Discovery of a CaPETase with high PET hydrolytic activity and thermostability
To discover a PET hydrolase, we performed a sequence homology analysis using the National Center for Biotechnology Information (NCBI) database and selected 10 PETase candidates (see Methods for details). A phylogenetic tree was constructed for the 10 selected PETase candidates and 17 reported PET hydrolases. The 27 enzymes were divided into two groups: one group contained mesophilic enzymes, such as IsPETase, and the other group contained thermophilic enzymes, such as TfCut2 and LCC (Fig.1a, Supplementary Fig.1 and Supplementary Table1). The phylogenetic tree was further separated into 10 subgroups, and 3 selected PETase candidates, namely, RZL00883.1, KOX11336.1, and SHM40309.1, formed discrete lineages with low phylogenetic relationships to the reported PET hydrolases (Fig.1a). To characterize the 10 selected PETase candidates (PCs), we first attempted to produce them in a signal peptide-truncated form. Eight of the 10 PETase candidates were successfully produced, except for KOX11336.1 and MAM88718.1. We also measured the melting temperature (Tm) of the eight PETase candidates to determine the thermostability of these enzymes. The candidates exhibited a range of Tm values from 38.6C to 70.5C (Fig.1b and Supplementary Fig.2). We then checked PET hydrolytic activity of the eight PETase candidates at a wide range of temperatures from 30 to 60C using several PET samples, such as post-consumer transparent PET powder (PC-PETTransparent), semi-crystalline PET powder (Cry-PET, Goodfellow Cambridge Ltd) (Cat. No. ES306000), and amorphous PET film (AF-PET, Goodfellow Cambridge Ltd) (Cat. No. ES301445). Among these candidates, PC3, PC7, PC8 and PC10 showed relatively low or undetectable levels of PET hydrolytic activity across the tested conditions (Fig.1c and Supplementary Fig.3). PC6, which have the highest Tm value of 70.5C, produced only negligible amounts of PET hydrolysis products at 30C, but showed the optimal PET hydrolytic activity at 50C (Fig.1b, c and Supplementary Figs.2, 3). PC4 and PC5 showed relatively high PET hydrolytic activity at 50C and 40C, respectively (Fig.1c and Supplementary Fig.3). Surprisingly, compared to other PETase candidates, PC2 exhibited significantly high PET hydrolytic activity across a broad range of reaction conditions. Notably, PC2 exhibited superior activity at 30C and produced the highest amount of PET hydrolysis products across all three PET substrates conditions compared to other candidates. (Fig.1c and Supplementary Fig.3). In addition, the pH profile results for the eight PETase candidates also showed that PC2 showed the highest level of PET hydrolytic activity (Supplementary Fig.4). It is noteworthy that PC2 showed remarkable PET hydrolytic activity compared with the other enzymes, and also exhibited high thermostability with a Tm value of 66.8C (Fig.1b, c and Supplementary Fig.2). Moreover, PC2 had the highest soluble expression level compared with the other enzymes (Fig.1b). These results indicate that PC2 has excellent properties for efficient PET degradation, including enzyme activity, thermostability, and protein expression levels. Thus, we selected PC2 (accession code: SHM40309.1, PETase from Cryptosporangium aurantiacum, CaPETase) as the most robust PET hydrolase among the eight PETase candidates tested. The measurements of changes of the Tm value and activity by addition of metal ions showed that PC2 is not a metal ion-dependent enzyme (Supplementary Fig.5).
a Maximum likelihood phylogenetic tree and percentage identity matrix of the 10 selected PETase candidate (PC1PC10) and 17 reported PET hydrolase sequences. Bootstrap values for 1000 replications are shown at the branching edges. The colored bar represents the level of the percent identity of the enzymes, and detailed percent identity values are listed in Supplementary Fig.31. b Protein yield and the Tm values of the 8 PCs. c PET hydrolytic activity of the eight PCs. The reaction was performed with post-consumer transparent PET powder (PC-PETTransparent, 15mgmL1 with 500nM enzyme), semi-crystalline PET powder (Cry-PET, 15mgmL1 with 2M enzyme), and amorphous PET film (AF-PET, 15mgmL1 with 2M enzyme) in 50mM Glycine-NaOH pH 9.0 buffer at various temperatures (30C, 40C, 50C, 60C) for 3 days. Reactions were performed in triplicate; Data are presented as mean valuesSD. d Comparison of the PET hydrolytic activity of CaPETase, IsPETase, LCC, and TfCut2. The reaction was conducted with Cry-PET (15mgmL1 with 2M enzyme) in 50mM Glycine-NaOH (pH 9.0) at various temperatures (30C, 40C, 50C, 60C) for 12h. Reactions were performed in triplicate; Data are presented as mean valuesSD.
Next, we compared the PET hydrolytic activity of CaPETase with that of well-known PET hydrolases, such as IsPETase, TfCut2, and LCC, over a broad temperature range from 30 to 60C using Cry-PET as a substrate. In reactions at 30C, CaPETase showed significantly higher PET hydrolytic activity than LCC and TfCut2 (Fig.1d and Supplementary Fig.3). Moreover, CaPETase exhibited 1.4-fold higher activity than IsPETase, which is known to have the highest PET hydrolytic activity at ambient temperature among the reported PET hydrolases (Fig.1d)17. In particular, the PET hydrolytic activity of CaPETase was 3.1-fold higher than that of IsPETase at 40C (Fig.1d), likely because CaPETase has much higher thermostability than IsPETase. However, the PET hydrolytic activity of CaPETase at 60C dramatically decreased and reversed compared with that of LCC at temperatures of 50C and 60C (Fig.1d). These results indicate that CaPETase is a promising PET hydrolase that exhibits high PET decomposition ability and thermostability. Considering that it is important to make improvements without the loss of enzyme activity and thermostability in the development of superior PET-degrading enzymes37, we propose that CaPETase represents a more efficient template enzyme for enzyme engineering than other enzymes with extreme mesophilic and thermophilic properties, such as IsPETase and LCC, respectively.
To provide a structural basis for high PET hydrolytic activity of CaPETase, we determined its crystal structure at a resolution of 1.36 (Supplementary Table2). CaPETase shows an / hydrolase fold and a nine-stranded -sheet at the center surrounded by six -helices and two 310-helices (Fig.2a and Supplementary Fig.6). Sequence-independent pairwise superposition of CaPETase with three distinctive PET hydrolases, namely, IsPETase, LCC, and TfCut2, generated global root mean square deviation values of 0.69, 0.65, and 0.53, respectively. Formation of one conserved disulfide bond (DS, C279/C297) and lack of an extended loop in the substrate binding site of CaPETase suggest that the enzyme originated from an ancestor of TfCut2 and LCC rather than IsPETase (Supplementary Fig.7). Interestingly, CaPETase exhibits a somewhat different backbone structure at the active site compared with other PET hydrolases (Supplementary Fig.8). Because the structural comparison using a Cartesian coordinate system is known to be subjective for distinguishing the detailed structural differences of the main chains38, we further analyzed the - torsion angles of the main chains of these four PETases (Supplementary Fig.9) and found that there were local differences in backbone torsion angles between CaPETase and other PET hydrolases (Fig.2a and Supplementary Fig.10). Interestingly, CaPETase also showed significant differences in the backbone torsion angles at the five connecting loops (31, 42, 67, 74, and 85) that form an active site, whereas comparisons of the corresponding loops between the other three PET hydrolases exhibited less differences (Fig.2a and Supplementary Fig.11), suggesting that CaPETase has a unique active site conformation. There were some differences in the network of residues extending from the active site to the nearby spatial environment compared with that of the other PET hydrolases. Among them, we observed unique differences affecting the backbone torsion angles of these loops. Near the 31 loop, distinct residues positioned in the 42 loop and a W105L108G124 network force, which form a unique side-chain internal network, appear to influence the conformation of the 31 loop (Supplementary Fig.12). In fact, the 31 loop has high root mean square fluctuation values near the active sites of other PET hydrolases in molecular dynamic simulations39,40. A unique A192G212F248 network is formed under the 85 loop, where catalytic H246 is located (Supplementary Fig.13). At the corresponding F248 position in CaPETase, LCC and TfCut2 have an alanine residue, whereas IsPETase has a cysteine residue that forms a second disulfide bond. Therefore, the positioning of a bulky F248 might cause significant torsional differences in the 85 loop and 74 loop of CaPETase (Supplementary Fig.13). Finally, an R176W200F209 network appears to trigger conformational differences in the 3-helix and 67 loop (Supplementary Fig.14). Importantly, the 3-helix contains the catalytic S169, and the 67 loop was previously annotated as a wobbling tryptophan-containing loop in PETase from Rhizobacter gummiphilus (Supplementary Fig.14)41. We further analyzed backbone fluctuations of these four PET hydrolases using molecular dynamic simulations and CaPETase exhibits quite unique backbone fluctuation profile (Fig.2b and Supplementary Fig.15). CaPETase has more stable 67 and 74 loops than mesophilic IsPETase, and particularly, the enzyme shows high stability at the front region of the 85 catalytic loop where H246 is located (Fig.2b). However, the end region of 85 which corresponds to the extended loop of IsPETase, and the front region of 31 loop showed the highest and lowest flexibility among the four homologs, respectively (Fig.2b). To our interest, the differences of the backbone fluctuation profile was localized exactly to the unique internal network affecting the backbone torsion angles of these loops. Thus, we believe that the unique backbone conformation of CaPETase allows the enzyme to maintain high activity while stabilizing several flexible loops of the mesophilic PET hydrolase.
a Comparison of the backbone torsion angle differences between CaPETase and IsPETase, LCC, and TfCut2. The structure of five connecting loops forming the active site of CaPETase is displayed as a putty tube representation of the same diameter in PyMoL. The structure is colored according to the Euclidean distance values between the two Ramachandran points of the aligned residues. Colors of white to red designate low to high Euclidean distance values, respectively. The catalytic triad of CaPETase is shown as a stick model with a cyan-color circle. b MD simulations show unique backbone fluctuation profile of CaPETase. C atom root-mean-square fluctuations (RMSF, ) of the CaPETase, IsPETase, TfCut2, LCC during MD simulations. c Comparison of the residues forming the substrate binding cleft of CaPETase, LCC, and TfCut2. The highlighted residues are shown as a stick model. d Distinct residues in the substrate binding site of CaPETase. Distinct and conserved residues are presented in magenta and light blue, respectively. e PET hydrolytic activity of the variants. PC-PETTransparent (15mgmL1) were incubated with 500nM enzymes at 40C for 24h in 50mM Glycine-NaOH buffer pH 9.0. Total amount of released products and the Tm value of the variants are shown as bars and red-colored dots, respectively. Reactions were performed in triplicate; Data are presented as mean valuesSD.
In addition to the unique backbone conformation at the active site, residues forming the substrate binding cleft of CaPETase showed significant differences compared with other thermophilic PET hydrolases (Fig.2c, d). In the vicinity of the wobbling W194, CaPETase possesses unique G196 and L133 residues, where highly conserved residues are located in other PET hydrolases (Fig.2c, d and Supplementary Fig.7). Mutating these residues to the corresponding residues in other PET hydrolases, such as G196L, L133Y, and L133Q, had a negative effect on enzyme activity and/or thermostability (Fig.2e). However, the G196T mutation exhibited enhanced thermostability (Fig.2e), which may result from the formation of a hydrogen bond between G196T and N195. CaPETase also contains a unique I102 residue in the 31 loop showing the largest torsion differences, whereas other PET hydrolases contain a highly conserved threonine residue at the corresponding position, which probably enables CaPETase to form a relatively wider substrate binding cleft (Fig.2c, d and Supplementary Fig.16). Replacement of I102 with threonine resulted in decreased enzyme activity, confirming that the residue contributes to high enzyme activity (Fig.2e). Furthermore, CaPETase has unique residues, such as Q107, W168, and T250, at the regions of the 31 loop, 85 loop, and 3, whereas most of the corresponding residues are highly conserved in other thermophilic PET hydrolases (Fig.2c, d and Supplementary Fig.7). Mutating these residues to the conserved residues in other thermophilic PET hydrolases decreased enzymatic activity and/or stability, indicating that the combined positioning of these residues is necessary to create an optimal substrate binding site for CaPETase with a unique shape and polarity (Fig.2e). One exception was the Q107S mutation, which resulted in no noticeable differences in enzyme activity or thermostability (Fig.2e). Taken together, we suggest that along with unique backbone torsion angles, the positioning of distinct residues at the substrate binding site enable CaPETase to form an optimal substrate binding site for high PET hydrolytic activity.
Although CaPETase has high PET hydrolytic activity and thermostability, its performance is still insufficient for industrial applications. We conducted rational protein engineering of CaPETase to further enhance the PET hydrolytic activity and thermostability of the enzyme using various strategies, such as introducing disulfide bonds and hydrogen bonds and modifying the protein surface charge (Supplementary Fig.17). The thermostability of the variants was monitored by measuring Tm values, and the PET hydrolytic activity of the variants was measured using post-consumer transparent PET powder (PC-PETTransparent) at ambient temperature (40C). We introduced four disulfide bonds, namely, G76C/A143C (DS1), L180C/A202C (DS2), T204C/R233C (DS3), and R242C/S291C (DS4). The DS2 and DS4 mutations increased the Tm value by approximately 3C, whereas the DS1 and DS3 mutations decreased the Tm value compared with CaPETaseWT (Fig.3a and Supplementary Fig.18). Moreover, the introduction of the DS2 and DS4 mutations increased PET hydrolytic activity by more than 20% compared with CaPETaseWT (Fig.3a and Supplementary Fig.18). These results indicate that the DS2 and DS4 mutations were successfully formed in CaPETaseWT and exerted positive effects on enzyme activity and thermostability. We also attempted to improve the thermostability of CaPETase by introducing noncovalent bonds, such as hydrogen bonds and salt bridges, and designed seven mutations, namely, V129T (NC1), P136S (NC2), A192T (NC3), R198K (NC4), V203T (NC5), A252N (NC6), and A257S(NC7). Of these, the NC1 and NC4 mutations increased Tm values by approximately 2C and enhanced PET hydrolytic activity by 30% compared with CaPETaseWT (Fig.3a and Supplementary Fig.18). Finally, in an attempt to improve the protein adsorption ability to the PET surface by modifying the protein surface charge, we designed five mutations to render the protein surface hydrophobic, i.e., N109A (HP1), R151A (HP2), R157A (HP3), R160A (HP4), and R233A (HP5), and four mutations to render the protein surface positive, i.e., T86R (SC1), A155R (SC2), T275R (SC3), and M294R (SC4). Unfortunately, most mutations did not show significant changes or even negative effects on thermostability or enzyme activity; however, the HP1 mutation increased the Tm value by 3.2C, and the SC2 mutation enhanced PET hydrolytic activity by 20% compared with CaPETaseWT (Fig.3a and Supplementary Fig.18). Taken together, we introduced eight point-mutations that resulted in improved thermostability and PET hydrolytic activity, i.e., DS2, DS4, NC1, NC4, HP1, and SC2, among the 20 rationally designed mutations tested (Fig.3a and Supplementary Fig.18). There were also ambiguous mutations that only improved enzyme activity or thermostability, such as NC2, HP2, and HP5. We excluded these mutations from the final selection to develop a much superior variant without compromising enzymatic activity or thermostability (Fig.3a and Supplementary Fig.18)37.
a Single-point mutations of CaPETase. Released PET hydrolysis products and the Tm values of the single-point mutations are presented. The reactions were performed using 500nM enzymes with post-consumer transparent PET powder (PC-PETTransparent, 15mgmL1) as the substrate in 50mM Glycine-NaOH buffer (pH 9.0) for 24h at 40C. Reaction was carried out in triplicate; error bars represent the s.d. of the replicate measurement. b Combinatorial mutations of CaPETase. PET hydrolytic activity of the variants generated using the combinatorial strategy. Released PET hydrolysis products per hour and the Tm values of the combinatorial variants are presented. The reactions were performed using 500nM enzymes with PC-PET (15mgmL1) as the substrate in 100mM Glycine-NaOH buffer (pH 9.0) at 40C for 24h and 60C for 6h, respectively. Reaction was carried out in triplicate; error bars represent the s.d. of the replicate measurement. c Comparison of PET hydrolysis activity between CaPETaseM9 and LCCICCG at various temperatures. The reactions were carried out at different temperatures using PC-PETTransparent (12.5mgmL1) with 1M enzyme and Cry-PET (12.5mgmL1) with 4M enzyme under the 200mM Glycine-NaOH buffer pH 9.0. Reactions were performed in triplicate; Data are presented as mean valuesSD.
We sequentially integrated the six mutations described above to develop a superior CaPETase variant with higher thermostability and PET hydrolytic activity. First, we combined the DS2 and DS4 mutations, and the resulting CaPETaseDS2/DS4 variant showed a synergistic effect on thermostability with a Tm value of 74.3C (Tm=7.4C) (Fig.3b and Supplementary Fig.19). Moreover, the variant enhanced PET hydrolytic activity by 1.35- and 4.45-fold at 40C and 60C, respectively, compared with CaPETaseWT (Fig.3b and Supplementary Fig.19). Next, we set up CaPETaseDS2/DS4 as a scaffold for the next combination. We integrated the NC1/NC4, HP1, and SC2 mutations individually into CaPETaseDS2/DS4 using our engineering strategy. The addition of the NC1/NC4 mutation increased the Tm value significantly by 3.9C and increased PET hydrolytic activity at both 40C and 60C (Fig.3b and Supplementary Fig.19). It showed 3.8- and 17-fold enhanced activity at 60C compared with CaPETaseDS2/DS4 and CaPETaseWT, respectively (Fig.3b and Supplementary Fig.19). The addition of the HP1 mutation increased the Tm value by 2.7C and enhanced PET hydrolytic activity by 1.7-fold at 60C compared with CaPETaseDS2/DS4 (Fig.3b and Supplementary Fig.19). When the SC2 mutation was integrated into the CaPETaseDS2/DS4 variant, we observed no noticeable improvements in activity and thermostability; however, there was a slight increase in PET hydrolytic activity at 60C (Fig.3b and Supplementary Fig.19).
These results suggest that all four mutations (NC1, NC4, HP1, and SC2) had a positive effect on the thermostability and activity of CaPETaseDS2/DS4; thus, we combined the four mutations into CaPETaseDS2/DS4 to generate CaPETaseDS2/DS4/NC1/NC4/HP1/SC2 (CaPETaseM8). Surprisingly, when all four mutations were added to CaPETaseDS2/DS4, a synergistic effect on thermostability and enzyme activity was observed, and CaPETaseM8 exhibited significantly enhanced thermostability with a Tm value of 80.7C and 1.5- and 25.8-fold enhanced PET hydrolytic activity at 40C and 60C, respectively, compared with CaPETaseWT (Fig.3b and Supplementary Fig.19).
As mentioned above, the G196T mutation resulted in positive effects on both thermostability and enzyme activity (Fig.2e); thus, we finally generated CaPETaseDS2/DS4/NC1/NC4/HP1/SC2/G196T (CaPETaseM9) by integrating the G196T mutation into CaPETaseM8. CaPETaseM9 exhibited a Tm value of 83.2C, which corresponds to a 16.7C increase in Tm compared with CaPETaseWT. Moreover, the PET hydrolytic activity of CaPETaseM9 increased by 1.7- and 31.2-fold at 40C and 60C, respectively, compared with CaPETaseWT (Fig.3b and Supplementary Fig.19). These results indicate a positive effect of G196T on CaPETaseWT was applied similarly to CaPETaseM8.
CaPETaseM9 showed much higher activity at all temperature conditions from 30 to 70C, and particularly, showed 41.7-fold higher specific activity at 60C than CaPETaseWT (Supplementary Fig.20). The result was also reproduced in a scale-up system of 50-mL shaking flasks (Supplementary Fig.21). These results demonstrated the improved enzyme activity and reinforced thermostability of CaPETaseM9. The improved thermostability of the variant was further verified through heat inactivation experiments, where CaPETaseM9 maintained its activity even after incubation at 60C for 12h, whereas CaPETaseWT showed complete loss of activity within an hour (Supplementary Fig.22).
We then compared the PET hydrolytic activity of CaPETaseM9 with LCCICCG towards PC-PET and Cry-PET at temperatures ranging from 30 to 60C. CaPETaseM9 showed significantly higher PET hydrolytic activity compared to LCCICCG, at 30C and 40C (Fig.3c). At 50C and 60C, CaPETaseM9 showed quite similar activity compared with LCCICCG (Fig.3c).
To provide structural insights into the enhanced PET-degrading capacity of CaPETaseM9, we determined its crystal structure at a resolution of 1.53 (Fig.4 and Supplementary Table2). The formation of the introduced DS2 and DS4 disulfide bonds was clearly observed in CaPETaseM9, and the SS interatomic length of both disulfide bonds was within the optimal disulfide bond length range (Fig.4 and Supplementary Fig.23). Interestingly, DS4 was located in the vicinity of one of the calcium binding sites of Cut190 and the mutation point of IsPETase R280A42,43, and the formation of DS4 also caused significant changes in the surface electrostatic potential and neighboring region conformation (Fig.4 and Supplementary Figs.23 and 24). The side chain of the mutated V129T was flipped to form a hydrogen bond with the adjacent T131 and D132 residues, thereby further stabilizing the 43 connecting loop (Fig.4 and Supplementary Figs.23 and 25). With respect to R198K, the mutated lysine residue moved inward to form hydrogen bonds with the main chains of N195 and D222, thereby stabilizing the wobbly tryptophan-containing loop (Fig.4 and Supplementary Fig.23). The mutated G196T formed a water-mediated hydrogen bond with the adjacent N195 residue, resulting in further stabilization of the wobbly tryptophan-containing loop (Fig.4 and Supplementary Fig.23). The A155R mutation changed the hydrophobic surface to a positive charge, which seems to increase the attachment of the enzyme to the PET surface, as suggested by a previous report (Fig.4 and Supplementary Fig.23)44. Finally, the N109A mutation appeared to strengthen internal hydrophobic interactions (Fig.4 and Supplementary Figs.23 and 26).
The crystal structure of CaPETaseM9 is shown as a cartoon diagram, and the mutated residues are shown as a stick or a surface electrostatic potential model.
To evaluate the industrial applicability of CaPETaseM9, we conducted a PET decomposition experiment in a pH-stat bioreactor using PC-PETTransparent as a substrate (Supplementary Fig.27). The bioreactor was operated at 55C using 2.70 mgenzymegPET1, and the pH was continuously titrated at 8.0 by adding NaOH. The decomposition rate was measured by monitoring released amounts of MHET and TPA. After a short lag phase of an hour, which was required for initial hydrophilization, the PET degradation rate increased exponentially, and 50% of PC-PETTransparent was depolymerized within 4h (Fig.5a). In the second half of the reaction, the degradation rate slightly decreased because of a decrease in the amount of substrate; however, a final degradation rate of 94.1% was achieved after 12h (Fig.5a). This was a significant result in terms of showing that a high depolymerization rate of 90% or more could be achieved even at 55C, which is a temperature condition relatively lower than the Tg temperature. This result also indicated that CaPETaseM9 has significant PET hydrolytic activity and thermostability comparable to other benchmark biocatalysts. We also performed decomposition of a post-consumer colored PET powder (PC-PETColored), which is known to be relatively difficult to recycle because of the presence of colors, additives, multilayer structure, labels and other complexities45. Interestingly, the depolymerization rate of PC-PETColored was almost identical to that of PC-PETTransparent, showing 50% depolymerization within 4h (Fig.5b); however, the final depolymerization rate of PC-PETColored was 89.2% at 12h, which was slightly lower than that of PC-PETTransparent (Fig.5b). This is probably due to the impurities present in PC-PETColored. These results demonstrate that unlike other recycling methods, biorecycling of PET plastic can be achieved regardless of the color of PET plastic.
Decomposition of post-consumer transparent PET powder (PC-PETTransparent) (a) and post-consumer colored PET powder (PC-PETColored) (b) in a pH-stat bioreactor using CaPETaseM9. Reactions were performed in triplicate independently; Data are presented as mean valuesSD. c Complete degradation of a post-consumer PET container using CaPETaseM9 at 60C. Reactions were performed in triplicate; Data are presented as mean valuesSD.
Finally, we determined whether untreated post-consumer PET containers can be depolymerized by CaPETaseM9. As depolymerization proceeded, the PET film became opaque and thin, and the PET film disappeared completely in 3 days (Fig.5c). These results suggest that CaPETaseM9 can be utilized for decomposing PET plastics with various physical properties.
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