Bioupcycling Technology for Sustainable Plastic Waste Management: History
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Subjects: Biotechnology & Applied Microbiology
Contributor:
cristiano varrone

Effective plastic bioupcycling processes that can act as a drive to increase waste removal from the environment and valorize post-consumer plastic streams, thus accelerating the implementation of a circular (plastic) economy.

  • bioupcycling
  • plastic waste
  • bioconversion
  • recycling
  • fermentation
  • biotechnology

1. Bioupcycling

To improve the circularity of the plastic sector, post-consumer plastic waste should be recycled at the highest possible level. However, currently implemented recycling technology mainly transforms the plastic waste into lower-value products (downcycling) or, at best, into the same level. There is an urgent need for new strategies that help improving the techno-economic feasibility of the recycled plastics (still not competitive with virgin fossil ones [12]) or that lead to improved value/properties of the new plastics (defined as “upcycling”) [13]. Besides biotechnological recycling, new upcycling processes are now extensively studied and under development. The possibility to convert post-consumer PET to polyhydroxyalkanoate (PHA) by enzymatic depolymerization and subsequent bacteria fermentation [1], for instance, allows researchers to obtain a bio-material with good technical substitution potential, novel properties (depending on the co-polymer) and biodegradability. From this perspective, post-consumer plastic can be upcycled rather than only recycled. Moreover, biological methods have the advantage that they can be applied to contaminated plastic waste (i.e., food or soil) and do not require previous separation of the different fractions. Furthermore, the high selectivity of enzymes could allow for a stepwise removal of specific components of the mixed-plastic waste, facilitating the downstream processing; thus, it can go beyond the limits of mechanical and chemical recycling [2].
Recent research in the biodegradation of plastic waste allowed for the establishment of plastic biodepolymerization processes for some plastic types, thus paving the path for greener plastic recycling processes [3][4][5][6][7][8][9][10].
Both natural and engineered enzymes for plastic depolymerization have been extensively studied [3][11][12][13][14][15][16]. Even though the cost of the enzymes is still of concern, a recent study on enzymatic recycling of PET has determined that it should only contribute to 4% of the overall operating costs [17]. The authors highlighted that an enzyme-based recycling process can be cost-competitive, and the constant development of enzyme performance and optimization of the process remains an opportunity to further improve the economic viability of this process. Once bioprocesses for bulk enzyme production have been established, the enzymatic degradation of plastic is a promising technology that will be implemented in the near future. Depolymerization enzymes can be utilized in multiple ways, including free enzymes, immobilized enzymes, extracellular enzymes of whole-cell biocatalysts, surface enzymes, and/or in the form of enzyme cocktails [18]. Moreover, synthetic biology can be applied to improve the catalytic activity of enzymes through protein engineering, e.g., direct evolution and rational protein design [19]. Plastic-degrading enzymes in microorganisms can evolve from their natural activity on recalcitrant biopolymers such as lignocellulose, chitin, and cutin. Polyester plastics could, for instance, be depolymerized by hydrolytic enzymes produced by bacteria or fungi, such as cutinases, esterases, lipases, ureases, and proteases, as they have hydrolyzable ester bonds in their backbone (Satti and Shah, 2020). An excellent example is the bacterial enzyme polyurethane hydrolases (PUase), capable of hydrolyzing polyurethane (PU) [20], or the well-known PETase and MHETase from Ideonella sakaiensis 201-F6, able to hydrolyze PET [15]. For plastics with a non-hydrolyzable C–C backbone, such as polyolefins (PE and PP), oxidative enzymes play a significant role in introducing active functional groups into the backbone, which consequently can undergo biodegradation [21]. Alkane hydroxylases from isolated Pseudomonas sp. E4 was also reported to play an important role in PE biodegradation [22]. This enzyme acts on the hydroxylation of C–C bonds to release primary or secondary alcohols, which are then further oxidized to ketones or aldehydes, and finally to carboxylic acids [23]. Thus, the identification and optimization of efficient enzymes represents a prerequisite for the development of bioupcycling processes. In fact, the enzymes are not going to use the polymers as a carbon source, as would happen with a microbial cell that degrades the polymers. This allows to fully recover the monomers for the subsequent upcycling step, for instance through the contribution of fermentation processes [24].
So, in conclusion, the established knowledge on biodepolymerization is expected to boost the development of new biocatalytic plastic upcycling processes, to produce value-added products and/or generate new (and more renewable) plastics with better properties, out of conventional plastic wastes. In this sense, the cost of renewable plastic production (which is still too high compared to conventional fossil-based plastics and related to feedstocks’ price and fluctuation) could be alleviated. Hence, renewable plastic will become more economically viable for general commodities, once large-scale production is reached. At present, bioupcycling with endogenous or engineered metabolic pathways has been demonstrated as a proof of concept and is a hot topic for researchers worldwide. The most relevant published studies have been reviewed in the sub-section below.

2. Bioupcycling of Polyethylene Terephthalate (PET)

PET is a petrochemical-based plastic produced on multimillion tons worldwide. It is a polyester made of the repeating units of ethylene glycol (EG) and terephthalic acid (TA). It is cheap and has a very low permeability to gas and moisture, making it an ideal material for single-use plastic bottles. The demand of PET worldwide is around 29 Mt in 2022 [25], while in Europe it reached 4.1 Mt in 2020 [26]. However, even if PET has a higher recycling rate than other plastic types (50% in Europe and 23% in the US for PET bottles), 69% of recycled PET was used for lower-grade applications such as trays, film, strapping, or fiber [27]. From these statistics, the PET management system clearly does not show a very high level of circularity; therefore, there is a clear need for new upcycling approaches to valorize PET into higher-value products or, at least, keep the material at the same level in the value chain.
Various upcycling strategies have been reported to valorize PET waste, including biotechnological processes and bioupcycling (Table 1). It typically combines depolymerization and fermentation/bioconversion strategies to produce new valuable products. Chemical processes (e.g., hydrolysis, alcoholysis, glycolysis, aminolysis, ammonolysis, and hydrogenolysis [28]) and thermal processes (e.g., hydrothermal liquefaction, pyrolysis, and microwave irradiation) are generally used for the depolymerization of PET. Interestingly, a considerable achievement through enzymatic or microbial degradation has also been reported, especially on PET, that can lead to the development of more sustainable bioupcycling processes. Since PET monomers are linked through hydrolyzable ester bonds, an increasing number of PET hydrolyzing (and/or surface modifying) enzymes have been reported recently. One of the major findings was, for instance, the discovery of Ideonella sakaiensis [15], a bacterium that can grow on PET as the only carbon source, thanks to the synergy of its enzymes IsPETase and MHETase that break down PET to mono-(2-hydroxyethyl)terephthalic acid (MHET), di-(2-hydroxyethyl)terephthalic acid (BHET), and finally to EG and TA. Subsequently, several studies reported the improvement of the mesophilic IsPETase through protein engineering, for example by increasing the hydrolytic activity [29] or the thermal stability, using rational protein engineering [3][13][30].
In order to boost enzyme production, several studies have been investigating the addition of signal peptides to the IsPETase gene to enhance its secretion during heterologous expression in Escherichia coli [31]. The use of heterogeneous immobilized biocatalysis through the use of different binding modules and linkers has also been extensively investigated [32][33]. Some of the most effective enzymes that act on PET hydrolysis are thermostable cutinases such as Humicola insolens cutinase (HiC) [4], Thermobifida fusca cutinase (TfCut2) [16], and leaf-branch compost cutinase (LCC) [34]. Improving their hydrolysis activity has also been reported using various methods, for example, by enhancing electrostatic interaction between TfCut2 and PET by cationic surfactant additive-based approach, and showed impressive biodegradation of PET by 97% within 30 h [5]. In general, the design of thermostable hydrolases has been intensively investigated during the last years [35] and resulted in the development of promising processes for enzymatic depolymerization. One of the biggest breakthroughs so far is probably represented by the study by Tournier and colleagues that engineered the LCC cutinase and increased its optimal reaction temperature up to the glass transition temperature (Tg) of PET [11]. Such bioprocesses can represent a valuable and sustainable alternative to thermochemical depolymerization and are laying the basis for the further conversion of plastic waste-derived monomers.
Bioupcycling post-consumer PET (or even polyolefins) to PHA has been gaining attention since this approach promotes the valorization of plastic waste into renewable biopolymers. PHA is a general term for microbial polyester of R-3-hydroxyalkanoic acids [36]. It is a promising substitute for several petroleum-based plastics, due to its superior thermal processability, biodegradability, and biocompatibility properties [37][38]. Kenny et al. reported using a two-step chemo-biotechnological process for upcycling PET to PHA [36]. First, the PET waste was hydrolyzed by pyrolysis at 450 °C at a feed rate of 1 kg/h, obtaining 77% of solid fraction (TA, oligomers, benzoic acid and others), 18% gas fraction (CO2, CO, H2, ethene, and others), and 6.3% liquid fraction (EG, acetic aldehyde and others). Second, the TA was dissolved in NaOH to generate sodium terephthalate, which was used as the sole carbon source to grow a PHA-accumulating strains. Pseudomonas putida GO16, P. putida GO19, and P. frederiksbergensis GO23 were found to consume sodium terephthalate and accumulated PHA up to 23–27% of CDW. GO19 was the most efficient at converting TA to PHA, with a productivity of 8.4 mgPHA/L/h. The research group continued to develop the bioprocess to enhance PHA production from TA by co-feeding with waste glycerol (WG) [39]. The fermentation was designed to have two distinct phases: the biomass growth phase and the PHA production phase. They found that when P. putida GO16 was fed with WG only during the growth phase (0–24 h) and WG and TA during the PHA production phase (24–48 h), the highest total PHA production was achieved (5.30 g/L). This strategy promoted a 2.0-fold higher PHA production than feeding with TA alone. This study showed that bioprocess engineering strategies are key to develop highly efficient bioupcycling of plastic waste.
Tiso et al. reported the bioupcycling of PET to PHA and hydroxyalkanoyloxyalkanoate (HAA) by using a combination of enzymatic hydrolysis and whole-cell biocatalyst [1]. PET was hydrolyzed by LCC, a polyester hydrolase capable of efficient PET depolymerization to TA and EG. Pseudomonas umsongensis GO16 KS3 was found to consume both TA and EG (the latter at a 3.5-fold lower rate than TA) within 23 h of cultivation, in a 5 L bioreactor producing PHA. The PHA production only reached 0.15 g/L, or 7% of CDW. Interestingly, HAA was produced from TA only, while EG was exclusively used for growth. A maximum HAA concentration of 35 mg/L was achieved with a production rate and yield of 5 mg/L/h and 0.01 gHAA/gTA, respectively [1]. Despite the low performance, this represents a highly interesting approach with potential industrial applications (once optimized). In fact, HAA can be directly polymerized with 4,4′-methyl diphenyl diisocyanate and butanediol (BDO) to form biopoly(amide urethane) (bio-PU).
Another study reported the bioconversion of TA, a monomer of PET, to muconic acid (MA), which is currently used to produce adipic acid (AA), an important monomer for various plastics [40]. PET was first depolymerized by microwave radiation at 230 °C for 50 min to TA and EG. Then, the E. coli strain CTL-1 (expressing TphAabc, TphB, and AroY, which is responsible for converting TA to catechol, via 1,2-dihydroxy-3,5-cyclohexadiene-1,4-dicarboxylate (DCD) and protocatechuic acid (PCA)) and the strain MA-1 (expressing CatA, which is a catechol 1,2-dioxygenase responsible for converting catechol to MA), were combined for MA synthesis. The MA concentration reported was 2.7 mM, accounting for a 85.4% molar yield (MA/TA). In the same study, gallic acid (GA), pyrogallol, vanillic acid (VA), and glycolic acid (GLA) were also produced from engineered stains, using TA. In addition, GLA was produced by EG-fermenting Gluconobacter oxydans KCCM 40109.
The recent study by Sadler and Wallace (2021) showed the development of a one-pot bioprocess to convert TA from PET waste into a value-added molecule, vanillin. PET from a post-consumer plastic bottle was firstly hydrolyzed to TA by semi-purified LCC at 72 °C. Then, the reaction was cooled down and the engineered strain, E. coli RARE_pVanX, was added to perform the bioconversion. E. coli RARE_pVanX was constructed with two plasmids that were encoded for different enzymes (terephthalate 1,2-dioxygenase, dihydroxy-3,5-cyclohexadiene-1,4-dicarboxylic acid dehydrogenase, carboxylic acid reductase, and catechol O-methyltransferase), which convert TA to vanillin via intermediates (PCA, VA, and dihydroxybenzaldehyde). The process optimization was performed by screening the protein-expression media (M9-glucose supplemented with L-Met and n-butanol (nBuOH)), increasing E. coli cell membrane permeability to TA (addition of 1% v/v n-BuOH), adjusting pH (5.5) and temperature (22 °C), and mitigating the toxicity of vanillin by in situ product removal, using oleyl alcohol. The final production of vanillin reached 789 μM (119 mg/L) or 79% conversion. [41].
The chemo-biological upcycling of PET to the multifunctional coating material, catechol, was also reported. First, PET waste was glycolyzed to a mixture of PET oligomers. Then, an enzymatic hydrolysis of the glycolyzed products was performed, turning the mixture without previous purification into TA, by Bacillus subtilis esterase (Bs2Est). The catechol production from TA was consequently conducted, using a catechol biosynthesis strain (obtained through the combination of the TA degradation module and catechol biosynthesis module in E. coli). The final titer of catechol was 5.97 mM, accounting for 99.5% conversion by mol of TA. Catechol shows great functions as a coating material without the need for an adherence layer, and its antibacterial activity is comparable to chitosan [42].
Table 1. PET bioupcycling.
Depolymerization Strategy Depolymerization
Products Used as a
Feedstock for
Fermentation Step		 Fermentation Strategy Products from Fermentation Titer Productivity Yield Ref.
Hydrolytic pyrolysis
at 450 °C		 Solid product mixture (terephthalic acid (TA), oligomers, benzoic acid, and others) Fermentation in shake flask containing 4.2 g/L of PET-derived sodium terephthalate and 67 mg/L of nitrogen at 30 °C for 48 h by Pseudomonas putida GO16 medium chain length PHA (mclPHA) 0.25 g/L 8.4 mgPHA/L/h 0.27 gPHA/gCDW [36]
Hydrolytic pyrolysis
at 450 °C		 Solid product mixture (TA, oligomers, benzoic acid, and others) Fermentation in shake flask containing 4.2 g/L of PET-derived sodium terephthalate and 67 mg/L of nitrogen at 30 °C for 48 h by P. putida GO19 mclPHA 0.25 g/L 8.4 mgPHA/L/h 0.23 gPHA/gCDW [36]
Hydrolytic pyrolysis
at 450 °C		 Solid product mixture (TA, oligomers, benzoic acid, and others) Fermentation in shake flask containing 4.2 g/L of PET-derived sodium terephthalate and 67 mg/L of nitrogen at 30 °C for 48 h by P. putida GO23 mclPHA 0.27 g/L 4.4 mgPHA/L/h 0.24 gPHA/gCDW [36]
Pyrolysis TA Fed-batch fermentation in 19.5 L-stirred tank reactor with controlled pH of 6.9 and dissolved oxygen (DO) level above 40% at 30 °C for 48 h by P. putida GO16 supplied with TA as the sole growth and PHA substrate mclPHA 2.61 g/L 0.05 g/L/h 0.30 gPHA/gCDW [39]
Pyrolysis TA Fed-batch fermentation in 19.5 L-stirred tank reactor with controlled pH of 6.9 and DO level above 40% at 30 °C for 48 h by P. putida GO16 supplied with waste glycerol (WG) as growth substrate and TA as PHA substrate mclPHA 5.22 g/L 0.11 g/L/h 0.36 gPHA/gCDW [39]
Pyrolysis TA Fed-batch fermentation in 19.5 L-stirred tank reactor with controlled pH of 6.9 and DO level above 40% at 30 °C for 48 h by P. putida GO16 supplied with TA as the sole growth and PHA substrate mclPHA 5.30 g/L 0.11 g/L/h 0.35 gPHA/gCDW [39]
Pyrolysis TA Fed-batch fermentation in 19.5 L-stirred tank reactor with controlled pH of 6.9 and DO level above 40% at 30 °C for 48 h by P. putida GO16 supplied with WG as growth and PHA substrate and TA as PHA substrate only mclPHA 4.98 g/L 0.10 g/L/h 0.35 gPHA/gCDW [39]
Pyrolysis TA Fed-batch fermentation in 19.5 L-stirred tank reactor with controlled pH of 6.9 and DO level above 40% at 30 °C for 48 h by P. putida GO16 supplied with WG and TA as both growth and PHA substrates mclPHA 4.42 g/L 0.09 g/L/h 0.36 gPHA/gCDW [39]
Enzymatic degradation by recombinant leaf-branch compost cutinase (LCC) TA, ethylene glycol (EG), mono-(2-hydroxyethyl)terephthalic acid (MHET), di-(2-hydroxyethyl)terephthalic acid (BHET) Fermentation in 5 L-stirred tank reactor with controlled pH of 7.0 and DO level above 20% at 30 °C for 28 h Pseudomonas umsongensis GO16 KS3 supplied with hydrolyzed PET at the amount to yield 40 mM of TA and EG and limited inorganic nutrient mclPHA 0.15 g/L NA 0.014 gPHA/gSubstrate [1]
Enzymatic degradation by recombinant LCC TA, EG, MHET, BHET Fermentation in shake flask containing Delf medium with diluted (1:20) hydrolyzed PET (TA and EG concentration of 15–18 mM) at 30 °C for 24 h by P. umsongensis GO16 KS3 pSB01 Hydroxyalkanoyloxy-alkanoate (HAA) 35 mg/L 5 mg/L/h 0.01 gHAA/gTA [1]
Microwave radiation for 50 min
at 230 °C		 TA Bioconversion by metabolically engineered E. coli strain PCA-1 and HBH-2 to convert TA to intermediate protocatechuic acid (PCA), and then to gallic acid (GA), at 30 °C and 250 rpm for 24 h in 50 mM Tris buffer (pH 7.0) containing 2% (w/v) glycerol GA 2.7 mM NA 0.925 MGA/MTA [40]
Microwave radiation for 50 min
at 230 °C		 TA Bioconversion by metabolically engineered E. coli strain PG-1a to convert TA to intermediate PCA, GA, and then pyrogallol, at 30 °C and 250 rpm for 6 h in 50 mM Tris buffer (pH 7.0) containing 2% (w/v) glycerol Pyrogallol 1.1 mM NA 0.327 MPyrogallol/MTA [40]
Microwave radiation for 50 min
at 230 °C		 TA Bioconversion for 6 h by metabolically engineered E. coli strain CTL-1 and MA-1 to convert TA to intermediate catechol, and then to muconic acid (MA) MA 2.7 mM NA 0.854 MMA/MTA [40]
Microwave radiation for 50 min
at 230 °C		 TA Bioconversion using double-catalyst VA-2a system for 48 h by metabolically engineered E. coli strain PCA-1 and OMT-2His to convert TA to intermediate PCA and then to vanillic acid (VA), in 50 mM Tris buffer (pH 7.0) containing 10% (w/v) glycerol, 10 g/L yeast extract, 20 g/L peptone, and 2.5 mM L-methionine VA 1.4 mM NA 0.416 MVA/MTA [40]
Microwave radiation for 50 min
at 230 °C		 EG Bioconversion by Gluconobacter oxydans KCCM 40109 using 10.7 mM of EG from PET hydrolysate as a feedstock, at 30 °C and 220 rpm in shake flask at the working volume of 1 L Glycolic acid (GLA) NA NA 0.986 MGLA/MEG [40]
- EG (mock substrate to study upcycling of PET-derived monomer) Fermentation in shake flask containing 10% (v/v) EG in 250 mM potassium phosphate buffer (pH 7.0) at 30 °C with gentle stirring and aeration at 1 VVM for 120 h by Pichia naganishii AKU 4267 GLA 105 g/L NA 0.880 MGLA/MEG [43]
- EG (mock substrate to study upcycling of PET-derived monomer) Fermentation in shake flask containing 10% (v/v) EG in 250 mM potassium phosphate buffer (pH 7.0) at 30 °C with gentle stirring and aeration at 1 VVM for 120 h by Rhodotorula sp. 3Pr-126 GLA 110 g/L NA 0.922 MGLA/MEG [43]
- EG (mock substrate to study upcycling of PET-derived monomer) Fermentation in shake flask containing 100 mM of EG in nitrogen limiting M9 medium (0.132 g/L of (NH4)2SO4) at 30 °C for more than 72 h by P. putida MFL185 (engineered strain that has the tac promoter inserted before the native glycolate oxidase operon and harbor overexpression) mclPHA NA NA 0.32 gPHA/gCDW and 0.06 gPHA/gEG [44]
- EG (mock substrate to study upcycling of PET-derived monomer) Anaerobic fermentation of 50 mM EG at 30 °C by acetogenic bacterium Acetobacterium woodii Acetate 10.4 mM 3.6 μmol/mg/h NA [45]
- EG (mock substrate to study upcycling of PET-derived monomer) Anaerobic fermentation of 50 mM EG at 30 °C by acetogenic bacterium A. woodii Ethanol 12.0 mM 4.8 μmol/mg/h NA [45]
Enzymatic degradation by semi-purified LCC (pH 10.0) at 72 °C for 48 h PET hydrolysate Bioconversion using metabolically engineered E. coli RARE_pVanX to convert TA to intermediate protocatechuate (PC), and then to vanillin using optimized condition: M9-glucose supplemented with L-Met and nBuOH as a protein expression media, pH 5.5, room temperature for 24 h, in situ product removal (ISPR) by oleyl alcohol Vanillin 300–400 μM NA NA [41]
- TA (mock substrate to study upcycling of PET-derived monomer) Bioconversion using metabolically engineered E. coli RARE_pVanX to convert TA to intermediate PC and then to vanillin using optimized condition: M9-glucose supplemented with L-Met and nBuOH as a protein expression media, pH 5.5, room temperature for 24 h, ISPR by oleyl alcohol Vanillin 789 μM NA 0.79 Mvanillin/MTA [41]
Chemical glycolysis
at 200 °C for 3 h		 Mixture of BHET, MHET, and PET oligomers at 84.8, 7.7, and 8.7%, respectively Enzymatic hydrolysis of the glycolyzed products (the mixture) into TA by Bacillus subtilis esterase (Bs2Est) (2 U/mL at 30 °C and 1000 rpm), following by producing catechol from PET hydrolysates using a catechol biosynthesis strain that was established using the combination of the TA degradation module and catechol biosynthesis module in E. coli (in 12 h) Catechol 5.97 mM NA 0.995 MCatechol/MTA [42]
Chemocatalytic glycolysis: PET was glycolyzed with EG as a solvent (1:4 w/w) and catalyzed by 1% (w/w) titanium (IV) butoxide at 220 °C overnight BHET Fermentation in 3 L-bioreactor with batch culture in the first 4 h fed with 4-hydroxybenzoic acid to induce the β-ketoadipate pathway, followed by fed-batch culture using BHET as a carbon source (pulse adding at 9.1, 23.3, 32.8, 48.2 and 73.9 h). The sequential metabolic engineered Pseudomonas putida KT2440 (constitutive expression of native genes for EG utilization, expression of gene for TA catabolism, expression of PETase and MHETase for BHET hydrolysis, and gene deletion to enhance β-ketoadipic acid production) was used for bioconversion. β-ketoadipic acid 15.1 g/L 0.16 g/L/h 0.76 Mβ-ketoadipic acid/MBHET [46]
Chemocatalytic glycolysis and enzymatic hydrolysis: PET was glycolyzed with EG as a solvent and catalyzed by betaine at 190 °C for 30–120 min, followed by enzymatic hydrolysis (PETase and MHETase) TA Whole cell bioconversion of TA (4.5 g/L) to protocatechuic acid by metabolically engineered E. coli PCA-1 was performed in shake flask at 30 °C and 250 rpm PCA 3.8 g/L - 0.904 MPCA/MTA [47]
Chemocatalytic glycolysis and enzymatic hydrolysis: PET was glycolyzed with EG as a solvent and catalyzed by betaine at 190 °C for 30–120 min, followed by enzymatic hydrolysis (PETase and MHETase) EG Whole cell bioconversion of EG (30.6 g/L) to GLA by Gluconobacter oxydan KCCM 40109 was performed in shake flask at 30 °C and 200 rpm. GLA 31.4 g/L - 0.916 MGLA/MEG [47]

3. Bioupcycling of Polyurethanes (PU)

PU is a general term used for a class of polymers typically derived from the polycondensation of (poly)isocyanates (−NCO) and polyols (exothermic reactions) [48]. There are three main types of PU: polyester, polycaprolactone, and polyether urethanes, depending on the type of long-chain diol used in PU production [49]. PUs are used in many fields of application, such as, building insulation, pillows and mattresses, and insulating foams for fridges [26]. In 2020, PU production was 3.81 Mt in Europe, representing 7.8% of total plastic production [26]. Since many PU types have a thermoset nature with covalent cross links, recycling is still extremely challenging. Almost 50% of PU produced in the European Union goes to a landfill, 45% being incinerated with energy recovery, and only 5% is mechanically recovered [50]. The statistics from the US show that the mechanical recycling of PU on a large scale has only been performed on flexible PU foams by shredding and mixing the scrap polymers with binders to produce padding-type products; such processes have the limitation that they can be repeated only 8–10 times, until reaching the end-of-life [51]; this represents a common problem with mechanical recycling. Chemical recycling methods used for PU may include hydrolysis, glycolysis, and aminolysis, which use water, glycol, and ammonia or ammonium hydroxide as a nucleophile, respectively [52]. Europe’s first chemical recycling plant for post-consumer PU foam was constructed very recently in Semoy, France, by the Dow Chemical Company and partners, employing glycolysis for recovering polyols [53]. Another method called acidolysis, based on a combination of two carboxylic acids and an unspecified non-metallic catalyst, has been successfully employed by H&S, a Poland-based company, to break up PU and recover polyols [54]. Even though these processes can recover chemical constituents from PU waste, the negative impact from the toxic isocyanides and the huge amount of chemical catalysts consumption cannot be neglected. Therefore, despite the recalcitrance of these polymers, biodegradation of PU using microorganisms (i.e., fungal species) or enzymes is gaining more interest lately, as a more environmentally friendly and sustainable alternative [55][56]. On the other hand, since PUs represent a large class of plastics, it is difficult to point to a single enzyme that can degrade them all. In the presence of hydrolase-type enzymes, e.g., lipase, esterase, and protease, PUs can be hydrolyzed by a three-step mechanism: (1) chemical dissolution of ester and amide bonds in the polymer chain; (2) decreasing molecular weight and viscosity; (3) cleaving all polymer chains [49]. However, urethane forms stable bonds that can be hydrolyzed at a slower rate than ester bonds [57]. Bacillus subtilis strain MZA-75 [58] and Pseudomonas aeruginosa strain MZA-85 [59] were reported for polyester-based PU degradation. These two strains were also studied in bacterial co-culture and found to utilize polyester PU as a carbon and energy source more efficiently, as compared to the individual strains. The main degradation products were BDO and AA [60]. Another study investigated PU degradation by the enriched mixed-microbial consortia and reported the presence of extracellular enzymes, such as esterases, proteases, and ureases, affecting ester, urethane, ether, aromatic, and aliphatic groups of the plastic [61]. Fungi, such as Aspergillus tubingensis [62], Penicillium sp. [63], Cladosporium cladosporioides complex [64], or fungal communities [65] were also reported for their ability to degrade PU.
Besides BDO and AA, other PU degradation products are EG, 2,4′-toluenediamine (TDA), and 4,4′-methylenedianiline (MDA). They can be used to synthesize new PU or other polyesters, e.g., PHA, polybutylene succinate (PBS), poly(1,3-propylene succinate-ran-1,4-butylene succinate) (PPBS), and poly(1,3-propylene adipate-ran-1,4-butylene adipate) (PPBA) [52].
Last but not least, a recent study reported the upcycling processes of PU waste to rhamnolipid biosurfactants, [48] (Table 2). They successfully showed the growth of a defined mixed culture composed of three Pseudomonas putida KT2440 strains obtained by adaptive laboratory evolution on mock PU hydrolysate. The highest specific growth rate was 0.21 h−1 on the mixture of AA, 1,4-BDO, and EG. After engineering of the mixed culture, 0.10 g/L of rhamnolipids were produced from the AA, BDO, and EG mixture.
Table 2. PU bioupcycling.
Depolymerization Strategy Depolymerization Products Used as a Feedstock for
Fermentation Step		 Fermentation Strategy Products from
Fermentation		 Titer Productivity Yield Ref.
Enzymatic degradation of polycaprolactone polyol-based PU by esterase (E3576) in 0.1 M phosphate buffer (pH 7.0). The enzyme solution was replaced every 3–4 d to overcome a loss of enzymatic activity. 6-hydroxycaproic acid (1 g/L) - - - - - [66]
- Adipic acid (AA) (mock substrate to study upcycling of PU-derived monomer) Bioconversion (at 30 °C and 200 rpm for 135 h) using metabolically engineered P. putida KT2440 A12.1p pPS05 to convert AA into HAA and then to rhamnolipid Rhamnolipid 0.02 g/L NA 0.014 gRhamnolipid/gSubstrate [48]
- 1,4-Butanediol (BDO) (mock substrate to study upcycling of PU-derived monomer) Bioconversion (at 30 °C and 200 rpm for 135 h) using metabolically engineered P. putida KT2440 B10.1 pPR05 to convert BDO into HAA and then to rhamnolipid Rhamnolipid 0.13 g/L NA 0.088 gRhamnolipid/gSubstrate [48]
- EG (mock substrate to study upcycling of PU-derived monomer) Bioconversion (at 30 °C and 200 rpm for 135 h) using metabolically engineered P. putida KT2440 ∆gclR ∆PP_2046 ∆PP_2662::14d to convert EG into HAA and then to rhamnolipid Rhamnolipid 0.07 g/L NA 0.038 gRhamnolipid/gSubstrate [48]
- AA + BDO + EG (mock hydrolysate to study upcycling of PU-derived monomers) Bioconversion (at 30 °C and 200 rpm for 210 h) using mixed culture of three metabolically engineered P. putida KT2440 to convert the mock hydrolysate into HAA and then to rhamnolipid Rhamnolipid 0.1 g/L NA 0.008 gRhamnolipid/gSubstrate [48]
  AA + BDO + EG + 2,4-toluenediamine (TDA) (mock hydrolysate to study upcycling of PU-derived monomers) Bioconversion (at 30 °C and 200 rpm for 210 h) using mixed culture of three metabolically engineered P. putida KT2440 to convert the mock hydrolysate into HAA and then to rhamnolipid without extraction of TDA Rhamnolipid 0.02 g/L NA 0.002 gRhamnolipid/gSubstrate [48]
  AA + BDO + EG + TDA (mock hydrolysate to study upcycling of PU-derived monomers) Bioconversion (at 30 °C and 200 rpm for 210 h) using mixed culture of three metabolically engineered P. putida KT2440 to convert the mock hydrolysate into HAA and then to rhamnolipid with extraction of TDA at pH 3.5 Rhamnolipid 0.07 g/L NA 0.005 gRhamnolipid/gSubstrate [48]
 
 
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This entry is adapted from the peer-reviewed paper 10.3390/polym14224996

References

  1. Tiso, T.; Narancic, T.; Wei, R.; Pollet, E.; Beagan, N.; Schröder, K.; Honak, A.; Jiang, M.; Kenny, S.T.; Wierckx, N.; et al. Towards bio-upcycling of polyethylene terephthalate. Metab. Eng. 2021, 66, 167–178.
  2. Vollmer, I.; Jenks, M.J.F.; Roelands, M.C.P.; White, R.J.; van Harmelen, T.; de Wild, P.; van der Laan, G.P.; Meirer, F.; Keurentjes, J.T.F.; Weckhuysen, B.M. Beyond Mechanical Recycling: Giving New Life to Plastic Waste. Angew. Chem. Int. Ed. 2020, 59, 15402–15423.
  3. Son, H.F.; Cho, I.J.; Joo, S.; Seo, H.; Sagong, H.Y.; Choi, S.Y.; Lee, S.Y.; Kim, K.J. Rational Protein Engineering of Thermo-Stable PETase from Ideonella sakaiensis for Highly Efficient PET Degradation. ACS Catal. 2019, 9, 3519–3526.
  4. Carniel, A.; Valoni, É.; Nicomedes, J.; Gomes, A.d.C.; de Castro, A.M. Lipase from Candida antarctica (CALB) and cutinase from Humicola insolens act synergistically for PET hydrolysis to terephthalic acid. Proc. Biochem. 2017, 59, 84–90.
  5. Furukawa, M.; Kawakami, N.; Tomizawa, A.; Miyamoto, K. Efficient Degradation of Poly(ethylene terephthalate) with Thermobifida fusca Cutinase Exhibiting Improved Catalytic Activity Generated using Mutagenesis and Additive-based Approaches. Sci. Rep. 2019, 9, 1–9.
  6. Yao, C.; Xia, W.; Dou, M.; Du, Y.; Wu, J. Oxidative degradation of UV-irradiated polyethylene by laccase-mediator system. J. Hazard. Mater. 2022, 440, 129709.
  7. Zhang, H.; Kong, D.; Wang, L.; Xia, W.; Yao, C.; Wu, J. Degradation of UV-pretreated polyolefins by latex clearing protein from Streptomyces sp. Strain K30. Sci. Total Environ. 2022, 806, 150779.
  8. Sanluis-Verdes, A.; Colomer-Vidal, P.; Rodríguez-Ventura, F.; Bello-Villarino, M.; Spinola-Amilibia, M.; Ruiz-López, E.; Illanes-Vicioso, R.; Castroviejo, P.; Cigliano, R.A.; Montoya, M.; et al. Wax worm saliva and the enzymes therein are the key to polyethylene degradation by Galleria mellonella. bioRxiv 2022, 13, 5568.
  9. Jeon, H.J.; Kim, M.N. Functional analysis of alkane hydroxylase system derived from Pseudomonas aeruginosa E7 for low molecular weight polyethylene biodegradation. Int. Biodeterior. Biodegrad. 2015, 103, 141–146.
  10. Yang, Y.; Yang, J.; Wu, W.M.; Zhao, J.; Song, Y.; Gao, L.; Yang, R.; Jiang, L. Biodegradation and Mineralization of Polystyrene by Plastic-Eating Mealworms: Part 1. Chemical and Physical Characterization and Isotopic Tests. Environ. Sci. Technol. 2015, 49, 12080–12086.
  11. Tournier, V.; Topham, C.M.; Gilles, A.; David, B.; Folgoas, C.; Moya-Leclair, E.; Kamionka, E.; Desrousseaux, M.L.; Texier, H.; Gavalda, S.; et al. An engineered PET depolymerase to break down and recycle plastic bottles. Nature 2020, 580, 216–219.
  12. Shi, L.; Liu, H.; Gao, S.; Weng, Y.; Zhu, L. Enhanced extracellular production of IsPETase in Escherichia coli via engineering of the pelB signal peptide. J. Agric. Food Chem. 2021, 69, 2245–2252.
  13. Rennison, A.; Winther, J.R.; Varrone, C. Rational Protein Engineering to Increase the Activity and Stability of IsPETase Using the PROSS Algorithm. Polymer 2021, 13, 3884.
  14. Nakamura, A.; Kobayashi, N.; Koga, N.; Iino, R. Positive charge introduction on the surface of thermostabilized PET hydrolase facilitates PET binding and degradation. ACS Catal. 2021, 11, 8550–8564.
  15. Yoshida, S.; Hiraga, K.; Takehana, T.; Taniguchi, I.; Yamaji, H.; Maeda, Y.; Toyohara, K.; Miyamoto, K.; Kimura, Y.; Oda, K. A bacterium that degrades and assimilates poly(ethylene terephthalate). Science 2016, 351, 1196–1199.
  16. Roth, C.; Wei, R.; Oeser, T.; Then, J.; Föllner, C.; Zimmermann, W.; Sträter, N. Structural and functional studies on a thermostable polyethylene terephthalate degrading hydrolase from Thermobifida fusca. Appl. Microbiol. Biotechnol. 2014, 98, 7815–7823.
  17. Singh, A.; Rorrer, N.A.; Nicholson, S.R.; Erickson, E.; DesVeaux, J.S.; Avelino, A.F.T.; Lamers, P.; Bhatt, A.; Zhang, Y.; Avery, G.; et al. Techno-economic, life-cycle, and socioeconomic impact analysis of enzymatic recycling of poly(ethylene terephthalate). Joule 2021, 5, 2479–2503.
  18. Ellis, L.D.; Rorrer, N.A.; Sullivan, K.P.; Otto, M.; McGeehan, J.E.; Román-Leshkov, Y.; Wierckx, N.; Beckham, G.T. Chemical and biological catalysis for plastics recycling and upcycling. Nat. Catal. 2021, 4, 539–556.
  19. Jenkins, S.; Quer, A.M.I.; Fonseca, C.; Varrone, C. Microbial Degradation of Plastics: New Plastic Degraders, Mixed Cultures and Engineering Strategies. In Soil Microenvironment for Bioremediation and Polymer Production; Jamil, N., Kumar, P., Batool, R., Eds.; Wiley Online Book: Hoboken, NJ, USA, 2019; pp. 215–238. ISBN 9781119592129.
  20. Wierckx, N.; Prieto, M.A.; Pomposiello, P.; de Lorenzo, V.; O’Connor, K.; Blank, L.M. Plastic waste as a novel substrate for industrial biotechnology. Microb. Biotechnol. 2015, 8, 900–903.
  21. Arkatkar, A.; Arutchelvi, J.; Bhaduri, S.; Uppara, P.V.; Doble, M. Degradation of unpretreated and thermally pretreated polypropylene by soil consortia. Int. Biodeterior. Biodegrad. 2009, 63, 106–111.
  22. Yoon, M.G.; Jeon, H.J.; Kim, M.N. Biodegradation of Polyethylene by a Soil Bacterium and AlkB Cloned Recombinant Cell. J. Bioremediat. Biodegrad. 2012, 3, 1–8.
  23. Mohanan, N.; Montazer, Z.; Sharma, P.K.; Levin, D.B. Microbial and Enzymatic Degradation of Synthetic Plastics. Front. Microbiol. 2020, 11, 2837.
  24. Lomwongsopon, P.; Varrone, C. Contribution of Fermentation Technology to Building Blocks for Renewable Plastics. Fermentation 2022, 8, 47.
  25. Tiseo, I. PET Demand Worldwide 2030. Available online: https://www.statista.com/statistics/1128658/polyethylene-terephthalate-demand-worldwide/ (accessed on 6 October 2022).
  26. PlasticsEurope The Facts 2021. Available online: https://plasticseurope.org/knowledge-hub/plastics-the-facts-2021/ (accessed on 16 February 2022).
  27. Zero Waste Europe New Report: PET is Far from Real Circularity. Available online: https://www.recycling-magazine.com/2022/02/16/new-report-pet-is-far-from-real-circularity/ (accessed on 16 February 2022).
  28. Hou, Q.; Zhen, M.; Qian, H.; Nie, Y.; Bai, X.; Xia, T.; Rehman, M.L.U.; Li, Q.; Ju, M. Upcycling and catalytic degradation of plastic wastes. Cell Rep. Phys. Sci. 2021, 2, 100514.
  29. Chen, C.C.; Han, X.; Li, X.; Jiang, P.; Niu, D.; Ma, L.; Liu, W.; Li, S.; Qu, Y.; Hu, H.; et al. General features to enhance enzymatic activity of poly(ethylene terephthalate) hydrolysis. Nat. Catal. 2021, 4, 425–430.
  30. Brott, S.; Pfaff, L.; Schuricht, J.; Schwarz, J.N.; Böttcher, D.; Badenhorst, C.P.S.; Wei, R.; Bornscheuer, U.T. Engineering and evaluation of thermostable IsPETase variants for PET degradation. Eng. Life Sci. 2022, 22, 192–203.
  31. Cui, L.; Qiu, Y.; Liang, Y.; Du, C.; Dong, W.; Cheng, C.; He, B. Excretory expression of IsPETase in E. coli by an enhancer of signal peptides and enhanced PET hydrolysis. Int. J. Biol. Macromol. 2021, 188, 568–575.
  32. Ferrario, V.; Todea, A.; Wolansky, L.; Piovesan, N.; Guarneri, A.; Ribitsch, D.; Guebitz, G.M.; Gardossi, L.; Pellis, A. Effect of Binding Modules Fused to Cutinase on the Enzymatic Synthesis of Polyesters. Catalyst 2022, 12, 303.
  33. Knott, B.C.; Erickson, E.; Allen, M.D.; Gado, J.E.; Graham, R.; Kearns, F.L.; Pardo, I.; Topuzlu, E.; Anderson, J.J.; Austin, H.P.; et al. Characterization and engineering of a two-enzyme system for plastics depolymerization. Proc. Natl. Acad. Sci. USA 2020, 117, 25476–25485.
  34. Sulaiman, S.; Yamato, S.; Kanaya, E.; Kim, J.J.; Koga, Y.; Takano, K.; Kanaya, S. Isolation of a novel cutinase homolog with polyethylene terephthalate-degrading activity from leaf-branch compost by using a metagenomic approach. Appl. Environ. Microbiol. 2012, 78, 1556–1562.
  35. Wei, R.; Haugwitz, G.V.; Pfaff, L.; Mican, J.; Badenhorst, C.P.S.; Liu, W.; Weber, G.; Austin, H.P.; Bednar, D.; Damborsky, J.; et al. Mechanism-Based Design of Efficient PET Hydrolases. ACS Catal. 2022, 12, 3382–3396.
  36. Kenny, S.T.; Runic, J.N.; Kaminsky, W.; Woods, T.; Babu, R.P.; Keely, C.M.; Blau, W.; O’Connor, K.E. Up-cycling of PET (Polyethylene Terephthalate) to the biodegradable plastic PHA (Polyhydroxyalkanoate). Environ. Sci. Technol. 2008, 42, 7696–7701.
  37. Saratale, R.G.; Cho, S.K.; Saratale, G.D.; Kadam, A.A.; Ghodake, G.S.; Kumar, M.; Bharagava, R.N.; Kumar, G.; Kim, D.S.; Mulla, S.I.; et al. A comprehensive overview and recent advances on polyhydroxyalkanoates (PHA) production using various organic waste streams. Bioresour. Technol. 2021, 325, 124685.
  38. Choi, S.Y.; Cho, I.J.; Lee, Y.; Kim, Y.J.; Kim, K.J.; Lee, S.Y. Microbial Polyhydroxyalkanoates and Nonnatural Polyesters. Adv. Mater. 2020, 32, 1907138.
  39. Kenny, S.T.; Runic, J.N.; Kaminsky, W.; Woods, T.; Babu, R.P.; O’Connor, K.E. Development of a bioprocess to convert PET derived terephthalic acid and biodiesel derived glycerol to medium chain length polyhydroxyalkanoate. Appl. Microbiol. Biotechnol. 2012, 95, 623–633.
  40. Kim, H.T.; Kim, J.K.; Cha, H.G.; Kang, M.J.; Lee, H.S.; Khang, T.U.; Yun, E.J.; Lee, D.H.; Song, B.K.; Park, S.J.; et al. Biological Valorization of Poly(ethylene terephthalate) Monomers for Upcycling Waste PET. ACS Sustain. Chem. Eng. 2019, 7, 19396–19406.
  41. Sadler, J.C.; Wallace, S. Microbial synthesis of vanillin from waste poly(ethylene terephthalate). Green Chem. 2021, 23, 4665–4672.
  42. Kim, H.T.; Ryu, M.H.; Jung, Y.J.; Lim, S.; Song, H.M.; Park, J.; Hwang, S.Y.; Lee, H.S.; Yeon, Y.J.; Sung, B.H.; et al. Chemo-Biological Upcycling of Poly(ethylene terephthalate) to Multifunctional Coating Materials. ChemSusChem 2021, 14, 4251–4259.
  43. Kataoka, M.; Sasaki, M.; Hidalgo, A.R.G.D.; Nakano, M.; Shimizu, S. Glycolic acid production using ethylene glycol-oxidizing microorganisms. Biosci. Biotechnol. Biochem. 2001, 65, 2265–2270.
  44. Franden, M.A.; Jayakody, L.N.; Li, W.J.; Wagner, N.J.; Cleveland, N.S.; Michener, W.E.; Hauer, B.; Blank, L.M.; Wierckx, N.; Klebensberger, J.; et al. Engineering Pseudomonas putida KT2440 for efficient ethylene glycol utilization. Metab. Eng. 2018, 48, 197–207.
  45. Trifunović, D.; Schuchmann, K.; Müller, V. Ethylene glycol metabolism in the acetogen Acetobacterium woodii. J. Bacteriol. 2016, 198, 1058–1065.
  46. Werner, A.Z.; Clare, R.; Mand, T.D.; Pardo, I.; Ramirez, K.J.; Haugen, S.J.; Bratti, F.; Dexter, G.N.; Elmore, J.R.; Huenemann, J.D.; et al. Tandem chemical deconstruction and biological upcycling of poly(ethylene terephthalate) to β-ketoadipic acid by Pseudomonas putida KT2440. Metab. Eng. 2021, 67, 250–261.
  47. Kim, D.H.; Han, D.O.; Shim, K.I.; Kim, J.K.; Pelton, J.G.; Ryu, M.H.; Joo, J.C.; Han, J.W.; Kim, H.T.; Kim, K.H. One-Pot Chemo-bioprocess of PET Depolymerization and Recycling Enabled by a Biocompatible Catalyst, Betaine. ACS Catal. 2021, 11, 3996–4008.
  48. Utomo, R.N.C.; Li, W.J.; Tiso, T.; Eberlein, C.; Doeker, M.; Heipieper, H.J.; Jupke, A.; Wierckx, N.; Blank, L.M. Defined Microbial Mixed Culture for Utilization of Polyurethane Monomers. ACS Sustain. Chem. Eng. 2020, 8, 17466–17474.
  49. Mahajan, N.; Gupta, P. New insights into the microbial degradation of polyurethanes. RSC Adv. 2015, 5, 41839–41854.
  50. Bisceglie, F.D.; Quartinello, F.; Vielnascher, R.; Guebitz, G.M.; Pellis, A. Cutinase-Catalyzed Polyester-Polyurethane Degradation: Elucidation of the Hydrolysis Mechanism. Polymer 2022, 14, 411.
  51. Liang, C.; Gracida-Alvarez, U.R.; Gallant, E.T.; Gillis, P.A.; Marques, Y.A.; Abramo, G.P.; Hawkins, T.R.; Dunn, J.B. Material Flows of Polyurethane in the United States. Environ. Sci. Technol. 2021, 55, 14215–14224.
  52. Liu, J.; He, J.; Xue, R.; Xu, B.; Qian, X.; Xin, F.; Blank, L.M.; Zhou, J.; Wei, R.; Dong, W.; et al. Biodegradation and up-cycling of polyurethanes: Progress, challenges, and prospects. Biotechnol. Adv. 2021, 48, 107730.
  53. Europur The Netherlands: A World-Leader in Mattress Recycling. Available online: https://www.europur.org/news-events/item/70-the-netherlands-a-world-leader-in-mattress-recycling (accessed on 18 February 2022).
  54. Robinson, S. Recycling Polyols Back into Flexible Foam. Available online: https://www.utech-polyurethane.com/node/807346/printable/print (accessed on 18 February 2022).
  55. Cregut, M.; Bedas, M.; Durand, M.J.; Thouand, G. New insights into polyurethane biodegradation and realistic prospects for the development of a sustainable waste recycling process. Biotechnol. Adv. 2013, 31, 1634–1647.
  56. Jin, X.; Dong, J.; Guo, X.; Ding, M.; Bao, R.; Luo, Y. Current advances in polyurethane biodegradation. Polym. Int. 2022, 71, 1384–1392.
  57. Gewert, B.; Plassmann, M.M.; Macleod, M. Pathways for degradation of plastic polymers floating in the marine environment. Environ. Sci. Proc. Impacts 2015, 17, 1513–1521.
  58. Shah, Z.; Krumholz, L.; Aktas, D.F.; Hasan, F.; Khattak, M.; Shah, A.A. Degradation of polyester polyurethane by a newly isolated soil bacterium, Bacillus subtilis strain MZA-75. Biodegradation 2013, 24, 865–877.
  59. Shah, Z.; Hasan, F.; Krumholz, L.; Atkas, D.; Shah, A.A. Degradation of polyester polyurethane by newly isolated Pseudomonas aeruginosa strain MZA-85 and analysis of degradation products by GC-MS. Int. Biodeterior. Biodegrad. 2013, 77, 114–122.
  60. Shah, Z.; Gulzar, M.; Hasan, F.; Shah, A.A. Degradation of polyester polyurethane by an indigenously developed consortium of Pseudomonas and Bacillus species isolated from soil. Polym. Degrad. Stab. 2016, 134, 349–356.
  61. Vargas-Suárez, M.; Fernández-Cruz, V.; Loza-Tavera, H. Biodegradation of polyacrylic and polyester polyurethane coatings by enriched microbial communities. Appl. Microbiol. Biotechnol. 2019, 103, 3225–3236.
  62. Khan, S.; Nadir, S.; Shah, Z.U.; Shah, A.A.; Karunarathna, S.C.; Xu, J.; Khan, A.; Munir, S.; Hasan, F. Biodegradation of polyester polyurethane by Aspergillus tubingensis. Environ. Pollut. 2017, 225, 469–480.
  63. Magnin, A.; Hoornaert, L.; Pollet, E.; Laurichesse, S.; Phalip, V.; Avérous, L. Isolation and characterization of different promising fungi for biological waste management of polyurethanes. Microb. Biotechnol. 2019, 12, 544–555.
  64. Álvarez-Barragán, J.; Domínguez-Malfavón, L.; Vargas-Suárez, M.; González-Hernández, R.; Aguilar-Osorio, G.; Loza-Tavera, H. Biodegradative activities of selected environmental fungi on a polyester polyurethane varnish and polyether polyurethane foams. Appl. Environ. Microbiol. 2016, 82, 5225–5235.
  65. Zafar, U.; Nzeram, P.; Langarica-Fuentes, A.; Houlden, A.; Heyworth, A.; Saiani, A.; Robson, G.D. Biodegradation of polyester polyurethane during commercial composting and analysis of associated fungal communities. Bioresour. Technol. 2014, 158, 374–377.
  66. Magnin, A.; Pollet, E.; Perrin, R.; Ullmann, C.; Persillon, C.; Phalip, V.; Avérous, L. Enzymatic recycling of thermoplastic polyurethanes: Synergistic effect of an esterase and an amidase and recovery of building blocks. Waste Manag. 2019, 85, 141–150.
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