Fungal laccases â occurrence and properties - Wiley Online Library

Fungal laccases ^ occurrence and properties Petr Baldrian Laboratory of Biochemistry of Wood-Rotting Fungi, Institute of Microbiology ASCR, Prague, Czech Republic

Correspondence: Petr Baldrian, Laboratory of Biochemistry of Wood-Rotting Fungi, Institute of Microbiology ASCR, V´ıdenˇska´ 1083, 14220 Prague 4, Czech Republic. Tel.: 1420 2410 62315; fax: 1420 2410 62384; e-mail: [email protected] Received 27 May 2005; revised 1 September 2005; accepted 1 September 2005. First published online 9 November 2005. doi:10.1111/j.1574-4976.2005.00010.x

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.

Editor: Jiri Damborsky Keywords biotechnology; ecology; humic substances; laccase; lignin; soil; wood-rotting fungi.

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 monopheFEMS Microbiol Rev 30 (2006) 215–242

nol 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 0 -bis(3,5-dimethoxy-4-hydroxybenzylidene hydrazine)] is often considered to be a unique laccase substrate (Harkin et al., 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 et al., 1992; Liu et al., 1994; Boudet, 2000; Ranocha et al., 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 et al., 2002; Claus, 2003; Givaudan et al., 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 2005 Federation of European Microbiological Societies Published by Blackwell Publishing Ltd. All rights reserved

c

216

point at pH 6.2 and the ability to oxidize syringaldazine (Rosconi et al., 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 et al., 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 et al., 1996, 2001). An overview of technological applications of oxidases including laccase was published by (Dura´ n & Esposito, 2000) and (Dura´ n et al., 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 ascomy2005 Federation of European Microbiological Societies Published by Blackwell Publishing Ltd. All rights reserved

c

P. Baldrian

cetes such as Gaeumannomyces graminis (Edens et al., 1999), Magnaporthe grisea (Iyer & Chattoo, 2003) and Ophiostoma novo-ulmi (Binz & Canevascini, 1997), as well as from Mauginella (Palonen et al., 2003), Melanocarpus albomyces (Kiiskinen et al., 2002), Monocillium indicum (Thakker et al., 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 et al., 1996; Scherer & Fischer, 1998), as well as some freshwater ascomycetes (AbdelRaheem & Shearer, 2002; Junghanns et al., 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 et al., 2000), only some strains of Trichoderma exhibit a low level of production of a syringaldazine-oxidizing enzyme (Assavanig et al., 1992), mainly associated with spores, which may act in the morphogenesis of this fungus (Assavanig et al., 1992; Holker et al., 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 et al., 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 et al., 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 et al., 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 et al., 2001; FEMS Microbiol Rev 30 (2006) 215–242

217

Fungal laccases – occurrence and properties

Ikeda et al., 2003). A homologous gene has also been demonstrated in Cryptococcus podzolicus but not in other heterobasidiomycetous yeasts tested (Petter et al., 2001) and there are some records of low laccase-like activity in some yeast species isolated from decayed wood (Jimenez et al., 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 Cu1 (Machonkin et al., 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 et al., 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 whiterot fungi was detected in Gloeophyllum trabeum. Oxidation of ABTS (2,2 0 -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 et al., 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 et al., 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. 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 woodrotting fungi have been found in several isolates of ECM species including Amanita, Cortinarius, Hebeloma, Lactarius, Paxillus, Piloderma, Russula, Tylospora and Xerocomus (Luis et al., 2004; Chen et al., 2003). In the case of Piloderma byssinum, transcription of putative laccase sequence was confirmed by RT-PCR (Chen et al., 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 syringaldaFEMS Microbiol Rev 30 (2006) 215–242

zine has never been detected (Günther et al., 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 et al., 1998, 1999). The gene sequences are not found very frequently either (Chen et al., 2003). Laccases have been purified from a few fungi-forming ectomycorrhiza: Cantharellus cibarius (Ng & Wang, 2004), Lactarius piperatus (Iwasaki et al., 1967), Russula delica (Matsubara & Iwasaki, 1972) and Thelephora terestris (Kanunfre & Zancan, 1998) or orchideoid mycorrhiza: Armillaria mellea (Rehman & Thurston, 1992; Billal & Thurston, 1996; Curir et al., 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 et al., 1999; Chen et al., 2003) and possibly also lignin peroxidase (Chen et al., 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 whiterot 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 et al., 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 et al., 1997) and Suillus granulatus (Günther et al., 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 et al., 1992, 1993; Palmieri et al., 2000). The extracellular laccase activity of Lentinula edodes was associated with a multicomponent protein complex of 2005 Federation of European Microbiological Societies Published by Blackwell Publishing Ltd. All rights reserved

c

c

Agaricus bisporus Agaricus bisporus Agaricus blazei Agrocybe praecox Albatrella dispansus Armillaria mellea Lac I Armillaria mellea Lac II Armillaria mellea Aspergillus nidulans II Botrytis cinerea Cantharellus cibarius Ceriporiopsis subvermispora L1 Ceriporiopsis subvermispora L2 Cerrena maxima Cerrena unicolor Cerrena unicolor Chaetomium termophilum Chalara paradoxa Colletotrichum graminicola Coniothyrium minitans Coprinus cinereus Coprinus friesii Coriolopsis fulvocinnerea Coriolopsis gallica Coriolopsis rigida I Coriolopsis rigida II Coriolus hirsutus Coriolus hirsutus Coriolus maxima Coriolus zonatus Cryptococcus neoformans Cyathus stercoreus Daedalea quercina Dichomitus squalens c1 Dichomitus squalens c2 Fomes fomentarius Ganoderma lucidum Ganoderma tsugae Gaeumannomyces graminis Hericium echinaceum

Species

2005 Federation of European Microbiological Societies Published by Blackwell Publishing Ltd. All rights reserved

190 63

67

80 80 74 92 71 68 57–67 66 58 77 67 85 74 58 60 54–65 84 66 66 55 78 57 60 77 70 69 66 66

96 65 66 66 62 59

MW (kDa)

5.6

3.5 3.0 3.5 3.6

4.6

4.0 4.0 3.5 3.5 4.2–4.3 3.9 3.9 4.0 4.2

5.1

3.4 4.8 3.5 4.0

4.0

3.1

4.1

4.0 4.0

pI

Table 1. Characteristics of laccases purified from fungi

5.0

4.5

4.0 3.0 3.0

3.0 3.0

3.0 2.5 2.5

4.8 2.0

8.0

3.5

4.5

4.0 4.0

6.5 3.5

3.5

5.5

DMP

4.0 5.0

4.5

4.0 3.0 3.0

4.0

2.0

ABTS

pH optimum

4.5

6.5

3.0 5.0

5.6

GUA

7.0

6.5

6.5 6.0

6.0

SYR

4

38

8

12 11

26 41

190 770

30 20

63

ABTS

26

48

328 348

100

96 14 720

2900 7700

100

178

1026

DMP

Km (mM)

510

93

70–90

400 10 230

1600 440 160–300

4307

GUA

131

34 3400 214

4

SYR

50

52 25

70, 55

55

45

70

60 60–70

60

50

55 57 50

70

Temperature optimum (1C) Reference

Wood (1980) Perry et al. (1993) Ullrich et al. (2005) Steffen et al. (2002) Wang & Ng (2004b) Rehman & Thurston (1992) Billal & Thurston (1996) Curir et al. (1997) Scherer & Fischer (1998) Slomczynski et al. (1995) Ng & Wang (2004) Fukushima & Kirk (1995); Wang & Ng (2004b) Fukushima & Kirk, (1995); Wang & Ng (2004b) Koroleva et al. (2001); Shleev et al. (2004) Bekker et al. (1990) Kim et al. (2002) Chefetz et al. (1998) Robles et al. (2002) Anderson & Nicholson (1996) Dahiya et al. (1998) Schneider et al. (1999) Heinzkill et al. (1998) Shleev et al. (2004); Smirnov et al. (2001) Calvo et al. (1998) Saparrat et al. (2002) Saparrat et al. (2002) Koroljova-Skorobogat’ko et al. (1998) Lee & Shin (1999) Smirnov et al. (2001) Koroljova et al. (1999) Williamson (1994) Sethuraman et al. (1999) Baldrian (2004) Perie et al. (1998) Perie et al. (1998) Rogalski et al. (1991) Lalitha Kumari & Sirsi (1972); Ko et al. (2001) Eller et al. (1998) Edens et al. (1999) Wang & Ng (2004c)

218 P. Baldrian

FEMS Microbiol Rev 30 (2006) 215–242

Junghuhnia separabilima Lactarius piperatus Lentinula edodes Lcc1 Lentinus edodes Magnaporthe grisea Marasmius quercophilus Marasmius quercophilusw Marasmius quercophilus Marasmius quercophilusz Marasmius quercophilus‰ Mauginiella sp. Melanocarpus albomyces Monocillium indicum Myrothecium verrucaria Neurospora crassa Ophiostoma novo-ulmi Panaeolus papilionaceus Panaeolus sphinctrinus Panus tigrinus Panus tigrinus Phanerochaete flavido-alba Phanerochaete chrysosporium Phellinus noxius Phellinus ribis Phlebia radiata Phlebia tremellosa Pholiota mutabilis Physisporinus rivulosus Lacc 1 Physisporinus rivulosus Lacc 2 Physisporinus rivulosus Lacc 3 Physisporinus rivulosus Lacc 4 Pleurotus eryngii I Pleurotus eryngii II Pleurotus florida Pleurotus ostreatus Pleurotus ostreatus POXA1b Pleurotus ostreatus POXA1w Pleurotus ostreatus POXA2 Pleurotus ostreatus POXA3a Pleurotus ostreatus POXA3b Pleurotus ostreatus POXC Pleurotus pulmonarius Lcc2 Pleurotus sajor-caju IV Podospora anserine


66 67 68 68 65 61 77 67 62 61 67 83–85 83–85 59 46 55 383

58–62 67 72 65 70 60 60 65 65 60 63 80 100 62 64 79 60 60 64 63 94 47 70 152 64 64

c

3.6

3.3 3.3 3.2 3.1 4.1 4.2 4.1 3.6 6.9 6.7 4.0 4.1 4.3 2.9

3.5

2.9–3.0

5.1

4.8–6.4 4.0

4.0–4.4 4.8–5.1 3.6

3.0 3.0

3.4–3.6

3.0 3.0 3.0 3.0

4.5 3.0–5.0 6.5 5.5 5.5 3.0–5.0

3.0 3.0 3.0 3.6 3.6 3.0 4.0–5.5 2.1

4.0–6.0

8.0 7.0

3.5

4.0

2.5 2.5 2.5 2.5 4.5 4.5

5.0

3.0

2.8 3.0 3.0

2.6 4.0 2.4 3.5

4.0

NA 6.0 6.2 6.2 6.0 6.0–8.0

3.5 3.5 3.5 3.5

6.0

4.0 5.0–7.5

4.0

6.0 6.2–6.5

5.8 6.0 6.0 6.0

3.5 3.5 3.5 3.5

6.0

6.0

6.0–7.0

6.0 5.0 5.0 4.5 6.2 4.5

370 90 120 70 74 280 210 92

207

51 32

8 113

108

260 2100 740 14 000 8800 230

1400 400

38

557

1200 550

NA 3100

7600 8000 30 000

917

220 130 140 36 79 20 12

11

7 50 4.2

118

45–65 25–35 35 35 50–60 50

50

55 55

30

65

30 80 80 75 80 80

40

Vares et al. (1992) Iwasaki et al. (1967) Nagai et al. (2002) Kofujita et al. (1991) Iyer & Chattoo (2003) Farnet et al. (2000) Farnet et al. (2000) Dedeyan et al. (2000) Farnet et al., (2002, 2004) Farnet et al. (2004) Palonen et al. (2003) Kiiskinen et al. (2002) Thakker et al. (1992) Sulistyaningdyah et al. (2004) Froehner & Eriksson (1974) Binz & Canevascini (1997) Heinzkill et al. (1998) Heinzkill et al. (1998) Maltseva et al. (1991) Leontievsky et al. (1997) Perez et al. (1996) Srinivasan et al. (1995) Geiger et al. (1986) Min et al. (2001) Vares et al. (1995) Vares et al. (1994) Leonowicz & Malinowska (1982) Hakala et al. (2005) Hakala et al. (2005) Hakala et al. (2005) Hakala et al. (2005) Munoz et al. (1997) Munoz et al. (1997) Das et al. (2000) Hublik & Schinner (2000) Giardina et al. (1999) Palmieri et al. (1997) Palmieri et al. (1997) Palmieri et al. (2003) Palmieri et al. (2003) Palmieri et al. (1993, 1997); Sannia et al. (1986) De Souza & Peralta (2003) Lo et al. (2001) Molitoris & Esser (1970); Durrens (1981)


219


c


58 66 63 81 70 170 55 60 63 62–64 55 86 67 66 66 60 60 64–68 63 64 65 62 70 68 63 63 62 71 43 58

Polyporus anceps Polyporus anisoporus Polyporus pinsitus Pycnoporus cinnabarinus Pycnoporus cinnabarinus Pycnoporus coccineus Rhizoctonia solani 4 Rigidoporus lignosus B Rigidoporus lignosus S Russula delica Schizophyllum commune Sclerotium rolfsii SRL1 Sclerotium rolfsii SRL2 Stropharia coronilla Stropharia rugosoannulata Thelephora terrestris Trametes gallica Lac I Trametes gallica Lac II Trametes hirsute Trametes multicolor II Trametes ochracea Trametes pubescens LAP 2 Trametes sanguinea Trametes trogii Trametes versicolor Trametes villosa 1 Trametes villosa 3 Trametes sp. AH28-2 A Trichoderma sp. Tricholoma giganteum Volvariella volvacea 3.7

3.5 6.0–6.5 4.2

3.1 3.0 3.7–4.0 3.0 4.7 2.6 3.5 3.3; 3.6

4.0 3.0

2.5 2.7 2.7

2.5 3.4 2.2 2.2

2.4

5.2 4.4

3.0 3.0

3.0

ABTS

3.7 3.1

3.0 3.7

3.4

pI

4.6

3.5

3.0 3.0

3.5

6.2 6.2

5.0

DMP

pH optimum

4.5

4.8 4.0 4.0

4.0–4.5 4.0

GUA

5.6

4.0 5.0–5.5 5.0–5.5

5.0

4.4–5.0

5.0–5.5

SYR

410 15

30 37

30

570

25

72

14

25

420 410

480 108

DMP

16 12 9

80 49

22

ABTS

Km (mM) GUA

420

90 360

121 405 400 63

330

10

6

3

30

SYR

70 45

50

55

45 70 70

62

Temperature optimum (1C)

Petroski et al. (1980) Vaitkyavichyus et al. (1984) Heinzkill et al. (1998) Schliephake et al. (2000) Eggert et al. (1996) Oda et al. (1991) Iwasaki et al. (1967) Bonomo et al. (1998) Bonomo et al. (1998) Matsubara & Iwasaki (1972) De Vries et al. (1986) Ryan et al. (2003) Ryan et al. (2003) Steffen et al. (2002) ¨ Schlosser & Hofer (2002) Kanunfre & Zancan (1998) Dong & Zhang (2004) Dong & Zhang (2004) Shleev et al. (2004); Vares & Hatakka (1997) Leitner et al. (2002) (Shleev et al. (2004) Galhaup et al. (2002) Nishizawa et al. (1995) Garzillo et al. (1998) ¨ Rogalski et al. (1990); Hofer & Schlosser (1999) Yaver et al. (1996) Yaver et al. (1996) Xiao et al. (2003) Assavanig et al. (1992) Wang & Ng (2004a) Chen et al. (2004)

Reference

ABTS, 2,2 0 -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. w Strain 17, induced with p-hydroxybenzoic acid. z Strain C7, constitutive form. ‰ Strain 19, induced with ferulic acid.

MW (kDa)

Species

Table 1. Continued.

220 P. Baldrian


221


660 kDa, which also exhibited peroxidase and b-glucosidase activities (Makkar et al., 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 (Vala´ sˇkova´ & Baldrian, 2005). Laccase activity is almost exclusively associated with cell walls in the white-rot basidiomycete Irpex lacteus (Svobodova´ , 2005), the yeast C. neoformans (Zhu et al., 2001) and in the spores of Trichoderma spp. (Holker et al., 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 et al., 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. 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 et al., 1986; Palmieri et al., 1993, 1997, 2003; Giardina et al., 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

Table 2. Properties of fungal laccases (data derived from Table 1) Property

n

Median

Q25

Q75

Min

Max

Molecular weight (Da) pI Temperature optimum ( 1C) pH optimum ABTS 2,6-Dimethoxyphenol Guaiacol Syringaldazine KM (mM) ABTS 2,6-Dimethoxyphenol Guaiacol Syringaldazine kcat (s 1) ABTS 2,6-Dimethoxyphenol Guaiacol Syringaldazine

103 67 39 49

66 000 3.9 55 3.0

61 000 3.5 50 2.5

71 000 4.2 70 4.0

43 000 2.6 25 2.0

383 000 6.9 80 5.0

36 24 31

4.0 4.5 6.0

3.0 4.0 4.7

5.5 6.0 6.0

3.0 3.0 3.5

8.0 7.0 7.0

36 30 23 21

39 405 420 36

18 100 121 11

100 880 1600 131

4 26 4 3

770 14 720 30 000 4307

12 12 10 4

24 050 3680 295 21 500

5220 815 115 18 400

41 460 6000 3960 25 500

198 100 90 16 800

350 000 360 000 10 800 28 000

n, number of observations; Q25, lower quartile; Q75, upper quartile.



c

222

cleaved by an extracellular protease (Palmieri et al., 1997; Giardina et al., 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 et al., 2000, 2003). Isoenzymes of laccase with different molecular weight and pI were also detected in the litter-decomposing fungus Marasmius quercophilus (Farnet et al., 2000, 2002, 2004; Dedeyan et al., 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 SDSPAGE) after the induction of laccase with different aromatic compounds (Farnet et al., 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 et al., 1997); however, since laccase gene was not found in the complete genome sequence of this fungus (Martinez et al., 2004), these are probably multicopper oxidases rather than true laccases (Larrondo et al., 2003). The molecular basis for the production of different isoenzymes is the presence of multiple laccase genes in fungi (see e.g. Chen et al., 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 et al., 2001), Pleurotus pulmonarius (De Souza & Peralta, 2003) and Trametes villosa (Yaver et al., 1996), the mycorrhizal fungus C. cibarius (Ng & Wang, 2004) and the ascomycete Rhizoctonia solani (Wahleithner et al., 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 et al., 1992): G. graminis produces a trimer of three 60-kDa subunits (Edens et al., 1999); P. anserina laccase is a heterooligomer (Molitoris & Esser, 1970); and one of the laccases purified from A. mellea has a heterodimeric structure (Curir et al., 1997). According to Wood (Wood, 1980), A. bisporus laccase consists of several polypeptides of 23–56 kDa. (Perry et al., 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 2005 Federation of European Microbiological Societies Published by Blackwell Publishing Ltd. All rights reserved

c

P. Baldrian

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 et al., 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 et al., 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 et al., 1995). Other preparations from the same species exhibited as much as 65–80% of saccharides including arabinose, xylose, mannose, galactose and glucose (Gigi et al., 1981; Marbach et al., 1984; Zouari et al., 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 et al., 1997). The glycans are N-linked to the polypeptide chain (Ko et al., 2001; Brown et al., 2002; Saparrat et al., 2002). The most detailed structure of laccase glycan is available for R. lignosus laccase, which is also glycosylated with N-bound mannose (Garavaglia et al., 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 et al., 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 et al., 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 type2 (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 FEMS Microbiol Rev 30 (2006) 215–242

223


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 et al., 2001). The ‘white’ laccase POXA1 from P. ostreatus contains only one copper atom, together with two zinc and one iron atoms per molecule (Palmieri et al., 1997). Future structural studies will probably show that laccases are a more structurally heterogeneous group of proteins than expected.

Catalytic properties

Fig. 1. Catalytic cycle of laccase.

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 et al., 1997, 1997). Until recently, the three-dimensional structure of five fungal laccases has been reported: Coprinus cinereus (in a copper type-2-depleted form) (Ducros et al., 1998), T. versicolor (Bertrand et al., 2002; Piontek et al., 2002), P. cinnabarinus (Antorini et al., 2002), M. albomyces (Hakulinen et al., 2002) and R. lignosus (Garavaglia et al., 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 et al., 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 et al., 2004). A detailed structural comparison between a low redox potential (E0) C. cinereus laccase and a high E0 T. versicolor laccase showed that structural differences of the Cu1 coordination possibly account for the different E0 values (Piontek et al., 2002). This was later confirmed by the study of R. lignosus laccase with a high redox potential (Garavaglia et al., 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 FEMS Microbiol Rev 30 (2006) 215–242

Laccase catalyses the reduction of O2 to H2O using a range of phenolic compounds (though not tyrosine) as hydrogen donors (Thurston, 1994; Solomon et al., 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 et al., 1974; Thurston, 1994). Unambiguous determination of laccase activity is best achieved by purification of the protein to electrophoretic homogeneity followed by determination of KM or kcat 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 et al., 1999; Shin & Lee, 2000). In common with catechol oxidase and tyrosinase, laccase catalyzes the fourelectron reduction of O2 to H2O. 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 et al., 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 et al., 1996; Xu, 1996; Xu et al., 2000) and the presence of four cupric ions, each co-ordinated to a single polypeptide chain, is an absolute requirement for optimal activity (Ducros et al., 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. 2005 Federation of European Microbiological Societies Published by Blackwell Publishing Ltd. All rights reserved

c

224

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 et al., 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. KM ranges from 10 s of mM for syringaldazine and ABTS to 100 s of mM for DMP and guaiacol. The catalytic performance expressed as kcat 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 KM constants higher. Low KM 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 KM value is also high for pentachlorophenol with T. versicolor laccase (Table 3). Some fungi produce isoenzymes with similar KM and kcat 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 KM and kcat 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 & 2005 Federation of European Microbiological Societies Published by Blackwell Publishing Ltd. All rights reserved

c

P. Baldrian

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; Scha´ neˇl & 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 et al., 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. In addition to the oxidation of phenols, laccases have also been recently demonstrated to catalyze the oxidation of Mn21 in the presence of chelators. Laccase from the whiterot fungus T. versicolor oxidized Mn21 to Mn31 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 Mn21/Mn31 couple. Mn21 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 et al., 2001). It was proposed that laccase and Mn-peroxidase can co-operate. In the presence of Mn21 and oxalate, laccase produces Mn31-oxalate. The latter initializes a set of followup reactions leading to H2O2 formation, which may initiate or support peroxidase reactions (Schlosser & Höfer, 2002). The production of H2O2 and Mn31 was also described in P. eryngii for the oxidation of hydroquinone (Munoz et al., 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 et al., 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 DE/ FEMS Microbiol Rev 30 (2006) 215–242

225


Table 3. Substrates and inhibitors of fungal laccases. The numbers in brackets indicate Michaelis constant (KM, mM) or rate constant (kcat, 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 (3,4-Dimethoxyphenyl)methanol (veratryl alcohol) (4-Hydroxy-3-methoxyphenyl)acetic acid 1,2,4,5-Tetramethoxybenzene 1,2,4-Benzenetriol 1,2-Benzenediol (catechol)

1,3-Dihydroxybenzene (resorcinol) 1,4-Benzohydroquinone

1-Naphthol 2-(3,4-Dihydroxyphenyl)-3,5,7-trihydroxy-4Hchromen-4-one 2-Chlorophenol 2,2 0 -Azinobis(3-ethylbenzothiazoline-6-sulfonic acid) 2,3-Dichlorophenol 2,3-Dimethoxyphenol 2,3,6-Trichlorophenol 2,4,6-Trichlorophenol 2,4,6-Trimethylphenol 2,4-Dichlorophenol 2,5-Dihydroxybenzoic acid 2,6-Dichlorophenol 2,6-Dimethoxy-1,4-benzohydroquinone 2,6-Dimethoxyphenol

2,7-Diaminofluorene 2-Amino-3-(3,4-dihydroxyphenyl)propanoic acid 2-Amino-3-hydroxybenzoic acid 2-Amino-4-methylphenol 2-Amino-4-nitrophenol 2-Aminophenol 2-Aminophenylamine 2-Chlorobenzene-1,4-diol 2-Chlorophenol 2-Methoxy-1,4-benzohydroquinone 2-Methoxy-4-[prop-1-enyl]phenol 2-Methoxy-4-methylphenol 2-Methoxyaniline 2-Methoxyphenol (guaiacol) 2-Methoxy-1,4-benzohydroquinone 2-Methyl-1,4-benzohydroquinone 2-Methylanthracene 2-Methylphenol 2-Naphthol 2,4-Dichlorophenol 2,4,6-Trichlorophenol 3-(3,4-Dihydroxyphenyl)acrylic acid (caffeic acid)


Ts, Tv Pe Cs (KM: 6900; kcat: 1680), Cs (KM: 900; kcat: 3360) Bc Ab, Am, Bc, Cf (KM: 85; kcat: 90), Ch, Cm (KM: 120; kcat: 320), Cn, Cr, Ds, Gg (KM: 250), Gl (KM: 55), Le (KM: 220), Le (KM: 22 400), Lp, Mi, Mq, Pc, Pe (KM: 2200), Pe (KM: 4100), Pr, Rl, Sr, Th (KM: 142; kcat: 390), To (KM: 110; kcat: 80), Tp (KM: 470; kcat: 27 600), Ts, Tt Cn, Ts Am, Bc, Cf (KM: 68; kcat: 110), Ch, Cn, Cm (KM: 100; kcat: 290), Cr, Ct (KM: 36), Ds, Gl (KM: 29), Lp, Le (KM: 110), Pc, Pe (KM: 2500), Pe (KM: 4600), Pi, Pn, Rl, Th (KM: 61; kcat: 450), To (KM: 74; kcat: 110), Tp (KM: 390; kcat: 19 200), Ts, Tt Ab, Bc, Gl Am Tv Al (kcat 21), Cr (kcat: 4680), Cr (kcat: 4620), Cs (kcat: 5760), Cs (kcat: 6060), Po (kcat: 16 000), Po (kcat: 350 000), Po (kcat: 90 000), Rl (kcat: 34 700), Rl (kcat: 32 100), Tp (kcat: 41 400), Tr (kcat: 41 520), Tt (kcat: 198) Tv Ds Tv Tv Pe Tt, Tv Pi Tt, Tv Cr (KM: 107; kcat: 8580), Cr (KM: 89; kcat: 11 220) Al (kcat: 15), Cr (kcat: 6360), Cr (kcat: 5640), Cs (kcat: 1380), Cs (kcat: 4560), Po (kcat: 100), Po (kcat: 250), Po (kcat: 360 000), Rl (kcat: 2800), Rl (kcat: 2000), Tp (kcat: 24 000), Tr (kcat: 4860), Tt (kcat: 109) Rl Am, Cn, Ct (KM: 100), Le (KM: 650), Lp, Ma, Pa (KM: 3300), Ts, Tv (KM: 15 600) Pc Tt Tt Tt Cn Tt Le (KM: 1350), Mq, Tt Cr (KM: 216; kcat: 7620), Cr (KM: 229; kcat: 6300), Pe Ts Tt Tt Al (kcat: 159), Cf (kcat: 95), Cm (kcat: 160), Cs (kcat: 3120), Cs (kcat: 3960), Po (kcat: 150), Th (kcat: 430), To (kcat: 90), Tp (kcat: 10 800), Tr (kcat: 4140), Tt (kcat: 115) Pe Pe (KM: 1600), Pe (KM: 2100) Cg (kcat: 0.082) Bc, Tt Bc, Gl Mq Am, Bc, Cs, Le (KM: 40), Mi, Mq, Ts, Tt


c

226

P. Baldrian

Table 3. Continued. Species 3-(4-Hydroxy-3,5-dimethoxyphenyl)acrylic acid

Cf (KM: 21, kcat: 140), Cm (KM: 24; kcat: 330), Cs, Ff, Le (KM: 110), Pn, Pr, Rl, Th (KM: 24; kcat: 580), To (KM: 11; kcat: 170), Tv Am, Cf (KM: 20), Ch, Cs, Ct (KM: 270), Ff, Le (KM: 240), Le (KM: 2860), Mi, Mq, Pc, Pn, Pr, Rl, Sr, Tt (KM: 40; kcat: 145), Tv Le (KM: 240), Pn, Rl, Tt Mi (KM: 25), Pr, Tc Ab, Am, Bc, Ct (KM: 130), Le (KM: 130), Mq, Nc, On Cr, Mi, Mq, Pe, Tt Pe Bc Am, Ct

3-(4-Hydroxy-3-methoxyphenyl)acrylic acid (ferulic acid) 3-(4-Hydroxyphenyl)acrylic acid 3,3 0 -Dimethoxy-1,1 0 -biphenyl-4,4 0 -diamine 3,4,5-Trihydroxybenzoic acid (gallic acid) 3,4-Dihydroxybenzoic acid 3,5-Cyclohexadiene-1,2-diol 3,5-Dimethoxy-hydroxy-benzaldazine 3-{[3-(3,4-Dihydroxyphenyl)prop-2-enoyl]oxy}1,4,5-trihydroxycyclohexanecarboxylic acid 3-Amino-4-hydroxybenzenesulphonic acid Tt 3-Methoxyphenol Pr 4-(Hydroxymethyl)-2-methoxyphenol Cs (KM: 1600; kcat: 2820), Cs (KM: 610; kcat: 2220), Pe 4-[3-Hydroxyprop-1-enyl]-2,6-dimethoxyphenol Mq 4-[3-Hydroxyprop-1-enyl]-2-methoxyphenol Pe, Rl (coniferyl alcohol) 4-[3-Hydroxyprop-1-enyl]-phenol Mq 4-Amino-2,6-dichlorophenol Bc, Tt 4-Aminophenol Pe (KM: 1000), Pe (KM: 800), Tt 4-Aminophenylamine Am (KM: 1690), Bc, Cn, Gl, Lp, Le (KM: 256), Pe, Tc, Ts 4-Hydroxy-3,5-dimethoxybenzaldehyde Cr 4-Hydroxy-3,5-dimethoxybenzaldehyde [(4-hydroxy3,5-dimethoxyphenyl)methylene] hydrazone (syringaldazine) Al (kcat: 5), Po (kcat: 23 000), Po (kcat: 28 000), Po (kcat: 20 000), Tp (kcat: 16 800) 4-Hydroxy-3,5-dimethoxybenzoic acid (syringic acid) Cr, Cs (KM: 100; kcat: 4680), Cs (KM: 130; kcat: 1860), Ds, Ff (KM: 30), Mi, Pe, Pr, Tv (KM: 60) 4-Hydroxy-3-methoxybenzaldehyde Cr, Cs (KM: 6300; kcat: 1560), Cs (KM: 9000; kcat: 600), Pe 4-Hydroxy-3-methoxybenzoic acid (vanillic acid) Cf (KM: 160), Cs (KM: 1000; kcat: 3960), Cs (KM: 1100; kcat: 2220), Ct (KM: 150), Ff (KM: 970), Mi, Pe, Pr, Tt, Tv (KM: 130) 4-Hydroxybenzoic acid Mi 4-Hydroxyindole Ch, Pc 4-Chlorophenol Le (KM: 1740), Tt 4-Methoxyaniline Cr, Mi, Pe (KM: 3100), Pe (KM: 3300), Tp (KM: 1600; kcat: 7800), Tt 4-Methoxyphenol Cr, Le (KM: 330), Pe (KM: 800), Pe (KM: 900), Pr, Tt 4-Methylbenzene-1,2-diol Am, Bc, Ch, Cy, Le (KM: 170), Mq, Pc, Rl 4-Methylphenol Bc, Le (KM: 2200), Tt 4-Nitrobenzene-1,2-diol Tt 5-(1,2-Dihydroxyethyl)-3,4-dihydroxyfuran-2-one Ab, Am, Bc, Lp, Mi, Nc, Pa (KM: 190) (ascorbic acid) 9-Methylanthracene Cg (kcat: 4) Acenaphthene Cg (kcat: 0.167) Anthracene Cg (kcat: 0.087), Po, Tv Benzcatechin Pa (KM: 2270) Benzene-1,2,3-triol (pyrogallol) Ab, Bc, Ch, Cy, Gg (KM: 310), Gl, Le (KM: 30), Le (KM: 417), Lp, Nc, Pc, Rl, Sr, Ts Benzene-1,3,5-triol (phloroglucinol) Mi Benzo[a]pyrene Cg (kcat: 1.38) Biphenylene Cg (kcat: 0.063) Fluoranthene Po K4[Fe(CN)6] Am (KM: 1720), Cf (KM: 170; kcat: 130), Cm (KM: 115; kcat: 450), Lp, Pa (KM: 1030), Pi, Th (KM: 180; kcat: 400), To (KM: 96; kcat: 150), Tp (KM: 43; kcat: 51 000) Mn21 St, Tv (KM: 186) N,N 0 -Dimethylbenzene-1,4-diamine Ab, Am, Bc, Cf, Pc o-Tolidine Gl (KM: 402) o-Vanillin Cf (KM: 3900) Pentachlorophenol Tv (KM: 3000; kcat: 0.023) Phenanthrene Cg (kcat: 0.013) Phenylhydrazine Rl


c


227

Fungal laccases – occurrence and properties Table 3. Continued Species Inhibitor Ca21 Cd21 Co21 Fe21 Hg21 Mn21 Rb1 Sn21 Zn21 1-Phenyl-2-thiourea 2-Mercaptobenzothiazole 2-Mercaptoethanol 3-(4-Hydroxyphenyl)acrylic acid 4-Nitrophenol 8-Hydroxyquinoline Ascorbic acid Cetylpyridinium bromide Cetyltriammonium bromide CNCysteine Diethyldithiocarbamic acid Dithiothreitol EDTA Glutathione Humic acid Hydroxylamine KCl Kojic acid NaCl NaF NaN3 Salicylaldoxime SDS Thiamine Thiogylcolic acid Thiourea Trifluoroacetic acid Tropolone

Le Dq, Le Dq Ct, Po, Lp, Tc Ct, Dq, Le, Pu Dq, Pu Le Le Le, Po Ct Ct Ct, Pu, Te Le Pz Lp, Te Ct, Tc Ab Ab, Tc Ab, Bc, Ct, Gl, Lp, Ma, Me, Mi, Pn, Po, Pz, Rl, Tg, Tr, Ts Ct, Ch, Dq, Gl, Le, Mq, Pc, Py, Sr, Te, Vv Bc, Ch, Gl, Lp, Mi, Pp, Pc, Ps, Sr Ch, Dq, Le, Pc, Py, Vv Ct, Ma, Mq, Vv Dq, Gl Pt Po Le Dq, Le, Po Sr Ds, Me, Sr, Tt 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 Gl Ds, Mq, Pu, Tr Sr Ct, Mi, Po, Pr, Sr, Vv Ct, Dq Tr Ch, Le, Pc

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



c

228

P. Baldrian

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 Species Substrate

Am

Ct

Ch

Cy

Ds

Ma

Me

Mv

Pr

2,6-Dimethoxyphenol 100.0 100.0 100.0 100.0 100.0 100.0 100.0 100.0 100.0 2-Amino-3-(3,4-dihydroxy55.9 phenyl)propanoic acid 4-Aminophenol 4-Aminophenylamine 62.9 14.7 4-Hydroxyindole 87.6 4-Methoxyphenol 7.3 4-Methylcatechol 69.3 31.4 ABTS 76.5 271.7 114.4 800.0 288.3 1.4 97.4 Caffeic acid 18.6 Catechol 74.2 44.9 59.2 34.9 33.1 Ferulic acid 111.8 48.4 8.5 Guaiacol 59.7 73.5 107.8 37.9 92.0 122.4 31.0 39.1 9.7 Hydroquinone 50.0 105.3 80.8 12.9 N,N-dimethyl-1,4-phenylenediamine 74.8 1.8 23.4 o-Anisidine 45.5 p-Anisidine Pyrogallol 24.0 11.8 12.3 Syringaldazine 51.6 120.6 79.2 131.7 126.5 Syringic acid 115.2 46.9 Vanillic acid 61.8 7.3

Pe

Pc

Sr

Ts

Tt

Tv

100.0 100.0 100.0 100.0 100.0 100.0 46.0 109.8 194.8

26.7 56.0 107.6

9.8

19.8

103.5 284.1 452.0 136.7 21.6

74.3 76.0 140.9 25.5 62.0 19.9 19.3 34.5

18.3 35.0

27.7 80.2 24.5 27.9 110.7 116.3 64.0 55.8 38.4 69.0 30.2 56.0 237.1 9.3 3.5 76.0 6.3 95.0 76.0

14.1 8.1

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

DpH = 59 mV at 25 1C, 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 et al., 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 et al., 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 501 and 70 1C (Table 2). Few enzymes with optima below 35 1C have been described, e.g. the laccase from G. lucidum with the highest activity at 25 1C (Ko et al., 2001). This has, however, no connection with the growth 2005 Federation of European Microbiological Societies Published by Blackwell Publishing Ltd. All rights reserved

c

optimum of the fungi. The temperature stability varies considerably. The half life at 50 1C ranges from minutes in B. cinnerea (Slomczynski et al., 1995), to over 2–3 h in L. edodes and A. bisporus (Wood, 1980; D’Annibale et al., 1996), to up to 50–70 h in Trametes sp. (Smirnov et al., 2001). While the enzyme from G. lucidum was immediately inactivated at 60 1C, 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 et al., 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 et al., 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 FEMS Microbiol Rev 30 (2006) 215–242

229


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 et al., 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 et al., 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 et al., 2005). (Eggert et al., 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 et al., 2001) as well as for the transformation of the fungicide cyprodinil (Kang et al., 2002) and hydrocarbon degradation (Johannes & Majcherczyk, 2000). A comprehensive screening for natural mediators was performed by (Camarero et al., 2005). Among 44 tested natural lignin-derived compounds they selected 10 phenolic compounds derived from syringyl, guaiacyl, and phydroxyphenyl 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 FEMS Microbiol Rev 30 (2006) 215–242

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 et al., 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 et al., 1997, 1998; Baldrian et al., 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 et al., 1991), extraction with surfactants or calcium chloride (Criquet et al., 1999) or the most widely used extraction with phosphate buffer (Lang et al., 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 et al., 1995; Criquet et al., 2000; Carreiro et al., 2000) and coniferous forests (Ghosh et al., 2003), where laccase is the dominant ligninolytic enzyme (Criquet et al., 2000; Ghosh et al., 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 et al., 2000) and the pattern of detected isoenzymes varied during the succession (Di Nardo et al., 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 et al., 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 et al., 2005). Along with the 2005 Federation of European Microbiological Societies Published by Blackwell Publishing Ltd. All rights reserved

c

230

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 et al., 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 et al., 2000; Gallo et al., 2004), with the simultaneous increase in the litter layer (Saiya-Cork et al., 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 et al., 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 et al., 2003). This is supported by the fact that humic acids induce laccase activity and mRNA expression (Scheel et al., 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 et al., 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 et al., 2000) and inhibits its activity (Claus & Filip, 1990). (Zavarzina et al., 2004) estimated inhibition constants for humic acids towards Panus tigrinus laccase. The Ki ranged from 0.003 mg mL 1 for humic acids from peat soils to 0.025 mg 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 et al., 1999). 2005 Federation of European Microbiological Societies Published by Blackwell Publishing Ltd. All rights reserved

c

P. Baldrian

Laccases are also actively produced during the composting process. Of 34 isolates of fungi from woody compost, 11 were able to oxidize syringaldazine (Chamuris et al., 2000). Laccase was isolated both from compost-specific fungi and the compost itself (Chefetz et al., 1998, 1998) and it seems that it participates in both degradation of lignin and humic acids and humic acid formation (Chefetz et al., 1998; Kluczek-Turpeinen et al., 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 et al., 1994; Williams et al., 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 et al., 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 et al., 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 et al., 1998; Savoie, 2001; Velazquez-Cedeno et al., 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 et al., 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 FEMS Microbiol Rev 30 (2006) 215–242

231


xenobiotic compounds including chlorinated phenolics (Royarcand & Archibald, 1991; Roper et al., 1995; Ullah et al., 2000; Schultz et al., 2001; Bollag et al., 2003), synthetic dyes (Chivukula & Renganathan, 1995; Rodriguez et al., 1999; Wong & Yu, 1999; Abadulla et al., 2000; Nagai et al., 2002; Claus et al., 2002; Soares et al., 2002; Peralta-Zamora et al., 2003; Wesenberg et al., 2003; Zille et al., 2003), pesticides (Nannipieri & Bollag, 1991; Jolivalt et al., 2000; Torres et al., 2003) and polycyclic aromatic hydrocarbons (Johannes et al., 1996; Collins et al., 1996; Majcherczyk et al., 1998; Majcherczyk & Johannes, 2000; Cho et al., 2002; Pozdnyakova et al., 2004). They can bleach Kraft pulp (Reid & Paice, 1994; Paice et al., 1995; Bourbonnais & Paice, 1996; Call & Mucke, 1997; Monteiro & de Carvalho, 1998; de Carvalho et al., 1999; Sealey et al., 1999; Balakshin et al., 2001; Lund et al., 2003; Sigoillot et al., 2004) or detoxify agricultural byproducts including olive mill wastes or coffee pulp (D’Annibale et al., 2000; Tsioulpas et al., 2002; Velazquez-Cedeno et al., 2002) (for review see Durán & Esposito, 2000; Durán et al., 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 et al., 2003; Law et al., 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 (4 1 V) only allows the direct degradation of low-redoxpotential compounds and not the oxidation of more recalcitrant aromatic compounds, including some synthetic dyes or polycyclic aromatic hydrocarbons (PAH) (Xu et al., 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 et al., 2004), whereas ‘blue’ laccases from other fungi were not capable of efficient oxidation (Johannes et al., 1996; Majcherczyk et al., 1998; Kottermann et al., 1998). The compounds with higher redox potential can only be transformed if the reaction product is subject to FEMS Microbiol Rev 30 (2006) 215–242

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 et al., 1998; Xu et al., 2000; Fabbrini et al., 2002; Baiocco et al., 2003), the search for new mediators (Bourbonnais et al., 1997; Fabbrini et al., 2002), and their applications in the degradation of aromatic xenobiotics (Bourbonnais et al., 1997; Johannes et al., 1998; Kang et al., 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 et al., 2005) or 20–40 (Eggert et al., 1996; Campos et al., 2001) increase the feasibility of the laccasemediator 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, 2,4,6trinitrotoluene or chlorinated phenols (Tatsumi et al., 1994; Dawel et al., 1997; Dec & Bollag, 2000; Ahn et al., 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 H2O2, 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 2005 Federation of European Microbiological Societies Published by Blackwell Publishing Ltd. All rights reserved

c

232

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.

References Abadulla E, Tzanov T, Costa S, Robra KH, Cavaco-Paulo A & Gubitz GM (2000) Decolorization and detoxification of textile dyes with a laccase from Trametes hirsuta. Appl Environ Microbiol 66: 3357–3362. Abdel-Raheem A & Shearer CA (2002) Extracellular enzyme production by freshwater ascomycetes. Fungal Diversity 11: 1–19. Ahn M Y, Dec J, Kim JE & Bollag JM (2002) Treatment of 2,4dichlorophenol polluted soil with free and immobilized laccase. J Environ Qual 31: 1509–1515. Alexandre G & Zhulin IB (2000) Laccases are widespread in bacteria. Trends Biotechnol 18: 41–42. Allen AC & Walker JRL (1988) The selective inhibition of catechol oxidase by salicylhydroxamic acid. Phytochemistry 27: 3075–3076. Anderson DW & Nicholson RL (1996) Characterization of a laccase in the conidial mucilage of Colletotrichum graminicola. Mycologia 88: 996–1002. Antorini M, Herpoel-Gimbert I, Choinowski T, Sigoillot JC, Asther M, Winterhalter K & Piontek K (2002) Purification, crystallisation and X-ray diffraction study of fully functional laccases from two ligninolytic fungi. Biochim Biophys Acta 1594: 109–114. Artigas J, Romani AM & Sabater S (2004) Organic matter decomposition by fungi in a Mediterranean forested stream: contribution of streambed substrata. Ann Limnol 40: 269–277. Assavanig A, Amornkitticharoen B, Ekpaisal N, Meevootisom V & Flegel TW (1992) Isolation, characterization and function of


c

P. Baldrian

laccase from Trichoderma. Appl Microbiol Biotechnol 38: 198–202. Baiocco P, Barreca AM, Fabbrini M, Galli C & Gentili P (2003) Promoting laccase activity towards non-phenolic substrates: a mechanistic investigation with some laccase-mediator systems. Org Biomol Chem 1: 191–197. Balakshin M, Capanema E, Chen CL, Gratzl J, Kirkman A & Gracz H (2001) Biobleaching of pulp with dioxygen in the laccase-mediator system–reaction mechanisms for degradation of residual lignin. J Mol Catal B-Enzym 13: 1–16. Baldrian P (2003) Interactions of heavy metals with white-rot fungi. Enzyme Microb Technol 32: 78–91. Baldrian P (2004) Purification and characterization of laccase from the white-rot fungus Daedalea quercina and decolorization of synthetic dyes by the enzyme. Appl Microbiol Biotechnol 63: 560–563. Baldrian P (2004) Increase of laccase activity during interspecific interactions of white-rot fungi. FEMS Microbiol Ecol 50: 245–253. Baldrian P, in der Wiesche C, Gabriel J, Nerud F & Zadrazil F (2000) Influence of cadmium and mercury on activities of ligninolytic enzymes and degradation of polycyclic aromatic hydrocarbons by Pleurotus ostreatus in soil. Appl Environ Microbiol 66: 2471–2478. Banerjee UC & Vohra RM (1991) Production of laccase by Curvularia sp. Folia Microbiol 36: 343–346. Barriere F, Ferry Y, Rochefort D & Leech D (2004) Targetting redox polymers as mediators for laccase oxygen reduction in a membrane-less biofuel cell. Electrochem Commun 6: 237–241. Bekker EG, Petrova SD, Ermolova OV, Eliasashvili VI & Sinitsyn AP (1990) Isolation, purification, and certain properties of laccase from Cerrena unicolor. Biochemistry (Mosc) 55: 1506–1510. Bertrand G (1896) Sur la presence simultanee de la laccase et de la tyrosinase dans le suc de quelques champignons. C. R. Hebd. Seances Acad Sci 123: 463–465. Bertrand T, Jolivalt C, Briozzo P, Caminade E, Joly N, Madzak C & Mougin C (2002) Crystal structure of a four-copper laccase complexed with an arylamine: Insights into substrate recognition and correlation with kinetics. Biochemistry 41: 7325–7333. Billal F & Thurston CF (1996) Purification of laccase II from Armillaria mellea and comparison of its properties with those of laccase I. Mycol Res 100: 1099–1105. Binz T & Canevascini G (1997) Purification and partial characterization of the extracellular laccase from Ophiostoma novo-ulmi. Curr Microbiol 35: 278–281. Blaich R & Esser K (1975) Function of enzymes in wood destroying fungi. 2. Multiple forms of laccase in white rot fungi. Arch Microbiol 103: 271–277. Boerner REJ & Brinkman JA (2003) Fire frequency and soil enzyme activity in southern Ohio oak-hickory forests. Appl Soil Ecol 23: 137–146.


233


Bollag JM (1991) Enzymatic binding of pesticide degradation products to soil organic-matter and their possible release. ACS Symp Ser 459: 122–132. Bollag JM, Chu HL, Rao MA & Gianfreda L (2003) Enzymatic oxidative transformation of chlorophenol mixtures. J Environ Qual 32: 63–69. Bollag JM & Myers C (1992) Detoxification of aquatic and terrestrial sites through binding of pollutants to humic substances. Sci Total Environ 118: 357–366. Bonomo RP, Boudet AM, Cozzolino R, Santoro E, Rizzarelli AM, Sterjiades R & Zapalla R (1998) A comparative study of two isoforms of laccase secreted by the ‘‘white-rot’’ fungus Rigidoporus lignosus, exhibiting significant structural and functional differences. J Inorg Biochem 71: 205–211. Boudet AM (2000) Lignins and lignification: selected issues. Plant Physiol Biochem 38: 81–96. Bourbonnais R, Leech D & Paice MG (1998) Electrochemical analysis of the interactions of laccase mediators with lignin model compounds. Biochim Biophys Acta 1379: 381–390. Bourbonnais R & Paice MG (1990) Oxidation of non-phenolic substrates. An expanded role for laccase in lignin biodegradation. FEBS Lett 267: 99–102. Bourbonnais R & Paice MG (1996) Enzymatic delignification of kraft pulp using laccase and a mediator. Tappi J 79: 199–204. Bourbonnais R, Paice MG, Freiermuth B, Bodie E & Borneman S (1997) Reactivities of various mediators and laccases with kraft pulp and lignin model compounds. Appl Environ Microbiol 63: 4627–4632. Brown MA, Zhao ZW & Mauk AG (2002) Expression and characterization of a recombinant multi-copper oxidase: laccase IV from Trametes versicolor. Inorg Chim Acta 331: 232–238. Burke RM & Cairney JWG (2002) Laccases and other polyphenol oxidases in ecto- and ericoid mycorrhizal fungi. Mycorrhiza 12: 105–116. Cairney JWG & Burke RM (1998) Do ecto- and ericoid mycorrhizal fungi produce peroxidase activity? Mycorrhiza 8: 61–65. Call HP & Mucke I (1997) History, overview and applications of mediated lignolytic systems, especially laccase-mediatorsystems (Lignozyms-process). J Biotechnol 53: 163–202. Call HP & Mucke I (1997) History, overview and applications of mediated lignolytic systems, especially laccase-mediatorsystems (Lignozyms-process). J Biotechnol 53: 163–202. Calvo A, Copapatino MJ, Alonso LO & Gonzalez AE (1998) Studies of the production and characterization of laccase activity in the basidiomycete Coriolopsis gallica, an efficient decolorizer of alkaline effluents. Arch Microbiol 171: 31–36. Camarero S, Ibarra D, Mart´ınez MJ & Mart´ınez AT (2005) Lignin-derived compounds as efficient laccase mediators for decolorization of different types of recalcitrant dyes. Appl Environ Microbiol 71: 1775–1784. Cambria MT, Cambria A, Ragusa S & Rizzarelli E (2000) Production, purification, and properties of an extracellular


laccase from Rigidoporus lignosus. Protein Expr Purif 18: 141–147. Campos R, Kandelbauer A, Robra KH, Cavaco-Paulo A & Gubitz GM (2001) Indigo degradation with purified laccases from Trametes hirsuta and Sclerotium rolfsii. J Biotechnol 89: 131–139. Carreiro MM, Sinsabaugh RL, Repert DA & Parkhurst DF (2000) Microbial enzyme shifts explain litter decay responses to simulated nitrogen deposition. Ecology 81: 2359–2365. Chambers SM, Burke RM, Brooks PR & Cairney JWG (1999) Molecular and biochemical evidence for manganesedependent peroxidase activity in Tylospora fibrillosa. Mycol Res 103: 1098–1102. Chamuris GP, Koziol-Kotch S & Brouse TM (2000) Screening fungi isolated from woody compost for lignin-degrading potential. Compost Sci Util 8: 6–11. Chefetz B, Chen Y & Hadar Y (1998) Purification and characterization of laccase from Chaetomium thermophilium and its role in humification. Appl Environ Microbiol 64: 3175–3179. Chefetz B, Kerem Z, Chen Y & Hadar Y (1998) Isolation and partial characterization of laccase from a thermophilic composted municipal solid waste. Soil Biol Biochem 30: 1091–1098. Chen DM, Bastias BA, Taylor AFS & Cairney JWG (2003) Identification of laccase-like genes in ectomycorrhizal basidiomycetes and transcriptional regulation by nitrogen in Piloderma byssinum. New Phytol 157: 547–554. Chen S, Ge W & Buswell JA (2004) Biochemical and molecular characterization of a laccase from the edible straw mushroom, Volvariella volvacea. Eur J Biochem 271: 318–328. Chen DM, Taylor AFS, Burke RM & Cairney JWG (2001) Identification of genes for lignin peroxidases and manganese peroxidases in ectomycorrhizal fungi. New Phytol 152: 151–158. Chivukula M & Renganathan V (1995) Phenolic azo dye oxidation by laccase from Pyricularia oryzae. Appl Environ Microbiol 61: 4374–4377. Cho SJ, Park SJ, Lim JS, Rhee YH & Shin KS (2002) Oxidation of polycyclic aromatic hydrocarbons by laccase of Coriolus hirsutus. Biotechnol Lett 24: 1337–1340. Claus H (2003) Laccases and their occurrence in prokaryotes. Arch Microbiol 179: 145–150. Claus H, Faber G & Konig H (2002) Redox-mediated decolorization of synthetic dyes by fungal laccases. Appl Microbiol Biotechnol 59: 672–678. Claus H & Filip Z (1990) Effects of clays and other solids on the activity of phenoloxidases produced by some fungi and actinomycetes. Soil Biol Biochem 22: 483–488. Claus H & Filip Z (1998) Degradation and transformation of aquatic humic substances by laccase-producing fungi Cladosporium cladosporioides and Polyporus versicolor. Acta Hydrochim Hydrobiol 26: 180–185.


c

234

Collins PJ, Kotterman MJJ, Field JA & Dobson ADW (1996) Oxidation of anthracene and benzo[a]pyrene by laccases from Trametes versiocolor. Appl Environ Microbiol 62: 4563–4567. Criquet S, Farnet AM, Tagger S & Le Petit J (2000) Annual variations of phenoloxidase activities in an evergreen oak litter: influence of certain biotic and abiotic factors. Soil Biol Biochem 32: 1505–1513. Criquet S, Tagger S, Vogt G, Iacazio G & Le Petit J (1999) Laccase activity of forest litter. Soil Biol Biochem 31: 1239–1244. Crowe JD & Olsson S (2001) Induction of laccase activity in Rhizoctonia solani by antagonistic Pseudomonas fluorescens strains and a range of chemical treatments. Appl Environ Microbiol 67: 2088–2094. Curir P, Thurston CF, Daquila F, Pasini C & Marchesini A (1997) Characterization of a laccase secreted by Armillaria mellea pathogenic for Genista. Plant Physiol Biochem 35: 147–153. Dahiya JS, Singh D & Nigam P (1998) Characterisation of laccase produced by Coniothyrium minitans. J Basic Microbiol 38: 349–359. Das N, Chakraborty TK & Mukherjee M (2000) Purification and characterization of laccase-1 from Pleurotus florida. Folia Microbiol 45: 447–451. Dawel G, Kastner M, Michels J, Poppitz W, Günther W & Fritsche W (1997) Structure of a laccase-mediated product of coupling of 2,4-diamino-6-nitrotoluene to guaiacol, a model for coupling of 2,4,6-trinitrotoluene metabolites to a humic organic soil matrix. Appl Environ Microbiol 63: 2560–2565. Dawley RM & Flurkey WH (1993) Differentiation of tyrosinase and laccase using 4-hexyl-resorcinol, a tyrosinase inhibitor. Phytochem 33: 281–284. Carvalho MEA, Monteiro MC & Sant’Anna GL (1999) Laccase from Trametes versicolor - stability at temperature and alkaline conditions and its effect on biobleaching of hardwood kraft pulp. Appl Biochem Biotechnol 77: 723–733. Souza CGM & Peralta RM (2003) Purification and characterization of the main laccase produced by the white-rot fungus Pleurotus pulmonarius on wheat bran solid state medium. J Basic Microbiol 43: 278–286. Vries OMH, Kooistra WHCF & Wessels GH (1986) Formation of an extracellular laccase by Schizophyllum commune dikaryon. J Gen Microbiol 132: 2817–2826. DeForest JL, Zak DR, Pregitzer KS & Burton AJ (2004) Atmospheric nitrate deposition, microbial community composition, and enzyme activity in northern hardwood forests. Soil Sci Soc Amer J 68: 132–138. DeForest JL, Zak DR, Pregitzer KS & Burton AJ (2004) Atmospheric nitrate deposition and the microbial degradation of cellobiose and vanillin in a northern hardwood forest. Soil Biol Biochem 36: 965–971. Dec J & Bollag JM (2000) Phenoloxidase-mediated interactions of phenols and anilines with humic materials. J Environ Qual 29: 665–676. Dedeyan B, Klonowska A, Tagger S, Tron T, Iacazio G, Gil G & Le Petit J (2000) Biochemical and molecular characterization of a


c

P. Baldrian

laccase from Marasmius quercophilus. Appl Environ Microbiol 66: 925–929. Nardo C, Cinquegrana A, Papa S, Fuggi A & Fioretto A (2004) Laccase and peroxidase isoenzymes during leaf litter decomposition of Quercus ilex in a Mediterranean ecosystem. Soil Biol Biochem 36: 1539–1544. Dittmer JK, Patel NJ, Dhawale SW & Dhawale SS (1997) Production of multiple laccase isoforms by Phanerochaete chrysosporium grown under nutrient sufficiency. FEMS Microbiol Lett 149: 65–70. Dong JL & Zhang YZ (2004) Purification and characterization of two laccase isoenzymes from a ligninolytic fungus Trametes gallica. Prep Biochem Biotechnol 34: 179–194. Ducros V, Brzozowski AM, Wilson KS, Brown SH, Ostergaard P, Schneider P, Yaver DS, Pedersen AH & Davies GJ (1998) Crystal structure of the type-2 Cu depleted laccase from Coprinus cinereus at 2.2 angstrom resolution. Nat Struct Biol 5: 310–316. Durán N & Esposito E (2000) Potential applications of oxidative enzymes and phenoloxidase-like compounds in wastewater and soil treatment: a review. Appl Catal B: Environ 28: 83–99. Durán N, Rosa MA, Dannibale A & Gianfreda L (2002) Applications of laccases and tyrosinases (phenoloxidases) immobilized on different supports: a review. Enzyme Microb Technol 31: 907–931. Durrens P (1981) The phenoloxidases of the ascomycete Podospora anserina: the three forms of the major laccase activity. Arch Microbiol 130: 121–124. D’Annibale A, Celletti D, Felici M, Dimattia E & GiovannozziSermanni G (1996) Substrate specificity of laccase from Lentinus edodes. Acta Biotechnol 16: 257–270. D’Annibale A, Stazi SR, Vinciguerra V & Sermanni GG (2000) Oxirane-immobilized Lentinula edodes laccase: stability and phenolics removal efficiency in olive mill wastewater. J Biotechnol 77: 265–273. D’Souza TM, Boominathan K & Reddy CA (1996) Isolation of laccase gene-specific sequences from white rot and brown rot fungi by PCR. Appl Environ Microbiol 62: 3739–3744. Edens WA, Goins TQ, Dooley D & Henson JM (1999) Purification and characterization of a secreted laccase of Gaeumannomyces graminis var. tritici. Appl Environ Microbiol 65: 3071–3074. Eggen T (1999) Application of fungal substrate from commercial mushroom production – Pleuorotus ostreatus – for bioremediation of creosote contaminated soil. Int Biodeter Biodegrad 44: 117–126. Eggert C, Temp U, Dean JFD & Eriksson KEL (1995) Laccasemediated formation of the phenoxazinone derivative, cinnabarinic acid. FEBS Lett 376: 202–206. Eggert C, Temp U, Dean JFD & Eriksson KEL (1996) A fungal metabolite mediates degradation of non-phenolic lignin structures and synthetic lignin by laccase. FEBS Lett 391: 144–148. Eggert C, Temp U & Eriksson KEL (1996) The ligninolytic system of the white rot fungus Pycnoporus cinnabarinus: purification


235


and characterization of the laccase. Appl Environ Microbiol 62: 1151–1158. Eller LR, Henderson RR, Slomczynski DJ & Eriksson KEL (1998) Isolation and characterization of laccase isozymes from Ganoderma tsugae. Abstr Pap Am Chem Soc 215: 401. Enguita FJ, Marcal D, Martins LO, Grenha R, Henriques AO, Lindley PF & Carrondo MA (2004) Substrate and dioxygen binding to the endospore coat laccase from Bacillus subtilis. J Biol Chem 279: 23472–23476. Enguita FJ, Martins LO, Henriques AO & Carrondo MA (2003) Crystal structure of a bacterial endospore coat component – a laccase with enhanced thermostability properties. J Biol Chem 278: 19416–19425. Fabbrini M, Galli C & Gentili P (2002) Comparing the catalytic efficiency of some mediators of laccase. J Mol Catal B 16: 231–240. Fahraeus G (1961) Monophenolase and polyphenolase activity of fungal laccase. Biochim Biophys Acta 54: 192–194. Fahraeus G & Ljunggren H (1961) Substrate specificity of a purified fungal laccase. Biochim Biophys Acta 46: 22–32. Fakoussa RM & Frost PJ (1999) In vivo-decolorization of coalderived humic acids by laccase-excreting fungus Trametes versicolor. Appl Microbiol Biotechnol 52: 60–65. Farnet AM, Criquet S, Cigna M, Gil G & Ferre E (2004) Purification of a laccase from Marasmius quercophilus induced with ferulic acid: reactivity towards natural and xenobiotic aromatic compounds. Enzyme Microb Technol 34: 549–554. Farnet AM, Criquet S, Pocachard E, Gil G & Ferre E (2002) Purification of a new isoform of laccase from a Marasmius quercophilus strain isolated from a cork oak litter (Quercus suber L.). Mycologia 94: 735–740. Farnet AM, Criquet S, Tagger S, Gil G & Le Petit J (2000) Purification, partial characterization, and reactivity with aromatic compounds of two laccases from Marasmius quercophilus strain 17. Can J Microbiol 46: 189–194. Farnet AM, Tagger S & Le Petit J (1999) Effects of copper and aromatic inducers on the laccases of the white-rot fungus Marasmius quercophilus. C R Acad Sci Series III 322: 499–503. Fioretto A, Papa S, Curcio E, Sorrentino G & Fuggi A (2000) Enzyme dynamics on decomposing leaf litter of Cistus incanus and Myrtus communis in a Mediterranean ecosystem. Soil Biol Biochem 32: 1847–1855. Freeman C, Ostle NJ, Fenner N & Kang H (2004) A regulatory role for phenol oxidase during decomposition in peatlands. Soil Biol Biochem 36: 1663–1667. Freitag M & Morrell JJ (1992) Changes in selected enzyme activities during growth of pure and mixed cultures of the white-rot decay fungus Trametes versicolor and the potential biocontrol fungus Trichoderma harzianum. Can J Microbiol 38: 317–323. Froehner SC & Eriksson KEL (1974) Purification and properties of Neurospora crassa laccase. J Bacteriol 120: 458–465. Fukushima Y & Kirk TK (1995) Laccase component of the Ceriporiopsis subvemispora lignin degrading system. Appl Environ Microbiol 61: 872–876.


Galhaup C, Goller S, Peterbauer CK, Strauss J & Haltrich D (2002) Characterization of the major laccase isoenzyme from Trametes pubescens and regulation of its synthesis by metal ions. Microbiology 148: 2159–2169. Galhaup C & Haltrich D (2001) Enhanced formation of laccase activity by the white-rot fungus Trametes pubescens in the presence of copper. Appl Microbiol Biotechnol 56: 225–32. Gallo M, Amonette R, Lauber C, Sinsabaugh RL & Zak DR (2004) Microbial community structure and oxidative enzyme activity in nitrogen-amended north temperate forest soils. Microb Ecol 48: 218–229. Garavaglia S, Cambria MT, Miglio M, Ragusa S, Lacobazzi V, Palmieri F, D’Ambrosio C, Scaloni A & Rizzi M (2004) The structure of Rigidoporus lignosus laccase containing a full complement of copper ions, reveals an asymmetrical arrangement for the T3 copper pair. J Mol Biol 342: 1519–1531. Garzillo AMV, Colao MC, Caruso C, Caporale C, Celletti D & Buonocore V (1998) Laccase from the white-rot fungus Trametes trogii. Appl Microbiol Biotechnol 49: 545–551. Geiger JP, Rio B, Nandris D & Nicole M (1986) Laccases of Rigidoporus lignosus and Phellinus noxius. Appl Biochem Biotechnol 12: 121–133. Ghosh A, Frankland JC, Thurston CF & Robinson CH (2003) Enzyme production by Mycena galopus mycelium in artificial media and in Picea sitchensis F-1 horizon needle litter. Mycol Res 107: 996–1008. Ghosh D & Mukherjee R (1998) Modeling tyrosinase monooxygenase activity. Spectroscopic and magnetic investigations of products due to reactions between copper(I) complexes of xylyl-based dinucleating ligands and dioxygen: aromatic ring hydroxylation and irreversible oxidation products. Inorg Chem 37: 6597–6605. Giardina P, Palmieri G, Scaloni A, Fontanella B, Faraco V, Cennamo G & Sannia G (1999) Protein and gene structure of a blue laccase from Pleurotus ostreatus. Biochem J 341: 655–663. Gigi O, Marbach I & Mayer AM (1981) Properties of gallic acidinduced extracellular laccase of Botrytis cinerea. Phytochemistry 20: 1211–1213. Givaudan A, Effosse A, Faure D, Potier P, Bouillant ML & Bally R (2004) Polyphenol oxidase in Azospirillum lipoferum isolated from rice rhizosphere: evidence for a laccase in non-motile strains of Azospirillum lipoferum. FEMS Microbiol Lett 108: 205–210. Gramss G, Günther T & Fritsche W (1998) Spot tests for oxidative enzymes in ectomycorrhizal, wood-, and litter decaying fungi. Mycol Res 102: 67–72. Gramss G, Kirsche B, Voigt KD, Günther T & Fritsche W (1999) Conversion rates of five polycyclic aromatic hydrocarbons in liquid cultures of fifty-eight fungi and the concomitant production of oxidative enzymes. Mycol Res 103: 1009–1018. Gramss G, Voigt KD & Bergmann H (2005) Factors influencing water solubility and plant availability of mineral compounds in the tripartite fairy rings of Marasmius oreades (BOLT.: FR.) FR. J Basic Microbiol 45: 41–54.


c

236

Günther H, Perner B & Gramss G (1998) Activities of phenol oxidizing enzymes of ectomycorrhizal fungi in axenic culture and in symbiosis with Scots pine (Pinus sylvestris L.). J Basic Microbiol 38: 197–206. Hakala TK, Lundell T, Galkin S, Maijala P, Kalkkinen N & Hatakka A (2005) Manganese peroxidases, laccases and oxalic acid from the selective white-rot fungus Physisporinus rivulosus grown on spruce wood chips. Enzyme Microb Technol 36: 461–468. Hakulinen N, Kiiskinen LL, Kruus K, Saloheimo M, Paananen A, Koivula A & Rouvinen J (2002) Crystal structure of a laccase from Melanocarpus albomyces with an intact trinuclear copper site. Nature Struct Biol 9: 601–605. Harkin JM, Larsen MJ & Obst JR (1974) Use of syringaldazine for detection of laccase in sporophores of wood rotting fungi. Mycologia 66: 469–476. Hatakka A (2001) Biodegradation of lignin. Lignin, Humic Substances and Coal (Hofrichter M & Steinbüchel A, eds), pp. 129–179. Wiley-VCH, Weinheim, Germany. Heinzkill M, Bech L, Halkier T, Schneider P & Anke T (1998) Characterization of laccases and peroxidases from woodrotting fungi (family Coprinaceae). Appl Environ Microbiol 64: 1601–1606. Hendel B & Marxsen J (2000) Extracellular enzyme activity associated with degradation of beech wood in a Central European stream. Int Rev Hydrobiol 85: 95–105. Holker U, Dohse J & Hofer M (2002) Extracellular laccases in ascomycetes Trichoderma atroviride and Trichoderma harzianum. Folia Microbiol 47: 423–427. Hoopes JT & Dean JFD (2004) Ferroxidase activity in a laccaselike multicopper oxidase from Liriodendron tulipifera. Plant Physiol Biochem 42: 27–33. Höfer C & Schlosser D (1999) Novel enzymatic oxidation of Mn21 to Mn31 catalyzed by a fungal laccase. FEBS Lett 451: 186–190. Hublik G & Schinner F (2000) Characterization and immobilization of the laccase from Pleurotus ostreatus and its use for the continuous elimination of phenolic pollutants. Enzyme Microb Technol 27: 330–336. Iakovlev A & Stenlid J (2000) Spatiotemporal patterns of laccase activity in interacting mycelia of wood-decaying basidiomycete fungi. Microb Ecol 39: 236–245. Ikeda R, Sugita T, Jacobson ES & Shinoda T (2003) Effects of melanin upon susceptibility of Cryptococcus to antifungals. Microbiol Immunol 47: 271–277. Ishigami T, Hirose Y & Yamada Y (1998) Characterization of polyphenol oxidase from Chaetomium thermophile, a thermophilic fungus. J Gen Appl Microbiol 34: 401–407. Itoh K, Fujita M, Kumano K, Suyama K & Yamamoto H (2000) Phenolic acids affect transformations of chlorophenols by a Coriolus versicolor laccase. Soil Biol Biochem 32: 85–91. Iwasaki H, Matsubara T & Mori T (1967) A fungal laccase, its properties and reconstitution from its protein and copper. J Biochem 61: 814–816.


c

P. Baldrian

Iyer G & Chattoo BB (2003) Purification and characterization of laccase from the rice blast fungus, Magnaporthe grisea. FEMS Microbiol Lett 227: 121–126. Jimenez M, Gonzalez AE, Martinez MJ, Martinez AT & Dále BE (1991) Screening of yeasts isolated from decayed wood for lignocellulose degrading enzyme activities. Mycol Res 95: 1299–1302. Johannes C & Majcherczyk A (2000) Laccase activity tests and laccase inhibitors. J Biotechnol 78: 193–199. Johannes C & Majcherczyk A (2000) Natural mediators in the oxidation of polycyclic aromatic hydrocarbons by laccase mediator systems. Appl Environ Microbiol 66: 524–528. Johannes C, Majcherczyk A & Hüttermann A (1996) Degradation of anthracene by laccase of Trametes versicolor in the presence of different mediator compounds. Appl Microbiol Biotechnol 46: 313–317. Johannes C, Majcherczyk A & Hüttermann A (1998) Oxidation of acenaphthene and acenaphthylene by laccase of Trametes versicolor in a laccase-mediator system. J Biotechnol 61: 151–156. Jolivalt C, Brenon S, Caminade E, Mougin C & Pontie M (2000) Immobilization of laccase from Trametes versicolor on a modified PVDF microfiltration membrane: characterization of the grafted support and application in removing a phenylurea pesticide in wastewater. J Membr Sci 180: 103–113. Junghanns C, Moeder M, Krauss G, Martin C & Schlosser D (2005) Degradation of the xenoestrogen nonylphenol by aquatic fungi and their laccases. Microbiology 151: 45–57. Kang KH, Dec J, Park H & Bollag JM (2002) Transformation of the fungicide cyprodinil by a laccase of Trametes villosa in the presence of phenolic mediators and humic acid. Water Res 36: 4907–4915. Kanunfre CC & Zancan GT (1998) Physiology of exolaccase production by Thelephora terrestris. FEMS Microbiol Lett 161: 151–156. Katase T & Bollag JM (1991) Transformation of trans-4hydroxycinnamic acid by a laccase of the fungus Trametes versicolor – its significance in humification. Soil Sci 151: 291–296. Keum YS & Li QX (2004) Copper dissociation as a mechanism of fungal laccase denaturation by humic acid. Appl Microbiol Biotechnol 64: 588–592. Kiiskinen LL, Viikari L & Kruus K (2002) Purification and characterisation of a novel laccase from the ascomycete Melanocarpus albomyces. Appl Microbiol Biotechnol 59: 198–204. Kim Y, Cho NS, Eom TJ & Shin W (2002) Purification and characterization of a laccase from Cerrena unicolor and its reactivity in lignin degradation. Bull Korean Chem Soc 23: 985–989. Kirk TK & Farrell RL (1987) Enzymatic ‘‘combustion’’: the microbial degradation of lignin. Annu Rev Microbiol 41: 465–505. Kluczek-Turpeinen B, Steffen KT, Tuomela M, Hatakka A & Hofrichter M (2005) Modification of humic acids by the


237


compost-dwelling deuteromycete Paecilomyces inflatus. Appl Microbiol Biotechnol 66: 443–449. Kluczek-Turpeinen B, Tuomela M, Hatakka A & Hofrichter M (2003) Lignin degradation in a compost environment by the deuteromycete Paecilomyces inflatus. Appl Microbiol Biotechnol 61: 374–379. Ko EM, Leem YE & Choi HT (2001) Purification and characterization of laccase isozymes from the white-rot basidiomycete Ganoderma lucidum. Appl Microbiol Biotechnol 57: 98–102. Kofujita H, Ohta T, Asada Y & Kuwahara M (1991) Purification and characterization of laccase from Lentinus edodes. Mokuzai Gakkaishi 37: 562–569. Koroleva OV, Yavmetdinov IS, Shleev SV, Stepanova EV & Gavrilova VP (2001) Isolation and study of some properties of laccase from the Basidiomycetes Cerrena maxima. Biochemistry (Mosc) 66: 618–622. Koroljova OV, Stepanova EV, Gavrilova VP, Biniukov VI, Jaropolov AI, Varfolomeyev SD, Scheller F, Makower A & Otto A (1999) Laccase of Coriolus zonatus – Isolation, purification, and some physicochemical properties. Appl Biochem Biotechnol 76: 115–127. Koroljova-Skorobogat’ko OV, Stepanova EV, Gavrilova VP, Morozova OV, Lubimova NV, Dzchafarova AN, Jaropolov AI & Makower A (1998) Purification and characterization of the constitutive form of laccase from the basidiomycete Coriolus hirsutus and effect of inducers on laccase synthesis. Biotechnol Appl Biochem 28: 47–54. Kottermann MJJ, Rietberg HJ, Hage A & Field JA (1998) Polycyclic aromatic hydrocarbon oxidation by the white rot fungus Bjerkandera sp. strain BOS55 in the presence of nonionic surfactants. Biotechnol Bioeng 57: 220–227. Lalitha Kumari H & Sirsi M (1972) Purification and properties of laccase from Ganoderma lucidum. Arch Microbiol 84: 350–357. Lang E, Eller G & Zadrazil F (1997) Lignocellulose decomposition and production of ligninolytic enzymes during interaction of white rot fungi with soil microorganisms. Microb Ecol 34: 1–10. Lang E, Nerud F & Zadrazil F (1998) Production of ligninolytic enzymes by Pleurotus sp. and Dichomitus squalens in soil and lignocellulose substrate as influenced by soil microorganisms. FEMS Microbiol Lett 167: 239–244. Larrondo LF, Salas L, Melo F, Vicuna R & Cullen D (2003) A novel extracellular multicopper oxidase from Phanerochaete chrysosporium with ferroxidase activity. Appl Environ Microbiol 69: 6257–6263. Lau KL, Tsang YY & Chiu SW (2003) Use of spent mushroom compost to bioremediate PAH-contaminated samples. Chemosphere 52: 1539–1546. Law WM, Lau WN, Lo KL, Wai LM & Chiu SW (2003) Removal of biocide pentachlorophenol in water system by the spent mushroom compost of Pleurotus pulmonarius. Chemosphere 52: 1531–1537.


Lee YJ & Shin KS (1999) Purification and properties of laccase of the white-rot basidiomycete Coriolus hirsutus. J Microbiol 37: 148–153. Lee KH, Wi SG, Singh AP & Kim YS (2004) Micromorphological characteristics of decayed wood and laccase produced by the brown-rot fungus Coniophora puteana. J Wood Sci 50: 281–284. Leitner C, Hess J, Galhaup C, Ludwig R, Nidetzky B, Kulbe KD & Haltrich D (2002) Purification and characterization of a laccase from the white-rot fungus Trametes multicolor. Appl Biochem Biotechnol 98: 497–507. Leonowicz A, Cho NS, Luterek J, Wilkolazka A, Wojtaswasilewska M, Matuszewska A, Hofrichter M, Wesenberg D & Rogalski J (2001) Fungal laccase: properties and activity on lignin. J Basic Microbiol 41: 185–227. Leonowicz A, Edgehill RU & Bollag JM (1984) The effect of pH on the transformation of syringic and vanillic acids by the laccases of Rhizoctonia praticola and Trametes versicolor. Arch Microbiol 137: 89–96. Leonowicz A & Malinowska M (1982) Purification of the constitutive and inducible forms of laccase of the fungus Pholiota mutabilis by affinity chromatography. Acta Biochim Polon 29: 219–226. Leontievsky AA, Myasoedova NM, Baskunov BP, Golovleva LA, Bucke C & Evans CS (2001) Transformation of 2,4,6trichlorophenol by free and immobilized fungal laccase. Appl Microbiol Biotechnol 57: 85–91. Leontievsky A, Myasoedova N, Pozdnyakova N & Golovleva L (1997) ‘‘Yellow’’ laccase of Panus tigrinus oxidizes nonphenolic substrates without electron-transfer mediators. FEBS Lett 413: 446–448. Leontievsky AA, Vares T, Lankinen P, et al. (1997) Blue and yellow laccases of ligninolytic fungi. FEMS Microbiol Lett 156: 9–14. Liers C, Ullrich R, Steffen KT, Hatakka A & Hofrichter M (2005) Mineralization of 14C-labelled lignin and extracellular enzyme activities of the wood-colonizing ascomycetes Xylaria hypoxylon and Xylaria polymorpha. Appl Microbiol Biotechnol. DOI 10.1007/s00253-005-0010-1. (in press). Liu L, Dean JFD, Friedman WE & Eriksson KEL (1994) Laccaselike phenoloxidase is correlated with lignin biosynthesis in Zinnia elegans stem tissue. Plant J 6: 213–224. Lo SC, Ho YS & Buswell JA (2001) Effect of phenolic monomers on the production of laccases by the edible mushroom Pleurotus sajor-caju, and partial characterization of a major laccase component. Mycologia 93: 413–421. Luis P, Walther G, Kellner H, Martin F & Buscot F (2004) Diversity of laccase genes from basidiomycetes in a forest soil. Soil Biol Biochem 36: 1025–1036. Lyons JI, Newell SY, Buchan A & Moran MA (2003) Diversity of ascomycete laccase gene sequences in a southeastern US salt marsh. Microb Ecol 45: 270–281. Lund M, Eriksson M & Felby C (2003) Reactivity of a fungal laccase towards lignin in softwood kraft pulp. Holzforschung 57: 21–26.


c

238

Machonkin TE, Quintanar L, Palmer AE, Hassett R, Severance S, Kosman DJ & Solomon EI (2001) Spectroscopy and reactivity of the type 1 copper site in Fet3p from Saccharomyces cerevisiae: correlation of structure with reactivity in the multicopper oxidases. J Am Chem Soc 123: 5507–5517. Majcherczyk A & Johannes C (2000) Radical mediated indirect oxidation of a PEG-coupled polycyclic aromatic hydrocarbon (PAH) model compound by fungal laccase. Biochim Biophys Acta 1474: 157–162. Majcherczyk A, Johannes C & Hüttermann A (1998) Oxidation of polycyclic aromatic hydrocarbons (PAH) by laccase of Trametes versicolor. Enzyme Microb Technol 22: 335–341. Makkar RS, Tsuneda A, Tokuyasu K & Mori Y (2001) Lentinula edodes produces a multicomponent protein complex containing manganese (II)-dependent peroxidase, laccase and beta-glucosidase. FEMS Microbiol Lett 200: 175–179. Maltseva OV, Niku-Paavola ML, Leontievsky AA, Myasoedova NM & Golovleva LA (1991) Ligninolytic enzymes of the white rot fungus Panus tigrinus. Biotechnol Appl Biochem 13: 291–302. Marbach I, Harel E & Mayer AM (1984) Molecular properties of extracellular Botrytis cinerea laccase. Phytochemistry 23: 2713–2717. Martinez D, Larrondo LF, Putnam N, et al. (2004) Genome sequence of the lignocellulose degrading fungus Phanerochaete chrysosporium strain RP78. Nature Biotechnol 22: 695–700. Martins LO, Soares CM, Pereira MM, Teixeira M, Costa T, Jones GH & Henriques AO (2002) Molecular and biochemical characterization of a highly stable bacterial laccase that occurs as a structural component of the Bacillus subtilis endospore coat. J Biol Chem 277: 18849–18859. Matsubara T & Iwasaki H (1972) Occurrence of laccase and tyrosinase in fungi of Agaricales and comparative study of laccase from Russula delica and Russula pseudodelica. Bot Mag Tokyo 85: 71. Mayer AM (1987) Polyphenol oxidases in plants – recent progress. Phytochemistry 26: 11–20. Mayer AM & Staples RC (2002) Laccase: new functions for an old enzyme. Phytochemistry 60: 551–565. Messerschmidt A (1997) Multi-Copper Oxidases. World Scientific, Singapore. Min KL, Kim YH, Kim YW, Jung HS & Hah YC (2001) Characterization of a novel laccase produced by the woodrotting fungus Phellinus ribis. Arch Biochem Biophys 392: 279–286. Molitoris HP & Esser K (1970) The phenoloxidases of the ascomycete Podospora anserine. V. Properties of laccase I after further purification. Arch Mikrobiol 72: 267–296. Monteiro MC & de Carvalho MEA (1998) Pulp bleaching using laccase from Trametes versicolor under high temperature and alkaline conditions. Appl Biochem Biotechnol 70: 983–993. Munoz C, Guillen F, Martinez AT & Martinez MJ (1997) Induction and characterization of laccase in the ligninolytic fungus Pleurotus eryngii. Curr Microbiol 34: 1–5.


c

P. Baldrian

Munoz C, Guillen F, Martinez AT & Martinez MJ (1997) Laccase isoenzymes of Pleurotus eryngii: characterization, catalytic properties, and participation in activation of molecular oxygen and Mn21 oxidation. Appl Environ Microbiol 63: 2166–2174. Nagai M, Sato T, Watanabe H, Saito K, Kawata M & Enei H (2002) Purification and characterization of an extracellular laccase from the edible mushroom Lentinula edodes, and decolorization of chemically different dyes. Appl Microbiol Biotechnol 60: 327–335. Nannipieri P & Bollag JM (1991) Use of enzymes to detoxify pesticide-contaminated soils and waters. J Environ Qual 20: 510–517. Nannipieri P, Gelsomino A & Felici M (1991) Method to determine guaiacol oxidase activity in soil. Soil Sci Soc Am J 55: 1347–1352. Neufeld HA, Latterell FM, Green LF & Weintraub RL (1958) Oxidation of meta-polyhydroxyphenols by enzymes from Piricularia oryzae and Polyporus versicolor. Arch Biochem 76: 317–327. Ng TB & Wang HX (2004) A homodimeric laccase with unique characteristics from the yellow mushroom Cantharellus cibarius. Biochem Biophys Res Commun 313: 37–41. Nicole M, Chamberland H, Geiger JP, Lecours N, Valero J, Rio B & Ouellette GB (1992) Immunocytochemical localization of laccase L1 in wood decayed by Rigidoporus lignosus. Appl Environ Microbiol 58: 1727–1739. Nicole M, Chamberland H, Rioux D, Lecours N, Rio B, Geiger JP & Ouellette GB (1993) A cytochemical study of extracellular sheaths associated with Rigidoporus lignosus during wood decay. Appl Environ Microbiol 59: 2578–2588. Nishizawa Y, Nakabayashi K & Shinagawa E (1995) Purification and characterization of laccase from white-rot fungus Trametes sanguinea M85-2. J Ferment Bioeng 80: 91–93. Nyanhongo GS, Gomes J, Gübitz GM, Zvauya R, Read J & Steiner W 2002 Decolorization of textile dyes by laccases from a newly isolated strain of Trametes modesta. Water Res 36: 1449–1456. Oda Y, Adachi K, Aita I, Ito M, Aso Y & Igarashi H (1991) Purification and properties of laccase excreted by Pycnoporus coccineus. Agric Biol Chem 55: 1393–1395. Paice MG, Bourbonnais R, Reid ID, Archibald FS & Jurasek L (1995) Oxidative bleaching enzymes – a review. J Pulp Paper Sci 21: 280–284. Palmieri G, Cennamo G, Faraco V, Amoresano A, Sannia G & Giardina P (2003) Atypical laccase isoenzymes from copper supplemented Pleurotus ostreatus cultures. Enzyme Microb Technol 33: 220–230. Palmieri G, Giardina P, Bianco C, Fontanella B & Sannia G (2000) Copper induction of laccase isoenzymes in the ligninolytic fungus Pleurotus ostreatus. Appl Environ Microbiol 66: 920–924. Palmieri G, Giardina P, Bianco C, Scaloni A, Capasso A & Sannia G (1997) A novel white laccase from Pleurotus ostreatus. J Biol Chem 272: 31301–31307. Palmieri G, Giardina P, Marzullo L, Desiderio B, Nitti G, Cannio R & Sannia G (1993) Stability and activity of a phenol oxidase


239


from the ligninolytic fungus Pleurotus ostreatus. Appl Microbiol Biotechnol 39: 632–636. Palonen H, Saloheimo M, Viikari L & Kruus K (2003) Purification, characterization and sequence analysis of a laccase from the ascomycete Mauginiella sp. Enzyme Microb Technol 33: 854–862. Peralta-Zamora P, Pereira CM, Tiburtius ERL, Moraes SG, Rosa MA, Minussi RC & Durán N (2003) Decolorization of reactive dyes by immobilized laccase. Appl Catal B-Environ 42: 131–144. Perez J, Martinez J & de la Rubia T (1996) Purification and partial characterization of a laccase from the white rot fungus Phanerochaete flavido-alba. Appl Environ Microbiol 62: 4263–4267. Perie FH, Reddy GVB, Blackburn NJ & Gold MH (1998) Purification and characterization of laccases from the whiterot basidiomycete Dichomitus squalens. Arch Biochem Biophys 353: 349–355. Perry CR, Matcham SE, Wood DA & Thurston CF (1993) The structure of laccase protein and its synthesis by the commercial mushroom Agaricus bisporus. J Gen Microbiol 139: 171–178. Petroski RJ, Peczynska-Czoch W & Rosazza JP (1980) Analysis, production, and isolation of an extracellular laccase from Polyporus anceps. Appl Environ Microbiol 40: 1003–1006. Petter R, Kang BS, Boekhout T, Davis BJ & Kwon-Chung KJ (2001) A survey of heterobasidiomycetous yeasts for the presence of the genes homologous to virulence factors of Filobasidiella neoformans, CNLAC1 and CAP59. Microbiology 147: 2029–2036. Pickard M, Roman AR, Tinoco R & Vazquez-Duhalt R (1999) Polycyclic aromatic hydrocarbon metabolism by white rot fungi and oxidation by Coriolopsis gallica UAMH 8260 laccase. Appl Environ Microbiol 65: 3805–3809. Pind A, Freeman C & Lock MA (1994) Enzymatic degradation of phenolic materials in peatlands – measurement of phenol oxidase activity. Plant Soil 159: 227–231. Piontek K, Antorini M & Choinowski T (2002) Crystal structure of a laccase from the fungus Trametes versicolor at 1.90angstrom resolution containing a full complement of coppers. J Biol Chem 277: 37663–37669. Pointing SB, Pelling AL, Smith GJD, Hyde KD & Reddy CA (2005) Screening of basidiomycetes and xylariaceous fungi for lignin peroxidase and laccase gene-specific sequences. Mycol Res 109: 115–124. Pozdnyakova NN, Rodakiewicz-Nowak J & Turkovskaya OV (2004) Catalytic properties of yellow laccase from Pleurotus ostreatus D1. J Mol Catal B 30: 19–24. Ranocha P, Chabannes M, Chamayou S, Danoun S, Jauneau A, Boudet AM & Goffner D (2002) Laccase down-regulation causes alterations in phenolic metabolism and cell wall structure in poplar. Plant Physiol 129: 1–11. Rehman AU & Thurston CF (1992) Purification of laccase-I from Armillaria mellea. J Gen Microbiol 138: 1251–1257.


Reid ID & Paice MG (1994) Biological bleaching of kraft pulps by white-rot fungi and their enzymes. FEMS Microbiol Rev 13: 369–376. Reinhammar B & Malmstrom BG (1981) Blue copper-containing oxidases. Copper Proteins. Metal Ions in Biology, Vol. 3 (Spiro TG, ed.), pp. 109–149. Wiley, New York, USA. Robles A, Lucas R, Martinez-Canamero M, Ben Omar N, Perez R & Galvez A (2002) Characterisation of laccase activity produced by the hyphomycete Chalara (syn. Thielavopsis) paradoxa CH32. Enzyme Microb Technol 31: 516–522. Rodriguez A, Falcon MA, Carnicero A, Perestelo F, Delafuente G & Trojanowski J (1996) Laccase activities of Penicillium chrysogenum in relation to lignin degradation. Appl Microbiol Biotechnol 45: 399–403. Rodriguez E, Pickard MA & Vazquez-Duhalt R (1999) Industrial dye decolorization by laccases from ligninolytic fungi. Curr Microbiol 38: 27–32. Rogalski J, Dawidowicz AL & Leonowicz A (1990) Purification and immobilization of the inducible form of extracellular laccase of the fungus Trametes versicolor. Acta Biotechnol 10: 261–269. Rogalski J, Wojtas-Wasilewska M, Apalovic R & Leonowicz A (1991) Affinity chromatography as a rapid and convenient method for purification of fungal laccases. Biotechnol Bioeng 37: 770–777. Roper JC, Sarkar JM, Dec J & Bollag JM (1995) Enhanced enzymatic removal of chlorophenols in the presence of cosubstrates. Water Res 29: 2720–2724. Rosconi F, Fraguas LF, Mart´ınez-Drets G & Castro-Sowinski S (2005) Purification and characterization of a periplasmic laccase produced by Sinorhizobium meliloti. Enzyme Microb Technol 36: 800–807. Rosenbrock P, Buscot F & Munch JC (1995) Fungal succession and changes in the fungal degradation potential during the initial stage of litter decomposition in a black alder Forest [Alnus glutinosa (L) Gaertn]. Eur J Soil Biol 31: 1–11. Royarcand L & Archibald FS (1991) Direct dechlorination of chlorophenolic compounds by laccases from Trametes (Coriolus) versicolor. Enzyme Microb Technol 13: 194–203. Ryan S, Schnitzhofer W, Tzanov T, Cavaco-Paulo A & Gubitz GM (2003) An acid-stable laccase from Sclerotium rolfsii with potential for wool dye decolorization. Enzyme Microb Technol 33: 766–774. Saiya-Cork KR, Sinsabaugh RL & Zak DR (2002) The effects of long term nitrogen deposition on extracellular enzyme activity in an Acer saccharum forest soil. Soil Biol Biochem 34: 1309–1315. Salas C, Lobos S, Larrain J, Salas L, Cullen D & Vicuna R (1995) Properties of laccase isoenzymes produced by the basidiomycete Ceriporiopsis subvermispora. Biotechnol Appl Biochem 21: 323–333. Sannia G, Giardina P, Luna M, Rossi M & Buonocore V (1986) Laccase from Pleurotus ostreatus. Biotechnol Lett 8: 797–800. Saparrat MCN, Guillen F, Arambarri AM, Martinez AT & Martinez MJ (2002) Induction, isolation, and characterization


c

240

of two laccases from the white rot basidiomycete Coriolopsis rigida. Appl Environ Microbiol 68: 1534–1540. Sassoon J & Mooibroek H (2001) A system of categorizing enzyme-cell wall associations in Agaricus bisporus, using operational criteria. Appl Microbiol Biotechnol 56: 613–622. Savoie JM (2001) Variability in brown line formation and extracellular laccase production during interaction between white-rot basidiomycetes and Trichoderma harzianum biotype Th2. Mycologia 93: 243–248. Savoie JM, Mata G & Billette C (1998) Extracellular laccase production during hyphal interactions between Trichoderma sp and Shiitake, Lentinula edodes. Appl Microbiol Biotechnol 49: 589–593. Scháneˇl L & Esser K (1971) The phenoloxidases of the ascomycete Podospora anserina. VIII. Substrate specificity of laccases with different molecular structure. Arch Mikrobiol 77: 111–117. Scheel T, Hofer M, Ludwig S & Holker U (2000) Differential expression of manganese peroxidase and laccase in white-rot fungi in the presence of manganese or aromatic compounds. Appl Microbiol Biotechnol 54: 686–691. Scherer M & Fischer R (1998) Purification and characterization of laccase II of Aspergillus nidulans. Arch Microbiol 170: 78–84. Schliephake K, Mainwaring DE, Lonergan GT, Jones IK & Baker WL (2000) Transformation and degradation of the disazo dye Chicago Sky Blue by a purified laccase from Pycnoporus cinnabarinus. Enzyme Microb Technol 27: 100–107. Schlosser D, Grey R & Fritsche W (1997) Patterns of ligninolytic enzymes in T. versicolor. Distribution of extra- and intracellular enzyme activities during cultivation on glucose, wheat straw and beech wood. Appl Microbiol Biotechnol 47: 412–418. Schlosser D & Höfer C (2002) Laccase-catalyzed oxidation of Mn21 in the presence of natural Mn31 chelators as a novel source of extracellular H2O2 production and its impact on manganese peroxidase. Appl Environ Microbiol 68: 3514–3521. Schneider P, Caspersen MB, Mondorf K, Halkier T, Skov LK, Ostergaard PR, Brown KM, Brown SH & Xu F (1999) Characterization of a Coprinus cinereus laccase. Enzyme Microb Technol 25: 502–508. Schultz A, Jonas U, Hammer E & Schauer F (2001) Dehalogenation of chlorinated hydroxybiphenyls by fungal laccase. Appl Environ Microbiol 67: 4377–4381. Sealey J, Ragauskas AJ & Elder TJ (1999) Investigations into laccase-mediator delignification of kraft pulps. Holzforschung 53: 498–502. Sethuraman A, Akin DE & Eriksson KEL (1999) Production of ligninolytic enzymes and synthetic lignin mineralization by the bird’s nest fungus Cyathus stercoreus. Appl Microbiol Biotechnol 52: 689–697. Shin KS & Lee YJ (2000) Purification and characterization of a new member of the laccase family from the white-rot basidiomycete Coriolus hirsutus. Arch Biochem Biophys 384: 109–115. Shleev SV, Morozova O, Nikitina O, Gorshina ES, Rusinova T, Serezhenkov VA, Burbaev DS, Gazaryan IG & Yaropolov AI


c

P. Baldrian

(2004) Comparison of physico-chemical characteristics of four laccases from different basidiomycetes. Biochimie 86: 693–703. Sigoillot C, Record E, Belle V, Robert JL, Levasseur A, Punt PJ, van den Hondel CAMJ, Fournel A, Sigoillot JC & Asther M (2004) Natural and recombinant fungal laccases for paper pulp bleaching. Appl Microbiol Biotechnol 64: 346–352. Slomczynski D, Nakas JP & Tanenbaum SW (1995) Production and characterization of laccase from Botrytis cinerea 61–34. Appl Environ Microbiol 61: 907–912. Smirnov SA, Koroleva OV, Vavrilova VP, Belova AB & Klyachko NL (2001) Laccases from basidiomycetes: physicochemical characteristics and substrate specificity towards methoxyphenolic compounds. Biochemistry (Mosc) 66: 774–779. Soares GMB, Amorim MTP, Hrdina R & Costa-Ferreira M (2002) Studies on the biotransformation of novel disazo dyes by laccase. Process Biochem 37: 581–587. Solomon EI, Chen P, Metz M, Lee SK & Palmer AE (2001) Oxygen binding, activation, and reduction to water by copper proteins. Angew Chem 40: 4570–4590. Solomon EI, Sundaram UM & Machonkin TE (1996) Multicopper oxidases and oxygenases. Chem Rev 96: 2563–2605. Srinivasan C, D’Souza TM, Boominathan K & Reddy CA (1995) Demonstration of laccase in the white rot basidiomycete Phanerochaete chrysosporium BKM-F1767. Appl Environ Microbiol 61: 4274–4277. Steffen KT, Hofrichter M & Hatakka A (2002) Purification and characterization of manganese peroxidases from the litterdecomposing basidiomycetes Agrocybe praecox and Stropharia coronilla. Enzyme Microb Technol 30: 550–555. Sterjiades R, Dean JFD & Eriksson KEL (1992) Laccase from sycamore maple (Acer pseudoplatanus) polymerizes monolignols. Plant Physiol 99: 1162–1168. Stoj C & Kosman DJ (2003) Cuprous oxidase activity of yeast Fet3p and human ceruloplasmin: implication for function. FEBS Lett 554: 422–426. Sulistyaningdyah WT, Ogawa J, Tanaka H, Maeda C & Shimizu S (2004) Characterization of alkaliphilic laccase activity in the culture supernatant of Myrothecium verrucaria 24G-4 in comparison with bilirubin oxidase. FEMS Microbiol Lett 230: 209–214. Svobodová K (2005) The implications of ligninolytic enzymes in the decolorization of synthetic dyes by the white-rot fungus Irpex lacteus. PhD Thesis, Charles University, Prague, Czech Republic. Tatsumi K, Freyer A, Minard RD & Bollag JM (1994) Enzymemediated coupling of 3,4-dichloroaniline and ferulic acid – a model for pollutant binding to humic materials. Environ Sci Technol 28: 210–215. Thakker GD, Evans CS & Rao KK (1992) Purification and characterization of laccase from Monocillium indicum Saxena. Appl Microbiol Biotechnol 37: 321–323. Thurston CF (1994) The structure and function of fungal laccases. Microbiology 140: 19–26.


241


Timonen S & Sen R (1998) Heterogeneity of fungal and plant enzyme expression in intact Scots pine Suillus bovinus and Paxillus involutus mycorrhizospheres developed in natural forest humus. New Phytol 138: 355–366. Torres E, Bustos-Jaimes I & Le Borgne S (2003) Potential use of oxidative enzymes for the detoxification of organic pollutants. Applied Catal B-Environ 46: 1–15. Tsioulpas A, Dimou D, Iconomou D & Aggelis G (2002) Phenolic removal in olive oil mill wastewater by strains of Pleurotus spp. in respect to their phenol oxidase (laccase) activity. Biores Technol 84: 251–257. Ullah MA, Bedford CT & Evans CS (2000) Reactions of pentachlorophenol with laccase from Coriolus versicolor. Appl Microbiol Biotechnol 53: 230–234. Ullrich R, Huong LM, Dung NL & Hofrichter M (2005) Laccase from the medicinal mushroom Agaricus blazei: production, purification and characterization. Appl Microbiol Biotechnol 67: 357–363. Vaitkyavichyus RK, Belzhite VA, Chenas NK, Banis RG & Kulis YY (1984) Isolation and kinetic parameters of laccase from Polyporus anisoporus. Biochemistry (Mosc) 49: 859–863. Valásˇková V & Baldrian P (2005) Estimation of bound and free fractions of lignocellulose-degrading enzymes of wood-rotting fungi Pleurotus ostreatus, T. versicolor and Piptoporus betulinus. Res Microbiol doi: 10.1016/j.resmic.2005.06.004 (in press). Vares T & Hatakka A (1997) Lignin-degrading activity and ligninolytic enzymes of different white-rot fungi: effects of manganese and malonate. Can J Bot 75: 61–71. Vares T, Kalsi M & Hatakka A (1995) Lignin peroxidases, manganese peroxidases, and other ligninolytic enzymes produced by Phlebia radiata during solid-state fermentation of wheat straw. Appl Environ Microbiol 61: 3515–3520. Vares T, Lundell TK & Hatakka A (1992) Novel heme-containing enzyme possibly involved in lignin degradation by the whiterot fungus Junghuhnia separabilima. FEMS Microbiol Lett 99: 53–58. Vares T, Niemenmaa O & Hatakka A (1994) Secretion of ligninolytic enzymes and mineralization of 14C-ring-labelled synthetic lignin by three Phlebia tremellosa strains. Appl Environ Microbiol 60: 569–575. Vasconcelos AFD, Barbosa AM, Dekker RFH, Scarminio IS & Rezende MI (2000) Optimization of laccase production by Botryosphaeria sp. in the presence of veratryl alcohol by the response-surface method. Process Biochem 35: 1131–1138. Velazquez-Cedeno MA, Farnet AM, Ferre E & Savoie JM (2004) Variations of lignocellulosic activities in dual cultures of Pleurotus ostreatus and Trichoderma longibrachiatum on unsterilized wheat straw. Mycologia 96: 712–719. Velazquez-Cedeno M A, Mata G & Savoie JM (2002) Wastereducing cultivation of Pleurotus ostreatus and Pleurotus pulmonarius on coffee pulp: changes in the production of some lignocellulolytic enzymes. World J Microbiol Biotechnol 18: 201–207. Wahleithner JA, Xu F, Brown KM, Brown SH, Golightly EJ, Halkier T, Kauppinen S, Pederson A & Schneider P (1996) The


identification and characterization of four laccases from the plant pathogenic fungus Rhizoctonia solani. Curr Genet 29: 395–403. Wang HX & Ng TB (2004a) Purification of a novel lowmolecular-mass laccase with HIV-1 reverse transcriptase inhibitory activity from the mushroom Tricholoma giganteum. Biochem Biophys Res Commun 315: 450–454. Wang HX & Ng TB (2004b) A novel laccase with fair thermostability from the edible wild mushroom (Albatrella dispansus). Biochem Biophys Res Commun 319: 381–385. Wang HX & Ng TB (2004c) A new laccase from dried fruiting bodies of the monkey head mushroom Hericium erinaceum. Biochem Biophys Res Commun 322: 17–21. Wesenberg D, Kyriakides I & Agathos SN (2003) White-rot fungi and their enzymes for the treatment of industrial dye effluents. Biotechnol Adv 22: 161–187. Williams CJ, Shingara EA & Yavitt JB (2000) Phenol oxidase activity in peatlands in New York State: response to summer drought and peat type. Wetlands 20: 416–421. Williamson PR (1994) Biochemical and molecular characterization of the diphenol oxidase of Cryptococcus neoformans – identification as a laccase. J Bacteriol 176: 656–664. Wong YX & Yu J (1999) Laccase-catalyzed decolorization of synthetic dyes. Water Res 33: 3512–3520. Wood DA (1980) Production, purification and properties of extracellular laccase of Agaricus bisporus. J Gen Microbiol 117: 327–338. Xiao YZ, Tu XM, Wang J, Zhang M, Cheng Q, Zeng WY & Shi YY (2003) Purification, molecular characterization and reactivity with aromatic compounds of a laccase from basidiomycete Trametes sp. strain AH28-2. Appl Microbiol Biotechnol 60: 700–707. Xu F (1996) Oxidation of phenols, anilines, and benzenethiols by fungal laccases: Correlation between activity and redox potentials as well as halide inhibition. Biochemistry 35: 7608–7614. Xu F (1997) Effects of redox potential and hydroxide inhibition on the pH activity profile of fungal laccases. J Biol Chem 272: 924–928. Xu F, Kulys JJ, Duke K, Li KC, Krikstopaitis K, Deussen HJW, Abbate E, Galinyte V & Schneider P (2000) Redox chemistry in laccase-catalyzed oxidation of N-hydroxy compounds. Appl Environ Microbiol 66: 2052–2056. Xu F, Shin WS, Brown SH, Wahleithner JA, Sundaram UM & Solomon EI (1996) A study of a series of recombinant fungal laccases and bilirubin oxidase that exhibit significant differences in redox potential, substrate specificity, and stability. Biochim Biophys Acta 1292: 303–311. Yaver DS, Xu F, Golightly EJ, Brown KM, Brown SH, Rey MW, Schneider P, Halkier T, Mondorf K & Dalboge H (1996) Purification, characterization, molecular cloning, and expression of two laccase genes from the white rot basidiomycete Trametes villosa. Appl Environ Microbiol 62: 834–841.


c

242

Yavmetdinov IS, Stepanova EV, Gavrilova VP, Lokshin BV, Perminova IV & Koroleva OV (2003) Isolation and characterization of humin-like substances produced by wooddegrading white rot fungi. Appl Biochem Microbiol 39: 257–264. Yoshida H (1883) Chemistry of lacquer (Urushi). Part 1. J Chem Soc 43: 472–486. Yoshitake A, Katayama Y, Nakamura M, Iimura Y, Kawai S & Morohoshi N (1993) N-Linked carbohydrate chains protect laccase-III from proteolysis in Coriolus versicolor. J Gen Microbiol 139: 179–185. Zavarzina AG, Leontievsky AA, Golovleva LA & Trofimov SY (2004) Biotransformation of soil humic acids by blue laccase of Panus tigrinus 8/18: an in vitro study. Soil Biol Biochem 36: 359–369.


c

P. Baldrian

Zhu XD, Gibbons J, Garcia-Rivera J, Casadevall A & Williamson PR (2001) Laccase of Cryptococcus neoformans is a cell wall-associated virulence factor. Infect Immun 69: 5589–5596. Zille A, Tzanov T, Gubitz GM & Cavaco-Paulo A (2003) Immobilized laccase for decolorization of Reactive Black 5 dyeing effluent. Biotechnol Lett 25: 1473–1477. Zouari H, Labat M & Sayadi S (2002) Degradation of 4chlorophenol by the white rot fungus Phanerochaete chryosporium in free and immobilized cultures. Biores Technol 84: 145–150. Zouari N, Romette JL & Thomas D (1987) Purification and properties of 2 laccase isoenzymes produced by Botrytis cinerea. Appl Biochem Biotechnol 15: 213–225.


Fungal laccases â occurrence and properties - Wiley Online Library

Fungal laccases â occurrence and properties - Wiley Online Library

Fungal laccases â occurrence and properties - Wiley Online Library

Fungal laccases â occurrence and properties - Wiley Online Library