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Broadening the scope of biocatalytic C–C bond formation

Abstract

Enzymes exercise impeccable control over chemoselectivity, site selectivity and stereoselectivity in reactions they mediate, such that we have witnessed a surge in the development of new biocatalytic methods. Although carbon–carbon (C–C) bonds are the central framework of organic molecules, biocatalytic methods for their formation have largely been limited to a select few lyase enzymes. Thus, despite several decades of research, there are not many biocatalytic C–C-bond-forming transformations at our disposal. This Review describes the suite of enzymes available for highly selective, biocatalytic C–C bond formation. We discuss each class of enzyme in terms of native activity, alteration of this activity through protein or substrate engineering, and its utility in abiotic synthesis.

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Fig. 1: Engineering enzymes for organic synthesis.
Fig. 2: Biocatalytic alkylation using activated electrophilic coenzymes or alkyl donors.
Fig. 3: Biosynthetic and biocatalytic ring-closing reactions.
Fig. 4: Site-selective acetylation of resorcinol derivatives catalysed by an acyltransferase.
Fig. 5: Intermolecular oxidative coupling reactions in nature.
Fig. 6: Cyclization reactions catalysed by native and engineered enzymes.
Fig. 7: Engineering non-natural carbene transferases for C–C bond formation.

References

  1. 1.

    Drew, S. W. & Demain, A. L. Effect of primary metabolites on secondary metabolism. Annu. Rev. Microbiol. 31, 343–356 (1977).

    CAS  PubMed  Google Scholar 

  2. 2.

    Williams, D. H., Stone, M. J., Hauck, P. R. & Rahman, S. K. Why are secondary metabolites (natural products) biosynthesized? J. Nat. Prod. 52, 1189–1208 (1989).

    CAS  PubMed  Google Scholar 

  3. 3.

    Maplestone, R. A., Stone, M. J. & Williams, D. H. The evolutionary role of secondary metabolites: a review. Gene 115, 151–157 (1992).

    CAS  PubMed  Google Scholar 

  4. 4.

    Devine, P. N. et al. Extending the application of biocatalysis to meet the challenges of drug development. Nat. Rev. Chem. 2, 409–421 (2018).

    Google Scholar 

  5. 5.

    Campos, K. R. et al. The importance of synthetic chemistry in the pharmaceutical industry. Science 363, eaat0805 (2019).

    CAS  PubMed  Google Scholar 

  6. 6.

    Bornscheuer, U. T. et al. Engineering the third wave of biocatalysis. Nature 485, 185–194 (2012). This paper provides an overview of biocatalysis and details the use of engineered enzymes as industrial biocatalysts.

    CAS  PubMed  Google Scholar 

  7. 7.

    Turner, N. J. & O’Reilly, E. Biocatalytic retrosynthesis. Nat. Chem. Biol. 9, 285–288 (2013).

    CAS  PubMed  Google Scholar 

  8. 8.

    Sheldon, R. A. & Woodley, J. M. Role of biocatalysis in sustainable chemistry. Chem. Rev. 118, 801–838 (2018).

    CAS  PubMed  Google Scholar 

  9. 9.

    Hönig, M., Sondermann, P., Turner, N. J. & Carreira, E. M. Enantioselective chemo- and biocatalysis: partners in retrosynthesis. Angew. Chem. Int. Ed. 56, 8942–8973 (2017).

    Google Scholar 

  10. 10.

    Schoemaker, H. E., Mink, D. & Wubbolts, M. G. Dispelling the myths — biocatalysis in industrial synthesis. Science 299, 1694–1697 (2003). This paper addresses common misconceptions regarding the availability, stability and reactivity of biocatalysts.

    CAS  PubMed  Google Scholar 

  11. 11.

    Hult, K. & Berglund, P. Enzyme promiscuity: mechanism and applications. Trends Biotechnol. 25, 231–238 (2007).

    CAS  PubMed  Google Scholar 

  12. 12.

    Brustad, E. M. & Arnold, F. H. Optimizing non-natural protein function with directed evolution. Curr. Opin. Chem. Biol. 15, 201–210 (2011).

    CAS  PubMed  Google Scholar 

  13. 13.

    Packer, M. S. & Liu, D. R. Methods for the directed evolution of proteins. Nat. Rev. Genet. 16, 379–394 (2015). This paper is an excellent resource for those seeking a more in-depth understanding of methods for the directed evolution of proteins.

    CAS  PubMed  Google Scholar 

  14. 14.

    Renata, H., Wang, Z. J. & Arnold, F. H. Expanding the enzyme universe: accessing non-natural reactions by mechanism-guided directed evolution. Angew. Chem. Int. Ed. 54, 3351–3367 (2015).

    CAS  Google Scholar 

  15. 15.

    Turner, N. J. Directed evolution drives the next generation of biocatalysts. Nat. Chem. Biol. 5, 567–573 (2009).

    CAS  PubMed  Google Scholar 

  16. 16.

    Fox, R. J. et al. Improving catalytic function by ProSAR-driven enzyme evolution. Nat. Biotechnol. 25, 338–344 (2007).

    CAS  PubMed  Google Scholar 

  17. 17.

    Fox, R. J. & Huisman, G. W. Enzyme optimization: moving from blind evolution to statistical exploration of sequence–function space. Trends Biotechnol. 26, 132–138 (2008).

    CAS  PubMed  Google Scholar 

  18. 18.

    Lutz, S. Beyond directed evolution: semi-rational protein engineering and design. Curr. Opin. Biotechnol. 21, 734–743 (2010).

    CAS  PubMed  PubMed Central  Google Scholar 

  19. 19.

    Currin, A., Swainston, N., Day, P. J. & Kell, D. B. Synthetic biology for the directed evolution of protein biocatalysts: navigating sequence space intelligently. Chem. Soc. Rev. 44, 1172–1239 (2015).

    CAS  PubMed  Google Scholar 

  20. 20.

    Yang, K. K., Wu, Z. & Arnold, F. H. Machine-learning-guided directed evolution for protein engineering. Nat. Methods 16, 687–694 (2019).

    CAS  PubMed  Google Scholar 

  21. 21.

    Huffman, M. A. et al. Design of an in vitro biocatalytic cascade for the manufacture of islatravir. Science 366, 1255–1259 (2019). This paper demonstrates the tunability of enzymes through directed evolution and shows how biocatalysts can successfully replace chemical steps on an industrial scale.

    CAS  PubMed  Google Scholar 

  22. 22.

    McLaughlin, M. et al. Enantioselective synthesis of 4ʹ-ethynyl-2-fluoro-2ʹ-deoxyadenosine (EFdA) via enzymatic desymmetrization. Org. Lett. 19, 926–929 (2017).

    CAS  PubMed  Google Scholar 

  23. 23.

    Schrittwieser, J. H., Velikogne, S., Hall, M. & Kroutil, W. Artificial biocatalytic linear cascades for preparation of organic molecules. Chem. Rev. 118, 270–348 (2018).

    CAS  PubMed  Google Scholar 

  24. 24.

    Rudroff, F. et al. Opportunities and challenges for combining chemo- and biocatalysis. Nat. Catal. 1, 12–22 (2018).

    Google Scholar 

  25. 25.

    Brovetto, M., Gamenara, D., Saenz Méndez, P. & Seoane, G. A. C−C bond-forming lyases in organic synthesis. Chem. Rev. 111, 4346–4403 (2011). This paper provides a thorough background on the use of lyases for C–C bond formation in organic synthesis.

    CAS  PubMed  Google Scholar 

  26. 26.

    Fessner, W.-D. Enzyme mediated C–C bond formation. Curr. Opin. Chem. Biol. 2, 85–97 (1998).

    CAS  PubMed  Google Scholar 

  27. 27.

    Machajewski, T. D. & Wong, C.-H. The catalytic asymmetric aldol reaction. Angew. Chem. Int. Ed. 39, 1352–1375 (2000).

    CAS  Google Scholar 

  28. 28.

    Schmidt, N. G., Eger, E. & Kroutil, W. Building bridges: biocatalytic C–C-bond formation toward multifunctional products. ACS Catal. 6, 4286–4311 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  29. 29.

    Pohl, M., Sprenger, G. A. & Müller, M. A new perspective on thiamine catalysis. Curr. Opin. Biotechnol. 15, 335–342 (2004).

    CAS  PubMed  Google Scholar 

  30. 30.

    Müller, M., Gocke, D. & Pohl, M. Thiamin diphosphate in biological chemistry: exploitation of diverse thiamin diphosphate-dependent enzymes for asymmetric chemoenzymatic synthesis. FEBS J. 276, 2894–2904 (2009).

    PubMed  Google Scholar 

  31. 31.

    Pohl, M., Lingen, B. & Müller, M. Thiamin diphosphate-dependent enzymes: new aspects of asymmetric C–C bond formation. Chem. Eur. J. 8, 5288–5295 (2002).

    CAS  PubMed  Google Scholar 

  32. 32.

    Effenberger, F., Förster, S. & Wajant, H. Hydroxynitrile lyases in stereoselective catalysis. Curr. Opin. Biotechnol. 11, 532–539 (2000).

    CAS  PubMed  Google Scholar 

  33. 33.

    Griengl, H., Schwab, H. & Fechter, M. The synthesis of chiral cyanohydrins by oxynitrilases. Trends Biotechnol. 18, 252–256 (2000).

    CAS  PubMed  Google Scholar 

  34. 34.

    Sukumaran, J. & Hanefeld, U. Enantioselective C–C bond synthesis catalysed by enzymes. Chem. Soc. Rev. 34, 530–542 (2005).

    CAS  PubMed  Google Scholar 

  35. 35.

    Schallmey, A. & Schallmey, M. Recent advances on halohydrin dehalogenases — from enzyme identification to novel biocatalytic applications. Appl. Microbiol. Biotechnol. 100, 7827–7839 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  36. 36.

    Miao, Y., Rahimi, M., Geertsema, E. M. & Poelarends, G. J. Recent developments in enzyme promiscuity for carbon–carbon bond-forming reactions. Curr. Opin. Chem. Biol. 25, 115–123 (2015).

    CAS  PubMed  Google Scholar 

  37. 37.

    Resch, V., Schrittwieser, J. H., Siirola, E. & Kroutil, W. Novel carbon–carbon bond formations for biocatalysis. Curr. Opin. Biotechnol. 22, 793–799 (2011).

    PubMed  PubMed Central  Google Scholar 

  38. 38.

    Cantoni, G. L. Biological methylation: selected aspects. Annu. Rev. Biochem. 44, 435–451 (1975).

    CAS  PubMed  Google Scholar 

  39. 39.

    Chiang, P. K. et al. S-Adenosylmethionine and methylation. FASEB J. 10, 471–480 (1996).

    CAS  PubMed  Google Scholar 

  40. 40.

    Fontecave, M., Atta, M. & Mulliez, E. S-adenosylmethionine: nothing goes to waste. Trends Biochem. Sci. 29, 243–249 (2004).

    CAS  PubMed  Google Scholar 

  41. 41.

    Lin, H. S-Adenosylmethionine-dependent alkylation reactions: when are radical reactions used? Bioorg. Chem. 39, 161–170 (2011).

    CAS  PubMed  PubMed Central  Google Scholar 

  42. 42.

    Yokoyama, K. & Lilla, E. A. C–C bond forming radical SAM enzymes involved in the construction of carbon skeletons of cofactors and natural products. Nat. Prod. Rep. 35, 660–694 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  43. 43.

    Schubert, H. L., Blumenthal, R. M. & Cheng, X. Many paths to methyltransfer: a chronicle of convergence. Trends Biochem. Sci. 28, 329–335 (2003).

    CAS  PubMed  PubMed Central  Google Scholar 

  44. 44.

    Struck, A.-W., Thompson, M. L., Wong, L. S. & Micklefield, J. S-Adenosyl-methionine-dependent methyltransferases: highly versatile enzymes in biocatalysis, biosynthesis and other biotechnological applications. ChemBioChem 13, 2642–2655 (2012).

    CAS  PubMed  Google Scholar 

  45. 45.

    Klimasauskas, S. & Weinhold, E. A new tool for biotechnology: AdoMet-dependent methyltransferases. Trends Biotechnol. 25, 99–104 (2007).

    CAS  PubMed  Google Scholar 

  46. 46.

    Dalhoff, C., Lukinavičius, G., Klimas;auskas, S. & Weinhold, E. Direct transfer of extended groups from synthetic cofactors by DNA methyltransferases. Nat. Chem. Biol. 2, 31–32 (2006).

    CAS  PubMed  Google Scholar 

  47. 47.

    Motorin, Y. et al. Expanding the chemical scope of RNA: methyltransferases to site-specific alkynylation of RNA for click labeling. Nucleic Acids Res. 39, 1943–1952 (2011).

    CAS  PubMed  Google Scholar 

  48. 48.

    Peters, W. et al. Enzymatic site-specific functionalization of protein methyltransferase substrates with alkynes for click labeling. Angew. Chem. Int. Ed. 49, 5170–5173 (2010).

    CAS  Google Scholar 

  49. 49.

    Deen, J. et al. Methyltransferase-directed labeling of biomolecules and its applications. Angew. Chem. Int. Ed. 56, 5182–5200 (2017).

    CAS  Google Scholar 

  50. 50.

    Stecher, H. et al. Biocatalytic Friedel–Crafts alkylation using non-natural cofactors. Angew. Chem. Int. Ed. 48, 9546–9548 (2009).

    CAS  Google Scholar 

  51. 51.

    Liang, P.-H., Ko, T.-P. & Wang, A. H.-J. Structure, mechanism and function of prenyltransferases. Eur. J. Biochem. 269, 3339–3354 (2002).

    CAS  PubMed  Google Scholar 

  52. 52.

    Tanner, M. E. Mechanistic studies on the indole prenyltransferases. Nat. Prod. Rep. 32, 88–101 (2015).

    CAS  PubMed  Google Scholar 

  53. 53.

    Epifano, F., Genovese, S., Menghini, L. & Curini, M. Chemistry and pharmacology of oxyprenylated secondary plant metabolites. Phytochemistry 68, 939–953 (2007).

    CAS  PubMed  Google Scholar 

  54. 54.

    Bandari, C. et al. FgaPT2, a biocatalytic tool for alkyl-diversification of indole natural products. MedChemComm 10, 1465–1475 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  55. 55.

    Elshahawi, S. I. et al. Structure and specificity of a permissive bacterial C-prenyltransferase. Nat. Chem. Biol. 13, 366–368 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  56. 56.

    Eliot, A. C. & Kirsch, J. F. Pyridoxal phosphate enzymes: mechanistic, structural, and evolutionary considerations. Annu. Rev. Biochem. 73, 383–415 (2004).

    CAS  PubMed  Google Scholar 

  57. 57.

    Fesko, K. Threonine aldolases: perspectives in engineering and screening the enzymes with enhanced substrate and stereo specificities. Appl. Microbiol. Biotechnol. 100, 2579–2590 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  58. 58.

    Chun, S. W. & Narayan, A. R. H. Biocatalytic synthesis of α-amino ketones. Synlett 30, 1269–1274 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  59. 59.

    Chun, S. W., Hinze, M. E., Skiba, M. A. & Narayan, A. R. H. Chemistry of a unique polyketide-like synthase. J. Am. Chem. Soc. 140, 2430–2433 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  60. 60.

    Miles, E. W. Tryptophan synthase: a multienzyme complex with an intramolecular tunnel. Chem. Rec. 1, 140–151 (2001).

    CAS  PubMed  Google Scholar 

  61. 61.

    Francis, D., Winn, M., Latham, J., Greaney, M. F. & Micklefield, J. An engineered tryptophan synthase opens new enzymatic pathways to β-methyltryptophan and derivatives. ChemBioChem 18, 382–386 (2017).

    CAS  PubMed  Google Scholar 

  62. 62.

    Dunn, M. F. Allosteric regulation of substrate channeling and catalysis in the tryptophan synthase bienzyme complex. Arch. Biochem. Biophys. 519, 154–166 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  63. 63.

    Buller, A. R. et al. Directed evolution of the tryptophan synthase β-subunit for stand-alone function recapitulates allosteric activation. Proc. Natl Acad. Sci. USA 112, 14599–14604 (2015).

    CAS  PubMed  Google Scholar 

  64. 64.

    Boville, C. E. et al. Engineered biosynthesis of β-alkyl tryptophan analogues. Angew. Chem. Int. Ed. 57, 14764–14768 (2018).

    CAS  Google Scholar 

  65. 65.

    Boville, C. E., Romney, D. K., Almhjell, P. J., Sieben, M. & Arnold, F. H. Improved synthesis of 4-cyanotryptophan and other tryptophan analogues in aqueous solvent using variants of TrpB from Thermotoga maritima. J. Org. Chem. 83, 7447–7452 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  66. 66.

    Murciano-Calles, J., Romney, D. K., Brinkmann-Chen, S., Buller, A. R. & Arnold, F. H. A panel of TrpB biocatalysts derived from tryptophan synthase through the transfer of mutations that mimic allosteric activation. Angew. Chem. Int. Ed. 55, 11577–11581 (2016).

    CAS  Google Scholar 

  67. 67.

    Dick, M., Sarai, N. S., Martynowycz, M. W., Gonen, T. & Arnold, F. H. Tailoring tryptophan synthase TrpB for selective quaternary carbon bond formation. J. Am. Chem. Soc. 141, 19817–19822 (2019).

    CAS  PubMed  Google Scholar 

  68. 68.

    Romney, D. K., Murciano-Calles, J., Wehrmüller, J. E. & Arnold, F. H. Unlocking reactivity of TrpB: a general biocatalytic platform for synthesis of tryptophan analogues. J. Am. Chem. Soc. 139, 10769–10776 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  69. 69.

    Nakamura, H., Schultz, E. E. & Balskus, E. P. A new strategy for aromatic ring alkylation in cylindrocyclophane biosynthesis. Nat. Chem. Biol. 13, 916–921 (2017).

    CAS  PubMed  Google Scholar 

  70. 70.

    Schultz, E. E., Braffman, N. R., Luescher, M. U., Hager, H. H. & Balskus, E. P. Biocatalytic Friedel–Crafts alkylation using a promiscuous biosynthetic enzyme. Angew. Chem. Int. Ed. 58, 3151–3155 (2019).

    CAS  Google Scholar 

  71. 71.

    Facchini, P. J. Alkaloid biosynthesis in plants: biochemistry, cell biology, molecular regulation, and metabolic engineering applications. Annu. Rev. Plant. Physiol. Plant Mol. Biol. 52, 29–66 (2001).

    CAS  PubMed  Google Scholar 

  72. 72.

    Suzuki, S. & Umezawa, T. Biosynthesis of lignans and norlignans. J. Wood Sci. 53, 273–284 (2007).

    CAS  Google Scholar 

  73. 73.

    Sheng, X. & Himo, F. Enzymatic Pictet–Spengler reaction: computational study of the mechanism and enantioselectivity of norcoclaurine synthase. J. Am. Chem. Soc. 141, 11230–11238 (2019).

    CAS  PubMed  Google Scholar 

  74. 74.

    Patil, M. D., Grogan, G. & Yun, H. Biocatalyzed C−C bond formation for the production of alkaloids. ChemCatChem 10, 4783–4804 (2018).

    Google Scholar 

  75. 75.

    Lichman, B. R., Zhao, J., Hailes, H. C. & Ward, J. M. Enzyme catalysed Pictet–Spengler formation of chiral 1,1ʹ-disubstituted- and spiro-tetrahydroisoquinolines. Nat. Commun. 8, 14883 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  76. 76.

    Sato, F., Inai, K. & Hashimoto, T. in Applications of Plant Metabolic Engineering (eds Verpoorte, R., Alfermann, A. W. & Johnson, T. S.) 145–173 (Springer, 2007).

  77. 77.

    Leonard, E., Runguphan, W., O’Connor, S. & Prather, K. J. Opportunities in metabolic engineering to facilitate scalable alkaloid production. Nat. Chem. Biol. 5, 292–300 (2009).

    CAS  PubMed  Google Scholar 

  78. 78.

    Galanie, S., Thodey, K., Trenchard, I. J., Filsinger Interrante, M. & Smolke, C. D. Complete biosynthesis of opioids in yeast. Science 349, 1095–1100 (2015).

    CAS  PubMed  PubMed Central  Google Scholar 

  79. 79.

    Diamond, A. & Desgagné-Penix, I. Metabolic engineering for the production of plant isoquinoline alkaloids. Plant. Biotechnol. J. 14, 1319–1328 (2016).

    CAS  PubMed  Google Scholar 

  80. 80.

    Thodey, K., Galanie, S. & Smolke, C. D. A microbial biomanufacturing platform for natural and semisynthetic opioids. Nat. Chem. Biol. 10, 837–844 (2014).

    CAS  PubMed  PubMed Central  Google Scholar 

  81. 81.

    Marienhagen, J. & Bott, M. Metabolic engineering of microorganisms for the synthesis of plant natural products. J. Biotechnol. 163, 166–178 (2013).

    CAS  PubMed  Google Scholar 

  82. 82.

    Du, J., Shao, Z. & Zhao, H. Engineering microbial factories for synthesis of value-added products. J. Ind. Microbiol. Biotechnol. 38, 873–890 (2011).

    CAS  PubMed  PubMed Central  Google Scholar 

  83. 83.

    Winkler, A. et al. A concerted mechanism for berberine bridge enzyme. Nat. Chem. Biol. 4, 739–741 (2008).

    CAS  PubMed  Google Scholar 

  84. 84.

    Resch, V. et al. Inverting the regioselectivity of the berberine bridge enzyme by employing customized fluorine-containing substrates. Chem. Eur. J. 18, 13173–13179 (2012).

    CAS  PubMed  Google Scholar 

  85. 85.

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

    CAS  PubMed  PubMed Central  Google Scholar 

  86. 86.

    Chang, W.-c, Yang, Z.-J., Tu, Y.-H. & Chien, T.-C. Reaction mechanism of a nonheme iron enzyme catalyzed oxidative cyclization via C–C bond formation. Org. Lett. 21, 228–232 (2019).

    CAS  PubMed  Google Scholar 

  87. 87.

    Martinez, S. & Hausinger, R. P. Catalytic mechanisms of Fe(II)- and 2-oxoglutarate-dependent oxygenases. J. Biol. Chem. 290, 20702–20711 (2015).

    CAS  PubMed  PubMed Central  Google Scholar 

  88. 88.

    Lazzarotto, M. et al. Chemoenzymatic total synthesis of deoxy-, epi-, and podophyllotoxin and a biocatalytic kinetic resolution of dibenzylbutyrolactones. Angew. Chem. Int. Ed. 58, 8226–8230 (2019).

    CAS  Google Scholar 

  89. 89.

    Li, J., Zhang, X. & Renata, H. Asymmetric chemoenzymatic synthesis of (−)-podophyllotoxin and related aryltetralin lignans. Angew. Chem. Int. Ed. 58, 11657–11660 (2019).

    CAS  Google Scholar 

  90. 90.

    Austin, M. B. & Noel, J. P. The chalcone synthase superfamily of type III polyketide synthases. Nat. Prod. Rep. 20, 79–110 (2003).

    CAS  PubMed  Google Scholar 

  91. 91.

    Hayashi, A. et al. Molecular and catalytic properties of monoacetylphloroglucinol acetyltransferase from Pseudomonas sp. YGJ3. Biosci. Biotechnol. Biochem. 76, 559–566 (2012).

    CAS  PubMed  Google Scholar 

  92. 92.

    Schmidt, N. G. et al. Biocatalytic Friedel–Crafts acylation and fries reaction. Angew. Chem. Int. Ed. 56, 7615–7619 (2017).

    CAS  Google Scholar 

  93. 93.

    Schmidt, N. G. & Kroutil, W. Acyl donors and additives for the biocatalytic Friedel–Crafts acylation. Eur. J. Org. Chem. 2017, 5865–5871 (2017).

    CAS  Google Scholar 

  94. 94.

    Żądło-Dobrowolska, A., Schmidt, N. G. & Kroutil, W. Thioesters as acyl donors in biocatalytic Friedel–Crafts-type acylation catalyzed by acyltransferase from Pseudomonas protegens. ChemCatChem 11, 1064–1068 (2019).

    PubMed  PubMed Central  Google Scholar 

  95. 95.

    Kozlowski, M. C. Oxidative coupling in complexity building transforms. Acc. Chem. Res. 50, 638–643 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  96. 96.

    Kozlowski, M. C., Morgan, B. J. & Linton, E. C. Total synthesis of chiral biaryl natural products by asymmetric biaryl coupling. Chem. Soc. Rev. 38, 3193–3207 (2009).

    CAS  PubMed  PubMed Central  Google Scholar 

  97. 97.

    Bringmann, G., Gulder, T., Gulder, T. A. M. & Breuning, M. Atroposelective total synthesis of axially chiral biaryl natural products. Chem. Rev. 111, 563–639 (2011).

    CAS  PubMed  Google Scholar 

  98. 98.

    Isin, E. M. & Guengerich, F. P. Complex reactions catalyzed by cytochrome P450 enzymes. Biochim. Biophys. Acta. 1770, 314–329 (2007).

    CAS  PubMed  Google Scholar 

  99. 99.

    Denisov, I. G., Makris, T. M., Sligar, S. G. & Schlichting, I. Structure and chemistry of cytochrome P450. Chem. Rev. 105, 2253–2278 (2005).

    CAS  PubMed  Google Scholar 

  100. 100.

    Guengerich, F. P. & Yoshimoto, F. K. Formation and cleavage of C–C bonds by enzymatic oxidation–reduction reactions. Chem. Rev. 118, 6573–6655 (2018).

    CAS  PubMed  Google Scholar 

  101. 101.

    Ikezawa, N., Iwasa, K. & Sato, F. Molecular cloning and characterization of CYP80G2, a cytochrome P450 that catalyzes an intramolecular C–C phenol coupling of (S)-reticuline in magnoflorine biosynthesis, from cultured Coptis japonica cells. J. Biol. Chem. 283, 8810–8821 (2008).

    CAS  PubMed  Google Scholar 

  102. 102.

    Pylypenko, O., Vitali, F., Zerbe, K., Robinson, J. A. & Schlichting, I. Crystal structure of OxyC, a cytochrome P450 implicated in an oxidative C–C coupling reaction during vancomycin biosynthesis. J. Biol. Chem. 278, 46727–46733 (2003).

    CAS  PubMed  Google Scholar 

  103. 103.

    Forneris, C. C. & Seyedsayamdost, M. R. In vitro reconstitution of OxyC activity enables total chemoenzymatic syntheses of vancomycin aglycone variants. Angew. Chem. Int. Ed. 57, 8048–8052 (2018).

    CAS  Google Scholar 

  104. 104.

    Gil Girol, C. et al. Regio- and stereoselective oxidative phenol coupling in Aspergillus niger. Angew. Chem. Int. Ed. 51, 9788–9791 (2012).

    CAS  Google Scholar 

  105. 105.

    Mazzaferro, L. S., Hüttel, W., Fries, A. & Müller, M. Cytochrome P450-catalyzed regio- and stereoselective phenol coupling of fungal natural products. J. Am. Chem. Soc. 137, 12289–12295 (2015).

    CAS  PubMed  Google Scholar 

  106. 106.

    Präg, A. et al. Regio- and stereoselective intermolecular oxidative phenol coupling in Streptomyces. J. Am. Chem. Soc. 136, 6195–6198 (2014).

    PubMed  Google Scholar 

  107. 107.

    Obermaier, S. & Muller, M. Biaryl-forming enzymes from aspergilli exhibit substrate-dependent stereoselectivity. Biochemistry 58, 2589–2593 (2019).

    CAS  PubMed  Google Scholar 

  108. 108.

    Zhao, B. et al. Binding of two flaviolin substrate molecules, oxidative coupling, and crystal structure of Streptomyces coelicolor A3(2) cytochrome P450 158A2. J. Biol. Chem. 280, 11599–11607 (2005).

    CAS  PubMed  Google Scholar 

  109. 109.

    Mate, D. M. & Alcalde, M. Laccase: a multi-purpose biocatalyst at the forefront of biotechnology. Microb. Biotechnol. 10, 1457–1467 (2017).

    CAS  PubMed  Google Scholar 

  110. 110.

    Azevedo, A. M. et al. Horseradish peroxidase: a valuable tool in biotechnology. Biotechnol. Annu. Rev. 9, 1387–2656 (2003).

    Google Scholar 

  111. 111.

    Jones, S. M. & Solomon, E. I. Electron transfer and reaction mechanism of laccases. Cell Mol. Life Sci. 72, 869–883 (2015).

    CAS  PubMed  PubMed Central  Google Scholar 

  112. 112.

    Davin, L. B. et al. Stereoselective bimolecular phenoxy radical coupling by an auxiliary (dirigent) protein without an active center. Science 275, 362–366 (1997).

    CAS  PubMed  Google Scholar 

  113. 113.

    Pickel, B. & Schaller, A. Dirigent proteins: molecular characteristics and potential biotechnological applications. Appl. Microbiol. Biotechnol. 97, 8427–8438 (2013).

    CAS  PubMed  Google Scholar 

  114. 114.

    Gasper, R. et al. Dirigent protein mode of action revealed by the crystal structure of AtDIR6. Plant. Physiol. 172, 2165–2175 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  115. 115.

    Liu, J., Stipanovic, R. D., Bell, A. A., Puckhaber, L. S. & Magill, C. W. Stereoselective coupling of hemigossypol to form (+)-gossypol in moco cotton is mediated by a dirigent protein. Phytochemistry 69, 3038–3042 (2008).

    CAS  PubMed  Google Scholar 

  116. 116.

    Obermaier, S., Thiele, W., Fürtges, L. & Müller, M. Enantioselective phenol coupling by laccases in the biosynthesis of fungal dimeric naphthopyrones. Angew. Chem. Int. Ed. 58, 9125–9128 (2019).

    CAS  Google Scholar 

  117. 117.

    Christianson, D. W. Structural and chemical biology of terpenoid cyclases. Chem. Rev. 117, 11570–11648 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  118. 118.

    Baunach, M., Franke, J. & Hertweck, C. Terpenoid biosynthesis off the beaten track: Unconventional cyclases and their impact on biomimetic synthesis. Angew. Chem. Int. Ed. 54, 2604–2626 (2015).

    CAS  Google Scholar 

  119. 119.

    Pronin, S. V. & Shenvi, R. A. Synthesis of highly strained terpenes by non-stop tail-to-head polycyclization. Nat. Chem. 4, 915–920 (2012).

    CAS  PubMed  Google Scholar 

  120. 120.

    Siedenburg, G. & Jendrossek, D. Squalene–hopene cyclases. Appl. Environ. Microbiol. 77, 3905–3915 (2011).

    CAS  PubMed  PubMed Central  Google Scholar 

  121. 121.

    Syrén, P. O., Henche, S., Eichler, A., Nestl, B. M. & Hauer, B. Squalene–hopene cyclases — evolution, dynamics and catalytic scope. Curr. Opin. Struct. Biol. 41, 73–82 (2016).

    PubMed  Google Scholar 

  122. 122.

    Hoshino, T., Kumai, Y., Kudo, I., Nakano, S.-i. & Ohashi, S. Enzymatic cyclization reactions of geraniol, farnesol and geranylgeraniol, and those of truncated squalene analogs having C20 and C25 by recombinant squalene cyclase. Org. Biomol. Chem. 2, 2650–2657 (2004).

    CAS  PubMed  Google Scholar 

  123. 123.

    Seitz, M. et al. Synthesis of heterocyclic terpenoids by promiscuous squalene–hopene cyclases. ChemBioChem 14, 436–439 (2013).

    CAS  PubMed  Google Scholar 

  124. 124.

    Hammer, S. C., Dominicus, J. M., Syrén, P.-O., Nestl, B. M. & Hauer, B. Stereoselective Friedel–Crafts alkylation catalyzed by squalene hopene cyclases. Tetrahedron 68, 7624–7629 (2012).

    CAS  Google Scholar 

  125. 125.

    Seitz, M. et al. Substrate specificity of a novel squalene–hopene cyclase from Zymomonas mobilis. J. Mol. Catal. B Enzym. 84, 72–77 (2012).

    CAS  Google Scholar 

  126. 126.

    Siedenburg, G. et al. Activation-independent cyclization of monoterpenoids. Appl. Environ. Microbiol. 78, 1055–1062 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  127. 127.

    Hammer, S. C., Marjanovic, A., Dominicus, J. M., Nestl, B. M. & Hauer, B. Squalene hopene cyclases are protonases for stereoselective Brønsted acid catalysis. Nat. Chem. Biol. 11, 121–126 (2015).

    CAS  PubMed  Google Scholar 

  128. 128.

    Degenhardt, J., Kollner, T. G. & Gershenzon, J. Monoterpene and sesquiterpene synthases and the origin of terpene skeletal diversity in plants. Phytochemistry 70, 1621–1637 (2009).

    CAS  PubMed  Google Scholar 

  129. 129.

    Ueda, D., Hoshino, T. & Sato, T. Cyclization of squalene from both termini: identification of an onoceroid synthase and enzymatic synthesis of ambrein. J. Am. Chem. Soc. 135, 18335–18338 (2013).

    CAS  PubMed  Google Scholar 

  130. 130.

    Siegel, J. B. et al. Computational design of an enzyme catalyst for a stereoselective bimolecular Diels–Alder reaction. Science 329, 309–313 (2010).

    CAS  PubMed  PubMed Central  Google Scholar 

  131. 131.

    Klas, K., Tsukamoto, S., Sherman, D. H. & Williams, R. M. Natural Diels–Alderases: elusive and irresistable. J. Org. Chem. 80, 11672–11685 (2015).

    CAS  PubMed  PubMed Central  Google Scholar 

  132. 132.

    Ohashi, M. et al. SAM-dependent enzyme-catalysed pericyclic reactions in natural product biosynthesis. Nature 549, 502–506 (2017).

    PubMed  PubMed Central  Google Scholar 

  133. 133.

    Cai, Y. et al. Structural basis for stereoselective dehydration and hydrogen-bonding catalysis by the SAM-dependent pericyclase LepI. Nat. Chem. 11, 812–820 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  134. 134.

    Dan, Q. et al. Fungal indole alkaloid biogenesis through evolution of a bifunctional reductase/Diels–Alderase. Nat. Chem. 11, 972–980 (2019).

    CAS  PubMed  Google Scholar 

  135. 135.

    Minami, A. & Oikawa, H. Recent advances of Diels–Alderases involved in natural product biosynthesis. J. Antibiot. 69, 500–506 (2016).

    CAS  PubMed  Google Scholar 

  136. 136.

    Kim, R.-R. et al. Mechanistic insights on riboflavin synthase inspired by selective binding of the 6,7-dimethyl-8-ribityllumazine exomethylene anion. J. Am. Chem. Soc. 132, 2983–2990 (2010).

    CAS  PubMed  PubMed Central  Google Scholar 

  137. 137.

    Kohli, R. M. & Massey, V. The oxidative half-reaction of old yellow enzyme. The role of tyrosine 196. J. Biol. Chem. 273, 32763–32770 (1998).

    CAS  PubMed  Google Scholar 

  138. 138.

    Winkler, C. K., Tasnádi, G., Clay, D., Hall, M. & Faber, K. Asymmetric bioreduction of activated alkenes to industrially relevant optically active compounds. J. Biotechnol. 162, 381–389 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  139. 139.

    Toogood, H. S. & Scrutton, N. S. Discovery, characterisation, engineering and applications of ene reductases for industrial biocatalysis. ACS Catal. 8, 3532–3549 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  140. 140.

    Heckenbichler, K. et al. Asymmetric reductive carbocyclization using engineered ene reductases. Angew. Chem. Int. Ed. 57, 7240–7244 (2018).

    CAS  Google Scholar 

  141. 141.

    Biegasiewicz, K. F. et al. Photoexcitation of flavoenzymes enables a stereoselective radical cyclization. Science 364, 1166–1169 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  142. 142.

    Black, M. J. et al. Asymmetric redox-neutral radical cyclization catalysed by flavin-dependent ‘ene’-reductases. Nat. Chem. 12, 71–75 (2020).

    PubMed  Google Scholar 

  143. 143.

    Sandoval, B. A., Meichan, A. J. & Hyster, T. K. Enantioselective hydrogen atom transfer: discovery of catalytic promiscuity in flavin-dependent ‘ene’-reductases. J. Am. Chem. Soc. 139, 11313–11316 (2017).

    CAS  PubMed  Google Scholar 

  144. 144.

    Ye, T. & McKervey, M. A. Organic synthesis with α-diazo carbonyl compounds. Chem. Rev. 94, 1091–1160 (1994).

    CAS  Google Scholar 

  145. 145.

    Doyle, M. P. & Forbes, D. C. Recent advances in asymmetric catalytic metal carbene transformations. Chem. Rev. 98, 911–936 (1998).

    CAS  PubMed  Google Scholar 

  146. 146.

    Davies, H. M. & Manning, J. R. Catalytic C–H functionalization by metal carbenoid and nitrenoid insertion. Nature 451, 417–424 (2008).

    CAS  PubMed  PubMed Central  Google Scholar 

  147. 147.

    Zhang, Y. & Wang, J. Recent development of reactions with α-diazocarbonyl compounds as nucleophiles. Chem. Commun. 5350–5361 (2009).

  148. 148.

    Ford, A. et al. Modern organic synthesis with α-diazocarbonyl compounds. Chem. Rev. 115, 9981–10080 (2015).

    CAS  PubMed  Google Scholar 

  149. 149.

    Franssen, N. M. G., Walters, A. J. C., Reek, J. N. H. & de Bruin, B. Carbene insertion into transition metal–carbon bonds: A new tool for catalytic C–C bond formation. Catal. Sci. Technol. 1, 153–165 (2011).

    CAS  Google Scholar 

  150. 150.

    Vaz, A. D. N., McGinnity, D. F. & Coon, M. J. Epoxidation of olefins by cytochrome P450: evidence from site-specific mutagenesis for hydroperoxo-iron as an electrophilic oxidant. Proc. Natl Acad. Sci. USA 95, 3555–3560 (1998).

    CAS  PubMed  Google Scholar 

  151. 151.

    Coelho, P. S., Brustad, E. M., Kannan, A. & Arnold, F. H. Olefin cyclopropanation via carbene transfer catalyzed by engineered cytochrome P450 enzymes. Science 339, 307–310 (2013). This groundbreaking study described one of the first examples of a new-to-nature transformation catalysed by enzymes.

    CAS  PubMed  Google Scholar 

  152. 152.

    Brandenberg, O. F., Fasan, R. & Arnold, F. H. Exploiting and engineering hemoproteins for abiological carbene and nitrene transfer reactions. Curr. Opin. Biotechnol. 47, 102–111 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  153. 153.

    Coelho, P. S. et al. A serine-substituted P450 catalyzes highly efficient carbene transfer to olefins in vivo. Nat. Chem. Biol. 9, 485–487 (2013).

    CAS  PubMed  PubMed Central  Google Scholar 

  154. 154.

    Heel, T., McIntosh, J. A., Dodani, S. C., Meyerowitz, J. T. & Arnold, F. H. Non-natural olefin cyclopropanation catalyzed by diverse cytochrome P450s and other hemoproteins. ChemBioChem 15, 2556–2562 (2014).

    CAS  PubMed  PubMed Central  Google Scholar 

  155. 155.

    Renata, H. et al. Identification of mechanism-based inactivation in P450-catalyzed cyclopropanation facilitates engineering of improved enzymes. J. Am. Chem. Soc. 138, 12527–12533 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  156. 156.

    Bordeaux, M., Tyagi, V. & Fasan, R. Highly diastereoselective and enantioselective olefin cyclopropanation using engineered myoglobin-based catalysts. Angew. Chem. Int. Ed. 54, 1744–1748 (2015).

    CAS  Google Scholar 

  157. 157.

    Sreenilayam, G., Moore, E. J., Steck, V. & Fasan, R. Stereoselective olefin cyclopropanation under aerobic conditions with an artificial enzyme incorporating an iron-chlorin e6 cofactor. ACS Catal. 7, 7629–7633 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  158. 158.

    Key, H. M. et al. Beyond iron: iridium-containing P450 enzymes for selective cyclopropanations of structurally diverse alkenes. ACS Cent. Sci. 3, 302–308 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  159. 159.

    Brandenberg, O. F. et al. Stereoselective enzymatic synthesis of heteroatom-substituted cyclopropanes. ACS Catal. 8, 2629–2634 (2018).

    CAS  Google Scholar 

  160. 160.

    Knight, A. M. et al. Diverse engineered heme proteins enable stereodivergent cyclopropanation of unactivated alkenes. ACS Cent. Sci. 4, 372–377 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  161. 161.

    Carminati, D. M. & Fasan, R. Stereoselective cyclopropanation of electron-deficient olefins with a cofactor redesigned carbene transferase featuring radical reactivity. ACS Catal. 9, 9683–9697 (2019).

    CAS  PubMed  Google Scholar 

  162. 162.

    Tinoco, A., Steck, V., Tyagi, V. & Fasan, R. Highly diastereo- and enantioselective synthesis of trifluoromethyl-substituted cyclopropanes via myoglobin-catalyzed transfer of trifluoromethylcarbene. J. Am. Chem. Soc. 139, 5293–5296 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  163. 163.

    Chandgude, A. L. & Fasan, R. Highly diastereo- and enantioselective synthesis of nitrile-substituted cyclopropanes by myoglobin-mediated carbene transfer catalysis. Angew. Chem. Int. Ed. 57, 15852–15856 (2018).

    CAS  Google Scholar 

  164. 164.

    Chandgude, A. L., Ren, X. & Fasan, R. Stereodivergent intramolecular cyclopropanation enabled by engineered carbene transferases. J. Am. Chem. Soc. 141, 9145–9150 (2019).

    PubMed  PubMed Central  Google Scholar 

  165. 165.

    Chen, K., Huang, X., Kan, S. B. J., Zhang, R. K. & Arnold, F. H. Enzymatic construction of highly strained carbocycles. Science 360, 71–75 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  166. 166.

    Bajaj, P., Sreenilayam, G., Tyagi, V. & Fasan, R. Gram-scale synthesis of chiral cyclopropane-containing drugs and drug precursors with engineered myoglobin catalysts featuring complementary stereoselectivity. Angew. Chem. Int. Ed. 55, 16110–16114 (2016).

    CAS  Google Scholar 

  167. 167.

    Wang, Z. J. et al. Improved cyclopropanation activity of histidine-ligated cytochrome P450 enables the enantioselective formal synthesis of levomilnacipran. Angew. Chem. Int. Ed. 53, 6810–6813 (2014).

    CAS  Google Scholar 

  168. 168.

    Key, H. M., Dydio, P., Clark, D. S. & Hartwig, J. F. Abiological catalysis by artificial haem proteins containing noble metals in place of iron. Nature 534, 534–537 (2016).

    CAS  PubMed  Google Scholar 

  169. 169.

    Dydio, P. et al. An artificial metalloenzyme with the kinetics of native enzymes. Science 354, 102–106 (2016).

    CAS  PubMed  Google Scholar 

  170. 170.

    Sreenilayam, G., Moore, E. J., Steck, V. & Fasan, R. Metal substitution modulates the reactivity and extends the reaction scope of myoglobin carbene transfer catalysts. Adv. Synth. Catal. 359, 2076–2089 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  171. 171.

    Zhang, R. K. et al. Enzymatic assembly of carbon–carbon bonds via iron-catalysed sp 3 C–H functionalization. Nature 565, 67–72 (2019).

    CAS  PubMed  Google Scholar 

  172. 172.

    Vargas, D. A., Tinoco, A., Tyagi, V. & Fasan, R. Myoglobin-catalyzed C−H functionalization of unprotected indoles. Angew. Chem. Int. Ed. 57, 9911–9915 (2018).

    CAS  Google Scholar 

  173. 173.

    Brandenberg, O. F., Chen, K. & Arnold, F. H. Directed evolution of a cytochrome P450 carbene transferase for selective functionalization of cyclic compounds. J. Am. Chem. Soc. 141, 8989–8995 (2019).

    CAS  PubMed  Google Scholar 

  174. 174.

    Zhang, J., Huang, X., Zhang, R. K. & Arnold, F. H. Enantiodivergent α-amino C–H fluoroalkylation catalyzed by engineered cytochrome P450s. J. Am. Chem. Soc. 141, 9798–9802 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  175. 175.

    Tyagi, V., Sreenilayam, G., Bajaj, P., Tinoco, A. & Fasan, R. Biocatalytic synthesis of allylic and allenyl sulfides through a myoglobin-catalyzed Doyle–Kirmse reaction. Angew. Chem. Int. Ed. 55, 13562–13566 (2016).

    CAS  Google Scholar 

  176. 176.

    Tyagi, V. & Fasan, R. Myoglobin-catalyzed olefination of aldehydes. Angew. Chem. Int. Ed. 55, 2512–2516 (2016).

    CAS  Google Scholar 

  177. 177.

    Weissenborn, M. J. et al. Enzyme-catalyzed carbonyl olefination by the E. coli protein YfeX in the absence of phosphines. ChemCatChem 8, 1636–1640 (2016).

    CAS  Google Scholar 

  178. 178.

    Clouthier, C. M. & Pelletier, J. N. Expanding the organic toolbox: a guide to integrating biocatalysis in synthesis. Chem. Soc. Rev. 41, 1585–1605 (2012).

    CAS  PubMed  Google Scholar 

  179. 179.

    Sheldon, R. A. & Brady, D. Broadening the scope of biocatalysis in sustainable organic synthesis. ChemSusChem 12, 2859–2881 (2019).

    CAS  PubMed  Google Scholar 

  180. 180.

    Sheldon, R. A., Brady, D. & Bode, M. L. The Hitchhiker’s guide to biocatalysis: recent advances in the use of enzymes in organic synthesis. Chem. Sci. 11, 2587–2605 (2020). This review serves an excellent primer on biocatalysis for synthetic chemists.

    PubMed  PubMed Central  Google Scholar 

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Acknowledgements

The authors are grateful for support from the National Institutes of Health (R35 GM124880 and T32 GM008353).

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L.E.Z. and A.R.H.N. contributed to the writing and editing of the article.

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Glossary

Lyases

A class of enzymes characterized by the ability to break or form a new chemical bond using mechanisms other than hydrolysis or oxidation.

Atroposelective

A preference for the formation of a specific stereoisomer (atropisomer) whose axis of chirality results from hindered rotation around a single bond that results in only one of two possible stable conformations.

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Zetzsche, L.E., Narayan, A.R.H. Broadening the scope of biocatalytic C–C bond formation. Nat Rev Chem 4, 334–346 (2020). https://doi.org/10.1038/s41570-020-0191-2

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