Enzymatic Polymerization of Tyrosine Derivatives. Peroxidase- and

Peroxidase- and Protease-Catalyzed Synthesis of Poly(tyrosine)s with .... The polymerization results of 2a were very close to those of 2b, suggesting ...
0 downloads 0 Views 103KB Size
Download PDF
Biomacromolecules 2002, 3, 768-774

768

Enzymatic Polymerization of Tyrosine Derivatives. Peroxidaseand Protease-Catalyzed Synthesis of Poly(tyrosine)s with Different Structures Tokuma Fukuoka, Yoichi Tachibana, Hiroyuki Tonami, Hiroshi Uyama, and Shiro Kobayashi* Department of Materials Chemistry, Graduate School of Engineering, Kyoto University, Kyoto 606-8501, Japan Received February 4, 2002; Revised Manuscript Received April 2, 2002

Polymerization of tyrosine derivatives has been carried out by using two enzymes, peroxidase and protease, as catalyst to give poly(tyrosine)s with different structures. Tyrosine ester hydrochlorides were oxidatively polymerized by a peroxidase in a buffer. Using a high buffer concentration produced the polymer in good yields. The resulting polymer was soluble in N,N-dimethylformamide, dimethyl sulfoxide, and methanol but was insoluble in acetone, tetrahydrofuran, and water. The ester moiety of the polymer was subjected to the alkaline hydrolysis, yielding a water-soluble polymer having the amino acid group in the side chain. The peroxidase also catalyzed the oxidative polymerization of N-acetyltyrosine to give the polymer soluble in water. The polymerization of tyrosine ester hydrochlorides proceeded in the presence of papain catalyst to give a polymer of R-peptide structure. The polymerization in the buffer of high phosphate concentration efficiently produced the polymer. On the other hand, the polymer formation was not observed in the low buffer concentration. The molecular weight was several thousands and almost constant during the reaction. The morphology of the precipitated polymer was examined. The product of the initial reaction stage was amorphous. After 24 h, the precipitates exhibiting clear birefringence were formed. Scanning electron microscopy observation of the polymer after 72 h showed the formation of a globular crystal in a diameter larger than 50 µm, which was not found by recrystallization of poly(tyrosine). Introduction Amino acid-based polymers including polypeptides have been remarkably developed owing to their wide potential applications for biocompatible materials as well as useful chemical materials.1 Some of these polymers have unique properties and functions derived from amino acid moiety. Modification and functionalization of natural proteins have been extensively studied in the standpoint of materials development from renewable resources. A thermally polymerized product from aspartic acid has received much attention as a new useful class of biodegradable, water-soluble polymeric materials.2 The polymers consisting of the amino acid moiety in the main chain often showed good biodegradability. Recently, multiblock copolymers of GlyAlaGlyAla and poly(ethylene oxide) were prepared as a model of silkbased materials, which formed nanostructures through β-sheet self-assembly.3 Polymerization of vinyl monomers possessing the amino acid group has been also extensively investigated.4 Enzymatic syntheses of polyphenols have received much attention as an alternative to preparation of conventional phenolic resins (novolak and resol resins), owing to no use of toxic formaldehyde, mild reaction conditions (in neutral solvents at room temperature), and facile procedures.5,6 Peroxidase catalyzed an oxidative polymerization of various phenol derivatives mainly with substituents at meta- and parapositions. The polymerization was often carried out in an

aqueous organic solvent. During the polymerization, the powdery polymer was precipitated. Most of the resulting polyphenols had structures of a mixture of phenylene and oxyphenylene units, which are formed by C-C and C-O couplings of phenols. Recent investigations revealed that the coupling selectivity (regioselectivity) could be controlled by changing the solvent composition, yielding a DMF-soluble polyphenol from unsubstituted phenol.7 The control of the molecular weight was achieved by using a small amount of 2,4-dimethylphenol as comonomer. Many studies concerning syntheses of biodegradable polymers by fermentation and chemical processes have been made in the viewpoint of biodegradable materials.8 Recently, another approach to synthesis of biodegradable polyesters and polycarbonates has been developed by lipase-catalyzed polymerizations.5,9 Lipase is an enzyme which catalyzes the hydrolysis of fatty acid esters normally in an aqueous environment in living systems. However, lipases are sometimes stable in organic solvents and can be used as catalyst for esterifications and transesterifications.10 By utilization of such catalytic specificities of lipase, functional aliphatic polyesters have been synthesized by various polymerization modes.5 Proteases are also known to catalyze reverse reactions, forming peptide bonds under selected conditions.11 Poly(amino acid)s were synthesized by protease-catalyzed polymerization of amino acid esters.5g In the case of diethyl L-glutamate hydrochloride, the polymerization regioselec-

10.1021/bm020016c CCC: $22.00 © 2002 American Chemical Society Published on Web 05/01/2002

Biomacromolecules, Vol. 3, No. 4, 2002 769

Enzymatic Polymerization of Tyrosine Derivatives Scheme 1

Table 1. Peroxidase-Catalyzed Polymerization of Tyrosine Ester Hydrochloridesa polymer

tively proceeded by papain catalyst to give the polymer consisting of R-peptide unit exclusively.12 In this study, we have investigated enzymatic polymerization of tyrosine derivatives by two different types of enzymes: peroxidase and papain. The former catalyzed the oxidative polymerization of tyrosine derivatives, in which a phenol moiety was subjected to an oxidative coupling. The polymerization of tyrosine esters, followed by alkaline hydrolysis of the ester group produced poly(tyrosine) having an amino acid moiety in the side chain. The latter catalyzed polycondensation of tyrosine ester hydrochloride to produce poly(tyrosine) with R-peptide structure. Results and Discussion Peroxidase-Catalyzed Polymerization of Tyrosine Ester Hydrochlorides. Phenol moiety of tyrosine in proteins is known to be subjected to enzymatic oxidative coupling (cross-linking), in which dityrosine linked with an aromatic carbon-carbon bond at the ortho position of phenol or isotyrosine with a bond between an aromatic carbon at the ortho position and phenoxy oxygen is formed.13 Biologically, the formation of dityrosine or isotyrosine is involved in an oxidative stress.14 For applications of food industry, a peroxidase-catalyzed cross-linking of the tyrosine residue in edible soy proteins and gliadins from wheat gluten was examined.15 The mechanical properties dramatically changed with the enzymatic cross-linking. In this study, horseradish peroxidase (HRP) was used as catalyst, which showed high catalytic activity for oxidative polymerization of phenol derivatives.5,6 At first, the HRPcatalyzed polymerization of L-tyrosine ethyl ester hydrochloride (1) was carried out using hydrogen peroxide as oxidizing agent in a Tris buffer (pH 7.6) at room temperature (Scheme 1). In the polymerization in 0.10 M Tris buffer, no polymeric precipitates were formed. The reaction mixture was poured into a large amount of acetone to give the powdery polymer in 19% yield. The molecular weight of the polymer, estimated by size exclusion chromatography (SEC) with DMF containing 0.10 M LiCl as eluent, was 2400. In distilled water, the formation of polymeric precipitates was not also observed during the polymerization. On the other hand, the powdery polymer was precipitated in high buffer concentration (1.0 M) during the reaction, which was collected by centrifugation after the reaction. The resulting polymer was soluble in N,N-dimethylformamide (DMF), dimethyl sulfoxide (DMSO), methanol, and acid and

entry

monomerb

yield (%)

Mnc

Mw/Mnc

1 2 3 4 5 6 7 8 9 10

1 (40) 1 (80) 1 (160) 1 (320) 2a (40) 2a (80) 2b (40) 2b (80) 2c (40) 2c (80)

35 74 82 24 17 64 19 64 62 63

1500 1700 1900 2200 3500 4000 2600 4000 4000 2500

1.8 2.5 3.6 1.7 2.9 3.4 2.5 3.1 3.3 3.4

a Polymerization of tyrosine ester hydrochloride using HRP catalyst (10 mg) in 1.0 M Tris buffer of pH 7.6 (25 mL) at room temperature for 3 h under air. b In parentheses, the monomer concentration (mM). c Determined by SEC using DMF containing 0.10 M LiCl as eluent.

basic aqueous solutions but insoluble in distilled water, acetone, acetonitrile, and tetrahydrofuran. The polymer structure was estimated to be of a mixture of phenylene and oxyphenylene units by NMR and IR analysis.5,6 Polymerization results are summarized in Table 1. The polymer yield strongly depended on the monomer concentration; there was a maximum point at the concentration of 160 mM (entry 3 in Table 1). The molecular weight was in the range of several thousands and increased as a function of the monomer concentration. To examine the effect of the stereoisomer on the present polymerization, L-, D-, and D,L-tyrosine methyl ester hydrochlorides (2a, 2b, and 2c, respectively) were polymerized by HRP in the Tris buffer. The solubility of 2a and 2b toward the buffer was not high; thus, the low concentration of the monomer was used. When the monomer concentration was 40 mM, the yield of the polymer from 2a or 2b was low (entries 5 and 7), whereas the moderate yield was achieved in using 2c (entry 9). The larger concentration of the monomer (80 mM) produced the polymer in higher yields in the case of 2a and 2b (entries 6 and 8), although the monomer was not perfectly soluble in the buffer before the reaction. The polymerization results of 2a were very close to those of 2b, suggesting that the stereoconfiguration of the tyrosine derivatives scarcely affected the present polymerization. The solubility of the polymer from 2 decreased, as compared with that from 1; the polymer was only soluble in DMF, DMSO, and acid and basic aqueous solutions. The HRP-catalyzed polymerization of 1 also took place in phosphate buffers. In the presence of the 0.10 M phosphate buffer of pH 7.0, the polymer precipitates were not formed. The reprecipitation using acetone as nonsolvent afforded the polymer with molecular weight of 2900 in 70% yield based on the starting monomer amount. In the polymerization in the 1.0 M phosphate buffer, the powdery polymer was precipitated during the polymerization (yield, 80%; molecular weight, 1600). Alkaline Hydrolysis of Poly(tyrosine ester) to a New Class of Poly(tyrosine). High molecular weight poly(tyrosine) with structure of R-peptide is commercially available. Here, poly(1) (entry 3) was converted to a new class of poly(tyrosine) (3) by alkaline hydrolysis. The

770

Biomacromolecules, Vol. 3, No. 4, 2002

Fukuoka et al. Scheme 2

Table 3. Peroxidase-Catalyzed Copolymerization of N-Acetyltyrosine and Arbutina copolymer Figure 1. FT-IR spectra of poly(1) (entry 3 in Table 1) and 3. Table 2. Peroxidase-Catalyzed Polymerization of N-Acetyltyrosinea polymer entry

solventb

yield (%)

Mnc

Mw/Mnc

1 2 3

distilled water 0.10 M phosphate buffer (7.0) 0.10 M Tris buffer (7.6)

20 29 32

850 1100 5700

1.6 1.4 1.1

a

Polymerization of N-acetyltyrosine (4) (80 mM) using HRP catalyst (10 mg) in aqueous solvent (25 mL) at room temperature for 3 h under b air. In parentheses, the buffer pH. c Determined by SEC using water containing 0.10 M NaCl as eluent.

resulting product was only soluble in water. The molecular weight of 3 was 2300, estimated by SEC with water containing 0.10 M NaCl as eluent, whose value was relatively close to that of poly(1). Figure 1 shows FT-IR spectra of poly(1) and 3. In the spectrum of poly(1), there was a characteristic peak at 1730 cm-1 due to the CdO vibration of the ester moiety, which completely disappeared after the reaction. A new strong peak at 1635 cm-1 ascribed to the carboxylate salt was seen in the FT-IR spectrum of 3. The ethyl ester group was not observed in the 1H NMR measurement of 3 in D2O. These data indicate that the ester moiety was hydrolyzed to form poly(tyrosine) having no peptide bond, whose structure was different from that of the commercially available poly(tyrosine). Thus, the present polymer is regarded as a new class of poly(amino acid). Relevant to this point, peroxidasecatalyzed polymerization tyrosine decyl esters in a micellar solution was reported,16 in which the view of production of new amino acid polymers was not pointed out. Enzymatic Polymerization and Copolymerization of N-Acetyltyrosine. HRP catalysis induced the oxidative polymerization of N-acetyltyrosine (4). The polymerization proceeded in the Tris and phosphate buffers as well as distilled water (Table 2). In all cases, the polymer precipitates were not formed during the polymerization. The polymer was isolated by dialysis of the reaction mixture (cutoff molecular weight, 1000) in water. The polymer yield was

entry

solventb

yield (%)

Mnc

Mw/Mnc

1 2 3

distilled water 1.0 M phosphate buffer (7.0) 1.0 M Tris buffer (7.6)

100 100 56

1900 15000 12000

1.1 1.2 1.2

a Copolymerization of N-acetyltyrosine (4) and arbutin (5) (each 100 mM) using HRP catalyst (2.5 mg) in aqueous solvent (12.5 mL) at room temperature for 3 h under air. b In parentheses, the buffer pH. c Determined by SEC using water containing 0.10 M NaCl as eluent.

not high, probably owing to the loss of the low molecular weight polymers during the dialysis. The molecular weight greatly depended on the solvent. The Tris buffer produced the polymer with the highest molecular weight. In other solvents, the molecular weight was around 1000. The HRP-catalyzed copolymerization of 4 with 4-hydroxyphenyl β-D-glucopyranoside (arbutin, 5) was carried out (Scheme 2). Copolymerization results are summarized in Table 3. Arbutin was reported to be polymerized by soybean peroxidase in a buffer to give a water-soluble polymer with molecular weight of several thousands.17 During the copolymerization, the polymeric precipitates were not formed. After the reaction, the copolymer was isolated by the reprecipitation using acetone as nonsolvent. The copolymer was soluble in water and insoluble in common organic solvents. In the phosphate buffer or distilled water, the copolymer was obtained quantitatively, whereas the polymer yield decreased in the Tris buffer, the behaviors of which were different from those of the enzymatic homopolymerization of 4. The molecular weight was estimated by SEC with 0.10 M NaCl solution as eluent. The molecular weight of the copolymer obtained in the Tris and phosphate buffers was higher than 1 × 104 and its index was small. The large difference of the yield and molecular weight between poly(4) and the product from a mixture of 4 and 5 suggest the formation of the copolymer of 4 and 5. The unit ratio of the copolymer could not be determined, owing to the overlapping of broad peaks of both units (see the Experimental Section). The copolymerization of 1 with 5 also proceeded in the presence of HRP catalyst; however, the resulting product was not perfectly soluble in water and organic solvents.

Biomacromolecules, Vol. 3, No. 4, 2002 771

Enzymatic Polymerization of Tyrosine Derivatives Scheme 3

Table 4. Protease-Catalyzed Polymerization of Tyrosine Ethyl Ester Hydrochloridea entry

buffer concn (M)

monomer (mM)

polymer yieldb (%)

1 2 3 4 5 6 7 8 9 10 11

0.50 0.50 0.50 1.0 1.0 1.0 2.0 2.0 2.0 2.0 2.0

80 100 200 80 100 200 40 60 80 100 200

0 10 20 0 30 37 0 25 40 59 64

Figure 2. FAB mass spectrum of the polymer obtained by the polymerization of 1 (200 mM) using papain catalyst (20 mg/mL) in phosphate buffer (1.0 M, pH 7.0) at 40 °C for 3 h.

a Polymerization of L-tyrosine ethyl ester hydrochloride (1) using papain catalyst (0.50 g) in phosphate buffer (pH 7.0, 25 mL) at 40 °C for 3 h under air. b Isolated yield of precipitated polymer based on the weight of the monomer.

Protease-Catalyzed Polymerization of L-Tyrosine Ethyl Ester Hydrochloride. Ester hydrochlorides of some L-amino acids were polymerized in a buffer by protease catalyst to give oligo(R-amino acid)s.5g,18 As for monomers of amino acids having an carboxylic acid group in the side chain, diethyl L-aspartate was polymerized by alkanophilic protease from Streptomyces sp. in bulk to give the polymer in a nonregioselective manner, having a mixed structure of R- and β-peptide linkages.19 Very recently, we achieved regioselective polymerization of diethyl L-glutamate hydrochloride by papain catalyst, leading to the exclusive formation of poly(R-peptide).12 In this study, we have examined the papain-catalyzed polymerization of 1 in an aqueous solution at 40 °C to give poly(tyrosine) (6) (Scheme 3). The polymerization proceeded in pH 7.0 buffers of relatively high phosphate concentration; during the polymerization, powdery precipitates were formed, which were collected by centrifugation. Results of the polymerization for 3 h are summarized in Table 4. In the case of the buffer concentration of 0.50 or 1.0 M, the polymer formation was observed at the monomer concentration larger than 100 mM; in the lower monomer concentration, the polymer precipitation was not seen during the reaction (entries 1 and 4). Using the higher buffer concentration (2.0 M) produced the polymer in higher yields than that of 0.50 or 1.0 M buffer. The resulting polymer was soluble in DMF, DMSO, and alkaline solution but insoluble in acetone, chloroform, methanol, THF, toluene, and water. The polymer structure was confirmed by 1H NMR. In all samples, molecular weights and their indexs, estimated by SEC using DMF containing 0.10 M LiCl as eluent, were about 2000 and 1.2, respectively. Fast atom bombardment (FAB) mass spectrum

Figure 3. Time-conversion curve in the polymerization of 1 (100 mM) using papain catalyst (20 mg/mL) in phosphate buffer (1.0 M, pH 7.0) at 40 °C.

of the polymer is shown in Figure 2. There were sets of two peaks with regular peak-to-peak distance (162 or 163) in the range of the degree of polymerization (DP) value from 4 to 9, whose value was slightly smaller than that determined by SEC (DP ca. 12). This may be due to the overestimate of the molecular weight by using polystyrene as standard in the SEC measurement. Figure 3 shows a time-conversion curve in the proteasecatalyzed polymerization of 1 in the 1.0 M phosphate buffer. The monomer was consumed relatively fast, and after 2 h, all the monomer disappeared. The yield of the precipitated polymer was not high in the same conditions (entry 5), suggesting that the soluble oligomers and/or the hydrolyzed monomer (tyrosine) remained in the medium. Figure 4 shows reaction time versus yield and molecular weight of the precipitated polymer. The polymer yield increased as a function of the time and reached 63% after 3 days. The molecular weights and their indexes (data not shown) were almost constant during the polymerization. These data suggest that the coupling of the soluble oligomers occurred after the complete consumption of the monomer. The constant molecular weight and its index may be due to the dependence of the solubility of 6 with different DP toward the buffer. Morphology Observation of Precipitated Poly(tyrosine). Poly(R-amino acid)s with high molecular weight are syn-

772

Biomacromolecules, Vol. 3, No. 4, 2002

Fukuoka et al.

Figure 4. Time versus yield and molecular weight of the precipitated polymer in the polymerization of 1 (100 mM) using papain catalyst (20 mg/mL) in phosphate buffer (1.0 M, pH 7.0) at 40 °C.

Figure 6. SEM images of (A) the polymer obtained by the polymerization of 1 (100 mM) using papain catalyst (20 mg/mL) in phosphate buffer (1.0 M, pH 7.0) at 40 °C for 72 h and (B) cross section of the fragment.

Figure 5. WAXD of (A) the polymer obtained by the polymerization of 1 (100 mM) using papain catalyst (20 mg/mL) in phosphate buffer (1.0 M, pH 7.0) at 40 °C for 72 h and (B) commercially available poly(tyrosine) with molecular weight of 1 × 104 to 4 × 104.

thesized by ring-opening polymerization of R-amino acid N-carboxylic anhydrides (NCAs). In some cases of the precipitation polymerization of NCAs, growth of the polypeptide crystals was observed during the reaction.20 For example, a ribbonlike fibril crystal with an antiparallel β-structure precipitated in the polymerization of D,L-Ala NCA in acetonitrile. From L-Pro NCA, a hexagonal lamellar crystal was formed. Recrystallization of synthetic polypeptides was also reported. Single crystals of polypeptides of glycine, L-alanine, and L-tyrosine were obtained from their dilute solution. From high molecular weight poly(L-tyrosine), small hexagonal platelets in the size of about 2 µm were precipitated from a dilute mixed solvent of DMF and 1-heptanol.21 Single crystals of poly(γ-methyl D-glutamate) with hexagonal platelets were also obtained by an isothermal crystallization from its dilute solution.22 Optical microscopic observation showed that the precipitates formed at initial stage (after 6 h) was of irregular shape in the papain-catalyzed polymerization of 1 (see, Supporting Information). Birefringence was not observed in the product. After 24 h, globular-shaped precipitates displaying strong birefringence in the diameter of ca. 60 µm were observed. Wide-angle X-ray diffraction (WAXD) of 6 (the product after 72 h) showed a relatively sharp spectral pattern (Figure 5A), which was somewhat different from that of the commercially

available poly(tyrosine) with molecular weight of 1 × 104 to 4 × 104 (Figure 5B). These data suggest the formation of higher crystalline poly(tyrosine) by the enzymatic polymerization of 1. Figure 6 shows scanning electron microscopy (SEM) images of the product after 72 h. The formation of the globular particle in the diameter larger than 50 µm was seen (Figure 6A). The cross section of the fragment showed that rodlike crystals radially originated from the center. This interesting morphology of poly(tyrosine) (6) is characteristic of the enzymatic polymerization of 1, since such a globular crystal was not observed in the recrystallization of poly(Ramino acid)s including poly(tyrosine).21 Conclusion Two different enzymes, peroxidase and protease, catalyzed the polymerization of tyrosine derivatives under mild reaction conditions. Peroxidase-catalyzed oxidative polymerization of tyrosine ester hydrochlorides produced a new class of poly(amino acid)s. The stereoconfiguration of the monomer scarcely affected the polymerization behaviors. The alkaline hydrolysis of the resulting polymer afforded water-soluble poly(tyrosine), whose structure was different from that of polypeptides. N-Acetyltyrosine was also polymerized by HRP catalysis, yielding the water-soluble poly(tyrosine) derivative. Papain catalyzed the polycondensation of L-tyrosine ethyl ester hydrochloride in the high buffer concentration to give poly(tyrosine) with R-peptide structure. The morphology observation of the precipitated product showed the formation of the globular crystal in the diameter larger than 50 µm, which may be useful as functional particle support for separation and immobilization. Further investigations on the enzymatic synthesis of new poly(amino acid)s are under way in our laboratory.

Enzymatic Polymerization of Tyrosine Derivatives

Experimental Section Materials. D-Tyrosine methyl ester hydrochloride was synthesized according to the modification of the literature.23 HRP (100 units/mg) and papain (origin: Carica papaya, 0.5 unit/g) were purchased from Wako Pure Chemical Industries, Inc. Enzymes and other reagents were commercially available and used as received. HRP-Catalyzed Polymerization of Tyrosine Ester Hydrochlorides. A typical run was as follows (entry 3 in Table 1). L-Tyrosine ethyl ester hydrochloride (0.98 g, 4.0 mmol) and HRP (10 mg) in 1.0 M Tris buffer of pH 7.6 (25 mL) were placed in a 50 mL flask. Hydrogen peroxide (5.0% aqueous solution, 2.7 mL, 4.0 mmol) was added dropwise to the mixture for 2 h at room temperature under air. After 3 h, polymer precipitates were collected by centrifugation. The polymer was washed with water, followed by drying in vacuo to give 0.80 g of the polymer (yield 82%).1H NMR (DMSO-d6): δ 1.0-1.5 (br, CH3), 2.7-3.2 (br, ArCH2), 3.7-4.0 (br, CH), 4.0-4.2 (br, OCH2), 6.5-7.5 (br, Ar); IR (KBr) 3200-3400 (ν(O-H)), 1731 (ν(CdO)), 1608, 1506 (ν(CdC) of Ar), 1214 (ν(C(Ar)-O-C(Ar) and C(Ar)OH)), 1028 cm-1 (ν(C(Ar)-O-C(Ar))). Alkaline Hydrolysis of Poly(tyrosine ethyl ester). Poly(L-tyrosine ethyl ester) (1.0 g) was kept in 1.0 M NaOH solution (25 mL) at 60 °C. After 24 h, the mixture was neutralized with 6.0 M hydrochloride solution, and subsequently, the polymer was purified by dialysis (cutoff molecular weight 100). The remaining solution was lyophilized to give 0.28 g of the polymer (yield 40%). 1H NMR (D2O): δ 2.7-3.2 (br, ArCH2), 3.6-4.0 (br, CH), 6.5-7.5 (br, Ar). HRP-Catalyzed Polymerization of N-Acetyltyrosine. A typical run was as follows (entry 3 in Table 2). NAcetyltyrosine (0.45 g, 2.0 mmol) and HRP (10 mg) in 1.0 M Tris buffer of pH 7.6 (25 mL) were placed in a 50 mL flask. Hydrogen peroxide (5.0% aqueous solution, 1.4 mL, 2.0 mmol) was added dropwise to the mixture for 2 h at room temperature under air. After 3 h, the mixture was subjected to dialysis (cutoff molecular weight 1000) in water. The remaining solution was lyophilized to give 0.14 g of the polymer (yield 32%).1H NMR (DMSO-d6): δ 1.5-2.2 (br, CH3), 2.5-3.2 (br, ArCH2), 4.0-4.5 (br, CH), 6.5-7.2 (br, Ar). HRP-Catalyzed Copolymerization of N-Acetyltyrosine and Arbutin. A typical run was as follows (entry 3 in Table 3). N-Acetyltyrosine (0.28 g, 1.25 mmol), arbutin (0.34 g, 1.25 mmol), and HRP (2.5 mg) in 1.0 M Tris buffer of pH 7.6 (12.5 mL) were placed in a 50 mL flask. Hydrogen peroxide (5.0% aqueous solution, 1.7 mL, 2.5 mmol) was added dropwise to the mixture for 2 h at room temperature under air. After 3 h, the reaction mixture was poured into a large amount of acetone. The precipitates were collected and dried in vacuo to give 0.35 g of the polymer (yield 56%).1H NMR (DMSO-d6): δ 1.5-2.2 (br, CH3), 2.5-3.2 (br, ArCH2), 4.0-5.5 (br, CH of tyrosine, H and OH of glucose moiety), 6.5-7.2 (br, Ar). Papain-Catalyzed Polymerization of L-Tyrosine Ethyl Ester Hydrochloride. A typical run was as follows (entry 5 in Table 4). In a 50 mL flask, a mixture of 1 (0.61 g, 2.5

Biomacromolecules, Vol. 3, No. 4, 2002 773

mmol), papain (500 mg), and 25 mL of 1.0 M phosphate buffer (pH 7) was placed. The mixture was kept at 40 °C under gentle shaking for 3 h at ambient pressure. The enzyme was deactivated by heating at 90 °C for 5 min. After the reaction mixture cooled to room temperature, the precipitates were collected by centrifugation and washed with water twice. The resulting white powder was dried in vacuo to give 0.18 g of the polymer (30% yield). 1H NMR (DMSO-d6): δ 1.1 (t, OCH2CH3), 2.8 (br, ArCH2), 4.0 (q, OCH2), 4.4 (br, NHCHCH2), 6.8 (m, Ar), 8.0 (br, NH), 9.2 (br, ArOH). Measurements. SEC analysis was carried out by using a Tosoh SC8010 apparatus equipped with refractive index (RI) detector under the following conditions: TSKgel R3000 column and DMF containing 0.10 M LiCl eluent at a flow rate of 0.50 mL/min at 60 °C or TSKgel R3000 column and distilled water containing 0.10 M NaCl eluent at a flow rate of 0.50 mL/min at 40 °C. The calibration curves were obtained using polystyrene (DMF eluent) or poly(ethylene oxide) (water eluent) as standard. NMR spectra were recorded on a Bruker DPX400 spectrometer. IR measurement was carried out with a Shimadzu IR-460 spectrometer. FAB mass measurement was carried out using a JEOL highperformance JMS-HX110 mass spectrometer. WAXD patterns were recorded on a Rigaku RINT-1400 (40 kV/200 mA) system with the Cu KR X-ray beams. Polarization microscope measurement was carried out using an Olympus IX70 optical microscope equipped with crossed polarizer. SEM measurement was performed on a Hitachi S-3000N scanning electron microscope. Acknowledgment. This work was supported by Program for Promotion of Basic Research Activities for Innovative Bioscience. Supporting Information Available. Optical electron microscopy image of the product after 6 h (A) and polarization microscopy image of the product after 24 h (B). This material is available free of charge via the Internet at http:// pubs.acs.org. References and Notes (1) (a) Erhan S. In Desk Reference of Functional Polymers, Syntheses and Applications; Arshady, R., Ed.; American Chemical Society: Washington, DC, 1997; pp 261-270. (b) Kaplan, D. L., Ed. Biopolymers from Renewable Resources; Springer: Berlin, 1998. (2) Gross, R. A.; Scholz, C., Eds. ACS Symp. Ser. 2001, No. 786. (3) (a) Rathore, O.; Winningham, M. J.; Sogah, D. Y. J. Polym. Sci., Polym. Chem. Ed. 2000, 38, 352. (b) Rathore, O.; Sogah, D. Y. Macromolecules 2001, 34, 1477. (4) Sanda, F.; Endo, T. Macromol. Chem. Phys. 1999, 200, 2651. (5) For recent reviews on enzymatic polymerizations, see: (a) Kobayashi, S.; Shoda, S.; Uyama, H. AdV. Polym. Sci. 1995, 121, 1. (b) Kobayashi, S.; Shoda, S.; Uyama, H. In Catalysis in Precision Polymerization; Kobayashi, S., Ed.; John Wiley & Sons: Chichester, 1997; Chapter 8. (c) Kobayashi S.; Uyama H. In Materials Science and Technology-Synthesis of Polymers; Schlu¨ter, A.-D., Ed.; WileyVCH: Weinheim, 1998; Chapter 16. (d) Gross, R. A., Kaplan, D. L., Swift, G., Eds. ACS Symp. Ser. 1998, No. 684. (e) Kobayashi, S. J. Polym. Sci., Polym. Chem. Ed. 1999, 37, 3041. (f) Gross, R. A.; Kumar, A.; Kalra, B. Chem. ReV. 2001, 101, 2097. (g) Kobayashi, S.; Uyama, H.; Kimura, S. Chem. ReV. 2001, 101, 3793. (h) Kobayashi, S.; Uyama, H.; Ohmae, M. Bull. Chem. Soc. Jpn. 2001, 74, 613. (i) Kobayashi, S.; Sakamoto, J.; Kimura, S. Prog. Polym. Sci. 2001, 26, 1525. (j) Kobayashi, S.; Uyama, H. In Encyclopedia of Polymer Science and Technology, 3rd ed.; Kroschwitz, J. I., Ed.; John Wiley & Sons: New York, in press.

774

Biomacromolecules, Vol. 3, No. 4, 2002

(6) For recent reviews and papers on enzymatic oxidative polymerizations, see: (a) Uyama, H.; Kobayashi, S. CHEMTECH 1999, 29 (10), 22. (b) Akkara, J. A.; Ayyagari, M. S. R.; Bruno, F. F. Trends Biotechnol. 1999, 17, 67. (c) Tonami, H.; Uyama, H.; Kobayashi, S.; Rettig, K.; Ritter, H. Macromol. Chem. Phys. 1999, 200, 1998. (d) Tonami, H.; Uyama, H.; Kobayashi, S.; Kubota, M. Macromol. Chem. Phys. 1999, 200, 2365. (e) Fukuoka, T.; Tonami, H.; Maruichi, N.; Uyama, H.; Kobayashi, S.; Higashimura, H. Macromolecules 2000, 33, 9152. (f) Reihmann, M. H.; Ritter, H. Macromol. Chem. Phys. 2000, 201, 798. (g) Tonami, H.; Uyama, H.; Kobayashi, S.; Fujita, T.; Taguchi, Y.; Osada, K. Biomacromolecules 2000, 1, 149. (h) Reihmann, M. H.; Ritter, H. Macromol. Chem. Phys. 2000, 201, 1593. (i) Liu, W.; Bian, S.; Li, L.; Samuelson, L.; Kumar, J.; Tripathy, S. Chem. Mater. 2000, 12, 1577. (j) Mandal, B. K.; Walsh, C. J.; Sooksimuang, T.; Behroozi, S. J. Chem. Mater. 2000, 12, 6. (k) Wu, X.; Kim, J.; Dordick, J. S. Biotechnol. Prog. 2000, 16, 513. (l) Reihmann, M. H.; Ritter, H. Macromol. Biosci. 2001, 1, 85. (m) Liu, W.; Cholli, A. L.; Kumar, J.; Tripathy, S.; Samuelson, L. Macromolecules 2001, 34, 3522. (n) Uyama, H.; Maruichi, N.; Tonami, H.; Kobayashi, S. Biomacromolecules 2002, 3, 187. (7) (a) Oguchi, T.; Tawaki, S.; Uyama, H.; Kobayashi, S. Macromol. Rapid Commun. 1999, 20, 401. (b) Oguchi, T.; Tawaki, S.; Uyama, H.; Kobayashi, S. Bull. Chem. Soc. Jpn. 2000, 73, 1389. (c) Mita, N.; Tawaki, S.; Uyama, H.; Kobayashi, S. Polym. J. 2001, 33, 374. (d) Kobayashi, S.; Uyama, H.; Tonami, H.; Oguchi, T.; Higashimura, H.; Ikeda, R.; Kubota, M. Macromol. Symp. 2001, 175, 1. (8) Mohanty, A. K.; Misra, M.; Hinrichsen, G. Macromol. Mater. Eng. 2000, 276/277, 1. (9) For recent reviews and papers on lipase-catalyzed polymerizations, see: (a) Kumar, A.; Gross, R. A. Biomacromolecules 2000, 1, 133. (b) Osanai, Y.; Toshima, K.; Matsumura, S. Chem. Lett. 2000, 576. (c) Nishida, H.; Yamashita, M.; Nagashima, M.; Endo, T.; Tokiwa, Y. J. Polym. Sci., Polym. Chem. Ed. 2000, 38, 1560. (d) Runge, M.; O’Hagan, D.; Haufe, G. J. Polym. Sci., Polym. Chem. Ed. 2000, 38, 2004. (e) Kumar, A.; Kalra, B.; Dekhterman, A.; Gross, R. A. Macromolecules 2000, 33, 6303. (f) Kikuchi, H.; Uyama, H.; Kobayashi, S. Macromolecules 2000, 33, 8971. (g) Namekawa, S.; Uyama, H.; Kobayashi, S. Macromol. Chem. Phys. 2001, 202, 801. (h) Uyama, H.; Inada, K.; Kobayashi, S. Macromol. Biosci. 2001, 1, 40. (i) Tsujimoto, T.; Uyama, H.; Kobayashi, S. Biomacromolecules 2001, 2, 29. (j) Kumar, A.; Garg, K.; Gross, R. A. Macromolecules 2001, 34, 3527. (k) Co´rdova, A. Biomacromolecules 2001, 2, 1347. (l) Kim, D.-Y.; Dordick, J. S. Biotechnol. Bioeng. 2001, 76, 200.

Fukuoka et al.

(10)

(11) (12) (13) (14) (15) (16) (17) (18)

(19) (20)

(21) (22) (23)

(m) Ebata, H.; Toshima, K.; Matsumura, S. Chem. Lett. 2001, 798. (n) Kobayashi, S.; Uyama, H. In AdVances in Biopolyesters, Babel, W., Steinbu¨chel, A., Eds.; Springer-Verlag: Heidelberg, 2001; pp 241-262. (o) Gross, R. A.; Karla, B.; Kumar, A. Appl. Microbiol. Biotechnol. 2001, 55, 655. (p) Uyama, H.; Kobayashi, S.; Morita, M.; Habaue, S.; Okamoto, Y. Macromolecules 2001, 34, 6554. (a) Klibanov, A. M. Acc. Chem. Res. 1990, 23, 114. (b) Santaniello, E.; Ferraboschi, P.; Grisenti, P.; Manzocchi, A. Chem. ReV. 1992, 92, 1071. (c) Wong, C.-H.; Whitesides, G. M. Enzymes in Synthetic Organic Chemistry; Pergamon: Oxford, 1994. Rozzell, J. D., Wagner, F., Eds. Biocatalytic Production of Amino Acids and DeriVatiVes; Hanser Publishers: Munich, 1992. Uyama, H.; Fukuoka, T.; Komatsu, I.; Watanabe, T.; Kobayashi, S. Biomacromolecules 2002, 3, 318. Michon, T.; Chenu, M.; Kellershon, N.; Desmadril, M.; Gue´guen, J. Biochemistry 1997, 36, 8504. Souza, J. M.; Giasson, B. I.; Chen, Q.; Lee, V. M.-Y.; Ichiropoulos, H. J. Biol. Chem. 2000, 275, 18344. (a) Stuchell, Y.; Krochta, J. M. J. Food Sci. 1994, 59, 1332. (b) Michon, T.; Wang, W.; Ferrasson, E.; Gue´guen, J. Biotechnol. Bioeng. 1999, 63, 450. (a) Sarma, R.; Alva, K. S.; Marx, K. A.; Tripathy, S. K.; Akkara, J. A.; Kaplan, D. L. Mater. Sci. Eng. 1996, C4, 189. (b) Marx, K. A.; Zhou, T.; Sarma, R. Biotechnol. Prog. 1999, 15, 522. Wang, P.; Martin, D.; Parida, S.; Rethwisch, D. G.; Dordick, J. S. J. Am. Chem. Soc. 1995, 117, 12885. (a) Sluyterman, L. A. E.; Wijdenes, J. Biochim. Biophys. Acta 1972, 289, 194. (b) Anderson, G.; Luisi, P. L. HelV. Chim. Acta 1979, 62, 488. (c) Jost, R.; Brambilla, E.; Monti, J. C.; Luisi, P. L. HelV. Chim. Acta 1980, 63, 375. (d) Aso, K.; Uemura, T.; Shiokawa, Y. Agric. Biol. Chem. 1988, 52, 2443. Matsumura, S.; Tsushima, Y.; Otozawa, N.; Murakami, S.; Toshima, K.; Swift, G. Macromol. Rapid Commun. 1999, 20, 7. (a) Komoto, T.; Kawai, T. Makromol. Chem. 1973, 172, 221. (b) Kim, K. Y.; Komoto, T.; Kawai, T. Makromol. Chem. 1977, 178, 2089. (c) Kim, K. Y.; Komoto, T.; Kawai, T. Makromol. Chem. 1979, 180, 465. Padden, F. J., Jr.; Keith, H. D. J. Appl. Phys. 1965, 36, 2987. Kumamaru, F.; Kajiyama, T.; Takayanagi, M. J. Macromol. Sci.Phys. 1978, B15, 87. Gmeiner, P.; Feldman, P. L.; Chu-Moyer, M. Y.; Rapoport, H. J. Org. Chem. 1990, 55, 3068.

BM020016C