Screening and characterization of novel polyesterases from

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Environmental Processes

Screening and characterization of novel polyesterases from environmental metagenomes with high hydrolytic activity against synthetic polyesters Mahbod Hajighasemi, Anatoli Tchigvintsev, Boguslaw P. Nocek, Robert Flick, Ana Popovic, Tran Hai, Anna N. Khusnutdinova, Greg Brown, Xiaohui Xu, Hong Cui, Julia Anstett, Tatyana N. Chernikova, Thomas Bruls, Denis Le Paslier, Michail M. Yakimov, Andrzej Joachimiak, Olga V. Golyshina, Alexei Savchenko, Peter N Golyshin, Elizabeth A. Edwards, and Alexander F. Yakunin Environ. Sci. Technol., Just Accepted Manuscript • DOI: 10.1021/acs.est.8b04252 • Publication Date (Web): 04 Oct 2018 Downloaded from http://pubs.acs.org on October 5, 2018

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Environmental Science & Technology

ACS Paragon Plus Environment


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Screening and characterization of novel polyesterases from environmental

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metagenomes with high hydrolytic activity against synthetic polyesters

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Mahbod Hajighasemi,1 Anatoli Tchigvintsev,1 Boguslaw Nocek,2 Robert Flick,1 Ana Popovic,1

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Tran Hai,3 Anna N. Khusnutdinova,1 Greg Brown,1 Xiaohui Xu,1 Hong Cui,1 Julia Anstett,1

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Tatyana N. Chernikova,3 Thomas Brüls,4 Denis Le Paslier,5 Michail M. Yakimov,6 Andrzej

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Joachimiak,2 Olga V. Golyshina,3 Alexei Savchenko,1 Peter N. Golyshin,3 Elizabeth A. Edwards,1

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and Alexander F. Yakunin1*

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Toronto, ON, M5S 3E5, Canada

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Department of Chemical Engineering and Applied Chemistry, University of Toronto,

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Midwest Center for Structural Genomics and Structural Biology Center, Biosciences Division, Argonne National Laboratory, Argonne, Illinois 60439, U.S.A.

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School of Biological Sciences, Bangor University, Gwynedd LL57 2UW, UK

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Commissariat à l’Energie Atomique et aux Energies Alternatives (CEA), Direction de

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la Recherche Fondamentale, Institut de Génomique, Université de d’Evry Val

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d’Essonne (UEVE), Centre National de la Recherche Scientifique (CNRS), UMR8030,

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Génomique métabolique, Evry, France

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Université de d’Evry Val d’Essonne (UEVE), Centre National de la Recherche

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Scientifique (CNRS), UMR8030, Génomique métabolique, Commissariat à l’Energie

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Atomique et aux Energies Alternatives (CEA), Direction de la Recherche

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Fondamentale, Institut de Génomique, Evry, France

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Institute for Coastal Marine Environment, CNR, 98122 Messina, Italy

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Corresponding author: Email [email protected]; phone 416-978-4013; fax 416-9788605

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ABSTRACT

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The continuous growth of global plastics production, including polyesters, has resulted in

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increasing plastic pollution and subsequent negative environmental impacts. Therefore, enzyme-

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catalyzed depolymerization of synthetic polyesters as a plastics recycling approach has become a

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focus of research. In this study, we screened over 200 purified uncharacterized hydrolases from

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environmental metagenomes and sequenced microbial genomes and identified at least 10 proteins

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with high hydrolytic activity against synthetic polyesters. These include the metagenomic

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esterases MGS0156 and GEN0105, which hydrolyzed polylactic acid (PLA), polycaprolactone,

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as well as bis(benzoyloxyethyl)-terephthalate. With solid PLA as a substrate, both enzymes

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produced a mixture of lactic acid monomers, dimers, and higher oligomers as products. The

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crystal structure of MGS0156 was determined at 1.95 Å resolution and revealed a modified α/β

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hydrolase fold, with a lid domain and highly hydrophobic active site. Mutational studies of

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MGS0156 identified the residues critical for hydrolytic activity against both polyester and

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monoester substrates, with two-times higher polyesterase activity in the MGS0156 L169A mutant

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protein. Thus, our work identified novel, highly active polyesterases in environmental

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metagenomes and provided molecular insights into their activity, thereby augmenting our

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understanding of enzymatic polyester hydrolysis.

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TOC Art

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INTRODUCTION

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Over the last 50 years, a tremendous increase in production of synthetic polymers and their

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persistence in the environment resulted in elevated levels of pollution.1-4 Most petroleum-based

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plastics (polyethylene, polypropylene, polyethyleneterephthalate) are remarkably stable in the

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environment, resulting in the accumulation of plastic waste and microplastic particles, which

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negatively affect marine ecosystems.4-6 The development of biodegradable plastics represents part

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of a solution that includes other approaches such as plastic recycling and polymer recovery.7, 8

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The most sustainable option for plastic waste treatment appears to be a closed-loop recycling

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process based on physical, chemical, and biocatalytic depolymerization and the recovery of

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chemical feedstocks for the synthesis of novel polymers (a circular economy).9-11 Compared to

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physical, thermal, and chemical plastics depolymerization, biocatalytic recycling has several

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advantages, including low energy consumption, mild reaction conditions, and the possibility for

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stereospecific degradation and enzymatic repolymerization.9, 12

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Most conventional polymers (polyethylene, polypropylene, polystyrene, poly(vinyl chloride), and poly(ethylene terephthalate)) exhibit limited or no biodegradability.2 The group of

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biodegradable plastics with commercial relevance is dominated by aliphatic polyesters such as

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polylactic acid (PLA) and polycaprolactone (PCL), whereas the aromatic-aliphatic co-polyester

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poly(ethylene terephthalate) (PET) is more resistant to microbial or enzymatic attack.13, 14 The

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complex process of polymer biodegradation in the environment is a combination of various

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abiotic and biotic factors.15, 16 In nature, different groups of anaerobic (Clostridium and

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Propionispora) and aerobic bacteria and fungi (including actinomycetes (Amycolatopsis, Lentzea,

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Pseudonocardia), Bacillaceae (Geobacillus, Brevibacillus, Paenibacillus), Pseudomonas, and

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Aspergillus)have been found to degrade polyesters.17, 18 Enzymatic hydrolysis of polyesters was

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demonstrated over 40 years ago using purified lipases and proteases from different fungi and

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Achromobacter sp., as well as hog liver esterase.13, 19 These enzymes belong to a large group of

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serine-dependent α/β hydrolases, which use the catalytic triad Ser-His-Asp (or Glu) for the

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hydrolysis of different monomeric and polymeric substrates. Furthermore, polyester degrading

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(polyesterase) activity was later demonstrated in cutinases (also serine-dependent α/β hydrolases)

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from fungal and bacterial plant pathogens, which secrete these enzymes to degrade the plant

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polyester cutin.20 Cutinases are esterase-like enzymes, hydrolysing not only cutin, but also water-

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soluble monoesters and synthetic polyesters, including PCL and PET.21

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A number of polyester degrading esterases and lipases have been characterized biochemically,

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including Paenibacillus amylolyticus PlaA, Thermobifida fusca TfH, ABO1197 and ABO1251

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from Alcanivorax borkumensis, several clostridial esterases (Chath_Est1, Cbotu_EstA,

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Cbotu_EstB), and the metagenomic polyesterases PlaM4, EstB3, and EstC7.22-27 These enzymes

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hydrolysed a broad range of both emulsified and solid polyester substrates, such as PLA, PCL,

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PET, poly(ethylene adipate), and poly(butylene adipate-co-butylene terephthalate) (PBAT).27-30

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Enzymatic activity of purified polyesterases was also characterized using the soluble monoesters

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p-nitrophenyl butyrate and p-nitrophenyl acetate, which are common substrates for

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carboxylesterases.24, 27, 30 Moreover, several purified cutinases from bacteria (Thermobifida),

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fungi (Humicola, Aspegillus, Fusarium), and environmental metagenomes have been shown to

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hydrolyze synthetic polyesters including PET and polyester polyurethane.14, 20, 31-33 These works

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also revealed several important factors limiting the biodegradation of synthetic polyesters,

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including the hydrophobicity, crystallinity, surface topography, and molecular size of synthetic

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polymers.2, 3, 34, 35 It has been shown that the hydrophobic polymer surface restricts effective

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adsorption and activity of polymer-degrading enzymes.36-38 To improve protein sorption and

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thereby polymer hydrolysis, the Thermobifida cellulosilytica cutinase Thc_Cut1 was fused to

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non-catalytic substrate-binding modules from the Hypocrea jecorina cellobiohydrolase I or

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Alcaligenes faecalis polyhydroxyalcanoate depolymerase.36 Both fusion enzymes demonstrated

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increased adsorption and hydrolytic activity on PET, likely due to enhanced hydrophobic

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interactions with the substrate. Crystal structures have been determined for several polyesterases, including several carboxyl

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esterases (Clostridium hathewayi Chath_Est1, C. botulinum Cbotu_EstA, and

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Rhodopseudomonas palustris RPA1511) and cutinases (Thermomyces (formerly Humicola)

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insolens HiC, Thermobifida alba Est119, and metagenomic LC-cutinase from leaf-branch

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compost).25, 30, 39-42 Polyesterase structures revealed the classical α/β hydrolase core domain with

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or without an α-helical lid (or cap) domain covering the active site, with the catalytic Ser residue

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positioned at the bottom. The polyesterase active site usually represents a wide-open cleft,

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directly accessible to polymeric substrates as revealed by the structures of RPA1511 and Est119

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in complex with polyethylene glycol bound close to the catalytic triad.41, 42 Interestingly, when the

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active site cleft of the Ideonella sakaiensis PETase (PET-degrading enzyme) was narrowed by

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site-directed mutagenesis (to make it more cutinase-like), the resulting engineered PETase

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outperformed the wild-type protein in degradation of both PET and polyethylene-2,5-

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furandicarboxylate.43 In addition, mutagenesis and protein engineering experiments with the

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Thermobifida cellulosilytica cutinases Thc_Cut1 and Thc_Cut2 confirmed an important role of

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enzyme surface and hydrophobic interactions for polyester hydrolysis.36, 44 In addition, the recent

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work by Zumstein et al. suggested that polyesterase activity of different carboxylesterases

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depends on the accessibility of their active sites.45

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Although recent studies have identified a number of polyester degrading enzymes, the

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continuously growing global demand for plastics and new polymers has also stimulated interest in

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novel enzymes and biocatalytic approaches for polymer recycling technologies. The discovery of

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novel polymer degrading enzymes, engineering of more active enzyme variants, as well as

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understanding of the molecular mechanisms of these enzymes represent the key challenges for the

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development of biocatalytic strategies for polymer hydrolysis and synthesis.3 To address these

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challenges, we have identified over 20 novel polyesterases using enzymatic screening, and

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biochemically characterized MGS0156 and GEN0105, which showed high hydrolytic activity

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against a broad range of polyesters (PLA, PCL, polyethylene succinate (PES), poly(butylene

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succinate-co-adipate) (PBSA), and 3PET). The crystal structure of MGS0156 was solved,

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revealing an open active site with hydrophobic surface, whereas structure-based mutagenesis

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studies identified amino acid residues critical for enzymatic activity, with the L169A mutant

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protein displaying two-times higher polyesterase activity.

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MATERIALS AND METHODS

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Reagents. All chemicals and substrates used in this study were of analytical grade unless

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otherwise stated. Polymeric substrates were purchased from Sigma-Aldrich (St. Louis, MO,

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USA) except poly (D,L-lactide) PLA2 (Mw 0.2 × 104), PLA70 (Mw 7.0 × 104), as well as poly (L-

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lactide) PLLA40 (Mw 4.0 × 104), that were obtained from PolySciTech (Akina Inc., West

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Lafayette, IN, USA). Commercial-grade PLA polymers (IngeoTM 4032D, and IngeoTM 6400D)

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were products of NatureWorks LLC (NE, USA), poly (D-lactide) PURASORBTM PD 24 of

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Corbion Purac (Amsterdam, The Netherlands), whereas polybutylene succinate (PBS)

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(BionolleTM 1001MD, and BionolleTM 1020MD) and polybutylene succinate-co-adipate (PBSA)

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(BionolleTM 3001MD, and BionolleTM 3020MD) were purchased from Showa Denko K.K., Japan.

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The PET model substrate, bis(benzoyloxyethyl) terephthalate (3PET), was synthesized by

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CanSyn (Toronto, ON, Canada) using a previously reported protocol.46 The surfactant Plysurf

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A210G was obtained from Dai-ichi Kogyo Seiyaku Co. (Tokyo, Japan) and used to emulsify the

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polymers.

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Gene cloning, protein purification, and mutagenesis. The coding sequences of selected

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hydrolase genes were PCR amplified using the genomic DNA of the host organism or the gene

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constructs from metagenomic libraries as the templates. The PCR products were cloned using a

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ligation-independent protocol into a modified pET15b (Novagen) expression vector containing an

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N-terminal 6His tag. The plasmids were transformed into Escherichia coli BL21 (DE3) Codon-

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Plus strain (Stratagene) as the expression host. Transformants were grown in shake flasks of

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Terrific Broth (1 L) at 37 °C to the optical density A600 ≈ 0.6 followed by an induction with 0.5

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mM (final concentration) isopropyl 1-thio-β-D-galactopyranoside (IPTG) and an overnight

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incubation at 16 °C. The cells were harvested by centrifugation, resuspended in binding buffer

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(50 mM HEPES pH 7.5, 0.25 M NaCl, 5 mM imidazole and glycerol 5% vol.) and disrupted by

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sonication followed by another round of centrifugation to remove cell debris. Recombinant

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proteins were purified to homogeneity (>95%) as described previously,47 using metal-chelate

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affinity chromatography on Ni-NTA Superflow (Ni2+ -nitrilotriacetate; Qiagen) resin as well as

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ion exchange chromatography on a Mono Q GL 10/100 column (GE Healthcare) or size

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exclusion chromatography on a Superdex 200 16/60 column (Amersham Bio- sciences)

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equilibrated with 10 mM HEPES (pH 7.5), 0.25 M NaCl and 1 mM TCEP [tris-(2-

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carboxyethyl)phosphine] using ÄKTA FPLC (Amersham Biosciences) where necessary.48 Site-

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directed mutagenesis of metagenomics esterases was performed using a QuickChange® kit

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(Stratagene) according to the manufacturer’s protocol. Wild-type MGS0156 and GEN0105 were

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used as the templates, and mutations were verified via DNA sequencing. The selected residues

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(Pro167, Leu169, Leu170, Glu172, Cys173, Val174, Ser175, Leu179, Leu197, Arg199, His231,

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Ser232, Lys233, Ser265, Phe271, Arg277, Glu280, Cys287, Leu296, Leu299, Glu330, Leu335,

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Phe338, Asp350, Leu352, Val353, Asp372, His373, Met378, Phe380 for MGS0156 and Ser168,

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Glu262, His292 for GEN0105) were mutated to Ala or Gly (for insoluble Ala mutant proteins).

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The amino acid numbering is based on the full-length protein. Mutant proteins were

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overexpressed and purified in the same manner as described for the wild-type proteins. Multiple

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sequence alignment was conducted by Clustal Omega v1.2.1 through EMBL-EBI server, whereas

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phylogenetic analysis was performed by MEGA v7.0 using the neighbor-joining method.49, 50

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Esterase assays with soluble substrates. Carboxylesterase activity was measured

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spectrophotometrically as described previously.47 Purified enzymes (0.05-10.0 µg

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protein/reaction) were assayed against α-naphthyl or p-nitrophenyl (pNP) esters of different

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saturated fatty acids (C2-C16; 0.25-2.0 mM) as substrates in a reaction mixture containing 50

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mM HEPES-K buffer (pH 8.0).47 The stock solutions of α-naphthyl (100 mM) and p-nitrophenyl

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(50 mM) ester substrates were prepared in acetone and isopropanol, respectively. The final

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concentration of substrates was maintained lower than their water solubility to yield a

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homogeneous solution. Reaction mixtures (200 µl, in triplicate) were incubated at 30 °C in a 96-

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well plate format. Enzyme kinetics were determined by substrate saturation curve fitting (non-

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linear regression) using GraphPad Prism software (version 7.0 for Mac, GraphPad Software, CA,

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USA).

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Polyester degradation (polyesterase) screens. Emulsified polyester substrates were prepared

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in 50 mM Tris-HCl buffer (pH 8.0) as described previously.41, 51 Polyester emulsions were

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solidified with agarose (1.5%, w/v) and poured into 150 mm cell culture dishes with 20 mm grids

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to make a uniformly opaque gel. Cylindrical wells (3 mm diameter) were aseptically punched in

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the assay plate and inoculated accordingly with purified enzyme solutions (20 µl containing 1.5

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nmol of protein/well). Sealed assay plates were incubated at 30 °C and monitored for 3 weeks.

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The presence of polyesterase activity was inferred from the formation of a translucent halo

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around the wells with purified proteins.41, 51 To compare the size of clear halos formed by

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different enzymes, image analysis was conducted using the histogram tool on Adobe Photoshop

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software and pixel counts were plotted on the graph with error bars representing measurements

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with varying pixel tolerance values.

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Analysis of the reaction products of solid PLA depolymerization. Purified enzymes (50 µg)

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and enzyme-free controls were incubated with PLA10 powder (10-12 mg, Resomer® R 202 H

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poly (DL-lactide); Mw 10-18K, melting point 58 °C) in a reaction mixture (1 ml) containing 0.4

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M Tris-HCl buffer (pH 8.0) for 18 hr at 30 °C with shaking. Supernatant fractions were collected

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at different time points, passed through centrifugal filters (MWCO 10 kDa), and the produced

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lactic acid was measured using a lactate dehydrogenase (LDH) assay which enabled the detection

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of both D- and L- enantiomers of lactic acid with high sensitivity.41, 52 Values for detected lactate

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in enzyme-free controls were subtracted from the reported results for all enzyme-containing

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reactions. The L-lactate dehydrogenase (PfLDH) from Plasmodium falciparum53 and the D-

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lactate dehydrogenase (D-LDH3) from Lactobacillus jensenii54 were heterologously expressed in

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E. coli and affinity purified to near homogeneity as described earlier. Both LDH enzymes were

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added to the reaction mixture in excess (total 500 µg/ml, 50/50) to maintain the reaction rate in

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the first order with lactate concentration. For the analysis of oligomeric PLA products in

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supernatant fractions (passed through 10 kDa filters), the flow-through aliquots (90 µl) were

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treated for 5 min at 95 °C with 1 M NaOH (final concentration) to convert oligomeric PLA

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products to lactic acid monomers prior to LDH assay (the data were corrected for the presence of

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monomeric lactic acid before alkaline treatment). In addition, the filtered supernatant fractions

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from solid PLA reactions were analyzed using reverse phase liquid chromatography55 coupled

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with mass spectrometry (LC-MS) to identify the water-soluble products of PLA hydrolysis. The

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platform configuration and methodology were as described previously.41

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Protein crystallization and crystal structure determination of MGS0156. Purified

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MGS0156 (75-421 aa) was crystallized at room temperature using the sitting drop vapor diffusion

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method by mixing 1 µl of the selenomethionine substituted protein (12 mg/ml) with 1 µl of

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crystallization solution containing 30 % (w/v) PEG 4k, 0.2 M ammonium acetate, 0.1 M sodium

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citrate (pH 5.6), and 1/70 chymotrypsin. Crystals were harvested using mounted cryo-loops and

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transfered into the cryo-protectant (Paratone-N) prior to flash-freezing in liquid nitrogen. Data

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collections were carried out at the beamlines 19-ID of the Structural Biology Center, Advanced

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Photon Source, Argonne National Laboratory.56 The data set was collected from a single crystal

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to 1.95 Å at the wavelength of 0.9794 Å and processed using the program HKL300057 (Table S1).

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The structure of MGS0156 was determined by the Se-methionine SAD phasing, density

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modification, and initial model building as implemented in the PHENIX suite of programs.58 The

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initial models (~90% complete) were further built manually using the program COOT59 and

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refined with PHENIX. Analysis and validation of structures were performed using

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MOLPROBITY60 and COOT validation tools. The final model was refined to Rwork/Rfree =

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0.1532/0.19, and it shows good geometry with no outliers in the Ramachandran plot. Data

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collection and refinement statistics are summarized in Table S1. Surface electrostatic charge

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analysis was performed using the APBS tool in Pymol on a model generated by the PDB2PQR

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server.61, 62 The topology diagram of MGS0156 was generated by HERA program63 through

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PDBsum server.64 The atomic coordinates have been deposited in the Protein Data Bank, with

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accession code 5D8M.

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RESULTS AND DISCUSSION Screening of purified microbial hydrolases for polyesterase activity. To discover novel

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polyesterases, 213 randomly selected purified uncharacterized hydrolases from sequenced

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microbial genomes and environmental metagenomes (Table S2, also ref. 65) were screened for

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hydrolytic activity against emulsified PLA10 [poly (DL-lactide); Mw 10K], PLLA40 [poly(L-

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lactide); Mw 40K], polycaprolactone PCL10 (Mw 10K), and bis(benzoyloxyethyl) terephthalate

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(3PET) using agarose-based screens. 24, 65 The experimental conditions used in these screens (30

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°C, pH 8.0) were within the activity range for most known polyesterases from different organisms

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(cold-adapted, mesophilic, and thermophilic). These screens revealed the presence of detectable

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polyesterase activity in 36 proteins, mostly from the α/β hydrolase superfamily (Table S3). Most

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of these proteins were active against poly(DL-lactide) (22 proteins), followed by 3PET (13

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proteins) and PCL (11 proteins), whereas nine proteins exhibited activity toward poly(L-lactide)

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(PLLA40). At least 10 identified polyesterases exhibited high activity against multiple substrates

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(e.g. MGS0156 and GEN0105) (Table S3). Thus, a significant number of microbial and

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metagenomic hydrolases exhibit hydrolytic activity against synthetic polyesters.

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Since the metagenomic polyesterases MGS0156 (GenBank AKJ87264) and GEN0105

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(GenBank AKJ87216) showed high hydrolytic activity against several polyesters (PLA10,

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PCL10, and 3PET), the present work was focused on the biochemical and structural

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characterization of these proteins, which were also compared with the recently published

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metagenomics polyesterases GEN0160 and MGS0084 65 (Figure 1).

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Figure 1. Polyesterase activity of purified metagenomic carboxylesterases. Agarose-based screen of purified proteins for the presence of polyesterase activity against emulsified PCL10. The presence of polyesterase activity is indicated by the formation of a clear zone around the wells (A) with purified proteins (50 µg of protein/well, 72 hours at 30 °C). Agarose (1.5%) plates contained 0.2% emulsified PCL10 in 50 mM Tris-HCl (pH 8.0) buffer. Previously characterized enzymes (metagenomic polyesterases PlaM4 26, GEN0160 65, MGS0084 65, and porcine liver esterase (PLE)) were used as controls. (B) The bar graph represents the polyesterase activity of tested enzymes as clear halo areas in pixels obtained via image scanning.

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Carboxylesterase activity of the selected enzymes was initially identified using tributyrin-based

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esterase screens of the metagenomic gene libraries from an anaerobic urban waste degrading

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facility (GEN0105) or paper mill waste degrading microbial community (MGS0156). 65 The

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MGS0156 gene encodes a protein comprised of 421 amino acids with a potential N-terminal

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signal peptide (1-75 aa), whereas the GEN0105 sequence (322 aa) appears to lack an obvious

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signal peptide (Figure S1). Based on sequence analysis, both MGS0156 and GEN0105 belong to

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serine dependent α/β hydrolases but share low sequence identity to each other (21.1%). Both

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enzymes represent metagenomic proteins as GEN0105 shares 61% sequence identity with the

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predicted esterase B0L3I1_9BACT from an uncultured bacterium, whereas the closest

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homologue of MGS0156 (DesfrDRAFT_2296 from Desulfovibrio fructosivorans) shows 70%

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sequence identity to this protein (Figure S1). When compared to experimentally characterized

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proteins (Swiss-Prot database), GEN0105 showed the highest similarity to the monoterpene

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epsilon-lactone hydrolase MlhB from Rhodococcus erythropolis (85% query cover, 35%

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sequence identity), whereas MGS0156 did not show sequence homology to any characterized

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hydrolase.66 Initially, carboxylesterases and lipases were classified by Arpigny and Jaeger (1999)

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into 8 families based on amino acid sequence homology and biochemical properties, which were

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later expanded to include novel enzymes (currently over 18 families, some of which containing

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only one family member).67-70 Phylogenetic analysis revealed that GEN0105 is associated with

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esterase family IV, whereas MGS0156, MGS0084, and GEN0160 showed no clustering with

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known families of lipolytic enzymes, suggesting that these proteins represent new esterase

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families (Figure 2). Thus, the type II (lipase/cutinase type) polyesterases, including PLA

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depolymerases, exhibit broad phylogenetic diversity and are associated with esterase families I,

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III, IV, V as well as with new esterase families.

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Figure 2. Phylogenetic analysis of selected metagenomic polyesterases. Phylogenetic tree of polyesterases showing their relatedness to known esterase families (I – VIII, based on Arpigny and Jaeger, 1999).67 The phylogenetic tree was generated by MEGA7 software71 using the neighbor-joining method. The numbers on the nodes correspond to the percent recovery from 1,000 bootstrap resamplings. The evolutionary distances were calculated using the Poisson correction method72 and are in the units of the number of amino acid substitutions per site. GenBank accession numbers or Uniprot IDs are shown in parentheses.

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Carboxylesterase activity of MGS0156 and GEN0105 against soluble monoester

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substrates. The acyl chain length preferences of purified recombinant MGS0156 (75-421 aa) and

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GEN0105 were characterized using spectrophotometric assays with α-naphthyl and p-nitrophenyl

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(pNP) monoesters (Figure 3). For these substrates, MGS0156 was most active against pNP -

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octanoate and pNP -decanoate (C8-C10 substrates), as well as against pNP-palmitate (C16;

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Figure 3), which is in line with the lipolytic activity of this protein against olive oil observed in

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agar-based screens,65 indicating that it is a lipase-like enzyme. Compared to MGS0156, the

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specific activity of GEN0105 was an order of magnitude lower with a preference for shorter

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substrates (C4 and C5 substrates; Figure 3). With monoester substrates, both enzymes

308

demonstrated saturation kinetics with MGS0156 showing high catalytic efficiencies with low Km

309

values toward different substrates (kcat 88.8 – 1,101 s-1, specific activities 170 – 1,700 U/mg)

310

(Table S4, Figure 3). Based on our data, both MGS0156 and GEN0105 are amongst the most

311

active known polyesterases, for which a broad range of specific activities against model

312

monoesters have been reported (4.4 – 1,150 U/mg, kcat 6 – 660 s-1). 26, 27, 29, 30, 37, 73

15


Page 17 of 38


313 314 315 316 317 318

Figure 3. Esterase activity of metagenomic polyesterases against soluble monoester substrates with different acyl chain length. The reaction mixtures contained 0.5 mM p-nitrophenyl (pNP)- or 1.5 mM α-naphthyl (αN) esters with different chain lengths and 0.01 µg of purified MGS0156 (A) or GEN0105 (B). The white bars show activity against α-naphthyl esters, whereas the gray bars represent activity against pNP- substrates.

319 320

Based on temperature profiles of esterase activity, both MGS0156 and GEN0105 are

321

mesophilic carboxylesterases showing maximal activity between 35-40°C and retaining

322

approximately 20% of maximal activity at 5°C (Figure S2). This is similar to mesophilic

323

metagenomic polyesterases MGS0169 and GEN0160, whereas the cold-resistant polyesterase

324

MGS0084 was most active at 20°C and retained almost 50% of its maximal activity at 5°C.65

16



325

While both enzymes showed significant activity at alkaline reactions (pH < 10.5), MGS0156 was

326

more tolerant to high acidity retaining approximately 50% of its optimal activity at pH 2

327

compared to GEN0105 which was inactive at pH < 4 (Figure S2). In addition, MGS0156 and

328

GEN0105 showed similar sensitivity to inhibition by detergents (Triton X-100 and Tween 20),

329

whereas MGS0156 retained higher residual activity (25 - 75%) in the presence of salts (0.5 – 2.5

330

M NaCl or KCl) (Figure S2). Thus, with monoester substrates, MGS0156 and GEN0105 exhibit

331

different acyl chain length preferences and salt resistances yet similar sensitivities to temperature

332

and detergents. The biochemical diversity of identified polyesterases presents potential

333

technological advantages for applications in enzymatic degradation of various polyesters under

334

different reaction conditions (temperature, salts, detergents).

335

Hydrolytic activity of metagenomics polyesterases against 22 polyester substrates. The

336

polyester substrate ranges of purified MGS0156 and GEN0105 were determined using agarose-

337

based assays with 22 emulsified synthetic polyesters, including PLA, PCL, 3PET as well as

338

copolymers of various composition (Table 1). Since melting temperature (Tm) of the polymers

339

highly affect their enzymatic degradability,74 polyester substrates from a range of molecular

340

weight were subjected to enzymatic hydrolysis. Polyesterase activity of these enzymes was

341

compared with the activity of the recently identified metagenomic esterases GEN0160 and

342

MGS0084.65 Based on the diameter of clear zones produced in agarose-based screens, the four

343

metagenomic esterases exhibited polyesterase activity against emulsified PCL10, which was

344

higher (GEN0105, GEN0160 and MGS0156) or comparable (MGS0084) to that of the previously

345

identified polyesterase PlaM4 from compost (Figure 1).26 When screened against 22 emulsified

346

polyesters, MGS0156 and GEN0105 degraded 13 and 17 substrates, respectively, including PLA,

347

poly(D,L-lactide-co-glycolide) (PLGA), PCL, PBSA, and 3PET (Table 1). Both enzymes

348

hydrolyzed the majority of the tested PLA polymers, with GEN0105 displaying activity against

349

poly(L-lactide) and neither enzyme displaying activity against poly(D-lactide). Previously, it has 17


Page 18 of 38

Page 19 of 38


350

been shown that type I (protease) PLA depolymerases are specific toward poly(L-lactide), as

351

opposed to type II (cutinase/lipase) PLA depolymerases, which show preference for poly(DL-

352

lactide).17, 75 Besides GEN0105, only the cutinase-like type II enzyme CLE from Cryptococcus

353

sp. strain S-2 has been shown to be able to hydrolyze poly(L-lactide).17, 76 PLA substrates with the

354

acid end protected by the addition of an ester group were also hydrolyzed by MGS0156 and

355

GEN0105, suggesting that these polyesterases can exhibit endo-type hydrolysis. In contrast,

356

GEN0160 and MGS0084 showed no polyesterase activity against PLA substrates (except for

357

MGS0084 toward PLA2) and 3PET (Table 1). Finally, the four metagenomic esterases showed

358

no hydrolytic activity toward poly(D-lactide), PHB and PBS. Thus, GEN0105 appears to be the

359

most versatile polyesterase from the four tested enzymes, being able to hydrolyze a copolymer of

360

hydroxybutyric acid and hydroxyvaleric acid (PHBV), as well as the commercial polymer Ingeo™

361

PLA6400 from NatureWorks (Table 1).

362

363 364 365 366

Table 1. Polyester substrate profile of purified GEN0105, GEN0160, MGS0084, and MGS0156. The presence of hydrolytic activity (+) was inferred from the formation of a clear halo around the agarose wells containing the indicated enzymes. Polyesterase activity against PLA (D,L; Mw 2K) and PCL (Mw 10K) was also reported in our previous work. 65

367

Substrate

GEN0105

GEN0160

MGS0084

MGS0156

1. PLAa (D,L); Mw 2K

+

−

+

+

2. PLA (D,L); Mw 10K

+

−

−

+

3. PLA (D,L); Mw 10K, ester terminated

+

−

−

+


+

−

−

+


+

−

−

+

6. PLA (L); Mw 40K

+

−

−

−

18



Page 20 of 38

7. PLA (L); ester terminated

+

−

−

−

8. PLA (D); Mw 124K

−

−

−

−

9. Ingeo™ PLA6400

+

−

−

−

10. Ingeo™ PLA4032

−

−

−

−

11. PLGA

+

−

+

+

12. PHB

−

−

−

−

13. PHBV

+

−

−

−

14. PCL; Mw 10K

+

+

+

+

15. PCL; Mw 45K

+

+

+

+

16. PCL; Mw 70K

+

+

+

+

17. Bionolle™ PBS 1001MD

−

−

−

−

18. Bionolle™ PBS 1020MD

−

−

−

−

19. Bionolle™ PBSA 3001MD

+

+

+

+

20. Bionolle™ PBSA 3020MD

+

+

+

+

21. PES

+

+

+

+

22. 3PET

+

−

−

+

368 369 370 371

a

PLA, polylactic acid; PLGA, poly(D,L-lactide-co-glycolide); PHB, poly[(R)-3-hydroxybutyric acid]; PHBV, poly(3-hydroxybutyric acid-co-3-hydroxyvaleric acid); PCL, polycaprolactone; PBS, polybutylene succinate; PBSA, poly(butylene succinate-co-adipate); PES, poly(ethylene succinate); 3PET, bis(benzoyloxyethyl) terephthalate

372

373

Analysis of the reaction products of solid PLA hydrolysis. To demonstrate hydrolytic

374

activity of the identified metagenomic polyesterases against solid PLA substrates, purified

375

MGS0156 and GEN0105 were incubated with solid poly(DL-lactide) (Mw 10K) powder (12 mg)

376

suspended in 1 ml of 0.4 M Tris-HCl buffer (pH 8.0, equivalent lactate concentrations ~ 135

19


Page 21 of 38


377

mM). At indicated time points (Figure 4), the enzyme and solid PLA particles were removed

378

from the reaction mixture using centrifugal filters (MWCO 10 kDa), and the production of

379

monomeric and oligomeric lactic acid products was analyzed using L- and D-lactate

380

dehydrogenases (as described in Materials and Methods). After 6 hours of incubation at 30 °C,

381

MGS0156 hydrolyzed approximately 80% of the solid PLA substrate producing a mixture of

382

oligomeric and monomeric products (Figure 4).

383 384 385 386 387 388

Figure 4. Production of lactic acid during incubation of solid PLA10 with purified metagenomic polyesterases: wild-type MGS0156 (A), GEN0105 (B) and MGS0156 L169A (C) and cutinase Cut_2 from Thermobifida fusca 77 (D). Monomeric and oligomeric lactic acid products were measured using D- and L-lactate dehydrogenases as described in Materials and Methods. Results are means ± SD from at least two independent determinations.

389 390

The proportion of monomeric lactic acid product increased with longer incubation times resulting

391

in almost full (95%) conversion of solid PLA substrate (monomeric + oligomeric products) after

392

overnight incubation (Figure 4). GEN0105 degraded ~70% of solid PLA after overnight

20



Page 22 of 38

393

incubation and was able to produce significant amounts of lactic acid within the first 30 min of

394

incubation (Figure 4). Under the same experimental conditions, the well-known polyester-

395

degrading cutinase Cut_2 from T. fusca 77 hydrolyzed 8.5% of solid PLA during the first six

396

hours of incubation with 30% conversion (monomeric + oligomeric products) after 18 h (Figure

397

4). Interestingly, these three enzymes showed no significant hydrolytic activity against the

398

oligomeric PLA products (obtained using centrifugal filters). In addition, the formation of

399

significant amounts of oligomeric and monomeric lactate products during incubation of

400

MGS0156 and GEN0105 with solid PLA (Figure 4) suggests that they can catalyze both endo-

401

and exo-esterase cleavage of solid PLA.

402

Liquid chromatography-mass spectrometry (LC-MS) was used for direct analysis of water-

403

soluble reaction products from solid PLA hydrolysis by MGS0156 and GEN0105 (Figure S3).

404

The soluble reaction products were separated using a C18 column and analyzed using mass

405

spectrometry. These analyses revealed that both enzymes produced mixtures of lactic acid

406

monomers and oligomers with different chain lengths (Figure S3 and Table S5). In line with the

407

results of LDH-based assays, GEN0105 showed a higher degree of monomeric products

408

compared to lactic acid oligomers suggesting a higher exo-esterase activity of this enzyme

409

compared to MGS0156 (Figure 4).

410

Recently, we have found that the purified polyesterase ABO2449 from Alcanivorax

411

borkumensis required the addition of detergents (e.g. 0.1% Plysurf A210G) for solid PLA

412

hydrolysis, suggesting that detergents can facilitate protein binding to solid PLA.41 However, in

413

this work detergents (0.1% Plysurf A210G or Triton X-100) significantly reduced hydrolytic

414

activity of MGS0156 against solid PLA, and had no effect on polyesterase activity of GEN0105

415

(data not shown). With monoester substrates, GEN0105 retained significant catalytic activity in

21


Page 23 of 38


416

the presence of up to 20% detergent, whereas MGS0156 was much more sensitive to detergents

417

(Figure S2). Thus, metagenomic polyesterases show different kinds of responses to detergents.

418

Crystal structure and active site of MGS0156. Purified metagenomic esterases (GEN0105,

419

GEN0160, MGS0084, and MGS0156) were submitted for crystallization trials, with only

420

MGS0156 (75-421 aa) producing diffracting crystals (Materials and Methods). The crystal

421

structure of the seleno-methionine-substituted MGS0156 was solved at 1.95 Å resolution (Table

422

S1), and revealed a protomer with an α/β-hydrolase fold comprised of a slightly twisted central β-

423

sheet with seven parallel β-strands (-5x, -1x, 2x, (1x)3) and 19 α-helices (Figure 5A and Figure

424

S4). The predicted catalytic nucleophile Ser232 is positioned on a short sharp turn (the

425

nucleophilic elbow) between the β4 strand and α8 helix. It is located at the bottom of the

426

MGS0156 active site, which is partially covered by a ring-shaped lid domain formed by seven

427

short α-helices (α4, α10, α11, α14, α15, α16, and α18) connected by flexible loops (Figure 5A).

428

Analysis of the MGS0156 crystal contacts using the quaternary prediction server PISA

429

suggested that this protein may form tetramers in solution through dimerization of dimers

430

(Figures 5B, C). The tetrameric state of MGS0156 is consistent with the results of size-exclusion

431

chromatography, which revealed a predominance for MGS0156 tetramers (70%), as well as the

432

presence of some octomeric (25%) and monomeric (5%) forms (151 kDa, 296 kDa, and 40 kDa;

433

predicted Mw 39 kDa). The tightly packed MGS0156 dimer is created through multiple

434

interactions between residues located on several α-helices (α1, α2, α10, α13, and α16) and the β1

435

strand (buried area 4,100 Å2, surface area 24,590 Å2). The two MGS0156 dimers are assembled

436

into a tetramer via interactions between the α11, α15, and α18 helices (surface area 47,980 Å2,

437

buried area 9,400 Å2) (Figure 5C). In the MGS0156 tetramer, the four active sites are not adjacent

438

to each other and are separated from the monomer interfaces with the two active site cavities open

439

on the wide sides of the oligomeric assembly (Figure 5C).

22



Page 24 of 38

440 441 442 443 444 445 446 447 448

Figure 5. Crystal structure of MGS0156. (A) Overall fold of the MGS0156 protomer shown in three views related by a 90° rotation. The protein core β-sheet is shown in cyan with α-helices colored in grey, and the lid domain in magenta. The position of the active site is indicated by the side chain of the catalytic Ser232. (B) Two views of the MGS0156 dimer related by a 90° rotation. The two protomers are colored in cyan and magenta. (C) Two surface presentations of the protein tetramer shown in two views related by 90° rotation. The protomers are shown in different colors, and the active site openings are indicated by arrows. A structural homology search of the DALI and PDBeFold databases revealed hundreds of

449

structurally homologous proteins, mostly lipases and carboxylesterases with low overall sequence

450

similarity to MGS0156 (

Screening and characterization of novel polyesterases from

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