Characterization of alginate lyase from Microbulbifer mangrovi sp. nov. DD-13 T
Thesis Submitted to Goa University For the degree of
Doctor of Philosophy in Biotechnology by
Ms. Poonam Vashist
Department Of Biotechnology Goa University
Taleigao- Goa
2014
Characterization of alginate lyase from Microbulbifer mangrovi sp. nov. DD-13 T
Thesis Submitted to Goa University For the degree of
Doctor of Philosophy in Biotechnology by
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-13
T” 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 strain DD-13
60- 121
Chapter 5.Optimization of the growth and culture conditions for enhancing production of alginate lyase from Microbulbifer mangrovi sp.nov. DD-13T
122- 148
Chapter 6. Purification and characterization of the alginate lyase from Microbulbifer mangrovi sp.nov. DD13T
149- 189
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……
1
CHAPTER 1:
INTRODUCTION
1 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
2 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)
3 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.
4 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
5 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.
6
RESEARCH GOAL AND
SIGNIFICANCE
6 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
7 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
8 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.
9
CHAPTER 2:
LITERATURE REVIEW
9 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.
10 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
11 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).
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
1H4 and14C 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 demonstrated that the G residues in homopolymeric blocks are arranged in 1C4
conformation whereas the mannuronate residues have the 4C1 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).
G G M M
α-1, 4 β-1, 4 β-1, 4
12 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.
13 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
14 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.
15 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..
16
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.
17 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.
18
• 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
19 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.
20 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
21 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,
22 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.
23 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.
(c) The replacement of the 4-O glycosidic bond by transferring the carboxyl electrons to make a C4-C5 double bond.
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).
Figure 2.2: Alginate lyase mechanism of action on alginate polymer
G G M M
M -Lyase G/M -Lyase
G- Lyase
Enzymatic Hydrolysis
Unsaturated Monomer
Unsaturated Monomer
CHO C C C C
H H H H H O COOH OH
OH
4- deoxy L-erythro-5- hexoseulose uronic acid Tautomeric conversion
25 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
26 (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,
27 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,
28 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
29 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.
30 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
31 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.
32 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
33 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).
34 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
35 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)
36
