Abstract

Laccases of fungi attract considerable attention due to their possible involvement in the transformation of a wide variety of phenolic compounds including the polymeric lignin and humic substances. So far, more than a 100 enzymes have been purified from fungal cultures and characterized in terms of their biochemical and catalytic properties. Most ligninolytic fungal species produce constitutively at least one laccase isoenzyme and laccases are also dominant among ligninolytic enzymes in the soil environment. The fact that they only require molecular oxygen for catalysis makes them suitable for biotechnological applications for the transformation or immobilization of xenobiotic compounds.

Introduction

Laccase is one of the very few enzymes that have been studied since the end of 19th century. It was first demonstrated in the exudates of Rhus vernicifera , the Japanese lacquer tree ( Yoshida, 1883 ). A few years later it was also demonstrated in fungi ( Bertrand, 1896 ). Although known for a long time, laccases attracted considerable attention only after the beginning of studies of enzymatic degradation of wood by white-rot wood-rotting fungi.

Laccase (benzenediol: oxygen oxidoreductase, EC 1.10.3.2) belongs to a group of polyphenol oxidases containing copper atoms in the catalytic centre and usually called multicopper oxidases. Other members of this group are the mammalian plasma protein ceruloplasmin and ascorbate oxidases of plants. Laccases typically contain three types of copper, one of which gives it its characteristic blue colour. Similar enzymes lacking the Cu atom responsible for the blue colour are called ‘yellow’ or ‘white’ laccases, but several authors do not regard them as true laccases. Laccases catalyze the reduction of oxygen to water accompanied by the oxidation of a substrate, typically a p -dihydroxy phenol or another phenolic compound. It is difficult to define laccase by its reducing substrate due to its very broad substrate range, which varies from one laccase to another and overlaps with the substrate range of another enzyme–the monophenol mono-oxygenase tyrosinase (EC 1.14.18.1). Although laccase was also called diphenol oxidase, monophenols like 2,6-dimethoxyphenol or guaiacol are often better substrates than diphenols, e.g. catechol or hydroquinone. Syringaldazine [N,N′-bis(3,5-dimethoxy-4-hydroxybenzylidene hydrazine)] is often considered to be a unique laccase substrate ( Harkin, 1974 ) as long as hydrogen peroxide is avoided in the reaction, as this compound is also oxidized by peroxidases. Laccase is thus an oxidase that oxidizes polyphenols, methoxy-substituted phenols, aromatic diamines and a range of other compounds but does not oxidize tyrosine as tyrosinases do.

Laccases are typically found in plants and fungi. Plant laccases participate in the radical-based mechanisms of lignin polymer formation ( Sterjiades, 1992 ; Liu, 1994 ; Boudet, 2000 ; Ranocha, 2002 ; Hoopes & Dean, 2004 ), whereas in fungi laccases probably have more roles including morphogenesis, fungal plant-pathogen/host interaction, stress defence and lignin degradation ( Thurston, 1994 ). Although there are also some reports about laccase activity in bacteria ( Alexandre & Zhulin, 2000 ; Martins, 2002 ; Claus, 2003 ; Givaudan, 2004 ), it does not seem probable that laccases are common enzymes from certain prokaryotic groups. Bacterial laccase-like proteins are intracellular or periplasmic proteins ( Claus, 2003 ). Probably the best characterized bacterial laccase is that isolated from Sinorhizobium meliloti , which has been described as a 45-kDa periplasmic protein with isoelectric point at pH 6.2 and the ability to oxidize syringaldazine ( Rosconi, 2005 ).

The chemistry, function and biotechnological use of laccases have recently been reviewed. The basic aspects of laccase structure and function were reviewed by ( Thurston, 1994 ), ( Leonowicz, 2001 ) focused on the functional properties of fungal laccases and their involvement in lignin transformation and ( Mayer & Staples, 2002 ) dealt with the latest results about the roles of laccases in vivo and its biotechnological applications. The physico-chemical properties of multicopper oxidases have been comprehensively reviewed by ( Solomon, 1996 , 2001 ). An overview of technological applications of oxidases including laccase was published by ( Durán & Esposito, 2000 ) and ( Durán, 2002 ) reviewed the literature concerning the use of immobilized laccases and tyrosinases.

The main aim of this work is to summarize the rich literature data that has accumulated in the last years from the studies of authors purifying the enzyme from different fungal sources. In addition to a generally low substrate specificity, laccase has other properties that make this enzyme potentially useful for biotechnological application. These include the fact that laccase, unlike peroxidases, does not need the addition or synthesis of a low molecular weight cofactor like hydrogen peroxide, as its cosubstrate – oxygen – is usually present in its environment. Most laccases are extracellular enzymes, making the purification procedures very easy and laccases generally exhibit a considerable level of stability in the extracellular environment. The inducible expression of the enzyme in most fungal species also contributes to the easy applicability in biotechnological processes. This review should help to define the common general characteristics of fungal laccases as well as the unique properties of individual enzymes with a potential biotechnological use and contribute to the discussion on the occurrence and significance of laccase in the natural environment.

Occurrence in fungi

Laccase activity has been demonstrated in many fungal species and the enzyme has already been purified from tens of species. This might lead to the conclusion that laccases are extracellular enzymes generally present in most fungal species. However, this conclusion is misleading as there are many taxonomic or physiological groups of fungi that typically do not produce significant amounts of laccase or where laccase is only produced by a few species. Laccase production has never been demonstrated in lower fungi, i.e. Zygomycetes and Chytridiomycetes ; however, this aspect of these groups has not as yet been studied in detail.

There are many records of laccase production by ascomycetes. Laccase was purified from phytopathogenic ascomycetes such as Gaeumannomyces graminis ( Edens, 1999 ), Magnaporthe grisea ( Iyer & Chattoo, 2003 ) and Ophiostoma novo-ulmi ( Binz & Canevascini, 1997 ), as well as from Mauginella ( Palonen, 2003 ), Melanocarpus albomyces ( Kiiskinen, 2002 ), Monocillium indicum ( Thakker, 1992 ), Neurospora crassa ( Froehner & Eriksson, 1974 ) and Podospora anserina ( Molitoris & Esser, 1970 ).

It is difficult to say how many ascomycete species produce laccases as no systematic search has been undertaken. In addition to plant pathogenic species, laccase production was also reported for some soil ascomycete species from the genera Aspergillus, Curvularia and Penicillium ( Banerjee & Vohra, 1991 ; Rodriguez, 1996 ; Scherer & Fischer, 1998 ), as well as some freshwater ascomycetes ( Abdel-Raheem & Shearer, 2002 ; Junghanns, 2005 ). However, the enzyme from Aspergillus nidulans was unable to oxidize syringaldazine ( Scherer & Fischer, 1998 ) and the enzymes from Penicillium spp. were not tested with this substrate, leaving it unclear if they are true laccases.

Wood-degrading ascomycetes like the soft-rotter Trichoderma and the ligninolytic Bothryosphaeria are ecologically closely related to the wood-rotting basidiomycetes producing laccase. Laccase activity has been described in both genera, but whereas Bothryosphaeria produces constitutively a dimethoxyphenol-oxidizing enzyme that is probably a true laccase ( Vasconcelos, 2000 ), only some strains of Trichoderma exhibit a low level of production of a syringaldazine-oxidizing enzyme ( Assavanig, 1992 ), mainly associated with spores, which may act in the morphogenesis of this fungus ( Assavanig, 1992 ; Holker, 2002 ). Although no enzyme purification has been reported so far, laccases are probably also produced by wood-rotting xylariaceous ascomycetes. Among the 20 strains tested, genes with sequences similar to basidiomycete laccases were detected in three strains, all of them Xylaria sp. Two strains of Xylaria sp. and one of Xylaria hypoxylon exhibited syringaldazine oxidation ( Pointing S, 2005 ). In complex liquid media, the fungi X. hypoxylon and Xylaria polymorpha produced appreciable titres of an ABTS oxidizing enzyme, also present in the extracts of colonized beech wood chips ( Liers, 2005 ). Furthermore, ascomycete species closely related to wood-degrading fungi which participate in the decay of dead plant biomass in salt marshes have been shown to contain laccase genes and to oxidize syringaldazine ( Lyons, 2003 ).

Yeasts are a physiologically specific group of both ascomycetes and basidiomycetes. Until now, laccase was only purified from the human pathogen Cryptococcus ( Filobasidiella ) neoformans . This basidiomycete yeast produces a true laccase capable of oxidation of phenols and aminophenols and unable to oxidize tyrosine ( Williamson, 1994 ). The enzyme is tightly bound to the cell wall and contributes to the resistance to fungicides ( Zhu, 2001 ; Ikeda, 2003 ). A homologous gene has also been demonstrated in Cryptococcus podzolicus but not in other heterobasidiomycetous yeasts tested ( Petter, 2001 ) and there are some records of low laccase-like activity in some yeast species isolated from decayed wood ( Jimenez, 1991 ). The production of laccase was not demonstrated in ascomycetous yeasts, but the plasma membrane-bound multicopper oxidase Fet3p from Saccharomyces cerevisiae shows both sequence and structural homology with fungal laccase. Although more closely related to ceruloplasmin, Fet3p has spectroscopic properties nearly identical to fungal laccase, the configuration of their type-1 Cu sites is very similar and both enzymes are able to oxidize Cu + ( Machonkin, 2001 ; Stoj & Kosman, 2003 ).

Among physiological groups of fungi, laccases are typical for the wood-rotting basidiomycetes causing white-rot and a related group of litter-decomposing saprotrophic fungi, i.e. the species causing lignin degradation. Almost all species of white-rot fungi were reported to produce laccase to varying degrees ( Hatakka, 2001 ), and the enzyme has been purified from many species ( Table 1 ). In the case of Pycnoporus cinnabarinus , laccase was described as the only ligninolytic enzyme produced by this species that was capable of lignin degradation ( Eggert, 1996 ). Although the group of brown-rot fungi is typical for its inability to decompose lignin, there have been several attempts to detect laccases in the members of this physiological group. A DNA sequence with a relatively high similarity to that of laccases of white-rot fungi was detected in Gloeophyllum trabeum . Oxidation of ABTS (2,2′-azinobis(3-ethylbenzathiazoline-6-sulfonic acid)) as an indirect indication of oxidative activity was also found in this fungus as well as in a few other brown-rot species ( D'Souza, 1996 ). Although no laccase protein has been purified from any brown-rot species, the oxidation of syringaldazine – a reliable indication of laccase presence – has recently been detected in the brown-rot fungus Coniophora puteana ( Lee, 2004 ) and oxidation of ABTS was reported in Laetiporus sulphureus ( Schlosser & Höfer, 2002 ). The occurrence and role of laccases in brown-rot decay of wood is still unclear but it seems to be rare.

Table 1

Characteristics of laccases purified from fungi

Species MW (kDa) pI pH optimum Km (μM) Temperature optimum (°C) Reference 
ABTS DMP GUA SYR ABTS DMP GUA SYR 
Agaricus bisporus 96    5.6       Wood (1980) 
Agaricus bisporus 65           Perry (1993) 
Agaricus blazei 66 4.0 2.0 5.5  6.0 63 1026 4307  Ullrich (2005) 
Agrocybe praecox 66 4.0          Steffen (2002) 
Albatrella dispansus 62  4.0        70 Wang & Ng (2004b) 
Armillaria mellea Lac I  59 4.1  3.5    178    Rehman & Thurston (1992) 
Armillaria mellea Lac II             Billal & Thurston (1996) 
Armillaria mellea 80 3.1          Curir (1997) 
Aspergillus nidulans II  80   6.5       55 Scherer & Fischer (1998) 
Botrytis cinerea 74 4.0  3.5    100   57 Slomczynski (1995) 
Cantharellus cibarius 92  4.0        50 Ng & Wang (2004) 
Ceriporiopsis subvermispora L1 71 3.4 3.0 4.0 3.0  30 2900 1600   Fukushima & Kirk (1995) ; Wang & Ng (2004b) 
Ceriporiopsis subvermispora L2  68 4.8 3.0 4.0 5.0  20 7700 440   Fukushima & Kirk, (1995) ; Wang & Ng (2004b) 
Cerrena maxima 57–67 3.5       160–300  50 Koroleva (2001) ; Shleev (2004) 
Cerrena unicolor 66 4.0          Bekker (1990) 
Cerrena unicolor 58           Kim (2002) 
Chaetomium termophilum 77 5.1     190 96 400 34 60 Chefetz (1998) 
Chalara paradoxa 67  4.5 4.5 6.5 6.5 770 14 720 10 230 3400  Robles (2002) 
Colletotrichum graminicola 85     6.0    214  Anderson & Nicholson (1996) 
Coniothyrium minitans 74 4.0  3.5    100   60 Dahiya (1998) 
Coprinus cinereus 58 4.0 4.0   6.5 26    60–70 Schneider (1999) 
Coprinus friesii 60 3.5 5.0 8.0   41     Heinzkill (1998) 
Coriolopsis fulvocinnerea 54–65 3.5       70–90   Shleev (2004) ; Smirnov (2001) 
Coriolopsis gallica 84 4.2–4.3 3.0        70 Calvo (1998) 
Coriolopsis rigida I  66 3.9 2.5 3.0   12 328    Saparrat (2002) 
Coriolopsis rigida II  66 3.9 2.5 3.0   11 348    Saparrat (2002) 
Coriolus hirsutus 55 4.0          Koroljova-Skorobogat'ko (1998) 
Coriolus hirsutus 78 4.2        45 Lee & Shin (1999) 
Coriolus maxima 57           Smirnov (2001) 
Coriolus zonatus 60 4.6         55 Koroljova (1999) 
Cryptococcus neoformans 77           Williamson (1994) 
Cyathus stercoreus 70 3.5 4.8         Sethuraman (1999) 
Daedalea quercina 69 3.0 2.0 4.0 4.5 7.0 38 48 93 131 70, 55 Baldrian (2004) 
Dichomitus squalens c1  66 3.5  3.0        Perie (1998) 
Dichomitus squalens c2  66 3.6  3.0        Perie (1998) 
Fomes fomentarius           52 Rogalski (1991) 
Ganoderma lucidum 67         25 Lalitha Kumari & Sirsi (1972) ; Ko (2001) 
Ganoderma tsugae            Eller (1998) 
Gaeumannomyces graminis 190 5.6  4.5    26 510   Edens (1999) 
Hericium echinaceum 63  5.0        50 Wang & Ng (2004c) 
Junghuhnia separabilima 58–62 3.4–3.6          Vares (1992) 
Lactarius piperatus 67           Iwasaki (1967) 
Lentinula edodes Lcc1  72 3.0 4.0 4.0 4.0  108 557 917  40 Nagai (2002) 
Lentinus edodes 65 3.0          Kofujita (1991) 
Magnaporthe grisea 70     6.0    118 30 Iyer & Chattoo (2003) 
Marasmius quercophilus 60 4.0–4.4    5.0     80 Farnet (2000) 
Marasmius quercophilus 60 4.8–5.1    5.0     80 Farnet (2000) 
Marasmius quercophilus 65 3.6    4.5    75 Dedeyan (2000) 
Marasmius quercophilus 65  2.6   6.2   50 80 Farnet, (2002 , 2004) 
Marasmius quercophilus 60  4.0   4.5 113   4.2 80 Farnet (2004) 
Mauginiella sp.  63 4.8–6.4 2.4 3.5 4.0       Palonen (2003) 
Melanocarpus albomyces 80 4.0 3.5  5.0–7.5 6.0–7.0     65 Kiiskinen (2002) 
Monocillium indicum 100           Thakker (1992) 
Myrothecium verrucaria 62           Sulistyaningdyah (2004) 
Neurospora crassa 64           Froehner & Eriksson (1974) 
Ophiostoma novo-ulmi 79 5.1 2.8  6.0 6.0      Binz & Canevascini (1997) 
Panaeolus papilionaceus 60  3.0 8.0   51     Heinzkill (1998) 
Panaeolus sphinctrinus 60  3.0 7.0   32     Heinzkill (1998) 
Panus tigrinus 64 2.9–3.0          Maltseva (1991) 
Panus tigrinus 63           Leontievsky (1997) 
Phanerochaete flavido-alba 94  3.0        30 Perez (1996) 
Phanerochaete chrysosporium 47           Srinivasan (1995) 
Phellinus noxius 70           Geiger (1986) 
Phellinus ribis 152  5.0 4.0–6.0  6.0 207 38  11  Min (2001) 
Phlebia radiata 64 3.5          Vares (1995) 
Phlebia tremellosa 64           Vares (1994) 
Pholiota mutabilis            Leonowicz & Malinowska (1982) 
Physisporinus rivulosus Lacc 1  66 3.3 2.5 3.0 3.5 3.5      Hakala (2005) 
Physisporinus rivulosus Lacc 2  67 3.3 2.5 3.0 3.5 3.5      Hakala (2005) 
Physisporinus rivulosus Lacc 3  68 3.2 2.5 3.0 3.5 3.5      Hakala (2005) 
Physisporinus rivulosus Lacc 4  68 3.1 2.5 3.0 3.5 3.5      Hakala (2005) 
Pleurotus eryngii I  65 4.1 4.5     1400 7600  55 Munoz (1997) 
Pleurotus eryngii II  61 4.2 4.5     400 8000  55 Munoz (1997) 
Pleurotus florida 77 4.1       30 000   Das (2000) 
Pleurotus ostreatus 67 3.6    5.8     50 Hublik & Schinner (2000) 
Pleurotus ostreatus POXA1b  62 6.9 3.0 4.5  6.0 370 260  220  Giardina (1999) 
Pleurotus ostreatus POXA1w  61 6.7 3.0 3.0–5.0 NA 6.0 90 2100 NA 130 45–65 Palmieri (1997) 
Pleurotus ostreatus POXA2  67 4.0 3.0 6.5 6.0 6.0 120 740 3100 140 25–35 Palmieri (1997) 
Pleurotus ostreatus POXA3a  83–85 4.1 3.6 5.5 6.2  70 14 000  36 35 Palmieri (2003) 
Pleurotus ostreatus POXA3b  83–85 4.3 3.6 5.5 6.2  74 8800  79 35 Palmieri (2003) 
Pleurotus ostreatus POXC  59 2.9 3.0 3.0–5.0 6.0 6.0 280 230 1200 20 50–60 Palmieri (1993 , 1997 ); Sannia (1986) 
Pleurotus pulmonarius Lcc2  46  4.0–5.5  6.0–8.0 6.2–6.5 210  550 12 50 De Souza & Peralta (2003) 
Pleurotus sajor-caju IV  55 3.6 2.1    92     Lo (2001) 
Podospora anserine 383           Molitoris & Esser (1970) ; Durrens (1981) 
Polyporus anceps      5.0–5.5      Petroski (1980) 
Polyporus anisoporus 58 3.4          Vaitkyavichyus (1984) 
Polyporus pinsitus 66  3.0 5.0   22     Heinzkill (1998) 
Pycnoporus cinnabarinus 63 3.0   4.0–4.5 4.4–5.0   330 30  Schliephake (2000) 
Pycnoporus cinnabarinus 81 3.7   4.0       Eggert (1996) 
Pycnoporus coccineus 70           Oda (1991) 
Rhizoctonia solani 4  170           Iwasaki (1967) 
Rigidoporus lignosus B  55 3.7 3.0 6.2   80 480    Bonomo (1998) 
Rigidoporus lignosus S  60 3.1 3.0 6.2   49 108    Bonomo (1998) 
Russula delica 63           Matsubara & Iwasaki (1972) 
Schizophyllum commune 62–64           De Vries (1986) 
Sclerotium rolfsii SRL1  55 5.2 2.4        62 Ryan (2003) 
Sclerotium rolfsii SRL2  86           Ryan (2003) 
Stropharia coronilla 67 4.4          Steffen (2002) 
Stropharia rugosoannulata 66  2.5 3.5        Schlosser & Höfer (2002) 
Thelephora terrestris 66  3.4  4.8 5.0 16  121 45 Kanunfre & Zancan (1998) 
Trametes gallica Lac I  60 3.1 2.2 3.0 4.0  12 420 405  70 Dong & Zhang (2004) 
Trametes gallica Lac II  60 3.0 2.2 3.0 4.0  410 400  70 Dong & Zhang (2004) 
Trametes hirsute 64–68 3.7–4.0       63   Shleev (2004) ; Vares & Hatakka (1997) 
Trametes multicolor II  63 3.0          Leitner (2002) 
Trametes ochracea 64 4.7       90    ( Shleev (2004) 
Trametes pubescens LAP 2  65 2.6     14 72 360  Galhaup (2002) 
Trametes sanguinea 62 3.5          Nishizawa (1995) 
Trametes trogii 70 3.3; 3.6     30 410    Garzillo (1998) 
Trametes versicolor 68  2.5 3.5  4.0 37 15   55 Rogalski (1990) ; Höfer & Schlosser (1999) 
Trametes villosa 1  63 3.5 2.7   5.0–5.5      Yaver (1996) 
Trametes villosa 3  63 6.0–6.5 2.7   5.0–5.5      Yaver (1996) 
Trametes sp. AH28-2 A  62 4.2   4.5  25 25 420  50 Xiao (2003) 
Trichoderma sp.  71           Assavanig (1992) 
Tricholoma giganteum 43  4.0        70 Wang & Ng (2004a) 
Volvariella volvacea 58 3.7 3.0 4.6  5.6 30 570  10 45 Chen (2004) 
Species MW (kDa) pI pH optimum Km (μM) Temperature optimum (°C) Reference 
ABTS DMP GUA SYR ABTS DMP GUA SYR 
Agaricus bisporus 96    5.6       Wood (1980) 
Agaricus bisporus 65           Perry (1993) 
Agaricus blazei 66 4.0 2.0 5.5  6.0 63 1026 4307  Ullrich (2005) 
Agrocybe praecox 66 4.0          Steffen (2002) 
Albatrella dispansus 62  4.0        70 Wang & Ng (2004b) 
Armillaria mellea Lac I  59 4.1  3.5    178    Rehman & Thurston (1992) 
Armillaria mellea Lac II             Billal & Thurston (1996) 
Armillaria mellea 80 3.1          Curir (1997) 
Aspergillus nidulans II  80   6.5       55 Scherer & Fischer (1998) 
Botrytis cinerea 74 4.0  3.5    100   57 Slomczynski (1995) 
Cantharellus cibarius 92  4.0        50 Ng & Wang (2004) 
Ceriporiopsis subvermispora L1 71 3.4 3.0 4.0 3.0  30 2900 1600   Fukushima & Kirk (1995) ; Wang & Ng (2004b) 
Ceriporiopsis subvermispora L2  68 4.8 3.0 4.0 5.0  20 7700 440   Fukushima & Kirk, (1995) ; Wang & Ng (2004b) 
Cerrena maxima 57–67 3.5       160–300  50 Koroleva (2001) ; Shleev (2004) 
Cerrena unicolor 66 4.0          Bekker (1990) 
Cerrena unicolor 58           Kim (2002) 
Chaetomium termophilum 77 5.1     190 96 400 34 60 Chefetz (1998) 
Chalara paradoxa 67  4.5 4.5 6.5 6.5 770 14 720 10 230 3400  Robles (2002) 
Colletotrichum graminicola 85     6.0    214  Anderson & Nicholson (1996) 
Coniothyrium minitans 74 4.0  3.5    100   60 Dahiya (1998) 
Coprinus cinereus 58 4.0 4.0   6.5 26    60–70 Schneider (1999) 
Coprinus friesii 60 3.5 5.0 8.0   41     Heinzkill (1998) 
Coriolopsis fulvocinnerea 54–65 3.5       70–90   Shleev (2004) ; Smirnov (2001) 
Coriolopsis gallica 84 4.2–4.3 3.0        70 Calvo (1998) 
Coriolopsis rigida I  66 3.9 2.5 3.0   12 328    Saparrat (2002) 
Coriolopsis rigida II  66 3.9 2.5 3.0   11 348    Saparrat (2002) 
Coriolus hirsutus 55 4.0          Koroljova-Skorobogat'ko (1998) 
Coriolus hirsutus 78 4.2        45 Lee & Shin (1999) 
Coriolus maxima 57           Smirnov (2001) 
Coriolus zonatus 60 4.6         55 Koroljova (1999) 
Cryptococcus neoformans 77           Williamson (1994) 
Cyathus stercoreus 70 3.5 4.8         Sethuraman (1999) 
Daedalea quercina 69 3.0 2.0 4.0 4.5 7.0 38 48 93 131 70, 55 Baldrian (2004) 
Dichomitus squalens c1  66 3.5  3.0        Perie (1998) 
Dichomitus squalens c2  66 3.6  3.0        Perie (1998) 
Fomes fomentarius           52 Rogalski (1991) 
Ganoderma lucidum 67         25 Lalitha Kumari & Sirsi (1972) ; Ko (2001) 
Ganoderma tsugae            Eller (1998) 
Gaeumannomyces graminis 190 5.6  4.5    26 510   Edens (1999) 
Hericium echinaceum 63  5.0        50 Wang & Ng (2004c) 
Junghuhnia separabilima 58–62 3.4–3.6          Vares (1992) 
Lactarius piperatus 67           Iwasaki (1967) 
Lentinula edodes Lcc1  72 3.0 4.0 4.0 4.0  108 557 917  40 Nagai (2002) 
Lentinus edodes 65 3.0          Kofujita (1991) 
Magnaporthe grisea 70     6.0    118 30 Iyer & Chattoo (2003) 
Marasmius quercophilus 60 4.0–4.4    5.0     80 Farnet (2000) 
Marasmius quercophilus 60 4.8–5.1    5.0     80 Farnet (2000) 
Marasmius quercophilus 65 3.6    4.5    75 Dedeyan (2000) 
Marasmius quercophilus 65  2.6   6.2   50 80 Farnet, (2002 , 2004) 
Marasmius quercophilus 60  4.0   4.5 113   4.2 80 Farnet (2004) 
Mauginiella sp.  63 4.8–6.4 2.4 3.5 4.0       Palonen (2003) 
Melanocarpus albomyces 80 4.0 3.5  5.0–7.5 6.0–7.0     65 Kiiskinen (2002) 
Monocillium indicum 100           Thakker (1992) 
Myrothecium verrucaria 62           Sulistyaningdyah (2004) 
Neurospora crassa 64           Froehner & Eriksson (1974) 
Ophiostoma novo-ulmi 79 5.1 2.8  6.0 6.0      Binz & Canevascini (1997) 
Panaeolus papilionaceus 60  3.0 8.0   51     Heinzkill (1998) 
Panaeolus sphinctrinus 60  3.0 7.0   32     Heinzkill (1998) 
Panus tigrinus 64 2.9–3.0          Maltseva (1991) 
Panus tigrinus 63           Leontievsky (1997) 
Phanerochaete flavido-alba 94  3.0        30 Perez (1996) 
Phanerochaete chrysosporium 47           Srinivasan (1995) 
Phellinus noxius 70           Geiger (1986) 
Phellinus ribis 152  5.0 4.0–6.0  6.0 207 38  11  Min (2001) 
Phlebia radiata 64 3.5          Vares (1995) 
Phlebia tremellosa 64           Vares (1994) 
Pholiota mutabilis            Leonowicz & Malinowska (1982) 
Physisporinus rivulosus Lacc 1  66 3.3 2.5 3.0 3.5 3.5      Hakala (2005) 
Physisporinus rivulosus Lacc 2  67 3.3 2.5 3.0 3.5 3.5      Hakala (2005) 
Physisporinus rivulosus Lacc 3  68 3.2 2.5 3.0 3.5 3.5      Hakala (2005) 
Physisporinus rivulosus Lacc 4  68 3.1 2.5 3.0 3.5 3.5      Hakala (2005) 
Pleurotus eryngii I  65 4.1 4.5     1400 7600  55 Munoz (1997) 
Pleurotus eryngii II  61 4.2 4.5     400 8000  55 Munoz (1997) 
Pleurotus florida 77 4.1       30 000   Das (2000) 
Pleurotus ostreatus 67 3.6    5.8     50 Hublik & Schinner (2000) 
Pleurotus ostreatus POXA1b  62 6.9 3.0 4.5  6.0 370 260  220  Giardina (1999) 
Pleurotus ostreatus POXA1w  61 6.7 3.0 3.0–5.0 NA 6.0 90 2100 NA 130 45–65 Palmieri (1997) 
Pleurotus ostreatus POXA2  67 4.0 3.0 6.5 6.0 6.0 120 740 3100 140 25–35 Palmieri (1997) 
Pleurotus ostreatus POXA3a  83–85 4.1 3.6 5.5 6.2  70 14 000  36 35 Palmieri (2003) 
Pleurotus ostreatus POXA3b  83–85 4.3 3.6 5.5 6.2  74 8800  79 35 Palmieri (2003) 
Pleurotus ostreatus POXC  59 2.9 3.0 3.0–5.0 6.0 6.0 280 230 1200 20 50–60 Palmieri (1993 , 1997 ); Sannia (1986) 
Pleurotus pulmonarius Lcc2  46  4.0–5.5  6.0–8.0 6.2–6.5 210  550 12 50 De Souza & Peralta (2003) 
Pleurotus sajor-caju IV  55 3.6 2.1    92     Lo (2001) 
Podospora anserine 383           Molitoris & Esser (1970) ; Durrens (1981) 
Polyporus anceps      5.0–5.5      Petroski (1980) 
Polyporus anisoporus 58 3.4          Vaitkyavichyus (1984) 
Polyporus pinsitus 66  3.0 5.0   22     Heinzkill (1998) 
Pycnoporus cinnabarinus 63 3.0   4.0–4.5 4.4–5.0   330 30  Schliephake (2000) 
Pycnoporus cinnabarinus 81 3.7   4.0       Eggert (1996) 
Pycnoporus coccineus 70           Oda (1991) 
Rhizoctonia solani 4  170           Iwasaki (1967) 
Rigidoporus lignosus B  55 3.7 3.0 6.2   80 480    Bonomo (1998) 
Rigidoporus lignosus S  60 3.1 3.0 6.2   49 108    Bonomo (1998) 
Russula delica 63           Matsubara & Iwasaki (1972) 
Schizophyllum commune 62–64           De Vries (1986) 
Sclerotium rolfsii SRL1  55 5.2 2.4        62 Ryan (2003) 
Sclerotium rolfsii SRL2  86           Ryan (2003) 
Stropharia coronilla 67 4.4          Steffen (2002) 
Stropharia rugosoannulata 66  2.5 3.5        Schlosser & Höfer (2002) 
Thelephora terrestris 66  3.4  4.8 5.0 16  121 45 Kanunfre & Zancan (1998) 
Trametes gallica Lac I  60 3.1 2.2 3.0 4.0  12 420 405  70 Dong & Zhang (2004) 
Trametes gallica Lac II  60 3.0 2.2 3.0 4.0  410 400  70 Dong & Zhang (2004) 
Trametes hirsute 64–68 3.7–4.0       63   Shleev (2004) ; Vares & Hatakka (1997) 
Trametes multicolor II  63 3.0          Leitner (2002) 
Trametes ochracea 64 4.7       90    ( Shleev (2004) 
Trametes pubescens LAP 2  65 2.6     14 72 360  Galhaup (2002) 
Trametes sanguinea 62 3.5          Nishizawa (1995) 
Trametes trogii 70 3.3; 3.6     30 410    Garzillo (1998) 
Trametes versicolor 68  2.5 3.5  4.0 37 15   55 Rogalski (1990) ; Höfer & Schlosser (1999) 
Trametes villosa 1  63 3.5 2.7   5.0–5.5      Yaver (1996) 
Trametes villosa 3  63 6.0–6.5 2.7   5.0–5.5      Yaver (1996) 
Trametes sp. AH28-2 A  62 4.2   4.5  25 25 420  50 Xiao (2003) 
Trichoderma sp.  71           Assavanig (1992) 
Tricholoma giganteum 43  4.0        70 Wang & Ng (2004a) 
Volvariella volvacea 58 3.7 3.0 4.6  5.6 30 570  10 45 Chen (2004) 

ABTS, 2,2′-azinobis(3-ethylbenzothiazoline-6-sulfonic acid); DMP, 2,6-dimethoxyphenol; GUA, 2-methoxyphenol (guaiacol); SYR. 4-hydroxy-3,5-dimethoxybenzaldehyde [(4-hydroxy-3,5-dimethoxyphenyl)methylene]hydrazone (syringaldazine).

The species are listed under the names used in the original references.

NA, not active.

*

Strain 17, constitutive form.

Strain 17, induced with p -hydroxybenzoic acid.

Strain C7, constitutive form.

§

Strain 19, induced with ferulic acid.

Several attempts have been undertaken to detect ligninolytic enzymes, including laccases in ectomycorrhizal (ECM) fungi ( Cairney & Burke, 1998 ; Burke & Cairney, 2002 ). Gene fragments with a high similarity to laccase from wood-rotting fungi have been found in several isolates of ECM species including Amanita, Cortinarius, Hebeloma, Lactarius, Paxillus, Piloderma, Russula, Tylospora and Xerocomus ( Luis, 2004 ; Chen, 2003) . In the case of Piloderma byssinum , transcription of putative laccase sequence was confirmed by RT-PCR ( Chen, 2003 ). However, a gene sequence does not necessarily correspond with the production of an enzyme. In Paxillus involutus , a species containing another putative laccase sequence, oxidation of syringaldazine has never been detected ( Günther, 1998 ; Timonen & Sen, 1998 ). It seems that tyrosinase is the major phenoloxidase of ECM, whereas syringaldazine oxidation has scarcely been reported ( Burke & Cairney, 2002 ) and the literature data reporting laccase activity in ECM fungi are usually based on the use of nonspecific substrates like ABTS or naphtol ( Gramss, 1998 , 1999 ). The gene sequences are not found very frequently either ( Chen, 2003 ). Laccases have been purified from a few fungi-forming ectomycorrhiza: Cantharellus cibarius ( Ng & Wang, 2004 ), Lactarius piperatus ( Iwasaki, 1967 ), Russula delica ( Matsubara & Iwasaki, 1972 ) and Thelephora terestris ( Kanunfre & Zancan, 1998 ) or orchideoid mycorrhiza: Armillaria mellea ( Rehman & Thurston, 1992 ; Billal & Thurston, 1996 ; Curir, 1997 ), as well as from the species of genera that contain both saprotrophic and mycorrhizal fungi Agaricus, Marasmius, Tricholoma and Volvariella ( Table 1 ). The activity of another ligninolytic enzyme, Mn-peroxidase, has thus far been confirmed only in Tylospora fibrillosa , a species containing also a putative sequence of laccase ( Chambers, 1999 ; Chen, 2003 ) and possibly also lignin peroxidase ( Chen, 2001 ).

Cellular localization

Due to the properties of their substrate, the enzymes participating in the breakdown of lignin should be exclusively extracellular. While this is without exception true for the lignin peroxidases and manganese peroxidases of white-rot fungi, the situation is not the same with laccases. Although most laccases purified so far are extracellular enzymes, the laccases of wood-rotting fungi are usually also found intracellularly. Most white-rot fungal species tested by Blaich & Esser (1975) produced both extracellular and intracellular laccases with isoenzymes showing similar patterns of activity staining after isoelectric focusing. When Trametes versicolor was grown on glucose, wheat straw and beech leaves, it produced laccases both in extracellular and intracellular fractions ( Schlosser, 1997 ). The majority of enzyme activity was produced extracellularly (98% and 95% on wheat straw and beech wood, respectively). Traces of intracellular laccase activity were found in Agaricus bisporus , but more than 88% of the total activity was in the culture supernatant ( Wood, 1980 ). The intra- and extracellular presence of laccase activity was also detected in Phanerochaete chrysosporium ( Dittmer, 1997 ) and Suillus granulatus ( Günther, 1998 ). A fraction of laccase activity in N. crassa, Rigidoporus lignosus and one of the laccase isoenzymes of Pleurotus ostreatus is also probably localized intracellularly or on the cell wall ( Froehner & Eriksson, 1974 ; Nicole, 1992 , 1993 ; Palmieri, 2000 ).

The extracellular laccase activity of Lentinula edodes was associated with a multicomponent protein complex of 660 kDa, which also exhibited peroxidase and β-glucosidase activities ( Makkar, 2001 ). Although laccase activity was not found in the cell wall fractions of the basidiomycete A. bisporus ( Sassoon & Mooibroek, 2001 ), a substantial part of T. versicolor and P. ostreatus laccase is associated with the cell wall ( Valášková & Baldrian, 2005 ). Laccase activity is almost exclusively associated with cell walls in the white-rot basidiomycete Irpex lacteus ( Svobodová, 2005 ), the yeast C. neoformans ( Zhu, 2001 ) and in the spores of Trichoderma spp. ( Holker, 2002 ). The localization of laccase is probably connected with its physiological function and determines the range of substrates available to the enzyme. It is possible that the intracellular laccases of fungi as well as periplasmic bacterial laccases could participate in the transformation of low molecular weight phenolic compounds in the cell. The cell wall and spores-associated laccases were linked to the possible formation of melanin and other protective cell wall compounds ( Eggert, 1995 ; Galhaup & Haltrich, 2001 ).

Structural properties

Current knowledge about the structure and physico-chemical properties of fungal laccase proteins is based on the study of purified proteins. Up to now, more than 100 laccases have been purified from fungi and been more or less characterized ( Table 1 ). Based on the published data we can draw some general conclusions about laccases, taking into account that most enzymes were purified from wood-rotting white-rot basidiomycetes; other groups of fungi-producing laccases (other groups of basidiomycetes, ascomycetes and imperfect fungi) have been studied to a much lesser extent. Typical fungal laccase is a protein of approximately 60–70 kDa with acidic isoelectric point around pH 4.0 ( Table 2 ). It seems that there is considerable heterogeneity in the properties of laccases isolated from ascomycetes, especially with respect to molecular weight.

Table 2

Properties of fungal laccases (data derived from Table 1 )

Property n Median  Q 25  Q 75 Min Max 
Molecular weight (Da) 103 66 000 61 000 71 000 43 000 383 000 
pI 67 3.9 3.5 4.2 2.6 6.9 
Temperature optimum (°C) 39 55 50 70 25 80 
pH optimum 49 3.0 2.5 4.0 2.0 5.0 
ABTS 
2,6-Dimethoxyphenol 36 4.0 3.0 5.5 3.0 8.0 
Guaiacol 24 4.5 4.0 6.0 3.0 7.0 
Syringaldazine 31 6.0 4.7 6.0 3.5 7.0 
K M (μM)  
ABTS 36 39 18 100 770 
2,6-Dimethoxyphenol 30 405 100 880 26 14 720 
Guaiacol 23 420 121 1600 30 000 
Syringaldazine 21 36 11 131 4307 
k cat (s −1 )        
ABTS 12 24 050 5220 41 460 198 350 000 
2,6-Dimethoxyphenol 12 3680 815 6000 100 360 000 
Guaiacol 10 295 115 3960 90 10 800 
Syringaldazine 21 500 18 400 25 500 16 800 28 000 
Property n Median  Q 25  Q 75 Min Max 
Molecular weight (Da) 103 66 000 61 000 71 000 43 000 383 000 
pI 67 3.9 3.5 4.2 2.6 6.9 
Temperature optimum (°C) 39 55 50 70 25 80 
pH optimum 49 3.0 2.5 4.0 2.0 5.0 
ABTS 
2,6-Dimethoxyphenol 36 4.0 3.0 5.5 3.0 8.0 
Guaiacol 24 4.5 4.0 6.0 3.0 7.0 
Syringaldazine 31 6.0 4.7 6.0 3.5 7.0 
K M (μM)  
ABTS 36 39 18 100 770 
2,6-Dimethoxyphenol 30 405 100 880 26 14 720 
Guaiacol 23 420 121 1600 30 000 
Syringaldazine 21 36 11 131 4307 
k cat (s −1 )        
ABTS 12 24 050 5220 41 460 198 350 000 
2,6-Dimethoxyphenol 12 3680 815 6000 100 360 000 
Guaiacol 10 295 115 3960 90 10 800 
Syringaldazine 21 500 18 400 25 500 16 800 28 000 

n , number of observations; Q 25 , lower quartile; Q 75 , upper quartile.

Several laccase isoenzymes have been detected in many fungal species. More than one isoenzyme is produced in most white-rot fungi. ( Blaich & Esser, 1975 ) performed a screening of laccase activity among wood-rotting fungi using staining with p -phenylenediamine after isoelectric focusing. All tested species, namely Coprinus plicatilis, Fomes fomentarius, Heterobasidion annosum, Hypholoma fasciculare, Kuehneromyces mutabilis, Leptoporus litschaueri, Panus stipticus, Phellinus igniarius, Pleurotus corticatus, P. ostreatus, Polyporus brumalis, Stereum hirsutum, Trametes gibbosa, T. hirsuta and T. versicolor , exhibited the production of more than one isoenzyme, typically with pI in the range of pH 3–5.

Several species produce a wide variety of isoenzymes. The white-rot fungus P. ostreatus produces at least eight different laccase isoenzymes, six of which have been isolated and characterized ( Sannia, 1986 ; Palmieri, 1993 , 1997 , 2003 ; Giardina, 1999 ). The main protein present in the cultures is the 59-kDa POXC with pI 2.7. The POXA2, POXB1 and POXB2 isoenzymes exhibit a similar molecular weight around 67 kDa, while POXA1b and POXA1w are smaller (61 kDa). The enzymes POXA3a and POXA3b are heterodimers consisting of large (61-kDa) and small (16- or 18-kDa) subunits. Although the POXC protein is the most abundant in cultures both extra- and intracellularly, the highest mRNA production was detected in POXA1b, which is probably mainly intracellular or cell wall-associated as it is cleaved by an extracellular protease ( Palmieri, 1997 ; Giardina, 1999 ). The production of laccase isoenzymes in P. ostreatus is regulated by the presence of copper and the two dimeric isoenzymes have only been detected in the presence of copper ( Palmieri, 2000 , 2003 ). Isoenzymes of laccase with different molecular weight and pI were also detected in the litter-decomposing fungus Marasmius quercophilus ( Farnet, 2000 , 2002 , 2004 ; Dedeyan, 2000 ). A study with 17 different isolates of this fungus showed that the isoenzyme pattern was consistent within different isolates. Moreover, all isolates showed the same isoenzyme pattern (one to three laccase bands on SDS-PAGE) after the induction of laccase with different aromatic compounds ( Farnet, 1999 ).

Some fungal species, e.g. Coriolopsis rigida, Dichomitus squalens, Physisporinus rivulosus and Trametes gallica , produce isoenzymes that are closely related both structurally and in their catalytic properties ( Table 1 ). Different properties of laccases purified from the same species and reported by different authors can be explained as a result of both the production of different isoenzymes and different laccase properties in different strains of the same fungus ( Table 1 ). In P. chrysosporium , production of different laccase isoenzymes was detected in cell extract and in the culture medium ( Dittmer, 1997 ); however, since laccase gene was not found in the complete genome sequence of this fungus ( Martinez, 2004 ), these are probably multicopper oxidases rather than true laccases ( Larrondo, 2003 ). The molecular basis for the production of different isoenzymes is the presence of multiple laccase genes in fungi (see e.g. Chen, 2003 ).

Most fungal laccases are monomeric proteins. Several laccases, however, exhibit a homodimeric structure, the enzyme being composed of two identical subunits with a molecular weight typical for monomeric laccases. This is the case of the wood-rotting species Phellinus ribis ( Min, 2001 ), Pleurotus pulmonarius ( De Souza & Peralta, 2003 ) and Trametes villosa ( Yaver, 1996 ), the mycorrhizal fungus C. cibarius ( Ng & Wang, 2004 ) and the ascomycete Rhizoctonia solani ( Wahleithner, 1996 ). The ascomycetes G. graminis, M. indicum and P. anserina also produce oligomeric laccases. In M. indicum a single band of 100 kDa after gel filtration resolved into three proteins (24, 56 and 72 kDa) on SDS-PAGE ( Thakker, 1992 ): G. graminis produces a trimer of three 60-kDa subunits ( Edens, 1999 ); P. anserina laccase is a heterooligomer ( Molitoris & Esser, 1970 ); and one of the laccases purified from A. mellea has a heterodimeric structure ( Curir, 1997 ). According to Wood ( Wood, 1980 ), A. bisporus laccase consists of several polypeptides of 23–56 kDa. ( Perry, 1993 ), on the basis of Western blot analyses, suggested that the native Lac2 of the same species is produced as a dimer of identical polypeptides, one of which is then partially proteolytically cleaved. SDS–PAGE and MALDI-MS analyses of purified POXA3a and POXA3b laccases from P. ostreatus reveal the presence of three different polypeptides of 67, 18 and 16 kDa, whereas the native proteins behave homogeneously, as demonstrated by the presence of a single peak or band in gel filtration chromatography, isoelectric focusing and native-PAGE analysis. All the other laccase isoenzymes isolated from P. ostreatus were characterized as monomeric proteins ( Palmieri, 2003 ).

Like most fungal extracellular enzymes, laccases are glycoproteins. The extent of glycosylation usually ranges between 10% and 25%, but laccases with a saccharide content higher than 30% were found: e.g. Coriolopsis fulvocinnerea −32% ( Shleev, 2004 ) and P. pulmonarius −44% ( De Souza & Peralta, 2003 ). Even higher saccharide contents were found in Botrytis cinnerea , the monomeric enzyme of the strain 61–34 containing 49% sugars ( Slomczynski, 1995 ). Other preparations from the same species exhibited as much as 65–80% of saccharides including arabinose, xylose, mannose, galactose and glucose ( Gigi, 1981 ; Marbach, 1984 ; Zouari, 1987 ). On the other hand, very low extent of glycosylation was detected in Pleurotus eryngii , where laccase I contained 7% and laccase II only 1% of bound sugars ( Munoz, 1997 ). The glycans are N-linked to the polypeptide chain ( Ko, 2001 ; Brown, 2002 ; Saparrat, 2002 ). The most detailed structure of laccase glycan is available for R. lignosus laccase, which is also glycosylated with N-bound mannose ( Garavaglia, 2004 ). The glycosylation of fungal laccases is one of the biggest problems for the heterologous production of the enzyme, which is extremely difficult to overcome. It was proposed that in addition to the structural role, glycosylation can also participate in the protection of laccase from proteolytic degradation ( Yoshitake, 1993 ).

Laccases belong to the group of blue multicopper oxidases (BMCO) that catalyze a one-electron oxidation concomitantly with the four-electron reduction of molecular oxygen to water ( Solomon, 1996 , 2001 ; Messerschmidt, 1997 ). The catalysis carried out by all members of this family is guaranteed by the presence of different copper centres in the enzyme molecule. In particular, all BMCO are characterized by the presence of at least one type-1 (T1) copper, together with at least three additional copper ions: one type-2 (T2) and two type-3 (T3) copper ions, arranged in a trinuclear cluster. The different copper centres can be identified on the basis of their spectroscopic properties. The T1 copper is characterized by a strong absorption around 600 nm, whereas the T2 copper exhibits only weak absorption in the visible region. The T2 site is electron paramagnetic resonance (EPR)-active, whereas the two copper ions of the T3 site are EPR-silent due to an antiferromagnetic coupling mediated by a bridging ligand. The substrates are oxidized by the T1 copper and the extracted electrons are transferred, probably through a strongly conserved His-Cys-His tripeptide motif, to the T2/T3 site, where molecular oxygen is reduced to water ( Messerschmidt, 1997 ) ( Fig. 1 ). Some enzymes lack the T1 copper and some authors hesitate to call them true laccases. Others use the term ‘yellow laccases’ because these enzymes lack the characteristic absorption band around 600 nm ( Leontievsky, 1997 , 1997 ).

Figure 1

Catalytic cycle of laccase.

Figure 1

Catalytic cycle of laccase.

Until recently, the three-dimensional structure of five fungal laccases has been reported: Coprinus cinereus (in a copper type-2-depleted form) ( Ducros, 1998 ), T. versicolor ( Bertrand, 2002 ; Piontek, 2002 ), P. cinnabarinus ( Antorini, 2002 ), M. albomyces ( Hakulinen, 2002 ) and R. lignosus ( Garavaglia, 2004 ), the latter four enzymes with a full complement of copper ions. Moreover, the three-dimensional structure of the CoA laccase from Bacillus subtilis endospore has also recently been published ( Enguita, 2003 , 2004 ). Despite the amount of information on laccases as well as other BMCO, neither the precise electron transfer pathway nor the details of dioxygen reduction in BMCO are fully understood ( Garavaglia, 2004 ). A detailed structural comparison between a low redox potential ( E0 ) C. cinereus laccase and a high E0T. versicolor laccase showed that structural differences of the Cu1 coordination possibly account for the different E0 values ( Piontek, 2002 ). This was later confirmed by the study of R. lignosus laccase with a high redox potential ( Garavaglia, 2004 ). However, more effort will be needed to elucidate the relation between the structure of the catalytic site and the substrate preference of different laccase enzymes.

Unlike the laccases described above, the enzyme from P. ribis with catalytic features typical for laccases does not belong to the blue copper proteins because it lacks Cu1 and contains one Mn atom per molecule. The structural differences are probably also responsible for the relatively high pH optimum for ABTS oxidation ( Min, 2001 ). The ‘white’ laccase POXA1 from P. ostreatus contains only one copper atom, together with two zinc and one iron atoms per molecule ( Palmieri, 1997 ). Future structural studies will probably show that laccases are a more structurally heterogeneous group of proteins than expected.

Catalytic properties

Laccase catalyses the reduction of O 2 to H 2 O using a range of phenolic compounds (though not tyrosine) as hydrogen donors ( Thurston, 1994 ; Solomon, 1996 ). Unfortunately, laccase shares a number of hydrogen donors with tyrosinase, making it difficult to assign unique descriptions to either enzyme. A further complication is the overlap in activity between monophenol monooxygenase and catechol oxidase (1,2-benzenediol: oxygen oxidoreductase, EC 1.10.3.1). The broad range of substrates accepted by laccase as hydrogen donors notwithstanding, oxidation of syringaldazine in combination with the inability to oxidize tyrosine, has been taken to be an indicator of laccase activity ( Harkin, 1974 ; Thurston, 1994 ). Unambiguous determination of laccase activity is best achieved by purification of the protein to electrophoretic homogeneity followed by determination of K M or k cat with multiple substrates. Ideally, these should include substrates such as syringaldazine, ABTS or catechol, for which laccase has a high affinity, and some (e.g. tyrosine) for which laccase has little or no affinity ( Edens, 1999 ; Shin & Lee, 2000 ). In common with catechol oxidase and tyrosinase, laccase catalyzes the four-electron reduction of O 2 to H 2 O. In the case of laccase, at least, this is coupled to the single-electron oxidation of the hydrogen-donating substrate ( Reinhammar & Malmstrom, 1981 ). Since four single-electron substrate oxidation steps are required for the four-electron reduction of water, the analogy of a four-electron ‘biofuel cell’ has been proposed to explain this complex mechanism ( Thurston, 1994 ; Call & Mucke, 1997 ; Barriere, 2004 ). Laccases are known to be highly oxidizing. E0 ranges from 450–480 mV in Myceliophthora thermophila to 760–790 mV in Polyporus pinsitus ( Solomon, 1996 ; Xu, 1996 ; Xu, 2000 ) and the presence of four cupric ions, each co-ordinated to a single polypeptide chain, is an absolute requirement for optimal activity ( Ducros, 1998 ). There have been few measurements of the redox potentials of tyrosinase or catechol oxidase; however, ( Ghosh & Mukherjee, 1998 ) estimated the E0 of a tyrosinase model system to be 260 mV, considerably lower than that reported for laccase, suggesting that this class of enzyme is much less oxidizing than laccase.

Due to the difficulties with distinguishing laccases from other oxidases, the data in this review are based exclusively on the reports concerning purified enzymes. However, the direct comparison of biochemical data reported for different fungal laccases that would be extremely important for the biotechnological applicability is difficult, as different test conditions have been used in different reports. There are only a few works comparing laccase properties of enzymes from different sources, e.g. the work of ( Shleev, 2004 ) focusing on physico-chemical and spectral characteristics of four different laccases. However, this comparison is rather limited.

A very wide range of substrates has been shown to be oxidized by fungal laccases ( Table 3 ) but the catalytic constants have been reported mostly for a small group of substrates – e.g. the non-natural test substrate ABTS and the phenolic compounds 2,6-dimethoxyphenol (DMP), guaiacol and syringaldazine. K M ranges from 10 s of μM for syringaldazine and ABTS to 100 s of μM for DMP and guaiacol. The catalytic performance expressed as k cat spans several orders of magnitude for different substrates and is usually characteristic for a specific protein ( Table 3 ). Laccases in general combine high affinity for ABTS and syringaldazine with high catalytic constant, whereas the oxidation of guaiacol and DMP is considerably slower and the respective K M constants higher. Low K M values are typical for sinapic acid, hydroquinone and syringic acid, whereas relatively high values were found for para -substituted phenols, vanillic acid or its aldehyde. For the species capable of oxidizing polycyclic aromatic hydrocarbons or pentachlorophenol, only very low catalytic constants were detected for these xenobiotic compounds; the K M value is also high for pentachlorophenol with T. versicolor laccase ( Table 3 ).

Table 3

Substrates and inhibitors of fungal laccases. The numbers in brackets indicate Michaelis constant (K M , μM) or rate constant (k cat , s −1 ), multiple values for the same species refer to different isoenzymes. Only compounds that undergo transformation without the presence of redox mediators are listed as substrates

 Species 
Substrate 
Substrate (3,4-Dimethoxyphenyl)methanol (veratryl alcohol) Ts, Tv 
(4-Hydroxy-3-methoxyphenyl)acetic acid Pe 
1,2,4,5-Tetramethoxybenzene  Cs (K M : 6900; k cat : 1680), Cs (K M : 900; k cat : 3360)  
1,2,4-Benzenetriol Bc 
1,2-Benzenediol (catechol)  Ab, Am, Bc, Cf (K M : 85; k cat : 90), Ch, Cm (K M : 120; k cat : 320), Cn, Cr, Ds, Gg (K M : 250), Gl (K M : 55), Le (K M : 220), Le (K M : 22 400), Lp, Mi, Mq, Pc, Pe (K M : 2200), Pe (K M : 4100), Pr, Rl, Sr, Th (K M : 142; k cat : 390), To (K M : 110; k cat : 80), Tp (K M : 470; k cat : 27 600), Ts, Tt  
1,3-Dihydroxybenzene (resorcinol) Cn, Ts 
1,4-Benzohydroquinone  Am, Bc, Cf (K M : 68; k cat : 110), Ch, Cn, Cm (K M : 100; k cat : 290), Cr, Ct (K M : 36), Ds, Gl (K M : 29), Lp, Le (K M : 110), Pc, Pe (K M : 2500), Pe (K M : 4600), Pi, Pn, Rl, Th (K M : 61; k cat : 450), To (K M : 74; k cat : 110), Tp (K M : 390; k cat : 19 200), Ts, Tt  
1-Naphthol Ab, Bc, Gl 
2-(3,4-Dihydroxyphenyl)-3,5,7-trihydroxy-4H-chromen-4-one Am 
2-Chlorophenol Tv 
2,2′-Azinobis(3-ethylbenzothiazoline-6-sulfonic acid)  Al (kcat 21), Cr (k cat : 4680), Cr (k cat : 4620), Cs (k cat : 5760), Cs (k cat : 6060), Po (k cat : 16 000), Po (k cat : 350 000), Po (k cat : 90 000), Rl (k cat : 34 700), Rl (k cat : 32 100), Tp (k cat : 41 400), Tr (k cat : 41 520), Tt (k cat : 198)  
2,3-Dichlorophenol Tv 
2,3-Dimethoxyphenol Ds 
2,3,6-Trichlorophenol Tv 
2,4,6-Trichlorophenol Tv 
2,4,6-Trimethylphenol Pe 
2,4-Dichlorophenol Tt, Tv 
2,5-Dihydroxybenzoic acid Pi 
2,6-Dichlorophenol Tt, Tv 
2,6-Dimethoxy-1,4-benzohydroquinone  Cr (K M : 107; k cat : 8580), Cr (K M : 89; k cat : 11 220)  
2,6-Dimethoxyphenol  Al (k cat : 15), Cr (k cat : 6360), Cr (k cat : 5640), Cs (k cat : 1380), Cs (k cat : 4560), Po (k cat : 100), Po (k cat : 250), Po (k cat : 360 000), Rl (k cat : 2800), Rl (k cat : 2000), Tp (k cat : 24 000), Tr (k cat : 4860), Tt (k cat : 109)  
2,7-Diaminofluorene Rl 
2-Amino-3-(3,4-dihydroxyphenyl)propanoic acid  Am, Cn, Ct (K M : 100), Le (K M : 650), Lp, Ma, Pa (K M : 3300), Ts, Tv (K M : 15 600)  
2-Amino-3-hydroxybenzoic acid Pc 
2-Amino-4-methylphenol Tt 
2-Amino-4-nitrophenol Tt 
2-Aminophenol Tt 
2-Aminophenylamine Cn 
2-Chlorobenzene-1,4-diol Tt 
2-Chlorophenol  Le (K M : 1350), Mq, Tt  
2-Methoxy-1,4-benzohydroquinone  Cr (K M : 216; k cat : 7620), Cr (K M : 229; k cat : 6300), Pe  
2-Methoxy-4-[prop-1-enyl]phenol Ts 
2-Methoxy-4-methylphenol Tt 
2-Methoxyaniline Tt 
2-Methoxyphenol (guaiacol)  Al (k cat : 159), Cf (k cat : 95), Cm (k cat : 160), Cs (k cat : 3120), Cs (k cat : 3960), Po (k cat : 150), Th (k cat : 430), To (k cat : 90), Tp (k cat : 10 800), Tr (k cat : 4140), Tt (k cat : 115)  
2-Methoxy-1,4-benzohydroquinone Pe 
2-Methyl-1,4-benzohydroquinone  Pe (K M : 1600), Pe (K M : 2100)  
2-Methylanthracene  Cg (k cat : 0.082)  
2-Methylphenol Bc, Tt 
2-Naphthol Bc, Gl 
2,4-Dichlorophenol Mq 
2,4,6-Trichlorophenol 
3-(3,4-Dihydroxyphenyl)acrylic acid (caffeic acid)  Am, Bc, Cs, Le (K M : 40), Mi, Mq, Ts, Tt  
3-(4-Hydroxy-3,5-dimethoxyphenyl)acrylic acid  Cf (K M : 21, k cat : 140), Cm (K M : 24; k cat : 330), Cs, Ff, Le (K M : 110), Pn, Pr, Rl, Th (K M : 24; k cat : 580), To (K M : 11; k cat : 170), Tv  
3-(4-Hydroxy-3-methoxyphenyl)acrylic acid (ferulic acid)  Am, Cf (K M : 20), Ch, Cs, Ct (K M : 270), Ff, Le (K M : 240), Le (K M : 2860), Mi, Mq, Pc, Pn, Pr, Rl, Sr, Tt (K M : 40; k cat : 145), Tv  
3-(4-Hydroxyphenyl)acrylic acid  Le (K M : 240), Pn, Rl, Tt  
3,3′-Dimethoxy-1,1′-biphenyl-4,4′-diamine  Mi (K M : 25), Pr, Tc  
3,4,5-Trihydroxybenzoic acid (gallic acid)  Ab, Am, Bc, Ct (K M : 130), Le (K M : 130), Mq, Nc, On  
3,4-Dihydroxybenzoic acid Cr, Mi, Mq, Pe, Tt 
3,5-Cyclohexadiene-1,2-diol Pe 
3,5-Dimethoxy-hydroxy-benzaldazine Bc 
3-{[3-(3,4-Dihydroxyphenyl)prop-2-enoyl]oxy}-1,4,5-trihydroxycyclohexanecarboxylic acid Am, Ct 
3-Amino-4-hydroxybenzenesulphonic acid Tt 
3-Methoxyphenol Pr 
4-(Hydroxymethyl)-2-methoxyphenol  Cs (K M : 1600; k cat : 2820), Cs (K M : 610; k cat : 2220), Pe  
4-[3-Hydroxyprop-1-enyl]-2,6-dimethoxyphenol Mq 
4-[3-Hydroxyprop-1-enyl]-2-methoxyphenol (coniferyl alcohol) Pe, Rl 
4-[3-Hydroxyprop-1-enyl]-phenol Mq 
4-Amino-2,6-dichlorophenol Bc, Tt 
4-Aminophenol  Pe (K M : 1000), Pe (K M : 800), Tt  
4-Aminophenylamine  Am (K M : 1690), Bc, Cn, Gl, Lp, Le (K M : 256), Pe, Tc, Ts  
4-Hydroxy-3,5-dimethoxybenzaldehyde Cr 
4-Hydroxy-3,5-dimethoxybenzaldehyde [(4-hydroxy-3,5-dimethoxyphenyl)methylene] 
hydrazone (syringaldazine)  Al (k cat : 5), Po (k cat : 23 000), Po (k cat : 28 000), Po (k cat : 20 000), Tp (k cat : 16 800)  
4-Hydroxy-3,5-dimethoxybenzoic acid (syringic acid)  Cr, Cs (K M : 100; k cat : 4680), Cs (K M : 130; k cat : 1860), Ds, Ff (K M : 30), Mi, Pe, Pr, Tv (K M : 60)  
4-Hydroxy-3-methoxybenzaldehyde  Cr, Cs (K M : 6300; k cat : 1560), Cs (K M : 9000; k cat : 600), Pe  
4-Hydroxy-3-methoxybenzoic acid (vanillic acid)  Cf (K M : 160), Cs (K M : 1000; k cat : 3960), Cs (K M : 1100; k cat : 2220), Ct (K M : 150), Ff (K M : 970), Mi, Pe, Pr, Tt, Tv (K M : 130)  
4-Hydroxybenzoic acid Mi 
4-Hydroxyindole Ch, Pc 
4-Chlorophenol  Le (K M : 1740), Tt  
4-Methoxyaniline  Cr, Mi, Pe (K M : 3100), Pe (K M : 3300), Tp (K M : 1600; k cat : 7800), Tt  
4-Methoxyphenol  Cr, Le (K M : 330), Pe (K M : 800), Pe (K M : 900), Pr, Tt  
4-Methylbenzene-1,2-diol  Am, Bc, Ch, Cy, Le (K M : 170), Mq, Pc, Rl  
4-Methylphenol  Bc, Le (K M : 2200), Tt  
4-Nitrobenzene-1,2-diol Tt 
5-(1,2-Dihydroxyethyl)-3,4-dihydroxyfuran-2-one (ascorbic acid)  Ab, Am, Bc, Lp, Mi, Nc, Pa (K M : 190)  
9-Methylanthracene  Cg (k cat : 4)  
Acenaphthene  Cg (k cat : 0.167)  
Anthracene  Cg (k cat : 0.087), Po, Tv  
Benzcatechin  Pa (K M : 2270)  
Benzene-1,2,3-triol (pyrogallol)  Ab, Bc, Ch, Cy, Gg (K M : 310), Gl, Le (K M : 30), Le (K M : 417), Lp, Nc, Pc, Rl, Sr, Ts  
Benzene-1,3,5-triol (phloroglucinol) Mi 
Benzo[a]pyrene  Cg (k cat : 1.38)  
Biphenylene  Cg (k cat : 0.063)  
Fluoranthene Po 
K 4 [Fe(CN) 6 ]   Am (K M : 1720), Cf (K M : 170; k cat : 130), Cm (K M : 115; k cat : 450), Lp, Pa (K M : 1030), Pi, Th (K M : 180; k cat : 400), To (K M : 96; k cat : 150), Tp (K M : 43; k cat : 51 000)  
Mn 2+  St, Tv (K M : 186)  
N,N′-Dimethylbenzene-1,4-diamine Ab, Am, Bc, Cf, Pc 
o -Tolidine   Gl (K M : 402)  
o -Vanillin   Cf (K M : 3900)  
Pentachlorophenol  Tv (K M : 3000; k cat : 0.023)  
Phenanthrene  Cg (k cat : 0.013)  
Phenylhydrazine Rl 
Inhibitor 
Ca 2+ Le 
Cd 2+ Dq, Le 
Co 2+ Dq 
Fe 2+ Ct, Po, Lp, Tc 
Hg 2+ Ct, Dq, Le, Pu 
Mn 2+ Dq, Pu 
Rb + Le 
Sn 2+ Le 
Zn 2+ Le, Po 
1-Phenyl-2-thiourea Ct 
2-Mercaptobenzothiazole Ct 
2-Mercaptoethanol Ct, Pu, Te 
3-(4-Hydroxyphenyl)acrylic acid Le 
4-Nitrophenol Pz 
8-Hydroxyquinoline Lp, Te 
Ascorbic acid Ct, Tc 
Cetylpyridinium bromide Ab 
Cetyltriammonium bromide Ab, Tc 
CN- Ab, Bc, Ct, Gl, Lp, Ma, Me, Mi, Pn, Po, Pz, Rl, Tg, Tr, Ts 
Cysteine Ct, Ch, Dq, Gl, Le, Mq, Pc, Py, Sr, Te, Vv 
Diethyldithiocarbamic acid Bc, Ch, Gl, Lp, Mi, Pp, Pc, Ps, Sr 
Dithiothreitol Ch, Dq, Le, Pc, Py, Vv 
EDTA  Ct, Ma, Mq * , Vv  
Glutathione Dq, Gl 
Humic acid Pt 
Hydroxylamine Po 
KCl Le 
Kojic acid Dq, Le, Po 
NaCl Sr 
NaF Ds, Me, Sr, Tt 
NaN 3 Ab, Bc, Ch, Ct, Dq, Ds, Gl, Le, Ma, Me, Mi, Pa, Pc, Po, Pp, Pr, Ps, Pu, Pz, Sr, Tc, Te, Tg, Tr, Ts, Vv 
Salicylaldoxime Gl 
SDS  Ds, Mq * , Pu, Tr  
Thiamine Sr 
Thiogylcolic acid Ct, Mi, Po, Pr, Sr, Vv 
Thiourea Ct, Dq 
Trifluoroacetic acid Tr 
Tropolone Ch, Le, Pc 
 Species 
Substrate 
Substrate (3,4-Dimethoxyphenyl)methanol (veratryl alcohol) Ts, Tv 
(4-Hydroxy-3-methoxyphenyl)acetic acid Pe 
1,2,4,5-Tetramethoxybenzene  Cs (K M : 6900; k cat : 1680), Cs (K M : 900; k cat : 3360)  
1,2,4-Benzenetriol Bc 
1,2-Benzenediol (catechol)  Ab, Am, Bc, Cf (K M : 85; k cat : 90), Ch, Cm (K M : 120; k cat : 320), Cn, Cr, Ds, Gg (K M : 250), Gl (K M : 55), Le (K M : 220), Le (K M : 22 400), Lp, Mi, Mq, Pc, Pe (K M : 2200), Pe (K M : 4100), Pr, Rl, Sr, Th (K M : 142; k cat : 390), To (K M : 110; k cat : 80), Tp (K M : 470; k cat : 27 600), Ts, Tt  
1,3-Dihydroxybenzene (resorcinol) Cn, Ts 
1,4-Benzohydroquinone  Am, Bc, Cf (K M : 68; k cat : 110), Ch, Cn, Cm (K M : 100; k cat : 290), Cr, Ct (K M : 36), Ds, Gl (K M : 29), Lp, Le (K M : 110), Pc, Pe (K M : 2500), Pe (K M : 4600), Pi, Pn, Rl, Th (K M : 61; k cat : 450), To (K M : 74; k cat : 110), Tp (K M : 390; k cat : 19 200), Ts, Tt  
1-Naphthol Ab, Bc, Gl 
2-(3,4-Dihydroxyphenyl)-3,5,7-trihydroxy-4H-chromen-4-one Am 
2-Chlorophenol Tv 
2,2′-Azinobis(3-ethylbenzothiazoline-6-sulfonic acid)  Al (kcat 21), Cr (k cat : 4680), Cr (k cat : 4620), Cs (k cat : 5760), Cs (k cat : 6060), Po (k cat : 16 000), Po (k cat : 350 000), Po (k cat : 90 000), Rl (k cat : 34 700), Rl (k cat : 32 100), Tp (k cat : 41 400), Tr (k cat : 41 520), Tt (k cat : 198)  
2,3-Dichlorophenol Tv 
2,3-Dimethoxyphenol Ds 
2,3,6-Trichlorophenol Tv 
2,4,6-Trichlorophenol Tv 
2,4,6-Trimethylphenol Pe 
2,4-Dichlorophenol Tt, Tv 
2,5-Dihydroxybenzoic acid Pi 
2,6-Dichlorophenol Tt, Tv 
2,6-Dimethoxy-1,4-benzohydroquinone  Cr (K M : 107; k cat : 8580), Cr (K M : 89; k cat : 11 220)  
2,6-Dimethoxyphenol  Al (k cat : 15), Cr (k cat : 6360), Cr (k cat : 5640), Cs (k cat : 1380), Cs (k cat : 4560), Po (k cat : 100), Po (k cat : 250), Po (k cat : 360 000), Rl (k cat : 2800), Rl (k cat : 2000), Tp (k cat : 24 000), Tr (k cat : 4860), Tt (k cat : 109)  
2,7-Diaminofluorene Rl 
2-Amino-3-(3,4-dihydroxyphenyl)propanoic acid  Am, Cn, Ct (K M : 100), Le (K M : 650), Lp, Ma, Pa (K M : 3300), Ts, Tv (K M : 15 600)  
2-Amino-3-hydroxybenzoic acid Pc 
2-Amino-4-methylphenol Tt 
2-Amino-4-nitrophenol Tt 
2-Aminophenol Tt 
2-Aminophenylamine Cn 
2-Chlorobenzene-1,4-diol Tt 
2-Chlorophenol  Le (K M : 1350), Mq, Tt  
2-Methoxy-1,4-benzohydroquinone  Cr (K M : 216; k cat : 7620), Cr (K M : 229; k cat : 6300), Pe  
2-Methoxy-4-[prop-1-enyl]phenol Ts 
2-Methoxy-4-methylphenol Tt 
2-Methoxyaniline Tt 
2-Methoxyphenol (guaiacol)  Al (k cat : 159), Cf (k cat : 95), Cm (k cat : 160), Cs (k cat : 3120), Cs (k cat : 3960), Po (k cat : 150), Th (k cat : 430), To (k cat : 90), Tp (k cat : 10 800), Tr (k cat : 4140), Tt (k cat : 115)  
2-Methoxy-1,4-benzohydroquinone Pe 
2-Methyl-1,4-benzohydroquinone  Pe (K M : 1600), Pe (K M : 2100)  
2-Methylanthracene  Cg (k cat : 0.082)  
2-Methylphenol Bc, Tt 
2-Naphthol Bc, Gl 
2,4-Dichlorophenol Mq 
2,4,6-Trichlorophenol 
3-(3,4-Dihydroxyphenyl)acrylic acid (caffeic acid)  Am, Bc, Cs, Le (K M : 40), Mi, Mq, Ts, Tt  
3-(4-Hydroxy-3,5-dimethoxyphenyl)acrylic acid  Cf (K M : 21, k cat : 140), Cm (K M : 24; k cat : 330), Cs, Ff, Le (K M : 110), Pn, Pr, Rl, Th (K M : 24; k cat : 580), To (K M : 11; k cat : 170), Tv  
3-(4-Hydroxy-3-methoxyphenyl)acrylic acid (ferulic acid)  Am, Cf (K M : 20), Ch, Cs, Ct (K M : 270), Ff, Le (K M : 240), Le (K M : 2860), Mi, Mq, Pc, Pn, Pr, Rl, Sr, Tt (K M : 40; k cat : 145), Tv  
3-(4-Hydroxyphenyl)acrylic acid  Le (K M : 240), Pn, Rl, Tt  
3,3′-Dimethoxy-1,1′-biphenyl-4,4′-diamine  Mi (K M : 25), Pr, Tc  
3,4,5-Trihydroxybenzoic acid (gallic acid)  Ab, Am, Bc, Ct (K M : 130), Le (K M : 130), Mq, Nc, On  
3,4-Dihydroxybenzoic acid Cr, Mi, Mq, Pe, Tt 
3,5-Cyclohexadiene-1,2-diol Pe 
3,5-Dimethoxy-hydroxy-benzaldazine Bc 
3-{[3-(3,4-Dihydroxyphenyl)prop-2-enoyl]oxy}-1,4,5-trihydroxycyclohexanecarboxylic acid Am, Ct 
3-Amino-4-hydroxybenzenesulphonic acid Tt 
3-Methoxyphenol Pr 
4-(Hydroxymethyl)-2-methoxyphenol  Cs (K M : 1600; k cat : 2820), Cs (K M : 610; k cat : 2220), Pe  
4-[3-Hydroxyprop-1-enyl]-2,6-dimethoxyphenol Mq 
4-[3-Hydroxyprop-1-enyl]-2-methoxyphenol (coniferyl alcohol) Pe, Rl 
4-[3-Hydroxyprop-1-enyl]-phenol Mq 
4-Amino-2,6-dichlorophenol Bc, Tt 
4-Aminophenol  Pe (K M : 1000), Pe (K M : 800), Tt  
4-Aminophenylamine  Am (K M : 1690), Bc, Cn, Gl, Lp, Le (K M : 256), Pe, Tc, Ts  
4-Hydroxy-3,5-dimethoxybenzaldehyde Cr 
4-Hydroxy-3,5-dimethoxybenzaldehyde [(4-hydroxy-3,5-dimethoxyphenyl)methylene] 
hydrazone (syringaldazine)  Al (k cat : 5), Po (k cat : 23 000), Po (k cat : 28 000), Po (k cat : 20 000), Tp (k cat : 16 800)  
4-Hydroxy-3,5-dimethoxybenzoic acid (syringic acid)  Cr, Cs (K M : 100; k cat : 4680), Cs (K M : 130; k cat : 1860), Ds, Ff (K M : 30), Mi, Pe, Pr, Tv (K M : 60)  
4-Hydroxy-3-methoxybenzaldehyde  Cr, Cs (K M : 6300; k cat : 1560), Cs (K M : 9000; k cat : 600), Pe  
4-Hydroxy-3-methoxybenzoic acid (vanillic acid)  Cf (K M : 160), Cs (K M : 1000; k cat : 3960), Cs (K M : 1100; k cat : 2220), Ct (K M : 150), Ff (K M : 970), Mi, Pe, Pr, Tt, Tv (K M : 130)  
4-Hydroxybenzoic acid Mi 
4-Hydroxyindole Ch, Pc 
4-Chlorophenol  Le (K M : 1740), Tt  
4-Methoxyaniline  Cr, Mi, Pe (K M : 3100), Pe (K M : 3300), Tp (K M : 1600; k cat : 7800), Tt  
4-Methoxyphenol  Cr, Le (K M : 330), Pe (K M : 800), Pe (K M : 900), Pr, Tt  
4-Methylbenzene-1,2-diol  Am, Bc, Ch, Cy, Le (K M : 170), Mq, Pc, Rl  
4-Methylphenol  Bc, Le (K M : 2200), Tt  
4-Nitrobenzene-1,2-diol Tt 
5-(1,2-Dihydroxyethyl)-3,4-dihydroxyfuran-2-one (ascorbic acid)  Ab, Am, Bc, Lp, Mi, Nc, Pa (K M : 190)  
9-Methylanthracene  Cg (k cat : 4)  
Acenaphthene  Cg (k cat : 0.167)  
Anthracene  Cg (k cat : 0.087), Po, Tv  
Benzcatechin  Pa (K M : 2270)  
Benzene-1,2,3-triol (pyrogallol)  Ab, Bc, Ch, Cy, Gg (K M : 310), Gl, Le (K M : 30), Le (K M : 417), Lp, Nc, Pc, Rl, Sr, Ts  
Benzene-1,3,5-triol (phloroglucinol) Mi 
Benzo[a]pyrene  Cg (k cat : 1.38)  
Biphenylene  Cg (k cat : 0.063)  
Fluoranthene Po 
K 4 [Fe(CN) 6 ]   Am (K M : 1720), Cf (K M : 170; k cat : 130), Cm (K M : 115; k cat : 450), Lp, Pa (K M : 1030), Pi, Th (K M : 180; k cat : 400), To (K M : 96; k cat : 150), Tp (K M : 43; k cat : 51 000)  
Mn 2+  St, Tv (K M : 186)  
N,N′-Dimethylbenzene-1,4-diamine Ab, Am, Bc, Cf, Pc 
o -Tolidine   Gl (K M : 402)  
o -Vanillin   Cf (K M : 3900)  
Pentachlorophenol  Tv (K M : 3000; k cat : 0.023)  
Phenanthrene  Cg (k cat : 0.013)  
Phenylhydrazine Rl 
Inhibitor 
Ca 2+ Le 
Cd 2+ Dq, Le 
Co 2+ Dq 
Fe 2+ Ct, Po, Lp, Tc 
Hg 2+ Ct, Dq, Le, Pu 
Mn 2+ Dq, Pu 
Rb + Le 
Sn 2+ Le 
Zn 2+ Le, Po 
1-Phenyl-2-thiourea Ct 
2-Mercaptobenzothiazole Ct 
2-Mercaptoethanol Ct, Pu, Te 
3-(4-Hydroxyphenyl)acrylic acid Le 
4-Nitrophenol Pz 
8-Hydroxyquinoline Lp, Te 
Ascorbic acid Ct, Tc 
Cetylpyridinium bromide Ab 
Cetyltriammonium bromide Ab, Tc 
CN- Ab, Bc, Ct, Gl, Lp, Ma, Me, Mi, Pn, Po, Pz, Rl, Tg, Tr, Ts 
Cysteine Ct, Ch, Dq, Gl, Le, Mq, Pc, Py, Sr, Te, Vv 
Diethyldithiocarbamic acid Bc, Ch, Gl, Lp, Mi, Pp, Pc, Ps, Sr 
Dithiothreitol Ch, Dq, Le, Pc, Py, Vv 
EDTA  Ct, Ma, Mq * , Vv  
Glutathione Dq, Gl 
Humic acid Pt 
Hydroxylamine Po 
KCl Le 
Kojic acid Dq, Le, Po 
NaCl Sr 
NaF Ds, Me, Sr, Tt 
NaN 3 Ab, Bc, Ch, Ct, Dq, Ds, Gl, Le, Ma, Me, Mi, Pa, Pc, Po, Pp, Pr, Ps, Pu, Pz, Sr, Tc, Te, Tg, Tr, Ts, Vv 
Salicylaldoxime Gl 
SDS  Ds, Mq * , Pu, Tr  
Thiamine Sr 
Thiogylcolic acid Ct, Mi, Po, Pr, Sr, Vv 
Thiourea Ct, Dq 
Trifluoroacetic acid Tr 
Tropolone Ch, Le, Pc 

Ab, Agaricus bisporus ( Wood, 1980 ); Al, Agaricus blazei ( Ullrich, 2005 ); Am, Armillaria mellea ( Rehman & Thurston, 1992 ; Curir, 1997 ); Bc, Botrytis cinerea ( Zouari, 2002 ); Cf, Coriolopsis fulvocinnerea ( Smirnov, 2001 ; Shleev, 2004 ); Cg, Coriolopsis gallica ( Pickard, 1999 ); Ch, Coriolus hirsutus ( Eggert, 1996 ); Cm, Cerrena maxima ( Shleev, 2004 ); Cn, Cryptococcus neoformans ( Williamson, 1994 ); Cr, Coriolopsis rigida ( Saparrat, 2002 ); Cs, Ceriporiopsis subvermispora ( Fukushima & Kirk, 1995 ; Salas, 1995 ); Ct, Chaetomium termophilum ( Chefetz, 1998 ; Ishigami, 1998 ); Cy, Cyathus stercoreus ( Sethuraman, 1999 ); Dq, Daedalea quercina ( Baldrian, 2004 ); Ds, Dichomitus squalens ( Perie, 1998 ); Ff, Fomes fomentarius ( Rogalski, 1991 ); Gg, Gaeumannomyces graminis ( Edens, 1999 ); Gl, Ganoderma lucidum ( Lalitha Kumari & Sirsi, 1972 ; Ko, 2001 ); Le, Lentinula edodes ( D'Annibale, 1996 ; Nagai, 2002 ); Lp, Lactarius piperatus ( Iwasaki, 1967 ); Ma, Mauginiella sp. ( Palonen, 2003 ); Me, Melanocarpus albomyces ( Kiiskinen, 2002 ); Mi, Monocillium indicum ( Thakker, 1992 ); Mq, Marasmius quercophilus ( Dedeyan, 2000 ; Farnet, 2004 ); Nc, Neurospora crassa ( Froehner & Eriksson, 1974 ); On, Ophiostoma novo-ulmi ( Binz & Canevascini, 1997 ); Pa, Podospora anserina ( Molitoris & Esser, 1970 ); Pc, Pycnoporus cinnabarinus ( Eggert, 1996 , 1995 ); Pe, Pleurotus eryngii ( Munoz, 1997 , 1997 ); Pi, Polyporus anisoporus ( Vaitkyavichyus, 1984 ); Pn, Phellinus noxius ( Geiger, 1986 ); Po, Pleurotus ostreatus ( Palmieri, 1997 ; Giardina, 1999 ; Pozdnyakova, 2004 ; Das, 2000 ); Pp, Panaeolus papilionaceus ( Heinzkill, 1998 ); Pr, Phellinus ribis ( Min, 2001 ); Ps, Panaeolus sphinctricus ( Heinzkill, 1998 ); Pt, Panus tigrinus ( Zavarzina, 2004 ); Pu, Pleurotus pulmonarius ( De Souza & Peralta, 2003 ); Py, Pycnoporus coccineus ( Oda, 1991 ); Pz, Pyricularia oryzae ( Neufeld, 1958 ); Rl, Rigidoporus lignosus ( Geiger, 1986 ; Bonomo, 1998 ; Cambria, 2000 ); Sr, Sclerotium rolfsii ( Ryan, 2003 ); St, Stropharia rugosoannulata ( Schlosser & Höfer, 2002 ); Tc, Trichoderma sp. ( Assavanig, 1992 ); Te, Thelephora terrestris ( Kanunfre & Zancan, 1998 ); Tg, Trametes gallica ( Dong & Zhang, 2004 ); Th, Trametes hirsuta ( Shleev, 2004 ); To, Trametes ochracea ( Shleev, 2004 ); Tp, Trametes pubescens ( Galhaup, 2002 ); Ts, Trametes sanguinea ( Nishizawa, 1995 ); Tr, Trametes sp. AH28-2 ( Xiao, 2003 ); Tt, Trametes trogii ( Garzillo, 1998 ); Tv, Trametes versicolor ( Bourbonnais & Paice, 1990 ; Rogalski, 1991 ; Salas, 1995 ; Johannes, 1996 ; Collins, 1996 ; Dawel, 1997 ; Höfer & Schlosser, 1999 ; Itoh, 2000 ; Ullah, 2000 ; Leontievsky, 2001 ); Vv, Volvariella volvacea ( Chen, 2004 ).

Some fungi produce isoenzymes with similar K M and k cat values. In wood-rotting basidiomycetes that are usually dikaryotic this fact probably indicates that allelic variability is responsible for the production of isoenzymes rather than the evolution of enzymes adapted to the special needs of the fungus. In the case of P. ostreatus , however, the isoenzymes show the K M and k cat values for 2,6-dimethoxyphenol or guaiacol differing by several orders of magnitude and the POXA1 isoenzyme is not active with guaiacol at all ( Table 3 ).

Even very early reports showed that different laccase enzymes differ considerably in their catalytic preferences. Laccases can be grouped according to their preference for ortho-, meta- or para- substituted phenols. Ortho -substituted compounds (guaiacol, o -phenylenediamine, caffeic acid, catechol, dihydroxyphenylalanine, protocatechuic acid, gallic acid and pyrogallol) were better substrates than para -substituted compounds ( p -phenylenediamine, p -cresol, hydroquinone) and the lowest rates were obtained with meta -substituted compounds ( m -phenylenediamine, orcinol, resorcinol and phloroglucinol) with crude laccase preparations from L. litschaueri and P. brumalis ( Blaich & Esser, 1975 ). Similar results were also obtained with T. versicolor and the ascomycetes P. anserina and Pyricularia oryzae , whereas laccase from Ganoderma lucidum catalyzed the oxidation of only ortho and para dihydroxyphenyl compounds, p -phenylenediamine and polyphenols, not the meta hydroxymethyl compounds or ascorbic acid ( Fahraeus, 1961 ; Fahraeus & Ljunggren, 1961 ; Scháněl & Esser, 1971 ; Lalitha Kumari & Sirsi, 1972 ). More than 70% oxidation of o -substituted compounds was obtained with laccase from M. indicum , whereas p -compounds and the m -phenol phloroglucinol were oxidized at a relatively low rate ( Thakker, 1992 ). The relative oxidation rates for different substrates in relation to the oxidation of 2,6-dimethoxyphenol are summarized in Table 4 . The data demonstrate the high activity with ABTS (with the exception of Myrothecium verrucaria ) and a generally high variation with other substrates.

Table 4

Reactivity of fungal laccases with different substrates. The numbers indicate the rate of substrate oxidation (%) compared to the oxidation of 2,6-dimethoxyphenol

Substrate Species 
Am Ct Ch Cy Ds Ma Me Mv Pr Pe Pc Sr Ts Tt Tv 
2,6-Dimethoxyphenol 100.0 100.0 100.0 100.0 100.0 100.0 100.0 100.0 100.0 100.0 100.0 100.0 100.0 100.0 100.0 
2-Amino-3-(3,4-dihydroxyphenyl)propanoic acid  55.9           46.0   
4-Aminophenol          109.8    26.7  
4-Aminophenylamine 62.9       14.7  194.8   56.0   
4-Hydroxyindole   87.6        107.6     
4-Methoxyphenol         7.3 9.8    19.8  
4-Methylcatechol   69.3 31.4       103.5     
ABTS  76.5 271.7 114.4  800.0 288.3 1.4 97.4 284.1 452.0 136.7   27.7 
Caffeic acid        18.6     95.0 80.2 24.5 
Catechol 74.2  44.9  59.2   34.9 33.1 21.6 74.3 18.3 76.0 27.9 110.7 
Ferulic acid  111.8 48.4      8.5  76.0   116.3  
Guaiacol 59.7 73.5 107.8 37.9 92.0 122.4 31.0 39.1 9.7  140.9 35.0 64.0 55.8 38.4 
Hydroquinone  50.0 105.3  80.8   12.9  25.5 62.0  69.0 30.2 56.0 
N,N-dimethyl-1,4-phenylenediamine 74.8  1.8     23.4   19.9    237.1 
o -Anisidine          45.5     9.3  
p -Anisidine           19.3    3.5  
Pyrogallol   24.0 11.8    12.3   34.5  76.0  6.3 
Syringaldazine 51.6 120.6    79.2 131.7  126.5       
Syringic acid     115.2    46.9 14.1      
Vanillic acid  61.8       7.3 8.1      
Substrate Species 
Am Ct Ch Cy Ds Ma Me Mv Pr Pe Pc Sr Ts Tt Tv 
2,6-Dimethoxyphenol 100.0 100.0 100.0 100.0 100.0 100.0 100.0 100.0 100.0 100.0 100.0 100.0 100.0 100.0 100.0 
2-Amino-3-(3,4-dihydroxyphenyl)propanoic acid  55.9           46.0   
4-Aminophenol          109.8    26.7  
4-Aminophenylamine 62.9       14.7  194.8   56.0   
4-Hydroxyindole   87.6        107.6     
4-Methoxyphenol         7.3 9.8    19.8  
4-Methylcatechol   69.3 31.4       103.5     
ABTS  76.5 271.7 114.4  800.0 288.3 1.4 97.4 284.1 452.0 136.7   27.7 
Caffeic acid        18.6     95.0 80.2 24.5 
Catechol 74.2  44.9  59.2   34.9 33.1 21.6 74.3 18.3 76.0 27.9 110.7 
Ferulic acid  111.8 48.4      8.5  76.0   116.3  
Guaiacol 59.7 73.5 107.8 37.9 92.0 122.4 31.0 39.1 9.7  140.9 35.0 64.0 55.8 38.4 
Hydroquinone  50.0 105.3  80.8   12.9  25.5 62.0  69.0 30.2 56.0 
N,N-dimethyl-1,4-phenylenediamine 74.8  1.8     23.4   19.9    237.1 
o -Anisidine          45.5     9.3  
p -Anisidine           19.3    3.5  
Pyrogallol   24.0 11.8    12.3   34.5  76.0  6.3 
Syringaldazine 51.6 120.6    79.2 131.7  126.5       
Syringic acid     115.2    46.9 14.1      
Vanillic acid  61.8       7.3 8.1      

Am, Armillaria mellea ( Rehman & Thurston, 1992 ); Ct, Chaetomium thermophilum ( Chefetz, 1998 ); Cy, Cyathus stercoreus ( Sethuraman, 1999 ); Ds, Dichomitus squalens c1 ( Perie, 1998 ); Ma, Mauginiella sp. ( Palonen, 2003 ); Me, Melanocarpus albomyces ( Kiiskinen, 2002 ); Mv, Myrothecium verrucaria ( Sulistyaningdyah, 2004 ); Pr, Phellinus ribis ( Min, 2001 ); Pe, Pleurotus eryngii ( Munoz, 1997 ); Pc, Pycnoporus cinnabarinus ( Eggert, 1996 ); Sr, Sclerotium rolfsii SRL1 ( Ryan, 2003 ); Ts, Trametes sanguinea ( Nishizawa, 1995 ); Tt, Trametes trogii ( Garzillo, 1998 ); Tv, Trametes versicolor ( Sulistyaningdyah, 2004 ).

In addition to the oxidation of phenols, laccases have also been recently demonstrated to catalyze the oxidation of Mn 2+ in the presence of chelators. Laccase from the white-rot fungus T. versicolor oxidized Mn 2+ to Mn 3+ in the presence of pyrophosphate ( Höfer & Schlosser, 1999 ). The same was also later demonstrated for the enzyme of the litter-decomposer Stropharia rugosoannulata with oxalic and malonic acids as chelators ( Schlosser & Höfer, 2002 ). The chelators probably decrease the high redox potential of the Mn 2+ /Mn 3+ couple. Mn 2+ oxidation involved concomitant reduction of laccase type-1 copper, thus providing evidence that it occurs via one-electron transfer to type-1 copper as usual for substrate oxidation by blue laccases ( Schlosser & Höfer, 2002 ). A P. ribis laccase devoid of type-1 copper was unable to catalyze the same reaction ( Min, 2001 ). It was proposed that laccase and Mn-peroxidase can co-operate. In the presence of Mn 2+ and oxalate, laccase produces Mn 3+ -oxalate. The latter initializes a set of follow-up reactions leading to H 2 O 2 formation, which may initiate or support peroxidase reactions ( Schlosser & Höfer, 2002 ). The production of H 2 O 2 and Mn 3+ was also described in P. eryngii for the oxidation of hydroquinone ( Munoz, 1997 ).

Fungal laccases typically exhibit pH optima in the acidic pH range. While the pH optima for the oxidation of ABTS are generally lower than 4.0, phenolic compounds like DMP, guaiacol and syringaldazine exhibit higher values of between 4.0 and 7.0 ( Table 2 ). pH optima of different fungal enzymes for hydroquinone and catechol are 3.6–4.0 and 3.5–6.2, respectively ( Lalitha Kumari & Sirsi, 1972 ; Shleev, 2004 ). It was proposed that the bell-shaped pH profile of phenolic compounds is formed by two opposing effects. The oxidation of phenols depends on the redox potential difference between the phenolic compound and the T1 copper ( Xu, 1996 ). The E0 of a phenol decreases when pH increases due to the oxidative proton release. At a rate of Δ E /ΔpH=59 mV at 25°C, a pH change from 2.7 to 11 would result in a E0 decrease of 490 mV for the phenol. However, over the same pH range, the E0 changes for the fungal laccases studied ( T. villosa, R. solani and M. thermophila ) were much smaller (≤100 mV) ( Xu, 1997 ). The enzyme activity at higher pH is decreased by the binding of a hydroxide anion to the T2/T3 coppers of laccase that interrupts the internal electron transfer from T1 to T2/T3 centres ( Munoz, 1997 ). Not only the rate of oxidation but also the reaction products can differ according to pH as pH may affect abiotic follow-up reactions of primary radicals formed by laccase. Laccases from Rhizoctonia praticola and T. versicolor formed different products from syringic and vanillic acids at different pH values, but both enzymes generated the same products at a particular pH ( Xu, 1997 ). The stability of fungal laccases is generally higher at acidic pH ( Leonowicz, 1984 ), although exceptions exist ( Mayer, 1987 ; Baldrian, 2004 ).

Temperature profiles of laccase activity usually do not differ from other extracellular ligninolytic enzymes with optima between 50° and 70°C ( Table 2 ). Few enzymes with optima below 35°C have been described, e.g. the laccase from G. lucidum with the highest activity at 25°C ( Ko, 2001 ). This has, however, no connection with the growth optimum of the fungi. The temperature stability varies considerably. The half life at 50°C ranges from minutes in B. cinnerea ( Slomczynski, 1995 ), to over 2–3 h in L. edodes and A. bisporus ( Wood, 1980 ; D'Annibale, 1996 ), to up to 50–70 h in Trametes sp. ( Smirnov, 2001 ). While the enzyme from G. lucidum was immediately inactivated at 60°C, the thermostable laccase from M. albomyces still exhibited a half life of over 5 h and thus a very high potential for selected biotechnological applications ( Lalitha Kumari & Sirsi, 1972 ; Kiiskinen, 2002 ).

A very wide range of compounds has been described to inhibit laccase ( Table 3 ). In addition to the general inhibitors of metal-containing oxidases like cyanide, sodium azide or fluoride, there are some selective inhibitors for individual oxidases. Carbon monoxide, 4-hexylresorcinol or salicylhydroxamic acid are examples of specific inhibitors of tyrosinases but not laccases ( Petroski, 1980 ; Allen & Walker, 1988 ; Dawley & Flurkey, 1993 ) that may facilitate estimation of laccase activity when protein purification is not successful. Inhibition by diethyl dithiocarbamate and thioglycolic acid could be supposed to be due to the presence of copper in the catalytic centre of the enzyme, and several sulfhydryl organic compounds have been described as laccase inhibitors: e.g. dithiothreitol, thioglycolic acid, cysteine and diethyldithiocarbamic acid. However, experiments with T. versicolor laccase showed that the inhibitory effect found with these compounds is probably due to the methodology using ABTS as the enzyme substrate ( Johannes & Majcherczyk, 2000 ) and that these compounds, contrary to sodium azide, do not decrease the oxygen consumption by laccase during the catalysis.

Given the natural occurrence of laccases in soil, the inhibition by heavy metals and humic substances must be taken into account ( Zavarzina, 2004 ). While some laccases exhibit a sensitivity towards heavy metals ( Table 3 ), others, e.g. the enzyme from G. lucidum , are completely insensitive ( Lalitha Kumari & Sirsi, 1972 ). In the complex environment of soil or decaying lignocellulosic material, many different substrates of laccase are usually present that can compete for the oxidation and thus competitively inhibit the transformation of other compounds ( Itoh, 2000 ). Thus it is difficult to estimate the transformation rates of laccase substrates in soils based on laboratory results and these rates can significantly differ in different soils.

Some low molecular weight compounds that can be oxidized by laccase to stable radicals can act as redox mediators, oxidizing other compounds that in principle are not substrates of laccase due to its low redox potential. In addition to enabling the oxidation of compounds that are not normally oxidized by laccases (e.g. the nonphenolic lignin moiety), the mediators can diffuse far away from the mycelium to sites that are inaccessible to the enzyme itself. Several compounds involved in the natural degradation of lignin by white-rot fungi may be derived from oxidized lignin units or directly from fungal metabolism and act as mediators ( Camarero, 2005 ). ( Eggert, 1996 ) proposed that 3-hydroxyanthranilate can be a mediator of lignin degradation by P. cinnabarinus , the fungus lacking ligninolytic peroxidases. Other naturally occurring mediators include phenol, aniline, 4-hydroxybenzoic acid and 4-hydroxybenzyl alcohol ( Johannes & Majcherczyk, 2000 ).

Recently, some phenols, including syringaldehyde and acetosyringone, have been described as laccase mediators for indigo decolorization ( Campos, 2001 ) as well as for the transformation of the fungicide cyprodinil ( Kang, 2002 ) and hydrocarbon degradation ( Johannes & Majcherczyk, 2000 ). A comprehensive screening for natural mediators was performed by ( Camarero, 2005 ). Among 44 tested natural lignin-derived compounds they selected 10 phenolic compounds derived from syringyl, guaiacyl, and p -hydroxyphenyl lignin units, characterized by the presence of two, one or no methoxy substituents, respectively (in ortho positions with respect to the phenolic hydroxyl). Syringaldehyde, acetosyringone, vanillin, acetovanillone, methyl vanillate and p -coumaric acid have been found to be the most effective for mediated oxidation using laccases of P. cinnabarinus and T. villosa . Among them, syringaldehyde and acetosyringone belong to the main products of both biological and enzymatic degradation of syringyl-rich lignins ( Kirk & Farrell, 1987 ).

Laccases in the natural environment

The considerable attention devoted to white-rot basidiomycetes and their ligninolytic system in the past might lead to the conclusion that decaying wood is the most typical environment for laccase production. The possible mechanisms involved in lignocellulose degradation by laccases have been studied in detail and a comprehensive recent review is available on this topic ( Leonowicz, 2001 ). Far less is known about the occurrence, properties and roles of laccases occurring in other types of natural lignocellulose-containing material like forest litter or soil. Compared to wood, soil or litter is a more complex and heterogeneous environment, which may hamper the detection and estimation of enzyme activities. Another problem is to link the enzyme activities in soil to a specific species producing it, if this is at all possible. Several works [e.g. ( Lang, 1997 , 1998 ; Baldrian, 2000 )] followed the production of enzymes by fungi introduced into soils and a number of protocols for laccase extraction have been proposed to optimize the extraction yield. These include direct incubation with guaiacol as laccase substrate ( Nannipieri, 1991 ), extraction with surfactants or calcium chloride ( Criquet, 1999 ) or the most widely used extraction with phosphate buffer ( Lang, 1997 ), depending on the nature of the substrate (forest litter, agricultural soil, compost).

Relatively high activities of laccase – compared to agricultural or meadow soils – can be detected in forest litter and soils in both broadleaved ( Rosenbrock, 1995 ; Criquet, 2000 ; Carreiro, 2000 ) and coniferous forests ( Ghosh, 2003 ), where laccase is the dominant ligninolytic enzyme ( Criquet, 2000 ; Ghosh, 2003 ). The laccase activity reflects the course of the degradation of organic substances and thus it varies with time. Laccase activity was found to increase during leaf litter degradation in Mediterranean broadleaved litter ( Fioretto, 2000 ) and the pattern of detected isoenzymes varied during the succession ( Di Nardo, 2004 ). In evergreen oak litter, laccase activity was found to reflect the annual dynamics of fungal biomass that is probably driven by the seasonal drying ( Criquet, 2000 ). The annual variation of laccase activity in temperate forests is also great and probably reflects the seasonal input of fresh litter (P. Baldrian, unpublished data). The activity of laccase also reflects the presence of fungal mycelia. Significantly increased laccase activity was detected in the fairy rings of Marasmius oreades along with the production of organic acids and a high concentration of available nitrogen and carbon due to the degradation of plant roots by the fungus ( Gramss, 2005 ). Along with the vertical gradient of fungal distribution in soil profiles, the laccase activity decreases with increasing depth. The decrease of laccase activity is also reflected in the decrease of laccase gene diversity with soil depth ( Chen, 2003 ).

Laccases as the most abundant ligninolytic enzymes in soil also attracted the attention of ecologists studying its role in the carbon cycle, especially with respect to the nitrogen input. Several studies documented a significant decrease of laccases and peroxidases in forest soils subjected to elevated nitrogen doses ( Carreiro, 2000 ; Gallo, 2004 ), with the simultaneous increase in the litter layer ( Saiya-Cork, 2002 ). This phenomenon was accompanied by the decrease of fungal biomass and the fungal: bacterial biomass ratio in soil as well as by increased incorporation of vanillin as a model lignin-derived substrate into fungal biomass; hence it seems that nitrate deposition directs the flow of carbon through the heterotrophic soil food web ( DeForest, 2004 , 2004 ). On the other hand, an increase of phenolic compounds in forest soil after burning increased laccase activity ( Boerner & Brinkman, 2003 ).

Similar to the situation in other lignocellulose-containing substrates, laccases probably also participate in the transformation of lignin contained in the forest litter. It is also generally presumed that laccases are able to react with soil humic substances that can be directly formed from lignin ( Yavmetdinov, 2003 ). This is supported by the fact that humic acids induce laccase activity and mRNA expression ( Scheel, 2000 ). However, the interaction of laccases with humic substances probably leads both to depolymerization of humic substances and their synthesis from monomeric precursors; the balance of these two processes can be influenced by the nature of the humic compounds ( Zavarzina, 2004 ). ( Fakoussa & Frost, 1999 ) observed the decolorization and decrease of molecular weight of humic acids, accompanied by the formation of fulvic acids during the growth of T. versicolor cultures producing mainly laccase, and humic acid synthesis was also documented in vitro using the same enzyme ( Katase & Bollag, 1991 ). Adsorption of laccases to soil humic substances or inorganic soil constituents changes their temperature and activity profiles ( Criquet, 2000 ) and inhibits its activity ( Claus & Filip, 1990 ). ( Zavarzina, 2004 ) estimated inhibition constants for humic acids towards Panus tigrinus laccase. The K i ranged from 0.003 μg mL −1 for humic acids from peat soils to 0.025 μg mL −1 for humic acids from chernozems. Recently, ( Keum & Li, 2004 ) demonstrated that humic substances do not strongly bind laccase and the inactivation is thus not due to binding but to the dissociation of copper that is chelated by humic substances. This introduces another difficulty for the determination of laccase activities in soil or forest litter, as different extraction methods extract different amounts of humic acids together with soil proteins – enzymes ( Criquet, 1999 ).

Laccases are also actively produced during the composting process. Of 34 isolates of fungi from woody compost, 11 were able to oxidize syringaldazine ( Chamuris, 2000 ). Laccase was isolated both from compost-specific fungi and the compost itself ( Chefetz, 1998 , 1998 ) and it seems that it participates in both degradation of lignin and humic acids and humic acid formation ( Chefetz, 1998 ; Kluczek-Turpeinen, 2003 , 2005 ). In water-saturated environments, laccase activity is driven by the concentration of oxygen. Laccase activity in peatlands is thus low due to low oxygen availability ( Pind, 1994 ; Williams, 2000 ) but increases dramatically when the oxygen concentration increases. The burst of laccase activity can lead to the depletion of phenolic compounds that inhibit organic matter degradation by oxidative and hydrolytic enzymes ( Freeman, 2004 ) and it can be assumed that the oxygen-regulated laccase activity plays an important role in carbon cycling in this environment. In the water environment, laccase was demonstrated to participate in the degradation of wood as well as humic substances ( Claus & Filip, 1998 ; Hendel & Marxsen, 2000 ). Its activity is dependent on the succession step of substrate decay and it can exhibit a seasonal pattern of activity dependent on the input of its substrate ( Artigas, 2004 ).

Although the breakdown of lignin and the metabolism of humic acids may be the main ecological processes where laccases are involved, there are probably more roles that these enzymes can play. One of them is the interaction of fungi with different microorganisms including soil fungi (e.g. Trichoderma sp.) and bacteria, a process usually accompanied by a strong induction of laccase ( Freitag & Morrell, 1992 ; Savoie, 1998 ; Savoie, 2001 ; Velazquez-Cedeno, 2004 ) that is probably general for laccase-producing basidiomycetes ( Iakovlev & Stenlid, 2000 ; Baldrian, 2004 ) but was also demonstrated in R. solani challenged with Pseudomonas strains producing antifungal compounds ( Crowe & Olsson, 2001 ). Since laccase and their products do not have a direct effect on soil bacteria or fungi ( Baldrian, 2004 ) it is probably involved in the passive defence by the formation of melanins or similar compounds ( Eggert, 1995 ; Baldrian, 2003 ). Laccase can probably also contribute to the degradation of phenolic antibiotics that inhibit fungal growth like 2,4-diacetylphloroglucinol. The role of laccases in the defence against heavy metals was also proposed in spite of the fact that different heavy metals induce its activity and is connected with the production of melanins ( Galhaup & Haltrich, 2001 ; Baldrian, 2003 ).

Laccases in environmental biotechnology

Laccases offer several advantages of great interest for biotechnological applications. They exhibit broad substrate specificity and are thus able to oxidize a broad range of xenobiotic compounds including chlorinated phenolics ( Royarcand & Archibald, 1991 ; Roper, 1995 ; Ullah, 2000 ; Schultz, 2001 ; Bollag, 2003 ), synthetic dyes ( Chivukula & Renganathan, 1995 ; Rodriguez, 1999 ; Wong & Yu, 1999 ; Abadulla, 2000 ; Nagai, 2002 ; Claus, 2002 ; Soares, 2002 ; Peralta-Zamora, 2003 ; Wesenberg, 2003 ; Zille, 2003 ), pesticides ( Nannipieri & Bollag, 1991 ; Jolivalt, 2000 ; Torres, 2003 ) and polycyclic aromatic hydrocarbons ( Johannes, 1996 ; Collins, 1996 ; Majcherczyk, 1998 ; Majcherczyk & Johannes, 2000 ; Cho, 2002 ; Pozdnyakova, 2004 ). They can bleach Kraft pulp ( Reid & Paice, 1994 ; Paice, 1995 ; Bourbonnais & Paice, 1996 ; Call & Mucke, 1997 ; Monteiro & de Carvalho, 1998 ; de Carvalho, 1999 ; Sealey, 1999 ; Balakshin, 2001 ; Lund, 2003 ; Sigoillot, 2004 ) or detoxify agricultural byproducts including olive mill wastes or coffee pulp ( D'Annibale, 2000 ; Tsioulpas, 2002 ; Velazquez-Cedeno, 2002 ) (for review see Durán & Esposito, 2000 ; Durán, 2002 ; Mayer & Staples, 2002 ). Unlike ligninolytic peroxidases they use molecular oxygen, which is usually available in the reaction system as the final electron acceptor, instead of hydrogen peroxide, which that must be produced by the fungus. Laccases are usually constitutively produced in at least some stages of the growth of a particular fungus. They can be extracted from lignocellulosic substrates colonized by fungi as well as from soil or forest litter, or used in the form of spent substrate from the cultivation of edible mushrooms ( Eggen, 1999 ; Lau, 2003 ; Law, 2003 ). The possibility of increasing the production of laccase by the addition of inducers to fungal cultures and a relatively simple purification process are other advantages. Last but not least, the considerable amount of data concerning the properties of fungal laccases accumulated in the past years allows us to select a protein suitable for a specific application (e.g. temperature-resistant or pH-stable).

However, the low redox potential of laccases (450–800 mV) compared to those of ligninolytic peroxidases (>1 V) only allows the direct degradation of low-redox-potential compounds and not the oxidation of more recalcitrant aromatic compounds, including some synthetic dyes or polycyclic aromatic hydrocarbons (PAH) ( Xu, 1996 ), although there is some evidence that PAH can be oxidized by some laccases to a considerable degree; yellow laccase from P. ostreatus (YLPO) degraded PAH anthracene (95% within 2 days) and fluoranthene (14% within 2 days) with an optimum pH of 6.0 without redox mediators ( Pozdnyakova, 2004 ), whereas ‘blue’ laccases from other fungi were not capable of efficient oxidation ( Johannes, 1996 ; Majcherczyk, 1998 ; Kottermann, 1998 ). The compounds with higher redox potential can only be transformed if the reaction product is subject to an immediately following reaction or when its redox potential is lowered, for instance by chelation.

Another possibility for the oxidation of compounds with high redox potentials is the use of redox mediators. From the description of the first laccase mediators, ABTS ( Bourbonnais & Paice, 1990 ), to the more recent use of the -NOH- type, synthetic mediators such as 1-hydroxybenzotriazole, violuric acid and N-hydroxyacetanilide or TEMPO, a large number of studies have been performed on the mechanisms of oxidation of nonphenolic substrates ( Bourbonnais, 1998 ; Xu, 2000 ; Fabbrini, 2002 ; Baiocco, 2003 ), the search for new mediators ( Bourbonnais, 1997 ; Fabbrini, 2002 ), and their applications in the degradation of aromatic xenobiotics ( Bourbonnais, 1997 ; Johannes, 1998 ; Kang, 2002 ; Keum & Li, 2004 ). Nevertheless, the laccase-mediator system has yet to be applied on the process scale due to the cost of mediators and the lack of studies that guarantee the absence of toxic effects of these compounds or their derivatives. The use of naturally occurring laccase mediators would present environmental and economic advantages. Their capability to act as laccase mediators has recently been demonstrated. The possibility of obtaining mediators from natural sources and the low mediator/substrate ratios of 1–4 ( Camarero, 2005 ) or 20–40 ( Eggert, 1996 ; Campos, 2001 ) increase the feasibility of the laccase-mediator system for use in biotechnology.

In addition to substrate oxidation, laccase can also immobilize soil pollutants by coupling to soil humic substances – a process analogous to humic acid synthesis in soils ( Bollag, 1991 ; Bollag & Myers, 1992 ). The xenobiotics that can be immobilized in this way include phenolic compounds and anilines such as 3,4-dichloroaniline, 2,4,6-trinitrotoluene or chlorinated phenols ( Tatsumi, 1994 ; Dawel, 1997 ; Dec & Bollag, 2000 ; Ahn, 2002 ). The immobilization lowers the biological availability of the xenobiotics and thus their toxicity.

The current development in laccase catalysis research and the design of mediators along with the research on its heterologous expression opens a wide spectrum of possible applications in the near future. Moreover, laccase can also offer a simple and convenient alternative to supplying peroxidases with H 2 O 2 , because laccases are available on an economically feasible scale.

Conclusions

This review summarizes the available data about the biochemical properties of fungal laccases, their occurrence under natural conditions and possible biotechnological use. However, it leaves many important questions open: Why do fungi produce laccase? What are the respective roles of different isoenzymes? Do their biochemical characteristics differ with respect to their function? The understanding of the physiological role of laccase has not increased significantly since it was reviewed by ( Thurston, 1994 ) and ( Mayer & Staples, 2002 ). The problem is its low substrate specificity and a very wide range of reactions that it can potentially catalyze. Despite many efforts to address the involvement of laccase in the transformation of lignocellulose, it is still not completely clear how important a role laccase plays in lignin degradation and if it contributes more to the formation or decomposition of humic substances in soils. In this sense it is even more difficult to estimate its involvement and role in carbon cycling or during interspecific interactions of soil fungi. Hopefully, these questions will attract more attention of researchers in the future.

Acknowledgements

This work was supported by the Grant Agency of the Czech Academy of Sciences (B600200516), by the Grant Agency of the Czech Republic (526/05/0168) and by the Institutional Research Concept no. AV0Z50200510 of the Institute of Microbiology, ASCR.

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