Download - 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
FEMS Microbiol Rev 30 (2006) 215–242 c� 2005 Federation of European Microbiological SocietiesPublished by Blackwell Publishing Ltd. All rights reserved
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)
FEMS Microbiol Rev 30 (2006) 215–242c� 2005 Federation of European Microbiological SocietiesPublished by Blackwell Publishing Ltd. All rights reserved
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)
FEMS Microbiol Rev 30 (2006) 215–242 c� 2005 Federation of European Microbiological SocietiesPublished by Blackwell Publishing Ltd. All rights reserved
219Fungal laccases – occurrence and properties
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.
FEMS Microbiol Rev 30 (2006) 215–242c� 2005 Federation of European Microbiological SocietiesPublished by Blackwell Publishing Ltd. All rights reserved
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.
FEMS Microbiol Rev 30 (2006) 215–242 c� 2005 Federation of European Microbiological SocietiesPublished by Blackwell Publishing Ltd. All rights reserved
221Fungal laccases – occurrence and properties
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
FEMS Microbiol Rev 30 (2006) 215–242c� 2005 Federation of European Microbiological SocietiesPublished by Blackwell Publishing Ltd. All rights reserved
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.
FEMS Microbiol Rev 30 (2006) 215–242 c� 2005 Federation of European Microbiological SocietiesPublished by Blackwell Publishing Ltd. All rights reserved
223Fungal laccases – occurrence and properties
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/
FEMS Microbiol Rev 30 (2006) 215–242c� 2005 Federation of European Microbiological SocietiesPublished by Blackwell Publishing Ltd. All rights reserved
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
FEMS Microbiol Rev 30 (2006) 215–242 c� 2005 Federation of European Microbiological SocietiesPublished by Blackwell Publishing Ltd. All rights reserved
225Fungal laccases – occurrence and properties
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
FEMS Microbiol Rev 30 (2006) 215–242c� 2005 Federation of European Microbiological SocietiesPublished by Blackwell Publishing Ltd. All rights reserved
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
FEMS Microbiol Rev 30 (2006) 215–242 c� 2005 Federation of European Microbiological SocietiesPublished by Blackwell Publishing Ltd. All rights reserved
227Fungal laccases – occurrence and properties
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).
FEMS Microbiol Rev 30 (2006) 215–242c� 2005 Federation of European Microbiological SocietiesPublished by Blackwell Publishing Ltd. All rights reserved
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
FEMS Microbiol Rev 30 (2006) 215–242 c� 2005 Federation of European Microbiological SocietiesPublished by Blackwell Publishing Ltd. All rights reserved
229Fungal laccases – occurrence and properties
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
FEMS Microbiol Rev 30 (2006) 215–242c� 2005 Federation of European Microbiological SocietiesPublished by Blackwell Publishing Ltd. All rights reserved
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
FEMS Microbiol Rev 30 (2006) 215–242 c� 2005 Federation of European Microbiological SocietiesPublished by Blackwell Publishing Ltd. All rights reserved
231Fungal laccases – occurrence and properties
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.
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