Lignin Degrading Fungal Enzymes

Kameshwar, Ayyappa Kumar Sista; Qin, Wensheng

doi:10.1007/978-981-10-1965-4_4

Ayyappa Kumar Sista Kameshwar⁴ &
Wensheng Qin⁴

Part of the book series: Biofuels and Biorefineries ((BIOBIO))

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10 Citations

Abstract

Lignin is a complex heterophenolic biopolymer present in plant cell walls. It supports plants growth by providing structural integrity, impermeability, protection against pathogenic infection and pest damage. Being the second most abundant polymer on the earth, it plays a major role in carbon recycling. Increases in greenhouse gas emission and decreases in fuel reserves have increased interest all over the world for the production of biofuels from plant biomass. Because of its complex structure, lignin has become one of the major hurdles for biofuels production. Research has been conducted on methods to separate lignin from lignocellulosic biomass by employing chemical, physical or mechanical methods. However, these methods tend to be expensive and require much energy and also pose risks to the environment. The polyphenolic structure of lignin has attracted interest for the production of renewable and commercially valuable platform chemicals, thus studies have been conducted on its degradation by enzymes. This chapter summarizes recent advances in lignin degrading enzymes (Lignin Oxidizing and Lignin Degrading Auxiliary enzymes) produced by wood degrading fungi and bacteria. Structural and functional aspects of lignin degrading and their auxiliary enzymes are covered with a short note being made on genomic studies of lignin degrading fungi.

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Fungal Diversity and Enzymes Involved in Lignin Degradation

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Keywords

1 Introduction

Lignin is the most complex and abundant naturally occurring biopolymer present in plant cell wall s. It forms a tight matrix around the carbohydrate s, and it is closely associated with cellulose , hemicellulose and pectin, forming an intricate structure . Lignin provides several advantages to the plant cell wall such as mechanical strength supporting large plant structures , protection against microbial infections as it is hard to degrade, impermeability and stability against chemical and mechanical attacks [1]. Lignin is an aromatic polymer made up of three basic units: p-coumaryl alcohol (4-hydroxycinnamyl alcohol), coniferyl alcohol and sinapyl alcohol which are collectively called monolignols that are derived from phenylalanine (aromatic amino acid) [2]. Monolignols produce three phenylproponoid units, p-hydroxyphenyl (H), syringyl alcohol (S), guaiacyl (G) later these units are collectively joined to form the lignin polymer [3]. The percentage of the different phenylproponoids present in lignin varies based on the type of cell, taxa, wood , environment and developmental conditions. Lignin present in dicotyledonous angiosperms contains mainly G and S, with traces of H units. Gymnosperms, contain high G and low H and, grasses (Monocots) contain high H units with, comparable G and S units [3]. Lignin contains several interionic bonds such as alkyl-alkyl, aryl-alkyl and aryl-aryl, lignin also associates with plant cell wall polysaccharides which makes the breakdown and separation of lignin very difficult [4]. Separation of lignin from lignocellulosic materials is not possible without partial disruption of the lignocellulosic network. During plant cell wall polymerization several intermediates are produced and, these intermediates react with oligolignols, carboxyl and hydroxyl groups of glucuronic acids in hemicellulose units resulting in ethers and esters [5–7]. Research groups have tried to efficiently separate lignin from lignocellulosic biomass (cellulose , hemicellulose ) for production of cellulosic ethanol (biofuel s). Biofuel and paper pulp industries make use of cellulose for the production of paper and cellulosic ethanol , leaving behind hemicellulose and lignin as industrial effluents. The depolymerization of the polyphenolic chemical structure of lignin offers many opportunities for producing conventional phenolic compounds [8].

Several studies have showed that lignin is resistant to the microbial attack however a few groups of microorganisms belonging to bacteria and fungi are able to efficiently degrade lignin [9]. It has been reported that anaerobic processes fail to attack aromatic rings while aerobic processes tend to degrade lignin [7, 9]. Fungi are the most studied organisms for lignin degradation . Basidiomycota phylum consists of a wide range of wood degrading fungi , thus it is the largest wood degrading fungal group [10, 11]. Wood degrading fungi belonging to Basidiomycota phylum can be further divided into white, brown and soft rotting fungi based on their wood decaying patterns. In the Basidiomycota phylum, most of the wood degrading fungi belong to the Agaricales and Aphyllophorale s orders [12]. White rot fungi are considered to be the most efficient lignin degraders, but their degradation rates for lignin and cellulose in wood tissue vary considerably [13]. Some white rot fungi are able to selectively degrade lignin without degrading much cellulose while other white rot fungi are able to attack both lignin and carbohydrate s. However, fungi which are able to selectively degrade lignin are of higher significance due to their commercial applications (e.g. paper and pulp industries) [10, 13]. White rot fungi secrete a wide range of wood degrading enzymes that are involved in the breakdown of carbohydrate components (cellulose and hemicellulose) and lignin. White rot fungi secrete different lignin degrading enzymes such as lignin peroxidase , manganese dependent peroxidase , laccase , horse radish peroxidase , and dioxygenases such as protocatechuate 3,4 dioxygenase, 1,2,4-trihydroxybenzene 1,2-dioxygenase, catechol 1,2-dioxygenase, superoxide dismutase, glyoxal oxidase , glucose 1-oxidase, aryl alcohol oxidase , veratryl alcohol oxidase, pyranose oxidase and, quinone oxidoreductase [13, 14]. When compared to white rot fungi , brown rot fungi are more efficient in degrading cellulose and hemicellulose than lignin, however these fungi potentially modify lignin [10]. Wood affected by brown rot fungi generally appears shrunken and dark in color with brick and cubical shaped fragments, these fragments can be easily broken down further to a brown color powder (modified lignin). Wood decaying fungi belonging to the phyla Ascomycota and Deuteromycota are mostly soft rot fungi , these fungi usually decay wood , causing a light brown color. Based on the patterns they form on the wood these fungi can be further classified into type I (form biconical or cylindrical cavities in secondary walls) and type II (this type of fungi cause erosion). When compared to white rot fungi , type II soft rot fungi do not attack the middle lamella (Table 4.1) [10]. Some basidiomycete fungi form unique symbiotic associations with wood degrading termites belonging to Termitomyces, Bacteroidetes and Firmicutes classes. Some bacteria and flagellated protists also resides in the termite hind gut [7]. Both lower and higher termites maintain a remarkable microbial diversity in their guts and, some bacteria were found to colonize in hindgut, fore and midgut. Termites depend on their gut microbiota for their nutrition, as most of their nutrition is derived from the downstream products of microbial metabolism [7]. The underlying mechanisms behind the utilization and degradation of lignocellulose biomass by termites was only revealed recently. Genome sequencing and metagenome sequencing (termite gut microbiome) studies conducted on different lignocellulose degrading fungi have revealed several interesting facts about lignin degradation [15].

Table 4.1 Illustrates potential wood degrading fungal phylum and their properties

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2 Carbohydrate Active Enzyme Database (CAZy)

CAZy is a sequence based classification of the enzymes which are involved in the formation, modification and breakdown of poly and oligosaccharides [16]. There are three major defining features underlying CAZyme classification, (a) Classification is based on significant similarity of amino acid sequences with a minimum of one biochemically characterized founding member (b) CAZymes generally are modular proteins with a catalytic module containing different discrete units, thus it is classified module by module [17]. Third important feature CAZyme classification is based on systematical protein sequences upon daily GenBank releases which avoids analyzing unfinished protein sequences with changing accession numbers. The CAZy database is currently divided into two main classes as Carbohydrate active enzymes and Carbohydrate binding modules (CBMs). Carbohydrate active enzymes are further divided into five classes they are (a) Glycoside hydrolases (GH) (b) Glycosyl Transferases (GT) (c) Polysaccharide Lyases (PL) (d) Auxiliary Activity enzymes (AA) (e) Carbohydrate Esterases [16]. Carbohydrate binding modules (CBMs) earlier known as Cellulose binding modules, recent discovery of several new binding modules which bind to carbohydrate s other than cellulose was the main reason behind the name change to carbohydrate binding modules. The occurrence of lignin in close associations with other polysaccharides (cellulose and hemicellulose ) in plant cell wall s led to, lignin degrading enzymes and lytic polysaccharide monoxygenases (LPMO) to be classified into “Auxiliary activity enzymes” among the large class of enzymes involved in the modification and breakdown of lignocellulose . Auxiliary activity enzymes are currently classified into three subfamilies of polysaccharide monoxygenases and eight classes of lignolytic enzymes , a total of 1045 enzymes were classified into 13 subfamilies and 304 non classified enzymes [16]. CAZy database can be used to analyze the genomes (Carbohydrate active enzymes encoded in the genome of an organism CAZome), and also to study the molecular details of substrate recognition. CAZome can provide significant insights about the nature and amount of metabolism of complex carbohydrate s by a species. At present the CAZy database covers 4623 genomes in the following kingdoms they are: bacteria (3946), archaea (220), eukaryota (180) and viruses (277) [16].

3 Fungal Oxidative Lignin Enzymes (FOLy)

When compared to other microorganisms , fungi are the most efficient lignin degraders. Fungi secrete a wide range of extra and intracellular enzymes for the degradation of lignin. Lignin degradation by fungi is a significant step during carbon recycling and for maintaining terrestrial ecosystems. Thus, understanding the fungal enzymes involved in the breakdown of lignin is important. FOLy database was developed to classify the enzymes involved in the breakdown of lignin, by retrieving publicly available information from GenBank, Uniprot PDB, EMP PMD, and Pubmed [18]. The structure of FOLy is similar to that of Carbohydrate Active Enzyme (CAZy) [16]. Degradation of carbohydrates by enzymes is dependent on cocktails of highly specific extracellular enzymes for breaking glycosidic bonds. While lignin depolymerization makes use of various extracellular oxidative enzymes , which are responsible for generating highly reactive free radicals that cause cleavage of carbon-carbon, and inter unit ether bonds. FOLy has classified lignin breaking enzymes into two major classes as (a) Lignin Oxidizing (LO) enzymes and (b) Lignin degrading auxiliary enzymes (LDA ’s) , based on their potential involvement in lignin breakdown [18].

4 Lignin Oxidizing Enzymes (LO)

Non-specificity and high oxidation potential are the main attributes of lignin oxidizing enzymes. Lignin oxidizing enzymes are categorized into three classes LO1 (Laccases ), LO2 (Lignin peroxidase s, Manganese peroxidases , Versatile peroxidase s and Chloroperoxidases) and LO3 (Cellobiose dehydrogenase ) (Fig. 4.1). The most thoroughly studied fungal enzymes involved in lignin attack are described below:

4.1 Laccases (EC 1.10.3.2, Benzenediol: Oxygen Oxidoreductase)

Laccases represents the largest sub group of blue multicopper oxidases (MCO) and are widely distributed among eukaryotes (fungi , plants) prokaryotes (bacteria ) [22] They perform varied functions based on the source organism [22]. Laccase was first discovered in the sap of the Japanese lacquer tree Rhus vernicifera [23] and then it was also demonstrated in fungi [24]. Although laccases were discovered during early nineteenth century they have received much attention during the last five decades for their application to biofuel and biorefinery fields. The involvement of laccase in the degradation of wood by fungal groups such as basidiomycete s, ascomycetes has attracted scientific communities to study the structure , function and mechanisms of laccases [25]. Many fungal species belonging to the basidiomycetes phylum such as Abortiporus biennis, Agaricus bisporus, Agaricus brunnescens, Armillaria mellea, Aspergillus nidulans, Botrytis cinerea, Ceriporiopsis subvermispora, Ganoderma lucidum, Lentinus edodes, Myceliophthora thermophile, Neurospora crassa, Penicillium crysogenum, Phanerochaete chrysosporium , Phlebia brevispora, Phlebia radiata, Pleurotus erygii, Pleurotus ostreatus, Pleurotus sojar-caju Polyporus species, Rhizoctonia Solani, Trametes hirsuta, Trametes versicolor and Trichoderma were reported to secrete laccase [26]. Laccases are widely studied for two major functions (a) there role in lignin polymerization (lignification) in plants, (b) lignin depolymerization by fungi [27]. The contrasting role of laccases on lignin depolymerization was proved in vitro by Hatakka 1994 and Youn et al. 1995, showing the oxidative reaction of laccases on lignin, resulting in loss of an electron from phenolic hydroxyl groups of lignin resulting in phenoxy radicals [28, 29]. These studies have also showed that these radicals can spontaneously reorganize leading to the cleavage of alkyl side chains of polymer. At the same time, the polymerizing activity of the laccase might result in the polymerization of low molecular weight compounds [26]. These studies suggested that lignin degradation by fungi in nature occurs by the synergistic effect of other lignin degrading enzymes and non-enzymatic components which establishes a balanced environment between lignin depolymerization and enzymatic polymerization [26]. Although studies have reported the involvement of laccases in both lignin polymerization and depolymerization, the exact role of laccases and other partnering enzymes in the degradation and modification of lignin were still under investigation [26, 27]. Apart from wood decay, laccases play important role in fungal physiological processes such as morphogenesis, fungal plant pathogen/host interactions , stress defense and lignin degradation [26, 30]. In fungi , laccases are expressed during different stages of fungal development (morphogenesis, growth of rhizomorphs, sporulation, pathogenesis and virulence). According to Leatham and Stahmann 1981, increased laccase activity was observed in the developing fruiting bodies of Lentinus edodes (a commercially cultivable mushroom) [31]. The role of laccases on mushroom development was proved by Ikegaya et al. (1993), in this study the developing fruiting bodies of L. edodes were treated with diethyldithiocarbamate (a potential inhibitor of laccase ) which resulted in the decreased growth of L. edodes fruiting bodies, thus proving the role of laccase in fungal development [32]. A similar study was conducted on Armillaria mellea by Worrall et al. (1986) which showed the requirement of laccase for the development and growth of rhizomorphs [33]. Laccases are also involved in imparting specific virulence properties to the fungi , Botrytis cinerea (common plant infecting fungi ) secretes laccases which causes infection in some plants especially carrot and cucumber by triggering plant toxins such as cucurbitacins and tetracyclic triterpenoids. However, the virulence of these laccases was inhibited in EDTA pretreated plant tissues [34]. Thus fungal laccases play three major functions: lignin degradation , detoxification and pigment formation. Industrially laccases are important in paper and pulp, bio bleaching, textile industries etc.

Structure

Several X-ray crystallography studies were conducted to determine the structural properties of laccases (Table 4.2), however in this chapter we are focusing on Trametes versicolor laccase . The structural analysis of laccase isolated from Trametes versicolor was first done by Klaus Pointek et al. (2002) [35]. Laccase is a monomeric protein ordered in three sequentially arranged domains with dimensions ranging 65 × 55 × 45 Å³, each domain consists of a β-barrel shaped architecture which is similar to other blue copper proteins such as azurin or plastocyanin [35]. With each domain different from the other domains in their structural composition. First domain comprises two four-stranded β-sheets and four 3₁₀-helices, of which three acts as a connecting peptide between β-strands and one helix forms the segments between first and second domains. The second domain consists of one six stranded and one five stranded β-sheets, with three 3₁₀-helices peptides connecting the individual β-strands and domains 1 and 3. A 3₁₀-helix forms a 40 amino acid long extended loop region between domains 2 and 3. The third domain contains two five stranded and a two stranded β-sheet, which together form a β-barrel. The β-barrel together with an α-helix and a β-turn, forms a cavity for a type-I copper . Compared to the other two domains, the third domain has the highest helical content with one 3₁₀-helix and two α-helices situated in between the connecting regions of different β-sheets. The completion of the protein fold involves the C-terminal end of domain 3 and three sequentially arranged α-helices. Two disulfide bridges (Cys85-Cys488 and Cys117-Cys205) were reported which connects domain 1 and 2 and stabilizes a 13 amino acid residue α-helix. An oxygen reducing site is present at the T2/T3 cluster which accesses the solvent through two channels, leading to type-III copper and to type-II copper sites. The type-II copper site is more exposed and easily altered, when compared to the other two copper sites present near the T3 site. Actually T2 copper site is deficient in copper in the copper depleted forms of both laccase and ascorbate oxidase [36, 62]. Electrostatic potential studies of Trametes versicolor laccase shows that it possess a high negative charge, which suggests the specific binding of the substrate to the negatively charged cavity near the T1 copper site. Negative charge occurring at binding site imparts functional significance to the enzyme by imparting stability to the radical cation products formed during catalytic cycle. The two channels near the copper sites provide access to molecular oxygen and allow water release from the T2/T3 cluster. At the same time a conserved His-Cys-His tripeptide is associated with the electron transfer pathway between the T1 copper and the trinuclear cluster. A two site ping pong bi-bi reaction mechanism was proposed for the catalytic mechanism of laccase enzyme [63], which means that products are released before binding of new substrates.

Table 4.2 Lists the catalytic mechanism and structural studies of different lignin oxidizing enzymes

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Mechanism of Action

Laccases use their distinctive redox ability of copper ions for catalyzing the oxidation of various aromatic substrates concurrently reducing the molecular oxygen to water [64]. Laccases are able to catalyze direct oxidation of ortho, para-diphenols, aminophenols, polyphenols, polyamines , aryl diamines and also some inorganic ion [26, 65–69]. Laccases depends on copper (Cu) for their catalytic action, based on the number of copper ions laccases can be classified as dimeric or tetrameric glycoproteins. In addition, based on the types of copper ion centers they are classified as: (a) Type-I (blue copper center) (b) Type-II (normal copper center) (c) Type-III (coupled binuclear copper center) that differ in their characteristic electronic paramagnetic resonance (EPR ) signals [70, 71]. Type-I copper coordinates with four amino acids as ligands: two histidines, one cysteine and one methionine. Type-I copper containing laccases are generally a deep blue color, which can be detected by its absorbance at 600 nm wavelength . However, laccases which fail to absorb at 600 nm were reported in Pleurotus ostreatus (called white laccase ) [72] Panus tirinus (called yellow laccases ) [73]. Type-II copper coordinates with two histidine and water as ligands, Type-III copper coordinates with three histidines and a hydroxyl bridge which imparts strong anti-ferromagnetic coupling between the type-III copper atoms [35]. Type-II copper atoms do not absorb in the visible spectrum, while type-III copper atoms have an electron absorption at a wavelength of 330 nm. Based on the structural properties of type of copper ions laccases are divided into high and low redox potential enzymes. Bacteria and plants secrete low redox potential laccases , whereas white rot fungi and some basidiomycete s secrete high-redox potential laccases [74, 75].

Different copper centers present in the laccase participate and complete the enzymatic reaction. Unlike peroxidases laccases does not require hydrogen peroxide for the oxidation of monolignols . Enzyme catalysis can be divided into three main stages: the copper ion of type-I is reduced by the reducing substrate followed by internal electron transfer between the type-I, type-II and type-III Cu clusters [22]. Finally the reduction of oxygen takes place at the type-II and III Cu’s resulting in water formation (Fig. 4.2). In vitro lignin degradation by laccase primarily oxidizes phenolic hydroxyl groups of lignin to form phenoxy radicals which further reorganize to cleave the alkyl side chains . Laccase can degrade β-1 and β-O-4 dimer linkages between Cα-Cβ and cause Cα oxidation and aryl-alkyl cleavage s [22]. Thus the generated reactive radicals further release monomers by breaking down covalent bonds [76]. Due to the steric hindrance of laccase it cannot directly contact large polymers, thus small organic compounds or metals such as veratryl alcohol , manganese and 3-hydroxy anthranilic acid are oxidized and further activated to mediate radical catalyzed depolymerization of lignin [9, 76].

4.2 Peroxidases (EC:1.11.1.x)

Peroxidases are large group of enzymes widely distributed among plants, animals and microbes. Peroxidases play a wide variety of activities based on the source of the organism. Peroxidases are involved in several physiological processes such as plants defense mechanisms (response to pathogens), wound healing, auxin catabolism, lignification and suberization [77]. Microbes such as fungi and bacteria are well known for their ability of delignification which is efficiently fulfilled by the different types of peroxidases such as (LiP, MnP and VP). Peroxidases can also efficiently decolorize synthetic dye s and bioremediation of waste water and degradation of several toxic chemicals such as phenolic contaminants, polychlorinated biphenyls , chlorinated alkanes and alkenes, chlorinated dioxins, chlorinated insecticides and removal of endocrine disruptive chemicals etc, thus playing variety of roles in the environment [78]. Molecular structures of lignin degrading peroxidases share several common characteristics such as [79], Ligninolytic peroxidases generally contain a haem cofactor located internally in a cavity (haem pocket), which is connected to the protein by two small access channels [79–83]. Larger channel are common among all haem peroxidases , they are required for the hydrogen peroxide to reach the haem and react with (Fe⁺³) forming an activated two electron enzyme form called compound I [79–83]. The entrance of this channel forms the substrate binding site in some peroxidases . A second channel extends to the heme propionate substrate where some specific lignolytic enzymes oxidize Mn²⁺ and Mn³⁺ which acts a diffusible oxidizers of phenolic lignin and other organic molecules [79–83]. In this section we will be focusing on the delignification mechanisms of lignin peroxidases , manganese peroxidases , and versatile peroxidase .

4.3 Lignin Peroxidases (E.C. 1.11.1.14)

Lignin peroxidases (LiP) the most studied lignin depolymerizing enzymes, LiP was first discovered in the extracellular medium of P. chrysosporium under nitrogen limited conditions [84]. Similar to classic peroxidases , LiP are dependent on hydrogen peroxide . The overall reaction mechanism of LiP is 1,2-bis(3,4-dimethoxyphenyl) propane-1,3-diol + H₂O₂ ⇌ 3,4-dimethoxybenzaldehyde + 1-(3,4-dimethoxyphenyl)ethane- 1,2-diol + H₂O [9]. LiP can oxidize a wide range of phenolic compounds, organic compounds and also different lignin model non-phenolic compounds by using hydrogen peroxide with a redox potential up to 1.4 V, thus showing its non-specificity towards substrates [85].

Structure

Structure of P. chrysosporium lignin peroxidase (LiP) was determined using protein crystallography by Edward, et al. (1992). These studies revealed that fungal LiPs are globular and mostly helical glycoproteins with a molecular size of 38-46 kDa with 343–344 amino acids depending on the isozyme with a reduction potential exceeding 1.4 V [9, 42]. Compared to other lignolytic peroxidases it has a characteristic low optimum pH 3 with pI values ranging between 3.2 and 4.0 [9]. LiP contains several N- and O- glycosylation sites, it was reported that N-glycosylated sites (Asn257) and O-glycosylated site (Ser334 and Thr320) were clustered at the end of the proximal domain [43]. LiP is a globular protein with proximal C-terminal and distal N-terminal domains, the C-terminal segment of about 50 amino acids is extended to pass through the surface with less contact to the protein core [9]. LiP consists of eight major and eight minor α-helices and limited β structure in the proximal domains, it also has eight Cys residues forming disulfide bonds. The calcium binding site present in each domain is involved in maintaining the topology of the active site. The peroxide binding site is located on the distal end of the heme with an extended channel towards the exterior of the protein . The negative charge developed as a result of peroxide cleavage is stabilized by the Arg43 residue , it also stabilizes the ferryl oxygen of compound I. At the same time His47 with Asn82 present on the distal end of the enzyme acts as a proton acceptor for the bound peroxide substrate [9]. The overall protein fold of LiP is similar to other typical haem peroxidases such as cytochrome c peroxidase, manganese peroxidase, horseradish peroxidase [42]. The haem moiety divides the protein structure of LiP into a proximal and a distal domain. The haem moiety is hidden inside the protein with limited access to the outer medium by a small channel [43]. Thus the crystal structure of LiP shows that the haem access channel is not sufficient to allow entry to large polymers like lignin, however this is the only channel to form a suitable binding site for the attachment of small molecule substrates [43]. Protein modeling studies have confirmed its suitability for binding of veratryl alcohol , however the exact binding site of veratryl alcohol was not determined. LiP possess two substrate binding sites for veratryl alcohol (VA), the first one is Trp 171 and the second one is the anionic substrate oxidation site [43]. To find the binding site of the VA on LiP, the molecular docking study was conducted by chemically modifying the surface of LiP enzyme with 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide (EDC) in the presence and absence of 2-aminoethanesulfonic acid for the introduction of N-acyl-urea groups in place of carboxyl groups. LiP was also modified by N-bromosuccinimide (NBS) to yield a Trp modified enzyme [86]. From these studies it was shown that VA probably binds to Trp171 or its surrounding area as a reducing substrate and enzyme bound mediator . At the same time it has been suggested that VA binds at different locations when it reacts with LiP compound III* for the reverse reaction. The major difficulty in determination of the VA binding site is the lack of inhibitors causing classical inhibition patterns like competitive or non-competitive inhibitions. Thus, studies should be conducted by using X-ray crystallography and NMR methods for the determination of the LiP binding site for VA [9, 86].

Mechanism

Lignin peroxidase resembles horse radish peroxidase (a classical peroxidase highly studied) by containing Fe (III) as a cofactor which is pentacoordinated to four heme tetrapyrrole nitrogen s and to a histidine residue [87]. Lignin peroxidases are dependent on H₂O₂ for their reaction. H₂O₂ oxidize LiP resulting a two electron-oxidized intermediate (Compound I) in which iron is present as Fe (IV) leaving a free radical on the tetrapyrrole ring or on a nearby amino acid . Compound I then oxidizes a donor substrate to form a second intermediate (Compound II) and a substrate free radical (Fig. 4.3) [87]. Later reduction of the enzyme to its resting state can be accomplished either by the same substrate molecule or with a second substrate molecule by giving off substrate-free radical [87]. An important functional difference between LiP and other classical peroxidases is that lignin peroxidases can oxidize aromatic rings that are moderately activated by electron donating substituents, at the same time classical peroxidases act only on strongly activated aromatic substrates. Therefore, LiP and horseradish peroxidase can oxidize 1, 2, 4, 5-tetra-methoxybenzene, phenols and anilines, at the same time LiP are capable of abstracting an electron from aromatic s that carry only two or three ether like the major nonphenolic structures of lignin [88]. Primary products of this oxidation are temporary cation radical intermediates which certainly breakdown. Majorly Cα-Cβ bonds of propyl side chains are broken down to give benzaldehydes which are the precursors of benzoic acid molecules, these benzoic acid molecules are mainly observed in lignin decaying white rot fungi (Fig. 4.4) [89]. The unusual activity of lignin peroxidases is due to two structural differences, an electron-deficient iron atom in the porphyrin compared to classical peroxidases which makes LiP a stronger oxidant [90] and an invariant Trp171 in the isozyme of LiPA. This residue is present on the enzyme surface and is known to participate in a wide range electron transfers from aromatic substrates since they cannot contact the oxidized haem directly [91]. This important feature of LiP is responsible for oxidizing complex lignin and its related substrates directly. This function of Trp171 in LiPA was proved by a site directed mutagenesis in which the Trp171 was replaced by serine, which resulted in the loss of activity [92]. It was shown that the efficiency of LiP catalyzed oxidation of lignin molecules markedly decreases with an increase in size of the lignin molecule. LiP catalyzed oxidation of lignin trimers was found to be only 4 % of the rate of oxidation of a monomer model [93]. Oxidation of lignin molecules by LiP takes place in the presence of veratryl alcohol , and the role of VA in oxidation by LiP are given below [87]. Studies have showed that the VA cation radical oxidizes has a long half-life of 40 ms even at acidic conditions [94, 95]. VA is the substrate of LiP. It was suggested that the VA cation radical oxidizes lignin molecules at remote locations [96]. VA acts as an efficient electron donor to protect LiP from oxidative inactivation by hydrogen peroxide . As LiP oxidizes large and complex lignin substrates, which is a slow reaction, VA prevents oxidization of LiP [97]. VA is also essential for the reduction of LiP compound II. Compound I is reduced by non-methoxylated lignin structures . As these lignin structures are difficult to oxidize since they carry only one electron donating ether group. Compound II of LiP is a comparatively weaker oxidant than compound I [98].

4.4 Manganese Peroxidase s (EC 1.11.1.13)

Wood decaying white rot fungus and other litter decomposing fungi efficiently degrade lignin in wood . These fungi secrete several non-specific oxidoreductases, among them manganese peroxidase plays an important role [99]. Manganese peroxidase (MnP ) was first discovered in P. chrysosporium two decades ago [100, 101], however it received less attention then lignin peroxidase in beginning. Later it was found that LiP is not produced by all white rot fungi [28, 102, 103]. Production of MnP is limited only to basidiomycete s. Mainly two ecophysiological groups of fungi i.e. wood degrading fungi causing white rot and soil litter decomposing fungi secrete manganese peroxidase [103]. Wood decaying fungi belonging to families such as Meruliaceae, Coriolaceae, Polyporaceae and soil litter decomposing fungi such as Strophoriaceae, Tricholomataceae are known fungal families, which secrete MnP . Some prominent MnP producing fungi are Abortiporus biennis, Agaricus bisporus, Armillaria mellea, Auricularia sp. M37, Bjerkandera adusta, Ceriporiopsis subvermispora, Coriolopsis polyzona, Dichomitus squalens, Ganoderma lucidum, Heterobasidion annosum, Hypholoma fasciculare, Lentinula (Lentinus) edodes, Panus tigrinus, Phaeolus schweinitzii, Phallus impudicus, Phanerochaete chrysosporium , Phanerochaete sordida, Phlebia brevispora, Phlebia radiata, Pleurotus enryngii, Pleurotus sajor-caju, Stropharia aeruginosa, Stropharia coronilla, Trametes hirsuta, Trametes versicolor [99].

Structure

The enzyme mechanism of MnP is similar to that of classical haem containing peroxidases , but this enzyme is unique by having Mn⁺² as a reducing substrate [49]. Crystal structure of P. chrysosporium MnP was demonstrated by Sundaramoorthy et al. (1994). MnP is an acidic glycoprotein with pI near to 4.5, it is often produced as a sequence of isozymes which are differentially regulated by different genes [104]. It contains one molecule of heme in iron protoporhyrin IX, showing a maximal activity at Mn (II) with concentrations above 100 μM [100]. The enzyme oxidizes Mn from Mn⁺² to Mn⁺³, it forms complexes with oxalate and acts as a diffusible redox mediator for the oxidization of lignin and other phenolic compounds. MnP is a haem containing glycoprotein with a molecular weight ranging between 38 and 62.5 kDa , however most of the purified enzymes have molecular weights around 45 kDa . About 43 % of the amino acid sequence of MnP is identical to LiP, overall protein folding of MnP is similar to that of other plant and fungal peroxidases [49, 50]. MnP consists of two domains and haem submerged between these domains, similar to that of LiP. MnP enzyme consists of ten major and one minor helix similar to that of LiP. Eight minor helices are present in MnP of which two minor helices are similar to that of LiP all are in the 3₁₀ helical confirmation. It contains five disulfide bonds of which Cys³-Cys¹⁵, Cys³³-Cys¹¹⁷, Cys¹⁴-Cys²⁸⁹, Cys²⁵³-Cys³¹⁹ are similar to that of LiP and only one disulfide bond is unique to MnP [49, 50]. The unique disulfide bond Cys³⁴¹-Cys³⁴⁸ is part of the long C-terminal tail in MnP which is partially responsible for forcing C-terminus away from the main body of the protein , which might be involved in manganese binding site formation. Arg⁴² and His⁴⁶ on the distal domain of MnP form the peroxide binding pocket, and histidine in the distal domain is required for the formation of compound I by acting as acid base catalyst [20, 49]. Superposition studies conducted on LiP and MnP have shown few significant facts such as differences occurring at the insertion sites of these enzymes. In LiP, the C-terminus lies between two heme propionate groups while in MnP the C-terminus is separated from the heme by the combined effects of a 7-residue insertion (Leu²²⁸-Thr²³⁴) in the loop between the G and H helices, Arg¹⁷⁷, Glu³⁵, and the Cys³⁴¹-Cys³⁴⁸ disulfide bond [20, 50]. It was also reported that the His46 and Asn80 residues present in the distal end are hydrogen bonded, which is required for Nε₂ of His46 to accept proton from the peroxide during acid-base catalysis . Similarly the hydrogen bond between His173 and Asp242 involved in increasing the anionic character of the ligand and further stabilizing the oxyferric iron in MnP -I. The cation binding site present on the surface of the protein is formed by several interactions of Mn (II) with the carboxylate oxygen groups of Glu35, Glu39, and Asp179, the heme propionate oxygen and two water. The oxygens impart flexibility and facilitate the binding of a wide range of metal ions . MnP also contains of two calcium ions bound tightly on the proximal and distal side chains of the heme that are involved in maintaining the thermal stability of the active site of MnP [9, 20, 49, 50].

Mechanism

MnP is different from other peroxidases as it uses Mn (II) as the reducing substrate. MnP oxidizes Mn (II) to Mn (III), which then catalyzes the oxidation of a wide range of monomeric phenols , lignin model phenolic compounds and dye s [100, 105, 106]. The reaction mechanism of MnP proceeds as: first oxidation of Mn (II) by compound I (MnP -I), followed by oxidation of compound II (MnP -II) yielding Mn (III). MnP is a strong oxidizing agent like LiP, it cannot oxidize nonphenolic lignin related compounds because it lacks the invariant Trp171 residue which is required for electron transfer to aromatic substrates [87]. MnP has a manganese binding site which contains many acidic amino acids and also a heme propionate group. Thus one electron transfers to compound I of MnP takes place from bound Mn⁺². Further Mn⁺³ is released from the active site in presence of the bidentate chelators such as oxalate, which helps prevent the disproportionation to Mn⁺² and insoluble Mn⁺⁴. This reaction is required for the transfer of oxidizing power of MnP to Mn³⁺, which diffuses into the lignified cell wall thus attacking it from inside [87]. An important feature of MnP is to oxidize the low permeable lignocellulose network making it different from other peroxidases [9]. Chelators such as oxalate increase the electron density on Mn⁺³ which makes it a weak oxidant, thus Mn⁺³ organic acid chelates produced by MnP cannot oxidize the nonphenolic substrates of lignin. Mn⁺³ chelates cannot cause extensive lignolysis as they can only attack rare phenolic structures of lignin, which often are the end groups of lignin. The catalytic cycle of MnP begins with the binding of hydrogen peroxide or an organic peroxide to the native ferric enzyme resulting in the formation of an iron-peroxide complex (Fig. 4.5). Further the breakdown of the oxygen-oxygen peroxide bond depends on a 2-electron transfer reaction from the heme resulting in the formation of MnP compound I (i.e. a Fe⁴⁺-oxo-porphyrin radical complex). The dioxygen bond is cleaved resulting in removal of water and further reduction proceeds via MnP compound II. The Mn²⁺ ion (monochelated) donates one electron to the porphyrin intermediate and is oxidized to Mn³⁺. Similarly compound II is reduced by releasing another Mn³⁺ and a second water molecule, thus leading to the resting state of the enzyme (Fig. 4.5) [107–109].

The oxidation of phenolic compounds by MnP occurs by Mn (III) chelator complexes, which diffuses and catalyzes one electron oxidation of phenolic compounds producing a phenoxy radical intermediate . The phenoxy radical intermediate undergoes bond cleavage s , rearrangements and degradation of compounds non-enzymatically to produce different breakdown products [106, 110, 111]. In contrast unchelated Mn (III) causes the formation of reactive radicals as second mediators for the oxidation of non-phenolic compounds. Oxidation of non-phenolic compounds by MnP is different from LiP, as LiP oxidizes by abstracting electrons from the aromatic ring resulting in a radical cation. In presence of thiols like glutathione , Mn (III) causes the oxidation of benzyl alcohol and diarylpropane structures to their corresponding aldehydes [112, 113].

4.5 Versatile Peroxidases

Versatile peroxidases a new family of lignolytic peroxidases were reported for the first time in P. chrysosporium along with other lignolytic enzymes such as LiP and MnP [79]. Several fungi belonging to genera such as Pleurotus, Bjerkandera, Lepista, Panus and Trametes species were reported to produce versatile peroxidase (VP). Versatile peroxidase have important properties which combines the substrate specificity characteristics of the three fungal peroxidases such as manganese peroxidase, lignin peroxidase and Coprinus cinereus peroxidase [79]. Two well-known studies have revealed the occurrence of versatile peroxidase s in nature, in the first study a Mn²⁺ binding site was introduced into the LiP of P. chrysosporium by site directed mutagenesis, the resulting enzyme had MnP activity [114]. In the second study a tryptophan residue similar to that in LiP was introduced into the MnP of P. chrysosporium and the enzyme acquired LiP activity [115]. Versatile peroxidase coding genes were first cloned and sequenced from Pleurotus eryngii. Studies of the catalytic properties of VP suggested that they were due to its hybrid molecular construction combining different oxidation and substrate binding sites [116, 117].

Structure

Crystallographic studies conducted by Boada et al. (2005) showed that the structures of VP and its variant W164S are very similar. A total of 319 (VP) and 320 (W164S) amino acid residues along with heme and other cations were shown in the structural studies [53, 79]. The VP structure includes 11 α-helices (Ala12- Asn27, Glu36-Ala49, Ser64-Glu72, Ile81-Lys94, Ala99-Ser112, Val145-Ala155, Pro159-Ile171, Gln196-Glu200, Gln229-Arg236, Ala241-Ser246 and Gln251-Ala266), four disulphide bridges (Cys3- Cys15, Cys14-Cys278, Cys34-Cys114 and Cys242- Cys307) and two structural Ca²⁺ ions [79]. The Mn²⁺ binding site of VP contains three amino acid residues : Glu36, Glu40 and Asp175, The binding site is formed by the carboxylate groups of these amino acids and also the carboxylate group of the heme propionate. It is known that oxidation of compounds by VP occurs through LRET (Long Range Electron Transfer ) pathways, structural studies reveal that three putative LRET pathways were present in the VPL isoenzyme for the oxidation of aromatic compounds. They are (i) His232-Asp231 via backbone atoms and H-bond from the carboxylate of Asp231 to the side chains of the proximal His169, (ii) backbone atoms between Trp164 and Leu165 and a van der Waals contact between C^β of Leu165 to the methyl group C of the heme, and (iii) Pro/His76-Ala77– Asn78 via backbone atoms and a H-bond from the side-chain oxygen atom of Asn78 to the distal His47 [79]. From site directed mutagenesis studies of the LRET pathways clearly show that the Trp164 pathway of VP is involved in the oxidation of high redox potential compounds such as veratryl alcohol and reactive black 5. Studies also showed that oxidation of veratryl alcohol and reactive black 5 and other high redox potential compounds by VP involves electron transfer from activated VP* through the Trp164 side chain to the backbone and then to the side chain of Leu165, whose Cβ is 3.66 A ° from the carbon of the methyl-C of heme (2.37 A° hydrogen distance) [79]. A comparison with LiP isoenzymes shows either Met (LiPH8) or Leu (LiPH2) are homologous with VP Leu165. Eight neighboring residues with three conserved regions are shown to illustrate the differences between the structures , as well as the position of the proximal histidine residues , whose Nε atoms act as ligand of heme iron at 2.11 A° in VP and 2.15 A° in LiP [53–56, 79].

Mechanism

Basic features of versatile peroxidase are similar to those of all other classical peroxidases , however it is unique as far as the substrates that it is able to oxidize. A complete catalytic cycle combining those of other fungal peroxidases such as LiP and MnP was proposed by Ruiz-Duenas et al. Similar to LiP, versatile peroxidase also initiates the LRET pathway (Long range electron transfer) at an exposed tryptophan residue [79, 118]. Studies have examined the catalytic mechanism of VP using veratryl alcohol (reducing substrate) and its transitory states in the catalytic cycle. On reaction with one molecule of hydrogen peroxide the ferric group of VP (resting state) was converted to Compound I (Fe⁴⁺-oxo-porphyrin ⁺ complex) causing spectral changes (Fig. 4.6) [54, 55, 79]. Compound I oxidizes a molecule of veratryl alcohol resulting in Compound II (Fe⁴⁺-oxo), which will further oxidize another molecule of veratryl alcohol further reducing the enzyme back to its resting state [54, 79]. VP can oxidize high redox potential dye s like reactive black 5 (RB5) and also can oxidize low redox potential compounds such as phenolic monomers , simple amines, Mn²⁺ etc [118]. Compared to LiP and MnP the oxidization capacity of VP is higher for phenolic compounds, this ability might be due to its relatively more accessible distal main solvent channel allowing a third lower redox potential substrate oxidation site as in CiP. A research study conducted by Parez-Boada et al. reported that the spectral changes occurring during the oxidation of phenolic compounds by VP shows that VP in its resting state has a higher absorbance at 407 nm. Similarly during charge transfer, the transient states such as Compound I and II have an absorbance at 505 nm and 637 nm respectively [54, 79]. Two major enzymes MnP and VP are known for their ability to oxidize Mn²⁺ to Mn³⁺, the Mn²⁺ oxidation site of P. eryngii VP is similar to that of P. chrysosporium MnP . In VP, the Mn²⁺ binding site is formed by the side chains of Glu36, Glu40 and Asp175 located in front of the internal propionate of heme. Carboxylate groups of the amino acids and heme propionate are responsible for Mn²⁺ binding and for succeeding electron transfer to the activated heme of VP compounds I and II. Studies of the VP crystal structure showed a variable orientation of the Glu36, and Glu40 sidechains by interaction with Asp175 [118]. The position of these amino acids in recombinant VP shows an open gate conformation before exposure to Mn²⁺, thus enabling the oxidation of the Mn²⁺. At the same time native P. eryngii VP shows that the two glutamate side chains are pointed towards the Mn²⁺ corresponding to a closed gate conformation. In this conformation the carboxylate groups of Glu36, Glu40, Asp 175 and the propionate heme groups are at a distance from Mn²⁺. VP also oxidizes high redox potential substrates similar to LiP (a classic ligninolytic enzyme ) through the LRET pathway. This pathway occurs in several redox proteins like cytochrome-c-peroxidase , which oxidizes cytochrome-c on its surface by transferring electrons to tryptophan residues [54, 55, 118]. The LRET pathway was known earlier for its involvement in lignin degradation by different ligninolytic enzymes, thus overcoming steric hindrance which prevent the direct interaction of the heme group and the lignin polymer. Structural studies of VP show that three possible LRET pathways are involved during the oxidation of aromatic substrates by VP [118]. Oxidation of aromatic substrates starts at Trp 164 or His232 of VPL and at His82 or Trp170 of VPS1. VP can also efficiently oxidize low reduction potential compounds like ABTS , p-hydroquinone and 2, 6 dimethoxy phenol . Enzyme kinetics studies have showed that VP has two independent oxidation sites characterized by high and low specificities. Site directed mutagenesis of VP Trp164 performed by Ruiz-Duenas et al. showed that in Trp164 mutants the high specificity active site was removed while the low specificity site remained intact [55, 118]. Studies have confirmed a similar effect of a W164S mutation on VP oxidation of phenols . Based on these studies we conclude that the catalytic features of VP are due to its hybrid molecular architecture which includes different oxidation sites for Mn2+, high redox potential substrates (aromatic compounds) and low redox potential substrates (phenols and dye s) [55, 118].

5 Cellobiose Dehydrogenase

Cellobiose dehydrogenase is an extracellular enzyme involved in carbohydrate metabolism that was shown to be involved in lignin degradation [119, 120]. It was first isolated from an imperfect form of P. chrysosporium (Sporotrichum pulverulentum) [121]. Cellobiose dehydrogenase is a flavocytochrome enzyme which can oxidize various carbohydrates such as cellobiose (major product of cellulose degradation ) and mannobiose (product of mannose degradation) [58]. Several fungi were reported to produce cellobiose dehydrogenase, mostly white rot fungi such as P. chrysosporium (Sporotrichum pulverulentum), Trametes versicolor, Pycnoporus cinnabarinus, Polyporus dichrous, Merulius tremellosus, Phlebia radiata, Pleurotus ostreatus and Fomes annosus. Coniophora puteana ( brown rot fungi ) soft rot fungi , such as Sporitrichum thermophile (Myceliophthore thermophile), Schizophyllum commune, Humicola insolens, Sclerotium Rolfsii, Chaetomium cellulolyticum, imperfect soft rot fungi such as Monilla sitophila, Agaricus bisporus (Mushroom, Stachybotrys (Mold), Cladodporium(Mold) [121]. CDH degrades cellobiose and mannobiose to lactones by removing two electrons, which can be further transported to electron acceptors such as quinones, phenoxyradicals and dioxygen [58]. In CDH two prosthetic groups, FAD and heme, makes the enzyme suitable for the reduction of one electron acceptors such as radicals and metal ions . CDH has a high specificity for amorphous cellulose and less towards microcrystalline cellulose a unique property among non-hydrolytic enzymes [58]. CDH can produce hydroxyl radicals by reducing Fe³⁺ to Fe²⁺ and O₂ to H₂O₂, These reactive species depolymerize cellulose, xylan and to some extent lignin polymers [58].

Structure

A crystallographic study of cellobiose dehydrogenase was performed by Hallberg et al. (2002) [21]. Cellobiose dehydrogenase is a 90 kDa protein consisting of 752 amino acid residues , with a MW of 80 kDa and 10 kDa due to mannose glycosylation [121]. CDH is a monomeric enzyme with a flavin domain containing FAD (60 kDa ) and a heme domain (30 kDa ) containing a cytochrome b type heme. These two domains are connected by a fifteen amino acid residue linker. The FAD-binding subdomain consists of 205 amino acids , in an α/β type fold containing a six stranded parallel β-pleated sheet which is between three anti parallel β-sheets (β meander) and three α-helices [21]. Its sequence and structural predictions suggests that the ADP binding moiety of the cofactor consists of a βαβ motif, which is usually seen in NAD or FAD dependent enzymes [21]. The F-subdomain also consists of three α-helices which together form the substructure on one side of the F-subdomain [21]. The substrate binding domain (S-subdomain) consists of 335 amino acid residues forming a central twisted seven stranded β-sheet (Sheet B), also containing three α-helices on one side of the sheet with the active site on the other side. The S-subdomain consists of a relatively long 3₁₀ helix and two α-helical regions, finally sheet B forms the bottom of the active site. The beginning of the S-subdomain forms a loop and lid structure that outlines the entrance of the active site and the outer wall of the FAD binding pocket [21]. The heme unit of CDH is highly glycosylated relative to the flavin domain. The isoelectric point of the flavin domain is 5.45, that of the heme domain is 3.42 and that of CDH is 4.2 [21, 121]. CDH is thermostable and pH stable and it has maximum activity at room temperature for 24 h in a pH range of 3–10. CDH is highly stable between pH 3–5 and its stability decreases with an increase or decrease from this range. CDH has highest specificity for sugars mainly disaccharides and oligosaccharides and the k_cat (enzyme turnover number) for cellobiose reduction by CDH is ten times higher than glucose [21, 121]. Cellobiose is the most effective electron donor for CDH but many other reducing sugars can be used for electron transfer. CDH can reduce a wide range of electron acceptors, which include oxidative radicals, transition metals and two electron acceptors (quinones). Studies have reported that one electron acceptors are reduced preferentially at the heme domain [121]. CDH has a strong binding specificity for microcrystalline cellulose, once bound it cannot be easily eluted even at high pH, high salt concentrations, detergents or by cellobiose. It was reported that CDH inhibits LiP (lignin peroxidases ) by reducing the veratryl alcohol and also can reduce compound II (intermediate formed during the catalytic cycle of LiP). CDH also reduces Mn³⁺ (redox mediator used by MnP ) and also compound II of MnP similar to LiP [21, 121].

Mechanism

CDH has the properties of a typical dehydrogenase with both oxidative and reductive reactions. CDH oxidizes the C1 position of a saccharide to a lactone which is spontaneously hydrolyzed to a carboxylic acid . The electrons taken up by the enzyme are later transferred to one or two electron acceptors [122, 123]. Substrate specificity of CDH is higher for cellobiose, cellodextrins, lactose, mannobiose and galactosylmannose. However, the later substrates have higher Km-values, the true substrates for CDH are di or oligosaccharides with reducing ends containing glucose or mannose residues . Monosaccharides such as glucose, mannose and maltose have very high Km values suggesting that there is binding of two glucose residues to the active site in separate subsites, at the same time monosaccharides have lower Kcat values then the di or oligo saccharides which suggests that binding of the β-dihexosides to the active site stimulates the catalysis creating an induced fit [124]. CDH also generates highly reactive hydroxyl radicals by a Fenton type reaction in the presence of an electron donor. Several studies were conducted to study the individual roles of the two prosthetic groups (flavin and heme domains) in the oxidation of compounds, which showed that oxidation of cellobiose (electron donor) is carried out by the FAD group which is further converted to FADH₂ and later transfers the electrons to the heme group (Fig. 4.7) [122, 123, 127]. Several groups have proposed a role for CDH in lignin depolymerization by reducing phenoxy radicals thus preventing repolymerization of the radicals. Studies conducted by Henriksson et al. (1995) have showed that the CDH could stop the repolymerization of lignin model compounds , this was showed by incubating CDH with cellobiose, ferric ions, hydrogen peroxide and lignin model compounds . CDH generates highly reactive hydroxyl molecules that depolymerized the polymers showing that CDH does not depolymerize lignin or its subunits directly but hydroxyl radical groups are involved in the degradation of lignin related compounds [119, 128–130].

Benzoquinone or quinone reductases are significant enzymes secreted by several fungi especially P. chrysosporium . 1,4-Benzoquinone reductase is a NADPH dependent intracellular enzyme , it contains flavin mononucleotide (FMN). 1,4-Benzoquinone reductase was active during both primary and secondary metabolism but the enzyme inducers are stronger during the primary metabolic processes [188–190]. Studies showed that when vanillate or methoxy-p-quinone are added to cells, carrying out primary metabolism, enzyme expression was increased. However, the effect was small when the same compounds were added to secondary metabolic cells, which suggests that quinone reductase is regulated independently of lignin and manganese peroxidase [188]. LiP and MnP are expressed only during the secondary metabolic stage of the growth and the there expression is not induced by aromatic substrates. The regulation of quinone reductases is similar to that of vanillate hydroxylase, which suggests its involvement in vanillate metabolism [188–190]. Quinone reductase was expressed during the lignolytic phase of P. chrysosporium , suggesting a role in the reduction of quinones generated during lignin degradation . It was reported that quinone reductases are induced upon quinone addition, suggesting the involvement of quinone reductase in lignin and quinone degradation [188–190]. Besides degrading of quinone and lignin derived compounds, it is also reported that quinone reductases protects P. chrysosporium from oxidative stress by acting as redox active toxins. Quinones obtained by metabolic conversion are reduced by one electron generating semiquinone radicals, which are then oxidized by oxygen generated super oxide anion, this superoxide anion is further converted to H₂O₂ through superoxide dismutase and later in the presence of suitable electron donors it results in production of highly reactive hydroxyl radicals from H₂O₂ [188–190]. The detailed mechanisms of the regulation and catalytic mechanism of quinone reductase need to be explored (Table 4.3).

Table 4.3 Catalytic mechanisms and structural studies of different lignin degrading auxiliary enzymes (LDA )

Full size table

7 A Short Note on Genome Sequencing Studies of Lignin Degrading Fungi

In 2004, Martinez et al., had sequenced the whole genome sequence of Phanerochaete chrysosporium [206]. As it was the first wood decaying fungus to be sequenced from the phylum Basidiomycota there were few difficulties in annotating the whole genome. In the year 2006, Vanden Wymelenberg et al. have improved the genome assembly and reannotated the P. chrysosporium genome [207]. P. chrysosporium genome has revealed a wide array of hydrolytic and oxidative enzymes mainly class-II peroxidases apparently supporting its lignocellulose degrading ability. Later in the year 2009, complete genome sequencing of Postia placenta was performed by Martinez et al, genome of P. placenta has revealed several interesting facts about its wood decaying patterns. P. placenta genome lacks genes coding for peroxidases , few genes coding for enzymes involved in degradation of crystalline cellulose (GH6, GH7 and GH61) [208]. However, these studies have reported that P. placenta genome codes for enzymes involved in generating Fenton reagents (reactive oxygen species). Whole genome sequences of prominent brown rot fungus such as Serpula lacrymans [209], Fibroporia radiculosa [210] have shown similar results such as lacking the genes coding for peroxidases , reduced expression of genes coding for CAZymes and possessing oxidoreductases involved in Fenton reactions. The genome sequence of the Schizophyllum commune was first wood decaying fungus to be sequenced from order Agaricales [211]. Though it was reported to degrade lignin, S. commune lacked the genes coding for peroxidases . While whole genome sequencing of several white rot fungus such as Ceriporiopsis subvermispora [212], Phanerochaete carnosa [213], Heterobasidion irregulare [214], Laccaria bicolor [215], Volvariella volvacea [216], Agaricus bisporus [217], Armillaria mellea [218] etc., have revealed several significant facts about lignocellulose degrading enzymes and their corresponding mechanisms [219]. Large scale genome comparison studies showed that whiterot fungi has wide range of genes coding for lignocellulolytic enzymes compared to brown rot fungi [220]. Especially enzymes involved in breakdown of crystalline cellulose such as GH6, GH7, GH61 and CBM1 are predominant in white rot fungi . Similarly whiterot fungi possess several genes coding for peroxidases , whereas in most of the brown rot fungi genes coding for peroxidases are mostly absent or reduced [220, 221].

These sequencing studies has dramatically raised the interest in sequencing projects of wood decaying fungi . The large scale sequencing projects like 1000 Fungal Genome Project, Joint Genome Institute (JGI) are playing a critical role in revealing and understanding the whole genome of sequences of several fungi (Figs. 4.12 and 4.13). The JGI have also developed a strong web based interactive and integrated genome analysis portal called MycoCosm [222]. As of 2016 JGI MycoCosm offers over 563 sequenced and annotated fungal whole genome sequences . As most of the wood decaying fungi were reported from the phylum Basidiomycota out of which 202 fungal genome sequences belongs to the phylum Basidiomycota. Among Basidiomycete s fungi 159 fungal genome sequences reported belongs to subphylum Agaricomycotina, 16 and 27 fungal genome sequences belong to Ustilaginomycotina and Pucciniomycotina respectively. Thus, the recent advancements in genomics field in the past decade has prominently improved our understanding the genes involved in coding for the lignocellulose degrading enzymes and their pathways [219].

8 Conclusion and Future Outlook

Depletion of fossil fuels in the world is one of the major reasons for increased biofuel research. Lignocellulose is one of the major alternative resource fulfill the current requirement for fuels and chemicals. The biofuel and paper pulp industries efficiently use cellulose in the production of paper and cellulosic ethanol , however lignin is left as a major industrial byproduct. Fungi are the most efficient lignin degrading microorganisms , producing several extra and intra cellular enzymes for breakdown of lignin. Understanding the lignin degrading mechanisms employed by white rot fungi is very important. Recent advances in the field of genomics and proteomics have revealed significant information about genes related to lignocellulose degradation and detoxification mechanisms . Whole genome sequences of Phanerochaete chrysosporium , Postia placenta, Gloeophyllum trabeum and Dichomitus squalens prove the occurrence of lignocellulolytic enzymes. Integrated computational analyses of proteome and secretome show that fungi undergo severe oxidative stress during the process of wood degradation , for which it employs various antioxidative and detoxification mechanisms . Application of advanced sequencing methodologies integrated with HPLC, GC and mass spectrometry techniques can help to elucidate the pathways involved in wood degradation.

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Department of Biology, Lakehead University, 955 Oliver Road, P7B 5E1, Thunder Bay, ON, Canada
Ayyappa Kumar Sista Kameshwar & Wensheng Qin

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Biomass Group, College of Engineering, Nanjing Agricultural University, Nanjing, Jiangsu, China
Zhen Fang
Research Center of Supercritical Fluid Technology, Graduate School of Environmental Studies, Tohoku University, Sendai, Japan
Richard L. Smith, Jr.

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Kameshwar, A.K.S., Qin, W. (2016). Lignin Degrading Fungal Enzymes. In: Fang, Z., Smith, Jr., R. (eds) Production of Biofuels and Chemicals from Lignin. Biofuels and Biorefineries. Springer, Singapore. https://doi.org/10.1007/978-981-10-1965-4_4

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DOI: https://doi.org/10.1007/978-981-10-1965-4_4
Published: 29 September 2016
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Print ISBN: 978-981-10-1964-7
Online ISBN: 978-981-10-1965-4
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Lignin Degrading Fungal Enzymes

Abstract

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Ligninolytic Fungi from the Indian Subcontinent and Their Contribution to Enzyme Biotechnology

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Keywords

1 Introduction

2 Carbohydrate Active Enzyme Database (CAZy)

3 Fungal Oxidative Lignin Enzymes (FOLy)

4 Lignin Oxidizing Enzymes (LO)

4.1 Laccases (EC 1.10.3.2, Benzenediol: Oxygen Oxidoreductase)

Structure

Mechanism of Action

4.2 Peroxidases (EC:1.11.1.x)

4.3 Lignin Peroxidases (E.C. 1.11.1.14)

Structure

Mechanism

4.4 Manganese Peroxidase s (EC 1.11.1.13)

Structure

Mechanism

4.5 Versatile Peroxidases

Structure

Mechanism

5 Cellobiose Dehydrogenase

Structure

Mechanism

6 Lignin Degrading Auxiliary Enzymes (LDA )

6.1 Aryl Alcohol Oxidase

Structure

Mechanism

6.2 Vanillyl Alcohol Oxidase

Structure

Mechanism

6.3 Glyoxal Oxidase

Structure

Mechanism

6.4 Pyranose Oxidase

Structure

Mechanism

6.5 Galactose Oxidase

Structure

Mechanism

6.6 Glucose Oxidase

Structure

Mechanism

6.7 Benzoquinone Reductase

Structure

Mechanism

7 A Short Note on Genome Sequencing Studies of Lignin Degrading Fungi

8 Conclusion and Future Outlook

References

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