Comparison of ligninase-I and peroxidase-M2 from the white-rot fungus Phanerochaete chrysosporium

ARCHIVES

OF BIOCHEMISTRY

Vol. 244, No. 2, February

AND

BIOPHYSICS

1, pp. 750-765,1986

Comparison of Ligninase-I and Peroxidase-M2 from the White-Rot Fungus Phanerochaete chrysosporkm’ ANDRZEJ

PASZCZYNSKI,*

VAN-BA

HUYNH,t

AND

RONALD

CRAWFORDp2

*University M. Curie-Sklodousku, Department of Biochemistry, 20-031 Dublin, Poland, and tGray Freshwater Biological Institute and Department of Microbiology, University of Minnesota, Navarre, Minnesota 55592 Received

July

1’7,1985,

and in revised

form

September

25,1985

Ligninase-I (Mr 42,000-43,000; carbohydrate, 21% ) and peroxidase-M2 (Mr 45,00047,000; carbohydrate, 17%), two representative, hydrogen peroxide-dependent extracellular enzymes produced by ligninolytic cultures of the white-rot fungus Phanerochuete chrgsosporium BKM-F-1767, were purified and their properties compared. Spectroscopic studies showed that both native enzymes are heme proteins containing protoporphyrin IX. EPR spectroscopy indicated that iron ions are coordinated with the enzymes’ prosthetic groups as high-spin ferriheme complexes. We confirmed reports of others that the ligninase-hydrogen peroxide complex (activated enzyme) reverts to its native state on addition of dithionite or one of the enzyme’s substrates (e.g., veratryl alcohol); however, we found that the peroxidase-M2-hydrogen peroxide complex required Mn2+ ions to accomplish a similar cycle. The peroxidase oxidized Mn2+ to a higher oxidation state, and the oxidized Mn acted as a diffusible catalyst able to oxidize numerous organic substrates. Unlike ligninase-I which is found free extracellularly, peroxidase-M2 appears to be associated closely with the fungal mycelium. In its peroxidatic reactions, ligninaseI oxidizes a variety of nonphenolic and phenolic lignin model compounds. In the presence of Mn2+, peroxidase-M2 oxidizes numerous phenolic compounds, especially syringyl(3,5dimethoxy-4-hydroxyphenyl) and vinyl side-chain substituted substrates. Also, the peroxidase-Mn2+ system (without hydrogen peroxide) expresses oxidase activity against NADPH, GSH, dithiothreitol, and dihydroxymaleic acid, forming hydrogen peroxide at the expense of oxygen. Both enzymes were believed to play roles in lignin degradation, and these are discussed. o 1386 Academic press, k.

Lignin is the earth’s second most abundant plant polymer, ranking only behind cellulose in quantity as a natural biopolymer (7). It is composed primarily of phenylpropanoid monomeric units, interconnected by a complex array of stable carboncarbon and carbon-oxygen (etheric) bonds (37). Lignin is both stereochemically complex and of high molecular weight. Rela-

tively few microorganisms can degrade this macromolecule. Recent studies have shown that woodrotting fungi of the white-rot variety synthesize a battery of extracellular heme proteins that act as nonspecific, hydrogen peroxide-dependent catalysts and probably comprise what is known as the ligninolytic system (2, 5, 10, 12, 16, 20, 26, 34, 43, 44). Two primary types of heme proteins have been identified in studies of the ligninolytic system of the fungus Phanerochaete chugsoap&urn. One type includes multiple forms of a diarylpropane oxygenase that is able to cleave side-chain carbon-carbon

r This work was supported by Grant PCM-8318151 from the National Science Foundation (R. Crawford and V. B. Huynh), and a postdoctoral fellowship from the 3M Company (St. Paul, Minn.) to A. Paszczynski. ’ To whom correspondence should be addressed.

0003-9861/86 Copyright All righta

$3.00

0 1986 by Academic Press, Inc. of reproduction in any form reserved.

750

PEROXIDASE

CHARACTERIZATION

bonds in lignin, and to perform a variety of other oxidative transformations of lignin and lignin model compounds (12, 19, 35, 38, 44). These glycoproteins have been referred to as “ligninases” by workers in the field. A second group of extracellular heme proteins includes two or more manganese-dependent peroxidases whose role in lignin degradation is less clear (16, 24, 34). These enzymes act as classical peroxidases, oxidizing low-molecular-weight phenols, thereby forming phenoxy radicals, but also catalyze additional reactions including hydrogen peroxide-independent oxidations of molecules like reduced glutathione and NADPH (34). Here we describe isolation and characterization of one representative of each class of lignin degradation-associated peroxidases from P. chrgsosporium. One enzyme is the predominant “ligninase” of strain BKM-F-1767. As it probably is the same enzyme studied by Tien and Kirk (44) and Gold et al. (12), we refer to this protein as ligninase-I. The second enzyme is the Mn2+-dependent peroxidase described in a preliminary publication by Paszczynski et aL (34). We refer to this enzyme as peroxidase-M2. We compare these two enzymes with respect to their physical and biochemical properties, providing more and different types of information than has been provided in previous publications. EXPERIMENTAL

PROCEDURES

Organism and culture conditions. P. chrysosporium Burds strain BKM-F-1767 (ATCC 24725) was maintained and spore inocula were prepared as described previously (16). The white-rot fungus was grown in a defined medium, modified slightly from that of Kirk et al. (23), that contained 2.2 mM nitrogen and 10 g/ liter glucose buffered at pH 4.5 with 10 mM Na-2,2dimethylsuccinate. Mycelium was cultivated attached to the roughened interior walls of a 20-liter carboy (polyethylene; 29 cm in diameter, 40 cm long) containing 1 liter of medium. The carboy was filled with pure oxygen, turned on its side, and rotated (1 rpm) in a 40°C incubator room. One day prior to harvest, 1 mM veratryl alcohol was added to the growth medium. Culture fluid was harvested when the ligninolytic enzyme system of the fungus was maximally aetive as determined by assaying the ligninase (hereafter referred to as ligninase-I) and the peroxidase

751

(hereafter referred to as peroxidase-M2; for assay conditions, see the following). Enzyme puri&ation. Harvested culture fluids were filtered through glass wool to remove mycelial fragments, and proteins in the filtrate were concentrated and purified essentially as described by Paszczynski et al (34). Briefly, these procedures involved ultrafiltration (10,000 il4, pore size membrane), dialysis against 0.025 M imidazole-HCl buffer (pH 7.4), and chromatofocusing column chromatography on Polybuffer Exchanger PBE-94 (Pharmacia, Uppsala, Sweden), eluting the column with simultaneous NaCl and pH gradients. Details of these procedures are outlined under Results. Enzyme assays. Peroxidase-MX was assayed as described previously using vanillylacetone [4-(4-hydroxy-3-methoxyphenyl)-3-buten-2-one] as substrate (16). Briefly, disappearance of vanillylacetone was monitored spectrophotometrically at 336 nm. Assay solutions contained 0.1 mM substrate, 0.1 mM manganese sulfate, 0.05 mM hydrogen peroxide, and 100 mM Na-tartrate (pH 5.0) plus enzyme and Hz0 to a final volume of 1 ml. Assays were initiated by addition of hydrogen peroxide. The amount of peroxidase required for oxidation of 1 rmol of vanillylacetone per minute was defined as one unit (U) of activity. In studies of substrate specificity of the peroxidase, assays were run similarly except that different substrates were employed and different wavelengths specific to the substrate being examined were monitored (see Results). For enzyme inhibitor studies, oxidation of vanillylacetone was monitored as usual, except that potential enzyme inhibitors were added to the assay mixture at 0.1 mM, and the reaction rate was compared to that obtained in the absence of inhibitor. Ligninase-I was assayed essentially as detailed by Tien and Kirk (44), except that Tween 80 was eliminated from the assay mixture and less hydrogen peroxide (50 pM) was employed. This assay involves monitoring at 310 nm the conversion of veratryl alcohol (3,4-dimethoxybenzylalcohol) to veratryl aldehyde in the presence of ligninase and hydrogen peroxide. Oxidase activity of peroxidase-M2 was measured by monitoring oxygen uptake in the presence of enzyme, substrate, and manganese ions. Oxygen concentration was determined using a Gilson Clark-type oxygen electrode (Gilson Medical Electronics, Middleton, Wise.). Reaction mixtures contained, in a total volume of 2.0 ml, 0.4 ml of 0.5 M Na-tartrate (pH 5.), 0.2 ml of 10 mM substrate, and 0.2 ml of 1.0 mM manganese sulfate at 37’C with water and enzyme to volume. The electrode was calibrated as described previously (36). Ckmihtion of Mn*’ by peroxiakse-M2. To examine the oxidation of Mn2+ by peroxidase-M2, enzyme, hydrogen peroxide, and manganese sulfate were mixed in 0.1 M

752

PASZCZtiSKI,

HUYNH,

Na-tartrate (pH 5.0) and incubated for 5 min at room temperature. The reaction mixture was then deproteinized by passing it through a 1 X lo-cm Sephadex G-25 column (Pharmacia PD-10). The deproteinized solution was yellow to brown in color. Spectroscopy. Ultraviolet/visible absorption spectra of ligninase-I and Peroxidase-M2 were obtained using either a Beckman Model 25 or a Hewlett-Packard Diode Array Model 345OA spectrophotometer. Spectra were obtained at room temperature in I-cm quartz cuvettes containing enzyme (0.2 mg/ml protein) dissolved in 50 mM Na-tartrate (pH 5.0). Other conditions were as described by Gold et al (12). Heme contents of the enzymes were determined by measuring the absorptivity of the heme-pyridine complex, prepared according to Pajot and Groundinsky (31). Absorptivity was measured at 557 nm using an extinction coefficient of 32,550 M-’ cm-’ for the pyridine-heme adduct. EPR spectroscopy was performed in 50 mM Na-tartrate buffer (pH 5.0). The peroxidase was added to give 0.35 mg/ml protein; ligninase-I was added to 0.4 mg/ml. Concentrated solutions of hydrogen peroxide and manganese sulfate were prepared in 100 mM phosphate buffer (pH 7.5); 10 pl of each was added to 0.3 ml of enzyme solution in an EPR tube with no attempt to keep the enzyme solution anaerobic. Dithionite was added as a small crystal, and then the solution was frozen rapidly after the crystal dissolved. Enzyme carbohydrate analyses. Carbohydrate contents of peroxidase-M2 and ligninase-I (dissolved in 50 mM Na-tartrate, pH 5.0) were determined by treating known amounts of the pure enzymes with phenolsulfuric acid reagent (8). Glucose dissolved in 50 mM Na-tartrate (pH 5.0) was used as a standard. Amino acid analyses. Proteins were hydrolyzed by heating known amounts of purified enzyme in 6 N HCl for 22 h at 1lO’C under nitrogen. For a cysteic acid determination, dimethylsulfoxide was added to the acid prior to hydrolysis (41). After hydrolysis, norleucine was added as an internal standard. The sample was dried to remove HCl and then diluted to a concentration of 0.2 mg/ml protein in buffer at pH 2.2. Fifty microliters of the diluted sample was applied to a Dionex DC-5A column on a Dionex D-330 amino acid analyzer. The initial column temperature was 45”C, and the first eluant was a pH 3.25 Na-citrate buffer. The temperature was then raised to ‘73°C while continuing the same buffer. A final eluant was pH 7.9, 1.1 N Na-citrate. The total elution time was about 1 h. Amino acid peaks were derivatized with ninhydrin reagent and quantified by measuring peak absorbances at 440 nm. whole-cell cxridations of 2,&dimethq&enol A small piece of fungal mycelium was obtained from a stationary, ligninolytic (gday-old) culture and washed with distilled water. It was then suspended in a small

AND

CRAWFORD

volume of 100 mM Na-tartrate (pH 5.0) containing 0.1 mM manganese sulfate, 0.05 mM hydrogen peroxide, and 1.0 mM 2,6-dimethoxyphenol. After 4 h at 37°C the mycelium was washed several times with 50 mM Na-tartrate (pH 5.0) containing 2% glutaraldehyde. After sitting overnight, the mycelium was examined for the presence of crystalline deposition products by optical microscopy using a Zeiss Model 18 phase-contrast scope. Electrophoresis. Purity and molecular weights of enzymes were examined using SDS’-polyacrylamide gel electrophoresie techniques. A discontinuous electrolyte system was used (25), employing a 3% stacking gel and a 10% running gel. Protein bands were stained with Coomassie brilliant blue R-250. For molecular weight determinations, gels were calibrated with a mixture of proteins of known molecular weight (lowand high-molecular-weight calibration kits; Pharmacia, Uppsala, Sweden). Protein determinations Protein concentrations were determined by the micro-Coomassie blue procedure (4), using the dye reagent supplied by Bio-Rad (Richmond, Calif.). Chemicals The following compounds were purchased from the Aldrich Chemical Company (Milwaukee, Wise.); catechol, guaiacol, 2,6-dimethoxyphenol, 3,5-di-tert-butyl-4-methoxyphenol, vanillin, syringaldehyde, vanillic acid, syringic acid, acetovanillone, acetosyringone, isoeugenol, ferulic acid, sinapic acid, 4,4’-dihydroxybiphenyl, vanillinazine, syringaldazine, glyoxal-bis(2-hydroxyanil), quercetin, rutin, chlorogenic acid, and coniferyl alcohol. Guaiacol glyceryl ether was purchased from K and K Laboratories, Inc. (Plainview, N. Y.). Vanillyl alcohol, syringyl alcohol, apocynol, and 1-[4-hydroxy-3,5-dimethoxylethanol were prepared by reduction of their carbonyl precursors (Aldrich Chemical Co.) with sodium borohydride in ethanol; product identities were confirmed by mass spectrometry (MS) and nuclear magnetic resonance (NMR) analyses (17,18). 3-m-ButylI-methoxyphenol was a gift from Dr. Luke K. T. Lam (Department of Laboratory Medicine and Pathology, University of Minnesota). Vanillylacetone and syringylacetone were prepared as follows: 0.1 M vanillin or syringaldehyde was dissolved in 50 ml of acetone. The solution was warmed to 60°C under nitrogen while stirring; 20 ml of 4 N aqueous KOH was added dropwise over 30 min, and the solution was refluxed (60°C) for 2 h. The solution was poured over ice and acidified to pH 2.5-3.0, which produced a precipitate of light yellow crystals which were collected and recrystallized from acetonezhexane (l:l), yielding about 90%.

* Abbreviations used: SDS, sodium dodecyl MS, mass spectrometry; DTT, dithiothreitol; dihydroxymaleic acid.

sulfate; DHM,

PEROXIDASE

CHARACTERIZATION

Vanillylacetone, ‘H-NMR (deuterochloroform): 2.30 (3H, s, CH3), 3.85 (3H, 8, OCH3), 6.51 (lH, d, vinyl), 7.41 (lH, d, vinyl), 6.82-7.08 (3H, aromatic); MS: m/z = 192 (M+, loo), 177 (97), 161 (8), 149 (18), 145 (58). Syringylacetone, ‘H-NMR: analogous to vanillylacetone except showing two methoxyls (6 H); MS: m/ z = 222 (M+, loo), 207 (47.6), 194 (24.1), 179 (3.4), 175 (23.8), 151 (8.4). Guaiacylglycerol-fi-guaiacyl ether [l-(l-hydroxy3-methoxyphenyl) -1,3- (2-methoxyphenoxy) -propanediol] was prepared by the procedure of Hosoya et al. (15). Disyringylmethane [l,l-&(3,5-dimethoxy-4-hydroxyphenyl)-methane] was synthesized as reported by Steelink (40). Identities of these two compounds were confirmed by MS and/or NMR analyses. 1,2,4-Trimethoxybenzene was prepared by methylation of 2,4-dimethoxyphenol (Aldrich Chemical Co.) with dimethylsulfate in 4 N KOH. MS: m/z = 168 (lOO), 153 (75), 139 (5.8), 135 (33), 110 (11). 4,4’-Dimethoxybiphenyl was obtained similarly by methylation of commercially available 4,4’dihydroxybiphenyl. MS: m/z = 214 (loo), 199 (55), 171 (14), 156 (35), 128 (2.7). Isoeugenol methyl ether was prepared by methylation of isoeugenol. MS: m/z = 178 (100). 163 (28), 147 (7.2), 135 (5.4), 117 (3.6), 107 (15.5). 1-(3,4-Dimethoxyphenyl)-2-(2,4-dichlorophenoxy)ethanol was obtained by reducing with borohydride its a-keto precursor [prepared by a reaction sequence analogous to that used for the synthesis of veratrylglycerol-6-guaiacyl ether (l)]. MS: m/z = 342 (M+, 5%),324 (lOO), 167 (76.5), 139 (24.2). Iso.!ntim and identification of reaction products. Substrates were incubated with particular enzymes under each enzyme’s usual assay conditions (cf. above), except that reaction volumes were increased to 50 ml. Reactions were run until no further changes were observed in the substrates’ uv-visible spectra. Products were recovered by extraction of acidified reaction solutions with chloroform (3 X 25 ml). They were identified by gas chromatography/MS techniques, comparing retention times and mass spectra of reaction products with those of authentic standards, or by MS fragmentation pattern interpretation alone where authentic standards were unavailable. Some products also could be tentatively identified by their very characteristic uv absorption spectra. In a number of instances, reaction products from particular substrates were not identified. In these cases it was simply noted that substrate transformation did or did not occur. RESULTS

Produh%ion of lipinase-I and per&eM2. Growth of P. chrysosopcwium as a mycelial mat on the inner surface of a slowly rotated carboy allowed efficient, continuous

753

production of extracellular enzymes. After the mycelium had been established, but prior to harvesting of the culture fluid, veratryl alcohol (1 mM) was added to the growth medium. This resulted in a threeto fourfold increase in activity of ligninaseI (10) and a twofold increase in activity of peroxidase-M2. Without addition of veratry1 alcohol, ligninase activity disappeared after a few replacements of the growth medium with fresh medium. PeroxidaseM2 activity, however, was produced each time the growth medium was replaced, even without addition of an inducer. When veratryl alcohol was included in replacement medium, we could harvest about 16 liters of medium containing both enzymes over a period of 1 month before fresh mycelium had to be grown. PurQkation of ligninase-I and peroxiduse-M2. Enzymes were purified simultaneously as outlined in Table I. LigninaseI was purified 2.5- to 3.5-fold and peroxidase-M2, 8.1- to Is-fold as compared to their specific activities in culture filtrates. Activities of the purified enzymes varied from 6.4-12.5 U/mg for the ligninase and 65-134 U/mg for the peroxidase, depending on culture age. The primary purification step involved chromatofocusing column chromatography. Figure 1 shows elution profiles observed for the two enzymes from a PBE-94 column. Numerous protein peaks were eluted. Two of these expressed ligninase activity (veratryl alcohol oxidation to veratryl aldehyde, in the presence of hydrogen peroxide), while three expressed peroxidase activity (oxidation of vanillylacetone in the presence of hydrogen peroxide and Mr?). Most of the enzymatic activity, however, was concentrated in two peaks (fractions 80-90, peroxidase-M2; fractions llO-123,ligninase-I). These peaks were pooled separately, and concentrated and desalted by ultrafiltration in 50 mM Na-tartrate, pH 5.0. Both peaks were subjected to SDS-polyacrylamide gel electrophoresis, and each gave only one protein band (even in heavily loaded gels; Fig. 2). Characterization of ligninase-I and peroxidase-M2. SDS-polyacrylamide gel electrophoresis of the two enzymes against

754

PASZCZYfiSKI,

HUYNH,

AND

TABLE PURIFICATION

1 2 3

Volume (ml) 10,000 75 5.5 (L) 4.5 (P)

a (1) Culture chromatography by ultrafiltration.

I

OF LIGNINASE-I AND PEROXIDASE-MB OF Phunerochaete chqsosporium

Units/ml Step”

CRAWFORD

Total

units

FROM THE GROWTH BKM-F-1767 Units/mg protein

P

L

P

L

0.006

0.016 2.0 9.1

61

163

3.0

8

32 5

151 41

4.2 6.4

20 65

filtrate; (2) ultrafiltration through an Amicon on PBE-94 (Pharmacia) polybuffer exchange L = ligninase-I; P = peroxidase-M2.

standard proteins of known molecular weight indicated molecular weights of 45,000-47,000 Da (peroxidase-M2) and 42,000-43,000 Da (ligninase-I) (Fig. 3). Acid hydrolysis of the two enzymes followed by determinations of reducing sugars released showed that both proteins contained appreciable amounts of carbohydrate (peroxidase-M2, 17% by weight; ligninase-I, 21%). In its purified state peroxidase-M2

0

20

40

60

60 Fraction

PM-10 resin

Purification

% Yield

L

0.42 0.9

MEDIUM

P

L

P

L

P

_

_

-

_

52 8

92 25

1.4 2.1

2.5 8.1

(10,000 M, cutoff) membrane; (3) column followed by desalting and concentration

was unstable at 5”C, but could be stored for at least 6 months in tartrate buffer, pH 4.5, if frozen. Ligninase-I also was unstable. In frozen crude concentrates of the growth medium, ligninase activity disappeared over a period of about 1 month. Purified ligninase-I was considerably more stable. Peroxidase-MB is a heme enzyme. The native enzyme shows a strong heme absorption at 406 nm, which shifts to 427 nm

100 Number

120

I40

160

FIG. 1. Chromatography of Phawochaete chrysospurium extracellular proteins on a PBE-94 ion exchange column. Bed height, 35 cm; flow rate, 20 ml/h; fraction volumes, 3.4 ml. Elution conditions: starting buffer, 0.025 M imidazole-HCl; elution buffer, Polybuffer 74-HCI (Pharmacia, diluted 1:8) at 280 nm; ---, absorbance at 409 nm; at pH 4 and containing up to 0.2 M NaCl.-, Absorbance -m-e, NaCl gradient; . * a, pH gradient; 0, peroxidase-M2 activity; 0, ligninase activity.

PEROXIDASE

A

B

755

CHARACTERIZATION

C

tains protoporphyrin IX at about 0.7 heme per enzyme molecule. Tien and Kirk (44) reported that their ligninase contained about 0.8 heme per protein molecule. Examination of Peroxidase-MB by EPR spectroscopic techniques (Figs. 5A-D) suggested that its active center contains a high-spin ferriheme complex. The native enzyme shows a g, value near 6 and a gll near 2, with E/A a little larger than 0, as expected for high-spin Fe3+ porphyrins (Fig. 5A) (3,42). The signal at 4.3 is due to contaminating Fe3+ not coordinated by heme. Signals are sharp with no signs of heterogeneity. When the enzyme is incubated in the presence of hydrogen peroxide, the signal at g = 6 nearly disappears, leaving the g = 4.3 signal (Fig. 5B). Hydrogen peroxide probably has oxidized the heme. After adding Mn2+, about one-third of the g = 6 signal returns (Fig. 5C), showing that the iron center has cycled back in part to Fe3+. In the presence of dithionite a large increase in EPR active Mn2+ is observed (Fig. 5D). EPR spectra of ligninase-I were not affected by Mn2+; these show g values

FIG. 2. Electrophoretogram of purified peroxidaseM2 and ligninase-I. (A) Ligninase-I (30 pg); (B) peroxidase-M2 (20 rg); (C) mixture of the two proteins (l:l, 50 fig).

after reduction and carbon monoxide saturation (Fig. 4A). Saturation of the native enzyme by CO without reduction resulted in no spectral shifts. The peroxidase/hydrogen peroxide complex absorbs at 421 nm, but this shifts back to 406 nm on addition of Mn2+ (Fig. 4B). When the peroxidase is reduced with dithionite, it absorbs at 440 nm, with a weaker band at about 560 nm (Fig. 4C). Spectral characteristics of ligninase-I (also a heme protein) were identical to those reported for this enzyme by other workers (12, 44), and are not reproduced here. The ratio of absorbance at 406 nm over that at 280 nm for the purified peroxidase was 3.13. Ligninase-I showed a ratio of 3.47. Estimation of the heme content of peroxidase-M2 by formation of the pyridine hemochromogen (30) indicated that the active center of the enzyme con-

105

!=

~hmpboryiosr

b (94.000)

FIG. 3. Molecular weight determinations for ligninase-I and peroxidase-M2. Enzymes and marker proteins were dissolved in 10 mM Tris-HCI, pH 8.0, containing 1 mM EDTA, 2.5% SDS, and 5% @-mercaptoethanol. After 5 min boiling and addition of 10% glycerol, samples were applied to the gel. Electrophoresis conditions: current, 1 mA for the first 0.5 h, then 3 mA for the next 4-6 h; marker, bromophenol blue. After electrophoresis gels were incubated for 2 h in 25% isopropanol/lO% acetic acid and then stained overnight in 0.1% Coomassie blue in 7% acetic acid/ 50% ethanol. Gels were destained electrophoretically at 25 V for 1 h in methanol/acetic acid/water (l/15/ 6, v/v/v).

‘756

PASZCZYrjSKI,

0.6 -

427 fi

HUYNH,

-

NATIVE

----

FERROUS-CO-COMPLEX

ENZYME

-

I 400

500 WAVELENGTH

600 (nm )

-

FERRIC-CO-COMPLEX- __.. -. -..

OL

I

700

400

500 WAVELENGTH

B

-

06-

CRAWFORD

0.6

01 A

AND

REDUCED

600 lnml

700

ENZYME

440

---------

------.-.-

01 400

C

500 WAVELENGTH

60’3 (nm)

700

FIG. 4. Absorption spectra of purified peroxidase-M2. Protein (0.2 mg/ml) was dissolved in 50 mM Na-succinate, pH 5, and spectra were run at room temperature. The reference cuvette contained only buffer. Procedures for forming the various protein complexes are described in Refs. (12,31).

near 6 and 2, with E/A close to 0 as expected for high-spin Fe3+ (Figs. 6A and B). The signal at 4.3 in the ligninase spectrum probably is adventitious iron. As with the peroxidase, EPR signals of the ligninase are sharp with no heterogeneity. In the presence of hydrogen peroxide, the ligninase g = 6 signal disappears leaving the 4.3 signal unaffected, probably the heme has been oxidized. Substrate spec@icitg of peroxiduse-MZ and ligninaae-I. As reported previously

(34), peroxidase-M2 shows peroxidatic activity against a wide variety of phenols and phenolic lignin model compounds. This activity is dependent on hydrogen peroxide and Mn2+. The peroxidase also shows oxidase activity against NADPH, GSH, dithiothreitol (DTT), and dihydroxymaleic acid (Table II). Oxidase activity does not require hydrogen peroxide, but produces hydrogen peroxide (34). This activity is Mn2+ dependent. Ligninase-I shows numerous activities (12, 44), including the

PEROXIDASE

757

CHARACTERIZATION 66.12

6.11

-G&F&r A

6.1

4.27

2.06

s

s 6.12

FIG. 6. EPR spectra of ligninase-I. Spectra were run in 50 mM Na-tartrate, pH 5. (A) Native enzyme, run at 5.5K, 0.2 mW, and 16,000 gain. (B) After addition of hydrogen peroxide, run at 5.5 K, 0.2 mW, and 32,000 gain. Scale for magnetic field as in Fig. 5A.

completely inactivated by boiling for 5 min, but about 20% of the original activity could be recovered after storing the heat-inactivated protein at 0°C for 1 h. Boiling for D

5. EPR spectra of peroxidase-M2. (A) Native enzyme in 50 mM Na-tartrate, pH 5; 0.35 mg/ml protein. (B) After oxidation with hydrogen peroxide. (C) After addition of Mn’+. (D) after dithionite reduction. Spectra were run at 6.2 K, 0.2 mW, and a gain of 20,000 (A) or 32,000 (B-D).

TABLE

II

FIG.

ability to oxidize nonphenolic lignin model compounds and the lignin polymer. Comparisons of the substrate specificities of peroxidase-M2, ligninase-I, and horseradish peroxidase are shown in Tables III and IV. Peroxidase-M2 will oxidize syringyl alcohol to syringyl aldehyde, while neither ligninase-I nor horseradish peroxidase will catalyze this reaction at a significant rate (Table III). Inhibitors of peroxidase-M2. PeroxidaseM2 is inhibited by metal ions (0.1 InM) such as Fe3+, Fe2+, Co2+, and Cu2+; these metals inhibit the peroxidase between 34 and 77% as compared to activities in the absence of inhibitors, using our standard assay of oxidation of vanillylacetone (Table V). Salicylic acid (0.1 mM) and ethanol do not inhibit the peroxidase. Ascorbic acid (0.1 mM) inhibits the enzyme 100% ,while superoxide dismutase (100 U) inhibits vanillylacetone oxidation by about 80%. The enzyme is

OXIDATIVE

ACTIVITIES

OF PEROXIDASE-MB Activity

Substrate NADPH GSH DTT DHM Syringyl

alcohol

Oxidase” 9.4 36.8 595.9 1444.6 -

Peroxidaticb 5.8'

a Activity was measured, in the absence of hydrogen peroxide, as oxygen consumption (pmol/min/mg protein) using an oxygen electrode. Reaction mixtures contained 1.6 ml of water, 0.4 ml of 0.5 M Na-tartrate (pH 5.0), 0.2 ml of 1 mM manganese sulfate, 0.1 ml of 10 mre substrate, and 25 ~1 of enzyme solution (0.175 mg protein/ml). Assays were performed at 30°C. b Activity was measured in the presence of hydrogen peroxide and Mns+, spectrophotometrically by monitoring conversion of the alcohol to its corresponding aldehyde (pmol/min/mg protein). Increase in absorbance at 307 nm was monitored using E = 12,500 for the aldehyde. Standard peroxidase-M2 assay conditions were employed (see Experimental Procedures). ’ Ligninase-I and horseradish peroxidase (HRP) oxidized syringyl alcohol to syringyl aldehyde, but at much slower rates (Ligninase-I, 0.1 pmol/min/mg, and HRP, 0.07 rmol/min/mg).

(catechol) (guaiacol)

OXIDIZED

.-

-

- -

-

.- . -

-

BY LIGNINASE-I,

2,6-Dimethoxyphenol 3-tert.-Butyl-4-methoxyphenol 3,5-Di-ted.-butyl-4-methoxyphenol 4-Hydroxy-3-methoxybenzaldehyde (vanillin) 4-Hydroxy-3,5-dimethoxybenzaldehyde (syringyl aldehyde) 4-Hydroxy-3-methoxybenzyl alcohol (vanillyl alcohol) 4-Hydroxy-3,5-dimethoxybenzyl alcohol (syringyl alcohol) 4-Hydroxy-3-methoxybenzoic acid (vanillic acid) 4-Hydroxy-3,5-dimethoxybenzoic acid (syringic acid) 1-[4-Hydroxy-3-methoxyphenyl] ethanone (acetovanillone) 1-[4-Hydroxy-3,5-dimethoxyphenyl] ethanone (acetosyringone) 1-[4-Hydroxy-3-methoxyphenyl] ethanol (apocynol) 1-[4-Hydroxy-3,5-dimethoxyphenyl ethanol 4-Hydroxy-3-methoxycinnamic alcohol (coniferyl alcohol) 2-Methoxy-4-propenylphenol (isoeugenol) 4-[4-Hydroxy-3-methoxyphenol]-3-buten-2-one (vanillylacetone) 4-[4-Hydroxy-3,5-dimethoxyphenyl]-3-buten-2-one (syringylacetone) 3-(4-Hydroxy-3-methoxyphenyl)-2-propenoic acid (ferulic acid) 3-(4-Hydroxy-3,5-dimethoxyphenyl)-2-propenoic acid (sinapic acid)

1,2-Dihydroxybenzene 1-Methoxyphenol

Substrate

SUBSTRATES

III

.- .- - -- -

I - - _

.-.

,

-t

+++ 310(d)

- .- . -

nd

nd

++

287/310(d)

++

+++

+++

nd nd nd nd nd

336(d)

nd ++ nd

nd -

+++t

++ +/+ +++ ++ +++

270-275(a)

285(a)/405(a)

288(t)/405(a)

nd + nd nd nd

+++

nd +/-

+++

+++ +++

HRP

336(d)

298(d) 278(d) 278/298 2f.Wa) 260(d)/298(d)

305(a) 2Wd) 260/285(a)

nd

+/+/+ + ++

+++ +/+++ + +++ t

3Wa)

258(a)

+++ ++ ++

470(a)

++t + ++ +

+ +++

265(d) 2856) 285(d) 298 306(d) 278(d) 268/280(d) 250/2ao =X4 275/305(d)

LigninaseI

(HRP)

PeroxidaseM2

398(a) 465(a)

Product

PEROXIDASE

275(d) 270(i)

Substrate

AND HORSERADISH

xmax”

PEROXIDASE-MZ,

TABLE

d co

PEROXIDASE

;+LWWWW: +++cFCEC+ +

+‘+$+ +Z+f$ ++++

759

CHARACTERIZATION

20 min inactivated the peroxidase which it could not be reactivated.

+

Hydrogen

peroxide

after

and Mn dependence

of peroxidase-M2. The influence of Mn2+ concentration on peroxidase activity in the presence of 0.1 mM hydrogen peroxide, using four substrates, is shown in Fig. 7. Mn2+ is maximally stimulatory at different concentrations depending on the substrate being oxidized. Maximal stimulation is seen at 1.0 mM when o-dianisidine, syringaldazine, and sinapic acid are substrates. When vanillylacetone is used, maximal stimulation by Mn2+ is seen at 0.05 mM. The influence of hydrogen peroxide concentration, in the presence of a standard Mn2+ concentration, on activity of the pure peroxidase against four of its substrates is shown in Fig. 8. Activity is maximal at about 0.1 mM hydrogen peroxide, and concentrations above 0.2 mM are inhibitory. When peroxidase-M2 was incubated with MnS04 and hydrogen peroxide, and then protein was removed over a Sephadex G25 column, the protein-free, yellow-brown solution was able to oxidize syringaldazine, o-dianisidine, and other phenolic compounds. This activity was unstable and disappeared in about 30 min. Hydrogen peroxide and Mn2+ in the absence of peroxidase-M2 (no enzymatic activation of Mn2+) were unable to carry out these oxidations. We immobilized the peroxidase on CNBr-activated Sepharose 4B. When Mn2+ and H202 were passed through a column of immobilized enzyme, a similar solution of protein-free, activated Mn was produced. The oxidative activities of these manganese solutions were unaffected by treatment with catalase to decompose residual H20z. This confirms that peroxidase-M2 is a Mnperoxidase, oxidizing Mn2+ (in the presence of HzOz) to a higher oxidation state. Cellular location ity. One substrate

of peroxiduse-M2

activ-

of the peroxidase, 2,6dimethoxyphenol, was readily oxidized to a purple-colored, crystalline product (2,5,2’,5’-tetramethoxy-p-dibenzoquinone) (11). When 2,6-dimethoxyphenol was oxidized by intact fungal mycelium, purple crystals were deposited on the mycelial surface (Fig. 9). Pure tetramethoxy-p-dibenzoquinone crystals prepared separately

760

PASZCZYP;ISKI,

HUYNH,

TABLE NONPHENOLIC

Substrate

1-(3,4-Dimethoxyphenyl)-lpropene (isoeugenol ether)

methyl

1-(3,4-Dimethoxyphenyl)-2-(2,4dichlorophenoxyl)-ethanol

SUBSTRATES OF LIGNINASE-I

285

328

Pink

2601288

398

p,p’-Dibenzoquinone

258 300 (sh.)

275 305 (sh.)

276,280

310

One product

318

Product phase

272

and then mixed with mycelium did not attach. We could not find peroxidase-M2 activity in cell-free extracts of washed mycelium. The enzyme, however, was readily extractable from intact mycelium by washing with 0.5 M NaCl or 0.1% Tween 80. Ligninase-I was free in the culture medium; additional activity was not extractable from mycelia with NaCl or Tween 80 nor was it found in mycelial cell-free extracts.

INHIBITION

Potential

None 0.1 mM Fen+ 0.1 mM Fez+ 0.1 mM Co2+ 0.1 mM Cu2+ 0.1 mM Salicylic acid 0.1 mM Ascorbic acid Superoxide dismutase Ethanol (5%)

Remarks product

(quinone

?)

is veratraldehyde

remains

in aqueous

Amino acid composition of peroxidaseMz. Peroxidase-M2 is rich in acidic amino acids, particularly aspartic acid and glutamic acid, these two amino acids comprise about 24% of the total amino acids present.

V

OF PEROXIDASE-M2 inhibitor

IV

Product

3-(o-Methoxyphenoxyl)-1,2propane-diol (guaiacyl glycerol ether)

TABLE

CRAWFORD

Substrate

1,2,4-Trimethoxybenzene 4,4’-Dimethoxybiphenyl

AND

ACTIVITY % Inhibition

(100 U)

0 34 77 6’7 62 0 100 80 0

Note. Peroxidase-MX activity was determined by monitoring oxidation of vanillylacetone in thre presence of hydrogen peroxide and Mnz+ as described under Experimental Procedures. Inhibitors were added at the concentrations indicated.

mM MnSO4

FIG. ‘7. Influence of Mn2+ concentration of peroxidase-M2 activity. Reaction mixtures contained in 1 ml: 0.2 ml of 0.5 M Na-tartrate (pH 5), 0.1 ml of 1 mM substrate, 0.1 ml of 1 mM hydrogen peroxide, 0.1 ml of an appropriate concentration of manganese sulfate, 25 pl(5 pg) of enzyme, and 0.4 ml of water. 0, Sinapic 0, o-dianisidine; A, syracid, -e-m, vanillylacetone; ingaldazine.

PEROXIDASE

.g

DISCUSSION

70

g

60

F ;

50

761

CHARACTERIZATION

d 5 40 :z a g 30 ii u-J 20

mM

Ii202

FIG. 8. Influence of hydrogen peroxide concentration on peroxide-M2 activity. Reaction mixtures contained 0.1 mM MnzC; other conditions as in Fig. ‘7.

Alanine, glycine, and proline comprise about 29% of the total (Table VI). These five amino acids comprise 53% of the protein component of the enzyme.

FIG. 9. Mycelium of Pkonerochaete Magnification: 1000X. Arrows indicate the crystals of tetramethoxy-pdibenzoquinone

Ligninase-I and peroxidase-M2 were purified from the growth medium of P. chrgsospwium by a combination of ultrafiltration and chromatofocusing chromatography. These methods yielded electrophoretically pure preparations of each enzyme, purified simultaneously and in only three steps. Peroxidase-M2 shows properties of an oxidase and a peroxidase. As an oxidase it behaves similarly to horseradish peroxidase (34), oxidizing substrates such as NADPH (to presumedly NADP+), GSH (to GSSG), DTT (to its disulfide derivative), and dihydroxymaleic acid (DHM; to its diketo derivative). These transformations require Mn2+. Electrons and protons removed from the oxidized substrate are accepted by oxygen, yielding hydrogen peroxide (34) (Fig. 10). An identical type of reaction has been described for the Mn2+dependent oxidations of DHM and DTT by

chrysospon'um after points where crystals were formed

incubation with 2,6-dimethoxyphenol. are observed attached to the mycelium; on the mycelial surface.

762

PASZCZYfiSKI,

HUYNH,

DTT GSH NADH DHM

FIG. 10. Reactions of peroxidase-M2 that generate hydrogen peroxide and involvement of this enzyme in oxidation of aromatic substrates (AH). Manganese ion serves as an electron carrier in peroxidatic reactions.

horseradish peroxidase (6, 30). Yamazaki and Piette (46) found that Mn II, though not required, promoted the oxidase reactions of horseradish peroxidase. They suggested that Mn II promotes HzOz formation by slowing the dismutation of HOa-radicals formed during the catalytic cycle. The physiological role of this oxidase activity is unclear in the case of horseradish peroxidase; however, in the case of peroxidaseM2, the production of hydrogen peroxide has important implications regarding the mechanisms of fungal lignin degradation (26, 34). The hydrogen peroxide produced may become available to ligninolytic enzymes (e.g., heme ligninases of the ligninase-I type, which are hydrogen peroxide dependent (12, 44)), or it may be used by the peroxidase-M2 itself when the enzyme acts as a peroxidase. It is interesting to speculate another potential role for peroxidase-M2 in fungal metabolism: if the enzyme is able to oxidize sulfhydryl groups in other proteins, it may function in regulating activities of extracellular enzymes by modifying their tertiary structures. Olsen and Davis (30) have postulated a similar regulatory role for horseradish peroxidase in the conversion of reduced galactose oxidase (sulfhydryl form, inactive) to oxidized galactose oxidase (disulfide form, active). Peroxidase-M2 also acts as a classic peroxidase. Hydrogen peroxide-oxidized peroxidase-M2 shows spectral characteristics similar to those of horseradish peroxidase compound II (9,28). This oxidized enzyme

AND

CRAWFORD

is able to extract electrons from Mn2+ (Fig. 10). The oxidized Mn apparently then may remove electrons from a phenolic substrate, oxidizing that substrate. These speculations are supported by our ultraviolet-visible and EPR spectroscopic experiments, and the observation that protein-free Mn ion solutions express peroxidatic activity after preincubation with peroxidase-M2. The lack of inhibition of peroxidatic activity by ethanol or salicylic acid implies that hydroxyl radicals ( * OH) are not involved in the peroxidative mechanism. Inhibition of peroxidase-M2 by superoxide dismutase indicates that superoxide free radical anion may be an electron acceptor during the peroxidatic process. The visible spectra of the native, oxidized, and reduced Mn’+-dependent peroxidase-M2 are very similar to analagous spectra of ligninase-I (2, 12, 44). These spectra along with EPR spectra and spectra of pyridine-heme complexes from both

TABLE AMINO

ACID

COMPOSITION

VI OF PEROXIDASE-M2 AA

composition Amino Cyst&

acid

(number/mol enzyme)

acid

2 49 28 24 37 32 33 42 19 5 14 24 17 8 14 10 40

ASP Thr Ser Glu Pro GlY Ala Vsl Met Ileo Leu Phe His LYS Arg Ammonia Weight of protohematin Weight of sugar

weight 63) 302 5,639 2,830 2,090 4,776 3,107 1,884 2,986 1,385 656 1,= 2,716 2,502 1,097 1,795 1,562 640 616

IX

7.800 46,458

Total Note. The amino under Experimental

Residue

acid analysis Procedures.

was carried

out aa described

PEROXIDASE

CHARACTERIZATION

enzymes suggest that all contain protohematin IX (this work and (2, 12, 40)). We found that ligninase-I from P. chrysospwium BKM-F-1767 contains about 21% by weight carbohydrate. Tien and Kirk (44) reported a carbohydrate content of 13% for what appears to be otherwise an almost identical enzyme. Their enzyme was isolated from the same strain of P. chrysosporium; thus the difference in carbohydrate contents of the two preparations may simply represent inherent variability. In any event, the amount of carbohydrate in a particular ligninase appears not to be very important regarding enzyme activity. Peroxidase-M2 also is a glycoprotein, containing about 1’7% carbohydrate. The peroxidase is rich in acidic amino acids, especially aspartic acid and glutamic acid (Table VI). In this regard it is similar to chloroperoxidase, horseradish peroxidase, Japanese radish peroxidase, and myeloperoxidase (27,39). The peroxidase is similar to plant peroxidases that have low lysine and cysteine contents (9). There are no precise methods for measuring protein contents of these acidic glycoproteins, which contain few aromatic residues. Thus, our estimations of heme and carbohydrate contents (which employed the micro-Coomassie blue protein determination) should be considered only as rough estimations. The role of Mn2+ in the catalytic action of peroxidase-M2 is of particular interest. EPR spectra show that divalent manganese is present only in small amounts in reaction mixtures (Fig. 5C), but after reduction anaerobically with dithionite, the g = 2 signal in the EPR spectrum shows a very large increase (Fig. 5D; Mn2+ regenerated). These observations indicate that most of the manganese is in the Mn3+ or higher oxidation state (EPR inactive). Mn2+ can reduce the peroxidase active center, but only after the enzyme has been oxidized previously by hydrogen peroxide (Figs. 4B and C). Our experiments indicate that peroxidase-M2 mediates electron transfer from Mn2+ (donor) to hydrogen peroxide (acceptor), and that a reaction product is Mn3+ (or higher oxidation state). This reactive Mn species will then oxidize various aro-

763

matic substrates, as shown in Fig. 10. Simple mixtures of hydrogen peroxide and Mn2+ or manganese dioxide do not oxidize our aromatic substrates-activation of Mn2+ by the enzyme is required. As shown in Figs. 6A and B, ligninase-I gave EPR spectra very similar to those of the peroxidase. Mn2+ had no discernable effect on the ligninase spectra. It is quite possible that manganese ions, which are not involved in the activity of ligninase-I, act in solution (rather than bound to the enzyme) as electron carriers in reactions catalyzed by peroxidase-M2. It is clear that activity of peroxidase-M2 is almost totally dependent on the activity of manganese ions acting as electron carriers, whether they be enzyme bound or not. Both peroxidase-M2 and ligninase-I oxidize a wide variety of substrates (this work and (12, 16, 20, 34, 44)). The peroxidase, however, appears to attack syringyl (3,5dimethoxy-4-hydroxyphenyl)-substituted compounds much more readily than guaiacyl(4-hydroxy-3-methoxyphenyl)-substituted compounds (Table III). Guaiacyl units are found primarily in lignins from conifers and grasses, while syringyl units predominate in lignins of hardwoods (7937).

It is interesting to note that phenol oxidizing enzymes are produced widely by ligninolytic fungi (37), and that white-rot fungi (including P. chrysosptium) often preferentially degrade lignin syringylpropane units as compared to their guaiacyl analogs (14, 22, 29). This preferential removal of lignin’s syringyl units by whiterotters recently was confirmed by Tai et al. (43) who used 13C-NMR spectroscopy and chemical fractionation techniques to examine white-rotted lignins. Indeed, birch lignin (largely syringyl units) long has been known to be more susceptible to white-rot decay than spruce lignin (largely guaiacyl units) (21). A reason may be that the 4hydroxy-3,5-dimethoxyphenylpropane moieties that comprise the syringyl structures are less susceptible to ortho coupling (condensation) reactions following their oxidation by phenol oxidases and/or peroxidases than are guaiacyl structures

764

PASZCZYtiSKI,

HUYNH,

which have an unoccupied 5 position on the ring where coupling may occur. Also, in the syringyl units the unpaired electron of a phenoxy radical produced by action of a peroxidase may be transmitted through the u electrons of the aromatic ring to the puru position where a side-chain displacement site may be created, leading to lignin depolymerization. Finally, lignin’s crossconjugation allows radicals to move from unit to unit within the lignin molecule, permitting long-range degradative effects. Thus, the oxidation of syringyl compounds catalyzed by peroxidase-M2 (Table III) may indicate an important role of this enzyme in mediating lignin degradation, especially of lignin’s syringyl units; however, a role for peroxidase-M2 in lignin degradation has not been proven. Future efforts will concentrate on this important question. Both ligninase-I and peroxidase-M2 show phenol oxidase activity; however, the ligninase shows more selectivity than the peroxidase. Both enzymes oxidize coniferyl alcohol, sinapic acid, and certain other substrates (Table III), but reaction rates often are very different for the two enzymes. The peroxidase shows no affinity for nonphenolic compounds, which are readily attacked by the ligninase (Table IV). Some substrates readily oxidized by the peroxidase (e.g., vanillylacetone) are not attacked by the ligninase. Thus, the two enzymes probably perform different roles in the lignin decay process. Our microscopy and enzyme assay experiments suggest that peroxidase-M2 activity is largely extracellular, but that most of the activity is associated with the outer mycelial surface, rather than being free in the growth medium. Unlike the purified peroxidase, the mycelial associated enzyme shows less dependence on Mn2+ and does not require exogenous hydrogen peroxide. These observations support earlier suggestions that enzymes involved in lignocellulose degradation may function in close association with the fungal mycelial surface or may even be complexed with extracellular mucilage sheaths (13, 32, 33). In addition, such a natural form of enzyme immobilization may limit competition of enzymes for low-molecular-weight sub-

AND

CRAWFORD

strates (e.g., NADPH, GSH, hydrogen peroxide, and Mn’+), prevent loss of enzymes by diffusion away from the mycelium, and allow some degree of structural organization of the ligninolytic system (45). ACKNOWLEDGMENTS We thank Pat Olson for invaluable technical assistance and Dr. Ed Brown for providing a rotating fermenter. EPR spectra were run by Dr. John Lipscomb. Discussions with Frank Rusnak and Dr. Eckard Miinck were helpful.

REFERENCES 1. ADLER,

E., AND ERIKSSON,

E. (1955)

Actu Chem.

Scand 9,341~348. 2. ANDERSSON, L. A., RENGANATHAN, V., CHIU, A. A., LOEHR, T. M., AND GOLD, M. H. (1985) J. Bid Chem 260,6080-608’7. 3. BENNET, J. E., INGRAM, D. J. E., GEORGE, P., AND GRIFFITH, J. S. (1955) Nature (London) 176,394. 4. BRADFORD, M. M. (1976) Anal Chem 72.248-254. 5. BUSWELL, J. A., MOLLET, B., AND ODIER, E. (1984)

FEMS Microbid

Lett. 25,295-299.

6. CHANCE, B. (1952) J. Biol. Chem. 19’7.577-589. ‘7. CRAWFORD, R. L. (1981) Lignin Biodegradation and Transformation, Wiley-Interscience, New York. 8. DUBOIS, M., GILLES, K. A., HAMILTON, J. K., REBERS, P. A., AND SMITH, F. (1956) Anal Chem. 28,350-356. 9. DUNFORD, H. B., AND STILLMAN, J. S. (1976) Coord Chem. Rev. 19.187-251. 10. FAISON, B. D., AND KIRK, T. K. (1983) Appl Environ Microbial. 46,1140-1145. 11. GIERER, J., AND OPARA, A. E. (1973) Acta Chem Stand. 27,2909-2922. 12. GOLD, M. H., KUWAHARA, M., CHIU, A. A., AND GLENN, J. K. (1984) Arch. Biochem Biophys. 234,353-362. 13. HIGHLEY, T. L., PALMER, J. G., AND MURMANIS, L. L. (1983) Holqfbrschung 37,179-184. 14. HIROI, T., AND TAMAI, A. (1983) in Recent Advances in Lignin Biodegradation Research (Higushi, T., Chang, H-M., and Kirk, T. K., eds.), pp. 3443, Uni Publ. Co., Tokyo. 15. HOSOYA, S., KAMAZAWA, K., KANEKO, H., AND NAKENO, J. (1980) Mokuzai Gakkaishi 26,118121. 16. HUYNH, V. B., AND CRAWFORD, R. L. (1985) FEMS MicmbioL L.&t. 28,119-X%. 17. HUYNH, V. B., ISHIZU, A., AND NAKANO, J. (1982)

Mokuzai Gakkaishi 28,129-136. 18. HUYNH,

V. B., ISHIZU,

Mokuazai

A., AND NAKANO,

Gakkaishi 28, X4-171.

J. (1982)

PEROXIDASE

765

CHARACTERIZATION

19. KERSTEN, P. J., TIEN, M., KALYANARAMAN, B., AND KIRK, T. K. (1985) J. BioL Chm 260,2609-2612. 20. KIRK, T. K., MOZUCH, D., AND TIEN, M. (1985) Biochxm. J. 226,455-460. 21. KIRK, T. K., CHANG, H-M., AND LORENZE, L. F. (1975) Wood Sci. TechnoL 9,81-86. 22. KIRK, T. K., AND LUNDQUIST, K. (1972) Svensk Papperstidn 73.294-306. 23. KIRK, T. K., SCHULTZ, E., CONNORS, W. J., LORENZ, L. F., AND ZEIKUS, J. G. (1978) Arch Microbial 117,277-285. 24. KUWAHARA, M., GLENN, J. K., MORGAN, M., AND GOLD, M. H. (1984) FEBS Lett. 169,247-250. 25. LAEMMLI, U. K. (1970) Nature @on&m) 227,680685. 26. EOBARZEWSKI, J., AND PASZCZYASKI, A. (1985) Enzyme MicrobioL Technd, in press. 27. MORRIS, D. R., AND HAGER, L. P. (1966) J. BioL Chew. 241,1763-1768. 28. MORTON, R. A. (1975) Biochemical Spectroscopy, pp. 308-357, Wiley, New York. 29. NOGUCHI, A., SHIMADA, M., HUGUCHI, T., AND AOSHIMA, K. (1978) Proceedings, 23rd Lignin Symposium, Ehime (Japan), pp. 9-12, Ehime University, Ehime. 30. OLSON, J., AND DAVIS, L. (1976) B&him. Biophys. Acta 445,324-329. 31. PAJOT, P., AND GROUNDINSKY, 0. (1970) Eur. J. B&hem. 12,158-164. 32. PALMER, J. G. (1983) Mycologia 75,995-1004. 33. PALMER, J. M., AND EVANS, C. S. (1983) in Proceedings of the International Symposium on Wood and Paper Chemistry, Vol. 3, pp. 19-24. 34. PASZCZY~SKI, A., HUYNH, V. B., AND CRAWFORD, R. L. (1985) FEMS Microbial Z&t. 29,37-41.

35. RENGANATHAN, V., MIKI, K., AND GOLD, M. H. (1985) Arch. B&hem. Biophys. in press. 36 ROBINSON, J., AND COOPER, J. M. (1970) Anal. Biochem. 33,390-399. 37. SARKANEN, K. V., AND LUDWIG, C. H. (19’71) Lignins: Occurrence, Formation, Structure, and Reactions, Wiley-Interscience, New York. 38. SCHOEMAKER, H. E., HARVEY, P. J., BOWEN, R. M., AND PALMER, J. M. (1985) FEMS I&t. 183, 716. 39. SCHULTZ, J., AND SHMUKLER, H. W. (1964) Birr chemistry 3,1234-1238. 40. STEELINK, C. (1966) in Lignin Structure and Reactions (Gould, R. F., ed.), pp. 51-64, Advances in Chemistry Series, Am. Chem. Sot., Washington, D. C. 41. SPENSER, R. L., AND WOLD, F. (1969) Anal Biochem. 32,185-190. 42. SUBRAMANIAN, J. (1975) in Porphyrins and Metaloporphyrins (Smith, K. M., ed.), pp. 555-589, Elsevier, New York. 43. TAI, D., TERASAWA, M., CHEN, C. L., CHANG, H-M., AND KIRK, T. K. (1983) in Recent Advances in Lignin Biodegradation Research (Higuchi, T., Chang, H-M., and Kirk, T. K., eds.), p. 44, Uni Pub. Co., Tokyo. 44. TIEN, M., AND KIRK, T. K. (1984) Proc. N&L Acad Sci USA 81,2280-2284. 45. WOOD, D. A. (1985) in Developmental Biology of Higher Fungi (Moose et al, eds.), British Mycol. SOC., pp. 375-387, Cambridge Univ. Press, London. 46. YAMAZAKI, Biophys.

I., AND PIETTE, Acta 77,47-64.

L. H. (1963)

Biochim