Biomacromolecules 2001, 2, 541-544
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Involvement of Catalytic Amino Acid Residues in Enzyme-Catalyzed Polymerization for the Synthesis of Polyesters Yoichi Suzuki,†,‡ Seiichi Taguchi,† Terumi Saito,§ Kazunobu Toshima,‡ Shuichi Matsumura,‡ and Yoshiharu Doi*,† Polymer Chemistry Laboratory, RIKEN Institute, 2-1, Hirosawa, Wako-shi, Saitama 351-0198, Japan; Faculty of Science and Technology, Keio University, 3-14-1, Hiyoshi, Kohoku-ku, Yokohama 223-8522, Japan; and Faculty of Science, Kanagawa University, 2946, Tsuchiya, Hiratsuka, Kanagawa 259-1293, Japan Received January 12, 2001; Revised Manuscript Received March 13, 2001
Recently, a variety of aliphatic polyesters have been synthesized using hydrolases such as lipases and PHB depolymerases, and the reaction mechanism for these enzyme-catalyzed polymerization has been discussed. In this paper, we have studied the involvement of the catalytic amino acid residues of the hydrolase in enzyme-catalyzed polymerization with an extracellular PHB depolymerase from Alcaligenes faecalis T1. A wild-type PHB depolymerase and three kinds of site-specific mutants (catalytic amino acids were substituted) were prepared and their polymerization activities for the ring-opening polymerization of (R)-β-butyrolactone (BL) were compared. BL was polymerized at 80 °C in bulk by the wild-type enzyme to yield polymers consisting of cyclic and linear structures in a high monomer conversion. In contrast, none of the mutant enzymes showed obvious polymerization activity. These results have clearly demonstrated that the catalytic triad is indeed responsible for the enzyme-catalyzed polymerization of BL. Introduction Polyhydroxyalkanoates (PHAs) are accumulated by a wide variety of microorganisms as intracellular carbon and energy storage compounds. Poly[(R)-3-hydroxybutyrate] [P(3HB)] and its copolyesters are the most frequently encountered PHAs, and their biodegradability and biocompatibility provide remarkable properties for next generation thermoplastic materials.1 The syntheses of these aliphatic polyesters have been extensively studied by both fermentation and chemical processes in the field of biodegradable materials science. Recently, in vitro enzymatic syntheses have become effective methods for designing and synthesizing environmentally acceptable polymeric materials. The characteristic features of the enzyme catalysis afforded novel reactions to produce polymers, which are often difficult to synthesize by conventional polymerizations. Enzymatic preparation of polyesters by condensation, transesterification, and ringopening polymerization with hydrolases such as lipases has already been demonstrated.2-7 Also, the reaction mechanism for the enzyme-catalyzed ring-opening polymerization of a lactone has been proposed.3,8 According to the proposed mechanism for the enzyme-catalyzed polymerization of lactones, the polymerization was initiated by the reaction of the enzyme with a lactone to form an acyl-enzyme complex, * To whom correspondence should be addressed. Telephone: +81-48467-9402. Fax: +81-48-462-4667. E-mail:
[email protected]. † RIKEN Institute. ‡ Keio University. § Kanagawa University.
or enzyme-activated monomer. This step involves the ringopening of the lactone, and the serine residue of the active site has been proposed to play an important role. The enzyme-activated monomer then reacts either with a nucleophile such as water, which is perhaps contained in the enzyme to accomplish the initiation, or with the hydroxy group of a growing polymer chain to continue the propagation. However, to our knowledge, there are no reports giving direct proof that the catalytic amino acid residues of the hydrolase are involved in the enzyme-catalyzed polymerization in nonaqueous media or under high reaction temperatures such as over 60 °C. Previously, we provided the first report on the polyhydroxybutyrate (PHB) depolymerasecatalyzed polymerization of β-butyrolactone into P(3HB).5c Extracellular PHB depolymerases isolated so far commonly have a catalytic triad (Ser-His-Asp) in their catalytic domains.9 In particular, Ser is the active site in a lipase box pentapeptide Gly-X-Ser-X-Gly,10a which has been found in all known serine hydrolases, such as lipase, esterase, and serine protease.11 Previously, the catalytic triad residues of PHB depolymerase from Alcaligenes faecalis T1 were identified to be 139Ser (Ser at the position of 139), 214Asp, and 273His by site-directed mutagenesis.10 To verify the involvement of the catalytic triad of PHB depolymerase in the enzyme-catalyzed polymerization of BL, three kinds of site-specific mutants (S139A, 139Ser was substituted to Ala; D214G, 214Asp to Gly; H273D, 273His to Asp) and the wild-type PHB depolymerase from A. faecalis
10.1021/bm015508o CCC: $20.00 © 2001 American Chemical Society Published on Web 04/06/2001
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Scheme 1
T1 (PhaZAfa) have been compared with respect to the ringopening polymerization of BL. Experimental Section Materials. BL having a 92% ee was kindly supplied by Takasago Koryo Kogyo Co., Ltd. (Hyogo, Japan) and used after drying with molecular sieves. The other chemicals were purchased from Kanto Chemicals (Tokyo) and Wako Chemicals (Osaka, Japan). All bacterial strains and plasmids used in this study were listed in previous reports.10 The PhaZAfas (wild-type, S139A, D214G, and H273D) were purified from recombinants of Escherichia coli JM109 according to the method described by Saito et al.12 with a slight modification [Q Sepharose HP column (Pharmacia Biotech, Uppsala, Sweden) was used instead of a (triethylaminoethyl)cellulose column]. Sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) was performed according to the procedure of Laemmli13 with a molecular weight calibration kit (Pharmacia Biotech). Then the enzyme solutions were dialyzed against 10 mM triethanolamine-HCl buffer (pH 7.0), and concentrated with a water-absorbent polymer. This procedure was repeated several times. Protein concentrations were determined by the method of Bradford14 with a protein assay kit (Bio-Rad Laboratories, Inc., Hercules, CA), and bovine serum albumin (BSA) was used as a standard. The P(3HB) hydrolysis activity of PhaZAfas was estimated based on the procedure described previously.15 The enzyme solutions were lyophilized to obtain PhaZAfas as a powder. For their circular dichroism (CD) measurement, an aliquot of each enzyme was dissolved in 10 mM phosphate buffer (pH 7.4) with a protein concentration of 0.2 mg/mL. CD spectroscopic analysis was carried out at 25 °C using a JASCO-J720WI CD spectrophotometer (JASCO Ltd., Tokyo). Polymerization. The enzyme-catalyzed ring-opening polymerization of BL was carried out as shown in Scheme 1. The general procedures were as follows. A mixture of BL (100 mg) and powdered enzyme was stirred in bulk under a nitrogen atmosphere in a capped vial placed in a thermostated oil bath. After the reaction, the reaction mixture was dissolved in chloroform (8 mL), and the insoluble enzyme was removed by filtration through a Celite pad. The chloroform was then evaporated under slightly reduced pressure to quantitatively obtain the polymer mixture. Measurements. All molecular weight data of the polymers were obtained by gel permeation chromatography (GPC) at 40 °C, using a Shimadzu 10A GPC System with a 10A refractive index detector (Shimadzu Corp., Kyoto, Japan) and GPC columns (Shodex K-802 and K-806 M, Showa Denko Co., Ltd., Tokyo). Chloroform was used as the eluent at a flow rate of 0.8 mL/min, and a sample concentration of 1.0 mg/mL was employed. The number-average molecular weight (Mn) and weight-average molecular weight (Mw) were calculated by using a Shimadzu Chromatopac C-R7A plus
Figure 1. CD spectra of the wild-type and site-specific mutants of PhaZAfa.
equipped with a GPC program. The system was calibrated with polystyrene standards with a narrow molecular weight distribution. Monomer conversion of BL to P(3HB) was determined by comparison of the 1H NMR spectral integration intensities for the δ ) 1.58 ppm peak corresponding to the methyl protons of monomeric BL with the corresponding methyl protons of the repeating unit of P(3HB) at δ ) 1.28 ppm. The polymer structure was analyzed by 1H and 13C NMR,16 and by matrix-assisted laser desorption ionization time-of-flight mass spectrometry (MALDI-TOF MS). The MALDI-TOF MS was measured with a Bruker Ploflex mass spectrometer which was equipped with a nitrogen laser. The detection was in the reflector mode. 2,5-Dihydroxybenzoic acid (DHBA) was used as the matrix, and positive ionization was used. Results and Discussion Preparation of Wild-Type and Site-Specific Mutants of PhaZAfa. The PhaZAfa recombinant proteins were purified to electrophoretic homogeneity, and the apparent molecular weights of PhaZAfas were about 47 kDa. Final yield was estimated to be about 5 mg from 1 L of culture of recombinant cells for all of the PhaZAfas. The P(3HB) hydrolysis activity of the PhaZAfa (wild-type) was estimated to be 75 U/mg of protein. On the other hand, the three mutant enzymes (S139A, D214G, and H273D) showed no P(3HB) hydrolysis activity, as reported.10 PhaZAfas were purified with same procedures from recombinant E. coli and the CD spectra showed no conformational differences between the wild-type and mutant enzymes (Figure 1). Therefore, the difference in the following polymerization results should be attributed only to the single amino acid substitutions at positions forming a catalytic triad of the enzyme. Polymerization of BL by Wild-Type PhaZAfa. It was found that BL was polymerized by wild-type PhaZAfa to yield P(3HB). Table 1 summarizes the typical ring-opening polymerization of BL with and without the wild-type PhaZAfa in bulk. The result of BL polymerization by wild-type PhaZAfa was dependent on the reaction temperature and enzyme concentration. In the absence of wild-type enzyme, no polymerization occurred at 60 °C, while at higher temperatures of 70 and 80 °C, BL was slightly oligomerized
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Catalytic Amino Acid Residues Table 1. Polymerization of (R)-β-Butyrolactone Using PHB Depolymerase from A. faecalis T1 (Wild-Type) entrya enzyme wt %b temp/°C time/h convnc/% 1 2 3 4 5 6 7 8 9 10 11 12
PhaZAfa PhaZAfa PhaZAfa PhaZAfa PhaZAfa PhaZAfa PhaZAfa PhaZAfa PhaZAfa
1 3 5 0 1 3 5 0 1 3 5 0
60 60 60 60 70 70 70 70 80 80 80 80
24 24 24 24 24 24 24 24 24 24 24 24
7 19 37 0 20 49 72 3 56 93 98 4
Mnd
Mwd
300 400 600 700 1100 1300 900 1600 2100 300 2100 3700 3200 400
1000 1900 2700 400 2800 5000 4100 500
a Entries 4, 8, and 12: blank tests. b Weight percent of enzyme to BL. Monomer conversion was determined by 1H NMR. d Molecular weights were determined by GPC system, calibrated with polystyrene standards.
c
Figure 3. 1H NMR spectrum of P(3HB) (mixture of two distinct mass series) obtained by the enzyme-catalyzed polymerization of BL using 3 wt % wild-type PhaZAfa for 24 h at 80 °C in bulk.
Figure 2. Typical MALDI-TOF mass spectrum of P(3HB) [M + Na]+ obtained by the bulk polymerization of BL using 3 wt % wild-type PhaZAfa for 14 h at 80 °C. DP: degree of polymerization. C: cyclic form, L: linear form.
thermally (Table 1, entries 8 and 12). However, both molecular weight and monomer conversion were significantly increased by the addition of wild-type enzyme, indicating that the PhaZAfa (wild-type) actually promoted the polymerization of BL. The polymer structure was analyzed by 1H and 13C NMR, and by MALDI-TOF MS. A typical MALDI-TOF mass spectroscopy of the P(3HB) produced by the ring-opening polymerization of BL using wild-type PhaZAfa is shown in Figure 2. Figure 2 also shows an expanded portion of the two distinct mass series. The product could be classified into two peak groups having mass differences of 18 m/z (Figure 2, C and L). The major series is due to the linear P(3HB) cationized with Na+ ions and terminated by carboxy and hydroxy end groups. The other might be due to two components, the cyclic form P(3HB) and crotonate terminus form P(3HB) each cationized with Na+ ions. The minor mass series increased gradually with reaction time, and they were contained approximately 20 mol % in the P(3HB) at high monomer conversion. 1H NMR spectrum of the product (mixture of two distinct mass series) supported that the minor mass series was to be the cyclic P(3HB) (Figure 3). In addition, the 1H and 13C NMR spectra of P(3HB) produced
at different reaction times had no peaks corresponding to crotonate terminus form (data not shown). These results suggested that the P(3HB) obtained from enzyme-cataylzed polymerization of BL by wild-type PhaZAfa were consisted of linear and cyclic P(3HB). It was found that the molecular weight of the resulting P(3HB) was significantly influenced by the enzyme concentration and the reaction temperature, and the polymerization with a 3 wt % concentration of wild-type enzyme at 80 °C seemed to be the best reaction conditions with respect to the molecular weight and the monomer conversion (Table 1, entry 10). Polymerization of BL by Site-Specific Mutants of PhaZAfa. Polymerizations of BL by site-specific mutants and wild-type PhaZAfa were compared under the same reaction conditions. Figure 4 shows the time-dependent changes in GPC profiles of the polymerization products of BL using 3 wt % wild-type and S139A PhaZAfa at 80 °C in bulk. It is notable that the S139A PhaZAfa showed no considerable polymerization of BL. A small and broad peak of low molecular weight showed almost no change with reaction time (Figure 4B). In contrast, the wild-type PhaZAfa showed a gradual increase in the monomer conversion with nearly unimodal peak, and it shifted toward high molecular weight with reaction time (Figure 4A). BL was also subjected to the polymerization using 5 wt % S139A PhaZAfa at 60 and 70 °C in bulk. The reactions were resulted in the same level to the blank tests at each temperature (Table 1, entries 4 and 8). That is, no oligomerization was observed by S139A PhaZAfa at 60 °C. It was directly confirmed that the catalytic center was responsible for polymerization activity. Figure 5 shows the time course of the polymerization of BL using 3 wt % of three different mutants (S139A, D214G, and H273D) together with wild-type PhaZAfa at 80 °C in bulk. There are obvious difference between the polymerization results of BL by site-specific mutants and wild-type PhaZAfa. In the case of using a wild-type enzyme, BL was gradually polymerized to produce P(3HB) with a maximum Mw greater than 5000 within a 48 h reaction time. A similar tendency
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Conclusion In conclusion, the involvement of the catalytic amino acid residues of the PHB depolymerase in the enzyme-catalyzed ring-opening polymerization of BL could be demonstrated using three kinds of PHB depolymerase site-specific mutants. Acknowledgment. This work was supported by a Grantin-Aid for Scientific Research on Priority Area, “Sustainable Biodegradable Plastics”, No 11217212 (1999) from the Ministry of Education, Science, Sports, and Culture (Japan). References and Notes
Figure 4. GPC profile changes of enzyme-catalyzed polymerization of BL using 3 wt % wild-type (A) and S139A (B) PhaZAfa at 80 °C in bulk.
Figure 5. Time course of enzyme-catalyzed polymerization of BL using 3 wt % PhaZAfa (wild-type and 3 mutants) at 80 °C in bulk: (b) wild-type; (2) S139A; (3) D214G; (]) H273D; (O) blank tests.
was observed for the monomer conversion. On the other hand, three kinds of mutants showed almost no significant polymerization of BL, at the same level to that of a blank test. These results clearly indicate that the amino acid residues forming the catalytic triad are indeed responsible for the enzyme-catalyzed ring-opening polymerization.
(1) (a) Doi, Y. Microbial Polyesters; VCH Publishers: New York, 1990. (b) Anderson, A. J.; Dawes, E. A. Microbiol. ReV. 1990, 54, 450. (c) Steinbu¨chel, A.; Valentin, H. E. FEMS Microbiol. Lett. 1995, 128, 219. (2) (a) Okumura, S.; Iwai, M.; Tominaga, Y. Agric. Biol. Chem. 1984, 48, 2805. (b) Matsumura, S.; Takahashi, J. Makromol. Chem., Rapid. Commun. 1986, 7, 369. (c) Margorin, A. L.; Crenne, J.-Y.; Klibanov, A. M. Tetrahedron Lett. 1987, 28, 1607. (d) Dordick, J. S. Trends Biotechnol. 1992, 10, 287. (e) Uyama, H.; Kobayashi, S. Chem. Lett. 1994, 1687. (f) Kobayashi, S.; Shoda, S.; Uyama, H. AdV. Polym. Sci. 1995, 121, 1. (3) (a) Uyama, H.; Takeya, K.; Kobayashi, S. Bull. Chem. Soc. Jpn. 1995, 68, 56. (b) MacDonald, R. T.; Pulapura, S. K.; Svirkin, Y. Y.; Gross, R. A.; Kaplan, D. L.; Akkara, J.; Swift, G.; Wolk, S. Macromolecules 1995, 28, 73. (4) Nobes, G. A. R.; Kazlauskas, R. J.; Marchessault, R. H. Macromolecules 1996, 29, 4829. (5) (a) Matsumura, S.; Beppu, H.; Tsukada, K.; Toshima, K. Biotechnol. Lett. 1996, 18, 1041. (b) Matsumura, S.; Suzuki, Y.; Tsukada, K.; Toshima, K.; Doi, Y.; Kasuya, K. Macromolecules 1998, 31, 6444. (c) Suzuki, Y.; Ohura, T.; Kasuya, K.; Toshima, K.; Doi, Y.; Matsumura, S. Chem. Lett. 2000, 318. (6) Kobayashi, S.; Uyama, H. Macromol. Symp. 1999, 144, 237. (7) Kumar, A.; Gross, R. A.; Jendrossek, D. J. Org. Chem. 2000, 65, 7800. (8) (a) Svirkin, Y. Y.; Xu, J.; Gross, R. A.; Kaplan, D. L.; Swift, G. Macromolecules 1996, 29, 4591. (b) Henderson, L. A.; Svirkin, Y. Y.; Gross, R. A.; Kaplan, D. L.; Swift, G. Macromolecules 1996, 29, 7759. (c) Xie, W.; Li, J.; Chen, D.; Wang, P. G. Macromolecules 1997, 30, 6997. (d) Kobayashi, S.; Uyama, H. In Enzymes in Polymer Synthesis; Gross, R. A., Kaplan, D. L., Swift, G., Eds.; ACS Symposium Series 684; American Chemical Society: Washington, DC, 1998; p 58. (9) (a) Jendrossek, D.; Backhaus, M.; Andermann, M. Can. J. Microbiol. 1995, 41, 160. (b) Jendrossek, D.; Frisse, A.; Behrends, A.; Andermann, M.; Kratzin, H.; Stanislawski, T.; Schlegel, H. G. J. Bacteriol. 1995, 177, 596. (10) (a) Shinohe, T.; Nojiri, M.; Saito, T.; Stanislawski, T.; Jendrossek, D. FEMS Microbiol. Lett. 1996, 141, 103. (b) Nojiri, M.; Saito, T. J. Bacteriol. 1997, 179, 6965. (11) (a) Schrag, J. D.; Li, Y. G.; Wu, S.; Cygler, M. Nature 1991, 351, 761. (b) Jeager, K. E.; Ransac, S.; Dijkstra, B. W.; Colson, D.; van Heuvel, M.; Misset, O. FEMS Microbiol. ReV. 1994, 15, 29. (c) Jaeger, K. E.; Steinbu¨chel, A.; Jendrossek, D. Appl. EnViron. Microbiol. 1995, 61, 3113. (12) Saito, T.; Suzuki, K.; Yamamoto, J.; Fukui, T.; Miwa, K.; Tomita, K.; Nakanishi, S.; Odani, S.; Suzuki, J.; Ishikawa, K. J. Bacteriol. 1989, 171, 184. (13) Laemmli, U. K. Nature 1970, 227, 680. (14) Bradford, M. M. Anal. Biochem. 1976, 72, 248. (15) Hiraishi, T.; Ohura, T.; Ito, S.; Kasuya, K.; Doi, Y. Biomacromolecules 2000, 1, 320. (16) 1H and 13C NMR spectra were recorded with a JEOL model ECP500 NMR spectrometer (JEOL Ltd., Tokyo) at 500 and 125 MHz, respectively. The spectral data of (R)-P(3HB) having an Mw of 5000 are shown (entry 10 in Table 1) to be representative. 1H NMR (CDCl3): δ ) 1.28 (d, 3H), 2.54 (m, 2H), 5.26 (m, 1H). 13C NMR (CDCl3): δ )19.8 (CH3), 40.8 (CH2), 67.6 (CH), 169.2 (ester CdO).
BM015508O
