Skip to main content

Thank you for visiting nature.com. You are using a browser version with limited support for CSS. To obtain the best experience, we recommend you use a more up to date browser (or turn off compatibility mode in Internet Explorer). In the meantime, to ensure continued support, we are displaying the site without styles and JavaScript.

  • Review Article
  • Published:

Enzymatic degradation of plant biomass and synthetic polymers

Nature Reviews Chemistry volume 4, pages 114–126 (2020)Cite this article

Subjects

Abstract

Plant biomass is an abundant renewable resource on Earth. Microorganisms harvest energy from plant material by means of complex enzymatic systems that efficiently degrade natural polymers. Intriguingly, microorganisms have evolved to exploit these ancient mechanisms to also decompose synthetic plastic polymers. In this Review, we summarize the mechanisms by which they decompose non-starch plant biomass and the six major types of synthetic plastics. We focus on the structural features of the enzymes that contribute to substrate recognition and then describe the catalytic mechanisms of polymer metabolism. An understanding of these natural biocatalysts is valuable if we are to exploit their potential for the degradation of synthetic polymers.

This is a preview of subscription content, access via your institution

Access options

Buy this article

  • Purchase on Springer Link
  • Instant access to full article PDF
Buy now

Prices may be subject to local taxes which are calculated during checkout

Fig. 1: Cutin and rubber structures and their enzymatic degradation.
Fig. 2: The structure of lignin and its enzymatic degradation.
Fig. 3: The structures of polysaccharide-based plant polymers.
Fig. 4: Glycoside hydrolases and polysaccharide lyases cleave sugars with acid–base pairs.
Fig. 5: Lytic polysaccharide monooxygenases cleave sugars oxidatively.
Fig. 6: Backbone structures of the most widely produced synthetic polymers.
Fig. 7: Hydrolytic reaction and structural feature of IsPETase.

Similar content being viewed by others

References

  1. Couto, S. R. & Herrera, J. L. T. Industrial and biotechnological applications of laccases: a review. Biotechnol. Adv. 24, 500–513 (2006).

    Article  CAS  Google Scholar 

  2. Pio, T. F. & Macedo, G. A. Cutinases: properties and industrial applications. Adv. Appl. Microbiol. 66, 77–95 (2009).

    Article  CAS  PubMed  Google Scholar 

  3. Mäkelä, M. R. et al. Fungal ligninolytic enzymes and their applications. Microbiol. Spectr. https://doi.org/10.1128/microbiolspec.FUNK-0017-2016 (2016).

  4. López-Mondéjar, R., Algora, C. & Baldrian, P. Lignocellulolytic systems of soil bacteria: a vast and diverse toolbox for biotechnological conversion processes. Biotechnol. Adv. 37, 107374–107396 (2019).

    Article  CAS  PubMed  Google Scholar 

  5. Lopes, A. M., Ferreira Filho, E. X. & Moreira, L. R. S. An update on enzymatic cocktails for lignocellulose breakdown. J. Appl. Microbiol. 125, 632–645 (2018).

    Article  CAS  PubMed  Google Scholar 

  6. Geyer, R., Jambeck, J. R. & Law, K. L. Production, use, and fate of all plastics ever made. Sci. Adv. 3, e1700782 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  7. Zheng, Y., Yanful, E. K. & Bassi, A. S. A review of plastic waste biodegradation. Crit. Rev. Biotechnol. 25, 243–250 (2008).

    Article  CAS  Google Scholar 

  8. Fich, E. A., Segerson, N. A. & Rose, J. K. C. The plant polyester cutin: biosynthesis, structure, and biological roles. Annu. Rev. Plant. Biol. 67, 207–233 (2016).

    Article  CAS  PubMed  Google Scholar 

  9. Roth, C. et al. Structural and functional studies on a thermostable polyethylene terephthalate degrading hydrolase from Thermobifida fusca. Appl. Microbiol. Biotechnol. 98, 7815–7823 (2014).

    Article  CAS  PubMed  Google Scholar 

  10. Egmond, M. R. & de Vlieg, J. Fusarium solani pisi cutinase. Biochimie 82, 1015–1021 (2000).

    Article  CAS  PubMed  Google Scholar 

  11. Holmquist, M. Alpha/beta-hydrolase fold enzymes: structures, functions and mechanisms. Curr. Protein Pept. Sci. 1, 209–235 (2000).

    Article  CAS  PubMed  Google Scholar 

  12. Ramos, M. V., Demarco, D., da Costa Souza, I. C. & de Freitas, C. D. T. Laticifers, latex, and their role in plant defense. Trends Plant. Sci. 24, 553–567 (2019).

    Article  CAS  PubMed  Google Scholar 

  13. Rose, K. & Steinbüchel, A. Biodegradation of natural rubber and related compounds: recent insights into a hardly understood catabolic capability of microorganisms. Appl. Environ. Microbiol. 71, 2803–2812 (2005).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  14. Hiessl, S. et al. Involvement of two latex-clearing proteins during rubber degradation and insights into the subsequent degradation pathway revealed by the genome sequence of Gordonia polyisoprenivorans strain VH2. Appl. Environ. Microbiol. 78, 2874–2887 (2012).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  15. Tsuchii, A. & Takeda, K. Rubber-degrading enzyme from a bacterial culture. Appl. Environ. Microbiol. 56, 269–274 (1990).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  16. Rose, K., Tenberge, K. B. & Steinbüchel, A. Identification and characterization of genes from Streptomyces sp. strain K30 responsible for clear zone formation on natural rubber latex and poly(cis-1,4-isoprene) rubber degradation. Biomacromolecules 6, 180–188 (2005).

    Article  CAS  PubMed  Google Scholar 

  17. Kasai, D. et al. Identification of natural rubber degradation gene in Rhizobacter gummiphilus NS21. Biosci. Biotechnol. Biochem. 81, 614–620 (2017).

    Article  CAS  PubMed  Google Scholar 

  18. Braaz, R., Armbruster, W. & Jendrossek, D. Heme-dependent rubber oxygenase RoxA of Xanthomonas sp. cleaves the carbon backbone of poly(cis-1,4-isoprene) by a dioxygenase mechanism. Appl. Environ. Microbiol. 71, 2473–2478 (2005).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  19. Seidel, J., Schmitt, G., Hoffmann, M., Jendrossek, D. & Einsle, O. Structure of the processive rubber oxygenase RoxA from Xanthomonas sp. Proc. Natl Acad. Sci. USA 110, 13833–13838 (2013).

    Article  PubMed  PubMed Central  Google Scholar 

  20. Schmitt, G., Seiffert, G., Kroneck, P. M., Braaz, R. & Jendrossek, D. Spectroscopic properties of rubber oxygenase RoxA from Xanthomonas sp., a new type of dihaem dioxygenase. Microbiology 156, 2537–2548 (2010).

    Article  CAS  PubMed  Google Scholar 

  21. Birke, J., Röther, W. & Jendrossek, D. RoxB is a novel type of rubber oxygenase that combines properties of rubber oxygenase RoxA and latex clearing protein (Lcp). Appl. Environ. Microbiol. 83, e00721-17 (2017).

    Article  PubMed  PubMed Central  Google Scholar 

  22. Birke, J., Röther, W. & Jendrossek, D. Latex clearing protein (Lcp) of Streptomyces sp. strain K30 is a b-type cytochrome and differs from rubber oxygenase A (RoxA) in its biophysical properties. Appl. Environ. Microbiol. 81, 3793–3799 (2015).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  23. Röther, W., Austen, S., Birke, J. & Jendrossek, D. Cleavage of rubber by the latex clearing protein (Lcp) of Streptomyces sp. strain K30: molecular insights. Appl. Environ. Microbiol. 82, 6593–6602 (2016).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  24. Ilcu, L. et al. Structural and functional analysis of latex clearing protein (Lcp) provides insight into the enzymatic cleavage of rubber. Sci. Rep. 7, 6179 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  25. Birke, J., Röther, W. & Jendrossek, D. Rhizobacter gummiphilus NS21 has two rubber oxygenases (RoxA and RoxB) acting synergistically in rubber utilisation. Appl. Microbiol. Biotechnol. 102, 10245–10257 (2018).

    Article  CAS  PubMed  Google Scholar 

  26. Rajan, V. V., Dierkes, W. K., Joseph, R. & Noordermeer, J. W. M. Science and technology of rubber reclamation with special attention to NR-based waste latex products. Prog. Polym. Sci. 31, 811–834 (2006).

    Article  CAS  Google Scholar 

  27. Holst, O., Stenberg, B. & Christiansson, M. Biotechnological possibilities for waste tyre-rubber treatment. Biodegradation 9, 301–310 (1998).

    Article  CAS  PubMed  Google Scholar 

  28. Bredberg, K., Persson, J., Christiansson, M., Stenberg, B. & Holst, O. Anaerobic desulfurization of ground rubber with the thermophilic archaeon Pyrococcus furiosus — a new method for rubber recycling. Appl. Microbiol. Biotechnol. 55, 43–48 (2001).

    Article  CAS  PubMed  Google Scholar 

  29. Li, Y., Zhao, S. & Wang, Y. Microbial desulfurization of ground tire rubber by Sphingomonas sp.: a novel technology for crumb rubber composites. J. Polym. Environ. 20, 372–380 (2012).

    Article  CAS  Google Scholar 

  30. Romine, R. A. & Romine, M. F. Rubbercycle: a bioprocess for surface modification of waste tyre rubber. Polym. Degrad. Stab. 59, 353–358 (1998).

    Article  CAS  Google Scholar 

  31. Sato, S. et al. Microbial scission of sulfide linkages in vulcanized natural rubber by a white rot basidiomycete, Ceriporiopsis subvermispora. Biomacromolecules 5, 511–515 (2004).

    Article  CAS  PubMed  Google Scholar 

  32. Sato, S. et al. Degradation of sulfide linkages between isoprenes by lipid peroxidation catalyzed by manganese peroxidase. Chemosphere 77, 798–804 (2009).

    Article  CAS  PubMed  Google Scholar 

  33. Boerjan, W., Ralph, J. & Baucher, M. Lignin biosynthesis. Annu. Rev. Plant. Biol. 54, 519–546 (2003).

    Article  CAS  PubMed  Google Scholar 

  34. Brown, M. E. & Chang, M. C. Exploring bacterial lignin degradation. Curr. Opin. Chem. Biol. 19, 1–7 (2014).

    Article  CAS  PubMed  Google Scholar 

  35. Manavalan, T., Manavalan, A. & Heese, K. Characterization of lignocellulolytic enzymes from white-rot fungi. Curr. Microbiol. 70, 485–498 (2015).

    Article  CAS  PubMed  Google Scholar 

  36. Piontek, K., Antorini, M. & Choinowski, T. Crystal structure of a laccase from the fungus Trametes versicolor at 1.90-Å resolution containing a full complement of coppers. J. Biol. Chem. 277, 37663–37669 (2002).

    Article  CAS  PubMed  Google Scholar 

  37. Solomon, E. I., Sundaram, U. M. & Machonkin, T. E. Multicopper oxidases and oxygenases. Chem. Rev. 96, 2563–2606 (1996).

    Article  CAS  PubMed  Google Scholar 

  38. Matera, I. et al. Crystal structure of the blue multicopper oxidase from the white-rot fungus Trametes trogii complexed with p-toluate. Inorg. Chim. Acta 361, 4129–4137 (2008).

    Article  CAS  Google Scholar 

  39. Solomon, E. I., Augustine, A. J. & Yoon, J. O2 reduction to H2O by the multicopper oxidases. Dalton Trans. 30, 3921–3932 (2008).

    Article  CAS  Google Scholar 

  40. Santhanam, N., Vivanco, J. M., Decker, S. R. & Reardon, K. F. Expression of industrially relevant laccases: prokaryotic style. Trends Biotechnol. 29, 480–489 (2011).

    Article  CAS  PubMed  Google Scholar 

  41. Hilgers, R., Vincken, J.-P., Gruppen, H. & Kabel, M. A. Laccase/mediator systems: their reactivity toward phenolic lignin structures. ACS Sustain. Chem. Eng. 6, 2037–2046 (2018).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  42. Hofrichter, M., Ullrich, R., Pecyna, M. J., Liers, C. & Lundell, T. New and classic families of secreted fungal heme peroxidases. Appl. Microbiol. Biotechnol. 87, 871–897 (2010).

    Article  CAS  PubMed  Google Scholar 

  43. Blodig, W., Smith, A. T., Doyle, W. A. & Piontek, K. Crystal structures of pristine and oxidatively processed lignin peroxidase expressed in Escherichia coli and of the W171F variant that eliminates the redox active tryptophan 171. Implications for the reaction mechanism. J. Mol. Biol. 305, 851–861 (2001).

    Article  CAS  PubMed  Google Scholar 

  44. Doyle, W. A., Blodig, W., Veitch, N. C., Piontek, K. & Smith, A. T. Two substrate interaction sites in lignin peroxidase revealed by site-directed mutagenesis. Biochemistry 37, 15097–15105 (1998).

    Article  CAS  PubMed  Google Scholar 

  45. Pérez-Boada, M. et al. Versatile peroxidase oxidation of high redox potential aromatic compounds: site-directed mutagenesis, spectroscopic and crystallographic investigation of three long-range electron transfer pathways. J. Mol. Biol. 354, 385–402 (2005).

    Article  CAS  PubMed  Google Scholar 

  46. Ruiz-Dueñas, F. J. et al. Substrate oxidation sites in versatile peroxidase and other basidiomycete peroxidases. J. Exp. Bot. 60, 441–452 (2009).

    Article  CAS  PubMed  Google Scholar 

  47. Hofrichter, M. Review: lignin conversion by manganese peroxidase (MnP). Enzyme Microb. Technol. 30, 454–466 (2002).

    Article  CAS  Google Scholar 

  48. Husain, Q. & Qayyum, S. Biological and enzymatic treatment of bisphenol A and other endocrine disrupting compounds: a review. Crit. Rev. Biotechnol. 33, 260–292 (2013).

    Article  CAS  PubMed  Google Scholar 

  49. Masran, R. et al. Harnessing the potential of ligninolytic enzymes for lignocellulosic biomass pretreatment. Appl. Microbiol. Biotechnol. 100, 5231–5246 (2016).

    Article  CAS  PubMed  Google Scholar 

  50. Wang, X., Yao, B. & Su, X. Linking enzymatic oxidative degradation of lignin to organics detoxification. Int. J. Mol. Sci. 19, 3373 (2018).

    Article  CAS  PubMed Central  Google Scholar 

  51. Burton, R. A., Gidley, M. J. & Fincher, G. B. Heterogeneity in the chemistry, structure and function of plant cell walls. Nat. Chem. Biol. 6, 724–732 (2010).

    Article  CAS  PubMed  Google Scholar 

  52. Scheller, H. V. & Ulvskov, P. Hemicelluloses. Annu. Rev. Plant. Biol. 61, 263–289 (2010).

    Article  CAS  PubMed  Google Scholar 

  53. Atmodjo, M. A., Hao, Z. & Mohnen, D. Evolving views of pectin biosynthesis. Annu. Rev. Plant. Biol. 64, 747–779 (2013).

    Article  CAS  PubMed  Google Scholar 

  54. Mohnen, D. Pectin structure and biosynthesis. Curr. Opin. Plant. Biol. 11, 266–277 (2008).

    Article  CAS  PubMed  Google Scholar 

  55. Cantarel, B. L. et al. The carbohydrate-active enzymes database (CAZy): an expert resource for glycogenomics. Nucleic Acids Res. 37, D233–D238 (2009).

    Article  CAS  PubMed  Google Scholar 

  56. Biely, P., Singh, S. & Puchart, V. Towards enzymatic breakdown of complex plant xylan structures: state of the art. Biotechnol. Adv. 34, 1260–1274 (2016).

    Article  CAS  PubMed  Google Scholar 

  57. Hemsworth, G. R., Déjean, G., Davies, G. J. & Brumer, H. Learning from microbial strategies for polysaccharide degradation. Biochem. Soc. Trans. 44, 94–108 (2016).

    Article  CAS  PubMed  Google Scholar 

  58. Moreira, L. R. S. & Filho, E. X. F. An overview of mannan structure and mannan-degrading enzyme systems. Appl. Microbiol. Biotechnol. 79, 165–178 (2008).

    Article  CAS  PubMed  Google Scholar 

  59. Patidar, M. K., Nighojkar, S., Kumar, A. & Nighojkar, A. Pectinolytic enzymes-solid state fermentation, assay methods and applications in fruit juice industries: a review. 3 Biotech 8, 199 (2018).

    Article  PubMed  PubMed Central  Google Scholar 

  60. Koshland Jr, D. E. Stereochemistry and the mechanism of enzymatic reactions. Biol. Rev. 28, 416–436 (1953).

    Article  Google Scholar 

  61. Chen, Z. et al. Tracing determinants of dual substrate specificity in glycoside hydrolase family 5. J. Biol. Chem. 287, 25335–25343 (2012).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  62. Wu, T.-H. et al. Diverse substrate recognition mechanism revealed by Thermotoga maritima Cel5A structures in complex with cellotetraose, cellobiose and mannotriose. Biochim. Biophys. Acta 1814, 1832–1840 (2011).

    Article  CAS  PubMed  Google Scholar 

  63. Yuan, S. F. et al. Biochemical characterization and structural analysis of a bifunctional cellulase/xylanase from Clostridium thermocellum. J. Biol. Chem. 290, 5739–5748 (2015).

    Article  CAS  PubMed  Google Scholar 

  64. Abbott, D. W. & Boraston, A. B. A family 2 pectate lyase displays a rare fold and transition metal-assisted β-elimination. J. Biol. Chem. 282, 35328–35336 (2007).

    Article  CAS  PubMed  Google Scholar 

  65. Charnock, S. J., Brown, I. E., Turkenburg, J. P., Black, G. W. & Davies, G. J. Convergent evolution sheds light on the anti-β-elimination mechanism common to family 1 and 10 polysaccharide lyases. Proc. Natl Acad. Sci. USA 99, 12067–12072 (2002).

    Article  CAS  PubMed  Google Scholar 

  66. Biely, P., Benen, J., Heinrichová, K., Kester, H. C. & Visser, J. Inversion of configuration during hydrolysis of α-1,4-galacturonidic linkage by three Aspergillus polygalacturonases. FEBS Lett. 382, 249–255 (1996).

    Article  CAS  PubMed  Google Scholar 

  67. Shimizu, T., Nakatsu, T., Miyairi, K., Okuno, T. & Kato, H. Active-site architecture of endopolygalacturonase I from Stereum purpureum revealed by crystal structures in native and ligand-bound forms at atomic resolution. Biochemistry 41, 6651–6659 (2002).

    Article  CAS  PubMed  Google Scholar 

  68. Mayans, O. et al. Two crystal structures of pectin lyase A from Aspergillus reveal a pH driven conformational change and striking divergence in the substrate-binding clefts of pectin and pectate lyases. Structure 5, 677–689 (1997).

    Article  CAS  PubMed  Google Scholar 

  69. Vitali, J., Schick, B., Kester, H. C., Visser, J. & Jurnak, F. The three-dimensional structure of Aspergillus niger pectin lyase B at 1.7-Å resolution. Plant. Physiol. 116, 69–80 (1998).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  70. van Alebeek, G. J., Christensen, T. M., Schols, H. A., Mikkelsen, J. D. & Voragen, A. G. Mode of action of pectin lyase A of Aspergillus niger on differently C(6)-substituted oligogalacturonides. J. Biol. Chem. 277, 25929–25936 (2002).

    Article  CAS  PubMed  Google Scholar 

  71. Yadav, S., Yadav, P. K., Yadav, D. & Yadav, K. D. S. Pectin lyase: a review. Process. Biochem. 44, 1–10 (2009).

    Article  CAS  Google Scholar 

  72. Lynd, L. R., Weimer, P. J., van Zyl, W. H. & Pretorius, I. S. Microbial cellulose utilization: fundamentals and biotechnology. Microbiol. Mol. Biol. Rev. 66, 506–577 (2002).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  73. Himmel, M. E. et al. Biomass recalcitrance: engineering plants and enzymes for biofuels production. Science 315, 804–807 (2007).

    Article  CAS  PubMed  Google Scholar 

  74. Horn, S. J., Vaaje-Kolstad, G., Westereng, B. & Eijsink, V. G. H. Novel enzymes for the degradation of cellulose. Biotechnol. Biofuels 5, 45 (2012).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  75. Levasseur, A., Drula, E., Lombard, V., Coutinho, P. M. & Henrissat, B. Expansion of the enzymatic repertoire of the CAZy database to integrate auxiliary redox enzymes. Biotechnol. Biofuels 6, 41 (2013).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  76. Harris, P. V. et al. Stimulation of lignocellulosic biomass hydrolysis by proteins of glycoside hydrolase family 61: structure and function of a large, enigmatic family. Biochemistry 49, 3305–3316 (2010).

    Article  CAS  PubMed  Google Scholar 

  77. Vaaje-Kolstad, G. et al. An oxidative enzyme boosting the enzymatic conversion of recalcitrant polysaccharides. Science 330, 219–222 (2010).

    Article  CAS  PubMed  Google Scholar 

  78. Song, B. et al. Real-time imaging reveals that lytic polysaccharide monooxygenase promotes cellulase activity by increasing cellulose accessibility. Biotechnol. Biofuels 11, 41 (2018).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  79. Quinlan, R. J. et al. Insights into the oxidative degradation of cellulose by a copper metalloenzyme that exploits biomass components. Proc. Natl Acad. Sci. USA 108, 15079–15084 (2011).

    Article  PubMed  PubMed Central  Google Scholar 

  80. Frandsen, K. E. H. et al. The molecular basis of polysaccharide cleavage by lytic polysaccharide monooxygenases. Nat. Chem. Biol. 12, 298–303 (2016).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  81. Vu, V. V., Beeson, W. T., Phillips, C. M., Cate, J. H. & Marletta, M. A. Determinants of regioselective hydroxylation in the fungal polysaccharide monooxygenases. J. Am. Chem. Soc. 136, 562–565 (2014).

    Article  CAS  PubMed  Google Scholar 

  82. Beeson, W. T., Phillips, C. M., Cate, J. H. D. & Marletta, M. A. Oxidative cleavage of cellulose by fungal copper-dependent polysaccharide monooxygenases. J. Am. Chem. Soc. 134, 890–892 (2012).

    Article  CAS  PubMed  Google Scholar 

  83. Phillips, C. M., Beeson, W. T. IV, Cate, J. H. & Marletta, M. A. Cellobiose dehydrogenase and a copper-dependent polysaccharide monooxygenase potentiate cellulose degradation by Neurospora crassa. ACS Chem. Biol. 6, 1399–1406 (2011).

    Article  CAS  PubMed  Google Scholar 

  84. Danneels, B., Tanghe, M. & Desmet, T. Structural features on the substrate-binding surface of fungal lytic polysaccharide monooxygenases determine their oxidative regioselectivity. Biotechnol. J. 14, e1800211 (2019).

    Article  CAS  PubMed  Google Scholar 

  85. Forsberg, Z. et al. Structural determinants of bacterial lytic polysaccharide monooxygenase functionality. J. Biol. Chem. 293, 1397–1412 (2018).

    Article  CAS  PubMed  Google Scholar 

  86. Vaaje-Kolstad, G., Forsberg, Z., Loose, J. S. M., Bissaro, B. & Eijsink, V. G. H. Structural diversity of lytic polysaccharide monooxygenases. Curr. Opin. Struct. Biol. 44, 67–76 (2017).

    Article  CAS  PubMed  Google Scholar 

  87. Leggio, L. L. et al. Structure and boosting activity of a starch-degrading lytic polysaccharide monooxygenase. Nat. Commun. 6, 5961-5969 (2015).

    Article  CAS  PubMed  Google Scholar 

  88. Vu, V. V., Beeson, W. T., Span, E. A., Farquhar, E. R. & Marletta, M. A. A family of starch-active polysaccharide monooxygenases. Proc. Natl Acad. Sci. USA 111, 13822–13827 (2014).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  89. Frommhagen, M. et al. Discovery of the combined oxidative cleavage of plant xylan and cellulose by a new fungal polysaccharide monooxygenase. Biotechnol. Biofuels 8, 101 (2015).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  90. Couturier, M. et al. Lytic xylan oxidases from wood-decay fungi unlock biomass degradation. Nat. Chem. Biol. 14, 306–310 (2018).

    Article  CAS  PubMed  Google Scholar 

  91. Simmons, T. J. et al. Structural and electronic determinants of lytic polysaccharide monooxygenase reactivity on polysaccharide substrates. Nat. Commun. 8, 1064 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  92. Bennati-Granier, C. et al. Substrate specificity and regioselectivity of fungal AA9 lytic polysaccharide monooxygenases secreted by Podospora anserina. Biotechnol. Biofuels 8, 90–103 (2015).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  93. Agger, J. W. et al. Discovery of LPMO activity on hemicelluloses shows the importance of oxidative processes in plant cell wall degradation. Proc. Natl Acad. Sci. USA 111, 6287–6292 (2014).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  94. Vu, V. V. & Marletta, M. A. Starch-degrading polysaccharide monooxygenases. Cell. Mol. Life Sci. 73, 2809–2819 (2016).

    Article  CAS  PubMed  Google Scholar 

  95. Raddadi, N. & Fava, F. Biodegradation of oil-based plastics in the environment: Existing knowledge and needs of research and innovation. Sci. Total Environ. 679, 148–158 (2019).

    Article  CAS  PubMed  Google Scholar 

  96. Yang, Y. et al. Biodegradation and mineralization of polystyrene by plastic-eating mealworms: part 2. Role of gut microorganisms. Environ. Sci. Technol. 49, 12087–12093 (2015).

    Article  CAS  PubMed  Google Scholar 

  97. Ho, B. T., Roberts, T. K. & Lucas, S. An overview on biodegradation of polystyrene and modified polystyrene: the microbial approach. Crit. Rev. Biotechnol. 38, 308–320 (2018).

    Article  CAS  PubMed  Google Scholar 

  98. Krueger, M. C., Harms, H. & Schlosser, D. Prospects for microbiological solutions to environmental pollution with plastics. Appl. Microbiol. Biotechnol. 99, 8857–8874 (2015).

    Article  CAS  PubMed  Google Scholar 

  99. Restrepo-Flórez, J.-M., Bassi, A. & Thompson, M. R. Microbial degradation and deterioration of polyethylene — a review. Int. Biodeterior. Biodegrad. 88, 83–90 (2014).

    Article  CAS  Google Scholar 

  100. Bonhomme, S. et al. Environmental biodegradation of polyethylene. Polym. Degrad. Stabil. 81, 441–452 (2003).

    Article  CAS  Google Scholar 

  101. Wei, R. & Zimmermann, W. Microbial enzymes for the recycling of recalcitrant petroleum-based plastics: how far are we? Microb. Biotechnol. 10, 1308–1322 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  102. Santo, M., Weitsman, R. & Sivan, A. The role of the copper-binding enzyme — laccase — in the biodegradation of polyethylene by the actinomycete Rhodococcus ruber. Int. Biodeterior. Biodegrad. 84, 204–210 (2013).

    Article  CAS  Google Scholar 

  103. Fujisawa, M., Hirai, H. & Nishida, T. Degradation of polyethylene and nylon-66 by the laccase-mediator system. J. Polym. Environ. 9, 103–108 (2001).

    Article  CAS  Google Scholar 

  104. Knyazev, V. D. Effects of chain length on the rates of C–C bond dissociation in linear alkanes and polyethylene. J. Phys. Chem. A 111, 3875–3883 (2007).

    Article  CAS  PubMed  Google Scholar 

  105. Huang, J.-b et al. Theoretical study of bond dissociation energies for lignin model compounds. J. Fuel Chem. Technol. 43, 429–436 (2015).

    Article  CAS  Google Scholar 

  106. Jeon, H. J. & Kim, M. N. Functional analysis of alkane hydroxylase system derived from Pseudomonas aeruginosa E7 for low molecular weight polyethylene biodegradation. Int. Biodeterior. Biodegrad. 103, 141–146 (2015).

    Article  CAS  Google Scholar 

  107. Yoon, M. G., Jeon, H. J. & Kim, M. N. Biodegradation of polyethylene by a soil bacterium and alkB cloned recombinant cell. J. Biorem. Biodegrad. 3, 145 (2012).

    CAS  Google Scholar 

  108. van Beilen, J. B. & Funhoff, E. G. Alkane hydroxylases involved in microbial alkane degradation. Appl. Microbiol. Biotechnol. 74, 13–21 (2007).

    Article  CAS  PubMed  Google Scholar 

  109. Nakamiya, K., Sakasita, G., Ooi, T. & Kinoshita, S. Enzymatic degradation of polystyrene by hydroquinone peroxidase of Azotobacter beijerinckii HM121. J. Ferment. Bioeng. 84, 480–482 (1997).

    Article  CAS  Google Scholar 

  110. Yang, Y. et al. Biodegradation and mineralization of polystyrene by plastic-eating mealworms: Part 1. Chemical and physical characterization and isotopic tests. Environ. Sci. Technol. 49, 12080–12086 (2015).

    Article  CAS  PubMed  Google Scholar 

  111. Wierckx, N. et al. in Consequences of Microbial Interactions with Hydrocarbons, Oils, and Lipids: Biodegradation and Bioremediation. Handbook of Hydrocarbon and Lipid Microbiology (ed. Steffan, R.) 1–29 (Springer, 2018).

  112. Russell, J. R. et al. Biodegradation of polyester polyurethane by endophytic fungi. Appl. Environ. Microbiol. 77, 6076–6084 (2011).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  113. Loredo-Treviño, A., Gutiérrez-Sánchez, G., Rodríguez-Herrera, R. & Aguilar, C. N. Microbial enzymes involved in polyurethane biodegradation: a review. J. Polym. Environ. 20, 258–265 (2011).

    Article  CAS  Google Scholar 

  114. Álvarez-Barragán, J. et al. Biodegradative activities of selected environmental fungi on a polyester polyurethane varnish and polyether polyurethane foams. Appl. Environ. Microbiol. 82, 5225–5235 (2016).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  115. Akutsu-Shigeno, Y. et al. Isolation of a bacterium that degrades urethane compounds and characterization of its urethane hydrolase. Appl. Microbiol. Biotechnol. 70, 422–429 (2006).

    Article  CAS  PubMed  Google Scholar 

  116. Akutsu, Y., Nakajima-Kambe, T., Nomura, N. & Nakahara, T. Purification and properties of a polyester polyurethane-degrading enzyme from Comamonas acidovorans TB-35. Appl. Environ. Microbiol. 64, 62–67 (1998).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  117. Nakajima-Kambe, T., Shigeno-Akutsu, Y., Nomura, N., Onuma, F. & Nakahara, T. Microbial degradation of polyurethane, polyester polyurethanes and polyether polyurethanes. Appl. Microbiol. Biotechnol. 51, 134–140 (1999).

    Article  CAS  PubMed  Google Scholar 

  118. Mahajan, N. & Gupta, P. New insights into the microbial degradation of polyurethanes. RSC Adv. 5, 41839–41854 (2015).

    Article  CAS  Google Scholar 

  119. Christenson, E. M., Dadsetan, M., Wiggins, M., Anderson, J. M. & Hiltner, A. Poly(carbonate urethane) and poly(ether urethane) biodegradation: in vivo studies. J. Biomed. Mater. Res. A 69, 407–416 (2004).

    Article  CAS  PubMed  Google Scholar 

  120. Santerre, J. P., Woodhouse, K., Laroche, G. & Labow, R. S. Understanding the biodegradation of polyurethanes: from classical implants to tissue engineering materials. Biomaterials 26, 7457–7470 (2005).

    Article  CAS  PubMed  Google Scholar 

  121. Taniguchi, I. et al. Biodegradation of PET: current status and application aspects. ACS Catal. 9, 4089–4105 (2019).

    Article  CAS  Google Scholar 

  122. Ribitsch, D. et al. Hydrolysis of polyethyleneterephthalate by p-nitrobenzylesterase from Bacillus subtilis. Biotechnol. Prog. 27, 951–960 (2011).

    Article  CAS  PubMed  Google Scholar 

  123. Spiller, B., Gershenson, A., Arnold, F. H. & Stevens, R. C. A structural view of evolutionary divergence. Proc. Natl Acad. Sci. USA 96, 12305–12310 (1999).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  124. Eberl, A. et al. Enzymatic surface hydrolysis of poly(ethylene terephthalate) and bis(benzoyloxyethyl) terephthalate by lipase and cutinase in the presence of surface active molecules. J. Biotechnol. 143, 207–212 (2009).

    Article  CAS  PubMed  Google Scholar 

  125. Zimmermann, W. & Billig, S. in Biofunctionalization of Polymers and their Applications. Advances in Biochemical Engineering/Biotechnology Vol. 125 (eds Nyanhongo, G., Steiner, W. & Gübitz, G.) 97–120 (Springer, 2010).

  126. Yoshida, S. et al. A bacterium that degrades and assimilates poly(ethylene terephthalate). Science 351, 1196–1199 (2016).

    Article  CAS  PubMed  Google Scholar 

  127. Han, X. et al. Structural insight into catalytic mechanism of PET hydrolase. Nat. Commun. 8, 2106 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  128. Austin, H. P. et al. Characterization and engineering of a plastic-degrading aromatic polyesterase. Proc. Natl Acad. Sci. USA 115, E4350–E4357 (2018).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  129. Fecker, T. et al. Active site flexibility as a hallmark for efficient PET degradation by I. sakaiensis PETase. Biophys. J. 114, 1302–1312 (2018).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  130. Joo, S. et al. Structural insight into molecular mechanism of poly(ethylene terephthalate) degradation. Nat. Commun. 9, 382 (2018).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  131. Chen, C.-C., Han, X., Ko, T.-P., Liu, W. & Guo, R.-T. Structural studies reveal the molecular mechanism of PETase. FEBS J. 285, 3717–3723 (2018).

    Article  CAS  PubMed  Google Scholar 

  132. Danso, D. et al. New insights into the function and global distribution of polyethylene terephthalate (PET)-degrading bacteria and enzymes in marine and terrestrial metagenomes. Appl. Environ. Microbiol. 84, e02773-17 (2018).

    Article  PubMed  Google Scholar 

  133. Sasoh, M. et al. Characterization of the terephthalate degradation genes of Comamonas sp. strain E6. Appl. Environ. Microbiol. 72, 1825–1832 (2006).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  134. Son, H. F. et al. Rational protein engineering of thermo-stable PETase from Ideonella sakaiensis for highly efficient PET degradation. ACS Catal. 9, 3519–3526 (2019).

    Article  CAS  Google Scholar 

  135. Longhi, S. et al. Crystal structure of cutinase covalently inhibited by a triglyceride analogue. Protein Sci. 6, 275–286 (1997).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  136. Yikmis, M. & Steinbüchel, A. Historical and recent achievements in the field of microbial degradation of natural and synthetic rubber. Appl. Environ. Microbiol. 78, 4543–4551 (2012).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  137. Sundaramoorthy, M., Gold, M. H. & Poulos, T. L. Ultrahigh (0.93 Å) resolution structure of manganese peroxidase from Phanerochaete chrysosporium: implications for the catalytic mechanism. J. Inorg. Biochem. 104, 683–690 (2010).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  138. Choinowski, T., Blodig, W., Winterhalter, K. H. & Piontek, K. The crystal structure of lignin peroxidase at 1.70 Å resolution reveals a hydroxy group on the Cβ of tryptophan 171: a novel radical site formed during the redox cycle. J. Mol. Biol. 286, 809–827 (1999).

    Article  CAS  PubMed  Google Scholar 

  139. McCarter, J. D. & Withers, S. G. Mechanisms of enzymatic glycoside hydrolysis. Curr. Opin. Struct. Biol. 4, 885–892 (1994).

    Article  CAS  PubMed  Google Scholar 

  140. Kim, H.-W. & Ishikawa, K. Functional analysis of hyperthermophilic endocellulase from Pyrococcus horikoshii by crystallographic snapshots. Biochem. J. 437, 223–230 (2011).

    Article  CAS  PubMed  Google Scholar 

  141. Seyedarabi, A. et al. Structural insights into substrate specificity and the anti β-elimination mechanism of pectate lyase. Biochemistry 49, 539–546 (2009).

    Article  CAS  Google Scholar 

  142. Hangasky, J. A., Detomasi, T. C. & Marletta, M. A. Glycosidic bond hydroxylation by polysaccharide monooxygenases. Trends Chem. 1, 198–209 (2019).

    Article  Google Scholar 

Download references

Acknowledgements

This work has been supported by the National Key Research and Development Program of China (2019YFA0706900) and Hubei Natural Science Foundation (grant 2017CFB158). We thank T.-P. Ko and X. Yu for helpful discussions.

Author information

Authors and Affiliations

  1. State Key Laboratory of Biocatalysis and Enzyme Engineering, Hubei Collaborative Innovation Center for Green Transformation of Bio-Resources, Hubei Key Laboratory of Industrial Biotechnology, School of Life Sciences, Hubei University, Wuhan, China

    Chun-Chi Chen, Longhai Dai, Lixin Ma & Rey-Ting Guo

Authors
  1. Chun-Chi Chen
    View author publications

    You can also search for this author in PubMed Google Scholar

  2. Longhai Dai
    View author publications

    You can also search for this author in PubMed Google Scholar

  3. Lixin Ma
    View author publications

    You can also search for this author in PubMed Google Scholar

  4. Rey-Ting Guo
    View author publications

    You can also search for this author in PubMed Google Scholar

Contributions

C.-C.C. and L.D. contributed to the writing and editing of this manuscript. L.M. and R.-T.G. contributed to the discussion of content and writing and reviewing/editing the manuscript before submission.

Corresponding authors

Correspondence to Lixin Ma or Rey-Ting Guo.

Ethics declarations

Competing interests

The authors declare no competing interests.

Additional information

Publisher’s note

Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Related links

CAZy database: http://www.cazy.org/

Statista: https://www.statista.com/

Rights and permissions

Reprints and permissions

About this article

Check for updates. Verify currency and authenticity via CrossMark

Cite this article

Chen, CC., Dai, L., Ma, L. et al. Enzymatic degradation of plant biomass and synthetic polymers. Nat Rev Chem 4, 114–126 (2020). https://doi.org/10.1038/s41570-020-0163-6

Download citation

  • Accepted:

  • Published:

  • Issue Date:

  • DOI: https://doi.org/10.1038/s41570-020-0163-6

This article is cited by

Search

Advanced search

Quick links

Nature Briefing

Sign up for the Nature Briefing newsletter — what matters in science, free to your inbox daily.

Get the most important science stories of the day, free in your inbox. Sign up for Nature Briefing