Peroxidase-Catalyzed Coupling of Phenol in the Presence of Model

to form phenoxy radicals and does so along a catalytic cycle involving .... and unlabeled phenol, 2-mM H2O2, a predetermined dosage of HRP ..... Scien...
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Environ. Sci. Technol. 2004, 38, 5238-5245

Peroxidase-Catalyzed Coupling of Phenol in the Presence of Model Inorganic and Organic Solid Phases QINGGUO HUANG AND WALTER J. WEBER, JR.* Department of Chemical Engineering, Energy and Environment Program, 4103 Engineering Research Building, The University of Michigan, Ann Arbor, Michigan 48109-2099

Peroxidase-catalyzed oxidative coupling reactions of phenol in aqueous systems variously containing silica sand, cellulose, lignin, and polymethylstyrene were investigated. These four solid phase materials represent a broad spectrum of different natural geosorbent types in terms of their physicochemical characteristics. Each solid was found to influence peroxidase-catalyzed phenol coupling, either by mitigation of enzyme inactivation, by participation in cross-coupling, or by a combination of these two activities. Mitigation of enzyme inactivation was observed for those three of the four model solids found to adsorb the enzyme effectively; i.e., cellulose, silica sand, and lignin. Two solids, polymethylstyrene and lignin, were found to participate significantly in cross-coupling reactions. It is postulated that relatively hydrophilic solids can mitigate peroxidase inactivation by forming enzymesolid associations. Aromatic structures or unsaturated C-C bonds were found to be features of the solid-phase materials that allowed them to participate in crosscoupling. The results have important implications for process feasibility assessment and the engineering design of soil/ sediment remediation systems employing enzymatic coupling schemes.

Introduction Organic contaminants of phenolic character are numerous and widely distributed, relatively hydrophilic and environmentally mobile, and exhibit a range of multiple toxicity effects (1-3). This broad class of organic contaminants represents substantial risks to both natural ecosystems and human populations and thus constitutes a focus of major environmental concern. One of the most important environmental transformation pathways for phenolic compounds in natural systems is oxidative coupling, a reaction mediated by such naturally occurring catalysts as extracellular oxidoreductases and mineral oxides (4-8). Peroxidases comprise a particularly important class of enzymes in that they are able to catalyze the oxidative coupling of a broad spectrum of phenolic compounds (9, 10). Within this class of enzymes, horseradish peroxidase (HRP) has been particularly well investigated with respect to structure and catalysis mechanisms (11-17). In the presence of hydrogen peroxide, HRP mediates single-electron oxidations of phenolic substrates to form phenoxy radicals and does so along a catalytic cycle * Corresponding author phone: (734)763-2274; fax: (734)936-4391; e-mail: [email protected]. 5238

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involving the three sequential steps represented schematically in Figure 1. The cycle is initiated by oxidation of native HRP by H2O2 to form an enzyme intermediate (i.e., Compound I,Ei). In the second step, Compound I abstracts an electron from a phenolic substrate to form another enzyme intermediate (i.e., Compound II,Eii) and a phenoxy radical. Compound II can in the third step abstract still another electron from a phenolic substrate to generate a second phenoxy radical and, in the process, revert to the native enzyme, thus completing the catalytic cycle. The radicalgeneration steps (the second and third steps) of the HRP catalytic cycle are essentially reversible (18); i.e., radicals may also abstract electrons from the enzyme intermediates via “reverse electron transfer” and thus revert to the original molecular state. Phenoxy radicals produced in the peroxidase-mediated enzymatic process can, in postenzymatic reactions, couple with each other (15-17) or with other reactive substances present in any given system (19, 20). Self-coupling of phenoxy radicals with each other dominates in systems that lack appropriate substrates to participate in cross-coupling reactions, leading to formation of polymeric precipitates that can be readily removed from water (21-28). Significant crosscoupling with other reactive substances usually occurs in systems containing soil or sediment materials (4, 19, 20, 29, 30), generally resulting in the incorporation of the phenolic substrates in soil/sediment organic matter (SOM). Both selfand cross-coupling reactions can be expected to decrease the environmental mobility and toxicity of the substrate compounds; i.e., by polymer precipitation or by binding to soil aggregates, respectively. It has been demonstrated that the extractability of phenolic contaminants from soil matrices is dramatically reduced as a result of oxidative coupling (3134). Indeed, enhancement of either or both types of oxidative coupling reactions by amendments of extracellular enzymes has been proposed as a potential strategy for remediation of soil/sediment environments contaminated by phenolic substances (4, 35, 36). Investigations of enzyme-mediated oxidative crosscoupling reactions of phenolic contaminants have focused principally on homogeneous aqueous systems, with dissolved humic precursors or humic substances serving as model cross-coupling substrates (30, 37-41). Although such studies have proved useful in elucidating mechanisms by which phenolic contaminants may bind to soil-derived organic matter, cross-coupling reactions so observed may not be used reliably to predict reactions occurring in soil/sediment environments involving solid-phase soil organic matter (SOM) matrices. These solid organic phases, in addition to potentially interacting with phenoxy radicals in crosscoupling reactions, may also influence the enzymatic coupling process by directly interacting with either the enzyme or the original phenolic substrate(s). Phenol coupling reactions in systems involving bulk soils have been investigated (29, 42, 43), but the complexities of the soil systems involved have precluded the drawing of insights regarding solid-phase effects. We reported in an earlier paper (29) that two different natural geosorbents, Chelsea soil and Lachine shale, were each found to enhance HRP-mediated phenol coupling reactions. They were found to do so to remarkably different extents, however, a fact that apparently relates to differences in the physicochemical properties of their respective associated soil organic matter phases. A systematic understanding of relationships between solid effects and the physicochemical characteristics of different 10.1021/es049826h CCC: $27.50

 2004 American Chemical Society Published on Web 08/27/2004

FIGURE 1. Schematic representation of HRP-mediated catalytic cycle.

FIGURE 2. Structural characteristics of organic polymers tested. solid is thus critically needed for predicting reaction efficiencies in natural systems involving soil/sediment materials having different properties and thus for guiding remediation practice. The work reported here is intended as an initial step in developing such a systematic understanding of these relationships. To this end, the horseradish peroxidase mediated oxidative coupling reactions of phenol were investigated in systems containing four model solid materials of different but well-defined physicochemical properties; specifically, silica sand, cellulose, lignin, and polymethylstyrene. The conditions under which the experiments were carried out involved solid/water ratios selected to ensure that any physical adsorption of phenol and its dissolved coupling products to the model solids would be negligible. Phenol transformation was found to be enhanced in the presence of each solid studied, but the relative effects were observed to vary greatly among the four different solids. Attempts were made in the study to elucidate mechanisms by which the solids affect the peroxidase-catalyzed phenol coupling process and to relate the observed effects to the physicochemical characteristics of the solids.

Experimental Section Materials. Silica sand, cellulose, lignin, and polymethylstyrene (PMS) were investigated as model solids. These materials represent a broad spectrum of natural geosorbent elemental composition and physicochemical characteristics. The structures of the three organic polymers are presented in Figure 2. Atomic O/C ratios of the three organic materials are 0.83, 0.33, and 0 for cellulose, lignin, and PMS, respectively, and their respective H/C ratios are 1.67, 0.98, and 1.11, for which the atomic ratios of cellulose and PMS are estimated from their structures and that of lignin is from ref 44. Silica sand samples were prepared by grinding Ottawa sand (Sil-CoSil-40, 99.8%SiO2, U.S. Silica, Ottawa, IL) and washing sequentially with 0.1-M hydrochloric acid, a 5% hydrogen peroxide solution, and milli-Q water to remove impurities. Cellulose and poly(4-methylstyrene) samples obtained from Scientific Polymer Products Inc. (Ontario, NY) were washed thoroughly by methanol followed by milli-Q water and dried

prior to use. Organosolv lignin obtained from Aldrich Chemical Co. (Milwaukee, WI) was washed sequentially with 50% methanol and milli-Q water and dried prior to use. The specific surface areas (m2/g), as measured by the N2-BET method, are respectively 1.2, 1.6, 1.8, and 0.19 m2/g for silica sand, cellulose, lignin, and PMS. Extracellular horseradish peroxidase (type-I, RZ)1.3), hydrogen peroxide (30.8%, ACS reagent), 2,2′-azino-bis(3ethylbenzthiazoline-6-sulfonic acid) (ABTS) (98%, in diammonium salt form), and phenol-UL-14C (51.4 mCi/mmol) were obtained from Sigma Chemical Co. (St. Louis, MO). Phenol (99+%, biochemical grade) was from Acros Organics (Belgium, NJ). ScintiSafe Plus 50% liquid scintillation cocktail and all other chemicals were obtained in the highest quality available from Fisher Scientific (Fairlawn, NJ). Enzymatic Coupling and Nonextractable Product Formation. Nonextractable products (NEP) are here defined as reaction products remaining in the solid phase after solvent extraction and phase separation, as measured collectively by the residual activity of 14C-labeled phenol employed in enzymatic coupling experiments. The experiments were performed at room temperature in 13 × 100-mm glass test tubes operated as completely mixed batch reactors (CMBRs). Each reactor contained 3 mL of 10-mM phosphate buffer (pH ) 7.0) solution comprised by a 500-µM mix of 14C-labeled and unlabeled phenol, 2-mM H2O2, a predetermined dosage of HRP, and a preselected amount of a model solid. The 2-mM H2O2 concentration was selected on the basis of our earlier investigations (45) to ensure enzyme saturation by H2O2 and avoid any significant side effects of hydrogen peroxide on HRP in the systems studied. Triplicate experiments were conducted for each reaction condition. After a 2-h reaction period during which enzyme activity was completely depleted, 3 mL of methanol was added to each reactor and mixed with the 3-mL reaction solution, essentially forming a 50% methanol solution. A 1.5-mL sample of the mixture was withdrawn from the reactor after 30 min of mixing and extraction and centrifuged at 14 000 rpm (20 800g) for 20 min to separate the liquid and solid phases. A 0.5-mL aliquot of the liquid phase was sampled and mixed with 3 mL of ScintiSafe Plus 50% liquid scintillation cocktail, and 14C radioactivity was then measured using a Beckman LS6500 liquid scintillation counter (Beckman Instruments, Inc.). The radioactivity remaining in the solid phase was calculated by mass balance and NEP concentration calculated and expressed in terms of molar equivalents of phenol in the original solution. Tests on blank control samples indicated that phenol losses during the reaction time and subsequent analytical processes were negligible. Sorption of Phenol and Dissolved Coupling Products. Sorption of phenol by the solid materials studied was evaluated by mixing a phosphate buffer solution having a 500-µM phenol concentration with the solids at predetermined solid/water ratios in the test tube reactors. A 1.5-mL sample of solution was taken from each reactor after overnight mixing and centrifuged at 14 000 rpm (20 800g) for 20 min. The supernatant was then sampled for analysis using an Agilent 1100 Series HPLC system equipped with a Phenomenex C18 column (250 × 2.0 mm, 5 µm particle size). The mobile phase, operated at 0.40 mL/min, comprised an acetonitrile component (35%) and an aqueous component (65%), each containing 1% acetic acid. Phenol concentration was determined by UV absorbance at 270 nm. The sorption of dissolved products from phenol-coupling reactions was also evaluated. The product solution employed in this experiment was prepared by carrying out the phenolcoupling reaction in a large vessel containing 500 mL of a reaction solution comprised by a 500-µM mix of radiolabeled and unlabeled phenol, 2-mM H2O2, and 0.5-unit/mL HRP. The product mixture was then centrifuged after a 2-h reaction VOL. 38, NO. 19, 2004 / ENVIRONMENTAL SCIENCE & TECHNOLOGY

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period, and the liquid phase was used for the sorption test. This solution contained a 320.1-µM concentration of phenol as measured by HPLC, and a 382.6-µM phenol equivalent of 14C as measured by radioactivity analysis. The difference between the phenol equivalent of 14C and the phenol concentration, i.e., 62.5 µM, represents the concentration of dissolved coupling products. Aliquots of the solution were then mixed with the solid at a series of predetermined solid/ water ratios in the test tube reactors. After 2 h of mixing, 1.5 mL of sample was taken and centrifuged at the same speed specified above. Concentrations of phenol and 14C remaining in the aqueous phase were then again measured, and the concentration of dissolved coupling products was calculated by difference. Enzyme Inactivation Rates. Enzyme inactivation rates during phenol coupling reactions were examined separately with nonradiolabeled phenol at selected conditions in the test tube reactor systems. Samples of 0.05-mL volume were taken at predetermined times and immediately measured for enzyme activity by the ABTS method described earlier (29, 46). Briefly, a 0.05-mL sample was added to a cuvette containing a 3-mL volume of phosphate buffer solution (pH ) 6.0), followed by addition of 0.3 mL of 20-mM ABTS and 0.3 mL of 10-mM hydrogen peroxide to start the assay. The absorbance change at 405 nm was monitored by a 6405 UV/ vis spectrophotomer (Jenway Inc., Princeton NJ). One unit of peroxidase activity is defined as that amount catalyzing the oxidation of 1 µmol of ABTS per minute. Because the procedure described above involves a dilution of a 0.05-mL sample to a 3-mL assay media, the interference of solidscontaining samples on absorbance measurements is largely reduced. Our control tests indicate that this method can reliably measure HRP activity in samples containing solids up to 100 g/L. Enzyme Sorption. As described earlier, HRP mediates phenol coupling reactions through a catalytic cycle that involves conversions of the enzyme among the native form and two intermediate forms (i.e. native HRP and Compound I and Compound II, respectively). It is thus necessary to evaluate the respective sorption of each HRP form by the solid materials studied for estimating enzyme sorption behaviors during HRP-mediated oxidative coupling processes. Because HRP intermediates are formed in a successive manner in the catalytic cycle shown in Figure 1, methods have been developed to prepare those intermediates by manipulating the stoichiometry of the enzyme substrates in a manner that allows the reaction to proceed precisely to a desired step in the catalytic cycle (14, 47). These methods were employed in the present study to prepare Compound I and Compound II samples from native HRP to evaluate their sorption by the solid materials independently. A 50unit/mL stock solution of the native HRP was first prepared by dissolving the enzyme in a 10-mM phosphate buffer solution (pH)7.0). The molar concentration of the enzyme in this solution was approximately 2.6 µM, as determined spectrophotometrically (6405 UV/vis spectrophotomer, Jenway Inc., Princeton NJ) at 403 nm and calculated using a molar absorptivity value of 1.02 × 105 M-1 cm-1 (47). This indicates that one enzyme unit is composed approximately of 0.052 nmole of HRP molecules. Compound I was formed by quantitative addition of hydrogen peroxide to a native HRP stock solution to yield a concentration equivalent to that of the enzyme (14, 47), the formation of Compound I being confirmed by observation of UV spectrum changes. This solution was then diluted by a factor of 100 to yield a solution of Compound I at a level of approximately 0.5-unit/ mL activity for subsequent use in related sorption experiments. For preparation of Compound II, hydrogen peroxide and K4Fe(CN)6 were both added to a sample of the 50-unit/ 5240

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mL native HRP stock solution in the concentrations equivalent to that of the enzyme (47). This solution was then diluted by a factor of 100 to obtain a Compound II solution of approximately 0.5-unit/mL activity. The 50-unit/mL stock solution of native HRP was directly diluted by a factor of 100 to obtain a solution of 0.5-unit/mL activity for use in the sorption experiments. Sample solutions of Compound I, Compound II, and native HRP were prepared immediately prior to the sorption experiments. A 7-mL sample of each enzyme solution was mixed by hand shaking for one minute with a solid at a predetermined solid/solution ratio in a 13 × 100-mm glass test tube. The mixture was then centrifuged at 3500 rpm (1300g) for 30 min, after which 0.05 mL of the supernatant was taken to measure the enzyme activity remaining in the aqueous phase by the ABTS method (46). This enzyme activity was divided by the enzyme activity measured for a solids-free control sample to calculate the percentage of enzyme remaining in the aqueous phase after sorption. Each condition was tested in triplicate. Enzyme activity reduction in the solids-free control was insignificant, generally less than 5% during the sorption test period.

Results and Discussion Enzymatic Coupling in the Presence of Solid Materials. As noted in the Introduction, the solid/water ratios used in the study were selected so that any direct sorption of the parent phenol and/or its dissolved coupling products would be insignificant. The results of the sorption experiments described in the Experimental Section indicate that phenol and its dissolved coupling products remained almost completely in the aqueous phase at the solid/water ratios studied. Percentages of the phenol remaining in the aqueous phases of nonreactive systems containing each of the solids at their respective highest solid/water ratios studied (i.e. cellulose, silica sand, and PMS at 200 g/L and lignin at 1 g/L) were 93.2 ( 0.7%, 100.8 ( 1.8%, 95.8 ( 1.1%, and 97.1 ( 0.5%, respectively. The sorption results for cellulose and lignin are consistent with those of a prior sorption study by Xing et al. involving these same two materials (44). The direct sorption of dissolved phenol-coupling products were similarly insignificant, the percentages remaining in aqueous phase at the highest solid/water ratios being 95.2 ( 1.3%, 96.8 ( 1.8%, 98.6 ( 2.6%, and 102.7 ( 3.5% for cellulose, silica sand, PMS and lignin, respectively. Preliminary tests on the 50% methanol extraction employed for NEP measurement in this study demonstrated essentially complete recoveries of phenol and dissolved phenol-coupling products from the solid/ solution systems investigated. Based on these several sets of results, we would not expect the solids to exert any meaningful influence on the extent of NEP formation in our experimental systems via direct sorption of phenol and/or its dissolved coupling products. HRP-mediated phenol coupling reactions were conducted in solids-free control systems and systems containing one of the four model solids studied. The yield of nonextractable products (NEP) formed in each system was then measured as described in the Experimental Section. Figure 3 presents molar equivalent phenol concentrations for NEP formed in systems having a constant reaction condition but containing different types of solids at varied solid/water ratios. As illustrated, NEP formation is enhanced by increasing solid/ water ratios in the case of each solid investigated. For cellulose and silica sand (Figure 3A), NEP formation initially increases sharply with increasing solid/water ratios up to a value of approximately 10 g/L and then either levels off or increases more modestly at higher solid/water ratios. For PMS (Figure 3A), NEP continues to increase significantly with increasing solid/water ratio until, at the highest ratio tested (200 g/L), a yield of 414 µM equivalents of NEP is obtained. This

FIGURE 3. NEP formation in systems containing cellulose, silica sand, PMS in part (A), and lignin in part (B) and having the same reaction condition (HRP dosage ) 0.5 unit/mL, initial phenol concentration ) 500 µM and initial H2O2 concentration ) 2 mM). Data points are the means of triplicate experiments and represent 1 SD error bars.

FIGURE 4. Enzyme inactivation rate coefficients for systems containing cellulose, silica sand, PMS in part (A), and lignin in part (B), all initially containing 500-µM phenol, 2-mM H2O2, and 0.5-unit/mL HRP. Data points are the means of triplicate experiments and represent 1 SD error bars. accounts for over 80% of the 500-µM phenol originally added to the system and represents nearly a 4-fold increase of NEP formation over that observed in a solid free reaction system, in which 116.4-µM of NEP was formed and a total of 179.9 µM of phenol was transformed to either dissolved or precipitated forms. Of the four solids studied, the influence of lignin on NEP formation is the most significant, as evidenced in Figure 3B. Comparison of parts A and B in Figure 3 shows that an increase in NEP production comparable to that of PMS was obtained by addition of a two-order-ofmagnitude lower mass of lignin. As noted above, the enhanced NEP formation in the presence of solids evident in Figure 3 cannot be attributed to physical-sorption phenomena. Mitigation of Enzyme Inactivation. The impacts of solids on the enzymatic-coupling reactions observed may be attributable in large measure to another important phenomenon noted in the experiments; namely, mitigation of enzyme inactivation. It is known that HRP inactivation during phenolcoupling essentially limits the extent of the reaction. Three potential mechanisms for such inactivation have been proposed; i.e., (i) attack by phenoxy radicals (1, 15), (ii) sideeffects caused by excess H2O2 (48, 49), and (iii) sorption/ occlusion of HRP by precipitated products formed in the coupling reactions (22). As noted earlier in the Experimental Section, our experimental conditions were selected at a region where H2O2-related side effects are insignificant. The third HRP-inactivation mechanism is also believed to be insig-

nificant because the amounts of precipitated product formed under the reaction conditions employed were of the milligram magnitude, and negligible HRP sorption/occlusion by this relatively low quantity of precipitate was verified in our earlier tests (45). Thus, the first of the three mechanisms mentioned above appears to have been the primary cause of HRP inactivation observed in the experimental systems described here. HRP activities were measured throughout the reactions carried out in systems containing different solids at varied solid/water ratios, as described in the Experimental Section. The time course of HRP activity decline in these systems was found to be reasonably well described by an apparent secondorder rate expression developed in one of our earlier studies (29); i.e.,

rin ) -

d[E] ) k′in[E]2 dt

(1)

in which [E] represents the active enzyme concentration and k′in the rate coefficient for apparent second-order enzyme inactivation. Values of k′in obtained from the fitting of eq 1 to the time course data of enzyme activities are presented in Figure 4. As illustrated, k′in values decrease as functions of solid/water ratios, indicating solids mitigation of enzyme inactivation. HRP inactivation was mitigated most significantly by lignin (Figure 4B) and to lesser extents by cellulose, silica sand, and PMS (Figure 4A). VOL. 38, NO. 19, 2004 / ENVIRONMENTAL SCIENCE & TECHNOLOGY

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FIGURE 5. Sorption of native HRP, Compound I, and Compound II on cellulose (A), silica sand (B), PMS (C), and lignin (D). [E]aq stands for enzyme concentration remaining in the aqueous phase. Data points are the means of triplicate experiments and represent 1 SD error bars. HRP Sorption on Solid Materials and Its Relationship to Enzyme Inactivation. As discussed in the Introduction, HRP mediates phenol oxidation via a catalytic cycle involving successive conversions of the enzyme to three different forms; i.e., native HRP is converted first to Compound I and then to Compound II, followed by reversion to the native HRP form. Because the three HRP forms differ from each other in molecular configuration and electronic charge (9), it can be expected that some differences exist in the sorption behaviors of the three enzyme forms. As delineated in the Experimental Section, sorption of each of the three forms individually on each of the four solids studied was evaluated in the current study. That data are presented in Figure 5 in terms of residual aqueous phase enzyme concentration ([E]aq) as a function of solid/water ratio. It should be noted that this figure represents the sorption resulting after only one minute of mixing of each enzyme solution with a solid. An important feature of the HRP catalytic cycle is that the formations of Compound I and Compound II occur much more rapidly than does the reversion of Compound II to native HRP, thus a pseudo-steady-state distribution of enzyme intermediates in which Compound II dominates is quickly developed and maintained during the enzymatic reaction (15, 16). The total sorption of all active forms of HRP existing during an enzymatic reaction should thus be reasonably represented by the sorption behavior of Compound II. Figure 5 clearly indicates that each of the three solid materials (i.e., cellulose, silica sand, and lignin) having the ability to mitigate enzyme inactivation sorbs HRP, whereas PMS does not. Each sorption represents an association 5242

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between the enzyme and the solid material. Such enzymesolid associations may have been formed in a way that reduces the susceptibility of the enzyme to radical attack by physically shielding the enzyme’s turn-off site or adjusting the enzyme conformation, thereby leading to mitigation of enzyme inactivation. It must be noted that the sorption of HRP on solid phases may not quantitatively account for the entire enzyme associations that are operative in a system to protect the enzyme. Such associations may also result from interactions of soluble substances dissolved from the solid materials to form complexes with the enzyme. It is interesting also to note that HRP transfer from the aqueous phase to the solid phase is not accompanied by a decrease in the catalytic efficiency of the enzyme. For example, the fraction of HRP remaining in the aqueous phase drops from around 86% to 1% when the cellulose/water ratio increases from 10 to 200 g/L, whereas the enzymatic reaction performance remains essentially stable for the same range of conditions, as evident in Figure 3. A similar phenomenon is observed also with silica sand, suggesting that the sorption of HRP by these solids imposes no discernible effect on the catalytic activity of the enzyme. It is not uncommon for an enzyme immobilized by physical sorption to retain its full catalytic activity if the sorption of the enzyme does not disturb the structure of its catalytic center (50). It should be reiterated, however, that the experimental conditions for this study were selected so that phenol sorption by the solids was insignificant. If the solids adsorb phenol strongly and reduce its aqueous concentration to a much lower level, the HRP may operate in a phenol-unsaturated mode, leading to a reduction

in performance. This may in fact have been the case in several earlier studies involving soil slurries at very high soil/water ratios (42, 43). Each of the three solids observed to sorb HRP and mitigate enzyme inactivation comprises a relatively hydrophilic polymeric matrix containing oxygen atoms. PMS, the only material not observed to exhibit such effects, is a hydrophobic polymeric matrix containing no oxygen atoms. This suggests that HRP may form associations most readily with hydrophilic materials containing oxygen atoms, which seems logical in that HRP is itself a polypeptide molecule containing abundant amino and carboxylic functionalities (9, 10). Mitigation of HRP inactivation has been observed also in previous studies with such organic additives as poly(ethylene glycol) (PEG) and gelatin (22, 27, 51, 52), both of which are oxygencontaining hydrophilic materials. In an earlier study of phenol coupling in the presence of different soils we further observed that relatively hydrophilic humic-containing soils mitigate HRP inactivation more effectively than kerogen-containing natural geosorbents (29). Cross-Coupling Reactions in PMS-Containing Systems. While mitigation of HRP inactivation may be partially responsible for increased NEP formation in the presence of some solids, it cannot account for all of the enhancement portrayed in Figure 3. NEP formation was enhanced significantly in the presence of PMS for example, while, as shown in Figure 4A, this solid exhibited no appreciable mitigation of HRP inactivation. These observations regarding the behavior of PMS are interesting indeed, for, as discussed below, they provide potential insight to the mechanisms involved in enzymatic coupling processes. The significant enhancement of NEP formation observed in PMS-containing systems may have resulted principally from reactive interactions between PMS and phenoxy radicals generated in the enzymatic processes. As discussed earlier, two sequential stages are involved in HRP-mediated phenol coupling reactions in solids-free systems. Phenol is first oxidized to form phenoxy radicals in an enzymatic reaction stage, and the radicals can then couple with each other in post enzymatic reactions to yield precipitable polymeric NEPs. In light of such a reaction scheme, PMS may hypothetically affect the process in either or both of the two reaction stages; i.e., (i) PMS may interact with the enzyme to increase its catalytic efficiency and thus generate more phenoxy radicals in the enzymatic reaction stage, and/or (ii) PMS may react with phenoxy radicals in the postenzymatic reaction stage to form covalently bonded products, thus providing an additional and alternative NEP formation pathway to that of phenol self-coupling. The first hypothesis can be ruled out for two reasons. First, no direct interaction between the enzyme and PMS was observed, as indicated by the lack of enzyme adsorption on PMS shown in Figure 5. Second, an enhanced radical generation process would logically lead to increased enzyme inactivation, which contradicts the fact that HRP inactivation remained essentially unchanged in the presence of PMS (Figure 4A). It is thus believed that PMS may have increased NEP formation primarily by postenzymatic reactions with phenoxy radicals. Phenoxy radicals generated in enzymatic reactions are highly reactive and thus able to attack and react with relatively inert materials, such as PMS, to form bound residuals. Such reactions may occur first by radical attack on and activation of the inert substrates through such mechanisms as hydrogen abstraction and free-radical addition followed by radical cross-coupling. Cross-coupling reactions between phenoxy radicals and such inert chemicals as polychlorinated biphenyls (PCBs) and polycyclic aromatic hydrocarbons (PAHs) have been reported previously for peroxidase-mediated phenol reaction systems (1, 28).

The enhancement of NEP formation by radical-scavenging organic solids such as PMS can be argued within the framework of enzymatic coupling mechanisms understood to date. As noted earlier, the HRP-mediated radical production process has recently been proven to be reversible (18); i.e., radicals generated in the second and third steps of the catalytic cycle can abstract electrons from enzyme intermediates via “reverse electron transfer” and thus revert to the original substrates (18). Thus, in a solids-free and selfcoupling dominated reaction system, radical-radical coupling essentially competes with the radical disappearance pathway of reverse electron transfer to form NEPs. When a reactive organic solid matrix such as PMS is included in the system, phenoxy radicals may react with this matrix, providing an additional pathway by which NEP formation can compete with reverse electron transfer, and enhancing NEP production. A total of three different pathways for radical disappearance are thus operative in a HRP-mediated phenol coupling reaction system containing a solid-phase radical scavenger such as PMS; i.e., reverse electron transfer, crosscoupling with PMS, and self-coupling of radicals. The fact that HRP inactivation by radical attack is not affected by the addition of PMS (Figure 4A) indicates that the pseudo-steadystate radical concentration remains unaffected. This suggests that a pathway other than cross-coupling dominates radical disappearance. The self-coupling pathway must be slower than that of cross-coupling, for otherwise the enhancement of NEP formation in the presence of PMS (Figure 3A) would not be clearly observed. It can thus be postulated that reverse electron transfer virtually dominates radical disappearance in a HRP-mediated phenol coupling reaction system, essentially controlling the pseudo-steady-state radical concentration. Quantitative Analysis. In light of the information presented above, we conclude that cross-coupling between phenol and PMS is responsible for essentially all of the NEP enhancement observed in the presence of this solid. It may be possible that such cross-coupling also occurred in the systems containing other solids and may to a certain extent have contributed to the increased NEP production observed in Figure 3. This possibility begs a more detailed quantitative analysis, as attempted below. NEP production in peroxidase-mediated phenol coupling systems that do not contain reactive solids can be attributed solely to phenol self-coupling. It has been observed under a variety of different reaction conditions, and is taken widely as a “rule of thumb”, that the outcomes of peroxidasemediated oxidative coupling in solids-free systems generally correlate directly to enzyme consumption during the course of the reaction (15, 17, 53). This suggests the relationship given in eq 2 between rates of radical self-coupling and enzyme inactivation

ΦTN ) -

d[NEP]s d[E]

)-

d[NEP]s/dt d[E]/dt

)

rs rin

(2)

where rs and rin respectively denote rates of self-coupling and enzyme inactivation and ΦTN represents a turnover capacity coefficient defined as the ratio between NEP formation by self-coupling (d[NEP]s) and enzyme consumption (- d[E]). As noted earlier, HRP inactivation was observed to occur relatively quickly in our study and was complete within the 2-h reaction period of the experiments. For this condition, integration of eq 2 over the reaction period gives

[NEP]s ) ΦTN[E]0

(3)

where [E]0 represents the initial enzyme concentration. As illustrated in Figure 6, the linear correlation between NEP formation and initial enzyme concentration indicated VOL. 38, NO. 19, 2004 / ENVIRONMENTAL SCIENCE & TECHNOLOGY

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FIGURE 6. NEP formation in solids free reaction systems having varied initial enzyme dosages, an initial phenol concentration of 500 µM, and an initial H2O2 concentration of 2 mM. Data points are the means of triplicate experiments and represent 1 SD error bars.

by ΦTN ) 382.7(k′in)-1 in Figure 7 represents an independent prediction from the inverse correlation between ΦTN and k′in, in which the coefficient 382.7 was estimated from the measured values of ΦTN ) 234.8 nmole/unit and k′in ) 1.63 mL/unit-min for the solids-free system. It is evident from Figure 7 that turnover capacities in systems containing cellulose and silica sand are predicted well by the correlation between ΦTN and (k′in)-1, indicating that cross-coupling between phenol and solids is insignificant in these systems and that NEP formation results principally from phenol selfcoupling reactions. Conversely, it is evident that both selfand cross-coupling reactions are operative and contribute directly to NEP formation in systems containing PMS and lignin. Cross coupling reactions between different soluble substrates have been studied extensively (19, 38-41), but the data presented in Figure 7 and associated analyses of reaction behaviors provide important new evidence that such reactions can occur as well between dissolved substrate and certain types of solid organic materials. The implications of such reactions for potential applications of enzymatic coupling for remediation of soil/sediment environments are significant. Among the four solids investigated, the structures of the two (PMS and lignin) that participated in cross-coupling each contain unsaturated aromatic moieties, whereas the other two do not. It may thus be inferred that aromatic structures or unsaturated C-C bonds must be present in the SOMs of soils and sediments for phenoxy radical attack and subsequent attachment in cross-coupling reactions to occur. A preliminary assessment of SOM aromaticity may therefore be used as an indicator of anticipated enzymatic crosscoupling feasibility in soil/sediment remediation applications.

Acknowledgments

FIGURE 7. Relationships between turnover capacity coefficients ΦTN and (k′in)-1 for systems containing different solids. in eq 3 is confirmed by experimental data collected for the solids-free system. The data regression represented by the solid line in Figure 6 corresponds to a ΦTN value of 234.8 nmole/unit. The spectroscopic measurements described in the Experimental Section indicate that one unit of enzyme molecularly comprises approximately 0.052 nmole of HRP, thus ΦTN can be expressed in dimensionless form as 4.5 × 103. Further analyses based on eqs 1, 2, and 3 can be used to evaluate more quantitatively the relationship between enzyme inactivation mitigation and NEP formation enhancement. Equations 1 and 2 infer that ΦTN is inversely related to the enzyme inactivation rate coefficient (i.e., ΦTN ∼ (k′in) - 1), indicating that turnover capacity is enhanced as enzyme inactivation is mitigated. Total NEP production can be attributed solely to phenol self-coupling if no cross-coupling occurs, and the effects of enzyme-inactivation mitigation can be predicted using the linear relationship between ΦTN and (k′in)-1. The data in Figures 3 and 4 can thus be reorganized in the form presented in Figure 7, in which turnover capacity ΦTN measured for different solids-containing systems are plotted against the inverse of the k′in values measured for those same systems. The linear trace defined 5244

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We thank Tighe B. Herren, Carl W. Lenker, and Sung Ho Kim for diligent efforts and important contributions in performance of the experimental work reported and Thomas Yavaraski for invaluable assistance with the instrumental analyses. The research was supported in part by Grant No. DE-FG07-02ER63488 from the Environmental Management Science Program of the United States Department of Energy (DOE) and in part by Research Grant P42ES04911-14 from the National Institutes for Environmental and Health Sciences. The content of the paper does not necessarily represent the views of either funding agency.

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Received for review February 2, 2004. Revised manuscript received July 12, 2004. Accepted July 13, 2004. ES049826H

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