Biocatalytic Recycling of Polyethylene Terephthalate: From Conventional to Innovative Routes for Transforming Plastic and Textile Waste into Renewable Resources

Abstract

The rapid accumulation of plastic and textile waste, particularly polyethylene terephthalate (PET), has emerged as a global challenge for sustainable resource management. Conventional recycling methods, including mechanical and chemical routes, recover limited value and often degrade material quality while consuming substantial energy. Biocatalytic recycling, by contrast, offers a resource-efficient alternative that transforms post-consumer PET into high-purity monomers under mild and environmentally benign conditions. This review examines advances in enzymatic PET depolymerization, focusing on hydrolases such as cutinases, PETases, MHETases, and lipases. The discussion highlights enzyme engineering, reactor design, and process integration that improve kinetics, thermostability, and yield. From a resource perspective, biocatalytic recycling redefines PET waste as a renewable carbon feedstock capable of re-entering industrial cycles, thereby reducing reliance on virgin petrochemicals and mitigating greenhouse gas emissions. Ultimately, this review positions biocatalytic PET recycling as a cornerstone technology for achieving circularity and advancing global resource sustainability.

1. Introduction

Plastics are usually man-made polymers chemically synthesized, with a backbone composed entirely of carbon–carbon bonds. Most of their raw materials come from fossil fuels. They have become essential in modern life due to their versatility, durability, and low cost, making them vital across industries such as packaging, textiles, electronics, and healthcare [1,2,3]. Global plastic production is estimated to reach around 1.1 billion tons by 2050 [4]. Its widespread use is driven by advantages like ease of processing, lightweight nature, and exceptional barrier properties, especially in food and beverage packaging [5]. However, the rapid growth of plastic consumption has led to severe environmental challenges, including plastic waste accumulation, greenhouse gas emissions, and threats to marine and terrestrial ecosystems. The majority of plastic waste originates from single-use packaging, which accounts for approximately 50% of global plastic waste [6]. In 2015, it was estimated that emissions from oil extraction to refining reached 68 million tons of CO₂ equivalents (CO₂e) [7]. This calculation was based on the weighted average carbon intensity of energy production from 8966 active oil fields across 90 countries, with approximately 4% of crude oil allocated for plastic manufacturing. In addition to the production and manufacturing stages, greenhouse gas emissions are also generated during the extraction and transportation of raw materials, as well as from plastic waste management and the release of plastics into the environment [8]. Emissions from production facilities are typically influenced by factors such as the efficiency, design, and lifespan of the equipment used. Even after plastics are discarded, their impact on the global climate continues, with most environmental effects occurring after their useful life has ended [9]. Implementing effective plastic waste management practices is essential for addressing sustainability challenges and mitigating environmental problems.

Apart from plastic waste, textiles that enhance our daily lives with beauty and comfort also pose a significant challenge for solid waste management. Garments typically have an average lifespan of around 5 years, after which most post-consumer textile waste is discarded, amounting to an estimated 65–92 million metric tons annually worldwide [10]. In 2018, global textile production reached 105 million metric tons [11]. The amount of textile waste ending up in landfills or incinerators continues to rise significantly [12], with less than 1% of the materials used in clothing production being recycled into new garments [13]. Currently, recycling technologies face five main challenges: (i) the absence of commercially viable methods for recycling low-grade textile fractions, (ii) limited large-scale processes and expertise for separating fiber types in mixed blends and composite structures, (iii) high costs associated with recovery processes, (iv) a recycling market dominated mainly by low-quality materials and blends, and (v) expensive logistics combined with a lack of sufficient textile recycling facilities at both local and regional levels [14,15]. To address these challenges, it is essential to deconstruct and separate the different fiber components into purified streams, enabling more efficient recycling and material recovery.

Both textile and plastic waste contain PET as a raw material. PET is widely used because of its durability, lightweight nature, and versatility, making it a popular choice in industries like packaging, especially for beverage bottles, and textiles, notably for synthetic fibers such as polyester. Compared to polyvinyl chloride (PVC), PET is considered a safer material since PVC in plastic waste can emit harmful chlorinated compounds, such as gaseous hydrogen chloride and chlorine-containing dioxins, during recycling or thermal energy recovery processes [16]. With the rapid growth of the plastic bottle market, PET consumption has been rising significantly to meet increasing demand [17]. PET also makes up a significant portion of consumer textile fibers, accounting for 88% of global textile fiber production when combined with other dominant fibers [18]. The primary approach to ensuring the sustainability of PET production is through recycling. Recycled PET is mainly processed into fibers (72%), followed by bottles (10%), sheets (10%), tapes (5%), and other products (3%), using chemical and mechanical recycling methods [19]. However, if the proportion of recycled PET in the production system is not optimal, the recycling process can still have negative environmental impacts [20]. To address this issue, efforts such as process optimization and enhancing recycling systems are crucial for further reducing greenhouse gas emissions and improving overall environmental performance.

The recycling of PET, derived from either plastic or textile waste, can be carried out through mechanical, chemical, or biological processes, with the primary goal of transforming PET into economically reusable forms. Mechanical recycling is typically categorized into primary and secondary recycling, while chemical and biological recycling are generally classified as tertiary and quaternary recycling. Each method offers distinct advantages and limitations, and the choice of approach is dependent on specific user requirements and application goals. Mechanical recycling transforms post-consumer plastics and textiles into raw materials for manufacturing new plastic or textile products [21]. Mechanical recycling represents the most widely used approach for PET products. The process involves collecting and shredding post-consumer PET into small fragments, washing and removing contaminants, and then melting the material into PET resin [22,23]. The purified PET flakes are then remelted and molded into new products, such as bottles, fibers, and films. Mechanical recycling offers a cost-effective, energy-efficient approach that helps reduce greenhouse gas emissions and conserve natural resources [24]. It also reduces the amount of PET products sent to landfills or oceans, aiding the shift toward a circular economy. However, this approach faces challenges such as inconsistent quality and availability of collected PET, contamination issues, and material property degradation. In such situations, alternative methods such as chemical recycling might offer better solutions.

Chemical recycling is an emerging technology that depolymerizes PET products into their constituent monomers, which can then be repolymerized to produce new PET materials [25,26]. Chemical recycling addresses the limitations of mechanical recycling by converting PET waste into high-quality raw materials, including terephthalic acid (TPA) and monoethylene glycol (MEG). This approach can also lower greenhouse gas emissions and reduce energy consumption by up to 50% compared to the production of virgin PET [27]. However, chemical recycling demands specialized equipment and technical expertise, making the process both costly and complex. In addition, the process is more expensive and intricate than mechanical recycling, and significant challenges remain regarding its scalability and commercial application [28]. Therefore, biological recycling could serve as an alternative to mechanical and chemical approaches.

Global plastic and textile waste has raised concerns about resource depletion and environmental pollution. Among synthetic polymers, PET plays a key role in both sectors due to its use in packaging and fibers. This dual purpose makes PET not just a pollutant but also a valuable, recoverable carbon resource. Traditional mechanical and chemical recycling methods, though common, often yield lower-quality materials and consume substantial energy. Therefore, developing sustainable recycling strategies that preserve monomeric value and conserve resources is crucial. Biocatalytic recycling—using enzymes to break down PET under mild conditions—offers an environmentally friendly and resource-efficient solution.

Biorecycling, or enzymatic recycling, is a growing technique that uses enzymes to break down PET products into their basic monomers. Usually, biological recycling relies on either microorganisms or synthetic enzymatic reactions [29]. The biorecycling of PET employs enzymes as biological catalysts to depolymerize the polymer into its constituent monomers [30]. The process generally consists of several key steps: collection and sorting, size reduction, enzymatic depolymerization, purification, and repolymerization. During collection and sorting, PET waste is separated according to color, type, and quality. Size reduction then involves shredding the waste into smaller fragments to increase surface area and enhance enzymatic degradation. Enzymatic depolymerization yields a mixture of monomers and oligomers, which are subsequently separated and purified. Finally, the purified monomers undergo polymerization to produce new PET, which can be used to manufacture bottles, films, fibers, and other products [31]. This process offers several benefits over traditional recycling methods and has the potential to help build a more sustainable future.

From a resource management perspective, biocatalytic PET recycling is more than just a technological innovation—it is a strategic approach for conserving resources and promoting a circular economy. Global PET production exceeds 30 million metric tons annually, using large amounts of petroleum-based feedstocks and creating significant post-consumer waste. By enzymatically transforming discarded plastics and textiles back into high-purity monomers like TPA and MEG, biocatalytic processes directly close the resource loop, decreasing reliance on virgin fossil materials and reducing waste buildup. Compared to traditional mechanical and chemical methods, enzymatic recycling operates under milder conditions, reducing energy use and greenhouse gas emissions. This shift turns PET waste from an environmental problem into a secondary raw material source, a renewable carbon supply that can be reused in production. Understanding biocatalytic PET recycling in this light supports broader goals of sustainable resource use, waste valorization, and low-carbon material management.

This review discusses current technologies for biological recycling of PET, focusing on its conversion and reuse to reduce CO₂ emissions. Among the various PET recycling approaches, biological recycling offers an environmentally friendly solution, relying on biocatalytic reactions. Several strategies have been developed for biocatalytic PET recycling, including the use of microorganisms and enzymes. The review also covers the process of obtaining new biologically recycled materials from PET, as well as the potential implementation of biocatalytic recycling on a large industrial scale. A deeper understanding of these biocatalytic methods is essential for policymakers to identify the most effective strategies to address the significant challenge of carbon emissions and promote sustainable recycling practices.

2. Resources: Plastic and Textile Waste to PET

The environmental impact of the textile industry is rising as global demand for textiles grows. Interestingly, 99% of polyester, a common synthetic fiber made from petroleum, is produced from post-consumer PET bottles, which make up 54% of the global fiber market. PET, the third most common polymer, is widely used in packaging and beverage bottles because of its durability, transparency, and lightweight qualities [32,33]. Global manufacturing of PET is predicted to reach 34 billion metric tons by 2050. PET is produced via a condensation process that utilizes TPA and EG, both sourced from petroleum feedstocks [34]. Furthermore, PET fiber was used worldwide to produce filament yarns and staple fibers. PET staple fibers are primarily used in the production of cotton-PET blended textiles; however, recycling these blended fabrics poses challenges due to the complex segregation of the mixed ingredients. Consequently, these textiles are predominantly discarded in landfills, resulting in environmental issues [35].

Converting plastic waste into PET usually starts with collecting and separating post-consumer plastics. Since waste streams often contain mixed polymers, proper sorting is essential to remove contaminants such as PVC, which can greatly reduce product quality during processing [36]. After sorting, size reduction methods such as shredding or cutting are used to increase the surface area, making subsequent cleaning and processing easier. Washing then removes labels, dirt, and organic residues, resulting in clean PET flakes ready for reprocessing. These initial steps are crucial for producing high-quality recycled PET, as impurities can harm the mechanical and thermal properties of the final product.

Once prepared, PET flakes can be melted down again and shaped into pellets or granules, which serve as raw materials for making new products. In mechanical recycling, these pellets are directly used to produce bottles, fibers, or films. In contrast, during chemical recycling, PET can be broken down into its monomers, such as TPA and MEG. These monomers can then be rebuilt into virgin-quality PET [37]. Such conversion routes not only extend the material life cycle but also reduce reliance on fossil-based raw materials, reduce greenhouse gas emissions, and promote circular-economy goals by reintegrating plastic waste into the production chain.

In textiles, converting waste into PET involves both mechanical and chemical methods. Mechanical recycling typically involves processes such as shredding and carding, where textile waste, primarily polyester-rich fabrics, is broken down into fibers that can be respun into yarn [33]. The mechanical approach has limitations, especially for post-consumer waste with blended or low-quality fibers, which often produces weaker fibers that need to be mixed with virgin materials to regain strength and durability [9]. Despite these issues, mechanical recycling remains a cost-efficient method and is commonly used to create insulation, padding, or nonwoven products from textile waste.

On the other hand, chemical recycling provides a more efficient pathway to recover PET from textile waste. This method depolymerizes polyester fibers into monomers such as PTA and MEG, which can then be purified and repolymerized into high-quality PET. Techniques like glycolysis, hydrolysis, and enzymatic hydrolysis are frequently applied, and advanced approaches such as using ionic liquids or subcritical water treatment are being developed to handle mixed fibers (e.g., polyester–cotton blends) [38,39]. Compared to mechanical methods, chemical recycling can regenerate PET with properties comparable to virgin plastic, creating new fibers, packaging materials, or even textiles [11]. This makes chemical recycling a key strategy for closing the loop in the textile-to-PET circular economy.

After plastic and textile waste are converted into PET, the material becomes suitable for biocatalytic degradation through enzymatic processes. In this approach, specific enzymes break down the PET polymer chains into their monomeric building blocks, primarily TPA and MEG [40]. This enzymatic degradation is a sustainable alternative to conventional recycling methods, as it operates under mild conditions, reduces energy consumption, and minimizes secondary pollution. The resulting monomers can be purified and repolymerized to produce new PET with properties comparable to virgin material, thereby closing the loop in a circular economy framework. In some cases, biodegradation can even occur directly on plastic or textile waste without prior conversion [10,41].

3. PET Characteristics and Recycling Context

Understanding the physical and chemical properties of PET is fundamental to optimizing recycling performance. The polymer’s high crystallinity and chemical resistance confer durability during use but hinder degradability post-consumption. These characteristics define PET as both an engineering success and a recycling challenge. In resource terms, the same molecular stability that ensures material longevity also creates kinetic barriers to depolymerization. Recognizing this paradox underscores the necessity for advanced recycling methods capable of reclaiming PET’s intrinsic carbon value.

PET’s lightweight, strong, and flexible characteristics make it a crucial material in today’s society [40,42]. Post-consumer and post-industrial PET waste differ from virgin PET due to their sources and contamination levels. Post-industrial waste from production is clean and readily recyclable, but post-consumer waste from consumer items might be polluted and difficult to recycle. Post-consumer PET waste contains soil, polymers, glue, labels, and small-molecule pollutants from bottles, packaging, and other uses [43]. PET is a semicrystalline polymer. The degree of crystallinity is determined by numerous criteria, including the molecular weight distribution, molecular weight, and crystallization temperature [44]. The crystallization rate of PET within the temperature range of 150–180 °C is based on the degree of chain orientation, molecular weight, and the characteristics of the polymerization catalyst. It is utilized in the process of producing PET [26,45]. Crystallinity is produced by heating until the glass transition temperature (Tg) is attained, which is followed by molecular orientation [46]. In addition to that, the crystalline microstructures demonstrate considerable resistance to the process diffusion of enzyme and water, which serves as a key component as a catalyst and reactant to biocatalytic depolymerization, respectively [47,48].

The melting temperature (Tm) of commercial PET is between 255 and 265 °C, with more crystalline PET reaching 265 °C. Virgin PET’s Tg ranges from 67 to 140 °C [26]. Furthermore, the proximate and ultimate analyses of PET for volatile matter, moisture, and ash were 88.2%, 0.5%, and 11.2%, respectively. On the other hand, the percentages of the ultimate analysis, such as carbon, hydrogen, nitrogen, and oxygen, are 64.2, 4.4, 0, and 31.4 wt%, respectively [49]. Furthermore, rheological studies are limited by crystallization at low temperatures and polymer breakdown at higher temperatures. PET rheological tests are typically performed at temperatures ranging from 260 to 290–300 °C. Above the latter temperature, rapid degradation occurs, limiting the practicality of the experiments [44]. In addition, the surface morphology of crystalline PET was studied by Daggubati. It compares SEM images of virgin and recycled PET to demonstrate the similarities in surface morphology between the two. Surface roughness in recycled PET is shown as holes, flakes, cracks, and pitting [50].

Kinetic studies of solid-state PET processes provide essential knowledge that can be used. Kinetic analysis aims to precisely predict the thermal behavior of a process under multiple circumstances that differ from those utilized in the PET studies [51]. In addition, Dubdud investigated various methods to determine the activation energy of PET. For instance, the activation energies of PET using Starnik, FWO, KAS, and Friedman were reported as 139.2, 142.8, 138.8, and 167.5 kJ/mol, respectively [49]. Furthermore, the physical and chemical properties were reached by thermogravimetric analysis, and derivative thermogravimetric analysis demonstrates that the thermal decomposition started at 370 °C, with the temperature maximum decomposition up to 441 °C, and the end of the degradation process of PET is 502 °C, with a heating rate of 10 °C/min [52].

Furthermore, the comparison with mechanical, chemical, and biological recycling approaches to the recycling of PET is listed in Table 1. The mechanical process is conducted by several methods, such as separation process, crushing, heat, and extraction. The waste from PET bottles/containers is melted to produce recycled PET pellets. On the other hand, the chemical process of PET recycling uses chemical and/or thermochemical methods to degrade PET waste into its constituent monomers, oligomers, and a mixture of solid, liquid, and gaseous hydrocarbons [53]. Furthermore, Biocatalysts and microorganisms capable of degrading plastics have received significant attention from the scientific community and the media, with news outlets presenting “plastic-eating” microorganisms and enzymes as potential solutions to the worldwide plastic crisis [54]. Enzymes facilitate rapid and selective chemical transformations in biological catalysis; however, they often exhibit limited tolerance to extreme processing conditions and additives, as their activity depends heavily on maintaining protein structure stability [55].

Table 1. Comparison of various technologies for the recycling of PET.

Approach	Mechanical Recycling	Chemical Recycling	Biological Recycling	Ref
Technology recycling of PET	Injection molding Extrusion molding Inflation molding Blow molding Vacuum molding	- Hydrolysis - Methanolysis - Ammonolysis - Glycolysis - Aminolysis - Methanolysis–Hydrolysis - Glycolysis–Methanolysis - Steam hydrolysis	Various microorganism is used to recycle PET	[26]
Product	New PET bottle	Dimethyl terephthalate, Bis(2-hydroxyethyl) terephthalate, TPA, MEG, and oligomers	Dimethyl terephthalate, Bis(2-hydroxyethyl) terephthalate, TPA, MEG	[26]
Rate of degradation	High speed	High speed	Moderate	[56]
Separation process	Applicable	Complicated to separate the product	Applicable	[56]
Temperature/Pressure	High	High	Moderate	[26]
Greenhouse gas	Low	High	Moderate
Advantages	- Cost-effective	- High yield - Simple method - Raw materials flexibility	- Environmentally friendly - High yield	[26,57]
Drawbacks	- Cleaning PET waste is a necessary initial step to eliminate contaminants. - Poor mechanical stress of PET - The life cycle of recycled PET will affect its quality	- High production cost - High corrosiveness for some chemical recycling of PET - Less environmentally friendly - Hazardous volatile organic substances for by-products - High consumption of energy	- Long reaction time - Enzyme cost	[57,58]

Table 1. Comparison of various technologies for the recycling of PET.

Approach	Mechanical Recycling	Chemical Recycling	Biological Recycling	Ref
Technology recycling of PET	Injection molding Extrusion molding Inflation molding Blow molding Vacuum molding	- Hydrolysis - Methanolysis - Ammonolysis - Glycolysis - Aminolysis - Methanolysis–Hydrolysis - Glycolysis–Methanolysis - Steam hydrolysis	Various microorganism is used to recycle PET	[26]
Product	New PET bottle	Dimethyl terephthalate, Bis(2-hydroxyethyl) terephthalate, TPA, MEG, and oligomers	Dimethyl terephthalate, Bis(2-hydroxyethyl) terephthalate, TPA, MEG	[26]
Rate of degradation	High speed	High speed	Moderate	[56]
Separation process	Applicable	Complicated to separate the product	Applicable	[56]
Temperature/Pressure	High	High	Moderate	[26]
Greenhouse gas	Low	High	Moderate
Advantages	- Cost-effective	- High yield - Simple method - Raw materials flexibility	- Environmentally friendly - High yield	[26,57]
Drawbacks	- Cleaning PET waste is a necessary initial step to eliminate contaminants. - Poor mechanical stress of PET - The life cycle of recycled PET will affect its quality	- High production cost - High corrosiveness for some chemical recycling of PET - Less environmentally friendly - Hazardous volatile organic substances for by-products - High consumption of energy	- Long reaction time - Enzyme cost	[57,58]

PET recycling routes can be categorized into mechanical, chemical, and biocatalytic pathways. Mechanical recycling remains the most accessible yet offers limited resource efficiency, as material properties deteriorate after repeated processing. Chemical recycling restores monomer quality but consumes high energy and often requires toxic reagents. Biocatalytic recycling, in contrast, converts PET waste into reusable monomers under mild conditions, enabling circular resource recovery at lower environmental cost. This progression from mechanical to biocatalytic routes reflects an evolving strategy—from waste disposal to resource regeneration.

4. Classes of PET-Hydrolyzing Enzymes

In the beginning, polyester hydrolases were discovered using traditional microbiological methods, such as isolating, cultivating, and enriching microbial strains capable of hydrolyzing natural polymers [48]. The following research resulted in the identification of bacterial and fungal species, which included those from Thermobifida and Fusarium species. It is found to degrade the synthetic polyesters, such as PET [59]. Furthermore, biocatalytic chemical recycling has the potential to revolutionize the commercial recycling industry by preserving the valuable properties of waste materials while remaining energy-efficient [60]. Enzyme-based PET depolymerization is becoming more popular as a possible alternative to other chemical recycling methods. This is mainly because (a) it requires lower temperatures, which use less energy; (b) it avoids the need for potentially hazardous substances like highly polar solvents and strong acids; and (c) enzymes have an excellent selectivity for polymer ester linkages, which helps recover monomers and allows the technique to be used with mixed materials, thus eliminating the need for difficult plastic sorting [61].

PET hydrolase (PETase) hydrolyzes PET via a classical serine-hydrolase mechanism, as shown in Figure 1 [47]. PETase uses a Ser–His–Asp catalytic triad, in which the activated serine attacks the ester bond in PET to form a tetrahedral intermediate and then an acyl–enzyme intermediate, releasing one fragment of the polymer. Next, a water molecule is activated by the His–Asp pair and attacks the acyl–enzyme, generating a second tetrahedral intermediate that collapses to release the carboxylate product and regenerate free serine. By repeating this two-step acylation–deacylation cycle along the polymer chain, PETase progressively depolymerizes PET into soluble oligomers and monomers such as MHET, TPA, and EG.

Currently, two types of enzymes have been identified that act on PET: (1) enzymes that attack the polymer surface, increasing its surface hydrophilicity, but having little or no effect on the polymer shape. The next category may include enzymes like nonspecific esterases, serine proteases, lipases, and certain genuine PETases. (2) This category includes genuine PET hydrolases, which are often derived from thermostable cutinases, as well as thermostable enzymes from compost metagenome libraries and variations of genuine PETase. Some of these enzymes could be useful for bio-recycling because they break down the inner blocks of PET and alter its structure [62]. Furthermore, PET can be degraded with several enzymes, mostly hydrolases, including PET hydrolase (PETase—EC 3.1.1.101, carboxyl ester hydrolases, and EC 3.1.1.2, arylesterase), cutinases (EC 3.1.1.74), carboxylesterases (EC 3.1.1.1), and lipases (triacylglycerol lipase, EC 3.1.1.3) [63,64]. Cutinases have a shallow, open active site on their surface, allowing them to hydrolyze water-insoluble, hydrophobic polyesters. In contrast, lipases and esterases feature an active site within a tunnel-like structure, enabling vertical hydrolysis. Research has progressed from screening natural strains to engineering highly active enzymes with improved thermostability and substrate affinity. This shift marks a move from environmental observation to intentional design for resource recovery.

4.1. Cutinase

Cutin is the insoluble, waxy polymer that is an essential part of the cuticle, which serves as a structural element in plants. It contains 12-monounsaturated 9,10,18-trihydroxy C18 acids, dihydroxy palmitic acids, fatty acids, and 12-monounsaturated 18-hydroxy 9,10-epoxy C18 acids. It shows a lesser quantity of phenolic groups compared to its similar, suberin, which exists in potatoes [65]. The initial enzyme identified for the degradation of PET fibers was a cutinase derived from Thermobifida fusca, which also possesses significant industrial potential for PET enzymatic degradation [66]. Furthermore, Cutinases (EC 3.1.1.74) are the only enzymes known to break down the inner block of PET films; nevertheless, lipases, esterases, and cutinases can also modify their surface [64,67].

Nevertheless, Cutinases are serine hydrolases found in numerous species, for instance Fusarium oxysporum, Saccharomonospora viridis, Thermobifida alba, Thermobifida cellulosilytica, Fusarium solani pisi, Thermobifida fusca, Humicola insolens, Kineococcus radiotolerans, Moniliophthora roreri [68,69]. These enzymes can digest cutin, a fatty acid polyester that forms the main part of the plant cuticle. Enzymes like these have become essential biocatalysts in breaking down synthetic polymers such as PET [70]. Compared to other enzymes that hydrolyze PET, cutinases can effectively break the ester bonds in PET because of the structural similarities between cutin and PET. Table 2 shows various comparisons of the degradation of PET by cutinase. In addition, cutinases have a greater attraction for esters of short-chain to medium-chain fatty acids [71]. The study by Won et al. focused on a cutinase enzyme found in Rhodococcus strains from the Ross Sea. It demonstrated outstanding activity under alkaline conditions. Additionally, it is possible to slowly break down PET with MEG-terephthalate ester bonds [72].

4.2. Carboxylesterases

Carboxylesterases (EC 3.1.1. x) have become common enzymes that exist in several species, including some viruses [73]. Carboxylesterases are a broad distribution of hydrolases that catalyze the breaking of ester bonds in a wide range of substrates. They successfully hydrolyze a wide range of molecules with particular functional groups, such as carboxylic acid esters, amides, and thioesters [74,75]. It follows that the taxonomy and categorization of bacterial carboxylesterases have been continuously revised and updated. Hitch and Clavel created a new classification system for bacterial lipolytic enzymes, dividing them into 35 families and 11 authentic lipase subfamilies based on their sequences and conserved features. The updated classification method, which relates to sequences for each category, was used for the study and categorization of new target esterases [76].

Previous research described the depolymerization of PET by carboxylesterases derived from Bacillota, namely the p-nitrobenzyl esterase (BsEstB) from Bacillus subtilis and the esterase (Cbotu_EstA) from Clostridium botulinum [60]. These types of enzymes, which are greater than cutinases, have a temperature optimum of 40 °C and demonstrate a minimal degree of PET hydrolysis. Consequently, they are classified as PET surface-modifying enzymes as compared to PET hydrolases [31]. In addition, carboxylesterases from I. sakaiensis and Thermobifida fusca KW3, identified as MHETase and TFCa, respectively, were demonstrated to assist in PET depolymerization by IsPETase and T. fusca cutinase. When degradation is conducted at elevated temperatures, inhibition mediated by MHET is alleviated [77]. Furthermore, terephthalate esters are generally deconstructed using two hydrolytic processes by carboxylesterases Figure 2; (1) PET is initially converted to mono(2-hydroxyethyl) terephthalic acid (MHET) by an aromatic polyester breaking down hydrolase, for instance PET digesting enzyme—also known as PETase, (2) MHETase, a second enzyme, breaks down MHET into TPA and MEG [78]. The combination of microorganisms and enzymes can enhance the depolymerization process of PET microplastics under elevated temperature [79].

Table 2. Comparison of the depolymerization of PET through Cutinase.

Substrate	Type of Enzyme	Source	Condition Operations			PET Degradation, %	Ref
			T, °C	t, h	pH
PET package	LC-cutinase	Metagenome from leaf branch compost	70	24	8	≤25	[80]
Amorphous PET film	Variant of TfCut2	T. f usca KW3	60	24	8	≤25	[81]
Amorphized and micronized PET	LC-cutinase variant	Metagenome from leaf branch compost	72	10	9	90	[82]
PET film	Cut190**SS	Escherichia coli	70	48	8.6	10.1	[83]
Amorphous PET film	Thermobifida fusca cutinase TfCut2	B. subtilis strain RH 11496	70	96	8	50	[84]
Textile PET fibres	Cutinase ICCGDAQI	E. coli BL21 (DE3)	70	14	9	97	[85]
PET-GF	Variant of Cut190	Saccharomonospora viridis AHK 90	63	NA	±8	33.6 ± 3.0	[86]
PET-GF film	HiC (Novo)	Humicola insolens	70	96	6.5–9.5	97 ± 3	[87]
PET	ThcCut1-G63A/F210I/D205C /E254C/Q93G (ThcCut1-AICCG)	NA	70	96	NA	96.2	[68]
Post-industrial PET fibres	Cutinase ICCGDAQI	E. coli	70	24	9	97.8	[88]
Post-consumer PET bottles	Cutinase Est1_5M	Thermobifida alba AHK119	65	24–72	8	90.8	[89]
PET fabrics	Tfu_0883	Thermobififida fusca	60	48	7.5	50	[90]

** thermostable mutant of cutinase.

Table 2. Comparison of the depolymerization of PET through Cutinase.

Substrate	Type of Enzyme	Source	Condition Operations			PET Degradation, %	Ref
			T, °C	t, h	pH
PET package	LC-cutinase	Metagenome from leaf branch compost	70	24	8	≤25	[80]
Amorphous PET film	Variant of TfCut2	T. f usca KW3	60	24	8	≤25	[81]
Amorphized and micronized PET	LC-cutinase variant	Metagenome from leaf branch compost	72	10	9	90	[82]
PET film	Cut190**SS	Escherichia coli	70	48	8.6	10.1	[83]
Amorphous PET film	Thermobifida fusca cutinase TfCut2	B. subtilis strain RH 11496	70	96	8	50	[84]
Textile PET fibres	Cutinase ICCGDAQI	E. coli BL21 (DE3)	70	14	9	97	[85]
PET-GF	Variant of Cut190	Saccharomonospora viridis AHK 90	63	NA	±8	33.6 ± 3.0	[86]
PET-GF film	HiC (Novo)	Humicola insolens	70	96	6.5–9.5	97 ± 3	[87]
PET	ThcCut1-G63A/F210I/D205C /E254C/Q93G (ThcCut1-AICCG)	NA	70	96	NA	96.2	[68]
Post-industrial PET fibres	Cutinase ICCGDAQI	E. coli	70	24	9	97.8	[88]
Post-consumer PET bottles	Cutinase Est1_5M	Thermobifida alba AHK119	65	24–72	8	90.8	[89]
PET fabrics	Tfu_0883	Thermobififida fusca	60	48	7.5	50	[90]

** thermostable mutant of cutinase.

4.3. Lipases

Lipases are triacylglycerol acylhydrolases (E.C.3.1.1.3) that are classified within the hydrolases category. In addition to that, Extracellular triacylglycerol acyl hydrolases are known as lipases, which could be a trigger as a catalyst for the hydrolysis of long-chain triglycerides into glycerol and fatty acids [91]. Lipases demonstrate unique properties that make them especially interesting for synthetic applications, such as versatility, stability, recyclability, extensive tolerance to hydrophobic substrates, activity in non-aqueous environments, and the capacity to operate independently of cofactors [92].

In addition to that, Lipases are water-soluble enzymes due to a lid domain that blocks the hydrophobic active site. Numerous fungal species produce lipases and can depolymerize PET, such as Humicola sp., Thermomyces lanuginosus, Candida sp., Pseudomonas sp., and Thermobidifida fusca [91,93]. At the same time, lipase (TfH) was mainly purified and isolated from Thermobifida fusca. It is possible to hydrolyze PET films by 40–50% within three weeks. Meanwhile, lipase has been purified from Thermomyces lanuginosus, demonstrating its ability to hydrolyze PET [94,95,96]. Furthermore, Lipase B (CALB) derived from yeast Candida antarctica, previously classified as Pseudozyma antarctica, is recognized for its exceptional selectivity and catalytic efficacy. The function of CALB resembles the activities of IsPETase and IsMHETase [97]. The kinetic parameters of PET degradation through several enzymes are shown in Table 3. Swiderek et al. studied the process of hydrolysis of PET oligomers catalyzed by the promiscuous lipase B from Candida antarctica using molecular dynamics simulations validated by experimental Michaelis-Menten kinetics. The selectivity of the CALB substrate in relation to the pH value. The k_cat values for BHET hydrolysis remain similar regardless of the pH of the reaction. However, the k_cat values for MHET hydrolysis decrease as the pH increases. The k_M of CALB toward both substrates, BHET and MHET, follows the same trajectory with regard to substrate affinity: k_M (pH 9); (61.0 mM ± 25.7 mM, and 100 mM) > (pH 5) (22.5 mM ± 9.6 mM, and 23.8 mM ± 8.8 mM) > (pH 7) (13.3 mM ± 4.3 mM, and 15.1 mM ± 3.2 mM), respectively [98]. Furthermore, the kinetic study of various cutinases is shown in Table 3. Acero et al. studied the kinetic properties of the cutinases from Thermobifida. The Thermobifida cellulosilytica DSM44535 (Thc_Cut1 and Thc_Cut2) and Thermobifida fusca DSM44342 (Thf42_Cut1) enzymatically hydrolyze PET. The three different Thermobifida cutinases exhibit significant similarity, with a maximum of 18 amino acid discrepancies; nonetheless, they have distinct kinetic characteristics on soluble substrates. Their k_cat and K_m values for pNP–acetate ranged from 2.4 to 211.9 s⁻¹ and 127 to 200 μM, respectively, whereas for pNP–butyrate, k_cat and K_m values ranged from 5.3 to 195.1 s⁻¹ and 1483 to 2133 μM, respectively. Thc_Cut1 generated the greatest amount of TPA and MHET from PET [59].

Furthermore, the PET macromolecular polymers cannot directly penetrate microbial cells; hence, the first breakdown of PET into monomeric products predominantly depends on extracellular enzymes released by the host microorganisms. As seen in Figure 2, lipases can utilize a variety of enzymatic systems to biodegrade PET into smaller, water-soluble monomeric compounds, including MHET to become MEG, and TPA [99]. Moreover, using CALB demonstrated highly efficient hydrolysis processes and polymer depolymerization, resulting in the buildup of terephthalic acid. Additionally, CALB and Humicola insolens cutinase achieved full PET depolymerization, with a mole fraction reaching 0.88 and a 7.7-fold enhancement in PET yield, generating TPA [93,100]. Furthermore, the hydrolytic efficacy of lipases on PET can be determined of their substrate-binding pocket. This is due mainly to the lid structure that blocks the active site, as a result inhibiting enzymes from directly interacting with the macromolecular substrates [95,101]. Safdar et al. studied the biodepolymerization of PET by the extracellular lipase of Aspergillus niger—the synthesis of lipase from Aspergillus niger MG654699.1 using agro-industrial byproduct (wheat bran) by solid-state fermentation. The synthesized lipase exhibited an activity of 176.55 U/mL, with optimum conditions of 37 °C and pH 7.0. The 3.6% weight loss of PET was as a result of the biocatalytic activity of 30 KDa lipase [102].

Nevertheless, low temperatures will affect the lower specific volume of polymeric materials, raising the thermodynamic and kinetic barriers to geometrical change, resulting in limited substrate deformability [103,104]. Directed evolution and genome mining research investigations have discovered mutant PETases that demonstrate improved thermostabilities and catalytic efficiencies, such as PHL7, HOT-PETase, LCCICCG, LCCWCCG, and Fast-PETase [105]. Interestingly, in less than three days, one such designed PETase can degrade 90% of 20 kg of PET waste that has been processed using extrusion and micronization [106]. Furthermore, PETase serves as the enzyme responsible for the complex process of continually breaking down the PET polymeric chain into MHET, with small amounts of TPA and BHET as byproducts [107].

The kinetic data summarized in Table 3 reveal clear performance differences among the principal classes of PET-hydrolyzing enzymes. Cutinases and PETases exhibit the highest overall catalytic turnover and substrate affinity under typical experimental conditions (40–70 °C, pH 7–9), followed by MHETase, which complements the degradation pathway by converting the intermediate mono(2-hydroxyethyl) terephthalate (MHET) to terephthalic acid (TPA) and ethylene glycol (EG). Lipases and carboxylesterases, while capable of hydrolyzing short-chain model esters, show limited activity toward crystalline PET due to steric hindrance and restricted active-site access. These distinctions underscore that effective PET depolymerization relies on both enzyme selection and synergistic catalytic cascades.

The apparent catalytic constants (k_cat) in Table 3 vary by two orders of magnitude across enzyme classes. Cutinases such as TfCut2 and HiC exhibit k_cat values around 10–40 s⁻¹, while wild-type PETase typically shows 2–10 s⁻¹. Substrate crystallinity significantly affects apparent kinetic performance. Enzymes achieve nearly complete hydrolysis (>90%) for amorphous or pretreated PET films, whereas only 10–25% conversion is observed for highly crystalline materials under identical conditions. This discrepancy supports the hypothesis that chain mobility, not just surface adsorption, governs reaction kinetics. Pretreatment methods such as thermal amorphization, glycol-assisted swelling, or surface etching can reduce activation energy barriers and enhance k_cat/km ratios across enzyme classes.

Table 3. Comparison of the Kinetic Depolymerization of PET by Lipase and Cutinase.

Substrate	Crystallinity, %	Type of Enzyme	Source	Condition Operations			Kinetic Parameter *			Product	Ref
				T, °C	t, h	pH	kcat	km	R2
Post-consumer PET	41.1	Humicola insolens cutinase	Novozymes	70	96	7	224.2 ± 34.0 (h−1)	0.0041 ± 0.0010 (L g−1)	0.981	TPA + MHET + BHET	[108]
Bis(benzoyloxyethyl) terephthalate (3PET) with (p-nitrophenyl acetate (PNPA))	NA	Tha_Cut1	Escherichia coli BL21-Gold(DE3)	50	NA	7	2.72 ± 0.2 (h−1)	213 19 (μmol L−1)	NA	MHET, TPA, Benzoic acid, 2-hydroxyethyl benzoate	[109]
Bis(benzoyloxyethyl) terephthalate (3PET) with (p-nitrophenyl butyrate (PNPB))	NA	Tha_Cut1	E.coli BL21-Gold(DE3)	50	NA	7	6.03 ± 0.59 (s−1)	1933 ± 306 (μmol L−1)	NA	MHET, TPA, Benzoic acid, 2-hydroxyethyl benzoate	[109]
PET nanoparticles	9.8	TfCut2	Thermobifida fusca	60	1	8.5	147.673 ± 4.928 (min−1)	0.010 ± 7.40 × 10−4 (mLcm−2)	0.998	BHET, MHET	[110]
Amorphous PET	NA	Cut190**SS	E. coli	37	24	7	24.9 (s−1)	0.082 (mM)	NA	NA	[111]
PET with (p-nitrophenyl acetate (PNPA))	37	Thf42_Cut1	Thermobifida fusca DSM44342	50	120	7	39.5 ± 3.0 (s−1)	167 ± 29 (μmol L−1)	NA	BHET, MHET, TPA, Benzoic acid, 2-hydroxyethyl benzoate	[59]
PET with (p-nitrophenyl butyrate (PNPB))	37	Thf42_Cut1	Thermobifida fusca DSM44342	50	120	7	30.9 ± 8.6 (s−1)	2100 ± 361 (μmol L−1)	NA	BHET, MHET, TPA, Benzoic acid, 2-hydroxyethyl benzoate	[59]

* k_cat: specific activity, km: Michaelis constant. ** thermostable mutant of cutinase.

Table 3. Comparison of the Kinetic Depolymerization of PET by Lipase and Cutinase.

Substrate	Crystallinity, %	Type of Enzyme	Source	Condition Operations			Kinetic Parameter *			Product	Ref
				T, °C	t, h	pH	kcat	km	R2
Post-consumer PET	41.1	Humicola insolens cutinase	Novozymes	70	96	7	224.2 ± 34.0 (h−1)	0.0041 ± 0.0010 (L g−1)	0.981	TPA + MHET + BHET	[108]
Bis(benzoyloxyethyl) terephthalate (3PET) with (p-nitrophenyl acetate (PNPA))	NA	Tha_Cut1	Escherichia coli BL21-Gold(DE3)	50	NA	7	2.72 ± 0.2 (h−1)	213 19 (μmol L−1)	NA	MHET, TPA, Benzoic acid, 2-hydroxyethyl benzoate	[109]
Bis(benzoyloxyethyl) terephthalate (3PET) with (p-nitrophenyl butyrate (PNPB))	NA	Tha_Cut1	E.coli BL21-Gold(DE3)	50	NA	7	6.03 ± 0.59 (s−1)	1933 ± 306 (μmol L−1)	NA	MHET, TPA, Benzoic acid, 2-hydroxyethyl benzoate	[109]
PET nanoparticles	9.8	TfCut2	Thermobifida fusca	60	1	8.5	147.673 ± 4.928 (min−1)	0.010 ± 7.40 × 10−4 (mLcm−2)	0.998	BHET, MHET	[110]
Amorphous PET	NA	Cut190**SS	E. coli	37	24	7	24.9 (s−1)	0.082 (mM)	NA	NA	[111]
PET with (p-nitrophenyl acetate (PNPA))	37	Thf42_Cut1	Thermobifida fusca DSM44342	50	120	7	39.5 ± 3.0 (s−1)	167 ± 29 (μmol L−1)	NA	BHET, MHET, TPA, Benzoic acid, 2-hydroxyethyl benzoate	[59]
PET with (p-nitrophenyl butyrate (PNPB))	37	Thf42_Cut1	Thermobifida fusca DSM44342	50	120	7	30.9 ± 8.6 (s−1)	2100 ± 361 (μmol L−1)	NA	BHET, MHET, TPA, Benzoic acid, 2-hydroxyethyl benzoate	[59]

* k_cat: specific activity, km: Michaelis constant. ** thermostable mutant of cutinase.

4.4. PETase

PETase (EC 3.1.1.101), is an enzyme known as PET hydrolase (PETase), is capable of converting PET into its monomers [112]. PET hydrolases can degrade PET, thereby offering an environmentally favorable solution to the buildup of PET in the environment [113]. A unique structural characteristic of PETase is its atypically shallow and solvent-exposed active-site cleft, with more hydrophobic pockets found in thermophilic homologs, such as leaf-branch compost cutinase (LCC) and Thermobifida fusca cutinase (TfCut2) [114]. In addition, PETase’s structural flexibility, activity at ambient temperature, and unique active-site structure make it an effective biocatalyst for the degradation of PET. Nonetheless, its drawbacks, especially regarding thermostability and efficiency in high-crystallinity PET, have prompted significant technological efforts to realize its full potential for environmental and industrial applications [114,115]. Meanwhile, PETase is heat-sensitive and will be inactivated at temperatures greater than 40 °C. At mild temperatures of 25−30 °C, PETase may lose its degrading activity after more than 24 h of incubation. PETase has lost efficiency for industrial applications due to the need for more extended incubation periods and elevated reaction temperatures of about 50−65 °C to increase the degradation rate [116,117]. The mesophilic PET depolymerase moves the degradation polymer of the system into the physiological temperature range, demonstrating a feasible strategy for the free biological transformation of PET into any product that an organism can generate [118].

Two α/β-hydrolase fold enzymes, PETase and MHETase, collaborate to degrade PET in a two-step process through MHET, resulting in TPA and MEG, which are the essential precursors for a new cycle of PET production [119], as seen in Figure 3. Moreover, determining the kinetic parameters of PETase is inherently difficult due to the insolubility and heterogeneity of PET [114], as listed in Table 4. Burgin et al. studied the kinetic reaction mechanism of the PETase enzyme. The reaction rate constant is predicted to be k = 0.82 ± 0.10 s⁻¹. As well as the reaction rate constant, Erickson et al. utilized the Michaelis-Menten equation to determine that the k_cat values for amorphous PET film and crystalline PET powder were 1.5 ± 0.5 and 0.8 ± 0.0 s⁻¹, respectively, at 30 °C [120,121].

Table 4. Comparison of kinetic degradation of PET through PETase, IsPETase, and IsMHETase.

Raw Materials	Type of Enzymes	Organism	Condition Operations		Kinetic Parameter *			Product	Ref
			T, °C	t, h	kcat	km	k
Waste PET bottle	IsPETase	E. coli BL21	40	24	11.59 ± 0.16 s−1	0.19 ± 0.01 mmol.L−1	NA	TPA, BHET,	[122]
Waste PET bottle	IsPETaseW159H/F229Y	E. coli BL21	40	24	9.64 ± 0.13	0.08 ± 0.01	NA	TPA, BHET,	[122]
PET sheet	LCCICCG	Escherichia coli	50	4.1	NA	0.11 ± 0.02 µM	NA	NA	[123]
Post-consumer recycled PET flakes	LCCICCG	E. coli BL21	65	48	4.66 g/(µmol⋅h)	5.39 g/L	0.036 h−1	TPA, EG, 1.3 Propanediol, 1.4 Butanediol	[124]
Post-consumer PET bottle	ICCG-GS4-αSP	E. coli Shuffle T7 and E. coli BL21 (DE3)	60	24	3.91 ± 0.81 s−1	5.8 ± 1.2 g/L	NA	TPA, BHET, MHET	[125]
PET Film	PETase	Polyester hydrolase TfCut2	60	1	0.31 ± 0.01 s−1	7.33 × 10−3 ± 3.62 × 10−4 mol L−1	147.673 ± 4.928 min−1	MHET, BHET	[110]
Semi-crystalline PET powder	HiC [AAE13316.1] and	Humicola insolens and	50	5	0.088 s−1	0.27 g/L	NA	BETEB, BHET, MHET	[126]
Semi-crystalline PET powder	TfC [AAZ54921.1]	Thermobifida fusca	50	5	0.015 s−1	1.2 g/L	NA	BETEB, BHET, MHET	[126]

* k_cat: specific activity, km: Michaelis constant, k: enzyme reaction stability coefficient.

Table 4. Comparison of kinetic degradation of PET through PETase, IsPETase, and IsMHETase.

Raw Materials	Type of Enzymes	Organism	Condition Operations		Kinetic Parameter *			Product	Ref
			T, °C	t, h	kcat	km	k
Waste PET bottle	IsPETase	E. coli BL21	40	24	11.59 ± 0.16 s−1	0.19 ± 0.01 mmol.L−1	NA	TPA, BHET,	[122]
Waste PET bottle	IsPETaseW159H/F229Y	E. coli BL21	40	24	9.64 ± 0.13	0.08 ± 0.01	NA	TPA, BHET,	[122]
PET sheet	LCCICCG	Escherichia coli	50	4.1	NA	0.11 ± 0.02 µM	NA	NA	[123]
Post-consumer recycled PET flakes	LCCICCG	E. coli BL21	65	48	4.66 g/(µmol⋅h)	5.39 g/L	0.036 h−1	TPA, EG, 1.3 Propanediol, 1.4 Butanediol	[124]
Post-consumer PET bottle	ICCG-GS4-αSP	E. coli Shuffle T7 and E. coli BL21 (DE3)	60	24	3.91 ± 0.81 s−1	5.8 ± 1.2 g/L	NA	TPA, BHET, MHET	[125]
PET Film	PETase	Polyester hydrolase TfCut2	60	1	0.31 ± 0.01 s−1	7.33 × 10−3 ± 3.62 × 10−4 mol L−1	147.673 ± 4.928 min−1	MHET, BHET	[110]
Semi-crystalline PET powder	HiC [AAE13316.1] and	Humicola insolens and	50	5	0.088 s−1	0.27 g/L	NA	BETEB, BHET, MHET	[126]
Semi-crystalline PET powder	TfC [AAZ54921.1]	Thermobifida fusca	50	5	0.015 s−1	1.2 g/L	NA	BETEB, BHET, MHET	[126]

* k_cat: specific activity, km: Michaelis constant, k: enzyme reaction stability coefficient.

4.5. IsPETase

IsPETase is a desirable option for a biocatalyst in PET recycling due to its unique catalytic characteristics. IsPETase was initially discovered in a novel microbial consortium named Ideonella sakaiensis 201-F6, which develops on PET film, as reported by Yoshida et al. [106]. IsPETase is a functional monomer which has a molecular mass of 30.1 kDa, consisting of 290 amino acid residues [127]. Due to its ease of cultivation and high yield efficiency, Escherichia coli (E. coli) is the most widely utilized host for the synthesis of PET hydrolases (such as BhrPETase, IsPETase, and their derivatives) [128] as shown in Table 4. According to structural studies, IsPETase and other previously identified PET-hydrolyzing enzymes, such as Cut190 [129]. Furthermore, the structural analyses of IsPETase have demonstrated that the enzyme’s mechanism of action is distinctive, exhibiting more activity on PET films compared to other hydrolases/esterases, and it can effectively use extremely massive and hydrophobic polymers [130]. It is believed that the interesting structural variations within homologous cutinases and IsPETase are responsible for this enhanced activity. The structural analyses indicate that IsPETase is a member of the α/β-hydrolase superfamily, consisting of a canonical α/β-hydrolase fold of a central β-sheet surrounded by α-helices [131]. Currently, it was recognized mechanistic hypothesis follows to the conventional serine hydrolase mechanism, which consists of a Ser–His–Asp (Ser160–His237–Asp206), catalytic triad stabilized by a two-residue oxyanion hole formed by the backbones of residues Tyr87 and Met161 [132]. Nevertheless, the wild-type enzyme is limited in terms of thermostability. Biotransformations must be conducted at ambient temperatures much below the T_g of PET, which causes problems with depolymerization rates [133]. In addition, IsPETase promotes the degradation of PET into the primary product MHET and the byproducts TPA, MEG, and BHET as shown in Figure 3 [134]. On the other hand, IsMHET can be hydrolyzed to MHET to become MEG and TPA [106].

4.6. IsMHETase

IsMHETase obtained from the MHETase Gram-negative bacteria from I. sakaiensis. It successfully depolymerized polymeric chains, yielding ecologically harmless substances such as carbon dioxide and water [100]. It was discovered that the catalytic machinery of IsMHETase and IsPETase was identical, as shown in Figure 3. There are four basic steps for both IsMHETase and IsPETase. (i) Nucleophilic attack introduced by Ser-His-Asp, (ii) The cleavage of the C−O bond, (iii) The nucleophilic assault by water molecules, and (iv) Deacylation of IsPETase/IsMHETase [135]. Furthermore, phylogenetic studies had identified IsMHETase as a member of the tannase enzyme family, consisting of fungal and bacterial tannases, along with feruloyl esterases [136]. Duan et al. demonstrated the IsMHETase’s lack of activity towards pNP-aliphatic esters and conventional aromatic ester compounds that can be catalyzed by enzymes belonging to the tannase family [137]. In addition to that, the initial stage of degradation involves the depolymerization of PET film, yielding the primary product MHET along with byproducts BHET, TPA, and MEG, facilitated by the IsPETase enzyme. IsMHETase afterwards continues to the second phase of degradation by hydrolyzing MHET to generate TPA and MEG [138].

Overall, Table 4 illustrates that achieving high depolymerization efficiency depends on balancing three factors: enzyme thermostability, substrate morphology, and controlled reaction chemistry. Systems that align these parameters not only exhibit superior kinetic constants but also demonstrate improved resource efficiency, as higher conversion at moderate energy input translates directly into reduced greenhouse-gas emissions and lower process costs. These findings reinforce that rational enzyme engineering and optimized pretreatment are complementary levers for advancing scalable, low-carbon PET recycling. Each enzyme has unique catalytic mechanisms and operational ranges. Cutinases demonstrate broad substrate tolerance and high activity against amorphous PET, while PETases are optimized for mild-temperature conditions. MHETase works with PETase by preventing the buildup of intermediates. Lipases and esterases assist in hydrolysis but are less effective alone. Comparative studies show that engineered cutinases and PETases surpass natural versions in both turnover rate and stability. These insights guide future enzyme development for sustainable PET recycling.

5. The Implementation of Biocatalytic PET for Large-Scale Industrial Applications

5.1. Process Engineering and Reactor Design

The process of biocatalytic PET with larger-scale production begins with melt-extrusion-based amorphization. For efficient biocatalytic depolymerization, the heated polymer extrudates must be quickly quenched in cold water or another solvent to achieve a low crystallinity, ideally less than 15% [139]. Nevertheless, the commercial PET has a large crystallinity of up to 30%. Therefore, it is crucial to have the pre-treatment process, such as micronization and extrusion, to enhance the amorphous region [140]. Furthermore, the process flow diagram for biocatalytic PET for large–scale industrial use is shown in Figure 4. After that, the amorphized PET generally undergoes a mechanical size-reduction process, which might include ambient temperature crushing or cryogenic grinding [48]. Brizendine et al. studied the effect of particle size reduction of PET during biocatalytic depolymerization. The reduction in the particle size of PET enhanced the maximum reaction rate for high crystallinity of PET. In contrast, the maximum hydrolysis rate for low crystallinity cryomilled PET demonstrated no significant changes across different particle sizes for every single substrate. Nonetheless, it indicates that the reduction in particle size has minimal impact on the total amount of conversion. The low-crystallinity cryomilled film achieved a mass loss of 99% within 48 h, whereas the high-crystallinity of PET powder attained only 23.5% conversion over 144 h [141].

The commercial potential of biocatalytic PET recycling has been previously explored. A dual enzyme reactor, combining a polyester hydrolase and a carboxyl esterase, could efficiently hydrolyze amorphous PET films. By converting the hydrolysis products of polyester hydrolase, MHET and BHET, into TPA and EG, the carboxyl esterase increases the rate of PET depolymerization. Additionally, a polyester hydrolase can effectively break down PET films in an enzyme reactor, with an ultrafiltration membrane continuously removing the hydrolysis byproducts [84,142]. Furthermore, Lu et al. studied the depolymerization of PET by using a rotating packed bed reactor. A rotating packed bed reactor may offer superior mass transfer and diffusion efficacy. Nevertheless, the shear strength intolerance of the enzyme prevents the direct application of a rotating packed bed reactor for the biocatalytic depolymerization process. A PET particle depolymerization rate up to 95% was attained under optimal circumstances after 24 h, representing a 30% enhancement compared to the rate obtained in a stirred tank reactor [143].

In addition, immobilizing the biocatalyst offers well-proven advantages, for instance, simplified product/biocatalyst separation, continuous processing, and, in some cases, enhanced stability [144]. Therefore, using immobilized biocatalysts in two-liquid-phase systems is an exciting method to combine two innovative technologies. Immobilization can increase the biocatalyst’s stability in these systems by protecting it from the phase-reducing effects of the organic solvent. However, issues may arise from unwanted interactions between the solvent and the immobilization support material. The type of support material influences both the activity and stability of the biocatalyst, as well as the hydrophilicity-hydrophobicity balance of natural or synthetic gels. The choice of solvent affects how substrates and products are distributed between the biocatalyst and the liquid phases [145,146].

Using the evaporative crystallizer process, several methods can enhance the separation of biocatalytic PET, including the TPA crystallizer. In this process, the solid product is TPA, and the remaining liquid product will continue to the membrane unit to remove the by-products. An ultrafiltration membrane can maintain insoluble substrate components and the enzyme within the reactor’s recycling loop, while allowing soluble reaction products to permeate the membrane and be separated from the reaction mixture [147,148]. Ultrafiltration technologies have effectively reduced product inhibition in enzymatic hydrolysis operations by eliminating soluble products and sustaining high hydrolysis yields [149]. Furthermore, the process will proceed in the salt crystallizer to separate the sodium sulfate and MEG. Then, the MEG and other byproducts will be separated at the distillation column.

5.2. Techno-Economic and Environmental Considerations

When assessing the viability of PET recycling strategies, it is crucial to not only analyze catalytic or mechanistic performance but also evaluate process-level economics, energy use, greenhouse gas emissions, and life-cycle environmental impacts. This section offers a comparative analysis of mechanical, chemical, and biocatalytic recycling pathways from techno-economic and environmental perspectives, highlights major cost or energy factors, and discusses the challenges and opportunities for improving resource efficiency in each approach.

(i)

Mechanical Recycling: Lowest Energy & Cost but Quality Loss and Limited Cycles

Mechanical recycling of PET is currently the most mature and widely implemented pathway. Because it avoids the need to break chemical bonds, its energy consumption and capital cost tend to be lower than those of more intensive methods. Several life-cycle assessment (LCA) studies indicate that mechanical recycling often yields lower environmental burdens (e.g., global warming potential, energy demand) than virgin PET production or more aggressive chemical routes, particularly when the recycled product can substitute for virgin resin [150].

However, mechanical recycling has inherent limitations: the recycled PET often suffers from degradation in molecular weight, coloration, contamination, and limited cycles of reuse. Over multiple reuse cycles, material properties deteriorate, constraining their application in high-value uses. Moreover, mechanical recycling is sensitive to feedstock purity; mixed, colored, or contaminated PET streams often cannot be recycled reliably by mechanical means and may be diverted to lower-value uses or incineration [40].

From a cost perspective, mechanical recycling typically features lower capital expenditures (CAPEX) and operating expenditures (OPEX), but its profitability is constrained by the market value of the recycled resin and the costs of sorting, washing, and quality control. Some techno-economic analyses suggest that in favorable local energy and logistics conditions, mechanical recycling can deliver recycled PET at a price comparable to virgin resin [151]. Mechanical recycling is currently the most resource-efficient route in many contexts (especially for clean PET streams), but its limitations in feedstock flexibility and property retention mean it cannot fully absorb all PET waste, particularly low-quality or mixed waste streams.

(ii)

Chemical Recycling: Flexibility and Quality at Cost of Energy Intensity

Chemical recycling (also called feedstock recycling or depolymerization) encompasses a variety of processes such as glycolysis, methanolysis, hydrolysis, pyrolysis, and solvolysis that break PET down into monomeric or oligomeric intermediates for repolymerization or chemical valorization. Its key advantage is that the recovered monomers can approach virgin-equivalent purity, offering higher substitutability and broader reuse scopes than mechanically recycled PET [150].

However, the trade-off lies in energy consumption, reagent costs, and emissions associated with breaking strong ester bonds. For example, Uekert et al. (2023) report that glycolysis-based chemical recycling and mechanical recycling show the best economic performance across many scenarios, with up to 9–73% lower cost than some alternative routes, and environmental impact reductions of 7–88% relative to baseline waste treatments (depending on system boundaries and assumptions) [152]. Nevertheless, in many LCAs chemical recycling still underperforms compared to mechanical recycling unless heat integration, renewable energy inputs, and efficient system design are implemented [153].

A further complication arises from the capital and operational costs associated with reactors, separation, purification, and catalyst recycling. The costs of chemical reagents (e.g., glycol, methanol, acids or bases), heating, and product recovery (distillation, separation) often dominate the cost structure, especially for smaller-scale plants or fragmented feedstocks. Moreover, the environmental footprint is sensitive to the quality of the feedstock (e.g., presence of additives, dyes, contaminants) and the efficiency of purification [26,154].

In comparison to mechanical recycling, chemical recycling offers higher flexibility and product quality, but at the expense of higher energy and capital demands. To make chemical recycling competitive, innovations in catalyst efficiency, heat recovery, process intensification, and integration with waste heat or renewable energy sources are essential.

(iii)

Biocatalytic Recycling: Promising Resource Efficiency, But High Uncertainty & Scale Challenges

Biocatalytic (enzymatic) PET recycling seeks to combine the resource attributes of chemical recycling (monomer-level recovery) with milder operating conditions and potentially lower energy input. Techno-economic and life-cycle studies to date suggest substantial upside potential, but also significant uncertainties and scaling challenges.

From the techno-economic perspective, Singh et al. (2021) built a process model for enzymatic PET depolymerization and estimated a minimum selling price for recycled TPA of about USD 1.93/kg under base-case assumptions, assuming effective enzyme reuse and optimized downstream purification [155]. Their model suggests that with further process optimization, especially reducing energy demand in pretreatment and downstream separation, enzymatic recycling can become cost-competitive with virgin production. On the environmental side, Singh et al. also report potential reductions in supply-chain energy consumption of 69–83% and greenhouse gas emissions of 17–43% per kg of TPA, compared to virgin TPA production [155].

However, more recent LCA work reveals a more nuanced picture. Uekert et al. (2022) conducted a detailed LCA of enzymatic PET recycling. They found that, in many impact categories and under current process assumptions, enzymatic recycling still performs worse than virgin PET production and chemical recycling—primarily due to high energy and chemical inputs in pretreatment, enzyme production, and product recovery steps [156]. Their sensitivity analysis shows that improved yields, elimination of harsh pre-treatment steps, and optimized pH control are crucial to bring enzymatic recycling’s environmental footprint down to parity with conventional routes [156].

Key cost and energy drivers include feedstock preparation (shredding, amorphization, micronization), buffer control and pH maintenance, enzyme production and recovery, purification of monomers (crystallization, filtration, distillation), and utilities (heating, cooling, electricity). In many process models, pretreatment and separation contribute the largest fraction of the energy and cost burden, limiting net benefits [156].

Despite these challenges, enzymatic recycling has attractive potential in certain niches: for low-quality, mixed, or contaminated PET streams not suitable for mechanical recycling; for modular or decentralized plants; and in systems with access to low-carbon electricity. Ongoing enzyme engineering (e.g., more thermostable and active variants) and process integration (e.g., combining biocatalysis with mild chemical steps) may help push the economics and environmental metrics toward competitiveness.

6. Limitations of Biocatalytic PET

Despite rapid progress, the scalability of biocatalytic PET recycling remains constrained by three key factors: (i) crystallinity-induced mass transfer barriers, (ii) enzyme stability under prolonged operation, and (iii) product inhibition by intermediates. Addressing these challenges requires both molecular-level and reactor-level strategies.

Overcoming the limitations of PET crystallinity and mass transfer is essential for large-scale industrial applications [157]. Table 5 shows the parameters and constraints of several biocatalysts for degrading PET. Solid porous supports may affect the efficacy of PET degradation, mainly because most proteins live within porous structures. Consequently, only enzymes exposed on the surface will initially engage with solid PET until the breakdown products are sufficiently small to pass through the pores. Moreover, enzymes attached to the carrier surface may exhibit inadequate interaction with the macromolecular PET substrate because of rigidification effects, which may result in considerable rate limitations related to the enzymes utilized in soluble form [85,158]. Nevertheless, the enzymatic depolymerization of amorphous PET has been significantly enhanced as the temperature increases, especially when approaching temperatures near the Tg of PET. The range of temperatures for depolymerization is typically 60–80 °C, depending on the substrate condition [123].

Table 5. The parameters and limitations of several biocatalysts for degrading PET.

Biocatalyst	Parameters	Limitations	Ref
Cutinase	The range of optimal temperature and pH for biocatalytic depolymerization is 60–70 °C and 8.0–9.0, respectively.	The smooth hydrophobic surface of PET inhibits the binding of cutinase to PET, consequently limiting the rate of the PET degradation process.	[159]
Carboxylesterases	The optimal temperature and pH degradation of biocatalytic are 50–70 °C and 8.0–9.0, respectively.	For applications in industrial polyester and plastic recycling, the stability and hydrolytic activity of known natural esterases toward synthetic polyesters are often inadequate.	[139,160,161]
Lipases	The desired temperature is usually lower than the glass transition temperature (Tg) of PET, and the ideal pH for the biocatalytic activity of PET is 7.	The challenges to depolymerization of PET by lipases in biocatalytic systems are the consistent product inhibition by hydrolysis byproducts, such as MHET, and another limitation is that it is tough to degrade the crystalline structure of PET	[162,163]
PETase	The ideal temperature for PETase to degrade the PET depends on the kind of enzyme and its engineering, occurring within the temperature range of 40–80 °C, with the optimal pH around 8–9.	PETase is heat sensitive, and lower temperatures might reduce the specific volume of polymeric materials. Meanwhile, it potentially reduces its degrading activity at mild temperatures.	[160,164]
IsPETase	IsPETase works efficiently to degrade the PET at temperatures and pH levels up to 30–60 °C and 7.5–8.0.	IsPETase’s poor catalytic effectiveness and inherent lack of thermostability are two of its drawbacks for PET degradation.	[114,116]
IsMHETase	The ideal temperature for IsMHETase in biocatalytic PET breakdown varies from 50–70 °C, and the ideal pH is around 9.0.	Low catalytic efficiency, poor thermostability, possible product inhibition from degradation intermediates, and the intrinsic stability and crystallinity of the PET polymer itself are the drawbacks of biocatalytic PET depolymerization employing enzymes such as IsMHETase.	[47,160]

Table 5. The parameters and limitations of several biocatalysts for degrading PET.

Biocatalyst	Parameters	Limitations	Ref
Cutinase	The range of optimal temperature and pH for biocatalytic depolymerization is 60–70 °C and 8.0–9.0, respectively.	The smooth hydrophobic surface of PET inhibits the binding of cutinase to PET, consequently limiting the rate of the PET degradation process.	[159]
Carboxylesterases	The optimal temperature and pH degradation of biocatalytic are 50–70 °C and 8.0–9.0, respectively.	For applications in industrial polyester and plastic recycling, the stability and hydrolytic activity of known natural esterases toward synthetic polyesters are often inadequate.	[139,160,161]
Lipases	The desired temperature is usually lower than the glass transition temperature (Tg) of PET, and the ideal pH for the biocatalytic activity of PET is 7.	The challenges to depolymerization of PET by lipases in biocatalytic systems are the consistent product inhibition by hydrolysis byproducts, such as MHET, and another limitation is that it is tough to degrade the crystalline structure of PET	[162,163]
PETase	The ideal temperature for PETase to degrade the PET depends on the kind of enzyme and its engineering, occurring within the temperature range of 40–80 °C, with the optimal pH around 8–9.	PETase is heat sensitive, and lower temperatures might reduce the specific volume of polymeric materials. Meanwhile, it potentially reduces its degrading activity at mild temperatures.	[160,164]
IsPETase	IsPETase works efficiently to degrade the PET at temperatures and pH levels up to 30–60 °C and 7.5–8.0.	IsPETase’s poor catalytic effectiveness and inherent lack of thermostability are two of its drawbacks for PET degradation.	[114,116]
IsMHETase	The ideal temperature for IsMHETase in biocatalytic PET breakdown varies from 50–70 °C, and the ideal pH is around 9.0.	Low catalytic efficiency, poor thermostability, possible product inhibition from degradation intermediates, and the intrinsic stability and crystallinity of the PET polymer itself are the drawbacks of biocatalytic PET depolymerization employing enzymes such as IsMHETase.	[47,160]

Thomsen et al. studied the effect of substrate crystallinity and glass transition temperature on the biocatalytic activity of PET by utilizing DuraPETase, which is derived from Ideonella sakaiensis PETase, and LCC_ICCG, a variation of the leaf-branch compost cutinase. The degree of crystallinity (X_C) influenced the rate of enzymatic product release, which stopped successfully around X_C 22–27% for LCC_ICCG compared to the degree of crystallinity of approximately 17% for DuraPETase. The Tg of amorphous PET disks declined from 75 °C to 60 °C after 24 h of pre-soaking in water at 65 °C, whereas the X_C remained constant. The enzymatic reaction on pre-soaked disks at 68 °C, more than the Tg, did not influence the removal rate of the enzymatic product catalyzed by LCC_ICCG [165]. Consequently, high-temperature resistant enzymes are required to facilitate the effective hydrolysis of high-crystalline PET. The inhibition caused by MHET or intermediate metabolites during the catalytic reaction stage is an additional constraint in enzymatic PET biodegradation [166].

Furthermore, the release of by-products from biodegradation or hydrolysis processes increases the acidity of the reaction mixture. Consequently, the wild-type enzyme is inactivated, slowing down the reaction rate. Native wild-type enzymes cannot operate effectively due to these limitations [67,167]. There are some limitations of biocatalytic PET because the enzymatic depolymerization of PET, a heterogeneous catalytic process at the solid–liquid interface, is significantly affected by mass transfer limitations, including insoluble substrates, enzyme adsorption and desorption, and the released products, especially at large solid loadings [157], whereas the mass transfer in solid–liquid heterogeneous processes during PET degradation is hindered by the poor diffusion rate of enzyme molecules across the liquid-solid boundary barrier, which blocks the effective transport of enzymes to the solid surface [143,168]. To eliminate mass transfer limitations and difficulties with accessibility for macromolecular enzymatic substrates, the agitation may significantly reduce the diffusional mass transfer boundary layer, enhancing mass transfer in the hydrolysis unit [169]. Continuous flow reactors are often much smaller than batch reactors; nonetheless, under optimal circumstances and perfect configurations, these bench-scale reactors can provide a greater quantity of product within specific periods compared to their batch counterparts [170]. In addition to that, an additional benefit is the direct transfer of flow systems to large-scale production without substantial further optimization, which sharply contrasts with the upscaling of batch operations [171]. The conventional upscaling of (bio)catalysis in large batch reactors has undergone substantial transformation in recent decades, with downsizing and numbering-up of catalytic processes in continuous flow emerging as a dominant framework [170].

Despite impressive progress in engineering PET-degrading enzymes, several knowledge gaps and practical challenges still hinder their industrial deployment [114,172,173]. Enzyme cost and stability remain critical: most high-performance PET hydrolases still require relatively high loadings, carefully controlled buffers, and stabilizing additives, and there is little long-term data on continuous operation under harsh industrial conditions (mixed plastics, dyes, additives, surfactants, shear, and fluctuating temperatures). Product inhibition and substrate heterogeneity are also not fully resolved; intermediates such as MHET and PET oligomers can accumulate on particle surfaces and slow further hydrolysis, while real waste streams contain multilayer packaging, pigments, and fillers that limit enzyme accessibility and complicate integration with downstream purification. In addition, mass-transfer and scalability issues present major bottlenecks: because PET is a solid, often semi-crystalline material, achieving efficient enzyme–substrate contact at the ton scale requires optimized pretreatment (e.g., grinding, crystallinity reduction) and reactor design, yet techno-economic and reactor-engineering data for slurry or packed-bed systems at high solids loading remain sparse. Finally, systems-level questions—including enzyme recycling, cofactor-free operation, life-cycle performance relative to mechanical and chemical recycling, and compatibility with existing mixed-polymer recycling infrastructure—are only beginning to be addressed and will be crucial for translating PET degradase enzymes from promising laboratory tools into truly competitive large-scale biorecycling technologies.

Recent advances in biocatalytic PET recycling have targeted overcoming the inherent limitations of native PET hydrolases—such as low thermostability, limited activity on crystalline PET, and quick activity loss—through enzyme engineering, directed evolution, and structure-guided design. Structure-based mutations of Ideonella sakaiensis PETase and cutinase-like enzymes have produced variants like DuraPETase, HotPETase, FAST-PETase, and ThermoPETase, which demonstrate significant increases in melting temperature (up to approximately 30 °C) and tens to hundreds of times higher PET hydrolysis rates compared to their wild types, allowing efficient depolymerization at 50–70 °C where PET chains are more mobile [107,163,174]. Directed evolution and semi-rational mutagenesis—often guided by molecular dynamics, free-energy calculations, or machine-learning models—have been used to optimize flexible loops, binding residues, and second-shell positions, balancing substrate affinity and catalytic turnover while enhancing expression and long-term stability under process conditions [175]. Together, these strategies are transforming PET hydrolases from fragile lab curiosities into robust biocatalysts capable of near-complete depolymerization of post-consumer PET, bringing enzymatic PET recycling much closer to industrial feasibility.

Despite remarkable progress in enzymatic PET depolymerization, several bottlenecks continue to limit translation from laboratory studies to full-scale operations. Future research should focus on three dimensions: (i) Enzyme innovation, utilizing machine-learning-guided protein engineering to enhance thermostability and reduce inhibition; (ii) Process integration, through hybrid chemo-enzymatic and continuous-flow systems that minimize energy loss; and (iii) System-level resource governance, promoting regulatory and logistic mechanisms to incorporate enzymatic recycling into national circular economy frameworks. Collectively, these efforts can transform PET waste management from end-of-pipe treatment to proactive resource recovery. Furthermore, Biocatalytic recycling of PET offers an emerging technological possibility. Focused on overcoming existing manufacturing sector difficulties through enzyme engineering and process optimization.

7. Conclusions

From the standpoint of sustainable resource utilization, biocatalytic recycling represents a paradigm shift in the management of PET waste. Rather than treating discarded plastics and textiles as pollutants, enzymatic depolymerization enables their conversion into valuable monomers, TPA and MEG, that can serve as renewable raw materials for new production cycles. This resource-centric approach contributes directly to a circular economy, in which carbon embedded in polymeric materials is conserved and continually reused, lowering dependence on fossil-based feedstocks.

Current research demonstrates that biocatalytic processes can achieve high conversion efficiency under moderate temperatures and without harsh reagents, reducing both energy consumption and environmental footprint compared to traditional chemical routes. Enzyme engineering has advanced rapidly, yielding variants and thermostable cutinases with improved catalytic efficiency and resilience to crystalline PET structures. When integrated with optimized pre-treatment and continuous-flow reactor systems, these enzymes can make closed-loop PET recycling technically and economically viable at an industrial scale.

However, several limitations remain—particularly regarding enzyme stability, mass transfer barriers, product inhibition, and cost-effective recovery of biocatalysts. Addressing these challenges will require interdisciplinary strategies that combine computational enzyme design, immobilization technologies, and life-cycle assessment–based optimization to maximize resource efficiency. Future work should also emphasize policy and infrastructure mechanisms to incorporate enzymatic recycling into broader resource management frameworks. Biocatalytic PET recycling embodies the convergence of biotechnology and resource science. By transforming waste polymers into renewable carbon feedstocks, it not only mitigates environmental pollution but also advances the global transition toward sustainable material circularity and responsible resource stewardship.