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    Recent declines in salmon body size impact ecosystems and fisheries

    Age-length (AL) datasets
    Alaska Department of Fish & Game (ADF&G) monitors the number, body size, sex, and age of Alaska salmon harvested in a variety of fisheries and on their return breeding migration from the ocean to freshwater. Age and body length (AL) data have been collected on mature adults from commercial, subsistence, and sport harvests, escapement (spawning population) projects, and test fisheries since the early 1900’s. ADF&G data has historically been archived in regional offices; however, for this project we were able to compile all available data from across the state (Supplementary Figs. S7–S10) into a single dataset, representing over 14 million raw AL samples.
    The majority of Alaska salmon fisheries target mature adults during their breeding migration into freshwater. Data from commercial harvests represent the largest proportion (57%) of measurements and are generally collected from marine waters and near river mouths. Although many Alaska salmon fishing districts are designed to operate as terminal fisheries, targeting fish destined for their river of origin, even terminal fisheries can intercept salmon returning to other Alaskan populations, and many other districts are non-terminal. Because most commercial salmon fisheries in Alaska catch a combination of fish from the target stock and intercepted fish returning to other populations, commercial samples often include a mix of fish from different populations within a river drainage and outside the drainage (e.g., Southeast Alaska troll fishery may be >80% non-local fish at times). Commercial samples from some fisheries targeting wild salmon could include a relatively low but unknown proportion of hatchery-origin salmon, which could not be excluded from our analyses without individual-level information on origin (hatchery or wild). Samples from escapement enumeration projects (sampling projects that count the number of mature adults that ‘escape’ the fishery and return to freshwater) make up the next highest proportion of AL measurements (33%). Escapement projects collect AL data from fish sampled in the freshwater environment, close to or on the spawning grounds, generally at counting towers, weirs, or fences. A variety of other sampling project types (test fishing, subsistence catch, sport catch) make up the remaining portion of these data, with no single project type representing more than 5% of the samples. ADF&G recorded the name of the sampling project, generally as the name of a given river (e.g., Fish Creek) or district (e.g., Togiak District), which we refer to as sampling locations. To ensure as much as possible that methods of data collection were consistent across locations and species, we excluded data collected from projects other than commercial harvest and escapement monitoring from statistical analyses.
    Age and length (AL) measurements were collected by ADF&G personnel using standard methods56. Briefly, fish length is collected to the nearest millimeter using a measuring tape or a manual or electronic measuring board, depending on project and year. Fish age was most commonly estimated by ADF&G scientists reading growth annuli on scales57. For many AL measurements, specimen sex was also recorded, predominantly using external characteristics for sex determination. Sex determination with external characteristics in ocean-phase fish is frequently unreliable58. Because most of our data come from commercial harvests that occur in ocean-phase fish prior to the development of obvious external secondary sexual characteristics, we did not analyze the sexes separately. However, other studies examining length at age with reliable sex determination have shown similar trends in size and age for males and females33,59. As in Lewis et al.19, we assume our results reflect similar trends in male and female salmon.
    To ensure data were of high quality, a number of quality assurance checks were established, and data failing those checks were excluded from analysis. These checks include ensuring that ages and lengths were within reasonable bounds for each species, that sample dates were reasonable, that data were not duplicated, and that data were all of the same length measurement type (mid-eye to fork of tail). Because mid-eye to fork length was by far the most commonly used length measurement type (85% of samples) within the data, and the vast majority of sample protocols use mid-eye to fork measurements, we assumed that observations where no length measurement type was reported (0.08% of samples) were mid-eye to fork. No other unique length measurement type accounts for more than 2% of samples. We also excluded any samples that measured fewer than ten fish for a given year/location combination. After these extensive checks, we were left with measurements on over 12.5 million individual salmon.
    A wide variety of gear types were used to collect samples. The three most common gear types included gillnet, seine, and weir. Sampling methods within projects did not change systematically over time; however, for at least some projects, changes did occur, such as changes in gillnet mesh materials and sizes (for commercial harvest60) or sampling location within a watershed (for escapement projects). Some of these methodology changes are sporadically reflected in the data (e.g., mesh size), whereas others are not included and difficult to capture (e.g., weir location changes). Given the inconsistency in data and metadata associated with these fine-scale methodology changes, and the spatial and temporal scale of this dataset, changes in mesh size, gear type, or fine scale location changes (movement of a project within the same river system) were not included in our analyses.
    Consistency in salmon size declines
    To quantify the spatial and temporal extent of body size change, we estimated the average length of fish for each species in each sampling location and return year (the year when the fish was caught or sampled on its return migration to freshwater), which we interpret as putative biological populations (henceforth referred to as populations). For each population, we averaged these annual means to find the mean body length during a baseline period before 1990 and recent period after 2010. The pre-1990 period included all data collected before 1990, though relatively little data was available before 1980. Comparing data from two discrete time periods avoids potential edge effects that would be introduced in dividing a consecutive time series. Only populations for which we had data in both periods were included (100 sockeye, 34 Chinook, 32 chum, and 13 coho salmon populations). We established a criterion of at least 3 years of data for each population during each time period for inclusion in this analysis. Although somewhat arbitrary, we chose 1990 as the end of the early period to ensure a large number of populations had sufficient data to be included, while still being early enough to provide a meaningful baseline for comparison with current data. Because our goal was to investigate trends experienced by resource users in Alaska, we included data from some stocks that are known to capture salmon that originated from areas other than Alaska. For example, estimates for Chinook salmon from Southeast Alaska are likely influenced by the inclusion of troll-caught Chinook salmon, which are largely composed of salmon originating from British Columbia (B.C.) and the U.S. West Coast. For visualization, the results of this analysis were then scaled up to the level of the fisheries management areas established by ADF&G (Fig. 1).
    To quantify and visualize continuous changes in body size across time, we fit general additive models (GAMs) to annual mean population body length for each species. To avoid convergence problems due to small sample sizes, data collected before 1975 were excluded from this analysis. In contrast to previous studies that assumed monotonic linear changes in size18,19, year was included as a nonlinear smoothed term because preliminary analyses suggested that the rate of length change varied through time. We included data from all populations for which observations from five or more years were available (276 sockeye salmon populations, 202 Chinook salmon populations, 183 chum salmon populations, 142 coho salmon populations). We knew a priori that salmon populations differ in average body size, so to preserve original units (mm) while controlling for variation in absolute body length among populations, we included two fixed factors: population and region. We assigned regions based on terrestrial biomes and the drainage areas of major watershed (shown numbered on Fig. 1, colored by ADF&G management region). Repeating these GAMs on escapement data alone provided equivalent results (Supplementary Fig. S11), which confirms that our results are not due to an artifact of sampling procedures through time.
    To visualize changes in age structure and size-at-age, we fit very similar GAMs to age and length-at-age data. As above we included fixed effects for population and region, as well as a nonlinear year effect. Using the same dataset as the previously described GAMs, we used either mean freshwater age, mean saltwater age, or mean length-at-age as the response variable. For length-at-age, we separately fit GAMs for the four most common age classes in each species, except coho salmon, for which sufficient data was available for only three age classes.
    To determine the extent to which patterns of body size change are consistent across space within a species, we re-fit these GAMs by replacing the main year effect by either a region-by-year or population-by-year interaction and compared model fit using AIC. These nonlinear interactions allow regions or populations to differ in their patterns of length change through time. These models are more data intensive than the previous GAMs, so we included data from all populations for which our time series consisted of any 20 or more years of data (123 sockeye salmon populations, 37 Chinook salmon populations, 38 chum salmon populations, 14 coho salmon populations).
    Contributions of declining age versus growth
    To partition the contribution of changes in population age structure versus size-at-age to changes in mean population length, we used the chain rule61. We used the discrete time analog of the chain rule

    $${Delta}left( {xy} right) = y{Delta}x + x{Delta}y,$$
    (1)

    and assume that change in mean length is a function of changes in population age structure, p(a), and mean length-at-age, x(a). For each species and population, age structure in year t was calculated as the proportion of individuals in each age a. Mean length in year t is given by

    $$x_t = {Sigma}_ap_tleft( a right)x_tleft( a right),$$
    (2)

    and the year-to-year change in length is given by

    $${Delta}x_t = x_{left( {t + 1} right)} – x_t = {Sigma}_ap_tleft( a right)x_tleft( a right) + {Delta}p_tleft( a right)x_tleft( a right),$$
    (3)

    where

    $$p_t(a) = 1/2left[ {p_{t + 1}(a) + p_t(a)} right],$$
    (4)

    and

    $${Delta}p_t(a) = left[ {p_{t + 1}(a) – p_t(a)} right].$$
    (5)

    Solving these formulas year-to-year for each species in each population, we estimated the proportion of change in mean length due to changes in age structure and size-at-age. We included all populations for which we had five or more years of data (though change can only be estimated for consecutive years of data) and averaged the results across populations in each region.
    Causes of age and size changes
    To identify potential causes of change in salmon body size, we quantified associations with a variety of indices describing physical and biological conditions in Alaska’s freshwater and marine salmon habitats. Each candidate explanatory variable was selected based on existing biological hypotheses or inclusion in previous analyses of salmon size or population dynamics.
    We considered several ocean climate indicators as potential causes of change in salmon size over time. Pacific Ocean conditions are often quantified using large-scale climate indices such as the Pacific Decadal Oscillation (PDO), El Niño Southern Oscillation (ENSO), and NPGO. These large-scale indices of ocean conditions, as proxies for climate and marine environment, have been shown to affect the survival and productivity of Pacific salmon in the North Pacific Ocean62,63. PDO, NPGO64, and MEI65,66 indices were all accessed and downloaded online (PDO, http://research.jisao.washington.edu/pdo/; NPGO, http://www.o3d.org/npgo/npgo.php, accessed 2018-02-07; MEI, https://www.esrl.noaa.gov/psd/enso/mei/, accessed 2018-02-08; MEIw, https://www.beringclimate.noaa.gov/, accessed 2018-02-08). In this analysis, winter means of NPGO and MEI were used in addition to an annual mean of MEI. Two ice cover metrics were also used to capture ocean climate conditions. Bering Sea ice cover and retreat were downloaded from https://www.beringclimate.noaa.gov/, originally derived from the National Snow and Ice Data Center data. Bering Sea ice cover index represents the winter anomaly, relative to 1981–2000 mean. Bering Sea ice retreat is an index representing number of days with ice cover after March 15.
    Sea surface temperature (SST) was also explored as a potential cause of the changes in salmon size and age. SST has proven to be closely linked to salmon productivity. Mueter et al.67 found that regional-scale SST predicted survival rates better than large-scale climate indices such as the PDO. They concluded that survival rates were largely driven by environmental conditions at regional spatial scales. SST was extracted from the Extended Reconstructed Sea Surface Temperature (ERSST) version 468. To approximate SST values close to the river mouths which juvenile salmonids are most likely to experience after ocean entry, a double layer of the grid cells tracing the coastline of Alaska were extracted and the mean summer SST was calculated for each region.
    Because in situ fluvial temperature measurements are sparse, both spatially and temporally, compared to the coverage of the AL dataset, air temperature was used as a proxy for temperature during the freshwater life stages. Air temperature data were extracted and sorted from remote-sensed satellite observations into multi-monthly regional means by season69.
    Finally, we considered the potential for competition with other salmon to influence salmon size by including the abundances of several highly abundant salmon species as explanatory covariates. Using data compiled by Ruggerone and Irvine39, we evaluated the abundance of adult pink, chum, and sockeye salmon returning to Asia and North America as a proxy for the abundance of adult salmon of each species in the North Pacific. In addition, we also considered the more localized abundance of pink, chum, and sockeye salmon returning to Alaska, because salmon body size has been shown to vary with salmon abundance in the year of return migration in some species70 at finer spatial scales. The abundances of coho and Chinook salmon were not included, because they occur at much lower abundance than sockeye, chum, and pink salmon.
    We also explored marine mammal abundances as potential predictor variables, but found that the data available precluded rigorous statistical comparison with our time series of salmon size and age structure. For example, the only estimates of orca abundance available for our study area (that from Southeast Alaska and Prince William Sound) show steady, near monotonic increases through our study period71,72. Statistically, this leads to insufficient replication and high collinearity with year effects. Although caution is warranted in interpretations of any models for which the assumptions are so obviously violated, we note that preliminary analyses including marine mammal abundance were not dramatically superior in terms of variance explained or model fit. Because of these limitations, we determined that a reliable test of the effect of marine mammal predation was not possible for Alaska.
    Ultimately, we only selected covariates with an absolute correlation among covariate time series of less than 0.61. By establishing this threshold for absolute pairwise covariate correlation we sought to include only covariates for which separate associations with salmon size could be identified. The final set of covariates included in our analyses were: (1) ocean climate indicators (PDO, NPGO, MEI, winter MEI (MEIw), and Bering Sea ice cover index); (2) sea surface temperature (SST); (3) air temperature as proxy for freshwater temperature; and (4) ocean salmon abundance (abundance of Alaska sockeye, pink, and chum salmon, and North Pacific wide abundance of sockeye, pink, and chum salmon).
    To test hypothesized associations between temporal trends in the average body size (length) of salmon and environmental conditions, we fit a series of Bayesian hierarchical models to data describing size trends across sampling locations for each species. Because the chain rule analysis showed that changes in age structure explained greater interannual body size variation than did changes in size-at-age, we analyzed age-aggregated mean body length. Time series, starting in 1975, of annual mean length by species for each sampling location (l) and environmental covariates were mean-variance (Z) standardized prior to model fitting. Models of the form

    $$L_{i,t} = mathop {{Sigma}}limits_c ( {beta _{l,c} * X_{t – delta _{c,}c}} ) + sleft( t right) + varepsilon _{l,t},$$
    (6)

    were fit to each salmon species separately using Bayesian methods, where Ll,t is the standardized length at each location (l) in each return or observation year (t), βl,c are coefficients describing the effect of each covariate (c) on average length at each location, and (X_{t – delta _{c,}c}) is the standardized value of each covariate in each year. The reference year for each covariate is specified relative to the return year, or year in which salmon length compositions are observed (t), by a species and covariate-specific offset δc that associates covariate effects with the hypothesized period of interaction in each species’ life history (Supplementary Table S2). Location-specific covariate effects are structured hierarchically such that parameters describing the effect of each covariate on observed changes in average length were subject to a normally-distributed prior whose hyperparameters (group-level means and standard deviations for each covariate) were estimated directly from the data:

    $$beta _{l,c} sim {mathrm{Normal}}left( {mu _c,tau _c ^{2}} right),$$
    (7)

    This hierarchical structure permitted us to quantify both the average (group-level) association between length observations at each sampling location (l) and hypothesized covariates (i.e., the hyperparameter μc), and the level of among-location variation in these effects (i.e., (τ_c^{2})). Prior distributions for model parameters were generally uninformative, with the exception of the prior on the group-level mean covariate effects (μc) which included a mild penalty toward zero,

    $$mu _c sim {mathrm{Normal}}left( {0,1} right).$$
    (8)

    The prior distribution of the group-level (hyper) standard deviation of covariate effects was broad and truncated at zero,

    $$tau _c sim {mathrm{Normal}}left( {0,10} right)left[ {0,} right],$$
    (9)

    allowing the model to freely estimate the appropriate level of among-location variability in covariate effects.
    Observation error was assumed to be normally distributed εl,t ~ Normal(0, σε2), with a common observation error variance (σε2) estimated as a free parameter and subject to a broad prior distribution

    $$sigma _varepsilon sim {mathrm{Normal}}left( {0,10} right)left[ {0,} right].$$
    (10)

    Each species-specific model also included a smoothed nonlinear year effect s(t) describing residual trends in length across time that were shared among sampling (observation) locations but were not explained by the covariates. The degree of nonlinearity for the univariate smooth s(t) quantifying the common residual trend in length is controlled by the variance term (σs) for the coefficients forming the spline73, for which a broad zero-truncated prior distribution was defined:

    $$sigma _s sim {mathrm{Normal}}left( {0,10} right)left[ {0,} right].$$
    (11)

    Hierarchical Bayesian models describing the temporal trend in location-specific salmon length were fit using the brms package73,74 in R (R Core Team 2018), which generates posterior samples using the No U-Turn Sampler implemented in the Stan software platform75. Three independent chains were run for 20,000 iterations with a 50% burn-in and saving every tenth posterior sample, resulting in 3000 posterior samples. Convergence of all chains was diagnosed by ensuring potential scale reduction factors (R̂) for each parameter were More

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    Presence of toxin-antitoxin systems in picocyanobacteria and their ecological implications

    1.
    Flombaum P, Gallegos JL, Gordillo RA, Rincón J, Zabala LL, Jiao N, et al. Present and future global distributions of the marine Cyanobacteria Prochlrococcus and Synechococcus. Proc Natl Acad Sci. 2013;110:9824–9.
    CAS  PubMed  Google Scholar 
    2.
    Li WKW, Url S. Primary production of prochlorophytes, cyanobacteria, and eucaryotic ultraphytoplankton: Measurements from flow cytometric sorting. Limnol Oceanogr. 1994;39:169–75.
    CAS  Google Scholar 

    3.
    Dvořák P, Casamatta DA, Poulíčková A, Hašler P, Ondřej V, Sanges R. Synechococcus: 3 billion years of global dominance. Mol Ecol. 2014;23:5538–51.
    PubMed  Google Scholar 

    4.
    Morel A, Ahn YH, Partensky F, Vaulot D, Claustre H. Prochlorococcus and Synechococcus: a comparative study of their optical properties in relation to their size and pigmentation. J Mar Res. 1993;51:617–49.
    CAS  Google Scholar 

    5.
    Partensky F, Blanchot J, Vaulot D. Differential distribution and ecology of Prochlorococcus and Synechococcus in oceanic waters: a review. Bull l’Institut océanographique. 1999;19:457–75.
    Google Scholar 

    6.
    Biller SJ, Berube PM, Lindell D, Chisholm SW. Prochlorococcus: the structure and function of collective diversity. Nat Rev Microbiol. 2015;13:13–27.
    CAS  PubMed  Google Scholar 

    7.
    Sun Z, Blanchard JL. Strong Genome-Wide Selection Early in the Evolution of Prochlorococcus Resulted in a Reduced Genome through the Loss of a Large Number of Small Effect Genes. PLoS ONE. 2014;9:e88837.
    PubMed  PubMed Central  Google Scholar 

    8.
    Larsson J, Nylander JAA, Bergman B. Genome fluctuations in cyanobacteria reflect evolutionary, developmental and adaptive traits. BMC Evol Biol. 2011;11:187.
    PubMed  PubMed Central  Google Scholar 

    9.
    Dufresne A, Garczarek L, Partensky F. Accelerated evolution associated with genome reduction in a free-living prokaryote. Genome Biol. 2005;6:R14.1–R14.10.
    Google Scholar 

    10.
    Scanlan DJ, Ostrowski M, Mazard S, Dufresne A, Garczarek L, Hess WR, et al. Ecological genomics of marine picocyanobacteria. Microb Mol Biol Rev. 2009;73:249–99.
    CAS  Google Scholar 

    11.
    Scanlan DJ. Marine Picocyanobacteria. In: Whitton B. (eds). Ecology of Cyanobacteria II: Their Diversity in Space and Time. Springer Netherlands: Dordrecht, Netherlands 2012, pp. 503–33.

    12.
    Sánchez-Baracaldo P, Hayes PK, Blank CE. Morphological and habitat evolution in the Cyanobacteria using a compartmentalization approach. Geobiology. 2005;3:145–65.
    Google Scholar 

    13.
    Wang K, Wommack KE, Chen F. Abundance and distribution of Synechococcus spp. and cyanophages in the Chesapeake Bay. Appl Environ Microbiol. 2011;77:7459–68.
    CAS  PubMed  PubMed Central  Google Scholar 

    14.
    Callieri C. Picophytoplankton in freshwater ecosystems: the importance of small-sized phototrophs. Freshw Rev. 2008;1:1–28.
    Google Scholar 

    15.
    Stockner JG. Phototrophic picoplankton: an overview from marine and freshwater ecosystems. Limnol Oceanogr. 1988;33:765–75.
    CAS  Google Scholar 

    16.
    Callieri C, Stockner JG. Freshwater autotrophic picoplankton: a review. J Limnol. 2002;61:1–14.
    Google Scholar 

    17.
    Honda D, Yokota A, Sugiyama J. Detection of seven major evolutionary lineages in cyanobacteria based on the 16S rRNA gene sequence analysis with new sequences of five marine Synechococcus strains. J Mol Evol. 1999;48:723–39.
    CAS  PubMed  Google Scholar 

    18.
    Rippka R, Deruelles J, Waterbury JB. Generic assignments, strain histories and properties of pure cultures of cyanobacteria. J Gen Microbiol. 1979;111:1–61.
    Google Scholar 

    19.
    Wilmotte AMR, Stam WT. Genetic relationships among cyanobacterial strains originally designated as ‘Anacystis nidulans’ and some other Synechococcus strains. J Gen Microbiol. 1984;130:2737–40.
    Google Scholar 

    20.
    Coutinho F, Tschoeke DA, Thompson F. Comparative genomics of Synechococcus and proposal of the new genus Parasynechococcus. PeerJ. 2016;4:e1522 1–18.
    Google Scholar 

    21.
    Robertson BR, Tezuka N, Watanabe MM. Phylogenetic analyses of Synechococcus strains (cyanobacteria) using sequences of 16S rDNA and part of the phycocyanin operon reveal multiple evolutionary lines and reflect phycobilin content. Int J Syst Evol Microbiol. 2001;51:861–71.
    CAS  PubMed  Google Scholar 

    22.
    Zwirglmaier K, Jardillier L, Ostrowski M, Mazard S, Garczarek L, Vaulot D, et al. Global phylogeography of marine Synechococcus and Prochlorococcus reveals a distinct partitioning of lineages among oceanic biomes. Environ Microbiol. 2008;10:147–61.
    PubMed  Google Scholar 

    23.
    Rocap G, Distel DL, Waterbury JB, Chisholm SW. Resolution of Prochlorococcus and Synechococcus ecotypes by using 16S-23S ribosomal DNA internal transcribed spacer sequences. Appl Environ Microbiol. 2002;68:1180–91.
    CAS  PubMed  PubMed Central  Google Scholar 

    24.
    Toledo G, Palenik B. Synechococcus diversity in the California Current as seen by RNA polymerase (rpoC1) gene sequences of isolated strains. Appl Environ Microbiol. 1997;63:4298–303.
    CAS  PubMed  PubMed Central  Google Scholar 

    25.
    Fuller NJ, Marie D, Partensky F, Vaulot D, Post AF, Scanlan DJ. Clade-specific 16S ribosomal DNA oligonucleotides reveal the predominance of a single marine Synechococcus clade throughout a stratified water column in the Red Sea. Appl Environ Microbiol. 2003;69:2430–43.
    CAS  PubMed  PubMed Central  Google Scholar 

    26.
    Dufresne A, Ostrowski M, Scanlan DJ, Garczarek L, Mazard S, Palenik BP, et al. Unraveling the genomic mosaic of a ubiquitous genus of marine cyanobacteria. Genome Biol. 2008;9:R90.
    PubMed  PubMed Central  Google Scholar 

    27.
    Huang S, Wilhelm SW, Harvey HR, Taylor K, Jiao N, Chen F. Novel lineages of Prochlorococcus and Synechococcus in the global oceans. ISME J. 2012;6:285–97.
    CAS  PubMed  Google Scholar 

    28.
    Callieri C, Coci M, Corno G, Macek M, Modenutti B, Balseiro E, et al. Phylogenetic diversity of nonmarine picocyanobacteria. FEMS Microbiol Ecol. 2013;85:293–301.
    CAS  PubMed  Google Scholar 

    29.
    Crosbie ND, Pockl M, Weisse T. Dispersal and phylogenetic diversity of nonmarine picocyanobacteria inferred from 16S rRNA gene and cpcBA-intergenic spacer sequence analyses. Appl Environ Microbiol. 2003;69:5716–21.
    CAS  PubMed  PubMed Central  Google Scholar 

    30.
    Jasser I, Królicka A, Karnkowska-Ishikawa A. A novel phylogenetic clade of picocyanobacteria from the Mazurian lakes (Poland) reflects the early ontogeny of glacial lakes. FEMS Microbiol Ecol. 2011;75:89–98.
    CAS  PubMed  Google Scholar 

    31.
    Ernst A, Becker S, Wollenzien UIA, Postius C. Ecosystem-dependent adaptive radiations of picocyanobacteria inferred from 16S rRNA and ITS-1 sequence analysis. Microbiology. 2003;149:217–28.
    CAS  PubMed  Google Scholar 

    32.
    Palenik B, Brahamsha B, Larimer FW, Land M, Hauser L, Chain P, et al. The genome of a motile marine Synechococcus. Nature. 2003;424:1037–42.
    CAS  PubMed  Google Scholar 

    33.
    Palenik B, Ren Q, Dupont CL, Myers GS, Heidelberg JF, Badger JH, et al. Genome sequence of Synechococcus CC9311: Insights into adaptation to a coastal environment. Proc Natl Acad Sci. 2006;103:13555–9.
    CAS  PubMed  Google Scholar 

    34.
    Stuart RK, Dupont CL, Johnson DA, Paulsen IT, Palenik B. Coastal strains of marine Synechococcus species exhibit increased tolerance to copper shock and a distinctive transcriptional response relative to those of open-ocean strains. Appl Environ Microbiol. 2009;75:5047–57.
    CAS  PubMed  PubMed Central  Google Scholar 

    35.
    Stuart RK, Brahamsha B, Busby K, Palenik B. Genomic island genes in a coastal marine Synechococcus strain confer enhanced tolerance to copper and oxidative stress. ISME J. 2013;7:1139–49.
    CAS  PubMed  PubMed Central  Google Scholar 

    36.
    Fucich D, Marsan D, Sosa A, Chen F. Complete genome sequence of Subcluster 5.2 Synechococcus sp. strain CB0101, isolated from the Chesapeake Bay. Microbiol Resour Announc. 2019;8:6–8.
    Google Scholar 

    37.
    Marsan D, Place A, Fucich D, Chen F. Toxin-antitoxin systems in estuarine Synechococcus strain CB0101 and their transcriptomic responses to environmental stressors. Front Microbiol. 2017;8:1–11.
    Google Scholar 

    38.
    Page R, Peti W. Toxin-antitoxin systems in bacterial growth arrest and persistence. Nat Chem Biol. 2016;12:208–14.
    CAS  PubMed  Google Scholar 

    39.
    Unterholzner SJ, Poppenberger B, Rozhon W. Toxin-antitoxin systems. Mob Genet Elem. 2013;3:e26219 1–13.
    Google Scholar 

    40.
    Makarova KS, Wolf YI, Koonin EV. Comprehensive comparative-genomic analysis of type 2 toxin-antitoxin systems and related mobile stress response systems in prokaryotes. Biol Direct. 2009;4:19.
    PubMed  PubMed Central  Google Scholar 

    41.
    Kaneko T, Nakamura Y, Sasamoto S, Watanabe A, Kohara M, Matsumoto M, et al. Structural analysis of four large plasmids harboring in a unicellular cyanobacterium, Synechocystis sp. PCC 6803. DNA Res. 2003;10:221–8.
    CAS  PubMed  Google Scholar 

    42.
    Chen Y, Holtman CK, Magnuseon RD, Youderian PA, Golden SS. The complete sequence and functional analysis of pANL, the large plasmid of the unicellular freshwater cyanobacterium Synechococcus elongatus PCC 7942. Plasmid. 2011;23:1–7.
    Google Scholar 

    43.
    Chen F, Wang K, Kan J, Suzuki MT, Wommack KE. Diverse and unique picocyanobacteria in Chesapeake Bay, revealed by 16S-23S rRNA internal transcribed spacer sequences. Appl Environ Microbiol. 2006;72:2239–43.
    CAS  PubMed  PubMed Central  Google Scholar 

    44.
    Xie Y, Wei Y, Shen Y, Li X, Zhou H, Tai C, et al. TADB 2.0: An updated database of bacterial type II toxin-antitoxin loci. Nucleic Acids Res. 2018;46:D749–D753.
    CAS  PubMed  Google Scholar 

    45.
    O’Leary NA, Wright MW, Brister JR, Ciufo S, Haddad D, McVeigh R, et al. Reference sequence (RefSeq) database at NCBI: Current status, taxonomic expansion, and functional annotation. Nucleic Acids Res. 2016;44:D733–D745.
    PubMed  Google Scholar 

    46.
    Agarwala R, Barrett T, Beck J, Benson DA, Bollin C, Bolton E, et al. Database resources of the national center for biotechnology information. Nucleic Acids Res. 2017;45:D12–D17.
    CAS  Google Scholar 

    47.
    Nordberg H, Cantor M, Dusheyko S, Hua S, Poliakov A, Shabalov I, et al. The genome portal of the Department of Energy Joint Genome Institute: 2014 updates. Nucleic Acids Res. 2014;42:26–31.
    Google Scholar 

    48.
    Shao Y, Harrison EM, Bi D, Tai C, He X, Ou HY, et al. TADB: a web-based resource for Type 2 toxin-antitoxin loci in bacteria and archaea. Nucleic Acids Res. 2011;39:606–11.
    Google Scholar 

    49.
    Marchler-Bauer A, Bo Y, Han L, He J, Lanczycki CJ, Lu S, et al. CDD/SPARCLE: Functional classification of proteins via subfamily domain architectures. Nucleic Acids Res. 2017;45:D200–D203.
    CAS  PubMed  Google Scholar 

    50.
    R Core Team. R: A Language and Environment for Statistical Computing. 2018. Vienna, Austria.

    51.
    Wickham H. ggplot2: elegant graphics for data analysis. New York: Springer-Verlag; 2009.
    Google Scholar 

    52.
    Bertelli C, Laird MR, Williams KP, Lau BY, Hoad G, Winsor GL, et al. IslandViewer 4: Expanded prediction of genomic islands for larger-scale datasets. Nucleic Acids Res. 2017;45:W30–W35.
    CAS  PubMed  PubMed Central  Google Scholar 

    53.
    Winther KS, Gerdes K. Enteric virulence associated protein VapC inhibits translation by cleavage of initiator tRNA. Prok Natl Acad Sci. 2011;108:7403–7.
    CAS  Google Scholar 

    54.
    Harms A, Brodersen DE, Matarai N, Gerdes K. Toxins,targets,and triggers: an overview of toxin-antitoxin biology. Mol Cell. 2018;70:768–84.
    CAS  PubMed  Google Scholar 

    55.
    Kopfmann S, Roesch S, Hess W. Type II toxin–antitoxin systems in the unicellular cyanobacterium Synechocystis sp. PCC 6803. Toxins. 2016;8:228.
    PubMed Central  Google Scholar 

    56.
    Leplae R, Geeraerts D, Hallez R, Guglielmini J, Drze P, Van Melderen L. Diversity of bacterial type II toxin-antitoxin systems: a comprehensive search and functional analysis of novel families. Nucleic Acids Res. 2011;39:5513–25.
    CAS  PubMed  PubMed Central  Google Scholar 

    57.
    Xia K, Bao H, Zhang F, Linhardt RJ, Liang X. Characterization and comparative analysis of toxin – antitoxin systems in Acetobacter pasteurianus. J Ind Microbiol Biotechnol. 2019;46:869–82.
    CAS  PubMed  Google Scholar 

    58.
    Biller SJ, Berube PM, Berta-Thompson JW, Kelly L, Roggensack SE, Awad L, et al. Genomes of diverse isolates of the marine cyanobacterium Prochlorococcus. Sci Data. 2014;1:1–11.
    Google Scholar 

    59.
    Palenik B, Barahamsha B, Larimer FW, Land M, Hauser L, Chain P, et al. The genome of a motile marine Synechococcus. Nature. 2003;424:1037–42.
    CAS  PubMed  Google Scholar 

    60.
    Scanlan DJ, West NJ. Molecular ecology of the marine cyanobacterial genera Prochlorococcus and Synechococcus. FEMS Microbiol Ecol. 2002;40:1–12.
    CAS  PubMed  Google Scholar 

    61.
    McWilliam H, Li W, Uludag M, Squizzato S, Park YM, Buso N, et al. Analysis tool web services from the EMBL-EBI. Nucleic Acids Res. 2013;41:W597–W600.
    PubMed  PubMed Central  Google Scholar 

    62.
    Sevin EW, Barloy-Hubler F. RASTA-bacteria: a web-based tool for identifying toxin-antitoxin loci in prokaryotes. Genome Biol. 2007;8:R155.1–R155.14.
    Google Scholar 

    63.
    Robson J, McKenzie JL, Cursons R, Cook GM, Arcus VL. The vapBC operon from mycobacterium smegmatis Is an autoregulated toxin-antitoxin module that controls growth via inhibition of translation. J Mol Biol. 2009;390:353–67.
    CAS  PubMed  Google Scholar  More

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    Mechanisms protect airborne green microalgae during long distance dispersal

    Isolation and identification of four airborne microalgal strains belonging to three genera from Dongsha Island in the South China Sea
    Four microalgal strains belonging to three genera were successfully isolated in the collection trip in December 2013 and all the four could not grow in 2f seawater medium10 (in both 33 and 22‰ salt levels), indicating they were freshwater species. As shown in Fig. 1, based on the morphology revealed by Scanning Electron Microscopy and the phylogenetic analysis, the four strains were named Scenedesmus sp. DSA1 (stands for Dongsha airborne #1), Coelastrella sp. DSA2, Coelastrella sp. DSA3, and Desmodesmus sp. DSA6 (hereafter referred to as DSA1, DSA2, DSA3 and DSA6, respectively). All four strains are members of the family Scenedesmaceae in Chlorophyta. Sequences of the 18S rDNA and ITS1-5.8S-ITS2 fragments of the four airborne microalgal strains were deposited into GenBank under the accession numbers KX818834–KX818841.
    Figure 1

    A Scanning electron microscopy (SEM) micrographs of the four airborne microalgae in the early stationary phase. Upper panel, left and right: Scenedesmus sp. DSA1 and Coelastrella sp. DSA2. Lower panel, left and right: Coelastrella sp. DSA3 and Desmodesmus sp. DSA6. B Phylogenetic analysis of the four airborne microalgal strains (underlined) and their related species based on the fusion sequences of ITS1 and ITS2 of each species. The numbers in the parentheses are accession numbers of each sequence in GenBank. The numbers at the nodes indicate bootstrap values (expressed as percentage) with 500 replicates.

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    The airborne green microalgae had better UV tolerance than the waterborne green microalgae
    When traveling in the air, airborne microalgae are inevitably exposed to much higher levels of UV radiation compared to microalgae in the aquatic environments. In order to survive aerial travel, these cells must have a mechanism(s) to protect themselves against the damaging effects of the radiation, a mechanism that is of less concern to waterborne microalgal populations. To verify whether the four airborne microalgae resisted UV radiation better than waterborne microalgae, these microorganisms were spread onto agar plates using the top agar method11, and exposed to UV-B radiation to determine their survival rates. The survival rates were compared to those of the two waterborne microalgae, Desmodesmus sp. F512 and Neodesmus sp. UTEX 2219-413 (hereafter referred to as F5 and 2219-4, respectively), both are members of Scenedesmaceae as well. As shown in Fig. 2A, the survival rates of the two waterborne microalgae were about 68 and 46%, respectively, after 1 min of the UV-B exposure, compared to more than 90% for all four airborne microalgae. After 3 min of the exposure, however, all of the waterborne cells died but the survival rates of the airborne cells were about 98, 73, 26, and 11% for DSA3, DSA2, DSA1 and DSA6, respectively. Significant differences could still be observed among the survival rates of DSA3, DSA2, and the other four species after 5 min of the UV-B exposure.
    Figure 2

    A UV-B stress tolerance of the four airborne microalgae and the two waterborne microalgae used as controls. About 500 cells in the early stationary phase were spread onto each agar plate and irradiated with 302 nm UV for the specified durations. The survival rate of each strain was defined as the colony numbers on the UV-treated plates compared to those on the non-treated plates (n = 6, mean ± SE). F5, Desmodesmus sp. F5; 2219-4, Neodesmus sp. UTEX 2219-4. B Different autofluorescence intensities from the cell wall of the six microalgal strains.

    Full size image

    For algal cells, cell wall is the first barrier to defend the cells against UV attack. It is well known that the cell walls of terrestrial plants are autofluorescent when excited by UV14. In this context, the four airborne microalgal strains would have better UV tolerance if their cell walls could absorb UV, and therefore are autofluorescent as well. To examine this possibility, the pigment-free cells were examined under an epifluorescence microscope. Indeed, as shown in Fig. 2B, the cell walls of the four airborne and the two waterborne green microalgae were able to emit autofluorescence when excited by UV under the microscope. Furthermore, the autofluorescence intensities emitted from the six microalgal strains varied, with DSA3 and DSA2 being the strongest, DSA1 and DSA6 in the middle, and F5 and 2219-4 the weakest as detected using the same parameters. The varied intensities suggested that these microalgae had different levels of UV tolerance because their cell walls absorbed different amounts of UV energy.
    The UV tolerance of the microalgae was positively correlated with their cell wall thickness
    There appeared to be a correlation between the autofluorescence intensities emitted by the cell walls and the survival rates of the six microalgal strains exposed to the UV-B radiation for 3 min. It was intriguing to investigate whether the cell wall thickness of these microalgae played a role in the survival rates. To measure the cell wall thickness of the six microalgal strains, these cells were fixed and sectioned, and then observed using Transmission Electron Microscopy (TEM). As shown in Fig. 3A, the cell wall thickness of the six strains varied, with DSA3 and DSA2 being the thickest, DSA1 and DSA6 intermediate, and F5 and 2219-4 the thinnest. This relation well-correlated with that of the autofluorescence intensities from their cell walls. When the survival rates under 3 min of the UV-B exposure were plotted against the cell wall thickness of the six strains, a good correlation (r = 0.99, p  More

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    A cosmopolitan fungal pathogen of dicots adopts an endophytic lifestyle on cereal crops and protects them from major fungal diseases

    Plant and fungal materials, maintenance, and preparation
    The winter wheat cultivar Zheng 9023 and the spring wheat cultivar Yongliang 4 were purchased from the commercial seed market in Wuhan City and Minqin County in China, respectively. The barley cultivar Huadamai 14 was provided from Prof. Dongfa Sun in Huazhong Agricultural University, Wuhan, China. The oat cultivar Mengmai 2 was donated by Prof. Jun Zhao in Neimeng Agricultural University. The maize cultivar Zhengdan 958 was purchased from the commercial seed market in Wuhan City. The rice cultivar LTH was donated by Prof. Youliang Peng of China Agricultural University. All seeds were surface-sterilized with a 0.5% sodium hypochlorite solution (NaClO) before sowing or S. sclerotiorum treatment.
    S. sclerotiorum strain DT-8, which was originally isolated from a sclerotium collected from a diseased rapeseed, is a hypovirulent strain infected by a DNA mycovirus. Strain DT-8VF, a virulent strain, is a virus-free derivative of DT-8 [21]. Strain DT-8VFRFP is a derivative of DT-8VF labeled with the mCherry fluorophore by DNA transformation; it shows normal virulence. The wheat Fusarium head blight (FHB) pathogen, F. graminearum strain PH-1, was used to inoculate wheat spikes. An S. sclerotiorum virulent strain Ep-1PNA367 and three hypovirulent strains, AH98, SCH941, and T1-1-20, were also used to investigate their potential endophytic growth on wheat. AH98 is infected by a negative-stranded RNA virus [22], while SCH941 and T1-1-20 are infected by various other mycoviruses. All S. sclerotiorum strains and F. graminearum strain PH-1 were grown on potato dextrose agar (PDA) or potato dextrose broth (PDB) at 20 °C; Magnaporthe oryzae strain 131 was grown on PDA at 28 °C, or grown on tomato–oat medium to produce conidia, and was stored on PDA slants at 4 °C.
    Microscopic observation
    To observe the growth of S. sclerotiorum in wheat by confocal microscopy, seeds were surface-sterilized with NaClO and sown on half-strength Murashige and Skoog (MS) agarose medium amended with 25 mM sucrose for 8 days, then root crowns were inoculated with mycelia of the strain DT-8VFRFP. Wheat seedlings were maintained at 20 °C for 4 days under a 12-h photoperiod. Seedlings were subsequently washed three times with PBS for 10 min each, and then root sections were separated and incubated in a 1:100 dilution of wheat germ agglutinin conjugated to FITC (Sigma) for 1 h at room temperature according to the manufacturer’s instructions. The roots were visualized with a LEICA confocal microscope (LEICA SP8) using the 488-nm line of a 25-mW Argon ion laser for FITC and the 561-nm line of a 20-mW solid-state laser for mCherry.
    For further observation with a transmission electron microscopy (TEM), 5-mm root segments from wheat seedlings grown on MS medium for 15 days after inoculation with strain DT-8VFRFP were fixed in 0.4% (v/v) glutaraldehyde solution overnight at 4 °C. After washing in PBS buffer, roots were dehydrated with a graded ethanol series. Samples were then embedded in Epon-821 and polymerized at 60 °C. Thin sections (50 nm) were cut using a Leica ULTRACUT UCT ultramicrotome with a diamond knife.
    For TEM immunodetection, wheat seedling roots were fixed in 4% (w/v) freshly depolymerized paraformaldehyde and 0.4% (v/v) glutaraldehyde in 1× PBS, pH 7.4, for 1 h at 4 °C. The samples were then embedded using an LR white embedding kit (Fluka) and polymerized at 50 °C for 24 h. Immunogold labeling specificity was detected by displacing the anti-mCherry antibodies with rabbit preimmune serum. The method for TEM immunodetection was performed as previously described [23].
    For scanning electron microscopy (SEM) observation, 2-mm root segments from the wheat, barley, oat, maize, and rice seedlings treated with different strains of S. sclerotiorum grown on MS medium for 15 days were used. All segments were fixed in a 0.4% (v/v) glutaraldehyde solution overnight at 4 °C. For SEM analysis, the sections were allowed to air-dry overnight in a desiccator at room temperature, sputter-coated with gold, and prepared for SEM analysis (EVO MA 10 Carl Zeiss SMT AG, Germany). Root segments from nontreated wheat were sampled and observed as a control comparison.
    For confocal microscopy observations, 45 days after the root crown inoculation with mycelia of the DT-8VFRFP strain, stems from wheat plants growing in soil in the greenhouse were carefully washed with distilled water and embedded in Tissue-Tek O.C.T. compound medium (Sakura Finetek USA, Inc., Torrance, CA) at −23 °C overnight. Microtome sections (25 µm thick) were sliced using a freezing microtome (LEICA SP8, Germany). The stem microtome sections were then visualized with a LEICA confocal microscope using the 561-nm excitation wavelength for mCherry.
    Re-isolation of S. sclerotiorum
    To further probe whether S. sclerotiorum can go to aerial parts of wheat when inoculated on root of wheat seedling, plants were grown in sterile nutrient soil for 45 days. The samples for re-isolation were taken from the second segment near the base of wheat stem of eleven individual DT-8VFRFP-treated wheat plants, and were cut into 5 mm long segments, then surface-sterilized by dipping them in 70% EtOH for 2 min and then in 0.5% NaClO for 2 min, followed by rinsing three times with sterile distilled water. The sterilized stem segments were placed on hygromycin-amended PDA medium plates (50 µg/mL) and incubated at 20 °C for 8 days. Then, the emerging colonies were identified as S. sclerotiorum based on colony morphology, and PCR amplification [24].
    PCR determination of S. sclerotiorum and mycoviruses
    DNA samples of either wheat or fungi were extracted using a cetyltrimethylammonium bromide method. A primer pair XJJ21/XJJ222 (GTTGCTTTGGCGTGCTGCTC/CTGACATGGACTCAATACCAATCTG) was used for detection of S. sclerotiorum [24], and a primer pair pRep-F/pRep-R (GTCACCACCCAAACATTACAAAGAGCGTATTCC/ACGTCA GGTGC) was used for detection of viral DNA of SsHADV-1. A procedure described by Yu et al. [21] was used for PCR amplification.
    Seed treatment of wheat with S. sclerotiorum and sowing
    To determine whether S. sclerotiorum could promote growth and disease resistance of wheat in the greenhouse, wheat seeds were washed with tap water and surface-sterilized with 0.5% NaClO for 10 min, then washed three times with sterile water. Surface-sterilized seeds were soaked in sterile water for 4 h, and then collected and blotted dry. Meanwhile, fresh mycelium of strains DT-8 and DT-8VF were collected from PDA medium and then cultured in PDB medium in 250-mL flasks in a shaker at 20 °C for 4 days to obtain hyphal fragment suspensions; the number of viable fungal fragments was adjusted to 1.4 × 105 cfu/mL before inoculation of wheat seeds. The hyphal fragment suspensions were then used to inoculate the prepared seeds (100 mL of hyphal fragment suspension/kg wheat seeds) by thoroughly mixing the hyphal fragment suspension and wheat seeds for 6 h. The inoculated seeds were further dried using an electric fan for 12 h at room temperature. Wheat seeds soaked only in sterile water for 6 h and then dried with the same method were used as a control.
    To test whether the treated seeds were successfully colonized by S. sclerotiorum, S. sclerotiorum-treated seeds were surface-sterilized with 0.5% NaClO for 10 min, rinsed three times with sterile distilled water, and then cut in half and placed on PDA plates and incubated at 20 °C for 7 days. All emerging colonies from S. sclerotiorum-treated seeds were confirmed as S. sclerotiorum based on colony morphology and PCR amplification. In addition, a small number of S. sclerotiorum-treated seeds were randomly picked and sown into soil taken from the field and grown in the greenhouse. The seedlings were then tested after 21 days for the presence of S. sclerotiorum by PCR amplification; all seedlings tested positive. Hence, the seed treatment was confirmed as an efficient method for inoculation of wheat seeds with S. sclerotiorum. Consequently, we used this method to treat wheat seeds for the rest of the study, and kept treated seeds under dry conditions at room temperature for up to 7 days before sowing.
    Seeds treated either with strains DT-8VF or DT-8 were sown in pots in a greenhouse. In a laboratory test, treated seeds were allowed to germinate in a Petri dish on a layer of wet filter paper. Germinating seeds were then sown either into soil that was taken from the field or sterile nutrient soil. Twelve plants were grown in each pot. Wheat seedlings were maintained under greenhouse conditions at 20 °C. Strain DT-8-treated seeds were also tested in the field. Those seeds were sown as rows in the wheat field in exactly the same way as farmers normally do in each sowing season. Field management was conducted as per normal farmer practice, except that no fungicide was applied.
    In order to investigate whether S. sclerotiorum colonization can spread to aerial parts of wheat plants originating from DT-8-treated seeds, ten plants from each plot were randomly sampled from the field. A total of 30 wheat plants from the strain DT-8-treated group and 20 plants from the nontreated group were sampled. First, the roots were rinsed thoroughly with tap water. Then the roots, flag leaves, and spikes were given three brief rinses in distilled water. Each wheat plant was given a number, from 1 to 30 for DT-8-inoculated plants, and from 1 to 20 for nontreated plants. DNA samples were extracted individually from the root, flag leaf, and spike of each plant and used to determine the presence of S. sclerotiorum and mycovirus.
    Evaluating the growth of treated wheat in the greenhouse and field
    To evaluate the growth of S. sclerotiorum-treated wheat in greenhouse experiments, plant height was measured at the seedling and anthesis stages, while flag leaf and spike lengths were evaluated only at the anthesis stage. There were 60 plants in each group treated with strain DT-8 and strain DT-8VF and 60 plants in the nontreated group. Determination of 1000-grain weight was repeated, four in each group. Measurement data for each group were calculated for statistical analysis.
    To investigate whether other strains could promote wheat growth, strains Ep-1PNA367, AH98, SCH941, and T1-1-20 were used instead of strains DT-8 and DT-8VF. Wheat seeds were treated and then sown in a sterile mixture of vermiculite and perlite at the ratio of 3:1 in pots and placed in greenhouse, with ~50 seedlings in each pot. Seedling shoot fresh weight was measured at 25 days after planting. There were 30 plants in each treatment and control, and the average weight of ten seedlings was calculated.
    For field tests, plant height and the length, width, and thickness of flag leaves at the early flowering stage were measured in the field at EZhou. Forty plants from each plot were randomly measured from a total of 120 plants in the DT-8-treated group and from 120 plants in the nontreated group. Measurement data for each group were calculated for statistical analysis. All the results were confirmed with independent lines and over two planting seasons.
    Field experiments and wheat yield tests
    To examine whether S. sclerotiorum treatment could enhance wheat yield under natural field conditions, DT-8-treated seeds were sown in a wheat field located at EZhou in late October of 2016 and harvested in mid-May of 2017. This experiment was repeated at EZhou, Jingmen City, and Xiangyang City in late October of 2017 and 2018 and harvested in mid-May of 2018 and 2019. Furthermore, seeds of the spring wheat cultivar Yongliang 4 were also treated with strain DT-8 and sown in mid-April of 2017 at Minqin and Tianzhu Counties in Gansu province and harvested in late July of 2017. All wheat was managed as per normal farmer practice, except that no fungicide was applied. The treatments with or without strain DT-8 were replicated four times at Tianzhu County and Jingmen City in 2017 and five times at other places and the wheat yield from 5 m2 was measured in each plot and used for statistical analysis.
    Analysis of chlorophyll content and photosynthetic rate in flag leaves
    To determine chlorophyll content, leaf tissues were harvested using a circular punch that yields 0.5-cm diameter leaf discs. There were four flag leaf replicates for each treatment. Chlorophyll was extracted from wheat flag leaves obtained from the field at EZhou using 95% (v/v) ethanol (analytically pure, Sinopharm Chemical Reagent Co., Ltd) and the extracted chlorophyll concentration was determined using a spectrophotometer (UV2102, Unico, Shanghai, China) [25].
    For the photosynthetic rate, flag leaf samples were obtained from the field at EZhou. Each treatment had three flag leaf replicates. Photosynthetic rate determination was performed as previously described [26].
    Assay of plant hormones
    Five frozen flag leaf and spike replicates from each treatment (~100 mg for each flag leaf and spike sample) were ground to a fine powder in liquid nitrogen using a mortar and pestle. Each sample was weighed into a 1.5-mL tube, mixed with 750 μL of cold extraction buffer (methanol: water: acetic acid, 80:19:1, v/v/v) supplemented with internal standards, 10 ng of 2H6ABA, 10 ng of DHJA, and 5 μg of NAA, vigorously shaken on a shaking bed for 16 h at 4 °C in the dark, and then centrifuged at 12,000 rpm for 15 min at 4 °C. Supernatant was carefully transferred to a new 1.5-mL tube and pellets remixed with 400 μL of extraction buffer, shaken for 4 h at 4 °C, and centrifuged. The two supernatants were combined and filtered using a syringe-facilitated 13-mm diameter nylon filter with a pore size of 0.22 μm (Nylon 66; Jinteng Experiment Equipment Co., Ltd, Tianjing, China). The filtrate was dried by evaporation under nitrogen gas flow for ~5 h at room temperature and then dissolved in 200 μL of methanol. Aliquots of dissolved samples were further diluted 40 times using methanol for jasmonic acid (JA), abscisic acid (ABA), and indole-3-acetic acid (IAA) quantification. Liquid chromatography was carried out using an ultrafast liquid chromatography with an autosampler (Shimadzu Corporation, Kyoto, Japan). The method used for hormone determination was as previously described [27].
    Inoculation of F. graminearum and rating of disease
    Infection assays on flowering wheat spikes were performed as previously described [28]. At the early flowering stage, a conidial suspension of F. graminearum strain PH-1 was collected from 5-day-old cultures growing in carboxymethylcellulose medium, then filtered through three layers of lens-wiping paper and then mixed with 0.01% (v/v) Tween 20. Ten microliters of 1 × 105 conidia mL−1 conidial suspension was inoculated individually onto the fourth spikelet from the bottom using a micropipette. The inoculated wheat spikes were maintained at a relative humidity of 95% for 72 h. Symptomatic spikes were examined and images captured after 14 days.
    For the greenhouse test, 15 spikes from each treatment were inoculated and the spikelet infection rate of each spike was calculated; then, the average spikelet infection rate for each treatment was calculated for statistical analysis. For the field test, ten spikes from each plot were inoculated from a total of 30 inoculated spikes in the strain DT-8-treated plots and 30 spikes in the nontreated plots. The spikelet infection rate for each plot was calculated and the average spikelet infection rate for strain DT-8 treatment and nontreated control were then calculated for statistical analysis. The field test was conducted twice, once in 2017 and repeated in 2018.
    FHB survey in a natural, noninoculated field
    To investigate natural FHB infection in strain DT-8-treated wheat, an FHB field survey was conducted in experimental fields located at EZhou City, Jingmen City, and Xiangyang City in 2018. The field survey protocol described by the National Agricultural Technology Extension Service Center of China was adopted for the FHB survey with minor modifications. A total of 500 spikes were sampled randomly in each plot; in total, 1500 spikes were collected from DT-8-treated plots, with the same number of spikes being collected from nontreated plots to calculate disease incidence (spikelet infection rate) and severity (disease index). The number of infected and noninfected spikelets on each spike was counted and the average spikelet infection rate for each plot was calculated and used for statistical analysis. To calculate the disease index, the infected spikes were divided into five grades, namely: grade 0, no spikelet was infected; grade 1, the spike was infected, but less than 25% of spikelets were infected; grade 2, more than 25%, but less than 50%, of the spikelets were infected; grade 3, more than 50%, but less than 75%, of the spikelets were infected; and grade 4, more than 75% of the spikelets were infected. Finally, the disease severity for each plot was calculated using a formula for disease index, DI = ∑(nX/4 N) × 100, where “X” is the scale value of each spike, “n” is the number of spikes in the category, and “N” is the total number of spikes assessed for each plot. The disease index for each group was used for statistical analysis.
    Inoculation of M. oryzae on barley and rice
    To probe if S. sclerotiorum could enhance resistance against the rice blast fungus (M. oryzae) in barley and rice, an inoculation test was carried out according to a method described by Kong et al. [29]. Conidia of M. oryzae were collected with sterile water from 4-day-old cultures growing on tomato–oat medium and then filtered through three layers of lens-wiping paper. Infection assays were performed in whole plant leaves by spray inoculation using an airbrush nebulizer compressor. Strain DT-8-treated seedlings, which were grown in a sterile mixture of vermiculite and perlite at the ratio of 3:1 in pots and placed in greenhouse for 9 days for barley at 20 °C and 20 days for rice at 28 °C, were sprayed with a conidial suspension [105 conidia/mL mixed with 0.02% (v/v) Tween 20], using 4 mL suspension for each pot. The plants were further incubated at 28 °C, 80% relative humidity, under a 16 h light/8 h darkness photoperiod. Then, we assessed the presence of S. sclerotiorum in aerial parts of DT-8-treated barley and rice by PCR amplification in greenhouse plants; 92% of samples were confirmed as positive for S. sclerotiorum. For barley, lesions of leaves with the same leaf age (bottom leaves) were examined and typical infected leaves were photographed with a digital camera at 5 days post inoculation (dpi); for rice, the leaves were examined and photographed at 7 dpi.
    Detection of toxins (DON)
    To assay point-inoculated spikelets from strain DT-8-treated and nontreated plants, each sample was placed in a 50-mL centrifuge tube and mixed with 400 μL of a mixed isotope internal standard. The solution was remixed with 20 mL of an acetonitrile–water solution after standing for 30 min, vigorously shaken on a shaking bed for 4 h at 4 °C, and then centrifuged at 10,000 rpm for 5 min. The supernatants were carefully transferred to a new tube. The supernatants were combined and filtered using a syringe-facilitated 13-mm diameter nylon filter with a pore size of 0.22 μm (Nylon 66; Jinteng Experiment Equipment Co., Ltd, Tianjing, China). The filtrate was dried by evaporation under the nitrogen gas flow for ~5 h at room temperature, and then dissolved in 200 μL of methanol. The method for DON determination was performed as described by the National Agricultural Technology Extension Service Center of China (GB5009.111-2016).
    RNA sequencing and analysis
    Sterilized wheat seeds were inoculated with strain DT-8, and then were sown in experimental fields located at EZhou City in 2017. Wheat flag leaves and spikes were collected from this field during the initial bloom stage, with three spikes and leaves being randomly sampled from each of the three replicate plots. The flag leaves and spikes of nontreated wheat plants in the same field were randomly taken from each of the three control replicate plots. The samples were immediately placed in liquid nitrogen and ground into powder. Total RNA samples were extracted with a TRIzol Plus RNA Purification Kit (Takara, Dalian, China) and treated with RNase-free DNase I (Takara, Dalian, China) according to the manufacturer’s instructions. The RNA quality was checked using a Nanodrop Spectrophotometer (Thermo Fisher Scientific Inc., Wilmington, DE, USA). Then, mRNA was enriched with magnetic beads Oligo (dT) (TransGen Biotech, Beijing, China). Subsequently, cDNA was synthesized using the mRNA as template. The cDNA fragments were linked with adapters, and suitable fragments were selected for PCR amplification. Agilent 2100 Bioanalyzer and ABI StepOnePlus Real-Time PCR Systems were used in the quantification and qualification of the sample library. Subsequently, the library was sequenced for raw data using an Illumina HiSeq X sequencer at BGI (The Beijing Genomics Institute, China). Then, adapters, low-quality sequences, and reads with high content of unknown base (N) reads were removed to obtain clean reads. The clean reads were then mapped to the wheat genome or S. sclerotiorum genome and the sequence results evaluated in terms of read quality, alignment, saturation, and the distribution of reads on reference genes [30]. Mismatches of no more than two bases were accepted in the alignment. Gene expression was calculated by the number of reads mapped to the reference genomes using the fragments per kilobase of transcript per million mapped reads method [31]. Subsequently, differentially expressed genes (DEGs) were selected with FDR  More

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    Hkakabo Razi landscape as one of the last exemplar of large contiguous forests

    1.
    FAO. Global forest resources assessment 2015: How are the world’s forests changing? 2nd edn, (Food and Agriculture Organization of the United Nations, 2015).
    2.
    Keenan, R. J. et al. Dynamics of global forest area: Results from the FAO global forest resources assessment. For. Ecol. Manag. 2015(352), 9–20 (2015).
    Google Scholar 

    3.
    Alkama, R. & Cescatti, A. Biophysical climate impacts of recent changes in global forest cover. Science 351, 600 (2016).
    ADS  CAS  PubMed  Google Scholar 

    4.
    Venter, O. et al. Sixteen years of change in the global terrestrial human footprint and implications for biodiversity conservation. Nat. Commun. 7, 12558–12558 (2016).
    ADS  CAS  PubMed  PubMed Central  Google Scholar 

    5.
    Duveiller, G., Hooker, J. & Cescatti, A. The mark of vegetation change on Earth’s surface energy balance. Nat. Commun. 9, 679 (2018).
    ADS  PubMed  PubMed Central  Google Scholar 

    6.
    Hansen, M. C. et al. High-resolution global maps of 21st-century forest cover change. Science 342, 850 (2013).
    ADS  CAS  PubMed  PubMed Central  Google Scholar 

    7.
    Song, X.-P. et al. Global land change from 1982 to 2016. Nature 560, 639–643 (2018).
    ADS  CAS  PubMed  PubMed Central  Google Scholar 

    8.
    Morales-Hidalgo, D., Oswalt, S. N. & Somanathan, E. Status and trends in global primary forest, protected areas, and areas designated for conservation of biodiversity from the global forest resources assessment 2015. For. Ecol. Manag. 352, 68–77 (2015).
    Google Scholar 

    9.
    Watson, J. E. M. et al. The exceptional value of intact forest ecosystems. Nat. Ecol. Evol. 2, 599–610 (2018).
    PubMed  Google Scholar 

    10.
    Lewis, S. L., Edwards, D. P. & Galbraith, D. Increasing human dominance of tropical forests. Science 349, 827 (2015).
    ADS  CAS  PubMed  Google Scholar 

    11.
    Potapov, P. et al. The last frontiers of wilderness: Tracking loss of intact forest landscapes from 2000 to 2013. Sci. Adv. 3, e1600821 (2017).
    ADS  PubMed  PubMed Central  Google Scholar 

    12.
    Curtis, P. G., Slay, C. M., Harris, N. L., Tyukavina, A. & Hansen, M. C. Classifying drivers of global forest loss. Science 361, 1108 (2018).
    ADS  CAS  PubMed  Google Scholar 

    13.
    Achard, F. et al. Determination of tropical deforestation rates and related carbon losses from 1990 to 2010. Glob. Change Biol. 20, 2540–2554 (2014).
    ADS  Google Scholar 

    14.
    Sloan, S. & Sayer, J. A. Forest Resources Assessment of 2015 shows positive global trends but forest loss and degradation persist in poor tropical countries. For. Ecol. Manag. 352, 134–145 (2015).
    Google Scholar 

    15.
    Leimgruber, P. et al. Forest cover change patterns in Myanmar (Burma) 1990–2000. Environ. Conserv. 32, 356–364 (2005).
    Google Scholar 

    16.
    Bhagwat, T. et al. Losing a jewel—Rapid declines in Myanmar’s intact forests from 2002–2014. PLoS ONE 12, e0176364 (2017).
    PubMed  PubMed Central  Google Scholar 

    17.
    FAO. Forests and tree supporting rural livelihoods: Case studies from Myanmar and Viet Nam by Kollert, W. Thuy, L.T.T., Voan, V.L, Oo, T.S. and Khaing, N. Planted Forests and Trees Working Paper FP/50/E. Rome, Italy (available at https://www.fao.org/3/a-i6710e.pdf) (2017).

    18.
    Kyaw, W. W., Sukchai, S., Ketjoy, N. & Ladpala, S. Energy utilization and the status of sustainable energy in Union of Myanmar. Energy Proc. 9, 351–358 (2011).
    Google Scholar 

    19.
    Mon, M. S., Mizoue, N., Htun, N. Z., Kajisa, T. & Yoshida, S. Factors affecting deforestation and forest degradation in selectively logged production forest: A case study in Myanmar. For. Ecol. Manag. 267, 190–198 (2012).
    Google Scholar 

    20.
    Woods, K. Timber trade flows and actors in Myanmar: The political economy of Myanmar’s timber trade. (2013).

    21.
    Lim, C. L., Prescott, G. W., De Alban, J. D. T., Ziegler, A. D. & Webb, E. L. Untangling the proximate causes and underlying drivers of deforestation and forest degradation in Myanmar. Conserv. Biol. 31, 1362–1372 (2017).
    PubMed  Google Scholar 

    22.
    Myers, N., Mittermeier, R. A., Mittermeier, C. G., da Fonseca, G. A. B. & Kent, J. Biodiversity hotspots for conservation priorities. Nature 403, 853–858 (2000).
    ADS  CAS  PubMed  PubMed Central  Google Scholar 

    23.
    Mittermeier, R. A., Turner, W. R., Larsen, F. W., Brooks, T. M. & Gascon, C. Global Biodiversity Conservation: The critical role of hotspot in Biodiversity Hotspots (eds F.E. Zachos & J.C. Habel) 3–22 (Springer, Berlin, 2011).

    24.
    Schaefer, H., Bartholomew, B. & Boufford, D. E. Indofevillea jiroi (Cucurbitaceae), a new floral oil producing species from Northeastern Myanmar. Bione 17, 323–332 (2012).
    Google Scholar 

    25.
    Hughes, M., Aung, M. M. & Armstrong, K. An updated checklist and new species of Begonia (B. rheophytica) from Myanmar. Edinb. J. Bot. 76, 285–295 (2019).
    Google Scholar 

    26.
    Rodda, M., Aung, M. M. & Armstrong, K. A new species, a new subspecies, and new records of Hoya (Apocynaceae, Asclepiadoideae) from Myanmar and China. Brittonia 71, 424–434 (2019).
    Google Scholar 

    27.
    Yang, B., Zhou, S.-S., Maung, W. & Tan, Y.-H. Two new species of Impatiens (Balsaminaceae) from Putao, Kachin State, northern Myanmar. Phytotaxa 321, 103–113 (2017).
    Google Scholar 

    28.
    Tong, Y. H. & Xia, N. H. New taxa of Agapetes (Ericaceae) from Myanmar. Phytotaxa 184, 39–45 (2014).
    Google Scholar 

    29.
    Rabinowitz, A., Amato, G. & Saw, T. K. Discovery of the black muntjac, Muntiacus crinifrons (Artiodactyla, Cervidae), in north Myanmar. Mammalia 62, 105–107 (1998).
    Google Scholar 

    30.
    Amato, G., Egan, M. G. & Rabinowitz, A. A new species of muntjac, Muntiacus putaoensis (Artiodactyla: Cervidae) from northern Myanmar. Anim. Conserv. 2, 1–7 (1999).
    Google Scholar 

    31.
    Soisook, P. et al. A new species of Murina (Chiroptera: Vespertilionidae) from sub-Himalayan forests of northern Myanmar. Zootaxa 4320, 159–172 (2017).
    Google Scholar 

    32.
    Rappole, J. H., Renner, S. C., Shwe, N. M. & Sweet, P. R. A new species of Scimitar-Babbler (Timaliidae: Jabouilleia) from the sub-Himalayan region of Myanmar. Auk 122, 1064–1069 (2005).
    Google Scholar 

    33.
    Rappole, J. H., Rasmussen, P. C., Aung, T., Milensky, C. M. & Renner, S. C. Observations on a new species: The Naung Mung Scimitar-Babbler Jabouilleia naungmungensis. Ibis 150, 623–627 (2008).
    Google Scholar 

    34.
    Renner, S. C., Rappole, J. H., Kyaw, M., Milensky, C. M. & Päckert, M. Genetic confirmation of the species status of Jabouilleia naungmungensis. J. Ornithol. 159, 63–71 (2018).
    Google Scholar 

    35.
    Päckert, M. et al. Pilot biodiversity assessment of the Hkakabo Razi passerine avifauna in northern Myanmar—implications for conservation from molecular genetics. Bird Conserv. Int. 30, 267–288 (2020).
    Google Scholar 

    36.
    Bates, P. et al. Intact forests of Hkakabo Razi Landscape are a hotspot of bat diversity in Southeast Asia. Oryx (In Press).

    37.
    Oo, S. S. L., Kyaw, M., Hlaing, N. M. & Renner, S. C. New to Myanmar: the Rosy Starling Pastor roseus (Aves: Passeriformes: Sturnidae) in the Hkakabo Razi Landscape. JoTT 12, 15493–15494 (2020).
    Google Scholar 

    38.
    Oo, S. S. L., Kyaw, M., Meyers, K. & Renner, S. C. Confirmation of the White-winged Duck from the Hkakabo Razi Landscape, Myanmar. BirdingASIA 30, 86–87 (2018).
    Google Scholar 

    39.
    Renner, S. C. et al. Land cover in the Northern forest complex of Myanmar: New insights for conservation. Oryx 41, 27–37 (2007).
    Google Scholar 

    40.
    Rao, M. et al. Biodiversity conservation in a changing climate: A review of threats and implications for conservation planning in Myanmar. Ambio 42, 789–804 (2013).
    PubMed  PubMed Central  Google Scholar 

    41.
    Webb, E. L., Phelps, J., Friess, D. A., Rao, M. & Ziegler, A. D. Environment-friendly reform in Myanmar. Science 336, 295–295 (2012).
    ADS  CAS  PubMed  Google Scholar 

    42.
    Prescott, G. W. et al. Political transition and emergent forest-conservation issues in Myanmar. Conserv. Biol. 31, 1257–1270 (2017).
    PubMed  Google Scholar 

    43.
    De Alban, D. J. et al. Integrating analytical frameworks to investigate land-cover regime shifts in dynamic landscapes. Sustainability 11, 1139 (2019).
    Google Scholar 

    44.
    Clifton, J., Hampton, M. P. & Jeyacheya, J. Opening the box? Tourism planning and development in Myanmar: Capitalism, communities and change. Asia Pac. Viewpoint 59, 323–337 (2018).
    Google Scholar 

    45.
    Belle, E., Shi, Y. & Bertzky, B. Comparative analysis methodology for World Heritage nominations under biodiversity criteria: A contribution to the IUCN evaluation of natural World Heritage nominations. 21 (UNEP-WCMC and IUCN, Cambridge, UK and Gland, Switzerland, 2014).

    46.
    Renner, S. C. et al. Avifauna of the Southeastern Himalayan mountains and neighboring Myanmar hill country. Bonn Zoological Bulletin—Supplementum 62, 1–75 (2015).
    Google Scholar 

    47.
    BirdLife International. Endemic Bird Area factsheet: Eastern Himalayas (130), (2015).

    48.
    BirdLife International. Endemic Bird Area factsheet: Yunnan mountains (139), (2015).

    49.
    BirdLife International. Endemic Bird Area factsheet: Northern Myanmar lowlands (s079), (2015).

    50.
    Karger, D. N. et al. Climatologies at high resolution for the Earth’s land surface areas. Sci. Data. 4, 170122 (2017).
    PubMed  PubMed Central  Google Scholar 

    51.
    Renner, S. C. & Rappole, J. H. Bird diversity, biogeographic patterns, and endemism of the eastern Himalayas and southeastern Sub-Himalayan mountains in Ornithological Monographs Vol. 70 (ed M. L. Morrison) Ch. 8, 153–166 (American Ornithologists’ Union, 2011).

    52.
    Dumbacher, J. P., Miller, J. R., Flannery, M. E. & Yang Xiaojun. Avifauna of the Gaoligong Shan mountains of western China: A hotspot of avian species diversity in Ornithological Monographs Vol. 70 (eds S.C. Renner & J.H. Rappole) Ch. 3, 30–63 (American Ornithologists’ Union, 2011).

    53.
    Rappole, J. H., Thein Aung, Rasmussen, P. C. & Renner, S. C. Ornithological exploration in the southeastern sub-Himalayan region of Myanmar in Ornithological Monographs Vol. 70 (ed M. L. Morrison) Ch. 2, 10–29 (American Ornithologists’ Union, 2011).

    54.
    Zhu, Z., Wang, S. & Woodcock, C. E. Improvement and expansion of the Fmask algorithm: cloud, cloud shadow, and snow detection for Landsats 4–7, 8, and Sentinel 2 images. Remote Sens. Environ. 159, 269–277 (2015).
    ADS  Google Scholar 

    55.
    Riano, D., Chuvieco, E., Salas, J. & Aguado, I. Assessment of different topographic corrections in Landsat-TM data for mapping vegetation types (2003). IEEE T Geosci. Remote 41, 1056–1061 (2003).
    ADS  Google Scholar 

    56.
    Farr, T. G. et al. The shuttle radar topography mission. Rev. Geophys. 45, 2 (2007).
    Google Scholar 

    57.
    Deng, Y., Chen, X., Chuvieco, E., Warner, T. & Wilson, J. P. Multi-scale linkages between topographic attributes and vegetation indices in a mountainous landscape. Remote Sens. Environ. 111, 122–134 (2007).
    ADS  Google Scholar 

    58.
    Guisan, A., Weiss, S. B. & Weiss, A. D. GLM versus CCA spatial modeling of plant species distribution. Plant Ecol. 143, 107–122 (1999).
    Google Scholar 

    59.
    Running, S. W. Estimating primary productivity by combining remote sensing with ecosystem simulation in Remote Sensing of Biosphere Functioning (eds R.J. Hobbs & H.A Mooney) 65–86 (Springer-Verlag, Berlin, 1990).

    60.
    Myneni, R. B., Hall, F., Sellers, P. & Marshak, A. The interpretation of spectral vegetation indexes. IEEE Trans. Geosci. Rem. Sens. 33, 481–486 (1995).
    ADS  Google Scholar 

    61.
    Gorelick, N. et al. Google Earth Engine: Planetary-scale geospatial analysis for everyone. Remote Sens. Environ. 202, 18–27 (2017).
    ADS  Google Scholar 

    62.
    Liaw, A. & Wiener, M. Classification and regression by random. Forest 2, 18–22 (2002).
    Google Scholar 

    63.
    Plummer, M.JAGS: A program for analysis of Bayesian graphical models using Gibbs sampling in Proceedings of the 3rd international workshop on distributed statistical computing. 125 (Vienna).

    64.
    R: A language and environment for statistical computing (R Foundation for Statistical Computing, Vienna, Austria, 2014).

    65.
    Rodriguez-Galiano, V. F., Ghimire, B., Rogan, J., Chica-Olmo, M. & Rigol-Sanchez, J. P. An assessment of the effectiveness of a random forest classifier for land-cover classification. ISPRS J. Photogramm. Remote Sens. 67, 93–104 (2012).
    ADS  Google Scholar 

    66.
    Connette, G., Oswald, P., Songer, M. & Leimgruber, P. Mapping distinct forest types improves overall forest identification based on multi-spectral landsat imagery for Myanmar’s Tanintharyi region. Remote Sens. 8, 2 (2016).
    Google Scholar 

    67.
    De Alban, J. D., Connette, G., Oswald, P. & Webb, E. Combined Landsat and L-Band SAR data improves land cover classification and change detection in dynamic tropical landscapes. Remote Sens. 10, 306 (2018).
    ADS  Google Scholar 

    68.
    Belgiu, M. & Drăguţ, L. Random forest in remote sensing: A review of applications and future directions. ISPRS J. Photogram. Sens. 114, 24–31 (2016).
    Google Scholar 

    69.
    Horning, N. Random Forests: An algorithm for image classification and generation of continuous fields data sets. (2010).

    70.
    SNAP – ESA Sentinel Application Platform v2.0 (2015).

    71.
    Olofsson, P. et al. Good practices for estimating area and assessing accuracy of land change. Remote Sens. Environ. 148, 42–57 (2014).
    ADS  Google Scholar 

    72.
    Colditz, R. R. et al. Potential effects in multi-resolution post-classification change detection. Int. J. Remote Sens. 33, 6426–6445 (2012).
    ADS  Google Scholar 

    73.
    Cuba, N. Research note: Sankey diagrams for visualizing land cover dynamics. Landsc. Urban Plan. 139, 163–167 (2015).
    Google Scholar 

    74.
    Riitters, K. H. et al. Fragmentation of continental United States forests. Ecosystem 5, 815–822 (2002).
    Google Scholar 

    75.
    Riitters, K. H. & Wickham, J. D. Decline of forest interior conditions in the conterminous United States. Sci. Rep. 2, 653 (2012).
    ADS  PubMed  PubMed Central  Google Scholar 

    76.
    Riitters, K. H., O’Neill, R. V. & Jones, K. B. Assessing habitat suitability at multiple scales: A landscape-level approach. Biol. Conserv. 81, 191–202 (1997).
    Google Scholar 

    77.
    McIntyre, S. & Hobbs, R. A framework for conceptualizing human effects on landscapes and its relevance to management and research models. Conserv. Biol. 13, 1282–1292 (1999).
    Google Scholar 

    78.
    Vogt, P. & Riitters, K. GuidosToolbox: universal digital image object analysis. Eur. J. Remote Sens. 50, 352–361 (2017).
    Google Scholar 

    79.
    Gillanders, S. N., Coops, N. C., Wulder, M. A., Gergel, S. E. & Nelson, T. Multitemporal remote sensing of landscape dynamics and pattern change: Describing natural and anthropogenic trends. Prog. Phys. Geogr. 32, 503–528 (2008).
    Google Scholar 

    80.
    Rubiano, K., Clerici, N., Norden, N. & Etter, A. Secondary forest and shrubland dynamics in a highly transformed landscape in the northern Andes of Colombia (1985–2015). Forest 8, 216 (2017).
    Google Scholar 

    81.
    IUSS, W. G. W. World Reference Base for Soil Resources 2014, update 2015. International soil classification system for naming soils and creating legends for soil maps, (2015).

    82.
    Oldeman, L., Hakkeling, R. & Sombroek, W. World map of the status of human-induced soil degradation: An explanatory note rev. (UNEP and ISRIC, Wageningen, 1991).
    Google Scholar 

    83.
    Fick, S. E. & Hijmans, R. J. Worldclim 2: New 1-km spatial resolution climate surfaces for global land areas. Int. J. Climatol. 37, 4302–4315 (2017).
    Google Scholar 

    84.
    Venables, W. N. & Ripley, B. D. Modern applied statistics with S 4th edn. (Springer, Berlin, 2002).
    Google Scholar 

    85.
    Greene, W. H. Econometric analysis (Prentice Hall, Pearson, 2000).
    Google Scholar 

    86.
    Songer, M., Aung Myint, S. B., DeFries, R. & Leimgruber, P. Spatial and temporal deforestation dynamics in protected and unprotected dry forests: A case study from Myanmar (Burma). Metrics 18, 1001–1018 (2008).
    Google Scholar 

    87.
    Reddy, C. S. et al. Quantifying and predicting multi-decadal forest cover changes in Myanmar: A biodiversity hotspot under threat. Metrics 28, 1129–1149 (2019).
    Google Scholar 

    88.
    Hall, C. A. S., Tian, H., Qi, Y., Pontius, G. & Cornell, J. Modelling spatial and temporal patterns of tropical land use change. J. Biogrph. 22, 753–757 (1995).
    Google Scholar 

    89.
    Di Lallo, G., Mundhenk, P., Zamora López, S., Marchetti, M. & Köhl, M. REDD+: Quick assessment of deforestation risk based on available data. Forests 8, 29 (2017).
    Google Scholar 

    90.
    Bax, V. & Francesconi, W. Environmental predictors of forest change: An analysis of natural predisposition to deforestation in the tropical Andes region, Peru. Appl. Geogr. 91, 99–110 (2018).
    Google Scholar 

    91.
    Pacheco, P. et al. Landscape transformation in tropical Latin America: Assessing trends and policy implications for REDD+. Forest 2, 1–29 (2010).
    Google Scholar  More

  • in

    Temperature increase altered Daphnia community structure in artificially heated lakes: a potential scenario for a warmer future

    1.
    IPCC. Summary for policymakers 1–32 (Cambridge, United Kingdom and New York, NY, USA, 2014).
    2.
    Carey, C. C., Ibelings, B. W., Hoffmann, E. P., Hamilton, D. P. & Brookes, J. D. Eco-physiological adaptations that favour freshwater cyanobacteria in a changing climate. Water Res. 46, 1394–1407 (2012).
    CAS  PubMed  Google Scholar 

    3.
    Daufresne, M., Lengfellner, K. & Sommer, U. Global warming benefits the small in aquatic ecosystems. Proc. Natl. Acad. Sci. USA 106, 12788–12793 (2009).
    ADS  CAS  PubMed  Google Scholar 

    4.
    Forster, J., Hirst, A. G. & Atkinson, D. Warming-induced reductions in body size are greater in aquatic than terrestrial species. Proc. Natl. Acad. Sci. USA 109, 19310–19314 (2012).
    ADS  CAS  PubMed  Google Scholar 

    5.
    Pörtner, H. O. & Farrell, A. P. Physiology and climate change. Science 322, 690–692 (2008).
    PubMed  Google Scholar 

    6.
    De Senerpont Domis, L. N., Bartosiewicz, M., Davis, C. & Cerbin, S. The effect of small doses of toxic cyanobacterial food on the temperature response of Daphnia galeata: is bigger better? Freshw. Biol. 58, 560–572 (2013).

    7.
    Magnuson, J. J. et al. Historical trends in lake and river ice cover in the northen hemisphere. Science 289, 1743–1746 (2000).
    ADS  CAS  PubMed  Google Scholar 

    8.
    Schoebel, C. N., Tellenbach, C., Spaak, P. & Wolinska, J. Temperature effects on parasite prevalence in a natural hybrid complex. Biol. Lett. 7, 108–111 (2011).
    PubMed  Google Scholar 

    9.
    Winder, M. & Schindler, D. E. Climate change uncouples trophic interactions in an aquatic ecosystem. Ecology 85, 2100–2106 (2004).
    Google Scholar 

    10.
    Verschoor, A. M., Van Dijk, M. A., Huisman, J. & Van Donk, E. Elevated CO2 concentrations affect the elemental stoichiometry and species composition of an experimental phytoplankton community. Freshw. Biol. 58, 597–611 (2013).
    CAS  Google Scholar 

    11.
    Zander, A., Bersier, L.-F. & Gray, S. M. Effects of temperature variability on community structure in a natural microbial food web. Glob. Change Biol. 23, 56–67 (2017).
    ADS  Google Scholar 

    12.
    Thomas, C. D. et al. Extinction risk from climate change. Nature 427, 145–148 (2004).
    ADS  CAS  PubMed  PubMed Central  Google Scholar 

    13.
    Urban, M. C. Accelerating extinction risk from climate change. Science 348, 571–573 (2015).
    ADS  CAS  PubMed  Google Scholar 

    14.
    Bellard, C., Bertelsmeier, C., Leadley, P., Thuiller, W. & Courchamp, F. Impacts of climate change on the future of biodiversity. Ecol. Lett. 15, 365–377 (2012).
    PubMed  PubMed Central  Google Scholar 

    15.
    Araújo, M. B. et al. Heat freezes niche evolution. Ecol. Lett. 16, 1206–1219 (2013).
    PubMed  Google Scholar 

    16.
    Meester, L. D., Stoks, R. & Brans, K. I. Genetic adaptation as a biological buffer against climate change: potential and limitations. Integr. Zool. 13, 372–391 (2018).
    PubMed  PubMed Central  Google Scholar 

    17.
    Scranton, K. & Amarasekare, P. Predicting phenological shifts in a changing climate. Proc. Natl. Acad. Sci. USA 114, 13212–13217 (2017).
    CAS  PubMed  Google Scholar 

    18.
    Hulme, P. E. Climate change and biological invasions: evidence, expectations, and response options. Biol. Rev. 92, 1297–1313 (2017).
    PubMed  Google Scholar 

    19.
    Parmesan, C. Ecological and evolutionary responses to recent climate change. Annu. Rev. Ecol. Evol. Syst. 37, 637–669 (2006).
    Google Scholar 

    20.
    Van Doorslaer, W. et al. Local adaptation to higher temperatures reduces immigration success of genotypes from a warmer region in the water flea Daphnia. Glob. Change Biol. 15, 3046–3055 (2009).
    ADS  Google Scholar 

    21.
    Bellard, C. et al. Will climate change promote future invasions?. Glob. Change Biol. 19, 3740–3748 (2013).
    ADS  Google Scholar 

    22.
    Holzapfel, A. M. & Vinebrooke, R. D. Environmental warming increases invasion potential of alpine lake communities by imported species. Glob. Change Biol. 11, 2009–2015 (2005).
    Google Scholar 

    23.
    Burns, C. W. Predictors of invasion success by Daphnia species: influence of food, temperature and species identity. Biol. Invas. 15, 859–869 (2013).
    Google Scholar 

    24.
    Spaak, P., Fox, J. & Hairston, N. G. Jr. Modes and mechanisms of a Daphnia invasion. Proc. Biol. Sci. 279, 2936–2944 (2012).
    PubMed  PubMed Central  Google Scholar 

    25.
    Wejnerowski, Ł., Sikora-Koperska, A. & Dawidowicz, P. Temperature elevation reduces the sensitivity of invasive cladoceran Daphnia lumholtzi to filamentous cyanobacterium Raphidiopsis raciborskii. Freshw Biol 935–946, https://doi.org/10.1111/fwb.13480 (2020).

    26.
    Wittmann, M. J., Gabriel, W., Harz, E.-M., Laforsch, C. & Jeschke, J. Can Daphnia lumholtzi invade European lakes?. NeoBiota 16, 39–57 (2013).
    Google Scholar 

    27.
    Keller, B., Wolinska, J., Manca, M. & Spaak, P. Spatial, environmental and anthropogenic effects on the taxon composition of hybridizing Daphnia. Philos. Trans. R. Soc. Lond, B Biol. Sci. 363, 2943–2952 (2008).
    Google Scholar 

    28.
    Petrusek, A. et al. A taxonomic reappraisal of the European Daphnia longispina complex (Crustacea, Cladocera, Anomopoda). Zool Scr. 37, 507–519 (2008).
    Google Scholar 

    29.
    Zeis, B., Horn, W., Gigengack, U., Koch, M. & Paul, R. J. A major shift in Daphnia genetic structure after the first ice-free winter in a German reservoir. Freshw. Biol. 55, 2296–2304 (2010).
    Google Scholar 

    30.
    Van Doorslaer, W., Stoks, R., Duvivier, C., Bednarska, A. & De Meester, L. Population dynamics determine genetic adaptation to temperature in Daphnia. Evolution 63, 1867–1878 (2009).
    PubMed  Google Scholar 

    31.
    Woszczyk, M. et al. Stable C and N isotope record of short term changes in water level in lakes of different morphometry: Lake Anastazewo and Lake Skulskie, central Poland. Org. Geochem. 76, 278–287 (2014).
    CAS  Google Scholar 

    32.
    Bernatowicz, P., Radzikowski, J., Paterczyk, B., Bebas, P. & Slusarczyk, M. Internal structure of Daphnia ephippium as an adaptation to dispersion. Zool Anz. 277, 12–22 (2018).
    Google Scholar 

    33.
    Moss, B. et al. Climate change and the future of freshwater biodiversity in Europe: a primer for policy-makers. Freshw. Rev. 2, 103–130 (2009).
    Google Scholar 

    34.
    Ma, X., Hu, W., Smilauer, P., Yin, M. & Wolinska, J. Daphnia galeata and D. dentifera are geographically and ecologically separated whereas their hybrids occur in intermediate habitats: A survey of 44 Chinese lakes. Mol. Ecol. 28, 785–802 (2019).

    35.
    Dzialowski, A. R., Lennon, J. T. & Smith, V. H. Food web structure provides biotic resistance against plankton invasion attempts. Biol. Invas. 9, 257–267 (2007).
    Google Scholar 

    36.
    Birks, H. H., Whiteside, M. C., Stark, D. M. & Bright, R. C. Recent paleolimnology of three lakes in Northwestern Minnesota. Quat. Res. 6, 249–272 (1976).
    Google Scholar 

    37.
    Tsugeki, N. K., Ishida, S. & Urabe, J. Sedimentary records of reduction in resting egg production of Daphnia galeata in Lake Biwa during the 20th century: a possible effect of winter warming. J. Paleolimnol. 42, 155–165 (2009).
    ADS  Google Scholar 

    38.
    Keller, B., Wolinska, J., Tellenbach, C. & Spaak, P. Reproductive isolation keeps hybridizing Daphnia species distinct. Limnol. Oceanogr. 52, 984–991 (2007).
    ADS  Google Scholar 

    39.
    Spaak, P. & Boersma, M. Predator mediated coexistence of hybrid and parental Daphnia taxa. Arch. Für Hydrobiol. 167, 55–76 (2006).
    Google Scholar 

    40.
    Kozłowski, J., Czarnołęski, M. & Dańko, M. Can optimal resource allocation models explain why ectotherms grow larger in cold?. Integr. Comput. Biol. 44, 480–493 (2004).
    Google Scholar 

    41.
    Angilletta, M. J. Jr., Steury, T. D. & Sears, M. W. Temperature, growth rate, and body size in ectotherms: fitting pieces of a life-history puzzle. Integr. Comput. Biol. 44, 498–509 (2004).
    Google Scholar 

    42.
    Brooks, J. L. & Dodson, S. I. Predation, body size, and composition of plankton. Science 150, 28–35 (1965).
    ADS  CAS  PubMed  Google Scholar 

    43.
    Gliwicz, Z. M. Relative significance of direct and indirect effects of predation by planktivorous fish on zooplankton. Hydrobiologia 272, 201–210 (1994).
    Google Scholar 

    44.
    Maszczyk, P. et al. Combined effects of elevated epilimnetic temperature and metalimnetic hypoxia on the predation rate of planktivorous fish. J. Plankton Res. 41, 709–722 (2019).
    PubMed  PubMed Central  Google Scholar 

    45.
    Świerzowski, A. & Godlewska, M. Effects of hydropower plant activities on fish population, abundance and distribution. Arch. Pol. Fish. 9, 157–172 (2001).
    Google Scholar 

    46.
    Thorslund, A. E. Potential uses of wastewaters and heated effluents. European Inland Fisheries Advisory Commission Occasional Paper No. 5. (Food and Agriculture Organization of the United Nations, 1971).

    47.
    Warren, G. J., Evans, M. S., Jude, D. J. & Ayers, J. C. Seasonal variations in copepod size: effects of temperature, food abundance, and vertebrate predation. J. Plankton Res. 8, 841–853 (1986).
    Google Scholar 

    48.
    Tunowski, J. Zooplankton structure in heated lakes with differing thermal regimes and water retention. Arch. Pol. Fish. 17, 291–303 (2009).
    Google Scholar 

    49.
    Tunowski, J. Changes in zooplankton abundance and community structure in the cooling channel system of the Konin and Pątnów power plants. Arch. Pol. Fish. 17, 279–289 (2009).
    Google Scholar 

    50.
    Stibor, H. & Lampert, W. Components of additive variance in life-history traits of Daphnia hyalina: seasonal differences in the response to predator signals. Oikos 88, 129–138 (2000).
    Google Scholar 

    51.
    Tereshchenko, V. G., Kapusta, A., Wilkońska, H. & Strelnikova, A. P. Long-term changes in 0+ fish assemblages in the littoral zone of heated lakes. I. Diversity, evennes and dynamic phase portrait of species structure. Arch Pol Fish 15, 415–430 (2007).

    52.
    Brzezinski, T. Filamentous cyanobacteria alter the relative fitness in a Daphnia hybrid species complex. Freshw. Biol. 60, 101–110 (2015).
    Google Scholar 

    53.
    Dziuba, M. K., Cerbin, S. & Wejnerowski, L. Is bigger better? A possibility for adaptation of Daphnia to filamentous cyanobacteria in the face of global warming. Hydrobiologia 798, 105–118 (2017).
    Google Scholar 

    54.
    Socha, D. & Hutorowicz, A. Changes in the quantitative relations of the phytoplankton in heated lakes. Arch. Pol. Fish. 17, 239–251 (2009).
    Google Scholar 

    55.
    Geerts, A. N. et al. Rapid evolution of thermal tolerance in the water flea Daphnia. Nat. Clim. Change 5, 665–668 (2015).
    ADS  Google Scholar 

    56.
    Van Doorslaer, W. et al. Experimental thermal microevolution in community-embedded Daphnia populations. Clim. Res. 43, 81–89 (2010).
    Google Scholar 

    57.
    Wolinska, J., Löffler, A. & Spaak, P. Taxon-specific reaction norms to predator cues in a hybrid Daphnia complex. Freshw. Biol. 52, 1198–1209 (2007).
    Google Scholar 

    58.
    Wolinska, J., Bittner, K., Ebert, D. & Spaak, P. The coexistence of hybrid and parental Daphnia: the role of parasites. Proc Biol Sci 273, 1977–1983 (2006).
    PubMed  PubMed Central  Google Scholar 

    59.
    Lindberg, R. T. & Collins, S. Quality–quantity trade-offs drive functional trait evolution in a model microalgal ‘climate change winner’. Ecol. Lett. 23, 780–790 (2020).
    PubMed  Google Scholar 

    60.
    Lampert, W. Daphnia: model herbivore, predator and prey. Pol. J. Ecol. 54, 607–620 (2006).
    Google Scholar 

    61.
    Bartosiewicz, M. et al. Hot tops, cold bottoms: Synergistic climate warming and shielding effects increase carbon burial in lakes. Limnol. Oceanogr. Lett. 4, 132–144 (2019).
    CAS  Google Scholar 

    62.
    Stawecki, K., Zdanowski, B. & Pyka, J. P. Long-term changes in post-cooling water loads from power plants and thermal and oxygen conditions in stratified lakes. Arch. Pol. Fish. 21, 331–342 (2013).
    CAS  Google Scholar 

    63.
    Bledzki, L. A. & Rybak, J. I. Freshwater Crustacean Zooplankton of Europe. Cladocera & Copepoda (Calanoida, Cyclopoida) Key to species identification, with notes on ecology, distribution, methods and introduction to data analysis. (Springer International Publishing Switzerland, 2016).

    64.
    Appleby, P. G. Chronostratigraphic techniques in recent sediments. In Last, W.M. and Smol, J.P., editors, Tracking environmental change using lake sediments volume 1: basin analysis, coring, and chronological techniques. (Kluwer Academic, London, 2001).

    65.
    Bruel, R. & Sabatier, P. Serac: a R package for ShortlivED RAdionuclide Chronology of recent sediment cores. J. Environ. Activity https://doi.org/10.31223/osf.io/f4yma (2020).
    Article  Google Scholar 

    66.
    Szczuciński, W. et al. Modern sedimentation and sediment dispersal pattern on the continental shelf off the Mekong River delta, South China Sea. Glob. Planet. Change 110, 195–213 (2013).
    ADS  Google Scholar 

    67.
    Dabert, M., Witalinski, W., Kazmierski, A., Olszanowski, Z. & Dabert, J. Molecular phylogeny of acariform mites (Acari, Arachnida): Strong conflict between phylogenetic signal and long-branch attraction artifacts. Mol. Phylogenet. Evol. 56, 222–241 (2010).
    PubMed  Google Scholar 

    68.
    Brede, N. et al. Microsatellite markers for European Daphnia. Mol. Ecol. Notes 6, 536–539 (2006).
    CAS  Google Scholar 

    69.
    Toonen, R. J. & Hughes, S. Increased throughput for fragment analysis on ABI Prism 377 automated sequencer using a membrane comb and STR and software. Biotechniques 31, 1320–1324 (2001).
    CAS  PubMed  Google Scholar 

    70.
    Alberto, F. MsatAllele: Visualizes the scoring and binning of microsatellite fragment sizes. R Package Version 104 (2013).

    71.
    Edgar, R. C. MUSCLE: multiple sequence alignment with high accuracy and high throughput. Nucl. Acids Res. 32, 1792–1797 (2004).
    CAS  PubMed  Google Scholar 

    72.
    Kumar, S., Stecher, G., Li, M., Knyaz, C. & Tamura, K. MEGA X: molecular evolutionary genetics analysis across computing platforms. Mol. Biol. Evol. 35, 1547–1549 (2018).
    CAS  PubMed  PubMed Central  Google Scholar 

    73.
    Pritchard, J. K., Stephens, M. & Donnelly, P. Inference of population structure using multilocus genotype data. Genetics 155, 945–959 (2000).
    CAS  PubMed  PubMed Central  Google Scholar 

    74.
    Earl, D. A. & von Holdt, B. M. STRUCTURE HARVESTER: a website and program for visualizing STRUCTURE output and implementing the Evanno method. Conserv. Genet Resour. 4, 359–361 (2012).
    Google Scholar 

    75.
    Evanno, G., Regnaut, S. & Goudet, J. Detecting the number of clusters of individuals using the software structure: a simulation study. Mol. Ecol. 14, 2611–2620 (2005).
    CAS  PubMed  Google Scholar 

    76.
    Jombart, T., Devillard, S. & Balloux, F. Discriminant analysis of principal components: a new method for the analysis of genetically structured populations. BMC Genet. 11, 94 (2010).
    PubMed  PubMed Central  Google Scholar 

    77.
    Chapuis, M.-P. & Estoup, A. Microsatellite null alleles and estimation of population differentiation. Mol. Biol. Evol. 24, 621–631 (2006).
    PubMed  Google Scholar 

    78.
    Bohonak, A. J. IBD (isolation by distance): a program for analyses of isolation by distance. J. Hered. 93, 153–154 (2002).
    CAS  PubMed  Google Scholar  More

  • in

    N2 fixation dominates nitrogen cycling in a mangrove fiddler crab holobiont

    1.
    Lee, S. Y. et al. Reassessment of mangrove ecosystem services. Glob. Ecol. Biogeogr. 23, 726–743 (2014).
    Google Scholar 
    2.
    Kathiresan, K. & Bingham, B. L. Biology of mangroves and mangrove ecosystems. Adv. Mar. Biol. 40, 81–251 (2001).
    Google Scholar 

    3.
    Dittmar, T., Hertkorn, N., Kattner, G. & Lara, R. J. Mangroves, a major source of dissolved organic carbon to the oceans. Glob. Biogeochem. Cycles 20(1), GB1012. https://doi.org/10.1029/2005GB002570 (2006).
    ADS  CAS  Article  Google Scholar 

    4.
    Kristensen, E., Bouillon, S., Dittmard, T. & Marchande, C. Organic carbon dynamics in mangrove ecosystems: A review. Aquat. Bot. 89, 201–219 (2008).
    CAS  Google Scholar 

    5.
    Reef, R., Feller, I. C. & Lovelock, C. E. Nutrition of mangroves. Tree Physiol. 30(9), 1148–1160 (2010).
    CAS  PubMed  Google Scholar 

    6.
    Woolfe, K. J., Dale, P. J. & Brunskill, G. J. Sedimentary C/S relationships in a large tropical estuary: evidence for refractory carbon inputs from mangroves. Geo-Mar. Lett. 15(3–4), 140–144 (1995).
    ADS  Google Scholar 

    7.
    Woitchik, A. F. et al. Nitrogen enrichment during decomposition of mangrove leaf litter in an east African coastal lagoon (Kenya): relative importance of biological nitrogen fixation. Biogeochemistry 39(1), 15–35 (1997).
    CAS  Google Scholar 

    8.
    Zuberer, D. & Silver, W. S. Biological dinitrogen fixation (acetylene reduction) associated with Florida mangroves. Appl. Environ. Microbiol. 35(3), 567–575 (1978).
    CAS  PubMed  PubMed Central  Google Scholar 

    9.
    Kristensen, E. et al. What is bioturbation? The need for a precise definition for fauna in aquatic sciences. Mar. Ecol. Prog. Ser. 446, 285–302 (2012).
    ADS  Google Scholar 

    10.
    Welsh, D. T. It’s a dirty job but someone has to do it: the role of marine benthic macrofauna in organic matter turnover and nutrient recycling to the water column. Chem. Ecol. 19, 321–342 (2003).
    CAS  Google Scholar 

    11.
    Stief, P. Stimulation of microbial nitrogen cycling in aquatic ecosystems by benthic macrofauna: mechanisms and environmental implications. Biogeosciences 10(12), 7829–7846 (2013).
    ADS  Google Scholar 

    12.
    Gilbertson, W. W., Solan, M. & Prosser, J. I. Differential effects of microorganism–invertebrate interactions on benthic nitrogen cycling. FEMS Microbiol. Ecol. 82, 11–12 (2012).
    PubMed  Google Scholar 

    13.
    Laverock, B., Gilbert, J. A., Tait, K., Osborn, A. M. & Widdicombe, S. Bioturbation: impact on the marine nitrogen cycle. Biochem. Soc. Trans. 39, 315–320 (2011).
    CAS  PubMed  Google Scholar 

    14.
    Magri, M. et al. Benthic N pathways in illuminated and bioturbated sediments studied with network analysis. Limnol. Oceanogr. 63, S68–S84. https://doi.org/10.1002/lno.10724 (2018).
    CAS  Article  Google Scholar 

    15.
    Kristensen, E. Mangrove crabs as ecosystem engineers; with emphasis on sediment processes. J. Sea Res. 59, 30–43 (2008).
    ADS  Google Scholar 

    16.
    Booth, J. M., Fusi, M., Marasco, R., Mbobo, T. & Daffonchioco, D. Fiddler crab bioturbation determines consistent changes in bacterial communities across contrasting environmental conditions. Sci. Rep. 9, 3749. https://doi.org/10.1038/s41598-019-40315-0 (2019).
    ADS  CAS  Article  PubMed  PubMed Central  Google Scholar 

    17.
    Cuellar-Gempeler, C. & Leibold, M. A. Multiple colonist pools shape fiddler crab-associated bacterial communities. ISME J. 12(3), 825–837 (2018).
    CAS  PubMed  PubMed Central  Google Scholar 

    18.
    Reinsel, K. A. Impact of fiddler crab foraging and tidal inundation on an intertidal sandflat: season-dependent effects in one tidal cycle. J. Exp. Mar. Biol. Ecol. 313, 1–17 (2004).
    Google Scholar 

    19.
    Nordhaus, I., Diele, K. & Wolff, M. Activity patterns, feeding and burrowing behaviour of the crab Ucides cordatus (Ucididae) in a high intertidal mangrove forest in North Brazil. J. Exp. Mar. Biol. Ecol. 374, 104–112 (2009).
    Google Scholar 

    20.
    Nordhaus, I. & Wolff, M. Feeding ecology of the mangrove crab Ucides cordatus (Ocypodidae): food choice, food quality and assimilation efficiency. Mar. Biol. 151, 1665–1681 (2007).
    Google Scholar 

    21.
    Fanjul, E., Bazterrica, M. C., Escapa, M., Grela, M. A. & Iribarne, O. Impact of crab bioturbation on benthic flux and nitrogen dynamics of Southwest Atlantic intertidal marshes and mudflats. Estuar. Coast. Shelf Sci. 92, 629–638 (2011).
    ADS  CAS  Google Scholar 

    22.
    Quintana, C. O. et al. Carbon mineralization pathways and bioturbation in coastal Brazilian sediments. Sci. Rep. 5, 16122. https://doi.org/10.1038/srep16122 (2015).
    ADS  CAS  Article  PubMed  PubMed Central  Google Scholar 

    23.
    Thongtham, N. & Kristensen, E. Physical and chemical characteristics of mangrove crab (Neoepisesarma versicolor) burrows in the Bangrong mangrove forest, Phuket, Thailand; with emphasis on behavioural response to changing environmental conditions. Vie et Milieu 53, 141–151 (2003).
    Google Scholar 

    24.
    De la Iglesia, H. O., Rodríguez, E. M. & Dezi, R. E. Burrow plugging in the crab Uca uruguayensis and its synchronization with photoperiod and tides. Physiol. Behav. 55(5), 913–919 (1994).
    PubMed  Google Scholar 

    25.
    Arfken, A., Song, B., Bowman, J. S. & Piehler, M. Denitrification potential of the eastern oyster microbiome using a 16S rRNA gene based metabolic inference approach. PLoS ONE 12(9), e0185071. https://doi.org/10.1371/journal.pone.0185071 (2017).
    CAS  Article  PubMed  PubMed Central  Google Scholar 

    26.
    Caffrey, J. M., Hollibaugh, J. T. & Mortazavi, B. Living oysters and their shells as sites of nitrification and denitrification. Mar. Pollut. Bull. 112(1–2), 86–90 (2016).
    CAS  PubMed  Google Scholar 

    27.
    Glud, R. N. et al. Copepod carcasses as microbial hot spots for pelagic denitrification. Limnol. Oceanogr. 60, 2026–2036 (2015).
    ADS  CAS  Google Scholar 

    28.
    Heisterkamp, I. M. et al. Shell biofilm-associated nitrous oxide production in marine molluscs: processes, precursors and relative importance. Environ. Microbiol. 15(7), 1943–1955 (2013).
    CAS  PubMed  Google Scholar 

    29.
    Ray, N. E., Henning, M. C. & Fulweiler, R. W. Nitrogen and phosphorus cycling in the digestive system and shell biofilm of the eastern oyster Crassostrea virginica. Mar. Ecol. Prog. Ser. 621, 95–105 (2019).
    ADS  CAS  Google Scholar 

    30.
    Stief, P. et al. Freshwater copepod carcasses as pelagic microsites of dissimilatory nitrate reduction to ammonium. FEMS Microbiol. Ecol. 94(10), fiy144. https://doi.org/10.1093/femsec/fiy144 (2018).
    CAS  Article  PubMed Central  Google Scholar 

    31.
    Wahl, M., Goecke, F., Labes, A., Dobretsov, S. & Weinberger, F. The second skin: ecological role of epibiotic biofilms on marine organisms. Front. Microbiol. 3, 292. https://doi.org/10.3389/fmicb.2012.00292 (2012).
    CAS  Article  PubMed  PubMed Central  Google Scholar 

    32.
    Foshtomi, M. Y. et al. The link between microbial diversity and nitrogen cycling in marine sediments is modulated by macrofaunal bioturbation. PLoS ONE 10, e0130116. https://doi.org/10.1371/journal.pone.0130116 (2015).
    CAS  Article  Google Scholar 

    33.
    Pelegri, S. P., Nielsen, L. P. & Blackburn, T. H. Denitrification in estuarine sediment stimulated by the irrigation activity of the amphipod Corophium volutator. Mar. Ecol. Prog. Ser. 105(3), 285–290 (1994).
    ADS  Google Scholar 

    34.
    Stief, P. & Beer, D. D. Probing the microenvironment of freshwater sediment macrofauna: Implications of deposit-feeding and bioirrigation for nitrogen cycling. Limnol. Oceanogr. 51, 2538–2548 (2006).
    ADS  Google Scholar 

    35.
    Pischedda, L., Cuny, P., Esteves, J. L., Pogiale, J. C. & Gilbert, F. Spatial oxygen heterogeneity in a Hediste diversicolor irrigated burrow. Hydrobiologia 680, 109–124 (2012).
    CAS  Google Scholar 

    36.
    Poulsen, M., Kofoed, M. V., Larsen, L. H., Schramm, A. & Stief, P. Chironomus plumosus larvae increase fluxes of denitrification products and diversity of nitrate-reducing bacteria in freshwater sediment. Syst. Appl. Microbiol. 37, 51–59 (2014).
    CAS  PubMed  Google Scholar 

    37.
    Petersen, J. M. et al. Chemosynthetic symbionts of marine invertebrate animals are capable of nitrogen fixation. Nat. Microbiol. 2, 16196. https://doi.org/10.1038/nmicrobiol.2016.195 (2016).
    CAS  Article  Google Scholar 

    38.
    Samuiloviene, A. et al. The effect of chironomid larvae on nitrogen cycling and microbial communities in soft sediments. Water 11, 1931. https://doi.org/10.3390/w11091931 (2019).
    CAS  Article  Google Scholar 

    39.
    Reis, C. R. G., Nardoto, G. B. & Oliveira, R. S. Global overview on nitrogen dynamics in mangroves nd consequences of increasing nitrogen availability for these systems. Plant Soil 410, 1–19 (2017).
    CAS  Google Scholar 

    40.
    Nagata, R. M., Moreira, M. Z., Pimentel, C. R. & Morandini, A. C. Food web characterization based on d15N and d13C reveals isotopic niche partitioning between fish and jellyfish in a relatively pristine ecosystem. Mar. Ecol. Progr. Ser. 519, 13–27 (2015).
    ADS  CAS  Google Scholar 

    41.
    Alfaro-Espinoza, G. & Ullrich, M. S. Bacterial N2-fixation in mangrove ecosystems: insights from a diazotroph–mangrove interaction. Front. Microbiol. 6, 445. https://doi.org/10.3389/fmicb.2015.00445 (2015).
    Article  PubMed  PubMed Central  Google Scholar 

    42.
    Jiménez, M.F.S.-S., Cerqueda-García, D., Montero-Muñoz, J. L., Aguirre-Macedo, M. L. & García-Maldonado, J. Q. Assessment of the bacterial community structure in shallow and deep sediments of the Perdido Fold Belt region in the Gulf of Mexico. PeerJ 6, e5583. https://doi.org/10.7717/peerj.5583 (2018).
    CAS  Article  Google Scholar 

    43.
    Wang, Y. et al. Comparison of the levels of bacterial diversity in freshwater, intertidal wetland, and marine sediments by using millions of Illumina tags. Appl. Environ. Microbiol. 78(23), 8264–8271 (2012).
    CAS  PubMed  PubMed Central  Google Scholar 

    44.
    Dias, A. C. F. et al. The bacterial diversity in a Brazilian non-disturbed mangrove sediment. Antonie Van Leeuwenhoek 98, 541–551 (2010).
    PubMed  Google Scholar 

    45.
    Grim, S. L. & Dick, G. J. Photosynthetic versatility in the genome of Geitlerinema sp. PCC (formerly Oscillatoria limnetica ‘Solar Lake’), a model anoxygenic photosynthetic cyanobacterium. Front. Microbiol. 7, 1546. https://doi.org/10.3389/fmicb.2016.01546 (2016).
    Article  PubMed  PubMed Central  Google Scholar 

    46.
    Zehr, J. P., Church, M. J. & Moisander, P. H. Diversity, distribution and biogeochemical significance of nitrogen-fixing microorganisms in anoxic and suboxic ocean environments. In Past and Present Water Column Anoxia. Nato Science Series: IV: Earth and Environmental Sciences (ed. Neretin, L.) 64, 337–369 (Springer, Berlin, 2006).
    Google Scholar 

    47.
    Brauer, V. S. et al. Competition and facilitation between the marine nitrogen-fixing cyanobacterium Cyanothece and its associated bacterial community. Front. Microbiol. 7, 795. https://doi.org/10.3389/fmicb.2014.00795 (2015).
    Article  Google Scholar 

    48.
    Beltrán, Y., Centeno, C. M., García-Oliva, F., Legendre, P. & Falcón, L. I. N2 fixation rates and associated diversity (nifH) of microbialite and mat-forming consortia from different aquatic environments in Mexico. Aquat. Microb. Ecol. 65, 15–24 (2012).
    Google Scholar 

    49.
    Wong, H. L., Smith, D.-L., Visscher, P. T. & Burns, B. P. Niche differentiation of bacterial communities at a millimeter scale in Shark Bay microbial mats. Sci. Rep. 5, 15607. https://doi.org/10.1038/srep15607 (2015).
    ADS  CAS  Article  PubMed  PubMed Central  Google Scholar 

    50.
    Rasigraf, O., Schmitt, J., Jetten, M. S. M. & Lüke, C. Metagenomic potential for and diversity of N-cycle driving microorganisms in the Bothnian Sea sediment. Microbiol. Open 6(4), 1. https://doi.org/10.1002/mbo3.475 (2017).
    CAS  Article  Google Scholar 

    51.
    Zhang, S. et al. Responses of bacterial community structure and denitrifying bacteria in biofilm to submerged macrophytes and nitrate. Sci. Rep. 6, 36178. https://doi.org/10.1038/srep36178 (2016).
    ADS  CAS  Article  PubMed  PubMed Central  Google Scholar 

    52.
    Holmes, A. J., Costello, A., Lidstrom, M. E. & Murrell, J. C. Evidence that particulate methane monooxygenase and ammonia monooxygenase may be evolutionarily related. FEMS Microbiol. Lett. 132(3), 203–208 (1995).
    CAS  PubMed  Google Scholar 

    53.
    Kraft, B. et al. Nitrogen cycling. The environmental controls that govern the end product of bacterial nitrate respiration. Science 345, 676–679 (2014).
    ADS  CAS  PubMed  Google Scholar 

    54.
    Jiang, X., Dang, H. & Jiao, N. Ubiquity and diversity of heterotrophic bacterial nasA genes in diverse marine environments. PLoS ONE 10(2), e0117473. https://doi.org/10.1371/journal.pone.0117473 (2015).
    CAS  Article  PubMed  PubMed Central  Google Scholar 

    55.
    Xu, T. et al. Genomic insight into Aquimarina longa SW024T: its ultra-oligotrophic adapting mechanisms and biogeochemical functions. BMC Genom. 16, 772. https://doi.org/10.1186/s12864-015-2005-3 (2015).
    CAS  Article  Google Scholar 

    56.
    Li, J. et al. Janibacter alkaliphilus sp. nov., isolated from coral Anthogorgia sp. Antonie Van Leeuwenhoek 102(1), 157–162 (2012).
    CAS  PubMed  Google Scholar 

    57.
    Zumft, W. G. Cell biology and molecular basis of denitrification. Microbiol. Mol. Biol. R. 61(4), 533–616 (1997).
    CAS  Google Scholar 

    58.
    Elifantz, H., Horn, G., Ayon, M., Cohen, Y. & Minz, D. Rhodobacteraceae are the key members of the microbial community of the initial biofilm formed in Eastern Mediterranean coastal seawater. FEMS Microbiol. Ecol. 85(2), 348–357 (2013).
    CAS  PubMed  Google Scholar 

    59.
    Glaeser, S. P. & Kämpfer, P. The family Sphingomonadaceae. In The Prokaryotes (eds Rosenberg, E. et al.) 641–707 (Springer, Berlin, 2014).
    Google Scholar 

    60.
    Katayama, Y., Hiraishi, A. & Kuraishi, H. Paracoccus thiocyanatus sp. nov., a new species of thiocyanate-utilizing facultative chemolithotroph, and transfer of Thiobacillus versutus to the genus Paracoccus as Paracoccus versutus comb. nov. with emendation of the genus. Microbiology 141, 1469–1477 (1995).
    CAS  PubMed  Google Scholar 

    61.
    Kraft, B., Tegetmeyer, H. E., Meier, D., Geelhoed, J. S. & Strous, M. Rapid succession of uncultured marine bacterialand archaeal populations in a denitrifying continuous culture. Environ. Microbiol. 16(10), 3275–3286 (2014).
    CAS  PubMed  Google Scholar 

    62.
    Härtig, E. & Zumft, W. G. Kinetics of nirS expression (cytochrome cd1 nitrite reductase) in Pseudomonas stutzeri during the transition from aerobic respiration to denitrification: evidence for a denitrification-specific nitrate- and nitrite-responsive regulatory system. J. Bacteriol. Res. 181(1), 161–166 (1999).
    Google Scholar 

    63.
    Marchant, H. K. et al. Denitrifying community in coastal sediments performs aerobic and anaerobic respiration simultaneously. ISME J. 11, 1799–1812 (2017).
    CAS  PubMed  PubMed Central  Google Scholar 

    64.
    Patureau, D., Zumstein, E., Delgenes, J. P. & Moletta, R. Aerobic denitrifiers isolated from diverse natural and managed ecosystems. Microb. Ecol. 39(2), 145–152 (2000).
    CAS  PubMed  Google Scholar 

    65.
    Ji, B. et al. Aerobic denitrification: a review of important advances of the last 30 years. Biotechnol. Bioproc. E 20(4), 643–651 (2015).
    CAS  Google Scholar 

    66.
    Strous, M. et al. Missing lithotroph identified as new planctomycete. Nature 400, 446–449 (1999).
    ADS  CAS  PubMed  Google Scholar 

    67.
    Luvizotto, D. M. et al. The rates and players of denitrification, dissimilatory nitrate reduction to ammonia (DNRA) and anaerobic ammonia oxidation (anammox) in mangrove soils. An. Acad. Bras. Ciênc. 91, e20180373. https://doi.org/10.1590/0001-3765201820180373 (2018).
    CAS  Article  PubMed  Google Scholar 

    68.
    Weihrauch, D., Sandra Fehsenfeld, S. & Quijada-Rodriguez, A. Nitrogen excretion in aquatic crustaceans. In Acid–Base Balance and Nitrogen Excretion in Invertebrate (eds Weihrauch, D. & O’Donnell, M.) 1–25 (Springer, Berlin, 2017).
    Google Scholar 

    69.
    Jiang, D.-H., Lawrence, A. L., Neill, W. H. & Gong, H. Effects of temperature and salinity on nitrogenous excretion by Litopenaeus vannamei juveniles. J. Exp. Mar. Biol. Ecol. 253(2), 193–209 (2000).
    CAS  PubMed  Google Scholar 

    70.
    Cardini, U. et al. Chemosymbiotic bivalves contribute to the nitrogen budget of seagrass ecosystems. ISME J. 13, 3131–3134 (2019).
    CAS  PubMed  PubMed Central  Google Scholar 

    71.
    Citadin, M., Costa, T. M. & Netto, S. A. The response of meiofauna and microphytobenthos to engineering effects of fiddler crabs on a subtropical intertidal sandflat. Aust. Ecol. 41(5), 572–579 (2016).
    Google Scholar 

    72.
    Dyea, A. H. & Lasiak, T. A. Assimilation efficiencies of fiddler crabs and deposit-feeding gastropods from tropical mangrove sediments. Comp. Biochem. Phys. Part A 87(2), 341–344 (1987).
    Google Scholar 

    73.
    Hopkins, P. Growth and regeneration patterns in the fiddler crab, Uca pugilator. Biol. Bull. 163, 301–319 (1982).
    Google Scholar 

    74.
    Masunari, S. Distribuição e abundância dos caranguejos Uca Leach (Crustacea, Decapoda, Ocypodidae) na Baía de Guaratuba, Paraná, Brasil. Rev. Bras. Zool. 23(4), 901–914 (2006).
    Google Scholar 

    75.
    Fusi, M. et al. Thermal sensitivity of the crab Neosarmatium africanum in tropical and temperate mangroves on the east coast of Africa. Hydrobiologia 803(1), 251–263 (2017).
    Google Scholar 

    76.
    Hemmi, J. M. & Zeil, J. Burrow surveillance in fiddler crabs I. Description of behaviour. J. Exp. Biol. 206, 3935–3950 (2003).
    PubMed  Google Scholar 

    77.
    Christy, J. H. Predation and the reproductive behavior of fiddler crabs (Genus Uca). In Evolutionary Ecology of Social and Sexual Systems—Crustaceans as Model Organisms (eds Duffy, E. J. & Thiel, M.) 211–231 (Oxford University Press, Oxford, 2007).
    Google Scholar 

    78.
    Teal, J. M. Respiration of crabs in Georgia salt marshes and its relation to their ecology. Physiol. Zool. 32, 1–14 (1959).
    Google Scholar 

    79.
    Michaels, R. E. & Zieman, J. C. Fiddler crab (Uca spp.) burrows have little effect on surrounding sediment oxygen concentrations. J. Exp. Mar. Biol. Ecol. 444, 104–113 (2013).
    Google Scholar 

    80.
    Alongi, D. M. Impact of global change on nutrient dynamics in mangrove forests. Forests 9(10), 596. https://doi.org/10.3390/f9100596 (2018).
    Article  Google Scholar 

    81.
    Barrera-Alba, J. J., Gianesella, S. M. F., Moser, G. A. O. & Saldanha-Corrêa, F. M. P. Bacterial and phytoplankton dynamics in a sub-tropical Estuary. Hydrobiologia 598, 229–246 (2008).
    Google Scholar 

    82.
    Bérgamo, A. L. Característica da hidrografia, circulação e transporte de sal: Barra de Cananéia, sul do Mar de Cananéia e Baía do Trapandé (Master in Physical Oceanography) (Universidade de São Paulo, São Paulo, Instituto Oceanográfico, 2000).
    Google Scholar 

    83.
    Cunha-Lignon, M. Dinâmica do Manguezal no Sistema Cananéia-Iguape, Estado de São Paulo—Brasil. Dissertação (Master in Biological Oceanography). Instituto Oceanográfico, Universidade de São Paulo, São Paulo (2001).

    84.
    Milani, C. et al. Assessing the fecal microbiota: an optimized ion torrent 16S rRNA gene-based analysis protocol. PLoS ONE 8, e68739. https://doi.org/10.1371/journal.pone.0068739 (2013).
    ADS  CAS  Article  PubMed  PubMed Central  Google Scholar 

    85.
    Callahan, B. J. et al. DADA2: high-resolution sample inference from Illumina amplicon data. Nat. Methods 13(7), 581–583 (2016).
    CAS  PubMed  PubMed Central  Google Scholar 

    86.
    Andrews, S. FastQC: A Quality Control Tool for High Throughput Sequence Data. Available online at https://www.bioinformatics.babraham.ac.uk/projects/fastqc (2010).

    87.
    Quast, C. et al. The SILVA ribosomal RNA gene database project: Improved data processing and web-based tools. Nucleic Acids Res. 41, D590–D596. https://doi.org/10.1093/nar/gks1219 (2012).
    CAS  Article  PubMed  PubMed Central  Google Scholar 

    88.
    Robertson, C. E. et al. Explicet: graphical user interface software for metadata-driven management, analysis and visualization of microbiome data. Bioinformatics 29(23), 3100–3101 (2013).
    CAS  PubMed  PubMed Central  Google Scholar 

    89.
    Langmead, B. & Salzberg, S. L. Fast gapped-read alignment with Bowtie 2. Nat. Methods 9, 357. https://doi.org/10.1038/nmeth.1923 (2012).
    CAS  Article  PubMed  PubMed Central  Google Scholar 

    90.
    Bolger, A. M., Lohse, M. & Usadel, B. Trimmomatic: a flexible trimmer for Illumina sequence data. Bioinformatics 30, 2114–2120 (2014).
    CAS  PubMed  PubMed Central  Google Scholar 

    91.
    Zhang, J., Kobert, K., Flouri, T. & Stamatakis, A. PEAR: a fast and accurate Illumina paired-end reAd mergeR. Bioinformatics 30, 614–620 (2014).
    CAS  PubMed  Google Scholar 

    92.
    Buchfink, B., Xie, C. & Huson, D. H. Fast and sensitive protein alignment using DIAMOND. Nat. Methods 12, 59–60. https://doi.org/10.1038/nmeth.3176 (2015).
    CAS  Article  PubMed  Google Scholar 

    93.
    Altschul, S. F., Gish, W., Miller, W., Myers, E. W. & Lipman, D. J. Basic local alignment search tool. J. Mol. Biol. Res. 215, 403–410 (1990).
    CAS  Google Scholar 

    94.
    Huson, D. H. & Mitra, S. Introduction to the analysis of environmental sequences: metagenomics with MEGAN. Methods Mol. Biol. 856, 415–429 (2012).
    CAS  PubMed  Google Scholar 

    95.
    Risgaard-Petersen, N. et al. Anaerobic ammonium oxidation in an estuarine. Aquat. Microb. Ecol. 36, 293–304 (2004).
    Google Scholar 

    96.
    Tréguer, P. & Le Corre, P. Manuel d’analysis des sels nutritifs dans l’eau de mer 2nd edn, 110 (Université de Bretagne Occidentale, Brest, 1975).
    Google Scholar 

    97.
    Kana, T. M. et al. Membrane inlet mass spectrometer for rapid high-precision determination of N2, O2, and Ar in environmental water samples. Anal. Chem. 66, 4166–4170 (1994).
    CAS  Google Scholar 

    98.
    Colt, J. Dissolved gas concentration in water: computation as functions of temperature, salinity and pressure 2nd edn. (Elsevier, Amsterdam, 2012).
    Google Scholar 

    99.
    De Brabandere, L. et al. Oxygenation of an anoxic fjord basin strongly stimulates benthic denitrification and DNRA. Biogeochemistry 126(1–2), 131–152 (2015).
    Google Scholar 

    100.
    Warembourg, F. R. Nitrogen fixation in soil and plant systems. In Nitrogen Isotope Techniques (eds Knowles, R. & Blackburn, T. H.) 127–156 (Academic Press, Cambridge, 1993).
    Google Scholar 

    101.
    Thamdrup, B. & Dalsgaard, T. Production of N2 through anaerobic ammonium oxidation coupled to nitrate reduction in marine sediments. Appl. Environ. Microbiol. 68(3), 1312–1318 (2002).
    CAS  PubMed  PubMed Central  Google Scholar 

    102.
    Bonaglia, S. et al. Denitrification and DNRA at the Baltic Sea oxic–anoxic interface: substrate spectrum and kinetics. Limnol. Oceanogr. 61(5), 1900–1915 (2016).
    ADS  CAS  Google Scholar  More

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    Publisher Correction: The tuatara genome reveals ancient features of amniote evolution

    Department of Anatomy, University of Otago, Dunedin, New Zealand
    Neil J. Gemmell, Kim Rutherford, Tim A. Hore, Nicolas Dussex, Helen Taylor, Hideaki Abe & Donna M. Bond

    LOEWE-Center for Translational Biodiversity Genomics, Senckenberg Museum, Frankfurt, Germany
    Stefan Prost

    South African National Biodiversity Institute, National Zoological Garden, Pretoria, South Africa
    Stefan Prost

    School of Life Sciences, Arizona State University, Tempe, AZ, USA
    Marc Tollis, Melissa Wilson & Shawn M. Rupp

    School of Informatics, Computing, and Cyber Systems, Northern Arizona University, Flagstaff, AZ, USA
    Marc Tollis

    School of Fundamental Sciences, Massey University, Palmerston North, New Zealand
    David Winter

    Peralta Genomics Institute, Oakland, CA, USA
    J. Robert Macey, Charles G. Barbieri & Dustin P. DeMeo

    School of Biological Sciences, The University of Adelaide, Adelaide, South Australia, Australia
    David L. Adelson, Terry Bertozzi, Lu Zeng, R. Daniel Kortschak & Joy M. Raison

    Department of Ecology and Genetics – Evolutionary Biology, Evolutionary Biology Centre (EBC), Uppsala University, Uppsala, Sweden
    Alexander Suh, Valentina Peona, Claire R. Peart & Vera M. Warmuth

    Department of Organismal Biology – Systematic Biology, Evolutionary Biology Centre (EBC), Uppsala University, Uppsala, Sweden
    Alexander Suh & Valentina Peona

    Evolutionary Biology Unit, South Australian Museum, Adelaide, South Australia, Australia
    Terry Bertozzi

    Amedes Genetics, Amedes Medizinische Dienstleistungen, Berlin, Germany
    José H. Grau

    Museum für Naturkunde Berlin, Leibniz-Institut für Evolutions- und Biodiversitätsforschung an der Humboldt-Universität zu Berlin, Berlin, Germany
    José H. Grau

    Department of Earth Sciences, Montana State University, Bozeman, MT, USA
    Chris Organ

    Department of Biochemistry, University of Otago, Dunedin, New Zealand
    Paul P. Gardner

    European Molecular Biology Laboratory, European Bioinformatics Institute, Hinxton, UK
    Matthieu Muffato, Mateus Patricio, Konstantinos Billis, Fergal J. Martin & Paul Flicek

    Section for Evolutionary Genomics, The GLOBE Institute, Faculty of Health and Medical Sciences, University of Copenhagen, Copenhagen, Denmark
    Bent Petersen

    Edward Via College of Osteopathic Medicine, Blacksburg, VA, USA
    Lin Kang & Pawel Michalak

    Center for One Health Research, Virginia–Maryland College of Veterinary Medicine, Blacksburg, VA, USA
    Pawel Michalak

    Institute of Evolution, University of Haifa, Haifa, Israel
    Pawel Michalak

    Manaaki Whenua – Landcare Research, Auckland, New Zealand
    Thomas R. Buckley & Victoria G. Twort

    School of Biological Sciences, The University of Auckland, Auckland, New Zealand
    Thomas R. Buckley & Victoria G. Twort

    School of Life and Environmental Sciences, The University of Sydney, Sydney, New South Wales, Australia
    Yuanyuan Cheng

    Biomatters, Auckland, New Zealand
    Hilary Miller

    Department of Vertebrate Zoology, National Museum of Natural History, Smithsonian Institution, Washington, DC, USA
    Ryan K. Schott

    The New Zealand Institute for Plant and Food Research, Auckland, New Zealand
    Melissa D. Jordan & Richard D. Newcomb

    Departamento de Ecología, Facultad de Ciencias Biológicas, Pontificia Universidad Católica de Chile, Santiago, Chile
    José Ignacio Arroyo

    Department of Ecology, Evolution, and Organismal Biology, Iowa State University, Ames, IA, USA
    Nicole Valenzuela, Valeria Velásquez Zapata & Zhiqiang Wu

    Instituto de Investigaciones Biomédicas ‘Alberto Sols’ CSIC-UAM, Madrid, Spain
    Jaime Renart

    Division of Evolutionary Biology, Faculty of Biology, Ludwig-Maximilian University of Munich, Planegg-Martinsried, Germany
    Claire R. Peart & Vera M. Warmuth

    Centre for Genomic Regulation (CRG), The Barcelona Institute for Science and Technology, Universitat Pompeu Fabra (UPF), Barcelona, Spain
    Didac Santesmasses, Marco Mariotti & Roderic Guigó

    School of Biological Sciences, University of Canterbury, Christchurch, New Zealand
    James M. Paterson

    Global Genome Initiative, National Museum of Natural History, Smithsonian Institution, Washington, DC, USA
    Daniel G. Mulcahy & Vanessa L. Gonzalez

    Austrian Institute of Technology (AIT), Center for Health and Bioresources, Molecular Diagnostics, Vienna, Austria
    Stephan Pabinger

    AgResearch, Invermay Agricultural Centre, Mosgiel, New Zealand
    Tracey Van Stijn & Shannon Clarke

    San Diego Zoo Institute for Conservation Research, Escondido, CA, USA
    Oliver Ryder

    Department of Organismic and Evolutionary Biology and the Museum of Comparative Zoology, Harvard University, Cambridge, MA, USA
    Scott V. Edwards

    Department of Biomedical Engineering, Johns Hopkins University, Baltimore, MD, USA
    Steven L. Salzberg

    School of Biological Sciences, Victoria University of Wellington, Wellington, New Zealand
    Lindsay Anderson & Nicola Nelson

    Ngatiwai Trust Board, Whangarei, New Zealand
    Clive Stone, Clive Stone, Jim Smillie & Haydn Edmonds More