Biodegradation of lignin

Biodegradation of lignin Ian D. Reid

Can. J. Bot. Downloaded from www.nrcresearchpress.com by Renmin University of China on 06/06/13 For personal use only.

Abstract: Lignin is an aromatic polymer forming up to 30% of woody plant tissues, providing rigidity and resistance to biological attack. Because it is insoluble, chemically complex, and lacking in hydrolysable linkages, lignin is a difficult substrate for enzymatic depolymerization. Certain fungi, mostly basidiomycetes, are the only organisms able to extensively biodegrade it; white-rot fungi can completely mineralize lignin, whereas brown-rot fungi merely modify lignin while removing the carbohydrates in wood. Several oxidative and reductive extracellular enzymes (lignin peroxidase, manganese peroxidase, laccase, and cel1obiose:quinone oxidoreductase) have been isolated from ligninolytic fungi; the role of these enzymes in lignin biodegradation is being intensively studied. Enzymatic combustion, a process wherein enzymes generate reactive intermediates, but do not directly control the reactions leading to lignin breakdown, has been proposed as the mechanism of lignin biodegradation. The economic consequences of lignin biodegradation include wood decay and the biogeochemical cycling of woody biomass. Efforts are being made to harness the delignifying abilities of white-rot fungi to aid wood and straw pulping and pulp bleaching. These fungi can also be used to degrade a variety of pollutants in wastewaters and soils, to increase the digestibility of lignocellulosics, and possibly to bioconvert lignins to higher value products. Key words: delignification, white-rot fungi, biobleaching, lignin peroxidase, manganese peroxidase, laccase.

RCsumC : La lignine est un polymbre aromatique qui constitue jusqu'i 30% des tissus ligneux des plantes, donnant de la rigidit6 et de la resistance a l'attaque biologique. Parce qu'elle est insoluble, chimiquement complexe, et sans lien hydrolysable, la lignine est un substrat difficile pour la dtpolymCrisation enzymatique. Seuls certains champignons, surtout des basidiomycbtes, peuvent la biodtgrader de f a ~ o nextensive; les champignons i pourriture blanche peuvent complbtement minCraliser la lignine, tandis que les champignons a pourriture brune la modifient 1Cgbrement tout en s'attaquant aux hydrates de carbone du bois. On a isole plusieurs enzymes extracellulaires, oxydatives (lignine peroxydase, manganbse peroxydase, laccase) et rCductives (cel1obiose:quinone oxydorCductase), partir des champignons ligninolytiques; le r61e de ces enzymes dans la biodigradation de la lignine fait l'objet de nombreuses Ctudes. La combustion enzymatique, un processus par lequel des enzymes gCnttrent des intermediaires rkactifs, sans diriger directement les reactions qui mbnent a la dCsintCgration de la lignine, a CtC suggCrCe cornme le mecanisme de la biodkgradation de la lignine. Parmi les consCquences Cconomiques de la biodegradation de la lignine, se trouvent la dCcomposition du bois et le recyclage biogCochimique de la biomasse ligneuse. On essaie de mettre les pouvoirs dklignificateurs des champignons a pourriture blanche au service d'une part de la fabrication des pltes a partir du bois et de la paille, et d'autre part, au service du blanchiment des pltes chimiques. Ces champignons sont aussi utiles pour dCgrader des polluants divers dans les eaux usCes et les sols, pour augmenter la digestibilitC des matCriaux lignocellulosiques, et possiblement pour la biovalorisation des lignines. Mots cltfs : dklignification, champignons a pourriture blanche, bioblanchiment, lignine peroxydase, manganbse peroxydase, laccase.

Lignin Lignin is the material that confers the qualities of rigidity and durability that make woody plants "woody." It makes u p about 3 0 % of softwood and about 2 0 % of hardwood, and is

found in the

in a

with

and

hemicellulosic polysaccharides (1). In this natural composite material, the cellulose fibrils provide tensile strength, and the hemicellulose and lignin provide cross-linlung, binding the

Pructure together. Lignin is biosynthesized by the polymerization of phenyl-

id (2). There are three of these precursors, .~ r o ~ a n oDrecursors A

Received August 15, 1994.

I.D. Reid. Pulp and Paper Research Institute of Canada, 570 St. John's Boulevard, Pointe-Claire, QC H9R 359, Canada.

, ,

differing in the number of methoxyl groups on the aromatic ring (Fig. 1). Softwood lignin contains mostly guaiacyl (monomethoxy) units. Hardwood lignin contains roughly equal amounts of guaiacyl and syringyl (dimethoxy) units. Grass lignins contain p-hydroxyphenyl (no methoxy 1) units a s well as the other two types. Lignin polymerization takes place in

Can. J. Bot. 73(Suppl. 1): S1011-S1018 (1995). Printed in Canada I Imprim6 au Canada

Can. J. Bot. Vol. 73 (Suppl. I ) , 1995


Fig. 1. Structures of the three lignin precursors, showing the conventional labelling of the carbons in lignin subunits.

cell walls after the polysaccharides have been deposited, and is initiated by enzymatic oxidation of the precursors to phenoxy radicals. These radicals can couple with each other and with the growing lignin polymer in numerous ways to form a complex cross-linked network (Fig. 2). The most common interunit linkage is an ether bond formed between the middle carbon of a propanoid side chain and the phenolic group of another unit (/3-0-4 bond). The benzyl (a) carbon of the side chain can also participate in ether bonds, and diary1 ethers form at lower frequency. Direct C-C bonds are formed by coupling between the /3-carbon and the 5- or 6-positions of a ring, another /3-carbon, or the 1-position of a ring (displacing the propanoid side chain). Direct C -C bonds also form between the 5-positions of two rings. Because the 5-position of syringyl units is occupied by a methoxyl group, these units cannot participate in any of the linkages involving that position. Some interunit linkages occur together, such as the combination of the /3-5 and a - 0 - 4 links to form a phenylcoumaran structure or the combination of a /3-/3 bond with 2 a - 0 - y links to form a pinoresinol structure. Occasional ether bonds to sugars in hemicelluloses are also present. Polymerization of the lignin precursors can be carried out in vitro to produce synthetic lignins, or dehydrogenative polymerizates (DHP) (3). Compared with natural lignins, DHPs have the advantages of being readily prepared free from carbohydrate, being soluble in some solvents, and being easily labelled with I4C or I3C. For these reasons DHPs have been used extensively in lignin biodegradation research, even though they are smaller and have different frequencies of interunit linkages than natural lignins. For detailed studies of the effects of lignin-degrading organisms and enzymes on the various types of interunit linkages, lignin model dimers have been used (4). Dimers containing the /3-0-4 and /3-1 linkages have received the most attention. Experiments with these models have shown the variety of reactions that can occur and led to the discovery of the extracellular peroxidases of white-rot fungi, but the task of demonstrating the relevance of the results obtained with dimers to the degradation of polymeric lignin has barely begun (5).

Lignin-degrading fungi The only organisms known to extensively degrade lignin are fungi (6, 7). For detailed recent reviews of lignin biodegradation, see Refs. 8 - 10. The white-rot fungi, the most effective lignin degraders, are basidiomycetes (1 I). They occur predominantly on hardwoods and, in the laboratory, they generally decay hardwood better

than softwood, apparently because of the presence of syringyl lignin in the hardwood (12, 13). White-rot fungi are able to completely mineralize both the lignin and carbohydrate components of wood. Some species (simultaneous rots) remove lignin and carbohydrate at the same proportional rate, others (selective white rots) remove lignin faster than cellulose (14). Many, but not all, white-rot fungi produce phenol oxidase activity in culture (15, 16). In this context, phenol oxidase activity is used loosely, to indicate the formation of extracellular coloured products from phenolic substrates; extracellular peroxidases as well as laccase may contribute to the reaction. Brown-rot fungi, in contrast to white rotters, depolymerize and remove the polysaccharides from wood while leaving the lignin as a crumbly brown residue. The lignin in brownrotted wood has been demethylated, and oxidized and depolymerized (17, 18), but not completely degraded. Brown rot is most common in softwood. Brown-rot fungi are basidiomvcetes. and are classified in the same families as white-rot genera. The brown-rot fungi, which generally lack phenol oxidase activity, apparently evolved from the whiterot fungi (11). A third type of wood decay is soft rot, caused by ascomycetous fungi (19). These fungi are thought to preferentially degrade the carbohydrates of the wood, although some of them have demonstrated significant ability to mineralize lignin (20). Unfortunately, lignin degradation by soft-rot fungi has been little studied.

Mechanism of lignin biodegradation Because lignin is an insoluble polymer, the initial steps in its biodegradation must be extracellular. The final steps in lignin mineralization, culminating in the release of COz, are likely to take place inside the fungal hyphae. Therefore, the extracellular reactions must break lignin into fragments that are able to diffuse to the hyphae and cross the cell membranes. Unlike other biopolymers, the monomers in lignin are joined by ether and C -C bonds which are not readily hydrolyzed. Chemical and physiological evidence shows that lignin degradation, like its biosynthesis, is predominantly oxidative, although reductive reactions may also participate (21). Oxidation of lignins that contain free phenolic groups would be expected to cause further polymerization and, indeed, this is the easiest effect to demonstrate in vitro (22). However, some low molecular weight fragments are also released (23, 24). Ligninolytic fungi seem to be able to tip the balance between polymerization and depolymerization in favour of fragmentation, possibly by removing the low molecular weight pieces from the reaction mixture (5) or by glycosylation (25) or methylation (21). DHPs added to ligninolytic cultures can be transiently polymerized before being depolymerized and mineralized (26, 27). Because of the random polymerization process that formed it, lignin has a complex and irregular structure. The diversity of the interunit linkages, magnified by the presence of both enantiomeric forms at the asymmetric a - and /3-carbons, and the irregularity of their arrangement make it difficult for a ligninolytic fungus to produce enzymes that could recognize and cleave all of them. The solution that has apparently evolved in the white-rot fungi is to produce enzymes of low specificity that initiate, but do not direct, oxidative reactions in lignin. Kirk and Farrell (6) have termed this process

F5.2. Reid


Fig. 2. Schematic structure of a portion of a guaiacyl lignin molecule.

enzymatic combustion: the enzyme activates the lignin to overcome an energy barrier and begin a thermodynamically favoured oxidative fragmentation without further control of the reaction pathway by the enzyme. Water-soluble intermediates of lignin degradation can be detected under either submerged or solid-state fermentation conditions (28-35). The water-soluble intermediates in the degradation of DHP have a broad molecular size distribution and are predominantly oligomeric (27). Partially degraded lignins can also be extracted from decayed wood with polar organic solvents (21, 36). Much of the extracted material is polymeric or oligomeric (37). Characterization of these polymers, and of products from DHPs specifically labelled with 13C(38), show that the chemical changes produced by whiterot fungi in the lignin polymer include oxidative cleavage of the propanoid side chains between the a- and /3-carbons and between the /3- and y-carbons, and demethylation and oxidative cleavage of aromatic rings (21). Lignin biodegradation does not proceed by an orderly removal of the peripheral subunits as single ring compounds; it also involves oxidation of the aromatic rings and side chains in the interior of the polymer, increasing the hydrophilicity and solubility of the polymer core at the same time as fragments of varying size are set free. The disorderly nature of this degradation agrees with the concept of enzymatic combustion.

Lignin-degrading enzymes A set of enzymes that will extensively degrade native lignin

and reproduce the effects of a white-rot fungus has not yet been identified. However, one reductive and three oxidative enzymes are commonly found extracellularly in ligninolytic cultures of white-rot fungi. These enzymes are active on lignin model dimers, and are generally believed to play a role in lignin biodegradation. Collecting irrefutable evidence for such a role, and identifying the agents still missing from a

complete ligninolytic enzyme system are the current challenges for lignin biodegradation research. It is already clear that different combinations of the known enzymes are produced by various lignin-degrading fungi (15), suggesting that there is more than one successful strategy for lignin biodegradation. Lignin peroxidase Once called ligninase, this enzyme is a heme peroxidase with an unusually high redox potential and low optimum pH (39-41). It shows little substrate specificity, reacting with a wide variety of lignin model compounds and even unrelated molecules (42). It has the distinction of being able to oxidize methoxylated aromatic rings without a free phenolic group, generating cation radicals that can react further by a variety of pathways, including C,-Co cleavage and ring opening. First discovered in Phanerochaete chrysosporium (43, 44), it is produced by many, but not all, white-rot fungi (15, 16). The enzyme is susceptible to inactivation by H202( 4 9 , and to inhibition or masking of its action by components of lignocellulosic media (46). Veratryl alcohol enhances its action on many substrates, including lignin (5, 24); the effect of veratry1 alcohol has been attributed to acting as a mediator (47), protecting against inactivation (48), or acting as a preferred substrate for the second one-electron reduction in the enzyme's catalytic cycle (49). The crystal structure of lignin peroxidase is known (50, 51). It is able to oxidize ferrocytochrome c (12 300 Da), suggesting that the enzyme could act directly on polymeric lignin (52). Lignin peroxidase can depolymerize DHP in vitro, if the DHP is presented in low concentration in a reaction mixture containing 30% or more of an organic solvent (5). Manganese peroxidase This is another heme peroxidase, but it shows a strong preference for Mn(I1) as its reducing substrate (53). The product


Can. J . Bot. Vol. 73 (Suppl. I ) , 1995

Mn(II1) forms a complex with organic acids and diffuses away from the enzyme to oxidize other materials, such as lignin. The redox potential of the Mn peroxidase - Mn system is lower than that of lignin peroxidase and it does not oxidize nonphenolic lignin models. Phenolic substrates are oxidized to phenoxy radicals, which can react further by demethylation, alkyl-phenyl cleavage, C, oxidation, or C,-CB cleavage (54). Manganese peroxidase is very widespread among whiterot fungi (15, 16) and is readily detected in cultures on real lignocellulosic substrates (55-57). It can depolymerize DHP in vitro (58).

Laccase This enzyme is a copper-containing oxidase; it does not require peroxide (59). Like Mn peroxidase, it normally oxidizes only those lignin model compounds with a free phenolic group, forming phenoxy radicals. However, in the presence of the artificial substrate 2,2'-azinobis(3-ethylbenzthiazoline5-sulphonate) (ABTS), laccase can also oxidize certain nonphenolic compounds (60), probably by hydrogen abstraction from benzyl carbons (6 1). ABTS also enhances the ability of laccase to degrade the residual lignin in kraft pulps (62); other synthetic mediators reportedly have a similar effect (63). Laccase is produced by most white-rot fungi (15, 16), with the notable exception of P. chrysosporium (6). Cel1obiose:quinone oxidoreductase (CBQase) This heme-flavin enzyme oxidizes cellobiose and some other carbohydrates, and reduces quinones and the radicals produced by the action of lignin peroxidase, Mn peroxidase, and laccase on lignin model compounds (64). It can also reduce Fe(II1) ions and ferricytochrome c. Suggested roles for CBQase in lignin biodegradation include preventing oxidative polymerization of phenols (65), supporting the production of hydroxyl radical by a Fenton cycle (66), and promoting the action of MnP by creating complexing agents for Mn(I1I) and by reducing Mn02 (67). Experimental evidence that CBQase enhances lignin degradation in an enzymatic or fungal system is still lacking. Other enzymes The activity of extracellular peroxidases depends on a supply of H202. Glucose-1 -oxidase (68), glucose-2-oxidase (69), glyoxal oxidase (70), aryl alcohol oxidase (l6), and methanol oxidase (71) have all been proposed as H202sources. Sugars needed as substrates for the glucose oxidases can be produced by the action of cellulase and hemicellulases on wood polysaccharides. Methanol can be released from methoxyl groups during lignin degradation (56). Glyoxal, aryl alcohols, and ethanol are excreted metabolites of white-rot fungi (16,70,72). A cycle involving intracellular reduction of the aldehyde products of aryl alcohol oxidase has been proposed (16, 73). Intracellular metabolism also supplies oxalate (74) and other organic acids which can chelate Mn(II1) (75). Two new types of extracellular peroxidase have been reported from Bjerkandera sp. and Junghuhnia separabilima (76, 77); the roles they might play in lignin biodegradation are still unknown. Enzyme access to lignin Native lignin is hydrophobic, insoluble, and sequestered in a dense matrix of polysaccharides, which restricts access of

enzymes to only the surface of the wood cell wall (78, 79). Physical contact between enzymes and lignin might be a ratelimiting factor in lignin biodegradation. For Mn peroxidase this may be less of a problem; complexed Mn(II1) ions can more readily diffuse into the lignocellulose complex than proteins can. For other enzymes, this problem seems to be resolved in two ways, depending on the type of decay. During simultaneous decay, the porosity of the cell wall does not increase enough to allow enzyme penetration into the ligninpolysaccharide matrix (78, 80). However, in this type of decay, polysaccharides are removed from the wall at the same rate as lignin, so that the zone of active lignin degradation remains directly accessible at the surface. In selective delignification, the cellulosic matrix of the secondary wall remains in place and lignin degradation takes place at a considerable distance from the surface of the fiber lumen. Irnrnunochemical labelling shows that lignin peroxidase can penetrate the selectively delignified cell walls (80), indicating that removal of lignin and hemicellulose creates channels wide enough to allow access of the enzymes to the sites of lignin degradation. Thus, in either case, the fungal enzymes can reach their substrate.

Economic consequences of lignin biodegradation Wood decay After cellulose, lignin is the second most abundant organic compound on earth; lignocellulose is the major storehouse of the carbon fixed by photosynthesis. Fungal lignin biodegradation is a key process in the biogeochemical cycling of this carbon, and this decomposer activity may be the most valuable service of lignin-degrading fungi to mankind (81). However, wood decay is sometimes inconvenient to us, when it occurs in timber that we want to harvest or lumber that we are using. It is estimated that 10% of the timber cut in the United States each year is to replace wood that decays in service (82). Pulping Making wood into pulp requires separating the fibers from each other and making them flexible, either by dissolving out the lignin in chemical pulping or by physically tearing apart the lignified fibers in mechanical pulping (83). Treating the wood with selectively lignin-degrading fungi decreases the energy input required for mechanical pulping (84, 85) and also improves the bonding ability of the fibers, leading to stronger paper. A similar treatment can also be used with rice straw (86) or bagasse (87). Biological delignification of wood is also reported to aid subsequent chemical pulping by the sulfite (88) and kraft (89) processes. Bleaching Kraft pulps contain up to 5 % of modified, dark-coloured residual lignin. Certain white-rot fungi can solubilize much of this lignin, decreasing the amount of chemicals required to bleach the pulp (90). Effective fungi include Phanerochaete chrysosporium (9 1), Phanerochaete sordida (92), Trametes versicolor (93), and several unidentified isolates (57, 94, 95). Even though direct physical contact between fungus and pulp is not required for limited delignification (92, 96), extensive bleaching by a cell-free enzyme system has been difficult to demonstrate. Pulp brightening is correlated with Mn peroxidase activity over a range of isolates and culture conditions

F5.2. Reid


(56, 57, 92, 93, 97). Manganese peroxidase (56), laccase plus ABTS (62), or other mediators (63) can partially reproduce the delignifying effect of the fungus and substantially increase the bleachability of kraft pulps. Recently Kondo et al. (98) have demonstrated that Mn peroxidase can also brighten kraft pulps if the surfactant Tween 80 is present. Either the detergent action of Tween 80 or the unsaturated fatty acids it contains (99) may be responsible for the enhancement of Mn peroxidase's bleaching effect.

Pollution abatement The low specificity and strong oxidative abilities of fungal lignin degradation systems allow them to be applied to the degradation of many organic pollutants. White-rot fungi or their enzymes can dechlorinate and decolorize the chlorolignin in effluents from chlorine bleaching of chemical pulps (100- 102). Selected white-rot fungi can also be used for bioremediation of soils contaminated with a wide variety of aromatic pollutants (42, 103). Digestibility enhancement Lignification protects polysaccharides from hydrolytic enzymes and limits the digestibility of lignocellulosic materials by ruminant animals or by cellulases applied in vitro. Delignification of the raw materials by solid-state fermentation with selective white-rot fungi increases their value as fodder for ruminant animals and as substrates for enzymatic saccharification and fermentation (104, 105). Some lignin-degrading fungi, notably Pleurotus spp. and Lentinus edodes, produce edible mushrooms and can directly convert lignocellulose into food for humans. Lignin bioconversion Large quantities of lignin are available as by-products from chemical pulping and more would become available if lignocellulose were utilized as a source of sugars for fermentation (106). The available markets for these lignins are mostly low valued. Using lignin-degrading fungi or their enzymes to tailor the properties of these lignins and thus increase their value is an appealing idea, although the low specificity of the known enzymes limits the control that can be exercised. In one successful application, laccase was used to polymerize lignin as a binder in particle boards (107) . Enzymatic polymerization of brown-rotted lignin with peroxidase and laccase was less promising (108). Laccase has also been used to incorporate organosolv and kraft lignins into graft copolymers (109).

Conclusion The biodegradation of lignin is a special talent of white-rot fungi, which possess a powerful extracellular oxidative system with low substrate specificity. Lignin peroxidase, Mn peroxidase, and laccase are components of this system in various fungal species; other constituents remain to be discovered. Lignin biodegradation as part of wood and litter decay is an important natural process, with positive and negative effects on human welfare. As our understanding of lignin biodegradation increases, we can apply these fungi and their enzymes in the pulp and paper, agricultural, and bioremediation industries.

References 1. Fengel, D., and Wegener, G. 1983. Wood: chemistry, ultrastructure, reactions. de Gruyter, New York. 2. Higuchi, T. 1985. Biosynthesis of lignin. In Biosynthesis and biodegradation of wood components. Edited b y T. Higuchi. Academic Press, Orlando, Fla. pp. 141- 161. 3. Kirk, T.K., and Brunow, G. 1988. Synthetic I4C-labeled lignins. Methods Enzymol. 161: 65-73. 4. Higuchi, T. 1985. Degradative pathways of lignin model compounds. bi Biosynthesis and biodegradation of wood components. Edited b y T. Higuchi. Academic Press, Orlando, Fla. pp. 557-578. 5. Hammel, K.E., Jensen, K.A. J., Mozuch, M .D., Landucci, L.L., Tien, M., and Pease, E.A. 1993. Ligninolysis by a purified lignin peroxidase. J. Biol. Chem. 268: 12 274 - 12 281. 6. Kirk, T.K., and Farrell, R.L. 1987. Enzymatic "combustion": the microbial degradation of lignin. Annu. Rev. Microbiol. 41: 465 -505. 7. Buswell, J.A., and Odier, E. 1987. Lignin biodegradation. CRC Crit. Rev. Biotechnol. 6: 1-60. 8. Eriksson, K.-E. L., Blanchette, R.A., and Ander, P. 1990. Microbial and enzymatic degradation of wood and wood components. Springer-Verlag, Berlin. 9. Buswell, J.A. 1991. Fungal degradation of lignin. In Handbook of applied mycology. Vol. 1. Edited b y D.K. Arora, B. Rai, K.G. Mukerji, and G.R. Knudsen. Marcel Dekker, New York. pp. 425-480. 10. Boominathan, K., and Reddy, C.A. 1992. Fungal degradation of lignin: biotechnological applications. In Handbook of applied mycology. Vol. 4. Edited b y D.K. Arora, R.P. Elander, and K.G. Mukerji. Marcel Dekker, New York. pp. 763-822. 11. Gilbertson, R.L. 1980. Wood-rotting fungi of North America. Mycologia, 72: 1-49. 12. Highley, T.L. 1982. Influence of type and amount of lignin on decay by Coriolus versicolor. Can. J. For. Res. 12: 435 -438. 13. Faix, O., Mozuch, M.D., and Kirk, T.K. 1985. Degradation of gymnosperm (guaiacyl) vs. angiosperm (syringyl/guaiacyl) lignins by Phanerochaete chrysosporiutn. Holzforschung, 39: 203 -208. 14. Blanchette, R.A. 1991. Delignification by wood-decay fungi. Annu. Rev. Phytopathol. 29: 381 -398. 15. Hatakka, A. 1994. Lignin-modifying enzymes from selected white-rot fungi: production and role in lignin degradation. FEMS Microbiol. Rev. 13: 125 - 135. 16. de Jong, E., Field, J.A., and de Bont, J.A.M. 1994. Aryl alcohols in the physiology of ligninolytic fungi. FEMS Microbiol. Rev. 13: 153- 188. 17. Kirk, T.K., and Adler, E. 1970. Methoxyl-deficient structural elements in lignin of sweetgum decayed by a brown-rot fungus. Acta Chem. Scand. 24: 3379-3390. 18. Jin, L., Schultz, T.P., and Nicholas, D.D. 1990. Structural characterization of brown-rotted lignin. Holzforschung, 44: 133- 138. 19. Blanchette, R.A. 1995. Degradation of the lignocellulose complex in wood. Can. J. Bot. 73(Suppl. 1). This issue. 20. Haider, K., and Trojanowski, J. 1980. A comparison of the degradation of '"-labeled DHP and corn stalk lignins by micro- and macrofungi and bacteria. In Lignin biodegradation: microbiology, chemistry, and potential applications. Vol. 1. Edited b y T.K. Kirk, T. Higuchi, and H.-m. Chang. CRC Press, Boca Raton, Fla. pp. 111 - 134. 21. Chen, C.-I., and Chang, H.-m. 1985. Chemistry of lignin biodegradation. In Biosynthesis and biodegradation of wood components. Edited b y T. Higuchi. Academic Press, Orlando, Fla. pp. 535-556.


Can. J. Bot. Vol. 73 (Suppl. I ) , 1995 22. Haemmerli, S.D., Leisola, M.S.A., and Fiechter, A. 1986. Polymerisation of lignins by ligninases from Phanerochaere chrysosporium. FEMS Microbiol. Lett. 35: 33-36. 23. Ishihara, T., and Miyazaki, M. 1972. Oxidation of milled wood lignin by fungal laccase. Mokuzai Gakkaishi, 18: 415-419. 24. Kurek, B., Monties, B., and Odier, E. 1990. Influence of the physical state of lignin on its degradability by the lignin peroxidase of Phanerochaete chrysosporium. Enzyme Microb. Technol. 12: 771 -777. 25. Kondo, R., Iimori, T., Imamura, H., and Nishida, T. 1990. Polymerization of DHP and depolymerization of DHP-glucoside by lignin oxidizing enzymes. Biol Technology, 13: 181 - 188. 26.Chua, M.G.S., Choi, S., and Kirk, T.K. 1983. Mycelium binding and depolymerization of synthetic I4C-labelled lignin during decomposition by Phanerochaete chrysosporiurn. Holzforschung, 37: 55 -61. 27. Reid, I.D. 1991. Intermediates and products of synthetic lignin (dehydrogenative polymerizate) degradation by Phlebia tremellosa. Appl. Environ. Microbiol. 57: 2834-2840. 28. Freer, S.N., and Detroy, R. W. 1982. Biological delignification of I4C-labelled lignocelluloses by basidiomycetes: degradation and solubilization of the lignin and cellulose components. Mycologia, 74: 943 -95 1. 29. Reid, I.D., Abrams, G.D., and Pepper, J.M. 1982. Water-soluble products from the degradation of aspen lignin by Phanerochaete chrysosporium. Can. J. Bot. 60: 2357 -2364. 30. Leisola, M., Ulmer, D., Haltmeier, T., and Fiechter, A. 1983. Rapid solubilization and depolymerization of purified Kraft lignin by thin layers of Phanerochaete chrysosporium. Eur. J. Appl. Microbiol. Biotechnol. 17: 117- 120. 31. Abbott, T.P., and Wicklow, D.T. 1984. Degradation of lignin by Cyathus species. Appl. Environ. ~ i c r o b i o l .47: 585-587. 32. Reid, I.D. 1985. Biological delignification of aspen wood by solid-state fermentation with the white-rot fungus Merulius tremellosus. Appl. Environ. Microbiol. 50: 133- 139. 33. Trojanowski, J., and Hiittermann, A. 1987. Screening of wood inhabiting fungi for their capacity to degrade and to solubilize '"-labelled lignin. Microbios, 50: 91 -97. 34. Suzuki, H., Iiyama, K., Yoshida, O., Yamazaki, S., and Yamamoto, N. 1990. Structural characterization of the immunoactive and anti-viral water-solubilized lignin in an extract of the culture medium of Lentinus edodes mycelium (LEM). Agric. Biol. Chem. 54: 479-487. 35.Boyle, C.D., Kropp, B.R., and Reid, I.D. 1992. Solubilization and mineralization of lignin by white rot fungi. Appl. Environ. Microbiol. 58: 3217-3224. 36. Kirk, T.K., and Chang, H.-m. 1974. Decomposition of lignin by white-rot fungi I. Isolation of heavily degraded lignins from decayed spruce. Holzforschung, 28: 217-222. 37.Tai, D.-s., Terazawa, M., Chen, C.-l., and Chang, H.-m. 1990. Lignin biodegradation products from birch wood by Phanerochaete chrysosporium. Part 1. Fractionation of methanol-extractive and characterization of ether-insoluble low-molecular-weight fraction. Holzforschung, 44: 185 - 190. 38. Ellwardt, P.-C., Haider, K., and Ernst, L. 1981. Untersuchungen des mikrobiellen Ligninabbaues durch 13C-NMR-Spektroskopie an spezifisch "C-angereichertem DHP-Lignin aus Coniferylalkohol. Holzforschung, 35: 103- 109. 39. Tien, M. 1987. Properties of ligninase from Pharlerochaete chrysosporium and their possible applications. CRC Crit. Rev. Microbiol. 15: 141 - 168.

40. Umezawa, T., and Higuchi, T. 1991. Chemistry of lignin degradation by lignin peroxidases. In Enzymes in biomass conversion. Edited by G.F. Leatham and M.E. Himmel. American Chemical Society, Washington, D.C. pp. 236-246. 41. Gold, M.H., and Alic, M. 1993. Molecular biology of the lignin-degrading basidiomycete Phanerochaete chrysosporium. Microbiol. Rev. 57: 605-622. 42. Barr, D.P., and Aust, S.D. 1994. Mechanisms white rot fungi use to degrade pollutants. Environ. Sci. Technol. 28: 78A-87A. 43. Tien, M., and Kirk, T.K. 1983. Lignin-degrading enzyme from the hymenomycete Phanerochaete chrysosporiurn Burds. Science (Washington, D.C.), 221: 661 -662. 44. Glenn, J.K., Morgan, M.A., Mayfield, M.B., Kuwahara, M., and Gold, M.H. 1983. An extracellular H702-requiring enzyme preparation involved in lignin biodegradation by the white-rot basidiomycete Phanerochaete chrysosporium. Biochem. Biophys. Res. Commun. 114: 1077 - 1083. 45. Wariishi, H., and Gold, M.H. 1990. Lignin peroxidase compound 111. Mechanism of formation and decomposition. J. Biol. Chem. 265: 2070-2077. 46. Bonnen, A.M., Anton, L.H., and Orth, A.B. 1994. Lignin-degrading enzymes of the commercial button mushroom, Agaricus bisporus. Appl. Environ. Microbiol. 60: 960-965. 47. Harvey, P.J., Schoemaker, H.E., and Palmer, J.M. 1986. Veratryl alcohol as a mediator and the role of radical cations in lignin biodegradation by Phanerochaete chrysosporium. FEBS Lett. 195: 242 -246. 48. Wariishi, H., and Gold, M.H. 1989. Lignin peroxidase compound 111-formation, inactivation, and conversion to the native enzyme. FEBS Lett. 243: 165- 168. 49. Koduri, R.S., and Tien, M. 1994. Kinetic analysis of lignin peroxidase-explanation for the mediation phenomenon by veratryl alcohol. Biochemistry, 33: 4225 -4230. 50. Poulos, T.L., Edwards, S.L., Wariishi, H., and Gold, M.H. 1923. Crystallographic refinement of lignin peroxidase at 2 A. J. Biol. Chem. 268: 4429 -4440. 51. Piontek, K., Glumoff, T., and Winterhalter, K. 1993. Low pH crystal structure of glycosylated ligni5 peroxidase from Phanerochaere chrysosporium at 2.5 A resolution. FEBS Lett. 315: 119- 124. 52. Wariishi, H., Sheng, D., and Gold, M.H. 1994. Oxidation of ferrocytochrome c by lignin peroxidase. Biochemistry, 33: 5545 -5552. 53. Glenn, J.K., and Gold, M.H. 1985. Purification and characterization of an extracellular Mn(I1)-dependent peroxidase from the lignin-degrading basidiomycete, Phanerochaete chrysosporium. Arch. Biochem. Biophys. 242: 329-341. 54.Tuor, U., Wariishi, H., Schoemaker, H.E., and Gold, M.H. 1992. Oxidation of phenolic arylglycerol 0-aryl ether lignin model compounds by manganese peroxidase from Phanerochaete chrysosporium: oxidative cleavage of an a-carbonyl model compound. Biochemistry, 31: 4986 -4995. 55. Forrester, I.T., Grabski, A.C., Mishra, C., Kelley, B.D., Strickland, W.N., Leatham, G.F., and Burgess, R.R. 1990. Characteristics and N-terminal amino acid sequence of a manganese peroxidase purified from Lentinula edodes cultures grown on a commercial wood substrate. Appl. Microbiol. Biotechnol. 33: 359-365. 56. Paice, M.G., Reid, I.D., Bourbonnais, R., Archibald, F.S., and Jurasek, L. 1993. Manganese peroxidase, produced by Trametes versicolor during pulp bleaching, demethylates and delignifies kraft pulp. Appl. Environ. Microbiol. 59: 260-265.


F5.2. Reid

57. Hirai, H., Kondo, R., and Sakai, K. 1994. Screening of lignin-degrading fungi and their ligninolytic enzyme activities during biological bleaching of kraft pulp. Mokuzai Gakkaishi, 40: 980-986. 58. Wariishi, H., Valli, K., and Gold, M.H. 1991. In vitro depolymerization of lignin by manganese peroxidase of Phanerochaete chrysosporium. Biochem. Biophys. Res. Commun. 176: 269 -275 59. Thurston, C.F. 1994. The structure and function of fungal laccases. Microbiology, 140: 19 -26. 60. Bourbonnais, R., and Paice, M.G. 1990. Oxidation of non-phenolic substrates. An expanded role for laccase in lignin biodegradation. FEBS Lett. 267: 99- 102. 61. Muheim, A,, Fiechter, A., Harvey, P.J., and Schoemaker, H.E. 1992. On the mechanism of oxidation of non-phenolic lignin model compounds by the laccase-ABTS couple. Holzforschung, 46: 121- 126. 62. Bourbonnais, R., and Paice, M.G. 1992. Demethylation and delignification of kraft pulp by Trametes versicolor laccase in the presence of 2,2'-azinobis-(3-ethylbenzthiazoline-6sulphonate). Appl. Microbiol. Biotechnol. 36: 823-827. 63. Call, H.P., and Miicke, I. 1994. State of the art of enzyme bleaching and disclosure of a breakthrough process. In International Non-Chlorine Bleaching Conference Proceedings. Miller Freeman, Inc., San Francisco. pp. 1-6. 64. Ander, P. 1994. The cellobiose-oxidizing enzymes CBQ and CbO as related to lignin and cellulose degradation-a review. FEMS Microbiol. Rev. 13: 297 -312. 65. Ander, P., Mishra, C., Farrell, R.L., and Eriksson, K.-E.L. 1990. Redox reactions in lignin degradation: interactions between laccase, different peroxidases, and ce11obiose:quinone oxidoreductase. J. Biotechnol. 13: 189-198. 66. Kremer, S.M., and Wood, P.M. 1992. Production of Fenton's reagent by cellobiose oxidase from cellulolytic cultures of Phanerochaete chrysosporium. Eur. J. Biochem. 208: 807-814. 67. Roy, B.P., Paice, M.G., Archibald, F.S., Misra, S.K., and Misiak, L.E. 1994. Creation of metal-complexing agents, reduction of manganese dioxide, and promotion of manganese peroxidase-mediated ~ n ( 1 i 1 )production by cel1obiose:auinone oxidoreductase from Trametes versicolor. . 19745 - 19750. J. Biol. ~ h k m 269: 68. Kelley, R.L., and Reddy, C.A. 1986. Identification of glucose oxidase activity as the primary source of hydrogen peroxide production in ligninolytic cultures of Phanerochaete chrysosporium. Arch. Microbiol. 144: 248-253. 69. Daniel, G., Volc, J., and Kubatova, E. 1994. Pyranose oxidase, a major source of H202 during wood degradation by Phanerochaete chrysosporium, Trametes versicolor, and Oudemansiella mucida. Appl. Environ. Microbiol. 60: 2524-2532. 70. Kersten, P.J., and Kirk, T.K. 1987. Involvement of a new enzyme, glyoxal oxidase, in extracellular H202production by Phanerochaete chrysosporium. J. Bacteriol. 169: 2195 -2201. 71. Nishida, A,, and Eriksson, K.-E. 1987. Formation, purification, and partial characterization of methanol oxidase, a H20,-producing enzyme in Phanerochaete chrysosporium. Biotechnol. Appl. Biochem. 9: 325 -338. 72. Reid, I.D., and Deschamps, A.M. 1991. Nutritional regulation of synthetic lignin (DHP) degradation by the selective white-rot fungus Phlebia (Merulius) tremellosa: effects of glucose and other cosubstrates. Can. J. Bot. 69: 147- 155. 73. GuillCn, F., and Evans, C.S. 1994. Anisaldehyde and

veratraldehyde acting as redox cycling agents for H,02 production by Pleurotus eryngii. Appl. Environ. Microbiol. 60: 281 1-2817. 74. Evans, C.S., Dutton, M.V., GuillCn, F., and Veness, R.G. 1994. Enzymes and small molecular mass agents involved with lignocellulose degradation. FEMS Microbiol. Rev. 13: 235 -240. 75. Roy, B.P., and Archibald, F. 1993. Effects of kraft pulp and lignin on Trametes versicolor carbon metabolism. Appl. Environ. Microbiol. 59: 1855- 1863. 76. de Jong, E., Field, J.A., and de Bont, J.A.M. 1992. Evidence for a new extracellular peroxidase. Manganese-inhibited peroxidase from the white-rot fungus Bjerkandera sp. BOS 55. FEBS Lett. 299: 107 - 110. 77. Vares, T., Lundell, T.K., and Hatakka, A.I. 1992. Novel heme-containing enzyme possibly involved in lignin degradation by the white-rot fungus Junghuhnia separabilima. FEMS Microbiol. Lett. 99: 53 -58. 78. Flournoy, D.S., Paul, J.A., Kirk, T.K., and Highley, T.L. 1993. Changes in the size and volume of pores in sweetgum wood during simultaneous rot by Phanerochaete chrysosporium Burds. Holzforschung, 47: 297 - 30 1. 79. Srebotnik, E., and Messner, K. 1990. Enzymatic attack of wood is limited by the inaccessibility of the substrate. In Biotechnology in pulp and paper manufacture. Applications and fundamental investigations. Edited by T.K. Kirk and H.-m. Chang. Butterworth-Heinemann, Boston. pp. 111-119. 80. Blanchette, R.A., Abad, A.R., Farrell, R.L., and Leathers, T.D. 1989. Detection of lignin peroxidase and xylanase by immunocytochemical labeling in wood decayed by basidiomycetes. Appl. Environ. Microbiol. 55: 1457- 1465. 8 1. Boddy , L. 1991. Importance of wood decay fungi in forest ecosystems. In Handbook of applied mycology. Vol. 1. Edited by D.K. Arora, B. Rai, K.G. Mukerji, and G.R. Knudsen. Marcel Dekker, New York. pp. 507-540. 82. Zabel, R.A., and Morrell, J.J. 1992. Wood microbiology. Decay and its prevention. Academic Press, San Diego, Calif. p. 6. 83. Reid, I.D. 1991. Biological pulping in paper manufacture. TIBTECH, 9: 262 - 265. 84. Akhtar, M., Attridge, M.C., Myers, G.C., and Blanchette, R.A. 1993. Biomechanical pulping of loblolly pine chips with selected white-rot fungi. Holzforschung, 47: 36-40. 85. Akhtar, M. 1994. Biomechanical pulping of aspen wood chips with 3 strains of Ceriporiopsis subvermispora. Holzforschung, 48: 199 -202. 86. Yu, H., Cai, Z.-y., Huang, X.-y., and Xu, H. 1990. Biological degradation and delignification of rice straw. In Biotechnology in pulp and paper manufacture. Applications and fundamental investigations. Edited by T.K. Kirk and H.-m. Chang. Butterworth-Heinemann, Boston. pp. 59-68. 87. Gutierrez, I., Fernandez, N., and Lopez, P. 1987. Production of pulp and fodder from bagasse. Int. Sugar J. 89: 157-159. 88. Messner, K., and Srebotnik, E. 1994. Biopulping-an overview of developments in an environmentally safe paper-making technology. FEMS Microbiol. Rev. 13: 351 -364. 89. Oriaran, T.P., Labosky, P., Jr., and Blankenhorn, P.R. 1990. Kraft pulp'and papermaking properties of Phanerochaete chrysosporium-degraded aspen. Tappi J. 73(7): 147- 152. 90. Reid, I.D., and Paice, M.G. 1994. Biological bleaching of kraft pulps by white-rot fungi and their enzymes. FEMS Microbiol. Rev. 13: 369 -376.


Can. J. Bot. Vol. 7 3 (Suppl. I ) , 1995 91. Kirk, T.K., and Yang, H.H. 1979. Partial delignification of unbleached Kraft pulp with ligninolytic fungi. Biotechnol. Lett. 1: 347-352. 92. Kondo, R., Kurashiki, K., and Sakai, K. 1994. In vitro bleaching of hardwood kraft pulp by extracellular enzymes excreted from white rot fungi in a cultivation system using a membrane filter. Appl. Environ. Microbiol. 60: 921 -926. 93. Addleman, K., and Archibald, F.S. 1993. Kraft pulp bleaching and delignification by dikaryons and monokaryons of Trametes versicolor. Appl. Environ. Microbiol. 59: 266-273. 94. Iimori, T., Kaneko, R., Yoshikawa, H., Machida, M., Yoshioka, H., and Murakami, K. 1994. Screening of pulp-bleaching fungi and bleaching activity of newly isolated fungus SKB- 1152. Mokuzai Gakkaishi, 40: 733-737. 95. Fujita, K., Kondo, R., Sakai, K., Kashino, Y., Nishida, T., and Takahara, Y. 1993. Biobleaching of softwood kraft pulp with white-rot fungus IZU-154. Tappi J. 76(1): 81 -84. 96. Archibald, F.S. 1992. The role of fungus-fiber contact in the biobleaching of kraft brownstock by Trametes (Coriolus) versicolor. Holzforschung, 46: 305 -3 10. 97. Kaneko, R., Iimori, T., Yoshikawa, H., Machida, M., Yoshioka, H., and Murakami, K. 1994. A possible role of manganese peroxidase during biobleaching by the pulp bleaching fungus SKB- 1152. Biosci. Biotechnol. Biochem. 58: 1517-1518. 98. Kondo, R., Harazono, K., and Sakai, K. 1994. Bleaching of hardwood kraft pulp with manganese peroxidase secreted from Phanerochaete sordida YK-624. Appl. Environ. Microbiol. 60: 4359-4363. 99. Moen, M.A., and Hammel, K.E. 1994. Lipid peroxidation by the manganese peroxidase of Pharzerochaete chrysosporiurn is the basis for phenanthrene oxidation by the intact fungus. Appl. Environ. Microbiol. 60: 1956- 1961. 100. Lackner, R., Srebotnik, E., and Messner, K. 1991. Oxidative degradation of high molecular weight chlorolignin

by manganese peroxidase of Phanerochaete chrysosporiurn. Biochem. Biophys. Res. Commun. 178: 1092- 1098. 101. Roy-Arcand, L., and Archibald, F.S. 1991. Direct dechlorination of chlorophenolic compounds by laccases from Trametes (Coriolus) versicolor. Enzyme Microb. Technol. 13: 194-203. 102. Yin, C.-f., Joyce, T.W., and Chang, H.-m. 1990. Dechlorination of conventional softwood bleaching effluent by sequential biological treatment. In Biotechnology in pulp and paper manufacture. Applications and fundamental investigations. Edited by T.K. Kirk and H.-m. Chang. Butterworth-Heinemann, Boston. pp. 231 -244. 103. Lamar, R.T., and Dietrich, D.M. 1990. In situ depletion of pentachlorophenol from contaminated soil by Phanerochaete spp. Appl. Environ. Microbiol. 56: 3093-3100. 104. Reid, I.D. 1989. Solid-state fermentations for biological delignification. Enzyme Microb. Technol. 11: 786 -803. 105. Mes-Hartree, M., Yu, E.K.C., Reid, I.D., and Saddler, J.N. 1987. Suitability of aspenwood biologically delignified with Phlebia tremellosus for fermentation to ethanol or butanediol. Appl. Microbiol. Biotechnol. 26: 120- 125. 106. Chum, H.L., Parker, S.K., Feinberg, D.A., Wright, J.D., Rice, P.A., Sinclair, S.A., and Glasser, W.G. 1985. The economic contribution of lignins to ethanol production from biomass. SERI, Golden, Colo. 107. Haars, A., Kharazipour, A., and Huttermann, A. 1986. Two component wood adhesives based on lignin and phenoloxidases. In Proceedings, 3rd International Conference on Biotechnology in the Pulp and Paper Industry. Vol. 1. pp. 117-1 19. 108. Jin, L., Nicholas, D.D., and Schultz, T.P. 1991. Wood laminates glued by enzymic oxidation of brown-rotted lignin. Holzforschung, 45: 467-468. 109. Milstein, O., Huttermann, A,, Frund, R., and Ludemann, H.-D. 1994. Enzymatic co-polymerization of lignin with low-molecular mass compounds. Appl. Microbiol. Biotechnol. 40: 760 -767.