Biodegradation of PET: Current Status and Application Aspects - ACS

Perspective Cite This: ACS Catal. 2019, 9, 4089−4105

pubs.acs.org/acscatalysis

Biodegradation of PET: Current Status and Application Aspects Ikuo Taniguchi,†,⊥ Shosuke Yoshida,‡,§,∥,# Kazumi Hiraga,‡,∇ Kenji Miyamoto,§ Yoshiharu Kimura,†,○ and Kohei Oda*,‡ †

Downloaded via UNIV OF CALIFORNIA SANTA BARBARA on April 8, 2019 at 22:49:15 (UTC). See https://pubs.acs.org/sharingguidelines for options on how to legitimately share published articles.

Department of Polymer Science, Faculty of Textile Science, Kyoto Institute of Technology, Matsugasaki, Sakyo-ku, Kyoto 606-8585, Japan ‡ Department of Applied Biology, Faculty of Textile Science, Kyoto Institute of Technology, Matsugasaki, Sakyo-ku, Kyoto 606-8585, Japan § Department of Biosciences and Informatics, Keio University, 3-14-1 Hiyoshi, Kohoku-ku, Yokohama, Kanagawa 223-8522, Japan ABSTRACT: Most petroleum-derived plastics, as exemplified by poly(ethylene terephthalate) (PET), are chemically inactive and highly resistant to microbial attack. The accumulation of plastic waste results in environmental pollution and threatens ecosystems, referred to as the “microplastic issue”. Recently, PET hydrolytic enzymes (PHEs) have been identified and we reported PET degradation by a microbial consortium and its bacterial resident, Ideonella sakaiensis. Bioremediation may thus provide an alternative solution to recycling plastic waste. The mechanism of PET degradation into benign monomers by PET hydrolase and mono(2-hydroxyethyl) terephthalic acid (MHET) hydrolase from I. sakaiensis has been elucidated; nevertheless, biodegradation may require additional development for commercialization owing to the low catalytic activity of these enzymes. Here, we introduce PET degrading microorganisms and the enzymes involved, along with the evolution of PHEs to address the issues that hamper microbial and enzymatic PET degradation. Potential applications of PET degradation are also discussed. KEYWORDS: poly(ethylene terephthalate) (PET), microbial consortium, Ideonella sakaiensis, PET hydrolase (PETase), mono(2-hydroxyethyl) terephthalic acid hydrolase (MHETase), PET hydrolytic enzyme (PHE)

1. INTRODUCTION

the plastic material demand in 2015, amounting to 18.8 million tons out of the total plastic production of 269 million tons.2 Most plastics, including PET, are difficult to degrade and tend to accumulate in the environment after use, causing serious environmental disruption, termed “plastic pollution”. In particular, the leakage of small pieces of waste plastics, called microplastics, into the oceans threatens marine life.3 A recent report from the Ellen MacArthur Foundation has warned that at least 8 million tons of plastics leak into oceans every year, adding to the current burden of 150 million tons.4 The economic impact of the wasted plastics also represents another major issue. As plastics are almost exclusively single use, 95% of plastic packaging material value is lost after a short first use.4 The total loss owing to this depreciation reaches up to 80−120 billion USD annually. To address these issues, various recycling programs have been launched in the past several decades, including recycling and reuse of the packaging plastic materials.

The development of synthetic plastics has effected a paradigm shift in products, packaging, and manufacturing. The low cost and usefulness of plastics have been well received in a wide variety of markets, such as packaging, building and construction, automotive, electrical and electronic, household, leisure and sports, and agriculture. Accordingly, the production amount of plastics increased by over 20-fold over the past half century, reaching 335 million tons in 2016.1 The majority of synthetic plastics are categorized as thermoplastics that can readily be molded by melt processing. Poly(ethylene terephthalate) (PET), developed by DuPont in the mid1940s, constitutes the most commonly manufactured thermoplastic because of its excellent mechanical and thermal properties. PET was first developed for use as a textile fiber (Dacron) and then applied to the fabrication of polymer films because of its transparent nature and good barrier and shatterproof properties. Subsequently, PET was processed into plastic bottles and jars by utilizing injection stretch blow molding methods. Patented in 1973, PET bottles and jars gained market acceptance and have been utilized worldwide as beverage containers. PET accounted for approximately 7% of © XXXX American Chemical Society

Received: December 29, 2018 Revised: March 12, 2019

4089

DOI: 10.1021/acscatal.8b05171 ACS Catal. 2019, 9, 4089−4105

Perspective

ACS Catalysis

Figure 1. Time course of PET collection, gross recycling, and utilization from 1995 to 2016 in the US. Source: The National Association for PET Container Resources Report on Postconsumer PET Container Recycling Activity in 2016.5

Table 1. Biochemically Characterized PET Hydrolytic Enzymes (PHEs) with Known Amino Acid Sequence enzyme

accession number

temp range (optimal temp) (°C)/substratea

organism

ref

PETase TfH

GAP38373.1 WP_011291330.1

Ideonella sakaiensis 201-F6 Thermobifida fusca DSM43793

20−45 (40)/ PET; 30/pNPB 30−60 (60)/PET; 20−70 (60)/pNPB

LCC FsC Thc_Cut1

AEV21261.1 1CEX ADV92526.1

30−80 (70)/PET; 30−80/pNPB 30−60 (50)/PET; 25−30/pNPB; 37/pNPP 50/PET; 25/pNPB

Thc_Cut2

ADV92527.1

50/PET; 25/pNPB

33

Thf42_Cut1 Tha_Cut1 Thh_Est

ADV92528.1 ADV92525.1 AFA45122.1

50/PET; 25/pNPB 50/PET; 25/pNPB 50/PET; 25/pNPB

33 34 35

Cut190 S226P/ R228S HiC BsEstB PET2

BAO42836.1 (wild type) 4OYY ADH43200.1 C3RYL0

uncultured bacterium Fusarium solani pisi Thermobifida cellulosilytica DSM44535 Thermobifida cellulosilytica DSM44535 Thermobifida fusca DSM44342 Thermobifida alba DSM43185 Thermobifida halotolerans DSM44931 Saccharomonospora viridis AHK190 Humicola insolens Bacillus subtilis 4P3-11 uncultured bacterium

17 14, 17, 18, 25, 26 17, 27−29 17, 18, 30−32 33

60−65/PET; 25−75 (65)/pNPB

36 32, 34 20 37

PET5 PET6

R4YKL9 UPI0003945E1F

Oleispira antarctica RB-8 Vibrio gazogenes

PET12 Tcur0390

A0A0G3BI90 CDN67546.1

Tcur1278

CDN67545.1

Polyangium brachysporum Thermomonospora curvata DSM 43183 Thermomonospora curvata DSM 43183

30−85 (80)/PET; 25/pNPB 40−45 (40)/PET; not described/pNPB 50/agar plate containing PET nanoparticles; 17−90 (60− 70)/pNPO 50/agar plate containing PET nanoparticles 50/agar plate containing PET nanoparticles; 17−90 (55)/ pNPO 50/agar plate containing PET nanoparticles 50/PET nanoparticles; 25−70 (55)/pNPB 50−60 (60)/PET nanoparticles; 25−70 (60)/pNPB

38

37 37 37 38

a Reported temperature range in which the enzymatic activity was detected against the indicated substrate. Abbreviations: pNPB, p-nitrophenol butyrate; pNPP, p-nitrophenol palmitate; pNPO, p-nitrophenol octanoate.

recycling market through recycling programs was only 795 kilotons in 2016, with a gross recycling rate of 28.4%. PET that was not recycled has been mostly disposed of by incineration, which includes thermal recycling for energy recovery at the expense of emitting a large amount of CO2.5 In particular, reclaimers regenerated approximately 562 kilotons of clean flake and pellet from collected PET bottles for reuse, from which the utilization rate was determined to be 20.1%. The recycled PET (RPET) was reprocessed in 2016 into fiber (43%), sheet and film (19%), strapping (8%), food

In the United States, the PET production amount has also grown with market demand in household goods and health and beauty sectors as well as in the bottled drink sector. As PET is highly resistant to natural degradation, recycling of PET has been encouraged. A report issued by the National Association for PET Container Resources relating to postconsumer PET recycling5 indicated that the total weight of PET bottles and jars available for recycling or on shelves reached 2.8 million tons in 2016. However, the amount of postconsumer PET bottles collected for recycling in the USA and sold to the 4090

DOI: 10.1021/acscatal.8b05171 ACS Catal. 2019, 9, 4089−4105

Perspective

ACS Catalysis

Figure 2. Unrooted phylogenetic tree of known PET hydrolytic enzymes (PHEs) described in detail in Table 1. Multiple amino acid sequences were aligned using ClustalW, and the guide tree was obtained on the basis of the neighbor-joining method using the p-distance model. The organism source of each protein is shown at the leaves. Bootstrap values are shown at the branch points. Scale bar: 0.1 amino acid substitution per single site. Bacterial PHEs are classified into Type I, Type IIa, and Type IIb on the basis of the phylogenetic analysis conducted by Joo et al.51

enzymatic action.10 In comparison to the C−C bonds, ester bonds in the polymer backbone are usually more susceptible to biodegradation.8 However, PET contains a high ratio of aromatic terephthalate units that reduces the chain mobility, resulting in extremely low hydrolyzability of the backbone ester linkages.6,10 Moreover, the majority of petroleum-based plastics, including PET, are semicrystalline, consisting of both crystalline and amorphous domains. Because the enzymes are generally able to attack the flexible amorphous domain, the biodegradation rate of plastics decreases with increasing crystallinity.6,11,12 Previous efforts have therefore focused on identifying hydrolases to cleave the ester linkages in the amorphous domain of PET.13,14 Microbes that can directly degrade PET have also been sought on the basis of the same concept. To date, numerous PET hydrolytic enzymes (PHEs) have been identified.15−17 Table 1 summarizes the biochemically characterized PHEs. The majority of this type of enzyme comprises cutinases, which are able to hydrolyze cutin, an insoluble aliphatic polyester excreted from the plant cuticle. Notably, the substrate specificity of cutinases is broad; these enzymes exhibit hydrolytic activities for both insoluble triglycerides (typical substrates for lipases) and soluble esters (substrates for esterases). Although several lipases are also able to hydrolyze PET, their activity is low.15,18,19 Esterases usually

and beverage bottles (25%), nonfood bottles (4%), and other uses (1%).5 Figure 1 depicts the transition of PET production, gross recycling rate, and utilization rate over the past two decades. Although the amount of RPET has increased concomitant with the escalation of virgin PET production, the gross recycling and utilization rates have remained at approximately 30% and 20% in recent years, respectively.5 In addition, the low cost of virgin PET affects the recycling activity. Thus, the research and development of economic, effective, and environmentally friendly measures to manage waste PET have been actively pursued. Other commodity plastics such as polyethylene (PE), polypropylene (PP), poly(vinyl chloride) (PVC), and polystyrene (PS) are petrochemical products, with extremely low biodegradability in the natural environment, which is similar to that for PET.6,7 Many factors restrict microbial (enzymatic) attack against these plastics: e.g., minimally reactive functional groups in the backbone, chain mobility, crystallinity, and surface hydrophobicity.6,8,9 In particular, the highly stable carbon−carbon (C−C) bonds of the polymer backbones along with the lack of enzymes that can directly cleave the C−C bonds in nature hamper the biodegradation of these olefinic polymers. Therefore, abiotic endotype activities such as ultraviolet (UV) irradiation and oxidation to form carbonyl groups in the polymer are required to aid the 4091

DOI: 10.1021/acscatal.8b05171 ACS Catal. 2019, 9, 4089−4105

Perspective

ACS Catalysis

microbial growth dependent on PET as a carbon source have not yet been reported. Having searched for such PET-degrading microorganisms for over a decade, we succeeded in isolating a microbial consortium No. 46 that degraded PET completely and assimilated the degradates into CO2 and water.17 In the early stage of research, we considered that the (1) crystallinity of PET (for efficient degradation), (2) shape of the PET specimen (film rather than fiber for stable attachment of cells), (3) isolation source, (4) culture medium, and (5) cultivation conditions (e.g., temperature, shaking speed, and pH) would be important for the isolation of PET-degrading microorganisms expressing superior PET-degrading enzymes. We used low-crystallinity PET films (crystallinity 1.9%, ca. 60 mg, 20 × 15 × 0.2 mm, Mw = 45 × 103, Mw/Mn = 1.9, Tg = 77 °C, Tm = 255 °C, density 1.3378 g/cm3) as the major carbon source for screening. We screened over 250 PET-debriscontaminated environmental samples including sediment, soil, wastewater, and activated sludge from a PET bottle recycling site. We isolated microbial consortium No. 46 from one of the sediment samples collected at Sakai-city, Osaka, Japan. The microbial consortium No. 46 adhered onto PET film during cultivation (Figure 3A), leading to drastic changes in the morphology of the PET film (Figure 3A, upper inset). Microscopic observation revealed that consortium No. 46 was composed of bacteria, yeast-like cells, and protozoa (Figure 3A, lower inset). The consortium degraded PET film at a rate of 0.13 mg/(cm2 day) at 30 °C, with 75% of the degraded PET film carbon being catabolized into CO2 at 28 °C.17 Thus, we succeeded in isolation of a microbial consortium (No. 46) that could degrade amorphous PET completely at ambient temperature. The stable adherence of microorganisms during shaking cultivation was confirmed. The PET degradation could be readily visualized as whitening of the film surface and/or decay of the PET film, such that lowcrystallinity PET film was effective for high-throughput screening. We confirmed that consortium No. 46 did not lose its PET degradation activity for at least 10 weeks. Furthermore, we were able to freeze-stock No. 46 and restore it for subsequent cultivation without loss of PET degradation activity.17 Therefore, the PET degradation capacity of No. 46 could be maintained and reproduced.

hydrolyze esters with a shorter chain aliphatic region in comparison to those hydrolyzed by lipases. In particular, only a few esterases such as p-nitrobenzylesterase from Bacillus subtilis (BsEstB) are known as PHEs20 (Table 1 and Figure 2). In contrast to the recent progress on PHEs, only a few reports exist regarding biological PET degradation by specific microorganisms. Known microorganisms with such degradative activity comprise the filamentous fungi Fusarium oxysporum and Fusarium solani, which can grow on a mineral medium containing PET yarns.21,22 However, the enzymatic system that enables these microorganisms to hydrolyze PET has not yet been well elucidated. For a potential solution toward these issues, we have developed the following three systems to degrade PET: (1) microbial consortium No. 46; (2) I. sakaiensis 201-F6, and (3) a system consisting of two novel enzymes.17,23,24 It should be emphasized that the degradation products, terephthalate (TPA) and ethylene glycol (EG), are environmentally benign monomers (Chart 1). All of these systems are applicable for Chart 1. Chemical Structures of PET, Its Analogue Polyethylene-2,5-furandicarboxylate (PEF), and Their Moietiesa

a

For PEF and 2,5-furandicarboxylate, please see section 8.7.

3. IDEONELLA SAKAIENSIS FROM CONSORTIUM NO. 46 CAN USE PET AS ITS MAJOR ENERGY AND CARBON SOURCE We found that one of the consortium No. 46 subcultures had lost its PET-degrading activity. Denaturing gradient gel electrophoresis (DGGE) analysis showed that several 16S rDNA bands disappeared after the activity loss of No. 46. To isolate the microorganism corresponding to the missing bands, limiting dilutions of No. 46 were performed. Each diluted sample was cultured with PET film to concentrate the PETlytic microorganisms. PET degradation activity was estimated from the change in film transparency consequent to the degradation. To monitor microbial members in the culture, the 16S rDNA V3−V5 region was amplified by PCR from genomic DNA extracted from the culture and analyzed by DGGE. Subsequently, we found a culture displaying a single band in the PCR-DGGE, which was then spread on a 0.5% soft agar plate containing minimal medium to obtain a single colony. As the isolated strain represented a new species of the genus Ideonella, it was subsequently named Ideonella sakaiensis 201-

biorecycling of waste PET. We hope that these applications may prove useful in environmental remediation and lead to the creation of a more sustainable society. With this perspective, in the following sections we discuss the current status of these systems for PET biodegradation and their application aspects.

2. MICROBIAL CONSORTIUM NO. 46, WHICH DEGRADES AND ASSIMILATES AMORPHOUS PET INTO CO2 AND WATER For screening of PET-degrading microorganisms, a reliable and high-throughput method for evaluating the PET degradability is needed; however, only a few such methods have been reported.39,40 Although cutinase, lipase, and esterase have been previously reported as PHEs, these enzymes can only accomplish limited degradation of PET.9,14,18,20,27−38,40−47 Thus, PET-degrading microorganisms producing superior PET-specific degrading enzymes that are able to support 4092

DOI: 10.1021/acscatal.8b05171 ACS Catal. 2019, 9, 4089−4105

Perspective

ACS Catalysis

Figure 3. Microbial PET degradation.17 (A) Growth of microbial consortium No. 46 on PET film in a test tube after 20 days. SEM image: the PET film degraded by No. 46 after 70 days. Light microscopy image: bacteria, yeast-like cells, and protozoa found in No. 46. (B) I. sakaiensis cell grown on PET film after 60 h. The SEM image of a degraded PET film surface by I. sakaiensis is given on the right, and the proposed PET degradation pathway is given at the bottom. Several images are reproduced with permission from Yoshida et al.17 Copyright American Association for the Advancement of Science, 2016.

F6.48 Notably, I. sakaiensis 201-F6 exhibited the unusual ability to degrade PET as a major carbon and energy source for its growth, although it could not utilize glucose. I. sakaiensis abolished PET film for the growth, with the effect being much greater in minimal medium including PET film as a carbon source in comparison to that on control medium without PET. This comparison clearly showed that I. sakaiensis degrades and assimilates PET. Conversely, in the culture fluid, we detected almost no PET hydrolysis products. These results indicated that, under aerobic conditions, I. sakaiensis degrades and

assimilates PET to produce CO2 as the complete oxidation product of PET. We thus successfully isolated a bacterium capable of degrading and assimilating PET (Figure 3B), representing, to our knowledge, the first report of a microorganism that completely degrades PET. I. sakaiensis 201-F6 cells adhered on PET film during growth and were connected to each other by appendages (Figure 3B). Numerous degradation traces were observed on PET film by SEM. The strain could degrade PET about twice as fast as the source microbial consortium No. 46. It was considered that the 4093

DOI: 10.1021/acscatal.8b05171 ACS Catal. 2019, 9, 4089−4105

Perspective

ACS Catalysis Table 2. PETase Site-Directed Mutants with Improved PET Degradation Activity mutation

noted substrate (PETa used in reaction)

Y87A

drinking bottle commercial PET film PET film

R90A L117F W159A

biaxially oriented PET film biaxially oriented PET film drinking bottle commercial PET film PET film

W159H

drinking bottle commercial PET film PET film

I208F A209I S214H R280A

biaxially oriented PET film drinking bottle drinking bottle commercial PET film

amplification of PET degradation activityb 3.1 0.4 (18 h), 0.5 (36 h) 0.1 (TPA production), 0.8 (MHET production) 1.4 2.1 1.3 0.2 (18 h), 0.2 (36 h) 0.1 (TPA production), 0.2 (MHET production) 2.4 0.5 (18 h), 0.6 (36 h) 0.1 (TPA production), 0.1 (MHET production) 2.5 1.3 1.9 1.2 (18 h), 1.3 (36 h)

assay conditions (temp, reaction time, pH)/activity detection

ref

30 °C, pH 9, 48 h/hydrolysis 30 °C, pH 9, 18 or 36 h/hydrolysis 30 °C, pH 9, 42 h/hydrolysis

53 51 50

30 30 30 30 30

°C, °C, °C, °C, °C,

8.5, 48 h/weight loss 8.5, 48 h/weight loss 9, 48 h/hydrolysis 9, 18 or 36 h/hydrolysis 9, 42 h/hydrolysis

55 55 53 51 50

30 °C, pH 9, 48 h/hydrolysis 30 °C, pH 9, 18 or 36 h/hydrolysis 30 °C, pH 9, 42 h/hydrolysis

53 51 50

30 30 30 30

°C, °C, °C, °C,

pH pH pH pH pH

pH pH pH pH

8.5, 48 h/weight loss 9, 48 h/hydrolysis 9, 48 h/hydrolysis 9, 18 or 36 h/hydrolysis

55 53 53 51

The crystallinities of substrate PET are estimated as follows: >30% for drinking bottles; 35% for biaxially oriented PET film;16 beyond our estimation from the noted “commercial PET film” and “PET film” in the corresponding papers. bRatio of value of the PET degradation activity of the PETase mutant to that of the wild type. a

around 75 °C), the chain mobility increases, resulting in more rapid enzymatic hydrolysis of this plastic.15,32 Accordingly, active PHEs that retain activity at over 75 °C, such as the thermostable LCC,28 T. fusca,26 H. insolens,32 PET2,37 and PET637 (Table 1), might be most useful in industrial applications such as surface modification of PET fibers in the textile industry19 and the breakdown process of PET polymers during chemical recycling.10 Recent studies determining the three-dimensional (3D) structures of PETase50−54 (section 5) have opened new prospects toward the rational redesign of this protein. Sitedirected mutagenesis studies of PETase have been carried out to uncover residues important for its catalysis and/or to overcome the low catalytic activities. All successful mutations that increase the activity are found in the PET-binding groove or close to this region (Table 2). Ma et al.55 mutated residues around the PET binding groove, which led to the creation of three mutants demonstrating improved PET (biaxially oriented) degradation. The mutant I208F (I179F in the original manuscript), which exhibits higher affinity to PET owing to enhanced hydrophobic interactions, showed the highest PET degradability, affording a 2.5-fold increase in comparison to wild-type PETase. Liu et al.53 also succeeded in creating several mutants with higher hydrolytic activity toward PET bottles. Mutating Trp159 (Trp130 in the original manuscript), located in the substrate binding pocket, to either alanine or histidine was effective in increasing the activity. Substitution of the Tyr87 residue (Tyr58 in the original paper, an oxyanion hole-forming residue17,50) with alanine showed 3.1-fold higher activity. However, previous analysis of these three mutants by other groups reported negative effects.50,51 As the assay conditions were similar among these studies, it is possible that differences such as the degree of crystallinity of PET used might have affected the results. A second enzyme identified from I. sakaiensis, which belongs to the tannase family,56 hydrolyzes MHET, the main PET hydrolysis product by PETase, into TPA and EG with a catalytic efficiency (kcat/Km) of 4200 ± 370 s−1 mM−1.17 This

appendages might assist in the delivery of PET-degrading enzymes onto the film.

4. IDENTIFICATION OF PET-DEGRADING ENZYMES IN I. SAKAIENSIS The genome sequence of I. sakaiensis revealed the presence of a single open reading frame that shares 51% amino acid sequence identity with a hydrolase from Thermobifida fusca (TfH), which exhibits PET-hydrolytic activity.14 The recombinant protein produced craterlike pitting on the film surface and released PET hydrolysis products into the aqueous medium. A comparison of the hydrolytic activities toward aliphatic esters and PET among this enzyme and the known selected PHEs: TfH, cutinase homologue from leaf-branch compost metagenome (LC cutinase, or LCC),27 and F. solani cutinase (FsC)31 from a fungus illuminated the highest catalytic preference toward PET of the I. sakaiensis enzyme. This led to assignment of this enzyme (namely, PET hydrolase or PETase) to the new EC number 3.1.1.101. PETase displays the highest PET hydrolytic activity at ambient temperature but is heat-labile, which is exceptional among the biochemically characterized PHEs that generally exhibit high thermostability (Table 1). Therefore, sequence alignment of PETase and the thermostable PHEs might identify amino acids that could impart higher thermostability to PETase. As another approach, a recent study by Shirke et al.29 might be applicable to the thermal stabilization of PETase. The authors expressed glycosylated LCC in the eukaryotic host Pichia pastoris, to circumvent protein aggregation at high temperature. This challenge succeeded in significantly improving LCC, in terms of both thermostability and PET hydrolysis. In addition, Furukawa et al.49 reported that the hydrolytic activity of PETase could be enhanced by surface coating of the PET film with anionic surfactants, allowing the anionized PET to recruit more cationic PETase. Anionic surfactants are also expected to act as protein aggregation inhibitors at higher temperatures. Notably, as the temperature approaches the glass transition temperature (Tg) of PET (i.e., 4094

DOI: 10.1021/acscatal.8b05171 ACS Catal. 2019, 9, 4089−4105

Perspective

ACS Catalysis

Figure 4. Proposed PET metabolic pathway by I. sakaiensis. Extracellular PETase hydrolyzes PET to produce MHET as the major product and TPA. The PET hydrolysis products are then transported into the periplasmic space through an outer membrane protein such as porin. MHETase, predicted by its signal peptide sequence17 to be an outer membrane anchored lipoprotein, hydrolyzes MHET into TPA and EG. I. sakaiensis harbors a gene cluster that is highly identical with two TPA degradation gene clusters identified in Comamonas sp. strain E6.57 Expression from this cluster in I. sakaiensis is dramatically upregulated under TPA.17 TPA is taken up into the cytoplasm through the TPA transporter coupled with TPA-binding protein58 and then integrated via protocatechuic acid (PCA) to the tricarboxylic acid (TCA) cycle.17,59,60 EG is metabolized via glyoxylic acid to the TCA cycle.61

various groups.50−54 All PETases were expressed in Escherichia coli. As PETase contains an N-terminal signal peptide, the codon-optimized gene for E. coli was synthesized without a signal peptide to promote efficient expression and cloned into pET-type expression vectors, as shown in Table 3. Various E.

enzyme showed no or very low activity for other tested ester compounds. The enzyme, designated as MHETase, was also assigned a new EC number (3.1.1.102). The biochemical properties of PETase and MHETase, along with their predicted localization, indicated a PET metabolic model by I. sakaiensis (Figures 3B and 4). Both enzymes act in their specific roles, with PETase being responsible for hydrolytic conversion of PET into oligomers that include MHET as their main component and MHETase further hydrolyzing MHET into PET monomers, TPA, and EG. Moreover, expression of the PETase gene was dramatically upregulated in the presence of PET (but not TPA) in the culture of I. sakaiensis, raising the question of which molecule(s) induces its expression.

Table 3. PDB Number, Resolution, and Expression System for PETase

5. THREE-DIMENSIONAL (3D) STRUCTURES OF PETase Structural analyses of the PETase produced by I. sakaiensis 201-F6, which uses PET as its major energy and carbon source, revealed that PETase has very unique characteristics in comparison with those of closely related cutinases.50−54,62 5.1. Expression of Recombinant PETase. Several crystal structures of recombinant PETases have been reported by

PDB ID

resolution (Å)

vector

host strain

ref

6EQE 5XJH 6ANE 5XG0 5YFE

0.92 1.54 2.02 1.58 1.39

pET21b pET15b pET24b pET32a pET21b

C41 (DE3) Rosetta gami-B (DE3) BL21 Gold (DE3) BL21trxB (DE3) BL21 CodonPlus (DE3) RIPL

54 51 52 50 53

coli strains such as C41 (DE3), BL21-gold (DE3), Rosetta gami-B (DE3), BL21trxB (DE3), and BL21CodonPlus (DE3) RIPL were used as expression hosts of the PETase gene. Among these, Rosetta gami-B (DE3) and BL21 trxB enable cytoplasmic disulfide bond formation. As PETase has two distinct disulfide bonds, these E. coli strains are advantageous for functional expression of PETase. In the case of using E. coli 4095

DOI: 10.1021/acscatal.8b05171 ACS Catal. 2019, 9, 4089−4105

Perspective

ACS Catalysis

Figure 5. 3D structure of PETase (PDB code 5XJH).51 The catalytic residues and disulfide bond formations are shown in the structures.

Figure 6. Catalytic mechanism of the α/β serine hydrolase family.

an acyl-enzyme intermediate. This intermediate is hydrolyzed to the product via the second tetrahedral intermediate. The role of the catalytic Ser160 has been validated by its mutation to an alanine.50 The other catalytic triad residues, Asp206 and His237, are located in two highly conserved reverse turns that follow strands β7 and β8. The shallow groove located above Ser160 constitutes the putative substratebinding pocket. Notably, the active site of PETase appears to be wider than that in related cutinases. PETase harbors two disulfide bonds (disulfide bonds 1 and 2, Figures 5 and 7), whereas homologous cutinases contain only disulfide bond 1. The PETase-specific disulfide bond 2 is located adjacent to the active site and connects the β7-α5 and β8-α6 loops that harbor the catalytic triad. Thus, the basic structure of PETase resembles that of other PHEs but includes some very unique structural features, described in further detail in subsequent sections. 5.3. Structures of PETase-Ligand Complexes and the Reaction Mechanism. To clarify the structural aspects of the active site of PETase, Han et al.50 attempted to obtain complexes of the wild-type enzyme with various ligands. As the expected complex was not obtained, inactive S160A and R132G/S160A mutants (numbered as in Joo et al.51) that had lost the hydrolytic activity were later used. The group obtained structures by soaking crystals of the variants with 1-(2-

BL21-gold (DE3), active enzymes were obtained by chemical refolding.52 Moreover, because PETase readily aggregates, 100−150 mM sodium chloride was added to the purified enzyme solution. 5.2. Overall Structure of PETase. Crystal structures of PETase were determined in several laboratories50−54 at high to atomic resolution (Table 3). The enzyme adopts the canonical α-/β-hydrolase fold, with a core consisting of seven α-helices and nine β-strands, as shown in Figure 5. Enzymes in this group include proteases, lipases, and esterases, all of which have a nucleophile-His-acid catalytic triad, evolved to efficiently operate on substrates with different chemical composition. The nucleophile is most commonly a side chain of serine or cysteine. For example, cutinase, which belongs to the α/β serine hydrolase family, employs a canonical catalytic triad consisting of Ser-His-Asp. The location of the PETase catalytic triad comprising Ser160His237-Asp206 was identified on the basis of the high sequence identity of that enzyme in comparison to cutinases (Figure 5). As shown in Figure 6, the residues (His and Asp) form a charge-relay network that activates the nucleophile (Ser). In turn, the nucleophile attacks an ester bond of the substrate, forming the first tetrahedral intermediate that is converted to 4096

DOI: 10.1021/acscatal.8b05171 ACS Catal. 2019, 9, 4089−4105

Perspective

ACS Catalysis

Figure 7. Amino-acid sequence alignments of PET hydrolytic enzymes (PHEs). Six PHEs of three phylogenetically different types (Figure 2) from I. sakaiensis (Is), Polyangium brachysporum (Pb), Vibrio gazogenes (Vg), Oleispira antarctica (Oa), Thermobifida fusca (Tf), and Saccharomonospora viridis (Sv) were aligned using ClustalW. Secondary structure elements of the Is enzyme are shown on the basis of the structural study by Joo et al.51 Arrows and helices indicate β-sheet and α-helix structures, respectively. Residues of the catalytic triad are highlighted in red. The extended loop is highlighted with a red box. The disulfide bonds 1 and 2 are marked with blue lines.

hydroxyethyl) 4-methyl terephthalate (HEMT, methyl ester of MHET) and p-nitrophenol. In the case of the R132G/S160AHEMT complex, the ligand was bound in a cleft containing Tyr87, Trp159, Met161, Trp185, and His237, as shown in Figure 8B. The residue Trp185 is T-stacked (face to edge) on the benzene ring of HEMT. The other residues provide hydrophobic interaction with HEMT. Han et al.50 also showed that the wobbling conformation of Trp185 is closely related to

the binding of its substrate and product. The backbone NH groups of Tyr87 and Met161 comprise an oxyanion hole. Han et al.50 and Joo et al.51 created these variants and demonstrated that these amino acid residues affect the activity. On the basis of these results, the reaction mechanism of PETase was proposed, as shown in Figure 9.50 Joo et al.51 attempted to obtain a complex with PETase and bis(2-hydroxyethyl) terephthalate (BHET, Chart 1). However, 4097

DOI: 10.1021/acscatal.8b05171 ACS Catal. 2019, 9, 4089−4105

Perspective

ACS Catalysis

binding cleft, which could be divided into two subsites (subsite I and II) as shown in Figure 10 and Table 4. W159H and S238F variants exhibited much lower activity in comparison with that of wild-type PETase.51 These residues are considered to be involved in the depth and breadth of subsite IIa (Figure 10 and Table 4). The decomposition process of the substrate can be divided into two steps, nick generation and terminal digestion. In the nick generation step, four MHET moieties bind to subsites I, IIa, IIb, and IIc. The ester bond is broken in the cleavage site in the vicinity of site I. As previously noted, MHET constitutes the major product in PET decomposition using PETase. For decomposition of PETn with a terminal 2-hydroxyethyl group (HE), four MHET moieties bind to subsites I, IIa, IIb, and IIc. The cleavage of the ester bond results in the production of one MHET and HEterminal PETn−1. Subsequent decomposition of the latter is expected to occur by a mechanism similar to the second cleavage process. Depending on the subsite binding, TPA and BHET may be generated in addition to MHET. The resulting BHET is further decomposed into MHET and EG by PETase. Consequently, MHET, TPA, and EG accumulate in the reaction system upon PET decomposition. To understand the reasons for the high activity of PETase, Joo et al.51 analyzed the amino acid sequences of 69 PETaselike enzymes. They classified the enzymes into two types, I and II, of which 57 enzymes, including Thermobifida fusca cutinase, belong to type I (Table 4 and Figure 2). Type II comprises the remaining 12 enzymes, including PETase. The type II enzymes were further divided into two subtypes, Types IIa (8 enzymes) and IIb (PETase and 3 other enzymes). All 69 enzymes contain the well-conserved catalytic triad consisting of Ser-HisAsp. The residues of subsite I (Tyr87, Gln119, Met161, and Trp185 in PETase) are highly conserved, as shown in Table 4. However, subsite II and the additional disulfide bond 2 markedly differ. No additional disulfide bonds (C203 and C239) or extended loops (Ser245, Asn246, and Gln247) were contained in the type I enzyme. In order to clarify the effect of the additional disulfide bond 2 near the active site, an alanine mutant (C203A/C239A) was prepared and evaluated. The activity of the double mutant and its Tm value were markedly decreased. These results supported the importance of the presence of disulfide bond 2 for the thermal stability of PETase. Moreover, these authors predicted that type I enzymes had lower PET-decomposition activity in comparison to that of PETase, whereas the other three enzymes of Type IIb were presumed to exhibit PET-degrading activity similar to that of PETase.51 In addition to the work of Han et al.50 and Joo et al.51 described above, Fecker et al.52 demonstrated by molecular dynamics simulations that the active site of PETase exhibits higher flexibility at room temperature in comparison to its thermophilic counterparts. This flexibility is controlled by a unique disulfide bond (disulfide bond 2) in its active site, the removal of which leads to destabilization of the catalytic triad and reduction of the hydrolase activity. Liu et al.53 proposed that a wide substrate-binding pocket of PETase is critical for its excellent ability to hydrolyze crystallized PET. Furthermore, Austin et al.54 demonstrated that PETase has a more open active-site cleft than homologous cutinases. By narrowing the binding cleft through mutation of two active-site residues to amino acids conserved in cutinases, they observed improved PET degradation, suggesting that PETase is not fully optimized for crystalline PET degradation. They also suggested that

Figure 8. Structure of the PETase active site. (A) Superposition of the structures of PETase (green) and cutinase from T. fusca KW3 (Tf Cut2, 4CG1) (gold).63 Residues constituting subsites I and II,51 the catalytic triad, and extended loop are shown inside red rings. (B) Electrostatic potential surface presentation of PETase with HEMT, as seen in the structure with PDB code 5XH3.50 Representative amino acid residues of the active site are shown in single-letter representation.

Figure 9. Reaction mechanism of PETase.

potentially owing to the low solubility of BHET, this complex could not be obtained. Therefore, they carried out docking calculations using the HEMT tetramer in order to estimate the binding mode of the substrate. From this, they estimated an elongated groove of approximately 40 Å as the substrate 4098

DOI: 10.1021/acscatal.8b05171 ACS Catal. 2019, 9, 4089−4105

Perspective

ACS Catalysis

Figure 10. Substrate binding mode of PETase and the PET decomposition process.

Table 4. Amino Acid Sequence Alignment of Key Residues in PHEs residue in PETase extended loop S242 Type I Type IIa Type IIb

G243

TSILMF GTSD ST

disulfide bond 2

N244

S245

ND YGNS N

SPVTIAE

GD G

N246

PSTIFLA

S residue in PETase

NYIS N

subsite I Type I Type IIa Type IIb

Q247

C203

C239

NEDG QA

GA C C

FA C C

subsite II

Y87

Q119

M161

W185

T88

A89

W159

S238

N241

YF YF YF

QY Q Q

M M M

W W W

TL VL T

AG SL A

H W W

F FY ST

NG N N

additional protein engineering to increase PETase performance is realistic and highlighted the need for further developments of structure/activity relationships for the biodegradation of synthetic polyesters.54 These new findings are of considerable importance for further in-depth research and engineering of PETase, and should advance the implementation of plastic biodegradation strategies. On the basis of these structural analyses of PETase and comparison with the available data for cutinases, the following unique features were demonstrated: (1) Subsite II of PETase clearly differs from the equivalent site in cutinase. PETase contains a pair of Trp159Ser238 residues at subsite II that create sufficient space to accommodate the substrate, PET, whereas the corresponding residues in TfCut 2, a pair consisting of His169-Phe249, would impede PET binding (Figure 8A and Table 4). (2) In addition, the extra three amino acids Ser245, Asn246, and Gln247 in the connecting loop of PETase provide sufficient space to accommodate PET, unlike that

available in the corresponding region of TfCut 2 (Figure 8A and Table 4). (3) PETase has an extra disulfide bond 2 at the vicinity of the active site that is important for its thermal stability and activity (Figures 5 and 7 and Table 4). PETase also exhibits higher flexibility and a wider active site cleft in comparison with cutinases, with the flexibility being controlled by the novel disulfide bond 2. Overall, such information will likely be very useful for protein engineering of PETase-like enzymes for their utilization in practical industrial applications.

6. EVOLUTION OF PETase AND MHETase A phylogenetic tree of PHEs based on the publicly available amino acid sequences (Table 1) reveals that they are distributed in both eukaryotes and bacteria (Figure 2). Although the amino acid identity of the fungal and bacterial enzymes is quite low,17 their 3D structures share the common α/β hydrolase fold.50,64 The bacterial domain is divided into actinobacterial and proteobacterial branches. Thermophilic 4099

DOI: 10.1021/acscatal.8b05171 ACS Catal. 2019, 9, 4089−4105

Perspective

ACS Catalysis

7.2. Ideonella sakaiensis 201-F6. I. sakaiensis 201-F6 may be used for complete PET degradation into CO2 and H2O or for monomer recycling after deletion of the degradation pathway for TPA and/or EG. Both the authentic strain and recombinant strains might be used for more rapid degradation of PET in comparison with that of microbial consortium No. 46. 7.3. PETase and MHETase. Recombinant and overexpressed PETase and MHETase may be used for more rapid degradation of PET or monomer recycling (TPA and EG) or for bioconversion to high-value compounds (e.g., PCA). Following improvement of the enzymatic activity, along with enhancement of the protein stability at high temperatures and concentrations of detergents by directed evolution and/or surface engineering, an enzyme mixture of PETase and MHETase may be used as additives in detergents for washing polyester-containing clothing. The enzyme mixture may be applied as well for rapid and efficient degradation and reduction strategies of microplastics and plastic microbeads in the environment. They might be also used for the degradation of another semiaromatic polyester, polyethylene2,5-furandicarboxylate (PEF).54 Overall, such technologies will likely contribute in the near future to biorecycling of used PET, bioconversion of the monomer TPA to PCA, PET-surface treatment, and degradation and reduction strategies for microplastics and plastic microbeads, as well as for the degradation of PEF. Such outcomes would be expected to provide clear benefits for both the economy and the environment.

actinomicetes Thermobifida strains have served as a major reservoir of PHEs since 2005,14 whereas recognition of the proteobacterial enzymes is relatively recent. These include PETase and three other enzymes identified through a search utilizing the hidden Markov model (HMM) constructed from the nine known polypeptide sequences of PHEs: PET5 from Oleispira antarctica RB-8, PET6 from Vibrio gazogenes, and PET12 from Polyangium brachysporum.37 The discovery of genes necessary for PET metabolism in I. sakaiensis 201-F6 raises the question of how this bacterium obtained the capacity to metabolize PET. To address the issue, we attempted to find organisms that harbored a set of gene homologues of the signature enzymes for PET metabolism (PETase, MHETase, TPA dioxygenase, and PCA dioxygenase) by analyzing the fully sequenced genome database. No single organism containing a full set could be found; however, 33 organisms were noted as carrying three enzymes (other than PETase), suggesting that a genomic basis to metabolize MHET analogues had been evolutionarily prepared prior to the participation of ancestral PETase proteins in the pathway. Protein evolution analysis on PETase conducted by Joo et al.51 suggested a further detailed model (Table 4 and Figure 2). A total of 69 phylogenetically diverse potential PET hydrolytic proteins selected through position-specific iterated BLAST (PSI-BLAST) were classified into two types (Type I and Type II), with all of the microbial isolates with Type I proteins (57 isolates) belonging to the phylum Actinobacteria and the other isolates with Type II proteins including PETase belonging to the phylum Proteobacteria. This correlation indicates that the recent evolution of PETase occurred in Proteobacteria. However, other Ideonella strains; i.e., I. dechloratans LMG 28178T and I. azotifigens JCM 15503T, displayed no growth on PET under the same conditions that allowed growth of I. sakaiensis.48 An HMM search of marine and terrestrial metagenomes in the public databases by Danso et al.37 to look for the presence of potential PHEs identified 349 putative PHE genes, originating mostly from Proteobacteria, Actinobacteria, and Bacteroidetes. The limited distribution of this type of enzyme at the phylum level might indicate the late appearance of this type of enzyme. Furthermore, although Bacteroidetes has been considered to be a minor host for PHEs according to BLASTp search, this phylum is likely to be the main host of PET hydrolytic enzymes, particularly in marine environments.

8. OUTLOOK PET, currently one of the most widely utilized synthetic plastics, is used in textiles, packaging, and other applications owing to its excellent durability, low price, and processability. However, after a single use, the majority of the waste causes severe environmental pollution, such as contamination of the oceans and land.65−71 Microplastic particles of PET and other plastics of less than 5 mm in size72−75 have also emerged as an environmental concern. Here we describe an outlook regarding these issues based on our findings. 8.1. Screening. In order to identify enzymes that can degrade plastics such as PET, screening of target microorganisms is crucial. In general, an appropriate substrate should be used as a sole carbon and energy source. Moreover, to degrade PET, it is necessary for enzymes or microorganisms to adhere onto the film surface. In our screening, we mostly utilized PET films. Samples were collected at a PET recycling facility, which led to the successful isolation of microbial consortium No. 46 and I. sakaiensis 201-F6, along with PETase and MHETase.17,23 Cutinase-like enzymes known to hydrolyze PET as well as their usual substrate, cutin, have also been screened by using composts or by metagenomic analysis.14,27,31 Nevertheless, it is essential to develop simpler and more reliable assay methods for effective screening. For example, although we prepared substrates harboring a chromophore, the limited solubility in water did not make screening feasible. The establishment of assay methods should be prioritized to allow the rapid identification of better plastic-degrading enzymes to facilitate the development and implementation of plastics recycling, as exemplified in the application of high-throughput analysis for the enzymatic hydrolysis of biodegradable polyesters.40

7. POSSIBLE APPLICATION ASPECTS We succeeded in finding a microbial consortium, No. 46, I. sakaiensis 201-F6, and PET-degrading enzymes that exhibit potential degradation of PET. Their possible application aspects are described below. 7.1. Microbial Consortium No. 46. The microbial consortium No. 46 contains several kinds of microorganisms, yet it retains PET-degrading ability even after subculture for more than several months. Thus, it has potential utility for the complete degradation of PET waste products (e.g., PET bottles, fibers, films, and sheets), with the generated CO2 applied for fertilization in greenhouses. Notably, this process does not require any specialized bioreactor or energy input for cultivation. The microbial consortium No. 46 might also be applied for the biodegradation of “PET oligomers” and “monomers”, which are found in large amounts in wastewater generated from factories during alkaline-washing processing of PET. 4100

DOI: 10.1021/acscatal.8b05171 ACS Catal. 2019, 9, 4089−4105

Perspective

ACS Catalysis 8.2. Microbial Consortium No. 46. Among the various kinds of bacteria, yeast-like cells, and protozoa that comprise microbial consortium No. 46, the roles of individual components from the approximately 20 kinds of identified bacteria have been investigated. Initially, Bacillus megaterium forms a biofilm on the film surface. In the biofilm, Rhizopus sp. cleaves the ester linkage of the PET chain into BHET, which is further decomposed into TPA and EG by Pseudomonas sp. The generated TPA and EG monomeric units are eventually assimilated by Pigmentiphaga sp. and Mycobacterium sp., respectively. Microbial consortium No. 46, which was isolated in 2001, was confirmed to exhibit consistent and reproducible PET degradation activity and thus may be suitable to degrade not only waste PET but also microplastics in sewage sludge.76 In general, the efficiency of substrate degradation increases when a microbial biofilm is formed on a substrate.77 Microbial populations that form biofilms display higher metabolic activity in comparison to microbes in the planktonic mode of growth.78 Similarly, Cavaliere et al.79 noted that the evolutionary resilience of microbial communities should be considered when designing robust biotechnological applications. In particular, De Tender et al.80 reported that biofilm formation is very important in the degradation of plastics. Mercier et al.81 investigated the fate of eight different polymers under uncontrolled composting conditions and proposed that biofilms might contribute to the alteration processes of all the polymers studied. In turn, through analysis of the community composition of microbes colonizing marine plastic debris, Oberbeckmann et al.82 postulated that biofilms could serve as target communities to probe for future discovery of plasticdegrading microbes, as well as individual genes involved in the enzymatic process. Thus, microbial consortia or biofilms may constitute effective solutions toward current plastic pollution issues, and scaling up of the degradation systems should be studied for commercial operations. 8.3. I. sakaiensis 201-F6. Although isolation of I. sakaiensis 201-F6 was serendipitous, it may be highly useful for bioremediation, as suggested by Atashgahi et al.83 Joo et al.51 further indicated that Acidovorax delafieldii,84 [Polyangium] brachysporum,85 and Burkholderiales bacterium86 should constitute good sources of PETase-like enzymes, as the membership of all of these bacteria in the order Burkholderiales suggests that they would have evolved in a similar manner. These findings also highlight the potential value in searching for other PETase-like enzymes in these families. In addition, I. sakaiensis 201-F6 may contribute to sustainable development goals (SDGs) in terms of system metabolic engineering.87,88 8.4. PETase and MHETase. The enzymatic properties of PETase17,23 have been elucidated and compared with those of cutinase-like enzymes from T. fusca (TfH),14 F. solani (FsC),31 and a cutinase homologue from leaf-branch compost metagenome (LC cutinase)27 that are distant from each other on the phylogenetic tree. These unique features are further supported by 3D structural analyses.50−54 However, the low thermal stability of PETase constitutes a bottleneck for the practical enzymatic degradation of PET. On the basis of structural information, rational protein engineering should be carried out to improve the thermal stability of these enzymes; such improvement might be also carried out by glycosylation.29 In the case of MHETase, unique features such as substrate specificity have been elucidated, although the 3D structure has not been determined.17 Owing to the novelty of both enzymes, they have been assigned EC numbers in the list of IUBMB

Enzyme Nomenclature in 2016: PETase (EC 3.1.1.101) and MHETase (EC 3.1.1.102). 8.5. Evolution. The comparative analyses of PETase raise the questions of how it evolved and the identity of the ancestral gene. A candidate for the predecessor of such rapid evolution over the past 50 years since the commercial synthesis of PET might be cutinase, an enzyme that degrades cutin from plant leaves. Examples of such rapid natural evolution are scarce; however, one prominent case consists of the evolution of atrazine chlorohydrase from melamine deaminase.89 Addressing these questions requires sufficient experimental results and data, including 3D structures of the phylogenetically related enzymes. 8.6. Possible Application Aspects. 8.6.1. Screening for Other PET-Degrading Microorganism(s). The same strategy as that used for screening of I. sakaiensis 201-F6 might be applied. The existence of such PET-degrading microorganisms as I. sakaiensis 201-F6 in a natural environment has been demonstrated. It is thus worthwhile to screen for other microorganisms, including (1) thermophilic microorganisms that degrade PET at high temperature and (2) salt-tolerant microorganisms that degrade microplastics, causing ocean pollution. 8.6.2. Bioprocess Production of PCA from Waste PET To Synthesize Value-Added Chemicals. In the cells of I. sakaiensis 201-F6, TPA is further degraded to PCA, which is a key compound in the synthesis of value-added chemicals such as catechol, adipic acid, and 6,6 nylon. PCA might be produced readily by using genetically modified I. sakaiensis 201-F6. This would represent a bioprocess production of PCA to develop an effective biorefinery system for early transition to a more sustainable society. 8.6.3. Treatment of PET Fibers To Produce Textures. A large portion of PET resin is manufactured into fibers and textiles. PET fiber is currently treated with a strong alkaline solution at high temperature to generate textures on the surface, although this is associated with high cost and may lead to severe water pollution. Treatment with PETase and MHETase may be an alternative to provide textures on the fibers. Through a more environmentally friendly manner in comparison to that of conventional alkaline treatment, a more silklike texture along with diversified, excellent color might be obtained via more irregular pit formation on the fiber surface. 8.6.4. Additives to Detergents. Cellulase is currently used as one of the additives to detergents for cotton fibers. In the same manner, PETase and MHETase might be used as additives to detergents for PET fibers. 8.6.5. CO2 Fertilization. PET waste such as bottles, fibers composed of PET/cotton, and films might be treated using consortium No. 46. CO2 generated from the degradation could be introduced into greenhouses or sugar cane farms to enhance plant growth. After numerous arguments regarding the effect of free-air concentration enrichment (FACE),90−92 Sakurai et al.93 reported in 2014 that soybean yields were enhanced by the increase of atmospheric CO2 concentration in three major soybean-producing countries. The increased yields during 2002−2006 in comparison with those in the 1980s in the US, Brazil, and China were +4.34, +7.57, and +5.10%, respectively. Thus, CO2 fertilization in agricultural production represents a promising strategy for use of the generated CO2 and reduction of CO2 emission into the environment. 8.6.6. Recovery of TPA or MHET for Resynthesis of PET. Recovery of the reaction intermediates TPA and MHET might 4101

DOI: 10.1021/acscatal.8b05171 ACS Catal. 2019, 9, 4089−4105

Perspective

ACS Catalysis be carried out by introducing gene modifications to I. sakaiensis 201-F6. Currently, only a limited number of collected PET bottles are reprocessed to new bottles (bottle-to-bottle recycling) owing to the ease of thermal degradation of the polymer chain during remolding, termed “material recycling”. However, if renewed PET can be synthesized from the TPA and EG obtained via the biodegradation processes of PET, such bottle-to-bottle recycling may be rendered more feasible through the methodologies of “chemical recycling”. This idea may substantially reduce the consumption of fossil resources. 8.6.7. Others. Microplastics currently represent a serious issue not only in oceans or rivers but also at the sites of wastewater treatment. Huge amounts of microplastics are accumulating in sewage, of which PET accounts for a notinsignificant percentage. In order to remove these microplastics, microbial consortium No. 46 might be used because these microorganisms can grow at ambient temperature by assimilating PET for growth. Furthermore, I. sakaiensis 201-F6, or PETase and MHETase, might be useful for the biofunctionalization of PET films or PET fibers.15,94−96 8.7. Poly(ethylene furanoate) (PEF) and Novel Plastics. Kucherov el al.97 recently published a perspective entitled “Chemical transformations of biomass-derived C6furanic platform chemicals for sustainable energy research, materials science, and synthetic building blocks”. In addition, Zhu et al.98 reported a polymer system based on γbutyrolactone (GBL) with a trans-ring fusion at the α and β positions. Notably, this plastic could be recycled repeatedly through chemical methods without loss of the properties. A review by Pellis et al.99 introduced the enzymatic hydrolysis of PEF, which is biorecyclable and is a substituent of PET. Similarly, hydrolysis of PEF by PETase from I. sakaiensis 201F6 has also been demonstrated.54 In summary, the presented versatilities and perspectives of our isolates (microbial consortium No. 46 and I. sakaiensis 201-F6) or enzymes (PETase and MHETase) regarding plastic, especially PET, degradation illustrate their potential in various fields to promote a new plastics economy. Although this may require substantial time for thorough implementation, the current advances support the value of prioritizing research and development in these areas.

■

Technology (NAIST), 8916-5 Takayama-cho, Ikoma, Nara 630-0192, Japan. E-mail: [email protected]. ∇ K.H.: Research Institute of Innovative Technology for the Earth (RITE), 9-2 Kizugawadai, Kizugawa-shi, Kyoto 6190292, Japan. E-mail: [email protected]. ○ Y.K.: Center for Fiber and Textile Science, Kyoto Institute of Technology, Matsugasaki, Sakyo-ku, Kyoto 606-8585, Japan. E-mail: [email protected]. Author Contributions ∥

I.T. and S.Y. contributed equally.

Notes

The authors declare no competing financial interest.

■

ACKNOWLEDGMENTS We acknowledge fruitful discussions with T. Takehana, H. Yamaji, Y. Maeda, and K. Toyohara. We thank A. Wlodawer for comments on this manuscript. This work was supported by a JSPS KAKENHI Grant-in-Aid for Scientific Research (B) (17H03794 to S.Y.) and a JSPS KAKENHI Grant-in-Aid for Scientific Research (A) (JP18H03857 to K.M.).

■

REFERENCES

(1) PlasticsEurope Plastics - the Facts 2017: An Analysis of European Plastics Production, Demand and Waste Data; https:// www.plasticseurope.org/application/files/5715/1717/4180/Plastics_ the_facts_2017_FINAL_for_website_one_page.pdf (accessed Jan 30, 2019). (2) PlasticsEurope World Plastic Production 1950−2015; https:// committee.iso.org/files/live/sites/tc61/files/ The%20Plastic%20Industry%20Berlin%20Aug%202016%20%20Copy.pdf (accessed Jan. 30, 2019). (3) Lebreton, L. C. M.; van der Zwet, J.; Damsteeg, J. W.; Slat, B.; Andrady, A.; Reisser, J. River plastic emissions to the world’s oceans. Nat. Commun. 2017, 8, 15611. (4) Ellen Macathur Foundation, The New Plastic Economy: Rethinking The Future of Plastics & Catalysing Action; https:// www.ellenmacarthurfoundation.org/assets/downloads/publications/ NPEC-Hybrid_English_22-11-17_Digital.pdf (accessed Jan 30, 2019). (5) The National Association for PET Container Resources Report on Postconsumer PET Container Recycling Activity in 2016; http:// www.plasticsmarkets.org/jsfcode/srvyfiles/napcor_2016ratereport_ final_1.pdf (accessed Jan. 30, 2019). (6) Tokiwa, Y.; Calabia, B. P.; Ugwu, C. U.; Aiba, S. Biodegradability of plastics. Int. J. Mol. Sci. 2009, 10, 3722−3742. (7) Devi, R. S. K.; Natarajan, K.; Nivas, D.; Kannan, K.; Chandra, S.; Antony, A. R. The role of microbes in plastic degradation In Environmental waste management; Chandra, R., Ed.; CRC Press: 2015; pp 341−370. (8) Webb, H. K.; Arnott, J.; Crawford, R. J.; Ivanova, E. P. Plastic degradation and its environmental implications with special reference to poly(ethylene terephthalate). Polymers 2013, 5, 1−18. (9) Shah, A. A.; Hasan, F.; Hameed, A.; Ahmed, S. Biological degradation of plastics: A comprehensive review. Biotechnol. Adv. 2008, 26, 246−265. (10) Wei, R.; Zimmermann, W. Microbial enzymes for the recycling of recalcitrant petroleum-based plastics: how far are we? Microb. Biotechnol. 2017, 10, 1308−1322. (11) Marten, E.; Müller, R. J.; Deckwer, W. D. Studies on the enzymatic hydrolysis of polyesters I. Low molecular mass model esters and aliphatic polyesters. Polym. Degrad. Stab. 2003, 80, 485−501. (12) Marten, E.; Müller, R. J.; Deckwer, W. D. Studies on the enzymatic hydrolysis of polyesters. II. Aliphatic-aromatic copolyesters. Polym. Degrad. Stab. 2005, 88, 371−381.

AUTHOR INFORMATION

Corresponding Author

*E-mail for K.O.: [email protected]. ORCID

Ikuo Taniguchi: 0000-0001-7644-8723 Shosuke Yoshida: 0000-0002-2831-9027 Kazumi Hiraga: 0000-0001-6980-3408 Kenji Miyamoto: 0000-0003-0080-4693 Yoshiharu Kimura: 0000-0003-4431-0193 Kohei Oda: 0000-0003-3243-6629 Present Addresses ⊥

I.T.: International Institute for Carbon-Neutral Energy Research (WPI-I2CNER), Kyushu University, 744 Moto-oka, Nishi-ku, Fukuoka, 819−0395, Japan. E-mail: ikuot@i2cner. kyushu-u.ac.jp. # S.Y.: Institute for Research Initiatives, Division for Research Strategy, Nara Institute of Science and Technology (NAIST), 8916-5 Takayama-cho, Ikoma, Nara 630-0192, Japan, and Division of Biological Science, Nara Institute of Science and 4102

DOI: 10.1021/acscatal.8b05171 ACS Catal. 2019, 9, 4089−4105

Perspective

ACS Catalysis (13) Vertommen, M. A.; Nierstrasz, V. A.; Veer, M.; Warmoeskerken, M. M. Enzymatic surface modification of poly(ethylene terephthalate). J. Biotechnol. 2005, 120, 376−386. (14) Müller, R. J.; Schrader, H.; Profe, J.; Dresler, K.; Deckwer, W. D. Enzymatic degradation of poly(ethylene terephthalate): Rapid hydrolyse using a hydrolase from T. f usca. Macromol. Rapid Commun. 2005, 26, 1400−1405. (15) Zimmermann, W.; Billig, S. Enzymes for the biofunctionalization of poly(ethylene terephthalate). Adv. Biochem. Eng./Biotechnol. 2010, 125, 97−120. (16) Wei, R.; Zimmermann, W. Biocatalysis as a green route for recycling the recalcitrant plastic polyethylene terephthalate. Microb. Biotechnol. 2017, 10, 1302−1307. (17) 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. (18) Eberl, A.; Heumann, S.; Bruckner, T.; Araujo, R.; CavacoPaulo, A.; Kaufmann, F.; Kroutil, W.; Guebitz, G. M. Enzymatic surface hydrolysis of poly(ethylene terephthalate) and bis(benzoyloxyethyl) terephthalate by lipase and cutinase in the presence of surface active molecules. J. Biotechnol. 2009, 143, 207−212. (19) Guebitz, G. M.; Cavaco-Paulo, A. Enzymes go big: surface hydrolysis and functionalization of synthetic polymers. Trends Biotechnol. 2008, 26, 32−38. (20) Ribitsch, D.; Heumann, S.; Trotscha, E.; Herrero Acero, E.; Greimel, K.; Leber, R.; Birner-Gruenberger, R.; Deller, S.; Eiteljoerg, I.; Remler, P.; Weber, T.; Siegert, P.; Maurer, K. H.; Donelli, I.; Freddi, G.; Schwab, H.; Guebitz, G. M. Hydrolysis of polyethyleneterephthalate by p-nitrobenzylesterase from Bacillus subtilis. Biotechnol. Prog. 2011, 27, 951−960. (21) Nimchua, T.; Punnapayak, H.; Zimmermann, W. Comparison of the hydrolysis of polyethylene terephthalate fibers by a hydrolase from Fusarium oxysporum LCH I and Fusarium solani f. sp. pisi. Biotechnol. J. 2007, 2, 361−364. (22) Nimchua, T.; Eveleigh, D. E.; Sangwatanaroj, U.; Punnapayak, H. Screening of tropical fungi producing polyethylene terephthalatehydrolyzing enzyme for fabric modification. J. Ind. Microbiol. Biotechnol. 2008, 35, 843−850. (23) Yoshida, S.; Hiraga, K.; Takehana, T.; Taniguchi, I.; Yamaji, H.; Maeda, Y.; Toyohara, K.; Miyamoto, K.; Kimura, Y.; Oda, K. Response to Comment on “A bacterium that degrades and assimilates poly(ethylene terephthalate). Science 2016, 353, 759. (24) Bornscheuer, U. T. Feeding on plastic. Science 2016, 351, 1154−1155. (25) Silva, C.; Da, S.; Silva, N.; Matama, T.; Araujo, R.; Martins, M.; Chen, S.; Chen, J.; Wu, J.; Casal, M.; Cavaco-Paulo, A. Engineered Thermobifida fusca cutinase with increased activity on polyester substrates. Biotechnol. J. 2011, 6, 1230−1239. (26) Chen, S.; Tong, X.; Woodard, R. W.; Du, G.; Wu, J.; Chen, J. Identification and characterization of bacterial cutinase. J. Biol. Chem. 2008, 283, 25854−25862. (27) 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. (28) Sulaiman, S.; You, D. J.; Kanaya, E.; Koga, Y.; Kanaya, S. Crystal structure and thermodynamic and kinetic stability of metagenome-derived LC-cutinase. Biochemistry 2014, 53, 1858−1869. (29) Shirke, A. N.; White, C.; Englaender, J. A.; Zwarycz, A.; Butterfoss, G. L.; Linhardt, R. J.; Gross, R. A. Stabilizing leaf and branch compost cutinase (LCC) with glycosylation: Mechanism and effect on PET hydrolysis. Biochemistry 2018, 57, 1190−1200. (30) Griswold, K. E.; Mahmood, N. A.; Iverson, B. L.; Georgiou, G. Effects of codon usage versus putative 5′-mRNA structure on the expression of Fusarium solani cutinase in the Escherichia coli cytoplasm. Protein Expression Purif. 2003, 27, 134−142.

(31) Silva, C. M.; Carneiro, F.; O’Neill, A.; Fonseca, L. P.; Cabral, J. S. M.; Guebitz, G.; Cavaco-Paulo, A. Cutinase - A new tool for biomodification of synthetic fibers. J. Polym. Sci., Part A: Polym. Chem. 2005, 43, 2448−2450. (32) Ronkvist, A. M.; Xie, W. C.; Lu, W. H.; Gross, R. A. Cutinasecatalyzed hydrolysis of poly(ethylene terephthalate). Macromolecules 2009, 42, 5128−5138. (33) Herrero Acero, E.; Ribitsch, D.; Steinkellner, G.; Gruber, K.; Greimel, K.; Eiteljoerg, I.; Trotscha, E.; Wei, R.; Zimmermann, W.; Zinn, M.; Cavaco-Paulo, A.; Freddi, G.; Schwab, H.; Guebitz, G. Enzymatic surface hydrolysis of PET: Effect of structural diversity on kinetic properties of cutinases from Thermobifida. Macromolecules 2011, 44, 4632−4640. (34) Ribitsch, D.; Acero, E. H.; Greimel, K.; Eiteljoerg, I.; Trotscha, E.; Fredd, G.; Schwab, H.; Guebitz, G. M. Characterization of a new cutinase from Thermobifida alba for PET-surface hydrolysis. Biocatal. Biotransform. 2012, 30, 2−9. (35) Ribitsch, D.; Herrero Acero, E.; Greimel, K.; Dellacher, A.; Zitzenbacher, S.; Marold, A.; Rodriguez, R. D.; Steinkellner, G.; Gruber, K.; Schwab, H.; Guebitz, G. M. A new esterase from Thermobif ida halotolerans hydrolyses polyethylene terephthalate (PET) and polylactic acid (PLA). Polymers 2012, 4, 617−629. (36) Kawai, F.; Oda, M.; Tamashiro, T.; Waku, T.; Tanaka, N.; Yamamoto, M.; Mizushima, H.; Miyakawa, T.; Tanokura, M. A novel Ca2+-activated, thermostabilized polyesterase capable of hydrolyzing polyethylene terephthalate from Saccharomonospora viridis AHK190. Appl. Microbiol. Biotechnol. 2014, 98, 10053−10064. (37) Danso, D.; Schmeisser, C.; Chow, J.; Zimmermann, W.; Wei, R.; Leggewie, C.; Li, X.; Hazen, T.; Streit, W. R. New insights into the function and global distribution of polyethylene terephthalate (PET)degrading bacteria and enzymes in marine and terrestrial metagenomes. Appl. Environ. Microbiol. 2018, 84, No. e02773-17. (38) Wei, R.; Oeser, T.; Then, J.; Kuhn, N.; Barth, M.; Schmidt, J.; Zimmermann, W. Functional characterization and structural modeling of synthetic polyester-degrading hydrolases from Thermomonospora curvata. AMB Express 2014, 4, 44. (39) Wei, R.; Oeser, T.; Billig, S.; Zimmermann, W. A highthroughput assay for enzymatic polyester hydrolysis activity by fluorimetric detection. Biotechnol. J. 2012, 7, 1517−1521. (40) Zumstein, M. T.; Kohler, H. E.; McNeill, K.; Sander, M. Highthroughput analysis of enzymatic hydrolysis of biodegradable polyesters by monitoring cohydrolysis of a polyester-embedded fluorogenic probe. Environ. Sci. Technol. 2017, 51, 4358−4367. (41) Wei, R.; Oeser, T.; Zimmermann, W. Synthetic polyesterhydrolyzing enzymes from thermophilic actinomycetes. Adv. Appl. Microbiol. 2014, 89, 267−305. (42) Ahmed, T.; Shahid, M.; Azeem, F.; Rasul, I.; Shah, A. A.; Noman, M.; Hameed, A.; Manzoor, N.; Manzoor, I.; Muhammad, S. Biodegradation of plastics: current scenario and future prospects for environmental safety. Environ. Sci. Pollut. Res. 2018, 25, 7287−7298. (43) Herrero Acero, E.; Ribitsch, D.; Dellacher, A.; Zitzenbacher, S.; Marold, A.; Steinkellner, G.; Gruber, K.; Schwab, H.; Guebitz, G. M. Surface engineering of a cutinase from Thermobifida cellulosilytica for improved polyester hydrolysis. Biotechnol. Bioeng. 2013, 110, 2581− 2590. (44) Ribitsch, D.; Herrero Acero, E.; Przylucka, A.; Zitzenbacher, S.; Marold, A.; Gamerith, C.; Tscheliessnig, R.; Jungbauer, A.; Rennhofer, H.; Lichtenegger, H.; Amenitsch, H.; Bonazza, K.; Kubicek, C. P.; Druzhinina, I. S.; Guebitz, G. M. Enhanced cutinase-catalyzed hydrolysis of polyethylene terephthalate by covalent fusion to hydrophobins. Appl. Environ. Microbiol. 2015, 81, 3586−3592. (45) Dimarogona, M.; Nikolaivits, E.; Kanelli, M.; Christakopoulos, P.; Sandgren, M.; Topakas, E. Structural and functional studies of a Fusarium oxysporum cutinase with polyethylene terephthalate modification potential. Biochim. Biophys. Acta, Gen. Subj. 2015, 1850, 2308−2317. (46) de Castro, A. M.; Carniel, A.; Nicomedes, J.; da Conceicao Gomes, A.; Valoni, E. Screening of commercial enzymes for 4103

DOI: 10.1021/acscatal.8b05171 ACS Catal. 2019, 9, 4089−4105

Perspective

ACS Catalysis poly(ethylene terephthalate) (PET) hydrolysis and synergy studies on different substrate sources. J. Ind. Microbiol. Biotechnol. 2017, 44, 835−844. (47) Gamerith, C.; Vastano, M.; Ghorbanpour, S. M.; Zitzenbacher, S.; Ribitsch, D.; Zumstein, M. T.; Sander, M.; Herrero Acero, E.; Pellis, A.; Guebitz, G. M. Enzymatic degradation of aromatic and aliphatic polyesters by P. pastoris expressed cutinase 1 from Thermobifida cellulosilytica. Front. Microbiol. 2017, 8, 938. (48) Tanasupawat, S.; Takehana, T.; Yoshida, S.; Hiraga, K.; Oda, K. Ideonella sakaiensis sp. nov., isolated from a microbial consortium that degrades PET. Int. J. Syst. Evol. Microbiol. 2016, 66, 2813−2818. (49) Furukawa, M.; Kawakami, N.; Oda, K.; Miyamoto, K. Acceleration of enzymatic degradation of poly(ethylene terephthalate) by surface coating with anionic surfactants. ChemSusChem 2018, 11, 4018−4025. (50) Han, X.; Liu, W.; Huang, J. W.; Ma, J.; Zheng, Y.; Ko, T. P.; Xu, L.; Cheng, Y. S.; Chen, C. C.; Guo, R. T. Structural insight into catalytic mechanism of PET hydrolase. Nat. Commun. 2017, 8, 2106. (51) Joo, S.; Cho, I. J.; Seo, H.; Son, H. F.; Sagong, H. Y.; Shin, T. J.; Choi, S. Y.; Lee, S. Y.; Kim, K. J. Structural insight into molecular mechanism of poly(ethylene terephthalate) degradation. Nat. Commun. 2018, 9, 382. (52) Fecker, T.; Galaz-Davison, P.; Engelberger, F.; Narui, Y.; Sotomayor, M.; Parra, L. P.; Ramirez-Sarmiento, C. A. Active site flexibility as a hallmark for efficient PET degradation by I. sakaiensis PETase. Biophys. J. 2018, 114, 1302−1312. (53) Liu, B.; He, L.; Wang, L.; Li, T.; Li, C.; Liu, H.; Luo, Y.; Bao, R. Protein crystallography and site-direct mutagenesis analysis of the poly(ethylene terephthalate) hydrolase PETase from Ideonella sakaiensis. ChemBioChem 2018, 19, 1471−1475. (54) Austin, H. P.; Allen, M. D.; Donohoe, B. S.; Rorrer, N. A.; Kearns, F. L.; Silveira, R. L.; Pollard, B. C.; Dominick, G.; Duman, R.; El Omari, K.; Mykhaylyk, V.; Wagner, A.; Michener, W. E.; Amore, A.; Skaf, M. S.; Crowley, M. F.; Thorne, A. W.; Johnson, C. W.; Woodcock, H. L.; McGeehan, J. E.; Beckham, G. T. Characterization and engineering of a plastic-degrading aromatic polyesterase. Proc. Natl. Acad. Sci. U. S. A. 2018, 115, E4350−E4357. (55) Ma, Y.; et al. Enhanced poly(ethylene terephthalate) hydrolase activity by protein engineering. Engineering. 2018, 4, 888−893. (56) Suzuki, K.; Hori, A.; Kawamoto, K.; Thangudu, R. R.; Ishida, T.; Igarashi, K.; Samejima, M.; Yamada, C.; Arakawa, T.; Wakagi, T.; Koseki, T.; Fushinobu, S. Crystal structure of a feruloyl esterase belonging to the tannase family: A disulfide bond near a catalytic triad. Proteins: Struct., Funct., Genet. 2014, 82, 2857−2867. (57) Sasoh, M.; Masai, E.; Ishibashi, S.; Hara, H.; Kamimura, N.; Miyauchi, K.; Fukuda, M. Characterization of the terephthalate degradation genes of Comamonas sp. strain E6. Appl. Environ. Microbiol. 2006, 72, 1825−1832. (58) Hosaka, M.; Kamimura, N.; Toribami, S.; Mori, K.; Kasai, D.; Fukuda, M.; Masai, E. Novel tripartite aromatic acid transporter essential for terephthalate uptake in Comamonas sp. strain E6. Appl. Environ. Microbiol. 2013, 79, 6148−6155. (59) Perez-Pantoja, D.; Donoso, R.; Agullo, L.; Cordova, M.; Seeger, M.; Pieper, D. H.; Gonzalez, B. Genomic analysis of the potential for aromatic compounds biodegradation in Burkholderiales. Environ. Microbiol. 2012, 14, 1091−1117. (60) Wilkes, R. A.; Aristilde, L. Degradation and metabolism of synthetic plastics and associated products by Pseudomonas sp.: Capabilities and challenges. J. Appl. Microbiol. 2017, 123, 582−593. (61) Muckschel, B.; Simon, O.; Klebensberger, J.; Graf, N.; Rosche, B.; Altenbuchner, J.; Pfannstiel, J.; Huber, A.; Hauer, B. Ethylene glycol metabolism by Pseudomonas putida. Appl. Environ. Microbiol. 2012, 78, 8531−8539. (62) Chen, C. C.; Han, X.; Ko, T. P.; Liu, W.; Guo, R. T. Structural studies reveal the molecular mechanism of PETase. FEBS J. 2018, 285, 3717−3723. (63) Roth, C.; Wei, R.; Oeser, T.; Then, J.; Follner, C.; Zimmermann, W.; Strater, N. Structural and functional studies on a

thermostable polyethylene terephthalate degrading hydrolase from Thermobif ida f usca. Appl. Microbiol. Biotechnol. 2014, 98, 7815−7823. (64) Longhi, S.; Czjzek, M.; Lamzin, V.; Nicolas, A.; Cambillau, C. Atomic resolution (1.0 Å) crystal structure of Fusarium solani cutinase: stereochemical analysis. J. Mol. Biol. 1997, 268, 779−799. (65) Debroas, D.; Mone, A.; Ter Halle, A. Plastics in the North Atlantic garbage patch: A boat-microbe for hitchhikers and plastic degraders. Sci. Total Environ. 2017, 599−600, 1222−1232. (66) Auta, H. S.; Emenike, C. U.; Fauziah, S. H. Distribution and importance of microplastics in the marine environment: A review of the sources, fate, effects, and potential solutions. Environ. Int. 2017, 102, 165−176. (67) Worm, B.; Lotze, H. K.; Jubinville, I.; Wilcox, C.; Jambeck, J. Plastic as a persistent marine pollutant. Annu. Rev. Environ. Resour. 2017, 42, 1−26. (68) Sharma, S.; Chatterjee, S. Microplastic pollution, a threat to marine ecosystem and human health: A short review. Environ. Sci. Pollut. Res. 2017, 24, 21530−21547. (69) Jambeck, J. R.; Geyer, R.; Wilcox, C.; Siegler, T. R.; Perryman, M.; Andrady, A.; Narayan, R.; Law, K. L. Marine pollution. Plastic waste inputs from land into the ocean. Science 2015, 347, 768−771. (70) Eriksen, M.; Lebreton, L. C.; Carson, H. S.; Thiel, M.; Moore, C. J.; Borerro, J. C.; Galgani, F.; Ryan, P. G.; Reisser, J. Plastic pollution in the world’s oceans: More than 5 trillion plastic pieces weighing over 250,000 tons afloat at sea. PLoS One 2014, 9, No. e111913. (71) Ng, E. L.; Lwanga, E. H.; Eldridge, S. M.; Johnston, P.; Hu, H. W.; Geissen, V.; Chen, D. L. An overview of microplastic and nanoplastic pollution in agroecosystems. Sci. Total Environ. 2018, 627, 1377−1388. (72) Suran, M. A planet too rich in fibre: microfibre pollution may have major consequences on the environment and human health. EMBO Rep. 2018, 19, No. e46701. (73) Anbumani, S.; Kakkar, P. Ecotoxicological effects of microplastics on biota: a review. Environ. Sci. Pollut. Res. 2018, 25, 14373− 14396. (74) Mason, S. A.; Welch, V. G.; Neratko, J. Synthetic polymer contamination in bottled water. Front. Chem. 2018, 6, 407. (75) Waring, R. H.; Harris, R. M.; Mitchell, S. C. Plastic contamination of the food chain: A threat to human health? Maturitas 2018, 115, 64−68. (76) Mahon, A. M.; O’Connell, B.; Healy, M. G.; O’Connor, I.; Officer, R.; Nash, R.; Morrison, L. Microplastics in sewage sludge: Effects of treatment. Environ. Sci. Technol. 2017, 51, 810−818. (77) Davey, M. E.; O’Toole, G. A. Microbial biofilms: from ecology to molecular genetics. Microbiol. Mol. Biol. Rev. 2000, 64, 847−867. (78) Orr, I. G.; Hadar, Y.; Sivan, A. Colonization, biofilm formation and biodegradation of polyethylene by a strain of Rhodococcus ruber. Appl. Microbiol. Biotechnol. 2004, 65, 97−104. (79) Cavaliere, M.; Feng, S.; Soyer, O. S.; Jimenez, J. I. Cooperation in microbial communities and their biotechnological applications. Environ. Microbiol. 2017, 19, 2949−2963. (80) De Tender, C.; Devriese, L. I.; Haegeman, A.; Maes, S.; Vangeyte, J.; Cattrijsse, A.; Dawyndt, P.; Ruttink, T. Temporal dynamics of bacterial and fungal colonization on plastic debris in the North Sea. Environ. Sci. Technol. 2017, 51, 7350−7360. (81) Mercier, A.; Gravouil, K.; Aucher, W.; Brosset-Vincent, S.; Kadri, L.; Colas, J.; Bouchon, D.; Ferreira, T. Fate of eight different polymers under uncontrolled composting conditions: Relationships between deterioration, biofilm formation, and the material surface properties. Environ. Sci. Technol. 2017, 51, 1988−1997. (82) Oberbeckmann, S.; Osborn, A. M.; Duhaime, M. B. Microbes on a bottle: Substrate, season and geography influence community composition of microbes colonizing marine plastic debris. PLoS One 2016, 11, No. e0159289. (83) Atashgahi, S.; Sanchez-Andrea, I.; Heipieper, H. J.; van der Meer, J. R.; Stams, A. J. M.; Smidt, H. Prospects for harnessing biocide resistance for bioremediation and detoxification. Science 2018, 360, 743−746. 4104

DOI: 10.1021/acscatal.8b05171 ACS Catal. 2019, 9, 4089−4105

Perspective

ACS Catalysis (84) Uchida, H.; Shigeno-Akutsu, Y.; Nomura, N.; Nakahara, T.; Nakajima-Kambe, T. Cloning and sequence analysis of poly(tetramethylene succinate) depolymerase from Acidovorax delafieldii strain BS-3. J. Biosci. Bioeng. 2002, 93, 245−247. (85) Tang, B.; Yu, Y.; Zhang, Y.; Zhao, G.; Ding, X. Complete genome sequence of the glidobactin producing strain [Polyangium brachysporum DSM 7029. J. Biotechnol. 2015, 210, 83−84. (86) Anantharaman, K.; Brown, C. T.; Hug, L. A.; Sharon, I.; Castelle, C. J.; Probst, A. J.; Thomas, B. C.; Singh, A.; Wilkins, M. J.; Karaoz, U.; Brodie, E. L.; Williams, K. H.; Hubbard, S. S.; Banfield, J. F. Thousands of microbial genomes shed light on interconnected biogeochemical processes in an aquifer system. Nat. Commun. 2016, 7, 13219. (87) Yang, D.; Cho, J. S.; Choi, K. R.; Kim, H. U.; Lee, S. Y. Systems metabolic engineering as an enabling technology in accomplishing sustainable development goals. Microb. Biotechnol. 2017, 10, 1254− 1258. (88) Dvorak, P.; Nikel, P. I.; Damborsky, J.; de Lorenzo, V. Bioremediation 3.0: Engineering pollutant-removing bacteria in the times of systemic biology. Biotechnol. Adv. 2017, 35, 845−866. (89) Seffernick, J. L.; de Souza, M. L.; Sadowsky, M. J.; Wackett, L. P. Melamine deaminase and atrazine chlorohydrolase: 98% identical but functionally different. J. Bacteriol. 2001, 183, 2405−2410. (90) Long, S. P.; Ainsworth, E. A.; Leakey, A. D.; Nosberger, J.; Ort, D. R. Food for thought: lower-than-expected crop yield stimulation with rising CO2 concentrations. Science 2006, 312, 1918−1921. (91) Schimel, D. Ecology Climate change and crop yields: beyond Cassandra. Science 2006, 312, 1889−1890. (92) Ewert, F.; Porter, J. R.; Rounsevell, M. D. Crop models, CO2, and climate change. Science 2007, 315, 459c. (93) Sakurai, G.; Iizumi, T.; Nishimori, M.; Yokozawa, M. How much has the increase in atmospheric CO2 directly affected past soybean production? Sci. Rep. 2015, 4, 4978. (94) del Hoyo-Gallego, S.; Perez-Alvarez, L.; Gomez-Galvan, F.; Lizundia, E.; Kuritka, I.; Sedlarik, V.; Laza, J. M.; Vila-Vilela, J. L. Construction of antibacterial poly(ethylene terephthalate) films via layer by layer assembly of chitosan and hyaluronic acid. Carbohydr. Polym. 2016, 143, 35−43. (95) Phaneuf, M. D.; Berceli, S. A.; Bide, M. J.; Quist, W. C.; LoGerfo, F. W. Covalent linkage of recombinant hirudin to poly(ethylene terephthalate) (Dacron): creation of a novel antithrombin surface. Biomaterials 1997, 18, 755−765. (96) Kulik, E. A.; Kato, K.; Ivanchenko, M. I.; Ikada, Y. Trypsin immobilization on to polymer surface through grafted layer and its reaction with inhibitors. Biomaterials 1993, 14, 763−769. (97) Kucherov, F. A.; Romashov, L. V.; Galkin, K. I.; Ananikov, V. P. Chemical transformations of biomass-derived C6-furanic platform chemicals for sustainable energy research, materials science, and synthetic building blocks. ACS Sustainable Chem. Eng. 2018, 6, 8064− 8092. (98) Zhu, J. B.; Watson, E. M.; Tang, J.; Chen, E. Y. A synthetic polymer system with repeatable chemical recyclability. Science 2018, 360, 398−403. (99) Pellis, A.; Haernvall, K.; Pichler, C. M.; Ghazaryan, G.; Breinbauer, R.; Guebitz, G. M. Enzymatic hydrolysis of poly(ethylene furanoate). J. Biotechnol. 2016, 235, 47−53.

4105

DOI: 10.1021/acscatal.8b05171 ACS Catal. 2019, 9, 4089−4105