Ecology

1.GESAMP. Global pollution trends: coastal ecosystem assessment for the past century. 101 (2018).2.Andrady, A. L. Microplastics in the marine environment. Mar. Pollut. Bull. 62, 1596–1605 (2011).CAS 
PubMed 

Google Scholar 
3.Hartmann, N. B. et al. Microplastics as vectors for environmental contaminants: Exploring sorption, desorption, and transfer to biota. Integr. Environ. Assess. Manag. 13, 488–493 (2017).PubMed 

Google Scholar 
4.Bour, A., Avio, C. G., Gorbi, S., Regoli, F. & Hylland, K. Presence of microplastics in benthic and epibenthic organisms: Influence of habitat, feeding mode and trophic level. Environ. Pollut. 243, 1217–1225 (2018).CAS 
PubMed 

Google Scholar 
5.Jambeck, J. R. et al. Plastic waste inputs from land into the ocean. Science (80-) 47, 768–770 (2015).ADS 

Google Scholar 
6.PlasticsEurope. Plastics—The Facts 2020. PlasticsEurope (2020).7.Sweet, M., Steifox, M. & Lamb, J. Plastics and Shallow Water Coral Reefs. Synthesis of the Science for Policy-Makers (2019).8.Stafford, R. & Jones, P. J. S. Viewpoint—Ocean plastic pollution: A convenient but distracting truth? Mar. Policy 19, 0–1 (2019).9.Backhaus, T. & Wagner, M. Microplastics in the environment: Much ado about nothing? A debate. Glob. Challenges 1900022, 1900022 (2019).
Google Scholar 
10.Browne, M. A., Niven, S. J., Galloway, T. S., Rowland, S. J. & Thompson, R. C. Microplastic moves pollutants and additives to worms, reducing functions linked to health and biodiversity. Curr. Biol. 23, 2388–2392 (2013).CAS 
PubMed 

Google Scholar 
11.Lebreton, L. C. M. et al. River plastic emissions to the world’s oceans. Nat. Commun. 8, 15611 (2017).CAS 
PubMed 
PubMed Central 
ADS 

Google Scholar 
12.Donovan, M. K. et al. Nitrogen pollution interacts with heat stress to increase coral bleaching across the seascape. Proc. Natl. Acad. Sci. USA. 117, 5351–5357 (2020).CAS 
PubMed 
PubMed Central 

Google Scholar 
13.Lamb, J. B. et al. Plastic waste associated with disease on coral reefs. Science (80-) 359, 460–462 (2018).CAS 
ADS 

Google Scholar 
14.Burke, L., Reytar, K., Spalding, M. & Perry, A. Reefs at Risk Revisited (2011).15.Tan, F. et al. Microplastic pollution around remote uninhabited coral reefs of Nansha Islands, South China Sea. Sci. Total Environ. 725, 138383 (2020).CAS 
PubMed 
ADS 

Google Scholar 
16.Bucol, L. A. et al. Microplastics in marine sediments and rabbitfish (Siganus fuscescens) from selected coastal areas of Negros Oriental, Philippines. Mar. Pollut. Bull. 150, 110685 (2020).CAS 
PubMed 

Google Scholar 
17.Galloway, T. S., Cole, M. & Lewis, C. Interactions of microplastic debris throughout the marine ecosystem. Nat. Ecol. Evol. 1, 1–8 (2017).
Google Scholar 
18.Lagarde, F. et al. Microplastic interactions with freshwater microalgae: Hetero-aggregation and changes in plastic density appear strongly dependent on polymer type. Environ. Pollut. 215, 331–339 (2016).CAS 
PubMed 

Google Scholar 
19.Cordova, M. R., Hadi, T. A. & Prayudha, B. Occurrence and abundance of microplastics in coral reef sediment: A case study in Sekotong, Lombok-Indonesia. AES Bioflux 10, 23–29 (2018).
Google Scholar 
20.Ogston, A. S., Storlazzi, C. D., Field, M. E. & Presto, M. K. Sediment resuspension and transport patterns on a fringing reef flat, Molokai, Hawaii. Coral Reefs 23, 559–569 (2004).
Google Scholar 
21.Bellwood, D. R. Direct estimate of bioerosion by two parrotfish species, Chlorurus gibbus and C. sordidus on the Great Barrier Reef, Australia. Mar. Biol. 121, 419–429 (1995).
Google Scholar 
22.Rosenfeld, M., Bresler, V. & Abelson, A. Sediment as a possible food source for corals. Ecol. Lett. 2, 345–348 (1999).
Google Scholar 
23.Rogers, C. S. Responses of coral reefs and reef organisms to sedimentation. Mar. Ecol. Prog. Ser. 62, 185–202 (1990).ADS 

Google Scholar 
24.Bastidas, C., Bone, D. & Garcia, E. M. Sedimentation rates and metal content of sediments in a Venezuelan coral reef. Mar. Pollut. Bull. 38, 16–24 (1999).CAS 

Google Scholar 
25.Smith, L. D., Negri, A. P., Philipp, E., Webster, N. S. & Heyward, A. J. The effects of antifoulant-paint-contaminated sediments on coral recruits and branchlets. Mar. Biol. 143, 651–657 (2003).CAS 

Google Scholar 
26.Stafford-Smith, M. Sediment rejection efficiency of 22 species of Australian scleractinian corals. Mar. Biol. 115, 229–243 (1993).
Google Scholar 
27.Junjie, R. K., Browne, N. K., Erftemeijer, P. L. A. & Todd, P. A. Impacts of sediments on coral energetics: Partitioning the effects of turbidity and settling particles. PLoS ONE 9, e107195 (2014).PubMed 
PubMed Central 
ADS 

Google Scholar 
28.Hall, N. M., Berry, K. L. E., Rintoul, L. & Hoogenboom, M. O. Microplastic ingestion by scleractinian corals. Mar. Biol. 162, 725–732 (2015).CAS 

Google Scholar 
29.Allen, A. S., Seymour, A. C. & Rittschof, D. Chemoreception drives plastic consumption in a hard coral. Mar. Pollut. Bull. 124, 198–205 (2017).CAS 
PubMed 

Google Scholar 
30.Mouchi, V. et al. Long-term aquaria study suggests species-specific responses of two cold-water corals to macro-and microplastics exposure. Environ. Pollut. 253, 322–329 (2019).CAS 
PubMed 

Google Scholar 
31.Tang, J., Ni, X., Zhou, Z., Wang, L. & Lin, S. Acute microplastic exposure raises stress response and suppresses detoxification and immune capacities in the scleractinian coral Pocillopora damicornis. Environ. Pollut. 243, 66–74 (2018).CAS 
PubMed 

Google Scholar 
32.Chapron, L. et al. Macro- and microplastics affect cold-water corals growth, feeding and behaviour. Sci. Rep. 8, 1–8 (2018).CAS 

Google Scholar 
33.Hankins, C., Moso, E. & Lasseigne, D. Microplastics impair growth in two atlantic scleractinian coral species, Pseudodiploria clivosa and Acropora cervicornis. Environ. Pollut. 275, 116649 (2021).CAS 
PubMed 

Google Scholar 
34.Rocha, R. J. M. et al. Do microplastics affect the zoanthid Zoanthus sociatus?. Sci. Total Environ. 713, 136659 (2020).CAS 
PubMed 
ADS 

Google Scholar 
35.Reichert, J. et al. Interactive effects of microplastic pollution and heat stress on reef-building corals. Environ. Pollut. 290, 118010 (2021).CAS 
PubMed 

Google Scholar 
36.Rotjan, R. D. et al. Patterns, dynamics and consequences of microplastic ingestion by the temperate coral, Astrangia poculata. Proc. R. Soc. B Biol. Sci. 286, 20190726 (2019).CAS 

Google Scholar 
37.Reichert, J., Schellenberg, J., Schubert, P. & Wilke, T. Responses of reef building corals to microplastic exposure. Environ. Pollut. 237, 955–960 (2018).CAS 
PubMed 

Google Scholar 
38.Hankins, C., Duffy, A. & Drisco, K. Scleractinian coral microplastic ingestion: Potential calcification effects, size limits, and retention. Mar. Pollut. Bull. 135, 587–593 (2018).CAS 
PubMed 

Google Scholar 
39.Reichert, J., Arnold, A. L., Hoogenboom, M. O., Schubert, P. & Wilke, T. Impacts of microplastics on growth and health of hermatypic corals are species-specific. Environ. Pollut. 254, 113074 (2019).CAS 
PubMed 

Google Scholar 
40.Mendrik, F. M. et al. Species-specific impact of microplastics on coral physiology. Environ. Pollut. 269, 116238 (2021).CAS 
PubMed 

Google Scholar 
41.Corona, E., Martin, C., Marasco, R. & Duarte, C. M. Passive and active removal of marine microplastics by a mushroom Coral (Danafungia scruposa). Front. Mar. Sci. 7, 1–9 (2020).ADS 

Google Scholar 
42.Martin, C., Corona, E., Mahadik, G. A. & Duarte, C. M. Adhesion to coral surface as a potential sink for marine microplastics. Environ. Pollut. 255, 113281 (2019).CAS 
PubMed 

Google Scholar 
43.Oldenburg, K. S., Urban-Rich, J., Castillo, K. D. & Baumann, J. H. Microfiber abundance associated with coral tissue varies geographically on the Belize Mesoamerican Barrier Reef System. Mar. Pollut. Bull. 163, 111938 (2021).CAS 
PubMed 

Google Scholar 
44.Axworthy, J. B. & Padilla-Gamiño, J. L. Microplastics ingestion and heterotrophy in thermally stressed corals. Sci. Rep. 9, 1–8 (2019).
Google Scholar 
45.Houlbrèque, F. & Ferrier-Pagès, C. Heterotrophy in tropical scleractinian corals. Biol. Rev. 84, 1–17 (2009).PubMed 

Google Scholar 
46.Borja, A. et al. Past and future grand challenges in marine ecosystem ecology. Front. Mar. Sci. 7, 362 (2020).
Google Scholar 
47.Rochman, C. M., Hentschel, B. T. & The, S. J. Long-term sorption of metals is similar among plastic types: Implications for plastic debris in aquatic environments. PLoS ONE 9, e85433 (2014).PubMed 
PubMed Central 
ADS 

Google Scholar 
48.Niu, Y., Ying, D., Li, K., Wang, Y. & Jia, J. Adsorption of heavy-metal ions from aqueous solution onto chitosan-modified polyethylene terephthalate (PET). Res. Chem. Intermed. 43, 4213–4245 (2017).CAS 

Google Scholar 
49.Frias, J., Sobral, P. & Ferreira, A. Organic pollutants in microplastics from two beaches of the Portuguese coast. Mar. Pollut. Bull. 60, 1988–1992 (2010).CAS 
PubMed 

Google Scholar 
50.Wright, S. L., Thompson, R. C. & Galloway, T. S. The physical impacts of microplastics on marine organisms: A review. Environ. Pollut. 178, 483–492 (2013).CAS 
PubMed 

Google Scholar 
51.Botterell, Z. L. R. et al. Bioavailability of microplastics to marine zooplankton: Effect of shape and infochemicals. Environ. Sci. Technol. 54, 12024–12033 (2020).CAS 
PubMed 
ADS 

Google Scholar 
52.Wild, C. et al. Coral mucus functions as an energy carrier and particle trap in the ecosystem. Nature 428, 66–70 (2004).CAS 
PubMed 
ADS 

Google Scholar 
53.Benson, A. & Muscatine, L. Wax in coral mucus: Energy transfer from corals to reef fishes. Limnol. Oceanogr. 19, 810–814 (1974).ADS 

Google Scholar 
54.Verla, A. W., Enyoh, C. E., Verla, E. N. & Nwarnorh, K. O. Microplastic–toxic chemical interaction: A review study on quantified levels, mechanism and implication. SN Appl. Sci. 1, 1–30 (2019).CAS 

Google Scholar 
55.Brown, B. E. & Bythell, J. C. Perspectives on mucus secretion in reef corals. Mar. Ecol. Prog. Ser. 296, 291–309 (2005).CAS 
ADS 

Google Scholar 
56.Weber, M., Lott, C. & Fabricius, K. E. Sedimentation stress in a scleractinian coral exposed to terrestrial and marine sediments with contrasting physical, organic and geochemical properties. J. Exp. Mar. Bio. Ecol. 336, 18–32 (2006).CAS 

Google Scholar 
57.Riegl, B. & Branch, G. Effects of sediment on the energy budgets of four scleractinian (Bourne 1900) and five alcyonacean (Lamoroux 1816) corals. J. Exp. Mar. Bio. Ecol. 186, 259–275 (1995).
Google Scholar 
58.Felsing, S. et al. A new approach in separating microplastics from environmental samples based on their electrostatic behavior. Environ. Pollut. 234, 20–28 (2018).CAS 
PubMed 

Google Scholar 
59.Bessell-Browne, P., Negri, A. P., Fisher, R., Clode, P. L. & Jones, R. Cumulative impacts: Thermally bleached corals have reduced capacity to clear deposited sediment. Sci. Rep. 7, 1–14 (2017).
Google Scholar 
60.Fitt, W. K. et al. Response of two species of Indo-Pacific corals, Porites cylindrica and Stylophora pistillata, to short-term thermal stress: The host does matter in determining the tolerance of corals to bleaching. J. Exp. Mar. Bio. Ecol. 373, 102–110 (2009).
Google Scholar 
61.Weber, M. et al. Mechanisms of damage to corals exposed to sedimentation. Proc. Natl. Acad. Sci. USA. 109, E1558 (2012).CAS 
PubMed 
PubMed Central 

Google Scholar 
62.Lear, G. et al. Plastics and the microbiome: Impacts and solutions. Environ. Microbiomes 16, 1–19 (2021).
Google Scholar 
63.Palardy, J., Rodrigues, L. & Grottolli, A. The importance of zooplankton to the daily metabolic carbon requirements of healthy and bleached corals at two depths. J. Exp. Mar. Bio. Ecol. 367, 180–188 (2008).CAS 

Google Scholar 
64.Jennings, H. Modifiability in behaviour, 1: Behavior of sea anemones. J. Exp. Zool. 4, 447–632 (1905).
Google Scholar 
65.Boschma, H. On the feeding reactions and digestion in the coral polyp Astrangia danae, with notes on its symbiosis with zooxanthellae. Biol. Bull. 49, 407–439 (1925).CAS 

Google Scholar 
66.Anthony, K. R. N. Coral suspension feeding on fine particulate matter. J. Exp. Mar. Bio. Ecol. 232, 85–106 (1999).
Google Scholar 
67.Schlekat, C., McGee, B. & Reinharz, E. Testing sediment toxicity in chesapeake bay with the amphipod Leptocheirus plumulosus: An evaluation. Environ. Toxicol. Chem. 11, 225–236 (1992).CAS 

Google Scholar 
68.Nie, H., Wang, J., Xu, K., Huang, Y. & Yan, M. Microplastic pollution in water and fish samples around Nanxun Reef in Nansha Islands, South China Sea. Sci. Total Environ. 696, 134022 (2019).CAS 
PubMed 
ADS 

Google Scholar 
69.Cai, M. et al. Lost but can’t be neglected: Huge quantities of small microplastics hide in the South China Sea. Sci. Total Environ. 633, 1206–1216 (2018).CAS 
PubMed 
ADS 

Google Scholar 
70.Zhu, L. et al. Microplastic pollution in North Yellow Sea, China: Observations on occurrence, distribution and identification. Sci. Total Environ. 636, 20–29 (2018).CAS 
PubMed 
ADS 

Google Scholar 
71.Saliu, F. et al. Microplastic and charred microplastic in the Faafu Atoll, Maldives. Mar. Pollut. Bull. 136, 464–471 (2018).CAS 
PubMed 

Google Scholar 
72.Saliu, F., Montano, S., Leoni, B., Lasagni, M. & Galli, P. Microplastics as a threat to coral reef environments: Detection of phthalate esters in neuston and scleractinian corals from the Faafu Atoll, Maldives. Mar. Pollut. Bull. 142, 234–241 (2019).CAS 
PubMed 

Google Scholar 
73.Bessa, F. et al. Occurrence of microplastics in commercial fish from a natural estuarine environment. Mar. Pollut. Bull. 128, 575–584 (2018).CAS 
PubMed 

Google Scholar 
74.Ivar Do Sul, J. A. & Costa, M. F. The present and future of microplastic pollution in the marine environment. Environ. Pollut. 185, 352–364 (2014).CAS 
PubMed 

Google Scholar 
75.Chubarenko, I., Bagaev, A., Zobkov, M. & Esiukova, E. On some physical and dynamical properties of microplastic particles in marine environment. Mar. Pollut. Bull. 108, 105–112 (2016).CAS 
PubMed 

Google Scholar 
76.Tang, J. et al. Differential enrichment and physiological impacts of ingested microplastics in scleractinian corals in situ. J. Hazard. Mater. 404, 124205 (2021).CAS 
PubMed 

Google Scholar 
77.Harrison, J. P., Schratzberger, M., Sapp, M. & Osborn, A. M. Rapid bacterial colonization of low-density polyethylene microplastics in coastal sediment microcosms. BMC Microbiol. 14, 1–15 (2014).
Google Scholar 
78.Veron, J. Corals of the World (2000).79.Benavides, M. et al. Diazotrophs: A non-negligible source of nitrogen for the tropical coral Stylophora pistillata. J. Exp. Biol. 219, 2608–2612 (2016).PubMed 

Google Scholar 
80.Einbinder, S. et al. Changes in morphology and diet of the coral Stylophora pistillata along a depth gradient. Mar. Ecol. Prog. Ser. 381, 167–174 (2009).ADS 

Google Scholar 
81.Erni-Cassola, G., Gibson, M. I., Thompson, R. C. & Christie-Oleza, J. A. Lost, but found with Nile red: A novel method for detecting and quantifying small microplastics (1 mm to 20 μm) in environmental samples. Environ. Sci. Technol. 51, 13641–13648 (2017).CAS 
PubMed 
ADS 

Google Scholar 
82.Swain, T. D., Schellinger, J. L., Strimaitis, A. M. & Reuter, K. E. Evolution of anthozoan polyp retraction mechanisms: Convergent functional morphology and evolutionary allometry of the marginal musculature in order Zoanthidea (Cnidaria: Anthozoa: Hexacorallia). BMC Evol. Biol. 15, 1–19 (2015).
Google Scholar 
83.Laissue, P. P., Gu, Y., Qian, C. & Smith, D. J. Light-induced polyp retraction and tissue rupture in the photosensitive, reef-building coral Acropora muricata. bioRxiv https://doi.org/10.1101/862045 (2019).Article 

Google Scholar 
84.Renegar, D. A. & Turner, N. R. Species sensitivity assessment of five Atlantic scleractinian coral species to 1-methylnaphthalene. Sci. Rep. 11, 1–17 (2021).
Google Scholar 
85.Armoza-Zvuloni, R., Schneider, A., Sher, D. & Shaked, Y. Rapid Hydrogen Peroxide release from the coral Stylophora pistillata during feeding and in response to chemical and physical stimuli. Sci. Rep. 6, 1–10 (2016).
Google Scholar 
86.Schneider, C. A., Rasband, W. S. & Eliceiri, K. W. NIH Image to ImageJ: 25 years of image analysis. Nat. Methods 9, 671–675 (2012).CAS 
PubMed 
PubMed Central 

Google Scholar 
87.Meijering, E., Dzyubachyck, O. & Smal, I. Methods for cell and particle tracking. In Methods in Enzymology (Elsevier, 2012).88.Sorgeloos, P., Dhert, P. & Candrevab, P. Use of the brine shrimp, Artemia spp., in marine fish larviculture. Aquaculture 200, 147–159 (2001).
Google Scholar 
89.R Core Team. R: A language and environment for statistical computing (2019).90.Wood, A. S., Scheipl, F. & Wood, M. S. Package ‘gamm4’. (2020).91.Zuur, A., Saveliev, A. & Ieno, E. A Beginner’s Guide to Generalized Additive Mixed Models with R. (Highland Statistics Ltd, 2014).92.Zuur, A. F., Hilbe, J. M. & Ieno, E. N. A Beginner’s Guide to GLM and GLMM with R (Highland Statistics Ltd, 2013).93.Pyke, A. & Thompson, J. Statistical analysis of survival and removal rate experiments. Ecology 67, 240–245 (1986).
Google Scholar 
94.Therneau, T. M. Mixed effects cox models. R-Package Description. https://doi.org/10.1111/oik.01149 (2015).Article 

Google Scholar 
95.Katki, H. A. & Mark, S. D. Survival analysis of studies nested within cohorts using the NestedCohort Package. 1–16 (2013).96.Bürkner, P. C. Advanced Bayesian multilevel modeling with the R package BRMS. R J. 10, 395–411 (2018).
Google Scholar 
97.Zuur, A. F., Ieno, E. N., Walker, N. J., Saveliev, A. A. & Smith, G. M. Mixed Effects Models and Extensions in Ecology with R. (Springer, 2009).98.Manly, B. F. J. Measuring selectivity from multiple choice feeding-preference experiments. Biometrics 51, 709 (1995).
Google Scholar 
99.Richardson, J. Package ‘ selectapref ’. Anal. F. Lab. Foraging 8–11 (2020). More

1.Mittermeier, R. A., Coimbra-Filho, A. F., Constable, I. D., Rylands, A. B. & Valle, C. Conservation of primates in the Atlantic forest region of eastern Brazil. Int. Zoo Yearb. 22, 2–17 (1982).
Google Scholar 
2.Mittermeier, R. A., Rylands, A. B. & Wilson, D. E. W. Handbook of the Mammals of the World, vol. 3, Primates. Lynx, Barcelona (2013).3.Rylands, A. B., Kierulff, M. C. M., Mendes, S. L. & Oliveira, M. M. Callithrix aurita. The IUCN Red List of Threatened Species 2008. IUCN Red List Threat. Species 8235, 1–7 (2008).
Google Scholar 
4.IUCN. The IUCN Red List of Threatened Species. Version 2020–2, Vol. 8235 (2020).5.Coimbra-Filho, A., Pissinatti, A. & Rylands, A. B. Experimental mutiple hybridism and natural hybrids among Callithrix species from easterna Brazil. In Marmosets and Tamarins Systematics Behaviour and Ecology (ed. Rylands, A. B.) 93–120 (Oxford University Press, 1993).
Google Scholar 
6.Mittermeier, R. A., Coimbra-Filho, A. F., Rylands, A. B. & Constable, I. D. Atlantic Forest region of eastern Brazil a top primate of conservation priority. IUCN / SSC Primate Spec. Gr. Newsl. 1, 9–11 (1981).
Google Scholar 
7.Carvalho, R. S. et al. Buffy-tufted-year marmoset Callithrix aurita É. Geoffroy Saint-Hilaire, 1812 Brazil. In Primates in Peril: The World’s 25 Most Endangered Primates 2018–2020 (eds. Schwitzer, C. et al.) 136 (IUCN SSC Primate Specialist Group, International Primatological Society, Global Wildlife Conservation, Bristol Zoological Society, 2019).8.ICMBio. Instituto Chico Mendes de Conservação da Biodiversidade. Livro Vermelho da Fauna Brasileira Ameaçada de Extinção. (2018).9.SMA-SP. Decreto no63.853, de 27 de novembro de 2018. (2018).10.Rylands, A. B. et al. An assessment of the diversity of New World Primates. Neotrop. Primates 8, 61–93 (2000).
Google Scholar 
11.Brandão, L. D. & Develey, P. F. Distribution and conservation of the buffy-tufted-ear marmoset, Callithrix aurita, in lowland coastal, Atlantic Forest, Southeast Brazil. Neotrop. Primates 6, 86–88 (1998).
Google Scholar 
12.Carvalho, R. S. et al. Callithrix aurita: a marmoset species on its way to extinctionin the Brazilian Atlantic Forest. Neotrop. Primates 24, 1–8 (2018).
Google Scholar 
13.Malukiewicz, J. et al. Natural and anthropogenic hybridization in two species of eastern Brazilian marmosets (Callithrix jacchus and C. penicillata). PLoS ONE 10, 1–22 (2015).
Google Scholar 
14.Ruiz-Miranda, C. R., Affonso, A. G., Martins, A. & Beck, B. Distribuição do sagui (Callithrix jacchus) nas áreas de ocorrência do mico-leão-dourado (Leontopithecus rosalia) no Estado do Rio de Janeiro. Neotrop. Primates 8, 98–101 (2000).
Google Scholar 
15.Mendes, S. L. Hybridization in free-ranging Callithrix flaviceps and the taxonomy of the Atlantic forest marmosets. Neotrop. Primates 5, 6–8 (1997).
Google Scholar 
16.Santos, C. V. et al. Ecologia, comportamento e manejo de primatas invasores e populações-problema. A Primatologia no Brasil 10, 101–118 (2007).
Google Scholar 
17.Rhymer, J. M. & Simberloff, D. Extinction by hybridization and introgression. Annu. Rev. Ecolology Evol. Syst. 27, 83–109 (1996).
Google Scholar 
18.Zinner, D., Arnold, M. L. & Roos, C. The strange blood: Natural hybridization in primates. Evol. Anthropol. 20, 96–103 (2011).PubMed 

Google Scholar 
19.Brumfield, R. T. Speciation genetics of biological invasions with hybridization. Mol. Ecol. 19, 5079–5083 (2010).PubMed 

Google Scholar 
20.Steeves, T. E., Maloney, R. F., Hale, M. L., Tylianakis, J. M. & Gemmell, N. J. Genetic analyses reveal hybridization but no hybrid swarm in one of the world’s rarest birds. Mol. Ecol. 19, 5090–5100 (2010).PubMed 

Google Scholar 
21.Allendorf, F. W., Leary, R. F., Spruell, P. & Wenburg, J. K. The problems with hybrids: Setting conservation guidelines. Trends Ecol. Evol. 16, 613–622 (2001).
Google Scholar 
22.Bechara, I. M. Abordagens metodológicas em Biogeografia da Conservação para avaliar risco de extinção de espécies: um estudo de caso com Callithrix aurita (Primates: Callitrichidae). (UFRJ, 2012).23.Braz, A. G., Lorini, M. L. & Vale, M. M. Climate change is likely to affect the distribution but not parapatry of the Brazilian marmoset monkeys (Callithrix spp.). Divers. Distrib. 25, 536–550 (2019).
Google Scholar 
24.Malukiewicz, J. A review of experimental, natural, and anthropogenic hybridization in Callithrix marmosets. Int. J. Primatol. 40, 72–98 (2019).
Google Scholar 
25.Pinto, L. F. G. & Voivodic, M. Reverse the tipping point of the Atlantic Forest for mitigation. Nat. Clim. Chang. 11, 364–365 (2021).ADS 

Google Scholar 
26.Seehausen, O., Takimoto, G., Roy, D. & Jokela, J. Speciation reversal and biodiversity dynamics with hybridization in changing environments. Mol. Ecol. 17, 30–44 (2008).PubMed 

Google Scholar 
27.Pereira, D. G., De Oliveira, M. E. A. & Ruiz-Miranda, C. R. Interações entre calitriquídeos exóticos e nativos no Parque Nacional da Serra dos Órgãos – RJ. Espaço e Geogr. 11, 87–114 (2008).
Google Scholar 
28.Aximoff, I., Soares, H. M., Pissinatti, A. & Bueno, C. Registros de Callithrix aurita (Primates, Callitrichidae) e seus híbridos no Parque Nacional do Itatiaia. Oecologia Aust. 20, 520–525 (2016).
Google Scholar 
29.Detogne, N. et al. Spatial distribution of buffy-tufted-ear (Callithrix aurita) and invasive marmosets (Callithrix spp.) in a tropical rainforest reserve in southeastern Brazil. Am. J. Primatol. 79, 1–11 (2017).
Google Scholar 
30.Carvalho, R. S. et al. Molecular identification of a buffy-tufted-ear marmoset (Callithrix aurita) incorporated in a group of invasive marmosets in the Serra dos Órgãos National Park, Rio de Janeiro – Brazil. Forensic Sci. Int. Genet. Suppl. Ser. 4, e230–e231 (2013).
Google Scholar 
31.Novaes, C. M. et al. Karyotypic characteristics of hybrid marmosets of the genus Callithrix (Erxeleben, 1777) suggest the participation of three parental species. Bol. do Mus. Biol. Mello Leitão 39, 11–21 (2017).
Google Scholar 
32.Pereira, D. G. Densidade, genética e saúde populacional como ferramentas para propor um plano de controle e erradicação de invasão biológica: o caso de Callithrix aurita (Primates) no Parque Nacional da Serra dos Órgãos, RJ, Brasil (Universidade do Estado do Rio de Janeiro, 2010).
Google Scholar 
33.Veracini, C., Galeni, L. & Forti, M. The concept of species and the foundations of biology, a case study: The Callithrix jacchus group (Primates-Platyrrhini). Riv. Biol. 95, 75–100 (2002).PubMed 

Google Scholar 
34.Campton, D. E. Natural hybridization and introgression in fishes: methods of detection and genetic interpretations. In Population Genetics and Fishery Management (eds Ryman, N. & Utter, F.) 16–192 (University of Washington Press, 1987).
Google Scholar 
35.Nagamachi, C. Y., Pieczarka, J. C., Schwarz, M., Barros, R. M. S. & Mattevi, M. S. Comparative chromosomal study of five taxa of genus Callithrix, group jacchus (Platyrrhini, Primates). Am. J. Primatol. 41, 53–60 (1997).CAS 
PubMed 

Google Scholar 
36.Mallet, J. Hybridization as an invasion of the genome. Trends Ecol. Evol. 20, 229–237 (2005).PubMed 

Google Scholar 
37.Ardito, G., Lamberti, L., Bigatti, P., Stanyon, R. & Govone, D. NOR distribution and satellite associations in Callithrix jacchus. Caryologia 40, 185–194 (1987).
Google Scholar 
38.Nagamashi, C. & Ferrari, I. Cytogenetic studies of Callithrix jacchus (Callitrichidae, Platyrrhini) from two different sites in Brazil. I. Morphological variability of Y chromosome. Rev. Bras. Genética 7, 497–507 (1984).
Google Scholar 
39.Nogueira, D. M. et al. Cytogenetic study in natural hybrids of Callithrix (Callitrichidae: Primates) in the Atlantic forest of the state of Rio de Janeiro, Brazil. Iheringia, Série Zool. 101, 156–160 (2011).
Google Scholar 
40.Moreira, M. A. M. SRY evolution in Cebidae (Platyrrhini: Primates). J. Mol. Evol. 55, 92–103 (2002).ADS 
CAS 
PubMed 

Google Scholar 
41.de Morais, M. M. OS SAGUIS (Callithrix spp., ERXLEBEN, 1777) Exóticos Invasores na bacia do Rio São João, Rio de Janeiro: Biologia populacional e padrão de distribuição em uma paisagem fragmentada. Uenf.Br (2010).42.Sweeney, C. G., Curran, E., Westmoreland, S. V., Mansfield, K. G. & Vallender, E. J. Quantitative molecular assessment of chimerism across tissues in marmosets and tamarins. BMC Genomics 13, 98. https://doi.org/10.1186/1471-2164-13-98 (2012).Article 
PubMed 
PubMed Central 

Google Scholar 
43.Buckner, J. C., Lynch Alfaro, J. W., Rylands, A. B. & Alfaro, M. E. Biogeography of the marmosets and tamarins (Callitrichidae). Mol. Phylogenet. Evol. 82, 413–425 (2015).PubMed 

Google Scholar 
44.Schneider, H. et al. A molecular analysis of the evolutionary relationships in the Callitrichinae, with emphasis on the position of the dwarf marmoset. Zool. Scr. 41, 1–10 (2012).
Google Scholar 
45.Perelman, P. et al. A molecular phylogeny of living primates. PLoS Genet. 7, 1–17 (2011).
Google Scholar 
46.Price, T. D. & Bouvier, M. M. The evolution of F1 postzygotic incompatibilities in birds. Evolution (N. Y). 56, 2083–2089 (2002).
Google Scholar 
47.Nievergelt, C. M., Mundy, N. I. & Woodruff, D. S. Microsatellite primers for genotyping common marmosets (Callithrix jacchus) and other callitrichids. Mol. Ecol. 7, 1432–1434 (1998).CAS 
PubMed 

Google Scholar 
48.Raveendran, M. et al. Polymorphic microsatellite loci for the common marmoset (Callithrix jacchus) designed using a cost- and time-efficient method. Am. J. Primatol. 70, 906–910 (2008).CAS 
PubMed 
PubMed Central 

Google Scholar 
49.Takabayashi, S. & Katoh, H. Noninvasive genotyping of common marmoset (Callithrix jacchus) by fingernail PCR. Primates 56, 235–240 (2015).PubMed 

Google Scholar 
50.Malukiewicz, J. et al. Mitogenomic phylogeny of Callithrix with special focus on human transferred taxa. BMC Genomics 22, 1–14 (2021).
Google Scholar 
51.Rozhnov, V. V. Extinction of the European mink: ecological catastrophe or a natural process?. Lutreola 1, 10–16 (1993).
Google Scholar 
52.Rosenthal, G. G. Individual mating decisions and hybridization. J. Evol. Biol. 26, 252–255 (2013).CAS 
PubMed 

Google Scholar 
53.Fundação SOS Mata Atlântica. Notícias. www.sosma.org.br/noticias/desmatamento-da-mata-atlantica-cresce-em-dez-estados/ (2021).54.Banks-Leite, C. et al. Using ecological thresholds to evaluate the costs and benefits of set-asides in a biodiversity hotspot. Science 345, 1041–1045 (2014).ADS 
CAS 
PubMed 

Google Scholar 
55.Oliveira, A. B. L. Presença ou Ausência do Callithrix aurita em Fragmentos de Mata Atlântica (Instituto Superior de Agronomia – Universidade Técnica de Lisboa, 2012).
Google Scholar 
56.Allendorf, F. W. et al. Intercrosses and the U.S. endangered species act: Should hybridized populations be included as westslope cutthroat trout?. Conserv. Biol. 18, 1203–1213 (2004).
Google Scholar 
57.Wayne, R. K. & Shaffer, H. B. Hybridization and endangered species protection in the molecular era. Mol. Ecol. 25, 2680–2689 (2016).PubMed 

Google Scholar 
58.U.S. Fish and Wildlife Service and National Oceanic and Atmospheric Administration. Endangered and threatened wildlife and plants: Proposed policy and proposed rule on the treatment of intercrosses and intercross progeny (the issue of “hybridization”). Fed. Regist. 67, 4710–4713 (1996).
Google Scholar 
59.Myers, N., Mittermeier, R. A., Mittermeier, C. G., Fonseca, G. A. B. & Kent, J. Biodiversity hotspots for conservation priorities. Nature 403, 853–858 (2000).ADS 
CAS 
PubMed 
PubMed Central 

Google Scholar 
60.Malukiewicz, J., Hepp, C. M., Guschanski, K. & Stone, A. C. Phylogeny of the jacchus group of Callithrix marmosets based on complete mitochondrial genomes. Am. J. Phys. Anthropol. 162, 157–169 (2016).PubMed 

Google Scholar 
61.Sambrook, J., Maniatis, T. & Fritsch, E. F. Molecular Cloning. A Laboratory Manual. Vol. 1 (Cold Spring Harbor Laboratory Press, 2001).
Google Scholar 
62.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 
63.Rozas, J. et al. DnaSP6: DNA sequence polymorphism analysis of large data sets. Mol. Biol. Evol. 34, 3299–3302 (2017).CAS 
PubMed 

Google Scholar 
64.Ronquist, F., van der Mark, P. & Huelsenbeck, J. P. Bayesian phylogenetic analysis using MRBAYES. Phylogenetic Handb. https://doi.org/10.1017/cbo9780511819049.009 (2012).Article 

Google Scholar  More

1.Graf, D. L. & Cummings, K. S. Review of the systematics and global diversity of freshwater mussel species (Bivalvia: Unionoida). J. Molluscan Stud. 73, 291–314. https://doi.org/10.1093/mollus/eym029 (2007).Article 

Google Scholar 
2.Graf, D. L. & Cummings, K. S. A “big data” approach to global freshwater mussel diversity (Bivalvia: Unionoida), with an updated checklist of genera and species. J. Molluscan Stud. 87, 034. https://doi.org/10.1093/mollus/eyaa034 (2021).Article 

Google Scholar 
3.Vaughn, C. C. Ecosystem services provided by freshwater mussels. Hydrobiologia 810, 15–27. https://doi.org/10.1007/s10750-017-3139-x (2018).Article 

Google Scholar 
4.Ożgo, M. et al. Lake-stream transition zones support hotspots of freshwater ecosystem services: Evidence from a 35-year study on unionid mussels. Sci. Total Environ. 774, 145114. https://doi.org/10.1016/j.scitotenv.2021.145114 (2021).ADS 
CAS 
Article 
PubMed 

Google Scholar 
5.Lopes-Lima, M. et al. Conservation of freshwater bivalves at the global scale: Diversity, threats and research needs. Hydrobiologia 810, 1–14. https://doi.org/10.1007/s10750-017-3486-7 (2018).Article 

Google Scholar 
6.Bolotov, I. N. et al. Climate warming as a possible trigger of keystone mussel population decline in oligotrophic rivers at the continental scale. Sci. Rep. 8, 35. https://doi.org/10.1038/s41598-017-18873-y (2018).ADS 
CAS 
Article 
PubMed 
PubMed Central 

Google Scholar 
7.Ferreira-Rodríguez, N. et al. Research priorities for freshwater mussel conservation assessment. Biol. Conserv. 231, 77–87. https://doi.org/10.1016/j.biocon.2019.01.002 (2019).Article 

Google Scholar 
8.Lundquist, S. P., Worthington, T. A. & Aldridge, D. C. Freshwater mussels as a tool for reconstructing climate history. Ecol. Ind. 101, 11–21. https://doi.org/10.1016/j.ecolind.2018.12.048 (2019).Article 

Google Scholar 
9.Sousa, R. et al. The role of anthropogenic habitats in freshwater mussel conservation. Glob. Change Biol. 27, 2298–2314. https://doi.org/10.1111/gcb.15549 (2021).ADS 
Article 

Google Scholar 
10.Bogan, A. E. Freshwater bivalve extinctions (Mollusca: Unionoida): A search for causes. Integr. Comp. Biol. 33, 599–609. https://doi.org/10.1093/icb/33.6.599 (1993).Article 

Google Scholar 
11.Lydeard, C. et al. The global decline of nonmarine mollusks. Bioscience 54, 321–330. https://doi.org/10.1641/0006-3568(2004)054[0321:TGDONM]2.0.CO;2 (2004).Article 

Google Scholar 
12.Hughes, J. et al. Past and present patterns of connectivity among populations of four cryptic species of freshwater mussels Velesunio spp (Hyriidae) in central Australia. Mol. Ecol. 13, 3197–3212. https://doi.org/10.1111/j.1365-294X.2004.02305.x (2004).CAS 
Article 
PubMed 

Google Scholar 
13.Martel, A. L. et al. Freshwater mussels (Bivalvia: Margaritiferidae, Unionidae) of the Atlantic Maritime Ecozone. In Assessment of Species Diversity in the Atlantic Maritime Ecozone (eds McAlpine, D. F. & Smith, I. M.) 551–598 (NRC Research Press, 2010).
Google Scholar 
14.Haag, W. R. North American Freshwater Mussels: Natural History, Ecology, and Conservation (Cambridge University Press, 2012).
Google Scholar 
15.Smith, C. H., Pfeiffer, J. M. & Johnson, N. A. Comparative phylogenomics reveal complex evolution of life history strategies in a clade of bivalves with parasitic larvae (Bivalvia: Unionoida: Ambleminae). Cladistics 36, 505–520. https://doi.org/10.1111/cla.12423 (2020).Article 
PubMed 

Google Scholar 
16.Sepkoski, J. J. Jr. & Rex, M. A. Distribution of freshwater mussels: Coastal rivers as biogeographic islands. Syst. Biol. 23, 165–188. https://doi.org/10.1093/sysbio/23.2.165 (1974).Article 

Google Scholar 
17.Haag, W. R. A hierarchical classification of freshwater mussel diversity in North America. J. Biogeogr. 37, 12–26. https://doi.org/10.1111/j.1365-2699.2009.02191.x (2010).Article 

Google Scholar 
18.Graf, D. L., Jones, H., Geneva, A. J., Pfeiffer, J. M. III. & Klunzinger, M. W. Molecular phylogenetic analysis supports a Gondwanan origin of the Hyriidae (Mollusca: Bivalvia: Unionida) and the paraphyly of Australasian taxa. Mol. Phylogenet. Evol. 85, 1–9. https://doi.org/10.1016/j.ympev.2015.01.012 (2015).Article 
PubMed 

Google Scholar 
19.Bolotov, I. N. et al. Ancient river inference explains exceptional Oriental freshwater mussel radiations. Sci. Rep. 7, 2135. https://doi.org/10.1038/s41598-017-02312-z (2017).ADS 
CAS 
Article 
PubMed 
PubMed Central 

Google Scholar 
20.Bolotov, I. N. et al. Integrative taxonomy, biogeography and conservation of freshwater mussels (Unionidae) in Russia. Sci. Rep. 10, 3072. https://doi.org/10.1038/s41598-020-59867-7 (2020).ADS 
CAS 
Article 
PubMed 
PubMed Central 

Google Scholar 
21.Lopes-Lima, M. et al. Diversity, biogeography, evolutionary relationships, and conservation of Eastern Mediterranean freshwater mussels (Bivalvia: Unionidae). Mol. Phylogenet. Evol. 163, 107261. https://doi.org/10.1016/j.ympev.2021.107261 (2021).Article 
PubMed 

Google Scholar 
22.Bolotov, I. N. et al. Eight new freshwater mussels (Unionidae) from tropical Asia. Sci. Rep. 9, 12053. https://doi.org/10.1038/s41598-019-48528-z (2019).ADS 
CAS 
Article 
PubMed 
PubMed Central 

Google Scholar 
23.Bolotov, I. N. et al. New freshwater mussel taxa discoveries clarify biogeographic division of Southeast Asia. Sci. Rep. 10, 6616. https://doi.org/10.1038/s41598-020-63612-5 (2020).ADS 
CAS 
Article 
PubMed 
PubMed Central 

Google Scholar 
24.Jeratthitikul, E., Paphatmethin, S., Zieritz, A., Lopes-Lima, M. & Bun, P. Hyriopsis panhai, a new species of freshwater mussel from Thailand (Bivalvia: Unionidae). Raffles Bull. Zool. 69, 124–136. https://doi.org/10.26107/RBZ-2021-0011 (2021).Article 

Google Scholar 
25.Jeratthitikul, E., Sucharit, C. & Prasankok, P. Molecular phylogeny of the Indochinese freshwater mussel genus Scabies Haas, 1911 (Bivalvia: Unionidae). Trop. Nat. Hist. 19, 21–36 (2019).
Google Scholar 
26.Jeratthitikul, E., Sutcharit, C., Ngor, P. B. & Prasankok, P. Molecular phylogeny reveals a new genus of freshwater mussels from the Mekong River Basin (Bivalvia: Unionidae). Eur. J. Taxon. 775, 119–142. https://doi.org/10.5852/ejt.2021.775.1553 (2021).Article 

Google Scholar 
27.Pfeiffer, J. M., Graf, D. L., Cummings, K. S. & Page, L. M. Taxonomic revision of a radiation of South-East Asian freshwater mussels (Unionidae: Gonideinae: Contradentini+ Rectidentini). Invertebr. Syst. 35, 394–470. https://doi.org/10.1071/IS20044 (2021).Article 

Google Scholar 
28.Zieritz, A. et al. A new genus and two new, rare freshwater mussel (Bivalvia: Unionidae) species endemic to Borneo are threatened by ongoing habitat destruction. Aquat. Conserv. https://doi.org/10.1002/aqc.3695 (2021).Article 

Google Scholar 
29.Smith, C. H., Johnson, N. A., Pfeiffer, J. M. & Gangloff, M. M. Molecular and morphological data reveal non-monophyly and speciation in imperiled freshwater mussels (Anodontoides and Strophitus). Mol. Phylogenet. Evol. 119, 50–62. https://doi.org/10.1016/j.ympev.2017.10.018 (2018).Article 
PubMed 

Google Scholar 
30.Inoue, K. et al. A new species of freshwater mussel in the genus Popenaias Frierson, 1927, from Gulf coastal rivers of central Mexico (Bivalvia: Unionida: Unionidae) with comments on the genus. Zootaxa 4816, 457–490. https://doi.org/10.11646/zootaxa.4816.4.3 (2020).Article 

Google Scholar 
31.Ortiz-Sepulveda, C. M. et al. Diversification dynamics of freshwater bivalves (Unionidae: Parreysiinae: Coelaturini) indicate historic hydrographic connections throughout the East African Rift System. Mol. Phylogenet. Evol. 148, 106816. https://doi.org/10.1016/j.ympev.2020.106816 (2020).Article 
PubMed 

Google Scholar 
32.Tomilova, A. A. et al. An endemic freshwater mussel species from the Orontes River basin in Turkey and Syria represents duck mussel’s intraspecific lineage: Implications for conservation. Limnologica 84, 125811. https://doi.org/10.1016/j.limno.2020.125811 (2020).CAS 
Article 

Google Scholar 
33.Tomilova, A. A. et al. Evidence for plio-pleistocene duck mussel refugia in the Azov Sea river basins. Diversity 12, 118. https://doi.org/10.3390/d12030118 (2020).Article 

Google Scholar 
34.Pfeiffer, J. M., Sharpe, A. E., Johnson, N. A., Emery, K. F. & Page, L. M. Molecular phylogeny of the Nearctic and Mesoamerican freshwater mussel genus Megalonaias. Hydrobiologia 811, 139–151. https://doi.org/10.1007/s10750-017-3441-7 (2018).CAS 
Article 

Google Scholar 
35.Bolotov, I. N. et al. A new genus and tribe of freshwater mussel (Unionidae) from Southeast Asia. Sci. Rep. 8, 10030. https://doi.org/10.1038/s41598-018-28385-y (2018).ADS 
CAS 
Article 
PubMed 
PubMed Central 

Google Scholar 
36.Konopleva, E. S. et al. New freshwater mussels from two Southeast Asian genera Bineurus and Thaiconcha (Pseudodontini, Gonideinae, Unionidae). Sci. Rep. 11, 8244. https://doi.org/10.1038/s41598-021-87633-w (2021).ADS 
CAS 
Article 
PubMed 
PubMed Central 

Google Scholar 
37.Lopes-Lima, M. et al. Freshwater mussels (Bivalvia: Unionidae) from the Rising Sun (Far East Asia): Phylogeny, systematics, and distribution. Mol. Phylogenet. Evol. 146, 106755. https://doi.org/10.1016/j.ympev.2020.106755 (2020).Article 
PubMed 

Google Scholar 
38.Rangin, C. Active and recent tectonics of the Burma Platelet in Myanmar. Geol. Soc. Lond. Mem. 48, 53–64. https://doi.org/10.1144/M48.3 (2017).Article 

Google Scholar 
39.Licht, A. et al. Magmatic history of central Myanmar and implications for the evolution of the Burma Terrane. Gondwana Res. 87, 303–319. https://doi.org/10.1016/j.gr.2020.06.016 (2020).ADS 
CAS 
Article 

Google Scholar 
40.Westerweel, J. et al. Burma Terrane part of the Trans-Tethyan arc during collision with India according to palaeomagnetic data. Nat. Geosci. 12, 863–868. https://doi.org/10.1038/s41561-019-0443-2 (2019).ADS 
CAS 
Article 
PubMed 
PubMed Central 

Google Scholar 
41.Morley, C. K., Chantraprasert, S., Kongchum, J. & Chenoll, K. The West Burma Terrane, a review of recent paleo-latitude data, its geological implications and constraints. Earth Sci. Rev. 220, 103722. https://doi.org/10.1016/j.earscirev.2021.103722 (2021).Article 

Google Scholar 
42.Martin, C. R. et al. Paleocene latitude of the Kohistan-Ladakh arc indicates multistage India-Eurasia collision. Proc. Natl. Acad. Sci. USA 117, 29487–29494. https://doi.org/10.1073/pnas.2009039117 (2020).ADS 
CAS 
Article 
PubMed 
PubMed Central 

Google Scholar 
43.Frisch, W., Meschede, M. & Blakey, R. C. Plate Tectonics: Continental Drift and Mountain Building (Springer Science & Business Media, 2010).
Google Scholar 
44.Ali, J. R. & Aitchison, J. C. Gondwana to Asia: Plate tectonics, paleogeography and the biological connectivity of the Indian sub-continent from the Middle Jurassic through latest Eocene (166–35 Ma). Earth Sci. Rev. 88, 145–166. https://doi.org/10.1016/j.earscirev.2008.01.007 (2008).ADS 
Article 

Google Scholar 
45.Chatterjee, S., Goswami, A. & Scotese, C. R. The longest voyage: Tectonic, magmatic, and paleoclimatic evolution of the Indian plate during its northward flight from Gondwana to Asia. Gondwana Res. 23, 238–267. https://doi.org/10.1016/j.gr.2012.07.001 (2013).ADS 
Article 

Google Scholar 
46.van Hinsbergen, D. et al. Greater India Basin hypothesis and a two-stage Cenozoic collision between India and Asia. Proc. Natl. Acad. Sci. USA 109, 7659–7664. https://doi.org/10.1073/pnas.1117262109 (2012).ADS 
Article 
PubMed 
PubMed Central 

Google Scholar 
47.van Hinsbergen, D. J. et al. Reconstructing Greater India: Paleogeographic, kinematic, and geodynamic perspectives. Tectonophysics 760, 69–94. https://doi.org/10.1016/j.tecto.2018.04.006 (2019).ADS 
Article 

Google Scholar 
48.Morley, C. K., Naing, T. T., Searle, M. & Robinson, S. A. Structural and tectonic development of the Indo-Burma ranges. Earth Sci. Rev. 200, 102992. https://doi.org/10.1016/j.earscirev.2019.102992 (2020).Article 

Google Scholar 
49.Poinar, G. Jr. Burmese amber: Evidence of Gondwanan origin and Cretaceous dispersion. Hist. Biol. 31, 1304–1309. https://doi.org/10.1080/08912963.2018.1446531 (2019).Article 

Google Scholar 
50.Zhang, X. et al. Tracing Argoland in eastern Tethys and implications for India-Asia convergence. GSA Bull. 133, 1712–1722. https://doi.org/10.1130/B35772.1 (2021).CAS 
Article 

Google Scholar 
51.Pfeiffer, J. M., Graf, D. L., Cummings, K. S. & Page, L. M. Molecular phylogeny and taxonomic revision of two enigmatic freshwater mussel genera (Bivalvia: Unionidae incertae sedis: Harmandia and Unionetta) reveals a diverse clade of Southeast Asian Parreysiinae. J. Molluscan Stud. 84, 404–416. https://doi.org/10.1093/mollus/eyy028 (2018).Article 

Google Scholar 
52.Whelan, N. V., Geneva, A. J. & Graf, D. L. Molecular phylogenetic analysis of tropical freshwater mussels (Mollusca: Bivalvia: Unionoida) resolves the position of Coelatura and supports a monophyletic Unionidae. Mol. Phylogenet. Evol. 61, 504–514. https://doi.org/10.1016/j.ympev.2011.07.016 (2011).Article 
PubMed 

Google Scholar 
53.Konopleva, E. S. et al. A new genus and two new species of freshwater mussels (Unionidae) from Western Indochina. Sci. Rep. 9, 4106. https://doi.org/10.1038/s41598-019-39365-1 (2019).ADS 
CAS 
Article 
PubMed 
PubMed Central 

Google Scholar 
54.Muanta, S., Jeratthitikul, E., Panha, S. & Prasankok, P. Phylogeography of the freshwater bivalve genus Ensidens (Unionidae) in Thailand. J. Molluscan Stud. 85, 224–231. https://doi.org/10.1093/mollus/eyz013 (2019).Article 

Google Scholar 
55.Zieritz, A. et al. Factors driving changes in freshwater mussel (Bivalvia, Unionida) diversity and distribution in Peninsular Malaysia. Sci. Total Environ. 571, 1069–1078. https://doi.org/10.1016/j.scitotenv.2016.07.098 (2016).ADS 
CAS 
Article 
PubMed 

Google Scholar 
56.Bolotov, I. N. et al. New taxa of freshwater mussels (Unionidae) from a species-rich but overlooked evolutionary hotspot in Southeast Asia. Sci. Rep. 7, 11573. https://doi.org/10.1038/s41598-017-11957-9 (2017).ADS 
CAS 
Article 
PubMed 
PubMed Central 

Google Scholar 
57.Subba Rao, N. V. Handbook. Freshwater Molluscs of India (Zoological Survey of India, 1989).58.Ramakrishna & Dey, A. Handbook on Indian Freshwater Molluscs (Zoological Survey of India, 2007).59.Prashad, B. The marsupium and glochidium of some Unionidae and on the Indian species hitherto assigned to the genus Nodularia. Rec. Indian Mus. 15, 143–148 (1918).
Google Scholar 
60.Burdi, G. H., Baloch, W. A., Begum, F., Soomro, A. N. & Khuhawar, M. Y. Ecological studies on freshwater bivalve mussels (Pelecypoda) of Indus River and its canals at Kotri Barrage Sindh, Pakistan. Sindh Univ. Res. J. 41, 31–36 (2009).
Google Scholar 
61.Nesemann, H. et al. Aquatic Invertebrates of the Ganga River System: Volume 1—Mollusca, Annelida, Crustacea (in part) (Hasko Nesemann and Chandi Press, 2007).62.Budha, P. B. A Field Guide to Freshwater Molluscs of Kailali, Far Western Nepal (Central Department of Zoology, Tribhuvan University, 2016).
Google Scholar 
63.Gittenberger, E., Leda, P., Gyeltshen, C. & Sherub, S. Distributional patterns of molluscan taxa in Bhutan (Mollusca). Biodiversität Naturausstattung Himalaya 4, 143–151 (2018).
Google Scholar 
64.Nanda, A. C., Sehgal, R. K. & Chauhan, P. R. Siwalik-age faunas from the Himalayan foreland Basin of South Asia. J. Asian Earth Sci. 162, 54–68. https://doi.org/10.1016/j.jseaes.2017.10.035 (2018).ADS 
Article 

Google Scholar 
65.Vredenburg, E. & Prashad, B. Unionidae from the Miocene of Burma. Rec. Geol. Surv. India 51, 371–374 (1921).
Google Scholar 
66.Prashad, B. On some Fossil Indian Unionidae. Rec. Geol. Surv. India 60, 308–312 (1928).
Google Scholar 
67.Modell, H. Paläontologische und geologische Untersuchungen im Tertiär von Pakistan. 4. Die tertiären Najaden des Punjab und Vorderindiens. Abhandlungen der Bayerischen Akademie der Wissenschaften, Mathematisch-naturwissenschaftliche Klasse, neue Folge 135, 1–49 (1969).68.Takayasu, K., Gurung, D. D. & Matsuoka, K. Some new species of freshwater bivalves from the Mio-Pliocene Churia Group, west-central Nepal. Trans. Proc. Paleontol. Soc. Jpn. New Ser. 179, 157–168. https://doi.org/10.14825/prpsj1951.1995.179_157 (1995).Article 

Google Scholar 
69.Gurung, D. Freshwater molluscs from the Late Neogene Siwalik Group, Surai Khola, western Nepal. J. Nepal Geol. Soc. 17, 7–28. https://doi.org/10.3126/jngs.v17i0.32095 (1998).Article 

Google Scholar 
70.Simpson, C. T. Synopsis of the naiades, or pearly fresh-water mussels. Proc. U.S. Natl. Mus. 22, 501–1044 (1900).
Google Scholar 
71.Madhyastha, N. A. & Mumbrekar, K. D. Two endemic genera of bivalves in the Tunga River of the Western Ghats, Karnataka, India. Tentacle 14, 23–24 (2006).
Google Scholar 
72.Prashad, B. Notes on lamellibranchs in the Indian Museum. Rec. Indian Mus. 19, 165–173 (1920).
Google Scholar 
73.Haas, F. Eine neude indische Najade, Trapezoideus prashadi. Senckenbergiana 4, 101–102 (1922).
Google Scholar 
74.Sowerby, G. B. Genus Unio. Conchologica Iconica 16, pls. 1, 61–96 (1868).75.Haas, F. Die Unioniden. H.C. Küster, Systematisches Conchylien-Cabinet von Martini und Chemnitz 9, 257–288 (1919).76.Hadl, G. Results of the Austrian-Ceylonese Hydrobiological Mission 1970 of the 1st Zoological Institute of the University of Vienna (Austria) and the Department of Zoology of the Vidyalankara University of Ceylon, Kelaniya. Part XVIII: Freshwater Mussels Bivalvia. Bull. Fish. Res. Stn. Sri Lanka (Ceylon) 25, 183–188 (1974).
Google Scholar 
77.Gittenberger et al. A Field Guide to the Common Molluscs of Bhutan (National Biodiversity Centre (NBC), Ministry of Agriculture and Forests, 2017).78.Annandale, N. & Prashad, B. The Mollusca of the inland waters of Baluchistan and of Seistan. Rec. Indian Mus. 18, 17–62 (1919).
Google Scholar 
79.Simpson, C. T. A Descriptive Catalogue of the Naiades, or Pearly Fresh-Water Mussels. Parts I-III (Bryant Walker, 1914).
Google Scholar 
80.Mörch, O. A. L. On the land and fresh-water Mollusca of Greenland. Am. J. Conchol. 4, 25–40 (1868).
Google Scholar 
81.Schröter, J. S. Die Geschichte der Flussconchylien: Mit vorzüglicher Rücksicht auf Diejenigen Welche in den Thüringischen Wassern Leben (Halle, bey Johann Jacob Gebauer, 1779).82.Spengler, L. Om Slaegterne Chaena Mya og Unio. Skrivter Naturhistorie-Selskabet 3, 16–69 (1993).
Google Scholar 
83.Haas, F. Bemerkungen über Spenglers Unionen. Videnskabelige Meddelelser fra Dansk naturhistorisk Forening i Kjøbenhav 65, 51–66 (1913).
Google Scholar 
84.Haas, F. Superfamilia Unionacea. Das Tierreich 88, 1–663 (1969).
Google Scholar 
85.Prashad, B. On some undescribed freshwater Molluscs from various parts of India and Burma. Rec. Geol. Surv. India 62, 428–433 (1930).
Google Scholar 
86.Conrad, T. A. A synopsis of the family of Naïades of North America, with notes, and a table of some of the genera and sub-genera of the family, according to their geographical distribution, and descriptions of genera and sub-genera. Proc. Acad. Natl. Sci. Phila. 6, 243–269 (1853).
Google Scholar 
87.Sowerby, G. B. Genus Unio. Conchol. Iconica 16, 31–54 (1866).
Google Scholar 
88.Frierson, L. S. A Classified and Annotated Check List of the North American Naiades (Baylor University Press, 1927).
Google Scholar 
89.Prashad, B. Studies on the anatomy of Indian Mollusca. The soft parts of some Indian Unionidae. Rec. Indian Mus. 16, 289–296 (1919).
Google Scholar 
90.Annandale, N. Further note on the burrows of Solenaia soleniformis. Rec. Indian Mus. 16, 205–206 (1919).
Google Scholar 
91.Godwin-Austen, H. H. Description of a new species of Margaritanopsis (Unionidae) from the Southern Shan States, with notes on Solenaia soleniformis. Rec. Indian Mus. 16, 203–205 (1919).
Google Scholar 
92.Pfeiffer, J. M., Breinholt, J. W. & Page, L. M. Unioverse: A phylogenetic resource for reconstructing the evolution of freshwater mussels (Bivalvia, Unionoida). Mol. Phylogenet. Evol. 137, 114–126. https://doi.org/10.1016/j.ympev.2019.02.016 (2019).Article 
PubMed 

Google Scholar 
93.Huang, X.-C. et al. Towards a global phylogeny of freshwater mussels (Bivalivia: Unionida): Species delimitation of Chinese taxa, mitochondrial phylogenomics, and diversification patterns. Mol. Phylogenet. Evol. 130, 45–59. https://doi.org/10.1016/j.ympev.2018.09.019 (2019).Article 
PubMed 

Google Scholar 
94.Bolotov, I. N., Kondakov, A. V., Konopleva, E. S. & Vikhrev, I. V. A new genus of ultra-elongate freshwater mussels from Vietnam and eastern China (Bivalvia: Unionidae). Ecol. Montenegrina 39, 1–6. https://doi.org/10.37828/em.2021.39.1 (2021).Article 

Google Scholar 
95.Pfeiffer, J. M. & Graf, D. L. Evolution of bilaterally asymmetrical larvae in freshwater mussels (Bivalvia: Unionoida: Unionidae). Zool. J. Linn. Soc. 175, 307–318. https://doi.org/10.1111/zoj.12282 (2015).Article 

Google Scholar 
96.Rafinesque, C. S. Continuation of a Monograph of the Bivalve Shells of the River Ohio and Other Rivers of the Western States. By Prof. C.S. Rafinesque. (Published at Brussels, September, 1820). Containing 46 species, from No. 76 to no. 121. Including an Appendix on Some Bivalve Shells of the Rivers of Hindostan, with a Supplement on the Fossil Bivalves of the Western States, and the Tulosites, A New Genus of Fossils (1831).97.Blanford, W. T. Contributions to Indian Malacology no VII. List of species of Unio and Anodonta described as occurring in India, Ceylon and Burma. J. Asiat. Soc. Bengal 35, 134–155 (1866).
Google Scholar 
98.Frierson, L. S. Remarks on classification of the Unionidae. Nautilus 28, 6–8 (1914).
Google Scholar 
99.Johnson, R. I. The types of Unionidae (Mollusca: Bivalvia) described by C. S. Rafinesque in the Museum national d’Histoire naturelle, Paris. J. Conchyliol. 110, 35–37 (1973).
Google Scholar 
100.Vanatta, E. G. Rafinesque’s types of Unio. Proc. Acad. Natl. Sci. Phila. 67, 549–559 (1915).
Google Scholar 
101.Baker, H. B. Some of Rafinesque’s unionid names. The Nautilus 77, 140–142 (1964).
Google Scholar 
102.Williams, J. D., Bogan, A. E. & Garner, J. T. Freshwater mussels of Alabama and the Mobile Basin in Georgia, Mississippi and Tennessee (University of Alabama Press, 2008).
Google Scholar 
103.Bogan, A. E. A resolution of the nomenclatural confusion surrounding Plagiola Rafinesque, Epioblasma Rafinesque, and Dysnomia Agassiz (Mollusca: Bivalvia: Unionidae). Malacol. Rev. 30, 77–86 (1997).
Google Scholar 
104.Graf, D. L. & Cummings, K. S. Palaeoheterodont diversity (Mollusca: Trigonioida+ Unionoida): What we know and what we wish we knew about freshwater mussel evolution. Zool. J. Linn. Soc. 148, 343–394. https://doi.org/10.1111/j.1096-3642.2006.00259.x (2006).Article 

Google Scholar 
105.Modell, H. Das natlirliche System der Najaden. Arch. Molluskenkunde 74, 161–191 (1942).
Google Scholar 
106.Starobogatov, Y. I. Fauna of Molluscs and Zoogeographic Division of Continental Waterbodies of the Globe (Nauka, 1970).
Google Scholar 
107.Bolotov, I. N. et al. Discovery of Novaculina myanmarensis sp. nov. (Bivalvia: Pharidae: Pharellinae) closes the freshwater razor clams range disjunction in Southeast Asia. Sci. Rep. 8, 16325. https://doi.org/10.1038/s41598-018-34491-8 (2018).ADS 
CAS 
Article 
PubMed 
PubMed Central 

Google Scholar 
108.Than, W. et al. Phylogeography and distribution of the freshwater razor clams Novaculina myanmarensis and N. gangetica in Myanmar, with notes on two doubtful nominal taxa described as Novaculina members (Bivalvia: Pharidae). Ecol. Montenegrina 40, 59–67. https://doi.org/10.37828/em.2021.40.4 (2021).Article 

Google Scholar 
109.Haas, F. Beiträge zu einer Monographie der asiatischen Unioniden. Abhandlungen der Senckenbergischen Naturforschenden Gesellschaft 38, 129–203 (1924).
Google Scholar 
110.Preston, H. B. Mollusca (Freshwater Gastropoda & Pelecypoda). Fauna of British India, including Ceylon and Burma (Taylor and Francis, 1915).
Google Scholar 
111.Prashad, B. A revision of the Burmese Unionidae. Rec. Indian Mus. 24, 91–111 (1922).
Google Scholar 
112.Theobald, W. Catalogue of the Recent Shells in the Museum of the Asiatic Society (Bengal Military Orphan Press, 1860).
Google Scholar 
113.Zieritz, A. et al. Diversity, biogeography and conservation of freshwater mussels (Bivalvia: Unionida) in East and Southeast Asia. Hydrobiologia 810, 29–44. https://doi.org/10.1007/s10750-017-3104-8 (2018).Article 

Google Scholar 
114.Konopleva, E. S. et al. A taxonomic review of Trapezidens (Bivalvia: Unionidae: Lamellidentini), a freshwater mussel genus endemic to Myanmar, with a description of a new species. Ecol. Montenegrina 27, 45–57. https://doi.org/10.37828/em.2020.27.6 (2020).Article 

Google Scholar 
115.Brandt, R. A. M. The non-marine aquatic mollusca of Thailand. Arch. Mollusckenkunde 105, 1–423 (1974).
Google Scholar 
116.Neumayr, M. Süsswasser-Mollusken. Die wissenschaftlichen ergebnisse der reise des grafen Béla Széchenyi in Ostasien 1877–1880(2), 637–662 (1899).
Google Scholar 
117.Tripathy, B. & Mukhopadhayay, A. Freshwater molluscs of India: An insight of into their diversity, distribution and conservation. In Aquatic Ecosystem: Biodiversity, Ecology and Conservation (eds Rawat, M. et al.) 163–195 (Springer, 2015).
Google Scholar 
118.Prashad, B. VIII—Some Noteworthy Examples of Parallel Evolution in the Molluscan Faunas of South-eastern Asia and South America. Proc. R. Soc. Edinb. 51, 42–53. https://doi.org/10.1017/s0370164600022987 (1932).Article 

Google Scholar 
119.Smith, E. A. Description of Mulleria dalyi, n. sp., from India. Proc. Malacol. Soc. Lond. 3, 14–16 (1898).
Google Scholar 
120.Bogan, A. E. & Hoeh, W. R. On becoming cemented: Evolutionary relationships among the genera in the freshwater bivalve family Etheriidae (Bivalvia: Unionoida). Geol. Soc. Lond. Spec. Publ. 177, 159–168. https://doi.org/10.1144/GSL.SP.2000.177.01.09 (2000).ADS 
Article 

Google Scholar 
121.Bogan, A. E. & Roe, K. J. Freshwater bivalve (Unioniformes) diversity, systematics, and evolution: Status and future directions. J. N. Am. Benthol. Soc. 27, 349–369. https://doi.org/10.1899/07-069.1 (2008).Article 

Google Scholar 
122.Hoeh, W. R., Bogan, A. E., Heard, W. H. & Chapman, E. G. Palaeoheterodont phylogeny, character evolution, diversity and phylogenetic classification: A reflection on methods of analysis. Malacologia 51, 307–317. https://doi.org/10.4002/040.051.0206 (2009).Article 

Google Scholar 
123.Woodward, M. F. On the anatomy of Mulleria dalyi, Smth. J. Molluscan Stud. 3, 87–91. https://doi.org/10.1093/oxfordjournals.mollus.a065152 (1898).Article 

Google Scholar 
124.Aravind, N. A. et al. The status and distribution of freshwater molluscs of the Western Ghats. In The Status and Distribution of Freshwater Biodiversity in the Western Ghats, India (eds Molur, S. et al.) 21–42 (IUCN and Zoo Outreach Organisation, 2011).
Google Scholar 
125.Madhyastha, N. A. Pseudomulleria dalyi (Acostea dalyi): A rare cemented bivalve of Western Ghats. Zoos’ Print J. 16, 573 (2001).
Google Scholar 
126.Loria, S. F. & Prendini, L. Out of India, thrice: Diversification of Asian forest scorpions reveals three colonizations of Southeast Asia. Sci. Rep. 10, 22301. https://doi.org/10.1038/s41598-020-78183-8 (2020).ADS 
CAS 
Article 
PubMed 
PubMed Central 

Google Scholar 
127.Köhler, F. & Glaubrecht, M. Out of Asia and into India: On the molecular phylogeny and biogeography of the endemic freshwater gastropod Paracrostoma Cossmann, 1900 (Caenogastropoda: Pachychilidae). Biol. J. Lin. Soc. 91, 627–651. https://doi.org/10.1111/j.1095-8312.2007.00866.x (2007).Article 

Google Scholar 
128.Dahanukar, N., Raut, R. & Bhat, A. Distribution, endemism and threat status of freshwater fishes in the Western Ghats of India. J. Biogeogr. 31, 123–136. https://doi.org/10.1046/j.0305-0270.2003.01016.x (2004).Article 

Google Scholar 
129.Britz, R. et al. Aenigmachannidae, a new family of snakehead fishes (Teleostei: Channoidei) from subterranean waters of South India. Sci. Rep. 10, 16081. https://doi.org/10.1038/s41598-020-73129-6 (2020).ADS 
CAS 
Article 
PubMed 
PubMed Central 

Google Scholar 
130.Hedges, S. B. The coelacanth of frogs. Nature 425, 669–670. https://doi.org/10.1038/425669a (2003).ADS 
CAS 
Article 
PubMed 

Google Scholar 
131.Dutta, S. K., Vasudevan, K., Chaitra, M. S., Shanker, K. & Aggarwal, R. K. Jurassic frogs and the evolution of amphibian endemism in the Western Ghats. Curr. Sci. 86, 211–216 (2004).CAS 

Google Scholar 
132.Roelants, K., Jiang, J. & Bossuyt, F. Endemic ranid (Amphibia: Anura) genera in southern mountain ranges of the Indian subcontinent represent ancient frog lineages: Evidence from molecular data. Mol. Phylogenet. Evol. 31, 730–740. https://doi.org/10.1016/j.ympev.2003.09.011 (2004).CAS 
Article 
PubMed 

Google Scholar 
133.Van Bocxlaer, I. et al. Mountain-associated clade endemism in an ancient frog family (Nyctibatrachidae) on the Indian subcontinent. Mol. Phylogenet. Evol. 62, 839–847. https://doi.org/10.1016/j.ympev.2011.11.027 (2012).Article 
PubMed 

Google Scholar 
134.Krishnan, R. M. & Ramesh, B. R. Endemism and sexual systems in the evergreen tree flora of the Western Ghats, India. Divers. Distrib. 11, 559–565. https://doi.org/10.1111/j.1366-9516.2005.00190.x (2005).Article 

Google Scholar 
135.Mörch, O. A. L. Catalogue des Mollusques terrestres et fluviatiles des anciennes colonies du golfe du Bengale. J. Conchyliol. 20, 303–345 (1872).
Google Scholar 
136.Graf, D. L. & Cummings, K. S. Freshwater mussel (Mollusca: Bivalvia: Unionoida) richness and endemism in the ecoregions of Africa and Madagascar based on comprehensive museum sampling. Hydrobiologia 678, 17–36. https://doi.org/10.1007/s10750-011-0810-5 (2011).Article 

Google Scholar 
137.Li, Z. et al. Kinematic evolution of the West Burma block during and after India-Asia collision revealed by paleomagnetism. J. Geodyn. 134, 101690. https://doi.org/10.1016/j.jog.2019.101690 (2020).Article 

Google Scholar 
138.Van Damme, D., Bogan, A. E. & Dierick, M. A revision of the Mesozoic naiads (Unionoida) of Africa and the biogeographic implications. Earth Sci. Rev. 147, 141–200. https://doi.org/10.1016/j.earscirev.2015.04.011 (2015).ADS 
Article 

Google Scholar 
139.Hall, R. Late Jurassic-Cenozoic reconstructions of the Indonesian region and the Indian Ocean. Tectonophysics 570, 1–41. https://doi.org/10.1016/j.tecto.2012.04.021 (2012).ADS 
Article 

Google Scholar 
140.Bosworth, W. Mesozoic and early Tertiary rift tectonics in East Africa. Tectonophysics 209, 115–137. https://doi.org/10.1016/0040-1951(92)90014-W (1992).ADS 
Article 

Google Scholar 
141.Guiraud, R., Bosworth, W., Thierry, J. & Delplanque, A. Phanerozoic geological evolution of Northern and Central Africa: An overview. J. Afr. Earth Sci. 43, 83–143. https://doi.org/10.1016/j.jafrearsci.2005.07.017 (2005).ADS 
Article 

Google Scholar 
142.Wilson, M. & Guiraud, R. Magmatism and rifting in Western and Central Africa, from Late Jurassic to Recent times. Tectonophysics 213, 203–225 (1992).ADS 

Google Scholar 
143.Chatterjee, S., Scotese, C. R. & Bajpai, S. Indian Plate and Its Epic Voyage from Gondwana to Asia: Its Tectonic, Paleoclimatic, and Paleobiogeographic Evolution (Special Paper 529, The Geological Society of America, 2017).144.Briggs, J. C. The biogeographic and tectonic history of India. J. Biogeogr. 30, 381–388. https://doi.org/10.1046/j.1365-2699.2003.00809.x (2003).Article 

Google Scholar 
145.Hartman, J. H., Erickson, D. N. & Bakken, A. Stephen Hislop and his 1860 Cretaceous continental molluscan new species descriptions in Latin from the Deccan Plateau, India. Palaeontology 51, 1225–1252. https://doi.org/10.1111/j.1475-4983.2008.00807.x (2008).Article 

Google Scholar 
146.Vandamme, D., Courtillot, V., Besse, J. & Montigny, R. Paleomagnetism and age determinations of the Deccan Traps (India): Results of a Nagpur-Bombay Traverse and review of earlier work. Rev. Geophys. 29, 159–190. https://doi.org/10.1029/91RG00218 (1991).ADS 
Article 

Google Scholar 
147.Bolotov, I. N. et al. Multi-locus fossil-calibrated phylogeny, biogeography and a subgeneric revision of the Margaritiferidae (Mollusca: Bivalvia: Unionoida). Mol. Phylogenet. Evol. 103, 104–121. https://doi.org/10.1016/j.ympev.2016.07.020 (2016).Article 
PubMed 

Google Scholar 
148.Lyubas, A. A. et al. A taxonomic revision of fossil freshwater pearl mussels (Bivalvia: Unionoida: Margaritiferidae) from Pliocene and Pleistocene deposits of Southeastern Europe. Ecol. Montenegrina 21, 1–16. https://doi.org/10.37828/em.2019.21.1 (2019).Article 

Google Scholar 
149.Campbell, D. C. et al. Phylogeny of North American amblemines (Bivalvia, Unionoida): Prodigious polyphyly proves pervasive across genera. Invertebr. Biol. 124, 131–164 (2005).
Google Scholar 
150.Lopes-Lima, M. et al. Revisiting the North American freshwater mussel genus Quadrula sensu lato (Bivalvia: Unionidae): Phylogeny, taxonomy and species delineation. Zool. Scr. 48, 313–336. https://doi.org/10.1111/zsc.12344 (2019).Article 

Google Scholar 
151.Aksenova, O. V. et al. Species richness, molecular taxonomy and biogeography of the radicine pond snails (Gastropoda: Lymnaeidae) in the Old World. Sci. Rep. 8, 11199. https://doi.org/10.1038/s41598-018-29451-1 (2018).ADS 
CAS 
Article 
PubMed 
PubMed Central 

Google Scholar 
152.Kosuch, J., Vences, M., Dubois, A., Ohler, A. & Böhme, W. Out of Asia: Mitochondrial DNA evidence for an oriental origin of tiger frogs, genus Hoplobatrachus. Mol. Phylogenet. Evol. 21, 398–407. https://doi.org/10.1006/mpev.2001.1034 (2001).CAS 
Article 
PubMed 

Google Scholar 
153.Sil, M., Aravind, N. A. & Karanth, K. P. Into-India or out-of-India? Historical biogeography of the freshwater gastropod genus Pila (Caenogastropoda: Ampullariidae). Biol. J. Lin. Soc. 129, 752–764. https://doi.org/10.1093/biolinnean/blz171 (2020).Article 

Google Scholar 
154.Sil, M., Aravind, N. A. & Karanth, K. P. Role of geography and climatic oscillations in governing into-India dispersal of freshwater snails of the family: Viviparidae. Mol. Phylogenet. Evol. 138, 174–181. https://doi.org/10.1016/j.ympev.2019.05.027 (2019).Article 
PubMed 

Google Scholar 
155.Garg, S. & Biju, S. D. New microhylid frog genus from Peninsular India with Southeast Asian affinity suggests multiple Cenozoic biotic exchanges between India and Eurasia. Sci. Rep. 9, 1906. https://doi.org/10.1038/s41598-018-38133-x (2019).ADS 
CAS 
Article 
PubMed 
PubMed Central 

Google Scholar 
156.Gorin, V. A. et al. A little frog leaps a long way: Compounded colonizations of the Indian Subcontinent discovered in the tiny Oriental frog genus Microhyla (Amphibia: Microhylidae). PeerJ 8, e9411. https://doi.org/10.7717/peerj.9411 (2020).Article 
PubMed 
PubMed Central 

Google Scholar 
157.Karanth, K. P. An island called India: Phylogenetic patterns across multiple taxonomic groups reveal endemic radiations. Curr. Sci. 108, 1847–1851 (2015).
Google Scholar 
158.Karanth, K. P. Out-of-India Gondwanan origin of some tropical Asian biota. Curr. Sci. 90, 789–792 (2006).
Google Scholar 
159.Datta-Roy, A. & Karanth, K. P. The Out-of-India hypothesis: What do molecules suggest?. J. Biosci. 34, 687–697. https://doi.org/10.1007/s12038-009-0057-8 (2009).Article 
PubMed 

Google Scholar 
160.Gower, D. J. et al. A molecular phylogeny of ichthyophiid caecilians (Amphibia: Gymnophiona: Ichthyophiidae): Out of India or out of South East Asia?. Proc. R. Soc. Lond. B 269, 1563–1569. https://doi.org/10.1098/rspb.2002.2050 (2002).CAS 
Article 

Google Scholar 
161.Kamei, R. G. et al. Discovery of a new family of amphibians from northeast India with ancient links to Africa. Proc. R. Soc. B 279, 2396–2401. https://doi.org/10.1098/rspb.2012.0150 (2012).Article 
PubMed 
PubMed Central 

Google Scholar 
162.Yamahira, K. et al. Mesozoic origin and ‘out-of-India’radiation of ricefishes (Adrianichthyidae). Biol. Let. 17, 20210212. https://doi.org/10.1098/rsbl.2021.0212 (2021).Article 

Google Scholar 
163.Klaus, S., Schubart, C. D., Streit, B. & Pfenninger, M. When Indian crabs were not yet Asian-biogeographic evidence for Eocene proximity of India and Southeast Asia. BMC Evol. Biol. 10, 287. https://doi.org/10.1186/1471-2148-10-287 (2010).Article 
PubMed 
PubMed Central 

Google Scholar 
164.Joshi, J., Karanth, P. K. & Edgecombe, G. D. The out-of-India hypothesis: Evidence from an ancient centipede genus, Rhysida (Chilopoda: Scolopendromorpha) from the Oriental Region, and systematics of Indian species. Zool. J. Linn. Soc. 189, 828–861. https://doi.org/10.1093/zoolinnean/zlz138 (2020).Article 

Google Scholar 
165.Foley, S., Krehenwinkel, H., Cheng, D. Q. & Piel, W. H. Phylogenomic analyses reveal a Gondwanan origin and repeated out of India colonizations into Asia by tarantulas (Araneae: Theraphosidae). PeerJ 9, e11162. https://doi.org/10.7717/peerj.11162 (2021).Article 
PubMed 
PubMed Central 

Google Scholar 
166.Dayanandan, S., Ashton, P. S., Williams, S. M. & Primack, R. B. Phylogeny of the tropical tree family Dipterocarpaceae based on nucleotide sequences of the chloroplast rbcL gene. Am. J. Bot. 86, 1182–1190 (1999).CAS 
PubMed 

Google Scholar 
167.Conti, E., Eriksson, T., Schönenberger, J., Sytsma, K. J. & Baum, D. A. Early Tertiary out-of-India dispersal of Crypteroniaceae: Evidence from phylogeny and molecular dating. Evolution 56, 1931–1942. https://doi.org/10.1111/j.0014-3820.2002.tb00119.x (2002).Article 
PubMed 

Google Scholar 
168.Chen, J. et al. Eurypterogerron kachinensis gen et sp nov, a remarkable minlagerrontid (Hemiptera, Cicadomorpha) in mid-Cretaceous Burmese amber. Cretaceous Res. 110, 104418. https://doi.org/10.1016/j.cretres.2020.104418 (2020).Article 

Google Scholar 
169.Rasnitsyn, A. P. & Öhm-Kühnle, C. Three new female Aptenoperissus from mid-Cretaceous Burmese amber (Hymenoptera, Stephanoidea, Aptenoperissidae): Unexpected diversity of paradoxical wasps suggests insular features of source biome. Cretac. Res. 91, 168–175. https://doi.org/10.1016/j.cretres.2018.06.004 (2018).Article 

Google Scholar 
170.Zhang, Q., Rasnitsyn, A. P., Wang, B. & Zhang, H. Hymenoptera (wasps, bees and ants) in mid-Cretaceous Burmese amber: A review of the fauna. Proc. Geol. Assoc. 129, 736–747. https://doi.org/10.1016/j.pgeola.2018.06.004 (2018).Article 

Google Scholar 
171.Bolotov, I. N. et al. A new fossil piddock (Bivalvia: Pholadidae) may indicate estuarine to freshwater environments near Cretaceous amber-producing forests in Myanmar. Sci. Rep. 11, 6646. https://doi.org/10.1038/s41598-021-86241-y (2021).ADS 
CAS 
Article 
PubMed 
PubMed Central 

Google Scholar 
172.Balashov, I. A., Perkovsky, E. E. & Vasilenko, D. V. A mid-Cretaceous land snail Burminella artiukhini gen. et. sp. nov. from Burmese amber: A “missing link” between Pupinidae and other Cyclophoroidea? (Caenogastropoda). Cretaceous Res. 118, 104941. https://doi.org/10.1016/j.cretres.2021.104941 (2021).Article 

Google Scholar 
173.Balashov, I. An inventory of molluscs recorded from mid-Cretaceous Burmese amber, with the description of a land snail, Euthema annae sp. nov. (Caenogastropoda, Cyclophoroidea, Diplommatinidae). Cretaceous Res. 118, 104676. https://doi.org/10.1016/j.cretres.2020.104676 (2021).Article 

Google Scholar 
174.Yu, T., Neubauer, T. A. & Jochum, A. First freshwater gastropod preserved in amber suggests long-distance dispersal during the Cretaceous Period. Geol. Mag. 58, 1327–1334. https://doi.org/10.1017/S0016756821000285 (2021).ADS 
Article 

Google Scholar 
175.Bingle-Davis, M. J. Systematics, diversity, and origins of Upper Cretaceous continental molluscan fauna in the infra- and intertrappean strata of the Deccan Plateau, central India (PhD Dissertation) (University of North Dakota, 2012).176.Huang, H. et al. At a crossroads: The late Eocene flora of central Myanmar owes its composition to plate collision and tropical climate. Rev. Palaeobot. Palynol. 291, 104441. https://doi.org/10.1016/j.revpalbo.2021.104441 (2021).Article 

Google Scholar 
177.Westerweel, J. et al. Burma Terrane collision and northward indentation in the Eastern Himalayas recorded in the Eocene-Miocene Chindwin Basin (Myanmar). Tectonics 39, e2020TC006413. https://doi.org/10.1029/2020TC006413 (2020).ADS 
Article 

Google Scholar 
178.Soe, T. T. & Watkinson, I. M. The Sagaing Fault Myanmar. Geol. Soc. 48, 413–441. https://doi.org/10.1144/M48.19 (2017).Article 

Google Scholar 
179.de Sena Oliveira, I. et al. Earliest onychophoran in amber reveals Gondwanan migration patterns. Curr. Biol. 26, 2594–2601. https://doi.org/10.1016/j.cub.2016.07.023 (2016).CAS 
Article 

Google Scholar 
180.Gustafson, L. L. et al. Evaluation of a nonlethal technique for hemolymph collection in Elliptio complanata, a freshwater bivalve (Mollusca: Unionidae). Dis. Aquat. Org. 65, 159–165. https://doi.org/10.3354/dao065159 (2005).Article 

Google Scholar 
181.Jaksch, K., Eschner, A., Rintelen, T. V. & Haring, E. DNA analysis of molluscs from a museum wet collection: A comparison of different extraction methods. BMC. Res. Notes 9, 348. https://doi.org/10.1186/s13104-016-2147-7 (2016).CAS 
Article 
PubMed 
PubMed Central 

Google Scholar 
182.Folmer, O., Black, M., Hoeh, W., Lutz, R. & Vrijenhoek, R. DNA primers for amplification of mitochondrial cytochrome c oxidase subunit I from diverse metazoan invertebrates. Mol. Mar. Biol. Biotechnol. 3, 294–299 (1994).CAS 
PubMed 

Google Scholar 
183.Graf, D. L. Patterns of freshwater bivalve global diversity and the state of phylogenetic studies on the Unionoida, Sphaeriidae, and Cyrenidae. Am. Malacol. Bull. 31, 135–153. https://doi.org/10.4003/006.031.0106 (2013).Article 

Google Scholar 
184.Nguyen, L.-T., Schmidt, H. A., Haeseler, V. A. & Minh, B. Q. IQ-TREE: A fast and effective stochastic algorithm for estimating maximum-likelihood phylogenies. Mol. Biol. Evol. 32, 268–274. https://doi.org/10.1093/molbev/msu300 (2015).CAS 
Article 
PubMed 

Google Scholar 
185.Ronquist, F. et al. MrBayes 3.2: Efficient Bayesian phylogenetic inference and model choice across a large model space. Syst. Biol. 61, 539–542. https://doi.org/10.1093/sysbio/sys029 (2012).Article 
PubMed 
PubMed Central 

Google Scholar 
186.Kalyaanamoorthy, S., Minh, B. Q., Wong, T. K. F., von Haeseler, A. & Jermiin, L. S. ModelFinder: Fast model selection for accurate phylogenetic estimates. Nat. Methods 14, 587–589. https://doi.org/10.1038/nmeth.4285 (2017).CAS 
Article 
PubMed 
PubMed Central 

Google Scholar 
187.Hoang, D. T., Chernomor, O., von Haeseler, A., Minh, B. Q. & Vinh, L. S. UFBoot2: Improving the ultrafast bootstrap approximation. Mol. Biol. Evol. 35, 518–522. https://doi.org/10.1093/molbev/msx281 (2017).CAS 
Article 
PubMed Central 

Google Scholar 
188.Trifinopoulos, J., Nguyen, L. T., von Haeseler, A. & Minh, B. Q. W-IQ-TREE: A fast online phylogenetic tool for maximum likelihood analysis. Nucleic Acids Res. 44, W232–W235. https://doi.org/10.1093/nar/gkw256 (2016).CAS 
Article 
PubMed 
PubMed Central 

Google Scholar 
189.Miller, M., Pfeiffer, W. & Schwartz, T. Creating the CIPRES Science Gateway for inference of large phylogenetic trees. In Gateway Computing Environments Workshop (GCE) 1–8 (IEEE, 2010).190.Kumar, S., Stecher, G. & Tamura, K. MEGA7: Molecular evolutionary genetics analysis version 7.0 for bigger datasets. Mol. Biol. Evol. 33, 1870–1874. https://doi.org/10.1093/molbev/msw054 (2016).CAS 
Article 
PubMed 
PubMed Central 

Google Scholar 
191.Kapli, P. et al. Multi-rate Poisson tree processes for single-locus species delimitation under maximum likelihood and Markov chain Monte Carlo. Bioinformatics 33, 1630–1638. https://doi.org/10.1093/bioinformatics/btx025 (2017).CAS 
Article 
PubMed 
PubMed Central 

Google Scholar 
192.Puillandre, N., Brouillet, S. & Achaz, G. ASAP: Assemble species by automatic partitioning. Mol. Ecol. Resour. 21, 609–620. https://doi.org/10.1111/1755-0998.13281 (2021).Article 
PubMed 

Google Scholar 
193.Villesen, P. FaBox: An online toolbox for fasta sequences. Mol. Ecol. Notes 7, 965–968. https://doi.org/10.1111/j.1471-8286.2007.01821.x (2007).CAS 
Article 

Google Scholar 
194.Bouckaert, R. et al. BEAST 2.5: An advanced software platform for Bayesian evolutionary analysis. PLoS Comput. Biol. 15, 1–28. https://doi.org/10.1371/journal.pcbi.1006650 (2019).CAS 
Article 

Google Scholar 
195.Bouckaert, R. et al. BEAST 2: A software platform for Bayesian evolutionary analysis. PLoS Comput. Biol. 10, e1003537. https://doi.org/10.1371/journal.pcbi.1003537 (2014).CAS 
Article 
PubMed 
PubMed Central 

Google Scholar 
196.Zieritz, A. et al. Mitogenomic phylogeny and fossil-calibrated mutation rates for all F-and M-type mtDNA genes of the largest freshwater mussel family, the Unionidae (Bivalvia). Zool. J. Linn. Soc. 193, 1088–1107. https://doi.org/10.1093/zoolinnean/zlaa153 (2020).Article 

Google Scholar 
197.Froufe, E. et al. Who lives where? Molecular and morphometric analyses clarify which Unio species (Unionida, Mollusca) inhabit the southwestern Palearctic. Org. Divers. Evol. 16, 597–611. https://doi.org/10.1007/s13127-016-0262-x (2016).Article 

Google Scholar 
198.Drummond, A. J., Suchard, M. A., Xie, D. & Rambaut, A. Bayesian phylogenetics with BEAUti and the BEAST 1.7. Mol. Biol. Evol. 29, 1969–1973. https://doi.org/10.1093/molbev/mss075 (2012).CAS 
Article 
PubMed 
PubMed Central 

Google Scholar 
199.Rambaut, A. et al. Posterior summarization in Bayesian phylogenetics using Tracer 1.7. Syst. Biol. 67, 901–904. https://doi.org/10.1093/sysbio/syy032 (2018).CAS 
Article 
PubMed 
PubMed Central 

Google Scholar 
200.Matzke, N. J. Model selection in historical biogeography reveals that founder-event speciation is a crucial process in island clades. Syst. Biol. 63, 951–970. https://doi.org/10.1093/sysbio/syu056 (2014).Article 
PubMed 

Google Scholar 
201.Matzke, N. J. Probabilistic historical biogeography: New models for founder-event speciation, imperfect detection, and fossils allow improved accuracy and model-testing. Front. Biogeogr. 5, 242–248. https://doi.org/10.21425/F5FBG19694 (2013).Article 

Google Scholar 
202.Yu, Y., Blair, C. & He, X. J. RASP 4: Ancestral state reconstruction tool for multiple genes and characters. Mol. Biol. Evol. 37, 604–606. https://doi.org/10.1093/molbev/msz257 (2020).CAS 
Article 
PubMed 

Google Scholar 
203.Ree, R. H. & Sanmartín, I. Conceptual and statistical problems with the DEC+ J model of founder-event speciation and its comparison with DEC via model selection. J. Biogeogr. 45, 741–749. https://doi.org/10.1111/jbi.13173 (2018).Article 

Google Scholar 
204.Yu, Y., Harris, A. J. & He, X. S-DIVA (Statistical Dispersal-Vicariance Analysis): A tool for inferring biogeographic histories. Mol. Phylogenet. Evol. 56, 848–850. https://doi.org/10.1016/j.ympev.2010.04.011 (2010).Article 
PubMed 

Google Scholar 
205.Müller, R. D. et al. GPlates: Building a virtual Earth through deep time. Geochem. Geophys. Geosyst. 19, 2243–2261. https://doi.org/10.1029/2018GC007584 (2018).ADS 
Article 

Google Scholar 
206.Müller, R. D. et al. A global plate model including lithospheric deformation along major rifts and orogens since the Triassic. Tectonics 38, 1884–1907. https://doi.org/10.1029/2018TC005462 (2019).ADS 
Article 

Google Scholar 
207.Cao, X. et al. A deforming plate tectonic model of the South China Block since the Jurassic. Gondwana Res. https://doi.org/10.1016/j.gr.2020.11.010 (2020).Article 

Google Scholar 
208.Young, A. et al. Global kinematics of tectonic plates and subduction zones since the late Paleozoic Era. Geosci. Front. 10, 989–1013. https://doi.org/10.1016/j.gsf.2018.05.011 (2019).ADS 
Article 

Google Scholar 
209.Torsvik, T. H. et al. Pacific-Panthalassic reconstructions: Overview, errata and the way forward. Geochem. Geophys. Geosyst. 20, 3659–3689. https://doi.org/10.1029/2019GC008402 (2019).ADS 
Article 

Google Scholar 
210.Nevill, G. List of the Mollusca brought back by Dr. J. Anderson from Yunnan and Upper Burma, with descriptions of new species. J. Asiatic Soc. Bengal 46, 14–41 (1877).
Google Scholar 
211.Bolotov, I. N. et al. Indonaia rectangularis (Tapparone-Canefri, 1889), comb. nov., a forgotten freshwater mussel species from Myanmar. ZooKeys 852, 23–30. https://doi.org/10.3897/zookeys.852.33898 (2019).Article 
PubMed 
PubMed Central 

Google Scholar 
212.Eydoux, F. Mollusques. Magasin Zool. 8, 181–192 (1838).
Google Scholar 
213.Lea, I. Observations on the Naïades, and descriptions of new species of that and other families. Trans. Am. Philos. Soc. 4, 63–121 (1831).
Google Scholar 
214.Nesemann, H. A., Sharma, S. U., Sharma, G. O. & Sinha, R. K. Illustrated checklist of large freshwater bivalves of the Ganga River system (Mollusca: Bivalvia: Solecurtidae, Unionidae, Amblemidae). Nachrichchtenblatt Ersten Vorarlberger Malakologischen Gesellschaft 13, 1–51 (2005).
Google Scholar 
215.Gmelin, J. F. Systema Naturae per Regna Tria Naturae, Secundum Classes, Ordines, Genera, Species, cum Characteribus, Differentiis, Synonymis, locis. Curt 1(6), 3021–3909 (1791).
Google Scholar 
216.Lea, I. Description of twenty-five new species of exotic uniones. Proc. Acad. Natl. Sci. Phila. 8, 92–95 (1856).
Google Scholar 
217.Martens, E. V. Binnen-Conchylien aus Ober-Birma. Arch. Nat. 65, 30–48 (1899).
Google Scholar 
218.Preston, H. B. A catalogue of the Asiatic naiades in the collection of the Indian Museum, Calcutta, with descriptions of new species. Rec. Indian Mus. 7, 279–308 (1912).
Google Scholar 
219.Annandale, N. & Prashad, B. XXVIII. The aquatic and amphibious Mollusca of Manipur. Rec. Indian Mus. 22, 529–631 (1921).
Google Scholar 
220.Annandale, N. & Prashad, B. Some freshwater molluscs from the Bombay Presidency. Rec. Indian Mus. 16, 139–152 (1919).
Google Scholar 
221.Philippi, R. A. Unio. Tab. I. Abbildungen und Beschreibungen neuer oder wenig gekannter Conchylien 1, 19–20 (1843).222.Hanley, S. Appendix, containing descriptions of the shells delineated in the plates, yet not described in the text; with a systematic list of the engravings, etc. In An Illustrated and Descriptive Catalogue of Recent Bivalve Shells 335–389 (Williams and Norgate, 1856).
Google Scholar 
223.Theobald, W. Descriptions of some new land and freshwater shells from India and Burmah. J. Asiatic Soc. Bengal 45, 184–189 (1876).
Google Scholar 
224.Lea, I. Observations on the Naïades; and descriptions of new species of that and other families. Trans. Am. Philos. Soc. 5, 23–119 (1834).
Google Scholar 
225.Hutton, T. Notices of some land and fresh water shells occurring in Afghanistan. J. Asiatic Soc. Bengal 18, 649–661 (1849).
Google Scholar 
226.Annandale, N. Aquatic molluscs of the Inlé Lake and connected waters. Rec. Indian Mus. 14, 103–182 (1918).
Google Scholar 
227.Gould, A. A. D. Gould described new shells, received from Rev Mr Mason, of Burmah. Proc. Boston Soc. Nat. Hist. 2, 218–221 (1847).
Google Scholar 
228.Benson, W. H. Descriptions of Indian and Burmese species of the genus Unio, Retz. Ann. Mag. Nat. Hist. 10, 184–195 (1862).
Google Scholar 
229.Lea, I. Description of new freshwater and land shells. Trans. Am. Philos. Soc. 6, 1–154 (1838).
Google Scholar 
230.Lamarck, J.-B. Histoire naturelle des animaux sans vertèbres. Vol. 6 (Chez l’Auteur, 1819).231.Müller, O. F. Vermivm Terrestrium et Fluviatilium, Seu Animalium Infusoriorum, Helminthicorum et Testaceorum, non Marinorum, Succincta Historia. Havniae Lisiae 2, 1–214 (1774).
Google Scholar 
232.Lea, I. Descriptions of three new species of exotic uniones. Proc. Acad. Natl. Sci. Phila. 11, 331 (1860).
Google Scholar 
233.Lea, I. Continuation of paper on fresh water and land shells. Proc. Am. Philos. Soc. 2, 30–34 (1841).
Google Scholar 
234.Benson, W. H. Descriptive catalogue of a collection of land and fresh-water shells, chiefly contained in the museum of the Asiatic Society. J. Asiatic Soc. Bengal 5, 741–750 (1836).
Google Scholar 
235.Hislop, S. Description of fossil shells, from the above-described deposits. Q. J. Geol. Soc. Lond. 16, 166–181 (1860).
Google Scholar 
236.Malcolmson, J. G. XXXVIII: On the Fossils of the Eastern portion of the Great Basaltic District of India. Trans. Geol. Soc. Lond. 5, 537–575 (1840).
Google Scholar 
237.Newbold, C. Summary of the Geology of Southern India. Part V. Fresh-water Limestones and Cherts. J. R. Asiatic Soc. Great Br. Irel. 8, 219–227 (1846).
Google Scholar 
238.Prashad, B. On a new fossil unionid from the intertrappean beds of Peninsular India. Rec. Geol. Surv. India 51, 368–370 (1921).
Google Scholar 
239.Lopes-Lima, M. et al. Phylogeny of the most species-rich freshwater bivalve family (Bivalvia: Unionida: Unionidae): Defining modern subfamilies and tribes. Mol. Phylogenet. Evol. 106, 174–191. https://doi.org/10.1016/j.ympev.2016.08.021 (2017).Article 
PubMed 

Google Scholar 
240.Bird, P. An updated digital model of plate boundaries. Geochem. Geophys. Geosyst. 4, 1–52. https://doi.org/10.1029/2001GC000252 (2003).Article 

Google Scholar 
241.Preece, R. C. et al. William Benson and the Golden Age of Malacology in British India. Trop. Nat. Hist. 22, 1–612 (2022).MathSciNet 

Google Scholar  More

1.IPCC. Shukla, P. et al. Climate Change and Land: an IPCC special report on climate change, desertification, land degradation, sustainable land management, food security, and greenhouse gas fluxes in terrestrial ecosystems. (2019).2.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 
3.Moritz, C. & Agudo, R. The future of species under climate change: resilience or decline?. Science (80-) 80(341), 504–508 (2013).ADS 

Google Scholar 
4.Gobiet, A. et al. 21st century climate change in the European Alps—A review. Sci. Total Environ. 493, 1138–1151 (2014).ADS 
CAS 

Google Scholar 
5.Damschen, E. I., Harrison, S., Ackerly, D. D., Fernandez-Going, B. M. & Anacker, B. L. Endemic plant communities on special soils: early victims or hardy survivors of climate change?. J. Ecol. 100(5), 1122–1130 (2012).
Google Scholar 
6.Essl, F. et al. Distribution patterns, range size and niche breadth of Austrian endemic plants. Biol. Conserv. 142, 2547–2558 (2009).
Google Scholar 
7.Hülber, K. et al. Uncertainty in predicting range dynamics of endemic alpine plants under climate warming. Glob. Change Biol. 22, 2608–2619 (2016).ADS 

Google Scholar 
8.Wershow, S. T. & DeChaine, E. G. Retreat to refugia: Severe habitat contraction projected for endemic alpine plants of the Olympic Peninsula. Am. J. Bot. 105, 760–778 (2018).
Google Scholar 
9.Dagnino, D. et al. Climate change and the future of endemic flora in the South Western Alps: relationships between niche properties and extinction risk. Reg. Environ. Change 20, 1–12 (2020).
Google Scholar 
10.Dirnböck, T., Essl, F. & Rabitsch, W. Disproportional risk for habitat loss of high-altitude endemic species under climate change. Glob. Chang. Biol. 17, 990–996 (2011).ADS 

Google Scholar 
11.Parmesan, C. & Hanley, M. E. Plants and climate change: complexities and surprises. Ann. Bot. 116, 849–864 (2015).PubMed 
PubMed Central 

Google Scholar 
12.Pauli, H., Gottfried, M., Dirnböck, T., Dullinger, S. & Grabherr, G. Assessing the long-term dynamics of endemic plants at summit habitats. in Alpine biodiversity in Europe 195–207 (Springer, 2003).13.Parolo, G. & Rossi, G. Upward migration of vascular plants following a climate warming trend in the Alps. Basic Appl. Ecol. 9, 100–107 (2008).
Google Scholar 
14.Dullinger, S. et al. Extinction debt of high-mountain plants under twenty-first-century climate change. Nat. Clim. Change 2, 619–622 (2012).ADS 

Google Scholar 
15.Scherrer, D. & Körner, C. Topographically controlled thermal-habitat differentiation buffers alpine plant diversity against climate warming. J. Biogeogr. 38, 406–416 (2011).
Google Scholar 
16.Randin, C. F. et al. Climate change and plant distribution: local models predict high-elevation persistence. Glob. Change Biol. 15, 1557–1569 (2009).ADS 

Google Scholar 
17.Patsiou, T. S., Conti, E., Zimmermann, N. E., Theodoridis, S. & Randin, C. F. Topo-climatic microrefugia explain the persistence of a rare endemic plant in the Alps during the last 21 millennia. Glob. Change Biol. 20, 2286–2300 (2014).ADS 

Google Scholar 
18.Suggitt, A. J. et al. Extinction risk from climate change is reduced by microclimatic buffering. Nat. Clim. Change 8, 713–717 (2018).ADS 

Google Scholar 
19.Körner, C. The alpine life zone. in Alpine Plant Life 9–20 (Springer, 2003).20.Badgley, C. et al. Biodiversity and topographic complexity: modern and geohistorical perspectives. Trends Ecol. Evol. 32, 211–226 (2017).PubMed 
PubMed Central 

Google Scholar 
21.Graae, B. J. et al. Stay or go–how topographic complexity influences alpine plant population and community responses to climate change. Perspect. Plant Ecol. Evol. Syst. 30, 41–50 (2018).
Google Scholar 
22.Dobrowski, S. Z. A climatic basis for microrefugia: the influence of terrain on climate. Glob. Change Biol. 17, 1022–1035 (2011).ADS 

Google Scholar 
23.Keppel, G. et al. Refugia: identifying and understanding safe havens for biodiversity under climate change. Glob. Ecol. Biogeogr. 21, 393–404 (2012).
Google Scholar 
24.Hülber, K. et al. Habitat availability disproportionally amplifies climate change risks for lowland compared to alpine species. Glob. Ecol. Conserv. 23, e01113 (2020).
Google Scholar 
25.Loarie, S. R. et al. The velocity of climate change. Nature 462, 1052–1055 (2009).ADS 
CAS 

Google Scholar 
26.Vittoz, P. & Engler, R. Seed dispersal distances: a typology based on dispersal modes and plant traits. Bot. Helv. 117, 109–124 (2007).
Google Scholar 
27.Sandel, B. et al. The influence of Late Quaternary climate-change velocity on species endemism. Science (80-) 80(334), 660–664 (2011).ADS 

Google Scholar 
28.Harrison, S. & Noss, R. Endemism hotspots are linked to stable climatic refugia. Ann. Bot. 119, 207–214 (2017).PubMed 
PubMed Central 

Google Scholar 
29.Pignatti, E. & Pignatti, S. Plant life of the Dolomites. (Springer, 2016).30.Pawlowski, B. Remarks on endemism in the flora of the Alps and the Carpathians. Vegetatio 21, 181–243 (1970).
Google Scholar 
31.Schönswetter, P., Stehlik, I., Holderegger, R. & Tribsch, A. Molecular evidence for glacial refugia of mountain plants in the European Alps. Mol. Ecol. 14, 3547–3555 (2005).PubMed 
PubMed Central 

Google Scholar 
32.Carton, A. & Soldati, M. Geomorphological features of the Dolomites (Italy). (1993).33.Bosellini, A., Gianolla, P. & Stefani, M. Geology of the Dolomites. Episodes 26(3), 181–185 (2003).
Google Scholar 
34.Gianolla, P., Panizza, M., Micheletti, C. & Viola, F. Nomination of the Dolomites for inscription on the World Natural Heritage list UNESCO, nomination document. Prov. di Belluno, Prov. Auton. di Bolzano—Bozen, Prov. di Pordenone, Prov. Auton. di Trento, Prov. di Udine (2008).35.Erschbamer, B. et al. Changes in plant species diversity revealed by long-term monitoring on mountain summits in the Dolomites (northern Italy). Preslia 83, 387–401 (2011).
Google Scholar 
36.Unterluggauer, P., Mallaun, M. & Erschbamer, B. The higher the summit, the higher the diversity changes–results of a long-term monitoring project in the Dolomites. Gredleriana 16, 5–34 (2016).
Google Scholar 
37.Guisan, A. & Zimmermann, N. E. Predictive habitat distribution models in ecology. Ecol. Modell. 135, 147–186 (2000).
Google Scholar 
38.Pearson, R. G. Species’ distribution modeling for conservation educators and practitioners. Synth. Am. Museum Nat. Hist. 50, 54–89 (2007).
Google Scholar 
39.Trivedi, M. R., Berry, P. M., Morecroft, M. D. & Dawson, T. P. Spatial scale affects bioclimate model projections of climate change impacts on mountain plants. Glob. Change Biol. 14, 1089–1103 (2008).ADS 

Google Scholar 
40.Lembrechts, J. J., Nijs, I. & Lenoir, J. Incorporating microclimate into species distribution models. Ecography (Cop.) 42, 1267–1279 (2019).
Google Scholar 
41.Perazza, G. & Lorenz, R. Le orchidee dell’Italia nordorientale. Atlante corologico e Guid. al riconoscimento. Ed. Osiride, Rovereto (2013).42.Prosser, F., Bertolli, A., Festi, F. & Perazza, G. Flora del Trentino. Fondazione Museo civico di Rovereto (2019)43.Bertolli A., Prosser F., Tomasi G., Argenti C., – Flora Dolomitica. 50 fiori da conoscere nel patrimonio Unesco. Edizioni Osiride, Rovereto, 68 pp. (2019)44.Guisan, A., Thuiller, W. & Zimmermann, N. E. Habitat suitability and distribution models: with applications in R (Cambridge University Press, Cambridge, 2017).
Google Scholar 
45.Rossi G., Orsenigo S., Gargano D., Montagnani C., Peruzzi L., Fenu G., Abeli T., Alessandrini A., Astuti G., Bacchetta G., Bartolucci F., Bernardo L., Bovio M., Brullo S., Carta A., Castello M., Cogoni D., Conti F., Domina G., Foggi B., Gennai M., Gigante D., Iberite M., Lasen C., Magrini S., Nicolella G., Pinna M.S., Poggio L., Prosser F., Santangelo A., Selvaggi A., Stinca A., Tartaglini N., Troia A., Villani M.C., Wagensommer R.P., Wilhalm T., Blasi C.,. Lista Rossa della Flora Italiana. 2 Endemiti e altre specie minacciate. Ministero dell’Ambiente e della Tutela del Territorio e del Mare (2020)46.Rossi G., Montagnani C., Gargano D., Peruzzi L., Abeli T., Ravera S., Cogoni A., Fenu G., Magrini S., Gennai M., Foggi B., Wagensommer R.P., Venturella G., Blasi C., Raimondo F.M., Orsenigo S. (Eds.), Lista Rossa della Flora Italiana. 1. Policy Species e altre specie minacciate. Comitato Italiano IUCN e Ministero dell’Ambiente e della Tutela del Territorio e del Mare (2013)47.Buffa G., Carpenè B., Casarotto N., Da Pozzo M., Filesi L., Lasen C., Marcucci R., Masin R., Prosser F., Tasinazzo S., Villani M., Zanatta K. Lista rossa regionale piante vascolari del Veneto. Regione Veneto (2016)48.Wilhalm, T. & Hilpold, A. Rote Liste der gefährdeten Gefäßpflanzen Südtirols (Naturmuseum Südtirols, Bozen, 2006).
Google Scholar 
49.Karger, D. N. et al. Climatologies at high resolution for the earth’s land surface areas. Sci. data 4, 1–20 (2017).
Google Scholar 
50.Schwalm, C. R., Glendon, S. & Duffy, P. B. RCP8 5 tracks cumulative CO2 emissions. Proc. Natl. Acad. Sci. 117(33), 19656–19657 (2020).ADS 
CAS 
PubMed 
PubMed Central 

Google Scholar 
51.Sanderson, B. M., Knutti, R. & Caldwell, P. A representative democracy to reduce interdependency in a multimodel ensemble. J. Clim. 28, 5171–5194 (2015).ADS 

Google Scholar 
52.Kassambara A., & Mundt F. factoextra: Extract
and Visualize the Results of Multivariate Data Analyses. R package
version 1.0.7. https://CRAN.R-project.org/package=factoextra (2020).53.Lenoir, J., Hattab, T. & Pierre, G. Climatic microrefugia under anthropogenic climate change: implications for species redistribution. Ecography (Cop.) 40, 253–266 (2017).
Google Scholar 
54.Araújo, M. B. & New, M. Ensemble forecasting of species distributions. Trends Ecol. Evol. 22, 42–47 (2007).
Google Scholar 
55.Thuiller, W. et al. Package ‘biomod2’. Species Distrib. Model. within an ensemble Forecast. Framew. (2016).56.Barbet-Massin, M., Jiguet, F., Albert, C. H. & Thuiller, W. Selecting pseudo-absences for species distribution models: how, where and how many?. Methods Ecol. Evol. 3, 327–338 (2012).
Google Scholar 
57.Elith, J. et al. Novel methods improve prediction of species’ distributions from occurrence data. Ecography (Cop.) 29, 129–151 (2006).
Google Scholar 
58.Allouche, O., Tsoar, A. & Kadmon, R. Assessing the accuracy of species distribution models: prevalence, kappa and the true skill statistic (TSS). J. Appl. Ecol. 43, 1223–1232 (2006).
Google Scholar 
59.Liu, C., Berry, P. M., Dawson, T. P. & Pearson, R. G. Selecting thresholds of occurrence in the prediction of species distributions. Ecography 28, 385–393 (2005).
Google Scholar 
60.Cao, Y. et al. Using Maxent to model the historic distributions of stonefly species in Illinois streams: the effects of regularization and threshold selections. Ecol. Modell. 259, 30–39 (2013).
Google Scholar 
61.R Core Team. R: A Language and Environment for Statistical Computing. (2020).62.Riley, S. J., DeGloria, S. D. & Elliot, R. Index that quantifies topographic heterogeneity. Intermt. J. Sci. 5, 23–27 (1999).
Google Scholar 
63.Irl, S. D. H. et al. Climate vs topography–spatial patterns of plant species diversity and endemism on a high-elevation island. J. Ecol. 103, 1621–1633 (2015).
Google Scholar 
64.Tarquini, S. & Nannipieri, L. The 10 m-resolution TINITALY DEM as a trans-disciplinary basis for the analysis of the Italian territory: Current trends and new perspectives. Geomorphology 281, 108–115 (2017).ADS 

Google Scholar 
65.Hamann, A., Roberts, D. R., Barber, Q. E., Carroll, C. & Nielsen, S. E. Velocity of climate change algorithms for guiding conservation and management. Glob. Chang. Biol. 21, 997–1004 (2015).ADS 

Google Scholar 
66.Dexter, F. Wilcoxon-Mann-Whitney test used for data that are not normally distributed. Anesth. Anal. 117, 537–538 (2013)67.Geppert, C. et al. Consistent population declines but idiosyncratic range shifts in Alpine orchids under global change. Nat. Commun. 11, 1–11 (2020).
Google Scholar 
68.Erfanian, M. B., Sagharyan, M., Memariani, F. & Ejtehadi, H. Predicting range shifts of three endangered endemic plants of the Khorassan-Kopet Dagh floristic province under global change. Sci. Rep. 11, 1–13 (2021).
Google Scholar 
69.Muñoz-Sáez, A., Choe, H., Boynton, R. M., Elsen, P. R. & Thorne, J. H. Climate exposure shows high risk and few climate refugia for Chilean native vegetation. Sci. Total Environ. 785, 147399 (2021).ADS 

Google Scholar 
70.Dullinger, S. et al. Post-glacial migration lag restricts range filling of plants in the European Alps. Glob. Ecol. Biogeogr. 21, 829–840 (2012).
Google Scholar 
71.Sedlacek, J. F., Bossdorf, O., Cortés, A. J., Wheeler, J. A. & van Kleunen, M. What role do plant–soil interactions play in the habitat suitability and potential range expansion of the alpine dwarf shrub Salix herbacea?. Basic Appl. Ecol. 15(4), 305–315 (2014).
Google Scholar 
72.Di Nuzzo, L. et al. Contrasting multitaxon responses to climate change in Mediterranean mountains. Sci. Rep. 11, 1–12 (2021).
Google Scholar 
73.Zecca, G., Casazza, G., Piscopo, S., Minuto, L. & Grassi, F. Are the responses of plant species to Quaternary climatic changes idiosyncratic? A demographic perspective from the Western Alps. Plant Ecol. Divers. 10, 273–281 (2017).
Google Scholar 
74.Dainese, M. et al. Human disturbance and upward expansion of plants in a warming climate. Nat. Clim. Chang. 7, 577–580 (2017).ADS 

Google Scholar 
75.Boisvert-Marsh, L., Périé, C. & de Blois, S. Divergent responses to climate change and disturbance drive recruitment patterns underlying latitudinal shifts of tree species. J. Ecol. 107, 1956–1969 (2019).
Google Scholar 
76.Malcolm, J. R., Liu, C., Neilson, R. P., Hansen, L. & Hannah, L. E. E. Global warming and extinctions of endemic species from biodiversity hotspots. Conserv. Biol. 20, 538–548 (2006).PubMed 
PubMed Central 

Google Scholar 
77.Casazza, G. et al. Climate change hastens the urgency of conservation for range-restricted plant species in the central-northern Mediterranean region. Biol. Conserv. 179, 129–138 (2014).
Google Scholar 
78.Körner, C. The use of ‘altitude’in ecological research. Trends Ecol. Evol. 22, 569–574 (2007).PubMed 
PubMed Central 

Google Scholar 
79.Engler, R. et al. Predicting future distributions of mountain plants under climate change: does dispersal capacity matter?. Ecography (Cop.) 32, 34–45 (2009).
Google Scholar 
80.Ozinga, W. A. et al. Dispersal failure contributes to plant losses in NW Europe. Ecol. Lett. 12, 66–74 (2009).
Google Scholar 
81.Morueta-Holme, N. et al. Strong upslope shifts in Chimborazo’s vegetation over two centuries since Humboldt. Proc. Natl. Acad. Sci. 112, 12741–12745 (2015).ADS 
CAS 
PubMed 
PubMed Central 

Google Scholar 
82.Niskanen, A. K. J., Niittynen, P., Aalto, J., Väre, H. & Luoto, M. Lost at high latitudes: Arctic and endemic plants under threat as climate warms. Divers. Distrib. 25, 809–821 (2019).
Google Scholar 
83.Trew, B. T. & Maclean, I. M. D. Vulnerability of global biodiversity hotspots to climate change. Glob. Ecol. Biogeogr. 30, 768–783 (2021).
Google Scholar 
84.Garcia, M. B. et al. Rocky habitats as microclimatic refuges for biodiversity. A close-up thermal approach. Environ. Exp. Bot. 170, 103886 (2020).
Google Scholar 
85.Tribsch, A. Areas of endemism of vascular plants in the Eastern Alps in relation to Pleistocene glaciation. J. Biogeogr. 31, 747–760 (2004).
Google Scholar 
86.Keppel, G. et al. The capacity of refugia for conservation planning under climate change. Front. Ecol. Environ. 13, 106–112 (2015).
Google Scholar 
87.Panizza, M. The geomorphodiversity of the Dolomites (Italy): a key of geoheritage assessment. Geoheritage 1, 33–42 (2009).
Google Scholar 
88.Santini, L., Benitez-López, A., Maiorano, L., Čengić, M. & Huijbregts, M. A. J. Assessing the reliability of species distribution projections in climate change research. Divers. Distrib. 27, 1035–1050 (2021).
Google Scholar 
89.Blois, J. L., Zarnetske, P. L., Fitzpatrick, M. C. & Finnegan, S. Climate change and the past, present, and future of biotic interactions. Science (80-) 341, 499–504 (2013).ADS 
CAS 

Google Scholar 
90.Meineri, E. & Hylander, K. Fine-grain, large-domain climate models based on climate station and comprehensive topographic information improve microrefugia detection. Ecography (Cop.) 40, 1003–1013 (2017).
Google Scholar 
91.Ferrarini, A. et al. Planning for assisted colonization of plants in a warming world. Sci. Rep. 6, 1–6 (2016).
Google Scholar 
92.Casazza, G. et al. Combining conservation status and species distribution models for planning assisted colonisation under climate change. J. Ecol. 109, 2284–2295 (2021) More

1.Gartman A, Findlay AJ. Impacts of hydrothermal plume processes on oceanic metal cycles and transport. Nat Geosci. 2020;13:396–402.CAS 

Google Scholar 
2.Sander SG, Koschinsky A. Metal flux from hydrothermal vents increased by organic complexation. Nat Geosci. 2011;4:145–50.CAS 

Google Scholar 
3.German CR, Casciotti KA, Dutay JC, Heimbürger LE, Jenkins WJ, Measures CI, et al. Hydrothermal impacts on trace element and isotope ocean biogeochemistry. Philos Trans R Soc A Math Phys Eng Sci. 2016;374:20160035.
Google Scholar 
4.Ardyna M, Lacour L, Sergi S, d’Ovidio F, Sallée JB, Rembauville M, et al. Hydrothermal vents trigger massive phytoplankton blooms in the Southern Ocean. Nat Commun. 2019;10:1–8.CAS 

Google Scholar 
5.McCollom TM. Geochemical constraints on primary productivity in submarine hydrothermal vent plumes. Deep Res Part I Oceanogr Res Pap. 2000;47:85–101.CAS 

Google Scholar 
6.Dick GJ, Tebo BM. Microbial diversity and biogeochemistry of the Guaymas Basin deep-sea hydrothermal plume. Environ Microbiol. 2010;12:1334–47.CAS 
PubMed 

Google Scholar 
7.Nakamura K, Takai K. Theoretical constraints of physical and chemical properties of hydrothermal fluids on variations in chemolithotrophic microbial communities in seafloor hydrothermal systems. Prog Earth Planet Sci. 2014;1:1–24.
Google Scholar 
8.Dick GJ. The microbiomes of deep-sea hydrothermal vents: distributed globally, shaped locally. Nat Rev Microbiol. 2019;17:271–83.CAS 
PubMed 

Google Scholar 
9.Sunamura M, Higashi Y, Miyako C, Ishibashi JI, Maruyama A. Two bacteria phylotypes are predominant in the Suiyo Seamount hydrothermal plume. Appl Environ Microbiol. 2004;70:1190–8.CAS 
PubMed 
PubMed Central 

Google Scholar 
10.Lavik G, Stührmann T, Brüchert V, Van Der Plas A, Mohrholz V, Lam P, et al. Detoxification of sulphidic African shelf waters by blooming chemolithotrophs. Nature. 2009;457:581–4.CAS 
PubMed 

Google Scholar 
11.Canfield DE, Stewart FJ, Thamdrup B, De Brabandere L, Dalsgaard T, Delong EF, et al. A cryptic sulfur cycle in oxygen-minimum-zone waters off the Chilean coast. Science. 2010;330:1375–8.CAS 
PubMed 

Google Scholar 
12.Callbeck CM, Lavik G, Ferdelman TG, Fuchs B, Gruber-Vodicka HR, Hach PF, et al. Oxygen minimum zone cryptic sulfur cycling sustained by offshore transport of key sulfur oxidizing bacteria. Nat Commun. 2018;9:1.CAS 

Google Scholar 
13.Glaubitz S, Kießlich K, Meeske C, Labrenz M, Jürgens K. SUP05 Dominates the gammaproteobacterial sulfur oxidizer assemblages in pelagic redoxclines of the central baltic and black seas. Appl Environ Microbiol. 2013;79:2767–76.CAS 
PubMed 
PubMed Central 

Google Scholar 
14.Pjevac P, Korlević M, Berg JS, Bura-Nakić E, Ciglenečki I, Amann R, et al. Community shift from phototrophic to chemotrophic sulfide oxidation following anoxic holomixis in a stratified seawater lake. Appl Environ Microbiol. 2015;81:298–308.PubMed 

Google Scholar 
15.Zhou K, Zhang R, Sun J, Zhang W, Tian RM, Chen C, et al. Potential interactions between clade SUP05 sulfur-oxidizing bacteria and phages in hydrothermal vent sponges. Appl Environ Microbiol. 2019;85:1–20.
Google Scholar 
16.Duperron S, Nadalig T, Caprais JC, Sibuet M, Fiala-Médioni A, Amann R, et al. Dual symbiosis in a Bathymodiolus sp. mussel from a methane seep on the Gabon Continental Margin (Southeast Atlantic): 16S rRNA phylogeny and distribution of the symbionts in gills. Appl Environ Microbiol. 2005;71:1694–700.CAS 
PubMed 
PubMed Central 

Google Scholar 
17.Ansorge R, Romano S, Sayavedra L, Porras MÁG, Kupczok A, Tegetmeyer HE, et al. Functional diversity enables multiple symbiont strains to coexist in deep-sea mussels. Nat Microbiol. 2019;4:2487–97.PubMed 

Google Scholar 
18.Anantharaman K, Breier JA, Sheik CS, Dick GJ. Evidence for hydrogen oxidation and metabolic plasticity in widespread deep-sea sulfur-oxidizing bacteria. Proc Natl Acad Sci USA. 2013;110:330–5.CAS 
PubMed 

Google Scholar 
19.Wang W, Li Z, Zeng L, Dong C, Shao Z. The oxidation of hydrocarbons by diverse heterotrophic and mixotrophic bacteria that inhabit deep-sea hydrothermal ecosystems. ISME J. 2020;14:1994–2006.CAS 
PubMed 
PubMed Central 

Google Scholar 
20.Spietz RL, Lundeen RA, Zhao X, Nicastro D, Ingalls AE, Morris RM. Heterotrophic carbon metabolism and energy acquisition in Candidatus Thioglobus singularis strain PS1, a member of the SUP05 clade of marine Gammaproteobacteria. Environ Microbiol. 2019;21:2391–401.CAS 
PubMed 

Google Scholar 
21.Marshall KT, Morris RM. Isolation of an aerobic sulfur oxidizer from the SUP05/Arctic96BD-19 clade. ISME J. 2013;7:452–5.CAS 
PubMed 

Google Scholar 
22.Shah V, Morris RM. Genome sequence of “Candidatus Thioglobus autotrophica” strain EF1, a chemoautotroph from the SUP05 clade of marine Gammaproteobacteria. Genome Announc. 2015;3:e01156–15.PubMed 
PubMed Central 

Google Scholar 
23.van Vliet DM, von Meijenfeldt FAB, Dutilh BE, Villanueva L, Sinninghe Damsté JS, Stams AJM, et al. The bacterial sulfur cycle in expanding dysoxic and euxinic marine waters. Environ Microbiol. 2021;23:2834–57.PubMed 

Google Scholar 
24.De Ronde CEJ, Baker ET, Massoth GJ, Lupton JE, Wright IC, Feely RA, et al. Intra-oceanic subduction-related hydrothermal venting, Kermadec volcanic arc, New Zealand. Earth Planet Sci Lett. 2001;193:359–69.
Google Scholar 
25.De Ronde CEJ, Baker ET, Massoth GJ, Lupton JE, Wright IC, Sparks RJ, et al. Submarine hydrothermal activity along the mid-Kermadec Arc, New Zealand: large-scale effects on venting. Geochem Geophys Geosyst. 2007;8:Q07007.
Google Scholar 
26.Kleint C, Bach W, Diehl A, Fröhberg N, Garbe-Schönberg D, Hartmann JF, et al. Geochemical characterization of highly diverse hydrothermal fluids from volcanic vent systems of the Kermadec intraoceanic arc. Chem Geol. 2019;528:119289.CAS 

Google Scholar 
27.Baker ET, Resing JA, Haymon RM, Tunnicliffe V, Martinez F, Ferrini V, et al. How many vent fields? New estimates of vent field populations on ocean ridges from precise mapping of hydrothermal discharge locations. Prog Earth Planet Sci. 2016;449:186–96.CAS 

Google Scholar 
28.Walker SL, Baker ET, Resing JA, Nakamura K, McLain PD. A new tool for detecting hydrothermal plumes: an ORP sensor for the PMEL MAPR. AGU Fall Meet Abstr. 2007;2007:V21D–0753.
Google Scholar 
29.Herlemann DPR, Labrenz M, Jürgens K, Bertilsson S, Waniek JJ, Andersson AF. Transitions in bacterial communities along the 2000 km salinity gradient of the Baltic Sea. ISME J. 2011;5:1571–9.CAS 
PubMed 
PubMed Central 

Google Scholar 
30.Reintjes G, Tegetmeyer HE, Bürgisser M, Orlić S, Tews I, Zubkov M, et al. On-site analysis of bacterial communities of the ultraoligotrophic South Pacific Gyre. Appl Environ Microbiol. 2019;85:e00184–19.CAS 
PubMed 
PubMed Central 

Google Scholar 
31.Martin M. Cutadapt removes adapter sequences from high-throughput sequencing reads. EMBnet J. 2011;17:10–2.
Google Scholar 
32.Schloss PD, Westcott SL, Ryabin T, Hall JR, Hartmann M, Hollister EB, et al. Introducing mothur: Open-source, platform-independent, community-supported software for describing and comparing microbial communities. Appl Environ Microbiol. 2009;75:7537–41.CAS 
PubMed 
PubMed Central 

Google Scholar 
33.Bushnell B. BBMap (version 35.14) [Software]. 2015. https://sourceforge.net/projects/bbmap/.34.Quast C, Pruesse E, Yilmaz P, Gerken J, Schweer T, Yarza P, et al. The SILVA ribosomal RNA gene database project: Improved data processing and web-based tools. Nucleic Acids Res. 2013;41:D590–6.CAS 
PubMed 
PubMed Central 

Google Scholar 
35.Ludwig W, Strunk O, Westram R, Richter L, Meier H, Yadhukumar A, et al. ARB: a software environment for sequence data. Nucleic Acids Res. 2004;32:1363–71.CAS 
PubMed 
PubMed Central 

Google Scholar 
36.Pernthaler A, Pernthaler J, Amann R.  Fluorescence in situ hybridization and catalyzed reporter deposition for the identification of marine bacteria. Appl Environ Microbiol. 2002;68:3094–101.CAS 
PubMed 
PubMed Central 

Google Scholar 
37.Andrews S. FastQC: a quality control tool for high throughput sequence data. Babraham Bioinformatics; 2010.38.Rodriguez-R LM, Gunturu S, Tiedje JM, Cole JR, Konstantinidis KT. Nonpareil 3: fast estimation of metagenomic coverage and sequence diversity. mSystems. 2018;3:e00039–18.PubMed 
PubMed Central 

Google Scholar 
39.Li D, Luo R, Liu CM, Leung CM, Ting HF, Sadakane K, et al. MEGAHIT v1.0: a fast and scalable metagenome assembler driven by advanced methodologies and community practices. Methods. 2016;102:3–11.CAS 
PubMed 

Google Scholar 
40.Strous M, Kraft B, Bisdorf R, Tegetmeyer HE. The binning of metagenomic contigs for microbial physiology of mixed cultures. Front Microbiol. 2012;3:410.PubMed 
PubMed Central 

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

Google Scholar 
42.Eren AM, Kiefl E, Shaiber A, Veseli I, Miller SE, Schechter MS, et al. Community-led, integrated, reproducible multi-omics with anvi’o. Nat Microbiol. 2021;6:3–6.CAS 
PubMed 
PubMed Central 

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

Google Scholar 
44.Bankevich A, Nurk S, Antipov D, Gurevich AA, Dvorkin M, Kulikov AS, et al. SPAdes: a new genome assembly algorithm and its applications to single-cell sequencing. J Comput Biol. 2012;19:455–77.CAS 
PubMed 
PubMed Central 

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

Google Scholar 
46.Bolger AM, Lohse M, Usadel B. Trimmomatic: a flexible trimmer for Illumina sequence data. Bioinformatics. 2014;30:2114–20.CAS 
PubMed 
PubMed Central 

Google Scholar 
47.Kopylova E, Noé L, Touzet H. SortMeRNA: fast and accurate filtering of ribosomal RNAs in metatranscriptomic data. Bioinformatics. 2012;28:3211–7.CAS 
PubMed 

Google Scholar 
48.Gomes AÉ, Stuchi LP, Siqueira NM, Henrique JB, Vicentini R, Ribeiro ML, et al. Selection and validation of reference genes for gene expression studies in Klebsiella pneumoniae using Reverse Transcription Quantitative real-time PCR. Sci Rep. 2018;8:1–4.
Google Scholar 
49.Kolde R. pheatmap: Pretty heatmaps. 2015. https://CRAN.R-project.org/package=pheatmap.50.Garnier S. viridis: Default Color Maps from’matplotlib’. 2017. https://CRAN.R-project.org/.51.R Core Team. R: a language and environment for statistical computing. Vienna, Austria: R Foundation for Statistical Computing; 2013.52.Oksanen J, Blanchet FG, Friendly M, Kindt R, Legendre P, McGlinn D, et al. Vegan: Community ecology package. 2020.53.Pena EA, Slate EH. gvlma: Global validation of linear models assumptions. R package version 1.0.0.3. 2019. https://CRAN.R-project.org/package=gvlma.54.Anderson MJ. A new method for non parametric multivariate analysis of variance. Austral Ecol. 2001;26:32–46.
Google Scholar 
55.Waite DW, Chuvochina M, Pelikan C, Parks DH, Yilmaz P, Wagner M, et al. Proposal to reclassify the proteobacterial classes Deltaproteobacteria and Oligoflexia, and the phylum Thermodesulfobacteria into four phyla reflecting major functional capabilities. Int J Syst Evol Microbiol. 2020;70:5972–6016.CAS 
PubMed 

Google Scholar 
56.Anantharaman K, Breier JA, Dick GJ. Metagenomic resolution of microbial functions in deep-sea hydrothermal plumes across the Eastern Lau Spreading Center. ISME J. 2016;10:225–39.CAS 
PubMed 

Google Scholar 
57.Biller SJ, Berube PM, Dooley K, Williams M, Satinsky BM, Hackl T, et al. Data descriptor: marine microbial metagenomes sampled across space and time. Sci Data. 2018;5:1–7.
Google Scholar 
58.Meier DV, Pjevac P, Bach W, Hourdez S, Girguis PR, Vidoudez C, et al. Niche partitioning of diverse sulfur-oxidizing bacteria at hydrothermal vents. ISME J. 2017;11:1545–58.CAS 
PubMed 
PubMed Central 

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

Google Scholar 
60.Zhou Z, Tran PQ, Kieft K, Anantharaman K. Genome diversification in globally distributed novel marine Proteobacteria is linked to environmental adaptation. ISME J. 2020;14:2060–77.CAS 
PubMed 
PubMed Central 

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

Google Scholar 
62.Blackburn NT, Clarke AJ. Identification of four families of peptidoglycan lytic transglycosylases. J Mol Evol. 2001;52:78–84.CAS 
PubMed 

Google Scholar 
63.Hashimoto W, Ochiai A, Momma K, Itoh T, Mikami B, Maruyama Y, et al. Crystal structure of the glycosidase family 73 peptidoglycan hydrolase FlgJ. Biochem Biophys Res Commun. 2009;381:16–21.CAS 
PubMed 

Google Scholar 
64.Ilbert M, Bonnefoy V. Insight into the evolution of the iron oxidation pathways. Biochim Biophys Acta Bioenerg. 2013;1827:161–75.CAS 

Google Scholar 
65.Barco RA, Emerson D, Sylvan JB, Orcutt BN, Jacobson Meyers ME, Ramírez GA, et al. New insight into microbial iron oxidation as revealed by the proteomic profile of an obligate iron-oxidizing chemolithoautotroph. Appl Environ Microbiol. 2015;81:5927–37.CAS 
PubMed 
PubMed Central 

Google Scholar 
66.Guo J, Bolduc B, Zayed AA, Varsani A, Dominguez-Huerta G, Delmont TO, et al. VirSorter2: a multi-classifier, expert-guided approach to detect diverse DNA and RNA viruses. Microbiome. 2021;9:1–13.67.Duarte CM. Seafaring in the 21st century: the Malaspina 2010 circumnavigation expedition. Limnol Oceanogr Bull. 2015;24:11–14.
Google Scholar 
68.Sheik CS, Anantharaman K, Breier JA, Sylvan JB, Edwards KJ, Dick GJ. Spatially resolved sampling reveals dynamic microbial communities in rising hydrothermal plumes across a back-arc basin. ISME J. 2015;9:1434–45.PubMed 

Google Scholar 
69.Konstantinidis KT, Rosselló-Móra R, Amann R. Uncultivated microbes in need of their own taxonomy. ISME J. 2017;11:2399–406.PubMed 
PubMed Central 

Google Scholar 
70.Murray AE, Freudenstein J, Gribaldo S, Hatzenpichler R, Hugenholtz P, Kämpfer P, et al. Roadmap for naming uncultivated Archaea and Bacteria. Nat Microbiol. 2020;5:987–94.CAS 
PubMed 
PubMed Central 

Google Scholar 
71.Shah V, Zhao X, Lundeen RA, Ingalls AE, Nicastro D, Morris RM. Morphological plasticity in a sulfur-oxidizing marine bacterium from the SUP05 clade enhances dark carbon fixation. MBio. 2019;10:e00216–19.CAS 
PubMed 
PubMed Central 

Google Scholar 
72.Yamamoto M, Takai K. Sulfur metabolisms in Epsilon- and Gammaproteobacteria in deep-sea hydrothermal fields. Front Microbiol. 2011;2:192.CAS 
PubMed 
PubMed Central 

Google Scholar 
73.White GF, Edwards MJ, Gomez-Perez L, Richardson DJ, Butt JN, Clarke TA. Mechanisms of bacterial extracellular electron exchange. Adv Micro Physiol. 2016;68:87–138.CAS 

Google Scholar 
74.Findlay AJ, Estes ER, Gartman A, Yücel M, Kamyshny A, Luther GW. Iron and sulfide nanoparticle formation and transport in nascent hydrothermal vent plumes. Nat Commun. 2019;10:1–7.CAS 

Google Scholar 
75.Gartman A, Luther GW. Oxidation of synthesized sub-micron pyrite (FeS2) in seawater. Geochim Cosmochim Acta. 2014;144:96–108.CAS 

Google Scholar 
76.Bonnefoy V, Holmes DS. Genomic insights into microbial iron oxidation and iron uptake strategies in extremely acidic environments. Environ Microbiol. 2012;14:1597–611.CAS 
PubMed 

Google Scholar 
77.Singh VK, Singh AL, Singh R, Kumar A. Iron oxidizing bacteria: insights on diversity, mechanism of iron oxidation and role in management of metal pollution. Environ Sustain. 2018;1:221–31.
Google Scholar 
78.He S, Barco RA, Emerson D, Roden EE. Comparative genomic analysis of neutrophilic iron(II) oxidizer genomes for candidate genes in extracellular electron transfer. Front Microbiol. 2017;8:1584.PubMed 
PubMed Central 

Google Scholar 
79.McAllister SM, Polson SW, Butterfield DA, Glazer BT, Sylvan JB, Chan CS. Validating the Cyc2 neutrophilic iron oxidation pathway using meta-omics of Zetaproteobacteria iron mats at marine hydrothermal vents. mSystems. 2020;5:e00553–19.PubMed 
PubMed Central 

Google Scholar 
80.Barco RA, Hoffman CL, Ramírez GA, Toner BM, Edwards KJ, Sylvan JB. In-situ incubation of iron-sulfur mineral reveals a diverse chemolithoautotrophic community and a new biogeochemical role for Thiomicrospira. Environ Microbiol. 2017;19:1322–37.CAS 
PubMed 

Google Scholar 
81.Lesniewski RA, Jain S, Anantharaman K, Schloss PD, Dick GJ. The metatranscriptome of a deep-sea hydrothermal plume is dominated by water column methanotrophs and lithotrophs. ISME J. 2012;6:2257–68.CAS 
PubMed 
PubMed Central 

Google Scholar 
82.Reed DC, Breier JA, Jiang H, Anantharaman K, Klausmeier CA, Toner BM, et al. Predicting the response of the deep-ocean microbiome to geochemical perturbations by hydrothermal vents. ISME J. 2015;9:1857–69.PubMed 
PubMed Central 

Google Scholar 
83.Maki JS. Bacterial intracellular sulfur globules: structure and function. J Mol Microbiol Biotechnol. 2013;23:270–80.CAS 
PubMed 

Google Scholar 
84.Neuholz R, Kleint C, Schnetger B, Koschinsky A, Laan P, Middag R, et al. Submarine hydrothermal discharge and fluxes of dissolved Fe and Mn, and He isotopes at Brothers Volcano based on radium isotopes. Minerals. 2020;10:969.CAS 

Google Scholar 
85.Waite DW, Vanwonterghem I, Rinke C, Parks DH, Zhang Y, Takai K, et al. Comparative genomic analysis of the class Epsilonproteobacteria and proposed reclassification to Epsilonbacteraeota (phyl. nov.). Front Microbiol. 2017;8:682.PubMed 
PubMed Central 

Google Scholar 
86.Waite DW, Vanwonterghem I, Rinke C, Parks DH, Zhang Y, Takai K, et al. Addendum: comparative genomic analysis of the class Epsilonproteobacteria and proposed reclassification to Epsilonbacteraeota (phyl. nov.). Front Microbiol. 2018;9:772.PubMed 
PubMed Central 

Google Scholar  More

1.Kiessling, W., Simpson, C. & Foote, M. Reefs as cradles of evolution and sources of biodiversity in the Phanerozoic. Science 327, 196–198. https://doi.org/10.1126/science.1182241 (2016).ADS 
CAS 
Article 

Google Scholar 
2.Kiessling, W. Geological and biologic controls on the evolution of reefs. Annu. Rev. Ecol. Evol. Syst. 40, 173–192. https://doi.org/10.1146/annurev.ecolsys.110308.120251(2009) (2009).Article 

Google Scholar 
3.Talent, J. A. Organic reef-building: Episodes of extinction and symbiosis?. Senckenb. Lethaea 69, 315–368 (1988).
Google Scholar 
4.Flügel, E. & Kiessling, W. Patterns of Phanerozoic reef crises. SEPM Spec. Public. 72, 691–733 (2002).
Google Scholar 
5.Pandolfi, J. M. & Kiessling, W. Gaining insights from past reefs to inform understanding of coral reef response to global climate change. Curr. Opin. Environ. Sustain. 7, 52–58. https://doi.org/10.1016/j.cosust.2013.11.020 (2014).Article 

Google Scholar 
6.Copper, P. Ancient reef ecosystem expansion and collapse. Coral Reefs 13, 3–11 (1994).ADS 
Article 

Google Scholar 
7.Copper, P. Silurian and Devonian reefs: 80 million years of global greenhouse between two ice ages. SEPM Spec. Public. 72, 181–238 (2002).
Google Scholar 
8.Copper, P. & Scotese, C. R. Megareefs in Middle Devonain supergreenhouse climates. Spec. Public. Geol. Soc. Am. 370, 209–230. https://doi.org/10.1130/0-8137-2370-1.209 (2003).Article 

Google Scholar 
9.Ries, J. B. Geological and experimental evidence for secular variation in seawater Mg/Ca (calcite-aragonite seas) and its effects on marine biological calcification. Biogeosciences 7(9), 2795–2849 (2010).ADS 
CAS 
Article 

Google Scholar 
10.Scotese, C. R., Song, H., Mills, B. J. & van der Meer, D. G. Phanerozoic paleotemperatures: The earth’s changing climate during the last 540 million years. Earth Sci. Rev. https://doi.org/10.1016/j.earscirev.2021.103503 (2021).Article 

Google Scholar 
11.Zapalski, M. K., Nowicki, J., Jakubowicz, M. & Berkowski, B. Tabulate corals across the Frasnian/Famennian boundary: architectural turnover and its possible relation to ancient photosymbiosis. Palaeogeogr. Palaeoclimatol. Palaeoecol. 487, 416–429. https://doi.org/10.1016/j.palaeo.2017.09.028 (2017).Article 

Google Scholar 
12.Mora, C. I., Driese, S. G. & Seager, P. G. Carbon dioxide in the Paleozoic atmosphere: Evidence from carbon-isotope compositions of pedogenic carbonate. Geology 19(10), 1017–1020 (1991).ADS 
CAS 
Article 

Google Scholar 
13.Foster, G. L., Royer, D. L. & Lunt, D. J. Future climate forcing potentially without precedent in the last 420 million years. Nat. Commun. 8(1), 1–8. https://doi.org/10.1038/ncomms14845 (2017).ADS 
CAS 
Article 

Google Scholar 
14.Kiessling, W., Flügel, E. & Golonka, J. Paleoreef maps: evaluation of a comprehensive database on Phanerozoic reefs. AAPG Bull. 83(10), 1552–1587 (1999).
Google Scholar 
15.Burchette, T. P. European Devonian reefs: a review of current concepts and models. SEPM Spec. Public. 30, 85–142 (1981).
Google Scholar 
16.Ziegler, A. M., Scotese, C. R., McKerrow, W. S., Johnson, M. E. & Bambach, R. K. Paleozoic paleogeography. Annu. Rev. Earth Planet. Sci. 7(1), 473–502 (1979).ADS 
Article 

Google Scholar 
17.Belka, Z. & Narkiewicz, M. Devonian. In: McCann, T. The Geology of Central Europe, 1: Precambrian and Palaeozoic. 383–410. The Geological Society of London (2008).18.Golonka, J. Plate-tectonic maps of the Phanerozoic. SEPM Spec. Public. 72, 21–75 (2002).
Google Scholar 
19.Oczlon, M. S. Ocean currents and unconformities: the north Gondwana Middle Devonian. Geology 18(6), 509–512 (1990).ADS 
Article 

Google Scholar 
20.Dopieralska, J. Reconstructing seawater circulation on the Moroccan shelf of Gondwana during the Late Devonian: Evidence from Nd isotope composition of conodonts. Geochem. Geophys. Geosyst. 10(3), Q03015. https://doi.org/10.1029/2008GC002247 (2009).ADS 
CAS 
Article 

Google Scholar 
21.Jakubowicz, M. et al. At the southern limits of the Devonian reef zone: Palaeoecology of the Aferdou el Mrakib reef (Givetian, eastern Anti-Atlas, Morocco). Geol. J. 54(1), 10–38. https://doi.org/10.1002/gj.3152 (2019).Article 

Google Scholar 
22.Wood, R. Reef evolution (Oxford University Press, 1999).
Google Scholar 
23.Raup, D. M. & Sepkoski, J. J. Mass extinctions in the marine fossil record. Science 215(4539), 1501–1503 (1982).ADS 
CAS 
Article 

Google Scholar 
24.McGhee, G. R. Jr., Sheehan, P. M., Bottjer, D. J. & Droser, M. L. Ecological ranking of Phanerozoic biodiversity crises: the Serpukhovian (early Carboniferous) crisis had a greater ecological impact than the end-Ordovician. Geology 40(2), 147–150. https://doi.org/10.1016/j.palaeo.2004.05.010 (2012).ADS 
Article 

Google Scholar 
25.Stanley, S. M. Estimates of the magnitudes of major marine mass extinctions in earth history. Proc. Natl. Acad. Sci. 113(42), E6325–E6334. https://doi.org/10.1073/pnas.1613094113 (2016).ADS 
CAS 
Article 
PubMed 
PubMed Central 

Google Scholar 
26.Zapalski, M. K., Wrzołek, T., Skompski, S. & Berkowski, B. Deep in shadows, deep in time: the oldest mesophotic coral ecosystems from the Devonian of the Holy Cross Mountains (Poland). Coral Reefs 36(3), 847–860. https://doi.org/10.1007/s00338-017-1575-8 (2017).ADS 
Article 

Google Scholar 
27.Zapalski, M. K., Baird, A. H., Bridge, T., Jakubowicz, M. & Daniell, J. Unusual shallow water Devonian coral community from Queensland and its recent analogues from the inshore Great Barrier Reef. Coral Reefs 40(2), 417–431. https://doi.org/10.1007/s00338-020-02048-9 (2021).Article 

Google Scholar 
28.Zapalski, M. K., Hubert, B. L., Nicollin, J. P., Mistiaen, B. & Brice, D. The palaeobiodiversity of stromatoporoids, tabulates and brachiopods in the Devonian of the Ardennes–changes through time. Bulletin de la Société Géologique de France 178(5), 383–390. https://doi.org/10.2113/gssgfbull.178.5.383 (2007).Article 

Google Scholar 
29.Zapalski, M., Pinte, E. & Mistiaen, B. Late Famennian? Chaetosalpinx in Yavorskia (Tabulata): the youngest record of tabulate endobionts. Acta Geol. Pol. 58(3), 321–324 (2008).
Google Scholar 
30.Zapalski, M. K. & Berkowski, B. The oldest species of? Yavorskia (Tabulata) from the upper Famennian of the Holy Cross Mountains (Poland). Acta Geol. Pol. 62(2), 197–204 (2012).
Google Scholar 
31.Zapalski, M. K., Berkowski, B. & Wrzołek, T. Tabulate corals after the Frasnian/Famennian crisis: a unique fauna from the Holy Cross Mountains, Poland. PLoS ONE 11(3), e0149767. https://doi.org/10.1371/journal.pone.0149767 (2016).CAS 
Article 
PubMed 
PubMed Central 

Google Scholar 
32.Stanley, G. D. Jr. & Lipps, J. H. Photosymbiosis: the driving force for reef success and failure. Paleontol. Soc. Paper 17, 33–60 (2011).Article 

Google Scholar 
33.Coates, A. G. & Jackson, J. B. C. Clonal growth, algal symbiosis, and reef formation by corals. Paleobiology 13, 363–378 (1987).Article 

Google Scholar 
34.Zapalski, M. K. Evidence of photosymbiosis in Palaeozoic tabulate corals. Proc. R. Soc B Biol. Sci. 281(1775), 20132663. https://doi.org/10.1098/rspb.2013.2663 (2014).CAS 
Article 

Google Scholar 
35.Zapalski, M. K. & Berkowski, B. The Silurian mesophotic coral ecosystems: 430 million years of photosymbiosis. Coral Reefs 38(1), 137–147. https://doi.org/10.1007/s00338-018-01761-w (2019).ADS 
Article 

Google Scholar 
36.Coates, A. G., & Oliver Jr, W. A. Coloniality of Coral Zoantharia: Animal Colonies.–3–29 (1973).37.Lipps, J. H., & Stanley, G. D. Photosymbiosis in past and present reefs. In Coral Reefs at the Crossroads (pp. 47–68). Springer (2016).38.Blieck, A., Brice, D., Fesir, R., Guillot, F., Majesté-Mejoulas, C., and Meillez, F., The Devonian of France and Belgium, in McMillan, A.F., Embry, A.F., and Glass, D.J., eds., Devonian of the world, Canadian Society of Petroleum Geologists, Calgary, 1, p. 359–400 (1988)39.Porter, J. W. Autotrophy, heterotrophy, and resource partitioning in Caribbean reef-building corals. Am. Nat. 110, 731–742 (1976).ADS 
Article 

Google Scholar 
40.McGhee, G. R. Jr., Clapham, M. E., Sheehan, P. M., Bottjer, D. J. & Droser, M. L. A new ecological-severity ranking of major Phanerozoic biodiversity crises. Palaeogeogr. Palaeoclimatol. Palaeoecol. 370, 260–270 (2013).Article 

Google Scholar 
41.Aboussalam, Z. S. & Becker, R. T. The global Taghanic Biocrisis (Givetian) in the eastern Anti-Atlas, Morocco. Palaeogeogr. Palaeoclimatol. Palaeoecol. 304(1–2), 136–164 (2011).Article 

Google Scholar 
42.Zambito, J. J., Brett, C. E., & Baird, G. C. The Late Middle Devonian (Givetian) Global Taghanic Biocrisis in its type area (northern Appalachian Basin): geologically rapid faunal transitions driven by global and local environmental changes. In Earth and Life (pp. 677–703). Springer (2012).43.Hughes, T. P. et al. Spatial and temporal patterns of mass bleaching of corals in the Anthropocene. Science 359(6371), 80–83. https://doi.org/10.1126/science.aan8048 (2018).ADS 
CAS 
Article 
PubMed 
PubMed Central 

Google Scholar 
44.Racki, G. A volcanic scenario for the Frasnian-Fammenian major biotic crisis and other Late Devonian global changes: More answers than questions?. Global Planet. Change 189, 103174 (2020).Article 

Google Scholar 
45.Kiessling, W. & Simpson, C. On the potential for ocean acidification to be a general cause of ancient reef crises. Glob. Change Biol. 17(1), 56–67 (2011).ADS 
Article 

Google Scholar 
46.Kowalewski, M. Time-averaging, overcompleteness, and the geological record. J. Geol. 104(3), 317–326 (1996).ADS 
Article 

Google Scholar 
47.Hubert, B. L., Zapalski, M., Nicollin, J. P., Mistiaen, B. & Brice, D. Selected benthic faunas from the Devonian of the Ardennes: an estimation of palaeobiodiversity. Acta Geol. Pol. 57(2), 223–262 (2007).
Google Scholar 
48.Zapalski, M. K. Tabulata (anthozoa) from the givetian and frasnian of the southern region of the holy cross Mts. (Poland). Spec. Pap. Palaeontol. 87, 1–100 (2012).
Google Scholar 
49.Nowiński, A. Tabulata and chaetetida from the devonian and carboniferous of southern Poland. Palaeontol. Pol. 35, 1–125 (1976).
Google Scholar 
50.McWilliam, M. et al. Biogeographical disparity in the functional diversity and redundancy of corals. Proc. Natl. Acad. Sci. 115(12), 3084–3089. https://doi.org/10.1073/pnas.1716643115 (2018).CAS 
Article 
PubMed 
PubMed Central 

Google Scholar 
51.Laliberté, E. & Legendre, P. A distance-based framework for measuring functional diversity from multiple traits. Ecology 91, 299–305 (2010).Article 

Google Scholar 
52.Laliberté, E., Legendre, P., & Shipley, B. (2014). FD: measuring functional diversity from multiple traits, and other tools for functional ecology. R package version 1.0–12.53.Mouillot, D., Graham, N. A. J., Villeger, S., Mason, N. W. H. & Bellwood, D. R. A functional approach reveals community responses to disturbance. Trends Ecol. Evol. 28, 167–177. https://doi.org/10.1016/j.tree.2012.10.004 (2013).Article 
PubMed 
PubMed Central 

Google Scholar 
54.Oksanen, Jari, F. Guillaume Blanchet, Michael Friendly, Roeland Kindt, Pierre Legendre, Dan McGlinn, Peter R. Minchin et al. “Package ‘vegan’.” (2020). More

Pollen collection and RNA extractionPollen is a microscopic and notoriously resistant plant product. Thus, methods to collect a sufficient and roughly equivalent volume of pollen per species, and to ensure RNA was collected from viruses both internal and external to pollen grains, were developed specifically for this work. At each of the four regions, we identified visually asymptomatic plants species that were in full flower and in high enough abundance to achieve our pollen sample minimum. Many of the pollen samples were collected from public roadsides. However, some from the California Grasslands were collected from the University of California’s McLaughlin Natural Reserve, and some from the Eastern Deciduous Agro-forest Interface were collected from the University of Pittsburgh’s Pymatuning Laboratory of Ecology. We had permission to sample in both places. In addition, we obtained permission from the USDA Forest Service to sample in the Till Ridge Cove area of the Chattahoochee-Oconee National Forest for sampling in Central Appalachia. None of the sampled plants displayed classic viral symptoms (e.g., leaf yellowing, vein clearing, leaf distortions, growth abnormalities). To achieve the broadest representation of plant species, we selected species in different families, where feasible. Also when possible, we focused primarily on perennial species to avoid any effects of life history variation. From these, we collected 30 to 50 mg of pollen from newly dehiscing anthers (3–967 fresh hermaphroditic flowers from 1–27 plants per species; Supplementary Table 3) in situ using a sterile sonic dismembrator (Fisherbrand Model 50, Fisher Scientific, Waltham, MA, USA) with a frequency of 20 Hz. We removed non-pollen tissues (e.g., anther debris) with sterile forceps. In addition to removing non-pollen debris that was visible to the naked eye in the field at the time of pollen sample collection, we conducted microscopic and gene expression analyses to confirm the purity of the pollen samples in the lab (Supplementary Methods). Visibly pure pollen from a single species was transferred to a 2-mL collection tube with Lysing Matrix D (MP Biomedicals, Irvine, CA, USA) and kept on dry ice until transported to and stored at −80°C at the University of Pittsburgh (Pittsburgh, PA, USA).Before extracting the total RNA, we freeze-dried the pollen samples (FreeZone 4.5 Liter Benchtop Freeze Dry System, Labconco Corporation, Kansas City, MO, USA) and lysed with a TissueLyser II (Qiagen, Inc., Germantown, MD, USA) at 30 Hz with varying times for different plant species (Supplementary Table 3). We confirmed via microscopy that this protocol resulted in the breakage of ≥50% of the pollen grains in a sample. The total RNA, including dsRNA, was extracted using the Quick-RNA Plant Miniprep Extraction Kit (Zymo Research Corporation, Irvine, CA, USA), following the full manufacturer’s protocol, including the optional steps of in-column DNA digestion and inhibitor removal.RNA sequencingWe assessed the quantity and quality of the total RNA extracted from each pollen sample with a Qubit 2.0 fluorometer (Invitrogen, ThermoFisher Scientific, Waltham, MA, USA) and with TapeStation analyses performed by the Genomics Research Core (GRC) at the University of Pittsburgh. Only samples with an RNA integrity value of ≥1.9 were used (Supplementary Table 3). Stranded RNA libraries were prepared by the GRC using the TruSeq Total RNA Library Kit (Illumina, Inc., San Diego, CA, USA), and ribosomal depletion was performed using a RiboZero Plant Leaf Kit (Illumina, Inc., San Diego, CA, USA). At the GRC, we pooled depleted RNA libraries from six species on a single lane of an Illumina NextSeq500 platform.Pre-virus detection stepsA sequencing depth of 117–260 million 75 bp paired-end reads was achieved per sample (Supplementary Table 3). Sequences were demultiplexed and trimmed of adapter sequences. We used the Pickaxe pipeline42,60,61 to detect known and novel pollen-associated viruses. First, Pickaxe removes poor-quality raw reads42,60,61 and aligns the quality-filtered reads using the Bowtie2 aligner with default parameters62 to a subtraction library. Each customized subtraction library contained the host plant species genome or the most closely related plant genomes in the National Center for Biotechnology Information (NCBI) database, if the host plant genome was not available (Supplementary Table 7), as well as other possible contaminant genomes (e.g., the human genome)42,60,61. The subtraction libraries with 1–8 closely related plant genomes, a bioinformatically tractable amount, were used to remove plant sequences, which allows for a conservative estimate of the viruses associated with pollen to be made. The size of the subtraction libraries did not influence the number of identified viruses, as there was no correlation between library size and either estimate of virus richness (conservative: r = 0.08, P = 0.75; relaxed: r = 0.06, P = 0.77). After subtraction, only non-plant reads remained and were used for viral detection.Known RNA virus detection, identity confirmationWith Pickaxe, we used the Bowtie2 aligner with default parameters62 (v2.3.4.2-3) to align viral non-plant reads to Viral RefSeq42,60,61 (hereafter, VRS; Index of /refseq/release/viral (nih.gov)). Each known virus reflects the top hit of an alignment to VRS42,60,61. Following Cantalupo et al.42, we considered a known virus to be present if the viral reads covered at least 20% of the top hit and aligned to it at least ten times. For viruses with segmented genomes, at least one segment was required to meet these criteria.Contig annotation and extension; novel RNA viral genome detection, identity confirmationViral reads were assembled into contigs using the CLC Assembly Cell (Qiagen Digital Insights, Redwood City, CA, USA), and Pickaxe was used to remove repetitive, short ( More

Female plants must not only allocate resources to flowering but also to producing seeds as well as fruits and/or cones. This suggests that the costs of reproduction are higher for female than male plants, or for female function of hermaphrodite plants. Some dioecious plants (i.e. separate male and female plants) are vegetatively very different (i.e. dimorphic) between the sexes, such as females having larger branch and leaf sizes. Differences between male and female resource allocation to reproduction and the possible consequences of this for vegetative dimorphism in dioecious plants, is a central issue in plant evolution but it is a controversial and difficult topic1,2. In the most highly cited paper on this topic, Obeso1 notes it is practically impossible to measure the direct costs of male and female allocation to sexual reproduction. For example, most vascular plant species (about 95%) are hermaphrodites which makes measuring direct allocation by the two sexes, difficult. Thus Paterno et al.3 used an indirect allometric method to measure sexual allocation in hermaphroditic inflorescences and concluded that larger flowers represent greater relative allocation to male function.The problem of shared sexual allocation to inflorescences is avoided in dioecious plants making them important tests for ideas of sexual allocation in plants. However, they are both rare as species and as individuals and are typically large, forest trees. For example, there are relatively few dioecious individual trees in the very large Barro Colorado forest tree data set4. Again, this large size makes direct measurement, such as of allocation to reproductive structures, difficult. The Cape Floral Region is a useful place to investigate sexual allocation in plants and its consequences, because dioecy is relatively common and vegetative dimorphism between the sexes can be extreme. Also, Cape plants are amenable to research being short (about 2–5 m), rapidly mature and short-lived (about 5–20 years). Thus, the large (about 85 spp.) Cape genus Leucadendron (Proteaceae) is probably the most researched genus globally for male and female differences5,6,7,8,9,10,11,12,13,14.Even in these dioecious plants it is difficult to directly measure allocation to male and female function because of the difficulty of finding a common currency to compare allocation. For example, comparing allocation differences in attractiveness, nectar, seeds, pollen, cones and fruits and differences in the timing of producing these structures1. In Leucadendron males are generally more visually attractive than females. This is achieved by the loss of photosynthetic capacity in floral leaves and bracts12,15. It would be difficult to directly compare this photosynthetic loss in males, with for instance, female allocation to cones and seeds. Despite the difficulties in directly measuring and comparing allocation to reproduction, the consensus is that female allocation to sexual reproduction typically exceeds male allocation1, including in Leucadendron2,9.Greater female allocation to reproduction is one of the suggested reasons for vegetative dimorphism between the sexes2. The three main hypotheses for sexual vegetative dimorphism are (i) greater female sexual resource allocation requires this to be balanced by having a more efficient physiology (resource use efficiency hypothesis), or (ii) greater female allocation requires females to be in the more optimum habitats (the sexual site dimorphism hypothesis) and this facilitates vegetative differences, such as larger female leaves in the more mesic habitats. Finally, (iii) vegetative dimorphism may be a consequence of selection on reproductive traits (reproductive traits hypothesis). In support of the resource use efficiency hypothesis in Leucadendron, Harris and Pannell9 argue that supplying water to live, closed cones in the canopy of serotinous Leucadendron females is a form of maternal care that non-serotinous species and males do not incur. To keep these cones from opening they need always to be hydrated and therefore serotinous females need to be more efficient in their water use than their males. They argued that fewer and thicker branches in females provides a hydraulic advantage. However, the data in Midgley8 and Roddy et al.14 showed no support for sexual differences in water use efficiency. Clearly, there are opposing views as to whether females allocate more to reproduction than males and whether females are eco-physiologically more efficient than males.The sexual site dimorphism hypothesis has not been tested for Leucadendron presumably because males and females co-occur on a small spatial scale16 but is tested in the present analysis of Aulax umbellata and A. cancellata. In support of the reproductive trait’s hypothesis, it was argued5 that in Leucadendron, vegetative dimorphism is an allometric consequence of selection for smaller male inflorescences. Smaller inflorescences are then associated with more, but narrower, stems and thus smaller leaves via Corners Rules5. Besides the evolutionary relevance for understanding sexual differences in allocation, it may also have conservation implications. For example, Hultine et al.17 argued that dioecious plants are under more threat than hermaphrodites because dioecious females are presumed to allocate more resources to reproduction than males. As global change progresses, females may suffer greater mortality and thus dioecious populations may have lower reproductive potential if they become more male biased.One way around the measurement problem of determining direct allocation to sexual reproduction is to use indirect methods based on trade-offs1 such as the influence of allocation to sexual reproduction, on sex ratios and sizes of co-occurring male and female plants. If for example, males allocated less to reproduction than co-occurring females, they should be relatively larger or live longer and this would impact size and sex ratios, especially as plants age and competition intensifies.
The Cape is uniquely suitable to consider allocation differences between the sexes because populations of dioecious Cape species are often large ( > 1000’s of plants ha−1) and with males and females co-existing at a fine spatial scale. The Cape Proteaceae grow in a stressful summer dry Mediterranean climate with nutrient-poor soils18. This provides strong selection on reproductive allocation to seeds (such as large size and high nutrient concentrations) to produce seedlings large enough to survive their first summer. The Cape Proteaceae are strongly fire-adapted. For example, many species are serotinous (canopy storage of seeds in live, closed cones which mainly open after fire)19. This too requires high female sex allocation to maintaining cones in the canopy. Most Cape Proteaceae species are post-fire re-seeders19 in that all plants die in fire. This results in single-aged populations of single-stemmed non-clonal individuals; adults die in fires and dense patches of seedlings establish in the first winter after the fire and die in the next fire. Co-occurring males and females have the same age and thus differences in size or sex ratios will mostly reflect allocation differences and competition rather than age or habitat. Also, because seedlings in the Cape grow up in an open post-fire environment, woody plants do not need to allocate specifically to height growth, to achieve full light. They are in full light their whole lives and therefore any sexual architectural differences do not reflect differences in habitat shadiness. Here we focused on Aulax umbellata, but also present sex ratios and size metrics for the congeneric A. cancellata. These are two common, single-stemmed strongly serotinous Cape species in the Proteaceae which are highly vegetatively dimorphic. Although both Leucadendron and Aulax are dioecious, a rare trait in the family, this represents independent evolution as the two genera are not close phylogenetically20. We test the hypothesis that vegetative sexual dimorphism in Aulax umbellata and Aulax cancellata can be explained by differences in allocation to growth. We predicted that co-occurring males and females would occur in equal sex ratios and be equal in size due to equal growth, despite vegetative dimorphism. More