei, J. et al. The E3 ligase AtCHIP positively regulates Clp proteolytic subunit homeostasis. J. Exp. Bot. 66, 5809–5820 (2015).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    56.Olinares, P. D., Kim, J., Davis, J. I. & van Wijk, K. J. Subunit stoichiometry, evolution, and functional implications of an asymmetric plant plastid ClpP/R protease complex in Arabidopsis. Plant Cell 23, 2348–2361 (2011).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    57.Sjögren, L. L., Stanne, T. M., Zheng, B., Sutinen, S. & Clarke, A. K. Structural and functional insights into the chloroplast ATP-dependent Clp protease in Arabidopsis. Plant Cell 18, 2635–2649 (2006).PubMed 
    PubMed Central 
    Article 
    CAS 

    Google Scholar 
    58.Dong, H. et al. A rice virescent-yellow leaf mutant reveals new insights into the role and assembly of plastid caseinolytic protease in higher plants. Plant Physiol. 162, 1867–1880 (2013).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    59.Nakabayashi, K., Ito, M., Kiyosue, T., Shinozaki, K. & Watanabe, A. Identification of clp genes expressed in senescing Arabidopsis leaves. Plant cell Physiol. 40, 504–514 (1999).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    60.Koussevitzky, S. et al. An Arabidopsis thaliana virescent mutant reveals a role for ClpR1 in plastid development. Plant Mol. Biol. 63, 85–96 (2007).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    61.Vierling, E. The roles of heat shock proteins in plants. Annu. Rev. Plant Physiol. Plant Mol. Biol. 42, 579–620 (1991).CAS 
    Article 

    Google Scholar 
    62.Guo, L. M., Li, J., He, J., Liu, H. & Zhang, H. M. A class I cytosolic HSP20 of rice enhances heat and salt tolerance in different organisms. Sci. Rep. 10, 1383 (2020).ADS 
    CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    63.Waseem, M., Rong, X. & Li, Z. Dissecting the role of a basic helix-loop-helix transcription factor, SlbHLH22, under salt and drought stresses in transgenic Solanum lycopersicum L. Front. Plant Sci. 10, 734 (2019).PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    64.De La Torre, A. R., Lin, Y. C., Van de Peer, Y. & Ingvarsson, P. K. Genome-wide analysis reveals diverged patterns of codon bias, gene expression, and rates of sequence evolution in Picea gene families. Genome Biol. Evol. 7, 1002–1015 (2015).Article 
    CAS 

    Google Scholar 
    65.Neale, D. B., Martínez-García, P. J., De La Torre, A. R., Montanari, S. & Wei, X. X. Novel insights into tree biology and genome evolution as revealed through genomics. Annu. Rev. Plant Biol. 68, 457–483 (2017).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    66.Nakashima, K., Yamaguchi-Shinozaki, K. & Shinozaki, K. The transcriptional regulatory network in the drought response and its crosstalk in abiotic stress responses including drought, cold, and heat. Front. Plant Sci. 5, 170 (2014).PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    67.Jiang, F. et al. The apricot (Prunus armeniaca L.) genome elucidates Rosaceae evolution and beta-carotenoid synthesis. Hortic. Res. 6, 128 (2019).PubMed 
    PubMed Central 
    Article 
    CAS 

    Google Scholar 
    68.Huo, H., Dahal, P., Kunusoth, K., McCallum, C. M. & Bradford, K. J. Expression of 9-cis-EPOXYCAROTENOID DIOXYGENASE4 is essential for thermoinhibition of lettuce seed germination but not for seed development or stress tolerance. Plant Cell 25, 884–900 (2013).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    69.Wang, H. et al. CG gene body DNA methylation changes and evolution of duplicated genes in cassava. Proc. Natl Acad. Sci. USA 112, 13729–13734 (2015).ADS 
    CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    70.Xu, J. et al. Single-base methylome analysis reveals dynamic epigenomic differences associated with water deficit in apple. Plant Biotechnol. J. 16, 672–687 (2018).CAS 
    PubMed 
    Article 

    Google Scholar 
    71.Friis, E. M., Pedersen, K. R. & Crane, P. R. Welwitschioid diversity in the early Cretaceous: evidence from fossil seeds with pollen from Portugal and eastern North America. Grana 53, 175–196 (2014).Article 

    Google Scholar 
    72.Damme, P. V. & Vernemmen, P. The natural environment of the Namib Desert. Afr. Focus 7, 355–400 (1992).
    Google Scholar 
    73.Siesser, W. G. Late Miocene origin of the Benguela upswelling system off northern Namibia. Science 4441, 283–285 (1980).ADS 
    Article 

    Google Scholar 
    74.Meyers, P. A., Brassell, S. C., Huc, A. Y., Barron, E. J. & Stradner, H. Organic geochemistry of sediments recovered by DSDP/IPOD Leg 75 from under the Benguela current. Volume 10, pp.14. (Plenum Press, 1983).75.Alzohairy, A. M., Yousef, M. A., Edris, S., Kerti, B. & Alzohairy, M. Detection of LTR retrotransposons reactivation induced by in vitro environmental stresses in barley (Hordeum vulgare) via RT-qPCR. Life Sci. J. 9, 5019–5026 (2012).
    Google Scholar 
    76.Morano, A. et al. Targeted DNA methylation by homology-directed repair in mammalian cells. Transcription reshapes methylation on the repaired gene. Nucleic Acids Res. 42, 804–821 (2014).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    77.Russo, G. et al. DNA damage and repair modify DNA methylation and chromatin domain of the targeted locus: mechanism of allele methylation polymorphism. Sci. Rep. 6, 33222 (2016).ADS 
    CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    78.Doerfler, W. The almost-forgotten fifth nucleotide in DNA: an introduction. Curr. Top. Microbiol. Immunol. 301, 3–18 (2006).CAS 
    PubMed 
    PubMed Central 

    Google Scholar 
    79.Nystedt, B. et al. The Norway spruce genome sequence and conifer genome evolution. Nature 497, 579–584 (2013).ADS 
    CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    80.Guignard, M. et al. Impacts of nitrogen and phosphorus: from genomes to natural ecosystems and agriculture. Front. Ecol. Evol. 5, 70 (2017).Article 

    Google Scholar 
    81.Drake, P. L., Froend, R. H. & Franks, P. J. Smaller, faster stomata: scaling of stomatal size, rate of response, and stomatal conductance. J. Exp. Bot. 64, 495–505 (2013).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    82.Massmann, U. Welwitschia: nach 90 jahren. Namib. und Meer 7, 45–46 (1976).
    Google Scholar 
    83.Ruan, J. & Li, H. Fast and accurate long-read assembly with wtdbg2. Nat. Methods 17, 155–158 (2020).CAS 
    PubMed 
    Article 

    Google Scholar 
    84.Hu, J., Fan, J., Sun, Z. & Liu, S. NextPolish: a fast and efficient genome polishing tool for long-read assembly. Bioinformatics 36, 2253–2255 (2020).CAS 
    PubMed 
    Article 

    Google Scholar 
    85.Li, H. et al. The sequence alignment/map format and SAMtools. Bioinformatics 25, 2078–2079 (2009).PubMed 
    PubMed Central 
    Article 
    CAS 

    Google Scholar 
    86.Xu, Z. & Wang, H. LTR_FINDER: an efficient tool for the prediction of full-length LTR retrotransposons. Nucleic Acids Res. 35, 265–268 (2007).Article 

    Google Scholar 
    87.Edgar, R. C. & Myers, E. W. PILER: identification and classification of genomic repeats. Bioinformatics 21, 152–158 (2005).Article 

    Google Scholar 
    88.Price, A. L., Jones, N. C. & Pevzner, P. A. De novo identification of repeat families in large genomes. Bioinformatics 21, 351–358 (2005).Article 

    Google Scholar 
    89.Benson, G. Tandem repeats finder: a program to analyze DNA sequences. Nucleic Acids Res. 27, 573–580 (1999).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    90.Stanke, M., Diekhans, M., Baertsch, R. & Haussler, D. Using native and syntenically mapped cDNA alignments to improve de novo gene finding. Bioinformatics 24, 637–644 (2008).CAS 
    PubMed 
    Article 

    Google Scholar 
    91.Korf, I. Gene finding in novel genomes. BMC Bioinformatics 5, 59 (2004).PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    92.Burge, C. & Karlin, S. Prediction of complete gene structures in human genomic DNA. J. Mol. Biol. 268, 78–94 (1997).CAS 
    Article 

    Google Scholar 
    93.Keilwagen, J. et al. Using intron position conservation for homology-based gene prediction. Nucleic Acids Res. 44, 89 (2016).Article 
    CAS 

    Google Scholar 
    94.Trapnell, C., Pachter, L. & Salzberg, S. L. TopHat: discovering splice junctions with RNA-Seq. Bioinformatics 25, 1105–1111 (2009).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    95.Haas, B. J. et al. Improving the Arabidopsis genome annotation using maximal transcript alignment assemblies. Nucleic Acids Res. 31, 5654–5666 (2003).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    96.Haas, B. J. et al. Automated eukaryotic gene structure annotation using EVidenceModeler and the program to assemble spliced alignments. Genome Biol. 9, 7 (2008).Article 
    CAS 

    Google Scholar 
    97.Mitchell, A. L. et al. InterPro in 2019: improving coverage, classification and access to protein sequence annotations. Nucleic Acids Res. 47, 351–360 (2019).Article 
    CAS 

    Google Scholar 
    98.Vanneste, K., Van de Peer, Y. & Maere, S. Inference of genome duplications from age distributions revisited. Mol. Biol. Evol. 30, 177–190 (2013).CAS 
    PubMed 
    Article 

    Google Scholar 
    99.Enright, A. J., Van Dongen, S. & Ouzounis, C. A. An efficient algorithm for large-scale detection of protein families. Nucleic Acids Res. 30, 1575–1584 (2002).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    100.Edgar, R. C. MUSCLE: multiple sequence alignment with high accuracy and high throughput. Nucleic Acids Res. 32, 1792–1797 (2004).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    101.Goldman, N. & Yang, Z. A codon-based model of nucleotide substitution for protein-coding DNA sequences. Mol. Biol. Evol. 11, 725–736 (1994).CAS 
    PubMed 

    Google Scholar 
    102.Yang, Z. PAML 4: phylogenetic analysis by maximum likelihood. Mol. Biol. Evolution. 24, 1586–1591 (2007).CAS 
    Article 

    Google Scholar 
    103.Guindon, S. et al. New algorithms and methods to estimate maximum-likelihood phylogenies: assessing the performance of PhyML 3.0. Syst. Biol. 59, 307–321 (2010).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    104.Proost, S. et al. i-ADHoRe 3.0–fast and sensitive detection of genomic homology in extremely large data sets. Nucleic Acids Res. 40, 11 (2012).Article 
    CAS 

    Google Scholar 
    105.Fostier, J. et al. A greedy, graph-based algorithm for the alignment of multiple homologous gene lists. Bioinformatics 27, 749–756 (2011).CAS 
    PubMed 
    Article 

    Google Scholar 
    106.Putnam, N. H. et al. Sea anemone genome reveals ancestral eumetazoan gene repertoire and genomic organization. Science 317, 86–94 (2007).ADS 
    CAS 
    PubMed 
    Article 

    Google Scholar 
    107.Gu, Z., Gu, L., Eils, R., Schlesner, M. & Brors, B. Circlize Implements and enhances circular visualization in R. Bioinformatics 30, 2811–2812 (2014).CAS 
    PubMed 
    Article 

    Google Scholar 
    108.Guy, L., Kultima, J. R. & Andersson, S. G. genoPlotR: comparative gene and genome visualization in R. Bioinformatics 26, 2334–2335 (2010).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    109.Moreno-Hagelsieb, G. & Latimer, K. Choosing BLAST options for better detection of orthologs as reciprocal best hits. Bioinformatics 24, 319–324 (2008).CAS 
    PubMed 
    Article 

    Google Scholar 
    110.Vanneste, K., Baele, G., Maere, S. & Van de Peer, Y. Analysis of 41 plant genomes supports a wave of successful genome duplications in association with the Cretaceous-Paleogene boundary. Genome Res. 24, 1334–1347 (2014).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    111.Ostlund, G. et al. InParanoid 7: new algorithms and tools for eukaryotic orthology analysis. Nucleic Acids Res. 38, D196–D203 (2010).PubMed 
    Article 
    CAS 

    Google Scholar 
    112.D’Hont, A. et al. The banana (Musa acuminata) genome and the evolution of monocotyledonous plants. Nature 488, 213–217 (2012).ADS 
    PubMed 
    Article 
    CAS 

    Google Scholar 
    113.Drummond, A. J., Suchard, M. A., Xie, D. & Rambaut, A. Bayesian phylogenetics with BEAUti and the BEAST 1.7. Mol. Biol. Evol. 29, 1969–1973 (2012).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    114.Group, A. P. An update of the angiosperm phylogeny group classification for the orders and families of flowering plants: APG IV. Bot. J. Linn. Soc. 181, 1–20 (2016).Article 

    Google Scholar 
    115.Gandolfo, M., Nixon, K. & Crepet, W. A new fossil flower from the Turonian of New Jersey: Dressiantha bicarpellata gen. et sp. nov. (Ceapparales). Am. J. Bot. 85, 964 (1998).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    116.Beilstein, M. A., Nagalingum, N. S., Clements, M. D., Manchester, S. R. & Mathews, S. Dated molecular phylogenies indicate a Miocene origin for Arabidopsis thaliana. Proc. Natl Acad. Sci. USA 107, 18724–18728 (2010).ADS 
    CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    117.Crepet, W. & Nixon, K. Fossil Clusiaceae from the late Cretaceous (Turonian) of new Jersey and implications regarding the history of bee pollination. Am. J. Bot. 85, 1122 (1998).CAS 
    PubMed 
    Article 

    Google Scholar 
    118.Xi, Z. et al. Phylogenomics and a posteriori data partitioning resolve the Cretaceous angiosperm radiation Malpighiales. Proc. Natl Acad. Sci. USA 109, 17519–17524 (2012).ADS 
    CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    119.Friis, E. M. Spirematospermum chandlerae sp. nov., an extinct species of Zingiberaceae from the North American Cretaceous. Tert. Res. 9, 7–12 (1988).
    Google Scholar 
    120.Janssen, T. & Bremer, K. The age of major monocot groups inferred from 800+rbcL sequences. Bot. J. Linn. Soc. 146, 385–398 (2004).Article 

    Google Scholar 
    121.Doyle, J. A. Early evolution of angiosperm pollen as inferred from molecular and morphological phylogenetic analyses. Grana 44, 227–251 (2005).Article 

    Google Scholar 
    122.Rydin, C., Pedersen, K. R. & Friis, E. M. On the evolutionary history of Ephedra: cretaceous fossils and extant molecules. Proc. Natl Acad. Sci. USA 101, 16571–16576 (2004).ADS 
    CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    123.Magallón, S. Using fossils to break long branches in molecular dating: a comparison of relaxed clocks applied to the origin of angiosperms. Syst. Biol. 59, 384–399 (2010).PubMed 
    Article 

    Google Scholar 
    124.Clarke, J. T., Warnock, R. C. & Donoghue, P. C. Establishing a time-scale for plant evolution. N. phytologist 192, 266–301 (2011).Article 

    Google Scholar 
    125.Heled, J. & Drummond, A. J. Calibrated tree priors for relaxed phylogenetics and divergence time estimation. Syst. Biol. 61, 138–149 (2012).PubMed 
    Article 

    Google Scholar 
    126.Llorens, C. et al. The Gypsy Database (GyDB) of mobile genetic elements: release 2.0. Nucleic Acids Res. 39, D70–D74 (2011).CAS 
    PubMed 
    Article 

    Google Scholar 
    127.Altschul, S. F., Gish, W., Miller, W., Myers, E. W. & Lipman, D. J. Basic local alignment search tool. J. Mol. Biol. 215, 403–410 (1990).CAS 
    Article 

    Google Scholar 
    128.Yu, X. J., Zheng, H. K., Wang, J., Wang, W. & Su, B. Detecting lineage-specific adaptive evolution of brain-expressed genes in human using rhesus macaque as outgroup. Genomics 88, 745–751 (2006).CAS 
    PubMed 
    Article 

    Google Scholar 
    129.Birney, E., Clamp, M. & Durbin, R. GeneWise and genomewise. Genome Res. 14, 988–995 (2004).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    130.Vilella, A. J. et al. EnsemblCompara geneTrees: complete, duplication-aware phylogenetic trees in vertebrates. Genome Res. 19, 327–335 (2009).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    131.Seberg, O. & Petersen, G. A unified classification system for eukaryotic transposable elements should reflect their phylogeny. Nat. Rev. Genet. 10, 276 (2009).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    132.Chen, Y. et al. SOAPnuke: a MapReduce acceleration-supported software for integrated quality control and preprocessing of high-throughput sequencing data. Gigascience 7, 1–6 (2018).ADS 
    PubMed 
    PubMed Central 

    Google Scholar 
    133.Krueger, F. & Andrews, S. R. Bismark: a flexible aligner and methylation caller for Bisulfite-Seq applications. Bioinformatics 27, 1571–1572 (2011).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    134.Quinlan, A. R. & Hall, I. M. BEDTools: a flexible suite of utilities for comparing genomic features. Bioinformatics 26, 841–842 (2010).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    135.Jühling, F. et al. Metilene: fast and sensitive calling of differentially methylated regions from bisulfite sequencing data. Genome Res. 26, 256–262 (2016).PubMed 
    PubMed Central 
    Article 
    CAS 

    Google Scholar 
    136.Xie, C. et al. KOBAS 2.0: a web server for annotation and identification of enriched pathways and diseases. Nucleic Acids Res. 39, W316–W322 (2011).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    137.Robinson, M. D., McCarthy, D. J. & Smyth, G. K. edgeR: a Bioconductor package for differential expression analysis of digital gene expression data. Bioinformatics 26, 139–140 (2010).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    138.Langfelder, P. & Horvath, S. WGCNA: an R package for weighted correlation network analysis. BMC Bioinformatics 9, 559 (2008).PubMed 
    PubMed Central 
    Article 
    CAS 

    Google Scholar 
    139.Chomczynski, P. & Sacchi, N. Single-step method of RNA isolation by acid guanidinium thiocyanate-phenol-chloroform extraction. Anal. Biochem. 162, 156–159 (1987).CAS 
    PubMed 
    Article 

    Google Scholar 
    140.Kechin, A., Boyarskikh, U., Kel, A. & Filipenko, M. CutPrimers: a new tool for accurate cutting of primers from reads of targeted next generation sequencing. J. Comput. Biol. 24, 1138–1143 (2017).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    141.Langmead, B., Trapnell, C., Pop, M. & Salzberg, S. L. Ultrafast and memory-efficient alignment of short DNA sequences to the human genome. Genome Biol. 10, 25 (2009).Article 
    CAS 

    Google Scholar 
    142.Friedländer, M. R., Mackowiak, S. D., Li, N., Chen, W. & Rajewsky, N. miRDeep2 accurately identifies known and hundreds of novel microRNA genes in seven animal clades. Nucleic Acids Res. 40, 37–52 (2012).PubMed 
    Article 
    CAS 
    PubMed Central 

    Google Scholar 
    143.Kozomara, A., Birgaoanu, M. & Griffiths-Jones, S. miRBase: from microRNA sequences to function. Nucleic Acids Res. 47, D155–d162 (2019).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    144.Li, Z. & He, Y. Roles of brassinosteroids in plant reproduction. Int. J. Mol. Sci. 21, 872 (2020).PubMed Central 
    Article 
    CAS 

    Google Scholar 
    145.Xin, P., Yan, J., Fan, J., Chu, J. & Yan, C. An improved simplified high-sensitivity quantification method for determining brassinosteroids in different tissues of rice and Arabidopsis. Plant Physiol. 162, 2056–2066 (2013).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    146.Li, L., Stoeckert, C. J. Jr. & Roos, D. S. OrthoMCL: identification of ortholog groups for eukaryotic genomes. Genome Res. 13, 2178–2189 (2003).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    147.Finn, R. D. et al. Pfam: the protein families database. Nucleic Acids Res. 42, D222–D230 (2014).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    148.Anders, S., Pyl, P. T. & Huber, W. HTSeq–a Python framework to work with high-throughput sequencing data. Bioinformatics 31, 166–169 (2015).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    149.Mortazavi, A., Williams, B. A., McCue, K., Schaeffer, L. & Wold, B. Mapping and quantifying mammalian transcriptomes by RNA-Seq. Nat. Methods 5, 621–628 (2008).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    150.Wang, Y. et al. MCScanX: a toolkit for detection and evolutionary analysis of gene synteny and collinearity. Nucleic Acids Res. 40, e49 (2012).ADS 
    CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar  More

  • in

    Dinophyceae can use exudates as weapons against the parasite Amoebophrya sp. (Syndiniales)

    1.Carlson CJ, Burgio KR, Dougherty ER, Phillips AJ, Bueno VM, Clements CF, et al. Parasite biodiversity faces extinction and redistribution in a changing climate. Sci Adv. 2017;3:e1602422.PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    2.Johnson PTJ, Preston DL, Hoverman JT, LaFonte BE. Host and parasite diversity jointly control disease risk in complex communities. Proc Natl Acad Sci USA. 2013;110:16916–21.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    3.Dougherty ER, Carlson CJ, Bueno VM, Burgio KR, Cizauskas CA, Clements CF, et al. Paradigms for parasite conservation: parasite conservation. Conserv Biol. 2016;30:724–33.PubMed 
    Article 

    Google Scholar 
    4.Paseka RE, White LA, Van de Waal DB, Strauss AT, González AL, Everett RA, et al. Disease-mediated ecosystem services: pathogens, plants, and people. Trends Ecol Evolut. 2020;35:731–43.Article 

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

    Google Scholar 
    6.Bjorbækmo MFM, Evenstad A, Røsæg LL, Krabberød AK, Logares R. The planktonic protist interactome: where do we stand after a century of research? ISME J. 2020;14:544–59.PubMed 
    Article 

    Google Scholar 
    7.Brussaard CPD. Viral control of phytoplankton populations-a review. J Eukaryot Microbiol. 2004;51:125–38.PubMed 
    Article 

    Google Scholar 
    8.Chambouvet A, Morin P, Marie D, Guillou L. Control of toxic marine dinoflagellate blooms by serial parasitic killers. Science. 2008;322:1254–7.CAS 
    PubMed 
    Article 

    Google Scholar 
    9.Vardi A, Van Mooy BA, Fredricks HF, Popendorf KJ, Ossolinski JE, Haramaty L, et al. Viral glycosphingolipids induce lytic infection and cell death in marine phytoplankton. Science. 2009;326:861–5.CAS 
    PubMed 
    Article 

    Google Scholar 
    10.Guillou L, Viprey M, Chambouvet A, Welsh RM, Kirkham AR, Massana R, et al. Widespread occurrence and genetic diversity of marine parasitoids belonging to Syndiniales (Alveolata). Environ Microbiol. 2008;10:3349–65.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    11.de Vargas C, Audic S, Henry N, Decelle J, Mahé F, Logares R, et al. Eukaryotic plankton diversity in the sunlit ocean. Science. 2015;348:1261605.PubMed 
    Article 
    CAS 
    PubMed Central 

    Google Scholar 
    12.Siano R, Alves-de-Souza C, Foulon E, Bendif EM, Simon N, Guillou L, et al. Distribution and host diversity of Amoebophryidae parasites across oligotrophic waters of the Mediterranean Sea. Biogeosciences. 2011;8:267–78.Article 

    Google Scholar 
    13.Park M, Cooney S, Yih W, Coats D. Effects of two strains of the parasitic dinoflagellate Amoebophrya on growth, photosynthesis, light absorption, and quantum yield of bloom-forming dinoflagellates. Mar Ecol Prog Ser. 2002;227:281–92.Article 

    Google Scholar 
    14.Velo-Suárez L, Brosnahan ML, Anderson DM, McGillicuddy DJ. A Quantitative assessment of the role of the parasite Amoebophrya in the termination of Alexandrium fundyense blooms within a small coastal embayment. PLoS ONE. 2013;8:e81150.PubMed 
    PubMed Central 
    Article 
    CAS 

    Google Scholar 
    15.Li C, Song S, Liu Y, Chen T. Occurrence of Amoebophrya spp. infection in planktonic dinoflagellates in Changjiang (Yangtze River) Estuary, China. Harmful Algae. 2014;37:117–24.Article 

    Google Scholar 
    16.Choi CJ, Brosnahan ML, Sehein TR, Anderson DM, Erdner DL. Insights into the loss factors of phytoplankton blooms: the role of cell mortality in the decline of two inshore Alexandrium blooms. Limnol Oceanogr. 2017;62:1742–53.PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    17.Coats DW, Park MG. Parasitism of photosynthetic dinoflagellates by three strains of Amoebophrya (Dinophyta); parasite survival, infectivity, generation time, and host specificity. J Phycol. 2002;38:520–8.Article 

    Google Scholar 
    18.Cai R, Kayal E, Alves-de-Souza C, Bigeard E, Corre E, Jeanthon C, et al. Cryptic species in the parasitic Amoebophrya species complex revealed by a polyphasic approach. Sci Rep. 2020;10:2531.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    19.Anderson SR, Harvey EL. Temporal variability and ecological interactions of parasitic marine Syndiniales in coastal protist communities. mSphere. 2020;5:e00209–20.PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    20.Alves-de-Souza C, Pecqueur D, Le Floc’h E, Mas S, Roques C, Mostajir B, et al. Significance of plankton community structure and nutrient availability for the control of dinoflagellate blooms by parasites: a modeling approach. PLoS ONE. 2015;10:e0127623.PubMed 
    PubMed Central 
    Article 
    CAS 

    Google Scholar 
    21.Alacid E, Park MG, Turon M, Petrou K, Garcés E. A game of russian roulette for a generalist dinoflagellate parasitoid: host susceptibility is the key to success. Front Microbiol. 2016;7:769.PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    22.Vincent F, Sheyn U, Porat Z, Schatz D, Vardi A. Visualizing active viral infection reveals diverse cell fates in synchronized algal bloom demise. Proc Natl Acad Sci USA. 2021;118:e2021586118.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    23.Chambouvet A, Alves-de-Souza C, Cueff V, Marie D, Karpov S, Guillou L. Interplay between the parasite Amoebophrya sp. (Alveolata) and the cyst formation of the red tide dinoflagellate Scrippsiella trochoidea. Protist. 2011;162:637–49.PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    24.Pelusi A, De Luca P, Manfellotto F, Thamatrakoln K, Bidle KD, Montresor M. Virus‐induced spore formation as a defense mechanism in marine diatoms. New Phytol. 2020;229:16951–2259.
    Google Scholar 
    25.Pouneva ID. Effect of abscisic acid and ontogenic phases of the host alga on the infection process in the pathosystem Scenedesmus acutus—Phlyctidium scenenedesmi. Acta Physiol Plant. 2006;28:395–400.CAS 
    Article 

    Google Scholar 
    26.Bai X, Adolf JE, Bachvaroff T, Place AR, Coats DW. The interplay between host toxins and parasitism by Amoebophrya. Harmful Algae. 2007;6:670–8.CAS 
    Article 

    Google Scholar 
    27.Place AR, Bai X, Kim S, Sengco MR, Wayne, Coats D. Dinoflagellate host-parasite sterol profiles dictate karlotoxin sensitivity. J Phycol. 2009;45:375–85.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    28.Rohrlack T, Christiansen G, Kurmayer R. Putative antiparasite defensive system involving ribosomal and nonribosomal oligopeptides in Cyanobacteria of the Genus Planktothrix. Appl Environ Microbiol. 2013;79:2642–7.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    29.Scholz B, Küpper F, Vyverman W, Ólafsson H, Karsten U. Chytridiomycosis of marine diatoms—the role of stress physiology and resistance in parasite-host recognition and accumulation of defense molecules. Marine Drugs. 2017;15:26.PubMed Central 
    Article 
    CAS 

    Google Scholar 
    30.Granéli E, Hansen PJ. Allelopathy in harmful algae: a mechanism to compete for resources? In: Granéli E, Turner JT, editors. Ecology of harmful algae. Springer Berlin Heidelberg; 2006. p. 189–201.31.Farhat S, Le P, Kayal E, Noel B, Bigeard E, Corre E, et al. Rapid protein evolution, organellar reductions, and invasive intronic elements in the marine aerobic parasite dinoflagellate Amoebophrya spp. BMC Biol. 2021;19:1.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    32.Chapelle A, Le Bec C, Amzil Z, Dreanno C, Klouch KZ, Labry C, et al. Etude sur la proliferation de la micro algue Alexandrium minutum en rade de Brest (2014).33.Chapelle A, Le Gac M, Labry C, Siano R, Quere J, Caradec F, et al. The Bay of Brest (France), a new risky site for toxic Alexandrium minutum blooms and PSP shellfish contamination. Harmful Algal News. 2015;51:4–5.34.Klouch KZ, Schmidt S, Andrieux-Loyer F, Le Gac M, Hervio-Heath D, Qui-Minet ZN. et al. Historical records from dated sediment cores reveal the multidecadal dynamic of the toxic dinoflagellate Alexandrium minutum in the Bay of Brest (France). FEMS Microbiol Ecol. 2016;92:fiw101PubMed 
    Article 
    CAS 
    PubMed Central 

    Google Scholar 
    35.Long M, Tallec K, Soudant P, Le Grand F, Donval A, Lambert C, et al. Allelochemicals from Alexandrium minutum induce rapid inhibition of metabolism and modify the membranes from Chaetoceros muelleri. Algal Res. 2018;35:508–18.Article 

    Google Scholar 
    36.Long M, Peltekis A, González-Fernández C, Bailleul B, Hégaret H. Allelochemicals of Alexandrium minutum: kinetics of membrane disruption and photosynthesis inhibition in a co-occurring diatom. Harmful Algae. 2021;103:101997.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    37.Starr RC, Zeikus JA. Utex—The culture collection of algae at the university of Texas at Austin 1993 List of cultures. J Phycol. 1993;29:1–106.Article 

    Google Scholar 
    38.Keller M, Selvin R, Claus W, Guillard RRL. Media for the culture of oceanic ultraphytoplankton 1, 2. J Phycol. 1987;23:633–8.39.Bigeard. Collect of Amoebophrya parasite (free-living stage) for genomic and transcriptomic analyses. 2019. Protocols.io.40.Kim S, Gil Park M, Yih W, Coats DW. Infection of the bloom-forming thecate dinoflagellates Alexandrium affina and Gonyaulax spinifera by two strains of Amoebophrya (Dinophyta). J Phycol. 2004;40:815–22.Article 

    Google Scholar 
    41.Kim S. Patterns in host range for two strains of Amoebophrya (Dinophyta) infecting thecate dinoflagellates: Amoebophrya spp. ex Alexandrium affine and ex Gonyaulax polygramma. J Phycol. 2006;42:1170–3.Article 

    Google Scholar 
    42.Kayal E, Alves-de-Souza C, Farhat S, Velo-Suarez L, Monjol J, Szymczak J, et al. Dinoflagellate host chloroplasts and mitochondria remain functional during Amoebophrya Infection. Front Microbiol. 2020;11:600823.PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    43.R Core Team. R: a language and environment for statistical computing. Vienna: R Foundation for Statistical Computing.44.John U, Tillmann U, Hülskötter J, Alpermann TJ, Wohlrab S, Van de Waal DB. Intraspecific facilitation by allelochemical mediated grazing protection within a toxigenic dinoflagellate population. Proc R Soc B. 2015;282:20141268.PubMed 
    PubMed Central 
    Article 
    CAS 

    Google Scholar 
    45.Lelong A, Haberkorn H, Le Goïc N, Hégaret H, Soudant P. A new insight into allelopathic effects of Alexandrium minutum on photosynthesis and respiration of the diatom Chaetoceros neogracile revealed by photosynthetic-performance analysis and flow cytometry. Microb Ecol. 2011;62:919–30.CAS 
    PubMed 
    Article 

    Google Scholar 
    46.Tillmann U, Alpermann T, John U, Cembella A. Allelochemical interactions and short-term effects of the dinoflagellate Alexandrium on selected photoautotrophic and heterotrophic protists. Harmful Algae. 2008;7:52–64.Article 

    Google Scholar 
    47.Durham WM, Stocker R. Thin phytoplankton layers: characteristics, mechanisms, and consequences. Annu Rev Mar Sci. 2012;4:177–207.Article 

    Google Scholar 
    48.Breier RE, Lalescu CC, Waas D, Wilczek M, Mazza MG. Emergence of phytoplankton patchiness at small scales in mild turbulence. Proc Natl Acad Sci USA. 2018;115:12112–7.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    49.Wheeler JD, Secchi E, Rusconi R, Stocker R. Not just going with the flow: the effects of fluid flow on bacteria and plankton. Annu Rev Cell Dev Biol. 2019;35:213–37.CAS 
    PubMed 
    Article 

    Google Scholar 
    50.Basterretxea G, Font-Muñoz JS, Tuval I. Phytoplankton orientation in a turbulent ocean: a microscale perspective. Front Mar Sci. 2020;7:185.Article 

    Google Scholar 
    51.Blossom HE, Markussen B, Daugbjerg N, Krock B, Norlin A, Hansen PJ. The cost of toxicity in microalgae: direct evidence from the dinoflagellate Alexandrium. Front Microbiol. 2019;10:1065.PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    52.Martens H, Van de Waal DB, Brandenburg KM, Krock B, Tillmann U. Salinity effects on growth and toxin production in an Alexandrium ostenfeldii (Dinophyceae) isolate from The Netherlands. J Plankton Res. 2016;38:1302–16.CAS 
    Article 

    Google Scholar 
    53.Long M, Holland A, Planquette H, González Santana D, Whitby H, Soudant P, et al. Effects of copper on the dinoflagellate Alexandrium minutum and its allelochemical potency. Aquat Toxicol. 2019;210:251–61.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    54.Brown ER, Kubanek J. Harmful alga trades off growth and toxicity in response to cues from dead phytoplankton. Limnol Oceanogr. 2020;65:1723–33.CAS 
    Article 

    Google Scholar 
    55.Selander E, Thor P, Toth G, Pavia H. Copepods induce paralytic shellfish toxin production in marine dinoflagellates. Proc R Soc B. 2006;273:1673–80.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    56.Lu Y, Wohlrab S, Groth M, Glöckner G, Guillou L, John U. Transcriptomic profiling of Alexandrium fundyense during physical interaction with or exposure to chemical signals from the parasite Amoebophrya. Mol Ecol. 2016;25:1294–307.CAS 
    PubMed 
    Article 

    Google Scholar 
    57.Seymour JR, Amin SA, Raina J-B, Stocker R. Zooming in on the phycosphere: the ecological interface for phytoplankton–bacteria relationships. Nat Microbiol. 2017;2:17065.CAS 
    PubMed 
    Article 

    Google Scholar 
    58.Place A, Harvey H, Bai X, Coats D. Sneaking under the toxin surveillance radar: parasitism and sterol content. Afr J Mar Sci. 2006;28:347–51.Article 

    Google Scholar 
    59.Ma H, Krock B, Tillmann U, Bickmeyer U, Graeve M, Cembella A. Mode of action of membrane-disruptive lytic compounds from the marine dinoflagellate Alexandrium tamarense. Toxicon. 2011;58:247–58.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    60.Deeds J, Place A. Sterol-specific membrane interactions with the toxins from Karlodinium micrum (Dinophyceae) — a strategy for self-protection? Afr J Mar Sci. 2006;28:421–5.Article 

    Google Scholar 
    61.Leblond JD, Sengco MR, Sickman JO, Dahmen JL, Anderson DM. Sterols of the Syndinian dinoflagellate Amoebophrya sp., a parasite of the dinoflagellate Alexandrium tamarense (Dinophyceae). J Eukaryotic Microbiol. 2006;53:211–6.CAS 
    Article 

    Google Scholar 
    62.Long M, Tallec K, Soudant P, Lambert C, Le Grand F, Sarthou G, et al. A rapid quantitative fluorescence-based bioassay to study allelochemical interactions from Alexandrium minutum. Environ Pollut. 2018;242:1598–605.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    63.Borcier E, Morvezen R, Boudry P, Miner P, Charrier G, Laroche J, et al. Effects of bioactive extracellular compounds and paralytic shellfish toxins produced by Alexandrium minutum on growth and behaviour of juvenile great scallops Pecten maximus. Aquatic Toxicol. 2017;184:142–54.CAS 
    Article 

    Google Scholar 
    64.Castrec J, Soudant P, Payton L, Tran D, Miner P, Lambert C, et al. Bioactive extracellular compounds produced by the dinoflagellate Alexandrium minutum are highly detrimental for oysters. Aquat Toxicol. 2018;199:188–98.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    65.Wang Y, Tang X. Interactions between Prorocentrum donghaiense Lu and Scrippsiella trochoidea (Stein) Loeblich III under laboratory culture. Harmful Algae. 2008;7:65–75.Article 

    Google Scholar 
    66.Tang YZ, Gobler CJ. Lethal effects of Northwest Atlantic Ocean isolates of the dinoflagellate, Scrippsiella trochoidea, on Eastern oyster (Crassostrea virginica) and Northern quahog (Mercenaria mercenaria) larvae. Mar Biol. 2012;159:199–210.Article 

    Google Scholar 
    67.Felpeto AB, Roy S, Vasconcelos VM. Allelopathy prevents competitive exclusion and promotes phytoplankton biodiversity. Oikos. 2018;127:85–98.Article 

    Google Scholar 
    68.Driscoll WW, Hackett JD, Ferrière R. Eco-evolutionary feedbacks between private and public goods: evidence from toxic algal blooms. Ecol Lett. 2016;19:81–97.PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    69.Driscoll WW, Espinosa NJ, Eldakar OT, Hackett JD. Allelopathy as an emergent, exploitable public good in the bloom-forming microalga Prymnesium parvum. Evolution. 2013;67:1582–90.PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    70.Rodríguez F, Figueroa RI. Confirmation of the wide host range of Parvilucifera corolla (Alveolata, Perkinsozoa). Eur J Protistol. 2020;74:125690.PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    71.Chambouvet A, Laabir M, Sengco M, Vaquer A, Guillou L. Genetic diversity of Amoebophryidae (Syndiniales) during Alexandrium catenella/tamarense (Dinophyceae) blooms in the Thau lagoon (Mediterranean Sea, France). Res Microbiol. 2011;162:959–68.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    72.Cosgrove S. Monitoring methods and bloom dynamic studies of the toxic dinoflagellate genus Alexandrium. 2014. Doctoral dissertation, National University of Ireland, Galway.73.Hutchinson GE. The Paradox of the plankton. Am Nat. 1961;95:137–45.Article 

    Google Scholar 
    74.Czaran TL, Hoekstra RF, Pagie L. Chemical warfare between microbes promotes biodiversity. Proc Natl Acad Sci USA. 2002;99:786–90.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    75.Garcés E, Alacid E, Reñé A, Petrou K, Simó R. Host-released dimethylsulphide activates the dinoflagellate parasitoid Parvilucifera sinerae. ISME J. 2013;7:1065–8.PubMed 
    PubMed Central 
    Article 
    CAS 

    Google Scholar 
    76.Fitzpatrick CR, Salas-González I, Conway JM, Finkel OM, Gilbert S, Russ D. et al. The plant microbiome: from ecology to reductionism and beyond. Annu Rev Microbiol. 2020;74:annurev-micro-022620-014327Article 
    CAS 

    Google Scholar 
    77.Carney LT, Lane TW. Parasites in algae mass culture. Front Microbiol. 2014;5:278. More

  • in

    Evolutionary loss of thermal acclimation accompanied by periodic monocarpic mass flowering in Strobilanthes flexicaulis

    1.Gunderson, C. A., O’Hara, K. H., Campion, C. M., Waler, A. V. & Edwards, N. T. Thermal plasticity of photosynthesis: The role of acclimation in forest responses to a warming climate. Glob. Change Biol. 16, 2272–2286 (2010).ADS 
    Article 

    Google Scholar 
    2.Sage, R. F., Way, D. A. & Kubien, D. S. Rubisco, Rubisco activase, and global climate change. J. Exp. Bot. 59, 1581–1595 (2008).CAS 
    PubMed 
    Article 

    Google Scholar 
    3.Sendall, K. M. et al. Acclimation of photosynthetic temperature optima of temperate and boreal tree species in response to experimental forest warming. Glob. Change Biol. 21, 1342–1357 (2015).ADS 
    Article 

    Google Scholar 
    4.IPCC AR5. Climate Change 2013: The Physical Science Basis. Contribution of Working Group I to the Fourth Assessment Report to the Intergovernmental Panel on Climate Change (Cambridge University Press, 2013).
    Google Scholar 
    5.Atkin, O. K. et al. 2008 Using temperature-dependent changes in leaf scaling relationships to quantitatively account for thermal acclimation of respiration in a coupled global climate–vegetation model. Glob. Change Biol. 14, 2709–2726 (2008).ADS 
    Article 

    Google Scholar 
    6.Reich, P. B. et al. Boreal and temperate trees show strong acclimation of respiration to warming. Nature 531, 633–636 (2016).ADS 
    CAS 
    PubMed 
    Article 

    Google Scholar 
    7.Hikosaka, H., Ishikawa, K., Borjigidai, A., Muller, O. & Onoda, Y. Temperature acclimation of photosynthesis: Mechanisms involved in the changes in temperature dependence of photosynthetic rate. J. Exp. Bot. 57, 291–302 (2006).CAS 
    PubMed 
    Article 

    Google Scholar 
    8.Ow, L. F., Griffin, K. L., Whitehead, D., Walcroft, A. S. & Turnbull, M. H. Thermal acclimation of leaf respiration but not photosynthesis in Populus deltoides × nigra. New Phytol. 178, 123–134 (2008).PubMed 
    Article 

    Google Scholar 
    9.Way, D. A. & Yamori, W. Thermal acclimation of photosynthesis: On the importance of adjusting our definitions and accounting for thermal acclimation of respiration. Photosynth. Res. 119, 89–100 (2014).CAS 
    PubMed 
    Article 

    Google Scholar 
    10.Heskel, M. A. et al. Convergence in the temperature response of leaf respiration across biomes and plant functional types. Proc. Natl. Acad. Sci. USA 113, 3832–3837 (2016).ADS 
    CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    11.Sage, R. F. & Kubien, D. S. The temperature response of C3 and C4 photosynthesis. Plant Cell Environ. 30, 1086–1106 (2007).CAS 
    PubMed 
    Article 

    Google Scholar 
    12.Scafaro, A. P. et al. Strong thermal acclimation of photosynthesis in tropical and temperate wet-forest tree species: the importance of altered Rubisco content. Glob. Change Biol. 23, 2783–2800 (2017).ADS 
    Article 

    Google Scholar 
    13.Higgins, S. I. & Richardson, D. M. Predicting plant migration rates in a changing world: the role of long-distance dispersal. Am. Nat. 153, 464–475 (1999).PubMed 
    Article 

    Google Scholar 
    14.Atkin, O. K. & Tjoelker, M. G. Thermal acclimation and the dynamic response of plant respiration to temperature. Trend. Plant Sci. 8, 343–351 (2003).CAS 
    Article 

    Google Scholar 
    15.Yamori, W., Hikosaka, K. & Way, D. A. Temperature response of photosynthesis in C3, C4, and CAM plants: temperature acclimation and temperature adaptation. Photosynth. Res. 119, 101–117 (2014).CAS 
    PubMed 
    Article 

    Google Scholar 
    16.Atkin, O. K. et al. Global variability in leaf respiration in relation to climate, plant functional types and leaf traits. New Phytol. 206, 614–636 (2015).CAS 
    PubMed 
    Article 

    Google Scholar 
    17.Slot, M. & Kitajima, K. General patterns of acclimation of leaf respiration to elevated temperatures across biomes and plant types. Oecologia 177, 885–900 (2015).ADS 
    PubMed 
    Article 

    Google Scholar 
    18.Atkinson, L. J., Hellicar, M. A., Fitter, A. H. & Atkin, O. K. Impact of temperature on the relationship between respiration and nitrogen concentration in roots: An analysis of scaling relationships, Q10 values and thermal acclimation ratios. New Phytol. 173, 110–120 (2007).CAS 
    PubMed 
    Article 

    Google Scholar 
    19.Wei, X. et al. Consistent leaf respiratory response to experimental warming of three North American deciduous trees: A comparison across seasons, years, habitats and sites. Tree Physiol. 37, 285–300 (2017).CAS 
    PubMed 

    Google Scholar 
    20.Kruse, J., Hopmans, P. & Adams, M. A. Temperature responses are a window to the physiology of dark respiration: Differences between CO2 release and O2 reduction shed light on energy conservation. Plant Cell Environ. 31, 901–914 (2008).CAS 
    PubMed 
    Article 

    Google Scholar 
    21.Kruse, J., Rennenberg, H. & Adams, M. A. Steps towards a mechanistic understanding of respiratory temperature responses. New Phytol. 189, 659–677 (2011).CAS 
    PubMed 
    Article 

    Google Scholar 
    22.Kakishima, S., Yoshimura, J., Murata, H. & Murata, J. 6-year periodicity and variable synchronicity in a mass-flowering plant. PLoS ONE 6, e28140. https://doi.org/10.1371/journal.pone.002814 (2011).ADS 
    CAS 
    Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    23.Kakishima, S. et al. Evolutionary origin of a periodical mass-flowering plant. Ecol. Evol. 9, 4373–4381 (2019).PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    24.Björkman, O. & Denning, B. Photon field of O2 evolution and chlorophyll fluorescence characteristics at 77K among vascular plants of diverse origins. Planta 170, 489–504 (1987).PubMed 
    Article 

    Google Scholar 
    25.Stirling, C. M., Aguilera, C., Baker, N. R. & Long, S. P. Changes in the photosynthetic light response curve during leaf development of field-grown maize with implications for modeling canopy photosynthesis. Photosynth. Res. 42, 217–225 (1994).CAS 
    PubMed 
    Article 

    Google Scholar 
    26.Ishida, A., Uemura, A., Koike, N., Matsumoto, Y. & Ang, L. H. Interactive effects of leaf age and self-shading on leaf structure, photosynthetic capacity and chlorophyll fluorescence in the rain forest tree, Dryobalanops aromatic. Tree Physiol. 19, 741–747 (1999).PubMed 
    Article 

    Google Scholar 
    27.Bouma, T. J. et al. Respiratory energy requirements and rate of protein turnover in vivo determined by the use of an inhibitor of protein synthesis and a probe to assess its effect. Physiol. Plant. 92, 585–594 (1994).CAS 
    Article 

    Google Scholar 
    28.Noguchi, K. et al. Costs of protein turnover and carbohydrate export in leaves of sun and shade species. Aust. J. Plant Physiol. 28, 37–47 (2001).CAS 

    Google Scholar 
    29.Stephanie, S. Y. et al. Respiratory alternative oxidase responds to both low- and high-temperature stress in Quercus rubra leaves along an urban-rural gradient in New York. Funct. Ecol. 25, 1007–1017 (2011).Article 

    Google Scholar 
    30.Reich, P. B., Walters, M. B. & Ellsworth, D. S. From tropics to tundra: Global convergence in plant functioning. Proc. Natl. Acad. Sci. USA 94, 13730–13734 (1997).ADS 
    CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    31.Wright, I. J. et al. The worldwide leaf economics spectrum. Nature 428, 821–827 (2004).ADS 
    CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    32.Ishida, A. et al. Coordination between leaf and stem traits related to leaf carbon gain and hydraulics across 32 drought-tolerant angiosperms. Oecologica 156, 193–202 (2008).ADS 
    Article 

    Google Scholar 
    33.He, P. et al. Leaf mechanical strength and photosynthetic capacity wary independently across 57 subtropical forest species with contrasting light requirements. New Phytol. 223, 607–618 (2019).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    34.Yamashita, N., Koike, N. & Ishida, A. Leaf ontogenetic dependence of light acclimation in invasive and native subtropical trees of different successional status. Plant Cell Environ. 25, 1341–1356 (2002).Article 

    Google Scholar 
    35.McDowell, N. et al. Mechanisms of plant survival and mortality during drought: Why do some plants survive while others succumb to drought?. New Phytol. 178, 719–739 (2008).PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    36.Sala, A., Piper, F. & Hoch, G. Physiological mechanisms of drought-induced tree mortality are far from being resolved. New Phytol. 186, 274–281 (2010).PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    37.O’Brien, M. J., Leuzinger, S., Philipson, C. D., Tay, J. & Hector, A. Drought survival of tropical tree seedlings enhanced by non-structural carbohydrate levels. Nat. Clim. Change 4, 710–714 (2014).ADS 
    Article 
    CAS 

    Google Scholar 
    38.Saiki, S.-T., Ishida, A., Yoshimura, K. & Yazaki, K. Physiological mechanisms of drought-induced tree die-off in relation to carbon, hydraulic and respiratory stress in a drought-tolerant woody plant. Sci. Rep. 7, 2995. https://doi.org/10.1038/s41598-017-03162-5 (2017).ADS 
    CAS 
    Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    39.Kono, Y. et al. Initial hydraulic failure followed by late-stage carbon starvation leads to drought-induced death in the tree Trema orientalis. Commun. Biol. 2, 8. https://doi.org/10.1038/s42003-018-0256-7 (2019).Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    40.Ebbert, V., Adams, W. W. III., Mattoo, A. K., Sokolenko, A. & Demming-Adams, B. Up-regulation of a photosystem II core protein phosphatase inhibitor and sustained D1 phosphorylation in zeaxanthin-retaining, photoinhibited needles of overwintering Douglas fir. Plant Cell Environ. 28, 232–240 (2005).CAS 
    Article 

    Google Scholar 
    41.Harayama, H., Ikeda, T., Ishida, A. & Yamamoto, S. I. Seasonal variations in water relations in current-year leaves of evergreen trees with delayed greening. Tree Physiol. 26, 1025–1033 (2006).PubMed 
    Article 

    Google Scholar 
    42.Yasumura, Y. & Ishida, A. Temporal variation in leaf nitrogen partitioning of a broad-leaved evergreen tree, Quercus myrsinaefolia. J. Plant Res. 124, 115–123 (2011).CAS 
    PubMed 
    Article 

    Google Scholar 
    43.Fowler, S. & Thomashow, M. F. Arabidopsis trancriptome profiling indicates that multiple regulatory pathways are activated during cold acclimation in addition in to the CBF cold response pathway. Plant Cell 14, 1675–1690 (2002).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    44.Oono, Y. et al. Monitoring expression profiles of Arabidopsis genes during cold acclimation and deacclimation using DNA microarrays. Funct. Integr. Genom. 6, 212–234 (2006).CAS 
    Article 

    Google Scholar 
    45.Nakaminami, K. et al. Analysis of differential expression patterns of mRNA and protein during acclimation and de-acclimation in Arabidopsis. Mol. Cell. Proteom. 13, 3602–3611 (2014).CAS 
    Article 

    Google Scholar 
    46.Wang, H. et al. Acclimation of leaf respiration consistent with optimal photosynthetic capacity. Glob. Chang Biol. 26, 2573–2583 (2020).ADS 
    Article 

    Google Scholar 
    47.Maseyk, K., Grünzweig, J. M., Rotenberg, E. & Yakir, D. Respiration acclimation contributes to high carbon-use efficiency in a seasonally dry pine forest. Glob. Change Biol. 14, 1553–1567 (2008).ADS 
    Article 

    Google Scholar 
    48.Jarvi, M. P. & Burton, A. J. Acclimation and soil moisture constrain sugar maple root respiration in experimentally warmed soil. Tree Physiol. 33, 949–959 (2013).CAS 
    PubMed 
    Article 

    Google Scholar 
    49.Smith, N. G., Li, G. & Dukes, J. S. Short-term thermal acclimation of dark respiration is greater non-photosynthetic than in photosynthetic tissues. AoBP Plants 11, 1–9 (2019).CAS 

    Google Scholar 
    50.Bennett, J. R. & Scotland, R. E. A revision of Strobilanthes (Acanthaceae) in Java. Kew. Bull. 58, 1–82 (2003).Article 

    Google Scholar 
    51.Wood, J. R. I. & Scotland, R. W. New and little-known species of Strobitlanthes (Acanthaceae) from India and South East Asia. Kew. Bull. 64, 3–47 (2009).Article 

    Google Scholar 
    52.Vongkamjan, S. & Sampson, F. B. Phenology, seed germination and some vegetative features of Strobilanthes fragrans (Acanthaceae), a recently described unusual species, found only in a single Forest Park in Thailand. Thai Forest Bull. Bot. 44, 6–10 (2016).Article 

    Google Scholar 
    53.Tsukaya, H., Kakishima, S., Hidayat, A., Murata, J. & Okada, H. Flowering phenology of the nine-year plant, Strobilanthes cernua (Acanthaceae). Tropics 20, 79–85 (2011).Article 

    Google Scholar 
    54.Sharma, M. V., Kuriakose, G. & Shivanna, K. R. Reproductive strategies of Strobilanthes kunthianus, an endemic, semelparous species in southern Western Ghats, India. Bot. J. Linn. Soc. 157, 155–163 (2008).Article 

    Google Scholar  More

  • in

    Tracking Chernobyl’s effects on wildlife

    Download PDF

    Thirty-five years after the explosion and meltdown at the Chernobyl Nuclear Power Plant in Ukraine, I study how amphibians in the region have changed, physically and genetically. In 2016, I joined an international research team to do this; since then, I have obtained various grants to continue the work. Chernobyl is a phenomenal place to study rapid evolution. I typically spend two to three weeks in the forests during the frogs’ spring breeding season.When I work in the ‘exclusion zone’, the 4,700 square kilometres around the reactor, I stay in a hostel in Chernobyl (20 kilometres from the reactor site), where we have a field laboratory inside an abandoned building. The radiation in the exclusion zone is roughly 1,000 times lower than at the time of the accident, and there are now two hostels, a bar, a couple of restaurants and a cash machine. In this image, I’m running a blood analysis on one of the tree frogs we have collected. The contamination maps on the wall behind me show that some hotspots of radiation persist.Around 8 p.m., we listen for male tree frogs calling in the field. Wearing chest waders and head lamps, we enter the ponds to gather frogs until 1 or 2 a.m.. Frogs in the exclusion zone are darker than those outside it, thanks to higher levels of melanin, which might be an adaptation that protects them from ionizing radiation. We analyse how much radiation their bodies contain, and tend to find damage to some, often to the liver.Once expected to become a wasteland, the Chernobyl area is now a nature reserve. New species have arrived, including European bison (Bison bonasus) and the wild Przewalski’s horse (Equus ferus przewalskii). We’re beginning to monitor these horses, originally from the Asian steppes: the effects on their health could be a proxy for what happens when humans return. The first 31 horses were released here in 1998, 12 years after the disaster, and it is one of the few places where they continue to live freely.

    Nature 595, 464 (2021)
    doi: https://doi.org/10.1038/d41586-021-01883-2

    Related Articles

    The zoologist tracking an island’s rebirth

    Extreme research

    Collection: Fieldwork

    Subjects

    Careers

    Ecology

    Evolution

    Latest on:

    Careers

    How a holistic research retreat can help our science
    Career Column 08 JUL 21

    Research managers are essential to a healthy research culture
    Editorial 07 JUL 21

    What polar researchers have learnt from the pandemic
    Career Feature 06 JUL 21

    Ecology

    Newfound ‘fairy lantern’ could soon be snuffed out forever
    Research Highlight 07 JUL 21

    Caution over the use of ecological big data for conservation
    Matters Arising 07 JUL 21

    How to buffer against an urban food shortage
    News & Views 07 JUL 21

    Evolution

    From the archive
    News & Views 06 JUL 21

    A piece of Triassic poo yields a beautifully preserved beetle
    Research Highlight 30 JUN 21

    Beyond coronavirus: the virus discoveries transforming biology
    News Feature 30 JUN 21

    Jobs

    Higher Scientific Officer – Cancer Organoids Facility

    Institute of Cancer Research (ICR)
    London, United Kingdom

    Research Fellow in Characterisation of Bioseparation Materials

    University College London (UCL)
    London, United Kingdom

    University of Surrey – Academic AI roles.

    University of Surrey
    Guildford, United Kingdom

    Research Assistant/Associate

    University of Glasgow
    Glasgow, United Kingdom

    Nature Briefing
    An essential round-up of science news, opinion and analysis, delivered to your inbox every weekday.

    Email address

    Yes! Sign me up to receive the daily Nature Briefing email. I agree my information will be processed in accordance with the Nature and Springer Nature Limited Privacy Policy.

    Sign up More

  • in

    Inferring the ecological niche of bat viruses closely related to SARS-CoV-2 using phylogeographic analyses of Rhinolophus species

    Genetic analyses of Rhinolophus species identified as reservoirs of viruses closely related to SARS-CoV-2Until now, SCoV2rCs have been found in four bat species of the genus Rhinolophus: R. acuminatus, R. affinis, R. malayanus, and R. shameli. The haplotype networks constructed using CO1 sequences of these four species are shown in Fig. 3. A star-like genetic pattern, characterized by one dominant haplotype and several satellite haplotypes was found for the two bat species endemic to Southeast Asia, i.e. R. acuminatus and R. shameli.Figure 3Haplotype networks based on CO1 sequences of the four Rhinolophus species found positive for viruses closely related to SARS-CoV-2 (SCoV2rCs). The networks were constructed with the median joining method available in PopART 1.513 and modified under Adobe Illustrator CS6 (version 16.0). The codes used for the countries are the following: B (Myanmar), C (Cambodia), Ch (China), I (Indonesia), L (Laos), M (Malaysia), T (Thailand), and V (Vietnam). Colours indicate the geographic origin of haplotypes according to Fig. 2 (see online supplementary Table S1). The circles indicate haplotypes separated by at least one mutation. The black lines on the branches show the number of mutations ≥ 2. Black circles represent missing haplotypes. Circle size is proportional to the number of haplotypes. Haplogroups separated by more than seven mutations (pairwise nucleotide distances  > 1%) are highlighted by dotted lines. The red arrows show the positions of the nine bats found positive for SCoV2rCs.Full size imageIn the network of R. acuminatus, the most common haplotype (named Rac1 in online supplementary Table S1) was found in northern Cambodia, southern Laos, eastern Thailand and southern Vietnam, indicating recent gene flow among these populations. Since a virus related to SARS-CoV-2 (91.8% of genome identity), named RacCS203, was detected in five R. acuminatus bats caught in eastern Thailand in June 20206, the genetic pattern obtained for this species suggests that viruses closely related to RacCS203 may have circulated in most southern regions of mainland Southeast Asia. In contrast, R. acuminatus bats collected in Borneo (M5) showed a divergent haplotype (separated by 12 mutations; haplogroup II), suggesting that the South China Sea between mainland Southeast Asia and Borneo constitutes a barrier to gene flow. Isolated populations of R. acuminatus described in northern Myanmar, Indonesia (Java and Sumatra) and the Philippines14 should be further studied.The network of R. shameli shows a typical star-like pattern, the most common haplotype (named Rsh1 in online supplementary Table S1) being detected in northern Cambodia and Laos. Since a virus related to SARS-CoV-2 (93.1% of genome identity), named RshSTT200, was recently discovered in two R. shameli bats collected in northern Cambodia in December 20107, the genetic pattern obtained for this species suggests that viruses closely related to RshSTT200 may have circulated, at least in the zone between northern Cambodia and central Laos. The bats sampled south to the Tonle Sap lake (n = 4; southern Cambodia and Vietnamese island of Phu Quoc) were found to be genetically isolated from northern populations (four mutations). However, further sampling in the south is required to confirm this result, as it may reveal CO1 sequences identical to the haplotypes detected in the north.For the two species distributed in both China and Southeast Asia, i.e. R. affinis and R. malayanus, the genetic patterns are more complex with different haplogroups showing more than 1% of nucleotide divergence. In the network of R. affinis, there are three major haplogroups (named I, II and III in Fig. 3) separated by a minimum of seven mutations. The results are therefore in agreement with those previously published using CO1 and D-loop mitochondrial sequences15. The CO1 haplotypes detected in the localities sampled in southern China (ch1, ch4, ch5) are distantly related to the single haplotype available for central China (ch6), but they are also found in Laos, northern and central Vietnam, northern Thailand and northeastern Myanmar. This result suggests recent gene flow between populations from southern Yunnan and those from northern mainland Southeast Asia. Since a virus related to SARS-CoV-2 (96.2% of genome identity), named RaTG13, was detected in one R. affinis bat captured in southern Yunnan in 20131, the genetic pattern obtained for this species suggests that viruses closely related to RaTG13 may have circulated in the zone comprising southern Yunnan and northern mainland Southeast Asia.In the network of R. malayanus, there are four major haplogroups (named I, II, III and IV in Fig. 3) separated by a minimum of seven mutations. The CO1 haplotypes detected in the localities sampled in southern China (ch2 and ch3) were also found in northern Laos (L1 and L3), suggesting recent gene flow between populations from these two countries. Since a virus related to SARS-CoV-2 (93.7% of genome identity), named RmYN02, was recently isolated from one R. malayanus bat collected in southern Yunnan in June 20195, the genetic pattern obtained for this species suggests that viruses closely related to RmYN02 may have circulated, at least between southern Yunnan and northern Laos. In contrast, the bats sampled in Myanmar were found to be genetically isolated from other geographic populations (haplogroup II in Fig. 3).Two different ecological niches for bat viruses related to either SARS-CoV or SARS-CoV-2In the wild, sarbecoviruses were generally detected after examining fecal samples collected on dozens of bats. For instance, two sarbecoviruses were found7 among the total 59 bats collected at the same cave entrance in northern Cambodia in 2010 (unpublished data). However, this does not mean necessarily that sarbecoviruses were absent in negative samples, as degradation of RNA molecules and very low viral concentrations may prevent the detection of RNA viruses. Despite these difficulties, full genomes of Sarbecovirus have been sequenced from a wide diversity of horseshoe bat species collected in Asia, Africa and Europe5,6,7,8,9,10. Therefore, there is no doubt that Rhinolophus species constitute the natural reservoir host of all sarbecoviruses3,8. The genus Rhinolophus currently includes between 9211 and 10916 insectivorous species that inhabit temperate and tropical regions of the Old World, with a higher biodiversity in Asia (63–68 out of the 92–109 described species) than in Africa (34–38 species), Europe (5 species) and Oceania (5 species). Although some Rhinolophus species are solitary, most of them are gregarious and live in large colonies or small groups generally in caves and hollow trees, but also in burrows, tunnels, abandonned mines, and old buildings11,16. However, they prefer large caves with total darkness, where temperatures are stable and less affected by diurnal and seasonal climatic variations. Importantly, all Rhinolophus species in which sarbecoviruses were detected in previous studies1,5,6,7,8,9,17 are cave species that form small groups or colonies (up to several hundreds)11,18,19.In China, many SCoVrCs were previously detected in several horseshoe bat species, including Rhinolophus sinicus, Rhinolophus ferrumequinum (currently R. nippon)16, Rhinolophus macrotis (currently R. episcopus)16, Rhinolophus pearsoni, and Rhinolophus pusillus, and it has been shown that they circulate not only among conspecific bats from the same colony, but also between bat species inhabiting the same caves17,20,21. The ecological niche predicted for bat SCoVrCs using a data set of 19 points (see online supplementary Table S2) is shown in Fig. 4. The AUC was 0.81. The value was  > 95% CI null-model’s AUCs (0.68), indicating that the model performs significantly better than a random model (see online supplementary Fig. S1). The highest probabilities of occurrence (highlighted in green in Fig. 4) were found in Nepal, Bhutan, Bangladesh, northeastern India, northern Myanmar, northern Vietnam, most regions of China south of the Yellow River, Taiwan, North and South Korea, and southern Japan.Figure 4Ecological niche of bat viruses related to SARS-CoV (SCoVrCs). The geographic distribution of suitable environments was predicted using the Maxent algorithm in ENMTools (see “Methods” section for details). AUC = 0.81. Black circles indicate localities used to build the distribution model (see geographic coordinates in online supplementary Table S2).Full size imageIn Southeast Asia and southern China, SCoV2rCs have currently been found in four Rhinolophus species (R. acuminatus, R. affinis, R. malayanus and R. shameli)1,6,7,8, but the greatest diversity of horseshoe bat species in mainland Southeast Asia (between 28 and 36 species)11,16 suggests that many sarbecoviruses will be discovered soon. Despite the limited data currently available on SCoV2rCs, several arguments support that bat intraspecific and interspecific transmissions also occur with SCoV2rCs. Firslty, recent genomic studies have revealed that SCoV2rCs circulate and evolve among horseshoe bats of the same colony, as five very similar genomes (nucleotide distances between 0.03% and 0.10%) were sequenced from five R. acuminatus bats collected from the same colony in eastern Thailand6, and as two genomes differing at only three nucleotide positions (distance = 0.01%) were sequenced from two R. shameli bats collected at the same cave entrance on the same night7. Secondly, the discovery of four viruses closely related to SARS-CoV-2 (between 96.2 and 91.8% of genome identity) in four different species of Rhinolophus is a strong evidence that interspecific transmission occurred several times in the past. As detailed in online supplementary Table S1, these species were collected together in several localities of Cambodia (three species in C1, C2, and C5; two species in C8), Laos (four species in L10; three species in L9; two species in L1, L5, L8, L11), and Vietnam (two speciess in V10, V9, V17, V18). These data corroborate previous studies suggesting that sarbecoviruses can be transmitted, at least occasionally, between Rhinolophus species sharing the same caves.The ecological niche of bat SCoV2rCs was firstly predicted using the four localities where bat viruses were previously detected1,6,7,8 (Fig. 5a). The highest probabilities of occurrence (highlighted in green in Fig. 5a) were found in Southeast Asia rather than in China. However, the AUC was only 0.58, and the value was  95% CI null-model’s AUCs (0.81), indicating that the model performs significantly better than a random model (see online supplementary Fig. S3). The areas showing the highest probabilities of occurrence (highlighted in green in Fig. 5b) include four main geographic areas: (i) southern Yunnan, northern Laos and bordering regions in northern Thailand and northwestern Vietnam; (ii) southern Laos, southwestern Vietnam, and northeastern Cambodia; (iii) the Cardamom Mountains in southwestern Cambodia and the East region of Thailand; and (iv) the Dawna Range in central Thailand and southeastern Myanmar.Figure 5Ecological niches of bat viruses closely related to SARS-CoV-2 (SCoV2rCs) predicted using 4 points (a) (AUC = 0.58) and 21 points (b) (AUC = 0.96). The geographic distributions of suitable environments were predicted using the Maxent algorithm in ENMTools (see “Methods” section for details). Black circles indicate localities used to build the distribution model (see geographic coordinates in online supplementary Table S1).Full size imageOur results show that bat SCoVrCs and SCoV2rCs have different ecological niches: that of SCoVrCs covers mainly China and several adjacent countries and extends to latitudes between 18° and 43°N, whereas that of SCoV2rCs covers northern mainland Southeast Asia and extends to latitudes between 10° and 24°N. Most Rhinolophus species involved in the ecological niche of SCoVrCs have to hibernate in winter when insect populations become significantly less abundant. This may be different for most Rhinolophus species involved in the ecological niche of SCoVrC2s. Since this ecological difference may be crucial for the dynamics of viral transmission among bat populations, it needs to be further studied through comparative field surveys in different regions of China and Southeast Asia. The ecological niches of SCoVrCs and SCoV2rCs slightly overlap in the zone including southern Yunnan, northern Laos, and northern Vietnam (Figs. 4, 5b). This zone corresponds to the northern edge of tropical monsoon climate23. Highly divergent sarbecoviruses of the two main lineages SCoVrCs and SCoV2rCs are expected to be found in sympatry in this area. This is confirmed by the discovery of both SCoVrCs and SCoV2rCs in horseshoe bats collected in southern Yunnan1,6,21. Collectively, these data suggest that genomic recombination between viruses of the two divergent lineages are more likely to occur in bats roosting, at least seasonally, in the caves of these regions. Since highly recombinant viruses can threaten the benefit of vaccination campaigns, southern Yunnan, northern Laos, and northern Vietnam should be the targets of closer surveillance.Mainland Southeast Asia is the cradle of diversification of bat SCoV2rCsChinese researchers have actively sought sarbecoviruses in all Chinese provinces after the 2002–2004 SARS outbreak. They found many bat SCoVrCs16,20,21 but only two SCoV2rCs1,5 and both of them were discovered in southern Yunnan, the Chinese province bordering Southeast Asia. The ecological niches predicted herein for bat sarbecoviruses suggest that SCoVrCs are dominant in China (Fig. 4) while SCoV2rCs are present mostly in Southeast Asia (Fig. 5). This means that viruses similar to SARS-CoV-2 have been circulating for several decades throughout Southeast Asia, and that different species of bats have exchanged these viruses in the caves they inhabit. The data available on human cases and deaths caused by the COVID-19 pandemic2 indirectly support the hypothesis that the cradle of diversification of bat SCoV2rCs is mainland Southeast Asia, and in particular the areas highlighted in green in Fig. 5b. Indeed, human populations in Cambodia, Laos, Thailand, and Vietnam appear to be much less affected by the COVID-19 pandemic than other countries of the region, such as Indonesia, Malaysia, Myanmar, and the Philippines (Fig. 6). This suggests that some human populations of Cambodia, Laos, Thailand, and Vietnam, in particular rural populations living in contact with wild animals for several generations, have a better immunity against SCoV2rCs because they have been regularly contaminated by bats and/or infected secondary hosts such as pangolins.Figure 6Number of COVID-19 patients per million inhabitants (in blue) and deaths per million inhabitants (in red) for the different countries of Southeast Asia. Data extracted from the Worldometers website2 on June 08, 2021. The figure was drawn in Microsoft Excel and PowerPoint (version 16.16.27).Full size imagePangolins contaminated by bats in Southeast AsiaApart from bats, the Sunda pangolin (Manis javanica) and Chinese pangolin (Manis pentadactyla) are the only wild animals in which viruses related to SARS-CoV-2 have been found so far. However, these discoveries were made in a rather special context, that of pangolin trafficking. Several sick pangolins were seized by Chinese customs in Yunnan province in 2017 (unpublished data), in Guangxi province in 2017–201824 and in Guangdong province in 201925. Even if the viruses sequenced in pangolins are not that close to SARS-CoV-2 (one was 85% identical and the other 90%), they indicate that at least two sarbecoviruses could have been imported into China well before the emergence of COVID-19 epidemic. Indeed, it has been shown that Sunda pangolins collected from different Southeast Asian regions have contaminated each other while in captivity on Chinese territory3. It has been estimated that 43% of seized pangolins were infected by at least one SARS-CoV-2-like virus3. Such a high level of viral prevalence and the symptoms of acute interstitial pneumonia detected in most dead pangolins24 indicate that captive pangolins are highly permissive to infection by SARS-CoV-2-like viruses. The question remained on how the Sunda pangolins became infected initially. Could it have been in their natural Southeast Asian environment, before being captured? The discovery of two new viruses close to SARS-CoV-2 in bats from Cambodia and Thailand7,8 supports this hypothesis, as Rhinolophus bats and pangolins can meet, at least occasionally, in forests of Southeast Asia, possibly in caves, tree hollows or burrows. Further substantiating this hypothesis, the geographic distribution of Manis javanica26 overlaps the ecological niche here predicted for bat SCoV2rCs (Fig. 5), and SARS-CoV-2 neutralizing antibodies have been recently detected in one of the ten pangolin sera sampled from February to July 2020 from three wildlife checkpoint stations in Thailand6. Collectively, these data strengthen the hypothesis that pangolin trafficking is responsible for multiple exports of viruses related to SARS-CoV-2 to China3. More

  • in

    Gene body DNA methylation in seagrasses: inter- and intraspecific differences and interaction with transcriptome plasticity under heat stress

    1.Merilä, J. & Hendry, A. P. Climate change, adaptation, and phenotypic plasticity: the problem and the evidence. Evol. Appl. 7, 1–14. https://doi.org/10.1111/eva.12137 (2014).Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    2.Reusch, T. B. Climate change in the oceans: evolutionary versus phenotypically plastic responses of marine animals and plants. Evol. Appl. 7, 104–122. https://doi.org/10.1111/eva.12109 (2014).Article 
    PubMed 

    Google Scholar 
    3.Pazzaglia, J., Reusch, T. B., Terlizzi, A., Marín‐Guirao, L. & Procaccini, G. Phenotypic plasticity under rapid global changes: the intrinsic force for future seagrasses survival. Evol. Appl. (2021).4.Lopez-Maury, L., Marguerat, S. & Baehler, J. Tuning gene expression to changing environments: from rapid responses to evolutionary adaptation. Nat. Rev. Genet. 9, 583–593 (2008).CAS 
    PubMed 
    Article 

    Google Scholar 
    5.Mäkinen, H., Papakostas, S., Vøllestad, L. A., Leder, E. H. & Primmer, C. R. Plastic and evolutionary gene expression responses are correlated in European grayling (Thymallus thymallus) subpopulations adapted to different thermal environments. J. Hered. 107, 82–89 (2016).PubMed 
    Article 
    CAS 

    Google Scholar 
    6.Alonso, C., Pérez, R., Bazaga, P., Medrano, M. & Herrera, C. M. MSAP markers and global cytosine methylation in plants: a literature survey and comparative analysis for a wild-growing species. Mol. Ecol. Resour. 16, 80–90 (2016).CAS 
    PubMed 
    Article 

    Google Scholar 
    7.Jeremias, G. et al. Synthesizing the role of epigenetics in the response and adaptation of species to climate change in freshwater ecosystems. Mol. Ecol. 27, 2790–2806 (2018).PubMed 
    Article 

    Google Scholar 
    8.Nicotra, A. B. et al. Adaptive plasticity and epigenetic variation in response to warming in an Alpine plant. Ecol. Evol. 5, 634–647 (2015).PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    9.Kelly, S., Panhuis, T. & Stoehr, A. (2012).10.Thorson, J. L. et al. Epigenetics and adaptive phenotypic variation between habitats in an asexual snail. Sci. Rep. 7, 1–11 (2017).CAS 
    Article 

    Google Scholar 
    11.Rey, O., Danchin, E., Mirouze, M., Loot, C. & Blanchet, S. Adaptation to global change: a transposable element–epigenetics perspective. Trends Ecol. Evol. 31, 514–526. https://doi.org/10.1016/j.tree.2016.03.013 (2016).Article 
    PubMed 

    Google Scholar 
    12.Law, J. A. & Jacobsen, S. E. Establishing, maintaining and modifying DNA methylation patterns in plants and animals. Nat. Rev. Genet. 11, 204–220 (2010).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    13.Zemach, A., McDaniel, I. E., Silva, P. & Zilberman, D. Genome-wide evolutionary analysis of eukaryotic DNA methylation. Science 328, 916–919 (2010).CAS 
    PubMed 
    Article 
    ADS 

    Google Scholar 
    14.Niederhuth, C. E. et al. Widespread natural variation of DNA methylation within angiosperms. Genome Biol. 17, 1–19 (2016).Article 
    CAS 

    Google Scholar 
    15.Zhang, X. et al. Genome-wide high-resolution mapping and functional analysis of DNA methylation in Arabidopsis. Cell 126, 1189–1201 (2006).CAS 
    Article 

    Google Scholar 
    16.Bewick, A. J. et al. On the origin and evolutionary consequences of gene body DNA methylation. Proc. Natl. Acad. Sci. 113, 9111–9116 (2016).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    17.Bewick, A. J. & Schmitz, R. J. Gene body DNA methylation in plants. Curr. Opin. Plant Biol. 36, 103–110 (2017).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    18.Sarda, S., Zeng, J., Hunt, B. G. & Yi, S. V. The evolution of invertebrate gene body methylation. Mol. Biol. Evol. 29, 1907–1916 (2012).CAS 
    PubMed 
    Article 

    Google Scholar 
    19.Takuno, S. & Gaut, B. S. Body-methylated genes in Arabidopsis thaliana are functionally important and evolve slowly. Mol. Biol. Evol. 29, 219–227 (2012).CAS 
    PubMed 
    Article 

    Google Scholar 
    20.Takuno, S. & Gaut, B. S. Gene body methylation is conserved between plant orthologs and is of evolutionary consequence. Proc. Natl. Acad. Sci. 110, 1797–1802 (2013).CAS 
    PubMed 
    PubMed Central 
    Article 
    ADS 

    Google Scholar 
    21.Takuno, S., Ran, J.-H. & Gaut, B. S. Evolutionary patterns of genic DNA methylation vary across land plants. Nat. Plants 2, 1–7 (2016).Article 
    CAS 

    Google Scholar 
    22.Wendte, J. M. et al. Epimutations are associated with CHROMOMETHYLASE 3-induced de novo DNA methylation. Elife 8, e47891 (2019).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    23.Aceituno, F. F., Moseyko, N., Rhee, S. Y. & Gutiérrez, R. A. The rules of gene expression in plants: organ identity and gene body methylation are key factors for regulation of gene expression in Arabidopsis thaliana. BMC Genomics 9, 438 (2008).PubMed 
    PubMed Central 
    Article 
    CAS 

    Google Scholar 
    24.Elango, N., Hunt, B. G., Goodisman, M. A. & Soojin, V. Y. DNA methylation is widespread and associated with differential gene expression in castes of the honeybee, Apis mellifera. Proc. Natl. Acad. Sci. 106, 11206–11211 (2009).CAS 
    PubMed 
    PubMed Central 
    Article 
    ADS 

    Google Scholar 
    25.Gavery, M. R. & Roberts, S. B. DNA methylation patterns provide insight into epigenetic regulation in the Pacific oyster (Crassostrea gigas). BMC Genomics 11, 1–9 (2010).Article 
    CAS 

    Google Scholar 
    26.Zilberman, D., Gehring, M., Tran, R. K., Ballinger, T. & Henikoff, S. Genome-wide analysis of Arabidopsis thaliana DNA methylation uncovers an interdependence between methylation and transcription. Nat. Genet. 39, 61–69 (2007).CAS 
    PubMed 
    Article 

    Google Scholar 
    27.Coleman-Derr, D. & Zilberman, D. in Cold Spring Harbor symposia on quantitative biology. 147–154 (Cold Spring Harbor Laboratory Press).28.Kim, M. Y. & Zilberman, D. DNA methylation as a system of plant genomic immunity. Trends Plant Sci. 19, 320–326 (2014).CAS 
    PubMed 
    Article 

    Google Scholar 
    29.Muyle, A. & Gaut, B. S. Loss of gene body methylation in Eutrema salsugineum is associated with reduced gene expression. Mol. Biol. Evol. 36, 155–158 (2019).CAS 
    PubMed 
    Article 

    Google Scholar 
    30.Roberts, S. B. & Gavery, M. R. Is there a relationship between DNA methylation and phenotypic plasticity in invertebrates?. Front. Physiol. 2, 116 (2012).PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    31.Dimond, J. L. & Roberts, S. B. Germline DNA methylation in reef corals: patterns and potential roles in response to environmental change. Mol. Ecol. 25, 1895–1904 (2016).CAS 
    PubMed 
    Article 

    Google Scholar 
    32.Dixon, G. B., Bay, L. K. & Matz, M. V. Bimodal signatures of germline methylation are linked with gene expression plasticity in the coral Acropora millepora. BMC Genomics 15, 1–11 (2014).Article 
    CAS 

    Google Scholar 
    33.Bird, A. P. DNA methylation and the frequency of CpG in animal DNA. Nucleic Acids Res. 8, 1499–1504 (1980).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    34.Sved, J. & Bird, A. The expected equilibrium of the CpG dinucleotide in vertebrate genomes under a mutation model. Proc. Natl. Acad. Sci. 87, 4692–4696 (1990).CAS 
    PubMed 
    PubMed Central 
    Article 
    ADS 

    Google Scholar 
    35.Suzuki, M. M., Kerr, A. R., De Sousa, D. & Bird, A. CpG methylation is targeted to transcription units in an invertebrate genome. Genome Res. 17, 625–631 (2007).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    36.Weber, M. et al. Distribution, silencing potential and evolutionary impact of promoter DNA methylation in the human genome. Nat. Genet. 39, 457–466 (2007).CAS 
    PubMed 
    Article 

    Google Scholar 
    37.Glastad, K., Hunt, B. G., Yi, S. & Goodisman, M. DNA methylation in insects: on the brink of the epigenomic era. Insect Mol. Biol. 20, 553–565 (2011).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    38.Aliaga, B., Bulla, I., Mouahid, G., Duval, D. & Grunau, C. Universality of the DNA methylation codes in Eucaryotes. Sci. Rep. 9, 1–11 (2019).CAS 
    Article 

    Google Scholar 
    39.Asselman, J., De Coninck, D. I., Pfrender, M. E. & De Schamphelaere, K. A. Gene body methylation patterns in Daphnia are associated with gene family size. Genome Biol Evol 8, 1185–1196 (2016).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    40.Park, J. et al. Comparative analyses of DNA methylation and sequence evolution using Nasonia genomes. Mol. Biol. Evol. 28, 3345–3354 (2011).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    41.Olsen, J. L. et al. The genome of the seagrass Zostera marina reveals angiosperm adaptation to the sea. Nature 530, 331–335. https://doi.org/10.1038/nature16548 (2016).CAS 
    Article 
    PubMed 
    ADS 

    Google Scholar 
    42.Costanza, R. et al. Changes in the global value of ecosystem services. Glob. Environ. Chang. 26, 152–158. https://doi.org/10.1016/j.gloenvcha.2014.04.002 (2014).Article 

    Google Scholar 
    43.Nordlund, L. M., Koch, E. W., Barbier, E. B. & Creed, J. C. Correction: Seagrass ecosystem services and their variability across genera and geographical regions. PLoS ONE 12, e0169942 (2017).PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    44.Orth, R. J. et al. A global crisis for seagrass ecosystems. Bioscience 56, 987–996. https://doi.org/10.1641/0006-3568(2006)56[987:agcfse]2.0.co;2 (2006).Article 

    Google Scholar 
    45.Waycott, M. et al. Accelerating loss of seagrasses across the globe threatens coastal ecosystems. Proc. Natl. Acad. Sci. 106, 12377–12381. https://doi.org/10.1073/pnas.0905620106 (2009).Article 
    PubMed 
    PubMed Central 
    ADS 

    Google Scholar 
    46.Koch, M., Bowes, G., Ross, C. & Zhang, X. H. Climate change and ocean acidification effects on seagrasses and marine macroalgae. Glob. Change Biol. 19, 103–132. https://doi.org/10.1111/j.1365-2486.2012.02791.x (2013).Article 
    ADS 

    Google Scholar 
    47.Marbà, N. & Duarte, C. M. Mediterranean warming triggers seagrass (Posidonia oceanica) shoot mortality. Glob. Change Biol. 16, 2366–2375. https://doi.org/10.1111/j.1365-2486.2009.02130.x (2010).Article 
    ADS 

    Google Scholar 
    48.Thomson, J. A. et al. Extreme temperatures, foundation species, and abrupt ecosystem change: an example from an iconic seagrass ecosystem. Glob. Change Biol. 21, 1463–1474. https://doi.org/10.1111/gcb.12694 (2014).Article 
    ADS 

    Google Scholar 
    49.Maxwell, P. S. et al. Phenotypic plasticity promotes persistence following severe events: physiological and morphological responses of seagrass to flooding. J. Ecol. 102, 54–64 (2014).Article 

    Google Scholar 
    50.Marín-Guirao, L., Ruiz, J. M., Dattolo, E., Garcia-Munoz, R. & Procaccini, G. Physiological and molecular evidence of differential short-term heat tolerance in Mediterranean seagrasses. Sci. Rep. 6, 28615. https://doi.org/10.1038/srep28615 (2016).CAS 
    Article 
    PubMed 
    PubMed Central 
    ADS 

    Google Scholar 
    51.Sandoval-Gil, J. M., Ruiz, J. M., Marin-Guirao, L., Bernardeau-Esteller, J. & Sanchez-Lizaso, J. L. Ecophysiological plasticity of shallow and deep populations of the Mediterranean seagrasses Posidonia oceanica and Cymodocea nodosa in response to hypersaline stress. Mar. Environ. Res. 95, 39–61. https://doi.org/10.1016/j.marenvres.2013.12.011 (2014).CAS 
    Article 
    PubMed 

    Google Scholar 
    52.Franssen, S. et al. Transcriptomic resilience to global warming in the seagrass Zostera marina, a marine foundation species. Proc. Natl. Acad. Sci. USA 108, 19276–19281 (2011).CAS 
    PubMed 
    PubMed Central 
    Article 
    ADS 

    Google Scholar 
    53.Jueterbock, A. et al. Phylogeographic differentiation versus transcriptomic adaptation to warm temperatures in Zostera marina, a globally important seagrass. Mol. Ecol. 25, 5396–5411 (2016).CAS 
    PubMed 
    Article 

    Google Scholar 
    54.Marín-Guirao, L., Entrambasaguas, L., Dattolo, E., Ruiz, J. M. & Procaccini, G. Molecular mechanisms behind the physiological resistance to intense transient warming in an iconic marine plant. Front. Plant Sci. https://doi.org/10.3389/fpls.2017.01142 (2017).Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    55.Lee, H. et al. The genome of a southern hemisphere seagrass species (Zostera muelleri). Plant Physiol. (2016).56.Greco, M., Chiappetta, A., Bruno, L. & Bitonti, M. B. Effects of light deficiency on genome methylation in Posidonia oceanica. Mar. Ecol. Prog. Ser. 473, 103–114 (2013).CAS 
    Article 
    ADS 

    Google Scholar 
    57.Greco, M., Chiappetta, A., Bruno, L. & Bitonti, M. B. In Posidonia oceanica cadmium induces changes in DNA methylation and chromatin patterning. J. Exp. Bot. 63, 695–709. https://doi.org/10.1093/jxb/err313 (2012).CAS 
    Article 
    PubMed 

    Google Scholar 
    58.Ruocco, M., De Luca, P., Marín-Guirao, L. & Procaccini, G. Differential leaf age-dependent thermal plasticity in the keystone seagrass Posidonia oceanica. Front. Plant Sci. https://doi.org/10.3389/fpls.2019.01556 (2019).Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    59.Ruocco, M., Marín-Guirao, L. & Procaccini, G. Within- and among-leaf variations in photo-physiological functions, gene expression and DNA methylation patterns in the large-sized seagrass Posidonia oceanica. Mar. Biol. 166, 24. https://doi.org/10.1007/s00227-019-3482-8 (2019).CAS 
    Article 

    Google Scholar 
    60.Ruocco, M. et al. A king and vassals’ tale: Molecular signatures of clonal integration in Posidonia oceanica under chronic light shortage. J. Ecol. (2020).61.Jueterbock, A. et al. The seagrass methylome is associated with variation in photosynthetic performance among clonal shoots. Front. Plant Sci. 11 (2020).62.Marín-Guirao, L. et al. Carbon economy of Mediterranean seagrasses in response to thermal stress. Mar. Pollut. Bull. 135, 617–629 (2018).PubMed 
    Article 
    CAS 

    Google Scholar 
    63.Beca-Carretero, P. et al. Effects of an experimental heat wave on fatty acid composition in two Mediterranean seagrass species. Mar. Pollut. Bull. 134, 27–37 (2018).CAS 
    PubMed 
    Article 

    Google Scholar 
    64.Angers, B., Castonguay, E. & Massicotte, R. Environmentally induced phenotypes and DNA methylation: how to deal with unpredictable conditions until the next generation and after. Mol. Ecol. 19, 1283–1295 (2010).CAS 
    PubMed 
    Article 

    Google Scholar 
    65.Dubin, M. J. et al. DNA methylation in Arabidopsis has a genetic basis and shows evidence of local adaptation. Elife 4, e05255 (2015).PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    66.Kawakatsu, T. et al. Epigenomic diversity in a global collection of Arabidopsis thaliana accessions. Cell 166, 492–505 (2016).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    67.Smith, Z. D. & Meissner, A. DNA methylation: roles in mammalian development. Nat. Rev. Genet. 14, 204–220 (2013).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    68.Serres-Giardi, L., Belkhir, K., David, J. & Glémin, S. Patterns and evolution of nucleotide landscapes in seed plants. Plant Cell 24, 1379–1397 (2012).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    69.Tatarinova, T., Elhaik, E. & Pellegrini, M. Cross-species analysis of genic GC3 content and DNA methylation patterns. Genome Biol. Evol. 5, 1443–1456 (2013).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    70.Vining, K. J. et al. Dynamic DNA cytosine methylation in the Populus trichocarpa genome: tissue-level variation and relationship to gene expression. BMC Genomics 13, 27 (2012).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    71.Lyko, F. et al. The honey bee epigenomes: differential methylation of brain DNA in queens and workers. PLoS Biol 8, 1506 (2010).Article 
    CAS 

    Google Scholar 
    72.Cortijo, S., Aydin, Z., Ahnert, S. & Locke, J. C. Widespread inter-individual gene expression variability in Arabidopsis thaliana. Mol. Syst. Biol. 15, e8591 (2019).PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    73.Procaccini, G., Olsen, J. L. & Reusch, T. B. H. Contribution of genetics and genomics to seagrass biology and conservation. J. Exp. Mar. Biol. Ecol. 350, 234–259. https://doi.org/10.1016/j.jembe.2007.05.035 (2007).CAS 
    Article 

    Google Scholar 
    74.Alberto, F. et al. Genetic differentiation and secondary contact zone in the seagrass Cymodocea nodosa across the Mediterranean-Atlantic transition region. J. Biogeogr. 35, 1279–1294 (2008).Article 

    Google Scholar 
    75.Becker, C. et al. Spontaneous epigenetic variation in the Arabidopsis thaliana methylome. Nature 480, 245–249 (2011).CAS 
    PubMed 
    PubMed Central 
    Article 
    ADS 

    Google Scholar 
    76.Schmitz, R. J. et al. Patterns of population epigenomic diversity. Nature 495, 193–198 (2013).CAS 
    PubMed 
    PubMed Central 
    Article 
    ADS 

    Google Scholar 
    77.Yi, S. V. Insights into epigenome evolution from animal and plant methylomes. Genome Biol. Evol. 9, 3189–3201 (2017).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    78.Jahnke, M. et al. Adaptive responses along a depth and a latitudinal gradient in the endemic seagrass Posidonia oceanica. Heredity https://doi.org/10.1038/s41437-018-0103-0 (2018).Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    79.Tuya, F. et al. Biogeographical scenarios modulate seagrass resistance to small-scale perturbations. J. Ecol. 107, 1263–1275 (2019).Article 

    Google Scholar 
    80.Gao, G. et al. Comparison of the heat stress induced variations in DNA methylation between heat-tolerant and heat-sensitive rapeseed seedlings. Breed. Sci. 64, 125–133 (2014).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    81.Dowen, R. H. et al. Widespread dynamic DNA methylation in response to biotic stress. Proc. Natl. Acad. Sci. 109, E2183–E2191 (2012).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    82.Wada, Y., Miyamoto, K., Kusano, T. & Sano, H. Association between up-regulation of stress-responsive genes and hypomethylation of genomic DNA in tobacco plants. Mol. Genet. Genomics 271, 658–666 (2004).CAS 
    PubMed 
    Article 

    Google Scholar 
    83.Yaish, M. W., Colasanti, J. & Rothstein, S. J. The role of epigenetic processes in controlling flowering time in plants exposed to stress. J. Exp. Bot. 62, 3727–3735 (2011).CAS 
    PubMed 
    Article 

    Google Scholar 
    84.Secco, D. et al. Stress induced gene expression drives transient DNA methylation changes at adjacent repetitive elements. Elife 4, e09343 (2015).PubMed Central 
    Article 
    PubMed 

    Google Scholar 
    85.Marín-Guirao, L., Entrambasaguas, L., Ruiz, J. M. & Procaccini, G. Heat-stress induced flowering can be a potential adaptive response to ocean warming for the iconic seagrass Posidonia oceanica. Mol. Ecol. 28, 2486–2501. https://doi.org/10.1111/mec.15089 (2019).Article 
    PubMed 

    Google Scholar 
    86.Nguyen, H. M. et al. Stress memory in seagrasses: first insight into the effects of thermal priming and the role of epigenetic modifications. Front. Plant Sci. 11, 494 (2020).PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    87.Pikaard, C. S. & Scheid, O. M. Epigenetic regulation in plants. Cold Spring Harbor Perspect. Biol. 6, a019315 (2014).Article 
    CAS 

    Google Scholar 
    88.Yu, Y. et al. Cytosine methylation alteration in natural populations of Leymus chinensis induced by multiple abiotic stresses. PLoS ONE 8, e55772 (2013).CAS 
    PubMed 
    PubMed Central 
    Article 
    ADS 

    Google Scholar 
    89.Liu, R. & Lang, Z. The mechanism and function of active DNA demethylation in plants. J. Integr. Plant. Biol. 62, 148–159 (2020).CAS 
    PubMed 
    Article 

    Google Scholar 
    90.Xu, X. et al. A CRISPR-based approach for targeted DNA demethylation. Cell Discovery 2, 1–12 (2016).
    Google Scholar 
    91.Arnaud-Haond, S. et al. Implications of extreme life span in clonal organisms: millenary clones in meadows of the threatened seagrass Posidonia oceanica. PLoS ONE 7, e30454. https://doi.org/10.1371/journal.pone.0030454 (2012).CAS 
    Article 
    PubMed 
    PubMed Central 
    ADS 

    Google Scholar 
    92.Mascaró, O., Romero, J. & Pérez, M. Seasonal uncoupling of demographic processes in a marine clonal plant. Estuar. Coast. Shelf Sci. 142, 23–31 (2014).Article 
    ADS 

    Google Scholar 
    93.Olesen, B., Enríquez, S., Duarte, C. M. & Sand-Jensen, K. Depth-acclimation of photosynthesis, morphology and demography of Posidonia oceanica and Cymodocea nodosa in the Spanish Mediterranean Sea. Mar. Ecol. Prog. Ser. 236, 89–97. https://doi.org/10.3354/meps236089 (2002).Article 
    ADS 

    Google Scholar 
    94.Ruocco, M. et al. Genomewide transcriptional reprogramming in the seagrass Cymodocea nodosa under experimental ocean acidification. Mol. Ecol. 26, 4241–4259. https://doi.org/10.1111/mec.14204 (2017).CAS 
    Article 
    PubMed 

    Google Scholar 
    95.Fraley, C. & Raftery, A. E. Model-based methods of classification: using the mclust software in chemometrics. J. Stat. Softw. 18, 1–13 (2007).Article 

    Google Scholar 
    96.R Core Team (ISBN 3-900051-07-0, 2012).97.Benaglia, T., Chauveau, D., Hunter, D., Young, D. mixtools: an R package for analyzing finite mixture models (2009).98.Langmead, B., Trapnell, C., Pop, M. & Salzberg, S. Ultrafast and memory-efficient alignment of short DNA sequences to the human genome. Genome Biol. 10, R25 (2009).PubMed 
    PubMed Central 
    Article 
    CAS 

    Google Scholar 
    99.Li, B. & Dewey, C. N. RSEM: accurate transcript quantification from RNA-Seq data with or without a reference genome. BMC Bioinformat. 12, 323 (2011).CAS 
    Article 

    Google Scholar 
    100.Robinson, M. D., McCarthy, D. J. & Smyth, G. K. edgeR: a bioconductor package for differential expression analysis of digital gene expression data. Bioinformatics 26, 139–140. https://doi.org/10.1093/bioinformatics/btp616 (2010).CAS 
    Article 
    PubMed 
    PubMed Central 

    Google Scholar  More

  • in

    Wet-dry cycles protect surface-colonizing bacteria from major antibiotic classes

    1.Or D, Smets BF, Wraith J, Dechesne A, Friedman S. Physical constraints affecting bacterial habitats and activity in unsaturated porous media–a review. Adv Water Resour. 2007;30:1505–27.Article 

    Google Scholar 
    2.Burkhardt J, Hunsche M. “Breath figures” on leaf surfaces—formation and effects of microscopic leaf wetness. Front plant Sci. 2013;4:422.PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    3.Wolf AB, Vos M, de Boer W, Kowalchuk GA. Impact of matric potential and pore size distribution on growth dynamics of filamentous and non-filamentous soil bacteria. PloS One. 2013;8:e83661.PubMed 
    PubMed Central 
    Article 
    CAS 

    Google Scholar 
    4.Forsberg KJ, Reyes A, Wang B, Selleck EM, Sommer MO, Dantas G. The shared antibiotic resistome of soil bacteria and human pathogens. Science. 2012;337:1107–11.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    5.Williams S, Vickers J. The ecology of antibiotic production. Microb Ecol. 1986;12:43–52.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    6.Raaijmakers JM, Weller DM, Thomashow LS. Frequency of antibiotic-producing Pseudomonas spp. in natural environments. Appl Environ Microbiol. 1997;63:881–7.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    7.Wells JS, Hunter JC, Astle GL, Sherwood JC, Ricca cM, Trejo WH, et al. Distribution of β-lactam and β-lactone producing bacteria in nature. The. J Antibiot. 1982;35:814–21.CAS 
    Article 

    Google Scholar 
    8.Kinkel LL, Schlatter DC, Xiao K, Baines AD. Sympatric inhibition and niche differentiation suggest alternative coevolutionary trajectories among Streptomycetes. ISME J. 2014;8:249–56.CAS 
    PubMed 
    Article 

    Google Scholar 
    9.Vetsigian K, Jajoo R, Kishony R. Structure and evolution of Streptomyces interaction networks in soil and in silico. PLoS Biol. 2011;9:e1001184.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    10.Traxler MF, Kolter R. Natural products in soil microbe interactions and evolution. Nat Prod Rep. 2015;32:956–70.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    11.Franklin AM, Aga DS, Cytryn E, Durso LM, McLain JE, Pruden A, et al. Antibiotics in agroecosystems: introduction to the special section. J Environ Qual. 2016;45:377–93.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    12.Jechalke S, Heuer H, Siemens J, Amelung W, Smalla K. Fate and effects of veterinary antibiotics in soil. Trends Microbiol. 2014;22:536–45.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    13.Mompelat S, Le Bot B, Thomas O. Occurrence and fate of pharmaceutical products and by-products, from resource to drinking water. Environ Int. 2009;35:803–14.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    14.Kelsic ED, Zhao J, Vetsigian K, Kishony R. Counteraction of antibiotic production and degradation stabilizes microbial communities. Nature. 2015;521:516–9.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    15.Cordero OX, Wildschutte H, Kirkup B, Proehl S, Ngo L, Hussain F, et al. Ecological populations of bacteria act as socially cohesive units of antibiotic production and resistance. Science. 2012;337:1228–31.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    16.Schlatter DC, Song Z, Vaz-Jauri P, Kinkel LL. Inhibitory interaction networks among coevolved Streptomyces populations from prairie soils. Plos One. 2019;14:e0223779.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    17.Abrudan MI, Smakman F, Grimbergen AJ, Westhoff S, Miller EL, Van Wezel GP, et al. Socially mediated induction and suppression of antibiosis during bacterial coexistence. Proc Natl Acad Sci. 2015;112:11054–9.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    18.Brauner A, Fridman O, Gefen O, Balaban NQ. Distinguishing between resistance, tolerance and persistence to antibiotic treatment. Nat Rev Microbiol. 2016;14:320.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    19.Andersson DI, Levin BR. The biological cost of antibiotic resistance. Curr Opin Microbiol. 1999;2:489–93.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    20.Handwerger S, Tomasz A. Antibiotic tolerance among clinical isolates of bacteria. Annu Rev Pharmacol Toxicol. 1985;25:349–80.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    21.Kester JC, Fortune SM. Persisters and beyond: mechanisms of phenotypic drug resistance and drug tolerance in bacteria. Crit Rev Biochem Mol Biol. 2014;49:91–101.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    22.Wood KB, Cluzel P. Trade-offs between drug toxicity and benefit in the multi-antibiotic resistance system underlie optimal growth of E. coli. BMC Syst Biol. 2012;6:1–11.Article 

    Google Scholar 
    23.Nguyen D, Joshi-Datar A, Lepine F, Bauerle E, Olakanmi O, Beer K, et al. Active starvation responses mediate antibiotic tolerance in biofilms and nutrient-limited bacteria. Science. 2011;334:982–6.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    24.Meredith HR, Srimani JK, Lee AJ, Lopatkin AJ, You L. Collective antibiotic tolerance: mechanisms, dynamics and intervention. Nat Chem Biol. 2015;11:182.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    25.Nagarajan R, Boeck LD, Gorman M, Hamill RL, Higgens CE, Hoehn MM, et al. beta.-Lactam antibiotics from Streptomyces. J Am Chem Soc. 1971;93:2308–10.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    26.Imada A, Kitano K, Kintaka K, Muroi M, Asai M. Sulfazecin and isosulfazecin, novel β-lactam antibiotics of bacterial origin. Nature. 1981;289:590–1.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    27.Sykes R, Cimarusti C, Bonner D, Bush K, Floyd D, Georgopapadakou N, et al. Monocyclic β-lactam antibiotics produced by bacteria. Nature. 1981;291:489.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    28.Wells JS, TREJO WH, PRINCIPE PA, Bush K, Georgopapadakou N, Bonner DP, et al. EM5400, a family of monobactam antibiotics produced by Agrobacterium radiobacter. J Antibiot. 1982;35:295–9.CAS 
    Article 

    Google Scholar 
    29.ThaKurIa B, Lahon K. The beta lactam antibiotics as an empirical therapy in a developing country: An update on their current status and recommendations to counter the resistance against them. J Clin Diagn Res. 2013;7:1207.PubMed 
    PubMed Central 

    Google Scholar 
    30.Russ D, Glaser F, Tamar ES, Yelin I, Baym M, Kelsic ED, et al. Escape mutations circumvent a tradeoff between resistance to a beta-lactam and resistance to a beta-lactamase inhibitor. Nat Commun. 2020;11:1–9.Article 
    CAS 

    Google Scholar 
    31.Grinberg M, Orevi T, Steinberg S, Kashtan N. Bacterial survival in microscopic surface wetness. eLife. 2019;8:e48508.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    32.Orevi T, Kashtan N. Life in a droplet: microbial ecology in microscopic surface wetness. Front Microbiol. 2021;12:797.Article 

    Google Scholar 
    33.Mauer LJ, Taylor LS. Water-solids interactions: deliquescence. Annu Rev food Sci Technol. 2010;1:41–63.CAS 
    PubMed 
    Article 

    Google Scholar 
    34.Wise ME, Martin ST, Russell LM, Buseck PR. Water uptake by NaCl particles prior to deliquescence and the phase rule. Aerosol Sci Technol. 2008;42:281–94.CAS 
    Article 

    Google Scholar 
    35.Burkhardt J, Koch K, Kaiser H. Deliquescence of deposited atmospheric particles on leaf surfaces. J Water, Air Soil Pollut: Focus. 2001;1:313–21.CAS 
    Article 

    Google Scholar 
    36.Beattie GA. Water relations in the Interaction of foliar bacterial pathogens with plants. Annu Rev Phytopathol. 2011;49:533–55.CAS 
    PubMed 
    Article 

    Google Scholar 
    37.Davila AF, Hawes I, Ascaso C, Wierzchos J. Salt deliquescence drives photosynthesis in the hyperarid A tacama D esert. Environ Microbiol Rep. 2013;5:583–7.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    38.Dai S, Shin H, Santamarina JC. Formation and development of salt crusts on soil surfaces. Acta Geotechnica. 2016;11:1103–9.Article 

    Google Scholar 
    39.Trechsel HR. Moisture control in buildings. ASTM International; West Conshohocken, PA19428-2959, USA; 1994.40.Schwartz-Narbonne H, Donaldson DJ. Water uptake by indoor surface films. Sci Rep. 2019;9:1–10.CAS 
    Article 

    Google Scholar 
    41.Patrick D, Findon G, Miller T. Residual moisture determines the level of touch-contact-associated bacterial transfer following hand washing. Epidemiol Infect. 1997;119:319–25.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    42.Tang IN, Munkelwitz HR. Composition and temperature dependence of the deliquescence properties of hygroscopic aerosols. Atmos Environ Part A Gen Top. 1993;27:467–73.Article 

    Google Scholar 
    43.Pöschl U. Atmospheric aerosols: composition, transformation, climate and health effects. Angew Chem Int Ed. 2005;44:7520–40.Article 
    CAS 

    Google Scholar 
    44.Tecon R. Bacterial survival: life on a leaf. eLife. 2019;8:e52123.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    45.Vejerano EP, Marr LC. Physico-chemical characteristics of evaporating respiratory fluid droplets. J R Soc Interface. 2018;15:20170939.PubMed 
    PubMed Central 
    Article 
    CAS 

    Google Scholar 
    46.Rubasinghege G, Grassian VH. Role (s) of adsorbed water in the surface chemistry of environmental interfaces. Chem Commun. 2013;49:3071–94.CAS 
    Article 

    Google Scholar 
    47.Campbell TD, Febrian R, McCarthy JT, Kleinschmidt HE, Forsythe JG, Bracher PJ. Prebiotic condensation through wet–dry cycling regulated by deliquescence. Nat Commun. 2019;10:1–7.Article 
    CAS 

    Google Scholar 
    48.Alsved M, Holm S, Christiansen S, Smidt M, Rosati B, Ling M, et al. Effect of aerosolization and drying on the viability of pseudomonas syringae cells. Front Microbiol. 2018;9:3086.PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    49.Xie X, Li Y, Zhang T, Fang HH. Bacterial survival in evaporating deposited droplets on a teflon-coated surface. Appl Microbiol Biotechnol. 2006;73:703–12.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    50.Runkel S, Wells HC, Rowley G. Living with stress: a lesson from the enteric pathogen Salmonella enterica. Adv Appl Microbiol. 2013;83:87–144.51.Amaeze N, Akinbobola A, Chukwuemeka V, Abalkhaila A, Ramage G, Kean R, et al. Development of a high throughput and low cost model for the study of semi-dry biofilms. Biofouling. 2020:36:403–15.52.Tuomanen E, Cozens R, Tosch W, Zak O, Tomasz A. The rate of killing of Escherichia coli byβ-lactam antibiotics is strictly proportional to the rate of bacterial growth. Microbiology. 1986;132:1297–304.CAS 
    Article 

    Google Scholar 
    53.Eng R, Padberg F, Smith S, Tan E, Cherubin C. Bactericidal effects of antibiotics on slowly growing and nongrowing bacteria. Antimicrobial Agents Chemother. 1991;35:1824–8.CAS 
    Article 

    Google Scholar 
    54.Lee S, Foley E, Epstein JA. Mode of action of penicillin: I. Bacterial growth and penicillin activity—Staphylococcus aureus FDA. J Bacteriol. 1944;48:393.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    55.Lopatkin AJ, Stokes JM, Zheng EJ, Yang JH, Takahashi MK, You L, et al. Bacterial metabolic state more accurately predicts antibiotic lethality than growth rate. Nat Microbiol. 2019;4:2109–17.56.Yoon H, Park B-Y, Oh M-H, Choi K-H, Yoon Y. Effect of NaCl on heat resistance, antibiotic susceptibility, and Caco-2 cell invasion of Salmonella. BioMed Res Int. 2013;2013:274096.57.Zhu M, Dai X. High salt cross-protects Escherichia coli from antibiotic treatment through increasing efflux pump expression. mSphere 3: e00095-18. mSphere. 2018;3:e00095–18.CAS 
    PubMed 
    PubMed Central 

    Google Scholar 
    58.Lee AJ, Wang S, Meredith HR, Zhuang B, Dai Z, You L. Robust, linear correlations between growth rates and β-lactam–mediated lysis rates. Proc Natl Acad Sci. 2018;115:4069–74.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    59.Loftin KA, Adams CD, Meyer MT, Surampalli R. Effects of ionic strength, temperature, and pH on degradation of selected antibiotics. J Environ Qual. 2008;37:378–86.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    60.Thonus IP, Fontijne P, Michel MF. Ampicillin susceptibility and ampicillin-induced killing rate of Escherichia coli. Antimicrobial Agents Chemother. 1982;22:386–90.CAS 
    Article 

    Google Scholar 
    61.Cho H, Uehara T, Bernhardt TG. Beta-lactam antibiotics induce a lethal malfunctioning of the bacterial cell wall synthesis machinery. Cell. 2014;159:1300–11.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    62.Yao Z, Kahne D, Kishony R. Distinct single-cell morphological dynamics under beta-lactam antibiotics. Mol Cell. 2012;48:705–12.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    63.Battesti A, Majdalani N, Gottesman S. The RpoS-mediated general stress response in Escherichia coli. Annu Rev Microbiol. 2011;65:189–213.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    64.Bernier SP, Lebeaux D, DeFrancesco AS, Valomon A, Soubigou G, Coppée J-Y, et al. Starvation, together with the SOS response, mediates high biofilm-specific tolerance to the fluoroquinolone ofloxacin. PLoS Genet. 2013;9:e1003144.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    65.Pu Y, Zhao Z, Li Y, Zou J, Ma Q, Zhao Y, et al. Enhanced efflux activity facilitates drug tolerance in dormant bacterial cells. Mol Cell. 2016;62:284–94.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    66.Martins D, McKay G, Sampathkumar G, Khakimova M, English AM, Nguyen D. Superoxide dismutase activity confers (p) ppGpp-mediated antibiotic tolerance to stationary-phase Pseudomonas aeruginosa. Proc Natl Acad Sci. 2018;115:9797–802.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    67.Page R, Peti W. Toxin-antitoxin systems in bacterial growth arrest and persistence. Nat Chem Biol. 2016;12:208–14.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    68.Liao X, Ma Y, Daliri EB-M, Koseki S, Wei S, Liu D, et al. Interplay of antibiotic resistance and food-associated stress tolerance in foodborne pathogens. Trends Food Sci Technol. 2020;95:97–106.CAS 
    Article 

    Google Scholar 
    69.Levin-Reisman I, Brauner A, Ronin I, Balaban NQ. Epistasis between antibiotic tolerance, persistence, and resistance mutations. Proc Natl Acad Sci. 2019;116:14734–9.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    70.Levin-Reisman I, Ronin I, Gefen O, Braniss I, Shoresh N, Balaban NQ. Antibiotic tolerance facilitates the evolution of resistance. Science. 2017;355:826–30.CAS 
    PubMed 
    Article 

    Google Scholar  More