Enzymes as Green Catalysts for Precision Macromolecular Synthesis

Review pubs.acs.org/CR

Enzymes as Green Catalysts for Precision Macromolecular Synthesis Shin-ichiro Shoda,† Hiroshi Uyama,‡ Jun-ichi Kadokawa,§ Shunsaku Kimura,∥ and Shiro Kobayashi*,⊥ †

Department of Biomolecular Engineering, Graduate School of Engineering, Tohoku University, Aoba-ku, Sendai 980-8579, Japan Department of Applied Chemistry, Graduate School of Engineering, Osaka University, Yamadaoka, Suita 565-0871, Japan § Department of Chemistry, Biotechnology, and Chemical Engineering, Graduate School of Science and Engineering, Kagoshima University, Korimoto, Kagoshima 890-0065, Japan ∥ Department of Material Chemistry, Graduate School of Engineering, Kyoto University, Katsura, Nishikyo-ku, Kyoto 615-8510, Japan ⊥ Center for Fiber & Textile Science, Kyoto Institute of Technology, Matsugasaki, Sakyo-ku, Kyoto 606-8585, Japan ‡

ABSTRACT: The present article comprehensively reviews the macromolecular synthesis using enzymes as catalysts. Among the six main classes of enzymes, the three classes, oxidoreductases, transferases, and hydrolases, have been employed as catalysts for the in vitro macromolecular synthesis and modification reactions. Appropriate design of reaction including monomer and enzyme catalyst produces macromolecules with precisely controlled structure, similarly as in vivo enzymatic reactions. The reaction controls the product structure with respect to substrate selectivity, chemo-selectivity, regio-selectivity, stereoselectivity, and choro-selectivity. Oxidoreductases catalyze various oxidation polymerizations of aromatic compounds as well as vinyl polymerizations. Transferases are effective catalysts for producing polysaccharide having a variety of structure and polyesters. Hydrolases catalyzing the bond-cleaving of macromolecules in vivo, catalyze the reverse reaction for bond forming in vitro to give various polysaccharides and functionalized polyesters. The enzymatic polymerizations allowed the first in vitro synthesis of natural polysaccharides having complicated structures like cellulose, amylose, xylan, chitin, hyaluronan, and chondroitin. These polymerizations are “green” with several respects; nontoxicity of enzyme, high catalyst efficiency, selective reactions under mild conditions using green solvents and renewable starting materials, and producing minimal byproducts. Thus, the enzymatic polymerization is desirable for the environment and contributes to “green polymer chemistry” for maintaining sustainable society.

CONTENTS 1. Introduction 2. Enzymes and Enzymatic Catalysis 3. Enzymes as Green Catalyst 3.1. Clean Reaction Processes Involving High Selectivity 3.2. Characteristics of Enzymes as Catalyst 3.3. Starting Raw Materials 3.4. Reaction Products 3.5. Reaction Solvents 3.6. Polymer Recycling and Degradation 4. Oxidoreductases 4.1. Peroxidases 4.1.1. Polymerization of Aromatic Compounds 4.1.2. Polymerization of Vinyl Monomers 4.2. Laccase, Tyrosinase, and Bilirubin Oxidase 4.2.1. Polymerization of Phenolic Compounds 4.2.2. Synthesis Approach to Artificial Urushi 4.2.3. Polymerization of Vinyl Monomers 4.2.4. Other Reactions 5. Transferases 5.1. Glycosyltransferases 5.1.1. α-Glucan Phosphorylase 5.1.2. Cellodextrin Phosphorylase 5.1.3. Glucansucrase and Fructansucrase © 2016 American Chemical Society

5.1.4. Branching Enzyme 5.2. Acyltransferases 5.2.1. Polyester Synthase 5.2.2. Transglutaminase 6. Hydrolases 6.1. General Introduction of Hydrolases 6.2. Glycosidases 6.2.1. Glycosidase Catalysis for Polycondensations 6.2.2. Cellulase 6.2.3. Xylanase, 4-Glucanohydrolase, and Amylase 6.2.4. Glycosidase Catalysis for Ring-Opening Polyadditions 6.2.5. Hyaluronidase 6.2.6. Keratanase 6.2.7. Endo-N-acetylglucosaminidase: Modification of Polypeptides 6.3. Lipases 6.3.1. Synthesis of Polyesters

2308 2308 2310 2310 2310 2310 2310 2310 2310 2311 2311 2311 2319 2319 2320 2323 2327 2328 2330 2330 2331 2340 2341

2342 2343 2343 2344 2344 2344 2345 2345 2348 2350 2351 2354 2354 2355 2355 2355

Special Issue: Frontiers in Macromolecular and Supramolecular Science Received: August 10, 2015 Published: January 21, 2016 2307

DOI: 10.1021/acs.chemrev.5b00472 Chem. Rev. 2016, 116, 2307−2413

Chemical Reviews 6.3.2. Synthesis of Other Polymers 6.3.3. Modification of Polysaccharides 6.4. Proteases 6.4.1. Synthesis of Polyesters 6.4.2. Synthesis of Polyamides 7. Concluding Remarks Author Information Corresponding Author Notes Biographies Acknowledgments References

Review

renewable biomass materials rather than fossil materials, nontoxic degradable products for the target, etc. We have been studying the enzyme-catalyzed polymer synthesis, “enzymatic polymerization”, as a new general method for macromolecular synthesis for more than two and half decades and have published comprehensive or short review papers and edited special issues from time-to-time.27−58 Also, other groups published related review papers or edited special issues in this area.59−66 From the environmental aspects, enzymes are natural and renewable and, hence, belong to a green catalyst. Thus, working on enzymatic polymerization contributes to conducting “green polymer chemistry”.34,40,41,46,50−53,67−72 All the above-mentioned literature shows that enzymes are powerful catalysts for the production of macromolecules. The present article comprehensively reviews the macromolecular synthesis using enzymes as catalysts, which allows the precise synthesis with controlling the structure of product macromolecules with respect to substrate selectivity, functional group selectivity, regio-selectivity, stereoselectivity, and choro-selectivty.73 These selectivities are very efficient also for the modification of macromolecules via end-group or side-chain group functionalizations.67 Enzymatic reactions representing polymerizations and modifications normally show high catalytic efficiency without producing side products. They proceed under mild reaction conditions (e.g., in a water-containing medium, at around neutral pH = ∼ 7, under ordinary temperature and pressure). Enzymes are renewable natural products, and hence, many cases of enzyme-catalyzed reactions are in the context of green chemistry. To date, in vitro enzymatic polymerizations utilized mainly enzymes from oxidoreductases, transferases, and hydrolases as catalysts. This article also contains the macromolecular synthesis using enzyme model compounds as catalysts but shall not mention in vivo enzymatic macromolecular syntheses (i.e., macromolecular productions in a living system like fermentation synthesis, microbial, or bacterial synthesis) or E. coli using processes. It is to be noted here that the terms “macromolecule” and “polymer” are used without conceptual discrimination throughout the article. At the early stage of the research in the macromolecular science, the former denoted a molecule having giant molecular weight according to Staudinger exemplified by proteins and DNA, whereas the structure of the latter is expressed by (M)n, in which M represents a monomer as well as a constitutional repeating unit and n is the degree of polymerization.74

2378 2381 2382 2382 2383 2385 2386 2386 2386 2386 2387 2387

1. INTRODUCTION Polymerization reactions to produce macromolecules normally need a catalyst or initiator to induce the reaction to proceed.1 Historically, in the polymerization chemistry, anionic, cationic, and radical catalysts (or initiators) belong to a classical entity known since the early stage of polymer science (i.e., for around 90 years).2 Among others, innovative and notable works are chronologically cited as follows. (1) Concept of macromolecules was proposed and established by H. Staudinger (1920s).3−7 (2) Radical chain mechanism in organic chemistry was proposed (1930).8 (3) W. H. Carothers created nylon via polycondensation (1930s).9,10 (4) Cationic ring-opening polymerization of tetrahydrofuran was disclosed (1937).11,12 (5) Ionic chain mechanism was also proposed (1940).13,14 (6) Discovery of Ziegler−Natta catalysts (1953 and 1955).15−17 (7) Living polymerization method was discovered (1956).18 (8) Finding of conducting polymers (1977).19 (9) Development of olefin metathesis catalysts (since 1950s).20,21 In addition, from the viewpoint of macromolecular and/or polymer synthesis, a new method of solid-phase synthesis contributed much, in particular to the peptide and nucleic acid synthesis.22 Also, the concept of supramolecular system is to be noted, which has often been utilized for polymer synthesis.23,24 It is important to note that new catalysts and/or new methods are often brought about to create new compounds and/or new materials, typically exemplified by Ziegler−Natta catalysts, which actually made possible the production of various polymers from α-olefins, dienes, and others in petroleum-based chemical industry worldwide. It can be regarded, therefore, that the catalyst is a key for developing new polymeric materials. In fact, the above-mentioned innovative works led to plenty new polymeric materials, many of them being currently used in modern society. Human beings enjoyed a daily life with various commodity materials as well as highly functional, value-added materials including medical and pharmaceutical objects, since the later half of the twentieth century. Many products could be obtained readily and relatively cheaply. However, since around the late twentieth century, such situation was considered not to be allowed, obviously due to change of the earth environment as assessed by the IPCC (Intergovernmental Panel on Climate Change) several times. In the chemistry area, a new concept, “green chemistry”, pointed out the important issues.25,26 The issues include green factors of synthesis processes (starting materials, catalysts, solvents, reaction temperature and pressure, reaction selectivity, catalytic reaction or molar reaction, atom economy, etc.), toxicity of the starting materials and products,

2. ENZYMES AND ENZYMATIC CATALYSIS Enzymes are indispensable substances in living cells, which catalyze all the in vivo metabolic reactions to produce necessary biomacromolecules for maintaining the living system. Enzymatic catalytic function attracted much interest to scientists, and the first finding was noted as diastase (amylase) in 1833 by A. Payen and J. F. Persoz.75 Since then, research study on enzymes and enzymatic functions has been one of the most actively investigated topics in science fields due to its mysterious and profound nature. To date, so many investigations have been accumulated on enzymatic functions, and over several thousand kinds of enzymes are known.76 Enzymatic catalysis is utilized not only for macromolecular synthesis but also for macromolecular modifications by using 2308


Chemical Reviews

Review

Table 1. Classification of Enzymes, Typical Examples, Macromolecule Examples Synthesized by Enzymatic Polymerization as well as Modified by Enzymatic Reaction example enzymes

macromolecules modified

class

enzymes

macromolecules synthesized

1.

oxidoreductases

peroxidase, laccase, tyrosinase, glucose oxidase

polyphenols, polyanilines, polythiophene, vinyl polymers

2.

transferases

phosphorylase, glycosyltransferase, acyltransferase

polysaccharides, cyclic oligosaccharides, polyesters

3.

hydrolases

glycosidase (cellulase, amylase, xylanase, chitinase, hyaluronidase), lipase, protease, peptidase

polysaccharides, polyesters, polycarbonates, polyamides, poly(amino acid)s, polyphosphates, polythioesters

4. 5. 6.

lyases isomerases ligases

decarboxylase, aldolase, dehydratase racemase, epimerase, isomerase ligase, synthase, acyl CoA synthase

polysaccharides, polypeptides (proteins) polysaccharides, polypeptides (proteins) polysaccharides, polypeptides (proteins)

selectivity perfectly. It is indispensable for the reaction to take place, therefore, that in vitro reactions the artificial substrate is to be designed so that the substrate may be recognized by the enzyme. Second, L. Pauling suggested in 1946 the specific reason why enzymes cause the catalysis under mild reaction conditions like in living cells.78,79 An enzyme (E) and a substrate (S) form a complex (ES) through a key and lock interaction, which activates the substrate to lead to a transition state ([ES]‡) for the reaction to proceed, where the activation energy (ΔGenz‡) is greatly lowered by the stabilizing action by the enzyme, in comparison with that (ΔGno‡) of a reaction without enzyme via a transition state [S]‡ (Scheme 1, Figure 2).52

selective reaction characters to convert to new functionalized macromolecules. Enzymes are divided into six main classes (Table 1).76 Many of them are currently utilized for industrial productions in the areas of food, pharmacology, medicine, textile industries, etc. The table includes enzyme examples of the class, typical macromolecules synthesized by enzymatic polymerization as well as macromolecules modified by enzymatic reaction, which are described in the present article. As to the enzymatic catalysis, there are two fundamental important issues. First, E. Fischer proposed “key and lock” theory in 1894, which refers to the specific relationships between enzyme and substrate.77 The relationships are presently recognized as the molecular recognition: in an in vivo reaction (cycle A, Figure 1), a natural substrate is

Scheme 1

Figure 1. Schematic expression for “key and lock” theory for in vivo enzymatic reactions via biosynthetic pathway (cycle A) and for in vitro enzymatic reactions via nonbiosynthetic pathway (cycle B).

Figure 2. Energy diagram for a chemical reaction. Comparison between an enzyme-catalyzed reaction and a reaction without enzyme. Reprinted from ref 52. Copyright 2009 American Chemical Society.

specifically recognized by an enzyme to form the enzyme− substrate complex. The complex formation is caused by supramolecular interaction and activates the substrate to lead to the reaction product with perfect selectivity control. Namely, the complex formation is essential for the reaction to occur, where the substrate must be recognized like a key and lock relationship. In an in vitro reaction (cycle B, Figure 1), on the other hand, an artificial substrate must be recognized by the enzyme to form the enzyme-artificial substrate complex, and then the substrate is activated for the reaction to proceed, giving rise to the product with controlling the reaction

The enzymatic catalysis normally brings about the rate acceleration of 106−1012 fold; however, a specific case reached even 1020 fold.80 The mechanism of in vivo enzymatic reaction shown in Scheme 1 and Figure 1 is generally well accepted.81 Then, deep consideration on the mechanistic characterizations between enzymes and antibodies brought about a new concept of catalytic antibody; the fundamental difference between them is that the former selectively binds transition states and the latter binds ground states.82 2309


Chemical Reviews

Review

chemo-enzymatic process. An immobilized enzyme can be recovered and repeatedly used. Moreover, enzyme catalyzes a very complicated reaction, when the substrate monomer is adequately designed, as seen below in the polysaccharide synthesis; the products like cellulose, hyaluronic acid, chondroitin, etc. are the macromolecules having the most complicated structure ever synthesized in vitro. The above characteristics are to be noted in particular from the viewpoint that nature is our good teacher. Enzymecatalyzed reactions often provide ideas for biomimetic chemistry, and then, this approach will lead to the development of new efficient reactions. It is fortunate that nowadays various enzymes become widely available, and the reaction mechanism has increasingly been elucidated due to extensive studies on the area. Hence, the enzymatic method will be extended more readily in the future.

An in vitro enzymatic reaction was noted in organic chemistry field; lipase was utilized in the 1930s for ester synthesis in an organic solvent by Sym,51,83,84 but much attention was not paid at that time. Using enzymes as the catalyst for organic synthetic reactions actively started by Klibanov et al, in the 1980s (e.g., in the esterification or transesterification reactions).85−87 These results indicate that the above key and lock theory does not necessarily mean that the enzyme and substrate relationship requires an absolutely strict combination, but the relationship involves flexibility to some extent. This is the fundamental reason why enzymes are able to catalyze in vitro reactions. So far as the substrate is recognized by enzyme to form an ES complex, the enzyme-catalyzed reaction is realized in vitro. It is to be paid attention that supramolecular chemistry operates very importantly during the course of reaction, in particular, in the transition states. These arguments are the cases for all the enzyme-catalyzed synthesis of macromolecules.

3.3. Starting Raw Materials

As starting materials, many renewable biobased chemicals are employed in place of fossil-based raw materials. They include important platform materials such as lactic acid, itaconic acid, and anhydride, succinic acid and anhydride, sebacic acid, several fatty acids, 1,4-butanediol, sorbitol, glycerol, cardanol, tuliparin, etc., which shall be mentioned in this article. All of these chemicals are derived from biobased corn, wheat, sugar cane, cassava, switch grass, etc. via fermentation and/or chemical/ physical treatments. The renewable starting materials are related to the concept of “carbon neutral”, not to increase the carbon dioxide emission. In addition, environmentally benign reagents can be employed; water, oxygen from air, hydrogen peroxide, carbon dioxide, etc. are applicable.

3. ENZYMES AS GREEN CATALYST The concept of “green chemistry” became well-known by a book published in 1998 by Anastas and Warner, and they pointed out our future direction of chemistry research and chemical industry.25 The book showed 12 philosophical principles to chemists for mitigating the environmental problems yet maintaining the sustainable society. In the macromolecular synthesis area, “green polymer chemistry” was noted in 199935 and in this context the green character of enzyme-catalyzed macromolecular synthesis has often been argued.35,41,42,46,48,50−54,58−72 It is worth noting in the area that a new journal, Green Chemistry, launched in 1999 from The Royal Society of Chemistry. From the American Chemical Society, a new journal, ACS Sustainable Chemistry & Engineering launched in 2013, and also, a journal, Environmental Science & Technology Letters newly appeared in 2014. The launch of these journals strongly suggests the importance of this direction from the environmental and sustainable society viewpoints. Here, we address the green polymer chemistry in using enzymes as catalyst for macromolecular synthesis. They include advantageous green aspects concerning clean-processes, reaction selectivity, energy savings, natural resource problems, carbon dioxide emission, etc.

3.4. Reaction Products

Product macromolecules are nontoxic, and almost all are biodegradable which are benign to nature. Functionalized polyesters and polysaccharides often provide with value-added products, which are applicable to biomedical and pharmaceutical areas. 3.5. Reaction Solvents

In vivo enzymatic reactions usually take place in a water solvent system. However, in vitro enzymatic reactions are sometimes robust enough to be carried out not only in an organic solvent but also in a green solvent, like water, supercritical carbon dioxide, ionic liquids, or in other green solvents.

3.1. Clean Reaction Processes Involving High Selectivity

In vivo, enzyme-catalyzed reactions normally proceed under mild conditions; at a lower temterature, at around neutral pH, under ordinary pressure, etc. The rate of these reactions is very large in terms of the turnover number. All the enzymatic reactions proceed under the high reaction selectivity. In vitro, enzyme-catalyzed macromolecular synthesis reactions are also possible by the appropriate reaction design to proceed in a similar way; they are highly selective with many respects, in enantio-, regio-, chemo-, and choro-selectivities,73,88 and hence, they give a clean reaction system with producing no- or minimal byproducts.31−35,40,41,46,48−71 These advantages contribute to energy savings by all means and are normally hard to be achieved by conventional catalytic reactions.

3.6. Polymer Recycling and Degradation

The first case is concerned with the lipase catalysis. The ester group is relatively easy for bond-forming and bond-breaking, which is a reversible process. A new method of chemical recycling of polyesters using lipase catalysis was reported.89−91 The principle lies in that the ring-opening polymerization (ROP) system of lactones by lipase catalysis is reversible between linear polymers and cyclic oligomers, which can be controlled by changing the reaction conditions. A continuous flow method combining degradation-repolymerization must be a good way of chemical recycling.92 Polyester degradation through lipase catalyst is specific to the stereochemistry of polyester backbone as well as branches.93 Second, enzymatic degradation behaviors of polysaccharides like cellulose by cellulase94 and chitin by chtinase,95 as well as various nylon polymers and copolymers by nylon hydrodase96 are well studied by using high-speed atomic force microscopy94,95 and gas cluster secondaly ion mass spectrometry.96 These studies

3.2. Characteristics of Enzymes as Catalyst

Enzyme is a nontoxic, renewable, natural catalyst, which is free from a metal in most cases. In addition to the high reaction selectivity, enzymatic catalytic activity is very high even in vitro. In some cases, enzyme is robust enough to be used in combination with other chemical catalysts, allowing a new 2310


Chemical Reviews

Review

4.1.1. Polymerization of Aromatic Compounds. As typical aromatic polymers, phenol-formaldehyde resins using prepolymers such as novolaks and resols are widely used in industrial fields. These resins show excellent toughness and thermal-resistant properties, but the general concern over the toxicity of formaldehyde has resulted in limitations on their preparation and use. Therefore, an alternative process for the synthesis of phenolic polymers avoiding the use of formaldehyde is strongly desired. 4.1.1.1. Polymerization of Phenol Compounds. HRPcatalyzed polymerization of unsubstituted phenol is first mentioned as the most thoroughly studied monomer. Unsubstituted and Substituted Phenols. Oxidative polymerization of phenol, the simplest and most important phenolic compound in industrial fields, proceeds in the presence of conventional polymerization catalysts; however, the product polymer was often insoluble in water and common organic solvents, and the structure was uncontrollable.103 Phenol was subjected to the oxidative polymerization using HRP or SBP as catalyst in a mixture of 1,4-dioxane and buffer, yielding a polymer consisting of phenylene and oxyphenylene units (Scheme 3). The polymer showed low solubility; it was partly

involve possible application of enzymatic biomass degradation to biofuel development, fabrication of fibers, and recycling.

4. OXIDOREDUCTASES Various oxidoreductases play an essential role in maintaining the metabolism in living cells. All enzymes of this class catalyze oxido-reduction reactions. The substrate that is oxidized is regarded as a hydrogen donor. The systematic name is based on donor:acceptor oxidoreductase. The recommended name will be dehydrogenase, wherever this is possible; as an alternative, reductase can be used. Oxidase is only used in cases where O2 is the acceptor.76 So far, several oxidoreductases, such as peroxidase, laccase, tyrosinase, bilirubin oxidase, have been reported to catalyze in vitro oxidative polymerization of aromatic compounds like phenol derivatives, and among them, peroxidase is most often used. 4.1. Peroxidases

Peroxidases include typically horseradish peroxidase (HRP) and soybean peroxidase (SBP) (both EC 1.11.1.7) having Fe at the active center. HRP is an enzyme whose catalysis is an oxidation of a donor to an oxidized donor by the action of hydrogen peroxide, liberating two water molecules. HRP is a single-chain β-type hemoprotein that catalyzes the decomposition of hydrogen peroxide at the expense of aromatic proton donors.97 HRP is active as a catalyst for the oxidative polymerization of phenolic compounds, whose catalytic cycle is shown in Scheme 2. The reaction proceeds via oxidative

Scheme 3

Scheme 2 soluble in DMF and DMSO and insoluble in other common organic solvents. On the other hand, aqueous methanol afforded the DMF-soluble polymer with molecular weight of 2100−6000 in good yields.100,104−108 The solubility of the resulting polymer strongly depended on the buffer pH and content of the mixed solvent. The resulting phenolic polymer showed relatively high thermal stability, and no clear glass transition temperature (Tg) was observed below 300 °C. The control of the polymer structure was achieved by solvent engineering. The ratio of phenylene and oxyphenylene units was strongly dependent on the solvent composition. In the HRP-catalyzed polymerization of phenol in a mixture of methanol and buffer, the oxyphenylene unit increased by increasing the methanol content, while the buffer pH scarcely influenced the polymer structure. The polymer solubility increased with increasing the oxyphenylene unit content.107 Molecular weight control of the polymer was achieved by the copolymerization with 2,4-dimethylphenol to give a soluble oligomer with a molecular weight of 500.109 The proposed polymerization mechanism is shown in Scheme 4. A phenoxy free radical is first formed, and then two molecules of the radical species dimerize via coupling. Since peroxidase often does not recognize larger molecules, a radical transfer reaction between a monomeric phenoxy radical and a phenolic polymer takes place to give the polymeric radical species. In the propagation step, such propagating radicals are subjected to oxidative coupling, producing polymers of higher molecular weight. The enzymatic polymerization of phenols in aqueous solutions often resulted in the low yield of the insoluble polymer. The peroxidase-catalyzed polymerization of phenol took place in the presence of 2,6-di-O-methyl-α-cyclodextrin (DM-α-CD) in a buffer. Only a catalytic amount of DM-α-CD

coupling between radical species.98−102 Peroxidase-catalyzed oxidative coupling of phenols proceeds fast in aqueous solutions, giving rise to the formation of oligomeric compounds. However, the resulting oligomers have not been well characterized, since most of them show low solubility toward common organic solvents and water. 2311


Chemical Reviews

Review

produced by the HRP-catalyzed dispersion polymerization of phenol in a mixture of 1,4-dioxane and phosphate buffer (3:2 v/ v) using poly(vinyl methyl ether) as a stabilizer.114,115 The particle size could be controlled by the stabilizer concentration and solvent composition. Thermal treatment of these particles afforded uniform carbon particles. The polymer microsphere was also prepared by HRP-catalyzed polymerization of phenol in the presence of PEG in an aqueous 1,4-dioxane, and palladium was loaded on the microsphere surface. This palladium complex showed good catalytic activity for Heck reactions of acrylic acid with aryl iodides.116 When sodium dodecyl sulfate was used for the enzymatic polymerization of phenol in a phosphate buffer, the resulting polymer was partly soluble in a common organic solvent.117 The modification of the product by epichlorohydrin, and the subsequent functionalization by triethylene tetramine and immobilization of palladium afforded the catalyst for the Heck reaction. A bienzymatic system was developed as a catalyst for the oxidative polymerization of phenol.118 The HRP-catalyzed polymerization of phenol in the presence of glucose oxidase, and glucose gave the polymer in a moderate yield, in which hydrogen peroxide was formed in situ by the oxidative reaction of glucose catalyzed by glucose oxidase. In this system, no successive addition of hydrogen peroxide was involved. A combination of the bienzymatic method and a reversible addition−fragmentation chain transfer polymerization of hydrophilic vinyl monomers provided the living polymerization system to produce the polymers with narrow molecular weight distribution and well-defined block copolymers.119 HRP-catalyzed polymerization of phenol was conducted in the presence of TEMPO-oxidized nanocellulose to produce the nanocomposites of the phenol polymer and cellulose.120 The phenol polymer in the nanocomposites existed as spherical or near-spherical clusters in the size of ca. 0.1 μm. FT-IR analysis showed the physical and chemical interaction between both components. Carbon nanotubes (CNTs) were used as a template for the HRP-catalyzed polymerization of phenol in water to functionalize the CNT surface. The polymerization was conducted in the presence of the p-hydroquinone (HQ)linked CNTs, in which phenolic polymer was grafted through the HQ moiety on the CNT surface. The phenol monomer was regioselectively polymerized to give mainly the thermally stable oxyphenylene unit.121 Phenol polymer-graf t-PEG was synthesized by the enzymatic polymerization of phenol, followed by anionic polymerization of ethylene oxide.122 The obtained graft copolymer was soluble in water, ethanol, DMF, THF, and methylene dichloride. The polymerization behaviors and properties of the phenolic polymers depended on the monomer structure, solvent composition, and enzyme origin. In the HRP-catalyzed polymerization of p-n-alkylphenols in aqueous 1,4-dioxane, the polymer yield increased as the chain length of the alkyl group increased from 1 to 5.123,124 HRP catalyzed the oxidative polymerization of all cresol isomers, whereas among o-, m-, and p-isopropylphenol isomers, only p-isopropylphenol polymerized by HRP catalysis.125 Poly(p-n-alkylphenol)s prepared in aqueous 1,4-dioxane showed low solubility toward organic solvents. On the other hand, a soluble oligomer with molecular weight lower than 1000 was formed from p-ethylphenol using aqueous DMF.126 Poly(p-t-butylphenol) enzymatically synthesized in aqueous 1,4-dioxane showed Tg and melting point (Tm) at 182 and 244 °C, respectively. The product structure from p-cresol and p-propylphenol was studied in detail by using

Scheme 4

was necessary to induce the polymerization efficiently.110 Poly(ethylene glycol) (PEG) was found to act as a template for an oxidative polymerization of phenol in water. The presence of the PEG template in an aqueous medium greatly improved the regioselectivity of the polymerization, yielding a phenol polymer with the phenylene unit content higher than 90% (Scheme 5).111,112 During the reaction, the polymer was Scheme 5

produced in high yields as precipitates in complexing with PEG. The molecular weight of PEG strongly affected the polymer yield. The unit molar ratio of the phenolic polymer and PEG was ca. 1:1. The FT-IR and DSC analyses exhibited the formation of the miscible complex of the phenolic polymer and PEG by a hydrogen-bonding interaction. PEG monododecyl ether, a commercially available nonionic surfactant, was also a good template for the polymerization of phenol in water. By using PEG-poly(propylene glycol) (PPG)-PEG triblock copolymer (Pluronic) with the high PEG content as the template, the phenolic polymer with ultrahigh molecular weight (Mw > 106) was formed.113 The regioselectivity was also high (the phenylene unit content of 86%). This polymerization method did not involve use of organic solvents; hence it is regarded as being an environmentally benign system. Morphology of the enzymatically synthesized phenolic polymers was controlled under the selected reaction conditions. Monodisperse polymer particles in the submicron range were 2312


Chemical Reviews

Review

NMR. A coupling mechanism of p-cresol was discussed from the structure of the dimers produced at the initial stage of polymerization.127,128 The structure and biodegradable properties of the polymers from various p-substituted phenols were examined.129−131 The polymer particles of submicron size from m-cresol, p-cresol, and p-phenylphenol were obtained in the dispersion system as described above.114,115 A natural phenol glucoside, 4-hydroxyphenyl β-D-glucopyranoside (arbutin), was subjected to regioselective oxidative polymerization using a peroxidase catalyst in a buffer solution, yielding the water-soluble polymer consisting of 2,6-phenylene units, in turn converted to poly(hydroquinone) by acidic deglycosylation (Scheme 6).132 Another route for the chemoenzymatic synthesis of poly(hydroquinone) was the SBPcatalyzed polymerization of 4-hydroxyphenyl benzoate, followed by alkaline hydrolysis.133

Scheme 7

Scheme 6

polymerization behaviors and the thermal properties of the resulting polymers. Coprinus cinereus peroxidase (CiP), a fungal peroxidase, was used as catalyst for the polymerization of bisphenol A in aqueous 2-propanol solution.140 The yield, molecular weight, and structure of the polymer depended on the solvent composition. The polymer was mixed with a diazonaphthoquinoe derivative to form a film and applied to a photoresist on the silicon wafer by UV irradiation. Sharply contrasted patterns were obtained from the polymer with a molecular weight of 3 × 103. α-Hydroxy-ω-hydroxyoligo(1,4-oxyphenylene)s were formed in the HRP-catalyzed oxidative polymerization of 4,4′-oxybisphenol in an aqueous methanol.141 During the reaction, hydroquinone was formed. Scheme 8 shows the postulated mechanism of the trimer formation; the redistribution and/or rearrangement of the quinone-ketal intermediate takes place, involving the elimination of hydroquinone. Oxidative polymerization of phenols was conducted not only in the monophasic solvents but also in interfacial solvents such as micelles, reverse micelles, and Langmuir trough systems.142 p-Phenylphenol was polymerized in an aqueous surfactant solution, yielding the polymer with relatively narrow molecular weight.143 For the HRP-catalyzed polymerization of phenol, sodium dodecylbenzenesulfonate was used to produce the polymer in high yields over a wide pH range from 4 to 10.144 The polymer showed good solubility toward organic solvents such as DMF, DMSO, acetone, and THF.145 Ionic liquids are effective as cosolvent for the enzymatic oxidative polymerization of phenols. In a mixture of a phosphate buffer and 1-butyl-3-methylimidazolium tetrafluoroborate or 1-butyl-3-methylpyridinium tetrafluoroborate, pcresol, 1-naphthol, 2-naphthol, and p-phenylphenol were polymerized by SBP to give the polymers in good yields.146,147 For p-phenylphenol, the polymerization proceeded regioselectively to exclusively produce 2,2′-bi-(4-phenylphenol). HRP-catalyzed oxidative polymerization of 4-hexyloxyphenol was conducted in an isooctane solvent. HRP was modified by

Chemoenzymatic synthesis of a new class of poly(amino acid), poly(tyrosine) containing no peptide bonds, was achieved by the peroxidase-catalyzed oxidative polymerization of tyrosine ethyl esters, followed by alkaline hydrolysis. The resulting poly(amino acid) is different from the peptide-type poly(tyrosine) and soluble only in water. The oxidative homopolymerization of N-acetyltyrosine and the copolymerization of N-acetyltyrosine with 4-hydroxyphenyl β-D-glucopyranoside (arbutin) catalyzed by HRP were reported.134 HRP catalysis induced a chemoselective polymerization of a phenol derivative having a methacryloyl group. Only the phenol moiety was polymerized to give a polymer having the methacryloyl group in the side chain. The resulting polymer was readily cured thermally and photochemically (Scheme 7).135 Phenolic monomers containing vinyl groups like 4′hydroxy-N-methacryloyl anilide, N-methacryloyl-11-aminoundecanoyl-4-hydroxy anilide, and 4-hydroxyphenyl-N-maleimide were also chemoselectively polymerized at the phenol moiety by HRP catalyst. The polymerization proceeded in a buffer in the presence of cyclodextrin. The resulting phenolic polymers having the vinyl group were copolymerized with methyl methacrylate or styrene.136,137 The polymers were subsequently cross-linked by a radical initiator.138 A phenolic polymer soluble in acetone, DMF, DMSO, and methanol was formed from bisphenol A via peroxidase catalysis.139 The polymer was produced in higher yields using SBP as a catalyst and showed Tg at 154 °C. Peroxidase also induced the polymerization of an industrial product, bisphenolF, consisting of 2,2′-, 2,4′-, and 4,4′-dihydroxydiphenylmethanes. Under the selected reaction conditions, the quantitative formation of a soluble phenolic polymer was achieved. Among the isomers, 2,4′- and 4,4′-dihydroxydiphenylmethanes were polymerized to give the corresponding polymers in high yields, whereas no polymerization of the 2,2′isomer occurred. In the case of 4,4′-dihydroxyphenyl monomers, the bridge structure enormously affected the 2313


Chemical Reviews

Review

Scheme 8

produced the corresponding copolymer, which was further modified for preparation of inclusion complexes with βcyclodextrin.153 Schiff bases containing a phenol group, 4-(benzylideneamino)-phenol and 4-[(anthracen-9-ylmethylene)-amino]-phenol, were polymerized by HRP in a mixture of 1,4-dioxane and phosphate buffer.154 Large red shift was observed for the polymer from the latter, suggesting the increase of the conjugation length. HRP also catalyzed the oxidative polymerization of 4-[(4-phenylazo-phenyimino)-methyl]phenol, which possesses a Schiff base and azo groups, yielding the polymer with the large potential for electronic and optic active materials.155 A phenolic polymer from 4-(2-aminoethyl)phenol (tyramine) was prepared in capsules.156 HRP-loaded capsules were first prepared via layer-by-layer assembly of polyelectrolites, poly(sodium 4-styrenesulfonate), and poly(allylamine hydrochloride). The selective permeability of the capsule wall allowed the monomer to penetrate, and the enzymatic oxidation polymerization of tyramine took place, while the product polymer and HRP remained in the capsule interior.

ion-pairing with an anionic surfactant (aerosol AT, AOT), and t-butyl hydroperoxide was used as an oxidant instead of hydrogen peroxide. The product polymer had the C−C coupled structure. The turnover number was about 5 × 103.148 p-Methoxyphenol was polymerized by HRP catalyst in an aqueous micelle system in the presence of sodium dodecyl sulfate. The resulting polymer showed good antioxidant properties.149 HRP-catalyzed polymerization of 4-hydroxybenzene-diazosulfonate monomer produced the water-soluble polymer.150 4-Aminophenol was polymerized by HRP in water, and the resulting polymer showed good adsorption capacity for silver ions, which were reduced to form silver nanoparticles with face-centered cubic structure.151 An urushiol analogue, an unsaturated higher alkyl groupcontaining phenolic monomer, was subjected to an oxidative polymerization by HRP or Fe-salen (a model complex of peroxidase) to produce a soluble prepolymer, which was further cured thermally or by a cobalt catalyst to yield the cross-linked film (artificial urushi, see also section 4.2.2) with a high gloss surface.152 HRP-catalyzed copolymerization of 4-tert-butylphenol and 4-ferrocenylphenol in an aqueous 1,4-dioxane 2314


Chemical Reviews

Review

linkable polymer was synthesized by the SBP-catalyzed polymerization of cardanol (Scheme 10). Fe-salen, a model

This biocatalytic method provided a new way to synthesize functional materials in the microcapsules and to modify permeation properties of the microcapsule wall. Enzymatically synthesized poly(tyramine) selectively adsorbed platinum and palladium in HCl solution.157 HRP also catalyzed the polymerization of 4-hydroxyphenylacetic acid (HPA) on the polyelectrolyte capsule wall, on which HRP was assembled by a layer-by-layer technique as both a catalyst and a coupling template.158 The phenolic polymer formed the layers of 70− 200 nm thickness and modified the permeability properties of the capsule wall. The efficient phenolic polymer production was achieved by the peroxidase-catalyzed polymerization of m-alkyl-substituted phenols in aqueous methanol. The mixed ratio of methanol and buffer greatly affected the yields and the molecular weight of the polymer. The enzyme source greatly affected the polymerization pattern of m-substituted monomers. Using SBP catalyst, the polymer yield increased as a function of the bulkiness of the substituent, whereas the opposite tendency was observed when HRP was the catalyst. These enzymatically synthesized phenolic polymers were applied to positive-type photoresists for printed wire boards because of their high solubility toward alkaline solution and high thermal stability.159,160 Various msubstituted phenols were oxidatively polymerized in an aqueous buffer in the presence of an equimolar amount of cyclodextrins to give the product polymers in high yields.161 CiP-catalyzed copolymerization of m-cresol and 2,2′-methylenebis[6-(2hydroxy-5-methylbenzyl)-p-cresol was examined in an aqueous acetone.162 The photoresist was prepared by mixing the copolymer with a diazonaphthoquinone derivative. The copolymer with high hydroxyl value showed remarkably improved dissolution characteristics. A phenol with an acetylenic substituent in the meta position was also chemoselectively polymerized by HRP to give a polymer bearing acetylenic groups (Scheme 9).163 For

Scheme 10

complex of HRP (see Scheme 13), efficiently catalyzed the polymerization of cardanol in organic solvents.164−168 The polymerization proceeded in 1,4-dioxane to give the soluble polymer with molecular weight of several thousands in good yields. The curing of the polymer took place in the presence of cobalt naphthenate catalyst at room temperature or thermal treatment (150 °C for 30 min) to form yellowish transparent films (“artificial urushi” in a broad sense, see also section 4.2.2), whose properties are similar to those of a traditional Japanese lacquer.169 The resulting cross-linked film exhibited good elastic properties comparable with natural urushi. FT-IR monitoring of the curing showed that the cross-linking mechanism is similar to that of the oil autoxidation. When HRP was used as a catalyst for the polymerization of cardanol, the reaction took place in the presence of a redox mediator (phenothiazine derivative) to give the polymer.170 CiP was also effective as a catalyst for the polymerization of cardanol.171 An equivolume mixture of tert-butanol and phosphate buffer afforded the highest yield.172 The curing of the polymer by using methyl ethyl ketone peroxide as an initiator and cobalt naphthenate as an accelerator was examined.173 Anacardic acid is another component of CNSL. SBP polymerized anacardic acid in an aqueous 2-propanol in the presence of phenothiazine-10-propionic acid (mediator).174 The polymer was cross-linked on a solid surface to form the coating film, which showed a good antibiofouling effect for Gram-positive and Gram-negative bacteria. Naphthol derivatives containing unsaturated fatty acid moieties were oxidatively polymerized by Fe-salen, and the curing of the obtained polymers afforded the hard films.152,175 HRP catalyzed the oxidative polymerization of 2-hydroxycarbazole in a mixture of 1,4-dioxane and phosphate buffer.176 The optical and electrochemical band gaps of the polymer were dramatically lower than those of 2-hydroxy-carbazole. The polymer showed good conductivity by doping with iodine vapor. ortho-Methoxyphenols (apocynin, vanillin, and 4-methylguaiacol) were polymerized by SBP catalyst in an aqueous buffer.177 A variety of oligophenols (dimers to pentamers) as well as some of their oxidation products including quinones and demethylated quinones were formed. The oxidation of guaiacol is often used as a tool of the enzyme assay. The structure of the product obtained by HRP-catalyzed polymerization of guaiacol was examined; trimers in addition to the hitherto known dimeric products were isolated and characterized by NMR.178 Micelle-templated polymeric nanowires were produced by HRP-catalyzed polymerization of guaiacol in a solution containing an ionic surfactant.179 The TEM image of the product showed the formation of the nanowire.

Scheme 9

comparison, the reaction of the monomer using a copper/ amine catalyst, a conventional catalyst for an oxidative coupling, was performed, producing a diacetylene derivative exclusively. The resulting polymer was converted to a carbon polymer in much higher yields than enzymatically synthesized poly(mcresol), suggesting a large potential as precursor of functional carbon materials. Cardanol, a main component obtained by thermal treatment of cashew nut shell liquid (CNSL), is a phenol derivative having mainly the meta substituent (R) of a C15 unsaturated hydrocarbon chain with one to three double bonds as the major. Since CNSL is nearly one-third of the total nut weight, a large amount of CNSL is obtained as byproducts from mechanical processes for the edible use of the cashew kernel. Only a small part of cardanol obtained in the production of cashew kernel is used in industrial fields, though it has various potential industrial utilizations such as resins, friction-lining materials, and surface coatings. Therefore, development of new applications for cardanol is very attractive.35,40,41 A new cross2315


Chemical Reviews

Review

coupling mechanism.188 CiP produced the copolymer in higher yield than HRP or SBP. Modification of sulfomethylated alkali lignin catalyzed by HRP was examined.189 By this enzymatic treatment, the carboxyl group content increased, while the contents of the phenolic group and methoxy group decreased. Peroxidase-catalyzed oxidative polymerization of 2-naphthol in a reverse micellar system gave the polymer in single and interconnected microspheres.190 The polymer showed the fluorescence characteristics of the 2-naphthol chromophore. The polymerization of 8-hydroxyquinoline-5-sulfonate was studied by in situ NMR spectroscopy, and the polymerization mechanism was discussed in detail.191 1,5-Dihydronaphtahalene (1,5-DHN) was polymerized by HRP to produce poly(1,5DHN) consisting of the 1,5-dihydroxynaphthalene unit, 1(or 5)-hydroxy-5(or 1)-oxynaphthalene unit, and 1,5-dioxynaphthalene unit.192 HRP-catalyzed polymerization of 2,6-DHN proceeded regioselectively at the aromatic ring to give the polymer exclusively with the 2,6-dihydroxynaphthalene unit. Reaction of poly(1,5-DHN) with Al(Et)(2-methyl-8-quinolinolato)2 afforded the Al−O bond formation, yielding a photoluminescent-aluminum complex. Fe-salen, a model complex of peroxidases, showing high catalytic activity for an oxidative polymerization of p-tbutylphenol and bisphenol A to produce soluble phenolic polymers,193 was also effective for the polymerization of 2,6dimethyl- and 2,6-difluorophenols. The PPO derivatives were formed, and the polymer from 2,6-difluorophenol showed crystallinity with a melting point higher than 250 °C.194,195 Fesalen catalyzed an oxidative cross-coupling of phenolic polymers onto a phenol-containing cellulose at room temperature under air to produce cellulose-graf t-phenolic polymers, a cellulose-phenolic polymer hybrid.196 Polyphenol Compounds. The compounds with more than two hydroxyl groups on the aromatic ring(s) are also subjected to oxidative couplings. During the reaction, an unstable oquinone intermediate is formed, which is converted to poly(catechol).197 Nanoscale polymer patterning was reported to be fabricated by the enzymatic oxidative polymerization of caffeic acid on a 4-aminothiolphenol-functionalized gold surface with dip-pen nanolithography technique.198 Bioactive polyphenols are present in a variety of plants and used as important components of human and animal diets. Flavonoids are a broad class of low molecular weight secondary plant polyphenolics, which are benzo-γ-pyrone derivatives consisting of phenolic and pyrane rings. Their biological and pharmacological effects including antioxidant, antimutagenic, anticarcinogenic, antiviral, and anti-inflammatory properties have been demonstrated in numerous human, animal, and in vitro studies.199,200 Enzyme-catalyzed oxidative polymerization of flavonoid compounds and evaluation of the biorelated properties of the products have been studied.101,201 Major components of polyphenols in green tea are flavanols, commonly known as catechins; the major catechins in green tea are (+)-catechin, (−)-epicatechin (EC), (−)-epigallocatechin (EGC), (−)-epicatechin gallate (ECG), and (−)-epigallocatechin gallate (EGCG) (Scheme 12). Numerous biological activities have been reported for green tea and its contents, among them, the preventive effects against cancer are most notable. Peroxidase catalyzed the oxidative polymerization of catechin to produce the oligomer with a degree of polymerization less than 5.202 Kinetic study on the oxidation of catechin by peroxidase from strawberries showed highly efficient catalytic activity of the enzyme at low concentration of

HRP-catalyzed oxidative polymerization of ortho-, meta-, and para-bromophenols were examined in water.180 During the polymerization, the precipitate including HRP was formed, which was due to the strong interaction between HRP and the product polymer. For the meta isomer, the lowest precipitation yield was observed. The addition of PEG suppressed the precipitation, leading to the enhancement of the oxidation, whereas the highest inactivation of HRP took place in the copolymerization. In the SBP-catalyzed polymerization of phenol, the enzyme activity in the precipitate remained.181 By the addition of a surfactant (Triton X-100), the adsorption of SBP was reversible. Poly(2,6-dimethyl-1,4-oxyphenylene) (poly(phenylene oxide), PPO) is a material widely used as high-performance engineering plastics, thanks to its excellent chemical and physical properties [e.g., a high Tg (ca. 210 °C)], and mechanically tough property103 PPO was first prepared from 2,6-dimethylphenol monomer using a copper/amine catalyst system. 2,6-Dimethylphenol was polymerized via HRP catalysis to give a polymer exclusively consisting of a 1,4-oxyphenylene unit,182 while small amount of Mannich-base and 3,5,3′5′tetramethyl-4,4′-diphenoquinone units are always contained in the chemically prepared PPO. HRP was immobilized on silica nanorods and used as a catalyst for polymerization of 2,6dimethylphenol.183 Substantially enhanced enzymatic activity and reusability were found in comparison with those with the free enzyme. Peroxidases (HRP and SBP) induced a new type of oxidative polymerization of the 4-hydroxybenzoic acid derivatives, 3,5dimethoxy-4-hydroxybenzoic acid (syringic acid) and 3,5dimethyl-4-hydroxybenzoic acid.184 The polymerization involved elimination of carbon dioxide and hydrogen from the monomer to give PPO derivatives with high molecular weight up to 1.5 × 104 (Scheme 11). Demethylation of the polymer from syringic acid by boron tribromide catalyst gave poly(2,6dihydroxy-1,4-oxyphenylene), which was thermally stable below 300 °C under nitrogen.185 Scheme 11

Coniferyl alcohol (4-hydroxy-3-methoxycinnamyl alcohol, CoA) is a phenolic lignin monomer (monolignol) contained in plant cell walls, whose dehydrogenated polymers (DHP = coniferyl alcohol polymers) are regarded as synthetic lignin, a model of cell walls. CoA was polymerized by HRP in a pectin solution in order to mimic the lignification that is the final step of biosynthesis of plant cell walls. The polymerization behaviors and product structures with cluster formation were examined in detail by various physical methods.186 CoA was also polymerized by HRP in the presence of α-cyclodextrin (αCD) in a phosphate buffer. The presence of α-CD led to DHP with 8-O-4′-richer linkages, compared with that prepared without any additives. This is probably because the inclusion complex formation between CoA and α-CD suppresses the formation of other linkages like 8−5′ and 8−8′ ones due to the steric hindrance of the complex.187 The peroxidase-catalyzed copolymerization of coniferyl alcohol and sinapyl alcohol in different ratios was conducted to examine the monolignol 2316


Chemical Reviews

Review

or electrochemical oxidation polymerization of aniline monomer. However, enzymatic polymerization provides an alternative method of a “green process”, which is usually carried out at room temperature, in aqueous organic solvents around neutral pH. Polyaniline is a typical example for the enzymatic polymerization using oxidoreductase enzymes.210 Bilirubin oxidase (EC 1.3.3.5, a copper-containing oxidoreductase)211 and horseradish peroxidase (an iron-containing oxidoreductase) together with using H2O2 oxidant212 were used. In the latter case, sulfonated polystyrene (SPS) acted as a polyanionic template, and the resulting polymer was complexed to the SPS.212 Lignosulfonate was also used as a natural polyelectrolyte.213 The reduction/ oxidation reversibility of the polyaniline/SPS complex was demonstrated. The conductivity of the polyaniline/SPS complex was measured to be 0.005 S/cm. The value increased to 0.15 S/cm with HCl doping. The enzymatic approach is claimed to offer unsurpassed ease of synthesis, processability, stability (electrical and chemical), and environmental compatibility.214 Instead of SPS, vesicles were employed as a soft material template.215 Micellar laccase,216 glucose oxidase,217 and a poly(ethylene glycol) modified hematin213,218 were used to catalyze the polymerization of aniline. Ionic liquid was used to immobilize, and the polymerization process took place at the ionic liquid/aqueous interface, where aniline monomer, H2O2 oxidant, and dodecylbenzenesulfonic acid were in the aqueous phase.219 The conductivity of polyaniline is related with the backbone structure of polyaniline.220 The branched structure lowered the conductivity,221 while a linear structure with the help of a template like SPS led to the conducting materials.222 Horseradish peroxidase-catalyzed oxidative polymerization of aniline was carried out in the presence of a template, poly(acrylic acid) (PAA), and of a chiral compound, 10camphorsulfonic acid (CSA) to generate a helical conformation.223 Chiral polyaniline was also prepared by micellar peroxidase-catalyzed synthesis in the presence of dodecylbenzenesulfonic acid.224 Poly(aniline-co-3-aminobenzeneboronic acid) was prepared via horseradish peroxidase-catalyzed oxidative copolymerization in the presence of SPS at pH 4.5 and was used as a boronic acid based sensor for saccharide molecules.225 Polyaniline colloid particles were prepared by enzymatic polymerization of aniline and were shown to be applicable for smart devices such as thermochromic windows, temperature-responsive electrorheological fluids, actuators, and colloids for separation technique.226 The enzyme-based imprinting approach using metal ions as the target analyte was tried. The horseradish peroxidasecatalyzed polymerization of various aromatic compounds227 was performed in the presence of metal ions Cu(II), Ni(II), and Fe(III) as imprinting templates.228 A new class of polyaromatics was synthesized by peroxidase-catalyzed oxidative copolymerization of phenol with o-phenylenediamine.105 Laccase (EC 1.10.3.2, a high redox potential)-mediated system based on potassium octocyanomalybdate (4+) (redox mediator) was first used for acceleration of the enzymatic aniline polymerization.229 The enzymatic reaction yielded oxidized octocyanomalybdate (5+) which can oxidize the aniline monomer to the aniline radical cation. This resulted in the formation of conducting polyaniline with the concomitant regeneration of the redox mediator. The presence of the mediator accelerated the polymerization of the monomer.

Scheme 12

hydrogen peroxide.203,204 The reaction products obtained by the HRP catalyst were analyzed by reversed-phase and sizeexclusion chromatographies.203,204 The products obtained by using HRP and polyphenol oxidase as catalysts were comparably studied. Both enzymes produced similar products.205 The formation of two unusual dimers were detected in the product produced by HRP; one was the dicarboxylic acid compound with C−C linkage between C-6′ of B-ring and C-8″ of D-ring, which was formed by an ortho cleavage of the E-ring, and another was the dimer of C−C linkage between C-2 of the C ring and C-8″ of the D ring. Similar to most of phenol and monosubstituted phenols, enzymatic polymerizations of natural polyphenols in a mixed solvent of polar solvent and buffer have been investigated. The HRP-catalyzed polymerization of catechin was in an equivolume mixture of 1,4-dioxane and buffer (pH 7); the polymer had a molecular weight of 3.0 × 103.206 Using methanol as a cosolvent improved the polymer yield and molecular weight. A water-soluble oligomer was formed by an HRP-catalyzed polymerization of catechin using a polyelectrolyte like a sulfonated polystyrene as a template and a surfactant like sodium dodecylbenzenesulfonate.207 Superoxide anion scavenging activity of the enzymatically synthesized poly(catechin) was evaluated. Poly(catechin), synthesized by a HRP catalyst, greatly scavenged superoxide anion in a concentration-dependent manner and almost completely scavenged at 200 μM of a catechin unit concentration.208 It was known that catechin showed a prooxidant property in concentrations lower than 300 μM. These results demonstrated that the enzymatically synthesized poly(catechin) possessed much higher potential for superoxide anion scavenging, compared with intact catechin. Furthermore, the enzymatically synthesized poly(catechin) also showed much greater inhibition activity against human low-density lipoprotein (LDL) oxidation in a concentration-dependent manner, compared to the catechin monomer. The chelating ability of polymers enzymatically synthesized from phenol, catechol, and pyrogallol.209 The binding capacity of the polymer from phenol for copper ion was larger than that from catechol or pyrogallol. Gold ion was selectively reduced by the phenol group in poly(pyrogallol) in acid media to form gold particles. 4.1.1.2. Polymerization of Other Aromatic Compounds. In addition to phenols, other aromatic compounds such as aniline, thiophene, and pyrrole compounds are available for oxidative polymerization. Polyaniline can be prepared by either chemical 2317


Chemical Reviews

Review

catalytic activity (k(cat)) was higher with chloroperoxidase than with horseradish peroxidase, but due to a higher affinity constant (KM ) the catalytic efficiency (k(cat)/K M ) of chloroperoxidase was lower than for horseradish peroxidase. A novel H2O2 biosensor based on horseradish peroxidase induced deposition of polyaniline on the designed graphenecarbon nanotube-nafion/gold−platinum alloy nanoparticles (GE-CNT-Nafion/AuPt NPs) modified glassy carbon electrode was constructed.237 The enzymatically induced deposition of polyaniline provides a general platform for the design of novel electrochemical biosensors. The enzymatic method realized highly efficient catalyzed deposition of polyaniline on the designed electrode, which improved the sensitivity and detection limit for H2O2 determination. Glucose oxidase (EC 1.1.3.4) from Penicillium vitale was immobilized on the carbon rod electrode by cross linking it with glutaraldehyde (glucose oxidase-electrode).238 Catalytic activity of immobilized glucose oxidase was used for polymerization of aniline by taking a high concentration of hydrogen peroxide produced during the catalytic action of immobilized glucose oxidase and locally lowered pH due to the formation of gluconic acid. The glucose oxidase layer, which was selfencapsulated within formed polyaniline matrix (glucose oxidase/polyaniline-electrode), showed an increase in the upper detection limit, optimal pH region for operation, and stability of glucose oxidase based electrode modified by polyaniline, when compared with an unmodified glucose oxidase electrode. Glucose oxidase was also implemented in the ELISA assay. The ELISA plate was subjected to the exposure with the primary antibody and subsequently with a biotinylated secondary antibody. Then, a glucose oxidase-avidin conjugate was connected to the secondary antibody via the biotin−avidin association. The adsorbed glucose oxidase finally catalyzed polymerization of rhodamine B-methacrylate and hydroxyethyl acrylate, etc. to yield a fluorescent polymer.239 Horseradish peroxidase enzyme was used as a catalyst for the polymerization process leading to a water-soluble PEDOT.240 Essential reaction conditions for the polymerization were performing the reaction at pH 2 at 4 °C. A similar horseradish peroxidase-catalyzed polymerization of EDOT gave a twophase reaction system. The enzyme and EDOT form droplets in the solution to trigger the polymerization in the interface, giving rise to water-soluble conducting polyEDOT. Toward the end of the polymerization, the polymer aqueous phase and the horseradish peroxidase/EDOT phase were separated and the latter phase acted as a biocatalyst, which can be recycled and reused.241 Enzymatically induced formation of the polythiophene layer over glucose oxidase modified electrode was constructed.242 KM(app.) of glucose oxidase/polythiophene-modified electrode increased by prolongation of the polymerization duration. The enzymatic thiophene polymerization could be applied in tuning the KM(app.) and other kinetic parameters in glucose oxidase based glucose biosensors. Poly(3,4-ethylenedioxythiphene):poly(styrenesulfonate) (PEDOT:PSS) was synthesized by denatured catalase (no enzymatic activity) or iron-containing protein, transferrin.243 Ion-containing proteins may be widely used as oxidans for the synthesis of conducting polymers. An ultrasensitive assay for electrical biosensing of a DNA was developed by using target-guided formation of polyaniline base on an enzymatically (horseradish peroxidase) catalyzed

Conducting polyaniline synthesized by the laccase-mediator method has the conductivity approximately five times as great as that in the case of the laccase-catalyzed method. Polyaniline/multiwalled carbon nanotubes (PANI/ MWCNT) composite was prepared by using fungal laccase, potassium octocyanomolibdate (4+), and atmospheric oxygen as catalyst, redox-mediator, and terminal oxidant, respectively.230 Enzymatic catalysis allowed conducting process of oxidative aniline polymerization at environmentally friendly and rather mild conditions (aqueous slightly acidic solutions and room temperature) without toxic byproducts formation such as benzidine. The obtained PANI/MWCNT composite with polyaniline content ca. 49 wt % had high specific capacitance of ca. 440 F/g measured by cyclic voltammetry technique with potential scan rate of 5 mV/s. The in situ enzymatic polymerization of aniline onto multiwalled carbon nanotubes (MWCNT) and carboxylated MWCNT (COOH-MWCNT) was reported.231 Polymerization was catalyzed with the enzyme horseradish peroxidase at room temperature in aqueous medium of pH 4. The synthesized nanocomposites showed higher conductivity than pure polyaniline, which may be due to the strong interaction between the polyaniline chains and the MWCNT. The enzymatic polymerization of aniline to polyaniline with Trametes versicolor laccase (TvL) as catalyst was investigated in an aqueous medium containing unilamellar vesicles with an average diameter of about 80 nm.232 The reaction yielded mainly overoxidized products, which had a lower amount of unpaired electrons if compared with the products obtained with horseradish peroxidase isoenzyme C/H2O2 (the emeraldine salt form of polyaniline). The color changed from green (emeraldine salt) to blue (emeraldine base) upon exposure to ammonia gas, demonstrating the expected ammonia sensing properties. Polyaniline/Ag nanocomposites were successfully synthesized via in situ enzymatic (horseradish peroxidase) polymerization of aniline based on 2-aminothiophenol (2-ATP)-capped Ag nanoparticles.233 Smaller particle sizes were obtained with the enzymatic method than chemical oxidation. The fabrication of cellulose/polyaniline composites has been also tried. For example, polyaniline-based aqueous suspensions containing a variety of polyaniline contents ranging between 5 and 80 wt % have been successfully developed through in situ polymerization of aniline in a nanocellulose suspension, which may find potential applications in flexible electrodes, antistatic coatings, and electrical conductors.234 The chemical and enzymatic deposition of polyaniline films by in situ polymerization was studied.235 The film formation and polymerization processes were simultaneously monitored by the evolution of the open circuit potential and quartz-crystal microbalance measurements. The mechanism of film formation on substrates by the enzymatic polymerization was proposed as follows. Before the hydrogen peroxide addition, some enzyme and aniline cations were adsorbed on the surface. After adding the hydrogen peroxide, polymerization started in the solution, and it is highly likely that the polyaniline film grew from aggregation of particles by electrostatic adsorption and weak Van der Waal forces. Fully sulfonated polyaniline was synthesized by the peroxidase-mediated polymerization of aniline-2-sulfonic acid.236 The sulfonated polyaniline from chloroperoxidase, horseradish peroxidase, and chemically catalyzed reactions showed no differences in their UV−vis and FTIR spectra. The 2318


Chemical Reviews

Review

method.244 Poly(aniline) deposition was performed using the hybridized DNA strands as templates. The hybridized DNA resulted in a negatively charged surface originated from the phosphate groups, that provides a local environment of high acidity facilitating a predominantly head-to-tail coupling during the polymerization. DL-Tyrosine was polymerized under environmentally friendly conditions using linear−dendritic [amphiphilic block copolymer of poly(ethylene glycol) and poly(benzyl ether) dendron] laccase complexes as initiators and water as solvent.245 The structure of the poly(tyrosine) was discovered that the polymer chains consist of phenol repeating units linked together by CC and CO bonds randomly distributed along the backbone of the polymers. It was also found that the linear−dendritic laccase complexes induced further chain growth upon addition of fresh monomer to the preformed poly(tyrosine) in a fashion resembling the classic living polymerization. 4.1.2. Polymerization of Vinyl Monomers. Polymerization of acrylic monomers through direct enzyme catalysis via in situ radical generation was reported.246 Free radicals were generated by the enzyme/substrate combinations of xanthine oxidase/hypoxanthine, horseradish peroxidase/H2O2, chloroperoxidase/t-butyl-hydroperoxide, and alcholol oxidase/methanol. All the combinations allowed polymerization of acrylamide and hydroxyethyl methacrylate. Free-radical polymerization of acrylamide with using horseradish peroxidase/ H2O2 was also carried out in the presence of β-diketone.247,248 Laccase alone could polymerize acrylamide and N,Ndimethylacrylamide.249 Horseradish peroxidase has been used for the free-radical polymerization of vinyl monomers in the presence of H2O2 and β-diketones.102,250 Polystyrene particles were prepared via the horseradish peroxidase-mediated emulsifier-free emulsion polymerization of styrene. The results indicate that the β-diketone moieties, which were located on the surface of the particles, influenced the particle formation and stability. The preparation of polystyrene particles by enzyme-mediated miniemulsion polymerization with a polymerizable surfactant [i.e., N,N-dimethyl-N-n-dodecyl-N-2-methacryloyloxyethylammonium bromide (C-12-DMAEMA)], using a horseradish peroxidase/H2O2/β-diketone system, in water was reported.251 The H2O2 to horseradish peroxidase initial concentration ratio and the β-diketone to H2O2 initial molar ratio were key parameters for the optimization of the reaction. The influence of surfactant type on monomer conversion and particle size of polystyrene particles was also studied.252 A recombinant catalase-peroxidase HPI from Escherichia coli was prepared, purified, and used in enzymatic polymerization reactions for phenol, 3-methoxyphenol, catechol, and aniline.253 HPI was shown to mimic the activity of HRP for the production of oligomeric products. Chemoselective polymerization of 2-(4-hydroxyphenyl)ethyl methacrylate was realized. The horseradish peroxidase/H2O2 catalyst oxidatively pholymerized at the phenol moiety without causing the vinyl polymerization, while a radical initiator of AIBN induced vinyl polymerization.135 The horseradish peroxidase mediated free radical polymerization of methyl methacrylate was possible and accelerated in the presence of surfactants of sorbitane monooleate, bis(2ethylhexyl)sodium sulfosuccinate (AOT), or cetyltrimethylammonium bromide (CTAB).254 The horseradish peroxidase mediated inverse emulsion polymerization of acrylamide was also shown in supercritical carbon dioxide (scCO2), where the

enzymatic polymerization took place within water droplets formed in scCO2.255 An immobilized horseradish peroxidase in semi-interpenetrating network of poly(N-isopropylacrylamide)/chitosan initiated the acrylamide polymerization by a redox system (H2O2/2,4-pentanedione) in water at room temperature.256 The polymerization of acrylamide was catalyzed by a laccase or sarcosine oxidase catalyst.257 The laccase/acetylacetone catalyst induced the polymerization of methyl methacrylate and styrene in a mixture of water and tetrahydrofuran. Laccase alone also acted as a catalyst for the vinyl polymerization of acrylamide and methyl methacrylate without acetylacetone. In the polymerization of methyl methacrylate using lipoxidase as the catalyst in the presence of acetylacetone, the reaction occurred in air. The so-called laccase mediator system (LMS), comprising laccase and βdiketones, polymerization reactions can be performed using molecular oxygen as a terminal electron acceptor.258 The enzymatic polymerization of vinylformamide and sodium vinylsulfonate in water was achieved by using horseradish peroxidase/H2O2/2,4-pentanedione.259 Starch-polyacrylamide graft copolymer was obtained by free-radical polymerization of acrylamide from the starch main chain in the presence of horseradish peroxidase catalyst/H2O2/2,4 pentanedione.260 When hematin was used as a catalyst for oxidative polymerization, hematin/H2O2 gave poly(methyl methacrylate), quantitatively, and 2,4-pentanedione was further needed as a radical mediator for the polymerization of styrene and acrylamide.261 Vitamin C functionalized poly(methyl methacrylate) was prepared by using horseradish peroxidase as a catalyst. The pendent antioxidant vitamin C retained an ability to scavenge radicals.262 Indeed, the functionalized polymer showed modulation of the proliferation and osteogenic differentiation of early and late-passage bone marrow-derived hut-nail mesenchymal stem cells.263 Similarly, L-ascorbyl-4vinylbenzoate was polymerized by horseradish peroxidase/ H2O2/2,4-pentanedione in water/methanol to obtain the antioxidant-functionalized polystyrene.264 Free-radical polymerization of sodium styrenesulfonate and sodium acrylate in water was possible with using pegylated-hematin as a catalyst, which was due to the improvement of the water-solubility of hematin by pegylation.265 The enzyme-initiated polymerization of hydrophobic vinyl monomer of styrene in water was demonstrated by the application of miniemulsion polymerization. Horseradish peroxidase/H2O2/2,4-pentanedione was applied to the miniemulsion of SDS/hexadecane/styrene to obtain a stable polystyrene latex with a particle size of ca. 50 nm.266 Three categories of functional polymer particles, (i) monodisperse polymer particles were prepared by enzymatic miniemulsion polymerization using surfmer (surfactant+monomer), (ii) polymer particles stabilized by β-diketone moieties were prepared by enzymatic emulsifier-free emulsion polymerization, and (iii) a novel method of surface-initiated enzymatic graft polymerization was used to synthesize core−shell particles were prepared by horseradish peroxidase-mediated vinyl polymerization in the presence of H2O2 and β-diketone.267 4.2. Laccase, Tyrosinase, and Bilirubin Oxidase

Laccase (EC 1.10.3.2: Scheme 13A), tyrosinase (polyphenol oxidase, EC 1.10.3.1), and bilirubin oxidase (EC 1.3.3.5) are the enzymes containing copper as active center(s). They catalyze oxidative polymerizations of aromatic compounds by using hydrogen peroxide or oxygen gas (air) as an oxidant. There is a 2319


Chemical Reviews

Review

Laccase or HRP enzyme was employed for the polymerization of syringic acid, a natural product, yielding a PPO-type polymer, poly(2,6-dimethoxy-1,4-phenylene oxide), while liberating carbon dioxide. The polymer had a molecular weight up to 1.8 × 104 (Scheme 16).184,272,273 It is notable that both laccase having a Cu-containing active site and HRP having a heme-containing Fe-active site showed a similar catalytic function.98,99,270 This type of polymerization was not possible by a conventional metal complex catalyst. The product polymer was acidically demethylated with boron tribromide catalyst, to produce a new polymer, poly(2,6-dihydroxy-1,4-oxyphenylene) (Scheme 16), being thermally stable under 300 °C under nitrogen.185 Laccase catalysis showed a similar function with a copper/amine catalyst for the oxidative polymerization of 2,6Me2P to produce 2,6-Me2PPO.182 Laccase-catalyzed oxidative polymerization of 1-naphthol took place in aqueous acetone to produce the polymer having the molecular weight of several thousands.274 Two product polymers, poly(3,5-dimethoxy-1,4phenylene oxide) and 2,6-Me2PPO, were prepared by laccasecatalyzed polymerization and analyzed by two different mass spectroscopic methods (MALDI-TOF MS and ESI-FTICR MS).275 Reaction behaviors in the laccase-catalyzed polymerization of coniferyl alcohol depended on the origin of the enzyme. Laccases from Pycnoporus coccineus (PCL) and Coriolus versicolor showed a high catalytic activity to produce the dehydrogenative insoluble polymer, whereas laccase from Rhus vernicifera showed a much-reduced activity.276 In the case of laccase from Trametes versicolor, the molecular weight increase was observed by the treatment of soluble lignin.277 Laccase-catalyzed oxidation of phenols was examined, where phenol substrates include phenol, m-cresol, bisphenol A, and 4t-butylphenol (4-TBP) (Scheme 17).278 The reaction was carried out in an aqueous alcoholic solvent (pH 5, acetate buffer) at room temperature with using PCL as a catalyst. The product polymers were obtained in high yields, 84−97% yields, the Mn values of which were 2300, 14800, 21300, and 1900, for phenol, m-cresol, bisphenol A, and 4-TBP, respectively. The polymer structure consisted of both a phenylene unit (Ph) and an oxyphenylene unit (Ox). The unit ratios of polymers (Ph:Ox) for these four substrates were 48:52, 46:54, 45:55, and 52:48, respectively. For 4-TBP, 2-propanol content in the solvent (from 30% to 60%) controlled the Ph:Ox ratio from 65:35 to 46:54. It was not easy, however, to gain the high regioselectivity reflected by the Ph:Ox ratio.278 In order to find the optimal operational conditions for maximum initial reaction rate in the laccase-catalyzed polymerization of bisphenol A (BPA), a multistep response surface methodology (RSM) was applied.279 The enzymatic polymerization rate of BPA was studied through RSM, based on the measurements of the initial dissolved oxygen consumption rate in a closed batch system. The optimal conditions were evaluated to be 748.46 mg/L, 32.24 °C, and 15.92% for monomer concentration, temperature, and solvent content, respectively. These results were in good agreement with the observed responses. Methoxyphenols (o-, m-, and p-) as well as 2,6-dimethoxyphenol were oxidatively polymerized by a fungal laccase catalyst using oxygen in air as an oxidant.280 From omethoxyphenol, polymers of molecular weight 7000−11000 were obtained. Polymerization rate was correlated with the energy of the highest occupied molecular orbitals (EHOMO) of the phenols.

Scheme 13

recent review paper available on tyrosinase enzyme from microbial origin with emphasis on their biochemical properties and discussing their current and potential applications for pharmaceutical, food bioprocessing, and environmental technology.268 4.2.1. Polymerization of Phenolic Compounds. 4.2.1.1. Substituted Phenol Compounds. In relation to the copper-containing enzyme catalysis, it is noted that a copper/ amine catalyst was found to cause an oxidative coupling polymerization of 2,6-disubstituted phenols with the use of oxygen gas as an oxidant in 1959 (Scheme 14).103,269 When R Scheme 14

= methyl, the product polymer, poly(oxy-2,6-dimethyl-1,4phenylene) (2,6-Me2PPO), had high molecular weight, Mn = 28000, with good properties. Owing to the pioneering work, 2,6-Me2PPO was commercialized from GE Co. and it is currently one of several important engineering plastics.98,99,270 As the mechanism to selectively form C−O coupling to the polymer 2,6-Me2PPO, three possible pathways were proposed as seen in Scheme 15: (i) coupling of free phenoxy radicals resulting from one-electron oxidation of 2,6-dimethylphenol (2,6-Me2P), (ii) coupling of phenoxy radicals coordinated to the catalyst complex, and (iii) coupling of phenoxonium cation formed by two-electron-oxidation of 2,6-Me2P.98,99,270 A theoretical calculation suggested that case (iii) would be most likely.271 The Cu/amine catalyst can be regarded as a Cucontaining oxidase model catalyst from its function. 2320


Chemical Reviews

Review

Scheme 15

Scheme 16

Scheme 18

It was extremely difficult to control the coupling regioselectivity of ortho-unsubstituted phenols by conventional catalysts. The reaction generates free phenoxy radical, in which ortho-positions readily take part in further reactions. Tyrosinase is a Cu-containing enzyme for catalyzing oxidation reactions like laccase. A tyrosinase model complex (Scheme 13B) catalyzed oxidative coupling polymerizations while suppressing the reaction at ortho-positions and produced unsubstituted PPO from unsubstituted 4-phenoxyphenol (Scheme 19A)40,282−286 and 2,5-Me2PPO from 2,5-dimethylphenol (Scheme 19B).40,287 Regioselectivity to give para-linkage was high (∼80%), and typically the product PPO was obtained in 77% yields at 40 °C for 24 h, having a Mn = 5400 (Mw = 29100).282,283 In these reactions, an intermediate phenoxy radical is considered not free, but complexed with Cu, and thus “controlled radical” species is involved in the coupling reaction, where the ortho-position is blocked from attacking by another radical species (“radical-controlled” oxidative polymerization).282,283,286 As a typical example of the polymerization of 2,5dimethylphenol (Scheme 19B), the reaction was carried out at 40 °C in toluene under oxygen gas (1 atm) for 24 h with 5 mol % of Cu-containing tyrosinase model complex to give a white polymer of 2,5-Me2PPO in 78% isolated yield.287 The polymer had values of Mn = 3900 (Mw = 19300). It is the first example showing heat-reversible crystallinity (Tm1 and Tm2 = 308 and 303 °C, respectively). Surprisingly, the isomeric polymer 2,6-Me2PPO showed a melting point at 237 °C (Tm1), but once the crystalline part was completely melted, recrystallization never occurred (Tm2 not detected) via slow cooling or annealing.288 Since thermoplastic polymers are mainly used as melt-moldings, 2,6-Me2PPO is generally considered as an amorphous polymer, while 2,5-Me2PPO can be practically regarded as a crystalline polymer. White polymers, 2,5-nPr2PPO and 2,5-Et2PPO, had Mw values, 23100 and 32000, respectively, and showed heat-reversible

Scheme 17

Linear−dendritic laccase complexes catalyzed the polymerization of DL-tyrosine, which is the enzyme-catalyzed “green” synthesis of an unnatural poly(amino acid) under environmentally friendly conditions carried out in water solvent.245 The poly(tyrosine) structure is of phenolic polymer type having C−O and C−C bonds via coupling in the main chain. Depending on the reaction conditions poly(tyrosine) with an Mw up to 82000 could be obtained in yields ranging between 45 and 69%. A similar type of poly(tyrosine) was obtained earlier by HRP-catalyzed polymerization.134 A new oxidative polymerization of 4-fluoroguaiacol (4-fluoro2-methoxyphenol) was achieved using the catalyst of laccase from Trametes versicolor in acetone or methanol and buffer (pH 5) mixture media with oxygen as oxidant to give poly(2methoxy-1,4-phenylene oxide) (Scheme 18).281 The polymer was derived by defluorination during a phenyl-oxy propagation, having low molecular weight up to 7 repeat units. The polymer is a novel photoluminescent material, displaying fluorescence with emissions in blue, green, and red upon applied UV-light frequencies. It is, therefore, potentially interesting for its use as a component of electronic devices. 2321


Chemical Reviews

Review

peroxo dicopper(II) complex (1), a copper−dioxygen model complex for tyrosinase.291 Then, complex 1 abstracts proton (not hydrogen atom) from phenol to give phenoxo−copper(II) complex (2), equivalent to phenoxy radical−copper(I) complex (3). Intermediate 3 is not a “free” radical but a “controlled” radical, which is identical with the coordinated radical described above. Thus, if a catalyst generates (and regenerates) only a nucleophilic active oxygen intermediate, followed by the reaction with phenols to give “controlled” radicals without formation of “free” radicals, the regioselectivity of the subsequent coupling will be entirely regulated. This view explains the exclusive formation of “controlled” phenoxy radicals, and hence, the new concept was termed “radicalcontrolled” oxidative polymerization.283 A computational approach was studied to confirm the mechanistic speculation.292 Just for reference, it is mentioned here that an HRP model complex, Fe-salen (Scheme 13C) shows similar catalysis as a tyrosinase model complex [e.g., for the oxidative polymerization of 2,6-difluorophenol to poly(2,6-difluoro-1,4-phenylene oxide) (2,6-F2PPO) (Scheme 19D)].195 Further, Fe-salen catalyzed the oxidative polymerization of 2,6-disubstituted phenols in 1,4-dioxane to produce 2,6-disubstituted PPO194 and of bisphenol A to give a soluble polyphenol.193 4.2.1.2. Polyphenol Compounds. Enzymatic polymerization is effective also for polyphenol compounds having more than two OH groups. The simplest polyphenol compounds are dihydroxy-benzenes (catechol, resorcinol, and p-hydroquinone), the polymerization of which was carried out using laccase catalyst. The mechanism of polymerization, and the structures of the polymers were evaluated in terms of UV−vis and FT-IR spectroscopy. It was estimated that polymers were formed; catechol is linked through C−O bond, resorcinol and p-hydroquinone through C−C bond, respectively. The numberaveraged molecular weights of the polymers ranged from 1000 to 1400.293 The formation of catechol oligomers, di-, tri-, and tetramers, was achieved via an oxidative polymerization catalyzed by synthetic water-soluble iron-porphyrin as an efficient alternative to biolabile natural peroxidase. It was demonstrated that the both C−C and C−O−C coupling mechanisms occurred.294 The oxidative coupling of catechol catalyzed by the similar catalyst formed three major phenylene dimers, which was elucidated by a number of NMR experiments.295 The laccase-catalyzed polymerization of a lignin-based macromonomer, lignocatechol, was carried out in ethanol− phosphate buffer solvent system to give cross-linked polymers in good yields. The copolymerization was also performed with

Scheme 19

crystallinity with a Tm2 at 276 °C, but the former did not show a detectable Tm2.40 Therefore, the recrystallization of poly(alkylated phenylene oxide)s after melting seems to be governed by the position as well as the nature of their alkyl substituents.270 Simple phenol derivatives, m-cresol289 and o-cresol,290 were regioselectively polymerized by using a μ−η2:η2 peroxo dicopper(II) complex, a tyrosinase model complex, as catalysts to give soluble 3-MePPO (Scheme 19C) and 2-MePPO, respectively, in good yields. Structure of 3-MePPO consists mainly of a 3-methyl-1,4-oxyphenylene unit as shown in (C) and a small amount of 3-methyl-1,2-oxyphenylene unit. The molecular weight of 3-MePPO was Mn = 3.95 × 104 and Mw = 2.15 × 10 5 , showing high thermal stability with Td 5 (temperature at 5% weight loss) 392 °C. Those values for 2MePPO were Mn = 3.8 × 103, Mw = 8.4 × 103, and high thermal stability value Td10 (temperature at 10% weight loss) 437 °C. On the basis of the above observations, a plausible explanation was made (Figure 3).98,99,270 First, a “nucleophilic” active oxygen species is generated in the form of a μ-η2:η2-

Figure 3. Plausible explanation for “radical-controlled” oxidative polymerization involving nucleophilic and electrophilic radical complexes. Reprinted with permission from ref 99. Copyright 2004 Wiley. 2322


Chemical Reviews

Review

low inhibition activity. The amplified activities might offer high potential as a therapeutic agent for prevention of various enzyme-related diseases.300 Further, flavonoid compounds were enzymatically polymerized primarily for the enhancement of biological activities such as antioxidant properties. The flavonoids polymerized include rutin (Scheme 20)301 and quercetin.302 The laccase-catalyzed

urushiol to afford the corresponding copolymers in high yields. The polymerization mechanism was estimated as that the polymerization proceeded through quinone radical intermediate at a catechol ring to give the cross-linked polymers.296 Laccase enzymes catalyzed the coupling short nonpolar chains containing aromatic groups onto flax fibers and nanofibrillated cellulose (NFC) with different lignin contents. Laccases, Trametes villosa, Pycnoporus cinnabarinus, and Myceliophthora thermophile, were used to facilitate surface coupling and to produce materials with different levels of hydrophobicity, increasing the resistance to water absorption. The highest hydrophobization levels of flax fibers was achieved by coupling dodecyl 3,4,5-trihydroxybenzoate, which yielded water contact angles (WCAs) of 80−96 degrees and water absorption times (drop tests) of ca. 73 min. In cases, WCAs in the range of 87− 104 degrees were achieved.297 Laccase catalyzed the polymerization of catechin, a polyphenol compound (Scheme 12). Soluble poly(catechin) with a Mn around 3 × 103 was obtained in a mixture of acetone and acetate buffer (pH 5). The polymer synthesized in 20% acetone showed low solubility toward N,N-dimethylformamide (DMF), whereas the polymer obtained in the acetone content less than 5% was completely soluble in DMF. In the UV−vis spectrum of poly(catechin) in methanol, a broad peak centered at 370 nm was observed. In alkaline solution, this peak red shifted and the peak intensity became larger than that in methanol. In the ESR spectrum of the poly(catechin), a singlet peak at g = 1.982 was detected, whereas the catechin monomer possessed no ESR peak.298 The polymerization of catechin greatly enhanced the antioxidant property; the poly(catechin) exhibited a much enhanced superoxide anion scavenging activity compared with the catechin monomer. Similarly, the poly(catechin) showed greatly amplified xanthine oxidase (XO) inhibitory activity (Figure 4).298

Scheme 20

polymerization of a water-soluble rutin derivative, a commercially available product, in an equivolume mixture of methanol and acetate buffer produced the polymer with a molecular weight of 1 × 104 in a high yield. Superoxide scavenging nature of poly(rutin) showed a much enhanced activity, compared with rutin itself. A similar result was observed for LDL oxidation.301 From poly(rutin) obtained by the laccase catalyst, rutin polymer fraction (RPF) was separated, which has 6−8 monomer units. Bioactivity of RPF showed more effectively suppressed adipogenesis in 3T3-L1 adipocytes compared to monomeric rutin, indicating that enzyme-catalyzed polymerization improved bioactivity of flavonoids.303 Laccase and tyrosinase from Ustilago maydis were used as catalysts for the oxidative polymerization of flavonoid compounds, quercetin and kaempferol. The former produced aggregates with relatively low molecular weight and higher antioxidant activity than the monomer quercetin. The product aggregates from kaempferol reached higher sizes, and the antioxidant activity increased in the beginning of the polymerization. Both product polymers showed strong scavenging effect on reactive oxygen species and inhibition of lipoperoxidation.304 The effect of different parameters on the enzymatic polymerization of sodium lignosulfonates (SLS) by laccase was compared with that of the chemical treatment by Mn(III). It was found that the enzymatic process is more efficient than a Mn(III) catalyst. SLSs of different initial molecular weights, commercial SLS (17800), F1 (4300), and F2 (2500) were employed. High commercial SLS (50 g/L) and laccase (30 U/ mL) concentrations led to the formation of polymers with increased molecular weight (1.086 × 105).305 4.2.2. Synthesis Approach to Artificial Urushi. Among many polymers from substituted catechol or phenol compounds, “urushi” is a very unique natural macromolecule. Urushi is the cured and cross-linked material of urushiol harvested from a special urushi tree (Rhus vernicifera). It is the only example in nature that is polymerized with the catalyst of natural enzyme of laccase and cured via oxidation in air. It is a traditional Japanese lacquer (more widely, oriental lacquer)

Figure 4. XO inhibition activity of poly(catechin), n = 3. ○, catechin; □, poly(catechin).298 Reprinted with permission from ref 298. Copyright 2003 Wiley-VCH.

A polymer conjugate was prepered by laccase-catalyzed oxidation polymerization of cathechin in the presence of poly(allylamine), leading to the poly(allylamine)-poly(catechin) conjugate. The conjugate showed a good antioxidant property against low-density lipoprotein (LDL) peroxidation induced by a free radical.299 The laccase-catalyzed conjugation of catechin on poly(ε-lysine), the product poly(ε-lysine)catechin conjugate, showed greatly improved inhibition effects against disease-related enzymes, collagenase, hyaluronidase, and xanthine oxidase. While, the catechin monomer showed very 2323


Chemical Reviews

Review

UA was possible when R is a diene or triene. The reaction was carried out in the presence of acetone powder (AP, an acetone insoluble part of the natural urushi sap containing mainly polysaccharide and glycoprotein) at 30 °C under 80% relative humidity for 24 h. AP is believed to act as an emulsifier of oily urushiol and aqueous laccase solution. The curing of the trienetype UA was measured by a dynamic microhardness tester (Figure 5, curve 1c). The curing was slow at the biginning, and

known from ancient time, going back to more than nine thousand years ago in Japan. This is understandable from the following view that Japan means the country name, while “japan” possesses meanings such as “urushi lacquer”, “urushi wares”, or “to coat urushi”. In Japan, there are many national treasures coated with urushi, which is brilliant and tough, and looks soft. Accordingly, chemical studies on structure of urushiol and its hardening mechanism to urushi have been performed by excellent chemists since more than a century ago.306−313 Urushiol is found not only in Japan but also in China and Korea, while laccol (from Vietnam, Taiwan) and thitsiol (from Thailand and Myanma) are the main components in other oriental lacquer trees. It is noted, by the way, that Yoshida discovered laccase enzyme in 1883.306 Natural urushi is formed by laccase-catalyzed polymerization of urushiol in air, oxygen gas as an oxidant, followed by curing via slow air oxidation of the polymerized urusiol.166,169,314,315 Urushiol is of catechol structure, having an unsaturated C15 hydrocarbon chain at the 3-position (Scheme 21A).169,314 Urushi sap is obtained from the urushi tree, which is known to cause allergy for human skin, like a so-called “poison ivy” tree often found in USA or European countries. Scheme 21

Figure 5. Time course of in hardening of artificial urushi films: curve 1c of the triene and curve 1b of the diene.315 Reprinted with permission from ref 315. Copyright 2000 Chemical Society of Japan.

after 2 weeks the hardness suddenly increased. Later, the value reached about 150 N mm−2. The pencil scratch hardness was H, which is hard enough for practical usages. The hardness and gloss values of the cured film were compatible with those to natural urushi coatings. UA from the diene, on the other hand, showed very low hardness (curve 1b). It is noted that the laccase catalysis was examined for three natural phenolic lipids obtained from oriental lacquer trees.317 Urushiol and laccol, both having an unsaturated hydrocarbon chain, C15 and C17, respectively, at 3-position of catechol, were effectively cured in the presence of laccase enzymes to produce the cross-linked polymeric films with a high gloss surface and high hardness properties. On the other hand, thitsiol having an unsaturated hydrocarbon chain (C17) at 4-position of catechol was slowly cured with laccase and the hardness attained was low. This different behavior comes from the different content of the unsaturated group in the hydrocarbon chain.317 As another approach to artificial urushi, cross-linkable polyphenols were prepared from a phenol derivative, lipasecatalyzed acylation of 4-hydroxyphenethyl alcohol with an unsaturated fatty acids (Scheme 22A). Compared with a catechol derivative (Scheme 21), the product phenol-type compound (also, urushiol analogue) was much less reactive, and hence, it was not polymerized by laccase catalyst. But, it was oxidatively polymerized by the iron-N,N′-ethylenebis(salicylidene-amine) (Fe-salen, shown in Scheme 13C) catalyst to produce a prepolymer.152,318 By the polymerization at room temperature in tetrahydrofuran under air, all three kinds of compounds produced soluble oily prepolymers with Mn between 4 × 103 and 1.6 × 104. During the polymerization, the unsaturated moiety did not react; the reaction took place on the phenol moiety. The curing of the prepolymers was carried out via two methods: the oxidation catalyzed by cobaltnaphthenate (3 wt %) in air and the thermal treatment (150 °C

Since natural urushi is very expensive and difficult to prepare in vitro, its chemistry was mimicked for approaching to “artificial urushi” (or “man-made urushi”),166 with hoping cheaper cost for practical applications. Artificial urushi is mentioned in section 4.1.1.1 but in more detail here. Natural urushiol analogues having a catechol structure were newly prepared for this purpose (Scheme 21B).315 4-Hydroxymethylcatechol was esterified by an unsaturated alkenyl carboxylic acid by the lipase catalyst to give 4-substituted catechol, which was called as an “urushiol analogue” (UA). Then, it was cured by laccase-catalyzed cross-linking reaction in air, giving rise to “artificial urushi”.315,316 This modeling is the first example of the single-step synthesis of urushi-like cured film from a monomeric phenol derivative. The enzymatic cross-linking of 2324


Chemical Reviews

Review

Scheme 22

for 2 h). The sample prepolymers from R = diene and triene were cured by both methods to give a hard film; however, the cured film was not obtained from R = monoene. Cardanol (Scheme 22B) is a main component obtained from cashew nut shell liquid (CNSL), byproducts out of mechanically treated cashew kernel for edible use. It is a phenol derivative having a meta-substituent of a C 15 unsuturated hydrocarbon chain, mainly with 1−3 double bonds. CNSL contained cardanol (83−84%), cardol (8− 11%), and 2-methylcardol (2%). It is said that recently the content of cardanol in the available CNSL is much increased. Cardanol is widely used practically for resins, laminations, surface coating and paints, electrical insulating, vanishes, brake lining, etc., as shown in recent studies.319,320 Thus, cardanol becomes available cheaply and a good candidate of a phenolic compound for artificial urushi. Laccase did not induce the cardanol polymerization, but Fesalen catalyzed the polymerization of cardanol.164,166,167,321−323 The polymerization was carried out by using H2O2 as an oxidant in an organic solvent like 1,4-dioxane, tetrahydrofuran, and toluene, at 30 °C for 2 h, giving rise to poly(cardanol), prepolymer, in 32−72% yields, with molecular weight Mn around several thousands. The prepolymer structure showed a mixture of phenylene and oxyphenylene units, without involving the reaction of unsaturated groups in the side chains. Curing of the prepolymer was conducted by a Co-naththenate catalyst (3 wt %) at ambient temperature in air or by the thermal treatment (150 °C for 30 min) in air. The cured films showed the hardness value of 80−100 N/mm2. This system was considered as an environmentally friendly process for polymer coatings, giving a good example to conduct “green polymer chemistry”.164 The film looked brilliant, having toughness and elasticity, and hence, it was named “man-made urushi”.166

It was found that soybean peroxidase (SBP) catalyzed the cardanol (CNSL) polymerization.165 The reaction catalyzed by SBP (10 mg for 0.60 g of monomer) in isopropyl alcohol/pH 7 phosphate buffer (50:50 in volume) at room temperature for 24 h in air gave a soluble cardanol prepolymer with molecular weight Mn = 6100 in 69% yields. During the reaction, only the phenolic moiety was involved in the polymerization. With an increased amount of SBP (50 mg), insoluble powdery materials were quantitatively obtained. In the similar reaction conditions, HRP did not cause the cardanol polymerization.159 It is a characteristic of SBP, therefore, that SBP allowed the polymerization of a phenolic monomer having a large metasubstituent like cardanol. The curing of the cardanol prepolymer was accomplished with a Co-naphthenate catalyst (3 wt %) at room temperature on the glass slide, and after 14 days, the film reached to H by the pencil scratch hardness test, which is hard enough for usual industrial use. The SBP catalyst was also employed, and product polymers showed a high potential as commercial coating materials and an excellent antifouling activity.168 It was found that HRP was inactive but became active for the cardanol polymerization when a redox mediator like N-ethyl phenothiazine is combined with HRP in water. The cardanol polymerization with this system afforded similar polymerization results as those of the SBP catalyst.170 Further, the oxidative polymerization of cardanol (CNSL) was examined in more detail by using various metal-containing catalysts to lead to the prepolymer and the curing behaviors of the prepolymers.167 Metal complexes with metal = Fe, Co, Cu, Mn, Ni, Zn, and VO having the salen-type ligand were employed for catalyst and Fe(salen) showed the strongest activity; Fe-salen produced the prepolymer with a Mn = 3.0 × 103 (Mw/Mn = 3.0) in 70% yields at room temperature for 2 h, 2325


Chemical Reviews

Review

above, a large number of chemicals and products have been developed starting from CNSL by taking advantage of the three reactive sites, namely, phenolic hydroxyl, aromatic ring, and unsaturation(s) in the alkenyl side chain.324 To search a wider possibility of artificial urushi, new crosslinkable polynaphthols were prepared by oxidative polymerization of naphthol monomers, in which an unsaturated group R from ROH is from linseed oil or fish oil (Scheme 23). Fesalen catalyst was employed for the polymerization, during which only the naphthol moiety was involved in the reaction to give a soluble naphthol prepolymer having Mn = 3.0−4.6 × 103 in high yields (81−87%). The thermally cured film showed the gloss hardness value over 150 N/mm2 after 1 day and around 200 N/mm2 after 1 week at 150 °C. With Co-naphthenate catalyst, the gloss value reached 210 N/mm2 after 1 week at 25 °C. The cured film was strong and brilliant, and those values were comparable to natural urushi.175 Cardol is one of major components derived from CNSL having a C15 unsaturated hydrocarbon chain with 1−3 double bonds at a meta position of resorcinol. Its oxidative polymerization was carried out for the first time using Coprinus cinerius peroxidase (Scheme 24). Under appropriate conditions, the polycardol was successfully obtained in 66% yield and a Mw of 13500. Compared to poly(cardanol), poly(cardol) was rapidly cured at room temperature within 4 h to give hardened dry and dark brown color coatings. The curing rate of polycardol was higher than that of polycardanol irrespective of curing methods. It was described that due to the TGA analysis poly(cardol) was more thermostable than poly(cardanol) when cured at room temperature.172 Natural anacardic acid (Scheme 25) was obtained from CNSL and enzymatically polymerized using soybean peroxidase (SBP) as catalyst, hydrogen peroxide as an oxidizing agent, and phenothiazine-10-propionic acid as a redox mediator, respectively. The polymer is of a structure having oxyphenylene and phenylene units through the phenol rings. Aqueous solvents played an important role in the polymer production yield and molecular weight. With 2-propanol, the highest production yield (61%) of polymer molecular weight 3900 was observed, and with methanol, higher-molecular-weight polymers (5000) were produced with lower production yields (43%). The resulting poly(anacardic acid) was cross-linked on a solid surface to form a permanent natural polymer coating. The FTIR analysis indicates the cross-linking reaction between the polymers through the unsaturated alkyl side chains.174 Copolymerization of urushiol with a lignin-based macromonomer (lignocatechol) was examined by laccase catalyst. Both urushiol and lignocatechol were soluble in the mixed solvent of ethanol and phosphate buffer in the proportion of

while other catalysts gave polymers with smaller molecular weight, in less yields, even at 80 °C for 24 h. Co-, Cu-, Fe-, and V-acetyl acetone complexes and Mn-, Cu-, and Fe-phthalocyanine complexes were also weaker than Fesalen in the activity. The curing behaviors of the prepolymer (sample 1, Mn = 5.9 × 103, Mw/Mn = 1.5), a commercially available CNSL-formaldehyde resin, and natural urushi sample are shown in Figure 6. The figure indicates that poly(cardanol)

Figure 6. Dynamic viscoelasticity of the cured film from (●) polyCNSL (cardanol) of sample 1; (■) CNSL-formaldehyde resin; and (▲) natural urushi.167 Reprinted with permission from ref 167. Copyright 2002 Elsevier.

afforded the cured film with good viscoelastic properties, comparable to other two sample films, in which Tg of the cured sample 1 was ca. 90 °C. Poly(cardanol) synthesized by enzymatic polymerization using fungal peroxidase as catalyst was partially or fully cured by using methyl ethyl ketone peroxide (MEKP) as an initiator and cobalt naphthenate (Co-Naph) as an accelerator. The curing behavior of poly(cardanol) was investigated in terms of curing temperature, curing time, concentration of initiator and accelerator, and the monomer-to-polymer conversion of poly(cardanol). The optimal conditions for fully curing poly(cardanol) were 1 wt % MEKP, 0.2 wt % Co-Naph, curing time 120 min, and curing temperature 200 °C. These results suggested that a poly(cardanol) with high monomer-topolymer conversion would be useful for processing a poly(cardanol) matrix composite under the optimal conditions of curing.173 A recent paper described general information on cardanol (CNSL), its purification and separation methods, reactivity, and applications in polymer chemistry, with emphasizing the importance of cardanol as biobased starting materials. As seen Scheme 23

2326


Chemical Reviews

Review

Scheme 24

HRP/H2O2 catalyst as seen above.327 This is a clear-cut evidence that the polymerization occurred through the laccase catalysis; however, the mechanism of how the initiating species is formed is not clarified. In the presence of AA, the polymerization of Am took place at room temperature. From the microstructure analysis of polyAm, the polymerization was of a free radical mechanism. Catalytic activity study of the Am polymerization revealed that in the presence of AA in water under argon at 65 °C for 4 h, laccases and sarcosine oxidase showed high activity; the polymerization produced polyAm with high molecular weight, Mn = 1.1 × 105−4.6 × 105 in high yields (>88%).257 Lipoxidase showed a little activity. Other oxidase enzymes, such as bilirubin oxidase, choline oxidase, glucose oxidase, and xanthine oxidase, did not show any activity. Methyl methacrylate (MMA) was polymerized by laccase/AA catalyst in water/ THF at 45−75 °C for 24 h to afford polyMMA with Mn = 1.3 × 105−2.7 × 105 in 77−97% yields. Laccase/AA catalyst induced the polymerization of styrene (St) at 60 °C, giving rise to polySt with a Mn = 8 × 104 in a low yield of 24% (Scheme 26). Stereoregulation of polyMMA and polySt was not observed, suggesting a free radical mechanism. AA was needed as a catalyst component.257 A laccase-mediator system induces the radical polymerization of vinyl monomers and is environmentally benign. Then, the practical feasibility of the Am polymerization using the mediator system with molecular oxygen as an oxidant was examined under various reaction conditions. Optimal conditions were slightly acidic reaction media at around 50 °C. The molecular weight of polyAm, Mn = 6−28 × 104 (Mw/Mn 2.5− 3.2), could be controlled via the ratio of monomer to enzyme. It was thought that at that time the system did not yet fulfill the requirements of economic feasibility on the industrial scale, but the biocatalytic polymerizations seemed to involve a big potential for a greener and more sustainable production of polymers.258 Xanthine oxidasae, chloroperoxidase, and alcohol oxidase catalyzed the Am polymerization in the presence of bisacrylamide and formed cross-linked products.246 In the radical polymerizations, most of the reactions were carried out at 60 °C under acidic conditions. However, laccase rapidly loses its native activity at 60 °C at pH 6.328 Vinyl polymerizations have been also described on not only HRP catalysts but also laccase catalyst.102

Scheme 25

70:30 vol %. The copolymerization was finished within 12 h to give the polymers with high thermal stabilities because of the cross-linked structures. The copolymers were obtained in good yields in all proportions of urushiol and lignocatechol.296 In addition, natural urushiol was separated and a thermally crosslinked urushi thin film was prepared by using iron(II) acetate as the additive. Iron(II) acetate could be regarded as a substitute of Fe-salen. Various properties of the film were examined, and the important function of the unsaturated long hydrocarbon chain of urushiol was pointed out.325,326 4.2.3. Polymerization of Vinyl Monomers. For the vinyl monomer polymerization, HRP is most often used as a catalyst among oxidoreductase enzymes.327 In the HRP-catalyzed polymerization of acrylamide (Am), using an HRP/additive catalyst was essential to generate initiating radical species, where the additive was a β-diketone like acetylacetone (AA) (Scheme 26). The polymerization gave high molecular weight polyAm in good yields.247 Scheme 26

On the other hand, laccase enzyme catalyst alone induced the polymerization of Am in water under argon at a temperature 50−80 °C to give high molecular weight polyAm having Mn up to 1 × 106 in good yields (up to 81%) (Scheme 26).249 The reaction did not occur at 65 °C under air, probably due to too much oxygen, a strong radical inhibitor of radical polymerization. The polymerization took place without an additive of mediator like AA, which is usually needed in the case of the 2327


Chemical Reviews

Review

Scheme 27

4.2.4. Other Reactions. Laccase was used to catalyze the curing of synthetic polymers having phenol moiety in the side chain.329 The polymer (3) was prepared via the copolymerization of phenoxy moiety-containing comonomer (1) and comonomer (2), having Mn ∼ 2.6 × 104 with a fraction of 1 as 12−100 mol % (Scheme 27). The sample film on a glass slide prepared from copopylmer 3 containing 12 and 23 mol % of monomer 1 became insoluble in any solvent after curing with laccase/O2 catalyst due to cross linking. The cross linking was confirmed by the UV−visible spectrum; a strong absorption at 278 nm before the curing shifted to 296 nm, indicating the C− C bond formation at the phenolic moieties. Also, a similar cross-linking reaction was reported by Fe(salen) catalyst.330 Phenol-containing precursor poly(amino acid)s, poly(αglutamine), poly(α/β-asparagine), and poly(γ-glutamine) derivatives, were subjected to oxidative coupling by using the Fesalen and HRP catalysts to produce a new class of soluble poly(amino acid)s without formation of insoluble gels. The process is suggested to be useful for development of new functional polymeric materials from renewable resources.331 Poly(ethylene oxide)-poly(propylene oxide)-poly(ethylene oxide) triblock copolymers were used as a mediator in the laccase-induced graft polymerization of diacrylic derivate of polyethylene glycols, giving rise to the formation of PEG-g-F68 hydrogels. The proper oxygen content in the reaction medium to obtain appropriate polymerization conversions (i.e., on the one hand, laccase needs oxygen as substrate, whereas on the other, oxygen is a strong inhibitor of radical polymerizations) was achieved by the use of an enzymatic scavenging system consisting of glucose oxidase and glucose. Eventually, laccase was immobilized within the resulting PEG-g-F68 hydrogel with full preservation of enzyme activity. Laccases have been used for bioremediation purposes because of their ability to degrade phenolic compounds. Thus, laccase-immobilized PEG-g-F68 hydrogels were submitted to the ISISA (ice segregation induced self-assembly) process for preparation of laccase-immobilized PEG-g-F68 cryogels which exhibited a macroporous structure where immobilized laccase preserved almost total activity (ca. 90%), showing outstanding adsorption capabilities to the cryogels (up to 235 mg g−1).332 A polymer-enzyme-multiwalled carbon nanotubes (MWCNTs) cast films was prepared for electrochemical biosensing and biofuel cell applications, which involved in situ laccase-catalyzed polymerization. The enzyme-catalyzed polymerization proposed as a new and efficient biomacromolecule-immobilization platform is considered promising in

preparing many other multifunctional polymeric bionanocomposites for wide applications.333 Tyrosinases were employed for modification of macromolecules. Mushroom tyrosinases (EC 1.14.18.1) are oxidation enzymes capable of converting low-molecular weight phenols and accessible tyrosyl residues of proteins like gelatin into oquinones. These o-quinones are reactive and undergo nonenzymatic reactions with a variety of nucleophiles. The primary amino groups of chitosan reacted with the quinone, giving rise to chitosan-natural phenol or chitosan−catechin conjugates via a Michael-type addition and/or a Schiff base formation (Scheme 28).334−336 Scheme 28

Tyrosinase effectively catalyzed several reactions of the conjugate-formation between chitosan and various proteins; cytochrome C,337 organophosphorus hydrolase (OPH),336 histidine-tagged chloramphenicol acetyltransferase (HisCAT),337 gelatin,338,339 green fluorescent protein,340 silk fibroin,341−343 and silk sericine.344 These tyrosinase-catalyzed reactions enabled proteins to be covalently tethered to a threedimensional chitosan gel network. This covalent coupling allowed the easy fabrication of biocatalytically active hydrogelbased membranes, films, and coating for a wide range of applications. Combining the biodegradable and biocompatible 2328


Chemical Reviews

Review

amine) for the oxidative polymerization in aqueous SDBS micellar solutions.349 The rate of the aniline dimer oxidation was markedly higher than the rate of the aniline oxidation under the similar conditions. Thus, aniline dimer formation may be the rate-limiting stage of the enzyme-catalyzed aniline polymerization. MALDI-TOF analysis showed the products of aniline oligomers with the polymerization degree of 4−22. TEM measurements showed that polyaniline nanoparticles have a granular shape. The aniline dimer PADPA was polymerized to poly(PADPA) at 25 °C with Trametes versicolor laccase (TvL)/O2 in the presence of vesicles formed from sodium bis(2-ethylhexyl) sulfosuccinate (AOT) as templates.350 The polymerization of PADPA is much faster, and considerably fewer enzymes were required for complete monomer conversion. Turbidity measurements indicate that PADPA strongly binds to the vesicle surface before oxidation and polymerization are initiated. Such binding is confirmed by molecular dynamics simulations, supporting the assumption that the reactions which lead to poly(PADPA) are localized on the vesicle surface. The poly(PADPA) obtained resembles the emeraldine salt form of polyaniline (PANI-ES) in its polaron state with a high content of unpaired electrons. There are, however, also notable spectroscopic differences between PANIES and the enzymatically prepared poly(PADPA). Poly(PADPA) appears to be similar to a chemically synthesized poly(PADPA) as obtained in a previous work with ammonium peroxydisulfate (APS) as the oxidant in a mixture of 50 vol % ethanol and 50 vol % 0.2 M sulfuric acid.351 ESI-MS measurements of early intermediates of the reaction with TvL and AOT vesicles indicated that the presence of the vesicles decreases the extent of formation of unwanted oxygencontaining species in comparison to the reaction in the absence of vesicles. Laccase-mediated system based on potassium octocyanomalybdate (4+) was used for acceleration of the enzymatic aniline polymerization to give conducting PANI. The enzymatic reaction yielded oxidized octocyanomalybdate (5+), which can oxidize the aniline monomer to the aniline radical cation. Comparison of reactions of the aniline oxidative polymerization with laccase and laccase-mediator system showed that the presence of the mediator accelerates the polymerization of the monomer. Conducting PANI synthesized by the laccasemediator method showed the conductivity approximately five times as great as that in the case of the laccase-catalyzed method and was obtained in a higher yield. PANI/SDBS complexes synthesized by the both methods have a granular structure but differ in the particle size.229 The aniline polymerization to PANI with Trametes versicolor laccase (TvL)/O2 was investigated in an aqueous medium containing unilamellar vesicles with an average diameter of about 80 nm formed from AOT. Compared to the same reaction carried out with HRP isoenzyme C (HRPC)/H2O2), notable differences were found in the kinetics of the reaction, as well as in the characteristics of the PANI obtained. Under comparable optimal conditions, the reaction with TvL/O2 was much slower than with HRPC/H2O2 (i.e., ≈27 days vs 1 day reaction time).232 The polymerization of pyrrole with the laccase (TvL) and dioxygen (O2) in aqueous solution at pH 3.5 was found to be regulated by anionic vesicles formed from AOT to afford polypyrrole (PPy).352−354 The polymeric products obtained in the presence of the vesicular templates have high absorption at λ ≈ 450 and 1000 nm, which is indicative for PPy in its

properties of chitosan, these novel hybrid biomaterials are good candidates as a promising material for gene and drug delivery as well as for regenerative medicine and tissue engineering. Also, tyrosinase catalyzed the oxidative coupling of soluble lignin fragments to afford the insoluble polymers.345 Further, designed peptides of Kcoil and Ecoil are known to heterodimerize in a highly specific and stable fashion to adopt a coiled-coil structure. Kcoil-functionalized chitosan was prepared using the tyrosinase-catalyzed oxidation of a tyrosine-containing Kcoil peptide. From the surface plasmon resonance (SPR) investigation of coil tagged epidermal growth factor (EGF)/ Kcoil chitosan interactions, it was found that the Kcoil chitosan conjugate could capture Ecoil-tagged EGF via coiled-coil mediated interaction. In this case, the coiled-coil interaction was relatively low and some improvements are essential for the general usage. This approach seems to provide with multiple added values to chitosan.346 As described in section 4.1.1.2, polyaniline (PANI) (Scheme 29) was extensively studied because of its wide potential for Scheme 29

many technological applications, such as organic lightweight batteries, microelectronics, optical displays, electromagnetic shielding, and anticorrosion coatings, etc. PANI is synthesized by not only a chemical method but also an enzymatic method; by the latter oxidative polymerization process, HRP/H2O2 was widely used as a catalyst. Besides HRP, bilirubin oxidase (BOD), a Cu-containing oxidoreductase, catalyzed the oxidative polymerization of aniline and 1,5-dihydroxynaphthalene. The aniline polymerization in the presence of BOD-solid matrix gave PANI film containing the active enzyme.211 The film was electrochemically reversible in the redox properties. The BOD-catalyzed polymerization of 1,5-dihydroxynaphthalene in aqueous organic solvents gave the insoluble polymer.347 The UV spectrum of the film exhibited a wide band from 300 to 470 nm, showing a long π-conjugation structure. After treatment of the polymer with HClO4, the polymer had an electro-conductivity of 10−3 S/cm. Laccase isolated from the fungi, Trametes hirsuta, catalyzed the oxidative polymerization of aniline in micellar solutions of the anionic surfactant sodium dodecylbenzenesulfonate (SDBS), in which the atmospheric oxygen served as an oxidizing agent. The polymerization gave a stable dispersion of electroactive PANI/SDBS complexes. It was shown that the laccase-catalyzed polymerization is kinetically controllable, and its mechanism is very distinctly different from that of chemical polymerization. The synthesized PANI/SDBS complexes were characterized using FT-IR and UV−vis spectroscopy, cyclic voltammetry, termogravimetry, transmission electron microscopy, and electron diffraction. The antistatic properties of the prepared PANI/SDBS composite were also studied. It was noted that laccase-catalyzed synthesis of conducting PANI has advantages as compared with both peroxidase-catalyzed and chemical methods due to the use of air oxygen as an oxidant in the reaction of aniline polymerization.348 The same laccase was used as a catalyst for the aniline dimer, PADPA (p-aminodiphenylamine = N-phenyl-1,4-phenylenedi2329


Chemical Reviews

Review

Figure 7. Stoichiometric equation for the polymerization of pyrrole into polypyrrole with the laccase (TvL) as a catalyst and O2 as an oxidant. In (A), three possible mesomeric structures of a tetrameric repeating unit of the bipolaron form of polypyrrole (PPy) are shown; two formal positive charges are localized either on two of the carbon atoms or on two of the nitrogen atoms of the repeating units. In (B), two possible polaron forms of PPy with the typical unpaired electrons are depicted, either as separated polarons (above) or as polaron pair (below).354 Repinted with permission from ref 354. Copyright 2015 Elsevier.

5.1. Glycosyltransferases

conductive bipolaron state. Absence of unpaired electrons in the bipolaronic PPy product obtained was supported by EPR measurements. Furthermore, the FTIR spectrum of isolated PPy is comparable with that reported in literature for chemically or electrochemically synthesized PPy. Structure of PPy along with its function is illustrated in Figure 7. These results were compared with those of PANI in regards to AOT vesicle function.

Polysaccharides are composed of one or several kinds of monosaccharide units linked through glycosidic linkages.356 The reaction for the formation of a glycosidic linkage, so-called glycosylation, is performed between two substrates, glycosyl donor and acceptor, in which an anomeric carbon at C-1 of the former is activated by introducing a leaving group, and a hydroxy group in the latter, which participates in the reaction, is employed as a free form, whereas other hydroxy groups in both the substrates are protected.357−359 The enzymatic synthesis of polysaccharides has been realized via the formation of glycosidic linkages by means of the enzymatic glycosylation approach.32,33,38,39,41,43,49,52,54−56,360,361 It has generally been accepted well that the enzymatic glycosylation is a very powerful tool for the stereo- and regioselective construction of glycosidic linkages under mild conditions, where a glycosyl donor and a glycosyl acceptor can be employed in their unprotected forms, leading to the direct formation of unprotected saccharide chains in aqueous media. In the reaction, first, the glycosyl donor is recognized by an enzyme to form a glycosyl-enzyme intermediate (or transition-state).

5. TRANSFERASES Transferases (EC class 2) are enzymes that catalyze to transfer a group from a donor substrate to an acceptor substrate. Although transferases have potential as catalysts to synthesize interesting polymeric materials, they are often very sensitive for isolation on the practical scale. Several transferases such as glycosyltransferases and synthases, however, have been found to be effective for catalyzing in vitro synthesis of polysaccharides and polyesters via greener processes mainly in aqueous media under mild conditions without use of harmful catalysts such as heavy metals and formation of byproducts.355 2330


Chemical Reviews

Review

Scheme 30

Scheme 31

practically synthesizing a wide variety of polysaccharides.368 Phosphorylases, sucrases, and branching enzyme, classified into non-Leloir glycosyltransferases, on the other hand, have been employed as catalysts for the practical synthesis of polysaccharides by both polymerization and modification.369−372 5.1.1. α-Glucan Phosphorylase. 5.1.1.1. Synthesis of Amylose and Other α(1→4)-Linked Polysaccharides. Various phosphorylases have been known, and all of them catalyze an exowise reversible phosphorolysis of the glycosidic linkage at the nonreducing end in the presence of inorganic phosphate to produce monosaccharide 1-phosphates (Scheme 30).373,374 Phosphorylases are generally classified by the anomeric forms of the glycosidic linkages in the substrates which are phosphorolyzed or by the anomeric forms of the monosaccharide 1-phosphates which are produced. The other way employed to classify phosphorylases is according to describing them in terms of the anomeric retention or inversion in the reaction. The stereo- and regiospecificities of phosphorylases are very strict, and they catalyze the phosphorolysis of the specific types of glycosidic linkages. Some phosphorylases catalyze the reactions for a synthetic way to polysaccharides or even oligosaccharides with relatively high degrees of polymerization (DPs) via reverse reactions of phosphorolysis, but other phosphorylases recognize only disaccharide substrates and

Then, the intermediate is attacked by the hydroxy group of the glycosyl acceptor, giving a glycoside (see also Figures 1 and 2). Enzymes involved in the synthesis of polysaccharides via enzymatic glycosylations are categorized into two main classes: glycosyltransferases (EC 2.4)362 and hydrolases (glycosidases, vide infra, 6.1).363 Glycosyltransferase catalyzes the transfer of a sugar moiety from an activated donor onto both saccharide and nonsaccharide acceptors. They are divided into the Leloir and non-Leloir types according to the types of glycosyl donors. Leloir glycosyltransferases are biologically important because they perform the role of synthesizing saccharide chains in vivo.364,365 Accordingly, natural polysaccharides are produced in vivo by Leloir glycosyltransferase catalysis via biosynthetic pathway. The enzymatic reactions are irreversible in the synthetic direction because of involving cleavage of the highenergy bond of the glycosyl-nucleotide of a substrate donor in the reaction. For example, natural cellulose is synthesized in vivo from uridine-5′-diphospho (UDP)-glucose via cellulose synthase (EC 2.4.1.12)-catalyzed reaction.366,367 However, Leloir glycosyltransferases are generally transmembrane-type proteins, present in very small amount in nature, and unstable for isolation and purification with difficulty in handling. Thus, the enzymes are expensive and hardly available and accordingly have not been employed as the catalyst for in vitro approach to 2331


Chemical Reviews

Review

Scheme 32

phosphorylase catalyzes enzymatic polymerization according to the reaction manner of successive glucosylations at the elongating nonreducing ends to produce a polysaccharide composed of α(1→4)-glucan chain, that is, amylose (Scheme 31).369,378−380 The polymerization manner catalyzed by αglucan phosphorylase belongs to chain-growth polymerization, and the glycosyl acceptor is often called a “primer” of the polymerization because the polymerization is exactly initiated at the nonreducing end of the acceptor. Therefore, the α-glucan phosphorylase-catalyzed enzymatic polymerization proceeds analogously as a living polymerization because of excluding the occurrence of termination and chain-transfer reaction. Accordingly, the molecular weights (or DPs) of the produced amyloses can be controlled by the Glc-1-P/primer feed ratios and their polydispercities are narrow (Mw/Mn < 1.2).384 Since the substrate, Glc-1-P for α-glucan phosphorylase is relatively expensive, to overcome this problem, coexistence systems using other enzymes that produce Glc-1-P has been reported. For example, the combined use of sucrose phosphorylase (EC2.4.1.7) with α-glucan phosphorylase was reported for the in situ production of Glc-1-P from sucrose as the substrate for the amylose synthesis.385,386 Use of cellobiose phosphorylase (EC 2.4.1.20) combined with α-glucan phosphorylase was also examined for the purpose to produce Glc-1-P in situ.379,380,387 Because of loose specificity for the recognition of substrates, α-glucan phosphorylase recognizes analogue glycosyl donors of Glc-1-P and catalyzes the corresponding glycosylations to produce oligosaccharides having the different monosaccharide residues at the nonreducing end.388,389 Potato α-glucan phosphorylase has been found to recognize several monosaccharide 1-phosphates as the analogue substrates of Glc-1-P, such as α-D-xylose (Xyl), α-D-mannose (Man), α-D-glucosamine (GlcN), and N-formyl-α-D-glucosamine (GlcNF) 1phosphates.390−393 Accordingly, potato α-glucan phosphorylase-catalyzed glycosylations using such substrates as glycosyl donors took place with Glc4, a smallest glycosyl acceptor of the enzyme, to produce non-natural pentasaccharides having the corresponding monosaccharide residue at the nonreducing end (Scheme 32). In these reactions, only a monosaccharide residue

catalyze the reversible phosphorolysis to produce the corresponding monosaccharide 1-phosphates and other monosaccharides. Only a few phosphorylases [i.e., α-glucan and cellodextrin phosphorylases (EC 2.4.1.1, and 2.4.1.49, respectively)] have been used in various investigations for the practical synthesis of poly- or oligosaccharides with relatively higher DPs, as well as the related poly- and oligosaccharidebased materials.56,372 Although it has been reported that several phosphorylases, such as kojibiose phosphorylase (EC 2.4.1.230), 3 7 5 β-1,3-oligoglucan phosphorylase (EC 2.4.1.30),376 and β-1,2-oligoglucan phosphorylase,377 recognize glucans with the higher DPs and catalyze their phosphorolysis, there have not been many studies on the synthesis of poly- or oligosaccharides catalyzed by these enzymes. Of the phosphorylases, α-glucan phosphorylase (glycogen phosphorylase, starch phosphorylase, or simply phosphorylase) is the most extensively studied.372 The role of α-glucan phosphorylase is considered to be in utilization of storage polysaccharides.373 This enzyme catalyzes the reversible phosphorolysis of α(1→4)-glucans at the nonreducing end, such as glycogen and starch, in the presence of inorganic phosphate, giving rise to α-D-glucose 1-phosphate (Glc-1-P) (Scheme 31). By means of the reversibility of the reaction, α(1→4)-glucosidic linkage can be constructed by the α-glucan phosphorylase-catalyzed glucosylation using Glc-1-P as a glycosyl donor (Scheme 31).369,378−380 As glycosyl acceptors, maltooligosaccharides with DPs higher than the smallest one, which recognized by the enzyme, are used. In the glucosylation, a glucose residue is transferred from Glc-1-P to a nonreducing end of the acceptor to form an α(1→4)-glucosidic linkage. The smallest substrates for the phosphorolysis and glucosylation recognized by potato α-glucan phosphorylase are maltopentaose (Glc5) and maltotetraose (Glc4), respectively. Recently, it has been reported that the smallest DPs of substrates accepted by α-glucan phosphorylase isolated from thermophilic bacteria sources (thermostable phosphorylase) for the former and latter reactions are one smaller than those by potato α-glucan phosphorylase (i.e., Glc4 and maltotriose (Glc3), respectively).379,381−383 When the excess molar ratio of Glc-1-P to the acceptor is present in the reaction system, α-glucan 2332


Chemical Reviews

Review

Scheme 33

Figure 8. Synthesis of GD by BE-catalyzed cyclization of (A) amylopectin and (B) synthesis of dendritic acidic and amphoteric α-glucans by thermostable α-glucan phosphorylase-catalyzed glucuronylation and subsequent glucosaminylation.

Furthermore, a fraction of higher molecular weight products with an average DP of 12 was also obtained. Thermostable α-glucan phosphorylase has shown the different recognition specificity for the glycosyl donor from that obtained from potato. For example, potato α-glucan phosphorylase does not recognize α-D-glucuronic acid 1phosphate (GlcA-1-P), whereas thermostable phosphorylase from Aquifex aeolicus VF5396 recognizes GlcA-1-P and catalyzed the glucuronylation of Glc3, which was a smallest glycosyl acceptor for this enzyme, to produce an acidic tetrasaccharide having a GlcA residue at the nonreducing end (Scheme 33).397 By means of this enzymatic reaction, dendritic acidic α-glucans have been synthesized.398 For this purpose, a highly branched cyclic dextrin (glucan dendrimer, GD) was employed as a multifunctional glycosyl acceptor. This material is a watersoluble dextrin that is produced from amylopectin by cyclization catalyzed by the branching enzyme (BE, EC 2.4.1.18, Bacillus stearothermophilus, vide infra) (Figure

is transferred from the donor to the acceptor, but successive glycosylations, that is, enzymatic polymerization does not occur. 2-Deoxy-α-D-glucose 1-phosphate (dGlc-1-P) was also reported to be recognized by potato α-glucan phosphorylase.394 Interestingly, this substrate could be prepared by two-step processes in situ.395 First, D-glucal was transferred as a 2-deoxyD-glucose unit to a nonreducing end of α(1→4)-glucan in the presence of inorganic phosphate. In the second step, 2-deoxy-Dglucose was released by the potato α-glucan phosphorylasecatalyzed phosphorolysis to produce dGlc-1-P, and consequently, α(1→4)-glucan remained as unchanged in the overall reaction. Accordingly, D-glucal was applied as a glycosyl donor in the potato α-glucan phosphorylase-catalyzed enzymatic 2deoxy-α-glucosylation. In the enzymatic reaction in the presence of D-glucal, Glc4, and only 0.05 equiv of inorganic phosphate, successive glucosylations took place to produce 2deoxy-α-D-glucosylated penta-, hexa-, and heptasaccharides. 2333


Chemical Reviews

Review

Scheme 34

8A).399−401 Because GD has a number of the nonreducing α(1→4)-glucan ends, this acts as a multifunctional glycosyl acceptor for the phosphorylase-catalyzed glycosylation. The thermostable α-glucan phosphorylase-catalyzed glucuronylation of GD (Mn = 1.25 × 105, the number of nonreducing ends = ca. 59) with GlcA-1-P was carried out to produce acidic α-glucans (Figure 8B), and the glucuronylation ratios of the GlcA residues to the nonreducing ends were 34.8, 50.5, and 64.4% in the products obtained by the donor/acceptor feed ratios of 0.5, 1, and 2, respectively. These results indicated that the glucuronylation ratios were controlled by the feed ratios. The enzymatic synthesis of dendritic amphoteric α-glucans having both GlcA and GlcN residues at the nonreducing ends was further conducted by the subsequent thermostable α-glucan phosphorylase-catalyzed glucosaminylation of the acidic products using GlcN-1-P (Figure 8B).402 The successive reactions in various glycosyl donor feed ratios of GlcA-1-P/GlcN-1-P gave the amphoteric materials having the different GlcA/GlcN ratios. Their inherent isoelectric points were calculated by ζpotential measurement, which were reasonably dependent upon the GlcA/GlcN ratios in the products. As aforementioned, potato α-glucan phosphorylase catalyzes the enzymatic polymerization as the successive glucosylation manner to produce the α(1→4)-glucan chain when the native substrate, Glc-1-P, is used as a glycosyl donor. The above results using the analogue substrates indicate, on the other hand, that if the transfer of a monosaccharide residue from the analogue substrates to the nonreducing end of the acceptor occurs once, further glycosylation was suppressed because the different structures from Glc residue at the nonreducing end are not recognized at the acceptor site of potato α-glucan phosphorylase. It has been found that thermostable phosphorylase catalyzes the different reaction manner from potato one when Man-1-P and GlcN-1-P were used as glycosyl donors.403 When the thermostable α-glucan phosphorylase-catalyzed reactions of Glc3 with 10 equiv of Man-1-P and GlcN-1-P were carried out, successive mannosylations and glucosaminylations took place to give non-natural heterooligosaccharides composed of α(1→4)-linked mannose and glucosamine chains at the nonreducing end of Glc3, respectively. The MALDI-TOF

MS of the crude products (feed ratio of donor to acceptor =10:1) showed several peaks corresponding to the molecular masses of tetra-octa-saccharides having one-five Man or GlcN residues with Glc3. The further progress of the enzymatic chain elongation has been inhibited by inorganic phosphate produced from the glycosyl donors because it is a native substrate for the phosphorolysis reaction of α-glucan phosphorylase. An attempt, thus, was made to remove inorganic phosphate from the enzymatic glucosaminylation system by precipitation in an ammonium buffer (0.5 M, pH 8.6) in the presence of MgCl2, on the basis of the fact that inorganic phosphate formed an insoluble salt with ammonium and magnesium.404 Consequently, the thermostable α-glucan phosphorylasecatalyzed enzymatic polymerization of GlcN-1-P from the Glc3 primer (30:1) took place according to the manner of successive glucosaminylations in the buffer system at 40 °C for 7 days to produce α(1→4)-linked glucosamine polymer with a Mn = 3760 with precipitation of phosphate salt, which corresponded to the structure of a chitosan stereoisomer, a non-natural aminopolysaccharide (Scheme 34).405 The isolated chitosan stereoisomer was then converted to a chitin stereoisomer, also a non-natural aminopolysaccharide, by Nacetylation using acetic anhydride in aqueous Na2CO3. The enzymatic copolymerization of Glc-1-P with GlcN-1-P from the Glc3 primer was also achieved by thermostable αglucan phosphorylase catalysis under the conditions of the removal of inorganic phosphate as the precipitate with ammonium and magnesium ions to obtain non-natural aminopolysaccharides composed of Glc/GlcN units, that is, α(1→4)-linked glucosaminoglucans.406 The analytical results suggested the relative random sequence of the two units in the products. 5.1.1.2. Synthesis of Amylose-Grafted Heteropolysaccharides and Polypeptides. As aforementioned, the reaction manner of α-glucan phosphorylase-catalyzed enzymatic polymerization belongs to chain-growth polymerization, in which the polymerization is initiated at the nonreducing end of the primer and propagation proceeds via successive transfers of glucose residues from the monomers to the propagating nonreducing end. Therefore, the α-glucan phosphorylase2334


Chemical Reviews

Review

polymerization using a primer having a polymerizable group at the reducing end.411 Radical polymerization of the macromonomer gave an amylose-grafted polystyrene (Scheme 35A). This type of the graft material was also synthesized by radical polymerization of a styrene-type macromonomer having a maltooligosaccharide, followed by the α-glucan phosphorylasecatalyzed enzymatic polymerization from the nonreducing end of the maltooligosaccharide primer on the product (Scheme 35B).411,412 By the similar approach, amylose-grafted dimethylsiloxane,413 polyacetylene,414,415 and poly(vinyl alcohol)416 have been synthesized. Amylose brushes were also fabricated by α-glucan phosphorylase-catalyzed enzymatic graft polymerization of Glc-1-P from maltoheptaose primers covalently attached to Au and Si surfaces.417 The detailed characterization of the products was conducted by X-ray photoelectron spectroscopy and spectroscopic ellipsometry measurements. Amylose-grafted heteropolysaccharides have been synthesized by chemoenzymatic approach including α-glucan phosphorylase-catalyzed enzymatic polymerization.56,372,410,418−420 To produce these types of materials, the maltooligosaccharide primer should be first introduced onto the main-chain polysaccharides by appropriate chemical reactions and then α-glucan phosphorylase-catalyzed enzymatic polymerization from the immobilized primer is performed (chemoenzymatic method). Two approaches for the chemical reactions have been employed to introduce the maltooligosaccharide primer onto the main-chain polysaccharides (i.e., reductive amination of maltooligosaccharide with aminated basic polysaccharides using reducing agent and condensation of an amine-functionalized maltooligosaccharide with carboxylated acidic polysaccharides using condensing agent) (Figure 10A). By means of the former approach using chitosan as a basic polysaccharide with amino groups, a maltooligosaccharidegrafted chitosan was synthesized, which was further converted into a maltooligosaccharide-grafted chitin by N-acetylation. Then, α-glucan phosphorylase-catalyzed enzymatic polymerization of Glc-1-P from the maltooligosaccharide nonreducing ends on the products was performed to give amylose-grafted chitin/chitosan (Figure 10B).421,422 An amylose-grafted cellulose was synthesized by the similar approach.423 A partially aminated cellulose at C-6 positions was used as the main-chain, and accordingly, which was reacted with a maltooligosaccharide

catalyzed enzymatic polymerization can be performed using the modified primer, where the reducing end that does not participate in the reaction, is immobilized on other materials (Figure 9).56,372,407−410 By means of this reaction characteristic,

Figure 9. Propagation manner in α-glucan phosphorylase-catalyzed enzymatic polymerization initiated from a modified primer.

a styrene-type macromonomer having an amylose chain was prepared by the α-glucan phosphorylase-catalyzed enzymatic Scheme 35

2335


Chemical Reviews

Review

Figure 10. Chemoenzymatic synthetic routes of (A) amylose-grafted polysaccharides and (B and C) their structures from basic and acidic polysaccharides.

indicating the occurrence of self-assembling nanofibrillation from the present material. After removal of the residual NaOH from the film by immersing in water, the SEM image showed that the nanofibers were merged at the interface, while the fiber arrangement was maintained. Glycogen is a water-soluble polysaccharide composed of α(1→4)-linked linear glucan chains interlinked by α(1→6)glucosidic linkages,428,429 which has the similar structure as that of GD. Besides it being the substrate for in vivo phosphorolysis by α-glucan phosphorylase (glycogen phosphorylase), glycogen has been used as a multifunctional polymeric primer for αglucan phosphorylase-catalyzed enzymatic polymerization because of the presence of a number of the nonreducing α(1→4)-glucan chain ends owing to the highly branched structure.430 When α-glucan phosphorylase-catalyzed enzymatic polymerization of Glc-1-P was carried out from glycogen as the primer and then the reaction mixture was left standing further at room temperature, it totally turned into a hydrogel form (Figure 11). The hydrogelation has reasonably been explained

by reductive amination to obtain a maltooligosaccharide-grafted cellulose. An amylose-grafted cellulose was then produced by αglucan phosphorylase-catalyzed enzymatic polymerization of Glc-1-P using the product (Figure 10B). On the other hand, the latter condensation approach has been performed to introduce maltooligosaccharide to acidic polysaccharides, such as alginate,424 xanthan gum,425 and carboxymethyl cellulose (CMC) (Figure 10C).426,427 Condensation of an amine-functionalized maltooligosaccharide at the reducing end with carboxylates in the acidic polysaccharides was carried out using condensing agent to produce maltooligosaccharide-grafted polysaccharides. Then, α-glucan phosphorylase-catalyzed enzymatic polymerization of Glc-1-P from the maltooligosaccharide nonreducing ends on the products was conducted to yield amylose-grafted alginate, xanthan gum, and CMC (Figure 10C). When a NaOH aqueous solution of the amylose-grafted CMC was casted on a glass plate, followed by drying, a film was formed. The SEM image of the film showed highly entangled nanofiber morphology, 2336


Chemical Reviews

Review

Figure 11. Preparation of glycogen hydrogel by α-glucan phosphorylase-catalyzed enzymatic polymerization.

Figure 12. Image of vine-twining polymerization and representative guest polymers.

ide-grafted poly(L-glutamic acid) formed a helical conformation at lower pH and a random coil conformation at higher pH. However, little α-helix content was observed for the amylosegrafted poly(L-glutamic acid), indicating that the helical formation was disturbed by the amylose chains, probably owing to steric hindrance. 5.1.1.3. Preparation of Amylose Supramolecules. An enzymatically synthesized pure amylose by α-glucan phosphorylase has been recognized as a candidate for functional polymeric materials because it acts as a host molecule owing to its left-handed helical conformation and forms supramolecular inclusion complexes with various guest molecules with a relatively low molecular weight.433 The driving force for inclusion of the guest molecules in the cavity is the host−guest hydrophobic interaction, as the inside of the amylose helix is hydrophobic. Therefore, the guest molecules, which are included by amylose, are typically hydrophobic. However, little has been reported regarding the formation of inclusion complexes between amylose and polymeric guest mole-

by the formation of cross-linking points based on the double helix conformation of the elongated amylose graft-chains among glycogen molecules. The gelation process by the α-glucan phosphorylasecatalyzed enzymatic polymerization was also conducted on amphoteric glycogens having GlcA/GlcN units at the nonreducing ends, which were prepared by the similar manner for the amphoteric GD, to obtain amphoteric glycogen hydrogels.431 The products exhibited a pH-responsive property. The synthesis of the polysaccharide−polypeptide conjugate [i.e., amylose-grafted poly(L-glutamic acid)] was also investigated by the similar chemoenzymatic method.432 Maltopentaosylamine was first condensed with the pendant carboxyl groups of poly(L-glutamic acid) using a condensing agent to give a maltooligosaccharide-grafted poly(L-glutamic acid). The α-glucan phosphorylase-catalyzed enzymatic polymerization of Glc-1-P from the maltooligosaccharide primers on the poly(Lglutamic acid) main chain was then performed to produce the amylose-grafted poly(L-glutamic acid). The maltooligosacchar2337


Chemical Reviews

Review

Scheme 36

Scheme 37

cules.434−443 It is considered that amylose does not have sufficient ability to directly include the long chain of the polymeric guest into its cavity only by weak hydrophobic interaction. An efficient method for the formation of amylosepolymer supramolecular inclusion complexes has been found by means of α-glucan phosphorylase-catalyzed enzymatic polymerization.56,372,410,444−449 When the enzymatic polymerization of Glc-1-P from a maltoheptaose (Glc7) primer is conducted in the presence of appropriate hydrophobic guest polymers dispersed in an aqueous polymerization solvent, the propagation is progressed with the inclusion of the guest polymers (Figure 12). The representation of this system for the complexation of amylose with the guest polymer is similar the way that vines of plants grow twining around a rod. Accordingly, it has been proposed that this polymerization method be named “vine-twining polymerization”. As the guest polymers for this polymerization system, hydrophobic poly-

ethers [e.g., polytetrahydrofuran (PTHF), polyoxetane (POXT)], 450,451 polyesters (e.g., poly(δ-varerolactone) (PVL), poly(ε-caprolactone) (PCL), poly(glycolic acid-co-εcaprolactone)),452−454 poly(ester-ether),453 and polycarbonates [e.g., poly(tetramethylene carbonate)]455 have been employed to construct supramolecular inclusion complexes with amylose (Figure 12). When the vine-twining polymerization was conducted using optically active polyesters, that is, poly(lactide)s (PLAs), which had three kinds of the stereoisomers [i.e., poly(L-lactide) (PLLA), poly( D -lactide) (PDLA), and poly( DL-lactide) (PLDLA)], as guest polymers, amylose, produced by the enzymatic polymerization, perfectly recognized the chirality in PLAs on complexation, and consequently, it only formed an inclusion complex with PLLA (Scheme 36).456 Amylose has also shown the selectivity toward polymers with resemblant chemical structures in the vine-twining polymerization. For 2338


Chemical Reviews

Review

Figure 13. Preparation of amylose supramolecular materials through formation of inclusion complexes by vine-twining polymerization.

ization progressed, the resulting inclusion complexes acted as cross-linking points for the formation of the supramolecular hydrogel. This vine-twining polymerization approach using another graft copolymer as the guest was also performed to produce an amylose supramolecular film through hydrogelation (Figure 13).464 A hydrogel was obtained according to the vine-twining polymerization using CMC-g-PCL. A film was fabricated by adding water to a powdered sample, obtained by lyophilization of the hydrogel, followed by drying. The supramolecular hydrogel showed the macroscopic healing behavior when poly(γ-glutamic acid-graft-ε-caprolactone) (PGA-g-PCL) was used for the vine-twining polymerization (Figure 13).465 The supramolecular hydrogel from PGA-g-PCL was cut into two pieces, and a sodium acetate buffer solution containing α-glucan phosphorylase and Glc-1-P was put on the surfaces of the two hydrogel pieces. After the surfaces were contacted, the gels were left standing at 40 °C for 6 h for the progress of the enzymatic polymerization. Consequently, the two pieces had cohered at the interface. The healing of the hydrogels on a macroscopic level was achieved by the complexation of the enzymatically produced amyloses with the PCL graft chains at the interface. 5.1.1.4. Self-Assembly of Amylose Block and Graft Copolymers. Amylose diblock copolymers were synthesized by α-glucan phosphorylase-catalyzed enzymatic polymerization of Glc-1-P using polymeric primers having a maltooligosaccharide moiety at the chain end. For example, an amylose-blockpolystyrene was produced by α-glucan phosphorylase-catalyzed enzymatic polymerization from the nonreducing end of a maltooligosaccharide covalently attached to the chain end of polystyrene.466−468 The polymerization could be initiated even though the primer-functionalized polystyrene was insoluble in aqueous polymerization solvent. When micellar aggregates of the products with various compositions were formed, the

example, amylose selectively included one side of the polyethers, that is, PTHF from a mixture of PTHF/ POXT.457 The selective inclusion by amylose was also found when the vine-twining polymerization was conducted in the presence of a mixture of two resemblant polyesters, PVL/PCL, in which PVL was selectively included by amylose.458 Furthermore, it was found that amylose selectively included a specific range of molecular weights in any PTHFs when the vine-twining polymerization was performed in the presence of PTHFs with different average molecular weights.459 The inclusion complexes composed of amylose and strongly hydrophobic polyesters were formed in a parallel enzymatic polymerization system. This was achieved by conducting αglucan phosphorylase-catalyzed polymerization of Glc-1-P from Glc7, giving rise to the host amylose, and lipase-catalyzed enzymatic polycondensation of dicarboxylic acids and diols,460,461 leading to the guest polyesters, simultaneously (Scheme 37).462 When the numbers of methylene units in a dicarboxylic acid and a diol as the monomers for the guest polyester were 8, the corresponding amylose-polyester inclusion complex was obtained. On the other hand, use of the monomers having methylene units of 10 and 12 hardly gave the inclusion complexes. The vine-twining approach by α-glucan phosphorylasecatalyzed enzymatic polymerization was applied to the architecture of amylose supramolecular materials such as gel and film using a designed guest polymer, such as poly(acrylic acid sodium salt-graf t-δ-varerolactone) (P(AA-g-VL); PAA acts as a main component of polymeric network in the hydrogel, and PVL is the guest polymer for the complexation with amylose. When the vine-twining polymerization was performed by α-glucan phosphorylase-catalyzed enzymatic polymerization of Glc-1-P with the Glc7 primer in the presence of the graft copolymer, the reaction mixture turned into a hydrogel (Figure 13).463 Because the produced amyloses included the PVL graft chains in the intermolecular guest copolymers as the polymer2339


Chemical Reviews

Review

Scheme 38

analytical data indicated the presence of unimers, oligomers, and large micellar species in THF. An amphiphilic amylose-block-methoxypoly(ethylene oxide) (MPEO) was synthesized by α-glucan phosphorylase-catalyzed enzymatic polymerization of Glc-1-P using a primer-MPEO conjugate.469,470 A sole amylose was insoluble in chloroform, while the amphiphilic product was slightly soluble in this solvent. It was confirmed that the complexation of MPEOblock-amylose with methyl orange was significantly enhanced in the amylose domain of the associate in chloroform. Maltopentaose-block-alkyl chain surfactants (octyl, C8Glc5; dodecyl, C12Glc5; hexadecyl, C16Glc5) formed micelles in water, which were dissociated upon α-glucan phosphorylasecatalyzed enzymatic polymerization (Scheme 38).471,472 Moreover, the micelle-to vesicle transition of the mixed lipid/primer systems was controllably caused by the enzymatic polymerization. An enzyme-responsive artificial chaperone system using C12Glc5 and α-glucan phosphorylase b was designed to enable protein refolding. Effective refolding of carbonic anhydrase B after both heat denaturation and guanidine hydrochloride denaturation was observed by controlled association between protein molecules and the C12Glc5 micelle through the enzymatic polymerization. When α-glucan phosphorylase-catalyzed enzymatic polymerization of Glc-1-P was conducted using a maltoheptaosefunctionalized PLLA (Glc7-PLLA) as a primer−guest conjugate, the propagation proceeded according to the vinetwining polymerization manner to form an inclusion supramolecular polymer composed of amylose and PLLA (Figure 14).473 The product formed a polymeric continuum of an inclusion complex in which PLLA is included by an amylose chain enzymatically elongated from the other Glc7-PLLA. The similar approach was also conducted using Glc7-PTHF to obtain an inclusion supramolecular polymer composed of amylose and PTHF.474 The relative chain orientation of amylose and PLA in inclusion complexes formed by α-glucan phosphorylase-catalyzed enzymatic polymerization was made clear by using four kinds of primer-guest conjugates, which had a maltoheptaosyl moiety at COOH- or OH-terminus of PLLA or PDLA (Glc7-PLLA, PLLA-Glc7, Glc7-PDLA, and PDLAGlc7).475 The enzymatic polymerization products from Glc7PLLA and PLLA-Glc7 were amylose-PLLA inclusion supramolecular polymers, indicating that the cavity of amylose included PLLA regardless of the chain orientation of PLLA. On the other hand, the products from Glc7-PDLA and PDLA-Glc7 were amylose-PDLA diblock copolymers due to the noninclusion owing to the recognition behavior on chirality by amylose, regardless of the chain orientation of PDLA.

Figure 14. Preparation of inclusion supramolecular polymer by vinetwining polymerization using primer−guest conjugate.

Accordingly, it was concluded that PLLA forms left-handed helix, the same as that of amylose, responsibly implying the inclusion complexation, whereas the directions of methyl substituents in PLA toward amylose, which are oppositely changed depending on the relative chain orientation, do not affect the complexation. The vine-twining polymerization using a branched Glc7PLLA2 conjugate produced a hyperbranched inclusion supramolecular polymer composed of amylose-PLLA inclusion complexes.476 The product formed an ion gel with an ionic liquid, 1-butyl-3-methylimidazolium chloride, which was further converted into a hydrogel by exchange of dispersion media. The lyophilization of the hydrogel resulted in the fabrication of a cryogel with a porous morphology. The poly(L-lysine) moiety with pendant maltooligosaccharide primers and cholesterols formed positively charged polypeptide nanogels (∼50 nm) via self-assembly in water.477 α-Glucan phosphorylase-catalyzed enzymatic polymerization of Glc-1-P on the nanogel produced amylose-conjugated nanogels (Scheme 39). The elongation of the saccharide chain shielded the positive charge of the nanogels. A series of amylose-based star polymers (1, 2, 4, and 8 arms) was synthesized by α-glucan phosphorylase-catalyzed enzymatic polymerization using Glc5-functionalized PEG.478 The 8-arm primer acted as a gelator when triggered enzymatically. Star polymers with DP = ca. 60 per arms served as allosteric multivalent supramolecular hosts. 5.1.2. Cellodextrin Phosphorylase. Cellodextrin phosphorylase is the enzyme that catalyzes phosphorolysis of oligoβ(1→4)-glucans (cellooligosaccharides) larger than cellobiose 2340


Chemical Reviews

Review

Scheme 39

Scheme 40

to produce Glc-1-P.479 In addition, the reversible cellodextrin phosphorylase-catalyzed glucosylation using Glc-1-P as a glycosyl donor and cellobiose derivatives as glycosyl acceptors successively occurs to obtain cellooligosaccharides (Scheme 40).480 When cellobiose was used as a glycosyl acceptor, various cellooligosaccharides ranging from water-soluble products to crystalline insoluble precipitates were yielded depending on the concentration of the acceptor. Although glucose had been believed to not act as a glycosyl acceptor for the cellodextrin phosphorylase catalysis, cellodextrin phosphorylase-catalyzed reaction using glucose as a glycosyl acceptor gave cellooligosaccharides with an average DP of 9.481 β(1→3,1→4)-Oligosaccharides and thiooligosaccharides were prepared by cellodextrin phosphorylase-catalyzed chainelongation using Glc-1-P donor and β(1→3)-linked oligosaccharide acceptors.482,483 Cellobiosylated dimer and trimer and polyamidoamine (PAMAM) dendrimers were also used as glycosyl acceptors for cellodextrin phosphorylase-catalyzed reaction to give the corresponding materials containing cellooligosaccharides at chain ends.484 5.1.3. Glucansucrase and Fructansucrase. The other glycosyltransferases, that have been used as catalyst for practical synthesis of polysaccharides, are glucansucrase and fructansucrase.370,371 These enzymes catalyze in transfer of either a

glucose or a fructose moiety of sucrose to produce glucans or frucutans of different types with respect to glycosidic linkages and side chains. The simplified reaction manners are represented as follows. Glucansucrases: n Sucrose → Glucan + n Fructose

Fructansucrases: n Sucrose → Fructan + n Glucose

In accordance with the reaction manner, typically dextran-, mutan-, alternan-, reuteran-, and amylosucrases (EC 2.4.1.5, 2.4.1.5, 2.4.1.140, 2.4.1.5, and 2.4.1.4) produce dextran (α(1→ 6)-glucosidic linkage), mutan (α(1→3)−), alternan (alternating α(1→3)- and α(1→6)−), reuteran (α(1→4)- and α(1→ 6)−), and amylose (α(1→4)−), respectively (Figure 15).485−488 Amylosucrase (EC 2.4.1.4) is the most extensively studied glucansucrase.489−491 For example, the catalytic properties of the highly purified amylosucrase from Neisseria polysaccharea were characterized.492 Consequently, it was revealed that in addition to the amylose synthesis, several reactions occurred by the enzymatic catalysis in the presence of sucrose alone, which are sucrose hydrolysis, maltose, and maltotriose synthesis by 2341


Chemical Reviews

Review

geothermalis, followed by spontaneous self-association.498 Single-walled carbon nanotubes (SWCNTs) were incorporated into the amylose microbeads during the enzymatic synthesis through a self-assembly process via hydrophobic interaction between amylose and SWCNT. Amylose magnetic microbeads were also prepared by the similar enzymatic approach in the presence of iron oxide nanoparticles.499 The produced microbeads had a well-defined spherical shape and exhibited excellent magnetic responses and dispersibility in aqueous solutions. Another glucansucrase, dextransucrase, was employed as the catalyst for the enzymatic synthesis of spherical dextran nanoparticles. Inclusion/complexation capabilities of the produced nanoparticles toward a hydrophobic nutraceutical, isoflavone genistein were evaluated.500 Two inclusion methods were employed; DMSO-water and acidification. Optimization of the inclusion processes led to the production of nanosized dextran particles containing genistein. Fructansucrases transfer the fructose units of sucrose onto polysaccharides or appropriate acceptors with release of glucose.370,371 Fructans, thus produced, are either inulin composed of β(2→1)-linked fructose residues by inulosucrase (EC 2.4.1.9) catalysis and levan composed of β(2→6)-linked fructose residues by levansucrase (EC 2.4.1.10) catalysis (Figure 15).487,488 Sucrose analogues with a similar glycosidic linkage to sucrose have been used for the synthesis of new polyand oligosaccharides by fructansucrase catalysis.370,501−504 For example, a wide range of fructansucrases recognize most of the sucrose analogues, such as those which were composed of galactose, mannose, xylose, fucose, and rhamnose in place of glucose, giving rise to novel poly- and oligosaccharides (Scheme 41).505,506 5.1.4. Branching Enzyme. The α(1→6)-glucosidic branches in amylopectin and glycogen are constructed by the action of BE catalysis. This enzyme catalyzes cleavage of a glucosidic linkage in an α(1→4)-glucan donor substrate and subsequently transfer of the nonreducing end of the produced fragment to the C-6 position of an internal glucose residue in an α(1→4)-glucan acceptor to form an α(1→6)-glucosidic branch.507 Depending on its source, BEs have a preference for transferring different lengths of glucan chains.508−510 As aforementioned, a highly branched cyclic dextrin (GD) was produced from amylopectin by the cyclization reaction of BE catalysis (Figure 8A).399−401 In this molecule, α(1→4)-glucan chains form highly branched structures by interlinked α(1→6)glucosidic linkages, which are mostly originated from the heavily branched region of amylopectin. Furthermore, one α(1→6)-glucosidic linkage takes part in the cyclic structure, which is newly formed by the BE-catalyzed reaction.

Figure 15. Typical polysaccharide structures obtained by glucan- and fructansucrases.

successive transfers of the glucose moiety of sucrose onto the released glucose, and finally turanose and trehalulose synthesis obtained by glucose transfer onto fructose. Three different glucosyltransferases, GTF-S1, GTF-S3, and GTF-S4, which were isolated from three strains of Streptococcus sobrinus-K1-R, −6715−13−201, and −6715−13−27, respectively, were used to synthesize, from sucrose, water-soluble α-glucans of quite different structure.493 When glycogen was used as a glycosyl acceptor for the amylosucrase catalysis, the sucrose hydrolysis decreased strongly with increasing the concentration of glycogen, as did oligosaccharide synthesis, by glucose transfer onto glucose and fructose.494 The glucose units consumed were then preferentially used for the chain-elongation from glycogen. Moreover, when various polysaccharides were tested as an acceptor for the amylosucrase catalysis, the chain-elongation proceeded only on the polysaccharides with α(1→4)- or α(1→ 4)- and α(1→6)-linkages.495 Recombinant amylosucrase was used to synthesize amylose from sucrose without use of a glycosyl acceptor.496 The products had the DP of 35−58. By changing only the initial sucrose concentration, it was possible to obtain amyloses with different morphology and structure. The recombinant amylosucrase was also used for the chain-elongation in the presence of glycogen as an acceptor.497 The morphology and structure of the resulting insoluble products were strongly dependent on the initial sucrose/glycogen ratio. For the lower ratio, all glucose molecules produced from sucrose transferred onto glycogen, giving rise to a slight elongation of the external chains and their organization into small crystallites at the surface of the glycogen particles. With a high initial sucrose/ glycogen ratio, the external glycogen chains were enzymatically extended, leading to dendritic nanoparticles with a diameter 4− 5 times that of the initial particle. Amylose microbeads were prepared by the enzymatic reaction of sucrose catalyzed by amylosucrase from Deinococcus Scheme 41

2342


Chemical Reviews

Review

The in vitro synthesis of amylopectin- or glycogen-like branched polysaccharides was performed by a tandem reaction by α-glucan phosphorylase and BE.369,511−517 For example, an enzymatically synthesized glycogen was obtained by the combined action of α-glucan phosphorylase and BE on Glc1-P in the presence of an adequate primer (Figure 16). The

Scheme 42

Figure 16. Enzymatic synthesis of glycogen by combined use of αglucan phosphorylase and BE.

molecular weight and branching pattern of the product are controlled by the Glc-1-P/primer ratio and by the relative BE/ α-glucan phosphorylase activity ratio, respectively. The branched glucans produced by high BE/phosphorylase activity ratios had more frequently branching points than those produced by low BE/phosphorylase ratios.

hydroxybutyryl-CoA (3HB-CoA) substrate by the (R)-specific acetoacetyl-CoA reductase with simultaneous oxidation of nicotinamide adenine dinucleotide phosphate (NADPH) to NADP+. This substrate is the direct precursor of P3HB and subject to the PHB synthase-catalyzed reaction. Gerngross and Martin first achieved and demonstrated the in vitro synthesis of P3HB in aqueous solution.523 The resulting P3HB had the higher molecular weight than that of in vivo synthesized P3HB. This is probably due to lack of a chain termination of the P3HB polymerization under the in vitro conditions, according to the manner of a living polymerization. The following studies also reported that the molecular weight of the P3HB produced by PHB synthase was inversely proportional to the monomer/ enxyme molar ratio, suggesting that the polymerization proceeds in a living fashion.524,525 Copolymerization of 3HBCoA and 3-hydroxypropyl-CoA catalyzed by PHB synthase was also conducted to produce random copolyesters.526 Sequential copolymerization in the same system produced the block copolymers. Kinetic studies in in vitro polymerizations catalyzed by PHA synthases have been demonstrated to discuss the detailed reaction mechanisms and fashions.527−529 PHA synthases using in vitro directed evolution and site-directed mutagenesis have further facilitated the synthesis of PHA and copolymers with novel material properties by broadening the spectrum of monomers available for PHA biosynthesis.530,531 For example, PHB synthase attached to gold nanoparticles (AuNP) produced PHB upon the addition of 3HB-CoA and then coalesced to form micrometer-sized AuNP-coated PHB granules.532

5.2. Acyltransferases

Acyltransferases (EC 2.3) are enzymes that catalyze reactions in which an acyl group is transferred from one compound (acyl donor) to another (acyl acceptor).355 The enzymes have been used to synthesize biological polyesters with properties comparable or even exceeding synthetic polymers. Crosslinking and modification of proteins are also available by transglutaminase-catalyzed acryl transfer reaction from glutamine residues of acyl donors to amino groups of acyl acceptors (transamidation). 5.2.1. Polyester Synthase. Polyhydroxyalkanoates (PHAs) are biological polyesters which are produced by a wide variety of bacteria as osmotically intercellular energy and carbon storage materials.518,519 PHAs are generated in almost all bacteria under nutrient-limited growth conditions when a carbon source is readily available. Polyhydroxyalkanoate synthase (PHA synthase, EC 2.3.1.class) is responsible for the polymerization in the production of PHAs in vivo because it catalyzes stereoselective conversion of (R)-hydroxyacyl-CoA substrates (monomers) to PHAs with the concomitant release of CoA.520 As the representative in vivo metabolic route (Scheme 42),521,522 the biosynthesis of poly(3-hydroxybutyrate) (P3HB) occurs through the condensation of two acetyl-CoA molecules by the β-ketothiolase catalysis, giving rise to the formation of acetoacetyl-CoA, which is then reduced to the (R)-32343


Chemical Reviews

Review

Scheme 43

improve functional properties and to provide tissue engineering materials.544−554 TGase has also been employed for improving the properties of protein-based fabrics such as wool.555,556 Moreover, TGase was employed for grafting/coating of wool fabrics with silk sericin or keratin, resulting in increased bursting strength and softness, and reduced felting shrinkage.557,558

Because of advantageous potentials of PLA, such as that is a biobased practical plastics used for commodity applications including food and stationary packaging, furniture, and clothing,533 the direct PLA production using the PHA biosynthetic system has attracted research interest. The possibility for the production of lactyl-containing PHA has been suggested by the examination of in vitro substrate specificities of natural PHA synthases using various HACoAs.534,535 Since natural PHA synthases have extremely low activity toward lactyl-CoA, screening for PHA synthases efficiently accepting lactyl-CoA as a substrate has been undertaken.536−538 For example, a copolymerization method using an LA-polymerizing enzyme (LPE) was developed, which was engineered from the PHA synthase from Pseudomonas sp. 61-3 by evolutionary engineering. 539 Two mutations, Ser324Thr and Gln480Lys, corresponding to those of LPE were introduced into PHA synthase (PhaC1SG) to evaluate the potential of the resulting protein as a “thermostable LPE”. The mutated PhaC1SG [PhaC1SG(STQK)] showed high thermal stability in synthesizing P(LA-co-3HB) in an in vitro reaction system under a range of high temperatures. 5.2.2. Transglutaminase. Transglutaminases (TGase, EC 2.3.2.13) belong to a class of enzymes known as aminoacyltranferases which catalyze calcium-dependent acyl transfer between peptide-bound glutamine residues as acyl donors and peptide-bound lysine residues as acyl acceptors, leading to the formation of intermolecular ε-(γ-glutamyl)lysine cross-links (Scheme 43).540 The transamidation is initiated by the nucleophilic attack from a thiol group of cysteine residue at the active site of TGase to the γ-carboxyamide group on the donor substrate, leading to a loss of ammonia and formation of a thioester intermediate. The acyl group of the thioester is then transferred to an amino group on the acceptor substrate to form an amide linkage.541 TGase recognizes a broad range of the primary amine-substituted acyl-acceptor substrates, whereas recognition of the acyl donor substrate is restricted to the γcarboxyamide of glutamine residue.542 For example, primary alkylamines can be used as acceptor substrates, which allowed the selective alkylation of proteins via their accessible glutamine residues.543 TGase has been used to catalyze cross-linking of several proteins, such as collagen, gelatin, casein, and their hybrid systems with poly/oligosaccharides, PLLA, and nanoclay, to

6. HYDROLASES 6.1. General Introduction of Hydrolases

Hydrolases (EC class 3) are naturally occurring macromolecular organic catalysts that cleave bonds of substrates in the presence of water molecule (H2O). The general scheme of hydrolase-catalyzed reactions can be described in Scheme 44, where substrate molecules and the corresponding hydrolyzates are denoted by A−B and A−OH and H−B, respectively (Scheme 44). Scheme 44

The cleavage is different from hydration reactions as in the addition of water to a carbon−carbon double (CC) bond to give a HOC−CH moiety because the hydrolysis of substrate A−B affords two chemical species, A−OH and H−B, by splitting parts A and B. Many naturally occurring organic compounds possess a carbon−oxygen bond or a carbon− nitrogen bond. For example, glycosides have an acetal structure [(RO)(R′O)CHC−] and triglycerides include carboxylic ester moieties (RCOOCH2−). These carbon-heteroatom bonds are readily hydrolyzed by an acid catalyst as the result of the protonation of the lone pair of electrons on the oxygen or nitrogen atom of the substrates. Thanks to these intrinsic structure designs by nature, many polymeric compounds can be decomposed by the action of hydrolases. A typical example can be seen in the biodegradation of naturally occurring polyacetals, polysaccharides, to monosaccharides catalyzed by glycosidases, which greatly contributes to the carbon cycle on earth.559 According to the principle of microscopic reversibility, hydrolases can catalyze not only hydrolysis but also the reverse reaction, dehydrative condensation, through the same transition 2344


Chemical Reviews

Review

concentration, (2) decreasing the amount of water, and (3) removing the final product from the reaction system, for example, by precipitation or by extraction.561 However, the yield of the dehydrative condensation products would normally be low even if the reactions were carried out under a high substrate concentration, which can be explained by the large negative values of standard Gibbs energy formation of hydrolysis. The addition of an organic solvent or the operation of an enzymatic reaction in organic solvents can enhance the dehydrative condensation toward the products by decreasing the amount of water. However, it is not recommended to use organic solvents that may increase the environmental stress of the synthetic process. The removal of the products by precipitation or extraction would also require special equipment such as packed columns with activated charcoal for absorption of the resulting products.562 In addition to these thermodynamic methods, there is another promising technique to increase product yield through kinetic control. One solution is to introduce a leaving group X on the substrate. By replacing the OH moiety of A−OH by an appropriate leaving group X, a variety of activated donor substrates A−X can be designed and chemically prepared for chemo-enzymatic synthesis of glycosides, esters, and amides, catalyzed by glycosidase, esterases, and proteases, respectively. The general scheme for hydrolase-catalyzed reactions by using the activated substrate is described by Scheme 47. In this

state. The use of hydrolase as catalysts for producing macromolecular organic materials is, therefore, commonly used in modern synthetic chemistry and engineering and has led to the creation of new chemical industries that contribute to our society from the viewpoint of green and sustainable process.52,71 Many organic compounds can be transformed by the action of hydrolases in nature. Thus, there are almost unlimited possibilities for the combination of A and B in Scheme 44. The combination of A and B is summarized in Table 2 together with the corresponding hydrolases. Table 2. Classification of Typical Substrates A−B in Scheme 44 by the Cleaved Moieties and the Corresponding Hydrolases A−B (cleaved moiety)

A

B

hydrolase

carboxylic ester phosphate glycoside nucleoside carboxylic amide

RCO ROPO(OH) sugar sugar RCO

OR′ OR′ OR NRR′ NHR′

esterase, lipase phosphatase glycosidase DNAglycosylase protease

In general, the hydrolysis of organic compounds A−B proceeds via a cationic chemical species denoted by A+ as a result of the elimination of B− from A−B. In other words, the hydrolytic reaction of A−B consists of the following two kinds of elemental reactions (1) and (2) (Scheme 45).

Scheme 47

Scheme 45 reaction, the water, which is the byproduct of dehydrative condensation in Scheme 46, is replaced by an acid XH. This is a typical example of transfer reactions where part A is transferred to part B, liberating XH. Another solution for shifting the equilibrium under kinetically controlled conditions is to use a substrate with higher potential energy. For example, by distorting the conformation of substrates toward the structure of the transition state, it becomes possible to lower the activation energy, which can promote the reaction effectively in the direction of product formation. In fact, many kinds of activated substrates have been developed and employed in the chemo-enzymatic synthesis of glycosides, esters, and amides.87 The following sections deal with synthesis of various macromolecules28,29,35,41,46,48,57,58,60 based on enzymatic condensations of activated substrates having a leaving group X as well as enzymatic addition of transition-state analogue substrates (TSAS) catalyzed by glycosidases,563 esterases,564 and proteases.565

Here, the water molecule (H2O) is divided into OH− and H , indicating that the overall hydrolysis reaction can be classified as an acid/base reaction. In fact, most reported mechanisms for hydrolyses catalyzed by hydrolases involve the formation of a cationic species A+ or its chemical equivalent as an intermediate. For example, the hydrolysis of esters (A = RCO, B = OR′) catalyzed by esterases takes place via an activated ester as an acyl cation equivalent (A−L, L = leaving group). Hydrolysis of glycosides (A = sugar, B = OR) catalyzed by glycosidases proceeds via an oxocarbenium ion or its chemical equivalent, a glycosyl ester intermediate.560 Hydrolase-catalyzed dehydrative condensations can be regarded as the reverse reaction of the hydrolytic process catalyzed by hydrolases (Scheme 46). +

Scheme 46

6.2. Glycosidases

6.2.1. Glycosidase Catalysis for Polycondensations. Glycosidase-catalyzed polycondensations are regarded as a repeating multistep process of a dehydrative condensation reaction between the hemiacetal moiety and one of the hydroxy groups in another saccharide unit. As mentioned in the previous section, in the synthesis of oligo- and polysaccharides, the direct dehydration between each monosaccharide unit in aqueous media is thermodynamically unfavorable. It is absolutely necessary to activate the anomeric carbon atom of the saccharide unit by introducing an appropriate leaving group. As a result of the introduction of the leaving group, the partial

Since enzymatic reactions are usually carried out in aqueous solutions to dissolve enzyme catalysts that are water-soluble polypeptides, the equilibrium of the reaction in Scheme 46 shifts toward the direction of starting materials, A−OH and H− B. Therefore, the synthesis of target molecules A−B based on the reverse reaction of hydrolysis catalyzed by hydrolases must be done under special reaction conditions to obtain the product A−B. Generally, the position of the equilibrium can be shifted toward product formation by (1) increasing substrate 2345


Chemical Reviews

Review

Scheme 48

Figure 17. Mechanism of hydrolase-catalyzed transglycosylation of glycosyl donor having a leaving group X to an alcohol ROH (glycosyl acceptor).

charge (δ+) of the anomeric carbon is increased and the nucleophilic attack of a hydroxy group of another monosaccharide unit occurs more smoothly. Glycosylation reactions are achieved by reacting a glycosyl donor with a glycosyl acceptor (Scheme 48). A glycosyl compound that provides the nonreducing end of the product by cleaving the bond between the anomeric carbon and the leaving group X is called a “glycosyl donor”. The saccharide moieties that include a nucleophilic hydroxy group and supply the reducing end of the product is called a “glycosyl acceptor”. Note that in glycosylation chemistry, the terms “donor” and “acceptor” are used in a different manner from those terms in organic chemistry; in organic chemistry, a donor and an acceptor correspond to a nucleophile and an electrophile, respectively, because the terms donor and acceptor are defined according to the provision of an electron pair. In Scheme 48, regio- and stereoselective formation of a disaccharide having a β(1 → 4) glycosidic linkage by the condensation of a glycosyl donor and a glycosyl acceptor is shown. This is a typical example of condensation reactions in which a small molecule XH is liberated during glycosidic bond formations. To achieve an efficient chemo-enzymatic glycosylation, it is necessary to select an appropriate glycosyl donor, a glycosyl acceptor, an enzyme catalyst, and reaction conditions that can include pH, temperature, and solvent. In glycosidase-catalyzed glycosylations, a glycosyl donor having a leaving group X is first incorporated into the catalytic site of the enzyme and then immediately converted into a glycosyl-enzyme intermediate by liberating the leaving group X (Figure 17). The structure of the intermediate can be expressed by either a covalent glycosyl

ester or an oxocarbenium ion (structure not shown). Two kinds of acidic amino acid side chains have been found to be critical in the formation of the intermediate. One of these acts as a proton donor, cleaving the C−X bond in the glycosyl donor, whereas another acidic amino acid acts as a nucleophile to generate a glycosyl ester intermediate or stabilizes the oxocarbenium ion. The acidic amino acid that triggers the cleavage of the C−X bond by protonation is normally glutamic acid or aspartic acid.566 If the resulting glycosyl enzyme intermediate reacts with a water molecule, the product will be a hemiacetal (hydrolyzate). In transglycosylation, the resulting glycosyl-enzyme intermediate is attacked by a hydroxy group of the glycosyl acceptor which is located in the acceptor site, leading to the formation of the glycosidic bond between the glycosyl donor and glycosyl acceptor. Glycosyl fluorides, which are sugar derivatives whose anomeric hydroxy group is replaced by a fluorine atom (X = F in Scheme 48 or Figure 17) are interesting sugar derivatives because fluorine has the smallest covalent radius among the halogens and the largest electronegativity among all elements.567 Glycosyl fluorides have mainly been studied in light of their physical and biological aspects.568 Ever since glycosyl fluorides were found to be recognized by glycosidase,569 numerous studies on the interaction of glycosyl fluoride and enzymes have been reported.570 On the other hand, in synthetic chemistry, the use of glycosyl fluoride as a glycosyl donor has the following advantages. First, the size of fluorine atom is comparable to that of a hydroxy group such that it can be accepted by the active site of a glycosidase enzyme. Second, of glycosyl halides, only the glycosyl fluoride is stable in an unprotected form due to the large bond-dissociation energy of 2346


Chemical Reviews

Review

the C−F bond (552 kJmol−1), which is necessary for most of enzymatic reactions to be carried out in aqueous media. The higher efficiency of utilizing glycosyl fluorides as glycosyl donors can also be rationalized by considering the number of lone pairs on the fluorine atom (Figure 18). The conventional

retained as the β-type via a doubly inverting process at the anomeric carbon atom of the β-glucosyl donor. Glycosidases are classified into two groups, exo- and endotype enzymes. Exo-type glycosidases have a pocket in their catalytic site and are used for enzymatic syntheses of glycosides having a small molecular structure size. Endo-type glycosidase, whose shape at the catalytic domain looks like a cleft, show high catalytic activity for enzymatic syntheses of polysaccharides. As mentioned above, the formation of only a single isomer can be realized perfectly regardless of the class of enzyme employed; both exo-type and endo-type glycosidases catalyze a completely stereoselective glycosylation. On the other hand, the regio-selectivity of enzyme-catalyzed reactions depends on the class of glycosidases. Exo-type glycosidases normally show lower regio-selectivity toward an acceptor, giving rise to a mixture of regio-isomers (Figure 19A).561 In the case of endo-type glycosidases, high regioselectivity can be achieved. Since both the reducing end of the glycosyl donor and the nonreducing end of the glycosyl acceptor are strictly recognized by the −1 subsite and +1 subsite of endo-type glycosidases, respectively, the hydroxy group that will be incorporated into the resulting glycosidic bond can be located in a suitable position at the +1 subsite of the enzyme to attack the anomeric center located in the −1 subsite (Figure 19B). Consequently, perfect regio-selectivity can be guaranteed concerning the resulting glycosidic bond. This is one of the main reasons why endo-type glycosidases have been extensively used as catalysts for synthesis of polysaccharides where perfectly controlled glycosylation reactions must occur repeatedly. Taking the characteristics of both glycosyl donor and hydrolase into consideration, the combined use of a glycosyl fluoride as monomer and an endo-type glycosidase as catalyst has now become the most favorable method to achieve a polysaccharide synthesis.360 Finding an appropriate combination of a glycosyl fluoride and an enzyme catalyst is, therefore, key in designing a method for preparation of polysaccharides. For the purpose of finding the best combination of glycosyl fluoride donor and the corresponding glycosidase, a novel enzyme assay for screening glycosidases has been developed by using glycosyl fluorides as enzyme substrates.571 The method is based on the color change caused by the complex formation of fluoride ion and lanthanum-alizarin complexone (La3+-ALC). The assay has a much higher sensitivity compared with the conventional methods using p-nitrophenyl glycoside as a screening substrate. According to this enzyme assay screening method, it is possible to find a suitable combination of glycosyl fluoride donor and the corresponding hydrolase by employing the glycosyl donor itself for an enzymatic transglycosylation.

Figure 18. (A) Visualization of the carbon−fluorine bond of βglucosyl fluoride from the front of C1. (B) Visualization of the carbon−oxygen bond of p-nitorophenyl β-glucoside from the front of C1. Only one conformation is shown for each compound.

glycosyl donors such as p-nitrophenyl glycosides have two lone pairs on the anomeric oxygen atom (Figure 18B), whereas the anomeric fluorine atom possesses three lone pairs (Figure 18A). Furthermore, there is the potential risk for the lone pairs of p-nitrophenyl derivatives not to be protonated effectively, because the position of the lone pairs may not be oriented in the proper position when influenced by the location of the bulky p-nitrophenyl moiety in the active site of the enzyme. The position of the lone pairs on the fluorine atom has considerable flexibility, which contributes to an efficient protonation from the acidic amino acid in the active site of the enzyme. In conclusion, the selection of glycosyl fluorides as glycosyl donors for glycosidase-catalyzed transglycosylation is quite reasonable, taking into consideration their reactivity and stability. Glycosidases show perfect stereoselectivity in transglycosylation reactions, giving rise to only one isomer (α-isomer or βisomer) because the formation of the undesired isomer becomes impossible due to steric hindrance caused by the wall of amino acid residues in the active site of the enzyme. For example, in the case of β-glucosidase, the attack of the hydroxy group takes place from the β-face of the glucose, leading to stereoselective formation of a β-glucosidic bond. Consequently, the stereochemistry of the anomeric center of the product is

Figure 19. (A) Exo-type glycosidases have a pocket in their catalytic site. (B) Endo-type glycosidases have a cleft where both a glycosyl donor and a glycosyl acceptor are strictly recognized by the corresponding subsites. 2347


Chemical Reviews

Review

In the following sections, several enzymatic polycondensation reactions catalyzed by glycosidases will be introduced by the type of target polysaccharides.360 Scheme 49 shows the Scheme 49

general scheme for the enzymatic polymerization of glycosyl fluoride monomers catalyzed by a glycosidase to give the corresponding polysaccharides having a β(1 → 4) glycosidic bond. 6.2.2. Cellulase. 6.2.2.1. Cellulose Synthesis. Cellulose, which is the most abundant organic compound on earth, has a linear structure of a β(1 → 4) linked D-glucose repeating units. It is one of the three major structural components of the primary cell walls of green plants, along with hemicellulose and lignin. In nature, cellulose is biosynthesized by the polycondensation of uridine diphosphate glucose (UDP-glucose) catalyzed by cellulose synthase mainly at the plasma membrane of plants. In spite of the importance of cellulose, an in vitro chemical synthesis of cellulose had not been achieved in the first challenge of the task in 1941; the in vitro cellulose synthesis has been a central problem in polymer chemistry.572 In 1991, however, the first in vitro synthesis of cellulose was achieved via the polymerization of β-cellobiosyl fluoride monomer catalyzed by cellulase, a hydrolysis enzyme (R1 = R3 = CH2OH, R2 = OH in Scheme 49).38,573,574 Cellulase catalyzed the in vitro bondformation reaction and produced “synthetic cellulose” having a perfectly controlled β(1 → 4) glycosidic structure with a degree of polymerization (DP) value around 22. During the polymerization, the glycosidic linkage forming reaction was repeated, and hence, the cellobiosyl moiety behaved as a glycosyl donor as well as a glycosyl acceptor. One of the most interesting differences between this enzymatic polymerization and the polymerization via biosynthetic path is that the cellulase-catalyzed polycondensation used a cellobiose derivative as a monomer, rather than a glucose derivative, taking the symmetry of cellulose crystal into consideration. The smallest unit of the cellulose repeating unit structure is “cellobiose” because cellulose has a 2-fold screw axis due to the two kinds of intramolecular hydrogen bonds between C3-OH with endocyclic oxygen and C6-OH with the C2-OH (Figure 20). It was postulated that cellobiose moiety would be a preferable substrate since it could be recognized by the catalytic site more strongly than a glucose derivative. Furthermore, from the viewpoint of supramolecular interaction between the substrate and amino acids, disaccharide substrates are preferable for the processive movement of chain elongation along the cleft of cellulase. In fact, it is to be noted that disaccharide fluoride monomers were widely used, and the disaccharide structure was confirmed effective.575,576 A monosaccharide fluoride was not a good substrate for the polysaccharide synthesis.577 The characteristic feature of synthetic cellulose formation in vitro is that a single glucan chain elongates by interacting with other growing glucan chains. These dynamic events via a nonbiosynthetic path can be realized only by the enzymatic polymerization technique where unprotected glucan chains propagate in aqueous media. These dramatic phenomena

Figure 20. Basic expression of 2-fold screw axis which corresponds to the crystal structure of cellulose.

prompted us to consider a polymer−polymer interaction during polymer chain elongation in addition to the conventional monomer-polymer interaction, monomer-catalyst interaction, and polymer-catalyst interaction in the field of supramolecular chemistry (Figure 21).56

Figure 21. Dynamic supramolecular polymer−polymer interaction during the chain elongation catalyzed by enzyme.

Cellulose forms typically two types of allomorphs of highorder molecular structure through self-assembly. One form is thermodynamically metastable cellulose I, in which cellulose chains are aligned in parallel. The other form is thermodynamically stable antiparallel cellulose II. Notably, naturally occurring cellulose forms the less stable cellulose I crystalline structure. In vitro, crystalline structure of cellulose synthesized via cellulasecatalyzed polymerization was of the cellulose II structure with crude enzyme578 and of cellulose I structure with purified enzyme surprisingly. Metastable cellulose I could be formed due to a kinetically controlled process.579,580 This was the first example of cellulose I formation via a nonbiosynthetic pathway. Such a control in high-order molecular assembly during polymerization was not reported before, therefore, a new concept of “choroselective polymerization” was proposed.73 The term “choros” has its origin in a Greek word which means “space”. The self-assembling process of synthetic cellulose during crude cellulase-catalyzed polymerization was investigated in detail in real time and in situ by a combined small-angle scattering (SAS) methods, together with wide-angle X-ray scattering (WAXS) and field-emission scanning electron microscopy (FE-SEM). The aggregation of the synthetic cellulose was observed and associated with characteristic lengths larger than 200 nm in aqueous media. Further, cellulose molecules created at each active site of enzymes associate 2348


Chemical Reviews

Review

cellulose by the immobilized EGII(core2H) was particularly high that can be attributed to the high density with the horizontally immobilized EGII(core2H) on the flat gold substrate. The welloriented endoglucanase should help crystallization of synthesized cellulose. EGII(core2H) was piled up on gold via NTA-NiHis-tag linkage, where EGII(core2H) was cross-linked with each other in the vertical direction on gold substrate.588 The highly crystalline cellulose was synthesized also by this cross-linked enzyme on gold substrate because of high local concentration of EGII(core2H) (Figure 22).

themselves around the enzyme associations into cellulose aggregates having surface fractal dimensions Ds, increasing from 2 (smooth surface) to 2.3 (rough surface with fractal structure) as the reaction progressed, extending over a surprisingly wide length scale ranging from ∼30 nm to ∼30 μm with 3 orders of magnitude. The construction of this unique self-assembly can be explained by an extremely large number of cellulose molecules repeatedly created at the active site of cellulase.581,582 Concerning the enzyme catalysts responsible to the polymerization of β-cellobiosyl fluoride, many kinds of cellulases have the possibility of possessing the glucan chain elongating ability. One of these is endoglucanase II (EGII) from Trichoderma viride. EGII is composed of three functional domains: the cellulose-binding domain, the linker, and the catalytic domain. Interestingly, for the enzymatic polymerization of β-cellobiosyl fluoride monomer, the catalytic domain was found to be sufficient. The cellulose-binding domain deficient enzyme is preferable for the enzymatic polymerization because the hydrolytic degradation of the crystalline product can be suppressed. The mutant EGII (EGIIcore), which is deficient of the cellulose-binding domain, was expressed in Saccharomyces cerevisiae.583 The mutant EGIIcore catalyzed the polycondensation of β-cellobiosyl fluoride to give crystalline cellulose of type II, which resisted against enzymatic degradation in the polymerization solution. Another mutant EGII having two sequential catalytic core domains (EGII(core)2) was prepared.584 This mutant EGII(core)2 catalyzed polymerization of β-cellobiosyl fluoride faster than EGIIcore, affording large spherulites. The resulting large spherulites were composed of platelike crystals radiating from the center of the spherulites. On the other hand, fibrous cellulose was produced from cross-linked mutant EGII(core2H) having two hexameric histidine residues (His-tags) in total on both enzyme chain terminals.585 The cross-linking molecule used is bisNTA, which has two nitrilotriacetic acid (NTA) moieties on both terminals of poly(ethylene oxide). EGII(core2H) mutant enzymes were cross-linked with the help of Ni ions through bisNTA. Using the cross-linked EGII(core2H) as a catalyst for enzymatic polymerization of β-cellobiosyl fluoride, the polymerization proceeded extremely fast, and the fibrous cellulose with high molecular weight was produced. Taken together, the configuration of enzymes in vitro synthesis of cellulose seems to influence the morphology of synthetic cellulose as is the case of in vivo synthesis. To obtain an in-depth understanding of in vitro synthesis of cellulose with regard to geometry of the mutant enzymes, EGII(core)2 was immobilized on gold and its hydrolytic activity586 and polymerization activity were analyzed.587 The linker molecules with thiol at one end and NTA at the other were self-assembled on gold, and EGII(core)2 having a His-tag was immobilized on the self-assembled monolayer (SAM) via Ni ion. The hydrolytic activity of the immobilized EGII(core)2 was nearly the same on either anchor molecule. The immobilized EGII(core)2 apparently retained the inherent hydrolytic activity similar to free EGII(core)2. The local high concentration of EGII(core)2 on gold probably promoted successive hydrolysis of the transient products, leading to high hydrolytic activity despite immobilization. Two kinds of the mutant enzymes, EGII(core)2 with one His-tag and EGII(core2H) with two His-tags on both terminals, were immobilized on the NTA-SAM. The crystallinity of the synthesized cellulose by the immobilized EGII(core)2 was higher than that by free EGII(core)2. The crystallinity of the synthesized

Figure 22. Schematic illustration of enzymatic polymerization with using cross-linked enzymes, immobilized enzyme on gold substrate, and cross-linked and immobilized enzymes.588

In principle, the products of hydrolase-catalyzed glycosylations suffer hydrolysis of the formed glycosidic bonds unless the products are removed from the reaction mixture. This inherent problem of product hydrolysis has been solved by using mutant glycosidases (glycosynthases).589 These mutants, which lack the catalytic nucleophile of an acidic amino acid, show no hydrolytic activities because the reactive glycosyl-enzyme intermediate cannot be formed. In these glycosynthasecatalyzed reactions, an α-glycosyl fluoride having the opposite anomeric configuration is used as the glycosyl donor. The glycosyl fluoride having α-configuration can be recognized by the catalytic site of the mutant enzyme as an analogue of the covalent glycosyl-enzyme intermediate. The transglycosylation reactions proceed in excellent yields because the mutant enzymes are hydrolytically inactive toward the glycosides formed. For example, the mutant enzyme from Humicola insolens was capable of transferring disaccharides from αcellobiosyl fluoride, giving rise to cellulose in good yield.575 Another approach of cellulose synthesis was achieved by utilizing a nonactivated disaccharide monomer and a cellulase/ surfactant (CS) complex as catalyst.590 In a nonaqueous medium of dimethylacetamide (DMAc)/LiCl, a cellulosesolubilizing solvent, polymerization occurred at 37 °C to produce cellulose as white powders. The DP value was high (over 100), and the product yield was low (up to 5%). This is a typical example of classical dehydrative polycondensation, affording H2O as a byproduct (X = OH in Scheme 48 and Figure 17). 6.2.2.2. Unnatural β-Glucan Synthesis. A synthetic strategy for hybrid-type oligosaccharides and polysaccharides having an alternating structure composed of a monosaccharide unit ○ and monosaccharide unit ● has been developed by chemoenzymatic procedures (Figure 23).43 In accordance with this principle, it is possible to design two kinds of disaccharide monomers, ○−● * and ●−○ * where ○ and ● denote different monosaccharide units and * means that the anomeric center of these disaccharides are activated by, for example, introducing a fluorine. Because of the wide spectrum in substrate recognition of glycoside hydrolases, synthesis of unnatural polysaccharides composed from different two polysaccharide components could be achieved.591 Such 2349


Chemical Reviews

Review

cellobiosyl fluoride in a buffer/organic solvent afforded a mixture of water-soluble hemithiocellodextrins with DP = 4− 14.596 Although the enzymatic glycosylation of glycosyl fluoride monomers has become a general strong synthetic method for polysaccharide synthesis, the preparation of glycosyl fluorides require several steps, including protection of the hydroxy groups, activation of the anomeric center by introducing chlorine or bromine, nucleophilic replacement by fluorine as a leaving group, and the removal of the protecting groups. These procedures eventually lower the total yields of the glycosyl donor syntheses. In addition, in case the of oligosaccharide donors, the cleavage of the inner glycosidic bonds occurs during the process of glycosyl donor synthesis, affording a mixture of oligosaccharide donors with different molecular weights.597 These demerits of the conventional glycosyl donor synthesis have hampered the application of enzymatic glycosylation to complex target molecules. Recently, one-step preparable glycosyl donors, 4,6-dimethoxy-1,3,5-triazin-2-yl glycosides (DMT-glycosides), have been developed as a new monomer for enzymatic polymerization (Scheme 50). A cellotetraose-backboned hepta-saccharide (XXXG) and a nona-saccharide (XLLG) have directly been converted to the corresponding 4,6-dimethoxy-1,3,5triazin-2-yl derivatives (DMT-β-XXXG and DMT-β-XLLG, respectively) by the action of 4-(4,6-dimethoxy-1,3,5-triazin-2yl)-4-methyl morpholinium chloride (DMT-MM).598 Note that the letter X represents a glucopyranose residue that is substituted with a xylopyranose through an α-1,6 glycosidic bond, and the letter G represents a nonsubstituted glucopyranose residue, and the letter L represents a glucopyranose residue that is substituted with a galactopyranoseβ(1 → 2)xylopyranose through an α(1 → 6) glycosidic bond. The resulting activated oligosaccharide derivatives were found to polymerize catalyzed by an endo-β-1,4-glucanase as catalyst. The polymerization took place in a complete regio- and stereoselective manner, affording non-natural polysaccharides having a XXXG-repeating unit and a XLLG-repeating unit, respectively, in the main chain. 6.2.3. Xylanase, 4-Glucanohydrolase, and Amylase. 6.2.3.1. Xylan Synthesis. Xylan is one of the most important components of hemicelluloses in plant cell walls, being the polycondensation products of xylose connected through a β(1 → 4) glycosidic linkage. A monomer, β-xylobiosyl fluoride, was subjected to polycondensation catalyzed by cellulase (R1 = R3 = H, R2 = OH in Scheme 49). The product was “synthetic xylan” having β(1 → 4) glycosidic linkage, having a Mn of 6.7 × 103

Figure 23. Design of two kinds of disaccharide monomers for synthesis of hybrid-type polysaccharides.

polysaccharides (hybrid polysaccharides) are difficult to synthesize via the biosynthetic path or by the conventional chemical synthesis. The 6-O-methyl-β-cellobiosy fluoride was found to be recognized by cellulase and polymerized, giving rise to the corresponding alternatingly C-6 methylated cellulose derivative (R1 = CH2OMe, R2 = OH, and R3 = CH2OH in Scheme 49).592 The Mn of the product was 3.9 × 103, corresponding to n ∼ 7. Disaccharide monomers, 2′-O-methyl-β-cellobiosyl fluoride and 4-O-(β-mannopyranosyl)-β-glucopyranosyl fluoride, have been polymerized using cellulase as a catalyst in a mixed solvent of acetonitrile-acetate buffer. The polymerization proceeded by a disaccharide unit, giving rise to alternatingly methylated cellooligosaccharides (R1 = R3 = CH2OH, R2 = OMe in Scheme 49) and oligosaccharide having a mannoseglucose unit, respectively.593 These results show that the 6OMe group, 2′-OMe group of the cellobiose moiety, and 2′OH group of the mannose unit do not have large steric hindrances for the substrate recognition. Cellulose−chitin hybrid polysaccharide has also been synthesized based on the same concept.594 GlcNAcβ(1 → 4)Glc-β-fluoride monomer was designed and prepared and polymerized by the action of cellulase (GlcNAc: Nacetylglucosamine). These results clearly show that the −2 subsite and +1 subsite of cellulase can accept even a nonglucotype monosaccharide unit like GlcNAc efficiently. The cellulase-catalyzed glycosylation reaction using glycosyl fluoride donors was extended to a stepwise elongation of glucose unit that was performed via combined use of cellulase and β-galactosidase catalysis. Repetitions of sequential manipulations using β-lactosyl fluoride allowed synthesis of the chainlength controlled unnatural oligosaccharides having a galactose unit at the nonreducing end as well as cello-oligosaccharides.595 The enzymatic polycondensation technique was further extended to a synthesis pseudo-oligosaccharides containing a sulfur atom. An enzymatic polycondensation of 4-thio-βScheme 50

2350


Chemical Reviews

Review

Scheme 51

stereoselective formation of a disaccharide having a β(1 → 4) glycosidic linkage is shown. Since the glycosyl donor possesses an intramolecularly dehydrated structure, the reaction proceeds smoothly via a nucleophilic attack of one of hydroxy groups in the glycosyl acceptor to the anomeric carbon atom of the glycosyl donor, accompanying a ring opening, without liberating any small molecules. This is a typical example of addition reactions where the O− and the H+ from the OH in the glycosyl acceptor binds to glycosyl donor’s anomeric carbon and the heteroatom unit Z, respectively. Sugar oxazolines, which are the intramolecularly dehydrated derivatives of 2-acetamido-2-deoxy sugars (Z = −NC(CH3)O− in Scheme 51), are potentially useful glycosyl donors for enzymatic glycosylation by addition reaction because the oxazoline ring can be opened by the attack of a hydroxy group at the anomeric position, regenerating an N-acetylglucosaminide moiety. The reasoning for using oxazoline derivatives can be shown in the retro-synthetic analysis of a N-acetylglucosaminide unit (Figure 24).604 An N-acetylgluco-

corresponding to the DP value of ∼23.599 The resulting xylan has the same structure as natural xylan extracted from esparto grass.600 6.2.3.2. Unnatural Xylose-Containing β-Glucan Synthesis. A cellulose−xylan hybrid polysaccharide has been prepared.601 There are two possible candidate monomers: Glcβ(1 → 4)Xylβ-fluoride and Xylβ(1 → 4)Glc-β-fluoride according to the concept introduced in section 6.2.2.2 (Figure 23). Both monomers were polymerized by xylanase (3.2.1.32) enzyme catalyst, giving rise to the corresponding cellulose-xylan hybrid type polysaccharide. 6.2.3.3. 1,3−1,4-D-Glucan Synthesis. The combination of a glycosyl fluoride and an endo-type glycosidase can be further applied to preparation of unnatural oligosaccharides having a β(1 → 3) glycosidic bond and β(1 → 4) glycosidic bond aternatingly. A 1,3−1,4-D-glucan 4-glucanohydrolase from Bacillus licheniformis has been shown to catalyze the polycondensation of β-laminaribiosyl fluoride (Glcβ(1 → 3)Glc-β−fluoride) and to lead to alternating 1,3−1,4-β-Dglucotetraose and -glucohexaose.602 6.2.3.4. Maltooligosaccharide Synthesis. Maltooligosaccharide is a glucose oligomer linked through α(1 → 4) glycosidic bonds. An α-amylase (EC 3.2.1.1)-catalyzed polycondensation has been demonstrated to give maltooligosaccharides. An activated substrate of α-D-maltosyl fluoride was designed as a monomer based on the double-displacement transfer mechanism. The reaction was carried out in a methanol−phosphate buffer, affording amylose oligomers up to heptasaccharide.603 The oligomer formation may be ascribed to steric hindrance caused by the helical structure of amylose molecules. 6.2.4. Glycosidase Catalysis for Ring-Opening Polyadditions. Addition reactions are organic transformations where two or more molecules combine to form larger adducts without liberating any small molecules like H2O and HX. The glycosidase-catalyzed polyaddition reaction is regarded as a repeating multistep process of an addition reaction of one of the hydroxy groups in a glycosyl monomer to the anomeric center of another monomer without liberating H2O or HX as a byproduct.52 In principle, it is impossible to construct dehydrated skeletons like O-glycosidic bonds through an addition reaction because the concept of elimination of H2O or HX is not included in any addition reactions. Therefore, to design a glycosylation based on an addition reaction, the use of a glycosyl donor having an already dehydrated moiety like an unsaturated bond or a heterocycle is indispensable. Scheme 51 shows a general scheme of glycosylation using a glycosyl donor containing a heterocyclic moiety denoted by Z (Scheme 51). By using such kind of already dehydrated glycosyl donor, the construction of a dehydrated moiety through an addition reaction becomes possible. In this scheme, a regio- and

Figure 24. (A) Retrosynthetic analysis of N-acetylglucosaminide to give 1,2-oxazoline synthon. (B) Synthesis of N-acetylglucosaminide by the addition of an alcohol to 1,2-oxazoline derivative.

saminide can be retrosynthetically disconnected to give the corresponding sugar oxazolines as a synthon (Figure 24A). On the basis of this retrosynthetic analysis, the real reaction for construction of the N-acetylglucosaminide would be the addition of an alcohol to the anomeric center of the sugar oxazoline derivative (Figure 24B). Another advantage of using sugar oxazolines is that the transglycosylation proceeds smoothly due to its lower activation energy. Sugar oxazolines are bicyclic molecules having a distorted structure where a six-membered pyranose ring and a five-membered oxazoline ring are fused sharing the anomeric and C2 positions. Consequently, sugar oxazolines 2351


Chemical Reviews

Review

Taking the above-mentioned requirements for glycosyl donor and enzyme catalysts into consideration, the use of sugar oxazolines as monomers and endo-type glycosidases that can act as a substrate-assisted catalyst has now become the most promising combination to achieve synthesis of N-acetylglucosamine-containing polysaccharides. In the following sections, several enzymatic polyaddition reactions catalyzed by Nacetylglucosaminidases will be introduced according to the type of target polysaccharides. The general scheme for the enzymatic polymerization of sugar oxazoline monomers catalyzed by an endo-type N-acetylglucosaminidase glycosidase is shown in Scheme 52.55 6.2.4.1. Chitin Synthesis. Chitin is a β(1 → 4) linked Nacetyl-D-glucosamine (GlcNAc) polysaccharide that can be biosynthesized by the polymerization of UDP-GlcNAc as substrate monomer with chitin synthase.606 Chitosan is an Ndeacetylated product of chitin. Chitin is one of the most abundant and widely found polysaccharides in the animal field. Chitin and chitosan show excellent characteristics of biodegradability, biocompatibility, and in particular low immunogenicity. In 1995, the first in vitro synthesis of chitin was accomplished via ring-opening polyaddition of a chitobiose oxazoline monomer catalyzed by chitinase and published in 1996 (R1 = R4 = OH, R2 = R5 = CH2OH, R3 = NHAc in Scheme 52).607−610 The reaction occurs as a ring-opening polyaddition mode and is promoted under weak alkaline conditions (pH 9.0−11.0). The yield of “synthetic chitin” in these works was almost quantitative because the resulting chitin was not hydrolyzed by the chitinase catalyst due to its lower hydrolytic activity under alkaline conditions. The DP value of synthetic chitin was evaluated as 10−20, depending on the reaction conditions.49 In the chitinase catalysis mechanism, two carboxylic acid residues are involved in the catalysis (Figure 26). It is considered that the oxazolinium ion formation is a transition state (or intermediate) in the hydrolysis catalyzed by chitinases in which a “substrate-assisted catalyst” mechanism is involved.605 In the polymerization, the monomer is first recognized, and the nitrogen atom is protonated by the carboxylic acid residue at the donor site of chitinase, forming the corresponding oxazolinium ion. This is because the monomer has already an oxazoline structure. Instead of the water molecule in the hydrolysis, C-4 hydroxyl group of sugar unit located at the acceptor site (ROH in Figure 26) acts as a nucleophile to attack the anomeric carbon from the β-side to open the ring, providing a new glycosidic linkage with βconfiguration. Thus, the monomer acted as a glycosyl donor as well as a glycosyl acceptor, and this glycosidic linkage formation reaction is repeated during the polymerization. The most important point is the structure resemblance of the transition state (or intermediate) involved in both hydrolysis and transglycosylation. From these considerations a new concept

possess higher potential energy compared with the conventional glycosyl donors such as p-nitrophenyl glycoside that decrease the activation energy between the starting glycosyl donor and the product of glycoside. The use of sugar oxazolines will, therefore, be very promising for an efficient chemoenzymatic glycosylation provided that such an artificial substrate with an oxazoline ring is recognized by a naturally occurring hydrolase. Since the target polysaccharides have Nacetylglucosaminde structure, enzymes that accept the Nacetylglucosamine unit at the −1 subsite are reasonable candidates as a catalyst. Chitinases (EC 3.2.1.14) hydrolyze the N-acetylglucosaminide unit of chitin, a β(1 → 4) linked polysaccharide composed of N-acetylglucosamines. Early comparisons of their amino acid sequences have revealed that their catalytic domains could be classified into two categories, families 18 and 19. A novel hydrolysis mechanism of family 18 chitinases that involves an oxazolinium ion intermediate named ‘‘substrate-assisted catalysis’’ has been proposed.605 In accordance with this mechanism, the glycosidic bond is cleaved as follows. First, the oxygen of the glycosidic bond is protonated by the carboxylic acid of an acidic amino acid followed by the formation of the oxazolinium ion intermediate by the nucleophilic attack of the amide carbonyl group to the anomeric center. The resulting oxazolinium ion intermediate is then attacked by water to give the hydrolyzate. The sugar oxazoline derivatives can be regarded as the transition state analogue for the above-mentioned hydrolysis reaction catalyzed by family 18 chitinases. By using these activated substrates as glycosyl donors, an efficient glycosylating process catalyzed by a chitinase can be designed. The activation energy of the conversion reaction from the oxazoline derivative to the glycoside becomes very low compared with the activation energy between the conventional substrate and the product (Figure 25). Therefore, the glycosylation reactions can proceed

Figure 25. Change from the conventional substrate to the transition state analogue substrate (TSAS) for enzymatic glycosylation.

efficiently even when a deactivated enzyme is employed. By using a deactivated enzyme, the side reaction to afford the hydrolyzate does not occur. It should be noted that the usage of the sugar oxazoline as a transition state analogue substrate (TSAS) allows the reaction to proceed only in the direction of the glycosylation reaction while suppressing hydrolysis of the product in aqueous media. Scheme 52

2352


Chemical Reviews

Review

Figure 26. Proposed mechanism for ring-opening addition of sugar oxazoline substrate in the catalytic site of chitinase. Note that two acidic amino acids synergetically act as proton donor and proton acceptor.49

of “transition state analogue substrate” (TSAS) was proposed.607 The organization process of crystalline chitin during the chitinase-catalyzed polyaddition of chitobiose oxazoline monomer was monitored by using phase contrast and polarization microscopy in combination with SEM and TEM (Figure 27).611 Under reaction conditions of pH 10.5 and 30 °C, 25 h were required for the complete consumption of the disaccharide monomer. During the first 30 min, a small number of rectangular platelike solids were observed whose width, height, and length of the plates were 25, 10, and 50 nm to 1 μm, respectively. The electron microdiffraction of the resulting plates clearly showed that the products were the thermodynamically stable form of chitin crystal, α-chitin. Therefore, single crystalline plates of α-chitin were formed, in which polysaccharide chains packed antiparallel and formed intra- and intermolecular hydrogen bonds. The crystal plates grew and stacked on each other as time elapsed (ca. 3h) and shaped into ribbons, followed by the formation of bundlelike assemblies. After 25 h, the texture of the synthetic chitin spherulites were observed by SEM that revealed the diameters of these spherulites to be 20−50 μm. A direct method for synthesis of sugar oxazolines from the corresponding N-acetyl-2-amino sugars in aqueous media has been developed with chloroformamidinium-type dehydrating reagents.612,613 This method was applied to one-pot regio- and stereospecific synthesis of chitoheptaose ((GlcNAc)7) by using chitopentaose ((GlcNAc)5) and chitobiose ((GlcNAc)2) as starting materials. The key intermediate, 1,2-oxazoline derivative of (GlcNAc)5 was transglycosylated to a (GlcNAc)2 acceptor catalyzed by a mutant chitinase with lower hydrolyzing activity.614 6.2.4.2. Unnatural Oligo- and Polysaccharide Synthesis. Thus far, it is known that oxazoline moieties at the reducing end of these disaccharide monomers are indispensable for polymerizations to occur that form oligo- and polysaccharides. However, it was postulated that the structure of the monosaccharide units at the nonreducing end would show

Figure 27. Hierarchy structures of synthetic chitin in vitro formed via enzymatic polymerization. Reprinted from ref 611. Copyright 2000 American Chemical Society.

some flexibility toward the −2 subsite of enzymes. On the basis of this hypothesis, several artificial disaccharide oxazoline substrates have been designed for chitinase-catalyzed synthesis of unnatural oligo- and polysaccharides having an Nacetylglucosamine unit.591 The stepwise elongation of the GlcNAc unit, which was performed via combined use of chitinase and β-galactosidase catalysis, was demonstrated. Repetitions of sequential manipulations, using N-acetyllactosamine (LacNAc) oxazoline derivative, allowed synthesis of unnatural chitooligosaccharides having a galactose at the nonreducing end and chain-length controlled chitooligosaccharides.615,616 Cellulose-chitin hybrid polysaccharide having an alternating Glc and GlcNAc was obtained using chitinase as an enzyme catalyst and Glcβ(1→4)GlcNAc oxazoline as a monomer (R1 = R3 = R4 = OH, R2 = R5 = CH2OH in Scheme 52) .594 The Mn values of the products were 4.0 × 103 and 2.8 × 103, respectively. Despite the high crystallinity of both cellulose and chitin homopolymers, the cellulose-chitin hybrid polysaccharides showed no crystalline structure. Similarly, chitin-xylan hybrid polysaccharide having an alternating GlcNAc and Xyl was prepared from Xylβ(1→4)GlcNAc oxazoline monomer with chitinase catalyst, which generated water-soluble polysaccharide of a mass weight larger than 1.0 × 104 (R1 = R3 = R4 = OH, R2 = CH2OH, R5 = H in Scheme 52).617 This sort of preparation of unnatural hybrid polysaccharide was extended to chitin-chitosan which has a regular alternating sequence of GlcNH2 and GlcNAc (R1 = R4 = OH, R2 = R5 = CH2OH, R3 = NH2 in Scheme 52).618 2353


Chemical Reviews

Review

A structurally well-defined synthetic chondroitin sulfate has been prepared using the hyaluronidase catalyst.631 Among three oxazoline monomers sulfated at C-4 of GalNAc, C-6 of GalNAc, and C-4,6 of GalNAc, the monomer sulfated at C-4 gave the chondroitin sulfate in good yields. The resulting synthetic chondroitin sulfate has the sulfonate group exclusively at C-4 of the GalNAc unit; the purity of the synthetic ChS-A 100% is to be compared with that of natural ChS-A ∼ 80%. The Mn value ranged from 4.0 × 103 to 1.8 × 104. 6.2.5.2. Unnatural Hybrid Type Glycosaminoglycan Synthesis. Hybrid glycosaminoglycans of hyaluronan-chondroitin and hyaluronan-chondroitin 4-sulfate can be obtained by enzymatic polymerization using a hyaluronidase catalyst (Scheme 54).632 N-Acetylhyalobiuronate (GlcAβ(1→3)-

Some chitin derivatives were prepared; new TSAS monomers, 3-O-methyl-chitobiose oxazoline (R1 = OMe, R2 = R5 = CH2OH, R3 = NHAc, R4 = OH in Scheme 52) or 3′-Omethyl-chitobiose oxazoline (R1 = OH, R2 = R5 = CH2OH, R3 = NHAc, R4 = OMe in Scheme 52) gave only oligosaccharides.619 6-O-Carboxymethylated chitobiose oxazoline (R1 = R4 = OH, R2 = CH2OCH2CO2Na, R3 = NHAc, R5 = CH2OH in Scheme 52) and 6′-O-carboxylmethylated chitobiose oxazoline (R1 = R4 = OH, R2 = CH2OH, R3 = NHAc, R5 = CH2OCH2CO2Na in Scheme 52) were polymerized by chitinase, indicating that both monomers were recognized by chitinase.620,621 Fluorine-substituted chitobiose oxazoline derivatives having C-6 fluorine (R1 = R4 = OH, R2 = CH2F, R3 = NHAc, R5 = CH2OH in Scheme 52) were polymerized by the chitinase catalyst, giving rise to white precipitates of structurally welldefined fluorinated-chitins with average molecular weights of Fchitins of 1400−1700.622,623 Chitinase-catalyzed polymerization of a TSAS monomer bearing a bulky N-sulfonate group at the C-2′ position (R1 = R4 = OH, R2 = R5 = CH2OH, R3 = NHSO3Na in Scheme 52) proceeded homogeneously, due to a good solubility of the resulting polysaccharide. Chitinase from Serratia marcescens provided a polysaccharide of Mn = 4180.624 With copolymerization of N-acetylchitobiose oxazoline monomer with N,N′-diacetylchitobiose oxazoline monomer, tailormade synthesis of a chitin derivative with controlled deacetylated extent ranging from 0% to 50% was achieved.625 6.2.5. Hyaluronidase. 6.2.5.1. Hyaluronan and Chondroitin Synthesis. Hyaluronidase (EC 3.2.1.35−36) is an endotype glycoside hydrolase, which hydrolyzes β(1→4) glycosidic linkage between GlcNAc and GlcA of hyaluronic acid. The hydrolysis mechanism of hyaluronidase is considered similar to that of chitinase due to the presence of GlcNAc at the −1 site of the catalytic center.626,627 Therefore, a novel GlcAβ(1→3)GlcNAc oxazoline monomer was designed and its hyaluronidase-catalyzed polymerization was examined (Scheme 53). The polymerization gave rise to hyaluronic acid with a high molecular weight Mn value of 1.74 × 104 at yields greater than 50%.628

Scheme 54

GlcNAc)-derived oxazoline was copolymerized with Nacetylchondrosine (GlcAβ(1→3)GalNAc)-derived oxazolines (R = H in Scheme 54) by the hyaluronidase catalysis at pH 7.5 and 30 °C, giving rise to the corresponding copolymer with a Mn = 7.4 × 103 in a 50% yield. Hyaluronidase-catalyzed copolymerization of monomer with N-acetylchondrosine oxazoline having a sulfate group at C4 on GalNAc (R = SO3− in Scheme 54) produces the corresponding copolymer with a Mn = 1.4 × 104 in a 60% yield. The copolymer compositions can be controlled by varying the comonomer feed ratio. Hyaluronidase catalyzes multiple enzymatic polymerizations with controlling regio- and stereoselectivity perfectly. This behavior, that is, the single enzyme being effective for multireactions and retaining the enzyme catalytic specificity, is unusual, and hence, hyaluronidase can be considered to be a “supercatalyst” (Figure 28).633 6.2.6. Keratanase. Keratanase II obtained from Bacillus sp. Ks36 has been used for the enzymatic polymerizations. Even though the details of the enzyme are still unknown, similar ones obtained from Bacillus circulans KsT202 have been reported.634 The enzyme was shown to hydrolyze keratan sulfate between the 4GlcNAcβ(1→3)Gal structure.635 Keratan sulfate is in the class of glycosaminoglycanes, which have repeated disaccharide structures of β(1→3) linked Galβ(1→4)GlcNAc (LacNAc). Galβ(1→4)GlcNAc(6S) and Gal(6S)β(1→4)GlcNAc(6S) oxazoline monomers and have been designed for polymerizations using keratanase II. These two sulfated monomers were polymerized by keratanase II (3.2.1.103), producing the corresponding keratin sulfate oligosaccharides.636 Notably, keratanase II catalyzed trans-

Scheme 53

Hyaluronidase-catalyzed copolymerizations of monomer combinations of 2-methyl/2-vinyl, 2-methyl/2-ethyl, 2-methyl/2-n-propyl, and 2-vinyl/2-ethyl oxazoline derivatives of hyalobiuronate GlcAβ(1→3)GlcN have been achieved.629 The PH-20 protein, which facilitates penetration for the sperm through the hyaluronan-rich matrix of the oocyte,627 is considered to be the primary enzyme for polymerization. Hyaluronidase is also known to be an in vivo hydrolysis catalyst of chondroitin, cleaving β(1→4) glycosidic linkage. A new oxazoline derivative of GlcAβ(1→3)GalNAc was designed, and its polymerization was examined using the hyaluronidase catalyst.630 The corresponding nonsulfated chondroitin of the Mn value of 5.0 × 103 was obtained. The monomers of 2-ethyl and 2-vinyl oxazoline derivatives or N-propionyl and N-acryloyl derivatives were also polymerizable with hyaluronidase. 2354


Chemical Reviews

Review

Figure 28. A hyaluronidase supercatalyst for the enzymatic polymerization to synthesize glycosaminoglycans. Reprinted with permission from ref 633. Copyright 2006 Wiley.

glycosylation to form a β(1→3)-glycosidic bond and required the 6-sulfate group in the GlcNAc residue. 6.2.7. Endo-N-acetylglucosaminidase: Modification of Polypeptides. The combined use of sugar oxazolines and the corresponding glycosidase has been further applied to the synthesis of glycoprotein having a definite oligosaccharide chain. Endo-N-acetylglucosaminidase (EC 3.2.1.50) from Mucor hiemalis (Endo-M) was found to recognize an oxazoline derivative of Man-GlcNAc disaccharide as a substrate for the transglycosylation reaction in the presence of a GlcNAc acceptor.637 On the basis of these findings, a transglycosylation reaction of disialo-oligosaccharide oxazolines to a glycopeptide and glycoprotein having a definite carbohydrate structure were synthesized catalyzed by a mutant enzyme of Endo-M N175Q (Figure 29).638−640 The preparation of human IgG having a

single glycoform from fucosylated IgG by remodeling using Endo-S has been achieved.641 Endo-N-acetylglucosaminidase from Streptococcus pneumoniae (Endo-D) also recognize fucosylated N-linked oligosaccharides. 6.3. Lipases

Lipase (EC 3.1.1.3) belongs to a hydrolase enzyme, which catalyzes the hydrolysis of fatty acid esters normally in an aqueous environment in living systems (see also the previous section 6.1 for the general introduction). In vitro lipase catalysis was noted in going back to the 1930s,51,83,84 and later it came to the attention of organic chemists.85−87 So far, lipase is the most frequently employed enzyme as catalyst for macromolecular synthesis, which constitutes the very actively studied area in the enzymatic polymerization field for around these three decades as typically seen in the reviews.41,52,60 It should be stressed that lipase-catalyzed macromolecular synthesis involves almost all the characteristics of “green nature” as discussed in section 3. A very important point is that lipase enzymes are readily and widely available; among others, a typical example is Novozym 435, which is commercially available and one of the most often studied enzymes as seen below. Furthermore, renewable resources like biomasses are often used as starting materials, leading to useful macromolecular products. 6.3.1. Synthesis of Polyesters. Since lipase cleaves the ester bond with hydrolysis, the reverse reaction can be catalyzed to form an ester bond. Typically, such reaction gives rise to produce a polyester via a repetitive cleavage-esterification of a cyclic ester monomer, which is a ring-opening polymerization, and via transesterification from two monomers with liberating the other molecule(s), which is a polycondensation.

Figure 29. endo-N-Acetylglucosaminidase-catalyzed synthesis of monodispersed glycoprotein by addition of oligosaccharide oxazoline donor to a protein having a GlcNAc scaffold. 2355


Chemical Reviews

Review

6.3.1.1. Ring-Opening Polymerization. Ring-opening polymerization (ROP) is one of the typical routes to polymers in chemical industries. Enzymatic catalysis has been used for ROP of various cyclic monomers, mainly cyclic esters (lactones) to polyester syntheses. Typical examples of cyclic monomers are shown in Figure 30.

Lipase catalysis also induced ring-opening copolymerization of cyclic monomers. The first example of the enzymatic ringopening copolymerization of lactones was a combination of δVL and ε-CL (Scheme 55B, m = 4); they were copolymerized by lipase PF catalyst to produce the copolyester with a molecular weight of 3.7 × 103. The resulting copolymer was of random structure having both units. Copolymerization of ε-CL with other lactones like 15-pentadecanolactone (PDL) (m = 14 in Scheme 55B), and D-lactide was also achieved.643 By using PPL as a catalyst and methanol as an initiator, ε-CL was polymerized in hexane at 45 °C for up to 26 days to complete the monomer conversion, affording a mixture of poly(ε-CL) and dilactone.644 So far, ROP of various unsubstituted and substituted lactones as well as other cyclic monomers has been extensively studied.35,37,40,41,44−46,60,646,647 Scheme 56 shows unsubstituted Scheme 56

lactones of different ring size which were reported to be polymerized by a lipase catalyst.648−652 The polymerization was conducted in bulk, an organic solvent, or a binary solvent system. ε-CL is a cheap monomer for polyester syntheses in industrial fields and shows high reactivity for ROP; thus enzymatic ROP of ε-CL (m = 5 in Scheme 56) has been extensively investigated. As mentioned above, various lipases were active for ROP of lactones. Lipases BC, CA, CC, CR, and MM, PPL, Aspergillus niger (lipase A), Penicillium roqueforti (lipase PR), and Rhizopus japanicus (lipase RJ) lipases also had catalytic activity for ROP of ε-CL.90,642−644,647−651,653−665 Recently, a novel thermophilic lipase from Fervidobacterium nodosum for ROP of ε-CL was developed.666 This enzyme could effectively catalyze the ROP at high temperatures and showed the highest activity at 90 °C. Yarrowia lipolytica lipase (lipase YL) immobilized on the selected supports catalyzed ROP of ε-CL to prepare poly(ε-CL) polyol for polyurethanes.667 In the case of crude industrial lipases such as PPL, lipases BC, CR, and PF, a large amount of catalyst (often more than 40 wt % for ε-CL) was required for the efficient production of the polymer. Lipase CA, a commercially available immobilized enzyme, on the other hand, showed the high catalytic activity toward the ε-CL polymerization; a very small amount of lipase CA (less than 1 wt % for ε-CL) was enough to induce the polymerization.668,669 Furthermore, poly(ε-CL) with the molecular weight close to 105 was obtained under the appropriate conditions. In the case of the polymerization in toluene at 70 °C, the polymer with the molecular weight of 2.5 × 10 4 was obtained. It was reported that covalently immobilized enzymes, Amberzyme-lipase CA and nanoPSGlipase CA, showed high catalytic activity for the ROP of εCL.670 Reaction parameters of enzymatic ROP of ε-CL with lipases of different origins were investigated by conducting the reaction in toluene at various temperatures. Lipase CA exhibited the

Figure 30. Examples of cyclic monomers for enzyme-catalyzed ringopening polymerizations.

Cyclic Ester (Lactone) Monomers and Lipase Catalyst. Various cyclic esters have been subjected to lipase-catalyzed ROP (Scheme 55A). Lipase catalyzed the ROP of 4- to 17Scheme 55

membered nonsubstituted lactones. In 1993, it was first demonstrated that medium-size lactones, δ-valerolactone (δVL, 6-membered), and ε-caprolactone (ε-CL, 7-membered), were polymerized by industrial lipases derived from Candida cylindracea (lipase CC), Burkholderia cepacia (lipase BC), Pseudomonas fluorescens (lipase PF), and porcine pancreas (PPL).642−644 For example, ROP of ε-CL by lipase PF in bulk at 75 °C for 10 days gave poly(ε-CL) with molecular weight of 7.7 × 103 in 92% yields, and ROP of δ-VL at 60 °C afforded the corresponding polyester with a molecular weight of 1.9 × 103.642 Candida antarctica (lipase CA), Candida rugosa (lipase CR), and Rhizomucor meihei (lipase RM) were also active for ROP of these monomers. The obtained polyesters possessed the terminal structure of a carboxylic acid group at one end and a hydroxyl group at the other, indicating that the ROP was initiated by a water molecule and terminated also by water without any additives. For ROP of ε-CL, the catalyst activity was examined by using the lipase catalyst in a supported form or a free form.645 2356


Chemical Reviews

Review

highest catalytic activity at an optimal temperature of 65 °C in an optimal toluene/monomer ratio of 50/50. Polymerization and degradation during the reaction are competitive, and hence, the molecular weight versus reaction time relationship was an important factor. A new model to account for simultaneous polymerization and degradation including various factors was developed.669 The lipase-catalyzed polymerization of ε-CL in bulk produced the linear polymer, whereas the main product obtained in organic solvents was of cyclic structure, suggesting that the intramolecular condensation took place during the polymerization. In the PPL-catalyzed ROP of ε-CL in the presence of butanol, both butanol and water were involved in the initiation reaction.653 Multifunctional initiators from polyglycidols having primary OH groups were employed for initiating ROP of ε-CL by enzymatic and chemical catalysts. Lipase CA and Zn(II) 2ethylhexanoate were used as enzymatic and chemical catalysts, respectively. The formed polymer architectures were of core− shell structure, and the structual difference was discussed for these processes.671 Poly(ε-CL) with molecular weight larger than 7.0 × 104 was enzymatically obtained from the corresponding oligomer at 70 °C in toluene.90,655 A cyclic dimer (14-memberd) of ε-CL was polymerized by lipase CA to produce a poly(ε-CL) molecular weight of 8.9 × 104 at higher temperature quantitatively, and another larger cyclic oligomer was also polymerized.90,672,673 Using a microbial telechelic polyester of low molecular weight, hydroxylated poly[(R)-3-hydroxybutylate] (PHB-diol), as initiator, lipase CA-catalyzed ROP of ε-CL in the presence of a microbial telechelic polyester of low molecular weight, hydroxylated poly[(R)-3-hydroxybutylate] (PHB-diol), was examined in toluene or 1,4-dioxane at 70 °C. The PHB-diol possessing primary and secondary hydroxyl groups at the terminal acted as an initiator. The initiation took place regioselectively at the primary hydroxyl group, affording the diblock polyester consisting of the PHB block with molecular weight of 2.4 × 103 and the poly(ε-CL) block with molecular weight of 1.5 × 103. Tg and Tm values of the block copolymers could be tuned by varying the content of the two blocks and their molecular weight.674 The cyclic oligomer prepared from 1,4-butanediol and dimethyl succinate was polymerized by the lipase catalyst to give poly(butylene succinate) (PBS) with a molecular weight of 1.3 × 105. Interestingly, the molecular weight of the polymer from the cyclic oligomer was higher than that of the polyester obtained by the direct polycondensation.675 Lipase CA was immobilized on a cross-linked macroporous copolymer from glycidyl methacrylate and ethylene glycol dimethacrylate. Effects of pore size, specific surface area, specific volume, and particle size on ROP of ε-CL were systematically investigated. Eighty percent of the enzyme was covalently immobilized on the macroporous resin. The decrease of the diameter, the increase of pore size, and specific surface area led to the improvement of the catalytic activity.676 Reuse of commercially available lipase CA for ROP of ε-CL was examined. Up to a 10 cycle, high molecular weight poly(εCL) with similar polyindex was obtained.677 Chemically immobilized lipase on photo-cross-linked chitin was developed for ROP of ε-CL.678 Lipase CA was also immobilized on montmorillonite and sepiolite nanoclays. An organo-modified clay catalyzed ROP of ε-CL to produce poly(ε-CL) and claynanohybrids, and the poly(ε-CL) chains were effectively grafted on the clay.679

Lipase-catalyzed ROP of ε-CL was examined to coat the hydrophilic cellulose-fiber surfaces with the hydrophobic poly(ε-CL) polyester chains by utilizing the cellulose-binding module-lipase CA conjugate as catalyst. The hydrophobicity of the surface did not arise from the covalently attached poly(εCL) to the surface hydroxyl groups but rather from the surfacedeposited polymers, which could be easily extracted.680 The lipase-catalyzed ROP reactivity of methyl-substituted εCL monomers was examined; ω-methyl ε-CL showed the least polymerizability among unsubstituted, α- and γ-substituted εCL monomers.664 Lipase CA induced the polymerization of αmethyl-substituted 6- and 7-membered lactones at 45 °C for 24 h to give the corresponding polyesters with molecular weight of 1.1 × 104 and 8.4 × 103, respectively.665 The microwave irradiation was applied for lipase CAcatalyzed ROP of ε-CL; the polymerization rate was decelerated in boiling nonpolar solvents such as toluene and benzene by the irradiation, whereas the rate was moderately accelerated in a boiling polar solvent, diethyl ether.681 The effects of the detailed reaction parameters such as reaction temperature, time, and microwave intensity were examined.682 The polymerization reaction under optimal conditions (90 °C, 240 min, and 50 W) afforded poly(ε-CL) with a molecular weight of 2.1 × 104. β-Propiolactone (β-PL, 4-membered) was polymerized by Pseudomonas family lipases as the catalyst in bulk, yielding a mixture of linear and cyclic oligomers with a molecular weight of several hundreds, whereas poly(β-PL) of high molecular weight (molecular weight > 5 × 104) was obtained by using lipase CR as a catalyst.683−687 Poly(malic acid) is a biodegradable and bioadsorbable watersoluble polyester having a carboxylic acid in the side chain. The chemoenzymatic synthesis of poly(malic acid) was achieved by the lipase-catalyzed polymerization of benzyl β-malolactonate followed by the debenzylation. The molecular weight of poly(benzyl β-malolactonate) increased by the copolymerization with a small amount of β-PL using the lipase CR catalyst. Propyl malolactonare (β-propyloxycarbonyl-β-PL) was also polymerized with lipase CR catalyst in toluene and in bulk to produce the polymer with a molecular weight of 5 × 103 quantitatively. The enzymatic polymerization took place much faster than the thermal polymerization.688 Lipase catalysis induced ROP of a 9-membered lactone (8octanolide, OL). The polymerization at 75 °C for 10 days produced the polymer with a molecular weight of 1.6 × 104.657,689 Lipases BC and CA showed the high catalytic activity for the polymerization. Macrolides, 11-undecanolide (12membered, UDL), 12-dodecanolide (13-membered, DDL), 15-pentadecanolide (16-membered, PDL), and 16-hexadecanolide (17-membered, HDL), were enzymatically polymerized.648−651,690−694 Various lipases catalyzed the polymerization of these macrolides. For the polymerization of DDL, the activity order of the catalyst was lipase BC > lipase PF > lipase CR > PPL. Lipases CC and PF produced polyUDL with high molecular weight, although the long reaction time was required. PolyPDL with a molecular weight of 6.2 × 104 was formed by the polymerization using lipase BC as a catalyst at 70 °C at a low reaction water level.693 The lipase CA-catalyzed polymerization of PDL proceeded fast in toluene to produce a high molecular weight polymer with the molecular weight higher than 8 × 104. The more amorphous phase was found in the polymerization at higher temperature.695 It is to be noted that a 2357


Chemical Reviews

Review

In the ring-opening copolymerization of ε-CL with DLLA by using lipase CA as catalyst, DLLA was consumed more rapidly than ε-CL in the initial reaction stage, which was incorporated into the LA dimer-wise. As the copolymerization proceeded, the relative amount of ε-CL in the copolymer increased. The nonrandom copolymer structure disappeared with time, probably due to the lipase-catalyzed transesterification reaction. The macrocyclic compound was formed during the copolymerization.706 The copolymerization of the cyclic dimeric monomers, DLLA, LLA and glycolic acid (GA) by using lipase BC catalyst were studied. The copolymerization of LA and GA (8:2 molar ratio) was conducted in bulk at 100 or 130 °C. Under these reaction conditions, the copolymer was formed without lipase (control experiment), the molecular weight of the copolymer was much smaller than that of the polymer obtained in the presence of the lipase catalyst. In the case of the copolymerization at 130 °C for 2 days, a cyclic random copolymer with molecular weight of 2.1 × 104 was formed, whereas the formation of the linear polymer with molecular weight of 1.2 × 104 was found at 100 °C for 7 days, suggesting that the complicated reactions took place.707 When multifunctional alcohols like ethylene glycol, pentaerythritol, inositol, and polyglyceline were used as initiator in the lipase PS-catalyzed ROP of LLA, DLA, and DLLA at 140 °C, various branched polymers were obtained.93 Generally, ring−chain (cyclic−linear structure) equilibrium exists in ROP of various cyclic monomers, and such an equilibrium was observed also for enzymatic ROP708 as well as enzymatic polycondensation.709 In some cases of lipasecatalyzed ROP of lactones such as ε-CL644 and β-PL,684 cyclic oligomers were formed in addition to major products of linear polyesters. Only a small amount of the cyclic dimer was formed in the lipase CA-catalyzed polymerization of ε-CL in bulk, whereas the polymerization in acetonitrile, THF, or 1,4-dioxane afforded the cyclic products. The formation of macrocycles up to the 23 monomer units were produced in 1,4-dioxane.708 These unique behaviors are explained by the backbiting reaction of a polymer acyl-enzyme intermediate. Lipase CA efficiently catalyzed the ring-opening polymerization of 1,5-dioxepan-2-one (DXO) to produce the corresponding poly(ester-alt-ether) (Scheme 57A).710,711 A

much higher ring-opening polymerizability of these macrolides than ε-CL was observed.657,689 Ring-opening copolymerization of PDL with four monomers, δ-VL, ε-CL, UDL, and DDL by lipases BC and PF at 60 °C in bulk afforded the corresponding copolymers, with structures not statistically random.648 Lipase CA catalyzed the ringopening copolymerization of PDL with 1,4-dioxan-2-one (DO) in toluene or diphenyl ether at 70 °C for 26 h gave the copolyester with a high molecular weight (>3 × 104), in which the enzymatic polymerizability of PDF was larger than that of DO.696 The largest-sized lactone monomer having a simple unsubstituted structure so far studied is 16-hexadecanolide (17-membered, HDL). ROP of HDL took place in the presence of lipase CA, CC, PC, PF, or PPL in bulk, efficiently producing polyHDL with a molecular weight of 5.8 × 103.694 A 24-membered lactone derived from natural sophorolipid was polymerized via lipase-catalyzed ROP to give a glycolipidbased polyester. The reaction proceeded in two modes, depending on the reacted position of a hydroxyl group with the formation of monoacylated products to oligomers and polymers.697 A 26-membered sophorolipid lactone having a double bond was enzymatically prepared and subjected to ringopening metathesis polymerization with a Ru catalyst, giving rise to poly(sophorolipid) with a molecular weight > 1 × 105.698 For the applications of glycolipid biomaterials, properties of the solid-state product were examined in detail.699 Macrocyclic esters having 24- to 84-ring atoms were polymerized by polymer-supported CALB catalyst. Typically, the polymerization was carried out in toluene at 70 °C for 20 h with 7.5% w/w of the CALB catalyst, affording polyesters with Mn up to 6.34 × 104 (PDI = 1.9) in 93% yields. The macrocyclic monomers can be cyclic oligomers of a smaller cyclic lactone and/or macrocyclic esters containing a functional moiety like steroid rings. These monomers are strainless, and hence, the polymerization was discussed in terms of a type of entropically driven ROPs (ED-ROPs).700 ROP of a bile acid (cholic acid) macrocyclic ester monomer was achieved by the CALB catalyst. The polymerization in toluene at 80 °C for 24− 72 h produced polymers with Mw = 10, 110−130, and 400 (PDI ∼ 1.20−1.94) in 29−67% yields. The reaction was understood to proceed via the transesterification mode. To obtain polymers efficiently, a high concentration of the starting monomer (∼30%) was chosen to favor the polymerization in the ring−chain equiliblium in the ROP.701 Lactide (LA), a six-membered cyclic dimer of lactic acid, is a very important starting monomer for industrial production of poly(lactic acid) (PLA), one of the most famous green plastics and bioabsorbable materials. Lactide was not enzymatically polymerized under mild reaction conditions; however, poly(lactic acid) with the molecular weight higher than 1 × 105 was formed using Pseudomonas species (lipase PS) as a catalyst at higher temperatures (80−130 °C), although the product yield was low. DL-LA (DLLA) gave the higher molecular weight in comparison with LL-LA (LLA) and DD-LA (DLA) monomers.702 Noticeably, lipase CA was reported not to catalyze ROP of LLA, whereas DLA was enantioselectively polymerized at 70 °C for 3 days to produce polyDLA with a molecular weight of 3.3 × 103.703 Mutants of lipase CA for ROP of DLA were created, and their activity was 90-fold higher than that of the wild type.704 Free lipase CA and immobilized one on chitin and chitosan were used for ROP of LLA to produce high molecular weight polyLLA.705

Scheme 57

linear relationship between the monomer conversion and the molecular weight of the polymer was observed. The monomer consumption followed a first-order rate law, suggesting no termination and chain-transfer reaction. The enzymatic polymerizability of DXO was much larger than that of ε-CL. In polyester synthesis via ROP, metal catalysts are often used. For medical applications of polyesters, however, there has been 2358


Chemical Reviews

Review

tetraoxacyclotetradecane-11,14-one, were used for varying the nature of the monomers. The lipase catalyst was located both in the water pool and in the bilayer and catalyzed the ROP of the lactones. OL and DDL were polymerized in the polymersome water pool similarly as in a free water system, but the resulting product had low molecular weight, probably due to sterically less accessibility of the monomer to the catalyst. scCO2, one of the most typical green solvents, could be used as a solvent for lipase-catalyzed ROP of lactones. The polymerization of ε-CL and the copolymerization of ε-CL and DDL using lipase CA as catalyst produced the polymer with molecular weight higher than 1 × 104.721 A further work reported the synthesis of poly(ε-CL) having a higher molecular weight reaching 7.4 × 104, and the repeated use of enzyme and scCO2 were achieved.722 The kinetic study showed that the polymerization was approximately first order with respect to monomer up to 80% conversion. Effects of water content in scCO2 on the reaction rate, and the molecular weight of the product were examined. The molecular weight reached 5 × 104 in the case of the dry reaction system, but the molecular weight could not be controlled owing to the large degree of transesterification, forming both linear polymers (intermolecular transesterification) and cyclic compounds (intramolecular transesterification), with behavior similarly observed in conventional solvents.723 Lipase catalysis also induced the degradation of poly(ε-CL) in scCO2 to form the oligomer of ε-CL.724,725 The oligomer with molecular weight less than 500 was polymerized by lipase CA to yield poly(ε-CL) with molecular weight higher than 8 × 104.725 scCO2 was used as solvent for chemoenzymatic synthesis of a block copolymer of ε-CL and MMA, which was obtained by the combination of lipase-catalyzed ROP of εCL and atom-transfer radical polymerization (ATRP) of MMA.726−728 Hyperbranched polyesters were prepared by lipase CA-catalyzed ROP of δ-VL or ε-CL in the presence of 2,2-bis(hydroxymethyl)butyric acid in 1,1,1,2-tetrafluoroethane (R-134a), a kind of green solvent, or scCO2.729 Room-temperature ionic liquids have received much attention as green designer solvents. Ionic liquids are nonvolatile, thermally stable, and highly polar liquids, which allow many organic, inorganic, metallo-organic compounds, and also polymeric materials to dissolve.730 Ionic liquid solvents such as 1-butyl-3-methylimidazolium salts ([bmim][X−]) have been used for lipase-catalyzed synthesis of polyesters. ε-CL was polymerized in [bmim][BF4],731 [bmim][PF6], or [bmim][(CF3SO2)2N] by lipase CA catalyst to produce poly(ε-CL) with molecular weight of several thousands in good yields.732 For lactide, high temperature was required in lipase CAcatalyzed ROP; PLA with a molecular weight of 3.6 × 104 was formed in the anhydrous ionic liquid, and the molecular weight decreased by the addition of water.733 Hyperbranched polyLLA was synthesized by lipase CA-catalyzed ROP in the presence of bis(hydroxymethyl)butyric acid in [bmim][PF6]. The degree of branching could be controlled by the reaction conditions.734 ROP of ε-CL also proceeded in various ionic liquids such as 1-ethyl-3-methylimidazolium tetrafluoroborate, 1-butyl-3-methylimidazolium tetrafluoroborate, 1-butylpyridinium tetrafluoroborate, 1-butylpyridinium trifluoroacetate, and 1-ethyl-3methylimidazoliumnitrate by using lipases YL and CR and PPL.735 Dicationic type of ionic liquids also promoted the lipase-catalyzed ROP of ε-CL.736 For the polymerization of εCL in an ionic liquid, the small amount was coated on an immobilized lipase CA.737 By selection of an appropriate ionic

concern about harmful effects of the metallic residues. Enzymatic synthesis of a metal-free polyester was demonstrated by the polymerization of 1,4-dioxan-2-one (DO) using lipase CA.712 Under appropriate reaction conditions, the high molecular weight polymer (molecular weight = 4.1 × 104) was obtained. Lipase CA-catalyzed the ring-opening copolymerization of DO with PDL in toluene or diphenyl ether at 70 °C and gave a copolyester of poly(DO-co-PDL) with a molecular weight higher than 3 × 104.696 Lipases CA, BC, and PF catalyzed the polymerization of cyclic diester type lactones, ethylene dodecanoate and ethylene tridecanoate to give the corresponding polyesters.713 The enzyme origin affected the polymerization behaviors; in using lipase BC catalyst, these bislactones polymerized faster than εCL and DDL, whereas the reactivity of these cyclic diesters was in the middle of ε-CL and DDL in using lipase CA. A cyclic dimer of ε-CL (14-membered) was polymerized by the lipase CA catalyst to yield poly(ε-CL) with a molecular weight of 8.9 × 104.90 A 12-membered lactone, 2-oxo-12-crown-4-ether (OC), showed high reactivity for lipase CA-catalyzed ROP to produce a water-soluble polyester with a molecular weight of 3.5 × 103 (Scheme 57B).714 In the copolymerization of OC with PDL, OC polymerized five times faster than PDL. Interestingly, the product copolymer was of the random structure. Reaction Solvents. Lipase-catalyzed ROP proceeded in bulk as well as organic solvents, typically toluene, heptane, 1,4dioxane, diisopropyl ether, and dibutyl ether. In addition, water, supercritical carbon dioxide (scCO2), and ionic liquids, which are regarded as typical “green solvents”, were also employed as solvent. In the first report in the lipase-catalyzed ROP in water, ε-CL, OL, UDL, DDL, and PDL were used as monomers (Scheme 56). Macrolides (UDL, DDL, and PDL) were polymerized by lipase in water to produce the corresponding polyesters with relatively low molecular weight in high yields. In terms of the yield and molecular weight of the product, lipase BC showed the highest activity.650,715 On the other hand, ε-CL and OL were not subjected to lipase-catalyzed ROP in water, due to no formation of an emulsion-like system. A mechanistic study on lipase-catalyzed ROP of PDL in aqueous biphasic medium was reported.716 A model was proposed assuming Michaelis− Menten interfacial kinetics followed by chain extension via lipase-catalyzed polycondensation. Polyester particles were formed by the lipase CA-catalyzed ROP of ε-CL, UDL, and PDL in water.717 Lipase-catalyzed ROP of lactones in a miniemulsion containing a surfactant was reported.718 Viscous mixing of PDL, a nonionic surfactant having a poly(ethylene glycol) chain (Lutensol AT50), water, and hexadecane afforded the miniemulsion, in which PDL was polymerized by addition of lipase PS at 45 or 60 °C. The PDL monomer was consumed quantitatively, and the polyPDL nanoparticles with diameter less than 100 nm were formed. PolyPDL showed a bimodal molecular weight distribution. In the polymerization in the presence of an unsaturated alcohol or acid such as linoleic acid, it was introduced at the polymer terminal via extensive esterifications to give a functionalized polyPDL. HDL was also polymerized in the miniemulsion system by lipase PS.719 Lipase CA-catalyzed ROP of lactones was examined in a stable polymersome with bilayer structure, which was formed by a polystyrene-polyisocyanopeptide block copolymer.720 Four lactones, ε-CL, OL, DDL, and a cyclic diester, 1,4,7,102359


Chemical Reviews

Review

alcohol, the macromonomer with various molecular weights was obtained. The amphiphilic nature of the macromonomer was controlled by the copolymerization to give an amphiphilic poly(ε-CL-co-DXO) macromonomer (Scheme 58B).711 4Pentene-2-ol could also initiate lipase CA-catalyzed ROP of ε-CL or DXO to give a macromonomer having a double bond. Propargyl alcohol and 4-dibenzocyclooctynol efficiently initiated ROP of ε-CL by lipase CA.742 The resulting alkyne group of the polymer was sequentially functionalized in a postpolymerization using metal-free and copper-catalyzed click chemistry. Poly(ethylene glycol) and poly(ε-CL)-diol were used as an initiator for ROP of ε-CL or DXO to give triblock polyesters with a hydroxyl group at both ends (Scheme 58C).743 The methacryl-type polyester macromonomer was produced via lipase-catalyzed ROP of PDL using HEMA as an initiator, which was subjected to radical polymerization, leading to polymers with brush structure.744 Lipase from Candida sp. 99− 125 catalyzed ROP of ε-CL in the presence of 6-mercapto-1hexanol to produce thiol-terminated poly(ε-CL). Interestingly, this enzyme remarkably showed chemoselectivity between hydroxyl and thiol groups.745 A porous material was obtained from a mixture of polyPDL and HEMA-initiated polyPDL by thermally induced phase separation method, which has a large potential as scaffolds for bone tissue engineering.746 Polyester-sugar or polyester-polysaccharide conjugates were regioselectively synthesized via enzyme catalysis (Scheme 59).

liquid, the polymer yield and molecular weight increased. It was reported that use of an ionic liquid as solvent and ultrasonic irradiation technique were combined in the lipase CA-catalyzed ROP of ε-CL. Compared to conventional nonsonicated operation, the sonication enhanced the polymerization rate by more than 2fold and the turnover number of the lipase catalyst by more than 3-fold.738 A continuous flow system for lipase-catalyzed ROP of ε-CL was developed by using a microreactor, in which an immobilized lipase CA was filled in the channel of the reactor. The large improvement of the polymerization rate was found in comparison with the conventional batch system. Furthermore, the molecular weight of the product was high.739 For lipase CAcatalyzed ROP of ε-CL in the presence of benzyl alcohol, the functionalization ratio of the polymer terminal was much higher than that in the batch system.740 End-Functionalized Polyesters. Since terminal-functionalized polymers such as macromonomers and telechelics are very important as prepolymer for construction of functional materials. Single-step functionalization of polymer terminal was achieved via lipase catalysis. A speculated mechanism of lipase catalysis involves a nucleophile like water and an alcohol to initiate ROP of lactones. Thus, alcohols could initiate lipase-catalyzed ROP of lactones to introduce the alcohol moiety at the polymer terminal. On the basis of this concept, enzymatic synthesis of various functional polyesters was reported (“initiator method”). Lipase CA-catalyzed ROP of ε-CL or DDL in the presence of a functional alcohol produced end-functionalized polyesters.741 Functional alcohols such as 2-hydroxyethyl methacrylate (HEMA), 5-hexen-1-ol, and 5-hexyn-1-ol afforded polyester macromonomers with methacryl, ω-alkenyl, and alkynyl groups, respectively. The quantitative introduction was achieved under the appropriate reaction conditions (Scheme 58A). Lipase CA-catalyzed ROP of ε-CL or 1,5-dioxepane-2-one (DXO) in the presence of HEMA produced the macromonomer. By changing the feed ratio of the monomer and

Scheme 59

Scheme 58 Lipase CA-catalyzed polymerization of ε-CL in the presence of alkyl glucopyranosides produced polyesters bearing a sugar at the polymer terminal.747,748 In the initiation step, the primary hydroxyl group of the glucopyranoside was regioselectively acylated. Enzymatic selective monosubstitution of a hydroxylfunctional dendrimer was demonstrated. Lipase CA-catalyzed polymerization of ε-CL in the presence of the first generation dendrimer gave the poly(ε-CL)-monosubstituted dendrimer.749 Ibuprofen, an important chiral 2-arylpropionic acid class of nonsteroidal anti-inflammatory drug, initiated lipase CAcatalyzed ROP of ε-CL, followed by polycondensation using alkaline protease from Bacillus subtilis to produce biodegradable polyesters with the drug pendants.750 The initiation process in the lipase-catalyzed ROP under various reaction conditions in bulk or in toluene strongly relates to synthesize end-functionalized polymers by introducing the functionality into polymers via ROP (initiator method).751 In the lipase-catalyzed ROP of lactones, an initiator nucleophile species can be water, an alcohol, an amine, or a thiol. Initiation reaction behaviors in lipase CA-catalyzed ROP of ε-CL using water, benzyl alcohol, and other Br-containing primary alcohols as initiators were investigated. Among these nucleophiles, water 2360


Chemical Reviews

Review

was the most reactive; thus the removal of water in the reaction system was important to achieve the high functionality to synthesized end-functionalized polymers. In the case of lipase CA-catalyzed ROP of ε-CL or PDL in the presence of HEMA in bulk at high temperature such as 80 °C, the complicated reaction behavior was found.752 Not only the HEMA-initiated ROP but also transesterification among the product polymers, monomer, and HEMA took place extensively after long reaction time. By utilizing the transesterification, dimethacrylated polyesters were obtained in onepot reaction. For the selective preparation of the desired macromonomer, the lipase-catalyzed ROP should be performed under mild conditions and terminated at a lowered monomer conversion. The effect of the initiator structure on the transesterification was examined. HEMA and 2-hydroxyethyl acrylate (HEA) were used as an initiator.753 In both cases, the transesterification occurred extensively at 80 °C in bulk, and the reaction rate was 15 times faster on the HEA-initiated system. Thiol end-functionalization was achieved by using lipase CA catalyst for ROP of ε-CL with 2-mercaptoethanol initiator (Scheme 58A).754 Enzymatic ROP was applied to synthesize hyperbranched aliphatic copolyesters by lipase CA-catalyzed copolymerization of ε-CL with 2,2-bis(hydroxymethyl)butylic acid. In preparing the AB2 polyesters, the degree of branching and the density of the functional end group were controlled.755 An alcohol attached on the gold surface was used as an initiator for lipase-catalyzed ROP of ε-CL and 1,4-dioxan-2-one (DO). The resulting system can be applied for biocompatible/biodegradable coating materials in biomedical areas such as passive or active coatings of stents. This method would be beneficial in the applications where the minimization of harmful species is critical.756 A similar surface-initiated polymerization was reported for the in situ solid-phase synthesis of biocompatible poly(3-hydroxybutyrate).757 A macroinitiator of a linear or a four arm star-shaped polyglycidol was used for synthesis of densely grafted poly(glycidol-graf t-ε-CL) and loosely grafted poly[(glycidol-graf t-ε-CL)-co-glycidol] copolymers via lipasecatalyzed ROP of ε-CL or Sn-catalyzed chemical process using the ε-CL monomer.758 In comparison with linear poly(ε-CL), the latter showed much enhanced degradability, probably due to high concentration of hydroxyl groups at the polyglycidol backbone. Terminator methods were also useful for enzymatic synthesis of end-functional polymers.759,760 A methacryl-type polyester macromonomer was synthesized by lipase PF-catalyzed polymerization of DDL using vinyl methacrylate as a terminator, in which the vinyl ester terminator was present from the beginning of the reaction. The polymerization in the presence of vinyl 10-undecanoate produced the ω-alkenyl-type macromonomer (Scheme 60A). In using divinyl sebacate as a terminator, the telechelic polyester having a carboxylic acid group at both ends was obtained (Scheme 60B).761 Telechelic polyesters containing a thiol group were prepared by an enzymatic one-pot system. The lipase CA-catalyzed ROP of PDL in the presence of 6-mercaptohexanol initiator and subsequent termination by γ-thiobutyrolactone produced a telechelic polyester with thiol groups at both ends (Scheme 60C).762 The obtained telechelic polymer was used for the preparation of semicrystalline polymer networks.763 When vinyl acrylate was used as a terminator, poly(PDL) with a thiol group at one end, an acrylate group at another end was formed.

Scheme 60

A combination of initiator and terminator method provided amphiphilic macromonomers containing poly(ε-CL); ROP of ε-CL was initiated via lipase catalysis from hydroxyl groupcontaining initiators like a hexahydroxyl dendrimer and ethyl glucopyranoside, followed by introduction of a (meth)acryloyl polymerizable group by termination with vinyl (meth)acrylate.764−766 Mechanism and Monomer Reactivity of ROP. It is wellknown that the catalytic site of lipase is a serine-residue, and lipase-catalyzed reactions proceed via an acyl-enzyme intermediate. The enzymatic polymerization of lactones is explained by considering the following reactions as the principal reaction course (Scheme 61).35,40,41,51,653,690 The key step is the reaction of lactone with lipase involving the ring-opening of the lactone to give an acyl-enzyme intermediate (“enzymeScheme 61

2361


Chemical Reviews

Review

Table 3. Comparison in Dipole Moment Values and Reaction Rate Values of Unsubstituted Lactones with Different Ring Size rate constant lactone (ring size)

dipole moment (C m)

δ-VL (6) ε-CL (7) HL (8) OL (9) NL (10) DL (11) UDL (12) DDL (13) PDL (16) HDL (17) butyl caproate

alkaline hydrolysisa (L mol

−1

−1

s , × 10 )

4.22 4.45 3.70 2.25 2.01 1.88 1.86 1.86 1.86

55000 2550 3530 116 0.22 0.53 3.3 6.0 6.5

1.75

8.4

4

relative rate of polymerization propagationb −1

enzymatic polymerization

(s , × 10 )

A

120

0.10 (0.14) 0.10 (0.14)

3

2.2 15

c

0.13 (0.18) 0.19 (0.26) 0.74 (1.0) 1.0 (1.35)

B

Zn-catalyzed polymerizatione

d

0.07 0.15 3.8 0.45 0.04 0.02 0.06 0.37 1.0

2500 (2800) 330 (370) 21 (23)

0.9 1.0 0.9 1.0

(1.0) (1.1) (1.0) (1.1)

With alkaline of NaOH in 1,4-dioxane/water at 0 °C.767 bNaOMe initiator in THF at 0 °C.768 cLipase PF as catalyst in i-propyl ether at 60 °C; the relative rate is given by normalizing the Vmax/Km values with respect to HDL and to PDL in parentheses.769 dLipase CA (immobilized) as catalyst in toluene at 45 °C; the relative rate is given by calculating the Vmax/Km values from Michaelis−Menten kinetics parameters and by normalizing the values with respect to PDL.652 eZn(Oct)2 initiator in bulk polymerization at 100 °C; the relative rate is given by normalizing the initial rate constants with respect to HDL and to PDL in the parentheses.770 Reprinted with permission from ref 52. Copyright 2009 American Chemical Society. a

mainly by the reaction rate (Vmax) but not by the binding abilities (i.e., the reaction process of the lipase-lactone complex to the acyl-enzyme intermediate is the key step of the polymerization). On the other hand, the Zn-catalyzed anionic polymerizability of lactones showed the reverse direction with large difference; for example, δ-VL and ε-CL showed 2500 and 330 times more reactive than the macrolides (Table 3), with behavior similar to that of alkaline hydrolysis. In the propagation reaction, the ratedetermining step is the nucleophilic attack of the propagating Zn-alkoxide species at the carbonyl carbon followed by the scission of the acyl-oxygen bond and the reformation of the alkoxide species.770 Immobilized lipase CA (Novozym 435) is the most used enzyme for enzymatic synthesis of polyesters. The structure of lipase CA was determined by X-ray crystallographic analysis.771 Lipase CA also exhibited higher catalytic activity than ε-CL for ROP of macrolides. Kinetic parameters as follows: initial rates (× 105, L mol −1 h−1 mg−1) were 2300 for ε-CL, 48 for OL, 1600 for DDL, and 4700 for PDL. For these four monomers, the corresponding relative values for lipase PF were 1.3, 1.8, 2.5, and 8.5, and those for lipase BC were 0.42, 2.2, 4.8, and 11.657,668,689 Catalysis of lipase CA for ROP of nine lactones was investigated according to Michaelis−Menten kinetics in detail.652 The values of Km were in a narrow range between 0.09 and 0.73 mol L−1, suggesting close affinities of lipase CA for all lactones. On the other hand, the values of Vmax varied between 0.07 and 6.10 mol L−1 h−1 and did not show a systematic trend. These behaviors are partially similar to those with lipase PF,650,651,769 suggesting that the key step in the lipase-catalyzed ROP of lactones is the process from the lipaselactone complex to form EM in Scheme 61. Column B in Table 3 shows the relative rate of the nine monomers derived from Vmax/Km values. The most reactive monomer (HL, 8membered) showed almost 200 times more reactive than the least reactive monomer (DL, 11-membered). The trend of the relative rates for the monomer ring sizes was complicated; there was no monotonous change depending on the ring size. These results suggest that various physical properties, dipole moment, ring strain, transoid and cisoid structure of lactones,767

activated monomer”, EM). The initiation is a nucleophilic attack of water, which is contained partly in the enzyme, onto the acyl carbon of the intermediate to produce ωhydroxycarboxylic acid (n = 1), the shortest propagating species. In the propagation stage, the intermediate is nucleophilically attacked by the terminal hydroxyl group of a propagating polymer to produce a one-unit-more elongated polymer chain. The kinetics of the polymerization showed that the rate-determining step of the overall polymerization is the formation of the enzyme-activated monomer. Thus, the polymerization proceeds via an “activated-monomer mechanism”. Reactivity of cyclic compounds generally depends on their ring size; small and intermediate ring-size compounds possess higher ring-opening reactivity than macrocyclic lactones (macrolides) due to their larger ring-strain. Table 3 summarizes dipole moment values and reactivities of lactones with different ring size. The dipole moment (indication of ring strain) of the macrolides is lower than that of δ-VL and ε-CL and close to that of an acyclic fatty acid ester (butyl caproate). The rate constant of the macrolides in anionic polymerization is much smaller than that of δ-VL and ε-CL, although the hydrolysis took a little peculiar behavior for medium-sized lactones, particularly NL and DL. These data clearly show that the macrolides have much lower ring strain and, hence, show less anionic reactivity and polymerizability than the medium-size lactones. As mentioned above, lipase catalysis provided the corresponding polyesters from macrolides with higher molecular weight in higher yields than that from ε-CL.648,690−692 These macrolides showed unusual enzymatic reactivity. Lipase PFcatalyzed polymerization of the macrolides proceeded much faster than that of ε-CL. The lipase-catalyzed polymerizability of lactones was quantitatively evaluated by Michaelis−Menten kinetics. For all monomers, linearity was observed in the Hanes-Woolf plot, indicating that the polymerization followed Michaelis−Menten kinetics. The Vmax(lactone) and Vmax(lactone)/ Km(lactone) values increased with the ring size of lactone, whereas the Km(lactone) values scarcely changed. These data imply that the enzymatic polymerizability increased as a function of the ring size, and the large enzymatic polymerizability is governed 2362


Chemical Reviews

Review

conformational strain, and transannular interactions should be taken into account for explanation of ROP behaviors with enzyme and chemical catalysts.652 Combination of docking and molecular dynamics studies provided information about the unique reactivity difference of lactones in lipase-catalyzed ROP. Cisoid or transoid conformation of the ester bond would govern the reactivity by the lipase catalysis.772 The turnover number (TON, mol of ε-CL converted to polymer per mol of enzyme per min) was examined for immobilized lipase CA and Humicola insolens cutinase (HIC). The value for lipase CA was close to that for HIC.773 A different approach of a copolymerization method gave some new insight into the ROP mechanism. Lipase catalysis is often used for enantioselective production of chiral compounds. The high enantioselectivity was achieved by the copolymerization with 7- or 13-membered nonsubstituted lactone using lipase CA catalyst (Scheme 62);774 the ee value reached ca. 70%

Figure 31. A general illustrative mechanism of lipase-catalyzed ROP of lactones.

plausible.648 Similar behaviors in the lipase PF-catalyzed copolymerization were found for a combination of δ-VL and ε-CL643 and of OL and ε-CL or DDL,689 in which the random copolymers were formed. Lipase CA-catalyzed ROP of 4-alkyl-substituted ε-CLs (methyl, ethyl, and propyl substituents) demonstrated the importance of the deacylating step in a mechanistic viewpoint (Scheme 63).777 The substituent size greatly affected the

Scheme 62

Scheme 63

in the copolymerization of β-BL with DDL. It is to be noted that in the case of lipase CA catalyst, the (S)-isomer was preferentially reacted to give the (S)-enriched optically active copolymer. The microstructure analysis of the copolymer by 13 C NMR showed that the diad unit of β-BL/β-BL was not included in the copolymer, indicating that the structure of the nucleophile greatly affects this copolymerization.774,775 The lipase CA-catalyzed copolymerization of δ-caprolactone (6membered) with DDL enantioselectively proceeded, yielding the (R)-enriched optically active polyester with ee of 76%. The above reports suggest that the rate-determining step is not always the formation of the acyl-enzyme intermediate (EM) in Scheme 61. It seems more general to consider a mechanism of the lipase-catalyzed ROP, in which the formation of EM (acylation of lipase) and/or the subsequent reaction of EM with monomer (deacylation of lipase) are operative depending on the monomer structure. In particular, the deacylation step becomes more important in the case of sterically bulky nucleophile of the propagating alcohol end, and this step determines the overall rate of reaction. Enantioselection is therefore induced possibly at both acylation and deacylation steps (Figure 31).775,776 The detailed microstructure of the copolymer from PDL and ε-CL obtained by lipase PF catalyst was analyzed in detail. In this case, the primary alcohol from the unsubstituted lactones is of the propagating chain-end structure. The competitive reaction rate ratio of PDL and ε-CL chain-ends toward EM of PDL was 1.04 and that of PDL and ε-CL chain ends toward EM of ε-CL was 0.89, suggesting that EM mainly governs the overall reactivity of the monomer, regardless of the propagating chain-end structure; the ROP mechanism involving the formation of the EM complex as a rate-determining step is

polymerization rate (the smaller one afforded the larger rate). Interestingly, the enantioselectivity changed from (S) for CH3 and C2H5 to (R) for C3H7, with great decreasing upon increasing the substituent size. For their enzymatic hydrolysis in water, on the other hand, the relative rate was close to each other and the enantioselectivity was (S) in all cases. These results suggest that the deacylation is a key step for the lipasecatalyzed ROP of 4-alkyl-substituted ε-CL; the reaction of EM with the nucleophile alcohol possessing different structures (bulky or less bulky alkyl) caused the polymerization rate difference and the shift of enantioselection. Transesterification also often took place in the lipasecatalyzed ROP under the severe reaction conditions besides the main reaction modes in Scheme 61 and Figure 31. In the lipase-catalyzed ROP of a macroride (DDL or PDL) in the presence of poly(ε-CL) or poly(1,4-butylene adipate)), a copolyester was formed due to the transesterification (Scheme 64).778,779 Lipase catalysis also induced the intermolecular transesterification between two different polyesters to produce a copolymer consisting of the two repeating units of the starting polyesters.778−780 Copolymerization of ε-CL with D,L-lactide was examined with lipase CA catalyst, and the product copolyester was structurally well characterized.706 Chemoselctive and Regioselective Polymerizations. Reactive polyesters were enzymatically synthesized. Lipase 2363


Chemical Reviews

Review

Scheme 64

Scheme 66

catalysis chemoselectively induced the ring-opening polymerization of a lactone having an exo-methylene group to produce a polyester having the reactive exo-methylene group in the main chain (Scheme 65). This is in contrast to the anionic Scheme 65 limited accessibility of the backbone. These molecular designs for selective functionalization of polymers demand high stereochemical regulation, and lipase catalysis is highly useful.787 Lipase-catalyzed graft copolymerization of ε-CL and β-BL onto chitin and chitosan was performed in a one-pot process. The hydroxyl group at the 6 position of chitin regioselectively initiated the ROP. For chitosan, the initiation took place at the hydroxyl group of the 6 position and amino group.788 Enantioselective Polymerization. As seen above, lipase enantioselectively catalyzed ROP of racemic cyclic monomers producing optically active polymers. Some examples are demonstrated to explain the mechanism of lipase-catalyzed ROP of lactones. Lipase PF catalyzed an enantioselective ROP of α-methyl-β-propiolactone in toluene to produce an optically active (S)-enriched (up to 75% (S)) polymer.685 Lipase BC induced the enantioselective polymerization of 3-methyl-4-oxa6-hexanolide (MOHEL). The initial reaction rate of the Sisomer was seven times larger than that of the R-isomer, indicating that the enantioselective polymerization of MOHEL took place through lipase catalysis.650 Lipase CA-catalyzed enantioselective ROP of substituted εCLs (racemic 4-MeCL and 4-EtCL) was achieved in bulk to give highly (S)-enriched poly(4-MeCL) and poly(4-EtCL) with an enantiomeric purity higher than 95%.789 Enantioselective ROP of methyl substituted ε-CLs were examined by using lipase CA and benzyl alcohol as the catalyst and initiator, respectively. For 3-MeCL, 4-MeCL, and 6-MeCL, (S)-selective polymerization proceeded, whereas (R)-selective behavior was found for 5-MeCL. The enantioselectivity for 4-MeCL was the highest; the enantiomeric excess (ee) value of the polymer = 0.88 (Scheme 67).790 Lipase CA also induced an enantiose-

polymerization; the vinyl polymerization of this monomer took place by a conventional anionic initiator or catalyst. The enzymatically synthesized polymer was readily subjected to the cross-linking with radical initiator to produce the insoluble polymer.781,782 Lipase CA induced chemoselective ROP of ambrettolide (Am) epoxide, a 17-membered macrolide having an epoxy group at the 10-position (Figure 30) to give the polyester with a molecular weight around 1 × 104. The epoxide group remained during the polymerization, which can be used for further modification.783 Globalide (Gl) and Am are industrial products used for fragrance. Gl is a 16-membered lactone having a double bond at the 11 or 12 position. These compounds were subjected to lipase CA-catalyzed ROP in toluene in the presence of molecular sieves, yielding polyGl and polyAm with molecular weight around 2.4 × 104. PolyGl and polyAm showed crystallinity, and their melting point (46−55 °C) was lower than that of the corresponding saturated polyester (around 95 °C). The unsaturated group in polyGl and polyAm was used for the radical cross-linking; the polymer was cured at 170 °C to give a transparent film, in which the endodouble bond was involved in the cross-linking.784 Regioselective initiation in lipase-catalyzed ROP of ε-CL was performed using a sugar molecule as an initiator. The primary hydroxyl group at the 6-position of methyl or ethyl glucopyranoside induced the ROP of ε-CL using CA and PPL as the catalysts.747−749 A sugar core-containing methacryltype macromonomer was also developed.766 Isosorbide initiated the ROP of ε-CL by using YLL immobilized on macroporous resins to produce an amphiphilic polymer consisting of the isosorbide headgroup and a hydrophobic polyester chain.785 A benzyl alcohol moiety in poly[styrene-co-(4-vinylbenzyl alcohol)] containing 10% hydroxyl group (m:n = 1:9) efficiently initiated lipase-catalyzed ROP of ε-CL to produce the corresponding graft copolymer (Scheme 66A).786 The enzymatic acetylation with vinyl acetate also proceeded for poly[styrene-co-(4-vinylbenzyl alcohol)] (Scheme 66B). In the case of the graft copolymerization, the grafting density of ε-CL was around 50−60%, whereas the acetylation ratio reached 95%. This is probably due to the sterical constraints, not to the

Scheme 67

lective ROP of fluorinated lactones in the ring size from 10 to 14. For contrast, the corresponding oxyacid gave an optically inactive polyester.791 Lipase CA-catalyzed ROP of ω-methylated six lactones was examined for elucidation of the relationships between the enantioselectivity of ROP and lactone structure (Scheme 68).792 The enantioselectivity was switched from (S)-selective 2364


Chemical Reviews

Review

opening of an (S) monomer with the benzyl alcohol initiator. For the lipase-catalyzed enantioselective propagation, the terminal (S) alcohol is less favored to react with the monomer via ring-opening, and hence, the Ru-catalyzed racemization of the terminal took place. Then, the terminal (R) alcohol was selectively reacted with the monomer by lipase catalysis to facilitate one monomer-unit elongation. This reaction cycle repeated to produce the (R) oligoester from the racemic monomer. This new approach can be regarded as a flexible and convenient tool for obtaining chiral macromolecules from racemic or prochiral monomers. A combination of simultaneous DKR of a secondary alcohol and ROP of ε-CL using lipase CA and Ru catalysts was reported.795 Racemic 1-phenylethanol (PhE) was used as a model alcohol and incorporated into the terminal of poly(εCL) under DKR conditions. The incorporation of 75% PhE was achieved, and the ee value of the product [(R)-PhE-poly(εCL)] was over 99%. This methodology is applicable for simple one-step production of polymers containing an optically pure group at the terminal, which have large application potentials for release of chiral species in medical fields.795 Chemoenzymatic Polymerization. Chemoenzymatic routes for production of functional polymers have been developed. Multiarm heteroblock star-type copolymers of PLA and poly(εCL) were prepared via a chemoenzymatic route. First, ROP of ε-CL was regioselectively initiated from the hydroxyl group at the 6 position of ethyl glucopyranoside by PPL catalyst, and the terminal hydroxyl group was acetylated with vinyl acetate by the same lipase catalyst. Second, Sn-catalyzed ROP of lactide was initiated from the remaining hydroxyl group of ethyl glucopyranoside to produce the block copolymer consisting of one poly(ε-CL) arm and three PLA arms.796 A different type of polymerization methods including enzymatic polymerization is useful for production of functional polymers.797 A combination of enzymatic ring-opening polymerization (eROP) of lactones and atom transfer radical polymerization (ATRP) was reported.798 Such a combination of two different consecutively proceeding reactions is referred to as cascade polymerization790 and allows a versatile synthesis of block copolymers consisting of a polyester chain and a vinyl polymer chain by using a designed initiator.790,797,799−801 Three

Scheme 68

for small (4-, 6-, and 7-membered) lactones to (R)-selective for large (8-, 9-, and 13-membered) lactones, and the kinetic study demonstrated the enantiomeric ratio was large for the large lactones. The unique behavior of the selectivity switch was supported by the molecular modeling studies on the free energy difference between the lactone structure and the active site cavity of the lipase. Lactone compounds take transoid and cisoid conformations, virtually small lactones for cisoid and large lactones for transoid. ROP of the small cisoid lactones was (S)-selective (3-MePL and 6-MeCL) or nonselective (5MeVL). ROP of the larger transoid lactones was (R)-selective with high enantioselectivity. For the lactones with intermediate ring size, 7-MeHL and 8-MeOL, the significant amount of cisoid conformers did not affect the enantioselectivity. These data suggest that the relationship between the lactone structure (especially lactone size) and the enantioselectivity of ROP is complicated.792 The similar behaviors of the selectivity switch was also found in the copolymerization from (S)- for 3-MePL (4-membered) to (R)-selectivity for 5-MeVL (6-membered) (Scheme 62),774 4-alkyl-substituted ε-CL (Scheme 63),777 and methyl-substituted ε-CL (Scheme 67).790 As mentioned above, lipase CA (immobilized) did not catalyze ROP of LLA, but ROP of DLA efficiently proceeded in the presence of lipase CA.703 Iterative tandem catalysis (ITC) was applied for lipasecatalyzed ROP of lactones. ITC means a polymerization in which the chain growth is effectuated by a combination of two different catalytic processes that are both compatible and complementary. By a combination of lipase CA (Novozym 435)-catalyzed ROP of 6-MeCL and Ru (2)-catalyzed racemization of a propagating secondary alcohol, optically pure (R) oligoesters (1a and 1b) were obtained (Scheme 69).793,794 This process is on the basis of dynamic kinetic resolution (DKR), and the polymerization was performed in one-pot. First, lipase CA enantioselectively catalyzed the ring-

Scheme 69. Reprinted from ref 793. Copyright 2005 American Chemical Society

2365


Chemical Reviews

Review

technique.727 A poly(ε-CL) macroinitiator was obtained via eROP using the bifunctional initiator similarly in Scheme 70. Subsequently, FOMA was polymerized from the macroinitiator to produce poly(ε-CL-block-FOMA). 2,2,2-Trichloroetahnol was used as an initiator for a combination of eROP and ATRP. Trichloromethyl groupterminated poly(ε-CL) was first prepared by lipase CAcatalyzed ROP of ε-CL, and subsequently, styrene (St) was polymerized with this macroinitiator via a carbon-chlorine bond cleavage via ATRP to produce poly(ε-CL-block-St).803 Instead of St, glycidyl methacrylate (GMA) was used to prepare an amphiphilic block copolymer. The resulting block copolymer formed polymeric micelles with hydrodynamic diameters around 100−200 nm.804 The trichloromethyl group-terminated polyDDL was similarly prepared by lipase catalyst, which was converted to a block copolymer of polyDDL and PSt via ATRP.805 Heterografted molecular bottle brushes (HMBB) were prepared via a chemoenzymatic process combining eROP and ATRP.806 Polyglycidol was used as multifunctional initiator for lipase CA-catalyzed ROP of ε-CL, followed by the selective enzymatic acetylation with vinyl acetate to form poly((glycidolgraf t-ε-CL-acetyl)-co-glycidol). The remaining hydroxyl group at the backbone was reacted with 2-bromo-2-methylpropionyl bromide to introduce the bromide group in the polymer. Then, MMA or n-butyl methacrylate was polymerized via ATRP with CuBr/2,2′-bipyridyl catalyst to produce HMBB with the designed polymer blocks. Liquid crystalline polymers were prepared via a chemoenzymatic route. Preparation of a monomer, 6-(4-methoxybiphenyl-4′-oxy)hexyl vinyl hexanedioate, was prepared with a lipase CA catalyst.807 For synthesis of multifunctional poly(meth)acrylates, functional monomers of (meth)acrylates were prepared by lipase CA-catalyzed transesterification of MMA and methyl acrylate with various functional alcohols. These monomers were radically polymerized by AIBN initiator to give poly(meth)acrylates via cascade reactions.808 New Monomer for Polyester Synthesis. A new monomer of the O-carboxylic anhydride derived from lactic acid (lacOCA) was found polymerized by lipase catalyst via ROP fashion with liberation of carbon dioxide (Scheme 71).809 Lipase PS and

plausible routes are demonstrated in Scheme 70. In route B, lipase CA-catalyzed ROP of ε-CL involving the initiation of the Scheme 70

hydroxyl group of the designed initiator produced Brterminated poly(ε-CL), followed by the Cu-catalyzed ATRP of styrene to produce poly(ε-CL-block-St).797 One-pot chemoenzymatic cascade synthesis of block copolymers combining lipase-catalyzed ROP of ε-CL and ATRP of alkyl methacrylate monomer was examined. ATRP system showed an inhibitory effect on the enzymatic activity. Methyl methacrylate interfered with eROP by transesterification, whereas t-butyl methacrylate was inert.802 This principle was extended to synthesis of optically active block copolymers. An enantioselective ROP from a combination of a bifunctional initiator having hydroxyl and bromide groups, an racemic monomer of 4-MeCL, and lipase CA produced an optically active polyester with the bromide terminal, which was subjected to ATRP of methyl methacrylate (MMA), yielding poly(4-MeCL-block-MMA) having a chiral polyester chain.790 By optimization of the reaction conditions, side reactions such as a homopolymer formation of poly(ε-CL) could be minimized to less than 5% with taking an optimized enzyme drying procedure.799 Branched polymers with or without polyMMA chain from a poly(ε-CL) macromonomer were produced via a chemoenzymatic pathway. 2-Hydroxyethyl α-bromoisobutyrate was used as a bifunctional initiator to synthesize a bromidecontaining poly(ε-CL) macromonomer, to which a polymerizable end group was introduced by in situ enzymatic acrylation with vinyl acrylate. Subsequent ATRP of the acrylate-type macromonomer afforded the branched polymer.800 Another living radical polymerization, a nitroxide-mediated radical process, was similarly used for one-pot chemoenzymatic synthesis of a block copolymer. Metal-free poly(ε-CL-block-St) was obtained in two consecutive polymerization steps via corresponding route B and in a one-pot cascade approach without intermediate transformation or workup step via corresponding route A. By combination of enantioselective ROP via lipase catalysis, a chiral block copolymer from 4-alkyl ε-CL and styrene with high enantiomeric excess was formed.801 scCO2 was used as a solvent for a spontaneous one-step chemoenzymatic synthesis of block copolymers via route A in Scheme 70.726 A combination of scCO2, lipase CA, ε-CL, MMA, CuCl, and 2,2′-bipyridine afforded poly(ε-CL-blockMMA) in a moderate yield. Each homopolymer was produced only in a small amount (less than 10%). An amphiphilic block copolymer was prepared from a fluorooctyl methacrylate (FOMA) in scCO2 by a sequential monomer addition

Scheme 71

Novozym 435 were used for the ROP of lacOCA to give poly(lactic acid) (PLA) having a relatively high Mn up to 38400 with Mw/Mn < 1.4 in high yields at 80 °C for a few hours. Both lipases showed similar catalytic acticity for ROP of lacOCA, and lacOCA is much more reactive with lipase catalysis than the lactide monomer. The lipase showed slightly higher catalyst activity for L-lacOCA than for D-lacOCA; enantioselection was scarcely observed. 6.3.1.2. Condensation Polymerization (Polycondensation). Following the ring-opening polymerization of cyclic compounds, condensation polymerization (polycondensation) is an important method as well for producing various polymers. General reactions for polyester synthesis are shown in Scheme 72. It is to be noted that enzyme-catalyzed polyester 2366


Chemical Reviews

Review

conditions.818 Immobilized Humicola insolens (HiC)- and Novozym 435 (N435)-catalyzed homopolymerizations of ωhydroxyalkanoic acids (ω-HA) were investigated. ω-HAs examined were adipic acid (C6), 10-hydroxydecanoic acid (C10-HA), 12-hydroxydodecanoicacid (C12-HA), 16-hydroxyhexadecanoic acid (C16-HA) (Scheme 74).819 HiC’s activity

Scheme 72

Scheme 74

synthesis was first achieved via polycondensation in the middle of the 1980s.810,811 Condensation reactions are basically reversible processes, and hence, it is necessary to remove or reduce a byproduct water or an alcohol from the reaction mixture for producing the product polymer efficiently. The catalysis of lipase in the dehydration polycondensation to produce polyester via bond formation is that of a reverse reaction of the hydrolysis of ester compounds via bond cleavage. Polycondensation of Oxyacid or Its Ester (Reaction 1 in Scheme 72). The simplest mode is a dehydration polycondensation of oxyacids. The first paper of this type was reported in 1985, describing a lipase-catalyzed polycondensation of an oxyacid monomer, 10-hydroxydecanoic acid.811 The degree of polymerization (DP) value of the product was small (≥5). A poly(ethylene glycol)-modified esterase also induced the oligomerization of glycolic acid, the smallest oxyacid (R = H, Scheme 73).811 A lipase-catalyzed polymerization of lactic acid gave a low molecular weight poly(lactic acid) (PLA) under various reaction conditions (R = CH3, Scheme 73).812,813

for ωHA substrates with 6, 10, 12, and 16 carbons was C16 > C12, where C10-ωHA and C6-ωHA were not polymerized. In contrast, N435’s activity for ωHA substrates was C16 = C12 > C10, where C6-ωHA was not polymerized. HiC-AO (amberzyme oxirane resin)- and N435-catalyzed C16-HA homopolymerization at 8 h gave polymers with Mn = 40.4 × 103 and 25.5 × 103, respectively. Novozym 435 was an efficient catalyst for the dehydration polycondensation of an oxyacid, cis-9,10-epoxy-18-hydroxyoctadecanoic acid isolated from the outer birch bark, carried out in toluene in the presence of molecular sieves at 75 °C for 68 h to give the polyester with the highest Mw = 20000 (Mw/Mn = 2.2).820 In addition to the dehydration polycondensation, the transesterification polycondensation is another reaction of producing polyesters, in which an ester compound is employed as a monomer. Polycondensation of methyl 6-hydroxyhexanoate, an oxyacid ester, with a lipase catalyst in hexane at 70 °C for more than 50 days gave the polyester with DP up to 100. PPL-catalyzed polymerization of methyl 5-hydroxypentanoate for 60 days produced the polymer with DP of 29.644 Various hydroxyesters, ethyl esters of 3- and 4-hydroxybutyric acids, 5and 6-hydroxyhexanoic acids, 5-hydroxydodecanoic acid, and 15-hydroxypentadecanoic acid, were polymerized by lipase PS catalyst at 45 °C to give the corresponding polyesters with a molecular weight of several thousands.821 The enzymatic polycondensation of methyl ricinoleate by lipase PC catalyst produced poly(ricinoleate) with high molecular weight (Mw > 1 × 105) in bulk in the presence of molecular sieves at 80 °C after 7 days. The polymer was a viscous liquid at room temperature with Tg of −74.8 °C and biodegradable, which was readily cured to give chloroform-insoluble cross-linked materials.815 Novozym 435 catalyst induced a regioselective polycondensation of isopropyl aleuriteate, where the only primary alcohol was reacted at 90 °C in a toluene/2,4-dimethyl-3-pentanol mixed solvent, giving rise to the polymer of Mn = 5600 in 43% yield (Scheme 75).783

Scheme 73

A dehydration polycondensation of ricinoleic acid (a main component from castor oil) was catalyzed by lipase at 35 °C in an organic solvent to give a polyester with a Mn ∼ 1000 in good yield.814 Ricinoleic acid was again polymerized via dehydration with immobilized lipase PC catalyst to give the polymer with a Mw up to 8500.815 Lipase of Novozym 435 catalyzed the polycondensation of several oxyacids such as 18-hydroxyoctadecanoic acid and 12hydroxydodecanoic acid. Oligomerization of cholic acid regioselectively acylated the hydroxy group at the 3-position to give the oligoester with molecular weight of 920.816 Polyesters of relatively high molecular weight were produced from 10-hydroxydecanoic acid and 11-hydroxyundecanoic acid using a large excess amount of lipase CR catalyst. In the case of 11-hydroxyundecanoic acid, the corresponding polymer with a molecular weight of 2.2 × 104 was obtained in the presence of activated molecular sieves.817 Lipase CA induced the polycondensation of hydrophobic oxyacids efficiently. In the polymerization of 16-hydroxyhexadecanoic acid, 12-hydroxydodecanoic acid, or 10-hydroxydecanoic acid under vacuum at a higher temperature (90 °C) in bulk for 24 h, the DP value of the product polymer was beyond 100, whereas the polyester with lower molecular weight was formed from 6-hydroxyhexanoic acid under similar reaction

Scheme 75

2367


Chemical Reviews

Review

cross-linked by a diisocyanate compound to give a biodegradable tough material of polyester.827 A new enzymatic oligomerization reaction of alkyl lactates (RLa) was disclosed, in which lipase828 and protease829 were employed as catalysts. These studies gave new information on the mechanism of enantioselection. Polycondensation of alkyl D-lactates (RDLa, an oxyacid ester) catalyzed by lipase (Novozym 435) at 50 °C produced oligo(D-lactic acid)s (oligoDLAs) up to 82% yields with a n value = 2−7 (Scheme 76). Primary alkyl lactates of R = Et-, Pr-,

Copolycondensation of 12-hydroxydodecanoic acid with methyl 12-hydroxystearate (both from seed oil) was catalyzed with N435 in toluene in the presence of molecular sieves at 90 °C. During the reaction, water as well as methanol liberated. After several days, the copolymer was obtained in good yields, having a Mw ∼ 1.0 × 105 showing elasticity and biodegradability.822 Terminal-functionalized polymers like macromonomers and telechelics are important and often used as prepolymers for synthesis of functional polymers. The lipase CA-catalyzed polycondensation of 12-hydroxydodecanoic acid in the presence of 11-methacryloylaminoundecanoic acid conveniently produced a methacrylamide-type polyester macromonomer, in which a β-cyclodextrin was attached in the side chain.823 Enatioselective polycondensation of racemic 10-hydroxyundecanoic acid by lipase CR catalyst gave an optically active oligoester. The resulting oligomer was enriched in the (S) enantiomer to a level of 60% enantio-excess (ee) and the residual monomer was recovered with 33% ee favoring the (R) enantiomer.824 Optically active oligomers (DP < 6) were also synthesized from racemic ε-substituted-ε-hydroxy esters using the PPL catalyst. The enantioselectivity increased as a function of bulkiness of the monomer substituent.825 A new method of synthesizing chiral polyesters was developed, which is called iterative tandem catalysis (ITC). In the transesterification polycondensation of racemic AB type monomers having a secondary hydroxy group and a methyl ester moiety (Figure 32), the concurrent actions of an

Scheme 76

and Bu- showed a higher reactivity than longer alkyl lactates like R = Pe-, Hx-, Hp-, and Oc-, and a secondary alkyl lactate of sBuDLa showed a reduced reactivity. L-Lactates did not show oligomarization reactivity (i.e., enantioselection for D-isomers is very strict).828 Michaelis−Menten eq 1 and for simplicity a pseudo-first order rate eq 2 were applied for the reaction analysis, k′

kcat

E + S ⇄ ES ⎯→ ⎯ P+E

−

(1)

d[S] = k′[E][S] = k[S](k′[E] = k) dt

(2)

where E, S, and P denote enzyme, substrate, and product, respectively. Plots of the integrated form of eq 2 gave k values ( × 104 s−1): MeDLa (3.7); EtDLa (4.4); PrDLa (3.7); and BuDLa (3.4). The inhibition function of EtLLa toward the oligomerization of EtDLa was found “competitive”. The results accorded with the following hydrolysis experimental results. Hydrolysis of BuDLa and BuLLa was conducted in THF (Scheme 77).828 In Scheme 77

Figure 32. Illustration of ITC method. Reprinted with permission of ref 826. Copyright 2008 Wiley.

contrast to the oligomerization, Novozym 435-catalysis induced the hydrolysis of both BuDLa and BuLLa substrates. The rough values (× 104 L mol−1 s−1) are k = 2.1 for BuDLa and k = 0.92 for BuLLa at 50 °C; the D-isomer proceeded about 2.3 times faster than the L-isomer. These observations led to the mechanistic aspects of lipase (Novozym 435) catalysis: enantioselection is operated by deacylation step as shown in Figure 33,828 where for simplicity only the dimer formation is shown. At first, the monomer (substrate) is to be activated by an enzyme with forming (R)acyl-enzyme intermediate in step (a), (enzyme-activated monomer: EM), (“acylation of lipase”). Onto the activated carbonyl carbon of EM, OH-group of the D-lactate nucleophilically attacks to form an ester bond with liberating lipase enzyme, giving rise to D,D-dimer in step (b) (“deacylation of lipase”). If, in place of the D-lactate monomer, the OH-group of D,D-dimer attacks EM, D,D,D-trimer will be formed, and the repetition of this type of reaction ends up with the formation of

enantioselective acylation catalyst (Novozym 435) and a racemization catalyst (Ru(Shvo)) brought about the high conversion of the racemic monomers to enantio-enriched polymers. AB type monomers used were typically methyl 6hydroxyheptanoate (Me-6HH), methyl 7-hydroxyoctanoate (Me-7HO), methyl 8-hydroxynonanoate (Me-8HN), and methyl 13-hydroxytetradecanoate (Me-13HT). The use of isopropyl esters and/or the strict exclusion of oxygen during the polymerization afforded chiral polymers with molecular weight (9.4 × 103) and high enantiomeric excess (92%).826 A chemoenzymatic method gave an oleic acid-based polyester: first, oleic acid (a C-18 monoene acid from vegetable oils) was epoxidized by lipase catalyst with H2O2 oxidant, and then the intermolecular ring-opening addition between the epoxide group and the CO2H group thermally took place to produce the poly(oleic acid)-based polyester. It was further 2368


Chemical Reviews

Review

Figure 33. Lipase-catalyzed oligomerization pathways of (A) D-lactates and (B) L-lactates: acyl-enzyme intermediate formation steps (a and e), subsequent dimer formation steps (b, c, f, and g), and hydrolysis steps (d and h). ○ denotes that the step takes place, whereas × denotes that the step does not take place. (In steps b, c, d, f, g, and h, the leaving group of lipase is omitted.) Reprinted from ref 828. Copyright 2010 American Chemical Society.

indicating that the lipase catalysis provides with a good enantiopurification method. It is to be added that a bacterial synthase or polymerase (PHA synthase; EC 2.3.1) was used as a catalyst for another in vitro enzymatic polyester synthesis using lipase catalyst. Poly(hydroxyalkanoate)s (PHAs) are well-known as biodegradable polyesters produced commercially via fermentation.831 Around 60 different PHA synthases were isolated and characterized.832 The reaction type of the PHA synthasecatalyzed polymerization is of polycondensation. There are some PHA examples; poly[(R)-3-hydroxybutylate] [P(3HB)], poly[(R)-3-hydroxyvalerate] [P(3HV)], poly[4hydroxybutylate] [P(4HB)], etc. The in vitro synthesis of a PHA via the polycondensation of (R)-3-hydroxybutyryl coenzyme A (3HB-CoA) was reported first using a PHB synthase from Ralstonia eutropha (R. eutropha) as a catalyst (a general scheme in Scheme 78; R = CH3, m = 1).523 The

higher D-oligomers. Since the L-lactate was not consumed, the reaction of EM with the OH group of L-lactate does not occur; reaction of step (c) does not take place. On the other hand, hydrolysis of D-lactate also needs activation to form EM. Then, EM reacts with water to give D-lactic acid shown in step (d). In the reactions of L-lactate monomers (B), alkyl L-lactates were not consumed in the oligomerization. But, in the hydrolysis, alkyl L-lactates were hydrolyzed to give L-lactic acid in step (h). This is a clear indication that step (e) actually took place to produce (S)-acyl-enzyme intermediate EM. However, neither the OH-group of D-lactate nor the OH-group of L-lactate was allowed to attack EM to give a L,D-dimer via step (f) or L,L-dimer via step (g). Hydrolysis steps (d) and (h) (both deacylations) are nonselective due to no chirality in water molecule, whereas esterification steps (b), (c), (f), and (g) (all deacylations) are enantioselective. The above results demonstrate that “the enantioselection is governed by the deacylation step”; among four steps, only step (b) is allowed to give a D,D-dimer. However, the EM formation, both steps (a) and (e), is possible from all alkyl D- and L-lactate monomers. The above D-selective reaction of alkyl lactates by lipase catalysis was applied for optical resolution of D,L-isomers.830 Typically, a mixture containing 90.4% of n-butyl L-lactate (BuLLa) and 9.6% of D-lactates (BuDLa) was incubated with an immobilized lipase for 72 h, during which D-selective oligomerization of BuDLa occurred. After the distillation of the reaction mixture, the purity of BuLLa was increased to 98.6%,

Scheme 78

monomers, CoA thioester derivatives of 3HB and 3HV, were chemically prepared and polymerized in an aqueous solution at room temperature by the catalysis of the purified, recombinant polymerase from R. eutropha.524,525 Homopolymerization of 3HB-CoA gave P(3HB) with high molecular weight Mn = 4.21 × 106 (Mw = 7.41 × 106), indicating a close value of the 2369


Chemical Reviews

Review

monomer-to-polymerase ratio (2 × 104). This suggests a “living nature” of the polymerization. Actually, a block copolymer of P(3HB)-b-P(3HV) was obtained by the two-step sequential copolymerizations of 3HB-CoA and then 3HV-CoA. In the batch copolymerization of the two monomers, random to somewhat blocky copolymers were obtained.524,525 A mechanistic study on PHB synthase from R. eutropha was examined in detail to lead to a revised model.524 Purified PHB synthase exists in equiliblium with monomer and dimer. The active enzyme is a homodimer; in spite of two identical thiol groups of Cys319 responsible for the catalysis, the dimer shows only a single catalytic site. PHA synthase from R. eutropha was immoibilized onto solid surfaces through a transition-metal (e.g., Ni) complex. The synthase catalyzed the surface-initiated enzymatic polymerization of 3HB-CoA, forming a polyester of P(3HB) film with a uniform thickness on the surface. Various substrate surfaces like agarose beads, Si/SiO2 glass, aluminum oxide, and gold can be modified.757 Surface modification of two hydrophobic substrates, highly oriented pyrolytic graphite (HOPG), and an alkanethiol self-assembled monolayer (SAM), was conducted through PHA synthase-catalyzed polymerization of (R)monomers of 3HB-CoA and 3HV-CoA. The resulted polyester-modified surfaces were analyzed by atomic force microscopy (AFM) and quarz crystal microbalance (QCM), revealing the PHA film thickness of 1−6 nm on HOPG and SAM surfaces. For giving thin film formation, PHA synthase enzyme is to be located near a hydrophobic substrate surface.833 Polycondensation of Dicarboxylic Acid or Its Ester and Diol or Polyol (Reaction 2 in Scheme 72). via Dehydration. Lipase-catalyzed polycondensation of this type reaction (2) in Scheme 72 was reported in 1984 on the dehydration reaction between a dicarboxylic acid and a diol to produce oligomers, the earliest paper on the lipase catalyzed polyester synthesis (Scheme 79).810 In the case of using a polyol with more than

polyester than a bulk reaction.837 Under vacuum, the molecular weight was greatly enhanced, leading the equilibrium toward the product polymer side; a polyester with molecular weight of 2 × 104 was obtained by the lipase-catalyzed polymerization of sebacic acid and 1,4-butanediol in diphenyl ether or veratrole under reduced pressure.838 Even in ionic liquids,730 lipase was active and dehydration polymerization took place to give effectively the product polyester.731 Lipase CA (CALB) efficiently catalyzed the polycondensation of dicarboxylic acids and glycols without solvent under mild reaction conditions at 60 °C. Methylene chain length of the monomers greatly affected the polymer yield and molecular weight. The polymer with molecular weight higher than 1 × 104 was obtained by the reaction under vacuum.839,840 CALB was covalently immobilized onto epoxy-activated macroporous poly(methyl methacrylate) amberzyme beads with a poly(glycidyl methacrylate) outer region. In bulk amberzyme-CALB catalyzed polycondensation between glycerol (0.1 equiv), 1,8octanediol (0.4 equiv), and adipic acid (0.5 equiv) at 90 °C for 24 h gave polyester of a Mw ∼ 4.0 × 104.670 For the polymerization of adipic acid and 1, 6-hexanediol, the enzymatic activity loss was small during the reaction, whereas less than half of the activity remained when using glycols with methylene chain length less than 4.841,842 A scale-up experiment produced the polyester in more than a 200 kg yield, claiming a large potential as an environmentally friendly synthetic process of polymeric materials owing to the mild reaction conditions without using organic solvents and toxic catalysts.841 To obtain information for commercialization of new enzymebased products, the scale-up experiments of the Novozym 435catalyzed polyester synthesis from adipic acid and glycerol (a triol) was performed on a 500 g scale. The reaction was carried out in a heated, solvent-free system and the influence of various reaction conditions (i.e., temperature 60−90 °C, reduced pressure, enzyme concentration, reactants ratio, stirrer type, stirring rate, and reaction time) on the substrate conversion and molecular weight of the product was investigated. Conversions were higher than 90%, and molecular weights were in the desired Mn range of 2000−3000. New function of the product polymer is expected due to high hydroxyl functionality.843 Direct polycondensation of adipic acid and 1,8-octanediol in the presence of L-malic acid (L-MA) in organic media was achieved with Novozym 435 catalyst. The molecular weight reached to a maximum of 17400 at 80 °C in isooctane at a LMA feed ratio in the diacids of 40 mol %. The Mw increased from 3200 to 16600 when the reaction time was extended from 6 to 48 h at 70 °C. The hydrophilicity, thermal stability, and crystallizability of the copolymer were also investigated.844 As a green polymerization cycle, the preparation of partially renewable aliphatic polyesters based on 1,8-octanediol and biobased long-chain diacids, namely 1,12-dodecanedioic and 1,14-tetradecanedioic acid, was performed. It involved CALBcatalyzed prepolymerization combined with low-temperature postpolymerization, in the melt or solid state.845 Dehydration reaction is generally achieved in nonaqueous media; the product water of the dehydration is in equilibrium with starting materials, and hence, the solvent water disfavors the dehydration to proceed in an aqueous medium due to the “law of mass action”. Nevertheless, lipase catalysis enabled a dehydration polycondensation of a dicarboxylic acid and a glycol in water at 45 °C, to afford a polyester in good yield.460,461 Lipases CA and other lipases were active for polycondensation of sebacic acid and 1,8-octanediol (p = q = 8,

Scheme 79

three OH groups, a product polyester is often of branched structure, depending on the reaction conditions. Typically, a dehydration polycondensation of adipic acid and 1,4-butanediol in diisopropyl ether gave a polyester with a degree of polymerization (DP) of 20 (p = q = 4).834 Higher molecular weight polyesters were enzymatically obtained by polycondensation of sebacic acid and 1,4-butanediol (p = 8, q = 4) under vacuum. In the lipase MM-catalyzed polymerization in hydrophobic solvents with high boiling point such as diphenyl ether and veratrole, the molecular weight of polyesters from various combinations of diacids and glycols reached the value exceeding 4 × 104.835,836 Effects of substrates and solvent on the ester-chain formation, polydispersity, and end-group structure were examined. Diphenyl ether was a preferred solvent for the polycondensation of adipic acid (p = 4) and 1,8-octanediol (q = 8) giving a Mn of 28500 (48 h, 70 °C). Monomers having longer alkylene chain length of diacids (sebacic and adipic acids) and diols (1,8octanediol and 1,6-hexanediol) showed a higher reactivity than the reactions of shorter chain derivatives. A reaction using diphenyl ether solvent gave a rather higher molecular weight 2370


Chemical Reviews

Review

Scheme 80

in H2O solvent, Scheme 79). In the polymerization of an α,ωdicarboxylic acid and a glycol, the polymerization behavior greatly depended on the methylene chain length of the monomers. The polymer was obtained in good yield from 1,10decanediol, whereas no polymer formation was observed from 1,6-hexanediol, suggesting that the combination of the monomers with appropriate hydrophobicity is needed for the polymer production by dehydration. This finding of dehydration in water is a new aspect and attracted organic chemists with much interest. Novozym 435-catalyzed dehydration polymerization was applied to use polyols including sugar components. The direct polycondensation between adipic acid and sorbitol in bulk was performed at 90 °C for 48 h. The product poly(sorbityl adipate) was water-soluble. The Mn and Mw values were ∼1.1 × 104 and ∼1.7 × 104, respectively. In the polymer, sorbitol was esterified at primary alcohol group of 1- and 6-positions with high regioselectivity (∼85%). In place of sorbitol, glycerol was employed; however, the Mn and Mw values were lower, 2500 and 3700, respectively. To obtain a water insoluble sorbitol copolyester, adipic acid, 1,8-octanediol, and sorbitol (molar ratio 50:35:15) were ter-polymerized at 90 °C for 42 h (Scheme 80A). The methanol-insoluble part (80%) had a Mw of 1.17 × 105. Another ter-polymerization of adipic acid, 1,8octanediol, and glycerol (molar ratio 50:40:10) was performed in bulk at 70 °C, to give a polyester (Scheme 80B). The product polymer showed the monomer ratio in the 50:41:9, respectively, and the values for Mw and Mw/Mn of 75600 and 3.1, respectively. The product contained 90% methanolinsoluble parts, showing few cross-links. The selectivity at glycerol primary alcohol sites was only 66%, therefore, the product was highly branched; 27% of glycerol units were for branched sites.846 Physical properties of the resulting polyesters containing sorbitol or glycerol were characterized in detail.847 Lipase CA catalyzed bulk dehydration polycondensation of the monomers, adipic acid (A2), 1,8-octanediol (B2), and glycerol (B′B2), gave hyperbranched polyesters at 70 °C for 42 h. With monomer feed molar ratio (A2 to B2 to B′B2) 1.0:0.8:0.2, linear copolyesters were formed during the first 18 h, and extending the reaction time to 42 h gave hyperbranched copolymers with dendritic glycerol units. The regioselectivity for esterification at the primary glycerol positions ranged from 77 to 82%. Variation of glycerol in the monomer feed gave copolymers with a degree of branching from 9 to 58%.848 A similar dehydaration polycondensation to produce terpolymers was conducted by

using monomers, adipic acid (A2), 1,8-octanediol (B2), and trimethyrolpropane (TMC, B3) with lipase CA catalyst in bulk at 70 °C for 42 h. As an example, a hyperbranched copolyester with 53% TMC adipate units was obtained in 80% yield, with a Mw = 14100 (Mw/Mn = 5.3).849 CALB (N435)-catalyzed onepot copolymerization of linoleic acid (LA), glycerol (G), and 1,18-cis-9-octanedecenedioic acid (oleic diacid, OD) yielded cross-linkable unsaturated polyesters. The reaction efficiently formed poly(OD-co-G-co-LA). For the comonomer feed ratio (OD:G:LA) 1:1:0.67, Mn reached ∼9500 in 8 h, trisubstituted G-units increased to 64%, and the monomer was well consumed. By varying the feed ratio of LA, polymeric triglyceride-type structures were formed with control of the chain length and trisubstituted G-unit content.850 Natural sugar polyols were used to prepare “sweet polyesters”. The polyols employed were C4-carbon, erythritol, C5-carbon, xylitol and ribitol, and C6-carbon, mannitol, glucitol, and galactitol. The ter-polymeriztion by Novozym 435 catalyst was performed in bulk for the combination of a polyol, adipic acid, and 1,8-octanediol under vacuum at 90 °C. The vacuum control during the reaction was very important to yield a high molecular weight polyester. The Mw value of the product polyol-polyester ranged from 1.1 × 104 (D-galactitol) to 7.3 × 104 (D-mannitol), having a branching structure. Primary alcohol groups are more reactive than the secondary groups.851 A dehydration polycondensation was effectively catalyzed by cutinase (EC 3.1.1.74). Glycols like 1,4-butanediol, 1,6hexanediol, 1,8-octanediol, and 1,4-cyclohexanedimethanol (1,4-CHDM), and diacids like adipic acid, succinic acid, suberic acid, and sebacic acid, were combined for the polycodensation with 1% w/w enzyme at 70 °C for 48 h under vacuum. In all reactions, the monomers were consumed quantitatively. With fixing the adipic acid component, polyesters from 1,4butanediol, 1,6-hexanediol, and 1,8-octanediol showed Mn values of 2700, 7000, and 12000, respectively. With fixing the 1,4-CHDM component, polyesters from succinic acid, adipic acid, suberic acid, and sebacic acid possessed Mn values of 900, 4000, 5000, and 19000 for these C4, C6, C8, and C10 diacids, respectively. For both glycols and diacids, there was a tendency that the higher the hydrophobicity, the higher the molecular weight of the product polyester.852 Lipases (immobilized Humicola insolens (HiC) and N435) on catalytic activity were studied with substrates varying in α,ω-n-alkane diol and α,ω-nalkane diacid chain length. Dehydration copolymerization of 2371


Chemical Reviews

Review

sebacic acid (C10-diacid) with α,ω-n-alkane diols with 3-, 4-, 5-, 6-, and 8-carbon chain lengths was C8 > C6, where C3, C4, and C5 diols were not polymerized. N435’s relative activity for diol substrates was in reactivity C8 = C6 = C5 > C4 > C3. HiC activity for copolymerizations of 1,8-octanediol with α,ω-nalkane diacids with 6-, 8-, 9-, 10-, and 13-carbon chain lengths was C13 = C10, where HiC showed little activity for C6, C8, and C9 diacid copolymerization. N435 displayed similar activity for all these diacid chain lengths. Thus, N435 has a broader substrate promiscuity than HiC.819 Biobased functional polyesters were synthesized from ωcarboxy fatty acid monomers 1,18-cis-9-octadecenedioic, 1,22cis-9-docosenedioic, and 1,18-cis-9,10-epoxy-octadecanedioic acids from oleic, erucic, and epoxy stearic acids by whole-cell biotransformations catalyzed by C. tropicalis ATCC20962.853 The conversion of ω-carboxy fatty acid and diol monomers to polyesters was carried out using the Novozym 435 catalyst, giving rise to corresponding polyesters with unsaturated and epoxidized repeat units and Mw values ranging from 25000 to 57000. The CALB-catalyzed synthesis of following aliphatic polyesters (PEs) was achieved, from diols (1,4-butanediol and 1,8-octanediol) and diacids or their derivatives (diethyl succinate, sebacic acid, 1,12-dodecanedioic acid, and 1,14tetradecanedioic acid), in order to produce poly(butylene succinate) (PE 4.4), poly(octylene sebacate) (PE 8.10), poly(octylene dodecanate) (PE 8.12), and poly(octylene tetradecanate) (PE 8.14).854 The two stage procedures were suggested, both sustainable and in accordance with the principles of “green” polymerization. The first comprised an enzymatic prepolymerization under vacuum, in the presence of diphenyl ether as solvent, whereas a low-temperature postpolymerization step [solid state polymerization (SSP)] followed in order to upgrade the PEs quality. In the first synthesized prepolymers, the Mn attained was from 3700 to 8000 with yields reaching even 97%. Next, SSP of PE 4.4 and PE 8.12 took place under vacuum or flowing nitrogen and lasted 10−48 h, at temperatures close to the prepolymer melting point (4∼14 °C). The solid state finishing led to an increase in the molecular weight, and it also contributed to the improvement of the physical characteristics and the thermal properties. The CALB-catalyzed synthesis of linear ester oligomers (LEOs) and cyclic ester oligomers (CEOs) from succinic acid in combination with dianhydro hexitols (DAH) in toluene were studied. The conversion is highest for isomannide and decreases in the order of isomannide (B) > isosorbide (A), isoidide (C) (Scheme 81). The maximum conversions under optimized conditions were 88.2% and 93.7% for succinic acid and isomannide, respectively. CEOs production was 32−48% yields.855

Several polyester thermosets based on photocurable prepolymers composed of itaconic acid (IA) and various polyols were developed by using CALB catalyst at 90 °C or by thermal dehydration reaction at 145 °C for future biomaterials. The reaction components were IA, a dicarboxylic acid like succinic acid and adipic acid, and a polyol like 1,4hexanedimethaol, a PEG-diol, and sorbitol. Dimethyl itaconate was an ideal monomer for enzymatic polymerization, as demonstrated by the synthesis of linear poly(1,4-cyclohexanedimethanol itaconate), poly(PEG itaconate), and poly(3-methyl-1,5-pentanediol itaconate-co-3-methyl-1,5-pentanediol adipate). Photolysis of the polyesters gave cured polyesters, whose physical properties were examined for proving the usefulness. It is claimed that itaconate-based polyesters are versatile and making them excellent candidates for future biomaterials.856 Starting from oleic diacid (with a reactive double bond) and glycerol, the synthesis and structure of poly(oleic diacid-coglycerol) that results by using Novozym 435 (N435) and dibutyl tin oxide (DBTO) as catalysts were compared. By using N435 catalysis and an oleic diacid to glycerol molar ratio of 1.0:1.0, the Mn values were 6000 at 6 h and 9100 at 24 h with a low-branching degree (% of glycerol 13%-16%). With N435 catalyst, resulting polyesters were not cross linked. In contrast, by using DBTO as catalyst and an oleic diacid to glycerol molar ratio of 1.0:1.0, polyester Mn of 1700 was obtained at 6 h and, thereafter, a gel was formed due to cross linking. Thus, due to the ability of N435 to deter cross linking owing to steric hindrance at the active site, soluble, hyperbranched copolyesters were obtained.857 In addition, the reaction of a linear polyanhydride, such as poly(azelaic anhydride), and a glycol, such as 1,8-octanediol, was induced with lipase CA catalyst at 30−60 °C in bulk or in toluene involving the dehydrating-insertion to give a polyester with molecular weight of several thousands (Scheme 82).858 Scheme 82

Via Transesterification. Polycondensation of this type normally needs activation of carboxylic acid groups. The activation is conducted ordinarily by esterification of the acid group as shown in Scheme 83. In the early studies, alkyl or Scheme 83

Scheme 81 haloalkyl esters and later vinyl esters have been often used. In 1989, a lipase-catalyzed high enantioselective polymerization was reported; the reaction of bis(2,2,2-trichloroethyl) trans-3,4epoxyadipate with 1,4-butanediol in anhydrous diethyl ether using porcine pancreas lipase (PPL) catalyst gave a highly optically active polyester. The feed molar ratio of the diester to the diol was adjusted to 2:1 so as to produce the (−)-polymer resulting in enantiomeric purity > 96%. The molecular weight 2372


Chemical Reviews

Review

Figure 34. Illustration of the synthesis processes. Reprinted from ref 860. Copyright 2015 American Chemical Society.

was estimated as 5300. The unchanged (+)-monomer was shown to have an enantiomeric purity higher than 95%.859 1,4-Butanediol (BD) and diethyl sebacate (DES) were copolymerized with bicyclic acetalized D-glucose derivatives (Glux) by polycondensation both in the melt at high temperature and in solution at mild temperature mediated by CALB (Figure 34). Two series of random copolyesters (PBxGluxySeb and PBSebxGluxy) were prepared differing in which D-glucose derivative (Glux diol or Glux diester) was used as the comonomer. Both methods were found to be effective for polymerization, although significant higher molecular weights were achieved by melt polycondensation. The thermal properties of the copolyesters were largely dependent on composition and also on the functionality of the replacing Glux unit.860 CALB-catalyzed polycondensation for the synthesis of polyester prodrugs of ketoprofen was studied, giving rise to the linear polyesters with pendent ketoprofen groups based on ketoprofen glycerol ester, poly(ethylene glycol), and divinyl sebacate. The polyester had an Mw reaching to 7000 and could be a promising prodrug with extended pharmacological effects by delayed release of ketoprofen.861,862 The ring−chain equilibrium of the product was noted, which is an important phenomenon. Polycondensation between dimethyl succinate and 1,6-hexanediol catalyzed by lipase CA in toluene at 60 °C reached the ring−chain (cyclic-linear structure) equilibrium of the product polymer. Adsorption of methanol by molecular sieves or elimination of methanol by nitrogen bubbling shifted to the thermodynamic equilibrium. Polyesters with the molecular weight about several thousands were produced from α,ω-alkylene dicarboxylic acid dialkyl esters, and regardless of the monomer structure, cyclic oligomers were formed (Scheme 84).709 In the polymerization of dimethyl terephthalate and diethylene glycol catalyzed by lipase CA in toluene for producing a terephthalate polymer, the distribution of the macrocyclic species obeyed the Jacobson− Stockmayer theory, in terms of ring−chain equilibrium.863 A cyclic dimer was selectively formed. The cyclic dimer content (%) in the products was dependent on reaction temperature and reaction time: at 50 °C, 2, 24, and 61% after 4, 8, and 24 h, respectively, and at 80 °C, 68, 80, and 99% after 4, 8, and 24 h, respectively. It was considered that among several factors the selective cyclic dimer formation is ascribed to the presence of a

Scheme 84

driving force because of a π−π stacking of the aromatic rings together with a relative flexibility of the diol segment.864 A chemo-enzymatic method was developed. Dicarboxylic acid dimethyl esters having unsaturated C18, C20, C26 alkylene chains were epoxidized via chemo-enzymatical oxidation with hydrogen peroxide/methyl acetate with lipase CA catalyst. Polycondensations of these dimethyl esters with a diol by the lipase catalysis gave the linear unsaturated and epoxidized polyesters with a molecular weight of 1950−3300 and a melting point of 47−75 °C from a 1,3-propanediol substrate and a molecular weight of 7900−11600 and melting point of 55−74 °C from a 1,4-butanediol substrate.865 Novozym 435-catalyzed synthesis of poly(butylene succinate) (PBS) via polycondensation was achieved using a monophasic reaction mixture of dimethyl succinate and 1,4-butanediol in bulk and in solution. Diphenyl ether was a preferred solvent to give a higher molecular weight PBS; at 60, 70, 80, and 90 °C after 24 h, Mn values of PBS were 2000, 4000, 8000, and 7000, respectively. The reaction at 95 °C after 21 h gave PBS with a Mn value of 38000.866 In an ionic liquid, a green solvent, such as 1-butyl-3-methyimidazolium tetrafluoroborate ([bmim][BF4]), a similar polycondensation between diethyl adipate or diethyl sebacate and 1,4-butanediol gave the polyester having Mn up to 1500 in good yields. Since the ionic liquid is nonvolatile, ethanol was removed under vacuum during the reaction. Lipase CAcatalyzed polycondensation of dimethyl adipate or dimethyl sebacate with 1,4-butanediol was performed in an ionic liquid such as [bmim][BF 4 ], [bmim][PF 6 ], and [bmim][(CF3SO2)2N] at 70 °C for 24 h to give a higher molecular 2373


Chemical Reviews

Review

weight polyester, Mn reaching several thousands. Using ionic liquids as solvents involves the wide range of tunability of solvent hydrophilicity and monomer solubility for the reaction.731,732 Concerning the structure of the alcohol component, transesterifications by lipase catalyst are affected and often very slow because of the reversible nature of the reactions. To shift the equilibrium toward the product polymer more effectively, activation of esters was conducted using a halogenated alcohol like 2-chloroethanol, 2,2,2-trifluoroethanol, and 2,2,2-trichloroethanol (Scheme 83). Compared with methanol or ethanol, they increased the electrophilicity of the acyl carbonyl and avoided significant alcoholysis of the products by decreasing the nucleophilicity of the leaving alkoxy group. Lipase PF-catalyzed reaction of bis(2-chloroethyl) succinate and 1,4-butanediol carried out in a mixed solvent of diisopropyl ether and chloroform at 37 °C gave the polyester with the highest Mn of 1570.867 Polycondensation of bis(2,2,2trichloroethyl) glutarate and 1,4-butanediol proceeded with the PPL catalyst at room temperature in diethyl ether to produce the polyesters with a molecular weight of 8.2 × 103.868 In the PPL-catalyzed polymerization of bis(2,2,2-trifluoroethyl) glutarate with 1,4-butanediol in 1,2-dimethoxybenzene, a periodical vacuum method for removing 2,2,2-trifluoroethanol from the reaction mixture increased the molecular weight to nearly 4 × 104.869 Also, the vacuum technique was effective to increase the molecular weight. The lipase-catalyzed polycondensation between bis(2,2,2-trifluoroethyl) sebacate and aliphatic diols was performed at 37 °C. The elimination of the product 2,2,2-trifluoroethanol was critical for obtaining the higher molecular weight polyesters; the polyester from 1,4butanediol reached the highest Mw of 46400.836 In a supercritical fluoroform solvent, polycondensation of bis(2,2,2-trichloroethyl) adipate with 1,4-butanediol using the PPL catalyst took place, giving rise to the polymer with a molecular weight of several thousands.870 A dynamic kinetic resolution (DKR) method was used to synthesize an optically active polyester from a racemic monomer via polycondensation.871 A mixture of stereoisomers of a secondary diol, α,α′-dimethyl-1,4-benzenedimethanol, was enzymatically polymerized with dimethyl adipate (Scheme 85).

days; during the reaction, molecular weight increased to 3000− 4000, and the optical rotation of the reaction mixture increased from −0.6° to 128°. Eventually, all product polymers will be end-capped with (R) stereocenters (chain-stoppers). An irreversible ester-formation process was disclosed; for this purpose, a divinyl ester was employed for the first time in 1994 as the activated acid form in the enzyme-catalyzed polyester synthesis. A vinyl ester proceeds much faster than an alkyl ester or a haloalkyl ester to form the desired product in higher yields, where the product of vinyl alcohol tautomerizes to acetaldehyde. Thus, the lipase PF-catalyzed polycondensation of divinyl adipate and 1,4-butanediol was performed at 45 °C in diisopropyl ether for 48 h to afford a polyester with the Mn of 6.7 × 103 with liberating acetaldehyde (Scheme 86), while the Scheme 86

use of adipic acid or diethyl adipate did not produce the polymeric materials under the similar reaction conditions. As a diol, ethylene glycol, 1,6-hexanediol, and 1,10-decanediol were also reacted to give the corresponding polyester with a molecular weight of several thousands.872 The same polycondensation of divinyl adipate and 1,4-butanediol with lipase PC (Pseudomonas cepacia) catalyst produced the polyester with a Mn of 2.1 × 104.873 By variation of the molar ratio of the divinyl ester and the glycol, a macromonomer having the glycol, the dicarboxylic acids, or the acid-alcohol end structure are to be obtained. During the lipase-catalyzed polymerization of divinyl esters and glycols, there was a competition between the enzymatic transesterification and hydrolysis of the vinyl end group, resulting in the limitation of polymer growth.874 A batch-stirred reactor was developed to minimize temperature and masstransfer effects. Using the reactor, the polycondensation became very fast; within 1 h at 60 °C, poly(1,4-butylene adipate) with the molecular weight of 2.3 × 104 was obtained.875 Aliphatic polyesters having pendant azide groups were prepared by CALB-catalyzed polycondensation. The grafting reaction to the N3-functional polyester was carried out quantitatively at room temperature using copper-catalyzed azide−alkyne cycloaddition (CuAAC, “click” reaction) with monoalkyne-functional poly(ethylene oxide) (alkyne-PEO, Mn = 750). Both enzymatic polycondensation and “click” reaction were carried out in a sequential one-pot reaction. The graft copolymer was surface-active and self-assembled in water.876 A combinatorial approach was applied for biocatalytic production of polyesters. A library of polyesters was synthesized in 96 deep-well plates from a combination of divinyl esters and glycols with lipases of different origin. In the screening, lipase CA was the most active biocatalyst for the polyester production. As an acyl acceptor, 2,2,2-trifluoroethyl esters and vinyl esters were examined and it was claimed that the former produced the polymer of higher molecular weight.877 Supercritical carbon dioxide (scCO2) was shown to be a good solvent for the lipase-catalyzed polycondensation of

Scheme 85

Because of the enantioselectivity of lipase CA, only the hydroxy groups at the (R) center preferentially reacted to form the ester bond with liberation of methanol. The reactivity ratio was estimated as (R)/(S) ∼ 1 × 106. In situ racemization from the (S) to the (R) configuration by Ru catalysis allowed the polymerization to high conversion, that is, the enzymatic polymerization and the metal-catalyzed racemization occurred concurrently. The DKR polymerization was carried out for 4 2374


Chemical Reviews

Review

bulk-polymerization of diesters and diols with various numbers of methylene groups in their chains. β-Cyclodextrin encircled the linear polymer chain and maintained the chain in a proper configuration to avoid its coagulation. Lipase initiated the polymerization and β-cyclodextrin threaded onto the polymer chain to control the structure for producing high molecular weight polyesters up to 62100 obtained at 70 °C. The corresponding polyesters showed an excellent thermal stability until 350 °C.883 Some of the above results accord with the frequent occurrence of an intermolecular transesterification between the resulting polyesters during the polymerization; from a mixture of two homopolyesters, a copolyester was obtained by lipase catalysis (Scheme 88).780

divinyl adipate and 1,4-butanediol. Quantitative consumption of both monomers was achieved to give the polyester with a Mn of 3.9 × 103.721 The lipase CA-catalyzed polycodensation of aromatic diacid divinyl esters was conducted to produce aromatic polyesters. Divinyl esters of isophthalic acid, terephthalic acid, and pphenylene diacetic acid were polymerized with various glycols to give aromatic polyesters with the highest Mn of 7200 in heptane.878 Enzymatic polycondensation of divinyl esters with aromatic diols also afforded the aromatic polyesters.879 Biodegradable and amorphous polyketoesters were prepared by the reaction of ketone-containing diesters and diacids, combined with di(ethylene glycol) through Novozym 435 catalyzed-polycondensation, achieving Mn values of up to 10.1 × 103. By including ketone groups in the repeat unit, facile postpolymerization modifications were possible by reaction with oxyamine-tethered ligands through the formation of an oxime linkage. Upon reaction with molecules that contain oxyamines, under mild conditions, these polymers can easily have a diverse set of side chains appended without coreagents or catalysts. The chemoselective oxime-forming coupling strategy is compatible with physiological conditions and can be done in the presence of a wide range of functional groups and biomolecules, including proteins and nucleic acids.880 CALB was used to synthesize silicone polyesters to give the polymers between 74 and 95% yields via polytransesterification (Scheme 87). A lipase from C. rugosa was only successful in

Scheme 88

The CALB catalyzed polymerization of various diacid ethyl esters with 2,5-bis(hydroxymethyl)furan, which is a highly valuable biobased rigid diol resembling aromatic monomers, via a three-stage method (Figure 35). Novel biobased furan

Scheme 87. Reprinted from ref 881. Copyright 2010 American Chemical Society

Figure 35. Synthesis of biobased furan polyesters. Reprinted from ref 884. Copyright 2014 American Chemical Society.

polyesters with Mn around 2000 were obtained. All product polyesters were semicrystalline. The degree of crystallinity ranged from 34 to 65%. The Tm was around 43−86 °C. These polyesters were thermally stable up to 250 °C, showing two decomposition steps. The Tmax at the major degradation step was around 276−332 °C. The Tg of 2,5-bis(hydroxymethyl)furan-based polyesters was around −38 to 4 °C, being significantly higher than that of the aliphatic counterparts.884 Isosorbide aliphatic polyesters were synthesized by the Novozym 435 catalyst. The use of diethyl ester derivatives of diacid comonomers gave copolyesters of isosorbide with highest isolated yield and molecular weights. The length of the diacid aliphatic chain was less restrictive but with a clear preference for longer aliphatic chains. The Mw values were in excess of 40000, which was unexpected, because isosorbide has two chemically distinct secondary hydroxyl groups. This is the first example in which isosorbide polyesters were synthesized by enzyme catalysis, opening large possibilities for the important class biomaterials.885 A cross-linkable group can be a mercapto group. Direct lipase CA-catalyzed polycondensation of 1,6-hexanediol and dimethyl 2-mercaptosuccinate at 70 °C in bulk gave an aliphatic polyester having free pendant mercapto groups with Mw =

performing esterifications using carboxy-modified silicones that possessed alkyl chains greater than three methylene units between the carbonyl and the dimethylsiloxy groups. The proteases α-chymotrypsin and papain were not suitable enzymes for catalyzing any polytransesterification reactions.881 Biobased commercially available succinate, itaconate, and 1,4butanediol were enzymatically copolymerized in solution via a two-stage method, using CALB as the biocatalyst. The chemical structures of the obtained products, poly(butylene succinate) (PBS) and poly(butylene succinate-co-itaconate) (PBSI) having reactive vinylene group, were confirmed. Values of Mn reached to ∼6500 in 90% yields.882 Diesters and diols were converted into aliphatic polyesters and polycarbonates by lipase Candida sp. 99−125 catalysis, with β-cyclodextrin acting as supporting architecture (in a similar way as chaperone proteins) without using organic solvents. The polytransesterification was a much greener process, being solvent-free and without metal residues. Lipase Candida sp. 99−125 showed a high catalytic activity for 2375


Chemical Reviews

Review

14000 in good yields. The mercapto group content could be controlled by copolymerization with other monomers. The polyester was readily cross-linked by the air-oxidation via the disulfide linkage formation in dimethyl sulfoxide.886 Reversibly cross-linkable aliphatic and aromatic polyesters with free pendant mercapto groups were prepared by lipase-catalyzed polycondensation of various diols and dimethyl mercaptosuccinate (Scheme 89). The CALB catalyzed reaction in an organic

Reactive polyesters were prepared by the lipase-catalyzed polycondensation of divinyl sebacate with glycerol in the presence of an unsaturated higher fatty acid (i) such as linoleic acid and linolenic acid obtained from renewable plant oils (route A, Scheme 90). Product polyester (iii) is biodegradable Scheme 90

Scheme 89

and contains an unsaturated fatty acid moiety in the side chain. Curing of (iii) induced by oxidation with cobalt naphthenate catalyst or thermal treatment gave a cross-linked transparent film. Biodegradability of the film was tested by biochemical oxygen demand (BOD) measurement.891,892 In addition, epoxide-containing polyesters were enzymatically synthesized via two routes, A and B. In route A, (iii) was enzymatically epoxidized to give (iv), and in route B the lipase-catalyzed epoxidization of the fatty acid (i) was first applied to give (ii) and lipase-catalyzed condensation polymerization of (ii) was performed to produce reactive polyester (iv). Curing of (iv) proceeded thermally, yielding transparent polymeric films with high gloss surface. Pencil scratch hardness of film (iv) from (ii) was higher than that from (iii). Both films showed good biodegradability.891−893 In a similar way, an unsaturated vegetable oil of oleic acid was epoxidized with lipase/H2O2, and the intermolecular ring-opening addition reaction between the carboxylic acid group−epoxy group gave poly(oleic acid), a thermally stable (up to 300 °C) material for general use.827 The CALB-catalyzed copolymerization of ethyl glycolate (EGA) with diethyl sebacate (DES) and 1,4-butanediol (BD) resulted in the formation of poly(butylene-co-sebacate-coglycolate) (PBSG) copolyesters with a Mw up to 28000 and typical polydispersity between 1.2 and 1.8 (Figure 36). This type of aliphatic copolyesters consisting of diester, diol, and glycolate repeat units were enzymatically synthesized for the first time. The synthesized copolymers contained 10−40 mol % glycolate (GA) units, depending on the monomer feed ratio. PBSG copolyesters were hydrolytically degradable and doxorubicin-encapsulated PBSG nanoparticles exhibited controlled release delivery of the drug.894 The CALB-catalyzed copolymerization of isosorbide (IS) or isomannide (IM) with diethyl adipate and fraction of different unsaturated diesters (diethyl itaconate, diethyl fumarate, diethyl glutaconate, and diethyl hydromuconate) gave unsaturated

solvent at 70 °C to produce the polyesters with Mw = 17000− 27000 in high yields without any reaction at the mercapto group. The polyesters were easily cross linked to form gels or films by air oxidation of the pendant free mercapto groups by forming disulfide bonds. The cross-linked polyesters were readily de-cross linked by reduction with tributylphosphine to regenerate polyesters with free mercapto groups almost quantitatively. For chemical recycling, the polyesters with free mercapto groups were depolymerized into cyclic oligomers, which were repolymerized to form polyesters with the same Mw and polymer structure as the initial polymers.887 Regioselctive polycondensation using polyols was achieved with divinyl esters. Triols such as glycerol were regioselectively polymerized at a primary hydroxy group with divinyl adipate by lipase catalyst to produce a linear polyester linked through mainly primary hydroxy group having also a secondary hydroxy group (5−10%) in the main chain. The polymer contained pendant hydroxy groups, with no evidence of network structure, having a Mw value from ∼3000 to 14000.888 The reaction of divinyl sebacate and glycerol with lipase CA catalyst produced water-soluble polyesters at 60 °C. From products, the chloroform-soluble part with a Mw of 19000 was isolated in 63%, which indicated the regioselectivity of primary OH/ secondary OH ratio of 74/26. At a lower temperature of 45 °C; however, the regioselectivity was perfectly controlled to give a linear polymer consisting exclusively of a 1,3-glyceride unit.889 The lipase CA catalysis gave a sugar-containing polyester regioselectively from divinyl sebacate and sorbitol, in which sorbitol was exclusively reacted at the primary OH group of 1and 6-positions in acetonitrile at 60 °C. Mannitol and mesoerythritol behaved similarly.890 2376


Chemical Reviews

Review

Figure 36. Illustration of the whole contents studied. Reprinted from ref 894. Copyright 2013 American Chemical Society.

postfunctionalize the polyester backbone. A change of the aza-Michael donor to polymeric amines, such as PEI, led to polymer−polymer coupling reactions. These resulted in material gelation and formation of hydrogels with tunable water uptake capacity.901 6.3.1.3. Ring-Opening Addition-Condensation Polymerization. A general reaction mode is given in Scheme 91, which is a hybrid type reaction of ring-opening polymerization and addition-condensation polymerization (ROACP).

polyesters in one-step. Unsaturated polyesters from IS and IM showed Mw values 4000−16000 when fumarate or glutaconate were used in 5 mol % for adipate.895 Amphiphilic dendronized polymers were synthesized by combining “chemo-enzymatic approach” and “click chemistry.” Dendronized and hyperbranched multifunctional amphiphilic polymers were prepared using varied amounts of propargylated alkyl chain with perfect and hyperbranched polyglycerol dendrons on the base copolymer of PEG (Mn = 1000) diethylester and 2-azidopropane-1,3-diol to examine their physicochemical properties.896 Modification of Monomers and Polymers. It is to be noted that an enzymatic reaction was utilized for the cascade reaction comprising: (1) Novozym 435-catalyzed transacylation reaction to prepare a new monomer, and (2) a free radical polymerization reaction (FRP) or a controlled radical polymerization (nitroxide-mediated polymerization, NMP). Various hydroxy functional polyacrylates were prepared by in situ lipasecatalyzed transacylation reactions of methyl acrylate (MA) with different diols and triols yielding a mixture of monoacrylates and bisacrylates. Up to 6 mol % of bisacrylate contained in the monomer mixture resulted to give soluble (noncross-linked) polymers by FRP, while bisacrylate concentrations of 10 and 12 mol % led to gel formation.897 The cascade reaction method was further developed and discussed. Transacylation of MA and methyl methacrylate with different functional alcohols, diols, and triols (1,2,6-hexanetriol and glycerol) in the presence of Novozym 435 led to functional (meth)acrylates. After the removal of the enzyme by filtration, removal of excess (meth)acrylate and/or addition of a new monomer, the (co)polymerization via FRP or NMP resulted in poly[(meth)acrylate]s with predefined functionalities. Hydrophilic, hydrophobic as well as ionic repeating units were assembled within the copolymer.898 Polymer end-groups were selectively modified via the green process.67 The transesterification of alkyl halo-esters with poly(ethylene glycol)s (PEGs) using CALB catalyst produced halo-ester-functionalized PEGs under the solventless conditions. Transesterifications of chlorine, bromine, and iodine esters with tetraethylene glycol monobenzyl ether were quantitative in less than 2.5 h.899 Chiral sec-alcohol groups in polymer side groups were enantioselectively esterified by CALB catalyst, which is strongly (R)-selective.900 CALB-catalyzed polycondensation between an itaconate and an oligoethylene glycol monomer gave a hydrophilic polyester backbone carrying an activated carbon−carbon double bond. These reactive sites served as aza-Michael acceptors. AzaMichael donors such as small molecular amines like piperidine, morpholine, imidazole, and diallylamine were used to

Scheme 91

Carboxylic Anhydride and Diol. It was reported in 1993 that a cyclic acid anhydride underwent ROACP with a glycol by lipase PF catalyst at room temperature, giving rise to a polyester having a Mn ∼ 2000 (Mw/Mn = 1.4) in good yields. The reaction involves the ring-opening addition as well as the dehydration (e.g., for R = −CH2CH2− and R′ = −(CH2)m− in Scheme 91.902 Various cyclic anhydrides, succinic, glutaric, and diglycolic anhydrides were polymerized by lipase CA catalyst with α,ω-alkylene glycols in toluene at 60 °C to give the polyesters with Mn reaching 1.0 × 104. This ROACP involving dehydration proceeded also in water and scCO2.903 This type of polymerization was recently extended to itaconic anhydride (IAn) as a new carboxylic anhydride monomer for lipase-catalyzed ROACP to produce reactive polyesters (Scheme 92).904 ROACP reaction of three components, IAn, succinic anhydride (SAn) or glutaric anhydride (GAn), and a diol at 25 °C in toluene, produced reactive polyesters in good to high yields. As diols, 1,4-butane, 1,6-hexane, 1,8-octane, and 1,10-decane diols were used. From the SAn reactions, polyesters with molecular weight (Mn) values of 650−3510 with 1.3−2.6 IAn units per molecule were obtained. From the GAn reactions, these values were 560−3690 and 1.2−3.1, respectively. Cross-linking reaction at the vinylidene group(s) of a product polyester showed reactive nature, giving a crosslinked hard solid polyester. These polyesters derived from renewable starting materials involve possible applications as macromonomer, telechelics, or cross-linking reagent, and the vinylidene group(s) can be used for further modification reactions. Regioselectivity at the carbonyl group of IAn was about the same (∼50%) for both α- and β-positions of IAn by lipase catalysis, whereas about 90% for β-selectivity with Sn(II) catalyst and without catalyst. ROACP between two components, IAn and a diol, did not give polyesters due to reduced reactivity of IAn.904 2377


Chemical Reviews

Review

Scheme 92

Cyclic Esters for Polycondensation. Terpolymerization of a lactone (in place of an acid anhydride), a divinyl ester, and a glycol produced an ester terpolymer with Mn higher than 1 × 104, which belongs to a ROACP (Scheme 93). Lipases showed

The monomer unit ratio in the copolymers could be controlled by adjusting monomer feed ratio, having nearly random distribution of PDL, butylene, and succinate repeat units, linked by ester groups along the macromolecule. Thermal stability of the copolymers is composition-dependent, increasing with increasing PDL unit content. All copolymers are highly crystalline irrespective of their composition. Mw values of the copolymers are 1.6−10.6 × 103 (Mw/Mn = 2.1−3.4).908 CALB was used to synthesize semicrystalline diepoxy functional macromonomers based on glycidol, pentadecalactone, and adipic acid. Macromonomers having epoxy-functions at both ends with molecular weight from 1400 to 2700 were obtained with conversion higher than 95% within 24 h at 60 °C. Macromonomers were used for photopolymerization, and copolymerized cationically with a cycloaliphatic diepoxide to afford a durable film with crystallinity.909 Cooperation between two hydrolases in the two-step tandem polymerization of bulky ibuprofen-containing hydroxyacid methyl ester (HAEP) and ε-CL catalyzed by CALB and Bacillus subtilis (BSP) was achieved (Figure 37) The tandem

Scheme 93

high catalytic activity for the terpolymerization involving both ring-opening and condensation polymerization simultaneously in one-pot to produce an ester terpolymer, without involving homopolymer formation.905 A similar terpolymerization was performed using three kinds of monomers, ω-pentadecalactone, diethyl succinate, and 1,4-butanediol, by CALB catalyst desirably at 95 °C via a two-stage vacuum technique. The polymerization was examined under various reaction conditions, and the product terpolyester reached a Mw = 77000 with Mw/Mn between 1.7 and 4.0.906 Copolymerization of isopropyl aleuriteate with ε-CL gave a random copolymer having a Mn up to 10600 in ∼70% yields in a way of ROACP by Novozym 435 catalyst.783 The CALB-catalyzed copolymerization of L-lactide (LLA) with diesters (diethyl adipate and dodecanedioate) and diols (1,6-hexanediol and 14-butanediol) was studied. Aliphatic lactate-bearing copolyesters were synthesized; the resultant copolymers had a Mw up to 38000, containing L-lactate units (up to 53 mol %), C6−C12 diester units, and C4−C6 alkylene units in the polymer chains. The lactate repeat units were present as lactate−lactate diads in the polymers. The LLAdiester-diol copolymers were purified in good yield (70−85%), and all purified copolymers were optically active. Hydrolytic degradation study showed that LLA-diethyl adipate-1,6hexanediol (LLA-DEA-HD) copolymers are degradable polymers as the molecular weight of the copolymer with 53% lactate units decreased by weight 70% upon incubation in phosphate buffer saline solution under physiological conditions for 80 days. The copolymers exhibit a wide range of physical properties (e.g., from white solid to wax and liquid) depending on their structure and composition.907 Poly(ω-pentadecalactone-co-butylene-co-succinate) copolymers with various compositions were synthesized via copolymerization of dialkyl succinate with 1,4-butandiol and ω-pentadecalactone (PDL, 16-membered) catalyzed by CALB.

Figure 37. Image of the synthesis procedures. Reprinted with permission from ref 750. Copyright 2014 Wiley-VCH.

process improved the molecular weight of the resultant ibuprofen-containing polyester from 3130 (CALB) and 720 (BSP) to 9200 (CAL-B/BSP). CALB mainly catalyzed ROP of ε-CL under the initiation of HAEP to form homopolymer of εCL with relatively low molecular weight, while BSP catalyzed the subsequent polycondensation of ROP product to yield copolymers with increased molecular weight. On the basis of the enzymatic cooperative tandem polymerization, valuable chiral (R)- or (S)-ibuprofen-containing polyesters were tailormade in high yields and ee values. The in vitro drug release behavior and degradation of the prepared chiral polymeric prodrug were also investigated.750 6.3.2. Synthesis of Other Polymers. In addition to polyesters, other polymers such as polyamides, polycarbonates, polyphosphates, polythioesters, etc. were synthesized by lipase catalysts. 6.3.2.1. Ring-Opening Polymerization. Lipase catalyzed the ring-opening polymerization of cyclic carbonates. Trimethylene 2378


Chemical Reviews

Review

chemically, was used as an initiator for lipase-catalyzed ROP of TMC to give a ABA type block copolymer. 2-Azetidinone (β-lactam) was polymerized via lipase CAcatalyzed ROP to afford linear poly(β-alanine) with a degree of polymerization = 8; and cyclic side products were formed,922 and ROP of substituted β-lactams was similarly performed with the lipase catalyst.923 The enzymatic ring-opening copolymerization of β-lactam and ε-CL by lipase CA catalyst was reported (Scheme 94B).924 A mixture of alternating and random copolymers as well as both homopolymers were detected in the product. A poly(ester-alt-amide) copolymer was prepared by the lipase-catalyzed ROP of a six-membered cyclic depsipeptide, 3(S)-isopropylmorpholin-2,5-dione (IPMD), in bulk (Scheme 94C). Lipase PS and PPL showed high catalytic activity to produce the polymer with a molecular weight up to 3 × 104, but lipase CA was almost inactive for ROP of IPMD. PPL catalyzed ring-opening copolymerization of IPMD with D,Llactide to produce a copolymer with a molecular weight of 1 × 104.925−927 A monomer of MD derivative, 6-(S)-methylmorpholin-2,5-dione, was also polymerized by lipase catalyst.928 ROP of a large cyclic ester-urethane oligomer was achieved by lipase CA to yield a poly(ester-urethane) with a molecular weight up to 1 × 105.673 A 5-membered cyclic phosphate was subjected to lipasecatalyzed ROP, yielding polyphosphate (Scheme 94D).929,930 A polymer with thioester linkage was synthesized by lipase CAcatalyzed ring-opening addition-condensation polymerization between ε-CL, and a mercaptoalkanoic acid under vacuum produced poly(ε-CL) containing thioester groups in the backbone.931,932 A 18-membered monomer of 1,6-hexanedithiol and sebacate was prepared and polymerized via ROP catalyzed by lipase CA in bulk in the presence of molecular sieves to give poly(thioester) with a molecular weight of 1.2 × 105 (Scheme 94E).933 This cyclic monomer was enzymatically copolymerized with the thioester monomer, giving rise to poly(thioester-co-ester). 6.3.2.2. Polycondensation. It is to be noted that lipase catalyzes the polycondensation to produce not only polyesters but also other polymers like polyamides, polycarbonates, and poly(thioester)s. Polyamide Synthesis. Lipases like Novozym 435 and Mucor miehei showed the high catalytic activities for polyamide synthesis.934 Typically, the reaction of dimethyl adipate and diethylene triamine produced the poly(aminoamide) with less branching at 50−110 °C with a Mn ∼ 3800 (Scheme 95). A

carbonate (6-membered, TMC) was polymerized by lipases BC, CA, PF, and PPL to produce the corresponding polycarbonate without involving elimination of carbon dioxide (Scheme 94A).910,911 The detailed tuning of the reaction Scheme 94

conditions enabled the increase of the molecular weight by adding a solvent and molecular sieves.912 High molecular weight poly(TMC) (Mw > 1 × 105) was obtained by using a small amount of PPL catalyst (0.1 or 0.25 wt % for TMC) at 100 °C.913 Cyclic dimers of carbonate were polymerized by lipase CA to produce polycarbonates with high molecular weight, which were also obtained by lipase-catalyzed polycondensation of 1,4butanediol or 1,6-hexanediol with diphenyl carbonate under the dilute conditions.914 Another cyclic carbonate dimer, 6,14dimethyl-1,3,9,11-tetraoxa-6,14-diaza-cyclohexadecane-2,10dione, was also subjected to lipase CA-catalyzed ROP to produce the polycarbonate with molecular weight up to 1.2 × 104.915 Lipase CA-catalyzed ROP of a 26-membered macrocyclic carbonate, cyclobis(decamethylene carbonate), was reported.916 Unlike in the case of lipase-catalyzed ROP of lactones with different ring size, the reactivity of cyclobis(decamethylene carbonate) was much lower than that of 6membered cyclic carbonate. An enantiomerically pure 7-membered cyclic carbonate having a ketal group, which was derived from L-tartaric acid, was polymerized by the lipase catalyst. Lipase CA showed efficient catalytic activity. Deprotection of the ketal group resulted in a hydroxyl group in the polycarbonate chain.917 ROP of cyclic carbonate oligomers and their ring-opening copolymerization with lactones were catalyzed by lipase CA, producing corresponding polycarbonates.918 Ring-opening copolymerization of a substituted TMC with 1,4-dioxan-2-one or with TMC by lipase catalyst was also reported to give a poly(carbonate-co-ester) or a substituted poly(carbonate).919,920 A degradable carbonate copolymer with micelle formation ability for pH-dependent controlled drug release was synthesized via a chemoenzymatic route.921 Poly(PEG-co-cyclic acetal) (PECA), an α,ω-glycol synthesized

Scheme 95

chemical method to prepare similar polymers needs a higher reaction temperature of ∼180 °C to give the polymer with more branching having a Mn ∼ 3000. Other carboxylic acids such as malonic acid and fumaric acid were used to prepare similar polymers via dehydration. As amines, triethylene tetramine, tetraethylene pentamine, and triethylene glycol diamine, were also employed. 2379


Chemical Reviews

Review

Figure 38. Outline of the lipase-catalyzed synthesis of aliphatic-aromatic polyamides. Reprinted from ref 936. Copyright 2013 American Chemical Society.

The CAL-B-catalyzed polycondensation between β-alanine esters gave poly-β-alanine. For the reaction, the effect of different solvents, reaction temperature, and substrate/enzyme concentration on polymer yield and DP was determined. Also the effect of Me substituents at the α and β position of βalanine was studied. Cross-linked enzyme aggregates of CALB converted β-alanine esters into polymers in high yield (up to 90%) and with high DP (up to 53 units). Optimum results were obtained when Me esters are used in Me t-Bu ether as a solvent, at 50 °C for 16 h.935 Enzyme-catalyzed polycondensation to lead to aliphatic−aromatic oligoamides was studied (Figure 38).936 The reaction of p-xylylenediamine and diethyl sebacate resulted in oligo(pxylylene sebacamide) with high melting temperatures (223−230 °C), and the enzymatic polycondensation of dimethyl terephthalate and 1,8-diaminooctane led to oligo-(octamethylene terephthalamide) with two melting temperatures at 186 and 218 °C. No oligoamides, but desired products, were formed from the enzymatic reaction of dimethyl terephthalate and p-xylylenediamine. All reactions were catalyzed by CAL-B, icutinase, or CLEA cutinase. All reactions catalyzed by CAL-B showed higher conversion than reactions catalyzed by icutinase or CLEA cutinase. The highest DPmax n = 15 was achieved in a one-step and two-step synthesis of oligo(p-xylylene sebacamide) catalyzed by CLEA cutinase.936 Polyamine Synthesis. CALB was found to be an efficient catalyst for copolymerization of diesters with amino-substituted diols to form poly(amine-co-esters) in one step via the ester linkage formation.934 Poly(thioester) Synthesis. Direct polycondensation of 11mercaptoundecanoic acid (11-MU) with lipase CA catalyst in bulk in the presence of molecular sieves gave an poly(thioester). The reaction at 110 °C for 48 h produced poly(11-MU) with a Mw of 34000 in high yields. The Tm value of 104.5 °C was about 20 °C higher than that of the corresponding polyoxyester. Poly(11-MU) was readily transformed by lipase into the cyclic oligomers mainly of the dimer, which were readily repolymerized by lipase via ROP as sustainable chemical recycling.932 A poly(thioester) was prepared by lipase CA catalyst via the direct transesterification of 1,6-hexanedithiol and a diester with eliminating ethanol in bulk (Scheme 96, m = 1−8). The product poly(thioester) possessed a Mw ∼ 1.0 × 104. Both the melting point and crystallization temperature were higher than those of the corresponding poly(oxyester)s.937 The lipase CA catalyzed ring-opening addition-condensation polymerization (ROACP) of ε-caprolactone (ε-CL) with 11mercaptoundecanoic acid or 3-mercaptopropionic acid under

Scheme 96

reduced pressure produced the copolymer of an ester-thioester structure with a molecular weight higher than 2 × 104. The transesterification between poly(ε-CL) and 11-mercaptoundecanoic acid or 3-mercaptopropionic acid was also observed.931 Polycarbonate Synthesis. The lipase catalyzed polycondensation of a carbonic acid diester and a glycol gave polycarbonates; the simplest reaction mode is given as Scheme 97. The reactions produced aromatic polycarbonates of DP Scheme 97

greater than 20879 and aliphatic polycarbonates of molecular weight higher than 4 × 104.938,939 Transesterification between diethyl carbonate and a diol to produce polycarbonates was investigated under various reaction conditions. It proceeded via two stages: the first to yield oligomers and the second to give higher molecular weight polymers.940 The CALB-catalyzed transesterification among three components, diethyl carbonate, a diester, and a diol, formed aliphatic poly(carbonate-co-ester)s with about a 1:1 molar ratio of the ester-to-carbonate repeat units. Molecular weight, Mw, value reached 59000 at a reaction temperature of 90 °C. A carbonate-ester transesterification reaction between poly(butylene carbonate) and poly(butylene succinate) was also catalyzed by CALB at 95 °C to give a copolymer of block type.941 Lipase CALB was used to promote synthesis of aliphatic poly(carbonate-co-ester) copolymers from dialkyl carbonate, diol, and lactone monomers via ROACP type reaction. The copolymerization was carried out in two stages: the first-stage oligomerization under low vacuum, followed by the secondstage polymerization under high vacuum. Therefore, copolymerization of ω-pentadecalactone (PDL), diethyl carbonate (DEC), and 1,4-butanediol (BD) yielded PDL-DEC-BD 2380


Chemical Reviews

Review

copolymers with a Mw of whole product up to 33000 and Mw/ Mn between 1.2 and 2.3, in up to 92% yields. Desirable reaction temperature was ∼80 °C. The copolymer compositions, in the range from 10 to 80 mol % PDL unit content versus total PDL carbonate units, were effectively controlled by adjusting the monomer feed ratio. The copolymers possessed near random structures with all possible combinations of PDL, carbonate, and butylene units via either ester or carbonate linkages and thus are more appropriately named as poly(PDL-co-butyleneco-carbonate) (Scheme 98).942

Enzymatic grafting of polymers on cellulose has also been performed to produce cellulose-graf t-polymer composites. An approach to introduce polymers to cellulosic materials was developed by using the ability of a cellulose binding moduleCALB conjugate to catalyze ring-opening polymerization of εCL in close proximity to cellulose fiber surfaces.680 ε-CL was introduced to the cellulose surfaces either by simple addition of liquid monomer or through the gas phase. The hydrophobicity of the resulted surface did not arise from the covalently attached poly(ε-CL) (PCL) to the surface OH groups but rather from surface-deposited polymers, which could be easily extracted. Graft composites of ethyl cellulose main chain with the P3HB side chain was synthesized through lipase-catalyzed esterification.947 SEM analyses of the composites revealed that the uniform P3HB dispersion in the main chain increased the area of P3HB-ethyl cellulose contact, which further contributed to the efficient functionality. Unreacted P3HB was hydrophobic in nature, whereas the composites attained higher levels of hydrophilicity. Water-soluble cellulose ethers, mostly carboxymethyl cellulose (CMC) and hydroxyethyl cellulose (HEC), have also been employed as substrates for lipase-catalyzed modification.948 For example, the enzymatic acetylation of CMC was conducted by Aspergillus niger lipase-catalyzed transesterification using vinyl acetate in both aqueous and organic media.949−951 The degree of acetylation was very low in aqueous buffer, whereas it was remarkably improved in a mixed solvent of DMSO and paraformaldehyde. The absence of bulk water and the better solubility of CMC against the organic solvents were considered to be a main reason for the improvement. The lipase-catalyzed synthesis of acetylated, stearated, and succinated HECs were performed using vinyl acetate, vinyl stearate, and succinic anhydride, respectively.952 The lipasecatalyzed enzymatic method was extended to graft alkyl ketene dimers onto HEC, which are widely used as sizing agents on paper.952 The lipase-catalyzed ring-opening graft polymerization of ε-CL was also carried out. When the HEC film was incubated with ε-CL and porcine pancreatic lipase at 60 °C for 3−5 days, PCL-grafted HECs with DSs between 0.10 and 0.32 in terms of per anhydroglucose unit were obtained.765 Lipase-catalyzed modification has extensively been demonstrated to produce various starch esters. Esterification of cassava starch with fatty acids, such as lauric and palmitic acids, catalyzed by lipase was carried out in a liquid state with DMSO/DMF solvents.953,954 For example, the liquid state esterification of cassava starch with palmitic acid using lipase obtained from Candida rugosa for 4 h at 70 °C in DMSO gave a degree of substitution of 1.05. Ionic liquid mixtures, 1-butyl-3methylimidazolium acetate and 1-butyl-3-methylimidazolium tetrafluoroborate, were also used as solvents for lipase-catalyzed synthesis of starch palmitate.955 The solvent-free esterification of starch with fatty acids, on the other hand, has been carried out by employing immobilized lipases such as Novozym-435 and CaCO3-immobilized lipase with and without microwaves.956,957 Microwave-assisted esterification of maize and cassava starches were also demonstrated using recovered coconut oil and microbial lipase.958 DSs for maize starch and cassava starch were 1.55 and 1.1, respectively. Rosin acid, a major component of natural resin, is a useful reagent for polymer modification. The enzymatic synthesis of rosin acid starch was carried out by lipase-catalyzed esterification of cassava starch with rosin acid in DMSO.959,960 Enzymatic modification of starch was also

Scheme 98

Lipase CALB-catalyzed condensations were reported to prepare high purity, metal-free, polycarbonate polyols. Terpolymerizations of diethyl carbonate (D) with 1,8octanediol (O) and tris(hydroxymethyl)ethane (T) were performed in bulk at a temperature of 80 °C using a reduced pressure. With D/O/T monomer feed ratio 3:0.9:0.1, the highest values of dendritic T-unit content (83%) and Mw (23900) by SEC-MALLS were attained. At short reaction times (e.g., 4 h), highly functional linear terpolymers were formed. Increase in reaction time from 4 to 8, 12, 24, and 30 h resulted in increased dendritic unit content (0−48%), Mw/Mn (1.5−5.6), and relative Mw (2100 to 39000).943 6.3.3. Modification of Polysaccharides. Enzymatic modification of hydroxy groups on cellulose and its derivatives have widely been investigated via hydrolase-catalyzed dehydrative condensation or transesterification. Immobilized lipase from Candida antarctica (Novozym 432)-catalyzed enzymatic modification of cellulose acetate and hydroxypropyl cellulose, for example, has been conducted to prepare various cellulosefatty acid esters.944,945 The acylation was progressed either in the free hydroxy groups of the cellulose acetate by direct esterification or in the acetyl groups via transesterification. A strategy for the modification of cellulose fiber surfaces was developed that used the ability of Candida antarctica lipase B (CALB) to allow the acylation of carbohydrates with high regioselectively, combined with the transglycosylating activity of the Populus tremula x P. tremuloides xyloglucan endotransglycosylase 16A (PttXET16A).946 Xyloglucan oligosaccharides (XGOs) prepared from tamarind xyloglucan were acylated with vinyl stearate or γ-thiobutyrolactone as an acyl donor in the presence of CALB to produce XGOs with hydrophobic alkyl chains or reactive thiol groups, respectively. The resulting XGO derivatives acted as glycosyl acceptors in the PttXET16Acatalyzed transglycosylation reaction. Therefore, the modified XGOs exhibited a high affinity for cellulose surfaces, which enabled the essentially irreversible introduction of fatty acid esters or thiol groups to cellulose fibers. 2381


Chemical Reviews

Review

conducted by lipase-coupling esterification with octenyl succinic anhydride in an aqueous slurry system since water activity surrounding starch granules decreased sharply due to its hydrating swell.961 The lipase-catalyzed reaction of starch and alkyl and alkenyl ketene dimers (AKD) was conducted to produce hydrophobically modified starches.962 The enzymatic reaction gave the products with higher DSs at lower temperatures. The properties of the AKD-starch adducts depended on the DSs. Regioselective modification of starch nanoparticles was performed using CALB.963 The starch nanoparticles were made accessible for acylation reactions by formation of AerosolOT [AOT, bis(2-ethylhexyl)sodium sulfosuccinate] stabilized microemulsions. Starch nanoparticles in microemulsions were reacted with vinyl stearate, ε-caprolactone, and maleic anhydride at 40 °C for 48 h to give starch esters with degrees of substitution of 0.8, 0.6, and 0.4, respectively. Substitution occurred regioselectively at the C-6 position of the repeating glucose units. Konjac glucomannan is a polysaccharide composed of β(1→ 4)-linked D-glucose and D-mannose, which has short side branches at the C-3 position of the mannose units and acetyl groups randomly at the C-6 position. In a solvent-free system with vinyl acetate, C-6 hydroxy groups of konjac glucomannan were acylated regioselectively by the catalysis of Novozym 435.964 Enzymatic synthesis of chitin- and chitosan-graf t-aliphatic polyesters was performed by lipase-catalyzed graft polymerization of β-butyrolactone and ε-caprolactone onto chitin and chitosan in bulk at 70 °C.788 The reactivity of ε-caprolactone was found to be higher than β-butyrolactone. All the graft polymers prepared were insoluble in common organic solvents.

Scheme 99

Scheme 100

tion is explained similarly to that of lipase as seen in Figure 39. These observations imply that the acylation step proceeds almost in the same rate both in the hydrolysis and oligomerization, and thus, in the oligomerization the deacylation step governs the preferential L-enantioselection. The opposite enantioselection between protease and lipase was argued in the PLA depolymerizing hydrolysis.965,968 These two enzymes belong to serine hydrolases, possessing a catalytic triad of serine, histidine, and aspartic acid; the catalytic-active site of the two class, however, is of topological mirror image structure.965,969−971 This catalyst site situation was considered responsible for the opposite selection, where protease is PLLApreferential and lipase is PDLA-specific.968 The enantioselection of Novozyme 435 was perfect. That of proteases was less selective. This selectivity difference is probably because in living systems the substrate of lipase is an ester having an ester linkage like RLa, whereas the substrate of protease is a protein having an amide linkage. Protease-catalyzed modification of polysaccharides were reported to give ester products. The regioselective ester-bond formation by the protease catalyst was noted for the modification of polysaccharides by vinyl esters to give acylated amylose at the 6-position.972 Transesterification of vinyl propionate and vinyl acrylate on cellulose solids was also performed in pyridine using protease type VIII (EC 3.4.21.62).973 The enzyme catalyzed the regioselective transesterification targeted to the primary C-6 hydroxy groups of repeating glucose units. The resulting cellulose esters were hydrolyzed partially by the same enzyme in aqueous media and were thus biodegradable. Surface grafting of the resulting cellulose acrylate was further demonstrated using free radical polymerization of acrylonitrile in DMF. The transesterification of vinyl acrylate on HEC was also achieved by protease catalysis.974 Dextrin acrylate with different degrees of substitution ranging from 10% to 70% was synthesized by enzymatic transesterification from vinyl acrylate to dextrin catalyzed by Bacillus subtilis protease Proleather FG-F.975 A free radical polymerization of dextrin-vinyl acryate initiated by the ammonium persulfate/N,N,N′,N′-tetramethylethylenediamine

6.4. Proteases

Proteases (EC 3.4) are hydrolysis enzymes, primarily catalyzing the hydrolysis of proteins for L-amino acid residues.965 They catalyze also the peptide bond formation, which catalyze both the bond cleavage via hydrolysis and the bond formation leading to polymer production.966 6.4.1. Synthesis of Polyesters. Some of proteases exhibited esterase activity to produce polyesters. Polycondensation of bis(2,2,2-trifluoroethyl) adipate with sucrose catalyzed by an alkaline protease from Bacillus sp. resulted in an alternating linear polyester soluble in water and polar organic solvents with a Mn = 1600 (Mw/Mn = 1.31). This enzyme allowed for polymers with molecular weight in excess of 13000. It was claimed that sucrose reacted regioselectively at 6- and 1′positions and behaved like a diol; cross-linking did not take place (Scheme 99).966 Protease was effective for aromatic polyester synthesis: a terephthalic acid diester and 1,4butanediol produced oligomers.967 Protease catalyzed the oligomerization of alkyl D- and Llactates (RDLa and RLLa) (Scheme 100).829 The examined four protease catalysts gave preferentially oligo(L-lactic acid)s (oligoLLa)s; dimer ∼ pentamer), with moderate to high yields. The enantioselection was L-/D-selective (56/28 to 25/4 in conversion % ratio). L-Enantioselection of the protease is opposite in that of the lipase (Scheme 76),828 but it is less strict compared with that of lipase, whose D-enantioselection is perfect. In the hydrolysis reaction of ethyl D- and L-lactates (EtLa)s catalyzed by protease, EtLLa was consumed a little faster than EtDLa. The mechanism of the protease-catalyzed oligomeriza2382


Chemical Reviews

Review

Figure 39. An illustrative mechanism of protease-catalyzed L-enantioselective oligomerization of alkyl L,D-lactates. Reprinted from ref 829. Copyright 2011 American Chemical Society.

(EC 3.4.22.2), bromelain (EC 3.4.22.4), and α-chymotrypsin (EC 3.4.21.1) showed a high catalytic activity. No γ-peptide unit was formed, indicating a perfect regioselectivity for αlinkage. For comparison, the protease-catalyzed polycondensation of γ-methyl L-glutamate was performed with expecting γlinkage formation under similar reaction conditions; however, no reaction took place (Scheme 101B). Again, protease catalysis was specific for forming α-linkage. Papain-catalyzed copolycondensation of L-glutamic acid diethyl ester with several L-amino acid esters like esters of L-alanine, L-tyrosine, and Laspartic acid produced copoly(amino acid)s in good yields. The copolymers had the perfect α-linkage structure with an average degree of polymerization (DP), n = 8−16 (Scheme 101C).978 It was also reported that papain-catalyzed polycondensation of Lglutamic acid diethyl ester gave similar results as above.978 The products polyamides having α-linkage structure showed n value less than 13, and mass spectroscopic analysis indicated the accumulation of dimer products at the initial stage.979 Polycondensation of L-tyrosine ethyl ester hydrochloride with the papain catalyst was conducted in a phosphate buffer (pH 7) at 40 °C for 3 h, to give a polymer of α-peptide structure, poly(tyrosine), with a molecular weight around 2000 (Scheme 101D).134 Poly(tyrosine) was soluble in DMF, DMSO, and in alkaline solution but insoluble in acetone, chloroform, toluene, THF, and water. The product poly(tyrosine) after 72 h showed the formation of the globular particle in the diameter larger than 50 μm. The cross section of the fragment showed that rodlike crystals radially originated from the center.976 This morphology of poly(tyrosine) is characteristic of the enzymatic polymerization of the monomer, since such a globular crystal was not observed before in the recrystallization of poly(α-amino acid)s including poly(tyrosine).980 Polycondensation of L-tyrosine ester was again catalyzed by papain in aqueous media to produce an oligotyrosine peptide, which showed angiotensin l-converting enzyme inhibitory activity. During the reaction, di-, tri-, and tetra-tyrosine accumulated in the soluble fraction, while oligomers of pentamer∼decamer were insoluble. The initial substrate concentration affected the yield of these fractions.981

system in water was further investigated to give dextrin hydrogels. It is to be noted that these protease-catalyzed reactions to give ester products are rather rare in organic chemistry.976 6.4.2. Synthesis of Polyamides. In nature, protease is an enzyme to catalyze hydrolysis of amide bonds of proteins.829,965,976 Proteases are also known to catalyze reverse reactions, forming peptide bonds under selected conditions.966 Thus, various poly(amino acid)s were synthesized by proteasecatalyzed polymerization of amino acids or their derivatives as seen below. Protease-catalyzed polycondensation of L-glutamic acid diethyl ester proceeded at 40 °C for 3 h in a phosphate buffer (pH 7) to give regioselectively an α-peptide with the n value = 7−9 in 61% yields and specific rotation [α]589 was −5.1° (DMSO. c = 10) (Scheme 101A).977 Proteases, such as papain Scheme 101

2383


Chemical Reviews

Review

polymerization by using papain as a catalyst. Products, block and random oligo(L-lysine-co-L-alanine), were obtained in a relatively short (30 min) and efficient reaction (over 40% yield). It is to be noted that the strong preference of papain for hydrophobic amino acids resulted in complete conversion of LAla-Et after only 15 min. Optical microscopy revealed differences in the crystalline morphology between random and block co-oligopeptides, showing possibilities to form materials for both tissue engineering and controlled drug/ gene delivery systems.988 An enzyme, α-chymotrypsin or poly(ethylene glycol) (PEG)modified papain, catalyzed the oligomerization of dialkyl Laspartate to produce α-linked poly(aspartate) (α-PAA), an αlinked polypeptide (Scheme 103, reaction A).989 Enzymatic

The protease-catalyzed synthesis of oligo(L-lysin) from Llysine (L-Lys) ethyl ester (L-Lys-Et) was achieved in an aqueous reaction medium (Scheme 102A). Four proteases (papain, Scheme 102

bromelain, α-chymotrypsin, and trypsin) were employed to determine their activity for L-Lys-Et oligomerization at pH values ranging from 6 to 11. Oligo(L-Lys)s were water-soluble. Bromelain was found to be preferred because it gave the highest values of oligo-(L-Lys) yield and optimal average chain length (DPavg); the DPavg and longest oligomer chain lengths reached maximum values of ∼3.6 and 12.0, respectively.982 The direct oligomerization of L-lys-OMe by bromelain catalysis gave oligo(L-lys) with a DPavg ∼ 3.6. For higher chain length oligo(Llys), reactive NH2 was protected with tert-butoxycarbonyl (Boc) or benzyloxycarbonyl groups. By using these monomers, the yield and DPavg of oligo(L-lysin) increased to 91% and to ∼7.5, respectively (Scheme 102B).983 L-Alanine ethyl ester was polymerized to poly(L-alanine) (polyL-Ala), one of the insoluble polypeptides by papain catalyst in aqueous buffer (Scheme 102C). At neutral pH, a maximum chain length of 11 was obtained. These polymers were dominated by random coiled structure and demonstrated a lack of patterned macromolecular assembly. Under alkaline conditions, longer polymer chain lengths were achieved, and the maximum chain length was 16 repeats. These longer chains showed distinct β-sheet formation and were capable of fibril assembly. These results showed the chemoenzymatic synthesis of a hydrophobic homopolypeptide under aqueous conditions as well as demonstrates a chain length dependency of secondary structure formation and macromolecular assembly of chemoenzymatically synthesized polyAla.984 In addition, three proteases were encapsulated in silica-nanoparticles, which showed an increased thermal stability in catalysis for both hydrolysis and aminolysis. They were employed as catalysts for poly(L-Ala), poly(L-Leu), and poly(L-Val) synthesis.985 The chemoenzymatic synthesis of oligo(L-phenylalanine) mediated by proteinase K from Tritirachium album was studied (Scheme 102D).986 The synthesized linear oligo-phenylalanine showed a unique self-assembly in aqueous solutions. To further functionalize linear oligo(L-phenylalanine) as a low-molecularweight gelator, it was cosynthesized with tris(2-aminoethyl)amine to obtain star-oligo(L-phenylalanine), which was bioconjugated to demonstrate its self-assembly into fluorescent fibers. The self-assembled fibers of star-oligo(L-phenylalanine) formed fibrous networks with various branching ratios. Four proteases (papain, bromelain, α-chymotrypsin, and protease SG) were used to catalyze co-oligomerizations of Lleucine ethyl ester (L-Et-Leu) with diethyl-L-glutamate (L(Et)2-Glu) to produce statistically random oligo(L-Glu-co-LLeu) of α-linkage with DPavg ∼ 7 units. There were not distinct differences of catalysis specificity (e.g., substrate selectivity) observed among four proteases.987 Amphiphilic diblock cooligopeptides were synthesized via one-pot, chemo-enzymatic

Scheme 103

polymerization of diethyl L-aspartate was catalyzed by alkalophilic proteinase from Streptomyces sp., giving rise to the polymer consisting of 88% α-linkage and 12% β-linkage (Scheme 103, reaction C).990 Exclusively α-linkage-containing polypeptide, α-PAA, was obtained by the catalyst of a protease from Bacillus subtilis (Scheme 103, reaction A).991 Papain catalyzed the polymerization of diethyl L-aspartate.992 Hydrolase-1 from PAA was modified with PEG to prepare an active catalyst; PEG-modified PAA hydrolase-1, which was well dispersed in an organic solvent. The catalyst facilitated to produce perfectly structure-controlled β-PAA consisting of exclusive β-amide linkage for the first time (Scheme 103, reaction B, R = Et). The product β-PAA had a molecular weight range of 750−2500. In contrast, native PAA hydrolase-1 did not cause the polymerization of diethyl L-aspartate, which was explained due to the low dispersibility of the enzyme in an organic solvent. Adhesive peptide, a blue mussel (Mytilus edulis) foot protein 5 (Mefp-5) composed mainly of glycine, L-lysine, and 3,4dihydroxy-L-phenylalanine (DOPA), was synthesized via two enzymatic reactions: chemo-enzymatic synthesis of copolypeptides of L-tyrosine and L-lysine by papain catalyst, followed by the enzymatic conversion from L-tyrosine to DOPA by tyrosinase catalyst (Figure 40).993 DOPA and lysine are essential for biomimetic adhesive design, and the multiple reaction steps was required for the synthesis of the copolymers. On the basis of the adhesion tests using the synthesized peptides, consisting DOPA, L-lysine, and L-tyrosine, at various pH, with different protonation/deprotonation states, was proposed a mechanism whereby deprotonated DOPA can interact with the surface materials, thus functioning as an adhesive molecule, while on the other hand, the primary amine 2384


Chemical Reviews

Review

Figure 40. Reaction schemes for the enzymatic synthesis of poly(L-tyrosine-γ-3,4-dihydroxy-L-phenylalanine-γ-L-lysine) [P(Tyr-DOPA-Lys)], via poly(L-tyrosine-γ-L-lysine) [P(Tyr-Lys)], from L-tyrosine ethyl ester (Tyr-Et) and L-lysine ethyl ester (Lys-Et). The images show the reaction intermediate, (a) P(75%Tyr-25%Lys), and products, (b) P(50%Tyr-25%Lys-25%DOPA) and (c) P(45%DOPA-30%Tyr-25%Lys). Reprinted from ref 993. Copyright 2014 American Chemical Society.

characteristics (value-added, biodegradable, etc.), reaction solvents to minimize damage of environment, as well as polymer recycling and degradation for reserving natural resources. It is to be noted that an enzyme catalyzes to induce a polymerization reaction in vitro, so far as a substrate, in most cases “monomer”, is recognized. The in vitro catalysis is often a reverse reaction of the inherent in vivo reaction as typically exemplified by a hydrolase catalysis. The repetition of the bondforming reaction ends up with the formation of a polymer. These observations are very specific to the enzymatic polymerization. The recognition of a substrate by enzyme brought about a new concept, a “transition-state analogue substrate” (TSAS) monomer as seen in the cellulose or chitin synthesis. The concept broadened the applicability of the enzymatic polymerization, giving rise not only to naturally occurring polymers but also to various non-natural polymers; hyaluronidase catalysis is another good example termed as a supercatalyst, showing a wide spectrum for monomer recognition to enable the preparation of a number of natural and non-natural polymers having very complicated structures. It is notable that hyaluronan, chondroitin, and their derivatives have one of the most complicated structures of the ever synthesized polymers in vitro. Further, almost all of polyester or aromatics polymer products are non-natural, which are quite common in variations of polymer structure by the enzyme catalyst. So far, research works of finding new reactions to afford new polymers, which otherwise were not possible to synthesize, have been actively developed. Mechanistic studies have also proceeded in accord with the developments of 3D-structure elucidation of enzymes in particular in the 1990s. Fundamental studies in this area have been widely accumulated. For the practical applications, an enzymatic polymerization process for polymeric materials productions has not been adopted, to our knowledge, on a definite commercial process, except for small scale commercial products of artificial urushi.998 This is probably due to a factor of enzyme cost or properties of the products; enzymes still seem not cheap enough for the products. However, factors of natural resources and environmental problems are increasingly operative; much hoped for will be the value-added functional polymer products derived from an enzymatic process. Various functional polymers

group of lysine induces molecular networks under deprotonated conditions. Previously, oligopeptides having various functions were synthesized using protease catalyst via condensation reactions in either a thermodynamically controlled or a kinetically controlled manner.994 For example, aspartame (Asp-Phe) by thermolysin,995 oxytocin (Cys-Tyr-Tyr-Ile-Pro-Leu-Leu-Gly) by papain, thermolysin, and chymotrypsin,996 and somatostatin (cyclic,-Ala-Gly-Cys-Lys-Phe-Phe-Trp-Lys-Thr-Phe-Thr-SerCys-) by thermolysin and chymotrypsin997 were prepared as typical biologically active peptides. Several proteases like α-chymotrypsin, trypsin, subtilisin, and papain were examined for catalysis of the polycondensation for the synthesis of various polyaminoamides from dicarboxylic acids and amines, giving rise to lower molecular weight polymers. This is rather interesting when compared with the results that lipase catalyst gave higher molecular weight polyamides because it is generally thought that protease is an appropriate catalyst for polyamide synthesis while lipase is for polyester synthesis.934

7. CONCLUDING REMARKS We have described a comprehensive review, with placing the enzyme as the first and the reaction or the product macromolecules as the second for the section formation axis. Three classes of enzymes, oxidoreductases, transferases, and hydrolases (EC 1, 2, and 3, respectively), among six classes have been used for the catalyst of in vitro “enzymatic polymerization” and, hence, constitute the main three axes. The present article is focused on recent developments of the enzymatic polymerization, covering research accomplishments mainly for these three decades. As seen above, the enzymatic polymerizations brought about remarkable achievements owing to various new findings of enzymatic catalysis. Enzyme catalysts are natural renewable and environmentally benign. Moreover, renewable resources like biomass can often be employed as a starting substrate for enzymatic polymerizations, producing useful polymeric materials. In this regard, the enzymatic polymerization is a valuable and promising method for conducting “green polymer chemistry”. The green nature of the reaction has been discussed from the viewpoints: clean reaction processes due to high selectivity without or minimal producing byproducts, starting raw materials, products 2385


Chemical Reviews

Review

mentioned in this article are expected as practically employed for biomedical, drug-delivery, pharmaceutical, and other application areas, which is realization of an important part of green polymer chemistry. Another possibility for the practical application is the development or finding of a new enzyme; for example, a highly active PHA synthase enzyme is probably derived by the modern biotechnology using the known DNA sequences. Then, the currently produced PHAs via fermentation in living cells will be produced instead via an in vitro enzymatic polymerization process. We believe that enzymatic processes shall become more important methods, especially for the production of valueadded functional macromolecules via an environmentally permissible method. In spite of frequent warning by IPCC of climate change like the global warming, the currently occurring processes of polymer production are not desirable in many cases, due to the carbon dioxide emission amount from the concept of the carbon neutral. In order to mitigate such environmental loading, replacement of the currently used petroleum-based processes with the enzymatic renewable resource-based processes and/or development of new enzymatic processes will definitely contribute to the sustainable society for the future.

Society of Japan Award (2002), and Synthetic Organic Chemistry

AUTHOR INFORMATION

Osaka University. He is the recipient of several awards and

Award, Japan (2013).

Hiroshi Uyama obtained his Master Degree of Engineering from Kyoto University in 1987 and subsequently Doctor Degree of Engineering from Tohoku University in 1991 under the direction of Professor Shiro Kobayashi. In 1987 he joined as an Assistant Professor in Tohoku University, thereafter moved to Kyoto University in 1996, and is presently Professor in the Department of Applied Chemistry, recognitions, including the Young Scientist Award (Chemical Society

Corresponding Author

of Japan) and the Highest Award of Japan Biotechnology Business

*E-mail: [email protected]. Fax/Tel: (+81)-75-724-7688.

Competition. His research interests include biomass plastics and

Notes

nanoprocessing of polymers.

The authors declare no competing financial interest. Biographies

Jun-ichi Kadokawa studied applied chemistry and materials chemistry at Tohoku University, where he received his M.S. degree in 1989 and Shin-ichiro Shoda received his Ph.D. degree from the University of Tokyo in 1981 under the supervision of Professor Teruaki Mukaiyama in the field of synthetic organic chemistry, where he developed the glycosyl fluoride method as a novel glycosylating technology. After spending three years working as an Assistant Professor at the University of Tokyo, he conducted his postdoctoral fellowship at ETH-Zurich (from 1984 until 1986) with Professor Dieter Seebach. In 1986, he moved to Tohoku University and joined the laboratories of Professor Shiro Kobayashi. In 1999, he was promoted to a Full Professor at Tohoku University (Functional Macromolecular Chemistry Laboratory). His research interests include synthesis of carbohydrates, the development of novel glycosylations, macromolecular architecture, and precision synthesis of well-designed functional oligo- and polysaccharides. He received the Award of the Chemical Society of Japan for Young Chemists (1986), the Cellulose

his Ph.D. in 1992 under the supervision of Prof. Shiro Kobayashi. He then joined Yamagata University as a Research Associate. From 1996 to 1997, he worked as a visiting scientist in the research group of Prof. Klaus Müllen at the Max-Planck-Institute for Polymer Research in Germany. In 1999, he became an Associate Professor at Yamagata University and moved to Tohoku University in 2002. He was appointed as a Professor of Kagoshima University in 2004. His research interests focus on enzymatic synthesis of non-natural polysaccharides, new polysaccharide-based materials, and control of the higher-ordered structures of polysaccharides. He received the Award for Encouragement of Research in Polymer Science (1998) and the Cellulose Society of Japan Award (2010). 2386


Chemical Reviews

Review

of Polymer Science, Japan (1986), Humboldt Research Award, Germany (1999), the Award of the Chemical Society of Japan (2001), the 30th John Stauffer Distinguished Lecture Award in the Sciences, University of Southern California (2002), Medal with Purple Ribbon (2007, Japanese Government), and The Order of the Sacred Treasure, Gold Rays with Neck Ribbon (2015, Japanese Government). He has been a Foreign Member of Northrhine Westfalian Academy of Sciences since 1999. He was a member of the Scientific Advisory Board for the Max Planck Institute for Polymer Research, Mainz (2001−2009). To date, he has served as an editor, regional editor, or honorary editor, and as a member of the (executive) advisory board or editorial (advisory) board for 24 international journals, and currently, he is serving for 8 journals. Shunsaku Kimura was born in Kyoto, Japan, in 1954. He received his B.S. (1976), M.S. (1978), and Ph.D. (1982, Professor Y. Imanishi) degrees from Kyoto University. He joined the Department of Polymer Chemistry, Kyoto University, as a Research Associate (1981), Lecturer (1992), and Associate Professor (1993). He moved to the Department of Material Chemistry, Kyoto University (1996), and in 1999 he was appointed Full Professor. He spent his postdoctoral career (1982− 1984, 1986, Professor R. Schwyzer) at ETH-Zurich, Switzerland. He received the Award of the Society of Polymer Science, Japan, in 1999. He currently serves as Associate Editor for Polymer Journal. His main interests are polymer supramolecular chemistry, peptide engineering, theranostic agents, immune-activating or suppressing materials, and optoelectronics devices.

ACKNOWLEDGMENTS S.S. acknowledges supports from the Ministry of Education, Sports, Science and Technology, and the International Center of Research & Education for Molecular Complex Chemistry from Tohoku University Global COE Program. J.K. acknowledges support from the Ministry of Education, Sports, Science and Culture, Japan, Asahi Glass Foundation, Sekisui Foundation, and Ezaki Glico Co. S.Ko. acknowledges various support particularly from the Ministry of Education, Science and Culture, Japan (especially, Granted for a Specially Promoted Research for “Enzymatic Polymerization”, 1996−1999), the Japan Society for the Promotion of Science, the 21st COE program of Kyoto University, NEDO for the project on the precision polymerization, the Mitsubishi Foundation, and several Japanese companies such as DIC Co., Toyo Ink Manufacturing Co., Sumitomo Chem. Co., Otsuka Chem. Co., and Toyota Motor Co. REFERENCES (1) Catalysis in Precision Polymerization; Kobayashi, S., Ed.; John Wiley & Sons: Chichester, England, 1997. (2) Advances in Polymer Science; Hierarchical Macromolecular Structures: 60 Years After the Staudinger Nobel Prize; Percec, V., Ed.; Springer: Cham; New York, 2013; Vol. 261-262. (3) Staudinger, H. Ü ber Polymerisation. Ber. Dtsch. Chem. Ges. B 1920, 53, 1073−1085. (4) Staudinger, H.; Fritschi, J. Ü ber Isopren und Kautschuk. 5. Mitteilung. Ü ber die Hydrierung des Kautschuks und Ü ber seine Konstitution. Helv. Chim. Acta 1922, 5, 785−806. (5) Staudinger, H. Ü ber die Konstitution des Kautschuks (6. Mitteilung). Ber. Dtsch. Chem. Ges. B 1924, 57, 1203−1208. (6) Staudinger, H. Hochpolymere Verbindungen. 5. Mitteilung. Ü ber die Konstitution der Poly-oxymethylene und Anderer Hochpolymerer Verbindungen. Helv. Chim. Acta 1925, 8, 67−80. (7) Staudinger, H.; Johner, H.; Singer, R.; Mie, G.; Hengstenberg, J. Polymerized Formaldehyde, A Model of Cellulose. Z. Phys. Chem. 1927, 126, 425−448. (8) Taylor, H. S.; Jones, W. H. The Thermal Decomposition of Metal Alkyls in Hydrogen-Ethylene Mixtures. J. Am. Chem. Soc. 1930, 52, 1111−1121. (9) Carothers, W. H. Polymerization. Chem. Rev. 1931, 8, 353−426. (10) National Historic Chemical Landmarks program of the American Chemical Society, Carothers, W. H.The Establishment of Modern Polymer Science, Commemorative booklet, 2000. (11) Meerwein, H. Ring-Opening Polymerization of Cyclic Ethers. Ger. Patent 741,478, 1937. (12) Meerwein, H.; Bodenbenner, K.; Borner, P.; Kunert, Fr.; Müller, K. W.; Sasse, H. J.; Schrodt, H.; Spille, J. Organic Ionic Reactions. Angew. Chem. 1955, 67, 374−380.

Shiro Kobayashi studied organic chemistry and polymer chemistry in Kyoto University, where he received his B.S. and M.S. from Prof. J. Furukawa and Ph.D. from Prof. T. Saegusa in 1969. Then, he worked as a postdoc with Prof. G. A. Olah at Case Western Reserve University for two years. In 1972, he joined Kyoto University as a research associate to start studying polymer synthesis. He stayed as a Humboldt fellow with Prof. H. Ringsdorf at University of Mainz in 1976. Following the lectureship in Kyoto University, he was appointed as a full professor at Tohoku University in 1986, starting research on enzymatic polymerization. In 1997, he moved to Kyoto University and officially retired in 2005, to become Emeritus Professor. Since then, he has been a distinguished professor at Kyoto Institute of Technology. His research interests include polymer synthesis and polymeric materials. In particular, his group developed the enzymatic polymerization, which enabled the first chemical synthesis of natural and unnatural polysaccharides like cellulose, xylan, chitin, hyaluronic acid, chondroitin, various functional polyesters, and new aromatics polymers. The method contributes to green polymer chemistry. He received verious awards among others: the Award of the Chemical Society of Japan for Young Chemists (1976), the Award of the Society 2387


Chemical Reviews

Review

(13) Williams, G. Kinetics of the Catalyzed Polymerization of Styrene. I. Experimental Methods and Some General Features of Stannic Chloride Catalysis. J. Chem. Soc. 1938, 246−253. (14) Williams, G. Kinetics of the Catalyzed Polymerization of Styrene. III. The Mechanism of the Metal Chloride Catalysis. J. Chem. Soc. 1940, 775−789. (15) Ziegler, K.; Holzkamp, E.; Breil, H.; Martin, H. The Mulheim Normal Pressure Polyethylene Process. Angew. Chem. 1955, 67, 541− 547. (16) Natta, G.; Pino, P.; Corradini, P.; Danusso, F.; Mantica, E.; Mazzanti, G.; Moraglio, G. Crystalline High Polymers of α-Olefins. J. Am. Chem. Soc. 1955, 77, 1708−1710. (17) Boor, J. Ziegler-Natta Catalysts and Polymerizations; Academic Press: New York, 1979. (18) Szwarc, M. Living Polymers. Nature 1956, 178, 1168−1169. (19) Shirakawa, H.; Louis, E. J.; Macdiarmid, A. G.; Chiang, C. K.; Heeger, A. J. Synthesis of Electrically Conducting Organic Polymers Halogen Derivatives of Polyacetylene, (CH)X. J. Chem. Soc., Chem. Commun. 1977, 578−580. (20) Trnka, T. M.; Grubbs, R. H. The Development of L2X2Ru = CHR Olefin Metathesis Catalysts: An Organometallic Success Story. Acc. Chem. Res. 2001, 34, 18−29. (21) Schrock, R. R. High Oxidation State Multiple Metal-Carbon Bonds. Chem. Rev. 2002, 102, 145−179. (22) Merrifield, R. B. Solid Phase Peptide Synthesis. I. The Synthesis of a Tetrapeptide. J. Am. Chem. Soc. 1963, 85, 2149−2154. (23) Lehn, J. M. Supramolecular Polymer Chemistry- Scope and Perspectives. Polym. Int. 2002, 51, 825−839. (24) Lehn, J. M. Dynamers: Dynamic Molecular and Supramolecular Polymers. Prog. Polym. Sci. 2005, 30, 814−831. (25) Anastas, P. T.; Warner, J. C. Green Chemistry: Theory and Practice; Oxford University Press, 1998. (26) Horvath, I. T.; Anastas, P. T. Innovations and Green Chemistry. Chem. Rev. 2007, 107, 2169−2173. (27) Kobayashi, S.; Shoda, S.; Kashiwa, K. The First in Vitro Synthesis of Cellulose via Enzymatic Polymerization and Characterization of the “Synthetic Cellulose”. In Cellulosics: Chemical, Biochemical and Material Aspects; Kennedy, J. F., Phillips, G. O., Williams, P. A., Eds.; Ellis Horwood Ltd.: London, 1993; pp 23−28. (28) Kobayashi, S.; Shoda, S.; Uyama, H. Enzymatic Polymerization and Oligomerization. Adv. Polym. Sci. 1995, 121, 1−30. (29) Kobayashi, S.; Shoda, S.; Uyama, H. Enzymatic Polymerization. In Polymeric Materials Encyclopedia; Salamone, J. C., Ed.; CRC Press Inc.: Boca Raton, FL, 1996; pp 2102−2107. (30) Uyama, H.; Kobayashi, S. Enzymatic Ring-Opening Polymerization of Macrolides to Polyesters. In Biomedical Functions and Biotechnology of Natural and Artificial Polymers; Yalpani, M., Ed.; ATL Press, Inc.: Mount Prospect, IL, 1996; pp 5−15. (31) Kobayashi, S.; Shoda, S.; Uyama, H. Enzymatic Catalysis. In Catalysis in Precision Polymerization; Kobayashi, S., Ed.; John Wiley & Sons: Chichester, England, 1997; pp 417−441. (32) Shoda, S.; Kobayashi, S. Recent Developments in the Use of Enzymes in Oligo- and Polysaccharide Synthesis. Trends Polym. Sci. 1997, 5, 109−115. (33) Shoda, S.; Fujita, M.; Kobayashi, S. Glycanase-Catalyzed Synthesis of Non-natural Oligosaccharides. Trends Glycosci. Glycotechnol. 1998, 10, 279−289. (34) Kobayashi, S. Enzymatic Polymerization: Synthesis of Artificial Macromolecules Catalyzed by Natural Macromolecules. Kobunshi 1999, 48, 124−127. (35) Kobayashi, S. Enzymatic Polymerization: A New Method of Polymer Synthesis. J. Polym. Sci., Part A: Polym. Chem. 1999, 37, 3041−3056. (36) Uyama, H.; Kobayash, S. Enzymatic Polymerization Yields Useful Polyphenols. CHEMTECH 1999, 29, 22−28. (37) Kobayashi, S.; Uyama, H. Biocatalytical Routes to Polymers. In Material Science and Technology-Synthesis of Polymers; Schlueter, A. D., Ed.; Wiley-VCH: Weinheim, 1999; Vol. 54, pp 549−569.

(38) Kobayashi, S.; Shoda, S.; Donnelly, M.; Church, S. P. Enzymatic Synthesis of Cellulose. In Methods in Biotechnology 10, Carbohydrate Biotechnology Protocols; Bucke, C., Ed.; Humana Press: Totowa, NJ, 1999; pp 57−69. (39) Kobayashi, S.; Kimura, S. In Vitro Biosynthesis of Natural and Unnatural Polysaccharides Catalyzed by Isolated Hydrolases. In Biochemical Principles and Mechanism of Biosynthesis and Biodegradation of Polymers; Steinbuechel, A., Ed.; Wiley-VCH: Weinheim, Germany, 1999; pp 161−167. (40) Kobayashi, S.; Uyama, H.; Ohmae, M. Enzymatic Polymerization for Precision Polymer Synthesis. Bull. Chem. Soc. Jpn. 2001, 74, 613−635. (41) Kobayashi, S.; Uyama, H.; Kimura, S. Enzymatic Polymerization. Chem. Rev. 2001, 101, 3793−3818. (42) Kobayashi, S.; Uyama, H. In Vitro Biosynthesis of Polyesters. In Advances in Biochemical Engineering/Biotechnology; Babel, W. S. A., Ed.; Springer: Heidelberg, 2001; pp 241−262. (43) Kobayashi, S.; Sakamoto, J.; Kimura, S. In vitro synthesis of cellulose and related polysaccharides. Prog. Polym. Sci. 2001, 26, 1525− 1560. (44) Kobayashi, S.; Uyama, H. In vitro Polyester Synthesis via Enzymatic Polymerization. Curr. Org. Chem. 2002, 6, 209−222. (45) Kobayashi, S.; Uyama, H. Enzymatic Polymerization to Polyesters. In Handbook of Biopolymers, Polyesters I; Doi, Y., Steinbüchel, A., Eds.; Wiley-VCH, 2002; Vol. 3a; pp 373−400. (46) Kobayashi, S.; Uyama, H. Enzymatic Polymerization. In Encyclopedia of Polymer Science and Technology, 3rd ed.; Kroschwitz, J. I., Ed.; Wiley: New York, 2003; pp 328−364. (47) Uyama, H.; Kobayashi, S. Enzymatic Synthesis of Polyphenols. Curr. Org. Chem. 2003, 7, 1387−1397. (48) Kobayashi, S.; Ritter, H.; Kaplan, D., Eds.; Enzyme-Catalyzed Synthesis of Polymers; Advances in Polymer Science; Springer: Berlin, 2006; Vol. 194. (49) Kobayashi, S. New Developments of Polysaccharide Synthesis via Enzymatic Polymerization. Proc. Jpn. Acad., Ser. B 2007, 83, 215− 247. (50) Kobayashi, S.; Ohmae, M. Polymer Synthesis and Modification by Enzymatic Catalysis. In Macromolecular Engineering: Precise Synthesis, Materials Properties, Applications; Matyjaszewski, K., Gnanou, Y., Leibler, L., Eds.; Wiley-VCH: Weinheim, 2007; pp 401−477. (51) Kobayashi, S. Recent Developments in Lipase-Catalyzed Synthesis of Polyesters. Macromol. Rapid Commun. 2009, 30, 237− 266. (52) Kobayashi, S.; Makino, A. Enzymatic Polymer Synthesis: An Opportunity for Green Polymer Chemistry. Chem. Rev. 2009, 109, 5288−5353. (53) Kobayashi, S. Lipase-Catalyzed Polyester Synthesis - A Green Polymer Chemistry. Proc. Jpn. Acad., Ser. B 2010, 86, 338−365. (54) Kadokawa, J.; Kobayashi, S. Polymer Synthesis by Enzymatic Catalysis. Curr. Opin. Chem. Biol. 2010, 14, 145−153. (55) Makino, A.; Kobayashi, S. Chemistry of 2-Oxazolines: A Crossing of Cationic Ring-Opening Polymerization and Enzymatic Ring-Opening Polyaddition. J. Polym. Sci., Part A: Polym. Chem. 2010, 48, 1251−1270. (56) Kadokawa, J. Precision Polysaccharide Synthesis Catalyzed by Enzymes. Chem. Rev. 2011, 111, 4308−4345. (57) Kobayashi, S. Enzymatic Polymerization. In Polymer Science: A Comprehensive Reference; Matyjaszewski, K., Moeller, M., Eds.; Elsevier: Amsterdam, 2012; Vol. 5, pp 217−237. (58) Kobayashi, S. Enzymatic Polymerization. In Encyclopedia of Polymer Science and Technology, 4th ed.; Seidel, A., Ed.; Wiley: Hoboken, 2014; pp 221−292. (59) Enzymes in Polymer Synthesis; Gross, R. A., Kaplan, D., Swift, G., Eds.; ACS Symposium Series 684; American Chemical Society: Washington, DC, 1998. (60) Gross, R. A.; Kumar, A.; Kalra, B. Polymer Synthesis by In Vitro Enzyme Catalysis. Chem. Rev. 2001, 101, 2097−2124. 2388


Chemical Reviews

Review

(61) Biocatalysis in Polymer Science; Cheng, H. N., Gross, R. A., Eds.; ACS Symposium Series 840; Americal Chemical Society: Washington, DC, 2003. (62) Polymer Biocatalysis and Biomaterials; Cheng, H. N., Gross, R. A., Eds.; ACS Symposium Series 900; American Chemical Society: Washington, DC, 2005. (63) Polymer Biocatalysis and Biomaterials II; Cheng, H. N., Gross, R. A., Eds.; ACS Symposium Series 999; Americal Chemical Society: Washington, DC, 2008. (64) Enzymatic Polymerisation; Palmans, A. R. A., Heise, A., Guebitz, G. M., Eds.; Advances in Polymer Science; Springer: Berlin, 2010; Vol. 237. (65) Biocatalysis in Polymer Chemistry; Loos, K., Ed.; Wiley-VCH: Weinheim, Germany, 2011. (66) Biocatalysis in Polymer Science; Gross, R. A., Cheng, H. N., Smith, P. B., Eds.; ACS Symposium Series 1144; American Chemical Society: Washington, D.C., 2013. (67) Puskas, J. E.; Sen, M. Y.; Seo, K. S. Green Polymer Chemistry Using Nature’s Catalysts, Enzymes. J. Polym. Sci., Part A: Polym. Chem. 2009, 47, 2959−2976. (68) Green Polymer Chemistry: Biocatalysis and Biomaterials; Cheng, H. N., Gross, R. A., Eds.; ACS Symposium Series 1043; American Chemical Society: Washington, DC, 2010. (69) Kobayashi, S. Green Polymer Chemistry: Recent Developments. Adv. Polym. Sci. 2013, 262, 141−166. (70) Green Polymer Chemistry: Biobased Materials and Biocatalysis; Cheng, H. N., Gross, R. A., Smith, P. B., Eds.; ACS Symposium Series 1192; American Chemical Society: Washington, DC, 2015. (71) Shoda, S.; Kobayashi, A.; Kobayashi, S. Production of Polymers by White Biotechnology. In White Biotechnology for Sustainable Chemistry; Coelho, M. A. Z., Ribeiro, B. D., Eds.; Royal Society of Chemistry: U.K., 2015; pp 274−309. (72) Kobayashi, S. Enzymatic Ring-Opening Polymerization and Polycondensation for the Green Synthesis of Polyesters. Polym. Adv. Technol. 2015, 26, 677−686. (73) Kobayashi, S.; Shoda, S.; Wen, X.; Okamoto, E.; Kiyosada, T. Choroselective Enzymatic Polymerization for Synthesis of Natural Polysaccharides. J. Macromol. Sci., Part A: Pure Appl. Chem. 1997, 34, 2135−2142. (74) Kobayashi, S.; Uyama, H. Biomacromolecules and Bio-related Macromolecules. Macromol. Chem. Phys. 2003, 204, 235−256. (75) Payen, A.; Persoz, J.-F. Memoir on Diastase, the Principal Products of Its Reaction, and Their Application to the Industrial Arts. Ann. Chim. Phys. 2nd Series 1833, 53, 73−92. (76) International Union of Biochemistry and Molecular Biology. Webb, E. C. Enzyme Nomenclature 1992: Recommendations of the Nomenclature Committee of the International Union of Biochemistry and Molecular Biology on the Nomenclature and Classification of Enzymes; International Union of Biochemistry and Molecular Biology by Academic Press: San Diego, 1992. (77) Fischer, E. Einfluss der Configuration auf die Wirkung der Enzyme. Ber. Dtsch. Chem. Ges. 1894, 27, 2985−2993. (78) Pauling, L. Molecular Architecture and Biological Reactions. Chem. Eng. News 1946, 24, 1375−1377. (79) Kollman, P. A.; Kuhn, B.; Donini, O.; Perakyla, M.; Stanton, R.; Bakowies, D. Elucidating the Nature of Enzyme Catalysis Utilizing a New Twist on an Old Methodology: Quantum Mechanical - Free Energy Calculations on Chemical Reactions in Enzymes and in Aqueous Solution. Acc. Chem. Res. 2001, 34, 72−79. (80) Borman, S. Much Ado about Enzyme Mechanisms. Chem. Eng. News 2004, 82, 35−39. (81) Alberts, B.; Bray, D.; Lewis, J.; Raff, M.; Roberts, K.; Watson, J. D. Mol. Biol. Cell; 3rd ed.; Newton Press: New York, 1994; Chapter 3. (82) Lerner, R. A.; Benkovic, S. J.; Schultz, P. G. At the Crossroads of Chemistry and Immunology - Catalytic Antibodies. Science 1991, 252, 659−667. (83) Sym, E. A. Ü ber die Esterasewirkung. III. Biochem. Z. 1933, 258, 304−324.

(84) Sym, E. A. A Method for Enzymic Ester Synthesis. Enzymologia 1936, 1, 156−160. (85) Jones, J. B. Enzymes in Organic-Synthesis. Tetrahedron 1986, 42, 3351−3403. (86) Klibanov, A. M. Asymmetric Transformations Catalyzed by Enzymes in Organic-Solvents. Acc. Chem. Res. 1990, 23, 114−120. (87) Wong, C. H.; Whitesides, G. M. P. Enzymes in Synthetic Organic Chemistry; Elsevier, Oxford, 1994. (88) Kobayashi, S.; Okamoto, E.; Wen, X.; Shoda, S. Chemical Synthesis of Native-type Cellulose and Its Analogues via Enzymatic Polymerization. J. Macromol. Sci., Part A: Pure Appl.Chem. 1996, A33, 1375−1384. (89) Kobayashi, S.; Uyama, H.; Takamoto, T. Lipase-Catalyzed Degradation of Polyesters in Organic Solvents, A New Methodology of Polymer Recycling Using Enzyme as Catalyst. Biomacromolecules 2000, 1, 3−5. (90) Ebata, H.; Toshima, K.; Matsumura, S. Lipase-Catalyzed Transformation of Poly(ε-caprolactone) into Cyclic Dicaprolactone. Biomacromolecules 2000, 1, 511−514. (91) Takahashi, Y.; Okajima, S.; Toshima, K.; Matsumura, S. LipaseCatalyzed Transformation of Poly(lactic acid) into Cyclic Oligomers. Macromol. Biosci. 2004, 4, 346−353. (92) Osanai, Y.; Toshima, K.; Matsumura, S. Enzymatic Degradation of Poly(R,S-3-hydroxybutanoate) to Cyclic Oligomers Under Continuous Flow. Green Chem. 2003, 5, 567−570. (93) Numata, K.; Srivastava, R. K.; Finne-Wistrand, A.; Albertsson, A. C.; Doi, Y.; Abe, H. Branched Poly(lactide) Synthesized by Enzymatic Polymerization: Effects of Molecular Branches and Stereochernistry on Enzymatic Degradation and Alkaline Hydrolysis. Biomacromolecules 2007, 8, 3115−3125. (94) Igarashi, K.; Uchihashi, T.; Koivula, A.; Wada, M.; Kimura, S.; Okamoto, T.; Penttila, M.; Ando, T.; Samejima, M. Traffic Jams Reduce Hydrolytic Efficiency of Cellulase on Cellulose Surface. Science 2011, 333, 1279−1282. (95) Igarashi, K.; Uchihashi, T.; Uchiyama, T.; Sugimoto, H.; Wada, M.; Suzuki, K.; Sakuda, S.; Ando, T.; Watanabe, T.; Samejima, M. Two-Way Traffic of Glycoside Hydrolase Family 18 Processive Chitinases on Crystalline Chitin. Nat. Commun. 2014, 5, 3975. (96) Negoro, S.; Shibata, N.; Tanaka, Y.; Yasuhira, K.; Shibata, H.; Hashimoto, H.; Lee, Y. H.; Oshima, S.; Santa, R.; Oshima, S.; Mochiji, K.; Goto, Y.; Ikegami, T.; Nagai, K.; Kato, D.; Takeo, M.; Higuchi, Y. Three-Dimensional Structure of Nylon Hydrolase and Mechanism of Nylon-6 Hydrolysis. J. Biol. Chem. 2012, 287, 5079−5090. (97) Pennerhahn, J. E.; Smith Eble, K.; McMurry, T. J.; Renner, M.; Balch, A. L.; Groves, J. T.; Dawson, J. H.; Hodgson, K. O. Structural Characterization of Horseradish Peroxidase Using EXAFS Spectroscopy. Evidence for FeO Ligation in Compounds I and II. J. Am. Chem. Soc. 1986, 108, 7819−7825. (98) Kobayashi, S.; Higashimura, H. Oxidative Polymerization of Phenols Revisited. Prog. Polym. Sci. 2003, 28, 1015−1048. (99) Higashimura, H.; Kobayashi, S. Oxidative Polymerization. In Encyclopedia of Polymer Science and Technology, 3rd ed.; Kroschiwitz, J. I., Ed.; John Wiley & Sons: New York, 2004; Vol. 10, pp 740−764. (100) Reihmann, M.; Ritter, H. Synthesis of Phenol Polymers Using Peroxidases. Adv. Polym. Sci. 2006, 194, 1−49. (101) Uyama, H.; Kobayashi, S. Enzymatic Synthesis and Properties of Polymers from Polyphenols. Adv. Polym. Sci. 2006, 194, 51−67. (102) Hollmann, F.; Arends, I. W. C. E. Enzyme Initiated Radical Polymerizations. Polymers 2012, 4, 759−793. (103) Hay, A. S. Polymerization by Oxidative Coupling: Discovery and Commercialization of PPO® and Noryl® Resins. J. Polym. Sci., Part A: Polym. Chem. 1998, 36, 505−517. (104) Dordick, J. S.; Marletta, M. A.; Klibanov, A. M. Polymerization of Phenols Catalyzed by Peroxidase in Nonaqueous Media. Biotechnol. Bioeng. 1987, 30, 31−36. (105) Uyama, H.; Kurioka, H.; Kaneko, I.; Kobayashi, S. Synthesis of a New Family of Phenol Resin by Enzymatic Oxidative Polymerization. Chem. Lett. 1994, 423−426. 2389


Chemical Reviews

Review

(106) Uyama, H.; Kurioka, H.; Sugihara, J.; Kobayashi, S. Enzymatic Synthesis and Thermal Properties of a New Class of Polyphenol. Bull. Chem. Soc. Jpn. 1996, 69, 189−193. (107) Oguchi, T.; Tawaki, S.; Uyama, H.; Kobayashi, S. Soluble Polyphenol. Macromol. Rapid Commun. 1999, 20, 401−403. (108) Oguchi, T.; Tawaki, S.; Uyama, H.; Kobayashi, S. Enzymatic Synthesis of Soluble Polyphenol. Bull. Chem. Soc. Jpn. 2000, 73, 1389− 1396. (109) Mita, N.; Tawaki, S.; Uyama, H.; Kobayashi, S. Molecular Weight Control of Polyphenols by Enzymatic Copolymerization of Phenols. Polym. J. 2001, 33, 374−376. (110) Mita, N.; Tawaki, S.; Uyama, H.; Kobayashi, S. Enzymatic Oxidative Polymerization of Phenol in an Aqueous Solution in the Presence of a Catalytic Amount of Cyclodextrin. Macromol. Biosci. 2002, 2, 127−130. (111) Kim, Y. J.; Uyama, H.; Kobayashi, S. Regioselective Synthesis of Poly(phenylene) as a Complex with Poly(ethylene glycol) by Template Polymerization of Phenol in Water. Macromolecules 2003, 36, 5058−5060. (112) Kim, Y. J.; Uyama, H.; Kobayashi, S. Peroxidase-Catalyzed Oxidative Polymerization of Phenol with a Nonionic Polymer Surfactant Template in Water. Macromol. Biosci. 2004, 4, 497−502. (113) Kim, Y. J.; Shibata, K.; Uyama, H.; Kobayashi, S. Synthesis of Ultrahigh Molecular Weight Phenolic Polymers by Enzymatic Polymerization in the Presence of Amphiphilic Triblock Copolymer in Water. Polymer 2008, 49, 4791−4795. (114) Uyama, H.; Kurioka, H.; Kobayashi, S. Preparation of Polyphenol Particles by Dispersion Polymerization Using Enzyme as Catalyst. Chem. Lett. 1995, 795−796. (115) Kurioka, H.; Uyama, H.; Kobayashi, S. Peroxidase-Catalyzed Dispersion Polymerization of Phenol Derivatives. Polym. J. 1998, 30, 526−529. (116) Nie, G.; Zhang, L.; Cui, Y. Synthesis of Polyphenol Microsphere-Supported Palladium Complex and Evaluation of its Catalytic Performance for Heck Reaction. Appl. Organomet. Chem. 2012, 26, 635−640. (117) Zhang, L.; Zhao, W.; Chen, H.; Cui, Y. Enzymatic Synthesis of Phenol Polymer and Its Functionalization. J. Mol. Catal. B: Enzym. 2013, 87, 30−36. (118) Uyama, H.; Kurioka, H.; Kobayashi, S. Novel Bienzymatic Catalysis System for Oxidative Polymerization of Phenols. Polym. J. 1997, 29, 190−192. (119) Zhang, B.; Wang, X.; Zhu, A.; Ma, K.; Lv, Y.; Wang, X.; An, Z. Enzyme-Initiated Reversible Addition−Fragmentation Chain Transfer Polymerization. Macromolecules 2015, 48, 7792−7802. (120) Li, Z.; Renneckar, S.; Barone, J. R. Nanocomposites Prepared by In Situ Enzymatic Polymerization of Phenol with TEMPOOxidized Nanocellulose. Cellulose 2010, 17, 57−68. (121) Peng, Y.; Liu, H. W.; Zhang, X. Y.; Li, Y. S.; Liu, S. Y. CNT Templated Regioselective Enzymatic Polymerization of Phenol in Water and Modification of Surface of MWNT Thereby. J. Polym. Sci., Part A: Polym. Chem. 2009, 47, 1627−1635. (122) Cui, Y.; Hu, A.; Lu, Z.; Zhang, L.; Luo, X.; Du, K. Synthesis of Water-Soluble Polyphenol-graft-poly(ethylene oxide) Copolymers via Enzymatic Polymerization and Anionic Polymerization. Polym. Int. 2009, 59, 676−679. (123) Kurioka, H.; Komatsu, I.; Uyama, H.; Kobayashi, S. Enzymatic Oxidative Polymerization of Alkylphenols. Macromol. Rapid Commun. 1994, 15, 507−510. (124) Uyama, H.; Kurioka, H.; Sugihara, J.; Komatsu, I.; Kobayashi, S. Oxidative Polymerization of p-Alkylphenols Catalyzed by Horseradish Peroxidase. J. Polym. Sci., Part A: Polym. Chem. 1997, 35, 1453− 1459. (125) Uyama, H.; Kurioka, H.; Sugihara, J.; Komatsu, I.; Kobayashi, S. Peroxidase-Catalyzed Oxidative Polymerization of Cresols to a New Family of Polyphenols. Bull. Chem. Soc. Jpn. 1995, 68, 3209−3214. (126) Ayyagari, M. S.; Marx, K. A.; Tripathy, S. K.; Akkara, J. A.; Kaplan, D. L. Controlled Free-Radical Polymerization of Phenol

Derivatives by Enzyme-Catalyzed Reactions in Organic-Solvents. Macromolecules 1995, 28, 5192−5197. (127) Sahoo, S. K.; Liu, W.; Samuelson, L. A.; Kumar, J.; Cholli, A. L. Biocatalytic Polymerization of p-Cresol: An In-Situ NMR Approach to Understand the Coupling Mechanism. Macromolecules 2002, 35, 9990−9998. (128) Wu, X. D.; Liu, W.; Nagarajan, R.; Kumar, J.; Samuelson, L. A.; Cholli, A. L. Role of Temperature in Suppression of the Formation of Pummerer’s Type Ketone in Enzymatic Polymerization of 4Propylphenol: An In-Situ Variable Temperature H-1 NMR Study. Macromolecules 2004, 37, 2322−2324. (129) Mita, N.; Tawaki, S.; Uyama, H.; Kobayashi, S. Structural Control in Enzymatic Oxidative Polymerization of Phenols with Varying the Solvent and Substituent Nature. Chem. Lett. 2002, 402− 403. (130) Mita, N.; Maruichi, N.; Tonami, H.; Nagahata, R.; Tawaki, S.; Uyama, H.; Kobayashi, S. Enzymatic Oxidative Polymerization of p-tButylphenol and Characterization of the Product Polymer. Bull. Chem. Soc. Jpn. 2003, 76, 375−379. (131) Mita, N.; Tawaki, S.; Uyama, H.; Kobayashi, S. Precise Structure Control of Enzymatically Synthesized Polyphenols. Bull. Chem. Soc. Jpn. 2004, 77, 1523−1527. (132) Wang, P.; Martin, B. D.; Parida, S.; Rethwisch, D. G.; Dordick, J. S. Multienzymic Synthesis of Poly(hydroquinone) for Use as a Redox Polymer. J. Am. Chem. Soc. 1995, 117, 12885−12886. (133) Tonami, H.; Uyama, H.; Kobayashi, S.; Rettig, K.; Ritter, H. Chemoenzymatic Synthesis of a Poly(hydroquinone). Macromol. Chem. Phys. 1999, 200, 1998−2002. (134) Fukuoka, T.; Tachibana, Y.; Tonami, H.; Uyama, H.; Kobayashi, S. Enzymatic Polymerization of Tyrosine Derivatives. Peroxidase- and Protease-Catalyzed Synthesis of Poly(tyrosine)s with Different Structures. Biomacromolecules 2002, 3, 768−774. (135) Uyama, H.; Lohavisavapanich, C.; Ikeda, R.; Kobayashi, S. Chemoselective Polymerization of a Phenol Derivative Having a Methacryl Group by Peroxidase Catalyst. Macromolecules 1998, 31, 554−556. (136) Reihmann, M. H.; Ritter, H. Oxidative Oligomerization of Cyclodextrin-Complexed Bifunctional Phenols Catalyzed by Horseradish Peroxidase in Water. Macromol. Chem. Phys. 2000, 201, 798− 804. (137) Reihmann, M. H.; Ritter, H. Enzymatically Catalyzed Synthesis of Photocrosslinkable Oligophenols. Macromol. Chem. Phys. 2000, 201, 1593−1597. (138) Pang, Y. J.; Ritter, H.; Tabatabai, M. Cyclodextrins in Polymer Chemistry: Enzymatically Catalyzed Oxidative Polymerization of paraFunctionalized Phenol Derivatives in Aqueous Medium by Use of Horseradish Peroxidase. Macromolecules 2003, 36, 7090−7093. (139) Kobayashi, S.; Uyama, H.; Ushiwata, T.; Uchiyama, T.; Sugihara, J.; Kurioka, H. Enzymatic Oxidative Polymerization of Bisphenol-A to a New Class of Soluble Polyphenol. Macromol. Chem. Phys. 1998, 199, 777−782. (140) Kim, Y. H.; An, E. S.; Park, S. Y.; Lee, J. O.; Kim, J. H.; Song, B. K. Polymerization of Bisphenol a Using Coprinus Cinereus Peroxidase (CiP) and its Application as a Photoresist Resin. J. Mol. Catal. B: Enzym. 2007, 44, 149−154. (141) Fukuoka, T.; Tonami, H.; Maruichi, N.; Uyama, H.; Kobayashi, S.; Higashimura, H. Peroxidase-Catalyzed Oxidative Polymerization of 4,4 ′-Dihydroxydiphenyl Ether. Formation of α,ω-Hydroxyoligo(1,4phenylene oxide) Through an Unusual Reaction Pathway. Macromolecules 2000, 33, 9152−9155. (142) Kommareddi, N. S.; Tata, M.; Karayigitoglu, C.; John, V. T.; McPherson, G. L.; Herman, M. F.; O'Connor, C. J.; Lee, Y. S.; Akkara, J. A.; Kaplan, D. L. Enzymatic Polymerizations Using Surfactant Microstructures and the Preparation of Polymer-Ferrite Composites. Appl. Biochem. Biotechnol. 1995, 51−52, 241−252. (143) Xu, P.; Kumar, J.; Samuelson, L.; Cholli, A. L. Monitoring the Enzymatic Polymerization of 4-Phenylphenol by Matrix-Assisted Laser Desorption Ionization Time-of-Flight Mass Spectrometry: A Novel Approach. Biomacromolecules 2002, 3, 889−893. 2390


Chemical Reviews

Review

(144) Zhang, L.; Zhao, W.; Ma, Z.; Nie, G.; Cui, Y. Enzymatic Polymerization of Phenol Catalyzed by Horseradish Peroxidase in Aqueous Micelle System. Eur. Polym. J. 2012, 48, 580−585. (145) Zhang, L.; Zhang, Y.; Xue, Y.; Duan, H.; Cui, Y. Enzymatic Synthesis of Soluble Phenol Polymer in Water Using Anionic Surfactant as Additive. Polym. Int. 2013, 62, 1277−1282. (146) Sgalla, S.; Fabrizi, G.; Cacchi, S.; Macone, A.; Bonamore, A.; Boffi, A. Horseradish Peroxidase in Ionic Liquids - Reactions with Water Insoluble Phenolic Substrates. J. Mol. Catal. B: Enzym. 2007, 44, 144−148. (147) Zaragoza-Gasca, P.; Villamizar-Galvez, O. J.; Garcia-Arrazola, R.; Gimeno, M.; Barzana, E. Use of Ionic Liquid for the EnzymeCatalyzed Polymerization of Phenols. Polym. Adv. Technol. 2009, 21, 454−456. (148) Angerer, P. S.; Studer, A.; Witholt, B.; Li, Z. Oxidative Polymerization of a Substituted Phenol with Ion-paired Horseradish Peroxidase in an Organic Solvent. Macromolecules 2005, 38, 6248− 6250. (149) Zheng, K.; Duan, H.; Zhang, L.; Cui, Y. Synthesis of Poly(4methoxyphenol) by Enzyme-Catalyzed Polymerization and Evaluation of Its Antioxidant Activity. New J. Chem. 2013, 37, 4185−4191. (150) Robert, J. P.; Uyama, H.; Kobayashi, S.; Jordan, R.; Nuyken, O. First Diazosulfonate Homopolymer by Enzymatic Polymerization. Macromol. Rapid Commun. 2003, 24, 185−189. (151) Duan, H.; Zheng, K.; Zhang, L.; Cui, Y. Synthesis of Poly(4aminophenol) by Horseradish Peroxidase and the Evaluation of Its Adsorptivity for Silver Ions. J. Appl. Polym. Sci. 2014, 131, 40361− 40366. (152) Tsujimoto, T.; Ikeda, R.; Uyama, H.; Kobayashi, S. Crosslinkable Polyphenols from Urushiol Analogues. Macromol. Chem. Phys. 2001, 202, 3420−3425. (153) Mondrzyk, A.; Mondrzik, B.; Gingter, S.; Ritter, H. New Enzymatically Polymerized Copolymers from 4-tert-Butylphenol and 4-Ferrocenylphenol and their Modification and Inclusion Complexes with β-Cyclodextrin. Beilstein J. Org. Chem. 2012, 8, 2118−2123, No. 2238.10.3762/bjoc.8.238 (154) Cui, Y.; Han, X.; Ding, Y.; Lu, Z. Enzymatic Synthesis of Polyphenols with Longer Conjugation Lengths. Polym. Bull. 2010, 64, 647−656. (155) Turac, E.; Sahmetlioglu, E. Oxidative Polymerization of 4-[(4Phenylazo-phenyimino)-methyl]-phenol Catalyzed by Horseradish Peroxidase. Synth. Met. 2010, 160, 169−172. (156) Ghan, R.; Shutava, T.; Patel, A.; John, V. T.; Lvov, Y. EnzymeCatalyzed Polymerization of Phenols within Polyelectrolyte Microcapsules. Macromolecules 2004, 37, 4519−4524. (157) Yoshimura, Y.; Khunathai, K.; Nozoe, A.; Ohto, K.; Kawakita, H. Precious Metal Recovery Using Poly(tyramine) Prepared by Radical Polymerization with Horseradish Peroxidase. J. Chem. Eng. Jpn. 2012, 45, 178−181. (158) Shutava, T.; Zheng, Z. G.; John, V.; Lvov, Y. Microcapsule Modification with Peroxidase-Catalyzed Phenol Polymerization. Biomacromolecules 2004, 5, 914−921. (159) Tonami, H.; Uyama, H.; Kobayashi, S.; Kubota, M. PeroxidaseCatalyzed Oxidative Polymerization of m-Substituted Phenol Derivatives. Macromol. Chem. Phys. 1999, 200, 2365−2371. (160) Kadota, J.; Fukuoka, T.; Uyama, H.; Hasegawa, K.; Kobayashi, S. New Positive-Type Photoresists Based on Enzymatically Synthesized Polyphenols. Macromol. Rapid Commun. 2004, 25, 441−444. (161) Tonami, H.; Uyama, H.; Kobayashi, S.; Reihmann, M.; Ritter, H. Enzymatic Polymerization of m-Substituted Phenols in the Presence of 2,6-Di-O-methyl-β-cyclodextrin in Water. e-Polym. 2002, 2, 1−7. (162) Kim, Y. H.; An, E. S.; Song, B. K. Co-polymerization of MTPC (Methylene tri p-cresol) and m-cresol Using CiP (Coprinus Cinereus Peroxidase) to Improve the Dissolution Characteristics of the EnzymeCatalyzed Polymer. J. Mol. Catal. B: Enzym. 2009, 56, 227−230. (163) Tonami, H.; Uyama, H.; Kobayashi, S.; Fujita, T.; Taguchi, Y.; Osada, K. Chemoselective Oxidative Polymerization of m-Ethynyl-

phenol by Peroxidase Catalyst to a Aew Reactive Polyphenol. Biomacromolecules 2000, 1, 149−151. (164) Ikeda, R.; Tanaka, H.; Uyama, H.; Kobayashi, S. A New Crosslinkable Polyphenol from a Renewable Resource. Macromol. Rapid Commun. 2000, 21, 496−499. (165) Ikeda, R.; Tanaka, H.; Uyama, H.; Kobayashi, S. Enzymatic Synthesis and Curing of Poly(cardanol). Polym. J. 2000, 32, 589−593. (166) Ikeda, R.; Tsujimoto, T.; Tanaka, H.; Oyabu, H.; Uyama, H.; Kobayashi, S. Man-Made Urushi - Preparation of Crosslinked Polymeric Films from Renewable Resources via Air-Oxidation Processes. Proc. Jpn. Acad., Ser. B 2000, 76, 155−160. (167) Ikeda, R.; Tanaka, H.; Uyama, H.; Kobayashi, S. Synthesis and Curing Behaviors of a Crosslinkable Polymer from Cashew Nut Shell Liquid. Polymer 2002, 43, 3475−3481. (168) Kim, Y. H.; An, E. S.; Song, B. K.; Kim, D. S.; Chelikani, R. Polymerization of Cardanol Using Soybean Peroxidase and Its Potential Application as Anti-Biofilm Coating Material. Biotechnol. Lett. 2003, 25, 1521−1524. (169) Kobayashi, S.; Uyama, H.; Ikeda, R. Artificial Urushi. Chem. Eur. J. 2001, 7, 4754−4760. (170) Won, K.; Kim, Y. H.; An, E. S.; Lee, Y. S.; Song, B. K. Horseradish Peroxidase-Catalyzed Polymerization of Cardanol in the Presence of Redox Mediators. Biomacromolecules 2004, 5, 1−4. (171) Kim, Y. H.; Won, K. H.; Kwon, J. M.; Jeong, H. S.; Park, S. Y.; An, E. S.; Song, B. K. Synthesis of Polycardanol from a Renewable Resource Using a Fungal Peroxidase from Coprinus cinereus. J. Mol. Catal. B: Enzym. 2005, 34, 33−38. (172) Park, S. Y.; Kim, Y. H.; Won, K.; Song, B. K. Enzymatic Synthesis and Curing of Polycardol from Renewable Resources. J. Mol. Catal. B: Enzym. 2009, 57, 312−316. (173) Zhou, Q.; Cho, D.; Song, B. K.; Kim, H.-J. Curing Behavior of Polycardanol by MEKP and Cobalt Naphthenate Using Differential Scanning Calorimetry. J. Therm. Anal. Calorim. 2010, 99, 277−284. (174) Chelikani, R.; Kim, Y. H.; Yoon, D.-Y.; Kim, D.-S. Enzymatic Polymerization of Natural Anacardic Acid and Antibiofouling Effects of Polyanacardic Acid Coatings. Appl. Biochem. Biotechnol. 2009, 157, 263−277. (175) Tsujimoto, T.; Uyama, H.; Kobayashi, S. Synthesis and Curing Behaviors of Cross-linkable Polynaphthols from Renewable Resources: Preparation of Artificial Urushi. Macromolecules 2004, 37, 1777−1782. (176) Bilici, A.; Kaya, I.; Yildirim, M.; Dogan, F. Enzymatic Polymerization of Hydroxy-Functionalized Carbazole Monomer. J. Mol. Catal. B: Enzym. 2010, 64, 89−95. (177) Antoniotti, S.; Santhanam, L.; Ahuja, D.; Hogg, M. G.; Dordick, J. S. Structural Diversity of Peroxidase-Catalyzed Oxidation Products of o-Methoxyphenois. Org. Lett. 2004, 6, 1975−1978. (178) Tonami, H.; Uyama, H.; Nagahata, R.; Kobayashi, S. Guaiacol Oxidation Products in the Enzyme-activity Assay Reaction by Horseradish Peroxidase Catalysis. Chem. Lett. 2004, 33, 796−797. (179) Nazari, K.; Adhami, F.; Najjar-Safari, A.; Salmani, S.; Mahmoudi, A. Biocatalytic Synthesis of Polymeric Nanowires by Micellar Templates of Ionic Surfactants. Biochem. Biophys. Res. Commun. 2011, 410, 901−903. (180) Cohen-Yaniv, V.; Dosoretz, C. G. Fate of Horseradish Peroxidase During Oxidation of Monobrominated Phenols. J. Chem. Technol. Biotechnol. 2009, 84, 1559−1566. (181) Feng, W.; Taylor, K. E.; Biswas, N.; Bewtra, J. K. Soybean Peroxidase Trapped in Product Precipitate During Phenol Polymerization Retains Activity and May Be Recycled. J. Chem. Technol. Biotechnol. 2013, 88, 1429−1435. (182) Ikeda, R.; Sugihara, J.; Uyama, H.; Kobayashi, S. Enzymatic Oxidative Polymerization of 2,6-Dimethylphenol. Macromolecules 1996, 29, 8702−8705. (183) Nanayakkara, S.; Zhao, Z.; Patti, A. F.; He, L.; Saito, K. Immobilized Horseradish Peroxidase (I-HRP) as Biocatalyst for Oxidative Polymerization of 2,6-Dimethylphenol. ACS Sustainable Chem. Eng. 2014, 2, 1947−1950. 2391


Chemical Reviews

Review

(184) Ikeda, R.; Sugihara, J.; Uyama, H.; Kobayashi, S. Enzymatic Oxidative Polymerization of 4-Hydroxybenzoic Acid Derivatives to Poly(phenylene oxide)s. Polym. Int. 1998, 47, 295−301. (185) Ikeda, R.; Uyama, H.; Kobayashi, S. Poly(2,6-dihydroxy-1,4oxyphenylene) - Synthesis of a New Poly(phenylene oxide) Derivative. Polym. Bull. 1997, 38, 273−277. (186) Lairez, D.; Cathala, B.; Monties, B.; Bedos-Belval, F.; Duran, H.; Gorrichon, L. Aggregation During Coniferyl Alcohol Polymerization in Pectin Solution: A Biomimetic Approach of the First Steps of Lignification. Biomacromolecules 2005, 6, 763−774. (187) Nakamura, R.; Matsushita, Y.; Umemoto, K.; Usuki, A.; Fukushima, K. Enzymatic Polymerization of Coniferyl Alcohol in the Presence of Cyclodextrins. Biomacromolecules 2006, 7, 1929−1934. (188) Moon, S.-J.; Kwon, M.; Choi, D.; Won, K.; Kim, Y. H.; Choi, I.-G.; Choi, J. W. In vitro Analysis of the Monolignol Coupling Mechanism Using Dehydrogenative Polymerization in the Presence of Peroxidases and Controlled Feeding Ratios of Coniferyl and Sinapyl Alcohol. Phytochemistry 2012, 82, 15−21. (189) Yang, D.; Chang, Y.; Wu, X.; Qiu, X.; Lou, H. Modification of Sulfomethylated Alkali Lignin Catalyzed by Horseradish Peroxidase. RSC Adv. 2014, 4, 53855−53863. (190) Premachandran, R. S.; Banerjee, S.; Wu, X. K.; John, V. T.; McPherson, G. L.; Akkara, J.; Ayyagari, M.; Kaplan, D. Enzymatic Synthesis of Fluorescent Naphthol-based Polymers. Macromolecules 1996, 29, 6452−6460. (191) Alva, K. S.; Samuelson, L.; Kumar, J.; Tripathy, S.; Cholli, A. L. Enzyme-Catalyzed Polymerization of 8-Hydroxyquinoline-5-sulfonate by in situ Nuclear Magnetic Resonance Spectroscopy. J. Appl. Polym. Sci. 1998, 70, 1257−1264. (192) Yamaguchi, I.; Yamamoto, T. Enzymatic Polymerization to Give Poly(dihydroxynaphthalene)s and their Photoluminescent Aluminum Complexes. React. Funct. Polym. 2004, 61, 43−52. (193) Tonami, H.; Uyama, H.; Oguchi, T.; Higashimura, H.; Kobayashi, S. Synthesis of a Soluble Polyphenol by Oxidative Polymerization of Bisphenol-A Using Iron-Salen Complex as Catalyst. Polym. Bull. 1999, 42, 125−129. (194) Tonami, H.; Uyama, H.; Kobayashi, S.; Higashimura, H.; Oguchi, T. Oxidative Polymerization of 2,6-Disubstituted Phenols Catalyzed by Iron-Salen Complex. J. Macromol. Sci., Part A: Pure Appl.Chem. 1999, A36, 719−730. (195) Ikeda, R.; Tanaka, H.; Uyama, H.; Kobayashi, S. Oxidative Polymerization of 2,6-Difluorophenol to Crystalline Poly(2,6-difluoro1,4-phenylene oxide). Macromolecules 2000, 33, 6648−6652. (196) Tonami, H.; Uyama, H.; Kobayashi, S. Oxidative CrossCoupling between Phenolic Polymer and Phenol-Containing Cellulose: Synthesis of a New Class of Artificial Wood Polymers. Macromolecules 2004, 37, 7901−7905. (197) Dubey, S.; Singh, D.; Misra, R. A. Enzymatic Synthesis and Various Properties of Poly(catechol). Enzyme Microb. Technol. 1998, 23, 432−437. (198) Xu, P.; Uyama, H.; Whitten, J. E.; Kobayashi, S.; Kaplan, D. L. Peroxidase-Catalyzed In Situ Polymerization of Surface Orientated Caffeic Acid. J. Am. Chem. Soc. 2005, 127, 11745−11753. (199) Bordoni, A.; Hrelia, S.; Angeloni, C.; Giordano, E.; Guarnieri, C.; Caldarera, C. M.; Biagi, P. L. Green Tea Protection of Hypoxia/ Reoxygenation Injury in Cultured Cardiac Cells. J. Nutr. Biochem. 2002, 13, 103−111. (200) Nakagawa, K.; Ninomiya, M.; Okubo, T.; Aoi, N.; Juneja, L. R.; Kim, M.; Yamanaka, K.; Miyazawa, T. Tea Catechin Supplementation Increases Antioxidant Capacity and Prevents Phospholipid Hydroperoxidation in Plasma of Humans. J. Agric. Food Chem. 1999, 47, 3967−3973. (201) Uyama, H. Artificial Polymeric Flavonoids: Synthesis and Applications. Macromol. Biosci. 2007, 7, 410−422. (202) Hamada, S.; Kontani, M.; Hosono, H.; Ono, H.; Tanaka, T.; Ooshima, T.; Mitsunaga, T.; Abe, I. Peroxidase-Catalyzed Generation of Catechin Oligomers that Inhibit Glucosyltransferase from Streptococcus sobrinus. FEMS Microbiol. Lett. 1996, 143, 35−40.

(203) López-Serrano, M.; Ros Barceló, A. Kinetic Properties of (+)-Catechin Oxidation by a Basic Peroxidase Isoenzyme from Strawberries. J. Food Sci. 1997, 62, 676−723. (204) Lopez-Serrano, M.; Ros Barceló, A. R. Reversed-Phase and Size-Exclusion Chromatography as Useful Tools in the Resolution of Peroxidase-Mediated (+)-Catechin Oxidation Products. J. Chromatogr. A 2001, 919, 267−273. (205) Lopez-Serrano, M.; Ros Barcelo, A. Comparative Study of the Products of the Peroxidase-Catalyzed and the PolyphenoloxidaseCatalyzed (+)-Catechin Oxidation. Their Possible Implications in Strawberry (Fragaria X ananassa) Browning Reactions. J. Agric. Food Chem. 2002, 50, 1218−1224. (206) Mejias, L.; Reihmann, M. H.; Sepulveda-Boza, S.; Ritter, H. New Polymers from Natural Phenols Using Horseradish or Soybean Peroxidase. Macromol. Biosci. 2002, 2, 24−32. (207) Bruno, F. F.; Nagarajan, S.; Nagarajan, R.; Kumar, J.; Samuelson, L. A. Biocatalytic Synthesis of Water-Soluble Oligo(catechins). J. Macromol. Sci., Part A: Pure Appl.Chem. 2005, 42, 1547− 1554. (208) Kurisawa, M.; Chung, J. E.; Kim, Y. J.; Uyama, H.; Kobayashi, S. Amplification of Antioxidant Activity and Xanthine Oxidase Inhibition of Catechin by Enzymatic Polymerization. Biomacromolecules 2003, 4, 469−471. (209) Kawakita, H.; Nakano, S.; Hamamoto, K.; Matsunaga, Y.; Yoshimura, Y.; Ohto, K.; Inoue, K. Copper-Ion Adsorption and GoldIon Reduction by Polyphenols Prepared by the Enzymatic Reaction of Horseradish Peroxidase. J. Appl. Polym. Sci. 2010, 118, 247−252. (210) Xu, P.; Singh, A.; Kaplan, D. L. Enzymatic Catalysis in the Synthesis of Polyanilines and Derivatives of Polyanilines. Adv. Polym. Sci. 2006, 194, 69−94. (211) Aizawa, M.; Wang, L. L.; Shinohara, H.; Ikariyama, Y. Enzymatic-Synthesis of Polyaniline Film Using a Copper-Containing Oxidoreductase - Bilirubin Oxidase. J. Biotechnol. 1990, 14, 301−310. (212) Samuelson, L. A.; Anagnostopoulos, A.; Alva, K. S.; Kumar, J.; Tripathy, S. K. Biologically Derived Conducting and Water Soluble Polyaniline. Macromolecules 1998, 31, 4376−4378. (213) Roy, S.; Fortier, J. M.; Nagarajan, R.; Tripathy, S.; Kumar, J.; Samuelson, L. A.; Bruno, F. F. Blomimetic Synthesis of a Water Soluble Conducting Molecular Complex of Polyaniline and Lignosulfonate. Biomacromolecules 2002, 3, 937−941. (214) Liu, W.; Kumar, J.; Tripathy, S.; Senecal, K. J.; Samuelson, L. Enzymatically Synthesized Conducting Polyaniline. J. Am. Chem. Soc. 1999, 121, 71−78. (215) Guo, Z. W.; Ruegger, H.; Kissner, R.; Ishikawa, T.; Willeke, M.; Walde, P. Vesicles as Soft Templates for the Enzymatic Polymerization of Aniline. Langmuir 2009, 25, 11390−11405. (216) Streltsov, A. V.; Shumakovich, G. P.; Morozova, O. V.; Gorbacheva, M. A.; Yaropolov, A. I. Micellar Laccase-Catalyzed Synthesis of Electroconductive Polyaniline. Appl. Biochem. Microbiol. 2008, 44, 264−270. (217) Kausaite, A.; Ramanaviciene, A.; Ramanavicius, A. Polyaniline Synthesis Catalysed by Glucose Oxidase. Polymer 2009, 50, 1846− 1851. (218) Ku, B. C.; Lee, S. H.; Liu, W.; He, J. A.; Kumar, J.; Bruno, F. F.; Samuelson, L. A. A New Approach to Catalyze Template Polymerization of Aniline Using Electrostatically Multilayered Hematin Assemblies. J. Macromol. Sci., Part A: Pure Appl.Chem. 2003, 40, 1335−1346. (219) Rumbau, V.; Marcilla, R.; Ochoteco, E.; Pomposo, J. A.; Mecerreyes, D. Ionic Liquid Immobilized Enzyme for Biocatalytic Synthesis of Conducting Polyaniline. Macromolecules 2006, 39, 8547− 8549. (220) Sahoo, S. K.; Nagarajan, R.; Samuelson, L.; Kumar, J.; Cholli, A. L.; Tripathy, S. K. Enzymatically Synthesized Polyaniline in the Presence of a Template Poly(vinylphosphonic acid): A Solid State NMR Study. J. Macromol. Sci., Part A: Pure Appl.Chem. 2001, 38, 1315−1328. (221) Alva, K. S.; Kumar, J.; Marx, K. A.; Tripathy, S. K. Enzymatic Synthesis and Characterization of a Novel Water-Soluble Polyaniline: 2392


Chemical Reviews

Review

Poly(2,5-diaminobenzenesulfonate). Macromolecules 1997, 30, 4024− 4029. (222) Liu, W.; Cholli, A. L.; Nagarajan, R.; Kumar, J.; Tripathy, S.; Bruno, F. F.; Samuelson, L. The Role of Template in the Enzymatic Synthesis of Conducting Polyaniline. J. Am. Chem. Soc. 1999, 121, 11345−11355. (223) Thiyagarajan, M.; Samuelson, L. A.; Kumar, J.; Cholli, A. L. Helical Conformational Specificity of Enzymatically Synthesized Water-Soluble Conducting Polyaniline Nanocomposites. J. Am. Chem. Soc. 2003, 125, 11502−11503. (224) Caramyshev, A. V.; Lobachov, V. M.; Selivanov, D. V.; Sheval, E. V.; Vorobiev, A. K.; Katasova, O. N.; Polyakov, V. Y.; Makarov, A. A.; Sakharov, I. Y. Micellar Peroxidase-Catalyzed Synthesis of Chiral Polyaniline. Biomacromolecules 2007, 8, 2549−2555. (225) Huh, P.; Kim, S. C.; Kim, Y.; Wang, Y.; Singh, J.; Kumar, J.; Samuelson, L. A.; Kim, B. S.; Jo, N. J.; Lee, J. O. Optical and Electrochemical Detection of Saccharides with Poly(aniline-co-3aminobenzeneboronic acid) Prepared from Enzymatic Polymerization. Biomacromolecules 2007, 8, 3602−3607. (226) Cruz-Silva, R.; Arizmendi, L.; Del-Angel, M.; Romero-Garcia, J. pH- and Thermosensitive Polyaniline Colloidal Particles Prepared by Enzymatic Polymerization. Langmuir 2007, 23, 8−12. (227) Kobayashi, S.; Kaneko, I.; Uyama, H. Enzymatic Oxidation Polymerization of Ortho-Phenylenediamine. Chem. Lett. 1992, 393− 394. (228) Cui, A.; Singh, A.; Kaplan, D. L. Enzyme-Based Molecular Imprinting with Metals. Biomacromolecules 2002, 3, 1353−1358. (229) Shumakovich, G.; Kurova, V.; Vasil'eva, I.; Pankratov, D.; Otrokhov, G.; Morozova, O.; Yaropolov, A. Laccase-mediated synthesis of conducting polyaniline. J. Mol. Catal. B: Enzym. 2012, 77, 105−110. (230) Otrokhov, G.; Pankratov, D.; Shumakovich, G.; Khlupova, M.; Zeifman, Y.; Vasil’eva, I.; Morozova, O.; Yaropolov, A. Enzymatic Synthesis of Polyaniline/Multi-Walled Carbon Nanotube Composite with Core Shell Structure and Its Electrochemical Characterization for Supercapacitor Application. Electrochim. Acta 2014, 123, 151−157. (231) Nabid, M. R.; Shamsianpour, M.; Sedghi, R.; Moghaddam, A. B. Enzyme-Catalyzed Synthesis of Conducting Polyaniline Nanocomposites with Pure and Functionalized Carbon Nanotubes. Chem. Eng. Technol. 2012, 35, 1707−1712. (232) Junker, K.; Kissner, R.; Rakvin, B.; Guo, Z. W.; Willeke, M.; Busato, S.; Weber, T.; Walde, P. The Use of Trametes Versicolor Laccase for the Polymerization of Aniline in the Presence of Vesicles as Templates. Enzyme Microb. Technol. 2014, 55, 72−84. (233) Nabid, M. R.; Asadi, S.; Sedghi, R.; Bayandori Moghaddam, A. Chemical and Enzymatic Polymerization of Polyaniline/Ag Nanocomposites. Chem. Eng. Technol. 2013, 36, 1411−1416. (234) Luong, N. D.; Korhonen, J. T.; Soininen, A. J.; Ruokolainen, J.; Johansson, L. S.; Seppala, J. Processable Polyaniline Suspensions Through in situ Polymerization onto Nanocellulose. Eur. Polym. J. 2013, 49, 335−344. (235) Carrillo, N.; Leon-Silva, U.; Avalos, T.; Nicho, M. E.; Serna, S.; Castillon, F.; Farias, M.; Cruz-Silva, R. Enzymatically Synthesized Polyaniline Film Deposition Studied by Simultaneous Open Circuit Potential and Electrochemical Quartz Crystal Microbalance Measurements. J. Colloid Interface Sci. 2012, 369, 103−110. (236) Roman, P.; Cruz-Silva, R.; Vazquez-Duhalt, R. PeroxidaseMediated Synthesis of Water-Soluble Fully Sulfonated Polyaniline. Synth. Met. 2012, 162, 794−799. (237) Sheng, Q. L.; Wang, M. Z.; Zheng, J. B. A Novel Hydrogen Peroxide Biosensor Based on Enzymatically Induced Deposition of Polyaniline on the Functionalized Graphene-Carbon Nanotube Hybrid Materials. Sens. Actuators, B 2011, 160, 1070−1077. (238) Kausaite-Minkstimiene, A.; Mazeiko, V.; Ramanaviciene, A.; Ramanavicius, A. Enzymatically Synthesized Polyaniline Layer for Extension of Linear Detection Region of Amperometric Glucose Biosensor. Biosens. Bioelectron. 2010, 26, 790−797. (239) Berron, B. J.; Johnson, L. M.; Ba, X.; McCall, J. D.; Alvey, N. J.; Anseth, K. S.; Bowman, C. N. Glucose Oxidase-Mediated Polymer-

ization as a Platform for Dual-Mode Signal Amplification and Biodetection. Biotechnol. Bioeng. 2011, 108, 1521−1528. (240) Rumbau, V.; Pomposo, J. A.; Eleta, A.; Rodriguez, J.; Grande, H.; Mecerreyes, D.; Ochoteco, E. First Enzymatic Synthesis of WaterSoluble Conducting Poly(3,4-ethylenedioxythiophene). Biomacromolecules 2007, 8, 315−317. (241) Sikora, T.; Marcilla, R.; Mecerreyes, D.; Rodriguez, J.; Pomposo, J. A.; Ochoteco, E. Enzymatic Synthesis of Water-Soluble Conducting Poly(3,4-ethylenedioxythiophene): A Simple Enzyme Immobilization Strategy for Recycling and Reusing. J. Polym. Sci., Part A: Polym. Chem. 2009, 47, 306−309. (242) Krikstolaityte, V.; Kuliesius, J.; Ramanaviciene, A.; Mikoliunaite, L.; Kausaite-Minkstimiene, A.; Oztekin, Y.; Ramanavicius, A. Enzymatic Polymerization of Polythiophene by Immobilized Glucose Oxidase. Polymer 2014, 55, 1613−1620. (243) Hira, S. M.; Payne, C. K. Protein-Mediated Synthesis of the Conducting Polymer PEDOT:PSS. Synth. Met. 2013, 176, 104−107. (244) Sheng, Q. L.; Wang, J.; Zheng, J. B.; Xu, Z. Q.; Zhang, H. F. Ultrasensitive Electrical Biosensing of Syphilis DNA Using TargetGuided Formation of Polyaniline Based on Enzyme-Catalyzed Polymerization. Biosens. Bioelectron. 2010, 25, 2071−2077. (245) Gitsov, I.; Wang, L. L.; Vladimirov, N.; Simonyan, A.; Kiemle, D. J.; Schutz, A. ″Green″ Synthesis of Unnatural Poly(Amino Acid)s with Zwitterionic Character and pH-Responsive Solution Behavior, Mediated by Linear Dendritic Laccase Complexes. Biomacromolecules 2014, 15, 4082−4095. (246) Derango, R. A.; Chiang, L. C.; Dowbenko, R.; Lasch, J. G. Enzyme-Mediated Polymerization of Acrylic Monomers. Biotechnol. Tech. 1992, 6, 523−526. (247) Emery, O.; Lalot, T.; Brigodiot, M.; Marechal, E. Free-radical polymerization of acrylamide by horseradish peroxidase-mediated initiation. J. Polym. Sci., Part A: Polym. Chem. 1997, 35, 3331−3333. (248) Teixeira, D.; Lalot, T.; Brigodiot, M.; Marechal, E. β-Diketones as Key Compounds in Free-Radical Polymerization by EnzymeMediated Initiation. Macromolecules 1999, 32, 70−72. (249) Ikeda, R.; Tanaka, H.; Uyama, H.; Kobayashi, S. LaccaseCatalyzed Polymerization of Acrylamide. Macromol. Rapid Commun. 1998, 19, 423−425. (250) Kohri, M.; Uzawa, S.; Kobayashi, A.; Fukushima, H.; Taniguchi, T.; Nakahira, T. Enzymatic Emulsifier-Free Emulsion Polymerization to Prepare Polystyrene Particles using Horseradish Peroxidase as a Catalyst. Polym. J. 2013, 45, 354−358. (251) Kohri, M.; Kobayashi, A.; Fukushima, H.; Kojima, T.; Taniguchi, T.; Saito, K.; Nakahira, T. Enzymatic Miniemulsion Polymerization of Styrene with a Polymerizable Surfactant. Polym. Chem. 2012, 3, 900−906. (252) Kohri, M.; Kobayashi, A.; Fukushima, H.; Taniguchi, T.; Nakahira, T. Effect of Surfactant Type on Enzymatic Miniemulsion Polymerization Using Horseradish Peroxidase as a Catalyst. Chem. Lett. 2012, 41, 1131−1133. (253) Di Gennaro, P.; Bargna, A.; Bruno, F.; Sello, G. Purification of Recombinant Catalase-Peroxidase HPI from E. coli and Its Application in Enzymatic Polymerization Reactions. Appl. Microbiol. Biotechnol. 2014, 98, 1119−1126. (254) Kalra, B.; Gross, R. A. HRP-Mediated Polymerizations of Acrylamide and Sodium Acrylate. Green Chem. 2002, 4, 174−178. (255) Villarroya, S.; Thurecht, K. J.; Howdle, S. M. HRP-Mediated Inverse Emulsion Polymerisation of Acrylamide in Supercritical Carbon Dioxide. Green Chem. 2008, 10, 863−867. (256) Zhao, Q.; Sun, J. Z.; Ren, H.; Zhou, Q. Y.; Lin, Q. C. Horseradish Peroxidase Immobilized in Macroporous Hydrogel for Acrylamide Polymerization. J. Polym. Sci., Part A: Polym. Chem. 2008, 46, 2222−2232. (257) Tsujimoto, T.; Uyama, H.; Kobayashi, S. Polymerization of vinyl monomers using oxidase catalysts. Macromol. Biosci. 2001, 1, 228−232. (258) Hollmann, F.; Gumulya, Y.; Tolle, C.; Liese, A.; Thum, O. Evaluation of the Laccase from Myceliophthora thermophila as 2393


Chemical Reviews

Review

Industrial Biocatalyst for Polymerization Reactions. Macromolecules 2008, 41, 8520−8524. (259) Nesterova, N. A.; Gavrilova, I. I.; Panarin, E. F. Enzymatic Polymerization of Vinyl Monomers. Russ. J. Appl. Chem. 2007, 80, 2129−2131. (260) Shogren, R. L.; Willett, J. L.; Biswas, A. HRP-Mediated Synthesis of Starch-Polyacrylamide Graft Copolymers. Carbohydr. Polym. 2009, 75, 189−191. (261) Kadokawa, J.; Kokubo, A.; Tagaya, H. Free-Radical Polymerization of Vinyl Monomers Using Hematin as a Biomimetic Catalyst in Place of Enzyme. Macromol. Biosci. 2002, 2, 257−260. (262) Singh, A.; Kaplan, D. L. Vitamin C Functionalized Poly(methyl methacrylate) for Free Radical Scavenging. J. Macromol. Sci., Part A: Pure Appl.Chem. 2004, 41, 1377−1386. (263) Wang, Y. Z.; Singh, A.; Xu, P.; Pindrus, M. A.; Blasioli, D. J.; Kaplan, D. L. Expansion and Osteogenic Differentiation of Bone Marrow-Derived Mesenchymal Stem Cells on a Vitamin C Functionalized Polymer. Biomaterials 2006, 27, 3265−3273. (264) Singh, A.; Kaplan, D. L. Biocatalytic Route to Ascorbic AcidModified Polymers for Free-Radical Scavenging. Adv. Mater. 2003, 15, 1291−1294. (265) Singh, A.; Roy, S.; Samuelson, L.; Bruno, F.; Nagarajan, R.; Kumar, J.; John, V.; Kaplan, D. L. Peroxidase, Hematin, and PegylatedHematin Catalyzed Vinyl Polymerizations in Water. J. Macromol. Sci., Part A: Pure Appl.Chem. 2001, 38, 1219−1230. (266) Qi, G. G.; Jones, C. W.; Schork, F. J. Enzyme-Initiated Miniemulsion Polymerization. Biomacromolecules 2006, 7, 2927−2930. (267) Kohri, M. Development of HRP-Mediated Enzymatic Polymerization Under Heterogeneous Conditions for the Preparation of Functional Particles. Polym. J. 2014, 46, 373−380. (268) Zaidi, K. U.; Ali, A. S.; Ali, S. A.; Naaz, I. Microbial Tyrosinases: Promising Enzymes for Pharmaceutical, Food Bioprocessing, and Environmental Industry. Biochem. Res. Int. 2014, 2014, 1− 16. (269) Hay, A. S.; Blanchard, H. S.; Endres, G. F.; Eustance, J. W. Polymerization by Oxidative Coupling. J. Am. Chem. Soc. 1959, 81, 6335−6336. (270) Higashimura, H. Oxidative Coupling Polymerization. In Polymer Science: A Comprehensive Reference; Möller, K. M., Ed.; Elsevier: Amsterdam, 2012; pp 141−173. (271) Baesjou, P. J.; Driessen, W. L.; Challa, G.; Reedijk, J. Ab initio Calculations on 2,6-Dimethylphenol and 4-(2,6-Dimethylphenoxy)2,6-dimethylphenol. Evidence of an Important Role for the Phenoxonium Cation in the Copper-Catalyzed Oxidative Phenol Coupling Reaction. J. Am. Chem. Soc. 1997, 119, 12590−12594. (272) Ikeda, R.; Uyama, H.; Kobayashi, S. Novel Synthetic Pathway to a Poly(phenylene oxide). Laccase-Catalyzed Oxidative Polymerization of Syringic Acid. Macromolecules 1996, 29, 3053−3054. (273) Uyama, H.; Ikeda, R.; Yaguchi, S.; Kobayashi, S. Enzymatic Polymerization of Natural Phenol Derivatives and Enzymatic Synthesis of Polyesters from Vinyl Esters. In Polymers from Renewable Resources; Biopolyesters and Biocatalysis; Scholz, C., Gross, R. A., Eds.; ACS Symposium Series 764; American Chemical Society: Washington, DC, 2000; pp 113−127. (274) Aktas, N.; Kibarer, G.; Tanyolac, A. Effects of Reaction Conditions on Laccase-Catalyzed α-Naphthol Polymerization. J. Chem. Technol. Biotechnol. 2000, 75, 840−846. (275) Marjasvaara, A.; Torvinen, M.; Kinnunen, H.; Vainiotalo, P. Laccase-Catalyzed Polymerization of Two Phenolic Compounds Studied by Matrix-Assisted Laser Desorption/Ionization Time-ofFlight and Electrospray Ionization Fourier Transform Ion Cyclotron Resonance Mass Spectrometry with Collision-Induced Dissociation Experiments. Biomacromolecules 2006, 7, 1604−1609. (276) Okusa, K.; Miyakoshi, T.; Chen, C. L. Comparative Studies on Dehydrogenative Polymerization of Coniferyl Alcohol by Laccases and Peroxidases. 1. Preliminary Results. Holzforschung 1996, 50, 15−23. (277) Milstein, O.; Hüttermann, A.; Fründ, R.; Lüdemann, H.-D. Enzymatic Co-polymerization of Lignin with Low-Molecular Mass Compounds. Appl. Microbiol. Biotechnol. 1994, 40, 760−767.

(278) Mita, N.; Tawaki, S.; Uyama, H.; Kobayashi, S. LaccaseCatalyzed Oxidative Polymerization of Phenols. Macromol. Biosci. 2003, 3, 253−257. (279) Yalcinkaya, Z.; Gun, S.; Sahan, T.; Birhanli, E.; Sahiner, N.; Aktas, N.; Yesilada, O. Influence of the Medium Conditions on Enzymatic Oxidation of Bisphenol A. Can. J. Chem. Eng. 2014, 92, 712−719. (280) Tanaka, T.; Takahashi, M.; Hagino, H.; Nudejima, S. I.; Usui, H.; Fujii, T.; Taniguchi, M. Enzymatic Oxidative Polymerization of Methoxyphenols. Chem. Eng. Sci. 2010, 65, 569−573. (281) López, J.; Alonso-Omlin, E. M.; Hernández-Alcántara, J. M.; Bárzana, E.; Gimeno, M. Novel Photoluminescent Material by Laccase-Mediated Polymerization of 4-Fluoroguaiacol Throughout Defluorination. J. Mol. Catal. B: Enzym. 2014, 109, 70−75. (282) Higashimura, H.; Fujisawa, K.; Moro-oka, Y.; Kubota, M.; Shiga, A.; Terahara, A.; Uyama, H.; Kobayashi, S. Highly Regioselective Oxidative Polymerization of 4-Phenoxyphenol to Poly(1,4-phenylene oxide) Catalyzed by Tyrosinase Model Complexes. J. Am. Chem. Soc. 1998, 120, 8529−8530. (283) Higashimura, H.; Kubota, M.; Shiga, A.; Fujisawa, K.; Morooka, Y.; Uyama, H.; Kobayashi, S. ″Radical-Controlled″ Oxidative Polymerization of 4-Phenoxyphenol by a Tyrosinase Model complex Catalyst to Poly(1,4-phenylene oxide). Macromolecules 2000, 33, 1986−1995. (284) Higashimura, H.; Fujisawa, K.; Namekawa, S.; Kubota, M.; Shiga, A.; Moro-Oka, Y.; Uyama, H.; Kobayashi, S. Coupling Selectivity in the Radical-Controlled Oxidative Polymerization of 4Phenoxyphenol Catalyzed by (1,4,7-Triisopropyl-1,4,7triazacyclononane)copper(II) Complex. J. Polym. Sci., Part A: Polym. Chem. 2000, 38, 4792−4804. (285) Higashimura, H.; Kubota, M.; Shiga, A.; Kodera, M.; Uyama, H.; Kobayashi, S. ″Radical-Controlled″ Oxidative Polymerization of 4Phenoxyphenol Catalyzed by a Dicopper Complex of a Dinucleating Ligand. J. Mol. Catal. A: Chem. 2000, 161, 233−237. (286) Higashimura, H.; Fujisawa, K.; Kubota, M.; Kobayashi, S. ″Radical-Controlled″’ Oxidative Polymerization of Phenol: Comparison with that of 4-Phenoxyphenol. J. Polym. Sci., Part A: Polym. Chem. 2005, 43, 1955−1962. (287) Higashimura, H.; Fujisawa, K.; Moro-oka, Y.; Namekawa, S.; Kubota, M.; Shiga, A.; Uyama, H.; Kobayashi, S. New Crystalline Polymers: Poly(2,5-dialkyl-l,4phenylene oxide)s. Macromol. Rapid Commun. 2000, 21, 1121−1124. (288) Cheng, S. Z. D.; Wunderlich, B. Glass-Transition and Melting Behavior of Poly(oxy-2,6-dimethyl-1,4-phenylene). Macromolecules 1987, 20, 1630−1637. (289) Higashimura, H.; Fujisawa, K.; Moro-oka, Y.; Kubota, M.; Shiga, A.; Uyama, H.; Kobayashi, S. ’Radical-controlled’ Oxidative Polymerization of m-Cresol Catalyzed by μ-η2:η2-Peroxo Dicopper(II) Complex. J. Mol. Catal. A: Chem. 2000, 155, 201−207. (290) Higashimura, H.; Fujisawa, K.; Moro-oka, Y.; Kubota, M.; Shiga, A.; Uyama, H.; Kobayashi, S. ’Radical-Controlled’ Oxidative Polymerization of o-Cresol Catalyzed by μ-η2:η2-Peroxo Dicopper(II) Complex. Appl. Catal., A 2000, 194-195, 427−433. (291) Kitajima, N.; Moro-oka, Y. Copper-Dioxygen Complexes Inorganic and Bioinorganic Perspectives. Chem. Rev. 1994, 94, 737− 757. (292) Kubota, M.; Shiga, A.; Higashimura, H.; Fujisawa, K.; Morooka, Y.; Uyama, H.; Kobayashi, S. Ab initio Calculation on the Reaction Mechanism of ″Radical-Controlled″ Oxidative Polymerization of Phenols. Bull. Chem. Soc. Jpn. 2004, 77, 813−818. (293) Sun, X.; Bai, R.; Zhang, Y.; Wang, Q.; Fan, X.; Yuan, J.; Cui, L.; Wang, P. Laccase-Catalyzed Oxidative Polymerization of Phenolic Compounds. Appl. Biochem. Biotechnol. 2013, 171, 1673−1680. (294) Smejkalova, D.; Piccolo, A.; Spiteller, M. Oligomerization of Humic Phenolic Monomers by Oxidative Coupling Under Biomimetic Catalysis. Environ. Sci. Technol. 2006, 40, 6955−6962. (295) Smejkalova, D.; Conte, P.; Piccolo, A. Structural Characterization of Isomeric Dimers from the Oxidative Oligomerization of 2394


Chemical Reviews

Review

Catechol with a Biomimetic Catalyst. Biomacromolecules 2007, 8, 737− 743. (296) Yoshida, T.; Lu, R.; Han, S. Q.; Hattori, K.; Katsuta, T.; Takeda, K. I.; Sugimoto, K.; Funaoka, M. Laccase-Catalyzed Polymerization of Lignocatechol and Affinity on Proteins of Resulting Polymers. J. Polym. Sci., Part A: Polym. Chem. 2009, 47, 824−832. (297) Garcia-Ubasart, J.; Vidal, T.; Torres, A. L.; Rojas, O. J. LaccaseMediated Coupling of Nonpolar Chains for the Hydrophobization of Lignocellulose. Biomacromolecules 2013, 14, 1637−1644. (298) Kurisawa, M.; Chung, J. E.; Uyama, H.; Kobayashi, S. LaccaseCatalyzed Synthesis and Antioxidant Property of Poly(catechin). Macromol. Biosci. 2003, 3, 758−764. (299) Chung, J. E.; Kurisawa, M.; Tachibana, Y.; Uyama, H.; Kobayashi, S. Enzymatic Synthesis and Antioxidant Property of Poly(allylamine)-Catechin Conjugate. Chem. Lett. 2003, 32, 620−621. (300) Ihara, N.; Schmitz, S.; Kurisawa, M.; Chung, J. E.; Uyama, H.; Kobayashi, S. Amplification of Inhibitory Activity of Catechin Against Disease-Related Enzymes by Conjugation on Poly(ε-lysine). Biomacromolecules 2004, 5, 1633−1636. (301) Kurisawa, M.; Chung, J. E.; Uyama, H.; Kobayashi, S. Enzymatic Synthesis and Antioxidant Properties of Poly(rutin). Biomacromolecules 2003, 4, 1394−1399. (302) Jiménez, M.; Garcia-Carmona, F. Oxidation of the Flavonol Quercetin by Polyphenol Oxidase. J. Agric. Food Chem. 1999, 47, 56− 60. (303) Jeon, J. K.; Lee, J.; Imm, J. Y. Effects of Laccase-Catalyzed Rutin Polymer Fraction on Adipogenesis Inhibition in 3T3-L1 Adipocytes. Process Biochem. 2014, 49, 1189−1195. (304) Desentis-Mendoza, R. M.; Hernandez-Sanchez, H.; Moreno, A.; Rojas del c, E.; Chel-Guerrero, L.; Tamariz, J.; Jaramillo-Flores, M. E. Enzymatic Polymerization of Phenolic Compounds Using Laccase and Tyrosinase from. Biomacromolecules 2006, 7, 1845−1854. (305) Madad, N.; Chebil, L.; Charbonnel, C.; Ioannou, I.; Ghoul, M. Enzymatic Polymerization of Sodium Lignosulfonates: Effect of Catalysts, Initial Molecular Weight, and Mediators. Can. J. Chem. 2013, 91, 220−225. (306) Yoshida, H. LXIII.Chemistry of Lacquer (Urushi). Part I. Communication from the Chemical Society of Tokio. J. Chem. Soc., Trans. 1883, 43, 472−486. (307) Bertrand, G. Sur le Latex de L’abre à Laque. Compt. Rend. 1894, 118, 1213−1218. (308) Majima, R. Ü ber den Hauptbestandteil des Japanlacks. (I. Mitteilung.) Ü ber Urushiol und Urushioldimethyläther. Ber. Dtsch. Chem. Ges. 1909, 42, 1418−1423. (309) Majima, R. Ü ber den Hauptbestandteil des Japan-Lacks, IX. Mitteilung: Chemische Untersuchung der verschiedenen natürlichen Lackarten, die dem Japan-Lack nahe verwandt sind. Ber. Dtsch. Chem. Ges. B 1922, 55, 191−214. (310) Symes, W. F.; Dawson, C. R. Poison Ivy “Urushiol. J. Am. Chem. Soc. 1954, 76, 2959−2963. (311) Loev, B.; Dawson, C. R. On the Geometrical Configuration of the Olefinic Components of Poison Ivy Urushiol. The Synthesis of a Model Compound. J. Am. Chem. Soc. 1956, 78, 1180−1183. (312) Rogue, B. V. A History of Japanese Lacquer Work; University of Toronto Press: Toronto, 1976. (313) Vogl, O. Oriental Lacquer, Poison Ivy, and Drying Oils. J. Polym. Sci., Part A: Polym. Chem. 2000, 38, 4327−4335. (314) Kumanodani, J. Oriental Lacquer. In Polymeric Materials Encyclopedia; Salamone, J. C., Ed.; CRC Press, Inc.: Boca Raton, FL, 1996; pp 4835−4842. (315) Kobayashi, S.; Ikeda, R.; Oyabu, H.; Tanaka, H.; Uyama, H. Artificial Urushi: Design, Synthesis, and Enzymatic Curing of New Urushiol Analogues. Chem. Lett. 2000, 1214−1215. (316) Ikeda, R.; Tanaka, H.; Oyabu, H.; Uyama, H.; Kobayashi, S. Preparation of Artificial Urushi via an Environmentally Benign Process. Bull. Chem. Soc. Jpn. 2001, 74, 1067−1073. (317) Tsujimoto, T.; Ando, N.; Oyabu, H.; Uyama, H.; Kobayashi, S. Laccase-Catalyzed Curing of Natural Phenolic Lipids and Product

Properties. J. Macromol. Sci., Part A: Pure Appl.Chem. 2007, 44, 1055− 1060. (318) Tsujimoto, T.; Ikeda, R.; Uyama, H.; Kobayashi, S. Synthesis and Curing of Crosslinkable Polyphenols from Urushiol Analogues. Chem. Lett. 2000, 1122−1123. (319) Souza, F. G.; Richa, P.; de Siervo, A.; Oliveira, G. E.; Rodrigues, C. H. M.; Nele, M.; Pinto, J. C. New in situ Blends of Polyaniline and Cardanol Bio-resins. Macromol. Mater. Eng. 2008, 293, 675−683. (320) Shibata, M.; Itakura, Y.; Watanabe, H. Bio-based Thermosetting Resins Composed of Cardanol Novolac and Bismaleimide. Polym. J. 2013, 45, 758−765. (321) Ikeda, R.; Kobayashi, S.; Uyama, H. Resin Composition and Process for Producing the Same. U.S. Patent US 6,344,516 B1, 2002. (322) Ikeda, R.; Kobayashi, S.; Uyama, H. Resin Composition and Process for Producing the Same. Jpn. Patent 3,760,406, 2006. (323) Ikeda, R.; Kobayashi, S.; Uyama, H. Resin Composition and Process for Producing the Same. Eur. Patent EP 1,114,836 B1, 2006. (324) Voirin, C.; Caillol, S.; Sadavarte, N. V.; Tawade, B. V.; Boutevin, B.; Wadgaonkar, P. P. Functionalization of Cardanol: Towards Biobased Polymers and Additives. Polym. Chem. 2014, 5, 3142−3162. (325) Watanabe, H.; Fujimoto, A.; Takahara, A. Characterization of Catechol-Containing Natural Thermosetting Polymer ’Urushiol’ Thin Film. J. Polym. Sci., Part A: Polym. Chem. 2013, 51, 3688−3692. (326) Watanabe, H.; Fujimoto, A.; Takahara, A. Surface Texturing of ̀ Natural Urushi’ Thermosetting Polymer Thin Films. Polym. J. 2014, 46, 216−219. (327) Singh, A.; Kaplan, D. L. In vitro Enzyme-Induced Vinyl Polymerization. In Advances in Polymer Science; Springer-Verlag: Berlin, 2006; Vol. 194, pp 211−224. (328) Kurniawati, S.; Nicell, J. A. Characterization of Trametes versicolor laccase for the transformation of aqueous phenol. Bioresour. Technol. 2008, 99, 7825−7834. (329) Ikeda, R.; Uyama, H.; Kobayashi, S. Laccase-Catalyzed Curing of Vinyl Polymers Bearing a Phenol Moiety in the Side Chain. Polym. J. 2001, 33, 540−542. (330) Ikeda, R.; Tanaka, H.; Uyama, H.; Kobayashi, S. A New Crosslinking Method of Vinyl Polymers Having a Phenol Moiety via Oxidative Coupling. Polym. J. 2001, 33, 959−961. (331) Fukuoka, T.; Uyama, H.; Kobayashi, S. Polymerization of Polyfunctional Macromolecules: Synthesis of a New Class of High Molecular Weight Poly(amino acid)s by Oxidative Coupling of Phenol-containing Precursor Polymers. Biomacromolecules 2004, 5, 977−983. (332) Nieto, M.; Nardecchia, S.; Peinado, C.; Catalina, F.; Abrusci, C.; Gutierrez, M. C.; Ferrer, M. L.; del Monte, F. Enzyme-induced Graft Polymerization for Preparation of Hydrogels: Synergetic Effect of Laccase-immobilized-cryogels for Pollutants Adsorption. Soft Matter 2010, 6, 3533−3540. (333) Tan, Y. M.; Deng, W. F.; Li, Y. Y.; Huang, Z.; Meng, Y.; Xie, Q. J.; Ma, M.; Yao, S. Z. Polymeric Bionanocomposite Cast Thin Films with In Situ Laccase-Catalyzed Polymerization of Dopamine for Biosensing and Biofuel Cell Applications. J. Phys. Chem. B 2010, 114, 5016−5024. (334) Payne, G. F.; Chaubal, M. V.; Barbari, T. A. Enzyme-Catalysed Polymer Modification: Reaction of Phenolic Compounds with Chitosan Films. Polymer 1996, 37, 4643−4648. (335) Kumar, G.; Smith, P. J.; Payne, G. F. Enzymatic Grafting of a Natural Product onto Chitosan to Confer Water Solubility Under Basic Conditions. Biotechnol. Bioeng. 1999, 63, 154−165. (336) Wu, L. Q.; Embree, H. D.; Balgley, B. M.; Smith, P. J.; Payne, G. F. Utilizing Renewable Resources to Create Functional Polymers: Chitosan-Based Associative Thickener. Environ. Sci. Technol. 2002, 36, 3446−3454. (337) Chen, T. H.; Vazquez-Duhalt, R.; Wu, C. F.; Bentley, W. E.; Payne, G. F. Combinatorial Screening for Enzyme-Mediated Coupling. Tyrosinase-Catalyzed Coupling to Create Protein-Chitosan Conjugates. Biomacromolecules 2001, 2, 456−462. 2395


Chemical Reviews

Review

(338) Chen, T. H.; Embree, H. D.; Wu, L. Q.; Payne, G. F. In vitro Protein-Polysaccharide Conjugation: Tyrosinase-Catalyzed Conjugation of Gelatin and Chitosan. Biopolymers 2002, 64, 292−302. (339) Chen, T. H.; Embree, H. D.; Brown, E. M.; Taylor, M. M.; Payne, G. F. Enzyme-Catalyzed Gel Formation of Gelatin and Chitosan: Potential for In Situ Applications. Biomaterials 2003, 24, 2831−2841. (340) Chen, T. H.; Small, D. A.; Wu, L. Q.; Rubloff, G. W.; Ghodssi, R.; Vazquez-Duhalt, R.; Bentley, W. E.; Payne, G. F. Nature-inspired Creation of Protein-polysaccharide Conjugate and Its Subsequent Assembly onto a Patterned Surface. Langmuir 2003, 19, 9382−9386. (341) Sampaio, S.; Taddei, P.; Monti, P.; Buchert, J.; Freddi, G. Enzymatic Grafting of Chitosan onto Bombyx mori Silk Fibroin: Kinetic and IR Vibrational Studies. J. Biotechnol. 2005, 116, 21−33. (342) Monti, P.; Freddi, G.; Sampaio, S.; Tsukada, M.; Taddei, P. Structure Modifications Induced in Silk Fibroin by Enzymatic Treatments. A Raman Study. J. Mol. Struct. 2005, 744-747, 685−690. (343) Freddi, G.; Anghileri, A.; Sampaio, S.; Buchert, J.; Monti, P.; Taddei, P. Tyrosinase-Catalyzed Modification of Bombyx mori Silk Fibroin: Grafting of Chitosan Under Heterogeneous Reaction Conditions. J. Biotechnol. 2006, 125, 281−294. (344) Anghileri, A.; Lantto, R.; Kruus, K.; Arosio, C.; Freddi, G. Tyrosinase-Catalyzed Grafting of Sericin Peptides onto Chitosan and Production of Protein-Polysaccharide Bioconjugates. J. Biotechnol. 2007, 127, 508−519. (345) Guerra, A.; Ferraz, A.; Cotrim, A. R.; da Silva, F. T. Polymerization of Lignin Fragments Contained in a Model Effluent by Polyphenoloxidases and Horseradish Peroxidase/Hydrogen Peroxide System. Enzyme Microb. Technol. 2000, 26, 315−323. (346) Demolliens, A.; Boucher, C.; Durocher, Y.; Jolicoeur, M.; Buschmann, M. D.; De Crescenzo, G. Tyrosinase-Catalyzed Synthesis of a Universal Coil-Chitosan Bioconjugate for Protein Immobilization. Bioconjugate Chem. 2008, 19, 1849−1854. (347) Wang, L.; Kobatake, E.; Ikariyama, Y.; Aizawa, M. Regioselective Oxidative Polymerization of 1,5-Dihydroxynaphthalene Catalyzed by Bilirubin Oxidase in a Water−Organic Solvent Mixed Solution. J. Polym. Sci., Part A: Polym. Chem. 1993, 31, 2855−2861. (348) Streltsov, A. V.; Morozova, O. V.; Arkharova, N. A.; Klechkovskaya, V. V.; Staroverova, I. N.; Shumakovich, G. P.; Yaropolov, A. I. Synthesis and Characterization of Conducting Polyaniline Prepared by Laccase-Catalyzed Method in Sodium Dodecylbenzenesulfonate Micellar Solutions. J. Appl. Polym. Sci. 2009, 114, 928−934. (349) Shumakovich, G.; Streltsov, A.; Gorshina, E.; Rusinova, T.; Kurova, V.; Vasil’eva, I.; Otrokhov, G.; Morozova, O.; Yaropolov, A. Laccase-Catalyzed Oxidative Polymerization of Aniline Dimer (NPhenyl-1,4-phenylenediamine) in Aqueous Micellar Solution of Sodium Dodecylbenzenesulfonate. J. Mol. Catal. B: Enzym. 2011, 69, 83−88. (350) Junker, K.; Luginbuhl, S.; Schuttel, M.; Bertschi, L.; Kissner, R.; Schuler, L. D.; Rakvin, B.; Walde, P. Efficient Polymerization of the Aniline Dimer p-Aminodiphenylamine (PADPA) with Trametes versicolor Laccase/O2 as Catalyst and Oxidant and AOT Vesicles as Templates. ACS Catal. 2014, 4, 3421−3434. (351) Ciric-Marjanovic, G.; Trchova, M.; Konyushenko, E. N.; Holler, P.; Stejskal, J. Chemical Oxidative Polymerization of Aminodiphenylamines. J. Phys. Chem. B 2008, 112, 6976−6987. (352) Dearmitt, C.; Armes, S. P. Colloidal Dispersions of SurfactantStabilized Polypyrrole Particles. Langmuir 1993, 9, 652−654. (353) Song, H. K.; Palmore, G. T. R. Conductive Polypyrrole via Enzyme Catalysis. J. Phys. Chem. B 2005, 109, 19278−19287. (354) Junker, K.; Zandomeneghi, G.; Schuler, L. D.; Kissner, R.; Walde, P. Enzymatic Polymerization of Pyrrole with Trametes versicolor Laccase and Dioxygen in the Presence of Vesicles Formed from AOT (Sodium bis-(2-ethylhexyl) sulfosuccinate) as Templates. Synth. Met. 2015, 200, 123−134. (355) van der Vlist, J.; Loos, K. Transferases in Polymer Chemistry. In Enzymatic Polymerisation; Advances in Polymer Science; Palmans,

A. R. A., Heise, A., Guebitz, G. M., Eds.; Springer: Berlin, 2010; pp 21−54. (356) Schuerch, C. Polysaccharides. In Encyclopedia of Polymer Science and Engineering, 2nd ed.; Mark, H. F., Bilkales, N., Overberger, C. G., Eds.; John Wiley & Sons: New York, 1986; Vol. 13, pp 87−162. (357) Paulsen, H. Advances in Selective Chemical Syntheses of Complex Oligosaccharides. Angew. Chem., Int. Ed. Engl. 1982, 21, 155−173. (358) Schmidt, R. R. New Methods for the Synthesis of Glycosides and OligosaccharidesAre There Alternatives to the Koenigs-Knorr Method? Angew. Chem., Int. Ed. Engl. 1986, 25, 212−235. (359) Toshima, K.; Tatsuta, K. Recent Progress in O-Glycosylation Methods and Its Application to Natural Products Synthesis. Chem. Rev. 1993, 93, 1503−1531. (360) Kobayashi, S.; Shoda, S. Enzymatic Synthesis of Polysaccharides: A New Concept in Polymerization Chemistry. In New Macromolecular Architecture and Functions; Kamachi, M., Nakamura, A., Eds.; Springer: Heidelberg, 1996; pp 171−180. (361) Shoda, S.; Izumi, R.; Fujita, M. Green Process in Glycotechnology. Bull. Chem. Soc. Jpn. 2003, 76, 1−13. (362) Blixt, O.; Razi, N. Enzymatic Glycosylation by Transferases. In Glycoscience Chemistry and Chemical Biology; 2nd ed.; Fraser-Reid, B. O., Tatsuta, K., Thiem, J., Eds.; Springer: New York, 2008; pp 1361− 1385. (363) Thiem, J.; Thiem, J. Enzymatic Glycosylation by Glycohydrolases and Glycosynthases. In Glycoscience Chemistry and Chemical Biology; 2nd ed.; Fraser-Reid, B. O., Tatsuta, K., Thiem, J., Eds.; Springer: Cham; New York, 2008; pp 1387−1409. (364) Taniguchi, N.; Honke, K.; Fukuda, M. Handbook of Glycosyltransferases and Related Genes; Springer: New York, 2002. (365) Murrell, M. P.; Yarema, K. J.; Levchenko, A. The Systems Biology of Glycosylation. ChemBioChem 2004, 5, 1334−1347. (366) Scheible, W. R.; Pauly, M. Glycosyltransferases and Cell Wall Biosynthesis: Novel Players and Insights. Curr. Opin. Plant Biol. 2004, 7, 285−295. (367) Somerville, C. Cellulose Synthesis in Higher Plants. Annu. Rev. Cell Dev. Biol. 2006, 22, 53−78. (368) Palcic, M. M.; Hindsgaul, O. Glycosyltransferases in the Synthesis of Oligosaccharide Analogs. Trends Glycosci. Glycotechnol. 1996, 8, 37−49. (369) Fujii, K.; Takata, H.; Yanase, M.; Terada, Y.; Ohdan, K.; Takaha, T.; Okada, S.; Kuriki, T. Bioengineering and Application of Novel Glucose Polymers. Biocatal. Biotransform. 2003, 21, 167−172. (370) Seibel, J.; Beine, R.; Moraru, R.; Behringer, C.; Buchholz, K. A new Pathway for the Synthesis of Oligosaccharides by the Use of NonLeloir Glycosyltransferases. Biocatal. Biotransform. 2006, 24, 157−165. (371) Seibel, J.; Jordening, H. J.; Buchholz, K. Glycosylation with Activated Sugars Using Glycosyltransferases and Transglycosidases. Biocatal. Biotransform. 2006, 24, 311−342. (372) Kadokawa, J.; Kaneko, Y. Engineering of Polysaccharide Materials-by Phosphorylase-Catalyzed Enzymatic Chain-Elongation; Pan Stanford Publishing Pte Ltd.: Singapore, 2013. (373) Kitaoka, M.; Hayashi, K. Carbohydrate-Processing Phosphorolytic Enzymes. Trends Glycosci. Glycotechnol. 2002, 14, 35−50. (374) Nakai, H.; Kitaoka, M.; Svensson, B.; Ohtsubo, K. Recent Development of Phosphorylases Possessing Large Potential for Oligosaccharide Synthesis. Curr. Opin. Chem. Biol. 2013, 17, 301−309. (375) Chaen, H.; Nishimoto, T.; Nakada, T.; Fukuda, S.; Kurimoto, M.; Tsujisaka, Y. Enzymatic Synthesis of Kojioligosaccharides Using Kojibiose Phosphorylase. J. Biosci. Bioeng. 2001, 92, 177−182. (376) Kitaoka, M.; Sasaki, T.; Taniguchi, H. Synthesis of Laminarioligosaccharides Using Crude Extract of Euglena-gracilis ZCells. Agric. Agric. Biol. Chem. 1991, 55, 1431−1432. (377) Nakajima, M.; Toyoizumi, H.; Abe, K.; Nakai, H.; Taguchi, H.; Kitaoka, M. 1,2-β-Oligoglucan Phosphorylase from Listeria innocua. PLoS One 2014, 9, e92353. (378) Ziegast, G.; Pfannemüller, B. Linear and Star-Shaped Hybrid Polymers. Phosphorolytic Syntheses with Di-Functional, Oligo-Func2396


Chemical Reviews

Review

tional and Multifunctional Primers. Carbohydr. Res. 1987, 160, 185− 204. (379) Yanase, M.; Takaha, T.; Kuriki, T. α-Glucan Phosphorylase and Its Use in Carbohydrate Engineering. J. Sci. Food Agric. 2006, 86, 1631−1635. (380) Ohdan, K.; Fujii, K.; Yanase, M.; Takaha, T.; Kuriki, T. Enzymatic Synthesis of Amylose. Biocatal. Biotransform. 2006, 24, 77− 81. (381) Boeck, B.; Schinzel, R. Purification and Characterisation of an α-Glucan Phosphorylase from the Thermophilic Bacterium Thermus thermophilus. Eur. J. Biochem. 1996, 239, 150−155. (382) Takaha, T.; Yanase, M.; Takata, H.; Okada, S. Structure and Properties of Thermus aquaticus α-Glucan Phosphorylase Expressed in Escherichichia coli. J. Appl. Glycosci. 2001, 48, 71−78. (383) Yanase, M.; Takata, H.; Fujii, K.; Takaha, T.; Kuriki, T. Cumulative Effect of Amino Acid Replacements Results in Enhanced Thermostability of Potato Type L α-Glucan Phosphorylase. Appl. Environ. Microbiol. 2005, 71, 5433−5439. (384) Kitamura, S. Starch Polymers, Natural and Synthetic. In The Polymeric Materials Encyclopedia, Synthesis, Properties and Applications; Salamone, C., Ed.; CRC Press: New York, 1996; Vol. 10, pp 7915− 7922. (385) Waldmann, H.; Gygax, D.; Bednarski, M. D.; Randall Shangraw, W.; Whitesides, G. M. The Enzymatic Utilization of Sucrose in the Synthesis of Amylose and Derivatives of Amylose, Using Phosphorylases. Carbohydr. Res. 1986, 157, C4−C7. (386) Fujii, K.; Iiboshi, M.; Yanase, M.; Takaha, T.; Kuriki, T. Enhancing the Thermal Stability of Sucrose Phosphorylase from Streptococcus mutans by Random Mutagenesis. J. Appl. Glycosci. 2006, 53, 91−97. (387) Ohdan, K.; Fujii, K.; Yanase, M.; Takaha, T.; Kuriki, T. Phosphorylase Coupling as a Tool to Convert Cellobiose into Amylose. J. Biotechnol. 2007, 127, 496−502. (388) Kadokawa, J. Facile Synthesis of Unnatural Oligosaccharides by Phosphorylase-Catalyzed Enzymatic Glycosylations Using New Glycosyl Donors. In Oligosaccharides: Sources, Properties and Applications; Gordon, N. S., Ed.; Nova Science Publishers, Inc.: Hauppauge, NY, 2011; pp 269−281. (389) Kadokawa, J. Synthesis of Non-natural Oligosaccharides by αGlucan Phosphorylase-Catalyzed Enzymatic Glycosylations Using Analogue Substrates of α-D-Glucose 1-Phosphate. Trends Glycosci. Glycotechnol. 2013, 25, 57−69. (390) Evers, B.; Thiem, J. Further Syntheses Employing Phosphorylase. Bioorg. Med. Chem. 1997, 5, 857−863. (391) Nawaji, M.; Izawa, H.; Kaneko, Y.; Kadokawa, J. Enzymatic Synthesis of α-D-xylosylated Maltooligosaccharides by PhosphorylaseCatalyzed Xylosylation. J. Carbohydr. Chem. 2008, 27, 214−222. (392) Nawaji, M.; Izawa, H.; Kaneko, Y.; Kadokawa, J. Enzymatic αGlucosaminylation of Maltooligosaccharides Catalyzed by Phosphorylase. Carbohydr. Res. 2008, 343, 2692−2696. (393) Kawazoe, S.; Izawa, H.; Nawaji, M.; Kaneko, Y.; Kadokawa, J. Phosphorylase-Catalyzed N-Formyl-α-glucosaminylation of Maltooligosaccharides. Carbohydr. Res. 2010, 345, 631−636. (394) Evers, B.; Mischnick, P.; Thiem, J. Synthesis of 2-Deoxy-α-Darabino-hexopyranosyl Phosphate and 2-Deoxy-maltooligosaccharides with Phosphorylase. Carbohydr. Res. 1994, 262, 335−341. (395) Percival, M. D.; Withers, S. G. Applications of Enzymes in the Synthesis and Hydrolytic Study of 2-Deoxy-α-D-glucopyranosyl Phosphate. Can. J. Chem. 1988, 66, 1970−1972. (396) Bhuiyan, S. H.; Rus’d, A. A.; Kitaoka, M.; Hayashi, K. Characterization of a Hyperthermostable Glycogen Phosphorylase from Aquifex aeolicus Expressed in Escherichia coli. J. Mol. Catal. B: Enzym. 2003, 22, 173−180. (397) Umegatani, Y.; Izawa, H.; Nawaji, M.; Yamamoto, K.; Kubo, A.; Yanase, M.; Takaha, T.; Kadokawa, J. Enzymatic α-Glucuronylation of Maltooligosaccharides Using α-Glucuronic Acid 1-phosphate as Glycosyl Donor Catalyzed by a Thermostable Phosphorylase from Aquifex aeolicus VF5. Carbohydr. Res. 2012, 350, 81−85.

(398) Takemoto, Y.; Izawa, H.; Umegatani, Y.; Yamamoto, K.; Kubo, A.; Yanase, M.; Takaha, T.; Kadokawa, J. Synthesis of Highly Branched Anionic α-Glucans by Thermostable Phosphorylase-Catalyzed αGlucuronylation. Carbohydr. Res. 2013, 366, 38−44. (399) Takata, H.; Takaha, T.; Okada, S.; Takagi, M.; Imanaka, T. Cyclization Reaction Catalyzed by Branching Enzyme. J. Bacteriol. 1996, 178, 1600−1606. (400) Takata, H.; Takaha, T.; Okada, S.; Hizukuri, S.; Takagi, M.; Imanaka, T. Structure of the Cyclic Glucan Produced from Amylopectin by Bacillus stearothermophilus Branching Enzyme. Carbohydr. Res. 1996, 295, 91−101. (401) Takata, H.; Takaha, T.; Nakamura, H.; Fujii, K.; Okada, S.; Takagi, M.; Imanaka, T. Production and Some Properties of a Dextrin with a Narrow Size Distribution by the Cyclization Reaction of Branching Enzyme. J. Ferment. Bioeng. 1997, 84, 119−123. (402) Takata, Y.; Shimohigoshi, R.; Yamamoto, K.; Kadokawa, J. Enzymatic Synthesis of Dendritic Amphoteric α-Glucans by Thermostable Phosphorylase Catalysis. Macromol. Biosci. 2014, 14, 1437− 1443. (403) Shimohigoshi, R.; Takemoto, Y.; Yamamoto, K.; Kadokawa, J. Thermostable α-Glucan Phosphorylase-Catalyzed Successive α-Mannosylations. Chem. Lett. 2013, 42, 822−824. (404) Borgerding, J. Phosphate Deposits in Digestion Systems. J.Water Pollut. Control Fed. 1972, 44, 813−819. (405) Kadokawa, J.; Shimohigoshi, R.; Yamashita, K.; Yamamoto, K. Synthesis of Chitin and Chitosan Stereoisomers by Thermostable αGlucan Phosphorylase-Catalyzed Enzymatic Polymerization of α-DGlucosamine 1-Phosphate. Org. Biomol. Chem. 2015, 13, 4336−4343. (406) Yamashita, K.; Yamamoto, K.; Kadokawa, J. Synthesis of NonNatural Heteroaminopolysaccharides by α-Glucan PhosphorylaseCatalyzed Enzymatic Copolymerization: α(1→4)-Linked Glucosaminoglucans. Biomacromolecules 2015, 16, 3989−3994. (407) Kitamura, S.; Yunokawa, H.; Mitsuie, S.; Kuge, T. Study on Polysaccharide by the Fluorescence Method II. Micro-Brownian Motion and Conformational Change of Amylose in Aqueous-Solution. Polym. J. 1982, 14, 93−99. (408) Kaneko, Y.; Kadokawa, J. Chemoenzymatic Synthesis of Amylose-Grafted Polymers. In Handbook of Carbohydrate Polymers: Development, Properties and Applications; Ito, R., Matsuo, Y., Eds.; Nova Science Publishers, Inc.: Hauppauge, NY, 2009; pp 671−691. (409) Izawa, H.; Kadokawa, J. Preparation of Functional Amylosic Materials by Phosphorylase-Catalyzed Polymerization. In Interfacial Researches in Fundamental and Material Sciences of Oligo- and Polysaccharides; Kadoakwa, J., Ed.; Transworld Research Network: Trivandrum, 2009; pp 69−86. (410) Kadokawa, J. Chemoenzymatic Synthesis of Functional Amylosic Materials. Pure Appl. Chem. 2014, 86, 701−709. (411) Kobayashi, K.; Kamiya, S.; Enomoto, N. Amylose-Carrying Styrene Macromonomer and its Homo- and Copolymers: Synthesis via Enzyme-Catalyzed Polymerization and Complex Formation with Iodine. Macromolecules 1996, 29, 8670−8676. (412) Narumi, A.; Kawasaki, K.; Kaga, H.; Satoh, T.; Sugimoto, N.; Kakuchi, T. Glycoconjugated Polymer 6. Synthesis of Poly[styreneblock-(styrene-graf t-amylose)] via Potato Phosphorylase-Catalyzed Polymerization. Polym. Bull. 2003, 49, 405−410. (413) von Braunmühl, V.; Jonas, G.; Stadler, R. Enzymatic Grafting of Amylose from Poly(dimethylsiloxanes). Macromolecules 1995, 28, 17−24. (414) Kadokawa, J.; Nakamura, Y.; Sasaki, Y.; Kaneko, Y.; Nishikawa, T. Chemoenzymatic Synthesis of Amylose-Grafted Polyacetylenes. Polym. Bull. 2008, 60, 57−68. (415) Sasaki, Y.; Kaneko, Y.; Kadokawa, J. Chemoenzymatic Synthesis of Amylose-Grafted Polyacetylene by Polymer Reaction Manner and its Conversion into Organogel with DMSO by CrossLinking. Polym. Bull. 2009, 62, 291−303. (416) Kaneko, Y.; Matsuda, S.; Kadokawa, J. Chemoenzymatic Synthesis of Amylose-Grafted Poly(vinyl alcohol). Polym. Chem. 2010, 1, 193−197. 2397


Chemical Reviews

Review

(417) Mazzocchetti, L.; Tsoufis, T.; Rudolf, P.; Loos, K. Enzymatic Synthesis of Amylose Brushes Revisited: Details from X-Ray Photoelectron Spectroscopy and Spectroscopic Ellipsometry. Macromol. Biosci. 2014, 14, 186−194. (418) Omagari, Y.; Kadokawa, J. Synthesis of Heteropolysaccharides Having Amylose Chains Using Phosphorylase-Catalyzed Enzymatic Polymerization. Kobunshi Ronbunshu 2011, 68, 242−249. (419) Kadoakwa, J. Synthesis of Amylose-Grafted Polysaccharide Materials by Phosphorylase-Catalyzed Enzymatic Polymerization. In Biobased Monomers, Polymers, and Materials; Smith, P. B., Gross, R. A., Eds.; ACS Symposium Series 1105; American Chemical Society: Washington, DC, 2012; pp 237−255. (420) Kadoakwa, J. Synthesis of New Polysaccharide Materials by Phosphorylase-Catalyzed Enzymatic α-Glycosylations Using Polymeric Glycosyl Acceptors. In Green Polymer Chemistry: Biocatalysis and Materials II; Cheng, H. N., Gross, R. A., Smith, P. B., Eds.; ACS Symposium Series 1144; American Chemical Society: Washington, DC, 2013; pp 141−161. (421) Matsuda, S.; Kaneko, Y.; Kadokawa, J. Chemoenzymatic Synthesis of Amylose-Grafted Chitosan. Macromol. Rapid Commun. 2007, 28, 863−867. (422) Kaneko, Y.; Matsuda, S.; Kadokawa, J. Chemoenzymatic Syntheses of Amylose-Grafted Chitin and Chitosan. Biomacromolecules 2007, 8, 3959−3964. (423) Omagari, Y.; Matsuda, S.; Kaneko, Y.; Kadokawa, J. Chemoenzymatic Synthesis of Amylose-Grafted Cellulose. Macromol. Biosci. 2009, 9, 450−455. (424) Omagari, Y.; Kaneko, Y.; Kadokawa, J. Chemoenzymatic Synthesis of Amylose-Grafted Alginate and its Formation of Enzymatic Disintegratable Beads. Carbohydr. Polym. 2010, 82, 394−400. (425) Arimura, T.; Omagari, Y.; Yamamoto, K.; Kadokawa, J. Chemoenzymatic Synthesis and Hydrogelation of Amylose-Grafted Xanthan Gums. Int. J. Biol. Macromol. 2011, 49, 498−503. (426) Kadokawa, J.; Arimura, T.; Takemoto, Y.; Yamamoto, K. SelfAssembly of Amylose-Grafted Carboxymethyl Cellulose. Carbohydr. Polym. 2012, 90, 1371−1377. (427) Hatanaka, D.; Takemoto, Y.; Yamamoto, K.; Kadokawa, J. Hierarchically Self-Assembled Nanofiber Films from Amylose-Grafted Carboxymethyl Cellulose. Fibers 2014, 2, 34−44. (428) Calder, P. C. Glycogen Structure and Biogenesis. Int. J. Biochem. 1991, 23, 1335−1352. (429) Manners, D. J. Recent Developments in Our Understanding of Glycogen Structure. Carbohydr. Polym. 1991, 16, 37−82. (430) Izawa, H.; Nawaji, M.; Kaneko, Y.; Kadokawa, J. Preparation of Glycogen-Based Polysaccharide Materials by Phosphorylase-Catalyzed Chain Elongation of Glycogen. Macromol. Biosci. 2009, 9, 1098−1104. (431) Takata, Y.; Yamamoto, K.; Kadokawa, J. Preparation of pHResponsive Amphoteric Glycogen Hydrogels by α-Glucan Phosphorylase-Catalyzed Successive Enzymatic Reactions. Macromol. Chem. Phys. 2015, 216, 1415−1420. (432) Kamiya, S.; Kobayashi, K. Synthesis and Helix Formation of Saccharide-Poly(L-glutamic acid) Conjugates. Macromol. Chem. Phys. 1998, 199, 1589−1596. (433) Sarko, A.; Zugenmaier, P. Crystal Structures of Amylose and Its Derivatives. In Fiber Diffraction Methods; French, A. D., Gardner, K. H., Eds.; ACS Symposium Series 141; American Chemical Society: Washington, DC, 1980; pp 459−482. (434) Shogren, R. L.; Greene, R. V.; Wu, Y. V. Complexes of Starch Polysaccharides and Poly(ethylene-co-acrylic acid) - Structure and Stability in Solution. J. Appl. Polym. Sci. 1991, 42, 1701−1709. (435) Shogren, R. L. Complexes of Starch with Telechelic Poly(εcaprolactone) Phosphate. Carbohydr. Polym. 1993, 22, 93−98. (436) Star, A.; Steuerman, D. W.; Heath, J. R.; Stoddart, J. F. Starched Carbon Nanotubes. Angew. Chem., Int. Ed. 2002, 41, 2508− 1512. (437) Ikeda, M.; Furusho, Y.; Okoshi, K.; Tanahara, S.; Maeda, K.; Nishino, S.; Mori, T.; Yashima, E. A Luminescent Poly(phenylenevinylene)-amylose Composite with Supramolecular Liquid Crystallinity. Angew. Chem., Int. Ed. 2006, 45, 6491−6495.

(438) Kida, T.; Minabe, T.; Okabe, S.; Akashi, M. PartiallyMethylated Amyloses as Effective Hosts for Inclusion Complex Formation with Polymeric Guests. Chem. Commun. 2007, 1559−1561. (439) Kaneko, Y.; Kyutoku, T.; Shimomura, N.; Kadokawa, J. Formation of Amylose-Poly(tetrahydrofuran) Inclusion Complexes in Ionic Liquid Media. Chem. Lett. 2011, 40, 31−33. (440) Rachmawati, R.; Woortman, A. J. J.; Loos, K. Facile Preparation Method for Inclusion Complexes between Amylose and Polytetrahydrofurans. Biomacromolecules 2013, 14, 575−583. (441) Kumar, K.; Woortman, A. J. J.; Loos, K. Synthesis of AmylosePolystyrene Inclusion Complexes by a Facile Preparation Route. Biomacromolecules 2013, 14, 1955−1960. (442) Rachmawati, R.; Woortman, A. J. J.; Loos, K. Tunable Properties of Inclusion Complexes Between Amylose and Polytetrahydrofuran. Macromol. Biosci. 2013, 13, 767−776. (443) Rachmawati, R.; Woortman, A. J. J.; Loos, K. SolventResponsive Behavior of Inclusion Complexes Between Amylose and Polytetrahydrofuran. Macromol. Biosci. 2014, 14, 56−68. (444) Kaneko, Y.; Kadokawa, J. Vine-twining Polymerization: A New Preparation Method for Well-Defined Supramolecules Composed of Amylose and Synthetic Polymers. Chem. Rec. 2005, 5, 36−46. (445) Kaneko, Y.; Kadokawa, J. Synthesis of Nanostructured BioRelated Materials by Hybridization of Synthetic Polymers with Polysaccharides or Saccharide Residues. J. Biomater. Sci., Polym. Ed. 2006, 17, 1269−1284. (446) Kaneko, Y.; Kadokawa, J. Preparation of Polymers with WellDefined Nanostructure in the Polymerization Field. In Modern Trends in Macromolecular Chemistry; Lee, J. N., Ed.; Nova Science Publishers, Inc.: Hauppauge, NY, 2009; pp 199−217. (447) Kaneko, Y.; Kadokawa, J. Preparation Method for Polysaccharide Supramolecules Using Amylose-Forming Polymerization Field: Vine-Twining Polymerization. Kobunshi Ronbunshu 2010, 67, 553−559. (448) Kadokawa, J. Preparation and Applications of Amylose Supramolecules by Means of Phosphorylase-Catalyzed Enzymatic Polymerization. Polymers 2012, 4, 116−133. (449) Kadokawa, J. Architecture of Amylose Supramolecules in Form of Inclusion Complexes by Phosphorylase-Catalyzed Enzymatic Polymerization. Biomolecules 2013, 3, 369−385. (450) Kadokawa, J.; Kaneko, Y.; Tagaya, H.; Chiba, K. Synthesis of an Amylose-Polymer Inclusion Complex by Enzymatic Polymerization of Glucose 1-Phosphate Catalyzed By Phosphorylase Enzyme in the Presence of PolyTHF: A New Method for Synthesis of Polymerpolymer Inclusion Complexes. Chem. Commun. 2001, 449−450. (451) Kadokawa, J.; Kaneko, Y.; Nagase, S.; Takahashi, T.; Tagaya, H. Vine-Twining Polymerization: Amylose Twines Around Polyethers to Form Amylose - Polyether Inclusion Complexes. Chem. - Eur. J. 2002, 8, 3321−3326. (452) Kadokawa, J.; Kaneko, Y.; Nakaya, A.; Tagaya, H. Formation of an Amylose-Polyester Inclusion Complex by Means of PhosphorylaseCatalyzed Enzymatic Polymerization of α-D-Glucose 1-Phosphate Monomer in the Presence of Poly(ε-caprolactone). Macromolecules 2001, 34, 6536−6538. (453) Kadokawa, J.; Nakaya, A.; Kaneko, Y.; Tagaya, H. Preparation of Inclusion Complexes Between Amylose and Ester-Containing Polymers by Means of Vine-Twining Polymerization. Macromol. Chem. Phys. 2003, 204, 1451−1457. (454) Nomura, S.; Kyutoku, T.; Shimomura, N.; Kaneko, Y.; Kadokawa, J. Preparation of Inclusion Complexes Composed of Amylose and Biodegradable Poly(glycolic acid-co-ε-caprolactone) by Vine-Twining Polymerization and their Lipase-Catalyzed Hydrolysis Behavior. Polym. J. 2011, 43, 971−977. (455) Kaneko, Y.; Beppu, K.; Kadokawa, J. Preparation of Amylose/ Polycarbonate Inclusion Complexes by Means of Vine-Twining Polymerization. Macromol. Chem. Phys. 2008, 209, 1037−1042. (456) Kaneko, Y.; Ueno, K.; Yui, T.; Nakahara, K.; Kadokawa, J. Amylose’s Recognition of Chirality in Polylactides on Formation of Inclusion Complexes in Vine-Twining Polymerization. Macromol. Biosci. 2011, 11, 1407−1415. 2398


Chemical Reviews

Review

Polymerization Using Primer−Guest Conjugates. Macromol. Chem. Phys. 2015, 216, 794−800. (476) Tanaka, T.; Gotanda, R.; Tsutsui, A.; Sasayama, S.; Yamamoto, K.; Kimura, Y.; Kadokawa, J. Synthesis and Gel Formation of Hyperbranched Supramolecular Polymer by Vine-Twining Polymerization Using Branched Primer-Guest Conjugate. Polymer 2015, 73, 9−16. (477) Morimoto, N.; Yamazaki, M.; Tamada, J.; Akiyoshi, K. Polysaccharide-Hair Cationic Polypeptide Nanogels: Self-assembly and Enzymatic Polymerization of Amylose Primer-Modified Cholesteryl Poly(L-lysine). Langmuir 2013, 29, 7509−7514. (478) Nishimura, T.; Mukai, S.; Sawada, S.; Akiyoshi, K. Glyco Star Polymers as Helical Multivalent Host and Biofunctional NanoPlatform. ACS Macro Lett. 2015, 4, 367−371. (479) Sheth, K.; Alexander, J. K. Purification and Properties of β-1,4Oligoglucan:Orthophosphate Glucosyltransferase from Clostridium thermocellum. J. Biol. Chem. 1969, 244, 457−464. (480) Samain, E.; Lancelonpin, C.; Ferigo, F.; Moreau, V.; Chanzy, H.; Heyraud, A.; Driguez, H. Phosphorolytic Synthesis of Cellodextrins. Carbohydr. Res. 1995, 271, 217−226. (481) Hiraishi, M.; Igarashi, K.; Kimura, S.; Wada, M.; Kitaoka, M.; Samejima, M. Synthesis of Highly Ordered Cellulose II In Vitro Using Cellodextrin Phosphorylase. Carbohydr. Res. 2009, 344, 2468−2473. (482) Moreau, V.; Viladot, J. L.; Samain, E.; Planas, A.; Driguez, H. Design and Chemoenzymatic Synthesis of Thiooligosaccharide Inhibitors of 1,3:1,4-β-D-Glucanases. Bioorg. Med. Chem. 1996, 4, 1849−1855. (483) Hrmova, M.; Fincher, G. B.; Viladot, J. L.; Planas, A.; Driguez, H. Chemoenzymic Synthesis of (1→3,1→4)-β-D-Glucooligosaccharides for Subsite Mapping of (1→3,1→4)-β-D-Glucan Endohydrolases. J. Chem. Soc., Perkin Trans. 1 1998, 3571−3576. (484) Choudhury, A. K.; Kitaoka, M.; Hayashi, K. Synthesis of a Cellobiosylated Dimer and Trimer and of Cellobiose-Coated Polyamidoamine (PAMAM) Dendrimers to Study Accessibility of an Enzyme, Cellodextrin Phosphorylase. Eur. J. Org. Chem. 2003, 2003, 2462−2470. (485) Remaud-Simeon, M.; Willemot, R. M.; Sarcabal, P.; Potocki de Montalk, G.; Monsan, P. Glucansucrases: Molecular Engineering and Oligosaccharide Synthesis. J. Mol. Catal. B: Enzym. 2000, 10, 117−128. (486) Remaud-Simeon, M.; Albenne, C.; Joucla, G.; Fabre, E.; Bozonnet, S.; Pizzut, S.; Escalier, P.; Potocki-Vèronèse, G.; Monsan, P. Glucansucrases: Structural Basis, Mechanistic Aspects, and New Perspectives for Engineering. In Oligosaccharides in Food and Agriculture; Eggleston, G., Côté, G. L., Eds.; ACS Symposium Series 849; American Chemical Society: Washington, DC, 2003; pp 90−103. (487) Korakli, M.; Vogel, R. F. Structure/Function Relationship of Homopolysaccharide Producing Glycansucrases and Therapeutic Potential of their Synthesised Glycans. Appl. Microbiol. Biotechnol. 2006, 71, 790−803. (488) van Hijum, S. A. F. T.; Kralj, S.; Ozimek, L. K.; Dijkhuizen, L.; van Geel-Schutten, I. G. H. Structure-Function Relationships of Glucansucrase and Fructansucrase Enzymes from Lactic Acid Bacteria. Microbiol. Mol. Biol. Rew. 2006, 70, 157−176. (489) Albenne, C.; Skov, L. K.; Mirza, O.; Gajhede, M.; Feller, G.; D’Amico, S.; Andre, G.; Potocki-Véronèse, G.; van der Veen, B. A.; Monsan, P.; Remaud-Simeon, M. Molecular Basis of the Amylose-Like Polymer Formation Catalyzed by Neisseria polysaccharea Amylosucrase. J. Biol. Chem. 2004, 279, 726−734. (490) van der Veen, B. A.; Potocki-Véronèse, G.; Albenne, C.; Joucla, G.; Monsan, P.; Remaud-Simeon, M. Combinatorial Engineering to Enhance Amylosucrase Performance: Construction, Selection, and Screening of Variant Libraries for Increased Activity. FEBS Lett. 2004, 560, 91−97. (491) van Leeuwen, S. S.; Kralj, S.; Eeuwema, W.; Gerwig, G. J.; Dijkhuizen, L.; Kamerling, J. P. Structural Characterization of Bioengineered α-D-Glucans Produced by Mutant Glucansucrase GTF180 Enzymes of Lactobacillus reuteri Strain 180. Biomacromolecules 2009, 10, 580−588.

(457) Kaneko, Y.; Beppu, K.; Kadokawa, J. Amylose Selectively Includes one from a Mixture of Two Resemblant Polyethers in VineTwining Polymerization. Biomacromolecules 2007, 8, 2983−2985. (458) Kaneko, Y.; Beppu, K.; Kyutoku, T.; Kadokawa, J. Selectivity and Priority on Inclusion of Amylose toward Guest Polyethers and Polyesters in Vine-Twining Polymerization. Polym. J. 2009, 41, 279− 286. (459) Kaneko, Y.; Beppu, K.; Kadokawa, J. Amylose Selectively Includes a Specific Range of Molecular Weights in Poly(tetrahydrofuran)s in Vine-Twining Polymerization. Polym. J. 2009, 41, 792−796. (460) Kobayashi, S.; Uyama, H.; Suda, S.; Namekawa, S. Dehydration Polymerization in Aqueous Medium Catalyzed by Lipase. Chem. Lett. 1997, 105−105. (461) Suda, S.; Uyama, H.; Kobayashi, S. Dehydration Polycondensation in Water for Synthesis of Polyesters by Lipase Catalyst. Proc. Jpn. Acad., Ser. B 1999, 75, 201−206. (462) Kaneko, Y.; Saito, Y.; Nakaya, A.; Kadokawa, J.; Tagaya, H. Preparation of Inclusion Complexes Composed of Amylose and Strongly Hydrophobic Polyesters in Parallel Enzymatic Polymerization System. Macromolecules 2008, 41, 5665−5670. (463) Kaneko, Y.; Fujisaki, K.; Kyutoku, T.; Furukawa, H.; Kadokawa, J. Preparation of Enzymatically Recyclable Hydrogels Through the Formation of Inclusion Complexes of Amylose in a VineTwining Polymerization. Chem. - Asian J. 2010, 5, 1627−1633. (464) Kadokawa, J.; Nomura, S.; Hatanaka, D.; Yamamoto, K. Preparation of Polysaccharide Supramolecular Films by Vine-Twining Polymerization Approach. Carbohydr. Polym. 2013, 98, 611−617. (465) Kadokawa, J.; Tanaka, K.; Hatanaka, D.; Yamamoto, K. Preparation of Multiformable Supramolecular Gels Through Helical Complexation by Amylose in Vine-Twining Polymerization. Polym. Chem. 2015, 6, 6402−6408. (466) Loos, K.; Stadler, R. Synthesis of Amylose-block-polystyrene Rod-Coil Block Copolymers. Macromolecules 1997, 30, 7641−7643. (467) Loos, K.; Müller, A. H. E. New Routes to the Synthesis of Amylose-block-polystyrene Rod-Coil Block Copolymers. Biomacromolecules 2002, 3, 368−373. (468) Loos, K.; Boker, A.; Zettl, H.; Zhang, A. F.; Krausch, G.; Müller, A. H. E. Micellar Aggregates of Amylose-block-polystyrene Rod-Coil Block Copolymers in Water and THF. Macromolecules 2005, 38, 873−879. (469) Akiyoshi, K.; Kohara, M.; Ito, K.; Kitamura, S.; Sunamoto, J. Enzymatic Synthesis and Characterization of Amphiphilic Block Copolymers of Poly(ethylene oxide) and Amylose. Macromol. Rapid Commun. 1999, 20, 112−115. (470) Akiyoshi, K.; Maruichi, N.; Kohara, M.; Kitamura, S. Amphiphilic Block Copolymer with a Molecular Recognition Site: Induction of a Novel Binding Characteristic of Amylose by SelfAssembly of Poly(ethylene oxide)-block-amylose In Chloroform. Biomacromolecules 2002, 3, 280−283. (471) Morimoto, N.; Ogino, N.; Narita, T.; Kitamura, S.; Akiyoshi, K. Enzyme-Responsive Molecular Assembly System with Amylose-Primer Surfactants. J. Am. Chem. Soc. 2007, 129, 458−459. (472) Morimoto, N.; Ogino, N.; Narita, T.; Akiyoshi, K. EnzymeResponsive Artificial Chaperone System with Amphiphilic Amylose Primer. J. Biotechnol. 2009, 140, 246−249. (473) Tanaka, T.; Sasayama, S.; Nomura, S.; Yamamoto, K.; Kimura, Y.; Kadokawa, J. An Amylose-Poly(L-lactide) Inclusion Supramolecular Polymer: Enzymatic Synthesis by Means of Vine-Twining Polymerization Using a Primer-Guest Conjugate. Macromol. Chem. Phys. 2013, 214, 2829−2834. (474) Tanaka, T.; Tsutsui, A.; Gotanda, R.; Sasayama, S.; Yamamoto, K.; Kadokawa, J. Synthesis of amylose-polyether inclusion supramolecular polymers by vine-twining polymerization using maltoheptaose-functionalized poly(tetrahydrofuran) as a primer-guest conjugate. J. Appl. Glycosci. 2015, 62, 135−141. (475) Tanaka, T.; Sasayama, S.; Yamamoto, K.; Kimura, Y.; Kadokawa, J. Evaluating Relative Chain Orientation of Amylose and Poly(L-lactide) in Inclusion Complexes Formed by Vine-Twining 2399


Chemical Reviews

Review

(492) Potocki de Montalk, G.; Remaud-Simeon, M.; Willemot, R. M.; Sarçabal, P.; Planchot, V.; Monsan, P. Amylosucrase from Neisseria polysaccharea: Novel Catalytic Properties. FEBS Lett. 2000, 471, 219− 223. (493) Cheetham, N. W. H.; Walker, G. J.; Pearce, B. J.; Fiala-Beer, E.; Taylor, C. Structures of Water-Soluble α-D-Glucans Synthesized from Sucrose by Glucosyltransferases Isolated from Streptococcus sobrinus Culture Filtrates. Carbohydr. Polym. 1990, 14, 3−16. (494) Potocki de Montalk, G.; Remaud-Simeon, M.; Willemot, R. M.; Monsan, P. Characterisation of the Activator Effect of Glycogen on Amylosucrase from Neisseria polysaccharea. FEMS Microbiol. Lett. 2000, 186, 103−108. (495) Rolland-Sabaté, A.; Colonna, P.; Potocki-Véronèse, G.; Monsan, P.; Planchot, V. Elongation and Insolubilisation of α-Glucans by the Action of Neisseria polysaccharea Amylosucrase. J. Cereal Sci. 2004, 40, 17−30. (496) Potocki-Véronèse, G.; Putaux, J. L.; Dupeyre, D.; Albenne, C.; Remaud-Simeon, M.; Monsan, P.; Buleon, A. Amylose Synthesized In Vitro by Amylosucrase: Morphology, Structure, and Properties. Biomacromolecules 2005, 6, 1000−1011. (497) Putaux, J. L.; Potocki-Véronèse, G.; Remaud-Simeon, M.; Buleon, A. α-D-Glucan-Based Dendritic Nanoparticles Prepared by In Vitro Enzymatic Chain Extension of Glycogen. Biomacromolecules 2006, 7, 1720−1728. (498) Lim, M. C.; Seo, D. H.; Jung, J. H.; Park, C. S.; Kim, Y. R. Enzymatic Synthesis of Amylose Nanocomposite Microbeads Using Amylosucrase from Deinococcus geothermalis. RSC Adv. 2014, 4, 26421−26424. (499) Lim, M. C.; Lee, G. H.; Ngoc Huynh, D. T.; Morales Letona, C. A.; Seo, D. H.; Park, C. S.; Kim, Y. R. Amylosucrase-Mediated Synthesis and Self-Assembly of Amylose Magnetic Microparticles. RSC Adv. 2015, 5, 36088−36091. (500) Semyonov, D.; Ramon, O.; Shoham, Y.; Shimoni, E. Enzymatically Synthesized Dextran Nanoparticles and Their Use as Carriers for Nutraceuticals. Food Funct. 2014, 5, 2463−2474. (501) Baciu, I. E.; Jördening, H. J.; Seibel, J.; Buchholz, K. Investigations of the Transfructosylation Reaction by Fructosyltransferase from B. subtilis NCIMB 11871 for the Synthesis of the Sucrose Analogue Galactosyl-Fructoside. J. Biotechnol. 2005, 116, 347−357. (502) Seibel, J.; Moraru, R.; Götze, S. Biocatalytic and Chemical Investigations in the Synthesis of Sucrose Analogues. Tetrahedron 2005, 61, 7081−7086. (503) Seibel, J.; Moraru, R.; Götze, S.; Buchholz, K.; Na’amnieh, S.; Pawlowski, A.; Hecht, H. J. Synthesis of Sucrose Analogues and the Mechanism of Action of Bacillus subtilis Fructosyltransferase (Levansucrase). Carbohydr. Res. 2006, 341, 2335−2349. (504) Seibel, J.; Hellmuth, H.; Hofer, B.; Kicinska, A. M.; Schmalbruch, B. Identification of New Acceptor Specificities of Glycosyltransferase R with the Aid of Substrate Microarrays. ChemBioChem 2006, 7, 310−320. (505) Beine, R.; Moraru, R.; Nimtz, M.; Na’amnieh, S.; Pawlowski, A.; Buchholz, K.; Seibel, J. Synthesis of Novel Fructooligosaccharides by Substrate and Enzyme Engineering. J. Biotechnol. 2008, 138, 33−41. (506) Homann, A.; Seibel, J. Towards Tailor-made Oligosaccharides - Chemo-Enzymatic Approaches by Enzyme and Substrate Engineering. Appl. Microbiol. Biotechnol. 2009, 83, 209−216. (507) Boyer, C.; Preiss, J. Biosynthesis of Bacterial Glycogen. Purification and Properties of the Escherichia coli B α−1,4-Glucan: α1,4-Glucan 6-Glycosyl transferase. Biochemistry 1977, 16, 3693−3699. (508) Takata, H.; Takaha, T.; Kuriki, T.; Okada, S.; Takagi, M.; Imanaka, T. Properties and Active-Center of the Thermostable Branching Enzyme from Bacillus Stearothermophilus. Appl. Environ. Microbiol. 1994, 60, 3096−3104. (509) Lerner, L. R.; Krisman, C. R. Differential Action of Branching Enzymes According to Its Source. Cell. Mol. Biol. 1996, 42, 599−608. (510) Palomo, M.; Kralj, S.; van der Maarel, M. J. E. C.; Dijkhuizen, L. The Unique Branching Patterns of Deinococcus Glycogen Branching Enzymes Are Determined by Their N-Terminal Domains. Appl. Environ. Microbiol. 2009, 75, 1355−1362.

(511) van der Vlist, J.; Palomo Reixach, M.; van der Maarel, M.; Dijkhuizen, L.; Schouten, A. J.; Loos, K. Synthesis of Branched Polyglucans by the Tandem Action of Potato Phosphorylase and Deinococcus geothermalis Glycogen Branching Enzyme. Macromol. Rapid Commun. 2008, 29, 1293−1297. (512) Kakutani, R.; Adachi, Y.; Kajiura, H.; Takata, H.; Ohno, N.; Kuriki, T. Stimulation of Macrophage by Enzymatically Synthesized Glycogen: The Relationship Between Structure and Biological Activity. Biocatal. Biotransform. 2008, 26, 152−160. (513) Kajiura, H.; Kakutani, R.; Akiyama, T.; Takata, H.; Kuriki, T. A Novel Enzymatic Process for Glycogen Production. Biocatal. Biotransform. 2008, 26, 133−140. (514) Takata, H.; Kajiura, H.; Furuyashiki, T.; Kakutani, R.; Kuriki, T. Fine Structural Properties of Natural and Synthetic Glycogens. Carbohydr. Res. 2009, 344, 654−659. (515) Takata, H.; Akiyama, T.; Kajiura, H.; Kakutani, R.; Furuyashiki, T.; Tomioka, E.; Kojima, I.; Kuriki, T. Application of Branching Enzyme in Starch Processing. Biocatal. Biotransform. 2010, 28, 60−63. (516) Kajiura, H.; Takata, H.; Akiyama, T.; Kakutani, R.; Furuyashiki, T.; Kojima, I.; Harui, T.; Kuriki, T. In vitro Synthesis of Glycogen: The Structure, Properties, and Physiological function of EnzymaticallySynthesized Glycogen. Biologia 2011, 66, 387−394. (517) Hernández, J. M.; Gaborieau, M.; Castignolles, P.; Gidley, M. J.; Myers, A. M.; Gilbert, R. G. Mechanistic Investigation of a StarchBranching Enzyme Using HydrodynamicVvolume SEC Analysis. Biomacromolecules 2008, 9, 954−965. (518) Anderson, A. J.; Dawes, E. A. Occurrence, Metabolism, Metabolic Role, and Industrial Uses of Bacterial Polyhydroxyalkanoates. Microbiol. Rev. 1990, 54, 450−472. (519) Lee, S. Y. Bacterial Polyhydroxyalkanoates. Biotechnol. Bioeng. 1996, 49, 1−14. (520) Rehm, B. H. A.; Steinbuchel, A. Biochemical and Genetic Analysis of PHA Synthases and Other Proteins Required for PHA Synthesis. Int. J. Biol. Macromol. 1999, 25, 3−19. (521) Rehm, B. H. A. Polyester Synthases: Natural Catalysts for Plastics. Biochem. J. 2003, 376, 15−33. (522) Rehm, B. H. A. Genetics and Biochemistry of Polyhydroxyalkanoate Granule Self-assembly: The Key Role of Polyester Synthases. Biotechnol. Lett. 2006, 28, 207−213. (523) Gerngross, T. U.; Martin, D. P. Enzyme-Catalyzed Synthesis of Poly[(R)-(−)-3-Hydroxybutyrate] - Formation of Macroscopic Granules In-Vitro. Proc. Natl. Acad. Sci. U. S. A. 1995, 92, 6279−6283. (524) Lenz, R. W.; Farcet, C.; Dijkstra, P. J.; Goodwin, S.; Zhang, S. M. Extracellular Polymerization of 3-Hydroxyalkanoate Monomers with the Polymerase of Alcaligenes eutrophus. Int. J. Biol. Macromol. 1999, 25, 55−60. (525) Su, L.; Lenz, R. W.; Takagi, Y.; Zhang, S. M.; Goodwin, S.; Zhong, L. H.; Martin, D. P. Enzymatic Polymerization of (R)-3Hydroxyalkanoates by a Bacterial Polymerase. Macromolecules 2000, 33, 229−231. (526) Song, J. J.; Zhang, S. M.; Lenz, R. W.; Goodwin, S. In Vitro Polymerization and Copolymerization of 3-Hydroxypropionyl-CoA with the PHB Synthase from Ralstonia eutropha. Biomacromolecules 2000, 1, 433−439. (527) Zhang, S. M.; Yasuo, T.; Lenz, R. W.; Goodwin, S. Kinetic and Mechanistic Characterization of the Polyhydroxybutyrate Synthase from Ralstonia eutropha. Biomacromolecules 2000, 1, 244−251. (528) Kamachi, M.; Zhang, S. M.; Goodwin, S.; Lenz, R. W. Enzymatic Polymerization and Characterization of New Poly(3hydroxyalkanoate)s by a Bacterial Polymerase. Macromolecules 2001, 34, 6889−6894. (529) Zhang, S. M.; Kolvek, S.; Lenz, R. W.; Goodwin, S. Mechanism of the Polymerization Reaction Initiated and Catalyzed by the Polyhydroxybutyrate Synthase of Ralstonia eutropha. Biomacromolecules 2003, 4, 504−509. (530) Nomura, C. T.; Taguchi, S. PHA Synthase Engineering Toward Superbiocatalysts for Custom-made Biopolymers. Appl. Microbiol. Biotechnol. 2006, 73, 969−979. 2400


Chemical Reviews

Review

ties of Fish Gelatin−Nanoclay Composite Film. Food Chem. 2009, 114, 180−189. (550) Yin, W.; Su, R.; Qi, W.; He, Z. A Casein-Polysaccharide Hybrid Hydrogel Cross-Linked by Transglutaminase for Drug Delivery. J. Mater. Sci. 2012, 47, 2045−2055. (551) Zhang, Y.-N.; Liu, N.; Zhao, X.-H. A Study on the Preparation and Some Functional Properties of a Cross-Linked Casein-Gelatin Composite by a Microbial Transglutaminase. Int. J. Food Sci. Technol. 2011, 46, 2641−2647. (552) De Colli, M.; Massimi, M.; Barbetta, A.; Di Rosario, B. L.; Nardecchia, S.; Conti Devirgiliis, L.; Dentini, M. A Biomimetic Porous Hydrogel of Gelatin and Glycosaminoglycans Cross-linked with Transglutaminase and Its Application in the Culture of Hepatocytes. Biomed. Mater. 2012, 7, 055005. (553) Niger, C.; Beazley, K. E.; Nurminskaya, M. Induction of Chondrogenic Differentiation in Mesenchymal Stem Cells by TGF-β Cross-Linked to Collagen-PLLA [Poly(L-lactic acid)] Scaffold by Transglutaminase 2. Biotechnol. Lett. 2013, 35, 2193−2199. (554) Song, C.-L.; Zhao, X.-H. The Preparation of an OligochitosanGlycosylated and Cross-linked Caseinate Obtained by a Microbial Ttransglutaminase and Its Functional Properties. Int. J. Dairy Technol. 2014, 67, 110−116. (555) Cortez, J.; Bonner, P. L. R.; Griffin, M. Application of Transglutaminases in the Modification of Wool Textiles. Enzyme Microb. Technol. 2004, 34, 64−72. (556) Cortez, J.; Bonner, P. L. R.; Griffin, M. Transglutaminase Treatment of Wool Fabrics Leads to Resistance to Detergent Damage. J. Biotechnol. 2005, 116, 379−386. (557) Cardamone, J. M. Enzyme-Mediated Crosslinking of Wool. Part 1: Transglutaminase. Text. Res. J. 2007, 77, 214−221. (558) Cortez, J.; Anghieri, A.; Bonner, P. L. R.; Griffin, M.; Freddi, G. Transglutaminase Mediated Grafting of Silk Proteins onto Wool Fabrics Leading to Improved Physical and Mechanical Properties. Enzyme Microb. Technol. 2007, 40, 1698−1704. (559) Falkowski, P.; Scholes, R. J.; Boyle, E.; Canadell, J.; Canfield, D.; Elser, J.; Gruber, N.; Hibbard, K.; Hogberg, P.; Seitzinger, S.; et al. The Global Carbon Cycle: A Test of Our Knowledge of Earth as a System. Science 2000, 290, 291−296. (560) Zechel, D. L.; Withers, S. G. Glycosidase Mechanisms: Anatomy of a Finely Tuned Catalyst. Acc. Chem. Res. 2000, 33, 11−18. (561) Shoda, S. Enzymatic Glycosylation. In Glycoscience Chemistry and Chemical Biology; Fraser-Reid, B. O., Tatsuta, K., Thiem, J., Eds.; Springer-Verlag: Berlin, Heidelberg, NY, 2001; Vol. II, pp 1465−1496. (562) Ajisaka, K.; Fujimoto, H. Regioselective Synthesis of Trisaccharides by Use of a Reversed Hydrolysis Activity of α-DGalactosidase and β-D-Galactosidase. Carbohydr. Res. 1989, 185, 139− 146. (563) Sinnott, M. L. Catalytic Mechanisms of Enzymatic Glycosyl Transfer. Chem. Rev. 1990, 90, 1171−1202. (564) Bornscheuer, U. T.; Kazlauskas, R. J. Hydrolases in Organic Synthesis: Regio- and Stereoselective Biotransformations; Wiley-VCH Verlag GmbH & Co. KGaA, 2006. (565) Barrett, A. J.; Rawlings, N. D.; Fred Woessner, J. Handbook of Proteolytic Enzymes; 3rd ed.; Academic Press, 2012. (566) Grisham, C. M.; Garrett, R. H. Mechanism of Enzyme Action. In Biochemistry; Thomson Brooks/Cole: Australia, 2007; pp 467−469. (567) Kerr, A. K. CRC Handbook of Chemistry and Physics; CRC: Boca Raton, FL, 1995. (568) Barnett, J. E. G.; Ralph, A.; Munday, K. A. Structural Requirements for Active Intestinal Transport. The Nature of the Carrier−Sugar Bonding at C-2 and the Ring Oxygen of the Sugar. Biochem. J. 1970, 118, 843−850. (569) Barnett, J. E. G.; Jarvis, W. T. S.; Munday, K. A. The Hydrolysis of Glycosyl Fluorides by Glycosidases. Biochem. J. 1967, 105, 669−672. (570) Hehre, E. J.; Brewer, C. F.; Genghof, D. S. Scope and Mechanism of Carbohydrase Action - Hydrolytic and Non-Hydrolytic Actions of β-Amylase on α-Maltosyl and β-Maltosyl Fluoride. J. Biol. Chem. 1979, 254, 5942−5950.

(531) Park, S. J.; Kim, T. W.; Kim, M. K.; Lee, S. Y.; Lim, S. C. Advanced Bacterial Polyhydroxyalkanoates: Towards a Versatile and Sustainable Platform for Unnatural Tailor-made Polyesters. Biotechnol. Adv. 2012, 30, 1196−1206. (532) Rey, D. A.; Strickland, A. D.; Kirui, D.; Niamsiri, N.; Batt, C. A. In vitro Self-assembly of Gold Nanoparticle-Coated Poly(3-hydroxybutyrate) Granules Exhibiting Plasmon-Induced Thermo-optical Enhancements. ACS Appl. Mater. Interfaces 2010, 2, 1804−1810. (533) Auras, R.; Harte, B.; Selke, S. An Overview of Polylactides as Packaging Materials. Macromol. Biosci. 2004, 4, 835−864. (534) Yuan, W.; Jia, Y.; Tian, J. M.; Snell, K. D.; Muh, U.; Sinskey, A. J.; Lambalot, R. H.; Walsh, C. T.; Stubbe, J. Class I and III Polyhydroxyalkanoate Synthases from Ralstonia eutropha and Allochromatium vinosum: Characterization and Substrate Specificity Studies. Arch. Biochem. Biophys. 2001, 394, 87−98. (535) Zhang, S.; Kamachi, M.; Takagi, Y.; Lenz, R. W.; Goodwin, S. Comparative Study of the Relationship Between Monomer Structure and Reactivity for Two Polyhydroxyalkanoate Synthases. Appl. Microbiol. Biotechnol. 2001, 56, 131−136. (536) Nduko, J. M.; Matsumoto, K.; Taguchi, S. Biological LactatePolymers Synthesized by One-Pot Microbial Factory: Enzyme and Metabolic Engineering. In Biobased Monomers, Polymers, and Materials; Smith, P. B., Gross, R. A., Eds.; ACS Symposium Series 1105; American Chemical Society: Washington, DC, 2012; pp 213−235. (537) Nduko, J. M.; Matsumoto, K.; Taguchi, S. Microbial Plastic Factory: Synthesis and Properties of the New Lactate-Based Biopolymers. In Green Polymer Chemistry: Biocatalysis and Materials II; Cheng, H. N., Gross, R. A., Smith, P. B., Eds.; ACS Symposium Series 1144; American Chemical Society: Washington, DC, 2013; pp 175−197. (538) Matsumoto, K.; Taguchi, S. Biosynthetic Polyesters Consisting of 2-Hydroxyalkanoic Acids: Current Challenges and Unresolved Questions. Appl. Microbiol. Biotechnol. 2013, 97, 8011−8021. (539) Tajima, K.; Han, X.; Satoh, Y.; Ishii, A.; Araki, Y.; Munekata, M.; Taguchi, S. In Vitro Synthesis of Polyhydroxyalkanoate (PHA) Incorporating Lactate (LA) with a Block Sequence by Using a Newly Engineered Thermostable PHA Synthase from Pseudomonas sp. SG4502 with Acquired LA-Polymerizing Activity. Appl. Microbiol. Biotechnol. 2012, 94, 365−376. (540) Hu, B. H.; Messersmith, P. B. Rational Design of Transglutaminase Substrate Peptides for Rapid Enzymatic Formation of Hydrogels. J. Am. Chem. Soc. 2003, 125, 14298−14299. (541) Folk, J. E. Mechanism of Action of Guinea Pig Liver Transglutaminase. J. Biol. Chem. 1969, 244, 3707−3713. (542) Zhu, Y.; Tramper, J. Novel Applications for Microbial Transglutaminase Beyond Food Processing. Trends Biotechnol. 2008, 26, 559−565. (543) Ohtsuka, T.; Sawa, A.; Kawabata, R.; Nio, N.; Motoki, M. Substrate Specificities of Microbial Transglutaminase for Primary Amines. J. Agric. Food Chem. 2000, 48, 6230−6233. (544) Crescenzi, V.; Francescangeli, A.; Taglienti, A. New GelatinBased Hydrogels via Enzymatic Networking. Biomacromolecules 2002, 3, 1384−1391. (545) McDermott, M. K.; Chen, T. H.; Williams, C. M.; Markley, K. M.; Payne, G. F. Mechanical Properties of Biomimetic Tissue Adhesive Based on the Microbial Transglutaminase-Catalyzed Crosslinking of Gelatin. Biomacromolecules 2004, 5, 1270−1279. (546) Chau, D. Y. S.; Collighan, R. J.; Verderio, E. A. M.; Addy, V. L.; Griffin, M. The Cellular Response to Transglutaminase-Cross-Linked Collagen. Biomaterials 2005, 26, 6518−6529. (547) O Halloran, D. M.; Collighan, R. J.; Griffin, M.; Pandit, A. S. Characterization of a Microbial Transglutaminase Cross-linked Type II Collagen Scaffold. Tissue Eng. 2006, 12, 1467−1474. (548) Kim, Y. J.; Uyama, H. Biocompatible Hydrogel Formation of Gelatin from Cold Water Fish via Enzymatic Networking. Polym. J. 2007, 39, 1040−1046. (549) Bae, H. J.; Darby, D. O.; Kimmel, R. M.; Park, H. J.; Whiteside, W. S. Effects of Transglutaminase-Induced Cross-Linking on Proper2401


Chemical Reviews

Review

(571) Shoda, S.; Shintate, K.; Ishihara, M.; Noguchi, M.; Kobayashi, A. Colorimetric Assay for Evaluating Glycosyl Fluoride-hydrolyzing Activity of Glycosidase by Using Alizarin Complexon Reagent. Chem. Lett. 2007, 36, 16−17. (572) Kobayashi, S. Challenge of Synthetic Cellulose. J. Polym. Sci., Part A: Polym. Chem. 2005, 43, 693−710. (573) Kobayashi, S.; Kashiwa, K.; Kawasaki, T.; Shoda, S. Novel Method for Polysaccharide Synthesis Using an Enzyme - The 1st In Vitro Synthesis of Cellulose via a Nonbiosyntehtic Path Utilizing Cellulase as Catalyst. J. Am. Chem. Soc. 1991, 113, 3079−3084. (574) Kobayashi, S.; Shoda, S. Chemical Synthesis of Cellulose and Cello-Oligomers Using a Hydrolysis Enzyme as a Catalyst. Int. J. Biol. Macromol. 1995, 17, 373−379. (575) Fort, S.; Boyer, V.; Greffe, L.; Davies, G.; Moroz, O.; Christiansen, L.; Schulein, M.; Cottaz, S.; Driguez, H. Highly Efficient Synthesis of β(1→4)-Oligo- and -Polysaccharides Using a Mutant Cellulase. J. Am. Chem. Soc. 2000, 122, 5429−5437. (576) Faijes, M.; Imai, T.; Bulone, V.; Planas, A. In Vitro Synthesis of a Crystalline (1→3,1→4)-β-D-Glucan by a Mutated (1→3,1→4)-β-DGlucanase from Bacillus. Biochem. J. 2004, 380, 635−641. (577) Saxena, I. M.; Brown, R. M.; Fevre, M.; Geremia, R. A.; Henrissat, B. Multidomain Architecture of β-Glycosyl Transferases Implications for Mechanism of Action. J. Bacteriol. 1995, 177, 1419− 1424. (578) Kobayashi, S.; Shoda, S.; Lee, J.; Okuda, K.; Brown, R. M.; Kuga, S. Direct Visualization of Synthetic Cellulose Formation via Enzymatic Polymerization Using Transmission Electron-Microscopy. Macromol. Chem. Phys. 1994, 195, 1319−1326. (579) Lee, J. H.; Brown, R. M.; Kuga, S.; Shoda, S.; Kobayashi, S. Assembly of Synthetic Cellulose-I. Proc. Natl. Acad. Sci. U. S. A. 1994, 91, 7425−7429. (580) Kobayashi, S.; Hobson, L. J.; Sakamoto, J.; Kimura, S.; Sugiyama, J.; Imai, T.; Itoh, T. Formation and Structure of Artificial Cellulose Spherulites via Enzymatic Polymerization. Biomacromolecules 2000, 1, 168−173. (581) Hashimoto, T.; Tanaka, H.; Koizumi, S.; Kurosaki, K.; Ohmae, M.; Kobayashi, S. Chemical Reaction at Specific Sites and ReactionInduced Self-Assembly as Observed by In Situ and Real Time SANS: Enzymatic Polymerization to Synthetic Cellulose. Biomacromolecules 2006, 7, 2479−2482. (582) Tanaka, H.; Koizumi, S.; Hashimoto, T.; Kurosaki, K.; Kobayashi, S. Self-Assembly of Synthetic Cellulose During In-Vitro Enzymatic Polymerization Process as Studied by a Combined SmallAngle Scattering Method. Macromolecules 2007, 40, 6304−6315. (583) Nakamura, I.; Yoneda, H.; Maeda, T.; Makino, A.; Ohmae, M.; Sugiyama, J.; Ueda, M.; Kobayashi, S.; Kimura, S. Enzymatic Polymerization Behavior Using Cellulose-Binding Domain Deficient Endoglucanase II. Macromol. Biosci. 2005, 5, 623−628. (584) Nakamura, I.; Makino, A.; Sugiyama, J.; Ohmae, M.; Kimura, S. Enzymatic Activities of Novel Mutant Endoglucanases Carrying Sequential Active Sites. Int. J. Biol. Macromol. 2008, 43, 226−231. (585) Nakamura, I.; Makino, A.; Horikawa, Y.; Sugiyama, J.; Ohmae, M.; Kimura, S. Preparation of Fibrous Cellulose by Enzymatic Polymerization Using Cross-Linked Mutant Endoglucanase II. Chem. Commun. 2011, 47, 10127−10129. (586) Nakamura, I.; Makino, A.; Ohmae, M.; Kimura, S. Immobilization of His-Tagged Endoglucanase on Gold via Various Ni-NTA Self-Assembled Monolayers and its Hydrolytic Activity. Macromol. Biosci. 2010, 10, 1265−1272. (587) Nakamura, I.; Horikawa, Y.; Makino, A.; Sugiyama, J.; Kimura, S. Enzymatic Polymerization Catalyzed by Immobilized Endoglucanase on Gold. Biomacromolecules 2011, 12, 785−790. (588) Nakamura, I.; Makino, A.; Ohmae, M.; Kimura, S. Enzymatic Polymerization to Cellulose by Crosslinked Enzyme Immobilized on Gold Solid Surface. Chem. Lett. 2012, 41, 37−38. (589) Mackenzie, L. F.; Wang, Q. P.; Warren, R. A. J.; Withers, S. G. Glycosynthases: Mutant Glycosidases for Oligosaccharide Synthesis. J. Am. Chem. Soc. 1998, 120, 5583−5584.

(590) Egusa, S.; Kitaoka, T.; Goto, M.; Wariishi, H. Synthesis of Cellulose in vitro by Using a Cellulase/Surfactant Complex in a Nonaqueous Medium. Angew. Chem., Int. Ed. 2007, 46, 2063−2065. (591) Ohmae, M.; Makino, A.; Kobayashi, S. Enzymatic Polymerization to Unnatural Hybrid Polysaccharides. Macromol. Chem. Phys. 2007, 208, 1447−1457. (592) Okamoto, E.; Kiyosada, T.; Shoda, S.; Kobayashi, S. Synthesis of Alternatingly 6-O-Methylated Cellulose via Enzymatic Polymerization of a Substituted Cellobiosyl Fluoride Monomer Catalyzed by Cellulase. Cellulose 1997, 4, 161−172. (593) Izumi, R.; Suzuki, Y.; Shimizu, Y.; Fujita, M.; Ishihara, M.; Noguchi, M.; Kobayashi, A.; Shoda, S. Synthesis of Artificial Oligosaccharides by Polycondensation of 2′-O-Methyl Cellobiosyl Fluoride and Mannosyl-Glucosyl Fluoride Catalyzed by Cellulase. In Interfacial Researches in Fundamental and Material Sciences of Oligo- and Polysaccharides; Kadokawa, J., Ed.; Transworld Research Network: Trivandrum, India, 2009; pp 45−67. (594) Kobayashi, S.; Makino, A.; Matsumoto, H.; Kunii, S.; Ohmae, M.; Kiyosada, T.; Makiguchi, K.; Matsumoto, A.; Horie, M.; Shoda, S. Enzymatic Polymerization to Novel Polysaccharides Having a Glucose-N-acetylglucosamine Repeating Unit, a Cellulose-Chitin Hybrid Polysaccharide. Biomacromolecules 2006, 7, 1644−1656. (595) Shoda, S.; Kawasaki, T.; Obata, K.; Kobayashi, S. A Facile Enzymatic-Synthesis of Cellooligosaccharide Derivatives Using βLactosyl Fluoride. Carbohydr. Res. 1993, 249, 127−137. (596) Moreau, V.; Driguez, H. Enzymic Synthesis of Hemithiocellodextrins. J. Chem. Soc., Perkin Trans. 1 1996, 525−527. (597) Saura-Valls, M.; Fauré, R.; Ragàs, S.; Piens, K.; Brumer, H.; Teeri, T. T.; Cottaz, S.; Driguez, H.; Planas, A. Kinetic Analysis Using Low-Molecular Mass Xyloglucan Oligosaccharides Defines the Catalytic Mechanism of a Populus Xyloglucan Endotransglycosylase. Biochem. J. 2006, 395, 99−106. (598) Tanaka, T.; Noguchi, M.; Ishihara, M.; Kobayashi, A.; Shoda, S. Synthesis of Non-natural Xyloglucans by Polycondensation of 4,6Dimethoxy-1,3,5-triazin-2-yl Oligosaccharide Monomers Catalyzed by Endo-β-1,4-glucanase. Macromol. Symp. 2010, 297, 200−209. (599) Kobayashi, S.; Wen, X.; Shoda, S. Specific Preparation of Artificial Xylan: A New Approach to Polysaccharide Synthesis by Using Cellulase as Catalyst. Macromolecules 1996, 29, 2698−2700. (600) Croon, I.; Timell, T. E. Distribution of Substitution in a Partially Methylated Xylan. J. Am. Chem. Soc. 1960, 82, 3416−3418. (601) Fujita, M.; Shoda, S.; Kobayashi, S. Xylanase-Catalyzed Synthesis of a Novel Polysaccharide Having a Glucose-xylose Repeating Unit, a Cellulose-Xylan Hybrid Polymer. J. Am. Chem. Soc. 1998, 120, 6411−6412. (602) Viladot, J. L.; Moreau, V.; Planas, A.; Driguez, H. Transglycosylation Activity of Bacillus 1,3−1,4-β-D-Glucan 4-Glucanohydrolases. Enzymic Synthesis of Alternate 1,3,-1,4-β-D-Glucooligosaccharides. J. Chem. Soc., Perkin Trans. 1 1997, 2383−2387. (603) Kobayashi, S.; Shimada, J.; Kashiwa, K.; Shoda, S. Enzymatic Polymerization of α-D-Maltosyl Fluoride Utilizing α-Amylase as the Catalyst - A New Approach for the Synthesis of Maltooligosaccharides. Macromolecules 1992, 25, 3237−3241. (604) Corey, E. J. Retrosynthetic Thinking - Essentials and Examples. Chem. Soc. Rev. 1988, 17, 111−133. (605) Tews, I.; Terwisscha van Scheltinga, A.; Perrakis, A.; Wilson, K. S.; Dijkstra, B. W. Substrate-Assisted Catalysis Unifies Two Families of Chitinolytic Enzymes. J. Am. Chem. Soc. 1997, 119, 7954−7959. (606) Merz, R. A.; Horsch, M.; Nyhlen, L. E.; Rast, D. M. Biochemistry of Chitin Synthase. In Chitin and Chitinase; Jolles, P., Muzzarelli, R. A. A., Eds.; Birkhauser Verlag: Basel, Switzerland, 1999; pp 9−37. (607) Kobayashi, S.; Kiyosada, T.; Shoda, S. Synthesis of Artificial Chitin: Irreversible Catalytic Behavior of a Glycosyl Hydrolase Through a Transition State Analogue Substrate. J. Am. Chem. Soc. 1996, 118, 13113−13114. (608) Kiyosada, T.; Takada, E.; Shoda, S.; Kobayashi, S. Hydrolysis and Polymerization of Novel Monomers Containing Amino Sugar. Polym. Prepr. Jpn. 1995, 44, 660. 2402


Chemical Reviews

Review

(609) Kiyosada, T.; Shoda, S.; Kobayashi, S. Synthesis of Artificial Chitin by Enzymatic Ring-Opening Polyaddition. Polym. Prepr. Jpn. 1995, 44, 1230−1231. (610) Sato, H.; Mizutani, S.; Tsuge, S.; Ohtani, H.; Aoi, K.; Takasu, A.; Okada, M.; Kobayashi, S.; Kiyosada, T.; Shoda, S. Determination of the Degree of Acetylation of Chitin/Chitosan by Pyrolysis Gas Chromatography in the Presence of Oxalic Acid. Anal. Chem. 1998, 70, 7−12. (611) Sakamoto, J.; Sugiyama, J.; Kimura, S.; Imai, T.; Itoh, T.; Watanabe, T.; Kobayashi, S. Artificial Chitin Spherulites Composed of Single Crystalline Ribbons of α-Chitin via Enzymatic Polymerization. Macromolecules 2000, 33, 4155−4160. (612) Noguchi, M.; Tanaka, T.; Gyakushi, H.; Kobayashi, A.; Shoda, S. Efficient Synthesis of Sugar Oxazolines from Unprotected N-Acetyl2-amino Sugars by Using Chloroformamidinium Reagent in Water. J. Org. Chem. 2009, 74, 2210−2212. (613) Noguchi, M.; Fujieda, T.; Huang, W. C.; Ishihara, M.; Kobayashi, A.; Shoda, S. A Practical One-Step Synthesis of 1,2Oxazoline Derivatives from Unprotected Sugars and Its Application to Chemoenzymatic β-N-Acetylglucosaminidation of Disialo-Oligosaccharide. Helv. Chim. Acta 2012, 95, 1928−1936. (614) Yoshida, N.; Tanaka, T.; Noguchi, M.; Kobayashi, A.; Ishikura, K.; Ikenuma, T.; Seno, H.; Watanabe, T.; Kohri, M.; Shoda, S. OnePot Chemoenzymatic Route to Chitoheptaose via Specific Transglycosylation of Chitopentaose-Oxazoline on Chitinase-Template. Chem. Lett. 2012, 41, 689−690. (615) Shoda, S.; Fujita, M.; Lohavisavapanichi, C.; Misawa, Y.; Ushizaki, K.; Tawata, Y.; Kuriyama, M.; Kohri, M.; Kuwata, H.; Watanabe, T. Efficient Method for the Elongation of the NAcetylglucosamine Unit by Combined Use of Chitinase and βGalactosidase. Helv. Chim. Acta 2002, 85, 3919−3936. (616) Kohri, M.; Kobayashi, A.; Noguchi, M.; Kawaida, S.; Watanabe, T.; Shoda, S. Stepwise Synthesis of Chitooligosaccharides Through a Transition-State Analogue Substrate Catalyzed by Mutants of Chitinase A1 from Bacillus circulans WL-12. Holzforschung 2006, 60, 485−491. (617) Kobayashi, S.; Makino, A.; Tachibana, N.; Ohmae, M. Chitinase-Catalyzed Synthesis of a Chitin-Xylan Hybrid Polymer: A Novel Water-Soluble β(1→4) Polysaccharide Having an N-Acetylglucosamine-Xylose Repeating Unit. Macromol. Rapid Commun. 2006, 27, 781−786. (618) Makino, A.; Kurosaki, K.; Ohmae, M.; Kobayashi, S. ChitinaseCatalyzed Synthesis of Alternatingly N-Deacetylated Chitin: A ChitinChitosan Hybrid Polysaccharide. Biomacromolecules 2006, 7, 950−957. (619) Sakamoto, J.; Kobayashi, S. Enzymatic Synthesis of 3-OMethylated Chitin Oligomers from New Derivatives of a Chitobiose Oxazoline. Chem. Lett. 2004, 33, 698−699. (620) Ochiai, H.; Ohmae, M.; Kobayashi, S. Enzymatic Glycosidation of Sugar Oxazolines Having a Carboxylate Group Catalyzed by Chitinase. Carbohydr. Res. 2004, 339, 2769−2788. (621) Ochiai, H.; Ohmae, M.; Kobayashi, S. Enzymatic Synthesis of Alternatingly 6-O-Carboxymethylated Chitotetraose by Selective Glycosidation with Chitinase Catalysis. Chem. Lett. 2004, 33, 694− 695. (622) Makino, A.; Ohmae, M.; Kobayashi, S. Synthesis of Fluorinated Chitin Derivatives via Enzymatic Polymerization. Macromol. Biosci. 2006, 6, 862−872. (623) Makino, A.; Sakamoto, J.; Ohmae, M.; Kobayashi, S. Effect of Fluorine Substituent on the Chitinase-Catalyzed Polymerization of Sugar Oxazoline Derivatives. Chem. Lett. 2006, 35, 160−161. (624) Makino, A.; Nagashima, H.; Ohmae, M.; Kobayashi, S. Chitinase-Catalyzed Synthesis of an Alternatingly N-Sulfonated Chitin Derivative. Biomacromolecules 2007, 8, 188−195. (625) Makino, A.; Ohmae, M.; Kobayashi, S. Chitinase-Catalyzed Copolymerization to a Chitin Derivative Having Glucosamine Unit in Controlled Proportion. Polym. J. 2006, 38, 1182−1188. (626) Stern, R.; Jedrzejas, M. J. Hyaluronidases: Their Genomics, Structures, and Mechanisms of Action. Chem. Rev. 2006, 106, 818− 839.

(627) El-Safory, N. S.; Fazary, A. E.; Lee, C. K. Hyaluronidases, a Group of Glycosidases: Current and Future Perspectives. Carbohydr. Polym. 2010, 81, 165−181. (628) Kobayashi, S.; Morii, H.; Itoh, R.; Kimura, S.; Ohmae, M. Enzymatic Polymerization to Artificial Hyaluronan: A Novel Method to Synthesize a Glycosaminoglycan Using a Transition State Analogue Monomer. J. Am. Chem. Soc. 2001, 123, 11825−11826. (629) Ochiai, H.; Ohmae, M.; Mori, T.; Kobayashi, S. HyaluronidaseCatalyzed Copolymerization for the Single-Step Synthesis of Functionalized Hyaluronan Derivatives. Biomacromolecules 2007, 8, 1327−1332. (630) Kobayashi, S.; Fujikawa, S.; Ohmae, M. Enzymatic Synthesis of Chondroitin and Its Derivatives Catalyzed by Hyaluronidase. J. Am. Chem. Soc. 2003, 125, 14357−14369. (631) Fujikawa, S.; Ohmae, M.; Kobayashi, S. Enzymatic Synthesis of Chondroitin 4-Sulfate with Well-Defined Structure. Biomacromolecules 2005, 6, 2935−2942. (632) Ochiai, H.; Fujikawa, S.; Ohmae, M.; Kobayashi, S. Enzymatic Copolymerization to Hybrid Glycosaminoglycans: A Novel Strategy for Intramolecular Hybridization of Polysaccharides. Biomacromolecules 2007, 8, 1802−1806. (633) Kobayashi, S.; Ohmae, M.; Ochiai, H.; Fujikawa, S. I. A Hyaluronidase Supercatalyst for the Enzymatic Polymerization to Synthesize Glycosaminoglycans. Chem.−Eur. J. 2006, 12, 5962−5971. (634) Yamagishi, K.; Suzuki, K.; Imai, K.; Mochizuki, H.; Morikawa, K.; Kyogashima, M.; Kimata, K.; Watanabe, H. Purification, Characterization, and Molecular Cloning of a Novel Keratan Sulfate Hydrolase, Endo-β-N-acetylglucosaminidase, from Bacillus circulans. J. Biol. Chem. 2003, 278, 25766−25772. (635) Kariya, Y.; Watabe, S.; Mochizuki, H.; Imai, K.; Kikuchi, H.; Suzuki, K.; Kyogashima, M.; Ishii, T. Modification of Di- and Tetrasaccharides from Shark Cartilage Keratan Sulphate by Refined Anhydromethanolic Hydrochloric Acid-Treatments and Evaluation of Their Specific Desulphation. Carbohydr. Res. 2003, 338, 1133−1138. (636) Ohmae, M.; Sakaguchi, K.; Kaneto, T.; Fujikawa, S.; Kobayashi, S. Keratanase II-Catalyzed Synthesis of Keratan Sulfate Oligomers by Using Sugar Oxazolines as Transition-State Analogue Substrate Monomers: A Novel Insight into the Enzymatic Catalysis Mechanism. ChemBioChem 2007, 8, 1710−1720. (637) Fujita, M.; Shoda, S.; Haneda, K.; Inazu, T.; Takegawa, K.; Yamamoto, K. A Novel Disaccharide Substrate Having 1,2-Oxazoline Moiety for Detection of Transglycosylating Activity of Endoglycosidases. Biochim. Biophys. Acta, Gen. Subj. 2001, 1528, 9−14. (638) Umekawa, M.; Higashiyama, T.; Koga, Y.; Tanaka, T.; Noguchi, M.; Kobayashi, A.; Shoda, S.; Huang, W.; Wang, L. X.; Ashida, H.; et al. Efficient Transfer of Sialo-Oligosaccharide onto Proteins by Combined Use of a Glycosynthase-Like Mutant of Mucor hiemalis Endoglycosidase and Synthetic Sialo-Complex-Type Sugar Oxazoline. Biochim. Biophys. Acta, Gen. Subj. 2010, 1800, 1203−1209. (639) Asahina, Y.; Kamitori, S.; Takao, T.; Nishi, N.; Hojo, H. Chemoenzymatic Synthesis of the Immunoglobulin Domain of Tim-3 Carrying a Complex-Type N-Glycan by Using a One-Pot Ligation. Angew. Chem., Int. Ed. 2013, 52, 9733−9737. (640) Tomabechi, Y.; Squire, M. A.; Fairbanks, A. J. Endo-β-Nacetylglucosaminidase Catalysed Glycosylation: Tolerance of Enzymes to Structural Variation of the Glycosyl Amino Acid Acceptor. Org. Biomol. Chem. 2014, 12, 942−955. (641) Goodfellow, J. J.; Baruah, K.; Yamamoto, K.; Bonomelli, C.; Krishna, B.; Harvey, D. J.; Crispin, M.; Scanlan, C. N.; Davis, B. G. An Endoglycosidase with Alternative Glycan Specificity Allows Broadened Glycoprotein Remodelling. J. Am. Chem. Soc. 2012, 134, 8030−8033. (642) Uyama, H.; Kobayashi, S. Enzymatic Ring-Opening Polymerization of Lactones Catalyzed by Lipase. Chem. Lett. 1993, 1149−1150. (643) Uyama, H.; Takeya, K.; Kobayashi, S. Synthesis of Polyesters by Enzymatic Ring-Opening Copolymerization Using Lipase Catalyst. Proc. Jpn. Acad., Ser. B 1993, 69, 203−207. (644) Knani, D.; Gutman, A. L.; Kohn, D. H. Enzymatic Polyesterification in Organic Media -Enzyme-Catalyzed Synthesis of Linear Polyesters. I. Condensation Polymerization of Linear 2403


Chemical Reviews

Review

Hydroxyesters. II. Ring-Opening Polymerization of ε-Caprolactone. J. Polym. Sci., Part A: Polym. Chem. 1993, 31, 1221−1232. (645) Foresti, M. L.; Ferreira, M. L. Synthesis of Polycaprolactone Using Free/Supported Enzymatic and Non-Enzymatic Catalysts. Macromol. Rapid Commun. 2004, 25, 2025−2028. (646) Albertsson, A. C.; Varma, I. K. Recent Developments in Ring Opening Polymerization of Lactones for Biomedical Applications. Biomacromolecules 2003, 4, 1466−1486. (647) Varma, I. K.; Albertsson, A. C.; Rajkhowa, R.; Srivastava, R. K. Enzyme Catalyzed Synthesis of Polyesters. Prog. Polym. Sci. 2005, 30, 949−981. (648) Uyama, H.; Kikuchi, H.; Takeya, K.; Kabayashi, S. LipaseCatalyzed Ring-Opening Polymerization and Copolymerization of 15Pentadecanolide. Acta Polym. 1996, 47, 357−360. (649) Uyama, H.; Namekawa, S.; Kobayash, S. Mechanistic Studies on the Lipase-Catalyzed Ring-Opening Polymerization of Lactones. Polym. J. 1997, 29, 299−301. (650) Kobayashi, S.; Uyama, H.; Namekawa, S. In Vitro Biosynthesis of Polyesters with Isolated Enzymes in Aqueous Systems and Organic Solvents. Polym. Degrad. Stab. 1998, 59, 195−201. (651) Kobayashi, S.; Uyama, H. Precision Enzymatic Polymerization to Polyesters with Lipase Catalysts. Macromol. Symp. 1999, 144, 237− 246. (652) van der Mee, L.; Helmich, F.; de Bruijn, R.; Vekemans, J.; Palmans, A. R. A.; Meijer, E. W. Investigation of Lipase-Catalyzed Ring-Opening Polymerizations of Lactones with Various Ring Sizes: Kinetic Evaluation. Macromolecules 2006, 39, 5021−5027. (653) MacDonald, R. T.; Pulapura, S. K.; Svirkin, Y. Y.; Gross, R. A.; Kaplan, D. L.; Akkara, J.; Swift, G.; Wolk, S. Enzyme-Catalyzed εCaprolactone Ring-Opening Polymerization. Macromolecules 1995, 28, 73−78. (654) Henderson, L. A.; Svirkin, Y. Y.; Gross, R. A.; Kaplan, D. L.; Swift, G. Enzyme-Catalyzed Polymerizations of ε-Caprolactone: Effects of Initiator on Product Structure, Propagation Kinetics, and Mechanism. Macromolecules 1996, 29, 7759−7766. (655) Matsumura, S.; Ebata, H.; Toshima, K. A New Strategy for Sustainable Polymer Recycling Using an Enzyme: Poly(ε-caprolactone). Macromol. Rapid Commun. 2000, 21, 860−863. (656) Ebata, H.; Toshima, K.; Matsumura, S. A Strategy for Increasing Molecular Weight of Polyester by Lipase-Catalyzed Polymerization. Chem. Lett. 2001, 798−799. (657) Kumar, A.; Kalra, B.; Dekhterman, A.; Gross, R. A. Efficient Ring-Opening Polymerization and Copolymerization of ε-Caprolactone and ω-Pentadecalactone Catalyzed by Candida antartica Lipase B. Macromolecules 2000, 33, 6303−6309. (658) Mei, Y.; Kumar, A.; Gross, R. A. Probing Water-Temperature Relationships for Lipase-Catalyzed Lactone Ring-Opening Polymerizations. Macromolecules 2002, 35, 5444−5448. (659) Cordova, A.; Iversen, T.; Hult, K. Lipase-Catalyzed Formation of End-Functionalized Poly(ε-caprolactone) by Initiation and Termination Reactions. Polymer 1999, 40, 6709−6721. (660) Dong, H.; Cao, S. G.; Li, Z. Q.; Han, S. P.; You, D. L.; Shen, J. C. Study on the Enzymatic Polymerization Mechanism of Lactone and the Strategy for Improving the Degree of Polymerization. J. Polym. Sci., Part A: Polym. Chem. 1999, 37, 1265−1275. (661) Divakar, S. Porcine pancreas Lipase Catalyzed Ring-Opening Polymerization of ε-Caprolactone. J. Macromol. Sci., Part A: Pure Appl.Chem. 2004, 41, 537−546. (662) Kumar, A.; Gross, R. A. Candida antartica Lipase B Catalyzed Polycaprolactone Synthesis: Effects of Organic Media and Temperature. Biomacromolecules 2000, 1, 133−138. (663) Kobayashi, S.; Takeya, K.; Suda, S.; Uyama, H. LipaseCatalyzed Ring-Opening Polymerization of Medium-Size Lactones to Polyesters. Macromol. Chem. Phys. 1998, 199, 1729−1736. (664) Kikuchi, H.; Uyama, H.; Kobayashi, S. Lipase-Catalyzed RingOpening Polymerization of Substituted Lactones. Polym. J. 2002, 34, 835−840. (665) Küllmer, K.; Kikuchi, H.; Uyama, H.; Kobayashi, S. LipaseCatalyzed Ring-Opening Polymerization of α-Methyl-δ-valerolactone

and α-Methyl-ε-caprolactone. Macromol. Rapid Commun. 1998, 19, 127−130. (666) Li, Q.; Li, G.; Yu, S.; Zhang, Z.; Ma, F.; Feng, Y. Ring-Opening Polymerization of ε-Caprolactone Catalyzed by a Novel Thermophilic Lipase from Fervidobacterium nodosum. Process Biochem. 2011, 46, 253−257. (667) Barrera-Rivera, K. A.; Peponi, L.; Marcos-Fernandez, A.; Kenny, J. M.; Martinez-Richa, A. Synthesis, Characterization and Hydrolytic Degradation of Polyester-Urethanes Obtained by Lipase Biocatalysis. Polym. Degrad. Stab. 2014, 108, 188−194. (668) Uyama, H.; Suda, S.; Kikuchi, H.; Kobayashi, S. Extremely Efficient Catalysis of Immobilized Lipase in Ring-Opening Polymerization of Lactones. Chem. Lett. 1997, 1109−1110. (669) Sivalingam, G.; Madras, G. Modeling of Lipase Catalyzed RingOpening Polymerization of ε-Caprolactone. Biomacromolecules 2004, 5, 603−609. (670) Chen, B.; Hu, J.; Miller, E. M.; Xie, W. C.; Cai, M. M.; Gross, R. A. Candida antarctica Lipase B Chemically Immobilized on EpoxyActivated Micro- and Nanobeads: Catalysts for Polyester Synthesis. Biomacromolecules 2008, 9, 463−471. (671) Hans, M.; Gasteier, P.; Keul, H.; Moeller, M. Ring-Opening Polymerization of ε-Caprolactone by Means of Mono- and Multifunctional Initiators: Comparison of Chemical and Enzymatic Catalysis. Macromolecules 2006, 39, 3184−3193. (672) Okajima, S.; Kondo, R.; Toshima, K.; Matsumura, S. LipaseCatalyzed Transformation of Poly(butylene adipate) and Poly(butylene succinate) into Repolymerizable Cyclic Oligomers. Biomacromolecules 2003, 4, 1514−1519. (673) Soeda, Y.; Toshima, K.; Matsumura, S. Synthesis and Chemical Recycling of Novel Poly(ester-urethane)s Using an Enzyme. Macromol. Biosci. 2005, 5, 277−288. (674) Dai, S. Y.; Li, Z. Enzymatic Preparation of Novel Thermoplastic Di-Block Copolyesters Containing Poly[(R)-3-hydroxybutyrate] and Poly(ε-caprolactone) Blocks via Ring-Opening Polymerization. Biomacromolecules 2008, 9, 1883−1893. (675) Sugihara, S.; Toshima, K.; Matsumura, S. New Strategy for Enzymatic Synthesis of High-Molecular-Weight Poly(butylene succinate) via Cyclic Oligomers. Macromol. Rapid Commun. 2006, 27, 203− 207. (676) Miletic, N.; Vukovic, Z.; Nastasovic, A.; Loos, K. Macroporous Poly(glycidyl methacrylate-co-ethylene glycol dimethacrylate) ResinsVersatile Immobilization Supports for Biocatalysts. J. Mol. Catal. B: Enzym. 2009, 56, 196−201. (677) Poojari, Y.; Beemat, J. S.; Clarson, S. J. Enzymatic Synthesis of Poly(ε-caprolactone): Thermal Properties, Recovery, and Reuse of Lipase B from Candida antarctica Immobilized on Macroporous Acrylic Resin Particles. Polym. Bull. 2013, 70, 1543−1552. (678) Omay, D. Immobilization of Lipase onto a Photo-Crosslinked Polymer Network: Characterization and Polymerization Applications. Biocatal. Biotransform. 2014, 32, 132−140. (679) Ozturk Duskunkorur, H.; Pollet, E.; Phalip, V.; Guvenilir, Y.; Averous, L. Lipase Catalyzed Synthesis of Polycaprolactone and ClayBased Nanohybrids. Polymer 2014, 55, 1648−1655. (680) Gustavsson, M. T.; Persson, P. V.; Iversen, T.; Hult, K.; Martinelle, M. Polyester Coating of Cellulose Fiber Surfaces Catalyzed by a Cellulose-Binding Module-Candida antarctica Lipase B Fusion Protein. Biomacromolecules 2004, 5, 106−112. (681) Kerep, P.; Ritter, H. Influence of Microwave Irradiation on the Lipase-Catalyzed Ring-Opening Polymerization of ε-Caprolactone. Macromol. Rapid Commun. 2006, 27, 707−710. (682) Matos, T. D.; King, N.; Simmons, L.; Walker, C.; McClain, A. R.; Mahapatro, A.; Rispoli, F. J.; McDonnell, K. T.; Shah, V. Microwave Assisted Lipase Catalyzed Solvent-Free Poly-ε-caprolactone Synthesis. Green Chem. Lett. Rev. 2011, 4, 73−79. (683) Matsumura, S.; Beppu, H.; Nakamura, K.; Osanai, S.; Toshima, K. Preparation of Poly(β-malic acid) by Enzymatic Ring-Opening Polymerization of Benzyl β-Malolactonate. Chem. Lett. 1996, 795− 796. 2404


Chemical Reviews

Review

(684) Namekawa, S.; Uyama, H.; Kobayashi, S. Lipase-Catalyzed Ring-Opening Polymerization and Copolymerization of β-Propiolactone. Polym. J. 1996, 28, 730−731. (685) Svirkin, Y. Y.; Xu, J.; Gross, R. A.; Kaplan, D. L.; Swift, G. Enzyme-Catalyzed Stereoelective Ring-Opening Polymerization of αMethyl-β-propiolactone. Macromolecules 1996, 29, 4591−4597. (686) Nobes, G. A. R.; Kazlauskas, R. J.; Marchessault, R. H. LipaseCatalyzed Ring-Opening Polymerization of Lactones: A Novel Route to Poly(hydroxyalkanoate)s. Macromolecules 1996, 29, 4829−4833. (687) Xie, W. H.; Li, J.; Chen, D. P.; Wang, P. G. Ring-Opening Polymerization of β-Butyrolactone by Thermophilic Lipases. Macromolecules 1997, 30, 6997−6998. (688) Panova, A. A.; Taktak, S.; Randriamahefa, S.; Cammas-Marion, S.; Guerin, P.; Kaplan, D. L. Polymerization of Propyl Malolactonate in the Presence of Candida rugosa Lipase. Biomacromolecules 2003, 4, 19− 27. (689) Kobayashi, S.; Uyama, H.; Namekawa, S.; Hayakawa, H. Enzymatic Ring-Opening Polymerization and Copolymerization of 8Octanolide by Lipase Catalyst. Macromolecules 1998, 31, 5655−5659. (690) Uyama, H.; Takeya, K.; Kobayashi, S. Enzymatic Ring-Opening Polymerization of Lactones to Polyesters by Lipase Catalyst. -Unusually High Reactivity of Macrolides. Bull. Chem. Soc. Jpn. 1995, 68, 56−61. (691) Uyama, H.; Takeya, K.; Hoshi, N.; Kobayashi, S. LipaseCatalyzed Ring-Opening Polymerization of 12-Dodecanolide. Macromolecules 1995, 28, 7046−7050. (692) Uyama, H.; Kikuchi, H.; Takeya, K.; Hoshi, N.; Kobayashi, S. Immobilized Lipase Showing High Catalytic Activity Toward Enzymatic Ring-Opening Polymerization of Macrolides. Chem. Lett. 1996, 107−108. (693) Bisht, K. S.; Henderson, L. A.; Gross, R. A.; Kaplan, D. L.; Swift, G. Enzyme-Catalyzed Ring-Opening Polymerization of ωPentadecalactone. Macromolecules 1997, 30, 2705−2711. (694) Namekawa, S.; Uyama, H.; Kobayashi, S. Lipase-Catalyzed Ring-Opening Polymerization of 16-Hexadecanolide. Proc. Jpn. Acad., Ser. B 1998, 74, 65−68. (695) Wilberth, H.-K.; Manuel, C.-U. J.; Tania, L.-C.; Manuel, A.-V. Effect of Reaction Temperature on the Physicochemical Properties of Poly(pentadecanolide) Obtained by Enzyme-Catalyzed Ring-Opening Polymerization. Polym. Bull. 2015, 72, 441−45210.1007/s00289-0141288-x. (696) Jiang, Z. Z.; Azim, H.; Gross, R. A.; Focarete, M. L.; Scandola, M. Lipase-Catalyzed Copolymerization of ω-Pentadecalactone with pDioxanone and Characterization of Copolymer Thermal and Crystalline Properties. Biomacromolecules 2007, 8, 2262−2269. (697) Hu, Y.; Ju, L.-K. Lipase-Mediated Deacetylation and Oligomerization of Lactonic Sophorolipids. Biotechnol. Prog. 2003, 19, 303−311. (698) Gao, W.; Hagver, R.; Shah, V.; Xie, W. C.; Gross, R. A.; Ilker, M. F.; Bell, C.; Burke, K. A.; Coughlin, E. B. Glycolipid Polymer Synthesized from Natural Lactonic Sophorolipids by Ring-Opening Metathesis Polymerization. Macromolecules 2007, 40, 145−147. (699) Zini, E.; Gazzano, M.; Scandola, M.; Wallner, S. R.; Gross, R. A. Glycolipid Biomaterials: Solid-State Properties of a Poly(sophorolipid). Macromolecules 2008, 41, 7463−7468. (700) Manzini, B.; Hodge, P.; Ben-Haida, A. Entropically-Driven Ring-Opening Polymerization of Macrocyclic Esters with up to 84Membered Rings Catalysed by Polymer-Supported Candida antarctica Lipase B. Polym. Chem. 2010, 1, 339−346. (701) Strandman, S.; Tsai, I. H.; Lortie, R.; Zhu, X. X. Ring-Opening Polymerization of Bile Acid Macrocycles by Candida antarctica Lipase B. Polym. Chem. 2013, 4, 4312−4316. (702) Matsumura, S.; Mabuchi, K.; Toshima, K. Lipase-Catalyzed Ring-Opening Polymerization of Lactide. Macromol. Rapid Commun. 1997, 18, 477−482. (703) Hans, M.; Keul, H.; Moeller, M. Ring-Opening Polymerization of DD-Lactide Catalyzed by Novozyme 435. Macromol. Biosci. 2009, 9, 239−247.

(704) Takwa, M.; Larsen, M. W.; Hult, K.; Martinelle, M. Rational Redesign of Candida antarctica Lipase B for the Ring Opening Polymerization of D,D-Lactide. Chem. Commun. 2011, 47, 7392− 7394. (705) Omay, D.; Guvenilir, Y. Synthesis and Characterization of Poly(D,L-lactic acid) via Enzymatic Ring Opening Polymerization by Using Free and Immobilized Lipase. Biocatal. Biotransform. 2013, 31, 132−140. (706) Wahlberg, J.; Persson, P. V.; Olsson, T.; Hedenstrom, E.; Iversen, T. Structural Characterization of a Lipase-Catalyzed Copolymerization of ε-Caprolactone and D,L-Lactide. Biomacromolecules 2003, 4, 1068−1071. (707) Huijser, S.; Staal, B. B. P.; Huang, J.; Duchateau, R.; Koning, C. E. Topology Characterization by MALDI-TOF-MS of Enzymatically Synthesized Poly(lactide-co-glycolide). Biomacromolecules 2006, 7, 2465−2469. (708) Cordova, A.; Iversen, T.; Martinelle, M.; Hult, K. LipaseCatalysed Formation of Macrocycles by Ring-Opening Polymerisation of ε-Caprolactone. Polymer 1998, 39, 6519−6524. (709) Berkane, C.; Mezoul, G.; Lalot, T.; Brigodiot, M.; Marechal, E. Lipase-Catalyzed Polyester Synthesis in Organic Medium. Study of Ring-Chain Equilibrium. Macromolecules 1997, 30, 7729−7734. (710) Srivastava, R. K.; Albertsson, A. C. High-Molecular-Weight Poly(1,5-dioxepan-2-one) via Enzyme-Catalyzed Ring-Opening Polymerization. J. Polym. Sci., Part A: Polym. Chem. 2005, 43, 4206−4216. (711) Srivastava, R. K.; Kumar, K.; Varma, I. K.; Albertsson, A. C. Chemo-Enzymatic Synthesis of Comb Polymers. Eur. Polym. J. 2007, 43, 808−817. (712) Nishida, H.; Yamashita, M.; Nagashima, M.; Endo, T.; Tokiwa, Y. Synthesis of Metal-Free Poly(1,4-dioxan-2-one) by EnzymeCatalyzed Ring-Opening Polymerization. J. Polym. Sci., Part A: Polym. Chem. 2000, 38, 1560−1567. (713) Müller, S.; Uyama, H.; Kobayashi, S. Lipase-Catalyzed RingOpening Polymerization of Cyclic Diesters. Chem. Lett. 1999, 1317− 1318. (714) van der Mee, L.; Antens, A.; van de Kruijs, B.; Palmans, A. R. A.; Meijer, E. W. Oxo-Crown-Ethers as Comonomers for Tuning Polyester Properties. J. Polym. Sci., Part A: Polym. Chem. 2006, 44, 2166−2176. (715) Namekawa, S.; Uyama, H.; Kobayashi, S. Lipase-Catalyzed Ring-Opening Polymerization of Lactones in Water. Polym. J. 1998, 30, 269−271. (716) Panlawan, P.; Luangthongkam, P.; Wiemann, L. O.; Sieber, V.; Marie, E.; Durand, A.; Inprakhon, P. Lipase-Catalyzed Interfacial Polymerization of ω-Pentadecalactone in Aqueous Biphasic Medium: A Mechanistic Study. J. Mol. Catal. B: Enzym. 2013, 88, 69−76. (717) Inprakhon, P.; Panlawan, P.; Pongtharankul, T.; Marie, E.; Wiemann, L. O.; Durand, A.; Sieber, V. Toward One-Oot LipaseCatalyzed Synthesis of Poly(ε-caprolactone) Particles in Aqueous Dispersion. Colloids Surf., B 2014, 113, 254−260. (718) Taden, A.; Antonietti, M.; Landfester, K. Enzymatic Polymerization Towards Biodegradable Polyester Nanoparticles. Macromol. Rapid Commun. 2003, 24, 512−516. (719) Malberg, S.; Finne-Wistrand, A.; Albertsson, A.-C. The Environmental Influence in Enzymatic Polymerization of Aliphatic Polyesters in Bulk and Aqueous Mini-emulsion. Polymer 2010, 51, 5318−5322. (720) Nallani, M.; de Hoog, H. P. M.; Cornelissen, J.; Palmans, A. R. A.; van Hest, J. C. M.; Nolte, R. J. M. Polymersome Nanoreactors for Enzymatic Ring-Opening Polymerization. Biomacromolecules 2007, 8, 3723−3728. (721) Takamoto, T.; Uyama, H.; Kobayashi, S. Lipase-Catalyzed Synthesis of Aliphatic Polyesters in Supercritical Carbon Dioxide. ePolym. 2001, 4, 1−6. (722) Loeker, F. C.; Duxbury, C. J.; Kumar, R.; Gao, W.; Gross, R. A.; Howdle, S. M. Enzyme-Catalyzed Ring-Opening Polymerization of ε-Caprolactone in Supercritical Carbon Dioxide. Macromolecules 2004, 37, 2450−2453. 2405


Chemical Reviews

Review

to Terminal-Functionalized and Triblock Polyesters. Macromolecules 2006, 39, 46−54. (744) Kalra, B.; Kumar, A.; Gross, R. A.; Baiardo, M.; Scandola, M. Chemoenzymatic Synthesis of New Brush Copolymers Comprising Poly(ω-pentadecalactone) with Unusual Thermal and Crystalline Properties. Macromolecules 2004, 37, 1243−1250. (745) Zhu, N.; Zhang, Z.-L.; He, W.; Geng, X.-C.; Fang, Z.; Li, X.; Li, Z.-J.; Guo, K. Highly Chemoselective Lipase from Candida sp. 99−125 Catalyzed Ring-Opening Polymerization for Direct Synthesis of ThiolTerminated Poly(ε-caprolactone). Chin. Chem. Lett. 2015, 26, 361− 364. (746) Korzhikov, V. A.; Gusevskaya, K. V.; Litvinchuk, E. N.; Vlakh, E. G.; Tennikova, T. B. Enzyme-Mediated Ring-Opening Polymerization of Pentadecalactone to Obtain Biodegradable Polymer for Fabrication of Scaffolds for Bone Tissue Engineering. Int. J. Polym. Sci. 2013, 2013, 1−10. (747) Bisht, K. S.; Deng, F.; Gross, R. A.; Kaplan, D. L.; Swift, G. Ethyl Glucoside as a Multifunctional Initiator for Enzyme-Catalyzed Regioselective Lactone Ring-Opening Polymerization. J. Am. Chem. Soc. 1998, 120, 1363−1367. (748) Cordova, A.; Iversen, T.; Hult, K. Lipase-Catalyzed Synthesis of Methyl 6-O-Poly(ε-caprolactone)glycopyranosides. Macromolecules 1998, 31, 1040−1045. (749) Cordova, A.; Hult, A.; Hult, K.; Ihre, H.; Iversen, T.; Malmstrom, E. Synthesis of a Poly(ε-caprolactone) Monosubstituted First Generation Dendrimer by Lipase Catalysis. J. Am. Chem. Soc. 1998, 120, 13521−13522. (750) Qian, X.; Wang, J.; Li, Y.; Lin, X.; Wu, Q. Two Enzyme Cooperatively Catalyzed Tandem Polymerization for the Synthesis of Polyester Containing Chiral (R)- or (S)-Ibuprofen Pendants. Macromol. Rapid Commun. 2014, 35, 1788−1794. (751) de Geus, M.; Peters, R.; Koning, C. E.; Heise, A. Insights into the Initiation Process of Enzymatic Ring-Opening Polymerization from Monofunctional Alcohols Using Liquid Chromatography Under Critical Conditions. Biomacromolecules 2008, 9, 752−757. (752) Takwa, M.; Xiao, Y.; Simpson, N.; Malmstrom, E.; Hult, K.; Koning, C. E.; Heise, A.; Martinelle, M. Lipase Catalyzed HEMA Initiated Ring-Opening Polymerization: In Situ Formation of Mixed Polyester Methacrylates by Transesterification. Biomacromolecules 2008, 9, 704−710. (753) Xiao, Y.; Takwa, M.; Hult, K.; Koning, C. E.; Heise, A.; Martinelle, M. Systematic Comparison of HEA and HEMA as Initiators in Enzymatic Ring-Opening Polymerizations. Macromol. Biosci. 2009, 9, 713−720. (754) Hedfors, C.; Ostmark, E.; Malmstrom, E.; Hult, K.; Martinelle, M. Thiol End-Functionalization of Poly(ε-caprolactone), Catalyzed by Candida antarctica Lipase B. Macromolecules 2005, 38, 647−649. (755) Skaria, S.; Smet, M.; Frey, H. Enzyme-Catalyzed Synthesis of Hyperbranched Aliphatic Polyesters. Macromol. Rapid Commun. 2002, 23, 292−296. (756) Yoon, K. R.; Lee, K. B.; Chi, Y. S.; Yun, W. S.; Joo, S. W.; Choi, I. S. Surface-Initiated, Enzymatic Polymerization of Biodegradable Polyesters. Adv. Mater. 2003, 15, 2063−2066. (757) Kim, Y. R.; Paik, H. J.; Ober, C. K.; Coates, G. W.; Batt, C. A. Enzymatic Surface-Initiated Polymerization: A Novel Approach for the In Situ Solid-Phase Synthesis of Biocompatible Polymer Poly(3hydroxybutyrate). Biomacromolecules 2004, 5, 889−894. (758) Hans, M.; Keul, H.; Moeller, M. Poly(ether-ester) Conjugates with Enhanced Degradation. Biomacromolecules 2008, 9, 2954−2962. (759) Uyama, H.; Kobayashi, S. Enzyme-Catalyzed Polymerization to Functional Polymers. J. Mol. Catal. B: Enzym. 2002, 19-20, 117−127. (760) Uyama, H.; Kikuchi, H.; Kobayashi, S. One-Shot Synthesis of Polyester Macromonomer by Enzymatic Ring-Opening Polymerization of Lactone in the Presence of Vinyl Ester. Chem. Lett. 1995, 1047−1048. (761) Uyama, H.; Kikuchi, H.; Kobayashi, S. Single-Step Acylation of Polyester Terminals by Enzymatic Ring-Opening Polymerization of 12-Dodecanolide in the Presence of Acyclic Vinyl Esters. Bull. Chem. Soc. Jpn. 1997, 70, 1691−1695.

(723) Thurecht, K. J.; Heise, A.; deGeus, M.; Villarroya, S.; Zhou, J. X.; Wyatt, M. F.; Howdle, S. M. Kinetics of Enzymatic Ring-Opening Polymerization of ε-Caprolactone in Supercritical Carbon Dioxide. Macromolecules 2006, 39, 7967−7972. (724) Takamoto, T.; Uyama, H.; Kobayashi, S. Lipase-Catalyzed Degradation of Polyester in Supercritical Carbon Dioxide. Macromol. Biosci. 2001, 1, 215−218. (725) Matsumura, S.; Ebata, H.; Kondo, R.; Toshima, K. Organic Solvent-Free Enzymatic Transformation of Poly (ε-caprolactone) into Repolymerizable Oligomers in Supercritical Carbon Dioxide. Macromol. Rapid Commun. 2001, 22, 1325−1329. (726) Duxbury, C. J.; Wang, W. X.; de Geus, M.; Heise, A.; Howdle, S. M. Can Block Copolymers Be Synthesized by a Single-step Chemoenzymatic Route in Supercritical Carbon Dioxide? J. Am. Chem. Soc. 2005, 127, 2384−2385. (727) Villarroya, S.; Zhou, J. X.; Duxbury, C. J.; Heise, A.; Howdle, S. M. Synthesis of Semifluorinated Block Copolymers Containing, Poly(ε-caprolactone) by the Combination of ATRP and Enzymatic ROP in ScCO2. Macromolecules 2006, 39, 633−640. (728) Zhou, J. X.; Villarroya, S.; Wang, W. X.; Wyatt, M. F.; Duxbury, C. J.; Thurecht, K. J.; Howdle, S. M. One-step Chemoenzymatic Synthesis of Poly(ε-caprolactone-block-methyl methacrylate) in Supercritical CO2. Macromolecules 2006, 39, 5352−5358. (729) Lopez-Luna, A.; Gallegos, J. L.; Gimeno, M.; Vivaldo-Lima, E.; Barzana, E. Lipase-Catalyzed Syntheses of Linear and Hyperbranched Polyesters Using Compressed Fluids as Solvent Media. J. Mol. Catal. B: Enzym. 2010, 67, 143−149. (730) Kubisa, P. Ionic Liquids in the Synthesis and Modification of Polymers. J. Polym. Sci., Part A: Polym. Chem. 2005, 43, 4675−4683. (731) Uyama, H.; Takamoto, T.; Kobayashi, S. Enzymatic Synthesis of Polyesters in Ionic Liquids. Polym. J. 2002, 34, 94−96. (732) Marcilla, R.; de Geus, M.; Mecerreyes, D.; Duxbury, C. J.; Koning, C. E.; Heise, A. Enzymatic Polyester Synthesis in Ionic Liquids. Eur. Polym. J. 2006, 42, 1215−1221. (733) Yoshizawa-Fujita, M.; Saito, C.; Takeoka, Y.; Rikukawa, M. Lipase-Catalyzed Polymerization of L-lactide in Ionic Liquids. Polym. Adv. Technol. 2008, 19, 1396−1400. (734) Mena, M.; Lopez-Luna, A.; Shirai, K.; Tecante, A.; Gimeno, M.; Barzana, E. Lipase-Catalyzed Synthesis of Hyperbranched Poly-Llactide in an Ionic Liquid. Bioprocess Biosyst. Eng. 2013, 36, 383−387. (735) Barrera-Rivera, K. A.; Marcos-Fernandez, A.; Vera-Graziano, R.; Martinez-Richa, A. Enzymatic Ring-Opening Polymerization of εCaprolactone by Yarrowia lipolytica Lipase in Ionic Liquids. J. Polym. Sci., Part A: Polym. Chem. 2009, 47, 5792−5805. (736) Wu, C.; Zhang, Z.; He, F.; Zhuo, R. Enzymatic Synthesis of Poly(ε-caprolactone) in Monocationic and Dicationic Ionic Liquids. Biotechnol. Lett. 2013, 35, 879−885. (737) Wu, C.; Zhang, Z.; Chen, C.; He, F.; Zhuo, R. Synthesis of Poly(ε-caprolactone) by an Immobilized Lipase Coated with Ionic Liquids in a Solvent-Free Condition. Biotechnol. Lett. 2013, 35, 1623− 1630. (738) Gumel, A. M.; Annuar, M. S. M.; Chisti, Y.; Heidelberg, T. Ultrasound Assisted Lipase Catalyzed Synthesis of Poly-6-hydroxyhexanoate. Ultrason. Sonochem. 2012, 19, 659−667. (739) Kundu, S.; Bhangale, A. S.; Wallace, W. E.; Flynn, K. M.; Guttman, C. M.; Gross, R. A.; Beers, K. L. Continuous Flow EnzymeCatalyzed Polymerization in a Microreactor. J. Am. Chem. Soc. 2011, 133, 6006−6011. (740) Bhangale, A. S.; Beers, K. L.; Gross, R. A. Enzyme-Catalyzed Polymerization of End-Functionalized Polymers in a Microreactor. Macromolecules 2012, 45, 7000−7008. (741) Uyama, H.; Suda, S.; Kobayashi, S. Enzymatic Synthesis of Terminal-Functionalized Polyesters by Initiator Method. Acta Polym. 1998, 49, 700−703. (742) Castano, M.; Zheng, J.; Puskas, J. E.; Becker, M. L. EnzymeCatalyzed Ring-Opening Polymerization of ε-Caprolactone using Alkyne Functionalized Initiators. Polym. Chem. 2014, 5, 1891−1896. (743) Srivastava, R. K.; Albertsson, A. C. Enzyme-Catalyzed RingOpening Polymerization of Seven-Membered Ring Lactones Leading 2406


Chemical Reviews

Review

(762) Takwa, M.; Simpson, N.; Malmstrom, E.; Hult, K.; Martinelle, M. One-Pot Difunctionalization of Poly(ω-pentadecalactone) with Thiol-Thiol or Thiol-Acrylate Groups, Catalyzed by Candida antarctica Lipase B. Macromol. Rapid Commun. 2006, 27, 1932−1936. (763) Simpson, N.; Takwa, M.; Hult, K.; Johansson, M.; Martinelle, M.; Malmstrom, E. Thiol-Functionalized Poly(ω-pentadecalactone) Telechelics for Semicrystalline Polymer Networks. Macromolecules 2008, 41, 3613−3619. (764) Cordova, A. Synthesis of Amphiphilic Poly(ε-caprolactone) Macromonomers by Lipase Catalysis. Biomacromolecules 2001, 2, 1347−1351. (765) Li, J.; Xie, W. H.; Cheng, H. N.; Nickol, R. G.; Wang, P. G. Polycaprolactone-Modified Hydroxyethylcellulose Films Prepared by Lipase-Catalyzed Ring-Opening Polymerization. Macromolecules 1999, 32, 2789−2792. (766) Kumar, R.; Gross, R. A. Biocatalytic Route to Well-Defined Macromers Built Around a Sugar Core. J. Am. Chem. Soc. 2002, 124, 1850−1851. (767) Huisgen, R.; Ott, H. Die Konfiguration der Carbonestergruppe und die Sondereigenschaften der Lactone. Tetrahedron 1959, 6, 253− 267. (768) Nomura, R.; Ueno, A.; Endo, T. Anionic Ring-Opening Polymerization of Macrocyclic Esters. Macromolecules 1994, 27, 620− 621. (769) Namekawa, S.; Suda, S.; Uyama, H.; Kobayashi, S. LipaseCatalyzed Ring-Opening Polymerization of Lactones to Polyesters and its Mechanistic Aspects. Int. J. Biol. Macromol. 1999, 25, 145−151. (770) Duda, A.; Kowalski, A.; Penczek, S.; Uyama, H.; Kobayashi, S. Kinetics of the Ring-Opening Polymerization of 6-, 7-, 9-, 12-, 13-, 16-, and 17-Membered Lactones. Comparison of Chemical and Enzymatic Polymerizations. Macromolecules 2002, 35, 4266−4270. (771) Uppenberg, J.; Hansen, M. T.; Patkar, S.; Jones, T. A. Sequence, Crystal-Structure Determination and Refinement of 2 Crystal Forms of Lipase B from Candida antarctica. Structure 1994, 2, 293−308. (772) Veld, M. A. J.; Fransson, L.; Palmans, A. R. A.; Meijer, E. W.; Hult, K. Lactone Size Dependent Reactivity in Candida Antarctica Lipase B: A Molecular Dynamics and Docking Study. ChemBioChem 2009, 10, 1330−1334. (773) Hunsen, M.; Abul, A.; Xie, W. C.; Gross, R. Humicola Insolens Cutinase-Catalyzed Lactone Ring-Opening Polymerizations: Kinetic and Mechanistic Studies. Biomacromolecules 2008, 9, 518−522. (774) Kikuchi, H.; Uyama, H.; Kobayashi, S. Lipase-Catalyzed Enantioselective Copolymerization of Substituted Lactones to Optically Active Polyesters. Macromolecules 2000, 33, 8971−8975. (775) Kobayashi, S. Enzymatic Ring-Opening Polymerization of Lactones by Lipase Catalyst: Mechanistic Aspects. Macromol. Symp. 2006, 240, 178−185. (776) Anderson, E. M.; Larsson, K. M.; Kirk, O. One Biocatalyst Many Applications: The Use of Candida antarctica B-Lipase in Organic Synthesis. Biocatal. Biotransform. 1998, 16, 181−204. (777) Peeters, J. W.; van Leeuwen, O.; Palmans, A. R. A.; Meijer, E. W. Lipase-Catalyzed Ring-Opening Polymerizations of 4-Substituted ε-Caprolactones: Mechanistic Considerations. Macromolecules 2005, 38, 5587−5592. (778) Namekawa, S.; Uyama, H.; Kobayashi, S. Lipase-Catalyzed Ring-Opening Polymerization of Lactones in the Presence of Aliphatic Polyesters to Ester Copolymers. Macromol. Chem. Phys. 2001, 202, 801−806. (779) Kumar, A.; Gross, R. A. Candida antarctica Lipase B-Catalyzed Transesterification: New Synthetic Routes to Copolyesters. J. Am. Chem. Soc. 2000, 122, 11767−11770. (780) Takamoto, T.; Kerep, P.; Uyama, H.; Kobayashi, S. LipaseCatalyzed Transesterification of Polyesters to Ester Copolymers. Macromol. Biosci. 2001, 1, 223−227. (781) Uyama, H.; Kobayashi, S.; Morita, M.; Habaue, S.; Okamoto, Y. Chemoselective Ring-Opening Polymerization of a Lactone Having Exo-Methylene Group with Lipase Catalysis. Macromolecules 2001, 34, 6554−6556.

(782) Habaue, S.; Asai, M.; Morita, M.; Okamoto, Y.; Uyama, H.; Kobayashi, S. Chemospecific Ring-Opening Polymerization of αMethylenemacrolides. Polymer 2003, 44, 5195−5200. (783) Veld, M. A. J.; Palmans, A. R. A.; Meijer, E. W. Selective Polymerization of Functional Monomers with Novozym 435. J. Polym. Sci., Part A: Polym. Chem. 2007, 45, 5968−5978. (784) van der Meulen, I.; de Geus, M.; Antheunis, H.; Deumens, R.; Joosten, E. A. J.; Koning, C. E.; Heise, A. Polymers from Functional Macrolactones as Potential Biomaterials: Enzymatic Ring Opening Polymerization, Biodegradation, and Biocompatibility. Biomacromolecules 2008, 9, 3404−3410. (785) Barrera-Rivera, K. A.; Martinez-Richa, A. One-Pot Biocatalytic Synthesis of Sugar Based Poly (ε-caprolactone). Macromol. Symp. 2009, 283−284, 144−151. (786) Duxbury, C. J.; Cummins, D.; Heise, A. Selective Enzymatic Grafting by Steric Control. Macromol. Rapid Commun. 2007, 28, 235− 240. (787) Padovani, M.; Hilker, I.; Duxbury, C. J.; Heise, A. Functionalization of Polymers with High Precision by Dual Regioand Stereoselective Enzymatic Reactions. Macromolecules 2008, 41, 2439−2444. (788) Fujioka, M.; Okada, H.; Kusaka, Y.; Nishiyama, S.; Noguchi, H.; Ishii, S.; Yoshida, Y. Enzymatic Synthesis of Chitin- and Chitosangraf t-Aliphatic Polyesters. Macromol. Rapid Commun. 2004, 25, 1776− 1780. (789) Al-Azemi, T. F.; Kondaveti, L.; Bisht, K. S. Solventless Enantioelective Ring-Opening Polymerization of Substituted εCaprolactones by Enzymatic Catalysis. Macromolecules 2002, 35, 3380−3386. (790) Peeters, J.; Palmans, A. R. A.; Veld, M.; Scheijen, F.; Heise, A.; Meijer, E. W. Cascade Synthesis of Chiral Block Copolymers Combining Lipase Catalyzed Ring Ppening Polymerization and Atom Transfer Radical Polymerization. Biomacromolecules 2004, 5, 1862−1868. (791) Runge, M.; O’Hagan, D.; Haufe, G. Lipase-Catalyzed Polymerization of Fluorinated Lactones and Fluorinated Hydroxycarboxylic Acids. J. Polym. Sci., Part A: Polym. Chem. 2000, 38, 2004− 2012. (792) van Buijtenen, J.; van As, B. A. C.; Verbruggen, M.; Roumen, L.; Vekemans, J.; Pieterse, K.; Hilbers, P. A. J.; Hulshof, L. A.; Palmans, A. R. A.; Meijer, E. W. Switching from S- to R-Selectivity in the Candida antarctica Lipase B-Catalyzed Ring-Opening of ω-Methylated Lactones: Tuning Polymerizations by Ring Size. J. Am. Chem. Soc. 2007, 129, 7393−7398. (793) van As, B. A. C.; van Buijtenen, J.; Heise, A.; Broxterman, Q. B.; Verzijl, G. K. M.; Palmans, A. R. A.; Meijer, E. W. Chiral Oligomers by Iterative Tandem Catalysis. J. Am. Chem. Soc. 2005, 127, 9964− 9965. (794) van Buijtenen, J.; van As, B. A. C.; Meuldijk, J.; Palmans, A. R. A.; Vekemans, J.; Hulshof, L. A.; Meijer, E. W. Chiral Polymers by Iterative Tandem Catalysis. Chem. Commun. 2006, 3169−3171. (795) Zhou, J. X.; Wang, W. X.; Thurecht, K. J.; Villarroya, S.; Howdle, S. M. Simultaneous Dynamic Kinetic Resolution in Combination with Enzymatic Ring-Opening Polymerization. Macromolecules 2006, 39, 7302−7305. (796) Deng, F.; Bisht, K. S.; Gross, R. A.; Kaplan, D. L. Chemoenzymatic Synthesis of a Multiarm Poly(lactide-co-ε-caprolactone). Macromolecules 1999, 32, 5159−5161. (797) Meyer, U.; Palmans, A. R. A.; Loontjens, T.; Heise, A. Enzymatic Ring-Opening Polymerization and Atom Transfer Radical Polymerization from a Bifunctional Initiator. Macromolecules 2002, 35, 2873−2875. (798) Matyjaszewski, K.; Xia, J. H. Atom Transfer Radical Polymerization. Chem. Rev. 2001, 101, 2921−2990. (799) de Geus, M.; Peeters, J.; Wolffs, M.; Hermans, T.; Palmans, A. R. A.; Koning, C. E.; Heise, A. Investigation of Factors Influencing the Chemoenzymatic Synthesis of Block Copolymers. Macromolecules 2005, 38, 4220−4225. 2407


Chemical Reviews

Review

(819) Feder, D.; Gross, R. A. Exploring Chain Length Selectivity in HiC-Catalyzed Polycondensation Reactions. Biomacromolecules 2010, 11, 690−697. (820) Olsson, A.; Lindstrom, M.; Iversen, T. Lipase-Catalyzed Synthesis of an Epoxy-functionalized Polyester from the Suberin Monomer cis-9,10-Epoxy-18-hydroxyoctadecanoic Acid. Biomacromolecules 2007, 8, 757−760. (821) Dong, H.; Wang, H. D.; Cao, S. G.; Shen, J. C. LipaseCatalyzed Polymerization of Lactones and Linear Hydroxyesters. Biotechnol. Lett. 1998, 20, 905−908. (822) Ebata, H.; Toshima, K.; Matsumura, S. Lipase-Catalyzed Synthesis and Properties of Poly[(12-hydroxydodecanoate)-co-(12hydroxystearate)] Directed Towards Novel Green and Sustainable Elastomers. Macromol. Biosci. 2008, 8, 38−45. (823) Noll, O.; Ritter, H. New Side-chain Poly(methacryl-rotaxanes) Bearing Cyclodextrins as Non-covalently Anchored Ring Components. Chemoenzymatic Synthesis and Degradation. Macromol. Rapid Commun. 1997, 18, 53−58. (824) O’Hagan, D.; Parker, A. H. Enzyme Mediated Polyester Synthesis with the Lipase from Candida rugosa - Preparation of an Enantiomerically Enriched Polymer from an A-B Monomer. Polym. Bull. 1998, 41, 519−524. (825) Knani, D.; Kohn, D. H. Enzymatic Polyesterification in Organic Media. II. Enzyme-Catalyzed Synthesis of Lateral-substituted Aliphatic Polyesters and Copolyesters. J. Polym. Sci., Part A: Polym. Chem. 1993, 31, 2887−2897. (826) Kanca, U.; Van Buijtenen, J.; Van As, B. A. C.; Korevaar, P. A.; Vekemans, J.; Palmans, A. R. A.; Meijer, E. W. Iterative Tandem Catalysis of Racemic AB Monomers. J. Polym. Sci., Part A: Polym. Chem. 2008, 46, 2721−2733. (827) Miao, S. D.; Zhang, S. P.; Su, Z. G.; Wang, P. Chemoenzymatic Synthesis of Oleic Acid-based Polyesters for Use as Highly Stable Biomaterials. J. Polym. Sci., Part A: Polym. Chem. 2008, 46, 4243−4248. (828) Ohara, H.; Onogi, A.; Yamamoto, M.; Kobayashi, S. LipaseCatalyzed Oligomerization and Hydrolysis of Alkyl Lactates: Direct Evidence in the Catalysis Mechanism That Enantioselection Is Governed by a Deacylation Step. Biomacromolecules 2010, 11, 2008−2015. (829) Ohara, H.; Nishioka, E.; Yamaguchi, S.; Kawai, F.; Kobayashi, S. Protease-Catalyzed Oligomerization and Hydrolysis of Alkyl Lactates Involving L-Enantioselective Deacylation Step. Biomacromolecules 2011, 12, 3833−3837. (830) Ohara, H.; Yamamoto, M.; Onogi, A.; Hirao, K.; Kobayashi, S. Optical Resolution of n-Butyl D- and L-Lactates Using Immobilized Lipase Catalyst. J. Biosci. Bioeng. 2011, 111, 19−21. (831) Doi, Y. Microbial Polyesters; VCH Publishers: New York, 1990. (832) Steinbüchel, A. Perspectives for Biotechnological Production and Utilization of Biopolymers: Metabolic Engineering of Polyhydroxyalkanoate Biosynthesis Pathways as a Successful Example. Macromol. Biosci. 2001, 1, 1−24. (833) Sato, S.; Ono, Y.; Mochiyama, Y.; Sivaniah, E.; Kikkawa, Y.; Sudesh, K.; Hiraishi, T.; Doi, Y.; Abe, H.; Tsuge, T. Polyhydroxyalkanoate Film Formation and Synthase Activity During In Vitro and In Situ Polymerization on Hydrophobic Surfaces. Biomacromolecules 2008, 9, 2811−2818. (834) Binns, F.; Roberts, S. M.; Taylor, A.; Williams, C. F. Enzymatic Polymerization of an Unactivated Diol Diacid System. J. Chem. Soc., Perkin Trans. 1 1993, 899−904. (835) Wang, Z. L.; Hiltunen, K.; Orava, P.; Seppälä, J.; Linko, Y. Y. Lipase-Catalyzed Polyester Synthesis. J. Macromol. Sci., Part A: Pure Appl.Chem. 1996, 33, 599−612. (836) Linko, Y. Y.; Wang, Z. L.; Seppälä, J. Lipase-Catalyzed Linear Aliphatic Polyester Synthesis in Organic-solvent. Enzyme Microb. Technol. 1995, 17, 506−511. (837) Mahapatro, A.; Kalra, B.; Kumar, A.; Gross, R. A. LipaseCatalyzed Polycondensations: Effect of Substrates and Solvent on Chain Formation, Dispersity, and End-Group Structure. Biomacromolecules 2003, 4, 544−551.

(800) Peeters, J. W.; Palmans, A. R. A.; Meijer, E. W.; Koning, C. E.; Heise, A. Chemoenzymatic Synthesis of Branched Polymers. Macromol. Rapid Commun. 2005, 26, 684−689. (801) van As, B. A. C.; Thomassen, P.; Kalra, B.; Gross, R. A.; Meijer, E. W.; Palmans, A. R. A.; Heise, A. One-Pot Chemoenzymatic Cascade Polymerization Under Kinetic Resolution Conditions. Macromolecules 2004, 37, 8973−8977. (802) de Geus, M.; Schormans, L.; Palmans, A. A.; Koning, C. E.; Heise, A. Block Copolymers by Chemoenzymatic Cascade Polymerization: A Comparison of Consecutive and Simultaneous Reactions. J. Polym. Sci., Part A: Polym. Chem. 2006, 44, 4290−4297. (803) Sha, K.; Li, D. S.; Wang, S. W.; Qin, L.; Wang, J. Y. Synthesis and Characterization of Diblock Copolymer by Enzymatic RingOpening Polymerization and ATRP from a Novel Bifunctional Initiator. Polym. Bull. 2005, 55, 349−355. (804) Sha, K.; Li, D. S.; Li, Y. P.; Liu, X. T.; Wang, S. W.; Guan, J. Q.; Wang, J. Y. Synthesis, Characterization, and Micellization of an EpoxyBased Amphiphilic Diblock Copolymer of ε-Caprolactone and Glycidyl Methacrylate by Enzymatic Ring-Opening Polymerization and Atom Transfer Radical Polymerization. J. Polym. Sci., Part A: Polym. Chem. 2007, 45, 5037−5049. (805) Sha, K.; Li, D. S.; Li, Y. P.; Ai, P.; Liu, X. T.; Wang, W.; Xu, Y. X.; Wang, S. W.; Wu, M. Z.; Zhang, B.; Wang, J. Y. Chemoenzymatic Synthesis of an AB-Type Diblock Copolymer Combining Enzymatic Self-Condensation Polymerization and AtomTtransfer Radical Polymerization. J. Polym. Sci., Part A: Polym. Chem. 2006, 44, 3393−3399. (806) Hans, M.; Keul, H.; Heise, A.; Moeller, M. Chemoenzymatic Approach Toward Heterografted Molecular Bottle Brushes. Macromolecules 2007, 40, 8872−8880. (807) Hao, X. J.; Albertin, L.; Foster, L. J. R.; Davis, T. P.; BarnerKowollik, C. A New Chemo-Enzymatic Route to Side-chain Liquidcrystalline Polymers: The Synthesis and Polymerization of 6-(4Methoxybiphenyl-4 ′-oxy)hexyl Vinyl Hexanedioate. Macromol. Biosci. 2003, 3, 675−683. (808) Popescu, D.; Keul, H.; Moeller, M. Highly Functional Poly(meth)acrylates via Cascade Reaction. Macromol. Chem. Phys. 2008, 210, 123−139. (809) Bonduelle, C.; Martin-Vaca, B.; Bourissou, D. Lipase-Catalyzed Ring-Opening Polymerization of the O-Carboxylic Anhydride Derived from Lactic Acid. Biomacromolecules 2009, 10, 3069−3073. (810) Okumura, S.; Iwai, M.; Tominaga, Y. Synthesis of Ester Oligomer by Aspergillius-niger Lipase. Agric. Biol. Chem. 1984, 48, 2805−2808. (811) Ajima, A.; Yoshimoto, T.; Takahashi, K.; Tamaura, Y.; Saito, Y.; Inada, Y. Polymerization of 10-Hydroxydecanoic Acid in Benzene with Polyethylene Glycol-Modified Lipase. Biotechnol. Lett. 1985, 7, 303−306. (812) Ohya, Y.; Sugitou, T.; Ouchi, T. Polycondensation of αHydroxy Acids by Enzymes of PEG-Modified Enzymes in Organic Media. J. Macromol. Sci., Part A: Pure Appl.Chem. 1995, 32, 179−190. (813) Kiran, K. R.; Divakar, S. Lipase-Catalysed Polymerization of Lactic Acid and Its Film Forming Properties. World J. Microbiol. Biotechnol. 2003, 19, 859−865. (814) Matsumura, S.; Takahashi, J. Enzymatic-Synthesis of Functional Oligomers. 1. Lipase Catalyzed Polymerization of HydroxyAcids. Makromol. Chem., Rapid Commun. 1986, 7, 369−373. (815) Ebata, H.; Toshima, K.; Matsumura, S. Lipase-Catalyzed Synthesis and Curing of High-Molecular-Weight Polyricinoleate. Macromol. Biosci. 2007, 7, 798−803. (816) Noll, O.; Ritter, H. Enzymes in Polymer Chemistry, 9. Polymerizable Oligoesters from Cholic Acid via Lipase Catalyzed Condensation Reactions. Macromol. Rapid Commun. 1996, 17, 553− 557. (817) O’Hagan, D.; Zaidi, N. A. Enzyme-Catalyzed Condensation Polymerization of 11-Hydroxyundecanoic Acid with Lipase from Candida-cylindracea. Polymer 1994, 35, 3576−3578. (818) Mahapatro, A.; Kumar, A.; Gross, R. A. Mild, Solvent-Free ωHydroxy Acid Polycondensations Catalyzed by Candida antarctica Lipase B. Biomacromolecules 2004, 5, 62−68. 2408


Chemical Reviews

Review

(838) Linko, Y. Y.; Seppälä, J. Producing High Molecular Weight Biodegradable Polyesters. CHEMTECH 1996, 26, 25−31. (839) Uyama, H.; Inada, K.; Kobayashi, S. Enzymatic Polymerization of Dicarboxylic Acid and Glycol to Polyester in Solvent-Free System. Chem. Lett. 1998, 1285−1286. (840) Binns, F.; Harffey, P.; Roberts, S. M.; Taylor, A. Studies of Lipase-Catalyzed Polyesterification of an Unactivated Diacid/Diol System. J. Polym. Sci., Part A: Polym. Chem. 1998, 36, 2069−2080. (841) Binns, F.; Harffey, P.; Roberts, S. M.; Taylor, A. Studies Leading to the Large Scale Synthesis of Polyesters Using Enzymes. J. Chem. Soc., Perkin Trans. 1 1999, 2671−2676. (842) Uyama, H.; Inada, K.; Kobayashi, S. Lipase-Catalyzed Synthesis of Aliphatic Polyesters by Polycondensation of Dicarboxylic Acids and Glycols in Solvent-Free System. Polym. J. 2000, 32, 440−443. (843) Korupp, C.; Weberskirch, R.; Muller, J. J.; Liese, A.; Hilterhaus, L. Scaleup of Lipase-Catalyzed Polyester Synthesis. Org. Process Res. Dev. 2010, 14, 1118−1124. (844) Yao, D.-H.; Li, G.-J.; Kuila, T.; Li, P.; Kim, N.-H.; Kim, S.-I.; Lee, J.-H. Lipase-Catalyzed Synthesis and Characterization of Biodegradable Polyester Containing L-Malic Acid Unit in Solvent System. J. Appl. Polym. Sci. 2011, 120, 1114−1120. (845) Vouyiouka, S. N.; Topakas, E.; Katsini, A.; Papaspyrides, C. D.; Christakopoulos, P. A Green Route for the Preparation of Aliphatic Polyesters via Lipase-Catalyzed Prepolymerization and Low-Temperature Postpolymerization. Macromol. Mater. Eng. 2013, 298, 679−689. (846) Kumar, A.; Kulshrestha, A. S.; Gao, W.; Gross, R. A. Versatile Route to Polyol Polyesters by Lipase Catalysis. Macromolecules 2003, 36, 8219−8221. (847) Fu, H. Y.; Kulshrestha, A. S.; Gao, W.; Gross, R. A.; Baiardo, M.; Scandola, M. Physical Characterization of Sorbitol or Glycerol Containing Aliphatic Copolyesters Synthesized by Lipase-Catalyzed Polymerization. Macromolecules 2003, 36, 9804−9808. (848) Kulshrestha, A. S.; Gao, W.; Gross, R. A. Glycerol Copolyesters: Control of Branching and Molecular Weight Using a Lipase Catalyst. Macromolecules 2005, 38, 3193−3204. (849) Kulshrestha, A. S.; Gao, W.; Fu, H. Y.; Gross, R. A. Synthesis and Characterization of Branched Polymers from Lipase-Catalyzed Trimethylolpropane Copolymerizations. Biomacromolecules 2007, 8, 1794−1801. (850) Zhang, Y.-R.; Spinella, S.; Xie, W.; Cai, J.; Yang, Y.; Wang, Y.Z.; Gross, R. A. Polymeric Triglyceride Analogs Prepared by EnzymeCatalyzed Condensation Polymerization. Eur. Polym. J. 2013, 49, 793− 803. (851) Hu, J.; Gao, W.; Kulshrestha, A.; Gross, R. A. ″Sweet Polyesters″: Lipase-Catalyzed Condensation - Polymerizations of Alditols. Macromolecules 2006, 39, 6789−6792. (852) Hunsen, M.; Azim, A.; Mang, H.; Wallner, S. R.; Ronkvist, A.; Xie, W. C.; Gross, R. A. A Cutinase with Polyester Synthesis Activity. Macromolecules 2007, 40, 148−150. (853) Yang, Y. X.; Lu, W. H.; Zhang, X. Y.; Xie, W. C.; Cai, M. M.; Gross, R. A. Two-Step Biocatalytic Route to Biobased Functional Polyesters from ω-Carboxy Fatty Acids and Diols. Biomacromolecules 2010, 11, 259−268. (854) Kanelli, M.; Douka, A.; Vouyiouka, S.; Papaspyrides, C. D.; Topakas, E.; Papaspyridi, L.-M.; Christakopoulos, P. Production of Biodegradable Polyesters via Enzymatic Polymerization and Solid State Finishing. J. Appl. Polym. Sci. 2014, 131, DOI: 4082010.1002/ app.40820. (855) Habeych, D. I.; Juhl, P. B.; Pleiss, J.; Vanegas, D.; Eggink, G.; Boeriu, C. G. Biocatalytic Synthesis of Polyesters from Sugar-based Building Blocks Using Immobilized Candida antarctica Lipase B. J. Mol. Catal. B: Enzym. 2011, 71, 1−9. (856) Barrett, D. G.; Merkel, T. J.; Luft, J. C.; Yousaf, M. N. OneStep Syntheses of Photocurable Polyesters Based on a Renewable Resource. Macromolecules 2010, 43, 9660−9667. (857) Yang, Y. X.; Lu, W. H.; Cai, J. L.; Hou, Y.; Ouyang, S. Y.; Xie, W. C.; Gross, R. A. Poly(oleic diacid-co-glycerol): Comparison of Polymer Structure Resulting from Chemical and Lipase Catalysis. Macromolecules 2011, 44, 1977−1985.

(858) Uyama, H.; Wada, S.; Kobayashi, S. Synthesis of Aliphatic Polyester by Insertion of Glycols into Polyanhydrides Using Lipase Catalyst. Chem. Lett. 1999, 893−894. (859) Wallace, J. S.; Morrow, C. J. Biocatalytic Synthesis of Polymers -Synthesis of an Optically-Active, Epoxy-Substituted Polyester by Lipase-Catalyzed Polymerization. J. Polym. Sci., Part A: Polym. Chem. 1989, 27, 2553−2567. (860) Japu, C.; Martı ́nez de Ilarduya, A.; Alla, A.; Jiang, Y.; Loos, K.; Munoz-Guerra, S. Copolyesters Made from 1,4-Butanediol, Sebacic Acid, and D-Glucose by Melt and Enzymatic Polycondensation. Biomacromolecules 2015, 16, 868−879. (861) Wang, H. Y.; Zhang, W. W.; Wang, N.; Li, C.; Li, K.; Yu, X. Q. Biocatalytic Synthesis and in Vitro Release of Biodegradable Linear Polyesters with Pendant Ketoprofen. Biomacromolecules 2010, 11, 3290−3293. (862) Wang, H.-Y.; Zhou, Y.-J.; Wang, Z.; Wang, N.; Li, K.; Yu, X.-Q. Enzyme-Catalyzed Synthesis of a Novel Thermosensitive Polyester with Pendant Ketoprofen. Macromol. Biosci. 2011, 11, 595−599. (863) Jacobson, H.; Stockmayer, W. H. Intermolecular Reaction in Polycondensations. 1. The Theory of Linear Systems. J. Chem. Phys. 1950, 18, 1600−1606. (864) Lavalette, A.; Lalot, T.; Brigodiot, M.; Marechal, E. LipaseCatalyzed Synthesis of a Pure Macrocyclic Polyester from Dimethyl Terephthalate and Diethylene Glycol. Biomacromolecules 2002, 3, 225−228. (865) Warwel, S.; Demes, C.; Steinke, G. Polyesters by LipaseCatalyzed Polycondensation of Unsaturated and Epoxidized Longchain α,ω-Dicarboxylic acid Methyl Esters with Diols. J. Polym. Sci., Part A: Polym. Chem. 2001, 39, 1601−1609. (866) Azim, H.; Dekhterman, A.; Jiang, Z. Z.; Gross, R. A. Candida antarctica Lipase B-Catalyzed Synthesis of Poly(butylene succinate): Shorter Chain Building Blocks Also Work. Biomacromolecules 2006, 7, 3093−3097. (867) Linko, Y. Y.; Wang, Z. L.; Seppälä, J. Lipase-Catalyzed Synthesis of Poly(1,4-butanediol succinate) in Organic-Solvent. Biocatal. Biotransform. 1994, 8, 269−282. (868) Wallace, J. S.; Morrow, C. J. Biocatalytic Synthesis of Polymers. 2. Preparation of [AA-BB]X Polyiesters by Porcine Pancreatic Lipase Catalyzed Transesterification in Anhydrous, Low Polarity Organic Solvents. J. Polym. Sci., Part A: Polym. Chem. 1989, 27, 3271−3284. (869) Brazwell, E. M.; Filos, D. Y.; Morrow, C. J. Biocatalytic Synthesis of Polymers. 3. Formation of a High-Molecular-Weight Polyester Through Limitation of Hydrolysis by Enzyme-Bound Water and Through Equilibrium Control. J. Polym. Sci., Part A: Polym. Chem. 1995, 33, 89−95. (870) Chaudhary, A. K.; Beckman, E. J.; Russell, A. J. Rational Control of Polymer Molecular-weight and Dispersity During EnzymeCatalyzed Polyester Synthesis in Supercritical Fluids. J. Am. Chem. Soc. 1995, 117, 3728−3733. (871) Hilker, I.; Rabani, G.; Verzijl, G. K. M.; Palmans, A. R. A.; Heise, A. Chiral Polyesters by Dynamic Kinetic Resolution Polymerization. Angew. Chem., Int. Ed. 2006, 45, 2130−2132. (872) Uyama, H.; Kobayashi, S. Lipase-Catalyzed Polymerization of Divinyl Adipate with Glycols to Polyesters. Chem. Lett. 1994, 1687− 1690. (873) Uyama, H.; Yaguchi, S.; Kobayashi, S. Lipase-Catalyzed Polycondensation of Dicarboxylic Acid-Divinyl Esters and Glycols to Aliphatic Polyesters. J. Polym. Sci., Part A: Polym. Chem. 1999, 37, 2737−2745. (874) Chaudhary, A. K.; Beckman, E. J.; Russell, A. J. Biocatalytic Polyester Synthesis: Analysis of the Evolution of Molecular Weight and End Group Functionality. Biotechnol. Bioeng. 1997, 55, 227−239. (875) Kline, B. J.; Lele, S. S.; Lenart, P. J.; Beckman, E. J.; Russell, A. J. Use of a Batch-Stirred Reactor to Rationally Tailor Biocatalytic Polytransesterification. Biotechnol. Bioeng. 2000, 67, 424−434. (876) Naolou, T.; Busse, K.; Kressler, J. Synthesis of Well-Defined Graft Copolymers by Combination of Enzymatic Polycondensation and ″Click″ Chemistry. Biomacromolecules 2010, 11, 3660−3667. 2409


Chemical Reviews

Review

tional Acrylates Prepared via Lipase−Catalyzed Transacylation Reactions. J. Polym. Sci., Part A: Polym. Chem. 2010, 48, 2610−2621. (898) Popescu, D.; Keul, H.; Moeller, M. Poly(meth)acrylates Obtained by Cascade Reaction. Macromol. Rapid Commun. 2011, 32, 559−572. (899) Castano, M.; Seo, K. S.; Kim, E. H.; Becker, M. L.; Puskas, J. E. Green Polymer Chemistry VIII: Synthesis of Halo-Ester-Functionalized Poly(ethylene glycol)s via Enzymatic Catalysis. Macromol. Rapid Commun. 2013, 34, 1375−1380. (900) Yeniad, B.; Koekluekaya, N. O.; Naik, H.; Fijten, M. W. M.; Koning, C. E.; Heise, A. Synthesis of Enantiopure Homo and Copolymers by RAFT Polymerization and Investigation of their Enantioselective Lipase-Catalyzed Esterification. J. Polym. Sci., Part A: Polym. Chem. 2013, 51, 84−93. (901) Hoffmann, C.; Stuparu, M. C.; Daugaard, A.; Khan, A. AzaMichael Addition Reaction: Post-polymerization Modification and Preparation of PEI/PEG-based Polyester Hydrogels from Enzymatically Synthesized Reactive Polymers. J. Polym. Sci., Part A: Polym. Chem. 2015, 53, 745−749. (902) Kobayashi, S.; Uyama, H. Enzymatic Polymerization of Cyclic Acid Anhydrides and Glycols by a Lipase Catalyst. Makromol. Chem., Rapid Commun. 1993, 14, 841−844. (903) Uyama, H.; Wada, S.; Fukui, T.; Kobayashi, S. LipaseCatalyzed Synthesis of Polyesters from Anhydride Derivatives Involving Dehydration. Biochem. Eng. J. 2003, 16, 145−152. (904) Yamaguchi, S.; Tanha, M.; Hult, A.; Okuda, T.; Ohara, H.; Kobayashi, S. Green Polymer Chemistry: Lipase-Catalyzed Synthesis of Bio-based Reactive Polyesters Employing Itaconic Anhydride as a Renewable Monomer. Polym. J. 2014, 46, 2−13. (905) Namekawa, S.; Uyama, H.; Kobayashi, S. Enzymatic Synthesis of Polyesters from Lactones, Dicarboxylic Acid Divinyl Esters, and Glycols Through Combination of Ring-opening Polymerization and Polycondensation. Biomacromolecules 2000, 1, 335−338. (906) Jiang, Z. Z. Lipase-Catalyzed Synthesis of Aliphatic Polyesters via Copolymerization of Lactone, Dialkyl Diester, and Diol. Biomacromolecules 2008, 9, 3246−3251. (907) Jiang, Z.; Zhang, J. Lipase-catalyzed Synthesis of Aliphatic Polyesters via Copolymerization of Lactide with Diesters and Diols. Polymer 2013, 54, 6105−6113. (908) Mazzocchetti, L.; Scandola, M.; Jiang, Z. Z. Enzymatic Synthesis and Structural and Thermal Properties of Poly(ωpentadecalactone-co-butylene-co-succinate). Macromolecules 2009, 42, 7811−7819. (909) Eriksson, M.; Fogelstrom, L.; Hult, K.; Malmstrom, E.; Johansson, M.; Trey, S.; Martinelle, M. Enzymatic One-Pot Route to Telechelic Polypentadecalactone Epoxide: Synthesis, UV Curing, and Characterization. Biomacromolecules 2009, 10, 3108−3113. (910) Kobayashi, S.; Kikuchi, H.; Uyama, H. Lipase-Catalyzed RingOpening Polymerization of 1,3-Dioxan-2-one. Macromol. Rapid Commun. 1997, 18, 575−579. (911) Bisht, K. S.; Svirkin, Y. Y.; Henderson, L. A.; Gross, R. A.; Kaplan, D. L.; Swift, G. Lipase-Catalyzed Ring-Opening Polymerization of Trimethylene Carbonate. Macromolecules 1997, 30, 7735− 7742. (912) Chen, R.-Y.; Zhang, Y.-R.; Wang, Y.-Z. Synthesis of Poly(1,4dioxan-2-one) Catalyzed by Immobilized Lipase CA. J. Mol. Catal. B: Enzym. 2009, 57, 224−228. (913) Matsumura, S.; Tsukada, K.; Toshima, K. Enzyme-Catalyzed Ring-Opening Polymerization of 1,3-Dioxan-2-one to Poly(trimethylene carbonate). Macromolecules 1997, 30, 3122−3124. (914) Yamamoto, Y.; Kaihara, S.; Toshima, K.; Matsumura, S. HighMolecular-Weight Polycarbonates Synthesized by Enzymatic ROP of a Cyclic Carbonate as a Green Process. Macromol. Biosci. 2009, 9, 968− 978. (915) Wang, H.-F.; Su, W.; Zhang, C.; Luo, X.-H.; Feng, J. Biocatalytic Fabrication of Fast-Degradable, Water-soluble Polycarbonate Functionalized with Tertiary Amine Groups in Backbone. Biomacromolecules 2010, 11, 2550−2557.

(877) Kim, D. Y.; Dordick, J. S. Combinatorial Array-based Enzymatic Polyester Synthesis. Biotechnol. Bioeng. 2001, 76, 200−206. (878) Uyama, H.; Yaguchi, S.; Kobayashi, S. Enzymatic Synthesis of Aromatic Polyesters by Lipase-Catalyzed Polymerization of Dicarboxylic Acid Divinyl Esters and Glycols. Polym. J. 1999, 31, 380−383. (879) Rodney, R. L.; Allinson, B. T.; Beckman, E. J.; Russell, A. J. Enzyme-Catalyzed Polycondensation Reactions for the Synthesis of Aromatic Polycarbonates and Polyesters. Biotechnol. Bioeng. 1999, 65, 485−489. (880) Barrett, D. G.; Yousaf, M. N. Preparation of a Class of Versatile, Chemoselective, and Amorphous Polyketoesters. Biomacromolecules 2008, 9, 2029−2035. (881) Frampton, M. B.; Subczynska, I.; Zelisko, P. M. Biocatalytic Synthesis of Silicone Polyesters. Biomacromolecules 2010, 11, 1818− 1825. (882) Jiang, Y.; Woortman, A. J. J.; van Ekenstein, G. O. R.; Loos, K. Enzyme-Catalyzed Synthesis of Unsaturated Aliphatic Polyesters Based on Green Monomers from Renewable Resources. Biomolecules 2013, 3, 461−480. (883) Liu, W.; Wang, F.; Tan, T.; Chen, B. Lipase-Catalyzed Synthesis and Characterization of Polymers by Cyclodextrin as Support Architecture. Carbohydr. Polym. 2013, 92, 633−640. (884) Jiang, Y.; Woortman, A. J. J.; Alberda van Ekenstein, G. O. R.; Petrovic, D. M.; Loos, K. Enzymatic Synthesis of Biobased Polyesters Using 2,5-Bis(hydroxymethyl)furan as the Building Block. Biomacromolecules 2014, 15, 2482−2493. (885) Juais, D.; Naves, A. F.; Li, C.; Gross, R. A.; Catalani, L. H. Isosorbide Polyesters from Enzymatic Catalysis. Macromolecules 2010, 43, 10315−10319. (886) Kato, M.; Toshima, K.; Matsumura, S. Direct Enzymatic Synthesis of a Polyester with Free Pendant Mercapto Groups. Biomacromolecules 2009, 10, 366−373. (887) Tanaka, A.; Kohri, M.; Takiguchi, T.; Kato, M.; Matsumura, S. Enzymatic Synthesis of Reversibly Crosslinkable Polyesters with Pendant Mercapto Groups. Polym. Degrad. Stab. 2012, 97, 1415−1422. (888) Kline, B. J.; Beckman, E. J.; Russell, A. J. One-Step Biocatalytic Synthesis of Linear Polyesters with Pendant Hydroxyl Groups. J. Am. Chem. Soc. 1998, 120, 9475−9480. (889) Uyama, H.; Inada, K.; Kobayashi, S. Regioselectivity Control in Lipase-catalyzed Polymerization of Divinyl Sebacate and Triols. Macromol. Biosci. 2001, 1, 40−44. (890) Uyama, H.; Klegraf, E.; Wada, S.; Kobayashi, S. Regioselective Polymerization of Sorbitol and Divinyl Sebacate Using Lipase Catalyst. Chem. Lett. 2000, 800−801. (891) Tsujimoto, T.; Uyama, H.; Kobayashi, S. Enzymatic Synthesis of Cross-linkable Polyesters from Renewable Resources. Biomacromolecules 2001, 2, 29−31. (892) Tsujimoto, T.; Uyama, H.; Kobayashi, S. Enzymatic Synthesis and Curing of Biodegradable Crosslinkable Polyesters. Macromol. Biosci. 2002, 2, 329−335. (893) Uyama, H.; Kuwabara, M.; Tsujimoto, T.; Kobayashi, S. Enzymatic Synthesis and Curing of Biodegradable Epoxide-Containing Polyesters from Renewable Resources. Biomacromolecules 2003, 4, 211−215. (894) Yang, Z.; Zhang, X. F.; Luo, X. G.; Jiang, Q.; Liu, J.; Jiang, Z. Z. Enzymatic Synthesis of Poly(butylene-co-sebacate-co-glycolate) Copolyesters and Evaluation of the Copolymer Nanoparticles as Biodegradable Carriers for Doxorubicin Delivery. Macromolecules 2013, 46, 1743−1753. (895) Naves, A. F.; Fernandes, H. T. C.; Immich, A. P. S.; Catalani, L. H. Enzymatic Syntheses of Unsaturated Polyesters Based on Isosorbide and Isomannide. J. Polym. Sci., Part A: Polym. Chem. 2013, 51, 3881−3891. (896) Kumari, M.; Gupta, S.; Achazi, K.; Böttcher, C.; Khandare, J.; Sharma, S. K.; Haag, R. Dendronized Multifunctional Amphiphilic Polymers as Efficient Nanocarriers for Biomedical Applications. Macromol. Rapid Commun. 2015, 36, 254−261. (897) Popescu, D.; Hoogenboom, R.; Keul, H.; Möller, M. Free Radical and Nitroxide Mediated Polymerization of Hydroxy−Func2410


Chemical Reviews

Review

Polymerization of β-Alanine Esters, a Sustainable Route Towards the Formation of Poly-β-alanine. Curr. Org. Chem. 2013, 17, 682−690. (936) Stavila, E.; van Ekenstein, G. O. R.; Loos, K. Enzyme-Catalyzed Synthesis of Aliphatic-Aromatic Oligoamides. Biomacromolecules 2013, 14, 1600−1606. (937) Kato, M.; Toshima, K.; Matsumura, S. Enzyme-Catalyzed Preparation of Aliphatic Polythioester by Direct Polycondensation of Diacid Diester and Dithiol. Macromol. Rapid Commun. 2006, 27, 605− 610. (938) Rodney, R. L.; Stagno, J. L.; Beckman, E. J.; Russell, A. J. Enzymatic Synthesis of Carbonate Monomers and Polycarbonates. Biotechnol. Bioeng. 1999, 62, 259−266. (939) Matsumura, S.; Harai, S.; Toshima, K. Lipase-Catalyzed Polymerization of Diethyl Carbonate and Diol to Aliphatic Poly(alkylene carbonate). Macromol. Chem. Phys. 2000, 201, 1632−1639. (940) Jiang, Z. Z.; Liu, C.; Xie, W. C.; Gross, R. A. Controlled LipaseCatalyzed Synthesis of Poly(hexamethylene carbonate). Macromolecules 2007, 40, 7934−7943. (941) Jiang, Z. Z.; Liu, C.; Gross, R. A. Lipase-Catalyzed Synthesis of Aliphatic Poly(carbonate-co-esters). Macromolecules 2008, 41, 4671− 4680. (942) Jiang, Z. Z. Lipase-Catalyzed Copolymerization of Dialkyl Carbonate with 1,4-Butanediol and ω-Pentadecalactone: Synthesis of Poly-(ω-pentadecalactone-co-butylene-co-carbonate). Biomacromolecules 2011, 12, 1912−1919. (943) Liu, C.; Jiang, Z. Z.; Decatur, J.; Xie, W. C.; Gross, R. A. Chain Growth and Branch Structure Formation during Lipase-Catalyzed Synthesis of Aliphatic Polycarbonate Polyols. Macromolecules 2011, 44, 1471−1479. (944) Sereti, V.; Stamatis, H.; Koukios, E.; Kolisis, F. N. Enzymatic Acylation of Cellulose Acetate in Organic Media. J. Biotechnol. 1998, 66, 219−223. (945) Sereti, V.; Stamatis, H.; Pappas, C.; Polissiou, M.; Kolisis, F. N. Enzymatic Acylation of Hydroxypropyl Cellulose in Organic Media and Determination of Ester Formation by Diffuse Reflectance Infrared Fourier Transform (DRIFT) Spectroscopy. Biotechnol. Bioeng. 2001, 72, 495−500. (946) Gustavsson, M. T.; Persson, P. V.; Iversen, T.; Martinelle, M.; Hult, K.; Teeri, T. T.; Brumer, H. Modification of Cellulose Fiber Surfaces by Use of a Lipase and a Xyloglucan Endotransglycosylase. Biomacromolecules 2005, 6, 196−203. (947) Iqbal, H. M. N.; Kyazze, G.; Tron, T.; Keshavarz, T. ″One-pot″ Synthesis and Characterisation of Novel P(3HB)-Ethyl Cellulose Based Graft Composites Through Lipase Catalysed Esterification. Polym. Chem. 2014, 5, 7004−7012. (948) Cheng, H. N.; Gu, Q.-M. Enzyme-Catalyzed Modifications of Polysaccharides and Poly(ethylene glycol). Polymers 2012, 4, 1311− 1330. (949) Yang, K.; Wang, Y. J. Lipase-Catalyzed Cellulose Acetylation in Aqueous and Organic Media. Biotechnol. Prog. 2003, 19, 1664−1671. (950) Yang, K.; Wang, Y. J.; Kuo, M. I. Effects of Substrate Pretreatment and Water Activity on Lipase-Catalyzed Cellulose Acetylation in Organic Media. Biotechnol. Prog. 2004, 20, 1053−1061. (951) Yang, K.; Wang, Y. Lipase-Catalyzed Transesterification in Aqueous Medium Under Thermodynamic and Kinetic Control Using Carboxymethyl Cellulose Acetylation as the Model Reaction. Enzyme Microb. Technol. 2004, 35, 223−231. (952) Gu, Q.-M.; Cheng, H. N. Enzyme-Catalyzed Condensation Reactions for Polymer Modification. In Polym. Biocatal. Biomater.; Cheng, H. N., Gross, R. A., Eds.; ACS Symposium Series 900; American Chemical Society: Washington, D.C., 2005; pp 427−436. (953) Rajan, A.; Abraham, T. E. Enzymatic Modification of Cassava Starch by Bacterial Lipase. Bioprocess Biosyst. Eng. 2006, 29, 65−71. (954) Rajan, A.; Sudha, J. D.; Abraham, T. E. Enzymatic Modification of Cassava Starch by Fungal Lipase. Ind. Crops Prod. 2008, 27, 50−59. (955) Lu, X. X.; Luo, Z. G.; Yu, S. J.; Fu, X. Lipase-Catalyzed Synthesis of Starch Palmitate in Mixed Ionic Liquids. J. Agric. Food Chem. 2012, 60, 9273−9279.

(916) Feng, J.; Wang, H.-F.; Zhang, X.-Z.; Zhuo, R.-X. Investigation on Lipase-Catalyzed Solution Polymerization of Cyclic Carbonate. Eur. Polym. J. 2009, 45, 523−529. (917) Wu, R.; Al-Azemi, T. F.; Bisht, K. S. Functionalized Polycarbonate Derived from Tartaric Acid: Enzymatic Ring-Opening Polymerization of a Seven-Membered Cyclic Carbonate. Biomacromolecules 2008, 9, 2921−2928. (918) Namekawa, S.; Uyama, H.; Kobayashi, S.; Kricheldorf, H. R. Lipase-Catalyzed Ring-Opening Polymerization and Copolymerization of Cyclic Dicarbonates. Macromol. Chem. Phys. 2000, 201, 261−264. (919) He, F.; Jia, H. L.; Liu, G.; Wang, Y. P.; Feng, J.; Zhuo, R. X. Enzymatic Synthesis and Characterization of Novel Biodegradable Copolymers of 5-Benzyloxy-trimethylene Carbonate with 1,4-Dioxan2-one. Biomacromolecules 2006, 7, 2269−2273. (920) Al-Azemi, T. F.; Harmon, J. P.; Bisht, K. S. Enzyme-Catalyzed Ring-Opening Copolymerization of 5-Methyl-5-benzyloxycarbonyl1,3-dioxan-2-one (MBC) with Trimethylene Carbonate (TMC): Synthesis and Characterization. Biomacromolecules 2000, 1, 493−500. (921) Kaihara, S.; Fisher, J. P.; Matsumura, S. Chemo-Enzymatic Synthesis of Degradable PTMC-b-PECA-b-PTMC Triblock Copolymers and Their Micelle Formation for pH-Dependent Controlled Release. Macromol. Biosci. 2009, 9, 613−621. (922) Schwab, L. W.; Kroon, R.; Schouten, A. J.; Loos, K. EnzymeCatalyzed Ring-Opening Polymerization of Unsubstituted β-Lactam. Macromol. Rapid Commun. 2008, 29, 794−797. (923) Park, S.; Forro, E.; Grewal, H.; Fulop, F.; Kazlauskas, R. I. Molecular Basis for the Enantioselective Ring Opening of β-Lactams Catalyzed by Candida antarctica Lipase B. Adv. Synth. Catal. 2003, 345, 986−995. (924) Stavila, E.; Alberda van Ekenstein, G. O. R.; Woortman, A. J. J.; Loos, K. Lipase-Catalyzed Ring-Opening Copolymerization of εCaprolactone and β-Lactam. Biomacromolecules 2014, 15, 234−241. (925) Feng, Y. K.; Knufermann, J.; Klee, D.; Höcker, H. EnzymeCatalyzed Ring-Opening polymerization of 3(S)-Isopropylmorpholine-2,5-dione. Macromol. Rapid Commun. 1999, 20, 88−90. (926) Feng, Y. K.; Klee, D.; Keul, H.; Höcker, H. Lipase-Catalyzed Ring-Opening Polymerization of Morpholine-2,5-dione Derivatives: A Novel Route to the Synthesis of Poly(ester amide)s. Macromol. Chem. Phys. 2000, 201, 2670−2675. (927) Feng, Y.; Klee, D.; Höcker, H. Lipase Catalyzed Copolymerization of 3(S)-Isopropylmorpholine-2,5-dione and D,L-Lactide. Macromol. Biosci. 2004, 4, 587−590. (928) Feng, Y. K.; Klee, D.; Höcker, H. Lipase-Catalyzed RingOpening Polymerization of 6(S)-Methyl-morpholine-2,5-dione. J. Polym. Sci., Part A: Polym. Chem. 2005, 43, 3030−3039. (929) Wen, J.; Zhuo, R. X. Enzyme-Catalyzed Ring-Opening Polymerization of Ethylene Isopropyl Phosphate. Macromol. Rapid Commun. 1998, 19, 641−642. (930) He, F.; Zhuo, R. X.; Liu, L. J.; Jin, D. B.; Feng, J.; Wang, X. L. Immobilized Lipase on Porous Silica Beads: Preparation and Application for Enzymatic Ring-Opening Polymerization of Cyclic Phosphate. React. Funct. Polym. 2001, 47, 153−158. (931) Iwata, S.; Toshima, K.; Matsumura, S. Enzyme-Catalyzed Preparation of Aliphatic Polyesters Containing Thioester Linkages. Macromol. Rapid Commun. 2003, 24, 467−471. (932) Kato, M.; Toshima, K.; Matsumura, S. Preparation of Aliphatic Poly(thioester) by the Lipase-Catalyzed Direct Polycondensation of 11-Mercaptoundecanoic Acid. Biomacromolecules 2005, 6, 2275−2280. (933) Kato, M.; Toshima, K.; Matsumura, S. Enzymatic Synthesis of Polythioester by the Ring-Opening Polymerization of Cyclic Thioester. Biomacromolecules 2007, 8, 3590−3596. (934) Gu, Q.-M.; Maslanka, W. W.; Cheng, H. N. Enzyme-Catalyzed Polyamides and Their Derivatives. In Polymer Biocatalysis and Biomaterials II; Cheng, H. N., Gross, R. A., Eds.; ACS Symposium Series 999; American Chemical Society: Washington, D.C., 2008; pp 309−319. (935) Steunenberg, P.; Uiterweerd, M.; Sijm, M.; Scott, E. L.; Zuilhof, H.; Sanders, J. P. M.; Franssen, M. C. R. Enzyme-Catalyzed 2411


Chemical Reviews

Review

(956) Horchani, H.; Chaabouni, M.; Gargouri, Y.; Sayari, A. SolventFree Lipase-Catalyzed Synthesis of Long-Chain Starch Esters Using Microwave Heating: Optimization by Response Surface Methodology. Carbohydr. Polym. 2010, 79, 466−474. (957) Wang, Y.; Xin, J. Y.; Shi, J.; Wu, W. L.; Xia, C. G. A Kinetic Study of Starch Palmitate Synthesis by Immobilized Lipase-Catalyzed Esterification in Solvent Free System. J. Mol. Catal. B: Enzym. 2014, 101, 73−79. (958) Rajan, A.; Prasad, V. S.; Emilia Abraham, T. Enzymatic Esterification of Starch Using Recovered Coconut Oil. Int. J. Biol. Macromol. 2006, 39, 265−272. (959) Lin, R. H.; Li, H.; Long, H.; Su, J. T.; Huang, W. Q.; Wang, S. Optimization of Lipase-Catalyzed Rosin Acid Starch Synthesis by Response Surface Methodology. J. Mol. Catal. B: Enzym. 2014, 105, 104−110. (960) Lin, R. H.; Li, H.; Long, H.; Su, J. T.; Huang, W. Q. Structure and Characteristics of Lipase-Catalyzed Rosin Acid Starch. Food Hydrocolloids 2015, 43, 352−359. (961) Xu, J.; Zhou, C. W.; Wang, R. Z.; Yang, L.; Du, S. S.; Wang, F. P.; Ruan, H.; He, G. Q. Lipase-Coupling Esterification of Starch with Octenyl Succinic Anhydride. Carbohydr. Polym. 2012, 87, 2137−2144. (962) Qiao, L.; Gu, Q.-M.; Cheng, H. N. Enzyme-Catalyzed Synthesis of Hydrophobically Modified Starch. Carbohydr. Polym. 2006, 66, 135−140. (963) Chakraborty, S.; Sahoo, B.; Teraoka, I.; Miller, L. M.; Gross, R. A. Enzyme-Catalyzed Regioselective Modification of Starch Nanoparticles. Macromolecules 2005, 38, 61−68. (964) Chen, Z. G.; Zong, M. H.; Li, G. J. Lipase-Catalyzed Modification of Konjac Glucomannan. J. Appl. Polym. Sci. 2006, 102, 1335−1340. (965) Hedstrom, L. Serine Protease Mechanism and Specificity. Chem. Rev. 2002, 102, 4501−4523. (966) Patil, D. R.; Rethwisch, D. G.; Dordick, J. S. EnzymaticSynthesis of a Sucrose-Containing Linear Polyester in Nearly Anhydrous Organic Media. Biotechnol. Bioeng. 1991, 37, 639−646. (967) Park, H. G.; Chang, H. N.; Dordick, J. S. Enzymatic-Synthesis of Various Aromatic Polyesters in Anhydrous Organic-Solvents. Biocatal. Biotransform. 1994, 11, 263−271. (968) Kawai, F.; Nakadai, K.; Nishioka, E.; Nakajima, H.; Ohara, H.; Masaki, K.; Iefuji, H. Different Enantioselectivity of Two Types of Poly(lactic acid) Depolymerases Toward Poly(L-lactic acid) and Poly(D-lactic acid). Polym. Degrad. Stab. 2011, 96, 1342−1348. (969) Wilmouth, R. C.; Clifton, I. J.; Robinson, C. V.; Roach, P. L.; Aplin, R. T.; Westwood, N. J.; Hajdu, J.; Schofield, C. J. Structure of a Specific Acyl-Enzyme Complex Formed Between β-Casomorphin-7 and Porcine pancreatic Elastase. Nat. Struct. Biol. 1997, 4, 456−462. (970) Ollis, D. L.; Cheah, E.; Cygler, M.; Dijkstra, B.; Frolow, F.; Franken, S. M.; Harel, M.; Remington, S. J.; Silman, I.; Schrag, J.; Sussman, J. L.; Verschueren, K. H. G.; Goldman, A. The α/βHydrolase Fold. Protein Eng., Des. Sel. 1992, 5, 197−211. (971) Borén, L.; Martín-Matute, B.; Xu, Y.; Córdova, A.; Bäckvall, J.E. (S)-Selective Kinetic Resolution and Chemoenzymatic Dynamic Kinetic Resolution of Secondary Alcohols. Chem. - Eur. J. 2006, 12, 225−232. (972) Bruno, F. F.; Akkara, J. A.; Ayyagari, M.; Kaplan, D. L.; Gross, R.; Swift, G.; Dordick, J. S. Enzymatic Modification of Insoluble Amylose in Organic Solvents. Macromolecules 1995, 28, 8881−8883. (973) Xie, J.; Hsieh, Y.-L. Enzyme-Catalyzed Transesterification of Vinyl Esters on Cellulose Solids. J. Polym. Sci., Part A: Polym. Chem. 2001, 39, 1931−1939. (974) Cheng, H. N.; Gu, Q.-M.; Qiao, L. Reactions of Enzymes with Non-substrate Polymers. In Polymer Biocatalysis Biomaterials; Cheng, H. N., Gross, R. A., Eds.; ACS Symposium Series 900; American Chemical Society: Washington, D.C., 2005; pp 267−278. (975) Carvalho, J.; Goncalves, C.; Gil, A. M.; Gama, F. M. Production and Characterization of a New Dextrin Based Hydrogel. Eur. Polym. J. 2007, 43, 3050−3059. (976) Bordusa, F. Proteases in Organic Synthesis. Chem. Rev. 2002, 102, 4817−4867.

(977) Uyama, H.; Fukuoka, T.; Komatsu, I.; Watanabe, T.; Kobayashi, S. Protease-Catalyzed Regioselective Polymerization and Copolymerization of Glutamic Acid Diethyl Ester. Biomacromolecules 2002, 3, 318−323. (978) Li, G.; Vaidya, A.; Xie, W.; Gao, W.; Gross, R. A. EnzymeCatalyzed Oligopeptide Synthesis: Rapid Regioselective Oligomerization of L-Glutamic Acid Diethyl Ester Catalyzed by Papain. In Polymer Biocatalysis and Biomaterials II; Cheng, H. N., Gross, R. A., Eds.; ACS Symposium Series 999; American Chemical Society: Washington, D.C., 2008; pp 294−308. (979) Narai-Kanayama, A.; Koshino, H.; Aso, K. Mass Spectrometric and Kinetic Studies on Slow Progression of Papain-Catalyzed Polymerization of L-Glutamic Acid Diethyl Ester. Biochim. Biophys. Acta, Gen. Subj. 2008, 1780, 881−891. (980) Padden, F. J.; Keith, H. D. Crystalline Morphology of Synthetic Polypeptides. J. Appl. Phys. 1965, 36, 2987−2995. (981) Narai-Kanayama, A.; Shikata, Y.; Hosono, M.; Aso, K. High level production of bioactive di- and tri-tyrosine peptides by proteasecatalyzed reactions. J. Biotechnol. 2010, 150, 343−347. (982) Qin, X.; Xie, W. C.; Su, Q.; Du, W. Z.; Gross, R. A. ProteaseCatalyzed Oligomerization of L-Lysine Ethyl Ester in Aqueous Solution. ACS Catal. 2011, 1, 1022−1034. (983) Qin, X.; Xie, W. C.; Tian, S.; Ali, M. A.; Shirke, A.; Gross, R. A. Influence of N-ε-Protecting Groups on the Protease-Catalyzed Oligomerization of L-Lysine Methyl Ester. ACS Catal. 2014, 4, 1783−1792. (984) Baker, P. J.; Numata, K. Chemoenzymatic Synthesis of Poly(Lalanine) in Aqueous Environment. Biomacromolecules 2012, 13, 947− 951. (985) Baker, P. J.; Patwardhan, S. V.; Numata, K. Synthesis of Homopolypeptides by Aminolysis Mediated by Proteases Encapsulated in Silica Nanospheres. Macromol. Biosci. 2014, 14, 1619−1626. (986) Ageitos, J. M.; Baker, P. J.; Sugahara, M.; Numata, K. Proteinase K-Catalyzed Synthesis of Linear and Star Oligo(Lphenylalanine) Conjugates. Biomacromolecules 2013, 14, 3635−3642. (987) Li, G.; Raman, V. K.; Xie, W. C.; Gross, R. A. ProteaseCatalyzed Co-oligomerizations of L-Leucine Ethyl Ester with LGlutamic Acid Diethyl Ester: Sequence and Chain Length Distributions. Macromolecules 2008, 41, 7003−7012. (988) Fagerland, J.; Finne-Wistrand, A.; Numata, K. Short One-Pot Chemo-Enzymatic Synthesis of L-Lysine and L-Alanine Diblock Cooligopeptides. Biomacromolecules 2014, 15, 735−743. (989) Uemura, T.; Fujimori, M.; Lee, H. H.; Ikeda, S.; Aso, K. Polyethylene-glycol Modified Papain Catalyzed Oligopeptide Synthesis from the Esters of L-Aspartic and L-Glutamic Acids in Benzene. Agric. Biol. Chem. 1990, 54, 2277−2281. (990) Matsumura, S.; Tsushima, Y.; Otozawa, N.; Murakami, S.; Toshima, K.; Swift, G. Enzyme-Catalyzed Polymerization of LAspartate. Macromol. Rapid Commun. 1999, 20, 7−11. (991) Soeda, Y.; Toshima, K.; Matsumura, S. Sustainable Enzymatic Preparation of Polyaspartate Using a Bacterial Protease. Biomacromolecules 2003, 4, 196−203. (992) Hiraishi, T.; Masuda, E.; Miyamoto, D.; Kanayama, N.; Abe, H.; Maeda, M. Enzymatic Synthesis of Poly(α-ethyl β-aspartate) by Poly(ethylene glycol) Modified Poly(aspartate) Hydrolase-1. Macromol. Biosci. 2011, 11, 187−191. (993) Numata, K.; Baker, P. J. Synthesis of Adhesive Peptides Similar to Those Found in Blue Mussel (Mytilus edulis) Using Papain and Tyrosinase. Biomacromolecules 2014, 15, 3206−3212. (994) Kumar, D.; Bhalla, T. C. Microbial Proteases in Peptide Synthesis: Approaches and Applications. Appl. Microbiol. Biotechnol. 2005, 68, 726−736. (995) Kuhn, D.; Durrschmidt, P.; Mansfeld, J.; Ulbrich-Hofmann, R. Boilysin and Thermolysin in Dipeptide Synthesis: a Comparative Study. Biotechnol. Appl. Biochem. 2002, 36, 71−76. (996) Rizo, J.; Gierasch, L. M. Constrained Peptides - Models of Bioactive Peptides and Protein Substructures. Annu. Rev. Biochem. 1992, 61, 387−418. 2412


Chemical Reviews

Review

(997) Bille, V.; Ripak, C.; van Aasche, I.; Forni, I.; Degelaen, L.; Searso, A. A Semi-enzymatic Synthesis of Somatostatin. In Proceeding of 21st European Peptide Symposium; Giralt, E., Andreu, D., Eds.; ESCOM: Leiden, the Netherlands, 1991; pp 253−254. (998) Kobayashi, S. Artificial Urushi. J. Adhesion Soc. Jpn. 2013, 49, 16−23.

2413


Enzymes as Green Catalysts for Precision Macromolecular Synthesis

Recommend Documents