v”..- «(.‘11‘7 "F 0 Q .- 9: 1:333 £13; 0 Q hf); STUDIES ON THE DEGRADATION 0F KRAFT LIGNIN BY BACTERIA By Larry J. Forney A THESIS Submitted to Michigan State University in partial fulfillment of the requirements for the degree of MASTER OF SCIENCE Department of Microbiology and Public Health 1978 005-" 756’ ABSTRACT STUDIES ON THE DEGRADATION 0F KRAFT LIGNIN BY BACTERIA By Larry J. Forney Twenty-two bacterial strains, tentatively identified as pseudomonads, were isolated from lignin enrichments inoculated with tropical soil samples and were shown to readily degrade 4-hydroxy- benxoic acid, 4-hydroxy-3-methoxybenzoic acid, and 4-hydroxy-3,5- dimethoxybenzaldehyde. In media containing only high molecular weight (>l500 MN) kraft lignin and minerals, a mixed culture of these 22 strains attained a maximum population density of 8.7 x l07 organisms/ ml and degraded 35-38% of the kraft lignin in 52 days as shown by UV spectroscopy. The infrared spectrum of the degraded kraft lignin showed a strong absorption maximum at ~1720 cm'], as compared to that of undegraded lignin, indicating a substantial increase in car- boxylic acid/ester functional groups resulting from oxidative degrada- tion. The mixed culture did not require added glucose for the meta— bolism of kraft lignin; supplementation of the medium with 0.01% (w/v) glucose actually resulted in a decrease in the extent of lignin degradation. The results suggest that during the initial 9 days of incubation the observed microbial growth, in a medium containing kraft lignin but no minerals or glucose, is primarily due to metabol- ism of side chains with little or no metabolism of aromatic rings. To Susan who through encouragement. reassurance, patience and undying love made this work possible. ii ACKNOWLEDGMENTS I wish to express my sincere appreciation to Dr. C. A. Reddy for the guidance which he provided me throughout my graduate studies. I am very grateful to Drs. J. M. Tiedje, T. K. Kirk and M. Zabik for their critical suggestions and assistance. I appreciate Drs. M. J. Klug and F. Dazzo serving on the advisory committee. I would especi- ally like to thank my friends and colleagues for many infbrmative, helpful discussions and for their support. This work was partially funded by Competitive Minigrant No. 1300 from the Michigan State Uni- versity Agriculture Experiement Station and their support is grate- fully acknowledged. I would like to thank the Department of Micro- biology and Public Health, Michigan State University for the finan- cial assistance received. TABLE OF CONTENTS Page LIST OF TABLES ........................ v LIST OF FIGURES ....................... vi CHAPTER I INTRODUCTION ..................... l 11 LITERATURE REVIEW ................... 3 Distribution, Synthesis and Importance ........ 3 Abundance ..................... 3 Biosynthesis and Structure of Lignin ....... 3 Synthetic Lignin Polymers ............. l2 Lignins Within Plant Tissues ........... 12 Utilization of Lignocellulosic Materials ..... l4 Lignin in the Environment ............. l5 Biological Degradation of Lignin ........... 16 Fungal Degradation of Lignin ........... l6 Role of Polysaccharides in Lignin Degradation by Fungi .................... 25 Role of Phenol Oxidases in Lignin Degradation . . . 26 Bacterial Degradation of Lignin .......... 29 Footnotes ....................... 37 Appendix ....................... 39 Literature Cited ................... 40 III MATERIALS AND METHODS ................. 50 IV RESULTS ........................ 62 V DISCUSSION ...................... 72 VI SUMMARY ........................ LITERATURE CITED ....................... 77 iv LIST OF TABLES Table ’ Page l. Elemental and functional group analyses of milled wood lignin, undegraded and degraded by Polyporus versi- color .......................... l8 2. Source and type of soil samples used as inocula fOr lignin enrichment cultures and the number of bacterial strains isolated from each culture ........... El 3. Composition of the media used fbr studying kraft lignin degradation by a mixed culture isolated from tropical 58 soils .......................... 4. Characteristics of bacteria isolated from tropical soils and selected for study of kraft lignin degrada- tion .......................... 64 5. Extent of kraft lignin degradation by a mixed culture in three different media .................. 68 LIST OF FIGURES Figure Page l. Structures of (a) p-coumaryl, (b) coniferyl, and (c) sinapyl alcohols .................. 5 2. Structures of the mesomeric forms of the free radical derived from the ion of coniferyl alcohol ....... 8 3. Schematic formula for a representative portion of a spruce lignin ..................... l0 4. A scheme for the enzymic degradation of the guaiacylglycerol-B-coniferyl ether units ........ 22 5. Structures of certain aromatic compounds discussed in the literature review ................. 39 6. A typical ultraviolet spectrum of undegraded and de- graded kraft lignin .................. 60 7. (a-d) Viable bacterial counts in various kraft lignin (MN > 1500) media ................... 55 8. (a-b) Infrared spectrum of degraded and undegraded kraft lignin ...................... 70 vi CHAPTER I INTRODUCTION Lignin constitutes up to 30% of the dry weight of all vascular plants (Sarkanen and Hergert, l97l) and, next to cellulose, is the most abundant organic polymer in nature. Lignin degradation represents one of the most important steps in the total biological deterioration of plant materials or of industrial byproducts from lignocellulosic materials. Kraft lignin is produced during the delignification of wood by the alkaline pulping process (Marton, l97l) and resembles lignin j__situ in that it is a complex, three dimensional aromatic polymer (Lai and Sarkanen, 1971). However, during the pulping pro— cess the three-carbon side chains of the polymer undergo a variety of reactions including a partial loss of the y carbons, cleavage of aryl- alkyl ether linkages between phenylpropane units resulting in depoly- merization and liberation of new phenolic units, and limited demethy- lation of methoxyl groups (Lundquist et al., l977; Marton, l97l). The ability of certain fungi to degrade lignins, including kraft lignin, has been clearly established (Kirk, l97l; Kirk et al., l975; Lundquist et al., 1977), but little is known about the role of bacteria in the decomposition of lignins in nature. Trojanowski et al. (l977) demonstrated that approximately 4% of a (ring-14C) coniferyl alcohol dehydropolymer was released as 14C02 during TS days of incuba- tion with a strain of Norcardia. Most recently, Crawford (1978) 2 reported that 3% of (U-14C)-fir lignocellulose was degraced by cer- tain strains of Streptomyces. Odier and Monties (l977) isolated 18 strains of aerobic bacteria which were capable of utilizing acidolysis lignin, isolated from wheat straw, as a sole carbon source. Of these strains, a strain of Xanthomonas was shown to be the most effective in utilizing 71% of the lignin carbon within 7 days. The severe struc- tural modifications that lignin undergoes during the acidolysis ex- traction process (Lai and Sarkanen, 1971) may partially explain this unusually rapid rate of degradation. Other investigators suggested that certain strains of Micrococcus (Jaschof, 1964), Arthrobacter (Cartwright and Holdom, 1973), Flavobacterium (Odier and Monties, 1977)l Sorensen, 1962), Aeromonas (Odier and Monties, 1977), and Pseudomonas (Ferm and Nilsson, 1969; Sorensen, 1962) as well as bac- terial mixed cultures (Sundman et al., 1968) are involved in lignin decomposition, but conclusive evidence is lacking. To the best of our knowledge, no definitive studies have been conducted on the degradation of kraft lignin by bacteria. Considering that literally millions of tons of kraft lignin are generated annually in the United States, it appeared important to study its utilization and/or degradation by bacteria. Presented here are the results showing the degradation of a high molecular weight fraction of kraft lignin by a mixed culture of twenty-two strains of Pseudomonas isolated from tropical soils. CHAPTER II LITERATURE REVIEW Distribution, Synthesis and Importance of Lignin Abundance Lignins rank among the most abundant and widely distributed organic polymers in nature, being exceeded in abundance only by cel- lulose, and constitute 20-30% of the dry weight of all vascular plants 10 (55, 103). It has been estimated that 1.3 x 10 metric tons of lig- nin carbon are biosynthesized annually (81). In addition, 1 x 1012 to 3 x 1012 metric tons of carbon is estimated to be present in the organic matter of soil, primarily as peat and humus which are in large part derived from lignins (60, 95, 119). Obviously, lignins represent a key intermediate in the carbon cycle in the biosphere. Biosynthesis and Structure of Lignin The structure of lignin is best understood by retracing the steps involved in its biosynthesis. The phenylprOpanoid amino acids, tyrosine and phenylalanine, are synthesized via the shikimic acid pathway from carbohydrates formed during photosynthesis (55, 59). Shown in Figure 1 are the cinnamyl alcohol derivatives, p-coumaryl, coniferyl, and sinapyl alcohols, which in turn are derived from the phenylpropanoid amino acids (55, 104). Within the plant tissues free phenoxy radicals are formed from any of the above three cinnamyl Figure 1. Structures of (a) p-coumaryl, (b) coniferyl, and (c) sin- apyl alcohols which are direct precursors of lignin. OI Iuo goo @ UI IU 18 I I O U---U Figure 1. 6 alcohol derivatives, either through the removal of a hydrogen from the phenolic hydroxyl group or a one electron transfer from the phenolate ions by a phenol oxidase enzyme such as laccase (EC 1.10.3.2; p-di- phenol:02 oxidoreductase) or peroxidase (EC 1.11.1.7; donor:hydrogen peroxide oxidoreductase) (8, 55). Due to the extended pi electron system of the molecules, the various free radicals thus fOrmed are stabilized through equilibrium with various mesomeric forms as exem- plified by the radicalization of coniferyl alcohol shown in Figure 2 (55). Radical forms analogous to 2a are referred to as o- and p- methylene quinonoid radicals, respectively. Radicals analogous to 2c are designated as p-quinone methide radicals and are relatively more stable than the other radical forms described above. Polymerization occurs when the free radicals derived from p- coumaryl, coniferyl, and sinapyl alcohols undergo condensation reac- tions to become covalently linked (55, 104). The relative abundance of each potential bonding arrangement is dictated by the relative concentration and stability of the different free radical species (55). Phenolic hydroxyl groups in the growing lignin polymer are oxidized continuously with the formation of free radicals and these form co- valent bonds with the incoming monomer radicals supplied by the bio- synthetic apparatus of the cell. The net result of these reactions is a three dimensional, amorphous, highly branched polymer, repre- sented schematically by the formula for spruce lignin shown in Figure 3, originally proposed by Freudenberg (37), and as modified by Harkin (55). Sarkanen and Ludwig (103) suggest that spruce lignin is repre- sentative of other gymnosperm lignins in general. The structure shown represents an average fragment of a larger lignin molecule and shows some of the types of monomer units and their linkages that are Figure 2. Structures of the mesomeric forms of the free radical de- rived from the ion coniferyl alcohol. (a) phenoxyl rad- ical, (b) o-methylene quinonoid radical, (c) p-methylene quinonoid radical, (d) pequinone methide radical. .020u.< {.2500 a nu :00 I © 8 o... __ :o _ zoom: Figure 2. Figure 3. Schematic formula for a representative portion of a spruce lignin. 10 OCH HZCOH [- 3 “£0” a CO HCO _ 2 HZC'OH H2COH H3CO o —?H a ’C we -0 ”c I w3co "2?°” (r-—-cH wc 0‘5 ' . 2 a . . H2$m 0CH3 . HC--O \ ' * ocw u H ,o. 1‘ ° 3 3 wcolc6 mos)" wzc 3:" o 2:" on I wc-cw “é._.o rs 1 .1 HC JCHZ O ,QD "29°" N c ~cw wzcow 0‘ 3 O on u OCH OCH -cw 3 0—04 wzclon 3 on HCOH HC—O ”2‘30" OCH: I" we 0 29°” 1 coa cu H3 OCH HCOH u \ o no . I I \OCH \OCH H CO_O OH 3 -O 3 3 Figure 3. 11 presently known to occur in lignin. Freudenberg has isolated and identified at least 27 different intermediate products formed during the jn_vitrg_polymerization of coniferyl alcohol catalyzed by horse— radish peroxidase (35) and thus the actual distribution of inter- monomer bondings may be much wider than is shown in Figure 3 (102). Of the intermonomer linkages shown some are known to occur with rela- tively higher frequency than others. For example, it has been es- timated that approximately 40% of the linkages in spruce wood lignin are B-aryl ether bonds (35, 36) such as those occuring between units 1 and 4, 4 and 5, 5 and 6, 7 and 8 in Figure 3. Such B-aryl ether linkages are formed by the coupling of two coniferyl alcohol radicals; one a p—quinone methide radical and the second a phenoxyl radical (55). Phenylcoumaran type linkages (between units 17 and 18, Fig. 3), which reportedly constitute about 20% of the linkages in spruce lig- nin (2), are formed through the coupling of two coniferyl alcohol radicals, a p-quinone methide radical and a second o-methylene quin- onoid radical followed by rearrangement and a reaction involving inter- molecular nucleophilic attack (55). The combination of two p-quinone methide radicals of coniferyl alcohol to form a double quinone methide followed by double ring closure results in the formation of a D,L- pinoresinol type of structure as shown between units 8 and 9 in Figure 3 (55). Approximately 10-12% of the bonds in spruce lignin are reported to be of this type (35, 36). Biphenyl linkages (units 12 and 13, Fig. 3) may constitute up to 25% of the intermonomer link- ages in conifer lignin and are formed by the coupling of two o-methy- lene quinonoid radicals of coniferyl alcohol (99). Intercombination of monolignol radicals derived from p-coumaryl, coniferyl, and sinapyl 12 alcohols increases the structural complexity of the lignin polymer many fold. Synthetic Lignin Polymers Synthetic lignin polymers ("dehydrogenative polymerizates"; "DHPs") can be generated through the polymerization of p-hydroxycin- namyl alcohol derivatives (79). Recently, Kirk gt_al, have published a procedure describing the synthesis of DHP using specifically labelled 14C-coniferyl alcohol. Kirk gt_al, and earlier workers (38, 79) have shown DHP to contain many of the structural features of natural lignin. 14C—DHP was shown by Kirt gt a1, (79) to have an average MW of 1545 corresponding to 8.1 C6C3 units per polymer and to contain less than 1% of material of MW less than 750. The DHP contained C, 63.2%; H, 5.8% and 0CH3, 16.3%, whereas spruce lignin contained C, 62.9%; H, 6.1%; and 0CH3, 15.1%. Since synthetic lignins have been shown to be polymeric and to contain essentially the same structural units as natural lignins they are considered a suitable model for use in struc- tural and biodegradation studies. Lignins Within Plant Tissues Coniferyl, p-coumaryl and sinapyl alcohols are present in dif- ferent ratios in lignins of different phylogenetic origin and often within different tissues of the same plant species (55, 104). Lig- nins within deciduous trees (syringyl lignins) are formed from nearly equal proportions of sinapyl and coniferyl alcohols with only about 5% of the subunits being derived from p-coumaryl alcohol and contain 1.20-1.59 methoxyl groups per 09] (37, 103). Conifer lignins (guaiacyl lignins), such as pine or spruce lignins, are biosynthesized from l3 primarily coniferyl alcohol (approximately 85% of the subunits) with» lower levels of p-coumaryl and sinapyl alcohols and contain 0.87 to 1.00 methoxyl groups per 09 (34, 37). Lignins within mosses are formed almost exclusively from p-coumaryl alcohol (37, 103). Within plant tissues, lignins occur as integral cell wall com- ponents and in the middle lamellae (59, 103) imparting structural rigidity and resistance to impact, compression and bending (102). Ultraviolet microscopy of thin sections of plant tissues reveals that 72-81% of the lignin is located within the secondary cell wass layers, and 19-28% is present in the middle lamellae (30). Whereas the middle lamellae consists largely of lignin (approximately 72%), 45-50% of the secondary cell wall is polysaccharides (117). Therefore, the largest portion of lignin in cell walls occurs in close physical association with and is probably chemically bonded to celluloses and hemicelluloses (32, 55, 61, 117). Based upon theoretical considerations the follow- ing bonding arrangements have been proposed between lignin and plant polysaccharides: a) hydrogen bonding, b) ether bonds between the alpha carbon of the side chain and the hydroxyl groups of the poly- saccharide subunits, c) ester bonds between alpha carbon and the gul- curonic acid residues, d) carbon-carbon and carbon-oxygen bonds which result from radical coupling, and e) phenyl-glucosidic bonds (32, 33, 83). Physically, lignins, in conjunction with hemicelluloses, sur- round cellulose microfibrils in a sheath-like manner although the ex- act nature of this assOciation is unclear at this time (59, 99). 14 Utilization of Lignicellulosic Materials A growing awareness of the need to utilize our natural re- sources efficiently and the search for alternative sources of food and energy has generated a new interest in the exploitation of ligno- cellulosic waste materials, one of the largest renewable resources on earth. 1.0 x 109 metric tons of such material are generated each year (13). Current technology would allow for the biological produc- tion of ethanol (25, 90), butanol (87), methane (87), ethylene (14, 44, 115), butadiene(44), fatty acids (13), levulinic acid (44), and single cell protein (25, 26, 67) among others from cellulose and hemi- celluloses found in lignocellulosic wastes. However, the intimate relationship of lignin and plant polysaccharides described above pre- sents both a physical and chemical barrier to enzymes capable of hy— drolyzing cellulose or hemicelluloses and thus limits their utiliza- tion (73, 100, 110, 114). It has been found that any treatment which depolymerizes, solubilizes, or otherwise removes lignin increases the availability and susceptibility of cellulose and hemicelluloses to enzymatic or chemical attack (16, 17, 87, 91, 115). Physical deligni- fication procedures include: irradiation with gamma rays or high en- ergy electrons (84), comminution (28, 93, 98), photolysis with ultra- violet light (42), and treatment at elevated temperature and/or pres- sure (92). The aforementioned procedures, though generally effective, are energy intensive and uneconomical (76). Chemical delignification procedures include: bleaching agents such as HClO (29), alcoholysis with dilute mineral acids (116), extraction with organic solvents to which mineral acids have been added (116), thioglycolic acid pulping (116), alkaline pulping (43), oxygenalkali pulping (12), soaking with 15 dilute alkaline solutions (53, 91, 100), or peracetic acid (93), and oxidation with hydrogen peroxide (93). These procedures are not feasible for many applications due to the large quantities of chemical wastes which must be efficiently recycled or disposed of (56, 76). A microbial delignification process would potentially have many advantages including low cost, no chemical wastes, high specificity, and the conservation of lignin carbon in the form of microbial cells. Lignin in the Environment Lignins found in the wastewater of pulp and paper plants have been shown to be environmental pollutants (6, 41, 48, 82). Griffin and West (48) have shown that 0.89% (v/v) sulfite waste liquor (SWL) in water results in 50% mortality (L050) among rainbow trout. 0f the constituents of SWL examined, lignin content was found to be the most highly correlated with LC50 (r = 0.96). The mean lignin concentration of SWL was 0.92 g/l. Thus even at low concentration SWL may be detri- mental to the environment. Little is understood about the decomposition of plant material, especially lignins, within the soil and the concomitant formation of humus (so, 89). Plant tissues, including lignins, are subject to microbial attack in the soil. Hacket §t_al,(49) estimated the rates of lignin degradation in various soils using (14C-ring) DHP. The ob- served rates, after 30 days of incubation, varied from no detectable degradation with a sample from Costa Rica to 34% of the label being recovered as 14 002 with a sample from Yellowstone National Park. Most soils examined showed 4-l3% degredation of the added lignin during a 30 day incubation period. Similarly, Crawford gt_al, (19) observed 16 5-40% degradation of oak and maple (140)-1ignocellulose when incubated with various soil samples. Haider gt_a1, (53) have studied the decom- 14C-DHP in soil and have estimated position of (14C-1ignin) maize and their total turnover time to be 20 and 30 years, respectively. These data suggested that the rates of degradation of the lignin components of lignocelluloses are generally higher than those observed using DHP, probably because in the former the lignin is complexed with poly- saccharides which may, by acting as a substrate and/or source of en- ergy, speed lignin degradation (24). The highly oxidized, condensed, lignin residues, which result from microbial degradation, complex with proteins, carbohydrates, minerals and phenolic compounds to form humus (50, 60, 89, 95). Haider gt_al, (52) linked (‘4C-ring) coumaryl alcohol and protocatechuic acid into a model humic polymer, determined their rates of degradation in soil, and calculated the total turnover time to be 326 and 76 years respectively. In contrast, the total turn- over time fbr (U-14C) glucose was calculated to be 8 years. This difference in turnover rates clearly shows that humic materials are degraded very slowly within soil. In fact, they are believed to be precursors of bituminous coal (l8) and have recently been found in silicified wood (106). Biological Degradation of Lignin Fungal Degradation of Lignin It has long been known that certain fungi, mainly white, brown, and soft rot fungi, are important in degradation of wood and leaf litter in nature (47). However, these groups differ in their mode and extent of degradation (72, 105). Several hundred species of 17 Hymenomycetes (e.g. Agaricaceae, Corticaceae, Hydenaceae, Polypor- aceae, and Thelephoraceae), commonly known as white rot fungi, are capable of effecting extensive degradation of all major components of wood (74, 75) including as much as 97% of the lignin (l5). Cer- tain holobasidiomycetes, classified as brown rot fungi, utilize large portions of the cellulose and hemicelluloses of wood (75). Although brown rot fungi were shown to be incapable of degrading lignin, they are believed to effect limited structural modifications in the polymer (72, 75). Soft rot fungi, which include certain ascomycetes and Fungi Imperfecti, are primarily cellulolytic (51, 86); however, some reports indicate that tney may degrade up to 45% of the lignin in wood (86). Certain other soil fungi including Alternaria and Peni- cillium reportedly degrade as much as 20% of the lignin within wheat straw (113). The degradation of lignin by Polyporus versicolor (Polystictus versicolor, Coriolus versicolor), a typical white rot fungus, has been extensively studies by Ishikawa gt_al, (61) and Kirk and Chang (77, 78). Ishikawa gt_al, inoculated P, versicolor into a medium contain- ing spruce milled wood lignin (MWL)2 and incubated for 28 days. They reported a C9 formula of C9H8 6102 93(0CH3)0 76 for the degraded lignin and C9H8.7802.78(0CH3)0.97 for the MWL control. In comparison, Kirk and Chang isolated the degraded lignin from spruce wood after incuba- tion for 45 d. and determined the C9 formula of the degraded lignin t° be C9H7.2603.95(OCH3)0.74 as C°mpaVEd t° C9H8.8602.75(0CH3)0.92 fOr the MWL control (see Table l). Ishikawa gt_al, observed a 21% decrease in the methoxyl content of degraded lignin while Kirk and Chang observed a 19% decrease in the methoxyl content. These results 18 op.o m¢._ 8A.oflm:oovmm.~o_o.m=mu mm cucuumuo mo.o om._ NQOAMESwZomgzmu o ememcmuucs A_o .cnsv nflm.mm uzmxwgmH Po.o Km.o uk.ofim:oovmm.momm.hzmu mu unnmcmuo mo.o os._ mm.ofim=uovm~.momm.mxmu o uncmsmnuca Am“ .Luuv mango use xcwx Am0\mmpoE~ Amu\mmpoev mp:ELow mu Amxmnv Fxxongmu pzxocuxg map» pmpop Page? Paupvcasm cowpmmaucm Lopoupmsm> mssomxhoa x5 vmumsmmn can umumcmcocca .cwcmwF too; umFPPE mo mmmxpocm azosm chowpucam new Popcmsmpm .P mpnmh 19 suggest that approximately 20% of the methoxyl groups are preferen- tially degraded during the first 28 days of incubation without a sub- stantial number of additional groups being degraded upon prolonged incubation. Ishikawa gt_al, found that during degradation the total hydroxyl content of lignin increased from 1.20 to 1.48 per C9 and the carboxyl content increased from 0.03 to 0.098 per Cg. In contrast, Kirk and Chang found that during degradation the total hydroxyl con- tent of lignin decreased from 1.16 to 0.87 per C9 and the carboxyl group content increase was more than six fold that observed by Ishi- kawa gt_al, These data suggest that during the initial stages of lignin degradation the aromatic nuclei within lignin are hydroxylated and upon further incubation are cleaved to produce an increased number of carboxyl groups (dicarboxylic acid structures are common products of ring cleavage; 26) with a concomitant decrease in hydroxyl content. In both studies the increase in oxygen atoms per C9 was supported by the increase in oxygen containing functional groups observed. Ishikawa gt_al, (61) found dehydrovanillin, vanillic acid, van- illin, p-hydroxycinnamic acid, p-hydroxycinnamaldehyde, and conifer- aldehyde to accumulate in the medium during the degradation of lignin by E, versicolor (see appendix). These workers suggested that initial action of extracellular enzymes produced by white rot fungi on spruce lignins results in the formation of water-soluble metabolites. .The C603 fragments were thought to be further degraded by oxidative short- ening of the side chain and destruction of the aromatic nuclei. In further studies Ishikawa et_al, (62) examined the degradation by P, versicolor and an Forme forentarius of the compounds listed above. Guaiacylglycerol-B-guaiacyl ether was shown to be metabolized to 20 guaiacylglycerol and guaiacol. This suggested that the fungi could degrade guaiacylglycerol units containing B-aryl ether linkages.' When incubated with guaiacylglycerol, 4-hydroxy-3-methoxypheny1pyruvic acid was produced, and it was shown that this product could be further metabolized to vanillin, vanillic acid, and dehydrovanillin. Coniferyl alcohol was shown to be readily oxidized to coniferaldehyde and ferulic acid which could be further metabolized through shortening of the side chain to vanillin, vanillic acid, and dehydrovanillin. Similar re- sults were obtained on incubation of these model compounds with lac- case preparations and horeradish peroxidase but not from commercially available tyrosinase. These data suggested that guaiacylglycerol-B- ether units within lignin were metabolized to vanillin, vanillic acid and dehydrovanillin via various intermediates by extracellular enzymes (presumably by phenol oxidases) as shown in Figure 4. This conclusion was further supported by a decreased vanillin yield upon nitrobenzene oxidation of the degraded lignin indicating a depletion of units con- taining a 4-hydroxy-3-methoxy substitution pattern. The general mech- anism of degradation of lignin by white rot fungi was thought to in- volve sequential cleavage of 09 units from the polymer. Kirk and Change (78) proposed that P, versicolor was able to degrade the aromatic rings of phenyl propanoid units while they are still bound to the polymer. These investigators observed a strong 1 in the infrared Spectrum of the de- absorption band near 1720 cm- graded Spruce lignin, which was not diminished upon reduction with NaBH4, which indicated an increase in carboxylic acid and/or ester groups. The presence of carboxyl groups was further supported by a broad hydroxyl absorption between_2900-3300 cm'] in the IR spectrum. 21 Figure 4. A scheme for the enzymic degradation of the guaiacyl~ glycerol-B-coniferyl ether units present in softwood lignin by white rot fungi. (Ishikawa gt_al,, Arch. Biochem. Biophys. 100:140-149) 22 moon cnzcn—cwpn “clog "con ".90" ucl—o O cn:cn—cn,ou HC—OR' OCH ’ 11,0 “,0 /k OCH, 1 OCH, 1 ’ OCH, OR. R'.R”:" or C" H 1 O" Guahcylglyceml-J- Gualacylclycerol-li- Gualacylglyceml coniferyl ether units coniferyl other 1 , l-Hp _ WQCHZCfl—Cflpfl :33)" :33" ”f: —1 101 0C". aca ,.____, "U Confleryl alcohol "—"‘_’ ecu. OCH. 0C". [o] 0!! I L. I _‘ tote-torn Incl-tor. .i-IlydroxyconUerl Fantasy-twig mum-wk acld alcnlml MQCfli’CH—Cflo _1 ll. Confleraldehyde 101 [all 1101 ”gnaw... J “W... Ferullc acid mow 0C". Oil OH OH Vanlllyl alcohol Dehydrodlvanlllln Figure 4. 23 The IR spectrum of methylated degraded lignin showed a methylester carbonyl stretch near 1723 cm.1 indicating that the bulk of the car- boxylic acid groups were conjugated as 0,8 unsaturated and/or adjacent to alpha keto groups. Since the carbonyl absorption band in the IR spectrum was not noticably shifted to the nonconjugated region (a higher wave number) upon reduction, it was unlikely that significant amounts of a keto groups were present. Therefore, most of the car- boxylic acid groups were apparently conjugated as 0,8 unsaturated moieties. If the a,8 unsaturated carboxylic acid groups were derived from the side chains of intact aromatic nuclei, the presence of cinn- amic acid residues would have been observed due to their strong UV absorbance near 330 nm, this was not the case. Furthermore, there was a decrease in the number of aromatic rings within the degraded lignin as evidenced by the decreased absorbance at 1515 cm'] in the IR spectrum relative to other absorption bands, and a decreased pro- portion of aromatic protons as seen in the proton magnetic resonance spectrum. In addition to the observed decrease in the total phenolic hydroxyl content as determined by acetylation, oxidative degradation of the degraded lignin revealed an absence Of o-diphenolic units. I Also, previous studies (26) have shown that the oxidation of low mol- ecular weight aromatic compounds by microorganisms may yieldmetabolic products containing a,8 unsaturated carboxylic acid groups. From these results Kirk and Chang proposed that white rot fungi attack and cleave the aromatic nuclei while bound within the polymer thus resulting in an increased number of a,8 unsaturated carboxylic acid groups. This degradative mechanism is in contrast to the sequential cleavage of C9 subunits proposed by Ishikawa et a1, (61). However, 24 differences in the mode of attack proposed as well as functional group analyses may reflect differences in experimental design. Kirk and Chang (77, 78) extracted the degraded lignin from wood after incuba- tion whereas Ishikawa gt_al, extracted the lignin from wood prior to incubation. Recently, procedures for the labelling of lignin in plant tis- 14C-labelled phenylalanine or sues through the administration of ferulic acid (20) and, as mentioned above, for the synthesis of spec- ifically labelled DHP have been developed (79). The assay of lignin degradation using radioactively labelled substrate is both specific and sensitive. Kirk gt_al, (80) studied the degradation of specif- ically labelled side chain-14C, methoxyl-14C, and ring-14C DHP by Coriolus versicolor, Phanerochaete chrysosporium, Gloephyllum trabeum, Poria cocos and in a fOrest soil. After 600 h of incubation with the white rot fungus Phanerochaete chrysosporium, 33% of the total meth- 14 14 was recovered as 14 14 oxyl-C C02 whereas only 20% of the side chain-C was recovered as 14 and 15% of the ring-C 002. The corresponding values for 9, trabeum, a brown rot fungus, were 8.5 and 2% respectively. These results supported earlier reports (9, 71, 75) that brown rot fungi are capable of limited demethylation of methoxyl groups in lignin, but show little or no capacity to degrade the lignin polymer. When specifically labelled DHP was incubated for 600 h with a forest soil, the extent of degradation was markedly less than that observed with P, chrysosporium. Only 3% of the 14C-ring labelled DHP was re- covered as 14002. Furthermore, in contrast to the pure culture studies, the side chains of the DHP were degraded more rapidly than the methoxyl groups. The rates of degradation obtained in this study 25 should be considered to be minimal rates since the assay measures only evolved 14 CO2 and does not measure lignin carbon which has been incorporated into cellular material, or which has undergone transform- ation into humus, or fragmentation of the polymer without further metabolism (79). Despite these studies, an integrated scheme of lignin degrada- I tion by fungi can not yet be proposed. Role of Polysaccharides in Lignin Degradation by Fungi The rate of lignin removal (based upon the percent of original present) by E, versicolor has been shown to be roughly proportional to the rate of removal of glucan, mannan, and xylan (75). However, since wood contains roughly twice as much polysaccharides as lignin, the actual amount of polysaccharide metabolized (by weight) is much greater than the amount of lignin degraded per unit time. It can be stated that, in general, lignin degradation by white rot fungi is accompanied by a substantial decrease in the carbohydrate content of the plant tissues being examined. Earlier workers have reported that certain white rot fungi are able to utilize lignin as a sole carbon source (46, 47, 97, 112). How- ever, Kirk gt_al, (80) were unable to demonstrate the degradation of 14C-ring DHP when the medium was not supplemented with carbohydrate 14 but showed 6.3-19.7% of the lignin being degraded to CO after 55 2 d.in the presence of cellulose, D-xylose, D-glucose, or D-cellobiose. The extent and rate of lignin degradation was shown to increase with increasing cellulose concentration. 26 Role of Phenol Oxidases in Lignin Degradation Earlier workers established a strong correlation between the ability of white rot fungi to produce phenol oxidases and their abil- ity to extensively degrade lignin (27, 61, 62, 68, 69, 72).3’4 It was generally believed that phenol-oxidizing enzymes, which are pres- ent in and presumably released from the hyphae of white rot fungi as they penetrate wood tissue, catalyze the oxidation of accessible phen- olic hydroxyl groups within lignin. The unstable free radicals thus produced become stabilized by reactions that condense some of the lignin structureslnxtsimultaneously result in the cleavage of bonds between phenylpropanoid units and the remaining polymer. Subsequent removal of side chains on the polymer exposes additional phenolic groups fOr the continued degradative enzymatic oxidation. Theoretic- ally, the overall effect of such processes would be fragmentation of some parts of lignin and condensation of the remainder. This pro- posal has been supported by several observations. First, several workers using electron spin resonance spectroscopy have observed an increased spin content in degraded lignin (ll, 57). This increase is thought to be due to "trapped" or stabilized free radicals within the degraded lignin. Such free radicals, it is hypothesized, are a result of the action of phenol oxidases. Secondly, lignin degradation re- sults in a highly oxidized condensed residue, which as a part of humus, is very resistant to further degradation (18, 50). Third, Ander and Eriksson (1) isolated a mutant of Sporotrichum pulverulentum, desig- nated Phe 3, incapable of synthesizing phenol oxidase. Phe 3 was shown to be unable to degrade kraft lignin. However, when incubated with purified laccase Phe 3 could degrade 20% of the kraft lignin 27 (kL)5 in 30 days whereas the wild type strain degraded 24% of the KL in the same period. These data were taken by these investigators to be strong evidence for an obligatory role of phenol oxidases in lignin degradation. Fourth, Ishikawa gt_al, (61) studied the degradation of lignin from conifers by the following fungi: Pglyporus versicolor, Polyporus hirsutus, two strains of Poria subacida, Fomes fomentarius, Fomes annosus, and Trameses pini. One strain of P, subacida and the latter three fungi showed weak phenol oxidase activity, whereas the remaining fungi showed strong phenol oxidase activity. When incubated for 13 d in a medium containing either native lignin (NL)6 or MWL, phenol oxidase-rich fungi grew faster and degraded 32-48% of the lige nin, whereas phenol oxidaselxxn~fungi grew poorly and degraded 18-25% of the lignin. These dats suggested that the ability of fungi to degrade lignin was correlated with the activity and/or level of phenol oxidase enzymes. Finally, many investigators have demonstrated the oxidation of lignin and certain lignin metabolites by phenol oxidases (39, 40, 61, 85, 101). For example, Leonowicz and Trojanowski (85) were able to demonstrate the cleavage of guaiacylglycerol-B-coniferyl ether to coniferyl alcohol and the corresponding ortho-quinone by purified laccase. Culture filtrates of Stereum frustulatum and P, versicolor have been shown to catalyze the cleavage fo the alkyl- phenyl ether linkages in the model compound syringylglycol-B-guaiacy1 ether to form guaiacoxyacetaldehyde, guaiacoxyacetic acid and 2,6- dimethoxy—p-benzoquinone (69, 70; see appendix). Similar types of cleavage products were produced upon incubation of the above compound with purified laccase suggesting that phenol oxidases produced by S, frustulatum and P, versicolor were involved in the cleavage of the 28 alkyl-phenyl ether linkage by the fungi. It has been proposed by Kirk gt 91, and other authors (61, 62, 69), that the oxidation of phenolic subunits in the lignin polymer by phenol oxidases followed by various rearrangements in “reverse synthesis" reactions may be im- portant in the depolymerization of lignin by white rot fungi. Con- sidering the diversity of linkages which occur in lignin, the produc- tion by an organism of relatively nonspecific enzymes capable of effect- ing the depolymerization of lignin would be energetically less costly and would therefore provide the organism a selective advantage. The contention that phenol oxidases have an integral role in the degradation of lignin has been questioned since not all white rot fungi produce phenol oxidase enzymes (27). Of 210 species of white and brown rot fungi examined (27), 96% of the white rot fungi and 20% of the brown rot fungi tested gave positive Bavendamm tests. This work was later corroborated by Kirk and Kelman (68). Therefore, since many of the brown and soft rot fungi, which have little or no lignin degrading ability gave positive Bavendamm tests, the presence or absence of phenol oxidases does not necessarily indicate an organ- ism's ability or inability to extensively degrade lignin. Several other functions of phenol oxidases have been suggested. Laccase may function as a part of an extracellular electron transport system. Sporotrichum pulverulentum, a white rot fungus, has been shown to produce an extracellular cellobiose: quinone oxidoreductase (cellobiose dehydrogenase) which catalyzed the oxidation of cellobiose to collobiono-d-lactone using any of several quinones as electron acceptors (118). It was proposed that in nature the quinones which participate in this reaction may be derived through the oxidation of 29 phenolic compounds by phenol oxidases to the corresponding quinones (118). A laccase type enzyme produced by Rhizoctonia praticola has been shown to catalyze the polymerization of four subunits of phenolic intermediates of herbicide degradation (107). Similarly, Kirk gt_al, (69) isolated a dimer of guaiacylglycerol~8-conifery1 ether from a culture of Polyporus versicolor containing the monomer as a substrate. Ishikawa fl al_. (62) found 3. versicolor capable of polymerizing all of the following compounds apparently by the activity of an extracel- lular phenol oxidase: p-hydroxycinnamic acid, p-hydroxyphenyl pyruvic acid, ferulic acid, 4-hydroxy-3-methoxypheny1 pyruvic acid, conifer- aldehyde, coniferyl alcohol, isoeugenol, and guaiacylglycerol (see appendix). Although this phenomenon has not bee investigated in de- tail with respect to lignin degradation, one could hypothesize that phenol oxidases serve to detoxify the environment of the fungus through the polymerization of potentially toxic products of lignin degradation. It is also possible that phenol oxidases serve any or all of the func- tions mentioned above. Clearly, further work is necessary before an understanding of the role of phenol oxidases in lignin degradation can be obtained. Bacterial Degradation of Lignin Certain bacteria have been shown to be capable of metabolizing a wide range of aromatic compounds (26), many of which are inter- mediates in the breakdown of lignin by white rot fungi, or which have ring substitution patterns similar to those found in lignin and lignin precursors. Crawford gt_al, (22) isolated a strain of Pseudomonas acidovorans which metabolized l-(3,4—dimethoxyphenyl)-2- 3O (o-methoxyphenyl)-1,3-propanediol, with the concomitant accumulation of guaiacol. After incubation with the dilignol mentioned above, the isolate utilized vanillic acid without an observable lag period. Thus based on the theory of simultaneous adaptation, vanillic acid was preposed to be in the pathway for the metabolism of 1-(3,4-dimeth- oxyphenyl)-2—(o-methoxyphenyl)-1,3-propanediol. The initial cleavage was thought to occur between the a and B carbons of 3-hydroxy-l-(4- hydroxy-3-methoxypheny1)-2-(o-methoxyphenoxy)-l-propanone which had been formed through the demethoxylation and oxidation of the a carbon to a carbonyl of the parent compound. The resultant ether was fur- ther metabolized to guaiacol (see appendix). Catechol oxidase activ- ity was shown to be present and presumably was involved in the fur- ther metabolism of guaiacol while protoacatechuate-4,5-dioxygenase, also produced by this organism, was thought to cleave the ring struc- ture of vanillic acid. This was the first reported metabolism of a compound containing a B-aryl ether linkage by a bacterium. Such linkages, as mentioned earlier, constitute a major portion of the linkages in lignin. Crawford et_al, (23) also demonstrated the mutualistic degra- dation of l-(3,4-dimethoxyphenyl)-2-(o-methoxyphenyl)-l,3-propanediol by strains of Nocardia corallina and Acinetobacter sp, When N, 99:: allina was incubated in a medium containing the above compound, guaiacol accumulated in the medium and was found to be toxic at con- centrations of 0.025% or higher. However, when N. corallina was grown in coculture with a strain of Acinetobacter capable of metabol- izing guaiacol, growth of the Nocardia strain was not inhibited and complete decomposition of the model compound was possible. In view 31 of these data one could suggest that other such mutualistic arrange- ments may be beneficial or indeed necessary for the bacterial decom- position of lignin. Kawakami showed that a strain of Pseudomonas ovalis (64) readily degraded compounds which contained B-aryl ether linkages, whereas a- benzyl ethers were not metabolized indicating a rather high degree of specificity of the enzymes capable of cleaving B-ether bonds. Com- pounds with phenylcoumaran and pinoresinol type linkages were not de- graded and, with few exceptions, compounds with substitutuents in the 5 position were not metabolized. Although numerous studies have implicated bacteria in the degra- dation of lignin (63, 96, 109) little conclusive evidence was avail- able until recently (21, 111). Ferm and Nilsson (31) isolated four strains of Pseudomonas, and one strain of Acetobacter which could metabolize lignosulfonate (LS).7 Pseudomonas strain Al was found to utilize only LS less than 500 MW and could not cleave the aromatic rings as evidenced by UV spectroscopy. Hataguchi and Morohoshi iso- lated a Streptomyces sp, capable of utilizing LS as a sole carbon source (54). After one month incubation in medium inoculated with Streptomyces sp, the degraded LS was recovered, fractionated on Sephadex G-25, and its elution pattern was compared with that of un- degraded LS. The amount of LS present in fractions corresponding to a molecular weight of 33,800 decreased, which indicated the organism was able to degrade high molecular weight LS. However, the relative overall amount of high molecular weight LS increased, suggesting the presence of a phenol oxidase. Experiments conducted with crude enzyme preparations demonstrated extensive polymerization of 32 guaiacyl-B-coniferyl ether and limited polymerization of dehydrocon- iferyl alcohol, again suggesting the presence of a phenol oxidase. In 1962 Sorensen (108) isolated several strains of Pseudomonas and Flavobacterium from enrichments inoculated with soil samples con- taining native lignin (NL) as a sole carbon source. The Flavobacter- jgm_isolates reportedly lost the ability to produce clear zones on silica gel plates containing NL as a sole carbon source. This obser- vation suggested that the genes coding for lignin degrading enzymes were located on plasmids and were probably lost during repeated sub- culture in the laboratory. Pseudomonas strain 14 degraded approxi- mately 25% of the added NL within 6 to 8 weeks (108). These data have not been accepted as conclusive evidence for bacterial degrada- tion of lignin because NL is not considered to be representative of the bulk of lignin within wood (4, 5, 37). It has been shown to be of relatively low molecular weight (4, 38) and to differ in functional groups (4), especially phenolic hydroxyl content, from other lignin preparations. The increased phenolic hydroxyl content of NL suggests that NL is less condensed and therefore easier to degrade (38). Kawakami and collaborators (65, 66) studied the degradation of Kraft lignin (KL)6 recovered from pulp mill effluents and milled wood lignin (MWL) by Pseudomonas ovalis. Higher molecular weight fractions of KL were degraded as evidenced by gel permeation chromatography when E, gyalj§_was incubated with pine and beech KL for 60 days. A marked decrease in the phenolic hydroxyl content and dihydroxylated ring structures upon chemical oxidation suggested that ring cleavage occurred. The empirical C9 formula of the degraded KL was found to be C9H9.8302.6l$0.10(0CH3)0.85 whereas that of undegraded KL was 33 c9H8.7002.6650.09(0CH3)0.87' Although the authors conclude, on the basis of an observed increase in yield of compounds containing p- hydroxyphenyl structures and 4-methoxyisophthalic acid upon perman— ganate oxidation, that demethoxylation of KL occurred, the methoxyl analysis does not agree with their conclusion. The degraded KL was 1.1 atoms richer per C9 with the oxygen content essentially unchanged. This indicated that the KL residue was more reduced than the uninoc- ulated control KL. This is contrary to what has been previously re- ported with other biologically degraded lignins (57, 61, 78). Fur- thermore, the IR spectrum of the degraded KL showed an increase in absorbance at 1720 cm"1 as compared to that of the undegraded KL, indicating an increase in carbonyl groups. The observed increase in oxygen containing groups conflicts with the reported increase in hy- drogen atoms per C9 unit observed. Similar results were obtained with MWL (66) except that the oxygen content of the degraded MWL increased from 3.13 per C9 to 3.86 per C9 and a 25% decrease in meth- oxyl content was observed. Examination of the culture filtrate showed accumulation of several low molecular weight intermediates of MWL degradation. Odier and Monties isolated 85 strains of bacteria from wheat straw in decomposition and examined their ability to degrade wheat straw "acidolysis" lignin (AL)8 (94). Eighteen isolates were able to utilize AL as a sole carbonsource. When the medium was supple- mented with glucose, 23-71% of the AL was degraded after 7 days. TWelve of these 21 strains biosynthesized peroxidase enzymes. The isolates were found to be of the fellowing genera: Xanthomonas, Flavobacterium, Aeromonas, Arthrobacter, Bacillus, and Cellulomonas. 34 A strain of Xanthomonas was fOund to be most active. However, AL undergoes a variety of reactions during the acidolysis extraction procedure (37, 116) and is considered by Freudenberg (37) and others (116) to be not representative of the bulk of lignin jn_vivg, and therefore unsuitable for structural and biodegradation studies. Trojanowski gt_al,-(lll) isolated a strain of Nocardia which was capable of degrading specifically ‘4 14 C-labelled maize lignin and C-DHP. In a medium containing minerals, yeast extract (0.01%) and vanillic acid (0.05%), approximately 15% of the side chain (3'140), approximately 13% of the methoxyl groups and approximately 14 14 5% of the ring label was released as CO2 within 15 days from C- maize lignin. In comparison only about 10% of the methoxyl groups, about 6% of the side chain (2"4C) and about 4% of the ring label 14C-DHP. In contrast to the 14 14 14 was released from C-DHP, the C-maize lignin showed the highest rate of CO2 release from the side chains. This difference may be due to the placement of the label in the v 14 14 (terminal) position in C-maize lignin, whereas in C-DHP the label was in the 8 position and therefore not as accessible. The overall rates of specifically labelled DHP degradation by this Nocardia s9, were comparable to those observed by Kirk gt_al, 14 (79) using the white rot fungus Polyporus versicolor. However, C0 14 2 release from C—ring labelled DHP ceased after approximately 6 days suggesting that this strain was limited in its ability to metabolize the ring structures within DHP and thus to substantially degrade lig- nin. In addition, the Nocardia sp, was shown to be capable of meta- olizing a wide range of methoxylated aromatic compounds, aromatic acids and 06C3 compounds. It was not established whether or not the 35 Nocardia s2, produces phenol oxidase enzymes. Recently, (21) three strains of Streptomyces isolated from en- richments containing newsprint as a primary carbon and energy source have been shown to completely oxidize l.5-3.0% of (140)-1ignin fir 14C-MWL in 1025 hours. (Since lignicellulose and as much as 17% of the purified MWL was more readily attacked that lignocellulose it is possible that certain eubacteria which have been shown to decompose purified lignins (94, 111) may not have readily attacked lignins ig_ gitg,) Upon incubation with (140)-glucan fir lignocellulose 27-40% 14 of the radioactivity was released as C02 in 1025 h by these Strep- tomyces. The Streptomyces isolates were concluded to degrade ligno- cellulose in a manner similar to soft; rot fungi wherein lignin is degraded only slightly and the organisms primarily deplete the carbo- hydrate portion. Between 4 and 8% of the total 14C-lignocellulose was shown to be solubilized during incubation with these organisms suggesting that 14 002 evolution data indicate minimal levels of lignin decomposition. Little is known about the role of bacteria in the degradation of lignin in nature (72). Further studies are necessary in order to determine the types of bacteria involved, the biochemical mechanisms employed, and the rates at which they degrade lignin. The regulation of lignin degradation and the location of the genes, i.e. whether located on the chromosome or located on plasmids needs to be examined. Further investigation as to the requirements of carbohydrate, the basis for such a requirement, and the cultural conditions needed for optimal rates of lignin degradation should be conducted. Studies of the type mentioned will lead to a greater understanding of the role 36 bacteria in the recycling of lignin carbon in nature and of the po- tential use of bacteria in industrial delignification processes. 37 Footnotes 1The core of the lignin precursors is a C5C3 (phenylpropanoid) structure. Frequently, elemental and functional group analyses are expressed on the basis of C9 units (38). 2Milled wood lignin (MWL) is obtained by the fine grinding of wood fOllowed by the extraction of the lignin from the wood in a soxhlet apparatus with 96% aqueous dioxane according to the procedure described by Bjorkman (4). The lignin is obtained in yields up to 50% or more depending upon the length of time the wood is ball-milled. This lignin is regarded by lignin chemists to be representative of the bulk of lignin in wood (37, 38) primarily due to the relatively high yields obtainable (4) and because the lignin has an average molecular weight of 11,000 (4) and has not been subjected to chemical processing. 3A simple test for differentiating fungi which produce phenol ox- idases from those which do not was developed by Bavendamm (3). The test involves growing the fungus to be tested on malt agar which has been supplemented with 0.5% gallic or tannic acid. The development of dark zones diffused away from the fungal growth indicates the produc- tion of phenol oxidases which oxidize the hydroxylated acid used as a supplement to the medium resulting in the production of dark colored quinones and/or polymers. 4Phenol oxidases have been shown to catalyze the oxidation of phenolic hydroxyl groups by the removal of one reducing equivalent, e.g. fungal p-diphenol oxidase (p-diphenol:oxidoreductase, E.C. 1.10.3.2, formerly known as laccase) and peroxidase, or two reducing equivalents, o-diphenol oxidase (o-diphenol:0 oxidoreductase, E.C. 1.10.3.1, formerly known as tyrosinase). Freg radicals can result directly from this oxidation as with p-diphenol oxidase and peroxi- dase, or by disproportionation as with o-diphenol oxidase (69). 5Kraft lignin (KL) is produced during the delignification of wood by the alkaline (kraft) pulping process (88). The pulping pro- cess involves heating the pulpwood and pulping liquor (containing primarily sodium hydroxide and sodium sulfide) to 150-175°C (88). During the pulping process the lignin undergoes a variety of reactions including partial loss of the v carbons, limited demethylation of methoxyl groups, and cleavage of alkyl-aryl ether linkages (88). The result is a depolymerization of the lignin molecule which then becomes soluble in basic solution. Kraft lignin is polydisperse (Mn = 1600) and retains many of the structural characteristics of lignin ig_situ (88). The empirical Cg formula is C8 35H7 3O2 1(OCH3)0 78' 6Native lignin (NL; Brauns' native lignin) is that portion of lignin which can be extracted from sawdust with ethanol according to the procedure described by Brauns (7). Although Brauns reported yields of 8-10% of the lignin in wood (7), later investigators norm- ally obtained yields of 0.1-3.3% of the wood (83). NL contains a higher hydroxyl content than MWL suggesting that it is less 38 polymerized than the bulk of lignin in wood and has an average mol- ecular weight of 600 (4). It is generally believed that NL is not representative of the bulk of lignin in wood (4). 7Lignosulfonate (LS) is produced during the delignification of wood by the sulfite pulping process (43). The pulping process in- volves heating the pulping liquor and pulpwood (containing dissolved $02) at pH 1-2 to 135°C (43). The lignin is sulfonated, hydrolyzed, and condensed with a variety of minor reactions occuring as well 343). {he)sulfonated lignin has an average molecular weight of 1-4 X 10 43 . 8Acidolysis lignin (AL) is prepared by refluxing sawdust in a mixture of diozane and dilute HCl (16). The resulting lignin is highly condensed and is not considered to resemble lignin jg_situ (37). 39 Appendix F HzfiOH COH L.’ " /\\ CH 1 HC HCKOH CH3 0 Q ‘ 401 " ©OCH OCH Hsc \O OCH:5 O 3 H 3 H H SYRINGYLGLYCOL—B- R: COOH; GUAIACOXYACETIC ACID B-HYDROXYCONIFERYL ALCOHOL UJA”£YL ETHER R:COH:GUAJACOXYACETALDEHYDE 8 B H COH H C CH 2l 2I l 2 HC——O-{@3> CH CH ' 05“ u H HCOH 3 R H H ©OCH ; OCH H o O OCH 6 3 H 3 3C 0 O 3 H H H CUAIACYLCLYCEROL—B— R=COOH:VANILL|C ACID OJALACYL ETHER R:COH:VAN|LL1N B|S~S-DEHYDROCONIFERYL RszGUAIACOL ALCOHOL RZCHzCH-COOHiFERULm ACE RzCHZCH'COHiCONWERALDEHYDE Figure 5. Structures of certain aromatic compounds discussed in the literature review. 10. 11. 12. 40 Literature Cited Ander, P. and K. E. Eriksson. 1976. The importance of phenol oxidase activity in lignin degradation by the white-rot fungus Sporotrichum pulverulentum. Arch. Microbiol. 109: Adler, E. and K. Lundquist. 1963. Spectrochemical estimation of phenylcoumaran elements in lignin. Acta. Chem. Scand. 17:13-26. Bavendamm, W. 1928. Uber das vorkommen und der Nachweis von oxydasen béi holzzerstorenden. Pilzen. Z. Pflanzenkrankh. 38:257-276. Bjorkman, A. 1956. Studies on finely divided wood. Part 1. Extraction of lignin with neutral solvents. Svensk. Papper- stidn. 59:477-485. Bjorkman, A. and 8. Person. 1957. Studies on finely divided wood. Part 2. The properties of lignins extracted with neutral solvents from softwoods and hardwoods. Svensk. Papperstidn. 60:158-169. Bouvenag, H. 0. and P. Solyom. 1973. Longterm stability of waste lignins in aquatic systems. Svensk. Papperstidn. 76:26-29. Brauns, F. E. 1939. Native lignin 1. Its isolation and meth- ylation. J. Am. Chem. Soc. 61:2120-2127. Brown, 8. R. 1967. Biochemical aspects of oxidative coupling of phenols, p. 167-201. IQ_W. 1. Taylor and A. R. Battersby (eds.), Oxidative coupling of phenols, Marcel Dekker, Inc., New York. Brown, W. E. B. Cowling, and S. I. Falkehag. 1968. Molecular size distributions of lignins enzymatically liberated from wood. Svensk. Papperstidn. 71:811-821. Cartwright, N. J. and K. S. Holdom. 1973. Enzymic lignin, its release and utilization by bacteria. Microbios 8:7-14. Caldwell, E. S. and K. Steelink. 1969. Phenoxy radical inter- mediates in the enzymatic degradation of lignin model com- ponents. Biochim. Biophys. Acta 184:420-431. Chang, H. M. and G. G. Allen. 1971. Oxidation, p. 43-94. In_ K. V. Sarkanen and C. H. Ludwig (eds.), Lignins occurrence, fbrmation, structure and reactions, Wiley-Interscience, New York. 13. 14. 15. 16. 17. 18. 19. 20. 21. 22. 23. 24. 25. 41 Compere, A. L. and W. L. Griffith. 1976. Fermentation of waste materials to produce industrial intermediates. Dev. Ind. Microbiol. 17:247-252. Considine, P. J., N. Flynn, J. W. Patching. 1977. Ethylene production by soil microorganisms. Appl. Environ. Microbiol. 33:977-979. Cowling, E. B. 1961. Comparative biochemistry of the decay of sweetgum sapwood by white and brown rot fungi. USDA Tech. Bull. 1258. 79 p. Cowling, E. B. 1975. Physical and chemical constraints in the hydrolysis of cellulose and lignicellulosic materials. Bio- technol. Bioeng. Symp. No. 5. p. 163-181. Cowling, E. B. and T. K. Kirk. 1976. Properites of cellulose and lignocellulosic materials as substrates for enzymatic conversion processes. Biotechnol. Bioeng. Symp. No. 6. p. 95-123. Cooke, W. B. 1957. Natural and induced fungal degradation of lignin. T.A.P.P.I. 40:301-306. Crawford, 0. L., S. Floyd, A. L. Pometto, and R. L. Crawfbrd. 1977. Degradation of natural and kraft lignins by the micro- flora of soil and water. Can. J. Microbiol. 23:434-440. Crawford, 0. L., R. L. Crawford, and A. L. Pometto. 1976. Preparation of specifically labeled 14C-(lignin) and 14C- (cellulose) lignocelluloses and their decomposition by the microflora of soil. Appl. Environ. Microbiol. 33:1247-1251. Crawford, 0. L. 1979. Lignocellulose decomposition by selected Streptomyces strains. Appl. Environ. Microbiol. 35:1041-1045. Crawford, R. L., E. McCoy, T. K. Kirk, and J. B. Obst. 1973. Bacterial cleavage of an arylglycerol-B-aryl ether bond. Appl. Microbiol. 25:322-324. Dence, C. W. 1971. Halogenation and nitration, p. 372-432. In_K. V. Sarkanen and C. H. Ludwig (eds.), Lignins occur- rence, formation, structure and reactions, Wiley-Interscience, New York. Crawford, R. L., D. L. Crawford, C. Olofsson, L. Wikstrom, and J. M. Wood. 1977. Biodegradation of natural and manmade recalcitrant compounds with particular reference to lignin. J. Agric. Food. Chem. 25:704-708. Cysewski, G. R. and C. R. Wilke. 1976. Utilization of cellu- losic materials through enzymatic hydrolysis. I. Fermenta- tion of hydrolysate to ethanol and single cell protein. Bio- technol. Bioeng. 28:1297-1313. 26. 27. 28. 29. 30. 31. 32. 33. 34. 35. 36. 37. 38. 39. 42 Dagley, S. 1967. The microbial metabolism of phenolics, p. 287- 317. Ig_A. D. McLaren and G. Peterson (eds.), Soil biochem- istry, Marcel Dekker, Inc., New York. Davidson, R. W., D. A. Campbell, and D. J. Blaisdell. 1938. Differentiation of fungi by their reactions on gallic or tannic acid medium. J. Agric. Res. 57:683-695. Dehority, B. A. and R. R. Johnson. 1961. Effect of particle size upon the in vitro cellulose digestibility of forages by rumen bacteFia. J. Dairy Sci. 44:2242-2249. Dence, C. W. 1971. Halogenation and Nitration. In; Ligins Occurrence, Formation, Structure, and Reactions. K. V. Sarkanen and C. H. Ludwig, eds., Wiley-Interscience. p. 372-432. Fergus, B. J., A. R. Procter, J. A. N. Scott, and D. A. I. Gor- ing. 1961. The distribution of lignin in spruce wood as determined by untraviolet microscopy. Wood Sci. Technol. 3:117-138. Ferm, R. and A. C. Nilsson. 1969. Microbiological degradation of a commercial lignosulfonate. Svensk. Paperstidn. 72: 531-138. Freudenberg, K., and G. Grione. 1959. Bietrag zum bildungsnec- anismus des lignins und der 1ignin-kohlendhydrat-bindung. Chem. Ber. 92:1355-1363. Freudenberg, K. and J. M. Harkin. 1960. Modelle fur die bind- ung des lignins an die kohlenhydrat. Chem. Ber. 93:2814- 2819. Freudenberg, K., C. L. Chen., and G. Cardinale. 1962. Die oxydation des methylation naturlichen und kunstlichen lig- nins. Chem. Ber. 95:2814-2828. Freudenberg, K. 1964. Entwurf eines konstitutionsschemas fur das lignin der fichte. Holzforschung. 18:3-9. Freudenberg, K., and J. M. Harkin. 1964. Erganzung des konstitutionsschemas fur das lignin der fichte. Holzfor- schung 18:166-168. Freudenberg, K. 1965. Lignin: Its constitution and formation from p-hydroxycinnamyl alcohols. Science. 148:596-600. Freudenberg, K. and A. C. Neish. 1968. Constitution and Bio- synthesis of Lignin. Springer-Verlag, NY. 129 p. Fukuzumi, T. 1960. Enzymic degradation of lignin. 1. Paper chromatographical separation of intermediate degradation 40. 41. 42. 43. 44. 45. 46. 47. 48. 49. 50. 51. 43 products of lignin by the wood rotting fungus Poria subacida (Peck) Sacc. Bull. Agric. Chem. Soc. Jap. 24:728-738. Fukuzumi, T. and T. Shibamato. 1965. Enzymatic degradation of lignin. IV. Splitting of veratrylglycerol~8-guaiacyl ether by enzymes of Poria subacida. J. Jap. Wood Res. Soc. 11: 248-252. Ganczarczyk, J. and T. Obiaga. 1974. Mechanism of lignin re- moval in activated sludge treatment of pulp mill effluents. Water Res. 8:857-862. Gellerstidt, G. and E. L. Pettersson. 1975. Light induced ox- idation of lignin. The behaviour of structural units con- taining a ring conjugated double bond. Acta. Chem. Scand. 829:1005-1010. Glennie, D. W. 1971. Reactions in sulfite pulping. In_K. V. Sarkanen and C. H. Ludwig (eds.), Lignins occurrence, forma- tion, structure and reactions, Wiley-Interscience, New York. Goldstein, I. S. 1976. Chemicals from lignocellulose. Bio- technol. Bioeng. Symp. No. 6. p. 293-301. Goring, D. A. I. 1971. Polymer properties of lignin and lignin derivatives, p. 695-768. Ig_K. V. Sarkanen and C. H. Ludwig (eds.), Lignins occurrence, formation, structure and reac- tions, Wiley-Interscience, New York. Gottlieb, S., W. C. Day, and M. J. Pelczar. 1950. The biologi- cal degradation of lignin. II. The adaptation of white rot fungi to growth on lignin media. Phytopathol. 40: 926-935. Gottlieb, S. and M. J. Pelczar. 1951. Microbiological aspects of lignin degradation. Bacteriol. Rev. 15:55-76. Griffin, J. M. and J. L. West. 1976. Acute toxicity of ammonia base neutral sulfite pulp mill waste liquor to rainbow trout. Bull Env. Contamin. Toxicol. 15:608-612. Hackett, W. F., W. J. Connors, T. K. Kirk, and J. G. Zeikus. 1977. Microbial decomposition of synthetic 14C-labelled lignins in nature: Lignin biodegradation in a variety of natural materials. Appl. Environ. Microbiol. 33:43-51. Haider, K., J. P. Martin and Z. Filip. 1975. Humus biochemis- try, p. 195-244. Ig_E. A. Paul and A. D. McLaren (eds.), Soil biochemistry, vol. 4, Marcel-Dekker Inc., New York. Haider, K. and J. Trojanowski. 1975. Decomposition of spec- ifically labeled phenols and dehydropolymers of coniferyl alcohol as models for lignin degradation by soft and white rot fungi. Arch. Microbiol. 105:33-41. 52. 53. 54. 55. 56. 57. 58. 59. 60. 61. 62. 63. 44 Haider, K., J. P. Martin, and E. Reitz. 1977. Decomposition in soil of 14C-labeled coumaryl alcohols: Free and linked into dehydropolymer and plant lignins and model humic acids. Soil Sci. Soc. Am. J. 41:556-561. Han, Y. W. and C. D. Callihan. 1974. Cellulose fermentation: Effect of substrate pretreatment on microbial growth. Appl. Microbiol. 27:159-165. Haraguchi, T. and N. Morohoshi. 1975. Degradation of Ligno- sulphonates by Streptomyces s2, Proc. Third Int. Biodegrad. Symp. p. 719-729. Harkin, J. M. 1967. Lignin: A natural product of phenol oxida- tion, p. 243-321. In_W. I. Taylor and A. R. Battersby, (eds.), Oxidative coupling of phenols, Marcel-Dekker Inc., New York. Harkin, J. M., D. L. Crawford, and E. McCoy. 1974. Bacterial progein from pulp and paper mill sludge. T.A.P.P.I. 57: 131-134. Hata, K. Investigations on lignins and lignification. XXXII. Studies on lignins isolated from spruce wood decayed by Poria subacida. Bll. Holzforschung. 20:142-147. Higuchi, T., I. Kawamura, and H. Kawamura. 1955. Properties of the lignin in decayed wood. J. Jap. Forest Soc. 37: 298-302. Higuchi, T. 1971. Formation and biological degradation of lignins. Adv. Enzymol. 34:207-283. Hurst, H. M. and N. A. Burges. 1967. Lignin in humic acids, p. 260-286. IQ_A. D. McLaren and G. Peterson (eds.), Siol Biochemistry, Marcel-Dekker, New York. Ishikawa, H., W. J. Schubert, and F. F. Nord. 1963. Investiga- tions on lignins and lignification. XXVII. The enzymic degradation of softwood lignin by white rot fungi. Arch. Biochem. Biophys. 100:131-139. Ishikawa, H., W. J. Schubert and F. F. Nord. 1963. Investiga- tions on lignins and lignification. XXVIII. The degrada- tion by Polyporus versicolor and Fomes fomentarius of aromatic compounds structurally related to softwood lignin. Arch. Biochem. Biophys. 100:140-149. Jaschof, H. 1964. Preliminary studies of the decomposition of lignin by bacteria isolated from lignite. Geochim. Cosmo- chim. Acta. 28:1623-1638. 64. 65. 66. 67. 68. 69. 70. 71. 72. 73. 74. 75. 76. 45 Kawakami, H. 1975. Bacterial degradation of lignin model com- pounds. III. On the degradation of model dimers and 5- position condensed type compounds of guaiacyl nuclei. J. Japan. Wood Res. Soc. 21:629-634. Kawakami, H., M. Sugiura, and T. Kanda. 1975. Biodegradation of components of pulp waste effluents by bacteria (1) on the degradation of kraft lignin. Japan T.A.P.P.I. 29:33-39. Kawakami, H. 1976. Bacterial degradation of lignin. l. Degra- dation of MWL by Pseudomonas ovalis. J. Japan. Wood Res. Soc. 22:252-257. Kihlberg, R. 1972. The microbe as a source of food. Ann. Rev. Microbiol. 26:427-457. Kirk, T. K. and A. Kelman. 1965. Lignin degradation as related to the phenoloxidases of selected wood decaying Basidiomy- cetes. Phytopathol. 55:739-745. Kirk, T. K., J. M. Harkin, and E. B. Cowling. 1968. Oxidation of guaiacyl- and veratrylglycerol-B-guaiacyl ether by Poly- pgrus versicolor and Stereum frustulatum. Biochim. Biophys. Acta. 165:134-144. Kirk, T. K., J. M. Harkin and E. B. Cowling. 1968. Degradation of the lignin model compound syringylglycol-8-guaiacyl ether by Polyporus versicolor and Stereum frustulatum. Biochim. Biophys. Acta. 165:145-163 Kirk, T. K. and E. Adler. 1970. Methoxyl-deficient structural elements in lignin of sweetgum decayed by a brown rot fungus. Acta Chem. Scand. 24:3379-3390. Kirk, T. K. 1971. Effects of microorganisms on lignin. Ann. Rev. Phytopathol. 9:185-210. Kirk, T. K. and W. E. Moore. 1972. Removing lignin from wood with white rot fungi and the digestibility of the resulting wood. Wood and Fiber. 4:72-79. Kirk, T. K. 1973. Polysaccharide integrity as related to the degradation of lignin in wood by white rot fungi. Phyto- pathol. 63:1504-1507. Kirk, T. K. and T. L. Highley. 1973. Quantitative changes in structural components of conifer woods during decay by white and brown rot fungi. Phytopathol. 63:1338-1342. Kirk, T. K. and J. M. Harkin. 1973. Lignin biodegradation and the bioconversion of wood. Forest Products and the Environ- ment. AICHE Symp. Series. 69:124-126. 77. 78. 79. 80. 81. 82. 83. 84. 85. 86. 87. 88. 89. 46 Kirk, T. K. and H. M. Chang. 1974. Decomposition of lignin by white rot fungi. 1. Isolation of heavily decayed lignins from decayed spruce woods. Holszrschung 28:217-222. Kirk, T. K. and H. M. Chang. 1975. Decomposition of lignin by white rot fungi. 2. Characterization of heavily degraded lignins from decayed spruce woods. Holszrschung 29:56-64. Kirk, T. K., W. J. Conners, R. D. Bleam, W. F. Hackett, and J. G. Zeikus. 1975. Preparation and microbial decomposition of synthetic 14c-lignins. Proc. Nat. Acad. Sci. USA 72: 2515-2519. Kirk, T. K., W. J. Conners, and J. G. Zeikus. 1976. Require- ment for a growth substrate during lignin decomposition by two wood-rotting fungi. Appl. Environ. Microbiol. 32:192- 194. Kirk, T. K. 1976. Biological delignification present status- future directions. Proc. Weyerhouser Symp. p. 31-54. Kushner, D. J., M. Vincent, and C. Novitsky. 1974. Degradation of cellulose and wood products in polluted and unpolluted rivers. Proc. Ninth Can. Symp. Water Poll. Res. Can. p. 149- 153. Lai, Y. Z. and K. V. Sarkanen. 1971. Isolation and structural studies. In; Lignins Occurrence, formation, Structure, and reactions. K. V. Sarkanen and C. H. Ludwig, eds., Wiley- Interscience, NY. p. 165-240. Lawton, E. J., W. D. Bellamy, R. E. Hungate, M. P. Bryant, and E. Hall. 1951. Some effects of high velocity electrons on wood. Science 113:167-174. Leonowicz, A. and J. Trojanowski. 1975. Induction of a new laccase form in the fungus Pleurotus ostreatus by ferulic acid. Microbios 13:167-174. Levi, M. P. and R. 0. Preston. 1965. A chemical and microscop- ic examination of the action of the soft rot fungus Chaetomium globosum on beechwood (Fagus sylv.) Holszrschung 19YT83-190. Lipinsky, E. s. 1978. Fuels from biomass: Integration with food and materials systems. Science 199:644-651. Marton, J. 1971. Reactions in alkaline pul ing, p. 639-694. In_K. V. Sarkanen and C. H. Ludwig (eds. , Lignins occur- rence, formation, structure and reactions, Wiley-Interscience, New York. Martin, J. P. and K. Haider. 1971. Microbial activity in rela- tion to soil humus formation. Soil Sci. 111:54-63. 90. 91. 92. 93. 94. 95. 96. 97. 98. 99. 100. 101. 102. 103. 47 Miller, D. L. 1976. Fermentation ethyl alcohol. Biotechnol. Bioeng. Symp. No. 6. p. 307-312. Millet, M. A., A. J. Baker, W. C. Fiest, R. W. Mellenberger, and L. D. Satter. 1970. Modifying wood to increase its jn_vitro digestibility. J. Anim. Sci. 31:781-788. Millett, M. A., A. J. Baker, and L. D. Satter. 1976. Physical and chemical pretreatments for enhancing cellulose saccar- ification. Biotechnol. Bioeng. Symp. No. 6. p. 125-153. Nesse, N., J. Wallick, and J. M. Harper. 1977. Pretreatment of cellulosic wastes to increase enzyme reactivity. Bio- technol. Bioeng. 29:323-336. Odier, E. and B. Monties. 1977. Activite ligninolytique jg. vitro de bacteries isolees de paille de ble en decomposition. C. R. Acad. Sc. Paris 284:2175-2187. Oglesby, R. T., R. F. Christman, and C. H. Driver. 1967. The biotransformation of lignin to humus. Adv. Appl. Microbiol. 9:171-184. Pandilla, M. M. 1973. Microorganisms associated with microbio- logical degradation of lingnosulphonatesz A review of literature. Pulp and Paper Mag. Can. 74:80-84. Pelczar. M. J., S. Gottlieb, and W. C. Day. 1950. Growth of Polyporus versicolor in a medium with lignin as the sole carbon source. Arch. Biochem. 25:449-451. Pew, J. C. and P. Weyna. 1962. Fine grinding, enzyme digestion and the lignin cellulose bond in wood. T.A.P.P.I. 45:247- 256. Pew, J. C. 1963. Evidence of a biphenyl group in lignins. J. Org. Chem. 28:1048-1054. Pigden, W. and D. P. Heaney. 1969. Lignicelluloses in ruminant nutrition. Adv. Chem. Series 95:245-261. Raiha, M. and V. Sundman. 1975. Characterization of ligni- sulfonate induced phenol oxidase activity in the atypical white rot fungus Polyporus dichrous. Arch. Microbiol. 105:73-76. Reddy, C. A. and L. Forney. 1978. Lignin Chemistry: A brief review. Dev. Ind. Microbiol. 19:27-34. Sarkanen, K. V. and H. L. Hergert. 1971. Classification and distribution, p. 43-94. Ig_K. V. Sarkanen and C. H. Ludwig (eds.), Lignins occurrence, formation, structure and reac- tions, Wiley-Interscience, New York. 104. 105. 106. 107. 108. 109. 110. 111. 112. 113. 114. 115. 116. 48 Sarkanen, K. V. 1971. Precursors and their polymerization, p. 95-163. ln_K. V. Sarkanen and C. H. Ludwig (eds.), Lignins occurrence, formation, structure and reactions, Wiley-Interscience, New York. Schubert, J. 1965. Lignin Biochemistry. Academic Press, N.Y. 131 p. Sigleo, A. C. 1978. Degraded lignin compounds identified in silicified wood 200 million years old. Science 200:1054- 1055. Sjoblad, R. D. and J. M. Bollag. 1977. Oxidative coupling of aromatic pesticide intermediates by a fungal phenol oxidase. Appl. Environ. Microbiol. 33:906-910. Sorensen, H. 1962. Decomposition of lignin by soil bacteria and complex formation between autooxidized lignin and or- ganic nitrogen compounds. J. Gen. Microbiol. 27:21-34. Sundman, V., T. Kuusi, S. Kuhanen, S. Kilpi, and H. Sedrholm. 1968. Observations on bacterial utilization of the lignin from brown-rotted Spruce wood and Brauns' native lignin. Medd. Finska. Kemsamf 77:70-86. Tarkow, H. and W. C. Fiest. 1969. A mechanism for improving the digestibility of lignocellulosic materials with dilute alkali and liquid ammonia. Adv. Chem. Series 95:197-218. Trojanowski, J., K. Haider, and V. Sundman. 1977. Decomposi- tion of VIC-labelled lignin and phenols by a Nocardia §p_. Arch. Microbiol. 114:149-153. Van Vliet, W. F. 1954. The enzymic oxidation of lignin. Bio- chim. Biophys. Acta 15:211-216. Waksman, S. A. and I. J. Hutchings. 1936. Decomposition of lignin by microorganisms. Soil Sci. 42:119-130. Wilson, R. K. and W. J. Pigden. 1964. Effect of sodium hy- droxide treatment on the utilization of wheat straw and poplar wood by rumen microorganisms. Can. J. Anim. Sci. 44:122-123. Wishart, R. S. 1978. Industrial energy in transition: A petrochemical perspective. Science 199:614-618. Wallis, A. F. A. 1971. Solvolysis by acids and bases, p. 345- 372. In_K. V. Sarkanen and C. H. Ludwig (eds.), Lignins occurrence, formation, structure and reactions, Wiley- Interscience, New York. 49 117. Wardrop, A. B. 1971. Occurrence and formation in plants, p. 19-41. IQ K. V. Sarkanen and C. H. Ludwig (eds.), Lignins occurrence, formation, structure and reactions, Wiley-Interscience, New York. 118. Westermark, U. and K. E. Eriksson. 1974. Cellobiose: quinone oxidoreductase, a new wood degrading enzyme from white rot fungi. Acta. Chem. Scand. 828:209-214. 119. Woodwell, G. M. 1978. The carbon dioxide question. Scientific American 238:34-44. CHAPTER III MATERIALS AND METHODS Enrichment Inocula. The soil samples used as inocula for the enrichment media (see Table 2) were kindly provided by Dr. James Tiedje (Department of Microbiology and Public Health, Michigan State University, East Lansing, MI). The sources of each soil sample is given in Table 2 and the characteristics of each soil sample have been published previously (Gamble et al., 1977). Primary Lignin Enrichments. The primary enrichment medium (PEM) contained per 1: KH P0 2.0 9; K HPO -2H 2 4’ o, 0.3 g; MnCl 2.0 9; CaCl 0, 2 4’ o, 1.32 mg; C001 2 2 0.3 9; MgCl -6H -4H °6H20, 2.0 mg; 2 2 0, 1.4 mg; FeSO 2 2 2 ZnSO -7H 5.0 mg; (NH4) $0 1.4 g; urea, 0.3 g; 4 2 4’ 2 4’ Indulin AT, 5.0 g; and cellulose (Whatman, W&R Ralston, England), 1.0 g. The kraft lignin (Indulin AT, Westvaco, North Charleston, SC) used throughout this study was derived from the alkaline pulping of primarily long needle pine wood and was free of extraneous carbo- hydrates (I. Falkehag, personal communication). The medium was adjusted to pH 6.8, dispensed into 250 ml Erlenmeyer flasks (25 m1/flask) stoppered with foam plugs and autoclaved at 121°C for 15 min. Each flask was inoculated with 5.0 g (wet wt.) of a given soil type. The flasks were incubated in a gyrotory shaker water bath (New Bruns- wick Scientific Inc., New Brunswick, NJ) at 25°C and 120 rpm. At 50 51 Table 2. Source and type of soil samples used aS inocula for lignin enrichment cultures, and the number of bacterial strains isolated from each culture. Soil . . 1 Total No. of Isolates No. Sampllng Sjte Crop cover isolates selected2 1 San Pedro, Argentina Corn field 29 3 2 Parana, Argentina Wheat/sweet clover field 27 2 3 Mocaca, Brazil Wheat field. 22 0 4 Ibadan, Nigeria Corn field 29 9 5 Ibadan, Nigeria Rice/corn field 30 4 6 Palmira, Colombia Rice paddy 27 3 7 Los Banos, Philippines Rice paddy 21 l 8 Taichung, Taiwan Rice paddy 14 0 9 San Carlos de Rio Tropical rain Negro, Venezuela forest 29 0 1 Characteristics of each soil type have been described pre- viously (Gamble et a1. 1977). 2Isolates selected for the Study of kraft lignin degradation based on their ability to grow rapidly on SM. See text for details. 52 two week intervals, 0.1 ml aliquots of the enrichments were serially diluted in sterile 0.85% (w/v) NaCl dilution blanks. One-tenth ml each of the 10'4 through 10'7 dulutions were plated onto Tryptose Agar (BBL, Cockeysville, MD) and Indulin agar (PEM minus cellulose and supplemented with 15 g/l Bacto-agar (Difco, Detroit, MI). After 48 h of incubation the total number of bacteria in each enrichment was estimated by counting the colonies on a Tryptose Agar (TA) plate containing between 30-300 colonies. The Indulin agar (IA) plates, on the other hand, were generally incubated two to four weeks at 23°C in plastic bags to maintain adequate humidity before the colonies were picked. At two week intervals, 5.0 m1 aliquots of each enrichment culture were aseptically inoculated into 25 m1 of fresh medium. About 107-108 bacteria per ml were found in each primary enrichment culture. Secondary Lignin Enrichments. Each primary enrichment culture after four successive transfers in PEM, was transferred in 5 m1 amounts to a secondary enrichment medium (SEM). This medium was identical to PEM except that the cellulose was omitted and the Indulin AT was washed, prior to addition, to remove water soluble contaminating materials, if any. The washing was accomplished by thoroughly mix- ing Indulin AT in distilled water (5 g/25 ml), centrifuging at 48,000 x g for 10 min and discarding the supernatant fluid. The pellet was resuspended in distilled water, and was collected by filtration on Whatman #1 filter paper. Lignin retained on the filter paper was washed with approximately 100 m1 of distilled water, gently scraped into a glass Petri dish, dried overnight at 100°C in a circulating 53 air oven and was then added to the enrichment medium. The enrichments were incubated as above, except at 30°C. At weekly intervals, 5.0 ml aliquots of the respective enrichment cultures were transferred to 25 m1 of fresh medium. At the time of each transfer, viable bac- terial counts in each enrichment culture were estimated as described 5 7 above. These secondary enrichment cultures contained 10 to 10 bac- teria per m1. Enrichment with Phenolic Compounds. To enrich for bacterial strains capable of degrading certain phenolic compounds a medium with the same mineral composition as PEM but containing 200 mg/l each of 4-hydroxybenzoic acid, 4-hydroxy-3-methoxybenzoic acid, and 4-hydroxy- e,5-dimethoxybenzaldehyde, instead of Indulin AT, was used. The medium (PCM) was adjusted to pH 6.8, sterilized by filtration through 0.2 pm pore size filter, and 18.0 ml of the medium was dispensed into foam plugged, sterile 125 m1 Erlenmeyer flasks. The flasks were in- oculated with 2.0 m1 of each respective primary enrichment culture and were incubated as described above for the secondary enrichments. Twice, at 10 day intervals during incubation, serial dilutions of these enrichments were prepared as described above. One-tenth ml of the appropriate dilution was plated Onto a sterile solid medium (PCMA) prepared by supplementing PCM with 15 g/l Bacto-agar (Difco). The plates were incubated for about two weeks at 30°C and subsequently stored at 4°C. Isolation of Bacteria. Based on gross differences in colonial morphology and pigmentation, representative colony types were picked from all the plates (TA, IA, PCMA) inoculated with the three different 54 enrichments throughout the enrichment period. A total of 228 isolates were obtained. The number of isolates obtained from each soil source is indicated in Table l. The purity of each isolate was confirmed by streaking each colony on Tryptose Agar plates several times. Stock cultures of the isolates were maintained through monthly transfer on Tryptose Agar slants. Screening of Isolates. All isolates were screened for their ability to degrade the three phenolic compounds present in PCM. To do this each isolate was grown in 3.0 ml of sterile Tryptose Phsophate Broth (TPB; Difco, Detroit, MI) contained in 13)(100 mm foam-stoppered tubes at 30°C until visibly turbid. A drop of each culture was then used to inoculate screening medium (SM; 3 ml per 13 x 100 mm tube) which was identical to PCM except that the medium minus minerals was sterilized by autoclaving and upon cooling, 10 ml of a filter ster- ilized mineral solution containing in g/l: NaH2P04, 1.5; NaZHP04, O, 0.01; and FeSO -7H 1.5; NaC1, 0.25; (NH4)ZSO4, 1.25; MgSO4°7H O; 2 4 2 0.002 was added to the medium. The cultures were incubated at 30°C until visibly turbid and the ultraviolet spectrum (400-200 nm) of a 10'2 dilution of the culture in distilled water was obtained using a Varian model 6345 UV-visible spectrophotometer (Varian Associates, Palo Alto, CA). Distilled water was used as the reference. The deg- radation of the phenolic compounds was evidenced by a decrease in absorbance at 254,286 and 308 nm which are the absorbance maxima for the coumpounds employed (Lemon, 1947). Seventy-two of the isolates tested metabolized the phenolic compounds to a substantial extent and the remainder of the isolates metabolized these compounds to 55 a lesser extent or not at all. Further screening of these 72 isolates was accomplished as follows. Each stock culture was aseptically transferred into 3.0 m1 of sterile TPB and incubated at 30°C. One drop of each culture was aseptically transferred to 10 ml of sterile SM contained in 18 x 150 mm foam-plugged tubes. The tubes were then incubated at 30°C. At zero time and periodically during the ensuing 90 h of incubation, the absorbance of these tubes at 600 nm was determined using a Bausch- Lomb Spectronic 20 spectrophotometer. Twenty-two of the cultures, which showed an absorbance greater than 0.24 A, were studies further to determine their ability to degrade kraft lignin. The degradation of the phenolic substrates by these isolates was confirmed by deter- mining the UV spectra of the spent medium as described above. Characteristics of the Isolates. The basal medium used for testing the utilization of various substrates was the same as SM ex- cept that the aromatic compounds were substituted with a given sub- strate at a concentration of 0.1% w/v. A control medium without added substrate was included in each case. A test was considered positive if the medium containing the Substrate was visiblv more turbid than the control medium without the substrate. Fluorescent pigment production was determined on medium 8 described by King et al. (1954). Flagella were stained according to the procedure of West et al. (1977). The presence of oxidase was determined accord- ing to Kovacs test (Kovacs, 1965) and the production of catalase by the method of Colwell and Wiebe (1970). Isolates were characterized to the genus level based on the criteria listed in Bergey's Manual 56 of Determinitive Bacteriology (Duodoroff and Palleroni, 1974). Prepgrationlerraft Lignin. Kraft lignin from pine wood was reported to be polydisperse and to have an average MW of 1600 (Marton, 1971). To remove any low molecular weight components pre— sent, the kraft lignin employed was purified by gel filtration chrom- atography, as described by Kirk et al. (1975), using Sephadex LH-20 (Pharmacia Fine Chemicals, Uppsala, Sweden) swollen in redistilled dimethylformamide (DMF). The bed size was 2.8 x 27 cm, the void volume was 52 ml and a flow rate of 0.5 m1/min was maintained with redistilled DMF as the eluting solvent. The projected exclusion limit of the column was approximately 1470 MW (Kirk et al., 1975). The kraft lignin solution in DMF (100 mg/ml) was applied in 2.0 ml aliquots. All fractions up to 52 ml were pooled and subsequently evaporated to approximately 15 ml in a flash evaporator at 55°C. The fractionated kraft lignin was then precipitated by adding it drop- wise to 200 m1 of redistilled tolune. The solvent was removed by evaporation under N2 and the lignin stored jg_vggug_over P205. This fractionated kraft lignin, referred to in the rest of the text as lignin, was used in all subsequent experiments. Growth Studies in Kraft Lignin Media. The twenty-two isolates (see screening of isolates) were aseptically transferred from stock slants to 10 ml of SM broth and incubated for 48 h at 30°C. After one additional subculture in this medium, 0.05 ml aliquots of each of the 22 cultures were aseptically pooled and this pooled culture was used to inoculate the four different kraft lignin media described below. 57 The fbur types of media employed to study the degradation of kraft lignin by the isolates are Shown in Table 3. White rot fungi were shown to be unable to utilize lignin as a sole carbon source (Kirk et al., 1976), unless a growth substrate such as glucose or cellulose was provided. In view of this, 0.01% glucose was added to medium I, in addition to lignin and minerals, while medium II con- tained lignin and minerals only. To examine the effects of minerals on viable counts and lignin degradation, the nfineral solution was replaced by distilled water in medium III. Medium IV served as a minus lignin control. Each medium was adjusted to pH 6.8, dispensed in Erlenmeyer flasks (20 ml per flask), foam-stoppered and autoclaved at 121°C for 15 min. One-tenth ml of the pooled culture described above was then used to inoculate each type of medium contained in duplicate flasks. The first set of flasks (series A) was incubated on a gyrotory shaker water bath at 30°C and 120 rpm. After eight days of incubation, 1.0 m1 of each culture was transferred to a sec- ond series of flasks (series B). Flasks in series A and B were then incubated as described above for up to 35 and 52 days, respectively. The bacterial numbers in both series of flasks were determined at zero time and periodically thereafter during incubation, as described above for the PEM and SEM enrichments. Flasks of uninoculated medium II served as the controls. After 52 days of incubation, 1.0 m1 of medium 11, series B, was used to inoculate two flasks of fresh medium III (series C). These flasks were incubated as above for 9 days and the bacterial numbers were determined, as described above, periodically during the incubation period. 58 Table 3. Composition of the media used for studying kraft lignin degradation by a mixed culture of bacteria isolated from tropical soils. Medium Substituent Quantity2 I II III IV Kraft lignin 10 mg + + + - (>1500 MW) Mineral solution1 20 m1 + + - + Distilled water 20 ml - - + - Glucose 2 mg + - - _ 1The composition of mineral solution was the same as that used in the screening medium described in the text. 2Amounts present in 20 ml of each medium contained in sterile 125 m1 Erlenmeyer flasks stoppered with form plugs. 59 Kraft Lignin Analysis. The extent of kraft lignin degrada- tion at the end of 9, 35 and 52 days of incubation in the three series of flasks described above was estimated by UV spectroscopy, a procedure which has been widely used for the quantitation of lig- nin in various preparations (Goldschmid, 1971). In this procedure, the absorbance at 283 nm of an appropriate dilution of each culture in 50% aqueous dioxane was measured against 50% aqueous dioxane in an identical reference cuvette. A typical UV spectrum of undegraded and degraded lignin is Shown in Figure 6. The amount of lignin re- maining in a culture flask was calculated by using an absorptivity coefficient of 23.7 l-g'lcm']. For infrared analysis, the remaining dioxane-culture solution above was evaporated to 2-3 ml in a rotary flash evaporator at 60°C. This solution was then added dr0pwise to 300 ml of distilled water with continuous stirring and the lignin precipitate was recovered by filtration through a 0.2 pm filter (Amicon Corp., Lexington, MA). The filter was placed in a Petri dish, allowed to air dry, and 1.5 mg of this dried lignin was then added to 200 mg KBr and its IR spectrum (4000-400 cm']) was obtained using a Perkin-Elmer Model 334 infrared spectrophotometer. 60 Figure 6. A typical ultraviolet spectrum of undegraded (solid line) and degraded (dashed line) kraft lignin 61 MG? thbmm V 0.00 b 300 325 WAVELENGTH (nm) 275 250 62 CHAPTER IV RESULTS Characteristics of the Isolates. Twenty-two of the 228 bac- terial strains isolated from various tropical soils readily degraded p-hydroxybenzoic acid, 4-hydroxy-3-methoxybenzoic acid, and 4-hydrosy- 3,5-dimethoxybenzaldehyde and only these isolates were selected for further study. When these isolates were grown in a medium contain- ing the above three phenolic compounds as substrates, maximum absorb- ance values of 0.24 to 0.38, with 90 h of incubation, were observed. In all cases the observed growth was roughly proportional to the decrease in the absorbance at 254,286, and 308 nm, the absorption maxima of these substrates. None of the isolates showed a preferen- tial metabolism of one compound over another. None of these 22 isolates were from soils 3, 8 and 9 and only one isolate was from soil 7 (Table 2). Thirteen of the 22 isolates were from agricultural soil samples from Nigeria (soils 4 and 5). None of the isolates were from the Venezuelan tropical rain forest soil sample (soil 9) which was somewhat unexpected since the rate of organic matter turnover is high in such soils. However, the condi- tions of the enrichment, or screening procedures may have precluded the selection of the responsible microorganisms from this soil. All the selected isolates were aerobic, Gram-negative, 63 nonsporefbrming, catalase positive, oxidase positive, relatively short rods showing polar, monotrichous flagellation. All isolates produced white to cream-colored, medium sized colonies on Tryptose agar. AS shown in Table 4, 8 and 22 strains exhibited fluorescent pigmentation. All the strains grew on glucose but no gas production was detectable. All but one strain used pyruvate, acetate, or suc- cinate as a sole carbon source. Based on these and other character- istics (Table 4) the isolates were tentatively classified as members of the genus Pseudomonas (Duodoroff and Palleroni, 1974) Growth Studies with Kraft Lignin. Viable cell counts for the fOur types of media employed (Table 3) in Series A and B and for med- ium III of Series C are shown in Fig. 7. In media I-III there was a rapid increase in the viable bacteria counts during the first few days fallowed by a less rapid increase in bacterial numbers for the remain- der of the experiment. except in medium I (Fig 7a) in which the bac- terial population declined slightly after the initial period of rapid growth; presumably the decline in numbers followed the exhaustion of glucose. Data in Fig 7b & c Show that viable counts of 8.7 x 107 bacteria/ml in lignin medium supplemented with minerals (medium II) were slightly higher than those in unsupplemented lignin medium (medium III). Viable counts in medium IV containing minerals only (no lignin present) were about 1/3 those observed in medium II (Fig. 7b & d). Furthermore, the results showed that in all the media the viable counts in flasks of series A were similar to those in flasks of series B. In flasks of medium III in series C, which were inoculated with 1.0 ml of culture from an identical medium in series Table 4. 64 Characteristics of bacteria isolated from various tropical soils and selected for study of kraft lignin degrradation.1 Strain no.2 Substrates utilized4 2 plgment production pyruvate acetate succinate glucose fucose mannitol trehalose L-tryptophan inositol 15223L 16223L 11324T 28223L 21319T 4149TA 4449TA 44324L 4138L 4238L 4538L 4338L 42324L 42223L 55223L 5238L 53223L 51223L 6249TA 6149TA 62223L 71319T l l+l++ l lll+l+ ll+lll I llll+l+ Illlll|l+l++l lll+lllll+l l+ll+ll++ll lllll lll|+ll I+I+I lllllllllllll+lllll I llll l |+llll+l + + + + + + + +.+ + + + + l + + + + + + + + + + + + + +-+ + + + + + + + + + + + + + + + + + + + + + + + + + + + + l + + + + + + + + + + + + + + + + + + + + + + + + + + + + + + l++l++l+l 1 the isolates were aerobic, Gram-negative, nonsporeforming, catalase positive, oxidase positive, relatively short rods showing polar, monotrichous flagellation. None of the strains produced gas from glucose. 2 The first digit of the strain number corresponds to the soil number from which the strain was isolated. 3 Fluorescent pigment production when grown on medium 8 of King et al. 1954; + = positive; - = negative. 4 + = growth; i = questionable growth; - = no growth. 65 Figure 7 (a-d). Viable bacterial counts in various kraft lignin (MW > 1500) media: a) medium I; b) medium II; c) medium III; d) medium IV. Viable counts were determined as de- scribed in the text. Vertical bars represent the standard deviation of each determination. Symbols: (0—0), series A: (om-4), series B; (Gm—0), series C. 66 q§\ht>3 g; x839 quhtg 95$ x839 TllVE (days) TIDE (don) VI. TVII. LplL PEP h P I Dbl-D b D h b DP... if b Pb.- 4 v w w w. ngSS 953$ c a .. z x a v v v ll # séxs 955 $39 “NEW 1M: (days) 67 b, the maximum population density was 4.7 x 107 organisms/ml. This suggested that even after three subcultures, a medium containing lignin as the sole carbon and energy source, and not containing any minerals, supports growth of relatively large bacterial populations. Glucose in the medium did not appear to have any stimulatory effect on lignin degradation (Table 5). The results showed that a maximum of 28% of the lignin was degraded in medium I (Series A), supplemented with glucose and minerals, as compared to 35% lignin degradation observed in medium 11. Analysis of culture samples from flasks of medium III in series C, after nine days incubation, showed that 10.3 i 0.3 mg of lignin was present in these flasks. The appar- ent increase in kraft lignin content might be due to the oxidation of the a-carbons of the side chains to carbonyl groups (Kirk and Chang, 1975). The introduction of a carbonyl group adjacent to an aromatic ring is known to increase the absorptivity coefficient of the resulting compound (Silverstein et al., 1974). These results indicated that although the bacterial population doubled three times (see Fig. 5 c) in 9 days, there was no detectable degradation of lignin based on UV spectral analysis. The IR spectra of the degraded kraft lignin recovered from medium III and medium II after 9 and 52 days of incubation, respec- tively, and the IR spectrum of the undegraded kraft lignin from un- inoculated control medium are presented in Fig. 7. Absorbance dif- ferences were evident in the spectra of both degraded lignins in the region of 1500-1800 cm"1 as compared to the spectrum of the undegraded lignin. The most pronounced difference between the spectra of the degraded (recovered from medium 11) and undegraded kraft lignin was 68 Table 5. Extent of kraft lignin degradation by a mixed bacterial culture in three different media in flasks of series A and B. Series A2 Series 82 Medium1 Lignin 3 % 4 Lignin 3 % 4 remaining Degradation remaining Degradation II (uninocula- ted control) 10.00 i 0.45 0.0 --- --- I 6.67 i 0.47 28.3 7.30 i 0.06 26.5 II 5.97 t 0.27 35.3 5.74 i 0.36 37.6 III 6.88 t 0.06 26.2 7.03 s 0.57 24.7 1Composition of different media is given in Table 2. 2Flasks in series A and B were incubated at 30°C for 35 and 52 days, respectively. See text for other details. 3Calculated by determining the absorbance at 283 nm of the cul- ture diluted in 50% aqueous dioxane and using an absorptivity coeffic- ient of 23.7 1 g'1 cm'l. Initially, 10 mg of kraft lignin was present per 20 m1 of medium in each flask. Values are expressed as mean i standard deviation. 4Values are corrected to account for losses due to sampling and withdrawl of the inoculum. 69 the appearance of a strong absorption band centered at «4720 cm-1 in the former but not in the latter (Fig. 8b). This absorbance is attributable to a stretching vibration of conjugated carboxyl car- bonyl groups (Hergert, 1971; Silverstein et al., 1974; Kirk and Chang, 1 1975). A comparable band at 1720 cm' was not evident in the sepctrum of the kraft lignin recovered from medium III (Series C) after nine days of incubation (Fig. 2a). The increased intensity of absorption 1 in the IR spectrum of the latter preparation centered near 3400 cm- suggested an increase in the number of hydroxyl groups (Hergert, 1971; Silverstein et al., 1974), but the reason for the increased absorbance 1 is not clear (Fig. 8a). Nonetheless, these between 1500-1800 cm- differences suggested that limited structural changes did occur in the kraft lignin during the nine day period of incubation. Figure 8: 7O 4 (a-b). Infrared spectra of degraded and undegraded kraft lignin. a) Lignin from uninoculated control medium (dashed line) and from medium III of series C, after 9 days of in- cubation (solid line); b) lignin from uninoculated control medium (dashed line) and lignin recovered from medium II of series B after 52 days of incubation (solid line). ABSORBANCE ABSORBANQ' 71 0Q) a. 1 T T fl 7 a” ’ ~ \ l \\ I’,"‘\\"’ \‘ \\ ’1”, \J“, 1‘ U~r\.\ . ‘v” 1 A 0.25 >- V g r 0.50 i 1 L ‘ ‘ 4000 3500 5000 2500 2000 I500 FREQUENCY (cm") 0.00 T T T I I . b. I-“ I.‘/’--”-‘~-----‘.. l.--— ”" \‘ '1’ 1‘ l' l’.‘ I \‘ \‘ /’ VI A 1' r \\d’/ V 0.25 - f 0.50 ‘ ‘ ‘ ‘ ‘ 4000 3500 3000 2500 2000 1500 FREQUENCY (cm'U CHAPTER V DISCUSSION The rate of organic matter turnover in tropical soils is gen- erally greater than that in temperate climates (Kalpage, 1974; Money, 1972). Therefore, it seemed reasonable to use soils from a number of tropical countries as inocula for enrichment cultures. The results show that of the 228 strains isolated from the kraft lignin enrichments, 22 strains rapidly degrade p-hydroxybenzoic acid, 4-hydr0xy-3-methoxybenzoic acid, and 4-hydroxy-3,5-dimethoxybenzalde- hyde. It was not very Surprising that all these 22 isolates, selected for further study, were found to be members of the genus Pseudomonas because pseudomonads, in general, are known to metabolize a large number of very diverse substrates, and are frequently involved in the biodegradation of recalcitrant compounds (Duodoroff and Palleroni, 1974; Stanier et al., 1966). Furthermore, the lack of vitamins or other growth factors in the isolation media may have given selective advantage to pseudomonads which are known to have Simple nutritional requirements. The results show no apparent correlation between viable bac- terial counts and the extent of lignin degradation in different media. In medium I containing lignin, minerals and glucose, the viable counts were considerably higher, but the extent of lignin degradation was much lower than that observed in an identical medium minus glucose. 72 73 The lower extent of lignin utilization seen in medium I may be due to the presence of glucose which may result in the selection of a population less effective in netabolizing lignin or the metabolic end products from glucose may repress the enzymes involved in lignin metabolism or other factors which are not clear at this time. Fur- thermore, these results indicate that, unlike Phanerochaete chryso- 14 spgrium which was reported to metabolize synthetic C-lignin only when the medium was supplemented with a growth substrate such as cellulose or glucose (Kirk et al., 1976), the bacterial mixed culture is able to degrade lignin, even when the latter was offered as the sole exogenously added carbon source. Although the extent to which lignin is utilized by the isolates as a source of carbon and/0r en- ergy is not clear, the viable counts in medium 11 (containing lignin plus minerals) were approximately 3 fold higher than those in medium IV (containing minerals only) suggesting that lignin was being used as a source of carbon/energy to a substantial extent. Sustained growth in medium IV suggests that certain strains in the mixed cul- ture are capable of autotrophic growth. However, other factors such as contaminating materials leaching from foam plugs and/or absorption of volatile organic compounds from the atmosphere and/or increase in cell numbers but not in cell mass may explain the observed in- crease in viable counts in medium IV. The level of growth observed in medium III was also unexpected in view of the fact that no min- erals were added to this medium. The source of nitrogen employed by the bacteria in this medium is not obvious at this time although at least a part of the nitrogen may be derived from dead cells. The data on viable bacterial counts in the f0ur different 74 lignin degradation media show that bacterial populations dramatically increase in these media during the first five days of incubation. For cultures of series A, this may be due to the transfer of small amounts of phenolic compounds present in the inoculum. However, similar results were obtained with the cultures of series 8 also, which were inoculated from cultures of series A, indicating that the transfer of phenolic compounds via the inoculum is not respon- sible for the initial sharp intrease in viable counts in series 8 flasks. Further results showed that little or no degradation of the ring structure of the kraft lignin occurred in medium III (series C) during the initial nine days of incubation. The sharp initial in- crease in cell population observed in the latter flasks may have been due to demethoxylation or metabolism of side chains of the lignin polymer. Similar observations were made by Ferm and Nilsson (1969) who were able to demonstrate the utilization of lignosulfonate as a sole source of carbon and energy by a Pseudomonas EB: but were unable to show disruption of the aromatic ring structure as evidenced by a decrease in absorbance at 280 nm. These data suggest that the degra- dation of lignin by the bacteria employed is a gradual process and does not occur to any large extent during the initial period of in- cubation. In this study, lignin in various preparations was quantitated by UV spectroscopy, a procedure previously utilized by other investi- gators (Johnson et al., 1961; Goldschmid, 1971; Odier & Monties, 1977). The absorbance maximum at 283 nm of lignin is attributed to the n-h* transition that occurs in the aromatic rings of the polymer (Aulin-Erdtman, 1968). Therefore, the decrease in abosrbance at 75 283 nm in the spectrum of degraded lignin as compared to that of the undegraded lignin can only be attributed to cleavage of the aromatic ring structures within the polymer. Furthermore, the lignin spectra in this study were obtained using 200-fold dilutions of cultures with no prior treatment to remove bacterial cells or extracellular protein. However, the presence of such materials would only con- tribute to the absorbance at 283 nm and thus to an overestimation of the amounts of lignin remaining in cultures. In other words, the percentages of lignin degradation reported in Table 4 may well be minimal values. Kirk and Chang (1975) observed an increase in absorbance cent- ered near 1720 cm'1 in the infrared spectrum of spruce lignin de- graded by Coriolus versicolor. These workers indicated that this absorption band was primarily due to carboxylic acid and/or ester groups within the degraded lignin. In agreement with the results of Kirk and Chang (1975), the degraded kraft lignin used in this study also showed a sharp absorption maximum at ml720 cm']. The marked increase in the carbonyl groups within the degraded kraft lignin residue, as revealed by IR spectroscopy, suggests that the degrada- tion of kraft lignin by the bacterial mixed culture is an oxidative process. The reuslts conclusively Show that a mixed culture of 22 strains of Pseudomonas, isolated from various tropical soils, are capable of metabolizing aromatic ring structures in the high nolec- ular weight kraft fraction of kraft lignin. In a medium containing lignin plus minerals, up to 37.6% lignin is degraded in 52 days. These results are comparable to those of Lundquist et a1. (1977) who 76 showed 41% degradation of (ring-‘40) kraft lignin in 67-78 days by a strain of E, chrysosporium. In comparison, Crawford et al. (1977) 14 reported that only 18% of the 14 C-maple kraft lignin and 7% of the C-cottonwood kraft lignin was degraded to 14 C02 after 31 and 25 days incubation, respectively, with a soil sample from the base of a wood chip pile. The present results, compared with those of Trojanowski et al. (1977) and Crawfbrd (1973), suggest that mutual- istic degradation of lignin by a consortium of bacteria results in increased level of degradation as compared with that observed by bacterial monocultures. To the best of our knowledge, the results of this study rep- resent the first demonstration of microbial degradation of lignin when the latter was offered as the sole source of added carbon/ energy in the medium. Glucose was neither required nor did it in- crease the extent of lignin degradation by the pseudomonads. LITERATURE CITED LITERATURE CITED Aulin-Erdtman, G. and R. Sanden. 1968. Spectrographic contributions to lignin chemistry. IX. Absorption properties of some 4-hydroxypheny1, guaiacyl, and 4-hydroxy-3,5-dimethoxyphenyl type model compounds for hardwood lignins. Acta Chem. Scand. 22:1187-1209. Cartwright, N. J. and K. S. Holdom. 1973. Enzymic lignin, its re- lease and utilization by bacteria. Microbios 8:7-14. Colwell, R. R. and W. J. Weibe. 1970. "Core" characteristics for use in classifying aerobic, heterotrophic bacteria by numer- ical taxonomy. Bull. Ga. Acad. Aci. 28:165-185. Crawford, 0. L., S. Flody, A. L. Palmetto, and R. L. Crawford. 1977. Degradation of natural and kraft lignins by the microflora of soil and water. Canad. J. Microbiol. 23:434-440. Crawford, 0. L. 1978. Lignocellulose degradation by selected streptomyces strains. Appl. Environ. Microbiol. 35:1041-1045. Duodoroff, M. and N. J. Palleroni. 1974. Pseudomonas, p. 217-243. Ig_R. E. Buchanan and N. E. Gibbons,’(eds.)l Bergey's manual of determinative bacteriology. Williams and Wilkins Co., Baltimore. Ferm, R. and A. C. Nilsson. 1969. Microbial degradation of a com- mercial lignosulfonate. Svensk Papperstidn. 72:531-536. Gamble, T., M. Betlach, J. M. Tiedje. 1977. Numerically dominant denitrifying bacteria from world soils. Appl. Environ. Microbiol. 33:926-939. Goldschmidt, 0. 1971. Ultraviolet spectra. Ig_K. V. Sarkanen and C. H. Lugwig, (eds.). Lignin: Occurrence, formation, struc- ture and reactions. Wiley-Interscience, New York. Hergert, H. L. 1971. Infrared spectra, p. 267-297. IQ_K. V. Sarkanen and C. H. Ludwig, (eds.). Lignins: Occurrence, formation, structure and reactions. Wiley-Interscience, New York. Jaschof, H. 1964. Preliminary studies of the decomposition of lignin by bacteria isolated from lignite. Geochim. Cosmochim. Acta 28:1623-1638. 77 78 Johnson, D. 8., W. E. Moore, L. C. Zank. 1961. The spectrophotometric_ identification of lignin in small wood samples. Tappi 44: 793-798. Kalpage, F.S.C.P. 1974. Tropical soils. Macmillan Co. of India Ltd. , New Delhi. King, E. 0., M. K. Ward, and D. E. Ramsey. 1954. Two simple media for the demonstration of pyocyanin and fluorescein. J. Lab. Clin. Med. 44:301-307. Kirk, T. K. 1971. The effects of microorganisms on Lignin. Ann. Rev. Phytopathol. 9:185-210. Kirk, T. K. and H. M. Chang. 1975. Decomposition of lignin by white rot fungi. II. Characteristics of heavily degraded lignins from decayed spruce. Holzforschung 29:56-64. Kirk, T. K., W. J. Bleam, W. F. Hackett and J. G. Zeikus. 1975. Preparation and microbial decomposition of synthetic 14C lignins. Proc. Nat. 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 by two wood- rotting fungi. Appl. Environ. Microbiol. 32:192-194. Kovacs, N. 1965. Identification of Pseudomonas pyocyanea by the oxidase reaction. Nature 178:903. Lai, Y. Z. and K. V. Sarkanen. 1971. Isolation and structural studies, p. 165-240. In K. V. Sarkanen and C. H. Ludwig, (eds). Lignins: Occurrence, formation, structure and reac- tions. Wiley-Interscience, New York. Lemon, H, W. 1947. The effect of alkali on the ultraviolet absorp- tion spectra of hydroxyaldehydes, hydroxyketones and other phenolic compounds. J. Amer. Chem. Soc. 69:2998-3000. Lundquist, K., T. K. Kirk, and W. J. Connors. 1977. Fungal degrada- tion of kraft lignin and lignin sulfonates prepared from syn- thetic 14C-1ignins. Arch. Mikrobiol. 112:291-296. Marton, J. 1971. Reactions in alkaline pulping, p. 639-694. Ifl_ K. V. Sarkanen and C. H. Ludwig, (eds.). Lignins: Occur- rence, formation, structure, and reactions. Wiley-Interscience, New York. Money, 0. C. 1972. Climate, soils and vegetation. University Tutorial Press, Ltd., London, England. Odier, E. and B. Monties. 1966. Activite lignionolytique in vitro de bacteries isolees de paille de ble en decomposition. C. R. Acad. Sci., Paris, Series D. 284:2175-2178. 79 Sarkanen, K. V. and H. L. Hergert. 1971. Classification and distri- bution, p. 19-94. IQ K. V. Sarkanen and C. H. Ludwig, (eds.). Lignins: Occurrence, formation, structure and reactions, Wiley-Interscience, New York. Silverstein, R. M., G. C. Bassler and T. C. Morrill. 1974. Spec- trophotometric identification of organic compounds. 3rd ed., John-Wiley and Sons, Inc., New York. Sorensen, H. 1962. Decomposition of lignin by soil bacteria and complex formation between autoxidized lignin and organic nitrogen compounds. J. Gen. Microbiol. 27:21-34. Stanier, R. Y., N. J. Palleroni, and M. Doudoroff. 1966. The aerobic pseudomonads: a taxonomic study. J. Gen. Microbiol. 43:159-271. Sundman, V., T. Kuusi, S. Kuhanen, S. Kilpi, H. Sederholm. 1968. Observations on bacterial utilization of the lignin from brown rotted spruce wood and Brauns' native lignin. Finska Kemists, Medd. 77:70-86. Trojanowski4 J., K. Haider, and V. Sundman. 1977. Decomposition of 1 C-labelled lignin and phenols by a Nocardia 3p. Arch. Mikrobiol. 114:149-153. West, M., N. M. Burdash and R. Freimuth. 1977. Simplified silver- plating stain for flagella. J. Clin. Microbiol. 6:414-419. nIcHIonN STATE uwfv. LIBRARIES 1|l11111111111111111111111111111111111111111111111 31293101757502