Ligninolysis by a Purified Lignin Peroxidase

The lignin peroxidases (Lips) of white-rot basidiomycetes are generally thought to catalyze the oxidative cleavage of polymeric lignin in vivo. However, direct evidence for such a role has been lacking. In this investigation, Cand C-labeled synthetic lignins were oxidized with a purified isozyme of Phanerochaete chrysosporium LiP. Gel permeation chromatography of the radiolabeled polymers showed that LiP catalyzed their cleavage to give soluble lower-Mr products. To a lesser extent, the enzyme also polymerized the lignins to give soluble higher-M r products. This result is attributable to the fact that purified LiP, unlike the intact fungus, provides no mechanism for the removal of lignin fragments that are susceptible to repolymerization. LiP catalysis also gave small quantities of insoluble, perhaps polymerized, lignin, but in lower yield than intact P. chrysosporium does. C NMR experiments with C-labeled polymer showed that LiP cleaved it between Cα and Cβ of the propyl side chain to give benzylic aldehydes at Cα , in agreement with the cleavage mechanism hypothesized earlier. The data show that LiP catalysis accounts adequately for the initial steps of ligninolysis by P. chrysosporium in vivo.

Despite much detailed work on the reaction mechanism of LiPs. there is little evidence that they can actually cleave polymeric lignin. Experiments with low M r lignin model compounds provide strong support for a ligninolytic role, but cannot prove it. Investigations with lignin itself as a LiP substrate have, excepting some preliminary data (21,22), failed to show ligninolysis (23-26), and some researchers have concluded that LiPs do not cleave lignin in vivo (26). Here we show that a purified isozyme of LiP from the ligninolytic basidiomycete Phanerochaete chrysosporium does, in fact, catalyze the reaction that it is named for.

MATERIALS AND METHODS
Synthetic Lignins-Unlabeled synthetic guaiacyl lignin was prepared by polymerizing coniferyl alcohol in the presence of horseradish peroxidase/H 2 O 2 as described (27). Synthetic [β-14 C]guaiacyl lignin with a specific activity equal to 0.01 mCi mol -1 of phenylpropane substructures was prepared by the same procedure from [β-14 C] coniferyl alcohol. [ 14 C]Coniferyl alcohol was synthesized from vanillin and [2-14 C]malonic acid diethyl ester (Sigma, diluted with unlabeled compound to 0.01 mCi mmol -1 ) (27,28). Synthetic [ring-14 C]guaiacyl lignin with a specific activity of 0.07 mCi mmol -1 of phenylpropane substructures (29) was obtained from T. K. Kirk of the Forest Products Laboratory. An exhaustively methylated [β-14 C]guaiacyl lignin was prepared by treating the unmodified synthetic polymer in N,N-dimethylformamide (DMF) with periodic additions of diazomethane for 4 weeks (30). The progress of methylation was monitored by determining the UV/visible difference spectrum of alkaline versus neutral lignin. The complete absence of a phenoxide absorbance peak at 300 nm after 4 weeks showed that this treatment gave >99% methylation of the polymer (31).
The lignins were sized by preparative gel permeation chromatography (GPC) on a 1.8 × 32.5 cm column of Sephadex LH60/LH20 (1:1 wt/wt, Pharmacia LKB Biotechnology Inc.) in DMF (24), and fractions (2.0 ml) were collected. Size-fractionated lignins were then selected for subsequent investigations of LiP-catalyzed ligninolysis. For GPC experiments with unmodified or methylated [β-13 C]lignin, four adjacent fractions with a V e /V o (elution volume/excluded volume) range of 1.5-1.7 were pooled. For GPC experiments with [ring-14 C]lignin, a single fraction with a V e /V o of 1.7 was selected. For NMR experiments with [α-13 C]lignin or unlabeled lignin, fractions with a V e /V o range of 1.1-2.2 were pooled. After preparative chromatography, some of the DMF eluant was removed from the selected samples by rotary vacuum evaporation at <30°C, and the lignin solutions were stored at -20°C. When lignin solutions were added to in vitro depolymerization reactions, the final DMF concentration was <0.25% v/v. added to the flask and the mixture was concentrated to <4 ml by rotary evaporation at <30°C, which removed most of the water and alcohol. The resulting mixture. followed by a 1-ml DMF rinse of the reaction flask. was transferred to a silanized glass centrifuge tube and centrifuged at 1.5.000 × g for 30 min. The supernatant fraction was carefully decanted and brought to 5.0 ml with DMF in a silanized graduated cylinder. The centrifugation pellet, a sample (1.0 ml) of the supernatant fraction. and a DMF/H 2 O (1:1, v/v) rinse of the reaction flask were then assayed for 14 C in Polyfluor mixture (Packard) in an LKB Wallac 1214 scintillation counter. α-13 C-Depleted Veratryl Alcohol-4-Bromoveratrole (Aldrich) was condensed with 12 CO 2 (Icon Services, Mt. Marion, NY; >99.98 atom % 12 C) in the presence of n-butyl lithium to give [α-12 C]veratric acid. which was recrystallized from H 2 O. This procedure is essentially the same as that used by Newman et al. (33) to prepare vanillic acid from 4-bromoguaiacol. [α-12 C]Veratric acid was then refluxed in methanol/ H 2 SO 4 to give [α-12 C]veratric acid methyl ester (34), which was extracted with CCl 4 and recrystallized from ethanol/H 2 O. [α-12 C] Veratric acid methyl ester was reduced with diisobutylaluminum hydride (DIBAL-H, Aldrich) (32), and the resulting [α-12 C]veratryl alcohol was purified by vacuum distillation (35).
Fungal Peroxidases-An overproducing strain of P. chrysosporium, PSBL-1 (36, 37) was used for production of LiP and manganese peroxidase (MnP). Extracellular fluid from day 5 cultures was harvested and fractionated by ion exchange high performance liquid chromatography on a Pharmacia FPLC Mono Q column as described previously (38, 39). The eluted peak containing LiP isozyme H1 (pI = 4.7) (38) was then further purified by preparative isoelectric focusing (IEF) on a Pharmacia LKB Multiphor II electrophoresis system, using Ultrodex (Pharmacia) as a support, according to the manufacturer's instructions. The resulting LiP H1 was homogeneous by analytical IEF and had an A 408 nm /A 280 nm ratio > 4. The quantitation of LiP H1 was based on a of 20 s -1 at pH 3.0 in the veratryl alcohol oxidation assay (39). The purified LiP H1 showed 30% of maximal activity in 30% v/v 2-methoxyethanol, and 20% of maximal The Mono Q chromatographic fraction containing MnP isozyme H4 (pI = 4.5) (38) was dialyzed against sodium succinate (50 mM, pH 4.5) and further purified by dye-ligand chromatography on blue agarose (Cibacron Blue 3GA type 3000L, Sigma) (40). This H4 fraction was then further purified by preparative IEF as described above for LiP. The resulting MnP H4 was homogeneous by analytical IEF, had an A 406 nm /A 280nm ratio > 4 and contained <0.l% mol/mol LiP. MnP H4 was quantitated spectrophotometrically at 406 nm nm = 127 mM -1 cm -i ). The purified MnP H4 showed 40% of maximal activity in 40% (v/v) 2-propanol.
Other Reagents-H 2 O 2 (30%, Fisher) was determined by titration with KMnO 4 , and stock solutions for peroxidase-catalyzed reactions were prepared daily. 2-Methoxyethanol was glass-distilled and stored for 1 week or less at -20°C under N 2 . Reagent grade 2-propanol was used as received. Veratryl alcohol was purified by vacuum distillation (35).
MnP-catalyzed lignin oxidations were performed in the same way, but with the following changes. The reaction flask contained MnSO 4 (8.0 µmol), no veratryl alcohol, and no enzyme. The enzyme feed solution contained MnP (1.2 nmol) without veratryl alcohol, and the peroxide feed solution contained 12 µmol of H 2 O 2 .
At the conclusion of each reaction, the contents of the reaction flask were adjusted to pH 7 with NaOH, and a few crystals of catalase were added to destroy any remaining H 2 O 2 . DMF (7.0 ml) was then M r Analysis of Enzyme-treated Lignins-The remainder (4.0 ml) of the supernatant fraction was applied to a 1.8 × 32.5 cm lipophilic Sephadex (Pharmacia) GPC column previously equilibrated with DMF that was 0.1 M in LiCl (41). Sephadex LH60 was used for β-14 C-labeled lignin, and Sephadex LH60/LH20 (1:1, w/w) (24) was used for ring-14 C-labeled lignin. DMF containing 0.1 M LiCl was used as the column eluant, and fractions of 1.5 ml were collected for scintillation counting, which was done as described above. Nominal M r calibrations of the GPC columns were based on polystyrene standards. Number-average M r values (M n values) based on these calibrations were calculated from the standard equation (42).
13 C NMR Analysis of Enzyme-treated Lignins-The LiP-catalyzed oxidation of lignin for 13 C NMR analysis was done in a l-liter roundbottom flask. maintained in the dark at room temperature, that initially contained sodium glycolate (10 mM, pH 4.5), 2-methoxyethanol (30% vol/vol), [α-12 C]veratryl alcohol (0.05 mmol), and α-13 C-labeled synthetic lignin (1.5 mg). The volume was 375 ml. Mariotte flasks were used to deliver two separate solutions to the reaction vessel. One solution (62.5 ml, maintained throughout at 0°C) contained sodium glycolate (10 mM, pH 4.5), [α-12 C]veratryl alcohol (0.15 mmol), and LiP (130 nmol of active enzyme). The second solution (62.5 ml) contained sodium glycolate (10 mM, pH 4.5), 2-methoxyethanol (60%, v/v), and H 2 O 2 (0.15 mmol). Two control reactions were run, one without LiP and one with natural abundance lignin in place of α-13 C-labeled lignin. The two solutions were added at 7.8 ml h -1 , and the reaction flask was stirred continuously. The total reaction time was 13 h, after which the mixture was adjusted to pH 7 with NaOH, and approximately 1 mg of catalase was added to destroy any remaining H 2 O 2 . The mixture was then concentrated to about 100 ml by rotary vacuum evaporation at <30°C. and precipitated salts were removed by filtration through glass wool. The filtrate was further concentrated to dryness, and dichloroethane/ethanol (2:1, v/v) was added to the residue. The dichloroethane/ethanol-soluble fraction, consisting of the recoverable soluble lignin, was then filtered through a 0.45-µpm nylon filter and concentrated to dryness. This dried lignin sample was acetylated for 8 h in 1 ml of acetic anhydride/pyridine (1:1, v/v) and again concentrated to dryness. Residual pyridine was removed by azeotropic distillation with toluene under reduced pressure on the rotary evaporator at <30°C, and toluene was then removed likewise by distillation with acetone. The resulting dry, acetylated lignin sample was dissolved in deuteroacetone and filtered through glass wool for 13 C NMR analysis. 13 C NMR data were obtained on a Bruker WM250 spectrometer controlled by an Aspect 2000A minicomputer. A 10-mm fixed frequency probe tuned to 62.9 MHz was utilized, and samples were analyzed in 5-mm sample tubes. DEPT (distortionless enhancement by polarization transfer) spectra were acquired with a standard Bruker microprogram. Acquisitions consisting of approximately 0.5-1.5 × 10 5 transients were generally required to obtain sufficient signal/ noise ratios. A line broadening of 6 Hz was applied prior to Fourier transformation.

LiP-mediated M r Changes in Lignin-
The treatment of synthetic lignins with LiP/H 2 O 2 gave the same yield of organic-soluble products that P. chrysosporium gives in whole culture experiments of similar duration and with similar quantities of lignin (43,44). GPC of this soluble recoverable lignin showed that LiP degraded the polymer (Figs. 1 and 2A). Some higher M r material was formed, but depolymerization was the predominant fate of the DMF-soluble lignin in vitro, in agreement with the GPC results obtained from in vivo studies (43, 44). The β-14 C-labeled polymer appeared to be more extensively depolymerized than the ring-14 C-labeled polymer was, which might simply reflect differences in the  Table I. A, M r distributions of β-14 C-labeled lignin. The following reactions shown in Table I were analyzed: 1,-; 2,---; 2, ---; 3, · · · · ·. B, M r distributions of ring-14 Clabeled lignin. The following reactions shown in Table I were analyzed: 4,-; 5, ---; 6, · · · · ·. The designations a-e correspond to the elution positions of polystyrene standards with the following M r values: 34,500 (excluded from gel) (a), 22,000 (b), 10,000 (c), 3,100 (d), 1,050 (e). Standard (f) was veratraldehyde (M r = 166). degree of condensation and accessibility to LiP of the two lignins. Alternatively, the apparent difference might be an artifact if the aromatic/aliphatic ratio of degraded lignin were to decrease with decreasing polymer M r , but we have no independent indication that this was the case.
These reactions also gave an insoluble lignin fraction, much of which probably consisted of noncovalently aggregated polymer because it occurred in controls as well as enzyme-treated reactions ( Table I). The differences in insolubles between LiP-treated and control reactions presumably represent higher-M, DMF-insoluble lignin that had been polymerized by LiP, but the yields of this fraction (< 16%) were considerably lower than the yields of insoluble lignin (30%) obtained in 12-h P. chrysosporium cultures (43,44).
Enzyme-treated reactions gave lower overall yields than control reactions did, presumably because volatile low M r fragments were lost during evaporation of the sample, as reported in preliminary studies with crude LiP (22). The most likely sources of missing 14 C in side chain-labeled lignin are vinylic (coniferyl alcohol) end groups and internal β-O-4 substructures (Fig. 3). both of which should be cleaved by LiP to release 14 C β in two-carbon fragments (5,6). The loss of 14 C from degraded ring-labeled lignin might involve the cleavage of phenolic substructures to give volatile methoxyquinones (45-47). We cannot rule out the possibility that some of the missing 14 C might have been polymerized and tightly bound to the wall of the reaction flask, but such a result would not affect our conclusion that the action of LiP resembles the initial attack by intact P. chrysosporium on lignin: even if it is assumed that all of the missing 14 C was polymerized to insolubility, the maximum amount of insoluble material attributable to LiP in any enzymatic experiment (28%) was still no greater than the yield of insoluble lignin (30%) produced by the fungus after 12 h in vivo (43,44).
Ligninolysis in vitro required the addition of an organic solvent to disperse the polymer. Degradation was somewhat more extensive in the 2-methoxyethanol-amended system than it was in the reactions amended with 2-propanol (Figs. 1A and 2A ). Another ingredient required in these reactions was veratryl alcohol, a natural P. chrysosporium metabolite (48) and LiP substrate that is produced in ligninolytic fungal cultures. Veratryl alcohol stimulates the oxidation of some lignin model compounds by LiP (49,50) and was also found to be necessary for LiP-catalyzed ligninolysis in preliminary experiments with crude enzyme (22). Since this alcohol is a LiP substrate, its inclusion in ligninolysis reactions necessitated the addition of a stoichiometric equivalent of H 2 O 2 . Ligninolysis was effective at a veratryl alcohol delivery of 375 µmol per liter of reaction. which corresponds well with the extracellular veratryl alcohol concentrations found in ligninolytic P. chrysosporium cultures (48, 51). An increase in veratryl alcohol and H 2 O 2 above this level gave little change in the extent of ligninolysis (Fig. 1).
Catalytic Role of LiP-Although it is qualitatively apparent from the results that LiP degraded the lignin, simple inspection of the GPC plots does not reveal whether the enzyme was capable of acting catalytically in this process. This information can be obtained by comparing the number-average M r (M n ) of the most extensively degraded lignin (Fig. 1A, dotted curve) versus the M n of its control (Fig. 1A, solid curve). An exact determination of M n for the LiP-treated lignin was not possible, because a small proportion of the loaded polymer (1.0%) was excluded from the GPC column in the first two collected fractions of the peak and was therefore of unknown M r . However, M n values are relatively insensitive to errors at the high end of M r distributions (42), and good estimates could therefore be obtained by omitting these fractions from the calculations. (M w , the weight-average M r , could not be estimated reliably because it is strongly influenced by high-M r contributions, but M n and not M w is the quantity applicable to enumeration of the polymer molecules in the samples.) This approach, when applied to the data of Fig. 1A, gave M n = 4200 for the control lignin and M n = 850 for the most thoroughly degraded lignin. Since the M r of a guaiacyl lignin C9 substructure is about 180, it follows that the numberaverage polymer in our initial lignin preparation was roughly a 20-mer, that it was reduced by LiP to about one-fifth of this size, and that it therefore had to be cleaved in four positions to account for the final M r distribution. From the phenylpropane content in these ligninolysis reactions (14 µM), we can further calculate that they were initially about 0.7 µM in lignin polymer, which was 3 times the concentration of LiP added. Therefore, each LiP molecule must have participated in at least 12 cleavage reactions.
This number is probably an underestimate, because the presence of some polymerized lignin in LiP-treated reactions indicates that multiple cleavage and recoupling reactions, as opposed to the most parsimonious number of cleavages, must have occurred during ligninolysis. There are uncertainties inherent in these calculations (due for example to nonideal behavior of the lignin on the GPC column, inconstant 14 C specific activity across the M r distribution, or errors in M r

Mass balances for synthetic lignin in enzymatic depolymerization reactions
Reported yields are for single reactions whose GPC analyses are shown in Figs. 1 and 2. The yields are typical for each reaction type.
Reaction numbers correspond to those assigned in the figure legends.
Reaction conditions 14 C in fraction Reaction Lignin Enzyne Co-reactants Co-solvent Soluble Pellet Glassware rinse Total standardization), but it is qualitatively evident that the lig-roughly equal amounts of higher-and lower-M, DMF-soluble ninolysis observed in this experiment cannot have resulted material were produced ( Fig. 2A ). The M r of methylated lignin from a stoichiometric reaction between lignin and LiP. In-was not changed by MnP, which shows that this enzyme, stead, multiple enzyme-catalyzed reactions must have been unlike LiP, was limited to the oxidation of phenolic lignin required. structures (Fig. 2B). Different Modes of Attack for LiP versus MnP-LiP has been considered a likely catalyst for fungal ligninolysis because it can oxidize and cleave certain nonphenolic alkylaromatics that represent common internal substructures of lignin. Conventional peroxidases are less powerful oxidants, limited to the oxidation of phenolic lignin structures (9). If attack on nonphenolic units is integral to LiP-catalyzed ligninolysis, the enzyme should be able to depolymerize a lignin in which the residual unpolymerized phenolic groups have been blocked. To determine whether this was the case, we treated a sample of synthetic [β-14 C]lignin with diazomethane, which derivatizes lignin phenolic hydroxyls to methyl ethers with good selectivity. This reagent does, however, modify lignin in one other way that has to be considered: it converts vinylic substructures to heterocyclic pyrazolines (52). 13 C NMR Demonstration of LiP-catalyzed C α -C β Cleavage in Lignin-The foregoing GPC experiments show that LiP depolymerized synthetic lignin, but they do not identify the sites of cleavage. If LiP acts on the polymer in the same way that it does on dimeric lignin model compounds, the expected reaction is cleavage of the propyl side chain between C α and C β , with consequent formation of a benzylic aldehyde at C α (6, 7). With a heterogeneous substrate such as lignin, 13 C NMR spectroscopy provides the most feasible way to look for the formation of these aldehydes. Fig. 2B shows that LiP degraded the exhaustively methylated lignin. Polymerization did not occur in this reaction, probably because blockage of the phenolic groups initially present in the lignin prevented their oxidation to phenoxy radicals. Recoveries were also higher with LiP-treated methylated lignin than with LiP-treated underivatized polymer (Table I). This result suggests that derivatization of vinylic end groups in the lignin to give pyrazolines prevented their cleavage to give two-carbon fragments.
These findings contrast with the results we obtained with P. chrysosporium MnP, an enzyme that acts by oxidizing Mn(II) to chelated Mn(III), which in turn functions as a relatively mild oxidant at locations remote from the enzyme active site (40, 53). MnP attacked underivatized lignin, but was less effective than LiP at ligninolysis: GPC showed that Before this approach could be attempted, it was necessary to address a technical problem: veratryl alcohol, a necessary ingredient in these reactions, is also oxidized by LiP to give the benzylic aldehyde veratraldehyde. The formation of veratraldehyde during ligninolysis in vitro would obviously obscure the region of the lignin 13 C NMR spectrum diagnostic for benzylic aldehydes at C α . To circumvent this problem, we performed enzymatic depolymerizations in vitro with (α-13 Cenriched lignin and α-13 C-depleted veratryl alcohol. Under these conditions, the benzylic CH 2 OH of veratryl alcohol was barely detectable (at 68.4 ppm, data not shown), although the aromatic and methoxyl carbons of this ingredient were prominent. Conversely, the only carbons observable in spectra of the α-13 C-enriched lignin were those located at C α in various environments (see Fig. 3 for representative structures). Fig. 4C shows the 13 C NMR spectrum of a control reaction in which α-13 C-enriched lignin and α-13 C-depleted veratryl alcohol were treated only with H 2 O 2 . The signals characteristic of benzylic alcohols and ethers at C α in lignin are seen at 75.5 and 74.7 ppm for β-O-4 structures, at 86.3 and 85.4 Table I. A, M r distributions of β-14 C-labeled lignin. The following reactions shown in Table I were analyzed: 7,-; 8, ---; 9, · · · · ·. B, M r distributions of methylated β-14 C-labeled lignin. The following reactions shown in Table I were analyzed: 10,-; 11, ---; 12, · · · · ·. M r standards were as indicated in Fig. 1.   FIG. 3. Representative subunit structures in lignin. LiP-catalyzed C α -C β cleavage to give a benzylic aldehyde at C α is shown for a β-O-4 substructure (e). Work with lignin model compounds has suggested that the two-carbon compound released in this reaction is glycolaldehyde (6). but the identification of this fragment remains hypothetical. Structures (b) and (c) are also expected to undergo cleavage and yield benzylic aldehydes at C α upon oxidation by LiP. The effect of LiP on structure (d) is unknown. The symbol L denotes lignin, and the alphabetical designations under each structure correspond to 13 C NMR assignments in Fig. 4 and Table II. ppm for pinoresinol structures, and at 88.6 and 88.2 ppm for phenylcoumaran structures. Resinol and phenylcoumaran structures are considerably more frequent in synthetic lignin than in the natural polymer. Also characteristic of synthetic lignin, but much less prominent in natural lignin (54), are the two signals at 134.7 and 134.2 ppm that indicate vinylic C α in coniferyl alcohol end groups on the polymer. The region at 192-190 ppm, which is diagnostic for benzylic aldehydes, shows only a trace of these substructures in the lignin. These features of the spectrum were the same in H 2 O 2 -treated controls as in fresh lignin (data not shown), i.e. no ligninolysis occurred in reactions without LiP. The remaining signals in Fig. 4C are attributable to nonbenzylic positions of veratryl alcohol, to natural abundance acetate groups introduced during acetylation of the 13 C-depleted CH 2 OH group in veratryl alcohol, or in a few instances to artifacts that were also found in spectra without added 13 C-labeled lignin (Table II). Fig. 4A shows results from a complete reaction with LiP, H 2 O 2 , lignin, and 13 C-depleted veratryl alcohol, in which natural abundance lignin rather than 13 C-enriched polymer was used. The lignin, although present, is thus silent in the NMR spectrum, and all of the signals are attributable to veratryl alcohol, to products from its oxidation, or again to ligninunrelated artifacts. This control confirmed that no benzylic CHO signal appeared at 192-190 ppm when 13 C-depleted veratryl alcohol was oxidized by LiP. Any signals appearing at this position in experiments with [α-13 C]lignin must therefore come from C α positions of the polymer. Fig. 4B shows the results of the complete experiment with α-13 C-enriched lignin. All of the major signals in this spectrum are also found in one or another of the two controls, but with one prominent exception: the LiP-treated [α-13 C]lignin displayed large signals at 192-190 ppm, the region diagnostic for benzylic CHO groups and therefore for C α -C β cleavage of the polymer. A DEPT experiment (not shown) confirmed that these signals were due to protonated, i.e. aldehydic, carbons. In addition, there were minor changes in α-CHOR groups (90-70 ppm), and some of the terminal coniferyl alcohol substructures were oxidized at C γ to give coniferaldehyde units, which resulted in a downfield shift of the vinylic C α in these structures (153.7 ppm). The coniferyl alcohol substructures were preferentially attacked by LiP, as shown by their relative depletion in the spectrum, and it is clear that C α -C β cleavage of vinylic end groups must account for some of the new benzylic aldehydes seen at 192-190 ppm. However, benzylic aldehydes are under-represented in these qualitative NMR experiments because -CHO groups exhibit longer relaxation times than -HC=CH-or -CHOR-groups do. Even without correction for this error, the benzylic CHO signals are too large to come only from the missing vinylic structures. Therefore, we conclude that LiP catalyzed the C α -C β cleavage of both internal and terminal regions of the lignin backbone.

DISCUSSION
Ligninolysis by white-rot fungi in wood is a spatially complex process. The fungal hyphae grow within the lumens of woody cells, secreting ligninolytic and polysaccharide-cleaving agents that progressively degrade the lignified cell wall from the inside outwards (2, 55). During ligninolysis, the fungus produces an extracellular mucilaginous sheath, composed largely of β-1,3-1,6-linked glucan, which is found closely associated both with the hyphae and with regions of the woody cell wall that are undergoing decay (56). Electron microscopic immunogold labeling studies have demonstrated that LiPs are present in this mucilage (57-61). At early stages of decay, the enzymes are found at the surface of the lignified cell wall, but are evidently unable to penetrate it. As decay progresses, the cell wall swells and LiPs are found within its degraded regions. These results can be interpreted in two ways. First, LiPs may depolymerize lignin at the cell wall surface, gradually eroding it until they can penetrate and degrade it from within. Alternatively, some low M r agent (e.g. Mn(III) generated by MnP) may be required to penetrate the intact wall and initiate ligninolysis, and LiPs may enter the partially degraded wall later to depolymerize the remaining lignin.
There are several features of this process that must be considered if enzymatic ligninolysis is to be reproduced in vitro. (a) Lignin is insoluble in water. Its degradation in vivo appears to occur not in a simple aqueous environment, but rather in a more hydrophobic polysaccharide gel, which may promote the swelling or partial dissolution of lignin in the cell wall. The idea that extracellular polysaccharides might play a role in lignin dispersal finds support in earlier work that showed lignin to be readily soluble in glycols (62,63). (b ) H 2 O 2 , the oxidant for LiP, is present at a low concentration in fungal cultures, as shown by the observation that lignin model compounds are not detectably oxidized by crude P. chrysosporium extracellular fluid unless exogenous H 2 O 2 is supplied. These assays are spectrophotometric and sufficiently sensitive to conclude that the concentration of H 2 O 2 in ligninolytic extracellular fluid is <1 µM. (c) Lignin fragments produced during decay are rapidly assimilated and oxidized to CO 2 by the fungus, as shown by the fact that the oxidation of lignin to CO 2 begins simultaneously with depolymerization in vivo (43,44). This uptake process is important because it removes phenolic ligninolysis products, e.g. those produced via LiP-catalyzed cleavage of β-O-4 substructures, that would otherwise be repolymerized by fungal peroxidases or oxidases.
Here we have modeled the first of these conditions, dispersal of the lignin, by conducting enzymatic reactions in aqueous/organic solvent mixtures. The requirement that oxidant be rate-limiting was duplicated by delivering H 2 O 2 slowly throughout the reaction. The third feature, provision of a sink for small lignin fragments to prevent their repolymerization, could not be attempted with the experimental methods presently available. However, it was approximated by performing these reactions at low lignin concentrations. Our experiments show that, under these conditions, ligninolysis was obtainable using LiP in vitro.
To assess the physiological relevance of our data, it is useful to compare them with the results obtained when synthetic lignins are degraded by P. chrysosporium in liquid culture. In such experiments, lignin has three major fates: some is recoverable as organic-soluble degraded polymer, some remains insolubly bound to the fungal mycelium, and some is mineralized (43,44).
Organic-soluble Lignin-With LiP as an oxidant in vitro, the yield of soluble degraded polymer was essentially the same as that recoverable from P. chrysosporium cultures incubated with lignin for 6-24 h (43,44). The soluble lignin was predominantly depolymerized in vitro, as it is in vivo, although some polymerization occurred in the reactions with purified LiP. There is chromatographic evidence that whole P. chrysosporium cultures may transiently polymerize lignin (43, 44), but polymerization was more apparent in the cell-free system. This result is not surprising, considering that the reactions with purified LiP contained no mechanism to remove phenolic low-M r products. are due to T. Kent Kirk, Ewald Srebotnik, and Mark A. Moen for valuable suggestions. Insoluble Lignin-Insoluble polymer was obtained both in complete reactions and in controls from which LiP was omitted. Therefore, although some of the insoluble lignin probably resulted from LiP-catalyzed polymerization, much of it appears to have been noncovalently aggregated polymer. Moreover, the yields of insoluble lignin attributable to LiP in vitro were lower than the yields of insoluble mycelium-bound polymer obtained when synthetic lignin is degraded in whole P. chrysosporium cultures (43, 44). That is, purified LiP did not differ significantly from the intact fungus with regard to the final proportion of lignin that was insoluble and therefore unavailable for M r analysis.
Mineralization -Lignin clearly was not oxidized to CO 2 in the cell-free system. LiP alone is not sufficient for mineralization, which requires the uptake of lignin fragments and participation of intracellular metabolism (1). In sum, LiP alone cannot duplicate the entire process by which P. chrysosporium degrades isolated lignins, but it accounts adequately for the initial steps of this process.
Previous failures to depolymerize lignins with LiP have led some authors to conclude that LiP does not catalyze ligninolysis in vivo (26). However, the reaction conditions in these studies were not optimal. Reactions were conducted without co-solvents (23-26) and lignin concentrations were generally high (23,25,26), leading to poor dispersal of the polymer. H 2 O 2 was added at concentrations >100 µM (23-26), which probably supported high transient concentrations of lignin phenoxy radicals (24) and, therefore, polymerization of the lignin. Veratryl alcohol was usually not included as a cosubstrate (23, 24, 26), whereas it is now clear that this compound plays an important role in LiP catalysis: it protects the enzyme from H 2 O 2 -mediated inactivation (64, 65) and may also act as a one-electron redox shuttle to promote certain LiP-catalyzed reactions (49,66,67). One recent, unsuccessful attempt to depolymerize lignin with LiP was conducted under anaerobic conditions (26), but ligninolysis in vivo is aerobic and is stimulated by increased O 2 concentrations (68). O 2 is an important participant in the oxidation of dimeric lignin model compounds by LiP in vitro: under anaerobic conditions, some of the initial carbon-centered radical cleavage products couple, whereas the rapid addition of O 2 to these radicals under aerobic conditions prevents coupling (14). It is therefore expected that LiP-catalyzed ligninolysis should be inhibited under anaerobic conditions. In short, the negative results obtained previously fail to rule out a ligninolytic role for LiP in vivo. To the contrary, our findings with the purified enzyme strongly support such a role. Important questions remain. Some white-rot fungi appear to degrade lignin without expressing LiP activity, which suggests that there are LiP-independent ligninolytic mechanisms. It is noteworthy in this regard that all such nonproducers yet examined do produce MnPs (69-72), that MnP catalyzes cleavage reactions in certain phenolic lignin model compounds (46,47), and that this enzyme has also been reported to generate low M r fragments from synthetic lignins (73). We have shown here that MnP, like LiP, both polymerized and depolymerized lignin in vitro. However, MnP gave more polymerization and less depolymerization than LiP did and was unable to degrade nonphenolic structures in the polymer. These limitations suggest that MnP is a less versatile degradative agent than LiP is, but it is evident that both peroxidases are fundamentally capable of ligninolysis.