Sumit Thesis

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



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


14 V14 Milky white Wrinkled Wavy Concavex

15 V15 Creamy white Round Smooth Concavex



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


23 V23 Creamy white Round Wavy Concavex

24 V24 Creamy white Wrinkled 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



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 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.





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


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


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


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:


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.


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.

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