Ecology

Publisher’s note Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.This is a summary of: Yin, J. et al. Future socio-ecosystem productivity threatened by compound drought– heatwave events. Nat. Sustain. https://doi.org/10.1038/10.1038/s41893-022-01024-1 (2023). More

Fan, Y. & Pedersen, O. Gut microbiota in human metabolic health and disease. Nat. Rev. Microbiol. 19, 55–71 (2021).Article 
CAS 
PubMed 

Google Scholar 
Schmidt, T. S. B., Raes, J. & Bork, P. The human gut microbiome: from association to modulation. Cell 172, 1198–1215 (2018).Article 
CAS 
PubMed 

Google Scholar 
David, L. A. et al. Diet rapidly and reproducibly alters the human gut microbiome. Nature 505, 559–563 (2014).Article 
CAS 
PubMed 

Google Scholar 
Tap, J. et al. Gut microbiota richness promotes its stability upon increased dietary fibre intake in healthy adults. Environ. Microbiol. 17, 4954–4964 (2015).Article 
CAS 
PubMed 

Google Scholar 
Smits, S. A. et al. Seasonal cycling in the gut microbiome of the Hadza hunter-gatherers of Tanzania. Science 357, 802–806 (2017).Article 
CAS 
PubMed 
PubMed Central 

Google Scholar 
De Filippo, C. et al. Impact of diet in shaping gut microbiota revealed by a comparative study in children from Europe and rural Africa. Proc. Natl Acad. Sci. USA 107, 14691–14696 (2010).Article 
PubMed 
PubMed Central 

Google Scholar 
Morrison, K. E., Jašarević, E., Howard, C. D. & Bale, T. L. It’s the fiber, not the fat: significant effects of dietary challenge on the gut microbiome. Microbiome 8, 15 (2020).Article 
CAS 
PubMed 
PubMed Central 

Google Scholar 
Maslowski, K. M. & Mackay, C. R. Diet, gut microbiota and immune responses. Nat. Immunol. 12, 5–9 (2011).Article 
CAS 
PubMed 

Google Scholar 
Reynolds, A. et al. Carbohydrate quality and human health: a series of systematic reviews and meta-analyses. Lancet 393, 434–445 (2019).Article 
CAS 
PubMed 

Google Scholar 
Slavin, J. Fiber and prebiotics: mechanisms and health benefits. Nutrients 5, 1417–1435 (2013).Article 
CAS 
PubMed 
PubMed Central 

Google Scholar 
Desai, M. S. et al. A dietary fiber-deprived gut microbiota degrades the colonic mucus barrier and enhances pathogen susceptibility. Cell 167, 1339–1353.e21 (2016).Article 
CAS 
PubMed 
PubMed Central 

Google Scholar 
Makki, K., Deehan, E. C., Walter, J. & Bäckhed, F. The impact of dietary fiber on gut microbiota in host health and disease. Cell Host Microbe 23, 705–715 (2018).Article 
CAS 
PubMed 

Google Scholar 
Cantu-Jungles, T. M. et al. Dietary fiber hierarchical specificity: the missing link for predictable and strong shifts in gut bacterial communities. mBio 12, e01028-21 (2022).
Google Scholar 
Murga-Garrido, S. M. et al. Gut microbiome variation modulates the effects of dietary fiber on host metabolism. Microbiome 9, 117 (2021).Article 
CAS 
PubMed 
PubMed Central 

Google Scholar 
Cantu-Jungles, T. M. & Hamaker, B. R. New view on dietary fiber selection for predictable shifts in gut microbiota. mBio 11, e02179-19 (2020).CAS 
PubMed 
PubMed Central 

Google Scholar 
Singh, V. et al. Dysregulated microbial fermentation of soluble fiber induces cholestatic liver cancer. Cell 175, 679–694.e22 (2018).Article 
CAS 
PubMed 
PubMed Central 

Google Scholar 
Terrapon, N., Lombard, V., Gilbert, H. J. & Henrissat, B. Automatic prediction of polysaccharide utilization loci in Bacteroidetes species. Bioinformatics 31, 647–655 (2015).Article 
CAS 
PubMed 

Google Scholar 
Terrapon, N. et al. PULDB: the expanded database of Polysaccharide Utilization Loci. Nucleic Acids Res. 46, D677–D683 (2018).Article 
CAS 
PubMed 

Google Scholar 
Lombard, V., Golaconda Ramulu, H., Drula, E., Coutinho, P. M. & Henrissat, B. The carbohydrate-active enzymes database (CAZy) in 2013. Nucleic Acids Res. 42, D490–D495 (2014).Article 
CAS 
PubMed 

Google Scholar 
Kouzuma, A., Kato, S. & Watanabe, K. Microbial interspecies interactions: recent findings in syntrophic consortia. Front. Microbiol. 6, 477 (2015).Article 
PubMed 
PubMed Central 

Google Scholar 
Faust, K. & Raes, J. Microbial interactions: from networks to models. Nat. Rev. Microbiol. 10, 538–550 (2012).Article 
CAS 
PubMed 

Google Scholar 
Rakoff-Nahoum, S., Coyne, M. J. & Comstock, L. E. An ecological network of polysaccharide utilization among human intestinal symbionts. Curr. Biol. 24, 40–49 (2014).Article 
CAS 
PubMed 

Google Scholar 
Luis, A. S. et al. Dietary pectic glycans are degraded by coordinated enzyme pathways in human colonic Bacteroides. Nat. Microbiol. 3, 210–219 (2018).Article 
CAS 
PubMed 

Google Scholar 
Cartmell, A. et al. A surface endogalactanase in Bacteroides thetaiotaomicron confers keystone status for arabinogalactan degradation. Nat. Microbiol. 3, 1314–1326 (2018).Article 
CAS 
PubMed 
PubMed Central 

Google Scholar 
Rakoff-Nahoum, S., Foster, K. R. & Comstock, L. E. The evolution of cooperation within the gut microbiota. Nature 533, 255–259 (2016).Article 
CAS 
PubMed 
PubMed Central 

Google Scholar 
Pichler, M. J. et al. Butyrate producing colonic Clostridiales metabolise human milk oligosaccharides and cross feed on mucin via conserved pathways. Nat. Commun. 11, 3285 (2020).Article 
CAS 
PubMed 
PubMed Central 

Google Scholar 
Rogowski, A. et al. Glycan complexity dictates microbial resource allocation in the large intestine. Nat. Commun. 6, 7481 (2015).Article 
CAS 
PubMed 

Google Scholar 
Feng, J. et al. Polysaccharide utilization loci in Bacteroides determine population fitness and community-level interactions. Cell Host Microbe https://doi.org/10.1016/j.chom.2021.12.006 (2022).Pollak, S. et al. Public good exploitation in natural bacterioplankton communities. Sci. Adv. 7, eabi4717 (2022).Article 

Google Scholar 
Cuskin, F. et al. Human gut Bacteroidetes can utilize yeast mannan through a selfish mechanism. Nature 517, 165–169 (2015).Article 
CAS 
PubMed 
PubMed Central 

Google Scholar 
Patnode, M. L. et al. Interspecies competition impacts targeted manipulation of human gut bacteria by fiber-derived glycans. Cell 179, 59–73.e13 (2019).Article 
CAS 
PubMed 
PubMed Central 

Google Scholar 
Walter, J., Maldonado-Gómez, M. X. & Martínez, I. To engraft or not to engraft: an ecological framework for gut microbiome modulation with live microbes. Curr. Opin. Biotechnol. 49, 129–139 (2018).Article 
CAS 
PubMed 

Google Scholar 
Jernberg, C., Löfmark, S., Edlund, C. & Jansson, J. K. Long-term ecological impacts of antibiotic administration on the human intestinal microbiota. ISME J. 1, 56–66 (2007).Article 
CAS 
PubMed 

Google Scholar 
Dethlefsen, L., Huse, S., Sogin, M. L. & Relman, D. A. The pervasive effects of an antibiotic on the human gut microbiota, as revealed by deep 16S rRNA sequencing. PLoS Biol. 6, e280 (2008).Article 
PubMed 
PubMed Central 

Google Scholar 
Becattini, S., Taur, Y. & Pamer, E. G. Antibiotic-induced changes in the intestinal microbiota and disease. Trends Mol. Med. 22, 458–478 (2016).Article 
CAS 
PubMed 
PubMed Central 

Google Scholar 
Shade, A. et al. Fundamentals of microbial community resistance and resilience. Front. Microbiol. 3, 417 (2012).Article 
PubMed 
PubMed Central 

Google Scholar 
Coyte, K. Z., Schluter, J. & Foster, K. R. The ecology of the microbiome: networks, competition, and stability. Science 350, 663–666 (2015).Article 
CAS 
PubMed 

Google Scholar 
Stone, L. The stability of mutualism. Nat. Commun. 11, 2648 (2020).Article 
CAS 
PubMed 
PubMed Central 

Google Scholar 
Ratzke, C., Barrere, J. & Gore, J. Strength of species interactions determines biodiversity and stability in microbial communities. Nat. Ecol. Evol. 4, 376–383 (2020).Article 
PubMed 

Google Scholar 
Butler, S. & O’Dwyer, J. P. Stability criteria for complex microbial communities. Nat. Commun. 9, 2970 (2018).Article 
PubMed 
PubMed Central 

Google Scholar 
Li, W. & Stevens, M. H. H. Fluctuating resource availability increases invasibility in microbial microcosms. Oikos 121, 435–441 (2012).Article 

Google Scholar 
Nobuhiko, K. et al. Regulated virulence controls the ability of a pathogen to compete with the gut microbiota. Science 336, 1325–1329 (2012).Article 

Google Scholar 
Maltby, R., Leatham-Jensen, M. P., Gibson, T., Cohen, P. S. & Conway, T. Nutritional basis for colonization resistance by human commensal Escherichia coli strains HS and Nissle 1917 against E. coli O157:H7 in the mouse intestine. PLoS ONE 8, e53957 (2013).Article 
CAS 
PubMed 
PubMed Central 

Google Scholar 
Leatham, M. P. et al. Precolonized human commensal Escherichia coli strains serve as a barrier to E. coli O157:H7 growth in the streptomycin-treated mouse intestine. Infect. Immun. 77, 2876–2886 (2009).Article 
CAS 
PubMed 
PubMed Central 

Google Scholar 
Venturelli, O. S. et al. Deciphering microbial interactions in synthetic human gut microbiome communities. Mol. Syst. Biol. 14, e8157 (2018).Article 
PubMed 
PubMed Central 

Google Scholar 
Clark, R. L. et al. Design of synthetic human gut microbiome assembly and butyrate production. Nat. Commun. 12, 3254 (2021).Article 
CAS 
PubMed 
PubMed Central 

Google Scholar 
Hromada, S. et al. Negative interactions determine Clostridioides difficile growth in synthetic human gut communities. Mol. Syst. Biol. 17, e10355 (2021).Article 
CAS 
PubMed 
PubMed Central 

Google Scholar 
MacArthur, R. Species packing and competitive equilibrium for many species. Theor. Popul. Biol. 1, 1–11 (1970).Article 
CAS 
PubMed 

Google Scholar 
Ndeh, D. et al. Complex pectin metabolism by gut bacteria reveals novel catalytic functions. Nature 544, 65–70 (2017).Article 
CAS 
PubMed 
PubMed Central 

Google Scholar 
Grondin, J. M., Tamura, K., Déjean, G., Abbott, D. W. & Brumer, H. Polysaccharide utilization loci: fueling microbial communities. J. Bacteriol. 199, e00860-16 (2017).Article 
CAS 
PubMed 
PubMed Central 

Google Scholar 
Haiser, H. J. et al. Predicting and manipulating cardiac drug inactivation by the human gut bacterium Eggerthella lenta. Science 341, 295–298 (2013).Article 
CAS 
PubMed 
PubMed Central 

Google Scholar 
Devendran, S. et al. Clostridium scindens ATCC 35704: integration of nutritional requirements, the complete genome sequence, and global transcriptional responses to bile acids. Appl. Environ. Microbiol. 85, e00052-19 (2019).Article 
CAS 
PubMed 
PubMed Central 

Google Scholar 
Rey, F. E. et al. Metabolic niche of a prominent sulfate-reducing human gut bacterium. Proc. Natl Acad. Sci. USA 110, 13582–13587 (2013).Article 
CAS 
PubMed 
PubMed Central 

Google Scholar 
Kaoutari, A. E., Armougom, F., Gordon, J. I., Raoult, D. & Henrissat, B. The abundance and variety of carbohydrate-active enzymes in the human gut microbiota. Nat. Rev. Microbiol. 11, 497–504 (2013).Article 
PubMed 

Google Scholar 
Pereira, F. C. & Berry, D. Microbial nutrient niches in the gut. Environ. Microbiol. 19, 1366–1378 (2017).Article 
PubMed 
PubMed Central 

Google Scholar 
Despres, J. et al. Xylan degradation by the human gut Bacteroides xylanisolvens XB1A(T) involves two distinct gene clusters that are linked at the transcriptional level. BMC Genomics 17, 326 (2016).Article 
PubMed 
PubMed Central 

Google Scholar 
Déjean, G. et al. Synergy between cell surface glycosidases and glycan-binding proteins dictates the utilization of specific beta(1,3)-glucans by human gut bacteroides. mBio 11, e00095-20 (2020).Article 
PubMed 
PubMed Central 

Google Scholar 
Hamaker, B. R. & Tuncil, Y. E. A perspective on the complexity of dietary fiber structures and their potential effect on the gut microbiota. J. Mol. Biol. 426, 3838–3850 (2014).Article 
CAS 
PubMed 

Google Scholar 
Bishop, C. M. Pattern Recognition and Machine Learning (Information Science and Statistics) (Springer, 2006).Wasserman, L. All of Statistics: A Concise Course in Statistical Inference (Springer Texts in Statistics) (Springer, 2003).Willing, B. P., Russell, S. L. & Finlay, B. B. Shifting the balance: antibiotic effects on host–microbiota mutualism. Nat. Rev. Microbiol. 9, 233–243 (2011).Article 
CAS 
PubMed 

Google Scholar 
Panda, S. et al. Short-term effect of antibiotics on human gut microbiota. PLoS ONE 9, e95476 (2014).Article 
PubMed 
PubMed Central 

Google Scholar 
Ng, K. M. et al. Recovery of the gut microbiota after antibiotics depends on host diet, community context, and environmental reservoirs. Cell Host Microbe 26, 650–665.e4 (2019).Article 
CAS 
PubMed 
PubMed Central 

Google Scholar 
Van der Waaij, D., Berghuis-de Vries, J. M. & Lekkerkerk-van der Wees, J. E. C. Colonization resistance of the digestive tract in conventional and antibiotic-treated mice. J. Hygiene 69, 405–411 (1971).Article 

Google Scholar 
Freter, R. In vivo and in vitro antagonism of intestinal bacteria against Shigella flexneri. II. The inhibitory mechanism. J. Infect. Dis. 110, 38–46 (1962).Article 
CAS 
PubMed 

Google Scholar 
Maldonado-Gómez, M. X. et al. Stable engraftment of Bifidobacterium longum AH1206 in the human gut depends on individualized features of the resident microbiome. Cell Host Microbe 20, 515–526 (2016).Article 
PubMed 

Google Scholar 
Sorbara, M. T. & Pamer, E. G. Interbacterial mechanisms of colonization resistance and the strategies pathogens use to overcome them. Mucosal Immunol. 12, 1–9 (2019).Article 
CAS 
PubMed 

Google Scholar 
Litvak, Y. & Bäumler, A. J. The founder hypothesis: a basis for microbiota resistance, diversity in taxa carriage, and colonization resistance against pathogens. PLoS Pathog. 15, e1007563 (2019).Article 
CAS 
PubMed 
PubMed Central 

Google Scholar 
Jenior, M. L., Leslie, J. L., Young, V. B. & Schloss, P. D. Clostridium difficile colonizes alternative nutrient niches during infection across distinct murine gut microbiomes. mSystems 2, e00063-17 (2017).Article 
PubMed 
PubMed Central 

Google Scholar 
Momose, Y., Hirayama, K. & Itoh, K. Competition for proline between indigenous Escherichia coli and E. coli O157:H7 in gnotobiotic mice associated with infant intestinal microbiota and its contribution to the colonization resistance against E. coli O157:H7. Antonie van Leeuwenhoek 94, 165–171 (2008).Article 
CAS 
PubMed 

Google Scholar 
Fabich, A. J. et al. Comparison of carbon nutrition for pathogenic and commensal Escherichia coli strains in the mouse intestine. Infect. Immun. 76, 1143–1152 (2008).Article 
CAS 
PubMed 
PubMed Central 

Google Scholar 
Shepherd, E. S., DeLoache, W. C., Pruss, K. M., Whitaker, W. R. & Sonnenburg, J. L. An exclusive metabolic niche enables strain engraftment in the gut microbiota. Nature 557, 434–438 (2018).Article 
CAS 
PubMed 
PubMed Central 

Google Scholar 
Jenior, M. L., Leslie, J. L., Young, V. B. & Schloss, P. D. Clostridium difficilealters the structure and metabolism of distinct cecal microbiomes during initial infection to promote sustained colonization. mSphere 3, e00261-18 (2018).Article 
CAS 
PubMed 
PubMed Central 

Google Scholar 
Li, S., Tan, J., Yang, X., Ma, C. & Jiang, L. Niche and fitness differences determine invasion success and impact in laboratory bacterial communities. ISME J. 13, 402–412 (2019).Article 
PubMed 

Google Scholar 
Deng, Y.-J. & Wang, S. Y. Synergistic growth in bacteria depends on substrate complexity. J. Microbiol. 54, 23–30 (2016).Article 
PubMed 
PubMed Central 

Google Scholar 
Deng, Y.-J. & Wang, S. Y. Complex carbohydrates reduce the frequency of antagonistic interactions among bacteria degrading cellulose and xylan. FEMS Microbiol. Lett. 364, fnx019 (2017).Article 
PubMed Central 

Google Scholar 
Wu, F. et al. Modulation of microbial community dynamics by spatial partitioning. Nat. Chem. Biol. 18, 394–402 (2022).Article 
CAS 
PubMed 
PubMed Central 

Google Scholar 
Åström, K. J. & Murray, R. Feedback Systems. An Introduction for Scientists and Engineers (Princeton Univ. Press, 2008).Hammarlund, S. P. & Harcombe, W. R. Refining the stress gradient hypothesis in a microbial community. Proc. Natl Acad. Sci. USA 116, 15760–15762 (2019).Article 
CAS 
PubMed 
PubMed Central 

Google Scholar 
Pacheco, A. R., Osborne, M. L. & Segrè, D. Non-additive microbial community responses to environmental complexity. Nat. Commun. 12, 2365 (2021).Article 
CAS 
PubMed 
PubMed Central 

Google Scholar 
Dal Bello, M., Lee, H., Goyal, A. & Gore, J. Resource–diversity relationships in bacterial communities reflect the network structure of microbial metabolism. Nat. Ecol. Evol. 5, 1424–1434 (2021).Article 
PubMed 

Google Scholar 
Magnúsdóttir, S. et al. Generation of genome-scale metabolic reconstructions for 773 members of the human gut microbiota. Nat. Biotechnol. 35, 81–89 (2017).Article 
PubMed 

Google Scholar 
Baranwal, M. et al. Recurrent neural networks enable design of multifunctional synthetic human gut microbiome dynamics. eLife 11, e73870 (2022).Article 
CAS 
PubMed 
PubMed Central 

Google Scholar 
Palleja, A. et al. Recovery of gut microbiota of healthy adults following antibiotic exposure. Nat. Microbiol. 3, 1255–1265 (2018).Article 
CAS 
PubMed 

Google Scholar 
Dethlefsen, L. & Relman, D. A. Incomplete recovery and individualized responses of the human distal gut microbiota to repeated antibiotic perturbation. Proc. Natl Acad. Sci. USA 108, 4554–4561 (2011).Article 
CAS 
PubMed 

Google Scholar 
Ramirez, J. et al. Antibiotics as major disruptors of gut microbiota. Front. Cell. Infect. Microbiol. 10, 572912 (2020).Article 
CAS 
PubMed 
PubMed Central 

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

Google Scholar 
Schloss, P. D. et al. Introducing mothur: open-source, platform-independent, community-supported software for describing and comparing microbial communities. Appl. Environ. Microbiol. 75, 7537–7541 (2009).Article 
CAS 
PubMed 
PubMed Central 

Google Scholar 
Raue, A. et al. Lessons learned from quantitative dynamical modeling in systems biology. PLoS ONE 8, e74335 (2013).Article 
CAS 
PubMed 
PubMed Central 

Google Scholar 
Babtie, A. C., Kirk, P. & Stumpf, M. P. H. Topological sensitivity analysis for systems biology. Proc. Natl Acad. Sci. USA 111, 18507–18512 (2014).Article 
CAS 
PubMed 
PubMed Central 

Google Scholar 
Munsky, B., Hlavacek, W. S. & Tsimring, L. S. Quantitative Biology. Theory, Computational Methods, and Models (MIT Press, 2018).Ashyraliyev, M., Fomekong-Nanfack, Y., Kaandorp, J. A. & Blom, J. G. Systems biology: parameter estimation for biochemical models. FEBS J. 276, 886–902 (2009).Article 
CAS 
PubMed 

Google Scholar 
Ravcheev, D. A., Godzik, A., Osterman, A. L. & Rodionov, D. A. Polysaccharides utilization in human gut bacterium Bacteroides thetaiotaomicron: comparative genomics reconstruction of metabolic and regulatory networks. BMC Genomics 14, 873 (2013).Article 
PubMed 
PubMed Central 

Google Scholar 
Salyers, A. A., Vercelloitti, J. R., West, S. E. & Wilkins, T. D. Fermentation of mucin and plant polysaccharides by strains of Bacteroides from the human colon. Appl. Environ. Microbiol. 33, 319–322 (1977).Article 
CAS 
PubMed 
PubMed Central 

Google Scholar 
Sun, X., Liu, Y., Jiang, P., Song, S. & Ai, C. Interaction of sulfated polysaccharides with intestinal Bacteroidales plays an important role in its biological activities. Int. J. Biol. Macromol. 168, 496–506 (2021).Article 
CAS 
PubMed 

Google Scholar 
Respondek, F. et al. Short-chain fructo-oligosaccharides modulate intestinal microbiota and metabolic parameters of humanized gnotobiotic diet induced obesity mice. PLoS ONE 8, e71026 (2013).Article 
CAS 
PubMed 
PubMed Central 

Google Scholar 
Schwiertz, A. et al. Anaerostipes caccae gen. nov., sp. nov., a new saccharolytic, acetate-utilising, butyrate-producing bacterium from human faeces. Syst. Appl. Microbiol. 25, 46–51 (2002).Article 
CAS 
PubMed 

Google Scholar 
Benítez-Páez, A., Moreno, F. J., Sanz, M. L. & Sanz, Y. Genome structure of the symbiont Bifidobacterium pseudocatenulatum CECT 7765 and gene expression profiling in response to lactulose-derived oligosaccharides. Front. Microbiol. 7, 624 (2016).Article 
PubMed 
PubMed Central 

Google Scholar 
Bernalier, A., Willems, A., Leclerc, M., Rochet, V. & Collins, M. D. Ruminococcus hydrogenotrophicus sp. nov., a new H2/CO2-utilizing acetogenic bacterium isolated from human feces. Arch. Microbiol. 166, 176–183 (1996).Article 
CAS 
PubMed 

Google Scholar 
Moshfegh, A. J., Friday, J. E., Goldman, J. P. & Ahuja, J. K. C. Presence of inulin and oligofructose in the diets of Americans. J. Nutr. 129, 1407S–1411S (1999).Article 
CAS 
PubMed 

Google Scholar 
Sonnenburg, E. D. et al. Specificity of polysaccharide use in intestinal bacteroides species determines diet-induced microbiota alterations. Cell 141, 1241–1252 (2010).Article 
CAS 
PubMed 
PubMed Central 

Google Scholar 
Devillé, C., Damas, J., Forget, P., Dandrifosse, G. & Peulen, O. Laminarin in the dietary fibre concept. J. Sci. Food Agric. 84, 1030–1038 (2004).Article 

Google Scholar 
Selvendran, R. R. The plant cell wall as a source of dietary fiber: chemistry and structure. Am. J. Clin. Nutr. 39, 320–337 (1984).Article 
CAS 
PubMed 

Google Scholar  More

Coleman, D. J. et al. Limnol. Oceanogr. Lett. 7, 140–149 (2022).Article 

Google Scholar 
Noyce, G. L. et al. https://doi.org/10.1038/s41561-022-01070-6 (2022).Kirwan, M. L. & Megonigal, J. P. Nature 504, 53–60 (2013).Article 

Google Scholar 
Morris, J. T., Sundareshwar, P. V., Nietch, C. T., Kjerve, B. & Cahoon, D. R. Ecology 83, 2869–2877 (2002).Article 

Google Scholar 
Noyce, G. L., Kirwan, M. L., Rich, R. L. & Megonigal, J. P. Proc. Natl Acad. Sci. 116, 21623–21628 (2019).Article 

Google Scholar 
Langley, J. A., McKee, K. L., Cahoon, D. R., Cherry, J. A. & Megonigal, J. P. Proc. Natl Acad. Sci. 106, 182–6186 (2009).Article 

Google Scholar 
Dean, J. F. et al. Rev. Geophys. 56, 207–250 (2018).Article 

Google Scholar 
IPCC Climate Change 2021: The Physical Science Basis (eds Masson-Delmotte, V. et al.) (Cambridge University Press, 2021).Lin, Y. et al. Water Res. 205, 117682 (2021).Article 

Google Scholar 
Zakharova, L., Meyer, K. M. & Seifan, M. Ecol. Modell. 407, 108703 (2019).Article 

Google Scholar  More

Moberg, F. & Folke, C. Ecological goods and services of coral reef ecosystems. Ecol. Econ. 29, 215–233 (1999).Article 

Google Scholar 
Plaisance, L., Caley, M. J., Brainard, R. E. & Knowlton, N. The diversity of coral reefs: What are we missing?. PLoS ONE 6, e25026 (2011).Article 
ADS 
CAS 

Google Scholar 
Frieler, K. et al. Limiting global warming to 2 °C is unlikely to save most coral reefs. Nat. Clim. Change 3, 165–170 (2013).Article 
ADS 

Google Scholar 
Hughes, T. P. et al. Climate change, human impacts, and the resilience of coral reefs. Science 301, 929–933 (2003).Article 
ADS 
CAS 

Google Scholar 
Carpenter, K. E. et al. One-third of reef-building corals face elevated extinction risk from climate change and local impacts. Science 321, 560–563 (2008).Article 
ADS 
CAS 

Google Scholar 
Lotze, H. K. et al. Global ensemble projections reveal trophic amplification of ocean biomass declines with climate change. Proc. Natl. Acad. Sci. 116, 12907–12912 (2019).Article 
ADS 
CAS 

Google Scholar 
Doney, S. C. et al. Climate change impacts on marine ecosystems. Annu. Rev. Mar. Sci. 4, 11–37 (2012).Article 
ADS 

Google Scholar 
Van Oppen, M. J., Oliver, J. K., Putnam, H. M. & Gates, R. D. Building coral reef resilience through assisted evolution. Proc. Natl. Acad. Sci. 112, 2307–2313 (2015).Article 
ADS 

Google Scholar 
Parrett, J. M. & Knell, R. J. The effect of sexual selection on adaptation and extinction under increasing temperatures. Proc. R. Soc. B. 285, 20180303 (2018).Article 

Google Scholar 
Hagedorn, M. et al. Assisted gene flow using cryopreserved sperm in critically endangered coral. Proc. Natl. Acad. Sci. 118, e2110559118 (2021).Article 
CAS 

Google Scholar 
Hughes, T. P. et al. Global warming and recurrent mass bleaching of corals. Nature 543, 373–377 (2017).Article 
ADS 
CAS 

Google Scholar 
Hughes, T. P. et al. Global warming transforms coral reef assemblages. Nature 556, 492–496 (2018).Article 
ADS 
CAS 

Google Scholar 
Epstein, N., Bak, R. & Rinkevich, B. Applying forest restoration principles to coral reef rehabilitation. Aquat. Conserv. Mar. Freshw. Ecosyst. 13, 387–395 (2003).Article 

Google Scholar 
West, J. M. & Salm, R. V. Resistance and resilience to coral bleaching: Implications for coral reef conservation and management. Conserv. Biol. 17, 956–967 (2003).Article 

Google Scholar 
Yeemin, T., Sutthacheep, M. & Pettongma, R. Coral reef restoration projects in Thailand. Ocean Coast. Manag. 49, 562–575 (2006).Article 

Google Scholar 
Anthony, K. et al. Operationalizing resilience for adaptive coral reef management under global environmental change. Glob. Chang. Biol. 21, 48–61 (2015).Article 
ADS 

Google Scholar 
Randall, C. J. et al. Sexual production of corals for reef restoration in the Anthropocene. Mar. Ecol. Prog. Ser. 635, 203–232 (2020).Article 
ADS 

Google Scholar 
Porter, J. W., Fitt, W. K., Spero, H. J., Rogers, C. S. & White, M. W. Bleaching in reef corals: Physiological and stable isotopic responses. Proc. Natl. Acad. Sci. 86, 9342–9346 (1989).Article 
ADS 
CAS 

Google Scholar 
Mendes, J. M. & Woodley, J. D. Effect of the 1995–1996 bleaching event on polyp tissue depth, growth, reproduction and skeletal band formation in Montastraea annularis. Mar. Ecol. Prog. Ser. 235, 93–102 (2002).Article 
ADS 

Google Scholar 
Grottoli, A., Rodrigues, L. & Juarez, C. Lipids and stable carbon isotopes in two species of Hawaiian corals, Porites compressa and Montipora verrucosa, following a bleaching event. Mar. Biol. 145, 621–631 (2004).Article 
CAS 

Google Scholar 
Rodrigues, L. J. & Grottoli, A. G. Energy reserves and metabolism as indicators of coral recovery from bleaching. Limnol. Oceanogr. 52, 1874–1882 (2007).Article 
ADS 

Google Scholar 
Levas, S. J., Grottoli, A. G., Hughes, A., Osburn, C. L. & Matsui, Y. Physiological and biogeochemical traits of bleaching and recovery in the mounding species of coral Porites lobata: Implications for resilience in mounding corals. PLoS ONE 8, e63267 (2013).Article 
ADS 
CAS 

Google Scholar 
Schoepf, V. et al. Annual coral bleaching and the long-term recovery capacity of coral. Proc. R. Soc. B. 282, 20151887 (2015).Article 

Google Scholar 
Dai, C., Fan, T. & Yu, J. Reproductive isolation and genetic differentiation of a scleractinian coral Mycedium elephantotus. Mar. Ecol. Prog. Ser. 201, 179–187 (2000).Article 
ADS 

Google Scholar 
Vargas-Ángel, B., Colley, S. B., Hoke, S. M. & Thomas, J. D. The reproductive seasonality and gametogenic cycle of Acropora cervicornis off Broward County, Florida, USA. Coral Reefs 25, 110–122 (2006).Article 
ADS 

Google Scholar 
Rosser, N. & Gilmour, J. New insights into patterns of coral spawning on Western Australian reefs. Coral Reefs 27, 345–349 (2008).Article 
ADS 

Google Scholar 
Szmant, A. M. & Gassman, N. J. The effects of prolonged “bleaching” on the tissue biomass and reproduction of the reef coral Montastrea annularis. Coral Reefs 8, 217–224 (1990).Article 
ADS 

Google Scholar 
Baird, A. H. & Marshall, P. A. Mortality, growth and reproduction in scleractinian corals following bleaching on the Great Barrier Reef. Mar. Ecol. Prog. Ser. 237, 133–141 (2002).Article 
ADS 

Google Scholar 
Levitan, D. R., Boudreau, W., Jara, J. & Knowlton, N. Long-term reduced spawning in Orbicella coral species due to temperature stress. Mar. Ecol. Prog. Ser. 515, 1–10 (2014).Article 
ADS 

Google Scholar 
Ward, S., Harrison, P. & Hoegh-Guldberg, O. Coral bleaching reduces reproduction of scleractinian corals and increases susceptibility to future stress. In Proc. 9th Int. Coral Reef Symp. 1123–1128 (2002).Johnston, E. C., Counsell, C. W., Sale, T. L., Burgess, S. C. & Toonen, R. J. The legacy of stress: Coral bleaching impacts reproduction years later. Funct. Ecol. 34, 2315–2325 (2020).Article 

Google Scholar 
Hirose, M. & Hidaka, M. Reduced reproductive success in scleractinian corals that survived the 1998 bleaching in Okinawa. Galaxea 2000, 17–21 (2000).Article 

Google Scholar 
Omori, M., Fukami, H., Kobinata, H. & Hatta, M. Significant drop of fertilization of Acropora corals in 1999: An after-effect of heavy coral bleaching?. Limnol. Oceanogr. 46, 704–706 (2001).Article 
ADS 

Google Scholar 
Hagedorn, M. et al. Potential bleaching effects on coral reproduction. Reprod. Fertil. Dev. 28, 1061–1071 (2016).Article 
CAS 

Google Scholar 
Bassim, K., Sammarco, P. & Snell, T. Effects of temperature on success of (self and non-self) fertilization and embryogenesis in Diploria strigosa (Cnidaria, Scleractinia). Mar. Biol. 140, 479–488 (2002).Article 

Google Scholar 
Kenkel, C. D. et al. Development of gene expression markers of acute heat-light stress in reef-building corals of the genus Porites. PLoS ONE 6, e26914 (2011).Article 
ADS 
CAS 

Google Scholar 
Louis, Y. D., Bhagooli, R., Kenkel, C. D., Baker, A. C. & Dyall, S. D. Gene expression biomarkers of heat stress in scleractinian corals: Promises and limitations. Comp. Biochem. Physiol. C Toxicol. Pharmacol. 191, 63–77 (2017).Article 
CAS 

Google Scholar 
Bonesso, J. L., Leggat, W. & Ainsworth, T. D. Exposure to elevated sea-surface temperatures below the bleaching threshold impairs coral recovery and regeneration following injury. PeerJ 5, e3719 (2017).Article 

Google Scholar 
Gierz, S., Ainsworth, T. D. & Leggat, W. Diverse symbiont bleaching responses are evident from 2-degree heating week bleaching conditions as thermal stress intensifies in coral. Mar. Freshw. Res. 71, 1149–1160 (2020).Article 

Google Scholar 
Baker, D. M., Freeman, C. J., Wong, J. C., Fogel, M. L. & Knowlton, N. Climate change promotes parasitism in a coral symbiosis. ISME J. 12, 921–930 (2018).Article 
CAS 

Google Scholar 
Yee, S. H. & Barron, M. G. Predicting coral bleaching in response to environmental stressors using 8 years of global-scale data. Environ. Monit. Assess. 161, 423–438 (2010).Article 

Google Scholar 
Lesser, M. P. Coral bleaching: causes and mechanisms. In Coral Reefs: An Ecosystem in Transition (eds Riegl, B. M. & Purkis, S. J.) 405–419 (Springer, 2011).Chapter 

Google Scholar 
Barber, J. & Andersson, B. Too much of a good thing: Light can be bad for photosynthesis. Trends Biochem. Sci. 17, 61–66 (1992).Article 
CAS 

Google Scholar 
Aro, E.-M., Virgin, I. & Andersson, B. Photoinhibition of photosystem II. Inactivation, protein damage and turnover. Biochim. Biophys. Acta Bioenergy 1143, 113–134 (1993).Article 
CAS 

Google Scholar 
Lesser, M. P. & Farrell, J. H. Exposure to solar radiation increases damage to both host tissues and algal symbionts of corals during thermal stress. Coral Reefs 23, 367–377 (2004).Article 

Google Scholar 
Salih, A., Hoegh-Guldberg, O. & Cox, G. Bleaching responses of symbiotic dinoflagellates in corals: the effects of light and elevated temperature on their morphology and physiology. In Proceedings of the Australian Coral Reef Society 75th Anniversary Conference (eds Greenwood, J. G. & Hall, N. R.) 199–216 (1998).Smith, D. J., Suggett, D. J. & Baker, N. R. Is photoinhibition of zooxanthellae photosynthesis the primary cause of thermal bleaching in corals?. Glob. Chang. Biol. 11, 1–11 (2005).Article 
ADS 

Google Scholar 
Downs, C. et al. Heat-stress and light-stress induce different cellular pathologies in the symbiotic dinoflagellate during coral bleaching. PLoS ONE 8, e77173 (2013).Article 
ADS 
CAS 

Google Scholar 
Banaszak, A. T. & Lesser, M. P. Effects of solar ultraviolet radiation on coral reef organisms. Photochem. Photobiol. Sci. 8, 1276–1294 (2009).Article 
CAS 

Google Scholar 
Jokiel, P. L. & York, R. H. Jr. Solar ultraviolet photobiology of the reef coral Pocillopora damicornis and symbiotic zooxanthellae. Bull. Mar. Sci. 32, 301–315 (1982).
Google Scholar 
Vareschi, E. & Fricke, H. Light responses of a scleractinian coral (Plerogyra sinuosa). Mar. Biol. 90, 395–402 (1986).Article 

Google Scholar 
Henley, E. M. et al. Reproductive plasticity of Hawaiian Montipora corals following thermal stress. Sci. Rep. 11, 12525 (2021).Article 
ADS 
CAS 

Google Scholar 
Wellington, G. & Fitt, W. Influence of UV radiation on the survival of larvae from broadcast-spawning reef corals. Mar. Biol. 143, 1185–1192 (2003).Article 
CAS 

Google Scholar 
Gleason, D. F. & Wellington, G. M. Ultraviolet radiation and coral bleaching. Nature 365, 836–838 (1993).Article 
ADS 

Google Scholar 
Courtial, L., Roberty, S., Shick, J. M., Houlbrèque, F. & Ferrier-Pagès, C. Interactive effects of ultraviolet radiation and thermal stress on two reef-building corals. Limnol. Oceanogr. 62, 1000–1013 (2017).Article 
ADS 

Google Scholar 
Bahr, K. D., Jokiel, P. L. & Rodgers, K. S. The 2014 coral bleaching and freshwater flood events in Kāneʻohe Bay. Hawaiʻi. PeerJ 3, e1136 (2015).Article 

Google Scholar 
Rodgers, K. S., Bahr, K. D., Jokiel, P. L. & Richards Donà, A. Patterns of bleaching and mortality following widespread warming events in 2014 and 2015 at the Hanauma Bay Nature Preserve, Hawai‘i. PeerJ 5, e3355 (2017).Article 

Google Scholar 
Ritson-Williams, R. & Gates, R. D. Coral community resilience to successive years of bleaching in Kāne‘ohe Bay, Hawai‘i. Coral Reefs 39, 757–769 (2020).Article 

Google Scholar 
Krupp, D. A. Sexual reproduction and early development of the solitary coral Fungia scutaria (Anthozoa: Scleractinia). Coral Reefs 2, 159–164 (1983).Article 
ADS 

Google Scholar 
Kramarsky-Winter, E. & Loya, Y. Reproductive strategies of two fungiid corals from the northern Red Sea: Environmental constraints?. Mar. Ecol. Prog. Ser. 174, 175–182 (1998).Article 
ADS 

Google Scholar 
Loya, Y. & Sakai, K. Bidirectional sex change in mushroom stony corals. Proc. R. Soc. B. 275, 2335–2343 (2008).Article 

Google Scholar 
Hagedorn, M. et al. Coral larvae conservation: Physiology and reproduction. Cryobiology 52, 33–47 (2006).Article 
CAS 

Google Scholar 
Jokiel, P. L. & Brown, E. K. Global warming, regional trends and inshore environmental conditions influence coral bleaching in Hawaii. Glob. Chang. Biol. 10, 1627–1641 (2004).Article 
ADS 

Google Scholar 
Tanaka, K., Guidry, M. W. & Gruber, N. Ecosystem responses of the subtropical Kaneohe Bay, Hawaii, to climate change: A nitrogen cycle modeling approach. Aquat. Geochem. 19, 569–590 (2013).Article 
CAS 

Google Scholar 
Couch, C. S. et al. Mass coral bleaching due to unprecedented marine heatwave in Papahānaumokuākea Marine National Monument (Northwestern Hawaiian Islands). PLoS ONE 12, e0185121 (2017).Article 

Google Scholar 
Coles, S. L. et al. Evidence of acclimatization or adaptation in Hawaiian corals to higher ocean temperatures. PeerJ 6, e5347 (2018).Article 

Google Scholar 
Barnhill, K. A. & Bahr, K. D. Coral resilience at Malaukaa fringing reef, Kāneʻohe Bay, Oʻahu after 18 years. J. Mar. Sci. Eng. 7, 311 (2019).Article 

Google Scholar 
Lesser, M., Stochaj, W., Tapley, D. & Shick, J. Bleaching in coral reef anthozoans: Effects of irradiance, ultraviolet radiation, and temperature on the activities of protective enzymes against active oxygen. Coral Reefs 8, 225–232 (1990).Article 
ADS 

Google Scholar 
Brown, B., Dunne, R., Scoffin, T. & Le Tissier, M. Solar damage in intertidal corals. Mar. Ecol. Prog. Ser. 219–230 (1994).Le Tissier, M. D. A. & Brown, B. E. Dynamics of solar bleaching in the intertidal reef coral Goniastrea aspera at Ko Phuket, Thailand. Mar. Ecol. Prog. Ser. 136, 235–244 (1996).Article 
ADS 

Google Scholar 
Lesser, M. P. Elevated temperatures and ultraviolet radiation cause oxidative stress and inhibit photosynthesis in symbiotic dinoflagellates. Limnol. Oceanogr. 41, 271–283 (1996).Article 
ADS 
CAS 

Google Scholar 
Takahashi, S., Nakamura, T., Sakamizu, M., Woesik, R. V. & Yamasaki, H. Repair machinery of symbiotic photosynthesis as the primary target of heat stress for reef-building corals. Plant Cell Physiol. 45, 251–255 (2004).Article 
CAS 

Google Scholar 
Coelho, V. et al. Shading as a mitigation tool for coral bleaching in three common Indo-Pacific species. J. Exp. Mar. Biol. Ecol. 497, 152–163 (2017).Article 

Google Scholar 
Marquis, R. J. Phenological variation in the neotropical understory shrub Piper arielanum: Causes and consequences. Ecology 69, 1552–1565 (1988).Article 

Google Scholar 
Bouwmeester, J. et al. Latitudinal variation in monthly-scale reproductive synchrony among Acropora coral assemblages in the Indo-Pacific. Coral Reefs 40, 1411–1418 (2021).Article 

Google Scholar 
Hagedorn, M. et al. Preserving and using germplasm and dissociated embryonic cells for conserving Caribbean and Pacific coral. PLoS ONE 7, e33354 (2012).Article 
ADS 
CAS 

Google Scholar 
Zuchowicz, N. et al. Assessing coral sperm motility. Sci. Rep. 11, 61 (2021).Article 
CAS 

Google Scholar 
Binet, M., Doyle, C., Williamson, J. & Schlegel, P. Use of JC-1 to assess mitochondrial membrane potential in sea urchin sperm. J. Exp. Mar. Biol. Ecol. 452, 91–100 (2014).Article 
CAS 

Google Scholar 
Jokiel, P., Maragos, J. & Franzisket, L. Coral growth: buoyant weight technique. In Coral Reefs: Research Methods Vol. 5 (eds Stoddart, D. R. & Johannes, R. E.) 529–542 (UNESCO, 1978).
Google Scholar 
R Core Team. R: A Language and Environment for Statistical Computing. https://www.R-project.org (R Foundation for Statistical Computing, 2019).Fox, J. & Weisberg, S. An R Companion to Applied Regression 3rd edn. (Sage Publications, 2019).
Google Scholar 
Wickham, H. ggplot2: Elegant Graphics for Data Analysis (Springer, 2016).Book 
MATH 

Google Scholar 
Kuznetsova, A., Brockhoff, P. B. & Christensen, R. H. B. lmerTest package: Tests in linear mixed effects models. J. Stat. Softw. 82, 1–26 (2017).Article 

Google Scholar 
Lenth, R. V. Least-squares means: The R package lsmeans. J. Stat. Softw. 69, 1–33 (2016).Article 

Google Scholar 
Hothorn, T., Bretz, F. & Westfall, P. Simultaneous inference in general parametric models. Biom. J. J. Math. Methods Biosci. 50, 346–363 (2008).MathSciNet 
MATH 

Google Scholar 
Graves, S., Piepho, H.-P. & Selzer, M. L. multcompView: Visualizations of paired comparisons. R package version 0.1-7. https://CRAN.R-project.org/package=multcompView (2015).Christensen, R. H. B. ordinal-Regression models for ordinal data. R package version 2019.4-25. https://cran.r-project.org/package=ordinal/. (2019).Mangiafico, S. rcompanion: functions to support extension education program evaluation. R package version 2.3.7. https://cran.r-project.org/package=rcompanion (2019).Hope, R. M. Rmisc: Ryan Miscellaneous. R package version 1.5. https://cran.r-project.org/package=Rmisc (2013).Hervé, M. RVAideMemoire: Testing and plotting procedures for biostatistics, R package version 0.9-73. https://cran.r-project.org/package=RVAideMemoire (2019).Callaghan, J. A short note on the intensification and extreme rainfall associated with Hurricane Lane. Trop. Cyclone Res. Rev. 8, 103–107 (2019).Article 

Google Scholar 
Guest, J. R., Baird, A. H., Goh, B. P. L. & Chou, L. M. Seasonal reproduction in equatorial reef corals. Invertebr. Reprod. Dev. 48, 207–218 (2005).Article 

Google Scholar 
Lotterhos, K. E. & Levitan, D. Gamete release and spawning behavior in broadcast spawning marine invertebrates. In The Evolution of Primary Sexual Characters (eds Leonard, J. & Córdoba-Aguilar, A.) 99–120 (Oxford Univ. Press, 2010).
Google Scholar 
Ims, R. A. The ecology and evolution of reproductive synchrony. Trends Ecol. Evol. 5, 135–140 (1990).Article 
CAS 

Google Scholar 
Shlesinger, T. & Loya, Y. Breakdown in spawning synchrony: A silent threat to coral persistence. Science 365, 1002–1007 (2019).Article 
ADS 
CAS 

Google Scholar 
Guest, J. R., Baird, A. H., Bouwmeester, J. & Edwards, A. J. To assess temporal breakdown in spawning synchrony requires comparable temporal data. https://doi.org/10.1126/comment.737627/full/ (2020).Hartmann, D. L. et al. Observations: atmosphere and surface. In Climate change 2013 The Physical Science Basis: Working Group I Contribution to the Fifth Assessment Report of the Intergovernmental Panel on Climate Change (eds Stocker, T. F. et al.) 159–254 (Cambridge University Press, 2013).Pörtner, H. et al. IPCC Special Report on the Ocean and Cryosphere in a Changing Climate (IPCC Intergovernmental Panel on Climate Change, 2019).
Google Scholar 
Cheng, L., Abraham, J., Hausfather, Z. & Trenberth, K. E. How fast are the oceans warming?. Science 363, 128–129 (2019).Article 
ADS 
CAS 

Google Scholar 
Gorbunov, M. Y. & Falkowski, P. G. Photoreceptors in the cnidarian hosts allow symbiotic corals to sense blue moonlight. Limnol. Oceanogr. 47, 309–315 (2002).Article 
ADS 

Google Scholar 
Boch, C. A., Ananthasubramaniam, B., Sweeney, A. M., Doyle Iii, F. J. & Morse, D. E. Effects of light dynamics on coral spawning synchrony. Biol. Bull. 220, 161–173 (2011).Article 

Google Scholar 
Sweeney, A. M., Boch, C. A., Johnsen, S. & Morse, D. E. Twilight spectral dynamics and the coral reef invertebrate spawning response. J. Exp. Biol. 214, 770–777 (2011).Article 

Google Scholar 
Nozawa, Y. Annual variation in the timing of coral spawning in a high-latitude environment: Influence of temperature. Biol. Bull. 222, 192–202 (2012).Article 

Google Scholar 
Babcock, R. C. et al. Synchronous spawnings of 105 scleractinian coral species on the Great Barrier Reef. Mar. Biol. 90, 379–394 (1986).Article 

Google Scholar 
Hunter, C. Environmental cues controlling spawning in two Hawaiian corals, Montipora verrucosa and M. dilatata. In Proc 6th Int Coral Reef Symp. vol. 2, 727–732.Levitan, D. R. et al. Mechanisms of reproductive isolation among sympatric broadcast spawning corals of the Montastraea annularis species complex. Evolution 58, 308–323 (2004).
Google Scholar 
Negri, A. P., Marshall, P. A. & Heyward, A. J. Differing effects of thermal stress on coral fertilization and early embryogenesis in four Indo Pacific species. Coral Reefs 26, 759–763 (2007).Article 
ADS 

Google Scholar 
Humanes, A., Noonan, S. H., Willis, B. L., Fabricius, K. E. & Negri, A. P. Cumulative effects of nutrient enrichment and elevated temperature compromise the early life history stages of the coral Acropora tenuis. PLoS ONE 11, e0161616 (2016).Article 

Google Scholar 
Lesser, M. P., Kruse, V. A. & Barry, T. M. Exposure to ultraviolet radiation causes apoptosis in developing sea urchin embryos. J. Exp. Biol. 206, 4097–4103 (2003).Article 

Google Scholar 
Häder, D.-P. et al. Effects of UV radiation on aquatic ecosystems and interactions with other environmental factors. Photochem. Photobiol. Sci. 14, 108–126 (2015).Article 

Google Scholar 
Albright, R. & Mason, B. Projected near-future levels of temperature and pCO2 reduce coral fertilization success. PLoS ONE 8, e56468 (2013).Article 
ADS 
CAS 

Google Scholar 
Espinoza, J., Schulz, M., Sanchez, R. & Villegas, J. Integrity of mitochondrial membrane potential reflects human sperm quality. Andrologia 41, 51–54 (2009).Article 
CAS 

Google Scholar 
Paoli, D. et al. Mitochondrial membrane potential profile and its correlation with increasing sperm motility. Fertil. Steril. 95, 2315–2319 (2011).Article 
CAS 

Google Scholar 
Gallo, A., Esposito, M. C., Tosti, E. & Boni, R. Sperm motility, oxidative status, and mitochondrial activity: Exploring correlation in different species. Antioxidants 10, 1131 (2021).Article 
CAS 

Google Scholar 
Schlegel, P., Binet, M. T., Havenhand, J. N., Doyle, C. J. & Williamson, J. E. Ocean acidification impacts on sperm mitochondrial membrane potential bring sperm swimming behaviour near its tipping point. J. Exp. Biol. 218, 1084–1090 (2015).Article 

Google Scholar 
Gulko, D. Effects of ultraviolet radiation on fertilization and production of planula larvae in the Hawaiian coral Fungia scutaria. In Ultraviolet Radiation and Coral Reefs Vol. 41 (eds Gulko, D. & Jokiel, P. L.) 135–147 (University of Hawai’i, 1995).
Google Scholar 
Pruski, A. M., Nahon, S., Escande, M.-L. & Charles, F. Ultraviolet radiation induces structural and chromatin damage in Mediterranean sea-urchin spermatozoa. Mutat. Res. Genet. Toxicol. Environ. Mutagen. 673, 67–73 (2009).Article 
CAS 

Google Scholar 
Dahms, H.-U. & Lee, J.-S. UV radiation in marine ectotherms: Molecular effects and responses. Aquat. Toxicol. 97, 3–14 (2010).Article 
CAS 

Google Scholar 
Nesa, B., Baird, A. H., Harii, S., Yakovleva, I. & Hidaka, M. Algal symbionts increase DNA damage in coral planulae exposed to sunlight. Zool. Stud. 51, 12–17 (2012).CAS 

Google Scholar 
Paxton, C. W., Baria, M. V. B., Weis, V. M. & Harii, S. Effect of elevated temperature on fecundity and reproductive timing in the coral Acropora digitifera. Zygote 24, 511 (2015).Article 

Google Scholar 
Jokiel, P. & Coles, S. Effects of temperature on the mortality and growth of Hawaiian reef corals. Mar. Biol. 43, 201–208 (1977).Article 

Google Scholar 
Cantin, N. E., Cohen, A. L., Karnauskas, K. B., Tarrant, A. M. & McCorkle, D. C. Ocean warming slows coral growth in the Central Red Sea. Science 329, 322–325. https://doi.org/10.1126/science.1190182 (2010).Article 
ADS 
CAS 

Google Scholar 
Cooper, T. F., De’Ath, G., Fabricius, K. E. & Lough, J. M. Declining coral calcification in massive Porites in two nearshore regions of the northern Great Barrier Reef. Glob. Chang. Biol. 14, 529–538 (2008).Article 
ADS 

Google Scholar 
Tanzil, J., Brown, B., Tudhope, A. & Dunne, R. Decline in skeletal growth of the coral Porites lutea from the Andaman Sea, South Thailand between 1984 and 2005. Coral Reefs 28, 519–528 (2009).Article 
ADS 

Google Scholar 
Tanzil, J. T. I. et al. Regional decline in growth rates of massive Porites corals in Southeast Asia. Glob. Chang. Biol. 19, 3011–3023 (2013).Article 
ADS 

Google Scholar 
Richmond, R. H., Tisthammer, K. H. & Spies, N. P. The effects of anthropogenic stressors on reproduction and recruitment of corals and reef organisms. Front. Mar. Sci. 5, 226 (2018).Article 

Google Scholar 
Chen, P.-Y., Chen, C.-C., Chu, L. & McCarl, B. Evaluating the economic damage of climate change on global coral reefs. Glob. Environ. Change 30, 12–20 (2015).Article 

Google Scholar 
Kaniewska, P., Alon, S., Karako-Lampert, S., Hoegh-Guldberg, O. & Levy, O. Signaling cascades and the importance of moonlight in coral broadcast mass spawning. Elife 4, e09991 (2015).Article 

Google Scholar 
Lin, C.-H., Takahashi, S., Mulla, A. J. & Nozawa, Y. Moonrise timing is key for synchronized spawning in coral Dipsastraea speciosa. Proc. Natl. Acad. Sci. 118, e2101985118 (2021).Article 
CAS 

Google Scholar 
Anthony, K. R. et al. Interventions to help coral reefs under global change—A complex decision challenge. PLoS ONE 15, e0236399 (2020).Article 
CAS 

Google Scholar 
Daly, J. et al. Cryopreservation can assist gene flow on the Great Barrier Reef. Coral Reefs 41, 455–462 (2022).Article 

Google Scholar  More

Berner, L. T. et al. Summer warming explains widespread but not uniform greening in the Arctic tundra biome. Nat. Commun. 11, 4621 (2020).Article 
ADS 
CAS 

Google Scholar 
Elmendorf, S. C. et al. Plot-scale evidence of tundra vegetation change and links to recent summer warming. Nat. Clim. Change 2, 453–457 (2012).Article 
ADS 

Google Scholar 
Overland, J. E. et al. Surface air temperature. In Arctic Report Card: Update for 2019 (eds Richter-Menge, J. et al.) (U.S. National Park Service, 2020).
Google Scholar 
Post, E., Steinman, B. A. & Mann, M. E. Acceleration of phenological advance and warming with latitude over the past century. Sci. Rep. 8, 3927 (2018).Article 
ADS 

Google Scholar 
Diepstraten, R. A. E., Jessen, T. D., Fauvelle, C. M. D. & Musiani, M. M. Does climate change and plant phenology research neglect the Arctic tundra?. Ecosphere 9, e02362 (2018).Article 

Google Scholar 
Flynn, D. F. B. & Wolkovich, E. M. Temperature and photoperiod drive spring phenology across all species in a temperate forest community. New Phytol. 219, 1353–1362 (2018).Article 
CAS 

Google Scholar 
Billings, W. D. & Bliss, L. C. An alpine snowbank environment and its effects on vegetation, plant development, and productivity. Ecology 40, 388–397 (1959).Article 

Google Scholar 
Billings, W. D. & Mooney, H. A. The ecology of arctic and alpine plants. Biol. Rev. 43, 481–529 (1968).Article 

Google Scholar 
Sørensen, T. Temperature relations and phenology of the northeast Greenland flowering plants. Meddr Gronland 1–305 (1941).Barrett, R. T. & Hollister, R. D. Arctic plants are capable of sustained responses to long-term warming. Polar Res. 35, 25405 (2016).Article 

Google Scholar 
Julitta, T. et al. Using digital camera images to analyse snowmelt and phenology of a subalpine grassland. Agric. For. Meteorol. 198–199, 116–125 (2014).Article 
ADS 

Google Scholar 
Petraglia, A. et al. Responses of flowering phenology of snowbed plants to an experimentally imposed extreme advanced snowmelt. Plant Ecol. 215, 759–768 (2014).Article 

Google Scholar 
Semenchuk, P. R. et al. High Arctic plant phenology is determined by snowmelt patterns but duration of phenological periods is fixed: An example of periodicity. Environ. Res. Lett. 11, 125006 (2016).Article 
ADS 

Google Scholar 
Hollister, R. D., Webber, P. J. & Bay, C. Plant response to temperature in northern Alaska: Implications for predicting vegetation change. Ecology 86, 1562–1570 (2005).Article 

Google Scholar 
Oberbauer, S. et al. Phenological response of tundra plants to background climate variation tested using the International Tundra Experiment. Philos. Trans. R. Soc. B Biol. Sci. 368, 20120481 (2013).Article 
CAS 

Google Scholar 
Tieszen, L. L. Photosynthesis in the principal Barrow, Alaska, species: A summary of field and laboratory responses. In Vegetation and Production Ecology of an Alaskan Arctic Tundra (ed. Tieszen, L. L.) 241–268 (Springer, 1978).Chapter 

Google Scholar 
Körner, Ch. CO2 exchange in the alpine sedge Carex curvula as influenced by canopy structure, light and temperature. Oecologia 53, 98–104 (1982).Article 
ADS 

Google Scholar 
Tieszen, L. L. Photosynthesis and respiration in arctic tundra grasses: Field light intensity and temperature responses. Arct. Alp. Res. 5, 239–251 (1973).Article 
CAS 

Google Scholar 
Huang, M. et al. Air temperature optima of vegetation productivity across global biomes. Nat. Ecol. Evol. 3, 772–779 (2019).Article 

Google Scholar 
Marchand, F. L., Mertens, S., Kockelbergh, F., Beyens, L. & Nijs, I. Performance of high arctic tundra plants improved during but deteriorated after exposure to a simulated extreme temperature event. Glob. Change Biol. 11, 2078–2089 (2005).Article 
ADS 

Google Scholar 
Yan, W. An equation for modelling the temperature response of plants using only the cardinal temperatures. Ann. Bot. 84, 607–614 (1999).Article 

Google Scholar 
Zhou, G. & Wang, Q. A new nonlinear method for calculating growing degree days. Sci. Rep. 8, 10149 (2018).Article 
ADS 

Google Scholar 
Kramer, K. Selecting a model to predict the onset of growth of Fagus sylvatica. J. Appl. Ecol. 31, 172 (1994).Article 

Google Scholar 
Nakano, Y., Higuchi, Y., Sumitomo, K. & Hisamatsu, T. Flowering retardation by high temperature in chrysanthemums: Involvement of FLOWERING LOCUS T-like 3 gene repression. J. Exp. Bot. 64, 909–920 (2013).Article 
CAS 

Google Scholar 
del Olmo, I., Poza-Viejo, L., Piñeiro, M., Jarillo, J. A. & Crevillén, P. High ambient temperature leads to reduced FT expression and delayed flowering in Brassica rapa via a mechanism associated with H2A.Z dynamics. Plant J. 100, 343–356 (2019).Article 

Google Scholar 
Wolkovich, E. M. et al. Warming experiments underpredict plant phenological responses to climate change. Nature 485, 494 (2012).Article 
ADS 
CAS 

Google Scholar 
Hollister, R. D. et al. A review of open top chamber (OTC) performance across the ITEX Network. Arct. Sci. https://doi.org/10.1139/AS-2022-0030 (2022).Article 

Google Scholar 
Bütikofer, L. et al. The problem of scale in predicting biological responses to climate. Glob. Change Biol. 26, 6657–6666 (2020).Article 
ADS 

Google Scholar 
Gu, S. Growing degree hours—A simple, accurate, and precise protocol to approximate growing heat summation for grapevines. Int. J. Biometeorol. 60, 1123–1134 (2016).Article 
ADS 
CAS 

Google Scholar 
Roltsch, W. J., Zalom, F. G., Strawn, A. J., Strand, J. F. & Pitcairn, M. J. Evaluation of several degree-day estimation methods in California climates. Int. J. Biometeorol. 42, 169–176 (1999).Article 
ADS 

Google Scholar 
Richardson, A. D. et al. Ecosystem warming extends vegetation activity but heightens vulnerability to cold temperatures. Nature 560, 368–371 (2018).Article 
ADS 
CAS 

Google Scholar 
Ettinger, A. K., Buonaiuto, D. M., Chamberlain, C. J., Morales-Castilla, I. & Wolkovich, E. M. Spatial and temporal shifts in photoperiod with climate change. New Phytol. 230, 462–474 (2021).Article 
CAS 

Google Scholar 
Seyednasrollah, B., Swenson, J. J., Domec, J.-C. & Clark, J. S. Leaf phenology paradox: Why warming matters most where it is already warm. Remote Sens. Environ. 209, 446–455 (2018).Article 
ADS 

Google Scholar 
Breshears, D. D. et al. Underappreciated plant vulnerabilities to heat waves. New Phytol. 231, 32–39 (2021).Article 

Google Scholar 
Chaudhry, S. & Sidhu, G. P. S. Climate change regulated abiotic stress mechanisms in plants: A comprehensive review. Plant Cell Rep. 41, 1–31 (2022).Article 
CAS 

Google Scholar 
Sun, X. et al. Global diurnal temperature range (DTR) changes since 1901. Clim. Dyn. 52, 3343–3356 (2019).Article 

Google Scholar 
Ballinger, T. J. NOAA Arctic Report Card 2021: Surface Air Temperature. https://doi.org/10.25923/53XD-9K68 (2021).Jagadish, S. V. K., Way, D. A. & Sharkey, T. D. Plant heat stress: Concepts directing future research. Plant Cell Environ. 44, 1992–2005 (2021).Article 
CAS 

Google Scholar 
Gilmore, E. C. Jr. & Rogers, J. S. Heat units as a method of measuring maturity in corn. Agron. J. 50, 611–615 (1958).Article 

Google Scholar 
Sánchez, B., Rasmussen, A. & Porter, J. R. Temperatures and the growth and development of maize and rice: A review. Glob. Change Biol. 20, 408–417 (2014).Article 
ADS 

Google Scholar 
Molitor, D., Junk, J., Evers, D., Hoffmann, L. & Beyer, M. A high-resolution cumulative degree day-based model to simulate phenological development of grapevine. Am. J. Enol. Vitic. 65, 72–80 (2014).Article 

Google Scholar 
CaraDonna, P. J., Iler, A. M. & Inouye, D. W. Shifts in flowering phenology reshape a subalpine plant community. Proc. Natl. Acad. Sci. 111, 4916–4921 (2014).Article 
ADS 
CAS 

Google Scholar 
Inouye, B. D., Ehrlén, J. & Underwood, N. Phenology as a process rather than an event: From individual reaction norms to community metrics. Ecol. Monogr. 89, e01352 (2019).Article 

Google Scholar 
Miles, W. T. S. et al. Quantifying full phenological event distributions reveals simultaneous advances, temporal stability and delays in spring and autumn migration timing in long-distance migratory birds. Glob. Change Biol. 23, 1400–1414 (2017).Article 
ADS 

Google Scholar 
Moussus, J.-P., Julliard, R. & Jiguet, F. Featuring 10 phenological estimators using simulated data. Methods Ecol. Evol. 1, 140–150 (2010).Article 

Google Scholar 
Dowle, M. & Srinivasan, A. data.table: Extension of ‘data.frame’ (2019).Auguie, B. egg: Extensions for ‘ggplot2’: Custom Geom, Custom Themes, Plot Alignment, Labelled Panels, Symmetric Scales, and Fixed Panel Size (2019).Wood, S. & Scheipl, F. gamm4: Generalized Additive Mixed Models using ‘mgcv’ and ‘lme4’ (2020).Auguie, B. gridExtra: Miscellaneous Functions for ‘Grid’ Graphics (2017).Hamner, B. & Frasco, M. Metrics: Evaluation Metrics for Machine Learning (2018).Gilli, M., Maringer, D. & Schumann, E. Numerical Methods and Optimization in Finance (Elsevier/Academic Press, 2019).MATH 

Google Scholar 
Garnier, S. viridis: Default Color Maps from ‘matplotlib’ (2018).Wickham, H. et al. Welcome to the Tidyverse. J. Open Source Softw. 4, 1686 (2019).Article 
ADS 

Google Scholar  More

Systematic paleontologySauropterygia Owen, 186038.Eosauropterygia Rieppel, 199439.Pachypleurosauroidea Huene, 195640.Pachypleurosauridae Nopcsa, 192841.Luopingosaurus imparilis gen. et sp. nov.EtymologyThe genus name refers to the Luoping County, at which the fossil site is located. Species epithet imparilis (Latin) means peculiar and unusual.HolotypeA ventrally exposed skeleton with a posterior part of the caudal missing, IVPP V19049.Locality and horizonLuoping, Yunnan, China; Second (Upper) Member of Guanling Formation, Pelsonian (~ 244 Ma), Anisian, Middle Triassic37.DiagnosisA pachypleurosaurid distinguishable from other members of this family by the following combination of features (those unique among pachypleurosaurids identified with an asterisk): snout (preorbital portion) long and anteriorly pointed, 55.0% of skull length (*); orbital length about one quarter of skull length; internal naris retracted, without contribution from premaxilla; nasal ending at level of anterior margin of prefrontal; dentary length 71.7% of mandibular length; hyoid length 9.7% of mandibular length; presence of entepicondylar foramen in humerus; 21 cervical and 27 dorsal vertebrae (*); distinct expansions of distal heads of posterior two sacral ribs; six pairs of caudal ribs; phalangeal formula 2–3-5–5-3 for manus and 2–3-4–6-4 for pes (*); Metatarsal I short and stout with expanded proximal end, 56.4% of Metatarsal V in length (*); and Metatarsal IV being longest phalange in pes.Comparative descriptionThe holotype and only currently known specimen of Luopingosaurus has a preserved length of 46.2 cm from the rostral tip to the 30th caudal vertebra (for measurements, see Table 1). The estimated total length of the body may have reached 64 cm, assuming similar tail proportions of pachypleurosaurids. As such, Luopingosaurus is longer than most of other pachypleurosauroids that are small-sized with a maximum total length rarely exceeding 50 cm4,9,10,11,12,14,15,16,18,23,25, although some pachypleurosaurids are notably larger (e.g., 88 cm in Diandongosaurus cf. acutidentatus22, ~ 120 cm in Neusticosaurus edwardsii8, and ~ 130 cm in Wumengosaurus delicatomandibularis13).Table 1 Measurements (in mm) of the holotype (IVPP V19049) of Luopingosaurus imparilis gen. et sp. nov. R, right.Full size tableThe pre-orbital portion, distinctly longer than the postorbital region, measures 55% of the total skull length (the premaxillary symphysis to the occipital condyle) and 51% of the mandibular length. The paired premaxillae form most of the snout anterior to the naris with a pointed anterior tip, similar to the conditions in Wumengosaurus13,30 and Honghesaurus23. By contrast, other pachypleurosauroids uniformly have a blunt rostrum4,6,7,8,9,10,11,12,14,15,16,18,22,25. The premaxilla bears a long posteromedial process inserting between the anterior parts of the elongate nasals (Fig. 3). The premaxillary teeth are homodont with a tall peduncle and a short, conical crown, but the tooth number is hard to estimate because of occlusion of jaws. The posterior parts of nasals contact each other medially, and posteriorly, they contact the frontals in an interdigitating suture at the level of the anterior margin of the prefrontal. In Honghesaurus23, Wumengosaurus30, Neusticosaurus8 and Serpianosaurus9, the even longer nasal extends posteriorly beyond this level and ends at the anterior portion of the orbit.Figure 3Skull and mandible of Luopingosaurus imparilis gen. et sp. nov., IVPP V19049. Head before (a) and after (b) dusted with ammonium chloride. (c), Line- drawing. (d, e), two computed laminography scanning slices. (f), reconstruction in ventral view. ac, acetabulum; an, angular; ar, articular; ax, axis; c, cervical vertebra; den, dentary; eo, exoccipital; f, frontal; hy, hyoid; in, internal naris; j, jugal; m, maxilla; n, nasal; p, parietal; par, prearticular; pof, postfrontal; prf, prefrontal; pt, pterygoid; q, quadrate; qj, quadratojugal; sa, surangular; sp, splenial; sq, squamosal; stf, supratemporal fossa; v, vomer.Full size imageThe orbit is oval and large, measuring 24.8% of the skull length (Fig. 3). The lateral margin of the frontal contacts the prefrontal anteriorly and the postfrontal posteriorly, and defines most of the medial border of the orbit. The L-shaped jugal, together with the posterolateral process of the maxilla, forms the lateral border of the orbit. No distinct lacrimal is discernable; the bone is probably absent as in other sauropterygians. The postfrontal contacts the dorsal process of the triradiate postorbital ventrally, and both bones define the posterior border of the orbit. Additionally, the posterior process of the postorbital contacts the anterior process of the squamosal, forming the bar between the supratemporal fossa and the ventrally open infratemporal fenestra. The jugal extends beyond the ventral margin of the postorbital and also contacts the anterior process of the squamosal, resembling the conditions in Wumengosaurus30, Honghesaurus23 and Diandongosaurus15. This contact is absent in other pachypleurosauroids4,6,7,8,9,10,11,12.A pair of vomers and pterygoids and a right palatine are discernable in the palate (Fig. 3a–c). The vomer is elongate and slender, extending anteriorly well beyond the nasal. The internal naris, partly covered by the detached splenial, is longitudinally retracted, corresponding to a retracted external naris (Fig. 3d–f). The medial margin of the naris is defined by the nasal, without contribution from the premaxilla. A retracted naris is otherwise present in Wumengosaurus13,30, Qianxisaurus16 and Honghesaurus23. Similar to the condition in Honghesaurus23, the retracted naris of Luopingosaurus is relatively short, having a longitudinal diameter distinctly less than half of the longitudinal diameter of the orbit. By contrast, other pachypleurosauroids4,6,7,8,9,10,11,12,25 generally have an oval-shaped naris. The elongate palatine has a slightly convex medial margin suturing with the pterygoid. Because of the coverage of the detached splenial, the lateral portion of the palatine is unexposed, and it is hard to know whether an ectopterygoid is present or not. The pterygoid is the largest and longest element of the palate, measuring 55.2% of the mandibular length. It has an anterior projection that contacts the vomer anteromedially, and does not participate in the margin of the internal naris. At the level of the posterior orbital margin, the pterygoid has a triangular lateral extension, which was termed as the ectopterygoid process of the pterygoid in Neusticosaurus8. The pterygoid extends back to the occipital condyle, and covers the basicranium and parietals in ventral view. Additionally, the bone has a broad posterolateral process that is set off from the palatal surface by a distinct ridge, resembling the conditions in Serpianosaurus9 and Neusticosaurus8. Posteriorly, the basioccipital is exposed in ventral view, showing the area for attachment to the right exoccipital.The left quadrate is exposed in lateral view with its dorsal process extending underneath the base of the descending process of the squamosal. The posterior margin of the quadrate is excavated, as in many other pachypleurosaurids (e.g., Serpianosaurus9 and Honghesaurus23). The quadratojugal is narrow and splint-like, flanking the anterior margin of the quadrate. A pair of hyoids are ossified. They are rod-like, slightly expanded at both ends. The dentary is wedge-shaped, being 71.7% of the mandibular length. Laterally, it bears a longitudinal series of pores and grooves parallel to the oral margin of the bone (Fig. 3a). The elongate angular tapers at both ends, contacting the dentary anterodorsally and the surangular dorsally in ventral view. The surangular, slightly shorter than the angular, contacts the articular posterodorsally, with a pointed anterior tip wedging into the notched posterior margin of the dentary. The retroarticular process of the articular is very short with a rounded posterior margin. Medially, the splenial and prearticular form most of the inner wall of the mandible. The splenial tapers at both ends and enters the mandibular symphysis anteriorly, having a length similar to the dentary. The relatively slender prearticular contacts the splenial anterodorsally, extends posteriorly and abuts the articular dorsally, measuring 41.1% of the mandibular length.The whole series of 21 cervical vertebrae (including the atlas-axis complex) is well exposed ventrally. The atlas centrum is oval, much smaller than the axis centrum (Fig. 3c). From the axis, the cervical vertebrae increase gradually in size toward the trunk vertebrae posteriorly. The bicephalous cervical ribs have typical free anterior and posterior processes as in other pachypleurosauroids8,9. The trunk is relatively long, including 27 dorsal vertebrae. The posterior dorsal ribs show certain pachyostosis (Fig. S1). Each gastralium consists of five elements (a short and more massive median element and two slender rods in line towards each side; Figs. 3, 4a, b, S1), similar to the conditions in most of other pachypleurosauroids9,11,18,25 (except Neusticosaurus8). Three sacral ribs are clearly revealed by X-ray computed microtomography (Fig. 4c–f). They are relatively short and stout, with the posterior twos bearing a distinct expansion on their distal heads. The distal expansion of the sacral rib is also present in Keichousaurus11, Prosantosaurus25, Qianxisaurus16 and Wumengosaurus13, but it is not pronounced in other pachypleurosauroids4,6,7,8,9,10. The caudal ribs are relatively few, six pairs in number. Additionally, several chevron bones are visible in the proximal caudal region, and they are gradually reduced in length posteriorly (Fig. 4d).Figure 4Girdles, limbs and vertebrae of Luopingosaurus imparilis gen. et sp. nov., IVPP V19049. Photo (a) and line-drawing (b) of pectoral girdle, forelimbs and anterior dorsal vertebrae. Photo (c), line-drawing (d) and two computed laminography scanning slices (e, f) of pelvic girdle, hind limbs and posterior vertebrae. as, astragalus; ca, caudal vertebra; cal, calcaneum; car, caudal rib; co, coracoid; d, dorsal vertebra; dltp, deltopectoral crest; enf, entepicondylar foramen; fe, femur; fi, fibula; h, humerus; il, ilium; int, intermedium; is, ischium; mc, metacarpal; mt, metatarsal; pu, pubis; s, sacral vertebra; sc, scapula; sr, sacral rib; ti, tibia; ul, ulna; uln, ulnare.Full size imageThe paired clavicles and the median interclavicle form a transverse bar at the 20th cervical vertebrae (Fig. 4a, b). The blade-like clavicle tapers posterolaterally with its distal projection overlapped by the scapula in ventral view. The left clavicle contacts the right one anterodorsally to the interclavicle. The interclavicle tapers laterally to a point at each end. The anterior margin of the interclavicle is convex and its posterior margin is slightly concave without a midline projection (contra the condition in Anarosaurus42). The scapula consists of a broad ventral portion and a relatively narrow and elongate dorsal wing that varies little through its length. The coracoid is hourglass-shaped with a slightly concave posterolateral margin and a conspicuously concave anteromedial margin. The medial margin is straight, along which the coracoids would articulate each other in the midline. The humerus is constricted at the middle portion with a nearly straight preaxial margin and a concave postaxial margin. A slit in the expanded distal portion of this bone indicates the possible presence of an entepicondylar foramen (Fig. 4a, b). The radius, slightly longer than the ulna, is more expanded proximally than distally. The ulna is straight with a slightly constricted shaft. In each forelimb, there is two nearly rounded carpals, ulnare and intermedium; the former is half the width of the latter. Five metacarpals are rod-like, slightly expanded at both ends. Among them, Metacarpal I is the shortest, 48% of the length of Metacarpal II. Metacarpal III is slightly shorter than Metacarpal IV, which is the longest. Metacarpal V is 71% of the length of Metacarpal IV. The phalangeal formula is 2–3–5–5–3 for the manus, indicating presence of hyperphalangy in Luopingosaurus (see Discussion below).In the pelvic girdle, the ilia, pubes and ischia are well exposed (Fig. 4c–f). The ilium is nearly triangular with a relatively long and tapering posterior process. The plate-like pubis is well constricted at its middle portion, with the medial portion nearly equal to the lateral portion. The obturator foramen is slit-like, located at the posterolateral corner of this bone (Fig. 4e). The ischium is also plate-like, having a relatively narrow lateral portion and an expanded medial portion that is notably longer than the medial portion of the pubis. The posterolateral ischial margin is highly concave. The posterior pubic margin and anterior ischial margin are moderately concave, and both together would enclose the thyroid fenestra. The femur is slightly longer than the humerus, with a constricted shaft and equally expanded ends (Fig. 4d). No internal trochanter is developed. The tibia is nearly equal to the fibula in length; the former is straight and thicker than the slightly curved latter. Two ossified tarsals, calcaneum and astragalus, are nearly rounded; the latter is significantly larger than the former. As in Honghesaurus23, the astragalus lacks a proximal concavity. The right metatarsals are well-preserved. Metatarsal I is the shortest and stoutest phalange with an expanded proximal end, and Metatarsal IV is the longest. Metatarsal II is nearly twice the length of Metatarsal I. Metatarsal III is slightly shorter than Metatarsal IV, and Metatarsal V is 76% of the length of Metatarsal IV. The phalangeal count is 2–3–4–6–4, which is complete judging from the appearance of the distal phalanges in the right pes (Fig. 4c). More

Naturalist Alfred Russel Wallace went on an expedition to Amazonas state in Brazil in 1848–52.Credit: Mondadori Portfolio via Getty

Best known for formulating the theory of evolution by natural selection, independently of Charles Darwin, Alfred Russel Wallace is an appealing if enigmatic figure. The appeal stems in part from his underdog status: poor and self-educated, Wallace had none of Darwin’s social and financial advantages. The enigma comes from his keen embrace of a range of eccentric non-scientific causes, including spiritualism, phrenology and anti-vaccination (for smallpox).Scientists do not like their scientific heroes to bear the taint of irrational thinking. Wallace’s enthusiasms have therefore contributed to him becoming marginalized in the history of evolutionary thought. Most people know about Darwin and the HMS Beagle. But what about Wallace and the Helen?The Helen story is worth revisiting because it shows Wallace at his resolute best. Despite numerous disastrous career setbacks — of which the Helen episode was the most severe — he persevered and eventually succeeded as a scientist.More than 150 years after Wallace’s experience on the Helen, doing science continues to be hard and can be disappointing. Wallace’s misadventure provides both perspective and an object lesson in how to navigate setbacks. His response to problems showcases his most inspiring traits: his commitment to science, his almost superhuman resilience and his refusal to mire himself in self-pity.Tropical explorationsIn his first job as a land surveyor, Wallace developed an interest in the plants he encountered as he tramped across the countryside. Then, in 1844 at the age of 21, he met Henry Walter Bates, who would later discover ‘Batesian mimicry’ (whereby members of a palatable prey species gain protection by mimicking an unpalatable one).Bates, two years Wallace’s junior, had a fixation with beetles, and he catalysed Wallace’s transformation from hobbyist naturalist to serious collector. Wallace’s new-found focus on beetles transcended mere entomological stamp-collecting; he developed an interest in some of the great scientific questions of the time. He was particularly inspired by the anonymously published Vestiges of the Natural History of Creation (1844) by Robert Chambers, which put forward a vision of a transmutational process, with life progressing from simple to complex.Without money or connections, Wallace and Bates aspired to careers in science at a time when the field was the preserve of the moneyed elite. They would have to fund their scientific explorations by collecting and selling specimens. After a hasty choice of destination — tropical South America — and a crash course in collecting methods, Wallace, aged 25, and Bates, aged 23, arrived in Belém, Brazil, in May 1848 (see ‘Doggedly determined’).
Doggedly determined

Alfred Russel Wallace tends to be unjustly relegated to a footnote in the Charles Darwin story. He was, in fact, a pioneering biologist who refused to let disadvantage or disaster prevent him from pursuing his scientific dreams.
January 1823: Alfred Russel Wallace is born in Usk in Wales.
May 1848: Wallace and Henry Walter Bates arrive in Belém, Brazil.
July 1852: Wallace boards the Helen, which catches fire three weeks later while at sea.
October 1852: Wallace reaches Deal, England, aboard the Jordeson.
March 1854: Wallace leaves Southampton for southeast Asia.
September 1855: Wallace’s first evolutionary paper describing his ‘Sarawak Law’ is published.
May 1856: Citing the Sarawak Law paper, geologist Charles Lyell alerts Darwin to the possibility that Wallace is developing ideas similar to Darwin’s.
February 1858: Wallace sends his paper on natural selection to Darwin from Ternate in the Maluku islands (Moluccas), Indonesia.
July 1858: The joint Darwin–Wallace paper is presented at the Linnean Society in London.
November 1859: Darwin’s On the Origin of Species is published.
March 1862: Wallace returns from southeast Asia.
November 1913: Wallace dies in Broadstone, England.

The two split up early on, with Wallace concentrating on the Amazon River’s northern tributary, the Rio Negro, and Bates on the southern fork, the Solimões.Collecting was challenging. The Amazon’s ubiquitous ants often deprived science of hard-won specimens. Crucial collecting materials also disappeared: Wallace once recovered from a bout of fever to discover that local people had drunk the cachaça (a Brazilian rum) he’d been using to pickle specimens. Transport was a constant headache, with travel upstream past rapids requiring unwieldy portages of canoes and cargo. And thanks to his collecting, the cargo became ever more voluminous and unwieldy.Wallace and Bates sporadically sent back shipments of material to their agent in London, Samuel Stevens, who publicized their adventures in scientific journals and sold their specimens, taking a 20% commission.
Escaping Darwin’s shadow: how Alfred Russel Wallace inspires Indigenous researchers
Wallace’s journeys on the Rio Negro and its tributaries took him into areas that had not yet been visited by Europeans. He saw (and collected) an extraordinary array of species, many of them new to science. He had a chance to observe and collect artefacts from several Indigenous groups with little or no previous contact with Europeans. As he travelled, Wallace capitalized on his surveying skills to map the terrain. But the remoteness took its toll. He made an “inward vow never to travel again in such wild, unpeopled districts without some civilised companion or attendant”1.Wallace was frequently ill, on one occasion nearly lethally so. His younger brother came out to join him as an assistant in 1849 but died of yellow fever two years later in Belém, on his way back to England. Wallace learnt that his brother was sick but had to wait many anxious months before news of his death made it upriver.In 1852, after four years of exploring and collecting, it was time for Wallace himself to head home. He envisaged a triumphant return. He would complement his collections of preserved organisms with a menagerie of living ones. Mr Wallace’s biological wonders would surely be the toast of scientific London.On 12 July in Belém, Wallace boarded the Helen, a freighter ship bound for London. The trip across the continent to Belém had not gone smoothly. The authorities in Manaus, Brazil, had had to be persuaded to release some of his earlier shipments meant for London, which they had impounded, making the final haul aboard the Helen even larger. But now all that remained was the long voyage back across the Atlantic. Wallace, who shared Captain Turner’s cabin, was the only passenger.Disaster strikesThree weeks into the voyage, Captain Turner interrupted Wallace’s morning routine to tell him that the ship was on fire.Friction caused by the rocking of the ship had ignited poorly stowed cargo. Attempts to intervene were counterproductive — removing the hold covers merely oxygenated the fire — and soon the ship became what Wallace later called “a most magnificent conflagration”1.Captain Turner gave the order to abandon ship, and the scramble to prepare two small wooden boats began. Having been stored on deck in the tropical sunshine, both boats leaked badly. The cook had to find corks to plug their hulls.Before he left the ship, Wallace “went down into the cabin, now suffocatingly hot and full of smoke, to see what was worth saving”1. He retrieved his “watch and a small tin box containing some shirts and a couple of old note-books, with some drawings of plants and animals, and scrambled up with them on deck”1. He tried to lower himself on a rope into one of the small boats, but fever-weakened, he ended up sliding down the rope, stripping the skin off his hands.

Some of Alfred Russel Wallace’s sketches were salvaged from the fire aboard the Helen on his return journey from South America in 1852.Credit: The Natural History Museum/Alamy

With fine weather, the best hope of rescue lay in other ships seeing the fire. The two boats duly circled the burning wreck for the next 24 hours, meaning that Wallace got to witness every moment of the tragedy. The animals he had brought with him on the long river journey across the continent, now free from their cages, sought refuge on the one part of the ship still untouched by the flames, the bowsprit. Wallace watched as the monkeys, parrots and more — his pets as well as his best hope of impressing London’s scientific elite — were incinerated.The hoped-for rescue did not immediately materialize, and Captain Turner turned the two open boats towards Bermuda, 1,100 kilometres away to the northwest.As the days ticked by, the situation became increasingly desperate. Water ran low and the tropical sun left Wallace’s “hands and face very much blistered”1. Wallace nevertheless remained upbeat, later recalling that during one night, he “saw several meteors, and in fact could not be in a better position for observing them, than lying on [his] back in a small boat in the middle of the Atlantic”1.Finally, ten days into the ordeal, salvation appeared on the horizon in the form of the Jordeson, a creaking and already overladen cargo ship bound for London.With the immediate crisis past, the magnitude of what had happened started to sink in. In a letter2 written aboard the Jordeson to botanist Richard Spruce (see go.nature.com/3prhbdk), Wallace tallied his catastrophic losses — “almost all the reward of my four years of privation & danger was lost” — and concluded with characteristic understatement, “I have some need of philosophic resignation to bear my fate with patience and equanimity.”
Evolution’s red-hot radical
The Jordeson finally limped into Deal, England, on 1 October 1852. Wallace had been at sea for 80 days. His outward voyage with Bates had taken only 29 days.Wallace added a PS to his letter to Spruce. First there was immediate exhilaration about the return — “Such a dinner! Oh! beef steaks & damson tart”. But then came thoughts about the future: “Fifty times since I left Pará [Belém] have I vowed if I once reached England never to trust myself more on the ocean.” Even then, he noted that “good resolutions soon fade”.Stevens had thoughtfully taken out insurance. So Wallace had £200 (US$980 at the time) — a fraction of his collections’ actual value — to cover his costs for a year in London while he tried to salvage what he could from the disaster and make future plans.He rushed out two books, one a travelogue, the other a more technical account of the palm trees of the Amazon. Neither did well — 250 copies remained unsold a decade later from the travel book’s print run of 750. But he was getting his name out there. Stevens, too, had a done a good job of publicizing Wallace’s discoveries while Wallace had been away.Perhaps most crucially, the positive response of the UK Royal Geographical Society to his mapping work of the Rio Negro yielded a free steamship ticket to Singapore.In March 1854, less than 18 months since the Jordeson’s bedraggled arrival at Deal, Wallace departed from Southampton in England for what he would call the “central and controlling incident”2 of his life.Eight more years of perilous travel awaited. So, too, did the discoveries of what came to be known as Wallace’s Line (a boundary between the Asian and Australasian biogeographic regions) and of the theory of evolution by natural selection3,4.The scientific acclaim that greeted Wallace’s return from southeast Asia in 1862 was a just reward both for his contributions and for that phenomenal doggedness — his determination, despite everything, to be a scientist. More

Material characterization resultsTo investigate the structural composition of NMC-2, XRD analysis plots were performed. Figure 1a shows the XRD pattern of the NMC-2 composite before adsorption. The XRD pattern shows the corresponding strong and narrow peaks, from which it can be seen that the peaks of broad diffraction NMC-2 can correspond to the standard cards of Fe, C, Fe7C3, Fe2C, and FeC, indicating that the synthesized adsorbent is an iron-carbon composite. It can be indicated that mesoporous nitrogen-doped composites were formed during the carbonization process. During the experiments, it was found that the materials are magnetic, probably because of the presence of Fe, FeC, Fe7C3, Fe2C. Due to the magnetic properties of this type of material, rapid separation and recovery can be obtained under the conditions of an applied magnetic field, which allows easy separation of the adsorbent and metal ions from the wastewater15.Figure 1XRD and nitrogen adsorption and desorption tests on materials: (a) XRD pattern of NMC-2 adsorbent before adsorption, (b) pore size distribution of NMC-2, (c) nitrogen adsorption–desorption curve of NMC-2 adsorbent.Full size imageFrom the adsorption–desorption curves of adsorbent N2 in Fig. 1b, it can be seen that the NMC-2 isotherm belongs to the class IV curve, and the appearance of H3-type hysteresis loops is observed at the medium pressure end, and H3 is commonly found in aggregates with laminar structure, producing slit mesoporous or macroporous materials, which indicates that N2 condenses and accumulates in the pore channels, and these phenomena prove that NMC-2 is a porous material16. Figure 1c shows the pore size distribution of the adsorbent NMC-2 obtained according to the BJH calculation method, from which it can be seen that the pore size distribution is not uniform in the range, and most of them are concentrated below 20 nm, while according to Table 1, the specific surface area of the original sample of Fenton sludge and fly ash is 124.08 m2/g and 3.79 m2/g, respectively, and the specific surface area of NMC-2 is 228.65 m2/g. The Fenton The pore volume of the original samples of Fenton sludge and fly ash were 0.18 cm3/g and 0.006 cm3/g respectively, while the pore volume of NMC-2 was 0.24 cm3/g. The pore diameters of the original sample of Fenton sludge and fly ash were 5.72 nm and 6.70 nm respectively, while the pore diameter of NMC-2 was 4.22 nm. The above data indicated that the synthetic materials have increased the specific surface area and pore volume compared with the original samples, indicating that the doping of nitrogen can increase the specific surface area of the material. Since the pore size of mesoporous materials is 2–50 nm, NMC-2 is a porous material with main mesopores. Thanks to the large specific surface area provided by the mesopores, the material has a large number of active sites, and in addition, the mesopores can store more Cr(VI)16, which contributes to efficient removal.Table 1 Total pore-specific surface area, pore volume, and pore size of BJH adsorption and accumulation of Fenton sludge, fly ash and NMC-2.Full size tableThe morphological analysis of the material surface using SEM can see the surface structure and the pore structure of NMC-2. And Fig. 2a–d shows the swept electron microscope image of NMC-2. Figure 2a shows that the surface of the material is not smooth, and there are more lint-like fiber structures. The fibers in Fig. 2b are loosely and irregularly arranged, which may be due to the irregular morphology caused by the small particles of the NMC-2 sample. As shown in Fig. 2c and Fig. 2a there are more pores generated on the surface of NMC-2, which may be due to the addition of K2CO3 to urea and, Fenton sludge solution to generate CO217.Figure 2SEM, TEM and EDS testing of materials: (a–d) SEM image of NMC-2 adsorbent, (e) TEM image of NMC-2; (g–i) TEM-EDS spectrum of NMC-2, (j) TEM-EDS spectra of NMC-2 obtained from.Full size imageThese pores can provide many active sites, which is consistent with the results derived in Fig. 1, where NMC-2 is a mesoporous-dominated porous material, and also demonstrates that the addition of urea can provide a nitrogen source for the material, providing abundant active sites. Figure 2j depicts the TEM of NMC-2. the TEM images show that the synthesized NMC-2 has a folded structure with a surface covered by a carbon film, and the HRTEM (Fig. 2e) also confirms this result with a lattice spacing of 0.13, 0.15, 0.20, 0.23, 0.24, and 0.25 nm, corresponding to the (4 5 2) and (1 0 2) of C, the (2 0 1) of FeC) surface, the (2 1 0) surface of Fe7C3, the (5 3 1) surface of Fe2C, and the (2 0 1) surface of FeC, which also confirms the synthesis of the above substances. The corresponding EDS spectra of the dark field diagram NMC-2 were obtained from Fig. 2j, and the EDS spectra proved the presence of various elements: carbon (C) (Fig. 2f) from fly ash, iron (Fe) (Fig. 2g) from Fenton sludge, nitrogen (N) (Fig. 2h) from urea, and the presence of (O) (Fig. 2i), further confirming the successful preparation of NMC-2.The type of functional groups and chemical bonding on the surface of the material can be analyzed by IR spectrogram analysis. Figure 3b shows the FTIR image of NMC-2 adsorbent 3440 cm−1 wide and strong absorption peak is due to the stretching vibration of –OH, there is a large amount of –OH present on the surface of the material; the peak appearing at 1640 cm−1 is –COOH. Characterization reveals that the –OH absorption peak is wider18,19. In addition, the absorptions at 1390 cm−1 and 1000 cm−1 were attributed to the bending of –OH vibrations of alcohols and phenol and the stretching vibration of C–O20. The above results indicate that the surface of NMC-2 contains a large number of oxygen-containing functional groups, and these functional groups can provide many active sites for the removal of Cr(VI). It was also found that the weak peaks corresponding to 573 cm−1 and 550 cm−1 were attributed to Fe–O groups21. The stretching of Fe–O may be due to the oxidation of loaded Fe0 and Fe2+ to Fe3+22. Figure 3a shows the Fenton sludge and fly ash FTIR images. It can be seen from the figure that the surfaces of Fenton sludge and fly ash contain a large number of oxygen-containing functional groups, the surface functional groups of the two raw materials are more abundant, and the functional groups of NMC-2 around 3441 cm−1, 1632 cm−1, and 1400 cm−1 are not significantly different from those of the raw materials, and the C–H stretching vibration peaks of NMC-2 around 873 cm−1 and 698 cm−1 is not obvious, which may be because the material the C–H bond on the surface of the raw material was oxidized to C–O in the process of synthesis.Figure 3FTIR testing of materials: (a) FTIR image of Fenton sludge, fly ash, (b) Ftir image of NMC-2 adsorbent.Full size imageCr(VI) adsorption experimentSelection of adsorbentTo select the best adsorbent, Cr(VI) adsorption tests were performed on four adsorbents. Figure 4a shows the effect of Fenton sludge and the urea addition on the adsorption efficiency. The Cr(VI) removal rates of the four adsorbents were ranked from low to high: MC-1  More