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Extremophilic microorganisms have developed a variety of physiological strategies that help them to survive on different ecological niche such as extreme temperature, pH, salt concentration and pressure. It has been demonstrated that these microorganisms produce extracellular isoenzyme capable to degrade the ligninocellulosic waste and other related compounds for their growth and survival. These are known as extremophilic ligninolytic enzyme. The extremophilic enzymes are considered superior than normal enzyme because they allow the performance of industrial processes even under adverse condition in which conventional proteins are completely denatured. The common extremophilic ligninolytic enzymes are manganese peroxidase (MnP), lignin peroxidase (LiP) and laccase. These enzymes predominantly have been reported in fungus but their occurrence and role for decolourisation and detoxification of various environmental pollutants also have been reported in bacteria and actinomycetes. Biochemically, MnP and LiP are glycosylated haem protein with molecular weight (MW) ranging from 38 to 62.5 kDa (MnP: 38–62.5 kDa; LiP: 38–46 kDa) while laccases are monomeric, dimeric and trimeric glycoprotein with MW range from 50 to 97 kDa. The optimum activity at pH range for MnP and LiP in fungus is 3.0–5.0 while in bacteria pH range for these enzymes ranges from pH 4.0 to 9.0. The optimum activity for laccase in fungus and bacteria are noted pH 4.0–10.0. The extremophilic activity of these enzymes is regulated due to presence of various salt bridge between amino acids to maintain their stability for catalytic function. Furthermore, the oxidation mechanism of these ligninolytic enzymes have revealed that MnP and lacasse require specific mediator (e.g. GST, tween 80, ABTS, HBT) while LiP does not require any mediator for oxidation of phenolics and non-phenolics compounds. The major biotechnological applications of these enzymes are decolourisation and detoxification of various lignin and ligninolytic waste. It has also scope for pulp biobleaching and ethanol production.
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Chapter 8
Extremophilic Ligninolytic Enzymes
Ram Chandra, Vineet Kumar, and Sheelu Yadav
What Will You Learn from This Chapter?
Extremophilic microorganisms have developed a variety of physiological strategies
that help them to survive on different ecological niche such as extreme temperature,
pH, salt concentration and pressure. It has been demonstrated that these microor-
ganisms produce extracellular isoenzyme capable to degrade the ligninocellulosic
waste and other related compounds for their growth and survival. These are known
as extremophilic ligninolytic enzyme. The extremophilic enzymes are considered
superior than normal enzyme because they allow the performance of industrial
processes even under adverse condition in which conventional proteins are
completely denatured. The common extremophilic ligninolytic enzymes are man-
ganese peroxidase (MnP), lignin peroxidase (LiP) and laccase. These enzymes
predominantly have been reported in fungus but their occurrence and role for
decolourisation and detoxification of various environmental pollutants also have
been reported in bacteria and actinomycetes. Biochemically, MnP and LiP are
glycosylated haem protein with molecular weight (MW) ranging from 38 to
62.5 kDa (MnP: 38–62.5 kDa; LiP: 38–46 kDa) while laccases are monomeric,
dimeric and trimeric glycoprotein with MW range from 50 to 97 kDa. The optimum
activity at pH range for MnP and LiP in fungus is 3.0–5.0 while in bacteria pH range
for these enzymes ranges from pH 4.0 to 9.0. The optimum activity for laccase in
R. Chandra (*)
Environmental Microbiology Division, Indian Institute of Toxicology Research, Post Box
No. 80, Mahatma Gandhi Marg, Lucknow 226001, UP, India
Department of Environmental Microbiology, Babasaheb Bhimrao Ambedkar Central
University, Vidya Vihar, Raebareli Road, Lucknow 226025, UP, India
e-mail: prof.chandrabbau@gmail.com;rc_microitrc@yahoo.co.in
V. Kumar • S. Yadav
Department of Environmental Microbiology, Babasaheb Bhimrao Ambedkar Central
University, Vidya Vihar, Raebareli Road, Lucknow 226025, UP, India
©Springer International Publishing AG 2017
R.K. Sani, R.N. Krishnaraj (eds.), Extremophilic Enzymatic Processing of
Lignocellulosic Feedstocks to Bioenergy, DOI 10.1007/978-3-319-54684-1_8
115
fungus and bacteria are noted pH 4.0–10.0. The extremophilic activity of these
enzymes is regulated due to presence of various salt bridge between amino acids to
maintain their stability for catalytic function. Furthermore, the oxidation mecha-
nism of these ligninolytic enzymes have revealed that MnP and lacasse require
specific mediator (e.g. GST, tween 80, ABTS, HBT) while LiP does not require any
mediator for oxidation of phenolics and non-phenolics compounds. The major
biotechnological applications of these enzymes are decolourisation and detoxifica-
tion of various lignin and ligninolytic waste. It has also scope for pulp biobleaching
and ethanol production.
8.1 Introduction
Generally life exist on the earth in the moderate environment means with neutral
pH, temperature between 22 and 40 C and normal atmospheric pressure (1.0 atm)
with humidity, nutrient and salt condition. However, the deviation beyond moder-
ate environmental condition may create the extreme environment with very low or
high pH, temperature, pressure, salinity, humidity depending upon the geographical
situation also. Thus, for the survival life has been evolved and adapted specialised
mechanism for growth in harsh climate such organisms are known as
extremophiles. There is variable environmental conditions world over for the
microbial growth which different for physiology and biochemical properties for
adaptation.
Low temperature (cold) environment is found in fresh and marine waters, polar
and high alpine soils and water ecosystem. Oceans represent 71% of earths surface
and 90% by volume, which are at 5 C or colder at altitudes >3000 m. As the
altitude increases, the temperature decreases at the rate of 6.4 C per km progres-
sively and even temperatures below –40 C have been recorded. Low temperatures
are characteristic of mountains where snow or ice remains year-round. The tem-
perature is cold, part of the year, on mountains where snow or ice melts. Thus, cold
environment dominates the biosphere. According to Morita (2000), cold environ-
ments can be divided into two categories (1) psychrophilic (permanently cold) and
(2) psychrotrophic (seasonally cold or where temperature fluxes into mesophilic
range) environments. Although habitats with elevated temperatures are not as
widespread as temperate or cold habitats, a variety of high temperature, natural
and man-made habitats exist in environment. These include volcanic and geother-
mal areas with temperatures often greater than boiling, sun-heated litter and soil or
sediments reaching 70 C, and biological self-heated environments such as com-
post, hay, saw dust and coal refuse piles. In thermal springs, the temperature is
above 60 C, and it is kept constant by continual volcanic activity. Besides
temperature, other environmental parameters such as pH, available energy sources,
ionic strength and nutrients influenced the diversity of thermophilic microbial
populations. The best known and well-studied geothermal areas are in North
America (Yellowstone National Park), Iceland, New Zealand, Japan, Italy and the
116 R. Chandra et al.
Soviet Union. Hot water springs are situated throughout the length and breadth of
India, at places with boiling water (e.g. Manikaran, Himachal Pradesh, India).
Geothermal areas are characterized by high or low pH.
Fresh water alkaline hot springs and geysers with neutral/alkaline pH are located
outside the volcanically active zones. Solfatara fields, with sulphur acidic soils,
acidic hot springs and boiling mud pots, characterized other types of geothermal
areas. These fields are located within active volcanic zones that are termed high
temperature fields. Because of elevated temperatures, little liquid water comes out
to the surface, and the hot springs are often associated with steam holes called
fumaroles. With increase in temperature, two major problems are encountered,
keeping water in liquid state and managing the decrease in solubility of oxygen.
Therefore, the microbes growing above boiling point of water have been isolated
from hydrothermal vents, where hydrostatic pressure keeps the water in liquid state;
however, majority of them are anaerobes. Thermal vent sites have recently been
found in Indian Ocean. The primary areas with pH lower than 3.0 are those where
relatively large amounts of sulphur or pyrite are exposed to oxygen. Both sulphur
and pyrite are oxidized abiotically through an exothermic reaction where the former
is oxidized to sulphuric acid, and the ferrous iron in the latter to ferric form. Both
these processes occur abiotically, but are increased 10
6
times through the activity of
acidophiles. Most of the acidic pyrite areas have been created by mining and are
commonly formed around coal, lignite or sulphur mines. All such areas have very
high sulphide concentrations and pH values as low as one (pH 1.0). These are very
low in organic matter, and are quite toxic due to high concentrations of heavy
metals. In all acidic niches, the acidity is mostly due to sulphuric acid. Due to
spontaneous combustion, the refuse piles are self-heating and provide the high-
temperature environment required to sustain thermophiles. The illuminated regions,
such as mining outflows and tailings dams, support phototrophic algae.
In alkaline environments such as soils, increase in pH is due to microbial
ammonification and sulphate reduction, and by water derived from leached silicate
minerals. The pH of these environments fluctuates due to their limited buffering
capacity and therefore, alkalitolerant microbes are more abundant in these habitats
than alkaliphiles. The best studied and most stable alkaline environments are soda
lakes and soda deserts (e.g. East African Rift valley, Indian Sambhar Lake). These
are characterized by the presence of large amounts of Na
2
CO
3
but are significantly
depleted in Mg
2+
and Ca
2+
due to their precipitation as carbonates. The salinity
ranges from 5% (w/v) to saturation (33%). Industrial processes including cement
manufacture, mining, disposal of blast furnace slag, electroplating, food processing
and paper and pulp manufacture produce man-made unstable alkaline environ-
ments. Environments with high hydrostatic pressures are typically found in deep
sea and deep oil or sulphur wells. Almost all barophiles isolated to date have been
recovered from the deep sea below a depth of approximately 2000 m. High-pressure
condition is also met within soils where factors such as high temperature, high
salinity and nutrient limitation may exert further stress on living species. Bacteria
adapted to such an extreme environment are able to grow around or beyond 100 C,
and 200–400 bar of hydrostatic pressure.
8 Extremophilic Ligninolytic Enzymes 117
The majority of extremophiles that have been identified to date belong to the
domain of Archaea. However, many extremophiles from the eubacterial and
eukaryotic domain have also been recently identified and characterised. Recent
study suggested that the diversity of organisms in extreme environments is far
greater than was initially suspected. Extremophiles are an important source of
stable and valuable enzymes present in specific environment. Their enzymes,
sometimes called “extremozymes”. Natural extremozymes have been isolated
from thermophiles, halophiles, psychrophiles, acidophiles and alkaliphiles. But
the structural feature of the enzyme from acidophiles and alkaliphiles is not much
known. Accordingly, biological system and enzyme can even function at temper-
ature between 5 and 130 C, pH 0–12, salt concentration of 3–35% and pressure
up to 1000 bar. In many cases, microbial biocatalyst, especially of extremophiles,
are superior to the traditional catalyst, because they allow the performance of
industrial processes even under extreme conditions where conventional proteins
are completely denatured. Extremophilic enzymes are able to compete for hydra-
tion via alterations especially to their surface through greater surface charges and
increased molecular motion. The unique structural characteristics of the archaeal
polar lipids, that is, the sn-glycerol-1-phosphate (G-1-P) backbone, ether linkages,
and isoprenoid hydrocarbon chains, are in striking contrast to the bacterial charac-
teristics of the sn-glycerol- 3-phosphate (G-3-P) backbone, ester linkages, and fatty
acid chains. The chemical properties and physiological roles of archaeal lipids are
often discuss in terms of the presence of the chemically stable ether bonds in
thermophilic archaea. However, based on the archaeal lipids analyzed thus far, as
shown by lipid component parts analysis, the mesophilic archaea possess essen-
tially the same core lipid composition as that of the thermophilic archaea. The ether
bonds therefore do not seem to be directly related to thermophily. The chemical
stability of lipids and the heat tolerance of thermophilic organisms exhibit because
of the ether bonds of archaeal lipids are for the most part not broken down under
conditions in which ester linkages are completely methanolyzed (5% HCl/MeOH,
100 C for 3 h), it is generally believed that the archaeal ether lipids are
thermotolerant or heat resistant. This implies that thermophilic organisms are able
to grow at high temperature due to the chemical stability of their membrane lipids.
As a matter of fact, all the thermophilic archaea possess ether lipids, but not all of
the organisms possessing the so-called “thermophilic”. The properties have enabled
some extremophilic enzymes to function in the presence of nonaqueous organic
solvents, with this potential properties we can design useful catalysts. Especially
lignocellulolytic, amylolytic, and other biomass processing extremozymes with
unique properties are widely distributed in thermophilic prokaryotes and are of
high potential for versatile industrial processes.
In environment, important extremophilic enzymes have been reported as
ligninolytic enzyme which constitutes manganese peroxidase (MnP), lignin perox-
idase (LiP) and laccases. These enzymes act on broad range of their substrate in
normal to diverse conditions. The demand of these enzymes have increased in
recent year due to their commercial prospect and industrial applications. Such
enzymes have also proven their utility in the pollution abatement, especially in
118 R. Chandra et al.
the treatment of industrial waste/wastewater containing hazardous compound like
phenols, chlorolignin, synthetic dyes, and polyaromatic hydrocarbons (PAHs) as
well as recalcitrant organic compounds structurally similar to lignin. Microorgan-
isms with systems of thermostable enzymes decrease the possibility of microbial
contamination in large scale industrial reactions of prolonged durations. The mech-
anisms for many thermotolerant enzymes have been reported due to their structural
properties i.e. presence of Ca
2+
, saturated fatty acid, α-helical structure etc. There-
fore, the present chapter has been focused on important group of extremophilic
ligninolytic enzymes which have tremendous commercial value for industrial
application but its distribution, mode of action still has to be understood much
more large scale for biotechnological application.
8.2 Manganese Peroxidase
MnP (EC 1.11.1.13) is a ligninolytic extracellular oxidoreductase enzyme belong to
class II fungal haem containing peroxidases produced by almost all wood coloniz-
ing white root and several litter decomposing basidiomycetes during secondary
metabolism in response to nitrogen or carbon starvation. It has also been produce by
some native bacterial strains (Bharagava et al. 2009; Yadav et al. 2011). MnP was
first discovered in the mid-1980s in white-rot fungus Phanerochaete chrysosporium
by two international research teams (M. Golds and R. Crawfords groups) and
characterised as another key oxidative enzyme for lignin degradation (Paszczyn
´ski
et al. 1985). After nearly simultaneous discovery, it has been reported in a large
number of ligninolytic fungi including Phlebia radiata,Pleurotus ostreatus
Bjerkandera adusta,Dichomitus squalens,Trametes versicolor,Lentinus edodes
and so on. The presence of MnP has been also reported in Aspergillus terrus strain
and Penicillium oxalicum isolates-1. MnP is classified in carbohydrate-active
enzymes (CAZy) database in auxiliary activities 113 families. It oxidised Mn
2+
to
Mn
3+
chelate with organic acid and then oxidised various phenolic as well as
non-phenolic compounds including model compounds viz. veratryl alcohol (VA:
3,4-dimethoxybenzyl) and benzyl alcohol. During the degradation process MnP
system generate highly-reactive and non-specific free radicals that cleave carbon–
carbon and ether inter-unit bonds of various phenolics and non-phenolics
compounds. Therefore, a wide range of substrate oxidizing capability renders
it an interesting enzyme for biotechnological applications in several industries.
Potential applications for MnP include biomechanical pulping, pulp biobleaching,
dye decolourisation, bioremediation of some recalcitrant organopollutants (i.e., high
MW cholorolignin, chlorophenols, polycyclic aromatic hydrocarbons, nitroaromatic
compounds) and production of high value chemical from residual lignin from
biorefineries and pulp and paper side stream. MnP has a high potential for penetrating
deep into the soil fines and in nature it catalysed plant lignin depolymerisation. MnP
has also been reported for the degradation/detoxification of triclosan, aflatoxin, nylon,
β-carotene, as well as lignite originated from humic acid.
8 Extremophilic Ligninolytic Enzymes 119
8.2.1 Molecular Structure
MnP is a glycosylated haem protein with MW ranging from 38 to 62.5 kDa, and
averaging at 45 kDa. It occurs as a series of isoforms (isozymes) encoded by a
family of closely related genes and the sequence of cDNA and genomic clones of
three different mnp genes (mnp1,mnp2, and mnp3) from P. crysosporium have been
determined. In P. crysosporium putative metal have been identified upstream of
mnp1 and mnp2 that are also involved in transcription regulation of these genes.
The expression of MnP in nitrogen limited culture of P. crysosporium is regulated
at the level of gene transcription by hydrogen peroxide (H
2
O
2
) and various
chemicals including ethanol, sodium arsenite, and 2,4-dichlorophenol as well as
by Mn
2+
and heat shock. Recently, 11 different isoform of MnP and their genes
have been characterised in Ceriporiopsis subvermispora. The amounts produced
and strengths of these enzymes are different for each type of white rot fungi
resulting in different oxidative activities. However, the regulation of different
MnPs isoform can be largely dependent on the inducing compound (e.g. Mn
2+
,
VA, tween and sodium malonate) and nutrients. Each isoforms of MnP contain
1 mol of iron per mol of protein and differ mostly in their isoelectric points
(pI) which are usually rather acidic (pH 3.0–4.0), through less acidic and neutral
isoforms have found in certain fungi. MnP differs from other peroxidases in the
structure of its substrate binding site. Recent evolutionary studies showed that MnP
evolved from fungal generic peroxidases (similar to plant peroxidases) by devel-
oping a Mn-binding site. MnP then gave rise to versatyl peroxidases (VPs) by
incorporating an exposed catalytic tryptophan and finally to LiPs by loss of the VP
Mn-oxidation site, with the presence of the exposed tryptophan being characteristic
of both LiP and VP crystal structures, and the latter also conserving the above
Mn-binding site.
The crystal structure of MnP (PDB Id: 3m5q) from P. chrysosporium has been
crystallized and subsequently analysed at different refined resolution and this was
the third peroxidase (after cytochrome c peroxidase and LiP) which crystal struc-
ture has been solved (Sundaramoorthy et al. 2010). The structure of MnP consisting
of two domains with heme sandwiched in between. Electronic absorption, electro-
paramagnetic resonance (EPR) and resonance Raman spectral evidence suggested
that the heme iron in native MnP is in high spin, pentacoordinate, ferric state with
histidine coordinated as the fifth ligand. The protein molecule of MnP contains ten
major helices and one minor helix. MnP having five rather than four disulfide bond
present in LiP and VP. The additional disulfide bond Cys341–Cys348 is located
near the C-terminus of the polypeptide chain aids in the formation of Mn
2+
binding
site and responsible for pushing the C-terminus segment away from the main body
of enzyme. The molecular structure of MnP and its Mn
2+
binding site are shown in
Fig. 8.1.
The active site of MnP consist of proximal His173 ligand H-bonded to a
conserved Asp242 residue which contribute to the low negative reduction potential
of the iron, and stabilisation of the oxidation states, compound-I (MnP-I) and
120 R. Chandra et al.
compound-II (MnP-II), and a distal side H
2
O
2
binding pocket consisting as two
conserved amino acid residues, His46 and Arg42. Arg42 implicated in stabilizing
the MnP-I and MnP-II intermediate by forming a hydrogen bond with oxferryl
oxygen. Crystal structure analysis of P. crysosporium MnP showed that Glu35,
Glu39, and Asp179 are forming a Mn
2+
binding site. This site has considerable
flexibility to accommodate the binding of a wide variety of metal ions. The metal
ligands, Glu35 and Glu39, move from their original Mn
2+
binding conformations
and this provides insights into the mechanism of MnP. Further, MnP crystal
structure shows that the Mn
2+
bind side chain of three amino acids, Glu35,
Glu39, Asp179, one heme propionate, as well as two water molecules. The metal
free high-resolution structures shows that Glu35 does not move there original
position in MnP structure, Glu35 and Glu39 to adopt two conformations—“closed”
conformations in the metal bound state and “open” conformations in the metal free
state, possibly acting as a “gate”, enabling a small carboxylic acid like oxalate or
malonate to remove Mn
3+
from the binding site. However, Cd
2+
is a reversible
competitive inhibitor of Mn
2+
and bond to Mn
2+
binding site on MnP, preventing
oxidation of Mn
2+
.
The substrate-bound MnP (Mn–MnP) contains about 357 amino acid residues, three
sugar residues (GlcNac, GlcNac at Asn131 and a single mannose at Ser336), one iron
(III) protoporphyrin IX prosthetic group, two calcium ions, a substrate Mn
2+
ion,
and 478 solvent molecules, including two glycerol molecules. High concentration of
Ca
2+
and Mg
2+
enhance the activity of MnP. However, the substrate-free MnP model
differs only in lacking the Mn
2+
ion in the Mn binding site and in the number of solvent
molecules, 549, which includes two glycerol molecules. A recent survey of over
30 fungal genomes provides evidence that MnP has three subfamilies (long, extralong
Fig. 8.1 Three-dimensional ribbon structure of Phanerochaete chrysosprorium MnP (a) MnP
manganese binding site (b) (Sundaramoorthy et al. 2010, PDB entry 3m5q)
8 Extremophilic Ligninolytic Enzymes 121
and short MnP) defined by the length of the C-terminus tail. The genome of
C. subvermispora is representative of the three MnP subfamilies.
8.2.2 Catalytic Cycle and Mode of Action
MnP catalyzes the oxidation of Mn
2+
to Mn
3+
in the presence of H
2
O
2
(a cosubstrate). Mn
2+
is a specific effector that induces MnP and represses LiP. In
the presence of 1 equiv H
2
O
2
, MnP forms MnP-I, a high valent oxo-Fe
4+
porphyrin
based (Pi) free radical cation (step 1 in Fig. 8.2) which is in turn reduced by a bound
Mn
2+
atom to form MnP-II, an oxo-Fe
4+
porphyrin without the associated porphyrin
(Pi) free radical (step 2 in Fig. 8.2). However, in the absence of Mn
2+
the addition of
2 equiv H
2
O
2
yields MnP-II. The conversion of MnP-I to MnP-II can also be
achieved by addition of other electron donors, such as phenols and amines includ-
ing p-cresol, guiacol, vanillyl alcohol, 4-hydroxy-3-methoxycinnamic acid,
isoeugenol, ascorbic acid, o-dianisidine, ferrocyanide and a variety of phenolic
compounds (Wariishi et al. 1988). MnP-II then oxidizes another Mn
2+
ion, driving
the enzyme back to the ground state Fe
3+
porphyrin (step 3 in Fig 8.2). In the
absence of substrate, the addition of excess H
2
O
2
(250 equiv) drives MnP into
compound-III (MnP-III) which can be further oxidized until bleaching and irre-
versible inactivation (step 4 & 5 in Fig. 8.2).
The Mn
3+
is a strong oxidizer (1.54V) and released from the MnP but it is quite
unstable in aqueous media. To overcome this drawback, white-rot fungi secrete
various organic acids such as oxalate or malate act as chelating agents enabling the
Fig. 8.2 Catalytic cycle of manganese peroxidase
122 R. Chandra et al.
formation of organic acid–Mn
3+
complex. However, the addition of fumarate,
malonate, tartrate, or lactate in the medium enhanced MnP production. The com-
plex formation stabilizes Mn
3+
so that bidentate ligated Mn
3+
usually have redox
potentials of around 0.7–0.9V and significantly lower oxidation capacities when
compared to non-chelated Mn
3+
. The redox potential of chelated Mn
3+
depends on
the chelator. The degradation of recalcitrant non-phenolic compounds has been
limited with MnP generated Mn
3+
chelates alone due to this lower oxidation power,
but in the presence of some mediators or co-oxidants such as glutathione (GSH),
polyoxyethylene sorbitan monoleate (tween 80), acetosyringone, methyl syringate,
3,5-dimethoxy-4-hydroxy-benzonitrile, linoleic acid, linolenic acids, it has effec-
tive in the oxidation of recalcitrant compounds. Mediators are easily oxidizable low
MW compounds that can act as redox intermediates between the active site of the
enzyme and a non-phenolic substrate. Mediators act also as electrons shuttles,
providing the oxidation of recalcitrant complex substrates that do not enter the
active site due to steric hindrances. It is therefore of primary importance to
understand the nature of the reaction mechanism operating in the oxidation of a
substrate by the oxidized mediator species derived from the corresponding mediator
investigated. In the MnP-dependent oxidation of non-phenolic substrates, previous
evidence suggests an electron-transfer (ET) mechanism with mediator syringl type
phenols, towards substrates having a low oxidation potential. Alternatively, a
radical hydrogen atom transfer (HAT) route may operate with ArOH type media-
tors, if weak C–H bonds are present in the substrate. A schematic pathway for the
oxidation of substrate in presence/absences of mediator as shown in Fig. 8.3.
MnP catalyses the oxidation of Mn
2+
to Mn
3+
chelate Mn
3+
to form stable
complexes that diffuses freely and oxidized phenolic substrate (e.g. simple phenol,
amines, dyes, phenolic lignin substructure and dimers) by one electron oxidation of
the substrate, yielding phenoxy radical intermediate, which under undergoes
rearrangement, bond cleavage, and non-enzymatic degradation to yield several
breakdown products. In the presence of Mn
2+
, malonate and H
2
O
2
, MnP from
P. chrysosporium was found to calalyse C
α
–C
β
cleavage, alkyl cleavage and C
α
-
O2
H2O
O2
Substrateoxd
Substrate red
Peroxidase/
Laccase
Oxidised
Peroxidase/
Laccase
Peroxidase/
Laccase
Oxidised
Peroxidase/
Laccase
H2O
Mediator oxd
Mediator red
Substrate oxd
Substrate re
d
(a)
(b)
Fig. 8.3 Schematic representation of peroxidase/laccase-catalyzed redox cycles for oxidation of
substrate in the absence (a) presence (b) of redox mediators
8 Extremophilic Ligninolytic Enzymes 123
oxidation of phenolic arylglycerol β-aryl ether lignin model compounds (Tuor et al.
1992). In the absence of exogenous H
2
O
2
, MnP also has an oxidase activity against
NADPH, GSH, dithiothreitol and dihydroxymaleic acid, forming H
2
O
2
at the
expense of oxygen. The oxidation of phenolic lignin model compound is shown
in Fig. 8.4a.
The Mn
3+
chelate only oxidising phenol portion of the lignin polymer under
physiological conditions and cannot independently oxidised the non-phenolic parts.
Therefore, alternative mechanism whereby MnP could oxidise non-phenolic com-
pounds have been sought. For oxidation of non-phenolic compounds by Mn
3+
involves the formation of reactive radical in the presence of mediators. It was
found that in the presence of glutathione (GSH) MnP could oxidize non-phenolic
lignin model compounds, veratryl, anisyl, and benzyl alcohol (Wariishi et al. 1989).
They demonstrated that the Mn
3+
formed oxidise thiol to thiyl radical that in turn
abstracts a hydrogen from substrate (veratryl, anisyl, and benzyl alcohol) to
forming a benzyl radical which react with another thiyl radical to yield an inter-
mediate which decomposed to the benzaldehyde products (veratraldehyde,
anisaldehyde and benzaldehyde).
Bao et al. (1994) reported that MnP from P. chrysosporium could oxidise a
non-phenolic β-O-4 lignin model compounds in the presence of tween 80, an
anionic surfactant made from an unsaturated fatty acid, oleic acid. They suggested
that MnP oxidised the carbon–carbon double bond (C¼C) in tween 80 to a peroxide
which is known as lipid peroxidation and subsequently turned into a peroxy radical.
As a result, the MnP-lipid system catalyse C
α
–C
β
cleavage, and β-aryl ether
cleavage of non-phenolic diarylpropane and β-O-4 lignin, respectively. In the
MnP-dependent peroxidation of unsaturated fatty acid, lipid free radicals also
produce superoxide radicals O
2.
The substrate oxidation mechanism involve benzyl
hydrogen abstraction from benzyl carbon (C
α
) via lipid peroxy radical followed by
O
2
addition to form peroxy radical, and subsequent oxidative cleavage and
non-enzymatic degradation as shown in Fig 8.4b. It was suggested that this process
might enable the white rot fungi to accomplish the initial delignification of wood.
Moreover, MnP has been reported to oxidised various halogenated compounds
by the oxidation mechanism is different to described above. Halide ions are
oxidized by a unique peroxidase mechanism, compared with other electron donors.
Most electron donors are oxidized by MnP-I via a single- electron mechanism with
the intermediate formation of MnP-II (see Fig. 8.2) whereas halides are oxidized by
MnP-1via a two-electron mechanism, yielding the native enzyme directly. A novel
MnP from P. chrysosporium exhibits haloperoxidase activity at low pH. In the
presence of H
2
O
2
MnP oxidizes bromide and iodide to tribromide and triiodide at
optimum pH 2.5 and 3.0, respectively.
124 R. Chandra et al.
8.2.3 Common Substrate and Microorganisms
Microorganisms have the ability to interact, both chemically and physically, with
substances leading to structural changes or complete degradation of the target
molecule. A huge number of MnP producing fungi and bacteria genera possess
the capability to degrade various organic substrates as a sole carbon, nitrogen and,
phosphorus for their growth and metabolism in natural and controlled environment.
A range of MnP producing fungi and bacteria and their substrate are listed in
Table 8.1.
Fig. 8.4 Manganese peroxidase catalysed oxidation mechanism of phenolic aryglycerol β-aryl
ether and (a) non-phenolic β-O-4 lignin model compound (b) [modified from Wong (2009), Tuor
et al. (1992) and Bao et al. (1994)]
8 Extremophilic Ligninolytic Enzymes 125
Table 8.1 Some important MnP producing microorganisms in extremophilic conditions and their
substrate
Microorganisms Substrate pH Mediator Temp.
Fungi
Irpex Lacteus CD2 Remazol brilliant violet 5R, direct red 5B,
remazol brilliant blue R, indigo camine,
methyl green
3.5–6.0 – 40–60
Phanerochaete
sordita YK-624
Reactive red 120, bleaching of hard wood
craft pulp
4.5 Tween
80
30
Bjerkandera
sp. BOS55
Orange II 4.5 20–30
Phanerochete
Chrysosporium
BKM-F-1767
Poly R-478 4.5
Coriolus hirsutus Melanoidins 4.5 –
Lactobacillus kefir Sucrose-glutamic acid, sucrose- aspartic
acid, glucose-glutamic acid
7.2–7.4 – 30
Phaneochete
sordida YK-624
Aflatoxin B1 4.5 Tween
80
30
Nematoloma
forwardii
[U-
14
C]pentachlorophenol, [U-
14
C] cate-
chol, [U-
14
C] tyrosine, [U-
14
C] trypto-
phan, [4,5,9,10-
14
C] pyrene, [U-
14
C]2
amino-4,6-dinitrotoluene
[
14
C] pyrene, [
14
C] anthracene, [
14
C]
benzo(a)pyrene, [
14
C]benz(a)pyrene, [
14
C]
phenantherene, [
14
C]- synthetic lignin
(DHP)
4.5 – 30
Phanerochete
crysosporium
Non-phenolic β-vanillyl alcohol;
nonphenolic β-vanillyl dimer;
Nonphenolic β-1 diarylpropane lignin
model dimers
4.5 GSH
Tween
80
30
Anthracophyllum
discolor
Pyrene, anthracene, fluranthene,
phenanthrene)
4.5 – 25
Ganoderma
lucidum
Crescent, magna, textile effluent 4.5 25
Pleurotus
ostreatus
2,2 -Bis- (4-hydroxyphenyl) propane
(Bisphenol A)
4.5 – 25
Clitocybula
dusenii b11
Lignite originated humic acid 4.0 GSH 37
White rot fungi
IZU-154
Nylon-66
Schizophyllum
sp. F17
Congo red, orange G, orange IV 4.0–7.4 25
Trichophyton
rubrum LSK-27
Plant derived lignin 4.5 30 or
40 C
Penicillium
oxalicum isolate 1
Plant derived lignin 4.5 37
Ceriporiopsis
subvermispora
Plant derived lignin 2.0–5.0 25
(continued)
126 R. Chandra et al.
8.2.4 Screening of MnP Producing Microorganisms Its
Substrate, Bioassay and Purification
MnP oxidised a wide range of phenolic and non-phenolic compounds as a common
substrate in the presence of H
2
O
2
. There are various phenolic compounds
(e.g. 2,2-azinobis (3-ethylthiazoline-6-sulfonate (ABTS), 2,6-dimethyloxyphenol
(DMP), vanillylacetone, ferulic acid (4-hydroxy-3-methoxycinnamic acid),
syringol, guaiacol, isoeugenol, p-methoxyphenol, syringaldazine,
divanillylacetone, phenol red and coniferyl alcohol, [3-methyl-2-benzothiazolinone
hydrazone (MBTH)], 3-(dimethylamino) benzoic acid (DMAB), p-cresol,
o-dianisidine, catechol, hydroquinone) and non-phenol compound (e.g. vanillyl
alcohol, VA and benzyl alcohol) in the presence used for in vitro MnP assay. The
most commonly used substrate for MnP assay is guaiacol, a natural phenolic
product first isolated from guaiac resin and the oxidation of lignin.
The isolated and purified microorganisms are screened for MnP enzyme activity
by plate assay method using phenol red or guaiacol as substrate in minimal salt
media agar plate where the screening of MnP is based on as colorless halo
formation around the microbial growth. The assay plate containing composition
(g/l) 3.0 peptone, 10.0 D-glucose, 0.6 KH
2
PO
4
, 0.001 K
2
HPO
4
, 0.4 ZnSO
4
, 0.0005
FeSO
4
, 0.05MnSO
4
, 0.5 MgSO
4
,H
2
O
2
, and 20.0 agar supplemented with 0.2%
guaiacol. After incubation, MnP activity are visualised on plate by formation of
reddish brown zone around the microbial growth due to the oxidative polymeriza-
tion of guaiacol as shown in Fig 8.5a. While in another method MnP producing
Table 8.1 (continued)
Microorganisms Substrate pH Mediator Temp.
White rot fungi
IZU-154
Polyethylene Tween
80
Bacteria
Bacillus
sp. IITRM7
Sucrose aspartic acid maillard product 7.0 35
Raoultella
planticola
IITRM15
Sucrose aspartic acid maillard product 7.0 35
Enterobacter
sakazakii
IITRM16
Sucrose aspartic acid maillard product 7.0 35
Bacillus
licheniformis
(RNBS1)
Melanoidins 7.3 – 35
Bacillus
sp. (RNBS3)
Melanoidins 7.3 – 35
Alcaligenes
sp. (RNBS4)
Melanoidins 7.3 – 35
All values of temperature (Temp.) are given in C
8 Extremophilic Ligninolytic Enzymes 127
bacteria has been screened on modified GPYM as well as in broth amended with
different concentration of sucrose aspartic acid-maillar product (SAA-MP) using
method describe by Yadav et al. (2011). In this method, MnP producing microbial
culture are inoculated on GPYM agar plates containing D-glucose, 1.0; peptone,
0.1; K
2
HPO
4
, 0.1 and MgSO
4
7H
2
O, 0.05 with different concentration of SAA-MP
(800–3600 mg/l) supplemented with 0.1% phenol red (w/v). After incubation the
potential bacteria developed changing the deep orange colour of phenol to light
yellow as shown in Fig 8.5b. Chandra and Singh (2012) studied the production of
ligninolytic enzyme during pulp paper mill effluent degradation by bacterial con-
sortium (Fig. 8.5c). They found that MnP activity are lower comparison to laccase
but higher than to LiP as shown in Fig. 8.5d.
The bioassay of MnP activity is carried out by measuring optical density (OD) at
270 nm (ε¼11.59 mM
1
cm
1
) due to the formation of Mn
3+
-malonate complex at
4.5 as shown in Fig. 8.6. This method is based on oxidation MnSO
4
. The assay
mixtures containing 50 mM sodium malonate buffer (pH 4.5), 0.5 mM MnSO
4
,
0.1 mM H
2
O
2
and 100 μl of enzyme solution. The reaction is initiated by adding
H
2
O
2
at 25 C and Mn
3+
-malonate complex measured spectrophotometrically at
270 nm. The MnP activity has also been measured by using ABTS, DMP or VA as
the substrate under the conditions as described above and the oxidation has been
followed by monitoring optical density at 414 nm (ε¼36 mM
1
cm
1
) for ABTS
or at 470 nm (ε¼49.6 mM
1
cm
1
) for DMP or at 310 nm (ε¼9.3 mM
1
cm
1
)
for VA, respectively. The MnP activity can also been determined spectrophoto-
metrically by using phenol red as substrate at 610 nm. This activity assay is based
on the oxidation of phenol red in the presence of H
2
O
2
and Mn
2+
and the oxidation
product is measured by spectrophotometrically. In this method, a reaction mixture
containing enzyme extract (700 μl), 0.2% phenol red, 2 mM sodium lactate (50 μl),
2.0 mM MnSO
4
, 0.1% egg albumin, 2 mM H
2
O
2
in 20 mM sodium succinate buffer
at 4.5 pH. The oxidation product is measure by spectrophotometer recording the
absorbance at 610 nm (ε¼22 mM
1
cm
1
). However, the manganese independent
peroxidase activity is determined using 2.0 mM EDTA instead of MnSO4 solution.
A another very sensitive bioassay method for measurement of MnP activity by
using 3MBTH)/DMAB as a substrate has also been developed. In this method, the
reaction mixture contained 0.07 mM MBTH, 0.99 mM DMAB, 0.3 mM MnSO
4
,
0.05 mM H
2
O
2
and the sample in a 100 mM succinic/lactic acid buffer (pH 4.5).
7
6
5
4
3
2
0
1
0244872
Incubation time(h)
Enzyme activity (IU/ml)
96 144 168192 216
Lac.
MnP
LiP
120
ab cd
Fig. 8.5 Plate Showing growth of MnP producing microorganisms by using different substrate (a)
guaiacol (b) phenol red (c) light microscopy (d) ligninolytic enzyme activity
128 R. Chandra et al.
This reaction mixture yields a deep purple color with a broad absorption band with a
peak at 590 nm (ε¼53.0 mM
–1
cm
–1
). All activities are expressed in international
unit (IU). IU defined as the amount of enzyme produced 1μmol of product per
minute under the assay condition used.
Various enzyme/protein purification techniques are frequently employed puri-
fying MnP from microbial culture. MnP has been purified from Lentinula edodes by
applying the crude extract on cold acetone (20 C) and Sephadex G-100 column,
respectively and characterized by denaturing sodium dodecyl sulfate-
polyacrylamide gel electrophoresis (SDS-PAGE). They obtained the terminal spe-
cific activity of 5496 U mg
1
with a 6.76 fold. Further, MnP has also been isolated
and purified from Irpex lacteus CD2 by using diethylaminoethyl cellulose (DEAE)
sepharose column and characterised by SDS-PAGE using 10% polyacrylamide gel.
They obtained MnP a terminal specific activity of 24.9 U/mg protein with 29.3-fold.
In this method, the liquid culture of microorganisms first centrifuged at 5000gfor
20 min. Then the culture supernatant has concentrated by 80% ammonium sulfate at
4C . The pellets have dissolved in sodium acetate buffer (20 mM, pH 4.8) then the
enzymatic crude extract has dialyzed against the same buffer to remove ammonium
sulfate and then applied to a DEAE sepharose column previously equilibrated with
sodium acetate buffer (20 mM, pH 4.8). The MnP has eluted with a linear gradient
of 0–1 M NaCl in the same buffer at a flow rate of 1 ml/min. The proteins in the
eluted fractions are detected by recording the absorbance at 280 nm as shown in Fig
8.6a. Further, active fractions containing MnP activity is recorded after gel docu-
mentation of native-PAGE as shown in Fig 8.6b. The purified MnP has been
verified by SDS-PAGE using 10% polyacrylamide gel. The MW of the purified
MnP has been estimated in comparison to standard MW marker.
Fig. 8.6 UV-Visible spectrum of purified native MnP enzyme (dashed line)(a) Native PAGE for
MnP (b) symbol S: Molecular weight standard marker; L-I crude MnP extract; L-2 partially
purified MnP after ammonium sulfate precipitation; L-3 Gel filtration chromatography of purified
MnP
8 Extremophilic Ligninolytic Enzymes 129
8.2.5 Effect of Environmental Parameters, Organic Solvent
and Heavy Metal on MnP Activity
The ability of MnP to tolerate high temperature, different metal ions and organic
solvents is very important for the efficient application of this enzyme in the
biodegradation and detoxification of industrial waste. The activity and stability of
MnP is strongly influenced by the pH, temperature, and time of incubation. The
effect of temperature and buffer type on the stability of MnP has been previously
investigated by various workers. Sutherland and Aust (1996) found that MnP from
P. crysosporium was most stable at pH 5.5 and temperature at of below 37 C. They
also found that MnP become inactive at high temperature due to loss of Ca
2+
,
required for the stability and activity. The engineering of a disulfide bond (A48C
and A63C) near the distal calcium binding site of MnP by double mutation showed
the improvement in thermal stability as well as pH (pH 8.0) stability in comparison
to native enzyme (MnP). The disulfide bond adjacent to the distal calcium ligand
Asp47 and Asp64 stabilizes the recombinantly expresses MnP against the loss of
calcium. The MnPs (G1 and G2) from Ganoderma sp. YK-505 exhibited 100%
MnP activity after treatment for 60 min at 60 C, but lower than 20% residual
activity with 10 mM H
2
O
2
. Recently, a thermostable and H
2
O
2
tolerant MnP
isolated and purified from the culture medium of Lenzites betulinus named as
L-MnP. The purified L-MnP has the highest H
2
O
2
tolerance among MnPs reported
so far. It retained more than 60% of the initial activity after thermal treatment at
60 C for 60 min, and also retained more than 60% of the initial activity after
exposure to 10 mM H
2
O
2
for 5 min at 37 C. Now a novel MnP (CD2-MnP) has
been purified and characterised from the white-rot fungus I. lacteus CD2. The CD2-
MnP has strong capability for tolerating different metal ions such as Ca
2+
,Cd
2+
,Co
2+
,
Mg
2+
,Ni
2+
and Zn
2+
as well as organic solvents such as methanol, ethanol, DMSO,
ethylene glycol, isopropyl alcohol, butanediol and glycerin. CD2-MnP exhibited
high stability in pH range from 3.5 to 6.0 and optimal temperature was determined
to be 70 C. All these purified MnP are known as wild MnP (wMnP). wMnP are not
well suited for industrial used, which often required particular substrate specificities
and application conditions (including pH, temperature and reaction media) in
addition to high production levels. Thermostable enzymes are typically tolerant to
many other harsh conditions often required in industry, such as the presence of
organic co-solvents, extreme pH, high salt concentrations, high pressures, etc.
Therefore, the development of recombinant MnP (rMnP) for industrial application
through protein engineering and heterologous expression is in process.
130 R. Chandra et al.
8.3 Lignin Peroxidase
LiP (EC 1.11.1.14) is an extracellular H
2
O
2
dependent heme containing glycopro-
tein, produced by white-rot fungi and similar to the lignin-synthesizing plant
peroxidases. It was first discovered in nitrogen and carbon limited cultures of
P. chrysosporium and since then has become one of the most studied peroxidases
(Glenn et al. 1983). It has also been reported to be produced by many white rot fungi
including Phlebia flavido-alba,Bjerkandera sp. strain BOS55, T. trogii,Phlebia
tremellosa and P. chraceofulva. Several LiP isozymes have also been detected in
cultures of P. chrysosporium,T. versicolor,B. adusta and Phlebia radiate and so
one. LiP possess high redox potential (700–1400 mV), low optimum pH 3.0 to 4.5,
ability to catalyze the degradation of a wide number of aromatic substrates such
VA, methoxybenzenes and also a variety of non-phenolic lignin model compounds
as well as a range of organic compounds with a redox potential up to 1.4 V (versus
normal hydrogen electrode) in the presence of H
2
O
2
.
LiPs can catalyse the oxidative cleavage of Cα–Cβlinkages, β-O-4 linkages, and
other bonds present in lignin and its model compounds. The enzyme also catalyzes
side-chain cleavages, benzyl alcohol oxidations, demethoxylation, ring-opening
reactions and oxidative de chlorination. Moreover, bacteria are worthy of being
studied for their ligninolytic potential due to their immense environmental adapt-
ability and biochemical versatility. There is wide range of examples where bacteria
like Pseudomonas aeruginosa,Serretia marcescens,Nocardia,Arthrobacter,
Flavobacterium,Micrococcus,Xanthomonas sp. have been identified as
lignocellulosic-degrading microorganisms. Therefore, identification of bacteria
having lignin oxidizing enzymes would be of significant importance. The LiP
activity is associated with primary growth of bacteria and thus the delignification
process is presumed to be the result of primary metabolic activity and not dependent
upon other factors such as stress to induce production.
8.3.1 Molecular Structure
LiP a monomeric glycoprotein of 38–46 kDa (and pI of 3.2–4.0) containing 1 mol
of iron protoporphyrin IX per 1 mol of protein, catalyzes the H
2
O
2
-dependent
oxidative depolymerization of lignin. LiP has a distinctive property of an unusually
low pH optimum near pH 3.0 in extremophilic environment. In general, LiP which
has MW 40 kDa contains 343 amino acids residues, 370 water molecules, a heme
group, four carbohydrates, and two calcium ions. The crystal structure of LiP (PDB
Id: 1LGA) isolated and purified from P. chrysosporium as shown in Fig. 8.7a. The
heme is embedded in a crevice between the two domains, but is accessible from the
solvent via two small channels. The secondary structure of enzyme molecule
contains eight major and eight minor α-helices and two anti-parallel β-sheet, and
it is organised in a proximal and a distal domain. The heme is embedded in a crevice
8 Extremophilic Ligninolytic Enzymes 131
between the domains: two small channels connect the prosthetic group to the
solvent. The LiP contains eight Cys residues, all forming four disulfide bridges.
There are two calcium-binding sites, one in each domain, with possible function of
maintaining the topology of the active site. The heme is embedded in a crevice
between the domains: two small channels connect the prosthetic group to the
solvent. The heme iron is predominantly high spin, pentacoordinated with
His176-N at the proximal side as the fifth ligand, and Wat339 H-bonded to the
distal His47-N (Fig. 8.7b). The His is associated with the high redox potential of
LiP. The enzymes redox potential rises when the His has a reduced imidazol
character. In addition, a greater distance between the His and the heminic group
increases the redox potential of the enzyme. Another characteristic related with
LiPs high redox potential is the invariant presence of a tryptophan residue
(Trp171) in the enzymes surface. Trp171 seems to facilitate electronic transference
to the enzyme from substrates that cannot access into the heminic oxidative group.
This Trp171 residue has been suggested an important role in the binding and
oxidation of VA, a fungal secondary metabolite produced by at the same time as
LiP and oxidised by LiP. VA participates in the oxidation of different aromatic
molecules.
8.3.2 Catalytic Cycle and Mode of Action
The catalytic cycle of LiP is similar to that of other peroxidases like MnP where
ferric enzyme is first oxidized by H
2
O
2
to generate the two-electron oxidized
intermediate, compound-I (LiP-I). In this reaction 2-electron oxidation of ferric [Fe
3+
]
Fig. 8.7 Three-dimensional ribbon structure of P. chrysosporium lignin peroxidase (a) detail of
the heme environment (b) (Poulos et al. 1993; PDB entry 1LGA)
132 R. Chandra et al.
LiP produces LiP-I intermediate, a oxoferryl iron porphyrin radical cation [Fe
4+
¼O]
with the reduction of H
2
O
2
. Next, LiP-I is reduced by one electron donated by a
substrate such as VA, yielding the 1-electron oxidized enzyme intermediate,
compound-II [Fe
4+
¼O] (LiP-II), and a free radical product (VA
•+
). VA
•+
acts as a
redox mediator in the oxidation of lignin. VA
•+
is capable of mediating oxidation of
secondary substrates typically not oxidized by LiP. The catalytic cycle is completed
by the one-electron reduction of LiP-II by a second substrate molecule. But, in the
absence of a reducing substrate, the enzyme can undergo a series of reactions with
H
2
O
2
to form compound-III (LiP-III), oxyperoxidase. LiP-III is stable, but
prolonged incubation of enzyme with H
2
O
2
in the absence of a reducing substrate
such as VA can cause irreversible inactivation of the enzyme. In the presence of
VA, however, LiP-II undergoes multiple turnovers without any detectable inacti-
vation. Because VA is normally produced by white rot fungus, some workers
conceptualized that VA protects the enzyme from the action of H
2
O
2
- dependent
inactivation and participate as a redox mediator between the enzyme and substrates
which cannot get inside the heminic center. Moreover, VA
•+
converts LiP-III to the
native enzyme via the formation of veratryaldehyde and H
2
O, potentially making
more enzymes active for the oxidation of lignin. The catalytic cycle of LiP as shown
in Fig. 8.8.
Some studies have observed that substrates are not oxidized by LiP such as
anisyl alcohol and 4-methoxymandelic acid they are oxidized in the presence of
VA. They proposed that the one-electron oxidized product of VA, the aryl cation
Fe
3+ Fe 4+
Fe 4+
LiP-II
(Compound- II)
Fe 3+
LiP-III
(Compound- III)
H2O2H2O
LiP-I
(Compound- I)
VA
VA+
H2O
H2O2
Lignin Peroxidase
(LiP)
VA+
VA
VA+
H2O
+
2 VAD
VA- Veratryl alcohol
VA- Veratryl free radical
VAD -Veratrylaldehyde
VA+-Veratryl radical cation
O
O
O2
+
Fig. 8.8 Catalytic cycles for lignin peroxidase
8 Extremophilic Ligninolytic Enzymes 133
radical, is able to mediate the oxidation of substrates typically not oxidized by the
enzyme. The aryl cation radical is a diffusible species, capable of acting at a
distance. In another studies concluded that the stimulation of 4-methoxymandelic
acid and anisyl alcohol oxidation is due solely to the ability of VA to prevent
inactivation of lignin peroxidase. They claimed that enzyme in the presence of
anisyl alcohol and excess H
2
O
2
leads to the formation of inactive LiP-III. LiP can
be inhibited by cyanide and chloride. Chloride is expected to be a competitive
inhibitor because it is demonstrated that chloride is not a substrate for LiP.
Like MnP, LiP is capable of oxidizing a wide variety of phenolic compounds
including ring- and N-substituted anilines. It oxidise a wide range of aromatic
compounds (guaiacol, vanillyl alcohol, catechol, syringic acid, acetosyringone,
etc.) preferentially at a much faster rate compared to non-phenolic substrates.
Using a lignin model dimer as the substrate, the cation radical decays with the
spontaneous C
α
–C
β
fission of the alkyl side chain, with the products resembling
those found when fungi degrade lignin. In the reduction of LiP-I and LiP-II,
phenolic substrates are converted to phenoxy radicals. In the presence of oxygen,
the phenoxy radical may react to form ring-cleavage products, or they may other-
wise also lead to coupling and polymerization. It has been reported that
LiP-catalyzed oxidation of the lignin model dimer compound 1,2-di (3,4-
methoxyphenyl)-1,3-propanediol results in C
α
–C
β
cleavage to yield veratraldehyde.
LiP-catalyzed oxidative reaction of phenolic compounds is typically associated
with rapid decrease in enzyme activity. The decrease is likely caused by the
accumulation of the inactive LiP-III during catalysis. Phenoxy radicals, unlike the
non-phenolic VA discussed below, are unable to revert LiP-III to the native
enzyme, although both substrates show similar rate constants for the reaction of
LiP-I. The reactivity of phenolic compounds with LiP-I is much higher than that
with LiP-II, and the rate constant decreases as the size of the substrate increases as
demonstrated in the oxidation of oligomers of phenolic β-O-4 lignin model
compounds.
LiP shows a very high redox potential (1.2 V at pH 3.0) compared to laccases
(~0.8 V at pH 5.5), horseradish peroxidases (0.95 V at pH 6.3) and MnP (0.8 V at
pH 4.5). This property enables LiP to catalyze the oxidation of non-phenolic
aromatic compounds, even in the absence of a mediator. The oxidative reaction
of non-phenolic diarylpropane and β-O-4 lignin model compounds of lignin
involves initial formation of radical cation via 1eoxidation, followed by side-
chain cleavage, demethylation, intramolecular addition, and rearrangements. Oxi-
dation of the A ring giving rise to C
α
–C
β
cleavage is the major route. In the
mechanism, only the formation of the radical cation is enzyme catalyzed, and
subsequent reactions of the substrate are nonenzymatic. Decay of the radical cation
depends on the nature of the substituents on the aromatic ring. Electron-donating
groups, such as alkoxy groups, on the aromatic ring favor the formation and
stabilization of the aryl radical cation. LiP also oxidised the VA, a metabolic
product at the same time as LiP produced by P. chrysosporium. The addition of
VA is known to cause an increase in LiP activity and the rate of lignin minerali-
zation. At pH 6.0, the second-order rate constant for LiP-catalyzed oxidation of VA
134 R. Chandra et al.
is similar to that of β-O-4 dimer (6.7 10
3
and 6.5 10
3
M
1
s
1
, respectively).
The radical cation formed in the first reduction by LiP-I exist as a complex with
LiP-II, which is catalytically active on a second VA molecule to form an aldehyde.
The VA
•+
generated in the reduction step decays by deprotonation at Ca, a typical
reaction of alky aromatic radical cations, to form veratraldehyde as shown in
Fig. 8.9. Under aerobic condition, however, additional oxidative pathways involv-
ing activated oxygen species occur leading to quinone formation and aromatic ring
cleavage.
8.3.3 Common Substrate and Microorganisms
The association of ligninolytic enzymes with lignin breakdown arises because the
enzyme can oxidize lignin-related aromatic compounds. Single-ring aromatic sub-
strates are frequently used, including phenolic compounds i.e. guaiacol, vanillic
acid, and syringic acid, and non-phenolic VA, and dimethylphenylenediamine. This
type of substrates have been useful for characterization of catalytic cycles, partic-
ularly on the formation and reactions of the oxidized enzyme intermediates.
Another group of substrates consists of lignin model dimers that are frequently
used to investigate specific bond cleavages. These compounds are synthetic mimics
of the common lignin substructures, such as diarylpropane and β-aryl ether dimer.
The β-O-4 lignin model compounds are the most important type for elucidating
lignin degradation, as arylglycerol β-aryl ether or β-O-4 bond is the most prevalent
linkage type in lignin, accounting for about 50% of the interunit connections in
gymnosperm and 60% in angiosperm. These model compounds can be synthesized
either as phenolic or non-phenolic in nature. Phenolic subunits are present only
about 10–20% in lignin. However, demethylation and ether cleavage reactions in
enzyme-catalyzed degradation of phenolic compounds generate phenolic products,
Fig. 8.9 Degradation mechanism of veratryl alcohol by lignin peroxidase
8 Extremophilic Ligninolytic Enzymes 135
which can in turn be the substrate for further breakdown. LiP producing microor-
ganisms and their substrate are listed in Table 8.2.
8.3.4 Screening of LiP Producing Microorganisms, Its
Substrate, Bioassay and Purification
The isolated and purified bacterial strain has been screened for LiP activity by plate
assay method. The plate assay is generally performed using different substrate such as
azure B (0.002%) or methylene blue (0.025%) in B & K agar medium plates
containing dextrose 1%, peptone 0.5%, NaCl 0.5%, beef extract 0.3% and CuSO
4
(1 mM) (Chandra and Singh 2012). The disappearance of blue colour of the media
confirmed the presence of LiP activity around the bacterial growth as shown in
Fig. 8.10a (Chandra and Singh 2012). Moreover, LiP activity is determined spectro-
photometrically during degradation of organic pollutants by recording the increase in
absorbance at 310 nm through the oxidation of VA to veratryl aldehyde (E
310
¼9300
M
1
cm
1
). The reaction mixture contained 100 mM sodium tartrate pH 3.0, 2 mM
Table 8.2 Different LiP producing microorganisms in extremophilic environment and their
substrate
Microorganisms Substrate pH Temp. (C)
Fungi
Phanerochaete
chrysosporium
Azo dyes, penta chlorophenol, veratryl alcohol,
anthracenes
339
Phanerochaete flavido-
alba
Synthetic dehydropolymerized lignins (DHPs),
veratryl alcohol
3–4 30
Bjerkandera sp. strain
BOS55
2-chloro-1,4-dimethoxybenzene, veratryl
alcohol
330
Trametes trogii Lignin, veratryl alcohol 3 30
Phlebia ochraceofulva Lignin, dimethyl succinate, veratryl alcohol 3 30
Phlebia tremellosa Lignin, dimethyl succinate, veratryl alcohol 3 30
Bacteria
Pseudomonas sp. SUK1 n-propanol 3 30
Pseudomonas
aeruginosa
Lignin, bromophenol blue, veratryl alcohol
Bacillus megaterium Lignin, veratryl alcohol 7 37
Serretia marcescens Lignin, bromophenol blue and veratryl alcohol 4 30
Bacillus subtilis Alkaline lignin, veratryl alcohol 6 37
Arthrobacter
globiformis
Lignin, veratryl alcohol 7 37
Actinomycetes
Streptomyces
viridosporus T7A
Vanillic acid, syringic acid 6 35
Streptomyces sp. AD001 2,4 dichlorophenol, 4-aminoantipyrene 7 35
136 R. Chandra et al.
VA, LiP, and the reaction mixture was incubated at initiated by the addition of 0.5 ml
H
2
O
2
(final concentration 0.5 mM). The formation of the predominant product
2-chloro-1,4-benzoquinone from 2-chloro-1,4-dimethoxybenzene is measured at a
wavelength of 255 nm using a molar extinction coefficient of 16,900 M
1
cm
1
(ten
Have et al. 1998).
The LiP activity can also been determined spectrophotometrically by monitoring
the oxidation of azure B as substrate in presence of H
2
O
2
at 610 nm. The reaction
mixture contained sodium tartrate buffer (50 mM, pH 3.0), azure B (32 μM), 500 μl
of culture filtrate 500 μlofH
2
O
2
(2 μM). OD is taken at 651 nm after 10 min. One
IU of LiP activity is defined as activity of an enzyme that catalyzes the conversion
of μmole of substrate per minute. The purification of LiP enzyme has been reported
by using the method described by Yadav et al. (2009). In this method, the partial
purification of enzyme separated from exhausted medium is usually done by 70%
ammonium sulphate saturation. The mixture is then stored in a cold room for 24 h to
precipitate all the proteins and the precipitation is separated by centrifugation for
10 min. The supernatant is discarded and the remaining precipitate is further
dissolved with 5 ml of 1M citrate phosphate buffer (pH 8.0) the concentrated
enzyme mixture is subjected to dialysis. Further, the dialyzed enzyme is loaded
onto a DEAE-cellulose column of size 1–16 cm, which is pre-equilibrated with the
same phosphate buffer. The adsorbed enzyme is washed with 50 mL of the same
buffer and is eluted by applying a linear gradient of NaCl (0–200 mM; 50 mL buffer
150 mL buffer containing 200 mM NaCl).The elution profile of the LiP activity
from the DEAE cellulose column is given in Fig. 8.10b. The purpose of dialysis is
to remove undesired small molecular weight molecules from a mixture in which the
desired species of molecules are too large to travel across the membrane. Ordinarily
this process is utilized during protein purification in which salting out procedure has
Fig. 8.10 (a) Plate assay of lignin peroxidase (b) elution profile from a DEAE column ( filled
triangle) activity profile ( filled circle) protein at 750 nm; (straight line) NaCl gradient. Fractions
of 5 mL were collected. (c) SDS-PAGE analysis of purified lignin peroxidase. Lane 1 contains the
MW markers (from top): phosphorylase (97.4 kDa), bovine serum albumin (68 kDa), ovalbumin
(43 kDa), carbonic anhydrase (29 kDa), soyabean trypsin inhibitor (20.1 kDa) and lysozyme (14.3
kDa). Lane 2 contains the purified lignin peroxidase. 50 mL was loaded (Yadav et al. 2009)
8 Extremophilic Ligninolytic Enzymes 137
been employed as the initial step with ammonium sulphate. After the protein is
precipitated from the initial source, it is re-dissolved in buffer and then poured into
a dialysis bag. The homogeneity of the enzyme preparation is checked by SDS-
PAGE. The separating gel contains 12% acrylamide in 0.375M Tris-HCl buffer
(pH 8.8) and the stacking gel remains 5% acrylamide in 0.063M Tris-HCl buffer
(pH 6.8). Proteins are visualized by silver staining as shown in Fig 8.10c.
8.4 Laccase
Laccases (EC 1.10.3.2) are multi copper-containing polyphenol oxidases that are
widely distributed in microorganisms, insects, and plants, showing a specific
function in each of them. From this group, white rot fungi are the most studied
laccases. It catalyze the oxidation of various aromatic compounds, particularly
those with electron-donating groups such as phenols (OH) and anilines (NH
2
),
by using molecular oxygen as an electron acceptor. In nineteenth century, laccases
was first isolated from exudates of the Japanese tree Rhus vernicifera. Laccases use
molecular oxygen to oxidize a variety of aromatic and non-aromatic hydrogen
donors via a mechanism involving radicals. These radicals can undergo further
laccases catalyzed reaction and/or non-enzymatic reaction such as polymerization,
and hydrogen abstraction. Therefore, laccase has also the ability to oxidize phenolic
and non-phenolic substrates. The phenolic substrate oxidation by laccases result in
formation of an aryloxyradicals an active species that is converted to a quinone in
the second stage of the oxidation. Though, typical substrate of laccases known to
be diphenol oxidase, monophenol e.g. sinapic acid or guaiacol can also oxidize
polyamines, aminophenols, lignin, aryl diamine, inorganic ions and it may mitigate
the toxicity of some polycyclic hydrocarbon. However, 2,20-azino-bis
(3-ethylbenzothiazoline-6-sulphonic acid) or ABTS is substrates which are most
commonly used, does not form quinone and is not pH dependent. Laccases have
been mostly isolated and characterized from plants and fungi, but only fungal
laccases are used currently in biotechnological applications for the detoxification
of complex industrial wastewater. Unfortunately, these enzymes usually work only
efficiently under mild acidic conditions (pH 4.0–6.0) whereas the temperature range
(30–55 C) for catalytic activity is suboptimal. In contrast, little is known about
bacterial laccases, which broad range substrate specificity for industrial application.
Most of study has been focused on white-rot fungus in which Phlebia floridensis
showed higher thermostability at pH 4.5 and T. versicolor is useful for dye
decolourization. Recently, in bacterial laccase polyphenol oxidase activity has
been reported in an Azospirillum lipoferum, phenol oxidases, laccase were isolated
from cell extracts of the soil bacterium Pseudomonas putida F6. Four strains of the
bacterial genus Streptomyces (S. cyaneus,S. ipomoea,S. griseus and
S.psammoticus) and the white-rot fungus T. versicolor were studied for their ability
to produce active extracellular laccase in biologically treated wastewater with
different carbon sources. Similarly, Proteus mirabilis, Bacillus sp., Raoultella
138 R. Chandra et al.
planticola and Enterobacter sakazakii are used for degradation of persistent organic
pollutants from biomethanated distillery spent wash, but it is still lacking for
industrial application (Chandra and Singh 2012). Laccases have low substrate
specificity make this interesting for degradation of several compounds with a
phenolic structure because they are extracellular and inducible, do not need a
cofactor, and have low specificity. Therefore, laccases have been employed in
several areas such as bioremediation of aromatic recalcitrant compounds, treatment
of effluents polluted with lignin, chemical synthesis, degradation of a wide number
of textile dyes, and biomass pretreatment for biofuel production.
Due to the properties of their substrate, the enzymes participating in the break-
down of lignin and other related compounds should be exclusively extracellular.
While this is without exception true for the LiP and MnP of white rot fungi and
bacteria, the situation is not the same with laccases. Although most laccases studied
so far are extracellular enzymes, while in wood-rotting fungi and some bacteria
species are usually also found intracellular laccase activity. Blaich and Esser (1975)
has been observed that most white-rot fungal species produced both extracellular
and intracellular laccases with isoenzymes showing similar patterns of activity.
T. versicolor was produced laccases both in extracellular and intracellular fractions
when grown on glucose, wheat straw, and beech leaves (Schlosser et al. 1997). The
intra and extracellular presence of laccase activity was also detected in
P. chrysosporium and Suillus granulates. The fraction of laccase activity in Neu-
rospora crassa,Rigidoporus lignosus and one of the laccase isoenzymes of
P. ostreatus is also probably localized intracellularly or on the cell wall. The
localization of laccase is probably connected with its physiological function and
determines the range of substrates available to the enzyme. It is possible that the
intracellular laccases of fungi as well as periplasmic bacterial laccases could
participate in the transformation of low MW phenolic compounds in the cell. The
cell wall and spores-associated laccases were linked to the possible formation of
melanin and other protective cell wall compounds. However, laccase has been
reported intracellularly as in A. lipoferum,M. mediterranea and in B. subtilis.
The bacterial cells must have some strategy to cope with the intracellular presence
of laccase and its toxic by-products. Rearrangement of the electron transport system
has been hypothesized to be one of the ways in which the laccase-positive cells
adapt to endogenous substituted quinones generated as products of laccase cata-
lyzed reaction. Because, in fungi, extracellular localization of the enzymes helps
them circumvent the problem of the reactive species, such as semiquinones and
quinines that are generated by laccases while oxidizing aromatic substrates. These
reactive species are powerful inhibitors of the electron transport system in both
bacteria and mitochondria. The loss of cytochrome c oxidase activity and
acquistion of resistance to quinone analogues has been demonstrated in a laccase-
positive variant of A. lipoferum.
8 Extremophilic Ligninolytic Enzymes 139
8.4.1 Molecular Structure
Laccases are monomeric, dimeric and tetrameric glycoproteins, generally having
fewer saccharide compounds (10–25%) in fungus and bacteria than in plant
enzymes. The carbohydrates, which are 10–45% of the total molecular mass, are
covalently linked, and due to this property of enzymes show high stability. Man-
nose is one of the major components of the carbohydrates attached to laccases. The
molecular weight of a laccase is determined to be in the range of 50–97 kDa from
various experimental reports. The structure of laccase from T. versicolor containing
approximately 500 amino acid residues organized in three β-barrel sequential
domains. The three domains are distributed in a first domain with 150 initial
amino acids, a second domain between the 150 and 300 residue, and a third domain
from the 300 to 500 amino acid. The structure is stabilized by two disulfide bridges
localized between domains I and II and between domains I and III. Figure 8.11a,b
shows the three dimensional ribbon structure of bacterial lacasse from Bacillus
subtilis XI based on the homology modelling of B. subtilis MB24 laccase PDB Id:
2x88A) (Guan et al. 2014) and fungal laccase from T. versicolor, respectively (PDB
Id: 1N68) (Robert et al. 2003).
Laccases of three molecular forms of isozyme have been reported namely, Lac I,
Lac II and Lac III. DSouza-Ticlo et al. (2009) reported that the MW of Lac II is
56 kDa from Cerrena unicolor. In another study the MW of laccase has been
reported ~97 kDa in T. versicolor. The biochemical properties of spore coat protein
Cot A of B. subtilis has been reported to be similar to multicopper oxidases
including laccases (Hullo et al. 2001). Further, the studies have revealed that Cot
A contains all the structural features of a laccase including the reactive surface-
exposed copper center (T1) and two buried copper centers (T2 and T3). The most
thermostable laccases have been isolated from Streptomyces lavendulae, with half-
life of 100 min at 70 C, and in B. subtilis, for which Cot A reported a half-life of
112 min at 80 C. The redox potential of any substrate also plays a very important
role in laccase activity. The redox potential of bacterial laccases ranges from 0.4 to
0.5 V, but they are active and stable at high temperatures (66 h at 60 C), at pH
7.0–9.0 and at high salt concentrations. The first gene and c-DNA sequences were
recorded for laccases from the fungi Neurospora crassa,Aspergillus nidulans,
Coriolus hirsutus and Phlebia radiata. Since, then the number of sequenced laccase
genes has considerably increased. The sequences mostly encode polypeptides of
approximately 500–600 amino acids in Bacillus subtilis. Cot A is 513 amino acids
long, and its MW is 65 kDa typical eukaryotic signal peptide sequences of about
21 amino acids are found at the N-terminal of the protein sequences. In addition to
the secretion signal sequence, laccase genes from N. crassa,Podospora anserina,
Myceliophthora thermophila and Coprinopsis cinerea contain regions that code for
N-terminal cleavable propeptides. These laccases also have C-terminal extensions
of controversial function, i.e., the last amino acids from the predicted amino acid
140 R. Chandra et al.
sequence are not present in the mature protein. Figure 8.11c, d shows the active site
of copper oxidase. The active sites of CueO (T1Cu, T2Cu, and T3Cu) in ribbon
form are shown in different colours (blue, green and orange, respectively), and the
figure also shows that Asp112 is located behind the tri-nuclear copper center.
Fig. 8.11 Three-dimensional ribbon structure (a)Bacillus subtilis X1 laccase based on the
Bacillus subtilis MB24 laccase crystal structure (PDB entry 2x88A) (Guan et al. 2014)(b) Laccase
of Trametes verssicola showing the two channels leading to the T2/T3 cluster (PDB entry 1GYC)
(Piontek et al. 2002)(c) Active site of CueO. Each Cu center is closely connected with each other.
Asp112 is located behind the trinuclear Cu center, forming hydrogen bonds with the imidazoles
coordinating to a T2Cu and a T3Cu directly and indirectly through a water molecule (PDB entry
1N68) (Robert et al. 2003)(d) The laccase active site showing the relative orientation of the Cu
atoms including interatomic distances among all relevant ligands
8 Extremophilic Ligninolytic Enzymes 141
8.4.2 Catalytic Cycle and Mode of Action
The catalytic mechanism of a laccase involves the reduction of the oxygen mole-
cule, including the oxidation by one electron of a wide range of aromatic com-
pounds, which include polyphenols, and methoxy-substituted monophenols and
aromatic amines. The use of molecular oxygen as the oxidant and water as gener-
ated by-product are very common catalytic features. Because, laccases have a four
copper (Cu) atom in their active site which participate in oxygen reduction and
water production (Fig. 8.11d). The laccases four Cu atoms are disseminated in
three types of cores or places: type 1 Cu (T1Cu), type 2 Cu (T2Cu), and type 3 Cu
(T3Cu). These cores are in two metallic active sites: the mononuclear location T1
and the trinuclear location T2/T3. It is believed that laccase catalysis involves the
following mechanism. Initially reduction of T1Cu by reducing substrate. Further,
Internal electron transfer from the T1Cu to T2Cu and T3Cu. Subsequently, reduc-
tion of oxygen to water at T2Cu and T3Cu site. The T1Cu gives the protein its blue
colour absorbance at about 600 nm depends on the intensity of the Scys–CuII bond
and the ligand field strength. T2Cu does not give any colour, but it is EPR
detectable, and T3Cu contains a pair of atoms in binuclear conformation that
gives a weak absorbance at 330 nm but is not detected by EPR. Spectroscopy
combined with crystallography has provided a detailed description of the active site
in a laccase. T2Cu and T3Cu form a trinuclear center, which is involved in the
catalytic mechanism.
In the catalytic mechanism, an oxygen molecule binds to the trinuclear cluster of
the asymmetric activation site, and it is postulated to restrict access of the oxidizing
agent. During the steady state, laccase catalysis indicates that O
2
reduction takes
place. The bonds of the natural substrate, lignin, are cleaved by laccase through C
α
C
β
cleavage and aryl cleavage. Subsequently, the lignin degradation is caused by
phenoxy radicals leading to the oxidation of the α-carbon or by the cleavage of the
bond between the α-carbon and β-carbon. This oxidation results in oxygen-centered
free radicals, which can be converted by a second enzyme catalyzed reaction to
quinone. The quinone and the free radicals can then undergo polymerization. The
T2Cu center coordinates with two His ligands and water as ligands. The Type
3 coppers are each 4-coordinate, having three His ligands and a bridging hydroxide
(Fig. 8.11d). The reduction of oxygen by a laccase appears to occur in two 2e
steps. The first is the rate determining step. In this, the T2/3Cu bridging mode is
reduced by the first 2e
. The peroxide-level intermediate facilitates the second 2e
reduction (from the T2Cu and T1Cu centers) in which the peroxide is directly
coordinated to the reduced T2Cu, and the reduced T1Cu is coupled to the T3Cu by
covalent Cys–His linkages.
The above mentioned information is described for blue copper oxidases, i.e. blue
laccases, but some authors have reported a small group of laccases that lack the
600 nm band and hence the blue color; some of these non-blue laccases (dubbed
“yellow” or “white”) feature a high redox potential allowing them to oxidize
non-phenolic compounds without any mediators. Both types of laccases (yellow
142 R. Chandra et al.
and blue) have similar MW (70 kDa) and specific activities, and it is observed that
the yellow laccases are obtained from the culture grown on solid state medium,
while the blue forms were isolated from culture grown on liquid medium without
lignin. The yellow laccases are formed by the modification of the blue forms with
low molecular weight lignin decomposition products, and some non-blue laccases
(yellow) have high redox potentials, allowing them to oxidize non-phenolic com-
pounds without any mediator (Pozdnyakova et al. 2006). Therefore, it is assumed
that the yellow form of a laccase is a result of binding of the aromatic product of
lignin degradation to the blue laccase. It has been postulated that a yellow laccase
might contain endogenous mediators derived from lignin, which perform the role of
exogenous mediator in the reaction of non-phenolic compounds. Due to insufficient
information about yellow laccases, more research is required in this field. In
Fig. 8.3a the simplest case is the one in which the substrate molecules are oxidised
to the corresponding radicals by direct interaction with the copper cluster. How-
ever, the substrates of interest cannot be oxidized directly by laccases, either
because they are too large to penetrate into the enzyme active site or because
they have a particularly high redox potential. By mimicking nature, it is possible
to overcome this limitation with the addition of “redox mediators”, which act as
intermediate substrates for laccases, whose oxidized radical forms are able to
interact with the bulky or high redox potential substrate targets Fig. 8.3b
Laccases catalyze the removal of one electron from the phenolic hydroxyl
groups of phenolic lignin model compounds, such as vanillyl glycol, 4,6-di
(t-butyl)guaiacol, and syringaldehyde, to form phenoxy radicals, which generally
undergo polymerization via radical coupling. The reaction is also accompanied by
demethylation, formation of quinone, resulting in ring cleavage. The degradation of
phenolic β-1 lignin substructure models occurs via the formation of phenoxy
radicals, which leads to C
α
–C
β
cleavage, C
α
oxidation, alkyl–aryl cleavage, and
aromatic ring cleavage. Laccase-catalyzed oxidation of phenols, anilines, and
benzenethiols correlates with the redox potential difference between the TiCu site
of the laccase and the substrate. The presence of electron withdrawing o- and
p-substituents reduces the electron density at the phenoxy group, and thus it is
more difficult to oxidize the phenolic substrate. Bulky substituents, which impose
steric interference with substrate binding, cause a decrease in reactivity.
Laccase has been found to oxidize non-phenolic model compounds and β-1
lignin dimers in the presence of a mediator, indicating that the enzyme plays a
significant role in the depolymerization of lignin and pulp delignification. The most
studied mediators for laccases are ABTS, 1-hydroxybenzotriazole (HBT), and
3-hyroxyanthranilic acid (HAA). The oxidation is different for ABTS and HBT,
involving a dication and a benzotriazolyl-1-oxide radical, respectively. Oxygen
uptake by the laccase is faster with ABTS than with HBT, but the oxidation of
non-phenolic substrate is comparable for both the mediators. The ABTS-mediated
oxidation of a non-phenolic substrate proceeds via an electron transfer mechanism.
In the oxidation process, the ABTS is first oxidized to a radical cation (ABTSc
+
),
and then to a dication (ABTS
2+
), with redox potentials of 472 mV (ABTS/ABTS
+)
and 885 mV (ABTS
+/ABTS
2+
), respectively. The dication is the active
8 Extremophilic Ligninolytic Enzymes 143
intermediate, which is responsible for the oxidation of the non-phenolic substrate.
The HBT
mediated oxidation of non-phenolic substrate involves the initial oxi-
dation of HBT to HBT
+
by laccase, followed by an immediate deprotonation to
form an N-oxy radical. The latter abstracts the benzylic H-atom from the substrate,
converting it to a radical. The oxidation of VA by laccase-ABTS and by laccase-
HBT (radical Habstraction mechanism) is presented in Fig. 8.12. HBT/HBT
+
, with
an E
o
value of 1.08 V, has a mediator efficiency with laccase that is higher than that
of ABTS. The use of laccase/HBT for the bleaching of paper pulp and for the
removal of lipophilic extractives has been described. Recently, two lignin-derived
phenols, syringaldehyde and acetosyringone, have been shown to act as effective
laccase mediators for the removal of lipophilic compounds from paper pulp.
The degradation of non-phenolic β-O-4 model compounds, which represent the
major substructure in lignin, has been studied using laccase-mediator systems. Four
types of reactions, β-ether cleavage, C
α
–C
β
cleavage, Cα-oxidation and aromatic
ring cleavage, are catalyzed by the laccase–BHT (butylated hydroxytoluene)
coupled system. In the oxidation of a non-phenolic β-O-4 lignin model dimer,
1-(4-ethoxy-3-methoxyphenyl)-1,3-dihydroxy-2-(2,6-dimethoxyphenoxy)propane,
the coupled enzyme/HBT system catalyzes the 1eoxidation of the substrate to
form a β-aryl radical cation or benzylic (Ca) radical intermediates. The electron
density of the aromatic ring affects the 1eoxidation by the laccase/1-HBT couple.
Substrates containing electron-donating groups favor aromatic ring cleavage prod-
ucts. The β-aryl radical cation is converted to the product via an aromatic ring
cleavage, and the benzylic radical is cleaved at the C
α
–C
β
bond, similar to a
Baeyer–Villiger reaction. The β-ether cleavage of the β-O-4 lignin substructure is
caused by reaction with the Ca-peroxy radical intermediate produced from the
benzylic radical. The rate of oxidation depends on the kcat of the laccase for
the mediator and the stability of the enzyme to inactivation by the free radical of
the mediator.
The ability of LiP, MnP and lacasses to degrade lignin has been studied in
diverse industrial processes and bioremediation of contaminated soils and water,
but this ability is non-identical between these three types of enzymes. This may be
due to that enzyme-substrate interactions are different. The study of the interactive
OCH3OCH3OCH3
CH2OH CH2OH CHOH
OCH3
CHO
ABTS++ ABTS+
ABTS+ABTS++
ABTS+ABTS
e- H
ABTS
-O3S-O3S-O3S
SO3-SO3-
C2H5
C2H5
C2H5
C2H5
SO3-
e-
e-
e-
e-
S
+
S
N
NN
C2H5
C2H5
SS
SN N
NN
N
NN
N
S
N
+
+
+
+
-
+
-
Fig. 8.12 Radical H-atom abstraction and electron transfer mechanism
144 R. Chandra et al.
mechanisms involved in ligninolytic enzyme and lignin is indeed important in
understanding enzyme reactions and may provide further insights to the develop-
ment of biodegradation technologies. Ligninolytic enzymes are reported
for degradation of lignin by direct interactions of ligninolytic enzymes in terms
of a long-range electron transfer process. However, little is known about the effect
of ligninolytic enzymes structures on the lignin biodegradation at the molecular
level. The molecular docking approach can be used to model the interaction
between a small molecule and a protein at the atomic level, which allow us to
characterize the behavior of small molecules in the binding site of target proteins as
well as to elucidate fundamental biochemical processes. Crystallographic water is a
major challenge in molecular docking. These molecules are strongly bound to the
receptor and observed across several crystallographic structures of a particular
protein. In approximately 65% of the crystallographic protein-ligand complexes,
at least one water molecule is involved in ligand-receptor recognition. The release
of a crystallographic water molecule from its binding site is entropically favorable;
however the process causes a simultaneous loss in enthalpy. To compensate for this
enthalpy loss, a specific moiety of the ligand can be designed to mimic the
interaction network of the displaced water through the formation of equivalent
hydrogen bonds with the protein. Alternatively, structural water can be explicitly
included in the docking experiments, allowing the formation of highly favorable
hydrogen-bonding networks between the ligand and the target binding site. In this
case, a variety of methods are available to evaluate which water molecules are
strongly bound and, therefore, suitable for this purpose. Among these strategies one
can highlight free energy perturbation calculations using Monte Carlo (MC) statis-
tical mechanics simulations, which estimate the binding free energy for a given
water molecule, allowing the discrimination between displaceable and strongly-
bound structural water. MC methods generate poses of the ligand through bond
rotation, rigid-body translation or rotation. The conformation obtained by this
transformation is tested with an energy- based selection criterion. If it passes the
criterion, it will be saved and further modified to generate next conformation. The
main advantage of MC is that the change can be quite large allowing the ligand to
cross the energy barriers on the potential energy surface, a point that isnt achieved
easily by molecular dynamics based simulation methods.
8.4.3 Common Substrate and Microorganisms
Laccases have a broad range of substrate specificity. Due to this, microorganisms
oxidize a wide range of environmental pollutants as a sole carbon or nitrogen source
for their growth and metabolism. Laccases have been mostly isolated and charac-
terized from plants and fungi, and only fungal laccases are currently used in
biotechnological applications for the detoxification of complex industrial wastewa-
ter. Laccase producing microorganisms and their some common substrate are listed
in Table 8.3.
8 Extremophilic Ligninolytic Enzymes 145
Table 8.3 Some important laccase producing microorganisms in extremophilic environment and
their substrate
Microorganisms Substrate pH Temp. (C)
Fungi
Penicillium pinophilum
TERI DB1
Distillery effluent decolourisation 8.5 25
Trametes versicolor Non-phenolic lignin model dimer 5.0 25
Polyporus pinisitus Direct red 28, Acid Blue 74 4–5 50
Myceliophthora
thermophila
Direct red 28, Acid Blue 74 6.0 70–80
Trametes trogii Nitrobenzene and anthracene 5.0 25
Trametes versicolor Direct red 28, Acid Blue 74 5.0 65
Pleurotus sp. 2,4 Dichlorophenol, Benzo(a)pyrene [B(a)P] 5.0 28
Trichoderma atroviride Catechol, o-cresol, 4.0–5.0 40–50
Phanerochaete
crysosporium
Lignin 5.0 25
Daedalea flavida Lignin 5.0 25
Pycnoporus coccineus Anthracene, pyrene, fluoranthene, benzo[a]
pyrene, phenanthrene
5.0 28
Phlebia sp. Lignin 5.0 25
Phlebia floridensis Lignin 5.0 25
Pseudochrobactrum
glaciale (FJ581024)
Pulp paper mill effluent 5.0 25
Providencial rettgeri
(GU193984)
Pulp paper mill effluent 5.0 25
Ganoderma lucidum Antracene, benzo[a]pyrene, fluorine,
acenapthene, acenaphthylene and benzo[a]
anthracene
––
Bacteria
Serratia marcescens
(GU193982)
Black liquor 5.0 25
Citrobacter
sp. (HQ873619)
Black liquor 5.0 25
Klebsiella pnumoniae
(GU193983)
Black liquor 5.0 25
Staphylococcus
saprophyticus
Brilliant blue, methyl orange, neutral red 3.0 32
Bacillus SF Mordant black 9, mordant brown 96 &
15, acid blue 74
8.0 60
γ-Proteobacteria JB Caramine 4–10 55
Bacillus subtilis Syringaldazine, ABTS 3.0–7.0 75
146 R. Chandra et al.
8.4.4 Screening of Laccase Producing Microorganisms, Its
Substrate, Bioassay and Purification
Laccases can oxidize a wide range of molecules more than hundred different types
of compound have been identified as substrate for laccase. There are various natural
and synthetic substrates which are mentioned in Table 8.4, used for laccase assay.
All substrates cannot be directly oxidized by laccases, either because of their large
size which inhibit their penetration into the enzyme active site or because of their
particular high redox potential. To overcome this hindrance, suitable chemical
mediators are used which are oxidized by the laccase and their oxidized forms
are able to interact with high redox potential substrate.
Although polyphenol oxidases copper proteins are able to oxidize aromatic
compounds with molecular oxygen as the terminal electron acceptor. Polyphenol
oxidases are associated with three types of activities:
(a) Catechol oxidase or o-diphenol: oxygen oxidoreductase (EC 1.10.3.1)
(b) Laccases or p-diphenol: oxygen oxidoreductase (EC 1.10.3.2)
(c) Cresolase or monophenols monooxygenase (EC 1.18.14.1)
Faure et al. (1995), compared commercial fungal laccase and catechol oxidase,
purified from Pyricularia oryzae and Agaricus bisporus, respectively, with bacte-
rial laccase from A. lipoferum by using several substrates and phenol oxidase
inhibitors. Five classes of chemical compounds were investigated as substrates
for laccase:
1. L-Tyrosine and several substituted monophenols such as p-coumaric and o-
hydroxyphenylacetic or salicylic acids;
Table 8.4 Natural and synthetic substrate of laccases
S. No. Natural substrate Synthetic substrate
1. Acetosyringone 1- Hydroxylbenzotrizole (HBT)
2. Syringaldehyde N- Hydroxyphthalimide (HPI)
3. Vanilin Violuric acid (VLA)
4. Acetovanillone N- Hydroxylacetanlide (NHA)
5. Sinapic acid 2,2,6,6-Tetramethylpiperidine- N-oxyl (TEMPO)
6. Ferulic acid Acetohydroxamic acid
7. p-Coumaric acid 2,2,5,5- Tetramethylpyrrolidine-N-oxyl (PROXYL)
8. Reduced glutathione 2,20-Azinobis-(3-ethylbenzothia- zoline 6- sulfonic acid)
(ABTS)
9. Cystine Guaicol
10. Aniline Methyl syringate
11. 4 hydroxybenzyl
alcohol
8 Extremophilic Ligninolytic Enzymes 147
2. o-Diphenols (catechol, pyrogallol, guaiacol, and protocatechic, gallic, and
caffeic acids), L-3, 4- dihydroxyphenylalanine and o-aminophenol, which
could be oxidized by both laccase and catechol oxidase;
3. p-Diphenol and p-substituted aromatic compounds as typical p-phenol oxidase
substrates such as hydroquinone, p-cresol, p-aminophenol and
p-phenylenediamine;
4. m-Diphenols such as resorcinol, orcinol, 4-hexylresorcinol, and
5-pentadecylresorcinol;
5. Other laccase substrates such as syringaldazine, 1-naphthol, ABTS, and 4- and
5-hydroxyindoles. The range of substrates used by A. lipoferum laccase was
similar to that used by P. oryzae laccase.
The screening methods of laccase producing microorganisms are similar to MnP
as described previously but the minimal media does not contained H
2
O
2
. Bioassay
of laccase activity is measured based on the oxidation of various substrate in
presence of suitable buffer (i.e. sodium acetate) in acidic pH (5.0). The oxidation
product is measured by spectrophotometer taking the absorbance at 420 nm or
450 nm depending upon substrate specificity. ABTS has been most commonly used
substrate for laccase bioassay because it acts as cooxidant that can interact with
laccase to accomplish electron transfer and it is chemically oxidized in two steps via
ABTS
+
and ABTS
2+
. Anisyl alcohol and benzyl alcohol can be better oxidized by
ABTS
2+
than by ABTS
+
. Laccase activity is assay through ABTS method. In this
method reaction mixture contained 600 μL sodium acetate buffer (0.1 M, pH 5.0 at
27 C), 300 μL ABTS (5 mM), 300 μL culture filtrate and 1400 μL distilled water.
The mixture is then incubated for 2 min at 30 C and the absorbance was measured
immediately in 1-min intervals. One unit of laccase activity has been defined as
activity of an enzyme that catalyzes the conversion of 1 mole of ABTS per minute
.
Laccase activity has been measured by another substrate guaiacol. The reaction
mixture contained 10 mM sodium acetate buffer (pH 5.0), 2 mM guaiacol and
0.2 ml of culture supernatant was incubated at 25 C for 2 h and the absorbance was
read at 450 nm. The relative enzyme activity has been expressed as colorimetric
units/ml (CU/ml).
In general plant lacasses are purified from sap or tissue extracts, whereas fungal
lacasses are purified from culture are purified from culture (fermentation broth) of
the selected organism. Various protein purification techniques are used for purifi-
cation of laccase. Typical purification protocols involve ultrafiltration,
ion-exchange, gel filtration and other electrophoretic and chromatographic tech-
niques. Purification may be single or multi-step process. A single step lacasse
purification from N. crassa are performed by using celite column chromatography
and 54 fold purification with specific activity of 333 U/mg. Laccase from T
versicolor are purified using ethanol precipitation, DEAE–sepharose, phenyl-
sepharose and sephadex G-100 chromatography. T. versicolor 951022 excrete a
single monomeric laccase showing a high specific activity of 91,443 U/mg for
ABTS as substrate. Laccase from T. versicolor is purified with ion exchange
chromatography followed by gel filtration with specificity activity of 101 U mL
1
148 R. Chandra et al.
with 34.8 fold purification. Laccase from Stereum ostrea and obtained up to 70-fold
purification from culture filtrate by a two step protocol-ammonium sulphate (80%
w/v) and sephadex G-100 column chromatography.
8.4.5 Thermostability of Bacterial Laccases
Laccases are highly stable, industrially important enzymes that are capable of
oxidizing a large range of substrates. Thermostability plays an important role in
enzyme catalysis; several sequence and structural factors are involved in this
phenomenon. Thermostable enzymes allow high process temperatures with higher
associated reaction rates and less risk of microbial contamination. Some of the
mechanisms/indicators of increased thermostability include: a more highly hydro-
phobic core, tighter packing (compactness), a deleted loop, greater rigidity
(e.g. through increased proline content in the loop), higher secondary structure
content, greater polar surface area, fewer thermolabile residues, increased
H-bonding, higher pI, a disulfide bridge, more salt bridges and buried polar
interactions. Moreover, the enhanced thermostability of a laccase from Bacillus
sp. HR03 using site directed mutagenesis of the surface loop was achieved, in which
glutamic acid (Glu188) was substituted with 2 hydrophilic(lysine and arginine) and
1 hydrophobic (alanine) residues. There are some bacterial species that show
thermostability at different temperatures:
(a) Cot A from B. subtilis at 75 C showed maximum activity and at 80 C wild
type Cot A has a life of 4 h, whereas recombinant Cot A from E. coli has a half-
life of 2 h.
(b) T. thermophilum laccase has optimum activity at 92 C with a half-life of 4 h at
80 C and attains 60% of its activity after incubation for 10 min at 100 C.
8.5 Industrial and Biotechnological Applications of MnP,
LiP and Laccase
Ligninolytic enzymes are involved in the degradation of the complex and recalci-
trant environmental pollutants. This group of enzymes is highly versatile in nature
and they find application in a wide variety of industries. The biotechnological
significance of these enzymes has led to a drastic increase in the demand for
these enzymes in the recent time because of their oxidative ability toward a broad
range of phenolic and non-phenolic compounds.
8 Extremophilic Ligninolytic Enzymes 149
8.5.1 Industrial Waste Detoxification and Bioremediation
MnP, LiP and laccase have used to detoxify or remove various aromatic compounds
found in industrial waste and contaminated soil and water. MnP have capability to
mineralised various PAHs such as anthracene, benzo[a]pyrene, benz[a]anthracene,
phenanthrene, [U-
14
C]pentachlorophenol, [U-
14
C]catechol, [U-
14
C]tyrosine, [U-
14
C]tryptophan, [4,5,9,10-14C]pyrene, [ring U-
14
C]2-amino-4-6-dinitrotolune and
1.1.1-trichloro-2-2-bis-(4-chlorophenyl) ethane (DDT). MnP has also been reported
for the elimination and detoxification of triclosan, an emerging persistent pollutant
with ubiquitous presence in aquatic environment. Further, MnP detoxified afla-
toxins B
1
, lignite originated humic acid, nylon, polyethylene and amino carbonyl
maillard products (melanoidins). LiP and MnP was effective in decolourizaing kraft
pulp paper mill effluent. LiP was reported for mineralisation of PAHs like naph-
thalene, phenanthrene, benzo[c]phenenthrene, benz[e]pyrene, benz[a]anthracene,
pyrene, chrysene, anthracene, benzo[a] pyrene, perylene. Laccase has been reported
for the degradation and decolourization of chlorophenol- and chlorolignin-
containing black liquor and pulp paper mill wastewater. However, laccase has
been reported to mineralised anthracene, benzo pyrene, fluorene and other
16 PAHs compounds which are listed by USEPA (United States Environmental
Protection Agency) as priority pollutants of environment
8.5.2 Decolourisation of Textile Dye, Pulp Paper Mill
and Distillery Effluent
MnP decolourise and degrade various types of synthetic dye; azo dye (reactive red
120, congo red, orange G and orange IV, remazol brilliant violet 5R, direct red 5B),
anthraquinone dye (remazol brilliant blue R), indigo dye (Indigo Carmine) and
triphenylmethane dye (Methyl Green) containing textile wastewater. LiP
decolourised various dyes e.g. bromophenol blue, congo red, methylene blue,
methyl green, methyl arrange, poly-R-478, poly S-119, poly T-128. Laccase have
capability to decolourised a wide range of dyes e.g. cibanon red 2B-MD, cibanon
golden yellow PK-MD, cibanon blue GFJ-MD, indanthrene direct black RBS,
remazol brilliant blue R, congo red etc. Laccase catalysed textile dye bleaching
may also be useful in finishing dyed cotton fabric. Under laccase catalysis, soluble
dye precursors could be absorbed, oxidise and polymerised to give the desired
tanning effect. LiP decolourised four distinct classes of dyes, they are-
(a) tryphenylmethane(bromophenol blue), (b) heterocyclic dye (methylene blue
and toluene blue O), (c) azo dye (congo red and methyl orange) and
(d) polymeric dyes (Poly R-478, Poly S-119, AND Poly T-128).
150 R. Chandra et al.
8.5.3 Production of Ethanol and Value Added Products
Ligninolytic enzymes play a central role in lignin degradation. Due to high redox
potential, MnP, LiP and laccase are of high industrial interest for delignification and
production of ethanol and other cellulosed based low molecular weight chemicals
such as vanillin, dimethoxy sulfoxide and phenol. Laccases have been used for the
synthesis of several anticancer drugs such as actinocin, vinablastine and other
pharmaceutical products. Laccase have been employed in several applications
organic synthesis as the oxidation of functional group, the coupling of phenols
and steroids, medical agent (anesthetics, anti-inflammatory, antibiotics and seda-
tives) and synthesis of complex natural products. MnP and LiP have potential to
produce natural aromatic flavours compound e.g. vanillin, β-ionone, β-cyclocitral,
dihydroactinidiolide, flavanoids and so one. Laccase can be applied to certain food
processes that enhance or modify the colour appearance of food or beverages for the
elimination of undesirable phenolic compounds, responsible for the browning haze
formation and turbidity in clear fruit juice, beer and wine. The uses of laccase in
baking process increase strength, stability and reduced thickness, thereby improv-
ing the machine ability of dough. Laccase have also been employed to sugar beet
pectin gelation, baking and ascorbic acid determination.
8.5.4 Development of Biosensors
MnP and LiP are known as redox enzyme with efficient direct electron transfer
(DET) properties with electrode. It may enable to use for development of biosensor
based on DET, effective biofuels cell. However, laccase catalysis could be useful as
biosensor for detecting oxygen and a wide variety of reducing substrate (phenols
and anilines). A large number of biosensors containing laccase have been devel-
oped for immunoassay, glucose determination, aromatic amines and phenolic
compounds determination. A carbon paste biosensor modified with a crude extract
of the P ostreatus as a source of laccase source is proposed for catecholamine
determination in pharmaceutical formulation.
8.5.5 Biomechanical Pulping and Pulp Biobleaching
MnP, LiP and laccase are the most important enzyme involved in biomechanical
pulping and kraft pulp bleaching. In the laboratory scale consumption of refining
energy in mechanical pulping was reduced with MnP pre-treatment. However, MnP
degraded residual lignin of kraft pulp and enhanced the pulp bleaching effect. The
laccases have also attracted considerable interest for pulp bio-bleaching. During
lignin degradation, laccases are thought to act on small phenolic lignin fragments,
8 Extremophilic Ligninolytic Enzymes 151
in which the substrate reacts with the lignin polymer, resulting in its degradation.
Laccase-catalyze textile dye bleaching may also be useful in finishing dyed cotton
fabrics.
Take Home Message
Lignin is amorphous complex polymer of phenylpropane units, which are cross-
linked to each other with a variety of different chemical bonds. It confers rigidity
and recalcitrant nature to the lignocellulosic biomass.
The common extremophilic ligninolytic enzymes are manganese peroxidase
(MnP), lignin peroxidase (LiP) and laccase. Such enzymes have also proven
their utility in the pollution abatement, especially in the treatment of industrial
waste/wastewater containing hazardous compound like phenols, chlorolignin
synthetic dyes, and polyaromatic hydrocarbons (PAHs) as well as recalcitrant
organic compounds structurally similar to lignin.
Microorganisms with systems of thermostable enzymes decrease the possibility
of microbial contamination in large scale industrial reactions of prolonged
durations. The mechanisms for many thermotolerant enzymes have been
reported due to their structural properties i.e. presence of Ca
2+
, saturated fatty
acid, α-helical structure etc.
The ability of extremophilic ligninolytic enzymes to tolerate high temperature,
different metal ions and organic solvents is very important for the efficient
application of this enzyme in the biodegradation and detoxification of industrial
waste.
The engineering of a disulfide bond (A48C and A63C) near the distal calcium
binding site of MnP by double mutation showed the improvement in thermal
stability as well as pH (pH 8.0) stability in comparison to native enzyme (MnP).
Some of the mechanisms/indicators of increased thermostability of laccases
include: a more highly hydrophobic core, tighter packing (compactness), a
deleted loop, greater rigidity (e.g. through increased proline content in the
loop), higher secondary structure content, greater polar surface area, fewer
thermolabile residues, increased H-bonding, higher pI, a disulfide bridge, more
salt bridges and buried polar interactions.
Moreover, the enhanced thermostability of a laccase from Bacillus sp. HR03
using site directed mutagenesis of the surface loop was achieved, in which
glutamic acid (Glu188) was substituted with 2 hydrophilic (lysine and arginine)
and 1 hydrophobic (alanine) residues.
MnP, LiP and laccase have been used to detoxify or remove various aromatic
compounds found in industrial waste and contaminated soil and water. It is used
for the decolourisation of textile dye, pulp paper mill and distillery effluent.
MnP, LiP and laccase are of high industrial interest for delignification and
production of ethanol and other cellulose based low molecular weight chemicals
such as vanillin, dimethoxy sulfoxide and phenol. Laccases have been used for
the synthesis of several anticancer drugs such as actinocin, vinablastine and
other pharmaceutical products. MnP, LiP and laccase are the most important
enzymes involved in biomechanical pulping and kraft pulp bleaching. MnP and
152 R. Chandra et al.
LiP are known as redox enzyme with efficient direct electron transfer (DET)
properties with electrode. It may enable to use for development of biosensor
based on DET, effective biofuels cell.
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... In previous studies, the molecular weight of peroxidases enzyme in Phanerochaete chrysosporium, Phlebia radiate IZU 154 and Bjerkandera adusta was approximately 42 kDa, 43 kDa, 49 kDa and 45 kDa, respectively, which supports with the current research (Chandra et al. 2017;Baik et al. 2021). The production of laccase by various fungal strains, including Phenerochaete flavidoalba (96 kDa), Ganoderma lucidum (75 kDa), Panaeolus sphincrinus (60 kDa) and Trametes gallica (60 kDa), has also been reported (Agunbiade et al. 2021). ...
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... They are extra-cellular and have an inducible nature, which results in limited substrate specificity for breakdown. Because of their very compact hydrophobic core, hydrogen bonds, and increased polar surface area, laccases are known to be very thermostable (Chandra et al. 2017). Through the method of site-directed mutagenesis, it has been possible to enhance specific laccase qualities like thermal stability, activations, and compactness. ...
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Bioremediation is the process by which the contaminants are degraded by microorganisms. It involves the growth of specific microbes that has the ability to digest the contaminants such as oil, solvent, pesticides, hydrocarbon, greenhouse gases, and radioactive materials. Radioactive contaminants are hazardous to living organisms and the environment, exposure to radiation causes mild to chronic diseases in humans and other organisms. Diseases such as cancer (lungs, thyroid, and skin), acute radiation syndrome, cardiac, reproductive, neurological, and various diseases have been reported in humans exposed to radiation. These microorganisms have the capability of decreasing the mobility of radioactive material by Precipitation or Crystallization. Also, these microbes increase the process of metal solubility and are also involved in chemical reactions like Oxidizing the metal ions or reduction of metal ions like Uranium reducing from U(VI) to U(IV) with the help of bacteria Desulfovibrio and Desulfuricans. Also, in the case of chromium, the metal reduces from Cr (VI) to Cr (III). Recent discoveries yield that, the microorganisms are engineered to accumulate a higher dose of radiation which will lead to radioactive resistance. This type of strain has the capability to convert the toxic, volatile metal species to less reactive and less toxic in nature. This methodology can be achieved by multiple remediation strategies with more genetic engineering clusters. Scientists have inferred that toluene and chlorobenzene metabolism with the help of engineered bacteria to remediate the multiple contaminants caused by the nuclear plants. In this review, we have discussed the overview of Radioactive contaminants and the effect of Bioremediating the radioactive contaminants using Microorganisms.
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The crystal structure of the major lignin peroxidase isozyme from Phanerocheate chrysosporium has been refined to an R = 0.15 for data between 8 A and 2.03 A. The refined model consists of 2 lignin peroxidase molecules in the asymmetric unit, 2 calcium ions per monomer, 1 glucosamine per monomer N-linked to Asn-257, and 476 water molecules per asymmetric unit. The model exhibits excellent geometry with a root mean square deviation from ideality in bond distances and angles of 0.014 A and 2.9 degrees, respectively. Molecule 1 consists of all 343 residues, while molecule 2 consists of residues 1-341. The overall root mean square deviation in backbone atoms between the 2 molecules in the asymmetric unit is 0.36 A. The refinement at 2.0 A confirms our conclusions based on the partially refined 2.6-A structure (Edwards, S. L., Raag, R., Wariishi, H., Gold, M. H., and Poulos, T. L. (1993) Proc. Natl. Acad. Sci. U.S.A. 90, 750-754). The overall fold of lignin peroxidase closely resembles that of cytochrome c peroxidase. A superimposition of alpha-carbons gives a root mean square deviation of 2.65 A between the two peroxidases and 1.66 A for the helices. The active sites also are similar since both contain a proximal histidine heme ligand hydrogen-bonded to a buried aspartate residue and both contain histidine and arginine residues in the distal peroxide binding pocket. The most obvious difference in the active site is that whereas cytochrome c peroxidase has tryptophan residues located in the proximal and distal heme pockets, lignin peroxidase has phenylalanines. There are four other especially noteworthy differences in the two structures. First, although the heme in cytochrome c peroxidase is recessed about 10 A from the molecular surface, the heme pocket is open to solvent. The analogous opening in lignin peroxidase is smaller which can explain in part the differences in reactivity of the two hemes. This same opening may provide the site for binding small aromatic substrates. Second, lignin peroxidase has a carboxylate-carboxylate hydrogen bond important for heme binding that is not present in cytochrome c peroxidase. Third, lignin peroxidase contains 2 structural calcium ions while cytochrome c peroxidase contains no calcium. The calciums in lignin peroxidase coordinate to residues near the C-terminal ends of the distal and proximal helices and hence are probably important for maintaining the integrity of the active site. Fourth, the extra 49 residues in lignin peroxidase not present in cytochrome c peroxidase constitutes the C-terminal end of the molecule with the C terminus situated at the “front” end of the molecule between the two heme propionates.
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Lignin peroxidase from the culture filtrate of Loweporus lividus MTCC-1178 has been purified to homogeneity using Amicon concentration and DEAE cellulose chromatography. The molecular weight of the purified lignin peroxidase using SDS-PAGE analysis has been found to be 40 kDa. The Km values for veratryl alcohol and H2O2 for the purified enzyme were 58 and 83 μM, respectively. The calculated kcat value of the purified enzyme using veratryl alcohol as the substrate was 2.5 s−1. The pH and temperature optima of lignin peroxidase have been found to be 2.6 and 24°C, respectively.
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Melanoidins, complex biopolymer of amino-carbonyl compounds are the major coloring and polluting constituents of distillery wastewaters. In this study, three aerobic melanoidin-degrading bacteria (RNBS1, RNBS3 and RNBS4) were isolated from soil contaminated with distillery effluent and characterized as Bacillus licheniformis (RNBS1), Bacillus sp. (RNBS3) and Alcaligenes sp. (RNBS4) by biochemical tests and 16S rRNA gene sequence analysis. The degradation of synthetic and natural melanoidins was studied by using the axenic and mixed bacterial consortium. Results have revealed that the mixed consortium was more effective compared to axenic culture decolorizing 73.79 and 69.83% synthetic and natural melanoidins whereas axenic cultures RNBS1, RNBS3 and RNBS4 decolorized 65.88, 62.56 and 66.10% synthetic and 52.69, 48.92 and 59.64% natural melanoidins, respectively. The HPLC analysis of degraded samples has shown reduction in peak areas compared to controls, suggesting that decrease in color intensity might be largely attributed to the degradation of melanoidins by isolated bacteria.
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