biosynthesis, characterisation, and design of bacterial

Research review paper

Biosynthesis, characterisation, and design of bacterial

exopolysaccharides from lactic acid bacteria

Andrew Laws*, Yucheng Gu, Valerie Marshall

School of Applied Sciences, University of Huddersfield, Queensgate, Huddersfield HD1 3DH, UK

Abstract

Lactic acid bacteria (LAB) are characterised by their conversion of a large proportion of their

carbon feed, fermentable sugars, to lactic acid. However, in addition to lactic acid production, the LAB

are able to divert a small proportion of fermentable sugars towards the biosynthesis of

exopolysaccharides (EPSs) that are independent of the cell surface and cell wall material. These

microbial EPSs when suspended or dissolved in aqueous solution provide thickening and gelling

properties, and, as such, there is great interest in using EPSs from food grade microorganisms (such as

the LAB that are traditionally used for food fermentations) for use as thickening agents. The current

review includes a brief summary of the recent literature describing features of the biosynthetic

pathways leading to EPS production. Many aspects of EPS biosynthesis in LAB are still not fully

understood and a number of inferences are made regarding the similarity of the pathway to those

involved in the synthesis of other cell polysaccharides, e.g., cell wall components. The main body of

the review will cover practical aspects concerned with the isolation and characterisation of EPS

structures. In the last couple of years, a substantial number of structures have been published and a

summary of the common elements of these structures is included as is a suggestion for a system for

representing structures. A brief highlight of the attempts that are being made to design ‘tailor’-made

polysaccharides using genetic modification and control of metabolic flux is presented. D 2001 Elsevier

Science Inc. All rights reserved.

Keywords: Exopolysaccharides; Lactic acid bacteria; Biothickener; Oligosaccharide; Bacterial polysaccharide;

NMR

0734-9750/01/$ – see front matter D 2001 Elsevier Science Inc. All rights reserved.

PII: S0734 -9750 (01 )00084 -2

* Corresponding author. Tel.: +44-1484-472-668; fax: +44-1484-472-182.

E-mail address: [email protected] (A. Laws).

Biotechnology Advances 19 (2001) 597–625

1. Exopolysaccharides of lactic acid bacteria

Exopolysaccharides (EPSs) are long-chain polysaccharides that are secreted mainly by

bacteria and microalgae (Sutherland, 1972, 1977) into their surroundings during growth and

that are not permanently attached to the surface of the microbial cell. The physical

characteristics of EPSs are responsible for the slime-forming or mucoid trait of many

microorganisms. A second group of polysaccharides that are structurally similar but that

are permanently attached to the cell surface are classified as capsular polysaccharides

(Sutherland, 1985). The current review will focus exclusively on EPSs and capsular

polysaccharides will not be discussed here.

The ability of microorganisms to synthesise EPSs has both positive and negative attributes.

The food industry uses polysaccharides from plant and seaweeds for their thickening and

gelling properties. However, existing supplies are either not sufficiently reliable, are of

variable consistency, or the quantities available are not sufficient to match demand. In

response a number of polysaccharides of microbial origin have been developed including

xanthan from Xanthomanas campestris (Garcia-Ochoa et al., 2000) and gellan from

Sphingomonas paucimobilis (Banik et al., 2000; Giavasis et al., 2000; Sutherland, 1999).

These can be prepared in reliable quantities using conventional biotechnological processes.

However, the physical properties of these polymers are such that they are not suited to all

applications and there is a demand for novel materials that impart improved rheological

characteristics. One of the main drawbacks with using microbial polysaccharides in food

formulations is the requirement for them to be considered as a food additive.

Many lactic acid bacteria (LAB) are routinely used in food preparations for their

preservative effects: the acidification, resulting from sugar metabolism, restricts further

microbial contamination. LAB isolated from Scandinavian ropy fermented milk products

are known to produce EPSs (Forsen and Myllymaa, 1974; Forsen and Pakkila, 1979; Macura

and Townsley, 1984). These EPSs provide thickening properties, and are also considered to

improve the texture and mouthfeel of dairy products (Marshall and Rawson, 1999). LAB are

generally regarded as safe and EPSs isolated from LAB offer an alternative source of microbial

polysaccharides for wider use in food formulations. In addition to their use in food production,

there are a number of reports of possible health benefits of EPSs, and especially in regard to the

immunostimulating properties of bacterial EPSs (Oda et al., 1983; Schiffrin et al., 1995).

The negative attributes of EPS synthesis are associated with their spoilage properties. The

synthesis of EPS by LAB during wine (Denadra and Desaad, 1995; Lonvaud-Funel, 1999)

and cider (Duenas et al., 1995) production leads to products having undesirable rheological

properties. The formation of dental plaques is related to EPS synthesis by LAB (Loesche,

1986; Rozen, 2001). The EPSs from LAB are responsible for biofilm formation (Oliveria,

1992; Poulsen, 1999) that can lead to biofouling (Azeredo and Oliveira, 2000). The most

notable examples of biofouling are associated with biofilm formation in the equipment used

for the processing of dairy products. Biofilms cause a significant number of technical and

hygiene problems for the dairy industry (Poulsen, 1999).

Before trying to understand the biosynthesis of bacterial EPSs, it is important to realise that

bacteria synthesise a number of different classes of polysaccharide and that a significant

A. Laws et al. / Biotechnology Advances 19 (2001) 597–625598

proportion of these polysaccharides are used for the construction of components of the cell

wall. It is worth pointing out at this early stage that many of the mechanisms involved in the

synthesis of EPS are either known, or predicted, to be shared with those of other cell wall

components. The biosynthesis and the functionality of the main cell wall components have

recently been reviewed (Delcour et al., 1999).

Microbial EPSs can be divided into two groups: homopolysaccharides (e.g., cellulose,

dextran, pullulan, levan, and curdlan) and heteropolysaccharides (e.g., gellan and xanthan).

Homopolysaccharides are constructed from monosaccharides joined by either a single linkage

type (e.g., 1–2 or 1–4) or by a combination of a limited number of linkage types (e.g., 1–2

and 1–4). The current review will focus on recent work related to the synthesis and structural

characterisation of heteropolysaccharides from LAB. Heteropolysaccharides are constructed

from multiple copies of an oligosaccharide. The oligosaccharide can contain between three

and seven residues, it possesses a variety of two or more different types of monosaccharides

and frequently has a range of different linkage patterns. Current interest in aspects of EPS

synthesis is large, and consequently, there have been a number of reviews covering work in

this area (de Vuyst and Degeest, 1999; Ricciardi and Clementi, 2000; Cerning and Marshall,

1999; Sutherland, 1998; Cerning, 1990). In the current review, many aspects of the early

‘groundbreaking’ studies will not be covered and the author apologises to the many

researchers in the field whose efforts may appear to be treated without due reverence. This

article will attempt to review more practical aspects of current research and will try to paint a

picture of the current knowledge in the area. Readers interested in further development of the

subject are directed to the reviews listed above and particularly to that of Cerning (1990).

2. Biosynthesis of heteropolysaccharides by LAB

The biosynthesis of bacterial EPSs is complex and involves the concerted action of a large

number of gene products. The genes coding for the enzymes and regulatory proteins required

for EPS synthesis are of plasmid origin in the mesophilic LAB strains, e.g., Lactococcus and

chromosomally based in the thermophilic strains of Streptococcus and Lactobacilli. The EPS-

producing ability of LAB is regarded as being unstable. For mesophilic LAB strains, the

unstable nature of EPS synthesis is consistent with the genes for EPS synthesis being plasmid

bound. For the thermophilic LAB strains, it has been proposed that the loss of EPS-producing

character is due to deletions and rearrangement resulting from genetic instability. Identifica-

tion of EPS gene clusters and suggestions for the functional role/s of the gene products have

been reported for a number of food grade microorganisms: Streptococcus thermophilus Sfi 6

(Stingele et al., 1996), Streptococcus thermophilus Sfi 39 (Germond et al., 2001), and

Lactococcus lactis NIZO B40 (van Kranenburg et al., 1997) (Fig. 1). In addition to EPS-

specific gene products, the biosynthetic pathway relies on a number of the so-called

housekeeping enzymes such as those required for the preparation of sugar nucleotides.

The biosynthetic pathway can be broken down into four separate reaction sequences. These

are the reactions involved with sugar transport into the cytoplasm, the synthesis of sugar-1-

phosphates, activation of and coupling of sugars, and the processes involved in the export of

A. Laws et al. / Biotechnology Advances 19 (2001) 597–625 599

the EPS. These are schematically represented in Fig. 2. Each of these aforementioned

sequences must be evaluated in developing strategies for the engineering of EPS products.

2.1. Sugar transport into the cytoplasm

The movement of carbon feeds, mainly monosaccharides and disaccharides, from the

surrounding growth medium into the cytoplasm is a carefully regulated process. A number of

different proteins control the internalisation of sugars. Sugar transport and its possible

influence on the engineering of EPS structure has been reviewed (de Vos, 1996). The most

frequently encountered sugar transport machinery is that of the bacterial phosphoenolpyr-

uvate (PEP)–sugar phosphotransferase system (PTS) (Postma et al., 1993). The PEP–PTS

system contains a group of proteins that are responsible for binding, transmembrane transport,

and phosphorylation of a variety of sugar substrates. In addition, a number of other proteins

exist that regulate the activity of each of the latter processes.

The first group of proteins includes enzyme I, the histidine-containing phosphocarrier

protein HPr and a carbohydrate-specific permease enzyme complex (enzyme II) (Viana et al.,

2000). These enzymes act in sequence to provide phosphorylated sugars in the cytoplasm.

The sequence is initiated when a phosphate group is transferred from PEP to enzyme I;

enzyme I subsequently phosphorylates a histidine residue of HPr to yield HPr(His-P) (Postma

et al., 1993). At the same time, a protein of the enzyme II complex binds the carbohydrate

(Sliz et al., 1996, 1997). The carbohydrate-specific enzymes (II) transport sugars across the

membrane and catalyse the transfer of the phosphate group from HPr(His-P) onto the sugar.

In the PEP–PTS transport system of L. lactis, three Type II enzymes are required for lactose

transport: enzymes IIA–C (Wang et al., 2000). Enzymes IIA and IIB are located in the

cytoplasm and enzyme IIC acts as a membrane channel. On first inspection, the requirement

for a number of enzymes for the transport of a phosphate group from PEP to a sugar appears

over elaborate; however, the complexity of the process is required by the need for tight

regulation of nutrient acquisition.

A second set of proteins is responsible for regulation of nutrient acquisition. These

regulatory proteins do not only activate and repress PEP–PTS systems, they also regulate

non-PTS uptake systems (Poolman et al., 1997; Stulke and Hillen, 1998; Gunnewijk and

Fig. 1. Organisation of the EPS gene cluster of (A) L. lactis ssp. cremoris NIZO B40 (plasmid localised) (Stingele

et al., 1999) and (B) S. thermophilus Sfi6 (chromosomally encoded) (van Kranenburg et al., 1997). The proposed

function of the different gene products is indicated.


Fig. 2. (A) Schematic representation of a number of possible pathways for sugar transport and metabolism in LAB. (B) Schematic representation of a

possible pathway for EPS biosynthesis in L. lactis NIZO B40 starting from glucose-6-phosphate (the site of polymerisation of the repeat unit has not been

established and may occur on either face of the membrane). Known enzymes are indicated using either accepted nomenclature for the enzyme or by reference

to their encoding genes and using established genetic nomenclature. The figure is a highly adapted one from those presented by (Kleerebezem, 2000).

A.Lawset

al./Biotech

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19(2001)597–625

601

Poolman, 2000a,b; Gunnewijk et al., 1999). Most notable of the regulators is the binary

complex formed between the catabolite control protein CcpA and the PTS protein HPr (van

den Bogaard et al., 2000). The histidine-containing phosphocarrier protein HPr has two

phosphorylation sites and His(Ser-P) has been shown to regulate sugar transport; CcpA in

combination with HPr(Ser-P) binds to the cis-acting DNA sequence termed the catabolite-

responsive element (cre). Jones et al. (1997) demonstrated that CcpA only binds phosphory-

lated HPr(Ser-P), which suggests that catabolite repression is linked to active sugar transport.

Alternative, non-PEP–PTS, transport systems exist for the import of sugars, e.g., primary

and secondary transport systems. A number of LAB do not have active PEP–PTS transport

system for all the sugars that they are able to internalise. In the absence of a sugar-specific

PEP–PTS transport system there is a requirement for active transport. In S. mutans, multiple

sugars are transported by a primary transport system (Tao et al., 1993; Sutcliffe et al., 1993).

In S. thermophilus, sugar transport is mainly by a secondary transport mechanism coded for

by lacS (Friesen et al., 2000a,b). lacS is able to import lactose in symport with protons or,

alternatively, lacS can function as a lactose–galactose antiport system.

2.2. Synthesis of sugar-1-phosphates

Once inside the cytoplasm, the fate of the carbon feed is determined by the state of

phosphorylation of the sugar: sugar-6-phosphates are consumed in catabolic pathways

whereas sugar-1-phosphates can participate in polysaccharide synthesis. As the majority of

sugars are transported into the cytoplasm by PEP–PTS systems, which generate sugar-6-

phosphates, a number of authors have pointed to the possible key role that phosphoglu-

comutases (PGMs) may play in diverting flux between the two pathways (Sjoberg and

Hahn-Hagerdal, 1989; Degeest and de Vuyst, 2000).

Early work on PGMs focused on the role they play in the catabolism of sugar-1-

phosphates (Qian et al., 1994). In the metabolism of maltose, a sugar that is transported

via a permease in L. lactis, the disaccharide is initially converted into one molecule of

glucose and a molecule of b-glucose-1-phosphate. If the b-glucose-1-phosphate is to be used

in the catabolic pathway, it must subsequently be converted to glucose-6-phosphate and this

reaction is catalysed by b-PGM (Qian et al., 1997). In a similar manner, it has been suggested

that a-PGMs may play exactly the opposite role in EPS biosynthesis, i.e., in diverting the

glucose-6-phosphate to a-glucose-1-phosphates and EPS biosynthesis (Sjoberg and Hahn-

Hagerdal, 1989; Degeest and de Vuyst, 2000). It has recently been demonstrated that for the

synthesis of EPS in L. lactis grown on glucose, a proportion of the carbon feed must be

converted to glucose-6-phosphate for EPS synthesis (Ramos et al., 2001).

A number of alternative pathways leading to a-glucose-1-phosphate have been suggested

(see bottom right-hand side portion of Fig. 2 and the discussion below). The mode of

synthesis of glucose-1-phosphate is dependent on a number of variables: most notably, the

carbon source on which the culture is grown and the transport system used for sugar import. It

has been proposed (Grobben et al., 1996) that for L. bulgaricus grown on fructose that

fructose is imported via a PEP–fructose PTS that specifically yields fructose-1-phosphate.

Fructose-1-phosphate is converted, via fructose-1,6-bisphosphate, to fructose-6-phosphate, to


glucose-6-phosphate, and finally to glucose-1-phosphate. A much shorter pathway to glucose-

1-phosphate is available for L. bulgaricus grown on glucose; glucose is internalised via a

PEP–glucose PTS yielding glucose-6-phosphate that is transformed directly into glucose-1-

phosphate (Grobben et al., 1996). When L. lactis is grown on lactose (de Vos and Vaughan,

1994), lactose is imported by a lactose-specific phosphotransferase transport system providing

internal lactose-6-phosphate. Lactose-6-phosphate is subsequently hydrolysed by a phospho-

b-galactosidase to generate galactose-6-phosphate and glucose. A glucokinase is required for

the synthesis of a-glucose-6-phosphate. In contrast, galactose-negative S. thermophilus

internalises lactose by coupling a lactose permease within an antiport secondary transport

system (Poolman, 1993). In the latter system, it is necessary to use a combination of a

glycosidase and kinase to generate glucose-6-phosphate.

Given the variety of routes leading to glucose-1-phosphate, it might be expected that a

pathway should be available for the synthesis and subsequent utilisation of galactose-1-

phosphate. This is indeed the case and utilises the enzymes of the Leloir pathway. The enzymes

of the Leloir pathway convert galactose to a-galactose-1-phosphate (GalK), a-galactose-1-phosphate to UDP-galactose (GalT) and UDP-galactose to UDP-glucose (GalE) (see Fig. 3). It

is worth noting that for galactose-grown cells, the presence of UDP-glucose pyrophosphorylase

and a supply of UTP would allow the reaction to be considered as catalytic in UDP-glucose.

The combined action of the enzymes could supply both UDP-glucose and UDP-galactose

without a requirement for a PGM. However, it is well known that a number of dairy strains of S.

thermophilus lack an available galactokinase; though, under appropriate conditions, mutants

that are able to ferment galactose are selected indicating that the galactokinase genes are present

but are not transcribed (Thomas and Crow, 1984; Hutkins et al., 1985).

2.3. Synthesis of sugar nucleotides and polymerisation into the EPS repeat unit

It is instructive, at this point in the reaction sequence, to split the genes coding for the

proteins required for EPS biosynthesis into two groups: genes required for the synthesis of

sugar nucleotides and EPS-specific genes. These two groups are physically separated in the

genome, and in the case of L. lactis, are further removed from each other as the genes for EPS

biosynthesis (upper part of Fig. 2) are extrachromosomal. The first group consists of the

Fig. 3. Conversion of galactose to UDP-galactose using enzymes of the Leloir pathway. Enzymes involved in the

conversions are indicated.


genes coding for enzymes and proteins required for the synthesis of sugar nucleotides from

which the repeat unit is constructed. Sugar nucleotides are needed for the synthesis of a range

of polysaccharides and are not specific to EPS biosynthesis and, as such, are frequently

termed ‘housekeeping’ enzymes (see middle of Fig. 2).

The sugar nucleotides required for the construction of the majority of EPS structures are

UDP-glucose, UDP-galactose, and dTDP-rhamnose: the precursors of the repeat unit. The

genes coding for the enzymes needed for the synthesis of the sugar nucleotides from glucose-

1-phosphate (galU, galE, rfbA, rfbB, rfbC, and rfbD) have been identified and cloned from L.

lactis strain MG1363 (Boels et al., 1988; Reeves, 1993; Kleerebezem et al., 1999). The

function of the respective gene products is illustrated in the middle section of Fig. 2. The first

enzyme in the sequence is GalU, a UDP-glucose pyrophosphorylase. It has been reported

(Kleerebezem et al., 1999) that the intracellular levels of UDP-glucose is determined by the

activity of the enzyme GalU; functional overexpression of the lactococcal galU gene results in

much larger UDP-glucose levels in L. lactis. The production of UDP-Gal is believed to be

derived principally from UDP-Glc through the action of GalE that catalyses the interconver-

sion of the two UDP-sugars. The requirement for GalE for EPS biosynthesis in L. lactis NIZO

B40 was demonstrated by Kleerebezem et al. (1999): a galE mutant was not able to synthesise

EPS when grown on glucose but EPS was produced when the mutant was grown on galactose.

The latter result implies that, in the absence of galactose, UDP-galactose required for EPS

synthesis is derived solely from UDP-glucose. Details of the nature of the enzymes needed to

produce dTDP-rhamnose were established in Gram-negative bacteria where rhamnose is a key

constituent of the O antigens of lipopolysaccharides (Reeves, 1993). Four enzymes, RfbA,

RfbB, RfbC, and RfbD, convert a-glucose-1-phosphate initially to dTDP-glucose then to

4-keto-6-deoxymannose and finally to dTDP-rhamnose.

The next stage in EPS biosynthesis uses the EPS-specific enzymes. The first gene clusters

coding for production of secreted EPSs to be identified and characterised were those from S.

thermophilus Sfi 6 (Stingele et al., 1996) and for L. lactis NIZO B40 (van Kranenburg et al.,

1997). In S. thermophilus Sfi 6, the gene cluster is located on the chromosome, is 14.5 kb in size,

and contains 13 genes. In contrast, the L. lactis gene cluster is located on a 40-kb plasmid, the

EPS gene cluster is 12 kb in size and contains 14 genes. The organisation of the gene clusters is

similar for both organisms and consists of four separate domains (see Fig. 1). A central core

coding for the glycosyl-transferases is flanked at the ends by genes coding for proteins having a

strong homology with enzymes used for polymerisation and export. A regulatory domain is

present at the start of the gene cluster. Stingele et al. (1999) have demonstrated that the gene

cluster contains EPS-specific enzymes by insertion of the cluster into the non-EPS-producing

heterologous host L. lactis MG1363 and demonstrating EPS synthesis.

In vitro experiments using [14C]-labelled sugar nucleotides have provided evidence that

the monosaccharide repeat unit is assembled on a lipid carrier, which is attached to the

cytoplasmic membrane (van Kranenburg et al., 1997, 1999). In O antigen synthesis, the

repeat unit construction occurs on the inside surface of the cytoplasmic membrane (see top of

Fig. 2). For L. lactis NIZO B40, the first sugar to be attached to the membrane-anchored

phosphorylated lipid is glucose. Significantly, the [14C] glucose is attached through its

anomeric carbon by a pyro-diester link. The mode of action of the ‘priming’ glycosyl-


transferase, a UDP-glucose transferase for L. lactis NIZO B40, requires donation of the sugar

and a phosphate group to the lipid. In L. lactis NIZO B40, the second sugar in the repeat unit

is glucose and this is added, with a b-glycosidic link, through the attachment of the anomeric

carbon of a-UDP-glucose to the 4-hydroxy group of the lipid-bound glucose. This process

requires two EPS gene products, EpsE and EpsF. The final backbone residue, a b(1–4)-linked-galactose, is derived from UDP-galactose and the coupling is catalysed by a single

gene product EpsG. van Kranenburg et al. (1999) suggest that the branches, a rhamnose and

a phosphogalactose, are added to the 2- and 3-positions of the repeat unit in successive

glycosyl/phosphoglycosyl-transferase-catalysed steps generating the required repeat unit

joined by a diphosphate link to the lipid. Whether the sugar is incorporated as an a- or

b-glycoside is dictated by the catalytic mechanism of the glycosyl-transferases. These

enzymes have different catalytic domains, and some may catalyse insertion such that there is

retention of the anomeric configuration whilst others will result in inversion of the

configuration (Saxena et al., 1995).

The assembly of the repeat unit on a lipid carrier is a process that is used for the synthesis

of excreted polysaccharides, for cell wall peptidoglycans and for cell surface oligosaccharides

and polysaccharides. There is evidence to suggest that the various oligosaccharide and

polysaccharide syntheses use the same building blocks (sugar nucleotides) and scaffolding

(lipid carrier). The latter may account for the close relationships between rates of EPS

synthesis and cell growth that have been observed by a number of authors (Garcia-Garibay

and Marshall, 1991; Cerning et al., 1992).

2.4. EPS polymerisation and export to the surrounding medium

Details relating to the events leading to the polymerisation of the repeat unit and its export

from the plasma membrane through the peptidoglycan layers of Gram-positive bacteria are

very scarce. Gonzalez et al. (1998) have studied the genes involved in polymerisation and

export of the EPS succinoglycan in Sinorhizobium melilot, formerly Rhizobium meliloti. These

authors concluded that the subunits are constructed on an undecaprenol lipid carrier on the

cytoplasmic face of the plasma membrane, which are polymerised in a block fashion. Details

of the mechanism by which blocks are polymerised in EPS biosynthesis are not known. In O

antigen synthesis, three gene products are required for polymerisation and export (Whitfield

and Valvano, 1993). The gene products code for a protein that catalyses the movement of the

lipid-bound material form the cytoplasmic face of the membrane to the periplasmic face

(flippase or translocase), a protein that catalyses the polymerisation of the blocks (polymerase)

and a protein responsible for controlling polymer chain length.

By analogy, a simple model for EPS polymerisation and export requires the action of a

flippase to translocate the lipid-bound repeat units, a polymerase to catalyse the coupling of

repeat units and finally an enzyme to catalyse the detachment of the lipid-bound polymer and

that will control chain length. Evidence for the identification of the proteins responsible for

the catalytic functions described above come from comparison of sequences of genes and

from hydrophobicity cluster analysis of the derived amino acid sequences of similar proteins.

In succinoglycan synthesis, there is strong evidence to suggest that the gene product from


exoQ is the polymerase (Gonzalez et al., 1998). ExoQ proteins show sequence and

topological similarities with the O antigen polymerase gene (Wzy(Rfc)) product (Whitfield

and Valvano, 1993). Becker et al. (1995) have suggested that the ExoP product of R. meliloti

is involved in chain length determination.

In L. lactis NIZO B40, the genes epsA and epsB show homology with the R. meliloti ExoP

genes and the gene products are proposed to be involved in chain length determination (one

protein may detach the polymer whilst the second may regulate the activity of the first). EpsK

and EpsI are homologous to the flippase and polymerases in Salmonella or Shigella O antigen

synthesis (Morona et al., 1994; Liu et al., 1996).

3. EPS structure and yield

In determining the structure and yields of the EPSs, care should be taken in the choice of

techniques used for the preparation, isolation, and characterisation of the EPS. Early EPS

literature inevitably contains many examples where the presence of contaminants, both low-

molecular mass material and high-molecular mass polysaccharides, are included in calcu-

lations of monomer compositions and of yield. Many concluding statements regarding EPSs

having variable composition, dependent on the carbon feed, may be false and are likely to

simply be a measure of the extent to which foreign polysaccharides have been isolated with the

EPS. Particular caution should be taken when the quantities of the EPS recovered are small and

are of an order of a few milligrams. At the same time, it is worth noting that many of the

physical and chemical processes involved in the isolation and characterisation of polysac-

charides can seriously influence structure and yield of the EPS (see discussion below). There

are a number of experimental practices that can be adopted in order to avoid some of the simple

errors that are frequently reported. This review will briefly cover those areas of concern.

3.1. Preparation of EPSs

The starting point for EPS production is the preparation of a culture inoculum and this is

the first point at which contaminants may be added to the system. Master cultures are

frequently prepared from broths that contain polysaccharides. Degeest and de Vuyst (1999)

and Marshall et al. (2001b) generate inocula for use in fermentations using a minimum of two

subculturing steps. The subculturing steps are necessary to remove, via dilution, unwanted

high-molecular mass material. Many of the reports of presence of trace amounts of sugars

other than rhamnose, galactose, glucose, and N-acetylglucosamine in EPS compositions are

most likely false: the sugars being derived from polysaccharides originally present in the

starting inoculum culture broth.

Fermentation conditions greatly influence EPS production. There are a number of reports

suggesting that for a variety of LAB strains, mainly mesophilic strains, that EPS production is

greatest under conditions that are not optimal for growth (Cerning et al., 1992; Gassem et al.,

1995; Gamar et al., 1997; Schellhass, 1983; Mozzi et al., 1995). The latter result is consistent

with both EPS and cell wall biosynthesis pathways utilising the same membrane lipid.


Optimum temperatures for EPS synthesis have been determined for a number of LAB.

Cerning et al. (1992) have shown that the optimum temperature for EPS synthesis is 25 �C for

L. lactis. For S. thermophilus temperatures less than 37 �C are optimal for EPS biosynthesis.

EPS production in Lb. rhamnosus is high between 20 and 25 �C whilst growth is optimal

between 30 and 37 �C (Gamar et al., 1997). For Lb. delbrueckii ssp. bulgaricus RR, EPS

synthesis is increased at temperatures above (45 �C) and below (30 �C) that at which

optimum growth is observed (Schellhass, 1983). In contrast, for Lb. acidophilus and Lb. casei

EPS production is at a maximum at the optimum growth temperature (Mozzi et al., 1995;

Petry et al., 2000).

The literature relating to the influence of pH and medium composition on EPS production

has recently been reviewed (Ricciardi and Clementi, 2000; Cerning and Marshall, 1999) and

only a brief summary will be presented here. In studies of the influence of pH fermentations

are performed either using controlled pH or the pH is allowed to fall. Optimum pH values

vary from one species of LAB to another and, for a specific species, the pH optimum is

specific to that organism.

One aspect of EPS production that has only recently received attention is the degradation

of EPS that is observed after extended fermentations. Glycosyl-hydrolase activities, that

reduce the molecular mass, have been observed in fermentations of Lb. rhamnosus R (Pham

et al., 2000). A rapid reduction in molecular mass would require an endoglucanase to be

active in the cell supernatant. The influence that the pH of the medium has on EPS yield will

be dependent on the pH optimum of the glycosyl-hydrolases. The pH optimum for EPS

production will be that pH at which the opposing effects of production and degradation are

balanced. The effect of medium composition on EPS production is very marked. The

influence of medium components on EPS yields has received a great deal of attention and

has recently been reviewed (Ricciardi and Clementi, 2000). In our laboratories, the most

convenient medium for production of EPS is either unsupplemented skim milk or skim milk

supplemented with small proportions of casein hydrolysate. Attempts to isolate and character-

ise EPS from broth media or media to which peptone or yeast have been added as sources of

added nitrogen have been problematic. Polymannans present in broth, yeast, and peptone are

extracted with the EPS providing enhanced yields and monomer ratios that include high

percentages of mannose.

Only a limited number of studies of the kinetics of EPS production have been carried out

(Gassem et al., 1997a,b; Bouzar et al., 1996; de Vuyst et al., 1998; Kimmel and Roberts,

1998; Gancel and Novel, 1994a,b). Production of EPS appears to occur during the

logarithmic phase and, for some LAB, continues into the stationary phase.

3.2. Isolation of polysaccharides

Isolation of polysaccharide is required for their characterisation, and the procedure should

not alter the chemical and physical properties of the polysaccharide. Two different isolation

procedures are in common use. The only substantial difference between the two methods is

the way in which milk proteins, caseins, are removed. The first method to be reported is due

to Cerning et al. (1986, 1988) and utilises pronase to hydrolyse caseins. In the second


procedure, cell material and milk caseins are precipitated from the culture medium on the

addition of trichloroacetic acid (Garcia-Garibay and Marshall, 1991). These extraction

procedures have not been optimised and a number of workers use slight variants:

particularly in regard to the final concentration of added trichloroacetic acid that varies

between 4% and 20%.

There are a number of reports of LAB that synthesise mixtures of EPSs. The EPSs can

have different structures: Duenas-Chasco et al. (1998) have supplied strong evidence for a

strain that secretes two homopolysaccharides having different structures. There is also the

possibility of recovering EPS samples that have identical structure but different molecular

masses (Degeest and de Vuyst, 1999). The literature available describing the ability of LAB

to secrete mixtures of heteropolysaccharides of different structure should be read carefully.

Frequently, the only evidence used to support such mixtures comes from analysis of the

monomer composition of samples isolated by fractionation. Great care should be taken when

interpreting such data. This is particularly true when small samples are being analysed (see

discussion above). The preparation and isolation procedures may well provide contaminating

polysaccharides that are either originally present in the medium, have been added as yeast or

peptone supplements or that have been removed from cell walls either as a result of

mechanical disruption or lysis during fermentations.

The results of monomer composition ratios that are determined in chemically defined

media are more reliable (Petry et al., 2000). Grobben et al. (1997) have used a defined

medium to grow Lb. delbrueckii ssp. bulgaricus NCFB 2772. When grown on glucose or

fructose, two EPSs are isolated (Grobben et al., 1996): high-molecular mass polysaccharide

and a low-molecular mass polysaccharide. The monomer composition and linkages reported

for the large molecular mass polysaccharides derived from glucose and fructose are, to a first

approximation ( ± 5%), the same (Grobben et al., 1997). However, the low-molecular mass

polysaccharides, which are produced in small amounts, do appear to have a different

monomer composition.

As stated earlier, evidence confirming that a single LAB culture produces two EPSs that

have different repeat unit structures is only available for Lactobacillus spp. G-77 (Duenas-

Chasco et al., 1998) an organism that secretes two homopolysaccharides. Both monomer

composition and NMR spectral data are available for each of the EPSs. NMR evidence to

confirm that a single LAB culture produces more than one heteropolysaccharide repeat unit is

currently not available. To date there have been no complete structural studies published that

show that LAB are able to secrete more than one heteropolysaccharide that have different

structures. On a number of occasions we have observed NMR spectra that indicate that more

than one type of EPS is present; however, by the use of careful controls (performing

fermentations and isolation procedures without bacteria), we have demonstrated that the

spectra contain resonances that are characteristic of the media (M17) used to culture the

bacteria (S. thermophilus).

In contrast, there is substantial evidence for the synchronous production of EPSs having

the same structure but of different molecular mass. Degeest and de Vuyst (1999) reported the

production of a high-molecular mass (1.8� 106) and a low-molecular mass EPS (4.1�105)

by S. thermophilus LY03. The production of two polysaccharides by Lb. rhamnosus has


recently been reported (Pham et al., 2000); however, the low-molecular mass material is

believed to be generated by the glycosyl-hydrolase-catalysed hydrolysis of high-molecular

mass products.

3.3. Characterisation of EPSs

Before a polysaccharide can be considered to be fully characterised, it is necessary to

determine information about the molecular mass of the material, to identify the composition

and absolute configuration of the monomers, and finally, to determine the linkage pattern of

the monomers.

A number of methods for determining the molecular mass of polysaccharides are available.

Historically, retention times, as determined by size exclusion chromatography using refractive

index detection, have been used to determine molecular mass (Cerning et al., 1986).

However, the majority of EPS samples have very large molecular masses and elute close

to the exclusion limits of the available stationary phases and very much in excess of the

highest molecular mass standards used to calibrate the columns. Combination detectors

(Williams et al., 1992; Tinland et al., 1988), using light scattering for measurement of

molecular mass (Wyatt, 1993), provide a more accurate average molecular mass.

A number of different methods are available for determining the monomer composition of

EPS samples. Methanolysis and per-trimethylsilylation provides samples that can be analysed

by GLC. A simpler method, requiring acid hydrolysis followed by monomer detection using

high-pressure anion exchange chromatography with pulsed ampometric detection, has

recently been introduced (Cataldi et al., 2000; Hanko and Rohrer, 2000). Absolute config-

urations of monomers are determined by preparation of their per-trimethylsilyl (� )butyl

glycosides using the procedures described by Gerwig et al. (1978, 1979).

The linkage pattern of the monomers is determined using a combination of ‘methylation’

analysis (Stellner et al., 1973; Sweet et al., 1975) and NMR spectroscopy. There have been a

very large number of reviews covering the use of NMR spectroscopy in carbohydrate

chemistry, and the most recent by Duus et al. (2000) lists the reviews published between 1992

and 1999. Many of these have collated reference spectroscopic data, chemical shifts, and

coupling constants, and are extremely useful for assigning spectra of complex carbohydrates.

A review describing the application of 2D-NMR in determining the primary structure of

bacterial EPSs has recently been published (Leeflang et al., 2000).

In summary, 1D-1H and 13C spectra are recorded for solutions of EPS samples in

deuterium oxide and spectra are recorded at temperatures of 70 �C or above: the high

sample temperature reduces the viscosity of the medium and shifts the residual proton solvent

signal up-field and away from important resonances. Vliegenthart et al. (1983) have

suggested that spectra should be viewed as having ‘structural reporter signals’ and a ‘bulk

region’ (3–4 ppm). In proton spectra the low-field reporter region includes resonances from

the anomeric and ring atoms shifted out of the bulk region as a consequence of glycosylation.

There is also a high-field reporter region including resonances from ring substituents such as

acyl, alkyl, and acetal, and ring substitutions such as N-acetylamino and H6 of 6-deoxy

sugars. Homonuclear 2D spectra are used to assign protons resonances to individual rings


Table 1

Structures of the EPS repeat unit isolated from the genus Lactococci (Lactococcus lactis)

Group Origin Author Structure

L-3A B891 van Casteren et al.

(2000a,b)

L-3B cremoris

H414

Gruter et al. (1992)

L-3C SBT0495 Nakajima et al. (1992)

B40 van Kranenburg et al.

(1997)

cremoris

ARH74

Yang et al. (1999)

L-5A B39 van Casteren et al.

(2000a)

(Ac)0.5#6

b-D-Galp-(1! 4)-b-D-Glcp1

#6

! 4)-b-D-Galp-(1! 4)-b-D-Glcp-(1! 4)-a-D-Glcp-(1!


#3

! 4)-b-D-Galp-(1! 3)-b-D-Galp-(1! 4)-a-D-Galp-(1!

a-D-Galp-1-phosphate#3

! 4)-b-D-Glcp-(1! 4)-b-D-Glcp-(1! 4)-b-D-Galp-(t!2

"1

a-L-Rhap


#4

! 2)-a-L-Rhap-(1! 2)a-D-Galp-(1! 3)-a-D-Glcp-(1! 3)-a-D-Galp-(1! 3)-a-L-Rhap-(1!

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(COSY and TOCSY) and heteronuclear 2D spectra, 13C–1H, are used to assign carbons

(HMQC, HMBC) and to obtain linkage information (HMBC). Further information about

linkage, using nonscalar coupling, is available from ROESY spectra.

A number of techniques that have recently been applied to carbohydrate analysis but that

have not yet been routinely applied to the analysis of EPSs are worth highlighting. Vincent

and Zwahlen (2000) have reported the use of spin diffusion to provide information relating to

linkage and the technique, which requires shaped pulses, is complementary to the hetero-

nuclear HMBC experiments. Navarini et al. (2001) have reported the use of deuterium-

induced differential isotope shifts in the determination of linkage type. The latter technique is

particularly valuable when 2D experiments and shaped pulses are not available. A method for

undertaking linkage analysis by direct inspection of the NMR of oligosaccharides that have

been per-acetylated using 13C (CO)-labelled acetic anhydride has been reported (Bendiak,

1999a,b). The latter experiment, if applicable to polysaccharides, may remove the need to

undertake classical methylation analysis.

At the time of writing, this review the complete structural analysis, for polymers that

have been described both as being secreted EPSs and as heteropolysaccharides, have been

reported for 6 lactococci (Table 1), 21 streptococci (Table 2), and 19 lactobacilli (Table 3).

A small but significant number of LAB secrete the same EPS and in total 25 unique

structures have been reported. Each genus, Lactobacillus, Streptococcus, and Lactococcus

can be divided into groups according to the EPS they synthesise (see Tables 1, 2, and 3,

respectively). A convenient method for assigning EPSs from the different taxa into groups

is based, in the first instance, solely on the number of monosaccharides present in the

backbone of the oligosaccharide repeat unit. Each grouping in the table has been coded

using initials that identify the genus (S for Streptococcus, L for Lactococcus, and LB for

Lactobacilli). For lactobacilli, species are identified by an additional initial, e.g., H for

helveticus. The initial is followed by a number that reflects the number of residues in the

main chain of the repeat unit. Similarities between the EPS structures can only be easily

identified by visual inspection if a common mode of writing structures is adopted. The

latter requires the adoption of a set of rules for presenting structures. In the author’s

laboratory, the following rules have been developed and are loosely derived from the

guidelines presented by the International Union of Pure and Applied Chemistry (IUPAC)

for the nomenclature of polysaccharides composed of more than one kind of residue

(2-Carb-39.7):

1. The repeat unit is drawn such that the sugar residues of the back bone (principal chain)

is on the horizontal axis and the residue of highest priority is on the right-hand side with

the anomeric carbon at the end (see below for suggestions for assigning priorities to

sugar residues). The principal chain is drawn using standard IUPAC abbreviated

nomenclature for naming residues and for identifying linkage patterns.

2. If a single branch is attached to a sugar residue it is drawn above the principal chain.

3. If more than one branch is attached to a sugar residue the highest priority branch is

drawn above the main chain and the lower priority branch is placed below (see below

for suggestions for assigning priorities to branches).


Table 2

Structures of the EPS repeat unit isolated from the genus Streptococcus

Family Origin Author Structure

S-3A CNCMI 733,

734, 735

Doco et al.

(1990)

IMDO 01,

02, 03

and NCFB

859

Marshall et al.

(2001a,b)

EU 21 Laws

(unpublished)

Sfi 6 Stingele et al.

(1999)

S-3B Sfi 39 Lemoine et al.

(1997)

SY 89 Marshall et al.

(2001a,b)

SY 102

CH101 Laws

(unpublished)

S-3C Sc136 Vincent et al.

(2001)

a-D-Galp1

#6

! 3)-a-D-GalpNAc-(1! 3)-b-D-Galp-(1! 3)-b-D-Glcp-(1!

b-D-Galp1

#6

! 3)-b-D-Galf-(1! 3)-a-D-Glcp-(1! 3)-b-D-Glcp-(1!

b-D-Galf-(1! 6)-b-D-Glcp-(1! 6)-b-D-GlcpNAc1

#3

! 4)-b-D-Glcp-(1! 4)-a-D-Glcp-(1! 4)-b-D-Galp-(1!

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S-5A Sfi 12 Lemoine et al.

(1997)

S-5B OR901 Bubb et al.

(1997)

Rs and

Sts

Faber et al.

(1998)

S-5C S3 Faber et al.

(2001b)

S-6A EU20 Marshall et al.

(2001a)

b-D-Galp-(1! 6)-b-D-Galp1

#4

! 2)-a-L-Rhap-(1! 2)-a-D-Galp-(1! 3)-a-D-Galp-(1! 3)-a-D-Galp-(1! 3)-a-L-Rhap-(1!

Ac0.4#2

b-D-Galf1

#6

! 3)-a-L-Rhap-(1! 2)-a-L-Rhap-(1! 2)-a-D-Galp-(1! 3)-b-D-Galp-(1! 3)-a-D-Galp-(1!

a-L-Rhap1

#2

! 4)-b-D-Glcp-(1! 6)-a-D-Galf-(1! 6)-b-D-Glcp-(1! 6)-b-D-Galp-(1! 6)-a-D-Galp-(1! 3)-b-L-Rhap-(1!

b-D-Galp1

#4

! 2)-a-L-Rhap-(1! 2)-a-D-Galp-(1! 3)-a-D-Glcp-(1! 3)-a-D-Galp-(1! 3)-a-L-Rhap-(1!A.Lawset

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Table 3

Structures of the EPS repeat unit isolated from the genus Lactobacillus

Group Origin Authors Structure

Lactobacillus

helveticus

LB-H-3A K16 Yang et al.

(2000)

LB-H-4A TN-4 Yamamoto

et al. (1995)

Lh-59 Stingele

et al. (1997)

LB-H-4B TY1-2 Yamamoto

et al. (1994)

LB-H-5A 2091 Staaf et al.

(1996)

b-D-Glcp1

#2


#6



#3

! 3)-a-D-Galp-(1! 3)-a-D-Glcp-(1! 3)-b-D-Glcp-(1! 5)-b-D-Galf-(1!


#6

! 6)-b-D-Glcp-(1! 3)-b-D-Glcp-(1! 6)-a-D-GalpNAc-(1! 3)-b-D-Galp-(1!4

"a-D-Galp0.8

b-D-Galp1

#6

! 4)-b-D-Glcp-(1! 6)-b-D-Glcp-(1! 6)-b-D-Galp-(1! 4)-a-D-Galp-(1! 3)-b-D-Galp-(1!

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LB-H-5B Lb 161 Staaf et al.

(2000)

LB-H-5C 766 Robijn

et al. (1995)

Lactobacillus

LB-A-4A Lb.

acidophilus

LMG 9433

Robijn et al.

(1996a)

LB-D-3A Lb.

delbrueckii

ssp.

bulgaricus

291

Faber et al.

(2001a,b)

LB-D-4A Lb.

delbrueckii

ssp.

bulgaricus

RR

Gruter et al.

(1993)

EU03,

EU24,

EU25

Marshall

et al.

(2001a,b)

(continued on next page)

b-D-Glcp b-D-Glcp1 1

# #3 3

! 3)-b-D-Glcp-(1! 4)-a-D-Glcp-(1! 4)-b-D-Galp-(1! 3)-a-D-Galp-(1! 2)-a-D-Glcp-(1!

b-D-Galf1

#3

! 6)-a-D-Glcp-(1! 6)-a-D-Galp-(1! 6)-a-D-Glcp-(1! 3)-b-D-Glcp-(1! 4)-b-D-Glcp-(1!

b-D-GlcpNAc1

#3

! 4)-b-D-Glcp-(1! 4)-b-D-GlcpA-(1! 6)-a-D-Glcp-(1! 4)-b-D-Galp-(1!

b-D-Galp-(1! 4)-b-DGlcp1

#6


b-D-Galp a-L-Rhap b-D-Galp1 1 1

# # #4 3 3

! 3)-b-D-Glcp-(1! 3)-b-D-Galp-(1! 4)-a-D-Galp-(1! 2)-a-D-Galp-(1!

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LB-P-4A Lb.

paracasei

34-1

Robijn et al.

(1996b)

LB-P-4B Lb.

paracasei

van Calsteren

(2001)

two strains

LB-R-6A Lb.

rhamnosus

RW 9595

M and R

van Calsteren

et al.

Table 3. (continued )

Group Origin Authors Structure

sn-glycerol-3-phosphate

#3

! 6)-b-D-Galp-(1! 6)-b-D-Galp-(1! 3)-b-D-GalpNAc-(1! 4)-b-D-Galp-(1!

a-D-Galp1

#6

! 6)-a-D-Galp-(1! 3)-b-L-Rhap-(1! 4)-b-D-Glcp-(1! 4)-b-D-GlcpNAc-(1!3

"1

a-L-Rhap

HOOC 4

R a-D-GalpH3C 6 1

#2

! 3)-a-L-Rhap-(1! 3)-a-L-Rhap-(1! 2)-a-D-Glcp-(1! 3)-a-L-Rhap-(1! 3)-b-D-Glcp-(1! 3)-a-L-Rhap-(1!

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LB-R-5A Lb.

rhamnosus

C83

Vanhaverbeke

et al. (1998)

LB-R-5B Lb.

rhamnosus

GG

Landersjo

et al. (2001)

LB-S-3A sake 0–1 Robijn et al.

(1996a,b)

! 6)-a-D-Galp-(1! 6)-a-D-Glcp-(1! 3)-b-D-Galf-(1! 3)-a-D-Glcp-(1! 2)-b-D-Galf-(1!

b-D-Galf1

#6

! 3)-a-L-Rhap-(1! 3)-a-D-Galp-(1! 3)-b-D-Galf-(1! 3)-b-D-Galp-(1! 4)-a-D-Glcf NAc-(1!

sn-glycerol-3-phosphate! 4)-a-L-Rhap(Ac)0.85 1

# #2 3

! 3)-b-L-Rhap-(1! 4)-b-D-Glcp-(1! 4)-a-D-Glcp-(1!6

"1

b-D-Glcp

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The different sugar residues of the principal chain are assigned priorities as follows:

1. The sugar residue (including its attendant substituents and branches) of highest mass has

highest priority.

2. For sugar residues that have equal priority, after application of Rule 1, furanose sugars

are assigned priority over pyranose.

3. For sugar residues that have equal priority, after application of Rules 1 and 2, the residue

having linkages (locants) closest to the anomeric carbon has highest priority.

The different branches are assigned priorities as follows:

1. The branch of highest mass has highest priority.

2. If two branches are of equal mass then the branch linked closest to the anomeric carbon

has highest priority.

Whilst it is appreciated that the guidelines presented above will not allow definitive

structures to be drawn for all the different possible combinations of EPS structures they are

extremely useful for observing similarities between the different EPS structures and for

attempting to group structures into ‘families’.

The structures presented in the tables in this article are drawn using the guidelines.

Inspection of the different repeat unit structures (Tables 1–3), allows a number of conclusions

to be drawn regarding the frequency at which structural elements are observed:

� Taking both furanose and pyranose rings into account, the monosaccharide present in

highest frequency is galactose followed closely by glucose and then, at a much lower

frequency, rhamnose.� GalNAc, GlcNAc, and GlcA are observed in a small number of structures; as are the

phospho-diester, acetyl-ester, phosphate-ester, and pyruvate-acetal substitutions.� Without exception glucose and galactose have D-absolute configuration whereas

rhamnose has L-absolute configuration.� For glucose and galactose, there is a small preference for the b-anomer; however, for

rhamnose there is a definite preference for a-rhamnose: the ratio of the two anomers is

approximately 4a to 1b.� The only sugar that is found to adopt a furanose ring is galactose.

The frequency with which the different linkages, anomeric configuration, and linkage type,

are found is worthy of comment and, again, a number of conclusions can be drawn:

� A large number of branches are terminated by the addition of galactose to the branch,

over 75%.� b-D-Gal is preferentially attached either as a branch terminus or linked to the chain via

its 3-, 4-, or 6-hydroxyl group. In contrast, a-D-Gal shows a strong preference for

attachment through its 3-hydroxyl group.


� b-D-Glc is preferentially attached via either its 3-, 4-, or 6-hydroxyl group whilst a-D-Glc is preferentially attached via either its 3- or 6-hydroxyl group.

� Rhamnose is frequently used as a branch junction.� In bonding with sugars of the main chain b-Rha is very similar to a-D-Gal and prefers

attachment via either the 2- or 3-hydroxyl group.

In viewing the structures represented in Tables 1–3, it is clear that several of the structures

possess backbones that are very closely related. The EPSs isolated from two distantly related

species of Lactobacillus have identical structures: the main chain for the EPS isolated from

species helveticusK16 (group LB-H-3A) is the same as that derived from species L. lactisB891

(group L-3A). The main chain for the EPS isolated from species S. thermophilus Sfi 12 (group

S-5A) is the same as that derived from species L. lactis B39 (group L-5A). A number of EPSs

have main chains that differ only in the substitution of single sugar residues, e.g., the EPSs from

S. thermophilus groups S-5A and S-5B differ by substitution of a glucose to galactose.

Marshall et al. (2001a,b) have recently reported the genetic typing of groups of strains that

produce the same EPS structure. Strains from S. thermophilus group S-3A (IMDO1, IMDO2,

IMDO3, NCFB859, and EU21) and strains from Lactobacilli group LB-D-4A (EU03, EU24,

and EU25) were genetically typed using restriction endonuclease analysis of total DNA and

random amplification of polymorphic DNA. A number of S. thermophilus strains producing

different EPS structures were included in the analysis for comparison (group S-3B SY89 and

SY102). The profiles showed that those LAB secreting the same EPS were also grouped by

the genetic typing.

4. EPS engineering and design

The commercial exploitation of EPSs, as materials for enhancing the texture and mouthfeel

of food, requires the synthesis of EPS having desirable physical properties and for the EPS to

be available in sufficient quantities to match demand. Given that in most cases the desired

contribution of the EPS is to add texture, i.e., to provide thickening properties, it is necessary

to understand how structural components such as linkage, monomer type, substituents, and

molecular mass influence intrinsic viscosity.

In aqueous solutions, most EPSs can be described as random coil polymers. Navarini et al.

(2001) have described experiments investigating the rheological properties of solutions of the

EPS from S. thermophilus SFi 20 and have characterised the EPS as being composed of

random coils as opposed to ordered or rigid polymers. Tuinier et al. (1999a) have reported

results of multiangle light scattering experiments that are consistent with solutions of EPS

derived from L. lactis ssp. cremoris being composed of random coils. As the name suggests,

random coil polymers have no fixed shape and have randomly fluctuating tertiary structure. It

has also been demonstrated that the viscosity of solutions of random coil EPS polymers is

related to the concentration of the EPS and the specific volume of the polysaccharide in

solution (Tuinier et al., 1999b). High specific volumes occur for EPSs having large chain

lengths and for EPSs that are composed of linkages that are ‘stiff’. The general features


relating to the flexibility of different linkages has been described (Rees, 1977; Lapasin and

Pricl, 1995) b(1–4) links impart stiffness compared to b(1–3) and b(1–2), a-linkages are

usually more flexible than b-linkages. It is worth noting at this point that the main chains

derived from strains of S. thermophilus having large repeat units appear to have been

constructed with a view to providing maximum flexibility, a(1–2) and a(1–3) links dominate.

The influence that chain stiffness has on the specific volume can be seen by

comparison of the EPSs from L. lactis NIZO B891 and L. lactis NIZO B40. An EPS

from L. lactis NIZO B891 that has an average molar mass of 2.4� 106 has a smaller

radius of gyration than that measured for an EPS of average molecular mass of 1.4� 106

derived from the L. lactis NIZO B40 (Tuinier, 1999). The structures of the two EPSs have

been described. The backbone of L. lactis NIZO B40 is composed entirely of b(1–4)linked residues and will have a rigid structure and a high specific volume. In contrast, the

back bone of L. lactis NIZO B891 has both an a-linked residue and a (1–6) link that will

impart a degree of flexibility to the backbone and hence the EPS would be expected to

have a smaller radius of gyration. The latter correlation is extremely crude: the structures

of the two EPSs are too varied and the influence that the flexibility that the main chain

might have on the radius of gyration cannot be proven with certainty. More work is

required in this area; knowledge of the structures of a larger number of EPSs will permit

studies of the effect that systematic variation of structural components, such as branches

and substituents, has on the radius of gyration of the EPS.

In considering intrinsic viscosity of solutions of EPSs in isolation, if the intention is to

increase their intrinsic viscosity, then an ideal polysaccharide would be one that has a high

molar mass and a rigid structure. In real systems, it is necessary to consider the influence that

other media components, e.g., proteins and salts, have on the conformation and distribution

of EPS.

4.1. Engineering of EPS structures

One of the main problems encountered in the use of EPSs in texture enhancement is related

to the quantity of material available: production levels are very variable ranging from a few

milligrams to approaching 1 g/l for high producers grown on supplemented media. Given that

the biosynthetic pathways leading to EPS production and for sugar metabolism are known,

given that many of the factors involved in controlling and in the regulation of flux through the

pathway are understood, opportunities arise for engineering of EPSs. Engineering has focused

on increased production of EPS and the production of ‘designer’ polysaccharides.

4.2. Production of designer EPSs

Almost all of the work directed at the engineering of EPS structures has focused on the

genes coding for the glycosyl-transferases. van Kranenburg et al. (1999) have reported the

production of a nonpolar disruption in the gene coding for a priming glycosyl-transferase,

the epsD gene of L. lactis NIZO B40, which results in loss of EPS production. They also

report that a homologous insertion of an epsD gene into the mutated L. lactis NIZO B40


results in recovered EPS production. Heterologous complementation with the gene

encoding for the primary glucosyltransferase involved in the synthesis of a capsular

polysaccharide of S. pneumonie serotype 14 (cps14E gene) restored EPS synthesis (van

Kranenburg et al., 1999). However, homologous or heterologous insertion of genes coding

for priming galactosyl-transferases resulted in failure to produce EPS synthesis. The failure

to observe EPS synthesis when complementation is with a galactosyl-transferase suggests

that the activities of the transferases, involved in synthesis of the oligosaccharide repeat

unit, is restricted to specific lipid-bound acceptor sugars.

In developing the use of heterologous gene insertion to effect EPS biosynthesis, Stingele

et al. (1999) have expressed the entire gene cluster from S. thermophilus SFi 6 into a non-EPS

producer L. lactis MG1363 and observed limited production of EPS. More importantly,

the structure of the EPS was different to that generated by SFi 6. An a 1! 6 linked branching

a-D-Galp was absent and a main chain a-D-galactose replaced the a-D-N-acetylgalactosamine.

It is worth noting at this point that heterologous exchange of EPS gene clusters from one Gram-

negative bacterium to another Gram-negative bacterium has also been performed. Pollock et al.

(1997) have placed those genes responsible for xanthan gum formation in X. campestris into

bacteria from the genus Sphingomonas and observed xanthan gum synthesis.

4.3. Increased production of EPS

Increased production of EPS can be brought about by either genetic manipulation or by

control of microbial physiology; rerouting carbon metabolism towards EPS biosynthesis. de

Vuyst and Degeest (1999) have recently reviewed the influence that microbial physiology has

on EPS production and, as such, this topic will not be covered here. In principle, genetic

manipulation can be used to modify the activities of any of the enzymes involved in the

biosynthetic pathway. In order to optimise EPS production, a detailed knowledge of the flux

of metabolites in each of the enzyme-catalysed transformations is required. A number of

workers are currently investigating if the flux of carbon feed to the sugar nucleotides can be

altered either by deletion of genes coding for key enzymes, e.g., the PGMs or by functional

overexpression of the UDP-glucophosphorylases and/or uridyl-transferases.

In conclusion, LAB bacteria provide scope as cell factories for the synthesis of novel

polysaccharides for use in the food processing industry. Further work in determining the

mechanisms responsible for the control and regulation of EPS biosynthesis, both at the

level of genes and of proteins, is required before EPS production can be optimised. Work

is currently in progress in these areas and it will not be long before new ‘designer’

polysaccharides are being reported. The synthesis of a ‘designer’ polysaccharide has

recently been reported (Colquhoun et al., 2001), they inactivated the aceP gene of

Acetobacter xylinum: the aceP gene codes for a glucosyl-transferase required in the

construction of a branching side chain. The EPS synthesised by the genetically modified

bacteria had a truncated branch. The results complement those of Stingele et al. (1999) and

suggest that the transferase enzymes constructing the main chain must either act before the

branch is added or, alternatively, the transferase enzymes do not recognise the presence of

the branch. They also indicate that the polymerase and export mechanisms are able to work


with oligosaccharides and polysaccharides having modified structures. These results are

extremely encouraging and provide strong impetus for further work to be undertaken in

this area.

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