Characterization of Alginate Lyase from Microbulbifer Mangrovi Sp. Nov

Characterization of alginate lyase from Microbulbifer mangrovi sp. nov. DD-13T

Thesis Submitted to Goa University

For the degree of

Doctor of Philosophy in Biotechnology

Ms. Poonam Vashist

Department Of Biotechnology Goa University Taleigao- Goa 2014

Characterization of alginate lyase from Microbulbifer mangrovi sp. nov. DD-13T

Thesis Submitted to Goa University

For the degree of

Doctor of Philosophy in Biotechnology

Ms. Poonam Vashist

Under the supervision of: Dr. S. C. Ghadi Department Of Biotechnology Goa University Taleigao- Goa 2014

CERTIFICATE

This is to certify that the thesis entitled “Characterization of alginate lyase from

Microbulbifer mangrovi sp.nov. DD-13T” submitted by Ms. Poonam Vashist, for the award of the Degree of Doctor of Philosophy in Biotechnology is based on original studies carried out by him under my supervision.

The thesis or any part thereof has not been submitted for any other degree or diploma in any university or institution.

Place : Goa University Date : 25/06/2014

Dr. S.C. Ghadi (Research Guide) Professor, Department of Biotechnology Goa University, Goa -403 206, India

STATEMENT

As required by the Goa university ordinance OB-09.9(ii), I state that the present thesis entitled “Characterization of alginate lyase from

Microbulbifer mangrovi sp.nov. DD-13T” is my original contribution and that the same has been submitted on any previous occasions for any degree.

To best of my knowledge, the present study is the first comprehensive work of its kind from the area mentioned. The literature related to the problem investigated has been cited. Due acknowledgments have been made wherever facilities and suggestions have been availed of.

Place: Goa, India

Date: 25/06/2014

Poonam Vashist

ACKNOWLEDGEMENT

It gives me immense pleasure and satisfaction to convey my sincere thanks and gratitude to Dr. S.C. Ghadi, my guide and patron for enlightening me into the world of research, for teaching me to see things from different perspective, for sharing his vast knowledge in microbial biotechnology and for his patience, persistence and tolerance. Each and every day was new door-step of adventure and significance. Dr. S.C. Ghadi has been inspiring me to face courageously and intelligently the usual ups and downs of voyage of research.

It’s my fortune to gratefully acknowledge the support and encouragement of Dr. Usha Muraleedharan, Dr. Urmila Barros, Dr. Savita Kerkar, Prof.

U.M.X. Sangodkar, Dr. S K Dubey (Dept.of Microbiology) and Dr. Shanta Nair,

Scientist (NIO, Goa).

I will be privileged to thank Dr. Yuichi Nogi (JAMSTEC, Japan) for their kind cooperation and collaboration during this study. I also thank Dr. Y.

Shouche and Mr. Pankaj Verma, National Centre for Cell Sciences, Pune, Mr.

Sanjay Singh, NIO, Goa.

I would also like to acknowledge the Vice-Chancellor and the Dean of life sciences, Goa University for providing necessary infrastructure to carry out my research.

I would also like to to thank Dr. Rahul Mohan Sharma from NCAOR ,

Goa; Mr. Khedekar and Mr. Arif from NIO in helping me with the SEM pictures. I wish to express my gratitude to Shayna and Mr. Suhas NCAOR.

Words are short to express my deep sense of gratitude towards my friends and colleague Lillian and Shahin for being with me as confidence in my good and bad time. I also sincerely convey my thanks to Late Shri Ravi Chand for their encouragement and being there as my friendly guide.

I am privileged to thank Dr. Kanchana, Sudheer, Asha, Kuldeep, Nirmal

Prasad, Tonima, Surya, Imran, Preethi, Judith, Priyanka, Samantha, Parantho, Alisha, Amruta, Michelle, Shuvankar, Kirtidas, Delicia, Dr. Tomchou Singh,

Hanumanth, Deepa and M.Sc (Biotechnology) students for sharing delightful moments, help and co-operation during the study.

I convey my sincere thanks to Mr. Martin, Serrao, Ulhas, Amonkar,

Ruby, Vandana, Sadanand, Anna, Concessa, Neelima, Tulsidas, Bharath, Sumati and Samir for their assistance and great help in day to day laboratory work;

Administrative staff and library staff for their help and providing necessary facilities; Goa University for their financial assistance for this study.

I would like to pay high regards to my parents S.S. Vashisth and

Shakuntala Vashisth for their sincere encouragement and inspiration throughout my research work and lifting me uphill this phase of life. I owe everything to them. I wish to express my gratitude to my husband Amir

Kumbhar, in-laws Somanath D Kumbhar & Jyoti Kumbhar and my little daughter Shaivi and son Neehan, whose constant patience helped me to excel through my degree. Without their enduring support and sacrifices, this journey would not have been possible. I would also like to convey my thanks to Asha

Gavali who has proved to be a great help during this phase of my life. I also express gratitude to those who have contributed to my research directly or indirectly even though they remain anonymous.

Above all I thank the ALMIGHTY for giving me this opportunity and helping me to be patient & optimistic throughout my struggles and failures.

Poonam

Table of contents

Chapter 1 General introduction 1- 5

Research goal and significance 6 – 8

Chapter 2 Literature review 9- 35

Chapter 3 Screening for multiple polysaccharide degrading bacteria 36- 59

Chapter 4 Identification of Multiple polysaccharide degrading bacterial 60- 121 strain DD-13 Chapter 5.Optimization of the growth and culture conditions for 122- 148 enhancing production of alginate lyase from Microbulbifer mangrovi sp.nov. DD-13T Chapter 6. Purification and characterization of the alginate lyase from 149- 189 Microbulbifer mangrovi sp.nov. DD13T Chapter 7 Applications of alginate lyase. 190- 216

Summary and Conclusions 217- 221

Future prospects 222

References 223- 275 Appendix Paper published

LIST OF ABBREVIATIONS

A Absorbance at the given wavelength APS Ammonium persulphate

BSA Bovine serum albumin

O.D. Optical Density

EDTA Ethylenediaminetetraacetic acid

PAGE Polyacrylamidegel electrophoresis rpm Revolutions per minute

SDS Sodium dodecyl sulphate

TEMED NNN'N'- tetramethyl ethylene diamine

TLC Thin layer chromatography

U Unit

SCD Single cell detritus

G Guluronic acid

M Mannuronic acid

DP Degree of polymerization

SCD Single cell detritus

CPC Cetylpyridinium chloride

LPS Lipoopolysaccharides

Dedicated to my family &

Almighty……

CHAPTER 1: INTRODUCTION

Polysaccharides also known as glycans are branched or linear carbohydrate polymeric structure composed of repeating monosaccharide units bond together by glycosidic linkages. Based on the structure the glycans are either homoglycans (single type monosaccharides units) or heteroglycans (different monosaccharide units).

Polysaccharides are known to be an important class of biological polymer which usually functional as structural and/ or energy reservoir in living organisms.

Based on the charge on the polysaccharides, these are divided into three categories i.e. neutral such as guar gum, amylopectin, amylase, cellulose etc.; anionic polysaccharides such as alginates, carrageenan, xanthan, gellan etc.; and cationic polysaccharides such as chitin. Due to solubility and various substituent functional groups, these polysaccharides are recalcitrant and also referred as insoluble complex polysaccharides (ICPs). ICPs are the most abundant renewable resources on the earth.

Marine ecosystem comprises a major part of the biosphere and annually produces more than 2 billion tons of ICPs. These ICPs are associated with biofilms, algal blooms, planktonic organisms and shells of marine invertebrates (Kloareg and Quatrano, 1988;

Pakuski and Benner, 1994; Kurita, 2006).

Degradation of ICPs is an important component within global nutrient recycling and act as major sink of carbon in nature (Arrigo, 2005). Microorganisms produce various extra and intracellular polysaccharide hydrolyzing enzymes. These enzymes depolymerize long chains of polysaccharides by hydrolyzing the glycosidic linkages.

Various mechanisms involved in ICPs degradation by bacterial systems has been extensively reviewed (Salyers, et al., 1996). The ability of these microorganisms to ferment ICPs to simple sugars using polysaccharase and other enzymes has made the

1 utilization of these recalcitrant compounds easier. Hydrolysis of ICPs by enzymes produced intermediates that can be used as the valuable feedstock in aquaculture, antioxidants, nutraceuticals, antitumor, antidiabetic, skin whitening agents, thickeners, gelling agent or stabilizers of emulsions and dispersions. With rising understanding of biological functions of these marine ICPs, the utilization of these sources in pharmaceuticals, aquaculture and other biotechnology related industries has been improved significantly.

Alginate is an uronic acid co-polymer comprising β- D- mannuronic acid (M) joined by (1- 4) linkage to α-L- guluronic acid (G) (Haug and Larsen, 1962). These moities can be arranged in homopolymeric (Poly G / Poly M) or heteropolymeric (Poly

MG) blocks (Haug, et al., 1966). Alginate isolated from various sources differs in properties and molecular structures. These variations are either because of the arrangement of moities or different substitutions such as galactose substituted alginate, propylene glycol alginate, sulphated alginate or methylated alginate etc. Alginate is found in cell walls and intracellular spaces in brown seaweeds. Commercially used alginate is generally obtained from Laminaria, Ascophyllum and Macrocystis (Skjak-Barek, et al.,

1991). Heterotrophic bacteria belonging to two families Azotobacteriaceae and

Pseudomonadaceae also produce alginate. Bacterial alginate differs from algal alginate by having O-acetyl groups on 2 and/or 3 positions of D-mannuronate (Skjak-Barek, et al.,

1985). The size and the arrangement of these alginate monomers in the polymer affect the viscosity and gel forming ability.

Cellulose is one of the major organic polymer observed as primary cell wall of green algae and plants. It is fixed during photosynthesis (Shively, et al., 2001)

2 and is a linear homopolymer of β linked D-glucose units. The other major component of plant cell wall is hemicelluloses, which is chemically complex and contains numerous heteropolysaccharides such as arabinan, galactan, glucan, mannan and xylan (O'Sullivan,

1997). Hemicellulose is a polymer comprised of 20% of the total biomass of most plants being closely related to cellulose.

The second most profuse polysaccharide xylan is an vital element of the hemicellulose part of the cell walls. Xylans are as ubiquitous as cellulose in plant cell walls. Xylan comprises of left handed helix with six β (1-4) linked xylanopyranosyl residues per helix turn. Xylan a heteroglycan which contains substitutent groups such as

4-O-methyl-D-glucopyranosyl, acetyl and α-arabinofuranosyl residues. Xylan and cellulose account for more than 50% of plant biomass (Subramaniyan and Prema, 2002).

Xylo-oligosaccharides produced from xylan are considered as "functional food" or dietary fibers.

Carrageenan is a water soluble linear sulfated glycans extracted from certain red edible seaweeds. This polysaccharide consists of D-galactose units joined by α-1-3 and β-

1-4 linkages. Three different types of carrageenans exists in nature, namely iota- carrageenan or carrageenose-2,4-disulfate, kappa- carrageenan or carrageenose-4-sulfate and lambda- carrageenan or carrageenose-2, 6, 2-trisulfate (Van de Velde, et al., 2002).

Carrageenan differ from agar wherein α-1-4-linked galactose units are in D- configuration, whereas in latter they are present in L-configuration (Rees, 1969). They are far and wide employed in the food industry (dairy and meat products), primarily for their excellent capability of gelling, thickening and their ability to strongly bind to food proteins, thus promoting stabilizing activity.

Chitin, a polymer of β (1-4)-linked 2-acetoamido-2-deoxyD-glucopyranose. It is a naturally present as a component of crustacean exoskeletons, diatoms, radulae of molluscs, fungi as well as the beaks and internal shells of cephalopods in marine environments. X-ray diffraction studies have shown that chitin, naturally appears in α, β and γ polymeric forms. α-chitin being the most abundant in nature, the arrangement of polypeptide chains are anti-parallel compared to parallel β-chitin whereas in γ-chitin, the chains are present in miscellaneous form (Peberdy, 1985). It has also been assessed as a fertilizer, which can improve overall crop yields, binder in dyes, fabrics, and adhesives.

Agar is unbranched polysaccharide present commonly in red algae cell wall and is made of linear agarose units and heterogeneous mixture of agaropectin molecules.

Agarose is a linear series of 4-O-linked-3, 6-anhydro-α-L-galactose and 3-O-linked-β-D- galactopyranose with low degree of sulphation (2%) whereas agaropectin is a substituted agarose containing sulfoxy, methoxy or pyruvate groups. In red agae, agar occurs in pseudo crystalline form along with cellulose in cell wall matrix (Kloareg and Quatrano,

1988). Agar-agar is a natural vegetable gelatin counterpart and is 80% fiber which can be used as gelling agent to make jellies, puddings, and custards.

The microorganisms producing polysaccharide degrading enzymes are widely present and can be obtained from coastal water, sediments and mangroves. ICPs degrading bacteria have evolved in many different phylogenetic groups and are accountable for reprocess of organic carbon (Weiner, et al, 1998). Beguin and Aubert

(1994); Wong, et al., (2000); Howard, et al., (2003) and Michel, et al., (2006) reviewed bacterial strains degrading these ICPs and their enzyme systems. Few of these

4 polysaccharide degrading enzymes from bacteria have been characterized (Salyers, et al.,

1996). These polysaccharide degrading enzymes have been reported to have many functions in food, pharmaceutical, agricultural, leather industries as well as in bioremediation of algal wastes.

Marine organisms are richly endowed with diverse enzymes having unique properties. Marine organisms degrading ICPs are also reported from extreme environmental conditions such as low nutrients, high temperatures, salinity, hydrostatic pressure and radiation. The gradual research has uplifted the marine microbial enzyme technology to a much higher level in recent years, offering valuable bio-products. These bacteria produce diverse enzymes with unique catalytic functions and stability whose potential are still not amply explored. Studying polysaccharide degrading marine bacteria and their polysaccharase will provide a valuable insight about the role of these bacteria in ecosystem. Further, since the polysaccharases actively participate in carbon recycling of the ICPs, studying the biochemical properties of polysaccharase enzyme will help biologist to design novel applications in bioremediation of ICPs as well as explore unchartered potential applications.

RESEARCH GOAL AND SIGNIFICANCE

Biotechnology has influenced almost every sector of our activities for social and environmental needs. Currently only 5% of chemical products are produced using biotechnological methods. The world wide industrial enzyme market distribution on the basis of applications show that 34% of market is for food and animal feed followed by detergents and cleaners (29%). Pulp and paper share 11% while 17% is captured by textile, leather and fur industries (Binod, et al., 2013).

The oceans being wider niche opens abundant scope for research and development. Although the potential of this niche for new applications in biotechnology remains mainly still unknown. Indeed, a wide range of marine microorganisms have yet to be identified for the research field. Characterization of these bacteria and their extracellular enzymes will have a many applications.

In the marine environment the most important source for organic carbon is polysaccharides obtained from degradation of the insoluble complex polysaccharides which play an important role in recycling carbon. Several seaweeds, microbes, phytoplankton are natural resource for complex polysaccharides such as carrageenan, chitin, xylan, laminarin, agar, mannan and alginate.

Marine organisms are richly endowed with diverse enzymes having unique properties. A number of bacterial genera are capable to digest various polysaccharide which is important for many ecosystems (Salyers, et al., 1996). Polysaccharides can be degraded by specific glycosidase corresponding to either hydrolases or lyases (e.g. agarase, chitinase, xylanase, carrageenase and laminarase). The microorganisms producing polysaccharide-degrading enzymes are widely present and can be obtained

6 from coastal water, sediments and mangroves. Polysaccharide degrading enzymes have been quarantined from bacteria like Azotobacter vinelandii (Davidson and Lawson,

1977), Pseudomonas atlantica (Morrice, et al., 1984), Bacillus circulans (Hansen, et al.,

1984), Clostridium thermocellum (Gilad, et al., 2003), Beneckea pelagia and

Pseudomonas sp. etc. (Fett, et al., 1986; Sutherland and Keen, 1981). Few of these polysaccharide-degrading enzymes from bacteria have been characterized (Salyers, et al.,

1996). Many of these bacteria are epiphytic and are found in close association with seaweeds. Further, polysaccharide degrading bacteria have been observed to establish a parasitic or saprophytic role in relation to seaweeds (Ensor, et al., 1999).

Almost 50% of polysaccharide degradation products obtained by enzymatic hydrolysis have been used widely in pharma sector as some of these products have demonstrated anti-tumor, anti-tyrosinase, skin whitening and can also be used for wound dressing, treatment of cystic fibrosis, induction of apoptosis etc (Higuchi et al., 2003).

The use of enzymatic degraded polysaccharide products in the field of nutraceuticals and agriculture is escalating rapidly as these products are proved to have antioxidative properties and promotes root-shoot elongation and germination (Xu, et al., 2003; Iwasaki, et al., 2000).

Our research group has been extensively working on polysaccharide degrading bacteria that have been screened from marine ecosystem. Agarolytic bacteria have been isolated from the coast of Lakshadweep as well as from decomposing seaweeds (Ghadi, et al., 1999; RaviChand, et al., 2009). Agarase from Microbulbifer has been purified and characterized (Ravichand and S.C. Ghadi, 2011). Although agarolytic bacteria have been

7 extensively screened and reported by several groups from India (Lakshmikanth, et al.,

2006; Khambhaty, et al., 2008; Lakshmikanth, et al., 2009), very few alginolytic bacteria have reported from international water in comparison to Indian ecosystem. Further predominant occurrence of brown seaweeds on the coast of Goa, conceive an attractive proposition to explore and study the unexplored culturable novel alginolytic bacteria.

Thus the main focus of the proposed study relates to polyphasic approaches to identify a novel multiple polysaccharide degrading bacterium isolated from the mangrove of Goa, purification and biochemical characterization of alginate lyase and the application of strain DD-13 and alginate lyase in biotechnology.

The objectives of the Ph.D. research work were:

. Screening for polysaccharide degrading bacteria from various marine niches.

. Biochemical characterization of the selected bacterial isolate.

. Molecular phylogenetic identification of the bacteria using 16S rDNA universal primers.

. Purification and characterization of the polysaccharide degrading enzyme from the selected bacterial isolate

. Applications of this bacterial polysaccharide-degrading enzyme in Biotechnology.

CHAPTER 2: LITERATURE REVIEW

This chapter elaborately describes the chemistry, properties and applications of alginate polysaccharides. Further a detailed survey of alginolytic bacteria, their occurrence, alginate lyase enzyme, classification and the mechanisms involved in degradation of alginate polysaccharide is presented. This chapter also introduces various methodologies adopted for purification of alginate lyase, biochemical properties of the purified alginate lyase and molecular structures unraveled so far. Finally this chapter ends with a detailed review on various applications of alginate lyase enzyme.

2.1 ALGINIC ACID:

Alginates are abundantly found in nature constituting 40% of the dry weight as cell wall component in marine algae, as well as component of capsular polysaccharides in few bacterial species. Brown seaweeds with-stand dehydration over a prolonged period during low tides. This is attributed to the presence of alginic acid which retains the water molecules and protects it from drying. The presence of alginate in cell wall of brown seaweed gives the seaweeds both plasticity and mechanical tenacity. (Andresen, et al.,

1977). The physiological properties of alginates in marine brown algae are comparable to those of cellulose present in terrestrial plants. Most of the alginate used commercially is obtained from biomass of marine macroalgae mainly belonging to three genera,

Ascophyllum, Laminaria, and Macrocystis for example Laminaria hyperborea,

Laminaria japonica, Laminaria digitata, Ascophyllum nodosum, Macrocystis pyrifera, other species such as Sargassum sp., Lessonia nigrescens, Durvillea antarctica, Eclonia maxima etc. Two families of heterotrophic bacteria namely, Azotobacteriaceae and

Pseudomonadaceae are also known to produce alginate.

2.2 HISTORICAL OUTLINE OF ALGINIC ACID:

In the late late nineteenth century, E. C. C. Stanford, a British chemist discovered alginic acid from seaweeds and a patent filed on 12 January 1881 (Stanford, 1881).

Stanford deciphered the chemical structure of alginic acid and believed that alginic acid is a nitrogenous viscous/ gelatinous polysaccharide extract.

The discovery that alginic acid is composed of uronic acid was independeltly discovered [Schmidt and Vocke, (1926) and Atsuki and Tomoda, (1926)]. Later, Nelson and Cretcher, (1929, 1930, 1932); Bird and Haas, (1931); Miawa, (1930) studied the nature of the uronic acids present in alginic acid and presented a very simple picture of alginic acid composed of D-mannuronic acid. Hirst, et al., (1939) demonstrated that D- mannuronic acid are coupled together by β-1, 4 bonds similar to those found in cellulose.

The structure of alginic acid was again reviewed by Fischer and Dorfel (1955). The binary structure of alginic acid was reported to consist of β-D-mannuronic and α-L- guluronic residues and a method for estimation of guluronic and mannuronic acid was determined. Later by partial acid hydrolysis of alginate, Haug and coworkers separated alginate into 3 fractions/ blocks of β-D-mannuronic and α-L-guluronic residues namely

GG, GM/ MG and MM blocks (Haug, 1964; Haug, et al., 1966; Haug and Larsen, 1966;

Haug, et al., 1967a; Haug and Smidsr˘d, 1965).

2.3 CHEMISTRY OF ALGINIC ACID:

The widely varying blocks of covalently (1-4) linked β-D-mannuronic acid (M) and α-L-guluronic acid (G) residues constitutes to give rise to unbranched dual copolymer called alginic acid (Figure 2.1). Haug and coworkers concluded that alginate

10 could be either homopolymer with regions of G or M, termed as GG or MM blocks, respectively; or a heteropolymeric regions of M and G termed as MG/ GM blocks (Fig.

2.1).

G G M M α-1, 4 β-1, 4 β-1, 4

Figure 2.1: Chemical structure of alginate representing linear chain of β-D-mannuronate

(M) and its C5 epimer, α-L-guluronate.

A precise structure of alginic acid was determined by employing high-resolution

1 14 H4 and C NMR-spectroscopy (Penman and Sanderson, 1972; Grasdalen, et al., 1977,

1979; Grasdalen, 1983).The basic structure of each monomer is the tetrahydropyran ring.

According to Atkins, et al., (1970) X-ray diffraction studies of alginate polymer

1 demonstrated that the G residues in homopolymeric blocks are arranged in C4

4 conformation whereas the mannuronate residues have the C1 conformation. Thus, the structure of alginate was maintained by diaxial (GG), diequatorial (MM), axial-equatorial

(GM) and equatorial-axial (MG) glocosidic associations (Fig. 2.1).

The prominent variation between bacterial and seaweed alginates is that the latter demonstrated the existence of O-acetyl groups at C2 and/or C3 (Skjak-Braek et al., 1986).

Acetylation disrupts the water-binding ion-binding selectivity properties of the polymer

(Skj˚ak-Bræk, et al., 1989; Geddie and Sutherland, 1994; Skj˚ak-Bræk, et al., 1989). The sensitivity of alginic acid towards degradation is affected by both arrangement and degree of O-acetylation,

2.4 PROPERTIES OF ALGINATE:

The use of alginate is primarily governed by composition of uronic acid that in turn would alter the characteristics. The alginate molecules physical properties were discovered mainly during 1960s and 70s and indicated new insights on its gel forming characteristics.

The viscosity of the alginate extract is dependent on the presence of the sequence of M and G blocks. Investigation on viscosity data demonstrated that elasticity of the chain blocks augmented in the order of GG< MM< MG/GM (Smidsr˘d, et al., 1973;

Grasdalen, et al., 1977). The di-axial linkage in the G-blocks causes a hindered rotation on the glycosidic bond leading to stiffness of GG block polymer. (Smidsr˘d, et al., 1973).

The composition and sequential structure of alginate varies as per the growth environment and seasonal variations (Indergaard and Skjak-Braek, 1987; Haug, 1964).

For example, the stripe and holdfast of L. hyperborean depict a high level of G content conferring an elevated mechanical firmness whereas lower G-content of leaves possess more supple texture. Further, alginates from L. japonica, Macrocystis pyrifera and A.

12 nodosum are reported to have low G-blocks content and hence little gel potency.

Bacterial alginates with 100% mannuronate have also been reported (Valla, et al., 1996).

2.4.1 Gel formation:

The alginate gel formation is based on the property of selective binding of cations and the sol/gel transition is partially independent of temperature variability (Haug, 1964;

Smidsrod and Haug, 1968b; Haug and Smidsrod, 1970; Smidsrod, 1973 and1974). Due to excellent capability of alginate to absorb water e.g one part of alginate can retain 300 parts of liquid leading to excellent gel forming capabilities and thus are exploited as thickener in food industry.

2.4.2 Stability:

The stability of alginates decreases with increases in viscosity. Low viscosity sodium alginates stored at 10- 20°C are more stable for an average of three years whereas as alginates with medium viscosity range (up to 400 mPa.s)demonstrated 10-45% loss in stability at 25 and 33° C respectively after a year. Highly viscous alginates are relatively highly unstable.

2.4.3 Molecular Mass:

Akin to other known polysaccharides, alginate are also polydispersed especially with respect to occurrence of G, M or GM blocks in the polymer. Thus molecular weight of alginate is calculated as an average of molecular weights determined from the presence of G,M and GM blocks. Thus alginate with low proportion of G blocks will not participate in formation of network leading to poor gel strength. Simultaneously, the use

13 of alginate with high proportion of M blocks in high end applications is equally not feasible due to possibility of leakage (Stokke, et al., 1991; Otterlei, et al., 1991). Thus a medium molecular weight alginate is the preferred choice in most applications.

2.4.4 Solubility:

Similar to other known polysaccharides, alginate is insoluble in water as well as organic solvents. However salts of alginate demonstrate variable properties. Sodium, magnesium or potassium salts of alginate are known to yield viscous solution, while calcium, iron or zinc salts of alginates are insoluble in water. The three crucial criterias namely pH, ionic strength and the concentration of alginate determine the alginate solubility in aqueous solution. A gradual and composed decrease in pH may result in the gel formation in the alginate solution whereas a sudden pH change due to addition of acid leads to alginate precipitation of alginate blocks (Haug, 1959a; Haug and Larsen, 1963;

Haug, 1964;; Myklestad and Haug 1966;). Alginate also starts precipitating at high ionic strength of inorganic salts such as KCl (Haug and Smidsrod, 1967; Haug, 1959a).

2.5 Applications of Alginic acid/ Alginate:

Alginate is considered as at multifunctional resourceful polysaccharide based on its applications in various fields including traditional technical utilization in various industries, food and biomedicine. Among many useful applications of alginates, some are listed below:

o Shear thining viscosifier in textile printing, resulting in good colour yield,

brightness and print levelness.

o Certified to produce uniform surface on paper by coating with alginate.

o As binding component for construction of welding rods. (Onsoyen, 1996).

o Wound dressing and as dental impression materials.

o Immobilization of living cells by entrapment of cells within Ca-alginate beads

that can be further used for production of bioethanol and monoclonal antibody

production using hybridoma cells and mass production of artificial seed

(Smidsrod and Skjak-Braek, 1990); as well as cell transplantation (Aebisher, et

al., 1993; Soon-Shiong, et al., 1993, 1994; Read, et al., 2000).

o The alginate fragments also triggers immune responses that have been studied

using in-vivo animal models (Stokke, et al., 1993; Espevik and Skjak- Braek,

1996).

o As food supplements to enhance, reform and preserve the taste as well as

consistency (Cottrell and Kovacs, 1980; Sime, 1990; Littlecott, 1982; McHugh,

1987).

o To make synergestic gels of propylene glycol alginate (rich in guluronate) and

pectins for production of fruit fillings, jellies, fruit pulp extract etc. (Toft, et al.,

1986)

Some of the alginate properties described below merit exclusive consideration

(http://seafarmacy.co.uk/alginate-and-alginic-acids):

. Alginates elicit antioxidant activity of lipid peroxidation facilitating revitalization

of the digestive tract..

. Alginate lowers the blood glucose level in diabetic patients by binding water in

the gut, prohibiting it to split and imbibe carbohydrates resulting in decrease

hyperglycaemia effect..

. It helps in body detoxification evacuating heavy metals (lead, mercury, etc.) such

as barium (Ba), lead (Pb), strontium (Sr), cesium (Cs) getting replaced with

sodium (Na), calcium (Ca), potassium (K), magnesium (Mg) salts of alginates.

. Alginic acid and its salts evacuate dangerous radionuclide like Sr-90 and Cs-137.

. It acts as natural enterosorbent by binding and removing the toxic products such

as cholesterol, bile acids, carcinogens and radionuclides.

. The dietary fibres are potent stimulators of intestinal motility.

2.6 GLYCOSIDE HYDROLASES:

The major structural linkage in polysaccharides is glycosidic bond. It is considered to be the most eternal bond present in natural occurring polymers. It is stronger than phosphodiester linkage in DNA and stable than peptide and phosphodiester linkage in RNA. The expected half-life for glycosidic bond hydrolysis of the polysaccharide like cellulose is reported to be roughly 5 million years (Vivian, et al.,

2004). The most natural way to cleave the glycosidic bond is via hydrolysis. Based on their mode of action glycosyl hydrolases are classified as endo- glycosyl hydrolases or exo - glycosyl hydrolases, depending on where the enzyme cleaves the polymer. The exo-acting enzymes release the sugar residue with inverted C-1 configuration. The difference between exo- and endo- polysaccharases is that exoenzymes attack the free end and endo- enzymes bind to the internal regions of polysaccharide molecule.

Elimination is another mechanism evolved to cleave the uronic acid containing polysaccharides like alginic acid. Glycoside hydrolase (GH) (E.C. 3.2.1. - 3.2.3.), cleaves the glycoside bond between carbohydrates- carbohydrates or inbetween a carbohydrate- non-carbohydrate moiety. Due to direct rapport between folding similarities and sequence, enzymes have been classified on the basis of amino acid sequences. This categorization reveals the structural features of this group of enzymes, substrate specificity, evolutionary relationships and mechanistic information. Along with sequence based classification, crystal structure analysis of mutated & wild type glycosidases enzyme and ligands complexes as well as the kinetics and structure of transition state

(TST) and kinetic isotope effects have contributed for enhancement of mechanism details.

Hydrolysis of glycoside bond in an enzyme is due to the action of two catalytic residues, a nucleophilic base and a proton donor. Based on their positions, hydrolysis occurs through inversion or retention of the anomeric configuration. Retaining glycosides are endo- acting enzymes, and use two step double displacement mechanisms, as proposed by Koshland to form covalent glycosyl-enzyme intermediate through oxo- carbenium ion like transition state.

Retaining mechanism involves .

• Binding of enzyme to the polysaccharide substrate

• Glycosylation i.e. Cleaving the glycosyl-enzyme bond and forming an intermediate through the inversion of C-1 atom configuration.

• Deglycosylation i.e. Cleaving the covalently linked glycosyl-enzyme bond causing a second inversion of C-l atom which involve water molecule and deprotonated carboxylate residue.

The anomeric configuration of the Cl atom of the substrate is inverted twice during the catalysis. It involves the formation of oxo-carbenium ion transition state.

(Jedrzejas, 2000).

Polysaccharide lyases, the other class of polysaccharide degrading enzymes, works by elimination mechanism and the presence of an extended substrate

(polysaccharide) binding site is a common feature shared with hydrolyases. There are two active carboxylic acids, one results in the formation of glycosyl enzyme products through a nucleophilic action at anomeric centre. The other carboxylic acid group functions as acid/base catalyst, firstly to cleave the glycosidic bond, also known as general aid catalysis and is much more important than the second step which involves hydrolysis of glycosyl- enzyme, commonly known as general base catalysis. General base catalysis contributes15-19 KJ/mol (300 to 2,000 fold) to transition state stabilization and general acid catalysis depends on substrate and contribute 38 KJ/mol to transition state stabilization (Lay and Withers, 1999).

2.7 CLASSIFICATION OF POLYSACCHARIDE DEGRADING ENZYMES:

Enzymes are categorized based on their substrate specificity, mode of action and reaction products. The polysaccharases are classified based on amino acid sequence with their available 3-D structure obtained because of the well developed techniques for protein and genome sequencing. The classification based on sequences of polysaccharide

18 degrading enzymes can be obtained from the Carbohydrate Active enZYme server

(CAZY) at www.cazy.org . More than 6,500 carbohydrate active enzymes have been reported in public domain and classified into 106 families.

On the basis of type of reaction performed, this server identifies these enzymes into four different classes;

 Glycosyl Transferases (GT's): they act in polysaccharide synthesis by

forming new glycosidic bond by transferring sugar molecule from an

activated carrier molecule such as uridine diphosphate to acceptor

molecule. They also function in phosphorolytic cleavage of cellobiose and

cellodextrins.

 Polysaccharide Lyases (PL's): they act through β-elimination mechanism

in alginate and pectin depolymerization.

 Carbohydrate Esterase (CE's): they deacetylates the O- or N-substituted

polysaccharides in chitin and xylan deacetylation.

 Glycosyl Hydrolases (GH's): they hydrolase the glycosidic bonds in

cellulose, agar etc.

The classification based on sequences of polysaccharases families are subclassified into "clans" or "superfamilies" according to their 3-D structural analysis.

Enzymes considered within the same superfamily share common catalytic domain arrangement although they may be unrelated by function and sequence.

2.8 ALGINATE DEGRADING/ ALGINOLYTIC ORGANISMS:

Alginate degrading organisms are wide spread among diverse ecosystems.

Majority of the alginolytic organisms are from aquatic, particularly the marine environment. The efficiency of the alginate hydrolysis depends on the properties and relative concentrations of alginate lyase enzyme produced by alginolytic organisms.

Alginate lyases have been reported from various sources like marine organisms such as mollusks, algae, and many microorganisms. Alginase activity have been also reported in extracts obtained from a number of brown algal species, like Pelvetia canaliculata

(Madgwick, et al., 1978), Laminaria digitata (Madgwick, et al., 1973)and Undaria pinnatifida (Watanabe and Nisizawa, 1982). Alginate lyases have also been isolated from the hepatopancreas, style or gut glands of various marine mollusks. Additionally, alginate lyase was detected and isolated from Turbo cornutus mid-gut gland (Muramatsu, et al.,

1977), Littorina spp. hepatopancreas (Favorov, 1973), and Dollabella auricola (

Nisizawa, et al., 1968), Haliotis spp., Spisula solidissima (Boyen, et al., 1990b; Jacober, et al., 1980; Muramatsu, et al., 1977), and Choromytilus meridionalis crystalline style and Perna perna (Seiderer, et al., 1982). The alginate lyase presence in the guts of different mollusks may encourage brown algal tissue digestion.

Alginase have also been isolated from marine as well as soil fungi and bacteria.

Many of these life-forms are capable to using alginates as carbon and energy sources.

Several Many species of Azotobacters and Pseudomonads produces alginates as well as alginate lyases. Thjotta and Kass (1945) designated alginolytic bacteria from sea water as members of the genus Alginovibrio, and those from soil as associated with the genera

Alginomonas and Alginobacter. This involves many species of Azotobacters and

Pseudomonads; Agarbacterium alginicums (William and Eagon, 1962), Alginovibrio aquatilis (Gacesa, 1992), Alteromonas sp. (Iwamoto, et al., 2001), Azotobacter vinelandii

(Davidson and Lawson, 1977), Bacillus circulans (Hansen, et al., 1984), Dendryphiella salina (Shimokawa, et al., 1997a, b), Enterobacter cloacae (Nibu, et al., 1995),

Flavobacterium multivolum (Takeuchi, et al., 1994), Klebsiella aerogenes (Boyd and

Turvey, 1977), Klebsiella pneumonia (Ostgaard, et al., 1993), Pseudomonas aeruginosa

(Linker and Evans, 1984), and Vibrio sp. (Li, et al., 2003; Chao, et al., 1992a; 1992b;

1992c), Pseudoalteromonas citrea (Alekseeva, et al., 2004), Azotobacter chroococcum

(Haraguchi and kodama, 1995). Alginate degrading enzymes have also been isolated from bacteriophages specific for Azotobacter vinelandii, P. aeruginosa and associated with a Chlorella virus (Bartell, et al., 1966; Davidson, et al., 1977; Suda, et al., 1999).

2.9 SUBSTRATE SPECIFICITIES OF ALGINATE LYASE:

Alginate lyases have been classified on the basis of cleaving preferences for the

M-rich or G-rich alginates blocks, and categorized as

. Poly mannuronic acid (M) lyase (EC 4.2.2.3): [(1-4)--D-mannuronan lyase]

. Poly guluronic acid (G) lyase (EC 4.2.2.11): [(1-4)--L-guluronan lyase].

The specificity of substrate for alginate lyase possibly depends on the environmental conditions in which the alginase-producing organism resides and the availability of types of alginate. Many reported lyases are poly (M) lyase (Boyen, et al.,

1990a; Nisizawa, et al., 1968; Elyakova and Favorov, 1974, Favorov, et al., 1979;

Shiraiwa, et al., 1975; Madgwick, et al., 1973; Sawabe, et al., 1992; Sawabe, et al.,

1998), however few G-specific lyases have also been reported (Sutherland and Keen,

1981; Brown and Preston, 1991; Preston, et al., 1985; Quatrano and Stevens, 1976;

Sawabe, et al., 1992; Sawabe, et al., 1998). Although alginate lyase has been classified as

G or M specific, each lyase usually shows low to moderate cleavage activity for the another type of homopolymer. This may be due to the property of the alginate used as substrates, because of the procedure followed for extracting poly G or poly M blocks which produces substrates rich in one of the homopolymer nevertheless not devoid of the another type completely.

However, some crude alginate lyase preparations have been reported which demonstrate many substrate specificities, depicting either activity of enzyme for multiple substrate or the life-form produces different types of alginate lyase. The preponderant known alginate lyase have endolytic activity which degrades long chains resulting in oligomers (Shiraiwa, et al., 1975; Madgwick, et al., 1973; Madgwick, et al., 1978;

Watanabe and Nisizawa, 1982; Davidson, et al., 1976; Kashiwabara, et al., 1969; Min, et al., 1977a, b, c; Muramatsu and Sogi, 1990). On the other hand a few exolytic alginate lyase have also been reported (Brown and Preston, 1991; Doubet and Quatrano, 1982;

Doubet and Quatrano, 1984; Nakada and Sweeny, 1967; Schaumann and Weide, 1990), that strip off alginate dimers or monomers from the ends of alginate polymer.

2.10 ALGINATE LYASE MECHANISM OF ACTION

Alginase cleaves the glycosidic β-1-4 O-linked bond between the two monomers of alginate polymer via β-elimination. A double bond at C4-C5 carbons of the hexa-ring replaces the 4-O-glycosidic bond, which produces a compound with nonreducing terminal as 4-deoxy-L-erythro-hex-4-enopyranosyluronic acid (Haug, et al., 1967a).

Gacesa (1987, 1992) proposed a three-step catalytic mechanism for alginate lyase to depolymerize alginate. The alginate lyase and epimerase share similar mechanism of action on the alginate polymer, differing only in the last step of depolymerization of alginate.

The steps include:

(a) Elimination or neutralizing the carboxyl anion negative charge with the help of a salt bridge (lysine could be the residue);

(b) A base-catalyzed withdrawal of the proton from C5 (histidine, cysteine, lysine, glutamic acid and aspartic acid could be optional for this task), wherein one of the residue act as the proton benefactor and other as withdrawal, however the protons are obtained from the surrounding solvent.

In the reported epimerase mechanism, the epimerization followed in step (c). The enzyme facilitates epimerization of M-units into G-units in various patterns. The A. vinelandii epimerases have been cloned and expressed for the tailoring of alginates

(Valla, et al., 1996).

G- Lyase G/M -Lyase M -Lyase

G G M M

Enzymatic Hydrolysis

Unsaturated Monomer Unsaturated Monomer Tautomeric conversion

CHO

OH C H OH C H C H C O H

COOH 4- deoxy L-erythro-5- hexoseulose uronicH acid

Figure 2.2: Alginate lyase mechanism of action on alginate polymer 2.11 ALGINATE LYASE ISOLATION AND PURIFICATION

Usually ammonium sulfate precipitation is the first step employed for the downstream process of alginate lyases out of a miscellaneous mixture of proteins

(Kennedy, et al., 1992; Kitamikado, et al., 1992; Lange, et al., 1989; Davidson, et al.,

1976; Cao, et al., 2007; Wang, et al., 2006; Stevens and Levin, 1977; Doubet and

Quatrano, 1984; Baron, et al., 1994; Boyd and Turvey, 1977; Dunne and Buckmire,1985;

Kaiser, et al., 1968; Kraiwattanapong, et al., 1999; Matsubara, et al., 1998; Nakagawa, et al., 1998; Nibu, et al., 1995; Peci˜na and Paneque, 1994). Alternatively, ultrafilteration has also been used for alginate lyase concentration from extracts/ culture supernatant

(Xiao, et al., 2007; Xiaoke, et al., 2006). This is generally followed by separation of alginate lyase on ion exchange chromatography such as cation exchanger to capture alginate lyase with alkaline pI (Davidson, et al., 1976; Cao, et al., 2007; Stevens and

Levin, 1977; Linker and Evans 1984; Lange, et al., 1989; Haugen, et al., 1990;

Matsubara, et al., 1998; Nibu, et al., 1995; Shimokawa, et al., 1997a; Yoon, et al., 2000) or anion exchangers for detain lower pI values alginate lyases from crude enzyme preparations (Nibu, et al., 1995; Matsubara, et al., 1998; Boyd and Turvey, 1977;

Kennedy, et al., 1992; Kraiwattanapong, et al., 1999; Matsubara, et al., 1998; Nakagawa, et al., 1998; Rehm, 1998; Sawabe, et al., 1997; Shimokawa, et al., 1997b). After ion exchange, gel filtration has been employed for many purification protocols (Baron, et al.,

1994; Brown and Preston, 1991; Fujiyama, et al., 1995; Nakagawa, et al., 1998; Nibu, et al., 1995; Shimokawa, et al., 1997a, b).

Affinity chromatography has been also reported as one-step purification of alginate lyase and involves downstream purification using hexa-histidine-tagged protein

(Chavagnat, et al., 1996; Suda, et al., 1999). Alginate-sepharose affinity resin chromatography (Boyd, et al., 1993; Kennedy, et al., 1992) or alginate-epoxy resin affinity chromatography (Eftekhar and Schiller, 1994) and fast protein flow chromatography (Svanem, et al., 1999) have also been used for alginate lyase partial purification.`

Hydroxyapatite has been employed for purification of alginase gene from marine bacterium ATCC 433367 (Malissard, et al., 1995), while hydrophobic interaction chromatography (HIC) has been reported for the K. pneumonia alginate lyase partial purification (Østgaard, et al., 1993) and A. vinelandii alginase cloned genes (Ertesv˚ag, et al., 1998). Both, HIC and hydroxyapatite have been employed to obtain purified recombinant alginase gene from Azotobacter chroococcum (Peci˜na, et al., 1999). E. coli has been used to obtain many purified cloned genes for alginase enzyme (Baron, et al.,

1994; Chavagnat, et al., 1996; Fujiyama, et al., 1995; Kraiwattanapong, et al., 1997;

Malissard, et al., 1995; Peci˜na, et al., 1999; Yoon, et al., 2000) and Bacillus subtilis

(Hisano, et al., 1994b).

2.12 ALGINATE LYASE CHARACTERIZATION

Most of the alginate lyases reported and isolated from marine sources either associated with marine algae or mollusks, the major source for alginate lyase being the marine bacteria that are easy and quick to propagate. Usually the production alginate lyase is stimulated in the environment containing alginate as substrate, as in A. aquatilis

(Stevens and Levin, 1977), marine bacteria A3 and W3 (Doubet and Quatrano, 1984), or

Sargassum associated marine bacteria (Brown and Preston, 1991; Romeo and Preston,

1986); on the other hand in a few bacteria (e.g. P. alginovora), alginase is produced constitutively (Boyen, et al., 1990). The optimal pH for most alginase enzyme reported to be in the range of 7.5 to 8.5, whereas optimal temperature ranged from 25 to 50˚C and molecular weight in the range of 24–110 kDa.

Most of the alginate lyase reported to date are extracellular or periplasmic, having endo-poly (M) lyase activity. reported A marine bacterium demonstrating exolytic poly

(M) lyase activity has been reported (Doubet and Quatrano, 1984). Comparatively a fewer endolytic poly (G) lyase and exo-poly (G) lyase have also been published.

Sargassum fluitans associated marine bacterium strain SFFB 080483, produced a 38-kDa exolytic G-specific lyase (Brown and Preston, 1991).

Some bacterial isolates produce many alginate lyase; for e.g. P. alginovora strain

XO17 shows activity for both poly-G as well as poly-M (Boyen, et al., 1990; Chavagnat, et al., 1996). Alteromonas sp. strain H-4 produces five different extracellular alginase, a number of them have activity for heterogenous substrates (Sawabe, et al., 1992; Sawabe, et al., 1997; Sawabe, et al., 1998). Strain H-4 of Alteromonas sp. also produced 4 types of intracellular alginase enzyme, viz. two poly (G), poly (MG) and poly (M) specific. The extracellular and intracellular alginate lyases were constitutively produced as the substrate alginate presence stimulated the secretion of extracellular alginase although the production of intracellular alginase was not affected considerably. (Sawabe, et al., 1998).

The pIs for several marine bacterial alginate lyase are in the range of 4.3 and 6.7, although a pI of 7.8 was also reported for the Halomonas marina alginase

(Kraiwattanapong, et al., 1999). For numerous marine isolates, the NaCl presence is essential for production and activity of lyase as reported in case of Vibrio harveyi (Tseng,

27 et al., 1992). The divalent cations presence like calcium and magnesium is also obligatory for enzyme activity optima.

Alginate lyase has also been reported from terrestrial bacteria. The G-specific extracellular lyase of E. cloacae depicted optimum activity at 30˚C and pH 7.8 (Nibu, et al., 1995), and it also produces an intracellular alginase with optimal activity at 40˚C with pH 7.5 (Shimokawa, et al., 1997a). Kennedy, et al., (1992) accounted the existence of periplasmic alginate lyase activity in A. chroococcum and A. vinelandii strains.

Additionally, 23 kDa A. chroococcum strain 4A1M extracellular poly (M) lyase, depicted temperature and pH optima of 60˚C and 6.0 respectively (Haraguchi & Kodama 1996).

A. vinelandii reported to have an pH optima ranged from pH 8.1 and 8.4

(Ertesv˚ag, et al., 1998). Both extracellular and intracellular alginate lyases from E. cloacae are accounted to have comparable molecular weight 38–39,000 with same pI 8.9

(Nibu, et al., 1995; Shimokawa, et al., 1997b). Most of the alginase from gram-negative soil bacteria are reported to have basic pI values with the exception of M-specific lyases of A. vinelandii (pI-5.1) and A. chroococcum (pI-5.6) (Ertesv˚ag, et al., 1998; Haraguchi and Kodama, 1996).

The G-specific lyase produced extracellularly (Nibu, et al., 1995) and the M- specific intracellular alginate lyase (Shimokawa, et al., 1997b) from E. cloacae demonstrated similar properties as both are inhibited completely by EDTA and need 2 mM calcium ions for reinstatement of utmost activity. On the other hand, A. vinelandii alginate lyase exhibit optimal activity in 0.35 M NaCl presence although the divalent cations were not required (Ertesv˚ag, et al., 1998). Alginase activity specific to poly (M) from A. chroococcum was enhanced by the presence of divalent cations like calcium

28 whereas strongly inhibited in the presence of mercury ions (Haraguchi and Kodama,

1996).

Kinoshita, et al., (1991) reported two extracellular and intracellular alginate lyases from Pseudomonas OS-ALG-9. The molecular mass of 45 kDa intracellular alginase was observed with a optimum activity at 45˚C with pH 7.5 (Kinoshita, et al.,

1991).

A Sphingomonas sp. strain was reported to produce 3 types of alginase: ALY1-I

(60 kDa), ALY1-III (38 kDa) and ALY1-II (25 kDa) (Murata, et al., 1993) which are programmed by singel gene resulting in a 69 kDa peptide (Hisano, et al., 1994b).

However they differ in their pIs as ALY1-I has 9.03, ALY1-III has 10.16 and ALY1-II has 6.82, but other characteristics were reported to be common for these three enzymes.

They are cytoplasmic, and endolytic; with pH optima of 7.5–8.5, and temperature optima of 70˚C (Hashimoto, et al., 1998; Hisano, et al., 1993; Yonemoto, et al., 1991;

Yonemoto, et al., 1993a).

Nakagawa et al (1998) isolated an 40 kDa extracellular alginate lyase from

Bacillus sp. strain ATB-1015 demonstrating activity specific for both poly(G) and poly(M) effective against strains of P. aeruginosa present in biofilms (Nakagawa, et al.,

1998). Hansen, et al., (1984) isolated 40 kDa endo-poly (M) lyase from Bacillus circulans strain JBH2 by providing alginate as the only energy and carbon source.

Wicker-B¨ockelmann, et al., (1987) isolated another 58 kDa lyase from Bacillus circulans strain JBH2 with similar optimal pH of 5.8.

Larsen, et al., (1993) isolated a weak nonspecific poly (G) and poly (M) alginate lyase of 34 kDa from strain 1351 of B. circulans although the poly (M) lyase activity was enhanced in 2 mM Ca2+ presence. The enhancement of lyase activity in the divalent cations presence, particularly Mg2+ or Ca2+, have also been reported for some gram- positive bacteria (Hansen, et al., 1984; Kaiser, et al., 1968; Larsen, et al., 1993;

Matsubara, et al., 1998; Nakagawa, et al., 1998). Although these divalent cations are not essentially required, their presence often enhances the lyase activity. The alginate lyase activity from Clostridium alginolyticum (Kaiser, et al., 1968), got doubled in 0.02 M

NaCl presence, acquiring utmost activity with 0.25 M whereas 2.0 M NaCl inhibited the lyase activity.

Although alginase activity have also been reported from several marine algae and invertebrates such as marine mollusks only a few lyases have been extensively studied.

Although all animals generate single alginate lyase, T. cornutus produces two isozymes of alginate lyase (Muramatsu and Egawa, 1980), whereas Haliotis sp. produce two different lyases (Nakada and Sweeny, 1967). Most of the endo-poly (M) lyases have pH optima in the range of 5.6 to 9.6. Nakada & Sweeney (1967) explained an abalone hepatopancreas exo-poly (G) lyase with pH optima of 4.0. Both alginate lyase from

Haliotis sp. shows a requirement of 0.05 to 0.075 M NaCl. Nakada & Sweeney (1967) hypothesized that this huge amount of ionic strength either disrupt bound water molecules surrounding the alginate or maintains uronic acid units at a nominal inter-unit distance for appropriate fit. The requirement of high NaCl concentrations (0.1–0.2 M) is also reported for Spisula solidissima, Haliotis tuberculata and T. cornutus lyase activity

(Boyen, et al., 1990; Jacober, et al., 1980; Muramatsu, et al., 1977). H. tuberculata

30 alginase shows a fondness for G-M and M-M linkages (Heyraud, et al., 1996), while T. cornutus lyase have a preference only for M-M (Muramatsu, et al., 1993).

Calcium ions were also reported for increasing the lyases activity from Undaria pinnatifida and Pelvetia canaliculata ( Watanabe and Nisizawa, 1982; Madgwick, et al.,

1978). Shiraiwa et al., (1975) reported seasonal variation in alginate lyase activity isolated from a several algae. Most of the alginate lyases isolated from marine algae are bifuntional i.e specific for both poly (M) as well as poly (G).

Some species of marine fungi associated with decaying seaweed, such as

Asteromyces cruciatus, Corollospora intermedia and two species of Dendryphiella, have also been reported for the production of alginate lyases (Schaumann and Weide, 1990;

Shimokawa, et al., 1997a, b; Wainwright, 1980; Wainwright and Sherbrock-Cox, 1981).

For many fungi the production of alginase was induced by growing them in the pressence of alginate. These alginate lyase produced are typically extracellular; however the alginase is mostly bound to the cells in Asteromyces sp (Schaumann and Weide,

1990).Schaumann & Weide (1990) reported alginate hydrolase and lyase presence in A. cruciatus. Research on Dendryphiella salina demonstrate a 35 kDa alginase with a pI of

3.65 and optimum pH between 5 and 6 at 45ºC which shows endolytic cleaving of poly

(M) substrates (Shimokawa, et al., 1997b). Enzyme activity was stimulated in 1%–3%

NaCl presence.

Few bacteriophages have been also reported for alginase production which are specific for Pseudomonas and Azotobacter sp (Barker, et.al., 1968; Bartell, et.al., 1966;

Davidson, et.al., 1977; Eklund and Wyss, 1962; Pike and Wyss, 1975) that facilitate the penetration of the phage via exopolysaccharides rich in acetylated poly (M). These alginase shows endolytic activity with 30 - 42 kDa molecular masses and optimal pH of 7.5 to 8.5.

An alginase gene has also recently been sequenced from a Chlorella virus. (Suda, et al., 1999). The 39-kDa mannuronate lyase had a optimum pH of 10.5, and required

Ca2+for activity (Suda, et al., 1999).

2.13.1.1 APPLICATION OF ALGINATE LYASE

Alginate lyase from various sources and different substrate specifities have been extensively used to engineer alginate polymer for application of the resulted alginate oligosaccharides in different field of biotechnology based industries such as agriculture, aquaculture, medicine, textiles, Food and cosmetics.

The co-administration of alginate lyase with various antibiotics play a crucial task for treatment of cystic fibrosis (CF) caused by Pseudomonas aeruginosa (Islan, et al.,

2013; Alkawash, et al., 2006; Hatch and Schiller, 1998). Tajima. et al., (1999) had also demonstrated the alginate oligosaccharides effects which are produced by the action of alginate lyase, for expression of collagen and cell proliferation in cultured skin fibroblasts. The alginate oligosaccharides were observed to suppress proliferation of fibroblast to half compared to control cultures accompanied by a change in shape of the cell. Additionaly alginate oligosaccharides treatment of confluent cells resulted in a reduction in synthesis of collagen and hence provides a effective tool for the medication

32 of abnormal collagen metabolism disorders. The alginate oligosaccharides treatment also reported to protect neuron-like PC12 cells against oxidative stress induced mitochondrial and endoplasmic reticulum (ER) dependent apoptotic cell death by promoting expression of Bcl-2, and blocking expression of Bax while inhibiting the activation of H2O2 induced caspase-3 (Tusi, et al., 2011).

Further the alginate oligosaccharides neuro-protective potential against Aβ- induced neural damage was also reported (Tusi, et al., 2011). Akiyama, et al., (1992) had reported utilizing alginate oligosaccharides prepared by alginate lyase isolated from bacterium strain A2 (Yonemoto, et al., 1991), for acceleration of the growth of

Bifidobacterium as intestinal flora.

A mixture of alginate oligosaccharides with guluronic acid at the reducing end cleaved from alginate, have been reported to encourage migration of human endothelial cells and VEGF-mediated growth which was comparable to activity of heparin (Kawada, et al., 1999). A mixture of alginate oligosaccharides with more G blocks in the reduced terminus, demonstrated stimulation of uptake of (3H) thymidine and human keratinocyte growth, in the epidermal growth factor (EGF) presence. Alginate oligosaccharides reported to be used as a substitute for bovine pituitary extract in keratinocyte cultures

(Kawada, et al., 1997).

Whereas oligosaccharides of alginate with lower G blocks percentage shows comparatively more potent for inducing production of cytokine. The study demonstrates that the residues of mannuronic acid proves to be active inducers for cytokine (Otterlei, et al., 1991; Iwamoto, et al., 2003).

Ariyo, et al., (1997) reported about 50% enhanced yield of penicillin G from P. chrysogenum P2 cultural biomass (high penicillin producer) and 150% yield in P. chrysogenum NRRL 1951 cultures (low penicillin producer), when compared with the control cultures without the treatment of oligosaccharides.

Sodium alginate oligosaccharides especially trisaccharides prepared using alginate lyase from Alteromonas macleodii demonstrated a growth promoting effect on the barley roots mainly that on radical (Tomoda, et al., 1994). Treatment with the alginate oligosaccharides have been reported to increase the alcohol dehydrogenase activity under hypoxic condition promoting certain resistance to the hypoxic stress or initiating certain signal-transduction pathways (Farmer, et al., 1991). Xu, et al., (2003) reported the root elongation activity of the bacterial alginate lyase digestion mixtures of poly guluronic acid on carrot and rice plants. Similar results were demonstrated by Iwasaki and

Matsubara, (2000) for root growth of lettuce seedlings. In another study promoting effect of alginate oligosaccharides on germination and shoot elongation has been reported

(Yonemoto, et al., 1993b).

There have been many reports for the use of mixture of the polysaccharide degrading enzyme such as mixture of alginate lyase and cellulase for isolation of brown algae protoplasts (Saga, 1984; Saga and Sakai, 1984; Saga, et al., 1986; Polne-Fuller and

Gibor, 1987; Kloareg and Quatrano, 1987a,b; Fisher and Gibor, 1987; Tokuda and

Kawashima, 1988; Ducreux and Kloareg, 1988; Kajiwara, et al., 1988; Kloareg, et al.

1989; Butler, et al., 1989; Sawabe, et al., 1993; Matsumura, et al.2000; Wakabayashi, et al., 1999). As alginate and cellulose are major components of Laminaria cell wall. Inoue, et al., (2011) had demonstrated the recombinant abalone alginate lyase (rHdAly) use

34 along with cellulose and protease K for viable protoplast isolation from mature sporophytes of Laminaria japonica blades. Cultured young and small thalli which are easy to degrade, of Laminaria japonica had also been studied for protoplast preparation

(Sawabe and Ezura, 1996; Matsumura, et al., 2000).

Depolymerized fractions of alginate obtained by alginate lyase (Laboratory of

Applied Microbiology, Ocean University of China) exhibited an inhibitory effect for pathogenic strains from marine such as V.Pelagius, V. ﬂuvialis,Vibrio harveyi,

V. vulniﬁcus, and V. alginolyticus which promotes the use of the depolymerized products of alginate in aquaculture farms (Xiaoke, et al., 2005)

CHAPTER 3: SCREENING FOR MULTIPLE POLYSACCHARIDE DEGRADING BACTERIA

This chapter elucidates and rationalizes the strategies undertaken to screen and isolate polysaccharide-degrading bacteria from marine eco-niches. Subsequently some of these polysaccharide degrading bacteria were reported to decompose more than one polysaccharide. As mentioned in ‘Research goal and significance’ in the present study, alginolytic bacteria were specifically screened with an objective to characterize the alginate lyase for various biotechnological applications.

In the following chapter, marine eco-niches such as sandy/ rocky coastal water/ sediments, seaweeds and mangroves were screened for isolation of alginolytic bacteria.

Different levels of screening (primary, secondary and tertiary) were followed with an objective to isolate a novel polysaccharide degrading bacteria. The results obtained following the screening strategies are also discussed in comparison to on polysaccharide degrading bacteria screened by various research groups.

MATERIALS:

Agar (bacteriological, purified), alginic acid (mixture of guluronic and mannuronic acid), cetylpyridinum chloride, Tris base, magnesium chloride, potassium chloride, di- ammonium hydrogen phosphate, sodium chloride, calcium chloride were purchased from

HiMedia labs. All other chemicals and reagents used in experiments were of analytical/ clinical grade.

3.1 SAMPLE COLLECTION:

Samples for screening polysaccharide degrading bacteria were collected from coast of Goa, mangrove and seaweeds. Coastal water samples along with sediments were collected in a sterile 250 ml polypropelene bottles from the coast of Goa (namely Siridao,

Bambolim, Cacra, Caranzalem, Calangute, Vagator, Anjuna) and Malvan (from

Maharashtra) in the month of November/ December 2007. Brown seaweeds such as

Sargassum tenerrimum and Fucus distichus were collected from the intertidal region of

Cacra and Anjuna coast, in November/ December 2007 in sterile disposable plastic bags.

The water sample from the mangroves of Divar Island (Goa) was collected in January

2008. All water/sediment samples were used within 24 h and subjected to primary screening for isolation of alginolytic bacteria. The geographical locations of sampling sites were determined using GPS 12 (Germin Inc., Kansas City, USA). Table 3.1 and Fig

3.1 depict the location of various sampling sites.

Table 3.1: Geographical locations of the sampling sites.

Sampling sites Niches Locations

Siridao Sandy Coastal water 15˚26’39.37”N 73˚51’20.79”E

Bambolim Coastal water 15˚26’49.76”N 73˚51’12.25”E

Caranzalem Coastal water 15˚’27’58.41”N 73˚48’16.88”E

Calangute Coastal water 15˚32’54.51”N 73˚45’12.15”E

Vagator Coastal water 15˚35’43.80”N 73˚44’00.66”E

Anjuna Coastal water 15˚34’33.77”N 73˚44’24.08”E

Cacra Coastal water 15˚27’04.52”N 73˚50’11.33”E

Malvan Coastal water 15˚34’33.77”N 73˚44’24.08”E

Divar Mangrove 15˚34’33.77”N 73˚44’24.08”E

Figure 3.1: Map of Goa showing the sampling site locations.

3.2 SEAWEED DECOMPOSITION:

Brown seaweeds such as Sargassum tenerrimum and Fucus distichus were collected from the intertidal region of Cacra and Anjuna, the coastal of Goa, in

November/ December 2007. Sea water sample was obtained from the vicinity of the algal sampling site was added to the seaweeds samples that were kept for decomposing. The seaweed samples were incubated at 30ºC for 45 days in a polypropylene container and stirred/ mixed occasionally.

3.3 MEDIUM USED FOR SCREENING AND ISOLATION OF

POLYSACCHARIDE DEGRADING BACTERIA:

Artificial sea water (ASW) was routinely used for isolation of polysaccharide degrading bacteria (Appendix I). Bacteriological agar (1.8%) was added for the preparation of ASW agar plates.

3.4 PRIMARY SCREENING

During primary screening, ASW agar medium co-supplemented with 1% alginate was used for isolation of alginolytic bacterial isolates. The plates were kept at 30 ± 2˚C.

Direct plating and enrichment technique were used for screening of alginolytic bacterial isolates. ASW medium broth containing alginate at a final concentration of

1.0% as a sole carbon source was used for enrichment culture technique.

3.4.1 DIRECT PLATE TECHNIQUE:

Water samples (collected from coast/ decomposing seaweeds/ mangroves) was serially diluted and directly spread plated on ASW agar plates containing 1.0% alginate, and incubated at 30ºC for 96 h.

3.4.2 ENRICHMENT TECHNIQUE:

1 ml of water sample (collected from coast/ decomposing seaweeds/ mangroves) was inoculated in 25 ml of ASW medium supplemented with 0.2% alginate as only carbon source and incubated at room temperature for 24 hrs on orbital shaker (130 rpm).1 ml of the inoculum from the first enrichment was added to 25 ml of ASW broth containing 0.2% alginate and incubated for 24 h on orbital shaker. The enriched culture was serially diluted and cultured on ASW agar plates containing 1% alginate and incubated at 30ºC for 96 h.

3.4.3 PLATE SCREENING METHOD FOR DETECTING DEGRADATION OF

ALGINATE BY ALGINOLYTIC BACTERIA:

In order to visualize and detect the bacterial isolate with alginolytic activity, the

ASW agar- alginate plates were flooded with 10% cetylpyridinium chloride. Clearance zone around the bacterial colonies indicated alginolytic activity. (Gacesa and Wusteman,

1990).

3.5 SECONDARY SCREENING:

The bacterial isolates obtained during primary screening were subjected to secondary screening to evaluate the potential of the bacterial isolates to degrade other polysaccharides such as carrageenan, xylan, carboxy methyl cellulose (CMC) and agar.

The ASW broth supplemented with 0.2% of any one polysaccharide such as alginate/

CMC/ xylan/ carrageenan was inoculated with the individual bacterial isolates obtained from primary screening. Further to check for agar degradation the bacterial isolates were also grown on ASW agar plates. A flask containing ASW with one of the above polysaccharide without any bacterial inoculums was used as control.

The ASW broth (containing either alginate/ CMC/ xylan/ carrageenan) and plates

(containing only agar) were inoculated with the bacterial isolates obtained from primary screening and incubated at 30 ºC for 36 h. The growth of bacteria due to polysaccharide utilization from the broth was determined spectrophotometerically (Shimadzu Co. Kyoto,

Japan) at OD600. Degradation of the agar was confirmed on ASW agar plates by adding

Lugol’s iodine which resulted in clearance zones around the agarolytic positive colonies

(Hodgson and Chater 1981).

3.6 TERTIARY SCREENING:

Bacterial isolates chosen from secondary screening were further subjected to tertiary screening. Tertiary screening was based on capability of bacterial isolates to degrade seaweed thalli under in-vivo condition. The brown seaweeds Sargassum tenerrimum and Fucus distichus previously collected during sampling in the month of

November/ December 2007 from Cacra and Anjuna coast of Goa, were washed systematically with filtered sea water to eliminate epiphytes and later air-dried

The semi-dried thalli of Sargassum tenerrimum and Fucus distichus were surface sterilized with 80% ethanol for 8 min and washed with sterile sea water. Sterilized pieces of Sargassum tenerrimum and Fucus distichus were added to sterile ASW medium without any carbon substrate. Later, the above medium was inoculated with the individual bacterial isolate obtained during secondary screening and incubated for 48 h at room temperature (30˚C) on orbital shaker. Growth of bacterial cells was measured spectrophotometerically at A600 (Shimadzu Co. Kyoto, Japan). Additionally, the effect of bacterial growth on thalli was observed under light microscope.

3.7 RESULTS:

3.7.1 PRIMARY SCREENING

During primary screening, alginolytic bacteria from various niches were isolated.

Direct plating and enrichment method were employed for isolating alginolytic bacteria.

The screening of alginolytic bacteria were done on ASW agar plates supplemented with

1% alginate. Almost all bacteria depicted the formation of pits (depressions) or clearance zones. Based on the visual differences in the morphology of the colonies, the bacterial isolates were individually purified and sub-cultured.

A total of 200 bacterial isolates (from three types of niches) were obtained on

ASW agar alginate plates during primary screening. Out of 200 bacterial colonies, 151 bacterial isolates were from coastal areas, 25 bacterial isolates were obtained from mangrove whereas 24 bacterial isolates were isolated from decomposing seaweeds. Fig.

3.2 depicts the distribution of polysaccharide degrading bacteria from various niches.

Fig. 3.3 represents the distribution profile of bacterial isolates obtained by direct plating and enrichment technique. As observed from the figure the numbers of bacterial isolates obtained by direct plating (66%) were greater than the number of isolates obtained by enrichment technique which resulted in only 34% of total number of isolates.

Figure 3.2: Distribution of bacterial isolates obtained from primary screening.

Figure 3.3: Bacterial isolates obtained from direct plating and enrichment technique.

In order to differentiate the alginolytic isolates from agarolytic, plate based screening method was used. The replication of 200 bacterial colonies was done on ASW agar and ASW agar plates containing alginate. Observation of clearance zone around the bacterial colonies when Lugol’s iodine was added to ASW agar plates, indicated agrolytic activity alternatively, alginolytic activity was observed by visualization of clearance zone around the colonies on adding cetylpyridinium chloride (CPC) to ASW agar- alginate plates (Fig. 3.4). Lugol’s iodine and CPC used for detection of the agarolytic and alginolytic activity respectively were polysaccharide specific and no cross reactivity was observed.

Out of 200 bacterial isolates that were obtained during primary screening, only 50 bacterial isolates were observed to be alginolytic based on plate assay (Table 3.2). The remaining 150 isolates demonstrated only agarolytic activity (Fig.3.5). Compared to the variation observed in terms of occurrence of agarolytic bacteria in various niches/ sampling sites, no significant distribution was observed for occurrence of alginolytic bacteria (Fig. 3.5).

Figure. 3.4: The clearance zone shown by various alginolytic bacterial isolates.

Agarolytic 75%

Figure 3.5: Distribution of the alginolytic bacterial isolates from the sampling sites

Table 3.2: Alginolytic bacteria isolated from various niches on ASW agar- alginate plates during primary screening.

Isolates of alginolytic bacteria Sampling obtained by Niches sites Direct Enrichment plating technique SD4 SD6 Siridao Coastal water Nil SD8 SD9 BE-1 Bambolim Coastal water BD-1 BE-2 CD-3 CD-6 CE-1 CD-7 Cacra Coastal water CE-3 CD-10 CE-5 CD-11 CD-12 ZE-1 ZE-3 ZD-2 Caranzalem Coastal water ZE-4 ZD-7 ZE-5 ZE-8 Calangute Coastal water LD-7 LE-1 VD-3 Vagator Coastal water VE-5 VD-5 JD-7 JE-2 Anjuna Coastal water JD-8 JE-3 JD-12 JE-4 DD-13 DE-3 Divar Mangrove DD-14 DE-4 ND-2 ND-3 NE-4 Malvan Coastal water ND-4 NE-5 ND-8 Epiphytic AD-1 Decomposing bacteria from AD-2 AE-1 seaweed decaying AD-3 AE-8 seaweeds AD-4

Thus out of 50 bacterial isolates (out of 200 isolates) obtained during primary screening , 25 bacterial isolates were screened from mangrove samples out of which only

04 were alginolytic either by direct plating/ enrichment method. Whereas 24 bacterial isolates were sreened from the decomposing seaweed out of which only 06 were confirmed to be alginolytic isolates. Majority of the bacterial colonies were small, entire and white in colour. Additionally, only 19 bacterial isolates were obtained from sandy shore coastal niches (Siridao, Bambolim, Caranzalem, Calangute and Vagator) wherein

21 bacterial isolates were acquired from rocky coastal niches (Anjuna, Cacra and

Malvan)

3.7.2 SECONDARY SCREENING:

The secondary screening was employed to assess the multiple polysaccharide degrading characteristic of the 50 alginolytic bacterial isolates selected from the primary screening experiments.

The alginolytic bacteria obtained were further checked for degradation of other polysaccharides like agar, carrageenan, CMC and xylan. Degradation of agar was determined using Lugol’s iodine by plate based assay. All alginolytic bacteria (50 bacterial isolates) were observed to be agarolytic as they depicted clearance zone on flooding with Lugol’s iodine. In contrast the agarolytic bacterial isolates (150 isolates) did not demonstrate alginolytic activity. Bacterial isolates such as SD-6, BD-1, ZE-8,

VD-3, JD-8, JD-12, AE-1 and AE-8 were exceptional as they depicts pits/craters on

ASW agar plates with clearance zone observed after spreading Lugol’s iodine. The other bacterial isolates did not show any craters/ depression in the agar.

Out of 50 alginolytic bacterial isolates from primary screening, 22 bacterial isolates were found to degrade more than one polysaccharides (alginate, xylan, carageenan and CMC) (Table 3.3).

Table 3.3: Growth of alginolytic bacterial isolates on other polysaccharides. ((± weak :

OD 600 < 0.05; + good growth: OD 600 0.05- 0.2; ++ excellent growth : OD 600 > 0.2).

Bacterial Alginate Xylan Carrageenan CMC isolates SD-4 ++ - ± ± *SD-6 ++ - ± ± SD-8 ++ - + ± SD-9 ++ + ± ± *BD-1 ++ + ± ± BE-1 ++ ± ± ± BE-2 ++ - + ± CD-3 ++ ± ± ± CD-6 ++ - - ± CD-7 ++ ± ± ± CD-10 ++ ± ± ± CD-11 ++ ± ± ± CD-12 ++ - ± ± CE-1 ++ ± ± + CE-3 ++ ± ± ± CE-5 ++ - + ± ZD-2 ++ ± ± + ZD-7 ++ - ± - ZE-1 ++ - ± ± ZE-3 ++ - ± ± ZE-4 ++ - ± + ZE-5 ++ ± ± - *ZE-8 ++ ± ± ± LD-7 ++ ± ± ± LE-1 ++ ++ + ± *VD-3 ++ - + ± VD-5 ++ - ± ± VE-5 ++ - - + JD-7 ++ + ± + *JD-8 ++ - ± - *JD-12 ++ ± ± + JE-2 ++ - ++ -

Bacterial Alginate Xylan Carrageenan CMC isolates JE-3 ++ - ± - JE-4 ++ - ± ± DD-13 ++ + ± + DD-14 ++ ± + ± DE-3 ++ - ± ± DE-4 ++ + ++ ± ND-2 ++ - ± - ND-3 ++ - - - ND-4 ± - - - ND-8 ++ - ± ± NE-4 ++ + ± ± NE-5 ++ ++ ± + AD-1 ++ + ± ± AD-2 ++ + ± ± AD-3 ++ + ± - AD-4 ++ - ± - *AE-1 ++ ++ ± - *AE-8 ++ - + -

For all the five tested polysaccharide, 44% of alginolytic bacterial isolates (50 isolates) demonstrated multiple polysaccharide degradation (Fig. 3.6). Maximum number of multiple polysaccharide degrading bacterial isolates were predominantely isolated from one of the coastal sites i.e. Cacra. Thus a total of 22 multiple polysaccharide degrading bacterial isolates were obtained during secondary screening and were further evaluated for tertiary screening.

Figure 3.6: Distribution of multiple polysaccharide degrading bacterial isolates.

3.7.3 TERTIARY SCREENING:

The alginolytic bacteria with multiple polysaccharide degrading activity was isolated during secondary screening were tested for the algal decomposing activity using

Sargassum tenerrimum and Fucus distichus thalli as algal source.

The presence of Sargassum tenerrimum thalli in ASW broth (no other added carbon source) supported the growth of all alginolytic strains except for isolates ZD-2 and NE-5

(Table 4). The thalli of Fucus distichus did not sufficiently support the growth of the alginolytic bacterial isolates. As seen from the Table 3.4, all the bacterial isolates obtained from Bambolim, Calangute, Malvan and decomposing seaweeds did not depict any growth/ degradation with Fucus distichus.

Table 4: In-vitro brown seaweed thalli decomposition by the selected bacterial isolates from secondary screening after 48 h (± weak : OD 600 < 0.05; + : OD 600 0.05- 0.1; ++ :

OD 600 0.1- 0.2; +++ : OD 600 > 0.2)

Sargassum tenerrimum ASW Fucus distichus ASW Bacterial isolates broth broth

SD-9 + + BD-1 ++ - BE-1 + - CD-3 ++ ± CD-7 ++ ± CD-10 ++ ± CD-11 ++ ± CE-1 ++ - CE-3 ++ ± ZD-2 - ± ZE-8 ++ + LD-7 ± - LE-1 ± - JD-7 ++ - JD-12 + + DD-13 +++ + DD-14 ++ + DE-4 ++ + NE-4 ++ - NE-5 - - AD-1 ± - AD-2 ++ -

3.7.4 MICROSCOPIC OBSERVATION OF DEGRADATION OF Sargassum tenerrimum

Out of 22 multiple degrading bacterial isolates, only bacterial isolate DD-13 was found to decompose and demonstrate growth when either of seaweed thalli were included in the culture medium. Fig. 3.7 depicts degradation profile of S. tenerrimum thalli by strain DD-13. Microscopic observation of the seaweed thalli indicated degradation of seaweed thalli (Fig 3.7)

A) B)

C) D)

Figure 3.7: Seaweed thalli degradation on inoculation with alginolytic bacterial strain

DD-13 (B, C and D); control (A).

Thus on the basis of primary, secondary and tertiary screening, bacterial isolate

DD- 13 was observed to degrade multiple polysaccharides and demonstrated the capability to degrade brown seaweeds by in-vitro experiments degradation. Hence bacterial isolate DD-13 was selected for further identification and characterization.

3.8 DISCUSSION:

In the marine environment polysaccharides are significant resource of organic carbon and their degradation plays major role for recycling of carbon. Several seaweeds, microbes, phytoplankton are natural resource for complex polysaccharides such as agar, chitin, xylan, carrageen, laminarin, mannan and alginate. Therefore bacterial degradation of polysaccharide is an important activity in marine ecosystem which can be performed by varous genera of bacteria (Salyers, et al., 1996).

It is a challenging task for screening of culturable polysaccharide decomposing bacteria from various niches of marine ecosystem. Polysaccharide degrading bacteria have been screened from various ecological niches such as sediment (Rees, et al., 1976), seaweeds (Wang, et al., 2006; Vera, et al., 1998; Ensor, et al., 1999), tarballs

(Naganuma, et al., 1994), soil (Ruiz, et al., 2005; Meskiene, et al., 2003), rivers (Van Der

Meulen and Harder 1976), deep sea (Miyazaki, et al., 2008; Yoon and Lee, 2012), sand

(Park, et al., 2013) and guts of animals (Yi and Shin, 2006).

Thus polysaccharide degrading bacteria are ubiquitous. The presence of polysaccharides in various organisms serves as a selection pressure leading to the dominance of polysaccharide degrading bacteria (Wang, et al., 2006; Vera, et al., 1998;

Enso,r et al., 1999) whereas in certain cases polysaccharide deomposing bacteria have been screen from niches where polysaccharides occurance have not been observed (Van

Der Meulen and Harder 1976; Naganuma, et al., 1994). Certain niches like fresh water river or tarballs etc. do not report polysaccharide degrading bacteria. Polysaccharide degrading bacteria have been isolated possibly because of seeding of these niches with polysaccharides by other sources

In the present study, the main objective was to isolate a novel alginolytic bacterium from marine niches with a potential to decompose/degrade brown seaweeds.

Furthermore the alginolytic bacteria and the alginate lyase would be evaluated for some of the applications related to Biotechnology.

Three levels of screening were engaged to screen and isolate the alginolytic bacteria. The primary screening involved the use of ASW agar alginate plates to isolate alginolytic isolates. The bacterial isolates obtained during primary screening method were further evaluated by secondary screening for their capability to degrade multiple polysaccharides. The bacterial isolates selected from the secondary screening were further tested for their capability to degrade brown seaweeds.

The direct plating method had been reported to be more sensitive and reliable for screening of selective bacterial population with required characteristic (Rennels, et al.,

1980; Gangarosa, et al., 1968; Ahuja, et al., 1951; Feeley, 1962; Sack and Barua; 1964).

Monfort, et al., (1988) compared direct plating with thioglycolate broth enrichment method for isolating Campylobacter jejuni from feces. They reported 1.4 fold increase in

55 the number of isolates obtained by direct plating method in comparison to enrichment method.

Direct plating of the collected samples results in the isolation of indigenous bacterial isolates that is natively present in the respective niches in contrast enrichment technique exhibits the isolation of dominant fast growing bacterial isolates over other slow growing subvarients (Jannasch, 1967). Thus enrichment leads to dilution or loss of native population of bacterial isolates as compared to the number of isolates obtained by direct plating. Hence direct plating and enrichment technique were employed for screening of alginolytic bacteria. Indeed the number of bacterial isolates obtained from primary screening by direct plating always surpassed the number of isolates obtained by enrichment method.

The bacterial isolates were initially differentiated and screened primarily on the basis of different colony morphologies such as size, colour, elevations, margins and texture. Some bacterial colonies depicted clearance zone and/or pits around them on the agar-alginate plates. The bacterial isolates obtained were either agarolytic and/ or alginolytic as they grew on alginate- agar plates. Although all the bacterial isolates that were confirmed as agarolytic during secondary screening were not necessarily alginolytic. However all bacterial isolates that were confirmed as alginolytic during secondary screening were observed to be agarolytic. Dye based screening method involving Lugol’s iodine and cetylpyridinium chloride were used to differentiate the agarolytic and alginolytic activities respectively (Gacesa and Wusteman, 1990; Hodgson

56 and Chater 1981). The dye based plate assay is based on the non- covalent interaction between polysaccharide and dye.

Eight coastal sites were screened for isolation of polysaccharides degrading bacteria on ASW agar- alginate plates. Coastal sites such as Cacra, Anjuna and Malvan have rocky shores with abundant growth of seaweeds such as Sargassum and Fucus species. Thus as seen from Table 3.2, more polysaccharide degrading bacterial isolates were obtained from rocky shores in comparison to other sandy shores (the other five niches) that do not report occurrence of seaweeds.

Out of 200 polysaccharide degrading bacterial isolates obtained on agar-alginate plates, 50 isolates were alginolytic whereas predominant (150 isolates) were agarolytic.

The agarolytic bacterial isolates degraded only agar whereas all the alginolytic bacterial isolates could degrade agar also. This could be attributed to the use of agar-alginate plates used which might have acted as selective medium for favoring the growth of agarolytic bacteria. ASW medium containing alginate alone could not be used as alginate was not a suitable solidifying agent for preparation of solid medium plates. Further, the use of gelrite was restricted as presence of divalent cations in ASW, prevented its use as a gelling agent in the present study.

In the present study, out of total alginolytic bacteria (50 isolates) screened for multiple polysaccharide degradation, only 44% (22 bacterial isolates) were observed to degrade multiple polysaccharides (all the five tested polysaccharides). Most of the multiple polysaccharide degraders were from seaweed associated coastal areas (Anjuna,

Cacra and Malvan).

In comparison to 16 polysaccharide degrading bacterial isolates were obtained on average from five sampling sites (sandy shore) in comparison to total of 22 bacterial isolates from rocky shores, 25 isolates from mangrove and 24 isolates from decomposing seaweeds. The econiches such as mangrove and decomposing seaweeds predominantly contained ICPs either in association with marine organisms or as a part of detritus.

Additionally, the seaweed decomposing experiment served as an enrichment technique for favoring the growth and enrichment of robust polysaccharide degrading bacteria.

Strain DD-13 obtained from mangrove was selected for further studies on the basis of multiple polysaccharide degrading ability and its ability to degrade Sargassum tenerrimum and Fucus distichus. Mangrove ecosystem acts as a bridge between marine and terrestrial ecosystems being rich in organic matter due to the composed vegetative remains and mangrove foliage. Besides this, the litter of vegetative remains as well as mangrove contributes to the increment of complex polysaccharides (CPs) such as pullulan, xylan, cellulose, pectin etc. The increment of CP decomposing micro-organisms population within this ecosystem to prepare detritus from litter is a significant ecological process which is very important for recycling of carbon.

Multiple polysaccharide degrading bacteria such as Saccharophagus degradans strain 2-40T degrading ten different polysaccharides chitin, cellulose, agar, fucoidan, alginate, laminarin, pullulan, pectin, xylan and starch (Ensor, et al., 1999; Howard, et al.,

2003; Andrykovich and Marx, 1988; Ivanova, et al., 2001; Romanenko, et al., 2003) was screened from decaying salt marsh cord grass. The principal cell wall components as polysaccharides in red seaweeds are agarans and carrageenans , those in green seaweeds

58 are ulvans and in brown seaweeds fucans and alginates as well as laminarin as storage polysaccharide (Jiao, et al., 2011; Rioux, et al., 2007). Therefore complex bacterial communities have been found to be associated with the seaweeds and serve as niches for polysaccharide degrading bacteria. Bacteria belonging to different genera such as

Sulfitobacter, Vibrio, Halomonas, Cinetobacter, Pseudomonas, Cytophaga, Planococcus, etc., exist in abundance on the surface of seaweeds (Nakanishi, et al., 1999; Matsuo, et al., 2005; Marshall, et al., 2006).

CHAPTER 4: IDENTIFICATION OF MULTIPLE POLYSACCHARIDE DEGRADING

BACTERIAL STRAIN DD-13

The following chapter illustrates the polyphasic strategies followed to complete the second and third objective defined in the „Research goal and significance‟.

As mentioned in previous chapter several bacterial isolates were screened from various niches of marine ecosystem. Alginolytic bacterial strain DD-13 isolated from sediments of mangrove located at Divar Island (15°30‟35”N and 73°52‟63‟‟E), Goa,

India was also observed to degrade multiple polysaccharides. Compared to all other bacterial isolates, strain DD-13 was observed to degrade eleven polysaccharides. Thus strain DD-13 was contemplated as a novel strain and hence attempts were made to identify the bacterial strain DD-13 up-to species level, polyphasic approaches involving phenotypic, genotypic and chemotaxonomic methods were executed that supported the designation of the strain as a novel species of Microbulbifer.

The steps and methodologies followed for this study and results procured therein are explained in this chapter.

MATERIALS:

Agar (bacteriological, purified), Simmon's Citrate Agar, Muller Hinton‟s agar, alginic acid (mixture of guluronic and mannuronic acid), Zobell marine agar and broth, cetylpyridinium chloride, Tris base, magnesium chloride, potassium chloride, di- ammonium hydrogen phosphate, sodium chloride, calcium chloride, xylan (from oat spelts), carboxymethyl cellulose (sodium salt), chitin, pectin, carrageenan (Irish moss), starch (amylum, potato starch), gelrite, Gelatin, Casein, magnesium, nickel, potassium, zinc, copper, iron, cobalt, lead, mercury, manganese, lithium, L-asparagine, L-alanine, L- aspartic acid, L-arginine hydrochloride, L-cysteine, L-cysteine free base, L-glutamine, L-

60 glutamic acid, glycine, 4-hydroxyl-L-proline, L-histidine hydrochloride, L-leucine, L- isoleucine, L-methionine, L-lysine hydrochloride, L-proline, L-phenylalanine, L-serine,

L-tyrosine, L-tryptophan, L-threonine, L-valine were obtained from HiMedia labs. 70% ethanol, Crystal violet solution, safranin, Gram's Iodine solution, pullulan, laminarin, β- glucan, antibiotics discs (DE015, HX006, HX007, HX036 HiMedia), HiMedia

Carbohydrate utilizing kit (KB002 and KB009- Part A, B and C) and all other reagents/chemicals were AR grade (analytical reagent) and double glass filtered distilled water was used.

4.1 BACTERIAL STRAINS USED:

Bacterial strain DD-13 isolated from the mangroves of Goa was selected for further study and polyphasic identification. Five Type strains of Microbulbifer closely related to strain DD-13 were used as control for comparison studies. M. salipaludis SM-

1T (obtained from Japan collection of microorganisms); M. celer ISL-39T (acquired from

Korean Collection for Type Cultures); M. elongatus ATCC 10144T (acquired from

German Collection of Microorganisms and Cell Cultures); M. agarilyticus JAMB-A3T and M. hydrolyticus DSM 11525T were kindly provided by Dr. Yuichi Nogi,

(JAMSTEC) Yokosuka, Japan and Professor Gonza´lez University of Georgia, Athens respectively.

4.2 MAINTAINANCE OF BACTERIAL STRAINS:

The bacterial cultures were maintained on slants/ stabs consisting of Zobell marine agar or ASW agar containing 0.2% alginate and stored at 4˚C. The bacterial

61 strains were revived at an interval of 3 to 4 months and fresh stabs/ slants were prepared.

Alternatively, glycerol stocks were also prepared and maintained at -80˚C.

4.3 CHARACTERIZATION OF STRAIN DD-13

4.3.1 COLONY MORPHOLOGY:

The bacterial strain DD-13 was grown on ZMA or ASW agar plates containing 1% alginic acid and routinely incubated at 30˚C for 24 and 48 h respectively for determining colony characteristics and morphology.

4.3.2 DETECTION OF ALGINATE LYASE ACTIVITY:

In order to visualize the alginolytic activity, strain DD-13 was streaked on ASW agar- alginate plates for 48 h and subsequently flooded with 10% cetylpyridinium chloride.

Appearance of clearance zone around the bacterial colonies after incubation for about 10 min is an indication of alginolytic activity (Gacesa and Wusteman, 1990)

4.3.3 GRAM STAINING:

The bacterial strain DD-13 was grown on MA for 24 h at 30˚C. Gram-staining was performed as per Gerhardt et al. (1994). E. coli was used as control.

4.3.4 ELECTRON MICROSCOPY:

The morphology of bacterial strain DD-13 was studied by scanning electron microscopy.

The strain DD-13 was grown in marine broth for 24h at 30°C. The culture suspension was centrifuged at 5000 rpm for 10 min. and aseptically washed thrice with 0.9% saline.

A diluted suspension of bacteria was prepared in 0.9% saline and 0.1ml of diluted

suspension was smeared on a glass slide (1cm2) or metal stud and air dried. The dried

smear was coated with fine gold particles with the help of a gold sputter coater according

to manufacturer‟s instructions (SPI Instruments, India). The coated smear was later

examined under scanning electron microscope (Joel, model JSM- 5800LV) at 12,000X

resolution.

4.3.5 PLATE SCREENING METHOD FOR DETERMINATION OF MULTIPLE

POLYSACCHARIDE DEGRADING ABILITY:

Polysaccharides such as xylan, carboxymethyl cellulose, chitin, pectin, carrageenan,

starch, alginate, agar, gelrite, pullulan, laminarin, and β-glucan were incorporated as a

single/ individual substrate in the ASW medium to determine the capability of strain DD-

13 to degrade polysaccharides. The bacterial strain DD-13 was streaked on the ASW agar

plates containing 0.2% of one of the above polysaccharides and incubated for 48 h at

30ºC.

The polysaccharide degradation by bacterial strain DD-13 was detected by plate

based screening method as mentioned below: a) Detection of xylan degradation: The bacterial strain grown on ASW agar plate

containing 0.2% xylan (from oat spelts, HiMedia) was flooded with 0.1% (w/v) congo

red for 1 to 2 min. andwashed with distilled water, followed by rinsing with 1M sodium

chloride for 1-2 min.(Gessesse and Gashe, 1997). b) Detection of carboxymethyl cellulose degradation: The bacterial strain grown on ASW

agar plate containing carboxymethyl cellulose (sodium salt, HiMedia) was flooded with

0.1% (w/v) congo red for 15 min., followed by flooding with sodium chloride (1 M) for

15 min. (Wood, et al., 1998). c) Detection of chitin degradation: The bacterial strain grown on ASW agar plate

containing chitin (poly-N-acetyl-1,4,β-D-glucopyranosoamine, HiMedia) was flooded

with 0.2% (w/v) congo red for 15 min., followed by flooding with 0.1 M sodium chloride

for 15 min. (Vannini, et al., 1987). d) Detection of pectin degradation: The bacterial strain grown on ASW plate containing

pectin (poly D-galacturonic acid methyl ester, HiMedia) was flooded with 5 N HCl for 15

min (Cabezas and Maeso, 1980). e) Detection of starch degradation: The bacterial strain grown on ASW plate containing

starch (amylum, potato starch, HiMedia) was flooded with Gram‟s iodine/ Lugol‟s iodine

for 15 min. (Ruijssenaars and Hartmans, 2001). f) Detection of alginate and carrageenan degradation: The bacterial strain grown on

ASW plate containing alginate (mixed polymer of guluronate and mannuronate,

HiMedia) or carrageenan (Irish moss, HiMedia) was flooded with10% cetylpyridinium

chloride (Gacesa and Wusteman, 1990; Ohta and Hatada, 2006). g) Detection of agar degradation: The bacterial strain grown on ASW agar plate was

flooded with Lugol‟s iodine for 15 min. (Hodgson and Chater, 1981; Stanier, 1942). h) Detection of Pullulan degradation: The bacterial strain grown on ASW plate containing

pullulan (Sigma) was flooded with 0.2% (w/v) congo red for 15 min. (Ruijssenaars and

Hartmans, 2001)

64 i) Detection of Laminarin and β-glucan degradation: The bacterial strain grown on

ASW plate containing Laminarin (from Laminaria digitata) or β-glucan (from barley,

Sigma) was flooded with 0.2% (w/v) congo red for 15 min. (Teather and Wood, 1982).

4.3.6 BIOCHEMICAL CHARACTERIZATION OF STRAIN DD-13:

Biochemical characterization of bacterial strain DD-13 was carried out according

to Simbert and Kreig (1994). HiMedia Carbohydrate utilizing kits (KB002 and KB009-

Part A, B and C) were used for substrate utilization studies. In all experiments, distilled

water was replaced with ASW and appropriate bacterial controls were used to preclude

interference of ASW on biochemical test. Five Microbulbifer species type strains were

taken as reference strains: M. salipaludis SM-1T; M. celer ISL-39T; M. elongatus ATCC

10144T; M. hydrolyticus DSM 11525T and M. agarilyticus JAMB-A3T

4.3.6.1 Gelatin degradation:

ASW gelatin agar stabs were prepared containing 12% gelatin. The gelatin stabs

was later incubated for 48 h and then placed in the refrigerator for approximately 1h and

observed medium solidification in the refrigerator (negative test result). The medium

remained liquid (partially) and does not solidify in the refrigerator (positive test result).

4.3.6.2 Casein hydrolysis:

Double strength ASW containing 1.8% agar was sterilized for 15 min at 121°C

and 0.75% skimmed milk powder in water was sterilized for 10 min at 121°C. Both

solutions were cooled to 45°C and later mixed to make a homogenous solution. Strain

DD-13 was allowed to grow on these plates at 35°C for 3 days. The clearance zone around the colonies was visualized by flooding the plates with 10% HC1.

4.3.6.3 Indole production:

10 ml of ASW containing 1% tryptone (free of carbohydrate, nitrate and nitrite) was prepared and filter sterilized with 0.22 µm filter membrane. After the addition of inoculums the tubes were kept for 48 h at 35°C. Kovac's reagent (0.5 ml) was added and observed for development of red colour.

4.3.6.4 Malonate utilization:

ASW Broth containing yeast extract (0.1%), 0.3% sodium malonate, 0.025% glucose and 0.002% bromothymol blue in ASW was prepared and bacterial strain DD-13 was inoculated. The tubes were kept at 35°C for 48 h. A colour transformation from green to deep blue indicates malonate utilization.

4.3.6.5 Citrate utilization:

Citrate utilization was checked with Simmon's citrate agar prepared as per the manufacturer's instructions in ASW. The strain DD-13 was inoculated and kept for 48 h at 35°C. The appearance of blue colour was taken as positive control.

4.3.6.6 Catalase test:

Bacterial strain DD-13 was smeared on a clean glass slide with a drop of ASW.

3% hydrogen peroxide (freshly prepared) was added on the wet smear of the strain DD-

13 and observed for formation of air bubbles on the glass slide to confirm the catalase activity.

4.3.6.7 Urease test:

ASW containing 0.01% yeast extract and 0.01% phenol red was autoclaved followed by cooling to 50°C. Urea (2 g/100 m1) was dissolved separately in distilled water and filtersterilized with a 0.22 µm filter membrane. Both were mixed thoroughly to form a homogenous solution. The medium prepared was inoculated with strain DD-13 and kept for 48 h at 35°C. A positive urease activity was observed as a change in the medium colour from red to purple colour.

4.3.6.8 DNase test:

ASW agar plate containing 0.2% DNA (from Herring sperm; Sigma Chemicals) was prepared. Bacterial strain DD-13 culture was inoculated on ASW agar-DNA plate and incubated at 35°C for 48 h. Clearance zone around the colonies observed after flooding the plate with 1N HC1 indicates the production of DNase enzyme.

4.3.6.9 Oxidase test:

1 ml of oxidase reagent was added to sterile Whatman filter paper No.1 and air dried. Loopful of bacterial culture was spread on the filter paper where oxidase reagent

67 was spotted. Development of purple blue colour within 10 s indicated a positive test for oxidase.

4.3.6.10 MR-VP test:

Buffered glucose broth medium supplemented with glucose was prepared in ASW and sterilized in a autoclave at 121°C for 15 min. The tubes were inoculated with strain

DD-13 and incubated at 35°C. Five to six drops of methyl red reagent was added to the tubes and development of bright pink to red color at the top of the medium was recorded as positive for MR test. VP positive reaction was confirmed by adding few drops of

Barritt's reagent.

4.3.6.11 Triple Sugar Iron (TSI) agar test:

TSI medium slants with a minimum of 2 cm butt at the bottom of the slant were prepared in ASW. Strain DD-13 was inoculated on the slant and stabbed in the butt. The tubes were incubated at 35°C. The change in colour of the slant medium (red to yellow) and formation of bubble in the medium after 48 h was taken as a positive control. The change in the colour of the stabbed butt (red to black) was taken as positive for H2S production.

4.3.6.12 Utilization of substrates using HiMedia Carbohydrate utilizing Kit:

The bacterial strain DD-13 was grown in 25 ml of ZMB at 30˚C for 24 h on orbital shaker (130 rpm). The culture supernatant was obtained by centrifuging at 7000 rpm for 10 min in aseptic conditions. The bacterial cell pellet was collected and washed

68 three times with sterile ASW followed by re-suspension of bacterial cell pallet. The cell suspension was inoculated in HiMedia Carbohydrate utilizing kit (KB002 and KB009-

Part A, B and C) to ascertain the substrate utilization. The kits were kept at 30˚C for 48 h and results were recorded.

4.3.6.13 Effect of pH, temperature and NaCl on growth of strain DD-13

The growth of bacterial strain DD-13 at various concentrations of NaCl was studied. The strain DD-13 was grown in a modified ZMB medium. Modified ZMB medium (inherent NaCl) was amended with NaCl at various concentrations (0, 2, 4, 6, 8 and 10%) and inoculated with strain DD-13. The culture was incubated at 30˚C for 24 h on orbital shaker with 130 rpm.

Alternatively modified ZMB was again prepared by adding buffers accustomed to pH 3, 4, 5, 6, 7, 8, 9, 10, 11; acetate (pH 3 and 4), citrate (pH 5 and 6), Tris-Cl (pH 7- 8) and borate (pH 9-10) buffer. The pH effect on growth of bacterial strain DD-13 was determined. The medium was inoculated and incubated at 30˚C for 24 h at 130 rpm.

The growth profile at various temperatures was studied by inoculating ZMB medium with strain DD-13 and incubating it at various temperatures (15, 20, 25, 30, 35,

37, 38, 39 and 40˚C) for 24 h at 130 rpm.

The growth of bacterium was monitored periodically by measuring O.D. at A600 at intervals of 6h.

4.3.6.14 Amino acids Utilization:

To study the amino acid utilization profile various amino acids (L-asparagine, L- alanine, L-aspartic acid, L-arginine hydrochloride, L-cysteine, L-cysteine free base, L- glutamine, L-glutamic acid, glycine, 4-hydroxyl-L-proline, L-histidine hydrochloride, L- leucine, L-isoleucine, L-methionine, L-lysine hydrochloride, L-proline, L-phenylalanine,

L-serine, L-tyrosine, L-tryptophan, L-threonine, L-valine) were used as substrate for growth of the bacterial strain DD-13. Individual amino acids (0.2%) were dissolved in

ASW and filter sterilized through 0.22 µm filter. The inoculum (0.1%) of bacterial strain

DD-13 was added to ASW broth containing 0.2 % of individual amino acid and incubated at room temperature on orbital shaker (130 rpm) for 48 h. Appearance of turbidity in the medium depicts bacterial growth. Medium with amino acid without inoculated bacterial culture was used as control during the experiment.

4.3.6.15 Antibiogram study:

The effect of diverse antibiotics on the bacterial strain DD-13 growth was studied.

Different antibiotic discs such as gentamycin, tetracycline, erythromycin, chloramphenicol, kanamycin, streptomycin, penicillin, neomycin etc. were used to determine their inhibitory effect on the bacterial strain DD-13 growth . Muller Hinton‟s agar plates were swabbed with 24 h old grown culture. The various antibiotics discs

(DE015, HX006, HX007, and HX036 HiMedia) were positioned on the swabbed agar plates and later kept at 30ºC for 48 h.

4.3.6.16 Effect of cations on growth of strain DD-13:

The effect of different cations on the growth profile of strain DD-13 was studied.

Four different concentrations (0.125, 0.25, 0.5 and 1 M) of various cations such as magnesium, nickel, potassium, zinc, copper, iron, cobalt, lead, mercury, manganese, lithium solutions were prepared. Discs with different concentrations of individual cations were used to determine the inhibitory effect on the bacterial strain DD-13 growth . ZMA plates were prepared and swabbed with the 24 h old grown culture. The various cation saturated discs were positioned on the swabbed agar plates and later kept at 30˚C for 48 h to observe the zones of inhibitions.

4.3.7 FAME ANALYSIS OF STRAIN DD-13

The most popular methodology followed for the identification of bacteria is to characterize the types and relative proportions of cellular fatty acid types of the bacteria, using Microbial Identification system (MIDI Inc. Newark, USA).

Strain DD-13 was grown on marine agar up to late-exponential phase at 30°C.

Sample for quantitative cellular fatty acid analysis was done using manufacturer‟s instructions. Several loop-full (approx 40 mg) of bacterial cells were added to 1 ml of saponification reagent (3.75 M NaOH in 50% aqueous methanol) and vigorously vortexed in a sealed sterile teflon lined cap glass tube. Later the tube was incubated in boiling water bath for 30 min, with occasional vortexing. The tube was allowed to cool.

The saponified mixture was later mixed with 6 M hydrochloric acid (1.08 ml) for acidification, further methylated by 0.92 ml of methanol and incubated at 80°C for 10 min in a sealed tube. After getting it cooled to room temperature the methyl esters of fatty

71 acid were extracted with the help of 1.2 ml hexane-diethyl ether (1: l) with gentle mixing.

The organic phase and aqueous phase was separated followed by washing with 3.0 ml of

0.3 M NaOH for 5 min 2/3 of washed organic phase was collected in a clean glass vial.

Quantitative estimation of bacterial cells fatty acids was carry through according to Sherlock Microbial Identification System (MIDI Inc., Newark, USA) instructions

(Sasser, 1990) using Finnigan TRACE DSQ GC–MS system (Thermo Fisher Scientific) containing DB-5 column (J&W Scientific) at 1.5 ml min-1 helium flow and gradual increasing temperature ranging from 140˚C - 280˚C (5 min each) at the rate of 4˚C min-1.

Fatty acid standards were prepared by Microbial ID Inc. and used for calibration of column. The computer generated chromatogram was then compared with internal library data base of Sherlock Microbial Identification System.

4.3.8 GENOMIC DNA ISOLATION:

Genomic DNA from bacterial strain DD-13 was isolated according to Maloy

(1990). Strain DD-13 was grown for 24 h in 25 ml of ZMB at 35°C at 130 rpm. Bacterial cell pellet obtained by centrifugation (8,000 rpm for10 min at 4°C ) was rinsed and resuspended in 1.5 ml of TE buffer (pH 8.0) . 0.9 ml of 10% SDS and 99.9µl of

Proteinase-K (20 mg m1-1 stock) was added to suspension of bacterial cell and later incubated for 60 min at 37°C. After incubation, equal volume of chloroform and Tris- saturated phenol (pH 8.0) (1:1) was tote-up and mixed gently. The aqueous clear layer was separated and collected after centrifugation and re-extracted with phenol: chloroform as described above. The clear aqueous phase was collected into a separate tube and mixed

72 with one-tenth volume of 5 M sodium acetate (pH 5.2) followed by addition of two volumes of chilled ethanol. The tube was incubated at -20°C for 60 min. The precipitated

DNA after centrifugation was quickly rinsed with l ml of ethanol (70%). The ethanol was removed by centrifugation for 10 min at 4°C at 10,000 rpm. The DNA pellet was air dried to remove traces of ethanol and was dispenced in 2 ml of TE buffer. Further, RNase with final concentration of 20 µg ml-1 (from 2 mg/m1 stock) was added and incubated at

30°C for 30 min to remove the RNA contamination. The aqueous phase was again extracted with the help of (1:1) phenol: chloroform mixture. The precipitation of DNA was repeated again as described above, washed, dried and dissolved in TE buffer. The concentration of DNA was estimated at A260 using TE buffer as blank. The DNA profile was analyzed by agarose gel electrophoresis as mentioned below.

4.3.8.1 AGAROSE GEL ELECTROPHORESIS:

Agarose gel electrophoresis of genomic DNA was executed using horizontal slab gel electrophoresis. 0.8 g of agarose (low EEO) was dissolved in 100 ml of TBE electrophoresis buffer (1X) by boiling. The agarose suspension was cooled to 45°C and ethidium bromide (EtBr) was added (0.5 µg ml-1). The agarose solution was poured into a gel casting tray. The gel was solidified by allowing it to stand at 30°C for 30 min. 10 µl of genomic DNA sample was mixed with 6 X gel loading dye and was loaded into the well. The electrophoresis was carried out at 80 V till the dye reached at end. After electrophoresis, the DNA was visualized using UV trans-illuminator.

4.3.8.2 DNA G+C CONTENT ANALYSIS OF STRAIN DD-13:

The genomic DNA G+C content was estimated as mentioned by Tamaoka &

Komagata (1984) with the help of reversed-phase HPLC.

4.3.9 AMPLIFICATION OF 16S rDNA BY POLYMERASE CHAIN REACTION

(PCR):

16S rDNA from chromosomal DNA of strain DD-13 was amplified using universal eubacterial primers 27F (AGAGTTTGATCMTGGCTCAG) and 1525R

(AAGGAGGTGATCCAGCC). These primers correspond to 27 and 1525 of 16S rDNA of E.coli (Brosius et al., 1978). PCR reaction was done in total volume of 25 µl containing genomic DNA (100 ng) , 1 X Taq buffer containing 15 mM MgC12, 200 µM of each of dATP, dCTP, dGTP and dTTP, 10 µM of reverse and forward primers and Taq

DNA Polymerase enzyme (1.25 units). Amplification was completed with the help of thermal cycler (Gene Amp PCR System 9700, Applied Biosystems, Foster City, CA,

USA). The amplification was carried out using following parameters: Initial denaturation at 95°C (5 min) followed by 35 cycles of denaturation at 95°C (l min), annealing at

55°C (1 min) and extension at 72°C (1 min) followed by a final cycle of extension at

72°C (10 min).

The amplified products of PCR from the sample was decontaminated using QIA quick PCR Purification Kit (Qiagen Inc, Hilden, Germany) as per the manufacturer's protocol. The purified DNA concentration was measured using spectrophotometer at A260 and the quality was checked on a 0.8% agarose gel as described previously. The purified

DNA was used for sequencing reaction.

4.3.9.1 AUTOMATED DNA SEQUENCING:

Sequencing reaction was carried out in a 96 well PCR plate using Big Dye

Terminator Cycle Sequencing Kit, Ver. 3.1 (Applied Biosystems, Foster City, CA, USA).

The sequencing reaction was carried out in a total volume of 5 µl containing purified

PCR (30 ng) product, 3 µM primer, 2.5 X Ready Reaction Premix (final concentration l

X), 5 X sequencing buffer (final concentration 1 X). The reaction was performed in

Gene Amp-PCR System 9700 (Applied Biosystems, Foster City, CA, USA). The parameters for the sequencing reaction consists of program for 1 min at 96°C (initial denaturation) followed by 25 cycles for 10 s at 96°C for denaturation followed by annealing for 5 s at 50°C and lastly extension for 4 min at 60°C.

After sequencing reaction was completed, 500 µl of sodium acetate (3 M pH 5.2) and ethanol (12.5 µl) was added to the each well of 96 well PCR plate. The components were mixed using plate vortexer. After incubation for 15 min at 30°C, the plate was centrifuged for 25 min at 4000 rpm at 30°C. The plate was inverted on to a tissue paper and rinsed with ethanol, later allowed to deplete completely. The plate was centrifuged again for 15 min at 300 rpm at 30°C. 75 µl of ethanol (70%) was added to each well of the plate and kept for 5 min at 30°C and centrifuged for 20 min at 25°C at 4000 rpm. The step was repeated again and the plate was allowed to dry in vacuum dryer for 2 min. l0 µl of Hi-Di formamide (Applied Biosystems, Foster City, CA, USA) was added to the each of the plate and incubated at 30°C for 3 min. The plate was later centrifuged for 2 min at

3000 rpm at 30°C. The plate was heat denatured for 3 min at 95°C and quickly cooled by ice- bath for 10 min.

In order to obtain the complete 16S rDNA 7 different internal primers 27F, 121F,

343R, 536F, 704F, 1488R and 1525R were used to obtain overlapping sequences. The sequencing reaction was carried out with above mentioned parameters using DNA

Analyzer 3730 (Applied Biosystems, Foster City, CA, USA) according to manufacturer's operating protocol.

4.3.9.2 PHYLOGENETIC ANALYSIS:

The 7 different overlapping DNA sequences obtained after sequencing by using above mentioned primers were compiled using Chromaspro, Ver. 1.34., to remove the overlap regions in the sequence. DNA sequences homologous to 16S rDNA sequence of strain DD-13 was obtained from Ribosomal database project (RDP II)

(http//:rdp.cme.msu.edu) using seqmatch program. Alternatively, blast analysis of 16S rDNA sequence of strain DD-13 against other 16S rDNA sequences present in other

DNA sequence databases (GenBank/DDBJ/ EMBL) was also performed (Altschul et al.,

1997). The 16S rDNA sequences that showed similarity to the query sequence were obtained in FASTA format from NCBI database (www.ncbi.nlm.nih.gov.in) and aligned using Clustal W, Ver. 1.83. (Thompson, et al., 1997). Vibrio communis R40496T

GU078672 was used as out-group sequence. The aligned sequences were saved in

FASTA format. The gaps from 5' and 3' end of the aligned sequences were removed by use of MEGA program, version 3.1., (Kumar et al., 2004). The sequences were aligned to remove the internal gaps and the final sequence was saved in PHYLIP format.

Phylogenetic tree construction was done using the PHYLIP, Ver. 3.67., (Felsenstein J,

2006). The sequences were analyzed for stability using SEQBOOT program of PHYLIP.

Bootstrapping was performed for 10000 replicates. DNA distances were analyzed by the program DNADIST, with Kimura-2 parameter for 1000 data sets. The transition and transversion rate was kept at 2.0. Phylogenetic trees construction was done using

DNAML (maximum likelihood), DNAPARS (parsimony) and NEIGHBOR algorithms.

Phylogenetic trees construction was done using DNAPARS and DNAML with 1000 data sets with constant variation among sites from SEQBOOT output data. Phylogenetic tree constructed using NJ was done with-out file from DNADIST data. The consensus phylogenetic tree was saved to out tree and tree was viewed with Tree View, Ver. 1.6.6.

4.3.10 DNA–DNA HYBRIDIZATION STUDY:

The 16S rDNA sequence similarity of different Microbulbifer was judged against with the strain DD-13 by blast analysis. DNA–DNA hybridization and study of relatedness was performed at 48˚C for 4 h and quantified fluorometrically (Ezaki et al.

1989).

According to the 16S rDNA taxonomical tree, the DD-13 is found to be closely related to the Microbulbifer salipaludis and Microbulbifer hydrolyticus therefore these three strains were made as labeled DNA probe for the relatedness comparison.

4.3.11 EXTRACTION AND ANALYSIS OF ISOPRENOIDS QUINONES:

Extraction and analysis of isoprenoids quinones was done as according to

Komagata and Suzuki (1987). Freeze-dried cells of DD-13, M. celer, M. elongatus, M.

77 salipaludis, M. hydrolyticus and M. agarilyticus (100-300 mg) were suspended in 20 ml of chloroform-methanol (2: 1,v/v) and gently stirred overnight. After filtration, the extract was evaporated to dryness and dissolved in a small amount of acetone. Acetone solution was applied to silica-gel TLC (Merck F254, 0.5 mm thickness) and developed with benzene. Vitamin K was used as control for ubiquinone. The quinone spots were visualized by UV light at 254 nm. Ubiquinones spots were observed with Rf ≈ 0.4. The corresponding spot was scraped off and extracted with acetone. Before concentration, the acetone solution was filtered by Column-guard SJFH L04 (Millipore Corp., Bedford,

Mass., USA). Samples were stored at low temperature in small brown glass tubes. The quinone samples were applied to reverse phase HPLC (e.g. HYPERSIL ODS).Elution solvent used was methanol-isopropanol (2:1, v/v). Ubiquinones were observed at 275nm.

4.4 RESULTS:

4.4.1 Phenotypic characteristics:

The bacterial strain DD-13 was observed as grayish yellow coloured colonies with slight depressions on marine agar medium. The circular smooth colonies obtained were 2 –3 mm in diameter, entire, glossy and depicted convex elevation after 48 h incubation at 35 ºC (Fig. 4.1).

When strain DD-13 was grown on ASW agar plates for 48 h, small (pin-point) pits/ depressions with slight turbidity and clearance zone were observed around the colonies (Fig. 4.1 a). Addition of 10 % CPC to ASW agar alginate plates resulted in formation of clearance zones around the colonies, with non-degraded polysaccharide

(alginate) turning opaque white (Fig. 4.1 b).

(a) (b)

Figure 4.1: Morphological characteristics of bacterial strain DD-13 a) Strain DD-13 on ASW alginate-agar plate. b) Strain DD-13 on ASW agar-alginate plate after flooding with 10% CPC.

4.4.1.1 Gram staining:

The bacterial cells were grown for 24 h at 35°C on marine agar subjected to Gram staining and examined under light microscope. Gram character of the strain DD-13 was negative which was evident by the observation of reddish-pink colour bacterial cells. The bacterial cells were either rods or cocci. The cells were predominantly rod shaped during early stages of growth that later changed to cocci during late phases of cell growth cycle

(Fig. 4.2)

Rod and cocci cells

Figure 4.2: Gram staining character of strain DD-13

Single bacterial cell

(a) (b)

Figure 4.3: SEM of bacterial strain DD-13

4.4.1.2 SEM of the bacterial strain DD-13

The electron microscopy for the bacterial strain DD-13 showed the presence of

long rods in chain. The bacterial cells were observed to be rod shaped during the earlier

growth phase and later converted to coccoid–ovoid cells during late stages of growth (i.e.

after 24 h) as observed from Fig. 4.4. The average length and width of the rod shaped

bacterial cell was determined as 1.2 × 0.3 µm ± 0.02. The average size of the coccus

bacterial cell was measured to be 0.54 µm ± 0.02 in diameter.

4.4.1.3 Bacterial strain DD-13 degrades multiple polysaccharides:

The strain DD-13 was observed to degrade multiple polysaccharides as shown in

Table 4.1 and Fig 4.4. The clearance zones against turbid/ opaque background observed

during plate based assay demonstrates degradative activity of the strain. The strain did

not show any growth or clearance zone on the gelrite plate.

Table 4.1: Degradation of multiple polysaccharides by bacterial strain DD-13

Polysaccharides Degradation Polysaccharides Degradation

Xylan + Alginate +

Carboxymethyl cellulose + Agar +

Chitin + β- glucan +

Pectin + Laminarin +

Carrageenan + Pullulan +

Starch +

Xylanase CMCase Chitinase Pectinase

Carrageenase Amylase Alginase Agarase

β- Glucanase Pullulanase Laminarinase

Figure 4.4: Plate based assay for detection of degradation of multiple polysaccharides.

4.4.2 BIOCHEMICAL PROPERTIES OF STRAIN DD-13:

Growth of bacterial strain DD-13 was observed from 20 to 38ºC, but not at 4, 15,

39 or 40ºC. The optimal temperature for growth was observed as 35ºC. The results of biochemical characterization for strain DD-13 is depicted in table 4.2.The strain DD-13 grew in pH ranging from 5 to 10with pH 7 being optimal for growth. Although, strain

DD-13tolerated and grew upto 10% (w/v) NaCl, it was not essential component for the growth. Anaerobic growth was not seen on MA. Xanthine and urea are not hydrolyzed whereas Tweens 20/40/60/80 are hydrolyzed. Acid production was observed during utilization of D-glucose, D-arabinose, D-cellobiose, esculin and succinate whereas D- fructose, sucrose, melibiose, D-mannose, D-mannitol, trehalose, D-melezitose, L- arabinose, D-raffinose, citrate, L-rhamnose, succinate, D-ribose, benzoate, D-sorbitol, L- malate, inositol, formate, salicin and L-glutamate were not utilized. Gas production was not observed with any of the tested carbohydrates. No hydrolysis of casein and gelatin was observed. Nitrate reduction and ONPG tests were negative. Strain DD-13 was positive for catalase and oxidase test. Indole production was negative and strain was weakly positive for H2S production. The results were negative for DNase and MR-VP tests.

4.4.2.1 UTILIZATION OF VARIOUS AMINO ACIDS:

Various amino acids were tested to determine the utilization by the strain DD-13.

The strain DD-13 does not essentially require any vitamins/ amino acids or any other growth factors for proliferation in the ASW medium alginate supplemented as the individual energy and carbon source. Strain DD-13 was found to utilize only few amino

83 acids like L- alanine, glycine, L-cysteine, L- histidine hydrochloride and L- proline as shown in Table 4.3.

Table 4.2: Biochemical characteristic of strain DD-13.

Tests Results

Morphology rods Gram character -ve Motility -ve Temp. range 20-38 Salinity (%) 0-10 pH range 5-10

Catalase + Oxidase + Citrate utilization - Lysine andornithine utilization + Phenylalanine deamination - Urease test - Nitrate reduction -

H2S production + Casein hydrolysis -

Carbohydrates

Glucose,Maltose, Xylose, D- Arabinose, Cellobiose, + Esculin, Succinate Raffinose,Trehalose, Melibiose, Inuline, Sodium gluconate, Salicin, Dulcitol, Inositol, Sorbitol, Mannitol, Adonitol, Ribose, Melezitose, - Xylitol, Malonate, Sorbose, ONPG, Mannose, Galactose, Lactose, α- Methyl-D- glucoside, α- Methyl-D- mannoside

Table 4.3: Utilization of various amino acids by strain DD-13

Amino acids Growth of strain DD-13 L-alanine + L-arginine hydrochloride - L-asparagine - L-aspartic acid - L-cysteine free base - L-cysteine + L-glutamic acid - L-glutamine - glycine + L-histidine hydrochloride + 4-hydroxyl-L-proline - L-isoleucine - L-leucine - L-lysine hydrochloride - L-methionine - L-phenylalanine - L-proline + L-serine - L-threonine - L-tryptophan - L-tyrosine - L-valine -

4.4.2.2 EFFECT OF VARIOUS ANTIBIOTICS ON THE GROWTH OF

THE BACTERIAL STRAIN DD-13:

Different antibiotics discs (DE015, HX006, HX007, HX036 HiMedia) were tested to determine the resistance and inhibitory effect on the growth of bacterial strain DD-13.

Strain DD-13 was found to be sensitive to all the tested antibiotics as per the HiMedia zone size interpretation chart (Table 4.4).

4.4.2.3 EFFECT OF CATIONS ON PROLIFERATION OF STRAIN DD-13

Whatman filter paper disc saturated with different concentrations of various cations/ metal salts were tested to determine the resistance and inhibitory effect on the growth of bacterial strain DD-13. Strain DD-13 was found to be resistant to magnesium, potassium and lithium salts at concentration range of 0.125 to 1.0 M whereas medium resistant for calcium and lead salts at concentrations ranging from 0.125 and 0.25 M was observed. The strain DD-13 was very sensitive for nickel, zinc, copper, iron, cobalt, mercury and manganese salt as depicted from Table 4.5.

Table 4.4: Antibiogram of strain DD-13.

Antibiotics Resistant / Sensitive

Gentamicin (10mcg) Sensitive Tetracycline (30mcg) Sensitive Erythromycin (15mcg) Sensitive Chloramphenicol (30mcg) Sensitive Kanamycin (30mcg) Sensitive Streptomycin (10mcg) Sensitive Penicillin G (10mcg) Intermediate Neomycin (30mcg) Sensitive Ampicillin (10mcg) Sensitive Amoxyclav (30mcg) Sensitive Cefotaxime (30mcg) Sensitive Co-trimoxazole (25mcg) Sensitive Tobramycin (10mcg) Sensitive Ciprofloxacin (5mcg) Sensitive Cephalexin (30mcg) Sensitive Amikacin (30mcg) Sensitive Nitrofurantoin (300mcg) Sensitive Netillin (30mcg) Sensitive Nalidixic acid (30mcg) Sensitive Clarithromycin (15mcg) Sensitive Oxytetracycline (30mcg) Sensitive Furaolidone (50mcg) Sensitive Ceftazidime (30mcg) Sensitive

Table 4.5: Effect of various cations on growth of strain DD-13.

Metals salts Molarity Resistant/ Sensitive

MgSO4 0.125 - 1.0 M Resistant

NiCl2 0.125 - 1.0 M Sensitive

KCl 0.125 - 1.0 M Resistant

ZnSO4 0.125 - 1.0 M Sensitive

CuCl2 0.125 - 1.0 M Sensitive

FeCl3 0.125 - 1.0 M Sensitive

0.5 – 1.0 M Sensitive CaCl2 0.125 to 0.25M Resistant

CoSo4 0.125M to 1.0M Sensitive

0.5 – 1.0 M Sensitive Pb(NO)3 0.125 to 0.25 Resistant

HgCl2 0.125M to 1.0M Sensitive

MnSO4 0.125M to 1.0M Sensitive

LiCl 0.125M to 1.0M Resistant

Note: Zone size > 0.5mm were considered as resistant.

4.4.3 FAME ANALYSIS OF THE BACTERIAL STRAIN DD-13:

The bacterial strain DD-13 cellular fatty acid composition was evaluated by

Microbial Identification system (MIDI Inc., Newark, USA). Fatty acid such asiso-C15 : 0 and C18:17cfatty acids were predominantly observed in strain DD-13. C12:03-OH fatty acid was conspicuously absent. The percentage of other resolved fatty acids are given in Table

4.6.

4.4.4 AGAROSE GEL ELECTROPHORESIS:

The genomic DNA isolated from strain DD-13 was observed by using agarose gel electrophoresis apparatus. After electrophoresis, the DNA was visualized under UV.

As no RNA contamination was observed, the genomic DNA was confirmed to be pure was used for sequencing (Fig. 4.5).

1 2

Figure 4.5: Genomic DNA of strain DD-13 (Lane 1 and 2)

Table 4.6: Percentage cellular fatty acid composition of strain DD-13.

Fatty acids %

C10:0 1

C14:0 1

chain - C15:0 1

C16:0 14 Straight

C17:0 2

C18:0 1

iso-C11:0 2

iso-C15:0 24 anteiso-C -

15:0

iso-C15:1 1

iso-C16:0 - Branched iso-C17:0 9

anteiso-C17:0 -

iso-C17:19c 16

iso-C18:1 -

C16:17c 5

C17:18c -

C18:17c 18

Unsaturated

iso-C11:03OH 3 Hydroxy

C17:0cyclo 2

Cyclo C19:0cyclo 1

4.4.4.1 ANALYSIS OF 16S rDNA GENE SEQUENCE OF THE SELECTED

BACTERIAL STRAIN DD-13:

A continuous stretch of 1,532-bp fragment of strain DD-13 16S rDNA gene was sequenced and is depicted in Fig. 4.6. The sequence was submitted to GenBank (NCBI) with accession No. HQ424446.1 and Ref seq. accession No. NR_109105.1. The strain

DD-1316S rDNA sequence was compared with other Type strains present in Ribosomal

Database Project to obtain the most similar sequence matches. The homology search at

RDP and NCBI (Figure 4.7) indicated that the sequence was homologous to the member of gamma subclass of Proteobacteria and in particular to the genus Microbubifer. It shared a highest degree of similarity with Microbulbifer hydrolyticus DSM 11525T with

99% identity (Table 4.7). The 16S rDNA sequence of DD-13 demonstrate 98% similarity with Microbulbifer salipaludis SM-1T, whereas Microbulbifer elongatus DSM 6810T,

Microbulbifer celer ISL-39Tand Microbulbifer agarilyticus JAMB A3T demonstrate a similarity of 97% (Table 4.7). These results depict that the bacterial strain DD13 is closely associated with the genus Microbulbifer. Thus strain DD-13 exhibited 16S rRNA gene sequence similarity levels of 97.1–98.9 % to the type strains of species of the genus

Microbulbifer and hence these type strains were chosen for further studies. The phylogenetic hierarchy was prepared using sequences of closely related type strain species and genera. Three alternate algorithms were used to prepare the phylogenetic tree

(Figure 4.8, 4.9 and 4.10). The phylogenetic tree constructed clearly shows that strain

DD-13 forms a common coherent cluster with the clade that comprises Microbulbifer. based on neighbor-joining method phylogenetic analysis depicted that strain DD-13 formed an evolutionarily distinct lineage by falling in the same clade within the cluster

91 comprising M. hydrolyticus IRE-31T, M. salipaludis SM-1T, M. agarilyticus JAMB A3T,

M. celer ISL-39Tand M. elongatus DSM 6810T, supported by a bootstrap re-sampling value of 100 % (Fig. 4.8). The same pattern was also depicted by phylogenetic hierarchy obtained through maximum-parsimony and maximum-likelihood methods (Fig. 4.9 and

4.10)

Figure4.6: 16S rDNA sequence of the strain DD-13(Accession No. HQ424446.1; Ref seq. Accession No. NR_109105.1)

: NCBI BLAST similarity index of 16S rDNA sequence of sequence strain DD rDNA 16S of similarity index NCBI : BLAST

4.7

ure Fig

Table 4.7: Similarity index of 16S rDNA sequence of strain DD-13

DDBJ BLASTn NCBI Strains SequenceAccessionnumber (ver.2.2.24) % BLAST %

Microbulbifer hydrolyticus AJ608704 98 99 DSM 11525T

Microbulbifer salipaludis AF479688 97 98 SM-1T

Microbulbifer celer EF486352 96 97 ISL-39T

Microbulbifer agarilyticus AB158515 96 97 JAMB-A3T

Microbulbifer elongatus AF500006 96 97 DSM 6810T.

T 100 Microbulbifervariabilis Ni2088 AB167354 94 T  Microbulbiferepialgicus F104 AB266054 T Microbulbifermaritimus TF17 AY377986 T 85  Microbulbiferdonghaiensis CN85 EU365694 51 Microbulbiferokinawensis ABABA23T AB500893

T 95 Microbulbifermarinus Y215 GQ262812 Microbulbiferthermotolerans JAMBA94T AB124836  Microbulbiferchitinilyticus ABABA212T AB500894 100 70  T 70 Microbulbiferhalophilus YIM91118 EF674853

Microbulbiferyueqingensis Y226T GQ262813 Microbulbiferceler ISL39T EF486352 82 Microbulbiferagarilyticus JAMBA3T AB158515  93 81  T Microbulbiferelongatus DSM6810 AF500006  T 75 Microbulbifersalipaludis SM1 AF479688  Microbulbifermangrovi DD-13THQ424446 60  T 90 Microbulbiferhydrolyticus DSM11525 AJ608704

Marinobacteroulmenensis Set74T FJ897726  Marinobacterguineae LMG24048T AM503093 100  T 99 Marinobactersedimentalis R65 AJ609270  T 62 Marinobacterflavimaris SW145 AY517632 Vibrio communis R40496T GU078672

0.02

Figure 4.8: Neighbor-joining phylogenetic tree based on 16S rDNA gene sequences, depicting the hierarchy of strain DD-13T, Microbulbifer species and some other related taxa. Bootstrap percentages (based on 10000 replications)>50% are shown at branch points. () indicates that the corresponding nodes were also recovered in the trees generated with the maximum-likelihood and maximum-parsimony algorithms. Vibrio communis R40496T was used as an out-group. Bar 0.02 substitutions per nucleotide position.

T 89 Microbulbifer elongatus DSM6810 (AF500006) Microbulbifer agarilyticus JAMBA3T (AB158515) 52 Microbulbifer salipaludis SM1T (AF479688)(AF479688) Microbulbifer hydrolyticus DSM11525T (AJ608704) 74 MicMicrobulbiferrobulbifer mangrovi DD13T (HQ424446) Microbulbifer celer ISL39T (EF486352) 98 Microbulbifer yueqingensis Y226T (GQ262813) Microbulbifer halophilus YIM91118T (EF674853) 65 MicrobulbiferMicrobulbifer chitinilyticus ABABA212T (AB500894) MicrobulbifMicrobulbiferer thermotolerans JAMBA94T (AB124836) 90 Microbulbifer marinus Y215T (GQ262812) Microbulbifer okinawensis ABABA23T (AB500893) 58 Microbulbifer donghaiensis CN85T (EU365694) 54 MicrobulbiferMicrobulbifer maritimus TF17T (AY377986) T 86 Microbulbifer epiepialgicusalgicus F104 (AB266054) 99 Microbulbifer variabilis Ni2088T (AB167354) Marinobacter oulmenensis Set74T (FJ897726) Marinobacter sedimentalis R65T (AJ609270) 100 T 70 Marinobacter guineae LMG24048 (AM503093) Marinobacter flavimaris SW14SW1455T (AY517632) Vibrio communis R40496T (GU078672)

Figure 4.9: Maximum-parsimony tree based on nearly complete 16S rDNA gene sequences, showing the phylogenetic relationships between strain DD-13T and members of related taxa. Bootstrap values are based on 1000 replicates; only values greater than 50

% are shown at branch points. The sequence of Vibrio communis R40496T (GU078672) was used as an outgroup. Bar, 20 % nt sequence divergence.

93 Microbulbifer elongatus DSM6810T (AF500006) Microbulbifer agarilyticus JJAMBA3AMBA3T (AB158515) 62 Microbulbifer salipaludis SM1 T (AF479688) Microbulbifer hydrolyticus DSM11525T (AJ608704) 60 77 Microbulbifer mangrovi DD13 T (HQ424446) MicrobulbiferMicrobulbifer celer ISL39T (EF486352) 100 Microbulbifer yueqingensis Y226T (GQ2(GQ262813)62813)

64 Microbulbifer halophilus YIM91118T (EF674853) 69 Microbulbifer chitinilyticus ABABA212 T (AB500894) 60 Microbulbifer thermotolerans JAMBA94 T (AB124836) MicrobulbiferMicrobulbifer marinus Y215T (GQ262812) 98 Microbulbifer okinawensis ABABA23 T (AB500893) 61 Microbulbifer donghaiensis CN85 T (EU365694) 53 Microbulbifer maritimus TF17 T (AY377986) T 90 Microbulbifer epialgicus F104 (AB266054) 100 Microbulbifer variabilisvariabilis Ni2088 T (AB167354) Marinobacter oulmenensis Set74T (FJ897726) Marinobacter guineae LMG24048T (AM503093) 100 T 95 Marinobacter flavimaris SW145 (AY517632) 58 Marinobacter sedimentalis R65 T (AJ609270) Vibrio communis R40496T (GU078672)(GU078672)

0.02

Figure 4.10: Maximum-likelihood tree based on nearly complete 16S rDNA gene sequences, depicting the phylogenetic relationships between strain DD-13T and members of related taxa. Bootstrap values are based on 1000 replicates; only values greater than 50

% are shown at branch points. The sequence of Vibrio communis R40496T (GU078672) was used as an out-group. Bar, 0.02 nt substitutions per site.

4.4.4.2 DNA %G+C CONTENT ANALYSIS OF THE Microbulbifer

STRAIN DD-13:

The chromosomal DNA was purified according to Maloy (1990) and was observed as a single band after agarose gel electrophoresis. The DNA G+C content of strain DD13 was observed to be 61.4%.

4.4.4.3 DNA–DNA HYBRIDIZATION STUDY:

DNA–DNA hybridization was performed for the investigation of DNA relatedness among the selected strains. The DNA of bacterial strain DD-13 was compared with the closely related type strains of Microbulbifer such as Microbulbifer hydrolyticus

DSM 11525T, Microbulbifer celer strain ISL-39T, Microbulbifer salipaludis strain SM-

1T, Microbulbifer elongatus DSM 6810T, Microbulbifer agarilyticus JAMB-A3T, as these strain were observed to have similarity above 96% with the DNA of bacterial strain DD-

13 as shown earlier (Table 4.8).

On the basis of blast analysis, strain DD-13 is found to be closely related to the

Microbulbifer salipaludis and Microbulbifer hydrolyticus (Table 4.8). Hence DNA from these three strains were labeled as probe for the DNA–DNA relatedness (%) studies and hybridized with other type strains of Microbulbifer. As seen in Table 4.9, experimental set A represent strain DD-13 as labeled DNA probe hybridized with all other type strains, experimental set B represent M. hydrolyticus DSM 11525T as the DNA probe and set C represent M. salipaaludis SM-1T as the DNA probe. Table 4.10 depicts the relatedness

DNA–DNA hybridization percentage.

Thus DNA–DNA hybridization tests of the genomic DNA of DD-13 with M. hydrolyticus IRE-31T, M. salipaludis JCM 11542T, M. celer KCTC 12973T, M. agarilyticus JAMB-A3Tand M. elongates DSM 6810T revealed similarities of 28, 33, 27,

26 and 32 %, respectively.

Table 4.9: DNA–DNA reassociation between the strain DD-13 and closely related

Microbulbifer species.

DNA–DNA relatedness (%)

Strains CategoryA Category B Category C

Microbulbifer sp. strain DD-13 100 35 28

Microbulbifer hydrolyticus DSM 11525T 28 100 30

Microbulbifer salipaludis strain SM-1T 33 32 100

Microbulbifer celer strain ISL-39T 27 32 25

Microbulbiferagarilyticus strain JAMB-A3T 26 24 26

Microbulbifer elongatus DSM 6810T. 32 33 34

4.4.5 Isoprenoid extraction and analysis:

The ubiquinone were extracted from the strain DD-13 as well as five type strains of Microbulbifer strains namely M. celer, M. elongatus, M. salipaludis, M. hydrolyticus and M.agarilyticus and analyzed according to Komagata, and Suzuki (1987).

As observed from the Fig. 4.11, similar profile was observed on TLC for all the tested Microbulbifer strains, the ubiquinone (Rf = 0.42) was present in all the bacterial

99 samples. The spots for ubiquinone for DD-13T and M. agarilyticus were eluted and further analyzed by reverse phase analysis and re-confirmed as to be the same ubiquinone with similar peaks at same retention time.

Solvent front (13 cm)

Metaquinone (very light band) (9.4cm) (Rf =0 .72)

5.5cm (Rf = 0.42)

3.7 cm (Rf =0.28)

2.5cm (Rf =0.192)

1.7cm (Rf = 0.131) (Red colour)

0.6 cm (Rf = 0.046)

Sample loaded

1 2 3 4 5 6 7 8

Fig. 4.11: TLC for separation of isoprenoid (1: Microbulbifer strain DD-13, 2: M. celer,

3: M. elongatus, 4:M. salipaludis, 5: M. hydrolyticus, 6:M. agarilyticus, 7:vitamin K, 8:

Microbulbifer strain DD-13

100

Reverse Phase HPLC of ubiquinone from strain DD-13 and M. agarilyticus

[Column used: Hypersil ODS 5.0µm, 4.6 x 250mm; Flow rate: 1ml/min; Two phase

solvent gradient: Methanol-isopropanol (2:1, v/v); Wavelength for detection: 275nm]

Strain DD-13T Ref-std (Microbulbifer agarilyticus) Injection Volume: 20 µl Injection Volume: 20 µl

Detector 1-275nm Detector 1-275nm Retention Time Retention Time

4000 4000

mAU mAU mAU mAU 2000 2000

2000 2000

6.062

4.260

3.797

1.658

1.835 6.035

0 0 1.943 2.873 2.912 0 0

0 5 10 15 20 25 30 35 40 0 5 10 15 20 25 30 35 40 Minutes Minutes

Retention Area Area Retention Area Area Time Percent Time Percent 2.91 133762060 99.383 2.87 116642188 99.956

101

4.5 DISCUSSION:

Mangrove ecosystems are known to be highly productive ecosystems. The salinity in mangroves varies in the range of 0 to 90 ppt facilitating growth of facultative halophytes .The mangrove ecosystems are important bridge between marine and terrestrial ecosystems. Litter from trees, subsurface root foliage and vegetative remains provide significant inputs of organic carbon in the form of complex polysaccharides

(CPs) such as pullulan, cellulose, pectin, xylan, etc. to mangrove sediments. The degradation of these organic matters and recycling of nutrients in mangrove sediments is mediated by both aerobic and anaerobic microorganisms. The colonization of CP decomposing micro-organisms within litter for the formation of detritus is a significant ecological process which is important for recycling of carbon which results in fertility of the mangrove ecosystem. Thus different microorganisms play important role in the mangrove ecosystems performing various activities such as photosynthesis, nitrogen fixation, methanogenesis, production of antibiotics and polysaccharide degradation etc.

Several groups have reported the isolation of ecologically and commercially important bacterial strains from mangrove ecosystem. Noraphat et al., (2013) isolated a bacteriocin-producing lactic acid bacteria Lactococcus lactis subsp. lactis KT2W2L from mangrove forests in southern Thailand that has potential to act as bio-control agent. A bacterium which degrades phenanthrene was screened and isolated from sediments of mangrove in Hong Kong (Tam et al., 2002). Halobacillus faecis sp. nov., a spore- forming bacterium was also screened from a mangrove area near Ishigaki Island, Japan

(Sun-Young, et al., 2007). A bacterium with antifungal activity, phosphate solubilizing capabilities, as well as amylase, protease and lipase producer was isolated from

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Bhitarkanika mangrove ecosystem of Orissa coast (Gupta, et al., 2007). Behra et al.,

(2014) isolated and characterized a sulphur oxidizing bacteria from mangrove soil of

Mahanadi river delta, Odisha. Holguin et al., (1992) isolated two new diazotrophic bacteria Listonella anguillarum and Vibrio campbellii, and one non-nitrogen-fixing bacterium Staphylococcus sp. from the rhizosphere of the mangrove trees. Doan and

Nguyen (2012) isolated polyhydroxy alkanoates releasing bacterium strain

QN271isolated from the mangrove soil samples from Quang Ninh province. In the present study a bacterial strain DD-13 was isolated from the sediments of mangrove located at Divar Island, Goa, India which can degrade a total of eleven different ICP.

The genus Microbulbifer was coined by Gonza´lez et al., (1997) to include a gram negative, strictly aerobic, rod-shaped, and biopolymer-decomposing marinebacterium.

Microbulbifer belongs to the phylum Proteobacteria and have numerous vesicles on the membrane surface. Additionally, the iso-C15 : 0 presence as the major fatty acid and Q-8 as the predominant isoprenoid quinone are major characteristic features for the genus

Microbulbifer. Furthermore, species of the genus Microbulbifer are also known to demonstrate a characteristic rod–coccus growth related cycle (Nishijima et al., 2009).

The association of resting coccoid cells and vegetative rod associated with the growth phase is the first such description for the Microbulbifer genera. Subsequently, manyl novel species belonging Microbulbifer genus having the characteristic to degrade many different polysaccharides have been reported from various habitats such as intertidal sediment, salt marsh, solar saltern, marine algae, sediments (marine and deep-sea) and mangrove forests (Yoon et al., 2003a, 2003b, 2004, 2007; Miyazaki et al., 2008;

Nishijima et al., 2009; Wang et al., 2009; Baba et al., 2011; Zhang et al., 2012).

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4.5.1 PHENOTYPIC, GENOTYPIC AND CHEMOTAXONOMIC

CHARACTERISTICS:

4.5.1.1 Phenotypic characters:

On the basis of 16S rDNA blast analysis of strain DD-13, M. celer, M. elongatus,

M. salipaludis, M. hydrolyticus and M. agarilyticus were observed to demonstrate greater than 96% identity and hence were chosen for comparison. The colony morphology is the first characteristics that differentiate a bacterial strain from another. The colony of bacterial strain DD-13 was entire, circular, convex, grayish yellow colour and 2 - 3 mm in diameter after an incubation of 48 h on ZMA. The colonies of Microbulbifer celer were bigger and more butyrous, campared to the colonies of strain DD-13 (Yoon et al.,

2007). M. elongatus colonies depicted yellowish-brown pigment that diffused into the medium in presence of peptone and sunken colonies were observed due to agar liquefaction (Palleroni, 1984). M. salipaludis colonies are cream coloured, smooth irregular and 2-4 mm in diameter after 3 days of incubation on ZMA plates (Yoon et al.,

2003).

The cells of strain DD-13 were rod shaped with dimensions of 0.3 × 1.2 µm and occasionally coccoid cells were also observed. The average size of the coccoid bacterial cells was measured to be 0.54 µm in diameter. The presence of rods as well as coccus cells, have been reported in other type strains of Microbulbifer species. The dimension of rod and coccus cells of M. viriabilis was 0.42-0.63 × 3.5-13.5 µm and 0.4-0.85 µm respectively (Nishijima, et al., 2009). As reported by Nishijima, et al., (2009) the species

104 of genera Microbulbifer exhibited a discernible cycle of rodo- coccus cell cycle related to their growth phase. Nearly all the rod cells seen throughout the growth cycle, gets converted to coccoid–ovoid cells after the end of proliferation. The coccoid cells have been reported to be optically much more denser and opaque compared to the rod cells having coarse surfaces, which might play a significant role in attachment to fresh substratum while drifting in the respective ecosystem of the bacteria.

The continues fragmentation with rounding of each unit or a successive division could be the reason for the transformation of rod cells to ovoid cells. These coccoid cells attain the rod shape when comes in contact with fresh substratum. It has been reported that the budding of rods from the coccoid cells are visualized only after 6–7 h incubation and thereafter they grow rapidly to form long and curved rod cells for 24 h hence accomplishing rod–coccus cell growth cycle as observed in the growth phase of strain

DD-13.

The coccoid cells of Microbulbifer strains are viable for months. The coccoid cells formed are heat resistant whereas the rods are heat labile, but the heat resistance of coccoid cells diminishes gradually over a period of time. All of the rod cells of

Microbulbifer sp. are transformed to circular cells to survive a long periods, considered to be a nutrient starvation period, as the isolates do not acquire other stress in their respective niches.

In the present study, the formation of spherical bodies at the tips of rod cells were also observed other then the rod–ovoid cells cycle in older cultures of strain DD-13.

Nishijima et al., (2009) has also reported the presence of spherical bodies in

Microbulbifer species. These were completely circular when get separated from the main

105 body of the cell and had a smooth surface.The number of these spherical bodies increases when grown on marine agar or in marine broth at 30˚C as also reported by Nishijima et al., (2009). They were formed by brusting out like buds from the tips or other parts of rod shaped cells. The integrity of the cell surface gets weak in the older cells which became puffed-up by internal cytoplasmic pressure appearing like a bud (Nishijima et al., 2009).

These types of formation of spherical or coccoid bodies in older cultures has been reported already for many Gram-negative bacteria (Krieg, 1976, 1984).

The microscopic and macroscopic morphological characteristics of a bacterium are the first characters thus forming the corner stone for most of the bacterial identification algorithms. In the present study, the strain DD-13 was found to be Gram negative indicating that the cell walls contain a thin layer of peptidoglycan network.

Gram negative bacteria are believed to be comparatively more resistant to antibiotics than

Gram positive bacteria, due to their relatively impermeable lipid based bacterial outer membrane.

According to the new ISO 20776-1 standards, which is valid and followed over the world, the effect of any antibiotic has been defined into three categories (Rodloff, et al., 2008):

 Susceptibility (S): A bacterial strain is called susceptible if it is inhibited by a

concentration of the drug (in-vitro) which can be coupled with high possibility of

therapeutic success.

 Intermediate (I): the sensitivity can be called as intermediate when the drug related

with an uncertain therapeutic effect shows in vitro inhibition of the strain.

106

 Resistant ®: A bacterial strain is known to be resistant if the drug related with high

likelihood of therapeutic failure shows in vitro inhibition of the strain

Unlike Gram negative bacteria, that are comparatively more resistant to antibiotics than Gram positive bacteria, strain DD-13 was found to be highly susceptible to most of the antibiotics tested according to the above defined ISO standards.Similarly

M. agarilyticus was sensitive for ampicillin (10µg), chloramphenicol (30µg), malidixic acid (30µg) and resistant for tetracycline (30µg), gentamicin (10µg), streptomycin (10µg)

(Miyazakiet al., 2008).

4.5.1.2 BIOCHEMICAL CHARACTERISTICS:

However the bacterial classification based on genetic discrepancy describes the evolutionary relationships, the biochemical and morphological features remains the most practical way for identification and classification. The first scheme for bacterial identification was presented in 1984 in Bergey‟s Manual of Systematic

Bacteriology. The morphological features, like colour and shape of bacterial colonies are not always same and get influenced by surroundings. The biochemical properties are specifically used to identify bacterial species using the software for “Probabilistic

Identification of Bacteria for Windows (PIBWin)” Ver. 1.9.2.

(http://www.som.soton.ac.uk/staff/tnb/pib.htm).

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4.5.1.3 MULTIPLE POLYSACCHARIDE DEGRADATION:

Diverse polysaccharides play structural and signaling roles in plant / algae and mammalian extracellular matrix as well as microbial biofilm. Polysaccharide degradation can be easily monitored by observing the growth of bacteria on solid medium plates containing homologous polysaccharides. However, presence of organic impurities such as proteins and other carbon compounds are very common in polysaccharide preparations and growth on polysaccharide plates would enable non-polysaccharide degraders to grow utilizing this contaminating proteins or impurities as carbon source, resulting in false positive results

Dye based plate screening methods are essentially based on the principle that polysaccharides interact non covalently with dye in the area where polysaccharide degradation occurs resulting in clearance zone in contrast to the dark polysaccharide-dye complex formation where no degradation is evident. Plate assay using chromogenic substrates are also often used to screen potential polysaccharide degrading strains

(Barasan et al., 2001). However, due to the exorbitant cost of chromogenic substrates, dye based identification is more economical and commonly used.

Congo red interacts and forms precipitates with polysaccharides containing (1, 4)-

β-D-glucans, (1, 3)-β- D-glucans, (1, 4)(1, 3)-β-D-glucans and (1, 4)-β-D-xylans. Congo red has been commonly used for the identification of cellulolytic bacteria from rumen and soil samples as well as for zymogram studies of xylanases (Teather and Wood, 1982;

Hendricks et al., 1995; Breccia et al., 1995). In the present study, congo red was used for identification of CMCase, xylanase, and chitinase activities of the strains by growing them in ASW agar-CMC, ASW agar-xylan and ASW agar-chitin plates respectively. As a

108 consequence of polysaccharide degradation by the strain, light coloured clearance zone was generated around the colonies against dark opaque background after spreading congo red. The clearance zone around the colonies could be intensified after flooding with 1 M

NaC1 solution. The clarity of the clearance zone was further enhanced by washing with

1M acetic acid (Hendricks et al., 1995). Congo red did not show any clearance zone when ASW agar alone was used, indicating the clearance zone observed were specific to degradation of cellulose, chitinase and xylanase respectively (Beguin, 1983; Samanta,

1999).

Agar degrading bacteria were detected on ASW agar plates by flooding lugol's iodine solution. This method is widely used for the detection of agarase producing bacterial strains. Development of light yellow colour around the agarolytic bacterial colony in contrast to dark brown background colour of the undegraded agar indicates loss of integrity in agar polysaccharide structure and partial degradation of agar. The failure to form the deep brown colour is due to loss of double helical structure of agar polysaccharide due to action of agarase enzyme (Hodgson and Charter, 1981).

Cetylpyridinium chloride (CPC) is a cationic polymer known to react with sulphated galactans resulting in precipitation of polysaccharides. Gacesa and Wustman

(1990) detected alginase activity by flooding CPC on solid medium plates containing alginate. Alternatively staining with ruthenium red followed by treatment with 95% alcohol, diluted HC1, diluted H2SO4 and CaC12 have been also been reported in literature for estimation of alginate lyase activity on medium alginate plates (Wong et al., 2000).

Further CPC has also been used for detection of carrageenase activity on medium plates as well as for zymogram studies of carrageenase as carrageenan contains highly sulphated

109 galactans in its structure (Ohta and Hatada, 2006). Alginolytic and carrageenolytic nature of polysaccharide degrading bacteria were detected by growing on ASW agar+ 0.2% alginate and ASW agar+ 0.2% carrageenan plates after which the plates were flooded with 10% CPC. The CPC did not precipitate ASW agar plates, indicating it specifically detects alginolytic and carrageenolytic activity.

During the present investigation, agar has been used as solidifying agent for detecting the multiple polysaccharase activities on plates containing any other polysaccharides. Gelrite, another solidifying agent is widely used as it is resistant to microbial decomposition (Shungu, et al., 1983).

Gelrite is a linear polysaccharide containing repeated units of tetrasaccharide -(3-

β-D-Glcp-(1, 4)-β-D-GlcpA-(1, 4)-β-D-Glcp-(1, 4)-α-L-Rhaρ-1) containing glucuronic acid, glucose, rhamnose and O-acetyl moieties. It forms a firm gel in divalent cations presence such as Mg+2 or Ca+2 . Similarly, approximately100 mM Na + and K+ form firm gel with gelrite (Kelco application bulletin). During all present study, ASW was used as basal medium which contain high molar concentration of Mg+2 and Na+2 ions. Addition of gelrite to ASW results in instantaneous solidification which cannot be controlled even at high temperatures. Hence gelrite could not be used for preparations of polysaccharide/s plates.

The bacterial strain DD-13 was found to be multiple polysaccharide degraders and degraded eleven different polysaccharides. It has been reported that Sacchrophagus degradens strain 2-40T, belonging to Gammaproteobacteria and related to genus

Microbulbifer, which degraded ten different complex polysaccharides (Ekborg, et al.,

2005). Kosugi et al., (2001) have reported that Clostridium cellulovorans have developed

110 enzyme systems to degrade multiple polysaccharides. When a bacterium degrade different polysaccharide to respective oligosaccharides, these oligomers enter the cell membrane into the cell and induces the expression of enzymes responsible for respective polysaccharide degradation or sometimes due to structural relatedness may cause cross induction of the unrelated polysaccharide degrading enzymes.

The strains of Microbulbifer genera are acknowledged to be a biopolymer- decomposing marine bacterium belonging to the phylum Proteobacteria. It has been reported thatM. hydrolyticus degraded Xylan, CMC and chitin (Yoon, et al., 2003a,

2003b; Miyazaki, et al., 2008; Nishijima, et al., 2009). M. salipaludis could degrade agar, xylan and CMC (Yoon, et al., 2003a, 2003b; Miyazaki, et al., 2008; Nishijima, et al.,

2009), whereas M. elongatus degraded agar, xylan, CMC and chitin (Yoon, et al., 2003b;

Miyazaki, et al., 2008; Nishijima, et al., 2009). Further M. agarilyticus degraded agar, xylan and chitin (Miyazaki, et al., 2008; Nishijima, et al., 2009) while M. celer had been reported for chitin degradation (Miyazaki, et al., 2008; Nishijima, et al., 2009). In the present study the Microbulbifer reference strains were tested for other polysaccharide degradation and the results are as shown in Table 4D.1.

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4.5.1.4 OLIGODYNAMIC EFFECT:

However a many of the metal ions are necessary for growth of bacterial cells but some can also show destructive effects. It is because the heavy metals develop complexes with enzymes that leads to inactivation. Many heavy metals at the low concentrations are unfavorable to microorganisms (Mills & Colwell 1977). There have been many reports for heavy metals adaptation by microorganisms (Azam, et al. 1977). This observable fact has significant applications for microbial ecology in polluted ecosystems

Heavy metals are generally found in bacteria which have several natural processes of anthropogenic nature. Genes related to tolerance mechanism have been found in chromosomes and plasmids. There are several mechanisms in bacteria that are responsible for heavy metal tolerance. One method involves transporting heavy metal out the cell whereas the other one employs heavy metals as terminal electron acceptors during anaerobic respiration. Zhang, et al. (2007) suggested that the lead tolerant strains are more predominant in lead contaminated soils compared to soil without lead. Lead

(Pb) is one of the most widely reported heavy toxic metals in the environment. Since it is used at the petroleum production, its abundance has been reported around the world. lead resistant bacteria could be utilized in bioremediation process for the removal of this toxic metal. It is note-worthy that, the bacterial strain DD-13 was observed to be resistant to lead (Pb) at a concentration range of 0.125- 0.25 M and thus a potential candidate for bioremediation of lead.

The strain DD-13 was found to be sensitive for nickel, zinc, copper, iron, cobalt, manganese and mercury at the concentration ranges of 0.125- 1.0 M whereas it demonstrated resistance for magnesium, potassium and lithium. The same pattern was

112 observed with the other tested reference strains of Microbulbifer, except for M. elongatus.

The bacterial cell wall have many types of cations such as Mg2+, Ca2+, Na+, and

K+, which are responsible for different bacterial activities like maintaining the integrity of the outer layers, metabolism and enzymatic actions. Mg2+ and Ca2+ ions play a significant role in stabilization of the outer structures (Ferrero, et al., 2007; Peshenko, et al., 2007).

The outer most membrane in gram-negative bacteria is made up of negatively charged lipopolysaccharide (LPS). Therefore divalent cations binds to the negative charged lipid molecules in between the adjacent LPS molecules, removing the repulsive forces and binding the LPS molecules together (Katowsky, et al., 1991; Raetz, et al., 2007).

Magnesium, potassium, lithium and calcium are present in marine water/ sediments readily available for the bacterial cells. Lithium is present in many pegmatitic minerals, but because of its solubility is found in ocean water

4.5.1.5 OTHER BIOCHEMICAL CHARACTERISTICS:

Microbulbifer species related to strain DD-13 were used for the comparison of the biochemical characteristics. The experiments were performed under the same conditions

(table 4D.1). All tested Microbulbifer species are Gram negative and rod shaped. Except for M. salipaludis and M. agarilyticus all the strains were non motile. All Microbulbifer species showed positive results for the following: oxidase and catalase activities, ornithine utilization

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Table 4D.2: Differential phenotypic characteristics of strain DD-13T and other related species of the genus Microbulbifer Strains: 1,

Microbulbifer strain DD-13T, data from this study; 2, M. hydrolyticus IRE-31T; 3, M. salipaludis JCM 11542T; 4, M. celer KCTC

12973T; 5, M. agarilyticus JAMBA3T; 6, M. elongates DSM 6810T.

(+, Positive reaction; -, negative reaction; W, weakly positive reaction; S, sensitive; R, resistant; *Data from present study, # Data obtained from respective literature.)}

Tests 1 2 3 4 5 6 Colony colour Grayish yellow Cream Grayish yellow Grayish yellow Cream Yellowish brown Motility - - + - + -

Growth at pH 5 + - -* -* - +* Growth at pH 10 + - -* +* - +*

Growth at 0% NaCl # + - - - - - characters Physiological Physiological Growth at 10% NaCl # + - + + - -

H2S production + - - - - -

Nitrate reduction - - + - + - mical Bioche Casein hydrolysis - + + + + + Alanine + + +* +* -* + Leucine - + -* +* -* +

Proline + + +* +* +* - acid Amino Serine - + - -* -* + Chitin + + - -* + + Pullulan + -* -* -* ND ND

Pectin + -* -* -* -* +*

ICP Alginate + + -* -* -* +* β Glucan W -* +* +* ND ND Laminarin + -* -* +* ND ND

Oligodynamic effect (250 Mm)* Carbohydrates ICP LiCl MnSO HgCl Pb CoSO CaCl FeCl CuCl ZnSO KCl NiCl MgSO Malonate Raffinose Salicin Sorbitol Inuline Melibiose Trehalose D Maltose Xylose Glucose Galactose CMC Xylan Starch Agar Carrageenan -

(NO Arabinose

3 2

2 2 2

4 4

)

R S S R S R S S S R S R ------+ + + + - + + + + +

+ +* W +* R R S R S R R S S R R R +* +* +* +* - +* +* - +* - + +* + * * *

+ - + W R R S R S R R S S R R R W* - - +* - - +* +* + W* + - + * * * *

+ + + - R R S R S R R S S R S R W* ------+* - - - - + * *

R R S R S R S S S R S R W* ------+ - + - - + + + +* * * * * *

S S S S S R R S S R S R - - - +* +* W* W* + + - + + + W + + +* * * *

and hydrolysis of cellobiose, , xylan, starch and agar. All the tested species of

Microbulbifer are non spore forming, and negative for α-methyl-D-mannoside, , inositol, dulcitol, urease test, sodium gluconate, methyl-α-D-glucoside, and sorbose utilization.

The strain DD-13, M. salipaludis and M. celer sustained and growth was observed up-to

10% NaCl whereas among the tested reference Microbulbifer strains only DD-13 could grow in NaCl deficiency confirming the non requirement of NaCl for the growth of strain

DD-13. The strain DD-13 was isolated from a mangrove region where the salinity (NaCl concentration) varies from 0-90 ppt, this might be the reason for the strain DD-13 to sustain its multiplication under the presence of NaCl in a range of 0-10%. Unlike strain

DD-13, all reference strain hydrolyzed casein.

4.5.1.6 CELLULAR FATTY ACID COMPOSITION (%) FOR STRAIN

DD-13 AND OTHER RELATED Microbulbifer TYPE STRAIN SPECIES.

All the other related species of Microbulbifer (Table 4.8) were analyzed for cellular fatty acid composition. The experiments were performed under the same environmental conditions in the lab. All the tested strain showed predominant of iso-C15 :

0 and C18:17c fatty acids as shown in table 4D.2.The presence of iso-C15 : 0 as the chief fatty acid is a characteristic feature of the Microbulbifer sp. (Yoon, et al., 2003, 2004,

2007; Miyazaki, et al., 2008; Nishijima, et al., 2009; Wang, et al., 2009; Baba, et al.,

2011; Zhang, et al., 2012).

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Table 4D.2: Percentage cellular fatty acid composition

(Strains: 1, Microbulbifer strain DD-13T, data from this study; 2, M. hydrolyticus IRE-

31T; 3, M. salipaludis JCM 11542T; 4, M. celer KCTC 12973T; 5, M. agarilyticus

JAMBA3T; 6, M. elongatus DSM 6810T.Fatty acids that represented < 0.5% in all strains are omitted.

Fatty acid 1 2 3 4 5 6

C10:0 1 - - 1.6 - -

C14:0 1 1 4 0.9 - - chain - C15:0 1 1 2 1.5 - 1

C16:0 14 5 17 12.6 8 5 Straight

C17:0 2 2 2 2.6 2 3

C18:0 1 1 1 0.8 2 1

iso-C11:0 2 3 1 6.7 3 4

iso-C15:0 24 23 23 21.7 23 23

anteiso-C15:0 - - - - - 2

iso-C15:1 1 2 1 0.4 1 1

iso-C16:0 - - - 0.3 - 2 Branched

iso-C17:0 9 12 7 10 17 13

anteiso-C17:0 - - - 0.3 - 4

iso-C17:19c 16 19 14 8.6 15 17

iso-C18:1 - 3 - - - -

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Fatty acid 1 2 3 4 5 6

C16:17c 5 3 9 - 2 3

C17:18c - 5 - 0.5 - 1

C18:17c 18 14 15 6.5 19 10

Unsaturated

iso-C11:03OH 3 3 2 8.9 3 3 Hydroxy

C17:0cyclo 2 2 1 6.3 1 4

C19:0cyclo 1 2 1 - 4 2 Cyclo

The strain DD-13 16S rDNA sequence was observed to be continuous stretch of

1536 bp. A comparison of 16S rDNA gene sequences present in DNA databases with strain DD-13sequence indicated that the strain DD-13 belonged to the class

Gammaproteobacteria and most strongly associated with the Microbulbifer strains. The degree of sequence similarity of strains DD-13 to the type strain of M. hydrolyticus and

M. salipaludis was 98% whereas M. celer, M. agarilyticus and M. elongatus was 97%.

The stain DD-13 formed a phylogenetic cluster and falls in the same clade with M. hydrolyticus and M. salipaludis. As the strain DD-13 is found to be closely related to the

Microbulbifer salipaludis and Microbulbifer hydrolyticus therefore these three strains including strain DD-13, were made as DNA probe for the relatedness comparison in

DNA-DNA hybridization study. Strain DD-13 showed DNA–DNA relatedness values of

28, 33, 27, 26 and 32% (Table 4.9) with respect to the reference type strains of

Microbulbifer species namely M. hydrolyticus, M. salipaludis, M. celer, M. agarilyticus

118 and M. elongatus. According to Wayne, et al., (1987) the accepted differentiation limit for species should be below 70 %. The similarity perentages of strain DD-13 are far below the accepted limit and therefore not related any of the species hence can be called as a novel species of the genus Microbulbifer.

Again the phylogenetic results are confirmed by chemotaxonomic properties of strain DD-13. The strain DD13 DNA G+C content was compared with other strains of

Microbulbifer like Microbulbifer salipaludis strain SM-1T, M. hydrolyticus DSM 11525T,

M. celer strain ISL-39T, M.agarilyticus strain JAMB-A3T, Microbulbifer elongatus DSM

6810T as shown in table 4D.3.The strain DD-13 DNA G+C content was 61.4 mol%.

Table 4D.3: Comparison of DNA G+C content of DD-13 with Microbulbifer strains.

Strains DNA G+C content

Microbulbifer Strain DD-13 61.4

Microbulbifer hydrolyticus DSM 11525T 58.2

Microbulbifer salipaludis strain SM-1T 59

Microbulbifer celer strain ISL-39T 57.7

Microbulbifer agarilyticus strain JAMB-A3T 55.2

Microbulbifer elongatus DSM 6810T. 58.2

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Therefore, on the basis of phenotypic characteristics, phylogenetic data and genomic distinctiveness, strain DD-13 proves to be a novel species o the genus

Microbulbifer for which the name Microbulbifer mangrovi sp. nov. with the type strain

DD-13 is proposed.

4.5.1.7 DESCRIPTION OF Microbulbifer mangrovi sp. nov. strain DD-13T

Based on the biochemical and chemo-taxonomical results the strain DD-13T has been classified as follows:

Domain : Bacteria

Phylum : Proteobacteria

Class : Gammaproteobacteria

Order : Alteromonadales

Family : Incertaesedis

Genus : Microbulbifer

Species : mangrovi

Strain : DD-13T

In the present study, a novel Microbulbifer species strain DD-13T isolated from mangrove sediments of North Goa, India has been described. The cells of the strain DD-

13Tare Gram-negative and non-spore forming, aerobic, non-motile, rod shaped. The rod cells are occasionally found in chains. The organism shows a rod–coccus cell growth cycle. The bacterial cells are flexible, straight, slenderial rods when young and every one gets converted to ovoid cells as the propagation stops and revert back to the original form

120 when placed to the fresh medium. Colonies on MA are slightly convex, circular, smooth, glossy, grayish yellow in colour with 2.0–3.0 mm diameter after 2 days of incubation at

30 ºC. Growth occurs at 30 to 38 ºC, but not at 4 or 40 ºC. The strain DD-13 could grow in the pH in the range of 5 to 10. NaCl is not an essential requirement for the growth and the strain grew upto 10% (w/v) NaCl. Anaerobic growth is not shown on MA. Tweens

20, 40, 60 and 80 are hydrolyzed but gelatin and casein are not hydrolyzed. Nitrate reduction and ONPG tests are negative. Positive results were obtained for catalase and oxidase tests (Table 4D.1). Negative for indole production and weakly positive for H2S production. The strain can hydrolyze multiple polysaccharides like alginate, agar, starch, chitin, carrageenan, cellulose, pullulan, pectin, laminarin, xylan and β-glucan. The cells are susceptible to gentamicin, tetracyclin, erythromycin, chloramphenicol, kanamycin, streptomycin, penicillin and neomycin. The ubiquinone Q-8 is found to be predominant

T and iso-C11:0 and iso-C15:0 are major fatty acids. The bacterial strain DD-13 has been reported to be a novel species of Microbulbifer and has been designated as Microbulbifer mangrovi sp. nov. DD-13T. The strain DD-13T deposited in the bacterial culture collections (KCTC 23483T, JCM 17729T).

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CHAPTER 5:

OPTIMIZATION OF CULTURE

CONDITIONS, FOR ENHANCING

ALGINASE PRODUCTION FROM

Microbulbifer mangrovi sp.nov. DD-13T

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This chapter describes the methodology adopted to determine the parameters required for enhancing the production of alginate lyase.

The optimized parameters will be used for growing Microbulbifer mangrovi strain

DD-13T with sole objective to maximally produce alginate lyase in culture supernatant.

The production of alginate lyase in culture supernatant will be the source for further protein purification.

MATERIALS:

Sodium alginate, alginic acid, marine broth medium, yeast extract were purchased from HiMedia Labs, Mumbai, India. All other chemicals were AR (analytical reagent) grade and double glass filtered distilled water was used.

5.1 DETERMINATION OF IDEAL CARBON SOURCES FOR GROWTH

AND PRODUCTION OF ALGINATE LYASE:

The bacterial strain DD-13T was grown in the ASW medium supplemented with one of the following as sole carbon substrates (0.2% w/v): glucose, fructose, sucrose, maltose, mannose, starch, alginate and CMC. The seed inoculum of Microbulbifer mangrovi strain DD-13Twas obtained from flask in which strain DD-13T was propagated aerobically in the ASW broth supplemented with one of the carbon substrates mentioned earlier. The flasks with the inoculated medium were incubated at 30ºC on orbital shaker

(130 rpm) for 48 h. A fraction of the culture was taken aseptically from the cultural flask at intervals of 12 h and growth was estimated spectrophotometrically at 600 nm, whereas alginate activity was quantified by DNSA method as described in later section. A

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medium flask without any inoculum was also maintained simultaneously as control during the experiment.

The carbon source which demonstrated maximum alginate lyase was selected for further studies. Growth of Microbulbifer mangrovi strain DD-13T and production of alginate lyase was evaluated in ASW medium containing different concentration of selected carbon source (0.2% to1.0% w/v).

5.2 EFFECT OF NITROGEN SOURCES FOR GROWTH AND

PRODUCTION OF ALGINATE LYASE

The effect of various nitrogen sources (0.2% w/v) such as beef extract/ yeast extract/ peptone/ ammonium nitrate/ tryptone/ urea/ casein hydrolysate/ ammonium sulfate/ cysteine/ phenylalanine or riboflavin on growth of strain DD-13T and production of alginate lyase was examined. The bacterial strain Microbulbifer mangrovi strain DD-

13T was inoculated at 0.1% (v/v) from the seed inoculums pregrown in ASW medium containing alginate supplemented with respective nitrogen source (0.2% w/v)as mentioned above. The flasks with the inoculated medium were incubated at 30 ± 2˚C on orbital shaker (130 rpm) for 48 h. A fraction of the culture was taken aseptically from the cultural flask at intervals of 12 h and growth was estimated spectrophotometrically at 600 nm, whereas alginate activity was quantified by DNSA method. A medium flask without any inoculum was also maintained simultaneously as control during the experiment.

The nitrogen source which demonstrated maximum alginate lyase was selected for further studies. Growth of Microbulbifer mangrovi strain DD-13T and production of

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alginate lyase was evaluated in ASW medium containing different concentration of

selected nitrogen source (0.05 to 0.6 % w/v).

5.3 EFFECT OF PHYSICAL PARAMETERS ON GROWTH OF BACTERIAL

STRAIN DD-13 AND ALGINATE LYASE PRODUCTION:

5.3.1 Effect of agitation on growth and alginate lyase production:

The seed inoculum was produced by growing Microbulbifer mangrovi strain DD-

13T in ASW broth containing 0.2% alginic acid for 24 h. 0.1% (v/v) from the seed

inoculum was aseptically inoculated in ASW medium supplemented with 0.2% (w/v)

alginate and kept at 30˚C in stationary condition or kept on orbital shaker (130 rpm) to

study the consequence of agitation on the alginate lyase production and bacterial growth.

A fraction of the culture was taken aseptically from the cultural flask at intervals of 8 h

and growth was estimated spectrophotometrically at 600 nm, whereas alginate activity

was quantified by DNSA method. A medium flask without any inoculum was also

maintained simultaneously as control during the experiment.

5.3.2 Effect of temperature on growth and alginate lyase activity:

0.1% (v/v) of bacterial strain DD-13T from 24 h old seed inoculums was

inoculated to ASW medium supplemented 0.2% alginate supplemented with 0.4% yeast

extract and incubated on a orbital shaker at 130 rpm and was maintained at 4 or 30 or 37

°C. At interval of 12 h, the bacterial growth was monitored by estimating the absorbance

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at 600 nm and alginate lyase activity from the culture supernatant was determined as described earlier.

5.4 EFFECT OF pH ON GROWTH AND ALGINATE LYASE ACTIVITY:

The bacteria was grown in the ASW medium [adjusted to various pH values (pH

3.0- 9.0) prepared with help of different 50 mM buffers like acetate (pH 3 and 4), citrate

(pH 5 and 6) and Tris-Cl (pH 7- 9) buffer] containing 0.2% alginate and with addition of

0.4% yeast extract. The bacterial growth and alginate lyase production was studied. The bacterial strain DD-13T was inoculated at 0.1% (v/v) from 24 h old seed inoculum into the medium and incubated on a orbital shaker at 130 rpm at room temperature. The simultaneous estimation of absorbance of the culture supernatant at 600 nm and determination of alginate lyase activity was done at the intervals of 12 h.

5.5 EFFECT OF SALT CONCENTRATION ON BACTERIAL GROWTH

AND ALGINATE LYASE ACTIVITY:

The effect of different ionic strengths (0- 8% NaCl, w/v) in ASW medium containing 0.2% alginate on bacterial growth and alginate lyase production was studied.

The bacterial strain DD-13T was inoculated at 0.1% (v/v) from 24 h old seed inoculum into the medium and incubated on a orbital shaker at 130 rpm at room temperature. The simultaneous estimation of absaorbance of the culture supernatant at 600 nm and determination of alginate lyase activity was done at the intervals of 12 h.

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5.6 EFFECT OF ASW MEDIUM AND MARINE BROTH ON GROWTH AND

ALGINATE LYASE PRODUCTION:

The optimized ASW broth containing carbon and nitrogen source was compared with marine broth (MB) with respect to the growth of strain DD-13Tand production of the alginate lyase. The ASW broth containing 0.2% alginate and 0.4% yeast extract as carbon and nitrogen source and Zobell marine broth prepared in D/W was inoculated with 0.1% of the seed inoculum of Microbulbifer mangrovi strain DD-13T. The culture flasks were incubated at 30 ºC for 48 h.A fraction of the culture was taken aseptically from the cultural flask at intervals of 12 h and growth was estimated spectrophotometrically at 600 nm, whereas alginate activity was quantified by DNSA method. A medium flask without any inoculum was also maintained simultaneously as control during the experiment.

5.7 ALGINATE LYASE ASSAY:

Alginate lyase activity (AL) was estimated using 3, 5-dinitrosalicyclic acid

(DNSA) method according to Miller, (1959). The culture supernatant was acquired by centrifuging at 10,000 rpm for 15 min at 4˚C. The assay mixture total volume of 1 ml contained, sodium alginate (0.1%, w/v prepared in 50 mM Tris HCl buffer pH 7.0) and the culture supernatant (enzyme). The reaction mixture was incubated at 30˚C for 90 min.

After incubation, 1ml of DNSA reagent was added and incubated again for 10 min in boiling water bath. The tubes were cooled and O.D. was determined at 575nm using a spectrophotometer (Shimadzu Co. Kyoto, Japan). The final O.D. was calculated after

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subtracting the O.D. of substrate and enzyme controls. The reducing sugars released were calculated with glucose as standard (0.05%). The activity was expressed as Units/ml.

One unit of the enzyme activity was defined as the amount which liberates 1 μg reducing sugar per minute under the assay conditions.

5.8 STATISTICAL METHODS

All results were represented as the means of triplicate analysis ± the standard deviation. One-way ANOVA test followed by Duncan’s multiple range tests were used to evaluate the implication of different experimental sets.

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5.9 RESULTS:

Predominantly, the polysaccharide degrading enzymes are produced extracellularly into the culture supernatant. The production of these polysaccharase is dependent on physical parameters such as temperature, pH, agitation etc. Additionally, chemical composition of the medium as well as carbon substrate in turn influences the growth of alginolytic bacteria and production of alginate lyase. Thus with an objective to purify alginate lyase from Microbulbifer mangrovi strain DD-13T, attempts were made to optimize the physical and chemical parameters that would support the optimized growth of strain DD-13T and maximize the production of alginate lyase.

5.9.1 EFFECT OF CARBON AND NITROGEN SOURCES ON PRODUCTION

OF ALGINATE LYASE

5.9.1.1 Effect of various carbon sources:

0.2% of various carbon sources like glucose, fructose, sucrose, maltose, mannose, starch, alginate, and CMC were used as carbon source for growth and alginate lyase production studies. Maximum alginate lyase production and bacterial growth was observed when 0.2% of alginate was provided as carbon source in the culture medium.

Although the strain DD-13T was found to utilize starch as carbon substrate for the growth, alginate lyase production was comparatively less when compared to the culture medium containing alginate as sole carbon source. Neither bacterial growth nor alginate lyase production was observed when sucrose, maltose and mannose were used as carbon substrate (Fig. 5.1).

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Figure 5.1: Alginate lyase production with different carbon substrate inMicrobulbifer mangrovi strain DD-13Tat 24 h. Standard deviation from triplicate experiments are represented by Bars. Different alphabet above the columns depicts significant differences

(P < 0.05).

5.9.1.2 Effect of different concentration of alginate:

Various concentrations of alginate (0.2, 0.4, 0.6, 0.8 and 1.0%) was individually added to ASW medium and propagated aerobically at 30 ± 2˚C on orbital shaker (130 rpm).The maximum alginate lyase activity and bacterial growth was observed in culture medium containing 0.2% of alginate concentration (Fig. 5.2). Alginate lyase production was maximum at 30 h. Thus 0.2% concentration of alginate was selected for further studies.

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Figure 5.2: Effect of different concentration of alginate on bacterial growth production of alginate lyase from Microbulbifer mangrovi strain DD-13Tat 30 h. Standard deviation from triplicate experiments are represented by Bars. Different alphabet above the columns depicts significant differences (P < 0.05).

5.9.1.3 Effect of various nitrogen sources on alginate lyase production:

ASW containing 0.2% alginate supplemented with yeast extract (YE), beef extract

(BE), peptone (Pep), ammonium nitrate (AN), tryptone (Try), urea (U), casein hydrolysate (CH), ammonium sulfate (AS), cysteine (Cys), phenylalanine (PA) and riboflavin (R) were used to determine bacterial growth and alginate lyase production. No

130

bacterial growth was evident in the culture medium supplemented with cysteine and phenylalanine. As seen from Fig. 5.3 maximum alginate lyase production was observed with medium amended with yeast extract followed by tryptone, urea, beef extract and ammonium sulfate. Hence yeast extract was selected as ideal nitrogen source for alginate lyase production.

Figure 5.3: Alginate lyase production with different nitrogen sources from Microbulbifer mangrovi strain DD-13T. Standard deviation from triplicate experiments is represented by

Bars. Different alphabet above the columns depicts significant differences (P < 0.05).

Yeast extract (YE), beef extract (BE), peptone (Pep), ammonium nitrate (AN), tryptone

(Try), urea (U), casein hydrolysate (CH), ammonium sulfate (AS), cysteine (Cys), phenylalanine (PA) and riboflavin (R)

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5.9.1.4 Effect of different concentration of Yeast extract on production of alginate lyase activity:

The ASW medium containing 0.2% alginate was co-supplemented with different concentrations of the yeast extract (0.05, 0.1, 0.2, 0.4 and 0.6%). The maximum alginate lyase activity and bacterial growth was observed when culture medium was supplemented with 0.4% of yeast extract after 24 hours as seen from Fig. 5.4.

Figure 5.4: Alginate lyase production with different concentration of yeast extract from

Microbulbifer mangrovi strain DD-13T at 24 h. Standard deviation from triplicate experiments is represented by Bars. Different alphabet above the columns depicts significant differences (P < 0.05).

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5.9.2 EFFECT OF TEMPERATURE, pH, IONIC STRENGTH AND

AGITATION ON GROWTH AND ALGINATE LYASE PRODUCTION:

The effect of agitation on growth and production of alginate lyase in

Microbulbifer mangrovi strain DD-13T was studied. As observed from Fig. 5.5, alginate lyase production was linked to growth curve. In static culture condition, growth of bacterial culture was initiated after 4 h of inoculation; however production of alginate lyase activity initiated only at 16 h. although strain DD-13T continued to grow after 56 h, however alginate lyase production was highest at 48 h after which there was rapid decrease in production of alginate lyase.

When the culture was grown on orbital shaker (130 rpm), the growth of strain

DD-13T as well as production of alginate lyase was initiated at 4 h. Although the bacterial growth continued beyond 48 h the highest peak production of alginate lyase was observed upto 48 h after which the production decreased rapidly.

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Figure 5.5: Alginate lyase production in Microbulbifer mangrovi strain DD-13T under static and shaking conditions.

Further the bacterial strain DD-13T was grown at 4, 30 and 37 °C to determine the optimal incubation temperature for growth and alginate lyase production. Microbulbifer mangrovi strain DD-13Twas inoculated in ASW medium containing 0.2% alginate and

0.4% yeast extract at pH 7.0 in above mentioned temperatures on orbital shaker 130 rpm.

As observed from Fig.5.6 the maximum production of alginate lyase was observed at

30˚C after 24 h incubation in comparison to alginate lyase production at 37˚C. No growth or alginate lyase activity was observed in culture medium flask kept at 4°C.

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Figure 5.6: Effect of temperature on the production of the alginate lyase from

Microbulbifer mangrovi strain DD-13T. Standard deviation from triplicate experiments is represented by Bars. Different alphabet above the columns depicts significant differences

(P < 0.05).

The bacterial strain DD-13T was grown in ASW medium adjusted to different pH

(3 to 9) with various buffers and containing 0.2% alginate and 0.4% yeast extract.

Although, the strain demonstrated the growth in medium with pH range for 5- 9, alginate lyase production was minimum at pH 9 (Fig. 5.7). The maximum production of alginate lyase was in culture medium adjusted to pH 7 where in the maximum growth of the strain

DD-13T was also depicted. 135

Figure 5.7: Effect of pH of culture medium on production of the alginate lyase from

Microbulbifer mangrovi strain DD-13Tat 24 h. Standard deviation from triplicate experiments is represented by Bars. Different alphabet above the columns depicts significant differences (P < 0.05).

The bacterial strain DD-13T was grown in presence of 0% to 8% NaCl for 36 h to determine the effect of NaCl concentration on alginate lyase production in Microbulbifer mangrovi strain DD-13T. No bacterial growth was evident in ASW medium amended with 5% and 8% NaCl. The alginate lyase activity was maximum when culture medium was amended with 1.5% NaCl concentration (Fig. 5.4). The growth and production of alginate lyase was also detected when culture medium was devoid of NaCl. 136

Figure 5.8: Effect of NaCl concentrations on culture growth and production of alginate lyase in Microbulbifer mangrovi strain DD-13T at 24 h. Standard deviation from triplicate experiments is represented by Bars. Different alphabet above the columns depicts significant differences (P < 0.05).

5.9.3 EFFECT OF THE ZOBELL MARINE BROTH V/S ASW MEDIA ON

PRODUCTION OF THE ALGINATE LYASE:

The ASW broth containing 0.4% yeast extract supplemented with 0.2% alginate was compared with Zobell marine broth for the production of the alginate lyase. As observed from Fig.5.9, the production of alginate lyase enzyme was approximately 5 fold higher in ASW medium comparison to that in Zobell marine broth after 24 h incubation.

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Thus, ASW broth containing 0.2% alginate and 0.4% yeast extract medium would be selected for large scale production of enzyme.

Figure 5.9: Effect of the growth media on production of the alginate lyase from

Microbulbifer mangrovi strain DD-13T at 24 h. Bars represent the standard deviation from triplicate experiments. Different letters above the columns indicate significant differences (P < 0.05).

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5.10 DISCUSSION:

The optimized conditions for maximum production of alginate lyase from Microbulbifer mangrovi strain DD-13T in the culture supernatant were determined. ASW medium with

0.2% alginate supplemented with 0.4% yeast extract, pH 7 incubated at 37˚C for 24 h on orbital shaker (130 rpm) were the optimal conditions for maximizing the production of alginate lyase in Microbulbifer mangrovi strain DD-13T by culture flask experiments.

5.10.1 EFFECT OF CARBON SOURCES ON ALGINATE LYASE

PRODUCTION:

With the exception of the starch and alginate (0.2%), none of the substrates (mannose, sucrose, maltose) sustained growth of strainDD-13T nor induced production of alginate lyase. Strain DD-13T was isolated from mangroves, where commonly metabolized substrates are not easily available and complex detritus materials are available. Alginate is well known substrate responsible for alginate lyase induction. Alginate lyase induction was observed in Sphingomonas sp. A1, wherein alginate induced the in vivo expression of alginate lyase which was subsequently secreted extracellularly (Murata et al., 2008;

Hashimoto, et al., 2005). The specific activity of alginate lyase was found to be reduced

15 fold when produced in the absence of alginate suggesting that alginate is a significant inducer of alginate lyase expression (Jin Yoon, et al., 2000; Doubet and Quatrano, 1982).

Aasen, et al. (1992) and Ostgaard, et al. (1993) produced Klebsiella pneumoniae alginate lyase using a semi-defined alginate medium and alginates of different compositions, respectively. But in contrast to above facts alginate was not required for the alginase expression in Azotobacter species (Kennedy, et al., 1992)

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It was reported that starch sustainably supported cell growth and stabilized pH turbulence and at the same time did not suppress enzyme production/ synthesis (Zhou, et al., 2014). In this current study, the presence of starch as the sole carbon source did not suppress the alginate lyase production, although the cell growth was low and alginase expression was comparatively less to the alginate lyase production in the culture medium with alginate as sole carbon source.

In another study with Vibrio sp. YKW-34 (Xiaoting, et al., 2008) used glucose as the only carbon and energy source but alginate lyase expression was not observed.

Further, medium lacking any carbon source or amended with CMC/fucoidan, supported the growth of Vibrio sp. YKW-34 with no production of alginate lyase. However, when

Laminaria powder and alginate were provided as carbon sources, the bacterial strain demonstrated expression of alginate lyase. Although, the glucose addition along with

Laminaria powder leads to 10-fold decrease in alginate lyase production (Alekseeva, et al.2004; Xiaoting, et al., 2008). Horn and Østgaard (2001) postulated that the substrate showing the rapid growth will be used initially compared to the substrate utilized for enzyme synthesis which will be inhibited due to catabolite repression phenomenon.

Though alginate was adequate substrate for the alginate lyase production but higher concentrations enhanced the thickness of the cultural suspension medium and affected the strain DD-13T growth in turn, reduced the production of enzyme. El-Katatny, et al. (2003) and Xiaoting, et al., (2008) reported similar effect to the present study, i.e. reduced enzyme production at higher alginate concentration. In the present study the bacterial strain DD13T could utilize 0.1-0.6% alginate for alginate lyase production. The maximum production of alginate lyase observed when 0.2% alginate was provided with

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carbon source similarly Pseudoalteromonas sp. Y-4 shows growth in 1- 4% sodium alginate and the maximum enzyme activity was shown in 2- 3% sodium alginate presence

(Hyeon-Ah, et al., 2012). Many reports on alginate-degrading bacteria describes the alginate lyase production in 0.5-1.0% alginate optimal range of (Kim, et al., 2010; Li, et al., 2011a, 2011b).

5.10.2 EFFECT OF NITROGEN SOURCES ON ALGINATE LYASE

PRODUCTION:

A small amount of specific organic compounds may be required by the bacterium for the growth as the bacterial cell is unable to synthesize them from the available nutrients. Such compounds which enhance the growth are known as growth factors however a very small amount of these growth factors are required by the bacterial cells.

For example for the synthesis of nucleic acids (DNA/RNA) pyrimidines and purines are required; vitamins are needed as coenzymes and functional groups of certain enzymes; for synthesis of proteins amino acids are used.

In the present study, the strain DD-13T did not require any vitamins or growth factors for propagation or induction of alginate lyase production. The culture medium,

ASW along with 0.2% alginate contained adequate amount of salt/ minerals to sustain growth and production of alginate lyase. Similar results of alginate lyase production was observed in case of Pseudoalteromonas sp. Y-4 (Hyeon-Ah, et al., 2012).

Few reports indicate that nitrogen sources influence the alginate lyase production by the bacterial cell. In Vibrio strain YKW-34, potassium nitrate was observed optimal sole nitrogen source for production of alginate lyase chased by yeast extract but synthesis

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of enzyme got reduced in multiple nitrogen sources the presence, although the bacterial growth was enhanced (Xiaoting, et al., 2008). In the present study, Microbulbifer mangrovi strain DD-13T, the production of alginate lyase was enhanced in the yeast extract presence followed by tryptone as the sole nitrogen source. This may be due to the fact that yeast extract is a hydrolysate of yeast which is rich in B-complex vitamins, amino acids, peptides and carbohydrates. Since they are water soluble form therefore readily assimilated for propagation and biosynthesis by bacterial cells. The yeast extract may contain a polysaccharide similar enough to alginate which acts as an inducer for production of alginate lyase by the bacterial cells (Doubet and Quatrano, 1982). Whereas tryptone is a mixture of peptides produced by the casein digestion by the protease trypsin, which is similar to casamino acids digestion. However casamino acids are formed by acid hydrolysis and typically only have few peptide chains and free amino acids, whereas tryptone is produced by incomplete enzymatic hydrolysis with some oligopeptides readily available to bacterial cells. Reports are available in support of use of tryptone for alginate lyase production with alginate as the carbon source (Dong, et al., 2012; Xiaoke, et al.,

2006).

Many reports also suggested the importance of peptone over other nitrogen sources for bacterial growth and carbohydrase production (Sugano, et al., 1995;

Alexeeva, et al., 2002). In contrast, peptone has also been referred to be unfavorable for alginase production (Xiaoting, et al., 2008). In the present study, peptone was not ideal for alginate lyase production from strain DD-13T. This can be attributed to the fact that peptone is acid hydrolysates of proteins generally milk protein and are rich in amino- acids. Metabolic consumption of amino acids is relatively a more energy consuming

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process to the bacterial cells and thus demonstrates low induction of alginate lyase source. Further, inorganic nitrogen source has also been reported to be the most advantageous for alginase production (Hu, 2004).

5.10.3 EFFECT OF AGITATION

Agitation speed during bacterial growth is a very important factor in the fermentation process as it increases the amount of dissolved oxygen in the cultivation medium. Darah and Ibrahim (1996; 1997) reported a maximum lignin peroxidation activity and maximum fungal growth when the optimal agitation speed of 150 rpm was used. However, excessive agitation are known to produce greater mechanical forces or hydrodynamic shear stresses that leads to cell destruction thus lowering the enzyme production (Porcel, et al., 2005; Darah and Ibrahim, 1996).

Papagianni, et al., (2001) also reported the effect of agitation on enzyme production. The aeration speed of the cultural broth had multiple effect on microorganisms including alteration in shape of filamentous cells, rupture of cell wall, change in the rate and efficiency of the growth and also change in the rate of synthesis of the byproducts or enzymes (Porcel, et al., 2005). Further the increased lipase production was attributed to greater transfer rate of oxygen, improved the surface area for better contact with the media and better spread of the oil substrate throughout fermentation under agitatation. Additionally optimum agitation speed of 100 rpm for strain Vibrio sp.

510-64 was required to produce extracellular alginate lyase (Xiaoke, et al., 2006).

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In the present study, the agitation or aeration was not a factor that influenced the production of alginate lyase from Microbulbifer mangrovi strain DD-13T as the later could grow and express the alginate lyase activity even under static conditions. However, the agitation of culture medium at130 rpm enhanced the growth and alginate lyase production from strain DD-13T.

5.10.4 EFFECT OF TEMPERATURE

Temperature is a critical parameter during growth and enzyme production and has to be controlled and the effect of temperature varies from organism to organism. The secretion of extracellular enzymes is influenced by temperature parameters by effecting the cell membrane.

An important indirect effect of temperature is the change in temperature is directly proportional to the affinity of enzyme systems as temperatures decreases the affinity of enzyme systems also dereases. (Zweifel, 1999; Pomeroy and Wiebe, 2001).

The activity of extracellular bacterial polysaccharases from arctic and marine sediments increased with the temperature, showing temperature optima higher than the ambient temperatures (King, 1986; Helmke and Weyland, 1991; Christian and Karl, 1995;

Arnosti, et al., 1998). It has been reported that the mesophilic and psychrotolerant marine bacterial transporters which are membrane bound, at the lower end of their optimal temperature range, shows less affinities for substrates (Wiebe, et al., 1993;

Nedwell and Rutter, 1994) Numerous studies have also described the effect of seasonal variation on extracellular enzymes temperature sensitivity (Fenner, et al., 2005; Koch, et al., 2007; Trasar-Cepeda, et al., 2007), which is because of the synthesis of different

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isoenzymes (Loveland, et al., 1994). There has been few reports for bio-geographical patterns for temperature sensitivity of the enzyme. For example, some studies showed that microbial enzymes isolated from cold environments show low temperature optima

(Huston, et al., 2000; Coker, et al., 2003; Feller, 2003).Optimum temperature for Vibrio sp. 510-64 to produce extracellular alginate lyase was 25˚C (Xiaoke, et al., 2006).

Generally for marine bacterial isolates, the optimum temperature for the alginate lyase extracellular expression varies in the range of 25 to 30˚C that corresponds to the temperature primarily in the environment (Kitamikado, et al., 1992; Xiaoke, et al., 2006;

Doubet and Quatrano, 1984). Although growth of Vibrio sp. 510-64 was enhanced at higher temperatures, it decreased the production of alginate lyase was observed

(Xiaoting, et al., 2008).

Microbulbifer mangrovi strain DD-13T was obtained from mangroves. Although, the optimal temperature for bacterial growth was observed to be 36˚C, the optimal production of alginate lyase in the culture supernatant was observed at 30˚C, which is the average temperature of water in the mangrove region of Goa.

5.10.5 EFFECT OF pH ON ALGINATE LYASE EXPRESSION:

Contrary to intracellular enzymes the extracellular enzymes are directly affected by the pH of the surroundings, as hydrogen ions modify the 3D structure of the active site of the enzyme and the amino acids ionization state (Tipton and Dixon, 1979). Deviations from the optimal pH causes decrease in enzyme activity (Chróst, 1991; King, 1986;

Münster, 1991).

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The Vibrio strain survived in a broad range of pH from 6 to 9 and secreted alginate lyase at neutral as well as alkaline medium. The acidity of the medium influence greatly the production of alginate lyase as compared to the cell growth. Various alginate lyase exhibited optimal stability and activity in the little alkalineand/ or neutral environment (Liu, et al., 2001; Kim, et al., 2010). Generally alginate lyase was expressed in the pH ranges from 5 to 9 (Hyeon-Ah, et al., 2012; Kim, et al., 2010; Xiaoting, et al.,

2008). Similarly the present study demonstrated, the growth as well as alginate lyase expression from strain DD-13T in the pH range of 5 to 9, whereas the optimum pH for the maximum production of alginase enzyme was found to be at pH 7.0.

Literature supports the requirement of neutral pH for optimal production of alginate lyase. The marine bacterium Vibrio sp. QY102 produced alginate lyase in the culture medium with initial pH 5.0 and pH 7.0 during fermentation (Zhou, et al., 2014).

The alginate lyase was also produced by a marine bacterium Bacillus subtilis in a LB medium composed of 1.0% peptone, 0.1% yeast extract, and 3.0% NaCl, pH 7.8

(Kitamikado, et al., 1992). Strain Vibrio sp. 510-64 produced extracellular alginate lyase at optimal pH 6.5 (Xiaoke, et al., 2006). Alginate lyase-producing strain AL -128 isolated from seawater of Japan and V. alginolyticus ATCC 17749 were propagated on liquid medium with pH 7.8 and containing 1.0% peptone, 0.1% yeast extract, 3.0% NaCl, and

0.5% sodium alginate (Kitamikado, et al., 1992)

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5.10.6 EFFECT OF NaCl ON ALGINATE LYASE PRODUCTION:

Not much is known on the role of salinity on microbial communities. The higher salinity decreases the extracellular enzymatic activity as more amount of energy is needed for the production of osmolites whereas comparatively less amount of energy is used for extracellular enzymes production (Hobbie, et al., 1977). However, inverse relations have also been reported (Taylor, et al., 2003; Rejmánková and Sirová, 2007).

EIAhwany and EIborai (2012) reported two factors namely, NaCl and alginate, significantly affected the alginate lyase specific activity, produced from a marine bacterium Bacillus subtilis. In many studies, NaCl has been reported as an important activator of bacterial alginate lyases (Rahman, et al., 2010; Remeo and Preston, 1986;

Min, et al., 1977). The promoting effect of high salt concentration on enzyme production may be due to the effects of charge on enzyme- alginate complex or elimination of bound water molecules from sodium alginate in the medium (Jacobson, 1955). The alginate lyase production in some bacterial strains is stimulated by the presence of high salt concentration in the growth/ culture medium which also stimulate alginate biosynthesis, which conclusively suggests that alginate production/ synthesis and alginate lyase expression are correlated (Schiller, et al., 1993). Penaloza-Vazquez et al. (1997) reported that the stimulation of alginate gene expression by the addition of NaCl is due to increased osmolarity rather than ionic effect.

In the present study, NaCl was not crucial for alginate lyase production from

Microbulbifer mangrovi strain DD-13T. The alginate lyase activity was observed in culture supernatant and was significantly similar in absence or presence of 1% NaCl in

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culture medium. However, the maximum activity was observed in the culture broth with

1.5% NaCl. ASW alginate broth amended with 5% NaCl or more inhibited the nacterial growth as well as alginate lyase expression.

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CHAPTER 6: PURIFICATION AND CHARACTERIZATION OF ALGINATE LYASE

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Alginate lyases can cleave alginate via β-elimination of the covalently linked (1-4) glycosidic bond resulting in different molecular sized oligomers with unsaturated uronic acid monomers with or without non-reducing terminal ends. Several alginate lyases have been reported from various niches. Depending on the type of substrtae recognized, alginate lyases can be classified as poly M-, poly G-, and poly MG-specific lyases.

The following chapter highlights the methodology and results obtained for isolation, purification and characterization of alginate lyase from Microbulbifer mangrovi sp.nov. DD-13T isolated from the mangroves of Goa.

MATERIALS:

Sodium alginate, alginic acid, marine broth medium, yeast extract, tris- base, Blue dextran, were purchased from HiMedia Labs Mumbai, India. Conalbumin (77 kDa)

(Sigma), ovalbumin (45 kDa) (SRL), carbonic anhydrase (29 kDa) (Sigma),

Chymotrypsinogen (25.7 kDa) (Sigma), RNase (13.7 kDa) (SRL), Acrylamide, bis- acrylamide, SDS- protein marker, protein standards, DEAE Sepharose CL-6B (FF) and

Seralose CL-6B Sigma Chemicals, St. Louis, USA. Other chemicals were of AR

(analytical reagent) grade and double glass distilled water was used.

6.1 IDEAL METHOD FOR PROTEIN CONCENTRATION:

The Microbulbifer mangrovi strain DD13T was grown in ASW broth containing

0.2% alginate and supplemented with 0.4% yeast extract at 30°C on orbital shaker (130 rpm) for 24 h for maximum production of alginate lyase. The culture supernatant was obtained by centrifugation at 10,000 rpm for 15 min at 4˚C.

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6.1.1 Ultrafiltration: The crude supernatant was passed through a 0.22 µm polysulphonate membrane filter (Advanced Micro-devices Pvt. Ltd. Ambala, India) at

4°C by applying gentle vacuum suction. The culture supernatant was 10 fold concentrated with 10 kDa membrane using ultrafiltration unit (Pellicon XL, Biomax 10,

Millipore Inc., Billerica, MA, USA). The ultrafiltration unit and membrane were washed with Milli-Q water. The filtration unit was connected to a nitrogen gas cylinder to apply pressure and membrane was equilibrated by passing 250 ml of Milli-Q water. The ultrafiltration unit was then later filled with the crude filtered culture supernatant. The permeate was collected separately whereas retentate was collected in the sample container. When the sample was 10 fold concentrated, the utrafiltration process was stopped and the concentrated sample was collected and dialyzed against 20 mM Tris Cl

(pH 7.0).

Alginate lyase activity in the dialyzed sample was measured and compared as described earlier according to Miller (1959).

6.2 PREPARATION OF ION EXCHANGE CHROMATOGRAPHY

COLUMNS:

DEAE Sepharose (Amersham Biosciences, Uppsala, Sweden) in 20% alcohol was activated by washing with 10 mM Tris Cl (pH 7.0) until the pH and conductivity values were similar to that of Tris Cl buffer. The Pharmacia XK 16/40 column was gradually packed with the DEAE Sepharose at a flow rate of 1.0 ml min-1 at 4˚C. The column was connected to the buffer reservoir and five column volumes (50 ml × 5) of 10 mM Tris CI

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(pH 7.0) was allowed to pass through the matrix at a flow rate of 0.8 ml min-1. The packed column was left at 4˚C for further use.

6.3 PREPARATION OF GEL FILTERATION CHROMATOGRAPHY

COLUMNS:

The Seralose CL-6B matrix was washed and equilibrated with 50 mM Tris CI (pH

7.0). The Pharmacia XK 26/100 column was gradually packed with the matrix at a flow rate of 0.5 ml/min at 4˚C. A column volume of 130 ml was prepared and three column volumes (130 ml × 3) of 50 mM Tris CI (pH 7.0) was allowed to pass through the matrix at a flow rate of 0.3 ml min-1.

6.4 PURIFICATION OF ALGINATE LYASE ENZYME FROM Microbulbifer mangrovi strain DD-13T.

The ultrafiltrate obtained from the ultrafiltration of culture supernatant, was subjected for further processing. The polyethyleneimine (PEI) precipitation method was used for precipitation of carbohydrates and alginate degradative products (Gegenheimer,

1990). A stock of 25% PEI was drop-wise added to ultrafiltred concentrated supernatant with stirring, till it attains a final concentration of 0.25 %.After 1 h, the precipitated impurities were removed by centrifugation (11,000 rpm, 15 min at 4˚C) and later dialyzed O/N against 10 mM Tris Cl pH 7.0 and subjected to DEAE-Sepharose column chromatography pre-equilibrated with 10 mM Tris Cl (pH 7.0). Four ml fractions were collected with the help of fraction collector (Frac- 920, GE Healthcare, Kowloon, Hong kong). The matrix was equilibrated with three bed volumes of 10 mM Tris-Cl (pH 7.0).

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The bound proteins were collected by passing buffer with with the help of 0 to 1 M NaCl linear gradient at flow rate maintained at 0.8 ml min-1. The collected fractions were checked for the presence of protein by measuring the absorbance at 280 nm. The fractions that depicted proteins existence were later assayed individually for detecting alginate lyase activity by DNSA method as described earlier (Miller, 1959). The active fractions demonstrating alginate lyase activity were pooled, concentrated by ultrafitration with a 10 kDa cut-off ultrafiltration membrane to a final volume of 1ml.

The concentrated partially purified enzyme preparation obtained from DEAE-

Sepharose chromatography was further purified by gel filtration on a Sepharose CL-6B column. 1 ml of concentrated enzyme by ultrafitration was added to a Sepharose CL-6B column and eluted with 50 mM Tris Cl buffer (pH 7.0). Two ml fractions were collected.

The absorbance of each fraction was measured at 280 nm and the fractions showing the presence of protein were assayed for alginate lyase activity by DNSA method as describe earlier (Miller, 1959). Fractions demonstrating alginate lyase activity were pooled, concentrated and used for further characterization of alginate lyase.

6.5 ALGINATE LYASE ASSAY AND PROTEIN DETERMINATION:

The purified alginate lyase activity was determined spectrometrically by Nelson method (Nelson, 1944). The reaction mixture contained 0.1% sodium alginate prepared in

50 mM citrate buffer (pH 6.0) and incubated with purified alginate lyase at 50˚C for 10 min. Total reaction volume was made to 0.7 ml. After 10 min of incubation, same amount of Nelson copper reagent (A) was added followed by heating in boiling water bath for 20 min. Later 0.35 ml of Nelson reagent (B) was added after immediate cooling, mixed for 3

152 min and incubated further for 10 min at 30˚C. The absorbance was estimated at 680 nm.

The reducing sugars released were calculated with maltose as standard (0.1%). The activity was expressed as Units/ml where one unit of the enzyme activity was defined as the amount which liberates 0.01 μmol reducing sugar per minute under the assay conditions.

The protein concentration was determined by Bradford’s method (Bradford, 1976) and Qubit® 2.0 fluorometer (InvitrogenTM, Life Technologies).

6.6 POLYACRYLAMIDE GEL ELECTROPHORESIS FOR ZYMOGRAM

STUDIES

Native and SDS-polyacrylamide (12%) gels were prepared according to the methodology mentioned (Protein Electrophoresis Technical Manual, Amersham

Biosciences, Uppsala, Sweden, 2009). Protein samples were mixed with 6X SDS-PAGE loading dye and heat treated at 100°C for 3 min before loading in duplicate on to the

SDS-Polyacrylamide gel. For native PAGE, protein samples were mixed with 6X native

PAGE buffer and loaded in duplicate on to the gel. After the end of electrophoresis, one part of the gel was subjected to coomassie staining whereas the other half of gel was washed three times with 50mM Tris-Cl (pH 7.0) for 20 min with gentle shaking. Agarose gel containing 1% sodium alginate was prepared and over-layered on the polyacrylamide gel and incubated O/N at 30˚C in a humid chamber. Alginate lyase activity was detected by spreading 5N H2SO4 to determine activity band on the polyacylamide gel.

6.7 DETERMINATION OF MOLECULAR WEIGHT:

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The purity and molecular mass of alginate lyase enzyme after the purification was revealed by sodium dodecyl sulfate (SDS) and Native polyacrylamide gel electrophoresis

(PAGE) according to Laemmli (1970).

The concentrated purified enzyme preparation obtained from gel filtration chromatography Seralose CL-6B that demonstrated alginate lyase activity, were electrophoresed in duplicates, on a 12% SDS-polyacrylamide gel and Native- PAGE.

After the electrophoresis, the gel was cut into two halves; one half of the gel was stained by coomassie brilliant blue R-250 with gentle shaking followed by methanol: acetic acid solution destaining to visualize the protein bands. The other half was used for zymogram studies as mentioned above. Medium range SDS protein molecular weight marker (6.5 to

97 kDa) (Genie Ltd, Bangalore) was used as standard protein marker during electrophoresis.

The native molecular weight was determined using gel filtration chromatography.

A range of molecular mass markers Conalbumin (77 kDa) (Sigma), ovalbumin (45 kDa)

(SRL), carbonic anhydrase (29 kDa) (Sigma), Chymotrypsinogen (25.7 kDa) (Sigma) and

RNase (13.7 kDa) (SRL) was loaded independently on the Seralose CL-6B column and the elution profile was prepared. The purified alginate lyase from Microbulbifer mangrovi strain DD-13T was also loaded separately on the column. The fraction was eluted with 50 mM Tris Cl buffer (pH 7.0). 2 ml fractions were collected with the help of fraction collector (Frac- 920, GE Healthcare, Kowloon, Hongkong). The OD280 of each fraction was measured to detect presence of protein. The molecular weight of the purified alginate lyase was obtained by plotting a graph of log of molecular weight of standard protein v/s elution volume.

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6.8 OPTIMUM CONCENTRATION OF PURIFIED ALGINASE ENZYME

FOR DETERMINING ALGINATE LYASE ACTIVITY:

0.1% of sodium alginate in 50 mM Tris Cl (pH 7.0) was incubated with different volumes of purified alginate lyase enzyme (0, 4.8, 9.6, 14.4, 19.2 and 24 µg) for 10 min at 50°C. Total reaction volume was made to 0.7 ml. After 10 min of incubation, the reducing sugar released was quantified by Nelson method as descried earlier. The reducing sugars released were estimated against maltose standard (0.05%) prepared in 50 mM Tris Cl (pH 7.0).

6.9 DETERMINATION OF OPTIMUM TIME FOR ALGINATE LYASE

ASSAY:

A fixed volume of purified alginate lyase was added to 0.1% sodium alginate in

50 mM Tris-Cl (pH 7.0) at 30°C in 10 individual test tubes and incubated at variable time intervals (5, 10, 15, 20, 25 and 30 min). Total reaction volume was made to 0.7 ml with 50 mM Tris-Cl (pH 7.0) buffer. Reducing sugars released were estimated by Nelson method as described above.

6.10 DETERMINATION OF OPTIMUM pH FOR ALGINATE LYASE

ACTIVITY:

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The alginate lyase activity of purified alginate lyase was estimated at different pH using various buffers. 50 mM citrate buffer (pH 4.0, 5.0 and 6.0), 50 mM Tris-Cl (pH 7.0 and 8.0) buffers were used for alginate lyase assay. Alginase activity was estimated in triplicates in a 0.7 ml of reaction volume containing purified enzyme and 0.1% sodium alginate, prepared in respective buffer and incubated for 10 min at 50°C. The released reducing sugars were measured by Nelson method as described above.

6.11 pH STABILITY OF ALGINATE LYASE ENZYME:

The purified alginate lyase enzyme was incubated for 60 min in various buffers adjusted to different pH such as 50 mM citrate buffer (pH 4.0, 5.0 and 6.0), 50 mM Tris-

Cl (pH 7.0 and 8.0) buffers in triplicates. After the incubation, 0.1% sodium alginate prepared in 50 mM Tris-CI (pH 7.0) was added to the reaction tubes containing the purified alginate lyase enzyme and the volume was made to 0.7 ml and incubated for 10 min at 50°C. The released reducing sugars were estimated by Nelson method as described before.

6.12 OPTIMUM TEMPERATURE FOR ALGINATE LYASE ACTIVITY:

Alginate lyase activity was measured in triplicates by incubating purified enzyme with 0.1% sodium alginate prepared in 50 mM Tris Cl (pH 7.0). Total reaction volume was made to 0.7 ml with 50 mM Tris CI (pH 7.0) buffer. The reaction mixture was incubated for 10 min at various temperatures (10, 30, 40, 50, 60 and 70°C). Reducing sugars released were quantified by Nelson method as described earlier.

6.13 THERMAL STABILITY OF ALGINATE LYASE ENZYME:

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The purified alginase enzyme was incubated at 30, 40, 50, 60 and 70°C for 60 min in triplicates. After the incubation, reaction tubes containing the enzyme were kept in ice for 10 min 0.1% sodium alginate prepared in 50 mM Tris CI (pH 7.0) was added to above enzyme fractions and the volume made to 0.7 ml with the same buffer. The reaction mixture was incubated at 50°C for 10 min. The released reducing sugars were estimated by Nelson method as described before.

6.14 EFFECT OF CATIONS OR CHEMICAL REAGENTS ON ALGINATE

LYASE ACTIVITY:

The purified enzyme was incubated with 1 mM EDTA for 1 h at 4°C. The enzyme was later dialyzed against 50 mM Tris C1 (pH 7.0) at 4°C for 24 h and was used in this study to determine the effect of various cations/ reagents. Alginase activity was measured with 0.1% sodium alginate prepared in 50 mM Tris C1 (pH 7.0) containing 2 mM of following metal salts/ reagents: CoSO4, CaC12 , MgSO4, FeSO4, ZnSO4, HgC12,

CuC12, NaCl, NiCl2, K2SO4, KCl and MnSO4. The 0.1% sodium alginate substrate amended with respective metal ions or chemical agents was incubated with purified alginase enzyme solution. The tubes were incubated at 50°C for 10 min. The released reducing sugars were estimated by Nelson assay as mentioned earlier. Alginate lyase activity observed with non- EDTA treated purified alginate lyase enzyme was considered as 100% and relative activity of EDTA treated alginate lyase and EDTA treated alginate lyase amended with metal/ reagents was determined and quantified.

6.15 PROCEDURE FOR PRODUCING URONIC ACID BLOCKS (M AND G

BLOCKS) FROM ALGINATE:

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The uronic blocks namely, guluronic acid and mannuronic acid were prepared as described by Haug, et al., (1966). 5 g sodium alginate was dispersed in 500 ml of 0.3 M

HCl and boiled at 100°C for 5 h with occasional stirring. The mixture was cooled and centrifuged. The acid hydrolysate, i.e. the supernatant after centrifugation, which contains

MG blocks, was collected and the precipitant (A) was kept for further processing. The supernatant containing MG blocks was neutralized with 5 M NaOH and centrifuged after treating with the ice cold 95% ethanol. The precipitate (B) which contains MG blocks was dialyzed and freeze-dried.

The precipitate (A) after first centrifugation, was neutralised with 5M NaOH to pH 7.0 containing both G and M block fractions, till the precipitate dissolved completely.

The alginate was adjusted to a final concentration of 1% by dilution. The pH 2.4 was attained by HCl titration. Concentrated acid was used to produce alginate concentration in the range 0.25-0.5 % at pH 2.4 and centrifuged at 4˚C for 15 min at 10,000 rpm. The precipitate (C) containing G blocks was dialysed and freeze-dried after neutralisation with 5M NaOH. The pH 1.3 was attained with 1 M HCl in the above supernatant and centrifuged. The precipitate (D) obtained containing the M block was neutralised with

5M NaOH and dialysed and lyophilized.

6.16 SUBSTRATE SPECIFICITY OF ALGINATE LYASE ENZYME

The purified alginase was incubated with guluronic acid (G) blocks or mannuronic acid (M) blocks to estimate the substrate specificity. The reaction was executed as described previously with 0.1% M or G block and the reducing sugar was quantified by Nelson method.

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6.17 TLC ANALYSIS OF ALGINATE HYDROLYSIS PRODUCTS:

Glucose, maltose raffinose and nystose were used as standards and dissolved in

50 µl of absolute ethanol. 50 µl of alginate lyase was added to 950 µl of 0.1% sodium alginate in 50 mM citrate buffer (pH 6.0). The reaction mixture was incubated for 15 min at 50°C followed by heating the reaction mixture tubes in boiling water bath for 5 min to stop the reaction. After cooling the tubes to 4°C, two volumes of chilled absolute ethanol was added. The samples were centrifuged for 10 min at 12,000 rpm at 4°C to remove the precipitate of polysaccharide. The supernatant was collected and subjected to vacuum concentration. The dried samples were reconstituted by adding 50 µl of absolute ethanol and analyzed by TLC.

Silica gel 60 sheets (Merck & Co Inc., NJ, USA) were activated at 85°C for 30 min. TLC chamber was saturated with the solvent system, n-butanol: acetic acid: water in

5: 2: 3 (v/v) ratio. The digested oligosaccharides which were concentrated earlier were spotted on TLC with the help of capillary tubes and allowed to air dry. The samples were resolved on TLC sheet, till the solvent front reaches the end of the sheet. The TLC sheet was removed from the chamber and dried in hot air oven for 15 min. The plates were sprayed with the developing reagent (Appendix) and heated at 85°C for 15 min to visualized the spots.

6.18 RESULTS

6.18.1 CONCENTRATION OF CULTURE SUPERNATANT:

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Initial experiments were conducted in which the culture supernatant from was saturated with 70% ammonium sulfate and the resultant protein precipitate collected after centrifugation was resuspended in 20 mM Tris Cl and dialyzed O/N against the same buffer. Additionaly alginate lyase activity was estimated by DNSA. However no alginase activity was observed after ammonium sulfate precipitation in comparison to the alginate lyase activity of the crude supernatant. Alternatively when culture supernatant was concentrated using lyophilization method the alginase activity decreased (1.7 fold).

However ultrafiltration method for concentrating the protein from culture supernatant increased the alginate lyase activity by 5.2 folds in comparison with the alginase activity from the culture supernatant (Fig 6.1). Alternatively culture supernatant was also lyophilized or subjected to ultrafiltration and dialyzed.

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Figure 6.1: Concentration of protein extract from the culture supernatant by different methods.

6.18.2 PURIFICATION OF ALGINATE LYASE ENZYME FROM

Microbulbifer STRAIN DD-13.

The alginase activity in the culture supernatant was determined as 3.55 Uml-1.

Protein concentration in the culture supernatant was estimated by Bradford’s method

(Bradford, 1976) and Qubit fluorometer and was quantified as 306 µg ml-1.

After the culture supernatant (1000 ml) was obtained from bacterial cells by centrifugation, and 10 fold concentrated by ultrafiltration using 10 kDa cut-off membrane at 4°C. 80 ml of the concentrated ultrafiltred enzyme obtained was stored at -20°C.

Alginate lyase activity and the protein concentration of the concentrated enzyme supernatant were estimated to be 9.13 U ml-1.and 0.57 mg ml-1 respectively (Table 6.1).

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The utrafiltred concentrated supernatant was immediately loaded and resolved on a DEAE-Sepharose FF matrix at 4°C. Elution was carried out at 0.8 ml min-1 flow rate and the bound proteins were eluted with 10mM Tris CI (pH 7.0) having 0-1 M NaCI linear gradient. 4 ml of each fractions were collected. Elution profile for proteins separated from DEAE-Sepharose FF matrix is as shown in Fig 6.3.

All the bound proteins were eluted in the salt concentration ranging from 0.125 -

0.5 M. The fractions that depicted the presence of proteins were assayed for the alginate lyase activity. The profile of protein purification depicted a single activity peak in the ion-exchange chromatogram and corresponded to 24 individual fractions. The fraction from tube number 13 to 37 (amounting to 110 ml) demonstrated alginate lyase activity

(Fig. 6.3). The fractions depicting alginate lyase activity was pooled and concentrated.

The protein concentration was determined as 62.4 µg ml-1 with specific alginate lyase activity of 2.12 U ml-1. The fractions were later concentrated to 1 ml by ultrafiltration.

The concentrated fraction demonstrating the alginate lyase activity obtained from ion exchange chromatography was further subjected to gel filtration chromatography using Sepharose CL-6B column (Fig 6.4). The bound proteins were eluted at the rate of

0.3 ml min-1 with 50 mM Tris Cl (pH 7.0). Two distinct protein peaks were depicted after measuring the O.D. of fractions at 280 nm. The protein peak that was detected at the end of the chromatogram depicted alginate lyase activity. The peak contained 12 individual fractions (tube number 17 to 29) and amounting to 63 ml was pooled and concentrated.

The protein concentration and the estimated alginate lyase activity of the pooled fraction was 0.02 mg ml-1 and 0.84 U ml-1 respectively. The pooled fraction was later concentrated and stored at -20˚C for further protein characterization.

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Protein AL activity NaCl gradient 8 0.18 Elution at 0.8ml/min 500ml of 0-1M NaCl Gradient 7 0.15 1

0.12 0.8

4 0.09 0.6

280 Abs Abs

3 Enyme activity (U/ml) Enyme 0.06 0.4 Gradient (M) NaCl

0.03 0.2 1

0 0 0 0 70 145 220 295 370 445 Volume (ml)

Figure 6.3: Alginate lyase elution profile through Ion exchange (DEAE Sepharose CL-6B)

Protein AL activity Elution at 0.4ml/min 0.7 0.14

0.6 0.12

0.5 0.1 (U/ml)

0.4 0.08

280

activity

Abs Abs 0.3 0.06 lyase

0.2 0.04 Alginate 0.1 0.02

0 0 4 27.5 51.5 76.5 101.5 126.5 151.5

Volume (ml)

Figure 6.4: Elution profile of proteins by Gel-filtration using Seralose CL-6B column

Table 6.1: Purification of alginate lyase enzyme from Microbulbifer mangrovi sp. nov. DD-13T

Total Total Total activity Specific Steps Yield (%) Fold (%) volume(ml) protein(mg) (U)* activity(U/mg)*

Crude Culture 1000 306 832.53 2.72 100 1 Supernatant

Ultra filtration 80 45.37 730.61 16.10 87.75 6

DEAE Sepharose CL- 110 6.86 233.11 33.97 28 13 6B

Seralose CL-6B 63 1.21 52.45 43.27 6.3 16

*One unit of alginate lyase activity was defined as the amount of enzyme required to release 1µM of reducing sugar at 50˚C min-1.

The purification table details depicting various parameters are as shown in Table

6.1. Alginate lyase enzyme was purified to 16 fold with a final yield of 6.5% and the final specific activity of the purified alginate lyase was observed to be 43.27 Umg-1.

6.18.3 ZYMOGRAM ANALYSIS OF ALGINASE ENZYME:

Purified alginate lyase (40 µl) was electrophoresed on the native and SDS- polyacrylamide (12%) gels. Zymogram of alginate lyase was performed empoying the overlay technique. The gel was incubated O/N at 30°C in a humid chamber. The alginate lyase activity band in polyacrylamide gel was visualized by adding 5N H2SO4 to the overlayered gel for 10 min.

Zymogram analysis on native page depicted presence of a single activity band that corresponded to the single band observed by coomassie blue staining of the native polyacrylamide gel (Fig 6.5a). No activity band was observed in zymogram analysis of

SDS-polyacrylamide gel.

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(a) (b)

Figure 6.5: Zymogram profile of purified alginate lyase from Microbulbifer mangrovi strain DD-13T on native PAGE (a) and coomassie blue staining of the native PAGE (b).

6.18.4 NATIVE AND SDS PAGE OF PURIFIED ALGINATE LYASE

FROM Microbulbifer mangrovi strain DD-13T

Purified alginate lyase was electrophoresed on a 12% of native and SDS polyacrylamide gels as per Laemmli (1970). 40 µl of the purified alginate lyase protein was loaded on both the gels separately. The purified alginate lyase from Microbulbifer mangrovi strain DD-13T showed a single band on native and SDS polyacrylamide gels as depicted in the Fig 6.6. The apparent molecular weight estimated from SDS-PAGE was

48.4 kDa. A single protein band was observed on native polyacrylamide gel indicating homogeneity of the purified alginate lyase.

167

A B M

Figure 6.6: Native and SDS- PAGE of purified alginate lyase from Microbulbifer mangrovi strain

DD-13T

A: Native –PAGE: alginate lyase from Microbulbifer mangrovi strain DD-13T.

B: SDS- PAGE: alginate lyase from Microbulbifer mangrovi strain DD-13T.

M: Molecular weight marker 14.4 to 97 kDa.

(Phosphorylase b 97 kDa; BSA 66 kDa; Ovalbumin 44.7 kDa; Carbonic anhydrase 31 kDa;

Trypsin inhibitor 20.1 kDa and α-Lactalbumin 14.4 kDa).

6.18.5 DETERMINATION OF NATIVE MOLECULAR MASS OF

ALGINATE LYASE:

The native molecular weight of purified alginate lyase was estimated using Gel filtration Seralose CL-6B column chromatography method. A range of standard molecular mass markers namely Conalbumin (77 kDa), ovalbumin (45 kDa),

Chymotrypsinogen (25.7 kDa), carbonic anhydrase (29 kDa), RNase (13.7 kDa,) were

168 loaded independently on the column. The fractions were eluted with 50 mM Tris Cl buffer (pH 7.0). 2 ml fractions were collected. The elution volume for each protein was plotted against logarithm of respective molecular weights shown in Fig 6.7. The purified alginate lyase was loaded on to the same column and eluted with similar elution parameters. On the basis of elution profile, the purified alginate lyase molecular mass was determined as 47.06 kDa (Fig. 6.7)

2.65 AgL from strain DD-13T

1.673

Figure 6.7: Determination of native molecular mass of purified alginate lyase.

169

6.18.6 CHARACTERIZATION OF PURIFIED ALGINATE LYASE:

The concentration of the purified alginase, incubation period for assay, optimum pH and stability, optimum temperature and stability, effect of various agents after EDTA treatment and substrate specificity for quantification of the reducing sugars released were estimated by Nelson and Somogyi estimation. One unit of the enzyme activity was defined as the amount which liberates 0.01 μmol reducing sugar per minute at 50°C and pH 6.0.

6.18.6.1 ALGINATE LYASE CONCENTRATION FOR ASSAY:

In order to determine an optimal concentration of alginate lyase for biochemical characterization various concentration (0, 4.8, 9.6, 14.4, 19.2 and 24 µg) of the purified alginate lyase were evaluated for the release of reducing sugar. As depicted from Fig 6.8 the purified alginate lyase activity profile showed linearity up to 80 µl corresponding to

19.2 µg of purified alginate lyase protein concentration. At higher concentration of the protein, the alginate lyase activity showed decreased activity. 30 µl of purified alginate lyase corresponding to 6 µg (early log phase), of the purified alginate lyase from

Microbulbifer mangrovi strain DD-13T was hence further selected for carrying biochemical characterization of purified alginate lyase.

170

Figure 6.8: Determination of optimum concentration of purified alginate lyase from

Microbulbifer mangrovi strain DD-13T

6.18.6.2 INCUBATION PERIOD FOR ASSAY

To determine the ideal time to carry-out alginate lyase assay, 6 µg of purified alginate lyase from Microbulbifer mangrovi strain DD-13T was incubated with the substrate at different incubation time with sodium alginate and reducing sugar released were quantified. The alginate lyase activity was observed to depict linear trend up to 15 min of incubation after which activity reached a plateau. As depicted from the Fig 6.9, 10 min was selected as an ideal incubation time for performing the alginate lyase assay.

171

Figure 6.9: Determination of optimum incubation time for purified alginate lyase activity from Microbulbifer mangrovi strain DD-13T

6.18.6.3 OPTIMUM pH AND STABILITY

Optimum pH for alginase activity was estimated using different buffer systems.

The alginate lyase activity profile at different pH as shown in Fig. 6.10. Alginate lyase activity was observed in the range of pH 4 to 7 and the enzyme showed almost no activity at pH 3 and 8. The optimum alginase activity was observed at pH 6.0. The purified alginate lyase from Microbulbifer mangrovi strain DD-13T was stable in the pH range of

5.0 - 7.0 for 1 h whereas more than 70% of the alginase activity was still retained after 1h at pH 4.0 and 7.0 (Fig 6.10).

172

Figure 6.10: Determination of optimum pH and pH stability studies of purified alginate lyase from Microbulbifer mangrovi strain DD-13T

6.18.6.4 OPTIMUM TEMPERATURE AND STABILITY

The profile of alginase activity at different temperatures is as shown in Fig.6.11.

The optimal temperature of this purified alginase was 50°C. As observed from Fig 6.11 the enzyme was partially active in the temperature range of 30 - 60°C. The alginate lyase from Microbulbifer mangrovi strain DD-13T did not show any activity at 70°C whereas very low activity was observed at 10 and 60˚C. The enzyme was observed to be 25% active at temperature below 30 ˚C.

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The enzyme stability was measured at different temperatures for 1h. The alginate lyase was observed to lose activity rapidly at 30 and 40°C and became totally inactive at

50°C at 1 h.

Figure 6.11: Determination of optimum temperature and thermal stability of the purified alginate lyase from Microbulbifer mangrovi strain DD-13T

174

6.18.6.5 EFFECT OF VARIOUS AGENTS AFTER EDTA TREATMENT:

Effects of metal ions and other reagents on the purified alginase activity is depicted in table 6.12. After 10 mM EDTA treatment, the purified alginate lyase from

Microbulbifer mangrovi strain DD-13T completely lost its activity in comparison to the control i.e. purified alginate lyase. All metal ions were tested at 2 mM concentration .

The alginate lyase activity was almost recovered in Na and Ca metal ions presence, whereas 50% activity was recovered in Ni and Zn metal ions presence. On the other hand

Mg, Fe, Mn, Hg, Co and Cu did not shown any effect on the alginase activity (Table 6.2).

0.1% SDS increased alginate lyase activity by 258%.

6.18.6.6 Substrate specificity

Fig. 6.13 shows the specificity for various substrates of the purified alginase.

When alginate as substrate was replaced with poly-guluronate or poly-mannuronate, the purified alginate lyase demonstrated activity against poly M as well as poly G. The relative activity of the purified alginase toward G and M blocks was estimated to be 115 and 25% respectively.

175

Table 6.2: Effect of various metal salts and other reagents on purified alginate lyase activity.

Reagents % Relative activity at 50˚C

Purified alginate lyase 100

EDTA 10mM 0

KCl (2 mM) 50 (± 0.6)

K2SO4 (2 mM) 91 (± 0.8)

MgSO4 (2 mM) 8.3 (± 0.1)

MnSO4.H2O (2 mM) 0

ZnSO4.7H20 (2 mM) 33 (± 0.3)

HgCl2 (2 mM) 0

NaCl (2 mM) 125 (±1.9)

FeSO4.7H2O (2 mM) 0

CoSO4.7H2O (2 mM) 0

CuCl2.2H2O (2 mM) 0

CaCl2 (2 mM) 58 (± 0.5)

NiCl2.6H2O (2 mM) 41 (± 0.3)

0.1% SDS 258 (± 2.1)

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Figure 6.12: Substrate specificity of the alginate lyase from Microbulbifer mangrovi strain DD-13T (M- mannuronic acid; G- guluronic acid).

6.18.6.7 TLC ANALYSIS OF INTERMEDIATES OF ALGINATE

HYDROLYSIS BY ALGINATE LYASE:

The hydrolysis products obtained after alginate hydrolysis by alginate lyase was resolved by TLC on silica gel 60 matrix. Four intermediates were observed as seen in figure 6.14.

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Mannuronic acid Lactone Decarboxylation product Mannuronic acid Guluronic acid

1 2 3 4 5 6 7 8 9 a b c

Figure 6.14: TLC profile of alginate oligosaccharide released during hydrolysis by alginate lyase from Microbulbifer mangrovi strain DD-13T [1: Glucose; 2:Maltose; 3: Raffinose; 4:Nystose; 5:Poly G; 6:Poly M; 7:Sodium alginate (HiMedia); 8:Alginic acid

(HiMedia); 9:Alginate Lyase + Poly G; a: Alginate Lyase + Poly M; b: Alginate Lyase + Sodium alginate; c: Alginate Lyase + alginic acid] 6.19 DISCUSSION:

The objective of a down-stream process is not only for removal of immpurities, but also to increase the desired protein concentration and give a stable and active environment to the enzyme where it is ready for application. Protein concentration and precipitation is the preliminary step in the downstream processing for purification of a protein.

Several workers successfully used solid ammonium sulfate for alginate lyase fractionation and precipitation in the range of 40-100% (Kennedy, et al., 1992;

Kitamikado, et al., 1992; Lange, et al., 1989; Davidson, et al., 1976; Cao, et al., 2007;

Wang, et al., 2006; Stevens and Levin, 1977; Doubet and Quatrano, 1984). However in this study, the alginase activity was found to be negligible or absent after ammonium sulfate precipitation. Conversely a single report by Linker and Evans (1984), reported that the loss of alginate lyase activity during ammonium sulfate precipitation and presence of 0.69 M sodium acetate during precipitation maintained the activity of alginate lyase.

Additionally lyophilizations of alginate lyase during present study indicated partial loss of alginate lyase activity with marginally increase in activity when compared to the ammonium sulfate precipitation method.

Several workers referred the use of ultrafiltration for the recovery of enzymes and other bio-molecules without altering their properties (Kim, et al., 1989; Sheu, et al.,

1987). Xiao, et al., (2007) and Xiaoke, et al., (2006) had successfully used ultrafiltration for concentration of alginate lyase during the downstream processing. Similarly, during present study, ultrafiltration using 10 kDa cut off filter successfully aided in

179 concentrating the protein from the crude culture supernatant and increased the alginate lyase activity by 5.2 folds in comparison to the alginate lyase activity in crude culture supernatant.

Polyethyleneimine (PEI) has been specifically used to eliminate carbohydrate impurities present in the crude ultrafiltered alginate lyase preparation. The carbohydrates in the supernatant, like undegraded alginate and degradative products of various molecular weight, were acidic (Romeo and Preston, 1986) and similar in charge and size to the alginate lyase affecting the purification of small protein from carbohydrates impurities.. Thus, PEI precipitation has been reported for removing the impurities like acidic carbohydrates and nucleic acids in the supernatant from plant sources

(Gegenheimer, 1990) and also for the purification of the extracelluar alginase in the culture supernatant of a bacterium from marine source (Romeo and Preston, 1986).

Further, Xiao, et al., (2007) have used PEI precipitation for the alginate lyase purification and also simultaneously reported a decrease in the yield of alginate lyase.

In this present study, the alginate lyase was successfully bound to the DEAE-

Sepharose CL-6B matrix and was recovered in presence of 0-1 M NaCl by gradient elution. All bound proteins were gradually eluted in 0- 0.5 M NaCl gradient range. The alginate lyase activity peak correlated with a protein peak. The alginate lyase was eluted from the column when the concentration of NaC1 in the buffer was approximately 0.11 -

0.38 M NaCl.

Alginate lyase purification from Azatobactor, Vibrio sp. YEW-34, Klebsiella aerogenes, Vibrio sp. 510-64 and Pseudoaltromonas citrea KMM 3297 have also been achieved using DEAE Sepharose (Kennedy, et al., 1992; Xiao, et al., 2007; Lange, et al.,

180

1989; Xiaoke , et al., 2006; Alekseeva, et al., 2004). Similarly the weak anion exchanger

DEAE cellulose, DEAE sephadex A-50, DEAE sephacel and cation exchanger CM sepharose CL-6B have also been successfully reported for the purification of alginase

(Davidson, et al., 1976; Cao, et al., 2007; Stevens and Levin, 1977; Linker and Evans

1984; Lange, et al., 1989 ). The strong anionic chromatography matrices such as SP-

Toyopearl 650 M, SP sepharose, Q sepharose FF have also been used for the purification of alginate lyase enzyme (Nibu, et al., 1995; Matsubara, et. al., 1998).

A few reports also describe the use of affinity and hydrophobic chromatography the most regular usedbeing the hexa- histidine- tagged protein purified using Ni+2 -chelate resin for one-step purification of alginate lyase production (Suda, et al., 1999). Tseng et al., (1992) had successfully reported the use of combination of hydrophobic (Phenyl sepharose CL-4B) and affinity chromatography (Blue sepharose CL-4B, Red sepharose

CL-6B and Hydroxyapatite) for the purification of alginate lyase from Vibrio sp. AL-9.

Alginate-Sepharose affinity resin chromatography has also been reported for partial purification of alginate lyase (Kennedy, et al., 1992) or alginate-epoxy resin affinity chromatography (Eftekhar and Schiller, 1994).

Gel filtration chromatography is often used as final step for the protein purification, where proteins move through the matrix based on their native molecular weight through a porous matrix beads. Purification of alginate lyase from Microbulbifer mangrovi strain DD-13T was achieved after DEAE ion exchange chromatography followed by chromatography through Seralose CL-6B. The partially purified fraction through DEAE sepharose CL-6B was fractionated on Seralose CL-6B and two protein

181 peaks were identified in the eluted fractions after gel filtration. One of the protein peaks depicted alginase activity.

Alginate lyase from Vibrio sp. YEW-34, 510-64, YWA, AL-9 (Xiao, et al., 2007;

Xiaoke, et al., 2006; Wang, et al., 2006; Tseng, et al., 1991), Bacillus sp. ATB-1015

(Xiaoke , et al., 2006) , Klebsiella aerogenes (Lange, et al., 1989), Enterobacter cloacae

(Nibu, et al., 1995), Pseudoalteromonas citrea KMM 3297 (Alikseeva, et al., 2004),

Pseudomonas aeroginosa (Linker and Evans, 1984), Sphingomonas sp. A1 (Hashimoto, et al., 2000), Alginovibrio aqualitis (Stevens and Levin, 1977), Streptomyces sp. A5 from banana rhizosphere (Cao, et al., 2007) have reportedly been purified by ion exchange followed by gel filtration.

The alginate lyase from Microbulbifer mangrovi strain DD-13T was purified to 16 fold from culture supernatant with a yield of 6.5% and a specific activity of 43.27 Umg-1.

Alginate lyase from Vibrio alginolyticus ATCC 17749 was purified to 175 fold with 1.93

U/mg specific activity whereas from Vibrio harveyi AL-128 alginate lyase was purified to 72.7 fold with specific activity of 1.89 U/mg (Kitamikado, et al., 1992); whereas alginate lyase from Vibrio sp was purified 25 fold with 55.93 U/mg specific activity

(Xiao, et al., 2007). Alginate lyase from Streptomyces sp. strain A5 was purified 14.2 fold with 101.6 U/mg specific activity (Cao, et al., 2007). Alginate lyase from

Corynbacterium sp. strain ALY-1 was purified 15 fold with 57 U/mg specific activity

(Matsubara, et al., 1998); whereas from Alginovibrio aqualitis, the alginate lyase was purified 61.29 fold with 57 U/mg specific activity (Stevens and Levin, 1977). Although there have been many characterization studies on partially purified or purified alginate

182 lyase but a few of these enzymes have been purified to high levels of purity and yield

(Sawabe, et al., 1997; Shimokawa, et al., 1997; Takeshita, et al., 1993).

The apparent molecular weight of purified alginate lyase from Microbulbifer mangrovi strain DD-13T, was determined as 48.4 kDa from SDS-PAGE studies. The native molecular weight determined by gel filteration chromatography was 47.06 kDa indicating that the purified alginate lyase from Microbulbifer mangrovi strain DD-13T is a monomer enzyme. The alginate lyase was purified to homogeneity as a single band was observed during PAGE in absence and presence of SDS. Most alginate lyase has been reported to be monomer enzymes (Nibu, et al., 1995; Nakagawa, et al., 1998) except for few bacterial alginate lyase that have been reported to consists of two subunits (Kaneko, et al., 1990).

Only few lyases have been reported to be purified to homogeneity. The molecular weight of alginate lyase ranging from 25 kDa to 110 kDa has been reported from microbial sources (Tseng, et al., 1992; Stevens and Levin, 1977). The alginate lyase from

Microbulbifer mangrovi strain DD-13T is closer in molecular weight (47 KDa) to alginate lyase reported from Vibrio alginolyticus ATCC 17749 (Kitamikado, et al., 1992).

Similarly alginate lyase from Azatobacter vinelandii and marine bacterium

Pseudomonads sp. produce alginate lyase having molecular weight of 49.4 kDa and 50 kDa respectively (Gimmestad, et al., 2009; Davidson, et al., 1976).

In the current study, in-situ activity of purified alginase was detected by native-

PAGE by overlayer method. 1% sodium alginate prepared in 50 mM Tris-Cl (pH 7.0) was overlayered onto the pre-electrophoresed polyacrylamide gel with the samples. After the electrophoresis the alginate lyase activity bands were visualized by spreading 5 N

183

H2SO4 for 10 min. The single activity band detected by H2SO4 staining correlated with the single purified protein band detected by coomassie brilliant blue staining on both native as well as SDS- polyacrylamide gel.

Alginate lyase from Microbulbifer mangrovi strain DD-13T has an optimum pH of

6.0 and lost 80% activity at pH 7.0. Azatobacter chroococcum and Pseudomonas aeuroginosa also produced alginate lyase with an optimum pH of 6 and 6.2 respectively

(Haraguchi and Kodama, 1996). The pH optimum for most alginate lyase is around neutral pH 7.0 to 8.5, for example Sphingomonas sp. at 7.5- 8.5 (Yonemoto, et al.,

1991), Pseudomonas sp. OS-ALG-9 at 7.5 (Kinoshita, et al., 1991), Pseudomonas maltophilia at 7.7 (Sutherland and Keen, 1981), Pseudomonas putida at 7.8 (Riesen,

1980), Klebsiella aerogenes at 7.0 (Lange, et al., 1989), Enterobactor cloacae M-1 at 7.8 and 7.5 (Nibu, et al., 1995; Shimokawa, et al., 1997), Azatobacter vinelandii at 7.5 and

7.2 (Davidson , et al., 1977; Kennedy, et al., 1992).

The purified alginate lyase from Microbulbifer mangrovi strain DD-13T was stable for 1 h in the pH range of 4.0 - 7.0 whereas more than 70% of the alginase activity still sustained during 1 h incubation at pH of 4.0 and 7.0. Wang, et al., (2006) had reported an alginase from Vibrio sp. that is stable in pH range of 6 – 7.5 for 2 h. Majority of purified alginate lyase are reported to be stable in alkaline pH (Nakagawa, et al., 1998;

Xiaoke , et al., 2006; Nibu, et al., 1995; Xiao, et al., 2007; Kitamikado, et al., 1992;

Tseng, et al., 1992). In contrast, the alginate lyase from Pseudomonas fluorescens was stable in the range of 5- 9 (Li, et al., 2011) whereas alginate lyase from and

Corynbacterium sp. strain ALY-1 was stable in the range of 4- 10 (Matsubara, et al.,

1998).

184

The optimum temperature of alginate lyase from strain DD-13T is 50°C. The enzyme was active in the temperature range of 30 to 60°C. The alginate lyase from

Microbulbifer mangrovi strain DD-13T completely lost its activity at 70°C whereas very low activity was observed at 10 °C. Alginate lyase from strain DD-13T retained 25% activity at 60°C. Majority of other alginate lyase have an optimum temperature range of

20- 40°C (Lange, et al., 1989; Nibu, et al., 1995; Linker and Evans, 1984; Davidson, et al., 1977; Sawabe, et al., 1997). Additionally, Matsubara et al., (1998) have reported optimum temperature of 55 °C for alginate lyase from Corynbacterium sp.

The EDTA treated and dialyzed enzyme of strain DD-13T lost 100% of activity indicating an essential requirement of metal ions. The enzyme conformation dependent on cations was presumably important for the lyase activity. Majority of the alginate lyase are dependent on metal ions for their activity. Thus, EDTA acts as an inhibitor for the alginate lyase activity as reported for alginate lyase from Bacillus sp. strain ATB-1015

(Nakagawa, et al., 1998), Pseudomonas atlantica strain AR06 (Matsushima, et al., 2009),

Sphingomonas sp. strain A-1 (Hashimoto, et al., 2000), Corynbacterium sp. strain ALY-1

(Matsubara, et al., 1998), Vibrio sp strain YEW-34 (Xiao, et al., 2007), Vibrio alginolyticus and Vibrio harveyi AL-128 (Kitamikado, et al., 1992).

In contrast, Wang, et al., (2006) and Cao, et al., (2007) reported enhancement of alginate lyase activity after EDTA treatment in Vibrio sp. strain YWA and Streptomyces sp. strain A5.

The dialyzed EDTA treated alginate lyase was further checked for the activity in the presence of various metal ions. Increase in activity was observed with Ca 2+, Mg+, Ni

2+, K+, Zn+ and Na+, which are commonly observed in marine environment. Na2+, K2+,

185

Ca2+ and Mg2+ are known to activate other alginate lyase enzymes (Tseng, et. al., 1992;

Nakagawa, et al., 1998; Matsushima, et al., 2009; Matsubara, et al., 1998; Hashimoto, et al., 2000). The effect of bivalent metal cations on the enzyme activity is closely related with their specificity towards poly M or poly G. Many of poly M specific bacterial or animal originating alginate lyases do not require Ca2+ (Wong, et al., 2000). Alginate lyase stimulating effect of Ca2+ is characteristic mainly for poly G specific alginate lyases as reported for poly-guluronate lyase from the F6 strain of the bacterium Pseudomonas sp. (Miyazaki, et al., 2001). The alginate lyase isolated from Clostridium alginolyticum was also activated in the presence of CaCl2 upto a concentration of 0.17 mM, but the further increase in Ca2+ concentration resulted in a gel formation that probably hampered the enzyme action (Saga, 1984). It is thought that Ca2+ bound to poly G blocks of alginic acid enhance the interaction of the carboxyl ion at C6 with the nucleophilic groups of the active site of the enzyme (Iwamoto, et al., 2002). The inhibiting effect of Mn2+ as well as other metal ions (Zn2+, Cd2+, Hg2+ ) have been demonstrated for many alginolytic enzymes of different specificities (Saga, 1984).

The presence of Hg2+, Cu2+, Mn+, Co+ and Fe 2+ did not increase alginate lyase activity in this study. The Hg2+, Fe2+ and Cu2+ inhibitory effect have been reported for other alginate lyase (Tseng, et al., 1992; Matsubara, et al., 1998; Cao, et al., 2007; Xiao, et al., 2007; Kitamikado, et al., 1992).

Alginate lyase from Microbulbifer strain DD-13T showed 25 % enhanced activity in NaCl presence. Nakada & Sweeney (1967) speculated that higher ionic strength can be obligatory for maintaining the minimum inter-unit distance for the enzyme to fit appropriately or organized or disrupt water molecules around alginate.

186

Alginate lyase activity from strain DD-13T was observed to increase in presence of 0.1% SDS. SDS-PAGE confirms the lack of alginate lyase quaternary structure; hence

SDS could not denature its tertiary structure (Hiramatsu and Yang, 1983). On the other hand, SDS has been reported to enhance the activity by 158%, demonstrating the significance of β-strands structure as SDS at low concentration has been found to stabilize β-strands (Zhong and Johnson, 1992). At very less concentrations, SDS molecules do not affect the protein core by binding at the outer surface whereas higher

SDS concentrations inactivate enzyme by changing the conformation. (Muga, et. al.,

1993).

Alginate lyases enzyme ha been classified, based on their substrate specificities, as EC 4.2.2.11, poly (G) lyase [(1-4)-L-guluronan lyase] or EC 4.2.2.3, poly (M) lyase

[(1-4)-D-mannuronan lyase]. Specificity for substrate mainly depends on the surrounding availability of the type of substrate in which the organism is found and produces the enzyme. Alginase are divided into 3 substrate specificities; mannuronate lyase (Romeo and Preston, 1986; Linker and Evans, et al., 1984), guluronate lyase (Haugen, et. al.,

1990; Lange, et al., 1989; Miyazaki, et al., 2001; Matsubara, et al., 1998; Shimokawa, et al., 1997) and dual M and G specific lyase (Iwamoto, et al., 2001; Alekseeva, et al.,

2004; Nakagawa, et al., 1998; Xiao, et al., 2007; Sawabe, et al., 1997). Most of the reported lyases studied are specefic for poly (M) substrate, however a a small number of

G-specific lyases have also been characterized. However a lyase when named as G or M specific, it usually shows low activity for the other homopolymer for example G-specific lyases mainly degrade poly (G-M) and poly (G) alginate and also shows less activity

187 towards poly (M). It is usually difficult to find out whether MG /G-M or both bonds are degraded.

The purified alginate lyase from Microbulbifer mangrovi strain DD-13T could degrade predominately poly G compared to poly M. It has been reported that endo poly G lyase also mildly attack poly M blocks. One of the reasons could be that the poly M preparation might have a significant proportion of G blocks as contamination. Another explanation could be that the alginate lyase preparation cleaved G-G linkage as well as

G-M linkages from the four glycosidic linkages present as alginate constituent (G-G, G-

M, M-M and M-G) (Haugen, et al., 1990). This explains the low activity of purified alginate lyase towards poly M preparation. Nibu, et al., (1995) had reported similar findings with alginate lyase in Enterobacter cloacae.

The most of enzymes reported degrades long chains into short hains or oligomers demonstrating endo-cleaving activity,. However, there are a few reports for exolytic enzymes. (Brown and Preston, 1991; Doubet and Quatrano, 1982; Doubet and Quatrano,

1984; Nakada and Sweeny, 1967; Schaumann and Weide, 1990), removing which removes dimers or monomers from the ends of polymers.

In the present study alginate oligosaccharides have been routinely separated by

TLC and detected with developing reagent. After incubation for 15 min, the observed four bands were identified as tri, tetra, penta and hexa uronides explaining the alginate lyase enzyme indicating endolytic nature of alginate lyase from Microbulbifer mangrovi sp. nov. DD-13T. Ando and Inoue (1961) reported similar results with an alginolytic enzyme produced by a Vibrio sp. Alginate degradation by G-lyase isolated from Vibrio sp. 510 generated a high content of di- to pentasaccharides, that was similar to that

188 observed in case of experiments related to G-lyase from Corynebacterium sp. ALY-1

(Zhang, et al., 2004, Matsubara, et al., 1998).The production of triuronides and diuronides by alginolytic enzymes of bacterial origin was also reported by Yoshikawa

(1961). Preiss and Ashwell (1962) isolated a partially purified alginate lyase from a

Pseudomonad that produced the hexuronic acid as the degradative product. More than one enzyme was found to be involved in the complete degradation of alginate. Ando and

Inoue (1961) reported that both intracellular and extracellular alginolytic enzymes from an Aeromonad were required to yield a monouronide from alginate.

189

CHAPTER 7:

APPLICATIONS OF ALGINATE

LYASE

190

Polysaccharase enzymes from bacterial sources have been widely exploited in diverse biotechnological applications related to pharmaceuticals, agriculture, textile and aquaculture industries etc. This chapter reports the methodology and results obtained on studying applications related to alginate lyase from Microbulbifer mangrovi strain

DD13T.

This chapter deals with the action of alginate lyase of Microbulbifer mangrovi strain DD-13T on alginate polysaccharide to produce alginate oligomers with the potential of being used as possible nutraceuticals. In the present study, concentrated culture supernatant of alginate lyase obtained by harvesting Microbulbifer mangrovi strain

DD13T in ASW broth containing alginate and yeast extract for 24 h that had revealed the presence of 48 kDa protein was used. The chapter provides evidence to demonstrate that the alginate oligosaccharides produced by enzymatic action have several antioxidative activities. Further the Microbulbifer mangrovi strain DD-13T was employed to produce single cell detritus from seaweeds and possibly can be used as potential biofeed material in aquaculture farm as well as its application in bioremediation especially for algal waste treatment.

MATERIALS:

Sodium alginate, agar (purified, bacteriological), potassium ferricyanide, trichloroacetic acid (TCA), ferric chloride, ferrous sulphate, heptahydrate, sodium salicylate, were obtained from HiMedia Labs, India. All other chemicals were AR

(analytical reagent) grade and double glass distilled water was used.

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7.1 PREPARATION OF ALGINATE LYASE EXTRACT FROM

Microbulbifer mangrovi sp.nov. DD-13T:

Microbulbifer mangrovi sp.nov. DD-13T was grown in ASW agar – alginate plates for 48 h at 30°C. A loopful of colonies were inoculated into 25 ml of ASW medium with 0.2% alginate and 0.4% yeast extract. The medium was incubated on an orbital shaker for 24 h at 30°C at 130 rpm. 0.1% of inoculum from the above flask was mixed to 100 ml of ASW medium having 0.2% alginate and 0.4% yeast extract. The culture was incubated on an orbital shaker for 24 h at 30°C at 130 rpm. The culture supernatant (obtained after centrifugation, as described previously) was concentrated by

10 kDa ultrafiltration unit and dialyzed against 50 mM Tris-C1 (pH 7.0) to 10 ml. The concentrated culture supernatant was stored at -20°C until further processing. A small fraction of the culture supernatant and concentrated culture supernatant was used for determining alginate lyase activity by DNSA method and the concentration of protein was estimated by Folin-Lowry method as described in the previous chapters.

7.2 PREPARATION OF ALGINATE OLIGOSACCHARIDES:

A stock of 0.5% sodium alginate in 50 mM citrate buffer (pH 6.0) was used as substrate for alginate lyase. 10 µl ml-1 of concentrated alginate lyase preparation was mixed to 50 ml of 0.5% sodium alginate stock solution and incubated at 50°C either for

3, 6 or 9 h with occasional mixing. The digestion was ceased by boiling the reaction mixture for 10 min. The sample was cooled to 4°C. The polysaccharide precipitate was removed by adding chilled ethanol followed by centrifugation for 15 min at 4°C at

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12,000 rpm. The supernatant was collected and concentrated using rotary evaporator

(Equitron Roteva 63, Mumbai, India) at 60°C and the concentrated sample was stored at -

80°C. The frozen oligosaccharide preparation was lyophilized and later resuspended in 50 mM citrate buffer (pH 6.0) followed by overnight dialyzed against the same buffer. The concentration of reducing alginate oligosaccharides was estimated by DNSA method with glucose as reference sugar as described earlier. Alginate oligosaccharides (50 and 100

µg) obtained by enzyme hydrolysis for 3, 6 and 9 h hydrolysis experiments were used for determining the antioxidant activities. Sodium alginate (0.5%) prepared in 50 mM citrate buffer (pH 6.0) was used as control for all the experiments.

7.3 TOTAL PHENOLIC CONTENT OF ALGINATE-OLIGOSACCHARIDES:

The total phenolic constituent of alginate oligosaccharides was determined using

Folin-Ciocalteu reagent (Slinkard and Singleton, 1977). Gallic acid was used as standard.

Alginate oligosaccharides at a concentration of 50 and 100 µg, were taken in a test tube,

1.58 ml d/w and 0.1 ml Folin-Ciocalteu reagent were mixed and vortexed . After incubation of 3 min , 0.3 ml of 20% Na2CO3 was followed by 30 min incubation with intermittent shaking at 25˚C. Optical density was estimated at 760 nm. The same protocol was followed for gallic acid standard (0.05%) and the standard curve obtained was used for determining total phenolic content in alginate oligosaccharide preparations.

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7.4 REDUCING POWER OF ALGINATE-OLIGOSACCHARIDES:

The reducing power of alginate oligosaccharides was estimated according to Wu et al, (2004). Alginate oligosaccharides (50 and 100 µg) were mixed to 1 ml of 0.2 M sodium phosphate buffer (pH 6.6). 1 ml of potassium ferricyanide (1%) was added. The reaction mixture was incubated for 20 min at 50°C in a water bath. 1 ml of 10% TCA was added and mixed gently. The samples were later centrifuged for 10 min at 3000 rpm and the upper aqueous layer was seperated. l ml of 0.1% FeCl3 diluted with 1 ml of d/w was added. The samples were mixed gently and O.D. was determined at A700. Ascorbic acid

(0.01%) was taken as positive control. The reducing power of agar oligosaccharides was expressed as µg/ml ascorbic acid equivalents.

7.5 HYDROXYL RADICAL SCAVENGING ACTIVITY OF

ALGINATE-OLIGOSACCHARIDES:

Hydroxyl radical scavenging activity was measured as described by Wang et al,

(2004). Alginate oligosaccharides (50 and 100 µg) were added separately to 3 ml of hydroxyl radical generation buffer (Appendix) followed by addition of 200 µl of H202.

The mixture was incubated for 10 min at 37°C and O.D510 was determined. Negative control was prepared by replacing H2O2 with sodium phosphate buffer. Positive control was prepared by replacing oligosaccharides with buffer (control group).

The antioxidant activities of the samples were calculated according to the inhibition percentage of free radical production as:

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Inhibition rate (%) = (O.D. of control group — O.D of test group) x 100

O.D. of control group

7.6 NITRIC OXIDE SCAVENGING ACTIVITY OF ALGINATE

OLIGOSACCHARIDES:

Nitric oxide produced from sodium nitroprusside was estimated by Greiss reaction and detrmined by spectrophotometric assay (Sumanont, et al., 2004). Sodium- nitroprusside (360 µl) and 216 µl Greiss reagent (2% H3PO4, 1% sulfanilamide and 0.1% napthylethylenediamine dihydrochloride) was added to 320 µl alginate oligosaccharides

(50 and 100 µg) and vortexed. The mixture was later incubated for 1 h at RT. Finally 2 ml of distilled water was added and vortexed for 2 min. The O.D. was taken at 546 nm.

Ascorbic acid (0.01%) was taken as positive control. The experiments were performed in triplicates. The nitric oxide scavenging activities of the samples were calculated according to the inhibition percentage of free radical production as:

Inhibition rate (%) = (O.D. of control group — O.D of test group) x 100

O.D. of control group

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7.7 METAL CHELATING ACTIVITY OF ALGINATE

OLIGOSACCHARIDES:

The chelating activity of alginate oligosaccharide, for Fe2+ ions was determined by the protocol of Dinis et al., (1994). To 0.25 ml of alginate oligosaccharides (50 and

100 µg), 1ml of distilled water with 0.05 ml of 2 mM FeCl2 was added. Vortexed and incubated for 30 s followed by addition of 0.1 ml ferrozine was (5 mM). A water soluble magenta colour stable complex species is formed when Ferrozine reacts with the divalent ferrous ion. After the incubation at 30°C for 10 min, the OD562 was determined. The experiments were performed in triplicate and % of scavenging activity was calculated as

Chelating rate (%) = (O.D. of control group — O.D of test group) x 100

O.D. of control group

7.8 ANTIOXIDANT ACTIVITY USING LINOLEIC ACID

PEROXIDATION METHOD:

Antioxidant activity of the oligosaccharides obtained from enzymatic hydrolysis of commercial alginate, was determined according to Yen and Hsieh (1998).

Linoleic acid (0.28 g) and 0.28 g of Tween- 40 was homogenized in 50 ml of 0.2 M phosphate buffer (pH 7.0) to prepare emulsion of.linoleic acid To 0.5 ml of alginate oligosaccharides (50 and 100 µg) linoleic acid emulsion (2.5 ml) and 0.2 M phosphate buffer (2.5 ml pH 7.0) were added and kept in dark at 40˚C for 24 h. The mixture without alginate oligosaccharides was taken as control. Aliquots of 0.1 ml were withdrawn from

195 incubated mixture and mixed with 0.1ml of 20 mM ferrous chloride in 3.5% HCl, 4.7 ml of ethanol (75%) and 0.1 ml of potassium thiocyanate (30%) followed by incubation at

30°C for 3 min. The optial density of the omplex formed was determined at A500 nm. The linoleic acid peroxidation was estimated as mentioned below (Pin-Der Duh, 1998). α-

Tocopherol (Vitamin E) was taken as standard antioxidant. The analyses were done in triplicates.

LPI (%) = (1 - A1- A2) x 100

LPI = Linoleic acid peroxidation Inhibition

A1 = Absorbance of control (without sample)

A2 = Absorbance of test sample

A0 = Absorbance of control without KSCN

7.9 STATISTICAL ANALYSIS:

The results have been calculated as the means of triplicate analysis ± the standard deviation. A one-way ANOVA test followed by Duncan’s multiple range tests were used to evaluate the significance of various experimental sets.

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7.10 PREPARATION OF SINGLE CELL DETRITUS FROM Sargassum tenerrimum USING Microbulbifer mangrovi sp.nov. DD-13T:

This study was intended to check the potential of Microbulbifer mangrovi sp.nov.

DD-13T for preparation of single cell detritus from Sargassum tenerrimum collected from coast of Anjuna, Goa, in the month of November 2011.

The dried thalli of Sargassum tenerrimum were surface sterilized with 80% ethanol for 8 min and washed with sterile sea water thoroughly. Thalli (1g) were aseptically cut into 3-4 mm pieces. The sterile ASW broth was inoculated with sterilized pieces of Sargassum tenerrimum. Thalli medium was inoculated with the loop-full of

Microbulbifer mangrovi sp.nov. DD-13T pre-grown on ZMA plates and incubated for 48 h at 30˚C on orbital shaker. Each after 8 h of incubation cells released from thalli were observed under compound microscope at 40 X resolution.

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7.11 RESULTS:

The applications studies related to alginate lyase have been divided into two groups, the use of alginate oligosaccharides obtained from digestion of sodium alginate using alginate lyase from Microbulbifer mangrovi strain DD-13T as antioxidants with potential as nutraceuticals in food industries whereas applications involves the use of whole cells of Microbulbifer mangrovi strain DD-13T to degrade multiple polysaccharides from seaweeds to generate SCDs (Single Cell Detritus) that can be used in the field of aquaculture industries.

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7.11.1 ANTIOXIDATIVE PROPERTIES OF ALGINATE

OLIGOSACCHARIDES OBTAINED FROM DIGESTION OF ALGINATE BY

ALGINATE LYASE OF Microbulbifer mangrovi strain DD-13T

7.11.1.1 TOTAL PHENOLIC CONTENT OF ALGINATE-

OLIGOSACCHARIDES:

Alginate oligosaccharides (50 and 100 µg) obtained at different period of hydrolysis (3, 6 and 9 h) were analyzed for total phenolic contents. The total phenolic content was highest in the sodium alginate sample that was digested for 3 h as compared to the phenolic content found in 6 and 9 h samples (Fig. 7.1). 0.5% of undigested sodium alginate did not depict presence of phenolic content.

7.11.1.2 REDUCING POWER OF ALGINATE-OLIGOSACCHARIDES:

50 or 100 µg of alginate oligosaccharides obtained by 3 h digestion of sodium alginate demonstrated more reducing power in comparison to the control. However, attenuation of reducing power was observed for 50 or 100 µg of alginate oligosaccharides obtained either by 6 or 9 h digestion of sodium alginate when compared to the reducing power of oligosaccharides obtained by 3 h (Fig. 7.2). 100 µg of 0.5% undigested sodium alginate did not demonstrate any reducing power when compared to the digested oligosaccharides.

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Figure 7.1: Total phenolic content of alginate oligosaccharides obtained after digestion of sodium alginate using alginate lyase of M. mangrovi sp. nov. DD-13T. Bars represent the standard deviation from triplicate experiments. Different letters above the columns indicate significant differences (P < 0.05).

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Figure 7.2: Reducing power of alginate oligosaccharides prepared by digestion of sodium alginate for 3, 6 and 9 h using alginate lyase from strain DD-13T. Bars represent the standard deviation from triplicate experiments. Different letters above the columns indicate significant differences (P < 0.05).

7.11.1.3 HYDROXYL RADICAL SCAVENGING ACTIVITY OF

ALGINATE OLIGOSACCHARIDES:

50 and 100 µg of alginate oligosaccharides obtained by 3 and 6 h digestion of sodium alginate showed similar hydroxyl radical scavenging activities that differed

201 significantly that of 9 h digestion of sodium alginate (Table 7.3). 0.5% undigested sodium alginate did not depict any hydroxyl radical scavenging activity.

Figure 7.3: Hydroxyl radical scavenging activity of alginate oligosaccharides prepared by digestion of sodium alginate using alginate lyase. Bars represent the standard deviation from triplicate experiments. Different letters above the columns indicate significant differences (P < 0.05).

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7.11.1.4 NITRIC OXIDE SCAVENGING ACTIVITY OF ALGINATE

OLIGOSACCHARIDES:

Nitric oxide scavenging activities of the samples were calculated according to the inhibition percentage of free radical production. 50 and 100 µg of alginate oligosaccharides obtained during 3 h digestion of sodium alginate demonstrated higher nitric oxide scavenging activity when compared to alginate oligosaccharides obtained from 6 and 9 h hydrolysis experiments (Fig. 7.4). The nitric oxide scavenging activity of undigested sodium alginate was negligible.

Figure 7.4: Nitric oxide scavenging activity of alginate oligosaccharides prepared by digestion of sodium alginate using alginate lyase. Bars represent the standard deviation from triplicate experiments. Different letters above the columns indicate significant differences (P < 0.05).

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7.11.1.5 METAL CHELATING ACTIVITY OF ALGINATE

OLIGOSACCHARIDES:

100 µg of alginate oligosaccharides from the 9 h hydrolysis experiments depicted the highest chelating activity towards ferrous ions in comparison to 50 µg oligosaccharides obtained from alginate. 100 µg 05% undigested sodium alginate depicted only 4% chelating activity

Figure 7.5: Ferrous ion chelating activity of alginate oligosaccharides prepared by digestion of sodium alginate using alginate lyase. Bars represent the standard deviation from triplicate experiments. Different letters above the columns indicate significant differences (P < 0.05).

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7.11.1.6 ANTIOXIDANT ACTIVITY USING LINOLEIC ACID

PEROXIDATION METHOD:

The undigested sodium alginate demonstrated 65% of peroxidation activity whereas 50 and 100 µg of alginate oligosaccharides obtained by 3 and 9 h digestion demonstrated above 90% of linoleic acid peroxidation (Table 7.6)

Figure 7.5: Lenoleic acid peroxidation activity of alginate oligosaccharides prepared by digestion of sodium alginate using alginate lyase. Bars represent the standard deviation from triplicate experiments. Different letters above the columns indicate significant differences (P < 0.05).

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7.11.2 PREPARATION OF SINGLE CELL DETRITUS FROM Sargassum tenerrimum USING Microbulbifer mangrovi sp.nov. DD-13T:

This study was intended to check the potential of whole cells of Microbulbifer mangrovi sp.nov. DD-13T for preparation of SCD from Sargassum tenerrimum. When observed under microscope, thalli degradation was evident from release of algal cells in the medium. It was also observed that release of algal cells left an empty mesh like structure in the thalli surface. Single cells detritus were observed as depicted from Fig.

7.7. When the control thalli sample was incubated under similar conditions without any inoculated culture, the surface of the thalli surface was intact and no SCD was observed

(Fig. 7.7).

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Figure 7.7: Microscopic observation of the seaweed thalli degradation by M. mangrovi strain DD-13T and release of irregular shaped cells from thalli of Sargassum tenerrimum.

7.12 DISCUSSION:

Oligosaccharides are intermediate carbohydrates of simple sugars and polysaccharides with low molecular weight. They can be directly extracted from natural sources, or can be chemically synthesized by hydrolysis. involving combination of disaccharides/ monosaccharides. The biodegradable, non-toxic and non-allergenic properties of oligosaccharides are of great scientific interest. Further the high-solvency and low-viscosity of oligosaccharide at pH 7.0 increased their applicability in various fields of Biotechnology. Recent advances report the use of oligosaccharides to enhance immunity for anticoagulation and as anti-tumor agents (Yuan, et al., 2006; Volpi and

Maccari, 2009; Zhao, et al., 2006; Fujihara and Nagumo 1992; Fujihara and Nagumo

1993). Oligosaccharides have also been reported to lower blood glucose, blood cholesterol and blood pressure (Fitton, et al., 2008; Hou, et al., 2009) as well as protect against peroxide and microbial infection (Sudarshan, et al., 1992; Alekshun and Levy,

2004; Xie, et al., 2001). Oligosaccharides also play an important role in plant systems and have been used as fertilizers, growth promoters as well as neuronal development, embryogenesis, metastasis and cell proliferation in animal system (Pospieszny, et al.,

1991; Yonemoto and Yamaguchi, 1992; Tomoda and Umemura, 1994).

In the present study, alginate oligosaccharides obtained by enzymatic hydrolysis demonstrated various antioxidant activities. An antioxidant is a molecule that inhibits free radical formation by interfering in a biochemical reaction. Although oxidation reactions

207 are crucial for life, the free radicals formed start chain reaction in cell leading to apoptosis or damage of the cell. Antioxidants being reducing agents remove free radical intermediates and stop these chain reactions or inhibit other oxidation reactions. Higher organisms like plants and animals are the examples for maintaining complex systems of numerous kinds of antioxidants. Inhibition of the antioxidant enzymes or insufficient levels can cause oxidative stress leading to destruction or apoptosis of cells in the organisms. Oxidative stress plays a significant role in many human diseases, including cancers. For these reasons, oxidative stresses are considered to be the cause as well as the consequence of various diseases.

Antioxidants have been explored for the treatment of diseases like coronary heart disease, cancer, acute sickness and hence are widely used as dietary supplements (Jha, et al., 1995; Baillie, et al., 2009; Bjelakovic, et al., 2007). Antioxidants have many other industrial uses, and have been employed asstabilizers in food, cosmetics and also prevents the gasoline and rubber degradation (Dabelstein, et al., 2007).

There have been many reports for identifying antioxidant activity of oligosaccharides from marine origin using various mechanisms including superoxide and hydroxyl radical scavenging, anti-lipid peroxidation, metal chelating and erythrocyte hemolysis inhibiting ativities (Kochkina and Chirkov, 2000a).

Phenolic compounds from plants or seaweeds such as flavonoids, phenolic acids, tannins etc., offer nutritional and quality value in terms of health, aroma, colour, flavor as well as taste benefits. They are important component of plant defense mechanism and prevent molecular damage to microorganisms and insects by counteracting reactive oxygen species (Vaya, et al., 1997). The phenolic compounds are considered to be

208 possessing antioxidant activities and reported to perform diverse natal activities such as anti-inflammatory, anti-apoptosis, anti-artherosclerotic, anti-ageing and anti-carcinogenic activities (Chung, et al., 1998; Han, et al., 2007). Due to antioxidant and chemoprotective activity of phenolic compounds, they are also referred as food additives. (Bravo, 1998;

Yingming, et al., 2004). The action is mainly because of the redox properties of these phenolic compounds, which play a significant role in decomposing peroxides, neutralizing and adsorbing free radicals or quenching singlet and triplet oxygen. (Louli, et al., 2004; Evas, et al., 2002).

Alginate oligosaccharides obtained by water / methanol extraction from marine organisms or acid/enzyme hydrolysis of polysaccharides produce different types of oligosaccharides with varying biological activities. In this study, the total phenolic content of digested commercially available sodium alginate preparation was compared with the undigested commercially available sodium alginate (HiMedia). In the present study, alginate lyase release hexa-, penta- or tetra- oligosaccharides (low DP) from sodium alginate by enzymatic action (Chapter IV).

The total phenolic content of digested sodium alginate was observed to be much higher whereas the undigested sodium alginate did not depict any phenolic content. This could be because the enzymatic treatment resulted in production of low molecular weight oligosaccharides exposing a large number of phenolic groups in the extract when compared to the non-hydrolyzed samples of sodium alginate polymers.

The most common method for the estimation for phenolic contents is based on spectroscopic method using Folin-Ciocalteau reagent (Slinkard and Singleton, 1977). The

Folin- Ciocalteau reagent is not specific and detects all phenolic groups. The Folin

209 reagent i.e. phosphomolybdic- phosphotungstic acid reduces to form a blue coloured complex in an alkaline solution that is measured spectro-photometrically.

The antioxidant activities of alginate oligosaccharides were assessed by estimating their reducing power, radical scavenging activities, metal chelating activities and linoleic acid peroxidation. During the present study, water soluble alginate oligosaccharides were obtained by hydrolysis of commercial sodium alginate by alginate lyase from Microbulbifer mangrovi strain DD-13T treatment. The oligosaccharides obtained demonstrated antioxidants activities such as reducing activity, metal chelating activity, hydroxyl and nitric acid radical scavenging activity and linoleic acid peroxidation inhibition.

The reducing activities of alginate oligosaccharides obtained by enzymatic hydrolysis serve as an indicator of potential anti-oxidant. The reducing activity was mainly due to transformation of Fe 3+ to Fe 2+ by oligosaccharides. In the presence of alginate oligosaccharides, potassium ferricyanide (Fe3+) is reduced to potassium ferrocyanide (Fe2+), which after reacting with ferric chloride forms ferric ferrous complex compound with maximum absorption at 700 nm.

In the present study, alginate oligosaccharides obtained by digestion for 3 h depicted 56.12 mg/g equivalents of ascorbic acid whereas 6 and 9 h oligosaccharides only

36.93 and 28.3 mg/g equivalents of ascorbic acid. Thus high molecular weight of oligosaccharides formed by 3 h hydrolysis depicted higher reducing power compared to low molecular weight oligosaccharides produced during 6 and 9 h hydrolysis. Hydrolysis of alginate also reported for reducing power of oligosaccharides that were obtained from

210 alginate hydrolysis by alginate lyase action (Wang et al., 2007). Rupérez et al., (2002) also reported reducing power in polymeric alginate-rich fractions of polysaccharides.

Hydroxyl radicals have very short life span but when they are generated, they hydroxylate nucleic acid, proteins and lipids. Hydrogen peroxide itself is not a free radical, however become harmful by crossing the biological membranes of the cells.

2+ 3+ - Fenton reaction (Fe +H202--*Fe +0H +OH*) synthesize the most reactive hydroxyl radical in the cells (Pryor, 1986; Wang, et al, 2007). These hydroxyl radicals attack biological molecules with a life time of 10 -7 seconds and mean diffusion distance of 4.5 nm (Lesser 2006).

In the present study, hydrogen peroxide promotes the formation of hydroxyl radicals in the reaction mixture. The alginate oligosaccharides digested for 3 and 6 h exhibited approximately 77-86 % of hydroxyl radical scavenging activity whereas the 9 h digested sample was much lower. The nonhydrolyzed alginate did not show any scavenging activity. Augmentation of hydroxyl radical inhibition by 3 and 6 h oligosaccharides is probably due to production of oligosaccharides with high DP and high molecular weight in comparison to the oligomers obtained during 9 h hydrolysis. No hydroxyl radical inhibition was associated with the non-hydrolyzed sample of sodium alginate. Zhao, et al., (2012) also reported the hydroxyl radical scavenging activity for low molecular weight alginate oligosaccharides. Additionally, Wang, et al., (2007) reported 60% hydroxyl radical inhibition by alginate oligosaccharides whereas chito- oligosaccharides and fucoidin oligosaccharides depicted 30% and 20% inhibition respectively.

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Nitric oxide has many physiological effects even at very low concentrations, including blood pressure control, inhibition of platelet aggregation, neuronal signaling, antimicrobial activity, antitumor activity, smooth muscle relaxation and regulation of cell mediated toxicity (Hagerman, 1998). Formation of NO is elevated during infections and inflammations and depicts undesired negative results (Marcocci, et al., 1994 a, b). The

NO molecule become very unstable in aerobic conditions and produced highly genotoxic intermediates in the presence of oxygen (Marcocci, et al., 1994a, b)

In the present study NO was formed from sodium nitroprusside at neutral pH

(Green et al., 1982; Marcoci et al., 1994a, b). Greiss reagent is used for the estimation of nitrite ions liberated from the reaction between the nitric oxide and oxygen. Nitric oxide scavengers rivalize oxygen leading to nitric oxide decreased generation (Marcocci et al.,

1994a, b). The optical density of the coloured complex produced by nitrite and sulphanilamide followed by pairing with napthylethylene-diamme was measured spectroscopically.

The nitric oxide scavenging activity of the alginate oligosaccharides produced by hydrolyzed sodium alginate were compared with the nonhydrolyzed sodium alginate lyase. The maximum activity of approximate 50% was depicted in the 3 h hydrolyzed sample whereas it was lower in 6 and 9 h alginate oligosaccharide samples. The non hydrolyzed sample exhibited negligible scavenging activity when compared to the digested oligosaccharides. Chen, et al., (2009) reported a dose dependent nitric oxide scavenging activities of low molecular weight agar, chitosan and starch oligosaccharides. the nitric oxide scavenging activity could be attributed to the negative charge on the oligosachharides (Tsai , et al., 2008; Liu and Ng, 1999)

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Earlier reports suggested that oligosaccharides from Laminaria japonica,

Porphyra haitanesis and Fucus vesiculosus demonstrated antioxidative properties. It is believed that these polysaccharides scavenge metal ions by chelation (Zhao, et al., 2002).

In this present study, the alginate oligosaccharide produced by enzymatic digestion exhibited the ferrous ion chelation activity whereas negligible activity was noticed in non-hydrolyzed alginate sample.

Oligosaccharides with different degree of sulphation and high molecular mass contribute differently to the antioxidative properties. However, low molecular mass polysaccharide preparations have been reported to demonstrate antioxidative properties

(Xue et al., 2001, Zhao, et al., 2012). Based on literature, it is expected that alginate oligosaccharides prepared by enzymatic digestion have antioxidant properties which varies with the molecular weight of alginate oligosaccharides. Zhao, et al., (2012) and

Wang, et al., (2007) have shown that enzymatically depolymerized alginate of varying molecular size can scavenge free radicals. It has been also reported that the low molecular weight oligosaccharides produced by enzymatic depolymerization (Zhao, et al., 2012) or by radiation-induced degradation (Sen, 2011) have higher antioxidant activity when compared to higher molecular weight fractions as observed in present study.

Poly G specific lyases mainly degrade poly (G/M) and poly (G) alginate and demonstrate less degradative activity for poly (M). Iwasaki and Matsubara, (2000) has articled that oligosaccharide generated by alginase from Corynebacterium sp. can enhance the elongation of lettuce and root. It has also been reported for root growth of carrot and rice with unsaturated oligoguluronate mixture whereas oligomannuronate

213 showed insignificant root elongation (Xu, et al., 2003). Therefore, the guluronate specific lyase of Microbulbifer mangrovi sp. nov. DD-13T can be potentially be used for production of oligosaccharides for agricultureapplications.

Thus the oligosaccharides obtained by alginate digested by alginate lyase of

Microbulbifer mangrovi strain DD-13T has the potential of being exploited as reducing agent, hydroxyl and nitric oxide scavenger as well as metal chelator. Alginate is generally a safe food additive as recommended by FDA. Thus alginate has been widely consumed as food additive without any side effects. Alginate has been used as thickening agents in food industry. They are also widely used in gelling and stabilizing agents. The alginate oligosaccharides prepared by alginate lyase digestion could be added as additives during preparation of jellies, candies, cakes and jam which in turn will improve the nutraceutical quality of edibles.

Algae and pytoplanktons are primary food source for herbivorous fish. As they are rich in dietary fibers, minerals, proteins, vitamins and carotenoids, they serve as rich nutritional source for aquaculture (Burtin, et al., 2003). The present aquaculture system needs large quantities of feeds in limited duration. This is a major concern for the future of the aquaculture industry. Studies have been under process to use plant proteins such as soybean meal and wheat flour to replace the conventional feeds. Algal biomass is another candidate to replace fishmeal. Algal diets have been extensively used for the cultivation and their effect on yield of aquaculture has been reviewed (Viera, et al., 2005). It is relatively a difficult task to utilize the seaweeds directly due to their high content of indigestible fiber.

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Both fungi and bacteria could play an important role in breaching the non digestible materials from seaweeds since many of them have the ability to digest cellulosic material resulting in formation of detritus. Detritus prepared with microbial consortia are readily utilized by marine animals for absorption of nitrogen content (Mann, et al., 1988). Rice (1982) showed that nitrogen material available after decomposition of

Spartina formed condensation products and produced precursors for complex biological processes. Hence provision of seaweed detritus to the artificial aquaculture ponds will certainly enhance the production and yield of commercially important marine macroorganisms.

Microorganisms with decomposing activity of seaweeds have been used in preparation of protoplasmatic detritus from Laminaria japonica (Uchida, 1996). The combined effect of enzymatic and microbial hydrolysis of the seaweed cell wall has been reported for improving protein digestibility. Uchida and Murata (2001) reported a successful preparation of SCD by enzymatic and microbial fermentation. Uchida, et al.,

(2004a) had reported the activity of both lactic acid bacteria (LAB) and yeast for the preparation of detritus to feed young pearl oysters. Further only enzymatic treatment had also been reported for the same (Fleurence, 1999; Nikolaeva, et al., 1999).

In the present study, Microbulbifer mangrovi strain DD-13T a multiple polysaccharide degrader, successfully releases single cell detritus from seaweeds without additional enzymatic treatment. The results clearly indicated that the Microbulbifer strain

DD-13T sequentially release algal cell clumps and single cells detritus into the surrounding medium. Such digestion of the seaweed cell wall would increase the bioavailability of nutrient to the marine organisms. The preliminary study on the

215 preparation of algal cell detritus from Sargassum tenerrimum thalli showed that

Microbulbifer strain DD-13T alone could be efficiently used for the preparation of algal cell detritus that can be used as a feeding material. Microbulbifer mangrovi strain DD-

13T produced multiple polysaccharide degrading enzyme and degrades eleven different polysaccharides such as CMC, agar, alginate, carrageenan, pullulan, laminarin, chitin, xylan, starch, β-glucan and pectin. The treatments with these multiple enzymes would break down the seaweeds to release cell clumps/single cells leaving the empty mesh aside. Other nutrients released into the medium during decomposition would also be made available to the feeding organisms.

These single cell detritus are natural food and would provide all the essential nutritional requirements for the growth of animals unlike artificial food which could cause proliferation of harmful bacteria and leading to loss in the quality and yield. Thus

Microbulbifer strain DD-13T has the potential for preparation of single cell detritus from various brown seaweeds.

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SUMMARY AND CONCLUSION

217

 Screening for alginolytic bacteria from four types of niches: coastal water/ sediments

(sandy shore and rocky shore), mangroves and decomposing seaweeds associated

microbial consortia, were used as study area.

 A total of 200 bacterial isolates were obtained on the ASW agar- alginate plates by

direct plate and enrichment technique. The direct plate method was found to be the

ideal method for isolating maximum number of isolates.

 Only 25% alginolytic bacterial isolates were obtained. The number of isolates

exceeds in seaweeds associated niches.

 Out of 25%, only 44% isolates were found to perform multiple polysaccharide

degradation (degrading five polysaccharides).

 The growth of the selected bacterial isolates with Fucus distichus is found to be very

poor when compared to the growth of bacterial isolates with Sargassum tenerrimum.

A total of 20 bacterial isolates were found to utilize Sargassum tenerrimum for their

growth.

 One of the alginate degrading isolate designated as isolate DD-13 was selected for

further studies based on its multiple polysaccharide degrading activity and extensive

degradation of brown seaweeds.

 Bacterial strain DD-13 is a Gram-negative, aerobic, non-motile and rod-shaped

bacteria isolated from sediments of mangrove located at Divar Island, Goa, India.

 Bacterial strain DD-13 possesses a rod–coccus cell cycle in association with the

growth phase.

 The strain DD-13 can degrade eleven tested insoluble complex polysaccharides (ICP)

such as agar, alginate, chitin, cellulose, laminarin, pectin, pullulan, starch,

217

carrageenan, β-glucan and xylan; the maximum number of polysaccharides reported

till date.

 The strain DD-13 was found to be sensitive for nickel, zinc, copper, iron, cobalt,

manganese and mercury whereas it demonstrated resistance for magnesium,

potassium, lithium and lead.

 The biochemical test did not aid in the identification of the bacterial strain DD-13.

 The strain depicted Q-8 as the major ubiquinone whereas iso-C15 : 0 and iso-C11 : 0 as

the major fatty acids.

 The DNA G+C content was 61.4 mol%.

 A phylogenetic analysis based on 16S rDNA gene sequences demonstrated that strain

DD-13 belonged to the genus Microbulbifer exhibiting sequence similarity values of

95.6–98.2% with respect to the type strains of five closely related Microbulbifer

species namely M. hydrolyticus, M. salipaludis, M. celer, M. agarilyticus and M.

elongatus.

 The sequence was submitted to GenBank (NCBI) with accession No. HQ424446.1

and Ref seq. accession No. NR_109105.1.

 Strain DD-13T showed DNA–DNA relatedness values of 28, 33, 27, 26 and 32% with

respect to the reference type strains of M. hydrolyticus, M. salipaludis, M. celer, M.

agarilyticus and M. elongatus respectively which is far below then the generally

accepted species differentiation limit of 70 %.

 Hence DNA–DNA relatedness data and the differential phenotypic properties and

phylogenetic distinctiveness of DD-13T make this strain distinguishable from other

recognized Microbulbifer species.

218

 On the basis of the phenotypic, phylogenetic and genetic data, strain DD-13T

represents a novel species of the genus Microbulbifer, for which the name

Microbulbifer mangrovi sp. nov. is proposed with the type strain DD-13T.

 The addition of glucose, fructose, sucrose, maltose, mannose, and CMC had negative

effect on alginate lyase production whereas alginate lyase expression was better

induced by alginate, followed by starch.

 The medium supplemented yeast extract demonstrated highest activity whereas other

nitrogen sources such as, beef extract, ammonium nitrate, tryptone, urea, casein

hydrolysate, ammonium sulfate and vitamin B2 had positive effect on the growth and

alginate lyase production.

 The activity of the alginate lyase, after being induced by alginate in the culture broth

of ASW containing 0.2% alginate supplemented with 0.4% yeast extract, increased 5

fold compared to the expression of alginate lyase in marine broth.

 The optimal temperature and harvest time for alginate lyase production was 30℃ for

24 h on orbital shaker (130 rpm)

 For the downstream processing of alginate lyase from strain DD-13T ultrafiltration of

crude culture supernatant to 10 fold with 10 kDa membrane cut-off was found to be

ideal method for protein concentration as it could increase the alginate lyase activity

by 5.2 fold.

 The alginate lyase from Microbulbifer mangrovi sp. nov. strain DD-13T was purified

from culture supernatant using a combination of anion-exchange (DEAE sepharose)

and gel filtration ( Seralose CL-6B) column chromatography.

219

 Alginate lyase enzyme was purified to 16 fold from culture supernatant with a yield

of 6.5% and the final specific activity of 43.27 Umg-1.

 The homogeneity of the purified alginate lyase was confirmed by PAGE as single

band was observed on native and SDS PAGE with a corresponding activity band in

Native zymogram.

 Molecular weight of the purified alginate lyase from SDS- PAGE (48.4 kDa) and gel

filtration (47.06 kDa) were obtained to be approximately same.

 The optimum pH and temperature of the purified alginate lyase activity was 6.0 and

50˚C respectively. The enzyme was stable at the pH range of 5.0 to 7.0.

 Cation such as potassium, sodium, zinc, copper, nickel at a concentration of 2 mM

and 0.1% SDS increased the activity of purified alginate lyase.

 The purified alginate lyase was observed to degrade poly M and poly G blocks but

predominately poly G.

 The TLC result depicts the purified alginate lyase to be an endolytic enzyme.

 Alginate oligosaccharides released from sodium alginate by the action of alginate

lyase from Microbulbifer mangrovi strain DD-13T show promising antioxidative

properties.

 The oligosaccharides produced exhibited reducing activity, inhibited hydroxyl radical

and nitric oxide formation and showed metal (ferrous ion) chelating activity.

 The oligosaccharides prepared by 3 h hydrolysis demonstrated better antioxidant

activity over 6 and 9 h hydrolysis products.

 Therefore the alginate oligosaccharides which are prepared by enzymatic hydrolysis

of commercially available sodium alginate could be added as additives during

220

preparation of jellies, candies and jam which in turn will increase the nutraceutical

quality of food.

 The strain DD-13T was successfully used for the preparation of algal cell detritus

from thalli of Sargassum tenerrimum.

 The single cell detritus was successfully released leaving an empty mesh like

structure in the thalli surface.

 Single cell detritus can be used as feed in aquaculture farms especially related to

prawn hatcheries. Unlike artificial food, the single cell detritus are natural food and

would provide all the essential nutritional requirements for the growth of animals in

aquaculture farms.

221

FUTURE PROSPECTS

222

 The oligosaccharides which are obtained by alginate lyase treatment have

immense and diverse applications. Oligosaccharides can be obtained from various

sources and their antioxidant activities could be examined.

 Further the use of alginate lyase to produce biofuels and extraction of bioactive

compounds from seaweed could also be explored.

 The use of purified alginate lyase for preparation of protoplast from seaweed

would be an additional application pursuing in the field of Biotechnology.

 Since Microbulbifer mangrovi sp. nov. strain DD-13T produces multiple

polysaccharase enzymes such as CMCases, alginases, agarase, chitinase,

laminarinase, pectinase, pullulanase, amylase, carrageenase, -glucanase and

xylanase could be purified and their biochemical properties as well as their

possible biotechnological applications could also be examined.

 Cloning of the alginate lyase gene and deciphering the protein structure will help

in classifying the alginate lyase to appropriate Glycosyl hydrolase family as well

as its genetic relatedness to other alginate lyase gene could be studied.

 Cloning of the gene of other polysaccharide degrading enzymes from

Microbulbifer mangrovi sp. nov. strain DD-13T.

222

APPENDIX

APPENDIX 1

1.1. Artificial Seawater (ASW) (g/L)

Tris base : 6.05

MgSO4 : 12.32

KCl : 0.74

(NH)2 HPO4 : 0.13

NaC1 : 17.52

CaC12 : 0.14

Dissolve in 900 ml of double distilled water and adjust the pH to 7.0 with con HCI immediately. Make up the volume to 1000m1 with double distilled water before autoclaving.

1.2. Lugol's Iodine (g/100m1)

Potassium Iodide : 1.66 (0.1M)

Iodine (crystals) : 1.26 (0.05M)

Add to 100 ml of distilled water in a amber bottle and stir at RT vigorously till Iodine crystals dissolves. Store in an amber colour bottle.

1.3. 3,5-dinitrosalicylic acid (DNSA reagent) (100m1 distilled water)

NaOH : 1 g

3, 5-dinitrosalicyclic acid : 1 g

Sodium Potassium Tartarate : 20 g

Phenol : 0.2g

Sodium bis-sulfite : 0.05g

Make volume to 100 ml with distilled water.

1.4. Nelson and Somogyi reagent

Solution A (100 ml)

Na2CO3 : 2.5 g

Sodium Potassium Tartarate : 2.5 g

Na2SO4 : 20 g

Solution B (100 ml)

CuSO4. 5H2O : 15 g

H2SO4 (conc) : 2 drops

Solution C (incubate at 37˚C for overnight) (250 ml)

Ammonium molybdate : 12.5 g Dissolved in 225 ml distilled water H2SO4 (conc) : 10.5 ml

Sodium arsenate heptahydrate: 1.5 g in 12.5 ml distilled water. Mix all and make the volume to 250 ml

Mix Solution A: B = 25:1 at the time of use.

1.6. Sodium Citrate buffer (pH 6.0) (50mM) (g/L)

Citric acid, Monohydrate : 10.5

Sodium citrate : 14.71

Dissolve in 900 ml of water and adjust the pH to 6.0 with 1 M NaOH. Make up the volume to 1000 ml. Sterilize by autoclaving at 121°C for 15 min.

1.7. Crystal Voilet (for Gram staining)

Crystal violet : 2 g in 20 ml of Ethyl alcohol (95%)

Ammonium Oxalate : 0.8 g in 80 ml of water

Mix both solutions to make homogenous solution and filter through Whatman no.1 filter.

1.8. Gram's Iodine (for Gram staining)

Iodine : 1 g

Potassium Iodide : 2 g

Dissolve in 300 ml of water by stirring at RT and stored at RT in amber colour bottle.

1.9. Decolurizing Agent (for Gram staining)

Absolute alcohol : 95 ml

Distilled water : 5 ml

1.10. Saffranin solution (for Gram staining)

Saffranin : 2.5 g

Ethyl alcohol (95%) : 100 ml

Dissolve and filter through Whatman no.1 filter. Dilute 10 ml of stock solution with 100 ml of water.

1.11. Kovac's Reagent

p- Dimethylaminobenzaldehyde : 3 g

Butanol : 75 ml

Con HCl : 25 ml

Dissolve p- Dimethylaminobenzaldehyde in butanol at 50°C. Cool to RT and add Con HCI. Store in amber colour bottle at 4°C.

1.12. Methyl Red Reagent

Methyl Red : 0.1 g

Prepare by dissolving in 300 ml of 95% ethanol (v/v). Make up the volume to 500 ml with water.

APPENDIX 2

2.1. TE Buffer (pH 8.0)

Tris-C1 (10mM) :0.157 g

EDTA (1mM) :0.028 g

Dissolve in 90 ml of water and adjust the pH to 8.0. Make up the volume to 100 ml.

2.2. Saturated Phenol (pH 8.0)

Phenol, which was stored at -20°C was thawed to room temperature and melted at 65°C in a water bath. To this 0.1% 8-hydroxyquinoline was added. To this, equal volume of 0.5M TrisCl (pH 8.0) was added and stirred for 15 min at RT. After stirring, the solution was allowed to settle and aqueous layer was removed as much as possible. To this 0.1M Iris-CI (pH 8.0) was added and repeated as above till the pH of phenol solution reaches 8.0. After pH was reached to 8.0, 0.1 volume of 0.1M Tris-Cl added and stored at 4°C.

2.3. 3M Sodium Acetate Buffer (pH 5.2)

Sodium acetate : 40.81 g

Distilled water : 80 ml

Adjust the pH to 5.2 with acetic acid and make up the volume to 100 ml. Sterilize by autoclaving.

2.4. 5X TBE buffer

Tris Base : 54 g

Boric Acid : 27.5 g

EDTA (0.5M) : 20 ml

(Dissolve 186.1 g of di sodium EDTA in 800 ml of water and adjust the pH to 8.0 with 1M NaOH) Dissolve in 800 ml of water and make up the volume to 1000 ml.

2.5. 50X TAE

Tris base : 242 g

Acetic acid (glacial) : 57.1 ml

0.5 M EDTA : 10 ml

Dissolve and mix the components in 800m1 of water and make up the volume to 1000 ml

2.6. 6X loading dye

Sucrose :40 g

Bromophenol Blue :0.25 g

Dissolve in 80 ml of water and make up to 100 ml with water.

2.7. 12% SDS-PAGE Composition

Resolving gel (12%) : 20 ml

Acrylamide Mix (30%) : 8.35 ml

4X Resolving gel buffer : 5 ml

10% SDS : 0.2 ml

10% Ammonium persulphate : 0.1 ml

TEMED : 6.65 µl

Water : 6.4 ml

Add TEMED before pouring into the glass plates with glass pipette.

Stacking gel (4%): 10 ml

Acryalamide : 1.33 ml

4X stacking gel buffer : 2.5 ml

10% SDS : 0.1 ml

10% APS : 0.05 ml

TEMED : 5 µl

Water : 6.0 ml

2.8. 6X SDS-PAGE Treatment Buffer

4X Tris-CI (pH 6.8) : 7 ml

Glycerol : 3 ml

SDS : 1 g

DTT : 0.93 g

BPB : 1.2 g

2.9. 6X Native PAGE Treatment Buffer

4X Tris-Cl(pH 6.8) : 7 ml

Glycerol :3 ml

BPB : 1.2 g

2.10. Coomassie Brilliant Blue stain

Coomassie Brilliant Blue R-250 : 0.25 g

Methanol : 50 ml

Acetic acid (glacial) : 10 ml

Water : 40 ml

Dissolve in 50 ml of Methanol and filter through Whatman no. 1. Volume was made tol 00 ml with 10 ml of acetic acid and 40 ml water.

APPENDIX 3

3.1. 10% Trichloroacetic acid (TCA)

Trichloroacetic acid : 500 g

Water : 227 ml to make 100% TCA.

10 ml of 100 % TCA and dilute to 100 ml with water.

3.2. Hydroxy radical generation buffer

Sodium PO4 Buffer (150 mM) : Made from 200 mM phosphate buffer

FeSO4 . 7H2O : 10 mM

EDTA disodium : 10 mM

Sodium Salicylate : 2 mM

3.3. TLC developing reagent:

Diphenylamine : 0.4 g

Aniline : 0.4 ml

Phosphoric acid : 3 ml

Acetone :20 ml

International Journal of Systematic and Evolutionary Microbiology (2013), 63, 2532–2537 DOI 10.1099/ijs.0.042978-0

Microbulbifer mangrovi sp. nov., a polysaccharide-degrading bacterium isolated from an Indian mangrove

Poonam Vashist,1 Yuichi Nogi,2 Sanjeev C. Ghadi,1 Pankaj Verma3 and Yogesh S. Shouche3

Correspondence 1Department of Biotechnology, Goa University, Taleigao Plateau, Goa, India Sanjeev C. Ghadi 2Extremobiosphere Research program, Japan Agency for Marine-Earth Science and Technology [email protected] (JAMSTEC), 2-15, Natsushima-cho, Yokosuka 237-0061, Japan 3Molecular Biology Unit, National Centre for Cell Science, Pune University, Pune, India

A rod-shaped, Gram-negative, non-motile, aerobic and non-endospore forming bacterium, designated strain DD-13T, was isolated from the mangrove ecosystem of Goa, India. Strain DD-13T degraded polysaccharides such as agar, alginate, chitin, cellulose, laminarin, pectin, pullulan, starch, carrageenan, xylan and b-glucan. The optimum pH and temperature for growth was 7 and 36 6C, respectively. The strain grew optimally in the presence of 3 % NaCl (w/v). The DNA G+C content was 61.4 mol%. The T predominant fatty acid of strain DD-13 was iso-C15 : 0. Ubiquinone-8 was detected as the major respiratory lipoquinone. Phylogenetic studies based on 16S rRNA gene sequence analysis demonstrated that strain DD-13T formed a coherent cluster with species of the genus Microbulbifer. Strain DD-13T exhibited 16S rRNA gene sequence similarity levels of 98.9–97.1 % with Microbulbifer hydrolyticus IRE-31T, Microbulbifer salipaludis JCM 11542T, Microbulbifer agarilyticus JAMB A3T, Microbulbifer celer KCTC 12973T and Microbulbifer elongatus DSM 6810T. However, the level of DNA–DNA relatedness between strain DD-13T and the five type strains of these species of the genus Microbulbifer were in the range of 26–33 %. Additionally, strain DD-13T demonstrates several phenotypic differences from these type strains of species of the genus Microbulbifer.ThusstrainDD- 13T represents a novel species of the genus Microbulbifer,forwhichthenameMicrobulbifer mangrovi sp.nov.isproposedwiththetypestrainDD-13T (5KCTC 23483T5JCM 17729T).

The genus Microbulbifer was originally reported by Gonza´lez associated with different stages of their growth stages et al. (1997) to describe strictly aerobic, rod-shaped (Nishijima et al., 2009). members of the phylum Proteobacteria having numerous The mangrove ecosystem, the connecting link between vesicles on the membrane surface. Subsequently, several terrestrial and marine ecosystems is rich in organic matter. novel species belonging to the genus Microbulbifer with the Diverse floral and faunal communities thrive in the capability to degrade several polysaccharides have been different niches existing in this ecosystem. Furthermore, isolated from various habitats such as salt marsh, intertidal the litter which is composed of mangrove foliage and sediment, solar saltern, sediments (marine and deep-sea), vegetative remains contributes to the induction of com- marine algae and mangrove forests (Yoon et al., 2003, 2004, plex polysaccharides (CPs) such as cellulose, xylan, pectin, 2007; Miyazaki et al., 2008; Nishijima et al., 2009; Wang et al., 2009; Baba et al., 2011; Zhang et al., 2012). pullulan etc. in this ecosystem. The colonization of CP- decomposing micro-organisms within litter to form the Additionally, the presence of iso-C15 : 0 as the major fatty acid and Q-8 as the predominant isoprenoid quinone are detritus is an important ecological process and plays an characteristic features of the genus Microbulbifer. important role in carbon recycling. In the present study, we Furthermore, species of the genus Microbulbifer are also report the taxonomic characteristics of a novel species known to demonstrate a characteristic rod–coccus cycle belonging to the genus Microbulbifer that was isolated from the mangrove ecosystem of Goa, India and is capable of Abbreviation: CP, complex polysaccharide. degrading several polysaccharides. T The GenBank/EMBL/DDBJ accession number for the 16S rRNA gene Bacterial strain DD-13 was isolated from a water sample sequence of Microbulbifer mangrovi DD13T is HQ424446. collected from mangroves of Goa, India (15u 309 350 N73u A supplementary figure and a supplementary table are available with the 529 630 E). The mangrove forest is situated along the online version of this paper. Mandovi estuary of Goa and detailed physico-chemical

2532 042978 G 2013 IUMS Printed in Great Britain Microbulbifer mangrovi sp. nov. characteristics of the water have been reported (Singh et al., 35 uC up to late-exponential phase. Fatty acid composi- 2009). The water sample was collected in sterile bottles and tions were determined following the instructions of the later serially diluted in an artificial sea water medium Sherlock Microbial Identification System (Sasser, 1990) (ASW) (Ghadi et al., 1997). An aliquot was spread plated using a Finnigan TRACE DSQ GC–MS system (Thermo on ASW containing 2 % agar supplemented with 1 % Fisher Scientific) equipped with a DB-5 column (J&W 2 sodium alginate. The plates were incubated at 30 uC for Scientific) under a helium flow of 1.5 ml min 1 and an 48–72 h. Single colonies displaying depression or clearance oven temperature program increasing from 140 uC (5 min) zones were picked and purity was confirmed by repeated to 280 uC (5 min) at 4 uC min21. Furthermore, strain DD- streaking. The alginolytic activity of the isolates was 13T cultivated in MB at 35 uC for 24 h was used for detected by flooding culture plates with 10 % cetyl isolation of genomic DNA (Maloy, 1990) and isoprenoid pyridinium chloride (Gacesa & Wusteman 1990). One of quinone analysis using reverse-phase HPLC (Komagata & the alginolytic isolates that demonstrated multiple poly- Suzuki, 1987). The DNA G+C content was determined saccharide degrading ability was designated strain DD-13T using the method of Tamaoka & Komagata (1984). DNA– and was subjected to polyphasic characterization to DNA hybridization (in triplicate) was measured fluorome- determine its exact taxonomic position. trically at 48 uC for 4 h as described by Ezaki et al. (1989). The morphological and growth characteristics of strain PCR amplification and sequencing of the 16S rRNA gene DD-13T were investigated by growth on marine agar 2216 were carried out as described by Hauben et al. (1997). The (MA; HiMedia) at 36 uC. Growth was determined at 16S rRNA gene sequence was determined using Big Dye different temperatures (4, 10, 15, 20, 25, 30, 32, 34–39 and terminator v3.1 Cycle Sequencing kit and the run was 45 uC). The pH range (3–12) for bacterial growth was carried out in an automated DNA sequencer, model assessed in customized marine broth (MB) whose pH was 3730XL (Applied Biosystems). The 16S rRNA gene T adjusted by adding respective buffers [50 mM of the sequence of strain DD-13 (1536 bp) was compared and respective buffers: citrate (pH 3–6); Tris base (pH 7–8); analysed with respective reference gene sequences by borate (pH 9); glycine–NaOH (pH 10); sodium bicarbon- NCBI-BLAST program for identification of closely related ate (pH 11); potassium chloride (pH 12)]. Modified MB type strains with validly published bacterial names. The 16S (devoid of NaCl) was supplemented with NaCl in the range rRNA gene sequences of the closest homologues were of 0–10 % (w/v) for investigating growth at different downloaded from GenBank and EMBL databases and concentrations of NaCl. multiple alignments were performed with CLUSTAL X program (version 1.83; Thompson et al., 1997). The Unless otherwise stated, biochemical studies of strain alignment gaps and missing data were edited. The T DD-13 were performed in ASW medium at 36 uC. evolutionary distance matrices were generated according Furthermore, whenever reference type strains of species of to Jukes & Cantor (1969). Phylogenetic trees were inferred the genus Microbulbifer were used for comparison, they were using the maximum-parsimony, maximum-likelihood and also grown in ASW at their respective optimum tempera- neighbour-joining methods with the MEGA program tures. Catalase and oxidase activity, hydrolysis of casein, (version 5.0; Tamura et al., 2011). The robustness of the starch and Tweens 20, 40, 60 and 80 were determined as topology of phylogenetic trees was evaluated by a bootstrap described by Cowan & Steel (1965). Hydrolysis of aesculin analysis with 10 000 replications. and urea, reduction of nitrate and carbohydrate utilization T profiles were analysed using HiAssorted biochemical test The 16S rRNA gene sequence of strain DD-13 (HQ424446) was a continuous stretch of 1536 bp. A BLAST analysis of the kit KB002 and HiCarbohydrate kit KB009 respectively T (HiMedia). The multiple polysaccharase activity was 16S rRNA gene sequence of strain DD-13 indicated that it detected by the plate screening method after 48 h on ASW belonged to the class Gammaproteobacteria and was closely related to the genus Microbulbifer. The sequence similar- agar plates or ASW agar plates supplemented with 0.2 % of T [ ity levels of 16S rRNA of strain DD-13 in compari- any one polysaccharide alginic acid (mixed polymer of T guluronate and mannuronate), chitin (poly-N-acetyl-1,4,b- son to Microbulbifer hydrolyticus IRE-31 , Microbulbifer salipaludis JCM 11542T, Microbulbifer agarilyticus JAMB D-glucopyranosoamine), CM-cellulose (sodium salt), lami- A3T, Microbulbifer celer KCTC 12973T and Microbulbifer narin (from Laminaria digitata), pectin (poly D-galacturonic T acid methyl ester), pullulan (standard), starch (from elongatus DSM 6810 were 98.9, 98.5, 97.5, 97.2 and potato), carrageenan (from Irish moss), b-glucan (from 97.1 %, respectively. The 16S rRNA gene sequence identity with other members of the genus Microbulbifer was in the barley) or xylan (from oat spelts)] (Stanier, 1942; Morgan range 96.7–93.6 %. The 16S rRNA phylogenetic tree based et al.,1979; Teather & Wood, 1982; Gacesa & Wusteman on the neighbour-joining algorithm, depicted strain DD-13T 1990; Gonza´lez-Candelas et al., 1995; Ruijssenaars & to be in the same clade as the species of the genus Hartmans, 2001). Laminarin, pullulan and b-glucan were Microbulbifer (Fig. 1). Phylogenetic analysis based on from Sigma whereas other polysaccharides were obtained neighbour-joining method showed that strain DD-13T from HiMedia. formed an evolutionarily distinct lineage within the cluster Bacterial strain DD-13T and type strains of closely related comprising M. hydrolyticus IRE-31T, M. salipaludis JCM species of the genus Microbulbifer were grown in MB at 11542T, M. agarilyticus JAMB A3T, M. celer KCTC 12973T http://ijs.sgmjournals.org 2533 P. Vashist and others

100 • Microbulbifer variabilis Ni-2088T (AB167354) 0.02 94• Microbulbifer epialgicus F-104T (AB266054) Microbulbifer maritimus TF-17T (AY377986) 85 Microbulbifer donghaiensis CN85T (EU365694) • • 51 Microbulbifer okinawensis ABABA23T (AB500893) 95• Microbulbifer marinus Y215T (GQ262812) Microbulbifer thermotolerans JAMB A94T (AB124836) • Microbulbifer chitinilyticus ABABA212T (AB500894) 100 70 • 70 Microbulbifer halophilus YIM 91118T (EF674853) Microbulbifer yueqingensis Y226T (GQ262813) Microbulbifer celer ISL-39T (EF486352) 82 JAMB A3T • 93 Microbulbifer agarilyticus (AB158515) 81 • Microbulbifer elongatus DSM 6810T (AF500006) • 75 Microbulbifer salipaludis SM-1T (AF479688) • Microbulbifer mangrovi DD-13T (HQ424446) 60 • 90 Microbulbifer hydrolyticus DSM 11525T (AJ608704) Marinobacter oulmenensis Set74T (FJ897726) • LMG 24048T (AM503093) 100 Marinobacter guineae • R65T (AJ609270) 99 • Marinobacter sediminum 62 Marinobacter flavimaris SW-145T (AY517632) Vibrio communis R-40496T (GU078672)

Fig. 1. Neighbour-joining phylogenetic tree based on 16S rRNA gene sequences, representing the position of strain DD-13T, species of the genus Microbulbifer and other related taxa. Bootstrap percentages based on 10 000 replications .50 % are shown at branch points. $ denotes that the corresponding nodes were also recovered in the trees generated with the maximum-likelihood and maximum-parsimony algorithms. Vibrio communis R-40496T was used as an outgroup. Bar, 0.02 substitutions per nucleotide position.

and M. elongatus DSM 6810T, supported by a bootstrap genus Microbulbifer, strain DD-13T is able to grow in MB resampling value of 100 % (Fig. 1). The same relationship without NaCl. Furthermore, strain DD-13T does not was also inferred from phylogenetic trees obtained using hydrolyse casein and produces H2S whereas the other maximum-parsimony and maximum-likelihood methods closely related species of the genus Microbulbifer used for (not shown). comparison in the present study hydrolysed casein and did T Strain DD-13T was isolated as a greyish yellow colony on not produce H2S (Table 1). Additionally, strain DD-13 MA and formed depressions when grown on ASW agar demonstrated several differences from closely related plates containing 1 % (w/v) sodium alginate. Strain DD- species of the genus Microbulbifer with regard to percentage + 13T is catalase and oxidase-positive and can degrade 11 G C content of DNA, ability to grow in MB amended with 10 % NaCl and ability to grow at pH 5 and 10 (Table different polysaccharides. Besides the presence of iso-C15 : 0 as the major fatty acid in strain DD-13T that confirmed 1). When compared with other species of the genus T its affiliation to the genus Microbulbifer, iso-C17 : 1v9c, Microbulbifer, strain DD-13 degrades 11 tested complex C18 : 1v7c and C16 : 0 were also detected as other prominent polysaccharides: agar, alginate, chitin, cellulose, laminarin, fatty acids. No significant differences were observed when pectin, pullulan, starch, carrageenan, xylan and b-glucan the fatty acid profile of strain DD-13T was compared with (Fig. S1). Saccharophagus degradans, isolated from salt the fatty acid profiles of related type strains of species of the marsh is the only other polysaccharide-degrading strain genus Microbulbifer (Table S1 available in IJSEM Online). reported to degrade 10 polysaccharides (Ekborg et al., Detailed results of phenotypic and biochemical studies of 2005). Amongst the species of the genus Microbulbifer strain DD-13T are shown in Table 1 or mentioned in the tested, only strain DD-13T and M. elongatus DSM 6810T species description. were observed to degrade pectin (Table 1). On the basis of physiological characteristics, strain DD-13T Strain DD-13T exhibited 16S rRNA gene sequence is easily differentiated from the closely related species of the similarity levels of 97.1–98.9 % to the type strains of genus Microbulbifer (Table 1). Unlike other species of the species of the genus Microbulbifer chosen in the present

2534 International Journal of Systematic and Evolutionary Microbiology 63 Microbulbifer mangrovi sp. nov.

Table 1. Differential phenotypic characteristics of strain DD-13T and other related species of the genus Microbulbifer

Strains: 1, DD-13T, data from this study; 2, M. hydrolyticus IRE 31T;3,M. salipaludis JCM 11542T;4,M. celer KCTC 12973T;5,M. agarilyticus JAMB A3T;6,M. elongatus DSM 6810T. All of the species are positive for the following: being rod-shaped, catalase and oxidase activities, ornithine utilization*, hydrolysis of starch, xylan, agar and cellobiose. All species are negative for spore formation, Gram-staining, urease testD, sodium gluconateD, dulcitolD, inositol, methyl-a-D-glucosideD, a-methyl-D-mannosideD and sorbose utilizationD. +, Positive reaction; 2, negative reaction; W, weakly positive reaction; ND, not determined.

Test 1 2 3 4 5 6

Colony colour Greyish yellow Cream Greyish yellow Greyish yellow Cream Yellowish brown Motility 22+ 2 + 2 DNA G+C (%)* 61.4 57.7 59 57.7 55.2 58.2 Growth at 0 % NaCl + 22222 Growth at 10 % NaCl + 2 ++22 Growth at pH 5 + 22D 2D 2 +D Growth at pH 10 + 22D +D 2 +D

H2S production + 22222 Nitrate reduction 22+ 2 + 2 Casein hydrolysis 2 +++++ Amino acid utilization L-Alanine +++D +D 2D + L-Leucine 2 + 2D +D 2D + L-Proline +++D +D +D 2 L-Serine 2 + 22D 2D + Polysaccharide degradation Chitin ++22D ++ Pullulan + 2D 2D 2D ND ND Pectin + 2D 2D 2D 2D +D Alginate + 2D 2D 2D 2D +D b-Glucan W 2D +D +D ND ND Laminarin + 2D 2D +D ND ND Carrageenan ++D W 2D +D +D Carbohydrate utilization D-Galactose 2 +D 222D + D-Glucose +++ 2 ++ D-Xylose + 2D WD 22D 2D Maltose ++D + 2 ++ D-Arabinose + 2D +D +D 2 + Trehalose 2 +D +D 22WD Melibiose 2 +D 2D 22D WD Inulin 22D 2D 2D 2D +D Sorbitol 2 +D +D 22+D Salicin 2 +D 2D 22D 2D Raffinose 2 +D 2222 Malonate 2 +D WD WD WD 2D

*Data for strains 2–6 is from Gonza´lez et al. (1997), Yoon et al. (2003), Yoon et al. (2007), Miyazaki et al. (2008) and Yoon et al. (2007), respectively. DData obtained from this study.

study. However, DNA–DNA hybridization tests of the differentiation limit of 70 % (Wayne et al., 1987). Thus genomic DNA of DD-13T with M. hydrolyticus IRE-31T, M. strain DD-13T is not related to these species and is a novel salipaludis JCM 11542T, M. celer KCTC 12973T, M. species of the genus Microbulbifer. agarilyticus JAMB-A3T and M. elongatus DSM 6810T Thus, based on chemotaxonomic characteristics, phylo- revealed similarities of 28, 33, 27, 26 and 32 %, respectively. genetic data and genomic distinctiveness, strain DD-13T These similarities are below the generally accepted species should be placed in the genus Microbulbifer as a novel http://ijs.sgmjournals.org 2535 P. Vashist and others species, for which the name Microbulbifer mangrovi sp. nov. Cowan, S. T. & Steel, K. J. (1965). Manual for the Identification of is proposed with strain DD-13T as the type strain. Medical Bacteria. London: Cambridge University Press. Ekborg, N. A., Gonza` lez, J. M., Howard, M. B., Taylor, L. 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