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    Beating in on a stable partnership

    1.Visick, K. L., Stabb, E. V. & Ruby, E. G. A lasting symbiosis: how Vibrio fischeri finds a squid partner and persists within its natural host. Nat. Rev. Microbiol. https://doi.org/10.1038/s41579-021-00557-0 (2021).Article 

    Google Scholar 
    2.Nyholm, S. V. & McFall-Ngai, M. J. A lasting symbiosis: how the Hawaiian bobtail squid finds and keeps its bioluminescent bacterial partner. Nat. Rev. Microbiol. https://doi.org/10.1038/s41579-021-00567-y (2021).Article 

    Google Scholar 
    3.Koch, E. J., Moriano-Gutierrez, S., Ruby, E. G., McFall-Ngai, M. & Liebeke, M. The impact of persistent colonization by Vibrio fischeri on the metabolome of the host squid Euprymna scolopes. J. Exp. Biol. 223, (2020).4.Schwartzman, J. A. et al. The chemistry of negotiation: rhythmic, glycan-driven acidification in a symbiotic conversation. Proc. Natl Acad. Sci. 112, 566–571 (2015).CAS 
    Article 

    Google Scholar 
    5.Brooks, J. F. & Mandel, M. J. The histidine kinase BinK is a negative regulator of biofilm formation and squid colonization. J. Bacteriol. 198, 2596–2607 (2016).Article 

    Google Scholar 
    6.Bultman, K. M., Cecere, A. G., Miyashiro, T., Septer, A. N. & Mandel, M. J. Draft genome sequences of type VI secretion system-encoding Vibrio fischeri strains FQ-A001 and ES401. Microbiol. Resour. Announc. 8, e00385-19 (2019).Article 

    Google Scholar 
    7.Guckes, K. R. et al. Incompatibility of Vibrio fischeri strains during symbiosis establishment depends on two functionally redundant hcp genes. J. Bacteriol. 201, e00221-19 (2019).Article 

    Google Scholar 
    8.Moriano-Gutierrez, S. et al. The noncoding small RNA SsrA is released by Vibrio fischeri and modulates critical host responses. PLoS Biol. 18, e3000934 (2020).CAS 
    Article 

    Google Scholar 
    9.Koehler, S. et al. The model squid–vibrio symbiosis provides a window into the impact of strain-and species-level differences during the initial stages of symbiont engagement. Environ. Microbiol. 21, 3269–3283 (2019).CAS 
    Article 

    Google Scholar 
    10.Bosch, T. C. G. & Hadfield, M. G. Cellular Dialogues in the Holobiont (CRC Press, 2020). More

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    Vocal universals and geographic variations in the acoustic repertoire of the common bottlenose dolphin

    1.Foster, S. A. & Endler, J. A. Geographic Variation in Behavior: Perspectives on Evolutionary Mechanisms 1–336 (Oxford University Press, 1999).Book 

    Google Scholar 
    2.Mundiger, P. C. Microgeographic and macrogeographic variation in the acquired vocalizations of birds. In Acoustic Communication in Birds 147–208 (Academic Press, 1982).
    Google Scholar 
    3.Green, S. Dialects in Japanese monkeys: Vocal learning and cultural transmission of locale-specific vocal behavior?. Z. Tierpsychol. J. Comp. Ethol. 38(3), 304–314 (1975).CAS 
    Article 

    Google Scholar 
    4.Hodun, A., Snowdon, C. T. & Soini, P. Subspecific variation in the long calls of the tamarin, Saguinus fusckollis. Z. Tierpsychol. 57, 97–110 (1981).Article 

    Google Scholar 
    5.Ford, J. K. B. & Fisher, H. D. Group-specific dialects of killer whales (Orcinus orca) in British Columbia. In Communication and Behavior of Whales 129–161 (Westview Press for the American Association for the Advancement of Science, 1983).
    Google Scholar 
    6.Filatova, O. A. et al. Call diversity in the North Pacific killer whale populations: Implications for dialect evolution and population history. Anim. Behav. 83, 595–603 (2012).Article 

    Google Scholar 
    7.Rendell, L. E. & Whitehead, H. Vocal clans in sperm whales (Physeter macrocephalus). Proc. Biol. Sci. R. Soc. 270, 225–231 (2003).CAS 
    Article 

    Google Scholar 
    8.Gero, S., Whitehead, H. & Rendell, L. Individual, unit and vocal clan level identity cues in sperm whale codas. R. Soc. Open Sci. 3, 1–12 (2016).
    Google Scholar 
    9.Cise, A. M., Van Mahaffy, S. D., Baird, R. W., Mooney, T. A. & Barlow, J. Song of my people: Dialect differences among sympatric social groups of short-finned pilot whales in Hawai’i. Behav. Ecol. Sociobiol. 72, 1–13 (2018).Article 

    Google Scholar 
    10.Podos, J. & Warren, P. S. The evolution of geographic variation in birdsong. Adv. Study Behav. 37, 403–458 (2007).Article 

    Google Scholar 
    11.Walker, T. J. Factors responsible for intraspecific variation in the calling songs of crickets. Evolution 16, 407–428 (1962).Article 

    Google Scholar 
    12.Velásquez, N. A. Geographic variation in acoustic communication in anurans and its neuroethological implications. J. Physiol. 108, 167–173 (2014).
    Google Scholar 
    13.Amorim, T. O. S., Andriolo, A., Reis, S. S. & dos Santos, M. E. Vocalizations of Amazon river dolphins (Inia geoffrensis): Characterization, effect of physical environment and differences between populations. J. Acoust. Soc. Am. 139, 1285–1293 (2016).ADS 
    PubMed 
    Article 

    Google Scholar 
    14.Moron, J. R. et al. Spinner dolphin whistle in the Southwest Atlantic Ocean: Is there a geographic variation?. J. Acoust. Soc. Am. 138, 2495–2498 (2015).ADS 
    PubMed 
    Article 

    Google Scholar 
    15.Bjørgesæter, A., Ugland, K. I. & Bjørge, A. Geographic variation and acoustic structure of the underwater vocalization of harbor seal (Phoca vitulina) in Norway, Sweden and Scotland. J. Acoust. Soc. Am. 116, 2459–2468 (2004).ADS 
    PubMed 
    Article 

    Google Scholar 
    16.Janik, V. & Slater, P. The different roles of social learning in vocal communication. Anim. Behav. 60, 1–11 (2000).CAS 
    PubMed 
    Article 

    Google Scholar 
    17.Lameira, A. R., Delgado, R. A. & Wich, S. A. Review of geographic variation in terrestrial mammalian acoustic signals: Human speech variation in a comparative perspective. J. Evol. Psychol. 8, 309–332 (2010).Article 

    Google Scholar 
    18.Janik, V. Acoustic communication networks in marine mammals. In Animal Communication Networks 390–415 (University Press, 2005).
    Google Scholar 
    19.Deecke, V. B., Ford, J. K. B. & Spong, P. Dialect change in resident killer whales: Implications for vocal learning and cultural transmission. Anim. Behav. 60, 629–638 (2000).CAS 
    PubMed 
    Article 

    Google Scholar 
    20.Weilgart, L. & Whitehead, H. Group-specific dialects and geographical variation in coda repertoire in South Pacific sperm whales. Behav. Ecol. Sociobiol. 40, 277–285 (1997).Article 

    Google Scholar 
    21.Azevedo, A. F. & Van Sluys, M. Whistles of tucuxi dolphins (Sotalia fluviatilis) in Brazil: Comparisons among populations. J. Acoust. Soc. Am. 117, 1456–1464 (2005).ADS 
    PubMed 
    Article 

    Google Scholar 
    22.Bazúa-Durán, C. & Au, W. W. L. Geographic variations in the whistles of spinner dolphins (Stenella longirostris) of the Main Hawaiian Islands. J. Acoust. Soc. Am. 116, 3757–3769 (2004).ADS 
    PubMed 
    Article 

    Google Scholar 
    23.Hawkins, E. R. Geographic variations in the whistles of bottlenose dolphins (Tursiops aduncus) along the east and west coasts of Australia. J. Acoust. Soc. Am. 128, 924–935 (2010).ADS 
    CAS 
    PubMed 
    Article 

    Google Scholar 
    24.Wang, D., Würsig, B. & Evans, W. Whistles of bottlenose dolphins: Comparisons among populations. Aquat. Mamm. 21, 65–77 (1995).
    Google Scholar 
    25.Connor, R. C., Wells, R. S., Mann, J. & Read, A. J. The bottlenose dolphin: Social relationships in a fission–fusion society. In Cetacean Societies: Field Studies of Dolphins and Whales 91–126 (The University of Chicago Press, 2000).
    Google Scholar 
    26.Costa, A. P. B. et al. Ecological divergence and speciation in common bottlenose dolphins in the western South Atlantic. J. Evol. Biol. 34, 16–32 (2021).PubMed 
    Article 

    Google Scholar 
    27.Hoelzel, A. R., Potter, C. W. & Best, P. B. Genetic differentiation between parapatric “nearshore” and “offshore” populations of the bottlenose dolphin. Proc. R. Soc. Lond. B 265, 1177–1183 (1998).CAS 
    Article 

    Google Scholar 
    28.Louis, M. et al. Habitat-driven population structure of bottlenose dolphins, Tursiops truncatus, in the North-East Atlantic. Mol. Ecol. 23, 857–874 (2014).PubMed 
    Article 

    Google Scholar 
    29.Wells, R. S., Natoli, A. & Braulik, G. Tursiops truncatus. The IUCN Red List of Threatened Species (2019).30.Marino, L. et al. Cetaceans have complex brains for complex cognition. PLoS Biol. 5, 966–972 (2007).CAS 
    Article 

    Google Scholar 
    31.Janik, V. & Slater, P. Context-specific use suggests that bottlenose dolphin signature whistles are cohesion calls. Anim. Behav. 56, 829–838 (1998).CAS 
    PubMed 
    Article 

    Google Scholar 
    32.Sayigh, L. et al. Individual recognition in wild bottlenose dolphins: a field test using playback experiments. Anim. Behav. 57, 41–50 (1999).CAS 
    PubMed 
    Article 

    Google Scholar 
    33.Au, W. W. L. Echolocation signals of wild dolphins. Acoust. Phys. 50, 454–462 (2004).ADS 
    Article 

    Google Scholar 
    34.Herzing, D. & dos Santos, M. E. Functional aspects of echolocation in dolphins. In Echolocation in Bats and Dolphins 386–393 (The University of Chicago Press, 2004).
    Google Scholar 
    35.Jensen, F. H., Bejder, L., Wahlberg, M. & Madsen, P. T. Biosonar adjustments to target range of echolocating bottlenose dolphins (Tursiops sp.) in the wild. J. Exp. Biol. 212, 1078–1086 (2009).CAS 
    PubMed 
    Article 

    Google Scholar 
    36.Diáz-López, B. & Shirai, J. Mediterranean common bottlenose dolphin’s repertoire and communication use. In Dolphins: Anatomy, Behavior, and Threats 129–148 (Nova Science Publishers, 2009).
    Google Scholar 
    37.Herzing, D. L. Acoustics and social behavior of wild dolphins: Implications for a sound society. In Hearing by Whales and Dolphins Springer Handbook of Auditory Research 225–272 (Springer, 2000).
    Google Scholar 
    38.dos Santos, M. E., Ferreira, A. J. & Harzen, S. Rhythmic sound sequences emitted by aroused bottlenose dolphins in the Sado estuary, Portugal. In Sensory Systems of Aquatic Mammals 325–334 (De Spil Publishers, 1995).
    Google Scholar 
    39.Luís, A. R., Alves, I. S., Sobreira, F. V., Couchinho, M. N. & dos Santos, M. E. Brays and bits: Information theory applied to acoustic communication sequences of bottlenose dolphins. Bioacoustics 28, 286–296 (2019).Article 

    Google Scholar 
    40.Jones, B., Zapetis, M., Samuelson, M. M. & Ridgway, S. Sounds produced by bottlenose dolphins (Tursiops): A review of the defining characteristics and acoustic criteria of the dolphin vocal repertoire. Bioacoustics 29(4), 399–440 (2020).Article 

    Google Scholar 
    41.May-Collado, L. J. & Wartzok, D. A. comparison of bottlenose dolphin whistles in the Atlantic ocean: Factors promoting whistle variation. J. Mammal. 89, 1229–1240 (2008).Article 

    Google Scholar 
    42.Jones, G. J. & Sayigh, L. S. Geographic variation in rates of vocal production of free-ranging bottlenose dolphins. Mar. Mamm. Sci. 18, 374–393 (2002).Article 

    Google Scholar 
    43.La Manna, G. et al. Assessing geographical variation on whistle acoustic structure of three Mediterranean populations of common bottlenose dolphin (Tursiops truncatus). Behaviour 154, 583–607 (2017).Article 

    Google Scholar 
    44.Papale, E. et al. Acoustic divergence between bottlenose dolphin whistles from the Central-Eastern North Atlantic and Mediterranean Sea. Acta Ethologica 17, 155–165 (2014).Article 

    Google Scholar 
    45.R Development Core Team. R: A Language and Environment for Statistical Computing (2018).46.Wickham, H. ggplot2: Elegant Graphics for Data Analysis. https://ggplot2.tidyverse.org (Springer, 2016).47.Mccomb, K. & Semple, S. Coevolution of vocal communication and sociality in primates. Biol. Lett. 1, 381–385 (2005).PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    48.Leighton, G. M. Cooperative breeding influences the number and type of vocalizations in avian lineages. Proc. R. Soc. B Biol. Sci. 284, 1–9 (2017).
    Google Scholar 
    49.Freeberg, T. M., Dunbar, R. I. M. & Ord, T. J. Social complexity as a proximate and ultimate factor in communicative complexity. Philos. Trans. R. Soc. B Biol. Sci. 367, 1785–1801 (2012).Article 

    Google Scholar 
    50.Pollard, K. A. & Blumstein, D. T. Evolving communicative complexity: insights from rodents and beyond. Philos. Trans. R. Soc. B Biol. Sci. 367, 1869–1878 (2012).Article 

    Google Scholar 
    51.Gustison, M. L., Le Roux, A. & Bergman, T. J. Derived vocalizations of geladas (Theropithecus gelada) and the evolution of vocal complexity in primates. Philos. Trans. R. Soc. B Biol. Sci. 367, 1847–1859 (2012).Article 

    Google Scholar 
    52.Augusto, J. F., Rachinas-Lopes, P. & dos Santos, M. E. Social structure of the declining resident community of common bottlenose dolphins in the Sado Estuary, Portugal. J. Mar. Biol. Assoc. U. K. 92, 1773–1782 (2012).Article 

    Google Scholar 
    53.Luís, A. R., Couchinho, M. N. & dos Santos, M. E. Changes in the acoustic behavior of resident bottlenose dolphins near operating vessels. Mar. Mamm. Sci. 30, 1417–1426 (2014).Article 

    Google Scholar 
    54.Ridgway, S. H., Moore, P. W., Carder, D. A. & Romano, T. A. Forward shift of feeding buzz components of dolphins and belugas during associative learning reveals a likely connection to reward expectation, pleasure and brain dopamine activation. J. Exp. Biol. 217, 2910–2919 (2014).CAS 
    PubMed 
    Article 

    Google Scholar 
    55.Luís, A. R., Couchinho, M. N. & dos Santos, M. E. A quantitative analysis of pulsed signals emitted by wild bottlenose dolphins. PLoS ONE 11, 1–11 (2016).
    Google Scholar 
    56.Nowacek, D. P. Acoustic ecology of foraging bottlenose dolphins (Tursiops truncatus) habitat-specific use of three sound types. Mar. Mamm. Sci. 21, 587–602 (2005).Article 

    Google Scholar 
    57.Caldwell, M. C., Caldwell, D. K. & Tyack, P. L. Review of the signature-whistle-hypothesis for the Atlantic bottlenose dolphin, Tursiops truncatus. In The Bottlenose Dolphin 199–234 (Academic Press, 1990).
    Google Scholar 
    58.Laland, K. N. & Janik, V. M. The animal cultures debate. Evolution 21, 542–547 (2006).
    Google Scholar 
    59.Kershenbaum, A., Sayigh, L. S. & Janik, V. M. The encoding of individual identity in dolphin signature whistles: How much information is needed?. PLoS ONE 8, e77671 (2013).ADS 
    CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    60.King, S. L. & Janik, V. M. Bottlenose dolphins can use learned vocal labels to address each other. Proc. Natl. Acad. Sci. U.S.A. 110, 13216–13221 (2013).ADS 
    CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    61.Sayigh, L., Esch, H., Wells, R. & Janik, V. Facts about signature whistles of bottlenose dolphins, Tursiops truncatus. Anim. Behav. 74, 1631–1642 (2007).Article 

    Google Scholar 
    62.Buckstaff, K. C. Effects of watercraft noise on the acoustic behavior of bottlenose dolphins, Tursiops truncatus, in Sarasota Bay, Florida. Mar. Mamm. Sci. 20, 709–725 (2004).Article 

    Google Scholar 
    63.Morisaka, T., Shinohara, M., Nakahara, F. & Akamatsu, T. Geographic variations in the whistles among three Indo-Pacific bottlenose dolphin. Fish. Sci. 71, 568–576 (2005).CAS 
    Article 

    Google Scholar 
    64.May-Collado, L. J. & Quiñones-Lebrón, S. G. Dolphin changes in whistle structure with watercraft activity depends on their behavioral state. J. Soc. Am. 135, EL193–EL198 (2014).ADS 

    Google Scholar 
    65.Garland, E. C. et al. Report dynamic horizontal cultural transmission of humpback whale song at the ocean basin scale. Curr. Biol. 21, 687–691 (2011).CAS 
    PubMed 
    Article 

    Google Scholar 
    66.Whitehead, H. & Rendell, L. The Cultural Lives of Whales and Dolphins (The University of Chicago Press, 2015).
    Google Scholar 
    67.Herzing, D. L. Vocalizations and associated underwater behavior of free-ranging Atlantic spotted dolphins, Stenella frontalis and bottlenose dolphins, Tursiops truncatus. Aquat. Mamm. 22, 61–79 (1996).
    Google Scholar 
    68.May-Collado, L. J. Changes in whistle structure of two dolphin species during interspecific associations. Ethology 116, 1065–742010 (2010).Article 

    Google Scholar 
    69.Catchpole, C. K. The evolution of bird sounds in relation to mating and spacing behavior. In Acoustic Communication in Birds 297–319 (Academic Press, 1982).
    Google Scholar 
    70.Herman, L. M. The multiple functions of male song within the humpback whale (Megaptera novaeangliae) mating system: Review, evaluation, and synthesis. Biol. Rev. 92, 1795–1818 (2017).PubMed 
    Article 

    Google Scholar 
    71.Janik, V. M. Food-related bray calls in wild bottlenose dolphins (Tursiops truncatus). Proc. R. Soc. B Biol. Sci. 267, 923–927 (2000).CAS 
    Article 

    Google Scholar 
    72.King, S. L. & Janik, V. M. Come dine with me: food-associated social signalling in wild bottlenose dolphins (Tursiops truncatus). Anim. Cogn. 18, 969–974 (2015).PubMed 
    Article 

    Google Scholar 
    73.Herzing, D. L. Synchronous and rhythmic vocalizations and correlated underwater behavior of free-ranging Atlantic Spotted Dolphins (Stenella frontalis) and Bottlenose Dolphins (Tursiops truncatus) in the Bahamas. Anim. Behav. Cogn. 2, 14–29 (2015).Article 

    Google Scholar 
    74.Pleslić, G. et al. The abundance of common bottlenose dolphins (Tursiops truncatus) in the former special marine reserve of the Cres-Lošinj Archipelago, Croatia. Aquat. Conserv. Mar. Freshwat. Ecosyst. 25, 125–137 (2015).Article 

    Google Scholar 
    75.Rako-Gospic, N., Radulovi, M., Vu, T., Plesli, G. & Mackelworth, P. Factor associated variations in the home range of a resident Adriatic common bottlenose dolphin population. Mar. Pollut. Bull. 124, 234–244 (2017).CAS 
    PubMed 
    Article 

    Google Scholar 
    76.Rako, N. et al. Leisure boating noise as a trigger for the displacement of the bottlenose dolphins of the Cres-Lošinj archipelago (northern Adriatic Sea, Croatia). Mar. Pollut. Bull. 68, 77–84 (2013).CAS 
    PubMed 
    Article 

    Google Scholar 
    77.Barragán-Barrera, D. C. et al. High genetic structure and low mitochondrial diversity in bottlenose dolphins of the Archipelago of Bocas del Toro, Panama: A population at risk?. PLoS ONE 12, 1–22 (2017).Article 
    CAS 

    Google Scholar 
    78.Ey, E. & Fischer, J. The, “Acoustic Adaptation Hypothesis”—A review of the evidence from birds, anurans and mammals. Bioacoustics 19, 21–48 (2009).Article 

    Google Scholar 
    79.Papale, E., Azzolin, M. & Giacoma, C. Vessel traffic affects bottlenose dolphin (Tursiops truncatus) behaviour in waters surrounding Lampedusa Island, south Italy. J. Mar. Biol. Assoc. U.K. 92, 1877–1885 (2012).Article 

    Google Scholar 
    80.Gridley, T., Nastasi, A., Kriesell, H. J. & Elwen, S. H. The acoustic repertoire of wild common bottlenose dolphins (Tursiops truncatus) in Walvis Bay, Namibia. Bioacoustics 24, 153–174 (2015).Article 

    Google Scholar 
    81.Au, W. W. L. & Hastings, M. C. Emission of social sounds by marine animals. In Principles of Marine Bioacoustics 401–499 (Springer, 2008).
    Google Scholar 
    82.Bázua-Duran, C. & Bazúa-Durán, C. Differences in the whistle characteristics and repertoire of Bottlenose and Spinner Dolphins. An. Acad. Bras. Ciênc. 76, 386–392 (2004).PubMed 
    Article 

    Google Scholar 
    83.Lammers, M. O., Au, W. W. L. & Herzing, D. L. The broadband social acoustic signaling behavior of spinner and spotted dolphins. J. Acoust. Soc. Am. 114, 1629–1639 (2003).ADS 
    PubMed 
    Article 

    Google Scholar 
    84.Simard, P. et al. Low frequency narrow-band calls in bottlenose dolphins (Tursiops truncatus): Signal properties, function, and conservation implications. J. Acoust. Soc. Am. 130, 3068–3076 (2011).ADS 
    PubMed 
    Article 

    Google Scholar 
    85.Luís, A. R., Couchinho, M. N. & dos Santos, M. E. Signature whistles in wild bottlenose dolphins: Long-term stability and emission rates. Acta Ethologica 19, 113–122 (2016).Article 

    Google Scholar 
    86.Ford, J. K. B. Vocal traditions among resident killer whales (Orcinus orca) in coastal waters of British Columbia. Can. J. Zool. 69, 1454–1483 (1991).Article 

    Google Scholar 
    87.Papale, E. et al. Biphonic calls as signature whistles in a free-ranging bottlenose dolphin. Bioacoustics 24, 223–231 (2015).Article 

    Google Scholar 
    88.Elliser, C. R. & Herzing, D. L. Long-term interspecies association patterns of Atlantic bottlenose dolphins, Tursiops truncatus, and Atlantic spotted dolphins, Stenella frontalis, in the Bahamas. Mar. Mamm. Sci. 32, 38–56 (2015).Article 

    Google Scholar 
    89.Hoffmann-Kuhnt, M., Herzing, D. L., Ho, A. & Chitre, M. A. Whose line sound is it anyway? Identifying the vocalizer on underwater video by localizing with a hydrophone array. Anim. Behav. Cogn. 3, 288–298 (2016).Article 

    Google Scholar 
    90.Lima, I. M. S. et al. Whistle comparison of four delphinid species in Southeastern Brazil. J. Acoust. Soc. Am. 139, EL124 (2016).ADS 
    PubMed 
    Article 

    Google Scholar  More

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    New evidence from exceptionally “well-preserved” specimens sheds light on the structure of the ammonite brachial crown

    1.Klug, C. & Lehmann, J. Soft part anatomy of ammonoids: reconstructing the animal based on exceptionally preserved specimens and actualistic comparisons. in Ammonoid Paleobiology: From Anatomy to Ecology 507–529 (Springer, 2015).2.Klug, C. et al. Anatomy and evolution of the first Coleoidea in the Carboniferous. Commun. Biol. 2, 1–12 (2019).Article 

    Google Scholar 
    3.Klug, C., Schweigert, G., Tischlinger, H. & Pochmann, H. Failed prey or peculiar necrolysis? Isolated ammonite soft body from the Late Jurassic of Eichstätt (Germany) with complete digestive tract and male reproductive organs. Swiss J. Palaeontol. 140, 1–14 (2021).PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    4.Maeda, H. & Seilacher, A. Ammonoid taphonomy. In Ammonoid paleobiology 543–578 (Springer, 1996).5.Wani, R. & Gupta, N. S. Ammonoid taphonomy. In Ammonoid Paleobiology: from Macroevolution to Paleogeography 5, 555–598 (2015).6.Klug, C. & Vallon, L. H. Regurgitated ammonoid remains from the latest Devonian of Morocco. Swiss J. Palaeontol. 138, 87–97 (2019).Article 

    Google Scholar 
    7.Hoffmann, R., Stevens, K., Keupp, H., Simonsen, S. & Schweigert, G. Regurgitalites—a window into the trophic ecology of fossil cephalopods. J. Geol. Soc. 177, 82–102 (2020).ADS 
    Article 

    Google Scholar 
    8.Gale, A. S., Kennedy, W. J. & Martill, D. Mosasauroid predation on an ammonite-Pseudaspidoceras-from the Early Turonian of south-eastern Morocco. Acta Geol. Pol. 67, 31–46 (2017).ADS 
    CAS 
    Article 

    Google Scholar 
    9.Vullo, R. Direct evidence of hybodont shark predation on Late Jurassic ammonites. Naturwissenschaften 98, 545–549 (2011).ADS 
    CAS 
    PubMed 
    Article 

    Google Scholar 
    10.Ibáñez, C. M. & Keyl, F. Cannibalism in cephalopods. Rev. Fish Biol. Fish. 20, 123–136 (2010).Article 

    Google Scholar 
    11.Lehmann, J., Solarczyk, A. & Friedrich, O. Belemnoid arm hooks from the Middle-Upper Albian boundary interval: taxonomy and palaeoecological significance. Paläontol. Z. 85, 287–302 (2011).Article 

    Google Scholar 
    12.Stevens, G. Palaeobiological and morphological aspects of Jurassic Onychites (cephalopod hooks) and new records from the New Zealand Jurassic. NZ J. Geol. Geophys. 53, 395–412 (2010).Article 

    Google Scholar 
    13.Klug, C., Davesne, D., Fuchs, D. & Argyriou, T. First record of non-mineralized cephalopod jaws and arm hooks from the latest Cretaceous of Eurytania, Greece. Swiss J. Palaeontol. 139, 1–13 (2020).Article 

    Google Scholar 
    14.Engeser, T. & Reitner, J. Beiträge zur Systematik von phragmokontragenden Coleoiden aus dem Untertithonium (Malm zeta,” Solnhofener Plattenkalk”) von Solnhofen und Eichstätt (Bayern). N. Jb. Geol. und Paläont. 527–545 (1981).15.Reitner, J. & Urlichs, M. Echte Weichteilbelemniten aus dem Untertoarcium (Posidonienschiefer) Südwestdeutschlands. N. Jb. Geol. Paläont. 165, 450–465 (1983).
    Google Scholar 
    16.Fuchs, D., Donovan, D. T. & Keupp, H. Taxonomic revision of “Onychoteuthis” conocauda Quenstedt, 1849 (Cephalopoda: Coleoidea). N. Jb. Geol. Pal. A. 270, 245–255 (2013).Article 

    Google Scholar 
    17.Donovan, D. T. & Crane, M. D. The type material of the Jurassic cephalopod Belemnotheutis. Palaeontology 35, 273–296 (1992).
    Google Scholar 
    18.Klug, C., Schweigert, G., Fuchs, D. & Dietl, G. First record of a belemnite preserved with beaks, arms and ink sac from the Nusplingen Lithographic Limestone (Kimmeridgian, SW Germany). Lethaia 43, 445–456 (2010).Article 

    Google Scholar 
    19.Hart, M. B., Hughes, Z., Page, K. N., Price, G. D. & Smart, C. W. Arm hooks of coleoid cephalopods from the Jurassic succession of the Wessex Basin, Southern England. Proc. Geol. Assoc. 130, 326–338 (2019).Article 

    Google Scholar 
    20.Doyle, P. & Shakides, E. V. The Jurassic Belemnite Suborder Belemnotheutina. Palaeontology 47, 983–998 (2004).Article 

    Google Scholar 
    21.Doguzhaeva, L. et al. An Early Triassic gladius associated with soft tissue remains from Idaho, USA—a squid-like coleoid cephalopod at the onset of Mesozoic Era. APP 63, 341–355 (2018).Article 

    Google Scholar 
    22.Doguzhaeva, L. A., Summesberger, H., Mutvei, H. & Brandstaetter, F. The mantle, ink sac, ink, arm hooks and soft body debris associated with the shells in Late Triassic coleoid cephalopod Phragmoteuthis from the Austrian Alps. Palaeoworld 16, 272–284 (2007).Article 

    Google Scholar 
    23.Engeser, T. S. & Clarke, M. R. Cephalopod hooks, both recent and fossil. in Paleontology and Neontology of Cephalopods 133–151 (Elsevier, 1988).24.Johnson, R. G. & Richardson, E. S. Ten-armed fossil cephalopod from the Pennsylvanian of Illinois. Science 159, 526–528 (1968).ADS 
    CAS 
    PubMed 
    Article 

    Google Scholar 
    25.Fuchs, D. & Hoffmann, R. Treatise Online no. 91: Part M, Chapter 10: Arm Armature in Belemnoid Coleoids. Treatise Online (2017).26.Fuchs, D., von Boletzky, S. & Tischlinger, H. New evidence of functional suckers in belemnoid coleoids (Cephalopoda) weakens support for the ‘Neocoleoidea’ concept. J. Molluscan Stud. 76, 404–406 (2010).Article 

    Google Scholar 
    27.Fuchs, D., Heyng, A. M. & Keupp, H. Acanthoteuthis problematica Naef, 1922, an almost forgotten taxon and its role in the interpretation of cephalopod arm armatures. N. Jb. Geol. Pal. A. 269, 241–250 (2013).Article 

    Google Scholar 
    28.Young, R. E., Vecchione, M. & Donovan, D. T. The evolution of coleoid cephalopods and their present biodiversity and ecology. S. Afr. J. Mar. Sci. 20, 393–420 (1998).Article 

    Google Scholar 
    29.Landman, N. H. & Waagé, K. M. Scaphitid ammonites of the Upper Cretaceous (Maastrichtian) Fox Hills Formation in South Dakota and Wyoming. Bull. AMNH 215, 257 (1993).
    Google Scholar 
    30.Kennedy, W. J., Landman, N. H., Cobban, W. A. & Larson, N. L. Jaws and Radulae in Rhaeboceras, a Late Cretaceous Ammonite. 20 (2002).31.Kruta, I., Landman, N., Rouget, I., Cecca, F. & Tafforeau, P. The radula of the Late Cretaceous scaphitid ammonite Rhaeboceras halli (Meek and Hayden, 1856). Palaeontology 56, 9–14 (2013).Article 

    Google Scholar 
    32.Kruta, I., Bardin, J., Smith, C. P. A., Tafforeau, P. & Landman, N. H. Enigmatic hook-like structures in Cretaceous ammonites (Scaphitidae). Palaeontology 63, 301–312 (2020).Article 

    Google Scholar 
    33.Miserez, A. et al. Microstructural and biochemical characterization of the nanoporous sucker rings from Dosidicus gigas. Adv. Mater. 21, 401–406 (2009).CAS 
    Article 

    Google Scholar 
    34.Kulicki, C. & Szaniawski, K. Cephalopod arm hooks from the Jurassic of Poland. Acta Palaeontol. Pol. 17, 379–419 (1972).
    Google Scholar 
    35.Jereb, P. & Roper, C. F. E. FAO Cephalopods of the World No. 4 Vol. 2, Oegopsid and Myopsid squids, 605 (Rome, 2010).36.Riegraf, W. v, Werner, G. & Lörcher, F. Der Posidonienschiefer: Biostratigraphie, Fauna und Fazies des Südwestdeutschen Untertoarciums, 1–195. (F. Enke, 1984)..37.Sasaki, M. A monograph of dibranchiate cephalopods of the Japanese and adjacent waters. J. Coll. Agric. Hokkaido Univ. 20, 1–357 (1929).
    Google Scholar 
    38.Evans, A. A systematic review of the squid family Cranchiidae (Cephalopoda: Oegopsida) in the Pacific Ocean. (PhD diss., Auckland University of Technology, 2018).39.Naef, A. Die fossilen Tintenfische. 322 pp. (1922).40.Kristensen, T. K. Scanning electron microscopy of hook development in Gonatus fabricii (Lichtenstein, 1818) (Mollusca: Cephalopoda). Vidensk. Meddel. Natuirist. Foren. Kjobenhavn. 140, 111–116 (1977).41.Hart, M. B., Arratia, G., Moore, C. & Ciotti, B. J. Life and death in the Jurassic seas of Dorset, Southern England. Proc. Geol. Assoc. 131, 629–638 (2020).Article 

    Google Scholar 
    42.Jenny, D. et al. Predatory behaviour and taphonomy of a Jurassic belemnoid coleoid (Diplobelida, Cephalopoda). Sci. Rep. 9, 1–11 (2019).ADS 
    CAS 
    Article 

    Google Scholar 
    43.Kröger, B., Vinther, J. & Fuchs, D. Cephalopod origin and evolution: a congruent picture emerging from fossils, development and molecules: Extant cephalopods are younger than previously realised and were under major selection to become agile, shell-less predators. BioEssays 33, 602–613 (2011).PubMed 
    Article 
    CAS 

    Google Scholar 
    44.Jereb, P. & Roper, C. F. Cephalopods of the world. An annotated and illustrated catalogue of cephalopod species known to date. Volume 1. Chambered nautiluses and sepioids (Nautilidae, Sepiidae, Sepiadariidae, Idiosepiidae and Spirulidae). 262 (2006).45.Bello, G., Potoschi, A. & Berdar, A. Adult of Ancistrocheirus lesueurii caught in the straits of Messina (Cephalopoda: Ancistrocheiridae). Bollettino Malacologico 29, 259–266 (1993).
    Google Scholar 
    46.Okutani, T. Rare and interesting squid from Japan V.: A gravid female of Ancistrocheirus lesueuri (D’ORBIGNY, 1839) Collected in the Kuroshio Area (Oegopsida: Enoploteuthidae). Venus (Japanese Journal of Malacology) 35, 73–81 (1976).47.Tsuchiya, K. Abralia fasciolata, a new species of enoploteuthid squid from the western Indian Ocean (Cephalopoda: Oegopsida). Bull. Natl. Sci. Museum 17, 69–79 (1991).
    Google Scholar 
    48.Hidaka, K. & Kubodera, T. Squids of the genus Abralia (Cephalopoda: Enoploteuthidae) from the western tropical Pacific with a description of Abralia omiae, a new species. Bull. Mar. Sci. 66, 417–443 (2000).
    Google Scholar 
    49.Bolstad, K. S. R. Systematics of the Onychoteuthidae Gray, 1847 (Cephalopoda: Oegopsida). Zootaxa 2696, 1–186 (2010).Article 

    Google Scholar 
    50.Hoffmann, R., Weinkauf, M. F. G. & Fuchs, D. Grasping the shape of belemnoid arm hooks—a quantitative approach. Paleobiology 43, 304–320 (2017).Article 

    Google Scholar 
    51.Mangold K. Les organes génitaux. In Traité de zoologie, Céphalopodes Tome V fascicule 4, Grassé, P. P (ed). 459–492. (Masson, 1989)52.Rosa, R. & Seibel, B. A. Voyage of the argonauts in the pelagic realm: physiological and behavioural ecology of the rare paper nautilus, Argonauta nouryi. ICES J. Mar. Sci. 67, 1494–1500 (2010).Article 

    Google Scholar 
    53.Jackson, G. D. & O’Shea, S. Unique hooks in the male scaled squid Lepidoteuthis grimaldi. J. Mar. Biol. Ass. 83, 1099–1100 (2003).Article 

    Google Scholar 
    54.Naglik, C., Tajika, A., Chamberlain, J. & Klug, C. Ammonoid locomotion. In Ammonoid Paleobiology: From anatomy to ecology 649–688 (Springer, 2015).55.Hoffmann, R., Lemanis, R., Naglik, C. & Klug, C. Ammonoid buoyancy. In Ammonoid paleobiology: From Anatomy to Ecology 613–648 (Springer, 2015).56.Ebel, K. Swimming abilities of ammonites and limitations. Paläontol. Z. 64, 25–37 (1990).Article 

    Google Scholar 
    57.Cobban, W. A., Walaszczyk, I., Obradovich, J. D. & McKinney, K. C. A USGS zonal table for the Upper Cretaceous middle Cenomanian-Maastrichtian of the Western Interior of the United States based on ammonites, inoceramids, and radiometric ages. U.S. Geol. Surv. Open-File Rep. 1250, 45 (2006).
    Google Scholar 
    58.Landman, N. H., Kennedy, W. J., Cobban, W. A. & Larson, N. L. Scaphites of the “Nodosus Group” from the Upper Cretaceous (Campanian) of the Western Interior of North America. Bull. Am. Mus. Nat. Hist. 342, 1–242 (2010).Article 

    Google Scholar 
    59.Lee, H., Chung, M. K., Kang, H., Kim, B.-N. & Lee, D. S. Computing the Shape of Brain Networks Using Graph Filtration and Gromov-Hausdorff Metric. in Medical Image Computing and Computer-Assisted Intervention–MICCAI 2011. 6892, 302–309 (Springer Berlin Heidelberg, 2011).60.Xia, K. & Wei, G.-W. Persistent homology analysis of protein structure, flexibility, and folding. Int. J. Numer. Methods Biomed. Eng. 30, 814–844 (2014).MathSciNet 
    Article 

    Google Scholar 
    61.Townsend, J., Micucci, C. P., Hymel, J. H., Maroulas, V. & Vogiatzis, K. D. Representation of molecular structures with persistent homology for machine learning applications in chemistry. Nat. Commun. 11, 1–9 (2020).
    Google Scholar 
    62.Xia, K. Persistent homology analysis of ion aggregations and hydrogen-bonding networks. Phys. Chem. Chem. Phys. 13, 13448–13460 (2018).Article 

    Google Scholar 
    63.Krishnapriyan, A. S., Montoya, J., Hummelshøj, J. & Morozov, D. Persistent homology advances interpretable machine learning for nanoporous materials. arXiv:2010.00532 [cond-mat, physics:physics] (2020).64.Fasy, B. T., Kim, J., Lecci, F. & Maria, C. Introduction to the R package TDA. arXiv preprint arXiv:1411.1830 (2014).65.Adler, D., Nenadic, O. & Zucchini, W. Rgl: A r-library for 3d visualization with opengl. in Proceedings of the 35th Symposium of the Interface: Computing Science and Statistics, Salt Lake City 35, 1–11 (2003).66.Roper, C. F., Sweeney, M. J. & Nauen, C. Cephalopods of the world. An annotated and illustrated catalogue of species of interest to fisheries, 277 (FAO Fish Synopsys, 1984). More

  • in

    Environmental and spatial risk factors for the larval habitats of Plasmodium knowlesi vectors in Sabah, Malaysian Borneo

    1.Fornace, K. M. et al. Exposure and infection to Plasmodium knowlesi in case study communities in Northern Sabah, Malaysia and Palawan, The Philippines. PLoS Negl. Trop. Dis. 12, e0006432 (2018).Article 

    Google Scholar 
    2.Singh, B. et al. A large focus of naturally acquired Plasmodium knowlesi infections in human beings. Lancet 363, 1017–1024 (2004).Article 

    Google Scholar 
    3.Chin, A. Z. et al. Malaria elimination in Malaysia and the rising threat of Plasmodium knowlesi. J. Physiol. Anthropol. https://doi.org/10.1186/s40101-020-00247-5 (2020).Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    4.Cooper, D. J. et al. Plasmodium knowlesi Malaria in Sabah, Malaysia, 2015–2017: Ongoing increase in incidence despite nearelimination of the human-only plasmodium species. Clin. Infect. Dis. 70, 361–367 (2020).Article 

    Google Scholar 
    5.William, T. et al. Increasing incidence of Plasmodium knowlesi malaria following control of P. falciparum and P. vivax malaria in Sabah, Malaysia. PLoS Negl. Trop. Dis. 7, e2026 (2013).Article 

    Google Scholar 
    6.Fornace, K. M. et al. Association between landscape factors and spatial patterns of Plasmodium knowlesi infections in Sabah, Malaysia. Emerg. Infect. Dis. 22, 201–208 (2016).CAS 
    Article 

    Google Scholar 
    7.Gunggut, H., Saufi, D. S. N. S. A. M., Zaaba, Z. & Liu, M.S.-M. Where have all the forests gone? Deforestation in land below the wind. Procedia Soc. Behav. Sci. 153, 363–369 (2014).Article 

    Google Scholar 
    8.Brock, P. M. et al. Predictive analysis across spatial scales links zoonotic malaria to deforestation. Proc. R. Soc. B Biol. Sci. 286, 20182913 (2019).Article 

    Google Scholar 
    9.World Health Organization. WHO|Larval Source Management: A Supplementary Measure for Malaria Vector Control (WHO, 2013).
    Google Scholar 
    10.Wong, M. L. et al. Incrimination of Anopheles balabacensis as the vector for simian malaria in Kudat Division, Sabah, Malaysia. J. Microbiol. Immunol. Infect. 48, S47–S48 (2015).Article 

    Google Scholar 
    11.Vythilingam, I. & Hii, J. Simian malaria parasites: Special emphasis on Plasmodium knowlesi and their anopheles vectors in Southeast Asia. in Anopheles mosquitoes: New insights into malaria vectors (InTech, 2013). https://doi.org/10.5772/54491.Article 

    Google Scholar 
    12.Loh, E., Murray, K., Nava, K., Aguirre, A. & Daszak, A. Evaluating the links between biodiversity, land-use change, and infectious disease emergence. in Tropical Conservation (eds. Aguirre, A. & Sukumar, R.) 79–88. (Oxford, 2016).
    Google Scholar 
    13.Brant, H. L. et al. Vertical stratification of adult mosquitoes (Diptera: Culicidae) within a tropical rainforest in Sabah, Malaysia. Malar. J. 15, 1–10 (2016).Article 

    Google Scholar 
    14.Chua, T. H., Manin, B. O., Vythilingam, I., Fornace, K. & Drakeley, C. J. Effect of different habitat types on abundance and biting times of Anopheles balabacensis Baisas (Diptera: Culicidae) in Kudat district of Sabah, Malaysia. Parasit. Vectors 12, 364 (2019).Article 

    Google Scholar 
    15.Wong, M. L. et al. Seasonal and spatial dynamics of the primary vector of Plasmodium knowlesi within a major transmission focus in Sabah, Malaysia. PLoS Negl. Trop. Dis. 9, e0004153 (2015).Article 

    Google Scholar 
    16.Brown, R. et al. Human exposure to zoonotic malaria vectors in village, farm and forest habitats in Sabah, Malaysian Borneo. PLoS Negl. Trop. Dis. 14, 1–18 (2020).Article 

    Google Scholar 
    17.Yasuoka, J. & Levins, R. Impact of deforestation and agricultural development on anopheline ecology and malaria epidemiology. Am. J. Trop. Med. Hyg. 76, 450–460 (2007).Article 

    Google Scholar 
    18.Manin, B. O. et al. Investigating the contribution of peri-domestic transmission to risk of zoonotic malaria infection in humans. PLoS Negl. Trop. Dis. 10, e0000506 (2016).Article 

    Google Scholar 
    19.Rohani, A. et al. Characterization of the larval breeding sites of Anopheles balabacensis (Baisas), in Kudat, Sabah Malaysia. Southeast Asian. J. Trop. Med. Public Health 49, 566–579 (2018).
    Google Scholar 
    20.Ageep, T. B. et al. Spatial and temporal distribution of the malaria mosquito Anopheles arabiensis in northern Sudan: Influence of environmental factors and implications for vector control. Malar. J. 8, 123 (2009).Article 

    Google Scholar 
    21.Roleček, J., Chytrý, M., Hájek, M., Lvončík, S. & Tichý, L. Sampling design in large-scale vegetation studies: Do not sacrifice ecological thinking to statistical purism!. Folia Geobot. 42, 199–208 (2007).Article 

    Google Scholar 
    22.Bellier, E., Monestiez, P., Durbec, J.-P. & Candau, J.-N. Identifying spatial relationships at multiple scales: Principal coordinates of neighbour matrices (PCNM) and geostatistical approaches. Ecography 30, 385–399 (2007).Article 

    Google Scholar 
    23.Brock, P. M. et al. Plasmodium knowlesi transmission: Integrating quantitative approaches from epidemiology and ecology to understand malaria as a zoonosis. Parasitology 143, 389–400 (2016).CAS 
    Article 

    Google Scholar 
    24.Fornace, K. M., Drakeley, C. J., William, T., Espino, F. & Cox, J. Mapping infectious disease landscapes: Unmanned aerial vehicles and epidemiology. Trends Parasitol. 30, 514–519 (2014).Article 

    Google Scholar 
    25.GES DISC. Tropical Rainfall Measurement Mission (TRMM). TRMM (TMPA) Rainfall Estimate L3 3 hour 0.25 degree x 0.25 degree V7, Greenbelt. https://doi.org/10.5067/TRMM/TMPA/3H/7 (2011).Article 

    Google Scholar 
    26.Didan, K. MOD13Q1 MODIS/Terra Vegetation Indices 16-Day L3 Global 250m SIN Grid V006 NASA EOSDIS Land Processes DAAC. USGS 5, 2002–2015 (2015).
    Google Scholar 
    27.Didan, K. MOD13Q1 MODIS/Terra Vegetation Indices 16-Day L3 Global 250m SIN Grid V006. NASA EOSDIS Land Processes DAAC. NASA EOSDIS Land Processes DAAC. 5, 2002–2015. https://doi.org/10.5067/MODIS/MOD13Q1.006 (2015).Article 

    Google Scholar 
    28.NASA/METI/AIST/Japan Spacesystems, and U. S. /Japa. A. S. T. ASTER Global Digital Elevation Model V003. NASA EOSDIS Land Processes DAAC. https://lpdaac.usgs.gov/products/astgtmv003 (2019).29.Fornace, K. M. et al. Environmental risk factors and exposure to the zoonotic malaria parasite Plasmodium knowlesi across northern Sabah, Malaysia: A population-based cross-sectional survey. Lancet Planet. Heal. 3, e179–e186 (2019).Article 

    Google Scholar 
    30.Stark, D. J. et al. Long-tailed macaque response to deforestation in a plasmodium knowlesi-endemic area. EcoHealth 16, 638–646 (2019).Article 

    Google Scholar 
    31.Davidson, G., Chua, T. H., Cook, A., Speldewinde, P. & Weinstein, P. Defining the ecological and evolutionary drivers of Plasmodium knowlesi transmission within a multi-scale framework. Malar. J. 18, 1–13 (2019).Article 

    Google Scholar 
    32.Diuk-Wasser, M. A. et al. Effect of rice cultivation patterns on malaria vector abundance in rice-growing villages in Mali. Am. J. Trop. Med. Hyg. 76, 869–874 (2007).Article 

    Google Scholar 
    33.Stefani, A., Roux, E., Fotsing, J. M. & Carme, B. Studying relationships between environment and malaria incidence in Camopi (French Guiana) through the objective selection of buffer-based landscape characterisations. Int. J. Health Geogr. 10, 65 (2011).Article 

    Google Scholar 
    34.Wang, X., Blanchet, F. G. & Koper, N. Measuring habitat fragmentation: An evaluation of landscape pattern metrics. Methods Ecol. Evol. https://doi.org/10.1111/2041-210X.12198 (2014).Article 

    Google Scholar 
    35.McGarigal, K., Cushman, S. & Ene, E. FRAGSTATS v4: Spatial Pattern Analysis Program for Categorical and Continuous Maps. Computer software program produced by the authors at the University of Massachusetts, Amherst. http://www.umass.edu/landeco/research/fragstats/fragstats.html. https://doi.org/10.1049/oap-cired.2017.1227 (2012).Book 

    Google Scholar 
    36.TuckerLima, J. M., Vittor, A., Rifai, S. & Valle, D. Does deforestation promote or inhibit malaria transmission in the Amazon? A systematic literature review and critical appraisal of current evidence. Philos. Trans. R. Soc. B. 372, 20160125 (2017).Article 

    Google Scholar 
    37.Sallum, M. A. M., Peyton, E. L. & Wilkerson, R. C. Six new species of the Anopheles leucosphyrus group, reinterpretation of An. elegans and vector implications. Med. Vet. Entomol. 19, 158–199 (2005).CAS 
    Article 

    Google Scholar 
    38.Stoops, C. A. et al. Remotely-sensed land use patterns and the presence of Anopheles larvae (Diptera: Culicidae) in Sukabumi, West Java, Indonesia. J. Vector Ecol. 33, 30–39 (2008).Article 

    Google Scholar 
    39.Singh, J. & Tham, A. S. Case history on malaria vector control through the application of environmental management in Malaysia. World Health Org. 88, 1–70 (1988).
    Google Scholar 
    40.Tangena, J. A. A., Thammavong, P., Wilson, A. L., Brey, P. T. & Lindsay, S. W. Risk and control of mosquito-borne diseases in southeast asian rubber plantations. Trends Parasitol. 32, 402–415 (2016).Article 

    Google Scholar 
    41.Kaewwaen, W. & Bhumiratana, A. Landscape ecology and epidemiology of malaria associated with rubber plantations in Thailand: Integrated approaches to malaria ecotoping. Interdiscipl. Perspect. Infect. Dis. 2015, 1–15 (2015).Article 

    Google Scholar 
    42.Foley, D. H., Torres, E. P. & Mueller, I. Stream-bank shade and larval distribution of the Philippine malaria vector Anopheles flavirostris. Med. Vet. Entomol. 16, 347–355 (2002).CAS 
    Article 

    Google Scholar 
    43.Service, M. W. & Service, M. W. Sampling the Larval Population. in Mosquito Ecology 75–209 (Springer, 1993). https://doi.org/10.1007/978-94-015-8113-4_2.Article 
    MATH 

    Google Scholar 
    44.Sallum, M. A. M., Peyton, E. L., Harrison, B. A. & Wilkerson, R. C. Revision of the Leucosphyrus group of Anopheles (Cellia) (Diptera, Culicidae). Rev. Bras. Entomol. 49, 1–152 (2005).Article 

    Google Scholar 
    45.Rattanarithikul, R., Harrison, B. A., Harbach, R. E., Panthusiri, P. & Coleman, R. E. Illustrated keys to the mosquitoes of Thailand IV. Anopheles. J. Trop. Med. Public Health 37, 1–26 (2006).
    Google Scholar 
    46.R Core Team. R: The R Project for Statistical Computing. https://www.r-project.org/ (2020).47.Borremans, B., Faust, C., Manlove, K. R., Sokolow, S. H. & Lloyd-Smith, J. O. Cross-species pathogen spillover across ecosystem boundaries: Mechanisms and theory. Philos. Trans. R. Soc. B https://doi.org/10.1098/rstb.2018.0344 (2019).Article 

    Google Scholar  More

  • in

    Integrating multiple chemical tracers to elucidate the diet and habitat of Cookiecutter Sharks

    1.Norse, E. A. et al. Sustainability of deep-sea fisheries. Mar. Policy 36, 307–320 (2012).Article 

    Google Scholar 
    2.Simpfendorfer, C. A. & Kyne, P. M. Limited potential to recover from overfishing raises concerns for deep-sea sharks, rays and chimaeras. Environ. Conserv. 36, 97–103 (2009).Article 

    Google Scholar 
    3.Kyne, P. & Simpfendorfer, C. In Sharks and Their Relatives II: Biodiversity, Physiology, and Conservation (eds Carrier, J. C. et al.) 37–113 (CRC Press, 2010).
    Google Scholar 
    4.Dunn, M. R., Szabo, A., McVeagh, M. S. & Smith, P. J. The diet of deepwater sharks and the benefits of using DNA identification of prey. Deep Sea Res. Part I 57, 923–930 (2010).CAS 
    Article 

    Google Scholar 
    5.Mauchline, J. & Gordon, J. Diets of the sharks and chimaeroids of the Rockall Trough, northeastern Atlantic Ocean. Mar. Biol. 75, 269–278 (1983).Article 

    Google Scholar 
    6.Cortes, E. Standardized diet compositions and trophic levels in sharks. ICES J. Mar. Sci. 56, 707–717 (1999).Article 

    Google Scholar 
    7.Peterson, B. J. & Fry, B. Stable isotopes in ecosystem studies. Annu. Rev. Ecol. Syst. 18, 293–320 (1987).Article 

    Google Scholar 
    8.Post, D. M. Using stable isotopes to estimate trophic position: Models, methods, and assumptions. Ecology 83, 703–718 (2002).Article 

    Google Scholar 
    9.Estrada, J. A., Rice, A. N., Lutcavage, M. E. & Skomall, G. B. Predicting trophic position in sharks of the north-west Atlantic Ocean using stable isotope analysis. J. Mar. Biol. Assoc. UK 83, 1347–1350 (2003).CAS 
    Article 

    Google Scholar 
    10.Hussey, N. E. et al. Stable isotopes and elasmobranchs: Tissue types, methods, applications and assumptions. J. Fish. Biol. 20, 1449–1484 (2012).Article 
    CAS 

    Google Scholar 
    11.Meyer, L., Pethybridge, H., Nichols, P. D., Beckmann, C. & Huveneers, C. Abiotic and biotic drivers of fatty acid tracers in ecology: A global analysis of chondrichthyan profiles. Funct. Ecol. 20, 20 (2019).
    Google Scholar 
    12.Munroe, S., Meyer, L. & Heithaus, M. Dietary biomarkers in shark foraging and movement ecology. Shark Res. Emerg. Technol. Appl. Field Lab. 20, 20 (2018).

    Google Scholar 
    13.Hobson, K. A., Barnett-Johnson, R. & Cerling, T. E. In Isoscapes: Understanding Movement, Pattern, and Process on Earth Through Isotope Mapping (eds West, J. B. et al.) 273–298 (Springer, 2010).
    Google Scholar 
    14.Michener, R. H. & Kaufman, L. In Stable Isotopes in Ecology and Environmental Science (eds Michener, R. & Lajtha, K.) 238–282 (Blackwell, 2007).
    Google Scholar 
    15.West, J. B., Bowen, G. J., Cerling, T. E. & Ehleringer, J. R. Stable isotopes as one of nature’s ecological recorders. Trends Ecol. Evol. 21, 408–414 (2006).PubMed 
    Article 

    Google Scholar 
    16.DeNiro, M. J. Influence of diet on the distribution of nitrogen isotopes in animals. Geochim. Cosmochim. Acta 45, 341–345 (1981).ADS 
    CAS 
    Article 

    Google Scholar 
    17.DeNiro, M. J. & Epstein, S. Influence of diet on the distribution of carbon isotopes in animals. Geochim. Cosmochim. Acta 42, 495–506 (1978).ADS 
    CAS 
    Article 

    Google Scholar 
    18.Tieszen, L. L., Boutton, T. W., Tesdahl, K. G. & Slade, N. A. Fractionation and turnover of stable carbon isotopes in animal tissues: Implications for δ13C analysis of diet. Oecologia 57, 32–37 (1983).ADS 
    CAS 
    PubMed 
    Article 

    Google Scholar 
    19.MacNeil, M. A., Skomal, G. B. & Fisk, A. T. Stable isotopes from multiple tissues reveal diet switching in sharks. Mar. Ecol. Prog. Ser. 302, 199–206 (2005).ADS 
    Article 

    Google Scholar 
    20.Kim, S. L., Martinez del Rio, C., Casper, D. & Koch, P. L. Isotopic incorporation rates for shark tissues from a long-term captive feeding study. J Exp Biol 215, 2495–2500 (2012).21.Madigan, D. J. et al. Tissue turnover rates and isotopic trophic discrimination factors in the endothermic teleost, Pacific bluefin tuna (Thunnus orientalis). PLoS One 7, e49220 (2012).ADS 
    CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    22.Carlisle, A. B. et al. Using stable isotope analysis to understand the migration and trophic ecology of northeastern Pacific white sharks (Carcharodon carcharias). PLoS One 7, 30492 (2012).ADS 
    Article 
    CAS 

    Google Scholar 
    23.Madigan, D. J. et al. Reconstructing transoceanic migration patterns of Pacific bluefin tuna using a chemical tracer toolbox. Ecology 95, 1674–1683 (2014).PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    24.Ackman, R.G. & Macpherson, E.J. Coincidence of cis-and trans-monoethylenic fatty acids simplifies the open-tubular gas-liquid chromatography of butyl esters of butter fatty acids. Food chem. 50(1), 45–52 (1994).25.Sargent, J., Bell, G., McEvoy, L., Tocher, D. & Estevez, A. Recent developments in the essential fatty acid nutrition of fish. Aquaculture., 177(1–4), 191–199 (1999).26.Tocher, D.R. Metabolism and functions of lipids and fatty acids in teleost fish. Rev. Fish. Sci. 11(2), 107–184 (2003).27.McMeans, B. C. et al. The role of Greenland sharks (Somniosus microcephalus) in an Arctic ecosystem: Assessed via stable isotopes and fatty acids. Mar. Biol. 160, 1223–1238. https://doi.org/10.1007/s00227-013-2174-z (2013).Article 

    Google Scholar 
    28.Pethybridge, H. R., Nichols, P. D., Virtue, P. & Jackson, G. D. The foraging ecology of an oceanic squid, Todarodes filippovae: The use of signature lipid profiling to monitor ecosystem change. Deep Sea Res. Part II 95, 119–128 (2013).CAS 
    Article 

    Google Scholar 
    29.Pethybridge, H. et al. Lipid and mercury profiles of 61 mid-trophic species collected off south-eastern Australia. Mar. Freshw. Res. 61, 1092–1108 (2010).CAS 
    Article 

    Google Scholar 
    30.Beckmann, C. L., Mitchell, J. G., Stone, D. A. & Huveneers, C. A controlled feeding experiment investigating the effects of a dietary switch on muscle and liver fatty acid profiles in Port Jackson sharks Heterodontus portusjacksoni. J. Exp. Mar. Biol. Ecol. 448, 10–18 (2013).CAS 
    Article 

    Google Scholar 
    31.Pethybridge, H. R., Choy, C. A., Polovina, J. J. & Fulton, E. A. Improving marine ecosystem models with biochemical tracers. Ann. Rev. Mar. Sci. 10, 199–228 (2018).PubMed 
    Article 

    Google Scholar 
    32.Belicka, L. L., Matich, P., Jaffé, R. & Heithaus, M. R. Fatty acids and stable isotopes as indicators of early-life feeding and potential maternal resource dependency in the bull shark Carcharhinus leucas. Mar. Ecol. Prog. Ser. 455, 245–256 (2012).ADS 
    CAS 
    Article 

    Google Scholar 
    33.Every, S. L., Fulton, C. J., Pethybridge, H. R., Kyne, P. M. & Crook, D. A. A seasonally dynamic estuarine ecosystem provides a diverse prey base for Elasmobranchs. Estuar. Coasts 42, 580–595 (2019).CAS 
    Article 

    Google Scholar 
    34.Ruppert, K. M., Kline, R. J. & Rahman, M. S. Past, present, and future perspectives of environmental DNA (eDNA) metabarcoding: A systematic review in methods, monitoring, and applications of global eDNA. Glob. Ecol. Conserv. 20, e00547 (2019).Article 

    Google Scholar 
    35.Soininen, E. M. et al. Shedding new light on the diet of Norwegian lemmings: DNA metabarcoding of stomach content. Polar Biol 36, 1069–1076 (2013).Article 

    Google Scholar 
    36.De Barba, M. et al. DNA metabarcoding multiplexing and validation of data accuracy for diet assessment: Application to omnivorous diet. Mol. Ecol. Resour. 14, 306–323 (2014).Article 
    CAS 

    Google Scholar 
    37.Deagle, B. E., Kirkwood, R. & Jarman, S. N. Analysis of Australian fur seal diet by pyrosequencing prey DNA in faeces. Mol. Ecol. 18, 2022–2038 (2009).CAS 
    PubMed 
    Article 

    Google Scholar 
    38.Bade, L. M., Balakrishnan, C. N., Pilgrim, E. M., McRae, S. B. & Luczkovich, J. J. A genetic technique to identify the diet of cownose rays, Rhinoptera bonasus: Analysis of shellfish prey items from North Carolina and Virginia. Environ. Biol. Fishes 97, 999–1012 (2014).Article 

    Google Scholar 
    39.Jensen, M. R., Knudsen, S. W., Munk, P., Thomsen, P. F. & Møller, P. R. Tracing European eel in the diet of mesopelagic fishes from the Sargasso Sea using DNA from fish stomachs. Mar. Biol. 165, 130 (2018).Article 
    CAS 

    Google Scholar 
    40.Compagno, L. FAO species catalogue. Vol. 4. Sharks of the world. An annotated and illustrated catalogue of sharks species known to date. Part 1. Hexanchiformes to Lammiformes. FAO Fish. Synop. 20, 1–249 (1984).
    Google Scholar 
    41.Jahn, A. & Haedrich, R. Notes on the pelagic squaloid shark Isistius brasiliensis. Biol. Oceanogr. 5, 297–309 (1988).
    Google Scholar 
    42.Nakano, H. & Tabuchi, M. Occurrence of the cookiecutter shark Isistius brasiliensis in surface waters of the North Pacific Ocean. Jpn. J. Ichthyol. 37, 60–63 (1990).
    Google Scholar 
    43.Hubbs, C. L., Iwai, T. & Matsubara, K. External and internal characters, horizontal and vertical distributions, luminescence, and food of the dwarf pelagic shark, Euprotomicrus bispinatus. (1967).44.Papastamatiou, Y. P., Wetherbee, B. M., O’Sullivan, J., Goodmanlowe, G. D. & Lowe, C. G. Foraging ecology of cookiecutter sharks (Isistius brasiliensis) on pelagic fishes in Hawaii, inferred from prey bite wounds. Environ. Biol. Fishes 88, 361–368 (2010).Article 

    Google Scholar 
    45.Feunteun, A. et al. First evaluation of the cookie-cutter sharks (Isistius sp.) predation pattern on different cetacean species in Martinique. Environ. Biol. Fishes 20, 1–11 (2018).
    Google Scholar 
    46.Jones, E. Isistius brasiliensis, a squaloid shark, probable cause of crater wounds on fishes and cetaceans. Fish Bull. 69, 791–798 (1971).
    Google Scholar 
    47.Strasburg, D. W. The diet and dentition of Isistius brasiliensis, with remarks on tooth replacement in other sharks. Copeia 20, 33–40 (1963).Article 

    Google Scholar 
    48.Widder, E. A. A predatory use of counter illumination by the squaloid shark, Isistius brasiliensis. Environ. Biol. Fishes 53, 267–273 (1998).Article 

    Google Scholar 
    49.Moore, M., Steiner, L. & Jann, B. Cetacean surveys in the Cape Verde Islands and the use of cookiecutter shark bite lesions as a population marker for fin whales. Aquat. Mamm. 29, 383–389 (2003).Article 

    Google Scholar 
    50.Muñoz-Chápuli, R., Salgado, J. R. & de La Serna, J. Biogeography of Isistius brasiliensis in the north-eastern Atlantic, inferred from crater wounds on swordfish (Xiphias gladius). J. Mar. Biol. Assoc. U K 68, 315–321 (1988).Article 

    Google Scholar 
    51.Murakami, C., Yoshida, H. & Yonezaki, S. Cookie-cutter shark Isistius brasiliensis eats Bryde’s whale Balaenoptera brydei. Ichthyol. Res. 65, 398–404 (2018).Article 

    Google Scholar 
    52.Castro, J., Anllo, T., Mejuto, J. & García, B. Ichnology applied to the study of Cookiecutter shark (Isistius brasiliensis) biogeography in the Gulf of Guinea. Environ. Biol. Fishes 101, 579–588 (2018).Article 

    Google Scholar 
    53.Kim, S. L. et al. Carbon and nitrogen discrimination factors for elasmobranch soft tissues based on a long-term controlled feeding study. Environ. Biol. Fishes 95, 37–52 (2012).Article 

    Google Scholar 
    54.Le Boeuf, B., McCosker, J. & Hewitt, J. Crater wounds on northern elephant seals: The Cookiecutter Shark strikes again. Fish Bull. 85, 20 (1987).
    Google Scholar 
    55.Niella, Y. et al. Cookie-cutter shark Isistius spp. predation upon different tuna species from the south-western Atlantic Ocean. J. Fish. Biol. 92, 1082–1089 (2018).CAS 
    PubMed 
    Article 

    Google Scholar 
    56.Manlick, P. J., Petersen, S. M., Moriarty, K. M. & Pauli, J. N. Stable isotopes reveal limited Eltonian niche conservatism across carnivore populations. Funct. Ecol. 33, 335–345 (2019).Article 

    Google Scholar 
    57.McMeans, B.C., Arts, M.T. & Fisk, A.T. Similarity between predator and prey fatty acid profiles is tissue dependent in Greenland sharks (Somniosus microcephalus): Implications for diet reconstruction. J. Exp. Mar. Biol. Ecol. 429, 55–63 (2012).58.Waugh, C.A., Nichols, P.D., Schlabach, M., Noad, M. & Nash, S.B. Vertical distribution of lipids, fatty acids and organochlorine contaminants in the blubber of southern hemisphere humpback whales (Megaptera novaeangliae). Mar. Environ. Res. 94, 24–31 (2014).59.Sigler, M. F. et al. Diet of Pacific sleeper shark, a potential Steller sea lion predator, in the north-east Pacific Ocean. J. Fish. Biol. 69, 392–405 (2006).Article 

    Google Scholar 
    60.Leclerc, L.-M. et al. Greenland sharks (Somniosus microcephalus) scavenge offal from minke (Balaenoptera acutorostrata) whaling operations in Svalbard (Norway). Polar. Res. 30, 7342 (2011).Article 

    Google Scholar 
    61.Yano, K., Stevens, J. & Compagno, L. Distribution, reproduction and feeding of the Greenland shark Somniosus (Somniosus) microcephalus, with notes on two other sleeper sharks, Somniosus (Somniosus) pacificus and Somniosus (Somniosus) antarcticus. J. Fish. Biol. 70, 374–390 (2007).Article 

    Google Scholar 
    62.Preti, A. et al. Comparative feeding ecology of shortfin mako, blue and thresher sharks in the California current. Environ. Biol. Fishes https://doi.org/10.1007/s10641-10012-19980-x (2012).Article 

    Google Scholar 
    63.Phillips, D. L. et al. Best practices for use of stable isotope mixing models in food-web studies. Can. J. Zool. 92, 823–835 (2014).Article 

    Google Scholar 
    64.Smith, J. A., Mazumder, D., Suthers, I. M. & Taylor, M. D. To fit or not to fit: Evaluating stable isotope mixing models using simulated mixing polygons. Methods Ecol. Evol. 4, 612–618 (2013).Article 

    Google Scholar 
    65.Hussey, N. E. et al. Rescaling the trophic structure of marine food webs. Ecol. Lett. https://doi.org/10.1111/ele.12226 (2013).Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    66.Childress, J. J. & Nygaard, M. H. Deep Sea Research and Oceanographic Abstracts 1093–1109 (Elsevier, 1973).
    Google Scholar 
    67.Childress, J., Price, M., Favuzzi, J. & Cowles, D. Chemical composition of midwater fishes as a function of depth of occurrence off the Hawaiian Islands: Food availability as a selective factor?. Mar. Biol. 105, 235–246 (1990).Article 

    Google Scholar 
    68.Choy, C. A., Popp, B. N., Hannides, C. C. & Drazen, J. C. Trophic structure and food resources of epipelagic and mesopelagic fishes in the North Pacific Subtropical Gyre ecosystem inferred from nitrogen isotopic compositions. Limnol. Oceanogr. 60, 1156–1171 (2015).ADS 
    Article 

    Google Scholar 
    69.Gloeckler, K. et al. Stable isotope analysis of micronekton around Hawaii reveals suspended particles are an important nutritional source in the lower mesopelagic and upper bathypelagic zones. Limnol. Oceanogr. 63, 1168–1180 (2018).ADS 
    CAS 
    Article 

    Google Scholar 
    70.Hannides, C. C., Popp, B. N., Choy, C. A. & Drazen, J. C. Midwater zooplankton and suspended particle dynamics in the North Pacific Subtropical Gyre: A stable isotope perspective. Limnol. Oceanogr. 58, 1931–1946 (2013).ADS 
    CAS 
    Article 

    Google Scholar 
    71.Dunstan, G.A., Sinclair, A.J., O’Dea, K. & Naughton, J.M. The lipid content and fatty acid composition of various marine species from southern Australian coastal waters. Comp. Biochem. Physiol. B: Comp. Biochem. 91(1), 165–169 (1988).72.Semeniuk, C.A., Speers-Roesch, B. & Rothley, K.D. Using fatty-acid profile analysis as an ecologic indicator in the management of tourist impacts on marine wildlife: a case of stingray-feeding in the Caribbean. Environ. Manag. 40(4), 665–677 (2007).73.Wai, T.C., Leung, K.M., Sin, S.Y., Cornish, A., Dudgeon, D. & Williams, G.A. Spatial, seasonal, and ontogenetic variations in the significance of detrital pathways and terrestrial carbon for a benthic shark, Chiloscyllium plagiosum (Hemiscylliidae), in a tropical estuary. Limnol. Oceanogr. 56(3), 1035–1053 (2011).74.Ebert, D. A., Fowler, S. L., Compagno, L. J. & Dando, M. Sharks of the World: A Fully Illustrated Guide (Wild Nature Press, 2013).
    Google Scholar 
    75.Vaudo, J. J., Matich, P. & Heithaus, M. R. Mother-offspring isotope fractionation in two species of placentatrophic sharks. J. Fish. Biol. 77, 1724–1727 (2010).CAS 
    PubMed 
    Article 

    Google Scholar 
    76.Olin, J. A. et al. Maternal meddling in neonatal sharks: Implications for interpreting stable isotopes in young animals. Rapid Commun. Mass Spectrom. 25, 1008–1016 (2011).ADS 
    CAS 
    PubMed 
    Article 

    Google Scholar 
    77.Grubbs, R. D. In Sharks and Their Relatives II: Biodiversity, Physiology, and Conservation (eds Carrier, J. C. et al.) 319–350 (CRC Press, 2010).
    Google Scholar 
    78.Yano, K. & Tanaka, S. Size at maturity, reproductive cycle, fecundity, and depth segregation of the deep sea squaloid sharks Centroscymnus owstoni and C. coelolepis in Suruga Bay Japan. Nippon Suisan Gakkaishi 54, 20 (1988).
    Google Scholar 
    79.Yano, K. & Tanaka, S. Review of the deep sea squaloid shark genus Scymnodon of Japan, with a description of a new species. Jpn. J. Ichthyol. 30, 341–360 (1984).
    Google Scholar 
    80.Munoz-Chapuli, R. Ethologie de la reproduction chez quelques requins de l’Atlantique Nord-Est. Cybium 8, 1–14 (1984).
    Google Scholar 
    81.Jakobsdóttir, K. B. Biological aspects of two deep-water squalid sharks: Centroscyllium fabricii (Reinhardt, 1825) and Etmopterus princeps (Collett, 1904) in Icelandic waters. Fish Res. 51, 247–265 (2001).Article 

    Google Scholar 
    82.Wetherbee, B. M. Distribution and reproduction of the southern lantern shark from New Zealand. J. Fish. Biol. 49, 1186–1196. https://doi.org/10.1111/j.1095-8649.1996.tb01788.x (1996).Article 

    Google Scholar 
    83.MacNeil, M. A., Drouillard, K. G. & Fisk, A. T. Variable uptake and elimination of stable nitrogen isotopes between tissues in fish. Can. J. Fish. Aquat. Sci. 63, 345–353 (2006).CAS 
    Article 

    Google Scholar 
    84.Logan, J. M. & Lutcavage, M. Stable isotope dynamics in elasmobranch fishes. Hydrobiologia 644, 231–244 (2010).CAS 
    Article 

    Google Scholar 
    85.Weidel, B. C., Carpenter, S. R., Kitchell, J. F. & Vander Zanden, M. J. Rates and components of carbon turnover in fish muscle: Insights from bioenergetics models and a whole-lake 13C addition. Can. J. Fish. Aquat. Sci. 68, 387–399 (2011).CAS 
    Article 

    Google Scholar 
    86.Carlisle, A. B. et al. Interactive effects of urea and lipid content confound stable isotope analysis in elasmobranch fishes. Can. J. Fish. Aquat. Sci. 74, 419–428 (2016).Article 
    CAS 

    Google Scholar 
    87.Kim, S. L. & Koch, P. L. Methods to collect, preserve, and prepare elasmobranch tissues for stable isotope analysis. Environ. Biol. Fishes 95, 53–63 (2012).Article 

    Google Scholar 
    88.Witteveen, B. H., Worthy, G. A. J. & Roth, J. D. Tracing migratory movements of breeding North Pacific humpback whales using stable isotope analysis. Mar. Ecol. Prog. Ser. 393, 173–183. https://doi.org/10.3354/meps08231 (2009).ADS 
    Article 

    Google Scholar 
    89.Parry, M. P. The trophic ecology of two ommastrephid squid species, Ommastrephes bartamii and Sthenoteuthis oualaniensis, in the North Pacific sub-tropical gyre Ph.D. thesis, University of Hawaii, (2003).90.Parry, M. P. Trophic variation with length in two ommastrephid squids, Ommastrephes bartramiii and Sthenoteuthis oualaniensis. Mar. Biol. 153, 249–256 (2008).Article 

    Google Scholar 
    91.Graham, B. S. Trophic dynamics and movements of tuna in tropical Pacific Ocean inferred from stable isotope analyses Ph. D. thesis thesis, University of Hawaii, (2007).92.Graham, B. S., Grubbs, D., Holland, K. & Popp, B. N. A rapid ontogenetic shift in the diet of juvenile yellowfin tuna from Hawaii. Mar. Biol. 150, 647–658 (2007).Article 

    Google Scholar 
    93.Carlisle, A. B. et al. Stable isotope analysis of vertebrae reveals ontogenetic changes in habitat in an endothermic pelagic shark. Proc. R. Soc. B-Biol. Sci. 282, 20141446. https://doi.org/10.1098/rspb.2014.1446 (2015).CAS 
    Article 

    Google Scholar 
    94.Stock, B. C. & Semmens, B. X. MixSIAR GUI user manual, version 1.0. http://conserver.iugo-cafe.org/user/brice.semmens/MixSIAR (2013).95.Folch, J., Lees, M. & Stanley, G.S. A simple method for the isolation and purification of total lipides from animal tissues. J. Biol. Chem. 226(1), 497–509 (1957).96.Kartikasari, L.R., Hughes, R.J., Geier, M.S., Makrides, M. & Gibson, R.A. Dietary alpha-linolenic acid enhances omega-3 long chain polyunsaturated fatty acid levels in
    chicken tissues. Prostaglandins Leukot. Essent. Fatty Acids. 87(4–5), 103–109 (2012).97.Froese, R. & D. Pauly. Editors. 2021. FishBase. World Wide Web electronic publication. https://www.fishbase.org, version (02/2021).98.Clarke, K. & Gorley, R. (PRIMER-E: Plymouth, 2006).
    99.Riaz, T. et al. ecoPrimers: Inference of new DNA barcode markers from whole genome sequence analysis. Nucleic Acids Res. 39, e145–e145 (2011).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    100.Kelly, R. P., Port, J. A., Yamahara, K. M. & Crowder, L. B. Using environmental DNA to census marine fishes in a large mesocosm. PLoS One 9, e86175 (2014).ADS 
    PubMed 
    PubMed Central 
    Article 
    CAS 

    Google Scholar 
    101.Stoeckle, M. Y., Soboleva, L. & Charlop-Powers, Z. Aquatic environmental DNA detects seasonal fish abundance and habitat preference in an urban estuary. PLoS ONE 12, e0175186 (2017).PubMed 
    PubMed Central 
    Article 
    CAS 

    Google Scholar 
    102.Port, J. A. et al. Assessing vertebrate biodiversity in a kelp forest ecosystem using environmental DNA. Mol. Ecol. 25, 527–541 (2016).CAS 
    PubMed 
    Article 

    Google Scholar 
    103.Shehzad, W. et al. Carnivore diet analysis based on next-generation sequencing: Application to the leopard cat (Prionailurus bengalensis) in Pakistan. Mol. Ecol. 21, 1951–1965 (2012).CAS 
    PubMed 
    Article 

    Google Scholar 
    104.Edgar, R. C. Search and clustering orders of magnitude faster than BLAST. Bioinformatics 26, 2460–2461 (2010).CAS 
    Article 

    Google Scholar 
    105.Martin, M. Cutadapt removes adapter sequences from high-throughput sequencing reads. EMBnet. J. 17, 10–12 (2011).Article 

    Google Scholar 
    106.Zhang, J., Kobert, K., Flouri, T. & Stamatakis, A. PEAR: A fast and accurate illumina paired-end reAd mergeR. Bioinformatics 30, 614–620 (2013).PubMed 
    PubMed Central 
    Article 
    CAS 

    Google Scholar 
    107.Schnell, I. B., Bohmann, K. & Gilbert, M. T. P. Tag jumps illuminated-reducing sequence-to-sample misidentifications in metabarcoding studies. Mol. Ecol. Resour. 15, 1289–1303 (2015).CAS 
    PubMed 
    Article 

    Google Scholar 
    108.Camacho, C. et al. BLAST+: Architecture and applications. BMC Bioinform. 10, 421 (2009).Article 
    CAS 

    Google Scholar 
    109.Oksanen, J. et al. Vegan: Community ecology package. R package version 1.17–4. http://cran.r-project.org. Acesso em 23, 2010 (2010). More

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    Climate change drives widespread shifts in lake thermal habitat

    OverviewWe used long-term time series of lake temperature profiles to determine the magnitude of thermal habitat change in 139 widely distributed lakes. Time series were interpolated across depth and season to generate data with consistent resolutions across lakes. To assess temperature change, we used a metric, ‘thermal non-overlap’, based on the percentage of two kernel density estimations of lake temperature which are non-overlapping. We calculated the metric for a range of plausible seasonal and depth habitat restrictions for aquatic species in the face of climate change. We used BRT to explain variability across lakes in their thermal habitat non-overlap as a function of lake characteristics (mean depth and latitude), characteristics of the time series for each lake (starting day of the year, ending day of the year, starting year and ending year, average number of sampling dates per year, long-term trend in the number of sampling dates per year, long-term trend in the yearly seasonal range of sampling dates), the habitat restriction values (season and depth) and the location of the time series delineation for thermal non-overlap calculations (30th, 50th and/or 70th quantiles of the years included in each lake’s time series).Study sitesWe compiled long-term lake temperature data from 139 lakes across the globe. Temperature variations in many of these lakes have already been linked to climate change1,2,19,20,57,58, but temperature change in at least one lake may be partially due to background climate variation in addition to anthropogenic climate change (Atlantic Multidecadal Oscillation in Lake Annie)59. The lakes included in our analysis represent a wide range of surface area (0.02 to 68,800 km2), maximum depth (2.3 to 1,642 m), latitude (60 °S to 69 °N) and elevation (−212 to 1,987 m above sea level) (see Supplementary Table 1 for more information).Temperature dataIn total, we used more than 32 million lake temperature measurements for our analyses. The number of observations per lake ranged from 368 (Lake Stensjon) to 7,636,767 (Lake Superior) with approximately 232,000 observations per lake on average. Temperature data from each lake came from in situ temperature profiles60,61,62,63,64 for lakes smaller than 169 km2 and from a combination of in situ temperature profiles and remotely sensed surface water temperatures for 21 larger lakes. Remote sensing data were used in recognition that temperature and warming rates can vary substantially across latitude and longitude for large lakes19,20,21.The mean length of the temperature time series was 36 years with a range from 15 to 101 years. All lakes had temperature data which started in the year 2000 or earlier and ended in 2000 or later. Lakes had on average 29 temperature profiles per year (inner quartile range: 7–26). In situ temperature data were measured using a wide variety of temperature sensors. Data collection methods included regularly collected discrete temperature profiles, high-resolution thermistor chains and other commonly accepted tools for measuring aquatic temperature. The in situ data are publicly available through the environmental data initiative60.Remotely sensed lake surface temperatures were measured using the Advanced Very High-Resolution Radiometer (AVHRR) and processed by the Group for High Resolution Sea Surface Temperature (GHRSST) project65. AVHRR data have been validated against buoy data from the North American Great Lakes and found to have a root mean squared error of 0.55 °C compared with in situ measurements2. AVHRR temperature data were included to capture horizontal variability in temperature and warming in 21 of the 139 lakes that would not be captured by temperature profiles from a single central location19,20,21. AVHRR data were pooled with in situ data for temperature interpolation.Temperature interpolationTemperature data were spatially and temporally interpolated for each lake. All temperature profile data were first linearly interpolated across depth because temperature variability with depth is highly constrained by lake physics and typically allows for robust interpolations. The largest data gap over which depth interpolation occurred was 0.1 × mean depth of each lake. Following interpolation across depth, data were interpolated across time using standard spline interpolation models with a Kalman filter66. The model output was used to fill data gaps to produce a continuous, daily time series over the day of the year range for which temperature profiles had been regularly measured. Some times of the year were excluded from specific lakes because they lacked regular measurements throughout the length of the long-term time series. Thus, the same starting and ending day of the year was used for each lake throughout its time series, and was often shorter than the full annual cycle (Supplementary Table 1). The largest gap in time over which interpolation occurred was 30 days and this included extrapolations for lakes with missing data at the beginning or end of seasonal coverage in a specific year. Years with longer gaps were omitted from the analysis and the length of the seasonal coverage was optimized to minimize the number of years that needed to be removed. For large lakes with many sampling points (for example, Baikal, Superior, Victoria), temperature data were divided into 1,000 km2 latitude–longitude bins and interpolated across depth and across time separately for each bin. The mean seasonal coverage of the interpolated lake time series was 245 days per year with a minimum of 17 days per year and a maximum of 365 days per year.The interpolated temperature output had a daily temporal resolution and a depth resolution which varied continuously over depth. At the lake surface, we interpolated temperatures every 0.1 m (for example, 0 m, 0.1 m, 0.2 m), to every 1 m starting at a depth of 10 m (for example, 10 m, 11 m, 12 m) and every 100 m starting at a depth of 1,000 m (for example, 1,000 m, 1,100 m, 1,200 m). These depth increments were used because they consistently gave good coverage over all major lake strata, regardless of each lake’s morphometric characteristics, while minimizing computational intensity by eliminating redundancy within lake strata.Thermal habitat non-overlap calculationsAfter interpolating the temperature data across depth and season for each lake, we bisected it into an early part (part a) and a later part (part b). Parts a and b were iteratively delineated at three points positioned serially along the time series—at the 30th, 50th and 70th quantiles. We averaged the final non-overlap values across these three delineations for each lake so that the results depended less on the somewhat arbitrary decision of where to split the time series. For each delineation, we randomly sampled 10,000 temperature values from each of parts a and b. This was repeated ten times resulting in a total of 300,000 temperature values across all three time series delineations and all ten repetitions for each lake (10,000 × 3 × 10). The sampling probability for temperature values in each comparison was weighted by the volume increment associated with each temperature value (depth increment (Id) × cross-sectional area at each depth (Cd)). Id was calculated as the difference between the depth of the sampled temperature value and the next depth in the depth resolution of the interpolated temperatures. Cd at each depth for each lake was calculated using standard, three-parameter models for estimating lake cross-sectional area based on surface area, maximum depth and mean depth67. For large lakes with temperature data at multiple locations across latitude and longitude, Cd was divided by the number of latitude–longitude bins used for each lake. Temperature values from large lakes were sampled regardless of their associated latitude–longitude bins. As a result of the volume-weighting procedure, temperature measurements were sampled in proportion to the volume of water represented by each value, with temperatures representing larger volumes being sampled more often. As a consequence of this volume-weighting procedure, the resulting temperature distributions were robust to moderate changes in the depths used for the temperature interpolation (Supplementary Fig. 1).We defined thermal non-overlap (TNO) as the symmetric difference (Ө) between the kernel density estimations of temperature values from parts a and b of the time series as a proportion of the union (∪) of both kernel density estimations, following an established method42. Conversely, we defined the thermal habitat overlap (as opposed to non-overlap) as the intersection (∩) of the kernel density estimations as a proportion of the union (∪) of both distributions. All values were converted to percentages by multiplying by 100.$${mathrm{TNO}}left( % right) = 100 times frac{{{{T}}_{{mathrm{recent}}},ominus,{{T}}_{{mathrm{baseline}}}}}{{{{T}}_{{mathrm{recent}}} cup {{T}}_{{mathrm{baseline}}}}} = 100 times left( {1 – frac{{{{T}}_{{mathrm{recent}}} cap {{T}}_{{mathrm{baseline}}}}}{{{{T}}_{{mathrm{recent}}} cup {{T}}_{{mathrm{baseline}}}}}} right)$$
    (1)
    We used simulations to test the sensitivity of TNO to changes in mean and s.d. of temperature. We primed these simulations with three baseline temperature distributions all with a mean of 15 °C but with varying s.d. (4, 6, 8 °C). We simulated a range of additional temperature distributions by increasing and decreasing the mean and s.d. of the baseline temperature distributions and then calculated the corresponding values of TNO. The simulated change in both mean and s.d. varied from −3 to +3 °C. We found that TNO was sensitive to changes in mean and s.d. but was slightly more sensitive to reductions in s.d. compared with increases. TNO values also depended on the baseline s.d., such that lower starting s.d. elevates values of non-overlap given an equivalent change in temperature (Extended Data Fig. 1).We also quantified null values of thermal non-overlap (TNOo) by repeating the thermal non-overlap calculations but where parts a and b were defined by randomly dividing the individual years of data into two separate groups as opposed to sequentially dividing them along the time series.$${mathrm{TNO}}_{mathrm{o}}(% ) = 100 times frac{{{{T}}_{{mathrm{random}},{{a}}},ominus,{{T}}_{{mathrm{random}},{{b}}}}}{{{{T}}_{{mathrm{random}},{{a}}} cup {{T}}_{{mathrm{random}},{{b}}}}}$$
    (2)
    To calculate standardized thermal non-overlap (TNOs), we subtracted TNOo from TNO thereby setting the null expectation to zero.$${mathrm{TNO}}_{mathrm{s}}left( {mathrm{% }} right) = {mathrm{TNO}} – {mathrm{TNO}}_{mathrm{o}}$$
    (3)
    In this case, if the temperature distributions in the recent and baseline time periods were identical, the TNOs would equal approximately zero. Values different from zero reflect a combination of random noise and long-term temperature change. All non-overlap values described in the main text and shown in Figs. 2–6 reflect values of TNOs. A comparison between raw values of TNO and TNOo can be found in Extended Data Fig. 5. Thermal non-overlap values and the null values were calculated using the ‘overlap’ function from the ‘overlapping’ package42 in the R environment for statistical computing and visualization. In the function, we set the number of equally spaced points at which the overlapping kernel density estimation is evaluated to 100 for all comparisons because it minimized the values of TNOo (we considered a range of values from 5 to 10,000).To assess the effect of seasonal habitat restrictions (Slimit) and volumetric habitat restrictions (Vlimit), we modified equations (1)–(3) by comparing temperature values only from a specified range of depths and/or days of the year. We considered a range of habitat restrictions scaled from 0 to 0.95, where 0.95 is the most restrictive (temperature values were compared from within bins equivalent to 1/20th of the available seasonal and volumetric habitat) and 0 is the least restrictive (temperature values were compared regardless of season and depth). We focused our interpretations on the unitless habitat restrictions (scaled from 0 to 0.95) instead of in units of days or m3 so that habitat restrictions could be more readily compared across lakes. Comparing a Vlimit value of 0.8 across lakes of different sizes assumes that a habitat restriction of 2 m3 in a 10 m3 lake would be comparable to a 20 m3 habitat delineation in a 200 m3 lake. The actual size of the seasonal habitat restrictions for each lake in units of days were calculated using the value of Slimit as follows:$$S = left( {mathrm{doy}}_{mathrm{max}} – {mathrm{doy}}_{mathrm{min}}right)left( {1 – S_{mathrm{limit}}} right)$$where S is the seasonal habitat restriction in units of days, doymax is the maximum day of the year of the lakes’ seasonal coverage, doymin is the minimum day of the year of the lakes’ seasonal coverage and Slimit is the seasonal habitat restriction scaled from 0 to 0.95. For example, in a lake with a seasonal coverage from day of the year 1 to day of the year 365, with an Slimit value of 0.75, we compared randomly selected temperatures from time periods a and b separately for four seasonal bins (days of the year 1–91, 92–183, 184–273 and 274–365). Similarly, the actual size of the volumetric habitat restrictions (V) for each lake in units of m3 were calculated using the value of Vlimit as follows:$$V = left( {mathrm{volume}} right) times left( {1 – V_{mathrm{limit}}} right)$$where V is the volumetric habitat restriction in units of m3, volume is the lake’s total volume and Vlimit is the volumetric habitat restriction value scaled from 0 to 0.95. For example, if a lake with a volume of 100 m3 had a Vlimit value of 0.75, we randomly selected temperature values from time periods a and b which were within four 25 m3 (100 m3 × (1 − 0.8)) bins. Volume bins were subsequently translated into sequential depth bins for the purpose of temperature value selection, making them functionally depth limits, and they are presented as such in the main text.We factorially combined a discrete series of values for Slimit and Vlimit (0, 1/2, 2/3, 5/6, 8/9, 12/13 and 19/20) to test a range of combined seasonal and volumetric habitat restrictions that do not require the overlap or truncation of bins. For reference, habitat restrictions are presented visually for hypothetical ‘Species 1’ (Slimit = 0, Vlimit = 0.8), ‘Species 2’ (Slimit = 0.8, Vlimit = 0) and ‘Species 3’ (Slimit = 0.8, Vlimit = 0.8) examples (Fig. 1). These limits reflect hypothetical restrictions in a species’ habitat due to ecological factors and approximate the habitat available for a low-light specialist phytoplankton (species 1), a spring migratory fish (species 2) and a diapausing benthic invertebrate (species 3). In Fig. 6, the species habitat restriction values for P. rubescens were Slimit = 0.74, Vlimit = 0.89 (Fig. 6).Explaining variability in thermal habitat non-overlapWe used BRT to explain lake-to-lake variability in thermal habitat change (percentage of non-overlap) while accounting for differences in the temporal coverage of each lake’s time series. The predictor variables in the BRT were the starting year of the time series, ending year of the time series, starting day of the year of the seasonal coverage, ending day of the year of the seasonal coverage, average number of sampling dates per year, linear trend (Theil–Sen slope) in the average number of sampling dates per year, linear trend (Theil–Sen slope) in the yearly extent of the time series’ seasonal coverage, lake mean depth, absolute latitude (degrees from the Equator), seasonal habitat restriction, depth habitat restriction and time series delineation. Geospatial and morphometric data for each lake is available from the previously published HydroLAKES database41. Of the available lake characteristics, we used latitude and mean depth because they were most strongly correlated to TNOs values and because they were least correlated to the other predictors in the model. We used a 100-fold cross-validation with a 70–30% split by lake (that is, 70% of lakes were used in each BRT). Model results were averaged to ensure that the patterns described therein were robust to the exclusion of some lakes. We optimized the learning rate for each BRT by iteratively running the model with smaller and smaller learning rates (from 0.8, 0.4, 0.2, 0.1, 0.05 to 0.025) until the number of trees in the model was greater than 1,000, as suggested in previous literature68. We found that the BRT performed well in cross-validation—the correlation between predicted and observed values in the test datasets from the 100-fold cross-validation was moderate on average across models (r = 0.56, Kendall’s rank correlation; see full goodness-of-fit summary statistics in Extended Data Fig. 6). The correlation between the predicted and the observed values was high (r = 0.76, Kendall’s rank correlation) when predictions were averaged across BRT. We found minimal patterning in the model residuals when comparing the model residuals with each predictor variable used in the BRT (Extended Data Fig. 7).To calculate lake-specific mean thermal non-overlap values and facilitate comparison across lakes, we used the BRT to remove the variation in thermal non-overlap attributable to the starting year of the time series, ending year of the time series, starting day of the year of the seasonal coverage, ending day of the year of the seasonal coverage, average number of sampling dates per year, linear trend (Theil–Sen slope) in the average number of sampling dates per year and the linear trend (Theil–Sen slope) in the yearly extent of the time series’ seasonal coverage of each lake’s time series, following previously published work24. We did this by setting the values for these variables to their median and using the BRT to make a prediction for each lake with these medians as predictors, along with each lake’s observed values for mean depth, absolute latitude, seasonal habitat restriction, depth habitat restriction and time series delineation. The residuals from the BRT were then added back to the predicted values used in further analyses and plotting. The mean lake-specific thermal dissimilarities were calculated as the average across all seasonal habitat restrictions (Slimit), depth habitat restrictions (Vlimit) (0, 1/2, 2/3, 5/6, 8/9, 12/13 and 19/20) and all three time series delineations. The statistical significance of these lake-specific thermal non-overlap values was estimated on a continuous gradient and calculated using a Wilcoxon signed-rank test. In the test, we compared TNO values to TNOo values separately for each combination of time series delineation, seasonal habitat restriction and depth habitat restriction (n = 108). The average P values from these tests for each lake are shown in Supplementary Table 1.We compared thermal non-overlap values to a more widely used metric of whole-lake thermal change—whole-lake temperature trends. Whole-lake temperature trends were calculated based on the annual averages of all temperature values sampled for the pairwise thermal non-overlap calculations to maximize the comparability of the resulting temperature trends and thermal non-overlap values. Due to the temperature sampling probability being volume-weighted, the temperature trend was also indirectly volume-weighted. Temperature trends were calculated using Theil–Sen slopes applied to annual mean temperatures and the statistical significance of each trend (P value) was calculated using a bootstrapped one sample Wilcoxon signed-rank test with 1,000 repetitions. The input data for the Wilcoxon signed-rank test were the complete list of all slopes derived from all pairwise combinations of points in the time series. The number of pairwise slopes used in each repetition of the Wilcoxon signed-rank test was equal to the number of years of temperature data for each lake. Whole-lake temperature trends and thermal non-overlap values were not strongly correlated (r = 0.10, Kendall’s rank correlation coefficient; Extended Data Fig. 4). All statistics and graphics were produced in the R statistical computing environment69.Reporting SummaryFurther information on research design is available in the Nature Research Reporting Summary linked to this article. More

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    First dynamics of bacterial community during development of Acropora humilis larvae in aquaculture

    1.Chavanich, S., Viyakarn, V., Loyjiw, T., Pattaratamrong, P. & Chankong, A. Mass bleaching of soft coral, Sarcophyton spp. in Thailand and the role of temperature and salinity stress. ICES J. Mar. Sci. 66, 1515–1519 (2009).2.Phongsuwan, N. et al. Status and changing patterns on coral reefs in Thailand during the last two decades. Deep Sea Res. Pt. II Top. Stud. Oceanogr. 96, 19–24 (2013).ADS 
    Article 

    Google Scholar 
    3.Gardner, T. A., Côté, I. M., Gill, J. A., Grant, A. & Watkinson, A. R. Long-term region-wide declines in Caribbean corals. Science 301, 958–960 (2003).ADS 
    CAS 
    PubMed 
    Article 

    Google Scholar 
    4.Bruno, J. F. & Selig, E. R. Regional decline of coral cover in the Indo-Pacific: Timing, extent, and subregional comparisons. PLoS ONE 2, e711. https://doi.org/10.1371/journal.pone.0000711 (2007).ADS 
    Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    5.De´ath, G., Fabricius, K. E., Sweatman, H. & Puotinen, M. The 27-year decline of coral cover on the Great Barrier Reef and its causes. PNAS 109, 17995–17999 (2012).6.Moberg, F. & Folke, C. Ecological goods and services of coral reef ecosystems. Ecol. Econ. 29, 215–233 (1999).Article 

    Google Scholar 
    7.Sheppard, C. et al. The Gulf: A young sea in decline. Mar. Pollut. Bull. 60, 13–38 (2010).CAS 
    PubMed 
    Article 

    Google Scholar 
    8.Cruz-Trinidad, A., Aliño, P. M., Geronimo, R. C. & Cabral, R. B. Linking food security with coral reefs and fisheries in the coral triangle. Coast Manag. 42, 160–182 (2014).Article 

    Google Scholar 
    9.Chavanich, S. et al. A tunicate from a Thai coral reef: A potential source of new anticancer compounds. Coral Reefs 24, 621. https://doi.org/10.1007/s00338-005-0036-y (2005).ADS 
    Article 

    Google Scholar 
    10.Rocha, J., Peixe, L., Gomes, N. & Calado, R. Cnidarians as a source of new marine bioactive compounds-an overview of the last decade and future steps for bioprospecting. Mar. Drugs 9, 1860–1886 (2011).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    11.Cooper, E. L., Hirabayashi, K., Strychar, K. B. & Sammarco, P. W. Corals and their potential applications to integrative medicine. Evid. Based Complement. Alternat. Med. 2014, 184959. https://doi.org/10.1155/2014/184959 (2014).Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    12.Petersen, D. et al. The application of sexual coral recruits for the sustainable management of ex situ populations in public aquariums to promote coral reef conservation-SECORE Project. Aquat. Conserv. 16, 167–179 (2006).Article 

    Google Scholar 
    13.Chavanich, S. & Viyakarn, V. Conservation and restoration of coral reefs under climate change: Strategies and practice. in The Cnidaria, Past, Present and Future. 787–792. (Springer, 2016).14.Boström-Einarsson, L. et al. Coral restoration–A systematic review of current methods, successes, failures and future directions. PLoS ONE 15, e0226631. https://doi.org/10.1371/journal.pone.0226631 (2020).CAS 
    Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    15.Webster, N. S. & Reusch, T. B. Microbial contributions to the persistence of coral reefs. ISME J. 11, 2167–2174 (2017).PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    16.van Oppen, M. J. & Blackall, L. L. Coral microbiome dynamics, functions and design in a changing world. Nat. Rev. Microbiol. 17, 557–567 (2019).PubMed 
    Article 
    CAS 

    Google Scholar 
    17.Lesser, M. P., Mazel, C. H., Gorbunov, M. Y. & Falkowski, P. G. Discovery of symbiotic nitrogen-fixing cyanobacteria in corals. Science 305, 997–1000 (2004).ADS 
    CAS 
    PubMed 
    Article 

    Google Scholar 
    18.Chimetto, L. A. et al. Vibrios dominate as culturable nitrogen-fixing bacteria of the Brazilian coral Mussismilia hispida. Syst. Appl. Microbiol. 31, 312–319 (2008).CAS 
    PubMed 
    Article 

    Google Scholar 
    19.Ceh, J. et al. Nutrient cycling in early coral life stages: Pocillopora damicornis larvae provide their algal symbiont (Symbiodinium) with nitrogen acquired from bacterial associates. Ecol. Evol. 3, 2393–2400 (2013).Article 

    Google Scholar 
    20.Gochfeld, D. J. & Aeby, G. S. Antibacterial chemical defenses in Hawaiian corals provide possible protection from disease. Mar. Ecol. Prog. Ser. 362, 119–128 (2008).ADS 
    Article 

    Google Scholar 
    21.Kirkwood, M., Todd, J. D., Rypien, K. L. & Johnston, A. W. The opportunistic coral pathogen Aspergillus sydowii contains dddP and makes dimethyl sulfide from dimethylsulfoniopropionate. ISME J. 4, 147–150 (2010).CAS 
    PubMed 
    Article 

    Google Scholar 
    22.Raina, J.-B. et al. Isolation of an antimicrobial compound produced by bacteria associated with reef-building corals. PeerJ 4, e2275. https://doi.org/10.7717/peerj.2275 (2016).CAS 
    Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    23.Lodwig, E. M. et al. Amino-acid cycling drives nitrogen fixation in the legume—Rhizobium symbiosis. Nature 422, 722–726 (2003).ADS 
    CAS 
    PubMed 
    Article 

    Google Scholar 
    24.Bourne, D., Iida, Y., Uthicke, S. & Smith-Keune, C. Changes in coral-associated microbial communities during a bleaching event. ISME J. 2, 350–363 (2008).CAS 
    PubMed 
    Article 

    Google Scholar 
    25.Mouchka, M. E., Hewson, I. & Harvell, C. D. Coral-associated bacterial assemblages: Current knowledge and the potential for climate-driven impacts. Integr. Comp. Biol. 50, 662–674 (2010).PubMed 
    Article 

    Google Scholar 
    26.Lema, K. A., Willis, B. L. & Bourne, D. G. Corals form characteristic associations with symbiotic nitrogen-fixing bacteria. Appl. Environ. Microbiol. 78, 3136–3144 (2012).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    27.Bourne, D. G., Morrow, K. M. & Webster, N. S. Insights into the coral microbiome: Underpinning the health and resilience of reef ecosystems. Ann. Rev. Microbiol. 70, 317–340 (2016).CAS 
    Article 

    Google Scholar 
    28.Lema, K. A., Bourne, D. G. & Willis, B. L. Onset and establishment of diazotrophs and other bacterial associates in the early life history stages of the coral Acropora millepora. Mol. Ecol. 23, 4682–4695 (2014).CAS 
    PubMed 
    Article 

    Google Scholar 
    29.Zhou, G. et al. Microbiome dynamics in early life stages of the scleractinian coral Acropora gemmifera in response to elevated pCO2. Environ. Microbiol. 19, 3342–3352 (2017).CAS 
    PubMed 
    Article 

    Google Scholar 
    30.Bernasconi, R. et al. Establishment of coral-bacteria symbioses reveal changes in the core bacterial community with host ontogeny. Front. Microbiol. 10, 1529. https://doi.org/10.3389/fmicb.2019.01529 (2019).Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    31.Damjanovic, K., Menéndez, P., Blackall, L. L. & van Oppen, M. J. H. Early life stages of a common broadcast spawning coral associate with specific bacterial communities despite lack of internalized bacteria. Microb. Ecol. 79, 706–719 (2020).CAS 
    PubMed 
    Article 

    Google Scholar 
    32.Miller, N., Maneval, P., Manfrino, C., Frazer, T. K. & Meyer, J. L. Spatial distribution of microbial communities among colonies and genotypes in nursery-reared Acropora cervicornis. PeerJ 8, e9635. https://doi.org/10.7717/peerj.9635 (2020).Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    33.Chamberland, V. F. et al. Four-year-old Caribbean Acropora colonies reared from field-collected gametes are sexually mature. Bull. Mar. Sci. 92, 263–264 (2016).Article 

    Google Scholar 
    34.Baria-Rodriguez, M. V., dela Cruz, D. W., Dizon, R. M., Yap, H. T. & Villanueva, R. D. Performance and cost-effectiveness of sexually produced Acropora granulosa juveniles compared with asexually generated coral fragments in restoring degraded reef areas. Aquat. Conserv. Mar. Freshwater Ecosyst. 29, 891–900 (2019).35.Henry, J. A., O’Neil, K. L. & Patterson, J. T. Native herbivores improve sexual propagation of threatened staghorn coral Acropora cervicornis. Front. Mar. Sci. 6, 713. https://doi.org/10.3389/fmars.2019.00713 (2019).36.Ligson, C. A., Tabalanza, T. D., Villanueva, R. D. & Cabaitan, P. C. Feasibility of early outplanting of sexually propagated Acropora verweyi for coral reef restoration demonstrated in the Philippines. Restor. Ecol. 28, 244–251 (2019).Article 

    Google Scholar 
    37.Tabalanza, T. D. et al. Successfully cultured and reared coral embryos from wild caught spawn slick in the Philippines. Aquaculture 525, 735354. https://doi.org/10.1016/j.aquaculture.2020.735354 (2020).Article 

    Google Scholar 
    38.Apprill, A., Marlow, H. Q., Martindale, M. Q. & Rappé, M. S. Specificity of associations between bacteria and the coral Pocillopora meandrina during early development. Appl. Environ. Microbiol. 78, 7467–7475 (2012).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    39.Kuanui, P., Chavanich, S., Viyakarn, V., Omori, M. & Lin, C. Effects of temperature and salinity on survival rate of cultured corals and photosynthetic efficiency of the zooxanthellae in coral tissues. Ocean Sci. J. 50, 263–268 (2015).ADS 
    CAS 
    Article 

    Google Scholar 
    40.Kuanui, P. et al. Effect of light intensity on survival and photosynthetic efficiency of cultured corals of different ages. Estuar. Coast Shelf Sci. 235, 106515. https://doi.org/10.1016/j.ecss.2019.106515 (2020).Article 

    Google Scholar 
    41.Marotz, C. et al. DNA extraction for streamlined metagenomics of diverse environmental samples. Biotechniques 62, 290–293 (2017).CAS 
    PubMed 
    Article 

    Google Scholar 
    42.Bulan, D. E. et al. Spatial and seasonal variability of reef bacterial communities in the upper Gulf of Thailand. Front Mar. Sci. 5, 441. https://doi.org/10.3389/fmars.2018.00441 (2018).Article 

    Google Scholar 
    43.Caporaso, J. G. et al. Ultra-high-throughput microbial community analysis on the Illumina HiSeq and MiSeq platforms. ISME J. 6, 1621–1624 (2012).CAS 
    PubMed 
    PubMed Central 
    Article 

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

    Google Scholar 
    45.Kozich, J. J., Westcott, S. L., Baxter, N. T., Highlander, S. K. & Schloss, P. D. Development of a dual-index sequencing strategy and curation pipeline for analyzing amplicon sequence data on the MiSeq Illumina sequencing platform. Appl. Environ. Microbiol. 79, 5112–5120 (2013).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    46.Pollock, J., Glendinning, L., Wisedchanwet, T. & Watson, M. The madness of microbiome: attempting to find consensus “best practice” for 16S microbiome studies. Appl. Environ. Microbiol. 84. https://doi.org/10.1128/AEM.02627-17 (2018).47.Bharti, R. & Grimm, D. G. Current challenges and best-practice protocols for microbiome analysis. Brief Bioinform. 22, 178–193. https://doi.org/10.1093/bib/bbz155 (2019).Article 
    PubMed Central 
    PubMed 

    Google Scholar 
    48.Kanehisa, M. & Goto, S. KEGG: Kyoto encyclopedia of genes and genomes. Nucleic Acids Res. 28, 27–30 (2000).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    49.RStudio Team. RStudio: Integrated Development for R. (RStudio, PBC, 2020).50.Olson, N., Ainsworth, T., Gates, R. & Takabayashi, M. Diazotrophic bacteria associated with Hawaiian Montipora corals: Diversity and abundance in correlation with symbiotic dinoflagellates. J. Exp. Mar. Biol. Ecol. 371, 140–146 (2009).CAS 
    Article 

    Google Scholar 
    51.Sharp, K. H., Sneed, J., Ritchie, K., Mcdaniel, L. & Paul, V. J. Induction of larval settlement in the reef coral Porites astreoides by a cultivated marine Roseobacter strain. Biol. Bull. 228, 98–107 (2015).CAS 
    PubMed 
    Article 

    Google Scholar 
    52.Sharp, K. H., Distel, D. & Paul, V. J. Diversity and dynamics of bacterial communities in early life stages of the Caribbean coral Porites astreoides. ISME J. 6, 790–801 (2012).CAS 
    PubMed 
    Article 

    Google Scholar 
    53.Apprill, A., Marlow, H. Q., Martindale, M. Q. & Rappé, M. S. The onset of microbial associations in the coral Pocillopora meandrina. ISME J. 3, 685–699 (2009).PubMed 
    Article 

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

    Google Scholar 
    55.Baquiran, J. I. P. et al. The prokaryotic microbiome of Acropora digitifera is stable under short-term artificial light pollution. Microorganisms 8, 1566. https://doi.org/10.3390/microorganisms8101566 (2020).CAS 
    Article 
    PubMed Central 
    PubMed 

    Google Scholar 
    56.Rohwer, F., Seguritan, V., Azam, F. & Knowlton, N. Diversity and distribution of coral-associated bacteria. Mar. Ecol. Prog. Ser. 243, 1–10 (2002).ADS 
    Article 

    Google Scholar 
    57.Pootakham, W. et al. High resolution profiling of coral-associated bacterial communities using full-length 16S rRNA sequence data from PacBio SMRT sequencing system. Sci. Rep. 7, 2774. https://doi.org/10.1038/s41598-017-03139-4 (2017).ADS 
    CAS 
    Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    58.Franco, Á. G., Cadavid, L. F. & Arévalo-Ferro, C. Biofilms and extracts from bacteria producing “quorum sensing” signaling molecules protomote chemotaxis and settlement behaviors in Hydractinia symbiolongicarpus (Cnidaria: Hydrozoa) larvae. Acta Biol. Colomb. 24, 150–162 (2019).Article 

    Google Scholar 
    59.Jayaprakash, N. et al. A marine bacterium, Micrococcus MCCB 104, antagonistic to vibrios in prawn larval rearing systems. Dis. Aquat. Org. 68, 39–45 (2005).CAS 
    Article 

    Google Scholar 
    60.Tsai, S., Chang, W.-C., Chavanich, S., Viyakarn, V. & Lin, C. Ultrastructural observation of oocytes in six types of stony corals. Tissue Cell 48, 349–355 (2016).PubMed 
    Article 

    Google Scholar 
    61.Lin, C., Kup, F.-W., Chavanich, S. & Viyakarn, V. Membrane lipid phase transition behavior of oocytes from three gorgonian corals in relation to chilling injury. PLoS ONE 9, e92812. https://doi.org/10.1371/journal.pone.0092812 (2014).ADS 
    CAS 
    Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    62.Shnit-Orland, M. & Kushmaro, A. Coral mucus-associated bacteria: A possible first line of defense. FEMS Microbiol. Ecol. 67, 371–380 (2009).CAS 
    PubMed 
    Article 

    Google Scholar 
    63.Leite, D. C., Salles, J. F., Calderon, E. N., van Elsas, J. D. & Peixoto, R. S. Specific plasmid patterns and high rates of bacterial co-occurrence within the coral holobiont. Ecol. Evol. 8, 1818–1832 (2018).PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    64.Rypien, K. L., Ward, J. R. & Azam, F. Antagonistic interactions among coral-associated bacteria. Environ. Microbiol. 12, 28–39 (2010).CAS 
    PubMed 
    Article 

    Google Scholar 
    65.ElAhwany, A. M., Ghozlan, H. A., ElSharif, H. A. & Sabry, S. A. Phylogenetic diversity and antimicrobial activity of marine bacteria associated with the soft coral Sarcophyton glaucum. J. Basic Microbiol. 55, 2–10 (2015).CAS 
    PubMed 
    Article 

    Google Scholar 
    66.Damjanovic, K., van Oppen, M. J., Menéndez, P. & Blackall, L. L. Experimental inoculation of coral recruits with marine bacteria indicates scope for microbiome manipulation in Acropora tenuis and Platygyra daedalea. Front. Microbiol. 10, 1702. https://doi.org/10.3389/fmicb.2019.01702 (2019).Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    67.Damjanovic, K., Blackall, L. L., Menéndez, P. & van Oppen, M. J. H. Bacterial and algal symbiont dynamics in early recruits exposed to two adult coral species. Coral Reefs 39, 189–202 (2020).Article 

    Google Scholar 
    68.Neave, M. J., Michell, C. T., Apprill, A. & Voolstra, C. R. Endozoicomonas genomes reveal functional adaptation and plasticity in bacterial strains symbiotically associated with diverse marine hosts. Sci. Rep. 7, 40579. https://doi.org/10.1038/srep40579 (2017).ADS 
    CAS 
    Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    69.Hernandez-Agreda, A., Leggat, W., Bongaerts, P., Herrera, C. & Ainsworth, T. D. Rethinking the coral microbiome: Simplicity exists within a diverse microbial biosphere. MBio 9, e00812. https://doi.org/10.1128/mBio.00812-18 (2018).Article 
    PubMed 
    PubMed Central 

    Google Scholar  More

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    Zinc isotopes from archaeological bones provide reliable tropic level information for marine mammals

    1.Horstmann‐Dehn, L., Follmann, E. H., Rosa, C., Zelensky, G. & George, C. Stable carbon and nitrogen isotope ratios in muscle and epidermis of arctic whales. Mar. Mamm. Sci. 28, E173–E190 (2012).Article 

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

    Google Scholar 
    3.Misarti, N., Finney, B. P., Maschner, H. & Wooller, M. J. Changes in northeast Pacific marine ecosystems over the last 4500 years: evidence from stable isotope analysis of bone collagen from archaeological middens. Holocene 19, 1139–1151 (2009).Article 

    Google Scholar 
    4.Szpak, P., Buckley, M., Darwent, C. M. & Richards, M. P. Long-term ecological changes in marine mammals driven by recent warming in northwestern Alaska. Glob. Chang. Biol. 24, 490–503 (2018).PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    5.Michener, R. H. & Kaufman, L. in Stable Isotopes in Ecology and Environmental Science (eds Michener, R. & Lajtha, K.), 238–282 (Oxford, 2007).6.Dunton, K. H., Saupe, S. M., Golikov, A. N., Schell, D. M. & Schonberg, S. V. Trophic relationships and isotopic gradients among arctic and subarctic marine fauna. Mar. Ecol. Prog. Ser. 56, 89–97 (1989).Article 

    Google Scholar 
    7.Ramsay, M. A. & Hobson, K. A. Polar bears make little use of terrestrial food webs: evidence from stable-carbon isotope analysis. Oecologia 86, 598–600 (1991).PubMed 
    Article 
    CAS 
    PubMed Central 

    Google Scholar 
    8.Hobson, K. A. & Welch, H. E. Determination of trophic relationships within a high Arctic marine food web using δ13C and δ15N analysis. Mar. Ecol. Prog. Ser. 84, 9–18 (1992).Article 
    CAS 

    Google Scholar 
    9.Evershed, R. P. et al. in Stable Isotopes in Ecology and Environmental Science (eds Michener, R. & Lajtha, K.) 480–540 (Oxford, 2007).10.Jaouen, K. et al. Exceptionally high δ15N values in collagen single amino acids confirm Neandertals as high-trophic level carnivores. Proc. Natl Acad. Sci. USA 116, 4928–4933 (2019).PubMed 
    Article 
    CAS 

    Google Scholar 
    11.Heuser, A., Tütken, T., Gussone, N. & Galer, S. J. Calcium isotopes in fossil bones and teeth − Diagenetic versus biogenic origin. Geochim. Cosmochim. Acta 75, 3419–3433 (2011).Article 
    CAS 

    Google Scholar 
    12.Martin, J. E., Vance, D. & Balter, V. Natural variation of magnesium isotopes in mammal bones and teeth from two South African trophic chains. Geochim. Cosmochim. Acta 130, 12–20 (2014).Article 
    CAS 

    Google Scholar 
    13.Jaouen, K., Beasley, M., Schoeninger, M., Hublin, J. J. & Richards, M. P. Zinc isotope ratios of bones and teeth as new dietary indicators: results from a modern food web (Koobi Fora, Kenya). Sci. Rep. 6, 26281 (2016).PubMed 
    PubMed Central 
    Article 
    CAS 

    Google Scholar 
    14.Martin, J. E., Tacail, T., Adnet, S., Girard, C. & Balter, V. Calcium isotopes reveal the trophic position of extant and fossil elasmobranchs. Chem. Geol. 415, 118–125 (2015).Article 
    CAS 

    Google Scholar 
    15.Jaouen, K., Szpak, P. & Richards, M. P. Zinc isotope ratios as indicators of diet and trophic level in arctic marine mammals. PLoS ONE 11, e0152299 (2016).PubMed 
    PubMed Central 
    Article 
    CAS 

    Google Scholar 
    16.Bourgon, N. et al. Zinc isotopes in Late Pleistocene fossil teeth from a Southeast Asian cave setting preserve paleodietary information. Proc. Natl Acad. Sci. USA 117, 4675–4681 (2020).PubMed 
    Article 
    CAS 

    Google Scholar 
    17.Jaouen, K. What is our toolbox of analytical chemistry for exploring ancient hominin diets in the absence of organic preservation? Quat. Sci. Rev. 197, 307–318 (2018).Article 

    Google Scholar 
    18.Minagawa, M. & Wada, E. Stepwise enrichment of 15N along food chains: further evidence and the relation between δ15N and animal age. Geochim. Cosmochim. Acta 48, 1135–1140 (1984).Article 
    CAS 

    Google Scholar 
    19.Vander Zanden, M. J. & Rasmussen, J. B. Variation in δ15N and δ13C trophic fractionation: implications for aquatic food web studies. Limnol. Oceanogr. 46, 2061–2066 (2001).Article 

    Google Scholar 
    20.Post, D. M. Using stable isotopes to estimate trophic position: models, methods, and assumptions. Ecology 83, 703–718 (2002).Article 

    Google Scholar 
    21.Moynier, F., Fujii, T., Shaw, A. S. & Le Borgne, M. Heterogeneous distribution of natural zinc isotopes in mice. Metallomics 5, 693–699 (2013).PubMed 
    Article 
    CAS 

    Google Scholar 
    22.Balter, V. et al. Contrasting Cu, Fe, and Zn isotopic patterns in organs and body fluids of mice and sheep, with emphasis on cellular fractionation. Metallomics 5, 1470–1482 (2013).PubMed 
    Article 
    CAS 

    Google Scholar 
    23.Mahan, B., Moynier, F., Jørgensen, A. L., Habekost, M. & Siebert, J. Examining the homeostatic distribution of metals and Zn isotopes in Göttingen minipigs. Metallomics 10, 1264–1281 (2018).PubMed 
    Article 
    CAS 

    Google Scholar 
    24.Jaouen, K. et al. Dynamic homeostasis modeling of Zn isotope ratios in the human body. Metallomics 11, 1049–1059 (2019).PubMed 
    Article 
    CAS 

    Google Scholar 
    25.Jaouen, K. et al. Zinc isotope variations in archeological human teeth (Lapa do Santo, Brazil) reveal dietary transitions in childhood and no contamination from gloves. PLoS ONE 15, e0232379 (2020).PubMed 
    PubMed Central 
    Article 
    CAS 

    Google Scholar 
    26.McMahon, K. W., Hamady, L. L. & Thorrold, S. R. Ocean ecogeochemistry: a review. Oceanogr. Mar. Biol. 51, 327–374 (2013).
    Google Scholar 
    27.Rau, G. H., Sweeney, R. E. & Kaplan, I. R. Plankton 13C:12C ratio changes with latitude: differences between northern and southern oceans. Deep Sea Res. Part I Oceanogr. Res. 29, 1035–1039 (1982).Article 
    CAS 

    Google Scholar 
    28.McMahon, K. W., Hamady, L. L. & Thorrold, S. R. A review of ecogeochemistry approaches to estimating movements of marine animals. Limnol. Oceanogr. 58, 697–714 (2013).Article 
    CAS 

    Google Scholar 
    29.Hedges, R. E., Clement, J. G., Thomas, C. D. L. & O’Connell, T. C. Collagen turnover in the adult femoral mid‐shaft: modeled from anthropogenic radiocarbon tracer measurements. Am. J. Phys. Anthropol. 133, 808–816 (2007).PubMed 
    Article 

    Google Scholar 
    30.Szpak, P., Savelle, J. M., Conolly, J. & Richards, M. P. Variation in late Holocene marine environments in the Canadian Arctic Archipelago: evidence from ringed seal bone collagen stable isotope compositions. Quat. Sci. Rev. 211, 136–155 (2019).Article 

    Google Scholar 
    31.Szpak, P. & Buckley, M. Sulfur isotopes (δ34S) in Arctic marine mammals: indicators of benthic vs. pelagic foraging? Mar. Ecol. Prog. Ser. https://doi.org/10.3354/meps13493 (2020).32.Reeves, R. R. in Ringed Seals in the North Atlantic (eds Heide-Jørgensen, M. P. & Lydersen, C.) 9–45 (NAMMCO Scientific Publications, 1998).33.Koehler, G., Kardynal, K. J. & Hobson, K. A. Geographical assignment of polar bears using multi-element isoscapes. Sci. Rep. 9, 9390 (2019).PubMed 
    PubMed Central 
    Article 
    CAS 

    Google Scholar 
    34.Moody, J. F. & Hodgetts, L. M. Subsistence practices of pioneering Thule–Inuit: a faunal analysis of Tiktalik. Arct. Anthropol. 50, 4–24 (2013).Article 

    Google Scholar 
    35.Dyke, A. S. et al. An assessment of marine reservoir corrections for radiocarbon dates on walrus from the Foxe Basin region of Arctic Canada. Radiocarbon 61, 67–81 (2019).Article 
    CAS 

    Google Scholar 
    36.Derocher, A. E., Wiig, Ø. & Andersen, M. Diet composition of polar bears in Svalbard and the western Barents Sea. Polar Biol. 25, 448–452 (2002).Article 

    Google Scholar 
    37.Hobson, K. A. et al. A stable isotope (δ13C, δ15N) model for the North Water food web: implications for evaluating trophodynamics and the flow of energy and contaminants. Deep Sea Res. Part II Top. Stud. Oceanogr. 49, 5131–5150 (2002).Article 
    CAS 

    Google Scholar 
    38.Iverson, S. J., Stirling, I. & Lang, S. L. C. in Top Predators in Marine Ecosystems (eds Boyd, I. L., Wanless, S. & Camphuysen, C. J.) 98–117 (Cambridge University Press, 2006).39.Thiemann, G. W., Iverson, S. J. & Stirling, I. Polar bear diets and arctic marine food webs: insights from fatty acid analysis. Ecol. Monogr. 78, 591–613 (2008).Article 

    Google Scholar 
    40.Stein, R. & MacDonald, R. W. The Organic Carbon Cycle in the Arctic Ocean (Springer, 2004).41.Lynch‐Stieglitz, J., Stocker, T. F., Broecker, W. S. & Fairbanks, R. G. The influence of air‐sea exchange on the isotopic composition of oceanic carbon: Observations and modeling. Glob. Biogeochem. Cycles 9, 653–665 (1995).Article 

    Google Scholar 
    42.Hobson, K. A., Ambrose, W. G. Jr & Renaud, P. E. Sources of primary production, benthic-pelagic coupling, and trophic relationships within the Northeast Water Polynya: insights from δ13C and δ15N analysis. Mar. Ecol. Prog. Ser. 128, 1–10 (1995).Article 

    Google Scholar 
    43.France, R., Loret, J., Mathews, R. & Springer, J. Longitudinal variation in zooplankton δ13C through the Northwest Passage: inference for incorporation of sea-ice POM into pelagic foodwebs. Polar Biol. 20, 335–341 (1998).Article 

    Google Scholar 
    44.Søreide, J. E., Hop, H., Carroll, M. L., Falk-Petersen, S. & Hegseth, E. N. Seasonal food web structures and sympagic–pelagic coupling in the European Arctic revealed by stable isotopes and a two-source food web model. Prog. Oceanogr. 71, 59–87 (2006).Article 

    Google Scholar 
    45.Saupe, S. M., Schell, D. M. & Griffiths, W. B. Carbon-isotope ratio gradients in western arctic zooplankton. Mar. Biol. 103, 427–432 (1989).Article 
    CAS 

    Google Scholar 
    46.Schell, D. M., Barnett, B. A. & Vinette, K. A. Carbon and nitrogen isotope ratios in zooplankton of the Bering, Chukchi and Beaufort seas. Mar. Ecol. Prog. Ser. 162, 11–23 (1998).Article 
    CAS 

    Google Scholar 
    47.Tamelander, T., Kivimäe, C., Bellerby, R. G., Renaud, P. E. & Kristiansen, S. Base-line variations in stable isotope values in an Arctic marine ecosystem: effects of carbon and nitrogen uptake by phytoplankton. Hydrobiologia 630, 63–73 (2009).Article 
    CAS 

    Google Scholar 
    48.Pomerleau, C. et al. Spatial patterns in zooplankton communities across the eastern Canadian sub-Arctic and Arctic waters: insights from stable carbon (δ13C) and nitrogen (δ15N) isotope ratios. J. Plankton Res. 33, 1779–1792 (2011).Article 
    CAS 

    Google Scholar 
    49.Pomerleau, C. et al. Pan-Arctic concentrations of mercury and stable isotope ratios of carbon (δ13C) and nitrogen (δ15N) in marine zooplankton. Sci. Total Environ. 551, 92–100 (2016).PubMed 
    Article 
    CAS 
    PubMed Central 

    Google Scholar 
    50.De la Vega, C., Jeffreys, R. M., Tuerena, R., Ganeshram, R. & Mahaffey, C. Temporal and spatial trends in marine carbon isotopes in the Arctic Ocean and implications for food web studies. Glob. Chang. Biol. 25, 4116–4130 (2019).PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    51.Goni, M. A., Yunker, M. B., Macdonald, R. W. & Eglinton, T. I. Distribution and sources of organic biomarkers in arctic sediments from the Mackenzie River and Beaufort Shelf. Mar. Chem. 71, 23–51 (2000).Article 
    CAS 

    Google Scholar 
    52.Parsons, T. R. et al. Autotrophic and heterotrophic production in the Mackenzie River/Beaufort Sea estuary. Polar Biol. 9, 261–266 (1989).Article 

    Google Scholar 
    53.Dehn, L. A. et al. Feeding ecology of phocid seals and some walrus in the Alaskan and Canadian Arctic as determined by stomach contents and stable isotope analysis. Polar Biol. 30, 167–181 (2007).Article 

    Google Scholar 
    54.Butt, C. M., Mabury, S. A., Kwan, M., Wang, X. & Muir, D. C. Spatial trends of perfluoroalkyl compounds in ringed seals (Phoca hispida) from the Canadian Arctic. Environ. Toxicol. Chem. 27, 542–553 (2008).PubMed 
    Article 
    CAS 

    Google Scholar 
    55.Dittmar, T. & Kattner, G. The biogeochemistry of the river and shelf ecosystem of the Arctic Ocean: a review. Mar. Chem. 83, 103–120 (2003).Article 
    CAS 

    Google Scholar 
    56.Pons, M. L. et al. A Zn isotope perspective on the rise of continents. Geobiology 11, 201–214 (2013).PubMed 
    Article 
    CAS 

    Google Scholar 
    57.Isson, T. T. et al. Tracking the rise of eukaryotes to ecological dominance with zinc isotopes. Geobiology 16, 341–352 (2018).PubMed 
    Article 
    CAS 

    Google Scholar 
    58.Samanta, M., Ellwood, M. J. & Strzepek, R. F. Zinc isotope fractionation by Emiliania huxleyi cultured across a range of free zinc ion concentrations. Limnol. Oceanogr. 63, 660–671 (2018).Article 
    CAS 

    Google Scholar 
    59.Köbberich, M. & Vance, D. Zn isotope fractionation during uptake into marine phytoplankton: implications for oceanic zinc isotopes. Chem. Geol. 523, 154–161 (2019).Article 
    CAS 

    Google Scholar 
    60.Maréchal, C. N., Nicolas, E., Douchet, C. & Albarède, F. Abundance of zinc isotopes as a marine biogeochemical tracer. Geochem. Geophys. Geosyst. 1, 1015 (2000).Article 

    Google Scholar 
    61.John, S. G. The Marine Biogeochemistry of Zinc Isotopes. [Doctoral Thesis]. (Massachusetts Institute of Technology, 2007).62.Conway, T. M. & John, S. G. The biogeochemical cycling of zinc and zinc isotopes in the North Atlantic Ocean. Glob. Biogeochem. Cycles 28, 1111–1128 (2014).Article 
    CAS 

    Google Scholar 
    63.Wyatt, N. J. et al. Biogeochemical cycling of dissolved zinc along the GEOTRACES South Atlantic transect GA10 at 40°S. Glob. Biogeochem. Cycles 28, 44–56 (2014).Article 
    CAS 

    Google Scholar 
    64.John, S. G. & Conway, T. M. A role for scavenging in the marine biogeochemical cycling of zinc and zinc isotopes. Earth Planet. Sci. Lett. 394, 159–167 (2014).Article 
    CAS 

    Google Scholar 
    65.Sieber, M. et al. Cycling of zinc and its isotopes across multiple zones of the Southern Ocean: insights from the Antarctic Circumnavigation Expedition. Geochim. Cosmochim. Acta 268, 310–324 (2020).Article 
    CAS 

    Google Scholar 
    66.Samanta, M., Ellwood, M. J., Sinoir, M. & Hassler, C. S. Dissolved zinc isotope cycling in the Tasman Sea, SW Pacific Ocean. Mar. Chem. 192, 1–12 (2017).Article 
    CAS 

    Google Scholar 
    67.Little, S. H., Vance, D., Walker-Brown, C. & Landing, W. M. The oceanic mass balance of copper and zinc isotopes, investigated by analysis of their inputs, and outputs to ferromanganese oxide sediments. Geochim. Cosmochim. Acta 125, 673–693 (2014).Article 
    CAS 

    Google Scholar 
    68.Zhao, Y., Vance, D., Abouchami, W. & De Baar, H. J. Biogeochemical cycling of zinc and its isotopes in the Southern Ocean. Geochim. Cosmochim. Acta 125, 653–672 (2014).Article 
    CAS 

    Google Scholar 
    69.Liao, W. H. et al. Zn isotope composition in the water column of the Northwestern Pacific Ocean: the importance of external sources. Glob. Biogeochem. Cycles 34, e2019GB006379 (2020).CAS 

    Google Scholar 
    70.Vance, D., de Souza, G. F., Zhao, Y., Cullen, J. T. & Lohan, M. C. The relationship between zinc, its isotopes, and the major nutrients in the North-East Pacific. Earth Planet. Sci. Lett. 525, 115748 (2019).Article 
    CAS 

    Google Scholar 
    71.Jensen, L. T. et al. Biogeochemical cycling of dissolved zinc in the Western Arctic (Arctic GEOTRACES GN01). Glob. Biogeochem. Cycles 33, 343–369 (2019).Article 
    CAS 

    Google Scholar 
    72.DeNiro, M. J. Postmortem preservation and alteration of in vivo bone collagen isotope ratios in relation to palaeodietary reconstruction. Nature 317, 806–809 (1985).Article 
    CAS 

    Google Scholar 
    73.Ambrose, S. H. Preparation and characterization of bone and tooth collagen for isotopic analysis. J. Archaeol. Sci. 17, 431–451 (1990).Article 

    Google Scholar 
    74.Muir, D. C. G. et al. Can seal eating explain elevated levels of PCBs and organochlorine pesticides in walrus blubber from eastern Hudson Bay (Canada)? Environ. Pollut. 90, 335–348 (1995).PubMed 
    Article 
    CAS 
    PubMed Central 

    Google Scholar 
    75.Young, B. G. & Ferguson, S. H. Seasons of the ringed seal: pelagic open-water hyperphagy, benthic feeding over winter and spring fasting during molt. Wildl. Res. 40, 52–60 (2013).Article 
    CAS 

    Google Scholar 
    76.Matley, J. K., Fisk, A. T. & Dick, T. A. Foraging ecology of ringed seals (Pusa hispida), beluga whales (Delphinapterus leucas) and narwhals (Monodon monoceros) in the Canadian High Arctic determined by stomach content and stable isotope analysis. Polar Res. 34, 24295 (2015).Article 
    CAS 

    Google Scholar 
    77.Michel, C., Ingram, R. G. & Harris, L. R. Variability in oceanographic and ecological processes in the Canadian Arctic Archipelago. Prog. Oceanogr. 71, 379–401 (2006).Article 

    Google Scholar 
    78.Tremblay, J. É., Gratton, Y., Carmack, E. C., Payne, C. D. & Price, N. M. Impact of the large‐scale Arctic circulation and the North Water Polynya on nutrient inventories in Baffin Bay. J. Geophys. Res. 107, 3112 (2002).Article 

    Google Scholar 
    79.Ingram, R. G., Bâcle, J., Barber, D. G., Gratton, Y. & Melling, H. An overview of physical processes in the North Water. Deep Sea Res. Part II Top. Stud. Oceanogr. 49, 4893–4906 (2002).Article 

    Google Scholar 
    80.Pauly, D., Trites, A. W., Capuli, E. & Christensen, V. Diet composition and trophic levels of marine mammals. ICES J. Mar. Sci. 55, 467–481 (1998).Article 

    Google Scholar 
    81.Woollett, J. Oakes Bay 1: a preliminary reconstruction of a Labrador Inuit seal hunting economy in the context of climate change. Geogr. Tidsskr. 110, 245–259 (2010).Article 

    Google Scholar 
    82.Stirling, I. & Archibald, W. R. Aspects of predation of seals by polar bears. J. Fish. Res. Board Can. 34, 1126–1129 (1977).Article 

    Google Scholar 
    83.Pilfold, N. W., Derocher, A. E., Stirling, I. & Richardson, E. Polar bear predatory behaviour reveals seascape distribution of ringed seal lairs. Popul. Ecol. 56, 129–138 (2014).Article 

    Google Scholar 
    84.Elorriaga-Verplancken, F., Aurioles-Gamboa, D., Newsome, S. D. & Martínez-Díaz, S. F. δ15N and δ13C values in dental collagen as a proxy for age-and sex-related variation in foraging strategies of California sea lions. Mar. Biol. 160, 641–652 (2013).Article 
    CAS 

    Google Scholar 
    85.Hauser, D. D., Laidre, K. L., Suydam, R. S. & Richard, P. R. Population-specific home ranges and migration timing of Pacific Arctic beluga whales (Delphinapterus leucas). Polar Biol. 37, 1171–1183 (2014).Article 

    Google Scholar 
    86.Harwood, L. A., Smith, T. G., Auld, J., Melling, H. & Yurkowski, D. J. Seasonal movements and diving of ringed seals, Pusa hispida, in the Western Canadian Arctic, 1999–2001 and 2010–11. Arctic 68, 193–209 (2015).Article 

    Google Scholar 
    87.Ferguson, S. H., Taylor, M. K., Born, E. W., Rosing-Asvid, A. & Messier, F. Activity and movement patterns of polar bears inhabiting consolidated versus active pack ice. Arctic 54, 49–54. (2001).Article 

    Google Scholar 
    88.Lunn, N. J. et al. Polar bear management in Canada 1997–2000. In: Proc. 13th Working Meeting of the IUCN/SSC Polar Bear Specialist Group, 23–28 June 2001, Nuuk, Greenland. Occasional Paper 26 (eds Lunn, N. J., Schliebe, S. & Born, E. W.) 41–52 (IUCN, 2002).89.Ronald, K. & Dougan, J. L. The ice lover: biology of the harp seal (Phoca groenlandica). Science 215, 928–933 (1982).PubMed 
    Article 
    CAS 
    PubMed Central 

    Google Scholar 
    90.Sergeant, D. E. Harp seals, man and ice. Can. Spec. Publ. Fish. Aquat. Sci. 114, (1991).91.Ogloff, W. R., Yurkowski, D. J., Davoren, G. K. & Ferguson, S. H. Diet and isotopic niche overlap elucidate competition potential between seasonally sympatric phocids in the Canadian Arctic. Mar. Biol. 166, 103 (2019).Article 
    CAS 

    Google Scholar 
    92.Mansfield, A. W. Seals of arctic and eastern Canada. Fish. Res. Board Canada Bull. 137 (1963).93.Sergeant, D. E. Migrations of harp seals Pagophilus groenlandicus (Erxleben) in the Northwest Atlantic. J. Fish. Res. Board Can. 22, 433–464 (1965).Article 

    Google Scholar 
    94.Richard, P. R., Heide-Jørgensen, M. P., Orr, J. R., Dietz, R. & Smith, T. G. Summer and autumn movements and habitat use by belugas in the Canadian High Arctic and adjacent areas. Arctic 54, 207–222 (2001).
    Google Scholar 
    95.Maréchal, C. N., Télouk, P. & Albarède, F. Precise analysis of copper and zinc isotopic compositions by plasma-source mass spectrometry. Chem. Geol. 156, 251–273 (1999).Article 

    Google Scholar 
    96.Moynier, F., Albarède, F. & Herzog, G. F. Isotopic composition of zinc, copper, and iron in lunar samples. Geochim. Cosmochim. Acta 70, 6103–6117 (2006).Article 
    CAS 

    Google Scholar 
    97.Toutain, J. P. et al. Evidence for Zn isotopic fractionation at Merapi volcano. Chem. Geol. 253, 74–82 (2008).Article 
    CAS 

    Google Scholar 
    98.Copeland, S. R. et al. Strontium isotope ratios (87Sr/86Sr) of tooth enamel: a comparison of solution and laser ablation multicollector inductively coupled plasma mass spectrometry methods. Rapid Commun. Mass Spectrom. 22, 3187–3194 (2008).PubMed 
    Article 
    CAS 

    Google Scholar 
    99.Brown, T. A., Nelson, D. E., Vogel, J. S. & Southon, J. R. Improved collagen extraction by modified Longin method. Radiocarbon 30, 171–177 (1988).Article 
    CAS 

    Google Scholar 
    100.Qi, H., Coplen, T. B., Geilmann, H., Brand, W. A. & Böhlke, J. K. Two new organic reference materials for δ13C and δ15N measurements and a new value for the δ13C of NBS 22 oil. Rapid Commun. Mass Spectrom. 17, 2483–2487 (2003).PubMed 
    Article 
    CAS 

    Google Scholar 
    101.Szpak, P., Metcalfe, J. Z. & Macdonald, R. A. Best practices for calibrating and reporting stable isotope measurements in archaeology. J. Archaeol. Sci. Rep. 13, 609–616 (2017).
    Google Scholar 
    102.R Core Team, R version 3.6.1 (R Foundation for Statistical Computing, Vienna, Austria, 2018).103.Haug, T. et al. Trophic level and fatty acids in harp seals compared with common minke whales in the Barents Sea. Mar. Biol. Res. 13, 919–932 (2017).Article 

    Google Scholar  More