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AN STATE UNIVERSITY LIBR Illl’llxlll m Hill I will 3 1293 00899 7656 This is to certify that the dissertation entitled The Involvement of Glucose oxidase in Lignin Degradation by the White-Rot Fungus Phanerochaete chysosporium presented by Robert L. Kelley has been accepted towards fulfillment of the requirements for Ph-D- degree in Micmhiolagy CHM Major professor C. A. Reddy Date F¢5ruwry /9z/77/ MS U i: an Affirmative Action/Equal Opportunity Institution 0-12771 i LIBRARY {Michigan State l University PLACE IN RETURN BOX to remove this checkout from your record. TO AVOID FINES return on or before date due. DATE DUE DATE DUE DATE DUE ":- ~; 1 q to, n l:__ ——l——l_ MSU is An Affirmative ActiorVEqual Opportunity Inflation chnS-r THE INVOLVEMENT OF GLUCOSE OXIDASE IN LIGNIN DEGRADATION BY THE WHITE-ROT FUNGUS BHAHEBQQEAEIE QHBXSQSBQBIHM BY Robert Lewis Kelley A DISSERTATION Submitted to Michigan State University in partial fulfillment of the requirements for the degree of DOCTOR OF PHILOSOPHY Department of Microbiology and Public Health 1990 my ~5730 ABSTRACT THE INVOLVEMENT OF GLUCOSE OXIDASE IN LIGNIN DEGRADATION BY THE WHITE-ROT FUNGUS EHAHEBQQHAEIE QHBXfiQfiBQBIHM BY Robert L. Kelley Previous studies have shown that the production of hydrogen peroxide (H202) plays an important role in lignin degradation by the white-rot fungus, Enangrggngete chrxfigspgzinm. However, the metabolic source of H202 in ligninolytic cultures of this fungus was not known. In this study, glucose oxidase was identified as the primary physiological source of H202 in cell extracts of ligninolytic cultures of E. ghrysggpgrinm. Polyacrylamide gel electrophoresis of extracts from ligninolytic cultures followed by the diaminobenzidine/horseradish peroxidase staining procedure showed the presence of a single protein band that exhibited glucose-dependent H202 production. Both glucose oxidase activity and lignin degradation were triggered in response to nitrogen (N) or carbohydrate starvation, and were repressed in media containing high levels of N (24 mM) and carbohydrate (56 mM) or on the addition of exogenous N sources, such as glutamate, to the low N medium (2.4 mM N). The results indicated that glucose oxidase activity was the primary source of H202 production in ligninolytic cultures of E. gnrysgspgrium and that nutritional parameters which affected lignin degradation had a parallel affect on glucose oxidase activity. Glucose oxidase from ligninolytic cultures of E. ghzysgspgzium was purified to homogeneity and was shown to be a flavoprotein with an apparent native molecular weight of 180,000 daltons and a denatured molecular weight of 80,000 daltons. It had optimal activity with D-glucose, which was stoichiometrically oxidized to D-gluconate. The Km values for glucose and 02 were 38 mM and 0.95 mM, respectively. The enzyme had a pH optimum of 4.5. It was inhibited by Ag+ and o-phthalate but not by Cu++, NaF or KCN. In an effort to better understand the involvement of glucose oxidase in lignin degradation, mutants deficient in this activity (ggx:) were isolated and characterized. The mutant strains had little ligninolytic activity (2-(14C) synthetic lignin == 14C02, were unable to decolorize poly R dye 481, and were also deficient in "ligninase" (a lignin-degrading enzyme) and peroxidase activity. 993+ revertants regained most of the lost activities including the ability to degrade lignin. The results suggest that the genetic lesion in ggx’ mutants affects the regulation of a set of secondary metabolic characteristics. To my parents, Erin, Jacki and Karin iv ACKNOWLEDGMENTS I would like to express my appreciation to Dr. C.A. Reddy for his valued counsel and patience during the course of this study. I am also grateful to Drs. H. L. Sadoff, N. E. Tolbert, P. T. Magee, J. M. Tiedje and J. Dodgson for serving on my guidance committee. The critical suggestions and assistance of numerous others throughout the university have been very useful and are gratefully acknowledged. Finally, I thank the Department of Microbiology and Public Health for the financial assistance provided to me. TABLE OF CONTENTS List of Tables List of Figures Introduction Literature Review A. C. D. The Structure and Chemistry of Lignin 1. Biosynthesis and Structure 2. Occurence and Function of Lignin in Nature Biodegradation of Lignin 1. Quantification and Chemical Characterization of Lignin Degradation 2. Lignin Degrading Organisms 3. Physiology of Lignin Biodegradation Biochemistry of Lignin Degradation 1. Enzymology of Lignin Degradation 2. Involvement of Reduced Oxygen Species in Lignin Degradation References Cited vi viii 11 12 12 15 19 24 26 29 37 Chapter I: Chapter II. Chapter III. Identification of Glucose Oxidase Activity as the Primary Source of Hydrogen Peroxide Production in Ligninolytic Cultures of CDIX§Q§DQILEE 55 Purification and Characterization of Glucose Oxidase From Ligninolytic Cultures of Enanezggnggte chrxscsncrium 86 Characterization of Glucose Oxidase- Negative Mutants of a Lignin Degrading Basidiomycete Enanerggnaete chrxscsncrium 119 vii Chapter I 1 Chapter II 1 LIST OF TABLES EQQQ Industrial Applications for Lignin 4 Major Intermonomer Linkages and Their Frequencies in Gymnosperm and Angiosperm Lignin 9 The Effect of Different Substrates on H202 Production by Cell Extracts of Glucose- Grown Ligninolytic Cultures of R. 77 The Effects of Different Co-Substrates (Growth Substrate) on Ligninolytic Activity and on H202 Production by Cultures of 2. Wm 78 The Effect of Nitrogen and Carbohydrate Concentrations on the Specific Activity for Lignin Degradation and Glucose Oxidase Activity 79 Purification Data for Glucose Oxidase From Ligninolytic Cultures of Enanezggnagte Win 106 Comparison of the Effects of o-Phthalate, KCN, NaF and Different Metal Ions on Glucose Oxidase From 2. ghzysgspgzium and Commercially Prepared A. niger Glucose Oxidase 107 viii 3 Substrate Specificity of Purified Glucose Oxidase 103 Chapter III 1 Characteristics of the Wild Type, Glucose Oxidase-Negative Mutants and Revertants of 2. Win 132 2 The Effects of Exogenous Addition of Ligninase and Glucose Oxidase on Lignin Degradation by the Wild Type and Glucose- 0xidase Negative Mutants of B. W 133 Chapter 1 1 Chapter 2 1 LIST OF FIGURES The Structure of the Immediate Precursors Used in the Biosynthesis of Lignin Schematic Representation of Spruce Lignin Degradation of Lignin Model Compounds by an Enzyme From 2 chrxsesncrium Polyacrylamide Gel Electrophoresis of Cell Extracts From 6 Day-01d Cultures of B. ghzysgspgrinm Grown in Low N Medium (Containing Glucose as the Substrate) Polyacrylamide Gel Electrophoresis of Cell Extracts of E cnrxsosncrium Polyacrylamide Gel Electrophoresis of Cell Extracts of Ligninolytic Cultures Grown on Various Growth Substrates A. Elution Profile of Protein and Glucose Oxidase Activity (nmole/min-mg) of Sephacryl S-300 Column B. Elution Profile of Protein and Glucose Oxidase Activity From a Column of DEAE- Sepharose Sodium Dodecyl Sulfate Polyacrylamide Gel Electrophoresis (SDS-PAGE) Preparation From Different Purification Steps (See Table 1) X 10 35 8O 82 84 109 111 Chapter 3 1 Molecular Weight Determination of Purified Glucose Oxidase by Gel Filtration Chromatography with a Sephacryl S-300 Column 113 Molecular Weight Determination and Staining for Carbohydrate and Protein After SDS- PAGE of Commercial Glucose Oxidase From A. niger and that from B chrxagsncrium 115 Lineweaver-Burk Plots Showing the Effect of Glucose and 0 Concentration on Glucose Oxidase Activ1ty 117 Release of 14C0 From 2'-14C-Synthet1c Lignin by the W1ld Type ( c) ),m Mutant (COX-10 ; A.) and LIG-5 Mutant ( to ) 134 xi Intreducticn Lignin is one of the most abundant biopolymers in nature constituting approximately 20-30% of the dry weight of all vascular plants (23,94,136). Hence, lignin biodegradation and its utilization as a renewable resource for the production of chemical feedstocks and other useful products is of great interest. Previous studies have shown that lignin biodegradation is an oxidative process (101) and that hydrogen peroxide (H202) plays an important role in this degradation (136,137). A temporal correlation between ligninolytic activity and H202 production by Ehgngggghggte gnzysgspgzium has been demonstrated (45,46). Growth under 100% 02 markedly increased both H202 production and ligninolytic activity, and lignin degradation was inhibited by catalase and a scavenger of H202 (38). Cytochemical diaminobenzidine (DAB) staining techniques revealed that H202 production in ligninolytic cultures of BL ghrysgspgzium, is localized in ovoid periplasmic microbody-like structures which are not found in non-ligninolytic 2 cells (46). Several HZOZ-dependent, lignin degrading enzymes, have been isolated from the extracellular fluid of P. ghrysgspgrium cultures. These enzymes were only found in ligninolytic cultures and have been shown to oxidize a variety of lignin model compounds (54,62,152,153), depolymerize lignin and decolorize the polymeric dye poly R481 (62,113). It became obvious from these studies that H202 plays an important role in lignin degradation by E; ghzysgspgrinm, however, the physiological source of H202 in ligninolytic cultures of this organism was not known. The objective of this research is to determine the physiological source of H202 in lignin-degrading cultures of 21 chrygsgpgrium, study the nutritional and physiological parameters regulating this activity. Also, the enzyme(s) involved in H202 production will be purified and characterized. 1.! ! E . Lignin is a three-dimensional, highly-branched, amorphous, aromatic polymer present in all vascular plants. Unlike other natural polymers, such as cellulose, starch and proteins, it has no precise structure and contains no readily hydrolyzable linkages that repeat at regular intervals (129,165). 3 Instead, lignin is an irregular polymer with many intermonomeric linkages recurring randomly throughout the molecule and thus, is one of the most resistant compounds to biological degradation. Lignin is one of the most abundant and widely distributed renewable organic polymers on earth. It constitutes 25% of the dry weight of the estimated 100 billion metric tons of biomass produced annually in the biosphere, and is second only to cellulose in its abundance (136). However, lignin contains 60% of the carbon and 40% of the fuel value of total biomass (88) and thus is more important than its weight would indicate. Furthermore, it has been estimated that 1-3 x 1012 metric tons of lignineous material, primarily as peat and humus (161), have accumulated in the soil. Because of its great abundance and recalcitrance to microbial attack, lignin biodegradation plays an important role in the terrestrial carbon cycle. In the past decade, the possible industrial uses of lignocellulosic materials for the production of various fuels, animal feeds and chemical feedstocks have been discussed in numerous reviews (20,28,34,87,95,107,136). A recent review by Janshekar and Fiechter (87) discussed in detail the possible applications and products which can be made from lignin (Table 1). It is estimated that 900-3,000 Table 1: Industrial Applications for Lignin. 1. Energy 2. Polymers and modified polymers filters, rubber reinforcments, carriers for controlled release in fertilizers and pesticides, dispersants, emulsion stabilizers, complexing agents 3. Prepolymers polyphenolics, resins and extenders, foams, adhesives a. Fragmentation and chemical conversion a. b. hydrogenation---phenols, hydrocarbons hydrolysis---phenols, catechols, substitued phenols . oxidation---vanillin, dinethyl sulfide . alkali fission---phenolic acids, catechols . pyrolysis?--acetic acid, phenols, CO, 602, CH4 . fast thermolysis---acetylene, ethylene Adapted from reference 87. 5 million tons (87) of lignocellulosic by-products are generated annually by forestry, agriculture, paper- making and lumbering industries in the U.S. alone. Lignin occurs in close chemical and physical association with cellulose and hemicellulose in these materials and limits the efficient utilization of these carbohydrate polymers into useful products. Numerous pretreatments which depolymerize, solubilize or otherwise remove lignin from lignocellulosic materials, have been developed (17,39); however, most of these processes are either too energy intensive, create large quantities of chemical wastes or cannot be used on an industrial scale (95). Thus, in recent years lignin biotransformation by microbes has been an area of great interest (17,95,98,121,154). It is hoped that controlled microbial treatment of lignocellulosic materials might be a cost effective and energy efficient pretreatment process for the optimum utilization of biomass as useful products. As fossil fuel feedstocks become more scarce, chemical feedstocks, fuels and solvents made from biomass are likely to become economically competitive with those currently produced from petroleum (73). In this review, I will discuss briefly the available literature on the biodegradation of lignin. I will focus on recent evidence for the involvement of 6 reduced oxygen species, in particular H202, in lignin degradation by certain wood—rotting fungi. S! ! l :1 i ! E I' . A number of recent reviews (1,65,72,134) and books (13,28,105,132,147) have covered the accumulated knowledge on the chemistry and biosynthesis of lignin and should be consulted for a more complete treatment of the subject. Bigsynthesi§_and_fitrugture. Lignin is a complex and variable biopolymer made of three cinnamyl alcohol derivatives; p-coumaryl, coniferyl and sinapyl alcohols (Figure 1). These precursors are synthesized by the general path: C02 ==> carbohydrates ==> phenyl propanoid amino acids ==> cinnamic acid derivatives ==> cinnamyl alcohol derivatives (28,77). The ratio of these three alcohols are known to vary between lignins from different plant species and with the age and type of tissue within a single species (145). Higuchi et a1. (77) defined three major groups of lignins based on the relative ratios of these derivatives: Softwood lignin or guaiacyl lignin (found in most conifers, lycopods and ferns) is composed mostly of coniferyl alcohol units with a small amount of coumaryl and synapyl alcohol units. The relative proportions in spruce lignin are 80:14:6, CH CH - ll ll CH cH. @ ©-CH3 CH, 'ti 0&1 P-COUMARYL COilIFERYL SINAPYL ALCOHOL ALCOHOL ALCOHOL Figure 1. The structure of the immediate precursors used in the biosynthesis of lignin. 8 respectively. Hardwood lignin or guaiacyl-syringyl lignin, found mostly in angiosperms, contains approximately equal amounts of coniferyl and synapyl alcohol units with only minor amounts of coumaryl alcohol units. The proportions in beechwood are 49:46:5, respectively. Grass and bamboo lignin (guaiacyl-syringyl-p-hydroxyphenyl lignin) are thought to be composed of approximately equal amounts of all three cinnamyl alcohols; however, the exact ratios are not known because a considerable amount of p-courmaric acid is bound as esters to grass lignins (80). Lignin is formed by the dehydrogenative polymerization of the above-mentioned cinnamyl alcohol derivatives (48,49,72,79,145). At the site of lignification these alcohols are oxidized by phenol- oxidases to yield phenoxy radicals which, because of their extended electron systems, are stabilized through equilibrium with several mesomeric forms. These different radical species derived from the three cinnamyl alcohols randomly condense with each other and with the radicals in the growing lignin polymer. A variety of intermonomeric linkages result (Table 2), including a number of C-C and c-o-c linkages, forming a complex, three-dimensional polymer. Several models of hard- and soft-wood lignin have been proposed (1,72,124), including a model by Harkin (Figure 2) of Table 2: Major intermomer linkages and their frequencies in gymnosperm (spruce) and angiosperm (birch) lignins. ': of total 09 Linkage type units in Figure 3 units . spruce _ birch fl-Aryl ether (fi-O-4) 1-4, 5-4 5-6, 7-8, 48 60 13b-l4b, 15-16 Phenylcourmaran 17-18 9-12 6 (19-5) Biphenyl (biaryl or 12-13a 9.5-11 4.5 5-5) 1,2-Diarylpropane 16-20 7 7 (19-1) Diphenyl ether 6-7 3.5-4 6.5 (diaryl or 4-0-5 a-Aryl ether (a-0-4) 11-12 6-8 6-8 Pinoresinol (fi-fl) 8-9 2 From reference 136. 10 Figure 2. Schematic representation of spruce lignin (from reference 136). 11 a spruce lignin. The molecular weight of purified lignin varies from a few thousand to more than 106 daltons, depending on the isolation method. Therefore, the polymer illustrated in Figure 2 and most other lignin models published represent only a portion of the total lignin molecule (28). O 1 E !' E I' . I H ! . Approximately 80% of lignin in plants is found within the secondary cell walls, closely associated with hemicellulosic and cellulosic components (145). However, the highest concentration of lignin is present in middle lamella, serving as an intercellular cementing substance that binds the cells together (44). Lignin is a recalcitrant polymer and thus protects plant cells against infection by plant pathogens. Lignin also minimizes water permeation across the cell wall of xylem tissue, imparts structural rigidity, and confers resistance to impact, compression and bending (145,146). A major portion of the carbohydrate components of the cell wall are quickly metabolized when the plant dies, but lignin, although structurally modified is degraded only to a limited extent and forms a major constituent of humus and peat (164,165), which increase the aeration and moisture holding capacity of the soil and serves as an ion exchange resin 12 sequestering important inorganic molecules that increase the general soil fertility (82). El: l!’ EI" i !°E° !' 1 ll . J :1 ! . !' E Lignin_negradatign. Major advances have been made in understanding the biological decomposition of lignin because of the development of specific and sensitive radioisotopic methods for assaying metabolism of lignin. These assays are based on the oxidative 14C-[lignin]-lignocelluloses or 14C02. metabolism of Both the rate and extent of lignin biodegradation can be determined in these radioisotopic assays by trapping and quantifying the 14C02 produced from these substrates. 14C-[ligninJ-lignocellulose can be selectively extracted from plants which have been fed labeled precursors of lignin biosynthesis (21,25,30,31,69), 14C-phenylalanine. 0n the other hand, 14C- such as DHPs are prepared by the dehydrogenative polymerization of chemically synthesized 14C-coniferyl alcohol in a peroxidase/H202 catalyzed reaction (70,103). The advantages of using DHPs are: 1) they do not contain extraneous materials, such as carbohydrate; 2) they can be specifically labeled in 13 the side chain, aromatic ring or methoxyl groups and, therefore, the specific (structural/ chemical) nature of the attack on the lignin polymer by a particular microorganism can be determined. It is important to note that there may be considerable differences between the amount of 14C02 detected when 14C- [1ignin1-lignocellulose and 14C-DHP are used as substrates. For example, Robinson et al. (144) found that a Bacillus 59. could degrade approximately 12% of 14C-[lignin]-lignocellulose, but only 0.3% of 14C—DHPs to 14002 in 20 days. Also, failure to detect 14C02 14 from C-lignins does not exclude the possibility of significant but partial degradation of the lignin polymer. Recently, Glenn and Gold (54) showed that ligninolytic activity by E. chrysgspgrium is closely correlated with its ability to decolorize of polymeric dyes (e.g. Poly B-411, Poly R-481 and Poly Y-606) and have suggested that decolorization of these readily soluble, stable, inexpensive substrates could be used as a simple screening test for the selection of non- lignin degrading mutants of E. ghzysgspgrium. Kelley and Reddy (91) showed a similar correlation between the production of ethylene from a-oxo-q- methylthiobutyric acid (KTBA) and ligninolytic activity in P. gnrysgspgrium and suggested that l4 ethylene production from KTBA can be used as a sensitive measure of ligninolytic activity by wild- type B- W. A variety of other chemical and physical methods of varying degrees of reliability have also been developed for the study of lignin degradation (28, 87). Spectroscopic methods, including UV, IR and NMR, have been used to monitor lignin degradation (15,89,115,118). Chromatographic techniques, such as gel permeation chromatography, high performance liquid chromatography, and gas chromatography have been used to determine changes in molecular weight distribution of the lignin polymer and to identify aromatic acids produced during its degradation (81,85,100,102,117,118). Electron microscopic studies 'have also been used to understand the morphological changes in wood components that have been subjected to attack by wood-rotting fungi (4,11,36,44). Functional group analysis and elemental composition have provided useful information about the chemistry of lignin degradation (101,102). However, because of the complexity and diversity of the lignin polymer, a combination of chemical and/or physical methods mentioned above should be used to obtain complete information about the wide variety of modifications that occur during lignin degradation (87). 15 Lignin_negrading_grgani§m§. Previous workers have shown that several different genera of bacteria and fungi are capable of metabolizing lignin and several reviews have been published on this subject (6,14,23,29,94,96,98,l35, 136,164). Of the ligninolytic microorgansisms described to date, a group of mostly basidiomycetes and a few ascomycetes, called white-rot fungi, are known to degrade lignin more rapidly and more extensively than other micro- organisms. Consequently, this group of microorganisms has been studied most thoroughly. White-rot fungi are recognized by their ability to deplete all the major wood components: cellulose, hemicellulose and lignin and cause a whitish coloration in the decayed wood (4,28,98). These organisms can be separated into two primary groups: 1) simultaneous rot fungi which degrade all wood components at approximately the same rate and 2) white-pocket rot fungi which preferentially degrade lignin compared to cellulose and hemicellulose (11,87). 2. ghrysgspgzinn, Coriglus mam. and Elements estreatus. which are simultaneous rot fungi, have all been shown to degrade 14C-DHPs labeled in the ring, side chain or methoxyl group to 14C02 (67,68). Chemical analyses, spectroscopic analyses and chemical degradation studies, of spruce lignin attached by E. 16 W,Wm.mw and szia_subagiga showed that lignin degradation by these fungi is largely oxidative as evidenced by the high oxygen content, and the low hydrogen and methoxyl content of the decayed polymers in comparison to sound lignin (101,102). Electron microscopic studies of white-rot decay have shown that erosion troughs are formed in the immediate vicinity of the fungal hyphae (4,6,115) which are believed to be caused by extracellular catalysts. SEM studies revealed selective removal of lignin, but not cellulose and hemicellulose. by lecbolus frustulans. znellinus Dial and Ionctus drxcnhilus (11.130)- Brown-rot fungi, which include numerous genera (53) that are taxonomically similar to white-rot fungi, characteristically attack the carbohydrate components of wood in preference to the lignin components and leave a brown residue in the decayed wood (8,97). Kirk and Adler (99) showed, by chemical analyses of sweetgum decayed by Lenzites grapes, that brown-rot fungi cause considerable alteration of the lignin polymer, including demethoxylation and ring hydroxylation, but little aromatic ring cleavage was detected. Thus, the principal difference between brown-rot and white-rot fungi appears to be the ability of the latter to metabolize aromatic rings 17 (97). The soft-rot fungi, which include a variety of ascomycetes and fungi imperfecti (37,67,68), principally attack the carbohydrate components of wood, and the decay is usually accompanied by a softening of the surfaces of the woody tissue. Eslyn et a1. (37) examined the degradation of specific wood components from various woods decayed by strains of Grannies. Moncdicxls. Baccilcmxces. Eanulsncra. Ihielayia and Allesgheria, and found that in alder and poplar wood the carbohydrate moieties were depleted faster than lignin by most of the species examined. Haider and Trojanowski (67) found that soft-rot Species, Ereussia. Cheetomium and Stachxhetrxs. were capable of releasing substantial amounts of 14 14 C02 from C-ring-, side chain-, and methoxyl-labeled DHP and from 14C-[lignin] lignocellulose. These studies show 1 that soft-rot fungi can metabolize 4C-lignin to 14C02, but the detailed chemical nature of soft-rotted lignin remains to be elucidated. Bacterial degradation of lignin has recently been reviewed by Crawford and Crawford (23). A number of bacteria including Nggazdia sp. (35), Pseudgmgnads sp. (47). Aercmonas sp- (126). Xanthononas sp- (128). Bacillus_msgaterium (144) and sggggggmyggg Sp, (22- 27), have been shown to degrade 14C-DHP and 14C- 18 [lignin]-lignocellulose to 14C02. For example, Nggardia sp. has been shown to convert 14C-ring-, side chain-, and methoxyl-labeled DHP (5, 15 and 13%, respectively) to 14C02 (153). A number of strains of streptomyces were shown to degrade lignin but apparently cannot use the latter as a sole source of carbon and energy (27). Streptomyges badius was shown to degrade 13% of 14 C-[lignin]-lignocellulose in 42 days and SEM studies showed that it can degrade lignified cell walls in the inner bark of Douglas fir (27,150). A strain of Xanthgmgnas sp. has been reported to degrade as much as 77% of dioxane lignin in 15 days when it is provided as the sole carbon and energy source (128): however, dioxane lignin is not considered to be reliable substrate for studies on lignin biodegradation (28). Pseudgmgnas sp. were shown to be capable of utilizing kraft lignin as a sole source of carbon and energy; in fact, Forney e; 31. (47) found that addition of 0.01% (w/v) glucose to mixed cultures of pseudomonads actually decreases the extent of degradation of kraft lignin. Information about the biochemical nature of lignin degradation by bacteria is sparse: however, Crawford et a1. (12,27,29) found that decay by streptomyces viridgspgzns T7A was oxidative and involved demethylations, ring cleavage reactions, and oxidative 19 attack on phenylpropanoid side chains, which is similar to what is observed during white-rot degradation of lignin. Until recently, anaerobic degradation of lignin was thought not to occur to any great extent (126,127,164). Brenner et a1. (10) recently reported 14C-lignins incubated with methanogenic degradation of anaerobic lake sediments (10). They have shown significant (17% in 294 days) anaerobic degradation or of 14C-[lignin]-lignocellulose‘inlaquatic sediments has been reported. However, the chemical nature of this degradation is not known. Also, a number of lignin model compounds, including 3,4,5 trimethoxybenzoic, syringic and 3-0-methylgallic acid as well as methoxylated and hydroxylated benzoate have been shown to be degraded by pure cultures of obligate anaerobic bacteria (74,163). 2].] EI"Eii 3!. Most of the recent investigations on the physiology and biochemistry of lignin involved a single species, B. ghrysgspgzium Burds (ATCC 34541: 16). The attractiveness of this organism lies in its rapid growth, extensive degradation of lignin, prolific conidiation, relatively high temperature optimum (40°C) and relatively low phenol oxidase 20 activity (which can cause repolymerization of degraded lignin fragments: 94,96). Also, optimal culture conditions for lignin degradation by this organism are known. These studies showed that lignin degradation is optimal at a pH of 4 to 4.5. (42,108). Sodium 2,2- dimethylsuccinate is the preferred buffer, whereas 0- phthalate inhibits lignin degradation (42). A mixture of inorganic nitrogen source (e.g. NH4NO3) and an organic nitrogen source (e.g. asparagine) promotes optimal growth and lignin degradation (93,108). Thiamine is required for growth and a mixture of trace metals stimulate growth and lignin degradation (88,103,140). In the past decade a wealth of information has become available on the physiology of lignin degradation by P. chrysgspgrium. These studies show that liginolytic activity by P. ghrysgspgrium is a secondary metabolic event which is derepressed following the cessation of primary growth in response to either nitrogen, carbohydrate, or sulfur starvation (88,108). Synthesis of lignin degrading system does not require the presence of lignin in the growth medium (93). Lignin does not serve as a sole source of carbon/energy for E. ghzysgspgrium or for a number of other white-rot fungi and that a co-substrate, such as glucose, xylose, cellulose, cellobiose or 21 succinate, is required for lignin degradation (3,104,108). Although the ligninolytic system is known to be synthesized in the absence of lignin, recent studies show that ligninolytic activity is several fold higher when E. ghrysgspgzium is grown in the presence of lignin (155) or veratryl alcohol (38). Nitrogen_£ffiegt: Lignin degradation by B. ghzysgspgrium is profoundly affected by the level of nutrient nitrogen provided. Cultures grown with limiting amounts of nitrogen (2.4 mM) rapidly deplete the N source and enter the stationary phase of growth (108,114,138,140,162) during which the ligninolytic system appears and secondary metabolites, such as veratryl alcohol (119,148), are produced. Furthermore, the addition of exogenous nitrogen, such as NH4+ or glutamate, to ligninolytic cultures inhibits lignin degradation (41,43). Glutamate, glutamine and histidine suppress ligninolytic activity, (83, 76 and 76%, respectively), compared to a control with no additions (41). The addition of an N source to ligninolytic, idiophasic cultures appears to cause the transition to primary growth, stimulating not only glucose and succinate metabolism but growth in general, and stopping secondary metabolic events, such as synthesis of veratryl alcohol and lignin degradation (43). Kirk gt _1. (94) hypothesized that 22 (nitrogen metabolism via) glutamate (metabolism) plays an important role in the initiation of the repression of secondary metabolism and in turn ligninolytic activity. In addition, Reid (140) has suggested that C/N ratio maybe more important to the control of lignin degradation than the absolute levels of nitrogen because abundant carbon causes the available nitrogen to be used in cell synthesis so that it does not repress ligninolytic activity. Recently, cyclic AMP levels have been shown to rise 10-fold in response to nitrogen starvation, and it has been speculated that cyclic AMP levels may be important in the regulation of secondary metabolism, including ligninolytic activity (120). Degradation of a lignin model compound, 4- hydroxy-3-methoxyacetophenone as well as other lignin model compounds with b-guaiacyl ether-linkages has also been shown to occur in nitrogen starved cultures and was inhibited in high N cultures or by the addition of exogenous NH4+ to nitrogen starved cultures (159). Interestingly, Odier and Roch (129) found that some white-rot fungi such as Dighgmitus squalens, showed no suppression of the ligninolytic system by high levels of nitrogen. 9W: Recently . Jeffries at al. (88) have shown that ligninolytic 23 activity (14C-synthetic lignin ==> 14C02) occurs in fungal cultures containing nonlimiting nitrogen (24 mM), but limiting glucose (8.8 mM). They have also shown that these culture conditions cause autolysis of the fungal cells and that autolysis ceases when ligninolytic activity does, suggesting that autolysis of some cells provides carbon and energy sources for other remaining cells to cause lignin degradation. Ligninolytic activity has also been reported to be triggered by limiting sulfur (20 mM) (88). However, Reid (139) in similar studies did not observe any stimulation of ligninolytic activity by E. chrysgspgrium under sulfur limiting conditions using 14C-(lignin) lignocellulose as the substrate. Qxygen_£ff§gt: Kirk et al. (108) observed a 2- fold increase in ligninolytic activity by E. gnzysgsgzium when grown under 100% 02 as opposed to air. Bar-Lev and Kirk (9) showed that 02 did not induce the ligninolytic system, but enhanced lignin metabolism after the system was formed. These results are consistent with the finding that lignin degradation is a highly oxidative process. However, 02 pressure above 1 atm inhibits the growth of E. enzysgspgzinm and does not increase the rate of lignin metabolism (142,143). Lignin degradation by 2. ghzysgspgrigm as well as 24 Coriglus ygrsigglgr has been shown to be strongly inhibited by agitation of the cultures (108,162). It is believed that agitation causes pellet formation which restricts not only the cell surface area exposed to the lignin polymer, but also limits the amount of 02 diffusion into the interior of the pellet. 'However, recent work by Reid (141) seems to contradict these findings. In these studies, agitation of ligninolytic cultures did not significantly affect the rate of extent of 14C-DHP degradation and in fact 1 increased the degradation of 4C-[lignin1- lignocellulose. E' l l ! E II l D 1 !' Our knowledge of the biochemical mechanism of lignin degradation has been advanced significantly in the past 5 years, and this knowledge and associated methodologies should contribute greatly to our understanding of lignin biodegradation in the coming few years. Recent studies have extended our knowledge on degradative pathways for several low molecular weight lignin model compounds used by white-rot fungi (87). The involvement of oxygen radicals (2,38,45,112) and extracellular lignin degrading enzymes (50,113, 151,152) in lignin degradation have been perhaps the most significant recent discoveries 25 in this area. Techniques for selecting mutants (58,59) and complementation analysis (56,59,60,61) have been developed. Finally, an autonomous replication sequence from E. chrysgspgrium has been characterized (133) and construction of lambda genomic library of this organism has been described and is being used for obtaining a better understanding of the genetics of lignin degradation (158). Lignin biodegradation, based on cumulative evidence in the past two decades, appears to be an extracellular, oxidative and nonspecific process. Because of the size of the lignin polymer, direct uptake by microbial cells seems unlikely, and therefore, at least the initial attack on the lignin polymer must be extracellular. Analyses of white- rotted lignins and the solubilized products have shown a substantial increase in carboxyl and carbonyl groups and a decrease in hydrogen content suggest the oxidative nature of lignin degradation (84,85,101,102). Oxidation of the a-carbon of the propyl side chain of lignin monomers to a carbonyl group and hydroxylation and oxidative cleavage of the aromatic rings (94) offers further support to this conclusion. The fact that lignin is extensively degraded despite the variety of intermonomeric linkages (94) and the racemic nature of the asymmetric 26 carbons in the polymer indicates that the attack on lignin is nonspecific and nonstereoselective (122,125,149). Also, a variety of polymers which are structurally different, such as polyguaiacol (32) and Poly—R dyes (54), are metabolized efficiently by white-rot fungi. Therefore, given the specific nature of most enzymes, it was first thought that there would be a wide variety of enzymes involved in lignin degradation. WW While most of the enzymes responsible for lignin degradation have not been determined, several types of enzymes from various microorganisms which are thought to participate in lignin degradation have been described: 1) phenoloxidases (laccase, peroxidase and tyrosinase), 2) mono- and di-oxygenases, 3) aromatic alcohol oxidase and moist recently, 4) "ligninase", a new class of lignin-degrading enzymes. All these enzymes could account for the oxidative nature of lignin degradation, were extracellular and were relatively non-specific as evidenced by their reactivity with several types of lignin model compounds. The phenol-oxidizing enzymes known collectively as phenol oxidases includes three distinct types of enzymes: laccase (02: p-diphenol oxidoreductase), 27 peroxidase (donor: H202 oxidoreductase). Reviews by Ander and Eriksson (7,8) present a detailed description of the reactions catalyzed by these enzymes. Phenol oxidases can cause limited structural changes in lignin, but their involvement in total degradation of lignin to C02 is unlikely (28,52,66,87,90,125). The functional roles attributed to phenol oxidases include: 1) detoxifying low molecular weight phenols released during lignin degradation (52), 2) performing reactions critical to the preparation of the lignin polymer for degradation, such as demethylation or removal of asymmetric centers making the polymer more desirable to stereospecific attack by enzymes (102,149) 3) limited depolymerization of lignin polymer and 4) regulation both lignin-degrading and polysaccharide-degrading enzymes (5). Mutants of E. ghzysgspgrium lacking phenol oxidase were unable to degrade lignin; however, these mutants were later shown to be pleiotropic for a variety of other enzymes (5,59). Furthermore, at least one of these mutants (phe3) was shown recently to extensively metabolize certain lignin preparations (Ander, personal communication). Thus, the actual function of phenol oxidases in lignin degradation is yet to be established. Both mono- and di-oxygenases can catalyze 28 reactions which would aid in the biodegradation of lignin. Mono-oxygenases by incorporating one atom of oxygen cause of hydroxylation of methylated or demethylated aromatic rings (18) which results in the formation of ortho-diphenols and an increase the solubility of lignin substrates. Recently, lignin- degrading oxygenases have been isolated (see below) which actually appear to be peroxidases. Dioxygenases incorporate both atoms of oxygen into the ring structure which helps in ring cleavage (78). Both mono- and di-oxygenases have been reported in a variety of white- and brown-rot fungi (78), but their exact involvement in lignin degradation is not known. An aromatic alcohol oxidase, isolated in culture filtrates of Eusarium_sglani (86) and 29125319335 yezsigglgr (40), oxidizes a wide variety of aromatic monomers and dimers as well as various lignin preparations having a, B-unsaturated alcohol groups in their side chains. This enzyme is extracellular, nonspecific and reacts with the lignin polymer and could be important in lignin degradation by this organism in preparing the phenylpropanoid side chains for further oxidation (28). Several other enzymes have been thought to be important in lignin degradation. Ander and Eriksson (8,160) proposed that cellobiose:quinone 29 oxidoreductase (CBQase) is involved in the degradation of both cellulose and lignin by Spgrgtrighum pulxerulentum, but its requirement for lignin degradation has not been shown. Greene (63) has postulated the involvement of glucose oxidase in lignin degradation by Eglypgrgus yersigglgr based on the rationale that it can: 1) prevent the toxic accumulation of quinoid intermediates and 2) improve the efficiency of lignol oxidations. He has suggested that glucose oxidase preferentially reacts with quinoid than 02. This suggestion would lead to the prediction that reduced oxygen tension should increase lignin degradation, when in fact the opposite is true. Also, experimental support for the proposed reactions with the lignin polymer is lacking. And in fact, no enzyme has been isolated by Greene that could catalyze many of the complex reactions that occur during lignin degradation by white-rot fungi, such as cleage of 8-0- CB side-chain bonds, 4 linkages, cleavage of Ca- oxidative fission of aromatic ring and demethylations (29). This led to the belief that the initial attack on lignin may not be enzymatic, but due to the action of oxygen radicals (71).. I J ! E E 3 i ; 5 l . Ii . Degradation. Amer and Drew (2) reported that superoxide anion (02') is produced by whole cells of 30 Coriglus yersigglg: and speculated that this anion may be involved in lignin degradation by this fungi. The observation that the addition of superoxide dismutase to cultures of E. ghrysgspgrium inhibits ligninolytic activity supports this idea (38). However, the relative non-reactivity of 02' and the growing belief that essentially all the manifestations of 02’ can be explained by its participation in generating hydroxyl radicals (-OH) in aqueous systems has led to the suggestion that 02' may not be directly involved in lignin degradation, but may serve as a reductant in the iron-catalyzed Haber-Weiss reaction (see below) for the production of -OH (146). Nakatsubo et al. (123) reported inhibition of lignin degradation by PL_ghrysg§pgrium using anthracene-9,10-bis ethanesulfonic acid (AES) as a scavenger of singlet oxygen ('02). However, Kirk gt :1. (106) questioned the involvement of '02 in lignin degradation, after finding that artificial '02 generating systems produce different degradative produces from B-l-lignin model compounds compared to those produced by 2. ghrysgspgzium. Forney et a1 (45) proposed that HZOZ-derived, oOH may be involved in the attack on the lignin polymer by E. ghrysgspgzium. This proposed was supported by the following observations: 1) -OH is highly reactive 31 with a variety of organic compounds (33,71), including rapid degradation/depolymerization of the lignin polymer (T. McFadden, L. Forney and C. Reddy, unpublished data); ii) its attack is nonspecific and nonstereoselective (157): and iii) it is generated in biological systems by a one-electron reduction of H202 (19,33,51: See Reaction 1 and 2 below) which is known to be produced by numerous wood rotting fungi (75,76,109-111). Eentcn_Beacticn 1) 3202 + Fe (II) ==> .0H- + Fe (III) I _: ! J 1 H l _” . E !° Fe(III) chelate + 02- ==> Fe(II) chelate + 02 Fe(II) chelate + H202 ==> -OH + OH- + Fe(III) chelate The involvement of H202 and -0H in lignin degradation by P. ghrysgspgrium has been examined in several studies. The production of H202 and -0H (as measured by ethylene production from a-oxo-q- methylthiobutyric acid) was shown to coincide with the appearance of ligninolytic activity (45,137). H202 production was markedly enhanced by growing the fungus under 100% 02, mimicking what is seen with 32 ligninolytic activity (38). The addition of either -OH scavenging agents (e.g. mannitol and benzoate) or catalase, a H202 scavenger, to ligninolytic cultures of B. gnrysgspgrigm not only inhibited -OH and H202 production, but also, inhibited lignin degradation (38,45). Hydroxyl radical was demonstrated in cell extracts of ligninolytic cultures by showing the oOH- dependent hydroxylation of p-hydroxybenzoic acid to form protocatechuic acid, and by detection of the nitroxide radical of 5,5-dimethyl-1-pyroline-N-oxide by EPR (45). Both the hydroxylation reaction and the formation of the nitroxide radicals were markedly stimulated by azide which is known to inhibit catalase (45). Electron microscopic studies showed that H202 production by ligninolytic cell of B. ghrysgspgrium is located in unique periplasmic microbodies which are not found in non-ligninolytic cells (46). The involvement of H202 and -OH in lignin degradation is in keeping with the earlier findings that lignin degradation is oxidative in nature and is mediated by non-specific, non—stereoselective extracellular agents (54,122). The involvement of -OH is also in agreement with reports that the aromatic rings of lignin are extensively hydroxylated during degradation by white-rot fungi (102). Several enzymatic activities which produce H202 in fungi are 33 known, including glucose oxidase, galactose oxidase, sorbose oxidase and carbohydrate oxides (83). Greene and Gould (64) have suggested that a peroxisomal fatty acyl coenzyme A oxidase activity may be an important source H202 in ligninolytic cultures of E. chrysgspgrium. However, the specific activity for this enzyme is very low and a temporal correlation between the onset of lignin degradation and fatty acyl CoA oxidase activity has not been shown. While many researchers were trying to determine the involvement of oxygen radicals in lignin degradation, a new class of enzymes have recently been isolated from E; ghzysgspgrinm which can account for many of the reactions involved in lignin degradation. Lignin-degrading enzymes ("ligninases") from concentrated extracellular fluid of ligninolytic cultures of B. chrysgspgrium have been reported by several investigators (50,55,62,113,151,152). These enzymes have been purified and characterized. They appear to be similar if not identical. Typical ligninase has a molecular weight of approximately 41,000 daltons and a heme IX prosthetic group. It has an obligatory requirement for H202 for activity, and shows little specificity or stereoselectivity (113,152). Tien and Kirk (152) reported Ca-CB cleavage of side chains of non-phenolic B-l lignin 34 model compounds (See Figure 3A) as well as oxidation of various 8-0-4 model compounds (See Figure 38) by their ligninase. They concluded that their enzyme has oxygenate activity (152). Kuwahara et_al. (113) 180 from 1802 during a showed incorporation of diarylypropane cleavage, suggesting that this enzyme is an oxygenase. Ligninase also has peroxidase activity and generate oOH (as evidenced by ethylene production from KTBA) in the presence of H202. These enzymes do oxidize spruce and birch lignin as well as the polymeric dye, Poly R. Kersten et a1. (92) have proposed a mechanism for this enzyme involving a cation radical intermediate which has been detected during the conversion of 1,4-dimethoxybenzene to p- benzoquinone and methanol. However, the extent of the modification of the lignin polymer by ligninase(s) has not been characterized in detail. As summarized in the scheme below, evidence to date clearly shows that H202 plays an integral role in lignin degradation and that -OH or equivalent reduced oxygen species is involved in lignin degradation. However, it is not clear whether -OH is produced chemically by Fenton-type reaction (45) or by peroxidases, such as ligninases, as shown by Palmer and Evans (131). Also, it remains to be seen whether the -OH or an equivalent species is bound to the 35 A 611,00: cu,ou o c .1 on "I ~33» o. .9 70°" ”0, icon.) HO‘ZC'D tooth). g ‘ ' o‘er) (mcmg (0cm) 1:) °" 3...". (moo: on race.) 8» OH [ad 0" H’ OH * 06"; a OCH, m ecu, m a Figure 3. Degradation of lignin model compounds by an enzyme from Phanerochaete chyrysosporium (from ref. 29) 36 active site of ligninase(s) or not. Research in progress in several laboratories should be able to answer these questions in the near future. Ligninolytic Culture_ H202 'Fe++' Lignin Degrading " ' _ Enzymes i -08 ('OH)? / Lignin Degradation 37 LIST OF REFERENCES 38 LIST OF REFERENCES Adler, E., 1977. Lignin Chemistry -- Past, Present and Future. Wood Sci. Technol. 11:169- 218. Amer, G. I., and S. W. Drew, 1981. The Concentration of Extracellular Superoxide Radical as a Function of Time During Lignin Degradation by the Fungus chiclus yersicelor Dev. Ind Microbiol. 22: 479- 484. Ander, P., and K. E. Eriksson, 1975. Influence of Carbohydrates on Lignin Degradation by the White-Rot Fungus Sperctrichum nulyerulentum Sven. Paperstid. 78: 643- 652. Ander, P., and K. E. Eriksson, 1977. Selective Degradation of Wood Components by White-Rot Fungi. Physiol. Plant. 41:239-248. Ander, P., and K. E. Eriksson, 1976. The Importance of Phenol Oxidase Activity in Lignin Degradation by the White-Rot Fungus Spgrgtrighum pulyernlentum. Arch. Microbiol. 109: 1- 8. Ander, P. A., Hatakka, and K. E. Eriksson, 1980. Degradation of Lignin and Lignin-Related Substances by Sncrotrichum pulyerulentum (Ehanerochaete chrysosncrium)- p. 1-14. In T. K. Kirk, T. Higuchi, and H. Chang (Eds.) Lignin Biodegradation: Microbiology, Chemistry and Applications, Vol. II, CRC Press, West Palm Beach, FL. 10. 11. 12. 13. 14. 15. 16. 17. 18. 39 Ander, P., A. Hatakka, and K. E. Eriksson, 1980. Vanillic Acid Metabolism by the White-Rot Fungus pulyerulentum. Arch. Microbiol. 125:189-202. Ander, P., and K. E. Eriksson, 1978. Lignin Degradation and Utilization by Microorganisms. Prog. Ind. Microbiol. 14:1-58. Bar-lev, 8.8., and T. K. Kirk, 1981. Effects of Molecular Oxygen on Lignin Degradation by Ehanecchaete chryscsncrium Biochem- BicthS- Res. Commun. 99: 373- 378. Benner, R., A. E. MacCubbin, and R. E. Hodson, 1984. Anaerobic Biodegradation of the Lignin and Polysaccharide Components of Lignocellulose and Synthetic Lignin by Sediment Microflora. App. Environ. Microbiol. 47:998-1004. Blanchette, R. A., 1980. Wood Decomposition by Ehellinns (Bones) pini: A Scanning Electron Microscopy Study. Can. J. Bot. 58:1496-1503. Borgmeyer, J. R., and D. L. Crawford, 1985. "Production and Characterization of Polymeric Lignin Degradation Intermediates From Two Different Streptomyges spp. Appl. Environ. Microbiol. 49:273-278. Baruns, F. E, 1952. The Chemistry of Lignin. Academic Press, New York. Broda, P., and A. Paterson, 1983. Lignin Biodegradation Becomes Biochemistry. Nature 306:737-738. Brunow, G., and D. Robert, 1983. Quantitatige Estimation of Hydroxyl Groups in Lignin by NMR and for the Study of Benzylic Reactivity. 1983 IWSPC Proceedings, 1:92-94. Burdsall, H. R., Jr., and W. E. Elsyn, 1974. A New Bhanerechaete With a chyrscsncrium Imperfect State. Mycotaxon 1:123-133. Bungay, H. R. 1982, Biomass Refining. Science 218:643-646. Cain, R. B., 1980. The Uptake and Catabolism of Lignin Related Aromatic Compounds and Their Regulation in Microorganisms. p. 21-60, in T. K. 19. 20. 21. 22. 23. 24. 25. 26. 27. 4O Kirk, T. Higuchi, and H.-m. Chang (eds.) Lignin Biodegradation: Microbiology, Chemistry and Potential Applications. Vol. I, CRC Press, West Palm Beach, FL. Cohen, G., 1978. The Generation of Hydroxyl Radicals in Biological Systems: Toxicological Aspects. Photochem. Photobiol. 28:669-675. Crawford, D. L., 1981. Microbial Conversions of Lignin to Useful Chemicals Using a Lignin- Degrading Streptomyggges. Biotechnol. Bioeng. Symp. 11:275—291. Crawford, D. L., 197714 Preparation of Specifically Labeled C-(lignin) and (cellulose) Lignocellulose and Their Decomposition by the Microflora of Soil. Appl. Environ. Microbiol. 33:1247:1251. 14c- Crawford, D. L., M. J. Barden, A. L. Pommetto, and R. L. Crawford, 1982. Chemistry of Softwood Lignin Degradation by Strentcmyces yiridcsncrus Arch. Microbiol. 131: 140- 145. Crawford, D. L., and R. L. Crawford. 1980, Microbial Degradation of Lignin. Enzyme Microb. Technol. 2:11-22. Crawford, D. L., A. L. Pometto III, and R. L. Crawford, 1983. Lignin Degradation by yiridgspgzus: Isolation and Characterization of a New Polymeric Lignin Degradation Intermediates. Appl. Environ. Microbiol. 45:898-904. Crawford, D. L., and B. L. Sutherland, 1979. The Role of Actinomycetes in the Decomposition of Lignocellulose. Dev. Ind. Microbiol. 20:133-191. Crawford, D. L., J. B. Sutherland, A. L. Pometto, and J. M. Miller, 1982. Production of an Aromatic Aldehyde Oxidase by yizidgspgrus. Arch. Microbiol. 131:351-355. Crawford, D. L., and J. B. Sutherland, 1979. Isolation and Characterization of Lignocellulose- Decomposing Actinomycetes. P. 95-102 in T. K. Kirk., T. Higuchi, and H. Chang (Eds.) 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Polyguaiacol: A Useful Model Polymer for Lignin Biodegradation Research. Appl. Environ. Microbiol. 41:1112-1116. Czapski, G., and Y. A. Ilam, 1978. On the Generation of the Hydroxylation Agent From the Superoxide Radical: Can the Haber-Weiss Reaction be the Source of OH radical? Photochem. Photobiol. 28:651-654. Drew, 3. W., K. L. Kadam, S. P. Shoemaker, G. W. Glasser, and P. Hall, 1978. Chemical Feedstock and Fuels From Lignin. A.I.Ch.E. Symp. Series No. 181. 74:21-25. Engelhardt, G., 1979. Degradation of Aromatic Carboxylic Acids by Nggardia sp. DSM. FEMS Microbiol. Lett. 5:245-251. Eriksson, K. E., 1978. A Scanning Electron Microscopy Study of the Growth and Attack by Three White-Rot Fungi and Their Cellulaseless Mutants. Holzforschung. 34:207-213. 37. 38. 39. 40. 41. 42. 43. 44. 45. 46. 42 Eslyn, W. E., T. K. Kirk, and M. J. Effland, 1975. Changes in the Chemical Composition of Wood Caused by Six Soft-Rot Fungi. Phytopathol. 65:473-476. Faison, B., and T. K. Kirk, 1983. Relationship Between Lignin Degradation and Production of Reduced Oxygen Species by Enanergghaete chrysgspgrium. Appl. Environ. Microbiol. 46:1140-1145. Farmer, R. H., 1967. Chemistry in the Utilization of Wood. Pergamon Press, New York, N.Y. Farmer, V. C., M. E. Henderson, and J. D. Russell, 1960. Aromatic Alcohol Oxidase Activity in the Growth Medium Of EQlYfiLiQLBS yersicclcr. Biochem. J. 74: 257- -262. Penn, P., S. Choi, and T. K. Kirk, 1981. Ligninolytic Activity of Phanerggha_te : Physiology of Suppression by NH4+ QhIYSQSDQIiBm and L-Glutamate. Arch. Microbiol. 130: 66- 71. Penn, P., and T. K. Kirk, 1979. Ligninolytic activity of Ehaner9_haete_chryscsncrium: Inhibition by O-Phthalate. Arch. Microbiol. 134:307-309. Fenn, P., and T. K. Kirk, 1980. Relationship of Nitrogen to the Onset and Suppression of Ligninolytic Activity and Secondary Metabolism in Bhanercchaete chrysosngrium Arch Microbiol. 130: 59- 65. Fergus, B. J., A. R. Proctor, J. A. Scott, and D.A.I. Goring, 1969. The Distribution of Lignin in Spruce Wood as Determined by Ultraviolet Spectroscopy. Wood Science Technol. 3:117-138. Forney, L. J., C. A. Reddy, M. Tien, and S. D. Aust, 1982. The Involvement of Hydroxyl Radical Derived From Hydrogen Peroxide in Lignin Degradation by the White-Rot Fungus Enanergchaete ghnysgspgrigm. J. Biol. Chem. 257: 1455- 1462. Forney, L. J., C. A. Reddy, and H. S. Pankratz, 1982. Ultrastructural Localization of Hydrogen Peroxide Production in Ligninolytic Ehanergghaete cells. Appl. Environ. Microbiol. 44:732-736. 47. 48. 49. 50. 51. 52. 53. 54. 55. 56. 43 Forney, L. J., and C. A. Reddy, 1979. Bacterial Degradation of Kraft Lignin. Dev. Ind. Microbiol. 20:163-175. Freudenberg, K., and A. C. Neish, 1968. Constitution and Biosynthesis of Lignin. Springer-Verlag, New York, NY. Freudengerg, K. 1965, Lignin: Its Constitution and Formation From p-Hydroxycinnamyl Alcohols. Science 148:596-6. Frick, T. D., and Crawford, R. L., 1983. Mechanisms of Microbial Demethylation of Lignin Model Polymers, p. 143-152, In T. Higuchi, H. Chang and T. Kirk (Eds.) Recent Advances in Lignin Biodegradation Research, Uni Publ. Co. Ltd. Tokyo. Fridovich, I., 1976. Oxygen Radicals, Hydrogen Peroxide and Oxygen Toxicity, p. 157-206, in W. A. Pryor (Ed.) Free Radicals in Biology. Vol. 1, Academic Press, New York, NY. Gieren, J., and A. E. Opara, 1973. Studies on the Enzymatic Degradation of Lignin. Acta Chem. Scandinavica 27:2909-2922. Gilbertson, R. L., 1981. North American Wood- Rotting Fungi That Cause Brown-Rots. Mycotaxon 12:372-416. Glenn, J. K., and M. H. Gold., 1983. Decolorization of Several Polymeric Dyes by the Lignin-Degrading Basidiomycete Bhanergghaete cnrysgspgzium. Appl. Environ. Microbiol. 45:1741-1747. Glenn, J. K., M. A. Morgan, M. B. Mayfield, M. Kuwahara, and M. H. Gold., 1983. An extracellular HZOZ-Requiring Enzyme Preparation Involved in Biodegradation by the White-Rot Basidiomycete Ehanercchaets_chryscsncrium Proceedings International Symposium on Wood Pulping Chemistry 4:165-168. Gold, M. H., and T. M. Cheng, 1979. Conditions for the Fruit Body Formation in the White Rot Basidiomycete, Bhanerochaete chrysosncrium Arch. Microbiol. 121: 37-41. 57. 58. 59. 60. 61. 62. 63. 64. 65. 44 Gold, M. H., H. Kutsuki, M. A. Morgan, and R. M. Kuhn, 1983. Degradation of Lignin and Lignin Model Compounds by Several Radical Generating Systems and by Ehanergchaete chryscsncriuml Proceedings International Symposium on Wood Pulping Chemistry 4:165-168. Gold, M. H., and T. M. Cheng, 1978. Induction of Colonial Growth and Replica Plating of the White- Rot Basidiomycete Ehanergchaete_chrxscspcrium Appl. Environ. Microbiol. 35: 1223-1225. Gold, M. H., M. B. Mayfield, T. M. Cheng, K. Krisnangkura, M. Shimada, A. Enoki, and J. K. Glenn. 1982. A Phanercchaete Mutant Defective in Lignin Degradation as Well as Several Other Secondary Metabolic Functions. Arch. Microbiol. 132:115-122. Gold, M. H., T. M. Cheng, and M. B. Mayfield, 1982. Isolation and Complementation Studies of Ausotrophic Mutants of the Lignin Degrading Basidiomycete Bhanercchaete chrysosncrium Appl- Environ. Microbiol. 44: 996- 1000. Gold, M. H., T. M. Cheng, and M. Alic, 1983. Formation, Fusion and Regeneration of Protoplasts From Wild-Type and Auxotrophic Strains of the White-Rot Basidiomycete Phanergghaete ghrysgspgrigm. Appl. Environ. Microbiol. 46:260- 263. Gold, M. H., M. Kuwahara, A. A. Chiu, and J. K. Glenn, 1984. Purification and Characterization of an Extracellular H Oz-Requiring Diarylpropane Oxygenase From the Wh1te Rot Basidiomycete Ehanercchaete_chryscsncrium. Arch. Biochem. Biophys. 234:353-362. Green, T. R., 1977. Significant of Glucose Oxidase in Lignin Degradation. Nature. 268:78- 80. Greene, R. V. and J. M. Gould, 1983. Fatty Acyl- Coenzyme A Oxidase Activity and H202 Production in Phan_rcchaete chrxscsncrium. Biochem- Biophys. Res. Commun. 118:437-443. Gross, G. G., 1977. Recent Advances in the Chemistry and Biochemistry of Lignin. Rec. Adv. Phytochem. 12:177-220. 66. 67. 68. 69. 70. 71. 72. 73. 74. 75. 76. 45 Haars, A., and A. Huttermann, 1980. Function of Laccase in the White-Rot Fungus Egnes gnngsns. Arch. Microbiol. 125:233-237. . Haider, K., and J. Trojanowski, 19 0. A Comparison of the Degradation of C-labeled DHP and Cornstalk Lignins by Mirco and Macrofungi and by Bacteria, p. 111-134. In T. K. Kirk, T. Higuchi, and H.-M. Chang (Eds.), Lignin Biodegradation: Microbiology, Chemistry and Potential Applications, Vol. 1. CRC Press, West Palm Beach, Fla. Haider, K. and J. Trojanowski,11975. Decomposition of Specifically C-Labeled Phenols and Dehydropolymers of Coniferyl Alcohol as Models for Lignins Degradation by Soft White-Rot Fungi. Arch. Microbiol. 105:33-41. Haider, K., J. Trojanowski, and V. Sundman, 1978. Scregning for Lignin-Degrading Bacteria by Means of C-Labeled Lignins. Arch. Microbiol. 119:103-106. Haider, K., J. P. Martin,12nd E. Reitz, 1977. Decomposition in Soil of C-Labeled Coumaryl Alcohols: Free and Linked Into Dehydropolymer and Plant Lignins and Model Humic Acids. Soil Sci. Soc. Am. J. 41:556-561. Hall, P., 1980. Enzymatic Transformations of Lignin. Enzyme Microb. Technol. 2:170-176. Harkin, J. M., 1967. Lignin-- A Natural Polymeric Product of Phenol Oxidation, p. 243- 321. In W. I. Taylor, and A. R. Battersby (Eds.) Oxidative Coupling of Phenols, Marcel-Dekker, New York, NY. Hartline, F. F., 1981. Lowering the Cost of Alcohol. Science, 206: 41-42. Healy, J. B., and L. Y. Young, 1979. Anaerobic Biodegradation of Eleven Aromatic Compounds to Methane. Appl. Environ. Microbiol. 38:84-89. Highley, T. L., 1973. Influence of Carbon Source on Cellulase Activity of White-Rot and Brown-Rot Fungi. Wood Fiber 5:50-58. Highley, T. L., 1974. Degradation of Cellulose by 29:1; plngentg in the Presence of Compounds 77. 78. 79. 80. 81. 82. 83. 84. 85. 86. 46 That Affect Hydrogen Peroxide. Material und Organismen. 15:81-90. Higuchi, T., 1980. Biochemistry of Lignin. Wood Res. 66:1-17. Higuchi, T., 1981. Biodegradation of Lignin. Wood Res. 67-47-58. Higuchi, T., 1971. Formation and Biological Degradation of Lignins. Adv. Enzymol. 34:207- 258. Higuchi, T., M. Shimada, F. Nakatsubo, and M. Tanahashi., 1977. Differences in Biosynthesis of Guaiacyl and Syringyl Lignins in Woods. Wood Sci. Technol. 11:153-167. Himmel, M. E., K. K. Oh, D. W. Sopher, and H. L. Chum., 1983. High-Performance Size Exclusion Chromatography of Low-Molecular-Weight Lignins and Model Compounds. J. Chromatography. 267:249-265. Hurst, H. M., and H. A. Burges, 1967. Lignin in Humic Acids, p. 260-286. In A. D. McLaren, and G. Peterson (Eds.), Soil Biochemistry. Marcel- Dekker, New York, NY. Ishihara, T., and A. Nishida, 1983. The Role of Oxidation and Reduction Enzymes in Lignin Biodegradation. P. 134-142. In T. Higuchi, H.- M. Chang, and T. K. Kirk (Eds.). Recent Advances in Lignin Biodegradation Resarch. Uni. Publishers Co., Ltd., Tokyo, Japan. Ishikawa, H., and T. Oki, 1966. The Oxidative Degradation of Lignin. IV. The Enzymatic Hydrolysis Linkages in Lignin. J. Jap. Wood Res. Soc. 12:101-107. Ishikawa, H., W. J. Schubert, and F. F. Nord, 1963. Investigations on Lignins and Lignification. XXVII. The Enzymatic Degradation of Soft-Wood Lignin by White-Rot Fungi. Arch. Biochem. Biphys. 100:131-139. Iwahara, S., 1983. Metabolism of Lignin-Related Aromatic Compounds by Engaginm sp. Proceedings International Symposium on Wood Pulping Chemistry. 3:1-6. 87. 88. 89. 90. 91. 92. 93. 94. 95. 96. 97. 47 Janshekar, H., and A. Fiechter, 1984. Lignin: Biosynthesis, Application and Biodegradation. Adv. Biochem. Eng. Biotechnol. 27:119-178. Jeffries, T. W., S. Choi, and T. K. Kirk, 1981. Nutritional Regulation of Lignin Degradation by Ehanercchaete chrxsosncrium- Appl- Environ- Microbiol. 42:290-296. Johnson, D. B., W. E. Moore, and L. C. Zank, 1961. The Spectrophotometric Determination of Lignin in Small Wood Samples. Tappi 44:793-798. Kaplan, D. L., 1979. Reactivity of Different Oxidases With Lignin and Lignin Model Compounds. Phytochem. 18:1917-1919. Kelley, R. L., and C. A. Reddy, 1982. Ethylene Production From a-oxo-q-Methylthiobutyric Acid is a Sensitive Measure of Ligninolytic Activity by Phanercchaete chryscsncrium Biochem. J. 206: 423- -425. Kersten, P. J., M. Tien, B. Kalyanaraman, and T. K. Kirk, 1985. The Ligninase of Rngngrggngete gnnysgspgrinm Generates Cation Radicals From Methoxybenzenes. J. Biol. Chem. 260:2609-2612. Keyser, T., T. K. Kirk, and J. G. Zeikus, 1978. Ligninolytic Enzyme System of Ennnerg_haete gnzysgspgzinm: Synthesized in the Absence of Lignin in Response to Nitrogen Starvation. J. Bacteriol. 135:790-797. Kirk, T. K., 1984. Degradation of Lignin. P. 399-437. In D. T. Gibson (Ed.) Biochemistry of Microbial Degradation. Marcel-Dekker, New York. Kirk, T. K., 1983. Biotechnology in Utilization of Wood. Biotechnol. 3:666-668. Kirk, T. K., 1981. Toward Elucidation of the Mechanism of the Ligninolytic System Basidiomycetes. P. 131-148. In A. Hollaendar (Ed.) Trends in the Biology of Fermentation for Fuels and Chemicals. Plenum Press, New York. Kirk, T. K., 1975. Effect of a Brown-Rot Fungus, Lenzites trgbea, on Lignin in Spruce Wood. Holzforschung 29: 99-107. 98. 99. 100. 101. 102. 103. 104. 105. 106. 107. 108. 48 Kirk, T. K., 1971. Effects of Microorganisms on Lignin. Ann. Rev. Phytopath. 9:185-212. Kirk, T. K., and E. Adler, 1969. Catechol Moieties in Enzymatically Liberated Lignin. Acta Chem. Scand. 23:705-707. Kirk, T. K., W. Brown, and E. B. Cowling, 1969. Preparative Fractionation of Lignin by Gel- Permeation Chromatography. Biopolymers 7:135- 153. Kirk, T. K., and H. -m. Chang, 1974. Decomposition of Lignin by White-Rot Fungi. 1. Isolation of Degraded Lignins From Decayed Spruce. Holzforschung 29:56-64. Kirk, T. K., and H. M. Chang, 1975. Decomposition of Lignin by White-Rot Fungi. II. Characterization of Heavily Degraded Lignins From Decayed Spruce. Holzforschung 29:56-64. Kirk, T. K., W. J. Connors, R. D. Bleam, W. F. Hackett, and J. G. Zeikus, 1975. Preparation and Microbial Decomposition of Synthetic C-lignins. Proc. Natl. Acad. Sci. USA 72:2515-2519. Kirk, T. K., W. J. Connors, and J. G. Zeikus, 1976. Requirement for a Growth Substrate During Lignin Decomposition Wood-Rotting Fungi. Appl. Environ. Microbiol. 32:192-194. Kirk, T. K., T. Higuchi, and H. Chang (Eds.), 1980. Lignin Biodegradation: Microbiology, Chemistry and Potential Applications. Vol. 1 and 2. CRC Press. West Palm Beach, FL. Kirk, T. K., Nakatsubo, and I. D. Reid, 1983. Further Study Discounts Role for Singlet Oxygen in Fungal Degradation of Lignin Model Compounds. Biochem. Biophys. Res. Commun. 111:200-204. Kirk, T. K., and W. E. Moore, 1972. Removing Lignin From Wood With White-Rot Fungi and the Digestibility of the Resulting Wood. Wood Fiber. 4:72-79. Kirk, T. K., E. Schultz, W. J. Connors, L. F. Lorenz, and J. G. Zeikus, 1978. Influence of Culture Parameters on Lignin Metabolism by Ehanercchaete chryscsncrium. Arch. Microbiol. 117:277-285. 109. 110. 111. 112. 113. 114. 115. 116. 117. 118. 119. 49 Koenigs, J. W., 1975. Hydrogen Peroxide and Iron: A Microbial Cellulolytic System? Biotechnol. Bioeng. Symp. No. 5:151-159. Koenigs, J. W., 1972. Production of Extracellular Hydrogen Peroxide and Peroxidase by Wood-Rotting Fungi. Phytopahtology 62:100-110. Koenigs, J. W., 1974. Production of Hydrogen Peroxide by Wood-Rotting Fungi in Wood Correlation With Weight Loss, Depolymerization and pH. Arch. Mikrobiol. 99:129-145. Kutsuki, H., and M. H. Gold, 1982. Generation of Hydroxyl Radical and Its Involvement in Lignin Degradation by Phanercchaete chrxscsncrium. Biochem. Biophys. Res. Commun. 109:320-327. Kuwahara, M., J. K. Glenn, M. A. Morgan and M. H. Gold, 1984. Separation and Characterization of Two Extracellular H202 Dependent Oxidases From Ligninolytic Cultures of znnnerggnnete chzysgspgrinim. FEBS Letters 169: 247-250. Leisola, M. S., D. Ulmer, and A. Fiechter, 1984. Factors Affecting Lignin Degradation in Lignocellulose by Ehanercchaete chrysosncrium. Arch. Microbiol. 137:171-175. Levi, M. P., and R. D. Preston, 1965. A Chemical and Microscopic Examination of the Action of the Soft-Rot Fungus gnnetgninm gIngsnm on Beechwood (Eagus 52121). Holzforschung 19: 183- 190. Liese, W., 1970. Ultrastructural Aspects of Woody Tissue Disintegration. Annu. Rev. Phytopathol. 8:231-258. Ludeman, H. D, 1973. Carbon-13 Nuclear Magnetic Resonance Spectra of Lignins. Biochem. Biophys. Res. Commun. 52:1162-1169. Lundquist, K., 1979. NMR Studies of Lignin. Acta. Chem. Scan. B33:27-30. Lundquist, K., 1978. De Novo Synthesis and Decomposition of Veratryl Alcohol by a Lignin- Degrading Basiciomycete. Phytochemistry 17:1676. 120. 121. 122. 123. 124. 125. 126. 127. 128. 129. 50 MacDonald, M. J., A. Paterson, and P. Broda, 1984. Possible Relationship Between Cyclic AMP and Idiophasic Metabolism in the White-Rot Rungus Ehanerochaete chryscsncrium. J. Bacteriol- 160:470-472. Martin, J., P. K. Haider, and G. Kassim, 1980. Biodegradation and Stabilization After 2 Years of Specific Crop, Lignin and Polysaccharide Carbons in Soils. Soil Sci. Soc. Amer. J. 44:1250:1255. Nakatsubo, F., 1.19. Reid, and T. K. Kirk, 1982. Incorporation of 02 and Absence of Sterospecificity in Primary Product Formation During Fungal Metabolism of a Lignin Model Compound. Biochim. Biophys. Acta 719:284-291. Nakatsubo, F., I. D. Reid, and T. K. Kirk, 1981. Involvement of Singlet Oxygen in the Fungal Degradation of Lignin. Biochem. Biophys. Res. Commun. 102:484-491. Nimz, H, 1974. Beech Lignin -- Proposal of a Constitutional Scheme. Angewandte Chemie. 86:336- 344. Noguchi, A., M. Shimada, and T. Higuchi, 1980. Studies on Lignin Biodegradation. I. Possible Role of Nonspecific Oxidation of Lignin by Laccase. Holzforshcung 34:86-89. Odier, E., G. Janin, and B. Monties, 1981. Poplar Lignin Decomposition by Gram-Negative Aerobic Bacteria. Appl. Environ. Microbiol. 41:337-341. Odier, E., and B. Monties, 1983. Absence of Microbial Mineralization of Lignin in Anaerobic Enrichment Cultures. Appl. Environ. Microbiol. 46:661-665. Odier, E., and B. Monties, 1978. Biodegradation of Wheat Lignin by Knntngmgnns 23. Ann. Microbiol. (Paris) 129A:361-377. Odier, E., and P. Roch, 1983. Factors Controlling Biodegradation of Lignin in Wood by Various White-Rot Fungi, p. 188-194, In T. Higuchi. H-m. Chang, and T., K. Kirk (Eds.). Recent Advances in Lignin Biodegradation Research, UNI Pub., Tokyo. 130. 131. 132. 133. 134. 135. 136. 137. 138. 139. 51 Otjen, L., and R. A. Blanchette, 1984. Xylgpglns fiznsgnlgns Decay of Oak: Patterns of Selective Delignification and Subsequent Cellulose Removal. Appl. Environ. MIcrobiol. 47:670-676. Palmer, J. M., and C. S. Evans, 1983. Extracellular Enzymes Produced by Qgriglns yezsigglgz in Relation to the Degradation of Lignin. Proceedings International Symposium on Wood Pulping Chemistry, 3:19-24. Pearl, I. A., 1967. The Chemistry of Lignin, Marcel-Dekker, Inc., New York. Rao, T. R., and C. A. Reddy, 1984. DNA Sequences From Ligninolytic Filamentous Fungus Bhanerochaete chrysosnorium capable of Autonomous Replication in Yeast. Biochem. Biophys. Res. Commun. 118:821-827. Reddy, C. A., and L. J. Forney, 1978. Lignin Chemistry and Structure: A Brief Review. Dev. Ind. Microbiol. 19:27-34. Reddy, C. A., 1978. Introduction to Microbial Degradation of Lignin. Dev. Ind. Microbiol. 19:23-26. Reddy, C. A., 1984. Physiology and Biochemistry of Lignin Degradation, p. 558-571. In M. J. Klug and C. A. Reddy (Eds.) Current Perspectives in Microbial Ecology. Amer. Soc. Microbiol. Washington, D.C. Reddy, C. A., L. J. Forney, and R. L. Kelley, 1983. Involvement of Hydrogen Peroxide-Derived Hydroxyl Radical in Lignin Degradation by the White-Rot Fungus Ehanerochaete chrysosnorium- P. 153-163. In T. Higuchi, H.-m Chang, and T. K. Kirk (Eds.), Recent Advances in Biodegradation Research, Uni Publishers Co. LTD., Tokyo, Japan. Reid, I. D., 1983. Effects of Nitrogen Sources on Cellulose and Synthetic Lignin Degradation by Ehanerochaete - Appl Environ- Microbiol. 45:838-842. Reid, 1. D., 1983. Effects of Nitrogen Supplements on Degradation of Aspen Wood Lignin and Carbohydrate Components by Enanepggngege gnrysgspgzinm. Appl. Environ. Microbiol. 45:830- 837. 140. 141. 142. 143. 144. 145. 146. 147. 148. 149. 52 Reid, I. D., 1979. The Influence of Nutrient Balance on Lignin Degradation by the White-Rot Fungus Ehanercchaete chrysosnorium. Can. J. Bot- 57:2050-2058. Reid, I. D., E. E. Chao, and P. Dawson, 1984. Lignin Degradation by Ehanerochaete in Agitated Cultures. Can. J. Microbiol. 31:88- 90. Reid, I. D., and K. A. Seifert, 1982. Effect of an Atmosphere of Oxygen on Growth, Respiration, and Degradation by White-Rot Fungi. Can. J. Bot. 60:252-260. Reid, I. D., and K. A. Seifert,1980. Lignin Degradation by Ehanerochaete chrysosnorium in Hyperbaric Oxygen. Can. J. Microbiol. 26: 1168- 1171. Robinson, L. E., and R. L. Crawford, 1978. Degradation of 14-C Labelled Lignins by Bagillns megggeninm. FEMS Microbiol. Lett. 4:301-302. Sarkanen, K. V., and Ludwig, C. H. (Eds.), 1971. Lignins: Occurrence, Formation, Structure and Reactions. Wiley-Interscience, New York. Sawyer, D. T., and J. S. Valentine, 1981. How Super is Superoxide? Acc. Chem. Res. 14:393-400. Schubert, W. J., 1965. Lignin Biochemistry. Academic Press, New York. Shimada, M., F. Nakatatsubo, T. K. Kirk, and T. Higuchi, 1981. Biosynthesis of the Secondary Metabolite Veratryl Alcohol in Relation to Lignin Degradation in Ehanerochaete chrysosnorium Arch. Microbiol. 129: 321- 324. Shimada, M., 1980. Stereobiochemical Approach to Lignin Biodegradation: Possible Significance of Nonstereospecific Oxidation Catalyzed by Laccase for Decomposition by White-Rot Fungi. In T. K. Kirk, T. Higuchi, and H. Chang (Eds.). Lignin Biodegradation: Microbiology, Chemistry and Potential Applications. CRC Press. West Palm Beach, FL. 150. 151. 152. 154. 155. 156. 157. 158. 159. 160. 161. 53 Schoemaker, H. E., P. J. Harvey, R. M. Bowen and J. M. Palmer, 1985. On the Mechanism of Enzymatic Lignin Breakdown. FEBS Lett. 183:7-12. Sutherland, J. B., 1979. Breakdown of Douglas- Fir Phloem by a Large Cellulose-Degrading Streptomyges. Current Microbiol. 2:123-126. Tien, M., and T. K. Kirk, 1983. Lignin-Degrading Enzyme From the Hymenomycete Phangrgghagte ghrysgspgrium: Purification, Characterization, and Catalytic Properties of a Unique H202- Requiring Oxygenase. Proc. Natl. Acad. Sci. USA 81:2280-2284. , Trojanowski, J. K14Haider, and V. Sundman, 1977. Decomposition of C-Labeled Lignin and Phenols by a Nggardia. Arch. Microbiol. 114:149-153. Ulmer, D. C., M. Leisola, B. Schmidt, and A. Fiechter, 1983. Rapid Degradation of Isolated Lignins by Ehanergghaete ghryaoanorium- Appl- Environ. Microbiol. 45:1795-1801. Ulmer, D. C., M. Leisola, and A. Fiechter, 1984. Possible Induction of the Ligninolytic System of Ehanergghaete enzyaosnorium J Biotechnol 1: 13- 24. Vance, C. P., T. K. Kirk, and R. T. Sherwood, 1980. Lignification as a Mechanism of Disease Resistance. Ann. Rev. Phytopathol. 18:259-280. Walling, C., 1975. Fenton's Reagent Revisited. Acc. Chem. Res. 8:125-131. Wallace, L., A. Paterson, A. McCarthy, U. Raeder, L. Ramsey, M. MacDonald R., and P. Broda, 1984. The Problem of Lignin Biodegradation. Biochem. Soc. Symp. 48:87-95. Weinstein, D. A., K. Krisnangkura, M. B. Mayfield, and M. H. Gold, 1980. Metabolism of Radiolabeled B-Guaiacyl Ether-Linked Lignin Dimeric Compounds by Enanerognaete ghrysosnorium. Appl. Environ. Microbiol. 39:535-540. Westermark, U., and K. E. Eriksson, 1974. Cellobiose-Quinone Oxidoreductase, A New Wood- Degrading Enzyme by White-Rot Fungi. Acta. Chem. Scand. 828:209-214. 162. 163. 164. 165. 166. 54 Woodwell, G. M., 1978. The Carbon Dioxide Question. Scientific American 238(1):34-44. Yang, H. H., M. J. Effland, and T. K. Kirk, 1980. Factors Influencing Fungal Degradation of Lignin in a Representative Lignocellulosic, Thermomechanical Pulp. Biotechnol. Bioeng. 22:65-77. Young, Ly., 1984. Anaerobic Degradation of Aromatic Compounds, pp. 487-523. In D. T. Gibson (Ed.) Microbial Degradation of Organic Compounds. Marcel-Dekker, Inc., New York. Zeikus, I. G., 1983. Lignin Metabolism and the Carbon Cycle: Polymer Biosynthesis, Biodegradation and Environmental Recalcitrance, p. 211-243. In M. Alexander (Ed.), Advances in Microbial Ecology, Vol. 5, Plenum Press, NY. Zeikus, J. G., A. L. Wellstein, and T. K. Kirk, 1982. Molecular Basis for the Biodegradative Recalcitrance of Lignin in Anaerobic Environments, FEBS Lett. 15:193-197. 21WP/RPP/glucose 55 Chapter one: Identification of Glucose Oxidase Activity as the Primary Source of Hydrogen Peroxide Production in Ligninolytic Cultures.of Phanerochaete chrysosporium 56 Abstract The primary enzymatic activity involved in H202 production by extracts of ligninolytic cultures of Phanerochaete chrysosporium was investigated. Glucose supported the highest level of oxygen-dependent H202 production by cell extracts compared to a number of other substrates tested. No H202 production was observed anaerobically under N2. Polyacrylamide gel electrophoresis of extracts from ligninolytic cultures followed by diaminobenzidine/horseradish perOXidase staining procedUre showed that only one protein band exhibited glucose- dependent H202 production. This protein band was not seen in extracts of non-ligninolytic cultures or in the extracellular fluid of ligninolytic cultures. Extracts of cells grown with either xylose, succinate or cellobiose showed the presence of only one glucose-dependent 8202- producing band, with electrophoretic mobility similar to that observed in extracts of glucose-grown cells. Both iglucoseoxidase activity and lignin degradation were triggered in response to nitrogen (N) or carbohydrate starvation, and were repressed in media containing high levels of N (24 mM) or carbohydrate (56 mM), or an addition of exogenous N sources such as glutamate to the low N medium (2.4 mM N). The results indicate that glucose oxidase activity is the primary source of 3202 in ligninolytic cultures of 2, chrysosporium and that nutritional parameters which affect lignin degradation have a parallel effect on glucose oxidase activity. 57 Keywords: Phanerochaete chrysosporium--lignin degradation--g1ucose oxidase--nutritional parameters-- hydrogen peroxide production--white-rot basidiomycete. Lignin is one of the most abundant biopolymers in nature constituting approximately 20-30Z of the dry weight of all vascular plants (Crawford 1981, Kirk 1984, Reddy 1984). Hence, lignin biodegradation and its utilization as a renewable resource for the production of chemical feedstocks and other useful products is of great interest. Previous studies have shown that lignin biodegradation is an oxidative process (Kirk and Chang 1974) and that H202 plays an important role in this degradation (Crawford and Crawford 1984, Reddy 1984, Reddy et a1. 1983; Patterson and Lundquist 1984). -A temporal correlation between ligninolytic activity and H202 production by 2. chrysosporium has been demonstrated (Forney et al. 1982a,b). (Growth under 100% 02 markedly increased both H202 production and ligninolytic activity, and lignin degradation was inhibited by catalase, a scavenger of H202 (Faison and Kirk 1983). Cytochemical diaminobenzidine (DAB) staining techniques revealed that H202 production in ligninolytic cultures of P, Chrysosporium, is localized in ovoid periplasmic microbody-like structures which are not found in non-ligninolytic cells (Forney et al. 1982b). Q Several HZOZ-dependent, lignin degrading enzymes, have been 58 isolated from the extracellular fluid of P, chrysosporium cultures. These enzymes were only found in ligninolytic cultures and have been shown to oxidize a variety of lignin model compounds (Tien and Kirk 1983, Tien and Kirk 1984, Glenn et al 1983, Gold et a1. 1984), depolymerize lignin (Tien and Kirk 1984) and decolorize the polymeric dye poly R-481 (Gold et al. 1984; Kuwahara et a1. 1984). Also, an HZOZ-dependent lignin demethylase has recently been described by Frick and Crawford (1983). These results indicated that H202 plays an important role in lignin. degradation by P, chrysosporium; however, the physiological source of H202 in ligninolytic cultures of this organism was not known. The results presented here show that glucose oxidase (E.C. 1.1.3.4) is the primary source of H202 in ligninolytic cultures of 2. chrysosporium. METHODS AND MATERIALS Organism, culture conditions and media. ‘2. Chrysosporium (ATCC 34541) was maintained and conidial inoculum was prepared and used as previously described (Kirk et a1. 1978). The composition of the low and high N media as well as the low and high carbohydrate media was previously described (Kelley and Reddy 1982). Inoculated media in foam-stoppered flasks were covered with plastic bags to minimize evaporation and were incubated at 37°C without agitation. Assay for ligginglytic activity. Ligninolytic activity was assayed as previously described (Forney et al. 1982s) by measuring the evolution of 14C02 from 2'-(14C)- 59 synthetic lignin (30,000 dpm/flask, unless otherwise mentioned). Assay for £292. Three 50 ml cultures were harvested by centrifugation and washed 3X with 50 ml Na 2fl2'- dimethylsuccinate buffer (DMS, pH 4.5). 'The washed mycelial pellet was resuspended in 10 m1 DMS buffer, mixed with 0.1 mm size glass beads in a 1:1 ratio (mycelial wet weight:glass beads), blended for 3 min at 4°C using an Omni-mixer (Sorval, Inc., Newton, CT) and centrifuged at 27,000 x g for 15 min at 4°C. The supernatant cell extract was used for the assay. H202 production was assayed spectrophotometrically by measuring the peroxidative oxidation of o-dianisidine (Decker 1977). The reaction mixture consisted of 1.0 ml of oxygen-saturated DMS buffer (10 mM, pH 4.5), 1.0 m1 of 0.31 mM o-dianisidine, 0.3 m1 of a 1.0 M solution of D-glucose or other substrates (0.1 M final concn.),CL1 ml peroxidase (60 U/ml: EJL 1.11Ju7; Sigma, St. Louis, MO), and 0.5 ml cell extract. After 5 min of incubation at 37°C, the change in absorbance at 460 nm, caused by peroxidative oxidation of o-diansidine was measured using a CARY 219 spectrophotometer (Varian Associated, Palo Alto, CA, USA) and the units of glucose oxidase activity were calculated as previously described (Decker 1977). In one experiment, the extracellular fluid from 6-day-old ligninolytic cultures of P. chrysosporium, (Tien and Kirk 1984), concentrated 20-fold by an Amicon ultrafiltration unit equipped with a PM-lO filter (Amicon 60 Co., Lexington, MA, USA), was used to assay for H202 production, to determine if glucose oxidase activity is extracellular or not. Electrophoresis 3; extracts. Polyacrylamide gel electrophoresis of cell extracts was performed on an 82 native gel with a 5% stacking gel as previously described (Melachouris 1968). Samples (50 ul) were placed on a 1.5 mm slab gel (16 x18 cm) or a 0.5 cm tube gel, and electrophoresed at constant current (30mA for slab gel and 2.5 mA/tube for tube gels) for 3.5 h at 4°C using a Buchler Model 3-1014A power supply (Buchler Instruments, NJ). The gels were stained for HZOZ-generating enzymes with a diaminobenzidine (DAB)/horseradish peroxidase (HRP) system (Cohen 1972). In certain experiments, parallel gels were stained for protein by the silver staining method of Morrissey (1981L Chemical assays. Protein content of the extracts was determined by the procedure of Lowry et a1. (1951) using bovine serum albumin (IV, Sigma Chemical Co., St. Louis, MO, USA) as the standard. Reducing sugar content in the crude cell extract was determined using the dinitrosalicylic acid procedure (Miller 1957) and the extent of glucose contamination in various assay substrates was determined using the procedure described by Raabo and Terkildsen (1960). RESULTS Substrate specificity. To identify the enzimatic activity responsible for hydrogen peroxide (H202) 61 ‘production in ligninolytic cultures of E, chrysosporium, we first investigated the ability of different substrates to support H202 production by cell extracts of 6-day-old cultures grown in low N medium with glucose as the growth substrate (Table 1). The highest specific activity for H202 production was observed with D-glucose as the substrate. L-sorbose, D-cellobiose, D-mannose, and D- maltose supported lower levels of H202 production as compared to glucose. Glucose-dependent H202 production was not observed when the reaction mixture was made anaerobic by sparging it with 02-free N2 for 10 min (Table 18). H202 production activity reappeared and was comparable to that observed before (Table 1A) when the reaction mixture was replaced with air. The above results show that glucose is the primary substrate for H202 production and that oxygen is required for H202 production in extracts of ligninolytic cultures of "E. CDFySOSQOriUm.' These results further suggest that either glucose oxidase (Decker 1977) and/or a carbohydrate oxidase (Janssen and Ruelius 1968) may be involved in H202 production in these extracts. Gel electrophoresis and identification 3; £292; producing activity. To determine if one or more enzymes were involved in H202 production, extracts of glucose-grown ligninolytic cultures were electrophoresed on a native polyacrylamide gel, and the protein bands positive for H202 62 generating activity were visualized by the diaminobenzidine (DAB)/horseradish peroxidase (HRP) staining procedure using D-glucose as the substrate (Fig. 1, gel 1). A single, intensely stained band of activity was observed. When parallel gels were stained separately for H202 generating activity with different substrates, L-sorbose, D-xylose, D- cellobiose, D-maltose and D-mannose were the only other substrates with which an HZOZ-positive protein band was observed (Fig. 1, gels 2-6). The protein band observed with each of these substrates had the same electrophoretic mobility as that of the glucose-dependent activity band (gel 1). The results presented in Table 1 and Fig. 1 together suggest that a single enzyme is primarily involved in H202 production in ligninolytic cells and that the enzyme either has a broad substrate specificity and/or some of the substrates have glucose contamination. Further studies showed no detectable glucose contamination in sorbose, but a low level of glucose contamination (0.8-1Ji mg/g) in xylose, cellobiose, maltose and mannose. These low levels of glucose contamination could explain the reactivity of the protein band in gels 2 to 6 (Fig. 1). These results support the view that glucose-dependent H202 production mediated by glucose oxidase is the primary source of H202 in ligninolytic cells of g. chrysosporium. Demonstration of a novel, extracellular, H202- dependent mono-oxygenases in ligninolytic cultures of 2. chrysosporium is an exciting recent discovery (Tien and Kirk 1984). Hence, we investigated the possibility that 63 glucose-dependent, HZOZ-producing enzymatic activity occurs extracellularly in lignin-degrading cultures. Also, previous demonstration of an extracellular glucose oxidase in cultures of Penicillium purpurogenum was consistent with this view (Nakamatsu 1975). Our results showed no detectable glucose-dependent H202-producing protein band in 20: concentrated extracellular culture fluid from 6-day- old, glucose-grown ligninolytic cultures (Fig. 2A, lane 2), whereas a distinct DAB—positive band was seen in cell extracts of the same culture (Fig. 2A, lane 1). No glucose oxidase activity was detectable in the concentrated extracellular culture fluid even by the spectrophotometric assay; however, ligninase activity was readily detectable in the same preparation using the assay procedure of Tien and Kirk (1984) suggesting that lack of sufficient protein is not the problem. Both nitrogen and carbohydrate concentration in the medium are known to have a profound effect on lignin degradation by 21 chyrsosporium. Cultures of this fungus grown in low N (2.4 mM) medium exhibit high levels of both ligninolytic activity and H202 production, whereas cultures grown under identical conditions in high N (24 mM) medium show little of these activities (Kirk et a1. 1978; Forney et al. 1982b; Kutsuki and Gold 1982; Greene and Gould 1983). Consistent with these earlier studies, the DAB- positive protein band (lane 1) was also not seen in extracts of non-ligninolytic cultures grown for 4 days in 64 low N medium (Fig 2A, lane 3) or for 14 days in high N medium (Fig. 2A, lane 4). In a parallel gel stained for protein (Fig. 28), a band corresponding to the glucose- dependent, HZOZ-generating band (Fig 2A, lane 1) was seen only in extracts of non-ligninolytic cells (Fig. 28, lanes 3 & 4) indicating that the HZOZ-producing activity is attributable to a specific protein band (Fig. 28, lane 1). These results indicate that the glucose oxidase activity is correlated with lignin degradation. Effect g£_different co-substrates. It has been known for some time that lignin does not serve as a sole source of carbon/energy for g; chrysosporium and that a co- substrate, such as glucose, xylose or succinate, is required for growth and lignin degradation (Kirk et a1. 1976, 1978). Each co-substrate is known to support different levels of lignin degradation and it is possible that each of the co-substrates induces different H202- producing oxidase. To test this possibility, glucose, xylose or succinate were added to parallel cultures to give equi-molar carbon concentrations, and were assayed for ligninolytic activity, and H202 production (Table 2). Lignin degradation with different co-substrates decreased in the order: D-glucose > D-xylose > succinate. Hydrogen peroxide production by cell extracts of cultures grown with each of the co-substrates was then determined using the respective growth substrate (D-glucose, D-xylose and succinate) or glucose in the assay (Table 2). Hydrogen peroxide producing activity was maximal when extracts of 65 glucose-grown cells were assayed with glucose as the substrate. In contrast, little or no 8202 production was seen when extracts of cells grown with succinate or xylose were assayed, respectively, with succinate or xylose as the substrate. It was of interest that even the extracts of cells grown with xylose or succinate as the growth substrate displayed high levels of specific activity for H202 production when glucose was employed as the assay substrate. These results suggest that regardless of the co-substrate used, glucose oxidase was the predominant activity generating H202. To further demonstrate that glucose oxidase is the primary source of H202 production in ligninolytic cultures of this organism, cell extracts from cultures grown with either D-glucose, D-xylose, succinate or D-cellobiose, as the co-substrate (growth substrate) were subjected to polyacrylamide gel electrophoresis. Each gel was then stained for HZOZ-generating activity, with D-glucose as the substrate. A single H202-generating protein band with a slightly different electrophoretic mobility was seen in each case (Fig. 3A; see discussion below). When parallel gels were stained for HZOZ-generating activity with D- xylose, D-cellobiose or succinate as the substrate, instead of D-glucose, no additional band of HZOZ-producing activity was seen (data not shown). Parallel gels stained for protein showed one protein band each corresponding to the respective H202-producing band (Fig.38). It is not clear ' 66 whether the minor differences in mobility of the protein bands in lanes 1 to 4 (Fig. 3A) represent slight differences in physical characteristics of the protein in the four extracts or are a reflection of unknown differences in the four cell extracts which are affecting the mobility of the H202-producing protein. The answer to this question should await purification of the H202- producing enzyme from cultures grown on each of the co- substrates. Regardless, glucose oxidase appears to be the predominant HZOZ-producing activity in ligninolytic cultures of P; chrysosporium, irrespective of the growth substrate used. Effect pf various culture parameters pg glucose oxidase activity. Culture parameters such as nitrogen and carbohydrate concentrations are known to influence ligninolytic activity of 2, chrysosporium (Keyser et a1. 1978; Kirk et al., 1978; Penn and Kirk 1980; Fenn et a1. 1981). Addition of either (N84)2804, L-glutamate, or certain other aminoacids to nitrogen-straved lignin- degrading cultures of Z; chgysospprium suppressed ligninolytic activity. Given the above evidence that glucose oxidase activity is the primary source of H202 in ligninolytic cultures of 2, chrysosporium, we studied the effect of nitrogen concentration on the specific activity of this enzyme. The results of this study confirm and extend the results of our preliminary studies (Reddy et a1. 1983) in demonstrating an inverse correlation between the nitrogen concentration in the medium and the specific 67 activity for glucose oxidase as well as ligninolytic activity in cultures of g, chrysosporium (Table 3A). Furthermore, the addition of exogenous N source, such as (NH4)2804 or L-glutamate resulted in parallel suppression of both glucose oxidase and lignin degradation activities (Table 38). Jeffries et a1. (1981) reported that lignin degradation is repressed in a high N medium (24 mM N) containing high levels (56 mM) of glucose, but not in an identical medium containing low levels (8.8 mM) of glucose. The results of this study show that glucose oxidase activity, akin to ligninolytic activity, was higher in the low carbohydrate medium compared to that observed in the high carbohydrate medium (Table 3C). Even so, glucose oxidase activity in low carbohydrate medium was considerably lower than that seen in the low nitrogen medium. This may perhaps reflect the fact that cell yield is extremely low in low carbohydrate medium and apparently ' glucose oxidase acc0unts fOr only a small portion of the total cellular protein under these conditions. Discussion Recent studies show that 8202 plays an integral role in lignin degradation (Forney et al., 1982a, Faison and Kirk 1983, Crawford and Crawford 1984). The results of the present study indicate that the primary source of H202 13 glucose oxidase. We have shown that extracts of glucose- grown, ligninolytic cultures of E, chrysosporium can 68 produce H202 by oxidizing D-glucose in the presence of 02. Polyacrylamide gel electrophoresis of the cell extract and visualization of HZOZ-producing protein band using DAB/HRP staining procedure revealed the presence of a single, glucose-dependent band of activity. This protein band was seen only in extracts of ligninolytic cultures. Substrates which supported low levels of H202 production by this protein include sorbose, maltose, mannose, xylose and cellobiose. Although we have shown that the latter four substrates were contaminated with glucose, recent data with.° the purified enzyme have shown that glucose oxidase from E. chrysosporium does exhibit low levels of activity with L- sorbose, D-xylose, and D-maltose. The results of this study show D-glucose supported the higher levels of H202 production and ligninolytic activity than the other co-substrates tested (Table 2). Xylose- grown and succinate-grown cultures degraded less lignin, and cell extracts of these cultures produced significant amounts of H202 only when glucose was supplied. When cell extracts from glucose-, xylose-, succinate- or cellobiose- grown cultures were electrophoresed and stained for 8202- generating activity, a single band of glucose-dependent activity was seen (Fig. 3). However, the electrophoretic mobility of the band in individual extracts was slightly different; the band in extracts of succinate grown cells showed the more pronounced change. Whether these differences in mobilities represent a significant difference in physical characteristics of the protein in a 69 given extract or a reflection of differences in the cell extracts which are affecting the protein mobility is not known. The answer to this question should await purification of the enzyme from cultures grown on each of the co-substrates. Regardless, glucose oxidase appears to be the predominant HZOZ—producing activity in ligninolytic cultures of g, chrysosporium, irrespective of the growth substrate used. Both nitrogen and carbohydrate concentration in the. medium are known to have a profound effect on lignin degradation by g. chrysosporium. Cultures of this fungus grown in low N (2.4 mM) medium exhibit high levels of not only ligninolytic activity but also 8202 and hydroxyl radical («”0 production, whereas cultures grown under identical conditions in high N (24 mM) medium show little or none of these activities (Kirk et a1. 1978, Forney et al. 1982, Kelley and Reddy 1982, Kutsuki and Gold 1982, Greene and Gold 1983). Consistent with these earlier studies, the results presented here show that nitrogen starvation also triggers glucose oxidase activity. 'Jeffries et al. (1981) showed that ligninolytic activity is triggered by carbohydrate-starvation even in the presence of high levels of N (24 mM). Our results show that glucose oxidase activity is detectable in carbohydrate-starved ’cultures, albeit at much lower levels compared to that seen in cultures grown in low N medium which contains non- limiting amounts of glucose. This is in agreement with the 70 previous finding that the extent of lignin degradation in low N medium depends on the amount of carbohydrate supplied (Jeffries et al., 1981). Fenn et a1. (1981, 1980) showed that the addition of either (NH4)2804, L-glutamate or certain other amino acids to nitrogen-starved ligninolytic cultures of'g. chrysosporium suppressed ligninolytic activity. Similarly, we found that the addition of exogenous N sources suppresses not only ligninolytic activity, but also glucose oxidase activity.~ Consistent with previous resultS,‘ . . glutamate was a more effective suppressor of both these activities than (NH4)2S04. Demonstration of glucose oxidase activity in g. chrysospbrium is consistent with the fact that in nature glucose derived from cellulose hydrolysis is beleived to be the major sugar substrate available to wood-degrading fungi. Green (1977) demonstrated high levels of glucose oxidase in crude extracts of another white-rot fungus, Polyporus versicolor and postulated that glucose oxidase in concert with laccase may play a role in lignin degradation: however, conclusive evidence in support of this hypothesis is lacking. Our results are also consistent with previous reports documenting the wide distribution of glucose oxidase in other fungi (Gancedo et al. 1967). Our results show that glucose oxidase plays an important role in H202 production in ligninolytic cells of z, chrysosporium. However, H202-generating oxidases for a variety of other substrates are known in eukaryotes 71 (Tolbert 1981) and contribution of one or more of these enzymes to 8202 production in.§§ chrysosporium cannot be ruled out. Greene and Gould (1984) reported fatty acyl-CoA oxidase activity in 2, chrysosporium which catalyzed oxygen-dependent H202 production, by starved mycelia of 2. chrysosporium. However, the HZOZ-producing activity observed by these authors was about three orders of magnitude less than the glucose-dependent H202 producing activity observed 97,95 in this study.‘ Furthermore, we did not detect fatty acyl CoA-dependent H202 production with the cell extracts. Therefore, as Greene and Gould (1984) pointed out, it is difficult to determine the role of fatty acyl CoA oxidase in lignin degradation by E. chrysosporium. In conclusion, glucose oxidase activity appears to be the primary physiological source of H202 in cell extracts of ligninolytic cultures of 2, chrysosporium. Polyacrylamide gel electrophoresis data show that a single protein band, found only in extracts of ligninolytic cultures and not in the extracellular fluid is responsible for this activity. Glucose oxidase activity, similar to ligninolytic activity, appears to be a secondary metabolic event and is triggered in response to nitrogen or carbohydrate starvation. Glucose oxidase from 2, chrysosporium has recently'been purified to homogeneity (Kelley RL and Reddy CA, unpublished data). In support of the results of this study, the purified enzyme appears to have a relatively low substrate specificity compared to that from other fungi. 72 It gave optimal activity with glucose, but exhibited only low levels of activity with L-sorbose, D-xylose and D- maltose. A recent study from our laboratory (Ramasamy et al., 1985) also showed that glucose oxidase-negative mutants of £5 chrysosporium are deficient in both glucose oxidase activity and lignin degradation, whereas both these activities reappeared in glucose oxidase positive revertants. These results provide additional support to the view that glucose oxidase activity plays an important role in lignin degradation by E} chrysosporium. 73 References Cohen HJ (1972) The use of diaminobenzidine for spectrophotometric and acrylamide gel detection of sulfite oxidase and its applicability to hydrogen generating enzymes. Anal Biochem 53:208-222 Crawford RL (1981) Lignin Biodegradation and Transformation. John Wiley and Sons, New York Crawford RL, Crawford DL (1984) Recent advances in studies of the mechanisms of microbial degradation of lignin. Enzyme Microb Technol 6:434-442 Decker LA (1977) Worthington Enzyme Manual. Worthington ? Biochemical Corp., Freehold, NH, pp. 37-39. Faison 8, Kirk TK (1983) Relationship between lignin a degradation and production of reduced oxygen species by- Phanerochaete chrysosporium. Appl Environ Microbiol 46:1140-1145 Fenn P, Choi S, Kirk TK (1981) Ligninolytic activity of Phanerochaete chrysosporium: physiology of suppression by NHAT and L-glutamate. Arch Michrobiol 130:66-71 Fenn P, Kirk TK (1980) Relationship of nitrogen to the onset and suppression of ligninolytic activity and secondary metabolism in Phanerochaete chrysosporium. Arch Microbiol 130:59-65 Forney LJ, Reddy CA, Tien M, Aust SD (1982a) The involvement of hydroxyl radical derived from hydrogen peroxide in lignin degradation by the white-rot fungus Phanerochaete chrysosporium. J Biol Chem 257:1455-1462 Forney-LJ, Reddy CA, Pankratz HS (1982b) Ultrastructural localization of hydrogen peroxide production in ligninolytic Phanerochaete chrysosporium cells. Appl Environ Microbiol 44: 732- 736 Frick TD, Crawford RL (1983) Mechanisms of microbial demethylation of lignin model polymers. In: Higuchi T, Chang HM, and Kirk TK (eds) Lignin biodegradation research, Uni Publishers, Tokyo pp. 143-152. Gancedo J, Gancedo M, Asensio C (1967) Widespread occurrence of galactose oxidase and glucose oxidase. Arch Biochem 119:588-590 Glenn JK, Morgan MA, Mayfield M8, Kuwahara M, Gold MH (1983) An extracellular H2 0 -requiring enzyme preparation involved in lignin biodegra ation by the white- rot basidiomycete Phanerochaete chrysosporium. Biochem Biophys Res Commun 114:1077-1083 74 Gold MH, Kuwahara M, Chiu AA, Glenn JK (1984) Purification and characterization of an extracellular H 0 -requiring diarylpropane oxygenase from the white-rot basidomycete Phanerochaete chrysosporium. Arch Biochem Biophys 234:353- 362 Green TR (1977) Significance of glucose oxidase in lignin degradation. Nature 268:78-80 Greene RV, Gould JM (1983) Substrate-induced H202 production in mycelia from the lignin-degrading fungus Phanerochaete chrysosporium. Biochem Biophys Res Commun 117:275-281 Greene RV, Gould JM (1984) Fatty acyl-coenzyme A oxidase activity and H2 0 production in Phanerochaete chrysosporium.281achem Biophys Res Commun 118:437-443 Janssen FW, Reulius HW (1968). Carbohydrate oxidase, a novel enzyme from Polyporus obtusus. Biochem Biophys Acta 167:501-510 Jeffries TW, Choi S, Kirk TK (1981) Nutritional regulation of lignin degradation by Eggnerochaete chrysosporium. Appl Environ Microbiol 42:290—296 Kelley RL, Reddy CA (1982) Ethylene production of a-oxo-y- methylthiobutyric acid is a sensitive measure of ligninolytic activity by Phanerochaete chrysosporium. Biochem J 206:423-425 Keyser T, Kirk TK, Zeikus JG (1978) Ligninolytic enzyme system of Phanerochaete chrysosporium: synthesized in the absence of lignin in response to nitrogen starvation. J Bacterial 135: 790- 797 ‘ Kirk TK, (1984) 'Degradation of lignin. In: DT Gibson (ed) Biochemistry of Microbial Degradation, Marcel Dekker, New York, pp 399-437 Kirk TK, Chang H-m (1974) Decompostion of lignin by white- rot fungi. I. Isolation of degraded lignins from decayed spruce. Holzforschung 28:218-222 Kirk TK, Connors WJ, Zeikus JG (1976) Requirement for a growth substrate during lignin decomposition wood-rotting fungi. Appl Environ Microbiol 32:192-194 Kirk TK, Connors WJ, Bleam RD, Hackett WF, Zeikus JG £1975) Preparation and microbial decomposition of synthetic ' lignins. Proc Nat Acad Sci USA 72:2515—2519 75 Kirk TK, Schultz E, Connors WJ, Lorenz LF, Zeikus JG (1978) Influence of culture parameters on lignin metabolism by Phanerochaete chrysosporium. Arch Michrobiol 117:277-285 Kutsuki H, Gold MH (1982) Generation of hydroxyl radical and its involvement in lignin degradation by Phanerochaete chrysosporium. Biochem Biophys Res Commun 109:320-327 Kuwahara M, Glenn JK, Morgan MA, Gold MH (1984) Separation and characterization of two extracellular HZOZ-dependent oxidases from ligninolytic cultures of Phanerochaete chrysosporium. FEBS Lett 169:247-250 Lowry 0H, Rosebrough NJ, Farr AL, Randall RJ (1951) Protein measurement with the folin phenol reagent. J Biol Chem 193:247-275 Melachouris N (1968) Discontinous gel electrophoresis of whey proteins, casein, and clotting enzymes. -J Dairy Sci-- 52:456-459 Miller GL (1957) Use of dinitrosalicylic acid reagent for determination of reducing sugar. Anal Chem 31:426-428 Morrissey JH (1981) Silver stain for proteins in polyacrylamide gels: a modified procedure with enhanced uniform sensitivity. Anal Biochem 117:307-310 Nakamatsu T, Akamatsu T, Miyajima R, Shio T (1975) Microbial production of glucose oxidase. Agr Biol Chem 39:1803-1811 Patterson A, Lundquist K (1985) Radical breakdown of lignin. Nature 316:575-576. Raabo E, Terkildsen TC (1960) On the enzymatic .determination of blood glucose. Scand J Clin Lab Invest 12: 402-405 ' Ramasamy K, Kelley RL, Reddy CA (1985) Lack of lignin degradation by glucose oxidase-negative mutants of Phanerochaete chrysosporium. Biochem Biophys Res Commun 131:436-441. Reddy CA (1984) Physiology and biochemistry of lignin degradation. In: Klug MJ and Reddy CA (eds), Current perspectives in microbial ecology, Am Soc Microbial, Washington, DC, pp. 558-571 Reddy CA, Forney LJ, Kelley RL (1983) Involvement of hydrogen peroxide- derived hydroxyl radical in lignin degradation by the white-rat fungus Phanerochaete chrysosporium. In: Higuchi T, Chang H- -m, Kirk TK (eds), Recent advances in biodegradation research, Uni Publishers Co LTD, Tokyo, Japan, pp. 153- 163 76 Tien M, Kirk TK (1983) Lignin-degrading enzyme from the Hymenomycete Phanerochaete chrysosporium Burds. Science 221:661-663 Tien M, Kirk TK (1984) Lignin-degrading enzyme from Phanerochaete chrysosporium: purification, characterization, and catalytic properties of a unique gégz-requiring oxygenase. Proc Nat Acad Sci USA 81:2280- Talbert NE (1981) Metabolic pathways in peroxisames and glyoxysomes. Ann Rev Biochem 50:133-157 77 Table l. The effect of different substrates on hydrogen peroxide (H202) production by cell extracts of glucose-grown ligninolytic cultures of Phanerochaete chrysosporiwn Substrate H202 production (nmol/min : mg) A D-Glucose 61.5 :L- 3.9 L-Sorbose 28.2 i 2.7 D-Cellobiose 12.5 :1; 3.4 D-Maltose 10.3 i 1.8 D-Mannose 3.5 -l_- 1.0 D-Xylose 0.5 i- 0.26 B i. Glucose-(anaerobically under N2 gas) trace _ ii. Glucose-(after replacing N2 in B.i. above with oxygen) 63.5 i 14.5 Cultures grown for 6d in low N basal medium were used for pre- paring the cell extracts. All substrates were used at 0.1 M final concentration. Values given are mean + standard deviation and have been corrected for the “no substrate” controls 78 Table 2. The effects of different co-substrates on ligninolytic activity and on hydrogen peroxide (H202) production by cultures of P. chrysosporium Co-Substrate Ligninolytic Assay Specific activity activity substrate for H202 production (dpm/day - flask) (nmol/min - mg) D-Glucose 604.2 i 149.1 glucose 80.7 i 0.2 D-Xylose 291.7 i 153.9 xylose 14.5 :1: 2.0 glucose 113.5 1 7.0 Succinate 216.4 -_l_- 141.1 succinate 0.0 glucose 57.0 i 9.5 Cultures were grown separately in a low nitrogen (2.4 mM N) medium containing equimalar carbon concentrations of one of the following substrates: 56 mM glucose, 66.7 mM xylose, or 83.0 mM dissodium succinate for 14 d at 39°C. All values given represent the mean -_1- standard deviation and have been corrected for the “no substrate” controls 79 Table 3. The effect of nitrogen and carbohydrate concentrations on the specific activity for lignin degradation and glucose oxidase activity Medium Ligninolytic Glucose oxidase activity activity (dpm/day - flask) (nmol/min - mg) A. Low nitrogen 330.1 -_1_- 164.0 118.4 : 1.3 High nitrogen 25.0 i- 13.4 0.6 i 0.2 B. Low nitrogen 466.1 i 134.8 84.5 i- 2.4 Law nitrogen + (NH4)ZSO4 (2.8 mM) 141.7 : 103.7 1.6 i 0.02 Low nitrogen + glutamate (2.6 mM) 55.4 i 48.2 1.1 :1; 0.3 C. Low carbohydrate 54.6 i- 9.3 2.1 i 1.5 High carbohydrate 14.4 i 0.5 0.9 :1; 0.4 Low nitrogen (2.4 mM N), high nitrogen (24 mM N), low carbohydrate (8.8 mM glucose) and high carbohydrate (56 mM glucose) media were described previously (Kelley and Reddy 1982). See methods for other details. Ligninolytic activity is based on the amount of 14C02 released by 6-day-old cultures. Cell extracts of 6 day old cultures was used for determining glucose oxidase activity. Additional N sources were added to cultures on the fourth day of incubation. 80 Figure l. Polyacrylamide gel electrophoresis of cell extracts from 6-day-old cultures of EL-chrysasporium grown~ in low N medium (containing glucose as the substrate). Tube gels loaded with 1.4 mg protein were stained overnight for 8202 generating activity (Cohen 1972) using the diaminobenzidine/horseradish peroxidase staining procedure, in the presence of D-glucose (l), L-sorbose (2), D- cellobiose (3), D-xylose (4), D-maltose (5), and D-mannose (6). In this procedure, HZOZ-generating protein bands will stain dark brown (i.e. DAB-positive) due to the 'peroxidative oxidation of diaminObenzidine. 81 Figure l. 82 Figure 2. Polyacrylamide gel electrophoresis of cell extracts of g; chrysosporium. Slab gel in panel A were stained for glucose oxidase activity using the procedure described in the legend for Fig. l. Glucose was used as the substrate. Gels in panel B were stained for protein by the silver staining procedure (Morrissey 1981). In each panel, lanes 1, 2, 3 and 4 represent cell extract from ligninolytic culture (grown for 6 days in low N medium: 70 ug protein), extracellular culture fluid (20X) from ligninolytic cultures (20ug protein), extracts of non- Tigninolytic cultures grown in low N medium for 4 days (40 ug protein) or in high N medium from 14 days (40 ug protein), respectively. 83 Figure 2. 84 IFigure 3. ‘Polyacrylamide gel electrophoresis of cell extracts of ligninolytic cultures grown on various growth substrates. Gels were stained for HZOZ-generating activity (panel A) or protein (panel B) using the procedures described in the legend for Fig. 1. Lanes 1, 2, 3 and 4, respectively, contained cell extracts from cultures grown in low N medium with glucose (70 ug protein), xylose (60 ug protein), succinate (50 ug protein), or cellobiose (60 ug protein) as the growth substrate for 9 d. 85 Figure 3. 86 Chapter two: Purification and Characterization of Glucose Oxidase from Ligninolytic Cultures of Phanerochaete chrysosporium 87 ABSTRACT Glucose oxidase, an important source of hydrogen peroxide in lignin degrading cultures of Phanerochaete chrysosporium, was purified to electrophoretic homogeneity by a combination of ion-exchange and molecular sieve chromatography. The enzyme is a flavoprotein with an apparent native molecular weight of 180,000 daltons and a denatured molecular weight of 80,000 daltons. This enzyme does not appear to be a glycoprotein unlike glucose. oxidases reported from other fungi. It gives optimal activity with D-glucose, which is stoichiometrically oxidized to D-gluconate; however, it also exhibits some activity with L-sorbose, D-xylose and D-maltose. The enzyme has a relatively broad pH optimum of 4 to 5. It is inhibited by Ag+ and o-phthalate, but not by Cu++, NaF or KCN (each 10 mM). INTRODUCTION Hydrogen peroxide (H202) plays an important role in the ligninolytic system of Phanerochaete chrysosporium, a basidiomycete extensively used in studies of lignin biodegradation (16,21). Forney g; 21, (9,10) showed a temporal correlation between ligninolytic activity and H202 production. Both H202 production and ligninolytic activity increased when cultures were incubated under 100% 02, and lignin degradation was inhibited by catalase, which metabolizes 8202 to yield 02 and H20 (7). Nutritional 88 parameters, which are known to affect ligninolytic activity, such as nitrogen and carbohydrate concentration, were shown to have a similar effect on H202 production (12.15.18: C. A. Reddy and R. L. Kelley, _i_n C. O'Rear and G. C. Llewellyn, ed, Biodegradation 6, in press). HZOZ-producing periplasmic microbodies were seen only in lignin degrading cultures but not in non-ligninolytic cultures (10). H202-dependent extracellular, lignin- degrading oxygenases (ligninases) have been demonstrated in ligninolytic cultures of 2, chrySosporium (2,11,15,24).' Glucose oxidase (EC 1.13L4) has been recently identified as the predominant source of H202 production in ligninolytic cultures of E. chrysosporium (Reddy and Kelley, in press) as evidenced by the following observations: Glucose supported the highest level of H202 production in cell extracts. Polyacrylamide gel electrophoresis of these extracts showed the presence of a single protein band that supported glucose-dependent H202 production; this protein band was missing in extracts of non-ligninolytic cultures. An H202-producing protein band with similar electrophoretic mobility was observed in cell extracts regardless of the fact whether glucose, cellobiose, xylose, or succinate was employed as the growth substrate. Both glucose oxidase activity and ligninolytic activity are secondary metabolic events and are triggered in response to nitrogen or carbohydrate starvation (Reddy and Kelley, in press). In this report, we have described 89 the characteristics of glucose oxidase purified to electrophoretic homogeneity from ligninolytic cultures of ‘2. chrysosporium. MATERIALS AND METHODS Qggenism and culture conditions. 2; chrysosporium (ATCC 34541) was maintained and conidial inoculum was prepared as previously described (17). Sterile media in foam-stoppered flasks were inoculated with conidial suspensions in water (1.25 x 106 conidia/ml, 0.5 m1 inoculum/10 ml medium) as previously described (17). The flasks were incubated at 37°C without agitation for 6 days. Assay for glucose oxidase. Glucose oxidase activity was determined at 37°C by monitoring the change in absorbance at 460 nm due to the peroxidative oxidation of o-dianisidine by horseradish peroxidase and using a molar extinction coefficient of 8.3 (5). The reaction mixture consisted of 1.5 ml of citrate-Na phosphate buffer (0.1 M, pH 4.5), 1.0 ml of o-dianisidine (0.3 mM), 0.3 ml of a 1 M solution of the substrate (such as D-glucose, L-sorbose, D- xylase or D-maltose) in water, 0.1 horseradish peroxidase (HRP; 60 U/ml; EC 1.11.1.7; Sigma Chemical Co., St. Louis, MO), and 0.1 ml glucose oxidase solution. The reaction mixture was bubbled with 100% 02 for 10 min prior to the addition of the glucose oxidase. Electrophoresis. Sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) of different glucose oxidase preparations was done according to Laemmli (19) using gel slabs (14 x18 cm x1.5 mm) with a 4X 90 stacking gel and a 10% running gel. Samples (50 ul) were placed on the slab gel in.CL05 M tris-HCL buffer (pH 6.7) containing 20% w/v glycerol, 4% SDS, and 10% 2- mercaptoethanol. Electrophoresis was performed at 30 mA for 3.5 h using a Buchler Model 3-1014A power supply (Buchler Instruments Div., Fort Lee, NuL) set at constant amperage. Gels were stained for protein with Coomassie brillant blue R250 as previously described (27). Gels were stained for protein-bound carbohydrate by using the dansy1_ hydrazine staining procedure as described by Eckhardt g; 3;. (6). ' Protein and flavin assays. Protein content was determined by the procedure of Lowry gg'gl. (20) using Bovine serum albumin (IV, Sigma Co., St. Louis, MO) as the standard. Flavin content was determined by the method of 1. ('4). Cerletti g; Purficatian a; glucose oxidase. All purficatian steps _were done at 4°C. The pH of the phOSphate buffer used was 6.8. Protein solutions were concentrated as needed by ultrafiltration using an Amican ultrafiltration unit (Amicon Corp., Lexington, MA) equipped with a PM-lO filter (10,000 daltons pore size). i. Preparation 2; cell extracts. Cultures of 3; chrysosporium grown in low nitrogen medium for 6 d at 37°C, were collected by filtration of six layers of gauze in a Buchner funnel. The mycelial mat (-15 g dry wt.) was washed 3 times with 200 ml of 0.1 Na phosphate buffer (P04 91 buffer). The washed mycelium was resuspended in 100 ml of the same buffer, mixed with glass beads (0.1 mm) in a 1:1 ratio (glass beads:mycelial wet wt.), and blended at 4°C using an Omni-mixer (Sorvall Inc., Norwalk, CT) for 15 min. The glass beads and unbroken mycelium were removed by centrifugation at 4080 x g for 10 min. The supernatant was saved and the pellet was resuspended in 10 ml of P04 buffer and blended for an additional 15 min. The supernatants were combined and frozen until needed. ii. DEAR-Sephadex Chromatography. The frozen cell extracts were thawed, clarified by centrifugation at 27,000 x g for 15 min at 40C, and diluted S-fold with distilled water. This protein solution was applied to a DEAE- Sephadex (A50; Pharmacia Fine Chemicals, Piscataway, NJ) column (50 m1 of gel, 2.4 x 16 cm gel bed) previously equilibrated with 0.01 M P04 buffer. The column was washed with 50 m1 cf the same buffer, and the protein was eluted stepwise from the column with 100 ml volumes of 10 mM P04 buffer containing 0.05, 0.10 and 0.25 M NaCl. Fractions (2.25 ml) were collected and tested for glucose oxidase activity as described above. Fractions with the highest activity were pooled and concentrated by ultrafiltration. iii. Sephacryl chromatoggaphy. The concentration protein from the DEAE-Sephadex step was loaded onto a Sephacryl S-300 column (Pharmacia, 170 ml of gel, 2.4 cm x 46 cm gel bed) equilibrated with 0.1 M P04 buffer. The column was eluted with the same buffer at a flow.rate of 0.25 ml/min. Fractions (1.5 ml) were collected and those 92 with the highest activity were pooled. iv. DEAE-Sepharose chromatogyaphy. The pooled fractions were applied to a DEAE-Sepharose CL-6B column (Pharmacia, 20 m1 of gel, 1.6 x 20 cm gel bed) previously equilibrated with 0.01 M P04 buffer. Protein was eluted from the column with a linear salt gradient (440 ml total vol., 0 to 100 mM NaCl) in 0.01 M P04 buffer. The NaCl concentration in each fraction was calculated by using conductivity values as compared to those of NaCl standards. The flow was 0.25 m1/min and fractions (1.5) were collected and tested for activity. Molecular weight determination. The molecular weight of the purified glucose oxidase was determined by gel filtration chromatography, using a Sephacryl S-300 column. The column was calibrated with Biorad gel filtration standard containing thryoglobulin (670,000), gammaglobulin (158,000), ovalbumin (44,000), myoglobin (17,000) and vitamin 812 (1,350) obtained from BioRad Laboratories, Richmond, CA. Molecular weight marker kit for SDS gel electrophoresis (MW-SDS-ZOO) was purchased from Sigma Chemical Co. St. Louis, MO. Effect pf 25. A citrate-Na phosphate buffer adjusted to pH 3 to 6 was used to determine the optimal pH for glucose oxidase activity. Because pH could affect both glucose oxidase activity as well as the peroxidatic oxidation of o-dianisidine, glucose oxidase activity in this experiment was determined by monitoring the production 93 of H202 directly as measured by the change in absorbance at 240 nm at 37°C. The reaction mixture consisted of 0.5 m1 of citrate-P04 buffer (0.1 M. pH 3 to 6), 0.1 ml glucose solution (1 M) and 0.4 ml of glucose oxidase (0.1 mg protein/ml). Enzyme inhibition. Different metal ions or o- phthalate citrate-P04 buffer (0.1 M, pH 4.5) were added to the glucose oxidase assay mixture described above. Since KCN is known to inhibit HRP, activity of purified glucose oxidase in the presence of KCN was determined by measuring the anaerobic reduction of 2,6-dichlorophenol-indaphenol (25). Determination g; apparent £2 and Vmax. The apparent Km for glucose of the purified glucose oxidase was determined by measuring the initial velocities over a range of glucose concentrations (2 to 125 mM) at an 02 concentration of 1.6 mM. For determining the Km for oxygen, the reaction mixtures, containing 0.1 M glucose, in stoppered cuvettes were bubbled with 100, 80, 60, 40, 20, or 10% 02 in N2, which was obtained by mixing the gases through a pair of calibrated flow-meters (model 7322, Matheson, East Rutherford, NJ). After bubbling for 10 min at 37°C, the reaction mixtures were then allowed to equilibrate for 15 min at 37°C. The initial oxygen concentration in each reaction mixture was determined by measuring the amount of dissolved 02 present in an identically treated parallel cuvette using a biological oxygen monitor (model 5331, Yellow Springs Instrument, Co” 94 Yellow Springs, Ohio) equipped with a Clark-type electrode (22). Apparent Km values were calculated from Lineweaver- Burk plots (Fig. 5 A & B).- Quantification 2; glucose and gluconate. D-glucose and D-gluconate were identified and quantified by gas- 1iquid chromatography. Purified enzyme (0.5 ml containing 0.35 U/ml) was added to a reaction mixture containing 200 ul catalase solution (1 mg/ml) and 0.5 m1 glucose solution (0.1 M in 0.1 M citrate-phosphate buffer) and incubated for 16 h at 37°C. An internal standard of L-erythritol was added to the reaction mixture prior to sialylation. The reaction mixture was evaporated to dryness and dissolved in methanol (acidified with 50 ul trifluoraacetic acid/ml). Undissolved material was removed by centrifugation and the supernatant was evaporated to dryness, dissolved in 0.5 ml acetonitrile plus 0.5 m1 N,0-bis-(trimethylsily1) trifluoraacetamide (Pierce Chemical Co., Rockford, IL), and - was heated for 30 min at 70°C. Samples were analyzed using a Varian 3700 gas chromatograph equipped with a flame ionization detector, a Hewlett-Packard 3390A digital integrator and a glass column (2 m x 0.3 cm) packed with 3% SE-30 on 80/100 mesh Chromosorb W(HPL. The carrier gas was He at 25 ml/min and the chromatograph oven was temperature programmed to hold at 140°C for 10 min and then to increase at 2°C/min to 220°C. D-glucose and D-gluconate were quantified from peak areas as compared to those of the standards. 95 RESULTS Purification 3; glucose oxidase. The purification of glucose oxidase from P. chrysosporium is summarized in Table 1. A purification of about 90-fold was routinely achieved. In the DEAR-Sephadex step, about 602 of the total glucose oxidase activity present in crude cell extracts was recovered in the 0.25 NaCl eluate, giving about a five-fold increase in specific activity. The subsequent Sephacryl S-300 step (Figr 1A) produced a 38- fold enrichment in specific activity and a 41% recovery of total activity. The elution profile from the DEAE- Sepharose column (Fig. 18) showed that a single protein peak had all the glucose oxidase activity with approximately 90-fold enrichment in specific activity and an enzyme recovery of 92. The DEAE-Sepharase protein fraction was found to be homogeneous based on SDS-PAGE analysis (Fig 2). Molecular weight. Based on gel filtration chromatography on Sephacryl S-300 column, the apparent molecular weight of purified glucose oxidase was estimated to be 180,000 daltons (Fig. 3). The denatured molecular weight, determined by SDS-PAGE, was estimated to be 80,000 daltons (Fig. 4A). Carbohydrate and flavin content. Staining of the purified glucose oxidase from E, chrysosporium for protein bound carbohydrate by the dansyl hydrazine method showed no detectable carbohydrate (Fig. 4B, lane 2), whereas an equal amount of commercially prepared glucose oxidase from 96 Aspergillus niger, which is known to be a glycoprotein (5,23), stained positive (Fig. 4B, lane 1). Flavin analysis indicated that the purified enzyme contains 1.5 moles of flavin per mole of protein. pH optimum. The purified enzyme had a pH optimum between 4.6 and 5.0. In comparison, glucose oxidase from P, natatum and A, giggg were reported to have a pH optima of 5.5 and 5.6, respectively (1,3). Enayme inhibition. Glucose oxidase fromfi. chrysosporium, similar to that from A, niger, was inhibited by Ag+, but was not inhibited by Cu++, KCN, or NaF; in fact, there was substantial stimulation of activity in the presence of the latter (Table 2). Also, glucose oxidase from y. chrysosporium was severely inhibited by a- phthalate, whereas the commercial glucose oxidase from A. glgg; showed only a limited inhibition of activity. Kinetic prpperties and substrate specificity. The apparent KIn values_for glucose and 02 for this enzyme were 38 mM and 0.95 mM respectively (Fig. 5A and B). The V data (Table 3) show that glucose is the primary max/Km substrate for the enzyme whereas the others are relatively minor substrates at best. Furthermore, compared to a specific activity of 12.27 (unol min/ min per mg) with glucose as the substrate, the following substrates had 5 1% activity: cellobiose, glycolate, mannose, gluconate, ethanol, acetate, lactate, succinate, pyruvate, D:galactose and D-gluconalactone. 97 D-glucose was stoichiometrically oxidized to D- gluconate: from 28.6 males of D-glucose oxidized, we obtained 26.1 males of gluconate which amounts to 91.21 recovery. These results are in agreement with the results obtained with glucose oxidases from other fungi (1,3,5). DISCUSSION Fungal glucose oxidase (B-D-glucose:oxygen oxidoreductase, EC 1.1.3.4) catalyzes the oxidation of D- glucose to 6-D-gluconolactone and 8202 in the presence of molecular oxygen (1.3.4.25; see the reaction sequence below). In a subsequent step 5-D-gluconolactone is nonenzymatically hydrolyzed to D-gluconic acid. This enzyme has been demonstrated in various Aspergillus and Penicillum species (3.23.28). B-D-glucose Glucose oxidase H202 (oxidized) I 6-D-gluconolacton Glucose oxidase 02 (reduced) - D-gluconic acid Certain enzymes in animal tissues also catalyze oxidation of D-glucose (or derivatives) to d-D-gluconolactone, but these are readily differentiated from glucose oxidase because they do not require molecular oxygen and H202 is not a product (28). Glucose oxidase from different fungi has a molecular weight range from 150,000-186,000 daltons and normally consists of two identical polypeptide chain subunits covalently linked by disulfide bonds (3.5.28). The glucose oxidase that we have isolated from g. chrysosporium is a flavoprotein with a native molecular 98 weight of 180,000 daltons and a denatured molecular weight of 80,000 daltons. Presumably this enzyme, similar to other glucose oxidases, consists of two identical polypeptides (MW of 80,000 daltons each). The overestimation of the native molecular weight may perhaps be due to hydrodynamic properties of this enzyme different from other glucose oxidases. Our flavin analysis data revealed 1.5 mol of flavins per mol of purified glucose oxidase from 2. chrysosporium. Using identical procedures, we showed that A, pigg£_glucase oxidase has 1.6 flavins per mole of protein. Since A, giggg enzyme has been shown to have two flavins per mole of protein by a number of earlier investigators (3.23.25), we believe that both E. chrysosporium and A, gigg; glucose oxidases actually contain two mol of flavins per mol of protein and the lower value of 1.5 to 1.6 we obtained experimentally is apparently due to a limitation of the analytical procedure _ employed by us. No carbohydrate was detectable in glucose oxidase from P; chrysospprium based on the dansyl hydrazine method (6) used in this study (Fig. 48). Under identical conditions, an equal amount of the enzyme from £1132, which has been reported to contain approximately 18% sugar residues (5), stained strongly positive. These results suggest either that E; chrysosporium glucose oxidase is not a glycoprotein or that it contains a very low level of carbohydrete which is not detectable by the procedure used (6). This finding 99 is of interest in light of the previous observations that a majority of peroxisomal proteins appear not to be glycosylated (25.26) and that 8202 production in ligninolytic cultures of P, chrysosporium. presumed to be due to glucose oxidase activity, has been shown to be localized in periplasmic, peroxisome-like structures (10). Glucose oxidases from other fungal sources have been shown to possess a relatively low affinity for glucose with Km values ranging from 0.11‘mM to 33 mM and a slightly higher affinity for 02 with KIll values from 0.2 to 0.83 mM (3,24). The Kmvalues for glucose and 02 (38 mM and 0.95 mM. respectively) for the enzyme isolated from 2. chrysosporium fall within the range of the values reported for previously described glucose oxidases. Glucose oxidase from Aspergillus and Penicillim was shown to be highly specific for B-D-glucose. Although D- mannose, D-galactose, 2-deoxy-D-glucose and D-xylose have been shown to exhibit low activities as substrates, no ‘greater than‘ZXof the activity found with glucose was found with these or 50 other carbohydrates tested (1,3). The enzyme from g, chrysosporium on the other hand appears to be less specific in that it gave 331, 132 and 72 specific activity, respectively, with sorbose, xylose and maltose compared with that seen with glucose as the substrate. However, a comparison of the V ratios for max/Km the different substrates clearly shows that glucpse is the primary substrate for this enzyme. Another HZOZ-producing enzyme, designated carbohydrate oxidase, has been partially 100 is of interest in light of the previous observations that a majority of peroxisomal proteins appear not to be glycosylated (25,26) and that H202 production in ligninolytic cultures of E, chrysosporium, presumed to be due to glucose oxidase activity, has been shown to be localized in periplasmic, peroxisome-like structures (10). Glucose oxidases from other fungal sources have been shown to possess a relatively low affinity for glucose with Km values ranging from‘0.ll mM to 33 mM and a slightly higher affinity for 02 with Km'va‘lues from 0.2 to 0.83 mM (3,24). The Km values for glucose and 02 (38 mM and 0.95 mM. respectively) for the enzyme isolated from P. chrysosporium fall within the range of the values reported for previously described glucose oxidases. Glucose oxidase from Aspergillus and Penicillim was shown to be highly specific for B-D-glucose. Although D- mannose, D-galactose, 2-deoxy-D-glucose and D-xylose have been shown to exhibit low activities as substrates, no greater than 2% of the activity found with glucose was found with these or 50 other carbohydrates tested (1,3). The enzyme from 2, chrysosporium an the other hand appears to be less specific in that it gave 33%, 13% and 7% specific activity, respectively, with sorbose, xylose and maltose compared with that seen with glucose as the substrate. However, a comparison of the V ratios for max/Km the different substrates clearly shows that glucose is the primary substrate for this enzyme. Another HZOZ-producing enzyme, designated carbohydrate oxidase, has been partially 101 purified from extracts of a white-rot fungus, Polyparus obtusus (13). This enzyme exhibited 59% and 38% actively, respectively, with L-sorbose and D-xylose compared to that observed with glucose. 2, obtusus enzyme was however, different from P, chrysosporium glucose oxidase in that it could utilize D-gluconate as the substrate (14% of the activity observed with glucose) and showed no activity with D-maltose. A third type of H202-generating oxidase, L- sorbase oxidase from Trametes sangpinea, which catalyzed the oxidation of L-sorbose, D-glucose, D-galactose, D- xylose and D-maltose, has been described (29). It has been suggested that this enzyme is similar to the E, obtusus carbohydrate oxidase (13). The apparent low substrate specificity of P, chrysosporium glucose oxidase described here may allow the organism to utilize sugars derived not only from cellulose but also from hemicelluloses found in woody material, its natural habitat, to produce H202 which is known to be important to the ligninolytic system. Inhibition studies showed that glucose oxidase from E. chrysosporium, similar to glucose oxidase from A, giggg, is inhibited by Ag+ but not by Cu‘H'. NaF, or KCN (Table 2). Earlier results showed inhibition of lignin degradation when o-phthalate was used as a buffer in the growth medium (8). The results of this study show that glucose oxidase from E, chrysosporium is severely inhibited by o-phthalate suggesting that the inhibition of lignin degradation by this compound may at least partially be due to its effect 102 on 8202 production by glucose oxidase. In this report we have described the purification, characterization and kinetic properties of glucose oxidase from P, chrysosporium. This enzyme is similar in its physical and kinetic properties to glucose oxidases isolated from other fungal sources, except that we were unable to demonstrate the presence of carbohydrate in this protein. The enzyme is severely inhibited by a-phthalate and Ag+. 103 LITERATURE CITED 10. 11. Adams, E., R. Mast, and A. Free. 1960. Specificity of glucose oxidase. Arch. Biochem. Biophys. 91:230-234. Anderson, L. A., V. Renganathan, A. A. Chiu, T. M. Loehr, and M. H. Gold. 1985. Spectral characterization of diarylpropane oxygenase, a novel peroxide-dependent, lignin-degrading heme enzyme. J. Biol. Chem. 260:6080- 6087. . Bentley, R. 1963. Glucose aerodehydrogenase (glucose oxidase). Methods Enzymol. 6:37-97. Cerletti, P., R. Strom and M. G. Giordano. 1963. Flavin peptides in tissues: the prosthetic group of- succinic dehydrogenase. Arch. Biochem. Biophys. 101:423-428. Decker, L. A. (edJ. 1977. Worthington enzyme manual, Worthington Biochemical Corp., Freehold, NJ. Eckhardt, A. E., C. B. Hayes, and I. J. Goldstein. 1976. A sensitive fluorescent method for the detection of glycoproteins in polyacrylamide gels. Anal. Biochem. 75:192-197. Faison, 8., and T. K. Kirk. 1983. Relationship between lignin degradation and production of reduced oxygen species by Phanerochaete chrysosporium. Appl. Environ. Microbial. 46:1140-1145. Fenn, P” andTL K.Kirk. 1979. ldgninolytic activity of Phanerochaete chrysosporium: inhibition by a- phthalate. Arch. Microbial. 134:307-309. Forney, L. J., C. A. Reddy, M. Tien, and S. D. Aust. 1982. The involvement of hydroxyl radical derived from hydrogen peroxide in lignin degradation by the white-rot fungus Phanerochaete chrysosporium. J. Biol. Chem. 257:1455-1462. Forney, L. J., C. A. Reddy, and H. S. Pankratz. 1982. Ultrastructural location of hydrogen peroxide production in ligninolytic Phanerochaete chrysosporium cells. Appl. Environ. Microbiol. 44:732-736. Frick,13 D” R.I“ Crawford. 1983. Mechanisms of microbial demethylation of lignin model polymers, pp 143-152. In: T. Higuchi, H. M. Chang, and T. K. Kirk gated, Lignin biodegradation research, Uni delishers. 0 yo. 12. 13. 14. 15. 16. 17. 18. 19. 20. 21. 22. 23. 104 Greene, R. V., and J. M. Gould. 1984. Fatty acyl- coenzyme A oxidase activity and H20 production in Phanerochaete chrysosporium mycelia. Biochem. Biophys. Res. Commun. 118:437-443. Janssen, F. W., and H. W. Ruelius. 1968. Carbohydrate oxidase, a novel enzyme from Polyporus obtusus. Biochem. Biophys. Acta 167:501-510. Kelley, R. L., and C. A. Reddy. 1982. Ethylene production from a-oxo-y-methylthiobutyric acid as a measure of ligninolytic activity by Phanerochaete chrysosporium. Biochem. J. 206:423-425. Kersten, P. J., M. .Tien, B. Kalyanaraman, and T. K. Kirk. 1985. The ligninase of Phanerochaete chrysosporium generated cation radicals from methoxybenzenes. J. Biol. Chem. 260:2609-2612. Kirk. T. K. 1984. Degradation of lignin, pp. 399-437. ‘lg D. T. Gibson (ed.), Biochemistry of microbial degradation. Marcel Dekker, New York, NY. Kirk, T. K., E. Schultz, W. J. Connors, L. F. Lorenz, and J. G. Zeikus. 1978. Influence of culture parameters on lignin metabolism by Phanerochaete chrysosporium. Arch. Micorbiol. 117:277-285. Kutsuki, H” and M.lL Gold. 1982. Generation of hydroxyl radical and its involvement in lignin degradation by Phanerochaete chrygosporium. Biochem. Biophys. Res. Commun. 109:320-327. Laemmli, U. K. 1970. Most commonly used discontinuous buffer system for SDS electrophoresis. Nature 227:680- 688. Lowry, 0. H., N. J. Rosebrough, A. L. Farr, and R. J. Randall. 1951. Protein measurement with the folin phenol reagent. J. Biol. Chem. 193:265-275. Reddy, C. A. 1984. Physiology and biochemistry of lignin degradation, pp. 558-571. 13 M. J. Klug and C. A. Reddy (eds.), Current perspectives in microbial ecology. American Society Microbiology. Washington, DC. Robinson,.1n and J.lL Cooper. 1970. Method of determining oxygen concentrations in biological media. suitable for calibration of the oxygen electrbde. Anal. Biochem. 33:390-399. Swoboda, B. E. P. and B. Massey. 1965. Purification 24. 25. 26. 27. 28. 29. 105 and properties of glucose oxidase from Aspergillus niger. J. Biol. Chem. 240:2209-2215. Tien, M., and T. K. Kirk. 1984. Lignin-degrading enzyme from Phanerochaete chrysosporium: purification, characterization, and catalytic properties of a unique H 0 -requiring oxygenase. Proc. Natl. Acad. Sci. USA. 81:2280-2284. Talbert, N. E. 1971. Leaf peroxisames. Meth. Enzymol. 23:679-680. Volkl, A., and P. B. Lazarow. 1982. Affinity chromatography of peroxisomal proteins on lectin- sepharose columns. Ann. N.Y. Acad. Sci. 386:504-506. Weber, K. and M. Osborn. 1969. Method for staining polyacrylamide gels for protein with Coomassie brillant blue (R250). J. Biol. Chem. 244:4406-4412. Whitaker, J. R. 1972. Principles of enzymology for the food sciences. pp. 561-571. Marcel Dekker, Inc., New York, NY. Yamada, Y., K. Iizuka, K. Aida, and T. Uemura. 1967. Enzymatic studies on the oxidation of sugar and sugar alcohol. III. Purification and properties of L-sorbose oxidase from Trametes sanguinea. J. Biochem. (Tokyo) 62:223-229. 106 TABLE 1. Purification data for glucose oxidase from ligninolytic cultures of P. chrysosporium Total Total . . . . Sp act . . . YlCld Purification Fraction 0 protein actmty (U/mg) (mg) (U) (%) (fold) Crude cell extract 0.17 285 48.4 100 1.0 DEAE—Sephadex 0.91 32 29.1 60 5.4 Sephacryl S-3OO 6.39 3.1 19.8 41 37.8 DEAE-Sepharose 15.1 0.28 4.2 9 89.3 ‘ Specific activity is defined in units per milligram of protein. One unit of activity represents the oxidation of 1 umol of o—dianisidine per min at 37°C and pH 4.5. 107 TABLE 2. Comparison of the effects of o-phthalate, KCN, NaF, and different metal ions on glucose oxidase from P. chrysosporium and commercially prepared A. niger glucose oxidase . - % Activity of glucose oxidase . . Final concn from: Addition (mM) A. niger“ P. chrysosporium None 100 100 KCN 10 ‘ 128 135 NaF . 10 101 111 Cqu 10 133 123 Agso. 10 36 25 o-Phthalate 50 86 12 ‘5 Commercial A. niger glucose oxidase was obtained from Sigma. The concentration of each enzyme preparation was adjusted to give 0.015 U of activity per ml. 108 TABLE 3. Substrate specificity of purified glucose oxidase“ Vmax Substrate (nmol/min (£122) V.,“,IK," 923p per ml) D-Glucose 15 38.0 0.395 100 L-Sorbose 5 217.4 0.023 5.8 D-Xylose 2 105.2 0.019 4.8 D-Maltose 1 55.5 0.018 4.5 " Initial velocity was determined by the o-dianisidine—horseradish peroxi- dase assay described in Materials and Methods. 109 .Fig. 1.A. Elution profile of protein and glucose oxidase activity (nmol/min/mg protein) on Sephacryl S-3OO column. Fractions of 1.5 ml were collected and assayed for activity and protein. B. Elution profile of protein and glucose oxidase activity from a column of DEAR-Sepharose. Protein was eluted from the column with a linear salt gradient (see Material and Methods). Each datum point for salt concentration ) was determined by conductivity measurements. Fractions (1.5 ml) were assayed for glucose oxidase activity and for protein content. .93 .15 .03. w w I 4 ... u ... o . . 0 0 A 00 o I o 7 . . . U . 0 . 8 . C C r U . . O M C 5 m C . . . m . i . O . . -. 4 n 1w 0 o o s A O t o O o c I A m _a n . .. l I F . o O . . a 2 . . . ie x O . . 1 A . ._o —‘ p p b p a . 4. a ... .. a _ O 0 «0.6.0 0 e ... a - e 4 ... . . oo~< . . A A 1.0-u. 0. o. o. w m 01 L . b b b «‘DQIVD b h 4 3 1 1 O U ..c h h ... 2 1. 1. 1. C E C “Iv A . x .05: v >2>=O( “6.630.063 2.2.0.11» Fraction Number Figure 1. 111 Fig. 2. Sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) of glucose oxidase preparation obtained from different purification steps (see Table 1). Molecular weight standards (lane MW), the crude cell extract (lane 1: 0.14 mg protein), the DEAR-Sephadex fraction (lane 2; 20 ug protein), the Sephacryl S-3OO (lane 3 9.5 ug protein), and DEAR-Sepharose fraction (lane 4; 1 ug protein) were stained with Coomassie blue as described in the text. 112 Figure 2 . 113 Fig. 3 Molecular weight determination of purified glucose oxidase by gel filtration chromatography with_a Sephacryl S-300 column. See the text for details. V Elution volume. Ve(m|) 200 150 Vitamin 8-12 114 Glucose Oxidase 100'- Myoglobulln Ovalbumin T Gamma Globulln 50 '- Thyroglobulin 1 11111111 1 11111111 1 11111111 4 6 10 105 10 Molecular Weight . Figure 3. 115 Fig. 4. Molecular weight determination and staining for carbohydrate and protein after SDS-PAGE of commercial glucose oxidase from A, 2135; and that from g. chrysosporium. In panel A, molecular weight standards (lane MW), A, gigs; glucose oxidase (lane 1) and glucose oxidase from £5 chrysosporium (lane 2) were stained with Coomassie blue. In panel B, A, gigg; glucose oxidase (panel B, lane 1) and glucose oxidase from E, chrysosporium (lane 2) were stained for carbohydrate by the dansylhydrazine method as described in the text. In each panel, 1 ug of the respective enzyme was used. 116 Figure 4. 117 Fig. 5. Lineweaver-Burk plots showing the effect of glucose and 02 concentration on glucose oxidase activity. Glucose oxidase was assayed as described in methods. (A) Glucose concentration was varied at an 02 concentration of 1.6 mM, (B) 02 concentration was varied at glucose concentration of 0.1 M. Each point is the mean of these three values. Each reaction contained 1.6 ug of enzyme. IN (nmolelmln x 10") UV (nmolelmln x 10”) 118 14«- 12'"- 10" a«- ‘11-. 4-- 2., j a 1 a _1 l 1 1 J 1 I Ufi 1 U 1 'U 1 ‘l I 2 4 C I 101214101820 "mucosa“! x 10.2) 14-- - O 12-- . 1o-- 6‘- ...- 4.... 1 2 <1- % l l . 1 1 2 3 4 5 llCOz] MM Figure 5. IJIOnw -.__ III. 119 Chapter three: Characterization of Glucose Oxidase-Negative Mutants of a Lignin Degrading Basidiomycete Phanerochaete chrysosporium 120 I ABSTRACT Previous studies have shown that H202 plays an important role in lignin degradation by Phanerochaete chrysosporium, and that g1ucose.oxidase.(EC 1.15%4) is the primary physidlogical source of M2 2 in ligninolytic cultures of this organism. The isolation and characterization of glucose oxidase-negative (325‘) mutants of B. chrysosporium is described. These mutants are deficient not only in their ability to produce H202 but also in lignin degradation (2'—14C-synthetic lignin — 14C02), ligninase and peroxidase activities, decolorization of the dye poly-R 481, and production of ethylene from - oxy- -methylthiobutyric acid (KTBA). The 331‘ mutants retained, albeit at a lower level, the capacity to produce veratryl alcohol, a typical secondary metabolite, and produced conidia at a level comparable to that of the wild type. The addition of ligninase and/or glucose oxidase to a 321‘ mutant (COX-10) did not enhance its capacity to degrade lignin. The Gox+ revertant strains regained glucose oxidase activity, the ability to degrade lignin, as well as the other characteristics that were missing in the gox‘ mutants. The results suggest that the genetic lesion 121 in these mutants affects the regulation of a set of secondary metabolic characteristics. INTRODUCTION Lignin degradation by the white-rot fungus, Phanerochaete chrysosporium, has been studied extensively (Crawford and Crawford, 1984; Kirk, 1984; Reddy, 1984). Several recent studies have shown that H202 plays an important role in lignin degradation (Reddy and Kelley, 1985).. Glucose oxidase (GOX) has been shown to be the predominant source of H202 in lignin degrading cultures of P. chrysosporium (Reddy and Kelley, 1985; Kelley and Reddy, 1985; Reddy et al. 1983). Preliminary studies have shown that glucose oxidase—negative (ggxf) mutants of this fungus are deficient in H202 production and in lignin degradation (Ramasamy et al. 1985). Lack of H202 production by the ggx‘ mutants supports earlier evidence that glucose oxidase is the primary source of H202 in P, chrysosporium cultures grown under ligninolytic conditions. Native polyacrylamide gel electrophoresis of the extracts of wild type, gggf mutant and 395+ revertant, followed by staining of the gels for GOX activity showed the presence of a single protein band capable of glucose-dependent H202 produciton in extracts of the wild type and 33+ revertant but not in those of the ggx' mutant. Parallel gels stained for protein showed a corresponding protein band in extracts of the wild type and the gox+_revertant but not in extracts of 122 the ggg‘ mutant. We have presented here the metabolic features of a number of 321’ mutants and £21? revertants, which should be useful for future studies on the genetics of lignin degradation by P, chrysosporium. MATERIALS AND METHODS Organism and culture conditions. 2, chrysosporium (ATCC 34541) was maintained through periodic transfer on malt extract agar slants previously described (Kirk et a1. 1978). Composition and preparation of the low N medium Ff- used in this study has been described (Kirk et al. 1978). Unless described otherwise, 0.5 cm mycelial disc from 7 d- old GOX plates (see below) served as the inoculum and cultures were incubated in air without agitation at 39°C. GOX medium used for the isolation of £25“ mutants has been described (Ramasamy et a1. 1985). Glucose oxidase- positive colonies produce a purplish or brownish violet discoloration (due to oxidation of orthoanisidine) of this medium in 5 to 8 days, whereas glucose oxidase-negative colonies show no discoloration. Poly-R medium (Glenn and Gold, 1983) is the same as the Low N medium of Kirk et al. (1978) with the following amendments per 100ml: 4 g sorbose, 10 mg sodium deoxycholate and 2 g agar (Bacto-Difco). After autoclaving and cooling to about 60°C, 1 ml of poly-R 481 (Aldrich Chemical Co.; 2% filter sterilized solution) was added. Ligninolytic colonies produce a decolorized zone under and around the colony in 10-15 days whereas non-ligninolytic 123 colonies show no decolorization. Mutagenisis and isolation g; mutants. Conidia from cultures grown on malt extract agar for 7d were collected and mutagenized as previously described (Ramasamy et a1. 1985). Mutagenized conidia were plated on GOX plates to screen for ggxf mutants. This mutation procedure yielded approximately one mutant colony for every 30 colonies screened, whereas the spontaneous mutation rate was 3.4 x 10’4. The glucose oxidase-negative mutants were also screened for ligninolytic activity by plating on poly-R medium. To produce 331+ revertants, conidia from the‘ggg' mutant (COX-10) were mutagenized as described above (Ramasamy et a1. 1985). A large number of 331+ revertants were isolated. The reversion frequency was approximately one to two revertants for every 100 colonies screened. Preparation gf cell extracts and assay gf glucose oxidase activity. Cultures grown in low N medium (3 foam- plugged Erlenmeyer flasks containing 50ml each) for 6 days were used for the preparation of cell extracts as described previously (Ramasamy et al. 1985). Glucose oxidase in cell extracts was assayed spectrophotometrically by measuring the peroxidatic oxidation of orthodianisidine through a horseradish peroxidase coupled system (Reddy and Kelley, 1985). Polyacrylamide gel electrophoresis of cell extracts, employed to demonstrate protein band(s) with glucose oxidase activity was performed as previously described 124 (Reddy and Kelley, 1985). Other assays. Extracellular culture fluid from 6-day- old cultures grown in low N medium in 100% 02 was concentrated 20-fold in an Amicon ultrafiltration unit equipped with a PM-lO filter (Amicon Co., Lexington, MA), and was used for determination of peroxidase and ligninase activity. Peroxidase and laccase activities were assayed spectrophotometrically as described by Harkin and Obst (1973). Ligninase activity was assayed spectrophotometrically by monitoring the conversion of veratryl alcohol to veratrylaldehyde (Tien and Kirk, 1984). Protein was assayed by the procedure of Lowry et al. (1951) using bovine serum albumin (IV, Sigma Chemical Co., St. Louis, MO, USA) as the standard. Veratryl alcohol synthesis. Veratryl alcohol was extracted from 7-day-old cultures, grown in low N medium with 100% 02, by the procedure of Shimada et al. (1981), and quantified by HPLC on a Micropac C18 reverse phase column (Varian Associates; 30 cm X 4mm; MCH-lO) with water/methanol (1:1) as the eluting solvent at 2.0 ml/min. The chromatograph system comprised a Rheodyne model 7125 injector (Rheodyne, Inc., Berkeley, CA, USA), a Milton-Roy pump (Milton-Roy Corp., Riveria Beach, FL, USA) and a Laboratory Data Control UV detector with a 254 nm filter (model LDC III, Laboratory Data Control, Riveria Beach, FL, USA). Veratryl alcohol produced was estimated by monitoring the absorption at 254 nm. An authentic sample 125 of veratryl alcohol (Aldrich Chemical Co., Milwaukee, WI, USA) served as the standard. Ligninolytic activity and hydroxyl radical production. Assay procedures for ligninolytic activity and hydroxyl radical production, as measured by ethylene produciton from KTBA, were performed as previously described (Forney et al. 1982; Kelley and Reddy 1982). Enumeration g: conidia. Conidial numbers were determined using 8—day-old cultures grown on malt extract plates. Conidia were suspended in 15 ml of sterile distilled water and their numbers determined using a hemocytometer. RESULTS AND DISCUSSION Several hundred presumptive ggx- mutants were isolated from the GOX plates. The lack of glucose oxidase in these mutants was confirmed by the spectrophotometric glucose oxidase assay and a small number of these were used for further study (Table 1). All the ggx‘ mutants tested were deficient in ligninolytic activity. Ethylene production from KTBA (believed to be a measure of hydroxyl radical production and the ability to decolorize poly-R dyes have been shown to be associated with ligninolytic activity in Phanerochaete chrysosporium (Forney et a1. 1982; Kelley and Reddy 1982; Glenn and Gold, 1983; Kutsuki and Gold, 1982). Consistent with this, the 33x" mutants showed depressed levels of ethylene production and were unable to decolorize poly—R 481 dye on plates (Table 1). Furthermore, Y. H. Ko 126 and C. A. Reddy (unpublished data) quantified poly-R dye decolorization in liquid cultures by measuring the A513/A362 absorption ratios as described by Glenn and Gold (1983), and showed that the mutants had less than 102 of the wild type's ability to decolorize poly-R 481. Cell yields (mg dry wt. of mycelium/ml culture) of the 325’ mutants in malt extract broth or in low N medium were 83.6 to 100% of the wild type cell yield (results not shown) indicating that the deficiency in glucose oxidase and ligninolytic activities in the mutants is not due to their. poor growth. An extracellular, lignin-degrading enzyme ("ligninase") has been purified and characterized from ligninolytic cultures of P, chrysosporium (Gold et al. 1984; Tien and Kirk, 1984). Ligninase activity was absent in wild type cultures grown under non-lignin degrading conditions as well as in cultures of a non-ligninolytic mutant of this organism. The enzyme is a heme protein and has been shown to be an unique HZOZ-requiring oxygenase that catalyzes the oxidation of a variety of lignin model compounds and the partial depolymerization of spruce and birch lignins (Tien and Kirk, 1984). Peroxidase activity was shown to be intrinsic to this enzyme. It was therefore of interest to determine whether the 335’ mutants are deficient in ligninase/peroxidase activity or not. Our results showed that these mutants had little or no ligninase/peroxidase activity (Table 1). Thus, the lack of 127 ligninolytic activity in ggx‘ mutants can not be solely attributed to their deficiency in glucose oxidase activity, but to their loss of several other enzyme activities including ligninase, peroxidase and perhaps other enzyme(s) in the lignin degradation pathway. Attempts to restore lignin degradation to 325' mutants by the addition of glucose oxidase purified from P, chrysosporium (R. L. Kelley and C. A. Reddy, unpublished data) and/or ligninase (a gift from T. K. Kirk, U.S. Forest Products Laboratory, WI, USA) to a 325' mutant (COX-10) did not enhance . ligninolytic activity (Table 2). Similar results were obtained when glucose oxidase from 2, chrysosporium was replaced in the above experiment by commercially available A, £133; glucose oxidase. Lignin degradation is known to be a secondary metabolic event (Kirk, 1984; Reddy, 1984). Evidence published to date shows that nutritional and physical factors which affect lignin degradation have a parallel effect on veratryl alcohol production. Gold et al. (1982) showed that a phenol oxidase-less mutant (phe’) of'fi. chrysosporium was defective not only in its ability to degrade lignin and various lignin model compounds, but also in its ability to produce fruiting bodies and synthesize veratryl alcohol, a typical secondary metabolite, and concluded that it is a pleiotropic mutant for a set of secondary metabolic characteristics. To determine the possibility that our ggx' strains are pleiotropic mutants defective in a set of secondary metabolic characteristics, similar to the Egg“ mutants of Gold et a1. (1982), we tested the ability of these strains to produce veratryl alcohol and to conidiate. The results (Table 1) showed that all the 335‘ mutants were able to produce substantial levels of veratryl alcohol, although at levels 2 to 3 times lower than those of the wild type. Conidiation was variable among the mutants; mutants COX-4 and COX-10 produced conidial numbers comparable to those of the wild type. The other three mutants also produced high numbers of conidia although less than those of the wild type. These results indicate that the 321' mutants studied lack several metabolic activities associated with lignin degradation, but unlike the pleiotropic mutants of the type described by Gold et a1. (1982), our mutants retained some of the major secondary metabolic characteristics. A number of ggx+ revertants were isolated by subjecting conidia from COX-10 to UV mutagenesis. These revertants, regained not only their ability to produce glucose oxidase but also most of the characteristics of the wild type that were lost in the 3395" mutants. These included: the ability to degrade lignin; decolorize poly-R 481; produce ligninase, peroxidase, veratryl alcohol and conidia; and to produce ethylene from KTBA (Table 1). The above results suggest the possibility that the structural genes for glucose oxidase, ligninase, peroxidase and perhaps other lignin degrading enzymes are regulated by a common regulatory gene which is inactivated in gox' mutants 129 and is reactivated in 3251 revertants. We have recently isolated another type of mutant (Lig- 5) which had 74% of the wild type's ability to produce glucose oxidase, but only 8% of wild type's ability to produce ligninase indicating that ligninase and glucose oxidase activities can be uncoupled. This mutant showed very little ability to degrade (14C) synthetic lignin to 14C02 until day 12, but after that the strain was somewhat erratic in its ability to degrade lignin (Fig. 1). Since we observed high reversion rates.with this mutant, the variability in ligninolytic activity observed after the 12th day with the Lig—S strain may be due to the emergence of wild type revertants. In conclusion, we have characterized a number of ggx‘ mutants which can be isolated and reverted with relative ease. These mutants are deficient not only in glucose oxidase activity but also in their ability to degrade 14C- synthetic lignin to 14C02, and produce ligninase and peroxidase activities. The mutants appear to be pleiotropic for a set of secondary metabolic characteristics, but have retained others such as the ability to synthesize veratryl alcohol and to form conidiospores. The evidence suggests that the primary lesion in these mutants affects the regulation of the onset of a variety of secondary metabolic characteristics. 130 REFERENCES Crawford RL, Crawford DL (1984) Recent advances in studies of the mechanisms of micorbial degradation of lignin. Enzyme Microb Technol 6:434-442. Forney LJ, Reddy CA, Tien M, Aust SD (1982) The involvement of hydroxyl radical derived from hydrogen peroxide in lignin degradation by the white-rot fungus Phanerochaete chrysosporium. J Biol Chem 257:1455-1462. Glenn JD, Gold MH (1983) Decolorization of several polymeric dyes by the lignin degrading basidiomycete Phanerochaete chrysosporium. Appl Environ Microbiol 45:1741-1747. Gold MH, Mayfield MB, Cheng TM, Krisnangkura K, Shimada M, Enoki A, Glenn JK (1982) A Phanerochaete chrysosporium mutant defective in lignin degradation as well as several other secondary metabolic functions. Arch Microbiol 132:115-122 Gold MH, Kuwahara M, Chiu AA, Glenn JK (1984) Purification and characterization of an extracellular HZOZ-requiring diarylpropane oxygenase from the white-rot basidiomycete Phanerochaete chrysosporium. Arch Biochem Biophys 234:353- 362. Harkin JM, Obst JR (1973) Syringaldazine: an effective reagent for detecting laccase and peroxidase in fungi. Experientia 29:381-387. Kelley RL, Reddy CA (1982) Ethylene production from a-oxo—y - methylthiobutyric acid is a sensitive measure of ligninolytic activity by Phanerochaete chrysosporium. Biocem J 206:423-425 Kelley RL, Reddy CA (1986) Identification of glucose oxidase activity as the primary source of hydrogen peroxide production in ligninolytic cultures of Phanerochaete chrysosporim. Arch Microbial 144:248-253 Kirk, TK (1984) Degradation of lignin, pp 399-437. In DT Gibson (ed.) Biochemistry of microbial degradation. Marcel Dekker, New York, NY. Kirk TK, Schultz E, Connors WJ, Lorenz LF, Zeikus JG (1978) Influence of culture parameters on lignin metabolism by Phanerochaete chrysosporium. Arch Microbiol 117:277-285. Kutsuki H, Gold MH (1982) Generation of hydroxyl radical and its involvement in lignin degradation on lignin metabolism by Phanergghaete chrysosporium. Biochem Biophys Res Commun 109:320-327. 131 Lowry OH, Rosebrough NJ, Farr AL, Randall RD (1951) Protein measurement with the folin phenol reagent. J Biol Chem 193:265-275. Ramasamy K, Kelley RL, Reddy CA (1985) Lack of lignin degradation by glucose oxidase-negative mutants of Phanerochaete chrysosporium. Biochem Biophys Res Commun 131:436-441. Reddy CA (1984) Physiology and biochemistry of lignin degradation. In Klug MJ and Reddy CA (eds.), Current perspectives in microbial ecology, Am Soc Microbiol, Washington DC, pp 558-571. Reddy CA, Kelley RL (1986) The central role of hydrogen peroxide in lignin degradation by Phanerochaete chrysosporium. In CYRear C and Llewellyn GC (eds.), Biodeterioration 6, Commonwealth Argricultural Bureaux,‘ London. Reddy CA, Forney LJ, Kelley RL (1983) Involvement of hydrogen peroxide-derived hydroxyl radicals in degradation by the white-rot fungus Phanerochaete chrysosporium. p. 153+163. Higuchi T, Chang H-m. and Kirk, TK (eds:), Recent Advances in Biodegradation Research, Uni Publishers Co. LTD” Tokyo,Japan. Shimada M, Nakatatsubo F, Kirk TK, Higuchi T (1981) Biosynthesis of the secondary metabolite veratryl alcohol in relation to lignin degradation in Phanerochaete chrysosporium. Arch Microbiol 129:321-324. Tien M, Kirk TX (1984) Lignin-degrading enyzme from Phanerochaete chrysosporium: purification, characterization, and catalytic properties of a unique gEgZ-requiring oxygenase. 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