Modern approaches to study plant–insect interactions in chemical ecology

Abstract

Phytochemical variation among plant species is one of the most fascinating and perplexing features of the natural world and has implications for both human health and the functioning of ecosystems. A key area of research on phytochemical variation has focused on insects that feed on plants and the enormous diversity of plant-derived compounds that reduce or deter damage by insects. Empirical studies on the ecology and evolution of these chemically mediated plant–insect interactions have been guided by a long history of theoretical development. However, until recently, such theory was substantially limited by inadequate data, a situation that is rapidly changing as ecologists partner with chemists utilizing the latest technological advances. In this Review, we aim to facilitate the union of ecological theory with modern chemistry by discussing important theoretical frameworks for studying chemical ecology and outlining the steps by which hypotheses on insect–phytochemical interactions can be advanced using current methodologies and statistical approaches. We highlight unique approaches to isolation, synthesis, spectroscopy, metabolomics and genomics relevant to chemical ecology and describe future areas for research that will bring an unprecedented understanding of phytochemical variation.

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Fig. 1
Fig. 2: Crude 1H NMR spectra from leaves of two Piper species.
Fig. 3: Periplanone and mandelalide A structure determination by total synthesis.
Fig. 4: Syntheses of methyl jasmonate and proposed modifications for asymmetric synthesis.
Fig. 5: Early approaches to the biomimetic synthesis of natural products.
Fig. 6: Building diversity via biosynthesis and diversity-oriented synthesis.

References

  1. 1.

    Fraenkel, G. S. The raison d’être of secondary plant substances. Science 129, 1466–1470 (1959).

  2. 2.

    Ehrlich, P. R. & Raven, P. H. Butterflies and plants: a study in coevolution. Evolution 18, 586 (1964).

  3. 3.

    Nicolaou, K. C., Snyder, S. A., Montagnon, T. & Vassilikogiannakis, G. The Diels–Alder reaction in total synthesis. Angew. Chem. Int. Ed. 41, 1668–1698 (2002).

  4. 4.

    Hay, M. E. & Fenical, W. Marine plant-herbivore interactions: the ecology of chemical defense. Annu. Rev. Ecol. Syst. 19, 111–145 (1988).

  5. 5.

    Felsenstein, J. Phylogenies and the comparative method. Am. Naturalist 125, 1–15 (1985).

  6. 6.

    Eisner, T. & Meinwald, Y. C. Defensive secretion of a caterpillar (Papilio). Science 150, 1733–1735 (1965).

  7. 7.

    Meinwald, J. Where might we go from here? J. Chem. Ecol. 40, 222 (2014).

  8. 8.

    Romeo, J. Perspectives in chemical ecology: Into the mainstream. Planta Medica 80, IL16 (2014).

  9. 9.

    Hay, M. E. Challenges and opportunities in marine chemical ecology. J. Chem. Ecol. 40, 216 (2014).

  10. 10.

    Dyer, L. A. New synthesis-back to the future: new approaches and directions in chemical studies of coevolution. J. Chem. Ecol. 37, 669–669 (2011).

  11. 11.

    Jones, C. G. & Firn, R. D. On the evolution of plant secondary chemical diversity. Phil. Trans. R. Soc. Lond. B Biol Sci. 333, 273–280 (1991).

  12. 12.

    Thompson, J. N. The Geographic Mosaic of Coevolution. (Univ. of Chicago Press, 2005).

  13. 13.

    Raguso, R. A. et al. The raison d’être of chemical ecology. Ecology 96, 617–630 (2015).

  14. 14.

    Rehr, S. S., Feeny, P. P. & Janzen, D. H. Chemical defence in Central American non-ant-acacias. J. Animal Ecol. 42, 405–416 (1973).

  15. 15.

    Romeo, J. T., Saunders, J. A. & Barbosa, P. Phytochemical Diversity and Redundancy in Ecological Interactions. (Plenum Press, 1996).

  16. 16.

    Berenbaum, M. & Neal, J. J. Synergism between myristicin and xanthotoxin, a naturally cooccurring plant toxicant. J. Chem. Ecol. 11, 1349–1358 (1985).

  17. 17.

    Hunter, M. D. The Phytochemical Landscape: Linking Trophic Interactions and Nutrient Dynamics. (Princeton Univ. Press, 2016).

  18. 18.

    Hilker, M. New synthesis: parallels between biodiversity and chemodiversity. J. Chem. Ecol. 40, 225–226 (2014).

  19. 19.

    Richards, L. A. et al. Phytochemical diversity and synergistic effects on herbivores. Phytochem. Rev. 15, 1153–1166 (2016).

  20. 20.

    Dyer, L. A. et al. Synergistic effects of three Piper amides on generalist and specialist herbivores. J. Chem. Ecol. 29, 2499–2514 (2003).

  21. 21.

    Richards, L. A., Dyer, L. A., Smilanich, A. M. & Dodson, C. D. Synergistic effects of amides from two Piper species on generalist and specialist herbivores. J. Chem. Ecol. 36, 1105–1113 (2010).

  22. 22.

    Tallarida, R. J. Drug Synergism and Dose-Effect Data Analysis. (Chapman & Hall, 2000).

  23. 23.

    Dweck, H. K. M. et al. Pheromones mediating copulation and attraction in Drosophila. Proc. Natl Acad. Sci. USA 112, E2829–E2835 (2015).

  24. 24.

    Ebrahim, S. A. M. et al. Drosophila avoids parasitoids by sensing their semiochemicals via a dedicated olfactory circuit. PLoS Biol. 13, e1002318 (2015).

  25. 25.

    Whitehead, S. R. & Bowers, M. D. Chemical ecology of fruit defence: synergistic and antagonistic interactions among amides from Piper. Funct. Ecol. 28, 1094–1106 (2014).

  26. 26.

    Jones, C. G., Firn, R. D. & Malcolm, S. B. On the evolution of plant secondary chemical diversity. Phil. Trans. R. Soc. Lond. B Biol Sci. 333, 273–280 (1991).

  27. 27.

    Firn, R. D. & Jones, C. G. Natural products — a simple model to explain chemical diversity. Nat. Prod Rep. 20, 382 (2003).

  28. 28.

    Becerra, J. X., Noge, K. & Venable, D. L. Macroevolutionary chemical escalation in an ancient plant-herbivore arms race. Proc. Natl Acad. Sci. USA 106, 18062–18066 (2009).

  29. 29.

    Kursar, T. A. et al. The evolution of antiherbivore defenses and their contribution to species coexistence in the tropical tree genus Inga. Proc. Natl Acad. Sci. USA 106, 18073–18078 (2009).

  30. 30.

    Moore, B., Andrew, R. L., Kulheim, C. & Foley, W. J. Explaining intraspecific diversity in plant secondary metabolites in an ecological context. New Phytol. 201, 733–750 (2014).

  31. 31.

    Fiehn, O. Metabolomics — the link between genotypes and phenotypes. Plant Mol. Biol. 48, 155–171 (2002).

  32. 32.

    Glassmire, A. E. et al. Intraspecific phytochemical variation shapes community and population structure for specialist caterpillars. New Phytol. 212, 208–219 (2016).

  33. 33.

    Richards, L. A. et al. Phytochemical diversity drives plant–insect community diversity. Proc. Natl Acad. Sci. USA 112, 10973–10978 (2015).

  34. 34.

    Janzen, D. H. When is it coevolution? Evolution 34, 611–612 (1980).

  35. 35.

    Carmona, D., Fitzpatrick, C. R. & Johnson, M. T. Fifty years of co-evolution and beyond: integrating co-evolution from molecules to species. Mol. Ecol. 24, 5315–5329 (2015).

  36. 36.

    Parsons, J. A. Isolationof two digitalis-like substances from glandular secretion of a poisonous grasshopper, Poekilocerus bufonius. Klug. J. Physiol. 169, 80 (1963).

  37. 37.

    Honda, K. Chemical basis of differential oviposition by lepidopterous insects. Arch. Insect Biochem. Physiol. 30, 1–23 (1995).

  38. 38.

    Despres, L., David, J.-P. & Gallet, C. The evolutionary ecology of insect resistance to plant chemicals. Trends Ecol. Evol. 22, 298–307 (2007).

  39. 39.

    Agrawal, A. A., Petschenka, G., Bingham, R. A., Weber, M. G. & Rasmann, S. Toxic cardenolides: chemical ecology and coevolution of specialized plant-herbivore interactions. New Phytol. 194, 28–45 (2012).

  40. 40.

    Spencer, K. C. in Chemical Mediation of Coevolution 1st edn (ed. Spencer, K. C.) 1–11 (Elsevier BV, 1988).

  41. 41.

    Brodie, E. D. et al. Parallel arms races between garter snakes and newts involving tetrodotoxin as the phenotypic interface of coevolution. J. Chem. Ecol. 31, 343–356 (2005).

  42. 42.

    Feldman, C. R., Brodie, E. D., Brodie, E. D. & Pfrender, M. E. The evolutionary origins of beneficial alleles during the repeated adaptation of garter snakes to deadly prey. Proc. Natl Acad. Sci. USA 106, 13415–13420 (2009).

  43. 43.

    Meiners, T. Chemical ecology and evolution of plant–insect interactions: a multitrophic perspective. Curr. Opin. Insect Sci. 8, 22–28 (2015).

  44. 44.

    Mithofer, A. & Boland, W. Plant defense against herbivores: chemical aspects. Annu. Rev. Plant Biol. 63, 431–450 (2012).

  45. 45.

    Thompson, J. N. The Coevolutionary Process (Univ. of Chicago Press, 1994).

  46. 46.

    Berenbaum, M. R. & Zangerl, A. R. Chemical phenotype matching between a plant and its insect herbivore. Proc. Natl Acad. Sci. USA 95, 13743–13748 (1998).

  47. 47.

    Zangerl, A. R., Stanley, M. C. & Berenbaum, M. R. Selection for chemical trait remixing in an invasive weed after reassociation with a coevolved specialist. Proc. Natl Acad. Sci. USA 105, 4547–4552 (2008).

  48. 48.

    Zangerl, A. R. & Berenbaum, M. R. Phenotype matching in wild parsnip and parsnip webworms: causes and consequences. Evolution 57, 806–815 (2003).

  49. 49.

    Althoff, D. M., Segraves, K. A. & Johnson, M. T. Testing for coevolutionary diversification: linking pattern with process. Trends Ecol. Evol. 29, 82–89 (2014).

  50. 50.

    Hembry, D. H., Yoder, J. B. & Goodman, K. R. Coevolution and the diversification of life. Am. Naturalist 184, 425–438 (2014).

  51. 51.

    Agrawal, A. A., Conner, J. K. & Rasmann, S. in Evolution After Darwin: the First 150 Years (eds Bell, M.A., Futuyma, D. J., Eanes, W. F. & Levinton, J. S.) 243–268 (Oxford Univ. Press, 2010).

  52. 52.

    Fry, J. D. The evolution of host specialization: are trade-offs overrated? Am. Naturalist 148 (Suppl.), S84–S107 (1996).

  53. 53.

    Poisot, T., Bever, J. D., Nemri, A., Thrall, P. H. & Hochberg, M. E. A conceptual framework for the evolution of ecological specialisation. Ecol. Lett. 14, 841–851 (2011).

  54. 54.

    Forister, M. L. et al. Revisiting the evolution of ecological specialization, with emphasis on insect-plant interactions. Ecology 93, 981–991 (2012).

  55. 55.

    Gompert, Z. & Messina, F. J. Genomic evidence that resource-based trade-offs limit host-range expansion in a seed beetle. Evolution 70, 1249–1264 (2016).

  56. 56.

    Futuyma, D. J. & Moreno, B. The evolution of ecological specialization. Annu. Rev. Ecol. Systemat. 19, 207–233 (1988).

  57. 57.

    Zorgo, E. et al. Life history shapes trait heredity by accumulation of loss-of-function alleles in yeast. Mol. Biol. Evol. 29, 1781–1789 (2012).

  58. 58.

    Gompert, Z. et al. The evolution of novel host use is unlikely to be constrained by trade-offs or a lack of genetic variation. Mol. Ecol. 24, 2777–2793 (2015).

  59. 59.

    Carrasco, D., Larsson, M. C. & Anderson, P. Insect host plant selection in complex environments. Curr. Opin. Insect Sci. 8, 1–7 (2015).

  60. 60.

    Janz, N. & Nylin, S. The role of female search behaviour in determining host plant range in plant feeding insects: a test of the information processing hypothesis. Proc. R. Soc. Lond. B Biol Sci. 264, 701–707 (1997).

  61. 61.

    Egan, S. P. & Funk, D. J. Individual advantages to ecological specialization: insights on cognitive constraints from three conspecific taxa. Proc. R. Soc. Lond. B Biol. Sci. 273, 843–848 (2006).

  62. 62.

    Baldwin, I. T. Jasmonate-induced responses are costly but benefit plants under attack in native populations. Proc. Natl Acad. Sci. USA 95, 8113–8118 (1998).

  63. 63.

    Massad, T. J., Dyer, L. A. & Vega, C. G. Costs of defense and a test of the carbon-nutrient balance and growth-differentiation balance hypotheses for two co-occurring classes of plant defense. PLoS ONE 7, e47554 (2012).

  64. 64.

    Stamp, N. Out of the quagmire of plant defense hypotheses. Quarterly Rev. Biol. 78, 23–55 (2003).

  65. 65.

    Cipollini, D., Walters, D. & Voelckel, C. in Annual Plant Reviews (eds Roberts, J. A., Evan, D., McManus, M. T. & Rose, J. K.) 263–307 (Wiley-Blackwell, 2014).

  66. 66.

    Smilanich, A. M., Fincher, R. M. & Dyer, L. A. Does plant apparency matter? Thirty years of data provide limited support but reveal clear patterns of the effects of plant chemistry on herbivores. New Phytol. 210, 1044–1057 (2016).

  67. 67.

    McKey, D. Adaptive patterns in alkaloid physiology. Am. Naturalist 108, 305–320 (1974).

  68. 68.

    Rosenthal, G. A. & Janzen, D. H. Herbivores: Their Interactions with Secondary Plant Metabolites. Vol. 1 (Academic Press, 1979).

  69. 69.

    Feeney, P. in Biochemical Interactions Between Plants and Insects (eds Wallace, J. W. & Mansell, R. L.) 1–40 (Springer, 1976).

  70. 70.

    Rhoades, D. F. & Cates, R. G. in Biochemical Interaction Between Plants and Insects (eds Wallace, J. W. & Mansell, R. L.) 168–213 (Springer, 1976).

  71. 71.

    Bryant, J. P., Chapin, F. S. & Klein, D. R. Carbon/nutrient balance of boreal plants in relation to vertebrate herbivory. Oikos 40, 357 (1983).

  72. 72.

    Coley, P. D., Bryant, J. P. & Chapin, F. S. Resource availability and plant antiherbivore defense. Science 230, 895–899 (1985).

  73. 73.

    Herms, D. A. & Mattson, W. J. The dilemma of plants: to grow or defend. Quarterly Rev. Biol. 67, 283–335 (1992).

  74. 74.

    Bezemer, T. M. & Jones, T. H. Plant-insect herbivore interactions in elevated atmospheric CO2: quantitative analyses and guild effects. Oikos 82, 212 (1998).

  75. 75.

    Koricheva, J., Larsson, S., Haukioja, E., Keinänen, M. & Keinanen, M. Regulation of woody plant secondary metabolism by resource availability: hypothesis testing by means of meta-analysis. Oikos 83, 212 (1998).

  76. 76.

    Zvereva, E. L. & Kozlov, M. V. Consequences of simultaneous elevation of carbon dioxide and temperature for plant-herbivore interactions: a metaanalysis. Global Change Biol. 12, 27–41 (2006).

  77. 77.

    Stiling, P. & Cornelissen, T. How does elevated carbon dioxide (CO2) affect plant–herbivore interactions? A field experiment and meta-analysis of CO2-mediated changes on plant chemistry and herbivore performance. Global Change Biol. 13, 1823–1842 (2007).

  78. 78.

    Schuman, M. C. & Baldwin, I. T. The layers of plant responses to insect herbivores. Annu. Rev. Entomol. 61, 373–394 (2016).

  79. 79.

    Duffey, S. S. & Stout, M. J. Antinutritive and toxic components of plant defense against insects. Arch. Insect Biochem. Physiol. 32, 3–37 (1996).

  80. 80.

    Price, P. W. et al. Interactions among three trophic levels: influence of plants on interactions between insect herbivores and natural enemies. Annu. Rev. Ecol. Systemat. 11, 41–65 (1980).

  81. 81.

    Rosenthal, G. A. & Berenbaum, M. R. Herbivores: Their Interactions With Secondary Plant Metabolites: Ecological and Evolutionary Processes. Vol. 2 (Academic Press, 2012).

  82. 82.

    Rascio, N. & Navari-Izzo, F. Heavy metal hyperaccumulating plants: How and why do they do it? And what makes them so interesting? Plant Sci. 180, 169–181 (2011).

  83. 83.

    Berenbaum, M. Coumarins and caterpillars: a case for coevolution. Evolution 37, 163 (1983).

  84. 84.

    Cornell, H. V. & Hawkins, B. A. Accumulation of native parasitoid species on introduced herbivores: a comparison of hosts as natives and hosts as invaders. Am. Naturalist 141, 847–865 (1993).

  85. 85.

    Jeschke, V., Gershenzon, J. & Vassão, D. G. in Advances in Botanical Research Vol. 80 (ed. Stanislav, S.) 199–245 (Academic Press, 2016).

  86. 86.

    Dyer, L. A. Tasty generalists and nasty specialists? Antipredator mechanisms tropical Lepidopteran larvae. Ecology 76, 1483–1496 (1995).

  87. 87.

    Ode, P. J. Plant chemistry and natural enemy fitness: effects on herbivore and natural enemy interactions. Annu. Rev. Entomol. 51, 163–185 (2006).

  88. 88.

    Smilanich, A. M., Dyer, L. A., Chambers, J. Q. & Bowers, M. D. Immunological cost of chemical defence and the evolution of herbivore diet breadth. Ecol. Lett. 12, 612–621 (2009).

  89. 89.

    Dyer, L. A. in Tropical Forest Community Ecology (eds Carson, W. P. & Schnitzer, S. A.) 275-293 (Blackwell Publishing, 2008).

  90. 90.

    Pearson, C. V., Massad, T. J. & Dyer, L. A. Diversity cascades in alfalfa fields: from plant quality to agroecosystem diversity. Environ. Entomol. 37, 947–955 (2008).

  91. 91.

    Martinsen, G. D., Driebe, E. M. & Whitham, T. G. Indirect interactions mediated by changing plant chemistry: beaver browsing benefits beetles. Ecology 79, 192 (1998).

  92. 92.

    Wimp, G. M. et al. Plant genetics predicts intra-annual variation in phytochemistry and arthropod community structure. Mol. Ecol. 16, 5057–5069 (2007).

  93. 93.

    Kessler, A. The information landscape of plant constitutive and induced secondary metabolite production. Curr. Opin. Insect Sci. 8, 47–53 (2015).

  94. 94.

    Turlings, T. C. J., Tumlinson, J. H. & Lewis, W. J. Exploitation of herbivore-induced plant odors by host-seeking parasitic wasps. Science 250, 1251–1253 (1990).

  95. 95.

    Muller, M. S. et al. Tri-trophic effects of plant defenses: chickadees consume caterpillars based on host leaf chemistry. Oikos 114, 507–517 (2006).

  96. 96.

    Poelman, E. H. et al. Hyperparasitoids use herbivore-induced plant volatiles to locate their parasitoid host. PLoS Biol. 10, e1001435 (2012).

  97. 97.

    Aldrich, J. R. et al. Insect chemical ecology research in the United States Department of Agriculture — Agricultural Research Service. Pest Management Sci. 59, 777–787 (2003).

  98. 98.

    Nicolaou, K. C. & Snyder, S. A. Chasing molecules that were never there: misassigned natural products and the role of chemical synthesis in modern structure elucidation. Angew. Chem. Int. Ed Engl. 44, 1012–1044 (2005).

  99. 99.

    Still, W. C. (.+-.)-Periplanone-B. Total synthesis and structure of the sex excitant pheromone of the American cockroach. J. Am. Chem. Soc. 101, 2493–2495 (1979).

  100. 100.

    Willwacher, J., Heggen, B., Wirtz, C., Thiel, W. & Fürstner, A. Total synthesis, stereochemical revision, and biological reassessment of mandelalide A: chemical mimicry of intrafamily relationships. Chemistry 21, 10416–10430 (2015).

  101. 101.

    Veerasamy, N. et al. Enantioselective total synthesis of mandelalide A and isomandelalide A: discovery of a cytotoxic ring-expanded isomer. J. Am. Chem. Soc. 138, 770–773 (2016).

  102. 102.

    Nguyen, M. H., Imanishi, M., Kurogi, T., Amos, B. & Smith, I. Total synthesis of (−)-mandelalide A exploiting anion relay chemistry (ARC): identification of a type II ARC/CuCN cross-coupling protocol. J. Am. Chem. Soc. 138, 3675–3678 (2016).

  103. 103.

    Snyder, K. M. et al. Towards theory driven structure elucidation of complex natural products: mandelalides and coibamide A. Org. Biomol. Chem. 14, 5826–5831 (2016).

  104. 104.

    Dunn, W. B. et al. Mass appeal: metabolite identification in mass spectrometry-focused untargeted metabolomics. Metabolomics 9, 44–66 (2013).

  105. 105.

    Kim, J. H., Lee, B. W., Schroeder, F. C. & Jander, G. Identification of indole glucosinolate breakdown products with antifeedant effects on Myzus persicae (green peach aphid). Plant J. 54, 1015–1026 (2008).

  106. 106.

    Dyer, L. A., Dodson, C. D., Beihoffer, J. & Letourneau, D. K. Trade-offs in antiherbivore defenses in Piper cenocladum: ant mutualists versus plant secondary metabolites. J. Chem. Ecol. 27, 581–592 (2001).

  107. 107.

    Dodson, C. D., Dyer, L. A., Searcy, J., Wright, Z. & Letourneau, D. K. Cenocladamide, a dihydropyridone alkaloid from Piper cenocladum. Phytochemistry 53, 51–54 (2000).

  108. 108.

    Dyer, L. A. et al. Ecological causes and consequences of variation in defensive chemistry of a neotropical shrub. Ecology 85, 2795–2803 (2004).

  109. 109.

    Stökl, J., Hofferberth, J., Pritschet, M., Brummer, M. & Ruther, J. Stereoselective chemical defense in the Drosophila parasitoid Leptopilina heterotoma is mediated by (−)-iridomyrmecin and (+)-isoiridomyrmecin. J. Chem. Ecol. 38, 331–339 (2012).

  110. 110.

    Rasmann, S. et al. Recruitment of entomopathogenic nematodes by insect-damaged maize roots. Nature 434, 732–737 (2005).

  111. 111.

    Beckett, J. S., Beckett, J. D. & Hofferberth, J. E. A divergent approach to the diastereoselective synthesis of several ant-associated iridoids. Org. Lett. 12, 1408–1411 (2010).

  112. 112.

    Richards, L. A. et al. Synergistic effects of iridoid glycosides on the survival, development and immune response of a specialist caterpillar, Junonia coenia (Nymphalidae). J. Chem. Ecol. 38, 1276–1284 (2012).

  113. 113.

    Azmir, J. et al. Techniques for extraction of bioactive compounds from plant materials: a review. J. Food Eng. 117, 426–436 (2013).

  114. 114.

    Armenta, S., Garrigues, S. & de la Guardia, M. The role of green extraction techniques in Green Analytical Chemistry. Trends Analyt. Chem. 71, 2–8 (2015).

  115. 115.

    Bucar, F., Wube, A. & Schmid, M. Natural product isolation — how to get from biological material to pure compounds. Nat. Prod. Rep. 30, 525–545 (2013).

  116. 116.

    Demole, E., Lederer, E. & Mercier, D. Isolement et détermination de la structure du jasmonate de méthyle, constituant odorant caractéristique de l’essence de jasmin. Helv. Chim. Acta 45, 675–685 (1962).

  117. 117.

    Sisido, K., Kurozumi, S. & Utimoto, K. Synthesis of methyl dl-jasmonate. J. Org. Chem. 34, 2661–2664 (1969).

  118. 118.

    Luo, F. T. & Negishi, E. Nickel- or palladium-catalyzed cross coupling. 27. Palladium-catalyzed allylation of lithium 3-alkenyl-1-cyclopentenolates-triethylborane and its application to a selective synthesis of methyl (Z)-jasmonate. Tetrahedron Lett. 26, 2177–2180 (1985).

  119. 119.

    Kataoka, H., Yamada, T., Goto, K. & Tsuji, J. An efficient synthetic method of methyl (±)-jasmonate. Tetrahedron 43, 4107–4112 (1987).

  120. 120.

    Yoshioka, A. & Yamada, T. Development of methyl (±)-jasmonate production process on an industrial scale. J. Synthet. Org. Chem., Japan 48, 56–64 (1990).

  121. 121.

    Farmer, E. E. & Ryan, C. A. Interplant communication: airborne methyl jasmonate induces synthesis of proteinase inhibitors in plant leaves. Proc. Natl Acad. Sci. USA 87, 7713–7716 (1990).

  122. 122.

    Posner, G. H. & Asirvatham, E. A short, asymmetric synthesis of natural (-)-methyl jasmonate. J. Org. Chem. 50, 2589–2591 (1985).

  123. 123.

    Weinges, K., Gethöffer, H., Huber-Patz, U., Rodewald, H. & Irngartinger, H. Chemie und Stereochemie der Iridoide, IX. EPC-Synthese von (1 R,2 R,2-Z)-(−)-Methyl-jasmonat aus Catalpol – Kristall- und Molekularstruktur von Methyl-dehydrojasmonat-semicarbazon. Liebigs Ann. Chem. 1987, 361–366 (1987).

  124. 124.

    Takeda, H., Watanabe, H. & Nakada, M. Asymmetric total synthesis of enantiopure (−)-methyl jasmonate via catalytic asymmetric intramolecular cyclopropanation of α-diazo-β-keto sulfone. Tetrahedron 62, 8054–8063 (2006).

  125. 125.

    Quinkert, G., Adam, F. & Dürner, G. Asymmetric synthesis of methyl jasmonate. Angew. Chem. Int. Ed. 21, 856–856 (1982).

  126. 126.

    Jansen, D. J. & Shenvi, R. A. Synthesis of (−)-neothiobinupharidine. J. Am. Chem. Soc. 135, 1209–1212 (2013).

  127. 127.

    Germain, N., Guénée, L., Mauduit, M. & Alexakis, A. Asymmetric conjugate addition to α-substituted enones/enolate trapping. Org. Lett. 16, 118–121 (2014).

  128. 128.

    Mase, N. et al. Organocatalytic enantioselective Michael additions of malonates to 2-cyclopentenone. Synlett 2010, 2340–2344 (2010).

  129. 129.

    Zheljazkov, V. D., Shiwakoti, S., Jeliazkova, E. A. & Astatkie, T. in Medicinal and Aromatic Crops: Production, Phytochemistry, and Utilization (eds Jeliazkov (Zheljazkov), V D. & Cantrell, C. L.) 145–166 (ACS Publications, 2016).

  130. 130.

    Liu, W. C., Gong, T. & Zhu, P. Advances in exploring alternative Taxol sources. RSC Adv. 6, 48800–48809 (2016).

  131. 131.

    Li, D., Baldwin, I. T. & Gaquerel, E. Beyond the canon: within-plant and population-level heterogeneity in jasmonate signaling engaged by plant-insect interactions. Plants 5, 14 (2016).

  132. 132.

    Chapuis, C. et al. Route scouting towards a methyl jasmonate precursor. Helvet. Chim. Acta 99, 95–109 (2016).

  133. 133.

    Robinson, R. LXIII. — A synthesis of tropinone. J. Chem. Soc., Trans. 111, 762–768 (1917).

  134. 134.

    Robinson, R. LXXV. — A theory of the mechanism of the phytochemical synthesis of certain alkaloids. J. Chem. Soc., Trans. 111, 876–899 (1917).

  135. 135.

    Yoder, R. A. & Johnston, J. N. A case study in biomimetic total synthesis: polyolefin carbocyclizations to terpenes and steroids. Chem. Rev. 105, 4730–4756 (2005).

  136. 136.

    Stork, G. & Burgstahler, A. W. The stereochemistry of polyene cyclization. J. Am. Chem. Soc. 77, 5068–5077 (1955).

  137. 137.

    Gamboni, G., Schinz, H. & Eschenmoser, A. Über den sterischen Verlauf der säurekatalysierten Cyclisation in der Terpenreihe. Cyclisation der cis-7-Methyl-octadien-(2,6)-säure-(1). Health Care Anal. 37, 964–971 (1954).

  138. 138.

    de la Torre, M. C. & Sierra, M. A. Comments on recent achievements in biomimetic organic synthesis. Angew. Chem. Int. Ed. 43, 160–181 (2004).

  139. 139.

    Nielsen, T. E. & Schreiber, S. L. Towards the optimal screening collection: a synthesis strategy. Angew. Chem. Int. Ed. 47, 48–56 (2008).

  140. 140.

    Lee, D., Sello, J. K. & Schreiber, S. L. Pairwise use of complexity-generating reactions in diversity-oriented organic synthesis. Org. Lett. 2, 709–712 (2000).

  141. 141.

    Schreiber, S. L. Target-oriented and diversity-oriented organic synthesis in drug discovery. Science 287, 1964–1969 (2000).

  142. 142.

    Burke, M. D. & Schreiber, S. L. A planning strategy for diversity-oriented synthesis. Angew. Chem. Int. Ed. 43, 46–58 (2004).

  143. 143.

    Kato, N. et al. Diversity-oriented synthesis yields novel multistage antimalarial inhibitors. Nature 538, 344–349 (2016).

  144. 144.

    Kurita, K. L., Glassey, E. & Linington, R. G. Integration of high-content screening and untargeted metabolomics for comprehensive functional annotation of natural product libraries. Proc. Natl Acad. Sci. USA 112, 11999–12004 (2015).

  145. 145.

    Wang, M. et al. Sharing and community curation of mass spectrometry data with Global Natural Products Social Molecular Networking. Nat. Biotechnol. 34, 828–837 (2016).

  146. 146.

    Kuhlisch, C. & Pohnert, G. Metabolomics in chemical ecology. Nat. Prod. Rep. 32, 937–955 (2015).

  147. 147.

    Poulson-Ellestad, K. L. et al. Metabolomics and proteomics reveal impacts of chemically mediated competition on marine plankton. Proc. Natl Acad. Sci. USA 111, 9009–9014 (2014).

  148. 148.

    Bais, P. et al. PlantMetabolomics.org: a web portal for plant metabolomics experiments. Plant Physiol. 152, 1807–1816 (2010).

  149. 149.

    Guo, A. C. et al. ECMDB: the E. coli metabolome database. Nucleic Acids Res. 41, D625–D630 (2012).

  150. 150.

    Jewison, T. et al. YMDB: the yeast metabolome database. Nucleic Acids Res. 40, D815–D820 (2011).

  151. 151.

    Mardis, E. R. Next-generation sequencing platforms. Annu. Rev. Anal. Chem. 6, 287–303 (2013).

  152. 152.

    Ellegren, H. Genome sequencing and population genomics in non-model organisms. Trends Ecol. Evol. 29, 51–63 (2014).

  153. 153.

    Koboldt, D. C., Steinberg, K. M., Larson, D. E., Wilson, R. K. & Mardis, E. R. The next-generation sequencing revolution and its impact on genomics. Cell 155, 27–38 (2013).

  154. 154.

    Rubin, C.-J. et al. Whole-genome resequencing reveals loci under selection during chicken domestication. Nature 464, 587–591 (2010).

  155. 155.

    McGettigan, P. A. Transcriptomics in the RNA-seq era. Curr. Opin. Chem. Biol. 17, 4–11 (2013).

  156. 156.

    Deagle, B. E., Jones, F. C., Absher, D. M., Kingsley, D. M. & Reimchen, T. E. Phylogeography and adaptation genetics of stickleback from the Haida Gwaii archipelago revealed using genome-wide single nucleotide polymorphism genotyping. Mol. Ecol. 22, 1917–1932 (2013).

  157. 157.

    Narum, S. R., Buerkle, C. A., Davey, J. W., Miller, M. R. & Hohenlohe, P. A. Genotyping-by-sequencing in ecological and conservation genomics. Mol. Ecol. 22, 2841–2847 (2013).

  158. 158.

    Adamski, J. Genome-wide association studies with metabolomics. Genome Med. 4, 34 (2012).

  159. 159.

    Carreno-Quintero, N., Bouwmeester, H. J. & Keurentjes, J. J. B. Genetic analysis of metabolome–phenotype interactions: from model to crop species. Trends Genet. 29, 41–50 (2013).

  160. 160.

    Keurentjes, J. J. B. et al. The genetics of plant metabolism. Nat. Genet. 38, 842–849 (2006).

  161. 161.

    Chan, E. K. F., Rowe, H. C., Hansen, B. G. & Kliebenstein, D. J. The complex genetic architecture of the metabolome. PLoS Genet. 6, e1001198 (2010).

  162. 162.

    Fiehn, O. et al. Metabolite profiling for plant functional genomics. Nat. Biotechnol. 18, 1157–1161 (2000).

  163. 163.

    Matsuda, F. et al. Dissection of genotype-phenotype associations in rice grains using metabolome quantitative trait loci analysis. Plant J. 70, 624–636 (2012).

  164. 164.

    Schauer, N. et al. Comprehensive metabolic profiling and phenotyping of interspecific introgression lines for tomato improvement. Nat. Biotechnol. 24, 447–454 (2006).

  165. 165.

    Riedelsheimer, C. et al. Genome-wide association mapping of leaf metabolic profiles for dissecting complex traits in maize. Proc. Natl Acad. Sci. 109, 8872–8877 (2012).

  166. 166.

    Wen, W. et al. Metabolome-based genome-wide association study of maize kernel leads to novel biochemical insights. Nat. Commun. 5, 3438 (2014).

  167. 167.

    Feng, J. et al. Characterization of metabolite quantitative trait loci and metabolic networks that control glucosinolate concentration in the seeds and leaves of Brassica napus. New Phytol. 193, 96–108 (2012).

  168. 168.

    Lau, W. & Sattely, E. S. Six enzymes from mayapple that complete the biosynthetic pathway to the etoposide aglycone. Science 349, 1224–1228 (2015).

  169. 169.

    Nützmann, H. W., Huang, A. & Osbourn, A. Plant metabolic clusters — from genetics to genomics. New Phytol. 211, 771–789 (2016).

  170. 170.

    Mitchell-Olds, T. Arabidopsis thaliana and its wild relatives: a model system for ecology and evolution. Trends Ecol. Evol. 16, 693–700 (2001).

  171. 171.

    Agrawal, A. A. Current trends in the evolutionary ecology of plant defence. Funct. Ecol. 25, 420–432 (2011).

  172. 172.

    Wilson, J. S. et al. Host conservatism, host shifts and diversification across three trophic levels in two Neotropical forests. J. Evol. Biol. 25, 532–546 (2012).

  173. 173.

    Coley, P. D. & Kursar, T. A. On tropical forests and their pests. Science 343, 35–36 (2014).

  174. 174.

    Coley, P. D. et al. Divergent defensive strategies of young leaves in two species of Inga. Ecology 86, 2633–2643 (2005).

  175. 175.

    Endara, M.-J. et al. Divergent evolution in antiherbivore defences within species complexes at a single Amazonian site. J. Ecol. 103, 1107–1118 (2015).

  176. 176.

    Brenes-Arguedas, T. et al. Contrasting mechanisms of secondary metabolite accumulation during leaf development in two tropical tree species with different leaf expansion strategies. Oecologia 149, 91–100 (2006).

  177. 177.

    Wu, S.-B., Meyer, R. S., Whitaker, B. D., Litt, A. & Kennelly, E. J. A new liquid chromatography–mass spectrometry-based strategy to integrate chemistry, morphology, and evolution of eggplant (Solanum) species. J. Chromatogr. A 1314, 154–172 (2013).

  178. 178.

    McKey, D., Rosenthal, G. A. & Janzen, D. H. in Herbivores: Their Interaction With Secondary Plant Metabolites Vol. 1 (Rosenthal, G. A. & Berenbaum, M. R.) 55–133 (Academic Press, 1979).

  179. 179.

    Ohnmeiss, T. E. & Baldwin, I. T. Optimal defense theory predicts the ontogeny of an induced nicotine defense. Ecology 81, 1765 (2000).

  180. 180.

    Bolker, J. Model organisms: There’s more to life than rats and flies. Nature 491, 31–33 (2012).

  181. 181.

    Hamilton, J. G., Zangerl, A. R., DeLucia, E. H. & Berenbaum, M. R. The carbon-nutrient balance hypothesis: its rise and fall. Ecol. Lett. 4, 86–95 (2001).

  182. 182.

    Watrous, J. et al. Mass spectral molecular networking of living microbial colonies. Proc. Natl Acad. Sci. USA 109, E1743–E1752 (2012).

  183. 183.

    Cordell, G. A. Biosynthesis of sesquiterpenes. Chem. Rev. 76, 425–460 (1976).

  184. 184.

    Tetzlaff, C. N. et al. A gene cluster for biosynthesis of the sesquiterpenoid antibiotic pentalenolactone in Streptomyces avermitilis. Biochemistry 45, 6179–6186 (2006).

  185. 185.

    De Kraker, J.-W., Franssen, M. C., Joerink, M., De Groot, A. & Bouwmeester, H. J. Biosynthesis of costunolide, dihydrocostunolide, and leucodin. Demonstration of cytochrome P450-catalyzed formation of the lactone ring present in sesquiterpene lactones of chicory. Plant Physiol. 129, 257–268 (2002).

  186. 186.

    Bülow, N. & König, W. A. The role of germacrene D as a precursor in sesquiterpene biosynthesis: investigations of acid catalyzed, photochemically and thermally induced rearrangements. Phytochemistry 55, 141–168 (2000).

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Acknowledgements

The authors dedicate this article to Professor Jerrold Meinwald for his transformative contributions to the field of chemical ecology. The authors thank M.A. Stanton and two anonymous reviewers for their excellent edits and suggestions to this manuscript. The authors’ research was funded by the National Science Foundation (DEB-1442103 and DEB-1638793), the FAPESP (São Paulo Research Foundation, 2014/50316-7) and by a generous donation from the Hitchcock Fund for Chemical Ecology Research.

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L.A.D., C.S.P., K.M.O., L.A.R., T.J.M., A.M.S., M.L.F., T.L.P., L.M.G., P.J.H., A.E.E., A.E.G., J.G.H., C.M., S.Y., N.A.P., N.D.M., J.P.J., H.L.S., O.S. and C.S.J. researched the data for the article and wrote the article. All authors contributed to discussion of the content and reviewed and/or edited the manuscript before revision.

Correspondence to Christopher S. Jeffrey.

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Glossary

Plant secondary metabolites

Organic compounds not associated with primary metabolic functions; in plants in particular, these compounds have been the subject of research in biomedical fields and in chemical ecology, in which they have been found to have largely defensive functions (for example, anti-herbivore and antibacterial functions).

Co-evolution

The evolution of reciprocal adaptation in response to reciprocal natural selection occurring with respect to a pair or complex of interacting species; often hypothesized to be associated with adaptive radiation and co-diversification.

Synergy

Combined effects of compounds in a mixture that are greater than the sum of effects for the individual compounds acting in isolation.

Speciation

The evolutionary process that results in the formation of new species by the divergence of an ancestral population into two genetically independent populations. This process is most often characterized by the evolution of reproductive isolation and the subsequent independent evolution of lineages.

Parasitoids

Organisms characterized by a unique form of parasitic lifestyle, in which the host is killed by the developing juvenile stage; the most diverse taxa, the wasps (insect order Hymenoptera) and flies (insect order Diptera), have a dramatic influence on the ecology of terrestrial ecosystems.

Antagonistic pleiotropy

A type of genetic architecture in which a single genetic locus affects more than one trait (which can include performance or fitness in more than one environment), with effects of one trait (or in one environment) being positive and effects of the other trait (or environment) being negative.

Dereplication methods

Fast identification of compounds using orthogonal physicochemical characteristics to compare spectroscopic data with molecular features gleaned from libraries of known compounds and to confirm identifications.

Macrolides

Phytochemicals that have antibacterial or antifungal properties.

Next-generation sequencing

Modern DNA sequencing platforms that leverage direct sequencing by synthesis technologies to simultaneously determine the DNA sequences of millions or hundreds of millions of DNA fragments. Also known as high-throughput or massively parallel sequencing, these methods have revolutionized genomics.

RNA sequencing

The use of next-generation DNA sequencing approaches to characterize and quantify RNA from biological samples. RNA extracted from tissue is converted into cDNA and directly sequenced on next-generation sequencing platforms such as Illumina. These approaches allow for efficient characterization of the coding regions of genomes (for example, transcriptome sequencing) and for analysis of differential gene expression.

Genome-wide association studies

Observational studies of a genome-wide set of genetic variants in a sample of phenotypically variable individuals aimed at detecting specific variants in which genotypic variation is associated with phenotypic variation.

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Dyer, L.A., Philbin, C.S., Ochsenrider, K.M. et al. Modern approaches to study plant–insect interactions in chemical ecology. Nat Rev Chem 2, 50–64 (2018). https://doi.org/10.1038/s41570-018-0009-7

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