FEMS Microbiology Reviews 23 (1999) 153^177
Heteropolysaccharides from lactic acid bacteria
Luc De Vuyst *, Bart Degeest
Division of Industrial Microbiology, Fermentation Technology and Downstream Processing (IMDO), Faculty of Sciences,
Vrije Universiteit Brussel (VUB), Pleinlaan 2, B-1050 Brussels, Belgium
Received 10 July 1998; received in revised form 14 September 1998; accepted 19 November 1998
Abstract
Microbial exopolysaccharides are biothickeners that can be added to a wide variety of food products, where they serve as
viscosifying, stabilizing, emulsifying or gelling agents. Numerous exopolysaccharides with different composition, size and
structure are synthesized by lactic acid bacteria. The heteropolysaccharides from both mesophilic and thermophilic lactic acid
bacteria have received renewed interest recently. Structural analysis combined with rheological studies revealed that there is
considerable variation among the different exopolysaccharides; some of them exhibit remarkable thickening and shearthinning properties and display high intrinsic viscosities. Hence, several slime-producing lactic acid bacterium strains and their
biopolymers have interesting functional and technological properties, which may be exploited towards different products, in
particular, natural fermented milks. However, information on the biosynthesis, molecular organization and fermentation
conditions is rather scarce, and the kinetics of exopolysaccharide formation are poorly described. Moreover, the production of
exopolysaccharides is low and often unstable, and their downstream processing is difficult. This review particularly deals with
microbiological, biochemical and technological aspects of heteropolysaccharides from, and their production by, lactic acid
bacteria. The chemical composition and structure, the biosynthesis, genetics and molecular organization, the nutritional and
physiological aspects, the process technology, and both food additive and in situ applications (in particular in yogurt) of
heterotype exopolysaccharides from lactic acid bacteria are described. Where appropriate, suggestions are made for strain
improvement, enhanced productivities and advanced modification and production processes (involving enzyme and/or
fermentation technology) that may contribute to the economic soundness of applications with this promising group of
biomolecules. z 1999 Federation of European Microbiological Societies. Published by Elsevier Science B.V. All rights
reserved.
Keywords : Exopolysaccharide; Lactic acid bacteria; Biothickener
Contents
1. Exopolysaccharides, biopolymers from microbial origin . . . . . . . . . . . . . . . . . . .
1.1. Microbial exopolysaccharides . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
1.2. Exopolysaccharides from food grade microorganisms . . . . . . . . . . . . . . . . . .
2. Exopolysaccharides from lactic acid bacteria: classi¢cation, chemical composition
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* Corresponding author. Tel.: +32 (2) 629-3245; Fax: +32 (2) 629-2720; E-mail: [email protected]
0168-6445 / 99 / $20.00 ß 1999 Federation of European Microbiological Societies. Published by Elsevier Science B.V.
PII: S 0 1 6 8 - 6 4 4 5 ( 9 8 ) 0 0 0 4 2 - 4
FEMSRE 643 16-4-99
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154
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154
L. De Vuyst, B. Degeest / FEMS Microbiology Reviews 23 (1999) 153^177
2.1. Classi¢cation of exopolysaccharides from lactic acid bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
2.2. Chemical composition of exopolysaccharides from lactic acid bacteria . . . . . . . . . . . . . . . . . . . . . .
2.3. Structure of exopolysaccharides from lactic acid bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
3. Biosynthesis and genetics of exopolysaccharide synthesis by lactic acid bacteria . . . . . . . . . . . . . . . . . . .
3.1. Biosynthesis of exopolysaccharides by lactic acid bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
3.2. Genetics of exopolysaccharide biosynthesis by lactic acid bacteria . . . . . . . . . . . . . . . . . . . . . . . . .
3.3. Molecular organization of genes involved in exopolysaccharide biosynthesis by lactic acid bacteria .
4. Microbial physiology and process engineering of exopolysaccharide production by lactic acid bacteria . .
4.1. Methodology used to study exopolysaccharide production by lactic acid bacteria . . . . . . . . . . . . . .
4.2. Exopolysaccharide yields produced by lactic acid bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
4.3. Nutrients enhancing growth and exopolysaccharide production of lactic acid bacteria . . . . . . . . . . .
4.4. In£uence of the carbon and nitrogen source on exopolysaccharide size and composition . . . . . . . . .
4.5. Optimal control of exopolysaccharide production with lactic acid bacteria . . . . . . . . . . . . . . . . . . .
4.6. Production kinetics of exopolysaccharides from lactic acid bacteria . . . . . . . . . . . . . . . . . . . . . . . .
4.7. Large-scale production and downstream processing of exopolysaccharides from lactic acid bacteria .
5. Applications of exopolysaccharides from lactic acid bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
5.1. Exopolysaccharides from lactic acid bacteria in current use . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
5.2. Unwanted exopolysaccharide production by lactic acid bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . .
5.3. Novel applications of exopolysaccharides from lactic acid bacteria . . . . . . . . . . . . . . . . . . . . . . . . .
5.4. Application of exopolysaccharides from lactic acid bacteria as food additives . . . . . . . . . . . . . . . . .
5.4.1. Use as viscosifying agent . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
5.4.2. Use as gelling agent . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
5.5. Application of slime-producing starter cultures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
5.5.1. Rheology versus exopolysaccharide concentration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
5.5.2. Yogurt . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
5.5.3. Viili . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
5.5.4. Ke¢r . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
6. Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Acknowledgments
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References
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1. Exopolysaccharides, biopolymers from microbial
origin
1.1. Microbial exopolysaccharides
Long-chain, high-molecular-mass polymers that
dissolve or disperse in water to give thickening or
gelling properties are indispensable tools in food
product formulation. Such food polymers are also
used for secondary e¡ects, which include emulsi¢cation, stabilization, suspension of particulates, control
of crystallization, inhibition of syneresis (the release
of water from processed foods), encapsulation, and
¢lm formation. Most of the biothickeners in current
use by the food industry are polysaccharides from
plants (e.g. starch, pectin, locust bean gum, guar
gum) or seaweeds (carrageenan, alginate). The animal proteinaceous hydrocolloids gelatin and casein
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155
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are used too. The functional properties in foods of
these polymers are determined by quite subtle structural characteristics. However, these polysaccharides
may not always be readily available in the quality
needed or their rheological properties may not exactly match those required. Most of the plant carbohydrates used are chemically modi¢ed to improve
their structure and rheological properties [1,2]. Their
use is hence strongly restricted. For instance, in the
European Union (EU), their addition is allowed only
in some food products. Those food products need to
be labeled with an E-number.
An alternative class of biothickeners is that of microbial exopolysaccharides (EPS). Microbial exopolysaccharides are extracellular polysaccharides which
are either associated with the cell surface in the form
of capsules or secreted into the extracellular environment in the form of slime. They are referred to as
FEMSRE 643 16-4-99
L. De Vuyst, B. Degeest / FEMS Microbiology Reviews 23 (1999) 153^177
capsular or slime exopolysaccharides, respectively
[3]. EPS occur widely among bacteria and microalgae and less among yeasts and fungi [3^7].
EPS in their natural environment are thought to
play a role in the protection of the microbial cell
against desiccation, phagocytosis and phage attack,
antibiotics or toxic compounds (e.g. toxic metal ions,
sulfur dioxide, ethanol), predation by protozoans,
osmotic stress, adhesion to solid surfaces and bio¢lm
formation, and also in cellular recognition (e.g. via
binding to a lectin). It is not likely that EPS serve as
a food reserve, since most slime-forming bacteria are
not capable of catabolizing the EPS they produce [8].
In pathogenic bacteria, such as Streptococcus pneumoniae and Streptococcus agalactiae, capsular EPS
and O-antigen lipopolysaccharides are involved in
the immune response.
Examples of industrially important microbial exopolysaccharides are dextrans, xanthan, gellan, pullulan, yeast glucans and bacterial alginates [1,9,10].
Novel microbial biopolymers may ¢ll a gap in the
market-available polymers or may replace a traditional product in terms of the improved rheological
and stability characteristics. Now, microbial polysaccharides represent only a small fraction of the current biopolymer market. Factors limiting the use of
microbial EPS are their production, which requires a
thorough knowledge of their biosynthesis and an
adapted bioprocess technology, and the high cost
of their recovery. Xanthan is a microbial EPS approved in the food industry, mainly because of its
unique rheological properties in foods and the possibility of low-cost production. It is produced in high
amounts by Xanthomonas campestris, a phytopathogenic bacterium that is not generally recognized as
safe (GRAS). Recently, gellan from the phytopathogen Sphingomonas elodea has been introduced on
the market too. Strains of GRAS, food grade microorganisms, in particular lactic acid bacteria, that are
able to produce EPS in large enough quantities could
be the solution to many of the above-mentioned disadvantages.
This review particularly deals with microbiological, biochemical and technological aspects of heteropolysaccharides from, and their production by, lactic
acid bacteria.
155
1.2. Exopolysaccharides from food grade
microorganisms
Many food grade microorganisms produce EPS, in
particular lactic acid bacteria (LAB) [8,11], propionibacteria [11] and bi¢dobacteria [12,13]. Most of the
EPS-producing LAB strains studied in more detail
were isolated from dairy products, e.g. Scandinavian
ropy fermented milk products [14^16], various yogurts [17^21] and fermented milks [22], and milky
[23,24] and sugary [25] ke¢r grains. Also cheese [26]
and fermented meat and vegetables [27] served as a
source of EPS-producing LAB strains.
2. Exopolysaccharides from lactic acid bacteria:
classi¢cation, chemical composition and structure
2.1. Classi¢cation of exopolysaccharides from lactic
acid bacteria
EPS from LAB can be subdivided into two
groups: (1) homopolysaccharides, consisting of four
subgroups, namely (a) K-D-glucans, i.e. dextrans
(Leuconostoc mesenteroides subsp. mesenteroides
and Leuc. mesenteroides subsp. dextranicum), mainly
composed of K-1,6-linked glucose residues with variable (strain speci¢c) degrees of branching at position
3, and less frequently at positions 2 and 4, and alternan (Leuc. mesenteroides) and mutans (Streptococcus
mutans and Streptococcus sobrinus), both composed
of K-1,3- and K-1,6-linkages; (b) L-D-glucans composed of L-1,3-linked glucose molecules with L-1,2branches, produced by Pediococcus spp. and Streptococcus spp.; (c) fructans, mainly composed of L2,6-linked D-fructose molecules, such as levan with
some L-2,1-branching through the Ol site (S. salivarius); (d) others, like polygalactan, composed of
structurally identical repeating units with di¡erent
glycosidic linkages; and (2) heteropolysaccharides
produced by mesophilic (Lactococcus lactis subsp.
lactis, L. lactis subsp. cremoris, Lactobacillus casei,
Lb. sake, Lb. rhamnosus, etc.) and thermophilic (Lb.
acidophilus, Lb. delbrueckii subsp. bulgaricus, Lb. helveticus and S. thermophilus) LAB strains. The latter
group of EPS receives renewed interest, since they
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L. De Vuyst, B. Degeest / FEMS Microbiology Reviews 23 (1999) 153^177
play an important role in the rheology, texture and
body, and mouthfeel of fermented milk drinks. For
instance, the creamy, smooth texture is one of the
aspects of the quality of yogurt which seems to be
improved by the ability of the yogurt bacteria to
produce EPS, even though only small amounts of
EPS are being produced. In addition, EPS from
LAB have one of the largest technical potentials
for development of novel and improved products
such as low-milk-solid yogurts, low-fat yogurts,
creamier yogurts, etc. [28]. Moreover, some polysaccharides may contribute to human health, either as
non-digestible food fraction [29] or because of their
antitumoral [30,31], antiulcer [32], immunomodulating [33,34] or cholesterol-lowering activity [35].
Therefore EPS from LAB have potential for development and exploitation as functional food ingredients with both health and economic bene¢ts.
2.2. Chemical composition of exopolysaccharides
from lactic acid bacteria
The chemical composition of EPS from LAB has
long been controversial. The nature of slime materials from LAB was ¢rst studied by Sundman [36] and
Nilsson and Nilsson [37], and it was found to be
protein-like material. Later, some authors suggested
that the ropy characteristic which developed in milk
during fermentation was due to the production of
a glycoprotein or carbohydrate^protein complex
[15,38,39]. Others isolated exopolymer material enriched in its carbohydrate content upon further puri¢cation [16,18,19,26,30,40^45]. Finally, there is
now agreement that the exopolymers from LAB
are polysaccharides composed of (branched) repeating units, containing K- and L-linkages, and that
many di¡erent types are secreted [11]. However, their
monomer composition seems to be remarkably similar. D-galactose, D-glucose and L-rhamnose are almost always present, but in di¡erent ratios [16,20^
22,46^65]. Whereas the EPS from Lb. acidophilus
LMG 9433 [56], Lb. helveticus TYl-2 [50], Lb. helveticus NCDO 766 [54], Lb. rhamnosus C83 [22], S.
thermophilus S¢20 [46,61], S. thermophilus S¢32 [63]
and S. thermophilus LY03, S. thermophilus BTC and
S. thermophilus 480 [21,64] lack rhamnose, Lb. paracasei 34-1 EPS only contains galactose [55], S. thermophilus OR 901 EPS only contains galactose and
rhamnose [20,62] and Lb. sake 0-1 EPS only consists
of glucose and rhamnose [53]. In contrast, the EPS
produced by Lb. delbrueckii subsp. bulgaricus CRL
420 contains glucose and fructose in a ratio of 1:2
[17] and the polymer produced by S. thermophilus
MR-1C consists of an octameric basic repeating
unit composed of D-galactose, L-rhamnose and L-fucose in a 5:2:1 ratio [66]. Other residues, such as snglycerol-3-phosphate, N-acetyl-aminosugars, and
phosphate and acetyl groups can also be present
[16,46,47,50^53,55,56,61]. Besides poor isolation
and puri¢cation, especially when complex media
are used, the possibility that more than one polysaccharide can be secreted by one strain may explain
some of the di¡erent compositions observed in the
past. For instance, Grobben et al. [57^59] reported
di¡erent compositions of exopolysaccharide material
from the same strain, depending on the fermentation
conditions used and the isolation and puri¢cation
techniques applied. Marshall et al. [52] recently isolated two di¡erent EPS from the fermentation medium of the same strain that have di¡erent monosaccharide composition and apparent molecular
mass. High-molecular-mass and low-molecular-mass
EPS fractions were also isolated from the same Lb.
delbrueckii subsp. bulgaricus [59] and S. thermophilus
(Degeest et al., unpublished results) strain, but which
do not di¡er in monomeric composition. The media
and culture conditions used may be one of the factors in£uencing monomeric composition and variations in glycosidic bonds (cf. infra).
2.3. Structure of exopolysaccharides from lactic acid
bacteria
EPS from LAB possess apparent molecular masses
that range from 4.0U104 to 6.0U106 . The molecular
mass is one of the factors determining the functional
properties of EPS [43,67] (Degeest et al., unpublished
results). However, the physical and rheological properties of a polysaccharide in solution are closely related to its three-dimensional structure or conformation. Also, other factors than the molecular shape,
such as the potential of the molecule to form intermolecular associations, can be important for a complete understanding of the solution behavior, since
polysaccharide chains have to undergo some topological rearrangements from disordered random coils
FEMSRE 643 16-4-99
Fig. 1. Primary structure of EPS produced by LAB: (1) homopolysaccharides : A, dextran; B, levan ; C, polygalactan, L. lactis subsp. cremoris H414 [48]; (2) heteropolysaccharides from mesophilic LAB: D, L. lactis subsp. cremoris SBT 0495 [16,83] ; E, Lb. sake 0-1 [53]; F, Lb. paracasei 34-1 [55]; and (3) heteropolysaccharides from thermophilic LAB: G, Lb. acidophilus LMG 9433 [56]; H, Lb. delbrueckii subsp. bulgaricus rr [49,57]; I, Lb. helveticus NCDO 766 [54]; J, Lb. helveticus TY1-2 [50] and K, its mutant TN-4 [51]; L, S. thermophilus S¢6 [46,61]; M, S. thermophilus OR 901 [62]; N, S. thermophilus S¢32 [63]; and O, S. thermophilus S¢l2 [63]. Glc, glucose ; Gal,
galactose ; Rha, rhamnose ; GlcNAc, N-acetyl-glucosamine ; GalNAc, N-acetyl-galactosamine ; Ac, acetyl. The D- (D) and L- (L) con¢guration, and pyranose (p) and furanose
(f) structure are indicated.
L. De Vuyst, B. Degeest / FEMS Microbiology Reviews 23 (1999) 153^177
FEMSRE 643 16-4-99
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L. De Vuyst, B. Degeest / FEMS Microbiology Reviews 23 (1999) 153^177
to more ordered conformations favorable to intermolecular interactions and associations.
The secondary and tertiary conformation of a
polysaccharide is strongly dependent on its chemical
(primary) structure. Relatively small changes in primary structure may have a tremendous e¡ect on the
conformation and properties of a polysaccharide [7].
The structure of the repeating unit of a LAB heteropolysaccharide (produced by S. thermophilus) was
¢rst determined by Doco et al. [46]. Other repeating
unit structures of branched EPS produced by LAB
have been elucidated recently through acid hydrolysis, methylation analysis, periodate oxidation, acetolysis, enzymatic digestion, Smith degradation and 1D
and 2D 1 H-NMR spectroscopy, etc. [46^51,53^
56,61^63] (Fig. 1). Their size may range from a disaccharide to a heptasaccharide. They further show
few common features which raises the question
about the relationship between these EPS structures
and the texturizing properties they confer. It would
be of interest to understand this relationship and to
have the means to modify the biopolymers to in£uence the properties of the native polysaccharides.
Enzymes having a speci¢c action on the EPS structure can be used for tailoring the chemical structure
and hence the functional properties. Finally, these
data in combination with additional biochemical
and molecular biological studies of EPS (cf. infra)
will form the basis for future `polysaccharide engineering' and perhaps `oligosaccharide engineering'.
The latter could be of great value in view of the
explosive functional foods market.
3. Biosynthesis and genetics of exopolysaccharide
synthesis by lactic acid bacteria
3.1. Biosynthesis of exopolysaccharides by lactic acid
bacteria
For a small number of homopolysaccharides, including dextrans, mutans, alternan and levans, the
biosynthesis process is extracellular and requires
the speci¢c substrate sucrose. A highly speci¢c glycosyl transferase enzyme (e.g. dextran or levan sucrase for dextran and levan biosynthesis, respectively) is involved in the polymerization reaction;
the energy needed for polymerization comes from
the hydrolysis of sucrose (Fig. 2). The polysaccharide
can be produced either using whole bacterial cell
cultures or cell-free (immobilized) enzyme preparations [8].
Heteropolysaccharides are made by polymerizing
repeating unit precursors formed in the cytoplasm
[8,11]. Several enzymes and/or proteins are involved
in the biosynthesis and secretion of heterotype EPS
which are not necessarily unique to EPS formation
(Fig. 3). The sugar nucleotides, derived from sugar1-phosphates, play an essential role in heteropolysaccharide biosynthesis due to their role in sugar activation, which is necessary for monosaccharide polymerization, as well as sugar interconversions
(epimerization, decarboxylation, dehydrogenation,
etc.). Together with the sugar activation and modi¢cation enzymes, they play a crucial role in the formation of the building blocks and thus the ¢nal EPS
composition. For instance, in glucose-grown Lb. delbrueckii subsp. bulgaricus NCFB 2772 cultures, the
activity of UDP-glucose pyrophosphorylase, leading
to the biosynthesis of UDP-glucose and UDP-galactose, was higher than that in fructose-grown cultures,
whereas in fructose-grown cultures, no enzyme activities were found that led to the biosynthesis of
dTDP-rhamnose [58]. UDP-glucose pyrophosphorylase was also associated with EPS production in a
ropy S. thermophilus strain and not in the non-ropy
strain [68]. Since fructose-grown cultures of Lb. delbrueckii subsp. bulgaricus NCFB 2772 produced EPS
with a lower level of galactose, while the activity of
UDP-galactose-4-epimerase was only slightly lower
in these cells, it appears that this enzyme does not
play an important role in the sugar composition of
the EPS produced [58]. However, a relationship between the activity of the enzyme UDP-galactose 4epimerase and EPS production was found in L. lactis
[69]. It was not associated with EPS biosynthesis
either in any S. thermophilus strain examined by Escalante et al. [68]. Consequently, glucose or the glucose moiety from lactose hydrolysis seems to be the
source of sugar for heteropolysaccharide biosynthesis in lactic acid bacteria. A high UDP-glucose pyrophosphorylase activity was found in these strains.
However, since S. thermophilus has a Gal3 phenotype (su¡ering from a defect in the induction mechanism for the rate-limiting key enzyme galactokinase), it has been suggested that the most likely
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159
Fig. 2. Biosynthesis of the homopolysaccharide dextran. The enzyme dextran sucrase is involved in the polymerization reaction. The energy needed for polymerization comes from the hydrolysis of sucrose.
function of both Leloir enzymes uridine diphosphogalactose 4-epimerase and galactose 1-phosphate uridyltransferase is the biosynthesis of precursors for
extracellular polysaccharides [70]. Detailed biochem-
ical and enzymatic studies may contribute to unravel
this complex system. Anyway, glucose-1-phosphate ^
derived from glucose-6-phosphate which is in turn an
important metabolic intermediate from sugar break-
Fig. 3. Schematic representation of pathways involved in lactose catabolism (left and upper right) and exopolysaccharide biosynthesis
(lower right) in lactose-fermenting Lactococcus lactis (lactose transport via a lactose-speci¢c phosphotransferase primary transport system)
and galactose-negative Lactobacillus delbrueckii subsp. bulgaricus and Streptococcus thermophilus (lactose transport via a lactose/galactose
antiport secondary transport system) strains. 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.
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Fig. 4. Model for EPS biosynthesis in Lactococcus lactis subsp. lactis NIZO B40 [85]. C55-P, lipid carrier ; Glc, glucose ; Gal, galactose ;
Rha, rhamnose; UDP-Glc, UDP-Gal and TDP-Rha are nucleotide sugars. For the abbreviations of the responsible enzymes: see Fig. 5A.
down ^ is most probably a precursor for polysaccharide formation [71]. Hence, phosphoglucomutase
could be a key enzyme linking the lactose degradation pathway (energy generation) to EPS biosynthesis (sugar nucleotide biosynthesis) (Fig. 3). Assuming
that linkage of sugar catabolism and sugar anabolism occurs at this branching point, it is tempting to
speculate that it is possible to engineer overproduction of EPS since the galactose moiety could be catabolized completely via the glycolysis, while the glucose moiety could be used for EPS production.
Therefore, the £ux via the phosphoglucomutase
should be made su¤ciently high. The question is
whether this can be realized [72]. One of the problems may be the inability of S. thermophilus to catabolize galactose. When lactose is used as the carbon source, S. thermophilus and Lb. delbrueckii
subsp. bulgaricus release the galactose moiety to the
medium via the lactose/galactose antiport transport
process, leaving only the glucose moiety as the energy and carbon source for the bacterial cell.
The EPS monomeric composition may not only be
dependent on the sugar nucleotide level inside the
cell, but probably also on the assembly of the EPS
repeating unit. Polymerization of some hundreds to
several thousands of the repeating units takes place
through sequential addition of sugar residues by speci¢c glycosyl transferases from nucleotide sugars to a
growing repeating unit that is coupled to the undecaprenyl phosphate carrier yielding the ¢nal EPS
(Fig. 4). This isoprenoid glycosyl lipid carrier located
in the cell membrane would act as the recipient molecule for the ¢rst sugar residue. In contrast with
Gram-negative bacteria [3^7], there is only preliminary evidence for the existence of this lipid carrier in
LAB [73]. Furthermore, as indicated by the structural diversity of EPS produced by LAB, these microorganisms must contain a vast pool of speci¢c glycosyl transferases, all involved in the assembly of the
repeating units, which have not been exploited so
far. The functional expression of desired combinations of glycosyl transferase genes from di¡erent origin opens the way to polysaccharide engineering.
As a last step of EPS biosynthesis, the synthesized
polysaccharide is translocated across the membrane
to the exterior of the cell and is excreted in the environment (i.e. slime EPS) or remains attached to the
cell (i.e. capsular EPS). Both polymerization and
transport may a¡ect the amount or the sugar composition of the EPS.
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The biosynthesis of polysaccharides is an energydemanding process. First, one ATP is required for
the conversion of each hexose substrate molecule to
a hexose phosphate. A further high-energy phosphate bond is needed for the synthesis of each sugar
nucleotide, and one ATP is required for the phosphorylation of the isoprenoid C55 lipid carrier. Finally, polymerization and transport presumably need
energy. Because energy generation in lactic acid bacteria is very limited in contrast with aerobic bacteria,
such as X. campestris, this will strongly limit the
amount of exopolysaccharide material produced.
Furthermore, since the isoprenoid glycosyl lipid carrier is also involved in the biosynthesis of cell wall
polymers (peptidoglycans, teichoic acids, lipopolysaccharides), there is competition for this facilitating
membrane component during di¡erent phases of
growth [3].
Thus, the nature and composition of EPS as capsular or slime material are in£uenced by both medium composition, biosynthetic pathways, growth
phase, and rate of microbial growth. These are important factors in the understanding of polysaccharide biosynthesis and secretion and need further
study.
3.2. Genetics of exopolysaccharide biosynthesis by
lactic acid bacteria
The problems associated with most ropy strains
used for fermentation are that their ability to produce EPS is often an unstable characteristic, at the
genetic level as well as the instability of the ropy
texture itself. As an example, a ropy strain of S.
thermophilus produced under the same experimental
conditions from one day to another viscosities varying from 41 to 240 mPa s and amounts of EPS between 45 and 340 mg l31 [19]. Loss of the slimeproducing trait may occur on repeated subculture
or after prolonged incubation especially at high temperatures [15,74]. Also, spontaneous mutation of the
producing cells may occur that results in colony variants, weaker production or even an altered composition of the EPS produced [45,51,75]. Consequently,
not all ropy strains are suitable for large-scale industrial fermentations, and ropy strains in use have to
be periodically reselected from the culture to conserve the EPS production characteristics in industrial
161
strains. Thus, at present yogurt manufacturers still
rely on prefermentation processing, such as increasing milk solids through the addition of milk powder,
whey powder, caseinate, etc. or the concentration of
milk (by evaporation, membrane ¢ltration, etc.), heat
treatment of the milk prior to inoculation, homogenization, incubation conditions and handling of the
ripened coagulum and/or addition of stabilizers
(modi¢ed starch, carrageenan, guar gum, pectin, gelatin, sodium caseinate) for product stability. However, these additives adversely a¡ect the true taste
and aroma of yogurt. Consequently, in situ production of EPS and exploitation of the EPS-producing
characteristic of thermophilic LAB has clear economic bene¢ts, especially in stirred yogurts, enhancing smoothness and avoiding syneresis.
The loss of the slime-producing trait from mesophilic LAB strains has been attributed to loss of
plasmids. Here, the ropy character is associated
with plasmid DNA [26,76^81]. For thermophilic
LAB, such as Lb. delbrueckii subsp. bulgaricus and
S. thermophilus, this may not be the explanation, as
these ropy strains have not, to date, been shown to
harbor plasmids encoding EPS production [8,79],
and, when the phenotype is lost, it can frequently
be recovered, for instance, by repeated passage
through milk [8,19,43,75,79]. The genes required for
EPS biosynthesis seem, therefore, to be chromosomally located in thermophilic LAB. The genetic instability could be due to mobile genetic elements or
to a generalized genomic instability, including DNA
deletions and rearrangements. Both phenomena were
observed for Lb. delbrueckii subsp. bulgaricus and S.
thermophilus [61,75].
3.3. Molecular organization of genes involved in
exopolysaccharide biosynthesis by lactic
acid bacteria
Sequence data of speci¢c genes involved in EPS
production are known for S. thermophilus S¢6
[61,82], S. thermophilus NCFB 2393 [83], S. thermophilus MR-1C [66], Lb. delbrueckii subsp. bulgaricus
[65] and L. lactis subsp. cremoris NIZO B40 [81]. A
complex genetic organization is responsible for EPS
production and secretion (Fig. 5). For instance, the
chromosomally located eps gene cluster of S. thermophilus S¢6, identi¢ed by Tn916 mutagenesis in com-
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Fig. 5. Organisation of the eps gene cluster involved in the EPS biosynthesis of : (A) L. lactis subsp. cremoris NIZO B40 (plasmid-localized) [81]; and (B) S. thermophilus S¢6 (chromosomally encoded) [61]. The (possible) functions of the di¡erent gene products are indicated.
bination with a plate assay based on contrast staining, contains at least 14.52 kb of DNA encoding the
13 genes epsA to epsM [61]. Expression of the genes
epsE, epsF, epsG and epsI in Escherichia coli and in
vitro glycosyl transferase reactions with 14 C-labeled
sugar nucleotides followed by analysis of the labeled
sugars by thin-layer chromatography, identi¢ed
EpsE, EpsG, EpsI as being the ¢rst L-galactose, an
K-1,3-N-acetylgalactosamine and a L-1,3-glucose
transferase, respectively. By exclusion, the K-1,6-galactose transferase should be EpsF since it is the glycosyl transferase gene left without a functional assignment [84]. In L. lactis subsp. cremoris NIZO
B40 the 14 eps genes (12-kb region) are preceded
by an iso-IS 982 insertion sequence element and located on the 40-kb plasmid, pNZ4000 [81]. Insertional inactivation by homologous recombination and
antisense approaches were used to inactivate these
plasmid-located eps genes. Heterologous expression
in E. coli of the epsDEFG genes showed that they
are all involved in the assembly of the complete
backbone of the EPS repeating unit, linking glucose
to a lipid carrier (EpsD), glucose to lipid-linked glucose (EpsE and EpsF) and galactose to lipid-linked
cellobiose (EpsG), respectively. Moreover, homologous expression indicated that the epsDEFG encoded glycosyl transferases have the same activity
in L. lactis as in E. coli resulting in a lipid-linked
trisaccharide intermediate [85]. In addition, Southern
blot analysis revealed that the gene encoding the
glucosyl transferase linking the ¢rst sugar of the repeating unit to the membrane-bound lipid carrier is
found only in EPS-producing strains of S. thermophilus [83]; the activity of its gene product would be
a rate-limiting step in the EPS biosynthesis pathway
in lactic acid bacteria [85].
The organization, transcriptional direction, and
deduced functions of the genes in the di¡erent eps
gene clusters appear to be highly conserved. According to homology searches, the genes seem to be organized in four functional regions [61,81,84]: a central region with genes showing homology with
glycosyl transferases speci¢cally required for the biosynthesis of the EPS-repeating unit; two regions
£anking the central region that show homology to
enzymes involved in polymerization and export
(chain-length control, export, polymerization); and
a regulatory region located at the beginning of the
gene cluster. Northern hybridizations have shown
that the streptococcal as well as the lactococcal eps
gene clusters are transcribed as one unit and, in both
cases, an E. coli consensus promoter was found upstream of the ¢rst gene.
More knowledge of the molecular organization
and of the factors regulating expression of EPS will
make it possible to enhance EPS production under
de¢ned growth or fermentation conditions, to increase the number of possibilities for modifying the
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structure and function of EPS, and to realize simultaneous production of di¡erent EPS. Changing production by cloning the eps genes in suitable heterologous hosts was realized with the S. thermophilus eps
gene cluster that could be expressed in a non-producing L. lactis strain [61]. However, only low
amounts of EPS were produced in this heterologous
host and the recombinant EPS had a similar high
molecular mass, but a di¡erent structure (no branching, no aminosugars) than the EPS from the parent
strain [84]. According to the latter ¢ndings, this
modi¢cation implies that: (1) the glycosyl transferases are not exclusively speci¢c for the donor and the
acceptor sugar molecule; and (2) the polymerase
from S. thermophilus S¢6 can recognize and polymerize a repeating unit which di¡ers in the backbone
as well as the sidechains from its normal substrate.
Even though an increased understanding of the
mechanism underlying these catalytic processes, including the sugar nucleotide pathways, is necessary,
these results o¡er good perspectives for future polysaccharide engineering. Another approach is to engineer the EPS production in the production host itself
or in related strains. This was realized with the plasmid-encoded lactococcal eps genes that could be conjugationally transferred to the genetically well-characterized and well-transformable L. lactis strain MG
1614 [81]. Also Vedamuthu and Neville [76] and von
Wright and Tynkkynen [77] were able to transfer the
mucoidness plasmid from L. lactis subsp. cremoris
ARH87 and MS, respectively, to non-mucoid L. lactis strains, thereby making them mucoid. In these
and other production hosts, inactivation by site-directed or random mutagenesis of eps genes or relevant household genes coding for speci¢c sugar activations, may a¡ect the incorporation of activated
nucleotide sugars or other steps in the polymerization process. For instance, UDP-galactose is one of
the precursors of the NIZO B40 EPS biosynthesis.
The role of the Leloir enzyme galactose epimerase
(GalE) that converts UDP-galactose into UDP-glucose and vice versa, was studied using GalE mutant
strains [86]. It could be shown that GalE activity is
essential for EPS production when cells are grown in
media with glucose as sole carbon source. A second
precursor in EPS biosynthesis is TDP-rhamnose. The
L. lactis MG1363 rfb gene cluster encoding enzymes
involved in TDP-rhamnose biosynthesis from glu-
163
cose-1-phosphate (rfbAC) have been cloned and
characterized. Disruption of this rfb gene cluster
could lead to a loss of EPS production or a changed
EPS composition [86]. Alternatively, existing or new
eps genes may be overexpressed. This could result in
higher EPS yields and mutant EPS that have altered
structures and properties. Such EPS engineering offers not only the possibility to study structure^function relationships of EPS, but also to create EPS
with novel properties that can be used as industrial
biothickeners. However, genetically modi¢ed microorganisms and their products will require legal approval and need to be accepted by both the food
processor and the consumer.
4. Microbial physiology and process engineering of
exopolysaccharide production by lactic acid
bacteria
4.1. Methodology used to study exopolysaccharide
production by lactic acid bacteria
Biosynthesis and secretion of EPS from LAB occur during di¡erent growth phases and both the
amount and type of polymer is in£uenced by growth
conditions. However, contradictory results have been
reported regarding the in£uence of physical and
chemical factors on EPS production by LAB. Since
no EPS production was initially observed in MRS or
synthetic media, milk was the medium studied
[18,19,21,38,43,45,60,64,87^90]. Also whey and
whey-based media have been studied [91]. Only recently have semi-synthetic and (complex) synthetic
media been investigated [21,38,44,67,89,92^97]. A
chemically de¢ned medium [22,26,52,57,98] containing a carbohydrate source, mineral salts, amino
acids, vitamins, and nucleic acid bases is more suitable to investigate the in£uence of nutrients on the
growth, the metabolic pathways, and the biosynthesis of EPS in LAB, i.e. the quantitative and qualitative production of EPS and the investigation of the
composition of the EPS produced. Furthermore,
most investigators focused their attention on viscosity measurements, since those were used traditionally
as an indication for EPS production in liquid media
[18,19]. There are fewer reports on the quanti¢cation
of EPS produced by LAB [8,11].
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4.2. Exopolysaccharide yields produced by lactic acid
bacteria
The total yield of EPS produced by LAB depends
on the composition of the medium (carbon and nitrogen sources) and the conditions in which the
strains grow, i.e. temperature, pH and incubation
time (cf. infra). The production of intracellularly
synthesized EPS by di¡erent LAB strains varies
roughly from 0.045 to 0.350 g l31 when the bacteria
are grown under non-optimized culture conditions.
Optimal culture conditions result in EPS yields
from 0.150 to 0.600 g l31 , depending on the strain
[8,11]. When a ropy strain of S. thermophilus is
grown in association with a non-ropy strain of Lb.
delbrueckii subsp. bulgaricus in milk, EPS production
can reach quantities of almost 0.800 g l31 [19,88]. An
optimal carbon/nitrogen ratio in both milk and MRS
media resulted in 1.1 g l31 with S. thermophilus
LY03 [21,64,95]. With Lb. sake O-1, EPS yields of
approximately 1.4 g l31 are achieved [67] (Janssens
and De Vuyst, unpublished results). These values
are, however, not comparable with the high yields
obtained with dextran-producing LAB and Gramnegative EPS producers such as X. campestris.
4.3. Nutrients enhancing growth and
exopolysaccharide production of lactic acid
bacteria
Enhanced EPS production and growth were initially obtained when (hydrolyzed) casein was added
to skim milk cultures of Lb. delbrueckii subsp. bulgaricus [38,88]. According to early investigations,
neither growth nor EPS production was speci¢cally
linked to the presence of casein or whey proteins in
the growth medium of LAB. However, Cerning et al.
[88] found that casein stimulates EPS production,
but not growth of Lb. delbrueckii subsp. bulgaricus.
On the other hand, addition of hydrolyzed casein to
MRS did not increase speci¢c polymer production
by Lb. delbrueckii subsp. bulgaricus [38]. It has also
been reported that Lb. delbrueckii subsp. bulgaricus
is able to produce the same amount of EPS in milk
and milk ultra¢ltrate, but that S. thermophilus is not
[88]. On the other hand, supplementation of milk
and milk ultra¢ltrate with glucose or sucrose stimulates EPS production by Lb. casei and even modi¢es
the monosaccharide composition of the EPS with
glucose becoming dominant. In the latter case, rhamnose as EPS constituting residue is no longer present
as compared to growth and EPS production in a
synthetic medium [26,43,44]. Lb. delbrueckii subsp.
bulgaricus NCFB 2772 produced considerably larger
amounts of EPS when grown on glucose or lactose
than when grown on fructose to equal cell densities,
and the EPS had a di¡erent monomeric sugar composition (cf. infra). Compared with growth on the
other carbohydrate sources, growth on mannose
led to much lower levels of growth and EPS production [57,58]. Mannose or a combination of glucose
and fructose were the most e¤cient carbon sources
for EPS production by Lb. rhamnosus C83 [22]. Not
only the nature of the carbon source and sometimes
the combination of monosaccharides, but also their
concentration can have a stimulating e¡ect on EPS
biosynthesis [22,43,44]. As an example, when Lb.
rhamnosus C83 was grown in a chemically de¢ned
medium on 4% mannose or 2% glucose and fructose
(ratio 1:1), it was found that EPS production increased by three or four times, whereas the ¢nal
biomass concentrations were identical [22].
For some EPS-producing bacteria, such as Xanthomonas, Pseudomonas and Rhizobium spp., nitrogen limitation results in increased EPS production
[99]. This seems not to be the case for LAB
[21,64,95^97]. Moreover, it has been shown that an
optimal balance between the carbon and nitrogen
source is absolutely necessary to achieve high EPS
yields [21,64,95].
The possibility exists that also other medium components such as minerals, some amino acids, and/or
some bases and vitamins, a¡ect the composition of
the EPS produced. For instance, mineral requirements would a¡ect EPS production [22,93]. Also,
vitamins seem to play an important role [98]. In contrast to the omission of single or multiple amino
acids, it was demonstrated that the omission of multiple vitamins a¡ected the production of EPS relative
to cell growth. However, it only a¡ected the total
amount of EPS produced and not the ratio of the
high-molecular-mass fraction and the low-molecularmass fraction of the EPS from Lb. delbrueckii subsp.
bulgaricus NCFB 2772 as was found when fructose,
instead of glucose, was used as the carbohydrate
source (cf. infra). The requirement observed for vi-
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tamins could, however, be dependent on the presence
of lipids in the medium, e.g. oleate [100].
4.4. In£uence of the carbon and nitrogen source on
exopolysaccharide size and composition
The in£uence of the carbon source on the nature
and distribution of sugars in the EPS produced has
long been debated, particularly because minor di¡erences in the composition of the polymer were often
found due to analytical artefacts (cf. infra). However, Petit et al. [101] showed that polysaccharide
production increased and that carbohydrate and uronic acid distribution in polysaccharides altered in
favor of galactose with decreasing lactose feed rate
in fed-batch cultivation with a galactose-fermenting
S. thermophilus strain. Unfortunately, no uronic
acids have been found in the EPS structures elucidated so far. In addition, the assays mainly used for
quantitative determination of uronic acids are not
su¤ciently accurate, insofar as hexoses and pentoses
interfere with their speci¢city [102]. Also the way of
routine quanti¢cation during fermentation of the
EPS produced from LAB may be questionable. It
is mainly based on indirect methods, either colorimetric techniques resulting in exopolysaccharide
yields expressed as glucose equivalents [44,101] or
dextran equivalents [38,92], results that are only valid if extensive dialysis of the samples has been performed, or viscosity measurements which are not
necessarily related to exopolysaccharide yields
[18,19,97]. Measuring the polymer dry mass, obtained after EPS precipitation and extensive washing
and deproteinization of the samples followed by drying is a promising alterative [21,64].
Marshall et al. [52] showed that L. lactis subsp.
cremoris LC 330 produces two exopolysaccharides
simultaneously: a neutral EPS with a molecular
mass higher than 1.0U106 and a charged phosphopolysaccharide with a molecular mass of approximately 1.0U104 . In contrast to the low-molecularmass EPS, the production of the high-molecularmass EPS was positively in£uenced by nitrogen limitation. Degeest et al. (unpublished results) observed
a `shift' from high-molecular-mass to low-molecularmass EPS production by S. thermophilus LY03 with
increasing initial concentrations of the complex nitrogen source. Finally, Grobben et al. [58] initially
165
showed that in continuous culture, when the Lb. delbrueckii subsp. bulgaricus NCFB 2772 strain was
grown on lactose, the amount and sugar composition
of the EPS produced was comparable with values for
glucose-grown cultures, i.e. galactose, glucose and
rhamnose in a ratio of 6.8:1.0:0.7; when grown
with fructose as the carbohydrate source, the amount
of EPS produced was substantially lower and the
EPS were di¡erent in composition, being composed
of galactose and glucose in the ratio 2.5:1.0. No
rhamnose residues were detected in those EPS.
They concluded that Lb. delbrueckii subsp. bulgaricus
NCFB 2772 produces two exopolysaccharide fractions with relative molecular masses of 1.7U106
and 4U104 . Later they reported on a di¡erent monomer composition, and found that the production of
the high-molecular-mass fractions (galactose, glucose
and rhamnose in the molar ratio of 5.0:1.0:1.0) appeared to be dependent on the carbohydrate source
(glucose versus fructose), whereas the low-molecularmass fractions (galactose, glucose, rhamnose in the
molar ratio of approximately 11.0:1.0:0.4) were produced more continuously [59]. Some authors did or
do not ¢nd a di¡erent sugar composition of the EPS
produced when growth on di¡erent energy sources is
compared [17,67]. Hence, additional information on
the e¡ects of varying growth conditions on the size
and composition of the EPS produced and, consequently, the functional and rheological properties of
EPS from LAB would be of signi¢cant bene¢t to the
food industry.
4.5. Optimal control of exopolysaccharide production
with lactic acid bacteria
Optimal conditions of temperature, pH and incubation time result in improved EPS yields
[21,22,60,64,67,75,89^95,98]. Several reports show
that low temperatures markedly induce slime production [22,26,43,52,60,67,74,90^92,103]. This e¡ect
has been explained, based on information for EPS
production from Gram-negative bacteria, by the fact
that slowly growing cells exhibit much slower cell
wall polymers biosynthesis, making more isoprenoid
lipid carrier molecules available for EPS biosynthesis
[3]. However, several investigators ¢nd higher EPS
production by LAB strains at higher cultivation temperatures [21,38,57,64,90,95] and under conditions
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Fig. 6. Batch fermentation pro¢le of Streptococcus thermophilus LY03 grown in 10% skimmed milk, 2% pepton, 1% yeast extract and
4.4% extra lactose under controlled conditions of temperature (42³C) and pH (6.2).
optimal for growth, for instance, with respect to pH
[21,64,89,94,95,98] and oxygen tension [21,64,95,98].
Aeration is not required, since higher EPS yields are
obtained with a lower oxygen tension [21,64,95] as
well as anaerobically [21,64,67,95]. Aeration may become a problem under conditions that EPS are produced using engineered strains, for instance S. thermophilus strains capable of catabolizing galactose
and producing additional ATP through the production of acetate via acetate kinase [72]. Optimal pH
conditions for production of EPS are often close to
pH 6.0 [21,64,67,89,91,94,95,98]. Van den Berg et al.
[67] postulated that conversion of sugar to EPS is
more e¤cient at pH 5.8, but sugar is more e¤ciently
converted to biomass at pH 6.2. Gassem et al. [91]
suggested that maintenance of a higher pH will result
in increased EPS production by increasing the time
the culture is in the exponential growth phase. Higher pH also results in a longer stationary phase, which
would decrease peptidoglycan and teichoic acid syntheses and could result in increased EPS production.
It has further been shown that EPS production
under growth conditions with continuously controlled pH is signi¢cantly higher than in acidi¢ed
batch cultures; moreover, it seems that the e¡ect of
pH adjustment is greater than that of supplementation with nutrients [21,57,64,91,94,95,98]. This could
be an important limiting factor when considering
industrial exploitation of EPS-producing strains in
fermented milks production since the latter takes
place under free pH conditions.
4.6. Production kinetics of exopolysaccharides from
lactic acid bacteria
Whereas mesophilic strains seem to produce maximal amounts of EPS under conditions not optimal
for growth, for instance low temperatures, EPS production from thermophilic lactic acid bacteria appears to be growth-associated, i.e. maximal production under conditions optimal for growth [64]. In the
case of growth-associated production, EPS biosynthesis generally starts almost simultaneously with
growth, shows a maximum rate when the culture is
in its exponential growth phase and reaches a maximum towards the end of the active growth, indicating primary metabolite kinetics (Fig. 6)
[12,17,21,22,57,64,67,95,96]. Marshall et al. [52] indicate that the onset of EPS biosynthesis from a strain
of L. lactis subsp. cremoris is observed toward the
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end of the exponential phase of growth. Other investigators observed continued EPS production beyond
or only in the stationary phase of growth
[17,26,45,92,98]. Such conclusions were however
often based on optical density measurements of microbial growth, which is not always a valid parameter when using complex growth media. Cell dry mass
determinations are preferred to describe growth kinetics [64]. Because of this growth-associated EPS
production with thermophilic LAB strains and because of the limited number of catabolic pathways
that provide energy in LAB (substrate level phosphorylation and secondary metabolic energy generation), cell synthesis is limited, and so is the energydemanding EPS biosynthesis. Consequently, improving EPS production must be sought in enhancing
biomass formation. Growth-association of EPS production does not occur at all with most EPS-producing Gram-negative strains such as Xanthomonas and
Alcaligenes spp. [7]; they are often produced under
nitrogen limitation during the stationary growth
phase. Also with mesophilic LAB, EPS production
may be further enhanced by varying the environmental factors once enough cells have been formed.
In addition, whereas Bouzar et al. [45] report that
the sugar composition of the EPS from Lb. delbrueckii subsp. bulgaricus CNRZ 1187 changes during the fermentation cycle, De Vuyst et al. [64] found
that EPS composition of S. thermophilus LY03 remains constant during the whole batch fermentation
process. In view of the biosynthesis mechanism involved, namely polymerization of building blocks,
the latter might be a¡ected by the activities of di¡erent sugar activation and modi¢cation enzymes,
which may be dependent on the available substrates
or speci¢c stimuli from the environment. Hence, a
variable EPS composition during fermentation
would be expected when varying substrates are fed
to the microorganism. Alternatively, in case of speci¢c stimuli from the environment di¡erent enzymes
involved in EPS biosynthesis, might be switched on
or o¡, independent of the substrate used, and, as a
result, the EPS composition might be a¡ected.
Finally, EPS degradation often takes place upon
prolonged incubation [60,64,91]. This may be due to
glycohydrolase activity [15,19,43,75,88]. However, a
marked reduction in the EPS yield upon prolonged
fermentation seems to be dependent on the strain
167
used and both chemical and physical conditions
(temperature, pH, etc.) [64,75,89^91]. Harvesting
the EPS at the appropriate time and under the appropriate conditions of pH and temperature during
isolation may avoid this problem.
4.7. Large-scale production and downstream
processing of exopolysaccharides from lactic
acid bacteria
EPS production by LAB strains can hardly compete with aerobic bacteria, such as X. campestris, as
is re£ected in the production levels of xanthan gum
(30^50 g l31 ) and those of EPS produced by LAB
(0.1^1.5 g l31 ). From an economic point of view, a
10-fold increase in EPS production by LAB, to obtain 10^15 g l31 , is required to use these EPS as a
food additive. However, a much smaller amount
would be enough to exploit in situ applications.
Since EPS production from thermophilic LAB seems
to be coupled to growth and since LAB growth is
presumably inhibited by the formation of lactate,
there is scope for productivity improvement by reducing the concentration of lactate in the culture
broth, either via fed-batch cultivation, extractive fermentation, etc. Additionally, a controlled feeding
strategy may make it possible to produce tailormade exopolysaccharides without the need of manipulating the producer strain genetically (Degeest and
De Vuyst, unpublished results). Finally, EPS generally cause (even at low levels) a signi¢cant viscosity
increase of the culture broth. This presents a di¡usion barrier or mass transfer problem for nutrients
and metabolites, such that in fact EPS biosynthesis is
stopped early by the producer strain. Also, cell separation, product recovery and puri¢cation are much
more complex as compared to conventional fermentation products [104]. The main problem in such
preparations is the high viscosity of the slime solutions which hinders deposition of the cells and EPS
separation from the carbohydrates and protein material from the (milk) medium itself. Addition of
electrolytes (salts) may be useful in EPS precipitation
by neutralizing charges on a charged polysaccharide.
If the polysaccharide is in capsule form it must ¢rst
be detached from the cells. Extracellular polymer
material is usually harvested after repeated trichloroacetic acid (to remove contaminating proteins) and
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alcohol or acetone (to isolate EPS) precipitation
steps [21,64,95]. Large-scale isolation of EPS has
been performed by Kimmel et al. [96]; 29 g of the
Lb. delbrueckii subsp. bulgaricus RR exopolysaccharide was isolated from centrifuged, ultra¢ltered fermentation broth by organic solvent precipitation.
An important loss may be ascribed to centrifugation
of a high-solids solution, so that only the bottom
layer of precipitated material is centrifuged following
organic solvent precipitation. The remaining EPS
may be present as ¢ne particles dispersed throughout
the organic solvent^retentate mixture. A ¢ltration
step to recover the EPS might result in higher recovery rates. Isolated EPS material is often found to
contain protein, which is likely carried over from
medium ingredients during recovery and is not
tightly bound to EPS [21,96]. Puri¢cation of EPS
from LAB has been successfully achieved by anion
exchange chromatography and/or gel permeation
[21,67,105]. Finally, one should also be aware that
the fermentation and downstream processing conditions may have an important impact on the conformation and physical properties of the EPS preparation.
5. Applications of exopolysaccharides from lactic acid
bacteria
5.1. Exopolysaccharides from lactic acid bacteria in
current use
EPS from LAB are not yet intentionally exploited
by industrial manufacturers. A few exceptions exist
among the homopolysaccharides produced by LAB
[1,2,7,9]. Dextran derivatives and activated dextrans
¢nd several commercial uses. Industrial dextrans are
used in the manufacture of gel ¢ltration products
and as blood volume extenders and blood £ow improvers. Further possible uses of dextrans are in paper and metal-plating processes and as food syrup
stabilizers. Also levan may ¢nd application in foods
as a biothickener. Alternan with the unique structure
of alternating K-1,6- and K-1,3-linkages thought to
be responsible for its distinctive physical properties
of high solubility and low viscosity, has potential
commercial applications as low-viscosity bulking
agent, extender, etc. in foods and cosmetics.
Scandinavian fermented milk drinks (viili,
laîngmjoëlk, laîng¢l, taette, taëtmjoëlk, piimaë, pitkaîpiimaë, ¢l, skyr) display a ¢rm, thick, slimy consistency. They rely upon the souring capacity of mesophilic ropy strains of L. lactis subsp. lactis and L.
lactis subsp. cremoris and the concomitant production of heterotype EPS for texture. Dairy starter
cultures that contain slime-forming LAB strains are
also commercially available in other parts of Europe
and the United States. Ropy, thermophilic LAB
starter cultures for yogurt production are largely
used in some countries of the EU because the addition of stabilizers is prohibited in yogurts. As mentioned above, a problem is that thermophilic LAB
produce less EPS and the ropy character is unstable.
Finally, ke¢r, an acidic, mildly alcoholic, e¡ervescent
drink fermented from grains, is a popular beverage
in Eastern Europe, where it is manufactured and
marketed on a large scale.
5.2. Unwanted exopolysaccharide production by lactic
acid bacteria
EPS from LAB may also be disadvantageous.
Some of the microbial extracellular EPS play an important (and fatal) role in in vivo bio¢lm formation
and biofouling processes. Bacterial bio¢lms that are
widespread in nature and industrial settings, are supported by a matrix partly composed of EPS. Also,
LAB contribute to this bio¢lm formation [106]. Dextrans and levans are known in the sugar industry as
microbial products that cause sucrose loss and ¢ltration problems. Mutans are considered to be critically
important in dental plaque formation and hence in
the pathogenesis of dental caries, because they are
water-insoluble and possess a marked ability to promote adherence when synthesized de novo on various solid surfaces [88]. Cell-bound glucan appears to
promote the establishment of S. mutans and other
glucan-producing streptococci on the heart valves
causing subacute bacterial endocarditis. Other human pathogens such as S. pneumoniae produce a
polysaccharide capsule which is the cause of virulence. Microbial bio¢lm formation on heat exchanger plates in cheese and liquid milk factories can be a
source of the bacterial contamination of dairy products, leading, for example, to an excessive openness
in cheese or taste deviations of milk. Adhesion of
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thermophilic strains to heat-exchanger plates on the
downstream side of the regenerator section of pasteurizers is especially troublesome, as this may cause
contamination of already pasteurized milk [107,108].
Slime-producing LAB strains are further known
from their spoilage of beers [109], wines [110], ciders
[111], sugar-salted herring [112], vacuum-packed
cooked meat products [113], fermented sausage
[113^118], etc. Typical slime-forming food spoilers
are P. damnosus (beer, wine and cider), P. dextrinicus
(beer), P. inopinatus (beer), Lb. sake (meat), etc.
5.3. Novel applications of exopolysaccharides from
lactic acid bacteria
The intentional and controlled use of EPS from
LAB as natural food additives or of functional starter cultures, i.e. strains producing interesting EPS,
could result in a safe, natural endproduct, and may
have an important impact on the development of
novel food products (both fermented and non-fermented food products), especially food products
with enhanced, rheological properties, improved texture and stability, and/or water retention capacity.
For instance, low viscosity, gel fracture or high syneresis (whey separation) problems, which may occur
during yogurt manufacture, can be solved in both
ways (cf. infra). An EPS-producing S. thermophilus
strain was also responsible for an increased moisture
level in low-fat mozzarella cheese [66,119]. Since the
use of LAB (GRAS microorganisms) is historically
safe, production of in situ novel functional EPS
means that toxicological testing will not be required
and the products can be brought to the market more
quickly.
5.4. Application of exopolysaccharides from lactic
acid bacteria as food additives
The discovery of the EPS-producer Leuc. mesenteroides in 1878, responsible for the thickening and
gelling of sugar beet and sugar cane syrup, can be
considered as the start of the possible use of EPS
from LAB in food products. The use of xanthan
from X. campestris, the ¢rst microbial polysaccharide
that was allowed in food products, has been approved by the US Food and Drug Administration
in 1969. Recently, gellan from S. elodea has been
169
commercialized too. EPS from LAB which are
GRAS bacteria may form a new generation of biothickeners. However, although in some food applications the function of a polysaccharide additive seems
straightforward, it has to be realized that in most
cases the consistency and texture of the ¢nal product
is the result of a complex set of interactions between
the di¡erent food components, which requires EPS
with the right structure, conformation and properties.
5.4.1. Use as viscosifying agent
To be used as biothickener in foods, for instance,
in yogurts and other fermented milks, dairy desserts,
soups, sauces and salad dressings, the hydrated polymers should exhibit thixotropic or pseudoplastic
(shear-thinning) properties, i.e. their rheology should
decrease markedly upon shaking, stirring, or pouring, but recover completely when shear is removed.
The polymers must be compatible with any other
food components present or food processing conditions applied, i.e. stability of function is often required over a wide range of pH, ionic strength,
and/or temperature. This is the reason for the widespread use of xanthan that displays such characteristics. However, the viscosity of the Lb. sake 0-1 EPS
was higher throughout a range of increasing shear
rates, while shear-thinning properties were as good
as those of xanthan gum [67] (Fig. 7). Good shearthinning properties, i.e. a (reversible) drop in viscosity with increasing shear rate, are important for food
applications when considering processing costs (manufacturer) and mouthfeel and texture/consistency
(consumer) of food products. Viscosity is further
highly dependent on the average molecular mass distribution. As the molecular mass of both xanthan
and the Lb. sake 0-1 EPS is of the same order of
magnitude, the high viscosity of the 0-1 EPS is an
intrinsic property and is not due to large di¡erences
in molecular mass [67]. Also other EPS display interesting intrinsic viscosities, indicating that the polymers have remarkable thickening properties
[11,21,44,46].
5.4.2. Use as gelling agent
The alternative application of many of the biopolymers is as gelling agents. To form a polysaccharide
gel a three-dimensional network in which the poly-
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Because these products require increasing thickening
properties, the use of slime-producing starter cultures
in their manufacture is necessary.
Fig. 7. Flow curves of a 1% (m/v) aqueous solution of puri¢ed
Lb. sake 0-1 EPS and xanthan gum. E, viscosity of xanthan; F,
viscosity of 0-1 EPS; a, shear stress of xanthan; b, shear stress
of 0-1 EPS. (From van den Berg et al. [67]).
saccharide chains are crosslinked, must be formed,
for instance via interactions between negatively
charged polymer chains and positively charged ions
or via interactions of neutral polymer chains through
hydrogen bonds and Van-der-Waals forces. So far,
no gelling properties or uses of heteropolysaccharides (which are neutral or slightly negatively
charged) from LAB have been reported.
5.5. Application of slime-producing starter cultures
An alternative way to improve yogurt viscosity
and decrease susceptibility to syneresis and graininess is by utilizing the slime-producing strains in
the starter culture. The ropy strains may further contribute to the consistency of stirred-type yogurt, produced on a large scale, because yogurt containing
viscosifying EPS is supposed to be less damaged mechanically from pumping, blending and ¢lling machines. In addition, the coagulum would be more
resistant to thermal and physical shocks. This ultimately leads to the manufacture of yogurt without
the addition of stabilizers. This type of production
process gains increased popularity in the Western
world, because of the increased desire of the consumer for 100% natural products. Finally, EPS may play
an important role in the production of other yogurt
drinks and low-milk-solid yogurts, as well as in the
production of creamier yogurts with low or no fat
content and an enhanced smoothness of mouthfeel.
5.5.1. Rheology versus exopolysaccharide
concentration
In general, it is found that the apparent viscosity
of stirred yogurt increases with the ropiness of the
culture [120^125]. However, no clear correlation between this apparent viscosity and the concentration
of EPS is found for stirred yogurt produced with
ropy, moderately ropy and non-ropy cultures
[18,19,45,87,97,122^124,126,127]. The concentration
of isolated EPS is often low or not detectable. It is,
however, obvious that EPS are necessary to increase
the viscosity, but the magnitude of increase varies
because of di¡erences in culture strains, incubation
conditions, total solids of the medium, and viscosity
measurements [8,11]. Moreover, viscosity may not
only be a¡ected by the amount of EPS released,
but also by an EPS with slightly di¡erent structure
and apparent molecular mass, resulting in di¡erent
rheological characteristics of the medium. It hence
appeared that EPS concentration cannot alone explain the e¡ect of ropiness on the apparent viscosity
of yogurt and that the e¡ect of EPS appeared to be
much more complex than formerly thought. Indeed,
in fermented milk products, such as yogurt, the
structure and consistency mainly originate from the
aggregated proteins. The microorganisms and/or the
EPS they produce may a¡ect the protein aggregation, thereby changing the physical properties of
the gel and the ¢nal product after stirring (cf. infra).
Interactions between EPS and milk constituents, in
particular caseins that are precipitated at low pH,
and between EPS and the bacterial cell surface,
have been reported [43,45,88,120,121,123,128^130].
The gel structure has mainly been studied by scanning electron microscopy. In milk fermented with a
ropy Lb. delbrueckii subsp. bulgaricus strain, EPS
strands are observed between the cells and the milk
protein network, while a uniform layer of EPS covers the cells. However, EPS-like strands could be
seen in gels made with a non-ropy starter culture.
Anyway, yogurts manufactured using EPS-producing strains have properties that are distinctly di¡erent from those of yogurts made using non-producers
with or without commercially available stabilizers
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[131]. In addition, yogurts made using di¡erent EPSproducing strains had di¡erent rheological characteristics, suggesting the possibility of developing cultures that produce yogurts with speci¢c rheological
properties.
5.5.2. Yogurt
Yogurt consists of a network of casein micelles
that associate to form a gel-like structure. This protein network has interstitial spaces that contain the
liquid phase as well as larger void spaces containing
starter bacteria. By trapping the whey, the casein
matrix creates the typical viscoelastic properties of
yogurt texture. Stirred yogurt is a product resulting
from further processing of set yogurt. The retention
of whey is vital to product integrity and not only the
contribution of acid secretion and microbial proteinases is essential for texture development, but also
exopolysaccharides secreted by the yogurt strains
contribute to this. The use of ropy cultures results
in yogurts with a more stable texture for shear.
When subjected to a shear force changes occur in
the microstructure of ropy yogurt. An increase of
shear rate ¢rst disrupts the attachment of polymer
to the bacterial surface, but the polysaccharide material remains incorporated with the casein where it
continues to in£uence viscosity of yogurt [121]. If
interactions of EPS and protein require more energy
to break than interactions among the protein that
are responsible for yogurt gel texture, an initial decrease in viscosity, when examining shear stress
against shear rate, could be due to the disrupted
interactions between proteins, and a reduced rate
of shear thinning observed later could be explained
by the increased energy required to disrupt interactions among the bacteria, EPS and protein [132].
Skriver et al. [133] obtained similar results. The
fact that in their experiments yogurt made at 32³C
was less shear-thinning than yogurt made at 43³C is
indicative of a greater number of interactions among
bacteria, EPS and protein at the higher temperature.
Rohm and Kovac [134] suggested that the ropy yogurts contained protein^polysaccharides bonds that
had shorter relaxation times than protein^protein
bonds contained in non-ropy yogurts, implying
that, in case of highly viscous products, the probability of certain bonds breaking within a period of
time is greater than in classical yogurts. They as-
171
sumed that protein strand formation and protein^
protein bond development is partly prevented by excessive formation of polysaccharide ¢laments attached to the protein matrix and thus reducing rigidity of the resulting yogurt gels. Prentice [135],
however, comments that there is likely to be a wide
spectrum of relaxation times, that, over time, bonds
that have broken will reform over di¡erent time periods. Also, yogurt made with ropy cultures has a
more open structure with larger pores than does yogurt made with non-ropy strains [136]. These void
spaces may a¡ect the integrity of the food matrix,
resulting in a coagulum that is less ¢rm [137]. Also
the data of Hess et al. [131] are consistent with a
mechanism for shear-induced structural degradation
of yogurt made using EPS-producing strains in
which polymer associated with the casein network
prevents disruption of portions of the network. Finally, Rawson and Marshall [125] con¢rmed via
rheology and penetrometry studies that inclusion of
a ropy strain will not always lead to improved texture attributes; while ropy strains may increase viscosity, they may not in£uence ¢rmness which would
be more in£uenced by protein^protein interactions.
Certainly, all stirred yogurt has an apparent viscosity
and a texture which is subject to shear rate-thinning
and which can recover, i.e. exhibits partial thixotropy, and it is generally accepted that the behavior of
stirred yogurt is dependent on its shear history.
However, the physical properties imparted to yogurt
systems by non-ropy and ropy starter cultures are
usually evaluated by means of `destructive' rheological testing such as rotational viscometry or penetrometry. These techniques fail to reveal many of
the structural characteristics of yogurt systems themselves, as they e¡ectively measure the remnants of
the gel network after disruption. Since the properties
of stirred yogurt will originate from the properties of
the gel before stirring, a detailed study of the gel
structure before stirring was performed, applying
dynamic shear and permeability measurements
[123,138]. It is suggested that the spatial structure
of the protein network of yogurt may be an important factor determining the properties of stirred lowfat yogurt [123]. Although ropiness is still supposed
to be attributable to the extracellular polysaccharide,
it has been shown that the macrostructure demonstrated an inverse relationship between ropiness and
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permeability of the gel and that the protein network
is equally important. EPS production in a gel with a
homogeneous protein structure resulted in a decrease
in permeability of the gels and an increase in viscosity of the stirred yogurt. However, if the culture
caused an inhomogeneous protein network, an inhomogeneous, low viscosity stirred yogurt was achieved
and the EPS were not able to compensate for that
[123]. Indeed, rheological measurements that are indicative of gel strength (i.e. penetration, consistency
coe¤cient and both storage and loss moduli) indicate that yogurts made with EPS-producing strains
form weaker coagula [131,138]. All the results suggest that the mechanisms of structural degradation
are di¡erent for yogurts made using EPS-producing
and non-producing strains. Thus, ropiness, a characteristic which is imparted to the product as a result
of fermentation with particular EPS-producing
strains, contributes to `adhesiveness', while `¢rmness'
and `elasticity' are likely to be in£uenced more by the
protein matrix of the yogurt than by secretion of the
EPS by the ropy strains [125].
5.5.3. Viili
Measurements of texture showed that a Nordic
milk gel prepared by the ropy strain of L. lactis
subsp. cremoris SBT 0495, isolated from Finnish
ropy sour milk `viili' starter culture, exhibited remarkably increased adhesiveness as compared to
that by the non-ropy variant. Milk gel prepared by
the ropy strain also exhibited decreased syneresis
(wheying-o¡) as compared to that by the non-ropy
variant. Scanning electron micrographs of milk gel
prepared by the ropy strain showed that slime was in
the form of a network attaching the bacterial cells to
the protein matrix. A thick network of slime attached the casein micelle clusters to each other to
make casein conglomerates, which is likely to result
in the characteristic consistency of `viili' [139].
5.5.4. Ke¢r
Ke¢r grains consist of a polysaccharide gel embedding LAB and yeasts [23,24]. Lb. ke¢ranofaciens is
an important organism associated with ke¢r grains.
It produces the ke¢ran polymer that forms the matrix of the ke¢r grains [140^143]. Other microorganisms associated with ke¢r are the homofermentative
strains Lb. acidophilus and Lb. ke¢rgranum, the ob-
ligately heterofermentative strains Lb. ke¢r and Lb.
parake¢r, and the yeast Candida ke¢r [144]. The
polysaccharide from Lb. ke¢ranofaciens seems to
play an important role in maintaining the ecological
niche, which is necessary as the grains are recovered,
dried and reused for many successive milk inoculations [145].
6. Conclusion
Such a large number of possible EPS structures
from LAB exists that more polymers of potential
industrial value will almost certainly be obtained. If
these biomolecules are to be developed commercially, however, they must be cost e¡ective. The use
of whey could be a means of upgrading this dairy
byproduct by lactose fermentation with concurrent
production of a useful EPS polymer for food applications. Thus, major improvements must be sought
in fermentation techniques to ensure that conversion
of substrate to product is maximal. Control of fermentation conditions must be ensured otherwise
product yield as well as quality cannot be guaranteed. Also, downstream processing to recover the
product can probably be improved and costs lowered. While some of these developments will be
achieved by the industrial producers, others may
well result from basic studies in academic laboratories. Knowing the environmental and genetic factors
regulating expression of the EPS, genetic approaches
can be designed which enhance expression of a desired EPS under de¢ned growth or fermentation conditions. Finally, both genetic approaches and enzyme
and fermentation technology will increase the number of possibilities for modifying the structure and
function of EPS. This polysaccharide engineering
may lead to the development of `designer polysaccharides' for applications that may or may not be
food related. Although a popular high-tech concept,
the technology for tailoring polysaccharides to speci¢c uses is still in its infancy.
Acknowledgments
The authors' research on exopolysaccharide production by lactic acid bacteria was ¢nancially sup-
FEMSRE 643 16-4-99
L. De Vuyst, B. Degeest / FEMS Microbiology Reviews 23 (1999) 153^177
ported by the Institute Danone by means of a `Navorsingskrediet voor Fundamenteel Voedingsonderzoek'. The authors also acknowledge ¢nancial supportfrom the European Commission (EU), the
Flemish Institute for Encouragement of Scienti¢c
and Technological Research in Industry (IWT), the
Fund for Scienti¢c Research (FWO ^ Flanders) and
the Research Council of the Vrije Universiteit Brussel (VUB).
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