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Shrivastava, Rohit S., 2005, “Enzymology of Degradation of Xenobiotic
Compounds by Lignin-Degrading Fungi”, thesis PhD, Saurashtra University
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© The Author
A Thesis
Submitted for the
Degree of
Doctor of Philosophy
In
Microbiology
ENZYMOLOGY OF
DEGRADATION OF XENOBIOTIC
COMPOUNDS BY
LIGNIN- DEGRADING F UNGI
Rohit S hrivastava
Registration No: 2594 Date: June 12, 2001
Department of Bioscience s, Saurashtra University,
Rajkot 360 005 , Gujarat, India.
January, 2005
Think no more, lad; laugh, be jolly:
Why should men make haste to die?
Empty heads and tongues a -talking
Make the rough road easy walking,
And the feather pate of folly
Bears the falling sky.
Oh, 'tis jesting, dancing, drinking
Spins the heavy world around.
If young hearts were not so clever,
Oh, they would be young for ever:
Think no more; 'tis only thinking
Lays lads underground.
A. E. Housman
A Shropshire Lad, (1896)
Dedicated to . . . . .
. . . . . . My Beloved Parents
SAURASHTRA UNIVERSITY
DEPARTMENT OF BIOSCIENCES
University Campus, RAJKOT – 360 005, Gujarat, India
Phone: (O) +91-281–2586419, (R) 2444295 Fax : 2586419
E-mail : brmvyas@hotmail.com
Dr. B.R.M. Vyas
Assistant Professor
Date: January , 2005
Ref: SU/BIO/
C ERTIFICATE
I take pleasure in forwarding the thesis entitled “Enzymology of
Degradation of Xenobiotic Compounds by Lignin-degrading
Fungi” of Mr. Rohit Shrivastava for the acceptance of the degree
of Doctor of Philosophy in Microbiology. Thesis presented here
embodies a record of the results of original investigation carried
out by him.
(B. R. M. Vyas)
Forwarded Through:
Dr. S. P. Singh
Professor & Head
Dept. of Biosciences
Saurashtra University
A CKNOWLEDGEMENTS
“Acknowledgement” – This is where one admits, the imminence,
involvement and dedication of others to your work. It is indeed a moment
of great pleasure to mention the names of those wonderful persons who
walked-along till my thesis.
Firstly, my Supervisor, Dr. B.R.M. Vyas, a niche of its own, whose
enthusiastic cry of “Any Progress” will haunt me for the rest of my days
- his unquenchable curiosity and love for the subject are probably the
most valuable lessons I have learnt from this PhD. I feel sacred, for He
provided the creative and stimulating atmosphere that kept me going over
the last four years and lent a hand to shape-up my research to the existing
dimension.
I am grateful to Prof. S.P. Singh ( Head, Department of Biosciences) and
Prof. P.P. Sood (Former Head, Department of Biosciences) for providing
me the laboratory, needed facilities and for extending overwhelming
enthusiasm during my course of research . Their ideology and approach
helped to prepare me for life, both Inside & Outside my profession.
I pay my sincere gratitude to Dr. S.J. Pathak and Dr. S.C. Bhatt for their
continual assistance, encouragement and whole-hearted support for
whenever I needed during the investigation.
My thanks are due to all the Non-Teaching Staff, Parekhbhai, Rajubhai,
Zalabhai, Thakrarbhai & Nitesh for their kind co-operation.
I gratefully acknowledge Department of Biotechnology
(DBT), Ministry
of Science & Technology, India, for the financial support for my Senior
Research Fellowship.
I salute my Guru-of-the-Field, Dr. P.S. Nagar (whom we use d-to call
“Baba” with love), sagacity personified. The first harvest with him taught
me the real importance of other living things, the work outside the fourwalls and to respect the work for them.
I gratefully acknowledge my Laboratory-Colleagues Pinalee, Jagruti,
Dharmendra , Padma & Charmy for their help that rendered the
completion of investigation.
No phrases can fully express my gratitude to My-Dear-Labmates
Kalpesh ( Batti), Vimesh (Vimli), & Dr. Rajendra (Chacha) for their
extraordinary support, especially during my experiments when we used to
move like Night-Hawkers to stay awaken through -out the completion of
experiment.
I owe my sincere debt to my
Small-F amily, Viral (Viraliya), Sudipta
(Misra), Mital (Mitu), & Rupal (Rupali) for realizing me, what it means to
be, home away from home that made my life, A –Phenomenon, at Rajkot.
The “Luncheon” was always exotic for the meals and the moments
shared. I run short of words to express my hearty-thanks for their selfless
dedication, untiring efforts and magical-warm-caress that made the rough
road easy walking en-route to the final milestone.
Thanks to my Friends, Nikhil
(Pange), Rahul (Baccha), Ajay
(Chowdhary), Nirav (Nirva), & Ranjeet (Paaji), for their invariable propup in organizing my administrative work at Ahmedabad.
Finally, my deepest gratitude with homage to my Loving-Family,
Dr. S.M.
Shrivastava (Papa), Mrs. Punam Shrivastava (Ma’) & Ms. Nupur
Shrivastava (Pinky),
who have been great over the years and never raised an
eyebrow when I claimed my thesis, would be finished in the “next two months” for
nearly a year.. .. .. .. .. .. ..
Rohit Shrivastava
C ONTENTS
Page No.
Chapter 1. Introduction
1-25
(A) White-Rot Basidiomycetes: Lignin Degradation
1-14
(B) White-Rot Basidiomycetes: Degradation of Dyes
15-25
Chapter 2. Screening of White-Rot Basidiomycetes: Decolorization
26-32
of Textile dyes
2.1 Introduction
2.2 Materials and methods
26
27
2.3 Results and discussion
28-32
Chapter 3 . Ligninolytic Enzymes of Irpex lacteus: Decolorization
33-41
of Textile and Triphenyl methane Dyes
3.1 Introduction
33
3.2 Materials and methods
3.3 Results and discussion
34-35
35-41
Chapter 4. Enzymatic decolorization of Sulfonphthalein Dyes
42-52
4.1 Introduction
4.2 Materials and methods
42
43-44
4.3 Results and discussion
44-52
Chapter 5. Horseradish Peroxidase: Decolorization of Textile And
53-59
Triphenyl methane Dyes
5.1 Introduction
53
5.2 Materials and methods
5.3 Results and discussion
54
54-59
Chapter 6. Ligninolytic Enzymes of Irpex lacteus : Decolorization
60-67
of Effluents
6.1 Introduction
60
6.2 Materials and methods
6.3 Results and discussion
61-62
62-67
Chapter 7. Decolorization of D yes and Effluents by Fenton
68-75
and Fenton-like R eagent
7.1 Introduction
7.2 Materials and methods
68
69
7.3 Results and discussion
70-75
BIBLIOGRAPHY
IMPLICATIONS AND SPECULATIONS OF
INVESTIGATIONS
APPENDIX
76-95
96-99
100-106
I.
Scholarships awarded
100
II.
List of Publications & Presentations
101-104
III. Conferences / Seminars / Workshops attended
105-106
CHAPTER 1a
WHITE-R OT BASIDIOMYCETES:
D EGRADATION OF LIGNIN
L
ignum in Latin means wood. It is found in higher plants,
including ferns, but not in liverworts, mosses and plants of
lower taxa. Lignin differs from plant to plant, in different parts
of plant and in different age groups of plant. Lignin is
therefore not a single compound but a group of compounds. It is estimated
that our planet contains 3x10 11 metric tones of lignin with an annual
biosynthesis rate of approximately 30% of the dry weight of softwoods and
20% of the weight of hardwoods. Lignification is associated with the
development of vascular system in plants.
Lignin biosynthesis: Lignin is built from phenyl propane units linked
together by several different inter-unit linkages having bio logically stable
structure. Biosynthesis of lignin initiates with the fixation of CO2 by plants
leading to the generation of phenylalanine from glucose via shikimic acid
pathway (1). Lignification proceeds with the enzymatic conversion of
phenylalanine to cinnamic acid which is hydroxylated to coumaric acid and
caffeic acid. Methylation of caffeic acid sequentially yields ferulic acid and
sinnapic acid. The synthesis of lignin precursors completes with the reduction
of coumaric acid, ferulic acid, and sinnapic acid to their respective alcohols
(Figure 1.1).
CH2OH
CH2OH
CH2OH
C
C
C
C
C
C
OH
Coumaryl
alcohol
OH
Coniferyl
alcohol
OCH3
OCH3
OCH3
OH
Sinnapyl
alcohol
Figure 1.1 Precursors of lignin biosynthesis
1
These monolignols are of low solubility and are stabilized in cell wall as their
glucosides.
The
polymerization
of
monolignols
initiates
with
the
dehydrogenation catalyzed by peroxidase-H2O2 or laccase-O2 system
generating
spontaneous
resonance
and
stabilized
random
radicals
polymerization
(1).
The
reactions
of
non-enzymatic,
enzymatically-
generated radicals completes the three dimensional network of lignin.
Lignin structure and its role: The structural and physico-chemical
properties (Table 1.1) make lignin a highly recalcitrant polymer that resists
degradation in nature.
Lignin is a natural bio-plastic that (i) provides mechanical rigidity and
strength to stem, (ii) water proofs cell wall and thereby reduces water-loss
occurring through evaporation, and also enables transport through xylem and
phloem, (iii) protects plants against UV-rays due to its ability to absorb UVradiations, (iv) protects plants against microbial attack, and (v) plays an
important role in healing as injury to plant induces lignification.
Table 1.1 Physical and chemical properties of lignin
Physical Properties
Water insoluble
High molecular weight
Highly branched 3D
structure
Compact
Random
Optically inactive
Stereo-irregular
Chemical Properties
Highly reduced form of carbon
Presence of chiral C -atoms
Linkages
Ø Variety of linkages
Ø Non-hydrolysable ether bonds (β-o-4, α-o-4, etc.)
Ø Multiple linkages (phenylcoumaran,
pinoresinol, etc.)
Ø Randomly distributed linkages
Various microbial groups possess the ability to degrade lignin but white
rot basidiomycetes comprise the only group of organisms known to degrade
lignin completely (2). The degradation of lignin by white rot basidiomycetes
has several unusual features which are desc ribed below.
Feature I: Lignin degradation is part of secondary metabolism:
Lignin degradation occurs only after primary growth is complete i.e. during
secondary metabolism. Production of ligninolytic enzymes in the cultures of
white rot fungi occur when the primary growth ceases in response to limitation
2
of nutrient C / N / SO4 but not PO4. Shift to secondary metabolism occurs to
accommodate the limitation (3). Secondary metabolic phase is characterized
by the production of secondary metabolites (e.g. antibiotics), a series of
developmental changes (sporulation) and demise of the mycelium (decline
phase) (4-6). Ligninolytic enzyme system is associated with the secondary
and perhaps decline phase. White-rot fungi do not use lignin as a sole source
of carbon and energy because lignin degradation does not occur during
primary growth. Little lignin carbon enters central metabolic pathway and is
evident from the regulatory features of the ligninolytic system that also
suggest that fungi do not depend on lignin as an important carbon and energy
substrate (6, 7).
Feature II: The ligninolytic system is not induced by lignin:
Cultures of white rot fungi produce ligninolytic enzymes in response to nutrient
C or N or SO4 limitation irrespective of the presence or absence of lignin (6).
Ligninolytic activity in the cultures does not increase in the presence of lignin.
Ligninolytic system is induced by molecular oxygen. Higher concentrations of
molecular oxygen increase the titer and shorten the time of appearance of
ligninolytic activity (8-11).
Feature III: Lignin polymer degradation involves oxidative
reactions: Degradation of lignin polymer initiates by oxidation occurs (8).
Presence of molecular oxygen is necessary for the degradation of lignin. A
subsequent increas e in the oxygen content and decrease in hydrogen content
of partially degraded lignin polymer provides strong proof of the oxidative
nature of lignin degradation (12, 13). Introduction of hydroxyl group at C? of
the side chains and cleavage of aromatic rin gs bound to lignin explains its
degradation as oxidative. Products of lignin degradation are largely watersoluble and carbonyl rich (14). Ligninolytic enzyme system consists of
peroxidases, oxidases, reductases, and dehydrogenases and the reactions
catalyzed by them are largely oxidative and charge-transfer reactions
including reductive.
Feature IV: The ligninolytic system is non-specific: Despite the
presence of stereo-irregular and heterogeneous structure, lignin is degraded.
Ligninolytic system degrades lignins of different compositions i.e. of grasses,
3
hard-woods, soft-woods, etc. The non specificity of system is evidenced from
the degradation of modified lignins like Kraft lignin, DHP (polyguaiacol), etc
(15).
Structurally
diverse
xenobiotic
compounds
like,
polyaromatic
hydrocarbons (PAH), (16-18), nitroaromatics and explosives (19, 20),
polychlorinated biphenyls (18, 21), and dyes (22-26) are degraded by the
ligninolytic cultures.
Feature V: Depolymerization is not an obligatory initial step in
lignin degradation: Degradation can also be initiated by modification of lignin
polymer (12). Degraded lignins exhibit substantial oxidative changes. C 9subunits in the lignin polymer undergo demeth(ox)ylation, side-chain oxidation
and ring cleavage reactions (13, 27). Some of these reactions may cause
partial depolymerization. This leads to the generation of low molecular weight
lignin degradation products suggesting that all aromatic nuclei are not cleaved
while bound in the polymer (28). Thus, lignin degradation can be initiated by
any of the two ways; lignin polymer modification and lignin depolymerization.
Ligninolytic System: Ligninolytic system is extracellular, highly nonspecific and oxidative in nature. It comprises of enzymatic and non-enzymatic
components (Table 1.2.)
Table 1.2. Composition of ligninolytic system
Enzymatic components
Non-Enzymatic components
Peroxidases
Manganese peroxidase
Lignin peroxidase
Versatile peroxidase
RBBR oxygenase
Oxidases
Laccase
Aryl alcohol oxidase
Cellobiose oxidoreductase
Dehydrogenases
Cellobiose dehydrogenase
Pyranose 2-dehydrogenase
H2O2-generating oxidases
Pyranose 2-oxidase
Glyoxal oxidase
Aryl alcohol oxidase
Reductases
Quinone reductases
4
Reactive oxygen species
Veratryl alcohol
Slime
MnII
Quinones
Organic acids
H2O2
Enzymatic components of ligninolytic system
Manganese peroxidase: Manganese peroxidase (E.C.1.11.13) was
discovered in Phanerochaete chrys osporium (29, 30). It is glycosylated,
haem-containing protein with a large isozymes family. The molecular weight is
in the range of 42-46 kDa (31). One mole of iron protoporphyrin IX per mole of
enzyme is present. The family of isozymes have identical catalytic properties
but vary in their kinetic constants, substrate specificities, isoelectric point,
carbohydrate content and amino acid composition.
The catalytic cycle of manganese peroxidase resembles to that of the
classical peroxidase, horseradish peroxidase. In the presence of H2O2, native
enzyme is converted to a 2e ¯ deficient compound I. Compound I oxidizes Mn II
to MnIII and gets converted to compound II which in turn oxidizes another Mn II
to return back to native state (Fig 1.2). The conversion of compound I to
compound II is also facilitated by phenolic substrates but the conversion of
compound II to native enzyme occurs only in the presence of MnII .
FERRIC ENZYME
H202
H20
Mn III
COMPOUND III
COMPOUND I
MnII
MnII
H20
excess H202
MnIII
COMPOUND II
Figure 1.2 Catalytic cycle of manganese peroxidase
Fe IV=O centre is partially buried which is accessible to small
compounds like Mn II but not organic substrates. Therefore, the completion of
catalytic cycle of manganese peroxidase necessitates the presence of Mn II
(32).
5
MnIII alone and chelates of Mn III with organic acids (oxalate, malonate,
malate etc.), can diffuse from their site of generation and cause one electron
oxidation of various phenolic compounds, amino aromatic compounds and
low redox potential non-phenolic compounds via hydrogen abstraction
generating corresponding radicals (32).
reductant
MnP( I / II )
MnII
HQ•
MnP( II / N )
MnIII
H2Q
Assist in maintaining H2Q pool
for MnP-catalyzed
reduction reactions
recycling of quinones
Substrate
Q
Substrate reduced
Cellobiose dehydrogenase
Pyranose-2-dehydrogenase
Cellobiose oxidoreductase
Quinone reductase
Figure 1.3 Reduction reactions catalyzed by manganese peroxidase
Manganese peroxidase can also catalyze reduction reactions (Fig 1.3).
Reduced quinones (H 2Q) are generated by various dehydrogenases (33) and
reductases secreted in the ligninolytic pool of enzymes. Mn III can oxidise
these reduced quinones to 1e ¯ oxidised quinones (HQ•). These 1e¯ oxidised
quinones are than stabilized by getting further oxidized to native quinones,
simultaneously, reducing range of other oxidised chemicals. The radical
(oxidants
and
reductants)
generating
characteristics
of
manganese
peroxidase underline its potency in bioremediation, to bring about the
degradation of xenobionts.
Lignin peroxidase : Lignin peroxidase (E.C.1.11.14) (34, 35) is a
heme-containing glycoprotein with a molecular weight of 41-42 kDa. It is
optimally active, but not stable at low pH (pH 2) (36). Lignin peroxidase is
represented by a family of isozymes that vary with strains and cultural
conditions (37).
6
The native enzyme reacts with H2O2 to generate compound I, a two
electron oxidized intermediate, which is converted to one electron-oxidized
com pound II via oxidation of veratryl alcohol (VA) generating VA cation radical
VA+•. VA+• can undergo a variety of reactions and can mediate oxidation
reactions. Compound II returns to native state by oxidizing another molecule
of VA to complete its catalytic cycle (Fig 1.4). In the presence of excess of
H2O 2, compound II can be converted to compound III, an oxyperoxide, which
is an inactive form of enzyme (38).
FERRIC ENZYME
VA
VA +•
H202
H 20
VA+•
COMPOUND III
COMPOUND I
VA
VA
H20
excess H202
VA +•
COMPOUND II
Figure 1.4 Catalytic cycle of lignin peroxidase
Lignin peroxidase can also catalyze reductions using secondary
metabolites that are produced by the fungus (Fig 1.5). The VA+• generated by
lignin peroxidase can oxidize oxalic acid producing oxalate anion radical
which in turn can reduce variety of chemicals (39). Oxalate and VA act as
redox mediators in lignin peroxidase catalyzed reduction reactions. The redox
reactions that lignin peroxidases can catalyze stress their potential in
bioremediation program.
LP( I / II )
LP( II / N )
VA
anion radical
CO 2
+
CO2 ¯ •
Substrate
CO 2
Substrate reduced
Organic
acid
Figure 1.5 Reduction reactions catalyzed by LP
VA+•
7
Versatile peroxidase : Versatile peroxidase, another novel peroxidase,
has recently been reported from the cultures of Pleurotus eryngii and
Bjerkandera adusta (40, 41). Versatile peroxidase catalyzes the reactions
catalyzed by both the ligninolytic fungal peroxidases, namely, manganese
peroxidase and lignin peroxidase. It has an ability to oxidize the substrates of
manganese peroxidase and lignin peroxidase. It oxidizes Mn II to Mn III and
veratryl alcohol to its cation radical. It also oxidizes hydroquinone and
substituted phenols, substrates of plant peroxidase and is able to degrade
lignin model dimers veratryl glycerol β-guaiacyl ethers (42). Versatile
peroxidase, produced by Pleurotus eryngii, can be considered as third type of
ligninolytic peroxidase.
RBBR Oxygenase: Pleurotus ostreatus during solid state fermentation
of wheat straw produced a H2O2-dependent enzymatic
activity that
decolorized Remazol brilliant blue R (26). This activity is different from
manganese peroxidase, laccase, and veratryl alcohol oxidase produced by
the fungus and from lignin peroxidase which is not produced by Pleurotus
ostreatus but are well known in other white rot fungi.
Laccase : Laccase (E.C.1.10.3.2) is a blue copper oxidase. It is a
glycoprotein having molecular weight of 60-80 kDa. It is capable of oxidizing
electron rich phenolic substrates. Laccase contains multiple copper atoms
which are reduced as substrates are oxidized. After four electrons are
received, laccase converts molecular oxygen to water (43). Laccase catalyzes
one electron abstraction from mono- and poly-phenolics and aromatic amines,
to generate free radicals capable of undergoing further depolymerizatioin,
repolymerization, demethylation and quinone formation (44). A new laccase,
yellow laccase has been discovered recently in the cultures of Panus tigrinus
and Phlebia radiata capable of oxidizing non-phenolic lignin m odels and
veratryl alcohol in the absence of any mediator (45). Substrate specificities of
laccase can be further expanded by adding redox mediator like 2,2’-azinobis(3-ethylbenzothiozoline-6-sulphate) but there is no report whether such
8
mediators function in vivo in lignin degradation, and thus demand further
research to elucidate the actual role of laccase (46).
Aryl alcohol oxidase: Aryl alcohol oxidase (E.C.1.1.3.7) was first
detected by Farmer et.al (1960) (47). Aryl alcohol oxidases are flavo -proteins
containing one molecule of FAD per molecule of enzyme. Aryl alcohol oxidase
has wide substrate specificity towards differently substituted methoxy phenol.
Aryl alcohol oxidase catalyzes the oxidation of aryl α and α−β-unsaturated-γalcohols to their respective aldehydes with simultaneous reduction of O2 to
H2O 2. It has been proposed that aryl alcohol oxidase and intracellular NADPH
dependent aryl alcohol dehydrogenases could constitute a redox system
involving aryl alcohol/ aldehydes produced in order to ensure a steady state
concentration of H2O2 for ligninolytic activities (48).
Glyoxal oxidase: Glyoxal oxidase (E.C.1.2.3.5) yields three molecules
of hydrogen peroxide from glyceraldehydes generated by lignin peroxidase
from Cα-Cβ cleavage of aryl glycerol-α -aryl ethers (49). The various substrates
for glyoxal oxidase are methyl glyoxal, glycolaldehyde, acetaldehyde,
formaldehyde, glyoxal, glyoxalic acid, dihydroxyacetone and glyceraldehydes.
Thus, the mutual working of glyoxal oxidase and lignin peroxidase leads to
controlled radical formation and limitation of radical coupling, driving lignin
oxidation towards depolymerization.
Pyranose 2-oxidase: Pyranose 2-oxidase (E.C.1.1.3.10) is homotetrameric protein that contains covalently bonded flavin adenine dinucleotide
(FAD) (50). It is localized in periplasmic space, but during autolysis it is
localized extracellularly. Pyranose oxidases are widely distributed among
wood
degrading
basidiomycetes
and
catalyze
the
conversions
of
aldopyranoses to corresponding 2-keto aldoses like glucose, galactose and
xylose to 2-ketoglucose, 2-ketogalactose and 2-ketoxylose (51). During this
conversion, the oxidation reaction, electrons are transferred to molecular
oxygen resulting in the formation of hydrogen peroxide; a must needed
substrate for ligninolytic peroxidases.
9
Glucose oxidase: Glucose oxidase (E.C.1.1.3.4) while oxidising
glucose, generates H2O2
from molecular oxygen (52). It also reduces
quinones and phenoxy radicals generated by la ccase during oxidation of
lignin (53). The excess of quinones inhibit laccase activity. Thus, glucose
oxidase, in turn acts as a regulator of enzymes involved in transformation of
lignocelluloses by reducing the quinones and by producing H 2O2.
Cellobiose
dehydrogenase:
Cellobiose
dehydrogenase
(E.C.1.1.99.18) is a high molecular weight, 90 kDa, flavo -hemo-protein. It is
having two prosthetic groups, FAD and heme at two different domains. Heme
free portion of cellobiose dehydrogenase was thought to be a separate
enzyme, Cellobiose:quinone oxidoreductase. Cellobiose dehydrogenase
oxidizes cellobiose to cellobionolactone and prevents the recondensation of
the glycosidic bond of the cellulose chain nicked by endoglucanases (54).
Cellobiose dehydrogenase reduces quinones to phenols that can be used as
redox mediator during ligninolysis and also reduces aromatic radicals,
generated as a result of lignin degradation by the ligninolytic enzymes (55).
Cellobiose dehydrogenase generates H2O2 and superoxide anion and
oxidizes cellobiose to transfer 2e − to cytochrome thus producing ample supply
of energy to the fungus. It assists the ligninolytic peroxidases by reducing
compound II and especially, manganese peroxidase by dissolving the
Mn(IV)O2 precipitates (56).
Pyranose 2-dehydrogenase: It is a sugar C2-oxidoreductase having
molecular weight of 79 kDa (57). A number of monosaccharides,
oligosaccharides and glucosides serve as substrates for Pyranose 2dehydrogenase. It degrades cell wall polysaccharides thus interconnecting
lignocellulose decomposition by ligninolysis. This is proposed to reduce
quinone thus facilitating the smooth oxidation of lignin (57).
10
Non-enzymatic component of ligninolytic system
Veratryl alcohol: Veratryl alcohol, generated as a part of sec ondary
metabolism from glucose is a product of lignin degradation. Veratryl alcohol
plays a very important role in lignin peroxidase-catalyzed reductive ligninolysis
(58, 59). It not only acts as an inducer but also as a substrate. It facilitates the
redox reactions, catalyzed by lignin peroxidase, by acting as a redox mediator
(59).
Though, lignin peroxidase can catalyze some reactions in the absence
of veratryl alcohol but veratryl alcohol serves as a mediator for many redox
reactions. Veratryl alcohol prevents the formation of compound III and
compound IV, an inactivated form of lignin peroxidase due to excess of H2O 2,
and can also cause reactivation of these intermediates (38).
MnII: MnII acts as a substrate and redox coupler for manganese
peroxidase. It also acts as an inducer (controlling at the transcriptional level)
for the synthesis of manganese peroxidase in ligninolytic cultures. Manganese
peroxidase catalyzed oxidation and reduction reaction demands the Mn II as a
mediator (60).
H2O 2: H2O2 formed simultaneously with ligninolytic system is an
inseparable part of lignin degradation. It serves as a co-substrate of
ligninolytic peroxidases. It is also helps in generation of •OH radical, a very
potent reactive oxygen species. The reaction by which •OH radical is
generated is termed as Fenton’s reaction (61).
FeII+ + H2O2?? ?•OH + FeIII+ OH–
Thus, H2O2 supports ligninolysis in two major ways: 1) activates
peroxidases 2) supports generation of •OH radical.
Organic acids: These are synthesized and secreted during secondary
metabolism. Organic acids generate microenvironment necessary for
11
ligninolytic peroxidase activity by lowering the pH. It can chelate Mn III and
stabilize it which can diffuse away from the mycelial growth and can bring
about oxidation of various compounds in the remote sites. Organic acids can
serve as precursors for the generation of anion radicals (COO-•) that in turn
act as reductants and may also assist in the generation of other radicals like
-
O2 • and •OH (Fig. 1.6).
Fenton reaction
anion radical
LP( I / II )
VA
LP( II / N )
VA+
•
•
CO2 + CO2¯
O2
Organic CO2
acid
O2¯
cation radical
(oxidant)
•
anion radical
(reductant)
Fe
II
Fe
III
H 2O 2
OH ¯ + OH
•
Hydroxyl radical
(oxidant)
Figure 1.6 Organic acids as precursor for generation of various radicals
Quinones: Quinones are degradation products of lignin. Oxidation of
methoxy hydroquinones during lignin degradation followed by auto-oxidation
of the resulting methoxy semiquinones results in the formation of super oxide
anion radicals that further serve as a member non-enzymatic component of
ligninolytic system (62). Excess production of quinones ceases the process of
ligninolysis by inactivating the oxidases. Presence of dehydrogenase prevents
the excess accumulation of quinones and thus regulates lignin degradation.
Slime: Initiation of lignin degradation by white rot fungi results in the
accumulation of slime. The slime generates the microenvironment for lignin
degradation that is controlled temperature and pH, dissolved oxygen
availability, and aqueous environment. It also serves to concentrate the
nutrients for growth and the enzymatic and non-enzymatic ligninolytic
components essential for ligninolysis.
12
Importance of lignin degradation by lignin degrading fungi
Lignin degradation helps to understand the system of lignin degrading
fungi. Since, Lignin degrading fungi play a vital role in catalyzing lignin
degradation, the rate-limiting step of the ecologically important earth’s
Carbon-cycle. Lignin is a physical barrier in lingo-cellulose decomposition.
The breakdown of lignin will therefore help in the bioconversion of
lignocellulosic in many ways; first, production of a ruminant animal feed,
where breakdown of lignin will simplify the lignocellulosic feed and thus, will
increase the access of hydrolytic enzymes such as cellulases and
hemicellulases to their substrates (63). This will help in feed production by
controlled cultivation of ligninolytic fungi on lignocellulosic substrates. Second,
manufacture of mechanical pulp, where simplification of lignocelluloses will
act as a process of pretreatment for pulping . This has led to concept of biopulping. Mechanical and chemical processes employed in pulping lead to
pollution and demands high energy. Thus, pretreatment could reduce the
energy demand for mechanical pulping. Third, white rot fungi are efficient
producers of cellulases, hemicellulases and ligninases and can be used in the
treatment of waste liquors generated by sulphate pulping (64); for converting
lignocellulose wastes to protein rich feed (65) and effluent decolorization of
kraft pulp mills (66).
Apart from this, lignin degrading fungi can be used for producing
commercially important fungal metabolites or lignin-derived chemicals.
Solubilized lignin can be used in adhesive formulation, as an immunoadjuvant (67) etc. Fruiting bodies of many lignin degrad ing fungi are edible
and can be directly used as a food for humans. Since, they can grow on
cheap lignocellulosic waste, generation of edible lignin degrading fungi will not
only supplement the human food supply but also the decomposed waste
which can be u sed for ruminant animal feed.
Thus in brief lignin degrading fungi represent a potentially important
group of organisms having a wide range of biotechnological applications
(Figure 1.7). The understanding of biochemistry and enzymolozy of lignin
13
degrading fungi has opened a new door for research in the field of
bioremediation. Whole cell cultures as well as crude and pure enzymes have
been shown to degrade wide variety of xenobiotics compounds and appear to
be potent bioremediation tools.
Ecologically important
rate-limiting step of C cycle
Treatment of lignin-containing
industrial wastes
Treatment of
industrial wastes
Biodelignification
Production
of
Biomass
Lignin
biodegradation
Production of modified
lignin polymers
Production of
microbial metabolites
Production of chemicals
from lignin
Figure 1.7 Significance of lignin biodegradation
14
CHAPTER 1b
WHITE-R OT BASIDIOMYCETES:
D EGRADATION OF D YES
P
ersistence of many organic molecules synthesized by man in
the environment provided the impetus for studies on the
metabolism of xenobiotics. Compounding the problem of
recalcitrance was the bio-concentration and subsequent bio-magnification of
many xenobiotic compounds. The ability of microorganisms to metabolize
xenobiotic compounds has received much attention due to the environmental
persistence and toxicity of these chemicals.
Recent advances on the bioremediation of environmentally relevant
chemicals has centered on four important aspects: first, the characterization
of the biodegradation process useful for the treatment of the xenobiotic
compounds; second, the development of technical protocols for increasing the
degradation rates and substrate ranges of microbial enzymes; third, the
design and engineering of bioreactor systems and bio-treatment strategies to
optimize biodegradation process; and fourth, the development of data-base of
the ecological and human health risks associated with exposure to the
chemicals (68).
The recognition that environmental pollution is a worldwide threat to
public health has given rise to a new, massive industry for environmental
restoration. For both economic and ecological reasons, biological degradation
has become an increasingly popular alternative for the treatment of hazardous
wastes. White -rot basidiomycetes (WRB) comprise the only group of
organisms
known to degrade wood completely and employ unique
biodegradation mechanisms which confer them with the ability to degrade a
wide variety of xenobionts (37, 69).
15
Lignin Degradation by White-rot Basidiomycetes
In nature, the white -rot basidiomycetes (WRB) play a significant role in
the recycling of lignin -containing plant tissues. They have evolved strategies
for breaking down the irregular aromatic structure of the lignin polymer to
access and utilize cellulose and hemi-cellulose as carbon sources. Although
the lignin degrading ability has been recognized for many years, investigators
have now begun to understand the mechanisms by which this degradation is
accomplished (70, 71). The lignin degrading enzyme system of WRB is
extracellular and unusually nonspecific (37). White -rot fungi utilize simple
radical-generating mechanisms for the degradation of lignin. Peroxidases and
hydrogen peroxide, which are secreted by WRB, catalyze reactions
generating free radicals that cause depolymerization and degradation of
lignin. Although lignin is naturally a highly reduced polymer, it is mineralized to
carbon dioxide and water by WRB.
Understanding how this is accomplished is central to understanding
degradation of environmental pollutants by the fungi. The extracellular
biodegradation system also explains why the fungi can be quite resistant to
toxic or mutagenic chemicals.
White -rot Basidiomycetes: Dyes Degradation
White-rot basidiomycetes (WRB) are able to degrade pesticides, polyaromatic hydrocarbons, PCBs and other halogenated aromatics (including
dioxins), dyes, nitro aromatics, and toxic chemicals such as cyanides, azides,
carbon tetrachloride, and pentachlorophenol. WRB are also nonselective in
degrading all the chemical components of complex mixtures. For example, all
the components of Aroclors, toxaphene, creosote, and coal tars are degraded
by the fungi (72, 73). This ability is related to the nonspecific nature of the
peroxidases secreted by the fungi and to the variety of mechanisms that the
fungi use to degrade chemicals.
16
Synthetic dyes are extensively used in textile, paper, photography,
cosmetics and leather industries. Between 2-50% of the total dye consumed
in dyeing process are found in waste-waters depending on the class of dye
application (74, 75). Dyes possess several structural varieties such as acidic,
reactive,
basic,
disperse,
azo,
sulfonphthalein,
triphenyl
methane,
anthraquinone-based and metal-complex etc. and thus represent an important
role model for studies on xenobionts degradation. Dyes are chemically stable
that resist degradation by microbial attack (22) and therefore, persist in
conventional biological wastewater treatments.
Glenn and Gold (1983) suggested that the decolorization is a
secondary metabolic activity linked to the fungus ligninolytic activity (76). Such
secondary metabolizing ligninolytic cultures of Phanerochaete chrysosporium
decolorize structurally diverse polymeric dyes (Poly B-411, Poly R-481, and
Poly Y-606), crystal violet, para-rosaniline, cresol red, bromophenol blue,
ethyl violet, malachite green, brilliant green and azo and heterocyclic dyes
(77, 78).
Thereafter, degradation of azo, anthraquinone, heterocyclic, triphenyl
methane and polymeric dyes by WRB has been intensively studied (73, 79,
80). Degradation of copper-phthalocyanine dyes by P. chrysosporium resulted
in copper and organo copper breakdown products, which were found in
culture supernatants (81). Jarosz-Wilkolazka et al. screened 115 fungi of
different physio-ecological group for their ability to decolorize azo and
heterocyclic dyes, concluding h
t at heterocyclic dyes are decolorized easier
and faster by fungi than azo dyes (8 2). Recent investigation has shown the
involvement of Thelephora sp. in the decolorization of azo dyes and dye
industry effluents (83). This emphasizes the need of large-scale screening
program for WRB to identify potent species on behalf of their ability to
degrade dyes. Dye decolorization can be used as an alternative tool to radiolabeled
lignins
for
lignin
degradation
bioremediation for other xenobionts.
17
studies
and
in
programming
Involvement of lignin-degrading enzymes in the decolorization of dyes
was investigated by the several workers (79, 84 ). The first report of aerobic
degradation of azo dyes by white-rot fungi appeared in 1990, when Cripps et
al reported decolorization of azo dye Acid Orange 7 by nitrogen limited
cultures of P. chrysosporium (78). Decolorization of methylene blue by lignin
peroxidase (LIP) of P. chrysosporium provided evidence the oxidative mode
of degradation (80). Spadaro et al reported mineralization of azo dyes by the
ligninolytic enzymes (85). Two groups reported degradation of different
sulfonated azo dyes by crude and purified peroxidase preparations,
respectively (73, 86). LIP activity of P. chrysosporium was analyzed for the
decolorization of 50 structurally different dyes (87). Oxidation of nonsulfonated azo dyes [1-(4’-acetamidophenylazo)-2-napthol] by LIP from P.
chrysosporium resulted in the formation of 1,2-napthaquinone and acetanilide
(88). It was believed that manganese peroxidase (MnP) and laccase convert a
limited variety of azo dyes and preferred dyes carrying a phenolic substituent
in para- position to the azo bond and additionally methyl- or methoxysubstituents in 2- or 2, 6- position in relation to hydroxyl- group (89). Later it
was shown that the MnP from of Bjerkandera adusta and Pleurotus eryngii
catalyses decolorization of structurally diverse dyes in a Mn 2+-independent
reaction (90). Recently Moreira et al. demonstrated decolorization of Poly R478 using MnP (91). Laccases from Pycnoporus cinnabarinus is able to
decolorize complex, industrially relevant azo dyes such as Reactive Blue 5
and Direct Blue 1 (92). Our group reported the decolorization of
sulfonphthalein dyes by ligninolytic enzymes of different WRB (23, 25). Vyas
and Molitoris reported decolorization of RBBR by a novel enzyme, RBBR
oxygenase, produced by Pl. ostreatus , which is different from MnP, LIP,
veratryl alcohol oxidase and laccase (26). They reported decolorization of
RBBR by WRB that undergoes sequential change of blue dye to colorless
through a rainbow of intermediates also shown by other researchers (93).
Such observations strengthen the fact that decolorization is not a single step
reaction and intermediates are involved during complete degradation.
Decolorization of polymeric dyes Poly R-478 and Poly S-119 have
been shown by the cultures of P. chrysosporium in the static conditions as
18
well as immobilized in a bioreactor (94). Palma et al. correlated the
decolorization of the anthraquinone dye Poly R-478 with the ligninolytic
activity of an immobilized P. chrysosporium in a continuous -flow pulsing
packed bed reactor (95).
Involvement of degradative mechanism other than ligninolytic enzyme
for dye decolorization has also been studied. Plasma membrane dependent
redox system, of the fungus, allows adjusting the pH of its nearby surrounding
environment and maintains proton gradient across the plasma membrane.
Pasti & Crawford proposed plasma membrane redox system of WRB for dye
decolorization (96). Though dye decolorization has not been studied by this
system still the report on the mineralization of TNT by cultures of P.
chrysosporium explains the mechanism employed (19). The investigation
reports the reduction of TNT to mono- and di-amino congeners by transmembrane redox potential, which are further oxidized by MnP and
subsequent evolution of CO2 is associated with the production of LIP.
Report on the new enzymes, RBBR oxygenase and versatile
peroxidase, of ligninolytic system and other redox system involved in
degradation and the inherent complexity of dye molecules makes it difficult to
identify the degradative pathways utilized by WRB. The promising challenge
is to synchronize the parameters, like, environmental and nutrients, for whole
cell cultures and to integrate the role of various enzymes and other redox
machineries to come up with a total picture of how xenobionts are degraded
by white -rot basidiomycetes.
White -rot basidiomycetes: Mechanisms of Dye Degradation
Conventionally, coagulation and flocculation using Fe, Al, and Mg salts
have been practiced for the decolorization of dye wastewaters. The problems
associated with such process es are (i) limitation of chemical coagulation–
flocculation to produce good quality effluent, (ii) sludge handlin g problems, (iii)
the technical and economical constraints of reverse osmosis due to short
19
membrane life, (iv) economical non-feasibility, and (v) possible detrimental
effects of chemical oxidation using oxidative agents (22, 97, 98).
White-rot basidio mycetes degrade PAH, PCB, benzo[α ]-pyrene and
benzo[π]dioxins (18, 21, 99, 100). Initial reports not only established the nonspecificity of the fungus but also showed its ability to oxidize highly chlorinated
chemicals. Such highly chlorinated chemicals are electron deficient due to the
high electro negativity of the chlorine. These chemicals must therefore be
reduced before they can be oxidized (101).
White rot fungi synthesize and secrete a substrate for its own
peroxidase: veratryl alcohol (3,4-dimethoxybenzyl alcohol). The partially
oxidized veratryl alcohol (the cation free radical) then oxidizes other chemicals
that are not directly oxidized by the peroxidases (58, 102). To accomplish
further degradation, the fungus produces organic acids such as, oxalates that
also inhibit veratryl alcohol oxidation (103). Why would fungus produce a
substrate and an inhibitor of its own enzymes? Oxalate is easily oxidized to
carbon dioxide, thus its oxidation is essentially irreversible. However, the
oxidation of oxalate is a two-electron oxidation, whereas the reduction of the
veratryl alcohol cation radical consumes only one electron (104). Therefore,
the odd electron left in oxalate is available for other reductions. A number of
electron acceptors can thus be reduced by LIP provided with hydrogen
peroxide, veratryl alcohol (which is now called a "mediator"), and oxalate
(which has been termed the "donor"). Though the degradation of dye has not
been studied by this system yet the reductive dechlorination of carbon
tetrachloride (a highly oxidized chemical) to the trichloromethyl radical was
demonstrated with this system (105). The reductive dechlorination process
accomplished using a peroxidase, thus opens up a new field of investigation
into the role of peroxidases in the degradation of dyes requiring reduction.
Molecular oxygen can also be reduced by the oxalate radical to give
superoxide, a species of oxygen that is useful for either oxidation or reduction.
At low pH, superoxide is an excellent oxidant. Alternatively, superoxide is a
20
good reductant, especially at higher pH, for reductive dechlorinations (106).
Superoxide is also useful to generate another powerful oxidant, the hydroxyl
radical. In the presence of transition metals, such as iron, superoxide can
catalyze the generation of hydroxyl radicals by a sequence of reactions called
the Haber-Weiss reaction (107). White-rot basidiomycetes produce hydrogen
peroxide which can react with Fe 2+, termed as Fenton's reagent (108), to
produce hydroxyl radical.
WRB produces oxidants, like superoxide, hydroxyl radical etc., and
reductants, like oxalate-VA system etc., which generates redox-pump. It can
prove to be one of the important machinery in degradation of dyes.
Interest in the pollution potential of synthetic dyes has primarily been
prompted by concern over their possible toxicity and carcinogenicity. Many
dyes are made from known carcinogens, such as benzidine and other
aromatic compounds, all of which might be reformed as a result of physicochemical transformation (109). It has been shown that azo dyes are reduced
in sediments resulting in the parent toxic amines (110). Eaton et al. reported
for the first time biological decolorization of lignin -containing waste-water from
pulp and paper industries by two white-rot basidiomycetes, P. chrysosporium
and Tinctoporia sp (111). The activities of both the fungi were clear examples
of color removal through microbial degradation of polymeric lignin molecules.
Since then, P. chrysosporium in particular has been the subject of intensive
research related to the degradation of a wide range of synthetic dyes. Table1.1b summarizes some of the reports on dye decolorization by the white-rot
fungi. Phanerochaete chrysosporium decolorizes azo and heterocyclic dyes,
such as Orange II, Tropaeolin O, Congo red, and Azure B (78). The extent of
color removal varies depending on the dye complexity, nitrogen availability in
media, and the ligninolytic activity of the culture. At low nitrogen
concentrations, 90% of the color was removed within the initial 6 h; however,
in excess nitrogen concentrations, 63–93% decolorization of the above
mentioned dyes took up to 5 days (78). For P. chrysosporium, acidic
conditions (pH 3.5–5) and concentration of VA > 1 mM and hydrogen peroxide
(of 0.2 or 0.8 mM, as per the dye structu re) are highly favorable (113).
21
Azo dyes are the most widely studied dyes for decolorization studies
using white-rot fungi. Decolorization of azo dyes by ligninolytic enzymes is an
oxidative process that can result in complete degradation of the dye mole cule
to CO 2 and H2O (85, 122). The presence of the secondary metabolite VA
stimulates the decolorization by the fungi, by inducing the LIP activity (79, 84,
123). LIP oxidizes azo dyes where VA serves as a redox mediator. (84).
Indigo carmine, an azo dye, is a good substrate of MnP and LIP, and it has
been reported that although the dye is successfully decolorized by both
groups of extracellular peroxidases, the reaction by MnP is faster (87).
Although no general pattern has been established between the structure of
dyes and decolorization, it can be inferred from the research so far that the
dyes that are structurally similar to the natural substrates will be decolorized
faster and more efficiently. Dyes with structural similarities may still be
differentially decolorized by the white-rot fungi, for the reason that electron
distribution and charge density along with stearic distribution may differ (124).
Decolorization of Poly R-478 was used by Freitage and Morrel for
screening 170 strains of white-, brown-, and soft-rot fungi, as well as
xylophilous fungi for their peroxidase and phenol oxidase activity (125). They
concluded that no correlation exists between the MnP or LIP and
decolorization of dyes. Heinfling et al. initially showed the decolorization of
phthalocyanines dyes Reactive blue 38 and Reactive blue 15 by B. adusta, P.
chrysosporium, and Trametes versicolor (119). Later they reported the role of
MnP isozymes involed in the decolorization of these dyes (120). The ability of
the WRB to degrade the synthetic dyes has been exploited for the potential
use of dyes as substrates for enzyme activity. Pasti-Grigsby et al. examined a
number of azo dyes and found that Orange I and 3,5-dimethyl-4hydroxyazobenzene-4'-sulfonic acid served as good substrates for MnP, while
Orange II and 3,5-difluoro -4-hydroxybenzene-4'-sulfonic acid were better for
LIP (126). Transformation and degradation of the disazo dye Chicago Sky
Blue by a purified laccase from Py . cinnabarinus opened the possible use of
laccase in the dye degradation studies (74). A Brazilian strain of Pl.
pulmonarius produces only laccase and no MnP, LIP, or aryl alcohol oxidase,
decolorizes dyes of different spectra (127).
22
Table 1.1b Dyes decolorized by some known white-rot basidiomycetes
Organism
Dye
Phanerochaete Bromophenol blue, Congo red, Methyl Green, Methyl
chrysosporium Orange, Poly R-478, RBBR, Toluidine Blue O, Poly
S-119, Poly T-128
Orange II
Methyle ne Blue
Crystal violet
Disperse Orange 3
Disperse Yellow 3
Indigo carmine, Cibacron Brilliant Red, Brilliant green
Amaranth, Remazol Black B, Remazol Orange,
RBBR, Reactive Blue, Tropaeolin O
Polymeric Dyes, B-411, R-481, Y-606
Acid Yellow 9
Trametes
versicolor
Remazol Black B, Remazol Orange, RBBR, Reactive
blue
Indigo carmine, Reactive blue 15, Acid Blue 25, Acid
Black 24
Crystal violet
Pigment Violet 12
Orange II
Reactive orange 96, Reactive violet 5, Reactive black
5, Reactive blue 38
Bjerkandera
adusta
Reactive blue 38, Reactive violet 5, Reactive black 5,
Reactive orange 96, Reactive red 198, Reactive blue
15
Irpex lacteus
RBBR
Bromophenol blue, Cu-phtha locyanine, Methyl Red,
Congo red
Pleurotus
ostreatus
Acid Black 194, Acid Blue 185, Disperse Blue 56,
Sulfur Black 1, Vat Blue 6, Orisol Blue BH, Disperse
Yellow 54, Reactive blue 19, Reactive blue 158, Vat
Red 10, Vat Yellow 46
RBBR
Bromophenol red, Phenol red, Bromophenol blue, mCresol purple, o-Cresol red, Bromocresol purple,
Bromocresol green
Reference
79
78
80
73
85
88
126
112
76
124
112
113
115
114
116
117
118
119
120
121
26
25
Soares et al. have reported decolorization of Remazol Brilliant Blue R
using a combination of redox mediator, laccase, and nonionic surfactant
23
(128). They postulate the necessity of a redox mediator for laccase-catalyzed
decolorization of dyes.
Summing up the mechanisms used by WRB in degradation of dyes
and other xenobionts, explains that enzymatic system together with redoxpumps provide powerful means for the degradation dyes. The varied
combination of the above mentioned system and followed by standardized
parameters (environmental and nutrient) for the cultivation of fungi is most
important to scale-up the results from laboratory to field level. Future work
needs to be focused on this area for a sustainable technology using white-rot
basidiomycetes.
White -rot basidiomycetes: A Promising Candidate for Xenobionts
Degradation
Biotechnological applications of white-rot basidiomycetes (WRB) and
their enzymes are being developed, as alternative methods for pulping and
bleaching and for removal of toxic pollutants from soil and water reservoirs.
White-rot
basidiomycetes
have
developed
unique
mechanisms
for
degradation of recalcitrant compounds such as lignin. It was not until the latter
part of the 1980s that the white-rot fungi's capability for degradation was
considered also for organo-pollutants. The first studies with WRB, P.
chrysosporium and P. sordida in soil were published in 1989-90. Even so P.
chrysosporium in liquid culture continued to be the main focus for several
years. Since soil is not the natural habitat for WRB, soil remediation requires
knowledge from several areas; physiology of WRB, soil chemistry, and soil
biology.
Future work directed towards identifying novel oxido-reductases that
may have auxiliary roles in transformation of dyes that are more recalcitrant
could lead to the development of a more effective enzymatic cocktail for dye
decolorization. The use of whole fungi, rather than enzymes might be
expected to be a more viable option from the point of view of the
decolorization process control at the industrial scale for the reason that
24
degradation process is not achieved by a single enzyme but with many
enzymes and the varied composition of these enzymes. Again, the
degradation is not related only to the enzymatic system but also to the nonenzymatic components that adds up to the degradation by the generation of
redox pump and also provides the needed microenviro nment for the
degradation process. Thus instead of acclimatizing the system with such
factors it become obvious to understand that the whole cell appears to be a
more viable option.
The failure of success is probably due to an engineering problem rather
than a fundamental limitation of the technology itself. Advantage of using
WRB is that they do not require preconditioning to particular pollutants and
can degrade xenobiotics present in a very low concentration to non-detectable
level and even complete elimination. Thus in brief lignin degrading fungi
represent a potentially important group of organisms having wide range of
biotechnological applications .
25
CHAPTER 2
SCREENING OF WHITE-R OT BASIDIOMYCETES:
DECOLORIZATION OF TEXTILE D YES
2.1 Introduction
W
hite rot basidiomycetes (WRB) are the only known organisms
to mineralize lignin, a recalcitrant biopolymer (2). The
ligninolytic system of WRB is (I) extracellular (II) non-specific,
as they are able to degrade lignins of different origins and
composition (15), (III) highly oxidative (37), and (IV) non inducible, as lignin
degradation is an idiophasic event (6-7). These properties allow the WRB to
degrade a wide variety of xenobionts.
Xenobionts are defined as the organic compounds that are foreign to
life in general. Such compounds are usually recalcitrant as the cells are never
exposed to these chemicals; as a consequence they resist biodegradation
and persist in nature. Dyes can be considered as role models of xenobionts
because of their; (I) structural diversity, (II) stability, and (III) complexity in
structure, which make them resemble diverse groups of xenobionts. It is easy
to follow their degradation because of their chromogenic nature.
Conventional mechanisms like, photocatalysis, ozonation etc. are not
efficient to remove recalcitrant dyes from environment (97,110,111). The
environment-friendly and economic option to known conventional systems is
biodegradation. Xenobionts that are structurally similar to lignin sub-structures
and the mode of action of ligninolytic system of WRB make them potential
candidates and provide an efficient alternative in the bioremediation program.
Several methods for screening of white -rot basidiomycetes have been
reported where dyes are used as substrates (26, 76). We report the
26
decolorization of 7 textile dyes by the white-rot basidiomycetes, Irpex lacteus
and strain F (collected from Kathiyawad peninsula).
2.2 Materials and Methods
2.2.1. Microorganisms: The ligninolytic fungus Irpex a
l cteus Fr. 238
617/93, originated from Culture Collection of Basidiomycetes Prague (kindly
gifted by Šašek, V.), and strain F were used. The strain F was cultivated from
the fruit-body collected from Kathiyawad peninsula and isolated in pure in our
laboratory. The strains were maintained on malt extract agar plates at 5ºC.
2.2.2. Isolation of Strain F: Lignin degrading white rot fungus F was
collected from Tamarindus indicus plant root from Abhapara, Barda Hills. A
piece of collected sample was surface ste rilized with 0.1% HgCl2, followed by
ethanol washing. In between this procedure sterile double distilled water wash
was given regularly. This piece was then placed on water agar and observed
for development of mycelia on plate. Mycelia were then transferred to fortified
malt extract agar (FMEA) plates. The culture was obtained in pure by making
serial transfers and maintained on FMEA at 4ºC.
2.2.3. Textile dyes: Remazol Turquoise Blue G (RTBG), Reactive
Orange 16 (RO16), Remazol Yellow FG (RYFG), Remazol magenta HB
(RMHB), Reactive blue 21 (RB21), Remazol red H8B (RRH8B), Reactive Red
6BX (RR6BX), Reactive orange 13 (RO13), Reactive brown 18 (RB18),
Reactive black 5 (R B5), Reactive golden HR (RGHR) and Remazol Brilliant
blue R (RBBR) were used for the decolorization studies.
2.2.4. Decolorization of textile dyes on solid media: The fungi were
screened for their ability to decolorize textile dyes using malt extract agar
medium (malt extract 30 g.l-1, mycological peptone 5 g.l-1 and agar 15 g.l-1)
containing the dyes (100 ppm). The plates were inoculated with a disk (8 mm)
punched from the cultures grown on malt extract agar plates and incubated at
28ºC. The plates were observed daily till day 9 for the zone of growth (mm)
and the zone of decolorization (mm) of the dyes.
27
2.3 Results and Discussion
Irpex lacteus and strain F were screened for their ability to decolorize
textile dyes on malt agar medium. The growth of the fungus on malt agar
medium with or without dye occurred equally well. None of the textile dyes
appeared to inhibit the fungal growth. Dye decolorization on solid media was
under older mycelia which always started from the point of inoculation and the
zone of decolorization followed the zone of growth. It indicates that the
process of decolorization which is associated with older mycelia is perhaps a
part of secondary metabolic event when the ligninolytic enzymes are secreted
that have been shown to degrade a variety of organo-pollutants.
Mycelial growth of strain F covered the entire plate by day 5. Strain F
growing on malt agar medium decolorized all the dyes. The decolorization of
dyes that initiated after day 2 continues till the dye in the entire plate is
decolorized, but decolorization of Reactive golden HR (RGHR) occurred only
till day 5. Thereafter no further decolorization occurs as observed from the
zone of decolorization. Decolorization of Remazol Turquoise Blue G (RTBG),
Reactive Orange 16 (RO3R), Remazol Yellow FG (RYFG) and Remazol
Brilliant Blue R (RRBR) was completed in betw een day 6 and 7 (Fig. 2.1).
Textile dyes, Reactive Brown 18 (RB18), Remazol Red H8B (RRH8B),
Reactive Orange 13 (ROH2R), Remazol Magenta HB (RMHB), Reactive Blue
21 (RB21) and Reactive Red 6BX (RR6BX) were decolorized completely on
day 9 (Fig. 2.1).
Both the red dyes, viz. RRH8B and RR6BX, were decolorized but the
decolorization of RRH8B was faster and was completed on day 8. Similar
results were obtained for the orange dyes, viz. RO13 and RO16, and blue
dyes, viz. RBBR and RB21, where decolorization of orange dye, RO16 and
blue dye, RBBR was completed on day 6.
28
100
100
(b )
(a)
75
75
50
50
25
25
0
0
100
100
(d)
(c)
75
50
50
25
25
0
0
100
100
(e)
75
Size of zone (mm)
75
50
50
25
25
0
0
100
100
(h)
(g)
75
75
50
50
25
25
0
0
100
100
(j)
(i)
75
75
50
50
25
25
0
0
100
100
(k)
75
(f)
75
(l)
75
50
50
25
25
0
0
0
3
6
9
0
3
6
9
Incubation time (days)
Figure 2.1 Decolorization and the growth of the white rot basidiomycete
Strain F on malt agar medium containing textile dyes (a) RTBG; (b)
RYFG; (c) RR6BX; (d) RRH8B; (e) RO13; (f) RO16; (g) RMHB; (h) RB5;
(i) RB21; (j) RBBR; (k) RGHR; (l) RB18
29
Growth of I. lacteus and associated decolorization of textile dyes was
faster when compared to the strain F. Irpex lacteus decolorized all the textile
dyes, under investigation. Decolorization initiated after day 2 and most of the
dyes were decolorized by day 5 when the mycelial growth covered the entire
plate (Fig. 2.2). Decolorization of textile dyes, RGHR and RYFG, initiated on
day 2, like other dyes, were decolorized only till day 5. There was no further
increase in the size of zone of decolorization. Decolorization of RTBG,
RR6BX, RRH8B, RO13, RB21, and RB18 followed the growth and were
decolorized completely on day 9. The decolorization of red dyes (RRH8B and
RR6BX) orange dyes (RO13 and RO16) and blue dyes (RBBR and RB21)
were varied (Fig. 2.2) and resembled the decolorization process observed in
the case of strain F.
It is interesting to note that decolorization of blue and black dyes, by
both the fungi, initiated by change in colors having absorbance towards lower
wavelength. The color of the dyes changed from blue/black to red/orange and
than to yellow before getting colorless (Plate 2.1). The same phenomenon of
hypsochromic shift has been reported in the in vitro degradation of dyes (26).
This clarifies that the degradation of dyes is not a single step reaction but
involves formation of several intermediates.
(a)
(b)
(c)
Plate 2.1 Decolorization of Reactive black 5 by Irpex lacteus , showing
hypsochromic shift; (a) medium with dye; (b) I. lacteus growing on
malt extract medium; (c) I. lacteus growing on medium containing dye
30
100
100
75
50
50
25
25
0
0
100
100
(c)
75
Size of zone (mm)
75
(a)
(d)
75
50
50
25
25
0
0
100
100
(e)
75
(b)
(f)
75
50
50
25
25
0
0
100
100
(g)
75
50
50
25
25
0
0
100
100
(i)
75
(h)
75
(j)
75
50
50
25
25
0
0
100
100
(k)
75
(l)
75
50
50
25
25
0
0
0
3
6
9
0
3
6
9
Incubation time (days)
Figure 2.2 Decolorization and the growth of the white rot basidiomycete
Irpex lacteus on malt agar medium containing textile dyes (a) RTBG; (b)
RYFG; (c) RR6BX; (d) RRH8B; (e) RO13; (f) RO16; (g) RMHB; (h) RB5;
(i) RB21; (j) RBBR; (k) RGHR; (l) RB18
31
The same color dyes (red, orange and blue) were decolorized variably
by both the basidiomycetes. It has been reported that the dyes that are
structurally similar may be differentially decolorized by white rot fungi. The
reason may be attributed to the differences in the electron distribution and
charge density along with the steric -distribution in different dyes (126).
Growth of both the fungi was similar on the malt agar plates; the
growth was slower till day 2 and thereafter increased sharply. The plates were
entirely covered by day 5. Both the fungi decolorized all the textile dyes
investigated. Irpex lacteus decolorized textile dyes faster as compared to
strain F.
The present work provided important lead regarding the ability of the
white rot fungus to produce some of the ligninolytic activities that need to be
purified and characterized. The wood-rotting white rot basidiomycete, I.
lacteus is a better alternative than strain F in the degradation study
concerning bioremediation program. The ability of I. lacteus to decolorize
various groups of dye and the correlation between dye decolorization and
ligninolytic activities should be addressed. Such work would allow comparison
between the ligninolytic activities of this fungus with others of its kind.
32
CHAPTER 3
LIGNINOLYTIC ENZYMES OF IRPEX LACTEUS:
D ECOLORIZATION OF TEXTILE A ND
TRIPHENYL METHANE D YES
3.1 Introduction
D
yes are tailor-made for their stability. It is known that 90% of
reactive textile dyes entering activated sludge sewage treatment
plants pass through unchanged and are discharged to rivers
(129). Not all dyes currently used could be degraded and/or removed with
physical and chemical processes, and sometimes the degradation products
are more toxic (130). A great variety of synthetic dyes are used for textile
dyeing and other industrial applications. The structural diversity of dyes
derives from the use of different chromophoric groups
(e.g., azo,
anthraquinone, triphenyl methane, and phthalocyanine groups) and different
application technologies (e.g., reactive,direct, disperse, and vat dyeing) (74).
Lignin-degrading white rot fungi have received worldwide attention for
their industrial use in bio -pulping, bio-bleaching, dye decolorization, and in
detoxifying recalcitrant environmental pollutants (16, 85, 131-133). The
ligninolytic system of these fungi is extracellular, non-specific and noninducible, which allows the fungi to mineralize lignin and variety of organic
chemicals. Most information on these fungi comes from studies with
Phanerochaete chrysosporium and few other species, thus representing only
a small part of many hundreds of species existing in nature. Irpex lacteus is a
cosmopolitan white rot basidiomycete typically inhabiting dead hard wood
trees (134). But, its role in degradation of xenobionts is largely unknown;
except for the report on transformation of pentachlorophenol from soil (135).
In our report, we investigated decolorization of textile dyes and triphenyl
methane dyes by the ligninolytic activities of Irpex lacteus.
33
3.2 Materials and Methods
3.2.1. Microorganism: Irpex lacte us Fr. 238 617/93 obtained fro m
Culture Collection of Basidiomycetes (CCBAS), Prague (Kind gift from V.
Šašek). The strain is maintained on malt extract agar plates at 4 ºC.
3.2.2. Textile Dyes : Reactive black 5, Remazol black HR , Reactive
orange 13, Reactive orange 16, Remazol red H8B, Remazol magenta HB,
Remazol turquoise blue G, Remazol lemon yellow F4, Remazol golden yellow
R, Reactive brown 18 and Remazol brilliant blue R were obtained from the
dye manufacturing and textile processing industries.
3.2.3. Triphenyl methane Dyes: Brilliant green, Methyl blue and Fast
green were procured from Hi-Media (India).
3.2.4. Production of ligninolytic enzymes by I. lacteus : Solid state
fermentation of wheat straw by I. lacteus was performed as described
previously [1 36]. To the culture flasks, harvested on 20th day, 100 ml of
phosphate buffer (0.1 M, pH 6.5) containing 0.1 M NaCl was added, contents
were gently beaten and incubated on rotary shaker for 1 h. Liquor obtained
was then filtered and spun (6000 rpm, 15 min, 5oC) before being subjected to
ammonium sulfate precipitation (80% saturation). Precipitates separated upon
centrifugation (6000 rpm, 15 min, 5 oC) were dissolved in the minimum amount
of phosphate buffer (0.1 M, pH 6.5). Enzyme extract thus obtained was
desalted using PD10 column (Pharmacia) and stored in small aliquots at -5ºC
for further biochemical analysis.
3.2.5. Enzyme assays: Manganese peroxidase (MnP), manganeseindependent peroxidase (MIP) and lignin peroxidase (LIP) activities were
analyzed using dyes as the substrate. Reaction mixture (2 ml) for MnP activity
contained 100 µmol sodium tartarate buffer (pH 2.5, 3.0 and 3.5), succinate
lactate buffer (pH 4.0 and 4.5) and sodium citrate buffer (pH 5.0 and 5.5), 0.2
µmol MnSO4, 150 ppm textile dyes or 75 ppm triphenyl methane dyes, 0.4
µmol H2O2 and the enzyme extract. Reaction mixture for MIP activity was
similar to that of MnP but MnSO4 was not added. Reaction mixture for LIP
34
activity was similar to that of MnP but veratryl alcohol (4 µmol) was added
instead of MnSO 4. One unit enzyme activity is defined as the amount of
activity that consumes 1 µmol of the triphenyl methane dye or produces a
change of one absorbance unit per min upon oxidation of textile dye at its
λmax.
3.3 Results and Discussion
Irpex lacteus , during solid state fermentation of wheat straw, produced
ligninolytic activities, namely, manganese peroxidase (MnP), manganeseindependent peroxidase (MIP) and lignin peroxidase (LIP). Decolorization of
11 textile dyes and 3 triphenyl methane dyes by the ligninolytic enzymes
produced by I. lacteus was investigated at varying pH (Fig 3.1 and 3.2). All the
dyes investigated were decolorized by one or the other ligninolytic activities
except remazol golden yellow R (RGYR) and brilliant green (BG).
3.3.1. Decolorization of textile dyes by MnP activities: MnP decolorized
two textile dyes, reactive orange 16 (RO16) and remazol magenta HB
(RMHB) (Fig 3.1). Dyes were decolorized over a broad pH range (2.5 -5.5) and
were best decolorized at the MnP pH optimum (4.5). Decolorization of textile
dyes increased rapidly above pH 3.0. Decolorization of RO16 occurred in the
pH range of 2.5 to 5.0 and RMHB in the pH range of 2.5 to 5.5. Decolorization
of RO16 declined sharply above pH 4.5. Dyes that contain free phenolic
moiety are better substrates of MnP as MnP prefers aromatic compounds
having free phenolic moiety (25, 32).
3.3.2. Decolorization of triphenyl methane dyes by MnP activities:
Ligninolytic enzymes catalyze decolorization of synthetic dyes (2 4-26, 137).
Decolorization of triphenyl methane dyes was only associated to the MnP
activities (Fig 3.2). MnP decolorized methyl blue (MB) and fast green (FG) but
was unable to decolorize brilliant green (BG).
35
30
20
20
10
10
0
0
45
30
30
15
15
0
0
RO13
30
% Decolorization
RBBR
40
20
0
21
BHR
45
RO16
60
% Decolorization
30
RBB
RTBG
20
10
0
60
RRH8B
14
40
7
20
0
0
21
15
LYF4
RMHB
GYR
14
10
7
5
0
0
2
15
3
4
5
6
pH
BrH4R
10
5
0
2
3
pH
4
5
6
Figure 3.1 Decolorization of textile dyes by manganese peroxidase (?),
manganese-independent peroxidase (? ), and lignin peroxidase (? ): (a)
Reactive black 5; (b) Remazol black HR; (c) Reactive orange16; (d)
Reactive orange 13; (e) Remazol brilliant blue R; (f) Remazol turquoise
blue G; (g) Remazol red H8B; (h) Remazol magenta HB; (i) Remazol lemon
yellow F4; (j) Remazol golden yellow R; (k) Reactive brown 18
36
Decolorization of RMHB, MB and FG, was associated only with
the MnP activity. This signifies that certain dyes are better substrates of MnP
and therefore can be employed to differentiate different ligninolytic activities
and for studying MnP catalyzed reactions. Dye decolorization has been used
as an indicator and measure of ligninolytic activity (35, 138).
3.3.3. Decolorization of textile dyes by LIP activity: Decolorization of
RO16 and reactive brown 18 (RB18) was associated also with the LIP activity
(Fig 3.1). Decolorization of RO16 occurred over a pH range of 2.5 to 5 but
was best at pH 2.5, the pH optimum of LIP. Further increase in the pH from
2.5 led to the slower decolorization. Reactive brown 18 was decolorized in the
pH range of 2.5 to 4. Decolorization of RB18 increased sharply above pH 2.5
and was best decolorized at pH 3.5. Above pH 3.5 the decolorization
declined.
15
BG
10
5
% Decolorization
0
45
MB
30
15
0
45
FG
30
15
0
2
3
4
5
6
pH
Figure 3.2 Decolorization of triphenyl methane by manganese peroxidase
(?), manganese-independent peroxidase (? ), and lignin peroxidase (? ): (a)
Brilliant green; (b) Methyl blue; (c) Fast green.
37
3.3.4. Decolorization of textile dyes by MIP activity: Decolorization of
most of the textile dyes was associated with the MIP activity. MIP activity did
not decolorize the triphenyl methane dyes. Anthraquinone, azo and
phthalocyanine dyes have been reported to be decolorized in a manganeseindependent enzymatic oxidation (26, 40). MIP showed the highest non
specificity in selection of textile dyes for decolorization and decolorized textile
dyes over a broad pH range (2.5 -5.5) (Fig 3.1). Unlike MnP, the pH optimum
for decolorization by MIP activity was dependent on the dyes. Remazol
brilliant blue R (RBBR), RB18 and remazol lemon yellow F4 (LYF4), were
decolorized best at pH 2.5 and 3.0. Reactive black 5 (RB5), remazol red H8B
(RRH8B), reactive orange 13 (RO13), and remazol black HR (BHR) were
decolorized best at 3.5, the pH optimum of MIP. Remazol turquoise blue G
(RTBG) was decolorized best at pH 4.0. The variation in the pH for the
decolorization of dyes by MIP is indicative of pH as the function of enzymecatalyzed dye decolorization.
3.3.5. Influence of pH on the decolorization of textile dyes: pH plays a
crucial role in the decolorization of textile and triphenyl methane dyes.
Decolorization by the concentrated crude enzyme extracts prepared from I.
lacteus-infested wheat straw occurred over a pH range of 2.5 to 5.5 (Fig 3.1
and 3.2). MnP and LIP decolorized the dyes most at their respective optimal
pH. Optimal pH for the decolorization associated to MIP activity varied with
the dye. Differences in the production of ligninolytic enzymes make WRF to
differ in their ability to degrade and decolorize various dyes. As a result of
nonspecificity of lignin-degrading enzymes, individual azo-, heterocyclic -,
phthalocyanine-, polymeric-, triphenyl methane- and anthraquinonic dyes
(120, 137-139) as well as complex industrial effluents are efficiently
decolorized by these fungi (140-140).
3.3.6. Structure function relationship of decolorization of dyes by
ligninolytic activities produced by I. lacteus: MIP is more versatile in the
decolorization of textile dyes and MnP activity is the only ligninolytic activity
that is associated with the decolorizatio n of triphenyl methane dyes. RO16 is
an azo dye with an intermediate compound vinyl sulphone azotized to
38
naphthalene (Fig. 3.3). RO13 is derived from an intermediate compound
sulpho taubias acid azotized to naphthalene moiety and an additional
intermediate, cyanuric chloride. Therefore, RO13 contains two naphthalene
moieties (Fig. 3.3). Diazotization of vinyl sulphone group leads to the
formation of RB5 (Fig 3.3). RBBR is an anthraquinone dye with a vinyl
sulphone group (Fig 3.3).
Presence of a halogenate d compound, cyanuric chloride, in the
structure of RO13 makes it a suitable substrate for MIP and not for MnP.
Whereas, MnP decolorizes RO16 that has naphthalene ring with hydroxyl
group, presenting the dye as a simpler phenolic moiety. Reports on
decolorization of sulfonphthalein dyes by ligninolytic enzymes of P. ostreatus
has shown the preference of halogenated dye as a substrate for MIP and the
non halogenated, simpler dyes as a preferential substrate for MnP (23, 25).
RB5 has an additional vinyl sulphone group and is decolorized only by the
MIP activity. The presence of additional vinyl sulphone group does not allow
MnP or LIP to decolorize it. Decolorization of an anthraquinonic dye, RBBR,
by I. lacteus suggests MIP as an important tool in degradation of xenobionts
having anthraquinonic intermediates.
MnP was the only ligninolytic activity associated to the decolorization of
triphenyl methane dyes. Among the three triphenyl methane dyes, BG, FG
and MB, BG resisted decolorization. The presence of two N, N-diethyl amine
group in the structure of BG prevents the ligninolytic activities to use it as a
substrate (Fig. 3.4). MB was decolorized better than the FG. Presence of
three 4-amino benzene sulfonate group in MB, makes these dyes better
substrates for Mn P (Fig. 3.4). FG has two 3-[2-(ethylamino) ethyl] benzene
sulfonate group with one hydroxyl group and one sulfonic acid group attached
to it. The order of preference of triphenyl methane dyes as substrate for MnP
mediated decolorization is MB > FG.
Since the effluents usually do not contain single dye but a concoction
of dyes, therefore, a single enzymatic activity cannot bring about the
degradation of effluents and thus there is a need to develop a blend of
39
enzymatic system that can act on wide varieties of dyes present in the
effluent.
NaO3SOCH2
N N
S
O
Remazol Orange 3R
C.I. Name: Reactive orange 16
C.I. No.: 17757
HO
O
CH3 C
SO3Na
SO3 Na
NH
OH
N N
N
NaO 3S
NaO3S
NH
N
N
CH3
N
Cl
Remazol Orange H2R
C.I. Name: Reactive orange 13
C.I. No.: 18270
OH
NaO3SOCH 2OS
N N
NH2
N N
NaO 3S
SO3Na
Remazol Black B
C.I. Name: Reactive black 5
C.I. No.: 20505
Remazol Brilliant Blue R
C.I. Name: Reactive Blue 19
FW : 624.52
Figure 3.3 Structure of Textile dyes
40
SO 2CH 2CH 2OSO2Na
CH3
CH3
N
N
CH3
Brilliant Green
C.I. No: 42040
F.W.: 482.65
CH3
OH
SO 3Na
CH3
CH3
N
N
Fast Green
C.I. No: 42053
F.W.: 808.86
SO3Na
SO3 Na
SO 3 Na
HN
HN
SO3 Na
NH
Methyl Blue
Acid Blue 93
F.W.: 799.8
SO 3 Na
Figure 3.4 Structure of Triphenyl methane dyes
41
CHAPTER 4
ENZYMATIC DECOLORIZATION OF
SULFONPHTHALEIN DYES
4.1 Introduction
C
chemical
olor removal in particular has recently become of major scientific
interest, as indicated by the multitude of related research reports (22,
122, 140, 143). Conventional waster water treatment is not efficient
to remove recalcitrant dye stuffs from effluents. Physical and
methods
(adsorption,
chemical
transformation,
incineration,
photocatalysis and ozonation etc.) for the removal of dyes are not suitable due to
high cost, low efficiency, and in-applicability to a wide variety of dyes (90, 111,
112, 144). Biodecolorization of paper pulp, paper waste-waters and synthetic
dyes as measured by the decrease in color absorption was reported as early as
1980 (113).
Lignin degrading fungi mineralize a number of important recalcitrant and
xenobiotic compounds under ligninolytic conditions (16, 18, 21, 145, 146). WRF
degrade xenobionts with varied capacity and thus researchers have stressed the
need to perform a large scale screening of the ligninolytic enzyme systems of
white rot fungi (68, 147, 148). Several screening methods involving dyes have
been reported (26, 76, 149). Advantage of using WRF is that they do not require
preconditioning to particular pollutants and its free radical based enzyme
catalysis can degrade variety of xenobionts present in a very low concentration to
non-detectable level and even complete elimination (150). The ability of these
fungi to degrade lignin and structurally diverse pollutants results from the
relatively non inducible and nonspecific nature of their extracellular enzymes (37,
151, 152). We tested a number of white rot fungi for their ligninolytic activity by
using SP dyes as indicator substrate. In our report, we present the results of our
42
investigations of SP-decolorizing activity produced by P. ostreatus during solid
state fermentation of wheat straw, a natural lignocellulosic substrate.
4.2 Materials and methods
4.2.1. Organism: Pleurotus ostreatus strain 3004 (ATCC 3545) originated
from Culture Collection of Basidiomycetes (CCBAS), Prague was used (Kindly
gifted by Šašek, V.). The strains were maintained on malt extract agar plates at 5
ºC.
4.2.2. Production of ligninolytic enzymes by P. ostreatus: Solid state
fermentation of wheat straw by P. ostreatus was performed as described
previously [12]. A set of 7 flasks was harvested on 5th day onwards. To the
culture flasks 100 ml of phosphate buffer (0.1 M, pH 6.5) containing 0.1 M NaCl
was added, contents were gently beaten and incubated on the rotary shaker for 1
h. Liquor obtained was then filtered and spun (6000 rpm, 15 min, 5oC) before
being subjected to ammonium sulphate precipitation (80% saturation).
Precipitates separated upon centrifugation (6000 rpm, 15 min, 5oC) were
dissolved in the minimum amount of phosphate buffer (0.1 M, pH 6.5). Enzyme
extract thus obtained was stored in small aliquots at -5ºC for further biochemical
analysis.
4.2.3. Enzyme assays: Manganese peroxidase (MnP) and manganeseindependent peroxidase (MIP) activities were analyzed by oxidation of guaiacol
to colored product using spectrophotometer. Reaction mixture (2 ml) for MnP
activity contained 100 µmol succinate lactate buffer (pH 4.0), 0.2 µmol MnSO4,
0.9 µmol guaiacol, 0.7 µmol H2O2 and the enzyme extract. Reaction mixtures for
MIP activity were similar to that of MnP but sodium tatarate buffer (pH 3.5) was
used and MnSO4 was not added. One unit enzyme activity is defined as the
amount of activity that produces a change of one A470 unit per min upon oxidation
of guaiacol.
43
Sulfonphthalein (SP) dyes decolorizing activities of MnP and MIP were
assayed using respective reaction mixtures as described above, wherein
guaiacol was replaced by SP dyes investigated. MnII was varied in the range of
10-400 nmol in the reaction mixture. The reaction was terminated after 5 min with
1.25 µmol NaOH. Sodium tartarate buffer pH 3.5 and succinate lactate buffer pH
4.0 was used for the enzyme assay. Absorbance was monitored at the ?max of
respective dyes. Dye decolorizing activity was calculated using molar
absorbance coefficient of the respective dye (Table 4.1). One unit enzyme
activity is defined as the amount of activity that decolorizes 1 nmol dye per min in
the reaction mixture.
4.3 Results and discussion
4.3.1. Production of ligninolytic activities: Extracellular enzyme extracts
prepared from Pleurotus ostreatus infested wheat straw possessed manganese
peroxidase (MnP) and manganese-independent peroxidase (MIP) activities (Fig
4.1). MnP and MIP activities were produced throughout the experimental run
since they were detectable even on day 1. These activities were maximal on day
7 and thereafter decreased gradually.
Enzyme activity (U/L)
9
6
3
0
3
6
9
12
Days
Figure 4.1 Manganese peroxidase activity at pH 3.5 (? ) and 4.0 (¦ ) and
manganese-independent peroxidase activity at pH 3.5 (?) and 4.0 (?) of
Pleurotus ostreatus growing on wheat straw
44
Manganese
peroxidase
(MnP;
EC
1.11.1.13)
was
discovered
in
Phanerochaete chrysosporium and was later shown to be produced by other
white rot fungi including P. ostreatus. The decolorization of dyes involving
ligninolytic enzyme system of white rot fungi was first reported in 1983 and was
used to measure ligninolytic activity of Ph. chrysosporium [76]. Along with the
production of the ligninolytic activities, P. ostreatus also produces different dye
decolorizing activities [26, 122]. We report the sulfonphthalein (SP) dyes
decolorizing activities associated with MnP and MIP produced by P. ostreatus.
Mechanisms involved in MnP-catalyzed reactions have been elucidated. It
oxidizes MnII to MnIII, which is stabilized by organic acid chelators and acts in turn
as a low molecular mass, diffusible, redox mediator that attacks organic
molecules nonspecifically via hydrogen and one electron abstraction [32, 153].
4.3.2. Decolorization of sulfonphthalein dyes: The SP dyes (Fig 4.2) were
decolorized at pH 4.0 in the absence of MnII and the decolorizing activity
increased concomitantly with the increase in the concentration of MnII in the
reaction mixture. Enhancement in the sulfonphthalein decolorizing activity (SPDCA) by MnII is attributed to the MnP activities. However, MnII was observed to
inhibit 5th and 7th day bromophenol red (BPR) decolorizing activity (Fig. 4.3 and
4.4), 7th day bromocresol green (BCG) decolorizing activity (Fig. 4.4), and 5th, 7th
and 11th day bromophenol blue (BPB) decolorizing activity (Fig 4.3, 4.4 and 4.6).
MnP of Bjerkandera adusta and Pleurotus eryngii decolorized azo and
heterocyclic dyes in a MnII-independent manner [90].
Two interesting observations in this context were made pertaining
enzymatic decolorization of BPB, BCG and BPR. First, decolorization occurred in
the absence of MnII. Second, the inhibition of the decolorization reaction was
proportionate to the concentration of MnII. Decolorization of BPB, BCG and BPR
in the absence of MnII and inhibition by MnII led us to investigate the role of MIP
in the SP-DCA.
45
Inhibitory influence of MnII on the decolorization of SP dyes was stronger
and prominent at pH 3.5 than at 4 and was a function of MnII conc (Fig 4.3-4.6).
Halogenated SP dyes (BCG, BCP and BPB) are poorer substrates of MnP and
the extent of inhibition in the presence of MnII increased with the degree of
halogenation of these dyes (Fig 4.3-3.6).
O
S
R2'
O
R1'
O
R1
R2
OH
OH
R3'
Sulfonphthalein
R3
R1=R 1'
R2=R 2'
R3=R 3'
Bromocresol green
CH3
Br
Br
Bromocresol purple
H
CH3
Br
Bromophenol blue
H
Br
Br
Bromophenol red
H
Br
H
CH3
H
H
o-Cresol red
H
H
CH3
Phenol red
H
H
H
dyes
m-Cresol purple
Figure 4.2 General structure of sulfonphthalein dyes
46
MIP decolorized BPB better than BCP and BCG. MIP -catalyzed
decolorization of phenol red (PR), ortho-cresol red (o-CR) and meta-cresol purple
(m-CP) was less than the halogenated SP chromophores at pH 3.5 (Fig 4.3-4.6).
The increase in the degree of halogenation of SP dyes makes them better
Sulfonphthalein dye decolorizing activity (U/L)
substrates for MIP.
Figure 4.3 Influence of MnII on the sulfonphthalein decolorizing activity at pH 3.5
and 4.0 extracted from 5-day old wheat straw infested by Pleurotus ostreatus
MnP-catalyzed decolori zation of o-CR is better than m-CP. This indicates
that methyl group in ortho position (o-CR) is favorable than in meta position (mCP) for the decolorization reaction (Fig 3.2). The apparent Km values (Table 4.1)
47
of o-CR (26 mM) and m-CP (40 mM) provided the evidence of influence of
methyl group on the decolorization of the two dyes by MnP. Thus position of
auxochrome on SP chromophore is important for the decolorization.
MnP-catalyzed decolorizing activity of BCP was less than o-CR. Bromine
group on ortho position (BCP) was less effective than methyl group in ortho
position (o-CR) on the chromophore (Fig 4.2) for the decolorization by MnP. This
Sulfonphthalein dye decolorizing activity (U/L)
is evident from the higher K m for BCP (36 mM) and lower K m for o-CR (26 mM).
Figure 4.4 Influence of MnII on the sulfonphthalein decolorizing activity a t pH 3.5
and 4.0 extracted from 7-day old wheat straw infested by Pleurotus ostreatus
48
An additional bromine group on SP chromophore further impeded the
decolorizing ability of MnP. BPB and BCG (having two bromine) were
decolorized less than BCP (having one bromine) as evident from the kinetic
constants (Table 4.1). Kinetic constants (V max and Km) indicated that PR which
has low values of Km is catalytically more favorable for decolorization by MnP
Sulfonphthalein dye decolorizing activity (U/L)
(Table 4.1). PR served as a poor substrate for MIP.
Figure 4.5 Influence of MnII on the sulfonphthalein decolorizing activity at pH 3.5
and 4.0 extracted from 9-day old wheat straw infested by Pleurotus ostreatus
The order of preference of SP dyes as substrate is reverse in case of MnPand MIP-catalyzed decolorization. Free phenolic group present on the
49
chromophore of SP dyes (Fig 4.2) make them susceptible to attack by MnP
whereas presence of halogen (bromine) group (Fig 4.2) on SP chromophore
formulate them as a better substrate for MIP. Thus position, type and numbers of
the auxochromes on SP chromophores (Table 3.1) affect the enzymatic
decolorization of SP dyes. MIP appears to be a better candidate than MnP for the
Sulfonphthalein dye decolorizing activity (U/L)
degradation of halogenated aromatic compounds.
Figure3.6 Influence of MnII on the sulfonphthalein decolorizing activity at
pH 3.5 and 4.0 extracted from 11-day old wheat straw infested by
Pleurotus ostreatus
50
Table 4.1 Kinetic constants for sulfonphthalein dyes of manganese peroxidase
produced by Pleurotus ostreatus during solid-state fermentation of wheat straw
Sulfonphthalein
λmax
ε
Km
Vmax
dyes
(nm)
(M-1 cm-1)
(mM)
(U/ml)
Bromocresol green (BCG)
621
39569
117
0.125
Bromocresol purple (BCP)
593
60865
36
1
Bromophenol blue (BPB)
583
52663
78
0.066
Bromophenol red (BPR)
564
10735
69
0.065
m-Cresol purple (m-CP)
582
32406
40
0.133
o-Cresol red (o-CR)
577
41931
26
0.286
Phenol red (PR)
564
30737
16
0.057
PR-DCA (U/L)
100
75
50
25
0
0
5
10
15
NaN3 (µM)
Figure 4.7 Influence of sodium azide on Phenol red decolorizing activity (PRDCA) by manganese-independent peroxidase at pH 3.5 (¦ ) & 4.0 (? ) and
manganese peroxidase at pH 3.5 (?) & 4.0 (?)
White rot fungi characteristically produce several isozymes of ligninolytic
enzymes [22]. Variations in the enzymatic decolorization of SP dyes by these
51
activities produced on different days reflect differences in the composition of
isozymes of MnP and MIP activities. Difference (kinetics constants and also
substrate specificity) in the decolorization of SP dyes by enzyme preparations of
various days substantiates the observation made above (Fig 4.3-4.6).
NaN3, an oxidase inhibitor, was employed for the assessment of the nature
of decolorizing reactions of SP dyes. NaN3 inhibited the phenol red decolorizing
activity (PR-DCA) by MnP at pH 4.0. PR proved to be a better substrate for MnP
(Fig 3.7). This explained the oxidative nature of SP-decolorization reaction
catalyzed by MnP and MIP.
Chapter Published: Shrivastava, R., Christian, V., and Vyas, B. R. M. (2005)
Enzymatic decolorization of sulfonphthalein dyes. Enz. Microb. Technol. 36: 333337
52
CHAPTER 5
HORSERADISH PEROXIDASE:
DECOLORIZATION OF TEXTILE AND
TRIPHENYL METHANE DYES
5.1 Introduction
T
he word "horse" (as applied in "horseradish") is believed to denote
large size and coarseness. "Radish" comes from the Latin radix
meaning root. Horseradish peroxidase (E.C.1.11.1.7) (HRP) is a
heme protein with molecular weight of 40,000 Daltons and optimal pH 7.0 (154).
Seven isozymes of HRP have been reported (155-157). All contain photohemin
IX as prosthetic group. Neutral and amino sugars account for approximately 18%
of the enzyme. Horseradish peroxidase catalyzes oxidation of ascorbate,
ferrocyanide, cytochrome C and the leuco form of many dyes; the reaction
mechanism observed is as follow
HRP + H2O2 ? Compound I
Compound I + AH2 (reduced substrate) ? Compound II + AH?
Compound II + AH2 ? HRP + AH
2AH ? Oxidized Product
Bioremediation is a pollution control technology that uses biological
systems to catalyze the degradation or transformation of various toxic chemicals
to less harmful forms. The general approaches to bioremediation are to enhance
natural biodegradation by native organisms (intrinsic bioremediation), to carry out
environmental modification by applying nutrients or aeration (bio-stimulation) or
through
addition of
microorganisms (bio-augmentation). Unlike conventional
technologies, bioremediation can be carried out on-site. Bioremediation is limited
in the number of toxic materials it can handle, but where applicable, it is cost-
53
effective (158). Horseradish peroxidase is known to degrade recalcitrant organic
compounds such as phenol, substituted phenols and various synthetic dyes
(159). Present study focuses on the ability of HRP to degrade dyes, model of
xenobionts, of varied classes (textile and triphenyl methane).
5.2 Materials and Methods
5.2.1. Enzyme: Horseradish peroxidase (HRP) was obtained from SRL
Pvt. Ltd. The purified enzyme was preserved at -20oC.
5.2.2. Textile Dyes: Reactive black 5, Remazol black HR, Reactive orange
13, Reactive orange 16, Remazol red H8B, Remazol magenta HB, Remazol
turquoise blue G, Remazol lemon yellow F4, Remazol golden yellow R, Reactive
brown 18 and Remazol brilliant blue R were obtained from the dye manufacturing
and textile processing industries.
5.2.3. Triphenyl methane Dyes: Brilliant green, Methyl blue and Fast green
were procured from Hi-Media (India).
5.2.4. Enzyme assays: Reaction mixture (2 ml) for HRP contained 100
µmol sodium tartarate buffer (pH 2.5, 3.0 and 3.5), succinate lactate buffer (pH
4.0 and 4.5) and sodium citrate buffer (pH 5.0 and 5.5), 150 ppm textile dye or 75
ppm triphenyl methane dyes, 0.4 µmol H2 O2 and the purified enzyme. One unit
enzyme activity is defined as the amount of activity that consumes 1 µmol of the
triphenyl methane dye or produces a change of one absorbance unit per min
upon oxidation of textile dye at its λmax.
5.3 Results and Discussion
Horseradish peroxidase (HRP) derived from Ipomea palmata and
Saccharum spontaneum has been shown to degrade wide variety of textile dyes
and anthraquinonic dye, RBBR (159, 160). We investigated the decolorization of
54
eleven textile dyes and three triphenyl methane dyes by commercially available
purified HRP.
5.3.1. Decolorization of triphenyl methane dyes: Triphenyl methane dyes,
methyl blue (MB) and fast green (FG), were decolorized by Horseradish
peroxidase (HRP). Brilliant green (BG) resisted decolorization by HRP (Fig 5.1).
Methyl blue was the best decolorized triphenyl methane dye by HRP.
Decolorization of MB occurred above pH 4 and decolorization remained constant
above pH 5. Decolorization of FG also occurred above pH 4 and it increased till
pH 5.5. Decolorization of FG suggests the decolorization process needs to be
studied over a broader pH range.
30
BG
20
10
% Decolorization
0
30
MB
20
10
0
15
FG
10
5
0
2
3
pH
4
5
6
Figure 5.1 Decolorization of triphenyl methane dyes Brilliant green (BG),
Methyl blue and (MB), and Fast green (FG) by horseradish peroxidase.
55
5.3.2. Decolorization of textile dyes: Horseradish peroxidase
decolorized five of the textile dyes investigated, namely, reactive black 5 (RB5),
reactive orange 16 (RO16), remazol brilliant blue R (RBBR), remazol turquoise
blue G (RTBG), and remazol red H8B (RRH8B). Remazol black HR (BHR),
reactive orange 13 (RO13), remazol magenta HB (RMHB), remazol lemon yellow
F4 (LYF4), remazol golden yellow R (GYR), reactive brown 21 (RB21), were not
decolorized by the HRP (Fig. 5.2).
Decolorization of RB5 occurred above pH 3.5 which increased beyond
pH 4.5 and remained constant till pH 5.5. Similar results were obtained with
RBBR (Fig 5.2). Remazol red H8B (RRH8B) was decolorized over the pH range
of 3.5 to 5.5. The increase in the decolorization of RRH8B was the function of
pH, with the rise in the pH the decolorization also increased substantially.
Decolorization of the dyes, RO16 and RTBG, occurred above pH 4.5 and 5,
respectively.
5.3.3. Influence of pH: Decolorization of textile dyes and triphenyl methane
dyes by HRP occurred over a broad range of pH 3.5-5.5. But the significant
decolorization, in most of the cases, was at the optimal pH of HRP. Textile dyes,
RO16 and RTBG, and triphenyl methane dye, FG, were strict in pH selection for
decolorization by HRP. Their decolorization occurred at particular pH (4 or 4.5).
This implies that in scheming bioremediation, recognizing optimal pH for certain
class of dyes is must and under defined pH only the system can be made
efficient.
One of the two black dyes, RB5 was decolorized but not the other, BHR.
Similarly, of orange dyes, RO16 and RO13, only RO16 was decolorized. Similar
observation was also made in the case of green triphenyl methane dyes, BG and
FG, where, FG was decolorized but BG resisted decolorization. The
investigations done by Pasti-Grigsby et. al. have attributed the reason to be the
56
30
30
RB5
20
20
10
10
0
0
30
30
RO16
20
20
10
10
0
0
80
BHR
RO13
30
RBBR
60
RTBG
20
40
10
20
0
0
30
30
RRH8B
20
20
10
10
0
0
30
RMHB
30
LYF4
20
20
10
10
0
0
GYR
2
30
3
pH
4
5
6
BrH4R
20
10
0
2
3
pH
4
5
6
Figure 5.1 Decolorization of textile dyes by horseradish peroxidase
(?):Reactive black 5 (RB5); Remazol black HR (BHR); Reactive orange16
(RO16); Reactive orange 13 (RO13); Remazol brilliant blue R (RBBR);
Remazol turquoise blue G (RTBG); Remazol red H8B (RRH8B); Remazol
magenta HB (RMHB); Remazol lemon yellow F4 (LYF4); Remazol golden
yellow R (GYR); Reactive brown 18 (RB18)
57
differences in the electron distribution and charge density along with the stericdistribution in different dyes (126).
5.3.4. Structure function relationship of decolorization of dyes by
Horseradish peroxidase: Among the three triphenyl methane dyes, FG and MB
was decolorized and BG resisted decolorization by HRP. MB has three 4-amino
benzene sulfonate group and presence of this group makes MB decolorize better
compared to FG by HRP (Fig. 3.4, Chapter 3). Presence of two 3-[2-(ethylamino)
ethyl] benzene sulfonate group with one hydroxyl group and one sulfonic acid
group in FG makes this dye a substrate for HRP. The presence of two N,Ndiethyl amine group in the structure of BG prevents the HRP to use it as a
substrate (Fig. 3.4, Chapter 3). The selection of triphenyl methane dyes as a
substrate for decolorization by HRP is similar to that of manganese peroxidase of
Irpex lacteus (Chapter 3). The order of preference of triphenyl methane dyes as
substrate for HRP mediated decolorization is MB > FG.
Among eleven textile dyes investigated for decolorization, HRP was able
to decolorize five of the dyes namely, reactive black 5 (RB5), reactive orange 16
(RO16), remazol brilliant blue R (RBBR), remazol turquoise blue G (RTBG), and
remazol red H8B (RRH8B). Between two orange dyes, RO16 and RO13, RO16
was decolorized whereas, RO13 resisted decolorization by HRP. RO16 has an
intermediate compound vinyl sulphone azotized to naphthalene and RO13 has
an intermediate compound sulpho taubias acid azotized to naphthalene moiety
and an additional intermediate, cyanuric chloride (Fig. 3.3, Chapter 3).
Azotization of vinyl sulphone intermediate to naphthalene is favourable for
decolorization by HRP than azotization of sulpho taubias acid. Diazotization of
vinyl sulphone group leads to the formation of RB5 (Fig. 3.3, Chapter 3).
Therefore, presence of vinyl sulphone group as an intermediate in dye (like, RB5
and RO16) and absence of any halogenated intermediate (for example cyanuric
chloride) favours the decolorization of dye by HRP.
58
RBBR is an anthraquinone dye with a vinyl sulphone group (Fig. 3.3,
Chapter 3). Decolorization of an anthraquinonic dye, RBBR, by HRP is
suggestive of HRP as an important tool in degradation of xenobionts having
anthraquinonic intermediates.
The selection of dye as a substrate for decolorization by HRP is similar to
that of manganese peroxidase of Irpex lacteus. HRP preferred the dyes with
simple structure and dyes that do not have halogenated intermediates. The dye
having the simple phenolic moiety is decolorized better by HRP. Thus, HRP and
its reaction mechanism in degradation of dyes need due attention as a candidate
and in defining the strategies for the development of promising industrial wastewater treatment system.
59
CHAPTER 6
LIGNINOLYTIC ENZYMES OF IRPEX LACTEUS:
DECOLORIZATION OF EFFLUENTS
6.1 Introduction
W
hite-rot basidiomycetes (WRB) produce various isoforms of
extracellular oxidases namely laccase, manganese peroxidase
(MnP) and lignin peroxidase (LIP ) that are involved in the
degradation of lignin in the natural lignocellulosic substrates
(37). This ligninolytic system of WRB is directly involved also in the degradation
of various xenobiotics compounds and dyes (145, 146, 151, 152).
The textile industry, by far the most avid user of synthetic dyes, is in need
of eco-efficient solutions for its colored effluents. Chemical products used in
textile processing affect the environment by their effluents that can be
voluminous, colored and varied (74, 109). Thoug h, the physical and chemical
methods offer some solutions to the problems, it is not affordable by the unit
operators. Biological degradation is recognized as the most effective and ecofriendly method for degrading the dyes present in the wastes.
Research over a period of two decades has provided insight into the nonspecific enzyme system of white rot fungi, which oxidizes the recalcitrant dyes
(162). The decolorization and detoxification potential of WRB can be harnessed;
thanks to the emerging knowledge of the physiology and enzymology of these
organisms. The knowledge gathered needs to be transformed into reliable and
robust waste treatment processes. This chapter evaluates prospective use of
ligninolytic enzymes of WRB, Irpex lacteus, for the treatment of industrial
effluents, particularly dye containing effluents.
60
6.2 Materials and Methods
6.2.1. Microorganism: Irpex lacteus Fr. 238 617/93 obtained from Culture
Collection of Basidiomycetes (CCBAS), Prague (Kind gift from V. Šašek). The
strain is maintained on malt extract agar plates at 4 ºC.
6.2.2. Effluents: Effluents, E1 and E2, were obtained from dyeing and
printing industries located in Jetpur, Gujarat.
6.2.3. Production of ligninolytic enzymes by I. lacteus: Solid state
fermentation of wheat straw by I. lacteus was performed as described previously
[136]. To the culture flasks, harvested on 18th day, 100 ml of phosphate buffer
(0.1 M, pH 6.5) containing 0.1 M NaCl was added, contents were gently beaten
and incubated on rotary shaker for 1 h. Liquor obtained was then filtered and
spun (6000 rpm, 15 min, 5oC) before being subjected to ammonium sulfate
precipitation (80% saturation). Precipitates separated upon centrifugation (6000
rpm, 15 min, 5oC) were dissolved in the minimum amount of phosphate buffer
(0.1 M, pH 6.5). Enzyme extract thus obtained was desalted using PD10 column
(Pharmacia) and stored in small aliquots at -5ºC for further biochemical analysis.
6.2.4. Enzyme assays: Manganese peroxidase (MnP) and manganeseindependent peroxidase (MIP) activities were determined as described by Vyas
et al. (136). Lignin peroxidase activity was estimated according to Tien and Kirk
(34). One unit of enzyme activity is defined as the amount of activity that will
produce one µmol of the product per min upon oxidation of the substrate in the
reaction mixture under the assay conditions.
6.2.5. Effluent decolorizing enzyme assays: Effluent decolorizing activity of
MnP, MIP and LIP was assayed using respective reaction mixtures. Reaction
mixture (2 ml) for MnP activity contained 100 µmol buffer, 0.2 µmol MnSO4 , 150
ppm effluents, 0.4 µmol H2O2 and the enzyme extract. Sodium tartarate buffer for
pH 2.5, 3.0 and 3.5, succinate lactate buffer for pH 4.0 and 4.5, and sodium
61
citrate buffer for pH 5.0 and 5.5 were used. Reaction mixture for MIP activity was
similar to that of MnP but MnSO4 was not added. Reaction mixture for LIP activity
was similar to that of MnP but veratryl alcohol (4 µmol) was added instead of
MnSO4 . One unit enzyme activity is defined as the amount of activity that
produces a change of one absorbance unit per min upon oxidation of textile dye
at its λ max.
6.3 Results and Discussion
The ligninolytic system of white-rot basidiomycetes (WRB) is
directly involved in the degradation of many xenobionts. The decolorization of
dyes as well as its use to measure ligninolytic activities was first reported for
Phanerochaete chrysosporium [76]. Thereafter, Pleurotus ostreatus has also
been shown to produce different dye decolorizing activities [25, 26, 122]. We
report the effluents decolorizing activities associated with manganese peroxidase
(MnP), manganese-independent peroxidase (MIP) and lignin peroxidase (LIP)
produced by Irpex lacteus.
6.3.1. Influence of pH on ligninolytic enzyme activities: Extracellular
enzyme extracts prepared from Irpex lacteus-infested wheat straw possessed
MnP MIP and LIP activities (Fig 6.1). The optimal pH of MnP was observed to be
4 and that of LIP was 3. MIP activity was maximal at pH 3.5 and thereafter
decreased gradually.
Enzyme activity (U/l)
300
200
100
0
2
2.5
3
pH
3.5
4
4.5
Figure 6.1 MnP activity (-?-), MIP activity (-?-) and LIP activity (-?-) at
varying pH of Irpex lacteus growing on wheat straw.
62
6.3.2. Influence of H2O2 on ligninolytic enzyme activities: All the ligninolytic
activities showed positive influence to the increasing concentration of hydrogen
peroxide (H2O2). MnP and MIP activities were maximum at the final concentration
of H2 O2 (100 µM) examined (Fig. 6.2). LIP activity was also maximal at 500 µM of
H2O2 (Fig. 6.2). Mechanisms involved in MnP-catalyzed and LIP -catalyzed
reactions have been elucidated (32, 105). H2O2 , the primary substrate of MnP,
generates oxidized species of MnP (2 electron deficient Compound I) that
oxidizes MnII to MnIII which is stabilized by organic acid chelators. Similarly LIP
Compound I generated in the presence of H2 O2 oxidizes veratryl alcohol to
veratryl alcohol cation radical. MnIII and veratryl alcohol cation radical in turn act
as low molecular mass, diffusible oxidants attacking organic molecules
nonspecifically via hydrogen and one electron abstraction [32, 105, 153].
MnP (U/ml)
1
0.75
0.5
0.25
0
0
25
50
H2 O2 (µM)
75
100
MIP (U/ml)
0.5
0.25
0
0
25
50
H2O2 (µM)
63
75
100
LIP (U/ml)
0.15
0.1
0.05
0
0
100
200
300
400
500
H2O2 (µM)
Figure 6.2: Influence of H2 O2 on ligninolytic enzyme activities produced by
Irpex lacteus growing on wheat straw
6.3.3. Decolorization of Effluent E1 by ligninolytic enzyme activities:
Ligninolytic activities investigated decolorized the effluent E1 completely. MnP
was found to be the best ligninolytic activity associated with decolorization of
effluent E1 (Fig. 6.3). MIP was better than LIP in the decolorization of effluent E1.
During the enzymatic decolorization absorbance in the UV-region decreased
drastically and ultimately resulted in the complete loss of the peak in the UVregion. Decolorization pattern did not change in the pH range 3-5.5. The pH of
the reaction mixture did not exert any influence on decolorization of E1 by the
ligninolytic activities. The physico-chemical properties of E1 are mentioned in
Table 6.1.
6.3.4. Decolorization of effluent E2 by ligninolytic enzyme activities:
Ligninolytic activities investigated also decolorized the effluent E2 completely.
MnP and MIP were found to be the better ligninolytic activities associated with
the decolorization of effluent E2 (Fig. 6.4). LIP also contributed to the
decolorization of effluent E2. Similar to the decolorization of E1, during the
decolorization of E2 simultaneous loss of the peak in UV-region was observed.
Decolorization of E2 by the ligninolytic activities, MnP and MIP , was not
influenced by the pH of the reaction mixture. LIP -associated decolorization of E2
64
showed loss in the efficiency with the increasing pH. The physico-chemical
properties of E2 are mentioned in Table 6.1.
4
pH 3
2
4
pH 4.5
0
2
4
Absorbance
Absorbance
pH 3.5
2
0
4
pH 5
0
2
4
pH 4
0
150
350
550
750
wavelength (nm)
2
0
150
350
550
750
wavelength (nm)
Figure 6.3 UV-visible spectra of effluent E1 before (??? ) and after incubation
with MnP (………), MIP (- - - -) and LIP (-·-·-·-)
65
4
pH 3
2
4
pH 4.5
0
2
4
Absorbance
Absorbance
pH 3.5
2
0
4
pH 5
0
2
4
pH 4
0
150
2
350
550
750
wavelength (nm)
0
150
350
550
750
wavelength (nm)
Figure 6.4 UV-visible spectra of effluent E2 before (??? ) and after incubation
with MnP (………), MIP (- - - -) and LIP (-·-·-·-)
66
Table 7.1 Physico-chemical characterization of effluents (E1 and E2) from
dyeing and printing industries
Physico-chemical
E-1
E-2
pH
6.45
6.8
Conductivity
5.79
5.24
Turbidity (NTU)a
0.4
1.3
solids
440
360
Copper (ppm)
0.022
0.081
Nitrite (ppm)
26
38
Zinc (ppm)
2.9
3.9
Chloride (ppm)
38
86
Sulfate (ppm)
666.4
780.9
parameter
Total dissolved
Ligninolytic activities investigated decolorized the effluents, E1 and E2
completely. The two major differences observed in the decolorization of E1 and
E2 were, firstly, the ligninolytic activities, MnP was the only activity that
decolorized E1 better than any other ligninolytic activities investigated, whereas,
in the case of E2, MnP and MIP were the activities that decolorized E2 better.
Secondly, the ligninolytic activities associated to decolorization of E1 showed no
influence of pH and decolorized the effluent over the range of pH of investigated,
whereas, efficiency of LIP to decolorize E2 decreased with the rise in pH.
The differences in the decolorization of E1 and E2 can be attributed to the
differences in their physico-chemical properties. Though the pH has high impact
on ligninolytic activities, as in case of MnP where the pH at which the enzyme is
stable is 7 and the pH at which it is highly active is 4 or 4.5 but the pH in MnP
and MIP associated decolorization of effluents, E1 and E2, had no influence and
the effluents were decolorized with same efficiency over the range of pH
investigated.
67
CHAPTER 7
D ECOLORIZATION OF DYES AND EFFLUENTS BY
FENTON AND FENTON-LIKE R EAGENTS
7.1 Introduction
M
any metals have special oxygen transfer properties which
improve the utility of hydrogen peroxide. By far, the most
common of these is iron which, when us ed in the prescribed
manner, results in the generation of highly reactive hydroxyl
radicals (•OH). The Fenton's reaction has been known since 1894 by its
inventor H.J.H. Fenton (61) and is currently one of the most powerful oxidizing
reactions available. The reaction involves hydrogen peroxide and a ferrous
iron catalyst. The peroxide is broken down into a hydroxide ion and a hydroxyl
free radical. The hydroxyl free radical is the primary oxidizing species and can
be used to oxidize and break apart organic molecules.
[1] Fe II+ + H2O2? OH· + Fe III+ OH–
The common pathways for the white-rot fungal production of Fenton’s
reagents are mentioned below (107).
[2] 2Fe III+ + –O2C·CO2 – ? 2FeII+ + 2CO2
[3] FeII+ + O 2 ? Fe III+ + O2·–
[4] Fe II++ O 2· – + 2H + ? Fe III+ + H2O 2
–
[5] O 2· + H2O2 ?
OH· + OH – + O2
Toxic and refractory organic pollutants in industrial wastewater can be
degraded by advanced oxidation processes alone or in combination with
physico-chemical and biological processes (163). Of these oxidation methods,
Fenton's reagent (FeII/H2O 2) and Fenton-like reagents (CuII/H2O2), are
effective oxidants of large variety of organic pollutants. Fenton reagents are
also produced from diverse routes by lignin-degrading fungi (107). The
mechanism of decomposition of H2O2 and of oxidation of organic solutes by
Fenton's and Fenton-like reactions has been the subject of numerous studies
68
(163, 164). A recent study however, reported decolorization of synthetic dyes
by Fenton system (165).
In our report, we present the results of our investigations of
decolorization of textile, triphenyl methane dye and effluents from dyeing and
printing industries by Fenton Reagent (FeII/H2O 2) and Fenton-like reagents
(CuII/H2O2).
7.2 Materials and Methods
7.2.1 Chemicals: FeII as FeSO4, CuII as CuSO4 and hydrogen peroxide
were added as aqueous solutions to the reaction mixture.
7.2.2 Textile Dyes : Reactive black 5, Remazol black HR , Reactive
orange 13, Reactive orange 16, Remazol red H8B, Remazol magenta HB,
Remazol turquoise blue G, Remazol lemon yellow F4, Remazol golden yellow
R, Reactive brown 18 and Remazol brilliant blue R were obtained from the
dye manufacturing and textile processing industries.
7.2.3. Triphenyl methane Dyes: Brilliant green, Methyl blue and Fast
green were procured from Hi-Media (India).
7.2.4. Effluents : Effluents, E1 and E2, were obtained from various
dyeing and printing industries.
7.2.5. Decolorization of dyes: Textile dyes (150 ppm), triphenyl
methane dyes (75 ppm) and effluents (3/4 th of effluent diluted to 1/4th by distill
water) were added to the reaction mixture as aqueous solutions. Reaction
mixture (2 ml) for Fenton reagent contained 1 mM FeSO4 and 10 mM
hydrogen peroxide. Reaction mixture for Fenton-like reagent was similar to
that of Fenton reagent but contained 1 mM CuSO4. The dyes were incubated
with Fenton and Fenton-like reagent and scanned in the range of 200-800 nm,
before and after decolorization using Shimadzu UV-1601 spectrophotometer.
The decolorization was initiated by the addition of hydrogen peroxide to the
reac tion mixture. The incubation proceeded at 18 °C in the dark.
69
7.3 Results and Discussion
Ligninolytic enzyme system of white-rot basidiomycetes (WRB)
generates hydroxyl radical, a reactive oxygen species, which plays important
role in wood decay. Hydroxyl radical is the strongest oxidant that occurs in
aqueous system. It reacts with all organic molecules by abstracting hydrogen
from aliphatic structures or by adding an electrophile to aromatic structures
(166). Hydroxyl radical is found to be generated in all the three major types of
wood decay caused by white -rot, brown-rot and soft-rot fungi (163, 167, 168).
7.3.1. Decolorization of textile dyes by Fenton reagent and Fenton-like
reagent: Textile dyes were completely decolorized by Fenton reagent (Fig.
7.1). Decolorization initiated immediately with the addition of hydrogen
peroxide. Fenton reagent-mediated decolorization of dyes led to the loss of
absorbance maxima in visible region and decrease in the absorbance in UVregion. Textile dye, Remazol turquois e blue G (RTBG), was decolorized less
as compared to the other dyes. Decolorization of dyes, remazol brilliant blue
R (RBBR), reactive orange 13 (RO13), remazol red H8B (RRH8B), RTBG,
and remazol black HR (BHR) resulted in the generation of new peak in the
UV-region.
Textile dyes were completely decolorized by Fenton-like reagent (Fig.
7.1). Dyes, RO13, Remazol lemon yellow F4 (LYF4), Reactive brown 18
(RB18) and BHR, resisted immediate decolorization by Fenton-like reagent.
Decolorization of textile dyes by Fenton-like reagent led to the generation of
new peaks in the visible as well as in the UV-region. The new peaks,
generated as a part of decolorization reaction, shifted towards the lower
wavelength. It is important to report that decolorization of the some of the
dyes, reactive orange 16 (RO16), RRH8B, and LYF4, initiated with the
increase in absorption, shifting the peak towards the higher wavelength. The
reports on decolorization of dyes by white-rot fungi have shown the process of
oligomerization before degradation (169).
70
4.5
RBB
RB 5
Absorbance
Absorbance
4.5
3.0
1.5
0.0
200
400
600
3.0
1.5
0.0
200
800
RBBR
400
RO13
RO3R
4.5
Absorbance
Absorbance
4.5
3.0
1.5
0.0
200
400
600
600
3.0
1.5
0.0
200
800
RO16
ROH2R
400
λ (nm)
4.5
RRH8B
3.0
1.5
0.0
200
400
600
800
4.5
Absorbance
Absorbance
800
1.5
400
λ (nm)
RTBG
1.5
400
600
3.0
0.0
200
3.0
0.0
200
800
RMHB
λ (nm)
4.5
600
λ (nm)
Absorbance
Absorbance
4.5
800
λ (nm)
λ (nm)
600
3.0
1.5
0.0
200
800
λ (nm)
LY
LYF4
400
600
λ (nm)
71
800
4.5
GYF4
GYR
Absorbance
Absorbance
4.5
3.0
1.5
0.0
200
400
600
3.0
1.5
0.0
200
800
BRH4R
RB18
400
4.5
Absorbance
600
800
λ (nm)
λ (nm)
BHR
3.0
1.5
0.0
200
400
600
800
λ (nm)
Figure 7.1 UV-visible spectra of textile dyes before (? ? ? ) and after 20 min of
incubation with Fenton reagent (- - - -) and Fenton-like reagent (………):
Reactive black 5 (RB5); Remazol brilliant blue R (RBBR); Reactive orange 13
(RO13); Reactive orange 16 (RO16); Remazol red H8B (RRH8B); Remazol
magenta HB (RMH B); Remazol turquoise blue G (RTBG); Remazol lemon
yellow F4 (LYF4); Remazol golden yellow R (GYR); Reactive brown 18
(RB18); Remazol black HR (BHR).
7.3.2. Decolorization of triphenyl methane dyes by Fenton reagent and
Fenton-like reagent: The three triphenyl methane dyes investigated were
completely decolorized by the Fenton reagent. The decolorization of the dyes,
brilliant green (BG), methyl blue (MB) and fast green (FG) was similar to the
most of the textile dyes (Fig. 7.2). Decolorization proceeded with the loss of
peaks in the visible as well as decrease in the absorbance in UV-region.
Among the triphenyl methane dyes, BG was the only dye that was
decolorized by the Fenton-like reagent. Decolorization of BG by Fenton-like
reagent proceeded slowly (Fig. 7.2). The phenomena of oligomerization as
72
4.5
BG
Absorbance
Absorbance
4.5
3.0
1.5
0.0
200
400
600
MB
3.0
1.5
0.0
200
800
400
λ (nm)
4.5
Absorbance
600
800
λ (nm)
FG
3.0
1.5
0.0
200
400
600
800
λ (nm)
Figure 7.2 UV-visible spectra of triphenyl methane dyes before (??? ) and
after 20 min of incubation with Fenton reagent (- - - -) and Fenton-like reagent
(………): Brilliant green (BG); Methylene blue (MB); Fast green (FG).
observed in the decolorization of some of the textile dyes, was also
experiential in the decolorization of BG. Therefore, Fenton reagent is more
viable option in degradation studies concerning triphenyl methane dyes.
7.3.3. Decolorization of effluents by Fenton reagent and Fenton-like
reagent: Despite of difference in the physico-chemical properties (Table 7.1)
of the effluents, E1 and E2, they were decolorized efficiently by both the
systems, Fenton reagent and Fenton-like reagent (Fig. 7.3). Decolorization of
the effluents followed by loss in absorbance in visible as well as UV-region.
Fenton-like reagent was more efficient in decolorization of effluent E2.
73
4.5
E1
Absorbance
Absorbance
4.5
3.0
1.5
0.0
200
400
600
3.0
1.5
0.0
200
800
E2
400
600
800
λ (nm)
λ (nm)
Figure 7.3 UV-visible spectra of effluents before (? ? ? ) and after 20 min of
incubation with Fenton reagent (- - - -) and Fenton-like reagent (………):
Effluent 1 (E1); Effluent 2 (E2).
Decolorization of E1 by both the systems investigated showed
resistance, but E2 was decolorized better when compared to the former. The
better decolorization of E2 can be attributed to its physico-chemical properties
(Table 7.1). The conductivity and total dissolved solids of E2 were lower when
compare to E1. The pH of the E2 was higher, towards neutral, and this
assisted in rapid decolorization of E2 by both the systems investigated. It has
been reported that pH plays a very vital role in decolorization by redox
coupling system like, Fenton reagent (108, 164).
Decolorization of dyes and effluents by Fenton reagent and Fenton-like
reagent initiates with the addition of hydrogen peroxide. Dyes and effluents,
under investigation, were decolorized completely by Fenton reagent and
Fenton-like reagent and showed hypsochromic shift as well as appearance of
new peak towards lower wavelength (Fig. 1-3). Shift in the absorption
maxima, in both the cases, during decolorization of dyes implicates the
process to be occurring in steps where intermediates are generated. Similar
results were obtained with decolorization by modified Fenton reagent using
Cu/pyridine/hydrogen peroxide (108). Hypsochromic shift is suggestive of
degradation of the molecule through a series of intermediates (26, 149).
74
Table 7.1 Physico-chemical characterization of effluents (E1 and E2) from
dyeing and printing industries
Physico-chemical
parameter
E-1
E-2
6.45
6.8
5.79
5.24
0.4
1.3
Total dissolved
solids
440
360
Copper (ppm)
0.022
0.081
Nitrite (ppm)
26
38
Zinc (ppm)
2.9
3.9
Chloride (ppm)
38
86
Sulfate (ppm)
666.4
780.9
pH
Conductivity
Turbidity (NTU)
a
Fenton reagent is better than Fenton-like reagent for the decolorization
of dyes; firstly, Fenton reagent decolorized the dyes rapidly compare to
Fenton-like reagent, secondly, Fenton reagent is more powerful redox system
as it was able to decrease the absorption even in the UV -region, thirdly,
Fenton reagent is very efficient in programming the remediation of textile
effluents as it was able to decolorize most of the dyes completely. Redox
system and mechanism of Fenton reagent needs more attention to develop it
as a potent tool against the water pollution abatement programs.
75
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IMPLICATIONS & SPECULATIONS
OF THE
INVESTIGATIONS
T
he investigations were aimed on the potential of the lignin
degrading basidiomycetes in the degradation of xenobion ts. Dyes
were selected as a model of xenobionts as they represent
structural diversity. Investigation was also focused on the dye decolorization
by Fenton reagents, Fenton-like reagents and Horseradish peroxidase for
considering an equivalent measure to evaluate the ligninolytic system of
white-rot basidiomycetes (WRB).
Irpex lacteus and Strain F were screened for their ability to decolorize
textile dyes on solid medium. The WRB, Strain F and I. lacteus were able to
decolorize the textile dyes under investigations. Variation in the decolorization
of similar color dyes was observed. This is indicative that structurally similar
dyes may be differentially decolorized by white rot fungi. Decolorization of
blue and black dyes was followed by the change in color of lower wavelength.
The degradation of dyes is not a single step reaction but follows generation of
many intermediates. Dye decolorization on solid media was under older
mycelia which always started from the point of inoculation and the zone of
decolorization followed the zone of growth. This suggests decolorization on
solid media as a part of secondary metabolic event. The correlation between
the zone of growth and zone of decolorization on solid medium can be
attributed to the involvement of ligninolytic activities in the decolorization.
Lignin-degrading white rot fungi have received worldwide attention for their
industrial use in bio-pulping, bio-bleaching, dye decolorization, and in
detoxifying
recalcitrant
environmental
pollutants.
Irpex
lacteus
is
a
cosmopolitan white rot basidiomycete typically inhabiting dead hard wood
trees. Decolorization of textile dyes and triphenyl methane dyes by ligninolytic
96
enzymes produced by I. lacteus during solid-state fermentation wheat straw
was studied. Irpex lacteus under identical conditions produced manganese
peroxidase (MnP), manganese-independent peroxidase (MIP), and lignin
peroxidase (LIP). One or the other ligninolytic activities decolorized all the
dyes under investigation. Manganese peroxidase was the only enzyme that
was
associated
to
the
decolorization
of
triphenyl
methane
dyes.
Decolorization of most of the textile dyes were attributed to MIP. Influence of
pH on dye decolorization by ligninolytic activities was studied. Decolorization
of textile dyes by MnP occurred at its optimal pH whereas, MIP associated
decolorization varied with respect to dyes. This suggests that in defining the
bioremediation program, pH must be given its due attention.
The white rot fungus Pleurotus ostreatus produced manganese peroxidase
(MnP) and manganese-independent peroxidase (MIP) activities during solid
state fermentation of wheat straw, a natural lignocellulosic substrate. Most of
the SP dyes were decolorized by MnP at pH 4.0. The higher Km for metacresol purple (40 µM) and lower Km for ortho-cresol red (26 µM) for MnP
activities explained the preference for the position of methyl group at ortho
than at meta on chromophore. Bromophenol blue decolorizing activity was
higher at pH 3.5 and decreased as the concentration of Mn II was increased.
SP-decolorizing activity was associated not only with MnP but also with MIP.
Additional bromine group along with the methyl group on SP chromophores
decreases the rate of decolorization. Bromination of sulfonphthalein
chromophore makes them the poorer substrate for MnP. This is evident from
the higher Km for bromocresol green (117 µM) when compared to bromocresol
purple (36 µM) and bromophenol blue (78 µM). The order of preference for
the SP dyes as substrate for the MnP- catalyzed decolorizing activity is
phenol red > ortho-cresol red > meta -cresol purple > bromophenol red >
bromocresol purple > bromophenol blue > bromocresol green and the order of
preference for the SP dyes as substrate for the MIP - catalyzed decolorizing
activity is bromocresol green > bromophenol blue > bromocresol purple >
bromophenol red > meta-cresol purple > ortho-cresol red > phenol red.
Inhibition of PR decolorizing activity by NaN3 provided the evidence of
decolorizing activity as an oxidative process.
97
Horseradish peroxidase (HRP) is known to degrade recalcitrant organic
compounds such as phenol, substituted phenols and various synthetic dyes
(159). Present study focuses on the ability of HRP to degrade dyes, model of
xenobionts, of varied classes (textile and triphenyl methane). We investigated
the decolorization of eleven textile dyes and three triphenyl methane dyes by
commercially available purified HRP. Horseradish peroxidase decolorized five
of the textile dyes investigated, namely, reactive black 5 (RB5), reactive
orange 16 (RO16), remazol brilliant blue R (RBBR), remazol turquoise blue G
(RTBG), and remazol red H8B (RRH8B). Influence of pH on decolorization
was also studied and HRP was able to decolorize dyes over broader pH
range. The selection of dye as a substrate for decolorization by HRP is similar
to that of manganese peroxidase of Irpex lacteus . HRP and its reaction
mechanism in degradation of dyes need due attention as a candidate and in
defining the strategies for the development of promising industrial waste-water
treatment system.
White-rot basidiomycetes (WRB) produce various is oforms of extracellular
oxidases namely laccase, manganese peroxidase (MnP) and lignin
peroxidase (LIP) that are involved in the degradation of lignin in the natural
lignocellulosic substrates. This chapter reports the effluents decolorizing
activities associated with manganese peroxidase (MnP), manganeseindependent peroxidase (MIP) and lignin peroxidase (LIP) produced by Irpex
lacteus. Influence of pH and H2O2 on the ligninolytic activities was studied.
Decolorization of effluents was studied by varying pH for each ligninolytic
activities . MnP was the only activity that decolorized E1 better than any other
ligninolytic activities investigated, whereas, in the case of E2, MnP and MIP
were the activities that decolorized E2 better. the ligninolytic activities
associated to decolorization of E1 showed no influence of pH and decolorized
the effluent over the range of pH of investigated, whereas, efficiency of LIP to
decolorize E2 decreased with the rise in pH. The differences in the
decolorization of E1 and E2 can be attributed to the differences in their
physico-chemical properties.
98
Fenton's reagent (FeII/H2O 2) and Fenton-like reagents (CuII/H2O2), are
effective oxidants of large variety of organic pollutants. Fenton reagents are
also produced from diverse routes by lignin-degrading fungi. The mechanism
of decomposition of H2O 2 and of oxidation of organic solutes by Fenton's and
Fenton-like reactions has been the subject of numerous studies. In our report,
we present the results of our investigations of decolorization of textile,
triphenyl methane dye and effluents from dyeing and printing industries by
Fenton
Reagent
(Fe II/H2O2)
and
Fenton-like
reagents
(CuII/H2O2).
Decolorization initiated immediately with the addition of hydrogen peroxide.
Dyes and effluents, under investigation, were decolorized completely by
Fenton reagent and Fenton-like reagent and showed hypsochromic shift. It is
interesting to report that decolorization of the some of the dyes, reactive
orange 16 (RO16), RRH8B, and LYF4, initiated with the increase in
absorption, shifting the peak towards the higher wavelength. The reports on
decolorization of dyes by white-rot fungi have shown the process of
oligomerization before degradation Fenton reagent is very efficient in
programming the remediatio n of textile effluents as it was able to decolorize
most of the dyes completely. Redox system and mechanism of Fenton
reagent needs more attention to develop it as a potent tool against the water
pollution abatement programs.
99
A PPENDIX I
SCHOLA RSHIPS A WARDE D
Senior
Research
Fellowship
from
Department
of
Biotechnology (DBT), Ministry of Science and Technology,
New Delhi, India (December, 2003 onwards).
100
APPENDIX II
LIST OF PUBLI CATIONS AND PRESENT ATIONS
PUBLICATIONS
1.
Shrivastava, R., Christian, V., and Vyas, B. R. M. (2004) Enzymatic
decolorization of sulfonphthalein dyes. Enz. Microb. Technol. 36:
333-337
2.
Christian, V., Shrivastava, R., Shukla, D., Modi, H. A., Vyas, B.R.M.
(2004) Mediator role of veratryl alcohol in the lignin peroxidasecatalyzed oxidative decolorization of Remazol Brilliant Blue R. Enz.
Microb. Technol. 36: 327-332
3.
Christian, V., Shrivastava, R., Shukla, D., Modi, H. A., Vyas, B.R.M.
(2004) Degradation of xenobiotic compounds by lignin-degrading
white-rot fungi: Enzymology and mechanisms involved. Ind. J. Exptl.
Biol. (In-Press).
4.
Christian, V., Shrivastava, R., Modi, H. A., Vyas, B.R.M. (2004)
Physiology and biochemistry of lignin degradation by white-rot fungi.
(Review article) J. Tissue Res. 3(2): 165-173 (2004).
5.
Christian, V.V., Shrivastava, R., Novotny, C., Vyas, B. R. M. (2003)
Decolorization of sulfonphthalein by manganese peroxidase activity
of white rot fungus Phanerochaete chrysosporium. Folia Microbiol.
48(6): 771-776
101
RESEARCH ARTICLES COMMUNICATED
1.
Shrivastava, R., Christian, V., Shukla, D., Modi, H.A., and Vyas,
B.R.M. (2004) White-rot basidiomycetes: degradation of dyes (Review
Article), Communicated to Rev. Env. Sci. Biotechnol.
2.
Shrivastava, R., Christian, V., Shukla, D., Modi, H.A., and Vyas,
B.R.M. (2004) Ligninolytic enzymes of Irpex lacteus : decolorization of
textile
dyes,
triphenyl
methane
dyes
and
textile
effluents,
Communicated to Folia Microbiol.
3.
Shrivastava, R., Christian, V., Shukla, D., Modi, H.A., and Vyas,
B.R.M. (2004) Decolorization of textile dyes, triphenyl methane dyes
and textile effluents by Fenton reagent and Fenton-like reagent,
Communicated to Chemosphere
4.
Shrivastava, R., Christian, V., Shukla, D., Modi, H.A., and Vyas,
B.R.M. (2004) White-rot basidiomycetes: degradation of lignin (Review
Article), Communicated to Mycologia
5.
Shrivastava, R., Christian, V., Shukla, D., Modi, H.A., and Vyas,
B.R.M. (2004) Horseradish peroxidase: decolorization of dyes and
textile effluents. Communicated to Enz. Microb. Technol.
PRESENTATIONS
1.
Mehta, F., Joshi, A.P., Shrivastava, R., Christian, V.V., and Vyas,
B.R.M. (2004) Decolorization of textile dyes by the wood-rotting
basidiomycetes. Presented in XVIII Science Congress, March 13,
Saurashtra University, R ajkot, India.
102
2.
Joshi, A.P., Mehta, F., Shrivastava, R., Christian, V.V., and Vyas,
B.R.M. (2004) Decolorization of textile dyes by the wood-rotting
basidiomycetes. Presented in XVIII Science Congress, March 13,
Saurashtra University, Rajkot, India.
3.
Shrivastava, R., Christian V.V., Ladhawala K.P. and Vyas B.R.M.
Influence of Mn II and NaN3 on dye decolorization capabilities of
peroxidases from Pleurotus ostreatus . National Symposium on
Prospecting of Fungal Diversity and Emerging Technologies & 29th
Annual meeting of Mycological Society of India organized by Agharkar
Research Institiute, Pune. Maharashtra. 6-7 February 2003.
4.
Christian V.V., Shrivastava, R., Ladhawala K.P. and Vyas B.R.M.
Decolorization of Sulphonphthalein dyes by manganese peroxidase
from lignin degrading basidiomycetes Phanerochaete chrysosporium.
National Symposium on Prospecting of Fungal Diversity and Emerging
Technologies & 29 th Annual meeting of Mycological Society of India
organized by Agharkar Research Institiute, Pune. Maharashtra. 6-7
February 2003.
5.
Dave M. N., Shrivastava, R., Christian V.V., and Vyas B.R.M.
Decolorization of textile dyes by the ligninolytic enzymes of white rot
basidiomycetes. National Symposium on Prospecting of Fungal
Diversity and Emerging Technologies & 29th Annual meeting of
Mycological Society
of India organized by Agharkar Research
Institiute, Pune. Maharashtra. 6-7 February 2003.
6.
Parekh J. T., Shrivastava, R., Christian V.V., and Vyas B.R.M.
Ligninolytic enzymes of white rot basidiomycetes: Decolorization of
textile dyes. National Symposium on Prospecting of Fungal Diversity
and Emerging Technologies & 29th Annual meeting of Mycological
Society
of India organized by Agharkar Research Institiute, Pune.
Maharashtra. 6 -7 February 2003.
103
7.
Kesharia M. H., Christian V. V., Shrivastava, R., and Vyas B.R.M.
Dye
decolorization
capabilities
of
peroxidases
from
Litter
Basidiomycetes. National Symposium on Prospecting of Fungal
Diversity and Emerging Technologies & 29th Annual meeting of
Mycological Society
of India organized by Agharkar Research
Institiute, Pune. Maharashtra. 6-7 February 2003.
8.
Shrivastava, R., Christian V.V., and Vyas B.R.M. Production profile of
peroxidases
and
dye
decolorization
capabilities
of
white
rot
basidiomycetes. National Conference on Environmental Biology,
Department of Biosciences, Saurashtra University, Rajkot, Gujarat.
October 17-18, 2002.
9.
Christian V.V., Shrivastava, R., Avlani V.A., and Vyas B.R.M.
Screening of ligninolytic enzymes of selected white rot fungi in the
decolorizatio n of dye. National Conference on Fungal Diversity and
Biotechnology organized by Mycological Society of India, Thane,
Maharashtra. February 2-4, 2002
104
A PPENDIX III
CONFER ENCES / SE MINARS / WORKSHOPS
ATTENDED
1. Workshop on Recombinant DNA Technology organized by Central
Facility for Biotechnology Teaching and Research (CFBTR), Madurai,
Tamilnadu. September 27- October 5, 2003.
2. National Symposium on Prospecting of Fungal Diversity and
Emerging Technologies & 29
th
Annual meeting of Mycological
Society of India organized by Agharkar Research Institiute, Pune,
Maharashtra. 6-7 February 2003.
3. National Science Symposium on Environmental Science and
Technology organized by Commission for Scientific and Technical
Terminology and Community Science Center, Rajkot, Gujarat. 6-7
January 2003. (In Hindi)
4. National Conference on Environmental Biology
organized by
Department of Biosciences, Saurashtra University, Rajkot, Gujarat. 1718 October 2002.
5. State level seminar on Frontiers and Advancements in Biological
Science in New Millennium organized by Shree M. & N. Virani
Science College, Rajkot, Gujarat. 11August 2002.
105
6. National Conference on Fungal Diversity and Biotechnology
organized by Mycological Society of India, Thane , Maharashtra. 2 – 4
February 2002
7. Workshop on Principles and Methods in Regulatory Toxicology
organized by Jai Research Foundation, Vapi, Gujarat. 28 – 31 May
2001.
8. National Workshop on Bioinorganic Chemistry organized by M.S.
University, Baroda, Gujarat.
106