Natural diversity screening, assay development, and characterization of nylon-6 enzymatic depolymerization – Nature Communications

Identification, expression, and amide bond hydrolysis assays of potential nylonases

We first constructed a diverse set of enzymes, initially focusing on biocatalysts with reported PA polymer or linear-oligomer deconstruction activity from peer-reviewed literature and patents (Fig. 1B, Supplementary Data 1–2). As polymer chains become more accessible near the glass transition temperature (T_g) of the substrate (for PA6, T_g ~ 50–55 °C), enzyme thermotolerance was chosen as a desirable characteristic, and hence, was considered when selecting candidates⁶². As demonstrated by Negoro et al., NylC_p2 and its homologs (Ntn hydrolases) have previously been shown to exhibit activity on PA6 cyclic oligomers, linear oligomers and solid PA6 (powder and thin film)^{24,37,49,53,63}, whilst NylB and its homologs have been shown to exhibit PA6 dimer hydrolysis activity^38,39. Amidases have also been used for PA6 modification. One of these enzymes, NfPolyA, an amidase signature (AS) family protein from Nocardia farcinica, had also been reported to exhibit polyurethane (PUR)-hydrolyzing activity, hence, we included some known polyurethanases and urethanases, hypothesizing that these enzymes may also exhibit activity on PA^23,64. Additional Ntn hydrolases and AS family enzyme candidates were selected following NylC_p2 and NfPolyA homolog searches, selecting enzymes from potentially thermotolerant sources. Numerous reports suggest serine hydrolases with the canonical Ser-His-Asp (SHD) catalytic triad including cutinases, and proteases using either serine or cystine catalytic nucleophiles, exhibit promiscuous activity for nylon surface modification^{44,45,46,47,48,50,65}. Thus, these enzymes were also added to the set. A number of the cutinases described also had PET depolymerization activity, hence, several other PET-active SHD hydrolases were added, including some previously engineered thermostabilized variants^21,58, with the hope that they may also exhibit PA depolymerization.

Following selection, enzymes were broadly categorized into six main groupings dependent on both their catalytic activities and previously described substrate preferences (Fig. 1B, Supplementary Data 1). Enzymes found by homology searches were put in the same group as the query enzyme, whilst those where no natural substrate was described in the original publications were assigned functionality using BLAST searches⁶⁶. The groupings used from here on are as follows: (1) NylC-type enzymes, Ntn-hydrolases found to hydrolyze PA6 trimers and longer PA oligomers, (2) NylB-type enzymes, serine hydrolases (Ser-Tyr-Lys catalytic triad) previously demonstrated to specifically hydrolyze PA dimers, (3) amidases, AS family enzymes that act on linear amides or urethane bond containing compounds, (4) Ser-His-Asp hydrolases (SHD-hydrolases), serine hydrolases with a substrate preference for ester-linked polymers, (5) proteases, serine and cystine proteases that cleave peptide bonds in proteins, and (6) miscellaneous (misc.), enzymes that do not fall into one of the other five main groupings. Manganese peroxidases (MnPs), which have previously been implicated in PA hydrolysis^30,35, were excluded from our study as their radical-based reaction leads to diverse product mixtures that may be less desirable for a closed-loop circular recycling process^6,22.

Inspired by the successful homology-based thermostability enhancement of NylC_p2 by Negoro et al. (NylC_p2 D122G/H130Y/D36A/E263Q, T_m increase of 36 °C, demonstrated hydrolytic activity on solid PA6 powder)⁴⁹, we also rationally mutated NylCs from Agromyces (NylC_A) and Kocuria (NylC_K) to increase their melting temperatures (T_ms)^37,49,67. In both enzymes, residues previously described as potentially destabilizing were substituted for their homologous counterparts in NylC_p2 (S111G and A137L), with E263Q additionally being introduced into NylC_A as a potential stabilizing mutation from NylC_K. These mutations increased protein T_m by 16.4 °C and 25.1 °C for NylC_K-TS and NylC_A-TS, respectively (NylC_K-TS, T_m = 87.4 °C, NylC_A-TS, T_m = 87.1 °C, Supplementary Fig. 1). As a three-point mutant of NylB’ (NylB homolog from Arthrobacter sp. K172, NylB’-SCY: R187S/F264C/D370Y)³⁹ was demonstrated to have enhanced PA6 dimer hydrolysis activity, we introduced the corresponding mutations into NylB (from Arthrobacter sp. K172), to generate NylB-SCY, hoping that the mutations would impart additional PA depolymerization activity. The resulting panel of WT and engineered enzymes comprised of 41 candidates.

Five enzymes were procured from commercial sources; of the 36 enzymes not available commercially, 35 were successfully expressed and purified from Escherichia coli (expression summaries detailed in Supplementary Data 2), to give a 40-member candidate pool. To test the ability of each enzyme for amide bond hydrolysis, candidates were incubated with N-(4-nitrophenyl)butanamide (N4NB), a small molecule surrogate that contains a butyramide motif akin to the amide bonds in PA6 (0.5 mM N4NB, 2 µM enzyme, 30 °C in reaction buffer (100 mM sodium phosphate buffer, pH 7.5, 150 mM NaCl), Fig. 2, Supplementary Fig. 2A-I). Successful hydrolytic cleavage of the amide bond in N4NB results in the release of p-nitroaniline (p-NA), which can be monitored spectrophotometrically. The amidases^{64,68,69,70,71} (Supplementary Fig. 2D–F) and NylB-type enzymes^39,72,73 (Supplementary Fig. 2A) exhibited the highest levels of amide hydrolysis activity, with the UMG-SP enzymes²³ (metagenomic amidases), NfPolyA⁶⁴ (polyamidase from N. farcinica), GatA⁶⁹ (amidase of unspecified origin), and MsmegA (metagenomic amidase) completely converting 500 µM N4NB in under five minutes, whilst ReU⁷⁰ (Rhodococcus equi TB-60 urethanase), OoH⁷¹ (ω-octalactam hydrolase), and NylB’-SCY³⁹ reached 100% conversion in under 30 minutes. The NylC-type enzymes^37,67,74,75 (Supplementary Fig. 2B, C) and SHD-hydrolases^{21,42,58,76,77,78,79,80,81,82} (Supplementary Fig. 2H, I) exhibited a wider range of activities. The highest performers in these two groups, NylC_A⁶⁷ and AoC⁷⁶ (Aspergillus oryzae cutinase), displayed 16.2 and 29.3 µM/h p-NA production, respectively, during the linear phase of the reaction, and the poorest performers, M–NylC (Microbacterium sp. NylC) and LCC-ICCG²¹ (engineered cutinase from leaf/compost metagenome), only had detectable activity after 8 h and 12 h, respectively. Interestingly, none of the proteases (α-chymotrypsin, papain, subtilisin, and trypsin⁴⁸, Supplementary Fig. 2G) had any detectable hydrolysis activity under the tested reaction conditions; this may be due to the difference in chemical properties between N4NB and the protease’s natural substrates.

All successfully expressed enzymes were tested for their ability to hydrolyze the amide bond in N4NB, leading to the release of p-nitroaniline (p-NA, monitored spectrophotometrically at 410 nm) and butyric acid (BA). The bar chart shows the conversion extent of N4NB to p-NA after 5 h of reaction using 0.5 mM N4NB and 2 µM enzyme at 30 °C in reaction buffer (100 mM sodium phosphate buffer (NaPi), pH 7.5, 150 mM NaCl). Enzymes are colored by their group from Fig. 1, with SHD-hydrolase and misc. group bars hashed to aid clarity; the proteases exhibited no detectable activity and thus are not shown. A double asterisk (**) indicates reactions with enzymes that went to completion in ≤ 5 min, a single asterisk (*) indicates reactions with enzymes which went to completion in ≤ 30 min. All reactions are ordered with respect to reaction rate, with the fastest enzymes to the left. A no enzyme control reaction led to no detectable release of p-NA. Data shown are baseline corrected. Reactions were carried out in triplicate (n = 3), error bars show the standard deviation of the replicate measurements, the error bar centers are the means of the replicate measurements, and the replicate measurements are represented as gray circles. References for the enzyme source and group are available in Supplementary Data 1, and the enzyme sequences, EC numbers, and expression conditions are in Supplementary Data 2. Source data are provided in the Source Data file.

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Development of a high-throughput solid PA6 depolymerization analysis method by LC-MS/MS

We next sought to thoroughly characterize the ability of the potential nylonase panel to deconstruct solid PA6. Due to the lack of a strong chromophore in the predicted products and the desire for sub part per million (ppm) detection limits, we developed a high-throughput liquid chromatography with tandem mass spectrometry (LC-MS/MS) analysis method (published on protocols.io, [https://doi.org/10.17504/protocols.io.6qpvr3k92vmk/v1])⁸³ to monitor the major predicted soluble released compounds following PA6 depolymerization reactions. Utilizing tandem mass spectrometry enabled rapid analyses. When operating in multiple reaction monitoring (MRM) mode, the instrumentation and detection is optimized for each compound, making the mass transitions highly selective and unique to each analyte, and thus eliminating the need for chromatographic peak separation or product derivatization, significantly reducing analysis time.

Using this ex-situ analysis method, linear-oligomers of 6-aminohexanoic acid (6-AHA) from monomer to pentamer and cyclic-oligomers from ε-caprolactam to trimer were all detected in under three minutes, representing a significant advancement in the speed and diversity of potential products quantified (Fig. 3A). The use of a multisampler allows eight 96-well plates to be analyzed without manual intervention, with around 20 samples being analyzed per hour. Reaction samples were arrayed across 96-well plates, with a calibration verification standard (CVS) evaluated every 20 wells to ensure accurate quantification and system stability, demonstrating the suitability of this method for high-throughput analyses⁸³.All analytes were detectable from 0.01–7.0 µg/mL, with samples above the upper quantitation limit further diluted to enable quantification in the calibration range. Calibration curves were run prior to each set of samples analyzed to account for mass spectrometry signal drift (Fig. 3B). The linear 6-AHA products from monomer to trimer and cyclic products from ε-caprolactam to 6-AHA cyclic-dimer were quantified using commercial standards. The linear 6-AHA tetramer and pentamer were quantified by applying the calibration curve from 6-AHA linear trimer; the 6-AHA cyclic-trimer was quantified utilizing the calibration curve from the 6-AHA cyclic-dimer responses.

A Representative LC-MS/MS chromatogram following enzymatic hydrolysis of PA6 (0.01 µM NylC_K-TS, 13 mg PA6, 60 °C), showing the eight compounds that were detected and quantified. B Representative calibration plot of standards run prior to analysis of samples by LC-MS/MS, showing the five chemical standards (analytes) that were available commercially, measured from 0.01-7.0 µg/mL. The 6-AHA tetramer and pentamer concentrations, and the 6-AHA cyclic-trimer concentrations were determined using the calibration curves from the 6-AHA trimer and 6-AHA cyclic-dimer, respectively, as no commercial standards were available. The dotted line represents the quadratic line of best fit for each analyte. Source data are provided in the Source Data file.

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Assessment of potential nylonases on a solid PA6 substrate

With a robust nylon depolymerization analysis method in hand, we next sought to examine the capabilities of our enzyme panel for PA6 deconstruction. For activity screens, commercially available PA6 film from Goodfellow was used as the substrate (13.2% crystallinity by differential scanning calorimetry (DSC), 0.2 mm thickness, full material characterization detailed in Supplementary Table 1, with DSC, gel permeation chromatography (GPC) and thermogravimetric analysis (TGA) plots shown in Supplementary Fig. 3A–C).

PA6 film was washed with DI water prior to use, as detailed in the Methods section, to remove surface-bound cyclic-oligomer by-products created during the nylon manufacturing process⁸⁴. Reactions comparing the enzyme panel were conducted for 10 days at temperatures from 40 to 70 °C, with time points taken at 24, 72, 168, and 240 h (Supplementary Fig. 4). Control assays without enzyme led to minimal release of linear 6-AHA oligomers, with the 6-AHA monomer below the limit of detection across all tested temperatures when no enzyme was present (Supplementary Fig. 5A). Enzyme reactions were carried out in 100 mM sodium phosphate buffer (NaPi), pH 7.5, 150 mM NaCl, and contained 2 μM enzyme and 13 mg PA6 (two squares of 0.5 × 0.5 cm PA6 film), representing a substrate loading and an enzyme/substrate loading in the reactions of 0.65 wt% and 0.15 mM enzyme/g PA6 film, respectively, unless otherwise stated. Although polymer deconstructions may be surface-area limited processes, the enzyme/substrate loading is referred to throughout as amount of enzyme per mass of PA6 as is good practice for enzymatic depolymerization reactions^54,56,85. Across all tested enzymes, the extent of ε-caprolactam release was consistent; for most enzymes assayed, concentrations of the 6-AHA cyclic-dimer and cyclic-trimer also remained constant during the reaction (Supplementary Fig. 5B). Hence, for clarity, only the linear products are presented graphically, with cyclic products mentioned only when their levels changed. The linear oligomers (from monomer to trimer) appear robust to incubation for extended time periods at 40–80 °C, with an average recovery (as measured by LC-MS/MS) across all time points and temperatures of 96.4% ± 9.2, 99.9% ± 6.3 and 104.3% ± 7.2 of individually incubated 10 µg/mL standards of 6-AHA monomer, 6-AHA dimer and 6-AHA trimer, respectively (Supplementary Table 2). These results indicate that changes in oligomer levels seen in reactions can be attributed mainly to enzyme action. As a note, although the nylonase deconstruction assays presented below were all conducted with purified proteins, the LC-MS/MS analytical method is unaffected by instead using crude cell lysate (Supplementary Fig. 6) as the biocatalytic agent, with a similar product release profile witnessed as compared to using purified protein (Supplementary Fig. 7). To aid comparisons of total PA6 depolymerization extent in each reaction, the detected soluble released products are presented as their 6-AHA equivalents.

Nylonase activity was observed across a range of enzymes from the different groups tested: overall, 31 enzymes demonstrated measurable activity, while nine enzymes exhibited minimal detectable additional soluble product release above background levels (Fig. 4, Supplementary Figs. 8–13). The temperatures for optimal activity ranged from 40 to 70 °C, with, in general, higher levels of soluble product release seen with increasing temperatures. Interestingly, there appears to be different distributions of products released across the different enzyme groups. Namely, NylB-type enzymes, GatA, and UMG-SP-1 mainly produce 6-AHA monomer, whilst most amidases and cutinases release a mixture of linear products, with very little 6-AHA pentamer seen for many of the amidases. The most active NylC-type enzyme reactions, however, are dominated by 6-AHA dimer. It is important to note that only one pH and buffer condition was tested, so it is possible that relationships between different enzyme activities seen here may be altered by more extensive optimization of reaction conditions for each individual enzyme.

A Heat map time-course profiles of the total released (sum) 6-AHA monomer equivalents following PA6 depolymerization reactions with the top performing NylC-type (yellow), NylB-type (blue), SHD-hydrolase (pink), and amidase (purple) enzymes, carried out from 40 to 70 °C over 10 days. The heat map gradient represents the released 6-AHA monomer equivalents from 0 to 360 µM, with each square representing the average of reactions carried out in triplicate (n = 3). (Full enzyme names and sources are in Supplementary Data 1.) B Representative depolymerization reactions with enzymes from each group at their optimal temperatures (reactions conducted between 40 and 70 °C, x-axis), with 6-AHA oligomers of different lengths represented as their 6-AHA monomer equivalents. Reactions were carried out in triplicate (n = 3), error bars show the standard deviation of the replicate measurements, the error bar centers are the means of the replicate measurements, and the replicate measurements are represented as gray circles. For all data in (A) and (B), the reactions contained 2 µM enzyme and 13 mg PA6 (0.15 mM enzyme/g PA6, 0.65 wt% substrate loading) and were incubated in reaction buffer (100 mM NaPi buffer, pH 7.5, 150 mM NaCl). Source data are provided in the Source Data file.

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By far the most active enzymes were from the NylC-type group, with NylC_p2, NylC_A, NylC_K, and their engineered variants identified as the top-performers, being between two and six-fold more active than the best enzymes from other groups. The significant accumulation of linear 6-AHA trimer in the reactions with the NylC variants found via homology searches is probably a result of their lower propensity for trimer hydrolysis. For instance, NylC_K-TS hydrolyzes 100% of 50 µM 6-AHA trimer to dimer and monomer in under 30 minutes at reaction conditions, while trimer hydrolysis is much slower with Tt-NylC (Thermocatellispora tengchongensis NylC), where 67.1% trimer remains after five hours (pH 7.5 NaPi buffer, 150 mM NaCl, 2 µM enzyme, 60 °C) (Supplementary Fig. 14A, B). As the 6-AHA dimer remains intact at reaction conditions in the presence of NylC_K-TS after 24 h (100 µM dimer, pH 7.5 NaPi buffer, 150 mM NaCl, 2 µM NylC_K-TS, Supplementary Fig. 14C), monomer accumulation in reactions containing NylC_p2, NylC_A, NylC_K, and their variants may be due to the fast hydrolysis of released trimers to 6-AHA dimer and monomer. Thermostabilizing mutations appeared beneficial as hypothesized, with all NylC-TS variants showing increased activity at 70 °C (Supplementary Fig. 8, Fig. 4), compared to 50-60 °C optimal temperatures for their wild-type (WT) counterparts. However, interestingly, these mutations also appeared to slow the hydrolysis of the 6-AHA cyclic-trimer, which is a known substrate for NylC-type enzymes (Supplementary Fig. 15)⁷⁴. The best NylC-type enzyme tested was the two-point mutant NylC_K-TS (T_m = 87.4 °C) (Figs. 4 and  5A), which was even active up to 80 °C (Supplementary Fig. 16). In comparative reactions of NylC_K-WT and NylC_K-TS at 40-60 °C the total released products and reaction profiles are highly comparable (1 µM enzyme and 13 mg PA6, 0.08 µM/g PA6, 0.65 wt% substrate loading, pH 7.5 NaPi buffer, 150 mM NaCl, Supplementary Fig. 16). However, from 70-80 °C, NylC_K-TS has a higher overall yield of soluble products after 10 days (Supplementary Fig. 16). Comparing both enzymes at the temperatures which led to the highest product release (50 °C and 80 °C, respectively), NylC_K-TS starts producing more 6-AHA equivalents from 3 h of reaction onwards, releasing 28.8% more 6-AHA equivalents over the course of 10 days (1 µM enzyme and 13 mg PA6, 0.08 mM enzyme/g PA6, 0.65 wt% substrate loading, pH 7.5 NaPi buffer, 150 mM NaCl, Fig. 5B). Despite the promise of the overall best performing enzyme NylC_K-TS, the highest soluble PA6 oligomer release seen (339.7 µM of 6-AHA equivalents) only equates to 0.67 wt% PA6 depolymerization extent.

A Mutational map of NylC_K-TS (NylC_K-S111G/A137L) using the crystal structure of the close homolog NylC_A (PDB: 3AXG [https://doi.org/10.2210/pdb3AXG/pdb]) as a scaffold. NylC_K-TS is an αβ heterodimer, which is represented by the light and dark blue ribbons; the catalytic residue (T267) is shown as a van der Waals surface colored by all atoms with orange carbon atoms; the thermostabilizing mutations are represented as pink spheres. Residues 261-266 are unmodelled in the PDB structure and are represented as a dashed line. B Total released linear 6-AHA oligomers following PA6 film depolymerization reactions with either 1 µM NylC_K or 1 µM NylC_K-TS and 13 mg PA6 (0.08 mM enzyme/g PA6, 0.65 wt% substrate loading), at the temperature where the highest depolymerization extent was seen (reactions conducted from 40 to 80 °C). The gray inset graph is an expansion of the early time points taken during the reaction. Reactions were carried out in triplicate (n = 3), error bars represent the standard deviation of the replicate measurements, and the error bar centers are the mean of the replicate measurements. Source data are provided in the Source Data file.

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Intriguingly, although NylB appears sparingly active for PA6 film deconstruction, NylB’, a NylB homolog with 88% sequency identity, released 72.9 µM 6-AHA equivalents over 10 days at 40 °C (Fig. 4B, Supplementary Fig. 9). The three-point NylB’ variant, NylB’-SCY, is even more active and additionally more thermostable, with optimal 6-AHA equivalent release at 50 °C (130.2 µM 6-AHA equivalents), corresponding to 0.26 wt% PA6 deconstruction after 10 days of reaction. Although the NylB’-WT scaffold appears to be the driving force for increased PA6 hydrolysis activity compared to NylB-WT, the R187S/F264C/D370Y mutations also appear to have an effect as NylB-SCY is three-fold more active than NylB-WT. However, as the total 6-AHA equivalents released over 10 days at 40 °C by NylB-SCY was <8.0 µM, the overall effect on PA6 depolymerization was negligible. NylB’-SCY readily hydrolyzes both 6-AHA dimer and trimer at reaction conditions (pH 7.5 NaPi buffer, 150 mM NaCl, 2 µM NylB’-SCY, 50 °C), with 100 µM dimer or 50 µM trimer completely converted to monomer in under 30 minutes (Supplementary Fig. 17A, B). Based on these experimental results, it is difficult to identify whether the accumulation of 6-AHA monomer is a product of longer oligomer release followed by fast hydrolysis to monomer, or whether NylB’-SCY exclusively releases 6-AHA monomer from polymer chain ends.

For the nylon-active SHD-hydrolases, Fusarium solani cutinase (FsC) and Thermobifida cellulosilytica cutinase 1 (ThcCut1) have been used previously to modify the surface of PA-6,6 fabric^42,47,50. The reaction profiles and product distributions of all of the biocatalysts tested from this group suggest a similar mode of action, with the mixture of soluble oligomers released indicating non-specific cleavage of surface residues, rather than coordinated, progressive depolymerization activity (Supplementary Fig. 10). Support for this mode of action comes from the most active SHD-hydrolase tested, LCC-ICCG (Fig. 4B), which produces only 49.8 µM of 6-AHA equivalents over the course of 10 days at 70 °C, corresponding to 0.09 wt% depolymerization of the PA6 film. The observation that mixtures of oligomers are retained over the 10 days of reaction additionally suggests that there is limited further deconstruction of these soluble linear products by SHD-hydrolases. Indeed, incubation of LCC-ICCG with 50 µM 6-AHA trimer for 24 h at reaction conditions (pH 7.5 NaPi buffer, 150 mM NaCl, 2 µM LCC-ICCG, 70 °C), leads to negligible turnover to dimer or monomer (Supplementary Fig. 17C).

Similarly, low extents of depolymerization are also observed for the amidases, potentially due to their low thermotolerance, with most becoming deactivated above 40 °C (Fig. 4A, Supplementary Fig. 11). The limited activity of these enzymes was surprising, as the amidases were among the most active enzymes from the initial screens on N4NB. However, as the active sites of NfPolyA and its homologs are buried within the enzyme core, their ability to bind and access amide bonds in polymer chains is likely more restricted (Supplementary Fig. 18). GatA and UMG-SP-2 produced the most 6-AHA equivalents (109.6 µM and 126.3 µM, respectively, over 10 days at 40 °C) amongst the amidases (Fig. 4B), corresponding to 0.2–0.25% PA6 film depolymerization. The interesting preference of GatA for 6-AHA monomer release is in part due to its nylon oligomer hydrolysis ability; GatA can hydrolyze linear products, with 67% of 100 µM 6-AHA dimer and 100% of 50 µM 6-AHA trimer hydrolyzed under reaction conditions (pH 7.5 NaPi buffer, 150 mM NaCl, 2 µM GatA, 40 °C) within 24 h (Supplementary Fig. 19A, B), but it also hydrolyzes cyclic oligomers. GatA completely depletes 6-AHA cyclic-trimer during PA6 film reactions (Supplementary Fig. 20), with 6-AHA cyclic-dimer also being a substrate for the enzyme (Supplementary Fig. 21A, B), indicating that a proportion of the monomer release observed is the product of hydrolysis of these surface contaminants. Interestingly, UMG-SP-1 also has a preference for monomer release (Supplementary Fig. 11). UMG-SP-1 is similarly able to hydrolyze small PA6 oligomers, hydrolyzing both 100 µM 6-AHA dimer and 50 µM 6-AHA trimer to produce a majority 6-AHA monomer product at reaction conditions (pH 7.5 NaPi buffer, 150 mM NaCl, 2 µM UMG-SP-1, 40 °C) within 24 h (Supplementary Fig. 18C, D), with 6-AHA cyclic-trimer also appearing to be a substrate for this enzyme (Supplementary Fig. 20). Hence, product preference for the amidases does not appear as consistent as with other enzyme groups tested here, with soluble oligomer distributions following PA6 depolymerizations being highly enzyme dependent. As with the SHD-hydrolases, the low rates of depolymerization for all the amidases again currently suggests a surface modification process rather than significant bulk PA6 depolymerization.

No measurable PA6 depolymerization activity was seen for any of the proteases or miscellaneous category enzymes tested here (Supplementary Figs. 12 and 13). A possibility is that these enzymes may cleave polyamides, but release products of a higher molecular weight than what our analysis method detects. However, a lack of subsequent depolymerization down to smaller molecular weight oligomers or monomers suggests that the usefulness of these six enzymes may be limited.

In-depth characterization of a selection of the best performing enzymes

We were interested to examine a subset of the best performing enzymes, including NylC_K-TS, Tt-NylC, and NylB’-SCY, to test if the rate and extent of PA6 depolymerization could be improved and the mode of action understood. NylB’-SCY reactions were carried out at 50 °C, its highest operating temperature, while NylC-type enzyme reactions were conducted at 60 °C to allow fair comparison between homologs. For all three enzymes, increasing the substrate loading from 0.32-1.6 wt% (one to five squares of PA6 film, keeping enzyme concentration constant at 1 µM enzyme, 100 mM NaPi buffer, pH 7.5, 150 mM NaCl) leads to greater product release, most likely due to an increase in available reactive surface area (Fig. 6A). Product distributions remain constant for both NylC_K-TS and NylB’-SCY at all substrate loadings, while the 6-AHA trimer accumulates in Tt-NylC depolymerization reactions.

A Total released linear 6-AHA oligomers, represented as 6-AHA monomer equivalents, in reactions with 1 µM NylC_K-TS (60 °C), Tt-NylC (60 °C), or NylB’-SCY (50 °C), over the course of 10 days, varying the substrate loading from 1 to 5 PA6 squares (6.5–32.5 mg PA6, substrate loading of 0.32-1.6 wt%, sq=squares). Pie-charts represent the proportion of different 6-AHA oligomers measured at that time point. Reactions were carried out in triplicate (n = 3), error bars show the standard deviation of the replicate measurements, the error bar centers are the means of the replicate measurements, and circle, square, and triangle points represent the mean value of the triplicate measurements. B PA6 depolymerization reactions with varying enzyme loadings of NylC_K-TS (60 °C), Tt-NylC (60 °C), or NylB’-SCY (50 °C). Reactions contained 13 mg PA6 substrate (0.65 wt% substrate loading) and were monitored for 10 days. Reactions were carried out in duplicate (n = 2), and the replicate measurements are represented as gray circles. For all data in panels A and B, the reaction buffer consisted of 100 mM NaPi buffer, pH 7.5, 150 mM NaCl. Source data are provided in the Source Data file.

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Increasing enzyme loading (keeping PA6 film mass constant at 13 mg PA6 film, 0.65 wt% substrate loading, 100 mM NaPi buffer, pH 7.5, 150 mM NaCl), surprisingly, had little effect on total PA6 film depolymerization (Fig. 6B). 6-AHA equivalent release never surpassed 300 µM under any condition, with improvements in total NylC_K-TS depolymerization stalling above 0.1 µM enzyme in the reaction (8 µM enzyme/g PA6). However, interestingly at low NylC_K-TS enzyme loadings, where the reaction rate will be slowed, a higher proportion of 6-AHA trimer to pentamer products was observed. A proportion of NylC_K-TS PA6 hydrolysis events may therefore be random, producing mixed length oligomers that are subsequently hydrolyzed to dimer and monomer. Slowed oligomer hydrolysis in low enzyme loading reactions additionally suggests a preference of NylC_K-TS for release of product from the polymer surface over hydrolysis of oligomers in solution. Reactions with Tt-NylC support this hypothesis as trimer levels were only reduced at very high enzyme loadings. These trends, however, are not observed for NylB’-SCY: even at the lowest enzyme loading, only 6-AHA is produced. As an important consideration for further application, significantly less NylC_K-TS was required to achieve the highest levels of total depolymerization (0.1 µM NylC_K-TS, 8 µM enzyme/g PA6, equivalent to 0.58 mg enzyme/g PA6), compared to both Tt-NylC or NylB’-SCY (10 µM enzyme, 0.77 mM enzyme/g PA6, equivalent to 54.6 mg enzyme/g PA6 and 66.9 mg enzyme g/ PA6, for Tt-NylC and NylB’-SCY, respectively).

Variation of reaction pH from pH 6-10 (1 µM enzyme and 13 mg PA6, 0.08 mM enzyme/g PA6, substrate loading 0.65 wt%, 100 mM buffer, 150 mM NaCl) also did not elicit higher levels of PA depolymerization; however, it did reveal differences in pH tolerance amongst the three enzymes (Supplementary Fig. 22). NylC_K-TS was mostly unaffected by pH changes, with Tt-NylC being similarly pH-robust with drops in activity only seen at the extremities of the pH range tested. Conversely, NylB’-SCY is particularly sensitive to pH changes: pH 7 is optimal with significant decreases in activity either side of this, and complete deactivation from pH 9-10.

Examination of the reaction profile of NylC_K-TS

We next sought to identify what was limiting the enzymatic depolymerization of PA6 and causing the observed asymptotic reaction profiles, using the most active enzyme, NylC_K-TS, as the test case. We identified slight crystallinity increases in the PA6 substrate in no enzyme control reactions in reaction buffer (100 mM NaPi buffer, pH 7.5, 150 mM NaCl) at 70 °C (maximum 3.2% increase in crystallinity by DSC over 10 days), hence, reactions were carried out at 60 °C where this effect was less noticeable (maximum 0.2% increase in crystallinity by DSC over 10 days) (Supplementary Table 1, Supplementary Fig. 23). Reactions contained 1 µM of enzyme and 13 mg of PA6 (0.08 mM enzyme/g PA6, substrate loading of 0.65 wt%), in 100 mM pH 7.5 NaPi buffer and 150 mM NaCl. To rule out effects of incubation-induced PA6 changes during the reaction, we conducted NylC_K-TS depolymerizations using PA6 film incubated in reaction buffer at 60 °C for either three or seven days prior to enzyme addition. As there was no significant difference between the product release of pre-incubated PA6 film versus non-incubated PA6, this effect could be discounted (Supplementary Fig. 24A).

Interestingly, including bovine serum albumin (BSA) in reactions allows for the same level of PA6 depolymerization with 100-fold less enzyme loading (0.01 µM NylC_K-TS, 0.8 µM enzyme/g PA6, substrate loading of 0.65 wt%, 0.5 µM BSA, 100 mM NaPi buffer, 150 mM NaCl, pH 7.5, 60 °C, Fig. 7A, Supplementary Fig. 24B). Increasing the concentration of BSA in PA6 deconstruction assays (from 0.5 to 2 µM), does not promote additional depolymerization, indicating that BSA is not promoting further PA6 hydrolysis, but is instead acting to enhance NylC_K-TS specific activity (Supplementary Fig. 24B). Additional evidence for this is that the promotion of activity only occurs at lower NylC_K-TS concentrations (0.01 µM enzyme), with addition of BSA to reactions containing 1 µM NylC_K-TS leading to no additional benefit (Supplementary Fig. 24C). As BSA is known to prevent non-specific binding in reactions, we believe these findings suggest that the activity of NylC_K-TS may be substantially impacted by non-productive PA6 binding events. Alternatively, BSA may coat the reaction vessel, reducing enzyme adsorption to these surfaces, hence promoting mass transfer of NylC_K-TS to the polymer surface. More extensive exploration of this phenomenon may be necessary to help guide future NylC_K-TS engineering efforts or may suggest new ways to promote faster or additional PA6 depolymerization.

A Total released linear 6-AHA oligomers following PA6 film depolymerization reactions (13 mg PA6, 0.65 wt% substrate loading), using either 0.01 µM or 1 µM NylC_K-TS enzyme alone (0.8 µM enzyme/g PA6 or 0.08 mM enzyme/g PA6, respectively) compared to a reaction containing 0.01 µM NylC_K-TS supplemented with 0.5 µM bovine serum albumin (BSA) over the course of 10 days. B Total released linear 6-AHA oligomers in reactions with 1 µM NylC_K-TS and 13 mg PA6 (0.08 mM enzyme/g PA6, 0.65 wt% substrate loading), where the reaction was either allowed to progress for 13 days with no interruption (Normal reaction), where fresh enzyme was added to the reaction after three days (1 µM NylC_K-TS, + Fresh enzyme), or where fresh substrate was added to the reaction after 3 days (13 mg PA6, + Fresh substrate), and the reactions monitored for a further 10 days. For (A) and (B), circle, square, and triangle points represent the mean value of the replicate measurements, error bars represent the standard deviation of the replicate measurements, and the error bar centers are the means of the replicate measurements. C PA6 depolymerization reactions with 1 µM NylC_K-TS and 13 mg PA6 (0.08 mM enzyme/g PA6, 0.65 wt% substrate loading), with either low crystallinity Goodfellow film (13.2% crystallinity by DSC), or high crystallinity PA6 film (23.0% crystallinity by DSC). Replicate measurements are represented as gray circles. For (A)–(C), reactions were carried out in triplicate (n = 3), error bars represent the standard deviation of the replicate measurements, and the error bar centers are the means of the replicate measurements. D SEM images of PA6 films (13 mg) incubated either without enzyme (no enzyme) or with 1 µM NylC_K-TS (0.08 mM enzyme/g PA6, 0.65 wt% substrate loading). Additional images are shown in Supplementary Fig. 27. Scale bar, 100 μm. For (A)–(D), all reactions were carried out in reaction buffer: 100 mM sodium phosphate buffer (NaPi), pH 7.5, 150 mM NaCl at 60 °C. Source data are provided in the Source Data file.

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Reaction progress was not recovered by supplementation with fresh NylC_K-TS after either three or seven days of enzyme reaction (Fig. 7B, Supplementary Fig. 24D). However, addition of new PA6 substrate (13 mg) to a NylC_K-TS depolymerization reaction that had been running for three days (the start of reaction plateau), led to additional product release of a similar magnitude as from the original substrate, with the same relationship seen after substrate addition to a seven-day reaction (Fig. 7B, Supplementary Fig. 24D). Reaction profiles following new substrate addition match 6-AHA equivalent release seen in standard reactions like those shown in Figs. 5 and 6, indicating that NylC_K-TS retains almost full activity after both three and seven days of reaction at 60 °C, and that there are no inhibitory compounds present which could explain reaction stalling. Characterization of the PA6 films by GPC suggests that there were no extensive changes in the number average molar mass (M_n) or molar mass dispersity (Đ) of the PA6 polymer chains following 10 days of enzyme incubation with NylC_K-TS at any temperature (Supplementary Table. 3, Supplementary Fig. 25A). There were also no significant differences in the percentage crystallinity of the PA6 substrate incubated with or without enzyme over the course of 10 days, as measured by DSC (Supplementary Fig. 25B). As a note, TGA analysis revealed that the PA6 film absorbed an average of 1.9 wt% water following 10 days incubation in reaction buffer across all temperatures (no enzyme controls), and there was no noticeable difference in substrate water absorption in reactions with NylC_K-TS (average of 1.7 wt% across all temperatures) (Supplementary Table. 3, Supplementary Fig. 25C).

Taken together, these results suggest that the reaction plateau for NylC_K-TS is a consequence of lack of remaining hydrolysable substrate for this enzyme following 10 days of reaction. Furthermore, typically for more extensive enzymatic plastic depolymerizations, both rate and extent of deconstruction are highly sensitive to substrate crystallinity⁶¹. However, for PA6 depolymerization with NylC_K-TS, using a more crystalline PA6 film substrate (23.0% crystallinity by DSC, prepared in house, full material characterization in Supplementary Table 1, Supplementary Fig. 26), leads to a comparable amount of 6-AHA equivalents release compared to reactions with the Goodfellow film that is more amorphous (13.2% crystallinity by DSC, Fig. 7C). Hence, we can conclude that NylC_K-TS is likely only working on a small amount of accessible nylon polymer on the film surface, and so does not reach the depolymerization extents at which substrate crystallinity would play a role in reaction progression. Indeed, SEM images of PA6 incubated with NylC_K-TS (1 µM enzyme, 0.08 mM enzyme/g PA6, 0.65 wt% substrate loading, 100 mM pH 7.5 NaPi buffer, 150 mM NaCl, 60 °C, 10 days), reveal a slight surface roughening, but no significant pits or features that are commonly associated with more extensive biocatalytic polymer deconstruction (Fig. 7D, Supplementary Fig. 27)^58,86. As these results indicate a surface area-limited process, we attempted to alter the surface area of the available PA6 substrate whilst retaining the same mass. Typically, this is done by using a powdered substrate; however, we found that enzymatic assays with PA6 powder (sourced from Goodfellow, particle size: 5–50 µm, full material characterization detailed in Supplementary Table 3) were very difficult to standardize due to reaction volume changes over the course of incubation (Supplementary Fig. 28). Additionally, the powder exhibited a high crystallinity (47.6% crystallinity by DSC), which may be undesirable. Hence, we conducted assays using the same mass of a thicker PA6 film (PA6 thick film, 0.5 mm thickness, 13 mg = two 0.32 ×0.32 cm squares, sourced from Goodfellow, full material characterization detailed in Supplementary Table 3), representing a ~ 50% reduction in PA6 surface area in each reaction compared to our standard 0.2 mm thickness film. Using the PA6 thick film, product release proceeded at a slower rate, with 34% fewer 6-AHA oligomers released over the course of 10 days, compared to the standard PA6 film (1 µM of enzyme, 0.08 mM enzyme/g PA6, 0.65 wt% substrate loading, 100 mM pH 7.5 NaPi buffer, 150 mM NaCl, 60 °C, Supplementary Fig. 29), confirming that surface area is indeed a factor controlling both rate of reaction and extent of depolymerization.

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• Source: https://www.nature.com/articles/s41467-024-45523-5