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Characterization of tyrosinase for the treatment of aqueous phenols
Bioresource Technology 74 (2000) 191±199
Characterization of tyrosinase for the treatment of aqueous phenols Keisuke Ikehata, James A. Nicell * Department of Civil Engineering and Applied Mechanics, McGill University, 817 Sherbrooke Street West, Montreal, Que., Canada H3A 2K6 Received 7 August 1999; received in revised form 24 January 2000; accepted 30 January 2000
Abstract Mushroom tyrosinase (polyphenol oxidase, EC 1.14.18.1) was investigated as an alternative to peroxidase enzymes for the catalytic removal of phenolic compounds from wastewaters. Maximum catalytic activity was observed at pH 7 and more than 50% of optimum activity was observed at pHs ranging between 5 and 8. Tyrosinase was unstable under acidic conditions and at elevated temperatures. The activation energy for thermal inactivation of tyrosinase at pH 7 was determined to be 1.85 kJ molÿ1 using L tyrosine as a substrate. Phenol was successfully transformed by tyrosinase over wide ranges of pH 5±8 and initial phenol concentration (0.5±10 mM, 47±940 mg/l). Several chlorinated phenols were also successfully transformed. Polyethylene glycol and chitosan did not protect tyrosinase from inactivation during the treatment of phenol; however, chitosan induced the precipitation of reaction products arising from phenol transformation. Ó 2000 Elsevier Science Ltd. All rights reserved. Keywords: Mushroom tyrosinase; Inactivation; Stability; Phenol; Treatment; PEG; Chitosan
1. Introduction Aromatic compounds are present in the wastewaters of a large number of industries including coal conversion, petroleum re®ning, resins and plastics, wood preservation, dyes, chemicals and textiles (Karam and Nicell, 1997). The use of horseradish peroxidase and hydrogen peroxide to accomplish the removal of phenols and aromatic amines from these wastewaters has attracted much attention since the initial work of Klibanov et al. (1980). Treatment of aromatic substrates was accomplished through the oxidation of substrates by hydrogen peroxide under the in¯uence of the biocatalyst. The free radical reaction products spontaneously react to form polymeric products of reduced solubility, which can be separated from solution by coagulation and subsequent sedimentation (Nicell et al., 1993). While peroxidase enzymes have the potential to treat a large variety of phenolic compounds over wide ranges of pH and temperature (Klibanov et al., 1980; Nicell et al., 1993), the major concern regarding this enzymatic treatment is the prohibitive cost of the enzyme and hydrogen peroxide. One possible alternative to peroxidases *
Corresponding author. Tel.: +1-514-398-6675; fax: +1-514-3987361. E-mail address: [email protected] (J.A. Nicell).
is tyrosinase (EC 1.14.18.1) which is widely distributed in bacteria, fruits, vegetables and seafood products (van Gelder et al., 1997). This type of enzyme, also known as polyphenol oxidase, catalyzes the oxidation of phenols but uses molecular oxygen as an oxidant. Tyrosinase catalyzes two distinct oxidation reactions as shown in Fig. 1. In cycle 1, tyrosinase accomplishes the oxidation of monophenols by oxygen as it passes through four enzyme states (Edeoxy , Eoxy , Eoxy-M and Emet-D ). In cycle 2, o-diphenols are oxidized as the enzyme passes through six enzyme states (Edeoxy , Eoxy , Eoxy-D , Emet and Emet-D ). The two cycles lead to the formation of o-quinones which spontaneously react with each other to form oligomers (Dec and Bollag, 1995; Naidja et al., 1998). Atlow et al. (1984) proposed the application of mushroom tyrosinase to the treatment of phenolic wastewaters. While a variety of aqueous phenolic compounds were successfully transformed with tyrosinase (Wada et al., 1995), the ability of the enzyme to accomplish this transformation and to retain its catalytic activity over a range of temperatures and pHs has not been demonstrated. It has been shown that chemical additives such as polyethylene glycol (PEG) and chitosan can prevent inactivation and prolong the catalytic life of peroxidases through interaction with reaction products (Nakamoto and Machida, 1992; Wu et al., 1993; Ganjidoust et al., 1996). However, PEG has not
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Fig. 1. Catalytic cycles for the: (1) hydroxylation of monophenols and (2) dehydrogenation of o-diphenols to o-quinones by tyrosinase. M monophenol and D diphenol bound forms (after Solomon et al., 1996).
yet been investigated for its ability to protect tyrosinases from inactivation. Sun et al. (1992) and Wada et al. (1993) reported that chitosan accelerated the rate of oxidation of several phenols. They concluded that chitosan interacts with reaction products which may otherwise have inhibited or inactivated tyrosinase. While they demonstrated that the rate of reaction was signi®cantly improved in the presence of chitosan, they did not quantify how much this eect resulted in a reduction in the quantity of tyrosinase required to achieve a given degree of transformation of phenolic substrates. Therefore, the objectives of this study were to: (1) characterize tyrosinase with respect to its catalytic activity, its stability and ability to catalyze the transformation of phenolic compounds; and (2) investigate the eect of the chemical additives including PEG and chitosan on the eciency of the catalytic transformation of phenol.
2. Methods Mushroom tyrosinase (EC 1.14.18.1) was obtained from Worthington Biochemicals of Lakewood, NJ and had a nominal activity of 500 units/mg according to the companyÕs speci®cations and based on the assay de-
scribed below. L -tyrosine was purchased from Sigma Chemicals of St. Louis, MO. Phenolic compounds were at least 98% pure and were purchased from either Fluka Chemicals of Ronkonkoma, NY or Aldrich Chemicals of Milwaukee, WI. ACS grade potassium ferricyanide and sodium bicarbonate were purchased from Fisher Scienti®c of Fair Lawn, NJ. Ninety eight percent pure 4-aminoantipyrine (4-AAP) was purchased from Aldrich Chemicals. A chitosan sample with a viscosity of 420 centepoise (cps) was obtained from Vanson of Redmond, WA. Polyethylene glycols (PEG) with average molecular weights of 20 000 and 35 000 were purchased from Fluka Chemicals. Other molecular weights of PEG (2000, 4600, 8000 and 10 000) were purchased from Aldrich Chemicals. ACS grade conjugate acids and bases were purchased from BDH Chemicals of Toronto, ON and were used to prepare buers in accordance with the methods of Gomori (1955). All aqueous solutions were prepared using distilled/deionized water obtained from a D4741 Nanopure Ultrapure Water System (Barnstead/Thermolyne, USA). Aqueous solutions of tyrosinase and 1 mM L -tyrosine were prepared using deionized water and stored at 4°C. Stock solutions of 1% (w/v) chitosan were prepared with 15% acetic acid and stored at 4°C. All other aqueous solutions were stored at room temperature. In colorimetric assays for tyrosinase and phenol the absorbance measurements of samples were performed at 25°C using 1.5 ml quartz and glass cuvettes and a Hewlett±Packard HP845x UV±Visible Spectrophotometer. Reaction pH was measured using an Orion SA520 pH meter with an Orion Ross 8102 multiple electrode from Orion Research. A RTE111 water bath from Neslab was used to maintain the temperature of enzyme solutions for the thermostability experiments conducted between 10°C and 50°C. Precipitates from the enzymatic transformation were removed by centrifugation at 3500 g for 15 min with an IEC Centra-8 centrifuge from International Equipment Company of Needham Height, MA. Tyrosinase activity was determined using L -tyrosine as a substrate in pH 6.5 sodium phosphate buer at 25°C (Worthington Enzyme Manual, 1977). One unit of enzyme results in an increase in absorbance at 280 nm of 0.001 AU per minute in a 3 ml reaction mixture containing 0.181 mg of L -tyrosine in a quartz cuvette with a path length of 1 cm. Phenolic compound concentrations were determined by a colorimetric assay (APHA et al., 1998) based on the absorbance at 510 nm caused by the reaction between the phenolic compound, 4-AAP and potassium ferricyanide. An aliquot of sample was diluted and brought to a volume of 800 ll with 0.25 mM sodium bicarbonate (pH 8.4). 100 ll of 20.8 mM 4-AAP and 100 ll of 83.4 mM potassium ferricyanide were added to the sample and the absorbance of the mixture at 510 nm was
K. Ikehata, J.A. Nicell / Bioresource Technology 74 (2000) 191±199
measured after 6 min. When a sample had a residual absorbance at 510 nm, the residual absorbance was subtracted from the absorbance developed during the colorimetric assay. The absorbance at 510 nm due to phenol was subsequently converted to phenol concentration using a calibration curve. The background color correction was especially signi®cant when a high degree of phenol conversion was achieved. For example, when 99% of 0.5 mM aqueous phenol was transformed, approximately 95% of the absorbance at the end of 6 min assay reaction period was due to treatment products and the remaining 5% was due to unreacted phenol that responded to assay reagents. However, this 5% additional color development due to residual phenol was of sucient magnitude that it could be measured reliably by spectrophotometry. All phenol treatment experiments except those conducted during the pH dependence study were carried out in 0.05 M sodium phosphate buer at pH 7. The reaction solutions were incubated while being stirred at 25°C. After a period of 3 h or more, the reaction solutions were centrifuged and the residual phenol concentrations of supernatants were measured by the colorimetric assay described above. Three hours was sucient to allow the reaction to go to completion. In cases where the phenol concentration was greater than 0.5 mM, the dissolved oxygen in the reaction was insucient to accomplish a complete reaction. Therefore, the reaction vials were left uncapped while being stirred to allow continuous replenishment of oxygen. The volatilization of phenol during the reaction was not signi®cant in the tested range of concentrations and reaction duration. 3. Results and discussion 3.1. Characterization of tyrosinase activity Tyrosinase activity was measured using L -tyrosine as a substrate. The choice of this substrate limits activity measurements to monophenolase activity which governs the monophenolase cycle shown in Fig. 1. As shown in Fig. 2, maximum activity was observed at neutral pH. This optimum pH was dierent from those reported for some tyrosinases obtained from other sources such as pear with an optimum pH of 4.3 using p-hydroxyphenyl propionic acid as a substrate (Espin et al., 1997a) and avocado with an optimum pH of 5 using 4-hydroxyanisole as a substrate (Espin et al., 1997b). Dierences in the reported values can be explained as being due to the nature of the source of enzyme, the substrate used for the activity measurement and the purity of the enzyme. Even though the reported optima range from pH 5 to 7, all results con®rm that tyrosinase is not signi®cantly active under basic conditions.
193
Fig. 2. Relative monophenolase activity of tyrosinase measured in various pH buers at 25°C using L -tyrosine as a substrate.
Based on the plots in Figs. 3 and 4, it is observed that tyrosinase was reasonably stable in neutral buer at room temperature but was quite unstable at pH > 8 or pH < 5 and at elevated temperatures. The stability of tyrosinase in basic buer solutions was higher than in acidic solutions, whereas the pH dependence of tyrosinase activity (see Fig. 2) showed that the enzyme was more active in acidic buers than basic buers. Therefore, tyrosinase is relatively stable under weakly alkaline conditions but is not substantially active. Interestingly, the stability of the enzyme declined sharply at pH 8, then improved somewhat at pH 9 and then declined again at pH 10. The reason for this minor stability optimum at pH 9 is unknown. Tyrosinase activity over time at ®ve dierent temperatures and pH 7 is presented in Fig. 5. It was observed that the thermal inactivation of tyrosinase was ®rst order and could be modeled using A t A0 10ÿt=D A0 eÿkt ;
1
Fig. 3. Stability of tyrosinase incubated at 25°C in various buers.
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Fig. 4. Stability of tyrosinase incubated at 40°C in various buers.
where A(t) is the activity at time t, A0 the activity at time t 0, D the inactivation decimal reduction value and k is the inactivation decay constant. The decimal reduction value is a measure of the time required for the activity to fall to 10% of its original value. As shown in Fig. 6, the relationship of the decimal reduction values to the incubation temperatures was linear when D was plotted on a logarithmic scale. According to DeCordt et al. (1992), decimal reduction values may be related to incubation temperature using
Fig. 6. Dependence of thermal inactivation decimal reduction value on temperature for tyrosinase in pH 7 sodium phosphate buer.
calculated from the data of Fig. 5. The activation energy, Ea , can be calculated using the Arrhenius equation k AeÿEa =RT ;
3
where DREF is the decimal reduction value at a temperature TREF and Z is the temperature change required to obtain a 10-fold increase or decrease in the decimal reduction value. The Z-value was calculated to be 10.4°C through a linear regression of data in Fig. 6. Fig. 7 is an Arrhenius plot showing the temperature dependence of thermal inactivation decay constants, k,
where A is a frequency factor which is inherent in the reaction, R the universal gas constant, and T is the absolute temperature. Based on a linear regression analysis of the data in Fig. 7, Ea was 1.85 kJ molÿ1 and A was 1:73 1019 minÿ1 . The temperature dependence of tyrosinase activity has been reported previously for several tyrosinases from other plants. The value obtained for the activation energy in this study was comparable to the value for palmito (Acanthophoenix rubra) polyphenol oxidase using 4-methylcatechol as a substrate (5.41 kJ molÿ1 ) (Robert et al., 1995); but much lower than the other values quoted in the same literature for potato polyphenol oxidase using pyrogallol as a substrate (54.5 kJ
Fig. 5. Thermal inactivation of tyrosinase in pH 7 sodium phosphate buer.
Fig. 7. Dependence of thermal inactivation decay constant on temperature at pH 7 for tyrosinase.
D DREF 10 TREF ÿT =Z ;
2
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molÿ1 ) and for banana polyphenol oxidase using catechol as a substrate (18.6 kJ molÿ1 ). The lower activation energy for inactivation implies a lower thermal stability of the enzyme used in this study. A comparison of the thermal inactivation parameters of tyrosinase and those of soybean peroxidase that was investigated earlier in our laboratory for the purpose of wastewater treatment (Wright and Nicell, 1999) is shown in Table 1. The decimal reduction value of tyrosinase at 50°C was approximately 104 times lower than that of soybean peroxidase. The activation energy also indicates a much lower thermal stability of tyrosinase than soybean peroxidase. However, the Z-value, which represents the susceptibility of the rate of enzyme inactivation to temperature change, of tyrosinase was quite similar to that of soybean peroxidase. The lower thermal stability of tyrosinase represents a drawback to its application to wastewater treatment, especially during the preparation and storage of stock solutions. 3.2. Tyrosinase catalyzed transformation of phenols After the addition of tyrosinase to oxygenated phenol solutions, mixtures became colored after a lag period and then gradually darkened over time. Precipitates did not form in any of the solutions that had initial phenol concentrations of 0.5±10 mM (47±940 mg/l) even when full transformation of phenol was achieved. This is in agreement with the observations of Wada et al. (1993) but not those of Atlow et al. (1984) who reported the gradual formation of a black precipitate. Wada et al. suggested that the lower purity enzyme used by Atlow et al. might have induced the precipitation of reaction products. However, the speci®c activity of tyrosinase used in this study (500 units/mg) implied that the enzyme stock was of lower purity than those used by Atlow et al. (2000 units/mg) and Wada et al. (3500 units/ mg), where all activities were measured using the same assay. Also, the concentrations and types of buers, which might have an in¯uence on precipitation, were the same in all of these studies (i.e., 50 mM sodium phosphate buer). It is also possible that dierences in the
195
oxygenation rates of reacting solutions could alter the nature of reaction products and induce their precipitation; however, there appears to be no dierence in oxygenation techniques between these three studies. Thus, there is currently no de®nitive explanation for the con¯icting observations. Irregardless, since the colored products are contaminants that may exert toxicity or oxygen demand on receiving water bodies, the removal of soluble and/or suspended reaction products from the euent must be studied further. The eects of pH and tyrosinase dose on phenol transformation were investigated. Parallel experiments were conducted in the presence and absence of chitosan to assess the eect of this additive on the transformation eciency of phenol. Fig. 8 presents the transformation of 0.5 mM phenol and the intensity of generated color (measured by absorbance at 510 nm) as a function of enzyme dose and pH, without the presence of chitosan. The incubation time was 3 h which was sucient to ensure that reactions had gone to completion. When limiting doses of enzyme were used (i.e., all enzyme is inactivated at the end of the reaction before 100% transformation of phenol is achieved), the pro®le of phenol transformation shown in Fig. 8(a) was similar to the relative activity of tyrosinase at dierent pHs shown in Fig. 2. Tyrosinase catalyzed the transformation of phenol very eectively at pHs ranging from 5 to 8 with an optimum at pH 7. This optimum is de®ned as the pH where a particular dose of tyrosinase achieves maximum transformation of phenol. Only minor transformations occurred at pH 4 and 9, and no phenol transformation was observed at lower and higher extremities of pH. Fig. 8(b) shows that the quantity of color generated at 510 nm follows the same trends as phenol transformation. As shown in Fig. 9, the intensity of color generated at 510 nm was independent of pH and was directly related to the quantity of phenol transformed. The dependence of phenol transformation on pH was also investigated using reactions conducted in the presence of 100 mg/l of chitosan. As shown in Figs. 8(a) and 10(a), the shapes of the curves for phenol transformation were slightly dierent depending on whether or not
Table 1 Comparison of thermal inactivation parameters between soybean peroxidase and mushroom tyrosinase (pH 7).
a
Enzyme
Temperature (°C)
D (min)
Ea (kJ molÿ1 )
Z (°C)
Mushroom Tyrosinasea
10 30 50
25800 2100 38.4
1.85
10.4
Soybean Peroxidaseb
50 70 80 91
4 105c 3974 300 30
2.46
This study. Wright and Nicell (1999). c Extrapolated from Wright and Nicell (1999). b
9.71
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Fig. 8. Eect of pH on the transformation of phenol catalyzed by tyrosinase in the absence of chitosan: (a) phenol transformation, (b) color generated at 510 nm (phenol0 0:5 mM, 3 h of reaction at 25°C).
Fig. 9. Relationship between the color generated at 510 nm and transformed phenol.
chitosan was present. Phenol transformation improved at pH 4 in the presence of chitosan. Dark-brown precipitates were observed at pHs between 5 and 8 and
Fig. 10. Eect of pH on the transformation of phenol catalyzed by tyrosinase in the presence of 420 cps chitosan: (a) phenol transformation, (b) color remaining at 510 nm after centrifugation (phenol0 0:5 mM, 3 h of reaction at 25°C).
residual color was depressed (see Fig. 10(b)) compared to treatments conducted without chitosan (see Fig. 8(b)). Since color generation is independent of pH (see Fig. 9), this depression of color is likely due to the ability of chitosan to act as a coagulant (Wada et al., 1995) which destabilizes the suspended polymer products. Color depression was eectively achieved between pH 5 and 7. No precipitation of colored products occurred at pH 4 where chitosan is highly soluble and cannot act as a coagulant. Chitosan was not eective as a coagulant above pH 7, presumably due to its lower solubility under basic conditions when the amino groups of chitosan molecules become deprotonated. This would prevent chitosan from eectively interacting with suspended reaction products, thereby destabilizing them and accomplishing their precipitation. Aqueous phenol solutions with concentrations ranging from 0.5 to 4.0 mM were treated with a range of doses of tyrosinase at pH 7 and 25°C with and without chitosan. The quantities of chitosan used were 100, 200, 400 and 800 mg/l for 0.5, 1.0, 2.0 and 4.0 mM phenol, respectively. These quantities of chitosan were sucient
K. Ikehata, J.A. Nicell / Bioresource Technology 74 (2000) 191±199
to maximally suppress color generation in the reaction solutions. An example of the relationship between phenol transformation and enzyme dose is presented in Fig. 11 for 1 mM phenol. Although the color residual was depressed (data not shown) and precipitates formed in the reaction solutions with chitosan, no signi®cant eect on the extent of phenol transformation was observed. The same observations were made for the full range of phenol concentrations studied. From graphs such as Fig. 11, the tyrosinase doses required to transform 95% of initial phenol concentrations ranging from 0.5 to 4 mM were interpolated and plotted in Fig. 12. A threshold of 95% was chosen because it is a representative of a high degree of removal of the aromatic substrate which would be desired in a treatment system. A linear relationship was shown to exist between initial phenol concentration and tyrosinase dose required to achieve 95% transformation. The slopes of the curves were 6.79 units/ml mM without chitosan and 6.40 units/ ml mM with chitosan. Although the addition of chitosan was very eective in inducing the precipitation of reaction products, the small dierence between these values indicates that chitosan did not signi®cantly reduce the quantity of enzyme required to treat the contaminant and, thus, was not able to protect tyrosinase from inactivation. This conclusion appears to contradict the results of Sun et al. (1992) and Wada et al. (1993) who demonstrated that the rate of reaction of several substrates including phenol were accelerated in the presence of chitosan. They concluded that the interaction of chitosan with reaction products resulted in the protection of tyrosinase from inhibition or inactivation. However, the current study indicates that the extent of transformation of phenol was not improved in the presence of chitosan. These studies taken together, indicate that even though chitosan can improve the rate of reaction, there appears
Fig. 11. Tyrosinase catalyzed transformation of phenol with and without 420 cps chitosan (phenol0 1 mM, 3 h of reaction in pH 7 sodium phosphate buer at 25°C).
197
Fig. 12. Amount of tyrosinase required to transform 95% of initial phenol (3 h of reaction in pH 7 sodium phosphate buer at 25°C).
to be a ®xed stoichiometry between tyrosinase and phenol. Thus, it appears that chitosan may protect the enzyme from inhibition (which limits the rate of reaction) during the treatment of phenol but not from inactivation (which limits the extent of transformation). This may be explained as being due to the in¯uence of a suicide inactivation mechanism that is independent of the mechanism of tyrosinase inhibition through interaction with aqueous reaction products. Further research is required to con®rm or dispel this hypothesis. An activity of 6.0 units/ml of tyrosinase was found to be sucient to accomplish almost 100% transformation of 0.5 mM phenol. This amount is 10-fold lower than that which was reported by Atlow et al. (1984) and threefold lower than that which was reported by Wada et al. (1993). The enzyme assay and reaction conditions used in this study, including pH, buer type and concentration, and temperature were the same as those used by both groups of researchers. However, the lower enzyme requirement reported here may be due to the signi®cant protein content of the low purity enzyme used in this study. That is, some amino acid residues on proteins can act as reducers to promote the monophenolase activity (see cycle I of Fig. 1) of tyrosinase (Cooksey et al., 1997). The application of the lower purity enzyme to wastewater treatment may be advantageous due to its improved performance and lower cost. Nakamoto and Machida (1992) had reported that the ability of PEG to protect peroxidase enzymes during the treatment of phenol was molecular weight dependent. Therefore, six types of PEG with average molecular weights ranging from 2000 to 35 000 were examined for their ability to improve the transformation of phenol. Although PEGs were used in concentrations as high as 400 mg/l, no signi®cant improvement in phenol transformation was observed (data not shown). In addition, no precipitates were observed in reaction solutions with
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and without PEG. The failure of PEG to protect tyrosinase, in contrast to its strong ability to protect peroxidase, may be due to the dierences in nature of the oxidized products of these two enzymes. Nakamoto and Machida (1992) proposed that the hydrogen bonding sites of PEG could interact with the hydroxyl groups of polymerized phenols. This interaction may minimize the entrapment and subsequent inactivation of peroxidase by precipitating polymers (Buchanan and Nicell, 1998). In contrast, the o-quinones formed during the tyrosinase-catalyzed oxidation of phenols react to form polymers of low molecular mass (Naidja et al., 1998). These polymers are substantially soluble and, thus, PEG may not have the opportunity to protect tyrosinase from precipitating products. The transformation of ®ve phenolic compounds (0.5 mM) with tyrosinase at pH 7 without chitosan is shown in Fig. 13. Phenol and 4-chlorophenol were fully transformed in less than 3 h using 8 and 48 units/ml, respectively. 3-Chlorophenol and 2-chlorophenol were also fully transformed using 48 units/ml but they required signi®cantly longer reaction times. 2,4-Dichlorophenol could not be fully transformed by 48 units/ml of tyrosinase and pentachlorophenol could not be transformed at all (data not shown). These observations con®rm that all phenols are not equally good substrates of tyrosinase and some cause more rapid inactivation of the enzyme than others. Thus, the quantity of tyrosinase required to achieve a given degree of transformation will vary substantially between substrates. In all cases, colored products were formed during the transformation of these phenols but no precipitates were observed. The impact of these colored products may represent a severe limitation to the application of tyrosinase for treating phenols in industrial wastewaters.
4. Conclusions Tyrosinase activity has been characterized using L tyrosine as a substrate. Maximum catalytic activity was observed at pH 7 with approximately 50% of the maximum was observed at pHs ranging from 5 to 8. Tyrosinase appeared to be unstable at low pH and at elevated temperature. The activation energy for thermal inactivation of tyrosinase at neutral pH was calculated to be 1.85 kJ molÿ1 . Tyrosinase was quite unstable in comparison to peroxidase enzymes which have also been examined for their potential application to the treatment of wastewaters. The optimum pH for phenol treatment was between pH 5 and 8. Tyrosinase was able to transform phenol over a wide range of initial phenol concentration (0.5±10 mM), but no precipitates were observed. Monochlorinated phenols were also successfully transformed with tyrosinase, but the quantity of the enzyme required to achieve a given amount of transformation was substrate dependent. While chitosan addition induced the precipitation of products and depressed color generation, it had no eect on the extent of phenol transformation within the tested range of phenol concentrations. PEG addition did not have any eect on the transformation of phenol. Even though it has been shown that tyrosinase can be used successfully to transform a variety of aqueous phenols, the generation of soluble reaction products is of signi®cant concern since they will impact upon the quality of the treated euent. In order to establish whether the use of tyrosinase is feasible for the treatment of phenolic wastewaters, the toxicity and treatability of these reaction products must be investigated further.
Acknowledgements This work was funded by the Natural Sciences and Engineering Research Council of Canada. References
Fig. 13. Tyrosinase catalyzed treatment of aqueous phenolic compounds as a function of time (phenols0 0:5 mM, tyrosinase dose 8 units/ml for phenol, 48 units/ml for chlorinated phenols, in pH 7 sodium phosphate buer at 25°C).
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Characterization of tyrosinase for the treatment of aqueous phenols Keisuke Ikehata, James A. Nicell * Department of Civil Engineering and Applied Mechanics, McGill University, 817 Sherbrooke Street West, Montreal, Que., Canada H3A 2K6 Received 7 August 1999; received in revised form 24 January 2000; accepted 30 January 2000
Abstract Mushroom tyrosinase (polyphenol oxidase, EC 1.14.18.1) was investigated as an alternative to peroxidase enzymes for the catalytic removal of phenolic compounds from wastewaters. Maximum catalytic activity was observed at pH 7 and more than 50% of optimum activity was observed at pHs ranging between 5 and 8. Tyrosinase was unstable under acidic conditions and at elevated temperatures. The activation energy for thermal inactivation of tyrosinase at pH 7 was determined to be 1.85 kJ molÿ1 using L tyrosine as a substrate. Phenol was successfully transformed by tyrosinase over wide ranges of pH 5±8 and initial phenol concentration (0.5±10 mM, 47±940 mg/l). Several chlorinated phenols were also successfully transformed. Polyethylene glycol and chitosan did not protect tyrosinase from inactivation during the treatment of phenol; however, chitosan induced the precipitation of reaction products arising from phenol transformation. Ó 2000 Elsevier Science Ltd. All rights reserved. Keywords: Mushroom tyrosinase; Inactivation; Stability; Phenol; Treatment; PEG; Chitosan
1. Introduction Aromatic compounds are present in the wastewaters of a large number of industries including coal conversion, petroleum re®ning, resins and plastics, wood preservation, dyes, chemicals and textiles (Karam and Nicell, 1997). The use of horseradish peroxidase and hydrogen peroxide to accomplish the removal of phenols and aromatic amines from these wastewaters has attracted much attention since the initial work of Klibanov et al. (1980). Treatment of aromatic substrates was accomplished through the oxidation of substrates by hydrogen peroxide under the in¯uence of the biocatalyst. The free radical reaction products spontaneously react to form polymeric products of reduced solubility, which can be separated from solution by coagulation and subsequent sedimentation (Nicell et al., 1993). While peroxidase enzymes have the potential to treat a large variety of phenolic compounds over wide ranges of pH and temperature (Klibanov et al., 1980; Nicell et al., 1993), the major concern regarding this enzymatic treatment is the prohibitive cost of the enzyme and hydrogen peroxide. One possible alternative to peroxidases *
Corresponding author. Tel.: +1-514-398-6675; fax: +1-514-3987361. E-mail address: [email protected] (J.A. Nicell).
is tyrosinase (EC 1.14.18.1) which is widely distributed in bacteria, fruits, vegetables and seafood products (van Gelder et al., 1997). This type of enzyme, also known as polyphenol oxidase, catalyzes the oxidation of phenols but uses molecular oxygen as an oxidant. Tyrosinase catalyzes two distinct oxidation reactions as shown in Fig. 1. In cycle 1, tyrosinase accomplishes the oxidation of monophenols by oxygen as it passes through four enzyme states (Edeoxy , Eoxy , Eoxy-M and Emet-D ). In cycle 2, o-diphenols are oxidized as the enzyme passes through six enzyme states (Edeoxy , Eoxy , Eoxy-D , Emet and Emet-D ). The two cycles lead to the formation of o-quinones which spontaneously react with each other to form oligomers (Dec and Bollag, 1995; Naidja et al., 1998). Atlow et al. (1984) proposed the application of mushroom tyrosinase to the treatment of phenolic wastewaters. While a variety of aqueous phenolic compounds were successfully transformed with tyrosinase (Wada et al., 1995), the ability of the enzyme to accomplish this transformation and to retain its catalytic activity over a range of temperatures and pHs has not been demonstrated. It has been shown that chemical additives such as polyethylene glycol (PEG) and chitosan can prevent inactivation and prolong the catalytic life of peroxidases through interaction with reaction products (Nakamoto and Machida, 1992; Wu et al., 1993; Ganjidoust et al., 1996). However, PEG has not
0960-8524/00/$ - see front matter Ó 2000 Elsevier Science Ltd. All rights reserved. PII: S 0 9 6 0 - 8 5 2 4 ( 0 0 ) 0 0 0 2 5 - 0
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Fig. 1. Catalytic cycles for the: (1) hydroxylation of monophenols and (2) dehydrogenation of o-diphenols to o-quinones by tyrosinase. M monophenol and D diphenol bound forms (after Solomon et al., 1996).
yet been investigated for its ability to protect tyrosinases from inactivation. Sun et al. (1992) and Wada et al. (1993) reported that chitosan accelerated the rate of oxidation of several phenols. They concluded that chitosan interacts with reaction products which may otherwise have inhibited or inactivated tyrosinase. While they demonstrated that the rate of reaction was signi®cantly improved in the presence of chitosan, they did not quantify how much this eect resulted in a reduction in the quantity of tyrosinase required to achieve a given degree of transformation of phenolic substrates. Therefore, the objectives of this study were to: (1) characterize tyrosinase with respect to its catalytic activity, its stability and ability to catalyze the transformation of phenolic compounds; and (2) investigate the eect of the chemical additives including PEG and chitosan on the eciency of the catalytic transformation of phenol.
2. Methods Mushroom tyrosinase (EC 1.14.18.1) was obtained from Worthington Biochemicals of Lakewood, NJ and had a nominal activity of 500 units/mg according to the companyÕs speci®cations and based on the assay de-
scribed below. L -tyrosine was purchased from Sigma Chemicals of St. Louis, MO. Phenolic compounds were at least 98% pure and were purchased from either Fluka Chemicals of Ronkonkoma, NY or Aldrich Chemicals of Milwaukee, WI. ACS grade potassium ferricyanide and sodium bicarbonate were purchased from Fisher Scienti®c of Fair Lawn, NJ. Ninety eight percent pure 4-aminoantipyrine (4-AAP) was purchased from Aldrich Chemicals. A chitosan sample with a viscosity of 420 centepoise (cps) was obtained from Vanson of Redmond, WA. Polyethylene glycols (PEG) with average molecular weights of 20 000 and 35 000 were purchased from Fluka Chemicals. Other molecular weights of PEG (2000, 4600, 8000 and 10 000) were purchased from Aldrich Chemicals. ACS grade conjugate acids and bases were purchased from BDH Chemicals of Toronto, ON and were used to prepare buers in accordance with the methods of Gomori (1955). All aqueous solutions were prepared using distilled/deionized water obtained from a D4741 Nanopure Ultrapure Water System (Barnstead/Thermolyne, USA). Aqueous solutions of tyrosinase and 1 mM L -tyrosine were prepared using deionized water and stored at 4°C. Stock solutions of 1% (w/v) chitosan were prepared with 15% acetic acid and stored at 4°C. All other aqueous solutions were stored at room temperature. In colorimetric assays for tyrosinase and phenol the absorbance measurements of samples were performed at 25°C using 1.5 ml quartz and glass cuvettes and a Hewlett±Packard HP845x UV±Visible Spectrophotometer. Reaction pH was measured using an Orion SA520 pH meter with an Orion Ross 8102 multiple electrode from Orion Research. A RTE111 water bath from Neslab was used to maintain the temperature of enzyme solutions for the thermostability experiments conducted between 10°C and 50°C. Precipitates from the enzymatic transformation were removed by centrifugation at 3500 g for 15 min with an IEC Centra-8 centrifuge from International Equipment Company of Needham Height, MA. Tyrosinase activity was determined using L -tyrosine as a substrate in pH 6.5 sodium phosphate buer at 25°C (Worthington Enzyme Manual, 1977). One unit of enzyme results in an increase in absorbance at 280 nm of 0.001 AU per minute in a 3 ml reaction mixture containing 0.181 mg of L -tyrosine in a quartz cuvette with a path length of 1 cm. Phenolic compound concentrations were determined by a colorimetric assay (APHA et al., 1998) based on the absorbance at 510 nm caused by the reaction between the phenolic compound, 4-AAP and potassium ferricyanide. An aliquot of sample was diluted and brought to a volume of 800 ll with 0.25 mM sodium bicarbonate (pH 8.4). 100 ll of 20.8 mM 4-AAP and 100 ll of 83.4 mM potassium ferricyanide were added to the sample and the absorbance of the mixture at 510 nm was
K. Ikehata, J.A. Nicell / Bioresource Technology 74 (2000) 191±199
measured after 6 min. When a sample had a residual absorbance at 510 nm, the residual absorbance was subtracted from the absorbance developed during the colorimetric assay. The absorbance at 510 nm due to phenol was subsequently converted to phenol concentration using a calibration curve. The background color correction was especially signi®cant when a high degree of phenol conversion was achieved. For example, when 99% of 0.5 mM aqueous phenol was transformed, approximately 95% of the absorbance at the end of 6 min assay reaction period was due to treatment products and the remaining 5% was due to unreacted phenol that responded to assay reagents. However, this 5% additional color development due to residual phenol was of sucient magnitude that it could be measured reliably by spectrophotometry. All phenol treatment experiments except those conducted during the pH dependence study were carried out in 0.05 M sodium phosphate buer at pH 7. The reaction solutions were incubated while being stirred at 25°C. After a period of 3 h or more, the reaction solutions were centrifuged and the residual phenol concentrations of supernatants were measured by the colorimetric assay described above. Three hours was sucient to allow the reaction to go to completion. In cases where the phenol concentration was greater than 0.5 mM, the dissolved oxygen in the reaction was insucient to accomplish a complete reaction. Therefore, the reaction vials were left uncapped while being stirred to allow continuous replenishment of oxygen. The volatilization of phenol during the reaction was not signi®cant in the tested range of concentrations and reaction duration. 3. Results and discussion 3.1. Characterization of tyrosinase activity Tyrosinase activity was measured using L -tyrosine as a substrate. The choice of this substrate limits activity measurements to monophenolase activity which governs the monophenolase cycle shown in Fig. 1. As shown in Fig. 2, maximum activity was observed at neutral pH. This optimum pH was dierent from those reported for some tyrosinases obtained from other sources such as pear with an optimum pH of 4.3 using p-hydroxyphenyl propionic acid as a substrate (Espin et al., 1997a) and avocado with an optimum pH of 5 using 4-hydroxyanisole as a substrate (Espin et al., 1997b). Dierences in the reported values can be explained as being due to the nature of the source of enzyme, the substrate used for the activity measurement and the purity of the enzyme. Even though the reported optima range from pH 5 to 7, all results con®rm that tyrosinase is not signi®cantly active under basic conditions.
193
Fig. 2. Relative monophenolase activity of tyrosinase measured in various pH buers at 25°C using L -tyrosine as a substrate.
Based on the plots in Figs. 3 and 4, it is observed that tyrosinase was reasonably stable in neutral buer at room temperature but was quite unstable at pH > 8 or pH < 5 and at elevated temperatures. The stability of tyrosinase in basic buer solutions was higher than in acidic solutions, whereas the pH dependence of tyrosinase activity (see Fig. 2) showed that the enzyme was more active in acidic buers than basic buers. Therefore, tyrosinase is relatively stable under weakly alkaline conditions but is not substantially active. Interestingly, the stability of the enzyme declined sharply at pH 8, then improved somewhat at pH 9 and then declined again at pH 10. The reason for this minor stability optimum at pH 9 is unknown. Tyrosinase activity over time at ®ve dierent temperatures and pH 7 is presented in Fig. 5. It was observed that the thermal inactivation of tyrosinase was ®rst order and could be modeled using A t A0 10ÿt=D A0 eÿkt ;
1
Fig. 3. Stability of tyrosinase incubated at 25°C in various buers.
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Fig. 4. Stability of tyrosinase incubated at 40°C in various buers.
where A(t) is the activity at time t, A0 the activity at time t 0, D the inactivation decimal reduction value and k is the inactivation decay constant. The decimal reduction value is a measure of the time required for the activity to fall to 10% of its original value. As shown in Fig. 6, the relationship of the decimal reduction values to the incubation temperatures was linear when D was plotted on a logarithmic scale. According to DeCordt et al. (1992), decimal reduction values may be related to incubation temperature using
Fig. 6. Dependence of thermal inactivation decimal reduction value on temperature for tyrosinase in pH 7 sodium phosphate buer.
calculated from the data of Fig. 5. The activation energy, Ea , can be calculated using the Arrhenius equation k AeÿEa =RT ;
3
where DREF is the decimal reduction value at a temperature TREF and Z is the temperature change required to obtain a 10-fold increase or decrease in the decimal reduction value. The Z-value was calculated to be 10.4°C through a linear regression of data in Fig. 6. Fig. 7 is an Arrhenius plot showing the temperature dependence of thermal inactivation decay constants, k,
where A is a frequency factor which is inherent in the reaction, R the universal gas constant, and T is the absolute temperature. Based on a linear regression analysis of the data in Fig. 7, Ea was 1.85 kJ molÿ1 and A was 1:73 1019 minÿ1 . The temperature dependence of tyrosinase activity has been reported previously for several tyrosinases from other plants. The value obtained for the activation energy in this study was comparable to the value for palmito (Acanthophoenix rubra) polyphenol oxidase using 4-methylcatechol as a substrate (5.41 kJ molÿ1 ) (Robert et al., 1995); but much lower than the other values quoted in the same literature for potato polyphenol oxidase using pyrogallol as a substrate (54.5 kJ
Fig. 5. Thermal inactivation of tyrosinase in pH 7 sodium phosphate buer.
Fig. 7. Dependence of thermal inactivation decay constant on temperature at pH 7 for tyrosinase.
D DREF 10 TREF ÿT =Z ;
2
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molÿ1 ) and for banana polyphenol oxidase using catechol as a substrate (18.6 kJ molÿ1 ). The lower activation energy for inactivation implies a lower thermal stability of the enzyme used in this study. A comparison of the thermal inactivation parameters of tyrosinase and those of soybean peroxidase that was investigated earlier in our laboratory for the purpose of wastewater treatment (Wright and Nicell, 1999) is shown in Table 1. The decimal reduction value of tyrosinase at 50°C was approximately 104 times lower than that of soybean peroxidase. The activation energy also indicates a much lower thermal stability of tyrosinase than soybean peroxidase. However, the Z-value, which represents the susceptibility of the rate of enzyme inactivation to temperature change, of tyrosinase was quite similar to that of soybean peroxidase. The lower thermal stability of tyrosinase represents a drawback to its application to wastewater treatment, especially during the preparation and storage of stock solutions. 3.2. Tyrosinase catalyzed transformation of phenols After the addition of tyrosinase to oxygenated phenol solutions, mixtures became colored after a lag period and then gradually darkened over time. Precipitates did not form in any of the solutions that had initial phenol concentrations of 0.5±10 mM (47±940 mg/l) even when full transformation of phenol was achieved. This is in agreement with the observations of Wada et al. (1993) but not those of Atlow et al. (1984) who reported the gradual formation of a black precipitate. Wada et al. suggested that the lower purity enzyme used by Atlow et al. might have induced the precipitation of reaction products. However, the speci®c activity of tyrosinase used in this study (500 units/mg) implied that the enzyme stock was of lower purity than those used by Atlow et al. (2000 units/mg) and Wada et al. (3500 units/ mg), where all activities were measured using the same assay. Also, the concentrations and types of buers, which might have an in¯uence on precipitation, were the same in all of these studies (i.e., 50 mM sodium phosphate buer). It is also possible that dierences in the
195
oxygenation rates of reacting solutions could alter the nature of reaction products and induce their precipitation; however, there appears to be no dierence in oxygenation techniques between these three studies. Thus, there is currently no de®nitive explanation for the con¯icting observations. Irregardless, since the colored products are contaminants that may exert toxicity or oxygen demand on receiving water bodies, the removal of soluble and/or suspended reaction products from the euent must be studied further. The eects of pH and tyrosinase dose on phenol transformation were investigated. Parallel experiments were conducted in the presence and absence of chitosan to assess the eect of this additive on the transformation eciency of phenol. Fig. 8 presents the transformation of 0.5 mM phenol and the intensity of generated color (measured by absorbance at 510 nm) as a function of enzyme dose and pH, without the presence of chitosan. The incubation time was 3 h which was sucient to ensure that reactions had gone to completion. When limiting doses of enzyme were used (i.e., all enzyme is inactivated at the end of the reaction before 100% transformation of phenol is achieved), the pro®le of phenol transformation shown in Fig. 8(a) was similar to the relative activity of tyrosinase at dierent pHs shown in Fig. 2. Tyrosinase catalyzed the transformation of phenol very eectively at pHs ranging from 5 to 8 with an optimum at pH 7. This optimum is de®ned as the pH where a particular dose of tyrosinase achieves maximum transformation of phenol. Only minor transformations occurred at pH 4 and 9, and no phenol transformation was observed at lower and higher extremities of pH. Fig. 8(b) shows that the quantity of color generated at 510 nm follows the same trends as phenol transformation. As shown in Fig. 9, the intensity of color generated at 510 nm was independent of pH and was directly related to the quantity of phenol transformed. The dependence of phenol transformation on pH was also investigated using reactions conducted in the presence of 100 mg/l of chitosan. As shown in Figs. 8(a) and 10(a), the shapes of the curves for phenol transformation were slightly dierent depending on whether or not
Table 1 Comparison of thermal inactivation parameters between soybean peroxidase and mushroom tyrosinase (pH 7).
a
Enzyme
Temperature (°C)
D (min)
Ea (kJ molÿ1 )
Z (°C)
Mushroom Tyrosinasea
10 30 50
25800 2100 38.4
1.85
10.4
Soybean Peroxidaseb
50 70 80 91
4 105c 3974 300 30
2.46
This study. Wright and Nicell (1999). c Extrapolated from Wright and Nicell (1999). b
9.71
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Fig. 8. Eect of pH on the transformation of phenol catalyzed by tyrosinase in the absence of chitosan: (a) phenol transformation, (b) color generated at 510 nm (phenol0 0:5 mM, 3 h of reaction at 25°C).
Fig. 9. Relationship between the color generated at 510 nm and transformed phenol.
chitosan was present. Phenol transformation improved at pH 4 in the presence of chitosan. Dark-brown precipitates were observed at pHs between 5 and 8 and
Fig. 10. Eect of pH on the transformation of phenol catalyzed by tyrosinase in the presence of 420 cps chitosan: (a) phenol transformation, (b) color remaining at 510 nm after centrifugation (phenol0 0:5 mM, 3 h of reaction at 25°C).
residual color was depressed (see Fig. 10(b)) compared to treatments conducted without chitosan (see Fig. 8(b)). Since color generation is independent of pH (see Fig. 9), this depression of color is likely due to the ability of chitosan to act as a coagulant (Wada et al., 1995) which destabilizes the suspended polymer products. Color depression was eectively achieved between pH 5 and 7. No precipitation of colored products occurred at pH 4 where chitosan is highly soluble and cannot act as a coagulant. Chitosan was not eective as a coagulant above pH 7, presumably due to its lower solubility under basic conditions when the amino groups of chitosan molecules become deprotonated. This would prevent chitosan from eectively interacting with suspended reaction products, thereby destabilizing them and accomplishing their precipitation. Aqueous phenol solutions with concentrations ranging from 0.5 to 4.0 mM were treated with a range of doses of tyrosinase at pH 7 and 25°C with and without chitosan. The quantities of chitosan used were 100, 200, 400 and 800 mg/l for 0.5, 1.0, 2.0 and 4.0 mM phenol, respectively. These quantities of chitosan were sucient
K. Ikehata, J.A. Nicell / Bioresource Technology 74 (2000) 191±199
to maximally suppress color generation in the reaction solutions. An example of the relationship between phenol transformation and enzyme dose is presented in Fig. 11 for 1 mM phenol. Although the color residual was depressed (data not shown) and precipitates formed in the reaction solutions with chitosan, no signi®cant eect on the extent of phenol transformation was observed. The same observations were made for the full range of phenol concentrations studied. From graphs such as Fig. 11, the tyrosinase doses required to transform 95% of initial phenol concentrations ranging from 0.5 to 4 mM were interpolated and plotted in Fig. 12. A threshold of 95% was chosen because it is a representative of a high degree of removal of the aromatic substrate which would be desired in a treatment system. A linear relationship was shown to exist between initial phenol concentration and tyrosinase dose required to achieve 95% transformation. The slopes of the curves were 6.79 units/ml mM without chitosan and 6.40 units/ ml mM with chitosan. Although the addition of chitosan was very eective in inducing the precipitation of reaction products, the small dierence between these values indicates that chitosan did not signi®cantly reduce the quantity of enzyme required to treat the contaminant and, thus, was not able to protect tyrosinase from inactivation. This conclusion appears to contradict the results of Sun et al. (1992) and Wada et al. (1993) who demonstrated that the rate of reaction of several substrates including phenol were accelerated in the presence of chitosan. They concluded that the interaction of chitosan with reaction products resulted in the protection of tyrosinase from inhibition or inactivation. However, the current study indicates that the extent of transformation of phenol was not improved in the presence of chitosan. These studies taken together, indicate that even though chitosan can improve the rate of reaction, there appears
Fig. 11. Tyrosinase catalyzed transformation of phenol with and without 420 cps chitosan (phenol0 1 mM, 3 h of reaction in pH 7 sodium phosphate buer at 25°C).
197
Fig. 12. Amount of tyrosinase required to transform 95% of initial phenol (3 h of reaction in pH 7 sodium phosphate buer at 25°C).
to be a ®xed stoichiometry between tyrosinase and phenol. Thus, it appears that chitosan may protect the enzyme from inhibition (which limits the rate of reaction) during the treatment of phenol but not from inactivation (which limits the extent of transformation). This may be explained as being due to the in¯uence of a suicide inactivation mechanism that is independent of the mechanism of tyrosinase inhibition through interaction with aqueous reaction products. Further research is required to con®rm or dispel this hypothesis. An activity of 6.0 units/ml of tyrosinase was found to be sucient to accomplish almost 100% transformation of 0.5 mM phenol. This amount is 10-fold lower than that which was reported by Atlow et al. (1984) and threefold lower than that which was reported by Wada et al. (1993). The enzyme assay and reaction conditions used in this study, including pH, buer type and concentration, and temperature were the same as those used by both groups of researchers. However, the lower enzyme requirement reported here may be due to the signi®cant protein content of the low purity enzyme used in this study. That is, some amino acid residues on proteins can act as reducers to promote the monophenolase activity (see cycle I of Fig. 1) of tyrosinase (Cooksey et al., 1997). The application of the lower purity enzyme to wastewater treatment may be advantageous due to its improved performance and lower cost. Nakamoto and Machida (1992) had reported that the ability of PEG to protect peroxidase enzymes during the treatment of phenol was molecular weight dependent. Therefore, six types of PEG with average molecular weights ranging from 2000 to 35 000 were examined for their ability to improve the transformation of phenol. Although PEGs were used in concentrations as high as 400 mg/l, no signi®cant improvement in phenol transformation was observed (data not shown). In addition, no precipitates were observed in reaction solutions with
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and without PEG. The failure of PEG to protect tyrosinase, in contrast to its strong ability to protect peroxidase, may be due to the dierences in nature of the oxidized products of these two enzymes. Nakamoto and Machida (1992) proposed that the hydrogen bonding sites of PEG could interact with the hydroxyl groups of polymerized phenols. This interaction may minimize the entrapment and subsequent inactivation of peroxidase by precipitating polymers (Buchanan and Nicell, 1998). In contrast, the o-quinones formed during the tyrosinase-catalyzed oxidation of phenols react to form polymers of low molecular mass (Naidja et al., 1998). These polymers are substantially soluble and, thus, PEG may not have the opportunity to protect tyrosinase from precipitating products. The transformation of ®ve phenolic compounds (0.5 mM) with tyrosinase at pH 7 without chitosan is shown in Fig. 13. Phenol and 4-chlorophenol were fully transformed in less than 3 h using 8 and 48 units/ml, respectively. 3-Chlorophenol and 2-chlorophenol were also fully transformed using 48 units/ml but they required signi®cantly longer reaction times. 2,4-Dichlorophenol could not be fully transformed by 48 units/ml of tyrosinase and pentachlorophenol could not be transformed at all (data not shown). These observations con®rm that all phenols are not equally good substrates of tyrosinase and some cause more rapid inactivation of the enzyme than others. Thus, the quantity of tyrosinase required to achieve a given degree of transformation will vary substantially between substrates. In all cases, colored products were formed during the transformation of these phenols but no precipitates were observed. The impact of these colored products may represent a severe limitation to the application of tyrosinase for treating phenols in industrial wastewaters.
4. Conclusions Tyrosinase activity has been characterized using L tyrosine as a substrate. Maximum catalytic activity was observed at pH 7 with approximately 50% of the maximum was observed at pHs ranging from 5 to 8. Tyrosinase appeared to be unstable at low pH and at elevated temperature. The activation energy for thermal inactivation of tyrosinase at neutral pH was calculated to be 1.85 kJ molÿ1 . Tyrosinase was quite unstable in comparison to peroxidase enzymes which have also been examined for their potential application to the treatment of wastewaters. The optimum pH for phenol treatment was between pH 5 and 8. Tyrosinase was able to transform phenol over a wide range of initial phenol concentration (0.5±10 mM), but no precipitates were observed. Monochlorinated phenols were also successfully transformed with tyrosinase, but the quantity of the enzyme required to achieve a given amount of transformation was substrate dependent. While chitosan addition induced the precipitation of products and depressed color generation, it had no eect on the extent of phenol transformation within the tested range of phenol concentrations. PEG addition did not have any eect on the transformation of phenol. Even though it has been shown that tyrosinase can be used successfully to transform a variety of aqueous phenols, the generation of soluble reaction products is of signi®cant concern since they will impact upon the quality of the treated euent. In order to establish whether the use of tyrosinase is feasible for the treatment of phenolic wastewaters, the toxicity and treatability of these reaction products must be investigated further.
Acknowledgements This work was funded by the Natural Sciences and Engineering Research Council of Canada. References
Fig. 13. Tyrosinase catalyzed treatment of aqueous phenolic compounds as a function of time (phenols0 0:5 mM, tyrosinase dose 8 units/ml for phenol, 48 units/ml for chlorinated phenols, in pH 7 sodium phosphate buer at 25°C).
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