2-Chloro-1,4-Dimethoxybenzene as a Novel Catalytic Cofactor for Oxidation of Anisyl Alcohol by Lignin Peroxidase

Abstract

2-Chloro-1,4-dimethoxybenzene (2Cl-14DMB) is a natural compound produced de novo by several white rot fungi. This chloroaromatic metabolite was identified as a cofactor superior to veratryl alcohol (VA) in the oxidation of anisyl alcohol (AA) by lignin peroxidase (LiP). Our results reveal that good LiP substrates, such as VA and tryptophan, are comparatively poor cofactors in the oxidation of AA. Furthermore, we show that a good cofactor does not necessarily serve a role in protecting LiP against H₂O₂ inactivation. 2Cl-14DMB was not a direct mediator of AA oxidation, since increasing AA concentrations did not inhibit the oxidation of 2Cl-14DMB at all. However, the high molar ratio of anisaldehyde formed to 2Cl-14DMB consumed, up to 13:1, indicates that a mechanism which recycles the cofactor is present.

White rot fungi are involved in the extensive degradation of lignin by means of their extracellular ligninolytic system (20). Key enzymes involved in the lignin-degrading system are extracellular peroxidases which are directly responsible for the oxidative depolymerization of lignin (12). Lignin peroxidase (LiP) plays an important role in the degradative ability. LiP can oxidize substrates with a higher ionization potential than other peroxidases (18). This capability enables the enzyme to oxidize nonphenolic methoxylated aromatic compounds (17). LiP also catalyzes the oxidation of recalcitrant xenobiotic compounds such as polycyclic aromatic hydrocarbons (10), dioxins (37), chlorophenols (11), and azo dyes (25, 26).

The catalytic cycle of LiP is like those of other peroxidases. LiP is activated in the presence of H₂O₂ to form compound I. This intermediate is able to catalyze a one-electron oxidation of numerous substrates, forming compound II. The cycle is completed by an additional one-electron oxidation of a limited number of substrates, causing the reduction of compound II back to the enzyme’s ferric state (31). However, in the absence of reducing substrate, compound II can undergo a series of reactions with H₂O₂ to form compound III, an inactive form of the enzyme (38).

In the presence of veratryl alcohol (VA), a secondary metabolite of several ligninolytic white rot fungi (7, 23), the formation of compound III is prevented (39). VA is a favorable substrate for compound II and converts it to the resting state, completing the catalytic cycle (21, 27). Nonphenolic monomethoxylated lignin model compounds, such as anisyl alcohol (AA), are poorly oxidized by LiP. Inclusion of VA in the reaction mixture accelerated the oxidation of AA (13). Koduri and Tien (21) showed that AA can be oxidized only by compound I. VA is required as an essential cofactor for oxidation by compound II, allowing the enzyme to return to its ferric state (21). Furthermore, a secondary consequence of the cofactor role is that VA prevents the inactivation of the enzyme by excess H₂O₂ (38).

VA has also been implicated as a redox mediator of LiP for substrates with a lower ionization potential than VA itself. VA is oxidized by one electron to form the VA cation radical (VA⁺·) (4, 13). VA⁺· was suggested to oxidize other substrates at a distance from the active site of the enzyme (13). However, the free VA cation radical is too short-lived (half-life, 0.5 ms) to diffuse away from the enzyme (16, 19). Therefore, it was suggested that an enzyme-bound radical was formed (4). Khindaria et al. (19) demonstrated the presence of a LiPII-VA⁺· complex, which has a half-life of 0.54 s, implying a more stable VA cation radical. The enzyme-bound radical could be more reactive because it is longer-lived or because it has a higher oxidation potential than the free VA⁺· (4).

It is possible that white rot fungi produce alternative metabolites which could serve as cofactors or mediators of LiP catalysis. Several other compounds were found to substitute for VA as reducing agents for compound II in AA oxidation: 3,4-dimethoxytoluene, 1,4-dimethoxybenzene (14DMB), and 3,4,5-trimethoxybenzyl alcohol (16, 21). Collins et al. (5) also introduced tryptophan (Trp) as an alternative cofactor for VA in LiP catalysis; Trp stabilizes the enzyme against H₂O₂ inactivation even better than VA. Moreover, 14DMB is oxidized to a cation radical, as was demonstrated by electron spin resonance spectroscopy, indicating the relative stability of this cation radical (17) and implying a possible function as a diffusible mediator in LiP oxidations (16). However, 14DMB is not naturally produced by white rot fungi.

Chlorinated 1,4-dimethoxybenzenes, structurally related to 14DMB, are naturally produced by several white rot fungi with ligninolytic activity. Examples of such metabolites include 2-chloro-1,4-dimethoxybenzene (2Cl-14DMB), 2,6-dichloro-1,4-dimethoxybenzene (26DCl-14DMB), tetrachloro-1,4-dimethoxybenzene (designated drosophilin A methylether [DAME]), and tetrachloro-4-methoxyphenol (designated drosophilin A [DA]) (8, 28–30). Consequently, chlorinated 1,4-dimethoxybenzenes can possibly serve the same function as VA in LiP catalysis. In this report we have identified 2Cl-14DMB as an alternative for VA in LiP catalysis. We demonstrate that 2Cl-14DMB is a cofactor superior to VA and 14DMB in the LiP-catalyzed oxidation of AA.

MATERIALS AND METHODS

Organism and media.

Bjerkandera sp. strain BOS55 (ATCC 90940) was isolated and maintained as previously described (6). Inoculum was prepared on malt extract plates containing, per liter, 5 g of glucose, 3.5 g of malt extract (Oxoid Ltd., Basingstoke, Hampshire, United Kingdom), and 15 g of agar. To obtain high LiP activity, Bjerkandera sp. strain BOS55 was grown in a high-nitrogen medium containing, per liter, 10 g of glucose, 5 g of mycological peptone (Oxoid), 1 g of yeast extract (Gibco BRL, Paisley, United Kingdom), 0.2 g of diammonium tartrate, 20 mmol of 2,2-dimethylsuccinate, 2 mmol of VA, 200 mg of thiamine, and 100 ml of modified BIII mineral medium (32). The modified BIII medium contained 54.4 g of KH₂PO₄ per liter and no manganese. After autoclaving, the pH was adjusted to 6.0 with 2 M autoclaved NaOH.

Cultivation conditions on liquid medium.

For the purification of LiP, Erlenmeyer flasks (5 liter) each containing 1 liter of standard medium were inoculated with five cylindrical agar plugs (diameter, 5 mm), which were taken from the outer periphery of a malt extract agar plate covered with mycelium of Bjerkandera sp. strain BOS55 incubated at 30°C for 6 days. The Erlenmeyer flasks were left to grow statically in the dark at 30°C for 20 days.

LiP preparations.

LiP was purified from the extracellular fluid of Bjerkandera sp. strain BOS55 cultures. The proteins were concentrated by ammonium sulfate precipitation (85% saturation). The concentrate was dialyzed against 10 mM sodium acetate buffer (pH 6.0). The dialyzed fraction was further purified on a Resource Q anion-exchange column (Pharmacia, Woerden, The Netherlands) with a gradient of 10 mM to 1 M sodium acetate (pH 6.0). The first LiP-containing fraction was further purified on a Source S cation-exchange column (Pharmacia) in order to separate LiP from aryl alcohol oxidase. The purity of the LiP isozyme was confirmed by sodium dodecyl sulfate-polyacrylamide gel electrophoresis. Also, a partially purified LiP preparation from Phanerochaete chrysosporium obtained from Tienzyme, Inc. (State College, Pa.) was used in several experiments. LiP activity is expressed in units. One U of LiP activity was defined as the amount of enzyme required to oxidize 1 μmol of VA per min.

Chlorinated compounds as substrates of LiP.

VA, 14DMB, 2Cl-14DMB, 26DCl-14DMB, DAME, and DA were tested as substrates for LiP. 14DMB and the chlorinated 14DMB derivatives were dissolved in acetone. The maximum acetone concentration in the final reaction mixture for the experiments was 5%. The reaction mixture was composed of the following: 500 μM substrate, 0.1 U of purified LiP of Bjerkandera sp. strain BOS55, and 0.25 mM H₂O₂ in 20 mM sodium succinate (pH 3.0). Assay volumes were adjusted to 1 ml with distilled H₂O. After 90 min of incubation at 30°C, the reaction was stopped with the addition of 1 ml of acetonitrile and the products were analyzed by high-pressure liquid chromatography (HPLC).

H₂O₂ inactivation.

The protective effects of VA and 2Cl-14DMB against LiP inactivation by high concentrations of H₂O₂ were examined. Assay mixtures were composed of the following: 2Cl-14DMB (25, 50, 100, 500, or 2,000 μM), LiP (initial activity, 0.1 U/ml), H₂O₂ (final concentration in assay mixture, 0.1 mM), and 100 mM sodium acetate buffer (pH 5.0). Assay volumes were adjusted to 1 ml with distilled H₂O. After 0, 2, 5, 8, 11, 14, 17, and 20 min of incubation at room temperature, 100-μl aliquots were removed from assay mixtures and their VA-oxidizing activities were measured (as described in “Enzyme assays”). Data are expressed as percentages of the initial LiP activities remaining.

AA oxidation.

The stimulating effect of 2Cl-14DMB on the oxidation of AA was examined. The assay mixture was composed of 500 μM AA, 20 mM sodium succinate (pH 3.0) (99% purity), 2 to 500 μM 2Cl-14DMB, 0.1 U of purified LiP from Bjerkandera sp. strain BOS55, and 250 μM H₂O₂. The assay volume was adjusted to 1 ml. The incubation time of the assay mixture was 90 min. The reaction was stopped by the addition of 1 ml of acetonitrile to the assay mixture, and the samples were analyzed by HPLC.

Cofactor recycling.

The effects of succinic acid, acetone, and bovine serum albumin (BSA) on the recycling of 2Cl-14DMB were examined. The assay mixture was composed of 500 μM AA, 0 to 40 mM sodium succinate (pH 3.0) (99% purity), 50 μM 2Cl-14DMB, 0.1 U of purified LiP from Bjerkandera sp. strain BOS55, and 250 μM H₂O₂. The reaction was stopped by the addition of 1 ml of acetonitrile to the assay mixture, and the samples were analyzed by HPLC for succinic acid elimination or by gas chromatography for CO₂ production. When acetone (0.05 to 5% of assay volume) or BSA (0 to 50 mg/liter) was added, 20 mM sodium succinate was used. The assay volume was adjusted to 1 ml. The incubation time of the assay mixture was 90 min. The reaction was stopped by the addition of 1 ml of acetonitrile to the assay mixture, and the samples were analyzed by HPLC.

Enzyme assays.

LiP activity was measured by monitoring the oxidation of VA to veratraldehyde (VAld) at 310 nm (ɛ = 9,300 M⁻¹ cm⁻¹) as described by Tien and Kirk (32) and corrected for background VA oxidase activity. VA oxidase activity was measured by monitoring the oxidation of VA to VAld at 310 nm without the addition of 0.4 mM H₂O₂.

Gas chromatographic analysis.

The methanol concentration was measured on a gas chromatograph. Samples were adjusted to pH 2.0 with 3% formic acid and centrifuged for 5 min (at 17,000 × g). Methanol was determined by using a Packard Becker model 417 (Delft, The Netherlands) gas chromatograph equipped with a 6-m by 2-mm glass column packed with a Supelco port (Bellefonte, Pa.), 100/120 mesh, coated with 10% Fluorad FC431 (3M, St. Paul, Minn.). The flow rate of the carrier gas (nitrogen saturated with formic acid) was 30 ml min⁻¹, and column pressure was ±3 × 10⁵ Pa. The column temperature was 70°C, and the injection port and detector were at 220 and 280°C, respectively.

CO₂ concentrations were determined by using a Packard model 427 gas chromatograph with a Hayesep Q column (Chrompack, Middelburg, The Netherlands). Headspace samples (50 μl) were analyzed.

HPLC analysis.

Fifty microliters of the incubation mixtures was analyzed for products by HPLC as described previously (30) with the column (200 by 3 mm) filled with ChromSpher C₁₈-PAH (5-μm particles) (Chrompack). Aromatic metabolites were analyzed with the following gradient (0.4 ml min⁻¹, 30°C): 90:10, 0:100, and 0:100 H₂O to CH₃CN at 0, 15, and 20 min, respectively. The UV absorbance was monitored at 2-nm wavelength intervals from 200 to 400 nm. Compound identification was carried out by matching UV spectra and the retention times of the observed products with their standards.

Organic acid determination.

Organic acid concentration was measured by HPLC. HPLC analysis was performed on an Aminex HPX-87H column (Bio-Rad, Veenendaal, The Netherlands). Samples were eluted with 5 mM H₂SO₄ at a flow rate of 0.6 ml/min at 40°C, and detection was at 210 nm. Compound identification was carried out by matching retention times of samples with their standards.

Reference compounds.

Standards for 26DCl-14DMB, DA, and DAME were kindly provided by J. Knuutinen, Department of Chemistry, University of Jyväskylä, Jyväskylä, Finland. 2Cl-14DMB and 14DMB were obtained from Janssen Chimica. VA was obtained from Aldrich.

Statistical procedures.

In all experiments, measurements were carried out in triplicate. Values reported are means with standard deviations.

RESULTS

Methoxybenzenes as substrates for LiP.

VA, 14DMB, and several chlorinated derivatives of 14DMB were compared as substrates of semipurified LiP from P. chrysosporium and purified LiP from Bjerkandera sp. strain BOS55 (Table 1). Of the nonphenolic compounds tested, VA, 14DMB, and 2Cl-14DMB were found to be substrates of LiP. VA and, to a lesser extent, 14DMB were observed to be relatively good substrates, whereas 2Cl-14DMB was a poor substrate. The more chlorinated nonphenolic compounds, 26DCl-14DMB and DAME, were not oxidized by the LiP preparations. Inclusion of VA (50 or 500 μM) in the reaction mixture did not enable these compounds to become oxidized by LiP, nor did VA alter the extent of 14DMB and 2Cl-14DMB oxidation (results not shown). The theoretical maximum conversion of the nonphenolic compounds (donating two electrons per molecule) was 50%, since they were supplied in a twofold excess of the H₂O₂ (accepting two electrons per molecule). The fact that VA was oxidized up to 63% indicates that there was probably either some endogenous production of peroxide or another electron acceptor during the reaction. The only phenolic compound tested, DA, was a good substrate for the LiP preparations.

TABLE 1.

Oxidation of VA, 14DMB, and several chlorinated derivatives of 14DMB by semipurified LiP from P. chrysosporium and a purified LiP isozyme of Bjerkandera sp. strain BOS55

Compound	% Oxidationa		Identified product(s)	Molar product yield (%)b
	Semipurified LiP	Purified LiP		Semipurified LiP	Purified LiP
VA	54	63	Veratraldehyde	100	83
14DMB	41	24	1,4-Benzoquinone	100	69
			Methanol	NM	160
2Cl-14DMB	13	11	2-Chloro-1,4-benzoquinone	100	35
			Methanol	NM	76
26DCl-14DMB	0	0c	Not detected
DAME	0	0		0	0
DA	73	53	Unidentified

^aCalculated as (initial concentration of substrate − final concentration of substrate/initial concentration of substrate) × 100. 

^bCalculated as (product concentration/initial concentration of substrate − final concentration of substrate) × 100. NM, not measured. 

^cIn some incubations, 2 to 5% 26DCl-14DMB was consumed. 

The major product of VA was VAld, whereas the major products of the 14DMB derivatives were the corresponding quinones (1,4-benzoquinone and 2-chloro-1,4-benzoquinone) and methanol. The molar ratio of methanol to quinone formed was approximately 2:1.

The molar product yields obtained from semipurified and purified LiP were different. The results with semipurified LiP showed a complete conversion of the consumed substrates to the identified products (molar product yield, 100%). The comparative product yield with purified LiP was lower, indicating the formation of other products besides those that were identified in these experiments. No products were detected in reactions lacking either enzyme or H₂O₂ or with boiled enzyme.

2Cl-14DMB in AA oxidation.

The abilities of LiP substrates to act as cofactors in the catalysis of purified LiP were examined. Relatively good substrates such as VA and 14DMB (Table 1), as well as tryptophan (5), were compared with the poor substrate 2Cl-14DMB. These compounds were tested at several concentrations in the range from 20 to 200 μM as shown in Fig. 1. Trp, which was clearly the best LiP substrate, had no role in improving the background level of AA oxidation. The next best substrate, VA, supported limited enhancement of AA oxidation up to 50 μM; however, there was no significant increase in the amount of anisaldehyde (AAld) formed as the VA concentration was increased further up to 200 μM. Both 14DMB and 2Cl-14DMB were better cofactors. The extent to which AA was oxidized increased with the cofactor concentration. 2Cl-14DMB was clearly the worst LiP substrate and the best cofactor, supporting the highest conversion of AA at any given cofactor concentration. 26DCl-14DMB, which is not a substrate of LiP, was also tested and was found not to have any cofactor effect (results not shown).

FIG. 1.

Effects of the concentrations of VA (⧫), 14DMB (▴), 2Cl-14DMB (▪), and Trp (×) on AAld formation (A) and the concomitant consumption of the cofactors (B) during AA oxidation.

The stoichiometric relationship between the AAld formed (corrected for the background formation) and the cofactor consumed is given in Table 2. This ratio reached a maximum of 2 for VA and 14DMB at the lowest cofactor concentration tested. But the ratio approached 1 or less at higher cofactor concentrations, indicating that VA and 14DMB are for the most part noncatalytic cofactors (just one turnover). On the other hand, this ratio was distinctly higher for 2Cl-14DMB, ranging between 3 and 13 in various experiments. Thus, 2Cl-14DMB is a cofactor which is catalytic; each molecule consumed supports multiple turnovers of the enzyme for AA oxidation.

TABLE 2.

Effects of increasing concentrations of VA, 14DMB, and 2Cl-14DMB on the oxidation of AA and the stoichiometric ratio of AAld to consumed cofactor

Cofactor and concn (μM)	% AA oxidation	% Oxidation of cofactor	Stoichiometric ratio, AAld formation/consumed cofactora
None	2
VA
20	3	50	2:1
50	4	60	1:1.5
200	10	80	1:3
14DMB
20	5	74	2:1
50	11	81	1.5:1
200	26	71	1:1
2Cl-14DMB
20	10	22	8:1 (11:1)
50	21	25	5:1 (13:1)
200	40	40	5:1 (3:1)

^aValues in parentheses are results observed in additional experiments. 

The cofactor role of 2Cl-14DMB was examined further by testing a larger range of concentrations and comparing the oxidation of substrates with the consumption of H₂O₂ (Fig. 2). AA oxidation increased with elevated 2Cl-14DMB concentrations up to 200 μM. Thereafter, the H₂O₂ supply became limiting and further increases in the cofactor concentration only had the effect of stealing H₂O₂ away from AA oxidation, thereby causing some decreases in the AAld formed. Throughout the entire cofactor concentration range considered, the sum of cofactor and AA oxidized was approximately 20% higher than the H₂O₂ consumption, suggesting that some endogenous production of H₂O₂ was occurring or that an alternative electron acceptor was present. The molar yield of 2-chloro-1,4-benzoquinone per mol of 2Cl-14DMB consumed was 100% at low cofactor concentrations but decreased to 70% at higher concentrations.

FIG. 2.

Effect of varying concentrations of 2Cl-14DMB on AAld formation (▪). Also shown are the concomitant consumption of 2Cl-14DMB (▴), the formation of the 2Cl-14DMB oxidation product 2-chloro-1,4-benzoquinone (2-Cl-BQ) (×), and the H₂O₂ used by the system (⧫).

The effect of increasing AA concentrations on the consumption of 50 μM 2Cl-14DMB by purified LiP was evaluated (Fig. 3). Irrespective of the AA concentration from 0 to 2,000 μM, AA had no effect on the extent to which the cofactor was consumed. Thus, AA did not inhibit 2Cl-14DMB oxidation by LiP. However, the production of AAld increased with increasing concentrations of AA.

FIG. 3.

Effect of varying concentrations of AA on the consumption of 2Cl-14DMB (▪) and the formation of AAld (⧫).

Effects of succinic acid, acetone, and protein on 2Cl-14DMB oxidation.

Assay components were examined for the ability to function as an electron donor for 2Cl-14DMB⁺·, reducing it back to 2Cl-14DMB. The effect of increasing succinic acid concentrations from 0 to 40 mM on the consumption of 50 μM 2Cl-14DMB was tested. Succinic acid had no effect on the extent to which 2Cl-14DMB was consumed. Furthermore, no CO₂ production or elimination of succinic acid was detected (results not shown). Acetone, which is present as a solvent for 2Cl-14DMB in the assay mixture, could possibly act as an electron donor. Therefore, increasing percentages of acetone from 0.05 to 5% of the assay volume were examined. However, none of the tested concentrations had an effect on the consumption of 2Cl-14DMB.

Noncatalytic parts of the protein could chemically interfere with the oxidation of 2Cl-14DMB. However, increasing concentrations of BSA, ranging from 0 to 50 mg/liter, added to the assay mixture did not affect the consumption of 2Cl-14DMB.

2Cl-14DMB as a protector against LiP inactivation by H₂O₂.

Only 2 mM 2Cl-14DMB partially protected LiP against inactivation by high concentrations of H₂O₂ (Fig. 4). However, the protective effect was not as good as that of 2 mM VA, which almost completely protected LiP from inactivation in the time period considered. Concentrations below 2 mM 2Cl-14DMB did not have a protective effect on LiP activity at all. On the contrary, 100 μM 2Cl-14DMB decreased LiP activity to a point below that observed when no 2Cl-14DMB was added to the reaction mixture. This was also observed for 25 and 50 μM 2Cl-14DMB. As shown in Fig. 5, when no 2Cl-14DMB or VA was added, 80% of the activity of purified LiP was gone after 8 min, but addition of 25, 50, or 100 μM 2Cl-14DMB caused a 90 to 100% inactivation of LiP activity. Similar results were obtained with semipurified P. chrysosporium LiP. Although the inactivation of LiP after 8 min did not proceed as far as that for purified Bjerkandera sp. strain BOS55 LiP, the same pattern of inactivation was observed. These results suggest that small amounts of 2Cl-14DMB stimulate the inactivation of LiP.

FIG. 4.

Protective effects of 100 μM (×) and 2 mM (▪) 2Cl-14DMB and of 2 mM VA (▴) against inactivation of purified LiP by 0.1 mM H₂O₂. ⧫, no 2Cl-14DMB added to reaction mixture.

FIG. 5.

Inactivation of purified (⧫) and semipurified (▪) LiP after 8 min of treatment with 0.1 mM H₂O₂ in the presence of varying concentrations of 2Cl-14DMB.

DISCUSSION

2Cl-14DMB is a 14DMB derivative which is produced de novo by Bjerkandera adusta and Lepista nuda (14, 28, 29). So far only trace amounts of the compound have been found in the ligninolytic cultures. In L. nuda growing on forest litter, up to 0.2 mg of 2Cl-14DMB/kg has been detected (14). In this report we show that this naturally produced chloroaromatic is a cofactor superior to VA and 14DMB in the oxidation of AA.

Naturally produced chlorinated 14DMB derivatives were compared as substrates for LiP with VA and 14DMB. LiP catalyzed the oxidation of VA, 14DMB, 2Cl-14DMB, and DA (Table 1). Oxidation of VA by LiP yielded VAld as the major product (83% of oxidized VA), as was also found by Joshi and Gold (16) and others (7). 14DMB and 2Cl-14DMB oxidation yielded the corresponding benzoquinones as major products (respectively, 69 and 35% of consumed substrates), although other products, which were not identified in this study, must have been formed as well. Joshi and Gold (16) reported the formation of 2-(2,5-dimethoxyphenyl)-1,4-benzoquinone as a major product of 14DMB, whereas Kersten et al. (17) only reported the formation of 1,4-benzoquinone. Previously, it was observed that 2Cl-14DMB was oxidized to 2-chloro-1,4-benzoquinone as a major product and that, to a lesser extent, 2,5-dimethoxy-1,4-benzoquinone and 3-chloro-4-methoxy-1,2-benzoquinone were formed (36). 26DCl-14DMB and DAME were not oxidized by LiP. Previous research showed that 2,5-dichloro-1,4-dimethoxybenzene also was not oxidized by LiP (15). Obviously, more chloro groups decrease the reactivity of the compound. The ionization potential of dimethoxybenzenes increases when more electron-withdrawing chloro groups are present; values of 8.55, 8.69, 8.81, and 8.95 were calculated for 14DMB, 2Cl-14DMB, 26DCl-14DMB, and DAME, respectively. These values indicate that the one-electron removal from highly chlorinated compounds becomes progressively more difficult. DA, however, is fairly well oxidized by LiP to a yet unidentified product. It has been shown that pentachlorophenol is also oxidized by LiP, with tetrachlorobenzoquinone as the major product (11, 24). Phenol oxidation proceeds via formation of a phenoxy radical, which occurs more easily than the formation of cation radicals from methoxybenzenes.

Although 2Cl-14DMB was the worst substrate for LiP, it was, surprisingly, found to be the best cofactor in AA oxidation. Our results reveal an inverse relation between the ability to be oxidized by LiP and the ability to act as a cofactor for the oxidation of the monomethoxylated lignin model substrate AA. The good substrates VA and 14DMB were worse cofactors than 2Cl-14DMB, whereas the excellent LiP substrate Trp (5) was not a cofactor at all (Fig. 1). VA was a good cofactor only at low concentrations. Although good substrates can close the catalytic cycle, because they can react with both compound I and compound II, they can also compete with AA oxidation by compound I. Previous research with VA showed that increasing VA concentrations compete with AA for oxidation with compound I, eventually leading to a decrease in AA oxidation (21). Trp is an excellent substrate for LiP; Collins et al. (5) suggested that Trp is even a better substrate for compound II than VA. Trp did not enhance AA oxidation at all, suggesting that AA oxidation was completely competitively inhibited by Trp oxidation. Probably 2Cl-14DMB cannot compete very well with AA for oxidation with compound I, but is primarily oxidized by compound II.

Good LiP substrates are more effective in protecting LiP against H₂O₂ inactivation. VA has previously been shown to extend the half-life of LiP in fungal cultures (34), whereas Collins et al. (5) showed the superior protective effect of Trp compared to VA. Low concentrations of 2Cl-14DMB, by comparison, did not protect LiP against high H₂O₂ concentrations; only 2 mM 2Cl-14DMB partially protected LiP (Fig. 4). Our results show that a good cofactor does not necessarily serve a role in protecting against H₂O₂ inactivation as proposed by Valli et al. (35).

In fact, we found that low 2Cl-14DMB concentrations even increased LiP inactivation by H₂O₂. One possible explanation for this phenomenon is that 2Cl-14DMB is a much better substrate for compound I than for compound II. At low concentrations, 2Cl-14DMB stimulates the formation of compound II, which in turn reacts with H₂O₂ to form compound III, whereas at high concentrations, 2Cl-14DMB progressively becomes a better reductant for compound II and likewise there is more cation radical available to restore compound III, as was shown for VA⁺· (2). VA⁺· can overcome compound III accumulation by converting it back to active ferric LiP (2). This was also shown for the 1,2,4,5-tetramethoxybenzene cation radical (3).

Although 2Cl-14DMB did not have a protective effect on LiP, the compound clearly stimulated AA oxidation. Unlike VA and 14DMB, 2Cl-14DMB is a catalytic cofactor; each molecule consumed supported multiple turnovers of the enzyme for AA oxidation. Our results show that 2Cl-14DMB is not a direct mediator in AA oxidation. If mediation had occurred, the presence of increasing AA concentrations should have completely inhibited the consumption of 2Cl-14DMB oxidation, as was found for VA in the oxidation of guaiacol, 4-methoxymandelic acid, and chlorpromazine (9, 22, 33). However, 2Cl-14DMB consumption was not inhibited by AA at all (Fig. 3). This result also suggests that 2Cl-14DMB and AA do not have the same binding site.

The molar ratio of AAld formed to cofactor consumed ranged from 3 to 13 (Table 2). Probably a mechanism is present which recycles the 2Cl-14DMB cation radical (2Cl-14DMB⁺·) back to 2Cl-14DMB. As indicated above, 2Cl-14DMB did not directly mediate the oxidation of AA. A second possibility is that H₂O₂ reacts with 2Cl-14DMB⁺·, as was described for VA⁺· (1). The one-electron reduction of the cation radical back to 2Cl-14DMB would result in net O₂ production from H₂O₂ (1). In such a case, the H₂O₂ consumption would exceed the sum of oxidized 2Cl-14DMB and AA; however, the H₂O₂ consumption was 20% lower than the sum of the oxidized compounds (Fig. 2). Consequently, other assay components which might be able to reduce 2Cl-14DMB⁺· back to 2Cl-14DMB were considered. However, none of these components, succinate, acetone, and noncatalytic protein, were found to affect the oxidation of 2Cl-14DMB, suggesting that the reduction of 2Cl-14DMB⁺· is carried out somewhere in the catalytic cycle of LiP or possibly by reduced oxygen radicals, which should be confirmed by further research.

In conclusion, this work demonstrates that 2Cl-14DMB is a catalytic cofactor superior to VA. Although so far only trace amounts of 2Cl-14DMB have been found in ligninolytic cultures (14, 28, 29), we showed that only small amounts of 2Cl-14DMB are necessary to exert a major increase in AA oxidation. As the molar ratio of AA oxidation compared to cofactor oxidation is so high, the cofactor must be recycled in the reaction.