• biotechnology;
  • fungi;
  • molecular engineering;
  • tyrosinases


  1. Top of page
  2. Abstract
  3. Introduction
  4. Characteristics of fungal tyrosinases
  5. Biotechnological applications
  6. Bioengineering advances
  7. Conclusions
  8. Acknowledgements
  9. References

Tyrosinases are type-3 copper proteins involved in the initial step of melanin synthesis. These enzymes catalyse both the o-hydroxylation of monophenols and the subsequent oxidation of the resulting o-diphenols into reactive o-quinones, which evolve spontaneously to produce intermediates, which associate in dark brown pigments. In fungi, tyrosinases are generally associated with the formation and stability of spores, in defence and virulence mechanisms, and in browning and pigmentation. First characterized from the edible mushroom Agaricus bisporus because of undesirable enzymatic browning problems during postharvest storage, tyrosinases were found, more recently, in several other fungi with relevant insights into molecular and genetic characteristics and into reaction mechanisms, highlighting their very promising properties for biotechnological applications. The limit of these applications remains in the fact that native fungal tyrosinases are generally intracellular and produced in low quantity. This review compiles the recent data on biochemical and molecular properties of fungal tyrosinases, underlining their importance in the biotechnological use of these enzymes. Next, their most promising applications in food, pharmaceutical and environmental fields are presented and the bioengineering approaches used for the development of tyrosinase-overproducing fungal strains are discussed.


3,4-dihydroxyphenyl L-alanine


  1. Top of page
  2. Abstract
  3. Introduction
  4. Characteristics of fungal tyrosinases
  5. Biotechnological applications
  6. Bioengineering advances
  7. Conclusions
  8. Acknowledgements
  9. References

Tyrosinases (monophenol, o-diphenol:oxygen oxidoreductase, EC belong to a larger group of proteins named type-3 copper proteins, which include the catecholoxidases from plants and the oxygen-carrier haemocyanins from molluscs and arthropods (Gerdemann et al. 2002). Tyrosinases are involved in the melanin pathway and are especially responsible for the first steps of melanin synthesis from L-tyrosine leading to the formation of L-dopaquinone and L-dopachrome (Sanchez-Ferrer et al. 1995). The particularity of tyrosinases is to catalyse the o-hydroxylation of monophenols (cresolase activity or ‘‘monophenolase’’) and the subsequent oxidation of the resulting o-diphenols into reactive o-quinones (catecholase activity or ‘‘diphenolase’’), both reactions using molecular oxygen. Subsequently, the o-quinones undergo non-enzymatic reactions with various nucleophiles, producing intermediates, which associate spontaneously in dark brown pigments (Soler-Rivas et al. 1999).

Tyrosinases are present in mammals, invertebrates, plants, and microorganisms in which they are involved in several biological functions and have considerable heterogeneity (Van Gelder et al. 1997). Tyrosinases were first characterized in mammals for their role in the development of melanomas, and for their implication in pigmentation troubles such as albinism and vitiligo (Riley 1997). In fungi, tyrosinases are mainly associated with browning and pigmentation (Kurahashi and Pontzen 1998, Soler-Rivas et al. 1999). Melanins in the fungal cell-walls are derived from L-tyrosine, γ-glutaminyl-3,4-dihyroxybenzene (GDHB), or catechol in the case of Basidiomycotina, and from 1,8-dihydroxynaphthalene (DHN) in the case of Ascomycotina (Bell and Wheeler 1986). Melanins constitute a mechanism of defence and resistance to stress such as UV radiations, free radicals, gamma rays, dehydratation and extreme temperatures, and contribute to the fungal cell-wall resistance against hydrolytic enzymes in avoiding cellular lysis (Bell and Wheeler 1986). Fungal pigments are also involved in the formation and stability of spores (Mayer and Harel 1979), in the defence and virulence mechanisms (Soler-Rivas et al. 1997, Jacobson 2000). Fungal tyrosinases were firstly characterized from the edible mushroom Agaricus bisporus (Nakamura et al. 1966, Robb and Gutteridge 1981, Wichers et al. 1996) because of enzymatic browning during development and postharvest storage, which particularly decreases the commercial value of the product (Jolivet et al. 1998). Further studies on tyrosinases from other fungi such as Neurospora crassa (Kupper et al. 1989), Lentinula edodes (Kanda et al. 1996), Aspergillus oryzae (Nakamura et al. 2000), and Pycnoporus sanguineus (Halaouli et al. 2005a) gave new insights into mushroom-tyrosinase biochemical and molecular characteristics. Because filamentous fungi are now recognized for their potential in biotechnological applications, this review is devoted to fungal tyrosinases with a special focus on recent biochemical and molecular biology advances. The exploitation of fungal tyrosinases for applications in food and nonfood industries will be discussed, especially the bioengineering approaches used to overcome the biotechnological limitations.

Characteristics of fungal tyrosinases

  1. Top of page
  2. Abstract
  3. Introduction
  4. Characteristics of fungal tyrosinases
  5. Biotechnological applications
  6. Bioengineering advances
  7. Conclusions
  8. Acknowledgements
  9. References

Biochemical and molecular features

Fungal tyrosinases are cytosolic enzymes, which have considerable heterogeneity compared to other copper-containing phenoloxidases such as laccases for example, and the first studies showed a range of various molecular weights (Van Gelder et al. 1997). The early assays for A. bisporus tyrosinase purification from crude carpophore extracts resulted in multiple forms, often polymeric. In 1963, Bouchilloux et al. highlighted the existence of several tyrosinase forms (α, β, γ, δ) with apparent molecular masses of 118–119 kDa by preparative electrophoresis and hydroxyapatite chromatography. These forms would result from the association of 30–34 kDa monomers. In 1969, five different forms (I, Ia, II–IV), ranging from monomers to octamers, were described by Jolley et al. (1969) who suggested that the predominant enzyme was tetrameric and composed of four identical sub-units of 32 kDa. Strothkamp et al. (1976) concluded that the α, β, γ and δ forms previously isolated (Bouchilloux et al. 1963) were in fact heteropolymers comprising two different polypeptidic subunits: a heavy chain (H) and a light chain (L) with sizes of 43 and 13·4 kDa respectively. In aqueous solution, the active form seemed to be the L2H form with an apparent molecular mass of 69 kDa. According to Strothkamp et al. (1976), the monomers of 30–34 kDa were the result of the aggregation of L chains in dimers. Two types of H chains were further identified (Robb and Gutteridge 1981): Hα (48 kDa) and Hβ (45 kDa) and two isoenzymes were thought to be present: αHinline imageL2 and γHinline imageL2. Thus, multiple forms could be attributed to conformational isomerism, differences in genetic expression, or limited proteolysis (Lerch 1981). Today, it is thought that the forms of high molecular-mass were artefacts in the preparation of tyrosinases from crude mushroom extracts. Indeed, fungi in general, but especially tyrosinase-containing strains, are rich in phenolic compounds and pigments which would aggregate with proteins in high molecular-mass complexes (Jolivet et al. 1998). Assays to eliminate these phenolics during A. bisporus tyrosinase purification using anion-exchange chromatography or specific adsorbents led to the disappearance of several isoforms (Robb 1984). Flurkey (1991) isolated two groups of A. bisporus tyrosinases based on their pI: one group with pI between 4–4·5 and the other with pI between 4·5–5. Later, Gerritsen et al. (1994) showed that the lower-pI tyrosinases were glycosylated whereas the higher-pI tyrosinases were not. Two monomeric tyrosinases were isolated from fruitbodies of the A. bisporus strain U1 (Wichers et al. 1996) and were encoded by two different genes: AbPPO1 and AbPPO2 (Wichers et al. 2003). Both tyrosinases, with pI ca 5·2 and 5·1, showed a molecular mass of ca. 43 kDa under reducing conditions and ca. 47 kDa under native conditions (Table 1). Both monomers were fully active (Wichers et al. 2003). The ORF for A. bisporus AbPPO1 and AbPPO2 cDNA were shown to code for tyrosinases of ca 64 kDa (Wichers et al. 2003) whereas the molecular mass of the mushroom extract proteins were ca. 45 kDa after activation by serine proteases (Espin et al. 1999). In the case of A. bisporus enzymes, tyrosine in position 381 (a chymotrypsin cleavage site) was postulated as the putative cleavage site in the C-terminus (Wichers et al. 2003). First described as tetrameric aggregates of active 47 kDa monomers (Robb and Gutteridge 1981), the N. crassa tyrosinase is now regarded as a 46 kDa monomer with full enzymatic activity, formed upon proteolytic processing of a 75 kDa precursor (Kupper et al. 1989). The mature form is 407 amino acids, which is derived from the cleavage of 213 amino acids (21 kDa) from the C-terminal end of the protyrosinase. The phenylalanine in position 407 is a chymotrypsin-like cleavage site as well (Kupper et al. 1989). The A. oryzae tyrosinase was described as a tetrameric protein consisting of 67 kDa subunits (Ichishima et al. 1984); the enzyme could be activated by acidic shock and the 67 kDa monomeric protein (Table 1) was active (Fujita et al. 1995). Six tyrosinase isoforms could be purified from the gill of L. edodes fruit-bodies and consisted of one catalytic subunit (54 or 55 kDa) and one hypothetic regulatory subunit (50 or 15 kDa) (Kanda et al. 1996). More recently, a new monomeric tyrosinase of 45 kDa was isolated from the intracellular medium of the white-rot fungus P. sanguineus (Table 1), and showed four isoforms after isoelectric focusing (Halaouli et al. 2005a). These isoforms might be explained by the different phosphorylation degree or different global charge due to the binding of phenolics (Halaouli et al. 2005a).

Table 1.  Main biochemical characteristics of fungal tyrosinases
FungiMW of the active form (kDa)pH optimumpIKm (mM)VmSpecific activity (U/mg protein)References
  1. *TS: thermostable isozyme; TL: thermolabile isozyme.

A. bisporus47nd5·1–5·20·44 (L-DOPA) 0·76 (catechol)24·5 μM/min (L-DOPA) 288·0 μM/min (catechol)ndEspin et al. 1997Wichers et al. 2003
A. bisporus (Portabella mushroom)487·05·1–5·35 (catechol) 9 (L-DOPA)ndndZhang et al. 1999Fan and Flurkey 2004
A. oryzae675·0-6·0ndndndndIchishima et al. 1984
L. edodes70–1056–6·54·3–4·7ndndndKanda et al. 1996
N. crassa46 (TS*)nd8·30·88 (L-DOPA)nd1340 (L-DOPA)Lerch 1983
N. crassa46 (TL*)nd8·50·95 (L-DOPA)nd1330 (L-DOPA)Lerch 1983
P. sanguineus456·5–7·04·5–5·01·0 (L-tyrosine) 0·9 (L-DOPA)54 (U/mg protein) (L-tyrosine) 112 (U/mg protein) (L-DOPA)30 (L-tyrosine) 84 (L-DOPA)Halaouli et al. 2005a

In micro-organisms, animals, plants and insects, tyrosinases were shown to exist in both latent (inactive) and active forms (Whitaker 1995). In A. bisporus carpophores, the latent form represented 98–99% of the total tyrosinase activity (Van Leeuwen and Wichers 1999). In vitro, latent fungal tyrosinases could be activated by acidic shocks (Ichishima et al. 1984), sodium dodecyl sulphate (Espin and Wichers 1999), and proteases (Espin et al. 1999). In vivo, tyrosinase activation was assumed to be the result of ageing (Burton et al. 1993), exposition to extreme environmental conditions or to pathogens (Soler-Rivas et al. 1997, Jolivet et al. 1998). The mechanism of fungal activation remains unclear and, today, it can only be supposed that it is the consequence of a conformational enzymatic change (Espin and Wichers 1999) or a simple protein solubilization (Mayer and Harel 1979). However, it is now thought that the activation process involves endogenous proteases such as the serine proteases isolated from A. bisporus during the postharvage mushroom senescence. Two such proteases could also be identified in A. oryzae (Ichishima et al. 1984).

Phylogenetic analysis, based on the whole tyrosinase protein sequences, showed that fungal tyrosinases clustered in groups for basidiomycetes, ascomycetes and deuteromycetes (Fig. 1). This observation was confirmed by the quantitative data on identity: for example, P. sanguineus tyrosinase showed the highest identity and similarity to ones from Polyporus arcularius (67% and 75%) (Morinaga 2003) and L. edodes (56% and 67%) (Sato 2001). Comparison between tyrosinases from the basidiomycetes P. sanguineus (Halaouli et al. 2005b), P. arcularius (Morinaga 2003), L. edodes (Sato 2001) and A. bisporus (Ebbelar et al. 1995, Wichers et al. 2003), the deuteromycetes Aspergillus fumigatus (Langfelder et al. 2000), and A. oryzae (Fujita et al. 1995), and the ascomycetes N. crassa (Schulte et al. 2003) and Podospora anserina (Marbach and Stahl 1996) highlighted an important similarity within the first 300 residues (Fig. 2), corresponding to the copper-binding domains, which is a common central domain highly conserved among the type-3 copper-containing metalloproteins (Seo et al. 2003). Since the last decade, several fungal tyrosinase-encoding genes have been isolated (Table 2), and share high heterogeneity concerning length, exon number and homology. For example, the P. sanguineus tyrosinase gene shared very low homology with other basidiomycete fungal tyrosinase nucleotide sequences: 51, 17 and 7% in the case of L. edodes, P. arcularius and A. bisporus, respectively.


Figure 1. Phylogenetic analysis of fungal tyrosinases. Distance analysis and tree construction of amino acid sequence data were done by the Neighbour-joining method. Numbers indicate the percent support for nodes after 1,000 replications of bootstrap analysis. The bar represents a 10% sequence difference.

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Figure 2. Amino acid sequence alignment of tyrosinases, haemocyanins and catechol oxidases. Conserved amino acids involved in the active site structure are shown in grey, conserved histidine residues are in bold, and copper-containing domains are boxed. The sequence accession numbers in NCBI Database are: Q00024 for Agaricus bisporus AbPPO1 (Wichers et al. 2003), O42713 for Agaricus bisporus AbPPO2 (Wichers et al. 2003), BAB71736 for Lentinus edodes (Sato 2001), CAC82195 for Aspergillus fumigatus (Langfelder et al. 2000), Q00234 for Aspergillus oryzae (Fujita et al. 1995), P00440 for Neurospora crassa TS (Kupper et al. 1989), Q92396 for Podospora anserina (Marbach and Stahl 1996), BAD51402 for Polyporus arcularius (Kanda et al. 2003), AY842859 for Pycnoporus sanguineus (Halaouli et al. 2005b), NP 826539 for Streptomyces avermitilis (Ikeda et al. 2003), CAB92266 for Streptomyces coelicolor (Bentley 2002), A24089 for Streptomyces glauscensens (Huber et al. 1985), NP035791 for Mus musculus (Lavado et al. 2005), AAH27179 for Homo sapiens (Strausberg 2002), CAA77764 for Vicia faba (Cary et al. 1992), CAA78300 for Lycopersicon esculentum (Newman 1996), P04254 for Panulirus interuptus (Bak et al. 1986), P56825 for Helix pomatia (Gielens et al. 1997).

Table 2.  Main features of fungal tyrosinase-encoding genes isolated from Deuteromycetes, Ascomycetes and Basidomycetes
FungiGeneGene length (bp)Coding region length (bp)Number of exonsAccession number (NCBI Database)References
A. bisporus U1AbPPO11916X85113Wichers et al. 2003
A. bisporus U1AbPPO21916X89382Wichers et al. 2003
A. fumigatusTyr 1230718858AJ293806Langfelder et al. 2000
A. oryzaeMel O175116962D37929Fujita et al. 1995
L. edodesgLeTyr231818489AB033993Sato 2001
N. crassa TST201618633M33271Kupper et al. 1989
N. crassa ORT201618633M32843Kupper et al. 1989
P. anserinaTyr189617114U66807Marbach and Stahl 1996
P. arcularius232818738AB120567Kanda et al. 2003
P. sanguineusgPycTyr185718537AY849378Halaouli et al. 2005b

The reactional mechanism

Depending on the copper-ion valence and linking with molecular oxygen, the active site of tyrosinases can exist in three intermediate states: deoxy (CuI–CuI), oxy (CuII–O2–CuII) and met (CuII–CuII) (Sanchez-Ferrer et al. 1995). The met-form is converted into deoxy-form in a two electron reduction, and the resulting deoxy-form is able to reversibly fix molecular oxygen leading to the oxy-form (Fig. 3). The oxy-form can also be obtained by direct addition of hydrogen peroxide to met-tyrosinase in the case of N. crassa (Lerch 1983) or by treatment of the met-form with reducing agents such as ascorbic acid, hydroxylamine, dithionite in the case of Aspergillus flavipes (Lerch 1988), Fe2+ (Gukasyan 2002), or dithiothreitol in the case of A. bisporus (Naish-Byfield et al. 1994). In vivo, most of the resting enzyme is in the met-form, which is unable to bind molecular oxygen (Lerch 1981). This predominant form does not act on monophenols, but has an important affinity for them and binds them through a dead-end pathway, resulting in a lag period (Cabanes et al. 2002). The exact mechanism of reaction of tyrosinases remains partially unclear, but it is generally admitted that ortho-diphenol oxidation by tyrosinases involves Michaelis-Menten kinetics, whereas monophenols hydroxylation is characterized by a phase of latency (Sanchez-Ferrer et al. 1995). The characteristic initial lag in the cresolase cycle can be abolished by addition of low amounts of ortho-diphenols as shown by Espin and Wichers (2001) with A. bisporus tyrosinase. Actually, it is recognized that, in vivo, DOPA is responsible for the recruitment of the resting enzyme by reducing met-forms to deoxy-forms (Solomon and Lowery 1993). A model for the reaction mechanism of tyrosinases (Fig. 3) including both cresolase (Fig. 3, ways 6,7,2) and catecholase (Fig. 3, ways 1–5) activities has been proposed (Solomon et al. 1996; Cabanes et al. 2002) taking into account the following characteristics: (i) both met- and oxy-forms can react with diphenols (Fig. 3, ways 1,4), whereas only oxy-forms can react with monophenols (Fig. 3, way 6), (ii) monophenols can compete with diphenols for binding to met-tyrosinase inhibiting its reduction to the deoxy-form (Fig. 3, way 8), (iii) a chemical step is necessary for the recycling of L-DOPA from ortho-DOPAquinones (Fig. 3, way 9). Both cresolase and catecholase cycles produce ortho-quinones, which are further spontaneously rearranged into polymeric pigments (Rodriguez-Lopez et al. 1992). Recently, Yamazaki et al. (2004) showed that mushroom tyrosinase could also have a catalase and peroxygenase (H2O2-dependent oxygenation of substrates) activity.


Figure 3. Catalytic cycle for fungal-tyrosinase monophenolase and diphenolase activities (according to Solomon et al. 1996, Cabanes et al. 2002). M: monophenol; D: diphenol; Q: quinone.

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Active site characteristics

Although tyrosinase was one of the first discovered monooxygenases, its crystallographic structure has not been yet elucidated. However, it can be assumed that tyrosinases, haemocyanins and catechol oxidases share similar binuclear copper sites considering the following features: (i) comparable valence and conformational change during oxygen binding process (Woolery et al. 1984, Della Longa et al. 1996), (ii) comparable spectroscopic and magnetic properties (Himmelwright et al. 1980), (iii) homologies in the primary sequence (Van Gelder et al. 1997), (iv) catechol oxidases showed monophenolase activity on 4-hydroxyanisole (Espin et al. 1998), (v) haemocyanins, which are oxygen-carriers, showed catecholase activity after in vitro sodium dodecyl sulphate or protease treatments (Decker and Rimke 1998, Jaenicke and Decker 2004), as in the case of fungal tyrosinases (Espin et al. 1999), probably resulting from a higher accessibility of phenolics to the active site.

The adjacent amino acid residues surrounding the copper ions are generally highly conserved among tyrosinases, catecholoxidases and mollusc haemocyanins (Fig. 2). Each of the two copper regions of these proteins contains three imidazole residues bound to copper ions (Lerch 1983, Jackman et al. 1992). All of the conserved copper ligands, namely HA1, HA2, HA3 for copper A, and HB1, HB2, HB3 for copper B, follow the general rules HA1-x(n)-HA2-x(8)-HA3 and HB1-x(3)-HB2-x(n)-HB3 (Fig. 2), where n is a variable number of residues (Garcia-Borron and Solano 2002). Considering crystallographic data for catecholoxidase (Klabunde et al. 1998) and structure predictions for N. crassa (Kupper et al. 1989) and A. bisporus (Van Gelder et al. 1997) tyrosinases, Garcia-Borron and Solano (2002) described the tyrosinases active site as a hydrophilic sphere delimited by a four-helix bundle containing the six imidazole residues. The hydrophilic sphere would be located in a hydrophobic shell which is aromatic and formed by highly conserved residues: F(HA3−4), Φ(HA3−1), and Φ(HA3+3) around CuA, F(HB3−4), and H(HB3−1) around CuB (with Φ meaning aromatic residues). In this arrangement, the configuration of the tyrosinases active site would essentially be maintained by electrostatic or cations-π interactions. According to Garcia-Borron and Solano (2002), these noncovalent binding forces are the result of interactions between several residues located at the proximity of the dicopper center: Φ(HA1−7), R(HA3+1), Φ(HA3+3), Φ(HA3+7), E(HA3+8), D(HB3−7), D(HB3+4), Φ(HB3+7), W(HB3+10). The comparison of fungal tyrosinases primary structures shows the conservation of the following amino acids (Fig. 2): Φ(HA1−7) (with Φ corresponding to F or Y), R(HA3+1), Φ(HA3+3) (with Φ corresponding to Y), E(HA3+8), D(HB3−7), D(HB3+4), and W(HB3+10). Another feature of the binuclear copper centre in fungal tyrosinases is the highly conserved C(HA2−2) which is involved in a covalent thio-ether bond with the copper A ligand HA2 (Lerch 1982). Such a covalent cysteine-histidine bridge was reported in molluscan haemocyanins subunits (Gielens et al. 1997) and in the catecholoxidase from Ipomea batatas (Klabunde et al. 1998). In the case of N. crassa tyrosinase, Lerch (1982) suggested that this unusual thio-ether linkage was formed during an intramolecular rearrangement of the enzyme, resulting in the activation of the protyrosinase, whereas Nakamura et al. (2000) suggested the involvement of the C(HA2−2) residue in the binding to the copper A atom in the A. oryzae tyrosinase. According to Klabunde et al. (1998), it seems likely that this structure optimizes the redox potential of the copper centre for ortho-diphenol oxidation, allowing for a rapid electron transfer in the redox processes.

Biotechnological applications

  1. Top of page
  2. Abstract
  3. Introduction
  4. Characteristics of fungal tyrosinases
  5. Biotechnological applications
  6. Bioengineering advances
  7. Conclusions
  8. Acknowledgements
  9. References

Until the last decade, studies on fungal tyrosinases, mostly from the edible mushroom A. bisporus, were essentially motivated by the enzymatic browning phenomenon during development and postharvest storage (Jolivet et al. 1998). Today, an increasing interest is dedicated to the potential of fungal tyrosinases in biotechnological and environmental applications. The ability of tyrosinases to convert monophenols into diphenols has first motivated studies concerning the production of antioxidant ortho-diphenols with beneficial properties as food additives or pharmaceutical drugs. The production of caffeic acid (3,4-dihydroxyphenyl-2-propenoic acid) using p-coumaric acid (4-hydroxyphenyl-2-propenoic acid) as the substrate, A. bisporus tyrosinase as the biocatalyst, and ascorbic acid as the inhibitor of quinone formation, was firstly reported by Sâto (1969). Hydroxytyrosol (2-(3,4-dihydroxyphenyl)ethanol) is another strong antioxidant which occurs naturally in olive oil and fruits, but which is not commercially available (Espin et al. 2000). Recently, the purified tyrosinases from A. bisporus (Espin et al. 2001) and P. sanguineus (Halaouli et al. 2005a) have demonstrated effectiveness in the biosynthesis of hydroxytyrosol from p-tyrosol (2-(4-hydroxyphenyl)ethanol), a monophenol found in agro-industrial by-products such as olive mill wastewaters (Lesage-Meessen et al. 2001). This type of enzymatic bioconversion can constitute an alternative to the chemical synthesis when ‘‘natural’’ compounds are required, and has also been investigated for the biosynthesis of useful drugs such as L-DOPA involved in the treatment of Parkinson's and myocardium's diseases (Raju et al. 1993). L-DOPA production has been achieved, using L-tyrosine as the substrate, with the A. oryzae (Haq et al. 2002) and Aspergillus flavus (Singh 1999) tyrosinases as the biocatalysts.

The quinones subsequently produced from ortho-diphenols oxidation by tyrosinases are reactive and can undergo non-enzymatic reactions with a variety of nucleophiles leading to the formation of biopolymers. Consequently, the potential use of fungal tyrosinase for enzymatic cross-linking has been investigated using several types of substrates such as polysaccharides and proteins. The A. bisporus tyrosinase has been used for the enzymatic grafting of phenolic moieties (Lenhart et al. 1998) or proteins (Aberg et al. 2004) onto chitosan, a polysaccharide biopolymer, obtained mainly from food-processing wastes. The A. bisporus tyrosinase catalyses the oxidation of phenols to electrophilic ortho-quinones which are able to freely diffuse and covalently bond with the nucleophilic amine groups of chitosan. Such modified chitosans provide a safe way to enlarge environmentally friendly polymers with useful viscoelastic properties such as hydrogels for skin substitutes (Kane et al. 1996), adhesives (Peppas and Sahlin 1996), and matrices for drug delivery and for tissue engineering (Mallapragada and Narasimhan 1998; Lee and Mooney 2001). The A. bisporus tyrosinase was also shown to induce rapid gelation of polyethyleneglycol conjugated with L-DOPA to form hydrogels (Lee et al. 2002). Mushroom tyrosinase was furthermore observed to catalyse protein-polysaccharide hydrogels formation with gelatine and chitosan (Chen et al. 2002). Enzymatic protein-protein cross-linking is also showing increasing interest, particularly in food technology. Until now, protein cross-linking has been described for only a few enzymes such as transglutaminase (for review, see Matheis and Whitaker 1987), based on the formation of cross-links between the glutamine and the lysine residues. However, the limited number and/or the stearic inaccessibility of the corresponding binding-sites limit the cross-linking of several proteins in the absence of reducing or denaturing agents (Færgemand et al. 1998). Fungal tyrosinases have been shown to in vitro polymerize several proteins such as lysozyme and ribonuclease (Leatham et al. 1980), casein (Hurrell et al. 1982), α-lactalbumin and β-lactoglobulin (Thalman and Lötzbeyer 2002), in the presence of low molecular-weight phenolics, such as chlorogenic and caffeic acids, as cross-linkers. In this process, the phenols acted as the substrate for the cross-linking process, being integrated into the polymerising complex as an interconnection between the proteins (Matheis and Whitaker 1984). Direct tyrosinase-induced protein cross-linking is also possible via tyrosine residues. Recently, the direct protein-protein polymerization of α-lactalbumin by A. bisporus tyrosinase (Thalman and Lötzbeyer 2002), and of casein by P. sanguineus tyrosinase (Halaouli et al. 2005a) has successfully been performed. Kato et al. (1986) showed that the ortho-quinone tyrosinase-oxidation products could covalently bind to proteins through sulphhydryl groups. Takasaki and Kawakishi (1997) reported that the A. bisporus tyrosinase catalysed the formation of gluten-bound 5-S-cysteinylDOPA and L-DOPA as cross-linkers of gluten, explaining its oxidant-like effect on the physical properties of dough.

For several years, the use of oxidizing enzymes from filamentous fungi has been known as a biological and friendly solution for the treatment of wastes containing toxic and pollutant phenolics (Karam and Nicell 1997, Burton 2003). Peroxidases and laccases have been studied extensively for the treatment of aromatics but the resulting processes generally need expensive chemical cofactors (such as manganese, malonate and hydrogen peroxide) for industrial purpose (Paice et al. 1995, Bourbonnais and Paice 1996). In a general manner, tyrosinases need only molecular oxygen as a cofactor for substrate oxidation (Burton 2003). Purified preparations of mushroom tyrosinase showed to be effective in the precipitation or biotransformation of aqueous phenols (Ensuncho et al. 2005), and chlorophenols (Ikehata and Nicell 2000). The extracellular tyrosine-inducible tyrosinase from the Zygomycete Amylomyces rousii allowed for the efficient degradation of pentachlorophenol, a recalcitrant xenobiotic used as a pesticide (Montiel et al. 2004).

Bioengineering advances

  1. Top of page
  2. Abstract
  3. Introduction
  4. Characteristics of fungal tyrosinases
  5. Biotechnological applications
  6. Bioengineering advances
  7. Conclusions
  8. Acknowledgements
  9. References

Native fungal tyrosinases are generally intracellular, and produced in low quantity. Consequently, several problems have to be overcome before using them in any future biotechnological application.

The first problem is the formation, during tyrosinase isolation and purification, of melanin pigments which often remain bound to the protein after extraction. Rescigno et al. (1997) suggested improving the purification of the A. bisporus tyrosinase by adding ascorbic acid in order to avoid phenolic oxidation. The removal of both pigments and phenolics kept in a reduced state was further achieved by diafiltration. The purification of tyrosinases from mycelia is somewhat more difficult because of the presence of a large number of proteases in the crude extracts of fungi. Protease inhibitors such as phenylmethanesulfonyl fluoride were added in crude extracts obtained from L. edodes gills in order to prevent protease activity (Kanda et al. 1996).

Secondly, the use of these enzymes remains hampered by their nonreusability, and their low stability. Assays of immobilization on different solid supports, extensively reviewed by Duran et al. (2002), were carried out in attempts to increase the stability of tyrosinases. Immobilization showed to protect tyrosinase from inactivation by reaction with quinones, and to preserve them from proteolysis (Wada et al. 1992). This process also improved the thermal stability of fungal tyrosinases (Munjal and Sawhney 2002), and activity yields (Hamann and Saville 1996) in comparison to soluble enzymes. Tyrosinase immobilization procedures were performed for drug production (Carvalho et al. 2000, Seetharam and Saville 2002), and removal of phenols from industrial wastewaters (Krastanov 2000, Ensuncho et al. 2005). Seo et al. (2003) detailed the development of biosensors with immobilized mushroom tyrosinase, enabling the quantification of phenols and the removal of related compounds in water and soil samples. More recently, tyrosinase biosensors have been used for the monitoring of alkaline phosphatase activity (Serra et al. 2005), a catechol-producing enzyme.

Lastly, one of the most important requirements to facilitate molecular studies and biotechnological applications is the development of expression systems for large-scale production, possibly in the extracellular medium. Until now, very few tyrosinase-promoters have been described (Ishida et al. 2001), and the regulation of fungal tyrosinases expression remains misunderstood. Nevertheless, it is widely admitted that fungal tyrosinases are naturally produced in response to starvation and, generally, in limited amounts. For instance, the N. crassa tyrosinase seems to be synthesized only during sexual differentiation (Lowry and Sussman 1958) or during starvation (Fling et al. 1963). Overproduction in a homologous host has been tried in the case of the tyrosinase-encoding gene from N. crassa, which codes for a protyrosinase of 75 kDa (Kupper et al. 1989). This gene was placed under the control of a strong and inducible metallothionein (MT) promoter in order to target the homologous expression of the protyrosinase gene in vegetative cultures of N. crassa. In tyrosinase-positive transformants, an intracellular totally active enzyme was produced, but weak specific signals were detected since 3·17 and 4·38 U mg−1 of proteins were respectively produced constitutively and after induction (Kupper et al. 1990). According to the authors, the low yields of recombinant tyrosinase obtained could possibly be the result of the toxicity of the high intracellular amounts of this enzyme. Kupper et al. (1990) also suggested that mycelia of the heterokaryotic transformants may carry nuclei that do not integrate the fusion construct, which resulted in lower fusion-gene expression per single cell. Several heterologous expression systems have also been tried in order to increase production yields. IPTG-induced expression of the A. bisporus AbPPO2 encoding-tyrosinase cDNA was carried out in Escherichia coli cells, and resulted in the production of an inactive protyrosinase of 64 kDa (Wichers et al. 2003). Similarly, heterologous expression of the A. oryzae melO-encoding tyrosinase cDNA in Saccharomyces cerevisiae resulted in the production of an intracellular inactive proenzyme (Fujita et al. 1995). However, reconstitution of the enzyme activity was possible after an acidic treatment of the cell homogenate at pH 3, leading to an average activity of 1 mU kg−1 of proteins (Fujita et al. 1995). More recently, the recombinant production of a P. sanguineus tyrosinase has been performed by heterologous expression in A. niger (Halaouli et al. 2005b) with an expression system already shown to be suitable for high-level protein production and exploited for various homologous and heterologous proteins (Van den Hondel et al. 1992). Encoding-tyrosinase cDNA was fused to the glucoamylase preprosequence in attempts to target the secretion in the extracellular medium, and was placed under the control of the strong and constitutive glyceraldehyde phosphate dehydrogenase promoter. The maturation process was shown to be effective in A. niger and the recombinant enzyme was fully active with a molecular mass of 45 kDa. Extracellular tyrosinase activities of 534 and 1,668 U l−1 for monophenolase and diphenolase, respectively, were obtained which corresponded to a protein yield of ca. 20 mg l−1 (Halaouli et al. 2005b), opening the way to higher yielding processes.


  1. Top of page
  2. Abstract
  3. Introduction
  4. Characteristics of fungal tyrosinases
  5. Biotechnological applications
  6. Bioengineering advances
  7. Conclusions
  8. Acknowledgements
  9. References

Research in the last decade has increased the knowledge on fungal tyrosinases, highlighting similarities and differences with other type-3 copper metalloproteins. Today, the detailed and complete understanding of their functioning still remains hampered by their cumbersome purification and the occurrence of several isoforms, either latent or active, in wild strains. However, this review shows that fungal tyrosinases remain an exciting field of research due to their vast importance in food, pharmaceutical and environmental industries. Homologous or heterologous gene expression in various hosts will be useful for improving, in the future, the enzymatic activity. More efforts in this direction will certainly help to explain the low expression yields, to design suitable expression constructs for extracellular production, and to improve productivity at the pilot scale.


  1. Top of page
  2. Abstract
  3. Introduction
  4. Characteristics of fungal tyrosinases
  5. Biotechnological applications
  6. Bioengineering advances
  7. Conclusions
  8. Acknowledgements
  9. References

We warmly thank D. Navarro (INRA, Marseilles, France) for phylogenetic analysis and C. Daniels for helpful comments on the manuscript. This work has been carried out with financial support from the Commission of the European Communities, specific RTD programme ‘‘Quality of Life and management of Living Resources’’, proposal number QLK1-2002-02208 ‘‘Novel cross-linking enzymes and their consumer acceptance for structure engineering of foods’’, acronym CROSSENZ. It does not reflect its views and in no way anticipates the Commission's future policy in this area.


  1. Top of page
  2. Abstract
  3. Introduction
  4. Characteristics of fungal tyrosinases
  5. Biotechnological applications
  6. Bioengineering advances
  7. Conclusions
  8. Acknowledgements
  9. References
  • Aberg, C.M., Chen, T., Olumide, A., Raghavan, S.R. and Payne, G.F. (2004) Enzymatic Grafting of Peptides from Casein Hydrosylates to Chitosan. Potential for Value-Added Byproducts from Food-Processing Wastes. J Agric Food Chem 52, 788793.
  • Bak, H.J., Neuteboom, B., Jekel, P.A., Soeter, N.M., Vereijken, J.M. and Beintema, J.J. (1986) Structure of arthropod hemocyanins. FEBS Lett 204, 141144.
  • Bell, A.A. and Wheeler, M.H. (1986) Biosynthesis and functions of fungal melanins. Ann Rev Phytopathol 24, 411451.
  • Bentley, S.D. (2002) NCBI Database. URL
  • Bouchilloux, S., McMahill, P. and Mason, H.S. (1963) The multiple forms of mushroom tyrosinase. Purification and molecular properties of the enzymes. J Biol Chem 238, 16991707.
  • Bourbonnais, R. and Paice, M.G. (1996) Enzymatic delignification of kraft pulp using laccase and a mediator. TAPPI J 79, 199204.
  • Burton, K.S., Love, M.E. and Smith, J.F. (1993) Biochemical changes associated with mushroom quality in Agaricus spp. Enzyme Microb Technol 15, 736741.
  • Burton, S.G. (2003) Oxidizing enzymes as biocatalysts. Trends Biotechnol 21, 543549.
  • Cabanes, J., Chazarra, S. and Garcia-Carmona, F. (2002) Tyrosinase kinetics: a semi-quantative model of the mechanism of oxidation of monohydric and dihydric phenolic substrates-reply. J Theo Biol 214, 321325.
  • Carvalho, G.M.J., Alves, T.L.M. and Freire, D.M.G. (2000) L-DOPA Production by immobilized tyrosinase. Appl Biochem Biotechnol 84–86, 791800.
  • Cary, J.W., Lax, A.R. and Flurkey, W.H. (1992) Cloning and characterization of cDNAs coding for Vicia faba polyphenol oxidase. Plant Mol Biol 20, 245253.
  • Chen, T., Embree, H.D., Wu, L.-Q. and Payne, G.F. (2002) In vitro protein-polysaccharide conjugation: tyrosinase catalysed conjugation of gelatin and chitosan. Biopolymers 64, 292302.
  • Decker, H. and Rimke, T. (1998) Tarantula hemocyanin shows phenoloxidase activity. J Biol Chem 273, 2588925892.
  • Della Longa, S., Ascone, I., Bianconi, A., Bonfigli, A., Castellano, A.C., Zarivi, O. and Miranda, M. (1996) The dinuclear copper site structure of Agaricus bisporus tyrosinase in solution probed by X-ray absorption spectroscopy. J Biol Chem 271, 2102521030.
  • Duran, N., Rosa, M.A., D'Annibale, A. and Gianfreda, L. (2002) Applications of lactases and tyrosinases (phenoloxidases) immobilized on different supports: a review. Enzyme Microb Technol 6179, 125.
  • Ebbelar, C.E.M., Wichers, H.J., van den Bosch, T., Oyevaar, J.I. and Recourt, K. (1995) NCBI Database. URL
  • Ensuncho, L., Alvarez-Cuenca, M. and Legge, R.L. (2005) Removal of aqueous phenol using immobilized enzymes in a bench scale and pilot scale three-phase fluidized bed reactor. Bioprocess Biosyst Eng 27, 185191.
  • Espin, J.C. and Wichers, H.J. (1999) Activation of a latent mushroom (Agaricus bisporus) tyrosinase isoform by sodium dodecyl sulfate (SDS). Kinetic properties of the SDS-activated isoform. J Agric Food Chem 47, 35183525.
  • Espin, J.C. and Wichers, H.J. (2001) Effect of captopril on mushroom tyrosinase activity in vitro. Biochim Biophys Acta 1544, 289300.
  • Espin, J.C., Morales, M., Garcia-Ruiz, P.A., Tudela, J. and Garcia-Canovas, F. (1997) Improvement of a continuous spectrophotometric method for determining the monophenolase and diphenolase activities of mushroom polyphenol oxidase. J Agric Food Chem 45, 10841090.
  • Espin, J.C., Tudela, J. and Garcia-Carnovas, F. (1998) 4-Hydroxyanisole: The most suitable monophenolic substrate for determining spectrophotometrically the monophenolase activity of polyphenol oxidase from fruits and vegetables. Anal Biochem 259, 118126.
  • Espin, J.C., Van Leeuwen, J. and Wichers, H.J. (1999) Kinetic study of the activation process of a latent mushroom (Agaricus bisporus) tyrosinase by serine proteases. J Agric Food Chem 47, 35093517.
  • Espin, J.C., Soler-Rivas, C. and Wichers, H.J. (2000) Characterization of the total free radical scavenger capacity of vegetables oils and oil fractions using 2,2-diphenyl-1-picrylhydrazyl radical. J Agric Food Chem 48, 648656.
  • Espin, J.C., Soler-Rivas, C., Cantos, E., Tomas-Barberan, F.A. and Wichers, H.J. (2001) Synthesis of the antioxidant hydroxytyrosol using tyrosinase as biocatalyst. J Agric Food Chem 49, 11871193.
  • Fan, Y. and Flurkey, W.H. (2004) Purification and characterization of tyrosinase from gill tissue of Portabella mushroom. Phytochem 65, 671678.
  • Færgemand, M., Otte, J. and Qvist, K.B. (1998) Cross-linking of whey Proteins by enzymatic oxidation. J Agric Food Chem 46, 13261333.
  • Fling, M., Horowitz, N.H. and Heinemann, S.F. (1963) The isolation and properties of crystalline tyrosinase from Neurospora. J Biol Chem 238, 20452053.
  • Flurkey, W.H. (1991) Identification of tyrosinase in mushrooms by isoelectric focusing. J Food Sci 56, 9395.
  • Fujita, Y., Uraga, Y. and Ichisima, E. (1995) Molecular cloning and nucleotide sequence of the protyrosinase gene, melO, from Aspergillus oryzae and expression of the gene in yeast cells. Biochim Biophys Acta 1261, 151154.
  • Garcia-Borron, J.C. and Solano, F. (2002) Molecular anatomy of tyrosinase and its related proteins: beyond the histidine bound metal catalytic center. Pigment Cell Res 15, 162173.
  • Gerdemann, C., Christoph, E. and Krebs, B. (2002) The crystal structure of catechol oxidase: New insight into the function of Type-3 copper proteins. Account Chem Res 35, 183191.
  • Gerritsen, Y.A.M., Chapelon, C.G.J. and Wichers, H.J. (1994) The low-isoelectric point tyrosinase of Agaricus bisporus may be a glycoprotein. Phytochem 35, 573577.
  • Gielens, C., DeGeest, N., Xin, X.-Q., Devreese, B., van Beeumen, J. and Préaux, G. (1997) Evidence for a cysteine-histidine thioether bridge in functional units of molluscan hemocyanins and location of the disulfide bridges in functional units d and g of the betaC-hemocyanin of Helix pomatia. Eur J Biochem 248, 879888.
  • Gukasyan, G.S. (2002) Study of the kinetics of oxidation of monophenols by tyrosinase. The effect of reducers. Biochem 67, 277280.
  • Halaouli, S., Asther, Mi., Kruus, K., Guo, L., Hamdi, M., Sigoillot, J.-C., Asther, M. and Lomascolo, A. (2005a) Characterization of a new tyrosinase from Pycnoporus species with high potential for food technological applications. J Appl Microbiol 98, 332343.
  • Halaouli, S., Record, E., Casalot, L., Hamdi, M., Sigoillot, J.-C., Asther, M. and Lomascolo, A. (2005b) Cloning and characterization of a tyrosinase gene from the white-rot fungus Pycnoporus sanguineus, and overproduction of the recombinant protein in Aspergillus niger. Appl Microbiol Biotechnol (in press, DOI 10.1007/s00253-005-0109-4).
  • Hamann, M.C.J. and Saville, B.A. (1996) Enhancement of tyrosinase stability by immobilization on Nylon-66. Food Bioprod Process 74, 4752.
  • Haq, I., Ali, S. and Qadeer, M.A. (2002) Biosynthesis of L-DOPA by Aspergillus oryzae. Biores Technol 85, 2529.
  • Himmelwright, R.S., Eickman, N.C. and Solomon, E.I. (1980) Chemical and spectroscopic studies of the binuclear copper site of Neurospora tyrosinase: comparison to hemocyanins. J Am Chem Soc 102, 73397344.
  • Huber, M., Hinterman, G. and Lerch, K. (1985) Primery structure of tyrosinase from Streptomyces glauscesens. Biochem 24, 60386044.
  • Hurrell, R.F., Finot, P.A. and Cuq, J.L. (1982) Protein-polyphenol reactions. 1. Nutritional and metabolic consequences of the reaction between oxidized caffeic acid and the lysine residues of casein. Br J Nutr 47, 191211.
  • Ichishima, E., Maeba, H., Amikura, T. and Sakata, H. (1984) Multiple forms of protyrosinase from Aspergillus oryzae and their mode of activation at pH 3.0. Biochim Biophys Acta 786, 2531.
  • Ikeda, H., Ishikawa, J., Hanamoto, A., Shinose, M., Kikuchi, H., Shiba, T., Sakaki, Y., Hattori, M. et al. (2003) Complete genome sequence and comparative analysis of the industrial microorganism Streptomyces avermitilis. Nat Biotechnol 21, 526531.
  • Ikehata, K. and Nicell, J.A. (2000) Color and toxicity removal following tyrosinase-catalysed oxidation of phenols. Biotechnol Progr 16, 533540.
  • Ishida, H., Matsumura, K., Hata, Kawatao, A., Suginami, K., Abe, Y., Imayasu, S. and Ichishima, E. (2001) Establishment of a hyper-protein production system in submerged Aspergillus oryzae culture under tyrosinase-encoding gene (melO) promoter control. Appl Microbiol Biotechnol 57, 131137.
  • Jackman, M.P., Huber, M., Hajnal, A. and Lerch, K. (1992) Stabilization of the oxy form of tyrosinase by a single conservative amino acid substitution. Biochem J 282, 915918.
  • Jacobson, E.S. (2000) Pathogenic roles for fungal melanins. Clin Microbiol Rev 13, 708717.
  • Jaenicke, E. and Decker, H. (2004) Conversion of crustacean hemocyanin to catecholoxidase. Micron 35, 8990.
  • Jolivet, J., Arpin, N., Wichers, H.J. and Pellon, G. (1998) Agaricus bisporus browning: a review. Mycol Res 102, 14591483.
  • Jolley, R.L., Robb, D.A. and Mason, H.S. (1969) The multiple forms of mushroom tyrosinase. association-dissociation phenomena. J Biol Chem 244, 15931599.
  • Kanda, K., Sato, T., Ishii, S., Enei, H. and Ejiri, S. (1996) Purification and properties of tyrosinase isozymes from gill of Lentinus edodes fruiting bodies. Biosci Biotechnol Biochem 60, 12731278.
  • Kanda, K., Aimi, T., Masumoto, S., Nakano, K., Kitamoto, Y. and Morinaga, T. (2003) NCBI Database. URL
  • Kane, J.B., Tompkins, R.G., Yarmush, M.L. and Burke, J.F. (1996) Burn dressings. In Biomaterials science: an introduction to materials in medicine ed. Ratner, B.D., Hoffman, A.S., Schoen, F.J. and Lemons, J.E. pp. 360370. San Diego: Academic Press.
  • Karam, J. and Nicell, J.A. (1997) Potential applications of enzymes in waste treatment. J Chem Technol Biotechnol 69, 141153.
  • Kato, T., Ito, S. and Fujita, K. (1986) Tyrosinase-catalysed binding of 3,4-dihydroxyphenylalanine with proteins through the sulfydryl group. Biochim Biophys Acta 881, 415421.
  • Klabunde, T., Eicken, C., Sacchettini, J.C. and Krebs, B. (1998) Crystal structure of a plant catechol oxidase containing a dicopper center. Nat Struct Biol 12, 10841090.
  • Krastanov, A. (2000) Removal of phenols from mixtures by co-immobilized laccase/tyrosinase and polyclar adsorption. J Ind Microbiol Biotechnol 24, 383388.
  • Kupper, U., Niedermann, D.M., Travaglini, G. and Lerch, K. (1989) Isolation and characterization of the tyrosinase gene from Neurospora crassa. J Biol Chem 264, 1725017258.
  • Kupper, U., Linden, M., Cao, K. and Lerch, K. (1990) Expression of tyrosinase in vegetative cultures of Neurospora crassa transformed with a metallothionein promoter/protyrosinase fusion gene. Curr Genetics 18, 331335.
  • Kurahashi, Y. and Pontzen, R. (1998) Carpropamid: A new melanin biosynthesis inhibitor. Pfanzenschutz-Nachrichten Bayer 51, 245256.
  • Langfelder, K., Glaser, P. and Brakhagee, A.A. (2000) NCBI Database. URL
  • Lavado, A., Olivares, C., Garcia-Borron, J.C. and Montoliu, L. (2005) Molecular basis of the extreme dilution mottled mouse mutation: a combination of coding and noncoding genomic alterations. J Biol Chem 280, 48174824.
  • Leatham, G.F., King, V. and Stahmann, M.A. (1980) In vitro protein polymerization by quinones or free-radicals generated by plant or fungal oxidative-enzymes. Phytopath 70, 11341140.
  • Lee, B.P., Dalsin, J.L. and Messersmith, P.B. (2002) Synthesis and gelation of DOPA-modified poly(ethylene glycol) hydrogels. Biomacromolecules 3, 10381047.
  • Lee, K.Y. and Mooney, D.J. (2001) Hydrogels for tissue engineering. Chem Rev 101, 18691879.
  • Lenhart, J.L., Chaubal, M.V., Payne, G.F. and Barbari, T.A. (1998) Enzymatic modification of chitosan by tyrosinase. In Enzymes in polymer synthesis ed. Gross, R.A., Kaplan, D.L. and Swift, G. pp. 188198. Washington DC: American Chemical Society Symposium Series 684.
  • Lerch, K. (1981) Copper monooxygenases: tyrosinase and dopamine-monooxygenase. In Metal ions in biological systems, Vol. 13 ed. Siegel, H. pp. 143186, Marcel Dekker, New York.
  • Lerch, K. (1982) Primary structure of tyrosinase from Neurospora crassa. II. Complete amino acid sequence and chemical structure of a tripeptide containing an unusual thioeher. J Biol Chem 257, 64146419.
  • Lerch, K. (1983) Neurospora tyrosinase: structural, spectroscopic and catalytic properties. Mol Cell Biochem 52, 125138.
  • Lerch, K. (1988) Protein and active-site structure of tyrosinase. In Advances in Pigment Cell Research. ed. Bagnara, J.T. pp. 8598, New York: Allan Liss Inc.
  • Lesage-Meessen, L., Navarro, D., Maunier, S., Sigoillot, J.-C., Lorquin, J., Delattre, M., Simon, J.-L., Asther, M. et al. (2001) Simple phenolic content in olive oil residues as a function of extraction systems. Food Chem 75, 501507.
  • Lowry, R.J. and Sussman, A.S. (1958) Ultrastructural changes during germination of ascospores of Neurospora tetrasperma. J Gen Microbiol 51, 403409.
  • Mallapragada, K. and Narasimhan, B. (1998) Drug delivery systems. In Handbook of biomaterials evaluation ed. Von Recum, A.F. pp. 415426, New York: Taylor and Francis.
  • Marbach, K. and Stahl, U. (1996) NCBI Database. URL
  • Matheis, M. and Whitaker, J.R. (1984) Modification of proteins by polyphenol oxidase and peroxidase and their products. J Food Biochem 8, 137162.
  • Matheis, M. and Whitaker, J.R. (1987) A review: enzymatic cross-linking of proteins applicable to foods. J Food Biochem 11, 309327.
  • Mayer, A.M. and Harel, E. (1979) Polyphenol oxidases in plants. Phytochem 31, 193215.
  • Montiel, A.M., Fernandez, F.J., Marcial, J., Soriano, J., Barrios-Gonzales, J. and Tomasini, A. (2004) A fungal phenoloxidase (tyrosinase) involved in pentachlorophenol degradation. Biotechnol Lett 26, 13531357.
  • Morinaga, T. (2003) NCBI Database. URL
  • Munjal, N. and Sawhney, S.K. (2002) Stability and properties of mushroom tyrosinase entrapped in alginate, polyacrylamide and gelatin gels. Enzyme Microb Technol 30, 613619.
  • Naish-Byfield, S., Cooksey, C.J. and Riley, P.A. (1994) Oxidation of monohydric phenol substrates by tyrosinase: effect of dithiothreitol on kinetics. Biochem J 304, 155162.
  • Nakamura, T., Sho, S. and Ogura, Y. (1966) On the purification and properties of mushroom tyrosinase. J Biochem 59, 481486.
  • Nakamura, M., Nakajima, T., Ohba, Y., Yamauchi, S., Lee, B.R. and Ichishima, E. (2000) Identification of copper ligands in Aspergillus oryzae tyrosinase by site-directed mutagenesis. Biochem J 360, 537545.
  • Newman, S.M. (1996) NCBI Database. URL
  • Paice, M.G., Bourbonnais, R., Reid, I.D., Archibald, F.S. and Jurasek, L. (1995) Oxidative bleaching enzymes: a review. J Pulp Paper Sci 21, 280284.
  • Peppas, N.A. and Sahlin, J.J. (1996) Hydrogels as mucoahesives and bioahesive materials: a review. Biomaterials 17, 15531561.
  • Raju, B.G.S., Rao, G.H. and Ayyanna, C. (1993) In Bioconversion of L-tyrosine to L-DOPA using Aspergillus oryzae, pp. 106110. Visakhapatnam India: CBS Publishers.
  • Rescigno, A., Sollai, F., Sanjust, E., Rinaldi, A.C., Curreli, N. and Rinaldi, A. (1997) Diafiltration in the presence of ascorbate in the purification of mushroom tyrosinase. Phytochem 46, 2122.
  • Riley, P.A. (1997) Molecules in focus: Melanin. Int J Biochem Cell Biol 29, 12351239.
  • Robb, D.A. and Gutteridge, S. (1981) The polypeptide composition of two fungal tyrosinases. Phytochem 20, 14811485.
  • Robb, D.A. (1984) Tyrosinase. In Copper proteins and copper enzymes ed. Lontie, R., pp. 207241. Boca Raton: CRC press.
  • Rodriguez-Lopez, J.N., Tudela, J., Varon, R., Garcia-Carmonas, F. and Garcia-Canovas, F. (1992) Analysis of a kinetic model for melanin biosynthesis pathway. J Biol Chem 267, 38013810.
  • Sanchez-Ferrer, A., Rodriguez-Lopez, J.N., Garcia-Canovas, F. and Garcia-Carmona, F. (1995) Tyrosinase: A comprehensive review of its mechanism. Biochim Biophys Acta 1247, 111.
  • Sâto, M. (1969) The conversion by phenolase of p-coumaric acid to caffeic acid with special reference to the role of ascorbic acid. Phytochem 8, 353362.
  • Sato, T. (2001) NCBI Database. URL
  • Schulte, U., Align, V., Hoheisel, J., Brandt, P., Fartmann, B., Holland, R., Nyakatura, G., Mewes, H.W. et al. (2003) NCBI Database. URL
  • Seetharam, G. and Saville, B.A. (2002) L-DOPA production from tyrosinase immobilized on zeolite. Enzyme Microb Technol 31, 747753.
  • Seo, S.-Y., Sharma, V.K. and Sharma, N. (2003) Mushroom tyrosinase: Recent prospects. J Agric Food Chem 51, 28372853.
  • Serra, B., Morales, M.D., Reviejo, A.J., Hall, E.H. and Pingarron, J.M. (2005) Rapid and highly sensitive electrochemical determination of alkaline phosphatase using a composite tyrosinase biosensor. Anal Biochem 336, 289294.
  • Singh, H. (1999) Mechanism of oxidation of L-tyrosine by fungal tyrosinases. J Chem 172, 83.
  • Soler-Rivas, C., Arpin, N., Olivier, J.M. and Wichers, H.J. (1997) Activation of tyrosinase in Agaricus bisporus strains following infection by Pseudomonas tolaasii or treatment with a tolaasin-containing preparation. Mycol Res 101, 375382.
  • Soler-Rivas, C., Jolivet, S., Arpin, N., Olivier, J.M. and Wihers, H.J. (1999) Biochemical and physiological aspects of brown blotch disease of Agaricus bisporus. FEMS Microbiol Rev 23, 591614.
  • Solomon, E.I. and Lowery, M.D. (1993) Electronic structure contributions to function in bioinorganic chemistry. Science 259, 15751581.
  • Solomon, E.I., Sundaram, U.M. and Machonkin, T.E. (1996) Multicopper oxidases and oxygenases. Chem Rev 96, 25632605.
  • Strausberg, R.L. (2002) NCBI Database. URL
  • Strothkamp, K.G., Jolley, R.L. and Mason, H.S. (1976) Quaternary structure of mushroom tyrosinase. Biochem Biophys Res Commun 70, 519524.
  • Takasaki, S. and Kawakishi, S. (1997) Formation of protein-bound 3,4-dihydroxyphenylalanine and 5-S-cysteinyl-3,4-dihydroxyphenylalanine as new cross-linkers in gluten. J Agric Food Chem 45, 34723475.
  • Thalman, C.R. and Lötzbeyer, T. (2002) Enzymatic cross-linking of proteins with tyrosinase. Eur Food Res Technol 214, 276281.
  • Van den Hondel, C.A., Punt, P.J. and van Gorcom, R.F. (1992) Production of extracellular proteins by the filamentous fungus Aspergillus. Anton Leeuwenhoek Int J Gen Microbiol 61, 153160.
  • Van Gelder, C.W.G., Flurkey, W.H. and Wichers, H.J. (1997) Sequence and structural features of plant and fungal tyrosinases. Phytochem 45, 13091323.
  • Van Leeuwen, J. and Wichers, H.J. (1999) Tyrosinase activity and isoform composition in seperate tissues during development of Agaricus bisporus fruit bodies. Mycol Res 103, 413418.
  • Wada, S., Ichikawa, H. and Tatsumi, K. (1992) Removal of phenols with tyrosinase immobilized on magnetite. Water Sci Technol 26, 20572059.
  • Whitaker, J.R. (1995) Polyphenol oxidase. In Food Enzymes Structure and Mechanism ed. Wong, D.W.S. pp. 271307. New York: Chapman Hall.
  • Wichers, H.J., Gerritsen, Y.A.M. and Chapelon, C.G.J. (1996) Tyrosinase isoforms from the fruitbodies of Agaricus bisporus. Phytochem 43, 333337.
  • Wichers, H.J., Recourt, K., Hendriks, M., Ebbelaar, C.E.M., Biancone, G., Hoeberichts, F.A., Mooibroek, H. and Soler-Rivas, C. (2003) Cloning, expression and characterization of two tyrosinase cDNAs from Agaricus bisporus. Appl Microbiol Biotechnol 61, 336341.
  • Woolery, G.L., Powers, L., Winkler, M., Solomon, E.I., Lerch, K. and Spiro, T.G. (1984) Extended X-Ray absorption fine structure study of the coupled binuclear copper active site of tyrosinase from Neurospora crassa. Biochem Biophys Acta 788, 155161.
  • Yamazaki, S.-I., Morioka, C. and Itoh, S. (2004) Kinetic evaluation of catalase and peroxygenase activities of tyrosinase. Biochem 43, 1154611553.
  • Zhang, X., van Leeuwen, J., Wichers, H.J. and Flurkey, W.H. (1999) Characterization of tyrosinase from the cap flesh of Portabella mushrooms. J Agric Food Chem 47, 374378.