This document appears to be a student's thesis proposal on isolating and characterizing lignin-degrading microbes. It includes an abstract stating the aim is to identify bacteria and fungi with high potential for degrading lignocellulose compounds. The introduction provides background on lignin and current lignin degradation research. The literature review covers the chemical composition of lignocellulose and its degradation, including details on cellulose, hemicellulose, and lignin breakdown enzymes like lignin peroxidase and manganese peroxidase produced by fungi. The proposal seems to focus on isolating microbes from paper/pulp industry effluent and soil that can degrade lignin as measured by laccase and lignin peroxidase assays.

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Isolation and characterization of Lignin-degrading microbes
to be submitted as Major Project in partialfulfilment of the
requirement for the degree of B.Tech.
Submitted by
Tarun Shekhawat
2K14/BT/027
Delhi TechnologicalUniversity, New Delhi, India
under the supervision of
Smita RastogiVerma
Assistant professor
Department of Biotechnology

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DECLARATION
I Tarun Shekhawat, hereby declare that the work entitled “Isolation and characterization of
Lignin-degrading microbes” has been carried out by me under the guidance of Dr. Smita Rastogi
Verma, in Delhi Technological University, New Delhi.
This Synopsis is a part partial fulfilment for the degree of B.Tech in Biotechnology. This is the
original work and has not been submitted for any other degree in any other university.

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CERTIFICATE
This is to certify that the dissertation entitled “Isolation and characterization of Lignin-degrading
microbes” in the partial fulfilment of the requirements for the reward of the degree of Bachelors of
Engineering, Delhi Technological University (Formerly Delhi College of Engineering, University
of Delhi), is an authentic record of the candidate’s own work carried out by him/her under my
guidance. The information and data enclosed in this thesis is original and has not been submitted
elsewhere for honoring of any other degree.
Dr. Smita Rastogi Verma
(Project Advisor)
Department of Bio-Technology
Delhi Technological University
(Formerly Delhi College of Engineering, University of Delhi)

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ACKNOWLEDGEMENT
I owe sincere thanks to Prof. D.Kumar, Head of Biotechnology department for his timely support
and inspiration.
I am extremely grateful to my guide Dr. Smita Rastogi Verma, Assistant Professor (Department of
Biotechnology) for her guidance, mentorship, motivation and valuable suggestions.
Last but not the least I would also like to thank my parents and my friends for giving me love and
support.
Tarun Shekhawat
2K14/BT/027

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CONTENTS
TOPIC PAGE NO
1) ABSTRACT ......................................................................................................6
2) INTRODUCTION ............................................................................................7
3) REVIEW OF LITERATURE 8-14
4) MATERIALS AND METHODS …………………………………………….15-24
5) RESULTS AND DISCUSSION.........................................................................25-28
6) CONCLUSION .................................................................................................29
7) REFERENCES....................................................................................................30-32

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ABSTRACT
Wood and other lignocellulosic biomass is a low cost and abundant resource that can be used in
the large scale production of fuels and chemicals. The presence of covalent lignin-carbohydrate
linkages between sugar hydroxyl of hemicelluloses and phenylpropane subunits in lignin gives
lignocellulose protection against degradation. Lignin, which is the nature’s plastic is the major
pollutant from paper-pulp mill effluent due to its intense unaesthetic brown color, hydrophobicity
and poor mechanical properties. Textile dye based industries release colored effluents due to
presence of large amount of mixture of dyes which is also hazardous. Microbial extracellular
lignin peroxidase enzymes have a potential to degrade lignin and a wide range of complex
aromatic dyestuffs. The aim of this project is to identify new strains of bacteria and fungi with
high potential in the degradation of lignocellulose compounds. This study focuses on isolation of
both bacteria and fungi from effluent of paper and pulp industries as well as from soil samples
collected from various locations. These microbes will be screened on the basis of Laccase and
Lignin peroxidase plate assay.

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1. INTRODUCTION
The lignocellulosic material of plants consists of three main components, namely cellulose,
hemicellulose and lignin. After cellulose, lignin is the second most abundant renewable
biopolymer in nature. It is the most abundant aromatic polymer in the biosphere. It is an essential
part of the plant cell wall, imparting rigidity and protecting the easily degradable cellulose from
attack by pathogens. Due to its complicated structure and non-hydrolysable bonds, lignin is more
difficult to break down than cellulose or hemicellulose. Lignin surrounds cellulose in the plant cell
wall forming a matrix, which itself is resistant to degradation. Lignin biodegradation is
responsible for much of the natural destruction of wood in use, and it may have an important role
in plant pathogenesis. On the other hand, potential applications utilizing lignin degrading
organisms and their enzymes have become attractive, because they may provide environmental
friendly technologies for the paperpulp and various other industries.
In biosphere, a wide variety of species are involved in lignin degradation including fungi and
bacteria. To date, only a few groups of organisms are capable of degrading complex lignin
polymers, and they are best exemplified by the white rot fungi and others such as Phanerochaete
chrysosporium, Streptomyces viridosporus, Pleurotus eryngii, Trametes trogii, Fusarium proliferatem,
Agaricus, Erwenia, Copricus, Mycema, Sterium. However, commercialization of lignin degradation
by fungi has disadvantages in the form of problems related to fungal protein expression and
genetic manipulations and showed a lack of stability under practical treatment condition
involving high pH, oxygen limitation and high lignin concentrations. For this reason, studies on
the bacterial degradation were more preferable for lignin and the production of bacterial
ligninolytic enzymes has seen increased in recent year.
Lignin degrading enzymes are essentially extracellular in nature due to the large and complex
structure of lignin which cannot enter the cell for intracellular action. Lignin peroxidase (LiP) is an
enzyme first discovered in 1983 was used to degrade lignin. Lignin peroxidase is useful in the
treatment of colored industrial effluents and other xenobiotic as it has bioremediation potential to
decolorize the effluents. Decolorization of methylene blue dye was also used as an indicator of the
oxidation ability of ligninolytic enzyme produced by the potential lignin degrading bacterial
strains. Lignocellulytic enzymes also have significant potential application in various industries
including chemicals, fuel, food, brewery and wine, animal feed, textile and laundry, pulp and
paper, and agriculture. In the present study, the isolation and biochemical characterization of
lignin-degrading microbes is proposed.

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REVIEW OF LITERATURE
This literature review contains information on the three main components of wood, their chemical
composition and degradation, strains of fungi and lignocellulose substrates that will be used in
this project
2.1 Chemical composition and degradation of lignocelluloses
Wood consists of about 45% of cellulose, 20 – 30 % hemicelluloses and 20 - 40 % lignin depending
on the wood species, their parts and age.
2.1.1 Cellulose
Cellulose, the major chemical component of the fiber wall, is a homopolysaccharide composed
entirely of D-glucose linked together by β-1,4-glycosidic bonds with degree of polymerization
ranging from 1,000 in bleached kraft pulps to 10,000 in native wood. Each glucose unit is rotated
180o relative to the adjacent one and the smallest repetitive unit of cellulose is cellobiose, two
glucose units. Cellulose is a linear structure that has a strong tendency to form intra- or
intermolecular hydrogen bonds resulting in the formation of cellulose microfibrils which promote
aggregation into crystalline, high order regions. The arrangement of cellulose and microfibrils in
plant cell wall is shown in Figure 1. The regions within microfibrils with less order are termed
amorphous. The arrangement of crystalline and amorphous cellulose results in interesting
properties of stiffness and rigidity on one hand and flexibility on the other hand. Moreover, the
structure of cellulose with its hydrogen bond makes it insoluble in most solvents and is partly
responsible for the resistance of cellulose against microbial degradation. The enzymes for cellulose
degradation belong predominantly to hydrolases, which cleave the glycosidic bonds. Cellulose is
hydrolyzed by cellulase (endoglucanase), 1,4-β-cellobiosidase and β-glucosidase.
Figure 1: Arrangement of fibrils, microfibrils and cellulose in plant cell wall

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The enzymes for the degradation of celluloses and hemicelluloses belong predominantly to the
hydrolases which cleave glycosidic bonds by hydrolysis. A cellulases system which hydrolyses
cellulose consists of cellobiohydrolases (exoglucanases, EC 3.2.1.91), endoglucanases (EC 3.2.1.4)
and β-glucosidase (EC 3.2.1.21). The cellobiohydrolase and endoglucanase work synergistically by
hydrolyzing 1,4-β-D-glycosidic linkages in cellulose, cello-oligomers and other β-D-glucans
releasing cellobiose from the non-reducing ends, which are then degraded by β-glucosidase to
glucose units as shown in Figure 2. Different microorganisms can produce variant of each enzyme
group. The presence of the different components of the cellulose enzymatic system in an
appropriate ratio is important in biorefineries to avoid, e.g., the repression of cellobiohydrolases
by the accumulation of cellobiose due to a low activity of β-glucosidases.
Figure 2: Diagram of enzymatic cellulose degradation
2.1.2 Hemicellulose
Hemicelluloses are complex heterogeneous polysaccharides made up of different monomeric
residues, such as D-xylose, D-glucose, D-arabinose, D-mannose and D-glucuronic acid.
Hemicelluloses have a lower degree of polymerization, (DP 50 – 300) compared to cellulose, they
have side chains that can be acetylated and are essentially amorphous. They are classified
according to the monomeric sugar in the backbone of the polymer, e.g. mannan (β-1,4-linked
mannose) or xylan (β-1,4-linked xylose) hemicelluloses. The main chain of glucose and mannose
residues are usually connected with β-(1,4) bond while the side chain is attached to main chain via
α-(1,6) bonds in galactoglucomannan as shown in Figure 3. Xylan can be degraded by endo-1,4-β-
xylanase (EC 3.2.1.8) and 1,4-β-xylosidase (EC 3.2.1.37) to xylose.

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Figure 3: Schematic structure of O-galacto-glucomannan
Due to the more heterogenous nature of hemicellulose compared to cellulose, a complex mixture
of enzymes is required for its degradation, such as endoxylanases, β-xylosidases,
endomannanases, β-mannosidases, α-L-arabinofuranosidases and α-galactosidases. The major
hemicellulose of hardwoods is xylan, whose contents vary from 15 to 35%, while birch wood
contains 22-30% xylan. The enzymatic xylan degradation is shown in Figure 4. The xylan
backbone is hydrolyzed by the endo-1,4-β-xylanase to oligomers, xylobiose and xylose.
Oligosaccharides are then degraded by xylan 1,4-β-xylosidase into xylose units from non-reducing
ends. The side groups, acetyl group and 4-Omethylglucuronic acid that are linked to xylan
backbone are also cleaved by acetylesterase (acetic-ester acetylhydrolase), and xylan α-D-1,2-(4-O-
methyl) glucuronohydrolase respectively. The degradation of hemicellulose is more common in
wood fungi than in bacteria.
Figure 4: Diagram of enzymatic xylan degradation. x xylose residue, Ac acetic acid residue, 4-O-
Me-GA 4-O-methylglucuronic acid residue, ⇑ degradation

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2.1.3 Lignin
The third main constituent in wood, lignin, is a complex macromolecule formed by the
dehydrogenative polymerization of three phenyl propane units (shown in Figure 5) namely
pcoumaryl alcohol, coniferyl alcohol and sinapyl alcohol joined through ether bonds. The
structure of lignin varies widely within species. It provides compressional strength to the cell wall
of plant while cellulose provides the plant with flexible strength. In contrary to cellulose, lignin is
a complex, three-dimensional macromolecule and is highly hydrophobic. It forms an amorphous
complex with hemicelluloses enclosing cellulose and thus prevents the microbial degradation of
accessible carbohydrates within wood cell wall. Moreover, its aromatic rings make it more
difficult to degrade (Schmidt 2006). Lignin also acts as a cementing component to connect cells
and harden the cell wall of xylem which helps the smooth transportation of water from roots to
leaves. Ester linkages occur between the free carboxy group in hemicellulose and the benzyl
groups in lignin, the lignin-carbohydrate complex (LCC), embeds the cellulose thus giving
protection against microbial and chemical degradation (Jeffries 1994). Cellulose and hemicellulose
are rather easily degradable, while lignin is resistant to degradation by most microorganisms due
to its phenylpropane units in the structure and the recalcitrant linkages between them (Schmidt
2006). Thus, there is great interest in finding organisms capable of breaking down lignin and in the
cleaving of the chemical bonds that exist between lignin and hemicelluloses (LCC). These
organisms might be producing enzymes that could be used in the modification of plant biomass.
Figure 5: Building unit of lignin. (A) p-coumaryl alcohol, (B) coniferyl alcohol, (C) sinapyl
alcohol
Lignin can be effectively broken down by a group of filamentous fungi known as white-rot fungi,
which are capable of producing a number of oxidoreductases, enzymes able to attack phenolic
structures. Lignin peroxidase (LiP), manganese peroxidase (MnP) and laccase are among them.

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Lignin peroxidase, LiP, EC 1.11.1.14 is a glycoprotein that contains heme, cleaves C-C bonds and
oxidizes benzyl alcohols to aldehydes or ketones. LiP attacks both phenolic and nonphenolic
lignin substructures by a one-electron oxidation reaction to generate unstable aryl radical cations.
It requires an extracellular hydrogen peroxide, H2O2, as an electron acceptor. LiP initiates
different non-enzymatic reactions such as the cleavage of Cα-Cβ bond in the side chain, β-O-4
bond between side chain and next ring and cleavage of aromatic ring as shown in Figure 6.
Figure 6: Reactions initiated by lignin peroxidase, (1) cleavage of Cα-Cβ bond in the side chain, (2)
β-O-4 bond between side chain and next ring and (3) cleavage of aromatic ring.
Figure 7: Catalytic cycle initiated by Manganese Peroxidase
Manganese peroxidase, MnP, EC 1.11.1.13, is a heme containing glycoprotein, involved in lignin
degradation by mainly attacking phenolic lignin component. The production of MnP is limited to
certain basidiomycetes and some wood decaying white-rot fungi, which secrete MnP in multiple

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forms into their environment. It oxidizes phenolic structures by oxidizing Mn2+ to Mn3+ in the
presence of H2O2. The catalytic cycle of MnP is initiated by binding of H2O2 to the native ferric
enzyme and the formation of iron-peroxide complex (Figure 7). The cleavage of oxygen-oxygen
bond of hydrogen peroxide or other peroxide to one water molecule requires two electron
transfers from the heme, resulting in the formation of MnP Compound I (Fe4+-oxo-porphyrin-
radical complex). MnP Compound I is then reduced to MnP Compound II (Fe4+-oxo-porphyrin
complex) by a monochelated Mn2+, which is oxidized to Mn3+. The native enzyme is then
regenerated from the further reduction of MnP Compound II by Mn2+ and the releasing of
another water molecule. The formed Mn3+, a strong oxidant, then oxidizes phenolic structures by
single electron oxidation.
The third enzyme involved in lignin degradation is laccase, EC 1.10.3.2, in the family of copper-
containing polyphenol oxidases, s also called a blue multicopper oxidase. It catalyzes the
oxidation of a wide range of substrates such as mono-, di- and poly-phenols, aromatic amines and
non-phenolic compound to free radicals and the fourelectron reduction of oxygen directly to
water. Then, the formed free radicals undergo numerous spontaneous reactions which in turn
result in various bond cleavages of aromatic rings and other bonds. However, laccase can’t
directly oxidize all potential substrates either due to the large size of the substrates which restrict
their penetration into the enzyme active site or due to their high redox potential. Thus, it is
required to have suitable chemical mediators that act as an intermediate substrate for laccase,
whose oxidized radical formed are then able to react with the high redox potential substrates.
2.2 Lignocelluloses degrading microorganisms
A wide range of microorganisms including bacteria and fungi are capable of producing cellulases
and hemicellulases but only a limited number of these microorganisms are capable of producing
lignin degrading enzymes. Among these microorganisms, filamentous fungi appear to be the most
efficient producers of enzymes degrading lignocellulose. The ability to produce specific enzymes
for degradation of different carbon and nitrogen sources are due to the diverse habitat they are
found.
White rot fungi, which mostly belong to the Basidiomycota phylum, have the ability to degrade
lignin completely to carbon dioxide and water. They secrete three classes of extracellular
ligninolytic enzymes: a phenol oxidase (laccase), two heme-containing peroxidases (lignin
peroxidase, LiP, and manganese peroxidase, MnP) and oxidases that produce H2O2 needed for

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peroxidase activities. LiP is capable of oxidizing phenolic and nonphenolics lignin substructures
directly while most of laccases and MnP are only capable of oxidizing phenolic substructures.
2.3 Lignocellulosic substrates
Two types of substrates are used in this study as a carbon source for the growth of the
microorganisms: birch wood and wheat bran. Birch is a hardwood tree that belongs to the genus
Betula sp. and is mainly used in pulp, paper industry and in furniture-making. The chemical
composition of birchwood differs from species to species. In general, it is composed of
approximately 14% non-cell wall materials, 42.7% cellulose, 26.4% hemicellulose and 16.9% lignin.
Wheat is a grass belonging to Triticum sp. which is cultivated worldwide mainly as a food crop. It
contains higher protein content than most of the cereals such as maize and rice. Wheat bran is the
hard outer layer of the wheat grain and it is particularly rich in essential fatty acids and dietary
fibers and contains significant amount of proteins, starch, vitamin and minerals. Differences in the
production of ligninolytic enzymes were expected by using different lignocellulosic substrates
which may contain significant concentrations of soluble carbohydrates and inducer of enzyme
expression.

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MATERIALS AND METHODS
Source of lignin-degrading microbes
Samples containing lignin-degrading microbes were taken from (a) Soil mixed with decaying
leaves; (b) Effluent from paper mills. Soil sample was taken 5 cm from the surface.
3.1 Isolation of microorganisms
The samples were screened for the presence of lignin-degrading bacteria and fungi.
3.1.1 Isolation of lignin-degrading bacteria
(i) Media preparation
Lignin-degrading bacteria were screened on the basis of laccase and lignin peroxidase plate assay.
For this nutrient agar medium (Table 1) was prepared and pH adjusted to 7.0. This medium was
complemented with agar 1.5% and autoclaved at 15 psi for 15 min. Autoclaved medium was
poured in sterile petriplates (25 ml/plate) under laminar flow hood and allowed to solidify.
This was followed by even spreading of 2 mM sterile guaiacol (in diethylether), or 0.05% azure B
for Lac and LiP plate assays, respectively.
(ii) Serial dilutions of samples
Soil and effluent samples from various locations were taken and serial dilutions were made. For
this 1 g or 1 ml sample was taken and put in tube containing saline and mixed thoroughly. This
represented 10-1 dilution. Under aseptic conditions, 10-2 to 10-9 dilutions of samples were
prepared.
Table 1: The composition of nutrient agar medium containing guaiacol or azure B

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(iii) Isolation of lignin-degrading bacteria by laccase and lignin peroxidase plate assays
100 µl of 10-6 to 10-9 dilutions were put on nutrient agar plates containing different indicator
compounds like guaicol or azure B for Lac or LiP plate assays, respectively. The samples were
spread evenly over the agar surface with the help of sterilized spreader. The plates were incubated
under inverted position for up to 72 hrs at 37°C. Positive cultures with lignin-degrading abilities
were screened by observing reddish brown zones on plates containing guaiacol (Lac-catalyzed
reaction) and clear zones with white colonies on plates containing azure B (LiP-catalyzed
reaction). The positive strains were subcultured.
3.1.2 Isolation of lignin degrading fungi
(i) Media preparation
Two media were used for isolation of lignin-degrading fungi
• Olga et al. medium (Olga et al. 1993) containing guaiacol for laccase plate assay and azure B for
lignin peroxidase plate assay.
• Crystal violet medium.
The Olga et al. medium (Olga et al. 1993) was prepared and autoclaved at 15 psi for 15 min (Table
2). This was followed by pouring of 25 ml medium per sterile petriplate and even spreading under
sterile conditions of 0.02% guaiacol (in diethylether) and 0.05% azure B on each plate containing
solidified medium.
The crystal violet medium (Swamy and Ramsay 1999; Gochev and Krastanov 2007) was prepared
(Table 3) and autoclaved at 15 psi for 15 min. The autoclaved media was poured in sterile
petriplate (25 ml/plate) and 2.5 µg/ml crystal violet dye was evenly spread over the solidified
medium.
(ii) Serial dilutions of samples
Serial dilution of various soil and effluent samples was done in the same manner as described
above for bacteria.
(iii) Isolation of lignin-degrading fungi by laccase and lignin peroxidase plate assays
100 µl of 10-6 to 10-9 dilutions were put on Olga et al. medium plates containing guaiacol and
azure B, the samples were spread evenly over the agar surface with sterile spreader.

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The plates containing Olga et al. medium were incubated for 7-10 days at 30°C and positive
cultures were screened by observing reddish brown color zones in the presence of guaiacol or
clear zones with white colonies in the presence of azure B.
Table 2: The composition of olga et al. medium containing guaiacol or azure B
Table 3: The composition of crystal violet medium
(iv) Isolation of lignin-degrading fungi by crystal violet medium
100 µl of 10-6 to 10-9 dilutions were put on separate plates containing crystal violet medium, and
the samples were spread evenly over the agar surface with sterile spreader.
The plates containing crystal violet medium were incubated for 15 days at 30°C and positive
cultures were screened by observing yellow green zones.

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(v) Storage of microbes
The microbes screened were stored as glycerol stocks at -80°C.
3.2 Morphological and biochemical characterization of lignin-degrading microorganisms
All the bacterial and fungal isolates tested positive in Lac and LiP plate assays were subjected to
morphological and biochemical characterization. All the experiments were performed in triplicate.
3.2.1Morphological characterization by microscopic examination
The purity and genuine identification of the isolated lignin-degrading bacteria were done by
Gram staining procedure and that of fungi by analyses of colony morphology on Sabouraud agar
medium and lactophenol cotton blue staining procedure.
3.2.1.1 Identification of bacteria by Gram staining procedure
The cell shape, structure, Gram positive and Gram negative behavior of bacteria screened by plate
assays were determined by Gram staining technique. Bacteria were smeared on slide and stained
with crystal violet for 1 to 2 min. The slide is then flooded with iodine for another 1 to 2 min. The
iodine was then poured off. The process was followed by decolorization process using acetone for
2 to 3 sec, and immediate washing with water. The slide was flooded with safranin counterstain
for 2 min and washed with water. The slide was air-dried and examined under the oil immersion.
Grampositive bacteria did not decolorize by the decolorization process and remained purple.
Gram-negative bacteria on the other hand were counterstained by safranin resulting in pink
coloration.
3.2.1.2 Identification of fungi on the basis of colony morphology
Sabouraud agar medium (Table 4) plates were prepared and inoculated with fungal cultures. The
plates were incubated at 25°C in inverted position and the growth patterns of different fungi were
analyzed after 5 to 7 days.

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Table 4: The composition of sabouraud agar medium
For microscopic examination, coverslips were inserted in the media in tilted position over the
fungi growing on Sabouraud agar medium plates. After growth, coverslips were lifted gently with
sterile forceps and placed over clean slides containing a drop of lactophenol cotton blue with the
fungi facing side down. The stained cultures were visualized and photographed under dissecting
microscope at 40X magnification.
3.2.2 Biochemical characterization of lignin-degrading microbes
All the bacterial and fungal isolates tested positive in Lac and LiP plate assay were subjected to
qualitative biochemical characterization by performing catalase production test, motility test,
methyl-red test (MR), indole test, nitrate reduction and gas production. Similarly, fungi were
analyzed qualitatively for their starch and cellulose hydrolyzing test.
3.2.2.1 Biochemical characterization of lignin-degrading bacteria
(i) Catalase production test
Catalase test was performed by addition of 3% hydrogen peroxide on the slide having few drops
of bacterial culture. The production of effervescence (bubbles) is indicative of positive catalase
reaction.
(ii) Motility test
A loopfull bacterial culture was added in the center of the tube containing nutrient media (agar
1%) and incubated in anaerobic incubator at 37°C for 72 hrs. After incubation, bacterial growth
pattern was observed.

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(iii) Methyl-red test (MR)
The isolates were incubated at 37°C for 72 hrs in a buffered glucose broth (Table 5). After
incubation, few drops of methyl red solution were added to the culture and read immediately for
change in color. Red color is indicative of positive MR test.
Table 5: The composition of buffered glucose broth
(iv) Indole test
The isolates were incubated at 37°C for 72 hrs in tryptone water. After 48 hrs, 1 ml Kovac’s reagent
was added to the culture and observed for a pink color. Bright pink color in the top layer is an
indication of positive indole test.
(v) Nitrate reduction test
The isolates were incubated at 37°C for 72 hrs in nitrate reduction broth (Table 6). After
incubation, 1 ml each of sulphanilic acid (0.8% in 5N acetic acid) and α-naphthylamine (0.5% in 5N
acetic acid) was added into the tubes. The appearance of red color is an indication of the positive
test for nitrate reduction test.
Table 6: The composition of nitrate reduction broth

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3.2.2.2 Biochemical characterization of lignin-degrading fungi
(i) Starch hydrolysis test
Starch agar medium (Table 7) was inoculated with isolated fungal cultures. The plates were
incubated at 25°C in inverted position for 5 to 7 days. The surface of the plates was flooded with
the iodine solution for 30 sec. Examine the disappearance of starch from the starch agar media
plates.
Table 7: The composition of starch agar medium
(ii) Cellulose hydrolysis test
Czapek-mineral salt agar medium (Table 8) was inoculated with isolated fungal cultures. The
plates were incubated at 35°C in inverted position for 2 to 5 days. The surface of the plates was
flooded with 1% aqueous solution of hexadecyltrimethyl ammonium bromide for 30 sec. The
plates were observed for the formation of a clear zone around the fungal growth.
Table 8: The composition of czapek-mineral salt agar medium
3.3 Methods used for analysis of lignin-degrading enzyme activities
Before analysis of lignin degrading enzyme activities, all the bacterial and fungal isolates tested
positive in plate assay were subjected to analysis of protein content.

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3.3.1 Analysis of protein content from bacteria
Bacterial colonies tested positive in plate assay were inoculated in nutrient broth medium (Table
9), poured in test tubes (10 ml/tube) and autoclaved at 15 psi for 15 min. The tubes were
incubated in incubator shaker at 120 rpm at 37°C for overnight.
Table 9: The composition of nutrient broth medium
Ten ml of above-grown bacterial culture was taken and filtered through Whatman no. 1 filter
paper. This was considered as crude protein or enzyme sample.
This sample was then subjected to Lowry’s method for the determination of protein content. 500
µl of bacterial culture was taken in microfuge tube and protein content was precipitated with
equal volume of ice-cold 20% trichloroacetic acid (TCA) and kept at 4°C overnight. The pellet was
recovered by centrifuging at 12,000 rpm for 5 min at room temperature and decanting the
supernatant. The pellet was washed with 0.1 ml icecold 10% TCA and ice-cold acetone. Depending
on the pellet size, it was dissolved in 0.5-1.0 ml of 0.1 N NaOH. The solution was subjected to
heating for 5 min in boiling water bath and vortexed vigorously. This was considered as protein
solution and the protein content was determined by Lowry’s method (Lowry et al. 1951).
For protein content determination, 0.5 ml of protein solution was taken in test tube and 2.5 ml of
alkaline solution [prepared by mixing 2% Na2CO3 solution (in NaOH), 2% sodium potassium
tartrate and 1% CuSO4.5H2O in 100:1:1] was added. The contents were mixed well and the tubes
were incubated at room temperature for 10 min. This was followed by addition of 0.25 ml of 1.0 N
Folin’s reagent. The contents in the tube were mixed thoroughly and after 10 min, absorbance at
660 nm against reagent blank was determined spectrophotometrically using bovine serum
albumin fraction V (BSA) standard. The protein concentration was determined using Lowry
standard curve (absorbance vs BSA concentrations).

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3.3.2 Analysis of lignin-degrading enzyme activities from bacterial isolates
All the bacterial isolates tested positive in plate assay were subjected to analyses of activities of
lignin-degrading enzymes, viz., Lac, LiP and MnP.
The composition of bacterial growth medium, incubation conditions and enzyme preparation
have already been described.
3.3.2.1Determination of laccase enzyme activity
Laccase activity was determined by monitoring the oxidation of ABTS (2,2'-azinobis-
3ethylbenzothiozoline-6-sulfonic acid) to blue coloration (Poppius-Levlin et al. 1997). Enzyme
samples were analyzed spectrophotometrically for Lac activity according to the protocol described
by Li et al. (2008). The reaction mixture comprised (total 3.0 ml) of 50 mM sodium tartrate buffer
(pH 4.5), 0.5 mM ABTS, 2.1 ml distilled water, 1.0 ml enzyme supernatant and 1.1 ml distilled
water. The contents were mixed well and the oxidation of ABTS was measured by an increase in
absorbance at 420 nm (extinction coefficient 36 x 10-4 mM-1cm-1) for 1 min against reagent blank.
One unit of laccase activity was defined as activity of an enzyme that catalyzes the conversion of 1
mole of ABTS per minute. The substrate-containing buffer, without supernatant, was used as a
standard. The experiment was performed in triplicates.
3.3.2.2 Lignin peroxidase enzyme assay
LiP enzyme activity was determined spectrophotometrically according to the protocol described
by Archibald (1992). Reaction mixture (3.0 ml) was prepared by mixing 50 mM sodium tartrate
buffer (pH 4.5), 100 µM Azure B, 0.1 mM H2O2, 1.0 ml enzyme and 0.88 ml water. The increase in
absorbance at 651 nm (millimolar extinction coefficient 48.8 mM) was recorded for 1 min against
reagent blank. One unit (U) of LiP activity was defined as activity of an enzyme that catalyzes the
conversion of 1 µmole of azure B per minute. The substrate-containing buffer, without
supernatant, was used as a standard. The experiment was performed in triplicates.
3.3.2.3 Analysis of manganese peroxidase enzyme activity
Enzyme samples were subjected to spectrophotometric analysis of MnP activity according to the
protocol described by Kuwahara et al. (1984). Experimental cuvette was prepared by adding 50
mM sodium tartrate buffer (pH 4.5), 0.0025% phenol red, 0.2 mM MnSO4, 0.1 mM H2O2, 1.0 ml
enzyme preparation and 1.57 ml distilled water. The contents were mixed well and increase in
absorbance was observed for 1 min at 520 nm (extinction coefficient 22 mM-1 cm-1) against

24. 24 | P a g e
reagent blank. Manganese-dependent activity was calculated by subtracting the absorbance in the
absence of MnSO4. The experiment was performed in triplicates. The absorbance was measured in
1 min intervals after addition of hydrogen peroxide. One unit of MnP activity was defined as
activity of an enzyme that catalyzes the conversion of 1 µmole of phenol red per minute.
3.3.3 Analysis of lignin-degrading enzyme activity from various fungal isolates
All the positive lignin-degrading fungal isolates from plate assay were subjected to analyses of
activities of lignin-degrading enzymes, viz., Lac, LiP and MnP.
3.3.3.1 Medium preparation for fungal culture
Inoculum from fungal colonies tested positive in Olga et al. plate assay were inoculated in LMM
broth medium (Table 10). The medium was poured in flask (50 ml/flask) and autoclaved at 15 psi
for 15 min. The flasks were incubated at 120 rpm, 25°C for one week.
3.3.3.2 Determination of Lac, LiP, MnP enzyme activities
Fifty milliliter of above-grown culture was taken and filtered through Whatman no. 1 filter paper.
The filtrate was preserved for the analyses of lignin-degrading enzyme activities and protein
content. The three lignin-degrading enzyme activities were determined as described.
Table 10: The composition of LMM broth

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2. RESULTS AND DISCUSSION
The source was a decaying wood from the garden with a black fungi growing over it.
Figure 8: Decaying wood
The nutrient agar medium was prepared to isolate lignin degrading bacteria. It was autoclaved and allowed
to cool down.
Figure 9: Nutrient agar
After incubation , growth was observed on the plates with serial dilution 10-5 and 10-7 which were labeled A
and B respectively.
Figure 10: plate with 10-5 dilution Figure 11: plate with 10-7 dilution

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The results of subcultured nutrient agar plates after streaking were observed as below.
Figure 12: Isolates obtained on plate 1 Figure 13: Isolates obtained on plate 2
Figure 14: Isolates obtained on plate 3
These isolates were then subjected to biochemical tests, the results of which are listed below.The first test
performed was catalase test, the colonies showing effervescence were positive while others tested as
negative.
Figure 15: H2O2 bubbles produced by colony 10

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Colony No.
Catalase test
Positive Negative
1 Negative
2 Positive
3 Negative
4 Positive
5 Negative
6 Positive
7 Negative
8 Positive
9 Negative
10 Positive
11 Negative
12 Positive
Table 6: Results of Catalase test
The results of indole test was negative for all the colonies.
The nitrate reduction broth test was performed on all the 12 colonies. Colonies giving bright pink color were
positive, others negative.
Figure 16(A) : Result of Nitrate reduction broth test for colonies 1,2,3,4,5,6

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Figure 16(B) : Result of Nitrate reduction broth test for colonies 7,8,9,10,11,12
Colony No. Nitrate reduction broth test
1 Positive
2 Positive
3 Negative
4 Negative
5 Positive
6 Positive
7 Negative
8 Negative
9 Positive
10 Positive
11 Negative
12 Negative
Table 7: Results of Nitrate reduction broth test

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CONCLUSION
Lignocellulolytic microorganisms, especially fungi, have attracted a great deal of interest as
potential biomass degraders for large-scale applications due to their ability to produce vast
amounts of extracellular lignocellulolytic enzymes. Lignin, the most recalcitrant component of
lignocellulosic material, acts as a barrier for any solutions or enzymes by linking to both
hemicelluloses and celluloses and prevents penetration of lignocellulolytic enzymes to the interior
lignocellulosic structure. It is primarily basidiomycetes white-rot fungi that are responsible for
efficient lignin degradation in wood decay processes. Their production of extracellular enzymes
known as lignin-modifying enzymes (LMEs) facilitates the degradation process. Based on their
activity on lignin and other lignocellulosic materials, white-rot fungi are categorized into two
groups, simultaneous and selective degraders. Selective lignin degraders white-rot fungi are most
attractive for their potential biotechnological applications in removing lignin, as in biopulping
processes, and for providing an unprotected carbohydrate for subsequent use, as in animal feed
and/or biofuel substrate. LMEs include mainly two ligninolytic enzyme families; i) phenol
oxidase (laccase) and ii) heme peroxidases (LiP, MnP and VP). LMEs, especially VPs require more
research to understand the efficiency of the enzymes on lignin oligomers and the mechanisms of
their action. In addition, accessory enzymes, such as oxidases that generate the H2O2 required by
peroxidases, have been found to be involved in lignin degradation.

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