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ISOLATION AND CHARECTERISATION OF AEROBIC HYDROLYTIC BACTERIA
A Project Report Submitted in partial fulfillment for the award of
Degree of M. Sc. Biotechnology
SUBMITTED BY
SUMIT KUMAR SINGH
UNDER THE SUPERVISION OF Dr. V. VASUDEVAN
SCIENTIST ‘B’
DIVISION OF BIOTECHNOLOGY DEFENCE RESEARCH AND DEVELOPMENT ESTABLISHMENT
GWALIOR
BOSTON COLLEGE FOR PROFESSIONAL STUDIES JIWAJI UNIVERSITY
GWALIOR-474006
CERTIFICATE
This is to certify that Mr SUMIT KUMAR SINGH, student of M. Sc (Biotechnology),
Department of Biotechnology, Boston College for Professional Studies, Jiwaji
University, Gwalior has completed his dissertation work entitled “ISOLATION AND
CHARECTERISATION OF AEROBIC HYDROLYTIC BACTERIA” under my
supervision from 24.02.2010 to 03.09.2010 towards the partial fulfillment of post
graduate degree in Biotechnology.
Gwalior Signature of Supervisor Date Dr V. Vasudevan
Scientist ‘B’ Defence R & D Estt, Gwalior – 474 002
Government of India Ministry of Defence Defence R&D Organization Defence R&D Establishment Biotechnology Division Jhansi Road, Gwalior- 474 002 (MP) INDIA
Gram : DEFRES Telex : 0786- 212 Phone : 0751-2233489, 2341848,
2230344 Ext 278 Fax : 91-751-2341148 Email : [email protected]
DECLARATION
I hereby declare that the dissertation work titled “ISOLATION AND CHARECTERISATION OF
AEROBIC HYDROLYTIC BACTERIA” submitted to the Department of Biotechnology, Boston
College For Professional Studies, Jiwaji University, Gwalior in partial fulfillment of requirement
for the award of Master of Science in Bio-Technology, is the dissertation work done by me under
the supervision and valuable guidance of Dr. V. Vasudevan, Scientist ’B’ Biotechnology
Division, at Defence Research Development & Establishment (D.R.D.E) Gwalior (M.P.)
Signature of the candidate
ACKNOWLEDGEMENTS
The present research work is brought to conclusion with the help of several people
who came on the way, selflessly lending support. I feel greatly privileged for having
worked at the Division of Biotechnology, Defence R & D Establishment, Gwalior.
It is my privilege to express my deep sense of gratitude to my supervisor Dr. V.
Vasudevan, Scientist ‘B’ Division of Biotechnology, DRDE, Gwalior, for his guidance
and innumerable supports. I am highly obliged for his encouragement every time when I
feel impossible.
I am deeply grateful to Dr. R. Vijayraghavan, Director, DRDE, Gwalior for
providing me this opportunity to pursue my dissertation in this reputed establishment. I
am also thankful to Dr. S.K. Raza, Head Technical Coordinator HRD, DRDE, Gwalior,
for introducing me this reputed research institute.
I am greatly thankful to Dr. Om Kumar Scientist ‘F’ Head of Division of
biotechnology DRDE Gwalior for his motivation and appropriate suggestions for my
dissertation work.
I am also thankful to Dr. D.V. Kamboj, Scientist, ‘E’, Dr. M.K. Agrawal
Scientist ‘D’, Dr. R.K. Dhakad, Scientist ‘D’, Dr. Imteyaz Alam, Scientist ‘D’, Dr.
A.K. Goal, Scientist ‘D’, Dr. S. Ponmoriappan, Scientist ‘C’, Mr. Arvind Tomar
Scientist ‘B’, Ms Pallvi Gupta, Scientist, ‘C’, Dr. D.P. Nagar, STA for their support
and guidance during my dissertation work.
I express my sincere gratitude to Dr. G. Tejowati, Head and Dean , Division of
Biotechnology, Boston College for Professional Studies, Jiwaji University, Gwalior, Dr
Meenu Rai, Principal, Boston College for Professional Studies, Jiwaji University,
Gwalior for providing me excellent opportunity and encouragement during course of
study.
My grateful thanks to Mr. Brijendra Kumar Kashyap, JRF, DRDE Gwalior who
help me out when I felt difficult without his kind co-operation and critical suggestion it
would have been impossible for me to get through this work.
I am extremely thankful to Mr. Pawan Kumar Singh, SRF, Smt. Meenu
Jain, SRF, Mr. Manglesh Kumar Singh, SRF, Ms. Swati Jain, Ms. Richa, Ms.
Preethika Arya Division of Biotechnology, DRDE Gwalior for their help and guidance
for my work.
I am extremely thankful to my close friends and colleagues Mr Rinchen Tshering
Lepcha, Mr Deben Patak, Ms. Sarita Dubey, Ms. Ruby Srivastava and Ms Radha
bhadoria, those who made my stay pleasant during this study through their Tea parties
and quarrels.
My heartful thanks to my parents, for their blessing, encouragement, affection,
inspiration and their emotional, financial and moral support in every step I took .
My heartful thanks to my brother Mr Sushil Kumar Singh, to my sister Ms
Priyanka Singh and my whole family for their love encouragement blessing, support for
my study.
(Sumit Kumar Singh)
Contents
Page No. 1. Introduction 1 - 3
2. Review of Literature 4 - 18
3. Materials and Methods 19 - 28
4. Results 29 - 35
5. Discussion 36 - 38
6. Reference 39 - 50
7. Appendix i - iv
1
INTRODUCTION
Nature is the very beautiful and scientifically balanced creation of nature itself by
evolution. Nature balance itself by creating and destroying the things, and then creating new one,
and so on. Destroying the things in the natural ways is now known as degradation or
biodegradation, because it is done by biological or biochemical ways.
Most of the things in the nature that create life are made up of complex hydrocarbon
molecules (organic molecules). The degradation of these hydrocarbons is generally done by the
microorganism present in the environment, by the hydrolysis of that complex hydrocarbon
molecule into simple small one. Now this simple molecule is available for the nature to create
the things again.
There are four types of complex hydrocarbons which are largely present in the nature, 1)
cellulose, 2) protein, 3) starch, and 4) lipid. The degradation i.e. hydrolysis of these complex
hydrocarbons into simple molecule is done by the microorganism, for the utilization of that
simple molecule as their source of nutrient. The degradation of the complex hydrocarbons is
extracellular, because of the size of these molecules is so big that these are unable to cross the
cell wall of the microbial cell, that’s why the microorganism produce extracellular enzymes that
hydrolyze these complex hydrocarbon into simple and small molecule, which can easily cross
through the cell wall of the microorganism. The degradation of these molecules is largely done
the bacteria by producing extracellular enzyme like cellulase, protease, amylase, and lipase, for
the degradation or hydrolysis of cellulose, protein, starch, and lipid respectively.
2
As the human developed and population increases day by day, the accumulation of solid
waste product increases, which cause pollution to for environment. These pollutants normally
have agro waste, and city waste etc. These wastes have the high content of complex hydrocarbon
molecule (organic molecules) that cause the depletion of the BOD and COD level of the water
bodies, which directly affect the aquatic micro flora and fauna. But one of the major pollutant of
water bodies human waste which is deposited in the chamber and after that through out to the
outside of the city by municipal system. But Human waste (night soil) is the one of the biggest
and dangerous pollutant for the water bodies, because it makes the water into store house of
pathogenic bacteria which cause water born disease to human beings, like diarrhea, typhoid,
cholera, etc.
Since biodegradation is the process by which these harmful complex organic compounds
(hydrocarbons) can broken-down into small and simple one by the extracellular enzyme
producing microorganisms. These organic compounds can be degraded by the two group of
microorganism 1) aerobic, and 2) anaerobic. Aerobic microorganism can degrade organic
compound with the help of oxygen while anaerobic on doesn’t use oxygen for the degradation of
it. The degradation of the organic compound anaerobiclly produces pungent gases and unable to
degrade the compound efficiently, but aerobic degradation of this have the high efficiency of
degradation, and not produce pungent gases. So aerobic degradation of these organic compounds
(pollutants) is one of the best method to overcome from this biggest problem called pollution.
Microbial diversity of these type microorganisms is not well known till now, so
biochemical characterization of this organism provides the better understanding patterns of
degradation and provides the manipulation of degradation of the organic compound.
3
With this background, the present study entitled “ISOLATION AND CHARECTERISATION
OF AEROBIC HYDROLYTIC BACTERIA” was undertaken with following objectives:
To elucidate qualitative enzymatic profiles of bacterial isolates.
To estimate the quantitative production of hydrolytic enzymes by efficient isolates.
molecular characterization and RAPD Profiling of bacterial isolates.
4
Review of Literature
Sustainable environmental management is recently becoming an
issue of global concern. Around the world, solid waste management
becomes an important issue in urban areas. Currently, the generation of
wastes has gained an important consideration in modern societies as a
result of changes in habits and lifestyle of consumers, along with economic
development. Due to the steady increase in population, urbanization, and
industrialization, Municipal Solid Waste generation has been increasing
over the last decade. For example, in Developing countries, about 38,000
ton/day of refuse was collected in the year 2002 as compared to 29,000
ton/day in 1992 (Chaya & Gheewala, 2006).
An important feature often cited when dealing with urbanization of
the developing world is the rapid growth of cities and metropolitan areas.
Since Asia urbanizes, solid waste production increased as urban residents
generate 2-3 times more solid waste than the rural counterparts. Urban
areas in Asia today generate about 760,000 tons of waste per day, and by
2025 will produce about 1.8 million tons per day (Chaya & Gheewala,
2006). Moreover, emerging mega cities will increase solid waste production.
By 2015, an estimation of 21 cities in the world will have populations of 10
million or more, ten of these cities will be in Asia (Wheler, 2004). The above
mentioned development will lead to the use of desperate measures in
5
urban solid wastes management unless a critical management and
disposal is considered.
During biodegradation pathways in aerobic digestion, hydrolysis
involve depolymerisation of the organic polymers by hydrolytic-fermentative
bacteria, through the action of extracellular hydrolytic enzymes namely,
cellulases, proteases and lipases, into oligomers and monomers (sugars,
amino acids, long-chain fatty acids and glycerol. Bacteria excrete enzymes
that hydrolyse particulate substrates to small molecules, which can pass
through the cell membrane. Once inside the cell, these simple molecules
are oxidized to provide energy and to synthesize cellular components.
When microbial process is completed, under ideal conditions, the
microbial cells will aggregate and form a settable floc structure (active
biomass) that is formed when organic matters is oxidized and degraded by
microorganism. In addition, the ratio of organic matters added to the active
microbial biomass can be varied. A lower rate system (low nutrient input
per unit of microbial biomass), with slower growing microorganism’s will
produce an effluent with low residue level of dissolved organic matter. A
high rate system (high nutrient input per unit of microbial biomass) with
faster growing microorganism will remove more dissolved organic carbon
per unit time, but produce a poor quality effluent. All aerobic process
produces excess microbial biomass, or sewage sludge, which contain
many recalcitrant organics.
Plant growth is important in terrestrial eco-system; equally important
are plant death, decomposition and recycling. This released soluble
material creates a residue sphere in an area between decaying plant
6
materials and the soil in the hot environmental conditions, which drives a
wide range of microbiological processes. After soluble materials have
leached from the plant, the remaining starch, cellulose, and proteins are
degraded under aerobic and anaerobic conditions. Lignin, a random
aromatic cycling polymer that contains number of N2 molecule is the one
exception. Its degradation requires O2. Some microbes secrete specific
enzymes used for degradation of lignin polymers as Laccases and
Phenoloxidase enzymes. Originally it was assumed that in given time by
almost infinite variety of microorganisms all organic compounds including
those synthesized in the laboratory, would eventually degrade in natural
environments. However, ecologists began to raise question about the
ability of microorganism’s to degrade these varied substrates, and the role
of the environment (clays, anaerobic condition) in protecting some
chemicals and the synthetic pesticides. It became distressingly evident
that not all organic compounds are immediately biodegradable.
Degradation of the complex compound takes place in the several
stages. Humic acids, brownish polymeric residues of lignin decomposition
that accumulate in the soil and water act as electron acceptors under
anaerobic process. Microbial communities change their characteristics in
response to the addition of inorganic or organic substrates. If a particular
compound, such as herbicide, is added repeatedly to a microbial
community, the community adapts and faster rate of degradation can occur.
Degradation process that occurs in soils also can be used in large
scale degradation of hydrocarbon or wastes from agricultural operations, in
7
a technique called land farming. The waste materials is incorporated in to
the soil or allowed to flow across the soil surface, where degradation
occurs. In fact, the partial degradation or modification of an organic
compound may not lead to decreased toxicity. Biodegradation also can
lead to wide spread damages and financial losses. Metal corrosion is a
particular important example. Microbial mediated corrosion of metals is
particularly critical where iron pipes are used in waterlogged anaerobic
environments or in secondary petroleum recovery process carried out at
older oil fields. Microorganism’s that use elemental iron as an electron
donor during the reduction of CO2 in methanogenesis has recently been
discovered.
The use of plants to stimulate the extraction, degradation, adsorption,
stabilization of contaminants is becoming an important part of
biodegradation technology. A plant provides nutrients that allow co-
metabolism to occur in the plant root zone or rhizosphere.
Phytoremediation also includes plant contribution to degradation,
immobilization, and volatilization processes. Psychrotrophic coliform was
also isolated from faecal polluted environments.
It has been well established that a primary role of bacteria in fresh
water environment is the degradation of particulate organics, recycling of
nutrients, the synthesis of vitamins and other growth factors. Temperature,
pH and dissolved oxygen were the most important parameters affecting
bacterial populations in a high nutrient rich lake, whereas in a nutrient
deficient lake, the level of particulate matter, pH and rainfall were the most
8
important governing factors. In certain lakes of temperate region, bacteria
have been found to posses the ability of adaptation and profounded growth
by utilizing the dissolved nutrient over the seasonal temperature ranges
(Boylen and Brock, 1973). Several different approaches were used to
increase biodegradation. These included nutrients additions, chemical
dispersants, biosurfactant additions, and the use of high pressure steam.
The capacity of microorganisms to assimilate organic matter
depends on their ability to produce the enzymes needed for degradation of
the substrate components e.g. starch, cellulose, hemicellulose and lignin.
The more complex the substrate, the more extensive and comprehensive is
the enzyme system required. Through the synergistic action of
microorganisms, complex organic compounds are degraded to smaller
molecules which can then be utilized by microbial cells (Golueke, 1991,
1992).
Amylase: Alpha-amylases (endo-1, 4-α- D-glucan glucanohydrolase EC 3.2.1.1) are
extra-cellular endoenzymes that randomly cleave α-1,4 linkages between
adjacent glucose units in the linear amylose chain and ultimately generate
glucose, maltose, and maltotriose units. This class of industrial enzymes
constitutes approximately 25% of the enzyme market covering many
industrial processes such as sugar, textile, paper, brewing, baking, distilling
industries, preparation of digestive aids, production of cakes, fruit juices,
starch syrups, and pharmaceuticals (Amoozegar et al., 2003; Ashger et al.,
2007). Demand for novel amylases worldwide is increasing day by day, as
these enzyme application spectra are spreading in various industrial
9
sectors. In view of this, researchers have diverted their attention for
isolation and characterization of enzymes from extremophiles. Amylase
production has been reported in eubacterial moderate halophiles such as
Acinetobacter, Micrococcus halobius, Micrococcus varians subsp.
halophilus, other Micrococcus isolates, and Halomonas meridian (Bernfeld,
1998).
Lipase:
The influence of lipid concentration on hydrolysis and
biomethanation of an artificial lipid rich (triolein) waste was evaluated.
When the effect of lipase addition on enzymatic hydrolysis of lipids was
studied, results showed that the higher the enzyme concentration, the more
accentuated was the inhibition of the methane production. The enzyme
seems to enhance the hydrolysis and produced intermediates are causing
inhibition of the later steps of the degradation process.
Lipid rich waste from food processing industry, slaughterhouse,
edible oil processing, dairy products industry, are attractive substrates for
anaerobic digestion due to the higher methane yield obtained when
compared to proteins or carbohydrates. Besides causing operational
problems in the anaerobic digesters due to clogging, these lipids may as
well lead to mass transfer problems for soluble substrates since they
adsorb on the microbial biomass surface. The flotation of biomass due to
fat adhesion may as well cause loss of active biomass through the outlet of
the digester (Cammarota et al., 2001).
Few studies have been conducted to investigate the influence of
lipid concentration on hydrolysis dynamics. Nevertheless, to study the
10
hydrolysis process for a wide range of concentrations of lipids would allow
better understanding of the process. Some studies have been reported on
the area but the, lipid amount is in most cases lower than 5% (w/v)
(Cammarota et al., 2001). Furthermore, a process configuration allowing
higher lipid concentration would improve the process economics.
Cellulase:
Cellulose is a linear condensation polymer consisting of D-
anhydroglucopyranose joined together by β-1,4- glycosidic bonds with a
degree of polymerization (DP) from 100 to 20,000 ( O'Sullivan, 1997;
Zhang and Lynd, 2004). Anhydrocellobiose is the repeating unit of cellulose.
Coupling of adjacent cellulose chains and sheets of cellulose by hydrogen
bonds and vander Waal's forces results in a parallel alignment and a
crystalline structure with straight, stable supra-molecular fibers of great
tensile strength and low accessibility (Demain et al., 2005; Nishiyama et al.,
2003; Notley et al., 2004; Zhang and Lynd, 2004).
The cellulose molecule is very stable, with a half life of 5–8 million
years for β-glucosidic bond cleavage at 25°C (Wolfenden and Snider,
2001), while the much faster enzyme-driven cellulose biodegradation
process is vital to return the carbon in sediments to the atmosphere (Berner,
2003; Cox et al., 2000; Falkowski et al., 2000).
Cellulases, responsible for the hydrolysis of cellulose, are composed of a
complex mixture of enzymes with different specificities to hydrolyse
glycosidic bonds. Cellulases can be divided into three major enzyme
classes (Rabinovich et al., 2002). These are:
11
- Endoglucanase or endo-1,4-β-glucanase (EC 3.2.1.4)
- Cellobiohydrolase (EC 3.2.1.91)
- β-glucosidase (EC 3.2.1.21)
Endoglucanases, often called carboxy methyl cellulases (CMCase), are
proposed to initiate attack randomly at multiple internal sites in the
amorphous regions of the cellulose fiber opening–up sites for subsequent
attack by the cellobiohydrolases (Wood, 1991). Cellobiohydrolase, often
called as exoglucanase, is the major component of the fungal cellulase
system accounting for 40-70% of the total cellulase proteins and can
hydrolyse highly crystalline cellulose (Esterbauer et al., 1991).
Cellobiohydrolases remove mono-and dimers from the end of the glucose
chain. β-glucosidase hydrolyze glucose dimers and in some cases cello-
oligosaccharides to glucose. These three hydrolysis processes occur
simultaneously as shown in below figure. The widely accepted mechanism
for enzymatic cellulose hydrolysis involves synergistic actions by
endoglucanase, exoglucanase or cellobiohydrolase, and β-glucosidase
(Lynd et al., 2002; Zhang and Lynd, 2004). Microorganisms generally
appear to have multiple distinct variants of endo-and exo-glucanases
(Beldman et al., 1987 and Shen et al., 1995).
12
Primary hydrolysis that occurs on the surface of solid substrates
releases soluble sugars with a degree of polymerization (DP) up to 6 into
the liquid phase upon hydrolysis by endoglucanases and exoglucanases.
The enzymatic depolymerization step performed by endoglucanases and
exoglucanases is the rate-limiting step for the whole cellulose hydrolysis
process. Secondary hydrolysis that occurs in the liquid phase involves
primarily the hydrolysis of cellobiose to glucose by β-glucosidases,
although some β-glucosidases also hydrolyze longer cellodextrins (Zhang
and Lynd, 2004). The combined actions of endoglucanases and
exoglucanases modify the cellulose surface characteristics (topography)
over time, resulting in rapid changes in hydrolysis rates.
There are many genera of bacteria which can degrade
lignocellulosic materials (Buswell and Odier, 1987). Li et al. (1997) studied
the production and properties of cellobiose oxidizing enzyme from
13
Cytophaga spp. LX-7. Haack and Breznak (1993) investigated the xylan-
degrading enzymes of Cytophaga xylanolytica that anaerobically
decompose the biopolymers. Many other bacterial species of
Pseudomonas, Acinetobacter, Bacillus and Clostridium degrade lignin
(Janshekar and Fietchter, 1983). Godden et al., (1992) reported that
cellulolytic Cytophaga and Sporocytophaga seem to play a role during
decomposition of cattle manure. Chang and Thayer (1975) had studied the
digestion of hardwood mesquite (Prosopis spp.) with cellulose
decomposing Cytophaga strain with the aim of improving the feed quality of
plant material for cattle.
Cellulomonas and Cytophaga are the mesophilic bacteria able to
degrade cellulose, while Clostridium thermocellum is a thermophilic
anaerobic cellulose degrader (Bhat and Bhat, 1997). Thanikachalam and
Rangarajan (2003) carried out protein enrichment of rice straw with a strain
of Cellulomonas (CBS 7) and reported that maximum cellulose utilization
(73.2%) was recorded with ammonium sulphate as nitrogen source.
The lignin degrading eubacteria can be divided into erosion,
cavitation and tunnelling bacteria (Eriksson et al., 1990 and Blanchette,
1995). Erosion bacteria grow towards the middle lamella of the wood cells
and cause erosion of the fibre wall, while tunnelling bacteria grow within the
cell wall. Bacteria of several genera such as Pseudomonas, Alcaligenes,
Arthrobacter can degrade single ring aromatic components. The role of
bacteria may be significant in consuming the small molecular weight
intermediate components produced by lignolytic fungi (Ruttimann et al.,
1991).
14
Kawakami and Shumiya (1983) studied the degradation of lignin
and lignin related compounds by alkalophilic bacteria. Haider et al. (1978)
showed that the Bacillus strain was able to convert 14C (side chain) lignin of
spruce to 14CO2. The rate was comparable to that of fungi upto 35 days.
Deschamps et al. (1981) demonstrated delignification of bark chips by a
mixed culture of Bacillus and Cellulomonas, which were effective singly.
Although aerobic microorganisms are generally lignin degraders in most
environments, it has been shown that anaerobic rumen microorganisms are
capable of degrading plant fibre cell walls (Kuhad et al., 1997).
Many bacterial strains especially actinomycetes, can solubilise and
modify the lignocellulosic structure extensively, but their ability to mineralize
lignin is limited (Ball et al., 1989, Eriksson et al., 1990 and Godden et al.,
1992). Thermophillic actinomycetes have been investigated for utilizing
paper mill fines and feed lot wastes but these processes were not found
feasible for commercial purpose, because of very slow growth rate and
requirement of sterile conditions and high pH etc. (Humphery et al., 1977).
Actinomycetes follow a characteristic pattern of lignocellulose
decomposition with the release of lignin rich, water soluble fragments that
are slowly metabolized thereafter (Vicuna, 1988).
Barder and Crawford (1981) reported that Streptomyces badius can
degrade milled wood lignin and it was enhanced when organic nitrogen and
organic carbon substrate were added to the medium. Antai (1985) selected
three Streptomyces strains which were lignolytic, out of which one was the
most rapid lignocellulose decomposer, depleting 42 per cent of lignin and
50 per cent of carbohydrate of lignocellulose after 12 weeks incubation.
15
Zimmermann and Broda (1989) had grown Streptomyces
viridosporus in lignocellulose supplemented medium, which released
coumaric acid and vanillic acid, intermediates of lignin degradation
(Donnelly and Crawford, 1988). In view of the energy and power crises
besides the high cost of agricultural inputs such as fertilizers, pesticides
and irrigation water, agricultural wastes are now considered quite an
important component of farming especially those based on organic systems
including crop residues. Organic amendments in the form of agricultural
wastes and crop residues activate the autochthonous microorganisms of
the soil, indirectly stimulate the biogeochemical cycles therein (Pascual et
al., 1997) and provide various minerals (e.g. N, P, and S) essential for plant
nutrition.
Molecular characterization:
An array of molecular techniques, such as amplified ribosomal DNA
(rDNA) sequencing, amplified rDNA restriction analysis, and temperature
and denaturing gradient gel electrophoresis (TGGE and DGGE) of rDNA,
has been applied to elucidate microbial population structures in the
environment (Borneman et al., 1996; Heuer et al., 1997; Hiorns et al., 1995;
Torsvik et al., 1996). Application of these molecular methods has led to a
tremendous increase in knowledge of microbial ecology. Ishii and Takii
(2003) analysed microbial communities during the composting of sewage
sludge and food-waste samples by DGGE and established that the
concentration of dissolved organic materials is very important in deciding
the microbial community composition of a compost microenvironment.
16
Ribosomal DNA genes are tandemly repeated multigene families
containing both genic and nongenic, or spacer, regions. Each repeat unit
contains a copy of the 18 s-, 5.8 s, and 28 s-like rDNA and two spacers, the
internal transcribed spacer (ITS) and an intergenic spacer (IGS). The 5.8s
rDNA gene is typically flanked by a bipartite ITS, the ITS1 and ITS2, which
separates the 5.8s rDNA from the 18s and 28s genes, respectively (Garber
et al., 1988).
Numerous studies have demonstrated that variable numbers of
tandemly repeated sequences, or subrepeats, within the IGS account for
the bulk of the length variation within the rDNA repeat unit (Rogers and
Bendich, 1987). Ribosomal RNAs (rRNAs) provide a powerful taxonomic
indicator, because they are highly conserved and are universally found in
living cells. The 5s rRNA was first used for this purpose (reviewed by Hori
and Osawa, 1987).
However, the 5s rRNA is so short and so conserved that it cannot be
used for studying closely related species; for such species one has to look
at larger rRNA molecules: 16s (Salim and Maden 1981; Woese et al., 1985)
and 28s (Qu et al., 1983). The development of a technique for rapid and
easy sequencing of large stretches of 18s or 28s rRNA opened the way for
systematic exploitation of the remarkable properties of these molecules as
phylogenetic indicators (Qu et al., 1988).
Tiquia and Michel Jr. (2002) determined the phylogenetic diversity of
bacterial communities in livestock manure compost on the basis of T-RFLP
of 16S rRNA genes and found a remarkable difference in the bacterial
community structure of in vessel and windrow composts from that of the
17
feedstock (fresh manure). Liang et al. (2006) analyzed the phylogenetic
relationships of 49 specimens comprising 14 morphologically similar
species of Pucciniastrum distributed in Japan based on the sequence data
of the large subunit (28S) rDNA, 5.8S rDNA, and internal transcribed
spacer (ITS) regions. The ITS regions were analyzed by Monreal et al.
(1999) with restriction enzymes that digested the amplified DNA into
discrete fragments.
Jackson et al. (1999) showed that polymorphism analysis of the
rDNA intergenic regions is a valuable technique both for strain typing and
species identification in the dermatophyte fungus Trichophyton rubrum and
the related fungi. Guadet et al. (1989) was able to evaluate the divergence
between fifty-two closely related strains from eight species of Fusarium by
sequencing two highly variable stretches (of 138 and 214 nucleotides
respectively) of the 5’ end of the 28S rRNA molecule and concluded that
this method is suitable for establishing a precise phylogeny between
closely related species within a genus.
Various nucleic acid fingerprinting methods are now available for
characterizing the diversity, structure and composition of microbial
communities. Since RNA is quite unstable in environmental matrices and
DNA can remain detectable in the environment long after cell death, RNA-
based community profiling is more reflective of metabolically active
bacterial communities than DNA-based analysis (Fey et al., 2004). The
growth rate of bacteria is usually proportional to the intracellular level of
ribosomal RNA (rRNA), and it is generally accepted that the amount of
rRNA per cell is roughly proportional to a cell’s metabolic activity (Wagner,
18
1994). Shifts in microbial community structure and metabolic activity have
been studied by comparative analysis of 16S rRNA gene (rDNA)- and 16S
rRNA-based fingerprints, respectively (Nicol et al., 2003; Eichler et al.,
2006).
19
MATERIALS AND METHODS
3.1 Collection of samples:
The soil samples were collected from the solid waste disposal site
and market waste disposal site in Gwalior. Flask which contains soil was
fed with human waste and aerated continuously for 3 months to enrich
aerobic bacteria. Bacterial cultures were isolated based on their qualitative
enzyme profile. These bacterial colonies were kept for further experiments.
3.2 Isolation of bacteria:
Bacteria were isolated from the slurry, samples were serially diluted
in 10 fold by sterilized normal saline and 100 µl of each dilution was plated
on amylase, protease, lipase and cellulose media. The inoculated plates
were incubated at 300C for 24 hrs. Desired colonies were picked and
purified on fresh media plates.
3.3 Qualitative screening of Bacterial Isolates for hydrolytic enzyme
activity:
Production of extracellular enzymes was recorded in a semi-
quantitative manner as reported earlier (Alam and Singh, 2002). Minimal
medium containing appropriate substrate and 1.5% agar (w/v) were
inoculated with freshly grown cultures and incubated at 30oC for 24 hr,
After incubation, plates were developed and clearing zone / color of
chromogenic substrate around the colonies were scored by measuring the
diameter of hydrolysis / relative colour intensity.
20
For protease activity, casein (0.4 % w/v) was used and after
incubation, plates were developed by staining with amido black (0.1%) for
15 min followed by destaining with 40 % methanol. Amylase activity was
screened by using 0.4 % soluble starch (w/v). After incubation plates were
developed by flooding the plates with iodine solution (1%) and washed with
normal saline. Extracellular β–galactosidase activity was confirmed by
incorporating 1 mM ortho-nitrophenyl-β-D-galactopyranoside (ONPG) into
the agar medium. The yellow zone of hydrolysis was measured and scored
in a semi-quantitative fashion. Lipase test was carried out with 0.4 % (w/v)
tributyrin and 0.01% (v/v) tween-80 and observing the zone of hydrolysis.
Cellulase activity was tested using 0.4% (w/v) carboxy methyl cellulose as
a substrate. After incubation, plates were treated with 0.1% aqueous
congo-red solution for 10 min. Plates were washed with 1M NaCl solution
and clearing zone was scored. Alkalline phosphatase test was performed
with the help of chromogenic substrate p-nitrophenyl phosphate (pNPP) at
a concentration of 500 mM. After incubation positive results were indicated
by a zone of yellow color.
3.4 Quantitative Estimation of Hydrolytic Enzymes:
3.4.1 Amylase Quantitative Estimation:
Reagents Required:
1. Soluble starch (1%).
2. 0.05 M K2HPO4 – NaOH Buffer – pH 5.0 Adjust pH with NaOH.
3. Working DNSA (Di-Nitro Salicylic Acid)*.
* Preparation of DNSA reagent:
21
Solution A: Sodium hydroxide (10 g) was dissolved in 1000 ml distilled
water.
Solution B:
Dinitrosalicylic acid : 10 g
Phenol (crystalline) : 2 g
Rochelle’s salt (sodium potassium tartarate) : 200 mg
All these ingredients were dissolved in 1000 ml of solution A.
Solution C: Sodium sulphite (5 g) was dissolved in 100 ml distilled water.
This solution was prepared fresh every time.
Preparation of working DNSA reagent
Ninety nine ml solution B and 1 ml solution C was mixed to prepare
working solution. It should be prepared fresh before use.
Estimation:
Culture filtrate was taken (amount may vary according to activity) in a test
tube. 2 tubes were made for each sample (Sample and Enzyme Blank). 0.5
ml of starch solution was added in sample tubes and 0.5 ml of DW in
enzyme blanks. The volume was made up to 1 ml. The tubes were
incubated in water bath at 50oC for 30 minutes. 3 ml of DNSA solution was
added in all the tubes. The tubes were incubated at 100 oC for 16 minutes
in water bath. OD was taken at 575 nm.
3.4.2 Protease Quantitative Estimation (Razak et al., 1994)
Reagents Required:
1. Tris Buffer.
2. Casein (1 %)
3. Tri-chloro Acetic Acid
4. Sodium Carbonate (0.4 M)
5. Folins reagent (1 N)
6. Whatman No: 1 Filter paper
Estimation:
Culture filtrate was taken in a test tube (amount may vary according
to activity) with that 0.5 ml of Tris buffer and 1 ml of casein solution were
22
added and the volume is made up to 2 ml by adding 0.4 ml of DW. For
each enzyme Blank preparation 1 ml of DW and no casein was added. The
tubes were incubated in water bath at 37oC for 20 minutes. 4 ml of TCA
solution was added in all the tubes to terminate the reaction. The tubes
were kept in room temperature for 1 hour. The contents were filtered with
whatman No: 1 filter paper.
Development of colour:
1 ml of aliquot was taken from the previous step separately, to this 5
ml of 0.4 M Na2CO3 and 0.5 ml Folins Reagent were added. The sample
was vortexed and incubated for 20 minutes. Then OD was taken at 660 nm.
3.4.3 Lipase Quantitative Estimation (Lambrechts and Galzy. 1995)
Reagents Required:
1. 0.05 M p-Nitro phenol in Acetonitrile. (Dissolve 0.094 g of p-Nitro
phenol in 5 ml of Acetonitrile).
2. Citrate Phosphate Buffer (0.1 M).
3. Solution A: Prepare 0.1 M Citric acid by dissolving 2.101 g of Citric
Acid in 100 ml of DW
4. Solution B: Prepare 0.2 M solution of Dibasic Sodium Phosphate
(2.8392 g of Na2HPO4 in 100 ml of DW)
5. Solution C: 0.1 M Citrate Phosphate Buffer (pH 7.0) containing 0.15
M NaCl and 0.5 % W/V Triton X– 100. (NaCl 8.775 g was dissolved
in 1 Litre 0.1 M Citrate Phosphate Buffer and 0.5 g of Triton X-100
was added to it).
Estimation:
Culture filtrate was taken in a test tube (amount may vary according to
activity). 1ml of Solution C and 40 µl of 0.5 M p-Nitro phenol were added.
The tubes were incubated for 30 min at 300C . Then OD was taken at 430
nm.
3.5 Estimation of Total soluble protein: (Lowry et al., 1951)
Reagents:
Alkaline copper reagent (ACR) [to be prepared freshly before use]:
23
Solution A:
Sodium carbonate solution (2% w/v) was prepared in dilute Sodium
hydroxide soln. (0.1N).
Solution B:
Sodium potassium tartarate solution (1.0% w/v) was prepared in Distilled
Water
Solution C:
CuSO4.7H
2O solution (1% w/v) was prepared in Distilled Water.
Working solution of ACR = Soln. A: Soln. B: Soln. C: 98: 1: 1
Folin Ciocalteu’s reagent:
Commercially available reagent (2N) was diluted to 1N with addition of
equal volume of distilled water.
Estimation method:
Culture filtrate sample (0.5 mL) was added to test tube containing 5 mL of
alkaline copper reagent followed by 10 min incubation at 280C. After 10 min,
0.5 mL of 1N Folin Ciocalteau’s reagent was added to the tubes and these
were again incubated for 30 min at 280C. A blank was also prepared with
0.5 mL distilled water instead of sample. Absorbance was taken at 750 nm.
The protein concentration was estimated by referring to standard prepared
with Bovine serum albumin (10-100 µg/mL).
3.6.1 Isolation of total cell DNA from the Bacterial isolates (Modified
method of Charles and Nester, 1993):
Method:
Total genomic DNA of bacteria was isolated by the method of Charles and
Nester (1993) with slight modifications. Pure cultures of bacteria were
raised in 5 mL of TY medium broth for 18 – 24 hrs to obtain cell O.D. of 0.6
at 600 nm. The culture broth (1.5 mL) was pelleted in a microfuge tube for
24
2 mins. at 12000 rpm. The bacterial pellet was washed in 1.5 mL of 0.85%
NaCl, centrifuged for 2 mins. at 12000 rpm and was resuspended in 0.4 mL
Tris-EDTA buffer (T10
E25
). Cell Lysis was done by adding 20 µL of 25%
SDS, 50 µL of 1% lysozyme and 50 µL of 5M NaCl followed by incubation
at 680C for 30 mins in a circulatory water bath. For protein precipitation,
260 µL of 7.5 M Ammonium acetate solution was added to the microfuge
tubes and the tubes were kept in ice for 20 mins followed by centrifugation
at 13000 rpm for 15 mins at 200C. Supernatant was carefully pipetted out in
another fresh, sterile microfuge tube in which 1µL RNase (4 mg mL-1
) was
added followed by Incubation at 370C for 20 mins. Equal vol. of chloroform
was added in the tubes and proper mixing was done by inverting the tube
up and down several times. RNA was precipitated by Centrifuging for 1 min.
at 12000 rpm. The top layer containing total cell DNA was pipetted out in
fresh microfuge tube and used for next step. DNA was precipitated by
adding 0.8 vol. of isopropanol followed by incubation on ice for 30 mins and
pelleted by centrifuging at 10000 rpm for 15 mins. DNA was further washed
with 0.5 mL of 70 % Ethanol and spun down at 10000 rpm for 1 min.
Traces of ethanol were removed by air drying the tubes in inverted position.
Pure DNA sample was then suspended in 20 µL Tris-EDTA buffer (T10
E1)
and stored at 40C.
3.6.2 Quantification of genomic DNA: Quantification through Spectrophotometric method:
The genomic DNA was diluted 100 times and quantified by
measuring the absorbance at 260 nm. The amount of DNA was estimated
using the relationship that O.D. of 1.0 corresponds to 50 µg mL-1. The
purity of DNA was assessed by measuring A260/A280 ratio.
3.6.3 Quantification through Agarose Gel Electrophoresis
Method:The genomic DNA samples of bacteria, Actinomycetes and fungi
were quantified through agarose gel electrophoresis by analyzing their
migration on 0.8% agarose gels prepared in 0.5 M Tris-borate-EDTA (TBE)
25
buffer and run in an Electrophoresis tank filled with the same conc. of TBE
buffer. The genomic DNA was diluted with Tris-EDTA (T10
E1) buffer so as
to achieve a concentration of 50 ng in 10 µL to be used as a template DNA
in PCR amplification reaction.
3.6.4 Molecular Typing (RAPD):
Two random primers (Operon Technologies, USA), each 10-mer
long, were used separately in the RAPD study. These primers were OPA
10: 5' -GGGTAACGCC-3' and OPA 13: 5'GTGATCGCAG-3'. In order to
determine the typeability, reproducibility and discrimination of the each
primer, separate amplifications of each primer were conducted (three trials
for each primer). The output of each experiment was compared to the
previous one (Arbeit, 1994). Amplification reactions were performed
according to Williams et al. (1990) in volumes of 25 µL containing 0.5 µmol
L -1 primer, IX PCR Buffer (MgCl2 free) (Promega, USA), 2 mmol L -1
MgC12 (Promega, USA), 100 µmol L-1 of each dNTP (Promega, USA), 0.5
U Taq DNA polymerase (Promega, USA), 0.3 mg template DNA.
Nucleases free water (Promega, USA) was used to bring the reaction
volume to 25 µL.
PCR amplification was carried out in 0.2 mL thin walled, nucleases
free PCR tubes (Treff Lab, Switzerland) using iCycler thermocycler (BIO-
RAD, USA) programmed as follows: Initial denaturating step at 95°C for 3
min, followed by 40 cycles of 95°C for 30 sec, annealing at 36°C for 30 sec,
extinction at noc for 1 min. Finally, extension at 72°C for 7 min, after that
tubes were held at 4°C for direct use, or stored at 20°C until needed.
26
Data analysis of RAPD profiles:
The RAPD photographs were analyzed under a magnifying lens
over an engineer's disk supplied with horizontal and vertical rulers. The
fingerprints were recorded in the binary form i.e., 1 in case of presence of a
band and 0 when there is no band, to generate a binary matrix (Demeke
and Adams, 1994; Sneath and Sokal, 1973) for each primer. These binary
matrices were used to calculate the similarities and the differences
between the isolates by the SPSS software, using the simple matching
coefficient (Sneath and Sokal, 1973).
3.6.4 PCR-Amplification of 16 S rRNA gene from the bacterial isolates
Solutions and buffer for polymerase chain reaction (PCR):
All the chemicals were obtained either from Fermentas, U.K. or Banglore
Genei Pvt. Ltd., Bangalore, India.
Method:
Amplification of 16S rDNA was carried out by polymerase chain
reaction using a thermal cycler (M.J. Research PTC-100). The PCRs were
carried out with 50-90 ng of pure genomic DNA. The amplification reactions
were performed in a 25 µL mixture containing 0.6 U of Taq DNA
Polymerase (Genei from 3U µL-1
), 2.5 µL of 10X Taq Polymerase buffer,
0.4 µL of dNTP mix and 0.3 µL each of the two primers described above.
The Following programme was used for the amplification of 16 S rDNA.
Step 1: 94 0 C for 5 minutes
Step 2: 94 0 C for 30 seconds
Step 3: 58 0 C for 40 seconds
Step 4: 72 0 C for 1.30 minutes
Repeat step 2 to 4, 40 times (40 cycles)
27
Step 5: 72 0 C for 10 minutes
Step 6: 4 0C forever.
For every PCR reaction, a negative control (no template DNA and no
primer) and a positive control (template DNA giving amplified product) were
invariably maintained. The amplified product was run on a 1.2 % agarose
gel along with 1 kb MW marker, at a constant voltage, and visualized under
UV light.
3.6.5 Purification of amplified PCR product
In order to remove traces of reagents used during amplification
reaction the amplified 16S rDNA obtained after PCR amplification were
purified and concentrated using Qiaquick PCR purification kit, obtained
from Qiagen (Germany). The kit included Qiaquick spin columns, Buffers
PB and PE, and elution buffer. The protocol designed for purification
renders DNA fragments ranging from 100 bp to 10 kb purified from primers,
nucleotides, polymerases, and salts. 100 µL of PCR reaction was mixed
with 500 µL of buffer PB. Qiaquick spin column were placed in a provided 2
ml collection tube. The total sample (600 µL) was applied to the column
and centrifuged at 13,000 rpm for 30-60 seconds. The flow-through was
discarded. The column was placed back on 2 mL collection tube and
washed with 750 µL of buffer PE and again centrifuged for 30-60 seconds.
The flow-through was discarded and the column was centrifuged for an
additional one minute at 13,000 rpm. The Qiaquick column was placed on a
clean 1.5 mL microfuge tube. To elute DNA, 50 µL of milli Q water was
applied at the centre of the Qiaquick membrane and the column was
28
centrifuged for 1 minute. The eluate (approximately 48 µL) was stored at
40C. The product was checked by horizontal electrophoresis.
29
Results
The present investigation was carried out to formulate
suitable consortia of microorganisms capable of rapid degradation
of human waste (Night Soil) under aerobic environment. The
experiments were divided into three parts: First experiment deals
with isolation and screening of microbial cultures for production of
hydrolytic enzymes with Starch / Casein / Tributyrin / cellulose as
sole carbon source. Second part deals with quantitative estimation
of hydrolytic enzymes produced by each culture. The selection of
microbes for biodegradation was made on the basis of high
hydrolytic enzyme activities. Third part deals with the study of
genetic diversity of bacterial isolates by Random Amplified
Polymorphic DNA.
4.1 Isolation of Aerobic Hydrolytic bacterial strains:
The soil samples were collected from solid waste
disposal and market waste disposal site in Gwalior. The soil
samples were aerated continuously and enriched with night soil for
30 days. In all, fifty cultures were isolated based on their colony
characteristics and growth in plates containing mineral media and
Starch, casein, tributyrin, CMC mixture (1%) as sole source of
carbon.
Table 3: Hydrolytic Enzyme activity profiles of selected bacterial isolates
Expressed in means of clear zones
Sl No Isolates Amylase Protease Lipase Cellulase
1 V 1 +++ ++ ++ - 2 V 2 + ++ ++ - 3 V 3 + ++ ++ - 4 V 4 + ++ ++ - 5 V 5 + ++ +++ - 6 V 6 ++ ++ ++ - 7 V 7 + ++ - - 8 V 8 - ++ ++ - 9 V 9 + ++ + -
10 V 10 +++ ++ +++ - 11 V 11 +++ ++ ++ - 12 V 12 +++ ++ +++ - 13 V 13 +++ ++ ++ - 14 V 14 +++ ++ ++ - 15 V 15 +++ ++ +++ - 16 V 16 ++ ++ +++ - 17 V 17 +++ ++ ++ - 18 V 18 + +++ +++ + 19 V 19 +++ +++ +++ + 20 V 20 ++ +++ ++ + 21 V 21 ++ ++ + - 22 V 22 ++ ++ +++ - 23 V 23 + ++ +++ - 24 V 24 ++ +++ +++ - 25 V 25 ++ ++ ++ - 26 V 26 ++ +++ ++ - 27 V 27 + ++ ++ - 28 V 28 ++ ++ ++ - 29 V 29 + ++ ++ - 30 V 30 ++ +++ ++ - 31 V 31 ++ ++ - - 32 V 32 ++ ++ ++ 33 V 33 ++ +++ ++ - 34 V 34 ++ +++ ++ - 35 L 11 + +++ ++ - 36 P 1 ++ +++ ++ - 37 P 2 ++ +++ +++ - 38 P3 + +++ +++ - 39 P 4 ++ +++ ++ - 40 P 5 +++ ++ ++ - 41 P 6 +++ ++ ++ + 42 P 7 ++ +++ ++ - 43 P 8 ++ +++ ++ - 44 P 9 ++ +++ + + 45 P 10 ++ +++ +++ - 46 P 11 ++ +++ +++ - 47 P 12 +++ +++ ++ - 48 P 13 ++ +++ ++ - 49 P 14 +++ +++ ++ - 50 P 15 ++ +++ ++ -
-Zone of hydrolysis (diameter) after 2 days at room temperature:
+/– = <5 mm, + = 5-10 mm, ++ = 10-20 mm, +++ = >20 mm.
Qualitative Profiling of bacterial isolates for hydrolytic enzymes.
Cellulase Protease
Amylase Lipase
30
4.2 Qualitative screening of hydrolytic enzymes:
The measurements of halo zones of clearance by the
hydrolytic enzyme activities by the fifty bacterial isolates after 2
days of growth were shown in Table 3 and Figure 1.
All the fifty isolates were positive for proteolytic activity,
produced halo (minimum of 10 – 20 mm diameter) zones resulting
in the utilization of casein as the sole source of carbon. Twenty two
bacterial cultures were exhibiting halos greater than 20 mm were
considered here as best protease producers. Rest all the cultures
were exhibited the halo diameter of 10 – 20 mm after 2 days of
growth.
For amylase activity, all 49 bacterial cultures (98%) species
were positive in utilizing the starch as the sole source of carbon
making unstained halos while the capacity of its formation
depended upon the individual species. Thirteen cultures were
considered as best amylase producers with a capacity to produce
halos of 20 mm diameter after 2 days of growth. Twenty four
cultures were shown intermediate ability to produce amylase
enzyme. They produced unstained halos of 10 – 20 mm diameter
after 2 days of growth. Twelve cultures were exhibited weak
amylase activity producing unstained halo of 5 -10 mm diameter
after 2 days of growth, while culture V8 found negative for amylase
activity.
For lipolytic activity 48 (96 %) bacterial isolates were found
positive exhibiting halo zones around the colonies. Cultures V31
31
and V7 found negative for the lipolytic activity. Fourteen cultures
were best amylase producers exhibiting halo zone diameter of
more than 20 mm after 2 days of growth. Culture V 21 has shown
weak lipolytic activity, while rest 33 cultures were exhibited
intermediate extracellular lipase production after 2 days of growth
in the presence of Soybean meal.
Bacterial isolates were screened for cellulae activity, only 4
(8 %) cultures were marked halos indicating the utilization of
substrate CMC. V18, V19, V20, P6, and P9 were exhibiting weaker
cellulase activity in the presence of CMC, while rest of the cultures
were found negative for cellulase production. This result indicates
bacterial isolates are selective in cellulase production.
Bacterial isolates V7, V8 and V9 were showing weaker
extracellular hydrolytic enzymatic activity, while 11 isolates (V10,
V12, V15, V18, V19, V24, P2, P3, P10, P11, and P13) showed
best extracellular enzyme production. They produced all three
extracellular enzymes, rest of the isolates showed intermediate
production. Based on qualitative assay 32 cultures were selected
for further studies.
4.3 Morphological and physiological characteristics of
bacterial isolates:
The preliminary Morphological and physiological characteristics of
bacterial isolates were examined for 50 isolates in terms of their
cell morphology, colony morphology, pigmentation, motility and
growth characteristic in nutrient broth media. There was large
Table 1: Morphological characters of bacterial isolates in broth
S. NO. STRAIN TURBIDITY VEIL SEDIMENTATION MOTILITY
1 P1 + - + +
2 P2 Less - - +
3 P4 + - + +
4 P5 + - - +
5 P7 + - + +
6 P8 + - + +
7 P13 + - + +
8 P14 + + + +
9 P15 + - + +
10 V1 + - + +
11 V10 - - + +
12 V11 - + + +
13 V12 + - + +
14 V13 + - - +
15 V14 + - - +
16 V15 + - - +
17 V16 + - + -
18 V17 + - - +
19 V18 + - - +
20 V19 - + + -
21 V20 + - + +
22 V21 + - + +
23 V22 + - + +
24 V23 + - + +
25 V24 + - + +
26 V25 + - + +
27 V26 - - + +
28 V27 + - + +
29 V30 + - + +
30 V31 + - + +
31 V33 + - + +
32 V34 + - + +
33 L11 + - - +
Table 2: Morphological Characterization of isolated bacterial colonies.
S. NO. STRAIN PIGMENT CONFIGURATION MARGIN ENLEVATIOS
1 P2 Milky white Round Smooth Concavex
2 P4 Milky white Wrinkled Smooth Flat
3 P5 Creamy white concentric Smooth Concavex
4 P7 Creamy white concentric Smooth Concavex
5 P8 Creamy white concentric Smooth Concavex
6 P13 Whitish grey concentric Smooth Flat
7 P14 Milky white Round Smooth Flat
8 P15 Creamy white Contractile Smooth Flat
9 V1 Creamy white concentric Smooth Concavex
10 V10 Creamy white concentric Smooth Concavex
11 V11 Creamy white Wrinkled Smooth Flat
12 V12 Milky white Irregular & spreading Lobate Concavex
13 V13 Creamy white Wrinkled Smooth Flat
14 V14 Milky white Wrinkled Wavy Concavex
15 V15 Creamy white Round Smooth Concavex
16 V16 Creamy white Round Smooth Concavex
17 V17 Creamy white Wrinkled Smooth Flat
18 V18 Milky white Irregular & spreading Lobate Flat
19 V19 Milky white Irregular & spreading Lobate Concavex
20 V20 Milky white Wrinkled Wavy Concavex
21 V21 Milky white concentric Smooth Concavex
22 V22 Creamy white Wrinkled Smooth Flat
23 V23 Creamy white Round Wavy Concavex
24 V24 Creamy white Wrinkled Smooth Concavex
25 V25 Creamy white Round Smooth Concavex
26 V26 Creamy white Round Smooth Flat
27 V27 Creamy white Round& concentric Smooth Flat
28 V30 Whitish grey concentric Lobate Concavex
29 V31 Milky white Wrinkled Wavy Flat
30 V33 Milky white Wrinkled Smooth Concavex
31 V34 Creamy white Round Smooth Flat
32 L11 Pink Round& concentric Wavy Concavex
32
variation in colony morphology – colour, shape, and motility.
Morphological features of the isolated bacterial strains were shown
in Table 1. Growth characteristics in broth were shown in Table 2.
Most of the isolates produced turbidity when grown in
nutrient broth, except V10, V11, V19 and V26. Bacterial isolate P2
has shown less turbidity, it exhibited thread like growth in broth.
Among 32 isolates studied, only 3 cultures (P14, V11 and
V19) were showed veil formation in broth condition. Bacterial
isolates P2, V13, V14, V15, V17, V18 and L11 showed
sedimentation in broth condition, while rest of the cultures did not
show sedimentation.
Isolate V16 did not show motility, while rest all isolates
showed motility after 24 hours of growth when spot inoculated on
the centre of the nutrient agar plates (0.2 % Agar).
Most of the bacterial isolates were white in colour, while
isolate L11 exhibited pink colour. All the bacterial isolates were
looking different with respect to its colony morphology (elevation,
margin, and configuration).
4.4 Quantitative screening of bacterial isolates for hydrolytic
enzyme production:
Extra cellular enzyme production potential of bacterial
isolates was determined quantitatively for amylase, protease and
lipase. Most of the bacterial isolates were selective in cellulase
production, so cellulase estimation was not carry out quantitatively.
The data were shown in Table 4 and figure 2.
Table 4: Quantitative enzyme profiles of isolated bacterial cultures.
Sl No Isolates Amylase Protease Lipase
1 V1 274.4 238.7 25.4 2 V2 63.6 241.4 55.3 3 V3 13.4 262.3 32.6 4 V4 36.8 246.6 55.9 5 V5 36.8 204.6 101.0 6 V6 120.5 228.2 61.7 7 V7 13.4 186.3 5.8 8 V8 1.7 306.9 40.3 9 V9 5.0 304.3 22.7 10 V10 289.4 251.9 158.0 11 V11 2111.0 304.3 40.3 12 V12 371.4 133.8 130.5 13 V13 378.1 170.5 61.7 14 V14 297.8 196.8 62.0 15 V15 247.6 238.7 125.8 16 V16 180.7 262.3 127.8 17 V17 209.1 251.9 51.9 18 V18 155.6 2251.9 29.7 19 V19 123.8 1909.6 34.9 20 V20 250.9 1994.8 33.6 21 V21 263.5 2011.9 24.8 22 V22 200.8 1622.3 35.4 23 V23 391.4 1943.7 123.2 24 V24 446.6 2474.8 93.9 25 V25 445.8 1732.5 13.1 26 V26 417.4 2196.8 144.4 27 V27 509.4 1998.7 30.9 28 V28 184.0 1979.1 69.2 29 V29 168.1 2476.1 87.1 30 V30 180.7 2224.3 93.4 31 V31 184.0 1506.9 0.7 32 V32 191.6 223.0 34.9 33 V33 271.8 2608.6 34.4 34 V34 214.1 2473.5 32.9 35 L11 1165.1 295.1 156.6 36 P1 208.3 1812.5 35.6 37 P2 169.0 1777.1 15.8 38 P3 169.8 1723.3 0.7 39 P4 164.0 1927.9 136.3 40 P5 147.2 1962.0 101.4 41 P6 58.6 1568.6 77.5 42 P7 145.6 1842.7 101.0 43 P8 114.6 2149.6 90.0 44 P9 215.0 1947.6 7.5 45 P10 77.0 1896.4 51.5 46 P11 363.0 2682.0 0.7 47 P12 503.5 1563.3 4.8 48 P13 356.3 1969.9 36.6 49 P14 373.0 2494.5 98.3 50 P15 519.4 2233.5 38.0
Figure 2: Quantitative
activities expressed in terms of IU / mg of
Quantitative Extracellular Hydrolytic Enzyme
activities expressed in terms of IU / mg of protein
M P12 V34 V11 V27 P15 V23 L11 P4 P1 V21 V33 P5 V31 P11 V17 V24 P13 P7 M
M V10 P8 V30 P14 V15 V18 V12 V19 V14 P2 V1 M
Figure 3: RAPD profile of PCR amplification with primer OPA 10
M V21 P12 V33 V11 P13 P15 V30 V31 V23 V16 V18 P5 P4 P1 P14 V34 P2 V10 M
M P8 V14 V27 V19 P11 V20 V17 V24 P7 V1 V12 L11 V15 NC M
Figure 4: RAPD profile of PCR amplification with primer OPA 13
Figure 3: RAPD profile of PCR amplification with primer OPA 10
M
P7
P
8
V1
4P
15
V
30
V
1
V3
3
V1
2
P1
L
11
P11
P
2
V1
0
V11
P
4
P1
3
V2
3
V2
0
M
M
V2
7
V1
5
V3
4
P5
V
21
P
14
V
31
V
17
P
12
V
18
V
24
V
19
V
16
M
1500 bp
Figure 5: RAPD profile of PCR amplification with primer OPA 16
Figure 9:16 S rDNA PCR amplification with universal primer
M
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
M
M – Marker1-20- Bacterial Isolates
33
Among the bacterial isolates V11 produced highest amount
of amylase (2111 IU / ml). Amylase production ranged between 1.7
– 2111 IU / ml. Isolates V23, V24, V26, V27, L11, P11, P12, and
P15 were found to be better producers of amylase enzyme.
Among the bacterial isolates P11 produced highest amount
of protease (2682 IU / ml). Protease production ranged between
133.8 – 2682 IU / ml. Isolates V18, V21, V24, V26, V29, V30, V33,
V34, P8, P11, and P14 were found to be better producers of
amylase enzyme. Protease production was found more when
compared to other extra cellular enzymes.
Among the bacterial isolates L11 produced highest amount
of amylase (156.6 IU / ml). Amylase production ranged between
0.7 – 156.6 IU / ml. Isolates V5, V10, V12, V16, V23, V26, L11, P4,
P5 and P7 were found to be better producers of amylase enzyme.
Bacterial isolate V26 was found to be the better isolate with
respect to all extra cellular enzyme production. Other isolates like
V23, V24, V27, L11, P11, P12 and P15 were also found better
isolates with respect to all extra cellular enzyme production.
4.5 Extraction of genomic DNA from bacterial isolates:
The extracted Genomic DNA was of a good quality as being
checked by agarose gel electrophoresis. The extracted DNA was
intact, clean and free of RNA (data not shown)
4.6 Molecular typing of bacterial isolates:
Total genomic DNA from thirty two colony variants were used as
templates for the RAPD reaction. DNA solutions with an
34
absorbance ratio at 260/280 nm of above 1.8 were used, because
it is important to use DNA of high purity in order to generate RAPD
profiles of high quality, as recommended by the manufacturer.
Six primers, RAPD Analysis Primers OPA 7, OPA 10, OPA
1 3, OPA 16, OPA 17, OPA 20 were examined for their ability to
produce potentially useful fragments for discrimination (data not
shown).
Primers OPA 10, OPA 13, and OPA 20 were used for
further studies, because they gave fingerprints that were
uncomplicated, yet had a sufficient number of fragments to be
discriminatory. The amplified products by RAPD reaction were
electrophoresed on agarose gels. The RAPD profiles obtained are
shown in Figures. 3, 4 and 5.
Apart from the colony variation, each strain showed an
individual RAPD profile. Identical profiles were shown respectively
by V1, V10, V12, V14, V16, V18, and V19. Isolates P8 and V23
were produced same profile when OPA 10 and OPA 13 primers
were used. Isolates V30 and P15 were produced same profile
when OPA 16 was used.
A total of 32 bacterial isolates were identified genetically
by the analysis of their RAPD fingerprints using three arbitrary
primers. The relatedness between bacterial isolates was evaluated
by comparison of their RAPD patterns generated by each primer.
The RAPD-fingerprints of the various strains differed in fragments
number, size and intensity depending on which primer being used
Figure 6: Dendogram obtained from the analysis of OPA-10
RAPD patterns. The scale indicates the similarity level.
Figure 7: Dendogram obtained from the analysis of OPA-13
RAPD patterns. The scale indicates the similarity level.
Figure 8: Dendogram obtained from the analysis of OPA-16
RAPD patterns. The scale indicates the similarity level.
35
in each run. Treecon dendrograms were derived from the
fingerprints generated by each primer.
Each RAPD experiment was repeated three times with
each primer and similar results were obtained. Based on that,
typing scheme as reported. The number of polymorphic bands
generated for each isolate was between 1 and 17 with a size
ranged from 250 bp to 2500 bp. The dendrograms presented in
Figures 6, 7 and 8 revealed that among the tested isolates V1,
V10, V12, V14, V16, V18, V19 were found to be similar
producing same profile. Strains P8 and V23 were found to be
similar. Both the primers produced the same results and
confirmed that all the isolates are similar and the difference in
their morphology is due to environment. .
4.7 16S r DNA Amplification of bacterial isolates:
The small subunit rRNA gene was amplified using the two
primers 16S1 & 16S2 corresponding to the nucleotide positions 9-
27 and 1477-1498, respectively, on E. coli 16S rRNA gene
nucleotide position. The purified DNA product, approximately 1.5
Kb in length, was run on agarose gel to check the quality of the
PCR product (Figure 9, 10). The products will be sequenced and
isolate will be identified.
36
Discussion
Ecological problems of health importance are extended
every year with the accumulation of different organic waste.
Biological factor is the one that could accomplish their
degradation without any implications. Bacteria are the group of
microorganisms with the greatest role in the biopurifying
processes. Their function is determined by the wide catabolic
potential and adapting abilities to assimilate different substrates
(Powlowski and shinger, 1994).
Aerobic, unlike anaerobic digestion, does not produce the
pungent gases. The aerobic process results in a more complete
digestion of waste solids reducing build up by more than 50% in
most cases. The aerobic process also improves the environment
of the workers and the animals and helps to keep pathogens in
check.
Aerobic biodegradation is the breakdown of organic
contaminants by microorganisms when oxygen is present. More
specifically, it refers to living only in the presence of oxygen;
therefore, the chemistry of the system, environment, or
organism is characterized by oxidative conditions. Many organic
contaminants are rapidly degraded under aerobic conditions by
aerobic bacteria called aerobes.
37
Microbes represent a good source of enzymes due to a
number of characteristics like their broad biochemical activity,
their rapid growth, the limited space required for cell cultivation
and the ease with which the enzymes can be genetically
manipulated to generate new enzymes for various applications
(Rao et al., 1998).
Four extracellular hydrolytic enzymes from fifty bacterial
isolate have been screened and identified by substrate
depended agar plate technique. The ability to produce number
of enzymes depended upon the isolates screened. The present
findings will form a base work on hydrolases of bacterial isolates
and their applications as biotechnological tools in the production
of services and products for human welfare.
This study contributes to catalogue of aerobic bacterial
isolates and provides additional information to support future
research about the industrial potential of these microorganisms.
Synergisms are expected from interchange of experiences
between crystallographers, biochemists, geneticists, and
enzyme kineticists, and food, chemical, and biochemical
engineers. Extensive and persistent screening for new
microorganisms and their hydrolytic enzymes will open new,
simple routes for synthetic processes and consequently, new
ways to solve environmental problems.
For a consortium of microorganisms to be used for rapid
degradation of night soil, it should be compatible with each other,
38
should have the ability to produce multiple hydrolytic enzymes in
addition to its ability to utilise and grow the mixed organic
sources.
The taxonomical determination of bacterial, exposed to
continuous pressure as a result of polluted environment, faces
difficulties using the classical methods. The adaptive changes of
these microorganisms give an advantage in evolutional aspect,
but differ them from the collections strains, cultivated at optimal
conditions. It is necessary to apply a number of modern
molecular genetic methods for determination that will be a future
task.
The results verified that RAPD is a powerful tool for
discrimination and can identify bacteria to the strain level, and
suggested that colony morphology and colour are not important
defining characteristics of strains. RAPD can verify the identity
of two strains genomically. Furthermore, RAPD is faster and
easier than the conventional methods. After preparation of DNA,
the RAPD profile can be obtained in one day. Our studies have
shown that RAPD fingerprinting is a valuable tool for molecular
profiling of bacterial cultures.
39
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i
APPENDIX – I
Culture Media:
Protease medium (components g / l ) KH2PO4 : 1.0
K2HPO4 : 1.0
(NH4)2SO4 : 2.0
MgSO4 : 0.8
CaCl2 : 0.1
Yeast Extract : 2.0
Sodium Citrate : 1.0
Casein : 4.0
Distilled Water : 1000 ml
pH : 7.4 ± 0.1
Amylase medium (components g / l )
KH2PO4 : 1.5
K2HPO4 : 1.0
(NH4)2PO4 : 2.0
MgSO4 : 0.8
CaCl2 : 0.1
Yeast Extract : 2.0
Sodium Citrate : 1.0
Starch Soluble : 4.0
Distilled Water : 1000 ml
pH : 7.4 ± 0.1
Lipase medium (components g / l)
Soyabean Meal : 2.0
Peptone : 1.0
Starch Soluble : 1.0
K2HPO4 : 0.2
MgSO4.7H2O : 0.1
CaCO3 : 0.5
Distilled Water : 1000 ml
pH : 7.4 ± 0.1
ii
Cellulase medium ( components g / l )
NaNO3 : 2.0
MgSO4 : 0.5
K2HPO4 : 0.05
FeSO4 : 0.01
CaCl2 : 0.02
MnSO4 : 0.02
CMC(Carboxy methyl cellulose) : 1.0
Distilled Water : 1000 ml
pH : 7.2 ± 0.1
Tryptone Yeast Extract Broth ( components g / l )
Tryptone : 1.0 g
Yeast Extract : 5.0 g
NaCl : 5.0 g
Glucose : 1.0 g
Distilled Water : 1000 mL
pH : 7.0 ± 0.1
iii
APPENDIX – II
Reagents:
Tris-HCl Stock Solution [1.0 M] (pH 7.5):Dissolve 12.1 g of Tris
(Hydroxymethyl aminomethane)-HCl in 75 mL of sterile double distilled
water. Add concentrated HCl dropwise till pH 7.5 is attained. Make up
the volume to 100 mL with double distilled water.
EDTA Stock solution (0.5M):Add 18.612 g of EDTA disodium salt in 40
mL of sterile double distilled water in a presterilized reagent bottle. Place
the bottle on a magnetic stirrer for uniform shaking and slowly add 40%
NaOH solution through the wall of reagent bottle till pH 8.0 is attained.
Now make the final volume up to 100 mL with sterile double distilled
water.
Sodium Dodecyl Sulphate solution (25%):Add 25 g Sodium dodecyl
sulphate (Sodium lauryl sulphate) in 75 mL of sterile double distilled
water in a clean, presterilized reagent bottle and dissolve by stirring
slowly avoiding formation of froth make the volume up to 100 mL.
Lysozyme stock solution (20 mg mL-1
): Dissolve 0.2 g of Lysozyme
powder in 10 mL of sterile milliq water and store at -200C for future use.
Ammonium acetate solution (7.5 M): Add 57.81 g of ammonium
acetate in 50 mL sterile double distilled water in a presterilized reagent
bottle, stir till homogeneous, transparent solution is obtained and make
the final volume up to 100 mL.
iv
Buffers:
Tris-EDTA buffer (T10
E25
) :For preparing 100 mL of T10
E25
buffer, add 2
mL of Tris-HCl stock and 5 mL of 0.5 mM EDTA stock and make the
volume up to 100 mL.
Tris-EDTA buffer (T10
E1):For preparing 100 mL of T
10E
1 buffer, add 2
mL of Tris-HCl stock and 0.2 mL of 0.5 mM EDTA stock and make the
volume up to 100 mL.
v
APPENDIX III
Quantification through Agarose Gel Electrophoresis:
Tris Borate EDTA (TBE) Buffer (5X Stock): Dissolve 54 g of Tris-HCl,
27.5 g of Boric acid and 20 mL of 0.5 M of EDTA (pH 8.0) to 500 mL of
sterile MilliQ water and make the final volume upto one liter sterile MilliQ
water.
TBE Buffer (0.5X working solution):Take 100 mL of 5X stock of TBE
buffer in 500 mL of sterile MilliQ water and make the final volume upto
one Liter with sterile MilliQ water.
Tracking dye (6X): Prepare 0.25% Bromophenol blue in 40% sucrose
(w/v).
Ethidium bromide stock: A stock solution of EtBr containing 5 mg mL-1
was prepared in sterile milliq water.
DNA molecular weight markers:
DNA ladders of 1 kb and 100 bp (Banglore Genei Pvt. Ltd.) were
used as molecular weight markers for visualising PCR amplification
product and RFLPs respectively.
Agarose (0.8%):
Add 0.8 g of Agarose to 100 mL of 0.5X TBE buffer in a 250 mL sterile
Erlenmeyer flask and dissolve to homogeneity by melting in a microwave
oven. When the temperature of the solution is about 50-55 0
C add 3 µL
of Ethidium Bromide (5 mg/mL) and pour into the gel casting tray with
combs in place. Allow the agarose to polymerize for about an hour.
vi
APPENDIX IV
PCR-Amplification of 16 S rRNA gene from the bacterial isolates
Solutions and buffer for polymerase chain reaction (PCR):
a) Polymerase reaction buffer (10X):
The polymerase reaction buffer commercially provided contains 100 mM
Tris. HCl (pH 9.0); 500 mM KCl, 15 mM MgCl2
and 0.1% gelatin.
b) Taq DNA polymerase :
Taq DNA polymerase was 5U µL-1
in a storage buffer consisting of 20
mM Tris-HCl (pH 8.0), 100 mM KCl, 0.1 mM EDTA, 1 mM DTT, 0.5%
Tween 20 (v/v), 0.5% Igepal and 50% Glycerol (v/v).
c) dNTP's mix: A stock of dNTP mix (dATP, dCTP, dGTP and DTTP)
has 2.5 mM concentration of each dNTP.
d) Primers:
The forward (PA) and reverse (PH) primers were custom synthesized
from Sigma, USA. The sequence of the oligonucleotide primers used for
amplification of 16S rDNA genes are:
16S1: 5' GAGTTTGATCCTGGTCA 3'
16S2: 5' ACGGCTACCTTGTTACGACTT 3'
The stock solution (100 ngmL-1
) of the primers was prepared by
reconstituting lyophilized primers in sterilized milli Q water and stored at
-200C.