CHEMBIOCHEM COMMUNICATIONS DOI: 10.1002/cbic.201402166
Structural Insights into Enzymatic Degradation of Oxidized Polyvinyl Alcohol Yu Yang,[a] Tzu-Ping Ko,[c] Long Liu,[a] Jianghua Li,[a] Chun-Hsiang Huang,[b] HsiuChien Chan,[b] Feifei Ren,[b] Dongxu Jia,[a] Andrew H.-J. Wang,[c] Rey-Ting Guo,*[b] Jian Chen,*[a] and Guocheng Du*[a] The ever-increasing production and use of polyvinyl alcohol (PVA) threaten our environment. Yet PVA can be assimilated by microbes in two steps: oxidation and cleavage. Here we report novel a/b-hydrolase structures of oxidized PVA hydrolase (OPH) from two known PVA-degrading organisms, Sphingopyxis sp. 113P3 and Pseudomonas sp. VM15C, including complexes with substrate analogues, acetylacetone and caprylate. The active site is covered by a lid-like b-ribbon. Unlike other esterase and amidase, OPH is unique in cleaving the CC bond of b-diketone, although it has a catalytic triad similar to that of most a/b-hydrolases. Analysis of the crystal structures suggests a double-oxyanion-hole mechanism, previously only found in thiolase cleaving b-ketoacyl-CoA. Three mutations in the lid region showed enhanced activity, with potential in industrial applications.
by Fusarium lini B was first reported.[1] More extensive characterization of PVA-degrading microorganisms and the responsible enzymes followed over the next 40 years.[2] Recently, as the environmental threat posed by PVA usage continues to increase, microbial degradation of PVA is receiving much attention world-wide, and new PVA-degrading species continue to be identified.[3] The chemical structure of PVA comprises mainly repeated 1,3-diol units. Microbial degradation of PVA entails a two-step metabolism (Scheme 1). In the first step two neighboring alcohols are oxidized to form a di-ketone structure in the polymer.
Introduction Polyvinyl alcohol (PVA) has various desirable properties including tensile strength and thermostability, and has found wide applications in many industrial processes such as fabric and paper manufacturing. The production of PVA is ever increasing in response to sustained demand. At the same time, large quantities of consumed PVA are poured into water systems. Although said to be nontoxic, PVA exhibits strong surfactant activity, which might cause serious environmental problems. On the other hand, PVA is one of the few biodegradable polymers that can be processed by microbes. Early studies on PVA biodegradation date back nearly 80 years, when the degradation [a] Y. Yang,+ Dr. L. Liu, Prof. Dr. J. Li, Dr. D. Jia, Prof. Dr. J. Chen, Prof. Dr. G. Du Key Laboratory of Industrial Biotechnology, Ministry of Education, Jiangnan University Lihu Ave. 1800, Wuxi 214122 (China) E-mail: [email protected] [email protected] [b] Dr. C.-H. Huang, Dr. H.-C. Chan, F. Ren, Prof. Dr. R.-T. Guo Industrial Enzymes National Engineering Laboratory Tianjin Institute of Industrial Biotechnology, Chinese Academy of Sciences 32 West 7th Avenue, Tianjin Airport Economic Area, Tianjin 300308 (China) E-mail: [email protected] [c] Dr. T.-P. Ko,+ Prof. Dr. A. H.-J. Wang Institute of Biological Chemistry, Academia Sinica 128 Academia Road Section 2, Nankang, Taipei 11529 (Taiwan) [+] These authors contributed equally to this work. Supporting information for this article is available on the WWW under http://dx.doi.org/10.1002/cbic.201402166.
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Scheme 1. PVA degradation. Two parallel pathways of PVA oxidation are catalyzed by SAO and PDH, and the resulting OPA can be cleaved by aldolase or OPH.
The enzyme can be extracellular PVA oxidase or the periplasmic PVA dehydrogenase (PDH). PVA oxidase is a secondary alcohol oxidase (SAO) that uses oxygen as the oxidant and produces hydrogen peroxide. PDH is a member of the quinohemoprotein family, and is dependent on pyrroloquinoline quinone (PQQ) for electron transfer to cytochrome. In the second step oxidized PVA (OPA) is hydrolyzed by OPA hydrolase (OPH, also known as b-diketone hydrolase, BDH), or, in the case of monoketone OPA, undergoes an aldolase-type cleavage reaction. The polymer is progressively degraded by these enzymes into smaller fragments, and eventually it is converted into acetic acid, which enters the central metabolic pathway. Several microorganisms are able to utilize PVA as an energy source, including Sphingopyxis sp. 113P3 and Pseudomonas sp. VM15C. The OPA hydrolase is encoded by the oph gene in Sphingopyxis and by bdh in Pseudomonas, and the corresponding enzymes (sOPH and pOPH) share 65 % amino-acid sequence identity ChemBioChem 0000, 00, 1 – 5
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(see Figure S1 in the Supporting Information), and belong to the a/b-hydrolase superfamily. Enzymes in the a/b-hydrolase superfamily include peptidases and lipases that use a Ser-His-Asp catalytic triad to hydrolyze amide, ester, and related chemical bonds.[4] Among these, the closest to OPH in sequence are the bacterial depolymerases for polyhydroxyalkanoates (PHA). Despite low homology (~ 15 % identity), sequence comparison allowed the active site of OPH to be correctly identified. In a previous study we expressed sOPH in Pichia pastoris and characterized the recombinant protein biochemically.[5] Here we crystallized the protein, and subsequent determination of its structure by X-ray diffraction was carried out in parallel with the highly homologous pOPH expressed in Escherichia coli. Based on the crystal structures, several mutants were constructed and characterized (details in the Supporting Information). The results not only help explain the catalytic mechanism but also encourage other modifications of OPH for industrial use.
tion).[7] The other OPH structures were then solved by using MR (Table 1).[8] The overall structures of sOPH and pOPH are similar (Figure 1; RMSD 0.59 between 322 pairs of matched Ca atoms). Four pairs of cysteine are conserved, as well as a cis-proline (Figure S1). The largest difference is found at resi-
Overall Structure Due largely to the lack of an effective search model, our initial attempts to solve the crystal structures by molecular replacement (MR) failed. We then attempted to obtain mercury derivatives by mutating active-site Ser156 of sOPH and Ser172 of pOPH to cysteines, as before.[6] However, this strategy did not work. The structure was eventually solved by using a rhenium (K2ReCl6) derivative of pOPH (see the Supporting Informa-
Figure 1. OPH structure. The ribbons drawings of pOPH and sOPH are colored red for a-helices and yellow for b-strands, except for the two strands (I and II) that forms the lid (colored as for the loops: cyan for pOPH and green for sOPH). The active site (red asterisk), is adjacent to the lid. The side chains of the catalytic triad are shown as stick models.
Table 1. Data collection and refinement statistics of the OPH crystals.
Data collection space group unit cell a, b, c [] a, b, g [8] resolution [][a] unique reflections average redundancy completeness [%] average hIi/hs(I)i Rmerge [%][b] Refinement no. of reflections[c] Rwork (95 % data) Rfree (5 % data) RMSD bonds [][d] RMSD angles [8] Ramachandran[e] favored [%] allowed [%] outliers [%] Baverage [2]/atoms protein ligands water PDB ID
pOPH native
S172C/ACA
S172A/PNPC
sOPH
P1 49.9, 58.3, 65.4 89.6, 73.2, 75.8 25–1.60 (1.66–1.60) 85 743 (8502) 3.2 (3.2) 95.6 (94.3) 22.1 (5.5) 5.3 (22.1)
P212121 58.6, 65.5, 84.4 90.0, 90.0, 90.0 25–1.67 (1.73–1.67) 37 918 (3681) 7.6 (6.9) 99.3 (98.1) 41.7 (8.0) 5.9 (18.9)
P212121 58.4, 65.3, 84.0 90.0, 90.0, 90.0 25–1.60 (1.66–1.60) 43 062 (4191) 6.4 (6.3) 99.7 (99.1) 23.5 (3.5) 8.2 (50.3)
P1 58.3, 75.7, 76.0 74.0, 88.7, 74.0 25–1.90 (1.97–1.90) 92 945 (9105) 3.0 (3.0) 97.8 (96.5) 26.2 (2.9) 5.6 (51.3)
83 750 (7572) 0.152 (0.209) 0.175 (0.224) 0.013 1.62
37 449 (3335) 0.158 (0.189) 0.190 (0.240) 0.020 1.88
41 325 (3752) 0.148 (0.224) 0.174 (0.255) 0.016 1.70
88 575 (7876) 0.193 (0.299) 0.239 (0.334) 0.018 1.84
97.7 2.3 0
97.0 3.0 0
97.3 2.7 0
96.8 2.9 0.3
16.1/5003 36.8/13 35.2/1052 3WL6
15.7/2580 29.2/20 26.9/362 3WL7
11.7/2577 19.6/23 30.8/688 3WL8
31.0/7542 31.7/20 40.8/834 3WLA
[a] Numbers in parentheses are for the outermost resolution shells. [b] Rmerge = hkli j Ii(hkl)hI(hkl)i j /hkliIi(hkl), in which the sum is over all the i measured reflections with equivalent miller indices hkl; hI(hkl)i is the averaged intensity of these i reflections, and the grand sum is over all measured reflections in the data set. [c] All positive reflections were used in the refinement. [d] According to Engh and Huber.[11a] [e] Calculated by using MolProbity.[11b]
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dues 239–249 in sOPH (255–268 in pOPH), a region that corresponds to a lid-like protrusion over the active site and comprises a b-ribbon (bI/bII). The OPH structures show similarly to other a/b-hydrolases. A DALI search returned a number of esterases with RMSD of 2.5–3.0 for 200–220 aligned residues in the catalytic domain with 14–16 % identity to OPH. Structural comparison with a PHA depolymerase that acts on polyhydroxybutyrate (PHB; PDB ID: 2D81)[9] showed an RMSD of 1.65 between 162 pairs of matched Ca atoms within 3.5 (Figure S2), among which 24 amino acids are identical. Unlike PHB depolymerase and other esterases, which cleave the CO or CN bond adjacent to a carbonyl group, OPH hydrolyzes the CC bond (Scheme 1). To investigate the catalytic mechanism, we obtained pOPH-complex crystals by using the substrate analogues acetylacetone (ACA) and p-nitrophenyl caprylate (PNPC). The PNP-derived ester had been used previously in activity measurement.[5] For clarity, residue numbers here refer to those of pOPH unless indicated otherwise. Active-site environment OPH has a more confined active-site cleft than does PHB depolymerase, mainly because of the presence of a covering lid structure. The catalytic triad of pOPH comprises Ser172, Asp253, and His298, which are embedded in this cleft (Figure 1). ACA was bound here, as was PNPC, although the PNP group was cleaved off and lost because the S172A mutant used for this crystallization retained ~ 20 % activity (Table 2; ligand electron densities in Figure S3). ACA stacks with Trp121 and its distal carbonyl group makes a hydrogen bond with Asn120. The other carbonyl is directed toward Tyr200 and the proximal methyl group is 3.5 from the sulfonate of Cys172 (Figure 2 A). The hydrocarbon moiety of caprylate also contacts the Trp121 side chain. The carboxyl group,
Figure 2. Ligand interactions. A) S172C/ACA crystal. The side chain of Cys172 is oxidized. B) S172A/PNPC crystal. Only the caprylate moiety (not PNP) was observed. Protein carbon atoms are colored green, bound ligands are in cyan, and hydrogen bonds are pink dashed lines.
3.1 from the Ala172 side chain, interacts with Ser66, Tyr270, and His298, and tends to stack Trp255 of the lid (Figure 2 B). Neither Trp121 nor Trp255 is found in PHB depolymerase, where the substrate is bound by a slightly different mode (Figure S2). The lid was disordered in a monomer (B) of the native pOPH crystal, which contained a citrate bound to the backbone nitrogens of Ser66 and Val67, whereas the sOPH crystal had a bound sulfate (Figure S4). The lid appears to be very flexible, without proper binding to substrate or analogue. The anion-binding propensity of the b3-a1 loop might account for the unexpected Cys172 oxidation. The loop also plays an important role in catalysis (see below). Mutant effects Next, some mutants in the lid region were produced and analyzed for OPH activity to investigate the catalysis. First, the authentic substrate OPA was used (Table 2; details in the Supporting Information). Trp255, which covers the active site, is
Table 2. Kinetics parameters with OPA as the substrate.[a]
sOPH wild-type S156C pOPH wild-type S172C S172A W255Y W255F W255A C257A/C267A R264A Y270F Y270A
Km [mm]
kcat [s1]
kcat/Km [s1 mm1]
Ratio [%]
0.42 0.01 8.7 0.2
24.5 0.6 51 2
58 3 5.8 0.3
100 9.9
0.52 0.01 2.82 0.08 1.25 0.02 0.73 0.01 1.15 0.03 1.95 0.07 1.04 0.03 0.40 0.01 0.48 0.01 2.42 0.03
31.7 0.6 15.0 0.4 15.9 0.4 70 1 21.2 0.3 53.4 0.7 12.4 0.3 35.3 0.7 41.6 0.7 41 1
62 2 5.3 0.3 12.7 0.1 96 4 18.5 0.6 27 1 11.9 0.6 88 4 87 1 16.8 0.6
100 8.6 20.7 156 30.1 44.6 19.3 142 141 27.3
[a] All activity measurements were carried out at 37 8C and pH 7.5 according to Klomklang et al.[12] Substrate concentrations were based on the estimated number of carbonyl groups in a fixed amount of polymer. Each value represents the mean of data obtained from independent Michaelis– Menten curve fitting calculations after triplicate measurements with various substrate concentrations.
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Table 3. Kinetics parameters with PNPC as the substrate.[a]
sOPH wild-type S156C pOPH wild-type S172C S172A W255Y W255F W255A C257A/C267A R264A Y270F Y270A
Km [mm]
kcat [s1]
kcat/Km [s1 mm1]
Ratio [%]
2.2 0.1 5.8 0.3
390 20 53 3
175 10 9.2 0.7
100 5.3
2.15 0.07 9.6 0.5 6.1 0.2 2.34 0.09 1.9 0.1 2.6 0.1 3.2 0.2 1.36 0.05 1.94 0.08 2.4 0.1
470 20 184 8 136 6 570 30 400 20 600 30 244 8 410 10 660 30 650 30
216 9 19.3 0.4 22.5 0.3 250 10 212 5 230 10 76 3 300 8 340 10 270 10
100 8.9 10.4 113 98 106 35 139 158 126
[a] All activity measurements were carried out at 37 8C and pH 8.0 according to Winkler et al.[5b] Each value represents the mean of data obtained from independent Michaelis–Menten curve fitting calculations after triplicate measurements with various substrate concentrations.
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important for substrate affinity. Mutations to Tyr, Phe, or Ala reing of the active site by Trp255, and consequently both Km and sulted in higher Km but with varied kcat. In particular, the kcat were not changed significantly when replacing or removing W255Y mutant showed activity 60 % higher than wild-type. its side chain. Y270F showed enhanced activity for PNPC, thus The amphipathic but less bulky side chain of Tyr255 would faagain suggesting that the phenolic OH of Tyr270 is not incilitate substrate binding and product release, yet it is still efvolved in catalysis. The retained esterase activity in Y270A indifective in shielding the active site from solvent. cates that “proper” formation of the OPA binding site is not reAlthough Tyr270 formed a hydrogen bond to caprylate in quired for PNPC binding. the S172A/PNPC structure, the side-chain hydroxyl group is not directly involved in catalysis, because mutant Y270F Substrate modeling and catalytic mechanism showed 40 % higher activity. However, the significant decrease in substrate affinity of Y270A suggests that the aromatic side Finally, based on the bound ACA and caprylate structures, chain of Tyr270 participates in maintaining the OPA binding a 13-carbon model of OPA was constructed to elucidate site. The Cys257–Cys267 disulfide bond is probably important enzyme–substrate interactions (see the Supporting Informain maintaining the lid structure, because mutating either resition). It contained six carbonyl (CO) groups (numbered 4, 3, 2, due to alanine had an adverse effect in terms of both Km and 1, 1’ and 2’ in Figure 3 A). Several hydrogen bonds to the COs are likely; CO4 and CO3 stack with Trp121 as does ACA. The kcat. Trp255, Tyr270, and Cys257–Cys267 are conserved, and cleavage site is between CO1 and CO1’; both have to be carthey presumably serve the same roles in pOPH and sOPH. With mutations further from the active site, pOPH R264A showed improved Km and kcat. Arg264 (not found in sOPH) is in the variable region of the lid. Apparently its positively charged side chain is dispensable for catalysis. Likewise, by mutating other nonessential lid residues, we might be able to obtain sOPH (or pOPH) with even better Km and kcat. In contrast, the active-site mutant S172C showed less than 10 % activity (higher Km and lower kcat). Presumably the oxidized Cys172 not only lost its function as a nucleophile but also hindered substrate binding. More activity was retained in mutant S172A, in which an activated water molecule comes directly into play. Both enzymes and the mutants were then analyzed for esterase activity with PNPC as the substrate (Table 3 and Supporting Information). The higher Km values indicate lower affinity to this artificial substrate, but the overall activity was enhanced (much higher kcat values). Apparently the bulky PNP head group is not well accommodated in the active-site cleft, but the ester bond is more readily cleaved. The mutants had generally similar effects, except for mutations Figure 3. Substrate binding and catalysis. A) The modeled OPA is shown as thick sticks, and the protein is shown to Trp255 and Tyr270. Hydrolysis as thin sticks (both in green). Potential hydrogen bonds are pink dashed lines. For comparison, the S172C/ACA of the ester CO bond in PNPC (cyan) and S172A/PNPC (gray) structures are also shown. B) Proposed catalytic pathway for OPH. The backbone probably does not need shield- NH of Ser66 might also interact with CO1. 2014 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
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CHEMBIOCHEM COMMUNICATIONS bonyls in order to be cleaved, whereas the others can be alcohols. In this model, the O atom of CO1 matches a sulfonate O of Cys172. It is hydrogen bonded to Ser66 and Ser173, which presumably constitute an oxyanion hole (Figure 3 A). The C atom is 2.7 from the OG atom of Ser172, which makes a nucleophilic attack when deprotonated by His298. The proton is then transferred to the methylene group, now connected only to CO1’. Finally, His298 deprotonates a water molecule to cleave the acyl–enzyme intermediate (Figure 3 B). Similar enzyme—substrate interactions are observed in PHB depolymerase.[9] In addition to the nucleophile Ser39, the base His155, and the oxyanion hole formed by Ser40 and Cys250, the side chain of Trp307 is also hydrogen bonded to a carbonyl group of PHB (equivalent to CO3). The reaction catalyzed by OPH is typical of a/b-hydrolases, except that the cleaved bond is between two carbon atoms. Here electron delocalization (in a way like enol–ketone equilibrium, which does occur in ACA), seems to play an essential role. Upon CC bond breaking, the negative charge on CO1’ is probably stabilized by hydrogen bonds to the backbone NH of Ser66 and Val67, thus forming a second oxyanion hole. As mentioned above, this anion-binding b3-a1 loop might promote spontaneous oxidation of Cys172 to a sulfonate. Although Ser66 and Ser173 in pOPH are Thr50 and Ala157 in sOPH, the potential to form an oxyanion hole is conserved. The employment of double oxyanion holes in catalysis has been observed in thiolase, which also cleaves the CC bond between two carbonyl groups.[10] Interestingly, the catalytic Cys123 in the native thiolase crystal was also found oxidized to a sulfonate.
Conclusion OPH crystal structures have allowed us to elucidate its catalytic mechanism. They also served as a basis for rational modifications (particularly in the lid region) to endow the enzyme with desired characteristics. The three lid mutants, W255Y, R264A, and Y270F, showed 40–60 % improvement in kcat/Km. The activity enhancement can be employed to facilitate bacterial assimilation of OPA. We are currently making double and triple mutants to see if there are synergistic effects. However, OPH is responsible for the second step of PVA degradation, which starts with a dehydrogenase or oxidase. Both remain to be further characterized, but PVA oxidase might find better applications because it is probably located on the cell surface and does not need the cofactor PQQ.[3a] Solving its crystal structure would thus be highly beneficial.
Acknowledgements This work was supported by the Program for Changjiang Scholars and Innovative Research Team in University (No. IRT1135), the National High Technology Research and Development Program
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www.chembiochem.org of China (863 Program, 2012AA022202), the Priority Academic Program Development of Jiangsu Higher Education Institutions, the 111 Project (111-2-06). We thank the National Synchrotron Radiation Research Center of Taiwan for beam-time allocation and data-collection assistance. Keywords: catalytic triad · beta-diketone hydrolases · double oxyanion holes · environmental chemistry · hydrolases · microbial assimilation
[1] F. F. Nord, Naturwissenschaften 1936, 24, 763. [2] a) T. Suzuki, Y. Ichihara, M. Yamada, K. Tonomura, Agric. Biol. Chem. 1973, 37, 747 – 756; b) Y. Watanabe, M. Morita, N. Hamada, Y. Tsujisaka, Agric. Biol. Chem. 1975, 39, 2447 – 2448. [3] a) A. Yamatsu, R. Matsumi, H. Atomi, T. Imanaka, Appl. Microbiol. Biotechnol. 2006, 72, 804 – 811; b) F. Kawai, X. Hu, Appl. Microbiol. Biotechnol. 2009, 84, 227 – 237. [4] A. Marchler-Bauer, C. Zheng, F. Chitsaz, M. K. Derbyshire, L. Y. Geer, R. C. Geer, N. R. Gonzales, M. Gwadz, D. I. Hurwitz, C. J. Lanczycki, F. Lu, S. Lu, G. H. Marchler, J. S. Song, N. Thanki, R. A. Yamashita, D. Zhang, S. H. Bryant, Nucleic Acids Res. 2013, 41, D348 – D352. [5] a) Y. Yang, L. Liu, J. Li, G. Du, J. Chen, Bioprocess Biosyst. Eng. 2014, 37, 777 – 782; b) U. K. Winkler, M. Stuckmann, J. Bacteriol. 1979, 138, 663 – 670. [6] a) Y.-S. Cheng, T.-P. Ko, T.-H. Wu, Y. Ma, C.-H. Huang, H.-L. Lai, A. H.-J. Wang, J.-R. Liu, R.-T. Guo, Proteins Struct. Funct. Bioinf. 2011, 79, 1193 – 1204; b) F. Ren, T.-P. Ko, X. Feng, C.-H. Huang, H.-C. Chan, Y. Hu, K. Wang, Y. Ma, P.-H. Liang, A. H.-J. Wang, E. Oldfield, R.-T. Guo, Angew. Chem. Int. Ed. 2012, 51, 4157 – 4160; Angew. Chem. 2012, 124, 4233 – 4236. [7] Phase angles were calculated by employing OASIS: a) T. Zhang, Y. He, J. W. Wang, L. J. Wu, C. D. Zheng, Q. Hao, Y. X. Gu, H. F. Fan, OASIS, Institute of Physics, Chinese Academy of Sciences, P. R. China, 2012, http:// cryst.iphy.ac.cn; SHELXD: b) T. R. Schneider, G. M. Sheldrick, Acta Crystallogr. Sect. D Biol. Crystallogr. 2002, 58, 1772 – 1779; SOLVE/RESOLVE: c) T. C. Terwilliger, Methods Enzymol. 2003, 374, 22 – 37. Initial model building used ARP/wARP: d) A. Perrakis, R. Morris, V. S. Lamzin, Nat. Struct. Biol. 1999, 6, 458 – 463. [8] Molecular replacement calculations used Phaser: a) A. J. McCoy, R. W. Grosse-Kunstleve, P. D. Adams, M. D. Winn, L. C. Storoni, R. J. Read, J. Appl. Crystallogr. 2007, 40, 658 – 674. Manual model adjustments used Coot: b) P. Emsley, K. Cowtan, Acta Crystallogr. Sect. D Biol. Crystallogr. 2004, 60, 2126 – 2132. Computational refinement employed CNS: c) A. T. Brnger, P. D. Adams, G. M. Clore, W. L. DeLano, P. Gros, R. W. GrosseKunstleve, J. S. Jiang, J. Kuszewski, M. Nilges, N. S. Pannu, R. J. Read, L. M. Rice, T. Simonson, G. L. Warren, Acta Crystallogr. Sect. D Biol. Crystallogr. 1998, 54, 905 – 921. [9] T. Hisano, K.-i. Kasuya, Y. Tezuka, N. Ishii, T. Kobayashi, M. Shiraki, E. Oroudjev, H. Hansma, T. Iwata, Y. Doi, T. Saito, K. Miki, J. Mol. Biol. 2006, 356, 993 – 1004. [10] R. K. Harijan, T. R. Kiema, M. P. Karjalainen, N. Janardan, M. R. N. Murthy, M. S. Weiss, P. A. M. Michels, R. K. Wierenga, Biochem. J. 2013, 455, 119 – 130. [11] a) R. A. Engh, R. Huber, Acta Crystallogr. Sect. A Found. Crystallogr. 1991, 47, 392 – 400; b) V. B. Chen, W. B. Arendall III, J. J. Headd, D. A. Keedy, R. M. Immormino, G. J. Kapral, L. W. Murray, J. S. Richardson, D. C. Richardson, Acta Crystallogr. Sect. D Biol. Crystallogr. 2010, 66, 12 – 21. [12] W. Klomklang, A. Tani, K. Kimbara, R. Mamota, T. Ueda, M. Shimao, F. Kawai, Microbiology 2005, 151, 1255 – 1262. Received: April 11, 2014 Published online on && &&, 0000
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COMMUNICATIONS Y. Yang, T.-P. Ko, L. Liu, J. Li, C.-H. Huang, H.-C. Chan, F. Ren, D. Jia, A. H.-J. Wang, R.-T. Guo,* J. Chen,* G. Du* && – && Structural Insights into Enzymatic Degradation of Oxidized Polyvinyl Alcohol
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Crystal-clear hydrolysis: The crystal structures of OPH with bound inhibitors from two PVA-degrading microbes were analyzed to elucidate the hydrolytic mechanism of oxi-PVA. Several site-directed mutants were also constructed to enhance the OPH activity.
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Structural Insights into Enzymatic Degradation of Oxidized Polyvinyl Alcohol Yu Yang,[a] Tzu-Ping Ko,[c] Long Liu,[a] Jianghua Li,[a] Chun-Hsiang Huang,[b] HsiuChien Chan,[b] Feifei Ren,[b] Dongxu Jia,[a] Andrew H.-J. Wang,[c] Rey-Ting Guo,*[b] Jian Chen,*[a] and Guocheng Du*[a] The ever-increasing production and use of polyvinyl alcohol (PVA) threaten our environment. Yet PVA can be assimilated by microbes in two steps: oxidation and cleavage. Here we report novel a/b-hydrolase structures of oxidized PVA hydrolase (OPH) from two known PVA-degrading organisms, Sphingopyxis sp. 113P3 and Pseudomonas sp. VM15C, including complexes with substrate analogues, acetylacetone and caprylate. The active site is covered by a lid-like b-ribbon. Unlike other esterase and amidase, OPH is unique in cleaving the CC bond of b-diketone, although it has a catalytic triad similar to that of most a/b-hydrolases. Analysis of the crystal structures suggests a double-oxyanion-hole mechanism, previously only found in thiolase cleaving b-ketoacyl-CoA. Three mutations in the lid region showed enhanced activity, with potential in industrial applications.
by Fusarium lini B was first reported.[1] More extensive characterization of PVA-degrading microorganisms and the responsible enzymes followed over the next 40 years.[2] Recently, as the environmental threat posed by PVA usage continues to increase, microbial degradation of PVA is receiving much attention world-wide, and new PVA-degrading species continue to be identified.[3] The chemical structure of PVA comprises mainly repeated 1,3-diol units. Microbial degradation of PVA entails a two-step metabolism (Scheme 1). In the first step two neighboring alcohols are oxidized to form a di-ketone structure in the polymer.
Introduction Polyvinyl alcohol (PVA) has various desirable properties including tensile strength and thermostability, and has found wide applications in many industrial processes such as fabric and paper manufacturing. The production of PVA is ever increasing in response to sustained demand. At the same time, large quantities of consumed PVA are poured into water systems. Although said to be nontoxic, PVA exhibits strong surfactant activity, which might cause serious environmental problems. On the other hand, PVA is one of the few biodegradable polymers that can be processed by microbes. Early studies on PVA biodegradation date back nearly 80 years, when the degradation [a] Y. Yang,+ Dr. L. Liu, Prof. Dr. J. Li, Dr. D. Jia, Prof. Dr. J. Chen, Prof. Dr. G. Du Key Laboratory of Industrial Biotechnology, Ministry of Education, Jiangnan University Lihu Ave. 1800, Wuxi 214122 (China) E-mail: [email protected] [email protected] [b] Dr. C.-H. Huang, Dr. H.-C. Chan, F. Ren, Prof. Dr. R.-T. Guo Industrial Enzymes National Engineering Laboratory Tianjin Institute of Industrial Biotechnology, Chinese Academy of Sciences 32 West 7th Avenue, Tianjin Airport Economic Area, Tianjin 300308 (China) E-mail: [email protected] [c] Dr. T.-P. Ko,+ Prof. Dr. A. H.-J. Wang Institute of Biological Chemistry, Academia Sinica 128 Academia Road Section 2, Nankang, Taipei 11529 (Taiwan) [+] These authors contributed equally to this work. Supporting information for this article is available on the WWW under http://dx.doi.org/10.1002/cbic.201402166.
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Scheme 1. PVA degradation. Two parallel pathways of PVA oxidation are catalyzed by SAO and PDH, and the resulting OPA can be cleaved by aldolase or OPH.
The enzyme can be extracellular PVA oxidase or the periplasmic PVA dehydrogenase (PDH). PVA oxidase is a secondary alcohol oxidase (SAO) that uses oxygen as the oxidant and produces hydrogen peroxide. PDH is a member of the quinohemoprotein family, and is dependent on pyrroloquinoline quinone (PQQ) for electron transfer to cytochrome. In the second step oxidized PVA (OPA) is hydrolyzed by OPA hydrolase (OPH, also known as b-diketone hydrolase, BDH), or, in the case of monoketone OPA, undergoes an aldolase-type cleavage reaction. The polymer is progressively degraded by these enzymes into smaller fragments, and eventually it is converted into acetic acid, which enters the central metabolic pathway. Several microorganisms are able to utilize PVA as an energy source, including Sphingopyxis sp. 113P3 and Pseudomonas sp. VM15C. The OPA hydrolase is encoded by the oph gene in Sphingopyxis and by bdh in Pseudomonas, and the corresponding enzymes (sOPH and pOPH) share 65 % amino-acid sequence identity ChemBioChem 0000, 00, 1 – 5
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(see Figure S1 in the Supporting Information), and belong to the a/b-hydrolase superfamily. Enzymes in the a/b-hydrolase superfamily include peptidases and lipases that use a Ser-His-Asp catalytic triad to hydrolyze amide, ester, and related chemical bonds.[4] Among these, the closest to OPH in sequence are the bacterial depolymerases for polyhydroxyalkanoates (PHA). Despite low homology (~ 15 % identity), sequence comparison allowed the active site of OPH to be correctly identified. In a previous study we expressed sOPH in Pichia pastoris and characterized the recombinant protein biochemically.[5] Here we crystallized the protein, and subsequent determination of its structure by X-ray diffraction was carried out in parallel with the highly homologous pOPH expressed in Escherichia coli. Based on the crystal structures, several mutants were constructed and characterized (details in the Supporting Information). The results not only help explain the catalytic mechanism but also encourage other modifications of OPH for industrial use.
tion).[7] The other OPH structures were then solved by using MR (Table 1).[8] The overall structures of sOPH and pOPH are similar (Figure 1; RMSD 0.59 between 322 pairs of matched Ca atoms). Four pairs of cysteine are conserved, as well as a cis-proline (Figure S1). The largest difference is found at resi-
Overall Structure Due largely to the lack of an effective search model, our initial attempts to solve the crystal structures by molecular replacement (MR) failed. We then attempted to obtain mercury derivatives by mutating active-site Ser156 of sOPH and Ser172 of pOPH to cysteines, as before.[6] However, this strategy did not work. The structure was eventually solved by using a rhenium (K2ReCl6) derivative of pOPH (see the Supporting Informa-
Figure 1. OPH structure. The ribbons drawings of pOPH and sOPH are colored red for a-helices and yellow for b-strands, except for the two strands (I and II) that forms the lid (colored as for the loops: cyan for pOPH and green for sOPH). The active site (red asterisk), is adjacent to the lid. The side chains of the catalytic triad are shown as stick models.
Table 1. Data collection and refinement statistics of the OPH crystals.
Data collection space group unit cell a, b, c [] a, b, g [8] resolution [][a] unique reflections average redundancy completeness [%] average hIi/hs(I)i Rmerge [%][b] Refinement no. of reflections[c] Rwork (95 % data) Rfree (5 % data) RMSD bonds [][d] RMSD angles [8] Ramachandran[e] favored [%] allowed [%] outliers [%] Baverage [2]/atoms protein ligands water PDB ID
pOPH native
S172C/ACA
S172A/PNPC
sOPH
P1 49.9, 58.3, 65.4 89.6, 73.2, 75.8 25–1.60 (1.66–1.60) 85 743 (8502) 3.2 (3.2) 95.6 (94.3) 22.1 (5.5) 5.3 (22.1)
P212121 58.6, 65.5, 84.4 90.0, 90.0, 90.0 25–1.67 (1.73–1.67) 37 918 (3681) 7.6 (6.9) 99.3 (98.1) 41.7 (8.0) 5.9 (18.9)
P212121 58.4, 65.3, 84.0 90.0, 90.0, 90.0 25–1.60 (1.66–1.60) 43 062 (4191) 6.4 (6.3) 99.7 (99.1) 23.5 (3.5) 8.2 (50.3)
P1 58.3, 75.7, 76.0 74.0, 88.7, 74.0 25–1.90 (1.97–1.90) 92 945 (9105) 3.0 (3.0) 97.8 (96.5) 26.2 (2.9) 5.6 (51.3)
83 750 (7572) 0.152 (0.209) 0.175 (0.224) 0.013 1.62
37 449 (3335) 0.158 (0.189) 0.190 (0.240) 0.020 1.88
41 325 (3752) 0.148 (0.224) 0.174 (0.255) 0.016 1.70
88 575 (7876) 0.193 (0.299) 0.239 (0.334) 0.018 1.84
97.7 2.3 0
97.0 3.0 0
97.3 2.7 0
96.8 2.9 0.3
16.1/5003 36.8/13 35.2/1052 3WL6
15.7/2580 29.2/20 26.9/362 3WL7
11.7/2577 19.6/23 30.8/688 3WL8
31.0/7542 31.7/20 40.8/834 3WLA
[a] Numbers in parentheses are for the outermost resolution shells. [b] Rmerge = hkli j Ii(hkl)hI(hkl)i j /hkliIi(hkl), in which the sum is over all the i measured reflections with equivalent miller indices hkl; hI(hkl)i is the averaged intensity of these i reflections, and the grand sum is over all measured reflections in the data set. [c] All positive reflections were used in the refinement. [d] According to Engh and Huber.[11a] [e] Calculated by using MolProbity.[11b]
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dues 239–249 in sOPH (255–268 in pOPH), a region that corresponds to a lid-like protrusion over the active site and comprises a b-ribbon (bI/bII). The OPH structures show similarly to other a/b-hydrolases. A DALI search returned a number of esterases with RMSD of 2.5–3.0 for 200–220 aligned residues in the catalytic domain with 14–16 % identity to OPH. Structural comparison with a PHA depolymerase that acts on polyhydroxybutyrate (PHB; PDB ID: 2D81)[9] showed an RMSD of 1.65 between 162 pairs of matched Ca atoms within 3.5 (Figure S2), among which 24 amino acids are identical. Unlike PHB depolymerase and other esterases, which cleave the CO or CN bond adjacent to a carbonyl group, OPH hydrolyzes the CC bond (Scheme 1). To investigate the catalytic mechanism, we obtained pOPH-complex crystals by using the substrate analogues acetylacetone (ACA) and p-nitrophenyl caprylate (PNPC). The PNP-derived ester had been used previously in activity measurement.[5] For clarity, residue numbers here refer to those of pOPH unless indicated otherwise. Active-site environment OPH has a more confined active-site cleft than does PHB depolymerase, mainly because of the presence of a covering lid structure. The catalytic triad of pOPH comprises Ser172, Asp253, and His298, which are embedded in this cleft (Figure 1). ACA was bound here, as was PNPC, although the PNP group was cleaved off and lost because the S172A mutant used for this crystallization retained ~ 20 % activity (Table 2; ligand electron densities in Figure S3). ACA stacks with Trp121 and its distal carbonyl group makes a hydrogen bond with Asn120. The other carbonyl is directed toward Tyr200 and the proximal methyl group is 3.5 from the sulfonate of Cys172 (Figure 2 A). The hydrocarbon moiety of caprylate also contacts the Trp121 side chain. The carboxyl group,
Figure 2. Ligand interactions. A) S172C/ACA crystal. The side chain of Cys172 is oxidized. B) S172A/PNPC crystal. Only the caprylate moiety (not PNP) was observed. Protein carbon atoms are colored green, bound ligands are in cyan, and hydrogen bonds are pink dashed lines.
3.1 from the Ala172 side chain, interacts with Ser66, Tyr270, and His298, and tends to stack Trp255 of the lid (Figure 2 B). Neither Trp121 nor Trp255 is found in PHB depolymerase, where the substrate is bound by a slightly different mode (Figure S2). The lid was disordered in a monomer (B) of the native pOPH crystal, which contained a citrate bound to the backbone nitrogens of Ser66 and Val67, whereas the sOPH crystal had a bound sulfate (Figure S4). The lid appears to be very flexible, without proper binding to substrate or analogue. The anion-binding propensity of the b3-a1 loop might account for the unexpected Cys172 oxidation. The loop also plays an important role in catalysis (see below). Mutant effects Next, some mutants in the lid region were produced and analyzed for OPH activity to investigate the catalysis. First, the authentic substrate OPA was used (Table 2; details in the Supporting Information). Trp255, which covers the active site, is
Table 2. Kinetics parameters with OPA as the substrate.[a]
sOPH wild-type S156C pOPH wild-type S172C S172A W255Y W255F W255A C257A/C267A R264A Y270F Y270A
Km [mm]
kcat [s1]
kcat/Km [s1 mm1]
Ratio [%]
0.42 0.01 8.7 0.2
24.5 0.6 51 2
58 3 5.8 0.3
100 9.9
0.52 0.01 2.82 0.08 1.25 0.02 0.73 0.01 1.15 0.03 1.95 0.07 1.04 0.03 0.40 0.01 0.48 0.01 2.42 0.03
31.7 0.6 15.0 0.4 15.9 0.4 70 1 21.2 0.3 53.4 0.7 12.4 0.3 35.3 0.7 41.6 0.7 41 1
62 2 5.3 0.3 12.7 0.1 96 4 18.5 0.6 27 1 11.9 0.6 88 4 87 1 16.8 0.6
100 8.6 20.7 156 30.1 44.6 19.3 142 141 27.3
[a] All activity measurements were carried out at 37 8C and pH 7.5 according to Klomklang et al.[12] Substrate concentrations were based on the estimated number of carbonyl groups in a fixed amount of polymer. Each value represents the mean of data obtained from independent Michaelis– Menten curve fitting calculations after triplicate measurements with various substrate concentrations.
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Table 3. Kinetics parameters with PNPC as the substrate.[a]
sOPH wild-type S156C pOPH wild-type S172C S172A W255Y W255F W255A C257A/C267A R264A Y270F Y270A
Km [mm]
kcat [s1]
kcat/Km [s1 mm1]
Ratio [%]
2.2 0.1 5.8 0.3
390 20 53 3
175 10 9.2 0.7
100 5.3
2.15 0.07 9.6 0.5 6.1 0.2 2.34 0.09 1.9 0.1 2.6 0.1 3.2 0.2 1.36 0.05 1.94 0.08 2.4 0.1
470 20 184 8 136 6 570 30 400 20 600 30 244 8 410 10 660 30 650 30
216 9 19.3 0.4 22.5 0.3 250 10 212 5 230 10 76 3 300 8 340 10 270 10
100 8.9 10.4 113 98 106 35 139 158 126
[a] All activity measurements were carried out at 37 8C and pH 8.0 according to Winkler et al.[5b] Each value represents the mean of data obtained from independent Michaelis–Menten curve fitting calculations after triplicate measurements with various substrate concentrations.
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important for substrate affinity. Mutations to Tyr, Phe, or Ala reing of the active site by Trp255, and consequently both Km and sulted in higher Km but with varied kcat. In particular, the kcat were not changed significantly when replacing or removing W255Y mutant showed activity 60 % higher than wild-type. its side chain. Y270F showed enhanced activity for PNPC, thus The amphipathic but less bulky side chain of Tyr255 would faagain suggesting that the phenolic OH of Tyr270 is not incilitate substrate binding and product release, yet it is still efvolved in catalysis. The retained esterase activity in Y270A indifective in shielding the active site from solvent. cates that “proper” formation of the OPA binding site is not reAlthough Tyr270 formed a hydrogen bond to caprylate in quired for PNPC binding. the S172A/PNPC structure, the side-chain hydroxyl group is not directly involved in catalysis, because mutant Y270F Substrate modeling and catalytic mechanism showed 40 % higher activity. However, the significant decrease in substrate affinity of Y270A suggests that the aromatic side Finally, based on the bound ACA and caprylate structures, chain of Tyr270 participates in maintaining the OPA binding a 13-carbon model of OPA was constructed to elucidate site. The Cys257–Cys267 disulfide bond is probably important enzyme–substrate interactions (see the Supporting Informain maintaining the lid structure, because mutating either resition). It contained six carbonyl (CO) groups (numbered 4, 3, 2, due to alanine had an adverse effect in terms of both Km and 1, 1’ and 2’ in Figure 3 A). Several hydrogen bonds to the COs are likely; CO4 and CO3 stack with Trp121 as does ACA. The kcat. Trp255, Tyr270, and Cys257–Cys267 are conserved, and cleavage site is between CO1 and CO1’; both have to be carthey presumably serve the same roles in pOPH and sOPH. With mutations further from the active site, pOPH R264A showed improved Km and kcat. Arg264 (not found in sOPH) is in the variable region of the lid. Apparently its positively charged side chain is dispensable for catalysis. Likewise, by mutating other nonessential lid residues, we might be able to obtain sOPH (or pOPH) with even better Km and kcat. In contrast, the active-site mutant S172C showed less than 10 % activity (higher Km and lower kcat). Presumably the oxidized Cys172 not only lost its function as a nucleophile but also hindered substrate binding. More activity was retained in mutant S172A, in which an activated water molecule comes directly into play. Both enzymes and the mutants were then analyzed for esterase activity with PNPC as the substrate (Table 3 and Supporting Information). The higher Km values indicate lower affinity to this artificial substrate, but the overall activity was enhanced (much higher kcat values). Apparently the bulky PNP head group is not well accommodated in the active-site cleft, but the ester bond is more readily cleaved. The mutants had generally similar effects, except for mutations Figure 3. Substrate binding and catalysis. A) The modeled OPA is shown as thick sticks, and the protein is shown to Trp255 and Tyr270. Hydrolysis as thin sticks (both in green). Potential hydrogen bonds are pink dashed lines. For comparison, the S172C/ACA of the ester CO bond in PNPC (cyan) and S172A/PNPC (gray) structures are also shown. B) Proposed catalytic pathway for OPH. The backbone probably does not need shield- NH of Ser66 might also interact with CO1. 2014 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
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CHEMBIOCHEM COMMUNICATIONS bonyls in order to be cleaved, whereas the others can be alcohols. In this model, the O atom of CO1 matches a sulfonate O of Cys172. It is hydrogen bonded to Ser66 and Ser173, which presumably constitute an oxyanion hole (Figure 3 A). The C atom is 2.7 from the OG atom of Ser172, which makes a nucleophilic attack when deprotonated by His298. The proton is then transferred to the methylene group, now connected only to CO1’. Finally, His298 deprotonates a water molecule to cleave the acyl–enzyme intermediate (Figure 3 B). Similar enzyme—substrate interactions are observed in PHB depolymerase.[9] In addition to the nucleophile Ser39, the base His155, and the oxyanion hole formed by Ser40 and Cys250, the side chain of Trp307 is also hydrogen bonded to a carbonyl group of PHB (equivalent to CO3). The reaction catalyzed by OPH is typical of a/b-hydrolases, except that the cleaved bond is between two carbon atoms. Here electron delocalization (in a way like enol–ketone equilibrium, which does occur in ACA), seems to play an essential role. Upon CC bond breaking, the negative charge on CO1’ is probably stabilized by hydrogen bonds to the backbone NH of Ser66 and Val67, thus forming a second oxyanion hole. As mentioned above, this anion-binding b3-a1 loop might promote spontaneous oxidation of Cys172 to a sulfonate. Although Ser66 and Ser173 in pOPH are Thr50 and Ala157 in sOPH, the potential to form an oxyanion hole is conserved. The employment of double oxyanion holes in catalysis has been observed in thiolase, which also cleaves the CC bond between two carbonyl groups.[10] Interestingly, the catalytic Cys123 in the native thiolase crystal was also found oxidized to a sulfonate.
Conclusion OPH crystal structures have allowed us to elucidate its catalytic mechanism. They also served as a basis for rational modifications (particularly in the lid region) to endow the enzyme with desired characteristics. The three lid mutants, W255Y, R264A, and Y270F, showed 40–60 % improvement in kcat/Km. The activity enhancement can be employed to facilitate bacterial assimilation of OPA. We are currently making double and triple mutants to see if there are synergistic effects. However, OPH is responsible for the second step of PVA degradation, which starts with a dehydrogenase or oxidase. Both remain to be further characterized, but PVA oxidase might find better applications because it is probably located on the cell surface and does not need the cofactor PQQ.[3a] Solving its crystal structure would thus be highly beneficial.
Acknowledgements This work was supported by the Program for Changjiang Scholars and Innovative Research Team in University (No. IRT1135), the National High Technology Research and Development Program
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www.chembiochem.org of China (863 Program, 2012AA022202), the Priority Academic Program Development of Jiangsu Higher Education Institutions, the 111 Project (111-2-06). We thank the National Synchrotron Radiation Research Center of Taiwan for beam-time allocation and data-collection assistance. Keywords: catalytic triad · beta-diketone hydrolases · double oxyanion holes · environmental chemistry · hydrolases · microbial assimilation
[1] F. F. Nord, Naturwissenschaften 1936, 24, 763. [2] a) T. Suzuki, Y. Ichihara, M. Yamada, K. Tonomura, Agric. Biol. Chem. 1973, 37, 747 – 756; b) Y. Watanabe, M. Morita, N. Hamada, Y. Tsujisaka, Agric. Biol. Chem. 1975, 39, 2447 – 2448. [3] a) A. Yamatsu, R. Matsumi, H. Atomi, T. Imanaka, Appl. Microbiol. Biotechnol. 2006, 72, 804 – 811; b) F. Kawai, X. Hu, Appl. Microbiol. Biotechnol. 2009, 84, 227 – 237. [4] A. Marchler-Bauer, C. Zheng, F. Chitsaz, M. K. Derbyshire, L. Y. Geer, R. C. Geer, N. R. Gonzales, M. Gwadz, D. I. Hurwitz, C. J. Lanczycki, F. Lu, S. Lu, G. H. Marchler, J. S. Song, N. Thanki, R. A. Yamashita, D. Zhang, S. H. Bryant, Nucleic Acids Res. 2013, 41, D348 – D352. [5] a) Y. Yang, L. Liu, J. Li, G. Du, J. Chen, Bioprocess Biosyst. Eng. 2014, 37, 777 – 782; b) U. K. Winkler, M. Stuckmann, J. Bacteriol. 1979, 138, 663 – 670. [6] a) Y.-S. Cheng, T.-P. Ko, T.-H. Wu, Y. Ma, C.-H. Huang, H.-L. Lai, A. H.-J. Wang, J.-R. Liu, R.-T. Guo, Proteins Struct. Funct. Bioinf. 2011, 79, 1193 – 1204; b) F. Ren, T.-P. Ko, X. Feng, C.-H. Huang, H.-C. Chan, Y. Hu, K. Wang, Y. Ma, P.-H. Liang, A. H.-J. Wang, E. Oldfield, R.-T. Guo, Angew. Chem. Int. Ed. 2012, 51, 4157 – 4160; Angew. Chem. 2012, 124, 4233 – 4236. [7] Phase angles were calculated by employing OASIS: a) T. Zhang, Y. He, J. W. Wang, L. J. Wu, C. D. Zheng, Q. Hao, Y. X. Gu, H. F. Fan, OASIS, Institute of Physics, Chinese Academy of Sciences, P. R. China, 2012, http:// cryst.iphy.ac.cn; SHELXD: b) T. R. Schneider, G. M. Sheldrick, Acta Crystallogr. Sect. D Biol. Crystallogr. 2002, 58, 1772 – 1779; SOLVE/RESOLVE: c) T. C. Terwilliger, Methods Enzymol. 2003, 374, 22 – 37. Initial model building used ARP/wARP: d) A. Perrakis, R. Morris, V. S. Lamzin, Nat. Struct. Biol. 1999, 6, 458 – 463. [8] Molecular replacement calculations used Phaser: a) A. J. McCoy, R. W. Grosse-Kunstleve, P. D. Adams, M. D. Winn, L. C. Storoni, R. J. Read, J. Appl. Crystallogr. 2007, 40, 658 – 674. Manual model adjustments used Coot: b) P. Emsley, K. Cowtan, Acta Crystallogr. Sect. D Biol. Crystallogr. 2004, 60, 2126 – 2132. Computational refinement employed CNS: c) A. T. Brnger, P. D. Adams, G. M. Clore, W. L. DeLano, P. Gros, R. W. GrosseKunstleve, J. S. Jiang, J. Kuszewski, M. Nilges, N. S. Pannu, R. J. Read, L. M. Rice, T. Simonson, G. L. Warren, Acta Crystallogr. Sect. D Biol. Crystallogr. 1998, 54, 905 – 921. [9] T. Hisano, K.-i. Kasuya, Y. Tezuka, N. Ishii, T. Kobayashi, M. Shiraki, E. Oroudjev, H. Hansma, T. Iwata, Y. Doi, T. Saito, K. Miki, J. Mol. Biol. 2006, 356, 993 – 1004. [10] R. K. Harijan, T. R. Kiema, M. P. Karjalainen, N. Janardan, M. R. N. Murthy, M. S. Weiss, P. A. M. Michels, R. K. Wierenga, Biochem. J. 2013, 455, 119 – 130. [11] a) R. A. Engh, R. Huber, Acta Crystallogr. Sect. A Found. Crystallogr. 1991, 47, 392 – 400; b) V. B. Chen, W. B. Arendall III, J. J. Headd, D. A. Keedy, R. M. Immormino, G. J. Kapral, L. W. Murray, J. S. Richardson, D. C. Richardson, Acta Crystallogr. Sect. D Biol. Crystallogr. 2010, 66, 12 – 21. [12] W. Klomklang, A. Tani, K. Kimbara, R. Mamota, T. Ueda, M. Shimao, F. Kawai, Microbiology 2005, 151, 1255 – 1262. Received: April 11, 2014 Published online on && &&, 0000
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COMMUNICATIONS Y. Yang, T.-P. Ko, L. Liu, J. Li, C.-H. Huang, H.-C. Chan, F. Ren, D. Jia, A. H.-J. Wang, R.-T. Guo,* J. Chen,* G. Du* && – && Structural Insights into Enzymatic Degradation of Oxidized Polyvinyl Alcohol
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Crystal-clear hydrolysis: The crystal structures of OPH with bound inhibitors from two PVA-degrading microbes were analyzed to elucidate the hydrolytic mechanism of oxi-PVA. Several site-directed mutants were also constructed to enhance the OPH activity.
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