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Degradation of phenol using tyrosinase immobilized on siliceous supports
Water Research 37 (2003) 436–440
Degradation of phenol using tyrosinase immobilized on siliceous supports Gayathri B. Seetharam, Bradley A. Saville* Department of Chemical Engineering and Applied Chemistry, University of Toronto, 200 College Street, Toronto,Ont., Canada M5S 3E5 Received 1 July 2000; received in revised form 1 January 2001; accepted 14 June 2002
Abstract The degradation of phenol by tyrosinase immobilized on chemically modified sodium aluminosilicate (NaA), calcium aluminosilicate (CaA), and silica gel was studied. Phenol conversion by immobilized tyrosinase ranged between B15% and 60%, depending upon the initial phenol concentration, pH, and enzyme loading. Tyrosinase immobilized on CaA and on NaA could be re-used repeatedly without any decrease in performance. However, in studies at pH 8.0, significant enzyme inhibition was observed, since phenol conversion was rapid for B20 min, then reached a plateau. The inhibition was reversible; activity was restored upon placing the immobilized enzyme in fresh substrate. Reducing the pH to 6.8 from 8.0 led to higher conversion of phenol, and decreased the inhibition of the immobilized enzyme. r 2002 Elsevier Science Ltd. All rights reserved. Keywords: Phenol; Tyrosinase; Wastewater; Immobilized enzymes
1. Introduction Phenol is present in the wastewater from a number of industries, including coal conversion, petroleum refinery, mining, and dressing operations, and facilities producing resins, plastics, dyes and other organic chemicals, textiles, timber, and pulp and paper [1–3]. Phenol ranks among the top 25 chemicals with the largest waste transfers in North America [4], and ranks among the 25 chemicals with the largest national pollutant release inventory and toxic release inventory (TRI) transfers. It is also among the top 10 TRI chemicals released to sewage or Publicly Owned Treatment Works. Virtually all phenols are toxic; furthermore, phenol and many of its derivatives are hazardous pollutants. *Corresponding author. Tel.: +1-416-978-7745; fax: +1416-978-8605. E-mail address: [email protected] (B.A. Saville).
Phenol concentrations greater than 50 ppb are toxic to some forms of aquatic life and ingestion of 1 g of phenol can be fatal in humans. Hence, due to their abundance and toxicity, removal of phenols from industrial aqueous effluents is an important practical problem. Conventional methods for dephenolization of industrial wastewaters include solvent extraction, deep-well injection, microbial degradation, adsorption on activated carbon and chemical oxidation [2,5]. Solvent extraction methods are expensive, and deep-well injection may lead to contamination of ground water. Adsorption and oxidation treatments become exceedingly expensive when low effluent concentrations must be achieved. Microbiological treatment has shown great promise, but for small volume wastes generated discontinuously, microbiological treatment has been plagued by instabilities caused by toxicity of these compounds to the microbial population. Therefore, alternative technologies have to be explored. Enzymatic treatment of phenolics was initially investigated by Munnecke [6] and Klibanov [7]. A
0043-1354/02/$ - see front matter r 2002 Elsevier Science Ltd. All rights reserved. PII: S 0 0 4 3 - 1 3 5 4 ( 0 2 ) 0 0 2 9 0 - 7
G.B. Seetharam, B.A. Saville / Water Research 37 (2003) 436–440
particular advantage of enzymatic treatment is the high specificity of enzymes for their substrates, unlike microbiological processes, which tend to be nonselective. However, enzymes can be readily inactivated, dramatically increasing the cost of an enzyme-based process for phenol removal. Tyrosinase, one enzyme suitable for treatment of phenolic wastes, catalyses two consecutive reactions via separate active sites: (1) hydroxylation of monophenols with molecular oxygen in the presence of a chemical reductor (AH2) to form ortho-diphenols, and (2) dehydrogenation of orthodiphenols with molecular oxygen to form ortho-quinones. Therefore, the overall process catalyzed by tyrosinase can be described by monophenol þ O2 þ AH2 -o-diphenol þ H2 O þ A;
ð1Þ
o-diphenol þ 1=2O2 -o-quinone þ H2 O:
ð2Þ
Quinones are usually formed rapidly, and undergo non-enzymatic conversion to form more stable intermediates. These intermediates subsequently undergo slow oligomerization reactions that ultimately yield high molecular weight, insoluble polyphenolics. Atlow et al. [8] used soluble tyrosinase to remove phenol from an aqueous synthetic waste solution; up to 99% conversion of the phenol was obtained, although, at higher concentrations (1.0 g/L), conversion was limited by inactivation of tyrosinase, likely by quinones formed during the reaction. Sun et al. [9] reported significant inactivation of soluble tyrosinase when phenol levels exceeded 0.05 g/L, likely because the quinones reacted with the free amino groups of the enzyme. Hence, they suggested the use of chitosan to adsorb quinones generated during phenol degradation. Given that immobilization has the potential to increase enzyme stability, various researchers have examined the effectiveness of immobilized tyrosinase for phenol degradation. Wada et al. [10] immobilized tyrosinase on magnetite and dianon WK-20, and obtained virtually complete conversion of a 0.02 g/L phenol solution. Kazandjian and Klibanov [11] and Estrada et al. [12] noted that phenols dissolved in chloroform could be degraded using tyrosinase immobilized on glass beads. Although enzyme stability was improved by immobilization or by incorporation in organic solvents (half-life of several hours versus several minutes for tyrosinase in aqueous solution), further improvements are required. The objectives of this study were, therefore, (1) to develop an immobilized form of tyrosinase which was stable, and (2) to establish the efficacy of the immobilized tyrosinase for removal of phenol from synthetic wastewater. The stability of the immobilized enzyme and
437
the kinetics of phenol removal were assessed via repeated batch experiments, over a range of phenol concentrations.
2. Materials and methods 2.1. Chemicals and equipment Mushroom tyrosinase (T7755), silica gel, and phenol were obtained from Sigma (Oakville, Ont.). Glutaraldehyde (GA) (25% w/v), sodium phosphate, and methanol (HPLC grade) were obtained from Fisher (Unionville, Ont.). Crystalline calcium aluminosilicate (CaA) and sodium aluminosilicate (NaA) were obtained from The PQ Corporation (Conshohocken, PA). All other chemicals used were of analytical grade. Stock enzyme solution was prepared by dissolving either 10,800 or 21,600 U of tyrosinase into 0.05 M phosphate buffer (pH 6.5) to make a 100 mL solution. 2.2. HPLC assay A Shimadzu liquid chromatograph was used for HPLC analysis. Samples were analyzed using an eluant flow rate of 1.5 mL/min, and UV detection at 280 nm. For elution, a mobile phase of 1% (v/v) acetic acid in methanol (35% v/v) and HPLC water (65% v/v) was prepared. A Supelcosil LC-8 column (5 mm; 15 cm 4.6 mm ID; Supelco, Oakville, Ont.) was used to separate phenol and reaction products from the internal standard, nitrophenol. Phenols and o-quinones were detected as separate peaks. The standard curve was linear ðr2 ¼ 0:99Þ for phenol concentrations up to 1.0 g/L. 2.3. Immobilization of tyrosinase onto zeolites The two supports used, CaA and NaA, are identical in their basic structures. Owing to the isomorphic substitution of aluminum by silicon, the three-dimensional oxygen framework carries an excess negative charge, compensated by cations. The only difference between these supports is this substituted cation, calcium in CaA and sodium in NaA. A previously patented procedure was adapted for immobilization of tyrosinase [13]. The supports were modified in GA/phosphate buffer solution; the concentration of GA and ratios of support mass to buffer volume were adjusted as required, to ‘‘optimize’’ the enzyme uptake. The mixture was gently stirred at room temperature, then washed with HPLC grade water. The modified support was recovered by vacuum filtration, and dried overnight before incubation in the tyrosinase stock solution. For example, 80 mg of modified support would be incubated with 20 mL of 108 U/mL enzyme
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solution. During incubation, the solution was gently mixed. The immobilized enzyme was recovered by vacuum filtration, then dried before use in studies of phenol degradation. Samples of soluble enzyme were collected from the enzyme solution before and after immobilization, and assayed for enzyme activity. The ratio of the activity in the ‘‘final’’ solution relative to that in the ‘‘initial’’ solution was used to estimate the enzyme uptake onto the support. The average uptake was 37% (range 33–45%). 2.4. Immobilization of tyrosinase onto silica gel Two grams of support was modified with either 100 or 250 mL of GA/phosphate buffer solution. The mixture was gently stirred, then washed with HPLC-grade water. The modified support was recovered by vacuum filtration, then dried. The modified support was subsequently added to a 3240 U tyrosinase stock solution and gently mixed for up to 48 h. The immobilized enzyme was recovered by vacuum filtration, then dried before use in studies of phenol degradation. Using samples collected from the enzyme solution before and after immobilization, the average enzyme uptake was 62%. 2.5. Investigations of phenol degradation A substrate solution was prepared by dissolving sufficient phenol in 0.05 M phosphate buffer (pH 8.0 or pH 6.8) to produce phenol solutions between 0.10 and 0.40 g/L. The immobilized enzyme was incubated in phenol solution at room temperature, and stirred gently to keep the immobilized enzyme suspended. Care was taken to avoid excessive shear. Three hundred microliter aliquots were collected at suitable intervals, and drawn through a 0.5 mm syringe filter to separate the support from the liquid. This prevented further reaction, and ensured that the immobilized enzyme did not interfere with the HPLC assay. An equal volume of 0.20 g/L 2-nitrophenol (internal standard) was added to the sample vial; samples were refrigerated, then assayed using HPLC.
concentration, confirming that conversion was solely due to immobilized tyrosinase.
3. Results and discussion 3.1. Dephenolization by tyrosinase immobilized to CaA Studies were conducted at pH 8.0, using initial (preimmobilization) enzyme loadings of either 1080 or 2160 U. With an average uptake of 37%, this corresponds to approximately 400 and 800 U of immobilized enzyme, respectively. In each study, four consecutive batches were conducted with the same enzyme; between batches, the immobilized enzyme was recovered by vacuum filtration. In studies with 400 U of tyrosinase on CaA, the initial phenol concentration was 0.40 g/L, and samples were collected every 10 min for 50 min. The average conversion for the 4 batches was 15%, ranging from 12.5% during batch 1, to 19% during batch 4. Thus, there was no evidence of loss of enzyme activity over the four consecutive uses of the enzyme. However, the fractional conversion was lower than desired, and thus, for the following study, the enzyme concentration was increased and the initial phenol concentration was reduced. In the second set of studies, 800 U of immobilized tyrosinase were used, and the initial phenol concentration was reduced to 0.10 g/L. The immobilized enzyme was again used four times, recovered by filtration following the first, second, and third batches. The average conversion over 4 batches was 25%; conversion profiles are shown in Fig. 1. The results from these studies also confirm that tyrosinase immobilized on CaA is stable. However, it is also noteworthy that much of the observed conversion occurs within the first 10–20 min of reaction; the conversion increases very
2.6. Control studies Control studies were undertaken to assess phenol degradation upon exposure to the support alone, and to determine if tyrosinase desorbed from the support, and contributed to phenol degradation. In the first case, no conversion of phenol was observed when phenol was incubated in the support alone. In the second control study, samples from an immobilized enzyme trial were processed in the usual way to remove immobilized enzyme, and assayed immediately. The samples were then re-analyzed 48 h later; there was no change in
Fig. 1. Phenol conversion at pH 8.0 using tyrosinase immobilized on CaA. The results from 4 consecutive batches, each in fresh substrate, are shown. The initial phenol concentration in each batch is 0.10 g/L.
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little between 20 and 50 min (Fig. 1). Since the enzyme activity is restored upon removing the enzyme from the substrate solution, it is likely that the observed ‘‘plateau’’ in the conversion profile is due to (reversible) enzyme inhibition, rather than irreversible inactivation. After filtration of the immobilized enzyme from the substrate solution, brown particles were noted on the recovered enzyme, consistent with the presence of quinones or polyphenolic species, which are known to be colored. The observed color change is also consistent with other studies with soluble tyrosinase, in which the originally clear solution turned brown as the reaction proceeded. 3.2. Comparison of tyrosinase immobilized to CaA and NaA In these studies, 800 U of immobilized tyrosinase (on either CaA or to NaA) were used in two repeated batch reactions in a 50 mL vessel, using 0.30 g/L phenol for each run. Samples were collected every 20 min for 160 min. The conversion profiles for these studies were qualitatively similar to the profiles in Fig. 1; most of the conversion occurred in the first 20–30 min. Essentially, the same final conversion was obtained for each batch, and for each support (35% and 36% for batches 1 and 2 with tyrosinase on CaA, vs. 34% and 34% for tyrosinase on NaA). Thus, tyrosinase immobilized on CaA and on NaA exhibits the same performance characteristics and enzyme stability. Thus, it has been demonstrated that tyrosinase is stable when immobilized on either CaA or NaA, and that repeated use is possible. However, the phenol conversion is less than desired likely limited by inhibition by the quinone intermediate/product. To reduce the effect of inhibition, the concentration of active enzyme must be increased relative to the concentration of inhibitor. This could be accomplished by adjusting the pH. 3.3. Effect of pH on phenol conversion In previous studies, Atlow et al. [8] observed significant conversion of phenol at pH 8.0, likely because quinone removal by polymerization is rapid at pH 8.0. However, the optimum activity for tyrosinase occurs near pH 6.5. If pH has a greater effect on tyrosinase activity than on quinone polymerization, some improvement in phenol conversion may be obtained by reducing the pH, since the active enzyme concentration may be enhanced relative to the concentration of quinone in the system. Phenol degradation was therefore studied at pH 6.8 and at pH 8.0. In each trial, 50 mL of 0.1 g/L phenol was incubated with 540 U of soluble enzyme for 5 h. The
439
reaction fluid turned brown or beige after approximately 1.5 h of processing at pH 6.8. A similar color, although less intense, was observed in the reactant fluid at pH 8.0. Both solutions turned dark brown, almost black, towards the end of reaction. The observed color changes are consistent with previous observations from reactions with immobilized tyrosinase. There was little difference in phenol conversion over the first 3 h, but by the end of the 5 h trial, 93% removal of phenol was observed at pH 6.8 versus 75% conversion at pH 8.0. Thus, it can be concluded that the performance of this form of tyrosinase is enhanced at pH 6.8. 3.4. Dephenolization using tyrosinase immobilized on silica gel In order to conduct studies with immobilized tyrosinase at pH 6.8, it was necessary to change supports, since NaA and CaA undergo gradual hydrolysis under (even slightly) acidic pH. Consequently, silica gel was used as the support for these studies; with an uptake of 62%, approximately 2000 U of tyrosinase were transferred to the support. Immobilized tyrosinase was incubated in a 25 mL solution of 0.1 g/L phenol at pH 6.8 for about 24 h. Approximately 14% conversion was obtained after 6 h; this increased to an average of 37% after 24 h. In a third trial conducted over 75 h (all other conditions the same), 58% conversion was obtained. These results suggest that conversion can be improved by operation at pH 6.8. Furthermore, increasing the reaction time led to increased conversion, unlike in previous studies with tyrosinase immobilized to either CaA or NaA, in which the conversion levelled off after 20 min of reaction. Thus, the effect of enzyme inhibition was reduced under these conditions. This suggests that the products of quinone polymerization may be responsible for tyrosinase inhibition, since these polymerization products are less prevalent at pH 6.8. Removal of these polymerization products may be the key to improving tyrosinase performance at pH 8.0. A combination of factors may therefore be responsible for the improved phenol conversion at pH 6.8. First, the optimum activity for tyrosinase occurs near pH 6.5, and the immobilized enzyme is expected to be more active at pH 6.8 than at pH 8.0. The reduction in enzyme inhibition observed at pH 6.8 would also improve performance, since there would be more enzyme freely available to react with phenol.
4. Conclusions Tyrosinase immobilized on these siliceous supports was able to remove between B15% and 60% of the phenol in solution, depending upon the initial phenol
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concentration, the enzyme loading, pH, and duration of reaction. The immobilized enzyme was stable. Tyrosinase immobilized on CaA and on NaA could be re-used repeatedly without any decrease in performance. However, in studies at pH 8.0, phenol degradation profiles consistently ‘‘leveled off’’ after approximately 20 min of rapid removal. Since the immobilized enzyme could be reused, it is apparent that this ‘‘leveling off’’ is due to strong inhibition, a reversible process. Removal of the inhibitor seems to be the key to ensuring higher conversion of phenol. In studies with soluble tyrosinase, phenol conversion reached 93% at pH 6.8, versus 75% at pH 8.0. The effect of enzyme inhibition was reduced by conducting reactions with immobilized tyrosinase at pH 6.8.
Acknowledgements The authors gratefully acknowledge financial support from NSERC, National Silicates Ltd., and from the University of Toronto.
References [1] Dailey NS. Process effluents: quantities and control technologies. In: Braumstein HM, Copenhaver ED, Pfuderer HA, editors. Environmental health and control aspects of coal conversion, vol. 1. TN: Oak Ridge National Laboratory, 1977 [Chapter 4].
[2] Lanouette KH. Treatment of phenolic wastes. Chem Eng 1977;84:99–106. [3] US Environmental Protection Agency, 1980. Industrial Wastewater, Report No. EPA-600/8-80-026. [4] Taking Stock, North American Pollutant Releases and Transfers. Commission for Environmental Cooperation, 1994. [5] Sittig M. How to remove pollutants and toxic pollutants from air and water. Noyes Data Corp.: Park Ridge, NJ, 1977. p. 430–3. [6] Munnecke DM. Enzymatic hydrolysis of organophosphate insecticides, a possible pesticide disposal method. Appl Environ Microbiol 1976;32:7–13. [7] Klibanov AM, Alberti BN, Morris ED, Felshin LM. Enzymatic removal of toxic phenols and anilines from wastewaters. J Appl Biochem 1980;2:414–21. [8] Atlow SC, Bonadonna-Aparo L, Klibanov AM. Dephenolization of industrial wastewaters catalyzed by polyphenol oxidase. Biotechnol Bioeng 1984;26:599–603. [9] Sun W-Q, Payne GF, Moas MSGL, Chu JH, Wallace KK. Tyrosinase reaction/chitosan absorption for removing phenols from wastewater. Biotechnol Prog 1992;8:179–96. [10] Wada S, Ichikawa H, Tatsumi K. Removal of phenols from wastewater by soluble and immobilized tyrosinase. Biotechnol Bioeng 1993;42:854–8. [11] Kazandjian RZ, Klibanov AM. Regioselective oxidation of phenols catalyzed by polyphenol oxidase in chloroform. J Am Chem Soc 1985;107:5448–50. [12] Estrada P, Sanchez-Munez R, Acebal C, Arche R, Castillon MP. Characterization and optimization of immobilized polyphenol oxidase in low water organic solvents. Biotech Appl Biochem 1991;14:12–20. [13] Le Fevre GN, Saville BA. Enzyme immobilization on a siliceous support with a polyaldehyde cross-linking agent. US Patent # 5,998,183, December 7, 1999.
Degradation of phenol using tyrosinase immobilized on siliceous supports Gayathri B. Seetharam, Bradley A. Saville* Department of Chemical Engineering and Applied Chemistry, University of Toronto, 200 College Street, Toronto,Ont., Canada M5S 3E5 Received 1 July 2000; received in revised form 1 January 2001; accepted 14 June 2002
Abstract The degradation of phenol by tyrosinase immobilized on chemically modified sodium aluminosilicate (NaA), calcium aluminosilicate (CaA), and silica gel was studied. Phenol conversion by immobilized tyrosinase ranged between B15% and 60%, depending upon the initial phenol concentration, pH, and enzyme loading. Tyrosinase immobilized on CaA and on NaA could be re-used repeatedly without any decrease in performance. However, in studies at pH 8.0, significant enzyme inhibition was observed, since phenol conversion was rapid for B20 min, then reached a plateau. The inhibition was reversible; activity was restored upon placing the immobilized enzyme in fresh substrate. Reducing the pH to 6.8 from 8.0 led to higher conversion of phenol, and decreased the inhibition of the immobilized enzyme. r 2002 Elsevier Science Ltd. All rights reserved. Keywords: Phenol; Tyrosinase; Wastewater; Immobilized enzymes
1. Introduction Phenol is present in the wastewater from a number of industries, including coal conversion, petroleum refinery, mining, and dressing operations, and facilities producing resins, plastics, dyes and other organic chemicals, textiles, timber, and pulp and paper [1–3]. Phenol ranks among the top 25 chemicals with the largest waste transfers in North America [4], and ranks among the 25 chemicals with the largest national pollutant release inventory and toxic release inventory (TRI) transfers. It is also among the top 10 TRI chemicals released to sewage or Publicly Owned Treatment Works. Virtually all phenols are toxic; furthermore, phenol and many of its derivatives are hazardous pollutants. *Corresponding author. Tel.: +1-416-978-7745; fax: +1416-978-8605. E-mail address: [email protected] (B.A. Saville).
Phenol concentrations greater than 50 ppb are toxic to some forms of aquatic life and ingestion of 1 g of phenol can be fatal in humans. Hence, due to their abundance and toxicity, removal of phenols from industrial aqueous effluents is an important practical problem. Conventional methods for dephenolization of industrial wastewaters include solvent extraction, deep-well injection, microbial degradation, adsorption on activated carbon and chemical oxidation [2,5]. Solvent extraction methods are expensive, and deep-well injection may lead to contamination of ground water. Adsorption and oxidation treatments become exceedingly expensive when low effluent concentrations must be achieved. Microbiological treatment has shown great promise, but for small volume wastes generated discontinuously, microbiological treatment has been plagued by instabilities caused by toxicity of these compounds to the microbial population. Therefore, alternative technologies have to be explored. Enzymatic treatment of phenolics was initially investigated by Munnecke [6] and Klibanov [7]. A
0043-1354/02/$ - see front matter r 2002 Elsevier Science Ltd. All rights reserved. PII: S 0 0 4 3 - 1 3 5 4 ( 0 2 ) 0 0 2 9 0 - 7
G.B. Seetharam, B.A. Saville / Water Research 37 (2003) 436–440
particular advantage of enzymatic treatment is the high specificity of enzymes for their substrates, unlike microbiological processes, which tend to be nonselective. However, enzymes can be readily inactivated, dramatically increasing the cost of an enzyme-based process for phenol removal. Tyrosinase, one enzyme suitable for treatment of phenolic wastes, catalyses two consecutive reactions via separate active sites: (1) hydroxylation of monophenols with molecular oxygen in the presence of a chemical reductor (AH2) to form ortho-diphenols, and (2) dehydrogenation of orthodiphenols with molecular oxygen to form ortho-quinones. Therefore, the overall process catalyzed by tyrosinase can be described by monophenol þ O2 þ AH2 -o-diphenol þ H2 O þ A;
ð1Þ
o-diphenol þ 1=2O2 -o-quinone þ H2 O:
ð2Þ
Quinones are usually formed rapidly, and undergo non-enzymatic conversion to form more stable intermediates. These intermediates subsequently undergo slow oligomerization reactions that ultimately yield high molecular weight, insoluble polyphenolics. Atlow et al. [8] used soluble tyrosinase to remove phenol from an aqueous synthetic waste solution; up to 99% conversion of the phenol was obtained, although, at higher concentrations (1.0 g/L), conversion was limited by inactivation of tyrosinase, likely by quinones formed during the reaction. Sun et al. [9] reported significant inactivation of soluble tyrosinase when phenol levels exceeded 0.05 g/L, likely because the quinones reacted with the free amino groups of the enzyme. Hence, they suggested the use of chitosan to adsorb quinones generated during phenol degradation. Given that immobilization has the potential to increase enzyme stability, various researchers have examined the effectiveness of immobilized tyrosinase for phenol degradation. Wada et al. [10] immobilized tyrosinase on magnetite and dianon WK-20, and obtained virtually complete conversion of a 0.02 g/L phenol solution. Kazandjian and Klibanov [11] and Estrada et al. [12] noted that phenols dissolved in chloroform could be degraded using tyrosinase immobilized on glass beads. Although enzyme stability was improved by immobilization or by incorporation in organic solvents (half-life of several hours versus several minutes for tyrosinase in aqueous solution), further improvements are required. The objectives of this study were, therefore, (1) to develop an immobilized form of tyrosinase which was stable, and (2) to establish the efficacy of the immobilized tyrosinase for removal of phenol from synthetic wastewater. The stability of the immobilized enzyme and
437
the kinetics of phenol removal were assessed via repeated batch experiments, over a range of phenol concentrations.
2. Materials and methods 2.1. Chemicals and equipment Mushroom tyrosinase (T7755), silica gel, and phenol were obtained from Sigma (Oakville, Ont.). Glutaraldehyde (GA) (25% w/v), sodium phosphate, and methanol (HPLC grade) were obtained from Fisher (Unionville, Ont.). Crystalline calcium aluminosilicate (CaA) and sodium aluminosilicate (NaA) were obtained from The PQ Corporation (Conshohocken, PA). All other chemicals used were of analytical grade. Stock enzyme solution was prepared by dissolving either 10,800 or 21,600 U of tyrosinase into 0.05 M phosphate buffer (pH 6.5) to make a 100 mL solution. 2.2. HPLC assay A Shimadzu liquid chromatograph was used for HPLC analysis. Samples were analyzed using an eluant flow rate of 1.5 mL/min, and UV detection at 280 nm. For elution, a mobile phase of 1% (v/v) acetic acid in methanol (35% v/v) and HPLC water (65% v/v) was prepared. A Supelcosil LC-8 column (5 mm; 15 cm 4.6 mm ID; Supelco, Oakville, Ont.) was used to separate phenol and reaction products from the internal standard, nitrophenol. Phenols and o-quinones were detected as separate peaks. The standard curve was linear ðr2 ¼ 0:99Þ for phenol concentrations up to 1.0 g/L. 2.3. Immobilization of tyrosinase onto zeolites The two supports used, CaA and NaA, are identical in their basic structures. Owing to the isomorphic substitution of aluminum by silicon, the three-dimensional oxygen framework carries an excess negative charge, compensated by cations. The only difference between these supports is this substituted cation, calcium in CaA and sodium in NaA. A previously patented procedure was adapted for immobilization of tyrosinase [13]. The supports were modified in GA/phosphate buffer solution; the concentration of GA and ratios of support mass to buffer volume were adjusted as required, to ‘‘optimize’’ the enzyme uptake. The mixture was gently stirred at room temperature, then washed with HPLC grade water. The modified support was recovered by vacuum filtration, and dried overnight before incubation in the tyrosinase stock solution. For example, 80 mg of modified support would be incubated with 20 mL of 108 U/mL enzyme
438
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solution. During incubation, the solution was gently mixed. The immobilized enzyme was recovered by vacuum filtration, then dried before use in studies of phenol degradation. Samples of soluble enzyme were collected from the enzyme solution before and after immobilization, and assayed for enzyme activity. The ratio of the activity in the ‘‘final’’ solution relative to that in the ‘‘initial’’ solution was used to estimate the enzyme uptake onto the support. The average uptake was 37% (range 33–45%). 2.4. Immobilization of tyrosinase onto silica gel Two grams of support was modified with either 100 or 250 mL of GA/phosphate buffer solution. The mixture was gently stirred, then washed with HPLC-grade water. The modified support was recovered by vacuum filtration, then dried. The modified support was subsequently added to a 3240 U tyrosinase stock solution and gently mixed for up to 48 h. The immobilized enzyme was recovered by vacuum filtration, then dried before use in studies of phenol degradation. Using samples collected from the enzyme solution before and after immobilization, the average enzyme uptake was 62%. 2.5. Investigations of phenol degradation A substrate solution was prepared by dissolving sufficient phenol in 0.05 M phosphate buffer (pH 8.0 or pH 6.8) to produce phenol solutions between 0.10 and 0.40 g/L. The immobilized enzyme was incubated in phenol solution at room temperature, and stirred gently to keep the immobilized enzyme suspended. Care was taken to avoid excessive shear. Three hundred microliter aliquots were collected at suitable intervals, and drawn through a 0.5 mm syringe filter to separate the support from the liquid. This prevented further reaction, and ensured that the immobilized enzyme did not interfere with the HPLC assay. An equal volume of 0.20 g/L 2-nitrophenol (internal standard) was added to the sample vial; samples were refrigerated, then assayed using HPLC.
concentration, confirming that conversion was solely due to immobilized tyrosinase.
3. Results and discussion 3.1. Dephenolization by tyrosinase immobilized to CaA Studies were conducted at pH 8.0, using initial (preimmobilization) enzyme loadings of either 1080 or 2160 U. With an average uptake of 37%, this corresponds to approximately 400 and 800 U of immobilized enzyme, respectively. In each study, four consecutive batches were conducted with the same enzyme; between batches, the immobilized enzyme was recovered by vacuum filtration. In studies with 400 U of tyrosinase on CaA, the initial phenol concentration was 0.40 g/L, and samples were collected every 10 min for 50 min. The average conversion for the 4 batches was 15%, ranging from 12.5% during batch 1, to 19% during batch 4. Thus, there was no evidence of loss of enzyme activity over the four consecutive uses of the enzyme. However, the fractional conversion was lower than desired, and thus, for the following study, the enzyme concentration was increased and the initial phenol concentration was reduced. In the second set of studies, 800 U of immobilized tyrosinase were used, and the initial phenol concentration was reduced to 0.10 g/L. The immobilized enzyme was again used four times, recovered by filtration following the first, second, and third batches. The average conversion over 4 batches was 25%; conversion profiles are shown in Fig. 1. The results from these studies also confirm that tyrosinase immobilized on CaA is stable. However, it is also noteworthy that much of the observed conversion occurs within the first 10–20 min of reaction; the conversion increases very
2.6. Control studies Control studies were undertaken to assess phenol degradation upon exposure to the support alone, and to determine if tyrosinase desorbed from the support, and contributed to phenol degradation. In the first case, no conversion of phenol was observed when phenol was incubated in the support alone. In the second control study, samples from an immobilized enzyme trial were processed in the usual way to remove immobilized enzyme, and assayed immediately. The samples were then re-analyzed 48 h later; there was no change in
Fig. 1. Phenol conversion at pH 8.0 using tyrosinase immobilized on CaA. The results from 4 consecutive batches, each in fresh substrate, are shown. The initial phenol concentration in each batch is 0.10 g/L.
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little between 20 and 50 min (Fig. 1). Since the enzyme activity is restored upon removing the enzyme from the substrate solution, it is likely that the observed ‘‘plateau’’ in the conversion profile is due to (reversible) enzyme inhibition, rather than irreversible inactivation. After filtration of the immobilized enzyme from the substrate solution, brown particles were noted on the recovered enzyme, consistent with the presence of quinones or polyphenolic species, which are known to be colored. The observed color change is also consistent with other studies with soluble tyrosinase, in which the originally clear solution turned brown as the reaction proceeded. 3.2. Comparison of tyrosinase immobilized to CaA and NaA In these studies, 800 U of immobilized tyrosinase (on either CaA or to NaA) were used in two repeated batch reactions in a 50 mL vessel, using 0.30 g/L phenol for each run. Samples were collected every 20 min for 160 min. The conversion profiles for these studies were qualitatively similar to the profiles in Fig. 1; most of the conversion occurred in the first 20–30 min. Essentially, the same final conversion was obtained for each batch, and for each support (35% and 36% for batches 1 and 2 with tyrosinase on CaA, vs. 34% and 34% for tyrosinase on NaA). Thus, tyrosinase immobilized on CaA and on NaA exhibits the same performance characteristics and enzyme stability. Thus, it has been demonstrated that tyrosinase is stable when immobilized on either CaA or NaA, and that repeated use is possible. However, the phenol conversion is less than desired likely limited by inhibition by the quinone intermediate/product. To reduce the effect of inhibition, the concentration of active enzyme must be increased relative to the concentration of inhibitor. This could be accomplished by adjusting the pH. 3.3. Effect of pH on phenol conversion In previous studies, Atlow et al. [8] observed significant conversion of phenol at pH 8.0, likely because quinone removal by polymerization is rapid at pH 8.0. However, the optimum activity for tyrosinase occurs near pH 6.5. If pH has a greater effect on tyrosinase activity than on quinone polymerization, some improvement in phenol conversion may be obtained by reducing the pH, since the active enzyme concentration may be enhanced relative to the concentration of quinone in the system. Phenol degradation was therefore studied at pH 6.8 and at pH 8.0. In each trial, 50 mL of 0.1 g/L phenol was incubated with 540 U of soluble enzyme for 5 h. The
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reaction fluid turned brown or beige after approximately 1.5 h of processing at pH 6.8. A similar color, although less intense, was observed in the reactant fluid at pH 8.0. Both solutions turned dark brown, almost black, towards the end of reaction. The observed color changes are consistent with previous observations from reactions with immobilized tyrosinase. There was little difference in phenol conversion over the first 3 h, but by the end of the 5 h trial, 93% removal of phenol was observed at pH 6.8 versus 75% conversion at pH 8.0. Thus, it can be concluded that the performance of this form of tyrosinase is enhanced at pH 6.8. 3.4. Dephenolization using tyrosinase immobilized on silica gel In order to conduct studies with immobilized tyrosinase at pH 6.8, it was necessary to change supports, since NaA and CaA undergo gradual hydrolysis under (even slightly) acidic pH. Consequently, silica gel was used as the support for these studies; with an uptake of 62%, approximately 2000 U of tyrosinase were transferred to the support. Immobilized tyrosinase was incubated in a 25 mL solution of 0.1 g/L phenol at pH 6.8 for about 24 h. Approximately 14% conversion was obtained after 6 h; this increased to an average of 37% after 24 h. In a third trial conducted over 75 h (all other conditions the same), 58% conversion was obtained. These results suggest that conversion can be improved by operation at pH 6.8. Furthermore, increasing the reaction time led to increased conversion, unlike in previous studies with tyrosinase immobilized to either CaA or NaA, in which the conversion levelled off after 20 min of reaction. Thus, the effect of enzyme inhibition was reduced under these conditions. This suggests that the products of quinone polymerization may be responsible for tyrosinase inhibition, since these polymerization products are less prevalent at pH 6.8. Removal of these polymerization products may be the key to improving tyrosinase performance at pH 8.0. A combination of factors may therefore be responsible for the improved phenol conversion at pH 6.8. First, the optimum activity for tyrosinase occurs near pH 6.5, and the immobilized enzyme is expected to be more active at pH 6.8 than at pH 8.0. The reduction in enzyme inhibition observed at pH 6.8 would also improve performance, since there would be more enzyme freely available to react with phenol.
4. Conclusions Tyrosinase immobilized on these siliceous supports was able to remove between B15% and 60% of the phenol in solution, depending upon the initial phenol
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concentration, the enzyme loading, pH, and duration of reaction. The immobilized enzyme was stable. Tyrosinase immobilized on CaA and on NaA could be re-used repeatedly without any decrease in performance. However, in studies at pH 8.0, phenol degradation profiles consistently ‘‘leveled off’’ after approximately 20 min of rapid removal. Since the immobilized enzyme could be reused, it is apparent that this ‘‘leveling off’’ is due to strong inhibition, a reversible process. Removal of the inhibitor seems to be the key to ensuring higher conversion of phenol. In studies with soluble tyrosinase, phenol conversion reached 93% at pH 6.8, versus 75% at pH 8.0. The effect of enzyme inhibition was reduced by conducting reactions with immobilized tyrosinase at pH 6.8.
Acknowledgements The authors gratefully acknowledge financial support from NSERC, National Silicates Ltd., and from the University of Toronto.
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