CROSS-REFERENCE TO RELATED APPLICATIONS This application claims the benefit of U.S. Provisional Application No. 62/636,594 filed on February 28, 2018, the contents of which are incorporated herein by reference in their entirety.

CONTRACTUAL ORIGIN

The United States Government has rights in this invention under Contract No. DE- AC36-08GO28308 between the United States Department of Energy and Alliance for Sustainable Energy, LLC, the Manager and Operator of the National Renewable Energy Laboratory .

REFERENCE TO SEQUENCE LISTING

This application contains a Sequence Listing submitted as an electronic text file entitled “NREL 18-54 Sequence Listing_ST25.txt,” having a size in bytes of 8 kb and created on February 6, 2019 Pursuant to 37 C.F.R. § 1.52(e)(5), the information contained in the above electronic file is hereby incorporated by reference in its entirety

BACKGROUND

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 to marine wildlife and ecosystems, because of the ultra-long lifetimes of most synthetic plastics in the environment. 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. 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.

Polyethylene 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. 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.

The foregoing examples of the related art and limitations related therewith are intended to be illustrative and not exclusive. Other limitations of the related art will become apparent to those of skill in the art upon a reading of the specification and a study of the drawings.

The following embodiments and aspects thereof are described and illustrated in conjunction with systems, tools, and methods that are meant to be exemplary and illustrative, not limiting in scope. In various embodiments, one or more of the above-described problems have been reduced or eliminated, while other embodiments are directed to other improvements.

Exemplary embodiments provide a modified poiytethylene terephthalate) (PET)- digesting enzyme (PETase) that exhibits improved polymer degradation capacity relative to wild-type PETase due to its narrowed binding cleft via mutation of two active-site residues is disclosed. In some embodiments, unmodified PETase is from a bacterium of the genus Ideonella. In some embodiments, the bacterium is a strain of Ideonella sakaiensis. In some embodiments, an amino residue at position 159 is mutated. In some embodiments, an amino residue at position 238 is mutated. In some embodiments, an amino residue at position 238 is mutated. In some embodiments, the modified PETases comprises the W159H/S238F double mutation.

In exemplary embodiments, a nucleic acid molecule encoding a modified PETase having mutations at two active-site residues is disclosed. In some embodiments, the amino acid residue at position 159 is mutated. In some embodiments, the amino acid residue at position 238 is mutated. In some embodiments, the modified PETase comprises the W159H/S238F double mutation. In exemplary embodiments, an expression vector comprising the nucleic acid molecule encoding a modified PETase having mutations at two active-site residues is disclosed.

Exemplary embodiments provide a nucleic acid encoding the enzyme comprising the amino acid sequence depicted in FIG. 2(B). In others, the cell that expresses the modified poly(ethylene terephthalate) (PET)-digesting enzyme (PETase) that exhibits improved polymer degradation capacity relative to wild-type PETase due to its narrowed binding cleft via mutation of two active-site residues.

Exemplary embodiments provide method for degrading a polymer comprising contacting the modified PETase of claim 1 or the cell of claim 13 with the polymer. In some methods, the polymer is a polyester. In other methods, the polymer is an aromatic polymer or a semi-aromatic polymer. In some methods, the polymer is polyethylene terephtha!ate (PET). In other methods, the polymer is poly ethyl enefuranoate (PEF). In certain methods the polymer is from a recycled plastic material.

In addition to the exemplary aspects and embodiments described above, further aspects and embodiments will become apparent by reference to the drawings and by study of the following descriptions.

BRIEF DESCRIPTION OF THE DRAWINGS

Exemplary embodiments are illustrated in referenced figures of the drawings. It is intended that the embodiments and figures disclosed herein are to be considered illustrative rather than limiting.

FIG. 1 shows the nucleotide (A) and amino acid (B) sequences of PETase from Ideonel la sakaiensi .

FIG. 2 shows the nucleotide (A) and amino acid (B) sequences of PETase from Ideonella sakaiensis containing the W159H and S238F mutations (bold and underlined).

FIG. 3 shows high resolution X-ray crystallography data collection and analysis of

PETase.

FIG. 4 illustrates the structure of PETase.

FIG. 5 shows PETase sequence analysis.

FIG. 6 illustrates the compari son of the active site cleft of PETase with eutinases FIG. 7 show's multiple sequence alignments of PETase with lipase and cutinase family members. FIG. 8 illustrates the structural and functional analysis of key residues in PETase.

FIG. 9 shows the chemical analysis of polymer substrates.

FIG. 10 illustrates a comparison of PETase and the engineered enzyme S238F/W159H with PET.

FIG. 1 1 shows the induced fit docking analysis of PETase and the engineered enzyme S238F/W159H with PET.

FIG. 12 show's degradation analysis of PBS and PLA by PETase.

FIG. 13 illustrates a comparison of PETase and the engineered enzyme S238F/W159H with PEF.

DETAILED DESCRIPTION

Disclosed herein are engineered enzymes capable of degrading polymers and plastics such as polyethylene terephthalate (PET). Such enzymes include PET-degrading enzymes (PETases) wherein certain amino acid residues are mutated to different amino acids to improve enzymatic activity. One example of an engineered PETase as disclosed herein is provided in FIG. 2, which provides nucleotide and amino acid sequences for a PETase from the bacterium Ideonella sakaiensis wherein the tryptophan residue at position 159 has been mutated to a histidine residue and the serine residue at position 238 has been mutated to a phenylalanine (W159H/S238F).

Ideonella sakaiensis 201-F6 is a bacterial strain with the ability to use PET as its major carbon and energy source for growth. Especially in the last decade, there have been multiple, foundational studies reporting such enzymes that can degrade PET, but previous work has not connected extracellular enzymatic PET degradation to catabolism in a single microbe. Previously, it had been demonstrated that an I. sakaiensis enzyme dubbed PETase converts PET to mono(2-hydroxyethyl) terephthalic acid (MHET), with trace amounts of terephthalic acid (TPA) and bis(2-hydroxyethyl)-TPA (BHET) as secondary products. A second enzyme, MHETase, 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 of PETase highlights similar to ab-hydrolase enzymes, including the cutinase and lipase families, which catalyze hydrolysis of cutin and fatty acids, respectively.

Beyond PET, humankind uses a wide range of polyesters, broadly classified by aliphatic and aromatic content. PET, for example, is a semi-aromatic polyester. Some aliphatic polyesters, such as polylactic acid (PL A), poly butylene succinate (PBS), or polyhydroxyalkanoates can be produced from renewable sources and are marked 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 semi-aromatic polyesters, conversely, often exhibit enhanced thermal and material properties, and accordingly, have reached substantially higher market use, but are typically not as biodegradable as their aliphatic counterparts. An emerging, bio-based PET replacement is polyethylene-2, 5-furandicarhoxylate (or polyethylenefuranoate, called PEF), which is based on sugar-derived 2, 5-fumandi carboxylic acid (FDCA). PEF exhibits improved gas barrier properties over PET and is being pursued industrially. Even though PEF is a bio based semi-aromatic polyester, which is predicted to offset greenhouse gas emissions relative to PET, its lifetime in the environment, like that of PET, is likely to be quite long. Given that PETase has evolved to degrade crystalline PET, it potentially may have promiscuous activity across a range of polyesters.

Exemplary modified enzymes are provided throughout this disclosure. Amino acids for modification may be selected from those found in or near a PETase active site - for example, as determined by reference to the PETase’ s crystal structure. For the PETase exemplified herein, active site residues include T88, S238, H237, S160, D206, W159 or W185.

In various embodiments, the modified enzymes may be from microorganisms such as bacteria, yeast, or fungi. Exemplary bacteria and fungi include species from the genera Ideonella (such as I. sakaiensi ), Thermohifida (such as T. fusca ) or Fusarium (such as F. solani). Though specific examples are provided herein, other examples of modified enzymes or PETases from microbial sources are within the scope of this disclosure.

Also presented are microorganisms engineered to express the modified enzymes disclosed herein and their use to degrade or depolymerize polymers. Polymer degradation/depolymerization may be carried out be culturing such microorganisms with a material containing a polymer and allowing the microorganisms to enzymatically degrade the conversion. Any microorganism capable of expressing the enzymes disclosed herein may be suitable. Exemplary microorganisms include bacteria, such as those from the genus Ideonella. Specific examples include strains of Ideonella sakaiensis, such as /. sakaiensis 201-F6. Polymers or polymer-containing materials (supplemented with media or nutrients as needed) may be contacted with organisms at a concentration and a temperature for a time sufficient to achieve the desired amount of degradation or depolymerization. Suitable times range from a few hours to several days and may be selected to achieve a desired amount of conversion. Exemplary reaction times include 1, 2, 3, 4, 5, 6, 7, 8, 9, 10, 11, or 12 hours; and

0.5, 1, 1.5, 2, 2.5, 3, 3.5, 4, 4.5, 5, 5.5, 6, 6.5, 7, 7.5, 8, 8.5, 9, 9.5, 10, 10.5, 1 1 , 11.5, 12, 12.5,

13, 13.5, 14, 14.5, or 15 days. In some embodiments, reaction times may be one or more weeks. Unpurified or semi-purified culture supernatants containing the modified enzymes may also be contacted with polymers or polymer-containing materials (e.g , in vitro) under similar reaction conditions suitable for allowing polymer degradation. In certain embodiments, the degradation products may be further converted to additional products by further contact with enzymes. For example, PET may be contacted with a modified PETase to generate mono(2- hydroxyethyl) terephthalic acid (MHET), with trace amounts of terephthalic acid (TP A) and bis(2-hydroxyethyl)-TPA (BHET) as secondary' products. These reaction products (or isolated

MHET derived therefrom) may be contacted with a second enzyme, MHETase, that further converts MHET into the two monomers, TPA and ethylene glycol (EG).

Methods of fractionating, isolating or purifying degradation products (or further upgraded products) include a variety of biochemical engineering unit operations. For example, the reaction mixture or cell culture lysate may be filtered to separate solids from products present in a liquid portion. Products may be further extracted from a solvent and/or purified using conventional methods. Exemplary methods for purification/i solation/ separation of products include at least one of affinity chromatography, ion exchange chromatography, solvent extraction, filtration, centrifugation, electrophoresis, hydrophobic interaction chromatography, gel filtration chromatography, reverse phase chromatography, chromatofocusing, differential solubilization, preparative disc-gel electrophoresis, isoelectric focusing, high performance liquid chromatography (HPLC), and/or or reversed-phase HPLC.

Exemplary' polymers include polyesters such as aromatic and semi-aromatic polyesters. Specific examples include polyethylene terephthalate (PET) or polyethylenefuranoate (PEF), which may be present in recycled materials such as beverage bottles, clothing, packaging, or earpetm The sequences disclosed herein provide nucleic acid and amino acid sequences for exemplary enzymes for use in the disclosed methods. "Nucleic acid” or "polynucleotide" as used herein refers to purine- and pyrimidine-containing polymers of any length, either polyribonucleotides or poiydeoxyribonucleotide or mixed polyribo-polydeoxyribonucleotides. This includes single-and double-stranded molecules (/.<?., DNA-DNA, DNA-RNA and RNA- RNA hybrids), cDNAs, as well as "protein nucleic acids" (PNA) formed by conjugating bases to an amino acid backbone. This also includes nucleic acids containing modified bases.

Nucleic acids referred to herein as "isolated" are nucleic acids that have been removed from their natural milieu or separated away from the nucleic acids of the genomic DNA or cellular RNA of their source of origin (e.g., as it exists in cells or in a mixture of nucleic acids such as a library) and may have undergone further processing. Isolated nucleic acids include nucleic acids obtained by methods described herein, similar methods or other suitable methods, including essentially pure nucleic acids, nucleic acids produced by chemical synthesis, by combinations of biological and chemical methods, and recombinant nucleic acids that are isolated.

Nucleic acids referred to herein as "recombinant" are nucleic acids which have been produced by recombinant DNA methodology, including those nucleic acids that are generated by procedures that rely upon a method of artificial replication, such as the polymerase chain reaction (PCR) and/or cloning or assembling into a vector using restriction enzymes. Recombinant nucleic acids also include those that result from recombination events that occur through the natural mechanisms of cells but are selected for after the introduction to the cells of nucleic acids designed to allow or make probable a desired recombination event. Portions of isolated nuclei c acids that code for polypeptides having a certain function can be identified and isolated by, for example, the method disclosed in U.S. Patent No. 4,952,501.

An isolated nucleic acid molecule can be isolated from its natural source or produced using recombinant DNA technology (e.g, polymerase chain reaction (PCR) amplification, cloning or assembling) or chemical synthesis. Isolated nucleic acid molecules can include, for example, genes, natural allelic variants of genes, coding regions or portions thereof, and coding and/or regulatory regions modified by nucleotide insertions, deletions, substitutions, and/or inversions in a manner such that the modifications do not substantially interfere with the nucleic acid molecule's ability to encode a polypeptide or to form stable hybrids under stringent conditions with natural gene isolates. An isolated nucleic acid molecule can include degeneracies. As used herein, nucleotide degeneracy refers to the phenomenon that one amino acid can be encoded by different nucleotide codons. Thus, the nucleic acid sequence of a nucleic acid molecule that encodes a protein or polypeptide can vary due to degeneracies.

Unless so specified, a nucleic acid molecule is not required to encode a protein having enzyme activity. A nucleic acid molecule can encode a truncated, mutated or inactive protein, for example. In addition, nucleic acid molecules may also be useful as probes and primers for the identification, isolation and/or purification of other nucleic acid molecules, independent of a protein-encoding function.

Suitable nucleic acids include fragments or variants that encode a functional enzyme or proteins disclosed herein. For example, a fragment can comprise the minimum nucleotides required to encode a functional PETase or component thereof. Nucleic acid variants include nucleic acids with one or more nucleotide additions, deletions, substitutions, including transitions and transversions, insertion, or modifications (e.g., via RNA or DNA analogs). Alterations may occur at the 5' or 3' terminal positions of the reference nucleotide sequence or anywhere between those terminal positions, interspersed either individually among the nucleotides in the reference sequence or in one or more contiguous groups within the reference sequence.

In certain embodiments, a nucleic acid may be identical to a sequence represented herein. In other embodiments, the nucleic acids mav be at least about 80%, 81%, 82%, 83%, 84%, 85%, 86%, 87%, 88%, 89%, 90%, 91%, 92%, 93%, 94%, 95%, 96%, 97%, 98% or 99% identical to a sequence represented herein, or 80%, 81%, 82%, 83%, 84%, 85%, 86%, 87%, 88%, 89%, 90%, 91%, 92%, 93%, 94%, 95%, 96%, 97%, 98% or 99% identical to a sequence represented herein. Sequence identity calculations can be performed using computer programs, hybridization methods, or calculations. Exemplary computer program methods to determine identity and similarity between two sequences include, but are not limited to, the GCG program package, BLASTN, BLASTX, TBLASTX, and FASTA. The BLAST programs are publicly available from NCBI and other sources. For example, nucleotide sequence identity can be determined by comparing query sequences to sequences in publicly available sequence databases (NCBI) using the BLASTN2 algorithm. Embodiments of the nucleic acids include those that encode the polypeptides that possess the enzymatic activities described herein or functional equivalents thereof. A functional equivalent includes fragments or variants of these that exhibit one or more of the enzymatic activities. As a result of the degeneracy of the genetic code, many nucleic acid sequences can encode a given polypeptide with a particular enzymatic activity. Such functionally equivalent variants are contemplated herein.

Nucleic acids may be derived from a variety of sources including DNA, cDNA, synthetic DNA, synthetic RNA, or combinations thereof. Such sequences may comprise genomic DNA, which may or may not include naturally occurring introns. Moreover, such genomic DNA may be obtained in association with promoter regions or poly (A) sequences. The sequences, genomic DNA, or cDNA may be obtained in any of several ways. Genomic DNA can be extracted and purified from suitable cells by means well known in the art. Alternatively, mRNA can be isolated from a cell and used to produce cDNA by reverse transcription or other means.

Also disclosed herein are recombinant vectors, including expression vectors, containing nucleic acids encoding enzymes. A“recombinant vector” is a nucleic acid molecule that is used as a tool for manipulating a nucleic acid sequence of choice or for introducing such a nucleic acid sequence into a host ceil. A recombinant vector may be suitable for use in cloning, assembling, sequencing, or otherwise manipulating the nucleic acid sequence of choice, such as by expressing or delivering the nucleic acid sequence of choice into a host cell to form a recombinant cell. Such a vector typically contains heterologous nucleic acid sequences not naturally found adjacent to a nucleic acid sequence of choice, although the vector can also contain regulatory _' nucleic acid sequences (e.g., promoters, untranslated regions) that are naturally found adjacent to the nucleic acid sequences of choice or that are useful for expression of the nucleic acid molecules.

The nucleic acids described herein may be used in methods for production of enzymes or proteins through incorporation into ceils, tissues, or organisms. In some embodiments, a nucleic acid may be incorporated into a vector for expression in suitable host cells. The vector may then be introduced into one or more host cells by any method known in the art. One method to produce an encoded protein includes transforming a host cell with one or more recombinant nucleic acids (such as expression vectors) to form a recombinant cell. The term “transformation” is generally used herein to refer to any method by which an exogenous nucleic acid molecule (i.e., a recombinant nucleic acid molecule) can be inserted into a cell but can be used interchangeably with the term "transfection."

Non-limiting examples of suitable host cells include cells from microorganisms such as bacteria, yeast, fungi, and filamentous fungi. Exemplary microorganisms include, but are not limited to, bacteria such as E. coir, bacteria from the genera Ideonella (e.g., I. sakaiensis ), Thermobifidki (e.g., T. fused), Pseudomonas (e.g., P. pulida or P. fluorescens), Acinetobacter (e.g., strains of A. baylyi such as ADP1), Bacillus (e.g., B. subtilis, B. megaterium or B. brevis), Caulobacter (e.g., C. crescentus), Lactoccocus (e.g, L. lactis), Streptomyces (e.g., S. coelicolor), Streptococcus (e.g., S. lividans), and Corynybacterium (e.g., C. glutamicum); fungi from the 5 genera Fusarium (e.g., F. soiani), Trichoderma (e.g., T. reesei, T. viride, I koningu or T. harzianum), PeniciUimn (e.g., P. fimiculosum), Humicola (e.g., H. insolens), Chrysosporium (e.g., C. luclmowense), Gliocladium, Aspergillus (e.g, A. niger, A. nididans, A. aw amor i, or A. aculeatus), Neurospora, Hypocrea (e.g., H. jecorind), and. Emericella; yeasts from the genera Saccharomyces (e.g., S. cerevisiae), Pichia (e.g., P. pastoris ), or Kluyveromyces (e.g, K. lactis). 10 Ceils from plants such as Arabidopsis, barley, citrus, cotton, maize, poplar, rice, soybean, sugarcane, wheat, switch grass, alfalfa, miscanthus, and trees such as hardwoods and softwoods are also contemplated herein as host cells.

Host cells can be transformed, transfected, or infected as appropriate by any suitable method including electroporation, calcium chloride-, lithium chloride-, lithium acetate/polyene glycol-, calcium phosphate-, DEAE-dextran-, liposome-mediated DNA uptake, spheroplasting, injection, microinjection, microprojectile bombardment, phage infection, viral infection, or other established methods. Alternatively, vectors containing the nucleic acids of interest can be transcribed in vitro , and the resulting RNA introduced into the host cell by well- known methods, for example, by injection. Exemplary embodiments include a host cell or population of cells expressing one or more nucleic acid molecules or expression vectors described herein (for example, a genetically modified microorganism). The cells into which nucleic acids have been introduced as described above also include the progeny of such cells.

Vectors may be introduced into host cells such as those from bacteria or fungi by direct transformation, in which DNA is mixed with the cells and taken up without any additional manipulation, by conjugation, electroporation, or other means known in the art. Expression vectors may be expressed by bacteria or fungi or other host cells episomally or the gene of interest may be inserted into the chromosome of the host cell to produce cells that stably express the gene with or without the need for selective pressure. For example, expression cassettes may be targeted to neutral chromosomal sites by recombination.

Host cells carrying an expression vector (i.e., transformants or clones) may be selected using markers depending on the mode of the vector construction. The marker may be on the same or a different DNA molecule. In prokaryotic hosts, the transformant may be selected, for example, by resistance to ampicillin, tetracycline or other antibiotics. Production of a particular product based on temperature sensitivity may also serve as an appropriate marker.

Host cells may be cultured in an appropriate fermentation medium. An appropriate, or effective, fermentation medium refers to any medium in which a host cell, including a genetically modified microorganism, when cultured, is capable of growing or expressing the polypeptides described herein. Such a medium is typically an aqueous medium comprising assimilable carbon, nitrogen and phosphate sources, but can also include appropriate salts, minerals, metals and other nutrients. Microorganisms and other cells can be cultured in conventional fermentation bioreactors and by any fermentation process, including batch, fed- batch, cell recycle, and continuous fermentation. The pH of the fermentation medium is regulated to a pH suitable for growth of the particular organism. Culture media and conditions for various host cells are known in the art. A wide range of media for culturing bacteria or fungi, for example, are available from ATCC. Media may be supplemented with aromatic substrates, or components of thermochemical waste streams as needed.

The nucleic acid molecules described herein encode the enzymes with amino acid sequences such as those presented herein. As used herein, the terms "protein" and "polypeptide" are synonymous. "Peptides” are defined as fragments or portions of polypeptides, preferably fragments or portions having at least one functional activity _' ^· as the complete polypeptide sequence. "Isolated" proteins or polypeptides are proteins or polypeptides purified to a state beyond that in which they exist in cells. In certain embodiments, they may be at least 10% pure; in others, they may be substantially purified to 80% or 90% purity' or greater. Isolated proteins or polypeptides include essentially pure proteins or polypeptides, proteins or polypeptides produced by chemical synthesis or by combinations of biological and chemical methods, and recombinant proteins or polypeptides that are isolated. Proteins or polypeptides referred to herein as "recombinant" are proteins or polypeptides produced by the expression of recombinant nucleic acids.

Proteins or polypeptides encoded by nucleic acids as well as functional portions or variants thereof are also described herein. Polypeptide sequences may be identical to the amino acid sequences presented herein or may include up to a certain integer number of amino acid alterations. Such protein or polypeptide variants retain enzymatic activity, and include mutants differing by the addition, deletion or substitution of one or more amino acid residues, or modified polypeptides and mutants comprising one or more modified residues. The variant may have one or more conservative changes, wherein a substituted amino acid has similar structural or chemical properties (e.g., replacement of leucine with isoleucine). Alterations may occur at the amino- or carboxy-terminai positions of the reference polypeptide sequence or anywhere between those terminal positions, interspersed either individually among the amino acids in the reference sequence or in one or more contiguous groups within the reference sequence.

In certain embodiments, the polypeptides may be at least about 70%, 71%, 72%, 73%,

74%, 75%, 76%, 77%, 78%, 79%, 80%, 81%, 82%, 83%, 84%, 85%, 86%, 87%, 88%, 89%, 90%, 91 %, 92%, 93%, 94%, 95%, 96%, 97%, 98%, or 99% identical to the amino acid sequences set forth in the sequences provided herein and possess enzymatic function. Percent sequence identity can be calculated using computer programs (such as the BLASTP and TBLASTN programs publicly available from NCBI and other sources) or direct sequence comparison. Polypeptide variants can be produced using techniques known in the art including direct modifications to isolated polypeptides, direct synthesis, or modifications to the nucleic acid sequence encoding the polypeptide using, for example, recombinant DNA techniques.

Polypeptides may be retrieved, obtained, or used in "substantially pure" form, a purity that allows for the effective use of the protein in any method described herein or known in the art. For a protein to be most useful in any of the methods described herein or in any method utilizing enzymes of the types described herein, it is most often substantially free of contaminants, other proteins and/or chemicals that might interfere or that would interfere with its use in the method (e.g., that might interfere with enzyme activity), or that at least would be undesirable for inclusion with a protein. Disclosed herein are multiple high-resolution X-ray crystal structures of PETase, which enable comparison to known cutinase structures. Based on differences between the PETase and a homologous cutinase active-site cleft, PETase variants were produced and tested for PET degradation, including a double mutant distal to the catalytic center that altered important substrate-binding interactions. In some embodiments, this double mutant may have W159H/S238F mutations. Surprisingly, this double mutant, inspired by cutinase architecture, exhibits improved PET degradation capacity relative to wild-type PETase. In silica docking and molecular dynamics (MD) simulations were deployed to characterize PET binding and dynamics, which provide insights into substrate binding and may suggest an explanation for the improved performance of the PETase double mutant. Additionally, incubation of wild-type and mutate PETase with several polyesters was examined using scanning electron microscopy (SEM), differential scanning calorimetry (DSC), and product release. These studies showed that not only can the PETase degrade crystalline PET, but that it can also depolymerize PEE. However, it does not appear PETase is capable of depoly merizing aliphatic polyesters, suggesting a broader role for PETase in degrading semi-aromatic 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 biotechnical strategy to help remediate the environmental scourge of plastic accumulation in nature. The present disclosure may enable PET and PEF to be degraded at faster rates than available using unmodified PETase.

The discovery of a bacterium that uses PET as a major carbon and energy source (i.e , PETase) 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 present disclosure demonstrates that a collection of subtle variations on the surface of a lipase/cutinase-like fold has the ability to endow a modified and/or mutated 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 bio-accumulation and bio-based industrial polyester recycling. The Examples discussed herein are provided for purposes of illustration and are not intended to be limiting. Still other embodiments and modifications are also contemplated. Example 1: PETase exhibits a canonical ab-hydrolase structure with an open active-site cleft.

The high-resolution X-ray crystal structure of the I. sakaiensis PETase was analyzed employing a synchrotron beamline capable of long-wavelength X-ray crystallography. Using single wavelength anomalous dispersion, phases were obtained from the native sulfur atoms present in the protein. The low background from the in vacuo setup and large curved detector resulted in exceptional diffraction data quality extending to a resolution of 0.92 A, with minimal radiation damage (Table l, FIG. 3).

Table 1 : Crystallographic data and refinement statistics including crystallization conditions.

As shown in Table 1 , the completeness and anomalous completeness are the signal to noise ratio of intensities, with the highest resolution bin in brackets/parenthesis. The Rmerge (Rm) of Table 1 was calculated using Equation 1 :

Equation 1

Where Ifh.ij is symmetry-related intensities and 1(h) is the mean intensity of the reflection with unique index h. In Table 1, Cc ½ is the correlation coefficient of the mean intensities between two random half-datasets. In Table 1 , multiplicity refers to multiplicity for unique reflections. The ratio Rwork/Rfree (expressed as a percentage) as shown in Table 1 w'as calculated by randomly selecting 5% of reflections for determination of the free R factor, prior to any refinement. In Table 1, the B average is the temperature factors averaged for all atoms. In Table I, the line“geometry bond, angles” show RMS deviations from ideal geometry for bond lengths and restraint angles (Engh and Huber). In Table 1, the Ramachandran is the percentage of residues in the‘most favored region’ of the Ramachandran plot and percentage of outliers (MOLPROBITY). In Table 1 , PDB ID refers to the protein data bank identifiers for coordinates. The crystallography conditions analyzed were: long l, 0.1M MIB (Malonate, Imidazole, Borate), pH 5.0, 25% polyethylene glycol (PEG) 1500. Native 1, 0.2 M MgC , 0.1 M MES (2-(N-Morpholino)ethanesulfonic acid), pH 6.0, 20% PEG 6000. Native 2, 0.2 M NHrCl, 0.1 M MES, pH 6.0, 20% PEG 6000. Native 3, 0.2 M LbSOr, 0.1 M Bis-Tris (pH 5.5), 25% PEG 3350. Native 4, 1.6 M MgSOr, 0.1M MES, pH 6.5.

FIG. 3 shows high resolution X-ray crystallography data collection and analysis for PETase. Panel A of FIG 3 illustrates a representative section of a diffraction image from the 0.92 A PETase structure determination. The mean counts in the boxes are A (low resolution): 0.3 counts, B (solvent ring): 1.6 counts and C (high resolution): 0 1 counts. The inset show's an enlarged portion around reflection -49 13 -11 at 1.0 A resolution (box D) where each box represents an individual pixel. Values range from zero (white) and 1 (light grey) background counts, through to the center of this refection at 16 counts. Panel B of FIG. 3 illustrates representative electron density from the high-resolution structure (6EQE) showing continuous density' across the disulfide bridge between Cys273 and Cys289. The 2Fo-Fc map is contoured at 1s. Panel C of FIG. 3 illustrates representative electron density quality is shown centered around Trp257. The 2Fo-Fc map is contoured at 1 s. As predicted from the sequence homology to the lipase and cutinase families, PETase adopts a classical ab-hydrolase fold, with a core consisting of 8 b-strands and 6 a-helices (Panel A of FIG. 4). It had been demonstrated previously that PETase has sequence identity substantially close to bacterial cutinases, with Thermobifida fit sea cutinase being the closest known structural representative with 52% sequence identity (Panel B of FIG. 4, Panel A of FIG. 5), which is an enzyme that also degrades PET. Despite a conserved fold, the surface profile is quite different between the two enzymes. PETase has a highly polarized surface charge (Panel C of FIG 4), creating a dipole across the molecule, and resulting in an overall isoelectric point (pi) of 9.6. In contrast, T. fiisca cutinase, in common with other cutinases, has a number of small patches of both acidic and basic residues distributed over the surface conferring a more neutral pi of 6.3 (Panel D of FIG. 4).

FIG. 4 illustrates the structure of PETase. Panel A of FIG. 4 illustrates a cartoon representation of the PETase structure at 0.92 A resolution (PDB ID: 6EQE). The active site cleft is oriented at the top and highlighted with a dashed circle. Panel B of FIG 4 illustrates a comparative structure of the T fiisca cutinase (PDB ID: 4CG1). Panel C of FIG. 4 illustrates the electrostatic potential distribution mapped to the solvent accessible surface of PETase compared to the T. fiisca cutinase as a shaded gradient from black (acidic) -7 kTle to a lighter gray (basic) 7 kTle. Panel D of FIG. 4 illustrates the T. fiisca cutinase in the same orientation. Panel E of FIG 4 shows the view along the active site cleft of PETase corresponding to the area highlighted with a dashed circle in A and C. The width of the cleft is shown between Thr88 and Ser238 Panel F of FIG. 4 shows the narrower cleft of T. fiisca cutinase active site is shown with the width between Thr61 and Phe209 in equivalent positions. Panel G of FIG. 4 illustrates a close-up view of the PETase active site with the catalytic triad residues, His237, Seri 60, and Asp206, shaded. Residues Trpl59 and Trpl85 are lightly shaded. FIG. 4H shows a comparative view' of the T. fusca cutinase active site with equivalent catalytic triad residues shaded. Residues Hisl29 and Trpl 55 are lightly shaded. The residues in PETase are also shaded that correspond to the site-directed mutagenesis targets S238F, W159H, and W185A.

FIG. 5 shows PETase sequence analysis. Panel A of FIG. 5 show's the sequence alignment of PETase (labelled PET, accession number: A0A0K8P6T7), against a PET- degrading cutinase from T. fiuscia (TFcut, accession number: AET05798). The signal sequences from both enzymes, as predicted by LipoP 1.0 Server, were excluded from the alignment for clarity. The squares shaded dark gray indicate areas of identity with residues in text indicating moderate conservation. The cartoons above the alignment denote secondary structure, with spirals representing alpha-helices, arrows representing beta-strands, and Ί’ indicating turns. Differences in key residues around the active site are highlighted with a box, with residues 159 to 162 denoting the position of the conserved lipase box. The box spanning residues 237-238 denotes the presence of serine in PETase next to the catalytic histidine, as compared to a phenylalanine in I fusca. The numbering below the sequences indicates the position of residues forming productive disulfide bridges as observed in the PETase crystal structure reported here. Panel B of FIG. 5 shows the nucleotide sequence of the synthetic DNA fragment containing the gene encoding the PETase from I. sakaiensis optimized for expression in E. coli. Overlaps added for assembly into pET-21b(+) are underlined. The initiating ATG and stop codon of the His-tagged gene are in bold text. Ndel and Xhol restriction sites are capitalized.

Another difference between PETase and the closest cutinase homologues is the broader active-site cleft, which was hypothesized might be necessary to accommodate crystalline semi aromatic polyesters. At its widest point, the cleft in PETase approaches three-times the width of the corresponding structure in the T. fusca cutinase. The expansion is achieved with minimal rearrangement of the adjacent loops and secondary structure (Panels E and F of FIG. 4). A single amino acid substitution from phenylalanine to serine in the lining of the active site cavity appears sufficient to cause this change, with the remaining deft formed between Trpl59 and Tip 185 (Panel G of FIG. 4) This relative broadening of the active site cleft is also observed in comparisons with other known cutinase structures (Panels A-D of FIG. 6).

FIG. 6 illustrates the comparison of the active site cleft of PETase with cutinases. Panel A of FIG. 6 show's a PETase cleft width calculated from the distance between the van der Waals surface of TBS and S238 PDB ID: 6EQE. Panel B of FIG. 6 illustrates Tfusica cutinase with cleft width calculated from the distance between the van der Waals surface of T61 and F209 (PDB ID: 4CG1) (12). Panel C of FIG. 6 show _' s Thermobifida cellulosilytica cutinase (PDB ID: code 5LUI) with cleft width calculated from the distance between the van der Waals surface of T63 and F212. Panel D of FIG. 6 shows leaf compost cutinase obtained from a metagenomics analysis of uncultured bacteria. Cleft width was calculated from the distance between the van der Waals surface of T61 and F210 (PDB ID: 4EBQ). Arrow's with labels denote the width of the active site as determined from equivalent residues in each enzyme. Panel E of FIG. 6 shows hydrophobic adaptations in the active site of PETase with the catalytic residue, Seri 60, darkly shaded. Residues in a lighter shade indicate the hydrophobic residues surrounding the active site triad. Panel F of FIG 6 shows a comparative view of the T. fusca hydrophobic distribution, with equivalent orientation and shading.

FIG. 7 show's multiple sequence alignments of PETase with lipase and cutinase family members. Panel A of FIG. 7 illustrates multiple sequence alignment of PETase against members of the lipase family. Signal sequences, as predicted by LipoP 1.0 Server, were excluded from the alignment. As in Panel A of FIG. 5, the box spanning residues 159 to 162 highlights the conserved lipase box, while the box spanning 237-238 indicates the serine in PETase, which is occupied by a phenylalanine or tyrosine in most lipases. Aligned sequences with accession numbers are PET (PETase, I. sakaiensis , A0A0K8P6T7), CTlip (lipase, Ca!dimonas taiwanensis WP 06219554 -I }, PSlip (lipase, Pseudomonas saudimassiliensis CEA05385), OAlip (lipase, Oleispira Antarctica, CCK74972.1), SSlip (lipase, Saccharothrix sp, OKI36883.1), PYlip (tricylglycerol lipase, Pseudomonas yangmingensis, SFM35944), YPlip (lipase, Vibro palustri , SJL84994), RGlip (lipase, Rhizobacier gumtniphilus , ARN20166), VSlip (Lipase, Vibro spartinae , 81095186), and HClip (lipase, Herbidospora cretacea , WP 061298849). FIG. 7B show's the multiple sequence alignment of PETase against members of the cutinase family with the same scheme. Aligned sequences are PET (PETase, Ideonella sakaiensis, A0A0K8P6T7), PBeut (cutinase, Pseudomonas bauzanensis, SER72431), Picut (cutinase, Pseudomonase litoralis , SDS35700), PS cut (cutinase, Pseudomonas sale ge s, 81)1 28434), PXcut (cutinase, Pseudomonas xinjiangensis, SDS09569), MEcut (cutinase, Micromonospora echinospora, SCF30318), TAcut (cutinase, Thermobifida alba, ADV 92525), TFcut (cutinase, T. iusca, AET05798), TCcut (cutinase, Thermobifida celhdosilytica , E9EUΉ8), and LCcut (cutinase, uncultured bacteria isolated from metagenomics analysis, AEV21261). Annotations are consistent with FIG. 5A.

In terms of the active site, the catalytic triad is conserved across the lipases and cutinase families. In PETase, the catalytic triad comprises Serl60, Asp206, and His237, suggesting a charge-relay system similar to that found in other a/b-fold hydrolases. The specific location and geometry between the active site found in cutinases is also conserved in PETase (Panels G and H of FIG. 7). In common with most lipases, the catalytic residues reside on loops, with the nucleophilic serine occupying a highly conserved position known as the nucleophilic elbow. The nucleophilic serine sits in the consensus sequence (Gly-Xl-Ser-X2-Gly), and while this "lipase box” is common to most lipases (Panel A of FIG. 7) and cutinases (Panel B of FIG. 7), the XI position, usually occupied by a histidine or phenylalanine in cutinases and lipases contains a tryptophan residue, TrpJ 59, in PETase (Panel G of FIG. 4). This residue has the effect of extending the hydrophobic surface adjacent to the active site (Panels E and F of FIG. 6). In common with the Fusarium solani cutinase, PETase has two disulfide bonds, one adjacent to the active site and one near the C-terminus of the protein. MD simulations have predicted that the active site disulfide in F. solani cutinase is important for active site stability, and it may play a similar role in PETase.

FIG. 8 illustrates the structural and functional analysis of key residues in PETase. Panel A of FIG. 8 shows the superposition of the PETase structures (PDB IDs: 6EQE, 6EQF, 6EQG, and 6EQH). The RMSD of chains is 0.28A ± 0.02. Overall, all the chains adopt the same fold with only slight variations in the loops. The sideehains of the three residues in the catalytic triad, Serl60, Asp206, and His237 in addition to Trpl59 and Trpl85 are shown. All side-chain adopt the same rotameric state with the exception of Trpl85, where two residues adopted slightly different c-2 orientations, possibly reflecting a degree of mobility. Panel B of FIG. 8 shows the superposition of the three PETase domains from 6EQG with residues and sideehains shaded according to B-factors All the residues in the catalytic triad and Trpl59 have considerably lower B-factors compared to Trpl85. The increased B-factors indicate a greater mobility of this sidechain with respect to all of the other residues in the hydrophobic cleft and the catalytic triad. Panels C-E of FIG. 8 show surface views of the binding cleft from three independently refined chains in the asymmetric unit of crystal form 4, in space group P2i (PDB ID: 6EQG). The position of the Trpl85 residue is highlighted in each case. Panel F of FIG. 8 illustrates a MD simulation demonstrating a wide range of movement for residue Trpl85 (shaded). The catalytic triad residues, Serl60, Asp206, and His 237, are highlighted as well. Panel G of FIG. 8 shows PET degradation after incubation with buffer only. Panel H of FIG. 8 shows PET degradation after incubation with the W185A mutant PETase. Panel I of FIG. 8 shows PEF degradation after incubation with buffer only. Panel J of FIG. 8 shows PEF degradation after incubation with the Wl 85 A mutant PETase To explore the potential effects of crystallization conditions and packing effects, three additional crystallography datasets ranging in resolution from 1.58 to 1.80 A provided a total of seven independent PETase chains (Table 1). All domains adopt the same fold (relative roo mean square deviation (RMSD) values are -0.28 A) and all the residues of the catalytic triad exhibit the same conformation (Panels A and B of FIG. 8), including Trpl59. In one crystal form, however, Trpl85 was present in three distinct conformations, ail with higher B-factors than other residues in the putative binding cleft; these results were corroborated by MD simulations of the wild-type PETase (Panels C-F of FIG. 8). In all crystal forms, the packing of PETase involves extensive packing interactions in and around the hydrophobic cleft, resulting in little space for interaction with putative ligands.

Example 2: Converting PETase to a cutinase-like active-site cleft enables improved crystalline PET degrada tion

From the PETase structure, it was originally hypothesized that changes in the active site relative to the T fitsca cutinase resulted from the evolution of I. sakaiensis in a PET- containing environment, thus enabling more efficient PET depolymerization. To test this hypothesis, the PETase active site was mutated to make it more cutinase-like. Specifically, a double mutant was produced, S238F/W159H, which based on homology modeling, was predicted to narrow the PETase active site, similar to the T. fusca cutinase. Additionally, we produced the W185A mutant to examine the role of this highly conserved, dynamic residue.

FIG. 9 shows the chemical analysis of polymer substrates. Panel A of FIG. 9 shows ¾ nuclear magnetic resonance (NMR) spectra of the lab synthesized PET used in the study. Panel B of FIG. 9 shows ^l H NMR spectra of the lab synthesized PEF used in the study. Panel C of FIG. 9 shows ¹ H NMR spectra of the lab synthesized PL A. Panel D of FIG. 9 shows ¹ H NMR spectra of poly(butylene succinate). Panel E of FIG. 9 shows the reduction in crystallinity in samples before and after digestion as determined by DSC. Panel F of FIG. 9 shows representative DSC trace for PET. After digestion, the melting transition is broadened indicating a reduction in both crystallinity and crystal domain size.

It has been demonstrated that PETase digestions of amorphous PET films with a crystallinity of 1.9%, which is lower than most PET samples that would be either encountered in the environment or in an industrial recycling context were examined. To examine the performance in the wild-type PETase relative to the two mutants, this present disclosure examined PET digestion with coupons of higher crystallinity. Specifically, PET coupons with an initial crystallinity of 14.8±0.2% (for reference, a commercial soft drink botle examined via the same methods exhibits a crystallinity of 15.7% as measured by DSC) were synthesized and characterized by nuclear magnetic resonance (NMR) spectroscopy to confirm their structure and DSC to determine their crystallinity (Panel A of FIG. 9). Digestions were conducted at pH 7.2 and monitored with DSC, NMR spectroscopy, and SEM, and reaction products were quantified by EIPLC and NMR spectroscopy. Panels A-D of Fig. 10 show' the results of PET degradation, including a buffer-only control, the wild-type PETase, and the double mutant. It is clear that PETase induces surface erosion and pitting of a PET film with a crystallinity of 13.3±0.2%, resulting in a 10.1% relative crystallinity reduction (absolute reduction of 1.5%; Table 2). Surprisingly, the PETase double mutant outperforms the wild- type PETase by both crystallinity reduction and product release. The absolute crystallinity loss is 4.13% higher, and the corresponding SEM images appear that slightly more surface ablation occurs (Panel C of FIG. 5). After incubation, the digested PET samples for both the wild-type PETase and the double mutant exhibit a lower melting temperature over a wider temperature range (Panel F of FIG. 9), indicating that the crystalline domain regions are reduced in size.

Table 2: Crystallinity determined by differential scanning calorimetry for plastics with PETase and mutations for the aromatic polyesters.

For the data shown in Table 2, the average cold crystallization for the PET samples was 24 J/g. For the crystallinity of PBS shown in Table 2, the error in PBS crystallinity was outside significant MG s Additionally, for the crystallinity of PBS shown in Table 2, the results with the PETase double mutant exhibited no change in majored properties.

FIG. 10 illustrates a comparison of PETase and the engineered enzyme S238F/W159H with PET. Panel A of FIG. 10 show's buffer-only control of PET coupon. Panel B of FIG. 10 illustrates PET coupon after incubation with wild-type PETase. Panel C of FIG. 10 shows PET coupon after incubation with the PETase double mutant, S238FVW 159H. Ail SEM images were taken after 96 h of incubation at a PETase loading of 50 nM, pH 7.2 in phosphate buffer, or a buffer-only control. Panel D of FIG. 10 show's percent crystallinity change and reaction product concentration after incubation with buffer, wild-type PETase, and the S238FAV159H engineered enzyme. Panel E of FIG. 10 shows predicted binding conformations of wild-type PETase from docking simulations demonstrate that PET is accommodated in an optimum position for the interaction of the carbon (black) with the nucleophilic hydroxyl group of Serl60, at a distance of 5.1 A (dark dashed line). His237 is positioned within 3.9 A of the Ser i 60 hydroxyl (lighter dashed line). Residues Trpl59 and Ser238 line the active site channel (both are shaded). Panel F of FIG 10 show's the double mutant S238F/WI 59H adopts a more productive interaction with PET. The S238 mutation provides new p-stacking and hydrophobic interactions to adjacent terephthalate moieties while the conversion to Hisl 59 from the bulkier Tip allows the PET polymer to sit deeper within the active site channel. TWO aromatic interactions of interest between PET and Phe238 are at optimal distance (each at 5.4 A).

FIG. 11 show's the induced fit docking analysis. All distances shown are in angstroms. Panel A of FIG 11 shows the lowest energy, catalytically competent predicted pose of PEET tetramer in wild-type PETase, XP score of -8.23 kcal/mol; catalytic triad is intact, W185 and W159 stablize PET at optimal distances through parallel displaced and edge-to-face aromatic interactions, respectively. Panel B of FIG. 11 show's the lowest energy, catalytically competent predicted pose of PEF tetramer in wild-type PETase, XP score of -9.07 kcal/mol; catalytic triad is in act, WI 85 and W159 stablize PEF at optimal distances through parallel displaced and edge-to-face aromatic interactions, respectively. Panel C of FIG. 11 show's the lowest energy, catalytically competent pose of PET tetramer in double mutant PETase, XP score of -11.25 kcal/mol; catalytic triad is intact. Here, PET is stabilized by four optimal aromatic contacts: edge-to-face to Trpl85, parallel displaced to Tyr87, point-to-face to Phe238, parallel displaced to Phe 238 (i.e., Phe238 participates in aromatic interactions from two terephthalate units). Panel D of FIG. 11 shows the lowest energy, catalyticaHy competent pose of PEF tetramer in double mutant PETase, XI ³ score of -10.07 kcal/mole. Here, PEF is stabilized through three optimal aromatic interactions: parallel displaced to Trpl85, parallel displaced to Phe238, and a point-to-face interaction with His237. Due to the occupation of His237 with aromatic stabilization, it is not oriented within the catalytic triad, but rather Hisl59 supports Seri 60 via hydrogen bonding. This potential role of Hi si 59 as secondary hydrogen bonding support w'as independently observed by us through molecular dynamics simulations of the apo PETase double mutant. It should also be noted that His237 is within single bond rotation of catalytic triad participation, thus one could hypothesize it may assume this catalytic position after initial acyl-enzyme formation and protonation of Hisl59. Panel E of FIG. 11 shows a surface view of PET docked in the PETase wild-type structure, amino acids are shaded according to their properties. Panel F of FIG. 11 shows a surface view of PETdocked in PETase double mutant structure. Panel G of FIG. 11 shows a surface view of PEF docked in PETase wild type. Panel H of FIG. 11 shows a surface view of PEF docked in PETase double mutant structure. Panels E-H of FIG 1 1 show how PET and PEF occupy the same binding channel in both wild type and mutant proteins; also the S238F mutation changes the nature of the binding cleft. Panel I of FIG 1 1 shows an overlay of Trpl85 position in PETase crystal structure (shown in white), when PET is flexibly docked (shaded) and PEF is flexibly docked (lightly shaded). In response to PET/PEF binding, Trpl 85 rotates to provide optimal aromatic interactions for stabilization. In PETase crystal structure, the N-Ca-Cp-C dihedral is -177.5°, whereas when PET is docked the same dihedral has a value of 98.4°, and -146.4° with PEF docked. This dihedral is also flexible in the double mutant structure, having values of 178.8 with PET and -155.5 with PEF flexibly docked. Trpl85 is not the only flexible residue in the binding site, such rotations were also seen for H237, W159 (WT), H159 (mut), and F238 (mut). The rotation of these residues illustrates the importance of modeling induced fit effects for predicting ligand binding modes. W185’s flexibility was also captured with molecular dynamics simulations.

Understanding how PET binds in the PETase catalytic site aided with developing the PETase double mutant as described in some embodiments herein and to understanding the improved performance of the PETase double mutant. Multiple trials were attempted to obtain a ligand-bound structure of PETase to no avail. Predicting PET-PETase binding modes by conducting Induced Fit Docking (IFD) was attempted (Panel A of FIG. 11). Multiple PET orientations were predicted by IFD in and around the active site of both the wild-type and double mutant enzyme. The orientation shown in Panel E of FIG. 10 (and Panel A of FIG. 11) is one of several to illustrate a productive PET binding event in the wild-type enzyme: a PET carbonyl carbon is at a chemically relevant distance (5.1 A) for nucleophilic attack from the Seri 60 hydroxyl group, His237 is at an ideal distance (3.9 A) to activate Seri 60, and Asp206 provides hydrogen bonding support to His237 (2 8 A). This binding mode is predicted to have binding affinity (estimated by the docking score with descriptors in Table 3) of -8.23 kcal/moi (Table 3). Thus, our IFD predicted binding modes are consistent with a productive Michaelis complex for PET chain cleavage. Additionally, with this low energy, catalytically competent pose generated from flexible docking (i.e., IFD), a marked difference in the position of Trpl85 compared to the crystal structure was observed (Panel I of FIG. 11). The N-Co-Cp-C dihedral in the crystal PETase structure is -177.5°, whereas the predicted catalytically competent binding mode of PET indicates W185 rotates to accommodate aromatic interactions with PET and thus adopts a dihedral angle of 98.4°. This dihedral rotation was observed to various extents in all docking results and in apo MD simulations (Panel F of FIG. 8), and thus illustrates the necessity for flexible protein treatment during ligand binding mode prediction, especially if binding and/or cataly tic hy potheses are to be posited Table 3 : XP Descriptors for each of the discussed binding modes.

As shown in Table 3, XP descriptors are an energetic decomposition of the Glide XP score, which itself is an estimate of the binding affinity. Thus, XP descriptors provide insight into which energetic terms contribute most significantly to binding free energy. All values shown in Table 3 are in kcal/mol. It may be seen E _vd w contributes most significantly to all poses and is even more favorable in mutant binding likely due to increased aromatic interactions with Phe238. Note that in Table 3“Mut” refers to the S238H/W159H double mutant PETase structure as described some embodiments herein.

IFD results also suggest potential reasons for the improved performance of PETase double mutant over wild type PETase, as the substrate may interact with Phe238 through several aromatic interactions, as shown in Panel F of FIG. 10, In this predicted pose (docking score -11.25 kcal/mol, with descriptors in Table 3), a PET carbonyl is in appropriate attack distance from Seri 60 (3.1 A), Seri 60 is in range for deprotonation by His237 (2.9 A, Panel C of FIG. 11), and Asp206 is ready to accept a proton in the shuttle (2.9 A). PET aromatic rings are within ideal p-stacking distances to several binding site residues (W185, Y87), and in particular two aromatic interactions are formed to Phe238 (point-to-face interaction at 5 4 A, and parallel displaced interaction at 5.4 A). The marked difference in predicted binding affinities between WT and double mutant enzymes for PET is consistent with the increased activity of the PETase double mutant on PET, as observed experimentally, and we can identify aromatic interactions supported by the S238F mutation as being integral to this enhancement All aromatic ring-ring distances for described binding modes are illustrated in Panels A and C of FIG. 11.

Conversely to the double mutant, the W185A mutant exhibits highly impaired performance relative to the wild-type PETase, as described in Table 2. These data confirm a critical role for this residue. From the IFD, Trpl 85 is predicted to play an important role by contributing p-stacking interactions to PET aromatic groups. Additionally, in all productive binding modes (i.e., when the carbonyl is oriented to be in the oxyanion hole, and the carbonyl carbon at a catalytic distance from Seri 60), Trpl85 is predicted to reorient relative to the crystal structure, suggesting its movement opens the active site cleft, allowing PET binding (Panel I of FIG. 11).

Example 3: PETase depolymerizes PEE, but not aliphatic polyesters.

Understanding the activity of wild-type PETase and the PETase double mutant on other polymeric substrates, including aliphatic and other semi-aromatic polyesters is also of interest. To that end, aliphatic polyesters, PBS and PEA, were synthesized, characterized (Panels C and D of FIG. 9), and conducted similar incubations. None of these samples showed visual differences between the control images and the PETase-treated samples (FIG. 12), suggesting that PETase and the double mutant are not active on aliphatic polyesters. FIG. 12 shows degradation analysis of PBS and PLA by PETase. Panel A of FIG 12 shows a PBS coupon before incubation. Panel B of FIG. 12 shows a PBS coupon in buffer only. Panel C of FIG. 12 shows a PBS coupon in buffer with PETase. Panel D of FIG. 12 show's PLA film before incubation. Panel E of FIG. 12 shows a PLA film after incubation in buffer only. Panel F of FIG 12 shows PLA after incubation in buffer with PETase. All images w ^? ere taken after 96 hours of incubation at a PETase loading of 50 nM, pH 7.2 in phosphate buffer, or a buffer-only control. No surface pitting was observed.

PEF is another semi -aromatic polyester marketed as a bio-based PET replacement. Given the structural similarity of PET and PEF, and recent studies on PEF degradation by cutinases, we hypothesized that PETase may also depolymerize this substrate. Accordingly, we synthesized PEF coupons, and Panels A-D of FIG. 3 shows the results of PEF incubations with the wild-type PETase enzyme and the PETase double mutant, alongside a buffer-only control. Visually, the surface morphology of PETase-treated PEF is even more modified than PET with SEM revealing the formation of large pits, suggesting that PETase is potentially much more active on this substrate than PET. The observation of enhanced PEF degradation by microscopy is corroborated by the DSC data for PEF, which show _' a reduction in relative crystallinity of 15.7% (absolute of 2.4%) compared to the relative reduction of 10 1% for PET (Table 2, Panel E of FIG 9).

To predict how' a PEF oligomer interacts with the wild-type and double mutant PETase active sites, IFD was again performed. The expected PETase activity was again captured from a structural standpoint, with the PEF ester oriented within nucleophilic attack distance of Serl60 (Panels E and F of FTG 13, Panel B of FIG 1 1). As with PET IFD results, we were able to identify interactions to support increased activity of the PETase double mutant enzyme. In the PEF-wild type binding mode (docking score -9.07 kcal/mol), two aromatic interactions are formed to Trpl85 and Trpl59 (Panel B of FIG. 11). However, in the PEF-double mutant binding mode (docking score -10.07 kcal/mol) three aromatic interactions were observed: parallel displaced to Trpl85 (5.7 A), point to face to His237 (5.1 A), and parallel displaced to Phe238 (5.2 A). Additionally, Tyr87 is within range for a potential aromatic interaction at 6.2 A. One interesting interaction w'as observed in the PEF-double mutant binding mode: His237 flipped "up", out of the catalytic triad, to play an aromatic stabilization role (replacing the wild- type Trpl59 stabilization), and instead 1 lis t 59 supported Serl60 via hydrogen bonding, 3 2 A. This interaction, Serl60-Hisl59, is also observed in apo MI) simulations of the double mutant structure. It could thus be postulated that Hisl59 senses an additional means for shuttling protons in the PETase double mutant, which will be examined in a future study. As seen with PET, docking scores predict increased binding affinity of PEF to the double mutant PETase (Table 3), and structurally we can relate this to aromatic interactions supported by F238 and a potential alternative pathway for proton shuttling during catalysis.

FIG. 13 illustrates a comparison of PETase and the engineered enzyme S238F/W159H with PEF Panel A of FIG. 13 shows a buffer-only control of PEF coupon. Panel B of FIG. 13 shows PEF coupon after incubation with wild-type PETase. Panel C of FIG. 13 illustrates PEF coupon after incubation with the PETase double mutant, S238F/W 159H. All SEM images were taken after 96 h of incubation at a PETase loading of 50 nM, pH 7.2 in phosphate buffer, or a buffer-only control. Panel D of FIG. 13 shows the percent crystallinity change, coupon mass loss, and reaction product concentration after incubation with buffer, wild-type PETase, and the S238FAV159H double mutant. Panel E of FIG. 13 illustrates the predicted binding conformations of wild-type PETase from docking simulations demonstrate that PEF is accommodated in an optimum position for the interaction of the carbon (black) with the nucleophilic hydroxyl group of Serl60, at a distance of 5.0 A (dashed line). His237 is positioned within 3.7 A of the Seri 60 hydroxyl (dashed line). Residues Trp 159 (lightly shaded) and Ser238 (more darkly shaded) line the active site channel. In contrast, the double mutant S238F/W159H significantly alters the architecture of the catalytic site for PEF binding.

Residue His237 rotates away from Seri 60 and instead forms an aromatic interaction with PEF chain, 5.1 A. Surprisingly, the mutated Hisl59 becomes an alternative productive H-bond partner, 3.2 A. Similar to interactions with PET, Phe238 also provides additional hydrophobic interactions to an adjacent furan ring of the extended PEF polymer, creating a more intimate binding mode with the cleft with a parallel displaced aromatic interaction at 5.2 A.

The high-resolution structure described here reveals the binding site architecture of the 1 sakaiensis 201-F6 PETase, while the IFD results provide a mechanistic basis for both the wild-type and PETase double mutant towards the crystalline semi-aromatic polyesters PET and PEF. Changes around the active site result in a widening of the cleft compared to structural representatives of three thermophilic cutinases (FIG 13), without other major changes in the underlying secondary or tertiary structure. Furthermore, w ^? e demonstrated that PETase is active on PET of approximately 15% crystallinity, and 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. Enzyme scaffolds capable of PET breakdown above the glass transition temperature (>70°C for PET) 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 merit investigation towards enzymatic solutions for PET and PEF recycling. Furthermore, in light of recent studies that demonstrate the impressive synergistic effect combining multiple PET-active lipases, we expect that incorporation of I. sakaiemis MHETase will further increase the performance, 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 homologues, 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 Trpl85, Tyr87, and Trpl59, and Phe238 in the PETase double mutant. Binding of PET and PEF induce conformational changes in these residues relative to the crystal structure, thus modeling protein flexibility in response to PET/PEF is critical to predict cataiytically relevant binding modes. Additionally, results of these flexible docking studies agree with experimentally observed trends in the wild type relative to the double mutant performance and provide structural insight to explain this enhancement.

PETase activity on both PET and PEF, but not on aliphatic polyesters such as PBS and PL A, provides the basis for characterizing this enzyme more broadly as an aromatic polyesterase, rather than solely a PETase. It is likely that the enhanced gas barrier properties of PEF will lead to its adoption for beer bottles, and 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 retained similar performance profiles after freeze drying - a positive feature for its potential use in applications that require enzyme production and use be distinct, as it would be for most bio-based recycling options. The problem of plastics depolymerization by enzymes closely mirrors that of enzymes that depolymerize polysaccharides, such as cellulose and chitin. Indeed, strategies that have been used to both understand and improve glycoside hydrolases, including the development of quantitative assays for measuring enzyme (or cocktail) performance on solid substrates, likely can serve as inspiration for more quantitative metrics for comparing plastics-degrading enzymes and enzyme cocktails, which will be reported in future studies. Moreover, the method of PETase action is of keen interest for further protein and enzyme cocktail 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. Beyond the active site, the enzyme may interact with and cleave the substrate in an endo- fashion cleaving PET (or PEE) chains internal to a polymer or in an exo-fashion 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. Lastly, at low substrate loadings, many polysaccharide-active enzymes rely on multi- modular architectures, with a carbohydrate-binding module attached to the catalytic domain. For polyesterase enzymes, hydrophobias, carbohydrate-binding modules, and PHA-binding modules have been used to increase the catalytic efficiency of cutinases for PET degradation. 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 put into widespread use in the 1970s, it is likely that the enzyme system in 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, both to increase the efficiency and substrate range of this polyesterase, but also to provide clues how to further engineer thermophilic cutinases to better incorporate aromatic polyesters, towards to the persistent challenge of highly crystalline polymer degradation. Examples:

Example 1. A modified polyethylene terephthaiate (PET)-digesting enzyme (PETase) comprising at least one amino acid mutation of an active site residue, wherein the modified PETase has a narrowed binding cleft compared to the unmodified PETase.

Example 2. The modified PETase of Example 1, wherein the unmodified PETase is from a bacterium of the genus Ideonella.

Example 3. The modified PETase Example 2, wherei n the bacterium is a strai n of Ideonella sakaiemis.

Example 4. The modified PETase Example 1, wherein an amino residue at position 159 is mutated.

Example 5. The modified PETase Example 4, wherein an amino residue at position 238 is mutated.

Example 6. The modified PETase Example 1, wherein an amino residue at position 238 is mutated.

Example 7. The modified PETase Example 1, wherein the modified PETases comprises the W159H/S238F double mutation.

Example 8. A nucleic acid molecule encoding a modified PETase having mutations at two active-site residues.

Example 9. The nucleic acid molecule of Example 8, wherein the amino acid residue at position 159 is mutated.

Example 10. The nucleic acid molecule of Example 8, wherein the amino acid residue at position 238 is mutated.

Example 1 1. The nucleic acid molecule ofExample 8, wherein the modified PETase comprises the W159H/S238F double mutation.

Example 12. An expression vector comprising the nucleic acid molecule of Example 8.

Example 13. A nucleic acid encoding the enzyme comprising the amino acid sequence depicted in FIG. 2(B).

Example 14. A cell that expresses the modified PETase of Example 1.

Example 15. A method for degrading a polymer comprising contacting the modified PETase of Example 1 with the polymer.

Example 16. The method ofExample 14, wherein the polymer is a polyester. Example 17. The method of Example 14, wherein the polymer is an aromatic polymer or a semi-aromatic polymer.

Example 18 The method of Example 14, wherein the polymer is polyethylene terephthalate Example 19. The method ofExample 14, wherein the polymer is polyethylenefuranoate (PEF). Example 20. The method of Example 14, wherein the polymer is from a recycled plastic material .

Example 21. The method of Example 14, wherein the PETase is expressed by a cell.

The foregoing discussion and examples have been presented for purposes of illustration and description. The foregoing is not intended to limit the aspects, embodiments, or configurations to the form or forms disclosed herein. In the foregoing Detailed Description for example, various features of the aspects, embodiments, or configurations are grouped together in one or more embodiments, configurations, or aspects for the purpose of streamlining the disclosure. The features of the aspects, embodiments, or configurations may be combined in alternate aspects, embodiments, or configurations other than those discussed above. This method of disclosure is not to be interpreted as reflecting an intention that the aspects, embodiments, or configurations require more features than are expressly recited in each claim. Rather, as the following claims reflect, inventive aspects lie in less than all features of a single foregoing disclosed embodiment, configuration, or aspect. While certain aspects of conventional technology have been discussed to facilitate disclosure of some embodiments of the present invention, the Applicants in no disclaim these technical aspects, and it is contemplated that the claimed invention may encompass one or more of the conventional technical aspects discussed herein. Thus, the following claims are hereby incorporated into this Detailed Description, with each claim standing on its own as a separate aspect, embodiment, or configuration.