ABSTRACT
This study was carried out to screen, partially purify and characterize Manganese peroxidase from Rigidoporuslignosus. This study started with the optimization of enzymes production in the laboratory scale of submerged fermentation system.A pilot study was carried out for eight days to determine the day of highest Manganese peroxidase activity of which Day 7 wasthe highest. The optimal yield of Manganese peroxidase (0.888 U/ml) was found to be produced under the conditions of 20 mL of synthetic medium (containing (g/L): glucose, 10.0; NH₄NO₃, 2.0; KH₂PO₄, 0.8; Na₂HPO₄, 0.4; MgSO₄ · 7H₂O, 0.5; yeast extract, 2.0. pH 6.1) with 5% of glucose as the carbon sources, and with microelements (ZnSO₄ · 7H₂O, 0.001 g/L; FeSO₄ · 7H₂O, 0.005 g/L; CaCl₂ · 2H₂O, 0.06 g/L; CuSO₄ · 7H₂O, 0.05 g/L; MnSO₄ · H₂O, 0.05 g/L) with an initial pH of 4.5, and 4 cork borer of the pure culture ofRigidiporuslignosus, The specific activities for the crude enzyme was found to be 0.399 U/mg. Ammonium sulphate (80%) saturation was found suitable to precipitate protein with highest MnP activity. After ammonium sulphate precipitation and gel filtration, the specific activity was found to increase from 3.178U/mg protein to 1.707U/mg protein for fraction A with the purification of 4.28, while that for fraction B increased from 3.178 to 4.04U/mg protein with purification fold of 10.14. The optimum pH and temperature were found to be 5.0 and 50°C respectively. The Michealis-Menten constant, Kmand maximum velocity, Vmax obtained from Line-Weaver- Burk plot of initial velocity data at different substrate concentrations were found to be 1.102mg/ml and 11.561 U/ml using H2O2, 0.76mg/ml and 19.65U/ml using phenol red as substrate.Kinetics of MnP inactivation was studied over temperature range of 30- 70°C. The inactivation kinetics followed a biphasic pseudo first-order model with k values between 4.2×10-3 – 1.79×10-2 min-1. The decreasing trend of k values with increasing temperature indicates a faster inactivation of manganese peroxidase from Rigidiporuslignosusat higher temperatures. The activation energy (Ea) of 28.43kJ/mol was calculated from the slope of Arrhenius plot. Thermodynamic parameters (∆H, ∆G, ∆S) for inactivation of manganese peroxidase at different temperatures (30-70°C) were studied in detail.In conclusion, the results of this present study indicates that manganese peroxidase will be a good enzyme for delignification with a high capacity to remove xenobiotic substances and produce polymeric products which are useful in bioremediation.
CHAPTER ONE
INTRODUCTION
The rubber tree (Hevea brasiliensis Muell-Arg.) belongs to the Family Euphorbiaceaeof laticiferous plants. The tree growth can reach a height of over 20 m.Rubber tree is believed to be a native of the tropical forests of South America, the introduction of the rubber tree to Malaysia was made in 1876 via the Wickham collection from the Amazon valley. What was once the object of idle curiosity to the eccentric botanist suddenly became a main source of industrial material of immense value. Rubber thus began to be planted on large scale and the record of its expansion in Malaysia was phenomenal, running a close parallel to the development of the automobile industry of the world (Lieberei, 2007).
Natural rubber while promoting export earnings and livelihood of people supplement thousands of hectares to the forest cover. Over the past decades, the rubber yield has significantly increased due to the cultivation of high yielding clones. Monoclonal Hevea brasiliensis is principally valued for its latex content, the latex or Natural Rubber (NR) is very significant in world’s industrialization. This importance has been expressly emphasized in the production of elastomers, the use of which is indispensable in space, water, and ship technologies (Jacob, 2006). The dependence of world industrialization on NR production is further underscored especially now considering the diminishing reserves of petroleum with increasing environmental hazards.However, latex production still faces more serious economic losses due to many biotic constraints which include significant losses caused by pathogenic fungi. Among them, White root disease (WRD) is very destructive in rubber plantations of Nigeria and in many other rubber growing countries. This disease has been identified as one of the major causes for the loss of plants during the first five years after planting resulting in low productivity levels. Inspite of the fact that disease management strategies have been clearly outlined by the Rubber Research Institute, the disease incidence is showing an increasing trend. One of the main reasons for this has been identified as the increment of the host range (Jayasuriya, 2004).
The rubber tree is subject to a plethora of economically important pathological problems, mainly of fungal origin (the basidiomycetes) (Igeleke, 1998). In Nigeria, the most serious diseases of rubber seedlings and budded plants in the nursery are leaf diseases (Begho, 1990), while in mature plantation, the most devastating leaf disease is the South American Leaf Blight (SALB),
and Corynespora Leaf Fall Disease (CLFD) appears to be next to SALB. In field plantations, root diseases pose a serious problem especially in the first few years after planting. In Nigeria, the white root rot disease of rubber is the most serious. It accounts for about 94% of incidences of all root diseases and kills up to five Hevea trees/ha (Otoide, 1978).
Over a period of time, half of the rubber trees in a plantation are lost to the disease. The infective fungal organism of the white root rot disease is Rigidoporus lignosus (klotzsch) Imazeki. The brown root rot disease (Phellinus noxius) Corner-Cunn., unlike the white root rot, is the most serious root disease in Hevea plantations in Liberia while R. lignosus and Armillaria root rots occur to a lesser extent (Nandris et al., 1987). Similarly, in Cote d´lvoire, R. lignosus is the main cause of Hevea tree losses with 40-60% of the trees destroyed over a period of up to 21 years (Nandris et al., 1987). The white root rot incidence is absent in India, however, it is serious in Malaysia, Sri Lanka and Congo, aside its severe occurrences in Nigeria and Cote d´lvoire.
Rubber tree exhibits natural resistance to invading root pathogens. Resistance often breaks down due to effect of pathogens that colonize living tissues of the tree to obtain nutrients as a result of the damaging and weakening of the plant with toxins or by preventing the plants defense mechanism (Jayasuriya, 2004). A number of certain defense mechanisms in Hevea against R. lignosus and P. noxius have been identified. These include cellular hypertrophy and hyperplasia, cambium activity stimulation, lignifications and suberification of certain cell walls (Jayasuriya,
2004.; Nicole et al., 1985).
The process of pre- infection involves pathogen breaking down the host cuticle and cell wall. Plants respond to infection process by producing anti-microbial compounds of low molecular weight (phytoalexins). Hevea plants produce anti-microbial phenolic compounds such as coumarins, flavonoids, triterpenes among others (Jayasuriya, 2004) that can partially or completely inhibit microbial infection.
The growth and spread of infective fungal pathogens from existing population have been on the increase with great virulency and inflicting damages even to resistant genotypes. The impact of fungal pathogens results in crop losses. The production of phenylalanine ammonia-lyase (PAL) is implicated as key enzyme in the plant phenyl propanoid pathway to catalyse the synthesis of phenyl lignin and phytoalexin from L-phenylalanine. The synthesis of these anti-microbial compounds and the subsequent increase in PAL concentration are often useful resistance indicator in the host (Nicholson and Hammerschmidt, 1992). Also, oxidases and peroxidases are
known to be actively involved in polymerization of phenolic compounds in the lignin formation. In resistance mechanism action of peroxidase is related to initiation of hypersensitive cell collapse.
Pathogenesis-related proteins(PR) is yet another defense response of rubber against pathogen infection. The PR protein induced by polyacrylic, acetylsalicylic (aspirin) and salicylic acids are known to increase resistance to pathogens. There is variation among high yielding genotypes in disease tolerance level. In this regard, some clones are resistant to virtually most of the diseases but are susceptible to few diseases (Ebrahimet al., 2011).
However, certain clones exhibit tolerance to few diseases but some others are susceptible to many diseases. There are few clones which tolerate characteristics of pathogens, and as a result of abiotic factors, produce new strains, and these strains can be more aggressive against rubber clones. Mutation is seen to be responsible for variability in pathogens, which involves changing sequence bases in the nuclear DNA, either by way of substitution or addition or deletion of one or many base pairs (Omorusi, 2012).
The Hevea root fungal parasites- R. lignosus, and P. noxius are polyporaceae major causes of rubber tree losses in plantation causing the decay of lignified root tissues (Geiger et al., 1986). R. lignosus is partially involved lignin consumption whereas P. noxius degrade the polysaccharide fraction of but not lignin (Cowling, 1961; Kirk, 1971), however, findings by Geiger et al. (1986) showed that R. lignosus and P. noxius degrade both the lignin and polysaccharide fractions of the wood. Although P. noxius exhibits preferential degradation of polysaccharides, whereas, R. lignosus degrades both the lignin and polysaccharides in a relatively balanced manner but with slight preference for lignin (Omorusi, 2012).
Rigidoporus lignosisforms many white, somewhat flattened mycelia strands 1–2 mm thick that grow on and adhere strongly to the surface of the root bark. These rhizomorphs grow rapidly and may extend several meters through the soil in the absence of any woody substrate. Thus, healthy rubber trees can be infected by free rhizomorphs growing from stumps or infected woody debris buried in the ground as well as by roots contacting those of a diseased neighboring tree.
The pathogen produces two ligninolytic enzymes namely Laccase (EC 1.10.3.2) and Manganese- dependent peroxidase(MnP, EC 1.11.1.13). These two enzymes act in a complementary way during lignin degradation (Galliano et al., 2006) where MnP is thought to play the most crucial role in lignin degradation (Wong, 2008). Two ecophysiological groups of basidiomycetes, wood-
decaying fungi causing white-rot as well as certain soil litter decomposing fungi, secrete MnP mostly in multiple forms into their microenvironments. Potential applications for MnP include biomechanical pulping, pulp bleaching, dye decolorization, bioremediation and production of high-value chemicals from residual lignin from biorefineries and pulp and paper side-streams (Järvinenet al., 2012).
1.1 Wood composition
Wood is a porous plant material made up of various types of xylem cells. Softwoods (in gymnosperm trees) consist mainly of long tracheids and smaller ray parenchyma cells. Water transport and stem strength are mainly sustained by the dead tracheid cells. In addition, longitudinal resin ducts exist (Kuhad et al., 1997). Hardwoods (in angiosperm plants and deciduous trees) have more diverse types of xylem cells including fibers, vessels, and ray parenchyma cells. These cells are responsible for support and nutrient storage as well as water and nutrient transport between plant roots and the photosynthesizing leaves or needles. Wood cell walls, in particular the long tracheids and fibers, consist of several layers which differ in their structure and chemical composition (Fig. 1a). The major components of the wood cell walls are three biopolymers, namely cellulose, hemicellulose, and lignin (Harris and Stone, 2008).
Lignin is an aromatic and amorphous polymer present in all layers of woody cell walls. In fibers and tracheids, the thin middle lamella has the highest lignin content, whilst most of the lignin exists in the thick secondary wall layers embedded with cellulose microfibrils and hemicellulose. Lignin mechanically strengthens vascular plants and aids in water transportation since it physically attaches the xylem cells together. At the same time, lignin protects the more easily degradable cellulose and hemicellulose polymers from microbial attack.The heterogenic lignin polymer is synthesized in the plant xylem cells from phenylpropanoid precursors i.e. p-coumaryl, coniferyl, and sinapyl alcohol. During lignin biosynthesis these monolignols are polymerized to p-hydroxyphenyl, guaiacyl, and syringyl type of lignin subunits by the action of laccases and peroxidases (Higuchi, 2006). Lignin subunits are joined together by diverse carbon-carbon and ether bonds of which the β-aryl-ether (β-O-4) bond is the most common. The composition and amount of lignin varies between softwood and hardwood, and between plant species. The lignin content of softwoods (25-33% of xylem dry weight) consists mainly of guaiacyl subunits while hardwood lignin (20-25% of xylem dry weight) contain both guaiacyl and syringyl subunits
(Higuchi, 2006). In grass plants, xylem cell wall lignin also contains p-hydroxyphenyl subunits and e.g. esterified ferulic acid. Knowledge of the chemical structure of diverse plant lignin is still incomplete, although several lignin models have been presented (Fig. 1b).
Most wood species contain approximately 40-45% (as dry weight) cellulose which is the main component of wood. In the linear cellulose polymer, repeating glucose units are linked together by β-1,4-glucosidic bonds and the degree of polymerization is up to about 15000 glucose units within one polymeric chain (Kuhad et al., 1997). In wood cell wall the long, contiguous cellulose chains are stabilized by hydrogen bonds into microfibrils and further into cellulose fibers. The majority of cellulose is situated in the thick S layer of xylem secondary wall (Fig. 1a) where its fibrillous structure gives mechanical strength to wood (Argyropoulos and Menachem, 1997). Usually, the highly organized crystalline cellulose dominates whereas only a small portion exists as amorphous non-organized cellulose which is more susceptible to enzymatic degradation (Kuhad et al., 1997).
Hemicelluloses are a diverse group of branched heteropolysaccharides consisting of different hexose, pentose, and sugar acid units. Most of the hemicelluloses act as a supporting material and usually comprise 20-30% of wood dry weight. Hemicelluloses are amorphous and have a moderate degree of polymerization (100-200 units) and thus are more easily biodegradable than cellulose. The composition and structure of hemicelluloses differ in softwood and hardwood: galactoglucomannans are the main hemicelluloses in softwood while glucuronoxylan dominates in hardwood. In the woody cell walls, hemicelluloses are bound to cellulose microfibrils via hydrogen bonds. Hemicelluloses and lignin are covalently linked forming the so called lignin- hemicellulose matrix (Fig. 1a, Kuhad et al., 1997; Harris and Stone, 2008). Depending on the wood species, 2-5% of the wood dry weight is made up of extractives. Extractives are non- structural constituents of wood. They may be broadly divided into terpenes, resins, and phenols (Kuhad et al., 1997). These various organic compounds have several roles, acting as a nutrition reserve for the living wood cells as well as giving protection against microbial degradation. In addition, low amounts of proteins and inorganic compounds are present in the wood.
Figure 1. A) Composition of wood showing 1) tracheids, 2) wood cell wall layers, 3) distribution of lignin-hemicellulose matrix (black), hemicellulose (white) and cellulose (grey) in the secondary cell wall. ML: middle lamella, P: primary wall, S1-S3: secondary cell wall layers. B) Structural model of lignin by Brunow et al.(2001).
1.2 Wood degradation by fungi
Wood, being poor in nutrients other than organic carbon, is a demanding growth environment for microorganisms. In addition, the lignocellulose complex efficiently hinders the access of microbes and their enzymes into wood cell walls. Other wood components, such as extractives, also restrict the growth of many microbes. Of all organisms, fungi are the most powerful degraders of the wood polymers (Carlile et al., 2001). As vascular plants form the vast reservoir of photosynthetically fixed carbon on earth, wood-decaying fungi have enormous ecological impact on the global carbon cycling. Three different types of wood decay caused by fungi can be distinguished: white- rot, brown-rot, and soft-rot. Basidiomycetous white-rot fungi are saprotrophs mostly living on dead wood. White-rot fungi have a unique ability to efficiently mineralize lignin to CO2 with their oxidative lignin-modifying enzymes (LMEs) (Kirk and Farrell, 1987; Hatakka, 2001; Hammel and Cullen, 2008). Also wood-colonizing ascomycetous fungi are capable of mineralizing lignin to some extent (Liers et al., 2006). Basidiomycetous litter-decomposing fungi have been reported to mineralize lignin as well, but their growth and degradation capacity is usually restricted to the soil environment (Steffen, 2003).
Following the action of white-rot fungi, the decayed wood is characteristically white and fibre- like. Most white-rot fungi are able to degrade all the wood polymers. However, there are so called selective white-rot fungi which preferentially degrade lignin and hemicellulose leaving cellulose polymer almost intact (Kuhad et al., 1997). These species include e.g. Dichomitus squalens, Physisporinus rivulosus, Ceriporiopsissubvermispora (Hakala et al., 2004; Fackler et al., 2006) and Rigidoporus lignosus . The model white-rot fungus, Phanerochaete chrysosporium, is efficient in wood and lignocellulose decay but less selective for depolymerization of lignin over cellulose utilization (Hatakka, 2001; Hakala et al., 2004).
Another group of wood-decaying basidiomycetes is the brown-rot fungi which rapidly depolymerize cellulose in the early stage of wood decay. As wood polysaccharides are degraded and some modification of lignin occurs, mostly by demethoxylation, the decayed wood remains brown and has lost its strength. The mechanisms which brown-rot fungi use in cellulose degradation are still poorly understood, and the first brown-rot fungal genome sequenced (Postia placenta) (Martinez et al., 2009) has offered new insights into future studies on the decomposition of wood polysaccharides.
Some ascomycetes (e.g. Trichoderma and Xylaria species) cause a third type of wood-rot known as soft-rot. These fungi typically attack wood in wet environments. As a result of soft-rot, cavities or complete erosion of tracheid secondary cell walls is detected, and the decayed wood loses its mechanical strength due to cellulose breakdown (Carlile et al., 2001). Some ascomycetes, so called sap-staining fungi, degrade mainly wood extractives. These species are primary wood colonizers that characteristically discolour the sapwood with their dark pigmented hyphae, which leads to mainly cosmetic rather than structural damage (Breuil et al., 1998).
Thefungievaluatedfortheproductionofextracellular peroxidaseandlaccase arelistedinTable1.
Table 1: Fungi that produce peroxidases and laccase
| Microorganism | Type | Enzyme |
| Bjerkanderasp. | Basidiomycete | LiP,MnP |
| Ceriporiopsis subvermispora Chaetomium thermophilium | Basidiomycete Ascomycete | LiP Lac |
| Chrysosporium lignorum Coriolushirsutus | Basidiomycete Basidiomycete | LiP,MnP MnP |
| Dichomitussqualens | Basidiomycete | MnP,Lac |
| Elfvingiaapplanata Flavodonflavus | Basidiomycete Marine basidiomycete | MnP,Lac MnP,Lac |
| Halosarpheia ratnagiriensis | Marine ascomycete | Lac |
| Halosarpheia ratnagiriensis Irpexflavus Irpexlacteus | Marine ascomycete Basidiomycete Basidiomycete | Lac MnP MnP,Lac |
| Lentinusedodes | Basidiomycete | LiP,MnP, Lac |
| Lentinussquarrosulus Nematolomafrowardii | Basidiomycete Basidiomycete | LiP LiP,MnP |
| Oxyporus latemarginatus Phanerochaete chrysosporium | Basidiomycete Basidiomycete | LiP,MnP LiP,MnP, Lac |
| Phanerochaetecrassa | Basidiomycete | MnP |
| Phanerochaeteflavido alba Phanerochaete magnolia | Basidiomycete Basidiomycete | LiP,MnP LiP,MnP |
| Phanerochaete sordida | Basidiomycete | LiP,MnP |
| Phellinuspini Phlebiaradiate | Basidiomycete Basidiomycete | LiP,MnP LiP,MnP, Lac |
| Phlebiasp. | Basidiomycete | MnP,Lac |
| Phlebiasubserialis Phlebiatremellosa | Basidiomycete Basidiomycete | MnP MnP |
| Pleurotuseryngii Pleurotusostreatus | Basidiomycete Basidiomycete | MnP,Lac MnP,Lac |
| Pleurotussajor-caju | Basidiomycete | MnP,Lac |
| Poluporussp. | Basidiomycete | LiP,MnP, Lac |
| Polyporussanguineus Psathyrella atroumbonata | Basidiomycete Basidiomycete | MnP LiP |
| Pycnoporus cinnabarinus Rigidosporuslignosus | Basidiomycete Basidiomycete | MnP,Lac MnP,Lac |
| Schizophyllum commune | Basidiomycete | Lac |
| Sordariafimicola Stereumannosum | Ascomycete Basidiomycete | Lac Lac |
| Stereumhirsutum | Basidiomycete | LiP,MnP, Lac |
| Trametes(CoriolusorPolyporus)versicolor Trametestrogii | Basidiomycete Basidiomycete | LiP,MnP, Lac LiP,MnP, Lac |
| Trichodermaatroviride | Ascomycete | Lac |
LiP:ligninperoxidase;MnP:manganese peroxidase;Lac:laccase (Steffen, 2003).
1.3 Lignin-modifying enzymes
Lignin-modifying enzymes (LMEs) considered to be involved in lignin biodegradation include oxidative enzymes that catalyze unspecific reactions, i.e.Laccase (benzenediol:oxygen oxidoreductase, EC 1.10.3.2), lignin peroxidase (LiP, diarylpropane peroxidase, EC 1.11.1.14), manganese peroxidase (MnP, EC 1.11.1.13), and versatile peroxidase (VP, EC 1.11.1.16) (Hammel and Cullen, 2008). Also several H2O2 -generating enzymes such as aryl alcohol oxidase (AAO, EC 1.1.3.7), glyoxal oxidase (GLOX), and pyranose-2 oxidase (EC 1.1.3.10) are regarded as members of white-rot fungal lignin-degrading machinery (Kersten and Cullen, 2007). Recently, oxidases potentially involved in the degradation of lignin and related aromatic compounds have been classified into enzyme families according to their protein sequence and biochemical properties, and integrated into FOL (Fungal Oxidative Lignin enzymes) database (Levasseur et al., 2008).
LMEs generate by oxidative reactions highly reactive free radicals due to which degradation of lignin by white-rot fungi is known as “enzymatic combustion” (Kirk and Farrell, 1987). LMEs are expressed by white-rot and litter-decomposing fungi in different combinations and typically, several LME isozymes are encoded by multiple genes and their alleles within one fungal species (Kersten and Cullen, 2007; Pezzella et al., 2009). It has been considered that the multiple, structurally related LME-encoding genes and heterogeneity of their regulation can provide flexibility which white-rot fungi need for adaptation to for example, changing environmental conditions and during growth on different wood species (Conesa et al., 2002; Kersten and Cullen,
2007). On the other hand, this genetic diversity may barely represent functional redundancy
(Hammel and Cullen, 2008).
Current knowledge of lignin-degradation supports the view that lignin is depolymerized outside the fungal hyphae by the combined oxidative action of LMEs, oxygen radicals, and small metabolites after which at least some of the resulting fragments are mineralized intracellularly (Hatakka 2001; Hammel and Cullen, 2008). The importance of lignin- modifying peroxidases and H2O2 production in lignin breakdown has been highlighted in recent transcriptome and proteome studies of the white-rot model fungus Phanerochaete chrysosporium (Sato et al., 2009; Vanden Wymelenberg et al., 2009). During the growth of P. chrysosporium in nutrient limited liquid cultures that mimic lignin-degrading conditions the increased expression of LiPs, MnPs, and various extracellular oxidases was observed (Vanden Wymelenberg et al., 2009). On the
stationary wood cultures, P. chrysosporium genes encoding LiPs and alcohol oxidase were reported to be highly expressed (Sato et al., 2009).
In addition to this, the whole genome sequence of P. chrysosporium revealed a large number of putative genes encoding extracellular oxidative enzymes which can also be connected to lignocellulose degradation. The transcriptome and secretome analyses under lignin-degrading conditions have showed a set of expressed genes and secreted proteins of P. chrysosporium with unknown function (Sato et al., 2009; Vanden-Wymelenberg et al., 2009). These data suggest that the whole complexity of the white-rot fungal process of lignin degradation is yet to be unraveled.
1.3.1 Lignin-modifying Peroxidases
Lignin-modifying heme peroxidases (lignin peroxidase, LiP; manganese peroxidase, MnP; versatile peroxidase, VP) are extracellular glycoproteins that belong to the class II heme- containing, fungal secretory peroxidases within the plant peroxidase superfamily. The class II peroxidases are structurally related globular proteins predominantly consisting of 11-12 α-helixes divided to two domains. These peroxidases carry Fe-containing heme (protoporphyrin IX) as their prosthetic group coordinated by two highly conserved histidine residues (distal and proximal histidines) in a central cavity between the two domains.
Recently, the significance of lignin-modifying peroxidases in lignin degradation has been supported by comparison of the genomes of Phanerochaete chrysosporium, a model white-rot fungus for lignin degradation, and Postia placenta, which has become a model brown-rot fungus for wood polysaccharide degradation (Martinez et al., 2009). The haploid genome of P. chrysosporium strain RP78, derived from dicaryotic strain BKM-F-1767 (ATCC 24725), contains multiple lignin-modifying- peroxidase-encoding genes (10 LiPand 5 MnP genes). In contrast, the dicaryotic genome of P. placenta totally lacks lignin-modifying-peroxidase-encoding genes, and contains only one putative class II-fungal secretory peroxidase-encoding gene. This P. placenta peroxidase gene possibly encodes a low-redox potential type of peroxidase related to the CIP- enzyme of the basidiomycete Coprinopsis cinerea (Coprinus cinereus) not capable of lignin degradation (Martinez et al., 2009).
The first class II heme peroxidase gene of a brown-rot fungus was recently cloned from Antrodia cinnamomea and the corresponding enzyme was reported to decolorize and oxidize some phenolic dyes (Huang et al., 2009). Although A. cinnamomea peroxidase may represent a new
group of extracellular class II heme peroxidases from previously unstudied brown-rot basidiomycetes (Huang et al. 2009), its role in lignin modification is still unclear. Preliminary genomic PCR data indicate that also some species of the ectomycorrhizal basidiomycetes may possess genes coding for class II heme peroxidases (Bödeker et al., 2009). However, so far no lignin-modifying-peroxidase-encoding genes have been annotated in the whole genome sequence of the ectomycorrhizal model fungus L. bicolor.
1.3.1.1 Occurrence and Properties of Lignin-modifying Peroxidases
Active LiP isozymes, first found in the cultures of P. chrysosporium have been described from only a few genera of white-rot fungi including Phlebia (e.g.P. radiata and Phlebia (Merulius) tremellosa, Trametes (e.g. T. versicolor and T. trogii), and Bjerkandera (e.g. B. adusta and Bjerkandera sp.). In contrast, MnPs are widespread among lignin-degrading fungi including both white-rot and litter-decomposing basidiomycetous species (Hatakka, 2001; Hofrichter, 2002; Steffen et al., 2002; Lankinen et al., 2005). The lignin-modifying peroxidase described latest is VP, an enzyme that combines the catalytic properties of LiP and MnP, while its 3D structure resembles more LiP than MnP. Currently, VPs are characterized from two genera, Pleurotus and Bjerkandera, and evidence for their production has been reported for Panus, Trametes, and Spongipellis species (Ruiz-Dueñas et al., 2009).
The molecular masses of the LiP, MnP, and VP proteins vary between 35-48 kDa, 38-62 kDa, and 42-45 kDa, respectively. White-rot fungal lignin-modifying peroxidases have typically acidic pI values of 3.0-4.0 (Hatakka, 2001), while also neutral MnPs have been detected from litter- decomposing fungi (Steffen et al., 2002).
The overall amino acid sequences of fungal class II secretory peroxidases are well conserved. For example, two Ca2+-binding sites and eight cysteine residues that form four disulfide bridges are present to stabilize the protein structure and active site (Fig. 2).
The crystal structure of P. chrysosporium LiP H8 has been described in detail. The outmost residue needed for LiP activity is an invariant tryptophan, Trp171 in the isozyme LiPA (LiP H8) of P. chrysosporium. Necessity of the residue has been confirmed by several site-directed mutagenesis studies (Mester and Tien, 2001).
Figure 2. 3D model of Phanerochaete chrysosporium lignin peroxidase (LiP415) (Choinowski et al. 1999). Secondary structure elements (α- and 3-helices, blue; β-strands, orange arrows), two Ca2+ ions (purple spheres), and N- and C-termina are indicated. Heme group with proximal and distal histidine residues, four carbohydrate groups, Trp171, and disulfide bridges (S atoms, yellow spheres) are represented as ball and stick models. Reprinted with permission from Elsevier.
This tryptophan residue is situated in an exposed region of the enzyme surface (Fig. 2) and therefore it is thought to participate in the so-called long-range electron transfer from bulky aromatic substrates that cannot directly contact the oxidized heme in the active centre of LiP.A similar solvent-exposed tryptophan residue is conserved in all the cloned VP-encoding genes of Pleurotus and Bjerkandera spp. and it is needed for the LiP-like activity of VPs (Pérez-Boada et al., 2005).
1.3.1.2 Catalytic reactions of Lignin-modifying Peroxidases
In a H2O2 -dependent reaction, LiPs catalyze the initial one-electron oxidation of both phenolic and non-phenolic aromatic compounds, including the substructures of lignin, and several other substrates like veratryl alcohol. This leads to C –C cleavage aromatic ring oxidation, and cleavage reactions within the dimeric lignin-like model compounds (Kirk and Farrell, 1987; Hammel and Cullen, 2008).MnP catalyzes the specific oxidation of Mn2+ to Mn3+in the presence of H2O2.
Mn3+ions are stabilized in chelated form to perform oxidative reactions that yield organic radicals
from several phenolic substrates, carboxylic acids, and unsaturated lipids (Hammel and Cullen,
2008). The natural chelators of Mn3+are thought to be dicarboxylic acids, e.g. oxalic acid which is a common extracellular metabolite of white-rot fungi. The chelated Mn3+can diffuse even into the intact wood cell wall, the low porosity of which hinders the access from enzyme molecules. The Mn3+ions produced in MnP catalysis are not able to directly oxidize non- phenolic structures that comprise approximately 90% of lignin subunits in wood (Hammel and Cullen, 2008). This may be avoided by the subsequent reactions of Mn3+, which can result with e.g. lipid peroxidation, the radical chain reaction that has been shown to generate peroxyl radicals from lipids and also lead to the cleavage of non- phenolic synthetic lignin (Mäkelä, 2010).
VPs share the Mn2+-oxidizing activity with MnPs. Both MnP and VP have three conserved acidic
amino acid residues, two glutamates and one aspartate, which together with one of the heme propionates are involved in Mn2+-binding. However, VP has been shown to efficiently oxidize Mn2+ in the presence of only two acidic amino acid residues reflecting certain differences between these two enzymes (Ruiz-Dueñas et al., 2009).
1.3.1.3 Evolutional relations of Lignin-modifying Peroxidases
Lignin-modifying peroxidases are evolutionarily closely related and phylogenetic analyses divide them into several clearly defined main groups or subfamilies discriminated by certain key amino acid residues (Martínez 2002; Hildénet al., 2005; Ruiz-Dueñas et al., 2009). The first phylogenetic main cluster includes the typical mnp genes that code for proteins with long C- terminal tails and are found in e.g. Ceriporiopsis subvermispora, Dichomitus squalens, Phlebia radiata, and Phanerochaete chrysosporium. The second main group is formed by short MnPs, from e.g. Trametes versicolor and Pleurotus species, and VPs. LiPs are closely related to the short
MnP-VP group reflecting similar structural features between the short MnPs and LiPs (Hildén et al., 2005). The third main cluster of fungal class II peroxidases are the non-lignin-modifying, CIP-like peroxidases (Hildénet al., 2005). Interestingly, the same white-rot fungal species can express functionally similar but evolutionarily divergent MnPs as shown with P. radiata (Hildénet al., 2005) and Physisporinus rivulosus (Hakala et al. 2006).
1.3.1.4 Regulation of Lignin-modifying Peroxidase expression
Expression of the lignin-modifying peroxidases of white-rot fungi is often divergently regulated. The effect of different culture conditions and various supplements has been thoroughly investigated both at protein and transcript level. Expression of lignin- modifying peroxidases is commonly triggered e.g. by depletion of nutrients, oxidative stress, and heat shock (Belinky et al.,
2003).
Substrate-dependent expression of P. chrysosporium LiP-encoding genes has been detected on aspen wood chips, in defined liquid medium and in soil cultures (Boganet al., 1996). Spruce sawdust was shown to have a distinct effect on the transcript levels of P. rivulosus mnp genes (Hakalaet al., 2006). In accordance, production of P. radiata LiP isozymes has shown to be dependent on the lignocellulose materials used as carbon source (Mäkelä, 2010).
Nitrogen concentration (Hakala, 2007) and source, i.e. organic or inorganic nitrogen (Kaalet al.,
1993) are factors that affect fungal lignin-modifying peroxidase expression. For example, Pleurotus eryngii expresses one VP-encoding gene in peptone-containing liquid cultures while two allelic variants encoding another VP isozyme are expressed on lignocellulose cultures (Mäkelä, 2010).
Regulation of MnP expression by Mn2+ has been observed repeatedly in white-rot fungi. The
levels of different mnp transcripts vary in response to Mn2+ e.g. in P. chrysosporium,Pleurotus ostreatus (Cohenet al., 2001), C. subvermispora (Manubenset al., 2003), P. radiata (Hildénet al.,
2005), P. rivulosus (Hakalaet al., 2006), and Phlebia sp. MG-60 (Kameiet al., 2008). Putative metal response elements (MREs) are present in the promoter regions of several white-rot fungal mnp genes (Hildénet al., 2005) although the functionality of these elements needs still to be proven. Also aromatic compounds, such as veratryl alcohol and syringic acid may promote MnP production in white-rot fungal cultures (Hofrichter, 2002; Manubenset al., 2003; Hakalaet al.,
2006).
One fungal species typically harbours several genes for the lignin-modifying peroxidases. Close genomic organization of eight LiP and two MnP -encoding genes in P. chrysosporium and tandemly arranged LiP and MnP -encoding genes in T. versicolor have been reported. However, the relationship between LME gene clustering and transcriptional regulation is not apparent (Vanden Wymelenberget al., 2009).
1.3.2 Manganese peroxidase
Manganese peroxidase [EC 1.11.1.13, Mn (II): hydrogen-peroxide oxidoreductase, MnP]
catalyzes the Mn-dependent reaction.
2Mn (II)+2H++H2O2=2Mn (III)+2H2O
The first extracellular MnP was purified from P. chrysosporium, with its expression and production shown to be regulated by the presence of Mn (II) in the culture medium. Mn (II) controls the mnp gene transcription that is growth and concentration dependent. MnP is also regulated at the level of gene transcription by heat shock and H2O2. In addition to the stimulatory effect of Mn (II), organic acids, such as glycolate, malonate, glucuronate, 2-hydroxybutyrate, added to the medium enhances them production of MnP by the white-rot fungus Bjerkandera sp. strain BOS55. Bjerkandera species also produces versatile peroxidase that possesses both Mn- mediated and Mn-independent activity as described in the later section. Mn (II) also downregulates LiP titer in white-rot fungus, because of its suppression of the production of veratryl alcohol, which has been postulated to function in protecting LiP from inactivation by high levels of H2O2 (Wong, 2009).
1.3.2.1 Molecular Structure of MnP
The P. chrysosporium enzyme is an acidic glycoprotein with a pI near 4.5 and a Mr of 46,000. MnP is produced as a series of isozymes often coded and differentially regulated by different genes. MnP contains one molecule of heme as iron protoporhyrin IX and shows a maximal activity at Mn (II) concentrations above 100 μM. The heme iron in the native protein is in the high-spin, pentacoordinate, ferric state with a Hisresidue coordinated as the fifth ligand. The overall structure of P. chrysosporium. MnP is similar to LiP, consisting of two domains with the heme sandwiched in-between (Fig. 3).
Figure. 3: Three dimensional structure of P. chrysosporiummanganese peroxidase
(Sundaramoorthyet al., 2005)
The protein molecule contains ten major helices and one minor helix as found in LiP. MnP has five rather than four disulfide bonds, with the additional bond,Cys341–Cys348, located near the C terminus of the polypeptide chain. This additionaldisulfide bond helps to form the Mn (II)- binding site and is responsible for pushing the C terminus segment away from the main body of the protein. The distal His46 is hydrogen bonded to Asn80 to ensure that Nε2 is available to accept a proton from the peroxide in acid-base catalysis. The H bond formed between the proximal His173 and the side chain of Asp242 increases the anionic character of the ligand and helps stabilizing the oxyferry iron in MnP-I. The Mn (II) is located in a cation-binding site at the surface of the protein and coordinates to the carboxylate oxygens of Glu35, Glu39, and Asp179, the heme propionate oxygen, and two water oxygens. The site has considerable flexibility to accommodate the binding of a wide variety of metal ions (Sundaramoorthyet al., 2005). Two heptacoordinate structural calcium ions, one tightly bound on the proximal side and the other bound on the distal side of the heme, are important for thermal stabilization of the active site of the enzyme.
1.3.2.2 The Catalytic Cycle of MnP
Mn-dependent peroxidases are unique in utilizing Mn (II) as the reducing substrate (Glennet al.,
1985; 1986). MnP oxidizes Mn (II) to Mn (III), which in turn oxidizes a variety of monomeric
phenols including dyes as well as phenolic lignin model compounds. The catalytic cycle thus entails the oxidation of Mn (II) by compound I (MnP-I) and compound II (MnP-II) to yield Mn
(III).
MnP + H2O2 → MnP-I + H2O MnP-I + Mn2+ → MnP-II + Mn3+
MnP-II + Mn2+ → MnP + Mn3+ + H2O Mn (III) in turn mediates the oxidation of organic substrates.
Mn3+ + RH → Mn2+ + R. + H+
The characteristics of the cycle are very similar to that of LiP. Addition of 1 equivalent of H2O2 to the native enzyme yields MnP-I, which is a Fe (IV)-oxo-porphyrin radical cation [Fe (IV) =O. +]. The peroxide bond of H2O2 is cleaved subsequent to a 2e− transfer from the enzyme heme- porphyrin. The formation of MnP-I is pH independent, with a second-order rate constant of
2.0×106 M−1 s−1. Addition of 1 equivalent to Mn (II) rapidly reduces compound I to compound II. The conversion of MnP-I to MnP-II can also be achieved by the addition of other electron donors, such as ferrocyanide and a variety of phenolic compounds. In the reduction of compound II to generate the native enzyme, however, Mn (II) is an obligatory redox coupler for the enzyme to complete its catalytic cycle (Wong, 2009).
1.3.2.3 Mn (III) Chelators
The Mn (III) formed is dissociated from the enzyme and stabilized by forming complexes with α- hydroxy acids at a high redox potential of 0.8–0.9 V. Oxalate and malonate are optimalchelators that are secreted by the fungus in significant amounts. It has also been shown that MnP reacts with oxalate–Mn (II) instead of free Mn (II) as the true substrate with the chelator involved in the redox reaction of the metal. Other physiological functions have been associated with these chelators, including the enhancement of enzyme activity by their ability to facilitate the dissociation of Mn (III) from the enzyme. Oxalate has been proposed to function as an extracellular buffering agent, allowing the fungus to control the pH of its environment (Wong,
2009). It may also act as calcium sequester to increase the pore size of the plant cell wall and to facilitate the penetration of enzyme molecules. Oxidation of oxalic acid by Mn (III) produces a formate radical (HCO2.−) that reacts with dioxygen to form superoxide (O2.−) and, subsequently,
H2O2(Urzuaet al., 1998). This process has also been implicated in contributing to the ability of the fungus to degrade lignin.
1.3.2.4 Oxidation of Phenolic Substrates
The Mn(III) chelator complex acts as a diffusible oxidant of phenolic substrates involving1e− oxidation of the substrate to produce a phenoxy radical intermediate, which undergoes rearrangements, bond cleavages, and nonenzymatic degradation to yield various breakdown products (Fig. 4). MnP-generated Mn(III) can catalyze the oxidation of phenolic substrates, including simple phenols, amines, dyes, as well as phenolic lignin substructure and dimers. Mn(III) chelator is a mild oxidant under physiological conditions limited to the oxidation of phenolic lignin structures and, by itself, is not capable of oxidizing non-phenolic compounds (Wong, 2009).
Figure 4. MnP-catalyzed oxidation of phenolic aryglycerol β-aryl ether lignin model compound
Figure 5. MnP-catalyzed oxidation of non-phenolic β-O-4 lignin model compound (Kapichet al.,
2005).
1.3.2.5 Oxidation of Non-phenolic Substrates
For non-phenolic substrates, the oxidation by Mn (III) involves the formation of reactive radicals in the presence of a second mediator. This is in contrast to that of LiP catalyzed reaction, which involves electron abstraction from the aromatic ring forming a radical cation. Mn (III) in the presence of thiols, such as glutathione, mediates the oxidation of substituted benzyl alcohols and diarylpropane structures to their respective aldehydes (Reddyet al., 2003). In these reactions, Mn (III) oxidizes thiols to thiyl radicals, which subsequently Fig. 4. MnP-catalyzed oxidation of phenolic aryglycerol β-aryl ether lignin model compound abstract hydrogen from the substrate to form a benzylic radical. The latter undergoes nonenzymatic reactions to yield the final products. The enzyme generated Mn (III) also couples with peroxidation of lipids to catalyze Cα -Cβ cleavage and β-aryl ether cleavage of non-phenolic diarylpropane and β-O-4 lignin structures, respectively (Fig. 5)(Dainaet al., 2002; Kapichet al., 2005).The mechanism involves hydrogen abstraction from the benzylic carbon (Cα) via lipid peroxy radicals, followed by O2 addition to form a peroxy radical, and subsequent oxidative cleavage and nonenzymatic degradation. In the absence of exogenous H2O2, the enzyme oxidizes nicotinamide adenine dinucleotide phosphate (reduced form), glutathione, dithiothreitol, and dihydroxymaleic acid, generating H2O2. This
oxidase activity of MnP has implications in fungal lignin degradation, because the H2O2 produced may become available for the enzyme to start the peroxidase cycle. The H2O2 could also be utilized by lignin peroxidase, which is H2O2 dependent for catalytic activity (Wong, 2009).
1.3.2.6 Compound III
Similar to LiP and other peroxidases, MnP-II reacts with additional H2O2 to form a Fe(III) superoxo complex as compound III (MnP-III), which can be further oxidized by H2O2 resulting in heme bleaching and irreversible inactivation of the enzyme. The rate of MnP inactivation is one order of magnitude lower than that of LiP. The reactivation of MnP-III is mediated by Mn (III), which interacts either by oxidizing the iron-coordinated superoxide or reacting with H2O2in a catalase-type activity. This mechanism is in contrast to LiP where LiP-III is oxidatively converted back to the native LiP by the radical cation of aromatic substrates (Wong, 2009).
1.4 Fungal degradation of wood polysaccharides
1.4.1 Enzymatic decomposition of cellulose
In general, white-rot fungi express a set of hydrolytic enzymes for the degradation of cellulose. Endoglucanases (endo-1, 4-β-glucanases, EC 3.2.1.4) hydrolyze internal glycosidic bonds of the cellulose polymer while cellobiohydrolases (exo-1, 4-β- glucanases, EC 3.2.1.91) cleave the ends of cellulose chains resulting in the release of cellobiose. Moreover, cellobiohydrolase I (Cel7A) – type enzymes act on non-reducing ends of the cellulose chains, while other cellobiohydrolase II (Cel6A) -type enzymes act on reducing ends. β-glucosidases (EC 3.2.1.21) finalize the concerted action of cellulases by cleaving the released disaccharides to glucose molecules (Baldrian and Valášková, 2008). Expression and production of fungal cellulases is controlled by induction and repression mechanisms, including carbon catabolite repression (Kuhadet al., 1997). For example, the white-rot fungi Phlebia radiata and Dichomitus squalens secrete endoglucanases, cellobiohydrolases, and β-glucosidases under cellulose-containing liquid cultures. Also, a full array of cellulases has been identified in the transcriptome and proteome of Phanerochaete chrysosporium, both on solid-state wood and in cellulose-grown cultures (Vanden Wymelenberget al., 2005; Satoet al., 2009). The ascomycete Trichoderma reesei (teleomorph Hypocrea jecorina) is the major model fungus for cellulose decomposition and soft-rot type of wood decay. Surprisingly, among the ascomycete whole genome sequences, the T. reesei genome
reveals the smallest set of genes encoding enzymes involved in the decomposition of plant cell wall polysaccharides. Furthermore, T. reesei harbours even fewer cellulolytic and hemicellulolytic enzyme-encoding genes than is recognized in the genome of the white-rot basidiomycete P. chrysosporium. Although, still hyphothetical, efficient production of cellulases and control of gene expression have been suggested to explain the ability of T. reesei to cause powerful breakdown of cellulose and hemicellulose in natural lignocelluloses, regardless of the relatively low number of carbohydrate-active-enzyme (CAZyme) -encoding genes in the genome (Mäkeläet al., 2010).
Basidiomycetes and ascomycetes produce an additional extracellular enzyme, cellobiose dehydrogenase (CDH, EC 1.1.99.18), which oxidizes cellobiose to the corresponding lactone. The enzyme is often expressed by white-rot fungi but is so far identified only in a single brown-rot fungal species, Coniophora puteana. Furthermore, CDH is believed to play a role in degradation and modification of cellulose, hemicelluloses, and lignin by generating hydroxyl radicals in a Fenton-type reaction. CDH has been shown to be expressed during the growth of P. chrysosporium on solid-state wood and cellulose medium (Vanden Wymelenberget al., 2005; Satoet al., 2009) further supporting the role of this particular enzyme in wood degradation (Mäkeläet al., 2010).
1.4.2 Non-enzymatic decomposition of cellulose
Wood-decaying fungi, especially brown-rot fungi, are believed to degrade cellulose oxidatively by the means of hydroxyl radicals generated in the Fenton reaction
H2O2 + Fe2+ + H+ → H2O + Fe3+ + OH
The importance of Fenton chemistry in brown-rot fungal wood decay is recently emphasized by the whole genome sequence of Postia placenta, which harbours only two putative endoglucanases and several β-glucosidases, and totally lacks cellobiohydrolases (Martinezet al., 2009). In contrast, the P. placenta genome revealed a large variety of genes potentially involved in the generation of extracellular reactive oxygen species (ROS). Furthermore, transcripts of several genes putatively involved in the extracellular generation of Fe2+ and H2O2were also highly expressed during the growth of P. placenta on cellulose media (Martinezet al., 2009).
Some white-rot fungi produce ROS-quenching metabolites that can prevent the oxidative damage caused by active oxygen species. This may furthermore explain why these fungi leave wood
cellulose more or less intact. In Ganoderma species, amino acids, polysaccharides, and methanolic extract from mycelia were observed to act as ROS-converting compounds (Leeet al.,
2001; Tsenget al., 2008). Ceriporiopsis subvermispora produces itaconic (ceriporic) acids, which may suppress the Fenton reaction leading to diminished cellulose depolymerization (Rahmawatiet al., 2005).
Various fungal extracellular iron-chelating metabolites, e.g. siderophores and glycopeptides are thought to play a role in Fenton chemistry by reducing Fe3+back to Fe2+(Kuhadet al., 1997, Xu and Goodell, 2001). Quinones produced by fungi are also able to reduce Fe3+and contribute to a complete Fenton system in the so-called quinone redox cycling (Baldrian and Valášková, 2008) that has been shown to be a significant mechanism for cellulose cleavage in the brown-rot fungus Gloeophyllum trabeum (Suzukiet al., 2006).
Oxalic acid, secreted in relatively high concentrations by brown-rot fungi, is also proposed to participate in decomposition of cellulose. Oxalic acid strongly chelates Fe3+into soluble complexes which predominate in brown-rot wood decay (Suzukiet al., 2006). Iron can be sequestered from Fe3+-oxalate complexes and reduced back to Fe2+thus promoting the continuation of Fenton reaction (Xu and Goodell, 2001; Varela and Tien, 2003). In addition, the autooxidation of Fe2+-oxalate complexes can lead to the slow production of hydroxyl ions even when quinones are not available (Suzukiet al., 2006). On the other hand, abundance of oxalic acid is believed to suppress Fenton reaction and protect the fungal hyphae from oxidative damage by scavenging hydroxyl ions.
1.4.3 Decomposition of hemicellulose by basidiomycetous fungi
Due to the heterogeneous structure and organization of hemicellulose, a number of different CAZymes are required for its degradation. White-rot fungi secrete various glycoside hydrolases that cleave glycosidic bonds in the hemicellulose polymers, as well as carbohydrate esterases that hydrolyze ester linkages of acetate and the ferulic acid side groups. Carbohydrate esterases include e.g. feruloyl esterases (EC 3.1.1.73) which catalyze the hydrolysis of ester bond between arabinose subunits and ferulic acid involved in cross-linking of xylan to lignin (Kuhadet al.,
1997; Shallom and Shola,. 2003). Endo-1, 4-β-xylanases (EC 3.2.1.8) and endo-1, 4-β- mannanases (EC 3.2.1.78) are the two main enzymes degrading the backbone of wood hemicelluloses. Several enzymes are responsible for further hydrolysis of the formed
oligosaccharides (e.g. β-1,4- xylosidase, EC 3.2.1.37) and side groups (e.g. α-L-arabinosidase, EC
3.2.1.55) (Kuhadet al., 1997).
The recent brown-rot fungal transcriptome and secretome analysis of Postia placenta grown on cellulose revealed the expression of several hemicellulases (Martinezet al., 2009). Hemicellulase activities in white-rot fungi have been detected for example in the cultures of Dichomitus squalensand have been studied in wheat bran cultures of Phlebia radiate.
In the cultures mimicking lignin-degrading conditions, the Phanerochaete chrysosporium secretome has been shown to contain several hemicellulases together with LMEs (Vanden Wymelenberget al., 2006). This may be related to the degradation of covalently linked lignin- hemicellulose matrix in the wood cell walls. However, a somewhat narrower selection of hemicellulases was shown in the proteome and transcriptome studies of P. chrysosporium when the fungus was cultivated on solid-state wood (Satoet al., 2009) as compared to cellulose- containing cultures (Vanden Wymelenberget al., 2005).
1.5 Fungal low molecular weight compounds and wood degradation
Wood-decaying fungi produce several chemically diverse low molecular weight compounds, which have an impact on lignocellulose degradation. Low molecular weight compounds, such as phenols synthesized by fungi may be oxidized as substrates by the fungal LMEs. In consequence, this may lead to formation of free radicals which furthermore transfer oxidative reactivity to the lignocellulose matrix. Low-molecular weight compounds may also promote LME activity by stabilizing the reactive oxidants formed during enzyme catalytic action. Small organic molecules which readily diffuse away from the fungal hyphae are suggested to be important especially in the beginning of wood decay since the extracellular LMEs (laccases, LiPs, MnPs, VPs) are too large in size in order to penetrate into the intact wood cell walls. Veratryl (3,4-dimethoxybenzyl) alcohol (VA), which is a substrate for LiP, is a natural metabolite of a few white-rot fungi, e.g. Phanerochaete chrysosporium, Pycnoporus cinnabarinus, and Phlebia radiata. VA cation radical is most likely too short-lived to act as a far-diffusing redox-mediator upon LiP-catalysis under natural wood-decaying conditions. However, updated with the current knowledge of the LiP 3D structure, VA is oxidized by LiP at a very specific site (i.e. exposed tryptophan residue) on the enzyme surface (Mester and Tien, 2001; Johjimaet al.,2002), which does not rule out the role of VA as a potential protector of LiP from an inactivation caused by H2O2. In addition to LiP, the
fungal H2O2-producing enzyme aryl-alcohol oxidase (AAO) may use VA as a reducing substrate. In the case of laccase, the natural redox- mediator is also an aromatic compound, 3- hydroxyanthranilate (3-HAA), which is found to occur in the cultures of P. cinnabarinus. 3-HAA expands the oxidation capacity of laccase to non-phenolic and polymeric compounds. Cultures of many white-rot fungi including Bjerkandera adusta, Pleurotus pulmonarius, and Phlebia radiata become accumulated with fatty acids generated by the fungus (Gutiérrezet al., 2002). Unsaturated fatty acids participate in MnP-catalyzed lipid peroxidation reactions resulting with oxidation and even carbon-carbon bond cleavage of non-phenolic lignin substructures (Mäkeläet al., 2010). Involvement of phospholipids and membrane-released fatty acids may have a versatile regulatory impact on fungal decay of wood and lignocellulose. Several alkyl- and alkenylitaconic acids (ceriporic acids), produced by the selective lignin-degrading white-rot fungus Ceriporiopsis subvermispora were shown to repress Fenton reaction and concomitantly diminish depolymerization of cellulose (Rahmawatiet al., 2005). Quenching of cellulose degradation may in turn explain why this fungus leaves most of the wood cellulose intact while decaying lignin and hemicelluloses (Fackleret al., 2006).
White- and brown-rot fungi produce various iron-chelating compounds, e.g. glycopeptides, siderophores, oxalic acid, phenolates, and other monomeric aromatic compounds which are important for example in Fenton-type reactions. Oxalic and other carboxylic acids are generally secreted metabolites of fungi, and diverse functions of oxalic acid in wood degradation (Mäkeläet al., 2010).
1.6 Organic acids secreted by wood-decaying fungi
Wood-decaying fungi typically acidify their growth environment quickly by secreting organic acids. Several organic acids have been detected on defined liquid media and in lignocellulose- containing cultures of white-rot fungi. Oxalic acid is the most commonly secreted fungal acid. In brown-rot fungi, production of other organic acids than oxalate has not so far been reported. The amount and diversity of organic acid production vary between fungal species, and secretion of carboxylic acids depends on the cultivation conditions (Aguiaret al., 2006). Organic acids originating from e.g. tricarboxylic (TCA) cycle are secreted as waste compounds of fungal cellular metabolism. The smallest organic acids, such as formic and oxalic acid, may also
accumulate in fungal cultures as by-products of the cleavage of lignin substructures, such as side- chains and aromatic rings (Hofrichter, 2002).
1.6.1 Oxalic acid
Oxalic acid is a compound that is toxic to almost all organisms. It is the strongest dicarboxylic acid and has two pK values at 1.23 and 4.26. Oxalic acid is a major chelator of metal cations, e.g. Fe2+, Mn2+, Ca2+, and Al3+, and participates in various environmental and biological processes. Interestingly, oxalic acid plays several important roles in fungal growth and metabolism and is also connected to biological mechanisms underlying fungal pathogenesis (Dutton and Evans, 1996).
1.6.1.1 Fungal synthesis of oxalic acid
Fungi synthesize oxalic acid as a metabolic waste compound through the TCA cycle in mitochondria and by the so called glyoxylate cycle that operates in glyoxysomes (Dutton and Evans, 1996, Munir et al., 2001b). More recently, the glyoxylate cycle has been proposed to take place in other organelles, the peroxisomes, after the studies of the brown-rot fungus, Fomitopsis (Tyromyces) palustris (Sakaiet al., 2006). The biosynthesis of oxalic acid is catalyzed by the intracellular enzymes oxaloacetase (EC 3.7.1.1), glyoxylate oxidase, and cytochrome c-dependent glyoxylate (Mäkeläet al., 2010).
Carbon catabolite repression of the glyoxylate cycle by glucose that is typically observed in bacteria does not seem to operate in wood-rotting basidiomycetes, thereby allowing fungi to secrete substantial amounts of oxalic acid. A unique metabolic linkage between the TCA and glyoxylate cycles has been shown to be central in the oxalic acid biosynthesis of F. palustris (Muniret al., 2001a) and the brown- rot model fungus Postia placenta. Furthermore, this metabolic shunt has been proposed to be a general feature of wood-rotting fungi, a means of acquiring energy for growth during wood decay by oxidizing released glucose to oxalic acid (Muniret al., 2001a). In contrast to this hypothesis, exposure of the white-rot fungus Phanerochaete chrysosporium to vanillin that is structurally related to lignin subunits, caused a drastic change from glyoxylate cycle to TCA cycle, and a flow of TCA cycle metabolites into the heme biosynthesis pathway was observed (Shimizuet al., 2005).
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EXTRACTION AND CHARACTERIZATION OF MANGANESE PEROXIDASE (MNP) FROM RIGIDOPORUSLIGNOSUS, A WHITE ROOT ROT FUNGI OF RUBBER TREE.
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