Successfully interfacing enzymes and biomachinery with polymers affords on-demand modification and/or programmable degradation during the manufacture, utilization and disposal of plastics, but requires controlled biocatalysis in solid matrices with macromolecular substrates1,2,3,4,5,6,7. Embedding enzyme microparticles speeds up polyester degradation, but compromises host properties and unintentionally accelerates the formation of microplastics with partial polymer degradation6,8,9. Here we show that by nanoscopically dispersing enzymes with deep active sites, semi-crystalline polyesters can be degraded primarily via chain-end-mediated processive depolymerization with programmable latency and material integrity, akin to polyadenylation-induced messenger RNA decay10. It is also feasible to achieve processivity with enzymes that have surface-exposed active sites by engineering enzyme–protectant–polymer complexes. Poly(caprolactone) and poly(lactic acid) containing less than 2 weight per cent enzymes are depolymerized in days, with up to 98 per cent polymer-to-small-molecule conversion in standard soil composts and household tap water, completely eliminating current needs to separate and landfill their products in compost facilities. Furthermore, oxidases embedded in polyolefins retain their activities. However, hydrocarbon polymers do not closely associate with enzymes, as their polyester counterparts do, and the reactive radicals that are generated cannot chemically modify the macromolecular host. This study provides molecular guidance towards enzyme–polymer pairing and the selection of enzyme protectants to modulate substrate selectivity and optimize biocatalytic pathways. The results also highlight the need for in-depth research in solid-state enzymology, especially in multi-step enzymatic cascades, to tackle chemically dormant substrates without creating secondary environmental contamination and/or biosafety concerns.
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Tokiwa, Y. & Suzuki, T. Hydrolysis of polyesters by lipases. Nature 270, 76–78 (1977).
Tournier, V. et al. An engineered PET depolymerase to break down and recycle plastic bottles. Nature 580, 216–219 (2020).
Kuchler, A., Yoshimoto, M., Luginbuhl, S., Mavelli, F. & Walde, P. Enzymatic reactions in confined environments. Nat. Nanotechnol. 11, 409–420 (2016).
Yang, Z. et al. Activity and stability of enzymes incorporated into acrylic polymers. J. Am. Chem. Soc. 117, 4843–4850 (1995).
Panganiban, B. et al. Random heteropolymers preserve protein function in foreign environments. Science 359, 1239–1243 (2018).
Ganesh, M., Dave, R. N., L’Amoreaux, W. & Gross, R. A. Embedded enzymatic biomaterial degradation. Macromolecules 42, 6836–6839 (2009).
DelRe, C. et al. Reusable enzymatic fiber mats for neurotoxin remediation in water. ACS Appl. Mater. Inter. 10, 44216–44220 (2018).
Khan, I., Nagarjuna, R., Dutta, J. R. & Ganesan, R. Enzyme-embedded degradation of poly(epsilon-caprolactone) using lipase-derived from probiotic Lactobacillus plantarum. ACS Omega 4, 2844–2852 (2019).
Huang, Q. Y., Hiyama, M., Kabe, T., Kimura, S. & Iwata, T. Enzymatic self-biodegradation of poly(l-lactic acid) films by embedded heat-treated and immobilized proteinase K. Biomacromolecules 21, 3301–3307 (2020).
Xu, F. F. & Cohen, S. N. RNA degradation in Escherichia coli regulated by 3′ adenylation and 5′ phosphorylatian. Nature 374, 180–183 (1995).
Wei, R. et al. Possibilities and limitations of biotechnological plastic degradation and recycling. Nat. Catal. 3, 867–871 (2020).
Ivleva, N. P., Wiesheu, A. C. & Niessner, R. Microplastic in aquatic ecosystems. Angew. Chem. Int. Ed. 56, 1720–1739 (2017).
Jambeck, J. R. et al. Plastic waste inputs from land into the ocean. Science 347, 768–771 (2015).
Haider, T. P., Volker, C., Kramm, J., Landfester, K. & Wurm, F. R. Plastics of the future? The impact of biodegradable polymers on the environment and on society. Angew. Chem. Int. Ed. 58, 50–62 (2019).
Roohi, et al. Microbial enzymatic degradation of biodegradable plastics. Curr. Pharm. Biotechnol. 18, 429–440 (2017).
Breyer, W. A. & Matthews, B. W. Structure of Escherichia coli exonuclease I suggests how processivity is achieved. Nat. Struct. Biol. 7, 1125–1128 (2000).
Pleiss, J., Fischer, M. & Schmid, R. D. Anatomy of lipase binding sites: the scissile fatty acid binding site. Chem. Phys. Lipids 93, 67–80 (1998).
Li, S. M. & McCarthy, S. Influence of crystallinity and stereochemistry on the enzymatic degradation of poly(lactide)s. Macromolecules 32, 4454–4456 (1999).
Flory, P. J. & Yoon, D. Y. Molecular morphology in semi-crystalline polymers. Nature 272, 226–229 (1978).
Tokiwa, Y., Calabia, B. P., Ugwu, C. U. & Aiba, S. Biodegradability of plastics. Int. J. Mol. Sci. 10, 3722–3742 (2009).
Ru, J. K., Huo, Y. X. & Yang, Y. Microbial degradation and valorization of plastic wastes. Front Microbiol 11, 442, (2020).
Tennakoon, A. et al. Catalytic upcycling of high-density polyethylene via a processive mechanism. Nat. Catal. 3, 893–901 (2020).
Horn, S. J. et al. Costs and benefits of processivity in enzymatic degradation of recalcitrant polysaccharides. Proc. Natl Acad. Sci. USA 103, 18089–18094 (2006).
Beckham, G. T. et al. Molecular-level origins of biomass recalcitrance: decrystallization free energies for four common cellulose polymorphs. J. Phys. Chem. B 115, 4118–4127 (2011).
Payne, C. M., Himmel, M. E., Crowley, M. F. & Beckham, G. T. Decrystallization of oligosaccharides from the cellulose Iβ surface with molecular simulation. J. Phys. Chem. Lett. 2, 1546–1550 (2011).
Klibanov, A. M. Improving enzymes by using them in organic solvents. Nature 409, 241–246 (2001).
Christensen, P. R., Scheuermann, A. M., Loeffler, K. E. & Helms, B. A. Closed-loop recycling of plastics enabled by dynamic covalent diketoenamine bonds. Nat. Chem. 11, 442–448 (2019).
Satchwell, A. J. et al. Accelerating the deployment of anaerobic digestion to meet zero waste goals. Environ. Sci. Technol. 52, 13663–13669 (2018).
Hobbs, S. R., Parameswaran, P., Astmann, B., Devkota, J. P. & Landis, A. E. Anaerobic codigestion of food waste and polylactic acid: effect of pretreatment on methane yield and solid reduction. Adv. Mater. Sci. Eng. 2019, 4715904 (2019).
Nordahl, S. L. et al. Life-Cycle Greenhouse Gas Emissions and Human Health Trade-Offs of Organic Waste Management Strategies. Environ. Sci. Technol. 54, 9200–9209 (2020).
Radi, R. Oxygen radicals, nitric oxide, and peroxynitrite: redox pathways in molecular medicine. Proc. Natl Acad. Sci. USA 115, 5839–5848 (2018).
Iiyoshi, Y., Tsutsumi, Y. & Nishida, T. Polyethylene degradation by lignin-degrading fungi and malaganese peroxidase. J. Wood Sci. 44, 222–229 (1998).
Yang, J., Yang, Y., Wu, W. M., Zhao, J. & Jiang, L. Evidence of polyethylene biodegradation by bacterial strains from the guts of plastic-eating waxworms. Environ. Sci. Technol. 48, 13776–13784 (2014).
Schneiderman, D. K. & Hillmyer, M. A. There is a great future in sustainable polymers. Macromolecules 50, 3733–3749 (2017).
Liu, L. J., Li, S. M., Garreau, H. & Vert, M. Selective enzymatic degradations of poly(l-lactide) and poly(epsilon-caprolactone) blend films. Biomacromolecules 1, 350–359 (2000).
Varma-Nair, M., Pan, R. & Wunderlich, B. Heat capacities and entropies of linear, aliphatic polyesters. J. Polym. Sci. Pol. Phys. 29, 1107–1115 (1991).
Hall, M., Bansal, P., Lee, J. H., Realff, M. J. & Bommarius, A. S. Cellulose crystallinity – a key predictor of the enzymatic hydrolysis rate. FEBS J. 277, 1571–1582 (2010).
Jiang, T. et al. Single-chain heteropolymers transport protons selectively and rapidly. Nature 577, 216–220 (2020).
Arnold, F. H. Directed evolution: bringing new chemistry to life. Angew. Chem. Int. Ed. 57, 4143–4148 (2018).
Bornscheuer, U. et al. Lipase of pseudomonas-cepacia for biotechnological purposes – purification, crystallization and characterization. Biochim. Biophys. Acta Gen. Subj. 1201, 55–60 (1994).
Wurm, A. et al. Crystallization and homogeneous nucleation kinetics of poly(epsilon-caprolactone) (PCL) with different molar masses. Macromolecules 45, 3816–3828 (2012).
Ajioka, M., Suizu, H., Higuchi, C. & Kashima, T. Aliphatic polyesters and their copolymers synthesized through direct condensation polymerization. Polym. Degrad. Stabil. 59, 137–143 (1998).
Smith, A. A. A., Hall, A., Wu, V. & Xu, T. Practical prediction of heteropolymer composition and drift. ACS Macro Lett. 8, 36–40 (2019).
This work was primarily supported by the US Department of Energy, Office of Science, Office of Basic Energy Sciences, Materials Sciences and Engineering Division (DOE-BES-MSE) under contract DE-AC02-05-CH11231, through the Organic−Inorganic Nanocomposites KC3104 programme (enzyme–plastic composite design and degradation mechanism; C.D., L.M., A.H., I.J., R.O.R., T.X.). Y.J. and T.P.R. acknowledge support from DOE-BES-MSE under the same contract number, through the Adaptive Interfacial Assemblies Towards Structuring Liquids KCTR16 programme (tensiometry studies on RHP/enzyme/polymer complexation). C.D.S. acknowledges Laboratory Directed Research and Development funding from Berkeley Lab (integration with organic waste infrastructure). The US Department of Defense, Army Research Office supported Z.R. (lipase stabilization) under contract W911NF-13-1-0232. C.D. was partially supported by a National Defense Science and Engineering Graduate (NDSEG) Fellowship (Bioremediation). T.X. also acknowledges support from a Bakar Fellowship at University of California Berkeley (compost tests). Scattering studies at the Advanced Light Source were supported by the US Department of Energy, Office of Science, Office of Basic Energy Science under contract DE-AC02-05CH11231.
T.X., C.D. and J.K. have filed a PCT patent application. A.H. is the founder and CEO of Intropic Materials.
Peer review information Nature thanks Jayati Ray Dutta, Mattheos Koffas and the other, anonymous, reviewer(s) for their contribution to the peer review of this work.
Publisher’s note Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
Extended data figures and tables
a, DLS results for RHP and purified BC–lipase in toluene (the solvent used to cast PCL) with an average hydrodynamic diameter of 285 nm ± 35 nm (n = 5) (the error indicates standard deviation). b, DSC results for PCL and PCL–RHP–BC-lipase as-cast films. c, SAXS curves of PCL and PCL–RHP–BC–lipase as-cast films.
Liquid chromatogram of the degradation by-products for degradation by confined and dissolved (surface erosion) BC–lipase.
a, GPC curve of the degradation of PCL–RHP–CA–lipase, showing a shift and broadening of the main peak, indicative of random chain scission. b, Zoomed-in version of a illustrating the peak shift and broadening.
a, PCL degradation by BC–lipase dissolved in solution (surface), nanoscopically embedded in PCL with RHP, and embedded with Tween 80, a small-molecule surfactant, as microparticles. b, Hydrolysis of 4-nitrophenyl butyrate, a small-molecule ester, by BC–lipase in solution or confined in PCL (error bars represent one standard deviation; n ≥ 3).
Extended Data Fig. 5 Model interfacial-tension experiment to explain intermolecular interactions among enzyme, protectant and matrix.
a–c, When all three components are initially mixed in toluene (a, left) and then a water interface is introduced (a, right), RHP–lipase complexes immediately interact with PCL at the interface, as shown by the fluorescence microscopy image taken ~20 s after shaking the vial to produce an emulsion (b) and the long delay time in interfacial-tension reduction that is seen only for PCL–RHP–lipase (c).
Extended Data Fig. 6 Characterization of semi-crystalline properties of melt-processed PCL–RHP–BC–lipase.
a, DSC curves of PCL–RHP–BC–lipase with different recrystallization conditions (the film with recrystallization temperature Tc = 49 °C has a crystallinity of 41% ± 1.2% compared to 39% ± 1.8% for the as-cast film). The increase in melting temperature from ~58 °C to ~64 °C indicates a substantial thickening in crystalline lamellae for Tc = 49 °C films, which was confirmed by SAXS. b, SAXS profiles of as-cast and Tc = 49 °C films of PCL–RHP–BC–lipase. The increase of long periods (shift to lower q), combined with the negligible difference in the bulk per cent crystallinity according to DSC data, confirms a thickening in crystalline lamellae after crystallizing at Tc = 49 °C.
Small-molecule ester hydrolysis by embedded BC–lipase as a function of temperature (red), overlaid with the PCL–RHP–BC–lipase degradation rate. The small-molecule activity remained high at 60 °C but was not quantified because the film shrivelled owing to melting, and thus was much thicker than films at lower temperatures, making quantification incomparable to all other temperatures.
a, Hydropathy plots for RHPs with 60:10 MMA:EHMA composition. b, Hydropathy plots for RHPs with 50:20 MMA:EHMA composition. c, Hydropathy plots for RHPs with 20:50 MMA:EHMA composition. d, Average segmental HLB value for each RHP composition. Error bars indicate standard deviation, n ≥ 3.
a, Crystal structure of proteinase K with the same colour-coding scheme as that used for lipases in the main text (Fig. 3). b, GPC curve of PLA–RHP–proteinase K (‘ProK”) as cast and after depolymerizing in buffer; c, Interfacial-tensiometry experiment results for a DCM–water interface with PLA, RHP and proteinase K in the DCM phase. d, Photograph of ABTS small-molecule assay in malonate buffer after ~10 min, demonstrating that laccase embedded in polystyrene (PS) retains the ability to oxidize a small molecule. Similar results were found for manganese peroxidase and for both enzymes embedded in polyethylene. e, Interfacial-tensiometry experiment results for a toluene–water interface with PS, RHP and either laccase or manganese peroxidase (‘MnP’) in the toluene phase.
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DelRe, C., Jiang, Y., Kang, P. et al. Near-complete depolymerization of polyesters with nano-dispersed enzymes. Nature 592, 558–563 (2021). https://doi.org/10.1038/s41586-021-03408-3
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