Polymer Biocatalysis and Biomaterials - American Chemical Society

Chapter 30

Enzyme-Catalyzed Condensation Reactions for Polymer Modifications 1,2

1,

Downloaded by NORTH CAROLINA STATE UNIV on September 7, 2012 | http://pubs.acs.org Publication Date: February 15, 2005 | doi: 10.1021/bk-2005-0900.ch030

Qu-Ming Gu and H . N . Cheng * 1

Hercules Incorporated Research Center, 500 Hercules Road, Wilmington, DE 19808-1599 Current address: National Starch and Chemical Company, 10 Finderne Avenue, Bridgewater, N J 08807

2

The enzyme-catalyzed condensation is reviewed in view of its use in polymer modification reactions. The formation of esters and amides is most facile when (1) the reaction is carried out in a non-aqueous medium, and (2) the acyl donor contains a good leaving group. However, it is sometimes possible to relax these requirements. Thus, screening and selection of an appropriate enzyme and optimization of the reaction conditions can facilitate esterification and amidation reactions. Water or methanol can be physically removed through the use of vacuum or molecular sieves in order to shift the equilibrium to products and to enhance reaction yield,. For a propitious enzyme-polymer pair, the reaction can proceed even in water without a good leaving group. As illustrations, lipase– catalyzed syntheses are described for the amide of carboxymethylcellulose, substituted acrylic monomers, fatty acid esters of cationic guar, and fatty acid diesters of poly(ethylene glycol).

The use of a lipase to carry out ester synthesis is one of the earliest examples of enzyme-catalyzed reactions in organic media (1). A voluminous amount of papers has been published in the literature, including several reviews and books (2). In the polymer and biomaterials areas, lipases and esterases are

© 2005 American Chemical Society In Polymer Biocatalysis and Biomaterials; Cheng, H., et al.; ACS Symposium Series; American Chemical Society: Washington, DC, 2005.

427

428


also well known and extensively employed (3). This is an active research area, and improved reactions and processes continue to be sought. Much of the earlier work (2-4) entailed acids having activated leaving groups X , where X can be vinyl, trihalomethyl, 2,2,2-haloethyl, or related structures. (R and R' are organic moieties that may be part of a polymer.) RCOOX (acyl donor) + R O H (acyl acceptor) ^ RCOOR' + HOX Thus, numerous papers have been published in the literature on lipasecatalyzed condensation polymerizations using activated diacids, such as vinyl esters (3,4). Whereas these polymerizations are facile, they are not commercially viable due to the cost of the activated diacids. The methyl ester has been attempted (X = CH ); however, the reaction tends to be sluggish. The reaction for the carboxylic acid (X = H) is even more sluggish and is only practical with the removal of water through evaporation, azeotropic distillation, or chemical drying (2a,3). Most of these reactions are carried out in non-aqueous media, e.g., in bulk (neat) or in polar aprotic solvents. Recently, much progress has been made on this type of polymerizations (5). Instead of polymerization reactions, we focus in this work only on modifications of polymers and monomers using enzyme catalysis (6-10). Selected aspects of these reactions are pointed out, especially in terms of yield improvement and process optimization. As in polymerization, the modification reactions are most facile in non-aqueous media when the acyl donor contains a good leaving group. Several cases are given herein to illustrate the scope and the applicability of these reactions. 3

Results and Discussions 1. Fatty acid ester of hydroxyethylcellulose (HEC) This synthesis has been reported earlier (7,8) and represents an optimal case of lipase-catalyzed condensation reaction. ο

HEC

Stearoyl-HEC

Figure 1. Lipase-catalyzed acylation of HEC

In Polymer Biocatalysis and Biomaterials; Cheng, H., et al.; ACS Symposium Series; American Chemical Society: Washington, DC, 2005.


429

Similar to Figure 1, the palmitoyl (C-16) group has also been enzymatically grafted onto HEC. The reaction entails the incubation of HEC, vinyl palmitate, and a suitable enzyme in Ν,Ν-dimethylacetamide (DMAc) at 50°C. Several enzymes were attempted; satisfactory results were obtained with Pseudomonas fluorescens lipase (Amano P-30), Pseudomonas capecia lipase (Lipase PS), or Alcalase® immobilized alkaline protease from Novozymes A/S as a catalyst. Note that in this case a polar aprotic solvent (DMAc) is used, and one of the reactants, vinyl palmitate, has a facile leaving group (vinyl alcohol which leaves as acetaldehyde). 2. The amide derivative of carboxymethylcellulose (CMC) A systematic study of the enzyme-catalyzed synthesis of CMC amides has been reported earlier (9) by incubating an enzyme with CMC and 1,6hexamethylenediamine or allylamine in Ν,Ν-dimethylformamide (DMF). Although a large number of lipases and proteases were screened, the yield of the amide was still found to be 5-25% based on IR analysis. OCHgCOOH

^

^

Ρ^™**^^^^

OH

^ OO-feCOOH

^

^

^ HO

OCH2OOO-

Figure 2. Hydrolase-catalyzed amidation of CMC Note that carboxyl group of CMC has an unfavorable leaving group (OH). As a result, the reaction did not proceed too far even in a non-aqueous solvent with the help of an enzyme. 3. Substituted acrylic monomers In the literature, the general chemical methodology for the synthesis of acrylic monomers is to react methyl acrylate with an alcohol or an amine that carries a desirable functionality X (Figure 3). In this way, the functionality can be incorporated into the monomer and (after polymerization) into a polymer. Enzymes can be useful catalysts for these reactions because the reaction conditions are usually mild, often give less colored products, and tend to generate less byproducts. However, it has been noted that methyl esters give low yields in such reactions (2a).


430 Ο

Ο

Χ ^ ^ O C H

+ 3

Η 0

χ

'

Ο

HOCH

3

+

HOCH

3

Ο

Χ


+

χ

Figure 3. Synthesis of substituted acrylic monomers The reactions shown in Figure 3 were attempted in our laboratory using Novozym® 435 lipase of Novozymes A/S as a catalyst. However, both reactions appeared to reach equilibrium at a certain point and did not proceed further, thereby leading to low yields. In this case the addition of 4A molecular sieves shifted the equilibrium toward product formation by removing methanol. Thus, after several hours of molecular sieve addition, the yield increased to 69%. In addition, the molecular sieves also eliminated the formation of byproducts due to Michael addition. Note that in these examples the enzymatic reactions are assisted by the removal of methanol. Whereas molecular sieves are satisfactory for this purpose, the reactions can be equally enhanced through the use of vacuum (vide infra). 4. Fatty acid diester of poly(ethylene glycol) The fatty esters of polyethers are known to be good surfactants, and many commercial products are available in the market place. Most people use chemical methods to achieve the required synthesis. Enzymatic methods can also be used; however, the reaction needs to be optimized. An example is the lipase-catalyzed synthesis of fatty acid diester of poly(ethylene glycol) (PEG). ο FV

ΌΗ

+

HO,

,OH

Lipase

Figure 4. Lipase-catalyzed synthesis of PEG diester To carry out this reaction, we first screened a number of commercial lipases for substrate selectivity towards the esterification of the fatty acid and


431


poly(ethylene glycol). It was found that lipases from Candida antarctica and Mucor miehei had a strong preference for the fatty acid. The Candida antarctica lipase was chosen because it gave faster rates. We carried out the reaction by adding the enzyme to bulk PEG and fatty acid (at a 1.0 : 1.9 mole ratio). Although we attempted different temperatures, the yield was low, and a mixture of monoesters and diesters was obtained. We discovered that by pulling a vacuum, we could achieve high yields at 50-60°C in 8-48 hours. The final product was isolated byfiltration,which also recovered the enzyme for reuse. The same enzymatic esterification reaction was applied to the enzymatic synthesis of three PEG fatty diesters having PEG molecular weights of 2000, 8000 and 35,000. PEG 2000 fatty ester and PEG 8000 fatty ester are water insoluble, but PEG 35,000 fatty ester is water-soluble. In addition, the PEG 35,000 fatty ester has a much higher solution viscosity than the unmodified PEG in water. At 2%, the Brookfield viscosity of the PEG 35,000 fatty ester is around 340 cps at 30 RPM while the corresponding value for unmodified PEG is less than 3 cps at 30 RPM. Thus, this compound may also be used as a thickener. 5. Fatty acid ester of cationic guar The enzymatic synthesis of this material was reported earlier using vinyl stéarate in a reaction similar to Figure 1 (7). The yield was high, more than 90%. Interestingly, this reaction could also proceed, albeit at lower yields, using palmitic acid and cationic guar in an aqueous buffer. The yield could be improved by adding a small amount of DMF to water f 10). The idea originated from our observation that the low-shear solution viscosity of 1% cationic guar increased threefolds when a catalytic amount of Novolipase was added. As we investigated this reaction, we realized that cationic guar contained 1-2% fatty acids, which became covalently grafted onto the guar molecules. In contrast, underivatized guar gave no reaction under the same reaction conditions (10). This synthesis entails the use of unmodified fatty acid and is done in water. Thus, this represents a normally unfavorable case. There is no convenient leaving group, and water is present in abundance. The reason for this reaction to be possible is the interaction between the enzyme and the cationic charge on guar, which favors the ester formation even in the presence of water.


432

UM

HO Ο

HO,

O H

o

Lipase OH

Ο

OH

Ο


Palmitic Acid

^^^^io^"""""^)

OH

Ο

Figure 5. Lipase-catalyzed esterification of cationic guar

Experimental CMC, HEC, cationic guar, and fatty acids were all products of Hercules Incorporated. The other chemicals used were from commercial sources (SigmaAldrich). The enzymes came variously from Novozymes A/S, Amano Enzyme USA Co. Ltd., Enzyme Development Corp. (EDC), and Sigma-Aldrich. Fatty acid ester of hydroxyethylceilulose (HEC). A slurry containing 40% HEC, 40% vinyl palmitate, and 10% Pseudomonasfluorescenslipase (P-30, Amano) in Ν,Ν-dimethylacetamide was incubated at 50°C for 1-2 days. The resulting material was treated with acetone followed by washing with isopropanol. The modified HEC was obtained as a white solid. The grafting of palmitoyl group was confirmed by Ht analysis (1750 cm" ). 1

The amide derivative of carboxymethylcellulose (CMC). The details of this reaction were reported earlier (9). Basically CMC and the amine were dissolved


433


in Ν,Ν-dimethylformamide (DMG) with a suitable lipase. Twelve lipases and eight proteases were screened. It was observed that Pseudomonas sp. lipase, Subtilisin Carlsberg, Papain, and protease from Aspergillus saitoi produced low levels of amides when the reactions were carried out with DMF as solvent at 50% concentration. The yields of the amides were estimated to be 5-25% based on IR analysis. No amides were observed with all the enzymes when CMC was suspended in toluene or t-butanol even at elevated temperatures (40-60 C). Substituted acrylic monomers. The reactants (methyl acrylate plus alcohol or amine) were added neat or in a non-aqueous solvent together with Novozym® 435 immobilized lipase from Candida antarctica as a catalyst. Molecular sieves (4A) were used to remove water in order to shift the reaction equilibrium to product formation, and also to eliminate side reactions due to Michael addition that was usually enhanced by the presence of water or methanol. Unreacted starting materials were removed by evaporation, and the monomer products obtained without further purification. TLC analysis indicated that the desired products had formed. The purity of the products was confirmed by NMR and IR analysis. The two monomers were successfully polymerized in a separate step. Fatty acid diester of poly(ethylene glycol). The enzymatic esterification was performed in bulk at 50-60°C under vacuum and completed in 8-48 hours. The enzyme use level was 0.1-0.5% based on the substrate. The acid numbers of the ester products depended on the enzyme type, the enzyme use level and the reaction temperature. Progress of each reaction was monitored by TLC analysis (eluted with EtOAc/Hex, 1:3; detected by the phosphomolybdic acid reagent), which showed the disappearance of fatty acid (R = 0.4) and the emergence of the PEG ester (R = 0-0.1). The enzyme was recovered by filtration. The product was obtained as a yellowish liquid and analyzed without further purification. All product structures were confirmed by H and C-NMR analysis. In a typical example, the fatty acid and the PEG were mixed at a molar ratio of 1.9:1. Novozym 435 lipase (0.5-1 gram per kg substrate) was added. At 60°C for 20 hours under vacuum (10-20mm Hg), the acid number was determined to be 8.5-10. C-NMR indicated the conversion of the acid carbon (178 ppm) to the ester carbon (174 ppm), and *H-NMR spectra were used to quantify the ester formation by integrating the proton signals of -0-C-CH OH (3.3-3.6 ppm), - O C-CH OCO-R (4.0-4.2 ppm) and C H - of fatty acids and fatty esters (0.70-0.90 ppm). f

f

l

13

13

2

2

3

Fatty acid ester of cationic guar. Palmitic acid and cationic guar were dissolved in water buffered at pH 6.0 at concentrations of 0.2% and 2.0%, respectively. The enzyme, Novolipase, was used as the biocatalyst. Under these



HEC + vinyl palmitate

HEC ester

2

H 0

DMAc

None

Lipase and protease

DMF

None

Neat or polar Mol sieves aprotic solvents Neat

Lipase

Lipase

Vacuum

Neat or polar Mol sieves aprotic solvents

Lipase

2

Cationic guar + fatty acid H 0

Lipase

2

2

H 0 , or None H Q-DMF (80:20)

Case 4. Mediocre leaving group, unfavorable reaction medium

2

H 0

PEG diester PEG + fatty acid

3

3

CH OH

CH OH

Guar ester

Lipase

Case 3. Mediocre leaving group, favorable reaction medium

Alcohol + Me acrylate

2

Substituted Amine + Me acrylate acrylamide

Substituted acrylate

2

CMC amide NH (CH )6NH 4- C M C

2

3

CH CHO

Case 2. Poor leaving group, favorable reaction medium

Reactants

Product

Enzyme Solvent Leaving Other Aids Group Case 1. Good leaving group, favorable reaction medium

Table 1. Summary of the enzyme-catalyzed condensation reactions


Moderate

High

High/Moderate

High/Moderate

Low

High

Yield

435


reaction conditions after 24 hrs, part of the palmitoyl group was grafted to the cationic guar. The formation of ester bond was confirmed by IR analysis. The yield, however, was only moderate. The yield could be improved by adding a polar aprotic solvent to the solvent medium. Thus, for example, at 40°C in a mixture of water-DMF (80:20) at pH 6.0, cationic guar and palmitic acid produced the corresponding ester in the presence of Novozym 435® lipase in 6-24 hours.

Conclusions In this work several hydrolase-catalyzed condensation reactions have been reviewed with respect to polymer modification reactions. In general, it is true that for these enzymatic reactions a non-aqueous medium and a good leaving group for the acyl donor can provide high yields of esters and amides. However, sometimes it is possible to achieve reasonable yields by employing appropriate reaction conditions and using suitable enzymes. At least four situations, as demonstrated, have helped to enhance these reactions: 1) screening and selection of an appropriate enzyme, 2) removal of one of the products such as water or methanol, 3) favorable enzyme-substrate interactions with less water or alcohol activities, and 4) change in the polarity of the solvent medium (e.g., the addition of DMF) that is favorable for the substrates and the products. A summary of the reactions is given in Table 1.

Acknowledgments Thanks are due to Sadhana Mital, Gordon F. Tozer, and Arleen J. Walton for technical assistance.

References 1. For example, (a) Therisod, M . ; Klibanov, A. M . J. Amer. Chem. Soc. 1986, 108, 5638. (b) Klibanov, A. M . CHEMTECH 1986, 16, 354. 2. For example, (a) Faber, K. Biotransformations in Organic Chemistry, 3 Ed.; Springer-Verlag: Berlin, 1997; p. 309 ff. (b) Biocatalysis for Fine Chemicals Synthesis; Roberts, S. M . , Ed.; Wiley: Chichester, 1999; module 1:9. (c) Engineering of/with Lipases; Malcata, F. X . , Ed.; Kluwer: Dordrecht, 1996. 3. (a) Gross, R. Α.; Kumar, Α.; Kalra, B. Chem. Rev. 2001, 101, 2097. (b) Kobayashi, S.; Uyama, H.; Kimura, S. Chem. Rev. 2001, 101, 3793. (c) Cheng, Η. N.; Gross, R. A. ACS Symp. Ser. 2002, 840, 1.

rd


436


4.

Some recent examples include: (a) Linko, Y.-Y.; Seppälä, J. CHEMTECH 1996, 26(8), 25., and references therein, (b) Wu, X.-Y.; Linko, Y.-Y.; Seppälä, J.; Leisola, M . ; Linko, P. J. Ind. Microbiol. Biotechnol. 1998, 20, 328. (c) Kline, B, J.; Beckman, E.J.; Russell, A. J. J. Amer. Chem. Soc. 1998, 120, 9475. (d) Uyama, H.; Inada, K.; Kobayashi, S. Macromol. Rapid Commun.1999, 20, 171. (e) Binns, F.; Harffey, P.; Roberts, S. M . ; Taylor, A. J. Chem. Soc., Perkin Trans.1 1999, 2671. (f) Park, O.-J.; Kim, D.-Y.; Dordick, J. S. Biotechnol. Bioeng. 2000, 70, 208. (g) Matsumura, S.; Harai, S.; Toshima, K. Macromol. Chem. Phys. 2000, 201, 1632. (h) Mesiano, A. J.; Beckman, E. J.; Russell, A. J. Biotechnol. Prog. 2000, 16, 64. (i) Uyama, H.; Inada, K.; Kobayashi, S. Polym. J. 2000, 32, 440. (j) Uyama, H.; Inada, K.; Kobayashi, S. Macromol. Biosci. 2001, 1, 40. (k) Takamoto, T.; Uyama, H.; Kobayashi, S. e-Polymers 2001, 4, 1. (1) Kim, D.-Y.; Dordick, J. S. Biotechnol. Bioeng. 2001, 76, 200. 5. For example, (a) Kulshrestha, A. S.; Kumar, Α.; Gao, W.; Gross, R. A. ACS Polymer Preprints 2003, 44(2), 635. (b) Mahapatro, Α.; Kumar, Α.; Kalra, B.; Gross, R. A. ACS Polymer Preprints 2003, 44(2), 595. (c) Tsujimoto, T.; Uyama, H.; Kobayashi, S. Biomacromolecules 2001, 2, 29. (d) Kline, B. J.; Lele, S. S.; Lenart, P. J.; Beckman, E. J.; Russell, A. J. Biotechnol. Bioeng. 2000, 67, 424. (e) Park, O.-J.; Kim, D.-Y.; Dordick, J. S. J. Polym. Sci., Polym. Chem. Ed. 2000, 70, 208. 6. For example, (a) Yahya, A. R. M . ; Anderson, W. Α.; Moo-Young, M . Enzyme Microbial Technol.1998, 23, 438. (b) Kitagawa, M . ; Tokiwa, Y. Biotechnology Lett. 1998, 20, 627. (c) Cordova, Α.; Hult, K.; Iversen, T. Biotechnology Lett. 1997, 19, 15. (d) Redmann, I.; Pina, M . ; Guyot, B.; Blaise, P.; Farines, M . ; Graille, J. Carbohydr.Research 1997, 300, 103. (e) Shibatani, S.; Kitagawa, M . ; Tokiwa, Y . Biotechnology Lett. 1997, 19, 511. (f) Bruno, F. F.; Akkara, J. Α.; Ayyagari, M . ; Kaplan, D. L.; Gross, R.; Swift, G.; Dordick, J. S. Macromolecules 1995, 28, 8881. (g) Chen, X . ; Martin, B. D.; Neubauer, T. K.; Linhardt, R. J.; Dordick, J. S.; Rethwisch, D. G. Carbohydr. Polym. 1995, 28, 15. (h) Faber, K.; Riva, S. Synthesis 1992, 895. (i) Hiratake, J.; Yamamoto, K.; Yamamoto, Y.; Oda, J. Tetrahedron Lett. 1989, 30, 1555. (j) Uemura, Α.; Nozaki, K..; Yamashita, J.I.; Yasumoto, M . ; Tetrahedron Lett. 1989, 30, 248. (k) Degueil-Castaing, M . ; De Jeso, B.; Drouillard, S.; Maillard, Β. Tetrahedron Lett. 1987, 28, 953. 7. Gu, Q.-M. ACS Symp. Ser. 2002, 840, 243. 8. Cheng, Η. N . ; Gu, Q.-M. ACS Symp. Ser. 2002, 840, 203. 9. Cheng, Η. N . ; Gu, Q.-M. ACS Polymer Preprints 2000, 41(2), 1873. 10. Gu, Q.-M., poster presented at the 220 National ACS Meeting in Washington, DC, August 20-24, 2000. th


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