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Feb 23, 2010 - Received November 5, 2009; Revised Manuscript Received January 20, ... describes how well lignosulfonates can act as plasticizers is th...
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Investigation of the Molecular Weight Increase of Commercial Lignosulfonates by Laccase Catalysis Dimitri Areskogh, Jiebing Li, Go¨ran Gellerstedt, and Gunnar Henriksson* Department of Fibre and Polymer Technology, Royal Institute of Technology, KTH, 100 44 Stockholm, Sweden Received November 5, 2009; Revised Manuscript Received January 20, 2010

Lignosulfonates are by-products from the sulfite pulping process. During this process, lignin is liberated from pulp fibers through sulfonation and washed away. As a consequence, the lignosulfonate molecules contain both hydrophobic and hydrophilic moieties. Lignosulfonates are low-value products with limited performance and are used as such as binders, surfactants, and plasticizers in concrete. Lignosulfonates face strong competition from synthetic petroleum-based plasticizers with superior quality. Therefore, increasing the performance of lignosulfonates is desirable not only from a sustainability point of view but also to expand their usage. One important aspect that describes how well lignosulfonates can act as plasticizers is the molecular weight. In this paper, the molecular weight of four commercial lignosulfonates is increased through oxidation by two laccases without utilization of mediators. Different parameters to obtain maximal molecular weight increase were identified and the technical significance of the experiments is discussed.

Introduction Approximately 20 - 30% of the wood matrix consists of the complex aromatic polymer lignin which is also present in lower amounts in many nonwoody plant tissues.1 Lignin has several biological functions in wood; it provides stiffness to the cell wall, water impermeability to the cell wall, which is necessary for the water transport in the lumen of the cells, it mediates the contact between the cells,2,3 and it serves as an obstacle for microbial degradation.4 The highly amorphous structure5 as well as the racemic nature of the lignin polymer6 makes it well suited for fulfilling these functions because it allows the lignin to fill the cavities between the polysaccharides, making the woody cell wall a highly compact structure. In addition, lignin forms multiple covalent bonds to polysaccharides.7 Studies have demonstrated that different types of polysaccharides are covalently cross-linked by lignin,8 which acts as a curing agent to lock the other wood polymeric material to each other and prevents extensive swelling when exposed to water. In many technical processes based on wood, the properties of lignin are beneficial, and here the lignin is left more or less intact. Examples of such processes are medium density fibreboards,9 newsprint, and other paper products based on mechanical pulp. However, other wood-based products require the lignin to be removed from the wood fiber to ensure a pure, flexible, and activated cellulose fiber. Examples of such products are high-quality papers, raw material for cellulose derivatives, and regenerated cellulose,10 as well as binders for pharmaceutical agents. Currently, two techniques are dominant for selective removal of lignin from wood: the kraft process where the majority of pulp for paper board products is produced and the sulfite process where pure cellulose, regenerated cellulose, and cellulose derivatives are produced. Large amounts of lignin are liberated from the wood raw material in these processes. In the kraft * To whom correspondence should be addressed. Phone: +46 8 790 6163. Fax: +46 8 790 6166. E-mail: [email protected].

process, the released lignin is heavily fragmented and the majority is insoluble in water at neutral pH. Normally it is concentrated and burned during the chemical recovery processes in a kraft mill, but recent techniques have been developed for recovering kraft lignin from the cooking liquids (black liquor) for application as fuel or other uses.11 In the sulfite process, a relatively high-molecular water-soluble lignin derivate, lignosulfonate, is produced. The lignosulfonate macromolecule differs greatly in molecular weight and contains different kinds of anionic groups, phenolic hydroxyl groups, sulfur-containing groups, carboxylic groups, and magnesium or sodium cations that are present as counterions. These groups provide lignosulfonates with good hydrophilic properties despite the hydrophobic phenylpropane backbone. The coexistence of hydrophilic and hydrophobic properties within the same compound has enabled lignosulfonates to be used in a great number of applications as technical surfactants and plasticizers with dispersing and stabilizing abilities in the oil drilling industry and concrete production.12,13 However, lignosulfonates as plasticizers are facing strong competition from petroleum-based synthetic equivalents with superior performance. The negative environmental impacts of the oil industry and its products are well documented and there is an increased demand to replace products based on a finite resource such as oil with products with equivalent properties from an infinite resource such as biomass. Thus, there is a potential interest in improving the quality of the lignosulfonates to enhance their performance in present applications and find novel uses. One interesting feature that can be improved is the molecular weight. On a molecular level, size of lignosulfonate molecules is an important aspect of their functioning as successful dispersants and plasticizers. The viscosity of a cement/concrete slurry is believed to be reduced by adsorption of the polymer onto the cement surface,14 and therefore, the difference in size of the polymer will affect its adsorption behavior. Studies have shown that increasing the molecular size of the polymer greatly improves the dispersion of cement particles.14

10.1021/bm901258v  2010 American Chemical Society Published on Web 02/23/2010

Commercial Lignosulfonates by Laccase Catalysis

Increase in molecular weight of lignosulfonates can be performed by introduction of radicals on the phenolic end groups of lignosulfonates and their subsequent coupling, which would lead to cross-linking of different lignosulfonate molecules, a process that mimics the natural lignin biosynthesis. Phenoxy radicals can be introduced to lignin in several ways. Radical formation occurs during the pulping process but is highly unordered and with low or no selectivity. Formation of radicals through manganese(III) oxidation of phenols in lignin under mild conditions is very selective and will lead to coupling products,15,16 but the inevitable formation of manganese(IV) oxide precipitation also makes this method unsuitable for large scale applications. More suitable options are enzymatic methods, due to their high specificity of the radical generation, mild reaction conditions and lack of undesired by-product. Both peroxidases and laccases are able to oxidize the phenolic structure to radicals with water as the sole byproduct.17,18 The advantage of using laccases over peroxidases in an industrial process lies in the nature of the oxidant. While peroxidases utilize hydrogen peroxide as oxidant, laccases require dioxygen. Several advantages of using dioxygen as oxidant over hydrogen peroxide are evident, such as higher stability, lower price, and no decomposition into radicals with subsequent inactivation of the enzymes. In nature, the relatively low oxidation potential of laccases, compared to other oxidoreductive enzymes, such as lignin peroxidase, poses problems. Laccases are only able to oxidize phenolic end groups in lignin, leaving nonphenolic end groups untouched. This is solved in lignin-degrading organisms by enabling the enzyme to utilize low-molecular diffusible redoxmediators with low molecular weight.19 Once activated by laccase, these mediators are small enough to penetrate the highly impermeable lignin structure to oxidize nonphenolic aromatic groups. This concept of a laccase-mediator system (LMS) is, however, not verified, although a number of naturally occurring components have been suggested as potential redox mediators in vivo. The discovery of synthetic compounds that could act as mediators20 has given rise to a renewed interest in laccases as a potentially new industrial catalyst, especially for the pulp and paper industry where it would aid in the delignification process. The LMS could provide an efficient, selective, and more notably, a completely chlorine-free bleaching process. To date, a number of synthetic mediators have been successfully used for pulp bleaching purposes.21,22 The significant cost of the redox mediators and the competition with improved traditional totally chlorine-free (TCF) and elemental chlorine-free (ECF) bleaching stages have proven to be significant obstacles and have to be overcome before full commercialization of LMS can occur. The effect of laccases on model structures representing phenolic and nonphenolic end groups has been studied previously.23 An array of different coupling reactions were observed some of which resulted in cross coupling of molecules (termed productive couplings) and others where a phenolic group was simply shifted from one end group to another (termed unproductive couplings) without any cross-linking.23 The unproductive couplings were shown to be absent if a sulfonic group is introduced at the R-carbon position,24 which is the case in lignosulfonates. The present study is focused on oxidation of lignin by laccase without mediators to achieve polymerization instead of degradation. By omitting the high-cost mediators and targeting at a new area where high-molecular lignins are desirable, it is likely that laccases can be utilized industrially. Three techniques were utilized to analyze the effect of laccases on lignosulfonates:

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molecular weight estimation with alkaline size exclusion chromatography (SEC) phenolic group determination with Folin and Ciocalteu (FC) phenol reagent in combination with UVspectroscopy, and FT-IR analysis. Analysis with FC phenol reagent has been applied on several lignins as well as pulps with reliable results in good agreement with those obtained by other conventional analytical methods for phenolic determination.25 The main advantage of the FC-method is its simplicity, which enables it to be used as a direct tool to determine the change in phenolic content during the reaction.

Materials and Methods Materials. Four lignosulfonate salts were kindly donated by Borregaard Lignotech (Sarpsborg, Norway). They were provided as a dried powder and were used without any further modifications. The characteristics of the salts are listed in Table 2, Supporting Information. 2,2′Azino-bis(3-ethylbenzthiazoline-6-sulfonate) (ABTS), Tris(hydroxymethyl)aminomethane (Trizma), Tris(hydroxymethyl)aminomethane hydrochloride (Trizma-HCl), acetic acid, and Folin and Ciocalteau phenol reagent were supplied by Sigma Aldrich (Stockholm, Sweden). All other chemicals were of analytical grade. Enzyme Assay. Laccase activity was determined by oxidation of 2,2′-azino-bis(3-ethylbenzthiazoline-6-sulfonate) (ABTS). A 200 mM stock solution was prepared by dissolving 1.1 mg ABTS in 25 mM Tris buffer at pH 7.5. A total of 240 µL of the stock solution was added to 760 µL of 25 mM buffer (chosen according to the enzyme) in a quartz cuvette (10 mm path length). An appropriate amount of laccase was added, and the oxidation of ABTS was followed by an absorbance increase at 725 nm (ε725nm ) 19000 M-1 cm-1). Enzyme activity was expressed in units (1 U ) 1 µmol ABTS oxidized per min at room temperature). Enzymes. Two different monocomponent laccases, NS51002 (Trametes Villosa) and NS51003 (Myceliophthora thermophila) were kindly donated by Novozymes (Bagsvaerd, Denmark). The laccases had slightly different oxidation potentials (approximately 0.7 V for NS51002 and 0.5 V for NS51003) at optimal pH and temperature. The pH and temperature optima for NS1002 and NS51003 were pH 5 and 50 °C and pH 7.5 and 40 °C, respectively. The activities of NS51002 and NS51003 at their respective optimal conditions were determined to 300 and 100 U/mL, respectively. The enzymes were used as supplied without any prior purification. Enzymatic Treatments. Lignosulfonates DP398, DP399, DP400, and DP401 were dissolved at different concentrations (1, 10, and 100 g/L) in 50 mL of 0.1 M buffer (acetate buffer with pH 5 for NS51002 or Tris-buffer with pH set to 7.5 for NS51003) in a reaction vessel prior to addition of laccases NS51002 or NS51003 (amounts corresponding to 50 or 500 U). The vessels were capped and saturated with pure oxygen during the reaction time. Samples were withdrawn from the reaction at 0, 4, and 24 h of reaction time and analyzed. A total of 18 experiments were conducted where the concentration of the lignosulfonates and the amount of added enzymes were altered. Determination of Phenolic Content. From the reaction vessel, 1 mL was transferred to a 50 mL volumetric flask to which 3 mL of FC reagent and 30 mL of distilled H2O were added and mixed thoroughly. After 5-8 min, 10 mL of 20% sodium carbonate solution was added and the volume was adjusted to 50 mL with distilled water. The mixture was kept with stirring for 2 h after which the absorbance was measured at 760 nm. The procedure was run in duplicate, and a phenol-free sample was also taken through the entire procedure as reference. The absorbance at 760 nm of the blue-colored samples was measured on a Varian (Palo Alto, CA) Cary 1E UV-visible spectrophotometer with the supplied Cary WinUV software. A calibration curve was set up where absorbance was plotted against concentration. For calibration purposes, a standard solution of 4.5 mM vanillin was used as the top level in a series of dilutions. The obtained calibration curve was linear, or nearly so, down to the zero-level. The amount of phenolic hydroxyl

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Figure 1. Increase of molecular weight of lignosulfonate salts DP398, DP399, DP400, and DP401 after oxidation by laccase NS51002. The salt concentration and enzyme dosage were as follows: 1 g/L and 50 U ((), 10 g/L and 50 U (9), 100 g/L and 50 U (2), 1 g/L and 500 U (×), 10 g/L and 500 U (/), 100 g/L and 500 U (b).

Figure 2. Increase of molecular weight of lignosulfonate salts DP398, DP399, DP400, and DP401 after oxidation by laccase NS51003. The salt concentration and enzyme dosage were as follows: 1 g/L and 50 U ((), 10 g/L and 50 U (9), 100 g/L and 50 U (2), 1 g/L and 500 U (×), 10 g/L and 500 U (0), 100 g/L and 500U (b).

groups was quantified by measuring the absorbance at 760 nm and fitting it to the calibration curve using a data-sampling software program. A phenol-free sample was used as a reference. FT-IR Analysis. Midinfrared spectra (600-4000 cm-1) were recorded with a Perkin-Elmer Spectrum 2000 FTIR spectrometer (Waltham, MA) equipped with an ATR system Specac MKII GoldenGate (Creecstone Ridge, GA). All spectra were obtained from lyophilized samples that were subjected to 32 scans at a resolution of 4 cm-1 and an interval of 1 cm-1 at room temperature. Before collection, background scanning was performed. Size Exclusion Chromatography. The water-soluble lignosulfonates were analyzed using a size exclusion chromatography system consisting of a Rheodyne 7725i (Rohnert Park, CA) manual injector, Waters 515 HPLC pump (Milford, MA), three TSK-gel columns (Tosoh Bioscience, Tokyo, Japan) coupled in a series, G3000PW (7.5 × 300 mm, 10 µm particle size), G4000PW (7.5 × 300 mm, 17 µm particle size), and

G3000PW, and a Waters 2487 dual wavelength absorbance detector (Milford, MA). The mobile phase during the analysis was 10 mM NaOH. A volume of 20 µL was injected and absorbance at 280 and 254 nm was recorded. The columns were calibrated with polyethylene glycol (PEG) and polyethylene oxide (PEO) standards with specific molecular weights ranging from 1500 to 400000. Integration and quantification of peaks were performed using Millenium 2 software.

Results and Discussion Two different laccases, NS51002 (Trametes Villosa) and NS51003 (Mycoliophthora thermophila) with different pH optima (pH 5 and 7.5) were used for the treatment of four lignosulfonate salts at three concentrations ranging from 1 to 100 g/L. In all cases, an increase in molecular weight and

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Figure 3. Polydispersity index values of lignosulfonate salts DP398-DP401 at 100 g/L during laccase oxidation. The denotions are as follows: 50 U NS51002 ((), 500 U NS51002 (9), 50 U NS51003 (2), and 500 U NS51003 (×).

Figure 4. Decrease of phenolic content of lignosulfonate salts DP398, DP399, DP400, and DP401 after oxidation by laccase NS51002. The salt concentration and enzyme dosage were as follows: 1 g/L and 50 U ((), 10 g/L and 50 U (9), 100 g/L and 50 U (2), 1 g/L and 500 U (×), 10 g/L and 500 U (/), 100 g/L and 500 U (b).

polydispersity, as well as a decrease in the phenolic content, was observed (Figures 1-5). The polymerization is likely to be explained by the mode of action of the laccase where the enzyme initiates oxidation of phenolic end groups into stabilized radicals that subsequently undergo radical-radical coupling through which phenyl ether-carbon and carbon-carbon bonds are formed. These bonds act to form both intramolecular linkages within the lignosulfonate macromolecule and to link one lignosulfonate macromolecule to another and thus yield the observed increase in molecular weight (Figures 1 and 2). Previous model compound experiments with NS51002 and NS51003 highlighted a distinct difference in their ability to

oxidize different end groups in lignin.23 While NS51002 was able to oxidize non-phenolic end groups due to the higher redox potential, NS51003 proved to be capable of oxidizing only phenolic end groups. These findings are supported in the experiments with lignosulfonates. Not only did the phenolic content of the salts decrease at all concentrations during oxidation (Figures 4 and 5), the samples treated with NS51002 displayed higher Mw after 24 h of oxidation (Figures 1 and 2). In terms of the concentration of the lignosulfonate salts, a clear trend is observed; polymerization is greatly improved at high concentrations and the majority of the molecular weight increase is achieved already after 4 h of reaction time. The most

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Figure 5. Decrease of phenolic content of lignosulfonate salts DP398, DP399, DP400, and DP401 after oxidation by laccase NS51003. The salt concentration and enzyme dosage were as follows: 1 g/L and 50 U ((), 10 g/L and 50 U (9), 100 g/L and 50 U (2), 1 g/L and 500 U (×), 10 g/L and 500 U (/), 100 g/L and 500 U (b).

extreme case was DP401 oxidized by NS51002 where a 23fold increase in Mw was observed already after 4 h (the increase dropped, however, to 20-fold after 24 h). The lowest MW increase was seen in DP400 with only a 2.6-fold increase. The molecular weight of the remaining salts increased by a factor of 6-12. At lignosulfonate concentrations below 100 g/L, the MW increase was much less apparent, but the conclusion that increased concentration results in higher molecular weight is still valid. Polydispersity index values were obtained from lignosulfonates at a concentration of 100 g/L during oxidation (Figure 3). As seen, the increase of polydispersity is intimately linked with the MW increase. This is expected due to the nature of the nonselective radical-radical polymerization caused by laccase oxidation of phenolic end groups in lignosulfonates. The coupling reactions linking lignosulfonate end groups to each other occur spontaneously with little or no control, and because the radicals to initiate the polymerization are readily available, the reaction propagates with subsequent polydispersity increase. The ability of NS51002 to oxidize the lignosulfonate salts at high concentrations was shown to be largely independent of the activity of the enzyme as only minor difference in molecular weight increase was observed when the enzyme activity was increased 10-fold (Figure 1). This was not the case with NS51003 (Figure 2). To approach the levels of the MW increase seen with NS51002, the dosage of NS51003 had to be increased 10-fold (Figure 2). In this case it is evident that low enzyme dosage does not yield any MW increase at any concentration. To establish whether this observation is pH-related, the optimal pH for the two enzymes was switched, that is, the enzyme NS51002 with an optimum at pH 5 was run at pH 7.5 and vice versa for the enzyme NS51003. The results (data not shown) clearly stated that only a minor molecular weight increase was observed when the enzymes were not allowed to operate at their optimal pH. This suggests that the outcome of the reaction is directed by the catalytic properties of the enzyme and not the pH. The redox potential difference between the two laccases is most likely the probable explanation to these observations. Contrary to NS51003, NS51002 can oxidize both

phenolic and nonphenolic end groups to achieve a higher degree of polymerization. As a consequence of increased MW, an inversely proportional decrease in phenolic hydroxyl group (Ph-OH) was observed in all experiments (Figures 4 and 5). This suggests a strong relationship between consumption of phenols and increase of molecular weight. It appears that the phenolic group content of the lignosulfonate salts is one of the major characteristics to achieve a strong MW increase. Another characteristic of the lignosulfonates is the degree of sulfonation. The lignosulfonates DP398-DP401 contained different amounts of organic sulfur (see Table 1, Supporting Information). Previous experiments with model compounds revealed that unproductive couplings of phenolic end groups occur after oxidation of laccase,23 and those are avoided if the R-carbon of the end group is sulfonated,24 which is the case in lignosulfonates. It appears that the low degree of sulfonation in DP400 plays a role here where the consumption of phenols caused by laccase oxidation leads to an insignificant molecular weight increase due to the possible formation of unproductive couplings. While the phenolic content of DP400 was reduced within the same range as the other salts, no significant increase in molecular weight was observed, most likely due to extensive formation of nonproductive bonds. FTIR-analysis of the lignosulfonates DP398-DP401 prior to and after oxidation by NS51002 (Figure 7) revealed only minor differences, suggesting that no structural changes were inflicted on the lignosulfonates other than the increased molecular weight and reduction of phenols. Experiments were conducted to examine whether any differences were observed if the laccases are added in a portionwise or a batchwise fashion. For these experiments, the concentration of lignosulfonate salts was set to 100 g/L, the total amount of laccase NS51002 to 50 U, and the reaction time to 4 h. Two separate experiments were conducted where the enzyme was added in equal amounts, 12.5 U each hour, and where the total 50 U was added at t ) 0. Only minor differences in the molecular weight increase and phenolic content decrease were observed, but there was a clear trend that a plateau was reached

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Figure 6. A comparison between single and continuous addition of NS51002 to lignosulfonate salts DP398 ((), DP399 (9), DP400 (2), and DP401 (×). The top and bottom figures depict the molecular weight increase and decrease in phenolic content, respectively.

Figure 7. FT-IR analysis of lignosulfonate salts prior to (solid) and after oxidation of 50 U NS51002 (dashed).

after the first hour in the experiments where 50 U of laccase was added at t ) 0. In the experiments where the enzymes were added in 12.5 U dosages each hour, the molecular weight continued to increase, while the phenolic content decreased with each addition of laccase (Figure 6). These differences, although small, are significant as they highlight an important observation; the molecular weight increase can be directed by the amount of enzyme added. The tendency of the MW increase and Ph-OH decrease to level out after approximately 4 h of reaction time was observed in all experiments. After a rapid MW increase and a simultaneous Ph-OH decrease, the two values reach a plateau where they stay for the remainder of the reaction time. An explanation to these observations lies in the macromolecular structure of the lignosulfonate molecule. It appears that the lignosulfonate molecule is able to form spherical microgels26 in water where the hydrophobic elements of the macromolecule are buried in the core, while the hydrophilic elements are displaced at the surface, much like a micelle. This suggestion fit with the

observed changes in molecular size, phenolic content, and polydispersity changes. Due to sterical constraints, the enzyme can only access the phenolic end groups located on the surface of the lignosulfonate micelle. It is therefore likely that the molecular weight increase levels out when the phenolic end groups, available to the enzyme for oxidation, are consumed while the remaining groups buried within the micelle remain untouched. At that point, the polymerization reactions are slowed down and the molecular weight increase levels out. It is striking that both the MW increase (Figures 1 and 2) and the consumption of phenols (Figures 4 and 5) levels out and reaches a plateau after approximately 4 h. After that period of time, approximately two-thirds of the phenols are consumed, while the remaining one-third is left untouched, probably buried in the micelle core. Formation of micelle-like structures can also explain the relationship between initial and increased MW of the lignosulfonates. The size of the lignosulfonate micelle in solution depends on how much the micelle can grow during laccase oxidation. Small initial MW (as in the case with DP401) results in a strong increase (Table 2, Supporting Information), suggesting that small micelles are more likely to grow in size more extensively than large micelles.

Conclusions The results above clearly display the possibility to increase the molecular weight of lignosulfonates with a factor ranging from 5 to 25 by treatment with commercially available laccase without the usage of any mediators. Several key factors were identified as necessary to obtain the maximal molecular weight increase. Among these were reaction time, enzyme redox potential, continuous addition of enzyme, phenolic and sulphonate content, and initial MW of the lignosulfonates. However, the most important factor is the concentration of lignosulfonates. The conditions used during the oxidation, temperature, pH, enzyme dosage, and incubation times, are far from extreme and are likely to be suitable for industrial application. This opens up for a novel process to adjust the molecular weight of

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lignosulfonates with an environmentally benign method. The tailored molecular weight should expand the usage of lignosulfonates for existing or novel applications. Acknowledgment. Financial support from the Biorenew program, EU Grant FP6-NMP2-CT 2006-26456, is gratefully acknowledged. Borreegaard LignoTech and Novozymes A/S are thanked for providing lignosulfonates and laccases for these experiments. Supporting Information Available. Tabulated data with characteristics of the lignosulfonate salts used in these experiments as well as relative molecular weight increase. This material is available free of charge via the Internet at http:// pubs.acs.org.

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Areskogh et al. (12) Chiwetelu, C.; Hornof, V.; Neale, G. H. Enhanced oil recovery using lignosulfonate/petroleum sulfonate mixtures. Trans. Inst. Chem. Eng. 1982, 60 (3), 177–82. (13) Yousuf, M.; Mollah, A.; Palta, P.; Hess, T. R.; Vempati, R. K.; Cocke, D. L. Chemical and physical effects of sodium lignosulfonate superplasticizer on the hydration of portland cement and solidification/ stabilization consequences. Cem. Concr. Res. 1995, 25 (3), 671–82. (14) Anderson, P. J.; Roy, D. M.; Gaidis, J. M. The effect of superplasticizer molecular weight on its adsorption on, and dispersion of, cement. Cem. Concr. Res. 1988, 18 (6), 980–6. (15) Landucci, L. L.; Geddes, S. A.; Kirk, T. K. Synthesis of carbon-14labeled 3-methoxy-4-hydroxy-R-(2-methoxyphenoxy)-β-hydroxypropiophenone, a lignin model compound. Holzforschung 1981, 35 (2), 67–70. (16) Önnerud, H.; Zhang, L.; Gellerstedt, G.; Henriksson, G. Polymerization of monolignols by redox shuttle-mediated enzymatic oxidation: a new model in lignin biosynthesis. Plant Cell 2002, 14 (8), 1953–1962. (17) Azevedo Ana, M.; Martins Veronica, C.; Prazeres Duarte, M.; Vojinovic, V.; Cabral Joaquim, M.; Fonseca Luis, P. Horseradish peroxidase: a valuable tool in biotechnology. Biotechnol. Annu. ReV. 2003, 9, 199–247. (18) Riva, S. Laccases: Blue enzymes for green chemistry. Trends Biotechnol. 2006, 24 (5), 219–226. (19) Bourbonnais, R.; Paice, M. G. Oxidation of non-phenolic substrates. An expanded role for laccase in lignin biodegradation. FEBS Lett. 1990, 267 (1), 99–102. (20) Sealey, J.; Ragauskas, A. J.; Elder, T. J. Investigations into laccasemediator delignification of kraft pulps. Holzforschung 1999, 53 (5), 498–502. (21) Camarero, S.; Ibarra, D.; Martinez, A. T.; Romero, J.; Gutierrez, A.; del Rio, J. C. Paper pulp delignification using laccase and natural mediators. Enzyme Microb. Technol. 2007, 40 (5), 1264–1271. (22) Ibarra, D.; Camarero, S.; Romero, J.; Martinez, M. J.; Martinez, A. T. Integrating laccase-mediator treatment into an industrial-type sequence for totally chlorine-free bleaching of eucalypt kraft pulp. J. Chem. Technol. Biotechnol. 2006, 81 (7), 1159–1165. (23) Areskogh, D.; Li, J.; Henriksson, G.; Nousiainen, P.; Gellerstedt, G.; Sipila¨, J. Polymerisation of lignin end group model compounds by laccase. Holzforschung 2010, 64, 21–34. (24) Areskogh, D.; Nousiainen, P.; Li, J.; Gellerstedt, G.; Henriksson, G.; Sipila¨, J., Sulfonation of phenolic end-groups in lignin direct laccase initiated reaction towards cross-linking. Ind. Biotechnol. 2010, accepted for publication. (25) de Sousa, F.; Reimann, A.; Marianne, B.; Nilvebrant, N.-O. In Estimating the Amount of Phenolic Hydroxyl Groups in Lignins, 11th International Symposium on Wood and Pulping Chemistry, Nice, France, 2001; Nice, France, 2001; pp 649-653. (26) Rezanowich, A.; Goring, D. A. I. Polyelectrolyte expansion of a lignin sulfonate microgel. J. Colloid Sci. 1960, 15, 452–71.

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