Fungal laccase: properties and activity on lignin.

J. Basic Microbiol. 41 (2001) 3–4, 185–227

Review

(Department of Biochemistry, Maria-Curie-Sklodowska University, Pl-20031 Lublin, Poland; 1Schoolof Forest Resources, Chungbuk National University, Cheongju 360-763, Korea and �� !!"# ��$��%

Fungal laccase: properties and activity on lignin

ANDRZEJ LEONOWICZ, NAM-SEOK CHO1, JOLANTA LUTEREK, ANNA WILKOLAZKA,MARIA WOJTAS-WASILEWSKA, ANNA MATUSZEWSKA, MARTIN HOFRICHTER3,DIRK WESENBERG2 and JERZY ROGALSKI

(Received 17 October 2000/Accepted 12 April 2001)

The sources of ligninocellulose that occur in various forms in nature are so vast that they can only becompared to those of water. The results of several, more recent experiments showed that laccaseprobably possesses the big ability for “lignin-barrier” breakdown of ligninocellulose. The degradationof this compound is currently understood as an enzymatic process mediated by small molecules,therefore, this review will focus on the role of these mediators and radicals working in concert withenzymes. The fungi having a versatile machinery of enzymes are able to attack directly the “lignin-barrier” or can use a multienzyme system including “feed-back” type enzymes allowing for simulta-neous transformation of lignin and carbohydrate compounds.

Fungal laccase (benzenediol:oxygen oxidoreductase, EC 1.10.3.2) is an enzyme secretedinto the medium by mycelia of Basidiomycetes, Ascomycetes and Deuteromycetes (BOLLAGand LEONOWICZ 1984, AGEMATU et al. 1993). In some fungal strains, laccase can be in-duced by anilines (FAHRAEUS et al. 1958, BOLLAG and LEONOWICZ, 1984), methoxyphe-nolic acids (ROGALSKI et al. 1992, LEONOWICZ et al. 1997a) lignin preparations (ROGALSKIet al. 1991) or heat shock (FINK-BOOTS et al. 1997 and 1999). However, the most effectiveinducer of laccase in fungi is 2,5-xylidine. The stimulating effect of this compound was firstdescribed in 1958 by FAHRAEUS et al. for the white-rot basidiomycete Trametes (Coriolus)versicolor. The authors selected 2,5-xylidine among various aniline derivatives and foundan over 160-fold stimulation of laccase activity when the compound was added to growingfungal cultures. In addition, it was shown that not only in Trametes versicolor (FAHRAEUSand REINHAMMAR 1967), but also in other lignin degrading fungi such as Fomes annosum,Pholiota mutabilis, Pleurotus ostreatus (AGEMATU et al. 1993) and Phlebia radiata(ROGALSKI and LEONOWICZ 1992), laccase levels are enhanced in the presence of 2,5-xylidine. Also certain phenolic compounds were found to stimulate laccase production, e.g.,ferulic acid which considerably induces the enzyme in the cultures of Pholiota mutabilis,Pleurotus ostreatus and Trametes versicolor (LEONOWICZ et al. 1978).

The highest amounts of laccase are produced by white-rot fungi (wood-decaying basidio-mycetes; LEONOWICZ et al. 1997a). Biochemically, laccase is an enzyme which oxidizes avariety of aromatic hydrogen donors. Thus, it catalyzes the removal of an electron and aproton from phenolic hydroxyl or aromatic amino groups to form free phenoxy radicals andamino radicals, respectively. Moreover, this group of copper-containing laccase having fourcopper atoms all in the 2+ oxidation state in the active site (i.e. blue oxidase) does not onlyoxidize phenolic and methoxyphenolic acids, but also decarboxylates them (AGEMATU et al.1993) and attacks their methoxyl groups through demethylation (LEONOWICZ et al. 1984) or

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186 A. LEONOWICZ et al.

demethoxylation (POTTHAST et al. 1995). These reactions may represent an important stepin the initial transformation of the lignin polymer (LEONOWICZ et al. 1984, AGEMATU et al.1993, POTTHAST 1995). It has also been reported that the enzyme is involved in the trans-formation and detoxification of reactive lignin degradation products (BOLLAG et al. 1988).Laccase reacts with polyphenols and other lignin-derived aromatic compounds, which, inturn, can be both polymerized and depolymerized or act as low-molecular weight mediators(BOURBONNAIS et al. 1995).&��'�� (�� )��

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Laccase is also involved in various physiological processes of economic and ecologicalrelevance. It is engaged in the development of fungal fruit bodies (LEATHAM andSTAHMANN 1981), pigmentation of fungal spores (CLUTTERBUCK 1972), pathogenicity ofmoulds (RIGLING and VAN ALFEN 1991), sexual differentiation (AISEMBERG et al. 1989),and rhizomorph formation (WORALL et al. 1986).

The biodegradation of the lignocellulose constituents, cellulose, hemicelluloses and lig-nin, is achieved by enzymatic activities. The conversion of cellulose and hemicellulosesinto simple sugars has been studied for a long time. A large number of micro-organisms(bacteria, fungi, protozoans) make use of a whole string of hydrolases which are able toproduce large quantities of mono- and disaccharides from all polysaccharide components inligninocellulose. The degradation, however, is effected by the occurrence of polysaccha-rides in a complex with lignin because the latter forms a barrier against the microbial attackby hydrolytic enzymes. The lignin barrier also complicates cellulose production in the pulpand paper industry. For these ecological, economic and other reasons, research into thebiotransformation of lignin has been carried out for decades.

Wood-rotting basidiomycetes penetrate wood tissues in order to come into contact withthe easily assimilable carbohydrate constituents of the lignocellulosic complex. The white-rot group of these fungi, which has a versatile machinery of enzymes co-operating withcertain secondary metabolites of fungi, is capable of attacking the lignin barrier efficiently.These fungi use a multi-enzyme system including the so-called “feed-back” type enzymesto transform and degrade all structural elements of the lignocellulosic complex (polysaccha-rides and lignin). The currently known enzymes of white-rot fungi involved in wood degra-dation can be divided into three groups. The first can attack the wood constituents or theirprimary degradation products directly; this group includes the cellulase and hemicellulasecomplexes, laccase, different peroxidases, protocatechuate-3,4 dioxygenase, etc. The sec-ond group of enzymes, comprising among others aryl alcohol oxidase and glyoxal oxidase,co-operates with the first group by providing H2O2 for the peroxidases, but these enzymesdo not attack wood components directly. The third enzyme group represented by glucoseoxidase and cellobiose:quinone oxidoreductase (cellobiose dehydrogenase) includes feed-back type enzymes which play a key role in joining the metabolic chains during the bio-

Fungal laccase: properties and activity on lignin 187

transformation of high-molecular mass wood constituents. All these enzymes, includinglaccase, can act separately or in co-operation.

Among the white-rot fungi, the polyporous fungus Trametes versicolor and the corticoidfungus Phanerochaete chrysosporium have been particularly studied in detail due to theirhigh ligninolytic activities. This makes them promising candidates for applications in thepulp and paper industry (e.g., bleaching, pulping, waste water treatment; MANZANARESet al. 1995, PASZCZYNSKI and CRAWFORD 1995). The production of high amounts of lac-case by Trametes versicolor has already been known for decades (FAHRAEUS et al. 1958)while the lignin modifying activity of this enzyme was detected relatively early(LEONOWICZ and TROJANOWSKI 1965). The situation is different in the case of Phanero-chaete chrysosporium, the most-investigated white-rot fungus, since laccase was discoveredin this fungus relatively late and is only produced in certain strains and under special condi-tions (SRINIVASAN et al. 1995). Therefore, ligninolytic activities of this Phanerochaetechrysosporium have mainly been attributed to lignin peroxidase and manganese peroxidase(KUHAD et al. 1997); its laccase has received more attention only recently (DITTMER et al.1997, RODRIGUEZ et al. 1997). Consequently, it has been suggested that two classes ofextracellular oxidative enzymes – peroxidases and laccases – are involved in ligninolysisowing to their ability to catalyze the cleavage of carbon-carbon or carbon-oxygen bonds incomplex lignin polymer or lignin model compounds (KIRK and HAMMEL 1992, HAMMELet al. 1993, MARZULLO et al. 1996).

Both extracellular peroxidases and laccases (also called ligninolytic or lignin-modifyingenzymes; HATAKKA 1994) have the ability to catalyze one-electron oxidations resulting inthe formation of radicals which undergo numerous spontaneous reactions. These, in turn,lead to various bond cleavages including aromatic ring fission (SHOEMAKER and LEISOLA1990, TUOR et al. 1992, AKTHAR et al. 1997, ZAPANTA and TIEN 1997). Manganese peroxi-dase attacks aromatic lignin structures indirectly via Mn3+-chelates acting as a low-molecular mass redox mediator while lignin peroxidase oxidizes aromatic lignin moietiesdirectly (WARIISHI et al. 1991, KUAN and TIEN 1993, ZAPANTA and TIEN 1997). Laccaseoxidizes preferentially phenolic lignin structures to phenoxy radicals which subsequentlyform quinones. This spontaneous rearrangement can also lead to the fission of carbon-carbon or carbon-oxygen bonds inside the lignin phenyl-propane subunits resulting either inthe degradation of both side chains (FUKUZUMI 1960) and aromatic rings (KAWAI et al.1988b), or in demethylation processes (HARKIN and OBST 1974, ANDER et al. 1983,LEONOWICZ et al. 1984, 1991a). In this respect, laccase can co-operate with various FAD-containing oxidases like glucose oxidase (SZKLARZ and LEONOWICZ 1986), veratryl alcoholoxidase (MARZULLO et al. 1995), cellobiose:quinone oxidoreductase (WESTERMARK andERIKSSON 1975), and cellobiose dehydrogenase (AYERS et al. 1978).

Although the actual role of laccase in lignin biodegradation is still under discussion andnot completely understood, several authors have reported that laccase acts on the ligninpolymer in the ways different from ligninolytic peroxidases. In order to present the role oflaccase more clearly several conceptions of biochemical routes in lignin biodegradationhave been reviewed, especially with respect to the co-operation of laccase with other bio-catalysts.

Hististorical outlook

The first attempts to delignify wood by means of white-rot fungi were made already in theearly 1930s. According to CAMPBELL (1930) and, independently, to WIERTELAK (1932),Trametes versicolor and Trametes pini respectively were shown to transform wood effi-ciently. Several years later, a more detailed analysis of the delignification process of beechsawdust by the white-rot fungi Polyporus abietinus, Stereum rugosum, and Marasmiusscorodonius was carried out (FAHRAEUS et al. 1949). However, due to the poor analyticalmethods at the time the analysis was extremely difficult. It was only the development of


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Fig. 1Hypothetical scheme of lignin degradation by Poria subacida (adapted from FUKUZUMI 1960)

novel chromatographic methods that helped to explain some stages of lignin biotransforma-tion by the characterization of certain degradation products. Thus, in the 1960s, the firstschemes of lignin degradation by white-rot fungi were published based on the data fromfungal decay of wood mill. Among others, a hypothetical mechanism was suggested byFUKUZUMI (1960), where laccase was presented as acting as a lignin-attacking enzyme(Fig. 1).

In this scheme, however, laccase appeared not at the beginning of but in the latter part ofthe degradation process. That is why the questions arose of how the lignin barrier could beinitially attacked and what the responsible attacking agent was. One of our first reportscovered the earliest attempt at explaining the initial microbial attack on lignin and postu-lated that it could indeed be catalyzed by fungal laccase (and also peroxidase) through thepreliminary demethylation of lignin. In addition, it was suggested that following the de-methylation, the break-down of ether bonds connecting lignin subunits probably occursspontaneously (non-enzymatically) and takes place as a result of the electronophilic effectof the ortho-quinones on the ether bonds between phenylpropane subunits.

The first indication of the reliability of this conception was to be found in the work ofHARKIN and OBST (1974). They showed that laccase and/or peroxidase generate a numberof reactive phenoxy radicals during the process of 2,4,6-trimethoxyphenol oxidation.


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Mesomeric forms of these phenoxy radicals couple with para or orto-quinal dimers whichsubsequently undergo demethylation and decomposition to monomers and are extremelyunstable, which triggers the polymerization effect (Fig. 2).

The same report also demonstrated the ability of laccase (and peroxidase) to cleave etherbonds linking most aromatic subunits in lignin, which pointed to a general significance ofdemethylation and quinone formation in lignin degradation. Subsequent research confirmedthe demethylating activity of laccase and showed the importance of pH on the conversionprocess (LEONOWICZ et al. 1984), e.g., by the use of syringic acid (ISHIHARA and ISHIHARA1976), vanillyl glycol (LUNDQUIST and KRISTERSSON 1985), and vanillic acid (ANDER et al.1983, ANDER and ERIKSSON 1985). The latter authors proposed a possible pathway of va-nillic acid metabolism via demethylation up to the cleavage of the aromatic ring and theformation of ketoacids (ANDER et al. 1985). Later, such an aromatic ring fission was, infact, demonstrated for Trametes (Coriolus) versicolor laccase (KAWAI et al. 1988b) (Fig. 3).The substrate (I) is oxidized by laccase to form phenoxy radicals, which are subsequentlyattached by molecular oxygen. The resulting hydroperoxide reacts with nucleophilic oxygenspecies forming substance (II) via incorporation with H2O (pathway B) or by forming acyclic peroxide (pathway A) which is then converted to a muconate derivative. Subse-quently, the muconate compound undergoes cyclization to yield lactone (II).

Fig. 2Transformation of dimers formed from 2,4,6-trimethoxyphenol during the action of laccase. Decom-position of dimers is accompanied by oxidation anddemethylation processes (adapted from HARKIN andOBST 1974)


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Furthermore, it was shown that a phenol oxidase-less (= laccase-less) mutant of Sporo-trichum pulverulentum obtained by UV-mutagenesis did not transform lignin while the wildtype decomposed the polymer effectively (ANDER and ERIKSSON 1976). This findingstrongly pointed to laccase as an important factor in lignin transformation and was laterconfirmed by the use of the regenerated protoplast technique to gain laccase-less mutants(EGUCHI et al. 1994).

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Fig. 4A possible pathway for the formation of carboxyl groups in lignin by Trametes (Coriolus) versicolorlaccase (adapted from KONISHI et al. 1974)


Significant contributions of laccase to lignin transformation were also reported by manyother authors: ISHIHARA and MIYAZAKI (1972), KONISHI et al. (1974; Fig. 4), ANDER andERIKSSON (1976), LEONOWICZ et al. (1985), KAWAI et al. (1986), SZKLARZ and LEONOWICZ(1986), KAWAI et al. (1988a; Fig. 5). KIRK et al. (1968) proposed a pathway leading toformation of 2,6-Dimethoxy-p-benzoquinone which was indeed found four years later byISHIHARA and MIYAZAKI (1972) as the result of the action of Trametes versicolor laccase onBJÖRKMAN’s lignin. In addition, we have shown in our laboratory that Trametes versicolorlaccase both depolymerizes and polymerizes polymeric lignosulphonates (adequate modelsof the macromolecular lignin structure), which, as soluble in water, are readily available forenzymes (LEONOWICZ et al. 1985; Fig. 6). In similar experiments, laccase also depolymer-ized – although not to a high degree – BJÖRKMAN’s lignin (LEONOWICZ et al. 1997b). Incontrast, GALLIANO et al. (1991), who used 14C-labelled KLASON lignin as the substrate forlaccase from Rigidosporus lignosus, found only slight solubilization of the polymer, proba-bly due to predominant repolymerization reactions.

KONISHI et al. (1974) put forward an interesting hypothesis that the formation of carboxylgroups in the side chains of lignin by laccase “solubilizes” the polymer. The dissolved car-boxylated products were proposed to adhere to the fungal cell-walls, where their conversionand subsequent uptake into the cells might be facilitated. However, this hypothesis – attrac-tive from the point of view of lignin utilization – could not be confirmed in subsequentstudies.

The above route demonstrates that the disproportion of the phenoxy radical of substrate I,formed by the action of laccase, occurs via a cation intermediate, which is further trans-formed to a diphenyl propanone III (pathway A) and to a hydroquinone/quinone (pathway

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Fig. 5Possible mechanism for Ca oxidation (A) and alkyl-aryl cleavage (B) of the lignin model dimer1-(3,5-dimethoxy-4-hydroxyhenyl)-2-(3,5-dimethoxy-4-ethoxyphenyl) propane-1,3-diol (I) by laccaseof Trametes (Coriolus) versicolor (adapted from KAWAI et al. 1988a)


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Fig. 6Depolymerization (A) and polymerization (B) of Na-lignosuphonate fraction with approximate mo-lecular weights of 97 (A) and 1 (B) kDa. Continuous line: elution profile on the Sephadex G-50 col-umn after treatment with laccase. Dashed line: elution profile of the control (adapted from LEONO-WICZ et al. 1985)

B). It was postulated that laccase is only responsible for the formation of the phenoxy radi-cal whereas both C� and alkyl-aryl cleavages are caused by spontaneous reactions. Thissupports our original hypothesis of the lignin degradation.

Last but not least, it should be mentioned that the so-called “yellow laccases” have beenrecently discovered in straw cultures of Panus tigrinus and Phlebia radiata. These enzymesseem to have a changed oxidation state of copper in the active center, probably caused bythe integration of aromatic lignin degradation products (LEONTIEVSKY et al. 1997a). Inter-estingly, evidence has been provided that yellow laccases are capable of oxidizing non-phenolic lignin models (�-O-4) and veratryl alcohol directly in the presence of O2, but inthe absence of any diffusible mediator (LEONTIEVSKY et al. 1997b).

Enzymes cooperating with laccase in ligninolysis

Extracellular phenoloxidizing activity which indicates the presence of ligninolytic enzymeswas discovered in white-rot fungi already in the 1930s (ERIKSSON et al. 1990). With regardto their typical production patterns of extracellular ligninolytic enzymes, white-rot fungimay be divided into three main groups; (i) LiP-MnP group, (ii) MnP-laccase group, and (iii)LiP-laccase group (HATAKKA 1994). However, several enzymes possessing other than di-rectly lignin-degrading activities, such as H2O2 production, are indispensable in ligninolysisand therefore also reviewed here.

Lignin peroxidase (LIP; EC 1.11.1.14): Lignin peroxidase (at first designated as “ligni-nase”) was discovered in cultures of Phanerochaete chrysosporium after many years ofresearch. The glycosylated enzyme was identified and described independently by twoAmerican teams (GLENN et al. 1983, TIEN and KIRK 1983) and a Japanese group (SHIMADAand HIGUCHI 1983) in supernatants of nitrogen and carbon-limited cultures of the corticoidbasidiomycete Phanerochaete chrysosporium that were flushed with pure oxygen. Later,the enzyme was also found in other wood colonizing, corticoid and polyporous basidiomy-


cetes, e.g., Phlebia radiata, Phlebia tremellosa, Trametes versicolor (HATAKKA et al. 1987,LEONOWICZ et al. 1991b, HATAKKA 1994). The only currently known agaric fungi produc-ing lignin peroxidase are Nematoloma frowardii and Clitocybula dusenii (HOFRICHTER andFRITSCHE 1997).

Lignin peroxidase (molecular mass varies between 38 and 47 kDa) necessarily requireshydrogen peroxide (H2O2) generated by other enzymes (e.g., oxidases) to be active. Theenzyme comprises hem in the active site while the catalytic cycle resembles that of horse-radish peroxidase (TIEN et al. 1986). Lignin peroxidase catalyses several oxidations in thealkyl side chains of lignin-related compounds, C-C cleavages in the side chains of ligninsubunits, oxidation of veratryl alcohols and related substances to aldehydes or ketones,intradiol cleavage of phenylglycol structures, and hydroxylation of benzylic methylenegroups (TIEN and KIRK 1983). In Phanerochaete chrysosporium, the enzyme is produced inat least six isozymes (TIEN and KIRK 1988), which are all capable of oxidizing recalcitrantnon-phenolic lignin model substrates (redox potential around 1.5 V) by one-electron ab-straction to form reactive aryl cation radicals, which commonly decay via pathways in-volving C–C and C–O bond-cleaving reactions (KERSTEN et al. 1985).

Reactions of lignin peroxidase have been thought to be mediated by veratryl alcohol, alow-molecular-mass fungal metabolite which is oxidized by the enzyme to veratraldehydethrough an aryl cation radical (KIRK and FARRELL 1987). The latter is highly reactive andshould, in turn, oxidize the lignin polymer. However, this role of veratryl alcohol has beenquestioned since the stability of the radical (its life span) is too short to enable long-distancecharge-transfer. If veratryl alcohol radicals possess mediating properties, this will only berelevant in short distance transfers when the aryl cation radical is somehow complexed tothe enzyme (LUNDELL 1993). Nevertheless, veratryl alcohol stimulates lignin peroxidaseactivity, probably by protecting the sensitive enzyme from the damaging effects of excessH2O2 or phenolics (AKTHAR et al. 1997).

Lignin peroxidase is also capable of cleaving aromatic rings via initial one-electron ab-stractions and subsequent incorporation of oxygen (UMEZAWA and HIGUCHI 1987). Theoptimum activity of lignin peroxidase is extremely low (between pH 2.5 and 3.0; TIEN andKIRK 1988) while the enzyme itself is acidic (pIs 3-5). TIEN and KIRK (1983) have reportedthat in a cell-free system (in vitro) the enzyme is particularly active toward highly-methylated lignins (e.g., lignin isolated from birch wood and additionally methylated withmethyl iodide), which are depolymerized into lower-molecular mass fragments. In contrast,natural lignins containing phenolic rings tend to re-polymerize in vitro in the presence oflignin peroxidase (SARKANEN et al. 1991).

Two years later, it was shown (using 14C-labelled synthetic lignin preparations) that puri-fied Phanerochaete chrysosporium lignin peroxidase is also capable of depolymerizinglignin, which contains phenolic rings, if specific reactions conditions are created: the addi-tion of only low amounts of lignin dissolved in methoxyethanol (around 5 mg/l) and thepresence of veratryl alcohol. Under these conditions, lignin peroxidase is capable of oxi-

Fig. 7Lignin peroxidase catalyzed Ca-Cb cleavage in 14C-labelled synthetic lignin to give a benzoic alde-hyde at the Ca-position (adapted from HAMMEL et al. 1993)


dizing the lignin polymer directly through cleaving Ca–b bonds yielding benzoic aldehydederivatives of b-O-4 substructures (HAMMEL et al. 1993; Fig. 7).

Since lignin, degraded by lignin peroxidase, provides a substrates for laccase, both en-zymes can be considered as “partners” in certain biotransformation routes of lignin.

Manganese peroxidase (MnP; EC 1.11.1.13): Manganese peroxidase was also discoveredin cultures of Phanerochaete chrysosporium (KUWAHARA et al. 1984). The enzyme resem-bles lignin peroxidase: it is extracellular, glycosylated and contains hem as the prostheticgroup (GLENN and GOLD 1985, PASZCZYNSKI et al. 1986). Manganese peroxidase is alsoexpressed in multiple forms with molecular weights from 40 to 48 kDa and pIs between 2.9and 7.0. Besides Phanerochaete chrysosporium, manganese peroxidase has also been foundin many other white-rot and soil litter-decomposing basidiomycetes, but not in any otherfungi or bacteria (e.g., Trametes versicolor, JOHANSSON and NYMAN 1987; Phlebia radiata,NIKU-PAAVOLA et al. 1988; Ceriporiopsis subvermispora, RÜTTIMANN et al. 1992; Agari-cus bisporus, BONNEN et al. 1994; Nematoloma frowardii, SCHNEEGASS et al. 1997; Stro-pharia rugosoannulata, SCHEIBNER and HOFRICHTER 1998).

The catalytic cycle of manganese peroxidase resembles that of lignin peroxidase, includ-ing native ferric enzyme as well as peroxidase Compound I and Compound II redox states(WARIISHI et al. 1988). However, significant differences appear in the reductive reactionswhere Mn2+ is a required electron donor. Both Compound I and Compound II are reducedby Mn2+ while the latter is oxidized to Mn3+. Mn3+ ions are stabilized to high redox poten-tials via chelation with organic acids such as oxalate, malonate, malate, tartrate, or lactate(WARIISHI et al. 1992, Fig. 8). Chelated Mn3+, in turn, acts as diffusible redox mediator thatoxidizes phenolic lignin structures (WARIISHI et al. 1992).

The oxidative strength of the manganese peroxidase system can be considerably en-hanced in the presence of appropriate additional cooxidants such as thiols (FORRESTER et al.1988, HOFRICHTER et al. 1998c) or lipids (BAO et al. 1994), which results in the cleavedchemical bonds. These bonds are not normally open to attack by manganese peroxidase(e.g., recalcitrant non-phenolic �-O-4 arylethers). Moreover, manganese peroxidase isunique in its ability to degrade aromatic substances, including lignin, directly to carbondioxide (CO2); this process (enzymatic combustion), which is considerably stimulated in thepresence of the above mentioned cooxidants, probably allows ligninolytic fungi to miner-alize lignin - at least in part - outside the cell (HOFRICHTER et al. 1998a, KAPICH et al.1999). The enzyme has been successfully applied to depolymerize 14C-labelled syntheticand natural lignins in vitro (WARIISHI et al. 1991); as a result, low-molecular mass, water-soluble lignin fragments and significant amounts of CO2 have formed (HOFRICHTER et al.1998a, 1998b, 1998c, 1998d, 1998e).

H O2 2

H O2 2

MnP MnP IMnP I

MnP II MnP IIMnP II

k3

k3

k2

k2

Kd

Kd

k1

k1

OMn3 +OMn3 +

OMn3 +OMn3 +

OMn2 +OMn2 +

OMn2 +OMn2 +

OMn2 +OMn2 +

OMn2 +OMn2 +

Fig. 8Mechanism of manganese peroxidasecatalyzed formation of Mn3+-oxalatecomplexes. Mn-oxalate complexes arerepresented by OMn2+ and OMn3+

(adapted from ZAPANTA and TIEN 1997)


Manganese peroxidase has been shown to generate H2O2 in the oxidation of certain thiols(e.g., glutathione) and NAD(P)H2; recently, evidence has been provided that the enzyme iseven capable of acting efficiently in the absence of external H2O2 by oxidizing organic acids(e.g., oxalate, malonate, malate) in “oxidase-like,” autocatalytic reactions involving thetransient formation of several radical species (HOFRICHTER et al. 1998e, URZÚA et al.1998).�� '��

2�� 5��(�� )��,"6�� 2�� 2�� )�� 2�� '�(��+GALLIANO �� ,--,%/� 7�� (�� '�� 8��/

In recent years, an extended role of manganese peroxidase in the lignin degradation hasbeen proposed following the discovery of novel hybrid forms of the enzyme and other per-oxidases in some fungi, e.g., in Pleurotus eryngii and Bjerkandera spp. (MARTINEZ et al.1996, MESTER and FIELD 1998). These enzymes are able to oxidize both Mn2+ and aromaticsubstrates (e.g., veratryl alcohol, various phenolics), thus combining the properties of man-ganese peroxidase and lignin peroxidase.

Other peroxidases (EC 1.11.1.7): A “classic” extracellular peroxidase (similar to horse-radish peroxidase) was identified for the first time in a growing culture of the agaric white-rot fungus Pholiota mutabilis by LEONOWICZ and TROJANOWSKI (1965). Together withlaccase, this peroxidase caused the demethylation of vanillic acid and BJORKMAN's lignin.The enzyme was later purified from culture liquids of two other white-rot fungi, Inonotusradiatus and Trametes versicolor, by LOBARZEWSKI (1981). Its constitution (e.g., molecularweight, hem in the catalytic center, H2O2 requirement) and other properties were similar tothose of plant horseradish peroxidase. Interestingly, the enzyme did not show any effect onveratryl alcohol. Apart from its existing repolymerizing activities, this type of peroxidasefrom Trametes versicolor depolymerized high-molecular weight lignin (water-soluble Na-lignosulphonates) at pH 3 but not to a large extent (Fig. 9).

0.5

1.0

200 200 200600 600 600 ml

A B C

Ab

so

rb

an

ce

at

28

0n

m

Fig. 9Effects of immobilized Trametes versicolor peroxidase on Na-lignosulphonates. The continuous lineindicates the elution profile of lignosulphonates after treatment with peroxidase (A, B and C refer todifferent incubation periods; 3, 12 and 24 hrs respectively). The dashed line indicates the elutionprofile of the control. Radicals and quinones generated by peroxidase (as in the case of laccase) mayreadily undergo polymerization and thus, the depolymerization effect is less clear (adapted fromLOBARZEWSKI et al. 1982)


A similar type of peroxidase was discovered by de JONG et al. (1992) in Bjerkandera sp.cultured in a nutrient- and nitrogen-rich glucose/yeast extract/peptone medium. Accordingto MESTER et al. (1996), this enzyme – described as manganese independent peroxidase –was inhibited by Mn2+ ions. The enzyme did not oxidize veratryl alcohol at all; however, itoxidized various phenolic substrates, e.g., 2,6-dimethoxyphenol that was also used for theenzyme assay. The essential cosubstrate of this enzyme was hydrogen peroxide and thecatalytic center was found to contain hem as the prosthetic group (KOTTERMAN et al. 1994).Extracellular peroxidases with properties similar to those of Trametes versicolor and Bjer-kandera sp. were also found in a range of other basidiomycetous fungi, e.g., Coprinus spp.(MORITA et al. 1988), Junghuhnia separabilima (VARES et al. 1992) and Panaeolus spp.(HEINZKILL et al. 1998).

�� Dioxygenases, widely distributed amongbacteria and microfungi, have also been identified in ligninolytic basidiomycetes.Protocatechuate 3,4-dioxygenase activity was first observed in Trametes (Polystictus)versicolor by HAIDER et al. (1962) and may serve as an example. In order to examine thecharacteristic steps of dioxygenase catalyzed reactions, radioactively-labelled 14C-vanillicacid was used as test substrate.

Before the discovery of ligninolytic peroxidases, scientists of KIRK’s team (CHEN et al.1983) formulated the hypothesis that among many other reactions occurring during thedecay of lignin by white-rotters, dioxygenase-catalyzed reactions inside the intact polymercould take place. As a result, the peripheric aromatic rings of lignin could be cleaved andaliphatic carboxylic acids formed as the products (Fig. 10).

Protocatechuate 3,4-dioxygenase was also later isolated from the cellulolytic ascomyceteChaetomium piluliferum and the ligninolytic basidiomycete Pleurotus ostreatus, and puri-fied to homogeneity in our laboratory (WOJTAS-WASILEWSKA and TROJANOWSKI 1980,WOJTAS-WASILEWSKA et al. 1983). The effects of this enzyme, including its immobilized

C C

C

C

C

OCH3

OH

COOHCOOH

COOHCOOH

COOHCOOH

OH

CHOCHO

COOHCOOH

COOHCOOH

OCH3

OCH3

OCH3

OCH3

OCH3

O O

O

O

O

O

29

13

O

OH

O

O

CH C

C

C

C

CHOH C

C

C

C

OH

OH

OH

OH

H O2

CH OH2

CH2

COOHCOOH

CHOCHO

CHOCHO

CHOCHO

CHOCHO

HO

L L

L

L

L

Fig. 10Cleavage of a peripheral ring of lignin by Phanerochaete chrysosporium growing on spruce woodchips (adapted from CHEN et al. 1983)


form on sodium lignosulphonates, were examined (WOJTAS-WASILEWSKA and LUTEREK1987, WOJTAS-WASILEWSKA et al. 1987). As the result, a product was obtained whichshowed a significant decrease in the absorbency around 280 nm indicating a loss of aro-matic rings. This may indicate that some peripheric aromatic rings of the tested lignosul-phonates were cleaved by 3,4-protocatechuate dioxygenase.

Other intracellular, ring-cleaving dioxygenases (1,2,4-trihydroxybenzene-1,2 dioxy-genase, catechol-1,2 dioxygenase) were detected in Phanerochaete chrysosporium andPhlebia radiata (RIEBLE et al. 1994, ROGALSKI et al. 1996).

Glucose 1-oxidase (GOD; EC 1.1.3.4): Glucose oxidase is a FAD-dependent oxidase thatcan be found inside and outside fungal cells. During the enzymatic catalysis, glucose – forexample, one originating from cellulose – is oxidized first to the corresponding glucono-lactone with the simultaneous reduction of FAD to FADH2. In the second step, FADH2 isoxidized back to FAD by dioxygen (O2) while hydrogen peroxide is formed (Alberti andKlibanov 1982).

�-D-Glucose + E-FAD � D-Glucono-1,5-lactone + E-FADH2 (1)

E-FADH2 + O2 � E-FAD + H2O2 (2)

Intensive screening studies in our laboratory have shown that glucose oxidase occurs inmany white-rot fungi which attack lignin effectively and secrete high amounts of laccase(LEONOWICZ et al. 1986, 1999d). RAMASAMY et al. (1985) have reported that glucose oxi-dase negative mutants of Phanerochaete chrysosporium exhibited little or no ability todegrade lignin whereas the wild type was able to do so. The same group has, for the firsttime, demonstrated that Phanerochaete chrysosporium also produces laccase. The follow-ing part of this review shows that glucose oxidase co-operates effectively with laccase inlignin degradation; thus, the results of RAMASAMY et al. alone may point to glucose oxidaseas an agent involved in lignin degradation (SRINIVASAN et al. 1995).

Another related sugar oxidizing, intracellular enzyme with FAD as the prosthetic group –pyranose 2-oxidase (EC 1.1.3.10) – was found in the growing mycelium of Phanerochaetechrysosporium. The ultrastructural and immunocytochemical characterization of this en-zyme was reported by DANIEL et al. (1992).

Cellobiose : quinone oxidoreductase (CBQ; EC 1.1.5.1): In 1974, when Trametes (Poly-porus) versicolor was cultured on lignin agar plates supplemented with glucose, cellobioseor cellulose, WESTERMARK and ERIKSSON (1974a) found indications of the existence of anew quinone-reducing enzyme. This enzyme, later designated as cellobiose : quinone oxi-doreductase possessed the ability to stop phenolic coupling during the laccase oxidation ofquaiacol. Further experiments also demonstrated the production of this enzyme by Chryso-sporium lignorum (= Sporotrichum pulverulentum or Phanerochaete chrysosporium;WESTERMARK and ERIKSSON 1974b) from which the enzyme was purified and character-ized as a flavoprotein with FAD as the prosthetic group. The molecular weight of the ho-mogeneous enzyme was 58 kDa (WESTERMARK and ERIKSSON 1975).

Except in Phanerochaete chrysosporium, cellobiose-quinone oxido-reductase was alsofound in 25 other white-rot fungi (ANDER and ERIKSSON 1977). The enzyme reduces a largenumber of quinones produced, for example, by laccase to the corresponding phenols whilecellobiose is simultaneously oxidized to cellobionolactone (WESTERMARK and ERIKSSON1974b). During this reaction hydrogen peroxide is gradually formed from oxygen (MORPETHand JONES 1986).


Cellobiose dehydrogenase (CDH; EC 1.1.99.18 – earlier called cellobiose oxidase): Theenzyme, for the first time isolated and purified from Sporotrichum pulverulentum byERIKSSON’s team, is a monomeric protein with a molecular mass around 90 kDa (AYERSet al. 1978). It is also a flavoprotein, but with cytochrome-b type heme in the catalytic cen-tre. This group probably transfers two electrons from the electron donor (e.g., cellobiose) toone electron acceptors, e.g., cytochrome-c, Fe3+ or dioxygen. Similarly to cellobi-ose:quinone oxidoreductase, cellobiose dehydrogenase reduces a range of quinones (e.g.,2,6-dichlorophenoloindophenol; SAMEJIMA and ERIKSSON 1991) and oxidizes cellobioseand related oligosaccharides. However, oxygen in this reaction (different from that in thecase of the cellobiose:quinone oxidoreductase), is probably not incorporated into the cello-biose, but reduced to superoxide and/or hydrogen peroxide (ANDER 1994, HENRIKSSON1995). Moreover, cellobiose dehydrogenase also oxidizes the reducing end groups of cello-dextrins and cellulose (AYERS et al. 1978, HABU et al. 1993). Similarities in basic activitiesof CBQ and CDH underlie the assumption that cellobiose : quinone oxidoreductase isprobably a product of the proteolytic cleavage of cellobiose dehydrogenase (HENRIKSSONet al. 1991, WOOD and WOOD 1992, HABU et al. 1993). Apart from the differences in themolecular masses, the enzymes can be distinguished by some differences in their specific-ity; for example, only cellobiose dehydrogenase utilizes cytochrome c as an electron ac-ceptor (SAMEJIMA and ERIKSSON 1991). Recently, it has been unequivocally shown thatboth enzymes reduce not only quinones but also phenoxyl and aryl cation radicals producedfrom aromatic substrates by laccase and peroxidases (ANDER et al. 1990, HENRIKSSON et al.1993, ANDER 1994, ANDER and MARZULLO 1997).

Aryl alcohol oxidases (AAOs; EC 1.1.3.7): Aryl alcohol oxidase is an extracellular, FAD-dependent enzyme, which was discovered in Trametes (Polystictus) versicolor by FARMERet al. (1960). Later the enzyme was also found in Phanerochaete chrysosporium (ASADAet al. 1995) as well as in the culture liquids of other white-rot fungi, e.g., Pleurotus sajor-caju (BOURBONNAIS and PAICE 1988, 1989), Pleurotus eryngii (GUILLÉN et al. 1990), Pleu-rotus ostreatus (SANNIA et al. 1991, PALMIERI et al. 1993) and Bjerkandera adusta(MUHEIM et al. 1990a, 1990b). Aryl alcohol oxidase from Pleurotus eryngii is a glycopro-tein with a molecular weight of 72.6 kDa which oxidizes a number of aromatic alcohols(aryl a- and a-b-unsaturated, g-aromatic alcohols) to the corresponding aldehydes whileoxygen is reduced to hydrogen peroxide (Fig. 11). The optimum pH of the reaction is abovepH 6 (MARZULLO et al., 1995) – this is different from lignin peroxidase which alsoproduces veratraldehyde from veratryl alcohol, but only at pHs below 4 (TIEN and KIRK1988).

Aryl alcohol dehydrogenase (AAD; EC 1.1.91): The NADPH-dependent aryl alcoholdehydrogenase was found as an intracellular enzyme in the white-rot fungus Pleurotuseryngii by GUILLEN and EVANS (1994). The enzyme, reducing aromatic aldehydes intoalcohols, supports the ligninolytic system by providing a suitable amount of aryl alcoholsacting as source of hydrogen peroxide (see above). It has also been suggested that the en-zyme is important for the stability of lignin peroxidase or mediators.

R1 R1R3 R3

R2 R2

CH OH2

CH OH2

CHOO

2O

2

AAO

H O2 2

H O2 2

Fig. 11Mechanism of aromatic alcohol oxida-tion by aryl alcohol oxidases. R1–R3represent various groups characterizingparticular alcohols. In the case of veratrylalcohol, R1 = R2 = OCH3, R3 = H.(adapted from de JONG et al. 1994)


Glyoxal oxidase (GLO; EC 1.2.3.5): Glyoxal oxidase is a further FAD-dependent, H2O2-generating enzyme discovered in growing cultures of Phanerochaete chrysosporium(KERSTEN and KIRK 1987). Later, its formation was also demonstrated for Phlebia radiata(VARES et al. 1995). Enzyme purification and characterization revealed that among others itoxidizes glyoxylic acid into oxalic acid while generating hydrogen peroxide. In this way,glyoxal oxidase supports peroxidase activities and supplies chelators for Mn3+ ions(KERSTEN et al. 1990).

Superoxide dismutases (SODs; EC 1.15.1.1): Superoxide dismutase (SODs) is an almostuniversal enzyme of aerobic micro-organisms (HASSAN 1989). It catalyzes the dismutationof the superoxide anion radical to molecular oxygen and hydrogen peroxide in all oxygenmetabolizing organisms and even in certain anaerobes (MCCORT et al. 1971). In thisway, SOD protects organisms from toxic effects of the reactive superoxide radical (O2

. –

= SOR).All currently known SODs are metal proteins containing iron (FeSOD), or manganese

(MnSOD), or copper plus zinc (Cu-ZnSOD) or nickel (NiSOD) as the prosthetic groups.FeSOD commonly occurs in prokaryotic organisms, e.g., Escherichia coli (YOST andFRIDOVICH 1973), but also in various fungi. The crystalline form of FeSOD was obtainedfrom Sulfolobus acidocaldarius by KARDINAHL et al. (1996). MALARCZYK et al. (1995)identified the FeSOD in the white-rot fungus Pleurotus sajor-caju and also in some otherPleurotus spp. Interestingly, the enzyme was produced in increased amounts in fungal fruitbodies and mycelia shortly after the appearance of laccase and peroxidases. MnSODs werepurified to homogeneity from the white-rot fungi Ganoderma microsporium (PAN et al.1997) and Phanerochaete chrysosporium (OZTURK et al. 1999). Recently a Cu-ZnSOD wasidentified in the laccase-producing mould Botrytis cinerea (CHOI et al. 1997). A novelNiSOD dismutase was isolated in 1996 from Streptomyces species (YOUN et al. 1996). Itwas homotetramer protein with little sequence similarity to known SODs (KIM et al. 1996).

The toxic and reactive SOR commonly appears during the quinone redox cycles, e.g., inthe laccase-producing, white-rot fungus Pleurotus eryngii (GUILLEN et al. 1997). During thecycle of this fungus, the cell-bound divalent reduction of quinones (Q) to hydroquinones(Q2–) is followed by the extracellular laccase-mediated oxidation of hydroquinones intosemiquinones (Q. –), which, in turn, are autoxidized to quinones. In both quinone oxidationphases production of SOR occurs:

Q2– + O2 � Q. – + O2. – (3)

Q. – + O2 � Q + O2. – (4)

Subsequently, SOR reacts as a reducing or oxidizing agent with other radicals produced byligninolytic enzymes, thus contributing to various lignin transformation processes, e.g.,aromatic ring fission (KAWAI et al. 1988b) or demethylation (POTTHAST et al. 1995). As anoxidant, SOR is able to generate highly reactive Mn3+ from Mn2+ and hydrogen peroxide:

O2. – + Mn2+ + 2H+ � Mn3+ + H2O2; (5)

as a reducing agent, SOR reacts with Fe3+ reducing it to Fe2+ yielding oxygen:

O2. – + Fe3+ � O2 + Fe2+; (6)

then, the reduced iron and hydrogen peroxide react with each other to form extremely reac-tive hydroxyl radicals via the FENTON reaction (FENTON 1894, WAILING 1975, GOLDSTEINet al. 1993, PRATAP and LEMLEY 1998):

Fe2+ + H2O2 � Fe3+ + HO. + OH– ; (7)


hydroxyl radicals are also produced from SOR and hydrogen peroxide in the Haber-Weissreaction (HABER and WEISS 1934):

O2. – + H2O2 � HO. + OH– + O2 (8)

As a one-electron reduction product of oxygen, SOR is not strong enough for ligninoly-sis, but produced with a share of oxygen, hydrogen peroxide or hydroxyl radicals, it mightbe an important agent in lignin transformation (FORNEY et al. 1982, GUILLÉN et al. 1997,PARK et al. 1997).

On the other hand, the highly reactive hydroxyl radicals can interact with DNA, proteins,lipids, or other biomolecules resulting in damage to the cells. To prevent the formation ofhydroxyl radicals from SOR, SOD is produced, which catalyzes its rapid dismutation to O2

and H2O2 (GUILLÉN et al. 1997, MCCORMICK et al. 1998) as follows:

2 O2. – + 2H.

� H2O2 + O2 . (9)

Hydrogen peroxide yielded here is either transformed to O2 and H2O by catalase or in-volved in peroxidative reactions. Consequently, the living cells (also of basidiomycetousfungi) commonly possess SOD, catalase and peroxidases, which are able to eliminate harm-ful oxygen species. According to our results, the maximum activity of SOD in lignin de-grading fungi appear when the activities of ligninolytic enzymes diminish indicating aregulative function of this enzyme in ligninolysis (Fig. 12).

Proteolytic activities in the white-rot Basidiomycete fungi: In the wood-decaying basidi-omycetes known for ligninolytic activities, proteolytic enzymes have been occasionallydetected and isolated. As the majority of these enzymes is extracellular (ERIKSSON andPETTERSSON 1982, STASZCZAK et al. 1996 and 2000), they might be of general importancewith respect to their effects on the lignin- or cellulolytic enzymes secreted by the same

25

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0M P1 P2 P3 F1 F2 F3c F3s Ma

Stages of development

En

zym

eact

ivit

ies

Legend

peroxidase Ulaccase (nkatal/l)

SOD (U)phenol.oxid.(A315nm)

Fig. 12Activity of SOD and of ligninolytic enzymes in Pleurotus sajor-caju during growth in the stages ofprimordia and fruiting bodies. P – primordia; 1,2,3 – stages from small to big forms, F – fruit bodies;c – cap; s – stem of fruit body (adapted from MALARCZYK et al. 1995)


fungi. Such a proteolytic enzyme, namely carboxyl proteinase (EC 3.4.23.6), was firstfound in Pycnoporus coccineus and its substrate specificity was determined (ICHISHIMAet al. 1980). Three years later, the same authors (ICHISHIMA et al. 1983) found another pro-teolytic enzyme – acidic carboxypeptidase (EC 3.4.16.1) – in the same fungal genus, but ina different species (Pycnoporus sanguineus). STASZCZAK and NOWAK (1984) pointed to themultiple, extra-and intracellular proteolytic activities of the white-rot fungus Trametes ver-sicolor. Purification and partial characterization of two acid proteases from Sporotrichumpulverulentum grown on cellulose were described by ERIKSSON and PETERSON (1982,1988). The authors stated that the enzymes play a regulatory role in the activation ofthe endo-1,4-�-glucanases. The regulatory effect of proteases on lignin peroxidase and�-amylase was also proposed (DOSORETZ et al. 1990, DEY et al. 1991). HABU et al. (1993)demonstrated the release of a FAD domain from cellobiose oxidase (CBO) and the conver-sion of CBO to cellobiose:quinone oxidoreductase (CBQ) by proteases from cellulolyticcultures of Phanerochaete chrysosporium. The decrease in MnP activity due to the actionof proteases was observed in Phanerochaete chrysosporium (CRUZ-CORDOVA et al. 1999).The same report showed that LiP was less sensitive to proteolytic activities than MnP. Inaddition, reports from our laboratory (STASZCZAK et al. 1996 and 2000, STASZCZAK andGRZYWNOWICZ 1997) suggest both inhibiting and activating functions of Trametes versi-color serine proteinases in the regulation of laccase activity.

The growth of wood decaying fungi, especially under natural conditions, requires controlof their nitrogen economy, which involves the regulation of proteolytic activities for theintracellular protein turnover, the extracellular digestion of protein sources, and the modifi-cation of proteins through limited proteolysis. Lignin modifying enzymes are mainly ex-pressed during the secondary metabolic phase of the growth and be accompanied by in-creased protein turnover. Moreover, ligninolytic enzymes acting outside mycelium aresusceptible to the fungus’ extracellular proteinases; this is why their activity is of signifi-cance when the process of ligninolysis is considered.

Low-molecular mass mediators

According to several publications, the molecular size of wood-attacking enzymes does notpermit penetration of the undegraded plant cell wall (EVANS et al. 1991, 1994, FLUORNOYet al. 1991). On the other hand, there have been studies presenting evidence that some ofthese enzymes (e.g., laccase) are able to bleach hard wood pulp by depolymerizing andsolubilizing lignin in the presence of so-called mediator compounds (BOURBONNAIS andPAICE 1992, BOURBONNAIS et al. 1995, CALL 1994, 1996, CALL and MUCKE 1997,MAJCHERCZYK et al. 1999). Therefore, in recent years research has been focused on suchpotential low-molecular mass mediators of internal or external origin, which possess highenough redox potentials (>900 mV) to attack lignin and can migrate from the enzymes intothe tight lignocellulose complex. Consequently, many low-molecular mass compounds havebeen suggested to permeate wood cell walls and initiate decay. Examples of such sub-stances include veratryl alcohol, oxalate, 3-hydroxyanthranilic acid, and Gt-chelators. Theyare produced as a result of fungal metabolism and their secretion enables fungi to colonizeand degrade the wood cell wall structure more effectively than other organisms. The syn-thesis of veratryl alcohol was first observed in Phanerochaete chrysosporium (LUNDQUISTand KIRK 1978); oxalate and other organic acids were already found in culture liquids of anumber of wood-rotting fungi in 1965 (e.g., Armillaria mellea, Fomes annosus, Pleurotusostreatus, (TAKAO 1965). Later, their secretion was also shown for solid-state cultures(GALKIN et al. 1998, HOFRICHTER et al. 1999). 3-Hydroxyanthranilic acid was isolated andidentified from Pycnoporus cinnabarinus (EGGERT et al. 1996 and 1997), and a specialphenolate derivative, the so-called Gt-chelator (molecular mass <1 kDa), has been isolatedfrom the brown-rot fungus Gleophyllum trabeum (GOODELL et al. 1996).


Veratryl alcohol as well as oxalate and other dicarboxylic acids are fungal metabolites ofmany white-rot fungi: the first one is involved in the lignin peroxidase catalysis as a pro-tecting substance and may be involved in short-distance electron transfers while dicarbox-ylic acids stabilize highly reactive Mn3+ generated by manganese peroxidase. Additionally,the Mn3+-chelates can react with each other resulting in the formation of CO2 and carbon-centred radicals which, in turn, can add dioxygen to form peroxyl radicals (KEREM andHADAR 1997, HOFRICHTER et al. 1998e). The latter have recently been proposed to be deci-sive oxidants in lignin degradation (HAMMEL et al. 1999). Also, oxalate chelates other ca-tions such as Ca2+, Fe2+ and NH4

+ (DUTTON et al. 1993). The removal of calcium wouldenlarge the cell wall pore size, which may permit access of enzyme molecules (GREEN et al.1991). Moreover, oxalate regulates the concentration of ferric ions for the FENTON’s reac-tion (FENTON 1894, KOENIGS 1972). This reaction supplies the degradation environmentwith highly reactive hydronium ions (H3O+) and hydroxyl radicals (HO. and HO–) whichmay initiate lignocellulose degradation by certain fungi.

Four groups of benzene derivatives were identified as the components of the Gt-chelatorfrom Gleophyllum trabeum (Fig. 13). The presence of the chelator in this brown-rot fungusallows for the reduction of iron, the generation of oxygen radicals, and the degradation ofcellulosic and phenolic compounds (GOODELL et al. 1996).

The first natural mediator of laccase – 3-hydroxyanthranilic acid (3-HAA) – supportingligninolysis in Pycnoporus cinnabarinus, a white-rot fungus that does not produce manga-nese and lignin peroxidase, was discovered by EGGERT et al. (1996). The laccase/3-HAA

OCH3OCH3

OCH3OCH3

OCH3OCH3

OCH3OCH3 OCH3OCH3

OCH3OCH3 OCH3OCH3

OCH3OCH3

OCH3OCH3 OCH3OCH3 OCH3OCH3 OHOH O

OHOH O

O

O

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HO

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H CO3H CO3

H C3H C3 H C3H C3

H CO3H CO3

H CO3H CO3 H CO3H CO3

H CO3H CO3

H CO3H CO3

H CO3H CO3

HydroxyphenylaceticHydroxyphenylacetic acid derivatives

Hydroxybenzoic acidacid derivatives

Hydroxybenzene derivativesderivatives

Dihydroxyphenylpentane - 1,4 - diolDihydroxyphenylpentane - 1,4 - diol

Fig. 13Structures of the selected phenol-ic compounds identified in theGt-chelator mixture of Gleophyl-lum trabeum (adapted fromGOODELL et al. 1996)


system mediates the oxidation of non-phenolic lignin model dimers whereas laccase alonecannot act on these compounds.

The dimeric ethers possessing phenolic groups are neither oxidized nor cleaved by thelaccase/3-HAA system. The laccase/3-HAA activity on the non-phenolic dimer, apart fromthe production of veratric acid and guaiacol, also results in a 6-electron oxidation of 3-HAA.This leads to the formation of a phenoxazinone ring containing compound from 3-HAA i.e.cinnabarinic acid (CA) which is a common secondary metabolite of Pycnoporus cinnabari-nus (EGGERT et al. 1995, 1996). However, TEMP and EGGERT (1999) have also demon-strated that in the presence of a suitable cellulose-derived electron donor, cellulose dehydro-genase (CDH) can regenerate the fungal mediator 3-HAA from cinnabarinic acid (Fig. 14).

Several years ago the delignification of kraft pulp by laccase was found to be supportedby a number of synthetic, low-molecular mass dyes or other aromatic hydrogen donors. Thefirst mediator that was shown to be effective in the delignification of kraft pulp and lignintransformation by laccase was ABTS [2,2�azinobis-(3-ethylbenzenthiazoline-6-sulfonicacid)] (BOURBONNAIS and PAICE 1992, 1996, BOURBONNAIS et al. 1997a).

The reaction mechanism mediated by ABTS seems to proceed as follows: oxygen acti-vates laccase and the mediator is oxidized by the enzyme. The oxidized mediator diffusesinto the pulp and oxidizes lignin disrupting it into smaller fragments which are easily re-moved from the pulp through the alkaline extraction. Superoxide dismutase (SOD) inhibitspolymerization of soluble lignin by laccase and accelerates the delignification process(adapted from SEALEY et al. 1997).

MAJCHERCZYK et al. (1999) demonstrated the oxidation of aromatic alcohols, used as non-phenolic lignin models, by laccase of Trametes versicolor in the presence of ABTS. Cationradical and dication formation were proposed to be involved in this laccase mediator system.

Apart from ABTS, 1-hydroxybenzotriazole (HBT) was recently introduced as an effec-tive laccase mediator in pulp processing (CALL 1994). An R-NO. radical is the active formof this mediator; it is also selective for lignin oxidation. The laccase-HBT couple can causeup to 50% of pulp delignification (SEALEY et al. 1997).

HBT is not less effective in the bleaching processes than ABTS. The discovery of HBTintroduced a new class of mediators with N-OH as the functional group, which is oxidizedto a reactive radical. Other mediators of this type, e.g., 4-hydroxy-3-nitroso-1-naphtalenesulfonic acid (HNNS), 1-nitroso-2-naphthol-3,6-disulfonic acid (NNDS), andRemazol brilliant blue (RBB), have also been shown to promote delignification, but theyare not as efficient as HBT and ABTS (BOURBONNAIS et al. 1997b) (Table 1).

2H O2

laccasered

O2

laccaseox

3-HAA

CDHred

CDHOX

cellobiose

induction

cellobiose

cellulose

cellobiono-

-lactone�

cinnabarinic acid

oxidation

intermediate

Fig. 14Model of interaction of CDHs and laccase during CA formation in P.cinnabarinus (TEMP and EGGERT1999)


Table 1Delignification of kraft pulp by Trametes versicolor laccase and various mediators (adapted fromBOURBONNAIS et al. 1997b)

Mediator O2 uptakenmole/min

Residual laccase%

Delig.%

Control – – 10.8ABTS 280.0 32.0 34.2HBT 1.5 2.0 37.8NNDS 129.0 76.0 26.4HNNS 130.0 50.0 30.1RBB 49.0 82.0 18.0PZ 165.0 52.0 6.3CPZ 78.0 50.0 10.8

Softwood kraft pulp: Initial kappa of 11.1Mediator: 1% on pulpLaccase: 0.5 U/l for all mediators except with HBT the amount of enzyme was 20 U/g

Recently, LI et al. (1997, 1999) have studied various organic substances for their medi-ating potential and found that two compounds, HBT and 3-hydroxy-1,2,3-benzotriazin-4(3H)-one (HBTO), were the most effective laccase mediators oxidizing a recalcitrant b-O-4lignin model dimer 1-(3,4-dimethoxyphenyl)-2-(2-methoxyphenoxy) propan-1,3-diol.

The ketone type product II can be readily degraded in the Dakin type reaction (HOSOYAand NAKANO 1980) by H2O2 in an alkaline solution, thus enabling an effective pulping oflignocelluloses. The laccase/ABTS;HBT systems also degraded other lignin model com-pounds as the b-1 dimer (Fig. 15).

OCH3OCH3

BenzaldehydeBenzaldehyde

Oxid

ati

on

pro

du

cts,

%of

init

ial

DII

Oxid

ati

on

pro

du

cts,

%of

init

ial

DII

DII + Laccase/HBT DII + Laccase/ABTSDII + Laccase/ABTS

VeratraldehydeVeratraldehyde

OCH3OCH3

OCH3OCH3

OCH3OCH3

OCH3OCH3

OCH3OCH3

HC

HCHC

HCOH

C

C

C

O

H

H

O

- carbonyl of D II- carbonyl of D II

- carbonyl of D II- carbonyl of D II

VeratraldehydeVeratraldehyde

BenzaldehydeBenzaldehyde

HBT or ABTSHBT or ABTS

HBTHBT

OHOH

OH

0

20

40

60

80

100

Fig. 15Transformation of b-1 dimer by Trametes versicolor laccase/ABTS;HBT system (adapted fromBOURBONNAIS et al. 1997a)


Various laccases obtained from six strains of white-rot fungi were compared for theirdelignifying activity with ABTS and HBT as mediators (Table 2).

As it can be observed, there are differences among these laccases both in the delignifica-tion degree and the recovery of residual enzyme activities. Although the authors did notexplain this phenomenon, it seems that such differentiation may be a result of the variedstatus of copper in the activity center of particular laccases as well as of the different redoxpotentials (LI et al. 1999).

Even better results were obtained in pulp delignification using the sequential xylanaseand laccase/mediator system treatments followed by the alkaline peroxide treatment. Inthese experiments, HBT was used as the mediator. The final pulp brightness and reductionof viscosity after the xylanase treatment followed by two “traditional” alkaline peroxidestages were higher than those for laccase/HBT alone followed by two peroxide stages. Thisresult has yielded the greatest brightness of pulp of all biochemical/traditional techniquesapplied so far. Nevertheless, in the case of laccase-containing systems, the darkening effectof the enzyme should be taken into consideration (POPPIUS-LEVLING et al. 1997). It isworth mentioning here that laccase alone, working in the pulp under high oxygen pressure

Table 2Delignification of the kraft pulp with various laccases and ABTS and HBT as mediators (adaptedfrom BOURBONNAIS et al. 1997b)

Pulp Treatment (Laccase source) Residual laccase%

Delig.,%

Phlebia radiata + ABTS+ HBT

32.011.0

34.520.9

Trametes versicolor + ABTS+ HBT

27.013.0

30.030.0

Pleurotus ostreatus + ABTS+ HBT

n.d.1.4

30.012.7

Merulius tremellosus + ABTS+ HBT

86.048.0

32.719.1

Fomes fomentarius + ABTS+ HBT

15.013.0

33.631.8

Ganoderma colossum + ABTS+ HBT

63.024.0

30.040.0

Pulp: 4 g of SW O2 (Kappa = 11.0) at 10% in 50 mM sodium acetate, pH 5Mediator: 1% on pulpTreatment: 2 h, 60 �C, 140 kP of 02 followed by alkaline extraction

SO Na3

SO Na3

COOHCOOHHOHO

SO CH CH OSO Na2 2 2 3

SO CH CH OSO Na2 2 2 3

NH2

NH2

CH3

CH3

OH

CH OH2

CH OH2

O

O

OHH

H HO

H

OH H

O

O

OHHOHO

NH

OH

Fig. 16The chemical structures of Remazol brilliant Blue R (RBBR, left) and carminic acid (CA, right)


(5 Atm), caused a pulp brightness comparable to that of the laccase/mediator systems with-out increased oxygen pressure (LEONOWICZ et al. 1993). Such an experiment was conducted5 years ago, when both ABTS and other laccase mediators were not known or had just beendiscovered.

Remazol brilliant Blue R (RBBR) and carminic acid (CA) were also used as laccasemediators for bleaching the pulp black liquor (kraft effluent) (Fig. 16).

In the experiment CA used as a mediator caused considerably higher brightness of theblack liquor originating from pulp chlorination than RBBR did (LEONOWICZ et al. 1993).On the other hand, inducible forms of laccase of Kuehneromyces mutabilis and Pleurotusostreatus decolorized both RBBR and CA, which could be considered the ligninolytic ac-tivity of the enzyme degrading the dyes (CHO et al. 1999b). Some authors reported earlierthat decolorization of several polymeric dyes by Phanerochaete chrysosporium usuallyappears under the conditions which favour lignin degradation, thus pointing to interrelationsof dye decolorization and ligninolysis (GLENN and GOLD 1983, SPANDARO et al. 1992).

In the presence of ABTS, Trametes versicolor laccase also oxidized veratryl alcohol toveratraldehyde through the hydrogen abstraction from the �-carbon of the side chain andthe formation of a benzyl radical.

The mechanism of laccase/ABTS electrochemical oxidation was further studied andconfirmed by cyclic voltammetry and bulk electrolysis (BOURBONNAIS et al. 1997b, 1998).The reactions involved are shown in Fig. 17.

Veratryl alcohol is oxidized by the dication of ABTS2+ formed from ABTS via theABTS.+ radical. Thus, the laccase/mediator system oxidizes veratryl alcohol to veratralde-hyde similar to lignin peroxidase, but not via a cation radical intermediate.

The laccase/ABTS couple also cleaved a recalcitrant lignin model dimer (BLANCHETTE1991) while hardwood kraft pulp was bleached by an in vitro laccase/ABTS complex to anextent similar to that in whole fungal cultures (BOYLE et al. 1992). However, since ABTSand other hitherto proposed mediating compounds do not appear in pulp, it is necessary tosearch for novel natural mediators originating from the mycelium or lignocellulose. 3-Hydroxyanthranilate (3-HAA) and syringaldehyde could perform this role. The former isproduced as a natural redox mediator by Pycnoporus cinnabarinus; the latter is a product oflignin degradation. 3-HAA has been reported to be capable of mediating the oxidation ofveratryl alcohol and 14C-ring labelled synthetic lignin (EGGERT et al. 1996).

It was also shown that the laccase:syringaldehyde couple can oxidize veratryl alcohol(KAWAI et al. 1989). The presence of syringaldehyde in the wood degrading environment(also in pulp) is possible since syringic acid (readily reduced by fungi) is a lignin degrada-tion metabolite (CHEN et al. 1982). Thus, syringaldehyde can be considered a naturalequivalent of the ABTS mediator of laccase in wood and pulp. On the other hand, the

-e-

ABTS

Veratryl alcohol Veratraldehyde

In solution

Elektrodesurface

E E472 mV 885 mV

-2H+

-2e-

ABTS ABTS

-e-

. 2 ++: :

Fig. 17Redox reactions of veratryl alcohol andABTS (adapted from BOURBONNAIS et al.1997b)


commercialization of these systems for pulp bleaching requires more effective and abun-dant mediators. The first attempt at using laccase/mediator couples for delignification in thepulp industry was made by the development of the LignozymR-Process, which is describedin detail in a review from 1997 (CALL and MUCKE 1997).

To sum up, ligninolytic enzymes – particularly laccase – often carry out their actualligninolytic activities by co-operating with low-molecular weight compounds acting asredox mediators. The ideal mediator should “form a high redox potential oxidation productin a highly reversible reaction” (BOURBONNAIS et al. 1997b). Cheaper bleaching of pulprequires optimization of both factors (BOURBONNAIS et al. 1997a and 1997b).

Cooperation of ligninolytic enzymes in the direct attack on the lignin polymer

Judging from what is already known about the very complex structure of the lignin macro-molecule, it is hard to imagine that this biopolymer can be effectively degraded by oneenzyme alone. It seems logical to accept a hypothesis suggesting the co-operation of severalindividual enzymes in the process of lignin degradation. It is well known, for instance, thatcellulose, a biopolymer of a relatively simple structure (compared to lignin), requires for itstransformation a number of enzymes acting consecutively when the process of hydrolysiscontinues. The situation may be similar in the case of the lignin polymer. As the ligninmacromolecule has a complex structure, researchers have seldom worked with natural lig-nin preparations and used relatively low-molecular weight methoxyphenolic dimers oroligomers which model parts of the lignin structure. After the discovery of lignin peroxi-dase, researchers studied its direct effect on the lignin macromolecule, but with time it hasbecome more and more evident that the reproduction of lignin depolymerization by onlyone enzyme (namely LiP) is a limited approach. On the other hand, it was known at thattime that LiP needs hydrogen peroxide for its activity. Accordingly, researchers searched

OCH3OCH3OCH3OCH3

OCH3OCH3OCH3OCH3

veratrylveratryl

cationradicalcationradical

GLYOXALGLYOXALglyoxic acidglyoxic acidglyoxalglyoxal

OXIDASEOXIDASE

SpontaneonusSpontaneonus

LMW ProductsLMW Products

alcoholalcohol

LIGNIN PEROXIDASELIGNIN PEROXIDASE

MN PEROXIDASEMN PEROXIDASE

OO

HOHOHOHO and Mn2 +and Mn2 +H O2 2H O2 2

H O2 2H O2 2

CO2CO2

H O2 2H O2 2

H O2H O2 O2O2

O2O2

LigninLigninO

OO

HO

OO

O

OHOH

OHOH

+.

Fig. 18Possible co-operation of three enzymes (glyoxal oxidase, manganese peroxidase and lignin peroxi-dase) in ligninolysis Phanerochaete chrysosporium (adapted from KERSTEN et al. 1990)


for enzymes comprising this compound. Interestingly, one of the enzymes that generateshydrogen peroxide is manganese peroxidase. For this reason, PASZCZYNSKI et al. (1985)considered the possibility of co-operation of both enzymes in the transformation of ligninrelated compounds.

Five years later, KERSTEN et al. (1990) introduced veratryl alcohol as a mediator in ligni-nolysis and suggested the possibility of co-operation of lignin peroxidase with two otherenzymes, namely glyoxal oxidase and manganese peroxidase. Such a system may work incultures of Phanerochaete chrysosporium. For this reason, the lignin depolymerization tolower-molecular weight products (LMW) goes through the cation radicals found in theirlaboratory (Fig. 18).

Glucose oxidase as the hydrogen peroxide donor can also co-operate with lignin peroxi-dase. Such a co-operation in the decomposition of aromatic dimers by a system of bothenzymes was shown by GLENN et al. (1983). Glucose oxidase also co-operates with laccase.As it has already been mentioned, laccase alone can directly depolymerize the lignin mac-romolecule, but the efficiency of depolymerization is low and depends on the rapid removalof reactive radicals or quinones from the media that are formed by the action of laccase(such phenoxy radicals are created, e.g., when laccase acts on 2,4,6-trimethoxyphenol).Spontaneous coupling of the radicals which are formed from phenolic lignin fragmentsleads to products of higher molecular weight (compare Fig. 2). It has turned out that glucoseoxidase is an efficient factor reducing radicals and quinones formed by the action of lac-case. It may function according to the mechanism proposed by GREEN (1977, Fig. 19).

The hydrogen acceptors in glucose oxidation are quinones or radicals (but not molecularoxygen), which are formed from the phenolic compounds by laccase. A similar course ofreaction was noticed by ALBERTI and KLIBANOV (1982) with regard to p-benzoquinone. Thequinoide compound was reduced to hydroquinone by glucose oxidase.

We have investigated the transformation of various phenolic compounds under the se-quential action of laccase and glucose oxidase (LEONOWICZ et al. 1997b). In all cases, thequinones formed by laccase disappeared from the reaction medium when glucose oxidasewas introduced into the reaction mixture. The oxidation of several phenolic compounds andBJÖRKMAN lignin by Pleurotus ostreatus laccase and the reduction of the correspondingquinones may serve as examples of this (Fig. 20).

OH OHOH

H O2 2H O2 2

O2O2

OHOH

OXIDIZEDDEGRADATION

PRODUCTS

OXIDIZEDDEGRADATION

PRODUCTS

POLYMERIZEDQUINOIDS

POLYMERIZEDQUINOIDS

GLUCONOLACTONE

GLUCOSEGLUCOSE

OHOHH HOHOHHO

O

O O OHOHO

+ O2+ O2

O

R R RR

I

IIII H

-2e-2e

R

Fig. 19Cyclic diagram of possible co-operation of laccase and D-glucose oxidase (adapted from GREEN1977)


0.0

0.2

0.4

0.6

0.8

1.0

10 20 30 40 50

Time (min.)

Ab

sorb

an

ce

(390

nm

)

More recently, MARZULLO et al. (1995, 1996) reported that not only glucose oxidase, butalso another FAD-dependent oxidase – veratryl alcohol oxidase – can co-operate withlaccase in the transformation of some phenol/quinone derivatives. The above mentionedenzyme is secreted by the white-rot fungus Pleurotus ostreatus. The authors found that theenzyme is able to reduce laccase-generated quinones and phenoxy radicals with concomi-

0.00

0.02

0.04

0.06

0.08

0.10

80 90 100 110 120 130 140

Fraction number

Ab

sorb

an

ce

(28

0n

m)

Fig. 21Elution pattern of Björkman lignin on a Sepharose Cl-6B column after treatment with both laccaseand glucose oxidase (adapted from LEONOWICZ et al. 1997b, 1999d)

Fig. 20Formation and reductionof quinones under se-quential action of lac-case and glucose oxi-dase on BJÖRKMAN lig-nin and several phenoliccompounds (adaptedfrom LEONOWICZ et al.1997b)


tant oxidation of veratryl alcohol to veratryl aldehyde. This system also works with syn-thetic quinones, e.g., 2,6-dichlorophenoloindophenol (DCIP). For this reason, DCIP wasused both to assay the quinone reduction and to test the effect of oxygen and veratryl alco-hol on the veratryl alcohol oxidase activity. The cycle given below demonstrates that vera-tryl alcohol oxidase reduces DCIP and consequently prevents the polymerization of guaia-col and ferulic acid by laccase.

This mechanism have regulatory properties, which may, in fact, be applicable in lignintransformation. The first attempt at applying laccase and glucose oxidase was made in ourlaboratory. Two fractions of lignosulphonates (Peritan Na) were applied as laccase sub-strates (SZKLARZ and LEONOWICZ 1986); it was demonstrated that glucose oxidase added tothe reaction mixtures effectively counteracted polymerization and even accelerated thedepolymerization process catalyzed by laccase.

This noticeable effect of laccase and glucose oxidase working in concert during ligni-nolysis was confirmed using Björkman lignin isolated from wheat straw (Fig. 21).

Similar results were reported by GALLIANO et al. (1991). The authors used 14C-labelledKLASON lignin isolated from 14C-labelled ligninocellulose. The latter biopolymer was takenfrom young Hevea twigs grown in a medium with 14C-phenylalanine. The lignin solubiliza-tion in the conditions containing the glucose/glucose oxidase system increases three-foldcompared to the conditions with laccase alone.

Analogously to glucose oxidase, veratryl alcohol oxidase may also co-operate with lac-case (MARZULLO et al. 1995). Its co-operative action prevented the polymerization of phe-nolic lignin fragments and reduced the molecular weight of soluble lignosulphonates to asignificant extent.

Time_(days)

! " # $ %

14C

Solu

biliz

ed(%

ofto

tal14

C)

"&

"!

&

!

Fig. 2214C lignin solubilization by laccase andmanganese peroxidase. (�) control with-out enzymes; (�) laccase and manganeseperoxidase together; (�) laccase, manga-nese peroxidase and glucose oxidasetogether (adapted from GALLIANO et al.1991)


POSSIBLEOSSIBLELACCASE AND SUPEROXIDE DISMUTASE (SOD) COOPERATIONLACCASE AND SUPEROXIDE DISMUTASE (SOD) COOPERATIONIN PRODUCTION OF LIGNINOLYTIC FACTORS AND RADICALSIN PRODUCTION OF LIGNINOLYTIC FACTORS AND RADICALS

LACCASELACCASE

SODSOD ( SOR dismutation )( SOR dismutation )

Lignin degradationLignin degradation

Lignin degradationLignin degradation

Q2Q2

Q2Q2 Fe3+Fe3+

Fe3+Fe3+

Fe+2Fe+2

Fe+2Fe+2

Q

QQ

Q

+

+

+

+

+

+

+

+

+

++

+

+

+

O2O2

O2O2

O2O2

O2O2

O2O2O2O2Haber - Weiss reaction:Haber - Weiss reaction:

hydroquinonehydroquinone

semiquinonesemiquinone

SOD

superoxide radical (SOR)superoxide radical (SOR)

superoxidesuperoxide dismutasedismutase

OHFenton reaction:Fenton reaction:

OH

OH

OH

( iron reduction )( iron reduction )O2O2

O2O2

O2O2

2O22O2 H O2 2H O2 2

H O2 2H O2 2

H O2 2H O2 2

O2O22H

=

=

=

=

=

=

=

=

=

=

Fig. 23The possible co-operation of laccase and superoxide dismutase in production of ligninolytic factorsand radicals (adapted from LEONOWICZ et al. 1999b)

Manganese peroxidase can also effectively co-operate with laccase, which was unequivo-cally stated in the report by GALLIANO et al. (1991). When manganese peroxidase and lac-case, both isolated from cultures of Rigidosporus lignosus, were added to the same reactionmixture containing radioactive KLASON lignin, lignin degradation was efficient. The addi-tion of glucose oxidase to such an enzyme system again increased the amount of water-soluble, low-molecular weight products. In the presence of either laccase or manganeseperoxidase alone, lignin solubilization was also observed, but at a relatively low level. Thiswas the first demonstration of the synergistic effect of two oxidizing enzymes involved inlignin biodegradation (Fig. 22).

In our more recent results, the maximum of SOD activity during lignin biodegradation bylaccase- and peroxidases-producing fungi appeared when the activities of ligninolytic en-zymes decreased (MALARCZYK et al. 1995, LEONOWICZ et al. 1999a, 1999c). This may becaused by the fact that when the ligninolytic enzymes are not active, SOD protects the or-ganisms from toxic, reactive radicals. On the other hand, during the high activity of theseenzymes, the radicals might be fully involved in the biodeterioration processes. SOD ismost likely involved in the production of these factors from the intermediates of quinoneredox cycling caused by laccase (see Fig. 23). This thesis, however, requires further ex-perimental confirmation.

Cooperation of laccase with other enzymes and mediating factorsin ligninocellulose degradation

It has been unequivocally stated that lignin degradation is accelerated in the presence ofcellulose or its oligomers (HATAKKA and UUSI-RAUVA 1983). The idea of a feedback typeinterdependence of delignification and cellulose degradation was already postulated by


WESTERMARK and ERIKSSON (1974a, b). This hypothesis is still valid as the reports byGOTTLIEB et al. (1950) concerning the possibility of fungal growth on lignin as the solesource of carbon and energy have not been confirmed; the lignin degradation is now gener-ally thought to be a cometabolic process. The processes of depolymerization of celluloseand of lignin are interrelated in certain points and even accelerate each other (WESTERMARKand ERIKSSON 1974a, 1974b, 1975). The authors discovered the enzyme cello-biose : quinone oxidoreductase, which co-operates with laccase and cellulose hydrolyzingenzymes in the depolymerization of both components of the ligninocellulose complex in afeedback fashion by removing decomposition products. The authors suggested that laccasemight function as a link in an extracellular “electron transport chain” in this system.

It is necessary to mention that ligninolytic peroxidases, nowadays regarded as the crucialenzymes in the disassembly of lignin, had not been known when cellobiose:quinone oxi-doreductase was discovered. For this reason, ANDER and MARZULLO (1997) modifiedWESTERMARK and ERIKSSON’s scheme by introducing the enzymes and radical intermedi-ates discovered in the meantime (Fig. 24).

The DCIP oxidizing and reducing laccase/aryl alcohol oxidase system described byMARZULLO et al. (1995) prevents lignin polymerization as well as considerably reduces themolecular masses. Aryl alcohol oxidase is a part of systems involving not only laccase butalso other enzymes and factors. The hypothetical relationship between aryl alcohol oxidase,laccase, and other enzymes and radicals in the process of ligninocellulose transformation isillustrated in Fig. 25.

In this proposed complex system, laccase, LiP and MnP are secreted by fungal hyphaeclose to the hyphae environment, where they co-operate with each other as well as withmediating factors. The yielded chelators and mediating radicals are exported further to thewood tissue where they work as enzyme “messengers” in ligninocellulose degradation.Therefore, it has been proposed that this feed-back type enzyme system works in lignino-cellulose transformation with a meaningful participation of different ligninolytic enzymes.

native

LiP, MnP, Laccase

native

LiP, MnP, Laccase

PhenolsPhenols

ProteaseProtease

Cellobiose + O2Cellobiose + O2 Cellobiono - 1,5 - lactoneCellobiono - 1,5 - lactone

Cellobionic acidCellobionic acid

+ heme+ hemeMn4+Mn4+ Mn2+Mn2+ Mn3+Mn3+Fe3+Fe3+

Fe2+Fe2+

native LiP, MnPnative LiP, MnPQuinones, radicals

Compound II

Quinones, radicals

Compound II

Lignin

CDH

Cellulose

CBQCBQ

H O or O2 2 2H O or O2 2 2

H O2 2H O2 2 OHOH

H O2H O2

, ,

+

Fig. 24Interaction of the cello-biose dehydrogenase/cellobiose: quinone oxi-doreductase system(CDH/CBQ) with otherenzymes, quinones andphenoxy radicals pro-duced by these enzymes(adapted from ANDERand MARZULLO 1997)


FADH2FADH2FADH2FADH2

Fe(II)Fe(II)

Mn(II) Mn(III)Mn(III)Mn(III) ( )Mn(III) ( )

Mn(III) ( )Mn(III) ( )

Mn(III) ( )Mn(III) ( )

Fe(III)Fe(III)

FADFAD

AADAAD

NADPH

NADP

LaccoseLiP or MnPLaccoseLiP or MnP

O2O2

H O2 2H O2 2

H O2 2H O2 2 H O2H O2

OH OH

OH

OHOHOH

MnPMnP

LiPLiP

H O2 2H O2 2

H O2 2H O2 2 H O2H O2

++

+

+ +

+ +

-

-

+

+ +

+

+ +

+

+

+

+

H

Hypho

LignocelluloseLignocellulose

( )

Fig. 25Interaction of aryl alcohol dehydrogenase and other fungal enzymes with mediators and mediatingradicals during transformation of ligninocellulose. From left to right: AAD – aryl alcohol dehydroge-nase with NADP as the prosthetic group; aryl alcohol; aryl aldehyde; LiP – lignin peroxidase; MnP –manganese dependent peroxidase; lignin derived radicals or quinones and their reduced forms, VAO –veratryl alcohol dehydrogenase with FAD as the prosthetic group; LiP with Fe; MnP with Mn; metalchelating agents, e.g. oxalic acid (adapted from ANDER and MARZULLO 1997)

Laccase oxidizes lignin-derived radicals to quinones. These can serve as the oxygensource for glucose oxidase (GOD, SZKLARZ and LEONOWICZ 1986) and aryl alcohol oxidase(MARZULLO et al. 1995) producing H2O2 and preventing polymerization (SZKLARZ andLEONOWICZ 1986, MARZULLO et al. 1995); H2O2 formed serves as the co-substrate forligninolytic peroxidases. The CDH/CBQ system (Fig. 24) requires the presence of dioxygenand hydrogen peroxide for functioning; dioxygen is supplied from air, but external hydro-gen peroxide (needed at least by lignin peroxidase) is only produced at a low rate, by CDHand CBQ, as suggested by the authors. The crucial radical for H2O2 production is superox-ide (O2

. –), probably yielded from quinone cycling (see Superoxide dismutase) by generatingH2O2 through the oxidation of Mn2+. Simultaneously, highly reactive Mn3+ is formed. It isknown from manganese peroxidase to be an efficient lignin attacking agent.

O2. – + Mn2+ + 2H+ � Mn3+ + H2O2 (11)

H2O2 produced here accelerates both the production of Mn3+ from Mn2+ by manganeseperoxidase and the oxidation of lignin and derived fragments (WARIISHI et al. 1992).

However, since the H2O2 production by this system is not sufficient for all H2O2-consuming reactions, it was necessary to look for other natural systems generating enoughH2O2 for the biodeterioration of the ligninocellulose complex. Various possibilities havebeen considered. First of all, the possible involvement of certain enzymes was proven, e.g.,glyoxal oxidase, veratryl alcohol oxidase, manganese peroxidase, or glucose oxidase. Someof these enzymes can function both intracellularly and extracellularly. Other enzymes gen-erating hydrogen peroxide, such as pyranose-2-oxidase (mentioned above), methanol oxi-dase (EC 1.1.3.13), and fatty-acyl-CoA oxidoreductase (EC 1.2.1.42), work only intracel-lularly (DE JONG et al. 1994). This is why they should not be considered as woodtransformation is an exclusively extracellular process. The enzymes listed in the first group


are relatively stable and resistant to the conditions outside the cells. Among them, only glu-cose oxidase can join both lignin and carbohydrate metabolic chains. This means that glucoseoxidase possesses two possible functions for the acceleration of ligninocellulose degrada-tion: it produces H2O2 by oxidation of glucose (required for peroxidases), and it reduces qui-noids and phenoxy radicals originating from the action of laccase during lignin oxidation.

The remaining above listed enzymes which also reduce quinones and produce H2O2 (e.g.,aryl/veratryl alcohol oxidases) have been taken into consideration, but to a smaller extentbecause their H2O2 does not result from the oxidation of glucose, which is a key metabolitein carbohydrates transformation. Accordingly, probably only glucose oxidase co-operateswith the cellulolytic system by using glucose as the substrate generated by hydrolytic en-zymes from cellulose. The enzyme is produced by all white-rot fungi tested, includingPhanerochaete chrysosporium (KELLEY and REDDY 1988), which secrete laccase(LEONOWICZ et al. 1986) and/or peroxidases (ROGALSKI and LEONOWICZ 1992) and whichshow strong ligninolytic abilities (LEONOWICZ et al. 1986). Phanerochaete chrysosporiumwhich has been deprived of glucose oxidase by metagenesis is not able to decompose ligninat all (RAMASAMY et al. 1985).

In summary, glucose oxidase is a feed-back type enzyme that might prove to be impor-tant in ligninocellulose transformation. When oxygen is indispensable in the second stage ofthe glucose oxidase-glucose reaction, it can be supplied by radicals or quinones generatedwhen phenolic or methoxyphenolic compounds are exposed to laccase (SZKLARZ andLEONOWICZ 1986). On the other hand, when excess quinones produced by laccase inhibitthe enzyme (SZKLARZ and LEONOWICZ 1986), it can be concluded that glucose oxidasecounteracts the poisonous level of quinones in the medium and enables laccase to continueits function. Taking all these reasons into consideration, already more than 10 years ago wepostulated the following hypothetical mechanism of ligninocellulose complex transforma-tion (LEONOWICZ et al. 1986; Fig. 26).

CH OH3CH OH3

O2O2

O2O2

O2O2

CO2CO2

HO2

2

HO2

2

METHOXYPHENYLMETHO YPHENYLDERIVDERIVATIVESTIVES

LIGNIN

DISACCHARIDESDISACCHARIDES

DIPHENOLSDIPHENOLS

KETOACIDSKETOACIDS

KREBSCYCLEKREBSCYCLE

GLUCOSE

( XYLOSE )

OXIDASE

GLUCOSE

( YLOSE )

OXIDASE

QUINONESQUINONES -D-GLUCONO (XYLONO)LACTONE

-D-GLUCONO ( YLONO)LACTONE

�

PENTOSEPATHWAYPENTOSEPATHWAY

GLUCOSE(XYLOSE)GLUCOSE(XYLOSE)

WOODPOLYSACCHARIDESWOODPOLYSACCHARIDES

Dioxygenase

Dioxygenase

Lignindegradingenzyme

Lignindegradingenzyme

Gluco (xylo)

nasesGluco (xylo)

nases

Gluco (xylo)

sidasesGluco (xylo)

sidases

LaccaseLaccase

Fig. 26Hypothetical mechanism of ligninocellulose transformation by enzymes of white-rot fungi. The initialproducts of partial wood hydrolysis distinctly induce enzymatic systems accelerating the degradationprocesses (LEONOWICZ et al. 1986, ROGALSKI 1986)


In these mechanisms, glucose oxidase co-operates with lignin and manganese peroxidaseproviding hydrogen peroxide as well as with laccase reducing quinones to phenols. For thisreason, glucose oxidase operates as a feedback system where lignin and manganese peroxi-dase function as the first lignin attacking enzymes and laccase as the demethylating factor.Glucose, which is produced as a result of cellulose hydrolysis by the cellulase complex,becomes the substrate for glucose oxidase. Oxygen needed by glucose oxidase can be re-placed by quinones produced by laccase from lignin oligomers. As a result of glucose oxi-dation, �-D-gluconolactone is formed, which, after certain transformations, reinforces themetabolism of the fungus in the pentose phosphate cycle or in glycolysis. Hydrogen perox-ide produced in the reaction catalyzed by glucose oxidase activates, in turn, lignin andmanganese peroxidase. Lignin exposed to peroxidases undergoes decomposition into lower-molecular weight fragments containing methoxyl groups. Laccase demethylates these oli-gomers while peroxidases degrade them further into even smaller fragments. Generation ofexcess quinones and resulting repolymerization is counterbalanced by glucose oxidase,which reduces them to the respective phenols. The phenols, in turn, are the substrates fordioxygenases present in all fungal cells resulting in ring cleavages; the products obtained inthe form of ketoacids easily find their way to the Krebs cycle.

The above described mechanism could occur in those fungi which possess all the abovementioned enzymes, e.g., Phanerochaete chrysosporium, Phlebia radiata, Trametes versi-color, or Nematoloma frowardii. The mechanism suggested by WESTERMARK and

CH OH3CH OH3

H O2 2H O2 2

H O2 2H O2 2

H O2 2H O2 2

O2O2

CO2CO2

O2O2

Ligm

inase

Ligm

inase

LigninaseLigninase

LigninaseLigninase

Dioxygenase

Ring - cleavageproductsRing - cleavageproducts

Energy +pentosepathway

Energy +pentosepathway

GluconoXylonoCellobionolactones

GluconoXylonoCellobionolactones

PhenolsPhenols

KrebscycleKrebscycle

Ring - cleavageproductsRing - cleavageproducts

de novo

de novo

GlucoseXyloseGlucoseXylose

Glucose(Xylose)oxidases

Glucose(Xylose)oxidases

CBQXBQCBQXBQ

QuinonesPhenoxy radicalsQuinonesPhenoxy radicals

Methoxyphenylderivatives

Methoxyphenylderivatives

Laccase

Peroxidases

Laccase

PeroxidasesO

HO

22

2

OH

O

22

2

Gly

co-

sida

ses

Gly

co-

sida

ses

LIGNIN

LIGNIN

CellobioseXylobioseCellobioseXylobiose

Partlydegradedlignin

Partlydegradedlignin

WOODPOLYSACCHARIDESWOODPOLYSACCHARIDES

Ver. alc. Ver. alc.alc.

Cel

lula

ses

Xyl

anas

es

Cel

lula

ses

Xyl

anas

es

+ .+ .

+

Fig. 27Hypothetical scheme for de-gradation of lignin, celluloseand xylan in wood (adaptedfrom ERIKSSON et al. 1990,ERIKSSON 1993)


ERIKSSON, where cellobiose : quinone oxidoreductase is the crucial feed back type enzyme,seems to apply quite well to those species in which the presence of lignin peroxidase hasnot been confirmed but which transform ligninocellulose. It is not out of the question thatboth systems also act in concert as it was shown by ERIKSSON et al. (ERIKSSON et al. 1990,LEONOWICZ et al. 1991b, ROGALSKI 1992, ERIKSSON, 1993, Fig. 27).

As it can be observed, our proposed method of ligninocellulose transformation has beenfurther developed by Eriksson’s team by supplementing cellobiose:quinone oxidoreductaseand mediating factors. Its areas of functioning have been greatly extended (not one, but twoenzymes with feedback activity, veratryl alcohol, phenoxy radicals). We are in full agree-ment with this proposition, particularly since it also takes into consideration the contribu-tion of fungal laccase and peroxidases to the demethylation of lignin, and methoxyphenylderivatives, which was postulated by our group many years ago (TROJANOWSKI et al. 1966,LEONOWICZ and TROJANOWSKI 1965). This means that our old conception of the initiallignin transformation by these enzymes still remains valid. Recent discoveries concerningthe role of aryl alcohol oxidase, cellobiose dehydrogenase, aryl alcohol dehydrogenase, and,especially, low-molecular weight mediators and radicals in wood biodeterioration shown inFig. 24 (ANDER and MARZULLO 1997) support the proposed routes by explaining how high-molecular weight enzymes can function in the wood environment which is normally toocompact to allow the penetration of microbial enzymes. However, the existence of other,still-to-be-discovered mechanisms as well as of enzymes also operating as feedback sys-tems in the process of ligninocellulose transformation cannot be ruled out. Detailed investi-gations into these processes would definitely help solve the problems of how to utilizewaste lignocelluloses are accumulated as a result of human activities or how to protect ourforests from damaging microbial attack.

Conclusions

Considering earlier and present results, we can conclude that cellulose and lignin are de-graded simultaneously and that the general outline of the complementary character of car-bohydrates and lignin decomposition as well as the existence of enzymatic systems com-bining these processes are still valid. Demethylation caused by laccase after the initialactivity of lignin and manganese peroxidase appears to be important as the depolymerizingactivity of lignin peroxidase is inhibited by the phenolic groups of native lignin (ANDER andMARZULLO 1997). Also, the role of laccase (induced by toxic lignin degradation products orstress conditions) lies probably in the “activation” of some low-molecular weight mediatorsand radicals produced by fungal cultures. Such activated factors, also produced in coopera-tion with other enzymes, are probably exported into the wood microenvironment wherethey work in degradation processes as the “enzyme messengers.”

In conclusion, it is worth mentioning that the depolymerizing activity of laccase is ratherunusual, because its routine enzyme activity comprises polymerization or coupling reac-tions. However, this routine activity is usually employed in laboratories. Under naturalconditions laccase works in the wood environment in concert with other fungal enzymes,mediators and mediating radicals. Apart from polymerization, splitting processes are alsopossible there. It is in these processes that laccase finds the proper position among otherligninolytic enzymes.

Acknowledgements

Parts of the works cited here were financially supported by the European Community (the RTDE5 ECContract ICA2-CT-2000-10050), KBN SPUB-M.-5PR-UE/DZ 280/2000 and BS/UMCS projects.


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Mailing address: Prof. Dr. ANDRZEJ LEONOWICZ, Department of Biochemistry, Maria-Curie-Sklodowska University, Pl-20031 Lublin, PolandTel.: +49-81-5375665; Fax: +48-81-5333669;E-mail: [email protected]

Fungal laccase: properties and activity on lignin.

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