Sustainable Enzymatic Preparation of Polyaspartate Using a Bacterial

Jan 21, 2003 - Biomacromolecules , 2003, 4 (2), pp 196–203 ... having an Mw of up to 3700 and a maximum polymer yield of 85%. The best ... Chemical ...
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Biomacromolecules 2003, 4, 196-203

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Articles Sustainable Enzymatic Preparation of Polyaspartate Using a Bacterial Protease Yasuyuki Soeda, Kazunobu Toshima, and Shuichi Matsumura* Faculty of Science and Technology, Keio University, 3-14-1, Hiyoshi, Kohoku-ku, Yokohama 223-8522, Japan Received April 29, 2002

Diethyl L-aspartate was polymerized by a bacterial protease from Bacillus subtilis (BS) in organic solvent at a temperature between 30 and 50 °C to yield R-linked poly(ethyl L-aspartate) having an Mw of up to 3700 and a maximum polymer yield of 85%. The best polymerization conditions were the 40 °C polymerization of diethyl L-aspartate using 30% protease BS containing 4.5 vol % water in acetonitrile for 2 days. Poly(ethyl L-aspartate) was readily depolymerized by the enzyme into the oligomeric and monomeric L-aspartate in aqueous acetonitrile. Poly(sodium aspartate) prepared by the saponification of poly(ethyl L-aspartate) was readily biodegradable by activated sludge obtained from the municipal sewage treatment plant. Also, poly(sodium aspartate) was depolymerized by the hydrolase enzyme into the monomeric aspartate. These results may indicate the sustainable chemical recycling and biorecycling of this polymer. Introduction Significant efforts have been made to develop watersoluble and biodegradable polycarboxylate-type polymers. Poly(malic acid)1-9 and poly(γ-glutamic acid)10 are known to be biodegradable; however, the former contains ester linkages in the backbone and is slowly hydrolyzed in slight alkaline media, which may restrict their practical applications. The latter is stable in aqueous media; however, the number of the functional carboxy groups per molecular weight is lower than that of polyaspartate. Poly(aspartic acid) is attractive as a water-soluble and environmentally acceptable functional polycarboxylate in both the industrial and biomedical fields, because aspartic acid as a raw material is industrially available from renewable and petroleum resources both by chemical and by fermentation methods.11-24 Poly(sodium aspartate) is generally resistant to hydrolysis in slightly alkaline medium. Polyaspartate has several unique characteristics that can be exploited for a range of uses, such as a detergent builder and cosmetic additives. A variety of methods for the synthesis of polyaspartate have been known. The most common method for the production of polyaspartate involves the polycondensation of the appropriate Ncarboxyanhydride, which may not be feasible on an industrial scale and also may not satisfy the concepts of green chemistry. Recently, polyaspartate has been produced on a large scale with dry aspartic acid using a thermal polymerization that yields a polysuccinimide, which can be hydrolyzed in the presence of a base to form polyaspartate. However, it is indicated that the thus-produced polyaspartate contains some complexing structures such as a racemic mixture of aspartate, unreacted imide moieties, branching,

and cross-linking, which might not be responsive to the requirements of biodegradability and biocompatibility. Biodegradation of thermally polymerized polyaspartate has been extensively studied using isolated microbes by Tabata et al. Some polyaspartate-assimilating microbes were isolated, and the biodegradation of the polymer was demonstrated using these isolated microbes.25-27 On the other hand, the enzyme-catalyzed polymerization of aspartic acid may become one of the attractive methods for obtaining homogeneous polyaspartate with respect to green chemistry. However, the enzymatic polymerization method of aspartic acid has so far been restricted only to oligomer formation except for our previous report.28 The oligomerization of dialkyl L-glutamate hydrochloride using protease was reported by Aso et al.29 The oligomerization of dialkyl L-aspartate in addition to L-glutamate has so far been carried out using poly(ethylene glycol)-modified papain as a catalyst to yield a heptamer to decamer in benzene.30 Very recently, Uyama et al. reported the protease-catalyzed regioselective polymerization and copolymerization of diethyl L-glutamate hydrochloride in organic solvent.31 They confirmed using H-H COSY NMR the exclusive formation of poly(R-peptide) by the polymerization of diethyl L-glutamate hydrochloride using papain and bromelain. We previously reported the enzymatic polymerization of diethyl L-aspartate in bulk using alkalophilic proteinase from Streptomyces sp. forming polyaspartate having 88% R-linkages.28 It is reported that this enzyme had relatively wide substrate specificity and catalyzed both esterification and transesterification reactions in addition to amidation reaction.32-35 To apply the enzymatic polymerization method to commodity chemicals, not to

10.1021/bm0200534 CCC: $25.00 © 2003 American Chemical Society Published on Web 01/21/2003

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Enzymatic Preparation of Polyaspartate Scheme 1

Figure 1. Concept of the sustainable production and chemical recycling of polyaspartate using an enzyme.

medical use, large-scale availability of the enzyme is one of the important criteria for the selection of the enzyme. Industrially available microbial protease may be suited for such purposes in place of the precious alkalophilic proteinase from Streptomyces sp. Therefore, in this report, industrially available microbial protease was evaluated with respect to the catalytic activity for polymerization of diethyl L-aspartate. One of the characteristic features of the enzymatic polymerization may include the reverse reaction that enables establishing a new route to sustainable chemical recycling of polyaspartate using an enzyme. Figure 1 shows the concept of a sustainable polymer producton system using an enzyme. Once a polymer was produced using renewable resources, the thus-obtained polymer should be recycled as much as possible by repetitive depolymerization and repolymerization using an enzyme in order to save production energy and resources. There are two routes for the recycling of polyaspartate. One is the enzymatic conversion of polyaspartate into oligomeric and monomeric aspartates by protease as the chemical recycling. The other is the biorecycling route to monomeric aspartate via biodegradation with a subsequent fermentation process. The former may be more straightforward for chemical recycling of polyaspartate. In this paper, the enzyme-catalyzed polymerization of diethyl L-aspartate using the industrially available microbial protease and microbial and enzymatic degradation of polyaspartate were studied with the objectives of the sustainable production and chemical recycling. Experimental Section Materials. Diethyl L- and D-aspartates were prepared by the esterification of L- and D-aspartic acids with ethanol in the presence of thionyl chloride followed by dehydrochlorination to yield diethyl aspartate in 70% yield. Optical purities of the aspartates as measured by high-performance liquid chromatography (HPLC) using chiral crown ether column (Crownpak CR(+), Daicel Chemical Industries, Ltd., Tokyo, Japan) were >99% L- or D-aspartate using aqueous HClO4 (pH 1.5) as the eluent at 0 °C. Protease from Bacillus subtilis (crude enzyme powder of trade name Bioprase, 74% protein content) was kindly supplied by Nagase Chemtex Corp. (Fukuchiyama City, Japan). The enzyme was used after dialysis against distilled water at 4 °C for 24 h. The enzyme gave a single band on SDS-PAGE that corresponded to about 30 kDa. The protein content was 95% as determined by the Lowry method. The other materials used were of the highest available purity.

Measurements. The weight-average molecular weight (Mw), number-average molecular weight (Mn), and molecular weight dispersion (Mw/Mn) of poly(ethyl aspartate) were measured by size exclusion chromatography (SEC) using SEC column (TSK-GEL R-3000, TOSOH Co, Ltd., Tokyo, Japan) at 40 °C with a refractive index detector. Dimethylformamide containing 10 mM LiBr was used as the eluent at 0.8 mL/min. The SEC system was calibrated with poly(ethylene oxide) standards having a narrow molecular weight distribution, hexa(ethylene glycol) and di(ethylene glycol). The molecular weight of poly(sodium aspartate) was measured by SEC using a SEC column (Asahipak GS 320, Asahi Chemical Industry Co., Ltd., Tokyo, Japan) at 30 °C with an UV detector (220 nm). Phosphate buffer (0.1 M) containing 0.3 M NaCl (pH 6.8) was used as the eluent. The SEC system was calibrated with poly(sodium acrylate) standards of a narrow molecular weight distribution and hexamer, tetramer, and dimer of sodium acrylate. 1H NMR spectra were recorded with a JEOL model Lambda 300 (300 MHz) spectrometer (JEOL Ltd., Tokyo, Japan). 13C NMR spectra were recorded with a JEOL Lambda 300 Fourier transform spectrometer operating at 75 MHz with complete proton decoupling. The molecular weight was also measured by matrix-assisted laser desorption ionization time-of-flight mass spectrometry (MALDI-TOF MS). The MALDI-TOF MS was measured with a Bruker Proflex mass spectrometer after treatment with lithium trifluoromethanesulfonate. The spectrometer 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. Infrared (IR) spectra were measured using a JASCO Fourier transform spectrometer model FT/IR-5200 (JASCO Ltd., Tokyo, Japan). Enzymatic Polymerization. The enzyme-catalyzed polymerization of diethyl L-aspartate was carried out in organic solvent containing a small amount of water using protease BS as shown in Scheme 1. A typical preparation of poly(ethyl L-aspartate) was carried out as follows. A mixture of diethyl L-aspartate (54.5 mg), protease BS (5.5 mg, 10% relative to monomer), and 1 mL of acetonitrile (MeCN) containing 45 µL of distilled water (4.5 vol %) was stirred under an argon atmosphere in a capped vial placed in a oil bath at 40 °C for 2 days. After the reaction, anhydrous sodium sulfate (1.5 g) was added and stirred to remove water, the reaction mixture was then dissolved in 5 mL of a mixed

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solvent of chloroform and trifluoroacetic acid (100:3, v/v), and the insoluble enzyme was removed by filtration. The chloroform was then evaporated under reduced pressure at 1 mmHg to obtain crude products consisting of poly(ethyl L-aspartate), unreacted diethyl L-aspartate, and β-ethyl Laspartate. The composition of these compounds was determined by 1H NMR. The molecular structure was analyzed by SEC and FT-IR, 1H NMR, and 13C NMR spectroscopies. The spectral data of poly(ethyl L-aspartate) are shown as being representative. IR (KBr): 3428 (NH), 2984 (CH2), 1738, 1188 (ester CdO), 1658, 1560 (CONH) cm-1. 1H NMR (300 MHz, DMSO-d6): δ 1.18 (m, 3H, CH3 of Et), 2.5-2.8 (m, 2H, CH2), 4.06 (m, 2H, CH2 of Et), 4.56 (m, 1H, R-linkage CH), 8.0-8.3 (m, 1H, CONH). 13C NMR (75 MHz, DMSO-d6): δ 14.1 (CH3 of Et), 36.0 (CH2), 49.7 (CH), 60.3 (CH2 of Et), 170.2 (R-R CdO). Saponification of Poly(ethyl L-aspartate). The resulting poly(ethyl L-aspartate) was saponified into the corresponding poly(sodium aspartate). A mixture of poly(ethyl L-aspartate) (55 mg) and 1.3 equiv of 0.5 N NaOH in methanol and water (3:1, v/v) (4.8 mL) was stirred at 25 °C for 4 h. The reaction mixture was neutralized to pH 7 using 1 N HCl and then dialyzed against distilled water to remove inorganic salts and monomeric disodium aspartate. Through these procedures, the R-linkage of polyaspartate was partially inverted through polysuccinimide to produce R- and β-linkage poly(sodium aspartate).28 The R/β ratio of poly(sodium aspartate) was determined by comparison of the 1H NMR spectral integration intensities of the δ ) 4.6 ppm peaks corresponding to the R-methyne proton in poly(ethyl L-aspartate) with the β-methyne proton at δ ) 4.3 ppm. The spectral data for poly(sodium aspartate) having an Mw of 2450 are shown as being representative. IR(KBr): 3450 (NH), 1639 (CONH), 1597, 1400 (COONa) cm-1. 1H NMR (300 MHz, D2O): δ 2.63.1 (m; 2H, CH), 4.53 (m, 0.81H, β-linkage CH), 4.70 (m, 0.19H, R-linkage CH). 13C NMR (75 MHz, D2O): δ 39.7 (CH2), 52.3 (CH), 172.9 (β-β CdO), 173.5 (R-β, β-R CdO), 174.0 (R-R CdO), 178.6 (COONa). Biodegradation. Biodegradability of the poly(sodium aspartate) was evaluated by biochemical oxygen demand (BOD) measurements. BOD was determined with a BOD tester (model 200F, TAITEC Corp., Koshigaya-shi) by the oxygen consumption method, according to the modified MITI test.36 The activated sludge was obtained from a municipal sewage plant in Yokohama City. BOD-biodegradation (BOD/ThOD) was calculated from the BOD values and the calculated theoretical oxygen demand (ThOD). Enzymatic depolymerization using cell-free extracts of polyaspartate-assimilating microbe as the polyaspartatehydrolyzing enzyme source was carried out as follows. BreVibacillus reuszeri KS018 capable of degrading poly(sodium aspartate) was isolated from activated sludge by an enrichment culture technique using poly(sodium aspartate) as the sole carbon source. BreVibacillus reuszeri KS018 was grown in an inorganic medium containing 0.01% yeast extract and 1.0% citric acid as the growing substrate in a shake flask with reciprocal shaking at 30 °C. The initial pH of the medium was adjusted to 7.0. After 3 days, the cells were harvested by centrifugation (12000 rpm, 30 min, 4 °C).

Soeda et al. Table 1. Effect of Solvents on the Polymerization of Diethyl L-Aspartate by Protease BS at 40 °C for 2 days entry 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15

vol % H2O in solvent

solvent DMF

1,4-dioxane

MeCN

ethanol

toluene

0 2 5 0 2 5 0 2 5 0 2 5 0 2 5

Mw

M h w/M hn

300 350

1.1 1.1

900 2500

1.7 2.2

1000 1500

1.9 2.1

1600 1800

1.6 1.5

polymer yield (%) 0 0 0 0 2 7 0 11 37 0 17 29 0 26 27

The wet cells were washed once with 0.8% aqueous NaCl (20 mL) and centrifuged (12000 rpm, 30 min, 4 °C) to obtain the wet cells (5.5 g from 1 mL of medium). The cells (2 g) were suspended in 100 mM phosphate buffer (pH 7.0, 10 mL) and 0.5% Tween 80 and then disrupted with an ultrasonic oscillator (19 kHz, 125 W, 50% pulse) for 30 min at 4 °C. The cell debris was removed by centrifugation at 12000 rpm for 30 min at 4 °C to obtain the cell-free extracts. These cell-free extracts were used for the depolymerization tests of poly(sodium aspartate). Enzymatic depolymerization of poly(sodium aspartate) into the corresponding monomeric aspartate was evaluated using the cell-free extracts of BreVibacillus reuszeri KS018. A mixture of polymer (5 mg), cell-free extracts (2 mL), and 0.8% aqueous NaCl (3 mL) was incubated at 30 °C with stirring for 2 days. The reaction mixture was directly analyzed by SEC, and the yield of the liberated monomeric aspartate was measured by SEC using the calibration curve. Results and Discussion Protease-Catalyzed Polymerization of Diethyl L-Aspartate. It was confirmed that diethyl L-aspartate was polymerized by protease BS in an organic solvent containing a small amount of water at a temperature between 30 and 50 °C to produce R-linked poly(ethyl L-aspartate). Without the enzyme, no polymerization of diethyl L-aspartate occurred as analyzed by SEC and 1H NMR. It was also confirmed that free L-aspartic acid and β-ethyl L-aspartate alone did not polymerize under the same conditions as shown in Scheme 1. It was also found that the R-hydrolyzed monomer, β-ethyl L-aspartate, was produced by the hydrolysis through the action of protease BS with water as analyzed by SEC and 1H NMR. This R-hydrolyzed monomer might influence further polymerization. Details of the polymerization mechanism were analyzed. The polymerization of diethyl L-aspartate was dependent on the solvent used. Table 1 shows the polymerization of diethyl L-aspartate (54.5 mg) using various organic solvents (1 mL) containing 0, 2, and 5 vol % water using 10% protease BS (relative to the monomer) at 40 °C for 2 days.

Enzymatic Preparation of Polyaspartate

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Figure 2. Effects of water content on polymerization of diethyl L-aspartate: b, molecular weight; 9, polymer yield; 2, β-ethyl L-aspartate. Diethyl L-aspartate (54.5 mg) was polymerized using protease BS (5.5 mg) in 1 mL of MeCN containing water at 40 °C for 2 days. Figure 4. Effects of temperature on polymerization of diethyl L-aspartate: b, molecular weight and molecular weight dispersion; 9, polymer yield; 2, β-ethyl L-aspartate. Diethyl L-aspartate (54.5 mg) was polymerized using protease BS (5.5 mg) in 1 mL of MeCN containing 4.5 vol % water for 2 days.

Figure 3. Effects of water content on molecular weight of poly(ethyl L-aspartate). polymerization of diethyl L-aspartate: b, 30% protease BS; 2, 10% protease BS. Diethyl L-aspartate (54.5 mg) was polymerized using 10 and 30% protease BS (relative to monomer) in 1 mL of MeCN containing water at 40 °C for 2 days.

It was found that the polymerization reaction was facilitated in acetonitrile (MeCN), toluene, and ethanol in that order. However, no significant polymerization was observed in dimethyl formamide (DMF) and 1,4-dioxane. Also, the water content was responsible for both polymer yield and the molecular weight of the resulting polymer. Without water, practically no polymerization occurred, irrespective of the solvent used. Therefore, further studies were carried out using MeCN containing water. Effects of Polymerization Conditions. It was found that a small amount of water was needed for the polymerization of diethyl L-aspartate by protease BS. Figure 2 shows the molecular weight of the resulting poly(ethyl L-aspartate) as a function of the water content in MeCN by the polymeri-

zation of diethyl L-aspartate (54.5 mg) using 10% protease BS (relative to monomer) in MeCN (1 mL) at 40 °C for 2 days. Figure 2 also shows the polymer yield and β-ethyl L-aspartate formation as a function of water content in MeCN as measured by SEC and 1H NMR. It was found that both the Mw and monomer conversion were significantly influenced by the water content in MeCN and similar tendencies were observed. Both the Mw of the resulting polymer and the polymer yield significantly increased with the addition of water from 0 to 4.5 vol %. However, the addition of an excess amount of water decreased in both Mw and polymer yield with the increasing R-hydrolyzed monomer, β-ethyl L-aspartate. Maximum Mw and polymer yield were obtained at 4.5 vol % water content in MeCN. On the other hand, the R-hydrolyzed monomer was increased with the increasing water content in MeCN. This is ascribed to the competitive reactions of hydrolysis of the R-ethyl ester of the monomer and polycondensation of diethyl L-aspartate by the action of enzyme, and the former was more promoted by the excess amount of water. It was also found that with the increasing enzyme concentration from 10 to 30%, the optimum water concentration of 4.5 vol % in MeCN remained unchanged as shown in Figure 3. Increasing the enzyme concentration from 10 to 30% caused an increase in molecular weight of the resulting polymer and the polymer yield. Figure 4 shows the Mw of polyaspartate and the polymer yield as a function of polymerization temperature for a 2-day

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Figure 5. Effects of enzyme concentration on polymerization of diethyl L-aspartate: b, molecular weight and molecular weight dispersion; 9, polymer yield; 2, β-ethyl L-aspartate. Diethyl L-aspartate (54.5 mg) was polymerized using protease BS in 1 mL of MeCN containing 4.5 vol % water at 40 °C for 2 days.

polymerization of diethyl L-aspartate using 10% protease BS and in 1 mL of MeCN containing 4.5 vol % water. It was confirmed that diethyl L-aspartate polymerized by protease BS at a temperature between 30 and 50 °C. From the 1H NMR analysis, no imide bond was detected in the resulting polymer at a polymerization temperature below 50 °C. From these results, it is regarded that the similar polymerization results were obtained at a polymerization temperature between 30 and 50 °C. The polymerization results for diethyl L-aspartate were influenced by the enzyme concentration. Figure 5 shows the molecular weight of the resulting poly(ethyl L-aspartate), polymer yield, and β-ethyl L-aspartate as a function of enzyme concentration by the polymerization of diethyl L-aspartate (54.5 mg) using protease BS in 1 mL of MeCN containing 4.5 vol % water at 40 °C for 2 days. It was observed that the Mw of the polymer increased gradually with the increasing enzyme concentration from 5 to 40%. This might be ascribed to the low catalytic efficiency in the polymerization system where the produced polymer was partially solidified. Therefore, a large amount of enzyme promoted the polymerization. However, the polymer yield remained around 50% at enzyme concentrations between 5 and 40%. This might be ascribed to the formation of the R-hydrolyzed monomer, β-ethyl L-aspartate, in a yield around 50%. The polymerization of diethyl L-aspartate was also influenced by the monomer concentration in MeCN solution.

Soeda et al.

Figure 6. Effects of monomer concentration on polymerization of diethyl L-aspartate: b, molecular weight and molecular weight dispersion; 9, polymer yield; 2, β-ethyl L-aspartate. Diethyl L-aspartate was polymerized using protease BS (5.5 mg) in 1 mL of MeCN containing 4.5 vol % water at 40 °C for 2 days.

Figure 6 shows the molecular weight of the resulting poly(ethyl L-aspartate), the polymer yield, and β-ethyl L-aspartate as a function of monomer concentration in MeCN by the polymerization of diethyl L-aspartate using 30% protease BS (relative to monomer) in MeCN containing 4.5 vol % water at 40 °C for 3 days. It was observed that the polymer yield increased gradually with the increasing monomer concentration. This is ascribed to the decreasing R-hydrolyzed monomer with increasing monomer concentration. At the monomer concentration of 260 g/L, the polymer yield exceeded 85%. Increasing polymer concentration caused the partial solidification of the polymerization system, thus inhibiting the further polycondensation with the monomer. Therefore, the Mw of the resulting polymer slightly decreased with the increasing monomer concentration from 50 to 260 mg/mL. Figure 7 shows the time course of the Mw of the resulting poly(ethyl L-aspartate), polymer yield, and β-ethyl L-aspartate by the polymerization of diethyl L-aspartate (54.5 mg) using 30% protease BS (relative to monomer) in MeCN (1 mL) containing 4.5 vol % water at 40 °C. Similar tendencies were observed for Mw and polymer yield in the course of polymerization time. That is, both Mw and polymer yield gradually increased within 1 day and the Mw and polymer yield exceeded 3000 and 70%, respectively. The Mw and polymer yield then remained almost constant. On the other hand, the R-hydrolyzed monomer, β-ethyl L-aspartate, was

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Enzymatic Preparation of Polyaspartate

Figure 8. Typical MALDI-TOF MS spectrum of poly(ethyl L-aspartate) obtained after 2 days of polymerization of diethyl L-aspartate (54.5 mg) using protease BS (5.5 mg) in 1 mL of MeCN containing 2.5 vol % water at 40 °C. Scheme 2

Figure 7. Time course of the polymerization of diethyl L-aspartate (163.5 mg) using protease BS (49.0 mg) in 1 mL of MeCN containing 4.5 vol % water at 40 °C: b, molecular weight and molecular weight dispersion; 9, polymer yield; 2, β-ethyl L-aspartate.

slightly decreased with polymerization time for 2 days, suggesting that β-ethyl L-aspartate acted as a nucleophile to slowly incorporate at the carboxy-terminal of the polymer chain. Thus-produced polymer species have a free carboxylic acid group at the carboxy terminal. These phenomena are also confirmed by MALDI-TOF MS. Details of the polymerization mechanism are discussed in a later section. Enantioselectivity of Protease BS for L-Aspartate Polymerization. Though diethyl L-aspartate was readily polymerized by protease BS, no polymerization occurred for diethyl D-aspartate using the enzyme under the same conditions. That is, a mixture of diethyl D-aspartate (54.5 mg) and protease BS (5.5 mg) was stirred in 1 mL of MeCN containing water (45 µL, 4.5 vol %) at 40 °C for 3 days. Only a few percent of the R-hydrolyzed monomer was produced without polymer formation. More than 90% diethyl D-aspartate was recovered unchanged after 3 days of incubation at 40 °C. Therefore, enantioselectivity of the polymerization by protease BS can be regarded as L-form specific. Proposed Polymerization Mechanism. It was found that poly(ethyl L-aspartate) prepared by polycondensation of diethyl L-aspartate using protease BS exclusively consisted of an R-linked linear polymer having either an ethyl ester or a free carboxy end group at the carboxy terminal of the polymer chain. The free carboxy end group tended to increase by decreasing the water concentration. Figure 8 shows the entire spectrum of the MALDI-TOF MS of poly(ethyl L-aspartate) obtained by the relatively low water concentra-

tion of 2.5 vol %. It was confirmed that the repeating units have a mass of m/z 146, which confirms the polymer structures shown in Scheme 1. Furthermore, each tall peak corresponded to the poly(ethyl aspartate) having an ethyl ester end group. The protease-catalyzed polymerization mechanism seems to be similar to the lipase-catalyzed polymerization of lactones.37,38 That is, the polymerization is initiated by the reaction of the serine residue at the active site of the enzyme with diethyl L-aspartate to form an acyl-enzyme intermediate as shown in Scheme 2. The initiation is nucleophilic attack by the amino group of diethyl aspartate on the acyl carbon of the acyl-enzyme intermediate to produce aspartate dimer. In the propagation state, the acyl-enzyme intermediate is nucleophilically attacked by the terminal amino group of a propagating oligomer/polymer to give one-unit-more elongated polymer. It was also found that β-ethyl L-aspartate was produced as the consequence of hydrolysis of the R-ethyl ester group of the monomer when water attacked as a nucleophile to the acyl-enzyme intermediate. The amino group of this β-ethyl aspartate also acted as a nucleophile to attack the acyl-enzyme intermediate, thus forming polyaspartate species having a terminal free carboxy group. That is, this β-ethyl L-aspartate might act only as a nucleophile at the terminal amino group because the free carboxylic acid terminal could not react with the protease to form acylenzyme intermediate. To further analyze the terminal group

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Figure 9. Expanded MALDI-TOF MS spectrum of poly(ethyl Laspartate) obtained after 4, 8, and 24 h. Polymerization of diethyl L-aspartate (163.5 mg) was carried out using protease BS (49.0 mg) in 1 mL of MeCN containing 4.5 vol % water at 40 °C.

of the resulting polymer, the MALDI-TOF MS of the resulting polymer was measured periodically. Figure 9 shows the expanded form of MALDI-TOF MS of poly(ethyl L-aspartate) produced after 4, 8, and 24 h. It was confirmed that the repeating units have a mass of m/z 146, which confirms the polymer structures shown in Scheme 1. After 4 h of polymerization, a mass of m/z (146n + 46 + Li+), which was equal to the structure of poly(ethyl L-aspartate) having an ethyl ester end group, was mainly obtained. In this case, the amino group of diethyl L-aspartate and poly(ethyl aspartate) having an ethyl ester end group acted as a nucleophile. A mass of m/z (146n + 18 + Li+), which was equal to the structure of poly(ethyl L-aspartate) having a free carboxy end group, was gradually increased with time. Biodegradation of Water-Soluble Poly(sodium aspartate). The saponification of poly(ethyl L-aspartate) was carried out using 0.5 M NaOH at 25 °C in order to obtain water-soluble poly(sodium aspartate). The ratio of R- and β-linkages, which were determined by the area ratio of the 1 H NMR spectrum, was almost constant at ca. 1/4, which indicates that the isomerization between R- and β-linkages occurred via imide intermediate by the alkaline hydrolysis.13,20 It is reported that polyaspartate obtained by the thermal polymerization of dry L-aspartic acid consisted of 30% R-linkages and 70% β-linkages.20 It was also confirmed that the partial epimerization of L-aspartate residue occurred during the saponification of poly(ethyl L-aspartate) using 0.5 M NaOH in aqueous methanol. That is, poly(sodium aspartate) was hydrolyzed by heating with 6 M HCl at 80 °C for 1 day to produce the monomeric aspartic acid. The optical purity of the obtained aspartic acid was measured by HPLC using chiral column. It was found that the D-aspartate unit was detected in the poly(sodium aspartate) and the D-form content was dependent on the saponification conditions. It was also confirmed that no epimerization was detected using optically pure diethyl L-aspartate under these saponification conditions. Poly(sodium aspartate) obtained by the enzyme-catalyzed polymerization of diethyl L-aspartate was readily biodegraded by the activated sludge freshly obtained from the municipal sewage treatment plant in Yokohama city as shown in Figure 10. That is, the biochemical oxygen demand (BOD) biode-

Soeda et al.

Figure 10. BOD biodegradation of poly(sodium aspartate) using activated sludge at 30 °C: b, poly(sodium aspartate); 9, aniline.

Figure 11. Typical SEC profile changes of poly(sodium aspartate) before and after the 2-day incubation with the cell-free extracts at 30 °C.

gradability was over 60% after a 2-week incubation using the oxygen consumption method, basically according to the OECD Guidelines for Testing of Chemicals (301C, modified MITI test) at 25 °C. These results may establish sustainable biorecycling of water-soluble polymer that may not be recovered after use. The enzymatic depolymerization of polyaspartate into monomeric aspartate was evaluated with respect to monomer recycling as shown in Figure 1 using the cell-free extracts of BreVibacillus reuszeri KS018 as the hydrolyzing enzyme source. Figure 11 shows the SEC profile changes of poly(sodium aspartate) before and after the 2-day incubation with the cell-free extracts. It was confirmed that the single peak for the monomeric asparte appeared by the enzymatic depolymerization and the major part of the polymer peak disappeared with the cell-free extracts in 2 days. These results may establish sustainable biorecycling for this polymer. The specific enzyme that catalyzed the depolymerization of poly(sodium aspartate) is now under study. Poly(ethyl L-aspartate) was also depolymerized into oligomeric and monomeric L-aspartate by protease BS. That

Enzymatic Preparation of Polyaspartate

is, depolymerization occurred using poly(ethyl L-aspartete) (1 mg) and protease BS (10 mg) in 1 mL of acetonitrile containing 10 vol % water with stirring at 40 °C. Details of the depolymerization and repolymerization will be analyzed in the subsequent paper. Conclusion It was found that diethyl L-aspartate was readily polymerized by microbial protease BS containing a small amount of water to produce all R-linked poly(ethyl L-aspartate) having an Mw of up to 3700 in a maximum yield of 85%. The best polymerization conditions were the 40 °C polymerization of diethyl L-aspartate using protease BS containing 4.5 vol % water in MeCN for 2 days. This poly(ethyl L-aspartate) was readily depolymerized into oligomeric and monomeric L-aspartate by the protease in aqueous acetonitrile solution. It was found that water-soluble poly(sodium aspartate) was readily biodegraded by activated sludge. Also, poly(sodium aspartate) was converted to monomeric aspartate by the cell-free extracts of the polyaspartate-assimilating bacteria as the hydrolyzing enzyme source. These results may establish sustainable chemical recycling and biorecycling for this polymer. Acknowledgment. We acknowledge the gift of protease from Bacillus subtilis from Nagase Chemtex Corp. (Fukuchiyama City, Japan). References and Notes (1) Vert, M.; Lenz, R. W. Polym. Prepr. (Am. Chem. Soc., DiV. Polym. Chem.) 1979, 20, 608. (2) Gue´rin, Ph.; Vert, M.; Braud, C.; Lenz, R. W. Polym. Bull. (Berlin) 1985, 14, 187. (3) Abe, Y.; Matsumura, S.; Imai, K. J. Jpn. Oil Chem. Soc. 1986, 35, 937; Chem. Abstr. 1987, 106, 33789x. (4) Arnold, S. C.; Lenz, R. W. Makromol. Chem., Macromol. Symp. 1986, 6, 285. (5) Ouchi, T.; Fujino, A. Makromol. Chem. 1989, 190, 1523. (6) Benevenuti, M.; Lenz, R. W. J. Polym. Sci., Polym. Chem. Ed. 1991, 29, 793. (7) Matsumura, S.; Beppu, H.; Nakamura, K.; Osanai, S.; Toshima, K. Chem. Lett. 1996, 795. (8) Matsumura, S.; Beppu, H.; Toshima, K. In Enzymes in Polymer Synthesis; Gross, R. A., Kaplan, D. L., Swift, G., Eds.; ACS Symp. Ser. 684; American Chemical Society: Washington, DC, 1998; p 74. (9) Matsumura, S.; Beppu, H.; Toshima, K. Chem. Lett. 1999, 249. (10) Kunioka, M.; Goto, A. Appl. Microbiol. Biotechnol. 1994, 40, 867. (11) Swift, G. Acc. Chem. Res. 1993, 26, 105. (12) Low, K. C.; Koskan, L. P. Polym. Mater. Sci. Eng. 1993, 69, 253.

Biomacromolecules, Vol. 4, No. 2, 2003 203 (13) Rao, V. S.; Lapointe, P.; McGregor, D. N. Makromol. Chem. 1993, 194, 1095. (14) Swift, G. Polym. Degrad. Stabil. 1994, 45, 215. (15) Alford, D. D.; AWheeler, A. P.; Pettigrew, C. A. J. EnViron. Polym. Degrad. 1994, 2, 225. (16) Cromwick, A.-M.; Gross, R. A. Int. J. Biol. Macromol. 1995, 17, 259. (17) Low, K. C.; Wheeler, A. P.; Koskan, L. P. In Hydrophilic Polymers; Glass, J. E., Ed.; Adv. Chem. Ser. 248; American Chemical Society: Washington, DC, 1996; p 99. (18) Freeman, M. B.; Paik, Y. H.; Swift, G.; Wilczynski, R.; Wolk, S. K.; Yocom, K. M. In Hydrogels and Biodegradable Polymers for Bioapplications; Ottenbrite, R. M., Huang, S. J., Park, K., Eds.; ACS Symp. Ser. 627; American Chemical Society: Washington, DC, 1996; p. 118. (19) Tomida, M.; Nakano, T.; Kuramochi, M.; Shibata, M.; Matsunami, S.; Kakuchi, T. Polymer 1996, 37, 19. (20) Roweton, S.; Huang, S. J.; Swift, G. J. EnViron. Polym. Degrad. 1997, 5, 175. (21) Swift, G.; Freeman, M. B.; Paik, Y. H.; Simon, E.; Wolk, S.; Yocom, K. M. Macromol. Symp. 1997, 123, 195. (22) Ross, R. J.; Batzel, D. A.; Meah, A. R.; Kneller, J. F. Macromol. Symp. 1997, 123, 235. (23) Swift, G.; Creamer, M.; Wei, X.; Yocom, K. M. Macromol. Symp. 1998, 130, 379. (24) Shibata, M.; Kusuno, A.; Kakuchi, T.; Nakato, T.; Yoshikawa, M.; Tomida, M. Macromol. Symp. 1998, 130, 229. (25) Tabata, K.; Kasuya, K.; Abe, H.; Masuda, K.; Doi, Y. Appl. EnViron. Microbiol. 1999, 65, 4268. (26) Tabata, K.; Abe, H.; Doi, Y. Biomacromolecules 2000, 1, 157. (27) Tabata, K.; Kajiyama, M.; Abe, H.; Yamato, I.; Doi, Y. Biomacromolecules 2001, 2, 1155. (28) Matsumura, S.; Tsushima, Y.; Otozawa, N.; Murakami, S.; Toshima, K.; Swift, G. Macromol. Rapid Commun. 1999, 20, 7. (29) Aso, K.; Uemura, T.; Shiokawa, Y. Agric. Biol. Chem. 1998, 52, 2443. (30) Uemura, T.; Fujimori, M.; Lee, H.-H.; Ikeda, S.; Aso, K. Agric. Biol. Chem. 1990, 54, 2277. (31) Uyama, H.; Fukuoka, T.; Komatsu, I.; Watanabe, T.; Kobayashi, S. Biomacromolecules 2002, 3, 318. (32) Tokiwa, Y.; Kitagawa, M.; Fan, H.; Yokochi, T.; Raku, T.; Hiraguri, Y.; Shibatani, S.; Maekawa, Y.; Kashimura, N.; Kurane, R. Biotechnol. Lett. 1999, 13, 563. (33) Kitagawa, M.; Fan, H.; Raku, T.; Shibatani, S.; Maekawa, Y.; Hiraguri, Y.; Kurane, R.; Tokiwa, Y. Biotechnol. Lett. 1999, 21, 355. (34) Kitagawa, M.; Tokiwa, T.; Fan, H.; Raku, T.; Tokiwa, Y. Biotechnol. Lett. 2000, 22, 879. (35) Kitagawa, M.; Fan, H.; Raku, T.; Kurane, R.; Tokiwa, Y. Biotechnol. Lett. 2000, 22, 883. (36) OECD Guidelines for Testing of Chemicals, 301C, Modified MITI Test, Organization for Economic Cooperation and Development (OECD), Paris, 1981. (37) Uyama, H.; Takeya, K.; Kobayashi, S. Bull. Chem. Soc. Jpn. 1995, 68, 56. (38) 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.

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