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CHAPTER – I Isolation and Characterization of Bacteria and Fungi from Soils and

Composts

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GENERAL INTRODUCTION

Man’s use of land has aggravated the loss of soil organic carbon from

cultivated soils. The practices and conditions that favor higher evolution of carbon

dioxide oppose maintenance of organic carbon also known as carbon sequestration in

soils and vice versa (Heenan et al., 1995, Probert et al., 2001). Depletion in soil

organic carbon is further accentuated by the activities of people, livestock,

deforestation, overgrazing, burning crop residues and vegetation. Disuse of organic

manure, removal of crop residues, and monoculture without cover crops, fallow

vegetation leading to low productivity. Several studies have concluded that low soil

fertility is a major constraint for production of food grains and natural vegetation.

Although yield increases can be achieved via application of chemical fertilizers; they

alone cannot sustain crop yields in the long run due to high costs and resultant

decreases in soil fertility and quality. Improving soil organic matter and soil minerals

are vital in maintaining soil quality and agricultural productivity. Management

strategies for improving soil organic matter and crop productivity include the use of

N-fixing cover crops, application of crop residues such as surface mulch, composts

and manures, and application of biofertilizers or microbial inoculants.

The motivation for shifting from chemically intensive management to

alternative practices include: i) concern for protecting soil, human and animal health

from the potential hazard of chemical fertilizers and pesticides, ii) concern for

protecting environmental and soil resources, iii) to lower the cost of agricultural

practices. Organic farming systems rely on legume-cropping containing crop

rotations, crop and animal residues, microbial inoculants for cultivation to maintain

soil nutrient supply and control weeds and pests. Plant growth promoting and

biocontrol agents such as Bacillus, Enterobacter, Pseudomonas and Streptomyces

sps., have been identified in compost amended substrates (Nelson, 1992). The diverse

microbial populations present produce plant growth hormones and stimulate plant

growth directly; others produce natural chelators called siderophores that chelate iron.

Beneficial plant microbe interactions in the soil are the determinants of plant health

and soil fertility (Jeffries, 2003).

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Soil plant microbe interactions are complex and there are many ways the

outcome can influence plant health and productivity. These interactions may be

detrimental, beneficial, or neutral to the plants. However, the focus of this thesis is to

exploit the beneficial bacteria to enhance plant growth. These are known as plant

growth promoting bacteria (PGPB), and represent a wide variety of soil bacteria

which, when grown in association with a host plant, result in stimulation of plant

growth. Direct growth promotion is by nitrogen fixation, solubilization of mineral

phosphates, production of plant growth hormones like IAA, Ethylene, Gibberellins

(De Freitas et al., 1997; Rodriguez and Fraga, 1999) and ability to produce 1 – amino

– cyclopropane – 1 – carboxylate (ACC) deaminase. Indirect mechanisms include

antagonism against plant pathogenic fungi by producing siderophores, chitinases,

beta- 1, 3 glucanases and/or antibiotics.

The plant response to PGPB (Plant growth promoting bacteria) is obviously a

very complex phenomenon resulting from a combination of mechanisms, which

affect several aspects of mineral nutrition, root development and colonization of

potential bacteria (Chebotar, 2001; Hontzeas, 2004). Interactions between plant

growth promoting and phosphate solubilizing bacteria with rhizobia may be exploited

to enhance biological nitrogen fixation and crop yield (Cavender et al., 2003).

Pigeon pea (Cajanus cajan) is an important food crop, a principal source of

protein in the Indian diet, and a very popular food in developing tropical countries.

India accounts for 90% of world’s pigeon pea production, where it is being cultivated

as a sole crop or intercrop. Legumes contribute to increased productivity of other

crops when incorporated into cropping systems as intercrops. Nevertheless, in cereal-

legume intercropping systems, the recommended amount of chemical fertilizer for the

main crop is still being applied under the assumption that the legume component can

fulfill its own requirement. However, supplying the recommended dose of

biofertilizer to both crops could increase the yield of the intercrop.

Extensive use of fungicides to manage soil borne plant pathogens has disturbed

the ecological balance of soils, leading to groundwater contamination and increased

health risks to humans. Biocontrol agents, especially of the genera Pseudomonas and

Bacillus may be an ecologically sound alternative to chemical pesticides to inhibit

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phytopathogens (Punja, 1985). Fusarium udum is a soil borne plant pathogenic

fungus that causes root rot, blights, wilts and damping off in pigeon pea crops

(Manjula, 2005). Macrophomina phaseolina is one of the most destructive, seed and

soil borne plant pathogen, causing charcoal rot, dry-root, wilt, leaf blight, stem blight

and damping off diseases in a wide range of host plants (Khara, 2008). Recently,

there has been a growing interest in combining biocontrol agents with other chemical

components to enhance their activity against certain phytopathogens (Singh, 2008).

Sustainable agriculture involves the successful management of agricultural

resources to satisfy human needs, while maintaining or enhancing the quality of the

environment and conserving natural resources. Application of low-cost inputs can be

made efficient by value-addition using the scientific knowledge to improve the crop

productivity. Soil quality can serve as an indicator of ongoing conservation and

degradation process (Parr et al 1992; Halvarson et al., 1996). It depends highly on the

nutrient content, biological and microbiological component of the soil ecosystem and

influences crop yield and quality. Soil microorganisms are potentially one of the most

sensitive biological markers available and should, therefore, be useful for clarification

of disturbed or contaminated systems (Powlson, 1987).

The present work focuses on the isolation of bacteria with PGPR and

phytopathogen protective traits for promotion of pigeon pea plant growth. The present

study is divided into five chapters. Chapter I focuses on isolation, enumeration and

characterization of bacteria and fungi from a) composts, e.g. farm yard manure, rice

straw compost, gliricidia vermicompost and b) e.g. soils rhizosphere soil, rhizoplane

soil, and cultivable field soil. Chapter II presents qualitative and quantitative

screening of PGPR traits of the selected bacteria, e.g. phosphate solubilization,

hormone production (IAA), and 1-Amino- cyclopropane-1-carboxylate (ACC)

deaminase production. Chapter III focuses on the qualitative antagonistic abilities of

the isolated bacteria e.g. HCN production, siderophore production and chitinase

production; and the effect of these bacteria on fungal phytopathogen growth. Chapter

IV focuses on evaluation of pigeon pea growth using these potential bacterial isolates

as amendments to soil. Chapter V presents the molecular identification and

phylogenetic relationships of the isolated bacteria using 16S rRNA sequencing

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OBJECTIVES IN GENERAL The objectives of the present study include

• Isolation of bacteria and fungi from the soils and compost samples by Nutrient Agar

Plate culturing and purification of the isolated bacterial cultures by Streak plate

method.

• To identify the selected bacterial isolates using morphological and biochemical

methods via Bergey’s Manual of Systematic Bacteriology.

• To screen for PGPR traits present among the isolated sixteen bacterial samples, e.g.

phosphate solubilization, indole acetic acid production, ACC deaminase production

by both qualitative and quantitative analysis.

• To screen antagonistic traits of the bacteria among the sixteen isolates e.g. HCN,

siderophore and chitinase productions.

• To study the antagonistic behaviour exhibited by the bacterial isolates towards the

fungal phytopathogens Macrophomina phaseolina and Fusarium udum in in vitro

techniques.

• To evaluate the germination % and seed vigour index of Cajanus cajan using the

potential bacterial isolates.

• To evaluate plant growth parameters of Cajanus cajan such as. root length, shoot

length, Plant biomass with the bacterial bioinoculants in both sterilized and

unsterilized conditions for a period of over 30 days.

• To study the levels of plant susceptibility and disease resistance exhibited by Cajanus

cajan towards phytopathogens upon amending the soils with the bacterial inoculants.

• To study the interaction of the bacterial isolates with the nodulating bacteria like

Sinorhizobium, Rhizobium and Bradyrhizobium sps.

• Molecular identification of the potential bacterial isolates and construction of

phylogenetic dendrograms.

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CHAPTER 1

ISOLATION, ENUMERATION AND CHARACTERIZATION OF

BACTERIA AND FUNGI FROM SOILS AND COMPOSTS

INTRODUCTION

Soil is an ecosystem that plays a key role in the availability of plant nutrients

and contains a diverse community of organisms differentiated by morphology and

physiology. A scientifically managed system of soil microbial plant association is

useful in reducing fertilizer requirement of field crops while sustaining the soil

productivity. Being ubiquitous in nature with extensive host range and plant growth

benefits, microbial inoculants can enhance the growth and yield of crops.

A soil aggregate is a naturally occurring cluster of bound soil particles. Soil

organic matter including humus, polysaccharides and polyuronides produced by soil

microorganisms help to cement soil particles together, while filamentous fungi

provide additional mechanical support. Soil fertility depends not only on chemical

composition but also on the qualitative and quantitative nature of microorganisms

inhabiting it. The distribution of organisms in soil are closely linked with the

occurrence of organic matter. Alexander (1977) noted that bacteria in soil are

generally present in the film of colloidal material coating the mineral particles. Some

of these bacteria show positive effects on plant growth and are termed plant growth

promoting rhizobacteria (PGPR), whereas others show deleterious effects and are

called deleterious microorganisms (DRMOs).

Soil microorganisms can be isolated and grown on artificial media. Different

media encourage the growth of different types of microorganisms through the use of

inhibitors and specialized growth substrates. Dilution plate technique is a useful tool

to study the relative abundance of soil bacterial types and changes in population

density (Dubey and Maheshwari, 2004). This technique is based on the principle that

complete detachment and dispersion of cells from the soil will give rise to discrete

colonies when incubated on a petri plate containing nutrient media. The assumptions

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that underlie this technique are 1) complete dispersion of sample, 2) suitable growth

media for the organisms and 3) no interaction between organisms on the media.

Soil bacteria and fungi mediate some important processes such as

decomposition, nutrient mobilization and mineralization, nitrogen fixation and

denitrification etc. The differences of the composition of soils together with

differences in their physical characteristics and agricultural practices result in

corresponding large differences in the microbial populations both in total numbers

and kind. The soil conditions vary with different types of nutrients, available moisture

content and degree of aeration, temperature, and pH. The root system of higher plants

also influences the numbers and the kind of microorganisms present.

Pure cultures of bacteria are required to study colony morphology,

microscopic characteristics and biochemical characteristics. Genera and species

identification of bacterial pure cultures can be performed based on the variable

characteristics that are exhibited by different bacteria.

The objective of the present study is to determine the number of bacteria

and fungi present in various composts and soils to characterize them based on the

morphological and biochemical parameters.

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REVIEW OF LITERATURE

Bacteria are the most abundant organisms in soil. They are free living and are

a critical component of soil microbial populations. They have been extensively

studied because of their involvement in maintaining nitrogen and carbon cycles, and

due to their vital transformations of the soil (Kloepper, et al., 1980). The soil bacteria

include proteobacteria a wide variety of pathogens such as Escherichia. Salmonella,

Vibrio, and many other notable genera those are free-living, and responsible for

nitrogen fixation (Vincent, 1970; Woese, 1992). Bacteria form loose associations in

soils and with plants living near, on or even inside roots (Larcher et al., 2003). The

availability of C, N, and P the soil is of paramount significance for bacterial growth

(Kwok et al., 1987, Boehm et al., 1993). Additional input either in the form of litter

or from chemical substances can rapidly enhance growth of bacterial populations in

the soil for varying periods, (Chebotar and Asis, 2001; Sundara et al., 2002). The

nutrient supply is exhausted or reduced; bacterial populations drop (Hontzeas et al.,

2004).

Plant derived nutrients and growth factors, attractants or even inducers of

enzymes can aid bacterial colonization of soils (Chen et al., 1998). In return, these

bacteria exhibit properties favoring plant growth and productivity; those that do are

termed plant growth promoting rhizobacteria PGPR (Subba Rao, 1999). Incorporation

of organic components such as composts, farming practices of soil and soil

amendments, can dramatically affect soil microbial activity, soil microbial diversity,

soil microbial biomass, soil respiration and soil fertility (Elliott and Lynch, 1994).

Furthermore, bacteria improving the physicochemical characteristics of soils

(Grayston et al., 2004).

Composts and Soils

Compost is prepared by the biological degradation of plant and animal residues

under controlled aerobic conditions (Eghball et al., 1997). The management of soil

organic matter requires inputs of organic manures, crop residues, green manures and

other organic wastes (Beri et al., 2003; Manici et al., 2004; Artursson et al., 2006).

Composting is one method of utilization of these organic wastes by microbes to

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produce manure rich in plant nutrients (Aira et al., 2002; Jeffries et al., 2003; Hussen

et al., 2003). Composts are known to be products rich in microorganisms that help

plants to mobilize and acquire nutrients (Postma et al., 2003). Composts have the

potential for plant growth when added to soil as was demonstrated by Atiyeh et al.,

(2000) in marigold plants.

Rice straw compost (RSC) enhanced enzyme activities and C, N content when

added to soil (Crecchio et al., 2001). In tropical soils, application of Farm Yard

Manure (FYM) or organic amendments stimulates proliferation of bacteria and fungi

(Harinikumar and Bagyaraj, 1989). Degraded products of FYM had a residual

stimulatory effect on growth and proliferation of nitrogen fixing bacteria (Saha et al.,

1995). Composts and soil amendments enhance total fungal and actinomycetes

counts (Kim, 1998a). The work on Gliricidia Vermicomposts (GVC) in maintaining

the soil fertility and improved microbial proliferation was emphasized by many

workers (Pandey et al., 1998; Aira et al., 2002; Arancon, 2004).

The rhizosphere region and rhizoplane provides better sites for the isolation of

bacteria than the bulk soil (Curl et al., 1986). Several studies indicated that structural

and functional diversity of rhizosphere populations is affected by plant species, root

exudations and rhizodepositions (Martin and Loper, 1999; Kent et al., 2002). Soil

types, growth stage of plant, cropping practices, and environmental factors also

influence the composition of microbial community in rhizosphere (Grayston et al.,

2004). Bacterial diversity of wheat rhizosphere was more diverse than chickpea

rhizosphere (Sarita et al., 2006).

Physicochemical properties

The physicochemical characteristics of soil can select for a succession of

microbial communities and thus can have profound effects on entire process. For

example, in composts physicochemical properties such as temperature, carbon

dioxide content, and C: N ratio of the substrates can vary greatly. A significant

change in chemical content (organic C% and N %) is observed during cultivation and

application of composts and fertilizers (Dick, 1992). The chemical indicators of soil

are organic carbon, pH, electrical conductivity, cation exchange capacity (CEC),

nitrogen, and phosphorus and potassium contents. An improvement in the buffering

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capacity of soil and increased organic carbon, electrical conductivity and available

nitrogen, phosphorus and potassium were observed with the addition of organic

amendments in the soil in the form of FYM, compost and plant residues (Nambiar et

al., 1989., Clement et al., 1998). The physicochemical properties of soils and

composts provide different growth conditions for microbes and for different microbial

communities (Elliott and Lynch, 1994).

Isolation and Enumeration of Bacteria and Fungi

Two longstanding challenges in soil microbiology are the development of

effective methods to 1) determine which microorganisms are present in soil and 2)

determine microbial function in situ (Hall et al., 2003). The microbial population of

soils is made up of five major groups including bacteria, actinomycetes, fungi, algae

and protozoa and among these groups bacteria are the most abundant group

(Alexander 1977). Microbial communities particularly bacteria and fungi constitute

an essential component of the biological characteristics in soil ecosystems (Kent and

Triplett, 2002).

Bacterial populations in different soil types are highly variable, both in terms

of identity and spatial distribution and are dependent upon soil amendments. Taylor et

al., (1990) used plate count method and bacterial biomass methods for direct counting

of bacteria, and most earlier studies on bacterial soil communities have been

conducted using cultivation based methods e.g. Grayston et al., (2004). Typically the

microorganisms are grown on select culture media depending on the physiological

suitability of particular growth substrates, however it is estimated that by using this

culture method only 1% of the species are isolated, thus presenting a skewed picture

of microbial diversity in soil (Weller et al., 1994). It is clear that the soil in general is

quite rich in bacteria and that many bacteria are slow or difficult to culture (Martin

and Loper, 1999). A number of factors likely influence bacterial proliferation, e.g.

moisture, aeration and temperature (Schomberg et al., 1994; Subler et al., 1998). To

decrease the number of bacterial counts to feasible levels, serial dilutions are

performed and plated through drop plate method (Somasegaran et al., 1994). The

numbers of organisms recovered after growth on a specific media are referred to as

colony forming units (CFUs) (Kogure et al., 1979).

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Soil fungal populations are largely dependent on availability of organic

matter (Girvan et al., 2004). Scientific knowledge concerning soil fungal community

is scarce when compared to bacteria (Subba Rao, 1982; Anderson and Cairney,

2004). The size, shape and color of conidia or spores of fungal populations vary

under different physiological culture conditions (Ellis, 1993). Artificial and natural

substrates in culture medium however can help provide taxonomic criteria for the

classification of fungal isolates into well defined genera and species.

Phenotypic & Biochemical characterization of bacteria

Selective procedures to differentiate organisms on morphological, biochemical,

and growth phenotypes coupled to molecular identification are important to identify

“exotic” types of bacteria. Thus identification of bacteria, based on the phenotypic

characteristics, can sometimes be achieved by direct comparison of unknown bacteria

with known type cultures (Holt and Krieg et al., 2000).

Characterization of bacteria is done on the basis of their cell structure e.g.

bacilli, cocci, spirilla (Christiansen & Weigner, 1991). According to Jordan and

Hungria, (2004) convex elevation of Rhizobium colonies and capsule forming

capacity of microbes is related to the exopolysaccharide gum secretions and can be

identified by capsular staining. Bacterial gram natures and motility characteristics can

be determined as described by Dubey & Maheshwari, (2004). Bacterial motility could

contribute to survival in soil and the initial phase of colonization where attachment

and movement to the root surface are important (Tumbull 2001).

Genus and species identification of a particular isolate can result via

performing a set of biochemical tests (Cappuccino and Sherman, 2006). Thus,

primary identification of bacteria can involve a few simple tests including catalase

and oxidase activity assays utilization of sugars, and indole; methyl red; veges-

proskuer, citrate tests (IMViC) (Dubey & Maheshwari, 2005).

Biochemical characterization of isolates from rhizosphere of desert plants was

performed in order to screen potential phosphate solubilizing bacteria by Gothwal et

al., (2006). Biochemical characterization of bacteria from solid waste degradation

from organic manure was also performed based on IMViC and catalase oxidase tests

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by Zaved et al., (2008). Neelam Yadav and V.K Yadav (2003) characterized bacteria

for growth parameters such as high salt tolerance to 6% NaCl from native soils of

Rajasthan.

The present work was carried out

i. To isolate and enumerate bacteria from various composts; FYM, RSC, and

GVC and soils e.g. rhizosphere, rhizoplane of Cajanus cajan, and cultivable field

soil.

ii. To determine the physicochemical properties of these soils and composts

samples.

iii. To study morphological and biochemical characteristics of selected bacterial

isolates for determining genera and species identities.

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MATERIALS & METHODS

Geographical distribution of Samples:

Soil samples & compost applications were collected from agricultural regions

of Samalkot, Pithapuram, Rajupalem, Kakinada areas of East Godavari District,

Andhra Pradesh, India.

Collection of Composts:

Composts, Farm Yard Manure (FYM) were collected from dairy farms in

Samalkot, RSC (Rice Straw Compost) was collected from heaps prepared for

composting in paddy fields of Rajupalem, Gliricidia Vermicompost (GVC) was

collected from the Athchyutha Ramayya Cottage Industry of Vermicomposting,

Kakinada, Andhra Pradesh. Samples were collected from three different areas in the

fields for every soil sample and from compost heaps for compost sample. The

samples, about 100gms each were collected in to sterile polythene biodegradable

black colored bags and were bought to the Microbiology laboratory and passed

though a 2mm sieve to remove hard and large soil particles. The collected and

processed compost samples were stored at 4oC in refrigerator for 7-10 days in the

sterile ziploc polythene bags.

Collection of soil samples:

Agriculturally cultivable field soils, paddy fields cultivated with pigeon pea,

were dug into 60 cms depth for collection of soil samples. The sub surface soils from

three different slots were collected and mixed in the sterile black polythene bags.

Large and hard soil particles were removed and padded through a 2mm sieve to

collect fine soil particles and stored at 4oC for 7-10 days in sterile ziploc polythene

bags.

Collection of rhizosphere soil:

The collection of soil samples was performed by using the method of Harley

& Waid (1975). Soil samples were dug 2-3 inches from the immediate vicinity of

plant root (Cajanus cajan). Soil was collected by withering the roots into black

polythene bags. Rhizosphere soil from three different pigeon pea plants was

collected. Collected soil was passed through 2mm sieve and fine soil collected;

samples were stored at 4oC for 7-10 days in ziploc bags.

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Physico-chemical characteristics of collected soil and compost samples

The physico- chemical characteristics of soils and compost samples were

analyzed at SIFT (State Institute of Fisheries Technology), Kakinada, Andhra

Pradesh. Sample pH was determined by the Potentiometric method, an electrically

operated pH meter (ELICO). Soil EC (Electrical conductivity) was determined using

Conductivity meter (Systronics). Available nitrogen was measured by the Kjeldhal

method (Jackson, 1973). Available phosphorus was estimated by Olsen’s method

(1982). Available potassium in soils was analyzed by flame photometry method

(Systronics). Organic carbon was estimated by the dry combustion method described

by Anderson and Ingram (1993).

Chemicals & Raw Materials:

The chemicals and raw materials used in this study were obtained from

different sources. General chemicals e.g. ammonium Chloride, ammonium sulphate,

calcium chloride, calcium phosphate, magnesium sulphate, and sodium chloride,

H2SO4, HCl, NaOH and ethanol were from Qualigens, India. ACC (1-Amino-

cyclopropane-1Carboxylic acid, chrome azurol S dye, PIPES buffer, tryptophan,

indole acetic acid from Sigma Chemicals, USA. All Media components e.g.

cellulose, glucose, nutrient broth, peptone, agar-agar etc., were from Hi-media, India.

Sterilization of Glassware:

Requirements: H2SO4-dichromate solution (2.5%) (For cleaning glassware),

Distilled water.

Procedure: Glassware, e.g. test tubes, Petri plates, flasks, and pipettes were washed

by dipping in 2.5% H2SO4-dichromate solution and followed by detergent and then

were washed several times with distilled water to remove traces of H2SO4 and

detergent. The glassware then oven dried at 70oC in a hot air oven and sterilized by

wrapping petri plates in paper, plugging test tubes with non absorbent cotton and

placing in hot air oven (Dalal) (1200C for 1 hr by presetting the oven temperature).

Laminar flow chamber (Yorco, New Delhi) was sterilized by wiping with cotton

dipped in ethanol, closing the door and switching on the U.V lamp for 15 minutes.

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Preparation of H2SO4-dichromate solution (2.5%): Potassium dichromate - 25gms.

Conc. H2SO4 - 1litre, Distilled water - 50ml.

Dichromate (25gms) is added into warm water (50ml) and stirred until the crystals

dissolve in it. After cooling, the solution was made up to 1 liter with concentrated

H2SO4. The solution was transferred into a wide mouthed glass chamber and was

covered with a glass slab. Glassware was soaked into this solution for cleaning.

Isolation and Enumeration of Bacteria

Serial dilution technique:

Requirements: Sterile saline (0.85%), Test tubes, (Borosil), Micro pipette

(Qualigens), and a Laminar air flow cabinet with UV lamp (Yorko, New Delhi) were

used in this study.

Preparation of Saline (0.85%): Sodium chloride-8.5gms; Distilled water – 1litre.

NaCl 8.5gms was dissolved in 1litre distilled water to obtain 0.85% Saline. The

solution was dispensed into culture flasks, plugged with non-absorbent cotton and

sterilized in an autoclave at 1210C for 15 minutes. Sterile saline was stored at 4oC and

used for serial dilutions and suspensions of bacterial inoculums.

Procedure: All soil samples and compost samples that were stored were brought to

room temperature (28±1oC). One gram of each of the collected soil and compost

samples were weighed and suspended in sterile saline in test tubes to ten ml and

labeled as 10-1(1:10). The tubes were gently shaken for 5 minutes using an electric

shaker (REMI) to obtain homogenous soil suspension. The Suspensions were made

for all the collected samples.

One ml of soil suspension collected in the supernatant was transferred aseptically

with a sterile pipette into sterile saline sample (9ml) of next test tube. This dilution

was 10-2 (1:100). Similarly, 10 fold dilutions of each sample was performed till the

dilutions reached 10-5 (1:100000). The serial dilutions of soil and compost samples

were performed in a Laminar flow chamber under aseptic conditions. Each dilution of

soil and composts was then used as the bacterial inoculum for culture on nutrient agar

plates.

15

Spread plate method:

Requirements: Nutrient Agar media (Hi media), Petri plates (Borosil), Micropipette

(Qualigens).

Procedure: Dilutions (10-3, 10-4, and 10-5) of serially diluted soil and compost

samples were used for bacterial isolation. Aliquots each of 0.1ml dilution were

transferred onto sterile nutrient agar plates using a micropipette (100-1000µl) with

sterile disposable micro pipette tips (Qualigens). The inoculum was spread evenly on

the plates using a sterile glass spreader (dipped in ethanol and flamed) until the

inoculum was absorbed in the nutrient agar media. This experiment was performed

3X with three replicates for each sample.

Preparation of Nutrient Agar (NA): (Hi-media)

Nutrient agar (28gms) was dissolved in 1000ml of distilled water. The pH of the

medium was adjusted to neutrality (7.2) with 0.1N HCl or 0.1M NaOH. Culture

media flask was plugged with non absorbent cotton and aluminum foil and sterilized

in an autoclave at 15 lbs for 15 minutes.

Preparation of NaOH (0.1M): 0.4gms of NaOH was weighed and dissolved in 10ml

of distilled water and was made up to 100ml with distilled water to obtain 0.1M

NaOH.

Preparation of HCl (0.1N): 0.3ml of concentrated HCl was taken into a measuring

jar and was made up to 100ml with distilled water to obtain 0.1M HCl.

Preparation of Nutrient Agar plates:

Sterilized Nutrient Agar (NA) medium was allowed to cool to 55oC and poured into

petri plates held open with the lid at an angle of 30o under aseptic conditions in a

laminar flow chamber. Three fourth of the Petri plate was filled with Nutrient media

and allowed to cool in the laminar flow chamber for the agar media to set as

semisolid media. The nutrient agar plates were placed one above the other to prevent

water condensation.

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Checking for sterility: NA plates prepared with sterile nutrient agar and sterile saline

(0.85%) were left in the incubator (Yorko Scientific Industries) un-inoculated for

24hr at 37 0 C. Sterilized media plates that did not visually show any microbial growth

or contamination perceived were preserved for future use at 4oC. Traces of turbidity

in saline or visible microbial colonies on NA plates if observed, were discarded. The

contaminated plates and saline were discarded by autoclaving them at 15 lbs for 15

minutes.

Incubation:

Requirements: Incubator (REMI).

Procedure: All the plates inoculated with each respective dilution (10-3, 10-4, and 10-

5) were labeled and incubated in an incubator (M.C.Dalal Agencies) upside down at

37oC for 24 hrs. Sterile saline without soil samples and dilutions were similarly plated

on NA plates and used as controls. The plates were further observed for the

development of well isolated colonies after 24 hours for enumeration.

Enumeration of bacteria (SPC-Standard Plate count):

Requirements: Colony counters (Dalal)

Procedure: The bacterial colonies obtained in different dilutions were counted using

a graduated Quebec colony counter (Dalal). The number of colonies were counted

using the Misra and Miles drop count method (1938) on plates containing between 30

-300 cfus. Serial dilutions were used to determine the number of bacteria in the

original sample and were expressed as colony forming units (cfus) per gram of soil.

Colony forming units = Dilution

coloniesofNumber x Volume factor

Bacterial enumeration was performed for all soil and compost samples that were

diluted and tabulated as the average cfu/gm sample.

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Study of colony characteristics of bacteria:

Requirements: Microbiological Nichrome wire loop (3mm in diameter), Sterile

Nutrient Agar plates.

Nutrient Agar plates: As described earlier.

Procedure: Bacterial cultures (0.01ml) were inoculated on one edge of sterile

nutrient agar plate under aseptic conditions. The inoculum was further streaked by the

quadrant streak plate method (Prescott and Harley, 2002). This method of streaking

was used to isolate individual colonies that express 90% of the preliminary culture

traits. The plates were incubated in an incubator maintained at 370C for 24 hours.

Colony characteristics including pigmentation, elevation, margin, texture, and size

were observed with a magnifying lens and traits were recorded. The experiment was

repeated 3x for all bacterial cultures.

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Bacterial Pure culture preparations:

Nutrient Agar (NA): Nutrient agar was prepared and sterilized as described earlier.

Procedure: Single colony isolated bacterial cultures were streaked onto the NA

slants and were incubated at 37oC for 24 hours. To maintain the viability and

physiology of bacteria the culture slants were stored at 40C by wrapping them in

aluminum foil. These were used as the pure bacterial stocks and were sub cultured

every 15 days on sterile NA plates.

Preparation of NA: As prepared before.

Preparation of NA Slants: NA (10ml) was dispensed aseptically into test tubes

(15ml) and allowed to solidify in a slanting position. Solidified nutrient agar slants

were wrapped in aluminum foil and stored in at 4oC until further use.

Thirty two bacterial cultures exhibiting diverse colony morphologies were selected on

random basis and were purified to obtain pure cultures. These 32 pure bacterial

cultures were characterized based on microscopic and biochemical characteristics.

Study of Microscopic Characteristics of Bacteria:

Various Stains and an Olympus compound microscope (45 x magnifications) were

used to visualize and study Gram natures and morphological characteristics of

bacterial pure cultures.

Gram Staining

Requirements: Crystal violet, saffranine, ethanol, Olympus compound microscope.

Procedure: A loopful of bacterial culture from a well isolated colony was taken and

was spread evenly onto a slide. The smear was allowed to dry and flooded with

crystal violet (Fischer scientific), stained for 1 minute, washed gently under tap water,

and then flooded with mordant iodine (Fischer Scientific) for 1 minute.

Decolourization of the dye was performed with 95% ethanol. In the final step,

saffranine (Fischer Scientific) was used as a counter stain, via incubation for one

minute, followed by a gentle wash under tap water. The stained smear was air dried

19

and observed using Olympus Compound Microscope with magnification 45X and

100X oil immersion lens. Violet colored bacteria were identified as gram positive and

pink colored bacteria, gram negative. Bacterial shape, i.e. short rods and long rods

were also characterized. Experiments were performed in triplicates.

Preparation of Stains: Crystal violet, saffranine, Grams iodine were obtained

commercially from Qualigens Scientifics.

Preparation of decolorizing agent (Ethanol 95%): 95ml of ethanol (100%) was

made up to 100ml with distilled water and stored in reagent bottle for future use.

Capsule staining (Jordan 1984)

Requirements: Nigrosin (10%)

Procedure: A bacterial culture was spread onto a slide and flooded with nigrosin

stain and observed under microscope (45 x magnifications) for the presence of a

capsule. The capsulated and non capsulated forms of bacteria were recorded. The

cells appear colored and capsule colorless. The experiment was repeated 3x for each

culture.

Preparation of Nigrosin (10%): Nigrosin (Water soluble) - 10.0 g, Distilled water -

100 ml, Formalin - 0.5 ml.

Nigrosin was dissolved in 50ml of distilled water and placed in boiling water bath for

30 minutes. Thereafter 0.5ml of formalin is mixed and the contents were filtered

using double filter paper. Filtered stain was made up to 100ml with distilled water.

Endospore stain

Requirements: Malachite green, saffranine, bacterial cultures

Procedure: After seventy two hours of growth bacterial cultures were tested for

sporulation by spreading onto the slides to form uniform smear. The smear was

covered with a filter paper strip and flooded with malachite green stain. The slide was

placed over a steam bath until the stain over the smear began to steam. The filter

paper was then removed gently and the slide was rinsed with water and counter

20

stained with saffranine. The slide was allowed to air dry and observed under

microscope (45 x magnifications). Green cells on a pink background indicated

endospore formation and the results were recorded. The staining procedure was

performed 3X for each culture.

Preparation of Malachite green (5%): Malachite green - 5g, Distilled water - 100

ml.

Malachite green (5gms) was weighed and dissolved in 100ml distilled water to obtain

5% stain and stored in stain bottle.

Saffranine (Aqueous): Obtained from Qualigens Scientifics.

Motility test:

Requirements: Nigrosin (10%)

Procedure: Bacterial motility was observed by the hanging drop method using a

cavity slide. A loopful of one day old bacterial culture in saline was suspended in 1ml

of nigrosin solution. A drop of the suspension was placed on a cover slip. The cover

slip was placed on the shallow side of the cavity slide and sealed with vaseline. The

slide was then observed under microscope to test the motility of bacteria. The

experiment was repeated 3X for each culture.

Preparation of Nigrosin (10%): As prepared before.

Biochemical characterization of the bacterial isolates:

Preparation of Bacterial pure cultures:

Requirements: Nutrient broth, bacterial cultures

Procedure: Sterile nutrient broth (NB) was prepared and 5 ml dispensed in sterile

tubes and plugged with non absorbent cotton. A loopful of the bacterial stock culture

was transferred into the broth aseptically in a laminar flow chamber and mixed by a

vortex mixer and incubated at 370C for 24 hours on a shaker cum incubator (120rpm).

The turbidity of the broth was allowed to reach 0.5 O.D at 620nm which represents

21

the log phase of bacterial growth (109-1010 cfu/ml). These cultures were used for

further biochemical characterization.

Preparation of Nutrient broth: (Hi-media): Nutrient broth (13gms) was dispensed

into flasks containing 100ml of distilled water. The media was boiled to dissolve the

contents. The pH was adjusted to 7.2 with 0.1M NaOH or 0.1N HCl. The media was

made up to 1 liter with distilled water, plugged with non-absorbent cotton and

sterilized in an autoclave at 15lbs for 15minutes.

Preparation of 0.1M NaOH: As prepared above.

Preparation of 0.1N HCl: As prepared above.

Carbohydrate fermentation tests:

Requirements: Peptone water, Andrade’s indicator (Qualigens). Durham tubes

(Qualigens) and sterile screw cap tubes (Borosil) sterilized in hot air oven at 100oC

for 20 minutes.

Procedure: Peptone water (5 ml) was dispensed into each of the test tubes with an

inverted Durham tube without any air bubble formation and plugged carefully. The

tubes were sterilized in an autoclave at 15 lbs pressure for 15 minutes. Each of the

sterilized carbohydrate solutions (0.5ml) were transferred separately using aseptic

conditions into the peptone broth tubes at a final concentration of 1% sugar in

peptone water tubes. The bacterial isolates (0.01ml) were inoculated using a

micropipette and incubated at 37oC for 24 hours. Peptone water tubes with sugar

solution and without bacterial culture served as controls. After incubation, one drop

of Andrade’s indicator was added to the fermented tubes. A change of Andrade’s

indicator color from yellow to pink indicated a positive test for acid formation and

bubble formation in the Durham tube indicated gas formation. The test was

performed in triplicate for all bacterial cultures with all carbohydrate solutions. The

experiment was repeated 3x and results were recorded as positive or negative for

bacterial acid production and gas production.

22

Preparation of Peptone water (1%): Bacteriological peptone (Hi-media)–10gm,

distilled water-1000ml.

Peptone (10gm) was dissolved in distilled water (1000ml) and pH adjusted to 7.2.

Preparation of carbohydrate stock solutions (1%): Glucose, sucrose, arabinose

and mannitol (Qualigens) were weighed ten grams each and dissolved separately in

100 ml of distilled water. These stock solutions were sterilized by Tyndallization

(intermittent heating) using the tyndallizer (100oC). Tyndallization was performed for

20 min. for each carbohydrate solution and the process was repeated for three

successive days to ensure complete sterilization. The sterile carbohydrate solutions

were stored at 4oC.

Andrade’s Indicator (Acid-base): Obtained from Qualigens.

Indole test

Requirements: Tryptone broth, Kovacs reagent, E.coli (MTCC 119) and

Enterobacter aerogenes (MTCC 111)

Procedure: Sterile tryptone broth (5ml) tubes were allowed to cool. Test cultures

(0.01ml) were transferred aseptically into the tryptone broth tubes using a sterile

micropipette and incubated in the incubator at 37oC for 48 hours on a shaker cum

incubator at 120rpm. Positive and negative controls were run by using E.coli (MTCC

119) and Enterobacter aerogenes (MTCC 111) test cultures. The formation of a

crimson red indole ring on the surface of the broth cultures after addition of the indole

reagent indicated a positive test. The test was repeated 3x and the results were

recorded as positive or negative for each bacterial culture.

Preparation of Tryptone broth: Tryptone (Hi-media) - 5.0g, NaCl - 5.0g, Distilled

water - 1000ml

23

Tryptone (5gms) and NaCl (5gms) were dissolved in distilled water and the solution

was made up to 1000ml. The pH was adjusted to 7.2. Tryptone broth (5 ml) was

dispensed into test tubes and then sterilized in an autoclave at 15lbs for 15 minutes

and cooled in laminar flow chamber.

Kovacs reagent: Obtained from Qualigens.

Methyl Red-Voges Proskuer test (MR- VP Test)

Requirements: Glucose phosphate peptone broth (GPP broth), methyl red indicator

(Qualigens), Barritts Reagent (VP reagent), screw cap tubes.

Procedure: Ten ml of glucose phosphate peptone (GPP) broth was dispensed into

sterile screw cap tubes (18x150mm) and 0.01 ml of the bacterial culture was

inoculated into the GPP broth tubes with a micropipette. The culture tubes were

incubated for 48 hours at 370C in a 120rpm shaker cum incubator. After incubation,

the 10ml culture broth was divided into two test tubes (5ml each) for MR and VP

tests. The MR test was performed by adding 1ml of methyl red indicator into the first

tube and the VP test by adding 1ml of Barritt’s Reagent into second tube. The change

of color to red with methyl red reagent indicated a positive test to MR and a color

change to pink with the VP reagent indicated positive for the VP test. The

experiment was repeated 3X and included positive and negative controls (MR +ve

control E.coli, MR –ve control Enterobacter aerogenes, VP +ve control Enterobacter

aerogenes, VP –ve control E.coli. The results were recorded as +/-ve for all the

isolates.

Preparation of Glucose phosphate peptone broth (GPP broth) or (MR-VP

Broth): Glucose - 5.0g, peptone - 5.0g, K2HPO4 - 5.0g, distilled water- 1000ml

Glucose (5gms), Peptone (5gms) and K2HPO4 (5gms) were dissolved in 100ml of

distilled water. The broth was made up to 1000ml with distilled water. The pH was

adjusted to 7.2. The broth was dispensed (5ml) into screw cap test tubes and sterilized

in an autoclave at 15 lbs for 15 minutes.

24

Methyl Red reagent: (Qualigens)

Preparation of Barritts reagent: VPI&VPII

VP Reagent I: α – naphthol (5%): 5gms of α-naphthol was dissolved in 10ml of

absolute ethanol and the solution was made up to 100 ml with absolute ethanol.

VP Reagent II: KOH (40%): 40% KOH solution was prepared by dissolving

40gms of KOH in distilled water and solution was made up to 100 ml with distilled

water.

Citrate utilization test:

Requirements: Simmons citrate agar (Hi-media)

Procedure: Test cultures of 0.01 ml (104cfus/ml) were streaked onto citrate agar

slants and incubated at 37oC for 24 hours. Enterobacter aerogenes was used as a

positive control and E.coli as a negative control. A positive test result was indicated

by a color change of the bromothymol blue indicator present in simmons citrate agar

medium from green to blue, indicating the ability of organism to use citrate as a sole

carbon source. The test was performed in triplicate for each culture and the

experiment was repeated 3x. The results recorded as +/-ve for all the cultures.

Preparation of Simmons citrate agar (Hi-media): Simmons citrate agar was

weighed (2.4gms) and dissolved in distilled water by slightly boiling the media. The

pH adjusted to 7.2 and the solution was dispensed (5ml) into test tubes and sterilized

in an autoclave at 15lbs for 15minutes. The citrate tubes were removed from the

autoclave and allowed to solidify in a laminar flow chamber in a slanting position.

Slants were stored at 4oC for further use.

Catalase test

Requirements: 30% H2O2 (Fischer’s Scientific)

Procedure: Bacterial colonies were taken using microbiological loop and spread onto

slides which were then flooded with 30% H202 to detect catalase activity. Formation

of cloudy air bubbles over the culture showed positive catalase production. The test

was compared with a positive control using Staphylococcus aureus (MTCC 87). The

test was performed 3x and results were recorded.

25

Preparation of H202 (30%): Commercial hydrogen peroxide (30%) was used for the

test.

Oxidase test

Requirements: Oxidase reagent (1%), Pseudomonas aeruginosa (MTCC 2581)

Procedure: Production of oxidase by bacterial cultures was detected using the redox

dye, tetra methyl- p-phenylene-diamine. Bacterial test inocula from culture plates,

were taken on slides and oxidase discs were placed over the culture. A change of

color to purple was recorded as a positive reaction for oxidase activity. Pseudomonas

aeruginosa (MTCC 2581) was used as a positive control. The test was performed 3x

for each culture and recorded +/- ve for the isolates.

Oxidase reagent (1%): Commercially available discs from Qualigens.

Starch hydrolysis

Requirements: Starch Agar medium, Iodine (3%)

Procedure: Starch agar plates were prepared and streaked with pure bacterial culture

and incubated at 37oC for 48 hours. After incubation, iodine was poured onto the

plates. Formation of a blue black color due to starch-iodine complex in the unutilized

places of starch in the agar plates was indicated. A clear halo surrounding the

bacterial colony on the starch agar medium indicated starch hydrolysis by the bacteria

via production of amylase. The test was repeated 3X for each culture and recorded.

Preparation of Starch Agar medium- Starch-20gms, Beef extract- 3gms, Peptone-

5g, Agar-15g, Distilled water-1litre.

Starch (20gms), beef extract (3gms), peptone (5gms) were boiled in 500ml of

distilled water to dissolve the contents. The pH was adjusted to 7.2. Agar (15gms)

was added to the media. The media was made up to 1 liter with distilled water and

sterilized in an autoclave at 15lbs pressure for 15minutes.

Preparation of Iodine (3%): Iodine-1gm, potassium iodide-2gms, 300ml distilled

water.

26

Iodine (1gm) and potassium iodide (2gms) were dissolved in 100ml of distilled water

after grinding in a mortar and pestle. The solution was made up to 300ml using

distilled water.

Gelatin hydrolysis

Requirements: Gelatin medium

Procedure: Bacterial cultures were streaked on gelatin stabs and incubated for 15

days at 37oC. A positive reaction occurs when gelatin becomes due to gelatin

liquefaction by bacterial enzymes. The test was repeated 3X and results were

recorded.

Preparation of Gelatin medium: Gelatin – 40.0gms, Tryptone – 17.0 gms, Soytone–

3.0gms, NaCl – 5.0gms, K2HPO4- 2.5gms, Distilled water – 1000ml, pH 7.0.

Gelatin, tryptone, soytone, sodium chloride and K2HPO4 were weighed and boiled in

1000ml distilled water. The pH adjusted to 7.0 and the solution was sterilized in an

autoclave at 15lbs for 15 minutes. The gelatin agar was dispensed into test tubes and

allowed to solidify in an erect manner. Solidified gelatin stabs were stored at 4oC.

H2S production

Requirements: Gelatin media enriched with (1%) FeCl3, bacterial culture of Proteus

vulgaris (MTCC 426).

Procedure: Each of the bacterial isolates was inoculated in gelatin stabs with a

microbiological needle. Tubes were incubated at 37oC for 24 hours and 48 hours in an

incubator for formation of a black precipitate during growth of the colonies. If the

sulphur containing amino acids in the protein rich media could be metabolized by the

test bacteria, H2S will be produced. H2S was then detected using FeCl3 which forms a

black color in the gelatin stab in the presence of H2S. Tests were run in triplicates

using Proteus vulgaris (MTCC 426) as a positive control. The test was performed 3X

for each bacterial culture and results were recorded as positive or negative for H2S

production.

27

Preparation of gelatin media enriched with 1% FeCl3: Gelatin media prepared as

above and 1gm of FeCl3 was supplemented to 100ml of gelatin media while preparing

and then sterilized in an autoclave at 15lbs for 15minutes.

Urease production

Requirements: 3% Christensen’s urea agar (Hi-media)

Procedure: Sterile urea agar slants were inoculated and streaked with 0.1 ml of

exponentially growing bacterial cultures and incubated at 37oC for 48 hours. The

production of urease by bacteria results in hydrolysis of urea which increases the pH

of the media. The color of the media changes from yellow to pink with the change in

pH due to the presence of phenol red indicator in Christensen’s urea agar (Hi-media).

Positive and negative controls were set up by using Proteus vulgaris and uninoculated

urea agar respectively. The experiment was repeated 3X for every culture isolate and

results were recorded.

Preparation of Christensen’s Urea agar (Hi-media): 3gms of urea agar was

weighed and dissolved in 100ml of distilled water. Urea agar (5ml) was dispensed

into test tubes and sterilized by autoclaving at 15lbs for 15minutes. The test tubes

with urea agar were allowed to cool and solidify in slanting position to obtain urea

slants.

Ammonification test:

Ammonification and nitrification abilities were tested by method described by Dubey,

(2004).

Requirements: Peptone broth, Nessler’s reagent

Procedure: Peptone broth was used as the suitable nitrogen source. Sterile peptone

broth (5ml) was dispensed into sterile test tubes and plugged with cotton plugs.

Exponentially growing bacterial cultures (0.1ml) were inoculated into the peptone

broth tubes and incubated at 25oC for 5 days. Nessler’s reagent (0.5ml) was added to

each tube and the formation of a yellow to brown precipitate indicated

ammonification by the culture isolate. The test was performed in triplicates and was

performed 3x for each isolate and the ammonifiers were recorded.

28

Preparation of Peptone broth: Prepared as before.

Nessler’s reagent: From Qualigens

Nitrification test:

Requirements: Ammonium sulfate broth, Nessler’s reagent, Tromsdorff’s reagent

Procedure: Bacterial test cultures were inoculated into ammonium sulphate broth

and incubated for 48-72 hours in an incubator at 25oC. A few drops of the culture

broth were placed on a watch glass and 1-2 drops of Tromsdorff reagent were added.

Formation of a blue black color indicated the presence of nitrate. If the test culture did

not show nitrate formation, a few drops of the same culture broth was taken and

Nessler’s reagent was added to it. No color formation was used to indicate unoxidized

ammonia and yellow color formation indicated nitrite formation. The test was

recorded as positive and negative for production of nitrate, nitrite and was repeated

3x.

Preparation of Ammonium sulfate broth: (NH4)2SO4 - 2gms, MgSO4.H2O-0.5gms,

FeSO4.7H2O-0.03gms, NaCl-0.3gms, K2HPO4-1.0gms, Distilled water-1litre.

(NH4)2SO4, MgSO4, FeSO4, NaCl were dissolved in 100ml of distilled water and the

solution was made up to 1000ml with distilled water. Ammonium sulfate broth was

sterilized by membrane filter method.

Nessler’s reagent: Qualigens

Preparation of Tromsdorff reagent:

Zinc chloride solution 20% (100ml), Starch - 4g, Potassium iodide- 2g, and Distilled

water - 100ml.

Starch (4gms), KI (2gms) were added to 10ml of 20% ZnCl2. The starch-iodine

solution was made up to 100 ml with the zinc chloride (20%) solution with constant

stirring. The contents were heated to dissolve the starch until a clear solution was

obtained.

Preparation of 20% ZnCl2: 20gms of zinc chloride was dissolved in 100ml of

distilled water.

29

Growth parameters of the bacterial isolates.

Tolerances of bacteria to acidity, alkalinity, salinity and varying temperatures were

performed using a turbidometric method.

Tolerance to pH:

Requirements: Nutrient Broth (NB)

Procedure: Nutrient broth adjusted to different pH ranging from 5-9 was prepared,

sterilized in an autoclave and 5ml each was dispensed into sterile test tubes. The tubes

were inoculated with 0.1ml of exponentially growing bacterial cultures and incubated

at 37oC for 24 hours. Bacteria inoculated into nutrient broth of neutral pH (pH 7)

were used as control. Bacterial growth was measured by turbidometric method (O.D

at 620nm) and the ability of isolates to grow at varied pH was compared with growth

at neutral pH. Experiment was repeated 3x and results were recorded.

Preparation of Nutrient Broth (NB): As prepared earlier.

NB with different pH (5, 8 and 9) was adjusted by using 0.1M NaOH or 0.1M HCl.

Preparation of NaOH (0.1M): 0.4gms of NaOH was weighed and dissolved in

100ml of distilled water to obtain 0.1M NaOH.

Preparation of HCl (0.1M): 0.3ml of concentrated HCl in measuring cylinder was

made up to 100ml with distilled water to obtain 0.1M HCl.

Tolerance to Salinity

Requirements: Nutrient Broth tubes with NaCl

Procedure: Nutrient broth tubes with different salinity ranging from 0.85%, 2% and

3% were prepared, sterilized in an autoclave at 15 lbs for 15 minutes and 5 ml each

was dispensed into sterile test tubes. Exponentially growing cultures of (0.1 ml) were

dispensed aseptically into the nutrient broth tubes in a laminar air flow chamber and

was incubated at 370C for 24 hours. Bacteria inoculated in nutrient broth at salinity

(0.85%) were used as a control. Bacterial tolerance to various NaCl concentrations

30

was measured by turbidity (O.D at 620nm) and compared with growth under control

conditions. The experiment was repeated 3x and results were recorded.

Preparation of NB: As prepared earlier.

Preparation of NaCl Nutrient broth: 0.5gms, 1gm and 2gms of NaCl were weighed

and dissolved in 100 ml of nutrient broths separately to achieve 0.5%, 1%, and 2%

concentrations of salinity. Concentration of 0.85% saline was used as control. The

nutrient broth was sterilized in an autoclave at 15lbs for 15minutes.

Temperature Tolerance:

Requirements: NB tubes, Incubators.

Procedure: Tolerance levels of the isolates to varying temperatures were tested with

0.1 ml of exponentially growing cultures that were dispensed aseptically into sterile

nutrient broth tubes and incubated at 370C, 40oC, 42oC, 45oC in temperature

controlled incubators. The growth rate was determined for each culture by

turbidometry (O.D at 620 nm). The optimal temperature for growth of bacterial

cultures was determined. The experiment was repeated 3x and results were recorded.

Isolation and Enumeration of Soil Fungi: Fungal isolation and enumeration was

performed by Martins Method (1960).

Serial dilution:

Requirements: Soil samples, compost samples

Procedure: Soil samples and compost samples that were stored in 4oC were brought

to room temperature and weighed; 1 gm each and dispensed into sterile saline. The

solution was made to 10ml with saline and was considered as the 10 -1 dilution (1:10).

This was gently mixed on an electric shaker (REMI) and left undisturbed. Serial

dilution was performed as done for bacterial isolations to a dilution of 10-3. The

supernatant was used for isolation of fungi.

31

Spread plate technique:

Requirements: 2% PDA (Hi-Media)

Procedure: Aliquots of 0.1ml from the serial dilutions were transferred onto sterile

dry potato dextrose agar plates and spread evenly using a sterile spreader and

incubated in an incubator at 25o C for 5 days. Fungal colonies of all the dilutions were

enumerated and tabulated. The fungal counts were performed by Misra and Miles

drop plate method. Each dilution was plated in triplicate and the experiment was

repeated 3x.

Preparation of potato dextrose agar media (PDA): (Hi-Media)

PDA (2gms) was weighed and dissolved in 100ml of distilled water and sterilized in

an autoclave at 15lbs for 20minutes. After sterilization 10mg of streptomycin was

added to the sterilized media in Laminar Flow chamber.

Enumeration of Fungi:

Requirements: Colony counter, fungal plates

Procedure: The fungal colonies were counted as per the dilutions and the number of

cfus /gm was calculated using the following formula.

Colony forming units = Dilution

coloniesofNumber x volume factor

Morphological characterization of fungi:

Fungal isolates exhibiting diverse colony characteristics were further observed by

lacto phenol cotton blue (LCB) staining method.

Requirements: Lacto phenol cotton blue stain (LCB).

Procedure: Fungal mycelia of well isolated fungal colonies was picked with a sterile

microbiological needle and placed on a clean slide. The slide was flooded with LCB

stain and a cover slip was placed carefully on top without air bubble formation. The

slide was observed using 10x compound Microscope and the fungal morphology was

recorded. Each fungal colony was examined 3x. Fungal genera were characterized by

32

morphological and sporulating traits and occurrence percentage of each fungus was

recorded using the following formula:

Average No. of a fungal species

Occurrence (%) of fungi = ----------------------------------------x 100

Total No. of fungi

Number of fungal colonies with their occurrence percentage were calculated and

recorded.

Requirements: Lacto phenol cotton blue (LCB): Lactophenol+cotton blue mix for

microscopy obtained from Kemphosol chemicals.

33

RESULTS

1.1 Physicochemical characteristics of soils and composts

Analysis of the physicochemical characteristics of the soil and compost

samples used for bacterial isolation is presented in Table 1.1. All soils and composts

tested showed nearly neutral pH, with the exception of RSC which is lowest. Slight

variations are recorded for electrical conductivity of composts and soils. Nitrogen

content was found more in Rice straw compost (RSC) among the composts and in

rhizoplane soil (RPS) among soils lowest was CFS. Available phosphorus levels were

found to be slightly more in compost samples than in soils, whereas potassium and

organic carbon levels were found to be slightly more in soils compared to composts

(Results compared with standard values obtained from Agriculture Research Station,

Kakinada). GVC was rich in all the three minerals (N, P, and K) that were tested.

Overall, the tested parameters of the samples are not very different from standard

values.

Table 1.1:

Physico-chemical characteristics of Soils & Compost samples

Physico-

chemical

properties

*FYM *RSC *GVC *RS *RPS *CFS

pH 6.8±0.03 6.2±0.02 6.4±0.03 7.1±0.02 6.9±0.03 6.8±0.01

Soil EC mM 0.02±0.01 0.03±0.01 0.05±0.01 0.04±0.01 0.05±0.00 0.03±0.01

Total N% 2.87±0.03 2.94±0.03 2.47±0.01 2.38±0.01 2.87±0.02 1.45±0.03

P205 kg/ha 12.74±0.12 12.38±0.11 13.48±0.01 11.88±0.13 12.14±0.11 11.16±0.11

K kg/ha 149.34±0.52 132.33±0.64 144.53±0.56 148.38±0.04 154.32±0.58 148.8±0.45

Organic C% 12.37±0.12 11.32±0.24 15.38±0.21 13.47±0.18 14.08±0.21 12.48±0.21

*Each value is a mean of triplicate values

FYM - Farm Yard Manure RSC - Rice straw compost

GVC - Gliricidia Vermicompost RS - Rhizosphere soil

RPS - Rhizoplane soil CFS - Cultivable Field Soil

34

1.2 Bacterial enumeration of composts and soils

Bacteria were enumerated and evaluated as colony forming units per gram of

soil (cfus/gm ) and were found to be highest in the Farm yard manure, moderate in

Rice Straw Compost and lowest in vermicompost. Rhizoplane soil showed higher

bacterial count compared to Rhizosphere and Cultivable field soils whixh had

moderate bacterial counts (Table 1.2). The distribution of bacterial genera most

numerous in all the composts and soils as recorded in Table 1.2 Bacillus and

Pseudomonas species were the most numerous in composts and soils were seen. In

soils, Bacillus sps., were predominant followed by Pseudomonas and Enterobacter.

The “normal” soil flora i.e. Azatobacter, Azospirillum, Rhizobium was recorded to be

lesser in distribution in this study. The gram natures and morphological distribution

of bacteria were recorded. Gram positive rods were most numerous (48%) and gram

negative cocci were least (8%) (Graph 1.a). The percentage distribution of bacterial

genera in soil and compost samples is shown in Graph 1.b.

35

Table 1.2: Gram nature and Genera wise distribution of enumerated bacterial

isolates from composts and soil samples

*Each value represents the Mean±SE of three replicates

Cfu = Colony forming units; Gm+ve = Gram positive; Gm –ve = gram negative;

FYM – Farm Yard Manure, RS – Rhizosphere soil,

RSC- Rice Straw Compost, RPS- Rhizoplane soil,

GVC – Gliricidia vermicompost CFS – Cultivable field soil

Bacteria

* Compost Samples *Soil Samples

FYM RSC GVC RS RPS CFS

*Bacteria

cfu/gm

3.8X106 3X106 2.26X106 3.5X106 4X106 2.4X106

Gm +ve rods 34±2.13 38±2.31 48±0.33 41±0.33 32±0.58 38±2.31

Gm –ve rods 26±2.31 22±0.71 10±2.26 26±1.75 28±0.23 22±1.76

Gm +ve cocci 20±3.53 26±0.71 8±3.53 14±0.06 26±1.33 22±1.15

Gm –ve cocci 17±1.33 14±0.33 12±2.31 18±2.67 10±2.67 14±0.93

Bacillus 30±1.33 24.5±0.53 24.6±1.22 19.1±0.58 36.1±0.67 31.2±0.9

Enterobacter 10.1±3.5 18.5±0.88 19.9±0.11 17.0±1.88 16.5±1.33 12.1±3.3

Serratia 12.6±0.4 14.0±2.67 7.2±0.09 10.7±3.23 10.0±0.16 8.6±0.11

Klebsiella 12.1±1.0 11.9±2.62 14.4±1.33 18.04±1.12 11.2±2.31 14.0±1.6

Pseudomonas 20.1±0.0 16.8±1.62 16.4±0.01 12.1±0.53 13.3±0.11 16.6±2.3

Azatobacter 4.1±1.01 5.2±0.53 4.8±2.33 4.4±2.53 4.6±0.03 5.9±0.03

Azospirillum 4.9±0.01 6.5±2.33 7.2±0.67 13.0±1.75 6.0±1.33 5.3±0.51

Rhizobium 8.0±0.01 4.3±0.03 9.8±1.33 7.8±2.21 4.6±0.01 8.2±0.03

36

Graph 1. a Percent distribution of different bacteria in soils and composts

Graph 1. b. Percent distribution of bacterial genera in soils and composts

37

1.3 Colony characteristics and Microscopic characteristics:

Colony characteristics and microscopic characteristics of thirty two bacterial

isolates were tabulated in Tables 1.3a and 1.3b. The colonies of bacterial isolates on

Nutrient Agar were found to be smooth textured but with varied colony margins,

pigments, and densities that were used to form the preliminary identification of the

genera of each isolate. The majority of bacterial colonies were cream colored

Serratia formed pink colored colonies on nutrient agar (Fig. a, Plate-1), Bacillus,

white colored colonies (Fig. b, Plate-1), Klebsiella, large light pink and cream

colored colonies (Fig. c, Plate-1), Pseudomonas, fluorescent green colonies (Fig. d,

Plate-1) and Azospirillum large white colonies with wavy margins.

Of 32 isolates examined five were identified as Bacillus sps. (RB1, RB6, RB8,

RB13, and RB21) and were found to be gram positive and the other twenty seven

isolates were gram negative (for example, see Fig. a & Fig.b, Plate-2). All isolates

were rod shaped but differed in size, seven were long rods and the rest were short rods.

More than half of the bacterial cultures were non-capsulated. Azorhizobium, Klebsiella,

Azatobacter cultures showed elevated colony morphology and were capsulated

(example, Fig.c, Plate-2). Certain species of Bacillus were sporulating (example, Fig.d,

Plate-2).

38

Table 1.3 a: Colony characteristics & Microscopic Characteristics of Bacterial Isolates from soils and composts

Characteristic feature

Bacterial Isolates

RB1 RB2 RB3 RB4 RB5 RB6 RB7 RB8 RB9 RB10

RB11 RB12

RB13 RB14

RB15 RB16

GS + - - - - + - + - - - - + - - - SS S S S LR NS S S NS NS NS NS NS S NS NS NS Cell shape R R LR R LR SR R SR SR LR R R R R SR SR Capsule NC NC NC C NC NC NC NC NC C NC C NC NC NC NC Density O O TI O O TI O O O O O O O O TI TI Elevation C C C Ra C C C C C Ra C C C C C C Surface Sm Sm Sm Sm Sm Sm Sm Sm Sm Sm Sm Sm Sm Sm Sm Sm Texture Margin W W W W W Ro I E E Ro W I I W Ro Ro Pigments W NP P Cr NP W NP W W Cr GY P W NP GY GY Possible genus A E C D E A F A E F H G A H H H

A – Bacillus E – Enterobacter C – Serratia D – Azospirillum

F – Azatobacter H - Pseudomonas G – Klebsiella

GS=Gram stain, + Gram Positive rods, -Gram negative rods

SS= Spore stain, S=sporulating, NS= non-sporulating, R= rods, LR = long road, SR = short rods

C=capsulated, NC=non-capsulated, O = opacity, Tl = translucent

C= convex, Ra = raised, Sm = smooth, W = wavy, Ro= round, E = entire, I = irregular,

NP=no pigment, GY = greenish yellow, P= Pink, Cr= Cream

39

Table 1.3 b: Colony characteristics and Microscopic Characteristics of Bacterial Isolates from soils and composts

Characteristic feature

Bacterial Isolates

RB17 RB18 RB19 RB20 RB21 RB22 RB23 RB24 RB25 RB26 RB27 RB28 RB29 RB30 RB31 RB32 GS - - - - + - - - - - - - - - - - SS NS NS NS S NS S S NS NS NS S S S S NS NS Cell shape R R LR R LR SR SR LR SR SR R R R R SR LR Capsule NC NC C NC NC NC C NC NC NC C C C NC NC C Density O O TI O O TI O O O O O O O O TI TI Elevation C C C C C C C C C C Ra Ra C Ra Ra C Surface Sm Sm Sm Sm Sm Sm Sm Sm Sm Sm Sm Sm Sm Sm Sm Sm Texture Margin W W W W W Ro I E E Ro W I I W Ro Ro Pigments NP P C GY NP NP P P W GY NP C NP NP GY W

Possible genus E C G H A H D C B H F G B B B D

A – Bacillus E– Enterobacter C – Serratia F – Azatobacter

H - Pseudomonas G - Klebsiella D – Azospirillum B - Rhizobium

GS=Gram stain, + Gram Positive rods, -Gram negative rods

SS= Spore stain, S=sporulating, NS= non-sporulating, R= rods, LR = long road, SR = short rods,

C= capsulated, NC= non capsulated, O = opacity, TI = translucent,

C= convex, Ra = raised, Sm = smooth, W = wavy, Ro= round, E = entire, I = irregular,

NP=no pigment, GY = greenish yellow, P= Pink, C= Cream

40

41

42

1.4 Biochemical characterization of bacteria

For species identification of bacteria, biochemical characteristics were

performed for thirty two isolates and were recorded in Tables 1.4a&1.4b. Bacteria

belonging to Bacillus sps. (RB1 RB6, RB8, RB13, and RB21) were gram positive,

motile, sporulating rods, and were catalase positive. The species were identified to

be (RB1) Bacillus subtilis( Indole+ve), (RB6) B.licheniformis (oxidase +ve, H2S

+ve), B.cereus (H2S –ve), B.pumilis (Indole+ve, H2S-ve) and B.megaterium (oxidase

-ve) respectively. Isolates identified as belonging to genera Enterobacter (RB2, RB5,

RB9, and RB17) were found to be motile, except for RB5 and was distinguished as

Enterobacter asburiae (non motile). RB9 belonged to Enterobacter agglomerans as it

showed an indole –ve reaction and RB17 was assigned as Enterobacter cancerogenus

as it was positive for gelatin liquefaction and catalase tests and negative for oxidase

activity. Bacteria belonging to genera Klebsiella were RB12, RB19, and RB28. RB12

as non motile, MR +ve, VP –ve, could not produce H2S and was determined to be

Klebsiella panticola. RB19 and RB28 were MR –ve & Indole +ve, hence matched to

K. oxytoca. Bacterial strains RB3, RB18, and RB24 matched the genus Serratia. RB3

was distinguished as Serratia proteamaculans, based on positive citrate and MR

test. RB18 & RB24 were identified as Serratia marcescens as they were MR-ve.

Pseudomonas strains (RB11, RB14, RB15, RB16, RB20, RB22, RB26), were

distinctive in having catalase and oxidase activities and all the species of

Pseudomonas were positive in nitrate reduction tests. Azatobacter (RB7, RB10,

RB27), Azospirillum (RB23, RB32) Sinorhizobium (RB25, RB30) were identified

based on nitrite reduction tests.

Carbohydrate fermentation profiles for each isolate (Andrades indicator test);

and gas production (Durham tubes) are shown in Tables 1.4a &1.4b. Examples of

biochemical tests and the control results are presented in Fig. a for indole rest, Fig. b,

for methyl red test, Fig. c for voges proskauer and Fig. d for citrate utilization in

Plate-3. Enzyme activities catalase, oxidase, and amylase tests, are shown in Fig.a,

Fig. b, Fig. c of Plate-4, and carbon source utilization, e.g. sugar fermentations is

shown in Fig. d, Plate 4.

43

Table 1.4 a: Biochemical characterization of the bacterial isolates from soils and composts

A - Bacillus subtilis, B - Enterobacter aerogens, C - Serratia proteamaculans, D – Azospirillum brasilense, E - Enterobacter

asburiae,

F - Bacillus licheniformis, G - Beijerinckia indica, H - Bacillus pumilus , I - Enterobacter agglomerans, J - Azatobacter chroococcum

K - Pseudomonas alkaligens, L - Klebsiella panticola, M - Bacillus cereus , N - Pseudomonas cepacia, O – Pseudomonas fluorescence

P- Pseudomonas

Name of the test RB1 RB2 RB3 RB4 RB5 RB6 RB7 RB8 RB9 RB10 RB11 RB12 RB13 RB14 RB15 RB16 Motility + + + + - + + + + + + - + + + + Catalase + + - + + + + + + + + - + + + + Vogues Proskeur + + + - + + + + - + D - + D D D Citrate utilization + + + - + + - + - + - - + - - - Starch hydrolysis - + + + + + + - + + + + + + + + Indole + + - + + + + + - + + - + + + + Gelatin hydrolysis - + + + + - - - + - - + - - - - Methyl red - + + - - - - - - - + + - - - - Nitrate reduction - + + - + + - - - + + - - + + + Ammonification - + - - + + + + + + - - + + + - Urease - + - - - - - - - + + - - + + + Oxidase - + - + - + + - + + + + + + + + H2S - + - + + + - - + - - - - + - + Acid production from different carbon substrates Glucose + + + + + + + + + + + + - + - - Arabinose + - + + + - - + + + + + + + + + Mannitol + - + + + - - + + + + + + - + + Sucrose + - + - + - - + - + - + + - - - identfication of Bacteria A B C D E F G H I J K L M N 0 P

44

Table 1.4 b: Biochemical characterization of the bacterial isolates. Name of the Test RB17 RB18 RB19 RB20 RB21 RB22 RB23 RB24 RB25 RB26 RB27 RB28 RB29 RB30 RB31 RB32

Motility + + + + + + + + + + + + + + + + Catalase + + + + + + + + + + + + + + + + Vogues Proskeur + + + - + + + + - + - - - - - - Citrate utilization + + - - + + - + - + - - - - - Starch hydrolysis + + + + + + + - + + + + + + + + Indole + + + + + + + + + + + - + - + + Gelatin hydrolysis + + + - - - - - - - - - - - - - Methyl red + - - - - + - - - - - - - - - - Nitrate reduction - + + + + + - - - - - - - - - - Ammonifiers - + + + - - + + + - - + + + - - Urease + - + + - - + + - - + + - - - - Oxidase - + D + - + + - + + + + + + + + Acid production from different carbon sources Glucose + - + + + - - + + + + + + + + + Arabinose + - + + + - - + + + + + + + + + Mannitol + - + - + - - + - + - + - + + +

Sucrose + + - + + - + + _ + + - - - - - identfication of Bacteria A B C D E F G H I J K L M O Q R

A - Enterobacter cancerogenus B - Serratia marcescens C - Klebsiella oxytoca D -Pseudomonas cepacia E - Bacillus megaterium F - Pseudomonas mallei G - Azospirillum brasiliense H - Serratia marcescens I - Sinorhizobium fredii J-Pseudomonas cepacia K - Azatobacter beijerinckii L - Klebsiella oxytoca M - Bradyrhizobium japonicum O - Sinorhizobium meliloti Q - Rhizobium leguminosarum, R - Azospirillum lipoferum

45

46

1.5. Growth Parameters of Bacterial Isolates: Thirty two bacterial isolates were

tested for their growth potential under varied pH, temperature, salinity and their

growth efficiencies was recorded (Table 1.5). There was no variation among

bacterial isolates for growth potential with varied physical parameters except Bacillus

sps., Enterobacter and Pseudomonas sps., which showed tolerances to acidic pH,

salinity and high temperatures.

47

Table 1.5 Growth tolerances of bacterial isolates at different pH, NaCl concentration, temperature

Physical

growth

parameter

Bacterial isolates

RB

1

RB

2

RB

3

RB

4

RB

5

RB

6

RB

7

RB

8

RB

9

RB

10

RB

11

RB

12

RB

13

RB

14

RB

15

RB

16

pH 5 + + + + + + - + + + + + - - + +

pH 8 - - + - + - + - - - - - - - - +

pH 9 - - + - - - - - - - - - + + - -

0.85% Nacl + + + - + + + + - + + + - + +

2% Nacl + + + - + + + + - + - - - - + +

3%Nacl + + + - + + - + - + - - + - - -

4 0c - - - - - - - - - - - - + + - -

15 0c + + - + + + + + + + + + - + - -

25 0c + + + + + + + + + + + + - + + +

37 0c + + + + + + + + + + + + + + + +

42 0c + + + + + + + + + + + + + + + +

55 0c - - - + + - - - + + + - - - + +

Contd….

48

+ : Luxuriant Growth(O.D. ≥ 0.5), - : Scanty/No Growth (O.D. < 0.5)

Table 1.5 Growth tolerances of bacterial isolates at different pH, NaCl concentration, temperature

Physical

growth

parameter

Bacterial isolates

RB

17

RB

18

RB

19

RB

20

RB

21

RB

22

RB

23

RB

24

RB

25

RB

26

RB

27

RB

28

RB

29

RB

30

RB

31

RB

32

pH 5 + + + + + + - + + + + + + + + +

pH 8 - - + - + - + - - - - - - + - -

pH 9 - - + - - - - - - - - - - + - -

0.85% Nacl + + + - + + + + - + + - - - - -

2% Nacl + + + - + + - + - + - - - - - -

3%Nacl - - - - - - - - - - - - - - - -

4 0c + + - + + + + + + + + + + - + +

150c + + + + + + + + + + + + + - + +

250c + + + + + + + + + + + + + + + +

370c + + + + + + + + + + + + + + + +

420c + - - + + - - - + + + - + + + +

550c - - - - - - - - - + - - - + + +

49

1.6 Enumeration, characterization and occurrence (%) of fungi

Fungi were enumerated from soils and composts and were tabulated as cfus/gm

(Table 1.6). Fungal samples were examined microscopically and morphological

phenotypes were recorded. Maximum fungal counts were observed in GVC in

composts. Mucor, Fusarium and Penicillium (Fig. a, Plate-5) were found to be more

prevalent in soils and composts compared to other fungal genera. The percentage

distribution of fungi is shown in Graph 1.c; and the fungal genera distribution in

Graph 1.d.

Table 1.6: Distribution of fungal genera in composts and soil samples

*Fungi

(cfus/gm)

Composts Soil

FYM RSC GVC RPS RS CFS

Fungi x103 60±1.2 40±2.11 80±1.22 60±2.2 85±1.2 80±1.32

Mucor 22±1.33 12±1.32 17±0.91 28±1.22 11±0.58 20±0.81

Rhizopus 18±0.98 22±2.14 19±0.97 11±1.56 27±0.67 7±1.2

Aspergillus 19±0.76 29±1.23 14±1.11 18±0.67 8±1.97 13±1.22

Fusarium 23±0.45 19±1.24 11±1.21 15±0.87 12±1.78 28±0.91

Macrophomina 14±1.23 24±2.12 15±1.23 12±1.24 25±1.67 10±0.51

Penicillium 16±1.43 13±1.33 28±2.11 22±2.07 11±1.44 11±0.58

Yeast 7±1.45 19±0.99 18±0.54 14±1.22 21±0.54 28±1.11

*Values are Mean±S.D of triplicate values

50

Graph 1.c Percentage distribution of fungi in soils and composts

Graph 1. d. Percentage distribution of fungal genera in soils and composts

51

52

DISCUSSION

Physicochemical characteristics

Soil chemical parameters varied with crop management practices and were

different in each location. According to Tisdale et al., (1993) soil amendments such

as composts, fertilizers, microbial inoculants, and cropping methods can significantly

alter pH and other physical and biological indicators. In our study all the soils and

composts showed near neutral pH, salinity, and electrical conductivity. Rice straw

compost had slightly lower pH values compared to other soils and composts. The

lower pH of RSC in our study might be due to higher oxidation of organic substrates

with release of acids. Bashan et al., (2000), described that changes in acidity or

alkalinity of composts could be due to organic amendments and organic degradations.

Composts and soils that did not show variation in acidity or alkalinity might be due to

the buffering capacity of soil during mineralization that prevents large changes in pH

(Grayston et al., 2004).

In our study there was a higher content of available nitrogen and phosphorus in

composts compared to soils, which may be due to oxidation of organic matter.

Organic carbon and phosphorus were also higher in composts than soils. Samples

that showed higher carbon concentrations were also higher in potassium, phosphorus,

and nitrogen. This result may indicate that the proportion of organic carbon is related

to the availability of soil minerals such as nitrogen, phosphorus and potassium.

Similar results were obtained by Goyal et al., (1999). They concluded that the higher

the value of soil organic carbon matter, the higher the levels of nutrient turnover. In

the present study, the rhizoplane soil was reported with the highest potassium levels

compared to other soils and composts and this may be due to the increased of cation

Exchange Capacity (CEC) of the rhizosphere region which can hold more

exchangeable K+ by mass action as proposed by Bibuthi B Das & M.S Dkhar

(2011). Increased N, P, and K concentrations in soils after application of composts

was shown in the studies of Saha et al., (1995) & Hontzeas et al., (2004). The mineral

contents of composts and soils may vary based on organic matter, carbon substrates

and microbial activity (Atiyeh et al., 2000a; Crecchio et al., 2004). Tharmaraj et al.,

(2011) also illustrated that compost applications improve N, P, and K concentrations

in soils depending on the substrates that were responsible for mineralizations. Bibhuti

53

et al., (2011) reported that application of composts enhanced various physicochemical

properties of soil and promoted microbial growth. Our results suggest that composts

(FYM, GVC, and RSC) may have substrates for mineralization or minerals directly in

their composition. RPS appears to be superior to other soils in mineral contents

which might be due to higher levels of organic acids and rhizo-depositions.

Bacterial isolation and enumeration

Of the composts that were enumerated FYM showed the highest bacterial

counts. The physicochemical characteristics appeared to complement these results as

it is as moderate in organic carbon and neutral in pH. Our data indicate that supported

the highest growth of bacteria (Cfus) of FYM compared to other composts although

differences were not that great. This is in correlation with the report of Korwar et al.,

(2006), who stated that increased soil organic carbon improves the physical, chemical

and biological properties of soil which would aid in microbial proliferation. Hameeda

et al., (2006) emphasized that application of recommended doses of N, P, K, and

fertilizers in agricultural systems likely influences microbial proliferation. Thus FYM

(3.8x106 cfus/gm) amongst composts and RPS (4x106 cfus/gm) amongst soils with

substantial amounts of N,P, and K likely contributes to proliferating bacterial counts

and diverse types of bacteria as compared to other soil and compost habitats.

Rhizosphere and rhizo-depositions affect microbial communities (Grayston et

al., 2004; Kloepper, 1993). Composts have shown to increase bacterial counts

compared to untreated soils in the studies of Atiyeh et al., (2001) and Arancon et al.,

(2004). Thus, there is a positive relationship between nutrient levels and microbial

proliferation in composts and soils (Jack and Thies, 2006). Incorporation of composts

enhances mineralization of soils, which in turn, improves availability of C, N, P; and

all these attributes support proliferation of diverse types of bacterial genera (Gupta

and Sharma, 1999; Gyaneshwar, 2001; Arancon et al., 2004).

Colony characteristics and Microscopic characteristics

All the strains described in this study were isolated by plating. Bacillus isolates

produced white pigmented colonies on nutrient agar plates and were widely

distributed in the soils and composts examined. In the present study bacterial genera

were identified based on cell shape, sporulation pattern, colony morphology, and

54

biochemical tests using keys published by Palleroni (1984). Klebsiella formed large

convex elevated colonies that were mucoid showing the characteristic of capsulated

organisms as described earlier by Jordan (1984). Serratia had a prominent feature

namely formation of pink colored colonies on nutrient agar plates. Species of

Enterobacter gave diffused cream colored colonies. Pseudomonas showed varied

colony characters including black to greenish colonies and some exhibited

fluorescence. Fluorescent Pseudomonas were found to be the predominant group in

soils in a study by Dey et al., (2004). In this study bacterial colonies were also seen

that secreted slimy, copious, extracellular polysaccharides, and were assigned to

Azatobacter. Hungria et al., (2000) also reported similar slimy copious extracellular

polysaccharides of Azatobacter in studies of symbiotic and non-symbiotic bacteria.

Rods were more prevalent than cocci. Spiral shaped bacteria were recorded

lesser in numbers than other bacteria in both composts and soils. Baudoin et al.,

(2002) also reported similar results i.e. that spiral bacteria were fewer in number in

soils than rods and cocci. Our results were similar with results of Joshi P, Bhatt, A.B.

(2011) on microbial enumeration and characterization in wheat rhizosphere.

Increased organic substrates increases mineralization, and this in turn increased

numerical bacterial shifts in gram negative and gram positive bacteria (Powlson et al.,

2003). In our studies populations of Bacillus were found to predominant in all soils

and composts with Enterobacter next in abundance Pseudomonads were less

abundant in general, but were found at their highest levels in FYM among composts

and cultivable field soil among soils. Pseudomonads were reported to be the

predominant group in rhizosphere and rhizoplane soils (Sharma, 2003; Sharma,

2005). But, Hameeda et al., (2006) reported predominance of Pseudomonas and

Bacillus in composts. Pseudomonads were found to be rapid colonizers of organic

matter and composts and are widely distributed in diverse agricultural ecosystems

(Jayashree, 2000). Bacilli were dominant (40%) in studies of bacterial enumerations

from composts and soils (Hoitink and Boehm, 1999; Joshi, 2011). Serratia formed a

10% of soil microbes but were found to be lesser in our studies. Most of the genera

isolated in our study were non symbiotic diazotrophs.

55

Biochemical characteristics

The major genera identified in our studies biochemically were Bacillus,

Pseudomonas, Serratia, Enterobacter and Klebsiella. These genera were also found

in composts and soils in the studies of Aira et.al, (2002). Klebsiella panticola was

positive for methyl red, while Klebsiella oxytoca was positive in urease tests.

Serratia, Bacillus and Pseudomonas were positive for enzyme activities including

catalase and oxidase. Bacillus and Enterobacter were positive for amylase

production. Thus, the biochemical tests allowed for a robust identification of bacterial

genera and often species, in our sample and others (Cavender et al., 2003). Bacteria

isolated from composts and soils were characterized using biochemical approaches

as by Boehm (1993), Rodriguez (2000), Aneja (2003), Hameeda et al., (2006). Tilak

et al., (2004) isolated Azatobacter chroococcum, Azospirillum brasilense,

Pseudomonas fluorescens, Pseuodomonas putida, Bacillus cereus from pigeon pea

soils and these bacteria were characterized by using biochemical tests. Gothwal,

(2006) characterized Pseudomonas cepacia biochemically and reported that the strain

is catalase positive. Similarly Bisen (1996) characterized bacteria that were able to

produce acid from sugars, were rod shaped, sporulating, gram positive, and catalase

positive related their isolates to the genus Bacillus. Motile, citrate negative and nitrate

positive bacteria were characterized by Neelam Yadav (2003) as Bacillus coagulans.

Growth parameters of bacteria

Bacteria can tolerate various pH ranges from acidity to alkalinity. Although

the optimum growth temperature for many bacteria is 37oC, Bacillus and

Pseudomonas can tolerate higher temperature range of 50-550C. Burges, (1977)

demonstrated similar tolerances of Bacillus species to high temperatures. Diverse

tolerance levels of Bacillus to high temperatures and salinity were also reported by

Bertrand (2001). Bacillus and Pseudomonas counts were high in composts when

compared to soils in our study. This can potentially be explained in terms of heat

tolerance heat, as high temperatures are generated during the composting process.

This is consistent with results of Eghball (1997), who showed that microbes growing

in composts are mesophilic and thermophilic and help organic matter breakdown.

In our study all the isolates showed profuse growth in slightly acidic and neutral pH

but gradually showed reduced growth at higher pH indicating that most of these

56

bacteria are acidophilic and not alkaliphilic. This is similar to studies by Atiyeh

(2000) who showed that thermophilic and acidophilic bacteria proliferate in

composts.

Bacillus, Enterobacter and Pseudomonas were halophilic, tolerating salinity

upto 2% in the present study. The bacterial strains that were reported to be

thermophilic (Bacillus, Pseudomonas, Enterobacter) also tolerated high salinity.

Rodriguez (2000) characterized bacterial growth parameters and identified

thermophilic and halophilic bacteria from soil. Species of Bacillus that were

halophilic and thermophilic were also isolated and characterized by Neelam Yadav

(2003) from soils of Rajasthan.

Overall, the genera isolated from soils and composts in this study were highly

diverse in their growth requirements, likely reflecting the habitats from which they

originated.

Isolation and characterization of fungi:

Microbial enumeration revealed that the bacterial population in all the

sources studied was high but that fungal counts were comparatively low. Lower

fungal counts probably reflected due to slower growth rates or may be due to the

neutral pH that supported bacterial growth, while slightly acidic pH could support

fungal growth. This was in accordance with the studies of Grappelli et al., (1987),

that indicated that fungi can withstand lower pH conditions due to higher oxidation of

organic matter and release of organic acids. In our observations, FYM that was high

in mineral content and organic content also had the most fungal counts. Evidence

from studies by Girvan et al., (2004) indicated that soil fungal populations are largely

favored by organic content. Higher numbers of fungi (Cfus/gm) was observed in

GVC and FYM than other composts and soils in this study. This may correlate to

high P content of these composts. Increased fungal counts in correlation with

available P, phospholipids and fatty acids was seen in the studies of Kandeler et al.,

(2003) and Swer et al., (2011). Greater number of fungi was observed in composts,

likely reflecting available organic matter that helps fungal growth (Widmer 1999).

57

Fungal populations can be characterized based on macroscopic and

microscopic features such as morphological observations and sporulation patterns.

Fungi were identified by Ellis et al., (1993); Watanabe (1994); Swer et al., (2011)

based on hyphae and sporulating structures.

Fungal phytopathogens Fusarium and Macrophomina are common in soils

and composts. In addition, Siddiqui (2002), reported the occurrence of

phytopathogens including Fusarium , Macrophomina, Aspergillus, Penicillium in

composts.

58

CONCLUSIONS

Ø Physicochemical characteristics of composts and soils did not vary greatly and could

support both bacterial and fungal growth.

Ø Gram positive rods were abundant bacteria in all the soils and composts in our study.

Bacillus and Pseudomonas were found to be the major genera found. The

thermophilic and halophilic natures of these bacteria may explain their abundance in

the composts and soils used in this study.

Ø Thirty two isolates belonging to eight genera were identified based on colony and

microscopic characteristics. The bacteria were further identified to the species level

using biochemical tests.

Ø Fungi including Fusarium and Macrophomina were predominant in all soils and

composts studied.