LIGNIN DEGRADING

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PROLOGUE

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Tropical soils are responsible for near complete decomposition of leaf plant litter in as little as 18 months (Parton et al., 2007). There is an apparent contradiction of tropical forest soils, where rapid and efficient lignocellulose mineralization proceeds rapidly under low or fluctuating redox conditions. Rapid decomposition may be fueled by fluctuating redox conditions that regenerate oxidized iron; up to 10% of tropical bacteria are capable of iron reduction (Dubinsky et al., 2010). Resident microbes are adapted to the low and fluctuating redox potential in the soil (Silver et al., 1999, in press; Pett-Ridge et al., 2006), in contrast to temperate systems where oxidative enzyme activities are rate-limiting for decomposition (Paul and Clark, 1996; Freeman et al., 2001; Fierer et al., 2009).

Plant biomass represents a renewable carbon feedstock that could potentially be used to replace a significant level of petroleum-derived chemicals. One major challengein its utilization is that the majority of this carbon istrapped in the recalcitrant structural polymers of the plant cell wall. Deconstruction of lignin is a key step

in the processing of biomass to useful monomers but remains challenging. Microbial systems can provide molecular information on lignin depolymerization as they have evolved to break lignin down using metalloenzyme-dependent radical pathways. Both fungi and bacteria have been observed to metabolize lignin; however, their differential reactivity with this substrate indicates that they may utilize different chemical strategies for its breakdown. This review will discuss recent advances in studying bacterial lignin degradation as an approach to exploring greater diversity in the environment.(http://www.sciencedirect.com/science/article/pii/S1367593113002342)

INTRODUCTION

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Lignin is an amorphic three-dimensional substance the molecular weight of which is difficult to determine because lignins are highly polydisperse materials ( Argyropoulos and Menachem, 1997).

The chemical structure of lignin has also been difficult to determine, and even very recently, new bonding patterns have been described in softwood lignin, e.g,. dibenzodioxocin structures. Also, the isolation of nativelignin is complicated if possible at all ( Buswell and Odier, 1987).

Lignin degradation is in a central position in the earth's carbon cycle, because most renewable carbon is either in lignin or in compounds protected by lignin fromenzymatic degradation (cellulose and hemicellulose)( Kirk, 1983).

Lignin biodegradation is also responsible for much of thenatural destruction of wood in use, and it may have an important role in plant pathogenesis. On the other hand, potential applications utilizing lignin-degrading organisms and their enzymes have become attractive, because they may provide environmentally friendly technologies for the pulp and paper industry and for the treatment of many xenobiotic compounds, stains, and dyes.(http://www.wiley-vch.de/books/biopoly/pdf/v01_kap05.pdf)

Over the past three decades, the activities of four kindsof enzyme have been purported to furnish the mechanistic foundations for macromolecular lignin depolymerization indecaying plant cell walls. The pertinent fungal enzymes comprise lignin peroxidase (with a relatively high redox potential), manganese peroxidase, an alkyl aryl etherase,and laccase. The peroxidases and laccase, but not the etherase, are expressed extracellularly by white-rot fungi. A number of these microorganisms exhibit a marked preference toward lignin in their degradation of lignocellulose. Interestingly, some white-rot fungi secrete both kinds of peroxidase but no laccase, while others that are equally effective express extracellular laccase activity but no peroxidases. Actually, none of these enzymes has been reported to possess significant depolymerase activity toward macromolecular lignin substrates that are derived with little chemical modification from the native biopolymer. Here, the assayscommonly employed for monitoring the traditional fungal peroxidases, alkyl aryl etherase, and laccase are described in their respective contexts. A soluble native polymeric substrate that can be isolated directly from a conventional milled-wood lignin preparation is characterized in relation to its utility in next-generation lignin-depolymerase assays.(http://www.ncbi.nlm.nih.gov/pubmed/22843404)

PATHWAYS

Pathways associated with (A) xylose degradation, (B) lignindegradation, the 4-hydroxyphenylacetate degradation pathway, apossible pathway of lignin catabolism, and (C) dissimilatorylignin reduction via the electron transport chain. For eachpathway, the number next to the protein ID denotes the fold-

level induction in lignin-amended compared to unamended growthconditions. All genes listed were statistically significantlyup-regulated in lignin-amended compared to unamended controls(http://www.frontiersin.org/files/Articles/59376/fmicb-04-00280-HTML/image_m/fmicb-04-00280-g002.jpg)

Lignin degradation pathways.

Enzymes involved in the degradation of lignin andtheir main reactions

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Enzyme Activity, Abbreviation Cofactor or Substrate,Main Effect or Reaction

™Mediator∫Lignin peroxidase, LiP H2O2, veratryl alcohol aromatic ring oxidized to cation radicalManganese peroxidase, MnP H2O2, Mn, organic acid as chela- Mn(II ) oxidized to Mn( III ); tor,thiols, unsaturated lipids chelated Mn( III ) oxidizes phenolic compounds to phenoxyl radicals; other reactions in the presence of additional compoundsLaccase, Lacc O2; mediators, e.g., hydroxyben-

phenols are oxidized to

zotriazole or ABTS phenoxyl radicals; other

reactions in the presence of

mediators

Glyoxal oxidase, GLOX glyoxal, methyl glyoxal glyoxal oxidized to glyoxylic acid; H2O2 productionAryl alcohol oxidase, AAO aromatic alcohols (anisyl, veratryl

aromatic alcohols oxidized toalcohol)

aldehydes; H2O2 productionOther H2O2 producing enzymes many organic compounds O2 reduced to H2O2

Lignin peroxidaseIn enzymology, a lignin peroxidase (EC 1.11.1.14) is an enzyme that catalyzes the chemical reaction 1,2-bis(3,4-dimethoxyphenyl)propane-1,3-diol + H2O2 \rightleftharpoons 3,4-dimethoxybenzaldehyde + 1-(3,4-dimethoxyphenyl)ethane-1,2-diol + H2O

Thus, the two substrates of this enzyme are 1,2-bis(3,4-dimethoxyphenyl)propane-1,3-diol and H2O2, whereas its 3 products are 3,4-dimethoxybenzaldehyde, 1-(3,4-dimethoxyphenyl)ethane-1,2-diol, and H2O.

This enzyme belongs to the family of oxidoreductases, specifically those acting on a peroxide as acceptor (peroxidases) and can be included in the broad category of ligninases. The systematic name of this enzyme class is 1,2-bis(3,4-dimethoxyphenyl)propane-1,3-diol:hydrogen-peroxide oxidoreductase. Other names in common use include diarylpropane oxygenase, ligninase I,

diarylpropane peroxidase, LiP, diarylpropane:oxygen,hydrogen-peroxide oxidoreductase (C-C-bond-cleaving). It employs one cofactor, heme.

Lignin peroxidase (LiP) plays a central role in the biodegradation of the plant cell wall constituent lignin.LiP is able to oxidize aromatic compounds with redox potentials higher than 1.4 V (NHE) by single electron abstraction, but the exact redox mechanism is still poorly understood. The finding in our laboratory that theCbeta-atom of Trp171 carries a unique modification led usto initiate experiments to investigate the role of this residue. These experiments, employing crystallography, site-directed mutagenesis, protein chemistry, spin-trapping and spectroscopy, yielded the following results:(i) Trp171 is stereospecifically hydroxylated at its Cbeta-atom as the result of an auto-catalytic process, which occurs under turnover conditions in the presence ofhydrogen peroxide. (ii) Evidence for the formation of a Trp171 radical intermediate has been obtained using spin-trapping, in combination with peptide mapping and proteincrystallography. (iii) Trp171 is very likely to be involved in electron transfer from natural substrates to the haem cofactor via LRET. (iv) Mutagenetic substitutionof Trp171 abolishes completely the oxidation activity forveratryl alcohol, but not for artificial substrates. (v) Structural changes in response to the mutation are marginal. Therefore the lack of activity is due to the absence of the redox active indole side chain.(http://www.ncbi.nlm.nih.gov/pubmed/11356137)

Manganese peroxidase

Lignin degradation by basidiomycetous fungi (e.g., Nematoloma frowardii, Phlebia radiata). Fungi utilize preferentially sugars and acetate residues from hemicelluloses as carbon and energy source, while lignin is unproductively degraded ("enzymatic combustion") and the major part of cellulose remains fairely unused giving white-rotted wood its characteristic appearance (modified after Hofrichter, M., 2000, BioSpektrum 6: 198-199)(http://www.kolumbus.fi/ilona.barlund/ilona.barlund/MartinsProjects.html)

Alkyl aryl etherase

Sphingomonas paucimobilis SYK-6 produces unique and specificenzymes, such asβ-etherases,O-demethylases, and ring fission dioxygenases, for lignin degradation. Cleavage ofarylglycerol-β-aryl ether linkage is the most important process in the lignin metabolic pathway ofS. paucimobilis SYK-6. We reported the genes (ligD, ligE, ligF) for enzymes that cleavedβ-aryl ether linkage of dimeric compounds in previous studies. In this study we synthesized the fluorescent high-molecular-weight lignin (UBE-DHP) by dehydrogenative polymerization. We investigated theβ-arylether cleavage ability of these enzymes produced in recombinantEscherichia coli. When UBE-DHP was incubated with LigF, 4-methylumbeliferone was released as a result ofβ-aryl ether cleavage of αO-methylumbelliferyl-β-hydroxypropiovanillone (compound III) incorporated in UBE-DHP. Here, we report thatβ-etherase ofS. paucimobilis SYK-6 can be expressed inE. coli and is able to cleave theβ-aryl ether linkage in synthetic high-molecular-weight lignin.(http://link.springer.com/article/10.1007%2FBF00770705)

Ligninase

Ligninase is the original term encompassing many different types of oxidative, extracellular fungal enzymes which catalyze the breakdown of Lignin which is commonly found in the cell walls of plants. Instead of the term ligninase, the term lignin-modifying enzymes (LMEs) should be used, since these enzymes are not hydrolytic but oxidative (electron withdrawing) by their enzymatic mechanisms. LMEs include peroxidases, such as Lignin peroxidase, Manganese peroxidase and Versatile peroxidase, and many phenol-oxidases of Laccase type. LMEs have been known to be produced by many species of so called white rot basidiomyceotus Fungi including: Phanerochaete chrysosporium, Ceriporiopsis subvermispora, Trametes versicolor, Phlebia radiata, Pleurotus ostreatus, and Pleurotus eryngii. LMEs are produced not only by wood-white rotting fungi

but also by litter-decomposing basidiomycetous fungi suchas Agaricus bisporus (common button mushroom), and many Coprinus and Agrocybe species. The brown-rot fungi, whichare able to colonize wood by degrading cellulose, are notable to produce LMEs. Some results on LME-type of peroxidases have also been reported for some species of filamentous bacteria such as Streptomyces viridosporus T7A, Streptomyces lavendulae REN-7 and Clostridium stercorarium. However, efficient lignin and lignin-like polymer degradation is only achieved by fungal LME peroxidases, and laccases in combinations with organic charge transfer mediator compounds. Laccases are more widely distributed enzymes belonging to the multicopper oxidase (MCO) superfamily encompassing all three domains of life (bacteria, archaea, eukarya).

Laccase

Laccase catalyzes the oxidation of phenol containing compounds, including lignin, through the reduction of oxygen to water. The presence of mediators will allow theoxidation of non-phenolic compounds as well. The primary function of laccase is to degrade lignin in fungi.

Laccases (EC 1.10.3.2) are copper-containing oxidase enzymesthat are found in many plants, fungi, and microorganisms.The copper is bound in several sites; Type 1, Type 2, and/or Type 3. The ensemble of types 2 and 3 copper is called a trinuclear cluster. Laccases act on phenols and similar molecules, performing a one-electron oxidations, which remain poorly defined. It is proposed that laccasesplay a role in the formation of lignin by promoting the oxidative coupling of lignols, a family of naturally occurring phenols. Laccases can be polymeric, and the enzymatically active form can be a dimer or trimer. Otherlaccases, such as ones produces by the fungus Pleurotus ostreatus, play a role in the degradation of lignin, and can therefore be included in the broad category of ligninases.Spectrophotometry can be used to detect laccases, using thesubstrates ABTS, syringaldazine, 2,6-dimethoxyphenol, and dimethyl-p-phenylenediamine. Activity can also be monitored with an oxygen sensor, as the oxidation of the substrate ispaired with the reduction of oxygen to water.Unit Definition One unit corresponds to the amount of enzyme which converts 1 μmole of catechol per minute at pH 6.0 and 25 °C

Form powderColor light brownStorage temp. 2-8°C

LIGNIN DEGRADING MICROORGANISMSLignin degradation by specialized microorganisms is relatively rapid. Ulmer and others (1983) found that lignin isolates were reduced to low molecular weight soluble products by the white-rot fungus Phanerochaete chrysosporium within 6 to 8 days. The enzymatic degradationof lignin by microorganisms generally involves both depolymerization and aromatic-ring cleavage (Crestini andothers, 1998). This degradation of lignin is brought about by extra-cellular oxidative enzymes. The action of these enzymes generally has been studied using lignin model compounds and lignin isolates that have been

mechanically and chemically freed from the surrounding hemicellulose matrix.

Crestini and others (1996) studied the degradation of lignin model compounds by the white-rot fungus Lentinus edodes. β–O-4 linkages were oxidized to arylglycerol compounds, and aromatic rings were cleaved. Cleavage of aromatic rings coupled with β–O-4 oxidation can lead to the formation of cyclic carbonate structures. The aromatic rings are cleaved by enzymatic systems that mostly follow the β–ketoadipate pathway. Harwood and Parales (1996) have reviewed the enzymatic reactions thattake place in the β–ketoadipate pathway. A wide variety of aerobic soil bacteria and fungi secrete enzymes that convert aromatic compounds to aliphatic carboxylic acids.The first step of the conversion involves mono- or di-oxygenation of the aromatic ring to produce a dihydroxylated phenyl ring. Fission of the ring can take place either between two adjacent hydroxyl groups (ortho-fission) or adjacent to a hydroxyl group (meta-fission). Examples of these reactions are shown in figure 2. Schemes I and II (ortho-fission) in figure 2 lead to the production of β–ketoadipate. Scheme III is the first stepof meta-fission, and Scheme IV is the first step in the gentisate pathway in which fission takes place between the carbon atom that is attached to a carboxylate group and an adjacent phenolic carbon. Schemes III and IV do not lead to the formation of β–ketoadipate (Fairley and others, 2002; Mohamed and others, 2001; Zaar and others, 2001). In addition to the degradation of lignin polymeric units,the microbial degradation of other phenolic compounds, aromatic hydrocarbons, aminoaromatic compounds, and chlorinated aromatic compounds often follow the β–ketoadipate pathway in soils. Fused ring aromatic compounds also can undergo degradation by the β–ketoadipate pathway (Hinter and others, 2001). Similarly,carboxylate-substituted aromatic compounds other than lignin units may be degraded by the gentisate pathway (Fairley and others, 2002; Mohamed and others, 2001; Zaarand others, 2001).

Lignin degradation has been extensively studied in fungi,which produce extracellular peroxidases/catalase that areable to degrade lignin (Wong, 2009). Similarly, several published studies also report soil bacteria that are ableto degrade lignin with the use of catalase or peroxidase enzymes. Streptomyces viridosporous, Nocardia autotrophica, and Rhodococcus sp. are well studied aerobic lignin degrading bacteria that produce extracellular peroxidase (Zimmermann, 1990). We found two peroxidase type proteins which are significantly up-regulated in lignin-amended sample: catalase/peroxidase HPI (Entcl_4301) and DypB-type peroxidase (Entcl_1327) (Figure 2B). The dyp type peroxidase protein family was identified in Rhodococcus jostii RHA1 (Ahmad et al., 2011) and was suggested for lignin degradation by β-aryl ether breakdown. This enzyme is activated by Mn2+ ions and was shown to degrade lignin andproduce monoaryl like 2, 6-dimethaoxybenzoquinone (Singh etal., 2013). However, the nature of the involvement of peroxide in anaerobic lignin degradation is still unclear.(http://journal.frontiersin.org/article/10.3389/fmicb.2013.00280/full)

LIGNINASEFungi Type I: Ceriporiopsis subvermispora, Phanerochaete chrysosporium, Phlebia radiata, Pleurotus eryngii.Pleurotus ostreatus, Pycnoporus cinnabarinusTrametes versicolor,

Fungi Type I: Agaricus bisporus (common button mushroom), Agrocybe sCoprinus

Filamentous bacteria Clostridium stercorarium

Streptomyces lavendulae NCIB 6959 NCIB 8235 ATCC 14158 Streptomyces viridosporus T7A ,

LACCASEFungi Type I: Pleurotus ostreatus

SUGGESTED SOLUTIONLive Spores of Phanerochaete chrysosporium NCIM1197Pleurotus ostreatus Streptomyces lavendulae NCIM 2498 NCIB 6959 NCIB 8235 ATCC 14158 Chaetomium cellulolytium (Moo-Young et al., 1988). Trametes versicolor NCIM 1086Sachharomyces boulardiiWith enzymes like Laccase, Ligininase, Manganese Peroxidase.

Suggested Level of Usage:1-2 g of this product / 20 Kg Biomass once; with about 16% Water addition daily for about 9 days.

ANALYTICAL METHODOLOGY

MoA_LIGNIN

Lignin monomeric units.

For lignin, better approach is to analyse the components in fresh control, exposed control, Treated samples using cupric Oxide Alkaline degradation (Hedges et al., 1988; Martinez et al., 1991).Products from the CuO degradation are separated and quantified using Gas Chromotography.With this technique, it can be determined of the relativeproportions (in Micromol per g of sample) of such lignin constituents such as guaiacyl, syringyl, cinnamyl and vanillyl phenols( Martinez et al., 1991; Alberts et al., 1992)Comparing molar ratios of different constituents shall beindicating the extent of oxidative degradation and the degree to which different constituents have been degraded.

Oxidative Enzyme AssaysTo perform measurements of oxidative enzyme activity, cells were grown as above in xylose minimal media, and then amended with L-3,4-dihydroxyphenylalanine (L-DOPA). L-DOPA is a lignin analog, where reduction causes a colorchange detectable colorimetrically (Saiya-Cork et al., 2002). For aerobic analysis, SCF-1 was grown in xylose minimal media broth for 12 h at 30°C with shaking at 200 RPM (foraerobic growth; no shaking for anaerobic growth) until an

average OD at 600 nm of 0.9 was reached, indicating late log phase based upon previous growth curves of this organism grown aerobically. For anaerobic analysis, SCF-1was grown anoxically in xylose minimal media broth for 24h until an average OD at 600 nm of 0.1 was reached, indicating late log phase based upon previous growth curves of this organism grown anoxically. For phenol oxidase and peroxidase assays, 25 mM L-DOPA substrate wasprepared the same day as analysis, with 3% H2O2 added for peroxidase assays. Phenol oxidase and peroxidase were also measured using 2,2′-azino-bis(3-ethylbenzothiazoline-6-sulphonic acid) (ABTS) based on a published protocol (Floch et al., 2007). The ABTS assays were prepared in the same way as for the L-DOPA assays, where 2 mM ABTS was prepared, and these assays performed only on aerobically grown cells. To measure enzyme activity, 500 uL of cell culture was combined with 500 uL of substrate. Time was recorded from the time substrate was added to cell culture. Measurements were made at absorbance at 460 nm. Each plate contained three biological replicates for each assay, with eight technical replicates (wells) for each. For each assay, negative controls included media, cell culture, and mediaand substrate, and signal OD was calculated as: [(Assay Value – Blank) – (Reference Standard – Blank)] where the blank was media only, and the reference standard was media + DOPA or ABTS. This accounted for any activity of trace metals in the media (i.e., Mn and Fe). ABTS rates are reported as mU (106 cells)−1, which is milliunits of ABTS (or 10−3 units) per million cells.(http://journal.frontiersin.org/article/10.3389/fmicb.2013.00280/full)

MoA of efficacy of Lignin degradation by Enzymesor Microbes:

The determination of 14CO2 evolution from 14 C-DHP (dehydrogenation polymer of coniferyl alcohol or other lignin precursors) or other 14C-(lignin)-lignocelluloses

has been the most often adopted method to determine lignin-degrading ability ( Haider and Trojanowski, 1975; Kirk et al., 1975, 1978).The method has been widely used for more than 20 years ( Buswell and Odier, 1987; Kirk and Farrell, 1987; Eriksson et al., 1990). Preparations from different laboratories have given comparable results ( Hatakka and Uusi-Rauva, 1983; Hatakka et al., 1983). Buswell and Odier (1987) compared different nonradioactive and 14C-labeled lignin preparations for microbiological studiesand concluded thatall lignin preparations have disadvantages in reproducibility in preparation, or altered structure and molecular weight compared to natural lignin. Nevertheless, the use of 14 C-labeled preparations can beconsidered the most reliable method. In addition, methodsto study the degradation of polymeric lignin by, e.g., NMR spectroscopy have been developed ( Davis et al., 1994; Gamble et al.,1994), but they are not easily amenable for detailed physiological studies with micro-organisms or biochemical studies with enzymes. Dimeric lignin model compoundsattached to a polymer backbone, e.g., polystyrene ( Lundell et al., 1992) or to moresoluble polymers, e.g., polyethylene glycol ( Kawai et al., 1995), have allowed to use efficient analytical tools (e.g., NMR ) for more defined structures.

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