Download - Roshan Project Final
AN INVESTIGATION PROBLEM
ON
“Biodegradation of phenol by native bacteria isolated from
various soil samples and industrial wastes”
For partial fulfillment of the requirement for the degree of
MASTER OF SCIENCE
IN
BIOTECHNOLOGY
Session :( 2009-2011)
SUBMITTED TO: SUBMITTED BY:
Mr. Anil Kumar Roshan Shrestha
Assistant Professor Reg. N0. 09092042
DEPARTMENT OF BIO & NANO TECHNOLOGY
GURU JAMBHESHWAR UNIVERSITY OF
SCIENCE AND TECHNOLOGY
Mr. Anil Kumar, Assistant Professor
No.........…….. Dated…………………
CERTIFICATE I
This is to certify that the investigation problem report entitled, “Biodegradation of
phenol by native bacteria isolated from various oil samples and industrial wastes”
submitted by Roshan Shrestha (Reg.No.-9092042) for the partial fulfilment of the
requirement of the degree of Master of Science in Biotechnology to Department of Bio &
Nano Technology, GJUS&T, Hisar, Haryana-125001 has been carried out under my
supervision and guidance. The assistance and help received during the work has been
fully acknowledged.
Advisor
DEPARTMENT OF BIO AND NANO TECHNOLOGY,
GURU JAMBHESHWAR UNIVERSITY OF SCIENCE AND TECHNOLOGY,
Mr. Anil Kumar, Assistant Professor
No.........…….. Dated…………………
CERTIFICATE II
This is to certify that the investigation problem (MBD 600) entitled “Biodegradation of
phenol by native bacteria isolated from various soil samples and industrial wastes”
submitted by Mr. Roshan Shrestha, Reg. No. 09092042 to the Department of Bio &
Nano Technology, Guru Jambheshwar University of Science & Technology, Hisar for the
partial fulfillment of the requirement of the degree of Master of Science in Biotechnology
has been approved after an oral examination of the same.
External examiner Chairperson
Department of Bio & Nano Technology,
GJU S&T, Hisar.
DEPARTMENT OF BIO AND NANO TECHNOLOGY,
GURU JAMBHESHWAR UNIVERSITY OF SCIENCE AND TECHNOLOGY,
DECLARATION
Certified that the project report entitled “Biodegradation of phenol by native bacteria
isolated from various soil samples and industrial wastes”, submitted by me in partial
fulfilment for the award of degree of M.Sc. in Biotechnology, Guru Jambheshwar
University of Science & Technology, Hisar-125001, Haryana, is a record of the original
work carried out by me. The matter embodied in this report has not been submitted for the
award of any other degree or diploma.
Roshan Shrestha
ACKNOWLEDGEMENT
At the outset, I would like to convey my deepest sense of gratitude and indebtedness to
almighty God and my parents. The happiness of success lies in the sharing it with all who
have helped and inspired me to attain the goal. I would like to express my gratitude to all
those people whose constant guidance and encouragement served as a beacon of light.
I feel privileged to express my heartiest thanks and gratitude to my esteemed advisors Mr.
Anil Kumar, Assistant Professor, Department of Bio and Nano Technology for their
precise and inspiring guidance, valuable and generous suggestions throughout my
project.
It gives me immense pleasure in bringing this acknowledgement to pay my sincere
regards to Prof. Ashok Chaudhury, Chairperson, Department of Bio & Nano Technology,
GJUS&T, Hisar for his valuable support.
I pay my true regards to all the faculty members of the department especially Dr. Vinod
Chhokar, Dr. Namita Singh and Dr. Neeraj Dilbaghi for their kind co-operation.
I recognize with gratitude the invaluable co-operation and immense support provided by
Mr. Vikas Beniwal, Mr. Mahesh Kumar, Mr. Narender Malik, Mr. Rakesh Yadav, Mrs.
Sarika Malik, Ms. Pooja Arora, Mrs. Manju Yadav,Mr. Himanshu Aggarwal, Mr. Rajesh
Yogi and Mr. Ravinder Mehra.
I humbly offer my thanks to the whole staff of the Department of Bio & Nano Technology,
for their help and support during the period.
I truly acknowledge the ready help and co-operation extended to me by all my seniors,
classmates and my friends specially.
Again I iterate a word of thanks to all those hands that helped me in some way or the
other in pursuing this elusive work.
June,2011 Roshan Shrestha
CONTENTS
List of figures……………………………………………………………………….
List of Tables………………………………………………………………………..
CHAPTERS
Chapter 1 INTRODUCTION
Chapter 2 REWIEW OF LITERATURE
Chapter 3 MATERIALS AND METHODS
3.1 Isolating of phenol degrading bacteria
3.2 Characterization of isolated strain by morphological and biochemical tests.
3.3 Immobilization of microbial cells
Chapter 4 RESULTS & DISCUSSIONS
4.1Morhology and biochemical test of isolated strains.
4.2 Residual phenol estimation
4.3 Comparative study of phenol degradation by immobilized and free cells.
Chapter 5 CONCLUSIONS
RERERENCES
List of Tables
Table 1 Microscopic, morphological and
biochemical test of various isolates
Table 2 Standard curve
Table 3 Residual phenol Estimation
Table 4 Calculation of residual phenol
concenteration at end of 6, 12, 18 &
24 hrs incubation by S1G
Table 5 Calculation of residual phenol
concenteration at the end of 6, 12, 18
& 24 hrs incubation by S4A
Table 6 Calculation of residual phenol
concenteration at the end of 6, 12, 18
& 24 hrs incubation by S5C
Table 7 Comparative study of phenol
degradation by immobilized and free
cells.
LISTS OF FIGURES
Fig. No. Content
Fig. 1 Standard Curve
Fig. 2 Residual phenol estimation
Fig. 3 Residual phenol concentration
Fig 4 Phenol degradation by encapsulated cells
and free cells
INTRODUCTION
1. INTRODUCTION
Environmental pollution is an emerging threat and great concern in today’s context
pertaining to its effect on the ecosystem. The worldwide rise in population and the
industrialization during the last few decades have resulted in ecological unbalance and
degradation of the natural resources. One of the most essential natural resources which
have been the worst victim of population explosion and growing industrialization is
water. In recent years, considerable attention has been paid to industrial wastes
discharged to land and surface water. Industrial effluents often contain various toxic
metals, harmful dissolved gases, and several organic and inorganic compounds. Organic
pollutants comprise a potential group of chemicals which can be dreadfully hazardous to
human health. Many of these are resistant to degradation. As they persist in the
environment, they are capable of long range transportation, bioaccumulation in human
and animal tissue and biomagnification in food chain. Huge quantity of waste water
generated from human settlement and industrial sectors accompany the disposal system
either as municipal wastewater of industrial wastewater.
Over 5 million chemical substances produced by industries have been identified and
about 12000 of these are marketed which amount to around half of the total production.
Contaminated water by pesticides, such as DDT, aldrin, dieldrin, heptachlor etc is harm
full for aquatic life and human beings as well. Discharge of cyanide-contained wastewater
to water mass may lead to death of fish and other aquatic life therein. Use of water
containing fluoride can causes mental disorders and stomach ailments and can also
reduces agricultural production.
Phenol along with other xenobiotic compounds is one of the most common contaminants
present in effluents from process industries.
Phenols are compounds with ArOH formula which are extremely toxic and found in
different form or together with other elements (EPA, 2004). Phenol is one of the 50
major bulk chemicals produced in the world and its annual production reached 6.6 billion
pounds in 2004 and expected to grow by 6% per year (CMR 2005). Phenol is considered
to be a toxic compound by the Agency for Toxic Substances and Disease Registry
(Agency for Toxic Substance and Disease Registry 2003) and death among adults has
been reported with ingestion of phenol ranging from 1 to 32 g. Phenol and its derivatives
1
are also generated by various industries, such as petroleum refining, petrochemical, coke
conversion, pharmaceutical, plastic, and resin manufacturing, coal gasification, coke-
oven batteries, and other industries, such as synthetic chemicals, herbicides, pesticides,
antioxidants, pulp-and-paper, photo developing chemicals, etc. (Marrot et al., 2006;
Bodalo et al., 2008; Jayachandran and Kunhi, 2008) in the waste effluents and its
concentration may vary from 1 to 15000 mg l-1. Natural sources of phenol include forest
fire, natural run off from urban area where asphalt is used as the binding material and
natural decay of lignocellulosic material. United States Environmental Protection Agency
(USEPA) and Central Pollution Control Board of India (CPCB) have prescribed
maximum permissible limits of 3.4 and 5.0 mg l-1, respectively in industrial waste
discharges. Now the associated problem due to phenol is that when it is present in waste
water even in low concentrations can be toxic to some aquatic species and causes taste
and odour problems in drinking water (Rittmann and McCarty, 2001). Inhalation and
dermal contact of phenol causes cardiovascular diseases and severe skin damage, while
ingestion can cause serious gastrointestinal damage and oral administration into
laboratory animals has also induced muscle tremors and death.
Phenol in solution form, easily passes through the skin, and its metabolism occurs in the
liver, although, it could occur in the lung and kidney too. Phenol is toxic in environment
and could decrease enzymatic activity as well. Also, it is toxic to fishes and is mortal
between 5 – 25 mg/l for them. Moreover, direct effect of phenol is a blocker for biologic
reaction. Phenolic compounds are serious pollutant for rivers (EPA, 2004) and they have
harmful effects such as growth inhibition, decrease of resistance against diseases, aquatic
mortality and increase in growth of weedy plants.
Acute exposure of phenol causes central nervous system disorders. It leads to collapse
and coma. Muscular convulsions are also noted. A reduction in body temperature is
resulted and this is known as hypothermia. Mucus membrane is highly sensitive to the
action of phenol. Acute exposure of phenol can result in myocardial depression.
Whitening and erosion of the skin may also result due to phenol exposure. Phenol has an
anaesthetic effect and causes gangrene. Renal damage and salivation may be induced by
continuous exposure to phenol. Exposure to phenol may result in irritation of the eye,
conjunctional swelling, corneal whitening and finally blindness. Other effects include
frothing from nose and mouth followed by headache. Phenol can cause hepatic damage
also. Chronic exposure may result in anorexia, dermal rash, dysphasia, gastrointestinal
2
disturbance, vomiting, weakness, weightlessness, muscle pain, hepatic tenderness and
nervous disorder. It is also suspected that exposure to phenol may cause paralysis, cancer
and genetofibre striation. Phenol and its derivatives are toxic and classified as hazardous
materials (Zumriye and Gultac, 1999). These phenolic compounds possess various
degrees of toxicity and their fate in the environment is therefore important (Bollag et al.,
1988).
In recent years, a great deal of research work has been directed toward the development
processes in which enzymes are used to remove phenolic contaminants (Ghioureliotis and
Nicell, 1999). Phenol is an antiseptic agent and is used in surgery, which indicates that
they are also toxic to many microorganisms (EPA, 1979). If phenolic pollution goes to
underground water, it causes serious ecological problems. Hence, allowable amount of
phenol in industrial outgoing must not be more than 0.5 mg/l. Considering the above
issue, the removal of such chemicals from industrial effluents is of great importance. The
conventional methods of treatment of phenolic and nitrate-nitrogen Waste water are
largely physical and chemical processes like hybrid process (Bodalo et al., 2008)
electrocatalytic degradation (Wang et al., 2009) adsorption on to different matrices,
chemical oxidation, solvent extraction or irradiation (Spiker et al., 1992) but these
processes led to secondary effluent problems due to formation of toxic materials such as
cyanates, chlorinated phenols, hydrocarbons, etc. These methods are mainly chlorination,
ozonation, solvent extraction, incineration, chemical oxidation, membrane process,
coagulation, flocculation, adsorption, ion exchange, reverse osmosis, electrolysis, etc.
Biological treatment is attractive due to the potential to almost degrade phenol and other
pollutant while producing innocuous end products, reduced capital and operating costs,
maintains phenol concentrations below the toxic limit. However difficulty arises in such
treatment due to the toxicity of phenol to the microbial population. The biological
degradation of phenol is accomplished through benzene ring cleavage using the enzyme
present in the microorganism. The bacteria express differently when exposed to different
initial phenol concentrations and other conditions. The most efficient Pseudomonas
Putida is capable of using phenol as the sole source of carbon and energy for cell growth
and metabolism degrade phenol via metapathway. That is the benzene ring of phenol is
dehydroxylated to form catechol derivative and the ring is then opened through meta-
oxidation. The final products are molecules that can enter the tricarboxylic acid cycle.
3
OBJECTIVES OF WORK DONE:
To isolate the potent phenol degrading Bacteria.
To characterize various bacterial isolates by morphological and biochemical test
To study phenol degradation by cell immobilization method.
To study the growth curve.
4
REVIEW OF LITERTURE
5
2. REVIEW OF LITERATURE
Today Bioremediation is considered as a new tool to eliminate environment pollutions
(EPA, 2004). Bioremediation is defined as the process whereby organic wastes are
biologically degraded under controlled conditions to an innocuous state, or to levels
below concentration limits established by regulatory authorities (Atlas and Bartha, 1998).
For bioremediation to be effective, microorganisms must enzymatically attack the
pollutants and convert them to harmless products. As bioremediation can be effective
only where environmental conditions permit microbial growth and activity, its application
often involves the manipulation of environmental parameters to allow microbial growth
and degradation to proceed at a faster rate. Bioremediation techniques are typically more
economical than traditional methods such as incineration, and some pollutants can be
treated on site, thus reducing exposure risks for clean-up personnel, or potentially wider
exposure as a result of transportation accidents. Most bioremediation systems are run
under aerobic conditions, but running a system under anaerobic conditions may permit
microbial organisms to degrade otherwise recalcitrant molecules also. Microbial growth
and activity are readily affected by pH, temperature, and moisture. Although
microorganisms have been also isolated in extreme conditions, most of them grow
optimally over a narrow range, so that it is important to achieve optimal conditions. If the
soil has too much acid it is possible to rinse the pH by adding lime. Temperature affects
biochemical reactions rates, and the rates of many of them double for each 10 °C rise in
temperature. Above a certain temperature, however, the cells die. Available water is
essential for all the living organisms, and irrigation is needed to achieve the optimal
moisture level.
Hydrocarbons are readily degraded under aerobic conditions, whereas chlorinated
compounds are degraded only in anaerobic ones. To increase the oxygen amount in the
soil it is possible to till or sparge air. In some cases, hydrogen peroxide or magnesium
peroxide can be introduced in the environment. Low soil permeability can impede
movement of water, nutrients, and oxygen; hence, soils with low permeability may not be
appropriate for in situ clean-up techniques.
6
MICROORGANISMS IN PHENOL BIODEGRADATION
Degradation of phenol occurs as a result of the activity of a large number of
microorganisms including bacteria, fungi and actinomycetes. Bacterial species include
Bacillus sp, Pseudomonas sp, Acinetobacter sp, Achromobacter sp etc. Fusarium sp,
Phanerocheate chrysosporium, Corious versicolor, Ralstonia sp, Streptomyces sp etc are
also proved to be efficient fungal groups in phenol biodegradation. (Prieto et al., 2002).
Many studies on biodegradation of phenol come from bacteria. The genus Pseudomonas
is widely applied for the degradation of phenolic compounds. These bacteria are known
for their immense ability to grow on various organic compounds. Phenol biodegradation
studies with the bacterial species have resulted in bringing out the possible mechanism
and also the enzyme involved in the process. The efficiency of the phenol degradation
could be further enhanced by the process of cell immobilization (Annadurai et al., 2000a,
b). Phenol and other phenolic compounds are common constituents of many industrial
effluents. Once a suitable micro organism based process is developed for the effective
degradation of phenol these phenolic effluents can be safely treated and disposed
(Borghei and Hosseini, 2004). Candida tropicalis RETLCrl from the effluent of the
Exxon Mobile Oil Refinery waste water treatment was investigated for phenol
degradation using batch and fed batch fermentation under aerobic condition (Mohd Tuah,
2006). Microbiological degradation of phenol and some of its alkyl derivatives (p-cresol,
4-n-propyl phenol, 4-i –propyl phenol, 4-n-butyl phenol, 4-sec-butyl phenol, 4-t-butyl
phenol and 4-t-octyl phenol) were examined under both
aerobic and anaerobic conditions in seven Japanese paddy soil samples (Atsushi et al.,
2006). The rate of biodegradation of phenol by Klebsiella oxytoca strain was studied. It
was found that K. oxytoca degraded phenol at elevated concentration where 75% of initial
phenol concentration at 100 ppm was degraded within 72 h (Shawabkeh et al., 2007).
Phenol was degraded by Actinobacillus species (Khleifat and Khaled, 2007). They found
that pH 7, the incubation temperature of 35 to 37°C, and the agitation rate of 150 rpm
were the optimal conditions for achieving the higher percentage of phenol degradation.
Succinic acid and glycine as respective carbon and nitrogen source were found to be the
most efficient co-substrates for the removal of phenol. Immobilized Alcaligenes sp d2
was successfully used for the effective treatment of phenolic paper factory effluent
(Nair and Shashidhar, 2007).
7
Many studies on biodegradation of phenol using pure and mixed cultures have been
reported (Collins et al., 2005; Dursun and Tepe, 2005; Marrot, 2006; Shen et al., 2009;
Laowansiri et al., 2008; Celik et al., 2008; Santos et al., 2009). In India also a
considerable amount of study on phenol biodegradation has been undertaken like the
studies of Dhagat et al. (2002); Chandra and Rathore (2002); Ambujom and Manilal
(2004); Shetty et al. (2007); Jayachandran and Kunhi (2008). Biological processes are
generally preferred due to their lower operational costs and the possibility of complete
mineralization (Agarry et al. 2008; Brar et al. 2006). A number of microorganisms have
been shown to utilize phenol and its derivatives as sole sources of carbon and energy at
varying concentrations, under aerobic conditions. They include several bacterial cultures
(Babu et al. 1995b; Chung et al. 2003; Karigar et al. 2006), a few yeast strains (Agarry et
al. 2008; and mycelial fungi (; Santos et al. 2003; Yordanova et al. 2009). Removal of
phenol also has been carried out using activated sludge systems for the last several years
(Amor et al. 2005; Marrot et al. 2008). But, high phenol loading rates and fluctuations in
phenol loads have been reported to cause the breakdown of these systems (Watanabe et
al. 2000).
The major difficulty encountered in the microbial treatment of phenol-containing
wastewater is the substrate inhibition of microbial growth and concomitant hindrance in
the biodegradation. To overcome these problems immobilization of the degrading
organism(s) has been proposed as an effective strategy (Keweloh et al. 1989). The main
advantages of using immobilized cells, rather than free suspended ones, include the
retention in the reactor of higher concentrations of microorganisms, protection of cells
against toxicity and prevention of suspended microbial biomass in the effluent. Moreover,
immobilization of microbial cells provides, in general, high degradation efficiency and
good operational stability. Aerobic granulation of the degrading organism(s) as a new
form of cell immobilization for exploitation in phenol wastewater treatment in sequencing
batch reactors also has been gaining importance, recently (Tay et al. 2004). The
aggregation of microbial cells into compact self-immobilized granules can serve as an
effective protection against phenol toxicity. However, the traditional immobilization
technique of entrapment of microorganisms in gel polymers has been shown to be more
efficient in resisting the phenol toxicity and improving the degradation rates. Several
different polymeric materials have been tried as matrices for immobilization. Ca-alginate
has been used as the gel matrix in several studies for encapsulating strains of bacteria,
8
yeasts, and mycelial fungi (Abd-El-Haleem et al. 2003; Santos et al. 2003). Other
polymers which were occasionally used for immobilizing phenol-degrading
microorganisms were agar, polyacrylamide (PAA), polyacrylamide hydrazide (PAAH),
polyvinyl alcohol (PVA), polysulfone hollow fiber membrane, and modified
polyacrylonitrile membrane (Chen et al. 2002; Yordanova et al. 2009). All the phenol-
degrading strains exhibited improved ability, to varying degrees, in the immobilized
systems than as freely suspended cells. However, studies on comparison of the suitability
and efficiency of different matrices for immobilization of phenol degrading organisms are
very rarely found in literature (Bettmann and Rehm 1984). Particularly, a comparative
appraisal of the calcium alginate and agar immobilized microorganisms has been very
scarcely found (Karigar et al. 2006). Pseudomonas sp. strain CP4, a laboratory isolate
was found to be a potent degrader of phenol, isomers of cresol, and other aromatics under
shake flask conditions (Ahamad and Kunhi 1999; Ahamad et al. 2001; Babu et al. 1995b)
and was an efficient partner of a mixed culture with Pseudomonas aeruginosa strain 3mT
in the degradation of mixtures of 3-chlorobenzoate (3-CBA) and phenol/cresols (Babu et
al. 1995a).
MECHANISM OF PHENOL BIODEGRADATION
Generally aromatic compounds are broken down by natural bacteria. However, polycyclic
aromatic compounds are more recalcitrant. Derivatisation of aromatic nuclei with various
substituents particularly with halogens makes them more recalcitrant. There are reports on
many microorganisms capable of degrading phenol through the action of variety of
enzymes. These enzymes may include oxygenases hydroxylases, peroxidases, tyrosinases
and oxidases.
The critical step in the metabolism of aromatic compounds is the destruction of the
resonance structure by hydroxylation and fission of the benzoid ring which is achieved by
dioxygenase-catalysed reactions in the aerobic systems. Based on the substrate that is
attacked by the ring cleaving enzyme dioxygenase, the aromatic metabolism can be
grouped as catechol pathway, gentisate pathway, and proto catechaute pathway. In all
these pathways, the ring activation by the introduction of hydroxyl groups is followed by
the enzymatic ring cleavage. The ring fission products, then undergoes transformations
leading to the general metabolic pathways of the organisms. Most of the aromatic
9
catabolic pathways converge at catechol. Catechols are formed as intermediates from a
vast range of substituted and nonsubstituted mono and poly aromatic compounds.
Aerobically, phenol also is first converted to catechol, and subsequently, the catechol is
degraded via ortho or meta fission to intermediates of central metabolism. The initial ring
fission is catalysed by an ortho cleaving enzyme, catechol 1, 2 dioxygenase or by a meta
cleaving enzyme catechol 2,3 dioxygenase, where the product of ring fission is a cis-
muconic acid for the former and 2-hydro Cis- muconic semi aldehyde for the latter
(Gurujeyalakshmi and Oriel, 1988). Streptomyces setonii (ATCC 39116) degraded
aromatic compounds such as phenol or benzoate via an ortho cleavage pathway using
catechol 1,2 dioxygenase (An et al., 2001). These dioxygenases are highly labile
enzymes and there requires a detailed investigation into its structural properties. A
bacterial strain, Serratia plymuthica was able to tolerate phenol up to a concentration of
1050 mg/L. Phenol was degraded through ortho pathway and the crude extract showed
the presence of ring cleaving enzyme catechol 1, 2-dioxygenase (Nilotpala and Ingle,
2007). Catechols are cleaved either by ortho-fission (intradiol, that is, carbon bond
between two hydroxyl groups or by a meta-fission (extra diol, that is, between one of the
hydroxyl groups and a non-hydroxylated carbon) as given in Figures 2 and 3. Thus the
ring is opened and the open ring is degraded (Cerniglia, 1984). As a general rule, most of
the halo aromatics are degraded through the formation of the respective halocatechols, the
ring fission of which takes place via ortho-mode. On the other hand, most of the non
halogenated aromatic compounds are degraded through meta pathway.
10
Figure 3. Meta pathway of phenol degradation. Figure 2. Meta pathway of phenol degradation.
The fission product of ortho-cleavage would be cis, cis muconic acid or its derivative
depending on whether the catechol is substituted or not. The meta-fission product of
catechol would be 2-hydroxy muconic semialdehyde and the products of both ortho and
meta pathways are further metabolized as intermediates of TCA cycle. Ortho pathway is
the most productive pathway for the organism as it involves less expenditure of energy.
Phenol hydroxylase (E. C 14. 1.3.7) catalyses the degradation of phenol via two different
pathways initiated either by ortho or meta cleavage. There are many reports on phenol
hydroxylase and catechol 2, 3 dioxygenase involved in the biodegradation of phenol
(Leonard and Lindley, 1999). Hublik and Schinner (2000) reported the characterization of
11
laccase from Penurious ostreatus. The enzyme was purified to homogeneity and was
characterized. It was a monomeric protein with a molecular weight of 67 KD and with an
isoelectric point of 3.6. They observed that the laccase retained most of its activity in high
ionic buffer, pH.10, 20°C temperature in the presence of 10 mM benzoic acid and with
35% ethylene glycol. The degradation of phenolic compounds by immobilised laccase
from Streptomyces psammoticus was evaluated and confirmed by thin layer
chromatography and nuclear magnetic resonance spectroscopy (Niladevi and Prema,
2007). Polyphenol oxidase is a (EC 1.14.18.1) monoxygenase which catalyses the O-
hydroxylation of phenols and the oxidation of O-dihydric phenols to O-quinones using
molecular oxygen. Laccase are phenol oxidases which utilize molecular oxygen. They are
known to have the ability to oxidize polyphenols; meta substituted phenols, diamines and
a variety of other components (Kadhim, 1999). The mechanism by which polyphenol
oxidase catalyses the conversion of monophenols to O-quinones involves the
hydroxylation of monophenols followed by dehydrogenation to form O-quinones. These
quinones undergo spontaneous nonenzymatic polymerization in water, eventually
forming water insoluble polymers which can be separated from water by filtration
(Edwards et al., 1999). There were various reports on the exploitation of polyphenol
oxidase in the detoxification of the phenols. The interest in polyphenol oxidase had been
fueled by their potential uses in detoxification of environmental pollutants (Bollag et al.,
1988). Production of useful chemicals from lignin (Burton et al., 1993) by polyphenol
oxidase was also reported. Garzillo et al. (1998) reported a polyphenol oxidase from the
white rot fungus Trametes trogii. It was an enzyme with molecular weight 70 KD. The
purified enzyme oxidised a number of phenolic compounds. This multicopper oxidases
had a wide range of substrate specificity. Coprinus macrorhizus and Arthromyces
ramosus were proved to be effective in removing phenol and phenolic compounds from
water (Wu et al., 1998). Of the various enzymes acting on phenol, polyphenol oxidase
was the most important one probably because of its increasing demand in lignin
degradation (Garzillo et al., 1998). The non specific nature of the polyphenol oxidase was
also discussed by Schneider et al. (1999). Immobilised polyphenol oxidase on chitosan
coated polysulphone capillary membranes were used for improved phenolic effluent
bioremediation (Edwards et al., 1999).
They also highlighted the removal of quinones and other polymerized products using
chitosan. Polyphenol oxidases were widely distributed in many plants and fungal species
(Robles et al., 2000). They suggested the possibility of using a polyphenol oxidase
12
producing strain of the hyphomycete Chalara paradoxa in the detoxification of olive mill
wastewater. Sakurai et al. (2001) showed that the peroxidase from Coprinus cinereus
could be used for the removal of Bisphenol. Polymerization of the bisphenol by the
enzyme was utilized here. Manophenols in aqueous solution could also be removed by
peroxidase catalysed oxidation (Xia et al., 2003). Certain actinomyces and Streptomyces
strains could produce tyrosinase enzyme, which oxidized halogen substituted phenols.
Peroxidases could catalyse the transformation of phenol and halogenated phenols.
Peroxidases such as those from Arthrobacter and Streptomyces strains were being
reported as the phenol degrading enzymes (Fetzner and Lingens, 1994). The peroxidase
catalysed polymerization process was proved to be very effective in eliminating phenol
and a variety of other aromatic pollutants from waste waters (Ghioureliotis and Nicell,
1999). Peroxidases can act on phenol and other aromatic compounds through oxidative
coupling. In presence of hydrogen peroxide two equivalents of phenol are converted by
each equivalent of enzyme into highly reactive radical species. Once they are formed,
they react with one another to yield phenolic polymers. Tyrosinase catalyzes the
oxidation of phenols involving the formation of orthoquinones. The mechanism of the
enzymatic action of tyrosinase on various phenols was discussed in detail by Siegbahn
(2003). The mechanism of degradation of an organic compound may be unusual (Jenisch-
Anton, 1999). The mechanism of degradation is generally decided by the nature of the
organic compound, its solubility, and nature of the organism, type of the enzyme and also
by the external factors affecting biodegradation. In some cases, through
the action of monooxygenase, aromatic com-pounds may be converted into gentisic acid.
The fission of this compound occurs between the hydroxyl and carboxyl groups, that is,
meta fission. It has been shown in some cases that chloroaromatic compounds such as 4-
chlorobenzoate, 4-chlorophenol and others may get dechlorinated during the
hydroxylation resulting in the formation of 4–hydroxy benzoates (4-HBA). This 4 HBA
on further hydroxylation will be converted to protocatechuate
acid (3,4-dihydroxy benzoic acid), which may be cleaved either through ortho or meta
mode.
Each of these factors should be optimized for the selected organism for the maximum
degradation of the organic compound of choice. The optimization of the substrate
concentration in phenol biodegradation is particularly important since it inhibits the
growth of the organism at higher concentrations. Biotechnology for hazardous waste
management involves the development of biological systems that catalyse the
detoxification, degradation or decontamination of environmental pollutants.
13
MATERIALS AND METHODS
14
3. MATERIALS AND METHODOLOGY
3.1 GLASSWARES:
Autoclaved beakers, Flasks, Petri plates, Glass rods, Pestle and mortar, Centrifuge tubes
and Eppendorf tubes
3.2 INSTRUMENTS AND EQUIPMENT REQUIRED:
BOD Incubator SONAR
Autoclave NSW, New Delhi
Laminar Air Flow Atlantis, New Delhi
Weighing Machine A&D Company Ltd
Centrifuge REMI, Mumbai
Vortex LABNET
Water Bath POPULAR TRADERS, Ambala Cantt
Microwave oven L.G.
Refrigerator L.G.
µpH system- 361 Systronics
15
3.3 METHODS
3.3.1 METHODS OF ISOLATING PHENOL DEGRADING BACTERIA
Soil samples were collected from various places of India, the most potent phenol degrader
was isolated by following procedure.
1 gm of soil sample was added to 9ml of autoclaved distilled water and the resulting
mixture was centrifuged. Then 5 ml of this media was inoculated to 250 ml of new BHM
broth supplemented with phenol at the rate of 600mg/L at 37°C for another one week.
Again, 10 ml of the second passage was inoculated into the new BHM broth media and
incubated in the above mentioned situation. These passages were repeated until turbidity
was obtained from bacteria growth, which was not due to mixed sedimentation with the
first media. After the last passage, it was cultured on phenol agar media as an isolate and
the bacterium was isolated as a colony alone. Regarding water samples, same method was
applied, except that 10 ml water was mixed with 100 ml phenol broth media and all
previously mentioned stages were repeated (Koutny et al., 2003).
The compositions of Bushnell’s Hass Medium in g l-1 were
K2HPO4 - 0.1
NaCl - 0.1
CaCl2.2H2O - (0.015)
MgSO4 - 0.2
FeCl3 - 0.02
NH4Cl - 2.00
Phenol - 0.66
3.4 EXPERIMENTAL PROCEDURE:
The isolated bacteria were suspended in Bushnell’s Hass broth medium supplemented
with 600mg/l phenol at pH 7. The suspension cultures of the isolated bacteria were
inoculated in BHM containing 600 mg l-1 initial concentration of phenol to compare their
phenol degrading efficiency. The line of experiment was designed according to a previous
16
study of Wael et al. (2003), where he had isolated six pure bacterial strains from a coke
processing unit waste water. The major modification was the initial phenol concentration
of 600 mg l-1, which he had earlier taken as 100 mg l-1. The residual phenol concentration
was monitored at different time intervals spectrophotometrically according to the method
described by Yang and Humphrey (1975). The strain degrading phenol to a greater extent
within a relatively short time was selected as efficient phenol degrader among the isolates
for the optimization studies of the physical environment.
3.5 ISOLATION AND IDENTIFICATION OF THE BACTERIA:
Eight bacterial strains (S1G, S2B, S1H, D1, D5, S4A, S4C, and S5C) were found to be
growing in Bushnell’s Hash Medium containing 600 mg l-1 of phenol during the early
screening for the investigation.
S1G and S1H are isolated from the soil sample of Soya Industry, Indore.
S2B was isolated from Auto market Hisar.
S4A was isolated from Pesticide factory Hisar.
S5C was isolated from Alcohol industry Hisar,
D1 was isolated from soil sample of Palwal and
D5 was isolated from underground water of Hisar.
Microscopic observation and growth characteristics as well as biochemical tests of the
isolated bacteria were studied.
3.5.1 GRAM’S STAINING:
Gram’s stain developed by Christian Gram in 1884 is the most widely employed staining
method in bacteriology. It is an example of differential staining. This is a useful method
of identifying and classifying bacteria in two major groups:
-Gram Positive
- Gram Negative
The method relies on the cell structure differences between the two groups. The Gram
positive cell wall consist of 20-80nm thick homogenous layer of peptidoglycan (murein)
lying outside the plasma membrane. The peptidoglycan itself is not stained instead it
seems to act as a permeability barrier preventing loss of Crystal Violet. In contrast the
Gram negative cell wall is quite complex. It has a 2-7nm peptidoglycan layer covered by
a 7-8 nm thick outer membrane.
17
During the procedure the bacteria are first stained with crystal violet and next treated with
iodine to promote dye retention. When Gram positive are then treated with ethanol, the
alcohol is thought to shrink the pores of thick peptidoglycan. Thus the dye iodine
complex is retained during this short decolorization step and the bacteria remain purple.
In contrast the Gram negative peptidoglycan is very thin, not as highly cross linked and
has larger pores. Alcohol treatment may also extract enough lipids from the Gram
negative outer membrane to increase its porosity further. For these reasons, alcohol more
readily removes the purple Crystal Violet Iodine complex from Gram negative bacteria.
Thus Gram negative bacteria are then easily stained red or pink by the counterstain
saffranin.
3.5.1.1 PROCEDURE:
1. Initially a smear of bacterial culture was prepared on a slide, air dried and then
heat fixed in order to immobilize the bacteria.
2. Smear was then placed in a small jar containing Crystal Violet solution and was
allowed to stand for 1min.
3. After 1min slide was washed with distilled water and then dipped in iodine
solution for 1min.
4. After 1min decolorizer was applied drop wise until no more color flowed from
smear.
5. Slide was then washed with distilled water and then stained with saffranin and
kept for 1min.
6. Slide was then washed with distilled water and made dry.
7. Finally, the slide was examined under oil immersion objective microscope for
staining.
3.5.1.2 INTERPRETATION:
After staining if the bacterial strain was found to be purple it is Gram positive otherwise
Gram Negative
3.5.2 CITRATE TEST (SIMMON’S CITRATE TEST)
The citrate test utilizes Simmon's citrate media to determine if a bacterium can grow
utilizing citrate as its sole carbon and energy source. Simmon's media contains
bromthymol blue, a pH indicator with a range of 6.0 to 7.6. Bromthymol blue is yellow at
18
acidic pH's (around 6), and gradually changes to blue at more alkaline pH's (around 7.6).
Uninoculated Simmon's citrate agar has a pH of 6.9, so it is an intermediate green color.
Growth of bacteria in the media leads to development of a Prussian blue color (positive
citrate). Enterobacter and Klebsiella are citrate positive while E.coli is negative.
3.5.2.1 PROCEDURE:
1. A loopful of bacteria was streaked onto a citrate agar slant.
2. It was incubated 24 to 48 hours with a loose cap.
3.5.2.2 INTERPRETATION:
After incubation a positive reaction is indicated by a slant with a Prussian blue color. A
negative slant will have no growth of bacteria and will remain green.
3.5.3 UREASE TEST
Urease broth is a differential medium that tests the ability of an organism to produce an
exoenzyme, called urease that hydrolyzes urea to ammonia and carbon dioxide. The
broth contains two pH buffers, urea, a very small amount of nutrients for the bacteria, and
the pH indicator phenol red. Phenol red turns yellow in an acidic environment and
fuchsia in an alkaline environment. If the urea in the broth is degraded and ammonia is
produced, an alkaline environment is created, and the media turns pink.
Many enterics can hydrolyze urea; however, only a few can degrade urea rapidly. These
are known as “rapid urease-positive” organisms. Members of the genus Proteus are
included among these organisms.
Urea broth is formulated to test for rapid urease-positive organisms. The restrictive
amount of nutrients coupled with the use of pH buffers prevent all but rapid urease-
positive organisms from producing enough ammonia to turn the phenol red pink.
3.5.3.1 PROCEDURE:
1. Inoculate a urea tube with 3 loopful of slant culture.
2. Incubate 24 hours, observe for reaction.
3. A pink color formation indicates the breakdown of urea to ammonia and CO2
3.5.3.2 INTERPRETATION
19
Phenol red indicator is added to the broth so that when the pH reaches 8.4 (due to
ammonia production) the tube will turn from an orange color to a pink color, indicating
the choice of organism is Urease positive.
3.5.4 EOSINE METHYLENE BLUE (EMB) AGAR
DESCRIPTION:
A selective medium for gram-negative bacteria Levine’s EMB agar contains methylene
blue, which inhibits gram-positive bacteria. Differential for enterics: will differentiate
lactose fermenters from nonfermenters.
3.5.4.1 PROCEDURE:
One loopful of culture was inoculated in EMB agar slant.
3.5.4.2 INTERPRETATION:
After incubation: Lactose nonfermenters will have cream colored colonies. Lactose
fermenters will have pinkish colonies, sometimes with dark centres. E. coli often has a
greenish metallic sheen.
3.6 Immobilization of Microbial Cells
20
Cells of mixed culture collected from soils containing pollutants or specific culture (pure)
isolated from the pollutant containing soil are immobilized in/on solid matrix.
Acclimization of microorganisms is done by increasing the pollutant concentration (say
of phenol) gradually during culture preparation. The acclimized culture is used for the
immobilization in/on the solid matrix. Immobilization of cells means that the cells have
been confined or localized so that it can be reused continuously. These exhibit totally
different hydrodynamic characteristics than surrounding environment. Living cells
produce enzymes (biological catalysts) to catalyze cellular reactions vital to the organism.
The microorganisms are normally immobilized on natural and synthetic supports. Various
types of solid matrices like polyacrylamide gel, Ca alginate, porous glass, plastic beads,
activated carbon, sand, charcoal, diatomaceous earth, cement balls made of coal ash,
cellulose, polymeric materials, polymeric ions, chitosan, lignins, chitins, coal, collagens
etc. have been used for immobilization of whole cells. In the recent years, the
immobilization of biocatalysts with polyvalent salts of alginic acids has received much
attention because of low cost of alginate and the mild conditions of immobilization.
Techniques of immobilization are broadly classified into four categories namely: covalent
bonding, cross-linking (chemical methods), entrapment and adsorption (physical
methods). Covalent binding most extensively used technique, where cells or enzymes are
covalently linked to the support through the groups in them or through the functional
groups in the support material. In the cross-linking technique, the cells are immobilized
through chemical cross-linking using homo as well as hetero-bifunctional cross-linking
agents. Adsorption is the simplest of all techniques and does not alter the activity of the
bound cells. Adsorption involves adhesion or condensation of the cells to the surface of a
carrier. The diving force causing immobilization is the combined hydrophobic
interactions, hydrogen bonding and salt bridge formation between the adsorbent and cells.
Entrapment within gels or fiber is a convenient method for reactions involving low
molecular weight substrates and mainly used for immobilization of whole cells. This
method is nothing but the polymerization of the unsaturated monomers in the presence of
cells results in the entrapment of the cells within the interstitial spaces of the gel
3.6.1 PROCEDURE OF CELL IMMOBILIZATION:
21
a. Alginate was dissolved in boiling water and autoclaved at 121°C for 15 min.
b. Cells were harvested during the mid-logarithmic growth phase by centrifugation
(5000 g, 10 min)
c. The Cells mass was resuspended in 2 ml of saline and added to 100 ml of
sterilized alginate solution. This alginate/cell mixture (with stirring) was extruded
drop by drop into a cold, sterile 0.2 M CaCl2 solution through a sterile 5 ml
pipette.
d. Gel beads of approximately 2 mm diameter were obtained.
e. The beads were hardened by resuspending into a fresh CaCl2 solution for 24 h at
4°C with gentle agitation.
f. Finally these beads were washed with distilled water to remove excess calcium
ions and unentrapped cells.
g. Then the beads were transferred to 50 ml production medium and cultivated for
the required time.
22
RESULTS AND DISCUSSION
4. RESULT & DISCUSSION
23
4.1 The morphology and biochemical test of the isolated bacteria were studied and
following results were obtained.
S1G was found to be Gram +ve with cocci shaped which could utilize lactose and urea
but was found –ve to citrate.
S2B was found to be gram +ve and oval shaped which could utilize lactose but was found
–ve to both urea and citrate.
S1H was found to be Gram –ve and rod shaped which could utilize lactose but was –ve to
both urea and citrate.
D1 was found to Gram –ve and rod shaped which can neither utilize lactose, citrate nor
urea.
D5 was found to Gram –ve cocci bacilli which could utilize lactose and citrate but was
found –ve to urea.
S4A was found to Gram –ve and rod shaped which can utilize lactose but was found –ve
to both urea and citrate.
S4C was found to Gram +ve and cocci shaped which is positive to citrate but was found –
ve to lactose and urea.
S5 was found to Gram +ve and rod shaped which could utilize lactose but was found –ve
to both urea and citrate.
The details of the above description are summarized in the Table no 1.
4.2 RESIDUAL PHENOL ESTIMATION
The standard curve was plotted (O.D Vs concenteration of phenol). O.D was obtained by
spectrophotometric measurement at 630nm. This standard curve (Figure no.1) was used
to calculate the residual phenol i.e. the remaining phenol after degradation by isolated
strains by using the formula Y=0.110x + 0.041, where Y=O.D. Now the value of X was
obtained which was again multiplied by the difference in concentration, here the
difference is 200mg/l
The 3 most potent bacteria capable of degrading phenol were found to be S1G, S4A and
S5C having the degradation rate of 97.27%, 89.39% and 84.54% respectively.
The residual phenol concenteration of all 8 isolates is shown in Table 3.
24
4.4 RESIDUAL PHENOL CONCENTERATION mgl-1 after 6, 12, 18 and 24 hr
incubation (wavelength 630nm)
The Strain S1G was found to degrade 21.67% after incubation of 6 hr further incubation
degrades 97.27% at the end of 24 hr.
The rate of degradation is shown in table no.4
The strain S4A was found to degrade 18.49% phenol after incubation of 6hr which further
increases to 89.39% at the end of 24 hr incubation.
The rate of degradation is shown in Table no.5.
The strain S5C was found to degrade 12.42% phenol after incubation of 6 hr which
further increases to 85.54% at the end of 24 hr incubation.
The rate of degradation is shown in table no.6.
4.5 COMPARATIVE STUDY OF PHENOL DEGRADATION BY IMMOBILIZED
CELLS ANF FREE CELLS.
To study the enhanced biodegradation of phenol the cells after centrifuged at 5000rpm for
10min. Were immobilized and incubated at 37oC and the broth containing live cells which
equals to the wet weight of centrifuged cells were incubated at 37oC and the degradation
studies were compared. The data is shown in table no.7 and graph is shown in Figure no.3
Table - 1: MICROSCOPIC, GRAM STAIN AND BIOCHEMICAL CHARACTERS
OF THE ISOLATED STRAINS:
Microscopic
Examination
Gram stain Simmonn
Citrate
Chrisren’s
urease test
Lactose test
25
S1G Cocci Gram +ve -ve Weakly +ve +ve
S2B Oval shaped Gram +ve -ve -ve +ve
S1H Rod shaped Gram –ve -ve -ve +ve
D1 Rod shaped Gram –ve -ve -ve -ve
D5 Cocci bacilli Gram –ve +ve -ve +ve
S4A Rod shaped Gram –ve -ve -ve +ve
S4C Cocci Gram +ve +ve -ve -ve
S5 Rod shaped Gram +ve -ve -ve +ve
Table 2: STANDARD CURVE AT 630nm
Concentration (mg/l) O.D
200 01.14
400 0.272
26
600 0.375
800 0.479
1000 0.589
Figure 1 : Standard Curve (O.D Vs Concenteration)
Table 3: RESIDUAL PHENOL ESTIMATION
The spectrophotometric measurement was done at wavelength 630nm
Strain O.D Residual Phenol (mg/l)
S1G 0.05 16.37
27
S2B 0.170 234.54
S1H 0.158 212.72
S4A 0.076 63.63
S4C 0.186 263.63
S5C 0.092 92.72
D1 0.180 252.72
D5 0.194 278.18
Figure 2: Residual Phenol Estimation
The 3 most potent bacteria capable of degrading phenol were found to be S1G, S4A and S5C having the degradation rate of 97.27%, 89.39% and 84.54% respectively.
28
Table - 4: RESIDUAL PHENOL CONCENTERATION mgl-1 after 6, 12, 18 and 24 hr incubation (Wavelength 630nm)
Strain Time(hrs) 6 12 18 24
O.D(630nm) 0.30 0.21 0.12 0.05
S1G Residual Phenol(mg/l) 470.90 307.27 143.63 16.37
% Removal of phenol. 21.67 48.17 76.06 97.27
Table - 5: RESIDUAL PHENOL CONCENTERATION mgl-1 AFTER 6, 12, 18 and 24 hr incubation (Wavelength 630nm)
Strain Time(hrs) 6 12 18 24
S4A O.D (630nm) 0.31 0.22 0.15 0.076
Residual phenol (gm/l) 489.09 325.45 198.18 63.63
% Removal of phenol 18049 45.375 66.97 89.39
29
Table - 6: RESIDUAL PHENOL CONCENTERATION mgl-1 AFTER 6, 12, 18 and 24 hr incubation (Wavelength 630nm)
Strain Time(hrs) 6 12 18 24
O.D (630nm) 0.33 0.24 0.17 0.092
S5C Residual phenol (gm/l) 525.45 361.81 234.54 92.72
% Removal of Phenol 12.425 39.69 60.91 85.54
Figure 3: Phenol degradation by three most potent isolated at different time
The above graph shows that S1G was the most potent phenol degrader with degradation
of 97.27% after the end of 24hrs while the strains S4A and S5C were found to degrade
89.39% and 84.54% respectively.
30
Table 7: COMPARATIVE STUDY OF PHENOL DEGRADATON BY IMMOBILIZED S1G AND FREE S1G CELLS (Wavelength 630nm).
Phenol degradation study by immobilized cells
Time(Hrs) 6 12 18
O.D 0.29 0.185 0.05
Residual
Phenol(mg/l)
452.72 261.81 16.63
Phenol Degradation Study by live cells
Time (Hrs) 6 12 18
O.D 0.30 0.21 0.12
Residual
Phenol(mg/l)
470.90 307.27 143.63
Figure 4: Comparative study of the phenol degradation by immobilized and free cells
31
CONCLUSION
32
CONCLUSION
Phenol and its components are extremely toxic and can easily be isolated from different
industrial sewage such as oil refinery, petrochemical industry and mines, especially
collier and chemical factories. Hence the presence of these compounds in the
environment could cause environmental pollution, especially in water resources. In the
past, physicochemical method was used for the elimination of phenol and its compounds,
but today, bioremediation is preferable. The aim of this study is to isolate and identify
phenol degrading bacteria from industrial effluents and study the phenol degradation rate.
Till date several works are in progress to isolate new and efficient microbial strain that
have ability to degrade phenol. We report here a new, isolate as a potential selected strain
to utilize phenol as sole source of carbon and energy. The degradation ability of isolate
was checked up to 600mg/l concentration. The determination of intermediates of phenol
degradation was also determined. This work has provided a useful guideline in evaluating
potential phenol biodegrades isolated from environment. Different methods have been
used for the elimination of phenol, but the use of bacteria can be one of the cheap and
secure methods. Bacteria with rapid reproduction in the presence of phenol and its
compound have shown extraordinary ability in phenol elimination. So with isolation,
purifying and growing of species which has high ability of phenol elimination, they can
be used in areas with phenol pollution. Different bacteria of different genus have been
isolated as phenol degrading. Most of them, which chiefly belong to family
Pseudomonaceae, are gram negative. Kounty et al. (2003) isolated phenol degrading
bacteria from Siberia soils. They found out that the permanent genus of phenol degrading
in these soils is Pseudomonas and especially, Pseudomonas putida. Although, this agrees
with the finding, their vast distribution in soils and the ability of phenol or phenol
compounds to be eliminated was done by Williams and Sayers (1994) and Torres et al.
(1999).
33
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