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LACCASES 473

5.8. Ethanol Production

Dilute acid hydroysis of lignocellulose yields sugar mixtures which can be fermented

by yeast to produce fuel alcohol. However, phenolic compounds present in the

hydrolysate inhibit yeast fermentation. Heterologous expression of the laccase from

Trametes versicolor in S. cerevisiae during fermentation has recently been demon-

strated to improve the production of fuel ethanol from this renewable raw material

by eliminating these compounds (Larsson et al., 2001).

5.9. Cosmetics

Bleaching and/or dyeing of hair usually involves the use of harsh chemicals which

can damage hair. Dye precursors can be oxidized to the effective colouring agent

using laccase instead of the chemical agent. Laccases have also been used in

the presence of hydroxy stilbenes as hair bleaches (Onuki et al., 2000; Pruche

et al., 2000). Pigments used in cosmetic applications as well as melanins could be

produced using laccase catalysed oxidative reactions. Conversely, laccase may also

find utility in the reduction of melanin-derived blemishes in topical skin applications

(Tetsch et al., 2005; Nagai et al., 2003).

5.10. Food Industry

Laccase can be used in beverage (wine, fruit juice and beer) processing, ascorbic

acid determination, sugar beet pectin gelation, baking, as a biosensor and to improve

food sensory parameters (Minussi et al., 2002).

Wine stabilization is one of the main applications of laccase in the food industry.

Laccase is used to selectively target specific polyphenols during the madeirization

process. The use of the laccase in must results in a stable wine with a good flavour.

Since the use of laccase as a food additive is still not permitted, this enzyme has

been used in wine production in an immobilized form to ensure its elimination from

the must and facilitate its reutilization (Servili et al., 2000).

The tendency for hazes to develop in some beers during long-term storage is

a persistent problem in the brewing industry. As an alternative to the traditional

treatment, laccase can be added to the wort or at the end of the process. Since

oxygen is not desirable in the finished beer, addition of laccase can remove any

excess oxygen whereby the storage life of beer is enhanced. At the same time the

laccase removes some of the polyphenols that may still remain in the beer. The

polyphenol complexes formed by laccase may be removed by filtration or other

means of separation (Mathiasen, 1995).

For apple and grape juices excessive oxidation of phenolics has almost always

been considered detrimental to the organoleptic quality of the product. These

beverages are typically stabilized to delay the onset of protein-polyphenol haze

formation. Various enzymatic treatment have been proposed for fruit juice stabi-

lization, among them the use of laccase (Minussi et al., 2002).

474 ALCALDE

REFERENCES

Alcalde, M., Bulter, T. and Arnold, F.H. (2002) Colorimetric assays for biodegradation of polycyclic

aromatic hydrocarbons by fungal laccases. J. Biomol Screening 7, 547–553

Alcalde, M., Bulter, T., Zumárraga, M., García-Arellano, H., Mencía, M., Plou, F.J. and Ballesteros, A.

(2005) Screening mutant libraries of fungal laccases in the presence of organic solvents. J. Biomol.

Screening. 10, 624–631.

Alexandre, G. and Zhulin, I.B. (2000) Laccases are widspread in bacteria. Trends Biotechnol. 18, 41–42

Almansa, E., Kandelbauer, A., Pereira, L., Cavaco-Paulo, A. and Guebitz, G.M. (2004) Influence of

structure on dye degradation with laccase mediator systems. Biocatalysis and Biotransformation 22,

315–324

Alves, A.M.C.R., Record, E., Lomascolo, A., Scholtmeijer, K., Asther, M., Wessels, J.G.H. and Wosten,

H.A.B. (2004) Highly efficient production of laccase by the basidiomycete Pycnoporus cinnabarinus.

App. Environ. Microbiol. 70, 6379–6384

Amitai, G., Adani, R., Sod-Moriah, G., Rabinovitz, I., Vincze, A., Leader, H., Chefetz, B., Leibovitz-

Persky, L., Friesem, D. and Hadar, Y. (1998) Oxidative biodegradation of phosphorothiolates by

fungal laccase. Febs Letters 438, 195–200

Bajpai, P. (2004) Biological bleaching of chemical pulps. Crti. Rev. Biotechnol. 24, 1–58

Bourbonnais, R. and Paice, M.G. (1990) Oxidation of nonphenolic substrates - an expanded role for

laccase in lignin biodegradation. Febs Letters 267, 99–102

Bulter, T., Alcalde, M., Sieber, V., Meinhold, P., Schlachtbauer, C. and Arnold, F.H. (2003a) Functional

expression of a fungal laccase in Saccharomyces cerevisiae by directed evolution. Appl. Environ.

Microbiol. 69, 987–995

Call, H.P. and Call, S. (2005) New generation of enzymatic delignification and bleaching. Pulp and

Paper-Canada 106, 45–48

Call, H.P. and Mucke, I. (1997) History, overview and applications of mediated lignolytic systems,

especially laccase-mediator-systems (Lignozym(R)-process). J. Biotechnol. 53, 163–202

Camarero, S., Ibarra, D., Martinez, M.J. and Martinez, A.T. (2005) Lignin-derived compounds as

efficient laccase mediators for decolorization of different types of recalcitrant dyes. Appl. Environ.

Microbiol. 71, 1775–1784

Chen, T, Barton SC., Binyamin G, Gao Z, Zhang Y, Kim HH and Heller A (2001) A miniature biofuel

cell. J. Am. Chem. Soc. 123, 8630–8631

Claus, H. (2003) Laccases and their occurrence in prokaryotes. Arch. Microbiol. 179, 145–150

Claus, H. (2004) Laccases: structure, reactions, distribution. Micron 35, 93–96

Collins, P.J., Kotterman, M.J.J., Field, J.A. and Dobson, A.D.W. (1996) Oxidation of anthracene and

benzo[a]pyrene by laccases from Trametes versicolor. Appl. Environ. Microbiol 62, 4563–4567

Davies, G.J.a.D. (2002) Laccase. Handbook of Metalloproteins. (ed A.Messerschmidt, et al.), pp. 1359–

1368. John Wiley and Sons, LTD, New York

Ducros, V., Brzozowski, A.M., Wilson, K.S., Brown, S.H., Ostergaard, P., Schneider, P.,

Yaver, D.S., Pedersen, A.H. and Davies, G.J. (1998) Crystal structure of the type-2 Cu depleted

laccase from Coprinus cinereus at 2.2 angstrom resolution. Nature Struct. Biol. 5, 310–316

Duran, N., Rosa, M.A., D’Annibale, A. and Gianfreda, L. (2002) Applications of laccases and tyrosinases

(phenoloxidases) immobilized on different supports: a review. Enzyme Microb. Technol. 31, 907–931

Enguita, F.J., Martins, L.O., Henriques, A.O. and Carrondo, M.A. (2003) Crystal structure of a bacterial

endospore coat component - A laccase with enhanced thermostability properties. J. Biol. Chem. 278,

19416–19425

Felby, C., Hassingboe, J. and Lund, M. (2002) Pilot-scale production of fiberboards made by laccase

oxidized wood fibers: board properties and evidence for cross-linking of lignin. Enzyme Microb.

Technol. 31, 736–741

Freire, R.S., Duran, N. and Kubota, L.T. (2002) Development of a laccase-based flow injection electro-

chemical biosensor for the determination of phenolic compounds and its application for monitoring

remediation of Kraft E1 paper mill effluent. Anal. Chim. Acta 463, 229–238

LACCASES 475

Freire, R.S., Ferreira, M.M.C., Duran, N. and Kubota, L.T.(2003) Dual amperometric biosensor device

for analysis of binary mixtures of phenols by multivariate calibration using partial least squares. Anal.

Chim. Acta 485, 263–269

Gianfreda, L., Xu, F. and Bollag, J. (1999) Laccases: a useful group of oxidoreductive enzymes.

Bioremed. J. 3, 1–25

Hakulinen, N., Kiiskinen, L.L., Kruus, K., Saloheimo, M., Paananen, A., Koivula, A. and Rouvinen, J.

(2002) Crystal structure of a laccase from Melanocarpus albomyces with an intact trinuclear copper

site. Nature Struct. Biol. 9, 601–605

Huttermann, A., Mai, C. and Kharazipour, A. (2001) Modification of lignin for the production of new

compounded materials. Appl. Microbiol. Biotechnol. 55, 387–394

Jauregui, J., Valderrama, B., Albores, A. and Vazquez-Duhalt, R. (2003) Microsomal transformation of

organophosphorus pesticides by white rot fungi. Biodegradation 14, 397–406

Johannes, C. and Majcherczyk, A. (2000) Natural mediators in the oxidation of polycyclic aromatic

hydrocarbons by laccase mediator systems. Appl. Environ. Microbiol. 66, 524–528

Johannes, C., Majcherczyk, A. and Huttermann, A. (1996) Degradation of anthracene by laccase of

Trametes versicolor in the presence of different mediator compounds. Appl. Microbiol. Biotechnol.

46, 313–317

Kiiskinen, L.L. and Saloheimo, M. (2004) Molecular cloning and expression in Saccharomyces cerevisiae

of a laccase gene from the ascomycete Melanocarpus albomyces. Appl. Environ. Microbiol. 70,

pp. 137–144

Kumar, S.V.S., Phale, P.S., Durani, S. and Wangikar, P.P. (2003) Combined sequence and structure

analysis of the fungal laccase family. Biotechnol. Bioeng. 83, 386–394

Larsson, S., Cassland, P. and Jonsson, L.J. (2001) Development of a Saccharomyces cerevisiae strain with

enhanced resistance to phenolic fermentation inhibitors in lignocellulose hydrolysates by heterologous

expression of laccase. Appl Environ Microbiol 67, 1163–1170

Luterek, J., Gianfreda, L., Wojtas-Wasilewska, M., Cho, N.S., Rogalski, J., Jaszek, H., Malarczyk, E.,

Staszczak, M. and Fink-Boots, M. (1998) Activity of free and immobilized extracellular Cerrena

unicolor laccase in water miscible organic solvents. Holzforschung 52, 589–595

Majcherczyk, A. and Johannes, C. (2000) Radical mediated indirect oxidation of a PEG-coupled

polycyclic aromatic hydrocarbon (PAH) model compound by fungal laccase. Biochim. Biophys.

Acta-Gen. Subj. 1474, 157–162

Majcherczyk, A., Johannes, C. and Huttermann, A. (1998) Oxidation of polycyclic aromatic hydrocarbons

(PAH) by laccase of Trametes versicolor. Enzyme Microb. Technol. 22, 335–341

Mathiasen, T.E. Laccase and beer storage. [PCT Int.Appl.WO 9521240 A2]. 1995. Ref Type: Patent

Mayer, A.M. and Staples, R.C. (2002) Laccase: new functions for an old enzyme. Phytochemistry 60,

551–565

Milligan, C. and Ghindilis, A. (2002) Laccase based sandwich scheme immunosensor employing media-

torless electrocatalysis. Electroanalysis 14, 415–419

Minussi, R.C., Pastore, G.M. and Duran, N. (2002) Potential applications of laccase in the food industry.

Trends Food Sci. Technol. 13, 205–216

Nagai, M., Kawata, M., Watanabe, H., Ogawa, M., Saito, K., Takesawa, T., Kanda, K. and Sato, T. (2003)

Important role of fungal intracellular laccase for melanin synthesis: purification and characterization

of an intracellular laccase from Lentinula edodes fruit bodies. Microbiology-Sgm 149, 2455–2462

Onuki, T., Nogucji, M. and Mitamura, J. Oxidative hair dye composition containing laccase. [WO

0037,030. Chem. Abstr. 133, 78994m]. 2000. Ref Type: Patent

Parkinson, N.M., Conyers, C.M., Keen, J.N., MacNicoll, A.D., Smith, I. and Weaver, R.J.

(2003) cDNAs encoding large venom proteins from the parasitoid wasp Pimpla hypochon-

driaca identified by random sequence analysis. Comp. Biochem. Physiol. C-Toxicol. Pharmacol.

134, 513–520

Pickard, M.A., Roman, R., Tinoco, R. and Vazquez-Duhalt, R. (1999) Polycyclic aromatic hydrocarbon

metabolism by white rot fungi and oxidation by Coriolopsis gallica UAMH 8260 laccase. Appl.

Environ. Microbiol. 65, 3805–3809

476 ALCALDE

Piontek, K., Antorini, M. and Choinowski, T. (2002) Crystal structure of a laccase from the fungus

Trametes versicolor at 1.90-angstrom resolution containing a full complement of coppers. J. Biol.

Chem. 277, 37663–37669

Pruche, F., Saint, L.P. and Bernards, B. Hair dye compositions containing hydroxystilbene. [Eur.Pat.

Appl. EP 1013,260. Chem. Abstr. 133,63587g.]. 2000. Ref Type: Patent

Rodakiewicz-Nowak, J., Kasture, S.M., Dudek, B. and Haber, J. (2000) Effect of various water-miscible

solvents on enzymatic activity of fungal laccases. J. Mol. Catal. B-Enzymatic 11, 1–11

Servili, M., De Stefano, G., Piacquadio, P. and Sciancalepore, V. (2000) A novel method for removing

phenols from grape must. Amer. J. Enol. Viticult. 51, 357–361

Shleev, S., Tkac, J., Christenson, A., Ruzgas, T., Yaropolov, A.I., Whittaker, J.W. and Gorton, L. (2005)

Direct electron transfer between copper-containing proteins and electrodes. Biosen. Bioelectron. 20,

2517–2554

Sigoillot, C., Camarero, S., Vidal, T., Record, E., Asther, M., Perez-Boada, M., Martinez, M.J., Sigoillot,

J.C., Asther, M., Colom, J.F. and Martinez, A.T. (2005) Comparison of different fungal enzymes for

bleaching high-quality paper pulps. J. Biotechnol. 115, 333–343

Tetsch, L., Bend, J., Janssen, M. and Holker, U. (2005) Evidence for functional laccases in the acidophilic

ascomycete Hortaea acidophila and isolation of laccase-specific gene fragments. FEMS Microbiol.

Letters 245, 161–168

Thurston, C.F. (1994) The structure and function of fungal laccases. Microbiology 140, pp. 19–26

Torres, E., Bustos-Jaimes, I. and Le Borgne, S. (2003) Potential use of oxidative enzymes for the

detoxification of organic pollutants. Appl. Catal. B-Environ. 46, 1–15

Ullah, M.A., Bedford, C.T. and Evans, C.S. (2000) Reactions of pentachlorophenol with laccase from

Coriolus versicolor. Appl. Microbiol. Biotechnol. 53, 230–234

Valderrama, B., Oliver, P., Medrano-Soto, A. and Vazquez-Duhalt, R. (2003) Evolutionary and structural

diversity of fungal laccases. Antonie Van Leeuwenhoek 84, 289–299

Widsten, P., Tuominen, S., Qvintus-Leino, P. and Laine, J.E. (2004) The influence of high defibration

temperature on the properties of medium-density fiberboard (MDF) made from laccase-treated

softwood fibers. Wood Sci. Technol. 38, 521–528

Xu, F., Berka, R.M., Wahleithner, J.A., Nelson, B.A., Shuster, J.R., Brown, S.H., Palmer, A.E. and

Solomon, E.I. (1998) Site-directed mutations in fungal laccase: effect on redox potential, activity and

pH profile. Biochem. J. 334, 63–70

Yaropolov, A.I., Skorobogatko, O.V., Vartanov, S.S. and Varfolomeyev, S.D. (1994) laccase - properties,

catalytic mechanism, and applicability. Appl. Biochem. Biotechnol. 49, 257–280

CHAPTER 27

HIGH REDOX POTENTIAL PEROXIDASES

ANGEL T. MARTÍNEZ

∗

Centro de Investigaciones Biológicas, CSIC, Madrid, Spain

∗

atmartinez@cib.csic.es

1. INTRODUCTION

High redox potential peroxidases secreted by some basidiomycete fungi are unique

enzymes that play a central role in the degradation of lignin (Kirk and Cullen,

1998). Plant biomass (bryophytes excluded) is mainly made up of two polysaccha-

rides, cellulose and hemicelluloses, and the complex aromatic polymer of lignin

(Fengel and Wegener, 1984). The main role of lignin in the plant cell wall is

the protection of polysaccharides against the action of hydrolytic enzymes. It also

contributes to stem rigidity and water transport, two important characteristics that

facilitated land colonization by vascular plants. Such characteristics derive from

the recalcitrance and structural complexity of lignin which includes up to three

types of subunit derived from three different p-hydroxycinnamyl alcohols (Higuchi,

1997; Boerjan et al., 2003). These phenylpropanoid units are linked together

by a variety of ether and C-C bonds forming a three dimensional network that

confers lignin its mechanical resistance as well as an extremely high resistance to

degradation.

Biodegradation of the lignin polymer has been described as an “enzymatic

combustion”, where the aromatic units are oxidized by the hydrogen peroxide

secreted by ligninolytic basidiomycetes in a reaction catalysed by high redox

potential peroxidases, enzymes which are unique to this group of organisms (Kirk

and Farrell, 1987). Due to the light colour of the wood decayed by ligninolytic basid-

iomycetes these organisms are also known as white-rot fungi (Martínez et al., 2005).

By contrast, the so-called brown-rot basidiomycetes do not produce ligninolytic

enzymes but instead transform wood into a lignin-enriched brown material. In

addition to peroxidases, other fungal enzymes such as laccases and H

-producing

oxidases are also involved in lignin biodegradation. Ligninolytic enzymes are of

477

J. Polaina and A.P. MacCabe (eds.), Industrial Enzymes, 477–488.

478 MARTÍNEZ

considerable importance for the natural degradation of lignocellulosic materials

in terrestrial ecosystems, enabling the recycling of the organic carbon fixed by

photosynthesis, and also for most industrial processes that utilise lignocellulosic

biomass, including paper pulp manufacturing and bioethanol production both of

which require the removal of lignin to exploit cellulose.

2. LIGNINOLYTIC ENZYMES

Two ligninolytic peroxidases, lignin peroxidase (LiP) and manganese perox-

idase (MnP), were discovered in 1983–84 in the white-rot fungus Phanerochaete

chrysosporium (order Aphyllophorales) and described as “ligninases” because of

their high redox potentials which enable the oxidation of dimeric lignin model

compounds (Tien and Kirk, 1983; Glenn et al., 1983; Kuwahara et al., 1984). LiP is

able to degrade non-phenolic lignin units (up to 90% of the polymer) or the simple

compound 3,4-dimethoxybenzyl (veratryl) alcohol (Tien and Kirk, 1983), whereas

MnP generates Mn

that, when chelated with oxalic or other organic acids, acts as a

diffusible oxidizer of the phenolic lignin units (Glenn et al., 1986) or the non-phenolic

units via lipid peroxidation reactions (Jensen et al., 1996). Versatile peroxidase (VP),

a third type of ligninolytic peroxidase, was more recently described in Pleurotus

and Bjerkandera species (from the orders Agaricales and Aphyllophorales, respec-

tively) (Martínez et al., 1996; Mester and Field, 1998; Ruiz-Dueñas et al., 1999) and

combines the catalytic properties of LiP, MnP and other peroxidases (from plants and

micro-organisms) that oxidise phenolic compounds (Heinfling et al., 1998).

Laccases are known in plants, fungi, bacteria and insects, where they play a

variety of roles including the synthesis of pigments, fruiting-body morphogenesis

and detoxification (Thurston, 1994; Mayer and Staples, 2002) (see chapter by

Alcalde in this book). Laccase production on solid media was considered to be

a characteristic of white-rot fungi (Käärik, 1965), although some brown-rot fungi

produce laccase in liquid cultures (Lee et al., 2004). However, these phenoloxi-

dases have low redox potentials that only permit the oxidation of phenolic lignin

units (often representing less than 10% of the polymer). Recently, biotechnological

interest in laccases has been spurred by the discovery of their ability to oxidize high

redox potential substrates in the presence of synthetic redox mediators, forming

stable free radicals which act as diffusible oxidizers (Bourbonnais and Paice, 1990).

The existence of natural mediators involved in lignin biodegradation has not been

demonstrated despite various attempts (Li et al., 2001), although some lignin-

derived phenols can act as laccase mediators (Camarero et al., 2005).

Other extracellular enzymes involved in lignin degradation are the H

generating oxidases and mycelium-associated dehydrogenases. The former include

aryl-alcohol oxidase (AAO), first described in Pleurotus and Bjerkandera species

(Bourbonnais and Paice, 1988; Muheim et al., 1990; Guillén et al., 1992), and

glyoxal oxidase found in P. chrysosporium and other basidiomycetes (Kersten,

1990). Fungal dehydrogenases, including aryl-alcohol dehydrogenase (AAD), are

also involved in lignin degradation (Gutiérrez et al., 1994; Brock et al., 1995).

HIGH REDOX POTENTIAL PEROXIDASES 479

Ligninolytic peroxidases (LiP, MnP and VP) oxidize the lignin polymer in a

one-electron reaction resulting in cation radical formation (phenoxy radicals from

phenolic units) (Kersten et al., 1985). The aromatic cation radicals formed evolve

through different non-enzymatic reactions resulting in breakdown of ether and C-C

inter-unit linkages, demethoxylation and aromatic ring cleavage (Schoemaker, 1990;

Martínez et al., 2005). The aromatic aldehydes released after oxidative degradation

of the lignin-unit side-chains, or synthesized de novo by fungi (Gutiérrez et al.,

1994), constitute the substrate for the generation of the H

required for lignin

degradation by peroxidases in Pleurotus species. In these and other white-rot fungi

is continuously formed in cyclic redox reactions involving both extracellular

AAO and mycelial AAD (Guillén et al., 1994). Some oxidases and laccases also

participate in the (direct or indirect) reduction of ferric to ferrous iron that reacts

with H

yielding the hydroxyl free radical, a very strong oxidizer involved in

wood biodegradation (Evans et al., 1994; Guillén et al., 2000; Hammel et al., 2002).

Ultimately, simple breakdown products from lignin degradation enter the fungal

hyphae and are incorporated into the intracellular catabolic routes.

3. STRUCTURE AND FUNCTION OF LIGNINOLYTIC

PEROXIDASES

Due to their uniqueness as industrial biocatalysts, the catalytic mechanisms of lignin-

degrading peroxidases have been extensively investigated at the molecular level

(Banci, 1997; Gold et al., 2000; Martínez, 2002). Phanerochaete chrysosporium

LiP and MnP were the second and third peroxidases (after yeast cytochrome c

peroxidase, CCP) whose crystal structures were solved, just ten years after their

discovery (Poulos et al., 1993; Piontek et al., 1993; Sundaramoorthy et al., 1994);

the crystal structure of the Pleurotus eryngii VP has been solved recently (Pérez-

Boada et al., 2005; Piontek et al., 2006). As mentioned above, these peroxidases

are able to catalyse the oxidation of the recalcitrant lignin units by H

. This is

made possible by the formation of a high redox state oxo-ferryl intermediate after

the two-electron reaction of the haem cofactor with H

. The activated enzyme

oxidizes two substrate units and is reduced back to the resting state that reacts

again with peroxide. This catalytic cycle, which includes the resting peroxidase

and the so-called compound I (two-electron oxidized enzyme) and compound II

(one-electron oxidized enzyme), is common to other peroxidases such as the well-

known horseradish peroxidase (HRP) (Dunford, 1999). However, two aspects of

their molecular structure provide the ligninolytic peroxidases their unique catalytic

properties, namely: i) a haem environment conferring high redox potential to the

oxo-ferryl complexes; and ii) specific binding sites (and mechanisms) for oxidation

of their characteristic substrates including non-phenolic aromatic compounds in the

case of LiP, manganous iron in the case of MnP, and both types of compounds in

the case of VP. The hybrid properties of Pleurotus eryngii VP are illustrated in the

basic catalytic cycle shown in Fig. 1A which combines the previously described

cycles of LiP and MnP.

480 MARTÍNEZ

Figure 1. Catalytic cycle of the model ligninolytic peroxidase VP. A, Basic cycle as described by Ruiz-

Dueñas et al. (1999), including two-electron oxidation of the resting peroxidase (VP) by hydroperoxide

to yield compound-I (C-I), whose reduction in two one-electron reactions results in the intermediate

compound-II (C-II) and then the resting form of the enzyme. VP can oxidize both aromatic substrates

(AH) and Mn

. B, Extended cycle adapted from Pérez-Boada et al. (2005), including C-I

and C-II

involved in oxidation of veratryl alcohol (VA) and other high-redox potential aromatic substrates, where

a tryptophan radical is present at position 164 corresponding to LiP W171 (low redox potential aromatic

compounds are probably oxidized by both the A and B forms but for simplicity they are not included)

Similarities in the haem environment (located in the central region of the protein)

between the three ligninolytic peroxidase types were evidenced by

H-NMR and

confirmed by X-ray diffraction of peroxidase crystals (Banci, 1997; Banci et al.,

2003). In NMR solution experiments it is possible to identify the signals of protons

of the haem cofactor as well as those of several amino acid residues in the haem

HIGH REDOX POTENTIAL PEROXIDASES 481

pocket. This is possible due to the paramagnetic effects caused by the haem

iron that displaces the signals of neighbouring protons outside the region where

most protein protons overlap. These signals are better assigned in the spectra of

peroxidase CN-adducts that yield hyperfine NMR signals. One of the main differ-

ences between the various peroxidases is the position of the proximal histidine

side-chain N (acting as the fifth iron ligand) which in the ligninolytic peroxidases

is displace away from the haem (compared to HRP or CCP) contributing to the

electropositivity of the oxo-ferryl complex (Poulos, 1993; Banci, 1997; Martínez,

2002). The NMR results also explain the high redox potential of MnP, despite

that this peroxidase is only able to steady-state oxidize Mn

, and suggest that

this is due to the absence of a specific site in MnP for the oxidation of aromatic

substrates, which is present in LiP and VP. Similarly, LiP lacks the Mn-binding

site described below.

4. SUBSTRATE OXIDATION SITES IN LIGNINOLYTIC

PEROXIDASES

Recent studies based on peroxidase molecular structures solved by crystal X-ray

diffraction are contributing to the identification of the different substrate oxidation

sites in ligninolytic peroxidases. The manganese and aromatic substrate binding-

sites were first identified in MnP and LiP respectively (Doyle et al., 1998; Gold

et al., 2000; Blodig et al., 2001), and then in VP after its crystal structure was

solved. Studies on the molecular structure of VP confirmed that its novel catalytic

properties are due to a hybrid molecular structure, as was suggested several years

ago (Ruiz-Dueñas et al., 1999; Camarero et al., 1999).

oxidation by VP occurs at a binding site similar to that of MnP situated

near one of the propionates of the haem group. The cation is bound by three

acidic residues (one aspartate and two glutamates) enabling direct electron transfer

to the cofactor (Fig. 2, left side). Initially it had been suggested that Mn

oxidized by MnP at the edge of the haem in front of the classical access channel

(-position following NMR nomenclature) (Harris et al., 1991). However, it has

since been demonstrated that this cation uses a second access channel delimited

by the three acidic residues mentioned above, situated in front of the most internal

haem propionate (with respect to the main channel) (Sundaramoorthy et al., 1997).

The functionality of this site in VP was confirmed by site-directed mutagenesis

and the NMR spectra obtained during Mn

titration of the enzyme suggest that a

unique Mn-binding site exists in the vicinity of the VP haem (Banci et al., 2003).

By contrast, it has been shown that veratryl alcohol and other lignin model

substrates are oxidized by the ligninolytic peroxidases at the surface of the protein

by a long-range electron transfer mechanism that initiates at an exposed tryptophan

residue (Fig. 2, right side). The rationale for this mechanism is that most LiP and

VP aromatic substrates, including the lignin polymer, cannot penetrate inside the

protein to transfer electrons directly to the cofactor in the haem pocket. Thus,

they are oxidized at the enzyme surface and the electrons travel to the haem by a

482 MARTÍNEZ

Figure 2. Detail of substrate oxidation sites in the model ligninolytic peroxidase VP including: i) Mn

oxidation site (left) where this cation is bound by the side-chains of E36, E40 and D175 and the internal

propionate of haem that directly receives one electron from the substrate; and ii) high redox potential

aromatic substrate (such as lignin and veratryl alcohol) oxidation site (right) where one electron travels

via a long-range electron transfer pathway formed by an exposed tryptophan residue (W164, in the form

of a tryptophanyl radical in the H

-activated VP) and a leucine residue connected to the haem (axial

view from the proximal side of the haem group)

protein pathway. The tryptophan residue where the protein radical is expected to

be produced was identified in LiP to be W171, which is involved in the oxidation

of veratryl alcohol (Doyle et al., 1998) and a tetrameric lignin model compound

(Mester et al., 2001). The formation of a protein radical was suggested by indirect

evidence (Blodig et al., 1999). The homologous residue in VP (W164) has been

identified, and two other putative long electron transfer pathways proposed for

LiP have been discounted by site-directed mutagenesis studies. Simultaneously,

the existence of a tryptophanyl protein radical was directly detected for the first

time in a ligninolytic peroxidase using low-temperature EPR of H

-activated

VP (Pérez-Boada et al., 2005; Pogni et al., 2005). Moreover, using a combination

of multi-frequency EPR and DFT (density-functional theory) calculations it has

been possible to identify the VP protein radical as a neutral tryptophanyl radical

and confirm that it corresponds to W164 as suggested by site-directed mutage-

nesis (Pogni et al., 2006). On the other hand, it has been found that W171 in

wild LiP (isoenzymes H2 and H8), as well as in Escherichia coli-expressed LiP

treated with several H

equivalents, is C



-hydroxylated (Blodig et al., 1998;

Choinowski et al., 1999, 2001). This finding was related to the formation of the

above-mentioned protein radical, however, no trace of W164 hydroxylation was

found in the atomic resolution structures of recombinant VP treated with H

in wild VP (Pérez-Boada et al., 2005). This result demonstrates that hydroxylation

is not an immediate consequence of the formation of the tryptophan radical in

ligninolytic peroxidases (the evolution of this reactive radical in the absence of

reducing substrate probably depends on its protein environment that is different in

VP and LiP).

Finally, several pieces of evidence suggest that similarly to HRP (Henriksen

et al., 1999) and the low redox potential fungal peroxidase from litter-decomposing

Coprinus species (Tsukamoto et al., 1999), some low redox potential substrates –