1. general introduction

1. GENERAL INTRODUCTION

Enzymes are proteins with highly specialized catalytic functions, produced by all living organisms. Enzymes are responsible for many essential biochemical reactions in microorganisms, plants, animals, and human beings. They are essential for all metabolic processes, but are not alive. Although like all other proteins, enzymes are composed of amino acids, they differ in function in that they have the unique ability to facilitate biochemical reactions without undergoing change themselves. Most enzymes today are produced by the fermentation of bio-based materials (Louwrier, 1998). Recent developments on engineering techniques make application of industrially important enzymes such as proteases and lipases in detergents, amylase and glucose isomerase in starch processing and also in the bioprocessing of raw materials (Cheetham et al., 1995). Microbial enzymes are often more useful than enzymes derived from plants or animals, because of the greater variety of catalytic activities available, the high yields possible, ease of genetic manipulation and rapid growth of microorganisms (Wiseman, 1995). 1.1. Lipase and its properties Lipases (triacyl glycerol acyl hydrolases) are one of the most important classes of industrial enzymes. They hydrolyze triglycerides into diglycerides, monoglycerides, glycerol and fatty acids (Sharma et al., 2001). Triglycerides are the natural substrates for lipases. These reactions are reversible and therefore, lipases also catalyse the formation of acyl glycerols from glycerol and fatty acids. During

1

hydrolysis, lipases pick acyl group from glycerides forming lipase acyl complex, which then transfers into acyl group to OH group of water. However, in non-aqueous conditions, these naturally hydrolytic enzymes can transfer acyl groups of carboxylic acids to nucleophiles other than water (Martinelle and Hult, 1995). The presence of lipase in bacteria had been observed as early as 1901 AD in Bacillus prodigiosus (now Serratia marcescens), B. pyocyaneus (now Pseudomonas aeruginosa), B. fluorescens (now P. fluorescens) and Staphylococcus pyogenesaucreus (now S. aureus) by the microbiologist Eijkmann (Jaeger and Eggert, 2003). Lipases are the enzymes catalyzing the hydrolysis of ester bonds and are widely distributed in animals, plants and microorganisms. In organic media, they catalyze reactions such as esterification, intesterification and transesterification (Kawamoto et al., 1987). Extracellular lipases have been proven efficient and selective biocatalysts in many relevant industrial applications from biosensors, chemicals, pharmaceuticals, pesticides, foods, leather, and cosmetics to detergents (Pandey et al., 1999). In 2000, enzyme market was one of the top in the biotechnology field and it was estimated at 1.6 billion US dollars (Demain, 2000). Many extensive work about lipases have previously been published, ranging from industrial applications, production, and immobilization to biocatalytical properties of pure enzymes (Kamini et al., 1998; Essamri et al., 1998; Nawani et al., 1998; Ramarethinam et al., 2002; Mahadik et al., 2002; Saxena et al., 2003; Ellaiah et al., 2004). Lipases possess a wide range of catalytic properties which are mostly strain-dependent. They have frequently been used in the form of a crude extract for synthesis of chiral building blocks and

2

enantiomeric

compounds.

So,

catalytical

properties

such

as

specificity,

enantioselectivity, and operational parameters like thermostability, and optimum pH, among others are relevant because they define the enzyme application range and type of process. Lipases from several microorganisms have been studied. Generally, bacterial lipases have neutral or alkaline pH value and show activity in a broad pH range (pH 4 to 11). The thermal stability of lipases ranging from 20 to 60oC. Stability of organic solvents is desirable in synthesis reaction. Most of the bacterial lipases are stable in organic solvents (Gupta et al., 2004). According to substrate specificity, microbial lipases are divided into three categories; such as nonspecific, regiospecific and fatty acid-specific. Nonspecific lipases behave randomly on the triacyglyceride molecule and produce complete breakdown of triacyglyceride to fatty acid and glycerol. Conversely, regiospecific lipases are 1,3 specific lipases which hydrolyze only primary ester bonds (ester bonds at atoms C1 and C3 of glycerol) and thus hydrolyze triacylglyceride to give fatty acids. Fatty acid specific lipases display activity in the presence of fatty acids (Thomson et al., 1999 and Gupta, et al., 2004). Lipases (EC.3.1.1.3, triacylglycerols acyl hydrolases) are group of enzymes, which have the ability to hydrolyze triacylglycerols in oil-water interface to release free fatty acids and glycerol. According to Joseph et al. (2008), lipases catalyze a wide range of reactions, including hydrolysis, inter-esterification, alcoholysis, acidolysis, esterification and aminolysis. In the presence of organic solvents, the enzymes are effective catalysts for various inter-esterification and trans-esterification

3

reactions. Especially microbial lipases have different enzymological properties and substrate specificities. Their biotechnological potential is relying on their ability to catalyze not only the hydrolysis of a given triglyceride, but also its synthesis from glycerol and fatty acids. Therefore, microbial lipases have many industrial applications (Jaeger et al., 1999). The temperature stability of lipases are the most important characteristic for industrial use (Choo et al., 1997). The extracellular bacterial lipases are commercially valuable, because their bulk production is much easier (Gupta et al., 2004). Lipase catalyzed reactions are widely used in the manufacture of

fats and oils, detergents and degreasing formulations, food

processing, the synthesis of fine chemicals and pharmaceuticals, paper manufacture, and production of cosmetics (Rubin and Dennis, 1997a,b; Kazlauskas and Bornscheuer, 1998). Lipases are also used to accelerate the degradation of fatty wastes (Masse et al., 2001) and polyurethane (Takamoto et al., 2001). Lipases catalyse both the hydrolysis and the synthesis of esters formed from glycerol and long-chain fatty acids. These reactions usually proceed with high regioand/or enantioselectivity, making lipases an important group of biocatalysts in organic chemistry. The reasons for the enormous biotechnological potential of microbial lipases include the facts that they are stable in organic solvents do not require cofactors (Rubin and Dennis, 1997a), possess broad substrate specificity and exhibit a high enantioselectivity (Kazlauskas and Bornscheuer, 1998). A number of lipases have been produced commercially, with the majority of them originating from fungi and bacteria. White and White (1997) reported the commercially

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available triglycerol lipases listed enzymes from 34 different sources including 18 from fungi and 7 from bacteria. These microbial lipases that are appeared to be the most widely used enzymes in biotechnology. 1.2. Microbial sources for lipase Lipases occur widely in nature, but microbial lipases are commercially significant because of low production cost, greater stability and wider availability than plant and animal lipases. They may originate from fungi, molds or bacteria and most of them are formed extracellularly. This ready availability has created an enormous spin-off with respect to the enantioselective hydrolysis and formation of carboxyl esters (IUPAC, 1997). Many authors evidenced that, the lipases are ubiquitous enzymes and have been found in most organisms from the microbial (Wohlfahrt and Jaeger, 1993; Jaeger et al., 1994), plant (Huang, 1993; Mukherjee et al. 1994) and animal kingdom (Carriere et al., 1991 & 1994). Wooley and Petersen (1994) reported that, the lipase was prepared either by extraction from animal or plant tissues, or by cultivation of microorganisms. Similarly, Boel et al. (1996) documented that the commercially available lipases are usually derived from microorganisms. Since the advent of genetic engineering techniques, an increasing number of lipases is being commercially manufactured from recombinant bacteria and yeasts.

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Most commercially useful lipases are of microbial origin. Many species of bacteria, fungi, yeast and actinomycetes are found to produce lipase (Liu et al., 2008). Roseau and Jaeger (2000) documented that, the bacteria produce different classes of lipolytic enzymes including carboxylesterases (EC 3.1.1.1) and hydrolyzing water-soluble esters and lipases (EC 3.1.1.3), which hydrolyze longchain triacylglycerol substrates. Literatures revealed that, some important lipase producing bacterial species are Bacillus sp. (Saxena et al., 2011), Pseudomonas sp. (Simpkin et al., 1991; Queiroz and Nascimento, 2002) and Burkholderia sp. (Liu et al., 2012). Some of the strains like Staphylococcus sp. (Rahman et al., 2006), Lactobacillus sp. (Padmapriya et al., 2011), Streptococcus sp. (Brault et al., 2012) and Propionibacterium sp. (Miskin et al., 1996) are reported to be the lipase producing Gram positive bacteria; whereas Chromobacterium sp. (Taipa et al., 1995), Acinetobacter sp. (Khoramnia et al., 2011), Vibrio sp. (Amoozear et al., 2008) and Aeromonas sp. (Charoenpanich et al., 2011) are such Gram-negative bacteria. The lipase producing fungi include Aspergillus sp. (Aulakh and Prakash, 2010), Rhizopus sp. (Kobliz and Pastore, 2006), Penicillum sp. (Rigo et al., 2009) and Mucor sp. (Abbas et al., 2003). While among yeasts, the common lipase producers are Candida sp. (Tan et al., 2006), Yarrowia sp. (Kamzolova et al., 2005), Saccharomyces sp. (Takahashi et al., 1998), and Trichosporon sp. (Suresh Kumar and Gupta, 2008). Further, Streptomyces sp. (Aly et al., 2012) has been identified as the lipase producing actinomycetes. On the other hand, some microbiologists have paid enough attention to screening few lipase producing strains from some oil sludge areas or from industrial wastes (Ertugrul et al., 2007; Kiran et al., 2008)

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The lipase producing microorganisms have been isolated from various sources, for e.g. from the gut sample of abalone Haliotis discus hannai (Sawabe et al.,1995), raw milk (Lotrakul and Dharmsthiti, 1997), raw meat (Labuschagne et al., 1997), spoiled coconut (Ray et al., 1999), hot springs (Lee et al., 1999; Markossian et al., 2000), contaminated water samples (Haba et al., 2000), soil samples (Cardenas et al., 2001), palm fruit (Abbas et al., 2002), crude oil contaminated soil samples (Kanwar et al., 2002), oil contaminated soils (Sirisha et al., 2010), tripod fish gut sample (Selva Mohan et al., 2012), etc. The bacteria present in the aquatic environment influence the composition of the gut biota and vice versa as the host and micro-organisms share the ecosystem (Verschuere et al., 2000). Gut micro flora play an important role in the digestive process, growth and disease of the host (Finegold et al., 1983). Several authors reported that the predominant bacterial species isolated from most of the fish digestive tracts have been reported to be aerobes or facultative anaerobes (Trust and Sparrow, 1974; Bairagi et al., 2002; Saha et al., 2006). However, information regarding the enzyme producing intestinal bacteria, their source and significance in fish is scarce (Kar and Ghosh, 2008). Further, Sivasubramanian et al. (2012) evidenced that, the intestinal isolates produced for extracellular enzyme and also examined for their antibacterial ability against fish and human pathogens. Great number of researchers have reported that the presence of a lot of bacteria in digestive tract especially in the intestines (Liston, 1956; Shewan, 1961; Matthes, 1966 a, b, c, d; Ozaki, 1972). In general as many

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number of research findings described that bacterial flora on the body surface and gills are unlike the specific bacteria to be found in the intestinal flora of fish (Liston, 1954 & 1957; Spencer, 1961; Aiso et al., 1968; Okuzumi and Horie, 1969; Chung and Kou, 1973; Trust, 1974). A simple and reliable method for detecting lipase activity in microorganisms has been described by Sierra (1957). Lipolysis is observed directly by changes in the appearance of the substrate such as tributyrin and triolein, which are emulsified mechanically in various growth media and poured into a Petri dish. Lipase production is indicated by the formation of clear halos around the colonies grown on tributyrin containing agar plates (Jaeger et al., 1994 and Ertugrul et al., 2007). There must be three factors to detect a lipase-positive bacterium by culturing it. These factors include (i) growth of the organism, (ii) production of lipase by that organism under suitable growth conditions and (iii) the presence of a sensitive method to detect lipase activity (Shelley et al., 1987). Lipase activity is identified by using triacylglycerols composed of long-chain fatty acids. Substrates like Tweens and tributyrin can also be used for the detection of lipases (Jensen, 1983 and Shelley et al., 1987). In direct observation methods, the formation of clear or turbid zones around colonies or the production of crystals on the agar surface display the presence of lipolytic activity. This method is highly useful for only screening of lipase production, but not to measure lipase activity (Shelley et al., 1987). Likewise, Kouker and Jaeger (1987) documented that, the microbial lipase activity can also be

8

identified by using fluorogenic dye Rhodamine B. The method containing Rhodamine B as an indicator of the presence of lipase and olive oil as lipid substrate. Lipase producing bacteria form orange fluorescent halos around their colonies under UV light, but lipase negative bacteria do not show orange fluorescence upon UV irradiation (Kouker and Jaeger, 1987). The fluorescence is related to the formation of a rhodamine B long chain fatty acid conjugate (Jaeger et al., 1994). Likewise, Wang et al. (1995) used plates of a modified Rhodamine B agar to screen lipase activity in a large number of microorganisms. On the other hand, the synthesis and secretion of lipases by isolated and screening of bacteria is influenced by a variety of environmental factors. The production of extracellular lipases from bacteria is often dependent on nitrogen and carbon sources, inorganic salts, presence of lipids, temperature and availability of oxygen. Bacterial lipases are mostly released outside of the cell that is called extracellular enzyme. In order to improve the whole-cell lipase catalytic ability in organic solvent, the effects on synthetic enzyme production must be studied systematically and the culture condition needs to be optimized (Teng and Xu, 2008). Likewise, Rosenau and Jaeger (2000) evidenced that, the bacterial lipases influenced by nutritional and physicochemical factors; such as temperature, pH, nitrogen and carbon sources, presence of lipids, inorganic salts, stirring conditions, dissolved oxygen concentration etc. Microbial lipases are produced mostly by submerged culture (Ito et al., 2001), but solid state fermentation (SSF) method can also be used. Generally the lipase production is organism specific and it is released during the late

9

logarithmic or stationary phase (Ghosh et al., 1996; Sharma et al., 2001). Many studies have been undertaken to define the optimal culture and nutritional requirements for lipase production by submerged culture method (Elibol and Ozer, 2001). Commercially useful lipases are usually obtained from microorganisms that produce a wide variety of extracellular lipases. The submerged fermentation (SmF) is widely used in the enzyme industry and has advantages in process control and good yield of enzymes, the products in fermentation are relatively dilute and therefore the downstream process results in high volume of effluents (Nagy et al., 2006). A number of reports exist on influence of various environmental parameters such as temperature, pH, carbon and lipid sources, nitrogen sources, agitation and dissolved oxygen concentration on lipase production (Watanabe et al., 1977; Omer et al., 1987; Suzuki and Takahiro, 1998; Nahas and de Assis, 1998). Similarly, Sharma et al. (2011) documented that, the lipase production is usually coordinated with and dependent on the availability of triglycerides. Besides this, free fatty acids, hydrolysable esters, bile salts and glycerol are also stimulated lipase production. According to Pogaku et al. (2010) starch, glucose and sucrose are the major carbon sources that upscale the production of lipase. Among organic nitrogen sources, peptone and yeast extract are the excellent nitrogen sources that significantly favoured maximum lipase production. Next to carbon and nitrogen sources, metal ions, fatty acids, hydrolysable esters, tweens and glycerol are the important components, which favour lipase production. Likewise, Janssen et al. (1994)

10

documented that, the lipase production by thermophilic Bacillus sp. was increased several fold, when magnesium, iron and calcium ions were added in to the production medium. In the same way, Wang et al. (1995) reported that, the lipase production by Bacillus sp. A30-1 (ATCC 5384) required a complex medium that contained Ca2+, Mg2+, Na+, CO2+, Cu2+, Fe2+, K+, Mn2+, MO2+ and Zn2+. On the other hand, Ghosh et al. (1996) reported that the optimization of lipase production by submerged fermentation reached its maximum after 2 days of fermentation and the maximum activity decreased rapidly after the 5th day of incubation. These results make solid state fermentation (SSF) is an interesting alternative for microbial production of lipase. The SSF offers lower production cost of lipases when compared to the submerged fermentation as the medium employed can be obtained at low cost and the down stream processing is also easier. Likewise, Jaeger et al. (1999) evidenced that; the lipases produced by bacteria can catalyze both the hydrolysis and the synthesis of long-chain acylglycerols. These reactions usually proceed with high regioselectivity and enantioselectivity; therefore, lipases have become very important stereoselective biocatalysts used in organic chemistry. High-level production of these biocatalysts requires the understanding of the mechanisms underlying gene expression, folding, and secretion (Jaeger et al., 1999). 1.3. Purification and characterization of lipase Novel purification technologies are available to obtain homogeneity of lipase from a large number of bacteria, fungi and from a few plant and animal sources. The purification of lipases normally involves several steps depending upon the purity

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desired for food application (Schimidt-Dannert et al., 1994). Various authors have investigated the purification of the enzymes is a must for understanding the 3-D structure and the structure function relationships of proteins (Taipa et al.,1992; Aires-Barros et al. 1994; Saxena et al., 2003). Since most of the microbial lipases are extracellular, the fermentation process is usually followed by the removal of cells from the culture broth, either by centrifugation or by filtration. Several authors reported that, the cell-free culture broth is then concentrated by ultrafiltration, ammonium sulphate precipitation or extraction with organic solvents. About 80% of the purification schemes attempted have used a precipitation step, 60% of which use ammonium sulphate and 35% use ethanol, acetone or an acid, followed by a combination of several chromatographic methods, such as gel filtration and affinity chromatography. The final step of gel filtration normally yields a homogenous product (Schimidt-Dannert et al., 1994; Krieger et al., 1997; Ferrer et al., 2000; Kojima and Shimizu, 2003). On the other hand Chartrain et al. (1993) evidenced that, the purified lipase from P. aeruginosa MB5001 using a three step procedure viz. concentration by ultra filtration, followed by ion exchange chromatography and gel filtration. The purified lipase had a molecular mass of 29 KDa by SDS-PAGE. Lipases are reported to be monomeric proteins having molecular weight with in the range between16, 000 and 670,000 Daltons (Sharma et al., 2011). A Pichia burtonii lipase was purified to homogeneity by a combination of DEAE-Sephadex A-50 ion exchange chromatography, Sephadex G-100 gel filtration, and isoelectric

12

focusing. The purified enzyme was monoeric and had a molecular mass of 51KDa by SDS-PAGE (Sugihara et al., 1995). Also, Kim et al. (1996) reported that, the highly alkaline extracellular lipase of Proteus vulgaris was purified by ion exchange chromatography. The purified lipase had a maximum hydrolytic activity at pH 10.0 and its molecular mass was 31 kDa by SDS-PAGE. Similarly, Lin et al. (1996) purified an alkaline lipase from P. pseudoalcaligenes F-111 to homogeneity. The apparent molecular mass by SDS-PAGE was 32 kDa and the isoelectric pH was 7.3. Most of the lipase purification schemes described in the literature focused on purifying small amounts of the enzyme to homogeneity to characterize it. Little information has been published on large-scale processes for commercial purification of lipase. Most commercial applications of lipases do not require highly pure enzyme. Excessive purification is expensive and reduces overall recovery of the enzyme (Chisti, 1998). Many lipases have been extensively purified and characterized in terms of their activity and stability profiles relative to pH, temperature, effects of metal ions and chelating agents (Guit et al., 1991 and Malcata et al., 1992). Great number of authors have reported that, the thermostable lipases have been isolated from many sources, including P. fluorescens (Kojima et al., 1994), Bacillus sp. (Wang et al., 1995 and Sidhu et al., 1998 a, b), B. coagulans and B. cereus (El-Shafei and Rezkallah, 1997), B. stearothermophilus (Kim et al., 1998), Geotrichum sp. and Aeromonas sobria (Lotrakul and Dharmsthiti, 1997 and Macedo et al., 1997) and P. aeruginosa (Sharon et al., 1998). Similarly, Wang et al. (1995) reported that one

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of the more notable thermostable enzymes was isolated from a Bacillus strain. Also, this enzyme had maximum activity at 60ºC and retained 100% of its original activity after being held at 75ºC for 30 min. Likewise, Sidhu et al. (1998 a, b) documented that, the extracellular lipase isolated from Bacillus strains had an optimum activity at 50°C. This enzyme had half- life of 15 min at 75ºC and it was stable at various oxidizing, reducing, chelating agents, surfactants and inorganic solvents. On the other hand, many researchers investigated that, the lipase hydrolyzed from fat or oil is dissolved in an organic solvent (Kosugi et al., 1990; Yang and Rhee, 1991; Yang and Rhee, 1992; Kim and Rhee, 1993; Kosugi and Tomizuka, 1995; Yang and Russell, 1995). Likewise, Kosugi et al. (1995) reported that, the eicosapentaenoic acid (EPA) and docosaheaenoic acid (DHA) were liberated continuously from sardine oil through hydrolysis process of immobilized Candida rugosa lipase. 1.4. Separation of PUFA compounds (EPA and DHA) from fish oil by lipase hydrolysis Lipases can be capable of catalysing the hydrolysis and synthesis of esters formed from glycerol and long-chain fatty acids (Svendsen et al., 2000; Sharma et al., 2001). Lipases are showing wide applications in organic chemical processing, detergent formulations, synthesis of biosurfactants, the oleochemical industry, the dairy industry, the agrochemical industry, paper manufacture, nutrition, cosmetics, and pharmaceutical processing. The only commercial sources of n-3 PUFA (EPA and DHA) and arachidonic (AA) acids are obtained from fish oils and animal viscera by hydrolysis of purified lipase (Shimizu et al., 1987). This poor

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availability can be compensated by the addition of n-3 long-chain polyunsaturated fatty acids (LC-PUFAs) from marine oil into food products. However, particular attentions have to be given at the quality of marine oil as the refining process involves thermal treatments which affect the integrity of LC-PUFAs (Fournier et al., 2006). Previously Klinkesorn et al. (2004) reported that, the enhancement of PUFA contents in fish oils by lipase and chemical modification. The specificities of lipase have proven useful allowing for the enhancement of the functionality of a variety of lipids via hydrolysis (Song et al., 2007; Yao et al., 2005). According to Valenzuela et al. (1993), the fish oils are good sources to obtain these long-chain polyunsaturated fatty acids due to the high concentration of EPA and DHA (20 to 28% EPA+DHA). Likewise, Nieto et al. (1999) reported that the preparation of sn-2 long-chain polyunsaturated monoacylglycerols from fish oil by hydrolysis with a stereospecific lipase from Mucor mietiei. Several researchers reported that, the lipases are known to catalyze hydrolysis reactions and have been shown as a good alternative for obtaining PUFA concentrates as acylglycerols (Shimada et al., 1994 and Okada et al., 2007). Modification of lipase from marine oil normally involves a lipase-catalyzed process. The concentration of both EPA and DHA may be prepared by selective hydrolysis of fish oils by using lipases, which discriminate against n-3 PUFA (Hoshino et al., 1990; Shimada et al., 1995; Wanasundara and Shahidi, 1998).

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Lipase- assisted n-3 PUFA enrichment of fish oil may also be obtained by tranesterification reactions (Haraldsson et al., 1997). Similarly, Breivik et al. (1997) pointedout that, the lipase-catalyzed enzymatic production of EPA and DHA concentrates from fish oil has shown potential in producing a high quality product, due to the mild conditions (e.g., neutral pH and low temperatures) of the process. Commercial lipases from Candida rugosa (cylindracea), Geotrichum candidum, Humicola lanuginose, Chromobacterium viscosum, and Rhizomucor miehei have been largely used for these purposes (Shimada et al., 1994; Wanasundara et al., 1998; Gunstone, 1999; Okada et al., 2007). Similarly, Carvalho et al. (2009) evidenced that the salmon oil (n-3 PUFA content of 30.1%) was hydrolyzed with three

kinds

of

native

microbial

lipases

(Aspergillus

niger,

Rhizopus

javanicus and Penicillium solitum). Among the high numbers of lipases described in the literature, only the enzymes belonging to a few species have been demonstrated to have adequate stability and selectivity to allow routine use for the concentration of PUFA. Some researchers have reported that lipases from Candida species are effective for the enrichment of PUFA via selective hydrolysis (Tanaka et al., 1992; Rahmatullah et al., 1994; Shimada et al., 1995; Huang and Huang, 1997). Few studies have been carried out to exploit the capacities of novel microbial lipases to concentrate the content of PUFA in oils (Shimada et al., 1994; Stránský et al., 2007). Unsaturated fatty acids have been recommended for replacement of saturated fatty acids in human diet, since saturated fats were demonstrated to be correlated with cardiovascular diseases (Keil, 2000). Similarly, Freese et al. (1973) evidenced that,

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the fatty acids function as the key ingredients of antimicrobial food additives due to their inhibitory action on undesirable microorganisms. 1.5. Antimicrobial activity of n-3 polyunsaturated fatty acid extracts from fish oil Lipid oxidation is a critical problem during food processing, distribution, storage, and consumption as it decreases food quality, stability, safety, and nutritive value (Luther et al., 2007). Several bioassay reports have indicated the presence of antimicrobial compounds among marine flora and fauna (Mariana et al., 2000; Constantina et al., 2004). EPA and DHA inhibit the growth of biohydrogenasing bacterial and other bacteria in pure culture (Maia et al., 2010). As a consequence, bacterial communities are changed significantly in response to dietary fish oil (Kim et al., 2008; Huws et al., 2009 b). The antibacterial activities of fatty acids in general have been noted in several bioengineering studies (Willet and Morse, 1966; Kabara et al., 1972; Miller et al., 1977; Sun et al., 1977). Fish oil from a sardine species Sardnops melanostica has also been shown to inhibit microbial growth (Rybin et al., 1999). However, studies related to antibacterial activity of PUFA extracts from fishes have been very few till date. Marine lipids, especially n-3 fatty acids such as eicosapentaenoic acid (EPA; 20:5 n-3) and docosahexaenoic acid (DHA; 22:6 n-3), have been documented as a major source of PUFA (Simopoulos, 1991; Narayan et al., 2006). Likewise, Shin et al. (2007) reported that, the antibacterial activity of EPA against food born and food spoilage microorganisms (Bacillus

subtilis

ATCC

6633,

Listeria

17

monocytogenes

ATCC

19166,

Staphylococcus aureus ATCC 6538, S. aureus KCTC 1916 and Pseudomonas aeruginosa KCTC 2004). Also, Shin et al. (2007) investigated the antimicrobial activity of bioconversion extracts of EPA and DHA against a range of food born pathogenic bacteria like B. subtilis, E.coli, E. coli 0157: H7 (Human), Salmonella enteritidis and S. typhimurium. According to Som and Radhakrishnan (2011), the DHA rich extract from Sardinella fimbriata has an overall higher inhibitory activity against four bacterial strains (S. aureus, E. faecalis, E. coli and Pseudomonas) as compared to the DHA extract of Sardinella longiceps. However, PUFA extracts from both S. fimbriata and S. longiceps species showed inhibitory activity against both gram-positive and gramnegative bacterial strains (Shin et al. 2007). Previously, Desbois et al. (2008) reported that, the results on EPA and DHA showing activity against a range of both gram-positive and gram-negative bacteria. Long chain fatty acids are well-known to be inhibitory on gram positive bacteria even at low concentrations (Kabara et al., 1977; Kodicek, 1949). According to Shin et al. (2006), EPA can reduce the viability of P. aeruginosa. In their experiment, scanning electron microscopy (SEM) study of bacterial cells clearly exhibited the antibacterial effect of EPA evidenced by the damages found in the outer membrane of the cells when treated with EPA. Shin et al. (2007) later opinioned that DHA is even more potent against this bacterium. High positive results in this study for inhibiting cultures of P. aeruginosa could also be due to high DHA and EPA concentrations in both the extracts. Higher DHA concentration of

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S. fimbriata correlates with greater inhibitory effect on this bacterium. On other hand, Bommarius et al. (1995) evidenced that, the Pseudomonas fluorescens strain B52 was isolated and the lipase produced by it displayed high level enantioselectivity with (R) - tert – leucinate, an important pharmaceutical intermediate. Many authors reported that, the fatty acids (FAs) play an important cellular role as energy substrates, membrane components and precursors of lipid mediators. They are also involved in the permeability of ionic channels, in synaptic transmission and signal transduction path way (Ordway et al., 1991; Grader et al., 1994). During the recent years, it has become increasingly clear that FAs can likewise exert critical regulatory functions in the cell through direct activation or inhibition of genes (Amri et al., 1991; Jump et al., 1993). D 1.6. Molecular biology of lipase gene the recent ye Detailed studies on the mechanism of synthesis of enzymes by microorganisms have been limited primarily to intracellular enzyme systems (Pardee, 1961; Halvorson, 1960). Among the extracellular enzymes, the effect of specific compounds on the production of proteinases has been most actively investigated (Castafieda-Agullo, 1956; Van der Zant, 1957; Hartman et al., 1957; Hagihara, 1960). Modern genetic engineering approaches with protein engineering may help in maximizing the enzyme production for specific industrial need. For the first time, Sanchez et al. (2002) expressed a bacterial lipase gene of B. subtilis in baker’s yeast, S. cerevisiae. Lipase B that exhibited unique substrate specificity for long-chain cis-9 unsaturated triacylglycerols were over-expressed in Pichia pastoris

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(Catoni et al., 1997). Jaeger et al. (1999) stated that more than 70 lipases including atleast 47 different lipases from bacteria have been cloned and sequenced. Therefore, prospecting for novel lipase genes is of interest for both academic and industrial reasons. However, novel lipase genes can be difficult to isolate due to several factors, including toxicity of expression to heterologous hosts and a requirement for helper proteins to achieve functional lipase expression. In addition, the low homology observed between different lipase genes makes them difficult targets for PCR cloning. Generally lipase genes of bacteria were isolated from genomic library (Chung et al., 1991; Oh et al., 1999; Rahman et al., 2003). Isolation and cloning of the gene is very important as it is essential to get information regarding the gene organization and other molecular characteristics. The industrial importance of lipase makes it essential to clone and multiply the gene and the technique followed is referred as molecular cloning. It is also an important tool to understand and alter the structure, function and regulation of individual genes and their products. A novel gene coding for a LipA lipase was isolated and characterized from a metagenomic library by Couto et al. (2010). RT–PCR amplicons of genomic DNA and cDNA of the lipase encoding gene from fungus A. tamari FS132 (ATL) was recently cloned successfully by Shi et al. (2010). During the last few years, much progress has been made by investigating lipase genes from bacteria, especially from Pseudomonas. Nucleotide sequences of lipase genes are already known for the following gram – negative bacterial species; Pseudomonas fragi IFO – 3458 (Kugimiya et al., 1986), P. fragi

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IFO – 1209 (Aoyama et al., 1988), P. cepacia (Jorgensen et al., 1991), P. fluorescens SIK W1 (Chung et al., 1991) and P. glumae (Frenken et al., 1991). Furthermore, lipase genes from two gram – positive species have been published; viz Staphylococcus hyicus (Gotz et al., 1985) and S. aureus (Lee and Iandolo, 1986). Based on the above facts and considering the importance of lipase in food processing and pharmaceutical industry, the present study was undertaken with the following key objectives: 1. To isolate, screen and identify lipase producing bacterial strains from the gut of a marine fish Sardinella longiceps. Then to test the efficiency of different culture conditions on lipase production by prominent lipase producing strain through submerged fermentation. 2. To purify the produced lipase through ion exchange (DEAE Cellulose) and size exclusion chromatography (Sephdex G-75) methods and to determine its molecular mass. 3. To confirm the lipase activity through Native PAGE analysis and to determine the stability of purified lipase by using different culture conditions (organic solvents, metal ions, detergents, temperature and pH). 4. To extract the liver oil from trash fish Odonus niger through solvent extraction method and to perform the enzymatic hydrolysis of purified liver oil for the separation of PUFA especially DHA and EPA.

21

5. To test the antibacterial and antifungal activity of DHA and EPA through agar well diffusion method and to perform the MIC and MFC through two fold dilution method. 6. To identify lipase producing gene from the promising strain through RT-PCR analysis.

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2. GENERAL MATERIALS AND METHODS

The present study deals with optimization and production of lipase from marine bacterium and its application on production of biologically active PUFA concentrates. In this regard, Chapter – I accounts on isolation, screening and identification of lipase producing bacteria from the gut of a marine fish Sardinella longiceps. Chapter - II describes the optimization and production of lipase from promising bacterium through submerged fermentation. Chapter – III represents the process of purification and characterization of lipase. Chapter – IV details the separation of PUFA compounds from marine trash fish Odonus niger liver oil through lipase hydrolysis. Chapter – V describes the antimicrobial activity of PUFA concentrate. Chapter – VI deals with the identification of lipase producing gene from the promising bacterium through RT-PCR analysis. 2.1. Isolation, screening and identification of lipase producing bacteria For the present investigation, marine fish S. longiceps was collected from Colachal coast, Kanniyakumari District, Tamil Nadu and transported to the laboratory in ice cold condition. One gram of gut sample of S. longiceps was weighed and aseptically transferred into Erlenmeyer flask containing 99 ml of sterile saline water and serially diluted up to 10-6. 0.1 ml of each aliquot was plated on Petridish containing Zobell marine agar (ZMA) and incubated at 37°C for 24h. Then the total viable count (TVC) was recorded. Morphologically distinct bacterial

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isolates were restreaked and purified in ZMA. Pure cultures obtained were then stored in slants and kept as stock for further study. Isolated gut bacterial strains were individually subjected to primary screening for their lipase producing ability through different plate assay methods viz. spirit blue agar, Rhodamine B agar, phenol red agar and Tween 80 agar. The bacterial strains were then identified up to genus level through morphological, physiological and biochemical characteristics and subjected to secondary screening for their lipase producing abilities through various production media. Thereafter the maximum lipase producing bacterial strain was identified up to species level by PCR amplification of the 16S rRNA gene. BLAST analysis was carried out and compared with sequences of Gen Bank nucleotide database. 2.2. Production and optimization of lipase through submerged fermentation To meet the increasing demand of lipase, feasible method that aids bulk and cost effective production from microorganisms need to be developed. This could be achieved by optimization of culture conditions. In this regard, media optimization for the producer strain was performed by changing various culture conditions such as incubation time (h), agitation speed (rpm), pH, temperature (°C), inoculum size (ml), carbon sources, nitrogen sources, metal ions, triglycerides, surfactants and different concentrations of NaCl (%). During each course of experiment, the lipase production and protein content of supernatant of the lipase production medium were determined through spectrophotometric assay method (Kicawley et al., 2002).

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2.3. Purification and characterization of lipase The producer stain was cultivated in the optimized medium under shaking condition (150 rpm) for 48h at 37°C and then the culture broth was centrifuged at 10,000 rpm for 30 min to remove the cell pellet. The supernatant obtained was then precipitated by using 75% ammonium sulphate. After precipitation process, the protein sample was filtered based on the charges (Positive or negative) by ion exchange chromatography. The fractions (elutions) obtained were further tested to determine the level of lipase and protein contents by spectrophotometric assay method (Ogino et al., 2000). The high lipase activity rendering column fraction was then subjected to Sephadex G-75 for separation of protein based on the size of the molecules. The separated column fractions were individually investigated for their lipase and protein contents through Spectrophotometric assay method (Syed et al., 2010). Then the molecular mass of the maximum lipase activity rendering column fraction was determined through SDS and Native page analysis (Laemmli, 1970). Further, the purified lipase was characterized in terms of their stability profile relative to organic solvents, metal ions, detergents, temperature (°C) and pH. The purified lipase stability was measured by Spectrophotometric assay method (Kicawley et al., 2002). 2.4. Production of PUFA concentrates from trash fish O. niger liver oil through lipase catalysis The oil from the liver sample of trash fish O. niger was extracted by following Bligh and Dyer method (Bligh and Dyer, 1959). Then the extracted oil was

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purified by following various steps (Degumming, neutralization, drying, bleaching and deodorization) and the fatty acid profile of crude and purified liver oils was analyzed through Gas Chromatography (GC) (Hartman et al., 1973). From the GC result, the PUFA concentrates, especially EPA and DHA contents were quantified. Likewise, peroxide, acid, saponification and free fatty acid values of crude and purified liver oil samples were determined through titration assay methods. Further, the purified liver oil was enriched with different concentrations of lipase (0.25, 0.50 and 0.75%) through hydrolysis process (Shimada et al., 1998) and also the fatty acid profiles of these hydrolysed oils were determined. Then the PUFA components, especially EPA and DHA from the hydrolysed oil sample were separated and collected by preparative HPLC (AOCS, 1998). 2.5. Determination of antimicrobial properties of EPA and DHA The EPA and DHA separated from the PUFA concentrates were tested individually for their antimicrobial activity against the selected pathogenic bacterial and fungal strains by disc diffusion method. The minimum inhibitory concentration (MIC) of EPA and DHA against pathogenic bacterial strains and minimum fungicidal concentration (MFC) against pathogenic fungal strains were determined by two fold dilution method as described by Murray et al. (1995) and Onyewu et al. (2003).

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2.6. Identification of lipase producing gene of B. cereus MSU AS through RT-PCR analysis The RNA of the lipase producing prominent bacterium B. cereus MSU AS was extracted by a standard alkaline lysis procedure described by Brillard et al. (2008). Then the RNA was converted to synthesis of cDNA by RT- PCR. The amplification process was carried out by mixing the template cDNA with three different types of lipase gene primers (Ptz lipase gene, LipA lipase gene and phospholipase C gene), each at one time. After that, the amplified products were used to determine the lipase specific gene of B. cereus MSU AS by using 0.7% agarose gel electrophoresis (Mir Mohammad Sadeghi et al., 2008). 2.7. Statistical analysis The data obtained in the present study were expressed as Mean ± SD and were analysed using ANOVA test, student‘t’ test and subsequently with post hoc multiple comparison with SNK test at 5% level of significance using computer software STATISTICA 6.0 (Statosoft, Bedford, UK).

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CHAPTER -1 3. ISOLATION, SCREENING AND IDENTIFICATION OF LIPASE PRODUCING BACTERIA

3.1. Introduction Lipases have been isolated from microorganisms especially from bacteria, fungi and yeasts (Saeed et al., 2005). The common lipase-producing bacterial strains are Pseudomonas aeruginosa, P. fluorescens, Bacillus coagulans, B. cereus, Staphylococcus aureus, S. hyicus, Burkholderia glumae, B. cepacia, etc. (Kim et al., 1998; Simons et al., 1998; Ito et al., 2001). Similarly, the lipase producing fungal species are also identified, which belong to the genera Geotrichum, Penicillium, Aspergillus, Rhizopus and Ophiostoma spp. (Long et al., 1996; Hiol et al., 2000). The yeast strains such as Yarrowia lipolytica, Candida rugosam, C. valida, Saccharomyces lipolytica, S. crataegenesis, Rhodotorula glutinis, Pichia bispora and P. sivicola were recorded as lipase producers (Ellouz et al., 2003). Several authors have reported the lipase producing microorganisms are basically diversified in different environments including marine environment (WHO, 2004). For example Lactobacillus plantarum, Bacillus sp., B. thermoleovorans ID-1, B. thermoleovorans IHI-91, Achremonium murosum, Monascus mucoroides, Pseudomonas sp., Staphylococcus sp. and B. cereus are the lipase producers were isolated from the intestine of marine fishes (Balcázar et al., 2008). Wang et al. (2007) screened out 427 lipase producing yeast strains from gut of marine fishes,

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marine algae, seawater, sediments, and mud of salterns. Some of the reports also documented that, certain lipase producing microbes are isolated from various sources like fermented sausages (Lopes et al., 2002), sediments of oil contaminated areas (Dharmsthiti and Luchai, 1999), hot springs (Lee et al., 1999; Markossian et al., 2000), soil samples (Cardenas et al., 2001), contaminated water samples (Haba et al., 2000), crude oil sample (Kanwar et al., 2002), spoiled coconut (Ray et al., 2009), etc. The biological diversity of marine and estuarine environment provides a wide array of enzyme producing microbes (Chi et al., 2007). However, the bacterial flora that thrive in the gastrointestinal tract with diversified enzymatic potential plays a vital role in major part of the metabolism of the host animals (Clemments, 1997). The major advantage of bacterial species present in the gut of fish can influence the health, robustness of the host and induce extracellular enzyme production (Sivasubramanian et al., 2012). The microbial ecology of the gastro intestinal tract of variety of freshwater and marine fishes has been investigated intensively by many researchers during the last decade (Spanggaard et al., 2000; Ahmed et al., 2004, 2005; Ringø et al., 2006, 2006a; Skrodenyte-Arbaciauskiene et al., 2006; Hovda et al., 2007; Kim et al., 2007; Yang et al., 2007; Zhou et al., 2009). There are evidences that the alimentary tract of fish is a complex ecosystem, containing a large number of microorganisms. Numerous surveys of the bacterial flora in the gastro intestinal tract of fish have been made during the last twenty years. The bacterial flora of the gastro intestinal tract of

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fishes in general, represent a very important and diversified enzymatic potential. It is capable of producing proteolytic, amylolytic, cellulolytic, lipolytic, and chitinolytic enzymes, which are important for digestion of proteins, carbohydrates, cellulose, lipids and chitin (Bairagi et al., 2002; Gutowska et al., 2004). Different species of fishes and crustaceans have a specific resident of enzyme producing gut microbiota, for eg. some of the bacterial species such as Acinetobacter sp., Enterobacter sp. and Pseudomonas sp. isolated from the gut of fish Oncorhynchus sp. (Kordel et al., 1991); Aeromonas sp. and Lactobacillus sp. from the gut of Salvelimus alpines (Ringo et al., 1995); Enterovibrio sp. from the intestinal tract of Scophthalamus maximums (Thompson et al., 2002); Vibrio sp., Bacillus sp., Photobacterium sp. and Plesiomonas sp. from the intestine of shrimp Penaeus indicus (Vine et al., 2006) and Shewanella sp. isolated from the gut of Haliotis discus hannai (Izvekova et al., 2007). Esakkiraj et al. (2010) reported that, the potent lipolytic bacterium Staphylococcus epidermidis CMST Pi 1 was isolated from the gut of shrimp P. indicus. Lipase producing microorganisms and their lipase activity can be identified through a simple and reliable methods. It has been well described by Sierra (1957), who used a surfactant Tween 80 in a solid medium to identify a lipolytic activity. The formation of opaque zones around the colonies is an indication of lipase production by the organisms. Also, screening of lipase producers on agar plates is frequently done by using tributyrin as a substrate and clear zones around the colonies indicate the production of lipase (Cardenas et al., 2001). Wang et al. (1995) used

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plates of a modified Rhodamine B agar to screen lipase activity in a large number of microorganisms. Several screening methods can be used for determination of the presence of extracellular lipases from microorganism. In this screening process, different types of solid media are used with inducer substrates such as vegetable oils, standard triglycerides (tributyrin & triolein), Tween 80, and dyes have been widely described in the literature (Wang et al., 1995; Cardenas et al., 2001; Ko et al., 2005). Screening method involves measurement of fluorescence caused by the fatty acid released due to the action of lipase on olive oil. A quantitative fluorescence lipase assay is based on the interaction of Rhodamine B with fatty acid released during the enzyme hydrolysis of olive oil (Kouker and Jaeger, 1987). Esakkiraj et al. (2010) stated that the lipase positive organism, S. epidermidis CMST- Pi1 isolated from the gut of shrimp P. indicus utilised sprit blue agar medium supplemented with inducer of tributyrin for lipase production. Gopinath et al. (2005) documented that the lipase producers of different fungal strains were screened for their lipolytic activity by using Rhodamine B agar medium with lipid substrate of olive oil. Similarly, Savitha et al. (2007) identified few fungal strains (Aspergillus sp., Penicillium sp. and Mucor sp.) produce inducible, extracellular and alkalophilic lipase, which was determined by using Rhodamine B agar plates with olive oil. Singh et al. (2006) showed the lipolytic activity of Mucor miehei was screened by using phenol red plate supplemented with tributyrin and olive oil. Gilbert et al. (1991) documented the lipase producing capability of Pseudomonas aeruginosa EF2 was determined by using Tween 80 agar plate with Tween 80 solution.

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Lipase isolates were initially evaluated by conventional tests (Satpal et al., 2011). Many authors reported that, the molecular approaches provide a more complete picture about bacterial community composition than do cultured-based methods. Molecular methods enable characterization and quantification of the intestinal microbiota, while also providing a classification scheme to predict phylogenetic relationships (Brunvold et al., 2007; Liu et al., 2008; Zhou et al., 2007, 2009). Akanbi et al. (2010) evidenced that, the lipases-producing Bacillus species were identified by the conventional Gram-staining technique, Biochemical tests and Biolog Microstation system by using 16S rRNA gene sequencing. Tambekar and Dhundale (2012) identified 13 lipolytic Bacillus sp through physiological and 16S rRNA gene sequencing analysis. The purpose of the present study is to isolate, screen and identify the lipase producing bacterial strains from the gut of a marine fish Sardinella longiceps. 3.2. Materials and Methods 3.2.1. Collection of samples The bacteria used in the present study were isolated from the intestine of marine fish Sardinella longiceps (Plate 3.1) collected from the fish landing centre of Colachel fisheries harbour, Kanyakumari District, Tamil Nadu. 3.2.2. Enumeration of total bacterial population About 1g of fish intestinal sample was aseptically transferred into a sterile mortar and made it into slurry with 1ml sterile water and made up to 100ml by adding 99 ml of sterile distilled water. After thorough mixing, the homogenate 32

decimal dilutions of 10-1 to 10-6 were prepared by transferring 1ml each of the sample to 9ml of the saline (0.85% NaCl). From each dilution, 0.1 ml of the sample was poured into already prepared Zobell marine agar plates. The inocula were spreaded properly by using an L-rod. Control plates were also maintained along with inoculated plates for testing the sterility of medium. The plates were incubated for 24h at 37ºC and after incubation; number of colonies in each plate were counted using a colony counter (Plate 3.2). Average of the count obtained was calculated and total bacterial count per gram was calculated as No. of colonies x Dilution factor No. of bacteria /g = Weight of the sample Using colony morphology, shape, size and colour, major dominant flora were selected and streaked onto already prepared Zobell marine agar plates. Then the plates were incubated for 24h at 37ºC. After incubation, the plates were observed and pure cultures were isolated and maintained as stock culture in slants at 4ºC for further use. 3.2.3. Primary screening of lipase producing bacteria using spirit blue agar plate (Ghanem et al., 2000) Based on the colour, morphology and appearance, totally 20 different bacterial strains were isolated from the gut of fish S. longiceps. All the isolates were individually subjected for primary screening to determine their lipase production capability using spirit blue agar (SBA) medium. Briefly, each gut bacterial isolate was streaked on to spirit blue agar plates and incubated for 48h at 37ºC. After the

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specified period of incubation, lipase positive bacterial strains were identified and recorded based on the zone formation. Among the tested 20 bacterial strains, only 13 strains showed lipase positive result. They were designated as AB, AC, AD, AF, AI, BB, CB, CF, OSA, OSE, OSC, OSF and OSG (Plate 3.3). 3.2.4. Secondary screening of lipase producing bacteria at different composition of lipase producing basal media through submerged fermentation 3.2.4.1. Seed culture preparation Two loops full of 24h lipase positive slant cultures (AB, AC, AD, AF, AI, BB, CB, CF, OSA, OSE, OSC, OSF and OSG) were inoculated individually in sterilized seed culture medium (peptone - 0.5g; yeast extract - 0.5g; NaCl - 0.5g; distilled water – 50 ml; at pH 7) in 250 ml culture flasks. The flasks were incubated for 16-18h on a rotary shaker (150 rpm) at 370C. 3.2.4.2. Screening of different composition of lipase producing basal media by the selected isolates Five different basal media (BM1 – BM5) were evaluated for the production of lipase. The compositions of the different basal media evaluated are summarized below: Basal medium 1: Dextrose - 2g, peptone - 3g, MgSO4 - 0.2g, FeSO4 - 0.1g, NaCl 1g and pH – 6.0 (100ml). Basal medium 2: Tryptone - 0.6g, yeast extract – 0.2g, olive oil - 1.5g, CaCl2.2H2O - 0.02g, MgSO4.7H2O - 1g, FeCl3.6H2O - 1g and pH - 7 (100ml).

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Basal medium 3: Glucose - 1.0g, yeast extract - 0.2g, peptone – 0.50g, KH2PO4 0.10g, MgSO4 – 0.02g, NaCl - 1.0g, olive oil -0.5 ml and pH - 7 (100ml). Basal medium 4: Peptone - 5g, glucose - 1g, NaNO3 - 0.1g, MgSO4 - 0.1g, sunflower oil -0.3 ml and pH - 6 (100ml). Basal medium 5: Sucrose - 10g, KH2PO4 - 1g, NH4NO3 - 2g, MgSO4 - 2g and pH 4.8 (100ml). All the basal media were sterilized at 121ºC for 15min. After sterilization, each basal medium was individually inoculated with 5ml each of seed cultures of all the 13 lipase positive (AB, AC, AD, AF, AI, BB, CB, CF, OSA, OSE, OSC, OSF and OSG) strains. Then the flasks were incubated for 48h in a rotatory shaker (150 rpm). After incubation, the individual filtrates (culture supernatant) was considered as crude enzymatic extract and were subjected for the determination of lipase production by Spectrophotometric assay method described by Kicawley et al. (2002). 3.2.4.3. Lipase assay Modification of lipase assay (Kicawley et al., 2002) system consists of the following ingredients: 1ml of Tris HCl buffer (pH 7.2), 1ml of P-nitro phenyl palmitate (10mM) and 0.5 ml of culture supernatant. Appropriate controls were also maintained. The mixture was incubated for 15min on a rotary shaker (150 rpm) at 37°C. Then the absorbance was read at 400nm using UV-Vis Spectrophotometer (Techcomb-8500), the amount of lipase production was found with the help of

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P-nitro phenol standard graph. One unit of lipase activity is equivalent to one microgram of P-nitro phenol released under standard assay condition. Calculation Lipase present in the supernatant of the sample (U/ml)

OD of the Sample =

x Conc. of std. OD of Standard

3.2.5. Identification of lipase positive bacteria In the present study all the 13 lipase positive strains (AB, AC, AD, AF, AI, BB, CB, CF, OSE, OSC, OSF and OSG) were initially identified up to genus level based on their morphological, physiological and biochemical characteristics (Holt et al., 1994). However, among the 13 strains, maximum lipase producing strain OSA alone was further identified up to species level through 16S rRNA sequencing. 3.2.6. Confirmation of lipase production by the promising bacterium Based on the primary screening, secondary screening as well as the identification results, the maximum lipase producing bacterial strain OSA (Bacillus cereus MSU AS) was alone selected and further its lipase producing ability was confirmed through various plate assay methods such as Rhodamine B agar, phenol red agar and Tween 80 agar plates. 3.2.6.1. On Rhodamine B agar plate (Kouker and Jaeger, 1987) Briefly the OSA strain (B. cereus MSU AS) was streaked at the centre of petridish containing Rhodamine B agar medium (Rhodamine B - 0.001% (w/v), Zobell marine agar - 0.8% (w/v), NaCl - 0.4% (w/v), agar - 1% (w/v), olive oil - 2%

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and pH 6.5) and incubated for 3 days at 37ºC. Then the lipase producing ability of the candidate bacterial strain OSA was identified based on the formation of clear orange halo around the colony under UV light at 350 nm (Plate 3.4). 3.2.6.2. On phenol red agar plate (Singh et al., 2006) Phenol red agar medium (phenol red - 0.01g, glycerol tributyrin - 1ml, 10mM CaCl2 - 0.01g, agar - 2g and pH 7.3 -7.4) was prepared and then the promising lipase positive bacterial strain OSA (B. cereus MSU AS) was streaked at the centre of the plate and incubated at 37ºC for 24h. After incubation, the lipase producing ability of the bacterial strain OSA was identified based on the formation of yellow colour zone around the colony (Plate 3.5). 3.2.6.3. On Tween 80 agar plate (Schoofs et al., 1997) Tween 80 agar plate method is one of the earliest methods to confirm lipase producing bacterial strains. The medium (1.5g of peptone, 0.5g of NaCl, 0.1g of CaCl2, 1.5g of agar, 100 ml distilled water at pH 7) was prepared and autoclaved. As when the medium reaches ear touchable temperature (37ºC), then 1% (v/v) of Tween 80 solution (Himedia) was added and poured into the petriplate. Then the OSA strain (Bacillus cereus MSU AS) was inoculated on to the prepared Tween 80 agar plate and incubated at 37ºC for 48h. After incubation, the lipase producing ability of the bacterial strain OSA was confirmed based on the white clear crystal formed around the colony (Plate 3.6).

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3.3. Results 3.3.1. Enumeration of total bacterial population Total bacterial population (CFU/g) in the gut sample of fish S. longiceps recorded was maximum of 375 ± 4.08 CFU/g at10 -1dilution. But, further increase in dilution of 10-2 to 10-6 gradual reduction in bacterial population from 150 ± 4.89 to 8 ± 0.416 CFU/g was noticed (Table 3.1 and Plate 3.2). 3.3.2. Primary screening of lipase producing bacteria using spirit blue agar plate Result on primary screening inferred that, out of the tested 20 gut bacterial strains, 13 strains were found to show lipase positive. Here OSA was the only strain, which showed maximum lipase production with the zone formation of 17mm by hydrolyzing 1% glycerol tributyrin in Spirit blue agar plate. However, the other bacterial strains such as AB, AC, AD, AF, AI, BB, CB, CF, OSE, OSC, OSF and OSG were also showed low and moderate level of lipase production with the zone formation ranged between 5 and 15 mm in spirit blue agar medium (Table 3.2). 3.3.3. Secondary screening of lipase production in different basal media Five different basal media (BM1 to BM5) were used for screening of lipase production by all the 13 lipase positive strains (AB, AC, AD, AF, AI, BB, CB, CF, OSE, OSA, OSC, OSF and OSG). From the result, it was understood that, medium BM3 gave maximum lipase production (462.95 ± 1.4245 U/ml) by OSA strain, however the same strain produced minimum (99.02 ± 0.1081 U/ml) amount of lipase in medium BM4. Followed by, the strain AI recorded with maximum (262.10 ±

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1.1632 U/ml) lipase production in medium BM3 and it produced minimum (111.47 ± 0.1077 U/ml) amount of lipase in medium BM5. Among all the tested strains, strain AC produced very low (6.364 ± 0.008 U/ml) amount of lipase in medium BM4 (Table 3.3). Two-way ANOVA test conducted on lipase production as a function of variation between bacterial strains as well as variation between production media were statistically significant (F = 2. 965752 to 2. 984552; P