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Characterisation of Tyrosinase
for the Treatment of Aqueous Phenols
Keisuke Ikehata
Department of Civil Engineering and Applied Mechanics
McGill University, Montreal
A thesis submitted to the Faculty of Graduate Studies and Research in partiai
fulfilrnent of the requirements of the degree of Master of Engineering
0 Keisuke Ikehata, 1999
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ABSTRACT
Mushroom tyrosinase (polyphenol oxidase, EC 1.14.18.1) was investigated as an
alternative to peroxidase enzymes for the catalytic removal of phmolic compounds £kom
wastewaters. The maximum catalytic activity was observed at pH 7; however, significant
activity was observed at pHs mghg fiom 5 to 8. Tyrosinase was unstable under acidic
conditions and at elevated temperanires. The activation energy for thermal inactivation of
tyrosinase was determiaed to be 1.85 kJ mol" at p H 7. The transfo-on of phenols
catalysed by tyrosiriase was investigated as a f ' c t ion of pH, initial phenol concentration,
and additives. Phenol was transfod successfuliy with tyrosinase over a wide range of pH
(pH 5 - 8) and a wide range of initial concentrations (0.5 mM - 10 mM). Some chlorinated
phenols were also successfully transformed with tyrosinase. Polyethylene glycol and
chitosan did not improve the transformation efficiency of phenol. However, chitosau was
successfully used to remove coloured products resulting from treatrnent Since coagulation
with aluminium sulfate failed, the colour removd i n d d by chitosan addition appeared to
be the result of simultaneous coagulation and adsorption mecbanisms. Minimum doses of
chitosan required to achieve 90% of colour removal were logarithmically related to the
quantity of phenol treated AU solutions of phenol and chlorophenols treated with
tyrosinase had substantiaily lower toxicities than their corresponding initial toxicities.
Chitosan addition e h c e d the reduction in toxicity very effectively. The toxicities of the
phenol solutions treated with tyosinase were markedly lower than previously reporied
toxicities of solutions treated with peroxidase enzymes.
Le tyrosinase (EC 1.14.18.1) a été étudié comme un alternatif au enzymes
peroxidase pour l'enlèvement catalytique des composés phenoliques des eaw usées. Le
rnaximurn d'activité catalytique a été observé au niveau pH 7; cependant, l'activité observé
au niveaux pH de 5 à 8 était considérable. Le tyrosinase n'était pas stable sous conditions
acidiques et en températures élevées. L'énergie d'activée nécéssaire pour l'inactivité
thermique a été déterminé d'être 1.85 W mol-' à pH 7. L a txansformation des phénols
catalysés par le tyrosinase a été examiné comme une fonction de pH, les premières
concentrations de phénols, et additifs. Le phénols a été tramforné avec succès avec le
tyrosinase dans un large domaine de premières concentrations (0.5 m M - 10 mM).
Quelques phénols chlorurés ont aussi été transformé avec succès avec le tyrosiaase. Le
polyethylene glycol et le chitosan n'ont pas améliorés l'efficacité transformationnel du
phénol. Cependant, le chitosan a réussi d'enlever les produits colorés qui résultaient du
traitement. Puisque la coagulation avec le sulhue d'aluminum n'a pas réussi,
I'enlèvement de couleur produit par l'addition le chitosan a@t être le résultat de la
coagulation simultanée a le méchanisme d'absorption. Les doses minimums de chitosan
nécéssaires pour réaliser 90% d'enlèvement des couleurs étaient apparenté
logarithmiquement à la quantité de phénol traitée. Toutes solutions de phénols et
chlorophénols traitées avec le tyros- avaient des niveaux de toxicités considérablement
plus bases que leurs premières toxicités co~espondantes. L'addition de chitosan augmenta
la réduction des toxicités avec très bons éffets. Les toxicités des solutions phénols traitées
avec le tyrosinase étaient visiblement plus bases que remarqué précédemment dans les
solutions traitées avec les entymes de peroxidase.
ACKNOWLEDGEMENTS
First of all, 1 would like to thank my supervisor, Dr. James A Niceil for giving me
the opportuuity to do this work, and for his encouragement and guidance throughout the
course of this research.
My gratitude is expressed to my coîleagues Monika Wagner, Guoping Zhang, and
Tomas Hoi for their assistance and suggestions in the lab over the course of this work 1
would also like to tbank Diana Brumelis for her tecbnical assistance in the lab. 1 would like
to thank Eric Mills and Anna Tan for their assistance with the translation of the abstract
into French as well as their fnendship. I would also like to express my gratitude to
Mitsuhiro Hishida, Wafa Sakr, Aiex Hiil, Alexandra M a s s a . Mohamed Sheriff, Keiko
Sonoda, Ai Sato and Dr. Daim Ihaku for their fnendship and encouragement.
Finally, 1 am indebted to my parents, M o Ikehata and Tenimi Ikehata, and my
sister, Miyuki Ikehata, for their tremendous supports and encouragement-
TABLE OF CONTENTS
ABSTRACT
L' ABSTRACm
ACKNOWLEDGEMENTS
LIST OF TABLES
LIST OF FIGURES
1 INTRODUCTION
2 LITERATURE REVIEW
2.1 General Characteristics and Structure
2.2 Catalytic Activity and Its Inhiiition
2.2.1 Catalybc Cycle
2.2.2 Kinetic Features of Monophenola~ Activity
2.2.3 Kinetic Features of Diphenolase Aftivity
2.2.4 Inhibition and Inactivation
3.2.4.1 Benzoic Acid
2.2.4.2 Cyanide
2.2.4.3 Carbon Monoxide
2.2.4.4 Reducing Agents
2.2.4.5 Amino Acids and Proteins
2.2 -4.6 Carbon Dioxide
2.2.4.7 mclodextrin
2 -2.4.8 Substrate-hduced Inactivation
2.2.4.9 Themal Inactivation
2.3 Tyrosinase Catalysed Removal of Phenolic Compounds fkom Wastewaters
2.3.1 Mechanisms of Olïgomerisation Reaction and Structures of Products
2.3.2 Factors Anectiag Transformaiion of Phenols Catalysed by Tyrosinase
2.3 -2.1 pH and Temperature
2.3.2.2 Enzyme Concentrations
2.3.2.3 Substrates
2.3.2 -4 Chernical Additives
2.3 -2.5 Immobilisation of Tyrosinase
2.3.3 Toxicity of Treated Phenol Solutions with Various Enzymes
3 MATERIALS AND METHODS
3.1 Materials and Equipment
3.1.1 General
3.1.2 Microtox Experiment
3 -2 Tyrosinase Activity Assay
3.2.1 Procedures
3.2.2 Calculation
3 -3 Tyrosinase Stability Experiments
3.4 Colourhetric Assay for the Measurement of Phenols
3 -5 Tyrosinase Caîalysed Transformation of Phenols
3 -6 Toxicity Measurement of the Transfomed Phenol Solution
3 -6.1 Procedures
3 -6.2 Colour Correction
4 RESULTS
4.1 Characterisation of Tyrosinase Activity . 4.1.1 EffectofpH
4.1.2 StabilityofTyrosinase
4.1.3 Thermal Inactivation of Tyrosinase
4 -2 Tyrosinase Catalysed Transformation of Phenol
4.2.1 EffectofpH
4.2 -2 Effect of Initial Phenol Concentration
4.2.3 Effect of Substrate Type
4.2 -4 Effect of Polyethylene Glycol (PEG)
4.2.5 Effect of Chitosan
4.3 Removal of Colour Remaining in Treated Solutions
4.3.1 EffectofAlum
4.3 -2 Effect of Cbitosan
4.4 Toxicity of the Treated Phenol Solutions with Tyrosinase
4.4.1 Effect of Chitosan
4.4.2 Toxicity of the Treated Chlorinateci Phenol Solutions
5 DISCUSSION
5.1 Characteristics of Tyrosinase Activity
5.2 Tyrosinase Cataiysed Transformation of Phenol
5.3 Colour Removal fiom the Treated Phenol Solutions
5 -4 Toxicity of Phenol Solutions Treated with Tyrosinase
6 CONCLUSIONS AND RECOMMENDATIONS
REFERENCES
LIST OF TABLES
Table 2.1
Table 2.2
Table 2 3
Table 2.4
Table 2.5
Table 3.1
Table 3.2
Table 4.1
Table 4.2
Table 4 3
Table 5.1
Table 5.2
Table 5 3
Sources of tyrosinase and literahirp sources describing methods of
enzyme preparation
Kinetic Parameters of monophenolase activity of tyrosinase.
Kinetic Parameters of diphenolase aaivity of tyrosinase.
Removal of various phenols and aromatic amines by the
tyrosinasecatalysed reaction.
Adsorption enthalpies of phenol, pyrocatechol, andpquinone with
activated charmai and pquinone with chitosan
pH buffers used
Molar extinction coefficient used in AAP assay of phenols.
Summary of inactivation d-y constants, k, and decimal reduction
values, D, calcdated fiom Figure 4.4.
Colour of the various phenolic solutions treated with tyrosinase
and the prrcipitates f o d when chitosan was added before the
initiation of reaction.
Initial toxicity of phenol and chlorophenols.
Cornparison of thermal inactivation parameters between soybean
peroxidase and rnushroom tyrosinase.
Cornparison of toxicities (in TU50) of treated phenoi solutions in
this study with published data
Cornparison of toxicities (in TUSO) of the treated phenol solutions
using different enzymes.
vii
LIST OF FIGURES
Figure 2.1
Figure 2.2
Figure 2 3
Figure 2.4
Figure 2.5
Figure 2.6
Figure 2.7
Figure 4.1
Figure 4.2
Figure 4 3
Figure 4.4
Figure 4.5
Figure 4.6
Figure 4.7
Figure 4.8
Denvatives of the coupleci binuclear copper active site of
tyrosinase.
Catalytïc cycle for the hydroxylaîion of monophenois and the
dehydrogenation of O-diphenols to +quinones by tyrosinase.
Mefanin synthesis pathway.
Structures of quinone and semiquinone radical.
Proposed dimer fomiation during transformation of Zchloro-
phenol in the presence of tyrosïnase.
Proposed possible dimer structure formed nom catechol
oidation with tyrosinase.
Chernical structure of chibsan.
Tyrosinase activity measured in various pH buffer-s at 25°C.
Stability of tyrosinase incubated at 25OC in various b a e n : (a)
linear plot, (b) semi-log plot.
Stability of tyrosinase incubated at 40°C in various buffers: (a)
linear plot, (b) semi-log plot.
Thennal inactivation of tyrosinase in pH 7 sodium phosphate
b a e r (a) linear plot, (b) semi-log plot
Dependence of thermal inactivation decimal reduction value on
temperature for tyrosinase in pH 7 sodium phosphate buffer.
Dependence of thermal inactivation decay constant on
temperature at pH 7 for tyrosinase (Arrhenius plot).
Effect of pH on the transfomation of phenol catdysed by
tyrosinase in the absence of chitosan: (a) phenol transformation,
(b) colour generated at 510 nm.
Relationship between the colour generated at 5 10 nm and phenol
Figure 4.9
Figure 4.10
Figure 4.11
Figure 4.12
Figure 4.13
Figure 4.14
Figure 4.15
Figure 4.16
Figure 4-17
Figure 4.18
transformation.
Effect of pH on the transformation of phenol catalysed by
tyrosinase in the presence of 420 cps chitouin: (a) phenol
transformation, (b) coiour remaining at 510 nm after
cenûifùgation.
Tyrosinase cataiysed transformation of phenol with and without
420 cps chitosan: (a) Iphenoll0 = 0.5 mM, @) [phen~l]~ = 1 mM,
(c) [ p h e ~ l ] ~ = 2 mM, (d) [phen~l]~ = 4 m M
Amount of tyrosinsse required to transfonn 95% of initial
phenol .
Tyrosinase cataiysbd treatment of aqumus phenolic compounds
as a function of tirne: (a) phenol remaining, (b) absorbance .
Effect of PEG on tyrosinase catalysed transfomation of phenol:
(a) phenol remaining, (b) absorbance remaining-
Effect of chitosan on tyrosinase catalysed transformation of
phenol: (a) phenol remaining, (b) absorbance remaining .
Relationship between intensity of the colour generated at 5 10 nm
by tyrosiaase catalysed phenol oxidation and quantity of phenol
transformed foliowing the complete treatment (transformation r
98%) of the solutions with initial concentration between 0.5 mM
and 10 m M
Effect of alum on colour removal at 5 10 nm fiom the fully
treated (transformation 2 98%) phenol solutions.
Effect of chitosan type on the removal of colour at 5 10 nm h m
fiilly treated (transformation 1 98%) phenal solutions.
Removal of the colour fiom the M y treated (transformation 1
98%) phenol solutions by the addition of 420 cps chitosan: (a)
[phe~>l ]~ = 0.5 rxA& tyrosinase dose = 6 uniWrnL, (ô) Iphenoll0
Figure 4.19
Figure 4.20
Figure 4-21
Figure 4-22
Figure 4.23
Figure 4.24
Figure 4.25
Figure 4.26
Figure 4.27
Figure 4.28
= 1 mM, tyrosinase dose = 12 units/mL, (c) [phen~l]~ = 2 mM,
tyrosinase dose = 24 units/mL, (d) ~henoll0 = 4 mM, tyrosinase
dose = 48 units/mL, (e) [phen~l]~ = 10 mM, tyrosinase dose = 96
uniWmL.
Linear regressions of the linear portion of curves for each inibal
phenol concentration: (a) 3 hours incubation the, (b) 18 hours
incubation tirne.
Amount of chitosan required to achieve 9û% colour removal
nom the fûliy trated phenol solutions: (a) linear plot, (b) semi-
log plot.
Linearised Langmuir isotherms: (a) linearised form #I , (b)
linearised form #2.
Linearised Freundlich isotherm
Cornpanson between modelied adsorption isotherms and experi-
mental data of colour removal fiom the treated phenol solutions
by the addition of chitosan
Toxicities of the various concentration of fiilly treated phenol
solutions (transfomation 1 98%) by tyrosinase with and without
420 cps chitosan in pH 7 sodium phosphate b s e r at 25OC .
Relationship between toxicity and colour of the treated phenol
solutions.
Effect of chitosan (added prior to reaction initiation) on the
toxicity and colout of fully treated phenol solutions (trans-
formation 2 98%) by tyrosinase.
Effect of chitosan (added afker the reaction was completed) on
the toxicity and colour of fùlly treatcd phenol solutions (trans-
formation 2 98%) by tyrosinase.
Toxicities of the various chlorophenol solutions treated by
1 INTRODUCTION
Phenolic compounds are present in the wastewaten of a number of industries such
as coal conversion, resim and plastics, petroleum rehing, textiles, dyes, iron and steel, and
pulp and papa (Klibanov et al., 1980). Nearly al1 of these wmpomds are considered to be
toxic and some are suspec?ed carcinogens Conventional methods to remove phenolic
compounds fiom wastewaters include extraction, adsorption on activated carbon, stearn
distillation, bacterial and chernical oxidation, electrochemical techniques and irradiation,
among others.
The process of enzyme catalysed polymerisation and precipitation of phewls and
aromatic amines has attracted much attention since 1980 when Klibanov et al. reported
their first application of horseradish peroxidase. Peroxidase enrymes are widely distributed
in plants such as soybeans, potatoes, cauliflower, and fun@; however, peroxidase fiom
horseradish has received greater attention fiom researchers because of its weil studied
characteristics and wide variety of applications (Nice11 et al., 1992). These enzymes
catalyse the oxidation of a variety of phenols and aromatic amines by hydrogen peroxide.
Phenolic and aromaîic amine radicais are generated during this oxidaîion process and
spontaneously polyrnerise. Since the polymen are less soluble in water, they precipitate
fiom solution and can be physically removed either by filtration or by sedimentation
(Klibanov et al., 1980). Many kïnds of hydroxyl or amino benzene derivatives, including
phenols, biphenols, anilines, benzidines are substrates of this enzyme (Josephy et al., 1982)
and can be treated in this manner.
The application of peroxidase enzymes for the removal of phenolic compounds
fiom wastewaters has a number of potential advantages over conventional biological
treatment. The advantages of this process are: (1) its application to a wide variety of
compounds including those that are biorefiactory and toxic to microorganisms; its ability to
accomplish treatment over wide range of conîaminaut conceniration, pH, and temperature;
and (3) very high reaction rates of the enzymatic reactions wmpared with conventional
microbiological processes.
In spite of these apparent merits, there are several potential disadvantages of this
process. First of all, disposal of darkcoloured, rehctory pipi ta tes is a major concem.
The residuai toxicity of the treated phenol solutions can also be considereâ to be one of the
most serious probiems. Heck et al. (1992) and Aitken et al. (1994) applied a . acute toxicity
assay to determine the toxicity of phenolic solutions treated by peroxidase enrymes and
tyrosinase. Ghioureliotis (1997) studied the toxicities of reaction solutions treated with
honeradish and soybean proXidase. Surprisingly, the toxicities of the solutions after
treatment with both peroxidases were higher than those before treatment. The toxicity of
the treated effluent is very critical, and therefore, it is necessary to fhd ways to reduce it.
The economic feasibility of the process 1s directly related to the msts of the
reagents. Currently, the prïce of horseradish peroxidase is very hi& In addition, the
peroxidase-catalysed oxidation of phenols requim hydrogen peroxide as an oxidant.
Seved efforts to lower the costs of phenol treatment with horseradish peroxidase have
been examined including optimisation of reactor cunEigurations (Nicell et al., 1992; 1993),
application of lower pwity enzymes (Cooper and Nicell, 1996), the use of alternative
peroxidases (Ghioureliotis, 1997; Kinsley, 1998) and the use of protective additives. The
presence of additives such as polyethylene glycol (PEG), gelatin or chitosan significantly
reduced the amount of peroxidase needed (Nakamachi and Machida, 1992; Nicell et al.,
1995). The additive might act as a protector of the enzyme against the fia radicals
produced by erizymatic reaction and entrapment by polymer products (Nicell et al., 1995).
Based on the shortcomings associated with peroxidase enzymes, an alternative
enzyme for this process should be examineci. Tyrosinase (EC 1.14.18.1), also known as
polyphenol oxidase, is a copper-contaking enzyme which catalyses a similar phenol
oxidation reaction to peroxidase. This enzyme 1s widely distributed in fhits, vegetables,
and s e a f d products such as mushmom, apple, avocado, banam, potato, pear, tobacco,
papaya, Florida spiny lobster, brown shrimp, and othm (Janovitz-Klapp et al., 1990, Espin
et al., 1997b, Chen et al., 1993). Tyrosinase is Iargely responsiôle for browning in these
food products (Kahn, 1985).
Tyrosinase catalyses two Merent reactions. The first rcaction is the hydroxylaîion
of monophenols leadhg to o-diphenols, often known as monophenolase or cresolase
(Duckworth and Coleman, 1970). The second r d o n is the oxidation of o-diphenols to O-
quiriones, often referred to as odiphenolase or catecholase. In the both of these oxidation
reactions, oxygen is used as an oxidant. Atiow et al. (19û4) demonsbated that this enzyme
could also be applicable for the oxidation of many types of phenolic wrnpounds such as
chlorophenols, methylphenols, diphenols, and naphthols. Aniline and chlorinated anilines
c m be also oxidised to some extent, and for these difficult-twxidise compounds, CO-
polymerisation with unsubstituted phenol can resuit in good removals (Wada et al., 1995).
However, d i k e to peroxidase enzymes and laccases, it has been reported that the
precipitation of oxidised products of phenols did not occur in the presence of tyrosinase
during the reaction (Wada et al., 1993; Sun et al., 1992). This may becorne fetters for the
practical application of this enzyme to the wastewater treatment. In addition, it is knowa
that tyrosinase was quickly inactivated in the aqueous solution (Wada et al., 1992),
however, protection of the enzymes by protective additives such as PEG has never been
reported in the literature.
Therefore, the objectives of this work were:
(1) to summarise the current level of knowledge reported in the literature for tyrosinase
dealing with its characteristics, its reaction mechanisms, and its applications towards
the treatment of phenolic wastewaters;
(2) to characterise tyrosinase with respoa to its catalytic activityy its stability, and its
ability to cataiyse the traasformation of phenolic compouads nom water,
(3) to examine the effed of the additives, PEG and chitosan, on protecting the catalytic
activity of the enzyme;
(4) to attempt the colour removal from phenol solutions treated with tyrosinase;
(5) to perform a toxicity assesunent of phenolic solutions treated with tyroshase; and
(6 ) to assess the competitiveness of tyrosinase with othex peroxidase enzymes in terrns of
its ability to treat phenolic compounds.
LITERATURE REVIEW
2.1 Geaeral Characteristics and Structure
Tyrosinase (polyphenol oxidase, EC 1.14.18.1) is a coppercontainiog enzyme,
which catalyses the oxidation of tyrosine in Liviag organisms, and is widely dimibuted in
bacteria, fkuits, vegetables, sea foods, and animals (Duckworth and Coleman, 1970; Chen et
al., 1993; Slominski and Cost.iiItjtino, 199 1). This enzyme catalyses two different oxidation
reactions including ortho-hydroxylation of monophenols (xheme 2.1) followed by
dehydrogenation of the o-diphenols to the conesponding quinones (scheme 2.2).
Tyrosinase has been isolateci and purified fiom a number of plant and animal sources.
However, few of these have been wellcharacterised, and preparations ofkm show a
significant degree of heterogeneity (Solomon et aL, 1996). The molecular weigbt of
tyrosinase ranges fiom 13.4 kDa to 128 D a depending on the saurces (DuckWorth and
Coleman, 1970; Solomon et al., 1996). Because of its heterogeneity, Solomon et al. suggested
that there might have k e n some confùsion as to whether the enzymes considered as
tyrosinase are actually tyrosinases, catechol oxidases, or even laccases.
The eorymatic structure of tyrosinases has been snidied by both bioiogical and
chernical approaches with respect to prirnary, secdndary, and tertiary structure, domain
structure, Cu binding sites, and activation mecbanism (Schoot Uiterkamp and Mason, 1973;
Casella et al., 1993; Getlichermau et a', 1 9%; van Gelder et al., 1997). It is widely accepted
that the active site of tyrosinase is quite similar to tbat of hemocyanin, which is a respiratory
copper protein, and contains coupleà binuclear coppers (Schoot Uiterkamp and Mason, 1973).
The simplified structures of the active site at difFerent oxidation states are summarised in
Figure 2.1.
Figure 2.1 Derivatives of the coupled binuclear copper active site of tyrosinase (L = exogenous ligand). Note that the axial nitrogen atoms coordinated to the coppers are omitted for clarity. (Solomon er al., 1996)
Mushroom Agaricm bispom tyrosinase is the only commercially available
tyrosinase and is considerd to be one of the most studied tyrosinases (Duclcworth and
Coleman, 1970; Kahn, 1985; Zhan and Flurkey, 1997; etc.). Although tyrosinase was isolated
and purifieci fiom various bacteria, h g i , plants, and anirnals, the objectives were mostly
aimed at preventing food browning. The sources of tyrosinase and the authors who describeâ
preparation rnethods of enqmes were summarïsed in Table 2.1.
2.2 Catalytic Acîivity and its Inhibition
2-2.1 Catalytic Cycle
The cataIytic mechanism of tyrosinase has been studied for a long time (Duckworth
Table 2.1 Sources of tyrosinase and literature sources describing methods of enzyme preparation.
Source Author(s) Apple Murata et ai. (1997)
Avocado Espin et al. (1 99%) Floriàa Spiny Lobster Chen et ai. (1 992)
Dogrose Fruit Sakiroglu et al. (1996) Mushroom Agaricus bispufus Albisu et al. (1 989)
Neurospora Lerch (1976) p a ~ a ~ a Cano et ai. (1998)
Pear Espin et ai. (1 997a) Potato Leaf Sanchez-Ferrer et al. (1 993)
Tobacco Richardson and McDougaU (1 997)
and Coleman, 1970; Makino and Mason, 1973). Because of its complexity, this mechanism
was believed to be an allosteric mechankm involving two distinct binding sites for oxygen
and aromatic compounds (Duckworth and Coleman, 1970; Jolley et al., 1974). However,
Wilcox er al. (1985) have suggested that the overall catalytic mechanisrn can be explaineci
with one common binding site for both substrates (Figure 2.2). The oxidation state of copper
in resting tyrosinase is mody met derivative, which has two cupric centers (Kerteu et al.,
1972; Makino et al., 1974). In order to initiate the catalytic cycle, a reducing agent rems with
the two copper@) atoms of met-tyrosinase and reduces hem to deoxy-tyrosinase (scheme
When odiphenol is presented in the reaction system, this step might produce a
corresponding O-quinone (see scheme 2.10 and 2.11). But if monophenol was used as a
substrate, the enzyme activation could not occur without an aid of reducer (Naish-Byfield and
Riley, 1992). Molecular oxygen binds with the deoxy-tyrosinase and oxidises it to oxy-
tyrosinase (scheme 2.4).
Figure 2.2 Caîaiytic cycle for the hydroxylation of monophenols and the dehydrogenation of O-diphenols to oquinones by tyrosinase. M = Monophenoi and D = Diphenol bound forms. Axial ligands at Cu are omitted for clarity. (Solomon et al., 1996)
The next step of the redox reaction is cornpetitive between monophenol and (F
a diphenol. When the monophenolic substrate is dominant in the reaction mixture, the oxy-
tyrosinase binds with monophend and oxidises it to d p h e n o l (scheme 2.5 and 2.6).
Consequently, O-diphenoi produceci in this cycle is oxidised further to oquinone (scheme
2.7).
E , + monophenol 4 E ,-M + H+
E ,-M E,-D
E,-D + H+ + Ed-+oguinone+H20
This cycle of reactions is called a monophenolase or cresolase cycle (Figure 2.2). In
practice, the resting form of tyrosinase contains 10% to 15% of oxy-tyrosinase and
monophenolic substrates can react with this oxyamponent (Jolley et al., 1974). But there is
slow and relatively long lag period pnor to the anainment of the steady staie of the reaction
when only monophewlic compounds are useci as substrates (Espin et al., 1997a; 1997b).
When the O-diphenolic cornpound is dominant in the reaction mixture, the oxy-
tyrosinase binds with odiphenol (scheme 2.8) and oxidises it to quinone. The oxy-form of
the enzyme is simultaneously transformeci to the met fonn (scheme 2.9).
E , odiphenol -14 E ,-D +
E ~ ~ - D + ~ H + e, E ~ + o q u i n o n e + H z O
The met-tyrosinase is reduced to deoxy-fom as described scheme 2.3, but another O-diphenol
is used as a reducer at this time and consequently released as second oquinow (scheme 2.10
and 2.1 1). The deoxy-tyrosïnase is oxidised to oxy-fonn with molecdar oxygen again.
This cycle is refened to as a diphenolase or catecholase cycle (Figure 2.2).
2.2.2 Kinetic Features of Monophenohse Activity
Accorduig to the diagram described in Figure 2.2, tyrosinase cataiyses the production
of oquinone fiom both monophenol and o-diphenols, thus O-diphenol is never released as a
product. The oxidation mechauism of L-tyrosine and tyramine which are most well-studied
monophenolic substrates of tyrosinase 1s more complicated than that of simple phenoiic
wmpounds because it encompasses cyclisation of dopaquinone and tautomerisation of
cyclodopa (Figure 2.3); however, an understanding of the O-diphenol production is very
critical to reduce the initial lag period of the oxidation cycle.
In the case of L-DOPA production, there are two main theories: (1) the direct
formation by the hydroxylation of L-tyrosine proposed by Rodngwz-Lopu et al. (1992) and
Ros et al. (1994) and (2) an indirect formation theory proposed by Naish-Byfield and Riley
(1992) and Cooksey et al. (1997). The indirect theory explains that the L-DOPA is not
released at the second step of the pathway described in Figure 2.3, but it is produced by the
attack of inter- or intra-mofecular nucleophiles on the dopaquinone (Cooksey et al,, 1997).
Tyrosine DOPA Dopaquinone
C y c l o ~ p a OopPchr-
Figure 23 Meianin synthesis pathway (simplified) (Naish-Byfïeld and Riley, 1 992).
The examples of nucleophiles are thiol groups on the cysteine midue of the protein
and the amino group in the subsaate for the case of L-DOPA In order to synthesise O-
diphenol fiom monophenol under tyrosinase catalysis, an equivaient reducing agent such as
ascorbic acid is needed to prevent the o-quinone formation and fùrther oxiâation (Piaiis et al.,
i 996).
The lag period prior to the initiation of monophenolase activity of tyrosinase is
affected by several factors. Naish-Byfield and Riely (1992) used an oximetric method to
monitor the oxygen consurnption and found that the lag period decreased non-linearly with
increasing tyrosinase dose. They also investigated the effect of pH over a range fkom pH 5 to
pH 7 and found that at lower pH the lag peziod was shortened. They explain it in ternis of
mass action due to excess protons which inhibit the recruitment of the met-tyrosine. Later Ros
et al. (1994) reported that the monophenol concentration also affectecl the lag p e n d The lag
period increased when monophenol concentration increased The effect of these f m were
mainly observed with mushroom tyrosinase. It is know that the tyrosinases fiom different
sources have different properties and monophenolase activities (Ros et al., 1994). Difierences
in oxy-tyrosinase content between the tyrosinases is considered as a factor for Merences in
the lag period Espin et aL(1997) suggested that the purification process of the enzyme wouid
influence the oxy-tyrosinase content. It is also hown that trace arnounts of reducing agent
such as O-diphenois, dithiothreitol, or ascorbic acid activates met-tyrosinase and shortens the
lag period (Naish-Byfield and Riley, 1992; Cookxy, et al., 1997; Escribano et uL , 1997).
Kineîic analyses of monophenolase activity have been carrieci out by several groups
of researchers. Their results are sutnmarised in Table 2.2. Relatively little information
concerning kinetic parameten of monophenolase activity is available compared to that of
diphenolase activity especially for simple monopbenol compounds.
0 Table 2 2 Kinetic parameters of monophenolase activity of tyrosinase
Substrate v r m u ~ PH Source of Referemce
L-Tyrosine 0.153 L-Tyrosine 0.272 L-Tyrosine 0.827 Tyramine 0.639 Tyramine 0.4 1 T yramine 9.88 Tyramine 0.253
4-Methylphenol 0.0863 4-Hydroxyani sol O. 02
/ ,uM min-' N/A 1.34 N/A 6.88 6.39 6.81 1.6 NIA N/A
N/A 6.5 7 6.5 6.5 6.5 6.8 5 NIA
h h s b ~ m Naish-Byfield & Riley ( 1992) Mushtoom Ros et al. (1994)
Dog-rose f i t s Sakirogiu et al. (1 996) Mushroom Ros et al. (1994)
Frog epidennis Ros et ai. (1 994) Gra~e Ros et al. (1994)
Table beet leaves Edbano et d (1997) Dog-rose fiuits Wroglu et al. (1 996)
M u h o m Naish-Byfield & Riley (1992) 4-Hydroxyanisol O. 3 0.6 5 Avocado Espin et al. (1 99%)
N/A = not available
2.23 Kinetic Features of Diphenoiase Activity
The oxidase reaction is much more rapid than the oxygenation reaction (kdipbcmluc = 7 1 10 s- , k m-k = 1 o3 s") (Solomon et al., 1996). AlthoUgh a nurnber of studies were
conducted many years ago (Yamaguchi et al., 1969; Duckworth and Coleman, 1970; Lerch
and Enlinger, 1972; Lemer and Mayer, 1974)- the kinetic modelling of diphenolase activity of
tyrosinase has not been perfomed very successfully because of its unusual, complicated, and
controvenial reaction rnechanisms. Moreover, as Janovitz-Klapp et al. (1990) suggested,
these studies were mostly c d e d out in air-sanirateci solutions, therefore, the eEect of oxygen
concentration is stili unclear.
Kinetic parameters of diphenolase activity of various tyrosinases and substrates are
summarised in Table 2.3. Duckworth and Coleman (1970) have suggested that the Km for
catechol was decreased by substitution the of pura-position with electron-withdrawing
functional groups. The effect of the ability of the substituent to withdraw electrons on the K,
followed the Hammett rule (Duckworth and Coleman, 1970).
Table 2 3 Kinetic parameters of diphenolase activity of tyrosinase
Substrate K m / vm~, PH Source of Reference m M /,uM min-' Enzyme
L-DOPA 0.263 N/A 7.02 Mushroom DuckWorth & Coleman (1970) L-DOPA O. 168 L-DOPA 0.606
Dopamine 0.42 Dopamine 0.36 Dopamine 0.5 17 Dopamine 0.229 Dopamine 9.32 Dopamine 2.82 Catechol O, 194 Catechol 0.11 Catechol 0.2 Catechol 7.4 1
4-??iiocyanatocatechol 0.08 1 4-Acetylcatechoi 0.027 4-Formylcatechol 0.G 1 5 4-Cyanocatechol 0.0 14 4-Nitrocatechol 0.043
4-Methylcatechol 5 -2
8.17 N/A 50 90 100 100 100 N/A N/A N/A N/A N/A N/A NIA NIA NIA N/A N/A
Mushroom Ros et al. (1994) Dog-rose fruit Sakiroglu et al. ( 1996)
Table beet leaves Emibano et al. (1997) Table beet leaves Escribano et al. (1 997)
Mushroom Ros et al. ( 1994) Frog epidermis Ros et al. (1 994)
G r a ~ e Ros et al. (1994) Dog-rose fruit Sakirogiu et al. (1 996)
Mushroom Duckworth & Coleman (1 970) Mushroom Ingraham ( 1957) Mushroom Yamaguchi er ai. (1969)
Dog-rose f i t Sakiroglu et al. (1996) Mushroom Duckworth & Coleman (1 970) Mushroom Duckworth & Coleman (1970) Mushroom DuckWorth & Coleman (1970) Mushroom Duckworth & Coleman (1 970) Musinmm Duckworth & Coleman (1970)
Apple Janovitz-Kiapp et al. (1 990) 4-Methylcatechol 7.41 NIA 8.5 Dog-rose fhit Sakirogiu et al. (1996)
N/A = not available
2.2.4 Inhibition and Inactivation
Since tyrosinase is believed to be a key enqme responsible for the browning of
many fniits and vegetables, the inhibition and inactivation of tyrosinase activity derived fkom
these plants have been studied in order to preserve these products. The inhibitory effect of an
agent against tyrosinase activity may be caused in at least two ways: by reacting with o-
quinone which is plymerised by itself and forms dark-coloured melanias; and by chelating
with coppers at the active site of the entyme (Kahn, 1985).
Both reversible and irreversible inhibitions of tyrosinase activity were oôsewed-
Reversible inhibitors, are divided into three types: cornpetitive inhibitors; noncornpetitive
inhibitors; and uncornpetitive inhiiitors (Conn et al., 1987). It is known that there is another
type of inhibitor, k, inhibiton, which f h t act as substrates and then are used to produce
compounds that have the abihty to inhr'bit the enzyme (COM et al., 1987). Some of the
inhibitors showed mixed inhibitory effécts involving a combination of competitive and
noncornpetitive inhibition Many chernicals have k e n reported as potentiai inbibitors of
tyrosinase activity. Thermal decay is also considered as a one key inactivation process of
tyrosuiase.
2.2-4.1 Benzoic acid
Duckworth and Coleman (1970) reprted that benzoic acid inhibited diphenolase
activity of tyrosinase and this inhibition was competitive with catechol and irreversible. They
suggested that benzoic acid bound to Cu@) which was associated with the deoxy-form of
tyrosinase. Other arornatic carboxylic acids like cinnamic acid and phenylacetic acid showed
the same inhibitory effect and longer alkyl carboxyl group and additionai bulky substihents
such as methyl groups decreased the effect (Kermsha et al., 1993). It is considered tbat die
accessibility to the active site copper is related to the degree of inbiiitory effect of the
compounds.
2.2.4.2 Cyanide
Cyanide is one of the most well-known and oxygen competitive inhibitors of
oxidoreductases including cytochrome c oxidase, ascotbate oxidase, and peroxidase (Lee et
al., 1 994; Meyer et al., 199 1 ; Sessa and Anderson, 198 1). Tyrosinase is not an exception and
also exhibits an inactivation sensitivity to cyanide (Duckworth and Coleman, 1970).
2.2.43 Carbon Monoxide
Carbon monoxide is a known inhibitor of many copper-contabing oxidases and may
be competitive with oxygen Albisu et al. (1989) have studied the inhibitory effect of carbon
monoxide which was bubbled through tyrosinase extract fiom mushrooms. The authors found
O
that the inhibition was reversible when air was bubbied through the extract which had been
exposed to carbon monoxide. They suggested that the carbon monoxide treatment could
prevent self-inactivation of tyrosinase and preserve the freshness of food prociucts.
2.2.4.4 Reducing Agents
As explained in section 2.2.1, reducing agents lïke asmrbic aciâ, suifite, and thiol
compounds such as reduced glutathione and dithiothreitol activate the met-form of tyrosinase.
However, excess amounts of these compounds react with o-quinone and fom coloiirless
complexes and consequently prevent M e r oxidation of o-diphenol to oquinone. Therefore,
they are considered to be inhibiton of the diphenolase activity of tyrosinase (Golan-Goidhinh
and UXtaker, 1984). Golan-Goldhrish and WhitaLer aiso reported that the reducing agents
inactivate tyrosinase irrevmibly and its lcinetic behaviour appeared to be first order. They
suggested that ascorbic acid undenvent a change to a more reactive species during the early
stage of inactivation and it was likely to be a k, type of inadvation
2.2.4.5 Amino Acids and Proteins
It is believed that amino acids and peptides can inhibit tyrosinase activity in at lest
two ways (Kahn, 1985). One is the &on between oquinone and a nucleophilic amino acid
residue such as the thiol group of cysteine, the thioester group of methionine, and the 6 amiw
group of lysine. These amino acids and peptides which contain these residues react with O-
quinone and fom covalent coupling compoimds. h example of a peptide is glutathion (y-
glutamylcystainylglysine, GSH), which is an oligopeptide also introduced as a reducer in
section 2.2.4.4. The other reaction is the chelation of the residue with active site coppen of
tyrosinase. LHistidine and L-cysteine have particuiarly high affinities for cu2' because of the
imidazole ring of histidine and thiol group of cysteine (Kahn, 1985). These two effécts
usually appeared in combination Kahn (1985) repoitad Glysine, glycine, L-histidine and L-
phenolyalanine inhibited O-diphenolase activity of tyrosuiase in increasing order of
effectiveness. Garcia-Carmona et al. (1988) have reported that L-proline acts as a weak
activator of the monophenolase activity. It is possible that the amino acid residue on the
enzyme which consists of a number of amino acids reacts with its own product oquinone.
This is part of the activation processes of tyrosinase proposed by Cooksey et al. (1997).
2.2.4.6 Carbon Dioxide
Carbon dioxide bas been reported to have an influence on many enzyme activities
including the inactivation of tyrosinase (Chen et al., 1993). Not only the CO2 gas,
supercritical carbon dioxide, which exhibits physicochemical properties intermediate between
those of liquids and gases, has also been known to inactivate several enzymes such as
peroxidase and pectinesterase (Chen er al., 1992). Chen et al. investigated both hi@-pressure
carbon dioxide as a supercritical fluid and CO2 modified air to evaluate the inactivation of
tyrosinase from Flonda spiny lobster (Chen et al., 1992 and 1993). Carbon dioxide is a very
appropriate chernical for this use because it is nontoxic, nonfiammable, inexpensive and
readi 1 y available. However, the inhibition or inactivation mechanisms have not been
described in detail so far.
2.2.4.7 wyclodextrin
Fayad (1997) reported a unique inhibition effect of #%cyclodexnin, which coasists of
seven glucopyranose uni6 linked by a (1-4) glycosidic bond, on tyrosinase catalysed phenol
oxidation It is said that fiyclodewin fomis complexes with phenols, hence it prevents
phenol oxidation catalysed by tyrosinase. The afkîties of phenols to &clodextrin depend
on the chemical structure of these compomds.
2.2-4.8 Su bstrate-Indnced Ina&ation
As fieqenntly mentioned above, since oquinones, the oxidised produa of phenols
with tyrosinase, are highly reactive, they can atîack a nucleophilic group in proximity to the
active site of enzyme @ietler and Lerch, 1982; Albisu et al., 1989)- This is so called "suicide
inactivation" (Garcia-Canovas et al., 1987). The inactivation process can be descrïbed by the
a following schemes which aré the modifications of scheme 2-9 and 2- 1 1.
E o*-D + 3 c E, + oquinone + Hfl
E ,-D + H? '' + E + oquinone + HzO
l b
Ein
where :
Eh = inactivated enqme, k& = inactivation rate constant
According to Dietler and Lerch, the inactivation reaction was fmt order with respect to the
enzyme concentration and higher wncentrations of substrate exerted a protective effect on the
inactivation
2.2.4.9 Thermal Inactivation
Heat treatment is the most utiliseci method to inactivate tyrosinase for stabilising
foods (Lopez et al., 1994). Aithough several reports dealing with themal inactivation are
available, most of them are exclusively for industrial purposes (Robert et al, 1995). Robert et
al. studied the kinetics of themial inactivation of tyrosinase fiom p a h t o (Acanthophoenix
rubra) and the influence of pH. They determined optimal temperature, optimal pH, and
themodynamic parameten (Km and V&) for diphenolase activity of tyrosinase when 4-
methylcatechol or pyrogailol were used as substrates. They ewmiwd the assay at
temperatures ranging from 1 to 50°C and over pHs ranging fiom 2.5 to 8 with 4-
methylcatechol and found that the optimum temperature and pH were approximately 30°C
and pH 5, respectively. They also suggested that purity of enzyme, i-e. the protein content of
a the enzyme preparation, &acted the stability of enryme (Robert et al., 1995).
2.3 Tyrosinase Catalysed Removal of Phenolic Compounds from Wastewaters
Atlow et al. first reported use of tyrosinase for the beatment of phenolic wastewaters
in 1984. Several monophenols and o-diphenols such as phenol, cresol, chlorophenol, and
catechol were removed very effectively with both wmercially obtained and laboratory
extracted tyrosinases. The optimum enzyme dose to treat 50 mg/L (approximately 0.53 mM)
phenol was detennined to be 60 units/mL (Note: the "units" expressed in this literature review
are consistent with the activity rneaswement describeci in chapter 3). Tyrosinase was e f f d v e
in removing phenol with initial concentrations ranging fiom 0.01 g/L to 1 g/L. The authors
also tned to treat real wastewaters obtained fiom a steel coke plant and a staufFer plant
producing triarylphosphates. They reportecl that the phenols were successfully removed by
tyrosinase treatment which resulted in precipitated produccts.
Wada et al. (1993) followed up with the tyrosinase catalysed phenol removal £iom
wastewaters according to the procedures of Atlow et al-; however, no precipitate was forme4
but the reaction solution changed fiom colourless to dark-brown. They assumed that the
enzyme purity might have an effect on the formation of precipitate: i.e. treatment with lower
purity enzyme resulted in the precipitation.
2.3.1 Mecbanisms of Oligomerisation Reaction and Structures of Products
It 1s believed that oquinone produceci by the oxidaîion of o-diphenol (scheme 2-2)
and other reactive intermediates transfomi spontaneously to coloured pigments (Atlow et al.,
1984; Wada et al., 1992; Payne et al., 1992). The oligomerisation reaction is probably
associated with intermolecular nucleophilic addition of the electron-rich oxygen groups (i.e.
carbonyl or hydroxyl of quinone and pbenol) to the 3- or 4- position of oquinone (Figure
2.4). Semiquinone radicais presented in Figure 2.4 may be involveci (Dec and Bollag, 1995;
Hart, 1983).
Figure 2.4 Structures of o-quinone and semiquinone radical. (Hart, 1983)
Figure 2.5 Proposed dimer formation during transformation of 2-chlorophenol in the presence of tyrosinase. @ec and Bollag, 1995)
diphenykmdioxiôe-2.3quinot-œ 2,3,2',36.trrhydroxydip)i.ny(
Figure 2.6 Proposed possible dimer structures formed fiom catechol oxidation with
tyrosinase. (Naidja et al., 1 998)
Dec and Bollag (1995) proposed some probable pathways of oligomerisation of 2-
chlorophenol (Figure 2.5), and Simmons et al. (1989) and Naidja et al. (1998) proposed some
probable coupled structures of oxidised aromatics (Figure 2.6); however, the daails of both
were not clear.
As Sun et al. (1992) suggened, non-enqmatic polymerisation of oquinone is
considered to be a slow reaction; therefore, fiee quinone in the reaction solution may be
converted by itself to more stable intemediates or attached to the enzyme by nucleophilic
reactions with amino acid residues. As a result, the polymers c m t grow sufficiently large ro
that they wodd tend to precipitate.
Naidja et ai. (1998) reporteci the mass spectrometry &ta of the products, which were
supposed to contain a variety of compounds resulting fiom tyrosinase catalyseci oxidation of
catechol. They showed that the molecular weights of these compounds were distributecl fiom
57 to nearly 900, and the most abundant molecular weight range was between 300 and 600.
This suggests that the products were mostly condensates of three to six catecholic molecules.
This is consistent with the hypothesis of other researchers as mentioned above.
2.3.2 Factors Affecthg Transformation of Phenols Catalysed by Tyrosinase
23.2.1 pH and Temperature
The optimum pH of tyrosinase activity depends on the substrates and the source of
enzyme (Espin et al., 1997a and 1997b). However, only one study involving the matment of
phenol has k e n reported so far. In practice, wmercially obtained or crude extract
mushroorn tyrosinase was used in al1 of the literature dealing with tyrosinase-catalysed
phenolic wastewater treatment. Atlow et al. (1984) treated 50 mg/L phenol with 30 units/mL
tyrosinase in different pH buffersers The best removal efficiency was achieved when 50 mM
sodium phosphate buEer at pH 8 was used
No temperature effect has been investigated for phenol treatment with tyrosinase. Xt
is probably because some thermal inactivation studies of tyrosinase have already indicated its
instability upon exposure to high temperature conditions (Robert et al., 1995).
2.3.2.2 Enzyme Concentrations
Atiow et al. (1984) investigated the relationship between concentration of phenol
and tyosinase dose required to achieve over 98% removal .The required tyrosinase dose was
directly proportional to initial phenol concentration over the range of 50 m@ to 1 g/L, and
the ratio was 1.2 units/ml of tyrosinase for each 1 mg/L of phenol. Wada et al. (1993)
followed up with experiments involving the same procedures (including the use of the same
tyrosinase activity assay); however they fouod that the optimum tyrosinase dose for 0.5 mM
was 20 unitdml, which was about three times as small as that reported by Atiow er ai..
2.3.2.3 Substrates
Tyrosinase can transform a variety of phenolic and other aromatic compounds such
as phenol, 2-methylphenol, 3-methylphenol, 2chioropheno1, 3-chloropheno1, 2-
rnethoxyphenol, catechol, resorcinol, 2,3-dimethylphenol, 1-naphthol (Atlow et al., 19&4), 4-
chlorophenol, 3-methoxy-phenol, 4-methoxyphenol, 4-methylphenol, hydroquinone, aniline,
0
0 4-chloroaniline, 3,4-dichloro-aniline (Wada et al., 1995), 2-hydroxyacetophenone, and 4-
hydroxyaceto-phenone (Lenhart et aL, 1997). The treatment r d t s for these compounds are
summarised in Table 2.4.
Table 2.4 Removal of various phenols and aromatic amines by the tyrosinase-catalysed reaction
Compounds Phenol
2-Chlorophenol (a)
3 -C hlorophenol (4)
4-C hlorophenol 2-Methy lphenol
3 -Methypheno1 4-Methylphenol
2-Meîhoxyphenol 3-Methoxyphenol 4-Methoxyphenol
Catechol Resorcinol
Hydroquinone 2,3-Dimethylphenol
Aniline 4-C hloroaniline
3,4-Chloroaailine 1 -Naphth01
Removal of Substrate (%) Atlow er ol., 19W (') Wada et al., 1995 (2) Wada et al-, 1995 O)
100 100
(1) 50 mgL phenols, 300 units/mL enzyme, 50 mM phosphate b e i x (pH 8.0), 2S°C, 5 hours of incubaiion (2) 0.5 m M phenols, 100 unidmL enzyme, 50 rnM phosphate buffkr (pH 7.0), 2S°C, 3 hours of incubaIion (3) The same conditions of (2) with 1 mM phenol used as a ç o - p o l y m ~ o n agent (4) 24 hours of incubation (5) 5 hours of incubation
Aniline was quite difncult to oxïdise by tyrosinase treatment if it was treated alone,
but in the presence of 2 molar quivalents of phenol, this compound could be transformeci to
high levels (Wada et al., 1995). As show in Table 2.4, 0.5 m M aniline was completely
removed fiom the solution in the presence of 1 rnM phenol, whereas only 28% of aniline was
removed when aniline was the only substrate. It is suggested that the removal of aniline was
c a w d by the CO-polyrnerisation reactïon of aniline with oquinone which was derived fiom
phenol oxidation with tyrosinase. & p l ymerisation with humic substances such as guaiacol
was also investigated in order to treat the les-active compounds (Simmons et al., 1989). in
the case of aromatic compounds which are not substrate of tyrosinase such as anisole and
benzyl alcohol, there was no oxidation and removal observed wen in the presence of phenolic
substrates (Payne et al., 1992).
The removal efficiency of substituted phenol was dependent on the type of
substituent group and its positions (Wada et al., 1992; Lenhart et al., 1997). Usdly, para-
substituted phenols were moa easily oxidised. then meto-substitut4 phewls were moderately
oxidised, followed by ortho-substituted phenols. It has been proposed that the O-substituent
interferes with the binding of the substrate to the active site of tyrosinase and results in
reduced levesl of oxidation The effect of the types of substituent foïlowed the Hammett rule
and its parameter, cq which is a standard masure of the electroa donating or withdrawing
capability of substituent groups (Lenhart et al., 1997). For example, wmpounds with high q
such as 3 -hydroxyaceto-phenone, are not oxidised by tyrosinase.
2.3.2.4 Chernical Additives
Unlike the studies involving other phewl oxidases, chernical additives were
examined mostly for the purpose of removal of wloured soluble rnatter when tyrosinase was
studied. Since precipitate has rarely been observed in phenol soluîions treated with tyrosinase,
one must remove the coloud product rern-ng in the solution. Chitosan and other natural
or synthetic cationic polymers were investigated to accomplish the removal of colour f b m
O solutions (Sun et al., 1992; Payne et of., 1992; Wada et al., 1993 and 1995). Chitosan (Figure
2.7) is a deacetylated prduct of chiM which is a polysaccharide found abundantly in nature
in materials such as crabshells (Wada et al. 1993).
Figure 2.7 Chernical structure of chitosan.
Although the mechanism of the reaction between chitosan and oxidised phenols has
not yet been clearly established, the 2-amino groups of chitosan are iikely to perform a
neucleophilic attack on oquinones to form covalent bonds (Albisu et al., 1989; Nithianandam
and Erhan, 1 99 1 ). In order to characterise chitosan adsorption, the adsorption enthalpy of p
quinone with chitosan was determined by Sun et al- (1992). The result strongly suggested that
diis adsorption of the quinone onto the chitosan was pmumably the result of covalent
interactions, which is refemd to as chemisorption (Table 2.5). On the other hand, the
interaction between phenol, pyrocatechol, or pquinone and activated charcoal were weak and
considered to be the result of low-energy physical forces such as hydrophobic interaction.
Table 2.5 Adsorption enthalpies of phenol, pyrocatechol, and pquinone with activated charcoal and pquinone with chitosan (Sun et al., 1992)
-- . -
Adsorption enthalpy, AH0 (kcal mol-') Solute Activated charcoal Chitosan Phenol -6.4 -
Wada et ai. (1993) investigated the use of cellulose, chitin, chitosan, hexarnethylene-
diamine-epichloro hidrin pol ycondensate and p l yethyleneimine to remove coloured products.
The first three were natural polymers and the last two were synthetic cationic polymers that
O
had amino groups.
At first, they added natural polymen to reaction mixtures containing 0.5 mM pbenol
in pH 7 phosphate buffer at 25OC (Wada et QI., 1993). The reaction was initiated by the
addition of 20 units/rnL of tyrosinase. Met a 2-hour p e n d of reaction with 1.4 m m of
chitosan, the colour was diminished. Chitin also removed colour effectively, but celiulose had
no effect on the colour removal. However, the amount of chitasan quoted above was
considered to be too hi& for practicai use.
Secondly, they added chitasan or two synthetic polymers to treated phenol solutions.
They considered the additives to be acting as coagulants in these experiments (Wada er ai.,
1995). The coloured products were successfùily removed using very small amomts of
chitosan The optimum dose range of chitosan for phenol was reported to be nom 40 to 90
mgL. It was 15-35 fold smaller than the requirement when the chitosan was added pnor to
the initiation of the reaction (Wada et al., 1993). The authors suggested that it is much more
effective to use chitosan as a coagulant rather than as an adsorbent.
Ln the case of other enzymes, such as peroxidases, chemical additives have been used
to prevent inactivation and prolong the catalytic life of the enzymes (Nakamoto and Machida,
1992; Wu et al., 1993; Kinsley 1998). Nakamoto and Machida (1992) showed that gelatin and
polyethylene glycol (PEG) with an average molecular weight larger than 1000 @mole were
very effective in suppressing the inactivation of horseradish peroxidase. In this way, the
amount of horseradish peroxidase required to treat phenol with concentraîions between 10
and 30 g/L was 200-fold less than that required without the additives. Wu et al. (1993) also
proved that the PEG addition could reduce the required enzyme by dose between 40- and 75-
foId when lower (1 and 10 mM) initial concentrations of phenol were treated. Mi& casein,
bovine senun albumin, polyvinyl alwhol and borate were also shown to be effèctive
(Nakamoto and Machida, 1992).
Polymerised phenols have a number of hydroxyl groups in their structure. Nakarnoto
and Machida (1992) suggested that these highly hydrophilic polymers may heract with the
enzyme and form hydrogen bonds. This results in inactivation of the enzyrnes. Additives such
as PEG and borate cau also interact with polymerised phenols and preveut the inactivation of
enzymes. The authors also suggmed that the suppression effe* of PEG depended on PEG
molecular weight. Kuisley (1998) reported that the suppression effect of PEG on soybean
peroxidase inactivation was highest d e n the highest molecdar weight of PEG was ured. He
concluded the mechanism of PEG protection of the peroxidase enzymes may be related to the
water binding properties of PEG.
2.3.2.5 Immobilisation of Tyrosinase
Since inactivation of tyrosinase was considered to be associated with the oquùione
attachment on the amino acid residue in proximity to the active site of enzyme (Dietler and
Lerch, 1982), tyrosinase was immobilised on several support materials in order to improve the
stability for storage, catalytic lifetime and reusability of the enzyme.
Sarkar et al. (1989) immobilised laccase, glucose oxidase, tyrosïnase, PD glucosidase, and acid phosphatase on severai types of clays and soils activated with 3-
aminopropyltriethoxy-silane and glutaraldehyde. The mils used for the enzyme support were
silt and sandy loam soil, clay, and commercially obtained bentonite and kaolinite powder.
The enrymes were successfully immobilised on the soils and clays and retained large amormts
of activities. In the case of tyrosinase, the retained activity was over 60% for ail types of soils
and clays. Resistance to heat and protease attack was slightly improved.
Wada et al. (1992) immobilised tyrosinase on magnetite (Fe304) activated by the
same chernicals which Sarkar et al. wd When 1 mg of tyrosinase was used for
O immobilisation, immobilisation yield was about 80% and retained activity 7040% with 500
mg of the support materid. M e r 15 days of storage, the loss of activity of immobilised
tyrosinase was about 5%. Three types of chlorophenols, methylphenols and methoxyphenols
were treated with this immobilised tyrosinase. The immobilised tyrosinase could be used 5
times without significant reduction of activity, whereas soluble tyrosinase was inactivated
rapidl y.
Wada et al. (1993) aIso used a weakly acidic cation exchange resin, Diaion WK-20,
as a support for the enzyme. Tyrosinase was immobilised on the resin using lethyl-3(3-
dimethyl-aminopropyl) carbodiimide hydtochionde as a crosslinking agent. 7000 units of
tyrosinase were immobilised on 500 mg of resk and immobilised tyrosinase actïvity was
1 6.3% of that added. After 96 hours of storage at 25°C in O. 1 M phosphate bufEer (pH 7.0),
50% of the initial activity of immobilised tyrosinase was retaind This immobilised
tyrosinase could be used over 10 tirnes to treat 0.2 m M of phenol solution.
Payen and co-workers imrnobilised tyrosinase into chitosan gel in several ways
(Patel et al., 1994; Sun and Payne, 1996; Lenhart et al., 1997). According to their newest
preparation of chitosan gel (Sun and Payne, 1996), chitosan was dissolveci in 8% (vh) acetic
acid solution and stirred overnight. After centrifugation of the solution to remove undissolveci
chitosaq the viscous chitosan solution was added through a syringe needle into an 8% NaOH
solution The chitosan gel formed in the NaOH solution was spread on square glass slides.
Highly concentrated tyrosinase was added to the central region of the square and then two gel
films were combined so as to containhg tyrosinase between them and were then sealed using
rubber cernent. Chitosan was used not only for the support of the enzyme, but aiso for the
sorbent, therefore, this immobilised tyrosinase could not be reused.
Pialis et al. (1996) immobilised tyrosinase on chernically modified nylon 6,6
membranes. A nylon disc was successively modified using 3,3',5,5'-tetramethyl-beIlZidine,
~N'4cyclo-hexylcarbodiimide, and glutaraldehyde. They used this immobilised tyrosinase
to produce L-DOPA fiom tyrosine.
2 Tosicity of Treated Phenol Solutions with Various Enqmes
The toxicity of phenol solutions treated with enzymes has attractd more and more
concern. Acute toxicity assessrnent using a Microtox assay of the soluble reaction products of
several enzymatic treatments was investigated by Aitken er a/. (1993 and 1994). The
Microtox assay is based on the acute toxicity eEect of aqueous substances on a
bioluminescent marine bacterium Vibrio fischeri. Toxicity is measured by meashg the
reduction of light output after a specified exposure time (usually 5 or 15 minutes).
Horseradish peroxidase, lignin peroxidase, chloroperoxidase, and tyrosinase were used to
oxidise eight phenolic cornpounds including phenol, 2-chlorophenol, khlorophenol, 2-
methylphenol, 4methylpheno1, 2-nitrophenol, 4-nitro-phenol, and pentachlorophenol. Most
of the treated solutions showed substantially higher toxicity than the solutions of their parent
cornpounds except for the case of 4methylphenol oxidised by horseradish peroxidase and 4-
chlorophenol oxidised by tyrosinase. It was aiso shown that the toxicities of treated phenol
solutions were affected by the pH of the reaction mixture.
Ghioureliotis (1997) investigated the toxicities of partiaily treated phenol solutions
as a fûnction of the fhction of phenol removal using horseradish peroxidase and s o y h
peroxidase as catalysts. Generally, the toxicities of the phenol solutions treated with soybean
peroxidase were slightly higher than horseradish peroxidase. The toxicity of the soluble
byproduct in the partially treated phenol solutions increased when the £tactions of treated
phenol were increased Polyethylene glycol addition did not significantly alter the toxicities of
the treated solutions. The toxicities of the treated solutions of various chlorinateci or
methylated phenols were aiso examined. In the case of phenol, 2-chlorophenol, anà 2-
methylphenol, the toxicities of the solutions incfead after beatment with both peroxidases.
For the rest of the phenols including 3-chlorophenol, 4-chIoropheno1, 2,4-dichlorophenol,
pentachiorophenol, 3 -methy lphenol and 4-methylphenol, the toxicities of the treated solutions
were lower than their comsponding initial toxicities.
A mutagenicity shdy of enzyme-treated aqueous solutions of various phenols was
reported by Massey et al. (1994) using the Ames Salmonella tjphimurim plate incorporation
assay with two different strains, TA 98 and TA 100. They examined the same classes of
enzymes and phenols as those used for the Microtox toxicity assay in their concurrent study
(Aitken et al., 1994). Al1 the a s t e d solutions except for those of 2-aitrophenol and 4-
nitrophenol treated with lignin peroxidase did not exert mutagenicity. The parent compounds
were also tested, and none of them were determined to be mutagenic over the concentration
ranges that were studied
3 MATERIALS AND METHODS
3.1 Materiais and Eqoipment
3.1.1 General
Mushroom tyrosinase (polyphewl oxidase), d o g u e code TY, was obtained
fkom Worthington Biochemid Corporation (Lakewood, New Jersey) and stored at 4°C.
The specific activity was quoted by the Company as 500 mits pet mg, where one unit of
activity corresponds an increase in absorbarice at 280 nm of 0.001 per minute in a reaction
mixture containhg 0.1 mM Ltyrosine at pH 6.5 at 2S°C. Aqueous solutions of tyrosinase
(6 g/L) were prepared ushg disti11ed-deionized water immediately before use, and were
stored at 4OC for several days. Deionùed water was supplieà using a W741 Nanopure
Ultrapure Water System manufacturecl by Ba~l~teadîThermolyne. L-Tyrosine was
purchased fkom Sigma Chernids (St Louis, Missouri). USP grade oxygen gas (99.5%
purity) was purchased fiom Praxair. Phenol (99.5%+ purity) was purchased fiom Fiuka
Chemical Corporation (Ronkonkona, New York). 2-Chlorophenol (990/0), 3-chlorophenol
(98%), khlorophenol (99%+), 2,4-dichiorophenol (98%) were purchased fiom Aldrich
Chemicals ~ l w a u k e e , Wisconsin). Stock solutions 1 mM, 2 mM, and 20 m M of phenol
were prepared using deionized water. Stock solutions of chlorinated phenols were prepared
at a concentration of 20 m M using 20% aquwus methanol. Polyettiylene glycol (PEG) with
average rno1ecula.r weights of 20000 and 35000 were purchased fiom Fluka Chemical
Corporation, and other molecular weights of PEG (2000, 4600, 8000, and 10000) were
purchased fiom Aldrich Chemicals. Al1 chitosan samples with viscosities of 10, 100, 420,
930, 2920, and 5700 centepoise (cps) were obtain fiom Vanson (Redmond, Washington).
Stock solutions of PEG were prepared at a concentration of 32 giL with deioaized water
and stored at 4°C. Stock solutions of f % W N chitosan were prepared with 15% acetic acid
and stored at 4°C. Monobasic and dibasic sodium phosphate were purchased nom
Anachernia Science (Rouses Point, New York). Citric acid and sodium citrate were
a purchased fiom Sigma Chemicals. Bonc acid, sodium borate, and sodium hydrate were
- purchased fkom Fisher Scientific. ACS grade sodium bicarbonate and potassium
ferricyanide were purchased nom Fisher Scientific (Montreal, Quebec). 4-Aminoantipyrine
(98%) was purchased nom Aldrich Chemicais. The preparation of the buEer stock
solutions used in this work is describeci in Table 3.1.
Table 3.1 pH buffers used.
BufTer 1 Conjugate Acid Conjugate Base Deionized
0.05 M Citrïc pH3 pH4 pH5
0.1 M Sodium Phosphate
(mL) (mL) Water (mL) 0.1 M Citric Acid O. 1 M Sodium Citrate
46.5 3.5 50.0 33.0 17.0 50.0 20.5 29.5 50.0
0.2 M NaH2P04 0 -2 M Na2HFQ4
PH 8 O. 1 M Borate
Colourimetric assays for tyrosinase and phenoi and the absorbante of samples
were monitored using an Hewlett-Packard HP845x UV-Visible Spectrophotometer. G l a s
and quartz crystal cuvettes with a 1.5 mL volume and an optical path length of 1 cm were
purchased n o m Hellma Ltd. (Concord, Ontario). Al1 pH measurements were perfonned
using an Orion SA520 pH meter with an Onon Ross 8102 multiple electrode fiom Orion
Research Inc. The p H meter was routinely dibrated using pH 4.0 and 7.0 standards.
Precipitates fiom the enzymatic transformation were removed by centrifûgation at 3500
RPM for 15 minutes with an IEC Centra-8 centrifuge nom International Equipment
Company (Needham Heights, Massachusetts). A RTE1 1 1 water bath Erom Nesiab was used
a to maintain the temperature of enzyme solutions for the thennostability experirnents
5.3 94.7 100.0 0.2 M Boric Acid 0 .O5 M Borax
PH 9 O. 1 M B o M a O H
50.0 4.9 54.9 0.05 M Borax 0.2 M Sodium Hvdrate
conducted between 10°C and 50°C. Madel 5OOO series micropipetters manufactureci by
Nichïryo Co. Ltd of Japan were used to deliver liquid volumes between 10 and 1,000 pi,.
Micropipetters were fitted with Fisherbrsnd Uni-tips purchased from Fisher Scientific.
3 . 1 Microtox Aaalysis
A Microtox mode1 500 analyser pirrchased fkom Microbics Corporation (Carlsbad,
California) was used to evaluate the toxicity of the treated phenolic solutions. The
instrument was interfhced to a cornputer running Microtox statistical analysis software
(version 7.84) for data collection and interpretatioa Microtox reagent (fieezedned strain
of marine bacterium Vibrio jischeri), reconstitution solution (non-toxic ultra pure water),
diluent (non-toxic 2% NaCl solution), and osmotic adjustment solution (non-toxic 22%
NaCl solution) were purchased from Microbics Corporation AU these reagents were stored
at room temperature, except for the Microtox reagent which was kept at -15°C. Deionized
water was used for primary sample dilution
3.2 Tyrosinase Acîivity Assay
Since tyrosiaase catalyses two different oxidation reacbons, the substrates used to
determine its activity are divided into two groups, which are monophenols and diphenols. A
continuous spectrophotometnc rate detemination method was used to monitor the change
of the absorbance due to the transfomation of the substrates to products. A number of
substrates can be used to detexmine p s i n a s e activities (DucWorth and Coleman, 1970,
Espin et al., 1997b). Since al1 of the substrates used in this work were monophenols, L-
tyrosine was selected as the basis of the activity assay. This activity assay consists of 1 rnM
L-tyrosine, 0.1 M pH 6.5 sodium phosphate bufTer, and 6 mg/mL tyrosinase reacting at
25OC and pH 6.5 (Worthington Manuai, 1977). Tyrosinase oxidises L-tyrosine to L-3,4-
dihydroxyphenylalanuie (L-DOPA) which in tum is oxidised to dopaquinone. The latter
reaction is accompanied by an increase in absorbance at a wavelength of 280 nm wbich
was monitored by a spectrophotometer.
3.21 Procedure
The procedure to masure tyrosinase activity was as follows: (1) preparation of
reaction cocktail wnsisting of 10 mL of 1 mM Gtyrosine, 10 mL of 0.1 M sodium
phosphate buffer, and 9 mL of deionid water, (2) oxygenation of the reaction cocktail for
5 minutes, and (3) mhhg of 33 pL of tyr osinase solution and %7 pi, of reaction cocktail
in a 1.5 mL quartz crystai cuvette with a 1 cm pathlength and (4) monitoring the change of
the absorbance at 280 nm for approximately 10 minutes. The enzyme solution was dilutad
to 500 - 1000 units/mL (see below for the unit definition) with deionized water, if necessary.
33 of deionized water was used as a control. Al1 enzyme assays were performed in
triplkate.
3.2.2 Calcuiation
The rate of increase of absorbance at 280 nm (Azcro nm) is proportional to enzyme
concentration. An initiai lag is obxrved for 1-3 minutes depnding on the concentration of
enzyme. One unit will remit in hcrease in om of 0.001 AU per minute at pH 6.5 at
25°C in a 3 mL reaction mixture (Worthington Manuai, 1977). Activity of the stock
enryme is calculateci according to equatioas 3.1 and 3.2 below:
- 1 Act,,, - x - x b
dt a
where: Act- = activity in the cuvette (units/rnL)
Azso - = maximum dsorbance change at 280 nm (AU)
a = 0.001 AUfmin
b = 1/3 units/mL
where: AC^^^ = activity of the stock solution ( u n i d d )
V, = volume of the total assay solution = 1 mL
V'lk = volume of the sample d = 0.033 mL
3 3 Tyrosinase Stabüity Experimenb
The stability of ty~osiniise was evaiuaad by incubating the enzyme at 2PC in
various pH buffers, and at various temperatures in pH 7.0 sodium phosphate b d e r . Initial
tyrosinase activity was 300 units/mL. Tyrosinase activity in the incubation mixture was
measured over time using the wntinuous spectrophotometric methad described in section
3.2.1.
3.4 Colourimetric Assay for the Measarement of Phenols
Phenolic cornpond concentrations were detennined by a colourimetrïc method
based on the absorbance at 510 nm caused by the reaction between phenoiic compounds
and 4-aminoantipyrine (4-AAP) and potassium ferricyanide under alkaline
conditions maintaineci using 0.25 mM sodium bicarbonate buffer. The colour intensity is
linear with respect to the concentration of phenolic compound with different molar
extinction coefficients (6) mmponding to phenolic compound (Table 3.2). The reagents
were added in a plastic tube in the following order
800 pL of sample diluted with 0.25 M sodium bicarbonate buffer (pH 8.4)
100 pL of 20.8 mM 4-AAP (2.08 mM in cuvette)
100 pL of 83 -4 mM K3Fe(CN)6 (8.34 mM in cuvette)
The mixture of the sample and reagena was transferred to a g las cuvette and the
absorbance at 5 10 nm was measured using a spectrophotometer over a six minute period
Table 3.2 Molar extinction coefficient used in AAP assay for each phenols (8 5 0.998)
Assay Substrate Range of Concentration (mM) E (L-mol-'-~rn-~)* Phenol O - 0.10 9.862
2-C hlorophenol O - 0.05 12.58 3-Chlorophenoi 0 - 0.05 12.72 4-Chlorophenol O - 0.10 4.257
2,4-Dichlorophenol 0-0.10 3.766 * extinction coefficient of the d o d product arising h m the coiorimctric assay.
following the addition of K*e(CN)o- Ml sarnples were analyseci in dupiicate.
This assay represents a modification of the direct photometric method, which is a
standard analytical procedure for phenols (Eaton et al., 1995). The modified assay employs
higher concentrations of AAP and potassium ferricyanide reagents, allowing the
measurement of higher phenol concentrations than under the standard method, while also
using smaller sarnple volumes (Buchanan, 1996).
Since the phenolic solutions treated with tyrosinase had colour whicb absorbed a
broad range of visible light and interfered with the results, the absorbane at 5 IO nm of the
original sample diluted to the same concentration by sodium bicarbonate buffer was
subtracted. The calculation of the phenol concentration is:
where: ph] = phenol concentration (M)
A5iOnm(u, = absorbance at 5 10 nm of the assay sample with reagents (AU)
ASIOmn(org) = absorbance at 510 nm of the original sample (AU)
E = molar extinction coefficient (~mol-'*crn-')
1 = pathlengh (1 cm)
df = dilution factor
3.5 Tyrosinase Catalysed Transformation of Phenols
Tyrosinase-catalysed transformation of phenols was conducted under several
conditions: 0.5 mM to 10 mM of initial phenol concentration, pH range of 2 to 10, various
doses of additives, several types of phenols, and various doses of tyrosinase. These
conditions were used to evaluate the abiliîy of tyrosiuase to catalyse phenol tramformaton
and precipitation fiom aqueous phenoi solutions. In a batch reaction, concentrated aqueous
solutions of tyrosinase, phenol, and additives were added to a 20 mL glass via1 and adjusted
to a desùed concentration with deionized water and bufTer. In al1 batch reactions, Teflon-
coated stir bars and magnetic stirrers were used and the temperature was controkd at
2511°C. M e r a period of several hours, the reaction solution was centrifuged and the
residual phenol concentraton of the supernatant was measured by wlouIixnetric assay.
In cases where the phenol concentration was p a t e r than 0.5 mM, the dissolved
oxygen in the reaction solution was insufncient to accomplish a cornplete reaction.
Therefore, the reaction vials were left uncapped to allow continuous replenishment of
oxygen. Control sarnples containing only phenol demonstrated that volatilisation did not
contribute to phenol losses during treatrnent. However, in order to compare the treatment
efficiency of ty~osinase for various phenols, a sample concentration of 0.5 m M was chosen.
This selection was made in order to allow the vids to be capped when treating phenols of
high volatility. In this study, the oxygen consumed by tyroskse relied on dissolved oxygen
in the reaction solution. Aithough the quantity of dissolved oxygen was sufficient to
accomplish the Mi transformation phenolic compounds, it should be noted that the use of
excess quantities of oxygen (e.g. through bubbling of air or O2 gas through the reacting
mixture) rnight alter the rate of reactions as well as the nature of products. This was not
done in this shidy due to the potential for the stripping of dissolved phenols into the gas
phase when bubbling is used to maintain hi& dissolveci oxygen levels in the reaction
mixture.
3.6 Toxicity Merisurement of the Transformed Phenol Solution
Solutions of 0.5 mM, 1 mM, 2 mM, and 4 mM phenol and 0.5 rnM solutions of
chlorinated phenols were treated according to the method descn'bed in section 3 S. For al1
phenol solutions, toxicity was rneasured at 3 hours and 20 hours after the reaction had been
started Because of their s l o w reaction rate, chloriuated phenois were incubated for
periods of 24 hours and 48 hom. The toxicity of ail solutions were measured before and
after treatment.
3.6.1 Procedures
The Microtox MSOO analyzer consists of 30 sample cuvette welis which are
configured into six rows of five wells each. The temperature inside these wells was
maintained at 15 + 0. 1°C. The measurement of the toxicity was performed using the
following steps:
1. The Microtox reagent was prepared with 1 mL reconstitution solution and store it at
5°C;
2. The sample salt concentration was adjusted to 2% NaCl using 10% of sampte volume
of osmotic adjustment solution (primary dilution may be required);
3 Four series of dilutions and one contra1 were made in the i5rst row of the cuvettes;
4. 50 pL of diluent in the second row of the cuvettes was added and waited until the
temperature reaches 15°C;
5. 10 of Microtox reagent was added in the second row of the cuvettes;
6. Mer 15 minutes addition of the reagent, the light intensity, h,, was measured;
7. The samples were transfered to the second row of the cuvettes;
0 8. After 5 minutes, the light intensity, was measured.
The ratio of light lost to Lght remaining (T) was calculated by the following expression:
where, Cr is the correction factor, which is the fraction of light remaining in the blank
sample after t = 5 minutes. Usually, the c o d o n fàctor was between 0.8 and 0.9. A log-
log plot of ï versus sample concentration provides a linear relationship. The 50% e f f d v e
concentration (EC50) was de- as the concentration where the Gamma value was quai
to 1. The toxicity unit (TU50) was obtained fiom the inverse of the EC50. TU50 is simpler
to interpret than EC50 because it is linear with respect to sample concentration, and a
larger TU50 corresponds to a larger toxicity.
3.6.2 Colour Correction
The phenol solutions treated with tyrosinase had a brown colour. The colour
attributed to an absorbance at 490 nm intefieres with the emission of the light fiom the
microorganism and results in an underestimation of TU50 (Dr& R & D Technical Report).
Therefore, a colour conection procedure was pdormed as follows:
1. Measure the light intensity of the solution (Io and I,) as usual;
2. Measure the absorbance at 490 nm of the solution by using a spectrophotometer;
3. Correct the initial light intensity of the solution with the measufed absorbance using
the following calculation;
where: ACb = absor&ncearrected lo
A490 nm = absorbace at 490 nrn
4. Calculate the ï value with this absorbance-corrected initial light intensity.
4.1 Characterisation of Tyrosinase Activity
4.1.1 EffkctofpH
The effm of pH on tyrosinase activity was measured by varying the pH bufFer
used in the activity assay. The types and range of pH bufEers used arc described in Table
3.1. As shown Figure 4.1, the maximum activity was achieved when pH 7 sodium
phosphate buffer was used an4 therefore, pH 7 was d e e A as optimum. At least 90% of
optimal activity was achieved between pH 6.2 and 7.3 and at least 10 % of optimal activity
was achieved between pH 4.2 to 8.8. It is observeci that tyrosinase was less active in weakly
basic bufFen than in w&y acidic buffers.
4.1.2 Stability of Tyrosinase
The stability of tyrosinase incubated at 25°C and 40°C in various pH buffers is
presented in Figure 4.2 and Figure 4.3, respectively. Calculations predicted that the pH of
the assay solution which was originally 6.5 would not be changed signincantly by the
addition of aliquots of tyrosinase at other pHs (e.g. pH = 6.49 &er pH 3 eatyme aliquots
were added and pH = 6.50 d e r pH 10 enqme aliquots were added).
Tyrosinase activity d e c r d with time in every pH b e e r and at both
temperatures. The rate of inactivation was much faster at 40°C than 2S°C. It is suggested
that tyrosinase is vexy unstable especially at high temperature. At 2S°C, the activity first
increased in mon pH solutions and then decreased. The optimum pH was 7 for tyrosimse
at 25"C, and its 65% of the initial actinty was retained after 33 hours of incubation.
Tyrosinase was inactivated very rapidly under acidic conditions; however, it was relatively
stable under basic conditions at both temperatures. The activity curves appeared to follow
first order kinetics (Figure 4.2 (b) and 4.3 (b)). Tyosinase was most stable at pH 6 and 7 at
40°C, but approximately 90% of the initial activity was gone after 5 hours of incubation in
Citrate Buffer Phosphate Buffer Borate Buffer NaOWBorax Buffer \
90% of Optimum
10% of Optimum
Figure 4.1 Tyrosinase activity measured in various pH buffers a? 2S°C.
O pH4(CitrateBuffer) r pH 5 (Citrate Buff.0
i pH 7 (Phosphate Buffer) pH 8 (Phaspham Buffer) pH 9 (Borate Bufier)
O pH 10 (Borax/NaOH Buffer)
Time (min.)
pH 3 (Citrate Buffer) î E
0 pH 4 (Citrate Buffer)
3 r pHS(CitrateBuffer) m g
c v pH 6 (Phosphate Buffer)
3 w
pH 7 (Phosphate Buffer)
E O pH 8 (Phosphate Butfer) m m > m œ
pH 9 (Borate Buffer) CI O pH 10 (BomxNaOH Buffer) ü
Tirne (min.)
Figure 4.2 Stability of tyrasinase incubateci at 2S°C in various buffen: (a) linear plot, (b) semi-log plot.
(a)
pH 3 ( C i t e Buffer) O pH4(CitrateBuffer) r pH 5 (Citrate Buffer) v pH 6 (Phosphate Buffer) i pH 7 (Phosphate Buffer) 6i pH 8 (Phosphate Buffer)
pH 9 (Borate Buffer) O pH 10 (Bomx/NaOH Bufhr)
Time (min.)
1 O0 î E 5 .I
r 3 u pH 3 (Citrate Buffer) B .I
\ O pH 4 (Citrate Buffer) >
a- * r pHS(CitrateBuffer)
Y v pH 6 (Phosphate Buffer) pH 7 (Phosphate Buffer)
Cl pH 8 (Phosphate Buffei) pH 9 (Borate 6-r)
O pH 10 (BomcNaOH Buffer)
I O 0 200 300 400
Time (min.)
Figure 4.3 Stability of tyroshase incubated at 40°C in various buffers: (a) linear plot, (b) semi-log plot
pH 6 sodium phosphate buffer.
4.13 Thermal Inactivation of Tyrosinase
The tyrosinase activity over time at five different temperatures at pH 7 is preseated
in Figure 4.4. The activity d e c d very quickly at 50°C and disappeared completely after
80 minutes of incubation. On the other hand, it decreased very slowiy at 10°C and
remained at airnost 95% of its initial value after 26 hours of incubation, It was observed
that the thermal inactivation of tyrosinase was first order and could be modeiied using:
where A(t) is the activity at time t, A. is the activity at t h e t = O, k is the inactivation decay
constant, and D is the inactivation decimal reduction value. The decimal reduction value is
a measure of the time required for the activity to fall to 10Y0 of its original value, and it uui
be calculated from the inactivation decay constant with following equation:
Calculated inactivation decay constants and inactivation decimal reduction values are
shown in Table 4.1 and Figure 4.5. The relationship of the log D and the incubation
temperatures was linear.
Decimai reduction values may be related to incubation temprrature using (De
Cordt et al., 1992):
where DREF is the decimal reduction value at temperature TREF and Z is the temperature
Time (min.)
Io00 1500
Time (min.)
Figure 4.4 Thermal inactivation of tyrosinase in pH 7 sodium phosphate b d e r (a) iinear plot, (b) semi-log plot.
1000000
100000
ioooo
1000
1 O0
10 20 30 40
Temperature, T ( O C )
Figure 4.5 Dependence of thermal inactivation decùaal reduction value on temperature for îyrosinase in pH 7 sodium phosphate b a e r .
0 change required to obtain a ten fold increase or decrease in the decimal reduction value.
The Z value was calcdated ta be 10.4OC using the iinear regression of chia in Figure 4.5.
Table 4.1 Summary of inactivation decay constants, k, and decimal reduction values, D, calculateci f?om Figure 4.4.
Temperatme (OC) Temperature (K) k (min-') ( m m 10 283.15 8.92~ 10" 2.58~ 1 o4 25 298.15 2.43 x 104 9.49~ 103 30 303.15 1.10~ 10-~ 2. 1ox103 40 3 13.15 8.64~ 10" 267 50 323.15 5.9% 10'~ 38.4
Figure 4.6 is an Arrhenius plot showing the temperature dependence of thermal
inactivation decay constants fiom the &ta shown in Table 4.1. The Arrhenius activation
energy, E,, can be calculated using the Ar~henius' equation:
where A is a frequency factor which is inherent in the reaction, R is the gas constant, and T
is the absolute temperature (Yoshioka and Ogino, 1976). B a d on a linear regression
analysis of the data in Figure 4.6, E. was 1.85 kJ mol-' and A was 1.73 x 10'' min.".
Figure 4.6 Dependence of thermal inactivation decay constant on tempe- at pH 7 for tyrosinase (Arrhenius plot).
4.2 Tyrosinase Catalyseci Transformation of Pknob
The transformation of phenol with tyrosinase was investigated as a fiinction of pH,
initial phenol concentration, entyme dose and additives. The transformation of some
chlorine-substituted phenols was also investigated The incubation the was usually 3 hours.
Reaction solutions which did not contain tyrosinese were prepared as controls during each
experiment in order to see if signifiant phenol removal by volatilisation occuned. There
was no signifiant decrease in the concentration of phenois in control samples for the cases
of phenol, 3-chloropheM>i, and 4-chlorophenoi; however, the concentration of 2-
chlorophenol and 2,~chlorophenol decreased significantly due to vaporisation a h one
day of incubation Therefore, the reaction vials were seaieci with screw caps when the
incubation time was longer than 3 houn or chlorinated phenols were treated- The ovgen
which was dissolveci in the solution and a d a b l e in the head space of the vials was
estimated to be sufncient to transform ail the phenols when the concentration was lower
than 1 m M
After the addition of the appropriate amount of enzyme solution, the reaction
solution became coloured after a lag period and then gradually darkened over the. ïhere
was no precipitate observed even after overnight incubation. The intensity of the colour of
treated solutions seemed to depend on the initial phenol concentration. The treated phenol
solutions absorbeci a broad range of UV-visible Light. This colour generation was mooitored
at 5 1 O nm throughout this study because the measurement of this wavelength was necessas,
for the colour compensation of the phenol assay. The colour appeared not to change with
pH or bmer used within the range of the enzyme doses examind
As reported in the literatue (Nakamoto and Machida, 1992; Ganjidoust et al.,
1996), some additives such as polyethylene glycol (PEG) and chitosan protect horseradish
peroxidase and other peroxidases nom inactivation and thereby improve the removal
efficiency of the phenolic substrates. In order to identify whether a similar e f k t occurs in
the case of tyrosinase, the transformation of phenol in the presence of PEG and chitosan
was investigateà
Since phewlic solutions treated with tyrosinase result in the formaton of caloured
compounds that may contribute to toxicity and must be removeci, extensive experiments
were conducted to convert soluble products to insoluble precipitates using coagulants
(chitosan and d m ) as d i s c d in the later sections. Chitosan has also been reporteci to be
able to adsorb the products of phewl oxidation (Ganjidoust et al., 19%). Therefore,
chitosan was investigated fïrst as an additive (i.e. added before reaction initiation) and then
as coagulant/adsorbent (Le. added afkr the reaction was completed).
4.2.1 Effect of pH
The effect of pH on the t r ans fodon of phenol with tyrosinase was investigated
in the pH range between 3 to 10 with four different doses of tyrosinase. The types of pH
buffers used are listed in Table 3.1. Figure 4.7 presents (a) the fransformation of phenol and
@) the intensity of generated colour (messured by absorbance at SlOnm) as a niaction of
pH When limiting doses of enzyme were used, the profile of phenol transformation is
similar to the relative activity of tyrosinase at different pHs that was shown in Figure 4.1.
The optimum pH for phenol transformation with tyrosinase was detennined to be pH 7. A
broad optimum can be observed between pH 5 and 8. However, only minor transformatons
occurred at pH 4 and pH 9, and no phenol transformation was observed at lower and higher
pHs. As shown in Figure 4.8, the generated colour intensity for al1 pHs was proportional to
the transformed phenol (regression coefficient was 0.980).
Experiments were aiso perfonned involving reactions conducted in the presence of
100 mg/L of chitosan under the same conditions as descn'bed above. As shown in Figure
4.9, the shape of the cmes for phenol transformation changed Although no significant
change was observed under basic conditions, the transformation improved at pH 4 but
Figure 4.7 Effect of pH on the transformaton of phenol catalyseà by tyrosinase in the absence of chitosan: (a) phenol transformation, (b) colour generated at 5 10 nm ([phenolJo = 0.5 mM, 3 hours of reaction at 25OC).
A ,, = a ~(Phenol Transfomed)
a = 1.67 A U * ~ M " 8 = 0.980
0.0 0.1 0.2 0.3 0.4 0.5
Phenol Transfomed (mM)
Figure 4.8 Relationship between the colour generated st 5 10 nm and transfomecl phenol (each data were taken fiom Figure 4.7).
Figure 4.9 Effect of pH on the transformation of phenol catalyseci by tyrosinase in the presence of 420 cps chitosan: (a) phenol transformation, @) colour remaining at 510 nm afier centrifugation ([phenoll* = 0.5 mM, 3 hours of reaction at 25°C).
dropped a? pH 5. Dark-brown precipitates were obsemed at pHs W e e n 5 and 7 and
colour generation was d e p r e d No precipitation occuned at pH 4, and the colour of the
solution was ligbt brown Chitosan wuld not be fully dissolved in the basic solutions, thus
the colour removal did not occur effectively. Although the optimum pH was identifhi as
pH 6 for both colour depression and phenol transformation when the reactions were
conducted in the presence of chitosan, pH 7 sodium phosphate buf5er was used in al1 the
subsequent experiments for the sake of cornparison with reactions cmducted without
chi tosan.
4.2.2 Effect of Initial Pbenol Concentration
The relationsbip between phenol removal and enzyme dose was enamined
Aqueous phenol solutions with wtlcentrations ranging fiom 0.5 mM to 4 mM were treated
with a range of doses of tyrosinase and with and without chitosan. The transformation of
phenol as a function of enzyme dose is presented in Figure 4.10. Tyrosinase doses required
to transform 95% of initial phenol for each concentration were interpolated and plotted in
Figure 4.11. There was a linear relationship between initial phenol concentntion and the
tyrosinase dose required to achiwe 95% transformation The minimum dose of enzyme to
transfonn 95% of initial phenol content could be expresed as:
Tyrosinase dose = a x IphenolIo (4.5)
where a = 6.79 units/mLmM without chitosan and a = 6.40 units/mL-mM with chitosan, It
is clear that chitosan did not signifkantly improve the transformation of phenol catalysed
by tyrosinase. Based on these results, the minimum enzyme dose to achieve 95%
transformation of 0.5 mM phenol ( ie . the concentration used as a standard for this study)
was determined to be around 4 units/mL. In practice, 8 uniWrnL was usually used to
accomplish full treatment (i-e. maximum transformation) of 0.5 mM phenol in subsequent
experïments.
without Chitosan O withChitosan(1WmgL)
Tymsinase Dose (unitslmL)
wittiout Chitosan O with Chitotpn (200 mglL)
10 15 20 25
Tyrosinase Dore (unitstml)
Figure 4.10 Tyrosinase catalysed transformation of phenol with and without 420 cps chitosan: (a) IphenolIo = 0.5 mM, (b) [phenol],, = 1 mM (3 hours of reaction in pH 7 sodium phosphate buf5er at 2S°C).
.........................
95%
0 withoutChitosan O with Chitoron (400 rngfl)
Tyrosinase Dose (unblmL)
0 without Chitosan O with Chitosan (800 mgR)
1 rn I I I m
Tyrosinase Dose (unWmL)
Figure 4.10 (continued) Tyrosinase catalysed transformation of phenot with and without 420 cps chitosan: (c) [phewlJ0 = 2 mM, (d) [ p h e ~ > l ] ~ = 4 m M (3 houn of reaction in pH 7 sodium phosphate buffer at 25OC).
Tyrosinase Dose = a x phenoll,
with Chitosan a = 6.40 units/mL=mM
a = 6.79 un'WmL.mM
0 without Chitosan O with Chitosan
O 5 10 15 20 25 30
Tyrosinase Dose (uniorlmL)
Figure 4.11 Amount of tyrosinase required to transforrn 95 % of initial phenol (3 hours of reaction in pH 7 sodium phosphate buffer at 25°C).
4.23 Effeet of Snbsmte Type
The transformation and colour generation of five different 0.5 mM phenols with
tyrosinase were measured with time (Figure 4.12). The observed order of the rate of
transformation in the tested phenols was:
phenol r 4-chlorophenol> 3chlorophenol> 2-chlorophewl» 2,4-dichlorophenol
Phenoi, 34dorophenoi, and khlorophenol had been transformeci to nearly 100% witbin
10 houn with 48 u.nits/mL of tyrosinase (8 units/mL for phenol), but the rest of the
chiorinated phenols were transformed much more slowly than the former three compounds.
in partïcular, 2,4-dichlorophenol undenveat inwmplete transformation with only 38% of
its initial concentration king transformecl even after 1 &y with 48 units/mL of tyrosinare.
Probably, the enzyme was inactivateci by the products.
The colour of chlorinateci phenol solutions treated with tyrosinase varied with the
type of substrate. The colours observed in various phenol solutions treatexi with tyrosinase
are summarised in Table 4.2. The rate of colour development appeared to be closely related
to the rate of h;iosformation. When chitosan was initially added in the reaction solution,
precipitates formed with tirne. The colours of the precipitates are dso summarised in Table
4.2.
Table 4.2 Colour of the various pbenolic solutions treated with tyrosinase and the precipitates formed when chitosan was added before initiation of reaction
Compound Colour of treated Colour of the Precipitates solution (with chitosan)
Phenol Brown Dark brown 2-Chlorophenol BcoWLZish yellow Brown 3-Chlorophenol Light brown Brown 4-Chiorophenol Light brown Brown
2,4dichlorophenol Greenish yellow Dark greenish yellow
Phenol O 2-Chlorophenol v 3-Chlorophenol v CChlorophenol W 2,4=0ichlorophenol
Time (Hours)
Phenol O 2-Chlorophenol v 3Chlorophenol v 4Chlorophenol I 2,443ichlorop henol
10 15
Time (Hours)
Figure 4.12 Tyrosinase catalyseci treatment of aqueous phenolic compounds as a function of t h e : (a) phenol remaining, @) absorbance (Iphen~ls]~ = 0.5 mhd, tyrosinase dose = 8 uniWmL for phenol, 48 unitslmL for chlorinated phenols, in pH 7 sodium phosphate buffer at 25OC).
59
43.4 Effect of Polyethylene Glycol (PEG)
PEG was examined for its ability to improve the transformation of phenol
catalysed by tyrosinase. Six types of PEG which have different average molecular weights
were added to reaction solutions containing 0.5 mM phenol in pH 7 sodium phosphate
buf3er. A limiting quantity of tyrosinase was thni added to each solution and the mimim
were stirred for 3 hours. A sample without PEG was also prepared as a control. No visual
diEerence was obsewed between solutions contiu'ning PEG and the wntrol. The residual
phenol concentration and the absorbante at 510 nm of the supcrnatants werc m e a s d
after cenîrifiigation. There was no precipitate in any of the samples. As shown in Figure
4.13 (a), no significant improvement in phenol transformation was observed in the presence
of PEG.
4.2.5 Effect of Cbitosan
Chitosan was also examineci in the same manner as PEG. Six different viscosity
gracies of chitosan were assessed by adding them to the reaction mixtures prior to reaction
initiation. The adsorption of phenol on chitosan could be neglected (Sun et al., 1992). The
concentration range of chitosan selected was between O and 400 mg&,. Except for the 420
cps chitosan, the solubility limit of each chitosan was exceeded d e n 400 mg/L were used.
Therefore, a precipitate formed instantly in these reaction mixtures. Dark-brown flocs were
foxmed in most of the reaction solutions with tirne. The dark-brown colour still remaineci in
some solutions after 3 hours (see Figure 4.14@)). The colour and the form of the
precipitates appeared to be very similar to that obtained d e n chitcsan was added after the
reaction had occurred (see section 4.3). The absorbante at 510 nm and the residual phenol
concentration of supernatants were measured after centnfbgation As shown in Figure 4.14
(a) and (b), while the colour generation was depressed due to the presence of chitom no
significant improvement on the transformation of phenol was observed. It should also be
noted that Figure 4.14 @) shows that the addition of too much chitosan aFpears to stabilise
the presence of colour in the solutions. Therefore, it is necessary to minimise the amount of
0
Without PEG
Average Molecular Weight of PEG (glmol)
(b)
Without PEG
IOOOO 20000 30000
Average Molecular Weight of PEG (glmol)
Figure 4.13 Effect of PEG on tyrosinase cataiysed transformation of phenol: (a) phenol remaining, (b) absorbance remaining ([phen~l]~ = 0.5 mM, tyrosinase dose = 1 unit/mL, PEG dose = 400 mgk, 3 hours of reaction in pH 7 sodium phosphate b&er at 25OC).
Viscosity of Chitosan (cm) - .
(b) -............-..-.....-...~......*......*........-..-........ O 1Oû mglL
2WmglL 400 mgîL
-.-.. without Chitosan
10 100 420 930 2920 5706
Viscosity of Chitosan (cps)
Figure 4.14 Effect o f chitosan on tyrosinase catalysed transformation of phenol:
a phenol remaining, (b) absorbante remaining ([phen~l]~ = 0.5 mM, tyrosinase dose = 1 unit/ml, 3 houn of reaction in pH 7 sodium phosphate b u f k at 25OC).
chitosan used to treat caloured solutions in order to (1) minimise reagent wsts and (2)
achieve complete wlour rernoval.
4.3 Removal of the Colour Remainhg in Treated Solutioas
Colour removal fiom the phenol solutions treated with tyrosinaw was atternpted
using coagulants. As shown in Figure 4.15, the intensity of the dark-brown wlour was
proportional to the quantiîy of phenol transformeci for the fidl range of initial phenol
concentrations selected (0.5 m M - 10 mM). Since the absorbante at 5 10 nm was very high
(beyond the measurab1e range by spctrophotornetry), sample dilution was required before
the absorbance was measured. Tbus, the absorbance values in Figure 4.15 reflect the
correction of the measufed absorbance for the dilution of the samples.
4 . 1 Effect of AIum
At first, concentrated aluminium sulfate (A12(S04)344H20, Alum) solution was
used as a coagulant Six different doses ranging nom 10 to 320 mgL of alum were added
to the fùlly treated (2 98%) 0.5 mM phenol solutions st pH 7 and quickly shed with a
magnetic stirrer for several seconds. The stirring speed was slowed down and the solution
was incubated for 3 hours at room temperature. Very smaii particles were suspended in the
solution when 150 mgR. and 300 mg& of alum were added. AU samples were centrifugai
after 3 hours and the absorbance was measured. Small amounts of grey precipitates were
obtained after the centxifùgation The absorbance of the supernatant as a fiinction of alum
dose was presented in Figure 4.16. The absorbance declined slightly with increasing alum
dose, but no signifiant effect of alum on the colour was obsnved over the tested range.
Subsequently, a dose of 1000 mgL of alum was used Although more precipitates were
obtained after centrifiigation, the absorbance was d e c r d by ody about 20% (data not
show).
O 2 4 6 8 10 12
Phenol Transformed (mM)
Figure 4.15 Relationship between intensity of the colour generated at 510 nm by tyrosinase cataiysed phenoi oxidation and quantity of phenol transformed following the complete treatment (transfomation 2 98%) of the solutions with initial concentrations between 0.5 mM and 10 mM (3 hours of reaction in pH 7 sodium phosphate bufïer at 25°C).
Initial Absorbance
O 50 100 156 200 250 300
Alum Dose (mg1L)
Figure 4.16 Effect of dm on wlour removal at 510 nm fiom the M y treated (transformation 2 98%) phenol solutions ([phenolJo = 0.5 mM, tyrosinase dose = 8 unitdml, 3 hours reaction followed by 3 houn of incubation with various doses of alum in pII 7 sodium phosphate bufEer at 25OC).
4.3.2 Effect of Chitosrin
Colour removai by six dinerent viscosity grades of chitosan was assessed Limiting
amounts of chitosan were added to the M y treated (2 98%) phenol solutions at pH 7 and
incubated for 3 hours with gentle stining at room temperature. Dark brown flocs appeared
in each of the solutions immediately afkr the addition of chitosan AAcr 3 homg al1 the
solutions were centrifbged and the absorbance at 5 10 n m of each supernatant was measrued
There were srnall differences in the absorbance among the chitosan types (Figure 4.17).
Chitosan grades with between 10 and 420 cps viscusïty removeci the colour from the
coloured treated solution most effectively. However, the 420 cps chitosan was most easily
dissolved during the preparation of stock solutions. Therefore, this chitosan wu used in ail
subsequent experiments.
In order to determine the optimum chitosan dose, the colour removal fiom five
different wncentrations of phenol solutions treated to 2 98% at pH 7 were exaxnined.
Increasing amounts of chitosan were added to each of the treated solutions which were
incubated at room temperature with stirring- After 3 houn and 18 hours of incubation,
samples were taken fiom each solution and the absorbance of the supernatants at 5 10 nm
was measured following cenûifiigation. The results are show in Figure 4.18 for a
concentration range of 0.5 rnM to 10 mU In each case, the absorbance fkst decreased in
proportion to the amount of chitosan added When the a b s o h c e was reduced by nearly
90 % of the initial intensity, the cuwe became flat and started increasing. At âûy phenol
concentration tested, the absorbance was lower after 18 hours than after 3 hours, and the
absorbance difference was nearly constant for dl doses of chitosan The amount of chitosan
required to achieve 90% colour removai was estimated fiom the linear regression resdts
taken from early liwar portions of each curve (Figure 4.19). The amoimts of chitosan
required to achieve 90% colour removai logarithmicaily increased with initial phenol
concentration (Figure 4.20). The relatioaship can be expresseci as:
without Chitasan
Viscosity of Chitosan (cps)
Figure 4.17 Effect of chitosan type on the removal of colour at 510 nm fiom fully treated (transformation 2 98%) phenol solutions (Iphenoll0 = 0.5 mM, tyrosinase dose = 8 units/mL, 3 hours of reaction followed by 3 hours of incubation with 40 mg/L chitosan in pH 7 sodium phosphate buffer at 2S°C).
1.0 a u Initial Aborbance
3Hours O 18 Hou=
a 0.0 . I 1 7p
Chitosan Dose (mgJL)
3 Hours 18 Hours
90% Removal \
O 50 100 150 200 250 300
Chitosan Dose (mglL)
Figure 4.18 Removal of the wlour fiom the fully treated (transformation > 98%) phenol solutions by the addition of420 cps chitosan: (a) IphenolIo = 0.5 mM, tyrosinase dose = 6 units/mL, (b) [phenolIo = 1 mM, tyrosinase dose 12 units/mL (3 hours of teaction and 3 or 18 hours of additional incubation with chitosan in pH 7 sodium phosphate buf5er at 2S°C).
68
Chitosan Dose (mgIL)
Chitosan Dose (mglL)
Figure 4.18 (continued) Removal of the colour fiom the W y treated (transformation 1 98%) phenol solutions by the addition of 420 cps chitosan: (c) [phenolIo = 2 mM, tyrosinase dose = 24 uniWrnL, (d) Iphen~l]~ = 4 mM, tyrosinase dose 48 units/mL (3 houn of reaction and 3 or 18 hours of additional incubation with chitosan in pH 7 sodium phosphate buffer at 25OC). 69
90% Removal
O 100 200 300 400 m 600
Chitosan Dose (mgll)
Figure 4.18 (continued) Removal of the colour fiom the fully treated (transformation 2 98%) phenol solutions by the addition of 420 cps chitosan: (e) [phenoll0 = 10.0 mM, tyrosinase dose = 96 units/rnL (3 hours of r d o n and 3 or 18 houn of additional incubation with chitosan in pH 7 sodium phosphate buffer at 2S°C).
Chitosan Dose (mg/L)
O 100 200 300 400 500
Chitosan Dose (mgll)
Figure 4.19 Linear regressions of the linear portion of c w e s for each initial phenol
a concentration: (a) 3 hours incubation tirne; (b) 18 hours incubation time.
Requind Chitosan Dore = a x log [Phenoll, + b (b)
Figure 4.20 Amount of chitosan required to achieve 90% colour removal fkom the Mly
a treated pheno! solutions: (a) linear plot, (b) semi-log plot (3 hours of reaction and 3 or 18 houn of additionai incubation with 420 cps chitosan in pH 7 sodium phosphate bufEer at 25°C).
72
Required chitosan dose = a x log IphenolJo + b (4-6)
The constants were calculated from the linear regession resdts as: a = 368 mg/L, b = 162
mg& for 3 hours of incubation and a = 353 mg& b = 156 mg/L for 18 hours of incubation.
The colour was not removed by the addition of dum, which is a major coaguiant
used in water treatment, but was removed by the addition of chitosm Therefore, this
colour removai is not likely only due to coagulation but also adsorption It is proposed that
the coloured products chemicaiiy bind to the dissolved chitosan molecules (chemisorption)
and aiter the characteristics and the solubility of chitosan. The result is the formation of
solid precipitates which flocculate and settie. Thus, the adsorption behaviour of chitosan
and the coloufed produts were examiined using the data show in Figure 4.18. It was
assumed that equilibrium was achieved after 3 hours. 18 hours data was not used since it is
observed that colour removal is also bccurring due to a chitosau independent process (Le.
Figure 4.18 shows that samples with no chitosan lost the same amouat of colour with time
as samples with chitosan).
Two adsorption isotherm models were used: the Langmuir model and the
Freundlich model. The Langmuir isotherm model is based on the assumption that
molecules are adsorbeci on definite sites on the surface of the adsorbent (Benefield et al.,
1982). The equilibnum isothexm equation is:
where x = amount of matenal adsorbed (AVL, in this case), m = mass of adsorbent (mg), C
= concentration of material remaining in solutions after adsorption is complete (AU), and a
(AU') and b (AU-Umg) are constants. Two linearised forms of equation 4.7 can be
conStNcted as follows:
1 I l -=-+- x l m b abc
These equations are called linearised Langmuir isotherms #1 and #2 in this thesis.
The Freundlich isotherm mode1 is based on an empirid equation that allows for the
heterogeneity of the srirface and the exponential distribution of sites and their energia
(Benefield et al,, 1 982). The equii~'brium isotherm equation is:
where x = amount of soiute acisorbed (AU-L), m = mass of adsorbent (mg), C =
concentration of solute remaining in solution &er adsorption is cornpiete (AU), and K
(AU-L/mg) and n (dimensionless) are constants that must be evaluated for each solute and
temperature. A linearised fonn of equation 4.10 was can be mnstnicted as follows:
Figures 4.21 and 4.22 present the experimental data and linear regressions
expressed in these three linearised foms. Al1 data points presented in Figure 4.18 were
used to evaluate the adsorption isotherm except for some points where stabilisation was
observed in Figure 4.18 (a) and (b). The calculated coefficients detemhed by linear
regressions are shown on each figure. It was found that the coefficients of two linearised
Figure 4.21 Linearised Langmuir isotherxus: (a) linearised form #1, @) linearised fonn
log (dm) = (1ln)-log C + log K
9 . K = 1.04~10' AU-Urn~. n = 4.792 . I
log C
Figure 4.22 Linearised Fremdlich isothenn.
Langmuir isotherrns were subsîantiaily dinaent even these iinearised equations were
denved fiom the same equation.
The fit of the modeiled isothenns and the experimental data is shown in Figure
4.23. The Langmuir isotherm drawn with the coefficients estimated nom linearised
Langmuir isotherm #1 deviated so much fiom the experimental data, that it is not sbown.
Although the regression result of linearised Langmuir isotherm #2 (Figure 4.2 1 (b)) met the
experimental data relatively well (8 = 0.89), the overlay of the modei resula on the data
showed substantial deviations (Figure 4.23)
4.4 Toxicity of the Treated Phenol Solutions with Tymsinase
A senes of Microtox acute toxicity assays were carried out to determine the
toxicity of phenol solutions treated with tyrosinase. Four different concentrations of phenol
solutions were treated at room temperature in pH 7 sodium phosphate buf5er. The toxicities
of these solutions were tested after 3 hours when completion of the transformation was
confirmed (1 98%), and also after 24 hours of incubation at room temperature. The
toxicity of phenol was also tested and determined to be 12.4 mg/L (toxicity expressed as an
EC50). Al1 samples were centrifuged before the toxicity tests. The toxicities of the initial
phenol and treated solutions are presented in Figure 4.24. The toxicities of al1 the treated
phenol solutions were lower than the initial phenol toxicity. In addition, the toxicities were
substantialiy decreased afîer 24 hours of incubation compare- to after 3 hours. The
toxicities of the solutions containing chitosan were much lower than those without chitosan
for both 3 and 24 hours of incubation. Previous experiments conducted in this laboratory
(data not shown) confirmed that the chitoran did not contribute to the toxicity of phenol
solutions.
The toxicities of the treated phenol solutions were plotted versus colour intensity
at 5 10 nm in Figure 4.25. While colour and toxicity are correlateci with each other for
Figure 4.23 Cornparison b e n modelled adsorption isotherms and experimental data of colour removal fiom the treated phenol solution by the addition of chitosaa.
0.018 - 0.016 -
n p 0.014 - I
m . . . . . -- F. . - -
.*-• -.---m- . . - - ...-
. - - _ _ - - - . . . - -
a 0.5 mM V O 1mM
2mM v 4mM
1OmM - Langmuir #2 0.002 - ..... Freundtich
Initial Phenol
25 - O 3 Houm without Chitoaan r 3 Hours with C h i t a n v 18 Hours without Chitosan
20 - 18 Hours with Chitosan
10 -
Fignre 4.24 Toxicities of the various concentration of Mly treated phenol solutions (transformation 2 98%) by tyrosinase with and without 420 cps chitosan in pH 7 sodium phosphate buffer at 25°C (chitosau dose = 100 mg& for 0.5 mM phenol solutions, 200 mgL for 1 m M phenol solutions, 400 mg& for 2 mM phenol solutions, 800 mg/L for 4 mhl phenot solutions).
+ 3 hours without chitosan - + 3 hours with chitosan 18 hours without chitosan
a
O 1 2 3 4 5
Absorbante at 5f0 nm (AU)
Figure 4.25 Relationship between toxicity and colour of the treated phenol solutions.
different experiments, there was no correlation between colour and toxicity for al1
experiments. This implies that the c o l o d proàucts are not necessarily the source of
toxicity; rather, toxic products and colour are both removed by chitosan.
4.4.1 Efféct of Chitosan
The effect of chitosan addition on the toxicity was investigatd Chitosan did not
contribute to the solution toxicity. 0.5 mM phenol solutions in pH 7 sodium phosphate
buffer were prepared and r d o m were carrieci out at room temperatrire. Various doses of
chitosan were added to the phenol solutions followed by the enzyme to initiate the reaction.
The reaction was c-ed out at room temperature for 3 hours. The toxicities of the treated
solutions were tested afkr centrifbgation. The colour and the toxicity as a function of
chitosan dose are presented in Figure 4.26. The toxicities of the treated solutions with
chitosan were always lower than that of those without chitosan. But the estimated toxicities
were Iow (2.e. TU50 < 1) in al1 the treated solutions, therefore, the effect of chitosan on the
toxicity could not be adequately qwtif ied
The effect of chitosan when it was added after the reaction was aiso investigated.
Fully treated (2 98% transformation) 0-5 m M phenol solutions in pH 7 sodium phosphate
buffer were prepared. Various doses of chitosan were added to the dark-brown treated
solutions and incubated for 3 hours with gentle stirring at room temperature. The toxicities
of the samples were tested after centrifugation The toxicity and the colour of the samples
are shown in Figure 4.27. The toxiciîies of ail the solutions were lower than the toxicities
tested before the chitosan addition. While the eEect on colour removal was significant,
there was no significant effect of chitosan addition on toxicity since the toxicities of ail
samples were very low.
4.4.2 Toxicity of the Treated Chlorinated Phenol Solutions
The toxicities of the chlorophenol soiutions treated with îyrosinase were aiso
Toxicity Colour
50 100 150
C h b a n Dose (mgIl)
Figure 4.26 Effect of chitosan (added prior to reacîion initiation) on the toxicity and colour of fûlly treated phenol solutions (transformation 2 98%) by tyrosinase (Iphen~l]~ = 0.5 mM, tyrosinase dose = 8 units/rnL, 3 hours of reaction with a range of doses of 420 cps chitosan in pH 7 sodium phosphate b a e r at 25OC).
\ Toxicity of Untreated Phenol
Toxicity before Chitosan Addition \
Chitosan Dose (mgfL)
Figure 4.27 Effect of chitosan (added after the reaction was completed) on the toxicity and colour of fully treated phenol soluîions (transformation 2 98%) by tyrosinase ([phenolIo = 0.5 mM, tyrosinase dose = 8 uniWml, 3 hours of reaction and 3 houn of additional incubation with a range of doses of 420 cps chitosan in pH 7 sodium phosphate b-er at 25°C).
investigated, 0.5 mM 2-chlorophenol, 3cbloropbenol,4-chiorophenol, and 2,4-dicholoro-
phenol were prepared in pH 7 sodium phosphate b a e r and treated with 64 units/mL of
tyrosioase in the presence and absence of chitospn at rmm temperature. The completion of
the transformation of chlorophenols were c o b e d after 1 day of incubation except for
2,4-dichlorophenol solution without chitosan (1 1% transformation). Haif of the volume of
each solution were centnfuged and anaiysed for toxicity. The r a t of the samples w m
incubated for another day, and then their toxicities were assesseci The transformation of
2,4dichlorophenol was stiil only 38% after 2 days. The d t s are pmsented in Figure 4.28.
The initial toxicities of the chlorophenols expressed in EC50 (mg@) are shown in Table
4.3.
Table 4.3 Initial tolricity of phenol and chtorophenols in ECSO (m*).
Compound This study Ghioureliotis (1997)- Phenol 12.4 11.0
The decreasing order of the initial toxicity for the phenols tested including phenol
4-chlorophenol> 2,4-dichlorophenol ~ 3-chlorophenol> phenol s 2-chlorophenol
The initial toxicities of these compounds were consistent with those previously reported by
our laboratory (Ghioureliotis, 1997).
The toxicities of all the treated chlorophenot solutions were substantiaily lower
than their correspondhg initial toxicities except for the case of 2,4-dichlorophenol. This
0 probably occurred because the transformation of this compound was not completed
1 day without Chitosan
2 days with Chitwan
O
5 +
3-CP 2,4-DCP
Figure 4.U): Toxicities of the various chiorophem1 solutions aated by tyrosinase in the presence and absence of chitosan in pH 7 sodium phosphate briner at 2S°C ([chlorophenolJo = 0.5 mM, tyrosinase dose = 64 units/mL, chitosan dose = 100 mg/L).
Note: Transformation rates of 2,4-dichlorophenol were 1 1% after 1 day and 38% afler 2 days. The other chlorophenols were fitlly treated (transfonaation 2 98%).
Chitosan had irnproved the removal of 2,4-dichiorophenol very efféctively (1 98% removaî
after 1 day), and wnsequently the toxicity of the solution diminished However, it is wt
clear if the chitosan contributeci to the removal of 2,4-diclorophenol through adsorption or
through enhanced transformation by tyrosinase. In addition, the toxkities of the other
chloruiated phenols treated in the presence of chitosaa were also significantly diminished
after 1 &y of incubation (i-e. TU50 < 1).
5 DISCUSSION
5.1 Characterbation of Tyrosinuc Activity
The tyrosinase activity measured in this midy ushg Ltyrosine as a substrate was
monophenolase activity which governs the monophenolase cycle describeci in Figure 2.2.
The pH optimum determined in this mdy was Merent from those reporteci for some
tyrosinases obtained fkom other sources (Espin et al., 1997a; 199%) but agreed with the
enryme supplier's products report (Worthington Enzyme Manual, 1977). This differene
may be due to the nature of the source of enzyme, the substrate used, the type of buffer, and
the purity of the enzyme (Robert et al., 1995). Even though the reported optima were
ranging fiom 5 (Espin et d., 199%) to 7 (this study, see Figure 4.1), al1 resulîs indicated
that tyrosinase is not active under basic conditions.
Tyrosinase appeared to be reasonably stable in neutral b a e r solution at room
temperature, however it is quite unstable at high (> pH 8) or low (< pH 5) pHs and at
higher temperatures (see Figures 4.2 and 4.3). The stability of tyrosinase in basic b a e r
solutions was higher than in acidic ones, wfrereas the pH dependence of îyrosinase activity
(see Figure 4.1) showed that the e q m e was more active in acidic buffers than basic
bufEers. Therefore, tyrosinase is relatively stable under weakly alkaline conditions but is
not catalytically active.
The temperature dependence of tyrosinase activity has been reported previously
for several tyrosinases nom other plants. The value obtained for the activation energy in
this study (1 -85 kJ mol-') was comparable to the value for palmito (Acanthophoenit mbra)
polyphenol oxidase using 4methylcatechol as a substrate (5.41 W mol-') (Robert et
a1.,1995); but much lower than the other values reported in the same article for potato
polyphenol oxidase using pyrogallol as a s u b t e (54.5 kJ mol-') and for banana
polyphenol oxïdase using catechol as a substrate (18.6 W mol"). The lower activation
a energy for inactivation implies a lower thermal stability of the enzyxne used in this study.
Although there was no literaîure reference available to compare results for mushroom
tyrosinase, the sources and the purity of the enzyme as weil as the substrates used may
innuence the activation energy for inactivation.
A cornparison of the decimal reduction value, D, between tyrosinase and soybean
peroxidase which was investigated earlier in our laboratory (for the purpose of wastewater
treatment) shows tyrosinase was much more unstable than soybean peroxidase (Table 5.1).
The decimal reduction value for tyrosinase at 50°C at pH 7 was 10000-fold lower îban that
of soybean peroxidase at 50°C (approximately 4 x 103 estimated fiom the dezimal
reduction value for soybean peroxidase at 70°C and 2-value at pH 7. The activation energy
also indicated a much lower stability of tyrosinase than soybean peroxidase. Interestingly,
the 2-vaiue, which represents the susceptibility of the rate of entyme inactivaîion to
temperature change, of mushroom tyrosinase was quite similar to that of soybean
peroxidase. The lower stability of the enzyme represents a drawback to the use of
tyrosinase for actual wastewater treatment not ody during the reaction but also during the
preparation and storage of stock solutions.
Table 5.1 Cornparison of thermal inactivation parameters between soybean peroxidase and mushroom tyrosinase.
Enzyme pH Temperature D E4 Z Source (OC) (min.) (kJ mol-') (OC)
Soybean 7.0 70.2 3974 246 9.71 Wright (1995) Peroxidase 80.3 300
90.8 30 Mushroorn 7.0 10.0 25800 1.85 10.4 This Work Tyrosinase 30.0 2100
50.0 38.4
5.2 Tyrosinase Catrlysed Tnnsformation of Pbenol
In conaast to the report of Atlow et ai. (1984), the transformation of phenol
catalysed by tyrosiaase resuited in no precipitation in this study when no additives (cg.
chitosan) were added Wada et ai. (1994) also muntered the same situation and
explained that precipitation might be causeci by a Iowa pur@ enzyme. However the
tyrosinase used in this study was 500 unitdmg and wu much less pure than both enzymes
used by Atlow et al. (2000 unitdmg) and Wada et al. (3500 UIUtdrng)). The activity assays
used in al1 of these studies were the same; however, the tyrosinase sarnpla were obtained
fiom a different company (Sigma) than that used in this study (Worthington Biochemical
Company). The concentrations of bu&r, which may have an influence on precipitation,
were the wune in al1 of these d e s (50 mM sodium phosphate buffer). It is known that the
precipitation of synthetic lignin fiom coniferyl, coumaryl or sinapyl alcohols (phenols with
3-C sidechah) using laccases or peroxidases was influenced by the enzyme: substrate ratio
(Kondo et al., 1990). Since the aqueous concentration of oqgen, which is also one of the
subsîrates for tyrosinase, was not controiled, the ratio of tyrosinase: oxygen was udcnown.
It might be possible to alter the nature of products and induce their precipitation when more
oxygen had k e n introduced in the solution during the reaction.
The optimum pH for phenol treatment was determined to be between pH 5 and pH
8. Solutions containing between 0.5 m M and 10 mM phenol were successfully treated
within 3 hours (transformation 1 98%) with tyrosinase. In addition, several 0.5 mM
solutions of chlorinateci phenols were successfully treated However, experiments with 2,4-
dichlorophenol demonstrateci that this cornpouad is not a good substrate for tyrosinase. In
addition, preliminary experiments with pentachlorophenol showed that this wmpound 1s
not oxidise in the presence of tyrosinase. Therefore, while tyrosinase can be applied to treat
a variety of phenolic pollutants, it is not applicable to al1 phenols.
The minimum dose of tyrosinase required to transform 95% of initial phenol was
detennined to be 6.79 UIÙts/mL pet 1 m . of phenol in the absence of chitosan (see Figure
4.1 1). In this work, 6 U12itslrnL of tyrosinase was used to achieve complete tr-ent
(transformation 2 98%) of 0.5 mM phenol. Although the reaction conditions were the same
including pH, the concentration and the type of buffer, temperature and the enzyme activity
assay, this amount was much smaller than those reporteci by other researchers: 60 units/mL
(Atlow et al., 1984) and 20 units/mL (Wada et al., 1993). It may k due to the higher
protein content derived fiom less pure enzyme used in this study because neucleophilic
amino acid residues on proteins c a . act as reducers to promote the monophenolw activity
of tyrosinase (see section 2.2.4.5). The application of the lower purity eozyme to
wastewater treatment may be advantageous due to its improved performance and lower
cost.
The observed low transformation of Zchlorophenol and 2,4-dichlorophenol may
be explaineci by the steric hindrance of o-substituted chiorine on these substrates toward the
active site of the enzyme because tyrosinase catalyses the o-monohydroxylation of phenols.
This hypothesis is also supporteci by the extremely low reactivity of pentachlorophenol
against tyrosinase-caîalysed oxidation (Aitken et al. 1994). An undentandhg of the
substituent effects on oxidation by tyrosinase may be critical for the dwelopment of
specific treatment processes for substituted phenols. Additionally, the molar ratio of the
enzyme and the substrates including oxygen aad phenolic compounds is considerd to be
significantly important in that it will influence the rate of reaction More extensive
researches should be carried out to examine the reaction kinetics of tyrosinase-catalysed
oxidation of phenols.
Nakamoto and Machida (1992) reported that honeradish peroxidase could be
protected by the addition of PEG. The authors proposed that the hydrogen bonding sites of
PEG could interact with the hydroxyl groups of the polyrncrwd phenols. This interaction
may minimise the enwpment and the subsequent inactivation of peroxidase enryme by the
rapidly formed precipitating polymen (Buchanan and Nicell, 1998). However, the
transformation of phenol with tyrosinase was not affecteci by the addition of PEG. This is
probably because the phenolic polymm generated by self-polymerisation of quinones
were of low rnolecular mass and did not effêctively interact with the PEG molecules. The
hi& solubility of the polymerised products might result in less entrapment of the enzyme
by the polymer precipitates (comparai to peroxidase) and help to retain the activity of
enzyme in the solution. Moreowr, there is no effective affinity between PEG and O-
quinone which is coasidered as a major scavenger of tyrosinase activity (Garcia-Canovas,
1987).
Although the addition of chitosan was very effective in inducing the precipitation
of the products of tyrosinase-catalysed phenol treatment, it did not improve the
transformation of phenol in the tested range of initial phenol concentrations. Sun et al.
(1992) and Wada et al. (1993) reporteci that chitosan could reduce tyrosinase inactivation
when 4-methylphenol @-cresol) was used as a substrate. Sun et al. (1992) also showed that
chitosan did not limit tyrosinase inactivation d e n catechol was used as a substrate. They
suggested that this discrepcy was due to the diffmnce between o-diphenol and
rnonophenol (Sun et al., 1992). However, in our study, no signifiant improvement in the
transformation was observed for the case of a monophenol. Therefore, their hypothesis
seems to be incorrect. Sun et al. (1992) m e a s d dissolved oxygen over the course of the
reaction and interpreted an increase in the rate of disappearance of oxygen as an increase in
catalytic activity. However, they negiected to measure the phenolic substrate concentration
over time and at the end of the reaaion period Therefore, they did not actually prove their
hypothesis conceming the protective effect of chitosan on the transformation of phenolic
substrates. Based on the results of the curent study (see Figure 4.1 l), it can be concluded
that the effect of chitosan is limited to the interaction with soluble products arising h m
the oxidation of phenolic substrates (see discussion below).
53 Colour Removal from the Treated Phenol Solutions
The wlour generated by phenol transfomition with tyrosinase has been
successfully removed by the addition of chitosam Chitosan couid be added to the solution
both before and f i e r the reaction. In both cases, the colour was removed through the
formation of dark-brown precipitates. This p i p i t a t e formation was proposeâ to be due to
chernical reactions between oquinone and the MI2- groups on the chitosan molecules (see
Figure 2.8) resulting in the formation of covalent bonds (Sun et al., 1992). The effect of
chitosan type shown in Figures 4.14 and 4.17 suggested that the him viscous chitosan
(5700 cps), which is the least deacetylated of ail chitosan samples used in this snidy, has
the worst colour removal ability because of the limited number of -NH2 groups in the
molecules. In addition, the solubility of chitosan increased with decreasing viscosity.
In colour removal experiments conducted with a range of doses of various
chitosan, there was a tendency for precipitates to form immediately after high doses of
chitosan were added to reaction solutions, except in the case of 420 cps chitosan This
precipitation appeared to reduce the efficiency of colour removal (see Figure 4.14). This
would create problems when it is necessary to remove large quantities of colour resulting
from the transformation of large quantities of phenol (see Figure 4.15). Chitosan grades
between 10 and 420 cps were equdy effective in removing colour as long as excess doses
were not used (see Figures 4.14 and 4.17). Therefore, the 420 cps chitosan was chosen
because it did not spontaneously precipitate when added to reaction mixtures and was very
effective in removing colour from a large range of treated solutions (see Figures 4.19 and
4.20). In addition, the 420 cps chitosan was easily dissolved in the stock solution of acetic
acid (see section 3.1.1) in cornparison to other viscosity grades of chitosan Thus, the 420
cps chitosan was the subject of al1 M e r colour removal studies.
There was a logarithmic relationship between treated phenol concentration and
required chitosan dose to achieve 90% absorbame removal at 510 nm fiom the solutions
when chitosan was added &er the reaction (equation 4.6). W h et al. (1995) reported that
approximately 40 mg/L was required to remove over 90% of the colour fiom a 0.5 m M
phenol solution treated by tyrosinase. This result is consistent with the 50 m@
requirement predicted by equation 4.6. Wada er ai- also studied the use of synthesised
polymer coagulants with amino groups and achieved higher colour removal efficiencia;
however synthesised coagulants are expensive compand to chitosan and may not be
commercially feasible for the treatment of wastewater.
Wada et ai. (1995) concluded that the addition of chitosan to the reaction solution
before the initiation of reaction was not appropriate because too much chitosan (1400
m a ) was required to completely remove the colour generated fiom the treatment of 0.5
mM phenol (Wada et al., 1993). In this study, the chitosan dose required to depress the
colour generation by up to 90% was about 200 m@L under the same reaction conditions
(see Figure 4.26). This resdt was much lower than the previous report; however, it was still
higher than that required for the chitosan addition after the reaction was completed. It is
probably due to the dinmnce in the affinity of the adsorbed products for the chitosan
molecules. When the transformation of phenol is completed, there may be several types of
oligomen dissolved in the solution. nese oligomers are considered to be more reactive
with chitosan than quinone because they have many Mctionai groups which may interact
with chitosan an4 consequently, the colour is removed more effdvely. The addition of
chitosan before the initiation of reaction may not be a good method in terms of the costs of
chitoçan, but it would reduce the number of treatment steps because chitosan and enzyme
can be introduced into the reactor at the sarne time.
Since coagulation with aluminium sulfate (alum) fded (see Figure 4.16), the
colour removal induced by the addition of chitosan appeared not to be the result of purely
~oa~guiation but probably also adsorption Thus, the adsorption of the coloured products on
chitosan was investigated. It appeared to be very cornplex Although there was a relatively
good agreement between the linear tegression and the experimental data in the lïnearid
Langmuir isotherm #2 shown in Figure 4.21 (b), the modelled isotherms and the
experimental data did not fit very well when they were compareci in Figure 4.23. The
failure of the data to fit on a siagie isothem (see Figure 4.23) is probably the result of
multiple mechanisms of colour removal. a t o s a n can contribute to the colour rernoval
initially through a chemisorption process involving the covalent ôonding of coloured
products with the chitosan (Sun et al., 1992). This appeared to result in the formation of
precipitated products. Chitosan, which is a lmown coagulant (Wada et ai., 1995), can then
act to enhance the destabilisation and flocculation of solid particles as they form.
It should aiso be noted that cblour removd also appuus to occw by a third and
very slow mechanism which is independent of the presence of chitosan. The absorbame
was monitored at two time intemals (Le. 3 hours and 18 hours) following chitosan addition
at the end of the reaction (see Figure 4.2 8). ï h e absorbame measured after 18 hours was
ahvays srnailer than the absorbance measured earlier even for the case in which no chifosan
was added. This may be due to some urhown degradation p r e s s (e.g. enzymatic or
spontaneous redox reaction) of the products. This colour removal mechanism is sufficiently
low (4.5% - 6.3% of initial absorbance was removed in 15 hours) that it is unlikely to be of
practid importance for waste treatment applications.
5.4 Toxicity of Pheaol Solutions Treated with Tyrosinase
The toxicities of the phenol solutions treated with tyrosinase were substantially
lower than the toxicities of the initial solutions. This suggested that the oxidised products
such as oquinones and their oligomen were less toxic than their parent molecules. Longer
incubation times might allow the o-quinones to oligomerise and result in further
detoxification (see Figum 4.24 and 4.28). Further characterisation of the oxidised products
is required to ver@ this hypothesis.
Chitosan addition accomplished the detoxification of the treated phenol s01utioas
especidly for the cases d e n higher initial phenol concentrations were d (see Figure
4.24). Chitosan can rract with uquinones at a rate much faster than the spontaneous
oligomerisation of vuinones (Sun er al., 1992). Therefore, detoxification of the solutions
containing chitosan was much faster than solutions without chitosan Chitosan seems to
have removed both the colour and the toxic products at the same tirne. But there was no
definitive relationship between the quantity of residual d o u r and toxicity (see Figure
4.25).
A cornparison between the toxicities of treated solutions evaluated in this study
and previously published values is presented in Table 5.2. Ail results obtained in this study
were slightly lower than those reported by Aitken et ai. (1994), even though they treated
iower initial concentrations of phenol. Howevez, it should be noted tbat the results shown
in Figure 4.28 demonstrate that the toxicities of treated phenolic solutions decrease with
time. Aitken et al- (1994) did not mention how long they waited before measuring the
toxicity of treated solutions. Therefore, it is possible that the higher toxicities reported by
Aitken et al. (1994) compared to this current study arise fiom a difference in the t h e
between treatment and toxicity d y s i s .
A cornparison of the toxicities of the treated phenol solutions resulting fiom the
use of different enzymes is shown in Table 5.3. Phenol solutions treated with tyrosinase
had the lowest toxicity of al1 solutions even in the absence of chitosan. This represents a
very strong advantage to the application of tyrosinase for the treatment of phenolic wastes.
- Table 5.2 Comparison of toxicities (in TUSO) of treated phenol solutions in this study
with pubiîshed &a
WA = not available
Compound
Phenol 2-Chlorophenol 3-Chlorophenol 4-Chiorophenol
2,4-Dichlorophmol
(1) Initial toxicities of the 0.5 mM phenois. (2) Initiai conceniration of each phenolic compound was 0.5 mM. Thc incubation time was 1 day except for phenol(3 hours). (3) Same condition as in (2) but 100 mg/L of chitosan was iMtiaUy rdded to the reaction soluboa (4) Initial concentradion of w h phenolic compound was 0.1 mM. The incubarion the was not indicated. (5) 1 1 % transformation of 2,4-dichlorophenol was accomplished without chitosan but complete removal of this substrate %as achicved m the piesence of chitosan,
Table 5.3 Comparison of toxicities (in Tü50) of the treated phenol solutions using
- Initiai toxicity "' This study
without chitosan with câitosan 3.96 1 .O2 0.08 5.79 5.03 O. 10
24.16 1.14 O. 17 86.57 O.% O. 13 59.1 1 59.5 0.06
di fferent
Aitken et al. ( 1994) (4)
1.75 5.88 N/A 1.58 N/A
enzymes.
[phen~l ]~ Enzyme pH Initial Final Source Toxicity Torricity
1mM Tyrosinase 7 8.22 6.7 This work 1 mM Tyrosinase with chitosan 7 8.22 1.3 This work 1mM Soybean peroxidase 7 8.75 19.5 Ghioureliotis(1997) 1 mM Horseradish peroxidase 7 8.75 19.5 Ghioureliotis (1997)
O. 1 mM Honeradish peroxidase 7 0.5 6.7 Aiîken er al. (1 994) O. 1 mM Chloroperoxidase 7 0.5 13 Aitken et al. (1994) 0.1 mM Lignin peroxidase 4 N/A 5 -3 Aitken er al. (1994)
NIA = not available
6 CONCLUSIONS AND RECOMMENDATIONS
This study was undertaken in order to characterise tyrosinase in terms of its
activity, stability, and potential for the treaîment of aqueous phenols. Two types of
additives, PEG and chitosan, were examined to imprbve the transformation of phenol.
Chitosan and alum were also examined to assess their ability to remove the wlour
generated by the reaction- Due to concerns over the quality of the treated phenol solutions,
acute toxicity tests of phenol solutions were dso conducted
Tyrosinase activity bas been characterised using L-tyrosine as a substrate. The
maximum catalytic activity was observed at pH 7; however, significant activity was
observed at pHs m g h g h m 5 to 8. The stability of tyrosinase at different pHs and at
different temperatures was measured T y r o s i appeared to be unstable at low pH and at
elevated temperature. The activation energy for thermal inactivation of tyrosinase was
calculated to be 1.85 kJ mol-'. It is suggested that tyrosinase is quite unstable compared to
peroxidase enzymes which have aiso been examinai for theV potential application to the
treatment of wastewaters.
The transformation of phenol with tyrosinase was ais0 investigated The optimum
pH for phenol treatment was between pH 5 and pH 8. Tyrosinase was able to transform
phenol over a wide range of initial phewl concentrations (0.5 mM - 10 mM), but no
precipitates were observed Monochlorinated phenols were also successfûily transformeci
with tyrosinase; however 2,4dichlorophenol showed less reactivity to the oxidation
catalysed by tyrosinase. The ortho-substituted chlorine may inhibit the interaction of
substrates with the active site of the enzyme,
The minimum dose of enzyme to transform 95% of 1 mM phenol was determined
to be 6.79 units/mL (when treatment was conducted without chitosan) and 6.40 units/mL
(with chitosan). Neither PEG nor chitosan showed a positive effect on the transformation of
phenol. However, the addition of chitosan was very effective in inducing the precipitation
of the coloured products generated by the transformation of phmol with tyrosinase.
Although chitosan could be added to the solution both before the initiation of the reaction
and after the transformation was completed, the required dose to accomplish 90% colour
removal was 3-fold higher when chitosan was added initially thau when chitosan was added
later. This may suggest that the growing oligomers of quinone interact with chitosan
more strongly than o-quinone.
Since coagulation of the coloured products with aluminium sulfate failed, the
precipitation induced by the addition of chitosan appeared not to be the result of purely
coagulation but probably also adsorption. The adsorption of products on chitosan was
modelled using the Langmuir and the Freundlich isothemis. Although there was a relatively
good agreement between the linear regression and experimental &ta for an heafised foxm
of the Langmuir isotherm, the modelled isotherms and the experimental &ta did not fit
very well when they were presented in non-linear f o m . It is suggested that the removal of
coloured products were govemed by the combination of the several phenornena such as
adsorption and coagulation.
The acute toxicity of treated phenol solutions with tyrosinase was investigated
using the Microtox Assay. Al1 the treated phenolic solutions with tyrosinase showed Iower
toxicity than the conesponding untreated solutions. The addition of chitosan enhanced the
detoxification of phenols induced by tyrosinase very effectively. Compared to other
enzymes (i-e. peroxidases), the toxicities of the phenol solutions treated with tyrosinase
were very low. This represents a very strong advantage when considering this enyme for
applications in wastewater treatment.
Based on the results of the w o k several areas have been identified for M e r
-
investigation. These include:
(1) the developmem of an assay technique that would be used for the measwememt of
enzyme activity during treatment in order to investigate the substrate-induced inactivation
of tyrosinase during phenol transfomation;
(2) the investigation of the effects of oxygen concentration on the reaction rate and the
nature of products, and optimisation of the molar ratio of the substrates and enryme for the
treatment of phenols with tyrosinase;
(3) the kinetic modelling of tyrosinase-cataiysed oxidation of phenols with respect to both
of the substrates: phenolic compounds and oxygen;
(4) the investigation of the effect of chernical structure of substrates including some
aromatic amines and poly-arornatic phenols on the protective effkct of chitosan on
tyrosinase;
(5) the characterisation and quantification of the products of phenol oxidation by
tyrosinase in order to undentand the removal mechanism of the products nom the solutions
and to remove the colour more efficiently;
(6) the investigation of the immobilisation of tyrosinase on chemically stable supports to
preserve the enzyme activity and encourage the reuse of enzyme;
(7) the investigation of the potential for the application of tyrosinase and chitosan for
industrial wastewater treatment;
(8) the exploration of alternative sources of tyrosinase to determine if they have
beneficial characteristics cumpared to the musbraom tyroshse used in this srudy (e-g.
thexmal stability, pH range of cataîytic activity and wst); and
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füngal laccases.
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