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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 (ΔH, ΔG, ΔS) 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.

TABLE OF CONTENTS

Title page i
Certification ii
Dedication iii
Acknowledgements iv
Abstract v
Table of contents vi
List of Tables ix
List of Figures x
List of Abbreviation xii
CHAPTER ONE: INTRODUCTION
1.1 Wood composition 4
1.2 Wood degradation by fungi 6
1.3 Lignin-modifying enzymes 9
1.3.1 Lignin-modifying peroxidases 10
1.3.1.1 Occurrence and properties of lignin-modifying peroxidases 13
1.3.1.2 Catalytic reactions of lignin-modifying peroxidases 13
1.3.1.3 Evolutional relations of lignin-modifying peroxidases 13
1.3.1.4 Regulation of lignin-modifying peroxidase expression 14
1.3.2 Manganese peroxidase 15
1.3.2.1 Molecular structure of mnp 15
1.3.2.2 The catalytic cycle of mnp 16
1.3.2.3 Mn(III) chelators 17
1.3.2.4 Oxidation of phenolic substrates 18
1.3.2.5 Oxidation of non-phenolic substrates 19
1.3.2.6 Compound III 20
1.4 Fungal degradation of wood polysaccharides 20
1.4.1 Enzymatic decomposition of cellulose 20
1.4.2 Non-enzymatic decomposition of cellulose 21
1.4.3 Decomposition of hemicellulose by basidiomycetous fungi 22
1.5 Fungal low molecular weight compounds and wood degradation 23
vii
1.6 Organic acids secreted by wood-decaying fungi 24
1.6.1 Oxalic acid 25
1.6.1.1 Fungal synthesis of oxalic acid 25
1.6.1.2 Roles of fungal-produced oxalic acid 25
1.6.2 Oxalic-acid degrading enzymes 27
1.7 Basidiomycete genomes and lignocellulose decay 28
1.8 White-rot fungi and their enzymes in biotechnological applications 30
1.9 Aims and objectives 33
1.9.1 Aim of the research 33
1.9.2 Objectives of the study 33
CHAPTER TWO: MATERIALS AND METHODS
2.1 Materials 34
2.1.1 Reagents 34
2.1.2 Apparatus 35
2.1.3 Plant material 35
2.2 Methods 35
2.2.1 Isolation andculture of the organism 35
2.2.1.1 Preparation of potatoes dextrose agar (PDA) for fungi isolation 35
2.2.1.2 Preparation of liquid broth for isolation of microorganisms 37
2.2.1.3 Inoculation of plates and sub-culturing 37
2.2.2 Fungi identification 37
2.2.3 Extraction of enzyme (manganese peroxidase) 37
2.2.3.1 Submerged fermentation of Rigidiporuslignosus using glucose as
carbon source 38
2.2.3.2 Mass production of enzymes 38
2.2.4.1 Manganese peroxidase assay using phenol red, and manganese as substrates 38
2.2.4.2 Protein determination using lowry method 38
2.2.5.1 Ammoniumsulphate precipitation 39
2.2.5.2 Gel filtration chromatography 39
2.2.6 Characterization of enzyme 39
2.2.6.1 Effect of pH on manganese peroxidaseactivity 39
2.2.6.2 Effect of temperature on manganese peroxidaseactivity 40
2.2.6.3 Effect of H2O2 on manganese peroxidase activity 40
viii
2.2.6.4Effect of phenol red on peroxidase activity 40
2.2.7 Inactivation of peroxidase by hydrogen peroxide 40
2.2.8 Calculation of kinetic parameters 40
CHAPTER THREE: RESULTS
3.1 Pure sub-cultured fungi species 42
3.2 Effect of pilot study days on manganese peroxidaseproduction 42
3.3 Purification of crude manganese peroxidase 42
3.3.1 Ammoniumsulphate precipitation 42
3.3.2 Gel filtration 42
3.3.3 Purification table 49
3.4 Characterization of partially purified enzyme 49
3.4.1 Effect of pH on manganese peroxidase activity 49
3.4.2 Effect of temperature 49
3.5 Variation of manganese peroxidase activity with different concentrations
hydrogen peroxide 49
3.5.1 Determination of Km and Vmax 49
3.6 The Lineweaver-Burk plot of effect of different concentrations of
phenol red onmn-peroxidase activity. 54
3.7 The inactivation of manganese peroxidase 54
3.8 Temperature dependence of the decimal reduction of peroxidase to
calculate Z-value 54
3.9 Kinetic parameters for heat inactivation of manganese peroxidasefrom
Rigidporuslignosus 59
3.10 Thermodynamic analysis of thermal denaturation 59
CHAPTER FOUR: DISCUSSION
4.1 Discussion 62
4.2 Conclusion 65
4.3 Suggestions for further studies 65
REFERENCES 66
APPENDICES 81

CHAPTER ONE

Project Topics

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),
2
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
3
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 Manganesedependent
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, wood4
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
5
(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 ligninhemicellulose
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 nonstructural
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.
6
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).
7
Following the action of white-rot fungi, the decayed wood is characteristically white and fibrelike.
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
8
Microorganism Type Enzyme
Bjerkanderasp. Basidiomycete LiP,MnP
Ceriporiopsis subvermispora Basidiomycete LiP
Chaetomium thermophilium Ascomycete Lac
Chrysosporium lignorum Basidiomycete LiP,MnP
Coriolushirsutus Basidiomycete MnP
Dichomitussqualens Basidiomycete MnP,Lac
Elfvingiaapplanata Basidiomycete MnP,Lac
Flavodonflavus Marine basidiomycete MnP,Lac
Halosarpheia ratnagiriensis Marine ascomycete Lac
Halosarpheia ratnagiriensis Marine ascomycete Lac
Irpexflavus Basidiomycete MnP
Irpexlacteus Basidiomycete MnP,Lac
Lentinusedodes Basidiomycete LiP,MnP, Lac
Lentinussquarrosulus Basidiomycete LiP
Nematolomafrowardii Basidiomycete LiP,MnP
Oxyporus latemarginatus Basidiomycete LiP,MnP
Phanerochaete chrysosporium Basidiomycete LiP,MnP, Lac
Phanerochaetecrassa Basidiomycete MnP
Phanerochaeteflavido alba Basidiomycete LiP,MnP
Phanerochaete magnolia Basidiomycete LiP,MnP
Phanerochaete sordida Basidiomycete LiP,MnP
Phellinuspini Basidiomycete LiP,MnP
Phlebiaradiate Basidiomycete LiP,MnP, Lac
Phlebiasp. Basidiomycete MnP,Lac
Phlebiasubserialis Basidiomycete MnP
Phlebiatremellosa Basidiomycete MnP
Pleurotuseryngii Basidiomycete MnP,Lac
Pleurotusostreatus Basidiomycete MnP,Lac
Pleurotussajor-caju Basidiomycete MnP,Lac
Poluporussp. Basidiomycete LiP,MnP, Lac
Polyporussanguineus Basidiomycete MnP
Psathyrella atroumbonata Basidiomycete LiP
Pycnoporus cinnabarinus Basidiomycete MnP,Lac
Rigidosporuslignosus Basidiomycete MnP,Lac
Schizophyllum commune Basidiomycete Lac
Sordariafimicola Ascomycete Lac
Stereumannosum Basidiomycete Lac
Stereumhirsutum Basidiomycete LiP,MnP, Lac
Trametes(CoriolusorPolyporus)versicolor Basidiomycete LiP,MnP, Lac
Trametestrogii Basidiomycete LiP,MnP, Lac
Trichodermaatroviride Ascomycete Lac
LiP:ligninperoxidase;MnP:manganese peroxidase;Lac:laccase (Steffen, 2003).
9
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
10
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 hemecontaining,
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 CIPenzyme
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
11
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 litterdecomposing
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).
12
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).
13
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 Cterminal
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
14
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).
15
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 Mnmediated
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).
16
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
17
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 hemeporphyrin.
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,
18
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
19
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
20
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
21
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
22
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
23
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 ligninhemicellulose
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 cellulosecontaining
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
24
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 lignocellulosecontaining
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
25
accumulate in fungal cultures as by-products of the cleavage of lignin substructures, such as sidechains
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
26
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).
1.6.1.2 Roles of fungal-produced oxalic acid
Fungal species belonging to Ascomycota, Basidiomycota, and Zygomycota are known to secrete
considerable quantities of oxalic acid. Factors that affect the fungal production of oxalic acid
include carbon and nitrogen sources, and pH of the growth environment (Dutton and Evans,
1996). Several plant pathogenic fungi secrete oxalic acid to aid in defeating their host plant. In
fact, oxalic acid secretion by Sclerotinia sclerotiorum was reported to induce a programmed cell
death response in plant tissue (Kim et al., 2008a). Calcium oxalate crystals formed by fungi are
frequently found in decayed wood and in soil. Oxalic acid may sequester Ca2+ from the middle
lamella of plant cell wall resulting with calcium oxalate crystal formation. During wood decay,
this can lead to increased pore size within the wood tracheids and fibers, which facilitates
penetration of fungal extracellular enzymes into the inner layers of wood cell wall.
Oxalic acid may inhibit the growth of more sensitive fungi, thus having an impact on competition
between fungal species. For plant pathogenic fungi, secretion of oxalic acid is one of the factors
promoting hyphal penetration and weakening of host defence. Secretion of oxalic acid by litterdecomposing
and mycorrhizal fungi also increases the availability of soil nutrients (Dutton and
Evans, 1996). Leaching by oxalate has been shown to have an important role in solubilization of
radioactive uranium oxides (Fominaet al., 2007). On the other hand, fungi can tolerate high
environmental concentrations of toxic metals by secreting oxalic acid for chelation of cationic
metals into insoluble form (Jarosz-Wilkolazka and Graz, 2006).
Oxalic acid is a common metabolic product of wood-rotting fungi, including both white- and
brown-rot fungal species (Green and Clausen, 2003; Hakalaet al., 2005, Aguiaret al., 2006).
White-rot fungi typically accumulate oxalic acid to their growth medium in millimolar quantities
whereas in the cultures of brown-rot fungi, even several ten folds higher oxalic acid quantities are
often detected (Dutton and Evans,1996). Previously, such high concentrations of oxalic acid were
explained by the lack of oxalate decarboxylase (ODC) enzyme in the brown-rot fungi. However,
at least inPostia placenta, production and expression of ODC has been shown (Martinez, 2009).
One explanation for the production of substantial amounts of oxalic acid by cellulose-degrading
brown-rot fungi is the fact that wood as growth substrate is rich with carbon and scarce with
27
nitrogen (Eaton and Hale, 1993). By secreting oxalic acid the wood-decaying fungi can get rid of
the excess wood carbon and keep their nutritional C/N ratio in balance. Also, the difference in
accumulation of extracellular oxalic acid between brown- and white-rot fungi may be a result of
the lignin-degrading activity of white-rot fungi in which oxalic acid is consumed (Mäkeläet al.,
2010).
A number of roles for oxalic acid in fungal degradation and conversion of lignin have been
proposed. Oxalic acid lowers the pH of fungal extracellular environment to the optimal levels (pH
2-5) that are usually needed for the activity of LMEs. Poppet al. (1990) demonstrated that oxalic
acid is capable of mediating the oxidation of Mn2+ to Mn3+ via LiP and veratryl alcohol, thus
enabling the oxidation of compounds that are not preferred substrates or directly oxidized by LiP.
On the other hand, oxalic acid may also inhibit the LiP-catalyzed oxidation of veratryl alcohol.
Physiological concentrations of oxalic acid have been shown to stimulate MnP activity by
chelating unstable Mn3+. MnP can generate H2O2 by oxidation of oxalic and also glyoxylic acid
thus providing an endogenous source for extracellular H2O2 (Mäkeläet al., 2010).
During biopulping process with Ceriporiopsis subvermispora the oxalic acid secreted by the
fungus forms oxalate esters that contribute to the softening of wood fibers. The same mechanism
has been suggested to be involved in naturally occurring white-rot decay process. Oxalic acid is
also shown to contribute to the decrease of wood carbohydrate content (Suzukiet al., 2006). As an
indication, the brown-rot model fungus Postia placenta accumulates oxalic acid when colonizing
wood, whereas non-decay isolate of P. placenta is unable to secrete oxalic acid (Mäkeläet al.,
2010).
1.6.2 Oxalic-acid degrading enzymes
Three types of enzymes that catalyze oxalic acid degradation have been described from microbes
and plants: oxalate decarboxylases (ODC, EC 4.1.1.2), oxalate oxidases (OXO, EC 1.2.3.4), and
oxalyl-CoA decarboxylases (OXC, EC 4.1.1.8) (Svedružićet al., 2005). ODC, isolated from fungi
and bacteria, decomposes oxalic acid to formic acid and CO2via electron withdrawal in a very
specific single-step reaction (Fig. 6a). The enzyme contains catalytical Mn2+ ions and requires O2
for catalysis, although the overall reaction does not stoichiometrically utilize oxygen (Fig. 7a)
(Justet al., 2004).
28
The evolutionarily closely related enzyme, OXO, is oxidized by O2 but cleaves oxalic acid to two
CO2molecules with generation of H2O2 (Fig. 5b). OXO is known mainly from plants, and the
only two fungal OXOs are from the white-rot fungi Ceriporiopsis subvermispora andAbortiporus
biennis (Grązet al., 2009). In fact, C. subvermispora is the first fungus in which both ODC and
OXO activities have been detected (Watanabeet al., 2005).
The third oxalate-cleaving enzyme, OXC, which is a bacterial enzyme, convertsactivated
oxalyl-CoA to formyl-CoA and CO2 (Fig. 6c). It has thiamin pyrophosphateas a cofactor. A
number of bacterial species like Bifidobacterium lactis, Lactobacillus acidophilus, Oxalobacter
formigenes, and Thiobacillus novellus use OXC for the breakdown of oxalate (Anatharamet al.,
1989; Federiciet al., 2004; Turroniet al., 2007), and the enzyme is connected to oxalatedependent
ATP synthesis at least in O. formigenes (Anatharam et al 1989).
Figure 6:Schematic presentation of oxalate-degrading reactionscatalyzedby A)oxalate
decarboxylase, B) oxalate oxidase, and C) oxalyl-CoA decarboxylase. ThDP, thiamin
pyrophosphate. The figure is modified from Svedružić et al. (2005) and reprinted with permission
from Elsevier.
1.7 Basidiomycete genomes and lignocellulose decay
As at August 2009, nine complete genome sequences of different basidiomycetous fungal species
are available. The representatives of divergent basidiomycetous subphyla and order have been
among the first fungi selected for whole genome sequencing. These include the crop plant
29
pathogens Ustilago maydis and Puccinia graminis and the coniferous tree pathogen
Heterobasidion annosum spp. which cause remarkable economic losses, as well as a human
pathogen, Cryptococcus neoformans. Also species which have been studied as model organisms
for fungal genetics and development, i.e. Coprinopsis cinerea (Coprinus cinereus) and
Schizophyllum commune have been targets of whole genome sequencing projects (Mäkeläet al.,
2010).
Current sequencing efforts are turning from yeasts and pathogens to other filamentous fungi due
to their applicability for diverse biotechnological processes, in particular in conversion of
lignocellulose and plant material for production of biofuels and sustainable energy. The whole
genome sequence of the ecologically interesting ectomycorrhizal symbiotic species Laccaria
bicolor has been annotated. The first published white-rot fungal and also basidiomycetous whole
genome sequence was from Phanerochaete chrysosporium, and it promoted vital progress in the
molecular genetics of lignin-degrading white-rot fungi. Very recently, genome sequence data of
two other wood-colonising, saprobic white- rot fungi, i.e. S. commune and Pleurotus ostreatus
came available. In addition, the first published whole genome sequence of a brown- rot fungus,
Postia placenta (Martinezet al., 2009), adds up to the pool of fungi which have biotechnological
interest e.g. for biomass conversion.
Present genomic data clearly suggests a specific role for the lignin-modifying peroxidases (LiPs,
MnPs, VPs) in decomposition of lignin since the corresponding genes are found solely in the
white-rot fungi. Laccases are absent from P. chrysosporium genome and therefore they might be
unessential for lignin degradation. Instead, the P. chrysosporium genome contains other types of
multicopper oxidases, such as Fe-oxidoreductases (Larrondoet al., 2007), which are essential for
other cellular functions. For laccases, roles in processes other than lignin decay are probable
while even up to 17 laccase genes are found in the genome of the non-lignin-decaying, soilinhabiting
basidiomycete C. cinerea (Kilaruet al., 2006). In another non-lignin-degrading
basidiomycete L. bicolor, 9 laccase-encoding genes have been annotated (Courtyet al., 2009).
However, the importance of laccases is emphasized also for lignin-degrading white-rot fungi that
harbour multiple laccase genes which is further confirmed by whole genome sequence data from
P. ostreatus with 12 putative laccase-encoding genes (Mäkeläet al., 2010).
According to the current genomic data, strategies for cellulose degradation obviously differ
between white- and brown-rot fungi. The genome of the brown-rot fungus P. placenta harbours
30
several β-glucosidase-encoding genes, but totally lacks the type of carbohydrate-active enzymes
(CAZymes) that contain cellulose-binding modules, and has only two putative endoglucanaseencoding
genes (Martinezet al., 2009). On the contrary, the whole repertoire of multiple
cellulose-decomposing enzymes (endoglucanases, cellobiohydrolases, β-glucosidases), with and
without cellulose- binding modules, is present in the P. chrysosporium genome. In this respect,
the P. chrysosporium genome resembles that of the efficient cellulose-decaying, soft-rotting
ascomycetous fungus Trichoderma reesei (Mäkeläet al., 2010).
Despite the accumulating genomic data, physiologically related but taxonomically divergent
white-rot fungal species are still needed to be sequenced in order to reveal more of the genetic
factors required for efficient lignin-degradation. With multiple whole genome sequences from
near- and far-related fungal taxons, comparative genome analyses may be carried out to
investigate fungal wood-decay strategies. In addition, more whole genome sequences of white-rot
fungi representing selective and non-selective degraders of lignin are needed in order to
understand the fundamental differences in decomposition of lignocelluloses. The forthcoming
whole genome sequence from the selective lignin-degrading white-rot fungus Ceriporiopsis
subvermispora , the brown- rot fungus Serpula lacrymans, and the litter-decomposing, edible
fungus Agaricusbisporus, are currently under refining and annotation
(http://www.jgi.doe.gov/genome- projects/). Together with the accumulating whole genome
sequence data, novel genetic transformation systems for basidiomycetous species together with
efficient and precise gene knock-out and silencing systems are needed to untangle the
mechanisms of lignin and lignocellulose degradation. In fact, encouraging progress in fungal gene
silencing by RNA interference technique was recently shown in P. chrysosporium (Matityahuet
al., 2008).
1.8 White-rot fungi and their enzymes in biotechnological applications
Several biotechnological applications take advantage of white-rot fungi and their LMEs. For
example the utilization of fungi in the pulp and paper industry has been intensively studied.
Biopulping has gained a lot of interest in the past decades. Selectively lignin-degrading white-rot
fungi, e.g. Ceriporiopsis subvermispora, Physisporinus rivulosus, and Dichomitus squalens, have
been regarded as the most suitable organisms for biopulping, to minimize cellulose loss during
the fungal pretreatment (Hakala, 2007).
31
The direct use of white-rot fungal LMEs in pulp and paper industry could result in easier
optimization and applicability as compared to fungal treatment. For example the use of MnP has
been demonstrated to improve pulp bleaching (Moreiraet al., 2003; Feijooet al., 2008) and
decrease consumption of refining energy during paper manufacturing. The unspecific nature and
high redox potential of the lignin-modifying peroxidases (LiPs, MnPs, VPs) enable them to
convert various recalcitrant compounds. Therefore lignin-modifying peroxidases have been
extensively studied in bioremediation of e.g. soil and industrial effluents contaminated with
various harmful compounds of natural and anthropogenic origin (Hofrichter, 2002; Husain, 2006;
Haritash and Kaushik, 2009). However, the industrial use of fungal peroxidases is still hindered
by their high cost and low yields so far gained using heterologousexpression systems. Also, the
limited availability and low stability of the enzymes aswell as their inactivation by H2O2 and
elevated temperatures are problems in large-scaleproduction and in industrial applications
(Martínezet al., 2009).
Laccases, with prospects in diverse industrial areas, are the most studied fungal oxidoreductases
for biotechnological applications. Laccases have potential e.g. in foodindustry to improve dough
properties, and in cosmetic industry laccases may be added inproducts intended for skin
lightening and hair dyeing. In biotechnological applicationslaccases have potential as biosensors
and as enzyme-electrodes in biofuel cells (Couto and Toca-Herrera, 2007). Attention is paid also
to the exploitation of laccasesin synthesis of new biomaterials and polymers (Mikolasch and
Schauer, 2009).
In forest products industry, laccases could be used in biografting of low molecularweight
compounds onto lignocellulosic materials, and in cross-linking of fibers and ligninmoieties for the
production of wood composite products (Widsten and Kandelbauer, 2008a). Furthermore,
laccases could be used in biopulping, pitch control, deinking,and process water treatment, among
many other applications (Widsten andKandelbauer, 2008b). In fact, commercial laccase and
laccase-mediator applications e.g.for pulp and paper (delignification) and textile industry
(bleaching) are already available(Morozovaet al., 2007b).Since specific enzyme properties are
needed for different applications, molecularcharacterization of new fungal laccases has been in
focus. One property often desiredin industrial processes is enzyme thermotolerance, and some
thermotolerant andthermostable laccases from basidiomycetes have been described (reviewed by
Hildénet al., 2009). The commercial use of laccase mediators is still often hindered by their
32
highcost and lack of information on their toxicological safety. Recently, efforts have beenmade to
find efficient naturally occurring and cost-effective laccase mediators, whichcould remove these
obstacles (Camarero et al., 2007).The ascomycetous fungal genera Aspergillus and Trichoderma
comprise severalisolates demonstrating excellent protein production and secretion capacity, and
are themost promising host organisms for the heterologous expression of
basidiomycetousenzymes, such as the LMEs (Mäkeläet al., 2010).
Furthermore, the use of fungal hosts has an advantageover bacterial systems because fungi
perform the correct post-translational modificationsneeded for enzyme activity. Although a few
studies show successful heterologousproduction of e.g. MnP in ascomycetous hosts, the main
problem is the incorporationof heme to achieve a reasonable yield of active recombinant
peroxidase (Conesaet al., 2002).Heterologous expression of laccases by filamentous fungi with
increased enzymeproduction levels has been demonstrated in some cases (Recordet al., 2002;
Kiiskinenet al., 2004). Still, the lack of an efficient heterologous production system for laccasesat
bioreactor scale is constraining more bulky and industrial applications (Couto andToca-Herrera,
2007). Fungal laccases with improved catalytic properties and increasedstability have been
achieved by the use of molecular evolution techniques (Huet al., 2007; Festaet al., 2008). Future
studies will concentrate on improvement of the catalyticproperties of fungal LMEs by
mutagenesis, and to increase the production yield of activerecombinant enzyme in order to fit the
requirements of large-scale production (Mäkeläet al., 2010).
The global demand for the use of renewable materials for production of energyand consumables
has been expanded. So called biorefinery concept aims at the coproductionof various value-added
end products like biofuels and chemicals inadvanced biotechnological processes (e.g. enzymatic
hydrolysis followed by microbialfermentation) from renewable biomass. Forest and agricultural
waste lignocellulosesform a massive source of renewable biomass that can be used as a feedstock
for biorefining (Kumaret al., 2008 and Foustet al., 2008). Since lignin in plant cell walls prevents
the efficient use of cellulose andhemicellulose, its removal is a key step for the use of cellulosic
biomass. This highlightsthe role of white-rot fungi and their enzymes as environmentally friendly
biocatalysts forthe pretreatment of lignocelluloses (Ruiz-Dueñas and Martínez, 2009).
In addition to the LMEs, other fungal enzymes are biotechnologically promisingas well. For
example, oxalic-acid degrading enzymes like ODC that catalyze highlyspecific reactions have
several potential and established uses in diverse biotechnologicalapplications. One common
33
problem in many industrial processes is the formationof oxalate salt deposits which may
harmfully clog pipeworks and filters. In order toprevent formation of calcium oxalate, the so
called scaling, fungal ODC has been testedfor removal of oxalic acid from the bleaching filtrates
of the pulp and paper factories(Sjödeet al., 2008).Commercial assays are available to use ODC
for determination of oxalic acidconcentrations in clinical and food samples. Excessive excretion
of urinary oxalate(hyperoxaluria) can lead to the formation of calcium oxalate precipitates which
end upwith the formation of kidney stones. To find the treatment for hyperoxaluria, oral
therapywith crystalline cross-linked formulation of ODC has been shown to reduce symptoms
inexperiments with mice (Gruijicet al., 2009). Furthermore, lactic acid bacteria
expressingheterologous ODC could be used as possible probiotics for depletion of intestinal,
dietaryoxalic acid (Kolandaswamyet al., 2009).
In diverse application studies on fungal ODC, one goal has been the constructionof odcexpressing
transgenic crop plants. Contributing to their reduced content ofoxalic acid, transgenic
plants have proved to be resistant to certain oxalic-acid secretingpathogenic fungi. These plants
are also less toxic to herbivores, which lack oxalatedegradingenzymes and thereby are dependent
on intestinal bacteria to catabolize thedietary oxalic acid (Diaset al., 2006).
1.9 Aim and Objectives
1.9.1 Aim of the study
MnP is thought to play the most crucial role in lignin degradationof which its potential
applications 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. This study is then aimed at screening, partial purification and characterization
of MnP from Rigidoporus lignosususing a submerged fermentation system and determining the
effect of different concentrations of hydrogen peroxides and Phenol red on the MnP.
1.9.1 Objectives of the study
The following outlined below are the specific objectives of this research work;
1. To extract MnP from Rigidoporus lignosus.
2. To determine the protein content of the enzyme.
3. To assay for the activity of the enzyme using Phenol red.
34
4. To partially purify the enzyme through ammonium sulphate precipitation and gel filtration
(Sephadex G 100).
5. To characterize the partially purified enzyme in respect to pH, temperature and substrate
concentration.
6. To determine the kinetic parameters such as the Km and Vmax of the enzyme catalysed
reaction.
7. To study the properties of MnP inactivation.

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