More stories

  • in

    A state-space model to derive motorboat noise effects on fish movement from acoustic tracking data

    1.
    Desiderà, E. et al. Acoustic fish communities: Sound diversity of rocky habitats reflects fish species diversity. Mar. Ecol. Prog. Ser. 608, 183–197 (2019).
    ADS  Article  Google Scholar 
    2.
    Iorio, L. D., Gervaise, C., Lossent, J., Valentini-Poirier, C.-A. & Boissery, P. Benthic biophonic assemblages, their environmental divers, eco-acoustic scores at the level of the Western Mediterranean basin, and their implications for large-scale ecosystem monitoring. J. Acoust. Soc. Am. 144, 1692–1692 (2018).
    ADS  Article  Google Scholar 

    3.
    Buscaino, G. et al. Temporal patterns in the soundscape of the shallow waters of a Mediterranean marine protected area. Sci. Rep. 6, 1–13 (2016).
    Article  CAS  Google Scholar 

    4.
    McNett, G. D., Luan, L. H. & Cocroft, R. B. Wind-induced noise alters signaler and receiver behavior in vibrational communication. Behav. Ecol. Sociobiol. 64, 2043–2051 (2010).
    Article  Google Scholar 

    5.
    Hildebrand, J. A. Anthropogenic and natural sources of ambient noise in the ocean. Mar. Ecol. Prog. Ser. 395, 5–20 (2009).
    ADS  Article  Google Scholar 

    6.
    Popper, A. N. & Hawkins, A. D. An overview of fish bioacoustics and the impacts of anthropogenic sounds on fishes. J. Fish Biol. 94, 692–713 (2019).
    PubMed  PubMed Central  Article  Google Scholar 

    7.
    Belkovich, V. M., Bibikov, N. G., Dubrovsky, N. A., Suhoruchenko, M. N. & Zhuravlev, V. A. Preliminary estimates of low-frequency sound effect on sea animals in the Eastern Arctic. (1994).

    8.
    Mate, B. R., Stafford, K. M. & Ljungblad, D. K. A change in sperm whale (Physeter macroephalus) distribution correlated to seismic surveys in the Gulf of Mexico. J. Acoust. Soc. Am. 96, 3268–3269 (1994).
    ADS  Article  Google Scholar 

    9.
    Dunlop, R. A. et al. The behavioural response of humpback whales (Megaptera novaeangliae) to a 20 cubic inch air gun. Aquat. Mamm. 41, 412–433 (2015).
    Article  Google Scholar 

    10.
    Kunc, H. P., McLaughlin, K. E. & Schmidt, R. Aquatic noise pollution: Implications for individuals, populations, and ecosystems. Proc. R. Soc. B Biol. Sci. 283, 20160839 (2016).
    Article  Google Scholar 

    11.
    Rako-Gospić, N. & Picciulin, M. Underwater noise: Sources and effects on marine life. in World Seas: An Environmental Evaluation Volume III: Ecological Issues and Environmental Impacts (ed. Sheppard, C.) vol. 3 367–389 (Elsevier, 2019).

    12.
    Duarte, C. M., Chapuis, L., Collin, S. P., Costa, D. P., Devassy, R. P., Eguiluz, V. M., Erbe, C., Gordon, T. A. C., Halpern, B. S., Harding, H. R., Havlik, M. N., Meekan, M., Merchant, N. D., Miksis-Olds, J. L., Parsons, M., Predragovic, M., Radford, A. N., Radford, C. A., Simpson, S. D., Slabbekoorn, H., Staaterman, E., Van Opzeeland, I. C., Winderen, J., Zhang, X. & Juanes, F. The soundscape of the Anthropocene ocean. Science 371(6529), eaba4658 (2021).
    CAS  PubMed  Article  Google Scholar 

    13.
    Cox, K., Brennan, L. P., Gerwing, T. G., Dudas, S. E. & Juanes, F. Sound the alarm: A meta-analysis on the effect of aquatic noise on fish behavior and physiology. Glob. Chang. Biol. 24, 3105–3116 (2018).
    ADS  PubMed  Article  Google Scholar 

    14.
    La Manna, G., Manghi, M., Perretti, F. & Sarà, G. Behavioral response of brown meagre (Sciaena umbra) to boat noise. Mar. Pollut. Bull. 110, 324–334 (2016).
    PubMed  Article  CAS  Google Scholar 

    15.
    Nedelec, S. L. et al. Motorboat noise impacts parental behaviour and offspring survival in a reef fish. Proc. R. Soc. B Biol. Sci. 284, 20170143 (2017).
    Article  Google Scholar 

    16.
    Maxwell, R. J. et al. Does motor noise from recreational boats alter parental care behaviour of a nesting freshwater fish?. Aquat. Conserv. Mar. Freshw. Ecosyst. 28, 969–978 (2018).
    Article  Google Scholar 

    17.
    Picciulin, M., Sebastianutto, L., Codarin, A., Calcagno, G. & Ferrero, E. A. Brown meagre vocalization rate increases during repetitive boat noise exposures: A possible case of vocal compensation. J. Acoust. Soc. Am. 132, 3118–3124 (2012).
    ADS  PubMed  Article  Google Scholar 

    18.
    de Jong, K., Amorim, M. C. P., Fonseca, P. J., Klein, A. & Heubel, K. U. Noise affects acoustic courtship behavior similarly in two species of gobies. Proc. Meet. Acoust. 27, 010018 (2016).
    Article  Google Scholar 

    19.
    Nedelec, S. L. et al. Motorboat noise disrupts co-operative interspecific interactions. Sci. Rep. 7, 1–8 (2017).
    ADS  CAS  Article  Google Scholar 

    20.
    Codarin, A., Wysocki, L. E., Ladich, F. & Picciulin, M. Effects of ambient and boat noise on hearing and communication in three fish species living in a marine protected area (Miramare, Italy). Mar. Pollut. Bull. 58, 1880–1887 (2009).
    CAS  PubMed  Article  Google Scholar 

    21.
    Holles, S., Simpson, S. D., Radford, A. N., Berten, L. & Lecchini, D. Boat noise disrupts orientation behaviour in a coral reef fish. Mar. Ecol. Prog. Ser. 485, 295–300 (2013).
    ADS  Article  Google Scholar 

    22.
    Pine, M. K., Jeffs, A. G., Wang, D. & Radford, C. A. The potential for vessel noise to mask biologically important sounds within ecologically significant embayments. Ocean Coast. Manag. 127, 63–73 (2016).
    Article  Google Scholar 

    23.
    de Jong, K., Amorim, M. C. P., Fonseca, P. J., Fox, C. J. & Heubel, K. U. Noise can affect acoustic communication and subsequent spawning success in fish. Environ. Pollut. 237, 814–823 (2018).
    PubMed  Article  CAS  Google Scholar 

    24.
    Montgomery, J. C., Jeffs, A., Simpson, S. D., Meekan, M. & Tindle, C. Sound as an orientation cue for the pelagic larvae of reef fishes and decapod crustaceans. Adv. Mar. Biol. 51, 143–196 (2006).
    PubMed  Article  Google Scholar 

    25.
    Hussey, N. E. et al. Aquatic animal telemetry: A panoramic window into the underwater world. Science 348, 1255642 (2015).
    PubMed  Article  CAS  Google Scholar 

    26.
    Simpfendorfer, C. A., Heupel, M. R. & Collins, A. B. Variation in the performance of acoustic receivers and its implication for positioning algorithms in a riverine setting. Can. J. Fish. Aquat. Sci. 65, 482–492 (2008).
    Article  Google Scholar 

    27.
    Abecasis, D. et al. A review of acoustic telemetry in Europe and the need for a regional aquatic telemetry network. Anim. Biotelemetry 6, 12 (2018).
    Article  Google Scholar 

    28.
    Kessel, S. T. et al. A review of detection range testing in aquatic passive acoustic telemetry studies. Rev. Fish Biol. Fish. 24, 199–218 (2014).
    Article  Google Scholar 

    29.
    Patterson, T. A., Thomas, L., Wilcox, C., Ovaskainen, O. & Matthiopoulos, J. State-space models of individual animal movement. Trends Ecol. Evol. 23, 87–94 (2008).
    PubMed  Article  Google Scholar 

    30.
    Alós, J., Palmer, M., Balle, S. & Arlinghaus, R. Bayesian state-space modelling of conventional acoustic tracking provides accurate descriptors of home range behavior in a small-bodied coastal fish species. PLoS ONE 11, 1–23 (2016).
    Article  CAS  Google Scholar 

    31.
    Jonsen, I. D. et al. State-space models for bio-loggers: A methodological road map. Deep Sea Res. Top. Stud. Oceanogr. 88, 34–46 (2013).
    ADS  Article  Google Scholar 

    32.
    Bauchot, M. . Serranidae. in Guide FAO d’identification des espèces pour les besoins de la pêche (eds. Fisher, W., Bauchot, M. L. & Scheneider, M.) 1301–1329 (Organisation des Nations Unies pour l’Alimentation et l’Agriculture, 1987).

    33.
    Börger, L., Dalziel, B. D. & Fryxell, J. M. Are there general mechanisms of animal home range behaviour? A review and prospects for future research. Ecol. Lett. 11, 637–650 (2008).
    PubMed  Article  Google Scholar 

    34.
    Alós, J. et al. Selective exploitation of spatially structured coastal fish populations by recreational anglers may lead to evolutionary downsizing of adults. Mar. Ecol. Prog. Ser. 503, 219–233 (2014).
    ADS  Article  Google Scholar 

    35.
    Palmer, M., Balle, S., March, D., Alós, J. & Linde, M. Size estimation of circular home range from fish mark-release-(single)-recapture data: Case study of a small labrid targeted by recreational fishing. Mar. Ecol. Prog. Ser. 430, 87–97 (2011).
    ADS  Article  Google Scholar 

    36.
    Mills, S. C. et al. Hormonal and behavioural effects of motorboat noise on wild coral reef fish. Environ. Pollut. 262, 114250 (2020).
    CAS  PubMed  Article  Google Scholar 

    37.
    Hitt, S., Pittman, S. J. & Nemeth, R. S. Diel movements of fishes linked to benthic seascape structure in a Caribbean coral reef ecosystem. Mar. Ecol. Prog. Ser. 427, 275–291 (2011).
    ADS  Article  Google Scholar 

    38.
    March, D., Palmer, M., Alós, J., Grau, A. & Cardona, F. Short-term residence, home range size and diel patterns of the painted comber Serranus scriba in a temperate marine reserve. Mar. Ecol. Prog. Ser. 400, 195–206 (2010).
    ADS  Article  Google Scholar 

    39.
    Codling, E. A., Plank, M. J. & Benhamou, S. Random walk models in biology. J. R. Soc. Interface 5, 813–834 (2008).
    PubMed  PubMed Central  Article  Google Scholar 

    40.
    Campos-Candela, A., Palmer, M., Balle, S., Álvarez, A. & Alós, J. A mechanistic theory of personality-dependent movement behaviour based on dynamic energy budgets. Ecol. Lett. 22, 213–232 (2019).
    PubMed  Article  Google Scholar 

    41.
    Gardiner, C. W. Handbook of Stochastic Methods for Physics, Chemistry and the Natural Sciences (Springer, Berlin, 1990).
    Google Scholar 

    42.
    Follana-Berná, G., Palmer, M., Lekanda-Guarrotxena, A., Grau, A. & Arechavala-Lopez, P. Fish density estimation using unbaited cameras: Accounting for environmental-dependent detectability. J. Exp. Mar. Bio. Ecol. 527, 151376 (2020).
    Article  Google Scholar 

    43.
    Klimley, A. P., Voegeli, F., Beavers, S. C. & Le Boeuf, B. J. Automated listening stations for tagged marine fishes. Mar. Technol. Soc. J. 32, 94–101 (1998).
    Google Scholar 

    44.
    Hedger, R. D. et al. The optimized interpolation of fish positions and speeds in an array of fixed acoustic receivers. ICES J. Mar. Sci. 65, 1248–1259 (2008).
    Article  Google Scholar 

    45.
    Merchant, N. D., Blondel, P., Dakin, D. T. & Dorocicz, J. Averaging underwater noise levels for environmental assessment of shipping. J. Acoust. Soc. Am. 132, 343–349 (2012).
    ADS  Article  Google Scholar 

    46.
    Plummer, M. rjags: Bayesian graphical models using MCMC. (2016).

    47.
    Su, Y. S. & Yajima, M. R2jags: Using R to run ‘JAGS’. (2015).

    48.
    R Development Core Team. R: A language and environment for statistical computing. (2017).

    49.
    Plummer, M., Best, N., Cowles, K. & Vines, K. CODA: convergence diagnosis and output analysis for MCMC. R News 6, 7–11 (2006).
    Google Scholar 

    50.
    Gelman, A. et al. Bayesian data analysis. (Taylor & Francis Group, 2013).

    51.
    Fagan, W. F. & Calabrese, J. M. The correlated random walk and the rise of movement ecology. Bull. Ecol. Soc. Am. 95, 204–206 (2014).
    Article  Google Scholar 

    52.
    Payne, N. L., Gillanders, B. M., Webber, D. M. & Semmens, J. M. Interpreting diel activity patterns from acoustic telemetry: The need for controls. Mar. Ecol. Prog. Ser. 419, 295–301 (2010).
    ADS  Article  Google Scholar 

    53.
    Burger, J. & Gochfeld, M. On developing bioindicators for human and ecological health. Environ. Monit. Assess. 66, 23–46 (2001).
    CAS  PubMed  Article  Google Scholar 

    54.
    Alós, J., March, D., Palmer, M., Grau, A. & Morales-Nin, B. Spatial and temporal patterns in Serranus cabrilla habitat use in the NW Mediterranean revealed by acoustic telemetry. Mar. Ecol. Prog. Ser. 427, 173–186 (2011).
    ADS  Article  Google Scholar 

    55.
    Alós, J., Palmer, M., Rosselló, R. & Arlinghaus, R. Fast and behavior-selective exploitation of a marine fish targeted by anglers. Sci. Rep. 6, 1–13 (2016).
    Article  CAS  Google Scholar 

    56.
    Campioni, L. et al. Individual and spatio-temporal variations in the home range behaviour of a long-lived, territorial species. Oecologia 172, 371–385 (2013).
    ADS  PubMed  Article  Google Scholar 

    57.
    Topping, D. T. & Szedlmayer, S. T. Home range and movement patterns of red snapper (Lutjanus campechanus) on artificial reefs. Fish. Res. 112, 77–84 (2011).
    Article  Google Scholar 

    58.
    Jorgensen, S. J. et al. Limited movement in blue rockfish Sebastes mystinus: Internal structure of home range. Mar. Ecol. Prog. Ser. 327, 157–170 (2006).
    ADS  Article  Google Scholar 

    59.
    Harding, H. R. et al. Fish in habitats with higher motorboat disturbance show reduced sensitivity to motorboat noise. Biol. Lett. 14, 20180441 (2018).
    PubMed  PubMed Central  Article  Google Scholar 

    60.
    Holmes, L. J., McWilliam, J., Ferrari, M. C. O. & McCormick, M. I. Juvenile damselfish are affected but desensitize to small motor boat noise. J. Exp. Mar. Bio. Ecol. 494, 63–68 (2017).
    Article  Google Scholar 

    61.
    Huntingford, F. A. et al. Current issues in fish welfare. J. Fish Biol. 68, 332–372 (2006).
    Article  Google Scholar 

    62.
    Sierra-Flores, R., Atack, T., Migaud, H. & Davie, A. Stress response to anthropogenic noise in Atlantic cod Gadus morhua L. Aquac. Eng. 67, 67–76 (2015).
    Article  Google Scholar 

    63.
    de Jong, K. et al. Predicting the effects of anthropogenic noise on fish reproduction. Rev. Fish Biol. Fish. 30, 1–24 (2020).
    Article  Google Scholar 

    64.
    McCormick, M. I., Fakan, E. P., Nedelec, S. L. & Allan, B. J. M. Effects of boat noise on fish fast-start escape response depend on engine type. Sci. Rep. 9, 1–10 (2019).
    Article  CAS  Google Scholar 

    65.
    Hawkins, A. D. & Popper, A. N. A sound approach to assessing the impact of underwater noise on marine fishes and invertebrates. ICES J. Mar. Sci. 74, 635–651 (2017).
    Article  Google Scholar 

    66.
    Soudijn, F. H., van Kooten, T., Slabbekoorn, H. & de Roos, A. M. Population-level effects of acoustic disturbance in Atlantic cod: a size-structured analysis based on energy budgets. Proc. Royal Soc. B. Sci. 287(1929), 20200490 (2020).
    Article  Google Scholar  More

  • in

    DNA traces the origin of honey by identifying plants, bacteria and fungi

    1.
    Bogdanov, S., Ruoff, K. & Persano Oddo, L. Physico-chemical methods for the characterisation of unifloral honeys: a review. Apidologie 35, S4–S17 (2004).
    Article  Google Scholar 
    2.
    Kwakman, P. H. S., te Velde, A. A., de Boer, L., Vandenbroucke-Grauls, C. M. J. E. & Zaat, S. A. J. Two major medicinal honeys have different mechanisms of bactericidal activity. PLoS ONE 6, e17709 (2011).
    ADS  CAS  PubMed  PubMed Central  Article  Google Scholar 

    3.
    Lu, J. et al. The effect of New Zealand Kanuka, Manuka and Clover Honeys on bacterial growth dynamics and cellular morphology varies according to the species. PLoS ONE 8, e55898 (2013).
    ADS  CAS  PubMed  PubMed Central  Article  Google Scholar 

    4.
    Salonen, A., Ollikka, T., Grönlund, E., Ruottinen, L. & Julkunen-Tiitto, R. Pollen analyses of honey from Finland. Grana 48, 281–289 (2009).
    Article  Google Scholar 

    5.
    Balkanska, R., Stefanova, K. & Stoikova-Grigorova, R. Main honey botanical components and techniques for identification: a review. J. Apic. Res. https://doi.org/10.1080/00218839.2020.1765481 (2020).
    Article  Google Scholar 

    6.
    Soares, S., Amaral, J. S., Oliveira, M. B. P. P. & Mafra, I. A comprehensive review on the main honey authentication issues: production and origin. Compr. Rev. Food Sci. Food Saf. 16, 1072–1100 (2017).
    CAS  PubMed  Article  Google Scholar 

    7.
    Beckmann, K., Beckh, G., Luellmann, C. & Speer, K. Characterization of filtered honey by electrophoresis of enzyme fractions. Apidologie 42, 59–66 (2011).
    CAS  Article  Google Scholar 

    8.
    Anklam, E. A review of the analytical methods to determine the geographical and botanical origin of honey. Food Chem. 63, 549–562 (1998).
    CAS  Article  Google Scholar 

    9.
    Von Der Ohe, W., Persano Oddo, L., Piana, M. L., Morlot, M. & Martin, P. Harmonized methods of melissopalynology. Apidologie 35, 18–25 (2004).
    Article  Google Scholar 

    10.
    Bell, K. L. et al. Pollen DNA barcoding: Current applications and future prospects. Genome 59, 629–640 (2016).
    PubMed  Article  PubMed Central  Google Scholar 

    11.
    Guertler, P., Eicheldinger, A., Muschler, P., Goerlich, O. & Busch, U. Automated DNA extraction from pollen in honey. Food Chem. 149, 302–306 (2014).
    CAS  PubMed  Article  Google Scholar 

    12.
    Hawkins, J. et al. Using DNA metabarcoding to identify the floral composition of honey: A new tool for investigating honey bee foraging preferences. PLoS ONE 10, e0134735 (2015).
    PubMed  PubMed Central  Article  CAS  Google Scholar 

    13.
    Valentini, A., Miquel, C. & Taberlet, P. DNA barcoding for honey biodiversity. Diversity 2, 610–617 (2010).
    CAS  Article  Google Scholar 

    14.
    Prosser, S. W. J. & Hebert, P. D. N. Rapid identification of the botanical and entomological sources of honey using DNA metabarcoding. Food Chem. 214, 183–191 (2017).
    CAS  PubMed  Article  Google Scholar 

    15.
    Olivieri, C., Marota, I., Rollo, F. & Luciani, S. Tracking plant, fungal, and bacterial DNA in honey specimens. J. Forensic Sci. 57, 222–227 (2012).
    CAS  PubMed  Article  Google Scholar 

    16.
    Snowdon, J. A. & Cliver, D. O. Microorganisms in honey. Int. J. Food Microbiol. 31, 1–26 (1996).
    CAS  PubMed  Article  Google Scholar 

    17.
    Manirajan, B. A. et al. Diversity, specificity, co-occurrence and hub taxa of the bacterial-fungal pollen microbiome. FEMS Microbiol. Ecol. 94, 1–11 (2018).
    Article  CAS  Google Scholar 

    18.
    Anderson, K. E. et al. Microbial ecology of the hive and pollination landscape: Bacterial associates from floral nectar, the alimentary tract and stored food of honey bees (Apis mellifera). PLoS ONE 8, e83125 (2013).
    ADS  PubMed  PubMed Central  Article  CAS  Google Scholar 

    19.
    Aizenberg-Gershtein, Y., Izhaki, I. & Halpern, M. Do honeybees shape the bacterial community composition in floral nectar?. PLoS ONE 8, e83125 (2013).
    ADS  Article  CAS  Google Scholar 

    20.
    Fridman, S., Izhaki, I., Gerchman, Y. & Halpern, M. Bacterial communities in floral nectar. Environ. Microbiol. Rep. 4, 97–104 (2012).
    PubMed  Article  PubMed Central  Google Scholar 

    21.
    Nevas, M. et al. High prevalence of Clostridium botulinum types A and B in honey samples detected by polymerase chain reaction. Int. J. Food Microbiol. 72, 45–52 (2002).
    CAS  PubMed  Article  PubMed Central  Google Scholar 

    22.
    Bonilla-Rosso, G. & Engel, P. Functional roles and metabolic niches in the honey bee gut microbiota. Curr. Opin. Microbiol. 43, 69–76 (2018).
    CAS  PubMed  Article  PubMed Central  Google Scholar 

    23.
    Engel, P., Martinson, V. G. & Moran, N. A. Functional diversity within the simple gut microbiota of the honey bee. Proc. Natl. Acad. Sci. U. S. A. 109, 11002–11007 (2012).
    ADS  CAS  PubMed  PubMed Central  Article  Google Scholar 

    24.
    Oksanen, J. et al. Package ‘vegan’ Title Community Ecology Package Version 2.5-6. (2019).

    25.
    Larsson, J. Area-Proportional Euler and Venn Diagrams with Ellipses [R package eulerr version 6.1.0].

    26.
    Warton, D. I. et al. So many variables: Joint modeling in community ecology. Trends Ecol. Evol. 30, 766–779 (2015).
    PubMed  Article  Google Scholar 

    27.
    Tjur, T. Coefficients of determination in logistic regression models—A new proposal: The coefficient of discrimination. Am. Stat. 63, 366–372 (2009).
    MathSciNet  MATH  Article  Google Scholar 

    28.
    Fielding, A. H. & Bell, J. F. A review of methods for the assessment of prediction errors in conservation presence/absence models (2020).https://doi.org/10.1017/S0376892997000088

    29.
    Guisan, A. et al. Measuring model accuracy: Which metrics to use? in Habitat Suitability and Distribution Models 241–269 (Cambridge University Press, 2017). doi:https://doi.org/10.1017/9781139028271.022.

    30.
    Tikhonov, G. et al. Joint species distribution modelling with the r-package HMSC. Methods Ecol. Evol. 11, 442–447 (2020).
    PubMed  PubMed Central  Article  Google Scholar 

    31.
    Moran, N. A., Hansen, A. K., Powell, J. E. & Sabree, Z. L. Distinctive gut microbiota of honey bees assessed using deep sampling from individual worker bees. PLoS ONE 7, 1–10 (2012).
    Google Scholar 

    32.
    Fünfhaus, A., Ebeling, J. & Genersch, E. Bacterial pathogens of bees. Curr. Opin. Insect Sci. 26, 89–96 (2018).
    PubMed  Article  Google Scholar 

    33.
    Fries, I. Nosema ceranae in European honey bees (Apis mellifera). J. Invertebr. Pathol. 103, (2010).

    34.
    Balvočiute, M. & Huson, D. H. SILVA, RDP, Greengenes, NCBI and OTT—how do these taxonomies compare?. BMC Genomics 18, 114 (2017).
    PubMed  PubMed Central  Article  Google Scholar 

    35.
    Meiklejohn, K. A., Damaso, N. & Robertson, J. M. Assessment of BOLD and GenBank—their accuracy and reliability for the identification of biological materials. PLoS One 14 (2019).

    36.
    Nilsson, R. H. et al. The UNITE database for molecular identification of fungi: handling dark taxa and parallel taxonomic classifications. Nucleic Acids Res. 47, 259–264 (2018).
    Article  CAS  Google Scholar 

    37.
    Sickel, W. et al. Increased efficiency in identifying mixed pollen samples by meta-barcoding with a dual-indexing approach. BMC Ecol. 15, 1–9 (2015).
    Article  CAS  Google Scholar 

    38.
    Cole, J. R. et al. The Ribosomal Database Project: Improved alignments and new tools for rRNA analysis. Nucleic Acids Res. 37, 141–145 (2008).
    Article  CAS  Google Scholar 

    39.
    Bell, K. L., Loeffler, V. M. & Brosi, B. J. An rbcL reference library to aid in the identification of plant species mixtures by DNA metabarcoding. Appl. Plant Sci. 5, 1600110 (2017).
    Article  Google Scholar 

    40.
    Edgar, R. C. & Flyvbjerg, H. Error filtering, pair assembly and error correction for next-generation sequencing reads. Bioinformatics 31, 3476–3482 (2015).
    CAS  PubMed  Article  Google Scholar 

    41.
    Callahan, B. J., McMurdie, P. J. & Holmes, S. P. Exact sequence variants should replace operational taxonomic units in marker-gene data analysis. ISME J. 11, 2639–2643 (2017).
    PubMed  PubMed Central  Article  Google Scholar 

    42.
    Rognes, T., Flouri, T., Nichols, B., Quince, C. & Mahé, F. VSEARCH: a versatile open source tool for metagenomics. PeerJ https://doi.org/10.7717/peerj.2584 (2016).
    Article  PubMed  PubMed Central  Google Scholar 

    43.
    Vesterinen, E. J., Kaunisto, K. M. & Lilley, T. M. A global class reunion with multiple groups feasting on the declining insect smorgasbord. Sci. Rep. 10, 16595 (2020).
    CAS  PubMed  PubMed Central  Article  Google Scholar 

    44.
    Barcaccia, G., Lucchin, M. & Cassandro, M. DNA barcoding as a molecular tool to track down mislabeling and food piracy. Diversity 8, 2 (2015).
    Article  CAS  Google Scholar 

    45.
    Zábrodská, B. & Vorlová, L. Adulteration of honey and available methods for detection—a review. Acta Vet. Brno 83, S85–S102 (2014).
    Article  Google Scholar 

    46.
    Hebert, P. D. N., Cywinska, A., Ball, S. L. & DeWaard, J. R. Biological identifications through DNA barcodes. Proc. R. Soc. B Biol. Sci. 270, 313–321 (2003).
    CAS  Article  Google Scholar 

    47.
    DeSalle, R. & Goldstein, P. Review and Interpretation of Trends in DNA Barcoding. Front. Ecol. Evol. 7, 302 (2019).
    Article  Google Scholar 

    48.
    Hawkins, J. et al. Using DNA metabarcoding to identify the floral composition of honey: A new tool for investigating honey bee foraging preferences. PLoS One 10 (2015).

    49.
    De Vere, N. et al. Using DNA metabarcoding to investigate honey bee foraging reveals limited flower use despite high floral availability. Sci. Rep. 7, 1–10 (2017).
    Article  CAS  Google Scholar 

    50.
    Lucek, K. et al. Metabarcoding of honey to assess differences in plant-pollinator interactions between urban and non-urban sites. https://doi.org/10.1007/s13592-019-00646-3.

    51.
    Bruni, I. et al. A DNA barcoding approach to identify plant species in multiflower honey. Food Chem. 170, 308–315 (2015).
    CAS  PubMed  Article  Google Scholar 

    52.
    Laha, R. C. et al. Meta-barcoding in combination with palynological inference is a potent diagnostic marker for honey floral composition. AMB Express 7, 132 (2017).
    PubMed  PubMed Central  Article  CAS  Google Scholar 

    53.
    Utzeri, V. J., Ribani, A. & Fontanesi, L. Authentication of honey based on a DNA method to differentiate Apis mellifera subspecies: Application to Sicilian honey bee (A. m. siciliana) and Iberian honey bee (A. m. iberiensis) honeys. Food Control 91, 294–301 (2018).
    CAS  Article  Google Scholar 

    54.
    Bovo, S. et al. Shotgun metagenomics of honey DNA: Evaluation of a methodological approach to describe a multi-kingdom honey bee derived environmental DNA signature. PLoS ONE 13, e0205575 (2018).
    PubMed  PubMed Central  Article  CAS  Google Scholar 

    55.
    Bovo, S., Utzeri, V. J., Ribani, A. & Cabbri, R. Shotgun sequencing of honey DnA can describe honey bee derived environmental signatures and the honey bee hologenome complexity. https://doi.org/10.1038/s41598-020-66127-1.

    56.
    Vesterinen, E. J., Puisto, A. I. E., Blomberg, A. S. & Lilley, T. M. Table for five, please: Dietary partitioning in boreal bats. Ecol. Evol. 8, 10914–10937 (2018).
    PubMed  PubMed Central  Article  Google Scholar 

    57.
    Vesterinen, E. J. et al. What you need is what you eat? Prey selection by the bat Myotis daubentonii. Mol. Ecol. 25, 1581–1594 (2016).
    CAS  PubMed  Article  Google Scholar 

    58.
    Functional Genomics Unit, University of Helsinki, Finland. www.helsinki.fi/en/infrastructures/genome-analysis/biomedicum-functional-genomics-unit.

    59.
    Zhang, J., Kobert, K., Flouri, T. & Stamatakis, A. PEAR: A fast and accurate Illumina Paired-End reAd mergeR. Bioinformatics 30, 614–620 (2014).
    CAS  PubMed  Article  Google Scholar 

    60.
    Schmieder, R. & Edwards, R. Quality control and preprocessing of metagenomic datasets. Bioinformatics 27, 863–864 (2011).
    CAS  PubMed  PubMed Central  Article  Google Scholar 

    61.
    Edgar, R. C. UPARSE: Highly accurate OTU sequences from microbial amplicon reads. Nat. Methods 10, 996–998 (2013).
    CAS  PubMed  PubMed Central  Article  Google Scholar 

    62.
    Wang, Q., Garrity, G. M., Tiedje, J. M. & Cole, J. R. Naïve Bayesian classifier for rapid assignment of rRNA sequences into the new bacterial taxonomy. Appl. Environ. Microbiol. 73, 5261–5267 (2007).
    CAS  PubMed  PubMed Central  Article  Google Scholar 

    63.
    Altschul, S. F., Gish, W., Miller, W., Myers, E. W. & Lipman, D. J. Basic local alignment search tool. J. Mol. Biol. 215, 403–410 (1990).
    CAS  Article  Google Scholar 

    64.
    Benson, D. A., Karsch-Mizrachi, I., Lipman, D. J., Ostell, J. & Sayers, E. W. GenBank. Nucleic Acids Res. 39, D28–D31 (2011).
    Article  CAS  Google Scholar 

    65.
    Huson, D. H., Auch, A. F., Qi, J. & Schuster, S. C. MEGAN analysis of metagenomic data. Genome Res. 17, 377–386 (2007).
    CAS  PubMed  PubMed Central  Article  Google Scholar 

    66.
    Lee, T., Alemseged, Y. & Mitchell, A. Dropping Hints: Estimating the diets of livestock in rangelands using DNA metabarcoding of faeces. Metabarcoding Metagenomics 2, e22467 (2018).
    Article  Google Scholar 

    67.
    Alberdi, A., Garin, I., Aizpurua, O. & Aihartza, J. The foraging ecology of the Mountain Long-eared bat Plecotus macrobullaris revealed with DNA mini-barcodes. PLoS One 7, (2012).

    68.
    Bolger, A. M., Lohse, M. & Usadel, B. Genome analysis Trimmomatic: A flexible trimmer for Illumina sequence data. Bioinformatics 30, 2114–2120 (2014).
    CAS  PubMed  PubMed Central  Article  Google Scholar 

    69.
    Wood, D. E., Lu, J. & Langmead, B. Improved metagenomic analysis with Kraken 2. Genome Biol. 20, 1–13 (2019).
    Article  CAS  Google Scholar 

    70.
    National Center for Biotechnology Information (NCBI); Bethesda (MD): National Library of Medicine (US). https://www.ncbi.nlm.nih.gov/ (1988).

    71.
    Lu, J., Breitwieser, F. P., Thielen, P. & Salzberg, S. L. Bracken: Estimating species abundance in metagenomics data. PeerJ Comput. Sci. 2017, e104 (2017).
    Article  Google Scholar 

    72.
    Breitwieser, F. P. & Salzberg, S. L. Pavian: Interactive analysis of metagenomics data for microbiome studies and pathogen identification. Bioinformatics 36, 1303–1304 (2020).
    CAS  PubMed  Article  Google Scholar 

    73.
    DIN (Deutsches Institut für Normung),. Untersuchung von Honig – Bestimmung der relativen Pollenhäufigkeit. DIN 10760, 2002–2005 (2002).
    Google Scholar 

    74.
    Persano Oddo, L. et al. Main European unifloral honeys: descriptive sheets 1. Apidologie 35, 38–81 (2004).
    Article  Google Scholar 

    75.
    Piper, A. M. et al. Prospects and challenges of implementing DNA metabarcoding for high-throughput insect surveillance. Gigascience 8, 1–22 (2019).
    Article  Google Scholar 

    76.
    Ovaskainen, O. & Abrego, N. Joint species distribution modelling joint species distribution modelling (Cambridge University Press, Cambridge, 2020). https://doi.org/10.1017/9781108591720.
    Google Scholar  More

  • in

    Gharial nesting in a reservoir is limited by reduced river flow and by increased bank vegetation

    1.
    Strayer, D. L. & Dudgeon, D. Freshwater biodiversity conservation: Recent progress and future challenges. Freshw. Sci. 29, 344–358 (2010).
    Google Scholar 
    2.
    Dudgeon, D. et al. Freshwater biodiversity: Importance, threats, status and conservation challenges. Biol. Rev. 81, 163–182 (2006).
    PubMed  Article  Google Scholar 

    3.
    Reid, A. J. et al. Emerging threats and persistent conservation challenges for freshwater biodiversity. Biol. Rev. 94, 849–873 (2019).
    PubMed  Article  Google Scholar 

    4.
    He, F. et al. Freshwater megafauna diversity: Patterns, status and threats. Divers. Distrib. 24, 1395–1404 (2018).
    Article  Google Scholar 

    5.
    He, F. et al. Disappearing giants: A review of threats to freshwater megafauna. WIREs Water 4, e1208 (2017).
    Article  Google Scholar 

    6.
    Nilsson, C. & Berggren, K. Alterations of riparian ecosystems caused by river regulation: Dam operations have caused global-scale ecological changes in riparian ecosystems. How to protect river environments and human needs of rivers remains one of the most important questions of our time. BioScience 50, 783–792 (2000).

    7.
    Nilsson, C., Reidy, C. A., Dynesius, M. & Revenga, C. Fragmentation and flow regulation of the world’s large river systems. Science 308, 405–408 (2005).
    ADS  CAS  PubMed  PubMed Central  Article  Google Scholar 

    8.
    Nilsson, C. & Svedmark, M. Basic principles and ecological consequences of changing water regimes: Riparian plant communities. Environ. Manag. 30, 468–480 (2002).
    Article  Google Scholar 

    9.
    Lytle, D. A. & Poff, N. L. Adaptation to natural flow regimes. Trends Ecol. Evol. 19, 94–100 (2004).
    PubMed  Article  Google Scholar 

    10.
    Junk, W. J. & Wantzen, K. M. The flood pulse concept: New aspects, approaches and applications-an update. in Proceedings of the Second International Symposium on the Management of Large Rivers for Fisheries (eds. Welcomme, R. L. & Petr, T.) 117–149 (Bangkok: Food and Agriculture Organization and Mekong River Commission, FAO Regional Office for Asia and the Pacific, 2004).

    11.
    Wiens, J. A. Riverine landscapes: Taking landscape ecology into the water. Freshw. Biol. 47, 501–515 (2002).
    Article  Google Scholar 

    12.
    Benda, L. et al. The network dynamics hypothesis: How channel networks structure riverine habitats. Bioscience 54, 413–427 (2004).
    Article  Google Scholar 

    13.
    Poff, N. L. Beyond the natural flow regime? Broadening the hydro-ecological foundation to meet environmental flows challenges in a non-stationary world. Freshw. Biol. 63, 1011–1021 (2018).
    Article  Google Scholar 

    14.
    Castro, J. M. & Thorne, C. R. The stream evolution triangle: Integrating geology, hydrology, and biology. River Res. Appl. 35, 315–326 (2019).
    Article  Google Scholar 

    15.
    Palmer, M. & Ruhi, A. Linkages between flow regime, biota, and ecosystem processes: Implications for river restoration. Science 365, eaaw2087 (2019).

    16.
    Van Looy, K. et al. The three Rs of river ecosystem resilience: Resources, recruitment, and refugia. River Res. Appl. 35, 107–120 (2019).
    Article  Google Scholar 

    17.
    Braulik, G. T., Arshad, M., Noureen, U. & Northridge, S. P. Habitat fragmentation and species extirpation in freshwater ecosystems; causes of range decline of the Indus River Dolphin (Platanista gangetica minor). PLoS ONE 9, e101657 (2014).
    ADS  PubMed  PubMed Central  Article  CAS  Google Scholar 

    18.
    Lang, J., Chowfin, S. & Ross, J. P. Gavialis gangeticus. in The IUCN Red List of Threatened Species 2019: e.T8966A3148543 (2019). https://doi.org/10.2305/IUCN.UK.2019-1.RLTS.T8966A3148543.en.

    19.
    He, F. et al. The global decline of freshwater megafauna. Glob. Change Biol. 25, 3883–3892 (2019).
    ADS  Article  Google Scholar 

    20.
    Khanal, G. et al. Irrigation demands aggravate fishing threats to river dolphins in Nepal. Biol. Conserv. 204, 386–393 (2016).
    Article  Google Scholar 

    21.
    Paudel, S., Timilsina, Y. P., Lewis, J., Ingersoll, T. & Jnawali, S. R. Population status and habitat occupancy of endangered river dolphins in the Karnali River system of Nepal during low water season. Mar. Mammal Sci. 31, 707–719 (2015).
    Article  Google Scholar 

    22.
    Whitaker, R. & Basu, D. The Gharial (Gavialis gangeticus): A review. J. Bombay Nat. Hist. Soc. 79, 531–548 (1983).
    Google Scholar 

    23.
    Vesipa, R., Camporeale, C. & Ridolfi, L. Effect of river flow fluctuations on riparian vegetation dynamics: Processes and models. Adv. Water Resour. 110, 29–50 (2017).
    ADS  CAS  Article  Google Scholar 

    24.
    Merritt, D. M. & Cooper, D. J. Riparian vegetation and channel change in response to river regulation: a comparative study of regulated and unregulated streams in the Green River Basin, USA. Regul. Rivers Res. Mgmt. 16, 543–564 (2000).
    Article  Google Scholar 

    25.
    Latterell, J. J., Bechtold, J. S., O’keefe, T. C., Pelt, R. V. & Naiman, R. J. Dynamic patch mosaics and channel movement in an unconfined river valley of the Olympic Mountains. Freshw. Biol. 51, 523–544 (2006).

    26.
    Braatne, J. H., Rood, S. B., Goater, L. A. & Blair, C. L. Analyzing the impacts of dams on riparian ecosystems: A review of research strategies and their relevance to the Snake River through Hells Canyon. Environ. Manag. 41, 267–281 (2008).
    ADS  Article  Google Scholar 

    27.
    Merritt, D. M., Scott, M. L., LeRoy, P. N., Auble, G. T. & Lytle, D. A. Theory, methods and tools for determining environmental flows for riparian vegetation: Riparian vegetation-flow response guilds. Freshw. Biol. 55, 206–225 (2010).
    Article  Google Scholar 

    28.
    Poff, N. L. & Zimmerman, J. K. Ecological responses to altered flow regimes: A literature review to inform the science and management of environmental flows. Freshw. Biol. 55, 194–205 (2010).
    Article  Google Scholar 

    29.
    Miller, K. A., Webb, J. A., de Little, S. C. & Stewardson, M. J. Environmental flows can reduce the encroachment of terrestrial vegetation into river channels: A systematic literature review. Environ. Manag. 52, 1202–1212 (2013).
    ADS  Article  Google Scholar 

    30.
    Tonkin, J. D., Merritt, D. M., Olden, J. D., Reynolds, L. V. & Lytle, D. A. Flow regime alteration degrades ecological networks in riparian ecosystems. Nat. Ecol. Evol. 2, 86–93 (2018).
    PubMed  Article  Google Scholar 

    31.
    Liro, M. Dam reservoir backwater as a field-scale laboratory of human-induced changes in river biogeomorphology: A review focused on gravel-bed rivers. Sci. Total Environ. 651, 2899–2912 (2019).
    ADS  CAS  PubMed  Article  Google Scholar 

    32.
    Volke, M. A., Johnson, W. C., Dixon, M. D. & Scott, M. L. Emerging reservoir delta-backwaters: Biophysical dynamics and riparian biodiversity. Ecol. Monogr. 89, e01363 (2019).
    Article  Google Scholar 

    33.
    Choudhury, S. Seasonal habitat use and resource partitioning between two sympatric crocodilian populations (Gavialis gangeticus & Crocodylus palustris) in Katerniaghat Wildlife Sanctuary, India. Master’s thesis submitted to Saurashtra University, Rajkot, Gujarat, India (2011)

    34.
    MacClune, K. et al. Urgent case for recovery: What we can learn from the August 2014 Karnali River floods in Nepal. in Technical Report. Zurich Insurance Group Ltd, Zurich, ISET-International, Boulder 1–44 (2015).

    35.
    Lang, J. W. & Kumar, P. Behavioral ecology of gharial on the chambal river, India. in Crocodiles. Proceedings of the 22nd Working Meeting of the IUCN-SSC Specialist Group. 42–52 (IUCN, Gland, 2013)

    36.
    Lang, J. W. & Kumar, P. Chambal gharial ecology project-2016 update. in Crocodiles. Proceedings of the 24th Working Meeting of the IUCN-SSC Specialist Group. 136–148 (IUCN, Gland, 2016)

    37.
    Gladfelter, S. R. Training rivers, Training people: Interrogating the making of disasters and the politics of response in Nepal’s lower Karnali River basin. Master’s thesis, University of Colorado (2017). https://floodresilience.net/resources/item/training-rivers-training-people-interrogating-the-making-of-disasters-and-the-politics-of-response-in-nepals-lower-karnali-river-basin.

    38.
    Kolbe, J. J. & Janzen, F. J. Impact of nest-site selection on nest success and nest temperature in natural and disturbed habitats. Ecology 83, 269–281 (2002).
    Article  Google Scholar 

    39.
    Brown, G. P. & Shine, R. Maternal nest-site choice and offspring fitness in a tropical snake (Tropidonophis mairii, Colubridae). Ecology 85, 1627–1634 (2004).
    Article  Google Scholar 

    40.
    López-Luna, M. A., Hidalgo-Mihart, M. G., Aguirre-León, G., González-Ramón, M. D. C. & Rangel-Mendoza, J. A. Effect of nesting environment on incubation temperature and hatching success of Morelet’s crocodile (Crocodylus moreletii) in an urban lake of Southeastern Mexico. J. Therm. Boil. 49, 66–73 (2015).
    Article  Google Scholar 

    41.
    Calverley, P. M. & Downs, C. T. The past and present nesting ecology of Nile crocodiles in Ndumo Game Reserve, South Africa: Reason for concern?. J. Herpetol. 51, 19–26 (2017).
    Article  Google Scholar 

    42.
    Somaweera, R., Brien, M. L., Platt, S. G., Manolis, C. & Webber, B. L. Direct and indirect interactions with vegetation shape crocodylian ecology at multiple scales. Freshw. Biol. 64, 257–268 (2019).
    Google Scholar 

    43.
    Lang, J. W. & Andrews, H. V. Temperature-dependent sex determination in crocodilians. J. Exp. Zool. 270, 28–44 (1994).
    Article  Google Scholar 

    44.
    Andrews, H. V. & Whitaker, N. Captive breeding and reproductive biology of the Indian Gharial Gavialis gangeticus (Gmelin). in Crocodiles. Proceedings of the 17th Working Meeting of the IUCN-SSC Crocodile Specialist Group. 401–411 (IUCN, Gland, 2004).

    45.
    Rhen, T. & Lang, J. W. Phenotypic effects of incubation temperature in reptiles. In Temperature-dependent sex determination in vertebrates (eds. Valenzuela, N. & Lance, V. A.) 90–98 (Smithsonian Books, Washington, 2004).

    46.
    Singh, V. P. Status of the gharial in Uttar Pradesh and its rehabilitation. J. Bombay Nat. Hist. Soc. 75(3), 668–683 (1979).
    ADS  Google Scholar 

    47.
    Basu, D. The gharial of Katerniaghat. Sanctuary 11, 36–43 (1991).
    Google Scholar 

    48.
    Srivastava, A. K. The biology of Indian gharial, Gavialis gangeticus, with special reference to its behaviour. PhD thesis submitted at University of Lucknow, Uttar Pradesh, India (1981).

    49.
    Singh, V. P. Evaluation of gharial rehabilitation U.P. forestry project. Report prepared for biodiversity research, aided by World bank. 1–49 (2003).

    50.
    Andrews, H. V. Status of the Indian gharial, conservation action and assessment of key locations in North India. Unpublished report to Cleveland Metro Park. 1–8 (2006).

    51.
    Whitaker, R. The gharial: Going extinct again. Iguana 14, 24–33 (2007).
    Google Scholar 

    52.
    Chaudhari, S. Gharial reproduction and mortality. Iguana 15, 150–153 (2008).
    Google Scholar 

    53.
    Converse L. Katerniaghat Gharial Project 2008–2009. Report of Preliminary Findings. A Report to GCA and James Cook University, Australia. 1–8 (2009).

    54.
    Das, A., Basu, D., Converse, L. & Choudhury, S. C. Herpetofauna of Katerniaghat Wildlife Sanctuary, Uttar Pradesh, India. JoTT. 4, 2553–2568 (2012).
    Google Scholar 

    55.
    Choudhary, S., Choudhury, B. C. & Gopi, G. V. Differential response to disturbance factors for the population of sympatric crocodilians (Gavialis gangeticus and Crocodylus palustris) in Katarniaghat Wildlife Sanctuary, India. Aquat. Conserv. 27, 946–952 (2017).
    Article  Google Scholar 

    56.
    Kuussaari, M. et al. Extinction debt: A challenge for biodiversity conservation. Trends Ecol. Evol. 24, 564–571 (2009).
    PubMed  Article  Google Scholar 

    57.
    Figueiredo, L., Krauss, J., Steffan‐Dewenter, I. & Sarmento Cabral, J. Understanding extinction debts: Spatio-temporal scales, mechanisms and a roadmap for future research. Ecography 42, 1973–1990 (2019).

    58.
    Bashyal, A. et al. Gharials (Gavialis gangeticus) in Bardiya National Park of Nepal: Population, habitat, and threats. Aquat. Conserv. (in press).

    59.
    Grill, G. et al. Mapping the world’s free-flowing rivers. Nature 569, 215–221 (2019).
    ADS  CAS  PubMed  Article  PubMed Central  Google Scholar 

    60.
    Jensen, J. R. Remote Sensing of the Environment: An Earth Resource Perspective (Pearson Prentice Hall, Upper Saddle River, 2007).
    Google Scholar 

    61.
    Cohen, J. A. Coefficient of agreement for nominal scales. Educ. Psychol. Meas. 20, 37–46 (1960).
    Article  Google Scholar 

    62.
    Killick, R., Haynes, K. & Eckley, I. A. Changepoint: An R package for changepoint analysis. R package version 2.2.2 (2016). https://CRAN.R-project.org/package=changepoint

    63.
    Carpenter, S. R. & Kinne, O. Regime Shifts in Lake Ecosystems: Pattern and Variation, Vol. 15 (International Ecology Institute, Oldendorf/Luhe, 2003).

    64.
    Whited, D. C. et al. Climate, hydrologic disturbance, and succession: drivers of floodplain pattern. Ecology 88, 940–953 (2007).
    PubMed  Article  Google Scholar 

    65.
    Heffernan, J. B. Wetlands as an alternative stable state in desert streams. Ecology 89, 1261–1271 (2008).
    PubMed  Article  Google Scholar 

    66.
    R Core Team. R: A Language and Environment for Statistical Computing. (R Foundation for Statistical Computing, Vienna, 2013). http://www.R-project.org/. More

  • in

    Ecology-guided prediction of cross-feeding interactions in the human gut microbiome

    Overview of the GutCP algorithm
    Our approach uses the idea that we can leverage cross-feeding interactions—which comprise knowing the metabolites that each microbial species is capable of consuming and producing—to mechanistically connect the levels of microbes and metabolites in the human gut. Several different mechanistic models in past studies have shown that this is indeed possible18,20,29,36,37. While GutCP is generalizable and can be used with any of these models, in this paper, we use a previously published consumer-resource model20. We use this model because of its context and performance: it is built specifically for the human gut and is best able to explain the experimentally measured species composition of the gut microbiome with its resulting metabolic environment, or fecal metabolome (compared with other state-of-the-art methods, such as ref. 29). To predict the metabolome from the microbiome, it relies on a manually curated set of known cross-feeding interactions9. It then uses these known interactions to follow the stepwise flow of metabolites through the gut. At each step (ecologically, at each trophic level), the metabolites available to the gut are utilized by microbial species that are capable of consuming them, and a fraction of these metabolites are secreted as metabolic byproducts. These byproducts are then available for consumption by another set of species in the next trophic level. After several such steps, the metabolites that are left unconsumed constitute the fecal metabolome.
    We hypothesized that adding new, yet-undiscovered cross-feeding interactions would improve our ability to predict the levels of metabolites with our mechanistic and causal model. Specifically, we predict that the set of undiscovered interactions resulting in the most accurate and optimal improvement in predictions would be the most likely candidates for true cross-feeding interactions. Inferring such an optimal set of new cross-feeding interactions or reactions is the main logic driving GutCP. In what follows, we sometimes refer to cross-feeding reactions (i.e., metabolite consumption or production by microbes) as “links” in an overall cross-feeding network of the gut microbiome, whose nodes are microbes and metabolites (Fig. 1a; metabolites in blue, microbes in orange); the links themselves are directed edges connecting the nodes. Links can be of two types: consumption or nutrient uptake reactions (from nutrients to microbes) and production or nutrient secretion reactions (from microbes to their metabolic byproducts).
    Fig. 1: Overview of the GutCP algorithm.

    a Schematic of the original set of known cross-feeding interactions (top) and bar plot of the prediction error for each metabolite and microbe (bottom). The cross-feeding interactions are represented as a network, whose nodes are either metabolites (cyan circles) or microbial species (orange ellipses), and directed links represent the abilities of different species to consume (red arrows) and produce (blue arrows) individual metabolites. b GutCP adds a new consumption link (red) and production link (blue) as added links reduce the prediction errors for metabolites and microbes.

    Full size image

    The salient aspects of our method are outlined in Fig. 1. We start with the known set of consumption and production links that were originally used by the model; these links are known from direct experiments and represent a ground-truth dataset or original cross-feeding network9. These are shown in Fig. 1a through the pink and blue arrows connecting nutrients 1 through 6 with microbes (a) through (c). For each sample, using only the species abundance from the microbiome, we use the model to quantitatively estimate the microbiome’s species and metabolomic composition. Briefly, we assume that a defined set of polysaccharides, common to human diets, are available as the nutrient intake to the gut (nutrients 1 and 4 in Fig. 1a). We calculate the microbiome and metabolome profiles separately for each individual, which contain a different set of microbial species in their guts. At the first trophic level, all microbial species that are capable of using the polysaccharides (indicated by the pink arrows in Fig. 1a) consume each of them in proportion to their abundances (microbes a, b, and c in Fig. 1a). They subsequently secrete a fixed fraction of the consumed nutrients as metabolic byproducts; every species at this trophic level secretes all the metabolic byproducts it is known to secrete (blue arrows in Fig. 1a) in equal proportion (nutrients 2–6 in Fig. 1a). At the next trophic level, all species detected in the individual’s gut which can consume the newly secreted byproducts consume them as nutrients, secreting a new set of byproducts, and this continues for four trophic levels (not shown in Fig. 1a for simplicity). At the end of this process, all metabolites which remain unconsumed by the community comprise the metabolome of the individual and the microbial species which consume nutrients and grow comprise the microbiome of the individual (for a complete description, see “Methods” and previous work20).
    For each metabolite and microbial species, there can be two kinds of prediction errors, or biases: individual (the sample-specific difference between predicted and measured levels) and systematic (average difference across all samples). We focused on the “systematic bias” for each metabolite and microbial species: the average deviation of the predicted levels from the measured levels across all samples in our dataset (Fig. 1a, bottom). The systematic bias for each metabolite and microbe tells us whether our model generally tends to predict their level to be greater than observed (overpredicted), less than observed (underpredicted), or neither (well-predicted). We assume that metabolites and microbes with a large systematic bias are most likely to harbor missing consumption or production links that are relevant across many samples. We prioritize adding links to them in proportion to their systematic biases.
    After measuring the systematic bias for each metabolite and microbe, GutCP proceeds in discrete steps (Fig. 1a, b). At each step, we attempt to add a new link to the current cross-feeding network. This new link is chosen randomly from the entire set of combinatorially possible links (see “Methods”; for S species, M metabolites, and two kinds of links (consumption and production), there are a total of 2SM combinatorially possible links). We accept this link—keeping it in the current network—if it leads to an overall improvement in the agreement between the predicted and measured levels of microbes and metabolites. We repeat the process of adding new links—accepting or rejecting them—until the improvements in the levels of metabolites and microbes became insignificant. Overall, GutCP can add several links to improve the agreement between the predicted and measured levels of microbes and metabolites (in Fig. 1a, b, bottom, adding the extra red and blue link at the top results in improved predictions for metabolite (1), metabolite (3), and microbe (b). Figure 2a shows how the cross-feeding network improves over a typical GutCP run via the red trajectory, starting from the original network (Fig. 2a, top left) to the final network state (Fig. 2a, bottom right). Trajectories from 100 other runs are shown in gray. GutCP repeatably reduces both the error of the metabolome predictions (y axis; measured as ({text{log}}_{10}(frac{,text{pred}-text{meas}}{text{measurement},}))) and improves the correlation between the predicted and measured metabolomes (x axis).
    Fig. 2: Improvement in predictions using GutCP.

    a Improvement in log error (({text{log}}_{10}(frac{,text{pred}-text{meas}}{text{measurement},}))) and the correlation between the prediction and measured fecal metabolome during 100 typical runs of the GutCP algorithm. The gray point at the top left indicates the performance of the original cross-feeding network of Ref. 9, and the black points at the bottom right, that of improved networks predicted using GutCP. A trajectory example, highlighting how performance improves over a GutCP run, is shown in red, and others are shown in gray. b Rarefaction curve showing the number of unique cross-feeding interactions discovered by GutCP over 100 runs of the algorithm. c Prevalence of links, i.e., the number of GutCP runs in which they repeatedly appeared (red dots; total 100 runs) and for comparison, a corresponding binomial distribution with the same mean (black dotted line). P values for different prevalences are estimated using the one-sided binomial test.

    Full size image

    Cross-validating the newly predicted interactions
    To test if the cross-feeding interactions predicted by GutCP are generalizable to unknown datasets, we performed fourfold cross-validation. We used a sample -omics dataset of the gut microbiome and metabolome sampled from 41 human individuals, comprising 221 metabolites and 72 microbial species (data from ref. 38). We split our -omics dataset into two subsets: training (three-fourths of the individuals) and test (one-fourth of the individuals) subsets. We then ran GutCP on the training subset to discover new interactions and added them to the ground-truth interactions taken from ref. 9. Doing so resulted in a network of cross-feeding interactions learned only from the training subset of the data. Finally, we evaluated the improvement in accuracy of metabolome predictions resulting from the trained network on the unseen, test subset of the data. We repeated this process three times, each time splitting the full dataset into a training subset (with a randomly chosen three-fourths of the individuals) and test subset (with the remaining one-fourth of the individuals); finally, we calculated the average improvement in prediction accuracy over all four splits.
    We found that both the training and test set performances after using the links predicted by GutCP were significantly better than the baseline given by the original cross-feeding network (Table 1). Specifically, both measures of model performance, namely the logarithmic error and the average correlation, improved by 64% and 20%, respectively, after adding GutCP’s discovered interactions. In addition, the test set performance was comparable to the training set performance (6% difference; Table 1). This suggests that the cross-feeding interactions inferred by GutCP are not likely to be a result of over-fitting.
    Table 1 Cross-validating the newly predicted interactions.
    Full size table

    Building a consensus-based atlas of predicted cross-feeding interactions
    Having confirmed that GutCP is unlikely to over-fit data, we pooled the entire sample dataset of 41 individuals and ran 100 independent instances of our prediction algorithm on it; we verified that incorporating more instances did not qualitatively affect our results (Fig. 2b shows a rarefaction curve, which highlights the number of new links discovered by GutCP as we perform more runs the algorithm). Each run of the algorithm resulted in an average of 140 newly predicted cross-feeding interactions. Then, based on consensus from many runs, we assigned a confidence level to each predicted interaction, namely what fraction of GutCP runs it was discovered in. By calculating a null distribution (Fig. 2c, black), which predicts the fraction of GutCP runs where a random link would be discovered by chance, we assigned a P value to each link and set a threshold at P = 10−3 (Fig. 3c, red; see “Methods” for details). Doing so finally resulted in a complete consensus-based atlas of 293 predicted cross-feeding interactions, which we have provided as a resource for experimental verification in Supplementary Table 1. Figure 3a shows a condensed version of these interactions obtained from the simulation with the best performance (the trajectory example in Fig. 2a with the lowest log error and highest correlation coefficient) in the form of a matrix; specifically, newly added interactions are in dark colors, and old interactions in faded colors. Supplementary Fig. 3 shows a complete version of this matrix. Note that some of the predicted interactions in Fig. 3a are unrealistic, e.g., the production of certain sugars like D-Fructose and D-Sorbitol. Such interactions are unlikely to be predicted in repeated simulations, and thus will not be part of the final consensus set. This illustrates the power of pooling results from several simulations to arrive at a set of highly probable predictions.
    Fig. 3: New cross-feeding interactions predicted by GutCP.

    a Concise matrix representation of the improved cross-feeding network of the gut microbiome predicted by GutCP (the trajectory example in Fig. 2a with the best performance). The rows are metabolites, and columns, microbial species. Faded cells represent the original, known set of cross-feeding interactions, both production (light blue), consumption (light red), and bidirectional links (gray). The new cross-feeding interactions predicted by GutCP are shown in dark colors: production links in dark blue, consumption links in dark red, and bidirectional links in black. b Network of 293 new links predicted by GutCP (with a P value  More

  • in

    Bacteria enhance the production of extracellular polymeric substances by the green dinoflagellate Lepidodinium chlorophorum

    1.
    Siano, R. et al. Citizen participation in monitoring phytoplankton seawater discolorations. Mar. Policy 117, 1–11. https://doi.org/10.1016/j.marpol.2018.01.022 (2018).
    Article  Google Scholar 
    2.
    Elbrächter, M. & Schnepf, E. Gymnodinium chlorophorum, a new, green, bloom-forming dinoflagellate (Gymnodiniales, Dinophyceae) with a vestigial prasinophyte endosymbiont. Phycologia 35, 381–393 (1996).
    Article  Google Scholar 

    3.
    Hansen, G., Botes, L. & De Salas, M. Ultrastructure and large subunit rDNA sequences of Lepidodinium viride reveal a close relationship to Lepidodinium chlorophorum comb. Nov. (=Gymnodinium chlorophorum). Phycol. Res. 55, 25–41. https://doi.org/10.1111/j.1440-1835.2006.00442.x (2007).
    CAS  Article  Google Scholar 

    4.
    Gavalás-Olea, A. et al. 19,19′-diacyloxy signature: An atypical level of structural evolution in carotenoid pigments. Org. Lett. 18, 4642–4645. https://doi.org/10.1021/acs.orglett.6b02272 (2016).
    CAS  Article  PubMed  Google Scholar 

    5.
    Jackson, C., Knoll, A. H., Chan, C. X. & Verbruggen, H. Plastid phylogenomics with broad taxon sampling further elucidates the distinct evolutionary origins and timing of secondary green plastids. Sci. Rep. 8, 1523. https://doi.org/10.1038/s41598-017-18805-w (2018).
    ADS  CAS  Article  PubMed  PubMed Central  Google Scholar 

    6.
    Kamikawa, R. et al. Plastid genome-based phylogeny pinpointed the origin of the green-colored plastid in the dinoflagellate Lepidodinium chlorophorum. Genome Biol. Evol. 7, 1133–1140. https://doi.org/10.1093/gbe/evv060 (2015).
    CAS  Article  PubMed  PubMed Central  Google Scholar 

    7.
    Chapelle, A., Lazure, P. & Ménesguen, A. Modelling eutrophication events in a coastal ecosystem. Sensitivity analysis. Estuar. Coast. Shelf Sci. 39, 529–548. https://doi.org/10.1016/S0272-7714(06)80008-9 (1994).
    ADS  CAS  Article  Google Scholar 

    8.
    Sournia, A. et al. The repetitive and expanding occurrence of a green, bloom-forming dinoflagellate (Dinophyceae) on the coast of France. Cryptogam. Algol. 13, 1–13 (1992).
    Google Scholar 

    9.
    Claquin, P., Probert, I., Lefebvre, S. & Veron, B. Effects of temperature on photosynthetic parameters and TEP production in eight species of marine microalgae. Aquat. Microb. Ecol. 51, 1–11. https://doi.org/10.3354/ame01187 (2008).
    Article  Google Scholar 

    10.
    Alldredge, A. L., Passow, U. & Logan, B. E. The abundance and significance of a class of large, transparent organic particles in the ocean. Deep-Sea Res. 40, 1131–1140. https://doi.org/10.1016/0967-0637(93)90129-Q (1993).
    CAS  Article  Google Scholar 

    11.
    Passow, U. Transparent exopolymer particles (TEP) in aquatic environments. Prog. Oceanogr. 55, 287–333. https://doi.org/10.1016/S0079-6611(02)00138-6 (2002).
    ADS  Article  Google Scholar 

    12.
    Verdugo, P. et al. The oceanic gel phase: A bridge in the DOM-POM continuum. Mar. Chem. 92, 67–85. https://doi.org/10.1016/j.marchem.2004.06.017 (2004).
    CAS  Article  Google Scholar 

    13.
    Azam, F. & Malfatti, F. Microbial structuring of marine ecosystems. Nature 5, 782–791. https://doi.org/10.1038/nrmicro1747 (2007).
    CAS  Article  Google Scholar 

    14.
    Bittar, T. B., Passow, U., Hamaraty, L., Bidle, K. D. & Harvey, E. L. An updated method for the calibration of transparent exopolymer particle measurements. Limnol. Oceanogr. Methods. 16, 621–628. https://doi.org/10.1002/lom3.10268 (2018).
    Article  Google Scholar 

    15.
    Mari, X., Passow, U., Migon, C., Burd, A. B. & Legendre, L. Transparent exopolymer particles: Effects on carbon cycling in the ocean. Prog. Oceanogr. 151, 13–37. https://doi.org/10.1016/j.pocean.2016.11.002 (2017).
    ADS  Article  Google Scholar 

    16.
    Passow, U. et al. The origin of transparent exopolymer particles (TEP) and their role in the sedimentation of particulate matter. Cont. Shelf. Res. 21, 327–346. https://doi.org/10.1016/S0278-4343(00)00101-1 (2001).
    ADS  Article  Google Scholar 

    17.
    Jenkinson, I. R. Oceanographic implications of non-newtonian properties found in phytoplankton cultures. Nature 323, 435–437. https://doi.org/10.1038/323435a0 (1986).
    ADS  Article  Google Scholar 

    18.
    Alldredge, A. L. & Gotschalk, C. C. Direct observations of the mass flocculation of diatom blooms: Characteristics, settling velocities and formation of diatom aggregates. Deep-Sea Res. 36, 159–171. https://doi.org/10.1016/0198-0149(89)90131-3 (1989).
    ADS  CAS  Article  Google Scholar 

    19.
    Schapira, M., McQuaid, C. D. & Froneman, P. W. Free-living and particle-associated prokaryote metabolism in giant kelp forests: Implications for carbon flux in a sub-Antarctic coastal area. Estuar. Coast. Shelf. Sci. 106, 69–79. https://doi.org/10.1016/j.ecss.2012.04.031 (2012).
    ADS  CAS  Article  Google Scholar 

    20.
    Schapira, M., McQuaid, C. D. & Froneman, P. W. Metabolism of free-living particle-associated prokaryotes: Consequences for carbon flux around a Southern Ocean archipelago. J. Mar. Syst. 90, 58–66. https://doi.org/10.1016/j.jmarsys.2011.08.009 (2012).
    Article  Google Scholar 

    21.
    Bhaskar, P.V. & Bhosle, N.B. Microbial extracellular polymeric substances in marine biogeochemical processes. Curr. Sci. 88, 45–53. http://drs.nio.org/drs/handle/2264/89 (2005).

    22.
    Passow, U. & Alldredge, A. L. A dye-binding assay for the spectrophotometric measurement of transparent exopolymer particles (TEP). Limnol. Oceanogr. 40, 1326–1335. https://doi.org/10.4319/lo.1995.40.7.1326 (1995).
    ADS  CAS  Article  Google Scholar 

    23.
    Gärdes, A., Iversen, M. H., Grossart, H. P., Passow, U. & Ullrich, M. S. Diatom-associated bacteria are required for aggregation of Thalassiosira weissflogii. ISME J. 5, 436–445. https://doi.org/10.1038/ismej.2010.145 (2011).
    CAS  Article  PubMed  Google Scholar 

    24.
    Nosaka, Y., Yamashita, Y. & Suzuki, K. Dynamics and origin of transparent exopolymer particles in the Oyashio region of the Western Subarctic Pacific during the spring diatom bloom. Front. Mar. Sci. 4, 1–16. https://doi.org/10.3389/fmars.2017.00079 (2017).
    Article  Google Scholar 

    25.
    Burns, W. G., Marchetti, A. & Ziervogel, K. Enhanced formation of transparent exopolymer particles (TEP) under turbulence during phytoplankton growth. J. Plankton Res. 41, 349–361. https://doi.org/10.1093/plankt/fbz018 (2019).
    CAS  Article  Google Scholar 

    26.
    Riebesell, U., Reigstad, M., Wassmann, P., Noji, T. & Passow, U. On the trophic fate of Phaeocystis pouchetii (hariot): Significance of Phaeocystis-derived mucus for vertical flux. Neth. J. Sea Res. 33, 193–203. https://doi.org/10.1016/0077-7579(95)90006-3 (1995).
    Article  Google Scholar 

    27.
    Alderkamp, A. C., Buma, A. G. J. & van Rijssel, M. The carbohydrates of Phaeocystis and their degradation in the microbial food web. Biogeochemistry 83, 1–3. https://doi.org/10.1007/s10533-007-9078-2 (2007).
    CAS  Article  Google Scholar 

    28.
    Grossart, H. P., Simon, M. & Logan, B. E. Formation of macroscopic organic aggregates (lake snow) in a large lake: The significance of transparent exopolymer particles, phytoplankton, and zooplankton. Limnol. Oceanogr. 42, 1651–1659. https://doi.org/10.4319/lo.1997.42.8.1651 (1997).
    ADS  CAS  Article  Google Scholar 

    29.
    Iuculano, F., Mazuecos, I. P., Reche, I. & Agusti, S. Prochlorococcus as a possible source for transparent exopolymer particles (TEP). Front. Microbiol. 8, 1–11. https://doi.org/10.3389/fmicb.2017.00709 (2017).
    Article  Google Scholar 

    30.
    Thornton, D. C. O. Dissolved organic matter (DOM) release by phytoplankton in the contemporary and future ocean. Eur. J. Phycol. 49, 20–46. https://doi.org/10.1080/09670262.2013.875596 (2014).
    CAS  Article  Google Scholar 

    31.
    Zhang, Z. et al. The fate of marine bacterial exopolysaccharide in natural marine microbial communities. PLoS One 10, 1–16. https://doi.org/10.1371/journal.pone.0142690 (2015).
    CAS  Article  Google Scholar 

    32.
    Xiao, R. & Zheng, Y. Overview of microalgal extracellular polymeric substances (EPS) and their applications. Biotechnol. Adv. 34, 1225–1244. https://doi.org/10.1016/j.biotechadv.2016.08.004 (2016).
    CAS  Article  PubMed  Google Scholar 

    33.
    Thavasi, R. & Banat, I. M. Biosurfactant and bioemulsifiers from marine sources. In Biosurfactants: Research Trends and Applications, ***Chap 5 (eds Mulligan, C. N. et al.) 125–146 (CRC Press, Boca Raton, 2014).
    Google Scholar 

    34.
    Decho, A. W. & Gutierrez, T. Microbial extracellular polymeric substances (EPSs) in ocean systems. Front. Microbiol. 8, 1–28. https://doi.org/10.3389/fmicb.2017.00922 (2017).
    Article  Google Scholar 

    35.
    Parker, C. The effect of environmental stressors on biofilm formation of Chlorella vulgaris. Master thesis Appalachian State University (2013).

    36.
    Zhou, J., Mopper, K. & Passow, U. The role of surface-active carbohydrates in the formation of transparent exopolymer particles by bubble adsorption of seawater. Limnol. Oceanogr. 43, 1860–1871. https://doi.org/10.4319/lo.1998.43.8.1860 (1998).
    ADS  CAS  Article  Google Scholar 

    37.
    Fukao, T., Kimoto, K. & Kotani, Y. Production of transparent exopolymer particles by four diatom species. Fish Sci. 76, 755–760. https://doi.org/10.1007/s12562-010-0265-z (2010).
    CAS  Article  Google Scholar 

    38.
    Seebah, S., Fairfield, C., Ullrich, M. S. & Passow, U. Aggregation and sedimentation of Thalassiosira weissflogii (diatom) in a warmer and more acidified Future Ocean. PLoS One 9, 1–9. https://doi.org/10.1371/journal.pone.0112379 (2014).
    CAS  Article  Google Scholar 

    39.
    Staats, N., Stal, L. J. & Mur, L. R. Exopolysaccharide production by the epipelic diatom Cylindrotheca fusiformis: Effects of nutrient conditions. J. Exp. Mar. Biol. Ecol. 249, 13–27. https://doi.org/10.1016/S0022-0981(00)00166-0 (2000).
    CAS  Article  PubMed  Google Scholar 

    40.
    Underwood, G. J. C., Boulcott, M., Raines, C. A. & Waldron, K. Environmental effects on exopolymer production by marine benthic diatoms: Dynamics, changes in composition, and pathways of production. J. Phycol. 40, 293–304. https://doi.org/10.1111/j.1529-8817.2004.03076.x (2004).
    CAS  Article  Google Scholar 

    41.
    Engel, A. et al. Impact of CO2 enrichment on organic matter dynamics during nutrient induced coastal phytoplankton blooms. J. Plankton Res. 36, 641–657. https://doi.org/10.1093/plankt/fbt125 (2014).
    CAS  Article  Google Scholar 

    42.
    Thornton, D. C. O. & Chen, J. Exopolymer production as a function of cell permeability and death in a diatom (Thalassiosira weissflogii) and a cyanobacterium (Synechococcus elongatus). J. Phycol. 53, 245–260. https://doi.org/10.1111/jpy.12470 (2017).
    CAS  Article  PubMed  Google Scholar 

    43.
    Sugimoto, K., Fukuda, H., Abdul Baki, M. & Koike, I. Bacterial contribution to formation of transparent exopolymer particles (TEP) and seasonal trends in coastal waters of Sagami Bay, Japan. Aquat. Microb. Ecol. 46, 31–41. https://doi.org/10.3354/ame046031 (2007).
    Article  Google Scholar 

    44.
    Gordillo, F. J. L., Jiménez, C., Chavarria, J. & Niell, F. X. Photosynthetic acclimation to photon irradiance and its relation to chlorophyll fluorescence and carbon assimilation in the halotolerant green alga Dunaliella viridis. Photosynth. Res. 68, 225–235. https://doi.org/10.1023/a:1012969324756 (2001).
    CAS  Article  PubMed  Google Scholar 

    45.
    Ekelund, N. G. A. & Aronsson, K. A. Changes in chlorophyll a fluorescence in Euglena gracilis and Chlamydomonas reinhardii after exposure to wood-ash. Environ. Exp. Bot. 59, 92–98. https://doi.org/10.1016/j.envexpbot.2005.10.004 (2007).
    CAS  Article  Google Scholar 

    46.
    Cole, J. J. Interactions between bacteria and algae in aquatic ecosystems. Ann. Rev. Ecol. Syst. 13, 291–314. https://doi.org/10.1146/annurev.es.13.110182.001451 (1982).
    Article  Google Scholar 

    47.
    Joint, I. et al. Competition for inorganic nutrients between phytoplankton and bacterioplankton in nutrient manipulated mesocosms. Aquat. Microb. Ecol. 29, 145–159. https://doi.org/10.3354/ame029145 (2002).
    Article  Google Scholar 

    48.
    Amin, S. A., Parker, M. S. & Armbrust, E. V. Interactions between diatoms and bacteria. Microbiol. Mol. Biol. Rev. 76, 667–684. https://doi.org/10.1128/MMBR.00007-12 (2012).
    CAS  Article  PubMed  PubMed Central  Google Scholar 

    49.
    Ramanan, R., Kim, B. H., Cho, D. H., Oh, H. M. & Kim, H. S. Algae-bacteria interactions: Evolution, ecology and emerging applications. Biotechnol. Adv. 34, 14–29. https://doi.org/10.1016/j.biotechadv.2015.12.003 (2016).
    CAS  Article  PubMed  Google Scholar 

    50.
    Ray, S. & Bagchi, S. N. Nutrients and pH regulate algicide accumulation in cultures of the cyanobacterium Oscillatoria laetevirens. New Phytol. 149, 455–460. https://doi.org/10.1046/j.1469-8137.2001.00061.x (2001).
    CAS  Article  Google Scholar 

    51.
    Oremland, R. S. & Capone, D. G. Use of “specific” inhibitors in biogeochemistry and microbial ecology. Adv. Microb. Ecol. 10, 285–383. https://doi.org/10.1007/978-1-4684-5409-3_8 (1988).
    CAS  Article  Google Scholar 

    52.
    Middelburg, J. J. & Nieuwenhuize, J. Nitrogen uptake by heterotrophic bacteria and phytoplankton in the nitrate-rich Thames estuary. Mar. Ecol. Prog. Ser. 203, 13–21. https://doi.org/10.3354/meps203013 (2000).
    ADS  CAS  Article  Google Scholar 

    53.
    Mulholland, M. R., Rocha, A. M. & Boncillo, G. E. Incorporation of leucine and thymidine by estuarine phytoplankton: Implications for bacteria productivity estimates. Estuar. Coasts 34, 310–325. https://doi.org/10.1007/s12237-010-9366-2 (2010).
    CAS  Article  Google Scholar 

    54.
    Prieto, A. et al. Assessing the role of phytoplankton–bacterioplankton coupling in the response of microbial plankton to nutrient additions. J. Plankton Res. 38, 55–63. https://doi.org/10.1093/plankt/fbv101 (2016).
    CAS  Article  Google Scholar 

    55.
    Dakhama, A., de la Noüe, J. & Lavoie, M. C. Isolation and identification of antialgal substances produced by Pseudomonas aeruginosa. J. Appl. Phycol. 5, 297–306. https://doi.org/10.1007/BF02186232 (1993).
    CAS  Article  Google Scholar 

    56.
    Bowman, L. P. Bioactive compound synthetic capacity and ecological significance of marine bacterial genus Pseudoalteromonas. Mar. Drugs 5, 220–241. https://doi.org/10.3390/md504220 (2007).
    CAS  Article  PubMed  PubMed Central  Google Scholar 

    57.
    Meseck, S. L., Smith, B. C., Wikfors, G. H., Alix, J. H. & Kapareiko, D. Nutrient interactions between phytoplankton and bacterioplankton under different carbon dioxide regimes. J. Appl. Phycol. 19, 229–237. https://doi.org/10.1007/s10811-006-9128-5 (2007).
    CAS  Article  Google Scholar 

    58.
    Guerrini, F., Mazzotti, A., Boni, L. & Pistocchi, R. Bacterial-algal interactions in polysaccharide production. Aquat. Microb. Ecol. 15, 247–253. https://doi.org/10.3354/ame015247 (1998).
    Article  Google Scholar 

    59.
    Lu, X. et al. A marine algicidal Thalassiosira and its active substance against the harmful algal bloom species Karenia mikimotoi. Appl. Microbiol. Biotechnol. 100, 5131–5139. https://doi.org/10.1007/s00253-016-7352-8 (2016).
    CAS  Article  PubMed  Google Scholar 

    60.
    Li, Y. et al. Chitinase producing bacteria with direct algicidal activity on marine diatoms. Sci. Rep. 6, 1–13. https://doi.org/10.1038/srep21984 (2016).
    CAS  Article  Google Scholar 

    61.
    Li, Y. et al. The first evidence of deinoxanthin from Deinococcus sp Y35 with strong algicidal effect on the toxic dinoflagellate Alexandrium tamarense. J. Hazard. Mater. 290, 87–95. https://doi.org/10.1016/j.jhazmat.2015.02.070 (2015).
    ADS  CAS  Article  PubMed  Google Scholar 

    62.
    Lovejoy, C., Bowman, J. P. & Hallegraeff, G. M. Algicidal effects of a novel marine Pseudomonas isolate (class Proteobacteria, gamma subdivision) on harmful algal bloom species of the genera Chattonella, Gymnodinium and Heterosigma. Appl. Environ. Microbiol. 64, 2806–2813 (1998).
    CAS  Article  Google Scholar 

    63.
    Honsell, G. & Talarico, L. Gymnodinium chlorophorum (Dinophyceae) in the Adriatic Sea: Electron microscopical observations. Bot. Mar. 47, 152–166. https://doi.org/10.1515/BOT.2004.016 (2004).
    Article  Google Scholar 

    64.
    Iriarte, J. L., Quiñones, R. A. & González, R. R. Relationship between biomass and enzymatic activity of a bloom-forming dinoflagellate (Dinophyceae) in southern Chile (41°S): A field approach. J. Plankton. Res. 27, 159–161. https://doi.org/10.1093/plankt/fbh167 (2005).
    CAS  Article  Google Scholar 

    65.
    Gárate-Lizárraga, I., Muñetón-Gómez, M. S., Pérez-Cruz, B. & Díaz-Ortíz, J. A. Bloom of Gonyaulax spinifera (Dinophyceae: Gonyaulacales) in Ensenada de la Paz Lagoon, Gulf of California. CICIMAR Oceán. 29, 1–18 (2014).
    Google Scholar 

    66.
    McCarthy, P.M. Census of Australian Marine Dinoflagellates. Australian Biological Resources Study, Canberra. http://www.anbg.gov.au/abrs/Dinoflagellates/index_Dino.html. Accessed 11 July 2013 (2013).

    67.
    Azam, F. & Smith, D. C. Bacterial influence on the variability in the ocean’s biogeochemical state: A mechanistic view. In Particle Analysis in Oceanography. NATO ASI Series (Series G: Ecological Sciences), ***27 (ed. Demers, S.) (Springer, Berlin, 1991). https://doi.org/10.1007/978-3-642-75121-9_9.
    Google Scholar 

    68.
    Smith, D. C., Steward, G. F., Long, R. A. & Azam, F. Bacterial mediation of carbon fluxes during a diatom bloom in a mesocosm. Deep-Sea Res. 442, 75–97. https://doi.org/10.1016/0967-0645(95)00005-B (1995).
    ADS  Article  Google Scholar 

    69.
    Schuster, S. & Herndl, G. J. Formation and significance of transparent exopolymeric particles in the northern Adriatic Sea. Mar. Ecol. Prog. Ser. 124, 227–236. https://doi.org/10.3354/meps124227 (1995).
    ADS  Article  Google Scholar 

    70.
    Engel, A. & Passow, U. Carbon and nitrogen content of transparent exopolymer particles (TEP) in relation to their Alcian Blue adsorption. Mar. Ecol. Prog. Ser. 219, 1–10. https://doi.org/10.3354/meps219001 (2001).
    ADS  CAS  Article  Google Scholar 

    71.
    Hasui, M., Matsuda, M., Okutani, K. & Shigeta, S. In vitro antiviral activities of sulfated polysaccharides from a marine microalga (Cochlodinium polykrikoides) against human immunodeficiency virus and other enveloped viruses. Int. J. Biol. Macromol. 17, 293–297. https://doi.org/10.1016/0141-8130(95)98157-T (1995).
    CAS  Article  PubMed  Google Scholar 

    72.
    Yim, J. H., Kim, S. J., Ahn, S. H. & Lee, H. K. Characterization of a novel bioflocculant, p-KG03, from a marine dinoflagellate, Gyrodinium impudicum KG03. Bioresour. Technol. 98, 361–367. https://doi.org/10.1016/j.biortech.2005.12.021 (2007).
    CAS  Article  PubMed  Google Scholar 

    73.
    Mandal, S. K., Singh, R. P. & Patel, V. Isolation and characterization of exopolysaccharide secreted by a toxic dinoflagellate, Amphidinium carterae Hulburt 1957 and its probable role in harmful algal blooms (HABs). Microb. Ecol. 62, 518–527. https://doi.org/10.1007/s00248-011-9852-5 (2011).
    CAS  Article  PubMed  Google Scholar 

    74.
    Kesaulya, I., Leterme, S. C., Mitchell, J. G. & Seuront, L. The impact of turbulence and phytoplankton dynamics on foam formation, seawater viscosity and chlorophyll concentration in the eastern English Channel. Oceanologia 50, 167–182 (2008).
    Google Scholar 

    75.
    Seuront, L. & Vincent, D. Increased seawater viscosity, Phaeocystis globosa spring bloom and Temora longicornis feeding and swimming behaviours. Mar. Ecol. Prog. Ser. 363, 131–145. https://doi.org/10.3354/meps07373 (2008).
    ADS  CAS  Article  Google Scholar 

    76.
    Seuront, L., Vincent, D. & Mitchell, J. G. Biologically induced modification of seawater viscosity in the Eastern English Channel during a Phaeocystis globosa spring bloom. J. Mar. Syst. 61, 118–133. https://doi.org/10.1016/j.jmarsys.2005.04.010 (2006).
    Article  Google Scholar 

    77.
    Seuront, L. et al. The influence of Phaeocystis globosa on microscale spatial patterns of chlorophylla and bulk-phase seawater viscosity. Biogeochemistry 83, 173–188. https://doi.org/10.1007/s10533-007-9097-z (2007).
    CAS  Article  Google Scholar 

    78.
    Seuront, L. et al. Role of microbial and phytoplankton communities in the control of seawater viscosity off East Antarctica (30–80° E). Deep-Sea Res. 57, 877–886. https://doi.org/10.1016/j.dsr2.2008.09.018 (2010).
    ADS  CAS  Article  Google Scholar 

    79.
    Stoderegger, K. E. & Herndl, G. J. Production of exopolymer particles by marine bacterioplankton under contrasting turbulence conditions. Mar. Ecol. Prog. Ser. 189, 9–16. https://doi.org/10.3354/meps189009 (1999).
    ADS  CAS  Article  Google Scholar 

    80.
    Alunno-Bruscia, M. et al. A single bio-energetics growth and reproduction model for the oyster Crassostrea gigas in six Atlantic ecosystems. J. Sea Res. 66, 340–348. https://doi.org/10.1016/j.seares.2011.07.008 (2011).
    ADS  Article  Google Scholar 

    81.
    Thomas, Y. et al. Global change and climate-driven invasion of the Pacific oyster (Crassostrea gigas) along European coasts: A bioenergetics modelling approach. J. Biogeogr. 43, 568–579. https://doi.org/10.1111/jbi.12665 (2016).
    Article  Google Scholar 

    82.
    Guillard, R. & Hargraves, P. Stichochrysis immobilis is a diatom, not a chrysophyte. Phycologia 32, 234–236. https://doi.org/10.2216/i0031-8884-32-3-234.1 (1993).
    Article  Google Scholar 

    83.
    Scholin, C. A., Herzog, M., Sogin, M. & Anderson, D. M. Identification of group- and strain-specific genetic markers for globally distributed Alexandrium (Dinophyceae). II. Sequence analysis of a fragment of the LSU rRNA gene. J. Phycol. 30, 999–1011. https://doi.org/10.1111/j.0022-3646.1994.00999.x (1994).
    CAS  Article  Google Scholar 

    84.
    Nunn, G. B., Theisen, B. F., Christensen, B. & Arctander, P. Simplicity-correlated size growth of the nuclear 28S ribosomal RNA D3 expansion segment in the crustacean order Isopoda. J. Mol. Evol. 42, 211–223. https://doi.org/10.1007/BF02198847 (1996).
    ADS  CAS  Article  PubMed  Google Scholar 

    85.
    Marie, D., Partensky, F., Jacquet, S. & Vaulot, D. Enumeration and cell cycle analysis of natural populations of marine picoplankton by flow cytometry using the nucleic acid stain SYBR Green I. Appl. Environ. Microbiol. 63, 186–193. https://doi.org/10.1128/aem.63.1.186-193.1997 (1997).
    CAS  Article  PubMed  PubMed Central  Google Scholar 

    86.
    Wood, A. M., Everroad, R. C. & Wingard, L. M. Measuring growth rates in microalgal cultures. In Algal Culturing Techniques (ed. Anderson, R. A.) 269–285 (Elsevier, Amsterdam, 2005).
    Google Scholar 

    87.
    Kromkamp, J. C. & Forster, R. M. The use of variable fluorescence measurements in aquatic ecosystems: Differences between multiple and single turnover measuring protocols and suggested terminology. Eur. J. Phycol. 38, 103–112. https://doi.org/10.1080/0967026031000094094 (2003).
    Article  Google Scholar 

    88.
    Aminot, A. & Kérouel, R. Dosage Automatique des Nutriments dans les Eaux Marines: Méthodes en flux Continu (in French) (Ed. Ifremer, Plouzané, 2007).
    Google Scholar 

    89.
    Smith, P. K. et al. Measurement of protein using bicinchoninic acid. Anal. Biochem. 150, 76–85. https://doi.org/10.1016/0003-2697(85)90442-7 (1985).
    CAS  Article  PubMed  Google Scholar 

    90.
    Kamerling, J. P., Gerwig, G. J., Vliegenthart, J. F. G. & Clamp, J. R. Characterization by gas-liquid chromatography mass spectrometry of pertrimethylsilyl methyl glycosides obtained in the methanolysis of glycoproteins and glycolipids. Biochem. J. 151, 491–495. https://doi.org/10.1042/bj1510491 (1975).
    CAS  Article  PubMed  PubMed Central  Google Scholar 

    91.
    Laemmli, U. K. Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 22, 680–685. https://doi.org/10.1038/227680a0 (1970).
    ADS  Article  Google Scholar 

    92.
    Rigouin, C., Delbarre Ladrat, C., Sinquin, C., Colliec-Jouault, S. & Dion, M. Assessment of biochemical methods to detect enzymatic depolymerization of polysaccharides. Carbohydr. Polym. 76, 279–284. https://doi.org/10.1016/j.carbpol.2008.10.022 (2009).
    CAS  Article  Google Scholar 

    93.
    Dubray, G. & Bezard, G. A highly sensitive periodic acid-silver stain for 1,2-diol groups of glycoproteins and polysaccharides in polyacrylamide gels. Anal. Biochem. 119, 325–329. https://doi.org/10.1016/0003-2697(82)90593-0 (1982).
    CAS  Article  PubMed  Google Scholar 

    94.
    Aminot, A. & Kérouel, R. Hydrologie des Écosystèmes Marins: Paramètres et Analyses (in French) (Ed Ifremer, Plouzané, 2004).
    Google Scholar 

    95.
    R Core Team R: A Language and Environment for Statistical Computing. R Foundation for Statistical Computing, Vienna. https://www.R-project.org (2018).

    96.
    Lê, S., Josse, J. & Husson, F. FactoMineR: An R package for multivariate analysis. J. Stat. Softw. 25, 1–18 (2008).
    Article  Google Scholar  More

  • in

    A DNA barcode-based survey of wild urban bees in the Loire Valley, France

    1.
    Hallmann, C. A. et al. More than 75 percent decline over 27 years in total flying insect biomass in protected areas. PLoS ONE 12, e0185809 (2017).
    PubMed  PubMed Central  Article  CAS  Google Scholar 
    2.
    Macgregor, C. J., Williams, J. H., Bell, J. R. & Thomas, C. D. Moth biomass increases and decreases over 50 years in Britain. Nat. Ecol. Evol. 3, 1645–1649 (2019).
    PubMed  Article  Google Scholar 

    3.
    Sánchez-Bayo, F. & Wyckhuys, K. A. G. Worldwide decline of the entomofauna: A review of its drivers. Biol. Conserv. 232, 8–27 (2019).
    Article  Google Scholar 

    4.
    Thomas, C. D., Jones, T. H. & Hartley, S. E. “Insectageddon”: A call for more robust data and rigorous analyses. Glob. Chang. Biol. 25, 1891–1892 (2019).
    ADS  PubMed  Article  Google Scholar 

    5.
    van Klink, R. et al. Meta-analysis reveals declines in terrestrial but increases in freshwater insect abundances. Science (80-) 368, 417–420 (2020).
    ADS  Article  CAS  Google Scholar 

    6.
    Potts, S. G. et al. Global pollinator declines: Trends, impacts and drivers. Trends Ecol. Evol. 25, 345–353 (2010).
    PubMed  Article  Google Scholar 

    7.
    Pérez-Méndez, N. et al. The economic cost of losing native pollinator species for orchard production. J. Appl. Ecol. 57, 599–608 (2020).
    Article  Google Scholar 

    8.
    Porto, R. G. et al. Pollination ecosystem services: A comprehensive review of economic values, research funding and policy actions. Food Secur. 12, 1425–1442 (2020).
    Article  Google Scholar 

    9.
    Winfree, R., Aguilar, R., Vázquez, D. P., LeBuhn, G. & Aizen, M. A. A meta-analysis of bees’ responses to anthropogenic disturbance. Ecology 90, 2068–2076 (2009).
    PubMed  PubMed Central  Article  Google Scholar 

    10.
    Godfray, H. C. J. et al. A restatement of the natural science evidence base concerning neonicotinoid insecticides and insect pollinators. Proc. R. Soc. B Biol. Sci. 281, 20140558 (2014).
    Article  Google Scholar 

    11.
    Fortel, L. et al. Decreasing abundance, increasing diversity and changing structure of the Wild Bee Community (Hymenoptera: Anthophila) along an urbanization gradient. PLoS ONE 9, e104679 (2014).
    ADS  PubMed  PubMed Central  Article  CAS  Google Scholar 

    12.
    Geslin, B. et al. The proportion of impervious surfaces at the landscape scale structures wild bee assemblages in a densely populated region. Ecol. Evol. 6, 6599–6615 (2016).
    PubMed  PubMed Central  Article  Google Scholar 

    13.
    Geslin, B., Le Féon, V., Kuhlmann, M., Vaissière, B. E. & Dajoz, I. The bee fauna of large parks in downtown Paris, France. Ann. la Société Entomol. Fr. 51, 487–493 (2015).
    Article  Google Scholar 

    14.
    Baldock, K. C. R. et al. A systems approach reveals urban pollinator hotspots and conservation opportunities. Nat. Ecol. Evol. 3, 363–373 (2019).
    PubMed  PubMed Central  Article  Google Scholar 

    15.
    Lerman, S. B., Contosta, A. R., Milam, J. & Bang, C. To mow or to mow less: Lawn mowing frequency affects bee abundance and diversity in suburban yards. Biol. Conserv. 221, 160–174 (2018).
    Article  Google Scholar 

    16.
    Kerr, J. T. et al. Climate change impacts on bumblebees converge across continents. Science 349, 177–180 (2015).
    ADS  CAS  PubMed  PubMed Central  Article  Google Scholar 

    17.
    Soroye, P., Newbold, T. & Kerr, J. Climate change contributes to widespread declines among bumble bees across continents. Science 367, 685–688 (2020).
    ADS  CAS  PubMed  Article  Google Scholar 

    18.
    McFrederick, Q. S. & LeBuhn, G. Are urban parks refuges for bumble bees Bombus spp. (Hymenoptera: Apidae)?. Biol. Conserv. 129, 372–382 (2006).
    Article  Google Scholar 

    19.
    Hall, D. M. et al. The city as a refuge for insect pollinators. Conserv. Biol. 31, 24–29 (2017).
    PubMed  Article  Google Scholar 

    20.
    Ropars, L., Dajoz, I. & Geslin, B. La ville un désert pour les abeilles sauvages? J. Bot. Soc. Bot. Fr. 79, 29–35 (2017).
    Google Scholar 

    21.
    Falk, S. et al. Evaluating the ability of citizen scientists to identify bumblebee (Bombus) species. PLoS ONE 14, e0218614 (2019).
    CAS  PubMed  PubMed Central  Article  Google Scholar 

    22.
    Bloom, E. H. & Crowder, D. W. Promoting data collection in pollinator citizen science projects. Citiz. Sci. Theory Pract. 5(1), 3 https://doi.org/10.5334/cstp.217 (2020).

    23.
    Levé, M., Baudry, E. & Bessa-Gomes, C. Domestic gardens as favorable pollinator habitats in impervious landscapes. Sci. Total Environ. 647, 420–430 (2019).
    ADS  PubMed  Article  CAS  Google Scholar 

    24.
    Mason, L. & Arathi, H. S. Assessing the efficacy of citizen scientists monitoring native bees in urban areas. Glob. Ecol. Conserv. 17, e00561 (2019).
    Article  Google Scholar 

    25.
    Sheffield, C. S. et al. Contribution of DNA barcoding to the study of the bees (Hymenoptera: Apoidea) of Canada: Progress to date. Can. Entomol. 149, 736–754 (2017).
    Article  Google Scholar 

    26.
    Sheffield, C. S., Hebert, P. D. N., Kevan, P. G. & Packer, L. DNA barcoding a regional bee (Hymenoptera: Apoidea) fauna and its potential for ecological studies. Mol. Ecol. Resour. 9, 196–207 (2009).
    CAS  PubMed  Article  Google Scholar 

    27.
    Schmidt, S., Schmid-Egger, C., Morinière, J., Haszprunar, G. & Hebert, P. D. N. DNA barcoding largely supports 250 years of classical taxonomy: Identifications for Central European bees (Hymenoptera, Apoidea partim ). Mol. Ecol. Resour. 15, 985–1000 (2015).
    CAS  PubMed  Article  Google Scholar 

    28.
    Packer, L. & Ruz, L. DNA barcoding the bees (Hymenoptera: Apoidea) of Chile: Species discovery in a reasonably well known bee fauna with the description of a new species of Lonchopria (Colletidae). Genome 60, 414–430 (2017).
    CAS  PubMed  Article  Google Scholar 

    29.
    Tang, M. et al. High-throughput monitoring of wild bee diversity and abundance via mitogenomics. Methods Ecol. Evol. 6, 1034–1043 (2015).
    PubMed  PubMed Central  Article  Google Scholar 

    30.
    Sonet, G. et al. Using next-generation sequencing to improve DNA barcoding: Lessons from a small-scale study of wild bee species (Hymenoptera, Halictidae). Apidologie 49, 671–685 (2018).
    CAS  Article  Google Scholar 

    31.
    Creedy, T. J. et al. A validated workflow for rapid taxonomic assignment and monitoring of a national fauna of bees (Apiformes) using high throughput DNA barcoding. Mol. Ecol. Resour. 20, 40–53 (2020).
    CAS  PubMed  Article  Google Scholar 

    32.
    Gueuning, M. et al. Evaluating next-generation sequencing (NGS) methods for routine monitoring of wild bees: Metabarcoding, mitogenomics or NGS barcoding. Mol. Ecol. Resour. 19, 847–862 (2019).
    CAS  PubMed  PubMed Central  Article  Google Scholar 

    33.
    Lanner, J., Curto, M., Pachinger, B., Neumüller, U. & Meimberg, H. Illumina midi-barcodes: Quality proof and applications. Mitochondrial DNA Part A 30, 490–499 (2019).
    CAS  Article  Google Scholar 

    34.
    González-Vaquero, R. A., Roig-Alsina, A. & Packer, L. DNA barcoding as a useful tool in the systematic study of wild bees of the tribe Augochlorini (Hymenoptera: Halictidae). Genome 59, 889–898 (2016).
    PubMed  Article  CAS  PubMed Central  Google Scholar 

    35.
    Gibbs, J. DNA barcoding a nightmare taxon: Assessing barcode index numbers and barcode gaps for sweat bees. Genome 61, 21–31 (2018).
    CAS  PubMed  Article  PubMed Central  Google Scholar 

    36.
    Dorey, J. P., Schwarz, M. P. & Stevens, M. I. Review of the bee genus Homalictus Cockerell (Hymenoptera: Halictidae) from Fiji with description of nine new species. Zootaxa 4674, 1–46 (2019).
    Article  Google Scholar 

    37.
    Williams, P. H. et al. Unveiling cryptic species of the bumblebee subgenus Bombus s. str. worldwide with COI barcodes (Hymenoptera: Apidae). Syst. Biodivers. 10, 21–56 (2012).
    Article  Google Scholar 

    38.
    Magnacca, K. N. & Brown, M. J. F. DNA barcoding a regional fauna: Irish solitary bees. Mol. Ecol. Resour. 12, 990–998 (2012).
    CAS  PubMed  Article  Google Scholar 

    39.
    de Waard, J. R. et al. A reference library for Canadian invertebrates with 1.5 million barcodes, voucher specimens, and DNA samples. Sci. Data 6, 308 (2019).
    Article  CAS  Google Scholar 

    40.
    Hua, F. et al. Opportunities for biodiversity gains under the world’s largest reforestation programme. Nat. Commun. 7, 12717 (2016).
    ADS  CAS  PubMed  PubMed Central  Article  Google Scholar 

    41.
    Gueuning, M., Frey, J. E. & Praz, C. Ultraconserved yet informative for species delimitation: UCEs resolve long-standing systematic enigma in Central European bees. Mol. Ecol. Mec. https://doi.org/10.1111/mec.15629 (2020).
    Article  Google Scholar 

    42.
    Phillips, J. D., French, S. H., Hanner, R. H. & Gillis, D. J. HACSim: An R package to estimate intraspecific sample sizes for genetic diversity assessment using haplotype accumulation curves. PeerJ Comput. Sci. 6, e243 (2020).
    Article  Google Scholar 

    43.
    Phillips, J. D., Gwiazdowski, R. A., Ashlock, D. & Hanner, R. An exploration of sufficient sampling effort to describe intraspecific DNA barcode haplotype diversity: Examples from the ray-finned fishes (Chordata: Actinopterygii). DNA Barcodes 3(1), 66–73 (2015).

    44.
    Phillips, J. D., Gillis, D. J. & Hanner, R. H. Incomplete estimates of genetic diversity within species: Implications for DNA barcoding. Ecol. Evol. 9, 2996–3010 (2019).
    PubMed  PubMed Central  Article  Google Scholar 

    45.
    Muséum national d’Histoire naturelle (ed). 2003-2020. Inventaire National du Patrimoine Naturel. https://inpn.mnhn.fr.

    46.
    Zayed, A., Constantin, ŞA. & Packer, L. Successful biological invasion despite a severe genetic load. PLoS ONE 2, e868 (2007).
    ADS  PubMed  PubMed Central  Article  CAS  Google Scholar 

    47.
    Lecocq, T. et al. The alien’s identity: Consequences of taxonomic status for the international bumblebee trade regulations. Biol. Conserv. 195, 169–176 (2016).
    Article  Google Scholar 

    48.
    Danforth, B. N. Phylogeny of the bee genus Lasioglossum (Hymenoptera: Halictidae) based on mitochondrial COI sequence data. Syst. Entomol. 24, 377–393 (1999).
    Article  Google Scholar 

    49.
    Hebert, P. D. N. et al. A Sequel to Sanger: Amplicon sequencing that scales. BMC Genom. 19, 219 (2018).
    Article  CAS  Google Scholar 

    50.
    Ratnasingham, S. & Hebert, P. D. N. A DNA-based registry for all animal species: The Barcode Index Number (BIN) system. PLoS ONE 8, e66213 (2013).
    ADS  CAS  PubMed  PubMed Central  Article  Google Scholar 

    51.
    Carolan, J. C. et al. Colour patterns do not diagnose species: Quantitative evaluation of a DNA barcoded cryptic bumblebee complex. PLoS ONE 7, e29251 (2012).
    ADS  CAS  PubMed  PubMed Central  Article  Google Scholar 

    52.
    Praz, C., Müller, A. & Genoud, D. Hidden diversity in European bees: Andrena amieti sp. n., a new Alpine bee species related to Andrena bicolor (Fabricius, 1775) (Hymenoptera, Apoidea, Andrenidae). Alp. Entomol. 3, 11–38 (2019).
    Article  Google Scholar 

    53.
    Pauly, A. Abeilles de Belgique et des régions limitrophes (Insecta: Hymenoptera: Apoidea) Famille Halictidae. (Institut royal des sciences naturelles de Belgique, 2019).

    54.
    Gonçalves, R. B. & Oliveira, P. S. Preliminary results of bowl trapping bees (Hymenoptera, Apoidea) in a southern Brazil forest fragment. J. Insect Biodivers. 1, 1–9 (2013).
    Article  Google Scholar 

    55.
    Buri, P., Humbert, J.-Y. & Arlettaz, R. Promoting pollinating insects in intensive agricultural matrices: Field-scale experimental manipulation of hay-meadow mowing regimes and its effects on bees. PLoS One 9(1), e85635 (2014).

    56.
    Rhoades, P. et al. Sampling technique affects detection of habitat factors influencing wild bee communities. J. Insect Conserv. 21, 703–714 (2017).
    Article  Google Scholar 

    57.
    Lettow, M. C. et al. Bee community responses to a gradient of oak savanna restoration practices. Restor. Ecol. 26, 882–890 (2018).
    Article  Google Scholar 

    58.
    Onuferko, T. M., Skandalis, D. A., Cordero, R. L. & Richards, M. H. Rapid initial recovery and long-term persistence of a bee community in a former landfill. Insect Conserv. Divers. 11, 88–99 (2018).
    Article  Google Scholar 

    59.
    Geroff, R. K., Gibbs, J. & McCravy, K. W. Assessing bee (Hymenoptera: Apoidea) diversity of an Illinois restored tallgrass prairie: Methodology and conservation considerations. J. Insect Conserv. 18, 951–964 (2014).
    Article  Google Scholar 

    60.
    Griffin, S. R., Bruninga-Socolar, B., Kerr, M. A., Gibbs, J. & Winfree, R. Wild bee community change over a 26-year chronosequence of restored tallgrass prairie. Restor. Ecol. 25, 650–660 (2017).
    Article  Google Scholar 

    61.
    Ropars, L., Dajoz, I. & Geslin, B. La diversité des abeilles parisiennes. Osmia 7, 14–19 (2018).
    Article  Google Scholar 

    62.
    Portman, Z. M., Bruninga-Socolar, B. & Cariveau, D. P. The state of bee monitoring in the United States: A call to refocus away from bowl traps and towards more effective methods. Ann. Entomol. Soc. Am. 113, 337–342 (2020).
    Article  Google Scholar 

    63.
    Magnacca, K. N. & Brown, M. J. Mitochondrial heteroplasmy and DNA barcoding in Hawaiian Hylaeus (Nesoprosopis) bees (Hymenoptera: Colletidae). BMC Evol. Biol. 10, 174 (2010).

    64.
    Ballare, K. M. et al. Utilizing field collected insects for next generation sequencing: Effects of sampling, storage, and DNA extraction methods. Ecol. Evol. 9, 13690–13705 (2019).
    PubMed  PubMed Central  Article  Google Scholar 

    65.
    Hill, G. E. Mitonuclear coevolution as the genesis of speciation and the mitochondrial DNA barcode gap. Ecol. Evol. 6, 5831–5842 (2016).
    PubMed  PubMed Central  Article  Google Scholar 

    66.
    Ascher, J. S. & Pickering, J.  Life bee species guide and world checklist (Hymenoptera Apoidea Anthophila). http://www.discoverlife.org/mp/20q?guide=Apoidea_species (2020).

    67.
    LaBerge, W. E. A revision of the bees of the genus Andrena of the western hemisphere. Part XI. Minor subgenera and subgeneric key. Trans. Am. Entomol. Soc. 111, 441–567 (1985).
    Google Scholar 

    68.
    Warncke, K. Die Untergattungen der westpalaarktischen Bienengattung Andrena F. Memorias e Estud Muséu Zool. da Univ. Coimbra 307, 1–110 (1968).
    Google Scholar 

    69.
    Amiet, F., Herrmann, M., Müller, A. & Neumeyer, R. Apidae 6: Andrena, Melitturga, Panurginus, Panurgus. Fauna Helv. 26, 1–317 (2010).

    70.
    Michener, C. The bees of the world. (Johns Hopkins University Press, Baltimore, 2000).
    Google Scholar 

    71.
    Michener, C. D. The Social Behavior of the Bees: A Comparative Study (Harvard University Press, Cambridge, 1974).
    Google Scholar 

    72.
    Pauly, A., Noël, G., Sonet, G., Notton, D. G. & Boevé, J.-L. Integrative taxonomy resuscitates two species in the Lasioglossum villosulum complex (Kirby, 1802) (Hymenoptera: Apoidea: Halictidae). Eur. J. Taxon. 541 (2019).

    73.
    Eberle, J., Ahrens, D., Mayer, C., Niehuis, O. & Misof, B. A plea for standardized nuclear markers in metazoan DNA taxonomy. Trends Ecol. Evol. 35, 336–345 (2020).
    PubMed  Article  Google Scholar 

    74.
    Roulston, T. H., Smith, S. A. & Brewster, A. L. A comparison of pan trap and intensive net sampling techniques for documenting a bee (Hymenoptera: Apiformes) Fauna. J. Kansas Entomol. Soc. 80, 179–181 (2007).
    Article  Google Scholar 

    75.
    Westphal, C. et al. Measuring bee diversity in different European habitats and biogeographical regions. Ecol. Monogr. 78, 653–671 (2008).
    Article  Google Scholar 

    76.
    Amiet, F., Herrmann, M., Müller, A. & Neumeyer, R. Apidae 5: Ammobates, Ammobatoides, Anthophora, Biastes, Ceratina, Dasypoda, Epeoloides, Epeolus, Eucera, Macropis, Melecta, Melitta, Nomada, Pasites, Tetralonia, Thyreus, Xylocopa. Fauna Helv. 20, 1–356 (2007).

    77.
    Amiet, F., Herrmann, M., Müller, A. & Neumeyer, R. Apidae 2: Colletes, Dufourea, Hylaeus, Nomia, Nomioides, Rhophitoides, Rophites, Sphecodes, Systropha. Fauna Helv. 4, 1–239 (1999).

    78.
    Amiet, F., Herrmann, M., Müller, A. & Neumeyer, R. Apidae 3: Halictus, Lasioglossum. Fauna Helv. 6, 1–208 (2001).

    79.
    Amiet, F., Herrmann, M., Müller, A. & Neumeyer, R. Apidae 4: Anthidium, Chelostoma, Coelioxys, Dioxys, Heriades, Lithurgus, Megachile, Osmia, Stelis. Fauna Helv. 9, 1–273 (2004).

    80.
    Folmer, O., Black, M., Hoeh, W., Lutz, R. & Vrijenhoek, R. DNA primers for amplification of mitochondrial cytochrome c oxidase subunit I from diverse metazoan invertebrates. Mol. Mar. Biol. Biotechnol. 3, 294–299 (1994).
    CAS  PubMed  Google Scholar 

    81.
    Ratnasingham, S. & Hebert, P. D. N. BOLD: The barcode of life data system. Mol. Ecol. Notes 7, 355–364 (2007).

    82.
    Katoh, K. MAFFT: A novel method for rapid multiple sequence alignment based on fast Fourier transform. Nucleic Acids Res. 30, 3059–3066 (2002).
    ADS  CAS  PubMed  PubMed Central  Article  Google Scholar 

    83.
    Kimura, M. A simple method for estimating evolutionary rates of base substitutions through comparative studies of nucleotide sequences. J. Mol. Evol. 16, 111–120 (1980).
    ADS  CAS  PubMed  Article  Google Scholar 

    84.
    Price, M. N., Dehal, P. S. & Arkin, A. P. FastTree 2—Approximately maximum-likelihood trees for large alignments. PLoS ONE 5, e9490 (2010).
    ADS  PubMed  PubMed Central  Article  CAS  Google Scholar 

    85.
    Letunic, I. & Bork, P. Interactive Tree Of Life (iTOL) v4: Recent updates and new developments. Nucleic Acids Res. 47, W256–W259 (2019).
    CAS  PubMed  PubMed Central  Article  Google Scholar 

    86.
    Wickham, H. ggplot2 (Springer, New York, 2009). https://doi.org/10.1007/978-0-387-98141-3.
    Google Scholar 

    87.
    Rozas, J. et al. DnaSP 6: DNA sequence polymorphism analysis of large data sets. Mol. Biol. Evol. 34, 3299–3302 (2017).
    CAS  PubMed  Article  Google Scholar 

    88.
    Nei, M. Molecular evolutionary genetics (Columbia University Press, New York, 1987). More

  • in

    Native and invasive ants affect floral visits of pollinating honey bees in pumpkin flowers (Cucurbita maxima)

    1.
    Vitousek, P. M., D’Antonio, C. M., Loope, L. L., Rejmánek, M. & Westbrooks, R. Introduced species: a significant component of human-caused global change. N. Z. J. Ecol. 21, 1–16 (1997).
    Google Scholar 
    2.
    Courchamp, F. et al. Invasion biology: specific problems and possible solutions. Trends Ecol. Evol. 32, 13–22 (2017).
    PubMed  Article  PubMed Central  Google Scholar 

    3.
    Sanders, N. J., Gotelli, N. J., Heller, N. E. & Gordon, D. M. Community disassembly by an invasive species. PNAS 100, 2474–2477 (2003).
    ADS  CAS  PubMed  Article  PubMed Central  Google Scholar 

    4.
    Christian, C. E. Consequences of a biological invasion reveal the importance of mutualism for plant communities. Nature 413, 635–639 (2001).
    ADS  CAS  PubMed  Article  PubMed Central  Google Scholar 

    5.
    Suarez, A. V., McGlynn, T. P. & Tsutsui, N. D. Biogeographic and taxonomic patterns of introduced ants. In Ant ecology (eds Lach, L. et al.) 233–244 (Oxford University Press, Oxford, 2010).
    Google Scholar 

    6.
    Moloney, S. D. & Vanderwoude, C. Potential ecological impacts of red imported fire ants in eastern Australia. J. Agric. Urban Entomol. 20, 131–142 (2003).
    Google Scholar 

    7.
    Rajesh, T. P., Ballullaya, U. P., Unni, A. P., Parvathy, S. & Sinu, P. A. Interactive effects of urbanization and year on invasive and native ant diversity of sacred groves of South India. Urban Ecosyst. 23, 1335–1348 (2020).
    Article  Google Scholar 

    8.
    Hoffmann, B. D., Luque, G. M., Bellard, C., Holmes, N. D. & Donlan, C. J. Improving invasive ant eradication as a conservation tool: a review. Biol. Conserv. 198, 37–49 (2016).
    Article  Google Scholar 

    9.
    Traveset, A. & Richardson, D. M. Biological invasions as disruptors of plant reproductive mutualisms. Trends Ecol. Evol. 21, 208–216 (2006).
    PubMed  Article  PubMed Central  Google Scholar 

    10.
    Carney, S. E., Byerley, M. B. & Holway, D. A. Invasive Argentine ants (Linepithema humile) do not replace native ants as seed dispersers of Dendromecon rigida (Papaveraceae) in California, USA. Oecologia 135, 576–582 (2003).
    ADS  PubMed  Article  PubMed Central  Google Scholar 

    11.
    Styrsky, J. D. & Eubanks, M. D. Ecological consequences of interactions between ants and honeydew-producing insects. Proc. R. Soc. B Biol. Sci. 274, 151–164 (2007).
    Article  Google Scholar 

    12.
    Lach, L. Invasive ants: unwanted partners in ant-plant interactions?. Ann. Mo. Bot. Garden 90, 91–108 (2003).
    Article  Google Scholar 

    13.
    Willmer, P. G. et al. Floral volatiles controlling ant behaviour. Funct. Ecol. 23, 888–900 (2009).
    Article  Google Scholar 

    14.
    Vilamil, N., Boege, K. & Stone, G. N. Testing the distraction hypothesis: do extrafloral nectaries reduce ant-pollinator conflict?. J. Ecol. 107, 1377–1391 (2019).
    Article  Google Scholar 

    15.
    Raine, N., Willmer, P. & Stone, G. Spatial structuring and floral avoidance behavior prevent ant-pollinator conflict in a Mexican antacacia. Ecology 83, 3086–3096 (2002).
    Google Scholar 

    16.
    Weber, M. G., Porturas, L. D. & Keeler, K. H. World list of plants with extrafloral nectaries. www.extrafloralnectaries.org (2015)

    17.
    Dutton, E. M. & Frederickson, M. E. Why ant pollination is rare: new evidence and implications of the antibiotic hypothesis. Arthropod-Plant Interact. 6, 561–569 (2012).
    Article  Google Scholar 

    18.
    Gonzálvez, F. G., Santamaría, L., Corlett, R. T. & Rodríguez-Gironés, M. A. Flowers attract weaver ants that deter less effective pollinators. J. Ecol. 101, 78–85 (2013).
    Article  Google Scholar 

    19.
    Cembrowski, A. R., Tan, M. G., Thomson, J. D. & Frederickson, M. E. Ants and ant scent reduce bumblebee pollination of artificial flowers. Am. Nat. 183, 133–139 (2014).
    PubMed  Article  PubMed Central  Google Scholar 

    20.
    Sinu, P. A. et al. Invasive ant (Anoplolepis gracilipes) disrupts pollination in pumpkin. Biol. Invasions 19, 2599–2607 (2017).
    Article  Google Scholar 

    21.
    Hanna, C. et al. Floral visitation by the Argentine ant reduces bee visitation and plant seed set. Ecology 96, 222–230 (2015).
    PubMed  Article  PubMed Central  Google Scholar 

    22.
    LeVan, K. E., Hung, K.-L.-J., McCann, K. R., Ludka, J. T. & Holway, D. A. Floral visitation by the Argentine ant reduces pollinator visitation and seed set in the coast barrel cactus, Ferocactus viridescens. Oecologia 174, 163–171 (2014).
    ADS  PubMed  Article  PubMed Central  Google Scholar 

    23.
    Fuster, F., Kaiser-Bunbury, C. N. & Traveset, A. Pollination effectiveness of specialist and opportunistic nectar feeders influenced by invasive alien ants in the Seychelles. Am. J. Bot. 107, 957–969 (2020).
    PubMed  Article  PubMed Central  Google Scholar 

    24.
    Bissessur, P., Baider, C. & Florens, F. B. V. Infestation by pollination-disrupting alien ants varies temporally and spatially and is worsened by alien plant invasion. Biol. Invasions 22, 2573–2585 (2020).
    Article  Google Scholar 

    25.
    Del-Claro, K., Rodriguez-Morales, D., Calixto, E. S., Martins, A. S. & Torezan-Silingardi, H. M. Ant pollination of Paepalanthus lundii (Eriocaulaceae) in Brazilian savanna. Ann. Bot. 123, 1159–1165 (2019).
    CAS  PubMed  PubMed Central  Article  Google Scholar 

    26.
    Kuriakose, G., Sinu, P. A. & Shivanna, K. R. Ant pollination of Syzygium occidentale, an endemic tree species of tropical rain forests of the Western Ghats, India. Arthropod-Plant Interact. 12, 647–655 (2018).
    Article  Google Scholar 

    27.
    Galen, C. & Cuba, J. Down the tube: pollinators, predators, and the evolution of flower shape in the alpine skypilot Polemonium viscosum. Evolution 55, 1963–1971 (2001).
    CAS  PubMed  Article  PubMed Central  Google Scholar 

    28.
    Tsuji, K., Hasyim Harlion, A. & Nakamura, K. Asian weaver ants, Oecophylla smaragdina, and their repelling of pollinators. Ecol. Res. 19, 669–673 (2004).
    Article  Google Scholar 

    29.
    Ness, J. H. A mutualism’s indirect costs: the most aggressive plant bodyguards also deter pollinators. Oikos 113, 506–514 (2006).
    Article  Google Scholar 

    30.
    Lach, L. Argentine ants displace floral arthropods in a biodiversity hotspot. Divers. Distrib. 14, 281–290 (2008).
    Article  Google Scholar 

    31.
    Hansen, D. M. & Müller, C. B. Invasive ants disrupt gecko pollination and seed dispersal of the endangered plant Roussea simplex in Mauritius. Biotropica 41, 202–208 (2009).
    Article  Google Scholar 

    32.
    Lach, L. Interference and exploitation competition of three nectar-thieving invasive ant species. Insectes Soc. 52, 257–262 (2005).
    Article  Google Scholar 

    33.
    Blancafort, X. & Gómez, C. Consequences of the Argentine ant, Linepithema humile (Mayr), invasion on pollination of Euphorbia characias (L.) (Euphorbiaceae). Acta Oecol. 28, 49–55 (2005).
    ADS  Article  Google Scholar 

    34.
    Holway, D. A. Competitive mechanisms underlying the displacement of native ants by the invasive Argentine ant. Ecology 80, 238–251 (1999).
    Article  Google Scholar 

    35.
    Holway, D. A., Lach, L., Suarez, A. V., Tsutsui, N. D. & Case, T. J. The causes and consequences of ant invasions. Annu. Rev. Ecol. Syst. 33, 181–233 (2002).
    Article  Google Scholar 

    36.
    Silverman, J. & Buczkowski, G. 13 Behaviours mediating ant invasions. in Biological Invasions and Animal Behaviour 221 (2016).

    37.
    Sinu, P. A. et al. Effect of flower sex ratio on fruit set in pumpkin (Cucurbita maxima). Sci. Hortic. 246, 1005–1008 (2019).
    Article  Google Scholar 

    38.
    Sinu, P. A., Pooja, A. R. & Aneha, K. Overhead sprinkler irrigation affects pollinators and pollination in pumpkin (Cucurbita maxima). Sci. Hortic. 258, 108803 (2019).
    Article  Google Scholar 

    39.
    Bharti, H., Guénard, B., Bharti, M. & Economo, E. P. An updated checklist of the ants of India with their specific distributions in Indian states (Hymenoptera, Formicidae). ZooKeys 551, 1–83 (2016).
    Article  Google Scholar 

    40.
    R Core Team. R: A Language and Environment for Statistical Computing (R Foundation for Statistical Computing, Vienna, 2018).
    Google Scholar 

    41.
    Anusree, T. et al. Flower sex expression in cucurbit crops of Kerala: implications for pollination and fruitset. Curr. Sci. 109, 2299–2302 (2015).
    Article  Google Scholar 

    42.
    das Vidal, M. G., de Jong, D., Wien, H. C. & Morse, R. A. Produção de néctar e pólen em abóbora (Cucurbita pepo L). Braz. J. Bot. 29, 267–273 (2006).
    Article  Google Scholar 

    43.
    Junker, R., Chung, A. Y. & Blüthgen, N. Interaction between flowers, ants and pollinators: additional evidence for floral repellence against ants. Ecol. Res. 22, 665–670 (2007).
    Article  Google Scholar 

    44.
    Ibarra-Isassi, J. & Sendoya, S. F. Ants as floral visitors of Blutaparon portulacoides (A. St-Hil.) Mears (Amaranthaceae): an ant pollination system in the Atlantic Rainforest. Arthropod-Plant Interact. 10, 221–227 (2016).
    Article  Google Scholar 

    45.
    Beattie, A. J., Turnbull, C., Knox, R. B. & Williams, E. G. Ant inhibition of pollen function: a possible reason why ant pollination is rare. Am. J. Bot. 71, 421–426 (1984).
    Article  Google Scholar 

    46.
    Hickman, J. C. Pollination by ants: a low-energy system. Science 184, 1290–1292 (1974).
    ADS  CAS  PubMed  Article  PubMed Central  Google Scholar 

    47.
    Galen, C. The effects of nectar thieving ants on seedset in floral scent morphs of Polemonium viscosum. Oikos 41, 245–249 (1983).
    Article  Google Scholar 

    48.
    Witte, V., Attygalle, A. B. & Meinwald, J. Complex chemical communication in the crazy ant Paratrechina longicornis Latreille (Hymenoptera: Formicidae). Chemoecology 17, 57–62 (2007).
    Article  Google Scholar 

    49.
    Wetterer, J. Worldwide spread of the ghost ant, Tapinoma melanocephalum (Hymenoptera: Formicidae). Myrmecol. News 12, 23–33 (2009).
    Google Scholar 

    50.
    Todd, B. D. et al. Habitat alteration increases invasive fire ant abundance to the detriment of amphibians and reptiles. Biol. Invasions 10, 539–546 (2008).
    Article  Google Scholar  More

  • in

    Mayetiola destructor (Diptera: Cecidmyiidae) host preference and survival on small grains with respect to leaf reflectance and phytohormone concentrations

    1.
    Wiseman, B. R. Plant-resistance to insects in integrated pest-management. Plant Dis. 78, 927–932. https://doi.org/10.1094/pd-78-0927 (1994).
    Article  Google Scholar 
    2.
    Painter, R. H. Insect resistance in crop plants. Soil Sci. 72 (1951).

    3.
    Orr, D. B. & Boethel, D. J. Influence of plant antibiosis through four trophic levels. Oecologia 70, 242–249. https://doi.org/10.1007/BF00379247 (1986).
    ADS  CAS  Article  PubMed  Google Scholar 

    4.
    Smith, C. M. & Clement, S. L. Molecular Bases of Plant Resistance to Arthropods. Annu. Rev. Entomol. 57, 309–328. https://doi.org/10.1146/annurev-ento-120710-100642 (2011).
    CAS  Article  PubMed  Google Scholar 

    5.
    Radcliffe, R. H. in Radcliffe’s IPM world textbook Vol. https://ipmworld.umn.edu/ratcliffe-hessian-fly (eds Radcliffe E.B. & Hutchison W.D.) (University of Minnesota, 1997).

    6.
    Kosma, D. K., Nemacheck, J. A., Jenks, M. A. & Williams, C. E. Changes in properties of wheat leaf cuticle during interactions with Hessian fly. Plant J 63, 31–43 (2010).
    CAS  PubMed  Google Scholar 

    7.
    Smiley, R. W., Gourlie, J. A., Whittaker, R. G., Easley, S. A. & Kidwell, K. K. Economic impact of Hessian fly (Diptera: Cecidomyiidae) on spring wheat in Oregon and additive yield losses with Fusarium crown rot and lesion nematode. J Econ Entomol 97, 397–408 (2004).
    Article  Google Scholar 

    8.
    Harris, M. O., Sandanayaka, M. & Griffin, A. Oviposition preferences of the Hessian fly and their consequences for the survival and reproductive potential of offspring. Ecol. Entomol. 26, 473–486. https://doi.org/10.1046/j.1365-2311.2001.00344.x (2001).
    Article  Google Scholar 

    9.
    Ganehiarachchi, G. A. S. M., Anderson, K. M., Harmon, J. & Harris, M. O. Why oviposit there? Fitness consequences of a gall midge choosing the plant’s youngest leaf. Environ Entomol 42, 123–130 (2013).
    CAS  Article  Google Scholar 

    10.
    Kanno, H. & Harris, M. O. Physical features of grass leaves influence the placement of eggs within the plant by the Hessian fly. Entomol. Exp. Appl. 96, 69–80. https://doi.org/10.1046/j.1570-7458.2000.00680.x (2000).
    Article  Google Scholar 

    11.
    Harris, M. O. & Rose, S. Chemical, color, and tactile cues influencing oviposition behavior of the Hessian fly (Diptera, Cecidomyiidae). Environ. Entomol. 19, 303–308. https://doi.org/10.1093/ee/19.2.303 (1990).
    Article  Google Scholar 

    12.
    Kanno, H. & Harris, M. O. Leaf physical and chemical features influence selection of plant genotypes by hessian fly. J. Chem. Ecol. 26, 2335–2354 (2000).
    CAS  Article  Google Scholar 

    13.
    Cervantes, D. E., Eigenbrode, S. D., Ding, H. J. & Bosque-Perez, N. A. Oviposition responses by Hessian fly, Mayetiola destructor, to wheats varying in surface waxes. J. Chem. Ecol. 28, 193–210 (2002).
    CAS  Article  Google Scholar 

    14.
    Morris, B. D., Foster, S. P. & Harris, M. O. Identification of 1-octacosanal and 6-methoxy-2-benzoxazolinone from wheat as ovipositional stimulants for Hessian fly, Mayetiola destructor. J. Chem. Ecol. 26, 859–873 (2000).
    CAS  Article  Google Scholar 

    15.
    Harris, M. O., Rose, S. & Malsch, P. The role of vision in the host plant-finding behavior of the Hessian fly. Physiol. Entomol. 18, 31–42. https://doi.org/10.1111/j.1365-3032.1993.tb00446.x (1993).
    Article  Google Scholar 

    16.
    Rohfritsch, O. A fungus associated gall midge, Lasioptera arundinis (Schiner), on Phragmites australis (Cav) Trin. Bull. Soc. Bot. France Lett. Bot. 139, 45–59. https://doi.org/10.1080/01811797.1992.10824942 (1992).
    Article  Google Scholar 

    17.
    Schmid, R. B., Knutson, A., Giles, K. L. & McCornack, B. P. Hessian fly (Diptera: Cecidomyiidae) biology and management in wheat. J. Integr. Pest Manag. 9, 12. https://doi.org/10.1093/jipm/pmy008 (2018).
    Article  Google Scholar 

    18.
    Gagné, R. J. & Hatchett, J. H. Instars of the Hessian Fly (Diptera: Cecidomyiidae). Ann. Entomol. Soc. Am. 82, 73–79. https://doi.org/10.1093/aesa/82.1.73 (1989).
    Article  Google Scholar 

    19.
    Lidell, M. C. & Schuster, M. F. Distribution of the Hessian fly and its control in Texas. Southwestern Entomologist 15, 133–145 (1990).
    Google Scholar 

    20.
    Morgan, G., Sansone, C. & Knutson, A. Hessian fly in Texas wheat. E-350 (Texas A&M, 2005).

    21.
    Flanders, K. L., Reisig, D. D., Buntin, G. D., Herbert, J. D. A. & Johnson, D. W. Biology and management of Hessian fly in the Southeast. ANR1069 (Alabama Cooperative Extension System, 2013).

    22.
    Wellso, S. G. Aestivation and Phenology of the Hessian Fly (Diptera: Cecidomyiidae) in Indiana. Environ. Entomol. 20, 795–801. https://doi.org/10.1093/ee/20.3.795 (1991).
    Article  Google Scholar 

    23.
    Boyd, M. L. & Bailey, W. C. Hessian fly management on wheat. G7180 (Missouri Extension, University of Missouri-Columbia, 2000).

    24.
    Ando, K. et al. Genome-wide associations for multiple pest resistances in a Northwestern United States elite spring wheat panel. PLoS One 13, e0191305/0191301-e0191305/0191325. https://doi.org/10.1371/journal.pone.0191305 (2018).

    25.
    Anderson, K. M. & Harris, M. O. Susceptibility of North Dakota Hessian Fly (Diptera: Cecidomyiidae) to 31 H Genes Mediating Wheat Resistance. J. Econ. Entomol. 112, 2398–2406 (2019).
    Article  Google Scholar 

    26.
    Sardesai, N., Nemacheck, J. A., Subramanyam, S. & Williams, C. E. Identification and mapping of H32, a new wheat gene conferring resistance to Hessian fly. Theor. Appl. Genet. 111, 1167–1173 (2005).
    CAS  Article  Google Scholar 

    27.
    Zhu, L., Liu, X. & Chen, M.-S. Differential accumulation of phytohormones in wheat seedlings attacked by avirulent and virulent Hessian fly (Diptera: Cecidomyiidae) larvae. J. Econ. Entomol. 103, 178–185 (2010).
    CAS  Article  Google Scholar 

    28.
    Mithöfer, A. & Boland, W. Recognition of Herbivory-Associated Molecular Patterns. Plant Physiol. 146, 825. https://doi.org/10.1104/pp.107.113118 (2008).
    CAS  Article  PubMed  PubMed Central  Google Scholar 

    29.
    Stuart, J. J., Chen, M.-S., Shukle, R. & Harris, M. O. Gall midges (Hessian flies) as plant pathogens. Annu. Rev. Phytopathol. 50, 339–357 (2012).
    CAS  Article  Google Scholar 

    30.
    Liu, X. et al. Gene expression of different wheat genotypes during attack by virulent and avirulent Hessian fly (Mayetiola destructor) larvae. J. Chem. Ecol. 33, 2171–2194 (2007).
    CAS  Article  Google Scholar 

    31.
    Subramanyam, S. et al. Expression of two wheat defense-response genes, Hfr-1 and Wci-1, under biotic and abiotic stresses. Plant Sci. 170, 90–103. https://doi.org/10.1016/j.plantsci.2005.08.006 (2006).
    CAS  Article  Google Scholar 

    32.
    Wu, J. et al. Differential responses of wheat inhibitor-like genes to Hessian fly, Mayetiola destructor, attacks during compatible and incompatible interactions. J. Chem. Ecol. 34, 1005–1012 (2008).
    CAS  Article  Google Scholar 

    33.
    Giovanini, M. P. et al. A novel wheat gene encoding a putative chitin-binding lectin is associated with resistance against Hessian fly. Mol. Plant Pathol. 8, 69–82 (2007).
    CAS  Article  Google Scholar 

    34.
    Liu, X. et al. Reactive oxygen species are involved in plant defense against a gall midge. Plant Physiol. 152, 985. https://doi.org/10.1104/pp.109.150656 (2010).
    CAS  Article  PubMed  PubMed Central  Google Scholar 

    35.
    Bari, R. & Jones, J. D. G. Role of plant hormones in plant defence responses. Plant Mol. Biol. 69, 473–488. https://doi.org/10.1007/s11103-008-9435-0 (2009).
    CAS  Article  PubMed  Google Scholar 

    36.
    Denancé, N., Sánchez-Vallet, A., Goffner, D. & Molina, A. Disease resistance or growth: the role of plant hormones in balancing immune responses and fitness costs. Frontiers in Plant Science 4. https://doi.org/10.3389/fpls.2013.00155 (2013)

    37.
    Dinh, S. T., Baldwin, I. T. & Galis, I. The HERBIVORE ELICITOR-REGULATED1 gene enhances abscisic acid levels and defenses against herbivores in Nicotiana attenuate plants. Plant Physiol. 162, 2106–2124 (2013).
    CAS  Article  Google Scholar 

    38.
    War, A. R., Paulraj, M. G., War, M. Y. & Ignacimuthu, S. Role of salicylic acid in induction of plant defense system in chickpea (Cicer arietinum L.). Plant signaling & behavior 6, 1787–1792. https://doi.org/10.4161/psb.6.11.17685 (2011).

    39.
    Nguyen, D., Rieu, I., Mariani, C. & van Dam, N. M. How plants handle multiple stresses: hormonal interactions underlying responses to abiotic stress and insect herbivory. Plant Mol. Biol. 91, 727–740. https://doi.org/10.1007/s11103-016-0481-8 (2016).
    CAS  Article  PubMed  PubMed Central  Google Scholar 

    40.
    Lee, A. et al. Inverse correlation between jasmonic acid and salicylic acid during early wound response in rice. Biochem. Biophys. Res. Commun. 318, 734–738. https://doi.org/10.1016/j.bbrc.2004.04.095 (2004).
    CAS  Article  PubMed  Google Scholar 

    41.
    Kunkel, B. N. & Brooks, D. M. Cross talk between signaling pathways in pathogen defense. Curr. Opin. Plant Biol. 5, 325–331. https://doi.org/10.1016/S1369-5266(02)00275-3 (2002).
    CAS  Article  PubMed  Google Scholar 

    42.
    Farmer, E. E., Alméras, E. & Krishnamurthy, V. Jasmonates and related oxylipins in plant responses to pathogenesis and herbivory. Curr. Opin. Plant Biol. 6, 372–378. https://doi.org/10.1016/S1369-5266(03)00045-1 (2003).
    CAS  Article  PubMed  Google Scholar 

    43.
    Loake, G. & Grant, M. Salicylic acid in plant defence—the players and protagonists. Curr. Opin. Plant Biol. 10, 466–472. https://doi.org/10.1016/j.pbi.2007.08.008 (2007).
    CAS  Article  PubMed  Google Scholar 

    44.
    Felton, G. W., Bi, J. L., Summers, C. B., Mueller, A. J. & Duffey, S. S. Potential role of lipoxygenases in defense against insect herbivory. J. Chem. Ecol. 20, 651–666. https://doi.org/10.1007/BF02059605 (1994).
    CAS  Article  PubMed  Google Scholar 

    45.
    Audenaert, K., De Meyer, G. B. & Höfte, M. M. Abscisic Acid Determines Basal Susceptibility of Tomato to Botrytis cinerea and Suppresses Salicylic Acid-Dependent Signaling Mechanisms. Plant Physiol. 128, 491. https://doi.org/10.1104/pp.010605 (2002).
    CAS  Article  PubMed  PubMed Central  Google Scholar 

    46.
    Mohr, P. G. & Cahill, D. M. Suppression by ABA of salicylic acid and lignin accumulation and the expression of multiple genes, in Arabidopsis infected with Pseudomonas syringae pv. tomato. Functional & Integrative Genomics 7, 181–191, https://doi.org/10.1007/s10142-006-0041-4 (2007).

    47.
    Harris, M. O., Dando, J. L., Griffin, W. & Madie, C. Susceptibility of cereal and non-cereal grasses to attack by Hessian fly (Mayetiola destructor (Say)). N. Zeal. J. Crop Hortic. Scie.ce 24, 229–238. https://doi.org/10.1080/01140671.1996.9513957 (1996).
    Article  Google Scholar 

    48.
    Gitelson, A. A. & Merzlyak, M. N. Signature analysis of leaf reflectance spectra: algorithm development for remote sensing of chlorophyll. J. Plant Physiol. 148, 494–500. https://doi.org/10.1016/S0176-1617(96)80284-7 (1996).
    CAS  Article  Google Scholar 

    49.
    Foster, S. P. & Harris, M. O. Foliar chemicals of wheat and related grasses influencing oviposition by Hessian fly, Mayetiola destructor (Say) (Diptera: Cecidomyiidae). J Chem Ecol 18, 1965–1980 (1992).
    CAS  Article  Google Scholar 

    50.
    Gagne, R. J., Hatchett, J. H., Lhaloui, S. & El Bouhssini, M. Hessian fly and barley stem gall midge, two different species of mayetiola (Diptera: Cecidomyiidae) in Morocco. Ann. Entomol. Soc. Am. 84, 436–443. https://doi.org/10.1093/aesa/84.4.436 (1991).
    Article  Google Scholar 

    51.
    Cherif, A., Kinoshita, N., Taylor, D. & Mediouni Ben Jemâa, J. Molecular characterization and phylogenetic comparisons of three Mayetiola species (Diptera: Cecidomyiidae) infesting cereals in Tunisia. Applied Entomology and Zoology 52, 543–551, https://doi.org/10.1007/s13355-017-0507-y (2017).

    52.
    Gould, F. Simulation models for predicting durability of insect-resistant germ plasm: hessian fly (Diptera: Cecidomyiidae)-resistant Winter Wheat. Environ. Entomol. 15, 11–23. https://doi.org/10.1093/ee/15.1.11 (1986).
    Article  Google Scholar 

    53.
    Chen, M.-S., Liu, X., Wang, H. & El-Bouhssini, M. Hessian fly (Diptera: Cecidomyiidae) interactions with barley, rice, and wheat seedlings. J Econ Entomol 102, 1663–1672 (2009).
    Article  Google Scholar 

    54.
    Ratcliffe, R. H., Safranski, G. G., Patterson, F. L., Ohm, H. W. & Taylor, P. L. Biotype status of Hessian fly (Diptera, Cecidomyiidae) populations from the eastern United-States and their response to 14 Hessian fly resistance genes. J. Econ. Entomol. 87, 1113–1121. https://doi.org/10.1093/jee/87.4.1113 (1994).
    Article  Google Scholar 

    55.
    Tooker, J. F. & Frank, S. D. Genotypically diverse cultivar mixtures for insect pest management and increased crop yields. J. Appl. Ecol. 49, 974–985. https://doi.org/10.1111/j.1365-2664.2012.02173.x (2012).
    Article  Google Scholar 

    56.
    Erb, M., Meldau, S. & Howe, G. A. Role of phytohormones in insect-specific plant reactions. Trends Plant Sci. 17, 1–20 (2012).
    Article  Google Scholar 

    57.
    Williams, C. E., Collier, C. C., Nemacheck, J. A., Liang, C. Z. & Cambron, S. E. A lectin-like wheat gene responds systemically to attempted feeding by avirulent first-instar Hessian fly larvae. J. Chem. Ecol. 28, 1411–1428. https://doi.org/10.1023/a:1016200619766 (2002).
    CAS  Article  PubMed  Google Scholar 

    58.
    Herrera-Vasquez, A., Salinas, P. & Holuigue, L. Salicylic acid and reactive oxygen species interplay in the transcriptional control of defense genes expression (vol 6, 171, 2015). Frontiers in Plant Science 8, https://doi.org/10.3389/fpls.2017.00964 (2017).

    59.
    Hatchett, J. H., Kreitner, G. L. & Elzinga, R. J. Larval Mouthparts and Feeding Mechanism of the Hessian Fly (Diptera: Cecidomyiidae). Ann. Entomol. Soc. Am. 83, 1137–1147. https://doi.org/10.1093/aesa/83.6.1137 (1990).
    Article  Google Scholar 

    60.
    Schotzko, D. J. & Bosque-Perez, N. A. Relationship between Hessian fly infestation density and early seedling growth of resistant and susceptible wheat. J. Agric. Urban Entomol. 19, 95–107 (2002).
    Google Scholar 

    61.
    Ratcliffe, R. H. et al. Biotype composition of Hessian fly (Diptera: Cecidomyiidae) populations from the southeastern, midwestern, and northwestern United States and virulence to resistance genes in wheat. J Econ. Entomol. 93, 1319–1328 (2000).
    CAS  Article  Google Scholar 

    62.
    Song, S., Gong, W., Zhu, B. & Huang, X. Wavelength selection and spectral discrimination for paddy rice, with laboratory measurements of hyperspectral leaf reflectance. ISPRS J. Photogram. Remote Sens. 66, 672–682. https://doi.org/10.1016/j.isprsjprs.2011.05.002 (2011).
    ADS  Article  Google Scholar 

    63.
    Dechant, B., Cuntz, M., Vohland, M., Schulz, E. & Doktor, D. Estimation of photosynthesis traits from leaf reflectance spectra: correlation to nitrogen content as the dominant mechanism. Remote Sens. Environ. 196, 279–292. https://doi.org/10.1016/j.rse.2017.05.019 (2017).
    ADS  Article  Google Scholar 

    64.
    Ollinger, S. V. Sources of variability in canopy reflectance and the convergent properties of plants. New Phytol. 189, 375–394. https://doi.org/10.1111/j.1469-8137.2010.03536.x (2011).
    CAS  Article  PubMed  Google Scholar 

    65.
    Almeida Trapp, M., De Souza, G. D., Rodrigues-Filho, E., Boland, W. & Mithöfer, A. Validated method for phytohormone quantification in plants. Frontiers in Plant Science 5, https://doi.org/10.3389/fpls.2014.00417 (2014).

    66.
    Davis, T. S., Bosque-Pérez, N. A., Popova, I. & Eigenbrode, S. D. Evidence for additive effects of virus infection and water availability on phytohormone induction in a staple crop. Frontiers in Ecology and Evolution 3, https://doi.org/10.3389/fevo.2015.00114 (2015). More