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Exopolysaccharides of Lactic Acid Bacteria for Food and Colon Health Applications

Written By

Tsuda Harutoshi

Submitted: November 20th, 2011 Published: January 30th, 2013

DOI: 10.5772/50839

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1. Introduction

Lactic acid bacteria (LAB) are used in many fermented foods, particularly fermented dairy products such as cheese, buttermilk, and fermented milks. LAB produce lactic acid, carbon dioxide, and diacetyl/acetoin that contribute to the flavor, texture, and shelf life of fermented foods. Some LAB produce exopolysaccharide (EPS), and generally, EPS play a major role as natural texturizer in the industrial production of yoghurt, cheese, and milk-based desserts. Recently, EPS produced by LAB have received increasing attention, mainly because of their health benefits. In particular, immune stimulation, antimutagenicity, and the antitumor activity of fermented dairy products prepared with EPS-producing LAB or EPS themselves have been investigated [1-4].

EPS are polysaccharides secreted from the cell, or produced on the outer cell by extracellular enzymes. EPS from LAB are divided into two classes, homo- and hetero-EPS. Homo-EPS are composed of one type of monosaccharide, whereas hetero-EPS consist of regular repeating units of 3-8 different carbohydrate moieties synthesized from intracellular sugar nucleotide precursors [5]. The biosynthesis of homo-EPS and hetero-EPS are different. Homo-EPS are made from sucrose using glucansucrase or levansucrase [6-7], and the synthesis of hetero-EPS involves four major steps, sugar transportation, sugar nucleotide synthesis, repeating unit synthesis, and polymerization of the repeating units [8]. The major physiological function of EPS is believed to be biological defenses against various stresses such as phage attack, toxic metal ions, and desiccation [9], and it is very unlikely that bacteria use EPS as an energy source. However, some potentially probiotic LAB strains have been reported to degrade EPS produced by the other LAB strains [10-11].

The term "probiotic" was first proposed by Fuller [12], and its definition was further refined to "Live microorganisms which when consumed in adequate amounts as part of food confer a health benefit on the host" [13]. Probiotic LAB thus represent a class of live food ingredients that exert a beneficial effect on the health of the host. Beneficial microorganisms in the intestine are enhanced by “prebiotics,” which are defined as "nondigestible food ingredients that beneficially affect the host by selectively stimulating the growth and activity of one or a limited number of bacterial species already resident in the colon, and thus improving host health" [14].

Most of the current prebiotics are low molecular weight except for inulin. As long carbohydrate chains are metabolized more slowly than the short ones, and polysaccharides thus exert prebiotic effects in more distal colonic regions compared to oligosaccharides, which are more rapidly digested in the proximal colon [15]. Therefore, EPS produced by LAB can be used as prebiotics. This chapter reviews the physicochemical properties, genetics, and bioactivities of the EPS produced by LAB.


2. Chemical composition of EPS

2.1. Homo-EPS

Some LAB can produce EPS that are either secreted to the environment or attached to the cell surface forming capsules. EPS are classified into two groups: homo-EPS, consisting of a single type of monosaccharide (α-D-glucans, β-D-glucans, fructans, and others represented by polygalactan) and hetero-EPS, composed of different types of monosaccharides, mainly D-glucose, D-galactose, L-rhamnose, and their derivatives [16].

The differences arise between the homopolysaccharides mainly because of the features of their primary structure such as the pattern of main chain bonds, molecular weight, and branch structure. Two important groups of homo-EPS are produced by LAB; (i) α-glucans, mainly composed of α-1,6- and α-1,3-linked glucose residues, namely dextrans, produced by Leuconostoc mesenteroides subsp. mesenteroides and Leuconostoc mesenteroides subsp. dextranicum and mutans produced by Streptococcus mutans and Streptococcus sobrinus; and (ii) fructans, mainly composed of β-2,6-linked fructose molecules, such as levan produced by Streptococcus salivarius [17].

The formation of dextran from sucrose has been recorded for Leuc. mesenteroides subsp. mesenteroides. However, the ability to form dextran is often lost when serial transfers are made in media with increasing salt concentrations. Nevertheless, non-dextran-producing strains of Leuconostoc sp. can revert to dextran production when they are inoculated into medium containing tomato or orange juice [18]. In the 1950s, the use of a cell-free enzyme solution permitted dextran synthesis under controlled conditions yielding a polymer of greater purity. A common feature of all dextrans is the preponderance of α-1,6-linkages with branch points at positions 2, 3, or 4 [17]. Some strains of Leuconostoc amelibiosum [19] and Lactobacillus curvatus [20] are reported to be dextran-producing strains.

Mutan is the glucan synthesized by various serotypes of Str. mutans, and differs from dextran in that it contains a high percentage of α-1,3 linkages. Differences in solubility result from the proportions of different types of linkages; water-soluble glucans are rich in α-1,6 linkages, while water-insoluble glucans are rich in α-1,3 linkages [17]. Ingestion of mutan has been linked with dental caries, as insoluble mutans can adhere to teeth, thus helping microorganisms adhere to the surface of teeth.

Homo-EPSMain linkage(branching linkage)Organism
Dextranα-1,6 (α-1,3)Leuc. mesenteroides subsp. mesenteroides, Leuc. mesenteroides subsp. dextranicum, Leuc. amelibiosum, Lb. curvatus
α-1,3 (α-1,6)
Str. mutans
Alternanα-1,3 and α-1,6Leuc. mesenteroides
Levanβ-2,6 (β-2,1)Leuc. mesenteroides, Lb. reuteri
β-2,1 (β-2,6)
Str. mutans

Table 1.

Homo EPS produced by LAB

Alternan has alternate α-1,6 and α-1,3 linkages, and this structure is thought to be responsible for its distinctive physical properties including high solubility and low viscosity. These characteristics provide this glucan with a potential commercial application as a low viscosity texturizer in foods. Leuc. mesenteroides NRRL B-1355 was first reported to be an alternan-producing strain [21]

Levan is an EPS produced from sucrose. It is fructan composed of β-2,6-linked fructose molecules with some β-2,1-linked branches. Incidentally, inulin is a fructan composed of β-2,1-linked fructose molecules with some β-2,6-linked branches. Str. salivarius, Leuc. mesenteroides, and Lactobacillus reuteri are known to be levan-producing LAB [22-23]. In addition, the EPS produced by Lactobacillus sanfranciscensis TMW 1.392 has been reported to be fructan [11].

2.2. Hetero-EPS

The chemical composition of hetero-EPS shows wide variablity. Hetero-EPS are polymerized repeating units mainly composed of D-glucose, D-galactose, and L-rhamnose. The composition of the monosaccharide subunits and the structure of the repeating units are considered not to be species-specific, except in case of Lactobacillus kefiranofaciens subsp. kefiranofaciens. This species, isolated from kefir grain, a fermented dairy food from the North Caucasus region, produces large amounts of polysaccharides [24]. Hetero-EPS-producing strains of Streptococcus thermophilus, Lactococcus lactis, Lactobacillus delbrueckii, and Lactobacillus helveticus, among others have been identified (Table 2) [25-49]. Heterofermentative LAB such as Leuc. dextranicum are well known homo-EPS producers, while homofermentative LAB are well-studied hetero-EPS producers. Heterofermentative in addition to homofermentative LAB can produce EPS. Lactobacillus fermentum is an EPS-producing heterofermentative LAB for which the EPS structure has been determined [50]. Figueroa et al. reported that Lactobacillus brevis and Lactobacillus buchneri showed ropiness on glucose- or sucrose-containing media, although they did not investigate whether such ropiness derived from hetero-EPS or from other slimy substances [51].

The quantities of hetero-EPS produced by LAB vary greatly. EPS production is 50-350 mg/l for Str. thermophilus, 80-600 mg/l for Lc. lactis subsp. cremoris, 60-150 mg/l for Lb. delbrueckii subsp. bulgaricus, 50-60 mg/l for Lactobacillus casei [52], and approximately 140 mg/l for Lactobacillus plantarum [45, 53]. The highest recorded yields of hetero-EPS are 2775 mg/l for Lactobacillus rhamnosus RW-9595M [54] and 2500 mg/l for Lb. kefiranofaciens WT-2B [55]. However, the quantities of EPS produced by LAB are much lower than the yields from other industrially important microorganisms such as Xanthomonas campestris, which produces 30-50 g/l xanthan gum [56]. Even so, amounts of EPS produced by LAB are sufficient to exploit for in situ applications. LAB are ‘generally recognized as safe’ (GRAS) microorganisms, and LAB strain culture would be a useful method to produce EPS for food applications if the LAB could be grown in edible and safe culture media such as whey, and if fermentation conditions were optimized to obtain a high yield.

Fermentation conditions using undefined media have been improved to maximize yields. However, a chemically defined medium containing a carbohydrate source, mineral salts, amino acids, vitamins, and nucleic acid bases is more suitable for investigating the influence of different nutrients on LAB growth and EPS biosynthesis. The total yield of EPS produced by LAB depends on the composition of the medium (carbon and nitrogen sources) and the growth conditions, i.e., temperature, pH, and incubation time.

Under conditions of higher temperatures and slower growth, the production of the polymer per cell in Lb. delbrueckii subsp. bulgaricus NCFB 2772 was greater in milk [57]. Another study investigated the optimum culture conditions for EPS production by Lb. delbrueckii subsp. bulgaricus RR in semidefined medium [58], and determined the optimum temperature and pH conditions for EPS production to be 36°C - 39°C and pH 4.5 - 5.5. The optimal temperature for EPS production was approximately 40°C for thermophilic LAB strains, and around 25°C for mesophilic LAB. Gamar et al. [59] reported increased slime production at lower incubation temperatures, and an increase in the final EPS concentration in Lb. rhamnosus following incubation at 25°C instead of 30°C. The effects of temperature on EPS production in whey were investigated in Lb. plantarum [53], and the yield was found to be higher at 25°C than at either 30°C or 37°C. Moreover, an inverse relationship was observed between EPS production per cell and the growth temperature for Lactobacillus sake [49], i.e. the lower the temperature, the higher the EPS production per cell. However, the growth rate in the exponential phase decreased at low temperatures. Therefore, the temperature for the maximal production of EPS is based on a balance of cell density and EPS production per cell. Maximal EPS production by Lb. sake was obtained under anaerobic conditions at 20°C, although EPS production per cell was higher at 10°C. Therefore, it is possible that severe environmental conditions trigger EPS production as a protective mechanism.

The effects of alterations to the nitrogen and carbon sources used in EPS production have also been investigated. According to early reports, neither LAB growth nor EPS production was specifically linked to the presence of casein or whey proteins in the growth medium. Garcia et al. [57] reported that EPS production by Lb. delbrueckii subsp. bulgaricus NCFB 2772 increased during the early growth pase in the presence of hydrolyzed casein in milk, while the addition of hydrolyzed casein to MRS medium did not increase EPS production. This strain produced 25 mg/l EPS when grown on fructose in a defined medium, and 80 mg/l EPS when grown on glucose [60]. The optimum Bacto-casitone concentration for EPS production by Lb. delbrueckii subsp. bulgaricus RR was investigated in semidefined medium [58]. In this study, there was a significant relationship between the Bacto-casitone concentration and EPS production; the higher the casitone concentration, the higher the EPS yield that was obtained. For Lb. plantarum grown in whey, yeast extract was a more effective nitrogen source for EPS production than soy peptide, tryptone, peptone, and Lab-Lemco powder, and glucose was a more effective carbon source than galactose, sucrose, maltose, fructose, and raffinose [53]. EPS production by Lb. casei CG11 was investigated in basal minimum medium containing galactose, glucose, lactose, sucrose, maltose, and melibiose; glucose was the most efficient carbon source, and lactose and galactose were the least efficient ones [61]. EPS production by Lb. rhamnosus C83 was investigated in a chemically defined medium containing different carbon sources (glucose, fructose, mannose, and maltose) at different concentrations. Mannose at 40 g/l was by far the most efficient carbon source. Furthermore, increased Mg, Mn and Fe concentrations stimulated EPS production in synthetic media [59]. In addition, Macedo et al. [54] reported about the importance of salts in culture media and the strong positive effect of salts and amino acids on Lb. rhamnosus RW-9595M growth and EPS production. The addition of salts and amino acids largely increased EPS production (to 2775 mg/l) in whey permeate supplemented with yeast extract, although the addition of amino acids alone had no effect on EPS production.

It has been shown that an optimal ratio between the carbon and nitrogen is absolutely necessary to achieve high EPS yields [62]. The production of EPS by Str. thermophilus LY03 is modulated by both the absolute quantities and the ratio of carbon to nitrogen (C/N ratio). The carbon source is converted into lactic acid to produce energy as well as to synthesize the cell wall and EPS, and nitrogen is necessary for the synthesis of essential cell components. Therefore, a higher C/N ratio and sufficient quantities of both carbon and nitrogen increase EPS production.


3. EPS biosynthesis by LAB

3.1. Homo EPS biosynthesis

Homo EPS are synthesized outside the cell by specific glycosyltransferase (GTF) or fructosyltransferase (FTF) enzymes (commonly named glucansucrases or fructan-sucrases). Homo-EPS producing LAB also use extracellular GTF enzymes to synthesize high-molecular mass α-glucans from sucrose. This process uses sucrose as a specific substrate, and the energy required for the process comes from sucrose hydrolysis. There is no energy requirement for EPS-production other than for enzyme biosynthesis because EPS synthesis by GTF or FTF does not involve active transport processes or the use of activated carbohydrate precursors. Therefore, large amounts of sucrose can easily be converted to EPS. Lb. sanfranciscensis produces up to 40 g/l levan and 25 g/l 1-kestose during growth in the presence of 160 g/l sucrose [63].

Glucan synthesis reactions catalysed by GTF can be written as follows (Fig. 1):

sucrose + H2O → glucose + fructose

sucrose + acceptor carbohydrate → oligosaccharide + fructose

sucrose + glucan (n) → glucan (n+1) + fructose

Although GTF enzymes have a high degrees of similarity, lactobacilli produce a broad spectrum of glucans, including polymers with α-1,6 linkages (dextran), α-1,3 linkages (mutan), and both α-1,6 and α-1,4 linkages (alternan). The relative molecular weight of glucans from lactobacilli range from 1 × 106 Da to 5 × 107 Da [6]. In addition, GTF enzymes are not saturated by their substrate, and transfer reactions exceed the sucrose hydrolysis under sucrose concentrations above 100 mM [64].

Figure 1.

The dextran synthesis by GTF (dextran sucrase).

The GTF enzymes of streptococci are generally produced constitutively. In contrast, the GTF enzymes of Leuconostoc species are specifically induced by sucrose. For example, GTF expression in Leuc. mesenteroides is low in the presence of carbon sources other than sucrose and is increased by the addition of sucrose [5]. GTF expression during sucrose fermentation is 10-15-fold higher than that measured during glucose fermentation in Leuc. mesenteroides Lcc4. In fed-batch fermentation with both glucose and sucrose, GTF activity was similar to that obtained with sucrose alone. These results show that GTF expression is low in the presence of glucose alone, and that GTF activity is significantly induced by sucrose. A sucrose concentration of 20 g/l is sufficient to ensure the induction of enzyme synthesis, and higher concentrations (up to 60 g/l) do not lead to a further increase in enzyme synthesis [65].

The fructan synthesis reaction catalyzed by FTF can be written as follows:

sucrose + H2O → fructose + glucose

sucrose + acceptor carbohydrate → oligosaccharide + glucose

sucrose + fructan (n) → fructan (n+1) + glucose

Fructans generally have a relative molecular weight exceeding 5 × 106 Da. Similar to GTFs, FTFs are not saturated by their substrate, namely, sucrose, and transfer reactions exceed the rate of sucrose hydrolysis for sucrose concentrations above 200 mM [5]. FTFs such as Lev, Inu, and LevS from lactobacilli exhibit pH optima of between 5.0 and 5.5. The optimum temperature for enzymes from the thermophilic Lb. reuteri is higher (50°C) than that of the Lb. sanfranciscensis enzyme (35°C – 40°C) [5].

3.2. Hetero EPS biosynthesis

Hetero EPS are not synthesized by extracellular enzymes, but are instead synthesized by a complex sequence of interactions involving intracellular enzymes. EPS are made by polymerization of repeating units, and these repeating units are built by a series of addition of sugar nucleotides at the cytoplasmic membrane. Sugars are the starting materials for the synthesize sequence. LAB strains can utilize various monosaccharides and disaccharides as energy sources, via some well-studied sugar uptake systems include primary transport systems, direct coupling of sugar translocation to ATP hydrolysis via a transport-specific ATPase; secondary sugar transport systems, coupling of sugar transport to the transport of ions or other solutes, both as symport and antiport transport systems; and group translocation systems, coupling of sugar transport to phosphorylation via the phosphoenolpyruvate (PEP)-dependent phosphotransferase system (PTS; Fig. 2) [8]. Polysaccharides must be hydrolyzed before uptake. For example, starch is hydrolyzed by α-amylase, and the raction products are subsequently hydrolyzed by the enzymes described above.

Lc. lactis strains possess a lactose-specific PEP-PTS sugar transport system that imports extracellular lactose, resulting in increased intracellular lactose-6-phosphate. Lactose-6-phosphate is then hydrolyzed, and the galactose-6-phosphate moiety is metabolized by the tagatose-6P pathway (Fig. 2).

Lb. delbrueckii subsp. bulgaricus and Str. thermophilus are generally galactose-negative and take up lactose via a lactose/galactose antiport transport system. The glucose moiety of imported lactose is fermented by these strains, while the galactose moiety is excreted via the lactose/galactose antiport system.

After the addition of a hetero-EPS repeating unit, the unit is exported through the cell membrane and becomes polymerized into the final hetero-EPS. Hence, several enzymes and proteins are involved in the biosynthesis and secretion of heterotype EPS, and the enzymes and proteins involved in these processes may not be unique to hetero-EPS anabolism.

Sugars taken into the cell are converted into sugar nucleotides. Iintracellular monosaccharides are converted to sugar nucleotide substrates for polymerization reactions, including UDP (uridine diphosphate), dNTP (thymidine diphosphate), and GDP (guanosine diphosphate). Such polymerization reactions are catalyzed by glycosyl pyrophosphorylases.

Glu-1P (Gal-1P) + UTP → UDP-Glu (UDP-Gal) + pyrophosphate

UDP-glucose is then converted to UDP-galactose by epimerases such as UDP-glucose-4-epimerase. This reaction is reversible.

UDP-glucose ↔ UDP-galactose

Glycosidic linkages are formed on membranes in the cytoplasm. A sugar moiety is transferred to C55-polyprenyl phosphate, a carrier lipid and component of the membrane, by priming glycosyl transferases. This transfer triggers the addition of a repeating unit to the hetero-EPS molecule. Disruption of the priming glycosyl transferase gene generates non-EPS-producing mutants [66]. Thus, priming glycosyl transferases are thought to be crucial for EPS biosynthesis. The addition of the repeating unit is completed by the action of glycosyl transferase on the sugar residue attached to C55-polyprenyl phosphate. Therefore, the type and number of glycosyl transferases available determine the range of repeating units in hetero-EPS. C55-polyprenyl phosphate is also involved in bacterial cell wall biosynthesis, and therefore, cell wall biosynthesis and EPS synthesis compete for this substrate. The repeating unit is exported through the bacterial membrane, and is polymerized to become a hetero-EPS (Fig. 3).

Figure 2.

Pathway of lactose fermentation in lactic acid bacteria.

Figure 3.

Outline of biosynthesis of hetero EPS.PGM: α-phosphoglucomutase, UGP: UDP-glucose pyrophospholyraseUGE: UDP-galactose 4-epimerase, TGP: dTDP-glucose pyrophospholyraseTRS: dTDP-rhamnose synthetic enzyme system, PMI: phosphomannoisomerasePMM: phosphomannomutase, GMP: GDP-mannose pyrophospholyrase

3.3. Instability of EPS production

The instability of hetero-EPS production has been reviewed by de Vuyst et al. [8]. Briefly, a loss in the ability to produce slime may be caused by repeated subculture of bacterial strains or incubation at high temperatures. The loss of plasmids from ropy mesophilic LAB strains is generally the reason for loss of slime production. On the other hand, thermophilic LAB, namely, Lb. delbrueckii subsp. bulgaricus and Str. thermophilus, have been shown to lack plasmids encoding components required for slime production. These species can usually recover the ability to produce slime following loss due to culture conditions. Thus, genetic instability could be a consequence of the actions of mobile genetic elements such as insertion sequences. Recently, the EPS gene cluster in Lb. fermentum TDS030603 was reported to be located in chromosomal DNA [67].

Priming glycosyl transferases are thought to be crucial for EPS biosynthesis and disruption of the priming glycosyl transferase gene generates non-EPS-producing mutants. Tsuda et al. generated the EPS-producing mutant strain 301102S from the non-EPS-producing Lb. plantarum 301102 following exposure to the mutagens acridine orange and novobiocin [4]. The activities of α-phosphoglucomutase (PGM), UDP-glucose pyrophosphorylase (UGP), and UDP-galactose 4-epimerase (UGE) were measured in parental and mutant strains by using the method of Mozzi [68], and were found to be almost the same for both [Tsuda & Miyamoto, unpublished data]. Next, priming glycosyl transferase genes in parental and mutant strains were amplified with the thermal cycler. Primers were designed to amplify a priming glycosyl transferase gene referring to complete Lb. plantarum WCFS1 genome sequenced [69]. PCR products were subjected to restriction digestion, which allowed identification of putative priming glycosyl transferase gene. PCR products were also applied to single strand conformation polymorphism (SSCP) analysis for detecting point mutations. However, both parental and mutant strains had the same priming glycosyl transferase gene sequence, and similar levels of activities of the PGM, UGP, and UGE enzymes. Thus, although priming glycosyl transferases are essential, other factors may also be necessary for EPS production, and a mutation affecting EPS production may occure in another gene. Morona et al. reported that an autophosphorylating protein-tyrosine kinase is essential for encapsulation in Streptococcus pneumoniae [70]. A point mutation in the gene encoding the autophosphorylating protein-tyrosine kinase affecting the ATP-binding domain resulted in loss of EPS production.


4. Polysaccharides and oligosaccharides for colon health

EPS produced by LAB have various functional roles in human or animal health including immunomodulatory properties, antiviral activity, antioxidant activity, and antihypertensive activity [1, 55, 71, 72], and have also been used as food additives for texture improvement. These properties have been extensively reviewed [8, 9, 56, 73, 74]. Besides these properties, prebiotics based on LAB and oligosaccharides have other health benefits. Prebiotics are usually non-digestible oligosaccharides that selectively stimulate the growth and activity of a limited number of bacterial species in the colon, such as bifidobacteria and lactobacilli, and therefore, improve host health. Detrimental bacteria may form substances such as ammonia, hydrogen sulfide, indles, and amines that are noxious to the host. However, beneficial bacteria such as bifidobacteria and lactobacilli inhibit the proliferation of detrimental bacteria, and their cell components stimulate the host immune system [75]. Gastrointestinal microflora consist of approximately 1014 colony forming units (cfu)/g of various types of both detrimental and beneficial bacteria, and the numbers and composition vary greatly along the gastrointestinal tract. The balance of the gastrointestinal micro flora influences different aspects of host health such as bowel movement, tympanites flatulence, and the absorption of nutrients. Many factors may upset this balance, including stress, consumption of antibiotics, infection, food poisoning, and the natural ageing process. To redress this balance, the growth and activities of beneficial bacteria may be enhanced by specific ingredients in foods.

SpeceisStrainGlcGalRhaFucNAc GalGlcAGlyReference
Streprococcus thermophilusCNCMI 733121[25]
Lactococcus lactis subsp. cremorisNIZO B89132[30]
NIZO B39232[32]
SBT 0495221[33]
Lactobacillus delbrueckii subsp. bulgaricusOLL 1073R-111.6[34]
NCFB 277212.4[35]
NCFB 2772170.8[35]
Lb. helveticusTN-411[39]
Lb. rhamnosusRW-9595M214[43]
Lb. plantarumEP56311[45]
Lb. pentosusLPS26122[46]
Lb. paracasei34-1311[47]
Lb. kefiranofaciensK111[48]
Lb. sake0-132[49]

Table 2.

Monosaccharide ratio in hetero EPSGlc: glucose, Gal: galactose, Rha: rhamnose, Fuc: fucose, NAc Glu: N-acetyl glucosamine, NAc Gal: N-acetyl galactosamine, GlcA: glucuronic acid.

Various oligosaccharides have been identified as prebiotics, that can increase the number of Bifidobacterium in the host colon. Galacto-oligosaccharides (GOS) and fructo-oligosaccharides (FOS) are considered important prebiotics. Other carbohydrates including gluco-oligosaccharides, isomalto-oligosaccharides, lactulose, mannan-oligosaccharides, and nigero-oligosaccharides are also considered prebiotics. Increased numbers of bifidobacteria and/or lactobacilli in the colon have been shown to have beneficial effects, although the specific mixtures of populations of these genera necessary to provide health-promoting effects has not yet been determined. This is because the beneficial effects are likely to be due to improvement in the balance of coloni micro flora. However, difference do exist in the micro flora among individuals. To function most effectively, prebiotics must be resistant to digestive processes in the stomach and small bowel, so that they can come into contact with the bacteria growing in the large intestine.

The food for specified health use (FOSHU) system was introduced in Japan in 1991. FOSHU refers to foods containing ingredients that provide health benefits and have officially approved physiological effects on the human body. FOSHU is intended to be consumed for the maintenance or promotion of health or for special health uses, for example, to control conditions such as blood pressure or blood cholesterol. To be defined as FOSHU, it is important to assess the safety of the food as well as the effectiveness of health promotion, and this assessment must be approved by the Ministry of Health, Labour and Welfare in Japan. At present (2012), 990 foods are recognized as FOSHU, and of these, 86 provide gastrointestinal health benefit. Foods for balancing gastrointestinal micro flora contain galactosylsucrose, soy oligosaccharides, lactulose, GOS, FOS, isomalto-oligosaccharides, raffinose, xylo-oligosaccharides, mannobiose, and brewer's yeast cell wall as functional ingredients.

4.1. GOS

GOS are well-known type of prebiotic oligosaccharides found in human milk. The concentration of oligosaccharides is 100 times higher in human breast milk than in bovine milk [76]. Many studies have shown that breast-fed infants have intestinal microflora dominated by bifidobacteria. The reason for this phenomenon is thought to be that the oligosaccharides in breast milk, including GOS, can reach the upper gut without being digested where the bifidobacteria can utilize them. At present, GOS is produced by the enzymatic treatment of lactose by β-galactosidase. GOS produced in this manner usually have degrees of polymerization (DP) between 2 and 10. Furthermore, the type of glycosidic linkage is determined by the reaction conditions: final products usually possess β-1,2, β-1,3, or β-1,4 linkages. GOS is given a caloric value of 2 kcal/g in Japan and Europe for food-labelling purpose.

The effect of GOS on defecation has been studied in healthy volunteers. Defecation frequency was significantly increased, and faeces became significantly softer after the subjects drank a beverage containing 5.0 g of GOS, on a daily basis. Therefore, consumption of a beverage containing 5.0 g of GOS can improve defecation in individuals with a tendency for constipation [77]. Ishikawa et al. reported that the number of faecal bifidobacteria increased significantly after subjects consumed 2.5 g of GOS/day for 3 weeks [78]. GOS utilization by enterobacteria was further investigated in vitro. The trisaccharide forms of GOS were utilized by Bifidobacterium, Lactobacillus acidophilus, Lb. reuteri, Bacteroides, Clostridium perfringens, Klebsiella pnumoniae, Enterococcus faecium, and the tetra-saccharide forms were utilized by Bifidobacterium adolescentis, Bifidobacterium breve, Bifidobacterium infantis, and Ent. faecium. These results suggest that a higher DP of GOS enhanced selectivity, and that the tetrasaccharide forms of GOS are specifically utilized by bifidobacteria. Similarly, Bifidobacterium lactis DR10 utilizes trisaccharide and tetra-saccharide forms of GOS, whereas Lb. rhamnosus DR20 prefers disaccharides and monosaccharides [79]. Barboza reported that Bif. breve and Bif. longum subsp. infantis can consume GOS with a DP ranging from 3 to 8 [80]. Furthermore, Bif. longum subsp. infantis preferentially consume GOS with a DP of 4, and Bif. adolescentis utilizes GOS with DP of 3. In addition, the structure of GOS influences its utilization by lactobacilli and bifidobacteria [81]. Trisaccharides of 4ʹ-GOS (β-1,4 linkage) and 6ʹ-GOS (β-1,6 linkage) can be used as the sole carbon source. Almost all lactobacilli and bifidobacteria tested preferred to utilize 4ʹ-GOS, while Lb. acidophilus, Lb. reuteri, and Lb. casei could utilize both 4ʹ- and 6ʹ-GOS. GOS are used to stimulate beneficial bacteria, but can also be utilized by bacteroides and clostridia [82]. GOS selectivity may be enhanced by altering the structure and increasing the DP.

The use of beneficial bacteria or their enzymes in the synthesis of prebiotics may be a good way to produce prebiotics with high specificity. Rabiu reported that five different GOS were produced using β-galactosidase extracted from five different Bifidobacterium species, and that each GOS showed an increased growth rate in producer strains, except for Bif. adolescentis [83]. The utilization of these GOS by faecal bacteria was investigated using commercial GOS as control. The number of Bacteroides was decreased with GOS from bifidobacteria, whereas both GOS extracts and commercial GOS increased the number of bifidobacteria, lactobacilli, and clostridia.

4.2. FOS

FOS is used as a generic term for all β-2,1 linear fructans with a variable DP. Inulin and oligofructose are common forms of FOS that are widely found in nature. Chicory inulin has a DP of 2-60, and the product of its partial enzymatic hydrolysis is oligofructose or FOS with a DP of 2-10.

The effect of FOS intake on intestinal microflora was studied in humans. The number of bifidobacteria in faeces was significantly increased during the FOS intake (1 g/d) period, and a significant increase in stool frequency and a softening effect on stool were observed [84]. FOS increased the level of bifidobacteria in faeces, whereas that of bacteroides, clostridia, and fusobacteria decreased in subjects that were fed FOS (15 g/d) for 15 days [85]. Another study measured the increase in number of Bifidobacterium species in faeces by using real-time PCR [86]. The composition of bifidobacteria in the gut microflora was studied by clone library analysis in ten volunteers. All ten volunteers carried Bif. longum, and nine of these also carried Bif. adolescentis. The consumption of inulin (10 g/d) increased the number of bifidobacteria in faeces with Bif. adolescentis showing the highest increase response among Bifidobacterium species. Rossi et al. reported that only 8 of 55 Bifidobacterium strains fermented inulin in pure cultures, although inulin increased the number of bifidobacteria in faecal culture [87]. They, therefore, suggested that most bifidobacteria were not able to utilize long fructans in the absence of other intestinal bacteria that can hydrolyze fructans, and that fermentation of oligosaccharides in the colon is the result of a complex metabolic sequence carried out by numerous species.

4.3. Selection of high-efficiency prebiotics

It is not clear which oligosaccharides are the most suitable substrates for the selective growth of specific beneficial species or strains. Several research group have suggested useful methods to investigate the potential prebiotic activity of oligosaccharides [88-92]. Potential prebiotic activities were determined on the basis of the changes in the growth of beneficial and undesirable bacteria, such as bifidobacteria, lactobacilli, clostridia, and bacteroides. Such methods can evaluate the ability of specific strains to utilize a particular prebiotic, and a comparison of the prebiotic activities of oligosaccharides by using these methods could help in the choice of prebiotics for improving the gastrointestinal microflora on an individual basis. However, it is important to understand that only a limited group of bacteria can be chosen from the gastrointestinal microflora by using these methods, and that polysaccharides and oligo-saccharides are fermented by numerous species in the gastrointestinal tract.

Oligosaccharides produced by beneficial bacteria or their enzymes may enhance the growth of beneficial bacteria. A novel GOS mixture produced using Bif. longum NCIBM 41171 galactosidases increased the proportion of bifidobacteria in faeces relative to commercial GOS [93]. In the above-described study, oligosaccharides synthesized by the enzymes from Bifidobacterium strains were favored by the producer strains [83]. These studies suggest that the oligosaccharides produced by beneficial bacteria are selectively utilized by the producer strain, because the enzymes required for their degradation are already available. In addition, glycosyltransferases may possess both hydrolytic and transglycosylation activities [94], and glycosidases and glycosyltransferases may coexist in the same strains. Schwab et al. reported the production of novel oligosaccharides [95]. Hetero-oligosaccharides were produced from lactose, mannose, fucose, and N-acetylglucosamine by using crude cell extracts and whole cells of LAB and bifidobacteria. These hetero-oligosaccharides contained mannose, fucose, and N-acetylglucosamine, and could be digested by LAB strains. The prebiotic activities of these oligosaccharides were not investigated; however, a similar approach using probiotic and intestinal beneficial bacteria may lead to the production of highly selective prebiotics.

The dietary fiber, arabinoxylan is the predominant hemicellulose from cereals and exhibits prebiotic activity [96]. The addition of water-unextractable arabinoxylans increased the population of bifidobacteria and bacteroides in a medium inoculated with faecal slurry. Polysaccharides are not usually utilized by microorganisms. Remarkably, however, Bifidobacterium bifidum DSM20456 can utilize the EPS produced by Pediococcus pentosaceus, Lb. plantarum, Weissella cibaria, and Weissella confusa, and some growth is observed in cas of Bif. longum, Bif. adolescentis, and Lb. acidophilus [97]. For EPS production by LAB, reduced yields were frequently observed after the maximal level had been reached, which might be caused by the enzymes produced by the bacteria [98]. Tsuda and Miyamoto investigated the prebiotic activity of EPS produced by Lb. plantarum 301102S [52], a mutant strain derived from Lb. plantarum 301102. Oral administration of the parental strain 301102 showed the survivability and proliferation in porcine gastrointestinal tract [99]. The potential prebiotic activities of EPS, GOS, and inulin were measured in 37 LAB strains, and the activity scores of EPS in the strains 301102 and 301102S were highest. This suggests that the EPS produced by the mutant strain is utilized by the same strain 301102S and the parental strain, and that the parental strain has enzymes that can degrade the EPS.


5. Conclusion

Poly- and hetero-oligosaccharides produced by LAB may be potential prebiotics. Studies on the production of polysaccharides and oligosaccharides by enzymes in beneficial microorganisms may lead to the production of highly selective prebiotics, although in vitro evaluation may be difficult because of degradation and utilization of polysaccharides by various microorganisms in the gastrointestinal tract. Administration of synbiotic food containing a combination of a probiotic bacterial strain and the prebiotic sugar produced by that strain could be effective in improving human health.


  1. 1. ChabotS.YuH. LeseleucL.CloutierD.Van CalsterenM. R.RoyD.LacroixM.OthD.2001Exopolysaccharides from Lactobacillus rhamnosus RW-9595M stimulate TNF, IL-6 and IL-12 in human and mouse cultured immunocompetent cells, and IFN-γ in mouse splenocytes. Lait 81683698
  2. 2. KitazawaH.HarataT.UemuraJ.SaitoT.KanekoT.ItohT.1998Phosphate group requirement for mitogenic activation of lymphocytes by an extracellular phosphopolysaccharide from Lactobacillus delbrueckii ssp. bulgaricus. Int. J. Food Microbiol. 40169175
  3. 3. SreekumarO.HosonoA.1998The antimutagenic properties of a polysaccharide produced by Bifidobacterium longum and its cultured milk against some heterocyclic amines. Can. J. Microbiol. 4410291036
  4. 4. TsudaH.HaraK.MiyamotoT.2008Binding of mutagens to exopolysaccharide produced by Lactobacillus plantarum mutant strain 301102S. J. Dairy Sci. 9129602966
  5. 5. GanzleM.MichaelG.SchwabC.2005Exopolysaccharide production by intestinal lactobacilli. In: Tannock G. W, editors. Probiotics & Prebiotics: Scientific Aspects. Norfolk: Caister Academic Press. 8396
  6. 6. KraljS.van Geel-SchuttenG. H.DondorffM. M. G.KirsanovsS.van der MaarelM.J. E. C., Dijkhuizen L (2004Glucan synthesis in the genus Lactobacillus: isolation and characterization of glucansucrase genes, enzymes and glucan products from six different strains. Microbiology 15036813690
  7. 7. van HijumS. A. F. T.SzalowskaE.van der MaarelM.J. E. C., Dijkhuizen L (2004Biochemical and molecular characterization of a levansucrase from Lactobacillus reuteri. Microbiology 150621630
  8. 8. de VinF.VaningelgemF.DegeestB.2001Recent developments in the biosynthesis and applications of heteropolysaccharides from lactic acid bacteria. Int. Dairy J. 11687707
  9. 9. Ruas-MadiedoP.HugenholtzJ.ZoonP.2002An overview of the functionality of exopolysaccharides produced by lactic acid bacteria. Int. Dairy J. 12163171
  10. 10. RuijssenaarsH.StingeleF.HartmansS.2000Biodegradability of food-associated extracellular polysaccharides. Curr. Microbiol. 40194199
  11. 11. KorakliM.GanzleM. G.VogelR. F.2002Metabolism by bifidobacteria and lactic acid bacteria of polysaccharides from wheat and rye, and exopolysaccharides produced by Lactobacillus sanfranciscensis. J. Appl. Microbiol. 92958965
  12. 12. FullerR.1989Probiotics in man and animals. J. Appl. Bacteriol. 66365378
  13. 13. FAO/WHO2002Guidelines for the evaluation of probiotics in food. Report of a joint FAO/WHO working group on drafting guidelines for the evaluation of probiotics in food. London, Ontario, Canada.
  14. 14. Gibson G.R, Roberfroid M.B1995Dietary modulation of the human colonic microbiota: Introducing the concept of prebiotics. J. Nutr. 12514011412
  15. 15. RastallR.2003Enhancing the functionality of prebiotics and probiotics. In: Mattila-Sandholm T, Saarela M, editors. Functional Dairy Products. Florida: CRC Press LLC. 301315
  16. 16. MayoB.Aleksandrzak-PiekarczykT.FernandezM.KowalczykM.Alvarez-MartinP.BardowskiJ.2010Updates in the metabolism of lactic acid bacteria. In: Mozzi F, Raya R. R, Vignolo G. M, editors. Biotechnology of Lactic Acid Bacteria. Iowa: Blackwell Publishing. 333
  17. 17. CerningJ.1990Exocellular polysaccharides produced by lactic acid bacteria. FEMS Microbiol. Lett. 87113130
  18. 18. PedersonC. S.AlburyM.1955Variation among the heterofermentative lactic acid bacteria. J. Bacteriol. 70702708
  19. 19. DellaglioF.DicksL. M. T.TorrianiS.1995The genus Leuconostoc. In: Wood B. J. B., Holzapfel W. H, editors. The genera of lactic acid bacteria. Glasgow: Blackie Academic and Professional. 235278
  20. 20. MinerviniF.AngelisM. D.SuricoR. F.CagnoR. D.GanzleM.GobbettiM.2010Highly efficient synthesis of exopolysaccharides by Lactobacillus curvatus DPPMA10 during growth in hydrolyzed wheat flour agar. Int. J. Food Microbiol. 141130135
  21. 21. Cote G. L, Robyt F. J1982Isolation and partial characterization of an extracellular glucansucrase from Leuconostoc mesenteroides NRRl B-1355 that synthesizes an alternating (1→6), (1→3)-α-D-glucan. Carbohydr. Res. 1015774
  22. 22. UchidaK.1996Nyuusannkinn no kouzou to kinntaiseibunn. In: Nyuusannkinn kennkyuu syuudannkai, editors. Nyuusannkinn no kagaku to gijutsu. Tokyo: Gakkai Syuppann Center. 5988In Japanese)
  23. 23. Van Geel-SchuttenG. H.FaberE. J.SmitE.BontingK.SmithM. R.TenBrink. B.KamerlingJ. P.VliegenthartJ. F. G.DijkhuizenL.1999Biochemical and structural characterization of the glucan and fructan exopolysaccharides synthesized by the Lactobacillus reuteri wild-type strain and by mutant strains. Appl. Environ. Microbiol. 6530083014
  24. 24. De VosP.GarrityG. M.JonesD.KriegN. R.LudwigW.RaineyF. A.SchleiferK. H.WhitmanW. B.editors2009Bergey’s Manual of Systematic Bacteriology Second Edition Volume Three. New York: Springer.
  25. 25. DocoT.WieruszeskiJ. M.FournetB.1990Structure of an exocellular polysaccharide produced by Streptococcus thermophilus. Carbohydr. Res. 198313321
  26. 26. LemoineJ.ChiratF.WieruszeskiJ. M.StreckerG.FavreN.NeeserJ. R.1997Structural characterization of the exocellular polysaccharides produced by Streptococcus thermophilus SFi39 and SFi12. Appl. Environ. Microbiol. 633512
  27. 27. VuystL.2000Correlation of activities of the enzymes a-phosphoglucomutase, UDP-galactose 4-epimerase, and UDP-glucose pyrophosphorylase with exopolysaccharide biosynthesis by Streptococcus thermophilus LY03. Appl. Environ. Microbiol. 6635193527
  28. 28. BubbW. A.UrashimaT.FujiwaraR.ShinnaiT.ArigaH.1997Structural characterization of the exocellular polysaccharide produced by Streptococcus thermophilus OR901. Carbohydr. Res. 3014150
  29. 29. LowD.AhlgrenJ. A.HorneD.Mc MahonD. J.ObergC. J.BroadbentJ. R.1998Role of Streptococcus thermophilus MR-1C capsular exopolysaccharide in cheese moisture retention. Appl. Environ. Microbiol. 6421472151
  30. 30. Van CasterenW. H. WaaedP.DijkemaC.ScholsH. A.VoragenA. G. J.2000Structural characterization and enzymic modification of the exopolysaccharide produced by Lactococcus lactis subsp. cremoris B891. Carbohydr. Res. 327411422
  31. 31. Knoshaug E. P, Ahlgren J. A, Trempy J. E2007Exopolysaccharide expression in Lactococcus lactis subsp. cremoris Ropy352: Evidence for novel gene organization. Appl. Environ. Microbiol. 73897905
  32. 32. van CasterenW. H. M.DijkemaC.ScholsH. A.BeldmanG.VoragenA. G. J.2000Structural characterization and enzymic modification of the exopolysaccharide produced by Lactococcus lactis subsp. cremoris B39. Carbohydr. Res. 324170181
  33. 33. NakajimaH.HirotaT.TobaT.ItohT.AdachiS.1992Structure of the extracellular polysaccharide from slime-forming Lactococcus lactis subsp. cremoris SBT 0495. Carbohydr. Res. 224245253
  34. 34. KitazawaH.IshiiY.UemuraJ.KawaiY.SaitoT.KanekoT.NodaK.ItohT.2000Augmentation of macrophage functions by an extracellular phosphopolysaccharide from Lactobacillus delbrueckii ssp. bulgaricus. Food Microbiol. 17109118
  35. 35. GrobbenG. J.SmithM. BontJ. A. M.1996Influence of fructose and glucose on the production of exopolysaccharides and the activities of enzymes involved in the sugar metabolism and the synthesis of sugar nucleotides in Lactobacillus delbrueckii subsp. bulgaricus NCFB 2772. Appl. Microbiol. Biotechnol. 46279284
  36. 36. PetryS.FurlanS.WaghorneE.SaulnierL.CerningJ.maguinE.2003Comparison of the thickening properties of four Lactobacillus delbrueckii subsp. bulgaricus strains and physicochemical characterization of their exopolysaccharides. FEMS Microbiol. Lett. 221285291
  37. 37. HardingL. P.MarshallV. M.ElvinM.GuY.LawsA. P.2003Structural characterization of a perdeuteriomethylated exopolysaccharide by NMR spectroscopy: characterization of the novel exopolysaccharide produced by Lactobacillus delbrueckii subsp. bulgaricus EU23. Carbohydr. Res. 3386167
  38. 38. GruterM.LeeflangB. R.KuiperJ.KamerlingJ. P.VliegenthartF. G.1993Structural characterization of the exopolysaccharide produced by Lactobacillus delbrueckii subspecies bulgaricus rr grown in skimmed milk. Carbohydr. Res. 239209226
  39. 39. YamamotoT.NunomeT.YamauchiR.KatoK.SoneY.1995Structure of an exocellular polysaccharide of Lactobacillus helveticus TN-4, a spontaneous mutant strain of Lactobacillus helveticus TY1-2. Carbohydr. Res. 275319332
  40. 40. RobijnG. W.ThomasJ. R.HaasH.van denBerg. D.J. R, Haas H, van den Berg D. J. C, Kamerling J. P, Vliengenthart J. F. G (1995The structure of the exopolysaccharide produced by Lactobacillus helveticus 766. Carbohydr. Res. 276137154
  41. 41. StaafM.WidmalmG.YangZ.HuttunenE.1996Structural elucidation of an extracellular polysaccharide produced by Lactobacillus helveticus. Carbohydr. Res. 291155164
  42. 42. StaafM.YangZ.HuttunenE.WidmalmG.2000Structural elucidation of the viscous exopolysaccharide produced by Lactobacillus helveticus Lb161. Carbohydr. Res. 326113119
  43. 43. Van CalsterenM. R.Pau-RoblotC.BeginA.RoyD.2002Structure determination of the exopolysaccharide produced by Lactobacillus rhamnosus strains RW-9595M and R. Biochem. J. 363717
  44. 44. LandersjoC.YangZ.HuttunenE.WidmalmG.2002Structural studies of the exopolysaccharide produced by Lactobacillus rhamnosus strain GG (ATCC 53103). Biomacromolecules 3880884
  45. 45. TallonR.BressollierP.UrdaciM. C.2003Isolation and characterization of two exopolysaccharides produced by Lactobaciilus plantarum EP56. Res. Microbiol. 154705712
  46. 46. Rodriguez-CarvajalM. A.SanchezJ. I.CampeloA. B.MartinezB.RodriguezA.Gil-SerranoA. M.2006Structure of the high-molecular weight exopolysaccharide isolated from Lactobacillus pentosus LPS26. Carbohydr. Res. 34330663070
  47. 47. RobijnG. W.WienkH. L. J.van denBerg. D.J. C, Haas H, Kamerling J. P, Vliengenthart J. F. G (1996Structural studies of the exopolysaccharide produced by Lactobacillus paracasei 34-1. Carbohydr. Res. 285129139
  48. 48. MukaiT.TobaT.ItohT.AdachiS.1990Structural investigation of the capsular polysaccharide from Lactobacillus kefiranofaciens K1. Carbohydr. Res. 204227232
  49. 49. van denBerg. D. J. C.RobijnG. W.JanssenA. C.GiuseppinM. L. F.VreekerR.KamerlingJ. P.VliegenthartJ. F. G.LedeboerA. M.VerripsC. T.1995Production of a novel extracellular polysaccharide by Lactobacillus sake 0-1 and characterization of the polysaccharide. Appl. Environ. Microbiol. 6128402844
  50. 50. LeoF.HashidaS.KumagaiD.UchidaK.MotoshimaH.AraiI.AsakumaS.FukudaK.UrashimaT.2007Studies on a neutral exopolysaccharide of Lactobacillus fermentum TDS030603. J. Appl. Glycosci. 54223229
  51. 51. FigueroaC.DavilaA. M.PourquieJ.1995Lactic acid bacteria of the sour cassava starch fermentation. Lett. Appl. Microbiol. 21126130
  52. 52. CerningJ.1995Production of exopolysaccharides by lactic acid bacteria and dairy propionibacteria. Lait, 75463472
  53. 53. TsudaH.MiyamotoT.2010Production of exopolysaccharide by Lactobacillus plantarum and the prebiotic activity of the exopolysaccharide. Food Sci. Technol. Res. 168792
  54. 54. MacedoM. G.LacroixC.GardnerN. J.ChampagneC. P.2002Effect of medium supplementation on exopolysaccharide production by Lacctobacillus rhamnosus RW-9595M in whey permeate. Int. Dairy J. 12419426
  55. 55. MaedaH.ZhuX.SuzukiS.SuzukiK.KitamuraS.2004Structural characterization and biological activities of an exopolysaccharide kefiran produced by Lactobacillus kefiranofaciens WT-2BT. J. Agric. Food Chem. 5255335538
  56. 56. de VuystL.DegeestB.1999Heteropolysaccharides from lactic acid bacteria. FEMS Microbiol. Rev. 23153177
  57. 57. Garcia-Garibay M, Marshall V. M. E1991Polymer production by Lactobacillus delbrueckii ssp. bulgaricus. J. Appl. Bacteriol. 70325328
  58. 58. Kimmel S. A, Roberts R. F, Ziegler G. R1998Optimization of exopolysaccharide production by Lactobacillus delbrueckii subsp. bulgaricus RR grown in a semidefined medium. Appl. Environ. Microbiol. 64659664
  59. 59. GamarL.BlondeauK.SimonetJ. M.1997Physiological approach to extracellular polysaccharide production by Lactobacillus rhamnosus strain C83. J. Appl. Microbiol. 83281287
  60. 60. GrobbenG. J.Chin-JoeI.KitzenV. A.BoelsI. C.BoerF.SikkemaJ.SmithM. BontJ. A. M.1998Enhancement of exopolysaccharide production by Lactobacillus delbrueckii subsp. bulgaricus NCFB 2772 with a simplified defined medium. Appl. Environ. Microbiol. 6413331337
  61. 61. CerningJ.RenardC. M. G. C.ThibaultJ. F.BouillanneC.LandonM.DesmazeaudM.TopisirovicL.1994Carbon source requirements for exopolysaccharide production by Lactobacillus casei CG11 and partial structure analysis of the polymer. Appl. Environ. Microbiol. 6039143919
  62. 62. de VuystL.VandervekenF.Van de VenS.DegeestB.1998Production by and isolation of exopolysaccharides from Streptococcus thermophilus grown in a milk medium and evidence for their growth-associated biosynthesis. J. Appl. Microbiol. 8410591068
  63. 63. KorakliM.PavlovicM.GanzleM. G.VogelR. F.2003Exopolysaccharide and kestose production by Lactobacillus sanfranciscensis LTH2590. Appl. Environ. Microbiol. 6920732079
  64. 64. KraljS.van Geel-SchuttenG. H.van der MaarelM.H. E. C, Dijkhuizen L (2004Biochemical and molecular characterization of Lactobacillus reuteri 121 reuteransucrase. Microbiology 15020992112
  65. 65. NeubauerH.BaucheA.MolletB.2003Molecular characterization and expression analysis of the dextransucrase DsrD of Leuconostoc mesenteroides Lcc4 in homologous and heterologous Lactococcus lactis cultures. Microbiology 149973982
  66. 66. DabourN.La PointeG.2005Identification and molecular characterization of the chromosomal exopolysaccharide biosynthesis gene cluster from Lactococcus lactis subsp. cremoris SMQ-461. Appl. Environ. Microbiol. 7174147425
  67. 67. DanT.FukudaK.Sugai-BannaiM.TakakuwaN.MotoshimaH.UrashimaT.2009Characterization and expression analysis of the exopolysaccharide gene cluster in Lactobacillus fermentum TDS030603. Biosci. Biotechnol. Biochem. 7326562664
  68. 68. MozziF.Savoy deGiori. G.Font deValdez. G.2003UDP-galactose 4-epimerase: a key enzyme in exopolysaccharide formation by Lactobacillus casei CRL 87 in controlled pH batch cultures. J. Appl. Microbiol. 94175183
  69. 69. al.2003Complete genome sequence of Lactobacillus plantarum WCFS1. Proc. Natl. Acad. Sci. USA. 10019901995
  70. 70. MoronaJ. K.PatonJ. C.MillerD. C.MoronaR.2000Tyrosine phosphorylation of CpsD negatively regulates capsular polysaccharide biosynthesis in Streptococcus pneumonia. Mol. Microbiol. 3514311442
  71. 71. NagaiT.MakinoS.IkegamiS.ItohH.YamadaH.2012Effects of oral administration of yogurt fermented with Lactobacillus delbrueckii ssp. bulgaricus OLL1073R-1 and its exopolysaccharides against influenza virus infection in mice. Int. Immunopharmacolgy 1122462250
  72. 72. KodaliV. P.SenR.2008Antioxidant and free radical scavenging activities of an exopolysaccharide from a probiotic bacterium. Biotechnol. J. 3245251
  73. 73. KleerebezemM.van KranenburgR.TuinierR.BoelsI. VosW. M.1999Exopolysaccharides produced by Lactococcus lactis: from genetic engineering to improved rheological properties? Ant. van Leeuwenhoek 76357365
  74. 74. O’ConnorE. B.BarrettE.FitzgeraldG.HillC.StantonC.RossR. P.2005Production of vitamins, exopolysaccharides and bacteriocins by probiotic bacteria. In: Tamime A. Y, editors. Probiotic Dairy Products. Oxford: Blackewell Publishing Ltd. 167194
  75. 75. MitsuokaT.1992Intestinal flora and aging. Nutr. Rev. 50438446
  76. 76. KunzC.RudloffS.BaierW.KeinN.StrobelS.2000Oligosaccharides in human milk: structure, functional, and metabolic aspects. Ann. Rev. Nutr. 20699722
  77. 77. DeguchiY.MatsumotoK.ItoA.WatanukiM.1997Effects of β 1-4 galactooligosaccharides administration on defecation of healthy volunteers with constipation tendency. Eiyougakuzasshi 551322in Japanese)
  78. 78. IshikawaF.TakayamaH.MatsumotoK.ItoM.ChonanO.DeguchiY.Kikuchi-HayakawaH.WatanukiM.1995Effects of β 1-4 linked galactooligosaccharides on human fecal microflora. Bifizusu 9518in Japanese)
  79. 79. Gopal P. K, Sullivan P. A, Smart J. B2001Utilisation of galacto-oligosaccharides as selective substrates for growth by lactic acid bacteria including Bifidobacterium lactis DR10 and Lactobacillus rhamnosus DR20. Int. Dairy J. 111925
  80. 80. BarbozaM.SelaD. A.PirimC.LoCascio. R. G.FreemanS. L.GermanJ. B.MillsD. A.LebrillaC. B.2009Glycoprofiling bifidobacterial consumption of galacto-oligosaccharides by mass spectrometry reveals strain-specific, preferential consumption of glycans. Appl. Environ. Microbiol. 7573197325
  81. 81. Cardelle-CobasA.CorzoN.OlanoA.PelaezC.RequenaT.AvilaM.2011Galactooligosaccharides derived from lactose and lactulose: Influence of structure on Lactobacillus, Streptococcus and Bifidobacterium growth. Int. J. Food Microbiol. 1498187
  82. 82. MacfarlaneG. T.SteedH.MacfarlaneS.2008Bacterial metabolism and health-related effects of galacto-oligosaccharides and other prebiotics. J. Appl. Microbiol. 104305344
  83. 83. Rabiu B. A, Jay A. J, Gibson G. R, Rastall R. A2001Synthesis and fermentation properties of novel galacto-oligosaccharides by β-galactosidases from Bifidobacterium species. Appl. Environ. Microbiol. 6725262530
  84. 84. TokunagaT.NakadaY.TashiroY.HirayamaM.HidakaH.1993Effects of fructooligosaccharides intake on the intestinal microflora and defecation in healthy volunteers. Bifizusu 6143150in Japanese)
  85. 85. GibsonG. R.BeattyE. R.WangX.CummingsJ. H.1995Selective stimulation of bifidobacteria in the human colon by oligofructose and inulin. Gastroenterology 108975982
  86. 86. Ramirez-FariasC.SlezakK.FullerZ.DuncanA.HoltropG.LouisP.2009Effect of inulin on the human gut microbiota: stimulation of Bifidobacterium adolescentis and Faecalibacterium prausnitzii. Br. J. Nutr. 101541550
  87. 87. RossiM.CorradiniC.AmarettiA.NicoliniM.PompeiA.ZanoniS.MatteuzziD.2005Fermentation of fructooligosaccharides and inulin by bifidobacteria: a comparative study of pure and fecal cultures. Appl. Environ. Microbiol. 7161506158
  88. 88. Olano-MartinE.GibsonG. R.RastallR. A.2002Comparison of the in vitro bifidogenic properties of pectins and pectic-oligosaccharides. J. Appl. Microbiol. 93505511
  89. 89. PalframanR.GibsonG. R.RastallR. A.2003Development of a quantitative tool for the comparison of the prebiotic effect of dietary oligosaccharides. Lett. Appl. Microbiol. 37281284
  90. 90. VulevicJ.RastallR. A.GibsonG. R.2004Developing a quantitative approach for determining the in vitro prebiotic potential of dietary oligosaccharides. FEMS Microbiol. Lett. 236153159
  91. 91. Sanz M. L, Gibson G. R, Rastall R. A2005Influence of disaccharide structure on prebiotic selectivity in vitro. J. Agric. Food Chem. 5351925199
  92. 92. HuebnerJ.WehlingR. L.HutkinsR. W.2007Functional activity of commercial prebiotics. Int. Dairy J. 17770775
  93. 93. DepeintF.TzortzisG.VulevicJ.I’AnsonK.GibsonG. R.2008Prebiotic evaluation of a novel galactooligosaccharide mixture produced by the enzymatic activity of Bifidobacterium bifidum NCIMB 41171, in healthy humans: a randomized, double-blind, crossover, placebo-controlled intervention study. Am. J. Clin. Nutr. 87785791
  94. 94. van denBroek. L. A. M.HinzS. W. A.BeldmanG.VinckenJ. P.VoragenA. G. J.2008Bifidobacterium carbohydrases- their role in breakdown and synthesis of (potential) prebiotics. Mol. Nutr. Food Res. 52146163
  95. 95. SchwabC.LeeV.SorensenK. I.GanzleM. G.2011Production of galactooligosaccharides and heterooligosaccharides with disrupted cell extracts and whole cells of lactic acid bacteria and bifidobacteria. Int. Dairy J. 21748754
  96. 96. VardakouM.PalopC. M.ChristakopoulosP.FauldsC. B.GassonM. A.NarbadA.2008Evaluation of the prebiotic properties of wheat arabinoxylan fractions and induction of hydrolase activity in gut microflora. Int. J. Food Microbiol. 123166170
  97. 97. HongpattarakereT.CherntongN.WichienchotS.KolidaS.RastallR. A.2012In vitro prebiotic evaluation of exopolysaccharides produced by marine isolated lactic acid bacteria. Carbohydr. Polym. 87846852
  98. 98. PhamP. L.DupontI.RoyD.LapointeG.CerningJ.2000Production of exopolysaccharide by Lactobacillus rhamnosus R and analysis of its enzymatic degradation during prolonged fermentation. Appl. Environ. Microbiol. 6623022310
  99. 99. TsudaH.HaraK.MiyamotoT.2008Survival and colonization of orally administered Lactobacillus plantarum 301102 in porcine gastrointestinal tract. Anim. Sci. J. 7274278

Written By

Tsuda Harutoshi

Submitted: November 20th, 2011 Published: January 30th, 2013