Biotechnology Advances 31 (2013) 1754–1767

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Research review paper Cutinase: Characteristics, preparation, and application

Sheng Chen, Lingqia Su, Jian Chen, Jing Wu ⁎

State Key Laboratory of Food Science and Technology, Jiangnan University, 1800 Lihu Ave., Wuxi, Jiangsu 214122, China School of Biotechnology and Key Laboratory of Industrial Biotechnology, Ministry of Education, Jiangnan University, 1800 Lihu Ave., Wuxi, Jiangsu 214122, China article info abstract

Article history: Cutinases (E.C. 3.1.1.74) belong to the α/β-hydrolase superfamily. They were initially discovered because they Received 15 May 2013 are secreted by fungi to hydrolyze the ester bonds of the plant polymer cutin. Since then, they have been Received in revised form 4 August 2013 shown to catalyze the hydrolysis of a variety of polymers, insoluble triacylglycerols, and low-molecular-weight Accepted 11 September 2013 soluble esters. Cutinases are also capable of catalyzing esterification and transesterification reactions. These Available online 19 September 2013 relatively small, versatile, secreted catalysts have shown promise in a number of industrial applications. This re- view begins by describing the characteristics of cutinases, pointing out key differences among cutinases, esterases Keywords: Cutinase and lipases, and reviewing recent progress in engineering improved cutinases. It continues with a review of the Identification methods used to produce cutinases, with the goal of obtaining sufficient quantities of material for use in indus- Crystal structure trial processes. Finally, the uses of cutinases in the textile industry are described. The studies presented here dem- Molecular modification onstrate that the cutinases are poised to become important industrial catalysts, replacing older technologies with Preparation more environmentally friendly processes. Textile industry © 2013 Elsevier Inc. All rights reserved.

Contents

1. Introduction...... 1754 2. Cutinase identification, structure and modification...... 1755 2.1. Identificationoftruecutinases...... 1755 2.2. Cutinaseassays...... 1756 2.3. Simplerassaysappliedtocutinases...... 1757 2.4. Cutinasestructureandfunction...... 1758 2.5. Modificationofcutinases...... 1760 3. Preparationofcutinase...... 1761 3.1. Preparationofcutinasesfromwild-typestrains...... 1761 3.2. Preparationofcutinaseusingengineeredstrains...... 1762 4. Applicationofcutinaseinindustry...... 1763 4.1. Cotton fibers...... 1763 4.2. Synthetic fibers...... 1763 4.3. Woolfabrics...... 1764 5. Conclusions...... 1764 Acknowledgments...... 1764 References...... 1764

1. Introduction Abbreviations: CBM, carbohydrate-binding module; DHA, docosahexanoic acid; EPA, eicosapentanoic acid; PBM, polyhydroxyalkanoate-binding module; PET, polyethylene The plant cuticle is a protective layer that coats the epidermis of terephthalate; RBB, Remazol Brilliant Blue R. leaves, shoots, and other tender, aboveground portions of terrestrial ⁎ Corresponding author at: State Key Laboratory of Food Science and Technology, plants. Composed of waxes and lipid polymers, the cuticle protects the Jiangnan University, 1800 Lihu Ave., Wuxi, Jiangsu 214122, China. Tel.: +86 510 85327802; fax: +86 510 85326653. plant from dehydration, and is a barrier to infection by pathogen. The E-mail address: [email protected] (J. Wu). major constituent of the cuticle is insoluble lipid polyester called cutin,

0734-9750/$ – see front matter © 2013 Elsevier Inc. All rights reserved. http://dx.doi.org/10.1016/j.biotechadv.2013.09.005 S. Chen et al. / Biotechnology Advances 31 (2013) 1754–1767 1755 which consists primarily of hydroxylated 16- and 18-carbon fatty acids 2. Cutinase identification, structure and modification that are linked together via ester bonds. Microorgaisms, principally fungi and bacteria, can hydrolyze this polymer by secreting enzymes 2.1. Identification of true cutinases called cutinases. Early work focused on the role of cutinase in the infection of plants Cutinase genes were originally identified by observing the effect of by fungi. Fungal spores landing on the plant cuticle were shown to their deletion upon the virulence of the expressing organism. Although respond to cutin monomers by expressing cutinase, and specificinhibi- originally found in pathogenic fungi, cutinases have subsequently been tion of this enzyme blocked infectivity in several pathogen/host systems isolated and characterized from bacteria, such as Thermobifida fusca (Kolattukudy et al., 1989). Subsequent work with Alternaria brassicicola, (Chen et al., 2008), and from a variety of both pathogenic and non- demonstrated that cutinases are a diverse group of enzymes, in which pathogenic fungi, including the yeast Cryptococcus (Masaki et al., some cutinases are essential for pathogenicity, while others are 2005). The microbial strains that have been shown to express cutinases expressed during saprophytic growth on cutin as a carbon source (Fan are identified in Table 1. As the number of sequenced microbial and Köller, 1998; St Leger et al., 1997). More recent research has genomes has increased, so has the number of open reading frames demonstrated that both pathogenic and saprophytic microorganisms annotated as cutinases. In several instances, these putative cutinase express cutinases, that the genomes of individual species may harbor enzymes have been isolated and assayed using simple substrates. Un- several different putative cutinases, and that these enzymes are fortunately, since many of these enzymes have not been assayed using expressed at different times during the microbial life cycle (Skamnioti cutin as a substrate, it is not possible to distinguish them from similar et al., 2008a, 2008b). There is also evidence suggesting that cutinase- esterases or lipases that can also hydrolyze these simple substrates. To containing saprophytic bacteria, such as Pseudomonas putida, may pro- appreciate the number and diversity of the open reading frames that vide a carbon source to a wider microbial community (Sebastian et al., have been annotated as cutinases, and appreciate how varied the activ- 1987). ities of the proteins expressed from these genes are likely to be, studies Cutinases (EC 3.1.1.74) are serine esterases that belong to the α/β hy- of the molecular taxonomy of cutinases must be reviewed. drolase superfamily. They possess a classical Ser–His–Asp catalytic triad, Relatively recent work on the molecular taxonomy of fungal in which the catalytic serine is exposed to solvent. Because cutinases lack cutinases (Skamnioti et al., 2008a, 2008b) has suggested that the fungal the hydrophobic lid that covers the active site serine in true lipases, the cutinases can be divided into two ancient subfamilies that have been in cutinase active site is large enough to accommodate the high- existence since before the Ascomycota and the Basidobycota diverged molecular-weight substrate cutin, and some of them can also hydrolyse nearly a billion years ago. Interestingly, no cutinase sequences were high-molecular-weight synthetic polyesters. Besides, they are able to found in the “true yeasts” (Saccharomycotina). A similar study of the hydrolyse a greater variety of substrates, including low-molecular- cutinase sequences from Phytophthora, a genus of phytopathogenic weight soluble esters, short- and long-chain triacylglycerols. Cutinases oomycetes (water molds) (Belbahri et al., 2008) identified related are also capable of catalyzing esterification and transesterification. cutinase sequences in three genera of Actinobacteria, and suggested First identified in the 1960's and characterized in the early 1970's, lateral gene transfer between ancient bacteria and oomycetes. the cutinase from the filamentous fungus Fusarium solani pisi rapidly be- Skamnioti et al. (Skamnioti et al., 2008a) recognized the existence of came a model system for the study of cutinase structure, function and mycobacterial cutinases, but rejected the idea of lateral gene transfer reactivity. This early work has been described in concise reviews to fungi, placing the bacterial cutinase sequences in a separate group. (Carvalho et al., 1998; Egmond and de Vlieg, 2000; Longhi and Both research groups recognized that a single species may contain Cambillau, 1999). Cutinase was found to be distinct among the α/β several putative cutinases, sometimes more than a dozen. Skamnioti hydrolases because, unlike the majority of esterases (Panda and et al. (Skamnioti et al., 2008a) also performed a transcriptional analysis Gowrishankar, 2005), cutinase is able to hydrolyse lipid substrates ofthe14putativecutinasesfromMagnaporthe grisea strain Guy11. and, unlike typical lipases (Sharma et al., 2001), cutinase activity is not They found four of them to be constitutively expressed, including the activated by interfacial effects—it efficiently hydrolyzes hydrophobic previously characterized cutinase CUT1 (Sweigard et al., 1992). One substrates in solution or in emulsions. These properties, which can be gene showed elevated expression only during early differentiation. Six used to identify the true cutinases, blur the distinction between genes, including the previously described CUT2 (Skamnioti and Gurr, esterases and lipases to the point that some have argued for a new 2007), showed elevated expression during plant penetration. Finally, bio-physico-chemical classification system (Ben Ali et al., 2012). This re- three genes showed elevated expression in planta.Sincetherewasno view will begin by presenting recent progress concerning the identifica- correlation between the phylogenetic closeness of the putative cutinases tion and characterization of cutinases, highlighting the structural and their expression patterns, these authors concluded that there has characteristics and catalytic activities that distinguish cutinases from been considerable subfunctionalization and neofunctionalization related enzymes. among the putative cutinases. It is possible, therefore, that the The versatile catalytic ability of cutinase led to the early recognition cutinase-like proteins from the important human bacterial pathogen that these enzymes were potentially useful for a variety of industrial Mycobacterium tuberculosis, which share the high sequence homology applications (Carvalho et al., 1999). This recognition of the potential with fungal cutinase and exhibit strong esterase activity, do not possess value of these enzymes has resulted in a significant amount of effort cutinase activity (West et al., 2009). being devoted to the discovery and characterization of cutinases The Genbank accession numbers of the genes that have been shown and “cutinase-like” enzymes, as well as systems suitable for the to encode true cutinases are presented in Table 1. Aligning all of the true industrial-scale production of these enzymes. Additional research has cutinase sequences allows the construction of a phylogenetic tree of been undertaken to alter the substrate specificity and improve the true cutinases (Fig. 1). This analysis reveals that the prokaryotic stability of these enzymes. A brief review of the production of cutinases cutinases are distinct from the eukaryotic cutinases. Their low sequence and their application to the detergent and laundry industry, the fruit identity indicates that they have undergone extensive evolutionary dif- industry, and the production of esters appeared relatively recently ferentiation. The eukaryotic family appears to fall into three subgroups, (Dutta et al., 2009). There have also been two monographs: one detail- two of which contain fungal sequences, and the other of which contains ing the use of cutinases in producing chemicals and biofuels (Baker the true cutinases from yeast-like fungi (Cryptococcus sp. and and Montclare, 2010), and the other giving a more general overview Pseudozyma antarctica). The two fungal subgroups share roughly 30% of the industrial applications of cutinases (Pio and Macedo, 2009). similarity, while the yeast-like sequences shares approximately 19% This review will focus on the applications of cutinases in the textile similarity with the two fungal groups. Although it is difficult to make industry. generalizations based upon a few sequences, it appears that the 1756 S. Chen et al. / Biotechnology Advances 31 (2013) 1754–1767

Table 1 Microbial cutinases.

Organism Putative cutinasesa Genbank accessionb References

Fungi Fusarium solani pisi 2 AAA33334.1 Lin and Kolattukudy (1980b), Martinez et al. (1992), Purdy and Kolattukudy (1975), Soliday et al. (1984), van Gemeren et al. (1995) Fusarium oxysporum 1 – Nimchua et al. (2007), Rocha et al. (2008) Fusarium roseum culmorum –– Lin and Kolattukudy (1980a), Soliday and Kolattukudy (1976) Fusarium roseum sambucinum –– Lin and Kolattukudy (1980a) Ulocladium consortiale –– Lin and Kolattukudy (1980a) Helminthosporium sativum –– Lin and Kolattukudy (1980a) Aspergillus nidulans 4 ABF50887.1 Castro-Ochoa et al. (2012) Aspergillus niger 5 CAL00335.1 Nyyssola et al. (2013) Monilinia fructicola 4 AAZ95012.1 Lee et al. (2010), Wang et al. (2000, 2002) Alternaria brassicicola 4 – Koschorreck et al. (2010), Trail and Köller (1993), Yao and Köller (1994), Yao and Köller (1995) Pyrenopeziza brassicae 1 CAB40372.1 Davies et al. (2000), Li et al. (2003) Botrytis cinerea 1 CAA93255.1 Shishiyama et al. (1970), van der Vlugt-Bergmans et al. (1997) Botrytis cinerea (ungerminated conidia) 1 – Gindro and Pezet (1997), Gindro and Pezet (1999) Magnaporthe grisea 5 EHA46959.1 Sweigard et al. (1992) Colletotrichum gloeosporioides 1 – Z. Chen et al. (2007), Dickman et al. (1982), Ettinger et al. (1987) Cochliobolus heterostrophus –– Trail and Köller (1990) Rhizoctonia solani –– Trail and Köller (1990) Venturia inaequalis –– Köller and Parker (1989) Penicillium sp. 1 – Nimchua et al. (2008) Penicillium citrinum –– Liebminger et al. (2007) Aspergillus oryzae 2 BAA07428.1 Liu et al. (2009), Ohnishi et al. (1995) Pseudozyma antarctica – GAC73680.1 Shinozaki et al. (2013a, 2013b), Suzuki et al. (2012) Thielavia terrestris –– Yang et al. (2013) Coprinopsis cinerea 3 EU435153.1 Kontkanen et al. (2009) Humilica insolens –– Ronkvist et al. (2009)

Bacteria Streptomyces scabies –– Lin and Kolattukudy (1980a) Streptomyces acidiscabies –– Fett et al. (1992a) Streptomyces badius –– Fett et al. (1992a) Thermobifida fusca 2 AAZ54920.1 Chen et al. (2008), Dresler et al. (2006), Fett et al. (1999), Kleeberg et al. (1998), AAZ54921.1 Kleeberg et al. (2005) Thermobifida alba 2 ADV92525.1 Hu et al. (2010), Thumarat et al. (2012) Thermobifida cellulosilytica 2 ADV92527.1 Herrero Acero et al. (2013), Herrero Acero et al. (2011) ADV92526.1 Thermoactinomyces vulgaris –– Fett et al. (2000) Pseudomonas putida –– Sebastian et al. (1987), Sebastian and Kolattukudy (1988) Pseudomonas aeruginosa –– Fett et al. (1992b) Pseudomonas mendocina –– Degani et al. (2006), Degani et al. (2002), Ronkvist et al. (2009)

Yeast Cryptococcus sp. 1 BAK82405.1 Masaki et al. (2005)

a Number of enzymes suggested to be cutinases, but not all have demonstrated cutinase activity. b Genbank accession numbers for enzymes with demonstrated cutinase activity. known true cutinases provide a diverse sample of the cutinase families with the chromogenic compound Remazol Brilliant Blue R (RBB). In identified through broad phylogenetic analysis of genome sequences. this assay, cutinase activity is quantified by measuring the increase in In summary, genome sequencing has led to the annotation of a sub- absorption at 590 nm caused by the release of RBB-labeled material stantial number of open reading frames as cutinases. Phylogenetic anal- from the insoluble substrate into the solution (van der Vlugt- ysis has revealed two fungal subfamilies and at least one bacterial family Bergmans et al., 1997). Because labeled cutin is difficult to obtain, a of cutinase or cutinase-like proteins. Given the diversity of transcrip- method that uses native cutin as a substrate was developed. In this tional regulation seen in M. grisea,itseemsthatitwouldbebestto method, the cutin monomers can be determined using thin-layer adopt a practical means of identifying a true cutinase—demonstrating chromatography combined with infrared spectroscopy (Castro-Ochoa the ability of an isolated protein to hydrolyse cutin, or a suitable et al., 2012), or the hydroxylated fatty acids released by cutinase can substitute. be determined using gas chromatography/mass spectrometry (Chen et al., 2008; Fett et al., 1992a, 1992b, 1999, 2000; Kontkanen et al., 2.2. Cutinase assays 2009). Unfortunately, natural cutin is not commercially available. It must be prepared by individual laboratories through a tedious process The classical method to measure cutinase activity is to use cutin as that involves organic solvent extraction, followed by treatment with the substrate and detect the cutin monomers released by hydrolysis. pectinase, cellulase, and lipase to remove contaminants (Walton and This has been done using radiolabeled fruit cutin as the substrate and Kolattukudy, 1972). determining the radioactive cutin monomers by thin-layer chromatog- Fortunately, there are synthetic alternatives that are similar enough raphy (Davies et al., 2000; Dickman et al., 1982; Gindro and Pezet, 1997; to the natural substrate to be used in its place, and are much easier to ob- Köller and Parker, 1989; Lin and Kolattukudy, 1980b; Purdy and tain. These include high-molecular-weight synthetic polyesters (weight- Kolattukudy, 1975; Sebastian et al., 1987; Soliday and Kolattukudy, average MW ≥1.0 × 105 g/mol), such as polyethylene terephthalate 1976; Sweigard et al., 1992; Trail and Köller, 1990; Wang et al., 2000; (PET) fibers (Nimchua et al., 2007, 2008), poly(ε-caprolactone) (Liu Yao and Köller, 1995) or liquid scintillation spectrometry (Sebastian et al., 2009), polylactic acid, and poly(butylene succinate) (Masaki et al., 1987). As an alternative, cutin can be labeled non-specifically et al., 2005; Thumarat et al., 2012). Using PET as a substrate, cutinase S. Chen et al. / Biotechnology Advances 31 (2013) 1754–1767 1757

Fig. 1. Phylogenetic analysis of cutinase genes. The species and Genbank accession numbers of the true cutinases from Table 1 are arranged on a phylogenetic tree determined using the program MEGA4.

activity can be quantified by measuring the fluorescence of terephthalic the putative cutinases from Fusarium oxysporum (ABR19840.1), acid released from the polymer (Nimchua et al., 2007). When using A. brassicicola (AAA03470.1) and Colletotrichum gloeosporioides poly(ε-caprolactone), polylactic acid, and poly(butylene succinate) as (AAA33042.1), were excluded from Table 1 because their cutinase activity substrates, cutinase activity must be evaluated by monitoring the de- has not been confirmed. Their absence from Table 1, however, does not crease in absorbance at 660 nm after the addition of the enzyme. The ac- mean that these enzymes are not potentially useful in industrial applica- tivity is analyzed by plotting the fraction of undegraded polymer (called tions, nor does it mean that the simple assays used to measure their activ- the ratio of undegraded polymer) versus time (Masaki et al., 2005). ity are not useful in characterizing the activity of true cutinases. Perhaps the simplest assay used with cutinases is measuring their 2.3. Simpler assays applied to cutinases esterase activity by monitoring the hydrolysis of 4-nitrophenyl esters. The reaction can be followed by measuring the absorbance at 405 nm, Some well-studied strains that produce enzymes with substantial which results from the release of 4-nitrophenol. Several of these sequence identity to known cutinases, including Colletotrichum 4-nitrophenyl esters are readily available from commercial sources, in- acutatum (Liao et al., 2012), Colletotrichum kahawae (Z. Chen et al., cluding 4-nitrophenyl acetate, butyrate, valerate, and octanoate, and 2007), Glomerella cingulata (Bakar et al., 2001, 2005; Nyon et al., many of these have been used to assay putative cutinases. An insoluble 2009), and Trichoderma harzianum (Rubio et al., 2008) are not included 4-nitrophenyl ester that is reported to be resistant to hydrolysis by por- in Table 1. The reason for their omission is that neither the strains nor cine liver esterase has also been designed and synthesized. This unique the putative cutinases from them have been assayed using cutin, or synthetic substrate may be helpful for detecting and assaying cutinase other polyesters, as the substrate. Some of the strains listed in Table 1 activity in mixed solutions (Degani et al., 2006). are known to secrete cutinases, but the number of cutinases they The kinetics of 4-nitrophenyl acetate, butyrate, valerate and secrete is unknown because of a lack of annotated sequence information. hexanoate hydrolysis by F. solani pisi and Aspergillus oryzae cutinases In these cases, the Genbank accession numbers of putative cutinases iso- (Liu et al., 2009) revealed that F. solani pisi cutinase prefers −1 −1 lated from the strains, if any, are absent because of a lack of appropriate 4-nitrophenyl acetate (kcat/Km = 42,200 M s ) over the others by experimental evidence. For example, the Genbank accession numbers of about an order of magnitude, while A. oryzae cutinase prefers 4- 1758 S. Chen et al. / Biotechnology Advances 31 (2013) 1754–1767

−1 −1 nitrophenyl butyrate (kcat/Km = 59,200 M s ) or 4-nitrophenyl val- of the gel. Screening of organisms that produce cutinases has been −1 −1 erate (kcat/Km = 55,300 M s ). The kinetics parameters of important performed in media containing cutin as the sole carbon source bacterial cutinases from Thermobifida have also been studied. The (Dickman and Patil, 1986). In this screening assay, plugs of agar contain- cutinases from T. cellulosilytica (Thc_Cut1 and Thc_Cut2), T. fusca ing samples of the fungi in question were transferred to Czapek-Dox DSM44342 (Thf42_Cut1) and T. alba all prefer 4-nitrophenyl acetate, medium containing papaya cutin as the sole carbon source and cresol −1 with Kms of 127, 200, 167, 213 μMandkcats of 211.9, 2.4, 39.5, 2.72 s , red or phenol red as pH indicator dyes. Zones of yellow color developed respectively (Herrero Acero et al., 2011, 2013; Ribitsch et al., 2012b). around agar plugs containing organisms that express cutinases. Among the 4-nitrophenyl esters, the butyrate seems to be the most wide- Adapting a similar strategy, organisms suspected of expressing extracel- ly used. It has, for example, been used as the substrate for a 96-well lular cutinases have been grown on agar plates containing the triacyl- screening assay for cutinase inhibition (Walz and Schwack, 2007a), glycerol derivative triacetin and fluorescent dye Rodamin B (Macedo which was developed as part of a program to measure the levels of organ- and Pio, 2005). Colonies expressing esterases develop a fluorescent ophosphorus insecticides in plant tissues (Walz and Schwack, 2007b). halo. The simple 4-nitrophenyl esters are convenient substrates, but they are highly susceptible to impurities in the enzyme preparation, since 2.4. Cutinase structure and function many esterases are capable of hydrolyzing them. A somewhat more se- lective alternative may be to use triacylglycerols as substrates. This sub- The first X-ray crystal structure of the cutinase from F. solani pisi strate is, however, typically used to characterize lipases. Therefore, (PDB ID: 1CUS) was described in 1992 (Martinez et al., 1992). This although these assays can be used to characterize the activity of structure revealed that F. solani pisi cutinase adopts an α/β fold that cutinases, they cannot be used to demonstrate that an enzyme is a exposes the catalytic serine (Ser 120) to the solvent, instead of burying cutinase. In addition, this type of assay is susceptible to lipases that the active site with a hydrophobic loop. Since the increased activity that may be impurities in the cutinase preparation. In these assays, commer- lipases display at aqueous/lipid interfaces (interfacial activation) in- cially available substrates like tributyrin (glycerol tributyrate) are volves the interaction of these hydrophobic loops with the interface hydrolyzed by the cutinase to produce butyric acid and a diacylglycerol. (Cambillau et al., 1996), their absence in cutinases was cited as the The liberated acid perturbs the pH, which is maintained using a pH-stat. reason for their lack of interfacial activation. Since then, X-ray crystal Enzyme activity is proportional to the amount of base added to maintain structures of a significant number of variants of this protein, including the pH, which is usually around pH 8.0. Since tracylglycerol substrates inhibited complexes, have been deposited. The structure–activity rela- have limited solubility, bile acids (e.g. sodium taruodeoxycholate) are tionships revealed by these many structures have been previously frequently added, and the assay is run as an emulsion rather than a so- reviewed (Egmond and de Vlieg, 2000; Longhi and Cambillau, 1999). lution. Early experiments using the simple triacylglycerols tributyrin, In 2009, the X-ray crystal structures of A. oryzae (PDB ID: 3GBS; (Liu trioctanoin and triolein (Rogalska et al., 1993) or other triacylglycerol et al., 2009)) and C. sp. (PDB ID: 2CZQ; (Kodama et al., 2009)) were analogs (Mannesse et al., 1995)assubstratesshowedthatF. solani pisi solved. This was followed in 2012 by the structure of the cutinase cutinase prefers substrates with short acyl groups. The specificactivity from the bacterium Thermobifida alba (PDB ID: 3VIS; (Kitadokoro of F. solani pisi cutinase with tributyrin, for example, was reported to et al., 2012)). Characteristics of these structures are presented in be twice its specific activity with trioctanoin (Kwon et al., 2009; Table 2. Additional structures of proteins annotated as cutinases are de- Rogalska et al., 1993). Tributryin, which is commercially available, is posited in the RCSB PDB, including three structures of a putative perhaps the best choice of substrate for assays of this type. cutinase from the pathogenic fungus G. cingulata (PDB IDs: 3DCN, Finally, there have been efforts to adapt forms of zymography devel- 3DD5 and 3DEA; (Nyon et al., 2009)), five structures containing F. solani oped to identify esterases and lipases for the identification of cutinases pisi cutinase that has been modified with organometallic complexes in PAGE gels or on agar plates. To detect the esterase activity of cutinases (PDB IDs: 3EF3, 3ESA - 3ESD; (Rutten et al., 2009)), the structure of an in an SDS-PAGE gel, the proteins can be renatured in a solution of buffer essential lipase from Mycobacterium smegmatis (PDB ID: 3AJA), and and Triton X-100, and then incubated with α-naphthyl acetate and Fast the structure of a “cutinase homolog” from an uncharacterized bacterial Red TR, both of which are commercially available. Esterases produce a species obtained from leaf-branch compost (PDB ID: 4EBO). purple-colored band in the gel (Castro-Ochoa et al., 2012; Karpushova Cutinases, like other α/β serine hydrolases, perform their catalysis in et al., 2005; Yang et al., 2013). The lipase activity of cutinases in SDS- two discrete steps, with a covalent intermediate that links the catalytic PAGE gels has been detected using the commercially available serine to the carbonyl group of the ester being hydrolyzed (Jaeger et al., fluorogenic reagents methyumbelliferyl butyrate or methyumbelliferyl 1994). Soon after the first structure of F. solani pisi cutinase was pub- oleate (Prim et al., 2003; Yang et al., 2013). Hydrolysis of these reagents lished, a second structure (PDB ID: 2CUT, (Martinez et al., 1994)) produces the fluorescent molecule methylumbeliferone (excitation appeared that contained the covalent intermediate from the hydrolysis wavelength 323 nm, emission wavelength 448 nm), which can be de- of diethyl-p-nitrophenyl phosphate. This structure (Fig. 2a) helped tected in the gel using a simple UV lamp. An alternative system uses identify the catalytic triad (S120, H188, D175) and key residues in the the ester synthesis activity of the cutinase to combine a fatty acid oxyanion hole (S42 and Q121) that are important for stabilizing the (oleic acid or caprylic acid) with an alcohol (dodecanol) to form an in- transitions states in the acylation/deacylation steps of the enzyme soluble ester (Kwon et al., 2011), which causes an opaque coloration mechanism (Baker and Montclare, 2010). Alignment of the structures

Table 2 Characteristics of available cutinase structures.

Organism PDB ID Catalytic triad Oxyanion binding residues Disulfide bonds Resolution (Å) References

F. solani pisi variousa S120/H188/D175 S42/Q121 C31–C109 various Egmond and De Vlieg (2000) C171–C178 C. sp. 2CZQ S85/H180/D165 T17/Q86 C6–C78 1.05 Kodama et al. (2009) C161–C168 A. oryzae 3GBS, 3PQD S126/H194/D181 S48/Q127 C63–C76 1.75 Liu et al. (2009) C37–C115 C177–C184 T. alba 3VIS S169/H247/D215 M179/Y99 C280–C298 1.76 Kitadokoro et al. (2012)

a F. solani pisi structures include PDB IDs: 1AGY, 1CEX, 1OXM, 1XZA–1XZM, 1CUA–1CUJ, 1CUS, 1CUU–1CUZ, 1FFA–1FFE, 2CUT, 3QPC. Reference is to review discussing these structures. S. Chen et al. / Biotechnology Advances 31 (2013) 1754–1767 1759

Fig. 2. Cutinase active sites. a) Fusarium solani pisi cutinase (PDB ID: 2CUT (Martinez et al., 1994); gray ribbon) with a molecule of diethyl phosphate (black) bound to the catalytic Ser (red). The side chains of the catalytic His (green) and Asp (blue), along with the anion hole Ser (cyan) and Gln (magenta) are rendered as sticks. Putative hydrogen bonds are shown as black dashed lines. b) Overlay of aligned structures of F. solani pisi (PDB ID: 1CUS (Martinez et al., 1992); slate blue ribbon) and Cryptococcus sp. (PDB ID: 2CZQ (Kodama et al., 2009); gray rib- bon) cutinases. Side chains of the catalytic Ser (red), His (green) and Asp (blue), as well as the oxyanion hole residues Ser (1CUS) or Thr (2CZQ) (cyan) and Gln (magenta) are rendered as sticks. Putative hydrogen bonds are shown as black dashed lines. c) Active site of Thermobifida alba cutinase (PDB ID: 3VIS (Kitadokoro et al., 2012)). Side chains of the catalytic Ser (red), His (green) anad Asp (blue) are rendered as sticks, as are the side chains of the putative anion hole residues Met and Tyr (on right). Putative hydrogen bonds are shown as dashed black lines. All renderings were performed using PyMol.

of the cutinases from F. solani pisi and A. oryzae, using the jFATCAT flex- cutinases (serine/threonine and glutamine). Of course, as with F. solani ible algorithm at the RSCB PDB, reveals that these two structures are ex- pisi cutinase, the main-chain amides are thought to be responsible for tremely similar; they share 50% identity, and their alpha carbons display stabilizing the oxyanion (Kitadokoro et al., 2012). In addition to the cat- a root mean square deviation of 0.83 Å. In contrast, comparison of the alytic triad and the oxyanion hole, all these cutinases contain at least F. solani pisi cutinase with the phylogenetically more distant cutinase one disulfide bond, which is assumed to play an important role in stabi- from C. sp. yields a much lower identity (18.9%) and much larger root lizing the structure. However, the thermostability is not dependent on mean square deviation (3.20 Å). However, overlay of the two structures the number of disulfide bonds. For example, the T. alba cutinase, (PDB IDs: 1CUS and 2CZQ) using PyMol (Fig. 2b) demonstrates that the which contains only one disulfide bond, is much more thermostable catalytic triad of the C. sp. enzyme (S85/H180/D165) overlays very than either the F. solani pisi or A. oryzae cutinase, which have two and closely with that of the F. solani pisi enzyme, as do the key residues of three disulfide bonds, respectively. The thermostability of the T. alba the oxyanion hole (T17 and Q86). Note that the oxygen atoms of the cutinase is considerably enhanced in the presence of Ca2+ (Thumarat serine and threonine residues in the oxyanion hole are not thought to et al., 2012). interact with the oxyanion. Instead, it is the main-chain amides that sta- The T. alba enzyme has been shown to hydrolyze synthetic polyesters, bilize this negative charge through hydrogen bonding. The cutinase including Ecoflex™, poly(caprolactone), poly(butylene succinate-co- from T. alba also adopts an α/β fold, but it is larger than the others. It adipate), poly(butylene succinate), poly(L-lactic acid) and poly(D-lactic contains nine sheets at the heart of the protein, two of which are anti- acid) but not poly(3-hydroxybutyric acid) (Hu et al., 2010; Thumarat parallel, rather than the five parallel sheets present in the fungal et al., 2012). Its closest structural homolog in the PDB is the Streptomyces enzymes. Like the fungal cutinases, it contains a serine–histidine– exfoliatus lipase (PDB ID: 1JFR). This points to the fact that cutinases, aspartic acid catalytic triad (Fig. 2c), and the catalytic serine is accessible though easy to distinguish from esterases because they can hydrolyze in- to the solvent. However, the residues found in the oxyanion hole are soluble substrates, are more difficult to distinguish from lipases. The dif- quite different (methionine and tyrosine) than those found in the fungal ference between cutinases and lipases lies in the preference of lipases

Fig. 3. Ribbon diagrams of Rhizomucor miehi lipase showing the conformational change involved in catalysis. a) Native enzyme structure (PDB ID: 5TGL (Brzozowski et al., 1991; Derewenda et al., 1992); slate blue ribbon) showing catalytic triad residues Ser (red), His (green), and Asp (blue). The helix capping the active site is depicted as a magenta ribbon. b) Structure of the enzyme (PDB ID: 4TGL (Brzozowski et al., 1991; Derewenda et al., 1992); slate blue ribbon) inhibited by diethyl-4-nitrophenyl phosphate in the activated (open) confor- mation, showing the diethyl phosphate inhibitor (black) and the catalytic triad's Ser (red), His (green) and Asp (blue). The helix that capped the active site in the native structure (magenta ribbon) has moved away from the active site, opening it to the solvent. All renderings were performed using PyMol. 1760 S. Chen et al. / Biotechnology Advances 31 (2013) 1754–1767 for much longer chain lengths, and the interfacial activation that is typical 2.5. Modification of cutinases of true lipases. On the structural level, this difference is evidenced by the exposure Molecular modification technology has been widely used to investi- of the active site of the cutinases to solvent, and the presence of a hydro- gate the catalytic mechanisms and structural specificities of α/β hydro- phobic lid structure that covers the active site serine of the lipases. The lases, reviewed in (Jochens et al., 2011), including cutinases. Many conformational change involved in interfacial catalysis has been docu- site-direct mutants were constructed in an effort to fully understand mented using a triacylglyceride lipase from the fungus Rhizomucor the catalytic mechanism of F. solani pisi cutinase. By combining kinetic miehi (Brzozowski et al., 1991; Derewenda et al., 1992). In the native experiments with structural analysis, the residues involved in the structure (Fig 3a), a helix blocks solvent access to the active site. Inhibi- chemical catalysis, substrate binding and the oxyanion hole were con- tion of the lipase with diethyl-4-nitrophenyl phosphate causes this helix firmed (Table 3). to move almost 12 Å, and leads to an open form (Fig. 3b) that exposes a Based upon this model, amino acids in the substrate-binding region hydrophobic area of 800 Å2. It is thought that interaction of this surface were replaced with hydrophobic residues to increase the interaction be- with the lipid interface allows the lipid substrates access to the active tween these hydrophobic residues and the substrate. The variants A85F, site. A85W, T173K, T179Y and L189F all showed higher enzyme activity with Finally, given these structural differences between cutinases and li- hydrophobic, low-molecular-weight substrates in olive oil emulsions pases, the structure of a putative cutinase from G. cingulata should be (Egmond and De Vlieg, 2000). Sequence alignment with the cutinase examined. Three structures of this enzyme have been deposited in the from F. solani pisi suggested that S103 and H173 from Monilinia RCSB PDB: two inhibited structures (PDB IDs: 3DEA and 3DD5), and fructicola cutinase play important roles in catalysis. Site-directed mu- one uninhibited structure (PDB ID: 3DCN) (Nyon et al., 2009). This en- tants S103A, S103T and H173L all exhibited slightly increased Km values, zyme shares 50% sequence identity with F. soliani pisi cutinase, and while S103A showed a 2.3-fold higher kcat, with the soluble substrate the three dimensional structures of the two enzymes are very similar. 4-nitrophenyl butyrate (Wang et al., 2002). Site-directed mutagenesis However, the structures of the putative cutinase from G. cingulata reveal studies were also performed on C. sp. cutinase, based upon a compari- asignificant difference between the two. The conformation of the son of C. sp and F. solani pisi cutinases. The results suggested that large G. cingulata catalytic triad appears to cycle between an inactive form and hydrophobic residues would be crucial to increase the specificity and an active form during catalysis (Nyon et al., 2009). In the uninhibit- towards long-chain, low-molecular-weight substrates, such as 4- ed structure, the histidine residue that forms the center of the catalytic nitrophenyl palmitate. Introducing the mutations F52W and L181F in- triad is positioned outside of the active site, and does not interact with creased activity with 4-nitrophenyl palmitate by 4.86-fold and altered the remainder of the triad, catalytic serine and catalytic aspartate substrate specificity toward substrates with longer chain lengths (Fig. 4a). In addition, there is a small helix in the vicinity of the active (Kodama et al., 2009). site that places the catalytic serine in a deep hole in a deep pocket within One strategy to increase the activity of cutinase toward high- the active site (Fig 4b). This placement of the catalytic serine would molecular-weight substrates, such as various synthetic polyesters, was seem to make interaction with a high-molecular-weight substrate to create more space in the active site. F. solani pisi cutinase was modi- more difficult. Consequently, further investigation of its catalytic activi- fied by substituting the residues near the active site with smaller resi- ty is necessary to determine if this enzyme is a true cutinase. dues, such as alanine, to enlarge the active site. Compared with the

Fig. 4. Comparison of the unliganded structures of Glomerella cingulata and Fusarium solani pisi cutinases. a) Overlay of G. cingulata (PDB ID: 3DCN (Nyon et al., 2009), magenta ribbon) and F. solani pisi (PBB ID: 1CUS (Martinez et al., 1992), blue ribbon) cutinases. The catalytic triad of the G. cingulata structure is portrayed in magenta spheres. The catalytic triad of the F. solani pisi structure is displayed as red spheres (catalytic Ser), green spheres (catalytic His) and blue spheres (catalytic Asp). b) A rendering of the solvent accessible surface of G. cingulata cutinase showing the catalytic His in magenta, the catalytic Ser in red, and the catalytic Asp in blue. c) A rendering of the solvent accessible surface of F. solani pisi cutinase with the catalytic Ser in red and the catalytic His in green. The catalytic Asp cannot be seen in this view. All renderings were performed using PyMol. S. Chen et al. / Biotechnology Advances 31 (2013) 1754–1767 1761

Table 3 two mechanisms by which binding domains enhance hydrolysis of Site-directed mutagenesis results with F. solani pisi cutinase. polymeric material. On the one hand, the binding module dramatically Mutation Mutation site Amino acid Relative activity (%) increases the amount of active enzyme on the polymer interface, in- classification substitution in olive oil creasing the effective concentration of the enzyme in the vicinity of Active center S120 A 0 the substrate. On the other hand, binding modules can also partially Substrate binding T45 A, K 98, 74 disrupt the structure of the polymer, making the targeted bonds more region T50 V 25 accessible to the active site of the catalytic domain. This innovative R78 N, L 34, 49 approach of fusing cutinase with binding modules has been employed A79 G 50 T80 D 32 to improve the adsorption to, and hydrolysis of, PET. A cutinase from D83 S 62 T. cellullosylitica was fused with binding modules from Hypocrea jecorina A85 F, W 136, 109 cellobiohydrolase I (CBM) and Alcaligenes faecalis polyhydro- R88 A 39 xyalkanoate depolymerase (PBM), respectively. The results showed N172 K 45 that the adsorption of the fusion enzymes to PET was increased, and T173 K 119 T179 Y 131 PET hydrolysis activity of one of the fusions (Thc_Cut1 + CBM) was en- L182 W 19 hanced 3.8-fold (Ribitsch et al., 2013). I183 F 25 Cotton is another important substrate of cutinase in the textile in- A185 L 96 dustry. Unlike synthetic polyester PET, cotton fiber has a multilayered L189 F 109 fi Oxyanion hole S42 A 0.22 structure (Degani et al., 2004). In addition to cutin, cotton bers contain N84 A, L, W, D 26.5, 3.0, 0.11, 0.16 pectin, protein, and the surface layer of the cotton fiber contains a large Charged residues R17 N, E 31, 34 amount of cellulose. Thus, it also seems reasonable that engineering the D33 S 74 enzyme to specifical carbohydrate-binding modules (CBMs) in order to R96 N 57 increase its concentration on the surface of the cotton fiber substantial- D111 N 39 D134 S 37 ly. Our laboratory designed such a fusion protein, fusing CBMs from K151 R 29 T. fusca cellulase Cel6A (CBMCel6A)andCellulomonas fimi cellulase R156 E, L, K 79, 71, 115 CenA (CBMCenA), separately, to T. fusca cutinase. Both fusion proteins K168 L 83 displayed catalytic properties and pH stabilities similar to those of R196 E, L, K 45, 44, 38 R208 A 64 T. fusca cutinase. However, the fusion proteins did not bind to, or cata- Other mutations T18 V 90 lyze the hydrolysis of, cotton fibers significantly better than wild-type T19 V 35 cutinase. Somewhat surprisingly, the addition of pectinase enhanced I24 S 4 the cotton fiber binding activities of cutinase-CBMCel6A and cutinase- G26 A 32 CBM by 40%, and 45%, respectively. Moreover, a dramatic increase A29 S 64 CenA Y38 F 62 of up to 3-fold was observed in the amount of fatty acids released S54 E, K, W 34, 96, 89 from cotton fiber by the combination of cutinase-CBM fusion proteins W69 Y 12 with pectinase. The lack of improvement in catalytic efficiency from S92 R 50 simply fusing the CBM to cutinase suggests that most of the cellulose M98 C 35 fi L99 K 78 on the surface of cotton ber is not well exposed to the solvent and is L114 Y 20 embedded in an epidermis full of pectins, proteins, and other compo- T144 C 54 nents, thus limiting the binding of the CBM domain to the cotton fiber. N161 D 63 When pectinase was added in the reaction mixture, removal of pectin A164 R 41 by this enzyme must have exposed the cellulose to the solvent, resulting T167 L 54 G192 Q 44 in increased adsorption of cutinase-CBMs, which eventually led to a A195 S 38 higher scouring efficiency of the cotton fiber (Zhang et al., 2010).

A199 C 0 To expand the utility of the cutinase-CBMCel6A construct, the CBM E201 K 54 domain was modified to accommodate the structure of PET (Zhang I204 K 66 et al., 2013). Mutants W86L and W86Y exhibited a noticeable improve- ment in binding and catalytic efficiency (1.4- to 1.5-fold) toward PET wild type enzyme, mutants L182A, L81A, V184A and N84A showed ac- fiber, compared with the native enzyme. The enhanced binding and tivity enhancements of 5-, 4-, 2- and 1.7-fold toward high-molecular- hydrolytic activity may be the result of creating a new hydrogen bond, weight PET fibers, respectively. This strategy was also successfully or it might result from improved hydrophobic interactions between used to enhance the activity of T. fusca cutinase toward PET. The mutant the enzyme and the PET fiber. I218A, which was designed to create space, and the double mutant Q132A/T101A, which was designed both to create space and to increase 3. Preparation of cutinase hydrophobicity, both hydrolysed PET with considerable efficiency. In addition, the activity of the double mutant Q132A/T101A with the solu- 3.1. Preparation of cutinases from wild-type strains ble substrate 4-nitrophenyl butyrate was 2-fold higher than that of the wild-type cutinase (Silva et al., 2011). The increased activity of these The production of cutinases from wild strains has been studied mutants may be explained by an increased stabilization of the tetrahe- extensively in the past decades. Several essential factors influence the dral intermediate and a better accommodation of the large substrate secretion of cutinases from wild-type strains, including the types and (Araújo et al., 2007). Substitution of positively charged amino acids concentrations of carbon and nitrogen sources, the culture temperature located on the enzyme surface with uncharged hydrophilic, or hydro- and pH, and the concentration of dissolved oxygen. Secreted cutinase phobic amino acids was successful in improving the activity of a activity can be dramatically increased by induction with cutin, cutin hy- cutinase from Thermobifida cellulosilytica. Mutants R19S and R29S, for drolysis monomers, and some lipids (Table 4), while cutinase secretion example, showed strongly increased PET hydrolysis activity (Herrero is sometimes repressed by glucose (Castro-Ochoa et al., 2012; Fett et al., Acero et al., 2013). 1999, 2000). In our laboratory, using a two-stage pH control strategy, More recently, substrate-binding modules have also been described the maximal cutinase activity secreted by the T. fusca strain WSH03- for enzymes hydrolyzing natural and synthetic polyesters. There are 11 reached 19.8 U/mL (Du et al., 2007). To further evaluate the use of 1762 S. Chen et al. / Biotechnology Advances 31 (2013) 1754–1767

Table 4 expressed in the Gram-negative bacterium Escherichia coli, the expres- Production of cutinases by wild-type strains. sion level was relatively low because the cutinase gene contains 18 co- Source Inductor Fermentation Assay Enzyme Reference dons that are used at a low frequency in E. coli genes (Griswold et al., period (h) substratea activity 2003). Expressing F. solani pisi cutinase in the Gram-positive bacterium b (U/mL) Bacillus subtilis allowed the recombinant cutinase to reach 60 mg/L T. vulgaris Tomato 96 pNPB 13.58 Fett et al. (2000) (11.5 U/mL) in the culture supernatant (Brockmeier et al., 2006a, peel 2006b). Saturation mutagenesis of the positively charged N-domain S. badius Apple 336 pNPB 0.160 Fett et al. (1992a) (positions 2–7) of the signal peptide revealed four point mutations cutin fi S. acidiscabies Apple 336 pNPB 0.22 Fett et al. (1992a) that resulted in the production of signi cantly increased amounts of cutin cutinase (Caspers et al., 2010). A. nidulans Olive oil 24 pNPL 1.624 Castro-Ochoa A variety of yeast systems have also been used to express the F. solani et al. (2012) pisi cutinase. When this cutinase was expressed in Saccharomyces T. fusca Tomato 168 pNPB 12.15 Fett et al. (1999) cerevisiae SU50, the activity in the culture medium was 123 U/mL peel (van Gemeren et al., 1995). By controlling the dissolved oxygen concen- a pNPB = 4-nitrophenyl butyrate; pNPL = 4-nitrophenly laurate. tration and pH, the cutinase activity could reach 170 U/mL (Calado et al., b One unit of esterase activity was defined as the amount of enzyme that liberated 1 μmol 4-nitrophenol per minute under assay conditions. 2002). Engineering a consensus N-glycosylation sequence into the N-terminal region of F. solani pisi cutinase increased secretion of cutinase from S. cerevisiae by 5-fold, while engineering an N- municipal sludge for T. fusca cutinase production, short-chain organic glycosylation site into the C-terminal region increased cutinase secre- acids were used as carbon sources. It was found that the optimum tion 1.8-fold (Sagt et al., 2000). In further attempts to construct an ratio of butyrate, acetate, and lactate was 4:1:3. Subsequently, industrial-scale process for the production of recombinant F. solani pisi two-stage batch and fed-batch cultivation strategies were investigated cutinase in S. cerevisiae, an expression strain was constructed in which and it was found that the cutinase activity reached a maximum of expression was induced through a galactose promoter. By adopting a 51.0 U/mL after 22 h of culture (He et al., 2009). The production of typical fed-batch fermentation strategy, a cutinase concentration of F. oxysporum cutinase by solid-state fermentation using Brazilian agri- 546 mg/L was obtained in the medium (Ferreira et al., 2003). The final cultural by-products has also been recently studied. The maximum system produced by these researchers integrated expression and purifi- yield observed was 21.7 U/mL after 120 h of fermentation at 28.3 °C cation into a system for the production of F. solani pisi cutinase from (Fraga et al., 2012). Through the use of response surface methodology, S. cerevisiae that may be useful on an industrial scale (Calado et al., the greatest cutinase activity from F. oxysporum (22.68 U/mL) was 2004). F. solani pisi cutinase has also been expressed in the yeast Pichia achieved after 48 h of fermentation in a liquid mineral medium pastoris (Kwon et al., 2009). Without any optimization, the concentra- supplemented with flaxseed oil to induce cutinase secretion (Pio and tion of cutinase in flasks reached about 340 mg/L, which suggests that Macedo, 2007, 2008). an optimized P. pastoris expression system could be very promising Production of cutinases from their wild-type strains suffers from the for F. solani pisi cutinase production. disadvantages of low productivity and long fermentation periods. Thus, Fungal systems have also been used to express F. solani pisi cutinase. although these systems allow the production of sufficient quantities of Using Aspergillus awamori as host, cutinase was secreted into culture cutinase to meet the needs of researchers, they do not produce sufficient medium to a concentration of 50 mg/L (35 U/ml) (van Gemeren et al., material to meet the needs of industrial applications. Some researchers 1995). The effect of multicopy integration of the expression cassette have turned to recombinant expression of cutinases in homologous and upon cutinase production in A. awamori was also investigated. When heterologous hosts, hoping to shorten fermentation times, improve the the copy number reached 5–10, cutinase activity reached 64 mg/L robustness of the fermentation, and increase both the scale and the (van Gemeren et al., 1996). In 2007, the expression of F. solani pisi yield. cutinase in Fusarium venenatum A3/5, a recently developed expression system, was investigated. The yield of secreted cutinase was 0.021 U/mL (Sorensen et al., 2007). 3.2. Preparation of cutinase using engineered strains The fungal cutinases from A. oryzae, Aspergillus niger and M. fructicola have also been cloned and expressed in P. pastoris using the methanol- With the advent of recombinant DNA technology, many cutinases induced AOX1 as promoter (Liu et al., 2009; Nyyssola et al., 2013; Wang have been produced through large-scale expression in both homolo- et al., 2002). A P. Antarctica cutinase gene was cloned under the control gous and heterologous hosts. Optimization of fermentation conditions of the GAL1 promoter and expressed in S. cerevisiae (Shinozaki et al., further improved the production of cutinases. 2013b). Furthermore, Trichoderma reesei was chosen as host to express F. solani pisi cutinase has been extensively studied and expressed in a thecutinasegenefromC. cinerea (Kontkanen et al., 2009). A production variety of heterologous hosts (Table 5). When F. solani pisi cutinase was yield of 1.4 g/L cutinase was obtained in a laboratory fermenter.

Table 5 Expression of F. solani pisi cutinase in heterologous hosts.

Espression host Signal peptide Promoter Vector Cutinase in culture mediuma Reference

E. coli W6 Alkaline phosphatase Ptac (IPTG inducible) pMal/c as 2.5 U/mL Griswold et al. (2003) (PhoA) signal peptide Bacillus subtilis LipA signal peptide Constitutive promoter P59 pBSMul2 60 mg/L (11.5 U/mL) Brockmeier et al. (2006a, 2006b) Saccharomyces cerevisiae SU50 Gal7 promoter pUR7320 175 mg/L (123 U/mL) van Gemeren et al. (1995) Pichia pastoris α-factor signal peptide Methanol-inducible alcohol pPICZaA 340 mg/L Kwon et al. (2009) oxidase promoter Aspergillus awamori F. solani cutinase signal Endoxylanase II promoter pAW14B12 50 mg/L (35 U/mL) van Gemeren et al. (1995) peptide Fusarium venenatum A. niger glucoamylase Co-transformed with 0.021 U/mL Sorensen et al. (2007) promoter plasmids pIGF and pAN8-1

a One unit of esterase activity was defined as the amount of enzyme that liberated 1 μmol 4-nitrophenol per minute under assay conditions. S. Chen et al. / Biotechnology Advances 31 (2013) 1754–1767 1763

Bacterial cutinases have been cloned and expressed using the Gram- produce dehydrated fruits, several dairy products, flavor compounds negative bacterium E. coli as host. In our laboratory, the T. fusca cutinase and some important fatty acids, including eicosapentanoic acid (EPA) genes Tfu_0882 (Gene bank accession no. AAZ54920.1) and Tfu_ and docosahexanoic acid (DHA) (Dutta et al., 2009). In the chemical 0883(Gene bank accession no. AAZ54921.1) were expressed in E. coli industry, cutinases can be employed for the production of biodiesel, phe- using a PelB signal peptide. The enzyme activity in the culture superna- nolic compounds, surfactants, and some chiral chemicals. Cutinases also tant was 9.5-fold and 3.6-fold higher, respectively, than that achieved have shown potential in the surface functionalization of polymers, as with cutin-induced T. fusca cells (Chen et al., 2008). Subsequently, trans- well as in the removal of coatings and recycling of polymers (Badenes lational initial region (TIR) degeneracy mutagenesis was carried out on et al., 2010, 2011; de Barros et al., 2009; Greimel et al., 2013; Guebitz the initial sequence of the pelB signal peptide in recombinant E. coli.One and Cavaco-Paulo, 2008; Korpecka et al., 2010; Mueller, 2006). In the de- of the mutants produced twice as much cutinase as the control con- tergent industry, cutinases have been evaluated as lipolytic enzymes in struct (Liu et al., 2011). The influence of the signal peptide cleavage laundry and dishwashing detergent formulations. Because the cutinases site number on extracellular production of recombinant cutinase was have good stability in the 20–50 °C temperature range, and at pH 8–11, also investigated. The results indicated that an increase in signal peptide and are also stable in the presence of other enzymes, and even H2O2, cleavage sites can improve the extracellular production of recombinant they are thought to be more beneficial than commercial lipase for remov- cutinase in E. coli (Chen et al., 2011a). Using the OmpA signal peptide, ing triacylglycerols and hydrolyzing fats without added calcium (Dutta concentrations of cutinase Tfu_0883 as high as 500 mg/L could be et al., 2009). In the environmental industry, cutinase is helpful in the bio- achieved (Dresler et al., 2006). Optimizing the culture conditions im- degradation of wastes, including polymer and pesticide wastes (Bhardwaj proved cutinase production, and yielded 145.2 U/mL (Li et al., 2010). et al., 2013; Dutta et al., 2009; Kleeberg et al., 1998). The applications of A highly hydrophobic cutinase from T. fusca KW3 (TfCa), fused to a cutinases in these industries have been thoroughly documented in previ- pelB leader sequence, was also expressed in E.coli BL21(DE3). By opti- ous reviews, so it is unnecessary to review these applications here. In re- mizing codon usage and expression conditions, the yield of cutinase cent years, cutinases have begun to play a significant role as biocatalysts was remarkably improved—to 4500-fold higher than the yield from in the textile industry (J. Chen et al., 2007). Therefore, the remainder of the wild-type strain (Oeser et al., 2010). In all the studies above, the review will focus on the uses of cutinases in the textile industry. T. fusca cutinase was expressed and secreted via the type II secretory system in E. coli. After fermentation, it was apparent that a large amount 4.1. Cotton fibers of cutinase had accumulated in periplasm of the E. coli. Although glycine and surfactant have been used to promote the escape of recombinant Cotton fiber is the most popular textile in the world. In an important cutinase from the periplasm (Chen et al., 2011b), the problem has not step of the process of making cotton fabrics, called cotton scouring, the been completely solved. Because the alpha-hemolysin secretion system cuticle layer around the cotton fiber is removed. This improves the wet- can export target proteins across both the inner and outer membranes tability of the fiber, which then facilitates uniform dyeing and finishing. of E. coli, directly into the culture medium, this system was used to at- Traditionally, this process has been performed using alkaline hydrolysis tempt secretion of T. fusca cutinase. The specific signal peptide of the at high temperature, which not only consumes large quantities of water alpha-hemolysin secretion system was fused to the N-terminus of and energy but also causes severe pollution and fiber damage. There- T. fusca cutinase. In addition, HlyB and HlyD, which are strain-specific fore, environment-friendly treatments based on biocatalysts have translocation components of the alpha-hemolysin secretion system, been actively sought. were coexpressed to facilitate enzyme transport. It was found that this The cuticle layer of a cotton fiber has a complicated composition that engineered strain was able to secrete substantial cutinase (334 U/mL) includes cutin, wax, pectin and protein. Both the wax and cutin can be into the culture medium—2.5 times the amount produced using the hydrolysed by the cutinases from Pseudomonas mendocina, F. solani pisi type II secretion pathway, under the same culture conditions (Su et al., and T. fusca,aswellasT. fusca cutinase fused with CBM. When these 2012). Surprisingly, when T. fusca cutinase without a signal peptide cutinases are combined with pectinase, the wettability of cotton fiber was expressed in E. coli strain BL21(DE3), the majority of the cutinase can be improved efficiently, at low temperature, without the addition activity was located in the culture medium. In a 3 L fermenter, the of alkali (Agrawal et al., 2008; Degani et al., 2002; Yan et al., 2011; cutinase activity in culture medium reached 1063.5 U/mL after 24 h, Zhang et al., 2010, 2011). However, at relatively low temperatures, the which represents a 4.4-fold increase over the highest previous yield. wax is only partially hydrolyzed and can't be removed completely, The underlying mechanism for extracellular expression of cutinase which can lead to uneven properties of the textile. In contrast, the wax without a signal peptide could be “cell leakage induced by cutinase's can be degraded completely and become soluble at higher temperatures limited phospholipid hydrolysis” (Su et al., 2013a). Co-expression (80–90 °C). However, most cutinases have limited stability at higher with mature cutinase that lacks a signal peptide may be a useful system temperatures. Some researchers have tried to enhance the stability of for the secretion of other useful recombinant proteins from E. coli cutinases using immobilization technology (Baptista et al., 2003; BL21(DE3) (Su et al., 2013b). Goncalves et al., 1999; Serralha et al., 2002) Unfortunately, the immobi- To summarize, the production of recombinant cutinases in heterolo- lization material interferes with the interaction between cutinase and gous hosts has been widely investigated. Expression of fungal cutinases the bulky polyesters it must hydrolyse. Therefore, immobilization is not (particularly F. solani pisi cutinase) in yeast expression systems, wheth- a viable alternative in textile industry. Recent research has been directed er S. cerevisiae or P. pastoris, has shown significant promise on the road toward the search for a thermostable cutinase that can be applied in the to an industrial-scale process. Bacterial cutinases, in contrast, have treatment of cotton at higher temperatures. In addition to the cotton cu- expressed well in E. coli systems. The work described here is a solid ticle described above, cotton fibers are also contaminated with the cotton foundation upon which industrial scale production of useful cutinase seed coat. The cotton seed coat is resistant to removal, and its incomplete enzymes can be built. removal during bioscouring can cause problems during the subsequent dyeing process. Fortunately, cutinase combined with alkaline pectinase 4. Application of cutinase in industry or xylanase, can improve the degradation of cotton seed coat during the cotton fabric bioscouring process (Yan et al., 2009a, 2009b). Cutinases are multifunctional enzymes that can catalyze hydrolysis re- actions, esterifications and transesterifications. As a result, they have sub- 4.2. Synthetic fibers stantial potential to be widely used in the food, chemical, detergent, environmental, and textile industries (Carvalho et al., 1998, 1999). In Synthetic fibers represent almost 50% of the worldwide market for the food industry, cutinases have been evaluated as potent catalysts to textile fibers. The process of synthetic fiber modification, which is 1764 S. Chen et al. / Biotechnology Advances 31 (2013) 1754–1767 similar to cotton scouring and results in the partial hydrolysis of the syn- Cutinases have immense potential as industrial enzymes, especially thetic fiber, improves its wettability and facilitates the uniform dyeing in the textile industry. Great progresses have been made toward realiz- and finishing of synthetic fabrics. The application of cutinase in synthet- ing this potential in recent years. However, most of the reports available ic fiber modification is a relatively recently phenomenon. It has been describe work on the laboratory scale, and scale-up of these applications reported that F. solani pisi and T. fusca cutinase can modify the surface for commercial use has yet to be done. of synthetic fibers, like polyesters, polyamides, acrylics, and cellulose acetate, and improve their wettability and dyeability (C.M. Silva et al., 5. Conclusions 2005). fi PET, one of the most widely used synthetic bers, is a polyester com- Cutinases have recently been the subject of intense and increasing posed of terephthalic acid and ethylene glycol. Cutinase is able to par- research because of their substrate diversity. A substantial number of fi tially hydrolyse the ester bonds on the surface of the ber, which cutinase genes have been identified and an increasing number of liberates hydrophilic hydroxyl and carboxylic acid groups. This treat- three-dimensional structures have been solved, establishing a firm ment enhances the hydrophilicity of PET fabrics without compromising basis for developing a deeper understanding of cutinases and related fi the ber strength (Alisch-Mark et al., 2006; Brueckner et al., 2008; Eberl enzymes. However, most of the genes identified have been those of et al., 2009; Fischer-Colbrie et al., 2004; O'Neill et al., 2007; Ribitsch fungal cutinases; identification of cutinase genes from bacterial and et al., 2011, 2012a; Ronkvist et al., 2009). The addition of Triton X-100 other sources have been relatively limited. With this growing body of facilitates cutinase hydrolysis of the surface of PET fabrics (Lee and knowledge, it has become clear that there are substantial differences Song, 2010). The ability of cutinase to hydrolyse surface ester bonds is in structure and function between cutinases from different sources. greatly affected by the crystallinity of the polymer, displaying relatively The continued mining of new genes from the diversity of nature is an fi high activity towards an amorphous polyester lm and low activity on a excellent strategy to discover novel and powerful cutinases. highly crystalline substrate (Vertommen et al., 2005). Cutinases can be distinguished from esterases by their ability to fi Polyamide, another important synthetic ber in textile industry, is hydrolyze high-molecular-weight polyesters. Cutinases can be distin- made from hexamethylenediamine and adipic acid through the creation guished from lipases by their ability to hydrolyze smaller, soluble sub- fi of amide bonds. Polyamide can be modi ed by proteases and strates, and their lack of interfacial activation. At this point in time, polyamidases, as well as cutinases (Almansa et al., 2008; C. Silva et al., meeting industrial requirements for the catalytic efficiency of cutinases 2005; Heumann et al., 2009). The activities of these enzymes showed with polyester substrates is still very challenging. Low thermostability additive effects with higher levels of mechanical agitation (Silva et al., remains a disadvantage with most cutinases. Rational engineering and 2007). directed evolution of known cutinase genes will be powerful tools to Acrylic is somewhat different than polyester and polyamide, because improve the properties of cutinases. The data derived from these the bond that can be hydrolysed by cutinase is not part of the backbone novel cutinases, along with the information obtained from the modifica- of the polymer. Acrylic is prepared through the polymerization of acry- tion of existing cutinases, will further enable efforts to identify cutinase lonitrile, with 7% vinyl acetate as a comonomer. Cutinase is able to enzymes that are optimal for industrial process. Existing production hydrolyse the ester bonds of the vinyl acetate units, releasing acetic techniques, along with new processes that are being developed today, fi acid and leaving hydroxyl groups at the ber surface. Since these groups will enable production of these enzymes on a scale sufficient to meet in- fi are hydrophilic, they improve the wettability of the ber (C.M. Silva dustrial needs. These new cutinases will help manufacture valuable ma- et al., 2005). terials using environmentally friendly processes. Cutinase also exhibits acetyl esterase activity on cellulose diacetate and triacetate, which are the building blocks of cellulose acetate fibers. Acknowledgments An increase in the number of hydroxyl groups at the diacetate and triac- etate fiber surface can be obtained through the action of cutinase. The This work was supported financially by the Science and Technology cutinase-CBM fusion protein, mentioned above, was found that has a Support Project of Jiangsu Province (nos. BE2012018 and BE2012019), greater effect on cellulose acetate fiber than native cutinase in terms the 111 Project (no. 111-2-06), and the High-end Foreign Experts of improving wettability and dyeability (Matama et al., 2010; Zhang Recruitment Program (Ronald W. Woodard). The authors gratefully et al., 2012). Model substrates have developed to screen for enzymes acknowledge the writing and editing assistance provided by Dr. Tod P. that can more efficiently modify synthetic fibers. 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