Fungal Laccases Occurrence and Properties

Fungal laccases^occurrenceandpropertiesPetr 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, Vıdenska

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

Editor: Jiri Damborsky

Keywords

biotechnology; ecology; humic substances;

laccase; lignin; soil; wood-rotting fungi.

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 transforma-

tion or immobilization of xenobiotic compounds.

Introduction

Laccase is one of the very few enzymes that have been

studied since the end of 19th century. It was first demon-

strated 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 monophe-

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 sub-

strates than diphenols, e.g. catechol or hydroquinone.

Syringaldazine [N,N0-bis(3,5-dimethoxy-4-hydroxybenzyli-

dene 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 dia-

mines 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 inter-

action, 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 de-

scribed as a 45-kDa periplasmic protein with isoelectric

FEMS Microbiol Rev 30 (2006) 215–242 c� 2005 Federation of European Microbiological SocietiesPublished by Blackwell Publishing Ltd. All rights reserved

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 prop-

erties 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 (Duran & Esposito, 2000) and (Duran et al.,

2002) reviewed the literature concerning the use of immo-

bilized 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 envir-

onment.

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 ascomy-

cetes. Laccase was purified from phytopathogenic ascomy-

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 (Abdel-

Raheem & 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 Tricho-

derma and the ligninolytic Bothryosphaeria are ecologically

closely related to the wood-rotting basidiomycetes produ-

cing 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 syringal-

dazine-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 xylar-

iaceous ascomycetes. Among the 20 strains tested, genes

with sequences similar to basidiomycete laccases were de-

tected in three strains, all of them Xylaria sp. Two strains of

Xylaria sp. and one of Xylaria hypoxylon exhibited syringal-

dazine 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 asco-

mycetes and basidiomycetes. Until now, laccase was only

purified from the human pathogen Cryptococcus (Filobasi-

diella) neoformans. This basidiomycete yeast produces a

true laccase capable of oxidation of phenols and amino-

phenols and unable to oxidize tyrosine (Williamson, 1994).

The enzyme is tightly bound to the cell wall and contri-

butes to the resistance to fungicides (Zhu et al., 2001;

FEMS Microbiol Rev 30 (2006) 215–242c� 2005 Federation of European Microbiological SocietiesPublished by Blackwell Publishing Ltd. All rights reserved

216 P. Baldrian

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 (Machon-

kin 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 white-

rot fungi was detected in Gloeophyllum trabeum. Oxidation

of ABTS (2,20-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 Con-

iophora puteana (Lee et al., 2004) and oxidation of ABTS was

reported in Laetiporus sulphureus (Schlosser & Hofer, 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 lignino-

lytic enzymes, including laccases in ectomycorrhizal (ECM)

fungi (Cairney & Burke, 1998; Burke & Cairney, 2002). Gene

fragments with a high similarity to laccase from wood-

rotting fungi have been found in several isolates of ECM

species including Amanita, Cortinarius, Hebeloma, Lactar-

ius, 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 produc-

tion of an enzyme. In Paxillus involutus, a species containing

another putative laccase sequence, oxidation of syringalda-

zine has never been detected (Gunther et al., 1998; Timonen

& Sen, 1998). It seems that tyrosinase is the major pheno-

loxidase 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 (Ka-

nunfre & Zancan, 1998) or orchideoid mycorrhiza: Armil-

laria 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 exclu-

sively extracellular. While this is without exception true for

the lignin peroxidases and manganese peroxidases of white-

rot fungi, the situation is not the same with laccases.

Although most laccases purified so far are extracellular

enzymes, the laccases of wood-rotting fungi are usually also

found intracellularly. Most white-rot fungal species tested by

Blaich & Esser (1975) produced both extracellular and

intracellular laccases with isoenzymes showing similar pat-

terns 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

(Gunther 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


217Fungal laccases – occurrence and properties

Tab

le1.

Char

acte

rist

ics

of

lacc

ases

purified

from

fungi

Spec

ies

MW

(kD

a)pI

pH

optim

um

Km

(mM

)Te

mper

ature

optim

um

(1C

)Ref

eren

ceA

BTS

DM

PG

UA

SYR

ABTS

DM

PG

UA

SYR

Agar

icus

bis

poru

s96

5.6

Wood

(1980)

Agar

icus

bis

poru

s65

Perr

yet

al.(1

993)

Agar

icus

bla

zei

66

4.0

2.0

5.5

6.0

63

1026

4307

4U

llric

het

al.(2

005)

Agro

cybe

pra

ecox

66

4.0

Stef

fen

etal

.(2

002)

Alb

atre

lladis

pan

sus

62

4.0

70

Wan

g&

Ng

(2004b)

Arm

illar

iam

elle

aLa

cI

59

4.1

3.5

178

Reh

man

&Th

urs

ton

(1992)

Arm

illar

iam

elle

aLa

cII

Bill

al&

Thurs

ton

(1996)

Arm

illar

iam

elle

a80

3.1

Curir

etal

.(1

997)

Asp

ergill

us

nid

ula

ns

II80

6.5

55

Scher

er&

Fisc

her

(1998)

Botr

ytis

ciner

ea74

4.0

3.5

100

57

Slom

czyn

skie

tal

.(1

995)

Can

thar

ellu

sci

bar

ius

92

4.0

50

Ng

&W

ang

(2004)

Cer

iporiopsi

ssu

bve

rmis

pora

L171

3.4

3.0

4.0

3.0

30

2900

1600

Fuku

shim

a&

Kirk

(1995);

Wan

g&

Ng

(2004b)

Cer

iporiopsi

ssu

bve

rmis

pora

L268

4.8

3.0

4.0

5.0

20

7700

440

Fuku

shim

a&

Kirk,

(1995);

Wan

g&

Ng

(2004b)

Cer

rena

max

ima

57–6

73.5

160–3

00

50

Koro

leva

etal

.(2

001);

Shle

evet

al.(2

004)

Cer

rena

unic

olo

r66

4.0

Bek

ker

etal

.(1

990)

Cer

rena

unic

olo

r58

Kim

etal

.(2

002)

Chae

tom

ium

term

ophilu

m77

5.1

190

96

400

34

60

Chef

etz

etal

.(1

998)

Chal

ara

par

adoxa

67

4.5

4.5

6.5

6.5

770

14

720

10

230

3400

Roble

set

al.(2

002)

Colle

totr

ichum

gra

min

icola

85

6.0

214

Ander

son

&N

ichols

on

(1996)

Conio

thyr

ium

min

itan

s74

4.0

3.5

100

60

Dah

iya

etal

.(1

998)

Coprinus

ciner

eus

58

4.0

4.0

6.5

26

60–7

0Sc

hnei

der

etal

.(1

999)

Coprinus

frie

sii

60

3.5

5.0

8.0

41

Hei

nzk

illet

al.(1

998)

Coriolo

psi

sfu

lvoci

nner

ea54–6

53.5

70–9

0Sh

leev

etal

.(2

004);

Smirnov

etal

.(2

001)

Coriolo

psi

sgal

lica

84

4.2

–4.3

3.0

70

Cal

voet

al.(1

998)

Coriolo

psi

srigid

aI

66

3.9

2.5

3.0

12

328

Sapar

rat

etal

.(2

002)

Coriolo

psi

srigid

aII

66

3.9

2.5

3.0

11

348

Sapar

rat

etal

.(2

002)

Coriolu

shirsu

tus

55

4.0

Koro

ljova

-Sko

robogat

’ko

etal

.(1

998)

Coriolu

shirsu

tus

78

4.2

845

Lee

&Sh

in(1

999)

Coriolu

sm

axim

a57

Smirnov

etal

.(2

001)

Coriolu

szo

nat

us

60

4.6

55

Koro

ljova

etal

.(1

999)

Cry

pto

cocc

us

neo

form

ans

77

Will

iam

son

(1994)

Cya

thus

ster

core

us

70

3.5

4.8

Seth

ura

man

etal

.(1

999)

Dae

dal

eaquer

cina

69

3.0

2.0

4.0

4.5

7.0

38

48

93

131

70,55

Bal

drian

(2004)

Dic

hom

itus

squal

ens

c166

3.5

3.0

Perie

etal

.(1

998)

Dic

hom

itus

squal

ens

c266

3.6

3.0

Perie

etal

.(1

998)

Fom

esfo

men

tarius

52

Rogal

skie

tal

.(1

991)

Gan

oder

ma

luci

dum

67

425

Lalit

ha

Kum

ari&

Sirs

i(1972);

Ko

etal

.(2

001)

Gan

oder

ma

tsugae

Elle

ret

al.(1

998)

Gae

um

annom

yces

gra

min

is190

5.6

4.5

26

510

Eden

set

al.(1

999)

Her

iciu

mec

hin

aceu

m63

5.0

50

Wan

g&

Ng

(2004c)


218 P. Baldrian

Junghuhnia

separ

abili

ma

58–6

23.4

–3.6

Var

eset

al.(1

992)

Lact

ariu

spip

erat

us

67

Iwas

akie

tal

.(1

967)

Lentinula

edodes

Lcc1

72

3.0

4.0

4.0

4.0

108

557

917

40

Nag

aiet

al.(2

002)

Lentinus

edodes

65

3.0

Kofu

jita

etal

.(1

991)

Mag

nap

ort

he

grise

a70

6.0

118

30

Iyer

&C

hat

too

(2003)

Mar

asm

ius

quer

cophilu

s�60

4.0

–4.4

5.0

80

Farn

etet

al.(2

000)

Mar

asm

ius

quer

cophilu

sw60

4.8

–5.1

5.0

80

Farn

etet

al.(2

000)

Mar

asm

ius

quer

cophilu

s65

3.6

4.5

775

Ded

eyan

etal

.(2

000)

Mar

asm

ius

quer

cophilu

sz65

2.6

6.2

850

80

Farn

etet

al.,

(2002,2004)

Mar

asm

ius

quer

cophilu

s‰60

4.0

4.5

113

4.2

80

Farn

etet

al.(2

004)

Mau

gin

iella

sp.

63

4.8

–6.4

2.4

3.5

4.0

Palo

nen

etal

.(2

003)

Mel

anoca

rpus

albom

yces

80

4.0

3.5

5.0

–7.5

6.0

–7.0

65

Kiis

kinen

etal

.(2

002)

Monoci

llium

indic

um

100

Thak

ker

etal

.(1

992)

Myr

oth

eciu

mve

rruca

ria

62

Sulis

tyan

ingdya

het

al.(2

004)

Neu

rosp

ora

cras

sa64

Froeh

ner

&Er

ikss

on

(1974)

Ophio

stom

anovo

-ulm

i79

5.1

2.8

6.0

6.0

Bin

z&

Can

evas

cini(

1997)

Panae

olu

spap

ilionac

eus

60

3.0

8.0

51

Hei

nzk

illet

al.(1

998)

Panae

olu

ssp

hin

ctrinus

60

3.0

7.0

32

Hei

nzk

illet

al.(1

998)

Panus

tigrinus

64

2.9

–3.0

Mal

tsev

aet

al.(1

991)

Panus

tigrinus

63

Leontiev

sky

etal

.(1

997)

Phan

eroch

aete

flav

ido-a

lba

94

3.0

30

Pere

zet

al.(1

996)

Phan

eroch

aete

chry

sosp

orium

47

Srin

ivas

anet

al.(1

995)

Phel

linus

noxi

us

70

Gei

ger

etal

.(1

986)

Phel

linus

ribis

152

5.0

4.0

–6.0

6.0

207

38

11

Min

etal

.(2

001)

Phle

bia

radia

ta64

3.5

Var

eset

al.(1

995)

Phle

bia

trem

ello

sa64

Var

eset

al.(1

994)

Pholio

tam

uta

bili

sLe

onow

icz

&M

alin

ow

ska

(1982)

Phys

isporinus

rivu

losu

sLa

cc1

66

3.3

2.5

3.0

3.5

3.5

Hak

ala

etal

.(2

005)

Phys

isporinus

rivu

losu

sLa

cc2

67

3.3

2.5

3.0

3.5

3.5

Hak

ala

etal

.(2

005)

Phys

isporinus

rivu

losu

sLa

cc3

68

3.2

2.5

3.0

3.5

3.5

Hak

ala

etal

.(2

005)

Phys

isporinus

rivu

losu

sLa

cc4

68

3.1

2.5

3.0

3.5

3.5

Hak

ala

etal

.(2

005)

Pleu

rotu

ser

yngii

I65

4.1

4.5

1400

7600

55

Munoz

etal

.(1

997)

Pleu

rotu

ser

yngii

II61

4.2

4.5

400

8000

55

Munoz

etal

.(1

997)

Pleu

rotu

sflorida

77

4.1

30

000

Das

etal

.(2

000)

Pleu

rotu

sost

reat

us

67

3.6

5.8

50

Hublik

&Sc

hin

ner

(2000)

Pleu

rotu

sost

reat

us

POX

A1b

62

6.9

3.0

4.5

6.0

370

260

220

Gia

rdin

aet

al.(1

999)

Pleu

rotu

sost

reat

us

POX

A1w

61

6.7

3.0

3.0

–5.0

NA

6.0

90

2100

NA

130

45–6

5Pa

lmie

riet

al.(1

997)

Pleu

rotu

sost

reat

us

POX

A2

67

4.0

3.0

6.5

6.0

6.0

120

740

3100

140

25–3

5Pa

lmie

riet

al.(1

997)

Pleu

rotu

sost

reat

us

POX

A3a

83–8

54.1

3.6

5.5

6.2

70

14

000

36

35

Palm

ieri

etal

.(2

003)

Pleu

rotu

sost

reat

us

POX

A3b

83–8

54.3

3.6

5.5

6.2

74

8800

79

35

Palm

ieri

etal

.(2

003)

Pleu

rotu

sost

reat

us

POX

C59

2.9

3.0

3.0

–5.0

6.0

6.0

280

230

1200

20

50–6

0Pa

lmie

riet

al.(1

993,1997);

Sannia

etal

.(1

986)

Pleu

rotu

spulm

onar

ius

Lcc2

46

4.0

–5.5

6.0

–8.0

6.2

–6.5

210

550

12

50

De

Souza

&Pe

ralta

(2003)

Pleu

rotu

ssa

jor-

caju

IV55

3.6

2.1

92

Loet

al.(2

001)

Podosp

ora

anse

rine

383

Molit

oris

&Es

ser

(1970);

Durr

ens

(1981)



Tab

le1.

Continued

.

Spec

ies

MW

(kD

a)pI

pH

optim

um

Km

(mM

)Te

mper

ature

optim

um

(1C

)Ref

eren

ceA

BTS

DM

PG

UA

SYR

ABTS

DM

PG

UA

SYR

Poly

poru

san

ceps

5.0

–5.5

Petr

osk

iet

al.(1

980)

Poly

poru

san

isoporu

s58

3.4

Vai

tkya

vich

yus

etal

.(1

984)

Poly

poru

spin

situ

s66

3.0

5.0

22

Hei

nzk

illet

al.(1

998)

Pycn

oporu

sci

nnab

arin

us

63

3.0

4.0

–4.5

4.4

–5.0

330

30

Schlie

phak

eet

al.(2

000)

Pycn

oporu

sci

nnab

arin

us

81

3.7

4.0

Egger

tet

al.(1

996)

Pycn

oporu

sco

ccin

eus

70

Oda

etal

.(1

991)

Rhiz

oct

onia

sola

ni4

170

Iwas

akie

tal

.(1

967)

Rig

idoporu

slig

nosu

sB

55

3.7

3.0

6.2

80

480

Bonom

oet

al.(1

998)

Rig

idoporu

slig

nosu

sS

60

3.1

3.0

6.2

49

108

Bonom

oet

al.(1

998)

Russ

ula

del

ica

63

Mat

subar

a&

Iwas

aki(

1972)

Schiz

ophyl

lum

com

mune

62–6

4D

eV

ries

etal

.(1

986)

Scle

rotium

rolfsi

iSRL1

55

5.2

2.4

62

Rya

net

al.(2

003)

Scle

rotium

rolfsi

iSRL2

86

Rya

net

al.(2

003)

Stro

phar

iaco

ronill

a67

4.4

Stef

fen

etal

.(2

002)

Stro

phar

iaru

goso

annula

ta66

2.5

3.5

Schlo

sser

&H

ofe

r(2

002)

Thel

ephora

terr

estr

is66

3.4

4.8

5.0

16

121

345

Kan

unfr

e&

Zanca

n(1

998)

Tram

etes

gal

lica

Lac

I60

3.1

2.2

3.0

4.0

12

420

405

70

Dong

&Zh

ang

(2004)

Tram

etes

gal

lica

Lac

II60

3.0

2.2

3.0

4.0

9410

400

70

Dong

&Zh

ang

(2004)

Tram

etes

hirsu

te64–6

83.7

–4.0

63

Shle

evet

al.(2

004);

Var

es&

Hat

akka

(1997)

Tram

etes

multic

olo

rII

63

3.0

Leitner

etal

.(2

002)

Tram

etes

och

race

a64

4.7

90

(Shle

evet

al.(2

004)

Tram

etes

pubes

cens

LAP

265

2.6

14

72

360

6G

alhau

pet

al.(2

002)

Tram

etes

sanguin

ea62

3.5

Nis

hiz

awa

etal

.(1

995)

Tram

etes

trogii

70

3.3

;3.6

30

410

Gar

zillo

etal

.(1

998)

Tram

etes

vers

icolo

r68

2.5

3.5

4.0

37

15

55

Rogal

skie

tal

.(1

990);

Hofe

r&

Schlo

sser

(1999)

Tram

etes

villo

sa1

63

3.5

2.7

5.0

–5.5

Yav

eret

al.(1

996)

Tram

etes

villo

sa3

63

6.0

–6.5

2.7

5.0

–5.5

Yav

eret

al.(1

996)

Tram

etes

sp.A

H28-2

A62

4.2

4.5

25

25

420

50

Xia

oet

al.(2

003)

Tric

hoder

ma

sp.

71

Ass

avan

iget

al.(1

992)

Tric

holo

ma

gig

ante

um

43

4.0

70

Wan

g&

Ng

(2004a)

Volv

arie

llavo

lvac

ea58

3.7

3.0

4.6

5.6

30

570

10

45

Chen

etal

.(2

004)

ABTS

,2,20 -

azin

obis

(3-e

thyl

ben

zoth

iazo

line-

6-s

ulfonic

acid

);D

MP,

2,6

-dim

ethoxy

phen

ol;

GU

A,2-m

ethoxy

phen

ol(

guai

acol);

SYR.4-h

ydro

xy-3

,5-d

imet

hoxy

ben

zald

ehyd

e[(

4-h

ydro

xy-3

,5-d

imet

hoxy

phe-

nyl

)met

hyl

ene]

hyd

razo

ne

(syr

ingal

daz

ine)

.

The

spec

ies

are

liste

dunder

the

nam

esuse

din

the

origin

alre

fere

nce

s.

NA

,not

active

.� S

trai

n17,co

nst

itutive

form

.w S

trai

n17,in

duce

dw

ith

p-h

ydro

xyben

zoic

acid

.z S

trai

nC

7,co

nst

itutive

form

.‰St

rain

19,in

duce

dw

ith

feru

licac

id.


220 P. Baldrian

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 (Valaskova & Baldrian, 2005). Laccase activity is almost

exclusively associated with cell walls in the white-rot basi-

diomycete 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 com-

pounds 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-chemi-

cal 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 character-

ized (Table 1). Based on the published data we can draw

some general conclusions about laccases, taking into ac-

count 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 fascicu-

lare, 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) 103 66 000 61 000 71 000 43 000 383 000

pI 67 3.9 3.5 4.2 2.6 6.9

Temperature optimum ( 1C) 39 55 50 70 25 80

pH optimum 49 3.0 2.5 4.0 2.0 5.0

ABTS

2,6-Dimethoxyphenol 36 4.0 3.0 5.5 3.0 8.0

Guaiacol 24 4.5 4.0 6.0 3.0 7.0

Syringaldazine 31 6.0 4.7 6.0 3.5 7.0

KM (mM)

ABTS 36 39 18 100 4 770

2,6-Dimethoxyphenol 30 405 100 880 26 14 720

Guaiacol 23 420 121 1600 4 30 000

Syringaldazine 21 36 11 131 3 4307

kcat (s�1)

ABTS 12 24 050 5220 41 460 198 350 000

2,6-Dimethoxyphenol 12 3680 815 6000 100 360 000

Guaiacol 10 295 115 3960 90 10 800

Syringaldazine 4 21 500 18 400 25 500 16 800 28 000

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



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 quer-

cophilus (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 SDS-

PAGE) 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, pro-

duce isoenzymes that are closely related both structurally

and in their catalytic properties (Table 1). Different proper-

ties 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 isoen-

zymes 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 iso-

enzymes 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 ascomy-

cetes 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

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 na-

tive-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 (Slomc-

zynski et al., 1995). Other preparations from the same

species exhibited as much as 65–80% of saccharides includ-

ing 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 produc-

tion 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 oxi-

dases (BMCO) that catalyze a one-electron oxidation con-

comitantly 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 character-

ized by the presence of at least one type-1 (T1) copper,

together with at least three additional copper ions: one type-

2 (T2) and two type-3 (T3) copper ions, arranged in a

trinuclear cluster. The different copper centres can be

identified on the basis of their spectroscopic properties.

The T1 copper is characterized by a strong absorption

around 600 nm, whereas the T2 copper exhibits only weak

absorption in the visible region. The T2 site is electron

paramagnetic resonance (EPR)-active, whereas the two

copper ions of the T3 site are EPR-silent due to an

antiferromagnetic coupling mediated by a bridging ligand.

The substrates are oxidized by the T1 copper and the


222 P. Baldrian

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 (Hakuli-

nen 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 lac-

case 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 (Gar-

avaglia 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 differ-

ences 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

belong to the blue copper proteins because it lacks Cu1 and

contains one Mn atom per molecule. The structural differ-

ences 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

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). Unfortu-

nately, 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 cate-

chol 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 syringal-

dazine 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 deter-

mination 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 four-

electron 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 Myce-

liophthora 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 measure-

ments 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, consider-

ably lower than that reported for laccase, suggesting that this

class of enzyme is much less oxidizing than laccase.

Fig. 1. Catalytic cycle of laccase.



Due to the difficulties with distinguishing laccases from

other oxidases, the data in this review are based exclusively

on the reports concerning purified enzymes. However, the

direct comparison of biochemical data reported for different

fungal laccases that would be extremely important for the

biotechnological applicability is difficult, as different test

conditions have been used in different reports. There are only

a few works comparing laccase properties of enzymes from

different sources, e.g. the work of (Shleev 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), guaia-

col 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). Lac-

cases 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 phe-

nols, vanillic acid or its aldehyde. For the species capable of

oxidizing polycyclic aromatic hydrocarbons or pentachlor-

ophenol, 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-substi-

tuted compounds (guaiacol, o-phenylenediamine, caffeic

acid, catechol, dihydroxyphenylalanine, protocatechuic

acid, gallic acid and pyrogallol) were better substrates

than para-substituted compounds (p-phenylenediamine,

p-cresol, hydroquinone) and the lowest rates were obtained

with meta-substituted compounds (m-phenylenediamine,

orcinol, resorcinol and phloroglucinol) with crude laccase

preparations from L. litschaueri and P. brumalis (Blaich &

Esser, 1975). Similar results were also obtained with

T. versicolor and the ascomycetes P. anserina and Pyricularia

oryzae, whereas laccase from Ganoderma lucidum catalyzed

the oxidation of only ortho and para dihydroxyphenyl

compounds, p-phenylenediamine and polyphenols, not the

meta hydroxymethyl compounds or ascorbic acid (Fahraeus,

1961; Fahraeus & Ljunggren, 1961; Schanel & 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 (Thak-

ker et al., 1992). The relative oxidation rates for different

substrates in relation to the oxidation of 2,6-dimethoxyphe-

nol 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 sub-

strates.

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 white-

rot fungus T. versicolor oxidized Mn21 to Mn31 in the

presence of pyrophosphate (Hofer & 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 & Hofer, 2002).

The chelators probably decrease the high redox potential of

the Mn21/Mn31 couple. Mn21 oxidation involved conco-

mitant 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 & Hofer, 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 follow-

up reactions leading to H2O2 formation, which may initiate

or support peroxidase reactions (Schlosser & Hofer, 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 differ-

ence 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/


224 P. Baldrian

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) Ts, Tv

(4-Hydroxy-3-methoxyphenyl)acetic acid Pe

1,2,4,5-Tetramethoxybenzene Cs (KM: 6900; kcat: 1680), Cs (KM: 900; kcat: 3360)

1,2,4-Benzenetriol Bc

1,2-Benzenediol (catechol) 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

1,3-Dihydroxybenzene (resorcinol) Cn, Ts

1,4-Benzohydroquinone 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

1-Naphthol Ab, Bc, Gl

2-(3,4-Dihydroxyphenyl)-3,5,7-trihydroxy-4H-

chromen-4-one

Am

2-Chlorophenol Tv

2,20-Azinobis(3-ethylbenzothiazoline-6-sulfonic

acid)

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)

2,3-Dichlorophenol Tv

2,3-Dimethoxyphenol Ds

2,3,6-Trichlorophenol Tv

2,4,6-Trichlorophenol Tv

2,4,6-Trimethylphenol Pe

2,4-Dichlorophenol Tt, Tv

2,5-Dihydroxybenzoic acid Pi

2,6-Dichlorophenol Tt, Tv

2,6-Dimethoxy-1,4-benzohydroquinone Cr (KM: 107; kcat: 8580), Cr (KM: 89; kcat: 11 220)

2,6-Dimethoxyphenol 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)

2,7-Diaminofluorene Rl

2-Amino-3-(3,4-dihydroxyphenyl)propanoic acid Am, Cn, Ct (KM: 100), Le (KM: 650), Lp, Ma, Pa (KM: 3300), Ts, Tv (KM: 15 600)

2-Amino-3-hydroxybenzoic acid Pc

2-Amino-4-methylphenol Tt

2-Amino-4-nitrophenol Tt

2-Aminophenol Tt

2-Aminophenylamine Cn

2-Chlorobenzene-1,4-diol Tt

2-Chlorophenol Le (KM: 1350), Mq, Tt

2-Methoxy-1,4-benzohydroquinone Cr (KM: 216; kcat: 7620), Cr (KM: 229; kcat: 6300), Pe

2-Methoxy-4-[prop-1-enyl]phenol Ts

2-Methoxy-4-methylphenol Tt

2-Methoxyaniline Tt

2-Methoxyphenol (guaiacol) 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)

2-Methoxy-1,4-benzohydroquinone Pe

2-Methyl-1,4-benzohydroquinone Pe (KM: 1600), Pe (KM: 2100)

2-Methylanthracene Cg (kcat: 0.082)

2-Methylphenol Bc, Tt

2-Naphthol Bc, Gl

2,4-Dichlorophenol Mq

2,4,6-Trichlorophenol

3-(3,4-Dihydroxyphenyl)acrylic acid (caffeic acid) Am, Bc, Cs, Le (KM: 40), Mi, Mq, Ts, Tt



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

3-(4-Hydroxy-3-methoxyphenyl)acrylic acid (ferulic

acid)

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

3-(4-Hydroxyphenyl)acrylic acid Le (KM: 240), Pn, Rl, Tt

3,30-Dimethoxy-1,10-biphenyl-4,40-diamine Mi (KM: 25), Pr, Tc

3,4,5-Trihydroxybenzoic acid (gallic acid) Ab, Am, Bc, Ct (KM: 130), Le (KM: 130), Mq, Nc, On

3,4-Dihydroxybenzoic acid Cr, Mi, Mq, Pe, Tt

3,5-Cyclohexadiene-1,2-diol Pe

3,5-Dimethoxy-hydroxy-benzaldazine Bc

3-{[3-(3,4-Dihydroxyphenyl)prop-2-enoyl]oxy}-

1,4,5-trihydroxycyclohexanecarboxylic acid

Am, Ct

3-Amino-4-hydroxybenzenesulphonic acid Tt

3-Methoxyphenol Pr

4-(Hydroxymethyl)-2-methoxyphenol Cs (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

(coniferyl alcohol)

Pe, Rl

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-hydroxy-

3,5-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

(ascorbic acid)

Ab, Am, Bc, Lp, Mi, Nc, Pa (KM: 190)

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,N0-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


226 P. Baldrian

Inhibitor

Ca21 Le

Cd21 Dq, Le

Co21 Dq

Fe21 Ct, Po, Lp, Tc

Hg21 Ct, Dq, Le, Pu

Mn21 Dq, Pu

Rb1 Le

Sn21 Le

Zn21 Le, Po

1-Phenyl-2-thiourea Ct

2-Mercaptobenzothiazole Ct

2-Mercaptoethanol Ct, Pu, Te

3-(4-Hydroxyphenyl)acrylic acid Le

4-Nitrophenol Pz

8-Hydroxyquinoline Lp, Te

Ascorbic acid Ct, Tc

Cetylpyridinium bromide Ab

Cetyltriammonium bromide Ab, Tc

CN- Ab, Bc, Ct, Gl, Lp, Ma, Me, Mi, Pn, Po, Pz, Rl, Tg, Tr, Ts

Cysteine Ct, Ch, Dq, Gl, Le, Mq, Pc, Py, Sr, Te, Vv

Diethyldithiocarbamic acid Bc, Ch, Gl, Lp, Mi, Pp, Pc, Ps, Sr

Dithiothreitol Ch, Dq, Le, Pc, Py, Vv

EDTA Ct, Ma, Mq�, Vv

Glutathione Dq, Gl

Humic acid Pt

Hydroxylamine Po

KCl Le

Kojic acid Dq, Le, Po

NaCl Sr

NaF Ds, Me, Sr, Tt

NaN3 Ab, Bc, Ch, Ct, Dq, Ds, Gl, Le, Ma, Me, Mi, Pa, Pc, Po, Pp, Pr, Ps, Pu, Pz, Sr, Tc, Te, Tg, Tr, Ts, Vv

Salicylaldoxime Gl

SDS Ds, Mq�, Pu, Tr

Thiamine Sr

Thiogylcolic acid Ct, Mi, Po, Pr, Sr, Vv

Thiourea Ct, Dq

Trifluoroacetic acid Tr

Tropolone Ch, Le, Pc

Ab, Agaricus bisporus (Wood, 1980); Al, Agaricus blazei (Ullrich 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).

Table 3. Continued

Species



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 prati-

cola and T. versicolor formed different products from syr-

ingic 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

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 applica-

tions (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 inhibi-

tors of metal-containing oxidases like cyanide, sodium azide

or fluoride, there are some selective inhibitors for individual

oxidases. Carbon monoxide, 4-hexylresorcinol or salicylhy-

droxamic acid are examples of specific inhibitors of tyrosi-

nases 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 success-

ful. 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 inhibi-

tors: e.g. dithiothreitol, thioglycolic acid, cysteine and

Table 4. Reactivity of fungal laccases with different substrates. The numbers indicate the rate of substrate oxidation (%) compared to the oxidation of

2,6-dimethoxyphenol

Substrate

Species

Am Ct Ch Cy Ds Ma Me Mv Pr Pe Pc Sr Ts Tt Tv

2,6-Dimethoxyphenol 100.0 100.0 100.0 100.0 100.0 100.0 100.0 100.0 100.0 100.0 100.0 100.0 100.0 100.0 100.0

2-Amino-3-(3,4-dihydroxy-

phenyl)propanoic acid

55.9 46.0

4-Aminophenol 109.8 26.7

4-Aminophenylamine 62.9 14.7 194.8 56.0

4-Hydroxyindole 87.6 107.6

4-Methoxyphenol 7.3 9.8 19.8

4-Methylcatechol 69.3 31.4 103.5

ABTS 76.5 271.7 114.4 800.0 288.3 1.4 97.4 284.1 452.0 136.7 27.7

Caffeic acid 18.6 95.0 80.2 24.5

Catechol 74.2 44.9 59.2 34.9 33.1 21.6 74.3 18.3 76.0 27.9 110.7

Ferulic acid 111.8 48.4 8.5 76.0 116.3

Guaiacol 59.7 73.5 107.8 37.9 92.0 122.4 31.0 39.1 9.7 140.9 35.0 64.0 55.8 38.4

Hydroquinone 50.0 105.3 80.8 12.9 25.5 62.0 69.0 30.2 56.0

N,N-dimethyl-1,4-phenylenediamine 74.8 1.8 23.4 19.9 237.1

o-Anisidine 45.5 9.3

p-Anisidine 19.3 3.5

Pyrogallol 24.0 11.8 12.3 34.5 76.0 6.3

Syringaldazine 51.6 120.6 79.2 131.7 126.5

Syringic acid 115.2 46.9 14.1

Vanillic acid 61.8 7.3 8.1

Am, Armillaria mellea (Rehman & Thurston, 1992); Ct, Chaetomium thermophilum (Chefetz 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).


228 P. Baldrian

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 & Majcherc-

zyk, 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 media-

tors 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 & Majcherc-

zyk, 2000). A comprehensive screening for natural media-

tors was performed by (Camarero et al., 2005). Among 44

tested natural lignin-derived compounds they selected 10

phenolic compounds derived from syringyl, guaiacyl, and p-

hydroxyphenyl lignin units, characterized by the presence of

two, one or no methoxy substituents, respectively (in ortho

positions with respect to the phenolic hydroxyl). Syringal-

dehyde, acetosyringone, vanillin, acetovanillone, methyl

vanillate and p-coumaric acid have been found to be the

most effective for mediated oxidation using laccases of P.

cinnabarinus and T. villosa. Among them, syringaldehyde

and acetosyringone belong to the main products of both

biological and enzymatic degradation of syringyl-rich lig-

nins (Kirk & Farrell, 1987).

Laccases in thenatural environment

The considerable attention devoted to white-rot basidiomy-

cetes 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 mechan-

isms 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 intro-

duced 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 agricul-

tural 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 lignino-

lytic 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 succes-

sion (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



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 transfor-

mation 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 depolymer-

ization of humic substances and their synthesis from mono-

meric precursors; the balance of these two processes can be

influenced by the nature of the humic compounds (Zavarzi-

na 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 constitu-

ents 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).

Laccases are also actively produced during the compost-

ing 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 concen-

tration 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 environ-

ment, laccase was demonstrated to participate in the degra-

dation 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 accom-

panied 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 bio-

technological applications. They exhibit broad substrate

specificity and are thus able to oxidize a broad range of


230 P. Baldrian

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 Duran &

Esposito, 2000; Duran 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 lignocel-

lulosic 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-redox-

potential compounds and not the oxidation of more recal-

citrant 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 (Jo-

hannes 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

an immediately following reaction or when its redox poten-

tial 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-hydroxyben-

zotriazole, violuric acid and N-hydroxyacetanilide or TEM-

PO, a large number of studies have been performed on the

mechanisms of oxidation of nonphenolic substrates (Bour-

bonnais et al., 1998; Xu et al., 2000; Fabbrini et al., 2002;

Baiocco et al., 2003), the search for new mediators (Bour-

bonnais 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 deriva-

tives. 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 laccase-

mediator system for use in biotechnology.

In addition to substrate oxidation, laccase can also

immobilize soil pollutants by coupling to soil humic sub-

stances – 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 com-

pounds and anilines such as 3,4-dichloroaniline, 2,4,6-

trinitrotoluene 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 bio-

chemical properties of fungal laccases, their occurrence

under natural conditions and possible biotechnological use.

However, it leaves many important questions open: Why do

fungi produce laccase? What are the respective roles of

different isoenzymes? Do their biochemical characteristics



differ with respect to their function? The understanding of

the physiological role of laccase has not increased signifi-

cantly 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 2,4-

dichlorophenol 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

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.


232 P. Baldrian

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-mediator-

systems (Lignozyms-process). J Biotechnol 53: 163–202.

Call HP & Mucke I (1997) History, overview and applications of

mediated lignolytic systems, especially laccase-mediator-

systems (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, Martınez MJ & Martınez 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 manganese-

dependent 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.



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, Gunther 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

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.

Duran 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.

Duran 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 & Giovannozzi-

Sermanni 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


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) Laccase-

mediated 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


234 P. Baldrian

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 coal-

derived 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.


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


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


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 acid-

induced 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, Gunther 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, Gunther 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.



Gunther 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


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 & Steinbuchel 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 wood-

rotting 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 laccase-

like multicopper oxidase from Liriodendron tulipifera. Plant

Physiol Biochem 42: 27–33.

Hofer 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.

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 & Dale 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 & Huttermann 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 & Huttermann 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-4-

hydroxycinnamic 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


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


236 P. Baldrian

compost-dwelling deuteromycete Paecilomyces inflatus. Appl


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


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


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


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 2,4,6-

trichlorophenol by free and immobilized fungal laccase. Appl


Leontievsky A, Myasoedova N, Pozdnyakova N & Golovleva L

(1997) ‘‘Yellow’’ laccase of Panus tigrinus oxidizes non-

phenolic 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) Laccase-

like 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.


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.



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 & Huttermann 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 wood-

rotting 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.

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


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


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, Gubitz 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


238 P. Baldrian

from the ligninolytic fungus Pleurotus ostreatus. Appl Microbiol


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 & Duran 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 white-

rot 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 1.90-

angstrom 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


Rodriguez E, Pickard MA & Vazquez-Duhalt R (1999) Industrial

dye decolorization by laccases from ligninolytic fungi. Curr


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 co-

substrates. Water Res 29: 2720–2724.

Rosconi F, Fraguas LF, Martınez-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



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.

Schanel 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 & Hofer 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

(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 litter-

decomposing 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.

Svobodova 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) Enzyme-

mediated 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.


240 P. Baldrian

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


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.

Valaskova 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 white-

rot 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


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) Waste-

reducing 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 low-

molecular-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


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.



Yavmetdinov IS, Stepanova EV, Gavrilova VP, Lokshin BV,

Perminova IV & Koroleva OV (2003) Isolation and

characterization of humin-like substances produced by wood-

degrading 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.

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 4-

chlorophenol 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.


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Fungal Laccases Occurrence and Properties

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