Abstract

Cellulose is the main polymeric component of the plant cell wall, the most abundant polysaccharide on Earth, and an important renewable resource. Basidiomycetous fungi belong to its most potent degraders because many species grow on dead wood or litter, in environment rich in cellulose. Fungal cellulolytic systems differ from the complex cellulolytic systems of bacteria. For the degradation of cellulose, basidiomycetes utilize a set of hydrolytic enzymes typically composed of endoglucanase, cellobiohydrolase and β-glucosidase. In some species, the absence of cellobiohydrolase is substituted by the production of processive endoglucanases combining the properties of both of these enzymes. In addition, systems producing hydroxyl radicals based on cellobiose dehydrogenase, quinone redox cycling or glycopeptide-based Fenton reaction are involved in the degradation of several plant cell wall components, including cellulose. The complete cellulolytic complex used by a single fungal species is typically composed of more than one of the above mechanisms that contribute to the utilization of cellulose as a source of carbon or energy or degrade it to ensure fast substrate colonization. The efficiency and regulation of cellulose degradation differs among wood-rotting, litter-decomposing, mycorrhizal or plant pathogenic fungi and yeasts due to the different roles of cellulose degradation in the physiology and ecology of the individual groups.

Introduction

Cellulose is the main polymeric component of the plant cell wall, the most abundant polysaccharide on Earth, and an important renewable resource. The chemical composition is simple, it consists of d-glucose residues linked by β-1,4-glycosidic bonds to form linear polymeric chains of over 10 000 glucose residues. Cellulose contains both highly crystalline regions where individual chains are linked to each other and less-ordered amorphous regions. Although chemically simple, the intermolecular bonding pattern can result in a very complex morphology (Hon, 1994). Basidiomycetes are the most potent degraders of this polymer because many species grow on dead wood or litter, in environment rich in cellulose. Fungal cellulolytic systems differ from the complexed cellulolytic systems of bacteria while the differences between individual taxonomic groups are less pronounced (Lynd et al., 2002). Although cellulose degradation by basidiomycetes has been studied extensively since the middle of the last century, e.g. (Reese & Levinson, 1952), the view of cellulose degradation changed in the last few years. The main reasons were the formulation of the contribution of oxidative systems to cellulose degradation, including the first attempts to quantify their importance (Suzuki et al., 2006), the detection of processive endoglucanases in brown rot fungi (Cohen et al., 2005; Yoon et al., 2007), as well as the appearance of the first genome sequence of a wood-rotting basidiomycete, Phanerochaete chrysosporium (Martinez et al., 2004), that greatly enhanced the power of proteomic and computational methods for detection of individual components of its model cellulolytic system (van den Wymelenberg et al., 2005, 2006; Kersten & Cullen, 2007; Sato et al., 2007). Last, but not least, basidiomycetes from habitats other than wood, for example, the litter-decomposers and mycorrhizal species, attracted more attention in the past years. All of the above achievements contribute to a better picture of the different processes participating in cellulose degradation by basidiomycetes. Previous reviews on cellulose degradation by fungi were usually limited to the description of the properties of enzymatic systems or focused only on one of the several-redox based systems active upon all plant cell wall components. The aim of this review is to present both the information about the composition and biochemical properties of enzymatic systems utilized by basidiomycetous fungi for cellulose degradation and the redox, radical-generating systems, and to point out the main differences. It should be made clear that the degradation of cellulose is a complex process where several components may be acting at the same time. We hope that it will further promote research in the physiology and ecology of lignocellulose degradation.

Degradation of cellulose using hydrolytic enzymes

The classical array of fungal cellulose-degrading enzymes is composed of endo-cleaving (endoglucanases) and exo-cleaving (cellobiohydrolases, exocellulases) enzymes acting on cellulose. The resulting cellobiose or cello-oligosaccharides are usually processed by extracellular or intracellular β-glucosidases or subject to dehydrogenation by cellobiose dehydrogenase.

Endo-1,4-β-glucanase (EC 3.2.1.4, endocellulase)

Endoglucanases (EGs) were isolated from several wood-rotting basidiomycetes, brown rot and white rot fungi and also from the plant pathogen Sclerotium rolfsii, the basidiomycetous yeast Rhodotorula glutinis and the symbiont of termites Termitomyces sp. (Table 1). Because endoglucanase activity was also documented in cultures of litter-decomposing basidiomycetes (Steffen et al., 2007; Valášková, 2007), ectomycorrhizal fungi (Maijala et al., 1991; Cao & Crawford, 1993) and wood-associated yeasts (Jimenez et al., 1991), it seems that this enzyme is common among basidiomycetes.

Table 1

Selected properties of isolated endoglucanases

Fungus and enzyme Group Molecular mass (kDa) pI KM (g L−1pH optimum References 
Coniophora cerebellaBR 42   4.7 Goksoyr & Eriksen (1980) 
Coniophora cerebellaBR 39   4.2 Goksoyr & Eriksen (1980) 
Coniophora puteana BR     Schmidhalter & Canevascini (1992) 
Gloeophyllum sepiarium BR 85    Bhattacharjee et al. (1993) 
Gloeophyllum sepiarium EGS BR 45 3.8 7.6 4.1 Mansfield et al. (1998) 
Gloeophyllum trabeum BR 29  13.1 4.4 Herr et al. (1978a) 
Gloeophyllum trabeum Cel12A BR 28 4.8   Cohen et al. (2005) 
Gloeophyllum trabeum Cel5A BR 42 4.9   Cohen et al. (2005) 
Gloeophyllum trabeum EGT BR 41 3.1 6.3 4.2 Mansfield et al. (1998) 
Fomitopsis palustris BR 40    Ishihara & Shimizu (1984) 
Fomitopsis palustris EG47 BR 47    Yoon et al. (2007) 
Fomitopsis palustris EG35 BR 35    Yoon et al. (2007) 
Irpex lacteus WR 65   4.0 Kanda et al. (1976) 
Irpex lacteus E2-A WR     Kubo & Nisizawa (1983) 
Irpex lacteus E2-B WR     Kubo & Nisizawa (1983) 
Irpex lacteus En-1 WR 36   4.0 Kanda et al. (1980) 
Phanerochaete chrysosporium EG 28 WR 28 5.2   Henriksson et al. (1999) 
Phanerochaete chrysosporium EG 38 WR 38 4.9   Uzcategui et al. (1991a) 
Phanerochaete chrysosporium EG 44 WR 44 4.3   Uzcategui et al. (1991a) 
Piptoporus betulinus EG1 BR 62 2.6–2.8 2.2 2.5 Valášková & Baldrian (2006b) 
Polyporus arculariusWR 39  0.35 4.4–4.6 Ishihara et al. (2005) 
Polyporus arcularius II WR 36  0.26 4.4–4.6 Ishihara et al. (2005) 
Polyporus arcularius IIIa WR 24  0.26 4.9 Ishihara et al. (2005) 
Polyporus schweinitzii WR 45   4.0 Bailey et al. (1969), Keilich et al. (1969) 
Postia placenta BR 35–40    Clausen et al. (1995) 
Rhodotorula glutinis 40 8.6 11 4.5 Oikawa et al. (1998) 
Schizophyllum commune WR 41    Willick & Seligy (1985) 
Schizophyllum commune WR 39    Willick & Seligy (1985) 
Sclerotium rolfsii EG A 78  2.5 4.0 Sadana et al. (1984) 
Sclerotium rolfsii EG B 52  4.8 2.8–3.2 Sadana et al. (1984) 
Sclerotium rolfsii EG C 28  2.2 4.0 Sadana et al. (1984) 
Serpula incrassata Cel 25 BR 25 <3.6  2.5–4.0 Kleman-Leyer & Kirk (1994) 
Serpula incrassata Cel 49 BR 49 <3.6  2.5–4.0 Kleman-Leyer & Kirk (1994) 
Serpula incrassata Cel 57 BR 57 <3.6  2.5–4.0 Kleman-Leyer & Kirk (1994) 
Termitomyces sp. 36  7.5 4.4 Rouland et al. (1988) 
Trametes versicolor WR 30   5.0 Pettersson & Porath (1963), Idogaki & Kitamoto (1992) 
Volvariella volvacea EG1 LD 42 7.7 7.5 Ding et al. (2001) 
Fungus and enzyme Group Molecular mass (kDa) pI KM (g L−1pH optimum References 
Coniophora cerebellaBR 42   4.7 Goksoyr & Eriksen (1980) 
Coniophora cerebellaBR 39   4.2 Goksoyr & Eriksen (1980) 
Coniophora puteana BR     Schmidhalter & Canevascini (1992) 
Gloeophyllum sepiarium BR 85    Bhattacharjee et al. (1993) 
Gloeophyllum sepiarium EGS BR 45 3.8 7.6 4.1 Mansfield et al. (1998) 
Gloeophyllum trabeum BR 29  13.1 4.4 Herr et al. (1978a) 
Gloeophyllum trabeum Cel12A BR 28 4.8   Cohen et al. (2005) 
Gloeophyllum trabeum Cel5A BR 42 4.9   Cohen et al. (2005) 
Gloeophyllum trabeum EGT BR 41 3.1 6.3 4.2 Mansfield et al. (1998) 
Fomitopsis palustris BR 40    Ishihara & Shimizu (1984) 
Fomitopsis palustris EG47 BR 47    Yoon et al. (2007) 
Fomitopsis palustris EG35 BR 35    Yoon et al. (2007) 
Irpex lacteus WR 65   4.0 Kanda et al. (1976) 
Irpex lacteus E2-A WR     Kubo & Nisizawa (1983) 
Irpex lacteus E2-B WR     Kubo & Nisizawa (1983) 
Irpex lacteus En-1 WR 36   4.0 Kanda et al. (1980) 
Phanerochaete chrysosporium EG 28 WR 28 5.2   Henriksson et al. (1999) 
Phanerochaete chrysosporium EG 38 WR 38 4.9   Uzcategui et al. (1991a) 
Phanerochaete chrysosporium EG 44 WR 44 4.3   Uzcategui et al. (1991a) 
Piptoporus betulinus EG1 BR 62 2.6–2.8 2.2 2.5 Valášková & Baldrian (2006b) 
Polyporus arculariusWR 39  0.35 4.4–4.6 Ishihara et al. (2005) 
Polyporus arcularius II WR 36  0.26 4.4–4.6 Ishihara et al. (2005) 
Polyporus arcularius IIIa WR 24  0.26 4.9 Ishihara et al. (2005) 
Polyporus schweinitzii WR 45   4.0 Bailey et al. (1969), Keilich et al. (1969) 
Postia placenta BR 35–40    Clausen et al. (1995) 
Rhodotorula glutinis 40 8.6 11 4.5 Oikawa et al. (1998) 
Schizophyllum commune WR 41    Willick & Seligy (1985) 
Schizophyllum commune WR 39    Willick & Seligy (1985) 
Sclerotium rolfsii EG A 78  2.5 4.0 Sadana et al. (1984) 
Sclerotium rolfsii EG B 52  4.8 2.8–3.2 Sadana et al. (1984) 
Sclerotium rolfsii EG C 28  2.2 4.0 Sadana et al. (1984) 
Serpula incrassata Cel 25 BR 25 <3.6  2.5–4.0 Kleman-Leyer & Kirk (1994) 
Serpula incrassata Cel 49 BR 49 <3.6  2.5–4.0 Kleman-Leyer & Kirk (1994) 
Serpula incrassata Cel 57 BR 57 <3.6  2.5–4.0 Kleman-Leyer & Kirk (1994) 
Termitomyces sp. 36  7.5 4.4 Rouland et al. (1988) 
Trametes versicolor WR 30   5.0 Pettersson & Porath (1963), Idogaki & Kitamoto (1992) 
Volvariella volvacea EG1 LD 42 7.7 7.5 Ding et al. (2001) 
*

BR, brown rot; LD, litter decomposer; P, phytopathogen; S, symbiotic; WR, white rot; Y, yeast.

Substrate, carboxymethyl cellulose.

The enzymes are monomeric, with molecular masses typically between 22 and 45 kDa but enzymes almost double the size were found in Sclerotium rolfsii and Gloeophyllum sepiarium (Sadana et al., 1984; Bhattacharjee et al., 1993). Class 7 endoglucanases produced e.g. by Phanerochaete chrysosporium contain a large catalytic domain and a 4-kDa cellulose-binding domain (CBD) that can be detached by papain cleavage (Uzcategui et al., 1991a), while some smaller endoglucanases lack the CBD (Henriksson et al., 1999). Although some enzymes are not glycosylated, endoglucanases typically contain a relatively low amount of carbohydrate ranging from 1 to 12% (Eriksson & Pettersson, 1975a, b; Kanda, 1976, 1980; Henriksson et al., 1999). Isoelectric points are usually acidic, between 2.6 and 4.9, but the enzymes isolated from Volvariella volvacea and Rhodotorula glutinis exhibited pI above 7 (Oikawa et al., 1998; Ding et al., 2001).

Multiple endoglucanases are produced by many fungi. At least three different enzymes were isolated from Gloeophy-llum trabeum, Phanerochaete chrysosporium, Sclerotium rolfsii and Serpula incrassata. The most studied endoglucanase system in Phanerochaete chrysosporium was originally described to contain five different enzymes (Eriksson & Pettersson, 1975a, b), but some of them were later identified as cleavage products of other endoglucanases. There are two typical endoglucanases with a CBD, EG38 and EG44 belonging to class 7 glycosyl hydrolases (Uzcategui et al., 1991a) and another 28 kDa endoglucanase belonging to family 12. The latter enzyme lacks the CBD; it induces the swelling of filter paper and was proposed to catalyze the fast cleavage of amorphous cellulose regions that are inaccessible to larger endoglucanases (Henriksson et al., 1999). All Phanerochaete chrysosporium enzymes exhibit endo–exo synergism with cellobiohydrolases.

Endoglucanases of basidiomycetes show catalytic optima at pH between 4.0 and 5.0, i.e. near to the pH values found in fungus-colonized wood (Suzuki et al., 2006; Valášková & Baldrian, 2006b). Only the Volvariella volvacea enzyme heterologously expressed in Pichia sp. exhibits a neutral pH optimum (Ding et al., 2002). Temperature optima are between 50 and 70 °C (Kanda et al., 1976, 1980; Herr et al., 1978a, b; Lachke & Deshpande, 1988; Valášková & Baldrian, 2006b), i.e. well above the values occurring under natural conditions.

Carboxymethylcellulose and amorphous cellulose (e.g. phosphoric acid swollen cellulose, PASC) are good – although not natural – substrates of most endoglucanases and indicate that the enzyme activity is mainly directed towards amorphous regions in the cellulose molecule. The KM values for carboxymethylcellulose are in the range from 0.26 in Polyporus arcularius to 13 g L−1 in G. trabeum (Herr et al., 1978a; Ishihara et al., 2005). Significant activity towards crystalline cellulose was found only with the Irpex lacteus 65 kDa cellulase, the Cel5A from G. trabeum and EG35 from Fomitopsis palustris (Kanda et al., 1976; Cohen et al., 2005; Yoon et al., 2007). The latter two enzymes belong to a group of processive endoglucanases, originally reported from cellulolytic bacteria. These enzymes cleave cellulose internally but also release soluble oligosaccharides before detaching from the polysaccharide and thus act as a combination of endoglucanase and cellobiohydrolase (Tomme, 1996; Gilad, 2003). Cel5A from G. trabeum hydrolyze Avicel to cellobiose as the major product while introducing only a small proportion of reducing sugars into the remaining, insoluble substrate. It produced up to 4.5 nmol glucose equivalents from Avicel per minute and milligram protein (Cohen et al., 2005). The other processive endoglucanase EG35 was recently isolated from another brown rot fungus F. palustris, which was previously reported to degrade crystalline cellulose. The partial amino acid sequence of the protein did not show any similarity to known glycosyl hydrolases (Yoon & Kim, 2005, 2007) and it can be considered for sure that there is no corresponding enzyme in the white rot fungus Phanerochaete chrysosporium. In brown rot fungi, processive endoglucanases could potentially substitute for the absence of cellobiohydrolases. However, because most brown rot fungi are reported to be unable to degrade crystalline cellulose the prevalence of these enzymes is questionable. Some processivity was also shown for the EGS and EGT enzymes from G. trabeum and EG1 from Piptoporus betulinus, which liberate cellobiose and glucose from amorphous cellulose, although all of them are inactive with Avicel (Mansfield et al., 1998; Valášková & Baldrian, 2006b).

EG28 from Phanerochaete chrysosporium and endoglucanases from Gloeophyllum spp., Irpex lacteus, Piptoporus betulinus, Sclerotium rolfsii and Trametes versicolor are also active on cello-oligomers, cellotetraose is cleaved to cellobiose; EG1 from Piptoporus betulinus can even produce glucose and cellobiose from cellotriose (Kanda et al., 1980; Lachke & Deshpande, 1988; Idogaki & Kitamoto, 1992; Mansfield et al., 1998; Henriksson et al., 1999; Valášková & Baldrian, 2006b). KM for p-nitrophenyl-β-d-cellobioside (pNPC) is between 7 and 16 mM (Mansfield et al., 1998; Henriksson et al., 1999; Valášková & Baldrian, 2006b).

Cellobiohydrolase (CBH, EC 3.2.1.91; exocellulase)

Cellobiohydrolases have so far been isolated from several white rot basidiomycetes, the plant pathogen Sclerotium rolfsii and from Termitomyces sp. (Table 2). They are apparently absent from most brown rot fungi, and also the genomes of the human pathogen Cryptococcus neoformans and the plant pathogen Ustilago maydis lack the corresponding genes (Loftus et al., 2005; Kamper et al., 2006). Cellobiohydrolase activity was also documented in litter-decomposing fungi (Steffen et al., 2007; Valášková, 2007) and some ectomycorrhizal fungi (Cao & Crawford, 1993; Burke & Cairney, 1998).

Table 2

Selected properties of isolated cellobiohydrolases

Fungus and enzyme Group Molecular mass (kDa) pI KM (mM) pH optimum References 
Coniophora puteana CBH I BR 52 3.6 6.8 5.0 Schmidhalter & Canevascini (1993a) 
Coniophora puteana CBH II BR 50 3.6 4.3 5.0 Schmidhalter & Canevascini (1993a) 
Dichomitus squalens Ex-1 WR 39 4.6  5.0 Rouau & Odier (1986) 
Dichomitus squalens Ex-2 WR 36 4.5  5.0 Rouau & Odier (1986) 
Fomitopsis palustris BR  2.3  4.0 Hishida et al. (1997) 
Irpex lacteus WR 65   5.0 Kanda & Nisizawa (1988) 
Irpex lacteus Ex-1 WR 53 4.5  5.0 Hamada et al. (1999) 
Irpex lacteus Ex-2 WR 56 4.8  5.0 Hamada et al. (1999) 
Phanerochaete chrysosporium CBH 50 WR 50 4.9   Uzcategui et al. (1991b) 
Phanerochaete chrysosporium CBH 58 WR 58 3.8 2.1  Uzcategui et al. (1991b) 
Phanerochaete chrysosporium CBH 62 WR 62 4.9 3.4  Uzcategui et al. (1991b) 
Schizophyllum commune WR 59    Willick & Seligy (1985) 
Schizophyllum commune WR 58    Willick & Seligy (1985) 
Sclerotium rolfsii 42   4.2–4.5 Patil & Sadana (1984) 
Termitomyces sp. 52   4.4 Rouland et al. (1988) 
Fungus and enzyme Group Molecular mass (kDa) pI KM (mM) pH optimum References 
Coniophora puteana CBH I BR 52 3.6 6.8 5.0 Schmidhalter & Canevascini (1993a) 
Coniophora puteana CBH II BR 50 3.6 4.3 5.0 Schmidhalter & Canevascini (1993a) 
Dichomitus squalens Ex-1 WR 39 4.6  5.0 Rouau & Odier (1986) 
Dichomitus squalens Ex-2 WR 36 4.5  5.0 Rouau & Odier (1986) 
Fomitopsis palustris BR  2.3  4.0 Hishida et al. (1997) 
Irpex lacteus WR 65   5.0 Kanda & Nisizawa (1988) 
Irpex lacteus Ex-1 WR 53 4.5  5.0 Hamada et al. (1999) 
Irpex lacteus Ex-2 WR 56 4.8  5.0 Hamada et al. (1999) 
Phanerochaete chrysosporium CBH 50 WR 50 4.9   Uzcategui et al. (1991b) 
Phanerochaete chrysosporium CBH 58 WR 58 3.8 2.1  Uzcategui et al. (1991b) 
Phanerochaete chrysosporium CBH 62 WR 62 4.9 3.4  Uzcategui et al. (1991b) 
Schizophyllum commune WR 59    Willick & Seligy (1985) 
Schizophyllum commune WR 58    Willick & Seligy (1985) 
Sclerotium rolfsii 42   4.2–4.5 Patil & Sadana (1984) 
Termitomyces sp. 52   4.4 Rouland et al. (1988) 
*

BR, brown rot; P, phytopathogen; S, symbiotic; WR, white rot.

Substrate: p-nitrophenyl cellobioside.

Substrate: p-nitrophenyl cellobioside.

§

Substrate: p-nitrophenyl lactoside.

Substrate: p-nitrophenyl lactoside.

Isolated from the termite Macrotermes muelleri, but apparently originating from its fungal symbiont.

The enzymes are monomeric with molecular masses typically between 50 and 65 kDa though Dichomitus squalens and Sclerotium rolfsii cellobiohydrolases are smaller (Rouau & Odier, 1986; Sadana & Patil, 1988b). Papain cleavage of family 7 cellobiohydrolases yields a large domain that is catalytically active with low molecular mass substrates and a 4–5 kDa CBD similar to that of Trichoderma reesei (Uzcategui et al., 1991b). A structural model for family 7 CBH58 from Phanerochaete chrysosporium is available (Munoz et al., 2001). Similar to endoglucanases, glycosylation of cellobiohydrolases is none or low, <12% (Eriksson & Pettersson, 1975a, b; Schmidhalter & Canevascini, 1993a; Hamada et al., 1999); and the isoelectric points are acidic, typically between 3.6 and 4.9 (but only 2.3 in F. palustris).

The necessity for at least two cellobiohydrolases in the ascomycete Hypocrea jecorina (anamorph Trichoderma reesei) has been attributed to their particular preference for the reducing (CBHI) and nonreducing (CBHII) ends of cellulose chains. This notion has also been supported by the exo–exo synergy observed between these two enzymes (Lynd et al., 2002). This is also the probable reason for the multiplicity of cellobiohydrolases detected in most basidiomycetes studied so far (Table 2). Phanerochaete chrysosporium produces three cellobiohydrolases, CBH58 (originally designated CBH I), CBH62 and CBH50 (Uzcategui et al., 1991b). CBH58 and CBH62 liberate cellobiose from reducing ends of cellulose and their CBDs are located at the C terminus while CBH50 cleaves from the nonreducing end and its CBD is located at the N terminus. All combinations of Phanerochaete chrysosporium cellobiohydrolases exhibit considerable synergistic action.

Similar to most endoglucanases, catalytic optima of cellobiohydrolases are situated in a narrow pH range between 4.0 and 5.0. Temperature optima are between 37 and 60 °C depending on the enzyme and the substrate (Rouau & Odier, 1986; Sadana & Patil, 1988b; Hamada et al., 1999). Cellobiohydrolases are typically active on crystalline cellulose, e.g. Avicel. Interestingly, CBH58 and CBH50 from Phanerochaete chrysosporium were not active on carboxymethylcellulose and CBH62 as well as both cellobiohydrolases from Pleurotus ostreatus exhibited only weak activity against this substrate (Uzcategui et al., 1991b; Garzillo et al., 1994). On the other hand, CBHI and CBHII from Coniophora puteana are both active on amorphous cellulose (Schmidhalter & Canevascini, 1993a). Only enzymes acting from the reducing ends are able to liberate cellobiose from pNPC or p-nitrophenyl-β-d-lactoside (pNPL), the KM values are between 2 and 7 mM. Cellobiohydrolases are also active on cellotriose, cellotetraose or higher cellodextrins (Kanda et al., 1989; Schmidhalter & Canevascini, 1993a; Hishida et al., 1997).

Not surprisingly, cellobiose acts as a competitive inhibitor of cellobiohydrolases. The Ki is 1.2–2.4 mM in Coniophora puteana but as low as 65 μM in Phanerochaete chrysosporium (Schmidhalter & Canevascini, 1993a; Igarashi et al., 1998). It is proposed that the positive effect of cellobiose dehydrogenase (CDH) on cellulose hydrolysis is due to relieving the product inhibition of cellobiohydrolases by cellobiose oxidation (Igarashi et al., 1998).

β-Glucosidase (EC 3.2.1.21)

Because cellobiose is a largely available substrate, β-glucosidases are produced by the majority of microorganisms (Lynd et al., 2002). Among basidiomycetes, enzymes were isolated from several wood-rotting fungi, both white rot and brown rot ones, the mycorrhizal fungi Pisolithus tinctorius and Tricholoma matsutake, the plant pathogen Sclerotium rolfsii and Termitomyces sp. (Table 3). β-Glycosidases were also isolated from, and their activity detected in, basidiomycetous yeasts, although some of the wood-associated yeasts were unable to use cellobiose as a substrate (Peciarova & Biely, 1982; Onishi & Tanaka, 1996; Middelhoven et al., 2006). Activity of β-glucosidase was also detected in pure cultures of litter-decomposing basidiomycetes (Steffen et al., 2007; Valášková, 2007) and ectomycorrhizal species (Burke & Cairney, 1998; Mucha et al., 2006).

Table 3

Selected properties of isolated β-glucosidases

Fungus and enzyme Group Molecular mass (kDa) pI KM (μM) pH optimum References 
Ceriporiopsis subvermispora WR 110  390 3.5 Magalhaes et al. (2006) 
Ceriporiopsis subvermispora WR 53    Magalhaes et al. (2006) 
Gloeophyllum trabeum BR 320  41 4.5 Herr (1978a, b) 
Phanerochaete chrysosporium WR 90  160 5.5 Smith & Gold (1979) 
Phanerochaete chrysosporium WR 114  96 4.0–5.2 Lymar et al. (1995) 
Phanerochaete chrysosporium WR 410  110 7.0 Smith & Gold (1979) 
Phanerochaete chrysosporium WR 45 4.7 5300 5.0 Copa-Patino & Broda (1994) 
Phanerochaete chrysosporium WR 116  3350  Igarashi et al. (2003) 
Phanerochaete chrysosporium A1 WR 165 4.8 150 4.0–4.5 Deshpande et al. (1978) 
Phanerochaete chrysosporium A2 WR 172 4.5 150 4.0–4.5 Deshpande et al. (1978) 
Phanerochaete chrysosporiumWR 165–182 4.6–5.2 210 4.0–4.5 Deshpande et al. (1978) 
Phanerochaete chrysosporium BGL1A WR 53  230  Tsukada et al. (2006) 
Phanerochaete chrysosporium BGL1B WR 60  620  Tsukada et al. (2006) 
Piptoporus betulinus 36 2.6 1800 4.0 Valášková & Baldrian (2006b) 
Pisolithus tinctorius 450 3.8 870 4.0 Cao & Crawford (1993) 
Pleurotus ostreatus WR 35 7.5 2300  Morais et al. (2002) 
Pleurotus ostreatus WR 50 7.3 2360  Morais et al. (2002) 
Pleurotus ostreatus WR 66 8.5 2430  Morais et al. (2002) 
Poria vailantii BR    4.2 Sison & Schubert (1958) 
Rhodotorula minuta 144 4.8 1200 4.7–5.2 Onishi & Tanaka (1996) 
Schizophyllum commune WR 94–96    Willick & Seligy (1985) 
Schizophyllum communeWR 110    Lo et al. (1988) 
Schizophyllum commune II WR 96    Lo et al. (1988) 
Sclerotium rolfsii BG1 90 4.1 6500 4.2 Shewale & Sadana (1981) 
Sclerotium rolfsii BG2 90 4.6 7600 4.2 Shewale & Sadana (1981) 
Sclerotium rolfsii BG3 107 5.1 6900 4.2 Shewale & Sadana (1981) 
Sclerotium rolfsii BG4 92 5.6 6400 4.2 Shewale & Sadana (1981) 
Sporobolomyces singularis 146  1960 3.5 Ishikawa et al. (2005) 
Termitomyces sp.   260  Osore & Okech (1983) 
Termitomyces clypeatus 450   5.0 Sengupta et al. (1991) 
Trametes gibbosa WR 640   3.5 Bhattacharjee et al. (1992) 
Trametes versicolor WR 300  276  Evans et al. (1985) 
Tricholoma matsutake LD 160  220 5.0 Kusuda et al. (2006) 
Volvariella volvacea BGL-1 LD 158 5.6 90 7.0 Cai et al. (1998) 
Volvariella volvacea BGL-2 LD 256 5.0–5.2 500 6.2 Cai et al. (1998) 
Fungus and enzyme Group Molecular mass (kDa) pI KM (μM) pH optimum References 
Ceriporiopsis subvermispora WR 110  390 3.5 Magalhaes et al. (2006) 
Ceriporiopsis subvermispora WR 53    Magalhaes et al. (2006) 
Gloeophyllum trabeum BR 320  41 4.5 Herr (1978a, b) 
Phanerochaete chrysosporium WR 90  160 5.5 Smith & Gold (1979) 
Phanerochaete chrysosporium WR 114  96 4.0–5.2 Lymar et al. (1995) 
Phanerochaete chrysosporium WR 410  110 7.0 Smith & Gold (1979) 
Phanerochaete chrysosporium WR 45 4.7 5300 5.0 Copa-Patino & Broda (1994) 
Phanerochaete chrysosporium WR 116  3350  Igarashi et al. (2003) 
Phanerochaete chrysosporium A1 WR 165 4.8 150 4.0–4.5 Deshpande et al. (1978) 
Phanerochaete chrysosporium A2 WR 172 4.5 150 4.0–4.5 Deshpande et al. (1978) 
Phanerochaete chrysosporiumWR 165–182 4.6–5.2 210 4.0–4.5 Deshpande et al. (1978) 
Phanerochaete chrysosporium BGL1A WR 53  230  Tsukada et al. (2006) 
Phanerochaete chrysosporium BGL1B WR 60  620  Tsukada et al. (2006) 
Piptoporus betulinus 36 2.6 1800 4.0 Valášková & Baldrian (2006b) 
Pisolithus tinctorius 450 3.8 870 4.0 Cao & Crawford (1993) 
Pleurotus ostreatus WR 35 7.5 2300  Morais et al. (2002) 
Pleurotus ostreatus WR 50 7.3 2360  Morais et al. (2002) 
Pleurotus ostreatus WR 66 8.5 2430  Morais et al. (2002) 
Poria vailantii BR    4.2 Sison & Schubert (1958) 
Rhodotorula minuta 144 4.8 1200 4.7–5.2 Onishi & Tanaka (1996) 
Schizophyllum commune WR 94–96    Willick & Seligy (1985) 
Schizophyllum communeWR 110    Lo et al. (1988) 
Schizophyllum commune II WR 96    Lo et al. (1988) 
Sclerotium rolfsii BG1 90 4.1 6500 4.2 Shewale & Sadana (1981) 
Sclerotium rolfsii BG2 90 4.6 7600 4.2 Shewale & Sadana (1981) 
Sclerotium rolfsii BG3 107 5.1 6900 4.2 Shewale & Sadana (1981) 
Sclerotium rolfsii BG4 92 5.6 6400 4.2 Shewale & Sadana (1981) 
Sporobolomyces singularis 146  1960 3.5 Ishikawa et al. (2005) 
Termitomyces sp.   260  Osore & Okech (1983) 
Termitomyces clypeatus 450   5.0 Sengupta et al. (1991) 
Trametes gibbosa WR 640   3.5 Bhattacharjee et al. (1992) 
Trametes versicolor WR 300  276  Evans et al. (1985) 
Tricholoma matsutake LD 160  220 5.0 Kusuda et al. (2006) 
Volvariella volvacea BGL-1 LD 158 5.6 90 7.0 Cai et al. (1998) 
Volvariella volvacea BGL-2 LD 256 5.0–5.2 500 6.2 Cai et al. (1998) 
*

BR, brown rot; LD, litter decomposer; M, mycorrhizal; P, phytopathogen; S, symbiotic; WR, white rot; Y, yeast.

Substrate: p-nitrophenyl glucoside.

Intracellular enzyme.

§

Also β-xylosidase activity.

Also 1,3-β-glucosidase activity.

Recombinant protein expressed in Escherichia coli.

††

Substrate: o-nitrophenyl glucoside.

β-Glucosidases isolated so far exhibit high structural variability, partly reflecting the intracellular/extracellular localization of the enzyme (Table 3). The detected molecular masses range from 35 to 640 kDa. While the small enzymes with molecular masses around 100 kDa are monomeric and usually extracellular, homo-oligomeric enzymes have also been isolated. The yeasts Rhodotorula minuta and Sporobolomyces singularis produce dimeric, cell wall-associated β-glycosidases with high affinity for p-nitrophenyl-β-d-glucoside (pNPG) and a transgalactosidase activity (Onishi & Tanaka, 1996; Ishikawa et al., 2005). The enzymes are also able to cleave galactose and fucose units and lactose disaccharides, but the KM for pNPG is the lowest and they can thus be designated as β-glucosidases. The cell wall-associated enzyme from Pisolithus tinctorius is a homotrimer (Cao & Crawford, 1993) and the 450-kDa β-glucosidase from Termitomyces clypeatus is composed of 110-kDa subunits (Sengupta et al., 1991). On the other hand, the large 300-kDa β-glucosidase from Trametes versicolor is a monomeric glycoprotein with 90% glycosylation. The quaternary structure of the large intracellular enzymes from Phanerochaete chrysosporium (410 kDa) and Volvariella volvacea (256 kDa) is not known (Smith & Gold, 1979; Cai et al., 1998). β-Glucosidases are typically glycosylated but the sugar unit content varies (Evans et al., 1985; Lymar et al., 1995). Isoelectric points of cell wall-associated and extracellular enzymes are acidic, typically between 3.5 and 5.2 while the intracellular enzymes have pIs from 6.2 to 7.0.

As already mentioned, β-glucosidases can be extracellular, cell wall-associated and intracellular. The mycelia-associated fraction in Pleurotus ostreatus, Trametes versicolor and Piptoporus betulinus accounted for 65%, 13%, and 35% of the total activity, respectively (Valášková & Baldrian, 2006a). Only 10% of β-glucosidase in Volvariella volvacea cultures was extracellular while 26% were associated with cell wall fraction and 64% were located intracellularly (Cai et al., 1999).

The intracellular localization of some β-glucosidases requires the transport of cellobiose into the cell. Although not proven, it is probable that a permease similar to the Trichoderma reesei diglucoside permease also operates in basidiomycetes (Kubicek et al., 1993). The presence of extracellular and intracellular enzymes is reflected in the production of enzyme molecules differing in structural and catalytic properties. In Phanerochaete chrysosporium, an array of enzymes with β-glucosidase activity was independently isolated by several groups (Table 3). The smallest 45 kDa protein is extracellular, and in addition to pNPG it can also cleave p-nitrophenyl-β-d-xylopyranoside (pNPX) and laminaribiose (β-1,3 linkages). It is almost inactive with polymeric cellulose and xylan (Copa-Patino & Broda, 1994). Three extracellular β-glucosidases were found in cellulose-grown cultures (Lymar et al., 1995). The 114 kDa enzyme contains a CBD that can be detached by papain treatment. The 96 and 98 kDa proteins that do not bind cellulose are probably fragments of this larger molecule. β-Glucosidase activity is competitively inhibited by glucose and gluconolactone, but also by cellobionolactone, the product of CDH oxidation of cellobiose (Lymar et al., 1995). The 116-kDa extracellular enzyme belongs to family 3 glycosidases (Igarashi et al., 2003). It has high affinity for cellobionolactone (KM 1.29 mM) and laminaribiose (2.03 mM) compared with cellobiose (3.35 mM), but the kcat for cellobionolactone is 76 × lower than that for cellobiose. Cellobionolactone produced in high amounts by CDH thus effectively inhibits its activity (Igarashi et al., 2003). An array of monomeric extracellular enzymes with 165–182 kDa were isolated from cultures growing on cellulose; cellobiose induced only the activity of cell-bound enzyme with lower affinity for pNPG –KM 2.0 mM compared with 0.15–0.21 mM reported for the extracellular enzymes (Deshpande et al., 1978). The 410-kDa intracellular enzyme was strongly induced by cellobiose but only weakly by cellulose (Smith & Gold, 1979). Glucose acted as a repressor of β-glucosidase synthesis in Phanerochaete chrysosporium (Smith & Gold, 1979). Two more intracellular β-glucosidases were recently characterized which belong to glycosyl hydrolase family 1 (Tsukada et al., 2006). Recombinant proteins expressed in Escherichia coli have 53 and 60 kDa; BGL1A is also active as β-xylosidase with KM similar to that for pNPG; its crystal structure was recently determined (Nijikken et al., 2007).

As already mentioned for Phanerochaete chrysosporium, also β-glucosidases from brown rot fungi are relatively nonspecific and can also cleave xylose, mannose and galactose units from corresponding oligosaccharides, although with higher KM values (Herr et al., 1978b; Valášková & Baldrian, 2006b). Cello-oligosaccharides are usually good substrates but the enzymes are inactive on crystalline cellulose and exhibit only low activity on amorphous high molecular mass cellulose (Herr et al., 1978b; Sadana et al., 1988; Valášková & Baldrian, 2006b). Activity is competitively inhibited by glucose (Ki 0.2–6 mM), gluconolactone and cellobionolactone. The pH optimum is usually between 3.5 and 5.5, but the intracellular enzyme from Phanerochaete chrysosporium has a neutral pH optimum (Smith & Gold, 1979); the temperature optima are between 45 and 75 °C.

Phosphorolytic degradation of cello-oligosaccharides

Although phosphorolytic degradation is more typical for polysaccharides with α-1,4 bonds, several bacterial species possess enzymes capable of cleaving cellobiose or cellodextrins by Pi-mediated (ATP-independent) phosphorolytic reactions of cellobiose and cellodextrin phosphorylases.  

formula

Owing to intracellular localization of the enzymes, the reaction is not involved in cellulose hydrolysis but is a part of the intracellular utilization of oligosaccharides (Kitaoka & Hayashi, 2002; Lynd et al., 2002). Cellobiose phosphorylase (EC 2.4.1.20) was not screened for in fungi but it was occasionally detected in the white rot root pathogen Heterobasidion annosum. This fungus exhibited better growth on cellobiose than on glucose due to the energy-saving phosphorolysis (Hüttermann & Volger, 1973).

Oxidative decomposition of cellulose

The hypothesis that the decomposition system used by wood-rotting basidiomycetes to degrade plant cell-wall polysaccharides also involves a nonenzymatic component, which was already formulated 40 years ago (Halliwell et al., 1965; Koenigs et al., 1974). The detection of hydrogen peroxide (H2O2) production by several fungi led to the proposal of a degradation pathway based on the Fenton reaction  

formula
This reaction is a well-recognized route of •OH production in biological systems. H2O2 is produced by white rot fungi by the action of enzymes such as glyoxal oxidase (Kersten & Kirk, 1987), glucose oxidase (Kelley & Reddy, 1986), and aryl alcohol oxidase (Guillén, 1990; Muheim et al., 1990), and also by brown rot fungi (Ritschkoff & Viikari, 1991). In addition to free •OH radicals, certain states of hypervalent iron where the radical remains associated with iron were also considered by some authors as potential oxidizing agents (Wood et al., 1994; Branchaud et al., 1999; Welch et al., 2002a, b). Although iron is generally sequestered in redox-inactive complexes in most biological systems to prevent oxidative damage (Halliwell & Gutteridge, 1999), this does not hold for wood, where sufficient iron concentrations make •OH generation feasible (Koenigs et al., 1974), provided that chelators or reductants are available to solubilize the metal. However, because Fe2+ is usually absent in oxygenated environment, there was a question as to how the necessary reduction of Fe3+ is achieved.

Several systems of polysaccharide decomposition have been proposed where enzymes do not directly react with cellulose undergoing oxidative cleavage. Three oxidative systems operated by wood-rotting basidiomycetes have already received sufficient experimental evidence. These include (1) CDH catalysed reactions (2) redox cycling by small-molecular mass quinones or other redox compounds and (3) •OH production catalysed by small glycopeptides (Hammel et al., 2002; Goodell et al., 2003; Tanaka et al., 2007).

The CDH-based decomposition differs from the other oxidative systems in two ways: (1) it depends on the presence of cellobiose and its degradation products and the action is more specific due to enzyme binding to cellulose and (2) in addition to oxidative cleavage of polysaccharides it also transforms cellobiose and cello-oligosaccharides, major products of cellulose hydrolysis.

In all three oxidative systems, •OH is responsible for polysaccharide scission. Hydroxyl radicals can abstract hydrogen atoms from the sugar subunits of cellulose or other polysaccharides with high rate constants around 109 M−1 s−1 (Ek et al., 1989). These reactions produce transient carbon-centred radicals that react rapidly with O2 to give peroxyl radical species. If the ROO• carries a hydroxyl group on the same carbon, it eliminates •OOH (Halliwell & Gutteridge, 1999). If there is no α-hydroxyl group present, the molecule undergoes a variety of oxidoreductions, some of which can result in the cleavage of the cellulose chain (Kirk et al., 1991).

Cellobiose dehydrogenase (CDH; EC 1.1.99.18)

CDH is an extracellular enzyme produced by basidiomycetes and ascomycetes. It efficiently oxidizes cellobiose but also soluble cellodextrins, mannodextrins and lactose to their corresponding lactones using a wide spectrum of electron acceptors including quinones, phenoxyradicals, Fe3+, Cu2+, cytochrome c or triiodide ion (Henriksson et al., 2000; Zamocky et al., 2006).

CDH activity was first discovered as a cellobiose-dependent reduction of quinones in white rot fungi (Westermark & Eriksson, 1974a, b), and the enzyme named cellobiose quinone oxidoreductase (CBQ) carrying a flavin group was isolated from Phanerochaete chrysosporium (Westermark & Eriksson, 1975). Subsequently, a form containing both flavin and heme, that is now named CDH, has been isolated from the same fungus and named cellobiose oxidase (CBO) because it was incorrectly assumed to prefer O2 as an electron acceptor (Ayers et al., 1978). CBQ was later identified as a catalytic active fragment of CBO appearing in the cultures of some fungi probably due to the action of proteases (Wood & Wood, 1992; Henriksson et al., 2000). The FAD and heme prosthetic groups are contained within two separable domains.

Typical CDH produced by basidomycetes is a monomeric protein of 90–110 kDa with glycosylation in the range of 10–20% (Schmidhalter & Canevascini, 1993b; Fang et al., 1998; Baminger et al., 2001); the pI is typically around 4.0 (Table 4). The enzyme has been isolated mainly from white rot basidiomycetes but its activity has also been detected in some other basidiomycetous species (Zamocky et al., 2006). The brown rot fungi do not produce CDH with the only known exception of Coniophora puteana (Schmidhalter & Canevascini, 1993b). It was also isolated from the soil plant pathogen Sclerotium rolfsii and detected in the ectomycorrhizal fungi Pisolithus tinctorius, Suillus variegatus and Cortinarius sp. (Burke & Cairney, 1998).

Table 4

Selected properties of isolated cellobiose dehydrogenases

Fungus Group Molecular mass (kDa) pI KM (μM) pH optimum References 
Coniophora puteana BR 111 3.9 46–84 4.0 Schmidhalter & Canevascini (1993b), Kajisa (2004) 
Irpex lacteus WR 97  34  Hai et al. (2000) 
Phanerochaete chrysosorium WR 89 4.2 16–110 5.0 Henriksson et al. (1995), Zamocky et al. (2006) 
Pycnoporus cinnabarinus WR 92  111 4.5 Sigoillot et al. (2002) 
Pycnoporus cinnabarinus WR 101 3.8  4.5 Temp & Eggert (1999) 
Schizophyllum commune WR 102  30 4.5 Fang et al. (1998) 
Sclerotium rolfsii 101 4.2–5.0 120  Sadana & Patil (1988a); Baminger et al. (2001) 
Trametes hirsuta WR 92 4.2  5.0 Nakagame et al. (2006) 
Trametes pubescens WR 90 4.2 210 4.5–5.0 Ludwig et al. (2004) 
Trametes versicolor WR 97 4.2 120 5.0 Roy et al. (1996) 
Trametes villosa WR 98 4.4 210 4.5–5.0 Ludwig et al. (2004) 
Fungus Group Molecular mass (kDa) pI KM (μM) pH optimum References 
Coniophora puteana BR 111 3.9 46–84 4.0 Schmidhalter & Canevascini (1993b), Kajisa (2004) 
Irpex lacteus WR 97  34  Hai et al. (2000) 
Phanerochaete chrysosorium WR 89 4.2 16–110 5.0 Henriksson et al. (1995), Zamocky et al. (2006) 
Pycnoporus cinnabarinus WR 92  111 4.5 Sigoillot et al. (2002) 
Pycnoporus cinnabarinus WR 101 3.8  4.5 Temp & Eggert (1999) 
Schizophyllum commune WR 102  30 4.5 Fang et al. (1998) 
Sclerotium rolfsii 101 4.2–5.0 120  Sadana & Patil (1988a); Baminger et al. (2001) 
Trametes hirsuta WR 92 4.2  5.0 Nakagame et al. (2006) 
Trametes pubescens WR 90 4.2 210 4.5–5.0 Ludwig et al. (2004) 
Trametes versicolor WR 97 4.2 120 5.0 Roy et al. (1996) 
Trametes villosa WR 98 4.4 210 4.5–5.0 Ludwig et al. (2004) 
*

BR, brown rot; P, phytopathogen; WR, white rot.

††

Substrate: cellobiose.

‡‡

Quinones as electron acceptors; other acceptors can exhibit different optima.

CDH is a typical oxidoreductase with oxidative and reductive half reactions that occur separately (Fig. 1). The oxidative half reaction represents an oxidation in the C1 position of a saccharide; the hemiacetal at this position is converted to a lactone that hydrolyzes spontaneously to a carboxylic acid. The two electrons taken up by the enzyme are later transferred further to one two-electron acceptor, or to two one-electron acceptors (Morpeth et al., 1985; Henriksson et al., 1993). All results indicate that the oxidation of an electron donor is carried out by the FAD group, which is converted to FADH2 (Henriksson et al., 1991). The reduction of cytochrome c and other electron acceptors by CDH is carried out by the heme domain following flavin-to-heme intramolecular electron transfer. However, direct reduction of electron acceptors by the FAD domain of the protein is also possible, although sometimes slower (Henriksson et al., 2000; Zamocky et al., 2006).

Figure 1

Reactions of cellobiose dehydrogenase based on Henriksson (2000). ‘Fe’ represents the heme iron, ‘A’ represents the one-electron acceptor.

Figure 1

Reactions of cellobiose dehydrogenase based on Henriksson (2000). ‘Fe’ represents the heme iron, ‘A’ represents the one-electron acceptor.

The substrate specificity has been investigated most carefully with the Phanerochaete chrysosporium enzyme (Zamocky et al., 2006). CDH readily oxidizes cellobiose and higher cellodextrins, as well as lactose, maltose mannobiose and galactosylmannose, although the latter substrates display 10–100 × higher KM values (Ayers et al., 1978; Morpeth et al., 1985; Bao et al., 1993; Henriksson et al., 1998; Zamocky et al., 2006). These ‘true’ substrates are all di- or oligosaccharides with β-1,4 bonds and a glucose or mannose residue at the reducing end. The monosaccharides glucose and mannose and the 1,4-α-diglucoside maltose have very high KM values. Monosaccharides and maltose also have substantially lower kcat values (Henriksson et al., 2000; Zamocky et al., 2006). The isolated enzymes show KM for cellobiose in the range of 10–200 μM and a pH optimum for quinone reduction between 4 and 5 (Table 4). Temperature optima are typically between 30 and 55 °C (Henriksson et al., 1995; Baminger et al., 2001), but can be as high as 75 °C as in Pycnoporus cinnabarinus (Temp & Eggert, 1999).

CDH was identified as the first nonhydrolytic enzyme binding to cellulose (Renganathan et al., 1990; Henriksson et al., 1991). Unlike many other cellulose-binding proteins, CDH binds specifically to cellulose, i.e. it does not bind to insoluble xylan, mannan, starch and chitin (Henriksson et al., 1997; Temp & Eggert, 1999), and this specificity can be a key to its biological function. In contrast to some CDHs from ascomycetes, basidiomycete enzymes do not possess a typical cellulose binding motif (Hallberg et al., 2002); the binding to cellulose is of a hydrophobic nature (Henriksson et al., 1997).

The biological function of CDH is not fully understood, although Phanerochaete chrysosporium produces relatively high levels of the enzyme, c. 0.5% of the secreted protein. CDH moderately enhances the activity of a crude mixture of cellulases and also of isolated cellobiohydrolase (Bao & Renganathan, 1992; Igarashi et al., 1998). CDH has been shown to degrade not only cellulose, but also xylan and lignin in the presence of H2O2 and chelated Fe ions (Henriksson et al., 1995). It is produced along with cellulases and hemicellulases under cellulolytic conditions, i.e. when cellulose is the major carbon source. It seems likely that CDH is induced by low concentrations of cellobiose, but it is also subject to catabolite repression by excessive concentrations of cellobiose or glucose (Schmidhalter & Canevascini, 1993b; Henriksson et al., 2000). mRNA of CDH was found together with mRNAs for endoglucanase and MnP in Phanerochaete chrysosporium growing on wood (Vallim et al., 1998).

Several hypothetical mechanisms of CDH involvement in the degradation of cellulose (but also hemicelluloses and lignin) have been proposed, e.g. the reduction of substrate inhibition by cellulolysis products, reduction of quinones to be used by ligninolytic enzymes or the support of a Mn-peroxidase reaction (Henriksson et al., 2000). The currently accepted hypothesis is that CDH degrades and modifies cellulose, hemicelluloses and lignin by generating hydroxyl radicals in a Fenton-type reaction (Kremer & Wood, 1992; Henriksson et al., 1995; Mansfield et al., 1997). The enzyme can reduce Fe3+ to Fe2+, or Cu2+ to Cu+ by oxidation of cellobiose. Subsequent reaction between the reduced species and H2O2 generates hydroxyl radicals that may modify and depolymerize plant cell wall polymers. The iron is present in wood and H2O2 is readily produced by CDH itself or by other extracellular fungal redox enzymes (Ander & Marzullo, 1997). Depolymerization with CDH, cellobiose, Fe3+ and H2O2 was demonstrated for carboxymethylated cellulose (CMC), water-soluble xylan, radioactively labelled synthetic lignin (Henriksson et al., 1995) and for insoluble cellulose in the form of kraft pulp (Mansfield et al., 1997). Some depolymerization occurred even without added H2O2 due to its formation by the enzyme itself (Nutt et al., 1997).

A mechanism of CDH participation in the Fenton reaction has been proposed for the brown rot fungus Coniophora puteana (Hyde & Wood, 1997) in wood containing oxalic acid which strongly chelates Fe3+ and Fe2+ (Espejo & Agosin, 1991). The redox properties of Fe-oxalate complexes can have a large influence on Fenton chemistry (Hyde & Wood, 1997; Park et al., 1997, 1999). CDH can reduce Fe3+-oxalate effectively only at pH values below c. 2.5, where it is present as Fe3+-dioxalate, because the reduction potential of the Fe3+-trioxalate complex that predominates above this pH is too negative. The reduction of the Fe3+-dioxalate by CDH results in uncomplexed Fe2+ or in the Fe2+-mono-oxalate complex. These Fe2+ species are relatively stable at pH 2.5, but as they diffuse away from the hyphae, they will encounter a region with lower oxalate concentration and higher pH, which will result in the formation of the Fe2+-dioxalate complex. Around pH 4, Fe2+-dioxalate complexes autoxidize rapidly and as a result peroxyl radical is produced. This •OOH is reduced by Fe2+ or dismutates, thus generating H2O2, the second substrate for Fenton reaction (Hyde & Wood, 1997; Park et al., 1997).

This model proposes that a complete Fenton system is formed only after Fe2+ has diffused some distance from the fungal hyphae. In this way, Fenton reagent might be produced by wood-rotting fungi within the secondary wood cell wall, where it is needed to initiate the degradation of lignin or polysaccharides. This mechanism would also protect the fungus from the oxidative damage that can occur if hydroxyl radicals are produced near the hyphae. The model assumes that H2O2 is produced only via Fe2+ autoxidation. However, if CDH also uses O2 as an electron acceptor, which seems possible given that Fe3+-oxalate complexes are relatively difficult to reduce, then H2O2 will be produced at the same site as Fe2+. In addition, in white rot fungi that secrete H2O2-producing enzymes it is unlikely that H2O2 production could depend only upon Fe2+ autoxidation. Another potential problem with this model is the pH below c. 2.5 required near the hyphae. Although some wood-rotting fungi have been shown to reduce pH as much as to 1.6–2.5 (Green et al., 1991), it is not clear whether these conditions generally occur.

Oxalic acid, the compound essential for the functioning of the proposed mechanism is secreted by many brown rot and white rot fungi (Takao, 1965; Dutton et al., 1993). White rot fungi are usually reported to produce less oxalate than brown rot fungi due to the fact that excess oxalate inhibits the activity of ligninolytic peroxidases (Akamatsu et al., 1994; Shimada et al., 1997). Because a stable concentration of oxalate is necessary for the functioning of the ligninolytic system of white rot fungi, its concentration is regulated by the production of oxalate decarboxylase (Shimada et al., 1997; Kurek & Gaudard, 2000). In addition to the co-operation with CDH, oxalate was also proposed to directly participate in cellulose hydrolysis in brown rot fungi by a Fenton-type mechanism while H2O2 is produced during the synthesis of oxalate (Shimada et al., 1997). Oxalate in a certain range of concentrations greatly enhanced cellulose degradation by Fenton oxidation in vitro (Tanaka et al., 1994) and the wild-type brown rotter Postia placenta caused higher wood mass loss than its less oxalate-producing mutant (Micales & Highley, 1991). However, because it is still unclear how the strict regulation of oxalate concentration can be achieved in vivo it seems more probable that oxalic acid is just a part of one of the more complex systems of cellulose hydrolysis discussed here.

Interestingly, the analysis of Phanerochaete chrysosporium genome has identified, in addition to a CDH gene, a separate gene encoding a heme domain similar to that of CDH fused to a highly conserved family 1 CBD (Kersten & Cullen, 2007). The predicted cytochrome b562 protein is 46% identical to the corresponding region of CDH, and the isolated recombinant enzyme has the expected electron transfer activity (Yoshida et al., 2005). The structure and regulation of this CDH-like protein are compatible with a role in Fenton chemistry similar to CDH (Kersten & Cullen, 2007).

Quinone redox cycling

Low-molecular-weight chelators of catecholate origin were isolated from white rot and brown rot fungi in the late 1980s and early 1990s (Fekete et al., 1989; Jellison et al., 1991; Chandhoke et al., 1992). The research was focused on the chelators produced by G. trabeum (‘Gt chelators’) that were of small molecular size (<1000 Da) and, unlike enzymes, could penetrate through the wood cell wall matrix (Jellison et al., 1991; Goodell et al., 1997; Filley et al., 2002; Goodell et al., 2002). Chelators with iron-reducing capacity have been also documented in rotted wood, e.g. palo podrido (Ferraz et al., 2001). It was discovered later that some of the participating compounds are probably quinones (Goodell et al., 1997; Goodell et al., 2003).

The principle of the quinone redox cycling mechanism is in the fugal reduction of quinones to the corresponding hydroquinones, which then react with Fe3+ to give Fe2+ and semiquinone radicals. The semiquinones can reduce O2 to give •OOH and the original quinones. Because •OOH is a source of H2O2, this cycle will generate a complete Fenton system (Kerem et al., 1999).

This mechanism requires the reduction of extracellular quinones to their hydroquinone forms by the fungus. Enzymes potentially capable of this reaction include intracellular benzoquinone reductases (Brock et al., 1995) and extracellular sugar dehydrogenases such as CDH, which have been shown to use quinones as alternate electron acceptors (Henriksson et al., 2000). This model also assumes an adequate source of quinones. It has been suggested that quinones can be found in wood extractives or generated during lignin transformation by white rot fungi (Guillén, 2000). It was also proposed that demethylated lignin, produced during brown-rot degradation of wood, may also function as a redox active compound, serving as an electron source for Fe3+ to Fe2+ reduction (Xu & Goodell, 2001; Filley et al., 2002). It has also been shown that iron reduction capacity in soils is dependent on the amount of soil organic matter (Keppler et al., 2000). Because the humic portion of soils contains breakdown products of lignin, it seems likely that compounds formed during wood degradation might also promote redox reactions in soils (Goodell et al., 2006).

Because the redox-active quinones and hydroquinones produced in decayed wood are also a subject of •OH damage, it seems likely that the fungus has to provide its own source of extracellular quinones. Their production was originally identified in the brown rot fungi of the genus Gloeophyllum that secrete two hydroquinones, 2,5-dimethoxyhydroquinone (2,5-DMHQ) and 4,5-dimethoxycatechol (4,5-DMC). These compounds are able to reduce Fe3+ to give Fe2+ and semiquinone radicals (Kerem et al., 1999; Paszczynski et al., 1999; Jensen et al., 2001; Newcombe et al., 2002). The semiquinones reduce both O2 and Fe3+, giving peroxyl radicals, additional Fe2+ and the two quinones 2,5-dimethoxy-1,4-benzoquinone (2,5-DMBQ) and 4,5-dimethoxy-1,2-benzoquinone (4,5-DMBQ). In subsequent oxidoreductions, Fe2+/Fe3+ couple equilibrates with the •OOH/O2 couple, while Fe2+ reduces •OOH to give H2O2 (Buettner et al., 1993; Halliwell & Gutteridge, 1999) (Fig. 2). Production of 2,5-DMHQ was also found during the growth of G. trabeum and Postia placenta on wood (Cohen et al., 2002) and during the degradation of cellulose by Serpula lacrymans, another brown rot fungus (Shimokawa et al., 2004). In G. trabeum that produced both 2,5-DMHQ and 4,5-DMC, 2,5-DMHQ/2,5-DMBQ was the more efficient hydroquinone/quinone couple. It was always present in higher concentrations, 2,5-DMHQ also reduced O2 more rapidly and 2,5-DMBQ was more rapidly reduced by the fungus (Jensen et al., 2001; Cohen et al., 2002; Suzuki et al., 2006). In addition to 2,5-DMHQ and 4,5-DMC, other unidentified compounds of phenolic origin were found in wood extracts that can potentially also participate in the redox chemistry (Goodell et al., 1997; Suzuki et al., 2006).

Figure 2

Reactions involved in the quinone redox cycling in the brown rot fungus Gloeophyllum trabeum (shown for 2,5-dimethoxyhydroquinone). Based on Jensen (2002) and Suzuki (2006).

Figure 2

Reactions involved in the quinone redox cycling in the brown rot fungus Gloeophyllum trabeum (shown for 2,5-dimethoxyhydroquinone). Based on Jensen (2002) and Suzuki (2006).

Due to high production of oxalate, pH in G. trabeum cultures can be near 4.1 and even lower values were found near the hyphae in wood (Jensen et al., 2001; Suzuki et al., 2006). 2,5-DMHQ is very stable in the absence of iron at pH 2–4 and it readily reduces Fe3+ with a rate constant of 4.5 × 103 M−1 s−1 at pH 4.0. Fe2+ is also very stable at low pH. H2O2 generation results from the autoxidation of the semiquinone radical and was observed only when 2,5-DMHQ was incubated with Fe3+. At low concentrations of oxalate, around 50 μM, ferric ion reduction and production of •OH is enhanced. The enhancement of both Fe3+ reduction and •OH production may be due to the promotion of the ferric ion solubility by oxalate. On increasing the oxalate concentration the oxalate/ferric ion ratio favours formation of the 2: 1 and 3: 1 complexes and results in slower Fe3+ reduction and •OH formation (Varela & Tien, 2003).

Recently, (Suzuki et al., 2006) tried to quantify the relative importance of quinone redox cycling in the decay of spruce wood by G. trabeum. They found the highest decrease of holocellulose polymerization degree in 1-week-old cultures. Highest concentrations of 2,5-DMHQ and 4,5-DMC (in total 40 μM) were found in the same time interval, but the concentrations decreased rapidly (7 μM in week 5) along with decreasing pH and increasing oxalate concentrations. The rate constants for the reactions of 2,5-DMHQ and 4,5-DMC with the Fe3+-oxalate complexes were determined as 43 and 65 L mol−1 s−1, respectively. Calculations showed that quinone cycling is responsible for a significant fraction of cellulose scissions (in average more than 25%). However, the results also showed that there are also hydroquinone-independent mechanisms for holocellulose cleavage during early decay (Suzuki et al., 2006).

Quinone redox cycling can only be efficient provided the quinones are rapidly reduced to hydroquinones. An intracellular NADH:quinone oxidoreductase capable of 2,5-DMBQ and 4,5-DMBQ reduction was originally isolated from G. trabeum (Jensen et al., 2002). The isolated enzyme has a pI of 3.3 and it is a homodimer of 22 kDa subunits, each with an FMN. The KM and kcat for 2,5-DMBQ and 4,5-DMBQ are 5–7 μM and 1100–1600 s−1, respectively. Because the sugar oxidases that can also use quinones as alternative electron acceptors (Leitner et al., 2001) were present in mycelial extracts, the enzyme is probably responsible for intracellular quinone reduction (Jensen et al., 2002). Later work showed that there are two quinone reductases of which QRD1 is produced during growth on wood when 2,5-DMBQ is present while QRD2 is probably involved in intracellular quinone detoxification (Cohen et al., 2004).

The main obstacle for the functioning of the proposed quinone cycling process can be the intracellular localization of quinone reductases (Fig. 2). For it to be efficient, rapid transfer from and into the fungal hyphae must be possible and the fungus has to deal with the toxic quinones in the cell. Because no quinone reduction was found in cell wall fraction of G. trabeum, one of the alternatives is that the redox potential of native cytoplasmic membrane can be responsible for quinone reduction. However, so far there are no clear indications that such a process really works (Qi & Jellison, 2004).

Extracts from wood colonized by several brown rot fungi showed high iron-reducing capability in a low-molecular-mass fraction (<5000 Da) that was significantly greater than in extracts from wood colonized by white rot or nondecay fungi (Goodell et al., 2006). Although the nature of these compounds is unclear, hydroquinones are among the potential candidates for the detected activity.

Although the relationship to cellulose degradation is not clear, the ability to reduce quinones was also detected in the white rot species. Phanerochaete chrysosporium produces intracellular benzoquinone reductases (Brock et al., 1995) and reduction of quinones by a plasma membrane redox system was also demonstrated in this species (Stahl et al., 1995). Furthermore, quinone redox cycling was demonstrated for Pleurotus eryngii in the context of superoxide anion radical production (Guillén, 1997). Potentially, this could lead to redox cycling-driven Fenton reactions in white-rot fungi.

Glycopeptide-catalysed Fenton reaction

Low molecular mass compounds termed ‘glycopeptides’ were first detected in brown rot fungi in 1980s (Enoki et al., 1989; Tanaka et al., 1991) and identified as compounds capable of catalyzing redox reactions between O2 and electron donors to produce •OH, reduce Fe3+ to Fe2+ and to strongly bind Fe2+ (Enoki et al., 1992; Tanaka et al., 1993, 1996; Hirano et al., 1995, 1997, 2000). These compounds were initially described as iron containing, with c. 22% neutral carbohydrate, 12% protein and a molecular mass of 1.5–5 kDa in F. palustris (Hirano et al., 1995), G. trabeum (Enoki et al., 1992) and Irpex lacteus (Tanaka et al., 1993). Later, larger glycopeptides have been described in white rot fungi Trametes versicolor and Phanerochaete chrysosporium where they were claimed to be the major source of •OH in wood degrading cultures (Tanaka et al., 1999a, b, c) and to act synergistically with phenol oxidases in lignin degradation (Tanaka et al., 1999b; Yamakawa et al., 2005).

The glycopeptide from F. palustris was reported to have a molecular mass between 7.2 and 12 kDa, and contain 54–61% protein (Enoki et al., 2003; Kaneko et al., 2004). Recently, the glycopeptide from Phanerochaete chrysosporium was characterized in more detail (Tanaka et al., 2007). The preparation had a molecular mass of about 14 kDa and contained 25% neutral carbohydrate and 0.04% Fe. Moreover, cDNAs and two putative genes encoding glycoproteins have been sequenced – the 875-bp glp1 and 864-bp glp2. The glycoprotein contained an 1-amino-1-deoxy-2-ketose (ketoamine) produced by the condensation of an amino acid side chain and a carbohydrate. By tautomerization, this structure can yield a 2,3-enediol (Fig. 3) that can reduce Fe3+ to Fe2+ and produce H2O2 from O2 (Oak et al., 2000).

Figure 3

Glycopeptide-catalysed Fenton reaction. Modified from Enoki (2003) and Yamakawa (2005).

Figure 3

Glycopeptide-catalysed Fenton reaction. Modified from Enoki (2003) and Yamakawa (2005).

The size of glycopeptides does not allow them to penetrate the intact wood cell wall (Flournoy et al., 1991), and the reduction of their substrates thus probably occurs close to fungal hyphae although some diffusion into the cell wall was demonstrated (Hirano et al., 2000). To perform the complete catalytic cycle, oxidized saccharidic moieties of the glycopeptides have to be reduced again. This can potentially be performed by a cell wall-associated reductase that is most probably NADH-dependent (Enoki et al., 2003). However, the identity of the electron donor is not yet known.

Degradation of other plant cell wall material by cellulolytic enzymes

Nonspecificity of oxidation by hydroxyl radicals that can lead to modification or cleavage of both polysaccharides and lignin was discussed above. Also, some of the enzymes of the cellulolytic system exhibit broader specificity and can thus act in the degradation of hemicelluloses. Several basidiomycete endoglucanases can also act on hemicelluloses, galactomannan (Keilich et al., 1969), galactoglucomannan (Mansfield et al., 1998), or mannan (Henriksson et al., 1999). Xylan can be cleaved by endoglucanases from G. trabeum, Piptoporus betulinus and Sclerotium rolfsii (Lachke & Deshpande, 1988; Mansfield et al., 1998; Henriksson et al., 1999; Valášková & Baldrian, 2006b). Endoglucanases EG28 and EG38 from Phanerochaete chrysosporium are active on xylan, the latter even with higher kcat than for carboxymethylcellulose. Both of them are inactive on mannan. EG44 is inactive on xylan but degrades mannan with kcat comparable to carboxymethylcellulose (Lawoko et al., 2000). The enzyme Xyn10A firstly isolated as endoglucanase from G. trabeum is actually more active on xylan (Cohen et al., 2005). Among cellobiohydrolases, the enzymes from Dichomitus squalens showed some activity with xylan and o-nitrophenyl-β-d-xylobioside (Rouau & Odier, 1986) while CBH50 and CBH58 from Phanerochaete chrysosporium are inactive on xylan or mannan (Lawoko et al., 2000). The participation of Cellobiohydrolases (and cellulose-binding endoglucanases) in hemicellulose hydrolysis is, however, limited because they specifically bind to cellulose. β-Glucosidases generally exhibit low specificity and are able to cleave mannose, xylose or galactose units from oligosaccharides with varying affinity and the extracellular enzymes can thus participate in hemicellulose degradation.

Saprotrophic basidiomycetes, in particular, those growing on wood or litter, produce a rich array of hemicellulose-degrading enzymes (Baldrian et al., 2008). It can be anticipated that hemicellulolytic enzymes can also exhibit some activity against cellulose and in fact only a few have been explicitly reported not to act on cellulose or cellobiose. Endoxylanase from Termitomyces sp. is also active on carboxymethylcellulose (Faulet et al., 2006) and the endoxylanases XynA and XynC from Phanerochaete chrysosporium slowly cleave pNPC, although they are not active on carboxymethylcellulose (Decelle et al., 2004). Some endoglucanase activity was also demonstrated for G. trabeum endoxylanase (Ritschkoff et al., 1994). Similar to β-glucosidases, a broader substrate range can be also anticipated in β-glycosidases of other basidiomycetes. β-Galactosidase of Cryptococcus laurentii can be an example of enzyme active on pNPG and cellobiose (Ohtsuka et al., 1990).

Enzymatic vs. oxidative degradation of cellulose

Current results indicate that wood-rotting fungi possess two independent types of systems capable of internal scission of cellulose molecules – enzymatic and radical-based ones. Probably all wood-rotting fungi produce endoglucanases and a radical-based system. The radical-based systems share several features: they are based on redox chemistry and the produced hydroxyl radicals nonspecifically cleave cellulose and hemicelluloses and also modify (or perhaps also cleave) lignin. The CDH system is different in that redox reactions with cellobiose can occur extracellularly and are not dependent on NADH or other energy equivalents. This system is, however, dependent on the production of cellobiose and thus dependent on cellobiohydrolase. This is the main reason why brown rot fungi except Coniophora sp. cannot use it. Owing to the diffusible nature of oxidants produced, cleavage by radical-based systems can occur at larger distances from hyphae. If the pH near hyphal surfaces is low, generation of hydroxyl radicals occurs only at some distance with a higher pH value. The endocleavage of cellulose by endoglucanases is specific for cellulose, energy independent and, due to the enzyme size, it probably occurs immediately near hyphae. Moreover, a significant part of cellobiohydrolase which continues cellulose hydrolysis is localized in association with fungal hyphae (Valášková & Baldrian, 2006a).

In the case of the brown rot fungus Postia placenta, endoglucanases were only active on cellulose after oxidative pretreatment (Ratto et al., 1997), which suggests that the radical generation precedes the action of endoglucanases. This is in agreement with the fact that quinone redox cycling that occurs in this species is most active during initial wood degradation (Suzuki et al., 2006). On the other hand, CDH and endoglucanase are produced concomitantly (Henriksson et al., 2000). There is also as yet no explanation as to why some fungi produce more than one oxidative system; Phanerochaete chrysosporium apparently produces both CDH and glycopeptides and G. trabeum produces redox-active quinones and glycopeptides.

The answer to the question of why white rot fungi produce endoglucanases and do not use the combination of radical-based cellulose scission and cellobiose production by cellobiohydrolase lies probably in the differences in localization of degradation processes. We propose that enzymatic hydrolysis of cellulose has mainly or exclusively a nutritive role because it is localized near the hyphae. Endoglucanases probably generate more cuts in the cellulose chains accessible to cellobiohydrolases than do the nonspecific oxidative systems. The main role of the radical-based mechanisms can thus be in the structural degradation of wood to promote fungal colonization and resource capture. This is in agreement with the fact that •OH production during the growth of brown rot fungi on wood or cellulose is directly proportional to substrate mass loss (Kaneko et al., 2005). Where present, processive endoglucanases are involved in oligosaccharide liberation by brown rot fungi.

Future perspectives in the research on cellulose degradation by basidiomycetes

The research on cellulose degradation by basidiomycetes that has continued for several decades has reached considerable achievements in the description of the degradation system of the white rot fungus Phanerochaete chrysosporium as well as in the significant contribution to the mechanisms of brown rot decay. There are, however, still many fields where more efforts are needed in order to increase the understanding of the composition of cellulolytic systems, their regulation and ecological significance. One of the most important questions is how the brown rot fungi cope with the degradation of crystalline cellulose. It is not clear as to how abundant is the production of processive endoglucanases and which mechanisms of carbon acquisition from cellulose are used by the species unable to produce these enzymes. The finalization of the Postia placenta genome project in the near future will hopefully bring the possibility of highly sensitive proteomic analysis of the cellulolytic system of brown rot fungi and help us to answer some of the above questions.

The understanding of the mechanisms of radical-based cellulose degradation should be extended by the study of the relative importance of enzymatic/oxidative degradation and their physiological and ecological significance. Such quantitative studies would, however, require a very complex experimental setup.

Compared with wood degradation, the research on cellulose degradation by other groups of basidiomycetes, e.g. the soil saprotrophic species is far less developed. There are as yet no isolated enzymes, not to speak about the functioning of a complete cellulolytic system in the model species. The potential role of radical-generating mechanisms in soil saprotrophs remains to be addressed as well as the existence of nonligninolytic ‘brown rot type’ basidiomycetes in soils. To understand the contribution of basidiomycetes to the carbon cycling in the environment, the contribution of different fungal/microbial groups to the utilization of cellulose in different habitats should be quantified. This is impossible without linking specific producers to measured enzyme activities in natural substrates, e.g. wood, litter or soils, perhaps by transcriptome and proteome analyses. The development of methodology in the last few years gives us hope that the study of cellulose degradation by basidiomycetes in the future will offer a challenging work that will greatly increase our knowledge in the fields ranging from fungal physiology to ecosystem processes.

Acknowledgements

This work was supported by the Ministry of Agriculture of the Czech Republic (QH72216), by the Ministry for Education, Youth and Sports of the Czech Republic (LC06066) and by the Institutional Research Concept No. AV0Z50200510 of the Institute of Microbiology of the ASCR, v.v.i.

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Editor: Jiri Damborsky