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CHAPTER II

REVIEW OF LITERATURE

2.1 Polymers and Microbial polymers

In recent years, the search for environmental friendly polymers has been an

inkling of biotechnologists and engineers. Globally, synthetic plastics production and

consumption have drastically increased to an alarming height (Figure 2.1). In India,

plastic consumption ascended exponentially in the 1990s. During the last decade, the total

consumption of plastics grew twice as fast (12% p.a.) as the Gross Domestic Product

growth rate based on purchasing power parities (6% p.a.). The current growth rate in

Indian polymers consumption (16% p.a.) is higher than that in China (10% p.a.) and

many other key Asian countries. The consumption of plastics will increase six folds

between 2000 and 2030.

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Synthetic plastics are highly resistant to degradation and littering. These polymers

are a major public concern and causes jeopardizing effects on environment. This drives

the researchers globally to develop biodegradable polymers (Kolybaba et al., 2003).

Polymers are solid, non metallic compounds of high molecular weights (Callister et al.,

1999), possessing repetitive units of macromolecules, with varying characteristics

depending on their composition. A variety of materials, both renewable and non

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renewable, are employed as feedstock source for modern plastic materials. Plastics that

are formed from non renewable feedstocks are generally petroleum based and reinforced

by glass/ carbon fibers (William et al, 2000). Renewable resource feedstocks include

microbially grown polymers and those extracted from starch. It is possible to reinforce

such materials with natural fibers from plants such as flax, jute, hemp and other cellulose

sources (Bismarck et al., 2002).

There are three primary classes of polymer materials which material scientists are

currently focusing on. These polymer materials are usually referred to in the general class

of plastic by consumers and industry. Conventional plastics are resistant to

biodegradation, as the surfaces in contact with the soil in which they are disposed, are

characteristically smooth (Aminabhavi et al., 1990). This group of materials usually has

an impenetrable petroleum based matrix, which is reinforced with glass fibers.

Microorganisms within the soil are unable to consume a portion of plastic which would in

turn cause the breakdown of the supporting matrix. Second class of polymers is partially

degradable. These are designed with the goal of more rapid degradation than that of

conventional synthetic plastics. Production of this class of materials typically includes

surrounding naturally produced fibers with a conventional petroleum based matrix. When

disposed off, microbes are able to consume the natural macromolecules with the plastic

matrix. This leavens the weakened materials with rough open edges. Further degradation

may then occur. The final class polymer material is the current attention. These are

classified to be completely biodegradable. The polymer matrix is derived from natural

sources (such as starch or microbially based polymers). Microorganisms are able to

consume these materials in their entirely, eventually leaving CO2 and H2O as byproducts.

2.2 Microbial polysaccharides

Microbial biopolymer feedstock produces biological polymers through microbial

fermentation. The products are naturally degradable, environmental friendly substitutes

for synthetic plastics (Chau et al., 1999). Polysaccharides are microbial polymeric

substances that are structurally diverse group of biological macromolecules of

widespread occurrence in nature (Chawla et al., 2009). They can be divided according to

their morphological localization as intracellular polysaccharide located inside, or as a part

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of the cytoplasmic membrane; cell wall polysaccharide forming the structural part; and

extracellular polysaccharide or exopolysaccharide (EPS) located outside the cell wall,

secreted into the environment. Other polymeric substances produced by microbes are

PHA (polyhydroxy alkonates). These are intracellular carbon reserves when nutrient

deficiencies occur (Lakshmanan et al., 2004). These are microbially produced polyesters

having the same thermoplastic and water resistant qualities as synthetic plastics.

2.3 Structure of Cell Wall

Figure 2.2 Cell wall structure of Gram positive and Gram negative bacteria locating

exopolysaccharides (EPS) (Adapted from Madiedo et al., 2005)

Gram positive and negative bacteria have similar internal, but very different

structure. The structure, components and functions of the cell wall distinguish Gram

positive from Gram negative bacteria. A Gram positive bacterium has a thick,

mutilayered cell wall mainly consisting of peptidoglycan (15-80 nm) a mesh-like

exoskeleton, surrounding the cytoplasmic membrane. Peptidoglycan is important for

structure, replication, survival in normal hostile conditions in which bacteria grow.

During infection, peptidoglycan can interfere with phagocytosis and pyrogenesis. Cell

wall also contains teichoic acid, lipoteichoic acid and complex polysaccharides. Teichoic

acids are water soluble polymers, generally, polyol phosphates, covalently linked with

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peptidoglycan. Lipoteichoic acids are teichoic acids with lipids linked to them and thus

anchored to cytoplasmic membrane.

Gram negative cell wall is more complex than that of Gram positive, both

structurally and chemically. Structurally, Gram negative cell wall contains two layers

external to cytoplasmic membrane, immediately after which is the thin layer of

peptidoglycan layer, accounting only for 5-10 % of total weight. No teichoic and

lipoteichoic acids are found in Gram negative cell wall. Exterior of peptidoglycan is the

outer membrane which is a unique feature of Gram negative cell wall. The area between

the external surface of cytoplasm and internal surface of outer membrane is referred to

periplasmic space, which contains components of sugar transport systems and binding

proteins to facilitate the uptake of metabolites and other compounds.

Gram positive or Gram negative are closely surrounded by polysaccharide or

protein layers called capsules. In cases in which it is loosely adherent or thin in nature, it

is called as slime or biofilm. The layer of capsular and slimy polysaccharide is generally

known as glycocalyx. The capsule is hard to be viewed through a microscope but can be

visualized by the exclusion of India ink particles. A true capsule is a discrete detectable

layer deposited outside the cell wall with polysaccharide. Some bacteria produce slime

materials to adhere and float themselves as colonial masses in their environment. Other

bacteria produce slime to attach themselves to a surface or substrate, divide, produce

micro colonies within slime and construct biofilm which become enriched and protected

for themselves and other microbes.

Capsular materials like dextran may be over produced when bacteria are fed as

sugars to become reserves of carbohydrates for subsequent metabolism. Capsules and

slimes are unnecessary for growth of microbes but are very important for the survival of

the host. Synthesis of capsule takes energy and will not be affected by bacteria after

continued growth under laboratory conditions away from the selective pressure of the

host. Glycocalyx is essential as it is a major virulent factor of microorganism and

capsules act as the barrier to toxic hydrophobic molecule like detergents and can promote

adherence to other bacteria or any host tissue (www.digitalproteus.com,

www.textbookofbcteriology.net).

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2.4 Exopolysaccharides

Exopolysaccharides (EPS) are a complex mixture of biopolymeric

macromolecules consisting of polysaccharides along with proteins, nucleic acids, lipids

and humic substances (Vu et al., 2009). They are generally produced by bacteria, fungi,

yeast, algae, plants (guar gum, pectin) and animals. Few bacterial EPSs have emerged as

industrially important biopolymers with specific characteristics and unique rheological

properties. EPS occurs in two forms: slimy EPS, which is non adherent to the cell and

non uniform in density and; and microcapsules or capsules which adhere firmly to cell

wall, which have definite form and boundary, being slowly extracted in the water or salt

solution. It is therefore possible to separate capsules and microcapsules from slime by

centrifugation (Chawla et al., 2009). Capsular polysaccharides (CPS) are highly hydrated

molecules that are over 95% water. They are often linked to the cell surface through

covalent bonds by the means of membrane anchors like phospholipid and lipid - A

molecules and certain CPSs do not require membrane anchors. Difference between the

forms of capsular and slimy exopolysaccharides are difficult, since CPS released from the

cell, gives the appearance of slime EPS. In turn, distinguishing between CPS and other

cell wall surface polysaccharides, such as O- antigenic lipopolysaccharides (LPS), may

be difficult, since CPS may be found associated with LPS (www.digitalproteus.com).

Capsular and slimy exopolysaccharides can protect bacteria and contribute to

their pathogenicity. Attachment of nitrogen fixing bacteria to plant roots and soil

particles, which is essential for the colonization of rhizosphere and roots, can be also

mediated by EPS and polymers of Pseudomonas sp. involve in this process. EPS from

certain strains of Bacillus act as biocontrolling agent in plants (Bais et al., 2004).

Extracellular polysaccharides from Streptococcus pneumoniae are involved vitally in

severe oral infections of humans, especially in dental plaques. Dextrans, one of the most

commonly available microbial polymers had been primarily used in baking industry.

(www.digitalproteus.com; Freitas et al., 2001).

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Table 2.1 Various kinds of EPS produced by microorganisms

EPS MICROORGANISM

REFERENCE

Dextran

(neutral

homosaccharide with

glucose units)

Lactobacillus

delbruckeii,

L.lactis,

Streptococcus

thermophilus,

Leuconostoc sp.,

Maina et al., 2008;

Barbara Vu et al., 2009

Levan

(neutral

homosaccharide with

fructose units)

Bacillus licheniformis,

Halomonas sp.

Ghaly et al., 2007;

Liu et al.,2010;

Kucukasik et al., 2011

Kefiran

(heteropolysaccharide

with glucose and

galactose residues)

Lactobacillus

rhamnosus, L.kefir,

L.kefiranofaciens

Frengova et al., 2002;

Habibi et al., 2011;

Cheirsilp, 2006

Xanthan

(anionic

heteropolysaccharide)

Xanthomonas

campestris

Freitas et al., 2011;

Moshaf et al., 2011

Pullulan

(homopolysaccharide )

Aureobasidium

pullulans

Goksungu et al., 2011;

Choudhury et al.,2012;

Sena et al., 2006

Glucomannan

(heteropolysaccharide)

Sporobolomyces

salmonicolor Videva et al., 2010

Heteroglycan Bacillus licheniformis Patil et al., 2011

Alginate Pseudomonas

aeruginosa

Bylund et al., 2006; Leid

et al., 2005; Owlia et al.,

2007

Curdlan

(neutral homosaccharide

with glucose units)

Agrobacterium sp.,

Agaricus brasiliensis,

Alcaligenes faecalis

Shivakumar et al., 2006;

Jung et al., 2001; Shu et

al., 2007

Welan

(anionic polysaccharide) Alcaligenes sp. Barbara Vu et al., 2009

Gellan

(anionic polysaccharide)

Sphingomonas

paucimobilis,

Pseudomonas sp.

Bajaj et al., 2006;

Banik et al., 2000

Heparin

(sulfated

heteropolysaccharide)

Escherichia coli Barbara Vu et al., 2009

Cellulose

(homopolysaccharide) Acetobacter xylinum Jonas et al., 1998

Glucan Bradyrhizobium Adebayo et al., 2012;

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(homopolysaccharide) japonica,

Pleurotus

pulmonarius,

Ophiocordyceps

diterigena

Louch et al., 2001;

Kanokarn et al., 2010

Fructan

(neutral homosaccharide

with fructose units)

Streptococcus mutans Ogawa et al., 2011

Exopolysaccharide

(acidic

heteropolysaccharide)

Rhizobium meliloti Leigh et al., 1985

Lipopolysaccharide Serratia marcescens Allam et al., 2011

Exopolysaccharide

(homosaccharide with

glucose units)

Pseudoalteromonas

sp. Al Nahas et al., 2011

Poly glutamic acid Bacillus subtilis

Wu et al., 2007; Morikawa

et al., 2006; Stanley et al.,

2005

HePS-7

(heteropolysaccharide) Beijerinckia indica Wu et al., 2006

Capsular polysaccharide

(homosaccharide with

rhamnose units)

Burkholderia gladioli Kaczynski et al., 2006

Marginalan Pseudomonas

fluorescens Fett et al., 1989

Heteropolysaccharide

with pyranose units Enterobacter cloacae Jin et al., 2010

Heteropolysaccharide

with mannose units Klebsiella K32 Bryan et al., 1986

Mannoglucan

(heteropolysaccharide) Stemphylium Banerjee et al., 2009

Acetan Acetobacter xylinum Kranenberg et al., 1999

Succinoglycan

(heteropolysaccharide)

Rhizobium,

Pseudomonas,

Alcaligenes,

Agrobacterium sp.

Freitas et al., 2011

Hyaluronan

(heteropolysaccharide)

Pesudomonas

aeruginosa,

Streptococci sp.

Freitas et al., 2011

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2.5 Properties

2.5.1 Biofilm formation

Bacteria are formed by bacterial cells that are able to adhere to the substratum and

also capable of constructing themselves as complex cellular communities (Kolter and

Greenberg, 2006). Biofilms assemble on solid surfaces or as pellicles at air/liquid

interfaces. The composition of biofilms depends upon the environment in which the

biofilms are formed. An extracellular matrix present in biofilms holds the cells together

(Branda et al., 2005). The matrix typically consists of exopolysaccharides and proteins

and sometimes nucleic acid (Whitchurch et al., 2002). The mechanisms of the production

of the matrix differ from bacterium to bacterium suggesting that the formation of biofilm

is independent (Davies et al., 1998; Branda et al., 2005).

Biofilm impact on variety of environments, biofilms accumulation might lead to

flow blockage in industrial pipelines, water pipelines, medical devices such as catheters

and ventilators (Costerton et al., 1999). The process of microbial surface colonization

was described by Zobell in 1943. Later Busscher and Weerkamp hypothesized the

relation between the bacterial adhesion mechanism and distance of bacteria from the

surface. Van der waals forces such as reversible operate at a distance greater than 50nm

from the surface, both Van der waals forces and electrostatic interactions occur together

between 10-20nm from the surface. Lastly for the distance lesser than 1.5 nm between the

bacteria and the surface Van der waals forces, electrostatic interaction and specific

interactions together lead to irreversible binding between the bacteria and the surface.

Such closed association leads to the production of adhesive materials like EPS.

2.5.2 Adhesion to substratum

EPS that forms the structural matrix, greatly aids to increase the strength of the

adhesive bond and along with offering specificity of attachment in pathogenic bacteria

also provides a protective coat around the bacteria (Read and Costerton, 1987). Evolution � � ! " # $ % � % $ & ' ( ) # ! % � * + # $ ) ' * % ! + ! % * ' , - ' . % $ # * / % 0 � * + # * . % ! 0 ' 0 # 1 " � ! ! % 2 3 4 2 # $ ' - ! # % .would be strategically disadvantageous for bacteria to be restricted to specific types of

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substrata were diversity in the norm. Other substances produced by bacteria, proteins e.g.

outer membrane proteins; fimbriae and flagella (Timmermam et al., 1991) and

lipopolysaccharides are also implicated in bacterial adhesion.

2.5.3 Rheological properties

The intrinsic viscosity is the key parameter that describes the general 6 7 8 6 9 8 : ; < : = 7 8 > 9 ? 9 8 > 9 8 6 9 7 @ < 8 A B 9 C = D E 9 ? 7 E A F < 6 6 G < ; = > 9 HEPS imparts a sticky

consistency to bacterial growth on a solid medium or an increased viscosity in a liquid

medium. Viscosity depends on the combination of the polymer size, usually expressed as

the radius of gyration and the molar mass. The relation between the radius of gyration

and the molar mass is determined by the chain sti ness which depends on sugar

composition, type of linkages, charged groups, and degree of branching (Boels et al.,

2001). During most EPS production, the rheology of the fermentation broth changes

drastically from an initial Newtonian fluid behavior, with a viscosity near that of water, to

highly viscous fluid with shear thinning behavior (Freitas et al., 2011). This increase in

viscosity frequently causes a loss of bulk homogeneity, which makes it very difficult to

maintain appropriate mixing, aeration or control of bioreactor parameters.

2.5.4 Composition and linkages

EPS are mostly acidic in nature (Strednasky et al., 1999; Majumdar et al., 1999).

EPS contain a variety of negatively charged functional groups (e.g., carboxyl,

phosphoric, sulfate, and hydroxyl), as well as positively charged (e.g., amino) groups,

which together endow EPS with mostly negatively charged groups at neutral or slightly

alkaline pH. The microbial EPS may be non-ionic or may contain cationic, anionic and/or

both charges (Garnier et al., 2005; Celik et al., 2008; Osman et al., 1997). Microbial

polysaccharides are water soluble polymers and may be ionic or non-ionic. EPS are long

chain of polysaccharides consisting of branched, repeating units of sugars or sugar

derivatives mainly glucose, galactose or rhamnose in different ratios. They are classified

into two groups. Homopolysaccharides are repeating units of only one type of

monosaccharides (D-Glucose and / or D- Fructose) joined by either a single type of

linkage (1® 4/1®2) or a combination of a limited number of linkage types. Examples of

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homopolysaccharides are cellulose, dextran, mutan, pullulan and curdlan.

Heteropolysaccharides like gellan, xanthan are saccharides with multiple copies of

oligosaccharides containing 3 to 8 residues of sugar moieties, produced by a wide variety

of microorganisms. They are very common than homopolysaccharides and are more

diverse because of copolymerization of various monomeric units resulting in various

possible types of linkage, various repeats of monomers in chain and then relative ratio of

combination. The repeating units of these exopolysaccharides are very regular, branched

or unbranched, and are connected by glycosidic linkages (Banik et al., 2000).

Homopolysaccharides are common in Gram positive bacteria than in Gram negative and

are produced using extracellular enzymes but heteropolyasccharides are synthesized

inside the cell then excreted outside the cell onto the outer membrane (Ahmad and

Muhammadi, 2007). In organisms like Pseudomonas sp., presence of acetyl, succinyl and

pyruvyl groups influence the chemical composition and properties of EPS (Gianni et al.,

1999; Osman et al., 1997).

The composition and linkages are observed and analysed by FTIR and NMR

spectroscopic methods. An exopolysaccharide, AspY16, from a mangrove endophytic

fungus, Aspergillus sp. Y16, was analysed for its structural composition by NMR which

showed that it was composed of mannose with small amount of galactose, substituted at

C-6 by (1®K L E = 8 M 9 > N

-D mannopyranose, 1- E = 8 M 9 > O

-D galactofuranose and 1- E = 8 M 9 > O

-

D mannopyranose units (Chen et al., 2011). Paper chromatographic method also was

employed to study the composition of an acidic heteropolysaccharide which revealed that

EPS was made up of glucouronic acid, galactose, mannose and rhamnose (Ahmad and

Muhammadi, 2007). Functional groups of EPS from Bacillus cereus crs01 were found

using FTIR spectroscopy that displayed peaks representing primary and secondary

amines and amide groups, alkenes, ketones, alcohols, esters, ethers, carboxylic acids and

phenols. MonoP 9 ; F 7 @ N-D galactopyranosyl (1®2) glycerol phosphate were found be

present as indicated by the regions in NMR spectrogram of EPS secreted by

B.licheniformis (Sayem et al., 2011). FTIR spectrum displayed absorption peaks of

alcohol, amines, carbonyl and carboxyl groups in EPS isolated from Bacillus subtilis

DYU1. 1H and

13C Q NMR spectroscopic results indicated that the polysaccharide was

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related to poly glutamic acid (Wu et al., 2007). Biopolymer from Pseudoalteromonas

sp.9913 was composed of 6- linked glucose, other sugar units were present including

terminal arabinofuranosyl and glucofuranosyl residues and small amounts of other sugar

derivatives (Qin et al., 2007). NMR spectroscopic results showed that the polymer

synthesized by Bacillus thermoantarticus S < F P < > 9 T ? 7 @ N-

U P < 8 8 7 F 9 < 8 > O-D glucose

(Sayem et al., 2011). HPLC was also used to detect the composition of exopolysaccharide

generated by Astragalus F ? H S G = 6 G F G 7 S 9 > ? 9 < M F 7 @ N V W

®4) D glucan with (1®6)

branches attached to O-6 of branch points (Rui et al., 2009). 1, 3 linked galactoglucan

with galactose an X E T 6 7 F 9 = 8 N < 8 > O < 8 7 P 9 ; = 6 6 7 8 @ = X T ; < : = 7 8 F S 9 ; 9 = > 9 8 : = @ = 9 > = 8 Y Z [ @ ; 7 P

Pseudomonas marginalis HT041B using NMR (Osman et al., 1989). Sorangium

cellulosum NUST06 produced EPS which was composed of D-glucose, D- mannose and

D-glucuronic acid that were detected by NMR spectroscopic studies (Zhang et al., 2003).

2.6 Biosynthesis of EPS

Most bacterial EPSs are produced within the cell and exported extracellularly as

macromolecules. In a number of bacterial species, the EPS synthesis is controlled through

megaplasmids rather than chromosomally. Enzymes needed for the formation of EPS

precursors appeared to be under separate control from mechanisms of gene expression

associated with the EPS synthesis (Czaczyk et al., 2007).

In Gram negative bacteria, EPS are synthesized intracellularly whereas in gram

positive bacteria, the EPS like levan, dextran are produced extracellularly involving

extracellular lipoprotein enzymes secreted at the cell surfaces of Gram positive bacteria

such as Leuconostoc sp. (Sutherland, 1982; Freitas et al., 2011).

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Figure 2.3 Biosynthetic pathways involved in Gram negative bacterial EPS

Biosynthesis of exopolysaccharide in Gram negative bacteria is shown in Figure

2. The pathway comprises of substrate uptake, central metabolite pathway and

polysaccharide synthesis. Based on the type of the substrate used in the production, it

can be taken up by active or passive transport system. Crossing the membrane, movement

of sugars is coupled to the proton motive force through ATP- driven transport systems,

wherein ATP hydrolysis provides energy to drive the substrate against concentration

gradient (a). The substrate in the cytoplasm is then catabolized through intracellular

phosphorylation by glycolytic pathways and enters tricarboxylic acid cycle. The primary

metabolites obtained from these pathways are used as precursors for the synthesis of

primary macromolecules like proteins, lipids, ribonucleotides and hexosamines (b).

Polysaccharides synthesis requires the biosynthesis of activated precursors that are

energy rich monosaccharides, mainly nucleotide di phosphate Q sugars (NDP-sugars),

which can be derived from phosphorylated sugars. These precursors are interconverted

through reactions of epimerization, oxidation, decarboxylation, reduction and

rearrangement (c). In most of the Gram negative bacteria, polysaccharide synthesis and

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polymerization. Follow one of these two mechanisms Q Wzx-Wzy system and ABC

transporter system (d). In Wzx Q Wzy dependent mechanism, repeat units of sugars is

formed by sequential transfer of monomers from NDP- sugars to polyprenyl phosphate

lipid carrier. These units are transferred to periplasm by flippase (Wzx), where with the

help of polymerase (Wzy), polymerization to polysaccharide occurs. Polysaccharide

copolymerase (PCP) that determines the length of polymer chain translocates across

periplasm to cell envelope through a channel formed by an outer membrane

polysaccharide export protein (OPX). In ABC transporter system, polysaccharide is

polymerized in cytoplasm and then exported across the inner membrane through an ABC

transporter, followed by translocation of the polymer to outer membrane by PCP and

OPX proteins (Freitas et al., 2012; Rehm et al., 2010).

In Gram positive bacterial cells, exopolysaccharide production happens

extracellularly. One of the most studied Gram positive bacterial genus for EPS synthesis

is lactic acid bacteria whose production steps are explained thus. Homopolysaccharides

produced by them namely dextran, mutan, alternan and levan are synthesized

extracellularly using the specific substrate, sucrose. For the polymerization reaction, the

sugars requires specified glycosyl transferase enzyme for example, dextran sucrose for

dextran polysaccharide.

On hydrolysis of sucrose, energy is released which is used for the process.

Heteropolysaccharides are produced from the repeating sugar unit precursors present in

cytoplasm, with several enzymes. The sugar nucleotides, UDP-glucose, UDP galatose,

from sugar-1-phosphates play major role in their synthesis, as there are responsible for

sugar activation, polymerization and interconvertions by epimerization, dehydrogenation

etc.

I ]

Figure 2.4 Synthesis of EPS in Gram positive bacterium, Lactococcus lactis.

The numbers refer to the enzymes involved: 1, phospho-L-galactosidase; 2, L-

galactosidase; 3, glucokinase; 4, phosphoglucomutase; 5, UDP-glucose

pyrophosphorylase; 6, UDP-galactose-4-epimerase; 7, dTDP-glucose

pyrophosphorylase; 8, dehydratase; 9, epimerase reductase; 10, phosphoglucose

isomerase; 11, 6-phosphofructokinase; 12, fructose-1,6-bisphosphatase; 13, fructose-

1,6-diphosphate aldolase; 14, galactose 6-phosphate isomerase; 15, tagatose 6-

phosphate kinase; 16, tagatose-1,6-diphosphate aldolase.

Glucose -1- phosphate derived from glucose- 6- phosphate, an essential

metabolite from sugar catabolism, is mostly the main precursor for polysaccharide

formation. Phosphoglucomutase could be the key enzyme linking lactose degradation to

EPS generation. When lactose is used as the carbon source, Streptococcus thermophilus

and Lactobacillus delbrukeii bulgaricus release galactose to the medium through

lactose/galactose antiport system, hence only glucose act as the energy and carbon source

for the bacteria. Polymerisation of repeating sugar moieties occurs through sequential

addition of residues by their specific glycosyl tansferase from nucleotide sugars to

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undecaprenyl phosphate carrier in the cell membrane yielding the final EPS. Ultimately,

the synthesized polymer is translocated out of the cell and is excreted as slime or remains

attached as capsules (Degeest et al., 1999).

2.7 Isolation/Extraction Techniques

EPS isolation from the microorganisms relies on the bacterial strain and the

fermentation medium composition. EPS could be obtained from the cell free supernatant.

Generally the procedure involves precipitation using an alcohol with overnight incubation

after which the polymer would be removed by centrifugation followed by lyophilisation.

Various solvents had been used in precipitating EPS. Most commonly ethanol was used

for the purpose (Kanmani et al., 2011; Li et al., 2010; Llamas et al., 2010; El Shamy et

al., 2010; Ogambide et al., 1991; Kim et al., 2005). Also different other alcohols like

methanol (Wang et al., 2011; Borgio et al., 2009; Ko et al., 2000), isopropanol (Kanmani

et al., 2011; Patil et al., 2011; Liu et al., 2010) and butanol (Dai et al., 2010) were used to

sediment EPS. Acetone was also implemented for the process (Fett et al., 1989; Parikh et

al., 2006). Based on the complexity and nature of the EPS, purification steps were

improved. Chloroform had been used in combination with butanol in various studies to

remove lipid particles from EPS (Ko et al., 2000; Dai et al., 2010; Jin et al., 2010). Since

the cell free supernatant may contain many impurities protein has to be removed for

which trichloroacetic acid at varying concentration (4-20%) was used. The protein

materials settle at the bottom when left for overnight incubation. After centrifugation,

EPS was followed by precipitation (Madiedo et al., 2005, Al Nahas et al., 2011; Edward

et al., 2011; Hernandez et al., 2008).

Dialysis was carried out with membrane with molecular mass cut off ranging

from 3 -14 kDa (Tuncturk, 2009, Bylund et al., 2006; Ahmad and Muhammadi, 2007;

Orsod et al., 2012; Henandez et al., 2008). Deproteination was performed by enzymatic

digestion. Proteolytic enzymes were added to the culture supernatant to remove proteins

serving a pure EPS to be obtained. Pepsin (20U/ml) was added to B.licheniformis culture,

during EPS extraction (Ruffing et al., 2006). Pronase E, protease type XIV, extracted

from S.griseus, was used to digest milk proteins in lactic acid bacteria (Cerning et al.,

1986; Cerning et al., 1988). After digestion, the enzymes were heat inactivated, so that

I _

the enzymes would not intervene with polymer isolation. After heating at higher

temperatures, crude EPS solution was ultrafiltered or evaporated (Beshay et al., 2009)

followed by precipitation using alcohol. Heating may also let the complete recovery of

the biopolymer, if EPS is heat labile.

Other membrane filtration techniques had also been used in many other

researches using lactic acid bacteria to further purify EPS samples. Membrane filtration

included diafiltration, ultrafiltration, microfiltration (Yang et al., 2000; Levander et al.,

2001; Tuinier et al., 2001). Low molecular weighing molecules could be separated from

high molecular weight EPS molecules by these commercially available membranes with

varying porosity. Microfiltration was carried out using a ceramic membrane (pore size-

1.4mm, concentration factor of 10) and diafiltration with polysulfonate membrane with

20 volumes of distilled water (Madiedo et al., 2005).

Final and intensive purification was performed using ion exchange

chromatographic columns, which worked based on the ionic strength or affinity of the

macromolecules (Mody et al., 1987; Zhang et al., 2003; Zhao et al., 2010; Liu et al.,

2010; Chen et al., 2011) and also using Sephadex columns. The quantity of the final

biopolymer extracted relies on the isolation or extraction procedures.

2.8 Quantification and Characterisation of EPS

The freeze dried powder or lyophilized EPS had to be weighed and analyzed for

its quantity and structural characteristics respectively. The features of the polymer depend

on the strains of bacteria and the environment in which the culture was grown. The

disparity in quantity of the extracted biopolymer could be influenced by the presence of

proteins and non carbohydrate substituents in certain EPSs. A general procedure to

quantify the polymer is the colorimetric method using spectrophotometer. It could

estimate the amount of sugar and its related compounds by phenol sulfuric method as

described by Dubois et al, (1956). On experimenting, the results gave an orange yellow

color when treated with phenol and concentrated sulfuric acid. Using this protocol, mono,

oligo- , polysaccharides and their derivatives could be determined (Zhang et al., 2011a;

Kanmani et al., 2011; Shu et al., 2007; Allam et al., 2011).While total sugars were

I `

estimated by phenol sulfuric acid method, reducing sugars were analysed by DNS

method (Lee et al., 1999; Samain et al., 1997; Shu et al., 2007). Proteins and the non

carbohydrate substituents like uronic acids, pyruvates, acetates, succinate accompanying

the polysaccharide were also being estimated by different standard procedures (Kanmani

et al., 2011; Ahmad and Muhammadi, 2007; Chen et al., 2011).

The molar mass or molecular weight of the biopolymer often range between 1 Q 3

x 105 Da and they can be determined by various techniques like high performance liquid

chromatography with gel permeation columns with multiangle light scattering detectors

with refractive index detector (Osman et al., 1997; Luengo et al., 2003; Hilliou et al.,

2009), sedimentation and viscosity estimation (Luengo et al., 2003; Tuncturk, 2009;

Morris et al., 2001; Al Nahas et al., 2011; Torres et al., 2011). The composition and

structure of the polysaccharide define the primary conformation of the polymer (Madiedo

et al., 2005).

The monosaccharide of EPS was initially determined qualitatively by thin layer

chromatography (TLC) but it proved to a low intense technique (Madiedo et al., 2005).

The sugars were later estimated by HPLC with gel permeation columns and detection by

refractive index. Monomer composition of EPS produced by Lactobacillus sp. and

Streptococcus sp. were checked by isocratic elution using sulfuric acid at different molar

concentrations and temperatures (Grobben et al., 1998; De Vuyst et al., 1999; Madiedo et

al., 2005). The disadvantage of HPLC technique is the low resolving power and low

sensitive detection. The advanced techniques to determine monomeric units of EPS are

gas chromatography- mass spectroscopy (GCMS) and nuclear magnetic resonance

(NMR) spectroscopy.

Generally for GC MS analysis, the samples were hydrolysed with trifluoroacetic

acid or by methanolysis or converting them to trimethyl silyl derivatives (Samain et al.,

1997; Jung et al., 2001; DeVuyst et al., 2003; Zhang et al., 2003; Madiedo et al., 2005;

Ismail et al., 2010; Vijayabaskar et al., 2011; Chen et al., 2011). The resulting

monosaccharides were derivatised to alditol acetates and checked using GC with gas

columns, different temperature setup and carrier gases. The sugar units were identified

\ a

using flame ionization detectors or mass spectroscopy (Vijayabaskar et al., 2011). To

avoid derivatization, the samples can be used directly for analysis by high pressure anion

exchange chromatography with pulse amperometric detection (HPAEC-PAD) and also

capillary electrophoresis could be carried out without modifications of carbohydrates

(Degeest et al., 1999; DeVuyst et al., 2003; Madiedo et al., 2005). NMR (12

C and 1H)

study was carried out to learn the structure and conformation of the molecules and also

helps in elucidating the type of glycosidic linkages, the repeating units of the polymer.

The sample had to be deuterised before utilizing it for the examination (Osman et al.,

1989; Izawa et al., 2009; Liu et al., 2010). Physical properties like melting temperature,

crystal structures, polydispersity, rheological properties and mechanical properties could

also be analysed using various procedures (Luengo et al., 2003; Ismail et al., 2010).

2.9 Fermentation of EPS

Fermentation is an important condition in the biotechnological processes. Mode

of fermentation is one of the most important criteria for an industrial and economical

production of EPS (Tang et al., 2011). EPS is synthesized in lab and large scales by

different modes of fermentation namely batch, fed-batch, immobilization and continuous.

2.9.1 Batch fermentation

Batch fermentation is the most utilized mode of fermentation since it could be

easily generated by cultivating microorganisms in shake flasks in a laboratory scale and

in huge bioreactors for a pilot scale production. The fundamental idea of the batch

reaction is

Feed flow rate (input) = Product flow rate (output) = 0

Growth of culture during batch cultivation occurs in the following order: (1) lag

phase, (2) accelerated phase, (3) exponential or logarithmic phase, (4) decelerated phase,

(5) stationary phase and (6) decline or death phase.

\ b

Figure 2.5 Batch growth curve of microorganisms

The growth phase of the microbial cells plays a vital role in the batch

fermentation. The growth stages of surface associated bacteria may influence the quality

and chemical composition of the EPS. It was usually reported that the production of EPS

happened in the late exponential growth and stationary phase (Majumdar et al., 1999;

Petry et al., 2000; Czaczyk et al., 2007). Along with the EPS Production, the growth (cell

dry weight) also reached a maximum at 72 h and was evident that the production reached

the maximum towards the end of the log-phase and decreased in the stationary and

declining phase (Ismail et al., 2010). Recently it was found that most bacteria release the

highest quantity of EPS in the stationary phase of growth, this finding might be justified

by the competition occurring during the growth phase between EPS and cell- wall

polymer biosynthesis (Sutherland, 1982), however there are microorganisms that produce

the maximum amount of EPS during the exponential phase (Poli et al., 2010). In certain

cases highest EPS produced during exponential growth period which implied a better

correlation between cell multiplication and EPS formation (Mende et al., 2012). Growth-

associated EPS production was observed with L.plantarum (DeVuyst et al., 2003). Aslim

et al. (2005) explained that the EPS production increased during the exponential growth

phase and no further production was observed in the stationary growth phase (Ismail et

al., 2010).

\ J

Batch fermentation could be carried out in two ways Q solid state fermentation

(SSF) and submerged fermentation (SmF). Solid state fermentation is the process of

generating biological products cultivating microbes on a solid support which acts as

support and nutrient material. Submerged fermentation utilizes liquid medium for the

growth and production of desired compounds. SmF is carried out for cultures requiring

high moisture content (Subramaniyam et al., 2012).

In solid state fermentation of exopolysaccharides environment with high oxygen

content prevails for the microbes and solid support provides a natural habitat for the

microorganisms isolated from soil resulting in high product yield and cell growth.

Xanthan from Xanthomonas campestris and succinoglycan from Agrobacterium

tumefaciens and Rhizobium hedysari had been generated by SSF. The polysaccharides

formed by this process, are gradually absorbed into the porous substrate particles,

allowing free oxygen transfer through interparticle space and its diffusion through water

films surrounding the fermenting mass particles. Thus the bacterial cells adhering to the

surface of substrate revive unlimited oxygen quantities to perform metabolic activities.

But the prevalence of high oxygen may not a mandatory and influential parameter for the

production of EPS (Stredansky et al., 1999). A disadvantage of this process is that the pH

could not be controlled or lower acidification occurs. Another limitation is the recovery

of the product, which is trapped inside the solid materials.

Submerged batch fermentation has emerged as the preferred method for the

production of microbial EPS. Over the past 20 years, a rapid increase in interest is

observed in the large scale production of fungal EPS through SmF. Vantage of using this

process is that the flexibility of manipulating and optimizing environmental parameters

for the production of particular product. The process is a simpler one when compared

with SSF and could be carried out in liquid medium which aids in easy recovery of

product by mere centrifugation. This process helps out in learning the kinetics of cell

growth and product formation. Larger yields of biomass and product are obtained (Banik

et al., 2000; Kim et al., 2005; Kim et al., 2006; Ray and Moorthy, 2007; Onbasli et al.,

2008; Feng et al., 2010; Li et al., 2010; Yuan et al., 2012; Gaur and Singh 2010;

Chimilovski et al., 2011; Abdel-Aziz et al., 2012). In submerged cultures, filamentous

c d

organism may occur in pellet to dispersed forms depending on the culture conditions and

the genotype of the strains used. Aureobasidium pullulans exhibited yeast like growth

and gave a larger yield in submerged batch process (Campbell et al., 2004). Submerged

cultures can produce higher biomass in a compact space and shorter time with less

contamination (Chen et al., 2008a; Feng et al., 2010). For submerged batch aerobic

cultivation, it requires high carbon and low nitrogen conditions (Banik et al., 2000).

Batch processes are carried out both in shake flasks and bioreactors. Shake flasks

are comfortable when using in a lab scale production. Bioreactors are utilized in

industrial production and also in laboratories which could be especially beneficial to

study the kinetics. Production of EPS was found to be more efficient in bioreactors than

in flasks since the fermentation conditions especially agitation imparts homogeneity,

oxygen, mass transfer and pH could be controlled. Reports mentioned that EPS synthesis

had been better when culture was cultivated in bioreactors (De Vuyst et al., 1998;

Degeest et al., 2001; Mozzi, 1996; Bergmaier et al., 2003). Major disadvantage is that

when B.subtilis is to be grown for EPS synthesis, it should maintained in non-agitated

conditions, i.e., in still medium, for they form biofilms which is rich in EPS (Morikawa et

al., 2006). Optimisation studies in laboratories and downstreaming, could be studied

easily when carried out in shake flasks.

Consolidating all the advantages of submerged liquid fermentation, it had been

employed in this entire research work on the production of EPS from B.subtilis and

P.fluorescens.

2.9.2 Fed-batch fermentation

The strategy of Fed-batch fermentation is to add one or more of nutrients during

fermentation, based on the possibility that the high concentration required for the final

growth and product yields might inhibit growth if added in total at the start of the

process. Fed-batch culture is a batch culture which is fed continuously or sequentially

with substrate without the removal of fermentation broth. The Fed-batch operation may

be better than batch especially when concentration variations of one or more nutrients

affect the yield or productivity of the desired metabolite (Ramos et al., 2005). Compared

c e

with conventional batch culture, Fed-batch culture has several advantages, including

higher dissolved oxygen in the medium, decreased fermentation time, higher productivity

and reduced toxic effects of the medium components that are present at high

concentration (Roukas et al., 1997). Submerged mode of Fed batch fermentation is

generally preferred to solid state fermentation for large and lab scale processes (Wagner

et al., 2003; Kim et al., 2006; Champagne et al., 2007; Tang et al., 2011; Chen et al.,

2008b). Growth and product formation can be extended for long periods during fed batch

when compared to normal batch fermentation (Wagner et al., 2003).

An enhanced EPS and mycelia biomass production by fed batch culture was

observed with Ganoderma lucidum which were probably due to a increase in osmotic

pressure of culture medium or by prevention of a catabolite regulating effect exerted by

the rapidly degradable carbon source, glucose (Kim et al., 2006). Fed batch technology

was also performed to reduce the viscosity of the medium. In this process of biopolymer

production from Lactobacillus rhamnosus, medium viscosity increased with increase in

cell counts, but as the fermentation ended the capillary relative viscosity decreased to

40% when compared to batch cultivation (Champagne et al., 2007). EPS production by

mushroom, Tremella mesenterica was increased two folds when compared to batch

cultivation (De Baets et al., 2002). Fed-batch fermentation with high carbohydrate level

was successful in enhancing EPS production (Cheirsilp et al. 2003). The fed-batch mode

allows more flexibility in the control of the substrate concentration as well as product

concentration in the culture medium (Cheirsilp et al., 2006). But in case of L.

kefiranofaciens it was also found that high concentration of lactose and galactose,

product of lactose hydrolysis also inhibit cell growth (Cheirsilp et al., 2001).

Though fed batch mode of fermentation was found to be beneficial in production

of EPS from many microbial cultures, it has its own disadvantages too. Optimisation of

medium and production is complicated through fed batch mode and is cost intensive

(Cheirsilp, 2006). Batch culture proved to be a better fermentation system for the

production of pullulan than the fed batch culture system (Youssef et al., 1999).

c c

2.9.3 EPS production by Immobilisation

Immobilisation is the technique of attaching or entrapping cells on or within the

soild supports. This type of production process is commonly applied in many fields like

pharmaceutical, food and environmental (Gorecka et al., 2011). This technique has been

used widely in immobilizing enzymes and cells of plant, animals and microbes for

generating various metabolic products (Gorecka et al., 2011). Immobilised cell systems

are widely used to produce enzymes, antibiotics, alcohols, complex polysaccharides,

surfactants and food additives (Venugopalan et al., 2005). Microbial polysaccharides of

economic interest are usually produced at the industrial level by fermentation. Problems

associated with the reproducibility of fermentation during the industrial scale up may

result in inconsistent productivity, yield and quality. Increase in viscosity may result in

hard recovery. Hence immobilization is a useful technique to overcome such

inconveniences (Ismail et al., 2010).

The predominant advantages of using immobilized systems are reusability,

increased shelf life of cells for prolonged usage in fermentation, increased cell mass,

protection from hostile environment and damage, and stability (El Gizawy et al., 2013).

One common drawback of cell immobilization is the increase in mass transfer resistance

due to the polymer matrix (Ozdemir et al., 2005). Supports could be natural (agarose,

alginate, starch etc.), synthetic (styrene, polyacrylamide, vinyl etc.) polymers and carriers

like minerals, glass beads (El Gizawy et al., 2013). The physical characteristics like pore

size, swelling nature, mechanical strength and compression behavior are vital in

immobilized cell performance and choosing reactor like fixed bed, fluidized or stirred

tank, for studies. Porous supports are preferred to non-porous ones due to high surface

area leading to increased loading capacity. It should possess good control over pore

distribution so that loading and production could be optimized (Gorecka et al., 2011).

Different methods are available for immobilization namely adsorption and

entrapment. Adsorption with support materials happens by weak forces or covalent

bonding. This method is the most common and simplest reversible method of cell

immobilization. Based on weak forces like Van f g h i j j k l m n o h p g m q r o s r p t o s f m q u v f h o w g s

c x

bonding, hydrophobic interactions, disulfide bonds, chelation or metal binding or affinity

binding, cells bind with the solid support. Though certain advantages like easy

preparation, low cost and high reloading ability, this procedure possesses certain

disadvantages too. Weak interactive bonds increase leakage and low binding capacity.

Low reproducibility, less loading control, susceptibility to various environmental

parameters like pH, temperature and ionic strength are also the reasons for unhealthy

usage of this process. Some of the solid materials used to adsorb cells are porous glass

beads, ceramics, silica gel, sepharose, aluminium oxide, diatomaceous earth, carboxy

methyl cellulose, starch etc. (Gorecka et al., 2011).

Adsorption by covalent bonding is also advantageous in attaching cells with

matrix. It is an irreversible method providing high stable bonding and good contact

between cells and support. Two ways to increase the coupling are to add reactive

functional groups to support or to modify the polymer to activate for binding. The

disadvantage of this procedure is the damage to cells that might mostly happen during

bond formation (Gorecka et al., 2011; Bena and Viera 2006). Common matrices used for

this method are agarose, alginate, cellulose, poly vinyl chloride, ion exchange resins and

porous glass beads.

Certain other significant methods of irreversible methods of cell immobilization

are cross linking, entrapment and encapsulation. Cross linking can be made possible by

using bi or multifunctional reagents like glutataraldehyde. Chemicals used as cross

linking agents impart toxicity to cells which is the main disadvantage of this process.

Other biological processes like flocculation and pelletisation can be used for

immobilization due to strong adherent forces between cells and agents.

Entrapment is one of the most commonly used methods for immobilization of

microbial cells. This is the method where the cells are firmly trapped inside the support

matrix, from cells from diffusing into the external surrounding. Matrices that are used for

this procedure are agar, alginate, gelatin, collagen polyacrylamide, polyvinyl alcohol,

cross linking resins, polyesters, silicon rubber, polyurethane, sponge, wood (Berekaa et

al., 2009; Gorecka et al., 2011). The immobilized particle size to pore size ratio is the

c y

major criterion to be followed when immobilizing by this technique. Demerits of

entrapment are immobilization cost, diffusion limitations, abrasions of cells during usage

and low loading capacity (Gorecka et al., 2011). Encapsulation is a similar irreversible

process to entrapment. Here cells are trapped inside a semi permeable capsule like

structure made of polymers but the cells inside can freely float, with easy transfer of

nutrients, keeping the cells within the matrix. This is the most widely used technique in

lab scale processes. But this technique has some limitations like strict pore size control,

similarity of cell size and product size which might lead to leakage and accumulation of

products may cause rupture of capsule and eventually burst. Certain matrices used for this z h o p g f { h g j h g j k w r s j | g q p u r | o m j s q p g k k { k o m g q z o k v } r s v k j k p o u o k q w g k j | r s q ~ -carrageenan

(Gorecka et al., 2011).

Many investigations had been carried out on use of immobilization for EPS

production from various microbial cultures. This technique aids in recovery of cells by

simple filtration from production medium even with a higher viscosity as they are

entrapped within polymer and this is the main advantage (Champagne et al., 2007). Most

commonly used polymer in laboratories and for industrial purposes are alginate. Alginate

is a natural polymer produced by various brown algae and some of bacterial species like

Pseudomonas and Azotobacter (Emtiazi et al., 2004; Dimitrijevic et al., 2011). The

polymer is converted to hydrogels by cross linking with Ca++

ions. It is widely preferred

due to its biodegradability, hydrophilicity, and natural origin. Presence of COO- group in

alginate is the reasons of preferring alginate to other polymers like poly vinyl alcohol,

polyacrylates. Higher concentration of alginate might lead to increased death rate of cells

and lower sugar diffusivity into beads. Higher time of curing would harden the beads in

turn EPS release would be affected. Hence optimal conditions have to be maintained

(Ismail et al., 2010)

Alginate was used to immobilize C.luteola whose EPS was involved in heavy

metal adsorption. Increase in metal adsorption happened when EPS was immobilized in

alginate beads than free alginate beads (Ozdemir et al., 2005). Lactobacillus rhamnosus

RW-9595M was immobilized on commercial solid porous supports, ImmobaSil® and was

compared with free cells for EPS production during batch cultures under controlled pH.

c �

High biomass productivity was observed (1.7gL-1

) which in turn helped in maximum EPS

volumetric productivity (250mgL-1

h-1

after 7h) when compared to free cells EPS volume

(110 mgL-1

h-1

after 18h culture) (Bergmaier et al., 2003). The behavior of entrapped cells

depends on the nature of biological material in beads (Ismail et al., 2010). EPS produced

from Lactobacillus plantarum was found to be 4.5 folds increased (0.9gL-1

) when

encapsulated in alginate when compared to free cells (0.2gL-1

) in batch fermentation.

This is apparently due to the lesser competition between cells when entrapped than in

case of free cells (Ismail et al., 2010). Some reports had also put up contrast results.

Champagne et.al (1992, 1993, 2007) had studied extensively on biomass by using

alginate trapped growing lactic acid cultures, especially for cells which cannot resist

oxygen and centrifugation stresses. Immobilisation gave lower biomass yields, which was

a major drawback. When the total yield of bioreactor was considered, the total population

from the beads represented only 40% of that obtained in free cells in bioreactor. Viscosity

was also lesser than the free cells (Champagne et al., 2007). This could be a problem of

using this procedure for growth associated EPS production.

For EPS producing bacteria, immobilization method of adsorption on porous

supports is useful, since EPS produced could serve in adsorption and colonisation of the

cells on the matrix. But the quantification of polysaccharide concentration could be

difficult, which is a major limitation. To determine this, disruption of matrix would pose

shear stress upon cells in turn increase their mortality (Bergmaier et al., 2003).

Immobilised cells of Bacillus licheniformis had been utilized for the production of

polymer, poly-glutamic acid (PGA). In this study, two methods of immobilization were

involved� entrapment using agar, alginate, alginate-j w j h j s f ~ -carrageenan, and

adsorption on luffa pulp, sponge, pumice and wood. A quantity of 36.75gL-1

PGA was

obtained from alginate-agar entrapped cells and by adsorption 50 gL-1

PGA was obtained.

The study showed that adsorption was the best process of immobilization for EPS

synthesis (Berekaa et al., 2009).

Although immobilized cells had attained lot of attention in production of various

biological products, it is not possible to make a general statement about the behavior of

microorganisms in gel matrix or adsorbed on support material. Immobilised cells for

c �

production of EPS are carried out in batch and semi-continuous or fed batch bioreactors.

Since close packing of cells at high cell densities occur in immobilized cell systems, high

volumetric rates happen. In case of xanthan gum production, reduction in production

using free cells might be due to the higher mobility between cells than in immobilized

cells (Rosalam et al., 2008). In a bioreactor operation, immobilized cells ensure efficient

biomass retention and minimize cell loss through washout, thus downstream processing

becomes easier (Venugopalan et al., 2005). In order to investigate the possible reduction

in costs of PGA production, B. licheniformis cells adsorbed on sponge cubes were

cultivated in a trickle flow column in semi continuous manner. Relative viscosity and

EPS synthesis increased till the third day of process, 2.4 and 37.5gL-1

respectively

(Berekaa et al., 2009). Aureobasidium pullulans was immobilized in 4% agar cubes and

5% alginate beads and EPS generation was performed in an aerated column bioreactor.

Maximum production was observed in agar than in alginate. It had been suggested that

calcium ions may stimulate the synthesis of an alternate polysaccharide by alginate-

immobilised cells of A. pullulans (West et al., 2001). The composition of culture medium

can be readily monitored via an external loop and the concentrations of oxygen, sugar

and other physical parameters can be adjusted (Tyler et al., 1995). A continuous recycled

packed fibrous bed bioreactor with membrane separation technique was used for

producing xanthan gum from Xanthomonas campestris. Cells were let to adsorb on the

bed through batch fermentation and then the continuous production was commenced

later. Xanthan gum produced was 18 gL-1

by continuous fermentation and product was

extracted by membrane filtration (Rosalam et al., 2008).

2.10 Factors influencing EPS production

2.10.1 Effect of Carbon Sources

Carbon sources are one of the most influential nutrients for EPS production.

Fermentation medium of EPS should contain higher carbon content and decreased

nitrogen quantity (16). Many common sugars like glucose, fructose, sucrose, lactose,

starch and certain other sugars namely sorbitol, mannose, xylose, mannitol are tested for

their effect on EPS production (Shu et al., 2007). Non sugar sources like glycerol, n �

x �

alkanes, methanol had been used as carbon sources (Sivakumar et al., 2012; Chowdhury

et al., 2011; Mahecha et al., 2012). Different carbon sources might have specific catabolic

roles to play during secondary cellular metabolic activities (Chowdhury et al., 2011).

Glucose at an effective concentration between 2 and 3 % gave the maximum yield of

0.895gL-1

of EPS from Bacillus megaterium (Chowdhury et al., 2011). Himanshu et al.

reported that ratios of carbon and nitrogen sources play the most important role in cellular

growth and exobiopolymer production (Gandhi et al., 1997). Sucrose at a concentration

of 100 gL-1

was also found to be the best source for EPS production from B. licheniformis

221a, at 13.57 g EPS/L of medium (Tharek et al., 2006). Sugars like fructose, lactose,

glucose, and sucrose were used for EPS production in Streptococcus thermophilus ST1

from skim milk, yielding 64.52 mgL-1

, 66.39 mgL-1

, 69.35 mgL-1

and 73.28 mgL-1

,

respectively (Zhang et al., 2011). Higher yield with sucrose was obtained, since sucrose

apparently acts as a precursor of EPS synthesis. As the concentration of the sugars

increased above optimal levels, the cell growth and the yield were found to decline. This

is mostly due to the elevation of osmotic pressure in the cellular system, thereby causing

plasmolysis, leading to cell death (Kuntiya et al., 2010). Bacillus thuringiensis yielded a

maximum of 20.19gL-1

of biopolymer on fermenting with 10.45 gL-1

maltose as the

carbon substrate (Wang et al., 2011). A concentration of 2% maltose in the production

medium was able to produce 3.5 g EPS/L from Cordyceps jiangxiensis (Xiao et al.,

2004). A maximum of 44.49 mg/L of EPS was produced from Lactobacillus fermentum

when the medium was supplemented with 2% glucose and 0.5% whey protein

concentrate (Zhang et al., 2011). When Lactobacillus rhamnosus C83 was grown in a

medium containing 4% mannose or 2% glucose and fructose, EPS production was

increased by three or four times, but biomass concentration remained constant (Degeest

et al., 2001).

2.10.2 Effect of nitrogen sources

Various nitrogen sources had been employed in the enhanced synthesis of EPS.

Fermentation of EPs involved various organic sources like peptone, casein, yeast extract,

beef extract, and inorganic nitrogen sources like ammonium nitrate, ammonium citrate,

sodium nitrate and urea. From an extensive literature survey, organic nitrogen sources

x �

were inferred to yield a higher amount of EPS than inorganic nitrogen substrates. It was

suggested that certain essential amino acids cannot be synthesized from inorganic

nitrogen components (Wu et al., 2008), because of which bacterial cells might neither

fully grow nor undergo metabolism, and hence the deterioration of EPS yield. It was also

reported that the primary role of heterotrophic bacteria is classically considered to be

decomposition and mineralization of dissolved particulate organic nitrogen (Al Nahas et

al., 2011). Reports suggest that nitrogen limitation and higher amounts of carbon in the

medium could yield a maximum amount of EPS (Degeest and DeVuyst, 1999). A study

showed that EPS production from Rhizobium meliloti was higher when the nitrogen

source was in minimal quantity (Lee et al., 1997). Similarly, pullulan was generated by

Aureobasidium pullulans when it was grown in a medium with lesser amounts of

nitrogen source (Catley, 1971). A maximum production of EPS, from

Pseudoalteromonas sp., was observed on using meat extract as the nitrogen element at a

concentration of 10.51gL-1

(Al Nahas et al., 2011). Brevibacillus thermoruber 438

yielded a highest polymer content of 78. 1mgL-1

when the production medium was

supplemented with 0.1% diammonium hydrogen phosphate (Radchenkova et al., 2011).

Hirsutella sp. required 0.5% peptone in the fermentation medium to produce a maximum

of 2.17 gL-1

exopolysaccharide (Li et al, 2010). Malt extract was an essential nitrogen

requirement for the production of exopolymer by Ophiocordyceps dipterigena BCC2073

to yield a maximum of 41.2 gL-1

(Kocharin et al., 2010).

2.10.3 Effect of Carbon / Nitrogen ratio

A medium containing a high carbon and low nitrogen content in the fermentation

medium may favor polysaccharide production. In a study on EPS production from batch

biofilm reactors, yield of EPS was found to deteriorate as the C:N ratio declined. It was

reported that when C:N ratio was high, the bacterial cells alter their growth pathway, as

nitrogen components would not be available for protein synthesis. When the medium is

deficit of nitrogen and contain excess of carbohydrate, cells utilize energy from it to

biosynthesize polysaccharides (Miqueleto et al., 2010). Increased EPS production by K.

aerogenes and A. radiobacter was observed when excessive carbohydrates were present

in the medium which was used for producing energy currency, ATP. It is further shown

x �

that S. thermophilus LY03 produces a high-molecular-mass and a low-molecular mass

EPS, the proportion of which is dependent on the carbon/ nitrogen ratio of the

fermentation medium (Degeest and DeVuyst, 1999).

2.10.4 Effect of Industrial agricultural wastes as carbon sources

Recent investigations were carried out to produce EPS biotechnological

applications at a lower cost. For cost effective production, agro industrial wastes are used

as substrates (Muthusamy et al., 2008). Molasses is the final effluent obtained in

production of sugar by repeated crystallization (Olbrich, 2006). Sugarcane molasses

could be a better source of carbon due its higher content of total sugars at 48.3%. Due to

many advantages like high sucrose and other nutrient contents, low cost, ready

availability, and ease of storage, molasses has been used as a substrate for fermentation

production of commercial polysaccharides like curdlan, xanthan, dextran, scleroglucan,

and gellan (Mao et al., 2011). A fungus, Mucor rouxii, produced 87% EPS in medium

with 3% beet molasses (Abdel Aziz et al., 2012). Azotobacter was able to produce 7.5 mg

EPS/ mL of medium with 2% beet molasses (Goksungur et al., 2004). Jaggery, rich in

sucrose, had been utilized as carbon source at a concentration of 1% yielding 231mg EPS

per 100ml of medium (Sivakumar et al., 2012). Cheese whey, a dairy by product, rich in

lactose and other nutritive components had been used by B.indica ATCC 21423 for

heteropolysaccharide production, yielding a maximum of 6.18gL-1

(Wu et al., 2006).

Cordyceps sinensis produced large quantity of exopolysaccharides when grown in a

medium containing 1.5% rice bran, 0.5% molasses as carbon substrates and 3% corn

steep liquor as the nitrogen source. On addition of citrus peel to the medium composition,

an enhanced EPS productivity was observed (Choi et al., 2006).

2.10.5 Effect of vitamins and trace elements

Medium components such as minerals, some amino acids, and/or some bases and

vitamins are also found to affect the composition of the EPS produced. Mineral

requirements in trace quantities would favor or affect EPS production. It was reported

that certain minerals (Ca, Co2+

, Fe2+

, K+, and Mn

2+) were favorable to the mycelial

growth and EPS production of P. sinclairii, and as the concentration was increased, EPS

x d

was found to be increasing (Kim et al., 2002). Also, vitamins play an important role.

Vitamins perform a typical catalytic function on cellular metabolism as coenzyme or

constituents of coenzyme (Christiana A.V.Torres, 2012). In few cases, it was

demonstrated that the omission of multiple vitamins affected the production of EPS

relative to cell growth.

Salinity was an essential culture parameter for the production of higher amounts

of EPS. Like that observed with the sugars, the changes in salt concentrations caused

instability of osmotic pressure that led to detrimental effects on bacterial cells (Tharek et

al., 2006). Sivakumar et al. (2012) reported that 2-3% of NaCl was required to obtain

maximum amount of EPS from F. aurentia. When NaCl concentration was increased

from 10 to 30 gL-1

, EPS yield and growth of Pseudoalteromonas sp. increased as well,

however further increase in NaCl concentration to 40 gL-1

showed nearly the same yields

(Al Nahas et al., 2011).

2.10.6 Effect of pH, temperature and time

Environmental parameters - temperature and pH play a vital role in the synthesis

of exopolysaccharide. The amount of EPS production and properties are greatly

dependent on the microorganisms and their culture conditions such as temperature, pH

and media composition (Kanmani et al, 2011). They are the major factors controlling

microbial growth and metabolite synthesis (Torres et al., 2012). pH is a significant factor

influencing the physiology of a microorganism by affecting nutrient solubility and

uptake, enzyme activity, cell membrane morphology, by product formation and

oxidative- reductive reactions (Bajaj et al., 2009). Higher temperature could not help the

growth of the organism, since the stability of the cell structure might be affected. Since

Bacillus subtilis is mesophilic, decrease in temperature did not enhance the multiplication

of cells and since the biosynthetic pathway of exopolysaccharides would be inhibited

below optimal temperature (Sutherland, 2001), lesser production of EPS was observed.

Ismail et al. reported the maximum EPS production by Lactobacillus plantarum MTCC

9510 at 35ºC. Several reports show that low temperatures markedly induce slime

production. This effect has been explained, based on information for EPS production

from Gram-negative bacteria, by the fact that slowly growing cells exhibit much slower

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cell wall polymers biosynthesis, making more isoprenoid lipid carrier molecules available

for EPS biosynthesis (Degeest and DeVuyst, 1999). Incubation time is an essential factor

determining the enhancement of EPS synthesis in the culture. As EPS is highly

synthesized during late exponential growth phase or in the stationary phase, decrease in

incubation time may lower the production. Higher incubation time might affect the yield

due to the production of certain enzymes, saccharases, along with EPS, might act upon

polysaccharides, thus deteriorating the product formation. B.subtilis produced

polysaccharide (1.58±0.13mg/100ml) at 72h when temperature was 37ºC when cultured

in basal medium (Vijayabaskar et al., 2011). Optimal pH was 7-8 for the batch

fermentation of EPS, for 31 h, produced by B. polymyxa (33 g L-1

) in sucrose containing

medium. Relatively low concentration of EPS was obtained, during this study, when pH

lowered from 7 to 4.5, since the medium turned acidic (Lee et al., 1997). A temperature

range of 30-35ºC and pH 7 were found to be optimum for the production of exopolymer

by Bacillus sp. (Gandhi et al., 1997). Enterobacter sp. synthesized fucose containing

polysaccharides at a temperature range of 30-35ºC and pH ranging between 6 and 8

(Torres et al., 2012).

2.11 Applications

Exopolysaccharides find their own niche in varied fields of biotechnology and

several other industries. Based on their unique and specific composition they are utilized

for different purposes (Freitas et al.2011; Degeest et al. 2001). EPS is used in various

industries like pharmaceutics, food, textile and leather.

EPS play a vital role in human beings possessing various biological activities like

antioxidant, antimicrobials and in regulating the immune system as an

immunostimulating agent (Li, 2010). EPS produced by marine bacterial cultures could

induce regulatory cytokines in leukocytes and tissue cells (Arena et al, 2006). Sulfated

EPS have wide applications in the field of medicine as an anticoagulant, antiangiogenic,

antiproliferative agent (Llamas et al, 2010). They involve in absorption and penetration of

viruses into host cell and inhibit reverse transcriptase of retroviruses (Arena et al, 2006).

Phosphated EPS could be essential in the activation of lymphocytes and in certain

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antitumoral processes (Llamas et al, 2010). Cordyceps sinensis produced EPS that could

be involved as an anti complementary and in radical scavenging activities (Choi et al,

2010). Biopolymer synthesized by Bacillus licheniformis was found to be a potent

antiviral and an immnuoregulator (Arena et al, 2006). EPS produced by Paenibacillus

polymyxa ETS-3 had the ability to act as an antioxidant (Liu et al, 2010).

Exopolysaccharides from Astragalus sp. exhibited several biological activities like

immunomodulatory effect, antiinflammatory property and spleen lymphocyte

proliferation (Rui et al, 2009). Exopolymer synthesized by Hirsutella sp. exhibited high

antibacterial activity against Micrococcus tetragenus and Bacillus subtilis (Rong Li et al,

2010). Antioxidant activity and immunomodulating effect were exhibited by EPS

synthesized by Enterobacter cloacae Z0206 (Jin et al, 2010). The polymer from

Rhizobium sp.7613 showed a significant antitumor effect on mice bearing cancer cell

lines (Zhao et al., 2010).

Over a few years, the exobiopolymers have driven attention to treating wastewater

by removing of toxic metals based on metal binding capacities of bacteria, fungi, yeast

and algae as biosorptive materials. Bioflocculation is one method of removing heavy

metals from waste waters, since chemical or synthetic flocculating agents are toxic and

non ecofriendly. Bacterial bioflocculants are recommended as surface active agents or

biosurfactants for heavy metal absorption (Lin and Harichund, 2012). Negatively charged

EPSs act as flocculating agents forming bridges with the positively charged metal ions

eventually settle down as aggregated flocs, clearing water (Martins et al, 2008). BM07,

an exopolymer from Pseudomonas fluorescens, showed high metal ion binding capacity,

especially uptaking cadmium and mercury and playing an important role in toxic metal

absorption (Noghabi et al, 2007). Cr (VI) containing wastewaters from plating industry

could be treated using polysaccharides synthesized by cyanobacteria namely Cyanothece,

Cyanospira and Nostoc (Colica et al, 2010). Azotobacter EPS from A. indicus ATCC

9540 showed high efficiency on treating wastewater from industries like woolen, starch,

sugar and dairy industries (Patil et al, 2011). Efficient uptake of lead, zinc and cadmium

was observed by a novel bacterial exopolysaccharide from Alteromonas macleodii

(Martine Loaec et al, 1997).

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Bioremediation of oil polluted soil or environment is processed by wider range of

microorganisms and some EPS are employed as biosurfactants and in detoxification

mechanism of petrochemical oil polluted areas (Poli et al, 2010). The organisms use the

hydrocarbons and carbon as the nutritive sources for their growth and emulsification-

abling-exopolymer production. Emulsifiers are produced by microorganisms to

breakdown the non soluble oil or hydrocarbons and facilitating their uptake (Toledo et al,

2008, Satpute et al, 2010) Species of Acenitobacter showed significant emulsifying

activity on oil in water emulsions. Emulsan are the most powerful emulsifiers secreted by

the bacterial species (Belsky et al, 1979; Kaplan et al, 1982). Pseudomonas nautica

generated EPS composed of carbohydrates and proteins exhibited effective emulsifying

activity (Husain et al, 1997). Emulsion stabilizing activity was displayed by

exopolysaccharide generated by Bacillus sp. CP912 (Yun et al, 2000). Kerosene and

crude oil were efficiently emulsified by EPS synthesized by Rhodotorula glutinis (Oloke

et al., 2005). Gordonia alkanovorans CC-JG39 produced a potent emulsifier which could

degrade diesel and also acted as biostimulant for bioremediation of oil contaminated

water or soil (Chen et al, 2008a). Calvo et al investigated that sulfated

heteropolysaccharide isolated from Halomonas eurihalina possessed increased

emulsification property (Calvo et al., 1998). EPS secreted by Brevibacillus brevis was

potentially involved in treating hydrocarbon pollution and used in microbially enhanced

oil recovery (Ebrahimi et al., 2008). Yansan, an exopolymer extracted from Yarrowia

lipolytica, emulsified oil in water emulsions effectively (Amaral et al., 2006).

Bioremediation of hydrocarbon imparted soil was carried out efficaciously by using

heteropolysaccharide secreted by Microbacterium testaceaum (Edward et al, 2011).

In food industries, EPS find many applications such as stabilising, thickening,

emulsifying, viscosifying and gelling agents (Torres et al., 2012). Xanthan, produced by

Xanthomonas campestris, is the most commonly available commercial and first

industrialized polymer used as a viscosity enhancer, since it is a highly viscous solution

with entangled polymer chains (Freitas et al., 2011; Moshaf et al., 2011). Fucose, one rare

sugar molecule is commercially used as FucoPol is used as source of valuable chemicals

(Freitas et al., 2011). Sphingomonas elodea, S. paucimobilis are some of the potent

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producers of gellan gum, which is most widely used as gelling agent, thickening and

viscosifiers. Gellan gum has distinctive advantage in fruit based products due to its acid

stability (Bajaj et al., 2006). Hyaluronic acid from Streptococcus equii, succinoglycan

from Rhizobium had also been applied in the food industries (Banik et al., 2000). Kefiran,

produced by Lactobacillus kefiranofaciens, L. delbrukeii, Streptococcus thermophilus are

in dairy industries as viscosifiers and to produce traditional carbonated, slightly alcoholic

fermented milk (Ginka et al., 2002; Cheirsilp, 2006; Habibi et al., 2011; Vu et al. 2009).

Alginate, carrargeenan and agar are extracellular polysaccharides, produced by brown

seaweeds and Pseudomonas aeruginosa. These find great applications in food industry as

food additives (Brownlee et al., 2009).