Significance Synthetic polymers are ubiquitous in the modern world but pose a global environmental problem. While plastics such as poly(ethylene terephthalate) (PET) are highly versatile, their resistance to natural degradation presents a serious, growing risk to fauna and flora, particularly in marine environments. Here, we have characterized the 3D structure of a newly discovered enzyme that can digest highly crystalline PET, the primary material used in the manufacture of single-use plastic beverage bottles, in some clothing, and in carpets. We engineer this enzyme for improved PET degradation capacity and further demonstrate that it can also degrade an important PET replacement, polyethylene-2,5-furandicarboxylate, providing new opportunities for biobased plastics recycling.

Abstract Poly(ethylene terephthalate) (PET) is one of the most abundantly produced synthetic polymers and is accumulating in the environment at a staggering rate as discarded packaging and textiles. The properties that make PET so useful also endow it with an alarming resistance to biodegradation, likely lasting centuries in the environment. Our collective reliance on PET and other plastics means that this buildup will continue unless solutions are found. Recently, a newly discovered bacterium, Ideonella sakaiensis 201-F6, was shown to exhibit the rare ability to grow on PET as a major carbon and energy source. Central to its PET biodegradation capability is a secreted PETase (PET-digesting enzyme). Here, we present a 0.92 Å resolution X-ray crystal structure of PETase, which reveals features common to both cutinases and lipases. PETase retains the ancestral α/β-hydrolase fold but exhibits a more open active-site cleft than homologous cutinases. By narrowing the binding cleft via mutation of two active-site residues to conserved amino acids in cutinases, we surprisingly observe improved PET degradation, suggesting that PETase is not fully optimized for crystalline PET degradation, despite presumably evolving in a PET-rich environment. Additionally, we show that PETase degrades another semiaromatic polyester, polyethylene-2,5-furandicarboxylate (PEF), which is an emerging, bioderived PET replacement with improved barrier properties. In contrast, PETase does not degrade aliphatic polyesters, suggesting that it is generally an aromatic polyesterase. These findings suggest that additional protein engineering to increase PETase performance is realistic and highlight the need for further developments of structure/activity relationships for biodegradation of synthetic polyesters.

In less than a century of manufacturing, plastics have become essential to modern society, driven by their incredible versatility coupled to low production costs. It is, however, now widely recognized that plastics pose a dire global pollution threat, especially in marine ecosystems, because of the ultralong lifetimes of most synthetic plastics in the environment (1⇓⇓⇓⇓⇓⇓⇓–9). In response to the accumulation of plastics in the biosphere, it is becoming increasingly recognized that microbes are adapting and evolving enzymes and catabolic pathways to partially degrade man-made plastics as carbon and energy sources (10⇓⇓⇓⇓⇓⇓⇓⇓–19). These evolutionary footholds offer promising starting points for industrial biotechnology and synthetic biology to help address the looming environmental threat posed by man-made synthetic plastics (19⇓⇓⇓–23).

Poly(ethylene terephthalate) (PET) is the most abundant polyester plastic manufactured in the world. Most applications that employ PET, such as single-use beverage bottles, clothing, packaging, and carpeting, employ crystalline PET, which is recalcitrant to catalytic or biological depolymerization due to the limited accessibility of the ester linkages. In an industrial context, PET can be depolymerized to its constituents via chemistries able to cleave ester bonds (24, 25). However, to date, few chemical recycling solutions have been deployed, given the high processing costs relative to the purchase of inexpensive virgin PET. This, in turn, results in reclaimed PET primarily being mechanically recycled, ultimately resulting in a loss of material properties, and hence intrinsic value. Given the recalcitrance of PET, the fraction of this plastic stream that is landfilled or makes its way to the environment is projected to persist for hundreds of years (1).

In 2016, Yoshida et al. (17) reported a newly discovered bacterium, Ideonella sakaiensis 201-F6, with the unusual ability to use PET as its major carbon and energy source for growth. Especially in the past decade, there have been multiple, foundational studies reporting enzymes that can degrade PET (10, 26⇓⇓⇓⇓–31), but, to our knowledge, previous work had not connected extracellular enzymatic PET degradation to catabolism (11) in a single microbe. As illustrated in Fig. 1, Yoshida et al. (17) demonstrated that an I. sakaiensis enzyme dubbed PETase (PET-digesting enzyme) converts PET to mono(2-hydroxyethyl) terephthalic acid (MHET), with trace amounts of terephthalic acid (TPA) and bis(2-hydroxyethyl)-TPA as secondary products. A second enzyme, MHETase (MHET-digesting enzyme), further converts MHET into the two monomers, TPA and ethylene glycol (EG). Both enzymes are secreted by I. sakaiensis and likely act synergistically to depolymerize PET. Sequence analysis and recent structural studies of PETase highlight similarities to α/β-hydrolase enzymes (17, 32, 33), including the cutinase and lipase families, which catalyze hydrolysis of cutin and fatty acids, respectively. This observation provides clues to the origin of PETase, but further insights into its structural and functional evolution are needed.

Fig. 1. PETase catalyzes the depolymerization of PET to bis(2-hydroxyethyl)-TPA (BHET), MHET, and TPA. MHETase converts MHET to TPA and EG.

Beyond PET, humankind uses a wide range of polyesters, broadly classified by aliphatic and aromatic content. PET, for example, is a semiaromatic polyester. Some aliphatic polyesters, such as polylactic acid (PLA) (34), polybutylene succinate (PBS) (35), or polyhydroxyalkanoates (36), can be produced from renewable sources and are marketed as biodegradable plastics, given their relatively low crystallinity and glass transition temperatures, in turn, providing relatively more direct enzymatic access to ester linkages. Aromatic and semiaromatic polyesters, conversely, often exhibit enhanced thermal and material properties and, accordingly, have reached substantially higher market volume but are typically not as biodegradable as their aliphatic counterparts. An emerging, biobased PET replacement is polyethylene-2,5-furandicarboxylate [or poly(ethylene furanoate); PEF], which is based on sugar-derived 2,5-furandicarboxylic acid (FDCA) (37). PEF exhibits improved gas barrier properties over PET and is being pursued industrially (38). Even though PEF is a biobased semiaromatic polyester, which is predicted to offset greenhouse gas emissions relative to PET (39), its lifetime in the environment, like that of PET, is likely to be quite long (40). Given that PETase has evolved to degrade crystalline PET, it potentially may have promiscuous activity across a range of polyesters.

In this study, we aimed to gain a deeper understanding of the adaptations that contribute to the substrate specificity of PETase. To this end, we report multiple high-resolution X-ray crystal structures of PETase, which enable comparison with known cutinase structures. Based on differences in the PETase and a homologous cutinase active-site cleft (41), PETase variants were produced and tested for PET degradation, including a double mutant distal to the catalytic center that we hypothesized would alter important substrate-binding interactions. Surprisingly, this double mutant, inspired by cutinase architecture, exhibits improved PET degradation capacity relative to wild-type PETase. We subsequently employed in silico docking and molecular dynamics (MD) simulations to characterize PET binding and dynamics, which provide insights into substrate binding and suggest an explanation for the improved performance of the PETase double mutant. Additionally, incubation of wild-type and mutant PETase with several polyesters was examined using scanning electron microscopy (SEM), differential scanning calorimetry (DSC), and product release. These studies showed that the enzyme can degrade both crystalline PET (17) and PEF, but not aliphatic polyesters, suggesting a broader ability to degrade semiaromatic polyesters. Taken together, the structure/function relationships elucidated here could be used to guide further protein engineering to more effectively depolymerize PET and other synthetic polymers, thus informing a biotechnological strategy to help remediate the environmental scourge of plastic accumulation in nature (19⇓⇓⇓–23).

Discussion The high-resolution structure described in the present study reveals the binding site architecture of the I. sakaiensis 201-F6 PETase, while the IFD results provide a mechanistic basis for both the wild type and PETase double mutant toward the crystalline semiaromatic polyesters PET and PEF. Changes around the active site result in a widening of the cleft compared with structural representatives of three thermophilic cutinases (SI Appendix, Fig. S3), without other major changes in the underlying secondary or tertiary structure. Furthermore, we demonstrated that PETase is active on PET of ∼15% crystallinity; while this observation is encouraging, it is envisaged that its performance would need to be enhanced substantially, perhaps via further active-site cleft engineering similar to ongoing work on thermophilic cutinases and lipases (26, 30, 53, 54). Enzyme scaffolds capable of PET breakdown above the glass transition temperature (≥∼70 °C for PET) (20) will also be pursued in future studies. Coupling with other processes such as milling or grinding, which can increase the available surface area of the plastic, also merits investigation toward enzymatic solutions for PET and PEF recycling. Furthermore, in light of recent studies that demonstrate the impressive synergistic effect of combining multiple PET-active lipases (26, 30, 53, 54), we expect that incorporation of I. sakaiensis MHETase will further increase the performance (55), and this will be pursued in future work. The highly basic surface charge of PETase requires further investigation since it is not observed in other close structural homologs, but it is noteworthy that the MHETase partner is predicted to be a fairly acidic protein, with a pI in the region of 5.2. Both the IFD results and MD simulations independently indicate the PETase binding site is characterized by highly flexible, large aromatic side chains, such as Trp185, Tyr87, and Trp159, and Phe238 in the PETase double mutant. Binding of PET and PEF induces conformational changes in these residues relative to the crystal structure; thus, modeling protein flexibility in response to PET/PEF is critical to predict catalytically relevant binding modes. Additionally, results of these flexible docking studies agree with experimentally observed trends in performance in the wild type relative to the double mutant, and provide structural insight to explain this enhancement. PETase activity on both PET and PEF, but not on aliphatic polyesters such as PBS and PLA, provides the basis for characterizing this enzyme more broadly as an aromatic polyesterase rather than solely as a PETase. It is likely that the enhanced gas barrier properties of PEF will lead to its adoption for beer bottles, and that this recalcitrant material will thus ultimately find its way to the environment. It is therefore encouraging that PETase is also natively capable of PEF degradation. It is also noteworthy that in this study, PETase was freeze-dried and shipped between continents, and that it retained similar performance profiles after freeze-drying, which is a positive feature for its potential use in applications that require enzyme production and use be distinct, as it would potentially be the case for most biobased recycling options. The problem of plastics depolymerization by enzymes closely mirrors that of enzymes that depolymerize polysaccharides, such as cellulose and chitin (56, 57). Indeed, strategies that have been used to understand and improve glycoside hydrolases, including the development of quantitative assays for measuring enzyme (or enzyme cocktail) performance on solid substrates, likely can serve as inspiration for more quantitative metrics for comparing plastics-degrading enzymes and enzyme mixtures, which will be reported in future studies. Moreover, the method of PETase action is of keen interest for further protein and enzyme mixture engineering studies. The direct catalytic mechanism could be studied with mixed quantum mechanical/molecular mechanics MD-based approaches similar to previous work on carbohydrate-active enzymes (58). Beyond the active site, the enzyme may interact with and cleave the substrate in an endofashion by cleaving PET (or PEF) chains internal to a polymer or in an exofashion by only cleaving PET from the chain ends. Methods employed in the cellulase and chitinase research community, such as substrate labeling with easily detected reporter molecules or examination of product ratios, could potentially shed light on this question, and will be pursued in future efforts (59). Lastly, at low substrate loadings, many polysaccharide-active enzymes rely on multimodular architectures, with a carbohydrate-binding module attached to the catalytic domain (57). For polyesterase enzymes, hydrophobins, carbohydrate-binding modules, and polyhydroxyalkanoate-binding modules have been used to increase the catalytic efficiency of cutinases for PET degradation (60, 61). Certainly, further opportunities exist for engineering or evolving for higher binding affinity of accessory modules to increase the overall surface concentration of catalytic domains on the PET surface. Given the fact that PET was only patented roughly 80 y ago and put into widespread use in the 1970s, it is likely that the enzyme system for PET degradation and catabolism in I. sakaiensis appeared only recently, demonstrating the remarkable speed at which microbes can evolve to exploit new substrates: in this case, waste from an industrial PET recycling facility. Moreover, given the results obtained for the PETase double mutant, it is likely that significant potential remains for improving its activity further. This enzyme thus provides an exciting platform for additional protein engineering and evolution to increase the efficiency and substrate range of this polyesterase, as well as to provide clues of how to further engineer thermophilic cutinases to better incorporate aromatic polyesters, toward to the persistent challenge of highly crystalline polymer degradation.

Conclusions The discovery of a bacterium that uses PET as a major carbon and energy source has raised significant interest in how such an enzymatic mechanism functions with such a highly resistant polymeric substrate that appears to survive for centuries in the environment. This work shows that a collection of subtle variations on the surface of a lipase/cutinase-like fold has the ability to endow PETase with a platform for aromatic polyester depolymerization. These findings open up the possibility to further utilize and combine the extensive platform of cutinase and lipase research over the past decades with directed protein engineering and evolution to adapt this scaffold further and tackle environmentally relevant polymer bioaccumulation and biobased industrial polyester recycling.

Methods Cloning and Protein Production. Codon optimized Escherichia coli expression clones were constructed for PETase as described in SI Appendix, Fig. S2B. Crystallization and Structure Determination. PETase was crystallized in five conditions, and long-wavelength sulfur–single-wavelength anomalous diffraction and high-resolution X-ray data collection was performed in vacuo at beamline I23. Standard X-ray data collection was performed at beamlines I03 and I04 at the Diamond Light Source. Detailed methods and statistics are provided in SI Appendix, Table S1. Substrate Docking. The PETase crystal structure, PETase double mutant, and PET and PEF tetramers were modeled using tools from Schrödinger. Protein preparation and ligand preparation where conducted using tools in Schrödinger, along with IFD, to predict PET and PEF binding modes to PETase wild type and double mutant. Additional details can be found in SI Appendix (62, 63). Polymer Synthesis. PET and PEF were produced via the polycondensation of EG with TPA and FDCA, respectively. Following polycondensation, the polymers were dissolved in trifluoroacetic acid, precipitated in methanol, and subsequently redissolved in trifluoroacetic acid for film casting. Following casting, the coupons were annealed in a vacuum oven at 90 °C (above their glass transition temperature). Additional details can be found in SI Appendix. PETase Digestion of Polymer Films. Coupons sized ∼6 mm in diameter of each polymer film were placed in a 1.5-mL Eppendorf tube with 500 μL of 50 nM PETase in 50 mM phosphate buffer at pH 7.2. The digestions were carried out at 30 °C. Analysis of the films and supernatant was done after 96 h of digestion. SEM. Polymer coupons sized ∼6 mm in diameter were examined by SEM, both before and after PETase treatment for 96 h. PETase-treated samples were rinsed with 1% SDS, followed by dH 2 O and then ethanol. Samples were sputter-coated with 8 nm of iridium. Coated samples were mounted on aluminum stubs using carbon tape, and conductive silver paint was applied to the sides of the samples to reduce charging. SEM imaging was performed using an FEI Quanta 400 FEG instrument under low vacuum (0.45 torr) operating with a gaseous solid-state detector. Imaging was performed with a beam-accelerating voltage of 15 keV.

Acknowledgments We thank the staff at the Diamond Light Source for their support and Simon Cragg for helpful comments on the manuscript. B.S.D., N.A.R., G.D., W.E.M., A.A., M.F.C., C.W.J., and G.T.B. thank the National Renewable Energy Laboratory (NREL) Directed Research and Development Program for funding. J.E.M. was supported by Grant BB/P011918/1 from the Biotechnology and Biological Sciences Research Council. H.P.A. was funded through an NREL subcontract and University of Portsmouth Faculty of Science bursary. R.L.S. was supported by a postdoctoral fellowship abroad (Grants 2013/08293-7, 2014/10448-1, and 2016/22956-7) from the Sao Paulo Research Foundation and Center for Computational Engineering & Sciences. H.L.W. was supported by Grant DE-SC0011297TDD from the US Department of Energy (DOE) and the National Science Foundation (NSF) under Grant CHE–1464946. F.L.K. acknowledges support from the NSF Graduate Research Fellowship Program (Grant 3900101301). Computer time was provided by Extreme Science and Engineering Discovery Environment (XSEDE) allocation MCB-090159 at the San Diego Supercomputing Center and the Texas Advanced Computing Center, and by the NREL Computational Sciences Center supported by the DOE Office of Energy Efficiency and Renewable Energy under Contract DE-AC36-08GO28308. The publisher, by accepting the article for publication, acknowledges that the US Government retains a nonexclusive, paid up, irrevocable, worldwide license to publish or reproduce the published form of this work, or allow others to do so, for US Government purposes.

Footnotes Author contributions: J.E.M. and G.T.B. designed research; H.P.A., M.D.A., B.S.D., N.A.R., F.L.K., R.L.S., B.C.P., G.D., R.D., K.E.O., V.M., A.W., W.E.M., A.A., A.W.T., C.W.J., and H.L.W. performed research; H.P.A., M.D.A., B.S.D., N.A.R., F.L.K., R.L.S., B.C.P., G.D., R.D., K.E.O., V.M., A.W., W.E.M., A.A., M.S.S., M.F.C., A.W.T., C.W.J., H.L.W., J.E.M., and G.T.B. analyzed data; and J.E.M. and G.T.B. wrote the paper with contributions from all authors.

Conflict of interest statement: H.P.A., M.D.A., B.S.D., N.A.R., C.W.J., J.E.M., and G.T.B. have filed a patent application on the PETase double mutant.

This article is a PNAS Direct Submission.

Data deposition: The atomic coordinates and structure factors have been deposited in the Protein Data Bank, www.wwpdb.org (PDB ID codes 6EQD, 6EQE, 6EQF, 6EQG, and 6EQH).

This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10.1073/pnas.1718804115/-/DCSupplemental.