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Maharshi, Anjisha R., 2012, “Studies on Phytohormone Producing Plant Pathogenic Fungi”, thesis PhD, Saurashtra University http://etheses.saurashtrauniversity.edu/id/eprint/575

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Acknowledgments

This thesis arose in part out of years of research that has been done since I came to PPMB Lab. By that time, I have worked with a great number of people whose contribution in assorted ways to the research and the making of the thesis deserved special mention. It is a pleasure to convey my gratitude to them all in my humble acknowledgment.

In the first place I would like to record my gratitude to Prof. Vrinda. S. Thaker for her supervision, advice, and guidance from the very early stage of this research as well as giving me extraordinary experiences throughout the work. Above all and the most needed, she provided me unflinching encouragement and support in various ways which made her a backbone of this research and so to this thesis. Her involvement with her originality has triggered and nourished my intellectual maturity that I will benefit from, for a long time to come. Her truly scientist intuition has made her as a constant oasis of ideas and passions in science, which exceptionally inspire and enrich my growth as a student, a researcher and a scientist want to be. I am indebted to her more than she knows.

I gratefully thank to head of the Department of Biosciences Prof. S. P. Singh for providing me all the necessary laboratory facilities. I am also very indebted to other faculty members and nonteaching staff of Department of Biosciences for their help in direct or indirect manner.

It is a pleasure to pay tribute also to the sample collaborators. I would like to thank to the Institute of Microbial Technology (IMTECH), Chandigarh, India, for providing me the fungal samples. I would also acknowledge to Vimal Research Society for Agro-biotech and Cosmic Powers (VIRSACO), Rajkot, Gujarat, India, for providing me the samples of essential oils. I convey special acknowledgement to Dr. Manish L. Vekaria (Managing Director, VIRSACO) and Dr. Virendra J. Patel (Director, VIRSACO) for endowing me with all the necessary laboratory facilities and instrument facilities at VIRSACO. I wish to acknowledge University Grant Commission (UGC), New Delhi, India for financial support as meritorious scholar, also thanks to Centre For advanced Studies in Plant Biotechnology & Genetic Engineering (CPBGE), state Government of Gujarat for providing lab facility.

I convey special thanks to my rabbits who might have suffered a bit to accomplish my work; I also thank Jivanbhai for taking care of my rabbits. I thank Satyendarbhai, Govindbhai and Joshibhai for their help. I am also thankful to Saurabhbhai, Nitheshbhai and Purohitbhai for their all-time helping hand.

Collective and individual acknowledgments are also owed to my colleagues Mr. Lalit Chariya whose present somehow perpetually refreshed, helpful, and memorable. I am also thankful to Ms. Kiran Chudasama for her company and the kind help. My special thanks go to my senior colleagues Dr. Snehal Bhagtaria, Dr. Vaishali Pawar, Dr. Rita Chudasama, Dr. Kunjal Bhatt, Dr. Madhavi Joshi, Dr. Rohan Pandya and Mr. Anil Nakum for their guidance and support. I am also grateful to all my junior colleagues especially Mr. Viral Mandaliya for all his skilful assistance and carefully reading my thesis script. I thank to Ms. Vibhuti Jhala and Ms. Dipika Kalariya for managing bioinformatics software for my work. I also thank to Bhavisha Sheth, Khyati Bhatt, Pooja Patel, Purvi Rakhasiya, Kausal Yadav and Smita Girnari for their assistant and help.

It is a pleasure to express my gratitude wholeheartedly to my friends Jalpa and her family, Sarita, Shreya, Dr. Vivek, Ravi, Dhawal, Jahnavi and Dr. Rajan. They have helped me stay sane through these years. Their support, understanding and care helped me to stay focused on my work. I greatly value their friendship and I deeply appreciate their belief in me. I am also grateful to all the research scholar‘s of Department of Biosciences, who had helped me during my research period.

Where would I be without my family? My parents deserve special mention for their inseparable support and prayers. It was not possible to complete this work without their support. My Father, R.B. Maharshi, in the first place is the person who put the fundament my learning character, showing me the joy of intellectual pursuit ever since I was a child. Words fail me to express my gratitude towards my Mother, Bhagyawanti Maharshi is the one who sincerely raised me with her caring and gently love. She always gives me strength, courage and managed my time. I am thankful to Ashish Maharshi and Avdhendra Maharshi for being supportive and caring siblings. I also thankful to my cousin Manoj Prabhakar for his kind support at each and every step of my life.

Finally, I would like to thank everybody who was important to the successful realization of thesis, as well as expressing my apology that I could not mention personally one by one.

Anjisha Maharshi Index

No. Chapter Page No.

1 General Introduction 1-13

Isolation, Growth and Identification of 2 Plant Pathogenic Fungi 14-33

3 Phytohormone Production by Plant Pathogenic Fungi

3A Indole Acetic Acid Production 34-51

Abscisic Acid and Gibberellic Acid 3B Production 52-62

4 Biocontrol Approach

Biological approach by Plant extract 4A and Essential Oils 63-77

Immunological Approach Against Cell 4B Wall Degrading Enzymes 78-90

5 Summary 91-102

References 103-173

Fungi form a large and heterogeneous eukaryotic group of living organisms characterized by their lack of photosynthetic pigment and their chitinous cell wall. It has been estimated that the fungal kingdom contains more than 1.5 million species, but only around 100,000 have so far been described, with yeast, mold, and mushroom being the most familiar (Hawksworth, 1991). Although the majority of fungal species are saprophytes, a number of them are parasitics, in order to complete their biological cycle in animals or plants. With around 15,000 of fungi, the majority was belonging to the Ascomycetes and Basidiomycetes, causing disease in plants (Reddy, 2001).

Fungal diseases are, in nature, more the exception than the rule. Thus, only a limited number of fungal species are able to penetrate and invade host tissues, avoiding recognition and plant defense responses, in order to obtain nutrients from them, causing disease and sometimes host death. In agriculture, annual crop losses due to pre- and post harvest fungal diseases exceed 200 billion euros, and in the United Stated alone, over $600 million are annually spent on fungicides (Gonzalez-Fernandez et al., 2010). Fungal pathogens have complicated life cycles, with both asexual and sexual reproduction and stages involving the formation of different infective, vegetative, and reproductive structures (Glawe, 2008).

Current taxonomic identification of filamentous fungi is based on micro- and macro-morphological characteristics, such as cultural morphologies including colony and color characteristics on specific culture media, the size, shape, and development of sexual and asexual spores and spore-forming structures, and/or physiological characteristics such as the ability to utilize various compounds as nitrogen and carbon sources. The advent of molecular methods has supplemented traditional taxonomic methods with DNA-based tools with which to examine phylogenetics and systematics of fungi (Bowman et al., 1992; Bruns et al., 1992; Guerra-García et al., 2008).

Accurate identification and early detection of pathogens is a crucial step in health care, agriculture and environmental monitoring including the fight against bioterrorism (Schaad and Frederick, 2002; Ivnitski et al., 2003; 1║ P a g e

Schaad et al., 2003) plant disease management. The failure to adequately identify and detect plant pathogens using conventional, culture based morphological techniques has led to the development of nucleic acid based molecular approaches (López et al., 2003a; Lievens et al., 2005). Molecular diagnostic began to develop a real momentum after the introduction of polymerase chain reaction (PCR) in the mid 1980s and the first PCR based detection of a pathogen in diseased plants was published in the beginning of 1990s (Rasmussen and Wulff, 1991).

In the last two decades, DNA marker technologies have been revolutionized the plant pathogen genomic analysis and have been extensively employed in many fields of molecular plant pathology (Bridge et al., 2003). Molecular markers offer also the possibility of faster and accurate identification and early detection of plant pathogen (Schaad et al., 2003; Michailides et al., 2005; Ma and Michailides, 2007). On the other hand, with the advances of molecular biology many complex questions could be answered such as the sources of inoculum, changes in their population‟s structures and the population dynamics of the disease they cause. Currently, numerous diagnostics laboratories are adopting molecular methods for plant pathogen detection (Lopez et al., 2003b; Schaad et al., 2003; Hernandez-Delgado, 2009).

Among others important applications of these technologies which are increasingly developed as resourceful tools for quickly and sometimes cheaply assessing diverse aspects of plant pathogen genomes. These include genetic variation characterization, genome fingerprinting, gene mapping and tagging, genome evolution analysis, population genetic diversity, taxonomy and phylogeny of plant pathogen taxa. Ever since the introduction of both the DNA polymerase and the primer sequences (Mullis and Faloona, 1987; Mullis, 1990), the use of PCR in research laboratories has increased tremendously. Advances in molecular biology techniques have provided the basis for virtually unlimited numbers DNA markers. Currently, several DNA markers systems are now commune in the study of plant pathology (Lopez et al., 2003b; Schaad et al., 2003; Ippolito et al., 2004; Singh and Hughes, 2006; Benali et al., 2011). 2║ P a g e

To date an increasing number of diagnostic laboratories is adapting molecular methods for routine detection of pathogens. Besides conventional PCR techniques advanced methodologies such as the second generation PCR known as the real time PCR and microarrays which allow unlimited multiplexing capability have the potential to bring pathogen detection to a new and improved level of efficiency and reliability (Mumford et al., 2006).

PCR assays for the detection of fungal pathogens are an appealing approach due to their potential for rapid, sensitive, and accurate diagnosis of fungal infections. rRNA genes are particularly attractive targets because multiple copies are present in each genome and the genes have conserved regions for designing broad-range primers and have more variable regions for identifying fungi. Most studies have focused on regions within the 18s rRNA gene (Einsele et al., 1997; van Burik et al., 1998a), internal transcribed spacers (ITS1 and ITS2), the 5.8s rRNA gene, and the 5‟ end of the 28s rRNA gene (D1-D2 hypervariable region) for developing broad-range PCR primers targeting human fungal pathogens. The remainder of the 28s rRNA gene spanning nearly 2,900 bp remains largely unexplored. Sequences of the 28s rRNA gene beyond the D1-D2 hyper variable region for many important human fungal pathogens are not available or are undefined in public databases (Khot et al., 2009).

The 28s ribosomal gene was selected as a detection target because its large size was expected to provide adequate species-specific differences to distinguish closely related organisms. Additionally, ribosomal genes in fungi are present at as many as 100 or more copies per genome (Maleszka and Clark-Walker, 1993; Sandhu et al., 1995), and this would provide good detection sensitivity. Usually, a conventional PCR-based method consists in the design of PCR primers that will only originate an amplification product in the presence of DNA from the target species. The process of designing species-specific primers is now straight forward due to the vast number of genomic sequences available and software programs that assists in primer designing (Pereira et al., 2008). In the present study, the primer designed for specific region of 28s rDNA was used to amplify the various plant pathogens. 3║ P a g e

According to the International Code of Botanical Nomenclature (ICBN), the binomial Macrophomina phaseolina is the valid name for the pycnidial stage of Rhizoctonia bataticola (Ashby, 1927; Dhingra and Sinclair, 1978) with the synonyms i.e. Botryodiplodia phaseoli (Maubl.) Thirum, Dothiorella cajani Syd., P. Syd. and E. J. Butl., Manophoma cajani Syd., P. Syd. and E. J. Butl., Macrophomina phaseoli Maubl., Rhizoctonia lamellifera Small, Sclerotium bataticola Taubenh (Prabhu, 2009). Goidanich (1947) examined the original material of M. phaseoli collected by Tassi and renamed it as Macrophomina phaseolina (Tassi) Goid. It has a world-wide distribution and has been reported on wide range of hosts causing charcoal rot (Sergeeva et al., 2005). They are sterile and not forming conidia.

The genus Antrodia P. Karst., in a wide sense includes species with a dimitic hyphal system, clamped generative hyphae and causing a brown rot (Bernicchia et al., 2010). Antrodia P. Karst. is a polypore genus with more than 40 species causing brown rot of wood (Yu et al., 2010). Postia placenta was also cause brown rot of wood. Postia placenta (Fr.) Cooke is closely associated with wood strength loss. Brown-rot fungi can partially decompose lignin by demethoxylating it, but preferentially they degrade celluloses and hemicelluloses without removing the surrounding lignin. There is little outward evidence of degradation in the early stages of brown-rot decay, although mass losses can be between 5 and 10% in brown rotted wood (Goodell, 2003).

Antrodia Karsten is a cosmopolitan genus with the characteristics of resupinate to effused-reflexed basidiocarp, dimitic hyphal system with clamped generative hyphae, white or pale context, oblong to ellipsoid, hyaline, thinwalled, smooth and non-amyloid spores, and brown rot. They selectively degrade cellulose and hemicelluloses from wood and significantly weaken its structural properties (Jang et al., 2011). Antrodia sitchensis on the basis of the shape and size of its basidia and basidiospores, was firstly reported in America as Poria sitchensis by Baxter (1938). A. sitchensis has been reported in Europe and North America as well as in Asia, particularly in China (Dai et al., 2002). The macroscopic features of A. sitchensis were similar to A. xantha 4║ P a g e

was reported by some researchers (Jung, 1994; Lee et al., 2002). Some Antrodia species have been reported to cause decay in wood products from playground (Kim et al., 2005) as well as in building materials (Schmidt, 2006).

Fusarium sp. is widely distributed in soil, colonizing living plant parts or plant residues within the soil or adjacent to the soil surface. They cause root, stem and ear rot, with severe reductions in crop yield, often estimated at between 10% and 40% (Bottalico and Perrone, 2002). In addition, certain strains are capable of producing mycotoxins which can be formed in pre-harvest infected plants, or in stored grains (Logrieco et al., 2002). Fusarium solani is a phytopathogenic fungus that causes potato dry rot (Olivieri et al., 2002). Fusarium sp. causes Fusarium wilt of many plants (Chakrabarti et al., 2001; Wu et al., 2008) it also caused head blight of small-grain cereals (Parry et al., 1995; Menniti et al., 2003).

Wilt caused by Fusarium udum Butler is disease of pigeonpea (Joshi, 2000). Fusarium wilt in solanaceous crops is caused by several different types of the fungus Fusarium oxysporum. These are: F. oxysporum f. sp. lycopersici (tomato), F. oxysporum f. sp. melongenae (eggplant) and F. oxysporum var. vasinfectum (pepper). Fusarium wilt in potato is caused by a complex of up to four different Fusarium sp. All of the Fusarium wilt pathogens are generally specific to their hosts and are soilborne (Miller et al., 2010). Fusarium is one of the common soil inhabiting plant pathogenic fungus which causes diseases such as wilt of brinjal, pigeon pea, guava, gram, tomato etc. Several others species of this genus are responsible for huge loses to their respective host crop (Joseph et al., 2008). Fusarium solani f. sp. melongenae as causal agent of brinjal wilt (Najar et al., 2011). Fusarium solani is reported for Guava wilt (Misra, 2006; Gupta et al., 2009).

Curvularia leaf spot affects many species of grasses worldwide and is commonly caused by Curvularia eragrostidis, C. geniculata, C. intermedia, C. inaequalis, C. lunata, C. pallescens and C. protuberate or C. trifolii (Huang et al., 2005). A relationship between human infection and other crops has been noted which was caused by C. lunata (Yau et al., 1994; Wilhelmus and Jones, 5║ P a g e

2001). Curvularia eragrostidis, a causal organism of head blight disease caused in Digitaria sanguinalis (Wang et al., 2006).

Alternaria solani, the causal agent of early blight disease and Fusarium oxysporum f. sp. lycopersici, the causal agent of wilt disease in tomato (Hassanein et al., 2008). Black spot caused by Alternaria alternata (Fr.) Keissler are among the main diseases found on pears and apples in storage (Maouni et al., 2007). Alternaria alternata can have a certain pathogenic effect; it has been recorded as a saprophytic or a weak pathogen causing so- called "indefinite or opportunistic disease" on a number of crops (Nishimura, 1980; Ibraheem et al., 1987; Abbas et al., 1995; Fawzi et al., 2009).

The Gibberella fujikuroi species complex includes important fungal pathogens of various crops, such as maize, rice, barley, sugarcane, pine, mango, pineapple, sorghum, and many more (Leslie, 1999). Moreover, members of these mating populations can be found preferentially on different host plants (Leslie and Plattner, 1991; Zeller et al., 2003) and differ in their ability to produce secondary metabolites, such as fusaric acid (Bacon et al., 1996), moniliformin (Leslie et al., 1996; Desjardins et al., 2000), fusarins (Wiebe and Bjeldanes, 1981; Song et al., 2004), and gibberellins (GAs) (El-Bahrawi, 1977; Tudzynski and Holter, 1998; Desjardins et al., 2000).

Microscopic fungi of the genus Aspergillus are typical microbial soil saprophytes. A. niger is a filamentous fungus with prominent applications in biotechnological processes. Products of the fungus have the generally recognized as safe (GRAS) status allowing them to be used as food additives. Important products that are made by fermentation of A. niger are organic acids, such as citric acid and gluconic acid. Citric acid is used for acidification and to enhance taste and flavour in food (Ruijter et al., 2000).

In the second part of the study some fungi were used to evaluate hormone production from them. Standardization of auxin production was carried out in A. niger with the help of tryptophan substrate. A detailed study was done on aldehyde dehydrogenase gene responsible for auxin production their 6║ P a g e

amplification and with help of some bioinformatics tool its protein prediction was also carried out. ABA and GA production was also seen in Macrophomina phaseolina and Fusarium moniliforme, respectively.

Plant hormones are a group of naturally occurring, organic substances which influence physiological processes at low concentrations. The processes influenced consist mainly of growth, differentiation and development, although other processes, such as stomatal movement, may also be affected. Plant hormones have also been referred to as phytohormones (Davies, 1995; Iten et al., 1999). A phytohormone is an organic substance synthesized in defined organs of the plant that can be translocated to other sites, where it triggers specific biochemical, physiological, and morphological responses. However, phytohormones are also active in tissues where they are produced. In addition, numerous soil bacteria and fungi also produce phytohormones. The commonly recognized classes of phytohormones, regarded as the “classical five”, are: the Auxins, Gibberellins, Cytokinins, Abscisic acid, and Ethylene (Baca and Elmerich, 2003).

Many plant-associated microorganisms are themselves capable of synthesizing phytohormones, which are necessary as mediators in communications between the plant host and its microflora. The ability of microorganisms to synthesize phytohormones may be viewed, first of all, as a sign of their pathogenicity, because many pathogens produce growth regulators in amounts exceeding the levels necessary for the plants. Plant pathogen interaction increases the hormonal level. Hypersynthesis of phytohormones by pathogenic microorganisms disrupts the hormonal balance of the plants, causing the appearance of a variety of diseases, including gall formation; boil smut, hernia, and vascular wilt (Shimada et al., 2000; Chung et al., 2003; Yurekli et al., 2003). Plant hormones, as we all know, are important substances for growth and development of plants, but they are also important factors for the interaction of plants with microorganisms during pathogenesis or symbiosis (Ludwig-Muller, 2004).

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Many industries use conventional methods to produce economical substances (enzymes, antibiotics, plant growth hormones, steroids and organic acids). In recent years, microorganisms have been immobilized in both laboratory and industrial processes. The production of patulin and penicilin, gibberellic acid and bikaverin by immobilized fungi has been extensively examined (Unyayar et al., 2000). However, production of IAA has not been examined yet.

The principle auxin in plants is indole-3-acetic acid (IAA). IAA is an important metabolite in plant growth from embryogenesis to senescence, and developments in biotechnology have led to an increased awareness of the importance of industrial fungi for the production of plant growth hormones (Berry, 1988). These fungi include white-rot species widely used in biotechnological and biochemical applications. IAA and ABA are synthesized in plants and fungi (Blakesley et al., 1991; Lopez-Carbonell et al., 1994; Unyayar et al., 1997).

Gibberellins are tetracyclic diterpenoid acids that are involved in a number of developmental and physiological processes in plants (Davies, 1995; Crozier et al., 2000). These processes include seed germination, seedling emergence, stem and leaf growth, floral induction and flower and fruit growth (Pharis and King, 1985; King and Evans, 2003; Sponsel, 2003). Gibberellins are also implicated in promotion of root growth, root hair abundance, inhibition of floral bud differentiation in woody angiosperms, regulation of vegetative and reproductive bud dormancy and delay of senescence in many organs of a range of plant species (Tanimoto, 1987; Bottini and Luna, 1993; Fulchieri et al., 1993; Reinoso et al., 2002; Bottini et al., 2004).

The scientific origins of abscisic acid (ABA) have been traced to several independent investigations in the late 1940s, but it was only in the 1960s that ABA was isolated and identified (Leung and Giraudat, 1998). ABA is a small, lipophilic plant hormone, originally believed to be involved solely in promoting seed maturation and response to water stress. ABA levels accumulate in the seed by mid to late embryogenesis, where it controls many processes, such

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as embryo morphogenesis, storage-protein synthesis, desiccation tolerance, and the onset and maintenance of dormancy (Giraudat et al., 1994).

Fungal pathogens of plants have developed many mechanisms to evade or overcome healthy host plant defenses. Although plants do not have circulating or phagocytic cells, their cells have a thick, complex wall that acts as a barrier to invasion. Plants display innate pathogen-specific resistance, genetically controlled via resistance genes. Additionally, plants display inducible systemic acquired resistance, which occurs when previous exposure to a pathogen activates signaling pathways acting via molecules such as jasmonate, ethylene, and salicylic acid (Sexton and Howlett, 2006). These small and sometimes volatile molecules spread throughout the plant or even the plant population. This triggers responses such as the expression of “pathogenesis related proteins,” including chitinases or glucanases, which can lead to the increased resistance of the whole plant against a subsequent pathogen attack (Pieterse and van Loon, 2004).

In the course of evolution, plants have developed chemical defense mechanisms against potential pathogens and pests. Society‟s dependence on intensive agriculture and horticulture for food production has accentuated the need to reduce crop losses. Environmental considerations have highlighted the requirement for sustainable solutions in agriculture and consumer pressures for green alternatives have accompanied a boom in the organic farming sector. This has awakened new interest in natural products as a source for novel industrial plant protection strategies (Slusarenko et al., 2008).

As long as agriculture exists, farmers have to battle against pathogens which infect crop plants and thereby cause considerable losses of yield and quality. To control plant diseases and pests, several strategies have been developed. These strategies are based on genetic, chemical, biological and cultural methods (Honee, 1999). The general strategy of biological control is to use one living organism to control another. The control agents may be antagonistic microorganisms or even natural plant and animal-derived compounds (Pal and Gardener, 2006; Chanchaichaovivat et al., 2007). 9║ P a g e

Recently, biological control has been developed as an alternative to synthetic fungicide treatment.

Every year a lot of crop damage is caused by various diseases and among them fungal diseases are very common. Although the use of synthetic fungicides in plant disease control have shown its fruitfulness in the improvement of agriculture, several of these have been found to display side- effects in the form of carcinogenicity, detrimental effects and other residual toxicities. The alternative choice therefore would be the use of botanical fungicides, which are advocated to be largely non-phytotoxic, systematic and easily biodegradable in nature (Begum et al., 2007). It is believed that antimicrobial agents are present in higher plants. Some recent researches on antifungal activity of extracts of several higher plants have indicated the possibility of their exploitation as natural antifungal agents for control of plant diseases (Miah et al., 1990; Anwar et al., 1994; Qureshi et al., 1997; Begum et al., 2007).

The plant kingdom represents an enormous reservoir of antibiotic principles that are distributed widely. These compounds are varying greatly in their potency and distribution within a plant (Grayer and Harbone, 1994). There are 2600 plant species of which more than 700 are noted for their uses as medicinal herbs (Ali-Shtayeh and Abu, 1999). In response to stimuli from microbial infections, plant produces antimicrobial phytochemicals known as phytoalexine for their defense (Dixon et al., 1983). Increasing emergence of resistance to the currently available antibiotic have necessitated continued search for new antimicrobial compounds having novel mechanism of action. A wide spectrum of organisms is being screened in search of useful antimicrobials (Srinivasan et al., 2009).

Agavaceae native to America is distributed in arid to semiarid environments; the majority of species are found in Mexico. Currently nine genera and ca. 300 species are recognized in the family (Bogler et al., 2006). Members of Agavaceae display a wide array of ecological, reproductive, and morphological adaptations to arid environments. The family has been 10║ P a g e

important for people living in the Americas since prehistoric times, with various species providing clothes, rope, food, and beverages (both nonalcoholic and alcoholic) to humans (Rocha et al., 2006). Agaves have been used by humans since pre-Columbian times and are still widely used as sources of food, beverages, fibers, construction materials, and natural medicines (Callen, 1965; Felger and Moser, 1970; Arizaga and Ezcurra, 2002).

Essential oils (EOs) have been shown to possess antibacterial, antifungal, antiviral insecticidal and antioxidant properties (Burt, 2004; Kordali et al., 2005). Some oils have been used in cancer treatment (Sylvestre et al., 2006). Some other oils have been used in food preservation (Faid et al., 1995) and aromatherapy (Buttner et al., 1996). EOs are rich source of biologically active compounds. There has been an increased interest in looking at antimicrobial properties of extracts from aromatic plants particularly essential oils (Milhau et al., 1997). Therefore, it is reasonable to expect a variety of plant compounds in these oils with specific as well as general antimicrobial activity and antibiotic potential.

EOs (also called volatile oils) are aromatic oily liquids obtained from plant materials (flowers, buds, seeds, leaves, twigs, bark, herbs, wood, fruits and roots). They can be obtained by expression, fermentation or extraction but the method of steam distillation is most commonly used for commercial production (Kokate et al., 2008). An estimated 3000 EOs are known, of which 300 are commercially important in fragrance market. EOs are complex mixers comprising many single compounds. Chemically they are derived from terpenes and their oxygenated compounds (Bakkali et al., 2008).

Another, approach was done in biocontrol as cell wall degrading enzyme inhibition with the help of polyclonal antibody produced against fungal spore. Antigenic crossreactivities have been detected in various proteins that are related to one another. The implications of this cross-reactivity of related protein molecules could be very significant. Certain enzymes could be

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inhibited by anti-hapten antibodies when the enzyme molecules were covalently bound to the corresponding hapten (Katiyar and Porter, 1983).

Considering the aforesaid following objectives were resolute and worked out for this thesis:

1. To collect the fungi from infected plant materials. Primary identification, culture maintenances and preservation, Potato Dextrose Agar (PDA) media at 25 - 28 ºC and in distilled water at room temperature. 2. Studying growth of these fungi on define Murashige and Skoog (MS) media and their molecular identification which includes; A. DNA isolation and standardizations for more quality and quantity of DNA B. Universal primer designing for specific region of 28s fragment of fungal DNA C. Identification of amplified sequence of fungi by these primers with the help of some bioinformatics tools and data bases 3. To evaluate phytohormone production by plant pathogenic fungi (I) Auxin Production A. Identification of the fungi using specific primer of 18s region B. Standardizations of optimum production of IAA by this fungus in concern with various factors like media, pH, substrate concentration, incubation time and temperature. C. Designing primer sets for aldehyde 3- dehydrogenase for this fungus. D. Standardization of polymerase chain reaction to amplify aldehyde 3- dehydrogenase gene and sequenced which were then submitted to NCBI database E. Its protein structure was then predicted by homology modeling F. A mutational study with the help of bioinformatics tool was also carried out.

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(II) Abscisic acid and Gibberellic acid production A. Abscisic acid production by Macrophomina phaseolina in define media was examine B. Gibberellic acid production by Fusarium moniliforme was also check in define media 4. Bio-control approach against fungal pathogens (I) Biological approach by plant extracts and Essential oils A. To evaluate antifungal activity of plant extracts of Agave sp. B. Study of 65 Essential oils against Fusarium moniliforme (II) Immunological and Enzymatic approach A. Raising of antibody against fungal spore B. Purification and Separation of IgG C. Studying the inhibition of cell wall degrading enzymes of fungi by this antibody.

It is hoped that this study may help in understanding the mechanism(s) of pathogenecity and biological control of these fungi for better anticipated agronomical practices.

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INTRODUCTION

Fungi rank second only to insects as a cause of plant diseases, which result in heavy loss of plant products. Pathogenic fungi alone cause nearly 20% reductions in the yield of major food and cash crops (Agrios, 2000). One-third global agricultural production is reportedly destroyed each year by different pests and diseases. To avoid the implication of yield losses due to plant diseases, variety of control measures presently are in use (Bajwa et al., 2003). They are one of the principal causes of reduced quality in stored grains and foods. Many fungal species produce mycotoxins, secondary metabolites which are highly toxic to animals and humans (Shapira et al., 1997).

Microorganisms need nutrients as a source of energy and certain environmental conditions in order to grow and reproduce. In the environment, microbes have adapted to the habitats most suitable for their needs. In laboratory, however these requirements must be met by a culture medium. To culture fungi in the laboratory, it is necessary to furnish in the medium all essential nutrients for the synthesis of protoplasm (Davis, 1978; Griffin, 1966). First attempt to obtain laboratory cultures of fungi was made by the great Italian botanist Micheli (1729). He could succeed in growing three different molds viz., Mucor, Aspergillus and Botrytis on freshly cut surface of melon and pear (Rani et al., 2007).

To cure the diseases caused by them or to inhibit the growth of pathogen there is a prior need to identify them. The fungi are the important plant pathogens and continue to be the focus of extensive research with a wide variety of methodologies. Enhancements in microscopy techniques have increased our ability to visualize the intimate interaction of fungi and their host plants. However, in some circumstances, phenotype-based identification is not helpful. If the isolate displays atypical morphology, fails to sporulate, requires lengthy incubation or incubation on specialized media not available to the laboratory, the lack of observable reproductive structures can potentially increase the time to the reporting of an inconclusive result, or if the phenotypic

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results are nonspecific or confusing, the isolate may be a candidate for DNA- based identification (Balajee et al., 2007).

Therefore, there is a clear need for alternative methods for the identification of fungi that do not produce morphologically distinguishing features. Sequencing of the ribosomal genes has emerged as a useful diagnostic tool for the rapid detection and identification of fungi, regardless of whether morphologically distinct structures are produced (White et al., 1990; Hoorfar et al., 2004; Borman et al., 2008). One of the most common ribosomal targets for sequence identification is the internal transcribed spacer (ITS) region. This region contains two informative regions, ITS1 and ITS2, which are located between the 18s and 28s ribosomal subunits and which are separated by the 5.8s ribosomal subunit (Chen et al., 2000; Chen et al., 2001).

A second variable site within the ribosomal DNA (rDNA) cluster, called the D1/D2 region, can also be amplified from a broad spectrum of fungi with primers NL-1 and NL-4, although it is usually less variable than the ITS region (Kurtzman and Robnett, 1997). The D1/D2 region is located toward the 5‟ end of the large ribosomal subunit (26s or 28s) and overlaps the ITS region at the ITS-4/NL-1 primer site. The combination of conserved and variable regions offers great flexibility for polymerase chain reaction (PCR) sensitivity and specificity. The conserved sequences at the flanking ends of the D1/D2 and ITS regions provide universal PCR priming sites, while the variable internal regions provide species-specific sequences in many cases (Bruns et al., 1991; Kurtzman and Robnett, 1997; Balajee et al., 2007; Romanelli et al., 2010).

The PCR is a powerful method with widespread applications in molecular biology. This enzymatic reaction allows in vitro amplification of specific DNA fragments from complex DNA samples and can generate microgram quantities of target DNA. Any nucleic acid sequence can be cloned, analyzed or modified, and even rare sequences can be detected by PCR amplification (Bruns et al., 1990). One of the first applications of PCR in mycology was described in 1990 by White et al. and concerned the amplification and direct 15║ P a g e

sequencing of ribosomal rDNA to establish the taxonomic and phylogenetic relationships of fungi. rDNA sequences are often used for taxonomic and phylogenetic studies because they are found universally in living cells in which they have an important function; thus, their evolution might reflect the evolution of the whole genome. These sequences also contain both variable and conserved regions, allowing the comparison and discrimination of organisms at different taxonomic levels. The nuclear rDNA in fungi is organized as an rDNA unit which is tandemly repeated. One unit includes three rRNA genes: the small nuclear (18s-like) rRNA, the 5.8s rRNA, and the large (28s-like) rRNA genes. In one unit, the genes are separated by two internal transcribed spacers (ITS1 and ITS2) and two rDNA units are separated by the intergenic spacer (IGS). The last rRNA gene (5s) may or may not be within the repeated unit, depending on the fungal taxa (Edel, 1998). The 18s rDNA evolved relatively slowly and is useful for comparing distantly related organisms (Shou et al., 2011).

The nuclear ribosomal coding cistron (rDNA) has been widely utilized for detecting variation among isolates of a fungus. PCR is an attractive method for the detection of fungi. Extensive applications have been found for PCR in many fields of mycology, including fungal genetics and systematics (Rossman and Palm-Hernández, 2008), ecology and soil microbiology, (Bej and Mahbubani, 1992), medical mycology (Mirhendi et al., 2001; Cruzado et al., 2004), plant pathology (Walsh et al., 1995; Martin et al., 2000) fungal biotechnology and many others. Moreover, it is certain that fungal studies will continue to progress with PCR, as numerous improvements, modifications and new applications are regularly reported. PCR can be used to detect groups of strains, pathotypes, species or higher taxa, provided that specific oligonucleotide primers for these taxa are available (Bagyalakshmi et al., 2008; Balau and Faretra, 2010).

Thus, the development of PCR-based detection procedures requires knowledge of sequences of at least a part of the target DNA region in order to 16║ P a g e

design specific primers. The principle of these methods is the alignment of the sequences from target and non-target organisms and the selection of primers with mismatches to the non-target organisms but sufficient homology for efficient priming and amplification of all target organisms (Dieffenbach et al., 1993).

PCR technology can directly detect the presence of fungi with high level of sensitivity and specificity (Embong et al., 2008; Henson and French, 1993; McCartney et al., 2003). Universal primers common to all fungi have been used as a promising approach in plant pathology (Volossiouk et al., 2010). Various techniques have been reported to separate different fungi detected by universal primers, including restriction fragment length polymorphism (Hopfer et al., 1993; Kemker et al., 1991), The random amplified length polymorphism (RFLPs) belongs to the widely used hybridization developed in Humans projects in 1980s (Botstein et al., 1980), and thereafter in plant pathology research (Leung et al., 1993; McDonald, 1997). The other markers were also studied in plant pathology like, Single-strand conformational polymorphism (SSCP), Random amplified polymorphism DNA (RAPD), Amplified Fragment Length Polymorphism (AFLP), Simple Sequence Repeats (SSR), Internal Transcribed Sequence (ITS) and Single Nucleotide Polymorphism (SNP) (Mirhendi et al., 2001; Benali et al., 2011).

Depending on the type and combination of nutrients, different categories of media can be made but in this study Murashige and Skoog (MS) (1962) liquid media was used for growth and development of fungi. There are several methods to identify the fungi like microscopical, chemical, immunological and molecular. From all of these molecular means DNA sequencing based identification is more reliable. Considering all above factor the present study was formulate to standardize DNA isolation protocol and amplify the 28s specific rDNA region of different plant pathogenic fungi by primer designed with the help of bioinformatics tool. These products were further purified and sequencing and post sequencing analysis was carried out.

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MATERIALS AND METHODS

Isolation of pathogens Infected plant materials were taken and for surface sterilization they were put under running tap water for 1-2 h in jar. Then it was washed with 0.1% HgCl2 for 10 min. The sample was washed with sterilized distilled water 3 to 4 times for removal the HgCl2. With the help of pre-sterilized scalpel and forceps infected part of plant material was cut. The cutting part was inoculated on Potato Dextrose Agar (PDA) plate in aseptic condition and incubated at 28 ± 2ºC for 24-48 h for growth of pathogens. After their adequate growth, isolation for pure culture was carried out. Pure culture of pathogen(s) was incubated at 28 ± 2ºC for seven days. Then with the help of (1cm diameter) borer fungal disc were transfer on other PDA plate for radial growth to obtain the same growth pattern. Further this pure culture was preserved for experimentation.

Culture preservation and maintenance All the cultures (isolated and existing in lab or isolated earlier in the lab) were preserved on PDA slant at 4ºC as well as in distilled water at room temperature. When needed they were grown on PDA plate at 28 ± 2ºC for seven days and the growing hyphae were used. List of the isolates were given in Table-2.1.

Preparation of inoculums First the fungi were revived from old culture on PDA plates incubated for seven days at 28 ± 2°C. Then with sterile cup borer (1cm diameter) disc was cut from the peripheral region of the plate and transfer to another plate for radial growth. A disc (1cm diameter) of fungal hyphae from these seven day old cultures was used as inoculum.

Media and culture condition A define media MS (1962) with 2% sucrose supplement was prepared and 50 ml of it filled in each of the 250 ml conical flask. The media was autoclaved for 15 min and allowed to cool to room temperature. The inoculum was added to each flask and maintained in a stationary condition at 28 ± 2°C for their 18║ P a g e

growth and development. For growth analysis fresh and dry weights and pH were measured with ± standard deviation (SD) at every 24 h intervals. For each experimental data, three replicates were taken.

Morphological and microscopical identification Morphological identification was carried out by observing the growth pattern, color of hyphae and their structure (Table-2.1). The microscopical examination was carried out by lactophenol cotton blue (LPCB) wet mount (Thomas et al., 1991) preparation. It is the most widely used method of staining and observing fungi and is simple to prepare. The preparation has three components: phenol, which will kill any live organisms; lactic acid which preserves fungal structures, and cotton blue which stains the chitin in the fungal cell walls. LPCB mount was carried out to observe the structure of fungi.

Molecular Identification DNA preparation For culturing the fungi Potato Dextrose Broth were used. 30-60 mg dry material or a corresponding amount of fresh weight were sufficient for micro- extraction of DNA by using Moller et al. (1992) method with some variation depending upon fungal strain.

Dry mycelium was crushed with fine sand in a mortar or a fresh material in liquid nitrogen. Powdered mycelium were placed into a micro-tube with 500 l TES (100 mM Tris, pH 8.0, 10 mM EDTA, 2% SDS); 50-100 g Proteinase K was added from an appropriate stock solution and incubated for 30 min (minimum) up to 1 h at 55-60°C with occasional gentle mixing. Then tubes were centrifuged on 12000 xg for 20 min at 4°C. The centrifugation time and force was same in all the following steps. The supernatant was transferred and 140 l of 5 M NaCl and 65 l of 10% CTAB were added. It was incubated for 10 min at 60-65°C and centrifuged.

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Supernatant was extracted twicely first with 700 l of Phenol: Chloroform: Iso amylalcohol (PCI) (25:24:1) and then with 700 l of Chloroform: Iso amylalcohol (CI) (24:1). In these steps tubes were incubated for 30 min at 0°C then centrifuged. In the supernatant 225 l 5 M NH4Ac was added, placed on ice for approximately 30 min with gentle mixing (the longer the better) then centrifuged to the supernatant 2-4 l of RNase was added (as required) and incubated for 30 min at 0°C and then 30 min at room temperature, then it was centrifuged.

To pellet out DNA from supernatant, 510 l of isopropanol was added, incubated for overnight and centrifuged. DNA pellet was washed (at 8000 xg for 15 min) twice with cold 70% ethanol, air dry and dissolves in 100 l TE (10mM Tris, 1mM EDTA).

DNA concentration and purity The DNA concentration was determined at 260 nm and purity was determined by calculating the ratio of absorbance at 260 nm to that of 280 nm (Matasyoh et al., 2008) using the Quant microplate reader, Bio-Tek instruments incorporation, USA.

DNA concentration was determined using following two equations. Water has virtually no absorbance between 200 and 900 nm, but it has a small but significant peak at 977 nm. The microplate reader determines the pathlength of the sample by measuring the absorbance of the water in the microplate sample at 977 nm. Blanks that value using a reference wavelength of 900 and compares it to the absorbance of water at 1 cm. The ratio of those two values will be the pathlength of the sample (Eq. 1).

Eq. 1 As background absorption of the microplate is generally quite high in the UV range (below 340 nm), it is particularly important with UV absorbance

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measurements to blank prior to pathlength correction in order to obtain an accurate corrected value. Correction to 1 cm for DNA samples, which is determined using the absorbance at 260 nm, would need to accommodate this (Eq. 2).

Eq. 2 In terms of quantization, different nucleic acid species have different average extinction coefficients. By converting these known extinction coefficients to a specific concentration for a particular absorbance value at a defined wavelength, the nucleic acid concentration of solutions can be determined by a simple absorbance measurement. For example, the extinction coefficient for dsDNA (1 mg/ml) at 260 nm is 20 OD units; this can be rearranged to mean that a DNA sample that returns an absorbance 1 OD unit has a concentration of 50 µg/ml.

Preparation of UV transparent microplate The 96-well clear UV transparent Microplate was used. Distilled water (in triplicate), TE buffer (in triplicate) and DNA sample solution, each 100 l aliquots were pipetted into microplate wells and absorbance measurements at 260, 280, 977, and 900 nm were then made using the Quant Microplate Spectrophotometer, Bio Tek Instruments Incorporation, Winooski, VT. The subsequent path length corrected absorbance was then transformed using the appropriate conversion factor for determination of the concentration, and purity for samples in Microplate.

Fragmentation state of DNA The fragmentation state and RNA contamination of the extracted DNA was evaluated by agarose gel electrophoresis (Chaudhry et al., 1999). The DNA extracts from fungal mycelia were resolved by electrophoresis in 1.0 % agarose gel (TAE buffer system). The bands were visualized using EtBr

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(Ethidium bromide) stain under UV (Ultra violet) transilluminator (Hasibe and Abdulkadir, 2005).

Primer Designed for 28s region With the help of bioinformatics tool like primer3plus, a primer pair was designed. The primer pair oligonucleotide sequence was not presented in this thesis, and it is under IPR (Intellectual Property Rights) protection.

PCR Amplification The DNA samples of Isolate-2 to 7 were amplified using designed PCR primer set. It was carried out in 3 stages. The PCR conditions were: initial denaturation at 94ºC temperature for 10 min, followed by stage two of 45 cycle in which denaturation at 94ºC for 30s, primer annealing 48.3 ºC for 30s and extension was done at 72ºC for 60s. The final extension was done at 72ºC for 10 min then being held at 4ºC. PCR was carried out in Verity, Applied Biosystem (ABi), USA. The chemicals used in this experiment were purchased from Bangalore Genei (India).

Purification of PCR product to remove excess primer and nucleotides was performed using Qiagen‟s QIAquick PCR purification kit and 5 µl of product was checked on 2 % agarose gel by electophoresis. The remaining product was used in cycle sequencing.

Cycle Sequencing Cycle sequencing was performed by using ABi BigDye Terminatior v1.1 Cycle sequencing kit. Total volume was prepared 10 µl, as per standard protocol of ABi. The reaction mixture contain either of the primer (forward/reverse), 5x sequencing buffer, the template DNA, BigDye terminator and Mili-Q water.

Cycle sequencing was carried out in Veriti, ABi (USA). Reaction conditions were: first initial denaturation at 94ºC temperature for 10 min, followed by second stage of 45 cycles includes 96ºC temperature for 10s, 48.3ºC temperature for 5s, and 60ºC temperature for 4 min then being held at 4ºC.

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Purification of cycle sequencing products was done by Dye Ex 2.0 spin kit of Qiagen as per manufacture‟s protocol and subjected to automated sequencing and analyzed using ABi 3130 Genetic Analyzer.

Sequencing analysis The obtained sequences were BLAST to perform individual nucleotide- nucleotide searches on BLASTn algorithm of NCBI (http://www.ncbi.nlm.nih.gov/BLAST/). The outputs from the BLAST searches were sorted on the basis of the maximum identity and were recorded as they appeared without modification of genus or species names that may have been synonyms of other genus or species names in other GenBank records. Sequence-based identities with a cutoff of 86 % or greater were considered significant in this study, and the best hit was defined as the sequence with the highest maximum identity to the query sequence.

Phylogenetic analysis The phylogenetic analysis was performed (Hu et al., 2011) on CLC Main Workbench5 to construct dendrograms and for confirmation of fungal isolates based on molecular characterization.

RESULTS AND DISCUSSION

Growth and Development Surveying the nutritional capabilities of fungi is an endless task since every chemical found in living organism and a wide variety of manufactured and inorganic materials are potentially useful for satisfying the need of a fungus (Griffin, 1966).

In this experiment, seven different fungal isolates were studied for growth in the terms of fresh and dry weights. The growth patterns of these fungi are divided into (i) hyphae initiation phase or lag phase (ii) rapid growth of mycelia or log phase and (iii) declined phase (Fig. 2.1a-g). From seven different fungi studied in this experiment, accumulation of fresh weight in Isolate-1 showed lag phase up to 24 h, the log phase from 24-72 h and declined thereafter. 23║ P a g e

Accumulation in dry weight showed increasing trend up to 72 h and stabilized in later phase (Fig. 2.1a). Maximum fresh weight (2.32 ± 0.30 g) and dry weight (27 ± 0.02 mg) accumulation was recorded at 72 h. From the initial 5.6 pH, it falls to 2.3 on the second day (48 h); gradually it was decreased and observed 1.6 at seventh day (168 h) (Fig. 2.1a). Besides drastic changes in pH, the accumulation of secondary metabolites was also considered being responsible for the inhibition of fungal growth (Cooke and Whipps, 1993; He and Suzuki, 2003).

On the other hand, growth rate of Isolate-2 and Isolate-3 was found slower amongst the all fungi studied (Fig. 2.1b, 2.1c). Both the fungi showed lag phase up to 96 h, the log phase was observed from 96 h onwards up to 288 h in Isolate-2 and in case of Isolate-3 it was noted from 120-216 h and started declining thereafter. Maximum fresh weight in the log phase was measured as 2.98 ± 0.04 g and 2.00 ± 0.01 g in Isolate-2 and Isolate-3, respectively. The pH of the medium in both the isolates was gradually increased ranging from 5.8-8.5 (Fig. 2.1b, 2.1c). Dry weight was also gradually increased in Isolate-2 (2-266 mg) and Isolate-3 (1-214 mg). Overall duration of growth and development was higher i.e. double (14 days) as compared to other fungi studied.

In this experiment, Isolate-4 showed lag phase till the 48 h followed by log phase up to 120 h then declined gradually (Fig. 2.1d). Maximum fresh weight was recorded 4.5 ± 0.1 g at 120 h. However, a gradual increased in dry weight (1-181 mg) was recorded though out the growth period. The changes in pH ranged from 5.9-7.2 (Fig. 2.1d). Somewhat similar pattern of growth was also observed in Isolate-5, lag phase was parallel to Isolate-4 and in comparison to that log phase was up to 96 h (Fig. 2.1e). Maximum fresh weight was recorded 3.49 ± 0.5 g followed by decline and the dry weight was continuously increased (1-155 mg) till maturation. The pH of media increased up to 96 h from 6.1-6.4 after that it was steady at 6.1(Fig. 2.1e). On the other hand, Isolate-7 showed lag phase same as the above isolates up till 48 h subsequently followed by log phase up to 168 h then declined (Fig. 2.1g). The maximum fresh weight was measured 2.87 ± 0.2 g. The pH falls to 2.8 at 72 h 24║ P a g e

then slowly turns to neutral at 192 h (7.1). The changes in dry weight measured from 1-274 mg. In this the three isolates showed slight variation in their growth. In case of Isolate-4 and Isolate-7 the pH turns to neutral at the end of growth phases. But in the isolate-4, 5 and 7 was observed that it grew best in pH range 6.4-6.8 (Fig. 2.1d, 2.1e, 2.1g).

Isolate-6 showed lag phase of two days (48 h) followed by log phase, gradually increased in fresh weights up to 144 h (Fig. 2.1f) and declined thereafter. Maximum fresh weight was recorded 2.67 ± 0.2 g and dry weight ranged from 2-207 mg. The pH was increased from 6.5-8.3 at end of growth (Fig. 2.1f). Growth may be profoundly affected by a number of physical factors like temperature, pH, light, aeration, pressure etc. The pH range (between minimum and maximum values) is greater in fungi than it is in bacteria. Most microorganisms grow best around neutrality (pH-7) and on the other hand, fungi generally prefer slightly acid conditions for their growth (Hogg, 2005), but some species favor neutral to slightly alkaline conditions (Yamanaka, 2003). Present study revealed that in this define media among the seven fungi, five (Isolate-1, 4, 5, 6 and 7) were grew well in slightly acidic condition and their growth was completed all most in 189 h. In case of remaining two, Isolate-2 and Isolate-3, the growth period was double in compared to the rest of the fungi; they grow well in neutral to slightly alkaline condition. On the basis of this study it can be concluded that media and pH have a great impact on growth of fungi.

Primary Identification Many techniques were used for identification of fungi. Mainly fungi were identified on the basis of morphological character and molecular techniques. Morphological character like fruiting body, spore, mycelia, and growing habits were included. Amongst them, spore considered as one of the most important morphological character. There were various types of spores including oospores, zygospores, ascospores, basidiospores, conidia, sporangiospores and chlamydospores, and based on spores morphology fungi were easily classified into Mastigomycetous, Zygomycetous, Ascomycetous, Basidiomycetous and Deutromycetous fungi (Watanabe, 2002). 25║ P a g e

In the present study, primary morphological identification of seven isolates was carried based on spore structure (Table-2.1, Plate 1) (Alexopoulos and Mims, 1979). Amongst the seven isolates, three bears spores. The microscopical examination was performed to illustrate spore structure for precised conformation. Isolate-1 was identified as Aspergillus niger, Isolate-2 as Fusarium solani and isolate-6 as Curvularia intermedia.

DNA Isolation Some modification was done in DNA isolation method described by Moller et al. (1992) to achieve good purity and concentration of DNA. It was a CTAB and SDS-based extraction procedure. The cetyltrimethylammonium bromide (CTAB) protocol was first developed by Murray and Thompson in 1980 and successively published by Wagner and co-workers in 1987. The method is appropriate for the extraction and purification of DNA and is particularly suitable for the elimination of polysaccharides and polyphenolic compounds. The purity achieved by the modified protocol was ranged between 1.6-1.9 (Table-2.2). The low quality of DNA may be due to their cell wall is not properly lyses. In some DNA extraction methods for fungi, such as grinding cells frozen with liquid nitrogen using a mortar and pestle and disrupting cell walls with a probe sonicator, work well for the large-scale preparation of fungal DNA from cultures (Haugland et al., 1999, van Burik et al., 1998b; Fredricks et al., 2005).

The PCR amplification can be performed directly on various microbial cultures, but for filamentous fungi prior DNA isolation is often preferred. The DNA extraction procedure eliminates many interfering substances like mineral salts, proteins, polysaccharides, and plays an important role in ensuring consistent results. Considerable efforts have been made to improve the preparation of DNA from fungi (Lecellier and Silar, 1994; Haugland et al. 1999; Al-Samarrai and Schmid, 2000; Akada et al., 2000; Manian et al., 2001; Griffin et al., 2002; Plaza et al., 2004). In the fungal filaments the polysaccharides contain is higher so to reach the higher DNA purity level it is necessary to remove it. CTAB binds with polysaccharides, cell wall debris and denatured proteins. After lysis, contaminants such as lipophilic molecules, 26║ P a g e

proteins and CTAB/polysaccharide complexes are removed by extraction with CIA and a crude DNA extract is generated by precipitation. At a concentration below 0.5 M NaCl, CTAB forms an insoluble complex with nucleic acids which can be precipitated by centrifugation while polysaccharides, phenolic components and other enzyme-inhibiting contaminants remain in solution (van den Eede, 2006).

Some modifications were made like increasing the incubation time in some step and addition of PCI step. Tris saturated PCI, which is one of the most commonly used method for de-proteinizing DNA extraction with phenol, efficiently denatures proteins and probably dissolves denatured protein (Kirby, 1957). Chloroform stabilizes the rather unstable boundary between an aqueous phase and a pure phenol layer. The use of a mixture of chloroform and phenol also reduces the amount of aqueous solution retained in the organic phase, compared to a pure phenol phase, to maximize the yield (Penman, 1966; Palmiter, 1974). Iso-amyl alcohol is added to prevent foaming of the solution and to aid in the separation of the organic and aqueous phases (Marmur, 1961; Moore and Dowhan, 2002). The denatured protein forms a layer at the interface between the aqueous and organic phases and is thus isolated from the bulk of the DNA in the aqueous layer. The PCI, CI extraction and IPA precipitation is performed to concentrate the DNA solution. In the DNA precipitation step the incubation was for overnight to maximize the final yield. The band of genomic DNA was given in Fig. 2.2.

Molecular Identification Nucleic acid detection methods such as PCR have become a common tool for identification and characterization of microbial communities. The exponential amplification of a target DNA strand was catalyzed by a thermostable DNA polymerase. Molecular identification has become the foundation of Nucleic Acid-based pathogen detection. Although culture-based methods and enzyme-linked immunosorbent assay (ELISA) are still fundamentally valuable techniques (Vincelli and Tisserat, 2008). PCR detection is also usually completed more rapidly than culture techniques (Harmon et al., 2003; Martin and Tooley, 2004; Peres et al., 2007; Zhang et al., 2006a). PCR is 27║ P a g e

increasingly being used as an alternative to culture-based methods for the detection and, in some cases, quantification of microorganisms in various environmental samples including air, soils, landfills, waters, etc. (Steffan and Atlas, 1991; Plaza et al., 2004). PCR is widely recognized as a highly sensitive technique; detection of the DNA from as little as a single spore is possible (Wiglesworth et al., 1994; McCartney et al., 2003). As rDNA sequences are present in high copy number in the fungal genome, their use generally increases the sensitivity of a detection assay. Sequences unique to the target organisms can be found by other approaches. Specific primers can be designed from cloned genomic DNA fragments (Ersek et al., 1994; Doss and Welty, 1995; Jiménez-Gasco and Jiménez-Díaz, 2003).

PCR offers several advantages compared to more traditional methods of diagnosis: organisms need not be cultured prior to their detection by PCR the technique possesses exquisite sensitivity, with the theoretical potential to detect a single target molecule in a complex mixture without using radioactive probes; and it is rapid and versatile (López et al., 2009). Similar to serology, both narrow and broad selectivities are possible and, depending on the choice of primers, the method facilitates the detection of a single pathogen or many members of a group of related pathogens. Unlike serology, the development of reagents with narrow or broad specificities is accomplished almost at will with lower cost. Synthesis of hundreds of different PCR primers generates costs comparable to those of developing only a few monoclonal antibodies (Henson and French, 1993).

The purpose of this study was to develop molecular tools for identification of fungi from the infected sample specifically by designing precise primer for them. General concepts for primer design have been described by many researchers (Dieffenbach et al., 1993; He et al., 1994; Rychlik, 1995). Classical PCR amplifications require minimal sequence information from which appropriate primers are selected. Primers are generally 18-28 nucleotides in length and are defined according to the DNA sequences flanking the region to be amplified. Two parameters are important for the primer design: the efficiency and the specificity of amplification. For instance, 28║ P a g e

an important rule is that no complementarities between 3‟ ends of the primers should exist, to avoid formation of primer-dimers that would decrease the yield of the amplified product (Abd-Elsalam, 2003).

The DNA sequence approach basically involves comparing nucleotide sequences of specific segments of DNA. The sequences are aligned to determine the numbers of differences between comparable molecules. Nucleic acid sequence data is the ultimate level for determining homology. A primer was designed with the help of primer3plus software. Single sharp fragments of particular region of 28s amplified with the help of designed primer and the amplified product was about 325bp (Fig. 2.3). These products was further purified and used in sequencing by ABi 3130 genetic analyzer. Their electropherogram obtained from genetic analyzer was shown in Fig. 2.4(a-f). These sequences were then identified by nucleotide blast (BLASTn tool) from GenBank database of NCBI. The BLASTn percent similarity results were given in Table-2.3. On the basis of similarity results Isolate-2 was identified as Fusarium sp., Isolate-3 as Antrodia sitchensis, Isolate-4, 5 and 7 as Macrophomina phaseolina and Isolate-6 as Curvularia intermedia. Thus, identified isolates belong to different class in fungal taxonomy.

DNA sequences which are polymorphic between fungal species, such as ITSs, are good candidates for the detection of a species to the exclusion of all other species. For example, differences in ITS sequences have been used to develop PCR-based assays for the detection of many phytopathogenic fungal species in host plants without previous isolation of the fungi (Moukhamedov et al., 1994; Beck and Ligon, 1995; Goodwin et al., 1995; Sreenivasaprasad et al., 1996). Other sequences of rDNA have been used to design specific primers, such as 18s rDNA (Gargas and Taylor, 1992; Nikkuni et al., 1998), 28s rDNA (Fell, 1995) and mitochondrial rDNA (Li et al., 1994). rDNA sequences have also been used to develop PCR assays for the detection of fungal species in clinical samples (Spreadbury et al., 1993; Holmes et al., 1994).

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PCR amplification methods with specific fungal primers are powerful tools not only in diagnostics but also in ecological studies for monitoring fungi in natural environments, such as water, soil, plant or clinical samples. Furthermore, the development of specific primers has greatly facilitated studies on obligate parasites and symbionts. For example, specific amplification of mycorrhizal fungal DNA can be performed from colonized plant roots (Di Bonito et al., 1995). Moreover, PCR is classically used to amplify target DNA sequences less than 4000 bp in length (Kainz et al., 1992), but PCR protocols have recently been developed to amplify longer DNA fragments (Cheng et al., 1994; de Vega et al., 2010; Keeney, 2011).

Phylogenetic analysis of 28s rDNA sequences The morphological traits can vary substantially from culture to culture (Seady, 1996). The taxonomic considerations, based solely on phenotype, may be subject to ambiguities induced by environmental conditions. Previously, the taxonomy and identification of fungi have been based mainly on morphological characteristics. Combining molecular and morphological data sets has been discussed by Doyle (1992).

Recent molecular techniques have been introduced to provide more objective criteria. Hu et al (2011) had used rDNA ITS sequences from 60 isolated strains and 9 reference strains to construct phylogenetic tree of dematicacious fungi. The neighbor-joining (NJ) tree indicated the relationships between genetic distances of strains, for example, the strains that can cause diseases in human and animal often group together; on the other hand, the strains that can cause diseases in plants usually get into other crowd. The strains within the crowd or with closer genetic distance would found to be clustered together according their potential pathogenicity. The present study includes genus Macrophomina, Curvularia, Fusarium, and Antrodia to develop the phylogenetic tree based on the 28s rDNA sequences (Fig. 2.5).

The fig. 2.6 shown that species belongs to genus Macrophomina and Rhizoctonia were divided into two groups. The Macrophomina phaseolina D1,

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Macrophomina phaseolina 4 and Macrophomina phaseolina 7 strain of this experiment when paired wise align to develop NJ dendrograms, two major groups A and B where developed. Among them group A shown species of Rhizoctonia and some species of Macrophomina, while group B shown mainly Macrophomina phaseolina stain or isolates. Thus, the strains identified as Macrophomina phaseolina species.

The phylogenetic tree constructed by NJ method for Fusarium species shown in Fig. 2.7. The group C includes two species of Fusarium and Gibberella, while group D includes various species of Fusarium. The group D was further divided into D1 that includes Fusarium oxysporum and D2. The subgroup of D2 includes D3 that contains Fusarium avenaceum and D4. The subgroup of D4 contains D5 comprised of Fusarium solani and D6 which includes Fusarium species. The isolates of present studies fall in D6. The morphological character described isolate-2 as Fusarium solani based on the spore structures (Plate 1). Thus, morphological and phylogenetic analysis confirmed that this isolate-2 was belongs to Fusarium solani.

The 28s rRNA based phylogenetic tree were constructed for Antrodia species (Fig. 2.8). Two major groups E and F were generated in a tree. The group F was divided into two subgroups F1 and F2. Group F1 contains Antrodia albida while group F2 was further divided into F3 and F4. The F3 group contains many subgroups and confirmed that isolate-3 was Antrodia sitchensis.

The phylogenetic tree constructed for Curvularia species was shown in Fig. 2.9. It includes major two groups G and H. The group G was further divided into various subgroups. The phylogentic tree shown isolate-6 was near to Curvularia lunata and Curvularia intermedia, while BLASTn result showed that it was near to Curvularia intermedia (Table-2.3). Thus, based on both morphological and molecular analysis it was confirmed that isolate-6 was Curvularia intermedia.

In the last two decades, DNA technologies have been revolutionized the plant pathogen genomic analysis and have been extensively employed in many 31║ P a g e

fields of molecular plant pathology. Molecular techniques offer also the possibility of faster and accurate identification and early detection of plant pathogen (Bridge et al., 2003; Schaad et al., 2003). Currently, numerous diagnostics laboratories are adopting molecular methods for plant pathogen detection (Lopez et al., 2003; Schaad et al., 2003; Hernandez-Delgado, 2009). Specific primers are derived from sequences of either amplified or cloned DNA (cDNA) or RNA from the microorganism to be detected. Several factors affect primer specificity for the target sequences, including primer length, annealing temperature, magnesium concentration, and secondary structure of target and primer sequences (Bej and Mahbubani, 1991; Erlich et al., 1991; Steffan and Atlas, 1991; Riedel et al., 1992; Shen and Hohn, 1992; Arnheim and Erlich, 1992; Henson and French, 1993 ; Singh and Kumar, 2001; Jensen et al., 2010).

The increasing popularity of molecular approaches for the identification of fungal pathogens reflects significant improvements in DNA analysis in recent years, including the development of broad-range (panfungal) primers, kit- based automated sample processing and DNA amplification systems, and expanding public and private DNA sequence databases. The most promising results were derived from ribosomal sequencing studies examining yeasts and dermatophytes; however, dimorphic fungi and moulds are also being investigated in order to overcome diagnostic problems encountered by extended incubation periods and the plasticity of traditional identification criteria (Reiss et al., 2000; Chen et al., 2001; Lindsley et al., 2001; Iwen et al., 2002; Hall et al., 2003; Hall et al., 2004; Loeffler et al., 2002; Hinrikson et al., 2005).

CONCLUSIONS

The metabolic versatility of fungi is exploited by the fermentation industry, to make antibiotics and other high value substances of interest to medicine, agriculture and the chemical industry, to produce enzymes and to carry out specific steps in chemical processes. This study may help to the fermentation industries in production of various metabolites by understanding growth and 32║ P a g e

development of fungi, in define media. Nucleic Acid based pathogen detection techniques offer an increasing selection of tools for addressing disease problems currently of concern in applied plant pathology. On the basis of results it concluded that the fungi identified based on designed primer of 28s region were belongs to different classes. The primer may be used as universal primer for identification of fungi either from the infected host or any environmental isolates. This primer could be used to develop a kit for molecular identification of broad ranged fungal specimens.

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Plate-1: Identification of fungi on the basis of their morphology and spore

structure

Legends of Microscopical Examination (Light microscopy)

a. Conidiophore with formation of Sterigmata b. Conidiophore with Sterigmata c. Formation of conidia (asexual spore) d. Fully developed structure of Aspergillus e. Conidiophore, Vesicle, Sterigmata, Conidia in chain on sterigmata was observed f. Round black conidia g. Hyphae h. Spore observed under 10x objective lens i. Spore observed under 45x objective lens j. Sickle shaped four celled macro conidia of Fusarium k. Hyphae structure l. Hyphae m. Septated Hyphae n. Hyphae o. Conidiophore with initial stage of spore formation p. Three celled curved shape conidia q. Chlamydospore r. Chlamydospore and initial stage of conidial spore formation s. Hyphae

Isolate-1 2.5

1.5

1 Weight (g) Weight

0.5

0 24 48 72 96 120 144 168

Time (h) Fresh Weight Dry Weight

10 Isolate-1 pH 9 8 7

6 5 pH 4 3 2 1 0 24 48 72 96 120 144 168 Time (h)

Figure-2.1a: Variation in fresh weight, dry weight and pH of Isolate-1 with age in MS media (2% sucrose)

Isolate-2 3.5

2.5

1.5

Weight (g) Weight 1

0.5

0 24 72 120 168 216 264 312 Time (h) Fresh Weight Dry Weight

10 Isolate-2 pH 9 8 7

6 5 pH 4 3 2 1 0 24 48 72 96 120 144 168 192 216 240 264 288 312 336 Time (h)

Figure-2.1b: Variation in fresh weight, dry weight and pH of Isolate-2 with age in MS media (2% sucrose)

2.5 Isolate-3

1.5

1 Weight (g) Weight

0.5

0 24 48 72 96 120 144 168 192 216 240 264 288 312 336 Time (h) Fresh Weight Dry Weight

Isolate-3 pH 10 9 8 7 6

5 pH 4 3 2 1 0 24 48 72 96 120 144 168 192 216 240 264 288 312 336 Time (h)

Figure-2.1c: Variation in fresh weight, dry weight and pH of Isolate-3 with age in MS media (2% sucrose)

6 Isolate-4

Weight (g) Weight 2

0 24 48 72 96 120 144 168 Time (h) Fresh Weight Dry Weight

Isolate-4 pH 10 9 8 7 6

5 pH 4 3 2 1 0 24 48 72 96 120 144 168 Time (h)

Figure-2.1d: Variation in fresh weight, dry weight and pH of Isolate-4 with age in MS media (2% sucrose)

Isolate-5 4.5 4 3.5

3 2.5 2

Weight (g) Weight 1.5 1 0.5 0 24 48 72 96 120 144 168 Time (h)

Fresh Weight Dry Weight

pH 10 Isolate-5 9 8 7

6 5 pH 4 3 2 1 0 24 48 72 96 120 144 168 Time (h)

Figure-2.1e: Variation in fresh weight, dry weight and pH of Isolate-5 with age in MS media (2% sucrose)

3.5 Isolate-6

2.5

1.5 Weight (g) Weight 1

0.5

0 24 48 72 96 120 144 168 Time (h) Fresh Weight Dry Weight

Isolate-6 pH 10 9 8 7

6 5 pH 4 3 2 1 0 24 48 72 96 120 144 168 Time (h)

Figure-2.1f: Variation in fresh weight, dry weight and pH of Isolate-1 with age in MS media (2% sucrose)

Isolate-7 3.5

3.0

2.5

2.0

1.5 Weight (g) Weight 1.0

0.5

0.0 24 48 72 96 120 144 168 192 Time (h)

Fresh weight Dry weight

pH 10 Isolate-7 9 8 7

6 5 pH 4 3 2 1 0 24 48 72 96 120 144 168 192 Time (h)

Figure-2.1g: Variation in fresh weight, dry weight and pH of Isolate-7 with age in MS media (2% sucrose)

Figure-2.2: Agarose gel electrophoresis of genomic DNA (isolate-1 to 7)

Figure-2.3: Agarose gel electrophoresis of designed primer for 28s gene

Figure-2.4a: Electropherogram of isolate-2 for 28s gene retrieved from ABi 3130 genetic analyzer

Figure-2.4b: Electropherogram of isolate-3 for 28s gene retrieved from ABi 3130 genetic analyzer

Figure-2.4c: Electropherogram of isolate-4 for 28s gene retrieved from ABi 3130 genetic analyzer

Figure-2.4d: Electropherogram of isolate-5 for 28s gene retrieved from ABi 3130 genetic analyzer

Figure-2.4e: Electropherogram of isolate-6 for 28s gene retrieved from ABi 3130 genetic analyzer

Figure-2.4f: Electropherogram of isolate-7 for 28s gene retrieved from ABi 3130 genetic analyze

Figure-2.5: Phylogenetic analysis of six isolates by neighbor-joining (NJ) method.

Figure-2.6: The phylogenetic analysis by neighbor-joining (NJ) method for Macrophomina species

D5 D2

D3 D D1

Figure-2.7: The phylogenetic analysis by neighbor-joining (NJ) method for Fusarium species

Figure-2.8: The phylogenetic analysis by neighbor-joining (NJ) method for Antrodia species

Figure-2.9: The phylogenetic analysis by neighbor-joining (NJ) method for Curvularia species

Table-2.1: Morphological characteristics of fungi on PDA

Fungi Infected host Hyphal Spore plant Morphology Morphology Present, black in Isolate-1 Allium cepa (Onion) White color, round shape Present, pink in color, Isolate-2 Cuminum cyminum (Cumin) Pink elliptical shape

Isolate-3 Cicer arietinum (Chick pea) White Absent

Isolate-4 Ricinus communis (Castor) Grayish Absent black

Isolate-5 Mangifera indica (Mango) Grayish Absent black Present, Pennisetum typhoides brown in Isolate-6 Brown (Bajra) color, curve shape

Isolate-7 Rosa indica (Rose) Grayish Absent black

Table-2.2: DNA Purity and concentration

DNA Purity Concentration Name of fungi (A260/A280) g/ml

Isolate-1 1.79 123.3

Isolate-2 1.60 230.3

Isolate-3 1.63 135.6

Isolate-4 1.82 137.3

Isolate-5 1.83 104.25

Isolate-6 1.70 193.3

Isolate-7 1.96 161.8

Table-2.3: Molecular identification of fungi from NCBI (Genbank)

Genbank Accession Fungi Query Isolate No. No. of Max identity identified as coverage Reference sequence

02 Fusarium sp. HQ026783.1 94% 98%

Antrodia 03 AY333829.1 79% 96% sitchensis Macrophomina 04 FJ395247.1 95% 98% phaseolina Macrophomina 05 FJ395247.1 83% 86% phaseolina Curvularia 06 AF163984.1 95% 97% intermedia Macrophomina 07 FJ395247.1 95% 98% phaseolina

INTRODUCTION

Aspergillus niger is a filamentous ascomycete fungus that is ubiquitous in the environment and has been implicated in opportunistic infections of humans (Perfect et al., 2001; Baker, 2006). There are some fungi causing post-harvest diseases include: Aspergillus spp, Penicillium spp, Alternaria spp, Botrytis cinerea, Monilinia lax and Rhizopus stolonifer (Ogawa et al., 1995). Black mould of onions is caused by the fungus A. niger. It is a high-temperature fungus, with an optimum temperature range for growth of 28-34°C. Warmth and moisture favour development of the disease (Tyson and Fullerton, 2004). Although the disease can occasionally be seen in the field at harvest, black mould is primarily a postharvest disorder and can cause extensive losses in storage under tropical conditions (Thamizharasi and Narasimham, 1992).

As a common member of the microbial communities found in soils, A. niger plays a significant role in the global carbon cycle. This organism is a soil saprobe with a wide array of hydrolytic and oxidative enzymes involved in the breakdown of plant lignocelluloses (Gautam et al., 2010). A variety of these enzymes from A. niger are important in the biotechnology industry. A. niger also known to produce highly toxic secondary metabolites. Toxic metabolites of Aspergillus form and accumulate in soil, cause toxic damage, and inhibit plant growth via the soil. Although soil is the main habitat of Aspergillus fungi, they can grow on various substrates and are sometimes abundant in grain, its processing products, and industrial substrates. Fodders and foodstuffs infected by Aspergillus cause aspergillosis and aspergillotoxicosis in man, domestic animals, and birds. In addition, toxins of Aspergillus are known to be toxic to plants (Vinokurova et al., 2003).

Aspergillus and other filamentous fungi are known for their ability to secrete a variety of biologically active chemical compounds. Formally, the term “secondary metabolite” is used to describe this low molecular weight, “nonessential” natural products usually produced after primary growth has stopped. These compounds include antibiotics, mycotoxins, immunosuppressants, and cholesterol-lowering agents. They are classified 34║ P a g e

chemically by their biosynthetic origin as polyketides, nonribosomal peptides, sesquiterpenes, and so forth (Keller et al., 2005; Coleman et al., 2011).

Phytohormones play an important role as regulators of growth and development of plants. According to the conventional classification, there are five groups of phytohormones: auxins, gibberellins, cytokinins, ethylene, and abscisic acid (Tsavkelova et al., 2006a). Phytohormones contribute to the coordination of diverse physiological processes in plants, including the regulation of quiescence and seed germination, root formation, florescence, branching and tillering, and fruit ripening. They increase plant resistance to environmental factors and induce or suppress the expression of genes and the synthesis of enzymes, pigments, and metabolites (Agrios, 1997; Kulaeva and Kuznetsov, 2002)

The ability of microorganisms to synthesize phytohormones is widely known (Costacurta and Vanderleyden, 1995). Producers with the highest activity enter into association interactions with plants, which may have pathogenetic or symbiotic consequences. Bacteria, micromycetes, and algae form phytohormones of auxin, cytokinin, or gibberellin nature (Tsavkelova et al., 2006a). On the other hand, microorganisms also synthesize other phytohormones and phytohormone-like substances, including ethylene and abscisic acid (ABA), brassinosteroids, oligosaccharines, salicylic acid and jasmonic acid (Tsavkelova et al., 2006b).

Many researchers determined the phytohormone content in plants during pathogenesis (Volynets et al., 1993; Prinsen and van Onckelen, 1994; Nikitina and Talieva, 2001; Kilaru et al., 2007; Bruce et al., 2011). These studies showed that the phytohormone content in infected plants depended on the pattern of fungus development. However, the detailed investigations were devoted only to characterizing the phytohormone balance in plants infected by pathogens, which cause diseases of only leaves, stems, or roots (local fungal development) (Maksimov et al., 2002).

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Many microorganisms associated with plants as symbionts or parasites directly alter the auxin content of the host; the auxin imbalance benefits the microorganisms (Gaudin et al., 1994; Tudzynski and Sharon, 2002). The auxins control plant growth, differentiation, structural organization, and the shift from vegetative to reproductive growth of pathogenic strain. Virulence directly correlated with the levels of IAA, suggesting that IAA has a positive function in the infection process (Ek et al., 1983). Plants pre-infected with a pathogen caused auxin production, which then overcame the hypersensitive defense response of plants subsequently infected by an incompatible pathogen (Robinette and Matthysse, 1990; Cohen et al., 2002).

While the discovery of IAA in yeast extracts goes back to more than half a century, its recognition in filamentous fungi is relatively new. There is little data on the actual quantity of IAA synthesized by fungi or flowering plants. Soon after the discovery of indole-3-acetic acid (IAA) in higher plants, auxin activity was also detected in fungi (Gruen, 1959); however, IAA biosynthetic pathways were identified in only a few fungi (Furukawa et al., 1996; Sosa et al., 1997). Although high IAA levels are often found in diseased plants, the role of IAA in fungus-plant interactions has not been determined (Robinson et al., 1998; Reineke et al., 2008; Fu and Wang, 2011).

Biosynthesis of IAA is not limited to higher plants. Organisms such as bacteria, fungi, and algae are able to make physiologically active IAA that may have pronounced effects on plant growth and development. Many bacteria isolated from the rhizosphere have the capacity to synthesize IAA in vitro in the presence or absence of physiological precursors, mainly tryptophan (Trp) (Costacurta and Vanderleyden, 1995; Dong et al., 1995). Microbial isolates from the rhizosphere of different crops appear to have a greater potential to synthesize and release IAA as secondary metabolites because of the relatively rich supply of substrates (Barazani and Friedman, 1999).

In addition, numerous pathogens are active producers of IAA and cause abnormal cell enlargement in infected plants (Patten and Glick, 1996; Arshad and Frankenberger, 1998). Production of IAA by microbial isolates varies 36║ P a g e

greatly among different species and strains and depends on the availability of substrate(s). Different biosynthetic pathways for IAA production exist, sometimes in parallel in the same organism (Patten and Glick, 2002). For many years it was assumed that Trp was the only precursor of IAA. However, work with Trp-auxotrophic mutants and isotope labeling has established that IAA biosynthesis can occur via a Trp-independent route (Prinsen et al., 1993; Normanly and Bartel, 1999).

Most phylogenetic analyses are focused on the nuclear ribosomal DNA rDNA), which contains tandem repeats of three rRNA genes (28s, 18s, and 5.8s). DNA sequence comparisons of the highly conserved nuclear 18s rRNA gene have been used to infer evolutionary relationships between different groups of fungi as well as their relationships to organisms in other kingdoms (Berbee et al., 1992; Meyer et al., 2010).

On the basis of these, objective of this study was evaluated as follows:

I. Screening of optimum time of high IAA production in define medium. II. Substrate dependent assay was used to detect the optimum production of IAA. III. Identification of fungi by 18s sequencing. IV. Primer designing for Aldehyde 3-dehydrogenase gene responsible for IAA production their amplification and sequencing analysis. V. Post sequencing analysis for their physic-chemical properties and PDB structure.

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MATERIALS AND METHODS

Isolation and Morphological identification Fungus was isolated from infected host Allium cepa and grown on PDA media. Its microscopical examination was also carried out for their preliminary identification (See chapter- 2).

Fungal Culture and Inoculum Preparation A. niger strain previously stored in distilled water was streaked on PDA plate and incubated for seven days for culture activation. From that plate fungal disc (1 cm diameter) was cut with the help of cup borer and placed in the center of another plate. The plate was also incubated for seven days for complete radial growth and it was further used for inoculums.

Molecular identification DNA preparation DNA extraction and visualization were performed as described in Chapter-2.

PCR Amplification for 18s sequencing The 18S fragment of this fungus was amplified with one set of primer as directed by Melchers et al. (1994). PCR based DNA was amplified in a 25 l reaction volume using Verity, ABi (USA).

Media and culture condition Optimization of media In the first part of experiment MS (1962) with certain alteration was used. Main four types of media were designed as: (i) MS full strength with sucrose 2% [MS (2)]; (ii) MS full strength with sucrose 4% [MS (4)]; (iii) MS half strength with sucrose 2% [MS (1/2) (2)]; (iv) MS half strength with sucrose 4% [MS (1/2) (4)] with 1 mM of Trp as substrate. The control kept for each set was without Trp.

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To avoid the photo oxidation of product (IAA) the culture was grown in dark and in static condition at 25 ± 2ºC. The initial pH of define medium was 5.6 ± 0.2. At interval of every 24 h some amount of filtrate was withdrawn and estimated the amount of IAA present in that by Calorimetric method with the help of Salkowasky reagent up to third days. The O.D. was measured at 520nm. These tests were performed in triplicate and mean values with ± SD were presented (Fig. 3A.2).

Optimization of time In second part of the experiment, a define media MS (1/2) (4) and 5 mM Trp as a substrate supplement were used. The controls and culture condition was same as above but the filtrate was withdrawn and estimated at an interval of every day up to six days (Fig. 3A.3). In second set of experiment the filtrate was withdrawn at an interval of every 8 h up to 3 days and estimated by calorimetric method as described above (Fig. 3A.4a). To find out the optimum time for IAA production the filtrate collection was further narrow down at interval of every hour in between 39 h to 48 h (Fig. 3A.4b). Entire tests were performed in triplicate and the concentration of IAA was calculated with ± SD using standard curve.

Effect of substrate concentrations and temperature Depending on the results of previous experiments in the third part of experiment a substrate dependent assay was carried out. A define media and culture condition was same as above but In this the concentration of Trp (precursor) was varies like 5, 10, 15, 20, 25, 50 mM/ml. Withdrawn of filtrate at optimum time i.e. 46 h was carried out and production of IAA was estimated. The above experiment was also checked out at 20 ± 2 ºC for evaluation of temperature effect on IAA production. The entire tests were performed in triplicate. The fresh weight, dry weight of filaments and pH of the medium was also measured. The produced IAA concentration was calculated by plotting the value in standard curve of IAA and expressed with ± SD (Fig. 3A.5a-c).

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Primer designing Aldehyed dehydrogenase (ALDH) primer was synthesized from the sequence of Ustilago maydis. For that the enzyme gene sequence were retrieved from Gene bank (Accession no: U74468) and blast with whole genome of A. niger (An08g10820) in The Central Aspergillus REsource (CADRE) (http://www.cadre-genomes.org.uk/index.html) Fig. 3A.6a. The primer for similar gene sequence obtained from blast in A. niger were designed with the help of Primer3plus online software (http://www.bioinformatics.nl/cgi- bin/primer3plus/primer3plus.cgi). Five pairs of primer was obtained are listed in Table-3A.1. The probable primer was further used in optimization of PCR for singal band product. The criteria that was set for primer design was primer length is 20 ± 2, target temperature is 59 ± 1°C. Details of the primers sequences, their expected amplicons sizes after PCR, etc. are summarized in Table-3A.1.

Target gene These oligonucleotide primers were used for amplification of approximately a 300bp PCR fragment of targeted aldh gene from genomic DNA of A. niger.

Polymerase Chain Reaction Optimization PCR is carried out with the designed primer set. A reaction tube contained 100 ng of template DNA, 1× PCR buffer, 1 units of Taq DNA polymerase, 200 mM each of dNTPs, for the optimization of PCR different MgCl2 concentrations (1-2.5 mM) and various nucleotide primers 10-75 pmol/ l concentration in 25 l final volume. The PCR conditions were: denaturation at 95ºC temperature for 5 min, followed by stage two of 45 cycle in which denaturation at 94ºC temps for 30 s primer annealing 50.3 - 52.3 ºC for 30 s and extension was done at 72ºC for 60 s. The final extension was done at 72ºC for 10 min then being held at 4ºC in Verity, ABi (USA).

Purification of PCR product to remove excess primer and nucleotides was performed using Qiagen‟s QIAquick PCR purification kit and 5 µl of product

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was checked on 2 % agarose gel by Electophoresis. The remaining product was used in cycle sequencing.

Cycle sequencing 18s sequencing Cycle sequencing was performed by using ABi BigDye Terminatior v3.1 Cycle sequencing kit. Total volume was prepared 10 µl, as per standard protocol of ABi. The reaction mixture contain either of the primer (forward/reverse), 5x sequencing buffer, the template DNA, BigDye terminator and Mili-Q water.

Cycle sequencing was carried out in Veriti, ABi (USA). Reaction condition were: first initial denaturation at 95ºC temperature for 2 min, followed by second stage of 45 cycles includes 95ºC temperature for 30 s, 42ºC temperature for 45 s, and 60ºC temperature for 3 min then being held at 4ºC. Purification of cycle sequencing products was done by Dye Ex 2.0 spin kit of Qiagen as per manufacture‟s protocol and subjected to automated sequencing and analyzed using ABi 3130 Genetic Analyzer. aldh gene sequencing Cycle sequencing was performed by using ABi BigDye Terminatior v1.1 Cycle sequencing kit. Total volume was prepared 10 µl, as per standard protocol of ABi. The reaction mixture contain either of the primer (forward/reverse), 5x sequencing buffer, the template DNA, BigDye terminator and Mili-Q water.

Cycle sequencing was carried out in Veriti, ABi (USA). Reaction condition were: first initial denaturation at 95ºC temperature for 10 min, followed by second stage of 45 cycles includes 95ºC temperature for 20 s, 52ºC temperature for 10 s, and 60ºC temperature for 4 min then being held at 4ºC. Purification of cycle sequencing products was done by Dye Ex 2.0 spin kit of Qiagen as per manufacture‟s protocol and subjected to automated sequencing and analyzed using ABi 3130 Genetic Analyzer.

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Post sequencing analysis using bioinformatics tool In present study, the data collected from the ABi 3130 Genetic Analyzer was blast in NCBI data base for similarity match and for identification of the sequence. The gene sequence was submitted in NCBI GeneBank data base with the help of BankIt tool. Other post sequencing analysis was also carried out using different bioinformatics tool to study mutation, physic-chemical properties and protein structure prediction of aldh gene sequence.

ABi Prism SeqScape software SeqScape software is one of a suite of Applied Biosystems Genetic Analyser software Application designed to control an instrument, collect date and manage automated analysis. Some of the common applications of the mentioned software include (1) SNP discovery and validation, (2) mutation analysis and heterozygote identification, (3) sequence confirmation for mutagenesis or clone-construct confirmation studies, (4) identification of the genotype, allele and haplotype from a library of known sequences.

It identifies variants, positions that differ from the reference sequence, and classifies those variants as known or unknown sequences. When the reference sequence, samples files and specimen files have been added to the software, it performs the identification of nucleotide and amino acid variants. The software identifies the positions that differ from the reference sequence.

ExPASy translate tool ExPASy is the new SIB Bioinformatics Resource Portal which provides access to scientific databases and software tools in different areas of life sciences including proteomics, genomics, phylogeny, systems biology, population genetics, transcriptomics etc. It is a tool which allows the translation of a nucleotide (DNA/RNA) sequence to a protein sequence (http://web.expasy.org/translate/). Six sequences are obtained and from that a potential sequence were find out with the help of protein blast search tool of NCBI. That potential sequence was further used for physico-chemical analysis and protein structure prediction.

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BLASTp on UniProtKB UniProt is provided to scientific community with a comprehensive, high-quality and freely accessible resource of protein sequence and functional information. The obtained potential protein sequence was blast in UniProtKb for similarity result of relative sequence (http://www.uniprot.org/).

Physico-chemical analysis by ProtParam tool ProtParam computes various physico-chemical properties that can be deduced from a protein sequence (http://web.expasy.org/protparam/). No additional information is required about the protein under consideration. The protein can either be specified as a Swiss-Prot/TrEMBL accession number or ID, or in form of a raw sequence (Gasteiger et al., 2005).

Protein prediction by SWISS-MODEL It is a fully automated protein structure homology-modeling server, accessible via the ExPASy web server (http://swissmodel.expasy.org/workspace/), or from the program DeepView (Swiss Pdb-Viewer). The purpose of this server is to make Protein Modelling accessible to all biochemists and molecular biologists worldwide. To carried out homology modeling for ALDH protein, PDB ID i.e. 1a4zA of known ALDH protein was used as template.

RESULTS AND DISCUSSION

Numerous molecular techniques have been employed in the detection, identification and phylogenetic analysis of fungal populations. Fragments of 18s rDNA has been used as effective markers in genotyping studies (Hernan- Gomez et al., 2000, Vainio and Hantula, 2000). In this study a fungi was amplified by using 18s primer (Fig. 3A.6b) and identified as A. niger its accession no. given in Table-3A.2 and its electropherogram shown in Fig. 3A.7a.

Many fungi can produce auxins in axenic cultures (Buckley and Pugh, 1971; Maor et al., 2004). Most species use Trp to produce indole-3-acetic acid

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(IAA), mainly through the indole-3-pyruvic acid and tryptamine pathways (Tudzynski and Sharon, 2002). Although in the presence of Trp microbes release greater quantities of IAA and related compounds. The pathways for conversion of Trp to IAA can involve deamination, decarboxylation, and/or hydrolysis reactions, in higher plants and most microorganisms; the indole-3- pyruvic acid (IpyA) pathway is the main one for IAA synthesis (Kawaguchi and Syono, 1996; Lee et al., 2004; Mashiguchi et al., 2011).

In microorganisms, the three known pathways of IAA biosynthesis are also related to Trp metabolism (Costacurta and Vanderleyden, 1995; Patten and Glick, 1996; Kameneva and Muronets, 1999) (Fig. 3A.1). IAA formation via indole-3-pyruvic acid and indole-3-acetic aldehyde is found in the phytopathogenic micromycetes of the genera Fusarium, Rhizoctonia, and Colletotrichum (Thakur and Vyas, 1983; Furukawa et al., 1996; Chung et al., 2003). The conversion of Trp into indole-3-acetic aldehyde may involve an alternative pathway in which tryptamine is formed (Kameneva and Muronets, 1999). This pathway is believed to operate in Pseudomonades and Azospirilla (Mordukhova et al., 1991) and an unidentified mycorrhizal fungus of orchid (Ophrys lutea Cav.) (Barroso et al., 1986). IAA biosynthesis via indole-3- acetamide formation takes place in phytopathogenic fungi of the genus Colletotrichum (C. gloeosporioides and C. acutatum) (Maor et al., 2004; Chung et al., 2003).

Trp as a precursor of IAA in fungi was first reported by Thimann (1935). Trp is one of the essential amino acids and the major contributor of the indole ring for the synthesis of many important compounds including auxins, glucosinolates, nicotinic acid, phytoalexins and alkaloids (Barone and Widholm, 2008). Trp is synthesized by a five-enzyme biosynthetic pathway from the branch point compound chorismic acid at the end of the shikimic acid pathway (Radwanski and Last, 1995).

Optimization of media Here, the production of this hormone was investigated in an industrially important fungi A. niger. The main objective was to determine the conditions 44║ P a g e

(both physical and chemical) required for the optimal production of IAA by this fungus. Salkowasky reagents have their own color and may give some color reaction with Trp. But with indole it gives pinkish color. This reagent was also used by some or worker to check the amount of IAA present in the filtrate (Fukuhara et al., 1994; Lee et al., 2004; Rahman et al., 2010).

Yeasts of the genus Saccharomyces and Micromycetes belonging to the genera Fusarium, Rhizoctonia, Rhizopus, Absidia, Aspergillus, Penicillium Monilia, Phoma, Pythium, Trichoderma, and Actinomucor also produce IAA (Tsavkelova et al., 2006a; Bilkay et al., 2010). The ability to synthesize IAA and other auxins is found in diverse species of phytopathogenic fungi causing boil smut, gall formation, and other plant diseases (Taphrina, Phytophthora, Ustilago, Alternaria, Fusarium, Plasmodiophora, Colletotrichum, Phymatotrichum, Lentinus, and Sclerotium) as well as in mycorrhizal fungi of the genera Laccaria, Pisolithus, Amanita, Rhizopogon, Paxillus, and Hebeloma (Gunasekaran and Weber, 1972; Shin et al., 1991; Gopinathan and Raman, 1992; Furukawa et al., 1996; Shimada et al., 2000; Laurans et al., 2001; Wu et al., 2002; Niemi et al., 2002; Yurekli et al., 2003; Tsavkelova et al., 2003; Rincon et al., 2003).

The optimum media for higher production of IAA found was MS (1/2)(4) from the various media tested. It can be state that higher concentration of sucrose may responsible for more production of IAA (Fig. 3A.2). In work of Lee et al. (1989) they reported that there was no influence of sucrose supply on IAA content of wheat grain or bracts of detached ears. However, at further incubation periods, sucrose containing medium showed higher IAA levels than carbohydrate free medium.

Optimization of time The IAA levels increased rapidly in cultures during the early stages of incubation. It reached a maximum peak of IAA at 2nd day and then IAA production was declined dramatically thereafter (Fig. 3A.3). When the precursor was incorporated with media supplement the indole level have drastically increased. The condition required for optimum production of indole 45║ P a g e

in vitro was investigated. Time courses were study at every interval of 8 h and this range were narrow down up to every hour. From that it reveals that amount of indole increases exponentially after 40 h and highest at 46 h (Fig. 3A.4b). Chung and Tzeng (2004) were also recorded somewhat similar kind of result, the amounts of IAA increased exponentially after 2nd day in the Czapek‟s solution amended with Trp.

Effect of substrate To determine the important of Trp as a precursor for IAA biosynthesis the fungus was grown in sucrose containing Ms broth supplemented with various concentration of Trp like 5mM, 10mM, 15mM, 20mM, 25mM and 50mM. Trp and indole compound were extracted from the medium and measured at 520 nm by developing the reaction. In the absence of tryptoophan very trace amount of indole were detected, whereas in the presence of Trp IAA accumulated to high levels (Fig. 3A.5a). The amounts of IAA increased with increasing Trp concentrations, providing further support that IAA biosynthesis is Trp dependent which was also reported by Chung and Tzeng (2004) in their work with fungus Ustilago esculenta. The maximum production of IAA was noted in 25 mM substrate concentration at 25 ± 2 °C i.e. 724.23 ± 0.04 mg/L.

The optimum pH for higher production was noted in between pH 2.5-3 in various substrates containing media (Fig. 3A.5b). Yurekli et al. (2003) were reported in their work that pH-3 in dark is optimum for IAA production in Lentinus sajor-caju which supported the finding of our work. In contradiction to the our results Strzelczyk et al. (1992) who reported that auxin biosynthesis was favored in media at pH 6-9 in mycorrhizal fungi.

The amount of IAA increased with increasing Trp concentration upto 25 mM but at 50 mM it was decreased (Fig. 3A.5a). It providing support that up to 25mM its substrate dependent but the high concentration of substrate have toxic or inhibitory effect on the enzyme so the concentration of product may fall down. As shown earlier, IAA biosynthesis requires external Trp. Maor et al., (2004) also describe that Trp increased IAA biosynthesis 2.7-fold in C. gloeosporioides and biosynthesis is enhanced by substrate availability. These 46║ P a g e

results are consistent with the hypothesis that the limiting factor in IAA biosynthesis is the amount of available Trp.

Effect of temperature The effect of temperature on IAA production was also determined. The results indicated that the optimum temperature for IAA production was around 25 ± 2 °C. If the temperature below the 20 ± 2 ºC the production of IAA falls 5-10 folds (Fig. 3A.5a). The fungal growth was slower at 20 ± 2 ºC and it was also shown that the optimum temperature for production of IAA was related with the optimum temperature of fungal growth (Fig. 3A.5c). It also observed that the growth of fungi was more in media containing Trp (Fig. 3A.5b). In the media without Trp the fresh weight of fungal mycelium was 0.200 g and in Trp containing media the fresh weight of fungal mycelium was around 2 g in all (Fig. 3A.5b). The growth rate in Trp containing media was observed 10 fold higher than control and the sporulation were also enhanced (Plate 2).

Primer designing ALDHs catalyze the oxidation of aldehydes to their corresponding carboxylic acids and occur throughout all phyla. Many disparate aldehydes are ubiquitous in nature and most are toxic at low levels because of their chemical reactivity. Thus, levels of metabolic-intermediate aldehydes must be carefully regulated (Perozich et al., 1999). For this, most well-studied organisms are known to have several distinct ALDHs, which take part in a variety of physiological roles. Some ALDHs are highly specific for a very limited range of substrates while others show broad substrate specificity. All ALDHs require either NAD or NADP as a cofactor (Lindahl, 1992; Yoshida et al., 1998; Mihasan et al., 2010).

As reported by Basse et al. (1996) that two aldehyde dehydrogenase was responcible for IAA production in Ustilago maydis. The sequence for aldehyde dehydrogenase for Aspergillus niger was obtained by blast in The Central Aspergillus REsource (CADRE). It supports the international Aspergillus research community by gathering all genomic information regarding this

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significant genus into one resource. CADRE facilitates visualization and analyses of data using the Ensembl software suite.

Oligonucleotide primers are widely used in various molecular biology techniques like DNA sequencing and PCR. Since a primer serves as the starting point for DNA replication, specific binding of the oligonucleotide to the target sequence on the template strand is essential for a successful experiment. The binding specificity of a primer is determined by several of its properties, like the melting temperature (Tm), GC-content and self complementarily. Designing primers is usually done with the help of computer programs, among which Primer3 is most widely used judging from the hundreds of citations of the primary publication (Rozen and Skaletsky, 2000; Untergasser et al., 2007).

Optimization of PCR Furthermore designed primer was used for optimization of PCR. Firstly, different primer concentration was used for optimization ranging from 10, 25, 50 and 75 pmol/ l in that 10 pmol minimum primer concentraton were also shown amplification (Fig. 3A.6c). Then, for the optimization MgCl2 concentrations range from 0.5 to 3.5 mM with increments of 0.5 mM were tested. The amplification yield decreased with an increase in magnesium ion concentration beyond 2 mM, and with the optimum concentration ranging between 1.0 and 2.0 mM (Fig. 3A.6d). The best and clear 300 bp DNA fragment was produced at 1.5 mM MgCl2 concentration of and this concentration was used in the optimization of primer concentration. Primer pair 4 and 5 gives the singal sharp band amongst the all (Fig. 3A.6e).

When the primer concentration was increased, intensity of large bands decreased, whereas intensity of smaller bands increased. Such differences in the amplification occur because as more primer is available and binds to more locations on the DNA, smaller fragments become more prominent, whereas larger fragments are less conspicuous. This may promote misprinting and accumulation of nonspecific products. On the other hand, when using low

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concentrations, the primer may be exhausted before the reaction is completed, resulting in lower yields of amplification bands (Ramírez et al., 2005). These variations make it necessary to determine the optimal primer concentration and this is an important element to keep in mind in order to obtain reproducible patterns with a large number of real bands.

The obtained fragment were purified and further used for cycle sequencing. The purified product of cycle sequencing was run in ABi 3130 genetic analyzer. The data were retrieved from it (Fig. 3A.7b) and blast in NCBI data base. When compared with sequences in the database, Iadl revealed the highest similarity 91 % to aldehyde dehydrogenases from Aspergillus niger (XM_001393149.1) CBS 513.88 aldehyde dehydrogenase, mRNA. Then the sequence was submitted to Genbank data base through BlankIt the accession no was shown in Table-3A.2.

Mutational analysis In mutational studies carried out with ABI prism SeqScap sowtware. 18 sequences of ALDHs of A. niger was retrieved from NCBI data base. Amongst them with the 2 sequences (XM_001398829.2) and (XM_001398166.2) mutation was observed. In case of XM_001398166.2 sequence sixteen missense, one frameshift insertion and one silent type of mutation was found. On the other hand in XM_001398829.2 sequence eighteen missense and two silent type of mutation was found (Fig. 3A.8a, b, Table-3A.4).

Protein sequence analysis The UniProt ID and detail of similar sequence obtained from the UniProt blast was given in Table-3A.3. It was found that the protein sequence was 100% identical A2QV34 and belongs to ALDH family (EC=1.2.1.3). The UniProt sequence A2QV34 showed similarity to indole-3-acetaldehyde dehydrogenase Iad1 of Ustilago maydis. On the basis of similarity results it can be concluded that the aldehyde dehydrogenase sequence obtained in this study was also have properties to convert indole acetic aldehyde into Indole- 3-acetic acid and responsible for their production via indole-3-acetic aldehyde pathway. 49║ P a g e

Physico-chemical analysis of obtained ALDH protein sequence was carried out using ExPASy-ProtParam analysis, number of amino acid residues, molecular weight, theoretical pI, total number of negatively (Asp+Glu) and positively charged residues (Arg+Lys), instability index, aliphatic index, grand average of hydropathicity (GRAVY) were found out through this. Output of all these parameters for the sequence was given in Table-3A.5.

The 3D structure of proteins is better conserved than their sequences, it is often possible to identify a homologous protein with a known structure (template) for a given protein sequence (target). In these cases, homology modelling has proven to be the method of choice to generate a reliable 3D model of a protein from its amino acid sequence as impressively shown in several meetings of the bi-annual CASP experiment (Tramontano and Morea, 2003; Moult, 2005).

Homology modelling is routinely used in many applications, such as virtual screening, or rationalizing the effects of sequence variations (Marti-Renom et al., 2000; Kopp and Schwede, 2004; Hillisch et al., 2004). Building a homology model comprises four main steps: (1) identification of structural template(s), (2) alignment of target sequence and template structure(s), (3) model building and (4) model quality evaluation. These steps can be repeated until a satisfying modeling result is achieved. Each of the four steps requires specialized software as well as access to up-to-date protein sequence and structure databases (Arnold et al., 2006). Structure prediction for obtained potential protein of ALDH sequence was carried out with the homology modeling by SWISS-MODEL and its 3D structure was given in Fig. 3A.9.

CONCLUSIONS

On the basis of above result we can conclude that the optimum temperature for IAA production is 25 ± 2ºC, pH in between 2.5-3, media half strength of MS and 4% sucrose, incubation time in dark was 46 h. The best condition for PCR amplification was 1.5mM MgCl2 and 10 pmol primers. The primer pair 5 gives the best result amongst the all. It can be concluded that the aldehyde 50║ P a g e

dehydrogenase gene amplified in A. niger was responsible for IAA synthesis but for more conformation protein profiling and gene expression studies yet need to carried out.

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Figure-3A.1: Tryptophan metabolism pathway in Aspergillus niger (KEGG: http://www.genome.jp/kegg/pathway/ang/ang00380.html)

MS(1/2)S(2) Media optimization MS(1/2)S(4)

25 MS S(2) MS S(4) 20

IAA Production (mg/L) Production IAA 0 1 2 3 Time (Days)

Figure-3A.2: IAA production in various combinations of salt concentration and media

Optimization of Time

120

100

IAA Production (mg/L) Production IAA 0 1 2 3 4 5 6 Time (Days) MS(1/2) S (4)

Figure-3A.3: Changes in IAA content with time (days)

Optimization of Time

100 90 80 70 60 50 40 30 20

IAA Production (mg/L) Production IAA 10 0 8 16 24 32 40 48 56 64 72 80 Time (h)

(a) Broad range

Optimization of Time 120

100

20 IAA Production (mg/L) Production IAA 0 39 40 41 42 43 44 45 46 47 48 Time (h)

(b) Narrow range

Figure-3A.4: Changes in IAA content with time (h)

Substrate Dependent 800 700 600 500 400 300 200 100 0 IAA Production (mg/L) Production IAA 5mM 10mM 15mM 20mM 25mM 50mM

Concentration (Trp)

Effect of Temperature

140 120 100 80 60 40 20

IAA Production (mg/L) Production IAA 0 5mM 10mM 15mM 20mM 25mM 50mM

Concentration (Trp)

Figure-3A.5a: Effect of substrate concentrations and temperature on IAA production

3 Substrate Effect Fresh Weight Dry Weight 2.5

1.5

Weight (g) Weight 1

0.5

0 Control 5 10 15 20 25 50 Concentration mM (Trp)

3.5 pH pH 3

2.5

1.5 Weight (g) Weight 1

0.5

0 Control 5 10 15 20 25 50

Concentration mM (Trp)

Figure-3A.5b: Change in fresh weight, dry weight and pH with different concentrations of substrate (25 ± 2°C)

Temprature Effect Fresh Weight 2 Dry Weight 1.8 1.6 1.4 1.2 1 0.8

Weight (g) Weight 0.6 0.4 0.2 0 Control 5 10 15 20 25 50 Concentration mM (Trp)

2.9 pH pH 2.8 2.7

2.6 2.5 2.4 Weight (g) Weight 2.3 2.2 2.1 Control 5 10 15 20 25 50 Concentration mM (Trp)

Figure-3A.5c: Change in fresh weight, dry weight and pH with different concentrations of substrate (20 ±2°C)

Figure-3A.6a: Location of aldehyde dehydrogenase gene in A. niger genome (http://www.cadre-genomes.org.uk/index.html)

Figure-3A.6b: Agarose gel electrophoresis of 18s gene

Figure-3A.6c: Optimization of primer concentrations for ALDH gene (pmol/l)

Figure-3A.6d: Optimization of MgCl2 concentrations for ALDH gene (10

pmol/l) [D1- Primer pair 5 (0.5 mM MgCl2 ), D2- Primer

pair 4 (0.5 mM MgCl2 ), C1- Primer pair 5 (1.0 mM MgCl2 ), C2- Primer pair 4 (1.0 mM MgCl2 ), A1- Primer pair 5 (1.5 mM MgCl2 ), A2- Primer pair 4 (1.5 mM MgCl2 ), B1- Primer pair 5 (2.0 mM MgCl2 ), B2- Primer pair 4 (2.0 mM MgCl2 )]

Figure-3A.6e: Agarose gel electrophoresis of aldehyde dehydrogenase gene from A. niger [M- Marker, 1- Ald 9, 2- Ald 10, 3- Ald 7, 4-Ald 8]

Figure-3A.7a: Electropherogram for 18s gene retrieved from ABi 3130 genetic analyzer

Primer ALD- 9

Primer ALD-10

Figure-3A.7b: Electropherogram of Primer pair 5 (ALD-9 and ALD-10) retrieved from ABi 3130 genetic analyzer

Figure-3A.8a: Mutational analysis of ALDH sequence using SeqScape (XM_001398166.2)

Figure-3A8b: Mutational analysis of ALDH sequence using SeqScape (XM_001398829.2)

Figure-3A.9: 3D structure prediction by homology modeling of aldehyde dehydrogenase from A. niger

Table-3A.1: Specification of primers designed with the help of Primer3plus software

Sr. Name Sequence Tm °C Product No. Size

5’ ATG ACC AAA CCC AGA CTT ALD1 60.0 Primer CG3’ 232bp pair 1 5’ CTT GAT GAC GATGGTGTT ALD2 60.1 GC3’

5’ CGG GTC CAA GTT TGA TCT ALD3 60.0 Primer GT3’ 245bp pair 2 5’ TGT CAC TGT AAG CGG GTT ALD4 59.8 TG3’

5’ CGG GTC CAA GTT TGA TCT ALD5 60.0 Primer GT 3’ 221bp pair 3 5’ TGC AAA TGG CAT CAA GGT ALD6 59.7 AG 3’

5’ GAG TTT GTC CCC TCC CTC ALD7 60.0 Primer TC 3’ 241bp pair 4 5’ TGC AAA TGG CAT CAA GGT ALD8 59.7 AG3’

5’ ATG ACC AAA CCC AGA CTT ALD9 60.0 Primer CG3’ 245bp pair 5 5’ CCT TTT CGG AGG ACT TGA ALD10 59.7 TG3’ Table-3A.2: Genebank accession number of 18s and aldehyde 3- dehydrogenase gene from A. niger

Sr. No. Gene fragment Accession No.

1 18 s JF925334

Aldehyde 3- 2 JF925335 Dehydrogenase

Table-3A.3: Blastp result of aldehyde dehydrogenase gene from A. niger. in UniPro

Entry Gene Organism Identity Score E-value UniProt name Name ID

Aspergillus niger (strain A2QSI3_ An08g10 A2QSI3 CBS 513.88 100% 153 9x 10-11 ASPNC 820 / FGSC A1513)

Table-3A.4: Sequences used in mutational studied of ALDHs by ABI prism SeqScap

Sr. No. Accession No.

1 XM_001393149.1

2 XM_001393671.1

3 XM_001396908.1

4 XM_001397981.1

5 XM_001398834.1

6 XM_001402439.1

7 XM_001388469.2

8 XM_001389309.2

9 XM_001392807.2

10 XM_001392927.2

11 XM_001400734.2

12 XM_001400875.2

13 XM_001396676.2

14 XM_001397081.2

15 XM_001398166.2

16 XM_001398289.2

17 XM_001398829.2

18 XM_001401670.2 Table-3A.5: Physico-chemical analysis of ALDH protein sequences by ExPASy-ProtParam tool

Total No. Total No. Grand of of No. of Average Mol. Theoretical Negatively Positively Instability Aliphatic Sequences amino of weight pI charged charged index index acid Hydropathicity residues Residues (GRAVY) (Asp+Glu) (Arg+Lys)

80.96 JF925335 75 8323.5 8.27 4 6 74.13 0.040 (unstable)

INTRODUCTION

The phytohormone ABA regulates many of the physiologically important events in the adaptation of vegetative tissues to abiotic stresses, such as drought and high salinity, as well as in seed maturation and dormancy (Osakabe et al., 2005). ABA synthesis and accumulation are promoted by water stress in plant cells, and ABA triggers stomatal closure in guard cells and regulates the expression of many genes whose products may function in dehydration tolerance of vegetative tissues and seeds (McCourt, 2001).

ABA can induce short-term responses such as stomatal closure-minimizing transpirational water loss or long-term responses involving changes in gene expression (Himmelbach et al., 1998; Finkelstein et al., 2002; Liang and Harris, 2005). It plays important roles in many aspects of plant development, in the regulation of stomatal aperture, and in the initiation of adaptive responses to various environmental conditions. Adaptation to drought, low temperature and salinity is regulated by the combinatorial activity of interconnected ABA-dependent and ABA-independent signaling pathways. By contrast, the plant hormones salicylic acid (SA), jasmonic acid (JA) and ethylene (ET) play major roles in disease resistance (Mauch-Mani and Mauch, 2005).

ABA is well known as a stress hormone that is active in the adaptation to various forms of environmental stress in higher plants It might acts as a stress signal compound in some physiological responses in higher plants (Bray, 2002; Sarmad et al., 2007). Recently, mass production of ABA was brought to success by liquid fermentation of fungi Botrytis cinerea strains (Tan and Li, 1998; Tudzynski and Sharon 2002). Although some other fungi can also produce ABA, so far the commercial production has not been reported (Deng et al., 2007).

Macrophomina phaseolina is an imperfect fungus which causes charcoal rot in common beans, mainly under the dry and hot conditions of arid regions and other tropical and subtropical areas. Drought stress may occur in any 52║ P a g e

developmental stage of common bean and increases crop pre-deposition to infection by facultative parasites such as M. phaseolina, due to negative effect on host physiology (Mayek-Perez et al., 2002). M. phaseolina (Tassi) Goid, a fungal phytopathogen, infects about 500 plant species worldwide (Sett et al., 2000). Tropical crop plants are seriously affected by this pathogen (Malaguti, 1990). It elaborates a number of phytotoxins, namely asperlin, isoasperlin, phomalactone, phaseolinic acid, phomenon and phaseolinone (Dhar et al., 1982; Mahato et al., 1987; Bhattacharya et al., 1992a). Phaseolinone appears to be most important of these and it induces disease symptoms in plants similar to those produced by the pathogen (Bhattacharya et al., 1992b).

Among various pathogens, fungi constitute an important group as they inflict damage to crop plant at different stages (Agrios, 2000). Among the fungal diseases, the root - rot caused by Macrophomina remains to be a challenging task in terms of management, since it is soil - borne in nature. It is distributed worldwide and is prevalent in arid, sub-tropical and tropical climate, especially in the areas with low rainfall and high temperature (Ramezani, 2008).

Fusarium moniliforme (Sheld.) emend. Snyd. & Hans (Snyder and Hansen, 1945) has a wide host range and is widespread throughout the world; only heterothallic types are known. The fungus not only causes considerable damage to many plants, but also is parasitic on plants without production of visible symptoms (Hsieh et al., 1977). The Gibberella stage of the fungus was found first by Wineland (1924) when she put two compatible mating types together in culture. The gibberellin-producing strains of Gibberella fujikuroi are pathogens of rice (Oryza sativa) where they cause the disease known in Japan as bakanae (Cerda-Olmedo et al., 1994). Sawada (1917) recognized the causal role of the species in bakanae disease of rice and gave it the specific epithet fujikuroi in recognition of his colleague Fujikuro. In 1931, Ito and Kimura modified the name to Gibberella fujikuroi and wrote “the fungus seems to be identical with or only a variety of Gibberella moniliformis (Sheldon) Wineland (= Fusarium moniliforme Sheldon)” (Desjardins, 2003).

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The gibberellins, hormones made in small amounts by plants, are produced in large amounts by some strains of the ascomycete fungus Gibberella fujikuroi. Gibberellins promote growth and regulate various stages in plant development, from seed to fruit. Gibberellins isolated from the culture medium of the fungus find many practical applications in agriculture and the brewing industry (Takahashi et al., 1991; Mander, 1992).

Gibberellic acid (GA) is a naturally occurring plant growth regulator which may cause a variety of effects including stem elongation (Berrios et al., 2004). GA3 is produced commercially as an agrochemical and production strains under optimized conditions may be expected to produce many times more (Jacklin et al., 2000). They are a large family if tetracyclic diterpenoid compounds. Through phenotypic analysis of mutants with reduced GA production or sensitivity, it has been found that bioactive gas such as GA1 and GA4 play crucial roles in plant growth and development, including seed germination, leaf expansion, stem elongation and flower and fruit development (Hooley, 1994; Wen and Chang, 2002).

There are 126 known Gibberellic acids, divided into two classes i) those with 20 carbone atoms, ii) those with 19 carbone atoms, and many more may be discovered in the future (Soliman et al., 2010). GAs are secondary metabolites of the fungus G. fujikuroi and are a high value industrially important biochemical selling at 26-37 $/g in the international market, depending upon its purity and potency (Mitter et al., 2001; Srivastava et al., 2003).

A number of micro organisms have been reported to produce GA3 and GA like substances. Among these, the fungal cultures are able to produce GAs like activities in higher yields. Several strains, isolated from a variety of fungus- affected plants, were tested by Borrow et al. (1955) and only 9 cultures showed GA likes activity. Sanchez-Morroquin (1963) tested 43 strains of Fusarium species and reported that F. moniliforme was able to give higher yields of GA3 on a variety of media. The reports on phytohormones secretion by fungi to damage host plant are rather scanty. 54║ P a g e

This study was designed to evaluate production of ABA and GA in define media by fungi Fusarium moniliforme and Macrophomina phaseolina, respectively.

MATERIALS AND METHODS

Fungal cultures and inoculum preparation Fusarium moniliforme (MTCC-156) and Macrophomina phaseolina strain previously stored in distilled water was streaked on PDA plate and incubated up to seven days for culture activation. From that plate 1 cm diameter of fungal disc were cut with the help of cup borer and placed in the center of another plate. The plate was further incubated for seven days for complete radial growth and peripheral part of fungal mycelia was used as inoculums.

Preparation of media MS (1962) was used as define media. In the first part of experiment MS media with certain alteration were used. Main four types of media were designed as: (i) MS full strength with sucrose 2% [MS (2)]; (ii) MS full strength with sucrose 4% [MS (4)]; (iii) MS half strength with sucrose 2% [MS (1/2)(2)]; (iv) MS half strength with sucrose 4% [MS (1/2)(4)]. The culture was grown in dark and in static condition at 28 ± 2 ºC. The initial pH of define medium was 5.6±0.2. At interval of seven days some amount of filtrate was withdrawn and measured ABA/GA concentration. Second part of experiment evaluate the higher production of product in optimum media [MS (2)] pH and culture condition was same as above but the drawn time of filtrate was different in both the culture.

In case of F. moniliforme collection of filtrate was carried out at every seven days interval and on the other hand culture filtrate of M. phaseolina drained at interval of 24 h, simultaneously fresh weight, dry weight of mycelia and pH of medium was also noted with ± SD. Whole the experimentation was performed in triplicates. The produced ABA/GA concentration was calculated by plotting the value in standard curve of ABA/GA and express with ± SD.

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Raising of antibodies against ABA/GA Preparation of ABA-BSA/Casein conjugate Synthesis of ABA-BSA/ casein conjugate was carried out according to the protocol of Gokani and Thaker (2001). ABA (132 mg) was dissolved in 3 ml of the mixture of water and DMF (2:1) and pH adjusted to 8.0 with 1N NaOH. The solution was then added drop wise with gentle stirring to 250 mg of BSA/Casein dissolved in water and adjusted to pH 8.5. N-ethyl-N (3- dimethylaminopropul) carbodimide hydrochloride [EDC, 210 mg] was added into the preparation in four part at interval of 30 min and stirred in dark for 19 h at 4ºC, conjugate was finally dialyzed against distilled water for 4 days; final volume was made with distilled water and stored at 0ºC.

Preparation of GA-BSA/ Casein conjugates Plant growth regulators are haptens; therefore it is necessary to conjugate them with protein molecules i.e. Bovine Serum Albumin (BSA) or casein to make them immunogenic. GA3 was conjugated with BSA and or casein according to Gokani and Thaker (2002). GA3 was coupled to free amino group of BSA via a mixed anhydride reaction. GA3 (106 mg) and 75 l tributylamide were dissolved in 2.5 ml di-methyle Formemide (DMF) and cooled to 0°C. To this solution, 40 ml isobutylchlorocarbonate was added and the mixture was allowed to proceed for 20 min then, the solution was added, with through stirring to an ice- cold solution of 420 mg BSA in 22ml DMF/water (1:1 v/v). After 60 min 0.2 ml 1M NaOH was added and the mixture was stirred for 3 h and finally purified by dialysis against 10% aqueous DMF and 5 days against distilled water.

Immunization Two rabbits were selected for each hormone ABA/GA conjugate was mixed with an equal amount of Freund‟s incomplete adjuvant and injected to the rabbit (Thaker, 1995). Immunization was followed by intramuscular root. Booster injections were given periodically to raise the titer. Rabbits were bled (10 ml) at particular interval of time from marginal ear vein and serums were

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allowed to separate at room temperature, which was further used for IgG purification.

Purification of IgG Purification was accomplished by ion-exchange chromatography. - immunoglobulin (IgG) against GA and ABA were collected by passing the serum through Diethylaminoethyl (DEAE)-cellulose column pre- equilibrated with 0.01M phosphate buffer pH (8.0).The purified IgG was concentrated to the one third volume of serum and preserved at 0ºC in eppendroff and used for estimation after appropriate dilution.

Extraction of GA and ABA from samples The filtrate collected from the broth (MS) mixed with 5 ml of 80 % methanol (v/v). This mixture was stirred for 10 min and incubated for 48 h at 4°C in dark. The mixture was centrifuged at 10,000 xg for 10 min and supernatant was collected and evaporated to make it concentrated.

Estimation of ABA/GAcontent The ABA estimation was carried out as directed by Gokani and Thaker (2001) and estimation of GA content was carried out with the help of Indirect ELISA (Gokani and Thaker, 2002). ABA/GA –casein conjugate (300 l) was coated on ELISA plate and incubated for overnight at 4°C, followed by washing with Phosphate buffer saline-Tween-20 (PBS-T). The next step involved was blocking of free protein binding sites of well with egg albumin and incubated for 1 h at 37°C then the plate was washed thrice with PBS-T. Antibodies against GA/ABA mixed with samples were coated and incubated for 3 h at 37°C. Finally, the plate was coated with anti-rabbit IgG tagged with peroxidise and the colour was developed using o-phenylene diamine as a substrate. The reaction was terminated by addition of (50 l) 6 N sulphuric acid. After each coating, the ELISA plate was wash thoroughly with PBS containing 0.05 % Tween-20.

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Calibration curve of ABA A standard curve of ABA was prepared in a range of 100-800 ng/well for each plate and a value falling on the curve was calculated for each data point. To test the sensitivity of the assay each sample was mixed with known amount of ABA (400 ng) as an internal standard before reacting with the antibodies. Data were taken in triplicate and mean values of three replicates was calculated with ±SD.

Calibration curve of GA

To test the sensitivity of the assay internal standards (150 ng GA3) were mixed with samples. For preparation of calibration curve, standard ranged from 100-500 ng/well GA3 was mixed with an optimum dilution of antibodies (1: 10,000) instead of samples prior to coating on the plate. All estimations were done in triplicates and reported as mean value with ±SD.

RESULTS AND DISCUSSION

ABA production Plant pathogenic isolates of M. phaseolina have shown variation in morphological and physiological characteristics (Pearson et al., 1986) as well as in the pathogenicity or host speciﬁcity (Diourte et al., 1995; Mayek-Perez et al., 2001; Su et al., 2001; Reyes-Franco et al., 2006). Genetic variation has also been detected among M. phaseolina isolates using random ampliﬁed polymorphic DNA (RAPD) (Arias et al., 2011). These variations may have distinct pathogenicity in the host.

Amongst the four media tested for ABA production in M. posellina in different strength of MS salts and sucrose concentration MS (2) was found to be the best (Fig. 3B.1a) and concentration found to be 4752 g/L. Further study was carried out in MS (2) with time (Fig. 3B.2), pH, fresh weight and dry weight measurements. The filtrates were harvested everyday and the quantity of ABA measured. It was found that the amount of ABA was gradually increased

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every day and a highest value was recorded at seventh day i.e. 4.6±0.01 mg/L (Fig. 3B.2).

The fresh weights of culture mycelia was found 0.010, 0.064, 0.270, 0.533, 1.117, 2.143, 2.877 g/50ml and their relative dry weights was 0.001, 0.008, 0.030, 0.076, 0.131, 0.211, 0.274 g/50ml, respectively from 1st to 7th day (Fig. 3B.3a). The initial pH of the media was 5.6 but it was turned acidic up to 5th day (pH- 3.25) but at 6th day it reached to pH 5.0 and it was turned to 6.89 on 7th day (Fig. 3B.3b). The fresh weights and dry weights of the fungal mycelia were gradually increased with incubation time.

The direct role of toxins in pathogenesis induced by M. phaseolina has been questioned by Chan and Sackston (1969, 1973) and supported by the important role of cell wall dissolving enzymes in disease initiation and progression However, several groups had reported that the virulence of M. phaseolina isolates is related to their ability to produce toxins (Bhattacharya et al., 1992b). The role of toxin(s) produced by M. phaseolina in disease initiation has not yet been reported (Sett et al., 2000).

It has been suggested that ABA is one of the most effective plant hormones in terms of promoting leaf senescence. Application of ABA has been found to promote leaf senescence in a wide range of plant species (Hsu and Kao, 2010). An increase in endogenous ABA has been shown to coincide with senescence of leaves (Yang et al., 2002). Abiotic stress has a strong effect on ABA accumulation and is known to influence the outcome of plant–pathogen interactions (Mayek-Perez et al., 2002). Increased endogenous ABA levels were observed in response to infection with viruses, bacteria, and fungi (Whenham et al., 1986; Kettner and Dorffling, 1995). In addition, it is generally found that application of exogenous ABA increases the susceptibility of plants to fungal pathogens (Henfling et al., 1980; Ward et al., 1989; McDonald and Cahill, 1999). ABA also seems to interact with pathogen associated plant defense (Audenaert et al., 2002; Yasuda et al., 2008).

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ABA has been shown to be produced by members of different divisions of filamentous fungi, such as the basidiomycete Rhizoctonia solani, the ascomycete Ceratocystis fimbriata, and the zygomycete Rhizopus nigricans (Dorffling et al., 1984; Crocoll et al., 1991; Siewers et al., 2006). Many of the above-described species are involved in symbiotic or pathogenic interactions with plants. Therefore, the ability of microorganisms to produce plant hormones is thought to play a role during the establishment of these interactions (Siewers et al., 2006). The ABA pathway in fungi has been biochemically studied in the phytopathogenic ascomycetes Botrytis cinerea and several Cercospora species, and a direct route from farnesyl diphosphate has been postulated (Oritani and Kiyota, 2003).

ABA has huge economic potential and can be applied in agriculture and forestry because it considered to be involved in plant resistance to stresses such as cold, heat, salinity, drought, pathogens and wounding (Tudzynski and Sharon, 2002). The production yields reported of B. cinerea ABA were 0-1.6 g/L in axenic culture (Wu and Zheng, 1997; Tan and Li, 1998; Wu and Shi, 1998; Tudzynski and Sharon, 2002; Liang et al., 2004; Zhao et al., 2009).

GA production

In the present study GA3 estimation was also carried out from F. moniliforme in different strength of MS salts and sucrose concentration, filtrate was withdrawn at seven day and it was found that in the media tested MS (2) was the best for growth as well as for GA production, maximum concentration was recorded as 496 ± 0.01 g/L (Fig. 3B.1b). Further, F. moniliforme was grown in MS (2) and filtrate was harvested at interval of seven day up to a month. The GA estimation was carried out from that and fresh weight and dry weight was measured with change in pH at every interval. The fresh weight of culture mycelia was found 0.613, 1.707, 2.657, 2.308 g/200ml and their relative dry weights i.e. 0.058, 0.163, 0.235 and 0.259 g/200ml, respectively at 7th, 14th, 21st and 28th day (Fig. 3B.4a).

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The concentration GA3 in culture filtrate was evaluated (468.55 g/L) at seven day, (1689 g /L) 14 day, (402.16 g/L) 21 day and (368.76 g/L) at 28 day (Fig. 3B.5). Similar result was reported by Rachev and their co-workers (1997) in their work, GA3 concentration increased at 144 h and maximal quantities were observed in the interval between 216 and 264 h, in F. moniliforme.

In case of F. moniliforme the pH of the culture media slowly changed with the time (Fig. 3B.4b); initial pH of the media was 5.6, which reached to 8.5 on 28th day. The pH value was 6-6.6 when GA3 production was higher suggesting th thereby that it can be the optimum pH range for production of GA3. After 14 days the pH changed may be due to decreased sugar concentration (Darken et al., 1959). Qian et al. (1994) was also recorded maximum GAs accumulation after 10 to 12 days of incubation and usually marked by a sharp change in pH.

F. moniliforme pathogenicity is due to its ability to produce large quantities of a plant growth hormone, GA3. Under laboratory conditions the amount of GA3 produced by wild type strains of F. moniliforme appears to vary somewhat, but is in the range of 150 mg/L to 1.6 g/L (Jefferys, 1970; Bu‟Lock et al., 1974;

Brückner and Blechschmidt, 1991). GA3 is produced commercially as an agrochemical and production strains under optimized conditions may be expected to produce many times more (Jacklin et al., 2000).

Mander (1992) has given the structure of the 86 free natural gibberellins known at the time: 66 found only in plants, 11 found only in Gibberella, and the rest of gibberellins found in both. Gibberellic acid, often called GA3 and it is the main product in Gibberella (Cerda-Olmedo et al., 1994). The ability to synthesize gibberellins was detected in many micromycetes (Aspergillus flavus, A. niger, Penicillium corylophilum, P. cyclopium, P. funiculosum, Verticillum sp., Rhizopus stolonifer, and Schizophillum commune) (Tsavkelova et al., 2006a). Although GA formation had also been observed in Sphaceloma manihoticola (Rademacher and Graebe, 1979), Neurospora crassa (Kawanabe et al., 1983), Phaeosphaeria sp. (Kawaide and Sassa,

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1993), Aspergillus niger (Bilkay et al., 2010), Rhizobium Phaseoli (Bhattocharyya and Pati, 2000) and some other plant pathogens (Tudzynski, 1997). G. fujikuroi is unique because of the enormous quantities of GAs that it can secrete (Linnemannstons et al., 1999).

Since GA3 is known to loosen plant cell walls via hydrolytic enzymes like glycosidases and glucanase or due to turnover of wall polysaccharides (Leubner-Metzger et al., 1995; Voegele et al., 2011), possibility of pathogenic fungi to damage host through production of excess phytohormones cannot be ruled out. In this experiment, studied fungi showed very high amount of GA3 and ABA production, which may be one of the mechanism(s) to enter into the host. It is surprising that so little is known about the diversity of these fungi and their mechanisms of pathogenicity, including the role of ABA and GA in the infection.

CONCLUSIONS

In the best of knowledge no such study was carried out for production of ABA by M. phaseolina and on the basis of results it can be concluded that higher yield was obtained at the end of seventh day. On the other hand in case of F. moniliforme the maximum production was found at 14th day. However in filtrate GA concentration was higher at 14th day and decreases after that may be due to degradation of mycelia or change in pH. The amount produced by the fungi in given condition, can be improved with variation in media composition for commercially exploitation of these fungi.

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ABA production in different media 6

1 Concentration mg/L Concentration

0 MS 2 MS 4 MS(1/2) 2 MS(1/2) 4 Different media composition (MS)

Figure-3B.1a: ABA production in different MS and sucrose concentrations

GA production in different media 600

500 g/L  400

300

200 Concentration Concentration 100

0 MS 2 MS 4 MS(1/2) 2 MS (1/2) 4 Different media composition (MS)

Figure-3B.1b: GA production in different MS and sucrose concentrations

ABA production in MS(2) 5 4.5 4 3.5 3 2.5 2 1.5 1 0.5 Concentration of ABA mg/L ABA of Concentration 0 1 2 3 4 5 6 7 Time (Days)

Figure-3B.2: ABA production by M. phaseolina with time in MS (2) media

Fresh Weight 3.5 M. phaseolina Dry Weight 3.0

2.5

2.0

1.5 Weight (g) Weight 1.0

0.5

0.0 1 2 3 4 5 6 7

Time (Days)

Figure-3B.3a: Changes in fresh weight and dry weight during ABA production in M. phaseolina mycelia

pH 8 M. phaseolina 7 6

5 4 3 Weight (g) Weight 2 1 0 1 2 3 4 5 6 7

Time (Days)

Figure- 3B.3b: Changes in pH during ABA production in M. phaseolina

Fresh Weight 3.0 F. moliniforme Dry Weight 2.5

2.0

1.5

Weight (g) Weight 1.0

0.5

0.0 7 14 21 28

Time (Days)

Figure-3B.4a: Changes in fresh weight and dry weight during GA production in F. moliniforme mycelia

pH 10 F. moliniforme 9 8 7 6 5 4

Weight (g) Weight 3 2 1 0 7 14 21 28

Time (Days)

Figure-3B.4b: Changes in pH during GA production in F. moliniforme

GA production by F. moniliforme 1800

1600

g/L) 1400  1200 1000 800 600

Concentration ( Concentration 400 200 0 7 14 21 28

Time (Days)

Figure-3B.5: GA production by F. moniliforme with time in MS (2) media

INTRODUCTION

Effective management of a plant disease is a key to save plants from diseases caused from microbes, as they are both economical and aesthetic. Now, quick and effective management of plant diseases and microbial contamination in several agricultural commodities is generally achieved by the use of synthetic pesticides (Agrios, 1997). Despite serious environmental implications associated with their use, chemical fungicides remain the first line of defense against effective and efficient management of fungal pathogens. Many of these chemical fungicides pour tremendous environmental load, thereby adversely affecting the agro-ecosystem (Zadoks, 1993; Guleria and Kumar, 2009).

The recurrent and indiscriminate use of fungicides have posed a serious threat to human health and to the existing human eco geographical conditions as some of them have already been proved to be either mutagenic, carcinogenic or tetratogenic. But many pathogenic micro-organisms and insect pests have developed resistance against these chemical pesticides (Williams and Heymann, 1998; Witte, 1998; Yadav, 2010).

Biological control offers a potential alternative to chemical fungicides (El- Ghaouth, 1997; Dev and Dawande, 2010). Due to increased awareness about the risks involved in use of chemical pesticides, much attention is being focused on the alternative methods of pathogen control. The spiraling up cost of chemical fungicides particularly in those countries by the accumulation of obnoxious chemicals residues due to continuous use of fungicides and development of resistance races to these chemicals is therefore now facing the scientists to look for methods, which are ecologically friendly, safe and specific for pathogens (Joseph et al., 2008; Mazid et al., 2011).

Keeping in view the drawback of chemicals, the use of plant extracts in the management of plant diseases is gaining importance. Various plant products like plant extracts, essential oils, gum, resins etc. were shown to exert biological activity in vitro and in vivo and are used as bio-fungicidal 63║ P a g e

compounds (El-Mougy and Alhabeb, 2009; Fawzi et al., 2009; Al-Askar and Rashad, 2010).

Biological approaches for the control of pathogens on aerial surfaces have been worked out extensively over the past 30 years (Blakeman and Fokkema, 1982; Spurr and Knudsen, 1985; Andrews, 1992; Elad, 1993; Fokkema, 1993; Wilson, 1997; Haikal, 2007; Sahayaraj et al., 2011). During this period, most approaches employed for the biological control of diseases of aerial plant surfaces have concentrated on the use of a single, empirically-selected biocontrol agent to antagonize a single pathogen (Wilson, 1997).

Plants have been known for their medicinal and antimicrobial properties since ancient times. For this reason attention has been diverted to alternative; safe and economic methods for the management of pathogenic microorganisms from plant products (Khallil, 2001; Begum et al., 2010). The potential of plant extracts to control plant diseases has long been recognized (Ark and Thompson, 1959). There are approximately 250,000 species of higher plants, of which only 5%-15% have been studied for their therapeutic value (Rojas et al., 2003). The use of plants for human disease control attracts more attention, compared with its use in that of plant and animal (Newton et al., 2002; Cano and Volpato, 2004). In crop protection studies, various natural plant products have been identified and employed to control postharvest diseases (Mekbib et al., 2007; Ilondu, 2011).

Aromatic plants have been widely used in folk medicine. It is also known that most of their properties are due to their volatile oils. EOs and extracts obtained from many plants have recently gained a great popularity and scientific interest. The antimicrobial activity of EOs from oregano, thyme, sage, rosemary, clove, coriander, cinnamon, garlic and onion have been investigated and reviewed (Gill et al., 2002; Burt, 2004; Holley and Patel, 2005; Tepe et al., 2010). A survey of literature reveals that there are many EOs which possess antifungal activity (Soliman and Badeaa, 2002; Kalemba and Kunicka, 2003; Pinto et al., 2006; Tullio et al., 2007; Bansod and Rai, 2008; Singh, 2010). Screening for antimicrobial activity has been the subject 64║ P a g e

of many investigations and oils with very potent antibacterial and antifungal activity could be promising agents for the future more extensive research and in vivo examination.

The activity of natural plant products on the host tissue may involve direct interaction with the pathogen or induction of host resistance; the mechanism involved in the former, however, is the less understood one. Host resistance induction, on the other hand, may involve several complex mechanisms including hypersensitive responses, buildup of cell wall barriers, increased production of phytoalexins, accumulation of pathogenesis-related (PR) proteins, and fungal cell wall hydrolases (Fallik and Grinberg, 1992; Porat et al., 2002).

Agave is a plant which produces medically useful substances such as vitamins and steroid precursors like saponins and fructans (Gentry, 1982; Sanchez et al., 2005; Guleria and Kumar, 2009). The antifungal activity of Agave is due to steroidal saponins (Yang et al., 2006). Saponins are secondary plant metabolites that occur in a wide range of plant species (Hostettmann and Marston, 1995). The natural role of saponins in plants is to provide protection against attack by pathogens and pests (Morrissey and Osbourn, 1999). The antifungal activity of steroidal saponins, particularly against agricultural pathogens, has been known for a long time (Wolters, 1966; Imai et al., 1967; Dimoglo et al., 1985; Sahu et al., 2008; Zwane et al., 2011).

While other reported activities for this class of compounds include antibacterial (Chattopadhyay et al., 2001), antiinsect (Kozukue et al., 2004) and antiyeast activities (Miyakoshi et al., 2000). The antifungal potency of these compounds is not associated with cytotoxicity to mammalians cells (Dini et al., 2001). Because of their anti-pest properties, saponins can be utilized for the development of insecticidal and fungicidal preparations (Sparg et al., 2004).

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Natural plant products are usually inherently less toxic than conventional fungicides, often are effective in very small quantities, and generally decompose quickly, thereby resulting in lower exposures and largely avoiding pollution problems (Huang and Chou, 2005; Gupta and Dikshit, 2010). Considering to these issues, present study was undertaken to evaluate the effectiveness of the extract from five species of Agave namely A. americana, A. ferox, A. montana, A. scabra and A. marginata on the growth of eight phathogenic fungi: Postia placenta, Macrophomina phaseolina, Alternaria porii, Alternaria alternata, Aspergillus awamorii, Aspergillus niger, Fusarium udum, and Fusarium solani. As EOs are eco-friendly and safe, another objective of present study was to screen the antifungal activity of some selected EOs to find a potent one against Fusarium moniliforme.

MATERIALS AND METHOD

Agave extract as biocontrol Preparation of fungal inoculum The fungus Macrophomina phaseolina, Alternaria porii, Aspergillus niger, Fusarium solani were isolated from their host. Postia placenta (MTCC-144), Aspergillus awamorii (MTCC-548), Alternaria alternata (MTCC-1362) and Fusarium udum (MTCC-2204) were obtained from from Microbial Type Culture Collection (MTCC), Institute of Microbial Technology (IMTECH), Chandigarh, India. The lyophilized culture was revived on suitable media than streak that activated culture on PDA plate. After seven days of growth a disc of fungal culture was cut with 1 cm borer diameter and place in center of the other PDA plate. A disc from the peripheral margin of the seven days old growth of all fungi was used as inoculum.

Preparation of leaves extract The leaves were collected and washed with cleaned water. Fresh weights, 250 g, were measured and kept to dry at 60°C temperature. Dried leaves grounded into fine powder and extracted with 80% methanol for two days. The plant debris was removed twice by centrifuging at 5000 xg for 20 min. The supernatants were collected and allowed it to evaporate, then dissolved in 66║ P a g e

final volume (i.e. 10 ml) of sterile distilled water. These stock solutions were used for an antifungal activity against all fungi by media poisoning method.

Preparation of media The extracts of all the plants to be tested was mixed with PDA at 0.5 ml/20 ml distilled water. The final volume was made up to 20 ml in glass tubes. The media in the tubes was autoclaved at 15 lbs pressure and 121 C temperatures for 15 min. The sterilized medium (20 ml) was then poured into the radiation sterile petri plates (90mm 15mm diameter) under aseptic condition.

Inoculation of fungi The centre of each poisoned PDA plate (with plant extracts) was inoculated with one fungal disc (10 mm diameter) of seven day old cultured plate. Control plates contained no plant extracts. The plates were incubated at 28 ± 2°C for seven days. The antifungal activities of plant extracts were observed at every 24 h and noted at 7th day of incubation by measuring the diameter of test and control in millimeter. The toxicity of plant extracts were recorded in terms of % mycelial inhibition (Khamis et al., 2005) against the test fungi. The percent inhibition of hyphal growth was calculated based on the following formula:

Percent inhibition = Diameter of fungal colony in treatments X 100 % Diameter of fungal colony in control

Essential oil as biocontrol Test Fungus The test fungus was obtained from IMTECH. A lyophilized culture of Fusarium moniliforme (MTCC-156) was activated on their required medium at 28 ± 2°C. Then this culture was maintained on PDA medium at 4°C.

Culture suspension A disc (1cm) from a one-week-old culture of the fungi was cut from periphery, and suspended in 10 ml of sterile distilled water. It was shaken vigorously and

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centrifuged to remove the agar particles. The supernatant was used as a culture suspension for spreading.

Essential oils All the EOs used in this experiment was obtained from Vimal Research Society for Agro-biotech and Cosmic Powers (VIRSACO), Rajkot, Gujarat, India. It is listed in Table-4A.1 with their scientific and common name.

Antifungal activity PDA medium was used for the culture maintenance and the bioassay. Antifungal activity testing was done using standard disc diffusion assays (Bauer et al., 1966). The prepared inoculum (0.1 ml) was spread on PDA plates. The test disc contain 5 l of essential oil pipette onto 5 mm filter paper discs made from Whatman filter paper no. 1, which were carefully transferred onto the surface of seeded agar plates. The plates were incubated at 28 ± 2°C for 72 h and the diameter of zone of inhibition were measured in mm after the incubation period. The antifungal activities of the oils reported were given in Table-4A.2. The experiment was performed in triplicate and the values expressed as mean ± SD.

Gas chromatography-mass spectrometry (GC-MS) analysis of essential oils The essential oil of Cinnamomum zeylanicum (bark) was subjected to gas chromatography-mass spectrometry (GC-MS) analysis on Perkin Elmer Auto System XL. Gas chromatograph equipped with on Perkin Elmer Turbo Mass spectrometer as a detector and the NIST data base. The capillary column was DB-1(30 m X 0.25 mm, 0.25 m film thickness) while helium was used as a carrier gas. Flow rate was 1 ml/min and temperature of injector and detector was 250 and 280°C, respectively. The oven temperature was programmed from 60°C to 250°C at 5°C /min. The amount of oil injected into the GC-MS was 0.4 l. The injector operated at split mode (1:40) was held at 250 °C. The sample transfer line was heated to 280°C. The identification of oil components was based on their retention time and by comparison of their mass spectral

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fragmentation pattern with NIST data base. The components from GC-MS analysis are listed in Table-4A.3, 4A.4, 4A.5 (only those components in order with the highest percentage area in the oil were subjected to mass spectral analysis) (Fig. 4A.5, 4A.6, 4A.7). The GC-MS analysis was accomplished in SIKART, Vallabh Vidyanagar, Gujarat, India.

RESULTS AND DISCUSSION

Agave extract as biocontrol The inappropriate use of agrochemicals especially fungicides were found to possess adverse effects on ecosystems and a possible carcinogenic risk than insecticides and herbicides together (Osman and Al-Rehiayam, 2003; Siva et al., 2008). Moreover, resistance by pathogens to fungicides has rendered certain fungicides ineffective (Zhonghua and Michailides, 2005; Aly et al., 2010). Suggesting, a need to develop new management systems to reduce the dependence on the synthetic agrochemicals.

Recent trends favor the use of alternative substances derived from natural plant extracts to control pests (Xan et al., 2003; Fawzi et al., 2009). Plants will continue to provide novel products as well as chemical models for new drugs in the future since the chemistry of the active extracts from the majority of plant species has yet to be characterized (Cox and Balick, 1994; Rout et al., 2009). The Agave species analyzed were potential candidates for broadly active antifungal compounds. In the present studies, the extracts of the five species of Agave showed antifungal activities against the six pathogenic fungi amongst the tested by media poisoning method (Plate 3-10) and they failed to inhibit two of them A. niger and A. alternata (Fig. 4A.3 and 4A.4, Plate 8 and 10), and the inhibitory levels were different in all cases.

The radial growth assay was performed and results were noted down. At the end of seventh day of growth period the highest percent inhibition was exhibited in P. placenta by A. montana (69.31%), followed by A. marginata (65.84%), A. americana (64.03%), A. ferox (63.89%), and A. scabra (56.67%)

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(Fig. 4A.1, Plate 3) compared to other fungi tested. So the five selected species of Agave showed more than 50% inhibition in P. placenta (Fig. 4A.1).

M. phaseolina was also exhibit the higer inhibition in all the extracts. The highest inhibition was observed in A. marginata (64.75%) followed by A. americana (61.69%), A. ferox (59.39%), A. montana (58.24%) and lowest as 48.66 % in A. scabra (Fig. 4A.1, Plate 4). Bobbarala et al. (2009) reported in their study that significant reduction in growth of M. phaseolina with extracts of forty nine plants. M. phaseolina suggests as non-host specific fungus. Physiological specialization of the fungus is not well demonstrated. High level of variation in morphology, physiology and pathogenesis has been reported even when isolated from different parts of the same plant (Dhingra and Sinclair, 1973; Khan, 2007).

After that A. awamorii showed higher growth inhibition by A. americana (34.44%), and A. montana (34.17%) followed by A. ferox (30.00%). There was no significant inhibition observed in A. marginata (18.06%) and A. scabra failed to cause inhibition in it. In A. niger no inhibition was observed in any extracts (Fig. 4A.3, Plate 7).

F. solani showed maximum inhibition in A. ferox (39.34%) followed by A. americana (38.86%), A. montana (35.55%), A. scabra (31.75%) and A. marginata (22.27%) (Fig. 4A.2, Plate 5). On the other hand F. udum showed less effect of these extracts, A. scabra extract found ineffective. Amongst these extract highest activity showed in A. marginata (27.61%), followed by A. montana (26.12%), A. ferox (23.88%), and minute in A. americana (8.00%) (Fig. 4A.2, Plate 6).

A. porii showed 29.05% inhibition in growth by A. ferox, A. americana only able to inhibit 10.14% of the hyphal growth and negligible inhibition was by A. marginata (5.41%). A. montana and A. scabra act as a growth promoting agents (Fig. 4A.4, Plate 9). On the other hand, in case of A. alternata none of the extract showed an inhibitory effect and has the promotery effect (Fig. 4A.4, Plate 10). Totally contradictory to our finding, Guleria and Kumar (2009) 70║ P a g e

has been reported that leaf extract of A. americana showed the antifungal activity against Alternaria brassicae, casual agent of Alternaria blight of Brassica juncea. The inhibitory effect of the plant extracts might be attributed to the presence of antifungal compounds. Currently, only limited information is available regarding the biological activity of compounds isolated from Agave sp. Many workers have reported the antifungal and antimicrobial activity of A. americana (Pandey et al., 1992; Jin et al., 2002; Guleria and Kumar, 2009; Chetan et al., 2010; Khan et al., 2010) while very few have been carried out on other species of Agave (Abdel-Khalik et al., 2002; Verastegui et al., 2008; Santos et al., 2009; Hammuel et al., 2011).

A. lophanta has steroidal saponins with activity against stomach ulcers and anti-inflammatory properties (Abdel-Khalik et al., 2002). Antifungal activity of steroidal saponins from A. americana has been reported by Yang et al. (2006) against Candida albicans, C. glabrata and Aspergillus fumigatus. The antimicrobial activity of A. lecheguilla against pathogens such as Clostridium perfringens, Salmonella enteritidis, Proteus vulgaris, Y. enterocolitica, Actinomycetes, and molds has been also reported (Verastegui et al., 1996). Verastegui et al. (2008) was also reported the antimicrobial activity of A. scabra and A. picta extracts in E. coli, L. monocytogenes, S. aureus and V. cholera.

Saponins also exhibit anticholesterolemic, anti-cancer (Haridas et al., 2001; Afrose et al., 2009), adjuvant (Behboudi et al., 1999; Ragupathi et al., 2010), and haemolytic (Oh et al., 2000; Hassan et al., 2010) activities. Some saponins have negative effects and were detrimental to human health; steroidal glycoalkaloids can be toxic when they were ingested (Friedman, 2002). Saponins produced complexes with sterols and cause sterol- dependent membrane permeabilization (Morrissey and Osbourn, 1999). The antifungal activity of saponins was generally attributed to this membrane permeabilizing property. Some fungi were resistant to the toxic effects of saponins because they have little or no sterols in their membranes, while others produce enzymes which specifically detoxify the saponins of their host plant (Osbourn et al., 1995; Ito et al., 2004; Coleman et al., 2010).

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Natural chemicals and their use for integrated plant protection is one of focus of the research workers all over the world (Kiran et al., 2006; Joseph et al., 2008; Chandler et al., 2011). The results of present investigation are clear indication for the potential of plant extracts and their compounds can be used to control fungal pathogens. It is evident from the results that all the plant extracts significantly inhibited the radial growth of isolated fungus. The formulation studies of the plant extracts can be successfully devised as fungicides using a simple process with minimum instrumentation and few chemical agents.

An interesting observation was that some of the test chemicals showed stimulation rather than inhibition. Stimulation of radial mycelial growth of fungi with test chemicals may be due to utilization of the chemicals or/of their degradative products (by fungal enzymes) by the fungus for their growth and development, for that the mode of action should be worked out. The mode of action of saponins in imparting antifungal activity is not fully understood, according to one possible mechanism their activity could be due to their ability to form complexes with sterol constituents of fungal cell membranes, leading to the loss of membrane structure (Keukens et al., 1995; Morrissey and Osbourn, 1999; Simons et al., 2006; Shukla et al., 2011).

In view of the harmful effects associated with the use of synthetic chemical fungicides, the worldwide trend calls for environmentally safe methods of plant disease control such as biological control, use of induced resistance by biotic and abiotic means (Lyon et al., 1995; Guleria et al., 2005; Guleria and Kumar 2006a) and the use of biodegradable natural products especially from medicinal plants (Prithiviraj and Singh, 1995; Guleria and Kumar, 2006b; Begum et al., 2010) in sustainable agriculture and the present work may add one more building block in this research fields.

Essential oil as biocontrol Fusarium moniliforme occurs on a great variety of crop plants and is one of the prevalent fungi associated with corn worldwide (Tseng, 1990). The production of maize (Zea mays L.) for grain is affected by many diseases, 72║ P a g e

where ear rot, caused by F. moniliforme, is one of the most destructive diseases because of its direct effects on grain yield losses. In addition, the ingestion of infected grain can cause severe adverse effects in both humans and animals due to the diverse and potent mycotoxins produced (Zhang et al., 2006b).

The EOs and extracts of many plant species have become popular in recent years and attempts to characterize their bioactive principles have recently gained momentum (Alim et al., 2009). EOs from plant edible parts which are ecofriendly in nature have been used by several workers for controlling pathogenic fungi, bacteria, viruses and insect pests (Singh and Upadhyaya, 1993; Singh, 1996; Singh, 2001; Samy, 2005; Wannissorna et al., 2005; Melendez and Capriles, 2006; Lalitha et al., 2011). The main reasons for using EOs as antifungal agents is their natural origin and low chance of pathogens developing resistance.

The complexicity of EOs are attributed to their terpene hydrocarbons and their oxygenated derivatives such as alcohols, aldehydes, ketones, acids and esters (Tzortazakis and Economakis, 2007). Phenolic compounds present in EOs have been recognized as the bioactive components for the antimicrobial activity (Chanthaphon et al., 2008). A good number of EOs are reported to be effective against many phytopathogenic fungi (Srivatsava and Singh, 2001; Zerroug et al., 2011). But only limited information exists about activity toward human fungal pathogens. They have been empirically used as antimicrobial agents, but the mechanisms of action are still unknown (Pinto et al., 2006; Shirzad et al., 2011).

It was observed that out of the sixty five EOs used to screen their antifungal activity; F. moniliforme showed inhibition by twenty four oils (Table-4A.2). Amongst them it found to be highly susceptible to the three oils, Cinnamomum zeylanicum (75.8 ± 3.1 mm), Cinnamomum cassia (72.5 ± 2.6 mm) Syzygium aromaticum (49.3 ± 6.2 mm). in case of Cedrus deodara, Apium graveolens , Thymus Vulgaris and Laurus nobilis zone of inhibition was in between (37-32 ± 1 mm) respectively (Table-4A.2) (Plate 11-14). 73║ P a g e

Other oils like Ocimum basilicum, Matricaria chamomilla, Cymbopogon citratus Pelargonium graveolens, Anethum sowa, Carum carvi, Angelica archangelica, Cymbopogon martini, Gaultheria frangrantissima, Myristica fragrans, Viola odorata, Alpinia galangal showed moderate activity in between 26-10 ± 2 mm. However, rest of the oils showed activity 10 mm or below than that and may consider as ineffective (Table-4A.2). From the sixty five oils tested for their antifungal activity Cinnamomum zeylanicum (bark) was found to be a potent inhibitor of F. moniliforme. Nanasombat and Wimuttigosol (2011) reported antimicrobial activity of certain EOs like anise, bastard cardamom, cinnamon, dill, mace, zedoary, prikhom, and bitter ginger oils against 16 microorganism and among them cinnamon oil exhibited complete inhibition towards F. moniliforme.

EOs such as aniseed, calamus, camphor, cedarwood, cinnamon, citronella, clove, eucalyptus, geranium, lavender, lemon, lemongrass, lime, mint, nutmeg, orange, palmarosa, rosemary, basil, vetiver and wintergreen have been traditionally used by people for various purposes in different parts of the world (Prabuseenivasan et al., 2006). Cinnamon, clove rosemary and lavender oils had shown antibacterial and antifungal activity (Ouattara et al., 1997; Wilkinson and Cavanagh, 2005; Rana et al., 2011); Anti-inflammatory activity has been found in basil (Singh and Majumdar, 1999; Thring et al., 2011). Lemon and rosemary oils possess antioxidant property (Altiok et al., 2010). Peppermint and orange oils have shown anticancer activity (Kumar et al., 2004). Lime oil has shown immunomodulatory effect in humans (Arias and Ramon-Laca, 2005). Lavender oil has shown antibacterial and antifungal activity; it was also found to be effective to treat burns and insect bites (Cavanagh and Wilkinson, 2002).

The inhibitory effects of sixty five EOs in their commercial forms against F. moniliforme were studied. In the present study three EOs exhibited the highest antifungal activity against the tested fungi. Both the Cinnamon oils were found the most effective oils and their effect was more pronounced than the other one. So further GC-MS analysis was carried out to know which compound were present in that (Table-4A.3, 4A.4). A chromatogram of 74║ P a g e

Cinnamon bark oil is shown in Fig. 4A.5. The highest % area of Cinnamon bark was occupied by Cinnamaldehyde (64.13) after that -linaloo (10.25), Benzyl benzoate (9.26), Eugenol (9.04), Cinnamyl acetate (4.47), others (2.1) and Eugenyl acetate (0.75) respectively its retention time was given in Table- 4A.3. In case of Cinnamon cassia the highest % area was also occupied by Cinnamaldehyde (66.36) after that -linaloo (9.16), Benzyl benzoate (9.26), Eugenol (7.74), Cinnamyl acetate (3.97), others (1.63) and Eugenyl acetate (0.52) respectively its retention time was given in Table-4A.4, Fig. 4A.6.

In both the aforesaid oils Cinnamaldehyde was found in higher percent. Recent study was suggested that the antifungal activity of C. zeylanicum and C. cassia was probably due to Cinnamaldehyde. Cinnamaldehyde and Eugenol exhibited strong antifungal activity against many white-rot fungi and other plant pathogenic fungi (Cheng et al., 2008; Cheng et al., 2011). The antimicrobial properties of some spices of Cinnamon and their components have been documented and the studies confirm that cinnamon inhibits the growth of both Gram positive and Gram negative food borne pathogens or spoilage bacteria, yeast and molds (Aneja et al., 2009).

Clove is found to be the third highest antifungal agent in this study GC-MS data shown in Table-4A.5 six peak was analyzed (Fig. 4A.7) that the highest percent area was covered by Eugenol (47.64), followed Benzyl alcohol (34.10), 3allyl-6-Methoxyphenol (4.98), Caryophyllene oxide (1.35), Caryophyllene (1.0) and Vanillin (0.91) (Table-4A.5). It is reported by some workers that eugenol is major components in that are known to be potent antifungal materials (Beg and Ahmad, 2002; Ranasinghe et al., 2002; Matan and Matan, 2007). Cedar wood posses the fourth position as an antifungal agent against F. moniliforme, the essential oil from this plant has been reported to possess antiinflammatory, antitumor, antioxidant, immunomodulatory properties and antifungal activity (Bouchra et al., 2003; Barrero et al., 2005; Gupta et al., 2011). Rest of the oils seems least effective to the test fungi although they may effective to other fungus or micro organisms.

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Recently fungal plant diseases are usually controlled by application of fungicides. Among the different management strategies used in controlling fungal plant diseases, chemicals rule the roost. However extensive use of chemicals as antifungal agents might lead to severe side effects such as carcinogenicity, terotogenecity, oncogenecity and other genotoxic properties. Further extensive use of chemicals leads to biohazardous effects on ecosystem (Prasad et al., 2010). Thus alternative approaches are preferred which are eco-friendly. More over, the essential oil as such may be exploited for their fungitoxic potency because of the synergistic activity of their different compounds (Tripathi et al., 2004; Sharma and Tripathi, 2006).

It is evident from the reports that plant extracts and plant EOs are effective antimicrobial agents foliar pathogens and do not produce any residual effects. They are non-pollutive, cost effective, non hazardous easily available and not disturbing the ecological balance. Due to their biodegradability and favorable safety profiles, these plant base antimicrobial agents can play a vital role in achieving evergreen revolution (Dubey et al., 2010).

CONCLUSIONS

The present results are important for the development of a bio-fungicide preparation from Agave species for the management of fungal pathogens of crop plants. However, this activity could differ depending on the method used to determine it and on its applicability in different processes. The biological activity exhibited by the Agave species and their wide geographic distribution suggest that, in the future, Agaves may have applications important to human health and medicine in addition to their traditional uses.

The results indicate that EOs could find a practical and rational use in the inhibition of fungal growth. Particularly, C. zeylanicum essential oil possesses strong antifungal activity against F. moniliforme. The broad inhibition of fungal growth by C. zeylanicum essential oil, in addition to its availability as natural volatile product, justifies its possible rational use as an alternative antifungal compound to control the growth and dissemination of pathogen and 76║ P a g e

proved to be an efficient bio-control agent. As they have complex structure the activity towards different organisms are differ because of the active component present in that. In future it will used to cure diseases caused in plants as well as in animals and humans. But for that further in vivo study was needed to check their side effects. Their mode of action towards pathogen, each compound present in that will investigate for their individual activity against the fungi. It may use as a bio control agent to cure diseases caused by the pathogenic fungi.

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% inhibition in hyphal growth 100 (Postia placenta) 90 80 70 60 50 40 30 % inhibition% 20 10 0

Agave species

% inhibition in hyphal growth 100 (Macrophomina phaseolina)

90 80 70 60 50 40 30 % inhibition% 20 10 0

Agave species

Figure-4A.1: Percent inhibition in hyphal growth of Postia placenta and Macrophomina phaseolina by different species of Agave

% inhibition in hyphal growth 100 (Fusarium soloni) 90 80 70 60 50 40 30

% inhibition% 20 10 0

Agave species

% inhibition in hyphal growth 100 (Fusarium udum) 90 80 70 60 50 40 30 20 10

% inhibition% 0

Agave species

Figure-4A.2: Percent inhibition in hyphal growth of Fusarium solani and Fusarium udum by different species of Agave

% inhibition in hyphal growth 100 (Aspergillus awamorii) 90 80 70 60 50 40 30 % inhibition% 20 10 0

Agave species

% inhibition in hyphal growth 100 (Aspergillus niger) 90 80 70 60 50 40 30 20

% inhibition% 10 0

Agave species

Figure-4A.3: Percent inhibition in hyphal growth of Aspergillus awamorii and Aspergillus niger by different species of Agave

% inhibition in hyphal growth 100 (Alternaria porii) 90

80 70 60 50 40 30 20 % inhibition% 10 0 -10 -20

Agave species

% inhibition of hyphal growth (Alternaria alternata) 0 -5 -10 -15 -20 -25

% inhibition% -30 -35 -40 -45 Agave species

Figure-4A.4: Percent inhibition in hyphal growth of Alternaria porii and Alternaria alternata by different species of Agave

Table-4A.1: List of the essential oils with their scientific and common name

No. Botanical Name Common Name

1 Citrus limon (Linn.) Burm.f. (Rutaceae) lemon

2 Moringa oleifera Lamk. (Moringaceae) Drumstick

Cinnamomum camphora (Linn.) Presl. 3 Camphor (Lauraceae) Cedrus deodara (Roxb. Ex D.Don) 4 Cedar wood G.Don (Pinaceae)

5 Rosmarinus officinalis L. (Lamiaceae) Rosemary

Cinnamomum zeylanicum Blume 6 Cinnamon bark (Lauraceae)

7 Cinnamomum cassia Blume (Lauraceae) Cassia

8 Citrus aurantium var.amara (Rutaceae) Petitgrain

Prunus armeniaca L. 9 Apricot (Rosaceae)

10 Thymus Vulgaris L. (Lamiaceae) Thyme

11 Coriandrum sativum L. (Apiaceae) Coriender

12 Zingiber officinale Rosc (Zingiberaceae) Ginger

Citrus bergamia Risso and Poit 13 Bergamoth (Rutaceae) Cupressus sempervirens L. 14 Cypress (Cupressaceae) Syzygium aromaticum (L.) 15 Clove bud Merrill & Perry (Myrtaceae) Pelargonium graveolens L'Hér. 16 Geranium (Geraniaceae) 17 Daucus carota L. (Apiaceae) Carrot

Nardostachys jatamansi (D.Don) DC. 18 Jatamashi (Valerianaceae)

19 Melaleuca alternifolia L. (Myrtaceae) Tea tree

20 Citrus paradisi Macfad. (Rutaceae) Grapefruit

21 Cuminum cyminum L. (Apiaceae) Cumin

22 Laurus nobilis L. (Lauraceae) Bay

23 Piper nigrum L. (Piperaceae) Black Pepper

24 Melaleuca viridiflora L. (Myrtaceae) Niaouli

Hamamelis virgianiana 25 Witch hazzel L.(Hamamelidaceae) Pogostemon cablin Benth. 26 Patchouli (Lamiaceae)

27 Angelica archangelica L.(Apiaceae) Angelica

28 Jasminum grandiflorum L. (Oleaceae) Jasmin

Cymbopogon citratus (DC.) 29 Lemon grass Stapf (Poaceae)

30 Matricaria chamomilla L. (Asteraceae) German Chamomile

31 Vitis vinifera L.(Vitaceae) Grapes

32 Anethum sowa Kurz. (Apiaceae) Anithi

33 Saussurea lappa Decne (Asteraceae) Upleth(Costus)

34 Carum carvi L. (Apiaceae) Caraway

35 Pimpinella anisum L. (Apiaceae) Anise

36 Myristica fragrans Houtt. (Myristicaceae) Mace

37 Papaver somniferum L. (Papaveraceae) Khash

38 Apium graveolens L. (Apiaceae) Ajamod 39 Cyperus rotundus L. (Cyperaceae) Nagarmoth

40 Myristica fragrans Houtt. (Myristicaceae) Nutmeg

41 Mentha piperita L. (Lamiaceae) Peppermint

Cananga odoranta (Lam.) Hook.f. and 42 Yalng ylang Thoms. (Annonaceae) Cymbopogon martini (Roxb.) Wats. 43 Palmarosa (Poaceae)

44 Litsea cubeba L. (Lauraceae) Cubeb

Barringtonia acutangula (Linn.) 45 Myrht Gaertn.(Barringtoniaceae)

46 Aniba rosaeodora Ducke (Lauraceae) Rose wood

47 Lavandula angustifolia Mill. (Lamiaceae) Lavender

Hedychium spicatum (Buch.) Ham 48 Kapura kachari (Zingiberaceae)

49 Daucus carota L. (Apiaceae) Carrot seed

50 Calendula officinalis L. (Asteraceae) Calendula

Gaultheria frangrantissima Wall. 51 Winter green (Ericaceae)

52 Viola odorata L. (Violaceae) Violet

Alpinia galangal L. Willd. 53 Ghodawaj (Zingiberaceae)

54 Foeniculum Vulgare Miller (Apiaceae) Fennel

55 Jatropha curcas L. (Euphorbiaceae) Nepalo

56 Salvia sclarea L. (Lamiaceae) Clary sage

Hydnocarpus laurifolia (Dennst.) 57 Mogra Sleummer (Flacourtiaceae)

58 Sesamum indicum L. (Pedaliaceae) Sesame 59 Juniperus communis L. (Cupressaceae) Juniperberry

60 Olea europaea L. (Oleaceae) Olive

Simmondsia chinensis (Link) C. K. 61 Jojoba Schneid.

62 Syzygium cumini L. Skeels. (Myrtaceae) Ravano

63 Ocimum basilicum L. (Lamiaceae) Basil

64 Persea Americana Mill. (Lauraceae) Avocado

65 Curcuma longa L. (Zingiberaceae) Turmeric

Table-4A.2: Antifungal zone of different essential oils against F. moniliforme

Common Zone of No. Botanical Name name inhibition (mm)

Cedrus deodara (Roxb. Ex D.Don) Cedar wood 4 37.5 ± 1.9 G.Don (Pinaceae) distilled Cinnamomum zeylanicum Blume 6 Cinamon bark 75.8 ± 3.1 (Lauraceae) Cinnamomum cassia Blume 7 Cassia 72.5 ± 2.6 (Lauraceae)

10 Thymus Vulgaris L. (Lamiaceae) Thyme 33.8 ± 4.3 Syzygium aromaticum (L.) 15 Clove bud 49.3 ± 6.2 Merrill & Perry (Myrtaceae) Pelargonium graveolens L'Hér. 16 Geranium 16.3 ± 2.6 (Geraniaceae) Melaleuca alternifolia L. 19 Tea tree 7.5 ± 1.3 (Myrtaceae)

21 Cuminum cyminum L. (Apiaceae) Cumin 8 ± 1.4 22 Laurus nobilis L. (Lauraceae) Bay 33.3 ± 3.3 Angelica archangelica L. 27 Angelica 12 ± 2.1 (Apiaceae) Cymbopogon citratus (DC.) 29 Lemon grass 21.8 ± 2.8 Stapf (Poaceae) Matricaria chamomilla L. German 30 22.3 ± 5.6 (Asteraceae) Chamomile

32 Anethum sowa Kurz. (Apiaceae) Anithi 19.3 ±3.3 34 Carum carvi L. (Apiaceae) Caraway 20.8 ± 2.5 38 Apium graveolens L. (Apiaceae) Ajamod 34 ± 3.9 Myristica fragrans Houtt. 40 Nutmeg 13 ±1.4 (Myristicaceae) Cananga odoranta (Lam.)Hook.f. 42 Yalng ylang 10.8 ±1.7 and Thoms. (Annonaceae) Cymbopogon martini (Roxb.) Wats. 43 Palmarosa 22.3 ± 5.7 (Poaceae) Lavandula angustifolia Mill. 47 Lavender 9 ± 0.8 (Lamiaceae) Gaultheria frangrantissima Wall. 51 Winter green 12 ± 2.2 (Ericaceae)

52 Viola odorata L. (Violaceae) Violet 11 ± 3.7 Alpinia galangal L. Willd. 53 Ghodawaj 17 ± 3.4 (Zingiberaceae)

63 Ocimum basilicum L. (Lamiaceae) Basil 26.3 ± 1.3 Hexaconazole 1 14.9 ± 0.75

Table-4A.3: Chemical composition of Cinnamomum zeylanicum determine by GC-MS analysis

Peak Retantion Area Essential oil no. Compounds time (%)

1 -linaloo 11.647 10.25

2 Cinnamaldehyde 15.277 64.13

3 Eugenol 15.919 9.04

Cinnamomum Cinnamyl zeylanicum (bark) 4 acetate 17.093 4.47

5 Eugenyl acetate 17.844 0.75

6 Benzyl benzoate 21.163 9.26

others 2.1

Figure-4A.5: Chromatogram of Cinnamomum zeylanicum Table-4A.4: Chemical composition of Cinnamomum cassia determine by GC-MS analysis

Peak Retantion Area Essential oil no. Compounds time (%)

1 -linaloo 11.666 9.16

2 Cinnamaldehyde 15.369 66.36 Cinnamomum Cassia 3 Eugenol 15.956 7.74

Cinnamyl 4 acetate 17.111 3.97

5 Eugenyl acetate 17.862 0.52

6 Benzyl benzoate 21.181 9.26

7 Benzaldehyde 8.916 0.15

8 Benzyl alcohol 10.474 0.23

other 1.63

Figure-4A.6: Chromatogram of Cinnamomum cassia Table-4A.5: Chemical composition of Syzygium aromaticum determine by GC-MS analysis

Peak Retantion Area Essential oil no. Compounds time (%)

1 Eugenol 16.182 47.64

2 Benzyl alcohol 11.104 34.10

Syzygium 3allyl-6- aromaticum 3 Methoxyphenol 17.961 4.98

Caryophyllene 4 oxide 18.896 1.35

5 Caryophyllene 16.732 1

6 Vanillin 16.604 0.91

others 10.02

Figure-4A.7: Chromatogram of Syzygium aromaticum

INTRODUCTION

Plants successfully defend themselves from the attack of a wide range of pathogenic microorganisms. Because they lack a system of circulating antibodies, their defense relies on the capability of each cell to recognize the presence of pathogens and subsequently activate defense responses. Many of the recognition events between plants and pathogens occur in the plant cell wall, which is the first barrier to come into contact with the invading organisms. The majority of microorganisms need to breach this barrier to gain access to the plant tissue and produce enzymes that degrade the wall polymers (De Lorenzo and Ferrari, 2002; Matteo et al., 2003). They have evolved other mechanisms of antimicrobial defense which are either constitutive or inducible (Odjakova and Hadjiivanova, 2001). They are resistant to most pathogens in their environment, as they are not host plants for particular pathogen or are host plants, but harbor resistance genes, allowing them to recognize specifically distinct pathogen races (Scheel, 1998).

Two types of plant resistance response can be distinguished: nonhost and host or race/cultivar specific resistance response. In both cases, the biochemical processes involved in pathogen resistance are very similar (Somssich and Hahlbrock, 1998). Resistance in plants is manifested by the inability of the pathogen to grow or multiply and spread and often takes the form of a hypersensitive reaction. The hypersensitive response is characterized by localized cell and tissue death at the site of infection (van Loon, 1997). As a result the pathogen remains confined to necrotic lesions near the site of infection. A ring of cells surrounding necrotic lesions become fully refractory to subsequent infection, known as localized acquired resistance (Hammon-Koasack and Jones, 1996; Baker et al., 1997; Fritig et al., 1998).

These local responses often trigger nonspecific resistance throughout the plant, known as systemic acquired resistance, providing durable protection against challenge infection by a broad range of pathogens (Ryals et al., 1996; Sticher et al., 1997; van Loon, 1997). The metabolic alterations in localized 78║ P a g e

acquired resistance include: cell wall reinforcement by deposition and cross linking of polysaccharides, proteins, glycoproteins and insoluble phenolics; stimulation of secondary metabolic pathways, some of which yield small compounds with antibiotic activity (the phytoallexins) but also defense regulators such as salicylic acid, ethylene and lipid-derived metabolites; accumulation of broad range of defense-related proteins and peptides (Hahn, 1996; Fritig et al., 1998).

Plant pathogenic fungi produce extra-cellular enzymes which can degrade the cell wall components of plants. These fungi do not only digest plant cell wall polymers to obtain an important nutrient source but also degrade the cell wall enabling cell penetration and spread through plant tissue (An et al., 2005). Thorough research has been focused on trying to determine the role and importance of extra-cellular cell wall-degrading enzymes (CWDE) related to the virulence of plant pathogenic fungi. In several cases conclusive evidence has been obtained for the role of enzymes in the infection process (Annis and Goodwin, 1997; Nightingale et al., 1999; Feng et al., 2005; Jenczmionka and Schäfer, 2005).

Fungal colonization of a plant brings cells of both organisms into close contact; therefore, molecules of the fungal cell wall play an important role in the recognition between fungi and plants. Some of these molecules have the ability to induce plant defense responses and are called elicitors. The term elicitor was originally used with reference to molecules and other stimuli that induce the synthesis and accumulation of antimicrobial compounds (phytoalexins) in plant cells, but is now commonly applied to molecules that stimulate any plant defense mechanism (Hahn, 1996). At present, elicitors are defined as pathogen molecules that trigger defense responses resulting in enhanced resistance to the invading pathogen (Kamoun, 2006).

It is well established that a variety of fungal products can elicit inducible defensive plant responses in both host and non-host plants, and that such responses can also be triggered by plant products released during cell-wall degradation (Heath, 2000). Understanding the mechanisms involved in 79║ P a g e

protection of susceptible plants is essential for successful disease control. Oligosaccharides from fungal cell walls that induce defense responses in plants have been characterized. Accumulation of phytoalexins by the plant and induction of -1,3 glucanase activity have been correlated with the defense response (Wolski et al., 2005).

Plant cell wall polysaccharides are the most abundant organic compounds found in nature. They make up 90% of the plant cell wall and can be divided into three groups: cellulose, hemicellulose, and pectin (McNeill et al., 1984). Cellulose represents the major constituent of cell wall polysaccharides and consists of a linear polymer of 1,4-linked D-glucose residues. The cellulose polymers are present as ordered structures (fibers), and their main function is to ensure the rigidity of the plant cell wall.

Hemicelluloses are more heterogeneous polysaccharides and are the second most abundant organic structure in the plant cell wall. The major hemicellulose polymer in cereals and hardwood is xylan. It consists of a -1,4- linked D-xylose backbone and can be substituted by different side groups such as L-arabinose, D-galactose, acetyl, feruloyl, p-coumaroyl, and glucuronic acid residues (De Vries and Visser, 2001). A second hemicellulose structure commonly found in soft and hardwoods is (galacto) glucomannan which consists of a backbone of -1,4-linked D-mannose and D-glucose residues with D-galactose side groups. Softwoods contain mainly galacto glucomannan, whereas in hardwoods glucomannan is the most common form (Moreira and Filho, 2008).

Xyloglucans are present in the cell walls of dicotyledonae and some monocotylodonae (e.g., onion). Xyloglucans consist of a -1,4- linked D- glucose backbone substituted by D-xylose. L-Arabinose and D-galactose residues can be attached to the xylose residues, and L-fucose has been detected attached to galactose residues in xyloglucan. They interact with cellulose microfibrils by the formation of hydrogen bonds, thus contributing to

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the structural integrity of the cellulose network (Carpita and Gibeaut, 1993; Kikot et al., 2009).

Among the cell wall-degrading enzymes produced by phytopathogenic fungi, an important role is played by the endopolygalacturonases (EC 3.2.1.15) that cleave the linkages between D-galacturonic acid residues in nonmethylated homogalacturonan, a major component of pectin (Matteo et al., 2003). Polygalacturonases (PGs) are among the first enzymes secreted by phytopathogenic fungi, and their action on the outer component of the cell wall is a prerequisite for further wall degradation by other degrading enzymes (De Lorenzo et al., 2001; Veronico et al., 2011).

Pectic polysaccharides make up approximately 30% of the drymass of the primary cell wall in flowering plants and a much greater proportion of the middle lamella (Carpita and Gibeaut, 1993). PG activity has been measured also in ripening fruit (Fischer and Bennett, 1991), root initials (Peretto et al., 1992), root-cap cells (Hawes and Lin, 1990), abscission zones of leaves and flowers (Tucker et al., 1984), pod dehiscence zones (Jenkins et al., 1996), pith cells undergoing autolysis (Huberman et al., 1993) and in tissues infected by plant pathogens (Hong and Tucker, 1998).

β-1,4 xylanases (1,4 β-D-xylan-xylanohydrolase, EC 3.2.1.8) catalyze the hydrolysis of xylan, the major component of hemicellulose of plant cell walls to xylo-oligosaccharides and xylose. A variety of microorganisms, including bacteria, yeasts and filamentous fungi have been reported to produce xylanases (Coughlan and Hazlewood, 1993; Haltrich et al., 1996; Reis et al., 2003).

β-Glucosidases (β-D-glucoside glucohydrolases; EC 3.2.1.21) catalyze the hydrolysis of alkyl and aryl β-glycosides as well as disaccharides and short chain oligosaccharides. -glucosidases (EC 3.2.121) hydrolyse cellobiose into two molecules of glucose (Chauve et al., 2010). They are ubiquitous and can be found in bacteria, fungi, plants and animals. Fungal β-glucosidases are

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parts of the cellulose degrading enzyme system working synergistically with endoglucanases and cellobiohydrolases (Krisch et al., 2010).

-Glucosidase ( -D-glucoside glucohydrolase, EC 3.2.1.20) hydrolyses 1,4- - glucosidic linkages in oligosaccharides rapidly, relative to such linkages in polysaccharides, which are hydrolysed either slowly or not at all (Sahab and Zeikus, 1991). -Glucosidase is the final enzyme involved in the metabolism of starch to glucose, and is used with -amylase for the saccharification of starch. -Glucosidases are produced by micro-organisms in response to growth on starch-containing medium, and vary widely in their substrate specificity. Bacterial and fungal -glucosidases are reported to be specific for maltose hydrolysis whereas yeast -glucosidases hydrolyse p -nitrophenyl- - D-glucoside at a higher rate than maltose (Gupta and Gautam, 1993).

Invertase (EC 3.2.1.26) is classified in the GH32 family of glycoside hydrolases, that includes over 370 members (Alberto et al., 2004) and has been reported in plant , bacteria , yeast and filamentous fungi, as Aspergillus ochraceus , Aspergillus niger , Aspergillus japonicas and Thermomyces lanuginosus (Alegre et al., 2009). Acid phosphatases (EC 3.1.3.2) have been reported to occur in fungi, such as, Aspergillus (Nozawa et al., 1998), Penicillium (Yoshida et al., 1989; Haas et al., 1991), Fusarium (Yoshida and Tamiya, 1971) and Neurospora (Nahas et al., 1982; Aleksieva and Radoevska, 2003).

Animals can produce several different types of immunoglobulins which bind, at a specific site (epitope), the antigen or hapten. These immunoglobulins are synthesized by several cell types of lymphocytes. Such immune serum contains polyclonal antibodies which generally are not mono-specific for the given antigen/hapten. Polyclonal antibodies are faster and cheaper to develop, often give higher sensitivity by recognizing multiple epitopes on the antigen and can be more versatile (Catimel et al., 2006).

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They are produced by different cells, and in consequence, are immunochemically dissimilar; they react with various epitopes on the antigen against which they are raised. Thus, these types of purified antibody can react or “cross-react” with some structurally related antigen (Michaud et al., 2003). By far, the most frequently used animal for the production of polyclonal antibodies is the rabbit, followed by goat, pig, sheep, horse, guinea pig and others. The popularity of rabbits for the production of polyclonal antibodies is attributed primarily to their easy maintenance (Boenisch, 2001).

Antibodies are bioactive molecules that, owing to their individual and specific binding properties, allow a large diversity of potential applications. These include medical diagnosis and therapy, the sensitive detection and removal of environmental contaminants, control of pathogens, and industrial purification processes. Antibodies provide an invaluable tool in fundamental research, because of their ability to interfere with metabolic processes within an organism (Stoger et al., 2002).

In a previous work of Pawar (2007) it was found that the antibody against A. niger (spore) gives cross reactivity with the other seven fungi tested. Present study was designed to evaluate the inhibitory effect of this antibody on different enzyme like α/β-D galactosidase, β-glucosidase, Polygalacturonase, Xylanase, Acid phosphatase, Invertase of six different fungi. Thus, the mechanism of polyclonal antibody for six different fungi was attemted.

MATERIALS AND METHODS

Fungal strains In this study six different fungi used were Aspergillus niger, Fusarium solani, Antrodia sitchensis, Macrophomina phaseolina, Curvularia intermedia and Alternaria porri.

Immunization and antibody production Fungal antigen was extracted from one week old A. niger culture growth. Spore were used as an antigen and immunized into two different rabbits. 83║ P a g e

Each rabbit received intramuscular injections of antigen (1 mg/0.5ml) emulsified with an equal volume of Freund‟s complete adjuvant (total 1 ml was administered) in one month interval. After seven days of antigen dose, blood was collected by ear vein puncture. The titer of raised antibodies was checked and it was found to 1:10000 after 7 months of regular immunization. When antibody titers had fallen, the animals received booster injections followed by the withdrawal of further blood samples and this cycle was continued to build up anti-sera stocks.

Separation of antibodies Serum was clarified from blood clot by centrifugation. The IgG molecules were separated from serum by Ion Exchange Chromatography using DEAE (Diethylaminoethyl) cellulose column. These IgG molecules (serum) were collected in sterile eppendorfs and were further used in the experimental purpose.

Preparation of inoculum The fungi were cultured on PDA and after seven days of inoculation, discs of 1 cm were cut from the periphery with the help of cup borer and suspended in 10 ml of sterile distilled water. It was shaked vigorously and left as such for 2 h and then centrifuged to remove the agar particles and other debris. The supernatant was used as a culture suspension for inoculation.

Media preparation MS media with 2% sucrose was prepared and 50 ml of it was filled in each of the 250 ml conical flask. The media was autoclaved for 15 min and allowed to cool at room temperature. The inoculum (0.2 ml) was added to each flask and maintained in a stationary condition at 28 ± 2°C for culture growth.

Extracellular and cytoplasmic enzyme extraction After 72 h culture media were collected from individual flasks under aseptic conditions and filtered through a disk of Whatman no.1 filter paper. This filtrate constituted extracellular fraction which was used for further assays. The mycelium remained after removing the filtrate was gently blotted over the 84║ P a g e

layers of filter paper and weighed. The mycelial mass was homogenized in chilled pestle-mortar with sterilized sand in sodium acetate buffer (100 mM, pH 5). This homogenate was centrifuged at 4 ºC at 10,000 xg for 30 min. The supernatant was collected, which constituted cytoplasmic fraction.

Inhibition of enzymatic activity by antibody The collected extracellular and cytoplasmic fractions of the respective fungi were mixed with antibody in proportion of 1:100 and incubated for 24 h at 4 ºC for the reaction. Then they were used for enzymatic assay determination. The differences in control and fraction mix with antibody were calculated and compare to check out reduction in their activity.

Glycosidases assay This activity was determined as described by Thaker et al. (1987). The assay medium consisted equal volume of enzyme extract and buffered substrate [1 mg/ml p-nitrophenyl α/β-D galactopyranoside or p-nitrophenyl β-D- glucopyranoside in 100 mM sodium acetate buffer (pH 5)]. After 60 min incubation at 30 ± 2 °C, the reaction was terminated by the addition of 1M

Na2CO3 solution in one and a half times the volume of the reaction mixture. In control reaction, Na2CO3 was added prior to the addition of enzyme. The absorbance of the yellow pNP released was measured at 405 nm using ELISA Reader (μ Quant, Bio-tek, USA). The quantity of pNP released was calculated from the calibration curve prepared using a range of pNP concentration from 0.036 to 0.7 mM prepared in same buffer. The activity was expressed as M pNP released/mg protien for all cytoplasmic and extracellular fractions. The assay was performed in triplicates for each enzyme extract and mean values with ± SD were calculated.

Polygalacturonase and Xylanase assay The assay medium consisted equal volume of buffered substrate [2 mg/ml] polygalacturonic acid or xylan in 100 mM sodium acetate buffer (pH 5)] and enzyme solution. After 60 min incubation at 30 ± 2 °C, the reaction was terminated by the addition of 3, 5-dinitrosalicylic acid (Miller, 1959) and

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reducing sugars were estimated. Absorbance was measured at 520 nm using ELISA Reader (μ Quant, Bio-tek, USA). The activity was calculated from the standard curve prepared by using glucose. The activity was expressed as M sugar released/mg protein for all cytoplasmic and extracellular fractions. Control was prepared by addition of Miller‟s reagent prior to the substrate. The assay was performed in triplicates for each enzyme extract and mean values with ± SD were calculated.

Acid phosphatase assay The assay medium consisted of 0.3 ml, 100 mM sodium acetate buffer (pH 5), 0.4 ml enzyme solution and 2.3 mM p-nitrophenyl phosphate (pNPP) (in buffer) in a final volume of 1 ml. The mixture was incubated at 30 ± 2 °C and after 15 min incubation it was terminated by the addition of 2 ml of 1M

Na2CO3. In control reaction, Na2CO3 was added prior to the addition of enzyme. The absorbance of the pNP released was measured at 405 nm using ELISA Reader (μ Quant, Bio-tek, USA). The quantity of pNP released was calculated from the calibration curve prepared using a range of pNP concentration from 0.036 to 0.7 mM prepared in same buffer. The acid phosphatase activity was expressed as M pNP released/mg protein for all cytoplasmic and extracellulat fractions. The assay was performed in triplicates for each enzyme extract and mean values with ± SD were calculated.

Invertase assay The reaction mixture contained 0.15 M sucrose and enzyme solution. After 60 min incubation at 30 ± 2 °C, the reducing sugars were estimated with the 3, 5- dinitrosalicylic acid method (Miller, 1959). Control was prepared by addition of Miller‟s reagent prior to the substrate. Absorbance was measured at 520 nm using ELISA Reader (μ Quant, Biotek, USA). The activity was calculated from the standard curve prepared by using glucose. The activity was expressed as M sugar released/mg protein for all cytoplasmic and extracellular fractions. The assay was performed in triplicates for each enzyme extract and mean values with ± SD were calculated.

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Protein determination Total protein was determined from extracellular and cytoplasmic fractions, by the method of Lowry et al. (1951) using BSA as the standard.

RESULTS AND DISCUSSION

In modern diagnostics, antibodies (Abs) have become key affinity ligands as they can be produced against almost any component, from small drug molecules to intact cells. Approaches for plantpathogen- specific Ab production are mainly based on injection of whole pathogens, surface fragments or soluble surface components into a suitable animal-host for polyclonal antibody (pAb) or monoclonal antibody (mAb) production (Werres and Steffens, 1994; Skottrup et al., 2008).

DeBary (1886) was the first to suggest that extracellular enzymes may be involved in the infection process of plant pathogenic fungi. Since then, much research has been focused on trying to determine the role and importance of extracellular CWDE to the virulence of plant pathogenic fungi (Annis and Goodwin, 1997). The removal of D-galactose residues from plant cell wall polysaccharides requires the action of -galactosidases and galactosidases (De Vries and Visser, 2001). Invertases ( -fructofuranosidases) hydrolyze sucrose into glucose and fructose (Lee and Sturm, 1996). Endo-β-1,4- xylanases and β-xylosidases work synergistically to degrade xylan to xylose (Rose and van Zyl, 2002). Fungi produce different types of pectinases that are classified by their substrates, type of lysis and mode of action on the pectin polymer. Unesterified pectate polymers can be degraded by polygalacturonase (PG) (Rexova-Benkova and Markovic, 1976).

In the present study, the inhibitory effect of antibody was evaluated against the cell wall degrading enzymes in different fungi and found that cytoplasmic fraction of A. niger showed reduction in enzyme activity (Fig. 4B.1a). Glucosidases were slightly inhibited while significant inhibition was found in xylanase and invertase after antibody treatment. On the other hand in case of

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extracellular fraction the activity of xylanase and invertase was increases and acid phosphatase activity inhibited after antibody exposure (Fig. 4B.1b).

In case of F. solani cytoplasnic fraction inhibition exhibited within / galactosidase, acid phospatase and invertase (Fig. 4B.2a). An extracellular fraction showed drastic inhibition in -glucosidase and xylanase (Fig. 4B.2b). On the other hand, no remarkable inhibition was found in both the fraction (Fig. 4B.3a, b) of A. sitchensis only -glucosidase was slightly reduced in cytoplasmic fraction (Fig. 4B.3a). Sorensen et al. (2004) reported that xylanase inhibitors are not only of technological relevance, as they inhibit xylanases used in cereal based biotechnological processes, some of them are also of significance in plant defense against pathogens (Bellincampi et al., 2004).

M. phaseolina exhibited drastic inhibition in all the enzyme tested of both cytoplasmic and extracellular fractions (Fig. 4B.4a, b). All the enzymes were inhibited in cytoplasmic fraction of A. porii (Fig. 4B.6a) but in case of extracellular only -galactocidae, acid phosphatase and polygalacturonas were inhibited (Fig. 4B.6b). C. intermedia showed more inhibition in - glucosidae, -galactosidae, invertase and polygalacturonas in their cytoplasmic fraction compared to extracellular one (Fig. 4B.5a, b).

The invertase inhibitory protein isolated from Cyphomandra betacea Sendt and Solanum tuberosum inhibited the invertase activity from different species, genera and even plant family. Furthermore, proteinaceous inhibitors were not invertase specific; fungal, bacterial and higher plant enzymes including polygalacturonase, pectinase, pectin lyase, alpha-L-arabinofuranosidase and beta-glucosidase were also shown to be inhibited (Isla et al., 2002).

Polygalacturonase-inhibiting proteins (PGIPs) have been identified in the cell walls of many plant species, including bean, soybean, tomato, pear, apple, raspberry, onion and leek (Arendse et al., 1999; De Lorenzo and Cervone 1997; Favron et al., 1997; Johnston et al., 1993; Stotz et al., 1994; Yao et al.,

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1995). Plant PGIPs interact with endo-polygalacturonases from fungi, but do not appear to have an effect on those of bacterial or plant origin (Cervone et al., 1990). Bean PGIP was initially considered to have broad-spectrum activity against polygalacturonases from a range of fungi, including C. lindemuthanium, F. oxysporum, F. moniliforme, and two species of Aspergillus (Berger et al., 2000).

Enzyme inhibitors, both low-molecular weight compounds and proteinaceous molecules, have emerged as important pharmaceutical agents. The generation of antibody-based molecules forms an obvious alternative of enzyme inhibitor. More specifically, the heavy-chain antibodies acquired the potential to recognize protein cavities and as such the ability to inhibit enzymes (Lauwereys et al., 1998). However, despite the omnipotence of the antibody repertoire, the number of conventional antibodies (i.e. heterotetramers of two light chains and two heavy chains) acting as competitive enzyme inhibitors remains disappointingly low. A satisfactory explanation for this scarce occurrence is given by the incompatible surface topography of the enzyme‟s active site and the antigen-binding site of conventional antibodies. From a recent survey of enzyme structures it appears that the active site is found almost exclusively in the largest cleft on the protein surface (Laskowski et al., 1996).

The importance of specificity of antibody in its inhibitory effect is still controversial. Kinetic studies have led to contradictory and inconclusive data on the relation between specificity and inhibitory capacity (Cinader, 1955). The foregoing evidence indicates that the inhibitory capacity of antibody is related to specificity. Obtaining pathogen-specific Abs is a challenging task and Abs are often found to be specific to the genus level only, as the surface of intact cells contains protein and carbohydrate epitopes that are present throughout species (Li et al., 2000; Skottrup et al., 2008). All most in all the cases glycosidases and acid phosphatase were inhibited so they may have same kind of epitops by which antibody binds and may alter their structure leads to inhibitory activity of them.

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CONCLUSIONS

On the basis of results it can be concluded that, for the inhibitory activity of polyclonal antibody shown in glycosidases and acid phosphatase, the following possibilities must be there : (i) the effect of antibody binding sites on the enzyme is additive with each binding site that becomes attached to the enzyme adding to the total inhibition produced by the other bound sites regardless of the point of attachment; (ii) inhibition is caused solely by specific binding of a single antibody binding site to one or more specific on the enzyme; and (iii) inhibition is produced by the cooperative effect of two or more antibody binding sites which together cause greater net inhibition when bound to the same enzyme molecule than when bound to separate enzyme molecules.

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(i)

A. niger 72h (cytoplasmic) Normal Ab(24h) 4 3.5 3 2.5 2 1.5 1 0.5

M pNP released/mg protein/h released/mg pNP M 0  1 2 3 Ap

Enzymes

(ii)

Normal

A. niger 72h (cytoplasmic) Ab(24h)

70 60 50 40 30 20 10 0 M sugar released/mg protein/h released/mg sugar M I X Pg  Enzymes

Figure-4B.1a: Comparative activity of normal and antibody treated enzymes in cytoplasmic fraction of A. niger

(i): 1--galactosidase, 2--glucosidase, 3--galactosidase, Ap- Acid phosphatase (ii): I- Invertase, X-Xylanase, Pg- Polygalactouranase.

(i)

Normal A. niger 72h (Extracellular) Ab(24h) 0.4 0.35 0.3 0.25 0.2 0.15 0.1 0.05

M pNP released/mg protein/h released/mg pNP M 0  1 2 3 Ap

Enzymes

(ii)

Normal A. niger 72h (Extracellular) Ab(24h) 45 40 35 30 25 20 15 10 5 0 M sugar released/mg protein/h released/mg sugar M I X Pg 

Enzymes

Figure-4B.1b: Comparative activity of normal and antibody treated enzymes in extracellular fraction of A. niger

(i): 1--galactosidase, 2--glucosidase, 3--galactosidase, Ap- Acid phosphatase (ii): I- Invertase, X-Xylanase, Pg- Polygalactouranase.

(i)

Normal F. solani 72h (cytoplasmic) Ab(24h) 0.35 0.3 0.25 0.2 0.15 0.1 0.05 0 1 2 3 Ap M pNP released /mg /mg protein/h released pNP M  Enzymes

(ii)

Normal

F. solani 72h (cytoplasmic) Ab(24h) 500 450 400 350 300 250 200 150 100 50 0

M sugar released /mg protein/h /mg released sugar M I X Pg  Enzymes

Figure-4B.2a: Comparative activity of normal and antibody treated enzymes in cytoplasmic fraction of F. solani

(i): 1--galactosidase, 2--glucosidase, 3--galactosidase, Ap- Acid phosphatase (ii): I- Invertase, X-Xylanase, Pg- Polygalactouranase.

(i)

Normal F. solani 72h(Extracellular) Ab(24h) 0.07 0.06 0.05 0.04 0.03 0.02 0.01 0 M pNP released /mg /mg protein/h released pNP M

 1 2 3 Ap

Enzymes

(ii)

Normal

F. solani 72h (Extracellular) Ab(24h)

90 80 70 60 50 40 30 20 10

M sugar released /mg protein/h /mg released sugar M 0

 I X Pg

Enzymes

Figure-4B.2b: Comparative activity of normal and antibody treated enzymes in extracellular fraction of F. solani

(i): 1--galactosidase, 2--glucosidase, 3--galactosidase, Ap- Acid phosphatase (ii): I- Invertase, X-Xylanase, Pg- Polygalactouranase.

(i)

A. sitchensis Normal 72h (cytoplasmic) Ab(24h) 8 7 6 5 4 3 2 1 0

M pNP released /mg /mg protein/h released pNP M 1 2 3 Ap  Enzymes

(ii)

Normal

A. sitchensis 72h(cytoplasmic) Ab(24h) 1800 1600 1400 1200 1000 800 600 400 200 0

M sugar released/mg protein/h released/mg sugar M I X Pg  Enzymes

Figure-4B.3a: Comparative activity of normal and antibody treated enzymes in cytoplasmic fraction of A. sitchensis

(i): 1--galactosidase, 2--glucosidase, 3--galactosidase, Ap- Acid phosphatase (ii): I- Invertase, X-Xylanase, Pg- Polygalactouranase.

(i)

Normal A. sitchensis 72h (Extracellular) Ab(24h) 0.25

0.2

0.15

0.1

0.05

M pNP released/mg protein/h released/mg pNP M 0  1 2 3 Ap

Enzymes

(ii)

A. sitchensis 72h (Extracellular) Normal Ab(24h) 450 400 350 300 250 200 150 100 50 0

M sugar released/mg protein/h released/mg sugar M I X Pg 

Enzymes

Figure-4B.3b: Comparative activity of normal and antibody treated enzymes in extracellular fraction of A. sitchensis

(i): 1--galactosidase, 2--glucosidase, 3--galactosidase, Ap- Acid phosphatase (ii): I- Invertase, X-Xylanase, Pg- Polygalactouranase.

(i)

M. phaseolina Normal

72h (cytoplasmic) Ab(24h) 5 4.5 4 3.5 3 2.5 2 1.5 1 0.5 0 M pNP released/mg protein/h released/mg pNP M

 1 2 3 Ap

Enzymes

(ii)

M. phaseolina 72h (cytoplasmic) Normal

Ab(24h) 600

500

400

300

200

100

M sugar released/mg protein/h released/mg sugar M I X Pg  Enzymes

Figure-4B.4a: Comparative activity of normal and antibody treated enzymes in cytoplasmic fraction of M. phaseolina

(i): 1--galactosidase, 2--glucosidase, 3--galactosidase, Ap- Acid phosphatase (ii): I- Invertase, X-Xylanase, Pg- Polygalactouranase.

(i)

M. phaseolina 72h (Extracellular) Normal

Ab(24h)

0.1 0.09 0.08 0.07 0.06 0.05 0.04 0.03 0.02 0.01 0 MpNP released/mg protein/h released/mg MpNP

 1 2 3 Ap

Enzymes

(ii)

M. phaseolina 72h (Extracellular) Normal Ab(24h)

0 M sugar released/mg protein/h released/mg sugar M

 I X Pg

Enzymes

Figure-4B.4b: Comparative activity of normal and antibody treated enzymes in extracellular fraction of M. phaseolina

(i): 1--galactosidase, 2--glucosidase, 3--galactosidase, Ap- Acid phosphatase (ii): I- Invertase, X-Xylanase, Pg- Polygalactouranase.

(i)

Normal C. intermedia 72h (cytoplasmic)

Ab(24h)

0.3

0.25

0.2

0.15

0.1

0.05

M pNP released/mg protein/h released/mg pNP M 0  1 2 3 Ap

Enzymes

(ii)

Normal C. intermedia 72h (cytoplasmic) Ab(24h)

800 700 600 500 400 300 200 100 0 M sugar released/mg protein/h released/mg sugar M

 I X Pg

Enzymes

Figure-4B.5a: Comparative activity of normal and antibody treated enzymes in cytoplasmic fraction of C. intermedia

(i): 1--galactosidase, 2--glucosidase, 3--galactosidase, Ap- Acid phosphatase (ii): I- Invertase, X-Xylanase, Pg- Polygalactouranase.

(i)

Normal C. intermedia 72h (Extracellular)

Ab(24h) 0.25

0.2

0.15

0.1

0.05

M pNP released/mg protein/h released/mg pNP M 1 2 3 Ap  Enzymes

(ii)

Normal C. intermedia 72h (Extracellular) Ab(24h)

450 400 350 300 250 200 150 100 50

M sugar released/mg protein/h released/mg sugar M 0  I X Pg

Enzymes

Figure-4B.5b: Comparative activity of normal and antibody treated enzymes in extracellular fraction of C. intermedia

(i): 1--galactosidase, 2--glucosidase, 3--galactosidase, Ap- Acid phosphatase (ii): I- Invertase, X-Xylanase, Pg- Polygalactouranase.

(i)

A. porri 72h (cytoplasmic) Normal

Ab(24h)

5 4.5 4 3.5 3 2.5 2 1.5 1 0.5

M pNP released/mg protein/h released/mg pNP M 0  1 2 3 Ap

Enzymes

(ii)

A. porri 72h (cytoplasmic) Normal

Ab(24h)

3000

2500

2000

1500

1000

500

M sugar released/mg protein/h released/mg sugar M 0

 I X Pg

Enzymes

Figure-4B.6a: Comparative activity of normal and antibody treated enzymes in cytoplasmic fraction of A. porri

(i): 1--galactosidase, 2--glucosidase, 3--galactosidase, Ap- Acid phosphatase (ii): I- Invertase, X-Xylanase, Pg- Polygalactouranase.

(i)

Normal A. porri 72h (Extracellular) Ab(24h)

0.16 0.14 0.12 0.1 0.08 0.06 0.04 0.02

M pNP released/mg protein/h released/mg pNP M 0  1 2 3 Ap

Enzymes

(ii)

Normal A. porri 72h (Extracellular)

Ab(24h)

100 90 80 70 60 50 40 30 20 10

M sugar released/mg protein/h released/mg sugar M 0

 I X Pg

Enzymes

Figure-4B.6b: Comparative activity of normal and antibody treated enzymes in extracellular fraction of A. porri

(i): 1--galactosidase, 2--glucosidase, 3--galactosidase, Ap- Acid phosphatase (ii): I- Invertase, X-Xylanase, Pg- Polygalactouranase.

Plant-pathogenic fungi include a very large and heterogeneous group of organisms that occupy positions of great importance in both agriculture and natural plant communities. They show an enormous diversity in life-history strategies and the ways in which they interact with their hosts. Many pathogenic fungi can survive for long periods of time on dead host tissue or saprophytically in soil; others rely entirely on living host cells for sustenance (Burdon and Silk, 1997). Collectively, the pathogens can attack virtually any plant part, although individually they may be highly specialized. As a consequence, the range of pathogens found on different hosts also shows considerable diversity that may be associated with the evolutionary history of their hosts (Clay, 1995).

Fungal physiology refers to the nutrition, metabolism, growth, reproduction and death of fungal cells. It also generally relates to interaction of fungi with their biotic and abiotic environment, including cellular responses to stress. The physiology of fungal cells impacts significantly on the environment, industry and human health. Fungal metabolism is also responsible for detoxification of organic pollutants and for bioremediation of heavy metals in the environment (Walker and White, 2005). The production of many economically important industrial commodities relies on exploitation of fungal metabolism and these include such diverse products as whole foods, food additives, fermented beverages, pharmaceuticals (Hamlyn, 1997), pigments (Velisek and Cejpek, 2011), alkaloids (Mahmood et al., 2010), biofuels (Xia et al., 2011), industrial antibiotics, enzymes, vitamins, organic and fatty acids (Carlile and Watkinson, 1995; Levinskaite, 2004) and sterols (Li et al., 2004).

Plant pathogenic fungi cause important losses in a number of crops, it is necessary to make high-throughput studies on these organisms. The spread of drug resistant pathogens is one of the most serious threats to successful treatment of microbial diseases. Down the ages essential oils and other extracts of plants have evoked interest as sources of natural products. They had been screened for their potential uses as alternative remedies for the treatment of many infectious diseases (Tepe et al., 2004). 91║ P a g e

Keeping all these the present study has been focused on majorly three aspects: 1. Identification of fungi morphologically and molecular means by designing primers. 2. Optimizations of indole-acetic acid (IAA), Abscisic acid (ABA) and Gibberellic acid (GA) production in define media by Aspergillus niger, Macrophomina phaseolina and Fusarium moniliforme, respectively. 3. Biocontrol approach had been made against some plant pathogenic fungi using plant extract of Agave, essential oils (EOs) and an attempt to illustrate the mechanism of action of antibody (Ab) on cell wall degrading enzymes.

Studies of fungal pathogens and their interactions with plants have been performed using several approaches, from classical genetic, cell biology, and biochemistry (Hardham and Mitchell, 1998; Howard, 2001; Farkas, 2003; Perez and Ribas, 2004; Koh and Somerville, 2006; Xu, 2006; Aguileta et al., 2009; Alves and Pozza, 2009; Poland et al., 2009;), to the modern, holistic, and high-throughput omic techniques (Choquer et al., 2007; Egan and Talbot, 2008) accompanied by proper bioinformatic tools (Hadwiger, 2009).

In recent years, the study of fungal plant pathogens has been greatly promoted by the availability of their genomic sequences and resources for functional genomic analysis, including transcriptomics, proteomics, and metabolomics (Tan et al., 2009), which in combination with targeted mutagenesis or transgenic studies, are unraveling molecular host-pathogen crosstalk, the complex mechanisms involving pathogenesis and host avoidance (Walter et al., 2010). The importance of a genetic data base for fungal systematics as well as for other organisms is widely known. Recently, such fields as phylogeny, systematics and quick identification of particular fungal taxa of different ranks on the basis of sequence comparison have been developed (Seifert et al., 2007).

In this study from the morphological and microscopical examination three of seven fungi were identified on the basis of their spore structure. Amongst 92║ P a g e

them one was identified as A. niger based on both, spore structure and molecular identification by 18s specific primer. Rest of the six fungi was identified by designed primer of 28s region, in that two were identified on the basis of their morphological and microscopical structure as Fusarium solani and Curvularia intermedia. The phylogenetic analysis of these six fungi was also carried out and it revealed the fungi as Fusarium solani, Antrodia sitchensis, (three fungi as) Macrophomina phaseolina and Curvularia intermedia.

Almost all processes in the life of a plant are directly or indirectly affected by both stresses and phytohormones. Plant-associated micro-flora is the richest source of microorganisms synthesizing phytohormones. Investigations on microbial production of IAA, ABA and GA have been focused primarily on plant pathogens. The microbial synthesis of plant growth regulators like GA an IAA is an important factor in soil fertility (Kampert and strzelczyk, 1975). GA and IAA are secondary metabolites, which are important biotechnological products, produce commercially from fungi for the agriculture and horticultural industry (Hasan, 2002).

IAA production by A. niger was optimized in define media. The higher production of IAA was found in MS half strength and 4% of sucrose media at 46 h and 25 ± 2°C with substrate dependency. Another attempt had been done in the present work to optimize production of IAA and studied at molecular level. For that aldehyde dehydrogenase gene primer was designed as it is a key enzyme for conversation of indole acetic aldehyde to IAA. It was found that it amplified in A. niger, obtained sequence were submitted in GenBank database of NCBI. In further study protein sequence analysis and protein structure prediction with the help of some bioinformatics tool was also carried out that confirmed it as aldehyde dehydrogenase. Mutational study was also carried out to check the relative variation in that.

The bulk of available evidence indicates that phytohormones formed by fungi, algae, and bacteria are structurally identical to their plant counterparts (Tsavkelova et al., 2006a). Moreover, epiphytic and rhizospheric micro floras 93║ P a g e

of plants are of greatest importance in the conversion of Trp into IAA. Omission of Trp from the culture medium decreases the level of IAA synthesis by the culture‟s microorganisms. The presence of L-Trp in the medium is a necessary condition of the biosynthesis of IAA. When the production of IAA by soil microorganisms is studied under laboratory conditions, Trp is deliberately added to the medium (Kravchenko et al., 2004). Exogenous Trp may augment auxin biosynthesis by an order of magnitude or higher, this being the reason why the yield of the phytohormone in the most active strains exceeds 80-100 μg IAA/ml culture medium (Mordukhova et al., 1991; Forni et al., 1992; Ivanova et al., 2001; Sergeeva et al., 2002; Tsavkelova et al., 2003).

The ability for auxin production has also been reported among various phototrophic and heterotrophic bacteria, not only pathogenic or symbiotic species but also free living microorganisms not associated with plants. IAA synthesis was also reported for the representatives of the genera Pseudomonas, Agrobacterium, Azospirillum, Azotobacter, Alcaligenes, Enterobacter, Acetobacter, Rhizobium, Bradyrhizobium, Bacillus, Arthrobacter, Herbaspirillum, Xanthomonas, Klebsiella, Methylobacterium, Methylovorus, Aminobacter, and Paracoccus (Cacciari et al., 1989; Mordukhova et al., 1991; Wilkinson et al., 1994; Kameneva et al., 1999; Ivanova et al., 2001). Achromobacter, Flavobacterium, Rhodococcus, Acinetobacter, Corynebacterium, and Micrococcus also produce this plant growth stimulator (Libbert and Risch, 1969; Barea et al., 1976; Belimov et al., 1999).

The higher production of GA and ABA was found in MS (2) media. It is a first account to observed ABA production in M. phaseolina. Assante et al. (1977) showed that Cercospora rosicola produces large amounts of ABA. Since then ABA has been detected in many species of ascomycetes, fungi imperfecti and basidiomycetes. A review on phytohormones in fungi has been published by Tudzynski and Sharon (2002) including a list of ABA producing fungi. Using phytopathogenic fungi of the genus Cercospora as an exemplary case, direct ABA biosynthesis (via mevalonic acid oxidation) was shown to involve ionylidene acetate as an immediate precursor (in C. rosicola and C. cruenta) 94║ P a g e

(Yamamoto et al., 2000). The operation of the mevalonate pathway of ABA formation in certain phytopathogenic fungi was confirmed by other researchers (Hirai et al., 2000). Although ABA does not seem to have any function in fungi, fungal ABA maybe of some importance for plants infected. Only few published data are available on the effects of external ABA on fungi (Hartung, 2010).

The filamentous fungus Gibberella fujikuroi is the main producer of gibberellins. These hormones are widely used to optimize growth in several products, particularly seedless grapes (Bruckner and Blechschmidt, 1991). Among the gibberellins, the most important from an industrial perspective is

GA3, which can be produced by fermentation at relatively high concentrations (Qian et al., 1994). Several papers describe moisture content and water activity (aw), for both Solid State Fermentation and submerged culture, as critical variables that limit microbial growth, metabolite production and product efficacy (Pascual et al., 1999; Valik and Pieckova, 2001).

The medicinal properties of plants have been investigated in the light of recent scientific developments throughout the world, due to their potent pharmacological activities and low toxicity (Sharma et al., 1992; Vaquero et al., 2010). Antimicrobial activity of herbs has been known and described for several centuries (Begamboula et al., 2003). Essential oils and various plant extracts have a broad spectrum activity against the bacterial pathogenesis and they also perform the antifungal activity (Sartoratto et al., 2004; Kotzekidou et al., 2008).

The present study has been also emphasis on effectiveness of plant extract as a biocontrol agent. Different five species of Agave plant extracts was evaluated for their antifungal activity against eight fungi. In that six fungi were inhibited and amongst them P. placenta and M. phaseolina found to be more susceptible towards all the extracts tested. It has been well documented that antimicrobial compounds are abundantly present in medicinal plants (Valarini et al., 1994, Fiori et al., 2000, Ghosh et al., 2002). These compounds are thought to be involved in the defense of plants against microbial pathogens. 95║ P a g e

Schwan-Estrada et al. (1998) reported that the crude extract of Eucalyptus citriodora inhibited the mycelial growth of Rhizoctonia solani, Sclerotium rolfsii, Phytophthora sp, Alternaria alternata and Colletotrichum graminicola. The extract of Cymbopogon citratus leaves can completely inhibit the mycelial growth of Fusarium solani f. sp phaseoli, Sclerotinia sclerotiorum and Rhizoctonia solani (Valarini et al., 1994).

Deepak et al. (2005) screened methanolic extracts of forty plant species for antisporulant activity against Sclerospora graminicola (Sacc.) Schroet., the causative agent of pearl millet downy mildew and found that extracts of 11 species (Agave americana, Artemisia pallens, Citrus sinensis, Dalbergia latifolia, Helianthus annus, Murraya koenigii, Ocimum basilicum, Parthenium hysterophorus, Tagetes erecta, Thuja occidentalis and Zingiber offinale) exhibited remarkable antisporulant effect. Satya et al. (2005) had been reported that the leaf extract of zimmu exhibited strong antifungal activity against Rhizoctonia solani, Aspergillus flavus, Curvularia lunata and Alternaria solani.

Several studies have pointed out the possibility to use essential oils and/or their components in medical and plant pathology (Cantore et al., 2009). EOs can be best used for postharvest or stored crop protection, for their qualities that they are effective at very low concentration and also due to their volatile nature; they are highly diffusible and hence react quickly. Considering this into the mind second approach had been made to evaluate the antifungal potential of sixty five EOs against F. moliniforme (a common pathogen of maize) and found that amongst them twenty four were exhibit the inhibitory effect. It was observed that F. moliniforme was highly suseptible to three essential oils Cinnamomum zeylanicum, Cinnamomum cassia and Syzygium aromaticum. Other oils i.e. Cedrus deodara, Apium graveolens, Thymus Vulgaris and Laurus nobilis also posses higher antifungal activity.

EOs are very complex natural mixtures which can contain about 20-60 components at quite different concentrations. They are characterized by two 96║ P a g e

or three major components at fairly high concentrations (20-70%) compared to others components present in trace amounts. Bansod and Rai (2008) evaluate the antifungal activities of fifteen plants EOs against Aspergillus fumigatus and A. niger. In work of Aoudou et al. (2010) they were repoted the antifungal activity of EOs from L. rogusa, P. glandulosus, C. anisata and V. heterophylla against nine fungal samples. In that L. rogusa and P. glandulosus have been shown to have strong antifungal properties; significantly reducing the mycological growth of F. moniliforme.

The GC-MS of these three potent EOs was accomplished to investigate the major constituents responsible for high antifungal activity. Cinnamaldehyde and eugenol in Cinnamon oils and eugenol in Clone oil may be responsible for such higher activity. The antimycotic activity of cinnamon bark due to presence of cinnamaldehyde is well known (Wang et al., 2005; Sivamani et al., 2010). The synergic reaction between some components of cinnamon oil and clove oil were reported by Matan and Matan (2007) that affects the growth of mould at ratios of cinnamon oil to clove oil higher than 5:1. It showed good potential to inhibit the growth of rubberwood moulds i.e. A. niger, P. chrysogenum, and Penicillium sp.

In previous work of Pawar (2007), it was reported that inhibition in growth of all the fungi tested by the polyclonal Ab produce against A. niger (spore). So it was inferred that there might be any mechanism common for hyphae vegetative growth and spore growth. The common mechanism of action of Abs might help in detecting the factor responsible for pathogenecity and growth of fungi. There might be some common metabolite or key enzyme which if blocked may help to inhibit fungal growth thereby decreasing its pathogenic activities. In consideration of this, the present study was designed to evaluate the mechanism of action of Ab on cell wall degrading enzymes of six fungi.

The secretion of enzymes to degrade biological polymers is a defining feature of the Kingdom Mycota. Plant cell walls constitute the largest single source of available reduced carbon in terrestrial environments, and fungi secrete a large 97║ P a g e

number and variety of enzymes that can act on the linkages found in plant cell walls, including cellulases, hemicellulases, pectinases, esterases, oxidoreductases, and proteases. Some filamentous fungi secrete more than two hundred glycosyl hydrolases alone (Cuomo et al., 2007; Dean et al., 2005; Nagendran et al., 2009).

The plant cell wall is the first barrier for the invasion of fungal pathogens. Therefore, during the invasion, the cell wall degrading enzymes (CWDEs), which is secreted by pathogens, is a crucial factor for the successful development of the disease in host. These related enzymes have a close relationship with pathogenesis (Zhao and Zhang, 2002; Tian et al., 2009). In the present study the inhibition of CWDEs by Ab was evaluated in six fungi and it was found that M. phaseolina exhibited drastic inhibition in the entire enzyme tested of both cytoplasmic and extracellular fractions. In case of A. porii all the enzymes of cytoplasmic fraction were inhibited. Overall in most of fungi glycosidases and acid phosphatase were inhibited. So they may have same kind of epitops by which antibody binds and showed inhibitory activity. Thus this study may help to revile the mechanism of action of Ab on the suppression of pathogenesis.

Recombinant antibodies were first expressed in plants by Hiatt et al. (1989). Antibodies were first investigated as a strategy to prevent fungal diseases by Peschen et al. (2004). They expressed a fusion protein comprising a recombinant scFv recognizing a surface protein present on Fusarium oxysporum f. sp. matthiolae joined to an antifungal protein from Aspergillus giganteus. The scFv was isolated from a phage display library derived from chickens immunized with fungal spores. Preliminary analysis confirmed that the scFv-AFP fusion bound to the fungus in vitro and inhibited its growth.

In previous studies by group of researchers, antibodies have proven a very powerful tool for the detection and quantification of pathogens (Biosca et al., 1997; Dewey et al., 1997; Karpovich-Tate et al., 1998; Mutasa-Gottgens et al., 2000), but now a day‟s antibody-mediated resistances against plant pathogens are in focus (Safarnejad et al., 2011). Antibody-mediated fungal 98║ P a g e

resistance in plants remains to be demonstrated, monoclonal antibodies have been identified that inhibit fungal growth (Hiatt et al., 2001). Immunomodulation is a powerful tool for studying or altering the function of an antigen in vivo. To this end, an antigen, which may be an enzyme or metabolite, can either be stabilized or blocked in its action (De Jaeger et al., 2000; Conrad and Manteuffel, 2001; Jain et al., 2011). Although polyclonal antibodies have often been produced, their use in ecological studies of fungi under field conditions has been restricted by poor specificity (Hitchcock et al., 1997).

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MAJOR FINDINGS

 This study will help to identify the pathogens and the designed primer was used to develop PCR kit for diagnosis of infected specimen.

 The optimum temperature for IAA production is 25 ± 2ºC, pH in between 2.5-3, media half strength of MS and 4% sucrose, incubation time in dark was 46 h.

 The best condition for PCR amplification of aldehyde 3-dehydrogenase

gene was 1.5mM MgCl2 and 10 pmol primers. The primer pair 5 gives the best result amongst the all. The present study revealed that the aldehyde 3-dehydrogenase gene amplified in A. niger was responsible for IAA synthesis.

 In this study it was first time reported the production of ABA by M. phaseolina.

 Studies on hormonal production and its optimization will also help to exploit these organisms in biotech industries.

 The present results are important for the development of a bio- fungicide preparation from Agave sp. for the management of fungal pathogens of crop plants.

 The results of present study indicate that EOs could find a practical and rational use in the inhibition of fungal growth. Particularly, C. zeylanicum essential oil possesses strong antifungal activity against F. moniliforme. The broad inhibition of fungal growth by C. zeylanicum, in addition to its availability as natural volatile product, justifies its possible use as an alternative antifungal compound to control the growth and dissemination of pathogen and proved to be an efficient bio-control agent.

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 For the inhibitory activity of antibody on glycosidases and acid phosphatase, the following possibilities must be there : (i) the effect of antibody binding sites on the enzyme is additive with each binding site that becomes attached to the enzyme adding to the total inhibition produced by the other bound sites regardless of the point of attachment; (ii) inhibition is caused solely by specific binding of a single antibody binding site to one or more specific on the enzyme; and (iii) inhibition is produced by the cooperative effect of two or more antibody binding sites which together cause greater net inhibition when bound to the same enzyme molecule than when bound to separate enzyme molecules.

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FUTURE STUDIES

 Protein profiling and gene expression studies of Aldehyde dehydrogenase yet need to carry out.

 In future with the variation or other supplement of basal media the yield of phytohormone like ABA from the M. phaseolina could be enhanced.

 In vivo study of essential oil was needed to check whether there is any side effect of it. The mode of action of each compound present in EOs towards pathogen will investigate for their individual activity against the fungi.

 Antibody characterization will help to understand the mode of action towards the fungal pathogens.

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