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

Transcription

1 CHAPTER I Isolation and Characterization of Bacteria and Fungi from Soils and Composts

2 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). 1

3 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 cereallegume 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 2

4 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 3

5 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. 4

6 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 5

7 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. 6

8 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 7

9 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 8

10 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). 9

11 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; vegesproskuer, 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 10

12 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. 11

13 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 4 o C 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 4 o C 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 4 o C for 7-10 days in ziploc bags. 12

14 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, H 2 SO 4, HCl, NaOH and ethanol were from Qualigens, India. ACC (1-Aminocyclopropane-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: H 2 SO 4 -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% H 2 SO 4 -dichromate solution and followed by detergent and then were washed several times with distilled water to remove traces of H 2 SO 4 and detergent. The glassware then oven dried at 70 o C 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) (120 0 C 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. 13

15 Preparation of H 2 SO 4 -dichromate solution (2.5%): Potassium dichromate - 25gms. Conc. H 2 SO 4-1litre, 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 H 2 SO 4. 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 C for 15 minutes. Sterile saline was stored at 4 o C 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±1 o C). 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. 14

16 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 ( µl) 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 55 o C and poured into petri plates held open with the lid at an angle of 30 o 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. 15

17 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 4 o C. 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 37 o C 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 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 = Number of colonies Dilution x Volume factor Bacterial enumeration was performed for all soil and compost samples that were diluted and tabulated as the average cfu/gm sample. 16

18 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 37 0 C 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. 17

19 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 37 o C for 24 hours. To maintain the viability and physiology of bacteria the culture slants were stored at 4 0 C 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 4 o C 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 18

20 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) g, Distilled water ml, Formalin 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 19

21 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 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 37 0 C 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 20

22 the log phase of bacterial growth ( 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 100 o C 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 37 o C 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. 21

23 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 (100 o C). 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 4 o C. 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 37 o C 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 ml 22

24 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 37 0 C 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, K 2 HPO 4-5.0g, distilled water- 1000ml Glucose (5gms), Peptone (5gms) and K 2 HPO 4 (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. 23

25 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 (10 4 cfus/ml) were streaked onto citrate agar slants and incubated at 37 o C 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 4 o C for further use. Catalase test Requirements: 30% H 2 O 2 (Fischer s Scientific) Procedure: Bacterial colonies were taken using microbiological loop and spread onto slides which were then flooded with 30% H 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. 24

26 Preparation of H (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 37 o C 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. 25

27 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 37 o C. 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, K 2 HPO 4-2.5gms, Distilled water 1000ml, ph 7.0. Gelatin, tryptone, soytone, sodium chloride and K 2 HPO 4 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 4 o C. H 2 S production Requirements: Gelatin media enriched with (1%) FeCl 3, 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 37 o C 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, H 2 S will be produced. H 2 S was then detected using FeCl 3 which forms a black color in the gelatin stab in the presence of H 2 S. 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 H 2 S production. 26

28 Preparation of gelatin media enriched with 1% FeCl 3 : Gelatin media prepared as above and 1gm of FeCl 3 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 37 o C 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 25 o C 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. 27

29 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 hours in an incubator at 25 o C. 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: (NH 4 ) 2 SO 4-2gms, MgSO 4.H 2 O-0.5gms, FeSO 4.7H 2 O-0.03gms, NaCl-0.3gms, K 2 HPO 4-1.0gms, Distilled water-1litre. (NH 4 ) 2 SO 4, MgSO 4, FeSO 4, 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% ZnCl 2. 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% ZnCl 2 : 20gms of zinc chloride was dissolved in 100ml of distilled water. 28

30 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 37 o C 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 37 0 C for 24 hours. Bacteria inoculated in nutrient broth at salinity (0.85%) were used as a control. Bacterial tolerance to various NaCl concentrations 29

31 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 37 0 C, 40 o C, 42 o C, 45 o C 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 4 o C 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 The supernatant was used for isolation of fungi. 30

32 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 25 o 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 = Number of colonies Dilution 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 31

33 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. 32

34 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 Physicochemical properties *FYM *RSC *GVC *RS *RPS *CFS ph 6.8± ± ± ± ± ±0.01 Soil EC mm Total N% P kg/ha K kg/ha Organic C% 0.02± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ±0.21 *Each value is a mean of triplicate values FYM - Farm Yard Manure GVC - Gliricidia Vermicompost RPS - Rhizoplane soil RSC - Rice straw compost RS - Rhizosphere soil CFS - Cultivable Field Soil 33

35 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. 34

36 Table 1.2: Gram nature and Genera wise distribution of enumerated bacterial isolates from composts and soil samples * Compost Samples *Soil Samples Bacteria FYM RSC GVC RS RPS CFS *Bacteria 3.8X10 6 3X X X10 6 4X X10 6 cfu/gm Gm +ve rods 34± ± ± ± ± ±2.31 Gm ve rods 26± ± ± ± ± ±1.76 Gm +ve cocci 20± ±0.71 8± ± ± ±1.15 Gm ve cocci 17± ± ± ± ± ±0.93 Bacillus 30± ± ± ± ± ±0.9 Enterobacter 10.1± ± ± ± ± ±3.3 Serratia 12.6± ± ± ± ± ±0.11 Klebsiella 12.1± ± ± ± ± ±1.6 Pseudomonas 20.1± ± ± ± ± ±2.3 Azatobacter 4.1± ± ± ± ± ±0.03 Azospirillum 4.9± ± ± ± ± ±0.51 Rhizobium 8.0± ± ± ± ± ±0.03 *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 35

37 Graph 1. a Percent distribution of different bacteria in soils and composts Graph 1. b. Percent distribution of bacterial genera in soils and composts 36

38 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). 37

39 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 RB1 0 RB1 1 RB12 RB1 3 RB14 RB1 5 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 38

40 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 39

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43 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, H 2 S +ve), B.cereus (H 2 S ve), B.pumilis (Indole+ve, H 2 S-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 H 2 S 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. 42

44 Table 1.4 a: Biochemical characterization of the bacterial isolates from soils and composts 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 H 2 S 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 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 43

45 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 44

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47 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. 46

48 Table 1.5 Growth tolerances of bacterial isolates at different ph, NaCl concentration, temperature Bacterial isolates Physical growth parameter RB1 RB2 RB3 RB4 RB5 RB6 RB7 RB8 RB9 RB10 RB11 RB12 RB13 RB14 RB15 RB16 ph ph ph % Nacl % Nacl %Nacl c c c c c c Contd. 47

49 Table 1.5 Growth tolerances of bacterial isolates at different ph, NaCl concentration, temperature Physical growth Bacterial isolates parameter RB17 RB18 RB19 RB20 RB21 RB22 RB23 RB24 RB25 RB26 RB27 RB28 RB29 RB30 RB31 RB32 ph ph ph % Nacl % Nacl %Nacl c c c c c c : Luxuriant Growth(O.D. 0.5), - : Scanty/No Growth (O.D. < 0.5) 48

50 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 Composts Soil (cfus/gm) FYM RSC GVC RPS RS CFS Fungi x ±1.2 40± ± ±2.2 85±1.2 80±1.32 Mucor 22± ± ± ± ± ±0.81 Rhizopus 18± ± ± ± ±0.67 7±1.2 Aspergillus 19± ± ± ±0.67 8± ±1.22 Fusarium 23± ± ± ± ± ±0.91 Macrophomina 14± ± ± ± ± ±0.51 Penicillium 16± ± ± ± ± ±0.58 Yeast 7± ± ± ± ± ±1.11 *Values are Mean±S.D of triplicate values 49

51 Graph 1.c Percentage distribution of fungi in soils and composts Graph 1. d. Percentage distribution of fungal genera in soils and composts 50

52 51