- freely available
Polymers 2012, 4(1), 759-793; doi:10.3390/polym4010759
Abstract: Biocatalysis is propagating into practically every area of organic chemistry, amongst them radical polymerizations. A review of the recent developments of this dynamic and quickly evolving area of research is presented together with a critical evaluation of its potential to yield novel polymers and/or environmentally more benign synthetic procedures.
Great aspirations have been put on biocatalysts to make the chemical industry environmentally more benign and sustainable [1,2,3]. Indeed, more and more examples impressively demonstrate the potential of White Biotechnology to reduce resource consumption and waste generation [1,4,5,6]. However, most of the ‘landmark’ examples deal with the synthesis of small, chiral molecules useful for pharmaceuticals etc. Traditionally, the wastes generated in this industry are enormous, leaving room for significant improvements . The global impact, however, is limited by the comparably low production volumes.
In contrast, polymers are bulk products produced in huge annual quantities. Here, though the saving potential might be lower due to highly optimized ‘traditional’ production schemes, even comparably small improvements may have a dramatic global impact. Hence, it is not astonishing that biocatalysis is also enjoying growing interest in the field of polymers . One field that has expanded tremendously in the past two decades is the enzyme-initiated radical polymerization of aromatic and vinyl monomers [8,9,10,11,12]. In this contribution, we summarize and critically evaluate the past and recent developments in this field also giving an outlook of the future perspectives.
2. Mechanism(s) of Enzyme Initiated Radical Polymerizations
Two enzyme classes dominate the field of biocatalytic radical polymerization: peroxidases (EC 1.11.1) and laccases (EC 126.96.36.199). Though quite different with respect to catalytic mechanism and active site structure, both enzyme classes predominantly catalyze hydrogen abstraction reactions yielding radical species to initiate the polymerization reaction. Other enzyme classes such as oxidases or lipoxygenases play only a minor role [13,14,15].
2.1. Peroxidase-Initiated Polymerizations
By far the most popular catalysts for the enzymatic initiation of radical polymerization (both of aromatic and vinyl monomers) are the so-called heme peroxidases. Particularly, the peroxidases from horseradish (HRP) and soybean (SBP) have been used most frequently.
During the catalytic mechanism (Figure 2) the water ligand (intermediate 1) is substituted by hydrogen peroxide (or other organic hydroperoxides) resulting in a peroxo complex (intermediate 2). Upon heterolytic cleavage of the O-O bond the so-called compound I (Cpd-I) is formed. Cpd-I returns into the resting state by two individual hydrogen abstractions from the reducing substrates (In-H) resulting in the formation of two radical species (In·) which then initiate the polymerization process. Overall, peroxidases function as ‘electron relays’ coupling a two electron transfer step (reduction of peroxides to water or alcohols, respectively) to two subsequent single electron transfer steps.
H2O2 plays an ambivalent role in the peroxidase-initiated polymerization reaction. On the one hand, it serves as oxidant and therefore is essential for the catalytic action. On the other hand, if [H2O2] is too high, polymerization is inhibited twofold: (1) H2O2 is a known inactivator of heme [18,19]. It was proposed that in the presence of an excess of H2O2 Cpd-II reacts with another equivalent of H2O2 instead of returning to the resting state. The resulting Cpd-III decomposes along various pathways leading to irreversible enzyme inactivation . (2) A catalase-like activity of peroxidases  at high [H2O2] results in the generation of O2 thereby inhibiting the chemical polymerization reaction. This effect has been reported especially for the polymerization of vinyl monomers in form of a lag-phase. During this lag phase, remaining O2 is consumed, probably by reaction with In. The resulting superoxide anion (O2·−) disproportionates quickly thereby overall accounting for a futile consumption of H2O2 and radical quenching. It should, however, be emphasized that the exact mechanism remains to be elucidated. Polymerization only proceeds if O2 formed by the catalase-activity has been consumed .
The usual way of avoiding excess H2O2 is to add H2O2 in several portions, which turns out to be tedious and also bears the danger of leading to a certain degree of irreproducibility. Therefore, a range of in situ H2O2 generation methods have been proposed, [22,23,24] which might be useful also for peroxidase-initiated polymerizations. Indeed Uyama et al. and others reported a bienzymatic system for the polymerization of phenols comprising peroxidase as the initiation catalyst and glucose oxidase as in situ H2O2 generation catalyst (Figure 3) [25,26].
Interestingly, this system, despite its simplicity, has not found broad proliferation. Possibly, inhibitory effects of O2 are quite severe in practice.
A rather unusual in situ generation of peroxo species was found accidentally by Samuelson and coworkers . They found that a dioxane stabilizer (acetyl acetal) spontaneously hydrolyzed under the reaction conditions and was oxidized to peracetic acid, which then served as oxidant to initiate the polymerization of aniline (Figure 4).
Nevertheless, portionwise addition of H2O2 remains the most popular procedure to promote peroxidase-initiated polymerizations.
2.2. Laccase-Initiated Polymerization
Laccases (E.C. 188.8.131.52) belong to the so-called blue-copper oxidases predominantly found in fungi but also in plants and insects [28,29,30,31]. Laccases catalyze hydrogen abstraction reactions from phenolic and related substrates resulting in corresponding phenoxy radicals. They contain four copper ions classified in one T1 copper ion and a T2/T3 cluster. It has been shown that the T1 site is the primary redox center accepting electrons from the electron donors. Thus, the fully oxidized laccase is transformed via four successive, fast single electron transfer (SET) steps into the fully reduced laccase. Molecular oxygen interacts with the fully reduced (T2/T3) cluster via a fast 2-electron-transfer process. The resulting peroxide is tightly bound so that release of H2O2 prior to the second 2-electron-transfer is efficiently prevented. As a result, the fully oxidized form of laccases comprising a μ3-oxo-bridged trinuclear (T2/T3) site is formed. This structure is thermodynamically relatively stable and provides the driving force for the overall process and also provides an efficient electron transfer bridge to re-generate the fully reduced state (Figure 5).
Interestingly enough, the number of publications dealing with laccase-initiated polymerization falls back significantly behind peroxidase-initiated polymerization studies. This is astonishing insofar as the laccase-based systems appear to be significantly easier. One apparent advantage is that laccases utilize molecular oxygen instead of hydrogen peroxide as oxidant, but inhibition may prevent widespread use.
2.3. Laccase-/Peroxidase-Mediator-Systems (LMS/PMS)
Often direct oxidation of the monomers is not possible due to unfavorable steric interaction with the enzymes’ active sites and/or due to unfavorable redox potentials. In such cases, initiation of the polymerization reaction may take place by so-called laccase- or peroxidase mediator systems (LMS or PMS, respectively). Here, enzymatic chain initiation does not proceed directly on a monomer molecule but indirectly via a small, oxidizable mediator (Figure 6). Two mechanisms can be distinguished: First, the mediator can serve as radical transfer catalyst without being incorporated into the final product. Phenothiazenes, for example, have been used to initiate the polymerization of sterically demanding phenols, [32,33] or ABTS to facilitate pyrrole polymerization [34,35]. Also transition metals (e.g., Mn2+/3+)  or polyoxometallates  have been reported. Second, the mediator radical itself can initiate chain growth thereby being incorporated into the polymer (see Section 2.4). This mechanism is the predominant pathway for the polymerization of vinyl monomers with β-diketones representing the mediators of choice (vide infra) but may also be applicable for the polymerization of non-phenolic aromatics such as thiophenes .
2.4. Other Enzyme Systems/Miscellaneous
Using laccases or heme-enzymes such as catalase of HRP, an ATRP-like polymerization was achieved with ascorbic acid as reductant and α-bromoesters as initiators. Despite the very early stage of development (particularly further mechanistic studies clarifying the initiation reaction are needed), this approach represents already now a very promising biocatalytic alternative to the existing toolbox . Exciting further developments are expected here in the near future. Noteworthy, this approach also represents the first example(s) of a reductively initiated radical polymerization.
Ximenes and coworkers reported on the H2O2-independent Acac oxidation by HRP in the presence of molecular oxygen . Intermediate formation of Acac. and superoxide was postulated, which might potentially be exploited for the initiation of radical polymerizations. If feasible, such a pathway might open up new possibilities while circumventing the limitations of H2O2-promoted chain initiation reactions mentioned above. It remains to be shown whether such a mechanism might be fruitfully exploited for peroxidase-initiated polymerization reactions.
3. Controlling the Structure of the Polymers
Unlike most enzymatic transformations, oxidoreductase-initiated radical polymerizations lack any influence of the biocatalyst on the structure of the final product. Hence, if control over the structure and morphology of the polymer formed is desired, this has to be achieved with a mechanism which does not directly involve the enzyme. In the following some of the strategies generally applied are discussed.
3.1. Controlling Vinyl Polymerization
The major concern in vinyl polymerization is to control the polymer weight and polydispersity of the final product. In this respect, most studies have focused on the influence of biocatalyst-, mediator-, and oxidant concentration.
The biocatalyst concentration directly correlates with the chain initiation rate and thereby influences the number of growing chains in the reaction mixture and competing for the remaining monomers. As a rule of thumb, the higher the biocatalyst concentration the lower the average polymer weight, as demonstrated for peroxidases  and laccases . At very high enzyme concentration, the initiation rate can be so dominant that only oligomers are formed. Under denaturizing conditions, too low enzyme concentrations manifest in low conversions due to enzyme inactivation .
Like enzyme concentration, choice and concentration of the mediator can have a major impact on the polymer properties. This was demonstrated by Kaplan and coworkers on the HRP-initiated polymerization of styrene and by Marechal and coworkers for the polymerization of acrylamide (Table 1) [47,48].
|Initiator||Yield [%]||MW [×10−3 g·mol−1]||PD|
|Styrene polymerization |
|Acrylamide polymerization |
So far, detailed studies clarifying the influence of ß-diketone structure on enzyme activity are missing. Most likely, both steric and electronic effects influence the rate of the enzymatic initiation reaction as well as the rate of the chain growth (at least at an early stage of the chain growth).
The state of the keto-enol equilibrium should determine the actual substrate concentration. Hence, high pH values should be beneficial as well as using elevated substrate concentrations. However, these measures may also be counteracted by a decreasing activity of the biocatalyst under these conditions. Principally, electron withdrawing substituents should increase the enzyme rate by shifting the keto-enol equilibrium and lowering the redox potential on the one hand. On the other hand, the resulting radical might be too stable due to very low lying SOMO energy.
Obviously, the ratio of monomer to initiator has a strong influence on the average size of the resulting polymer. The number of growing polymer chains, competing for the remaining monomer reservoir, increases with initial initiator concentration (Figure 8) .
There is an ongoing discussion about the necessity of ß-diketones to initiate the polymerization reaction. Earlier reports on ß-diketone-free polymerization of acrylamide [13,50] could not be reproduced by others [48,51]. One potential solution of this apparent discrepancy may be found in the very high H2O2 concentration in the mediator-free polymerizations resulting in ‘unusual’ heme-iron species . Thus, highly reactive peroxidase-species might initiate the polymerization reaction by direct H-atom abstraction from the monomer. Alternatively, oxidative degradation of heme resulting of Fe-release into the reaction medium might account for catalytic, Fenton-like generation of reactive oxygen species (ROS) initiating the polymerization reaction.
Kobayashi and coworkers reported on the mediator-free direct polymerization of acryl amide using the laccase from Pycnoporus coccineus suggesting a simplified polymerization scheme . Using the laccase from Myceliophthora thermophilia, this was not achieved [14,45] and a follow-up study validating Kobayashi’s initial study is still missing.
Particularly when using laccases, molecular oxygen plays an ambivalent role. On the one hand, O2 serves as stoichiometric oxidant to initiate the polymerization reaction. On the other hand, excess O2 can also efficiently quench the radical polymerization [45,53].
Recently, Nieto et al. reported on an interesting approach to balance the in situ O2 concentration for laccase-initiated polymerization by co-application of glucose oxidase as additional O2-consuming reaction .
3.2. Controlling Polymerization of Aromatics
In contrast to the above-mentioned polymerization of vinyl monomers, regioselectivity also plays an important role in the polymerization of phenols and anilines. Hence, many efforts have been directed towards gaining control over the chemo- and regioselectivity of the enzymatic polymerization of phenols and anilines.
Also due to the poor aqueous solubility of most aromatic substrates, the influence of co-solvents is frequently addressed [55,56]. In some cases significant control over the polymer size and polydispersity has been achieved with organic solvents. For example, Dordick et al. observed changes in the average polymer weight of poly(phenol) from 1,000 to over 26,000 g·mol−1 upon varying the 1,4-dioxane content in the polymerization reactions . Oguchi et al. reported on the HRP-initiated polymerization of phenol in aqueous methanol solutions yielding number average molecular weights up to 5200 g·mol−1 . Mita et al. have investigated various polar solvents and their influence on the chemoselectivity of the peroxidase-initiated polymerization of phenols (Figure 9) [59,60]. Also the hydrophobicity of the starting material had an influence on the C-C vs. C-O selectivity: the higher the hydrophobicity, the higher was the C-C- selectivity. Later on the same authors substantiated these results also using laccase as catalyst [61,62].
In addition to classical organic solvents, also ionic liquids have received some attention as cosolvents to increase the solubility of the phenolic monomers. Interestingly, while Sgalla et al. . reported the exclusive formation of dimers using HRP in [BMIM][BF4], Eker et al. later on reported on the generation of comparably large polymers using the same IL as cosolvent . Obviously, more research will be necessary to fully understand the effect of ILs as cosolvents to support enzymatic polymerizations .
Ionic liquids can also be used as liquid support for enzyme immobilization as demonstrated by Rumbau et al. . HRP immobilized in [BMIM][PF6] was applied in an emulsion polymerization of aniline (using dodecylbenzensulfonate as anionic template, vide infra). After the reaction, the aqueous product phase was separated from the enzyme-containing IL which could be reused at least 5 times without significant changes in the product properties (Figure 10).
Of course, (almost) non-aqueous reaction media would solve the solubility issue of phenols. Furthermore, anhydrous conditions can significantly alter the outcome of a given enzyme reaction. For example, as early as 1986, Dordick et al. reported that HRP catalyzes the depolymerization of lignin model compounds in 95% dioxane whereas under aqueous conditions no depolymerization activity was detectable [57,70]. Of course the solubility of native polypeptides in non-aqueous reaction media is practically zero, leading to severe diffusion limitations when using immobilized biocatalysts. However, under certain conditions enzymes can be made organosoluble e.g., by chemical modification (acetylation), [57,70] or ion pairing with anionic surfactants  or calixarenes . For example, Li and coworkers could make HRP soluble in isooctane by ion pairing it with the anionic surfactant aerosol OT (AOT) (Figure 11). The resulting organosoluble HRP was used in the tBuOOH-driven polymerization of hexylphenol .
There are a number of investigations in which templates have been used for controlling the chemical structure of polymers synthesized by oxidoreductase-initiated polymerization. Very recently, Walde and Gou gave an excellent overview over the present state of the art , which is why here only a few representative examples are discussed.
Poly(styrenesulfonate) (PSS) represents one of the first an most widely applied templates to control polymerizations. Special interest has been paid to the polymerization of aniline to the conducting polymer form (emeraldine). In the absence of a suitable template aniline polymerizes with poor regioselectivity yielding complex, insoluble and non conducting polymers. However, e.g., in the presence of negatively charged PSS (and in the presence of protonated aniline, i.e., at suitable pHs) formation of the conducting emeraldine form is strongly favored (Figure 12) [27,73,74,75,76,77,78,79].
Other negatively charged polymers  such as poly(vinylphosphonic acid)  or DNA [82,83,84] have been reported as templates. The templates promote the formation of the electrically conducting emeraldine form of poly(aniline) and significant branching (as in case of in the absence of the templating agent) does not occur. The template is thought to at least partially lead to a proper alignment of the monomers and the growing chain; another role of the negatively charged template is to serve as counter ion (dopant) influencing the charge and conformation of the polyaniline and thereby its electronic properties. In addition, anionic surfactants under emulsion conditions have been reported as templates [85,86]. Using PSS as template also proved to be suitable to produce poly(pyrroles) with improved conductiveness .
Templated polymerization of phenols can be achieved also using uncharged templates such as poly(ethylene glycol) and others [88,89,90,91,92]. Probably due to H-bond-mediated phenol-template interactions, a certain degree of regioselectivity favoring phenylene connections over oxyphenylenes was observed [89,90,91]
Interestingly, using carbon nanotubes as polymerization templates, Liu and coworkers observed oxyphenylene selectivity in the polymerization of phenol .
Another very interesting templating effect was reported by Xu et al. who caffeic acid onto a p-aminothiophenol-modified gold surface . Oriented adsorption of the phenolic monomer was achieved via inonic and H-bond interactions of the substrate with the amino-modified surface. As a result, the laccase- or HRP-initiated polymerization proceeded with very high phenylene selectivity whereas under ‘normal aqueous’ polymerization conditions a mix of phenylene and oxyphenylene connections was observed (Figure 13).
3.2.3. Substrate Engineering
Also the structure of the monomers themselves can significantly influence the selectivity of the polymerization reaction. For example Mita et al.  showed that the phenylene content of poly(p-substituted phenols) increased with the hydrophobicity of the substituent. Blocking the o-position of aniline enabled selective polymerization of anilines yielding conductive emeraldines without the need for a template .
Polymerization of catechols and hydroquinones can be tedious due to facile oxidation to the corresponding quinones. This can be overcome by blocking of one of the OH-functionalities as demonstrated by Dordick and coworkers (Figure 15) . As a result, not only was quinine formation efficiently suppressed but also a high level of o-phenylene selectivity achieved. In a similar approach 4-amino phenol was selectively polymerized by reversible blocking of the amino group via imine formation .
4. Selected Examples
There is an enormous amount of structurally diverse phenols, anilines and other aromatic monomers, which have been subjected to enzyme-initiated radical polymerization. A representative, but by far not exhaustive selection of the different monomers is given in Figure 16 and Figure 17. For a detailed discussion about polymer properties and potential uses, the interested reader is referred to some excellent previous overviews [8,9,10,11,12].
Flavonoids represent a class of natural secondary metabolites that have found numerous applications in human health products. Especially their antioxidant and radical scavenging activity but also anti microbial properties make this product class interesting. The group around Kobayashi has investigated the production and properties of polymerized polyphenols . Interestingly, in the majority of these examples, the antioxidant activity (and quite frequently also other health properties) are significantly increased as compared to the monomers. A unifying explanation for this observation is missing so far.
Early reports dealing with modified flavonids concern the direct laccase- and/or peroxidase-mediated polymerization of the monomers. More recently, oxidative conjugation to other polymers has gained considerable interest (Figure 18). Table 2 gives an overview over the polymeric products reported.
As mentioned above, the oxidative grafting of flavonoids to polymers is enjoying increased attention especially aiming at functionalized textiles or food applications .
One of the first examples for this was reported by Kobayashi and coworkers who oxidatively grafted poly(allylamine) with catechin (Figure 19) , which was followed up more recently by Rao and coworkers [132,133].
Instead of poly(allylamine) also natural polymeric support can be used resulting in a composite with similarly superior antioxidant activity as compared to the catechin monomer. For example gelatin was grafted with catechin . Wool can also be grafted e.g., with gallates resulting in composites with antioxidant and antimicrobial activity . When using gallic acid esters, increased water repellence was achieved. Chitosan-composites with gallic acid (derivates) [136,137,138,139], ferulic acid , flavonoids  exhibit increased antioxidant activity. An interesting extension of this concept is to use amine-functionalized inorganic supports .
A practical application of this grafting technology may be in the textile and food industry yielding products with antimicrobial activity and modified rheological properties [130,138,139,143,144,145,146].
4.2. Polymer Modification
Oxidoreductase-initiated covalent linkage of phenols has also received considerable interest for the cross-linking of polymers to form biocompatible hydrogels. For example, proteins presenting tyrosine moieties at their surface can be gelled in the presence of laccases or peroxidases/H2O2 (Figure 20) [147,148,149,150].
Next to proteins also natural polymers exhibiting phenolic side chains can be gelled enzymatically. For example pectins contain some ferulic acid which can be used for oxidative crosslinking of the pectin chains for irreversible gellation [151,152,153]. However, even if no phenolic side chain is present in the polymer, it may be introduced chemically. In that respect, tyramine has gained some popularity e.g., for the crosslinking of carboxymethyl cellulose , alginate , hyaluronic acid , dextran  or artificial polymers such as Tetronic .
NH2-containing polymers (e.g., chitosan) can be modified with phenolic carboxylic acids via amidation  or directly act in Michael-fashion on enzyme-generated quinones (Figure 21) [146,160]. For example, tyrosinases generate reactive quinones from phenols, which can react with chitosan [141,145,160,161]. Similarly, reactive quinones can be generated using laccases [140,146] of peroxidases [136,139].
A very creative application of the system HRP/H2O2/Acac for graft polymerization of acryl amide on starch was reported recently by Biswas and coworkers . A biocatalytic alternative route to the established cerium(IV)-based routes to poly(acrylate)-grafted starch with applications as superabsorbersor performance additives in paper making or textile sizing was reported. Under non-optimized conditions grafting efficiencies of up to 65% are reported. The authors suggest covalent attachment of the poly(acrylamide) chain to the starch backbone via H-abstraction at the glycosidic C-atom (Figure 22).
Poly(acrylamide) grafting onto lignin has been reported in the presence of organic hydroperoxides [162,163]. The proposed mechanism (Figure 23.) comprises laccase-catalyzed H-atom abstraction from phenolic lignin residues. The resulting phenoxy radicals interact with the peroxides generation alkoxy- and hydroperoxy-radicals, which are supposed to initiate the acrylamide polymerization process. Grafting of poly(acrylamide) onto lignin is proposed to occur through recombination of the respective radicals.
4.3. Artificial Urushi
Urushiols are a class of natural alkyl-substituted phenols and catechols (Figure 24), which in the presence of laccase can polymerize to form Urushi, a natural lacquer used e.g., in traditional Japanese artwork.
Due to its potential as ‘fully renewable and benign’ alternative to chemical formaldehyde-phenol resins, the enzymatic polymerization of urushiols has been thoroughly investigated by the Kobayashi group [164,165,166,167].
Synthetic, tailored urushiols have also been proposed by lipase-catalyzed acylation of benzylic OH-groups followed by peroxidase- or laccase-initiated polymerization of the phenol/catechols moiety (Figure 25).
4.4. Selected Examples for Chemoselective Polymerization
One advantage of oxidoreductase-catalysis that is frequently mentioned is its high selectivity. As mentioned above, so far no examples of stereospecificity have been reported in enzyme initiated polymerization, and are not very likely to come up in the near future. However, there is a range of examples demonstrating chemoselective radical formation, which will be shortly outlined below.
The selective polymerization of bifunctional monomers containing vinyl and phenol moieties. Generally, laccases and peroxidases act on the phenol part selectively leading to vinyl-substituted poly(phenols) which then are available for further modification (Figure 26) [68,101,171].
A very nice example for the potential of biocatalysis as complementary tool to established chemocatalysis was reported using m-ethinylphenol as monomer . Subjecting this compound to HRP/H2O2, poly(phenol) formation was the only reaction observed wile leaving the ethinyl group untouched. In contrast, CuI catalysts lead to dimerization via the ethinyl function (Figure 27).
Another example for chemoselectivity of enzymatic polymerization was reported by Bilici et al. . Here, an aldehyde-bearing catechols was polymerized enzymatically without oxidizing the aldehyde moiety.
4.5. Phenol Detoxification
A good portion of the laccase- and peroxidase literature deals with bioremediation and pollutant control [130,173,174,175]. The removal of phenolics is rather straightforward as these compounds are generally oxidized by the laccases or peroxidases directly . In most of the cases, the detoxification mechanism is based on the formation of insoluble oligomeric or polymeric products which can then be removed by filtration.
Hence, various cresols and derivates  can be oxidatively polymerized and thereby reduced in toxicity. Likewise halophenols [178,179,180,181,182] bisphenol A, [183,184,185] various hormones and so-called endocrine disrupting compounds [186,187,188] or even TNT [189,190] have been reported. Non-phenolic compounds may be ‘inactivated’ using the LMS or PMS strategy [36,191,192,193,194,195,196,197,198].
Enzymatic reactions are enjoying increasing attention as alternatives to classical chemical routes. ‘The hallmark of enzyme catalysis is its superior catalytic power and high selectivity under mild reaction conditions’ . Furthermore, enzyme catalysis generally exhibits high chemo-, regio-, and enantioselectivity. In the context of oxidoreductase initiated radical polymerizations these advantages come into play only to a limited extent. Mostly the role of the enzyme is confined to the generation of a starter radical. Thus, enzyme-initiated polymerizations comprise all characteristics of a classical chemical polymerization.
Admittedly, enzymatic routes offer various handles to control polymer weight with enzyme concentration being the most efficient one. Furthermore, enzymes are derived from renewable feedstock and therefore can be considered more benign than many classical initiators. Overall, enzyme-initiated polymerization represents a promising alternative approach to the established chemical routes. Implementation on industrial scale will not occur on a short-term due to cost limitations but may eventually result in significantly greener production routes (though a full life cycle analysis will be necessary as enzymatic reactions are not per se greener than their chemical counterparts). Furthermore, it represents another example that enzyme catalysis is not confined to its traditional playgrounds such as chiral molecules.
This work was supported by the Marie Curie ITN ‘Biotrains’ (grant agreement number 238531) and the Deutsche Bundesstiftung Umwelt, ChemBioTec (grant agreement number 13253).
- Wenda, S.; Illner, S.; Mell, A.; Kragl, U. Industrial biotechnology—The future of green chemistry? Green Chem. 2011, 13, 3007–3047. [Google Scholar] [CrossRef]
- Hollmann, F.; Arends, I.W.C.E.; Buehler, K.; Schallmey, A.; Buhler, B. Enzyme-mediated oxidations for the chemist. Green Chem. 2011, 13, 226–265. [Google Scholar] [CrossRef]
- Hollmann, F.; Arends, I.W.C.E.; Holtmann, D. Enzymatic reductions for the chemist. Green Chem. 2011, 13, 2285–2313. [Google Scholar] [CrossRef]
- Kuhn, D.; Kholiq, M.A.; Heinzle, E.; Bühler, B.; Schmid, A. Intensification and economic and ecological assessment of a biocatalytic oxyfunctionalization process. Green Chem. 2010, 12, 815–827. [Google Scholar] [CrossRef]
- Thum, O.; Oxenbøll, K.M. Biocatalysis: A sustainable process for production of cosmetic ingredients. In Proceedings of the International Federation of Societies of Cosmetic Chemists, IFSCC Congress 2006, Osaka, Japan, 16–19 October 2006.
- Henderson, R.K.; Jiminez-Gonzalez, C.; Preston, C.; Constable, D.J.C.; Woodley, J.M. EHS & LCA assessment for 7-ACA synthesis A case study for comparing biocatalytic & chemical synthesis. Ind. Biotechnol. 2008, 4, 180–192. [Google Scholar] [CrossRef]
- Sheldon, R.A. E factors, green chemistry and catalysis: An odyssey. Chem. Commun. 2008, 3352–3365. [Google Scholar] [CrossRef]
- Kobayashi, S.; Makino, A. Enzymatic polymer synthesis: An opportunity for green polymer chemistry. Chem. Rev. 2009, 109, 5288–5353. [Google Scholar] [CrossRef]
- Uyama, H.; Kobayashi, S.; Ritter, H.; Kaplan, D. Enzymatic synthesis and properties of polymers from polyphenols. In Enzyme-Catalyzed Synthesis of Polymers; Springer: Berlin, Germany and Heidelberg, Germany, 2006; Volume 194, pp. 51–67. [Google Scholar]
- Singh, A.; Kaplan, D.L. Enzyme-based vinyl polymerization. J. Polym. Environ. 2002, 10, 85–91. [Google Scholar] [CrossRef]
- Uyama, H. Artificial polymeric flavonoids: Synthesis and applications. Macromol. Biosci. 2007, 7, 410–422. [Google Scholar] [CrossRef]
- Reihmann, M.; Ritter, H. Synthesis of phenol polymers using peroxidases. In Enzyme-Catalyzed Synthesis of Polymers; Kobayashi, S., Ritter, H., Kaplan, D., Eds.; Springer-Verlag: Berlin, Germany, 2006; Volume 194, pp. 1–49. [Google Scholar]
- Derango, R.; Chiang, L.-C.; Dowbenko, R.; Lasch, J. Enzyme-mediated polymerization of acrylic monomers. Biotechnol. Technol. 1992, 6, 523–526. [Google Scholar]
- Tsujimoto, T.; Uyama, H.; Kobayashi, S. Polymerization of vinyl monomers using oxidase catalysts. Macromol. Biosci. 2001, 1, 228–232. [Google Scholar] [CrossRef]
- Kausaite, A.; Ramanaviciene, A.; Ramanavicius, A. Polyaniline synthesis catalysed by glucose oxidase. Polymer 2009, 50, 1846–1851. [Google Scholar] [CrossRef]
- Hofrichter, M.; Ullrich, R.; Pecyna, M.J.; Liers, C.; Lundell, T. New and classic families of secreted fungal heme peroxidases. Appl. Microbiol. Biotechnol. 2010, 87, 871–897. [Google Scholar] [CrossRef]
- Van Rantwijk, F.; Sheldon, R.A. Selective oxygen transfer catalysed by heme peroxidases: Synthetic and mechanistic aspects. Curr. Opin. Biotechnol. 2000, 11, 554–564. [Google Scholar] [CrossRef]
- Hanson, R.L.; Howell, J.M.; LaPorte, T.L.; Donovan, M.J.; Cazzulino, D.L.; Zannella, V.; Montana, M.A.; Nanduri, V.B.; Schwarz, S.R.; Eiring, R.F.; et al. Synthesis of allysine ethylene acetal using phenylalanine dehydrogenase from Thermoactinomyces intermedius. Enzym. Microb. Technol. 2000, 26, 348–358. [Google Scholar] [CrossRef]
- Valderrama, B.; Ayala, M.; Vazquez-Duhalt, R. Suicide inactivation of peroxidases and the challenge of engineering more robust enzymes. Chem. Biol. 2002, 9, 555–565. [Google Scholar] [CrossRef]
- Hiner, A.N.P.; Hernández-Ruiz, J.; Williams, G.A.; Arnao, M.B.; García-Cánovas, F.; Acosta, M. Catalase-like oxygen production by horseradish peroxidase must predominantly be an enzyme-catalyzed reaction. Arch. Biochem. Biophys. 2001, 392, 295–302. [Google Scholar] [CrossRef]
- Collinson, E.; Dainton, F.S.; McNaughton, G.S. The polymerization of acrylamide in aqueous solution. Part 1.The X- and g-ray initiated reaction. Trans. Faraday Soc. 1957, 53, 476–488. [Google Scholar] [CrossRef]
- Hollmann, F.; Arends, I.W.C.E.; Buehler, K. Biocatalytic redox reactions for organic synthesis: Nonconventional regeneration methods. ChemCatChem 2010, 2, 762–782. [Google Scholar] [CrossRef]
- Churakova, E.; Kluge, M.; Ullrich, R.; Arends, I.; Hofrichter, M.; Hollmann, F. Specific photobiocatalytic oxyfunctionalization reactions. Angew. Chem. Int. Ed. 2011, 50, 10716–10719. [Google Scholar]
- Perez, D.I.; Mifsud Grau, M.; Arends, I.W.C.E.; Hollmann, F. Visible light-driven and chloroperoxidase-catalyzed oxygenation reactions. Chem. Commun. 2009, 6848–6850. [Google Scholar]
- Uyama, H.; Kurioka, H.; Kobayashi, S. Novel bienzymatic catalysis system for oxidative polymerization of phenols. Polym. J. 1997, 29, 190–192. [Google Scholar] [CrossRef]
- Taboada-Puig, R.; Junghanns, C.; Demarche, P.; Moreira, M.T.; Feijoo, G.; Lema, J.M.; Agathos, S.N. Combined cross-linked enzyme aggregates from versatile peroxidase and glucose oxidase: Production, partial characterization and application for the elimination of endocrine disruptors. Bioresour. Technol. 2011, 102, 6593–6599. [Google Scholar]
- Kim, S.C.; Huh, P.; Kumar, J.; Kim, B.; Lee, J.O.; Bruno, F.F.; Samuelson, L.A. Synthesis of polyaniline derivatives via biocatalysis. Green Chem. 2007, 9, 44–48. [Google Scholar] [CrossRef]
- Kunamneni, A.; Camarero, S.; Garcia-Burgos, C.; Plou, F.; Ballesteros, A.; Alcalde, M. Engineering and applications of fungal laccases for organic synthesis. Microb. Cell Factories 2008, 7. [Google Scholar] [CrossRef]
- Witayakran, S.; Ragauskas, A.J. Synthetic applications of laccase in green chemistry. Adv. Synth. Catal. 2009, 351, 1187–1209. [Google Scholar] [CrossRef]
- Riva, S. Laccases: Blue enzymes for green chemistry. Trends Biotechnol. 2006, 24, 219–226. [Google Scholar] [CrossRef]
- Cañas, A.I.; Camarero, S. Laccases and their natural mediators: Biotechnological tools for sustainable eco-friendly processes. Biotechnol. Adv. 2010, 28, 694–705. [Google Scholar] [CrossRef]
- 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. [Google Scholar] [CrossRef]
- 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. [Google Scholar] [CrossRef]
- Song, H.K.; Palmore, G.T.R. Conductive polypyrrole via enzyme catalysis. J. Phys. Chem. B 2005, 109, 19278–19287. [Google Scholar]
- Cruz-Silva, R.; Amaro, E.; Escamilla, A.; Nicho, M.E.; Sepulveda-Guzman, S.; Arizmendi, L.; Romero-Garcia, J.; Castillon-Barraza, F.F.; Farias, M.H. Biocatalytic synthesis of polypyrrole powder, colloids, and films using horseradish peroxidase. J. Colloid Interface Sci. 2008, 328, 263–269. [Google Scholar] [CrossRef]
- Eibes, G.; Debernardi, G.; Feijoo, G.; Moreira, M.T.; Lema, J.M. Oxidation of pharmaceutically active compounds by a ligninolytic fungal peroxidase. Biodegradation 2011, 22, 539–550. [Google Scholar] [CrossRef]
- Kim, S.; Silva, C.; Evtuguin, D.V.; Gamelas, J.A.F.; Cavaco-Paulo, A. Polyoxometalate/laccase-mediated oxidative polymerization of catechol for textile dyeing. Appl. Microbiol. Biotechnol. 2011, 89, 981–987. [Google Scholar] [CrossRef]
- Nagarajan, S.; Kumar, J.; Bruno, F.F.; Samuelson, L.A.; Nagarajan, R. Biocatalytically synthesized poly(3,4-ethylenedioxythiophene). Macromolecules 2008, 41, 3049–3052. [Google Scholar]
- Sigg, S.J.; Seidi, F.; Renggli, K.; Silva, T.B.; Kali, G.; Bruns, N. Horseradish peroxidase as a catalyst for atom transfer radical polymerization. Macromol. Rapid Commun. 2011, 32, 1710–1715. [Google Scholar] [CrossRef]
- Ng, Y.H.; di Lena, F.; Chai, C.L.L. PolyPEGA with predetermined molecular weights from enzyme-mediated radical polymerization in water. Chem. Commun. 2011, 47, 6464–6466. [Google Scholar]
- Ng, Y.H.; di Lena, F.; Chai, C.L.L. Metalloenzymatic radical polymerization using alkyl halides as initiators. Polym. Chem. 2011, 2, 589–594. [Google Scholar] [CrossRef]
- Tsarevsky, N.V.; Matyjaszewski, K. “Green” atom transfer radical polymerization: From process design to preparation of well-defined environmentally friendly polymeric materials. Chem. Rev. 2007, 107, 2270–2299. [Google Scholar] [CrossRef]
- Rodrigues, A.P.; da Fonseca, L.M.; de Faria Oliveira, O.M.; Brunetti, I.L.; Ximenes, V.F. Oxidation of acetylacetone catalyzed by horseradish peroxidase in the absence of hydrogen peroxide. Biochim. Biophys. Acta 2006, 1760, 1755–1761. [Google Scholar]
- Durand, A.; Lalot, T.; Brigodiot, M.; Maréchal, E. Enzyme-mediated radical initiation of acrylamide polymerization: Main characteristics of molecular weight control. Polymer 2001, 42, 5515–5521. [Google Scholar] [CrossRef]
- Hollmann, F.; Gumulya, Y.; Toelle, C.; Liese, A.; Thum, O. Evaluation of the laccase from myceliophthora thermophila as industrial biocatalyst for polymerization reactions. Macromolecules 2008, 41, 8520–8524. [Google Scholar] [CrossRef]
- Qi, G.G.; Jones, C.W.; Schork, F.J. Enzyme-initiated miniemulsion polymerization. Biomacromolecules 2006, 7, 2927–2930. [Google Scholar] [CrossRef]
- Singh, A.; Ma, D.; Kaplan, D.L. Enzyme-mediated free radical polymerization of styrene. Biomacromolecules 2000, 1, 592–596. [Google Scholar] [CrossRef]
- Teixeira, D.; Lalot, T.; Brigodiot, M.; Marechal, E. β-diketones as key compounds in free-radical polymerization by enzyme-mediated initiation. Macromolecules 1999, 32, 70–72. [Google Scholar] [CrossRef]
- Baader, W.J.; Bohne, C.; Cilento, G.; Dunford, H.B. Peroxidase-catalyzed formation of triplet acetone and chemiluminescence from isobutyraldehyde and molecular oxygen. J. Biol. Chem. 1985, 260, 10217–10225. [Google Scholar]
- Parravano, G. Chain reactions induced by enzymic systems. J. Am. Chem. Soc. 1951, 73, 183–184. [Google Scholar] [CrossRef]
- Emery, O.; Lalot, T.; Brigodiot, M.; Maréchal, E. Free-Radical polymerization of acrylamide by horseradish peroxidase-mediated initiation. J. Polym. Sci. A 1997, 35, 3331–3333. [Google Scholar] [CrossRef]
- Ikeda, R.; Tanaka, H.; Uyama, H.; Kobayashi, S. Laccase-catalyzed polymerization of acrylamide. Macromol. Rapid Commun. 1998, 19, 423–425. [Google Scholar] [CrossRef]
- 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. [Google Scholar] [CrossRef]
- 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. [Google Scholar]
- Ayyagari, M.S.R.; Kaplan, D.L.; Chatterjee, S.; Walker, J.E.; Akkara, J.A. Solvent effects in horseradish peroxidase-catalyzed polyphenol synthesis. Enzym. Microb. Technol. 2002, 30, 3–9. [Google Scholar] [CrossRef]
- Akkara, J.A.; Ayyagari, M.S.R.; Bruno, F.F. Enzymatic synthesis and modification of polymers in nonaqueous solvents. Trends Biotechnol. 1999, 17, 67–73. [Google Scholar] [CrossRef]
- Dordick, J.S.; Marletta, M.A.; Klibanov, A.M. Polymerization of phenols catalyzed by peroxidase in nonaqueous media. Biotechnol. Bioeng. 1987, 30, 31–36. [Google Scholar] [CrossRef]
- Oguchi, T.; Tawaki, S.; Uyama, H.; Kobayashi, S. Soluble polyphenol. Macromol. Rapid Commun. 1999, 20, 401–403. [Google Scholar] [CrossRef]
- Mita, N.; Maruichi, N.; Tonami, H.; Nagahata, R.; Tawaki, S.; Uyama, H.; Kobayashi, S. Enzymatic oxidative polymerization of p-t-butylphenol and characterization of the product polymer. Bull. Chem. Soc. Jpn. 2003, 76, 375–379. [Google Scholar]
- 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. [Google Scholar]
- Mita, N.; Tawaki, S.; Uyama, H.; Kobayashi, S. Precise structure control of enzymatically synthesized polyphenols. Bull. Chem. Soc. Jpn. 2004, 77, 1523–1527. [Google Scholar] [CrossRef]
- Mita, N.; Tawaki, S.-I.; Uyama, H.; Kobayashi, S. Laccase-catalyzed oxidative polymerization of phenols. Macromol. Biosci. 2003, 3, 253–257. [Google Scholar] [CrossRef]
- 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 2007, 44, 144–148. [Google Scholar] [CrossRef]
- Eker, B.; Zagorevski, D.; Zhu, G.Y.; Linhardt, R.J.; Dordick, J.S. Enzymatic polymerization of phenols in room-temperature ionic liquids. J. Mol. Catal. B 2009, 59, 177–184. [Google Scholar] [CrossRef]
- Zaragoza-Gasca, P.; Villamizar-Gálvez, O.J.; García-Arrazola, R.; Gimeno, M.; Bárzana, E. Use of ionic liquid for the enzyme-catalyzed polymerization of phenols. Polym. Adv. Technol. 2010, 21, 454–456. [Google Scholar]
- 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. [Google Scholar] [CrossRef]
- 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. [Google Scholar] [CrossRef]
- 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. [Google Scholar] [CrossRef]
- 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. [Google Scholar] [CrossRef]
- Dordick, J.S.; Marletta, M.A.; Klibanov, A.M. Peroxidase depolymerize lignin in organic media but not in water. Proc. Natl. Acad. Sci. USA 1986, 83, 6255–6257. [Google Scholar]
- 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. [Google Scholar]
- Oshima, T.; Sato, M.; Shikaze, Y.; Ohto, K.; Inoue, K.; Baba, Y. Enzymatic polymerization of o-phenylendiamine with cytochrome c activated by a calixarene derivative in organic media. Biochem. Eng. J. 2007, 35, 66–70. [Google Scholar] [CrossRef]
- Walde, P.; Guo, Z.W. Enzyme-catalyzed chemical structure-controlling template polymerization. Soft Matter 2011, 7, 316–331. [Google Scholar] [CrossRef]
- 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. [Google Scholar]
- Nabid, M.R.; Taheri, S.S.; Sedghi, R.; Rezaei, S.J.T. Synthesis and characterization of chemiluminescent conducting polyluminol via biocatalysis. Macromol. Res. 2011, 19, 280–285. [Google Scholar] [CrossRef]
- Nabid, M.R.; Sedghi, R.; Entezami, A.A. Enzymatic oxidation of alkoxyanilines for preparation of conducting polymers. J. Appl. Polym. Sci. 2007, 103, 3724–3729. [Google Scholar] [CrossRef]
- 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-3-am nobenzeneboronic acid) prepared from enzymatic polymerization. Biomacromolecules 2007, 8, 3602–3607. [Google Scholar] [CrossRef]
- Karamyshev, A.V.; Shleev, S.V.; Koroleva, O.V.; Yaropolov, A.I.; Sakharov, I.Y. Laccase-catalyzed synthesis of conducting polyaniline. Enzym. Microb. Technol. 2003, 33, 556–564. [Google Scholar]
- Tewari, A.; Kokil, A.; Ravichandran, S.; Nagarajan, S.; Bouldin, R.; Samuelson, L.A.; Nagarajan, R.; Kumar, J. Soybean peroxidase catalyzed enzymatic synthesis of pyrrole/EDOT copolymers. Macromol. Chem. Phys. 2010, 211, 1610–1617. [Google Scholar]
- Vasil’eva, I.; Morozova, O.; Shumakovich, G.; Yaropolov, A. Synthesis of electroconductive polyaniline using immobilized laccase. Appl. Biochem. Microbiol. 2009, 45, 27–30. [Google Scholar] [CrossRef]
- Nagarajan, R.; Tripathy, S.; Kumar, J.; Bruno, F.F.; Samuelson, L. An enzymatically synthesized conducting molecular complex of polyaniline and poly(vinylphosphonic acid). Macromolecules 2000, 33, 9542–9547. [Google Scholar]
- 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. [Google Scholar]
- Nagarajan, R.; Liu, W.; Kumar, J.; Tripathy, S.K.; Bruno, F.F.; Samuelson, L.A. Manipulating DNA conformation using intertwined conducting polymer chains. Macromolecules 2001, 34, 3921–3927. [Google Scholar]
- Ma, Y.; Zhang, J.; Zhang, G.; He, H. Polyaniline nanowires on si surfaces fabricated with DNA templates. J. Am. Chem. Soc. 2004, 126, 7097–7101. [Google Scholar]
- 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. [Google Scholar]
- 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. [Google Scholar] [CrossRef]
- Bouldin, R.; Ravichandran, S.; Kokil, A.; Garhwal, R.; Nagarajan, S.; Kumar, J.; Bruno, F.F.; Samuelson, L.A.; Nagarajan, R. Synthesis of polypyrrole with fewer structural defects using enzyme catalysis. Synth. Met. 2011, 161, 1611–1617. [Google Scholar]
- 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. [Google Scholar] [CrossRef]
- Kim, Y.J.; Uyama, H.; Kobayashi, S. Enzymatic template polymerization of phenol in the presence of water-soluble polymers in an aqueous medium. Polym. J. 2004, 36, 992–998. [Google Scholar] [CrossRef]
- 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. [Google Scholar] [CrossRef]
- 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. [Google Scholar] [CrossRef]
- Bruno, F.F.; Nagarajan, R.; Stenhouse, P.; Yang, K.; Kumar, J.; Tripathy, S.K.; Samuelson, L.A. Polymerization of water-soluble conductive polyphenol using horseradish peroxidase. J. Macromol. Sci. Pure Appl. Chem. 2001, 38, 1417–1426. [Google Scholar] [CrossRef]
- 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. Pol. Chem. 2009, 47, 1627–1635. [Google Scholar] [CrossRef]
- 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. [Google Scholar]
- 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. [Google Scholar] [CrossRef]
- 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. [Google Scholar]
- Reihmann, M.H.; Ritter, H. Regioselective HRP-catalyzed polymerization of 4-amino-phenol. J. Macromol. Sci. Pure Appl. Chem. 2002, A39, 1369–1382. [Google Scholar]
- Reihmann, M.H.; Ritter, H. Oxidative copolymerization of para-functionalized phenols catalyzed by horseradish peroxidase and thermocrosslinking via Diels-Alder and (1+3) cycloaddition. Macromol. Biosci. 2001, 1, 170–176. [Google Scholar]
- Turac, E.; Sahmetlioglu, E. Oxidative polymerization of 4-[(4-phenylazo-phenyimino)-methyl]-phenol catalyzed by horseradish peroxidase. Synth. Met. 2010, 160, 169–172. [Google Scholar] [CrossRef]
- Uyama, H.; Kurioka, H.; Sugihara, J.; Komatsu, I.; Kobayashi, S. Oxidative polymerization of p-alkylphenols catalyzed by horseradish peroxidase. J. Polym. Sci. Pol. Chem. 1997, 35, 1453–1459. [Google Scholar] [CrossRef]
- 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. [Google Scholar]
- Robert, J.P.; Uyama, H.; Kobayashi, S.; Jordan, R.; Nuyken, O. First diazosulfonate homopolymer by enzymatic polymerization. Macromol. Rapid Commun. 2003, 24, 185–189. [Google Scholar] [CrossRef]
- 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. [Google Scholar] [CrossRef]
- Zaragoza-Gasca, P.; Gimeno, M.; Hernandez, J.M.; Barzana, E. Novel photoconductive polyfluorophenol synthesized by an enzyme. J. Mol. Catal. B 2011, 72, 25–27. [Google Scholar] [CrossRef]
- Bilici, A.; Kaya, Ä.; Yildirim, M.; Dogan, F. Enzymatic polymerization of hydroxy-functionalized carbazole monomer. J. Mol. Catal. B 2010, 64, 89–95. [Google Scholar] [CrossRef]
- Tonami, H.; Uyama, H.; Kobayashi, S.; Fujita, T.; Taguchi, Y.; Osada, K. Chemoselective oxidative polymerization of m-ethynylphenol by peroxidase catalyst to a new reactive polyphenol. Biomacromolecules 2000, 1, 149–151. [Google Scholar] [CrossRef]
- 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 2007, 44, 149–154. [Google Scholar] [CrossRef]
- 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. [Google Scholar] [CrossRef]
- Wang, P.; Dordick, J.S. Enzymatic synthesis of unique thymidine-containing polyphenols. Macromolecules 1998, 31, 941–943. [Google Scholar] [CrossRef]
- Durand, A.; Lalot, T.; Brigodiot, M.; Maréchal, E. Enzyme-mediated initiation of acrylamide polymerization: Reaction mechanism. Polymer 2000, 41, 8183–8192. [Google Scholar]
- Lalot, T.; Brigodiot, M.; Maréchal, E. A kinetic approach to acrylamide radical polymerization by horse radish peroxidase-mediated initiation. Polym. Int. 1999, 48, 288–292. [Google Scholar] [CrossRef]
- 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. Pol. Chem. 2008, 46, 2222–2232. [Google Scholar] [CrossRef]
- Kalra, B.; Gross, R.A. HRP-mediated polymerizations of acrylamide and sodium acrylate. Green Chem. 2002, 4, 174–178. [Google Scholar] [CrossRef]
- Shogren, R.L.; Willett, J.L.; Biswas, A. HRP-mediated synthesis of starch-polyacrylamide graft copolymers. Carbohyd. Polym. 2009, 75, 189–191. [Google Scholar] [CrossRef]
- Kalra, B.; Gross, R.A. Horseradish peroxidase mediated free radical polymerization of methyl methacrylate. Biomacromolecules 2000, 1, 501–505. [Google Scholar] [CrossRef]
- Shan, J.; Kitamura, Y.; Yoshizawa, H. Emulsion polymerization of styrene by horseradish peroxidase-mediated initiation. Colloid Polym. Sci. 2005, 284, 108–111. [Google Scholar] [CrossRef]
- Singh, A.; Roy, S.; Samuelson, L.; Bruno, F.; Nagarajan, R.; Kumar, J.; John, V.; Kaplan, D.L. Peroxidase, hematin, and pegylated-hematin catalyzed vinyl polymerizations in water. J. Macromol. Sci. A 2001, 38, 1219–1230. [Google Scholar] [CrossRef]
- Singh, A.; Kaplan, D.L. Vitamin C functionalized poly(methyl methacrylate) for free radical scavenging. J. Macromol. Sci. Part A 2004, 41, 1377–1386. [Google Scholar] [CrossRef]
- Singh, A.; Kaplan, D.L. Biocatalytic route to ascorbic acid-modified polymers for free-radical scavenging. Adv. Mat. 2003, 15, 1291–1294. [Google Scholar] [CrossRef]
- Bruno, F.F.; Trotta, A.; Fossey, S.; Nagarajan, S.; Nagarajan, R.; Samuelson, L.A.; Kumar, J. Enzymatic synthesis and characterization of poly quercetin. J. Macromol. Sci. Part A 2010, 47, 1191–1196. [Google Scholar] [CrossRef]
- Desentis-Mendoza, R.M.; Hernandez-Sanchez, H.; Moreno, A.; Emilio, R.D.C.; Chel-Guerrero, L.; Tamariz, J.; Jaramillo-Flores, M.E. Enzymatic polymerization of phenolic compounds using laccase and tyrosinase from Ustilago maydis. Biomacromolecules 2006, 7, 1845–1854. [Google Scholar]
- Ma, H.-L.; Kermasha, S.; Gao, J.-M.; Borges, R.M.; Yu, X.-Z. Laccase-catalyzed oxidation of phenolic compounds in organic media. J. Mol. Catal. B 2009, 57, 89–95. [Google Scholar] [CrossRef]
- Kurisawa, M.; Chung, J.E.; Uyama, H.; Kobayashi, S. Laccase-catalyzed synthesis and antioxidant property of poly(catechin). Macromol. Biosci. 2003, 3, 758–764. [Google Scholar] [CrossRef]
- Bruno, F.F.; Nagarajan, S.; Nagarajan, R.; Kumar, J.; Samuelson, L.A. Biocatalytic synthesis of water-soluble oligo(catechins). J. Macromol. Sci. 2005, A42, 1547–1554. [Google Scholar]
- Anthoni, J.; Chebil, L.; Lionneton, F.; Magdalou, J.; Humeau, C.; Ghoul, M. Automated analysis of synthesized oligorutin and oligoesculin by laccase. Can. J. Chem. 2011, 89, 964–970. [Google Scholar] [CrossRef]
- Anthoni, J.; Humeau, C.; Maia, E.R.; Chebil, L.; Engasser, J.M.; Ghoul, M. Enzymatic synthesis of oligoesculin: Structure and biological activities characterizations. Eur. Food Res. Technol. 2010, 231, 571–579. [Google Scholar] [CrossRef]
- Kurisawa, M.; Chung, J.E.; Uyama, H.; Kobayashi, S. Enzymatic synthesis and antioxidant properties of poly(rutin). Biomacromolecules 2003, 4, 1394–1399. [Google Scholar] [CrossRef]
- Uzan, E.; Portet, B.; Lubrano, C.; Milesi, S.; Favel, A.; Lesage-Meessen, L.; Lomascolo, A. Pycnoporus laccase-mediated bioconversion of rutin to oligomers suitable for biotechnology applications. Appl. Microbiol. Biotechnol. 2011, 90, 97–105. [Google Scholar] [CrossRef]
- Kurisawa, M.; Chung, J.E.; Uyama, H.; Kobayashi, S. Oxidative coupling of epigallocatechin gallate amplifies antioxidant activity and inhibits xanthine oxidase activity. Chem. Commun. 2004, 294–295. [Google Scholar]
- Kudanga, T.; Nyanhongo, G.S.; Guebitz, G.M.; Burton, S. Potential applications of laccase-mediated coupling and grafting reactions: A review. Enzym. Microb. Technol. 2011, 48, 195–208. [Google Scholar] [CrossRef]
- 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. [Google Scholar] [CrossRef]
- Gogoi, P.; Hazarika, S.; Dutta, N.N.; Rao, P.G. Kinetics and mechanism on laccase catalyzed synthesis of poly(allylamine)-catechin conjugate. Chem. Eng. J. 2010, 163, 86–92. [Google Scholar] [CrossRef]
- Gogoi, P.; Hazarika, S.; Dutta, N.N.; Rao, P.G. Laccase catalysed conjugation of catechin with poly(allylamine): Solvent effect. Chem. Eng. J. 2009, 155, 810–815. [Google Scholar] [CrossRef]
- Chung, J.E.; Kurisawa, M.; Uyama, H.; Kobayashi, S. Enzymatic synthesis and antioxidant property of gelatin-catechin conjugates. Biotechnol. Lett. 2003, 25, 1993–1997. [Google Scholar] [CrossRef]
- Gaffar Hossain, K.M.; Díaz González, M.; Monmany, J.M.D.; Tzanov, T. Effects of alkyl chain lengths of gallates upon enzymatic wool functionalisation. J. Mol. Catal. B 2010, 67, 231–235. [Google Scholar] [CrossRef]
- Vachoud, L.; Chen, T.; Payne, G.F.; Vazquez-Duhalt, R. Peroxidase catalyzed grafting of gallate esters onto the polysaccharide chitosan. Enzym. Microb. Technol. 2001, 29, 380–385. [Google Scholar] [CrossRef]
- Cho, Y.S.; Kim, S.K.; Ahn, C.B.; Je, J.Y. Preparation, characterization, and antioxidant properties of gallic acid-grafted-chitosans. Carbohydr. Polym. 2011, 83, 1617–1622. [Google Scholar] [CrossRef]
- Vartiainen, J.; Rättö, M.; Lantto, R.; Nättinen, K.; Hurme, E. Tyrosinase-catalysed grafting of food-grade gallates to chitosan: Surface properties of novel functional coatings. Packag. Technol. Sci. 2008, 21, 317–328. [Google Scholar] [CrossRef]
- Pasanphan, W.; Buettner, G.R.; Chirachanchai, S. Chitosan gallate as a novel potential polysaccharide antioxidant: An EPR study. Carbohydr. Res. 2010, 345, 132–140. [Google Scholar] [CrossRef]
- Aljawish, A.; Chevalot, I.; Piffaut, B.; Rondeau-Mouro, C.; Girardin, M.; Jasniewski, J.; Scher, J.l.; Muniglia, L. Functionalization of chitosan by laccase-catalyzed oxidation of ferulic acid and ethyl ferulate under heterogeneous reaction conditions. Carbohydr. Polym. 2012, 87, 537–544. [Google Scholar]
- Sousa, F.; Guebitz, G.M.; Kokol, V. Antimicrobial and antioxidant properties of chitosan enzymatically functionalized with flavonoids. Proc. Biochem. 2009, 44, 749–756. [Google Scholar] [CrossRef]
- Ihara, N.; Kurisawa, M.; Chung, J.E.; Uyama, H.; Kobayashi, S. Enzymatic synthesis of a catechin conjugate of polyhedral oligomeric silsesquioxane and evaluation of its antioxidant activity. Appl. Microbiol. Biotechnol. 2005, 66, 430–433. [Google Scholar] [CrossRef]
- Kim, S.; Cavaco-Paulo, A. Laccase-catalysed protein-flavonoid conjugates for flax fibre modification. Appl. Microbiol. Biotechnol. 2012, 93, 585–600. [Google Scholar] [CrossRef]
- Silva, C.; Matama, T.; Kim, S.; Padrao, J.; Nugroho Prasetyo, E.; Kudanga, T.; Nyanhongo, G.S.; Guebitz, G.M.; Casal, M.; Cavaco-Paulo, A. Antimicrobial and antioxidant linen via laccase-assisted grafting. React. Funct. Polym. 2011, 71, 713–720. [Google Scholar] [CrossRef]
- Chen, T.; Kumar, G.; Harris, M.T.; Smith, P.J.; Payne, G.F. Enzymatic grafting of hexyloxyphenol onto chitosan to alter surface and rheological properties. Biotechnol. Bioeng. 2000, 70, 564–573. [Google Scholar] [CrossRef]
- Brzonova, I.; Steiner, W.; Zankel, A.; Nyanhongo, G.S.; Guebitz, G.M. Enzymatic synthesis of catechol and hydroxyl-carboxic acid functionalized chitosan microspheres for iron overload therapy. Eur. J. Pharm. Biopharm. 2011, 79, 294–303. [Google Scholar] [CrossRef]
- Matheis, G.; Whitaker, J.R. Peroxidase-catalyzed cross linking of proteins. J. Protein Chem. 1984, 3, 35–48. [Google Scholar] [CrossRef]
- Oudgenoeg, G.; Hilhorst, R.; Piersma, S.R.; Boeriu, C.G.; Gruppen, H.; Hessing, M.; Voragen, A.G.J.; Laane, C. Peroxidase-mediated cross-linking of a tyrosine-containing peptide with ferulic acid. J. Agric. Food Chem. 2001, 49, 2503–2510. [Google Scholar]
- Mattinen, M.L.; Hellman, M.; Permi, P.; Autio, K.; Kalkkinen, N.; Buchert, J. Effect of protein structure on laccase-catalyzed protein oligomerization. J. Agric. Food Chem. 2006, 54, 8883–8890. [Google Scholar]
- Minamihata, K.; Goto, M.; Kamiya, N. Site-specific protein cross-linking by peroxidase-catalyzed activation of a tyrosine-containing peptide tag. Bioconjugate Chem. 2011, 22, 74–81. [Google Scholar] [CrossRef]
- Kuuva, T.; Lantto, R.; Reinikainen, T.; Buchert, J.; Autio, K. Rheological properties of laccase-induced sugar beet pectin gels. Food Hydrocoll. 2003, 17, 679–684. [Google Scholar] [CrossRef]
- Micard, V.; Thibault, J.F. Oxidative gelation of sugar-beet pectins: Use of laccases and hydration properties of the cross-linked pectins. Carbohydr. Polym. 1999, 39, 265–273. [Google Scholar]
- Chen, B.; McClements, D.J.; Gray, D.A.; Decker, E.A. Stabilization of soybean oil bodies by enzyme (laccase) cross-linking of adsorbed beet pectin coatings. J. Agric. Food Chem. 2010, 58, 9259–9265. [Google Scholar]
- Ogushi, Y.; Sakai, S.; Kawakami, K. Synthesis of enzymatically-gellable carboxymethylcellulose for biomedical applications. J. Biosci. Bioeng. 2007, 104, 30–33. [Google Scholar] [CrossRef]
- Sakai, S.; Hashimoto, I.; Ogushi, Y.; Kawakami, K. Peroxidase-catalyzed cell encapsulation in subsieve-size capsules of alginate with phenol moieties in water-immiscible fluid dissolving H2O2. Biomacromolecules 2007, 8, 2622–2626. [Google Scholar] [CrossRef]
- Lee, F.; Chung, J.E.; Kurisawa, M. An injectable enzymatically crosslinked hyaluronic acid-tyramine hydrogel system with independent tuning of mechanical strength and gelation rate. Soft Matter 2008, 4, 880–887. [Google Scholar] [CrossRef]
- Jin, R.; Hiemstra, C.; Zhong, Z.Y.; Feijen, J. Enzyme-mediated fast in situ formation of hydrogels from dextran-tyramine conjugates. Biomaterials 2007, 28, 2791–2800. [Google Scholar] [CrossRef]
- Park, K.M.; Shin, Y.M.; Joung, Y.K.; Shin, H.; Park, K.D. In situ forming hydrogels based on tyramine conjugated 4-Arm-PPO-PEO via enzymatic oxidative reaction. Biomacromolecules 2010, 11, 706–712. [Google Scholar] [CrossRef]
- Sakai, S.; Yamada, Y.; Zenke, T.; Kawakami, K. Novel chitosan derivative soluble at neutral pH and in situ gellable via peroxidase-catalyzed enzymatic reaction. J. Mater. Chem. 2009, 19, 230–235. [Google Scholar]
- Jayakumar, R.; Prabaharan, M.; Reis, R.L.; Mano, J.F. Graft copolymerized chitosan—Present status and applications. Carbohydr. Polym. 2005, 62, 142–158. [Google Scholar] [CrossRef]
- 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. [Google Scholar] [CrossRef]
- Mai, C.; Milstein, O.; Hüttermann, A. Fungal laccase grafts acrylamide onto lignin in presence of peroxides. Appl. Microbiol. Biotechnol. 1999, 51, 527–531. [Google Scholar] [CrossRef]
- Mai, C.; Milstein, O.; Hüttermann, A. Chemoenzymatical grafting of acrylamide onto lignin. J. Biotechnol. 2000, 79, 173–183. [Google Scholar]
- Tsujimoto, T.; Ando, N.; Oyabu, H.; Uyama, H.; Kobayashi, S. Laccase-catalyzed curing of natural phenolic lipids and product properties. J. Macromol. Sci. A 2007, 44, 1055–1060. [Google Scholar] [CrossRef]
- Tsujimoto, T.; Ikeda, R.; Uyama, H.; Kobayashi, S. Crosslinkable polyphenols from urushiol analogues. Macromol. Chem. Phys. 2001, 202, 3420–3425. [Google Scholar] [CrossRef]
- Kobayashi, S.; Uyama, H.; Ikeda, R. Artificial urushi. Chem. Eur. J. 2001, 7, 4755–4760. [Google Scholar]
- Ikeda, R.; Tanaka, H.; Oyabu, H.; Uyama, H.; Kobayashi, S. Preparation of artificial urushi via an environmentally benign process. Bull. Chem. Soc. Japan 2001, 74, 1067–1073. [Google Scholar] [CrossRef]
- Park, S.Y.; Kim, Y.H.; Won, K.; Song, B.K. Enzymatic synthesis and curing of polycardol from renewable resources. J. Mol. Catal. B. 2009, 57, 312–316. [Google Scholar] [CrossRef]
- Kim, Y.H.; Won, K.; 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 2005, 34, 33–38. [Google Scholar] [CrossRef]
- 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. [Google Scholar] [CrossRef]
- Reihmann, M.H.; Ritter, H. Enzymatically catalyzed synthesis of photocrosslinkable oligophenols. Macromol. Chem. Phys. 2000, 201, 1593–1597. [Google Scholar] [CrossRef]
- Bilici, A.; Kaya, I.; Yildirim, M. Peroxidase-catalyzed synthesis of polyphenols bearing aldehyde units. Des. Monomers Polym. 2011, 14, 353–366. [Google Scholar] [CrossRef]
- Majeau, J.A.; Brar, S.K.; Tyagi, R.D. Laccases for removal of recalcitrant and emerging pollutants. Biores. Technol. 2010, 101, 2331–2350. [Google Scholar]
- Husain, Q. Peroxidase mediated decolorization and remediation of wastewater containing industrial dyes: A review. Rev. Environ. Sci. Bio Technol. 2010, 9, 117–140. [Google Scholar] [CrossRef]
- Minussi, R.C.; Pastore, G.M.; Duran, N. Potential applications of laccase in the food industry. Trends Food Sci. Technol. 2002, 13, 205–216. [Google Scholar] [CrossRef]
- Lopez, C.; Moreira, M.T.; Feijoo, G.; Lema, J.M. Economic comparison of enzymatic reactors and advanced oxidation processes applied to the degradation of phenol as a model compound. Biocatal. Biotransf. 2011, 29, 344–353. [Google Scholar]
- Kadhim, H.; Graham, C.; Barratt, P.; Evans, C.S.; Rastall, R.A. Removal of phenolic compounds in water using Coriolus versicolor grown on wheat bran. Enz. Microb. Technol. 1999, 24, 303–307. [Google Scholar] [CrossRef]
- Uhnakova, B.; Ludwig, R.; Peknicova, J.; Homolka, L.; Lisa, L.; Sulc, M.; Petrickova, A.; Elzeinova, F.; Pelantova, H.; Monti, D.; et al. Biodegradation of tetrabromobisphenol A by oxidases in basidiomycetous fungi and estrogenic activity of the biotransformation products. Bioresour. Technol. 2011, 102, 9409–9415. [Google Scholar]
- Gaitan, I.J.; Medina, S.C.; Gonzalez, J.C.; Rodriguez, A.; Espejo, A.J.; Osma, J.F.; Sarria, V.; Almeciga-Diaz, C.J.; Sanchez, O.F. Evaluation of toxicity and degradation of a chlorophenol mixture by the laccase produced by Trametes pubescens. Bioresour. Technol. 2011, 102, 3632–3635. [Google Scholar]
- Lisov, A.V.; Pozhidaeva, Z.A.; Stepanova, E.V.; Koroleva, O.V.; Leontievsky, A.A. Conversion of polychloronhenols by laccases with 1-hydroxybenzotriazole as a mediator. Appl. Biochem. Microbiol. 2007, 43, 616–619. [Google Scholar] [CrossRef]
- Leontievsky, A.A.; Myasoedova, N.M.; Baskunov, B.P.; Evans, C.S.; Golovleva, L.A. Transformation of 2,4,6-trichlorophenol by the white rot fungi Panus tigrinus and Coriolus versicolor. Biodegradation 2000, 11, 331–340. [Google Scholar] [CrossRef]
- Schultz, A.; Jonas, U.; Hammer, E.; Schauer, F. Dehalogenation of chlorinated hydroxybiphenyls by fungal laccase. Appl. Environ. Microbiol. 2001, 67, 4377–4381. [Google Scholar] [CrossRef]
- Watanabe, C.; Kashiwada, A.; Matsuda, K.; Yamada, K. Soybean peroxidase-catalyzed treatment and removal of BPA and bisphenol derivatives from aqueous solutions. Environ. Prog. Sustain. Energy 2011, 30, 81–91. [Google Scholar] [CrossRef]
- Fukuda, T.; Uchida, H.; Suzuki, M.; Miyamoto, H.; Morinaga, H.; Nawata, H.; Uwajima, T. Transformation products of bisphenol A by a recombinant Trametes vilosa laccase and their estrogenic activity. J. Chem. Technol. Biotechnol. 2004, 79, 1212–1218. [Google Scholar] [CrossRef]
- Tsutsumi, Y.; Haneda, T.; Nishida, T. Removal of estrogenic activities of bisphenol A and nonylphenol by oxidative enzymes from lignin-degrading basidiomycetes. Chemosphere 2001, 42, 271–276. [Google Scholar] [CrossRef]
- Lloret, L.; Hollmann, F.; Eibes, G.; Feijoo, G.; Moreira, M.; Lema, J. Immobilisation of laccase on Eupergit supports and its application for the removal of endocrine disrupting chemicals in a packed-bed reactor. Biodegradation 2012, 1–14. [Google Scholar]
- Lloret, L.; Eibes, G.; Feijoo, G.; Moreira, M.T.; Lema, J.M.; Hollmann, F. Immobilization of laccase by encapsulation in a sol-gel matrix and its characterization and use for the removal of estrogens. Biotechnol. Prog. 2011, 27, 1570–1579. [Google Scholar] [CrossRef]
- Lloret, L.; Eibes, G.; Lu-Chau, T.A.; Moreira, M.T.; Feijoo, G.; Lema, J.M. Laccase-catalyzed degradation of anti-inflammatories and estrogens. Biochem. Eng. J. 2010, 51, 124–131. [Google Scholar] [CrossRef]
- Wang, C.J.; Thiele, S.; Bollag, J.M. Interaction of 2,4,6-trinitrotoluene (TNT) and 4-amino-2,6-dinitrotoluene with humic monomers in the presence of oxidative enzymes. Arch. Environ. Contam. Toxicol. 2002, 42, 1–8. [Google Scholar] [CrossRef]
- Thiele, S.; Fernandes, E.; Bollag, J.M. Enzymatic transformation and binding of labeled 2,4,6-trinitrotoluene to humic substances during an anaerobic/aerobic incubation. J. Environ. Qual. 2002, 31, 437–444. [Google Scholar] [CrossRef]
- Collins, P.J.; Kotterman, M.J.J.; Field, J.A.; Dobson, A.D.W. Oxidation of anthracene and benzo[a]pyrene by laccases from Trametes versicolor. Appl. Environ. Microbiol. 1996, 62, 4563–4567. [Google Scholar]
- Zeng, J.; Lin, X.G.; Zhang, J.; Li, X.Z.; Wong, M.H. Oxidation of polycyclic aromatic hydrocarbons by the bacterial laccase CueO from E. coli. Appl. Microbiol. Biotechnol. 2011, 89, 1841–1849. [Google Scholar] [CrossRef]
- Wang, J.Q.; Ogata, M.; Hirai, H.; Kawagishi, H. Detoxification of aflatoxin B(1) by manganese peroxidase from the white-rot fungus Phanerochaete sordida YK-624. FEMS Microbiol. Lett. 2011, 314, 164–169. [Google Scholar] [CrossRef]
- Eibes, G.; Cajthaml, T.; Moreira, M.T.; Feijoo, G.; Lema, J.M. Enzymatic degradation of anthracene, dibenzothiophene and pyrene by manganese peroxidase in media containing acetone. Chemosphere 2006, 64, 408–414. [Google Scholar] [CrossRef]
- Eibes, G.; Lu-Chau, T.; Feijoo, G.; Moreira, M.T.; Lema, J.M. Complete degradation of anthracene by Manganese Peroxidase in organic solvent mixtures. Enzym. Microb. Technol. 2005, 37, 365–372. [Google Scholar] [CrossRef]
- Mielgo, I.; Lopez, C.; Moreira, M.T.; Feijoo, G.; Lema, J.M. Oxidative degradation of azo dyes by manganese peroxidase under optimized conditions. Biotechnol. Prog. 2003, 19, 325–331. [Google Scholar] [CrossRef]
- Minussi, R.C.; Miranda, M.A.; Silva, J.A.; Ferreira, C.V.; Aoyama, H.; Marangoni, S.; Rotilio, D.; Pastore, G.M.; Duran, N. Purification, characterization and application of laccase from Trametes versicolor for colour and phenolic removal of olive mill wastewater in the presence of 1-hydroxybenzotriazole. Afr. J. Biotechnol. 2007, 6, 1248–1254. [Google Scholar]
- Suda, T.; Hata, T.; Kawai, S.; Okamura, H.; Nishida, T. Treatment of tetracycline antibiotics by laccase in the presence of 1-hydroxybenzotriazole. Bioresour. Technol. 2012, 103, 498–501. [Google Scholar]
- Gross, R.A.; Kumar, A.; Kalra, B. Polymer synthesis by in vitro enzyme catalysis. Chem. Rev. 2001, 101, 2097–2124. [Google Scholar]
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