Patent Application
20240026114
Successfully interfacing enzymes and biomachineries with polymers affords on-demand modification and/or programmable plastic degradation during manufacture, utilization, and disposal, but requires controlled biocatalysis in solid matrices with macromolecular substrates.1-7 Embedded enzyme microparticles have sped up polyester degradation, but compromise host properties and unintentionally accelerate microplastics formation with partial polymer degradation.6,8,9
We disclose that by nanoscopically dispersing enzymes with deep active sites, semi-crystalline polyesters can be degraded primarily via chain-end mediated processive depolymerization with programmable latency and material integrity, akin to polyadenylation-induced mRNA decay.10 We also disclose how to realize the processivity with enzymes having surface-exposed active sites by engineering enzyme/protectant/polymer complexes. By example, polycaprolactone and poly(lactic acid) containing less than 2 wt. % enzymes are depolymerized in days with up to 98% polymer-to-small molecule conversion in standard soil composts or household tap water, completely eliminating current needs to separate and landfill their products in compost facilities. Furthermore, oxidases embedded in polyolefins retain activities. However, the hydrocarbon polymers do not closely associate with enzymes like their polyester counterparts and the reactive radicals generated cannot chemically modify the macromolecular host. The disclosed molecular guidances provide enzyme/polymer pairing and enzyme protectants' selection to modulate substrate selectivity and optimize biocatalytic pathways.
The invention provides systems and methods for depolymerization of polyesters with nano-dispersed enzymes.
In an aspect the invention provides a system for programmable degradation of a plastic, comprising a plastic comprising a nanoscopic dispersion of enzymes and configured to exploit enzyme active sites and enzyme-protectant interactions to provide processive depolymerization as the primary degradation pathway with expanded substrate selectivity to effect substantially complete depolymerization without substantial microplastics formation with partial polymer degradation.
In an aspect the invention provides a method of programmable degradation of a plastic, comprising providing a plastic comprising a nanoscopic dispersion of enzymes and configured to exploit enzyme active sites and enzyme-protectant interactions to provide processive depolymerization as the primary degradation pathway with expanded substrate selectivity to effect substantially complete depolymerization without substantial microplastics formation with partial polymer degradation.
In embodiments:
The invention encompasses all combinations of the particular embodiments recited herein, as if each combination had been laboriously recited.
Main Figure Legends
Unless contraindicated or noted otherwise, in these descriptions and throughout this specification, the terms “a” and “an” mean one or more, the term “or” means and/or. It is understood that the examples and embodiments described herein are for illustrative purposes only and that various modifications or changes in light thereof will be suggested to persons skilled in the art and are to be included within the spirit and purview of this application and scope of the appended claims. All publications, patents, and patent applications cited herein, including citations therein, are hereby incorporated by reference in their entirety for all purposes.
We envy nature's ability to program complex processes to achieve system-wide, long-term sustainability.11-14 The key bottleneck is molecularly interfacing bio-elements with synthetic counterparts and, for enzyme-based plastic modification/degradation, how to manipulate biocatalysis with macromolecules being both the reaction substrates and host matrices.2,3,8,15 Enzymatic activity depends on the protein structure, substrate binding, and reactivity at the active site16-18. In semi-crystalline polymers, which represent the majority of plastics,13 substrate accessibility can be rate-limiting due to the reduced mobilities of the confined enzyme3,4,7 and polymer matrix19 (
By nanoscopically confining enzymes in semi-crystalline polyesters and exploiting enzyme-active-site features and enzyme-protectant interactions, we show that processive depolymerization can be enabled as the primary degradation pathway with expanded substrate selectivity. Nanoscopic dispersion of a trace amount of enzyme, e.g., ˜0.02 wt. % lipase (<2 wt. % total additives) in poly(caprolactone), PCL, or ˜1.5 wt. % proteinase K (<5 wt. % total additives) in poly(lactic acid), PLA, leads to near-complete conversion to small molecules, eliminating microplastics in a few days using household tap water and standard soil composts. The programmable degradation overcomes their incompatibility with industrial compost operations, making them viable polyolefin substitutes.28-30 Analysis on the effects of polymer conformation and segmental cooperativity guide the thermal treatment of the polyester to spatially and temporally program degradation, while maintaining latency during processing and storage. The protectants are designed to regulate biocatalysis and stabilize enzymes during common plastic processing. Furthermore, with embedded oxidases such as laccase and manganese peroxidase, the enzymatically generated reactive radicals cannot oxidize the host polyolefins. There is a need to understand the biocatalytic cascades to design enzyme/host interactions and to enhance reactivity, diffusion, and lifetimes of reactive species without creating biohazards.
Biodegradable plastics PCL and PLA are market-ready alternates to many commodity plastics with increasing production and cost reduction.34 However, they are indifferentiable in landfills.14 Typical residence times are not adequate to allow for full breakdown even in thermophilic digesters operating at 48-60° C., 28,29 resulting in operational challenges and a financial burden to minimize contamination in organic waste.30 Burkholderia cepacia lipase (BC-lipase) and Candida Antarctica lipase (CA-lipase) were embedded in PCL and proteinase K was embedded in PLA given their known hydrolysis ability in solution.15 A previously developed four-monomer random heteropolymer (RHP) was added to nanoscopically disperse the enzymes.5,7 RHPs adjust the segmental conformations to mediate interactions between enzymes and local microenvironments.5 Extended Data Table 1 details the compositions of all blends.
Nano-Dispersed Lipase Accelerates PCL Degradation
At 0.02-2 wt. % enzyme loading, RHP-lipase nanoclusters are uniformly distributed throughout (
The overall PCL crystallinity in PCL-RHP-BC-lipase does not change when the degradation weight loss increased from 20% to 80% (
Design Enzyme/Polymer Blends to Realize Processive Depolymerization
When BC-lipase nanoclusters are embedded in pure PLA or a PCL/PLA blend, no PLA hydrolysis is observed even though lipase catalyzes a broad range of hydrolysis reactions.35 However, when the host matrix is a PCL-b-PLA diblock copolymer (40-b-20 kDa), both the PCL and PLA block depolymerize into small-molecules in a similar molar ratio as the parent copolymer (
BC-lipase shares common traits with processive enzymes.23,24 It has a deep (up to 2 nm), narrow (4.5 Å at the base) hydrophobic cleft from its surface to the catalytic triad,17 which may facilitate substrate polymer chain sliding while preventing dissociation. Opposite to the hydrophobic binding patch are six polar residues, providing a potential driving force to pull the remaining chain forward after hydrolysis (
Without nanoscopic confinement, BC-lipase degrades PCL via random chain scission in solution. When BC-lipase is embedded as micron-sized aggregates, the host degradation stops after ˜40% mass loss and leads to highly crystalline, long-lasting microplastics (
The turnover rate for embedded BC-lipase is ˜30 s−1 for 0-3 hours and ˜12 s−1 after 3 hours. The turnover rates of BC-lipase are ˜200 s−1 in solution with small molecule substrate, ˜19 s−1 in solution with a PCL film as substrate and ˜120 s−1 in PCL-RHP-BC-lipase with a small molecule substrate (
Therefore, to realize chain-end mediated processive depolymerization, the enzyme should be nanoscopically confined to co-reside with the polymer chain ends, exclude the middle segments from reaching the catalytic site, and have attractive interactions with the remaining chain end to slide the polymer chain without dissociation. With processive depolymerization, the host degrades with near-complete polymer-to-small molecule conversion, eventually eliminating highly crystalline microplastic particles. Kinetically, the apparent degradation rate benefits from substrate shuttling and catalytic latency can be regulated by thermal treatment and/or operation temperature.
Enzyme Protectants (RHPs) Modulate Enzyme Stability
RHPs assist nanoscopic dispersion of enzymes and affect the local micro-environment, substrate accessibility, and possibly the degradation pathway. A model experiment at the solvent/water interface was designed where the interfacial tension is used to monitor molecular associations of the enzyme, RHP, and polymer (
The RHPs modulate enzymes' micro-environment and provide entropic stabilization, enabling scalable processing of enzyme-embedded plastics using melt extrusion. PCL-RHP-BC-lipase containing ˜0.1 wt. % lipase was extruded at 85° C. to produce ˜1.5 mm diameter filament, which degraded completely over 36 hours in buffer by the same processive depolymerization mechanism (
Program Catalytic Latency
Polymer degradation can be programmed by thermal treatments. As the BC-lipase pulls the segments in the PCL stem spanning the crystalline lamellae, the competing force is governed by multiple pair-wise interactions between chains and degradation should not occur above a critical lamellae thickness. Indeed, PCL-RHP-BC-lipase films with thicker crystalline lamellae (crystallized at 49° C.) undergo negligible degradation over 3 months in 37° C. buffer, while films with thinner crystalline lamellae (crystallized at 20° C.) degrade over 95% in 24 hours (
Operation temperature is another handle to program degradation latency. There is a much lower conformational entropic penalty for a crystallized chain segment to bind to an enzyme than a completely amorphous chain.36 The high entropic penalty for enzyme binding overtakes the effects of increased chain mobility, leading to large reductions in degradation rates at higher temperatures (>43° C.) (
Enzyme Protectants (RHPs) Modulate Catalytic Kinetics and Pathway
Proteinase K readily degrades PLA but the active site is highly surface-exposed, such that partial PLA degradation occurs with random chain scission, leaving highly crystalline microplastics behind. We hypothesize that modulating interactions between proteinase K binding site and RHPs may create an RHP-covered active site to achieve the characteristics of processive enzymes without protein engineering. We experimentally screened RHPs guided by the analysis of RHP segmental hydrophobicity38 (
Hydrocarbon Substrate is Inaccessible to Embedded Oxidases
Besides synthetic catalysts, 22 biocatalysis of hydrocarbons is highly desirable due to its known efficiency, selectivity, and programmability.31 However, polyolefin degradation has mainly been reported using microbes, as opposed to enzymes.21 Polyolefin degradation is often initiated by side-chain modification, such as oxidation. To probe the bottlenecks, manganese peroxidase from white rot fungus and laccase from Trametes versicolor were embedded either in polyethylene or polystyrene with and without mediators (Tween 80 for manganese peroxidase and hydroxybenzotriazole for laccase). After two weeks in malonate buffer at 30° C. or 60° C., no changes are observed for any enzyme-polyolefin blends by infrared spectroscopy and gel permeation chromatography. For biosafety, these results are reassuring and expected with known longevity of plastic wastes. However, both enzymes remain highly active inside the plastics based on colorimetric assays, confirming formation of diffusive reactive radicals (
Our technology enables fabrication of functional plastics with programmable life cycles compatible with plastic melt processing. Considering recent developments in synthetic biology and biodegradable plastic production, 14,34,39 modulating biocatalysis of embedded enzymes can provide molecular control over reaction pathway, kinetics, latency, and production of high value by-products.
Section M1. Embedding Random Heteropolymer-Enzymes in Polyesters
Amano PS Lipase from Burkholderia cepacia (BC-lipase), Candida Antarctica Lipase B (CA-Lipase), and proteinase K from Tritirachium album were purchased from Sigma Aldrich. The BC-enzyme solution was purified following established procedure.40 Proteinase K was purified by using a 10,000 g/mole molecular weight cutoff filter by spinning in a centrifuge at 6,000 rcf for 3 total cycles. The concentration of the purified lipase and proteinase K stock solution was determined using UV-vis absorbance at 280 nm. Detailed information for all samples is listed in Table 51.
The random heteropolymer (RHP) (70 KDa, PDI=1.55) was synthesized.5 The monomer molar composition used, unless otherwise specified, was 50% methyl methacrylate (MMA), 20% 2-ethylhexyl methacrylate (EHMA), 25% oligo(ethylene glycol methyl ether methacrylate) (OEGMA; Mn=500 g/mole), and 5% 3-sulfopropyl methacrylate potassium salt (SPMA). The RHP is referred as MMA:EHMA:OEGMA:SPMA=0.5:0.2:0.25:0.5. Two RHP variants were used to perform experiments described in
RHP and enzymes were mixed in aqueous solution, flash-frozen in liquid nitrogen, and lyophilized overnight. The dried RHP-enzyme mixture was resuspended directly in the specified polymer solutions or melts. RHP was mixed with purified BC-lipase in a mass ratio of 80:1 (total polymer matrix mass=98.4%). For commercial BC-lipase and CA-lipase blends, the RHP to blend weight ratio was kept at 2:1 (total polymer matrix mass=95.5%). For proteinase K in PLA, a 2:1 RHP:enzyme ratio was used (total polymer matrix mass=95.5%).
PCL (80 KDa) and PLA (85-160 KDa) were purchased from Sigma Aldrich and used without further purification. To prepare solution-cast films, PCL (or PLA) was dissolved in toluene (or dichloromethane) at 4 wt. % concentration and stirred for at least 4 hours to ensure complete dissolution. The dried RHP-enzyme complexes were resuspended at room temperature directly in the polymer solution at the specified enzyme concentration. Mixtures were vortexed for ˜5 mins before being cast directly on a glass plate. PCL films were air dried and PLA films were dried under a glass dish to prevent rapid solvent evaporation given the volatility of dichloromethane.
To probe enzyme distribution, lipase was fluorescently labeled. NHS-Fluorescein (5/6-carboxyfluorescein succinimidyl ester) was used to label lipase and remove excess dye by following manufacturer's procedure. A U-MWBS3 mirror unit with 460-490 nm excitation wavelengths was used to take the fluorescence microscopy images. TEM images were taken on a JEOL 1200 microscope at 120 kV accelerating voltage. Vapor from a 0.5 wt. % ruthenium tetroxide solution was used to stain the RHP-lipase and the amorphous PCL domains.
Section M2. Characterization of as-Cast Plastics
Dynamic light scattering (DLS) was used to obtain the complex's particle size in toluene. Crystallinity and mechanical properties of enzyme-embedded polyesters were probed via differential scanning calorimetry (DSC) and tensile testing, respectively. For DSC, ˜5 mg PCL films were pressed into aluminum pans and heated from 25° C. to 70° C. at a 2° C./min scan rate. To quantify percent crystallinity, the sample's enthalpy of melting was normalized by 151.7 J/g, enthalpy of melting for 100% crystalline PCL.41 For uniaxial tensile tests, PCL solutions were cast directly in custom-designed Teflon molds with standard dog-bone shapes. For small angle x-ray scattering (SAXS) studies, ˜300 μm thick films were cast in Teflon beakers. Samples were vacuum dried after degradation for 16 hours prior to running SAXS at beamline 7.3.3 at the Advanced Light Source (ALS). X-rays with 1.24 Å wavelength and 2 s exposure times were used. The scattered X-ray intensity distribution was detected using a high-speed Pilatus 2M detector. Images were plotted as intensity (I) vs. q, where q=(4π/λ) sin(θ), λ is the wavelength of the incident X-ray beam, and 2θ is the scattering angle. The sector-average profiles of SAXS patterns were extracted using Igor Pro with the Nika package. The same SAXS method was used to analyze the nanoporous structure of samples at different time points of the degradation process, as shown in
Section M3. Characterization of Enzyme-Embedded PCL Degradation
Degradation was carried out in sodium phosphate buffer (25 mM, pH 7.2) at temperature specified. Mass loss was determined by drying the remaining film and measuring mass on a balance. After 24 hours, mass loss was estimated by integrating gel permeation chromatography (GPC) peaks. The microplastic experiment shown in
At each timepoint from 0-5 hours, PCL-RHP-BC-lipase remaining films were dried and analyzed via DSC to determine crystallinity. To analyze degradation by-products, vials were lyophilized overnight before resuspending in the proper solvent for GPC or LCMS. GPC measurements were run using a total concentration of 2 mg/mL of remaining film and by-product in THF. 20 uL of solution was injected into an Agilent PolyPore 7.5×300 mm column; GPC spectrum for BC-lipase in solution was normalized to the solvent front. Liquid chromatography-mass spectrometry (LC-MS) measurements were obtained by resuspending degradation supernatant in acetonitrile/water (67/33 vol %), using an Agilent InfinityLab EC-C18, 2.7 μm column. Control experiments for surface erosion were run with ˜0.15 mg/mL total BC-lipase blend concentration. The mass spectrum shown in
Section M4. Enzyme Active Site Affects Degradation by Confined Enzymes
RHP-BC-lipase was embedded in a PCL-b-PLA deblock copolymer blended with pure PLA for the testing because the diblock on its own was too brittle to form a freestanding film after drying. The film was cast from a solution of 9 wt. % PCL-b-PLA (purchased from Polymer Source)+4 wt. % pure PLA in dichloromethane. The film was allowed to degrade at 40° C. buffer for 24 hours, and the by-products were analyzed using NMR. Similar results were obtained for homemade PCL-b-PLA diblock copolymer without any blended pure PLA homopolymer (10k-b-8k based on NMR analysis).
Crystal structures of BC-lipase and CA-lipase are taken from entries SLIP and 1TCA in protein data bank, respectively. Analysis of proteinase K active site was carried out using entry 3PRK. Hydrophobic residues (gray) are defined as the following amino acids: alanine, glycine, valine, leucine, isoleucine, phenylalanine, methionine, and proline. Aspartic acid and glutamic acid are defined as negative residues (red), while lysine, arginine, and histidine are defined as positive residues (blue). The remaining residues are considered polar uncharged residues (purple). GPC on PCL-RHP-CA-lipase films (degraded in 37° C. buffer) was carried out following the same procedure as for BC-lipase-embedded films.
Section M5. Confinement Affects Degradation Pathway: Nanoscopic Vs. Microscopic Vs. Surface Erosion
Degradation was run in a 1 mL and 1 L container while shaking the container every few hours to demonstrate the effects of enzyme leaching and diffusion. PCL-RHP-BC-lipase degrades similarly in both volumes (≥95% degradation in 24 hours), consistent with internal degradation and limited enzyme leaching.
Pure PCL films were placed in 1 L buffer with an equivalent mass of total lipase as was present in the PCL-RHP-lipase films. Pure PCL films exhibited negligible degradation in 1 L buffer over a week, whereas pure PCL films in 1 mL buffer with the same enzyme mass lost ˜80% mass in 1 day. This buffer volume dependence is expected, because enzyme must diffuse to plastic surface in order to hydrolyze the plastic.
To simulate experiments detailed in previous literature for comparison, 6,8 Tween80 was mixed with purified lipase in a 1:1 mass ratio, lyophilized, and resuspended in PCL/toluene to cast films. In 1 L buffer, films with Tween80-embedded enzyme at the same enzyme loading as PCL-RHP-BC-lipase degraded by ˜40% in 1 day and then stopped degrading (monitored over 1 week), whereas in 1 mL buffer the small molecule-embedded film degraded similarly to RHP-embedded film (≥95% in 24 hours). This reliance on buffer volume suggests that small molecule surfactant-embedded enzyme experiments previously reported in literature exhibit significant leaching, and in large volumes this enzyme leaching prevents complete polymer degradation.
Section M6. Kinetic Analysis of BC-Lipase in Different Environments with Different Substrates
M6.1 Confined BC-lipase with PCL substrate: The slope of the degradation plot shown in
M6.2 Dissolved BC-lipase with PCL substrate: Pure PCL films (˜5 mg each) were placed in 1 mL buffer (37° C.) containing ˜1 lag of lipase to mimic concentrations from degradation experiments of confined lipase. The turnover rate provided in the text was determined by also assuming a trimer by-product, which may represent an upper bound since surface erosion can occur by random scission (larger oligomers generated per bond cleavage would serve to reduce the apparent turnover rate since more mass is lost per bond cleavage).
M6.3 Dissolved and confined BC-lipase with small molecule substrate: The same small molecule assay was used to quantify activity of dissolved and confined BC-lipase. 4-nitrophenyl butyrate was dissolved in buffer at each substrate concentration prior to running the assay to rule out interfacial effects of soluble lipase. Activity was quantified via UV-vis to monitor the absorbance over 10 mins of the hydrolyzed by-product at 410 nm. Extinction coefficient for by-product was estimated as 16,500 M−1 cm−1. PRISM software was used to fit the activity as a function of substrate concentration in order to obtain Vmax, the theoretical maximum reaction rate at saturated substrate concentration. Vmax was converted to a turnover rate by converting per-mass to per-lipase molecule. The same small molecule assay was used to quantify activity of confined lipase in PCL.
Section M7. Dynamic Interfacial Tension Experiments to Probe PCL-RHP-Lipase Interactions
Interfacial tension between a toluene and water phase was used to probe the blends. A MilliQ water droplet was dispensed by a 1 mL syringe through a 1.27 mm-diameter needle and immersed in toluene. The droplet shape was captured by a CCD camera every second and fitted by Young-Laplace equation to obtain interfacial tension. For each sample, the measurement was repeated three times and showed good consistency and reproducibility.
RHP-lipase were mixed in a 10-1 mass ratio and lyophilized to remove the aqueous solvent. A different ratio was used here compared to actual degradation studies because 80-1 RHP-lipase resulted in unstable droplets due to high RHP interfacial activity, preventing accurate measurement. PCL was dissolved first in toluene at a 0.5 mg/mL concentration. The PCL/toluene solution was then used to directly disperse RHP-lipase, giving a final concentration of 0.005 mg/mL for RHP and 0.0005 mg/mL for lipase in toluene. The concentration of each component was fixed across all groups. The water droplet was immersed in toluene after all three components (PCL, RHP, and lipase) were dispersed in toluene.
To determine whether PCL alone could disperse lipase in toluene, fluorescently labeled lipase was dissolved in the water phase (0.75 mg/mL concentration) while PCL was dissolved in the toluene phase (0.5 mg/mL). The fluorescence intensity of both phases did not change over a 3-hour period (data not shown), indicating the inability of PCL alone to disperse lipase in toluene via the water/toluene interface.
Section M8. Melt Processing, Thermal Treatment, and Operating Temperature to Program Degradation
PCL (10,000 g/mole) was first ground into a fine powder using a commercial grinder. RHP-lipase dried powder (1-1 mass ratio) was mixed with PCL powder and all three components were again passed through the commercial grinder. The PCL-RHP-lipase powder was then placed in a single-screw benchtop extruder, with a rotating speed of 20 RPM and an extrusion temperature of 85° C. Melt-extruded PCL-RHP-lipase filaments degrade with the same processive mechanism, as confirmed by GPC and LCMS.
For thermal treatment, PCL-RHP-lipase films were cast on microscope slides, placed on a hot plate at 80° C. for 5 mM to ensure complete melting, and crystallized at the specified temperature for up to 3 days to ensure complete recrystallization.
To determine the dependence of degradation on operating temperature, PCL-RHP-BC-lipase solution-cast films were placed in buffer at specified temperatures. For as-cast films, ramping temperature from 20° C. to ˜43° C. results in increased degradation rates. Further increases in temperature, however, result in degradation rate decreases. To rule out enzyme denaturation, the same small molecule assay described in section MS was employed at the given temperatures. Controls of just the 0.5 mM ester solution were run at each temperature to ensure that the ester was not self-hydrolyzing over the given measurement time period. The activity toward the small molecule significantly increases above 43° C., ruling out denaturation as the cause for reduced PCL degradation at high temperatures.
Section M9. RHPs with Different Compositions Enable PLA Depolymerization and Regulation of Embedded Enzyme Activity
RHPs' compositions were screened to determine the effects of RHP-enzyme interactions on depolymerization by embedded enzymes. Three compositions were chosen based on the segmental hydrophobicity, which was determined by simulating RHP sequences. Briefly, RHP sequences were generated using Compositional Drift.43 The hydrophile-lipophile balance (HLB) value was used to evaluate the solubility of monomer side-chains through group contribution theory. Using the equation HLB=7+Σi ni HLBi, where ni is the number of the ith chemical group in the molecule with corresponding value HLBi. The HLB value for each monomer side chain was estimated as: HLB(MMA)=8.45, HLB(EHMA)=5.12, HLB(OEGMA)=11.4 and HLB(SPMA)=18.5. A lower HLB value denotes higher hydrophobicity and a higher value means greater hydrophilicity. A Python program was created to continuously calculate the average segmental HLB values for a window sliding from the alpha to the omega ends of the simulated RHP chains. The window advanced by one monomer each time. We used a span containing odd numbers of monomers and assigned the average HLB value of that span to its middle monomer. Window size of 9 was used as an intermediate segmental region size. Hydropathy plots were generated to visualize randomly sampled sequences for each RHP composition and window size. An HLB-threshold=9 was set to distinguish hydrophobic and hydrophilic segments. The sequences are then averaged both across positions along the chain as well as across all 15,000 sequences in a simulated batch, to make batch-to-batch comparisons on the average segmental (window) hydrophobicity.
Similar tensiometry experiments as those outlined in section M7 were carried out using RHP (0.005 mg/mL)-proteinase K (0.0025 mg/mL), PLA and dichloromethane. PLA showed little interfacial activity. For the 20:50 MMA:EHMA RHP, addition of PLA measurably reduced the interfacial activity of the RHP. The 50:20 MMA:EHMA RHP had similar interfacial activity with or without PLA.
Section M10. Depolymerization in ASTM Composts or Tap Water
PCL-RHP-BC-lipase films were placed in tap water or an at-home compost setup. For water, films were submerged in 100 mL of tap water from a sink, and degradation proceeded identically over 24 hours (<95%) at the specified temperature. Soil was purchased from a local composting facility. The total dry organic weight of the soil was determined by leaving a known soil mass in an oven set to 110° C. overnight and then weighing the remaining material mass. Water was added to the soil to achieve a total moisture content of 50 or 60%, consistent with ASTM standards. For PCL-RHP-BC-lipase, up to 40% mass loss and 70% mass loss was observed after 2 and 4 days, respectively, in the compost setup at 40° C. For PLA-RHP-proteinase K, ˜34% mass loss occurred for 40 KDa PLA and ˜8% mass loss occurred for 85-160 KDa PLA after 5 days in a 50° C. soil compost.
Section S11. Oxidative Enzymes Embedded in Polyolefins
Manganese peroxidase from white rot fungus and laccase from Trametes versicolor were purchased from Sigma and used as purchased. RHP (50:20 MMA:EHMA) was mixed with either enzyme in a 4:1 ratio. Both enzymes were embedded in polyethylene (Mw=35 KDa) or polystyrene (Mn=260 KDa). For polyethylene, enzymes were embedded by solution casting from a 5 wt. % solution in toluene or melt pressing at 95° C. from polyethylene powder. For polystyrene, enzymes were embedded by resuspending directly in a 10 wt. % polystyrene in dichloromethane solution. Enzymes were embedded with and without mediators (Tween 80 for manganese peroxidase and hydroxybenzotriazole for laccase). The films were then placed in 30° C. or 60° C. malonate buffer (pH 4.5) for up to two weeks. After drying the films, infrared spectroscopy and GPC were used and no changes were observable for any enzyme-polyolefin system.
To confirm that enzymes were still active after embedding inside polyolefins, the films were submerged in a 1 mM solution of the small molecule 2,2′-Azino-bis(3-ethylbenzothiazoline-6-sulfonic acid) diammonium salt (ABTS) in malonate buffer. The solution turned dark blue for both manganese peroxidase and laccase, demonstrating that the embedded enzymes retained a high portion of activity. Tensiometry tests were carried out using RHP-manganese peroxidase or RHP-laccase with or with PS in toluene in the same setup and concentrations outlined for PCL/lipase. RHP-enzyme clusters with both enzymes achieved the same final interfacial tension with or without PS present and no lag phase or change in final interfacial tension, suggesting that the PS chains do not strongly interact with the enzymes.
Additional References Used in the Methods Section:
1:1‡
This invention was made with government support under the DA Army Research Office, contract number W911NF-13-1-0232, and the Department of Energy, grant number DE-AC02-05-CH11231. The government has certain rights in the invention.
| Number | Date | Country | |
|---|---|---|---|
| 63173507 | Apr 2021 | US |
| Number | Date | Country | |
|---|---|---|---|
| Parent | PCT/US22/24171 | Apr 2022 | US |
| Child | 18473252 | US |
