HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS		QUICK SEARCH:   [advanced] Author: Keyword(s): Year:  Vol:  Page:

This Article Abstract Full Text (PDF) Interpretive Summary Alert me when this article is cited Alert me if a correction is posted Services Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Citing Articles Citing Articles via HighWire Citing Articles via Google Scholar Google Scholar Articles by Ruas-Madiedo, P. Articles by de los Reyes-Gavilán, C. G. Search for Related Content PubMed PubMed Citation Articles by Ruas-Madiedo, P. Articles by de los Reyes-Gavilán, C. G.

Invited Review: Methods for the Screening, Isolation, and Characterization of Exopolysaccharides Produced by Lactic Acid Bacteria

Instituto de Productos Lácteos de Asturias, CSIC, Carretera de Infiesto s/n, 33300 Villaviciosa, Asturias, Spain

Corresponding author: Patricia Ruas-Madiedo; e-mail: ruas-madiedo{at}ipla.csic.es.

	ABSTRACT

TOP ABSTRACT INTRODUCTION DETECTION OF EPS-PRODUCING... SCREENING OF EPS-PRODUCING LAB... EPS ISOLATION METHODS QUANTIFICATION OF EPS YIELD PRIMARY AND 3-D STRUCTURE... EPS MOLECULAR PARAMETERS RELATED... ACKNOWLEDGEMENTS REFERENCES

 
The ability to produce exopolysaccharides (EPS) is widespread among lactic acid bacteria (LAB), although the physiological role of these molecules has not been clearly established yet. Some EPS confer on LAB a "ropy" character that can be detected in cultures that form long strands when extended with an inoculation loop. When EPS are produced in situ during milk fermentation they can act as natural biothickeners, giving the product a suitable consistency, improving viscosity, and reducing syneresis. In addition, some of these EPS may have beneficial effects on human health. The increasing demand by consumers of novel dairy products requires a better understanding of the effect of EPS on existing products and, at the same time, the search for new EPS-producing strains with desirable properties. The use of genetically modified organisms capable of producing high levels of EPS or newly designed biopolymers is still very limited. Therefore, exploration of the biodiversity of wild LAB strains from natural ecological environments is currently the most suitable approach to search for the desired EPS-phenotype. The screening of ropy strains and the isolation and characterization of EPS responsible for this characteristic have led to the application over the past years of a wide variety of techniques. This review summarizes the available information on methods and procedures used for research on this topic. The information provided deals with methods for screening of EPS-producing LAB, detection of the ropy phenotype, and the physicochemical and structural characterization of these molecules, including parameters related to their viscosifying properties. To our knowledge, this is the first compilation of methods available for the study of EPS produced by LAB.

Key Words: exopolysaccharide • screening • isolation • characterization

Abbreviation key: CSLM = confocal scanning laser microscopy, ESM = EPS selection media, EPS = exopo-lysaccharides, GCMS = gas chromatography/mass spectrometry, HePS = heteropolysaccharides, HoPS = homopolysaccharides, HPAEC-PAD = high-performance anion-exchange chromatography pulse ampero-metric detection, LAB = lactic acid bacteria, MRS = De Man, Rogosa, Sharpe, NIRS = near infrared spectroscopy, Rg = radius of gyration, RI = refractive index.

	INTRODUCTION

TOP ABSTRACT INTRODUCTION DETECTION OF EPS-PRODUCING... SCREENING OF EPS-PRODUCING LAB... EPS ISOLATION METHODS QUANTIFICATION OF EPS YIELD PRIMARY AND 3-D STRUCTURE... EPS MOLECULAR PARAMETERS RELATED... ACKNOWLEDGEMENTS REFERENCES

 
The ability to produce polysaccharides is widely spread among bacteria. Microorganisms can synthesize storage polysaccharides such as glycogen that are located in the cytoplasm, cell wall structural polysaccharides such as peptidoglycan and lipoteichoic acids of gram-positive bacteria, and the lipopolysaccharides anchored in the outer membrane of gram-negative bacteria (Figure 1). Moreover, some bacteria can secrete polysaccharide layers on their surface, which together with a few glycoproteins, are grouped within the general term "glycocalyx". These exocellular polymers comprise the capsular polysaccharides, which form a cohesive layer or capsule covalently linked to the cell surface, and the exopolysaccharides (EPS), which form a slime layer loosely attached to the cell surface or secreted into the environment (Madigan et al., 1997). Although it is generally recognized that exocellular polysaccharides are not used as energy and carbon sources by the producer microorganism, the physiological role of these molecules has yet to be established.

View larger version (22K): [in this window] [in a new window]   Figure 1. Cell location of polysaccharides produced by gram-positive and gram-negative bacteria. CPS = Capsular polysaccharides (capsule), EPS = exopolysaccharides (slime layer).

	DETECTION OF EPS-PRODUCING PHENOTYPES

TOP ABSTRACT INTRODUCTION DETECTION OF EPS-PRODUCING... SCREENING OF EPS-PRODUCING LAB... EPS ISOLATION METHODS QUANTIFICATION OF EPS YIELD PRIMARY AND 3-D STRUCTURE... EPS MOLECULAR PARAMETERS RELATED... ACKNOWLEDGEMENTS REFERENCES

 
The nomenclature used to describe the different EPS-producing phenotypes of LAB is confusing, and terms such as "ropy", "mucoid", and "slime" have been indistinctly used. However, not all mucoid or slime-producing strains are ropy. The mucoid colonies have a glistening and slimy appearance on agar plates, but are not able to produce strands when extended with an inoculation loop, whereas the ropy colonies form a long filament by this method (Vescovo et al., 1989; Dierksen et al., 1997; Figure 2). The ropy characteristic can be detected in liquid cultures of EPS-producing bacteria that show high resistance to flow through serological pipettes, and form viscous strands during free fall from the pipette tip (Vedamuthu and Neville, 1986). In sensory evaluations on fermented milks, ropy strains confer smoother consistency, higher viscosity, and lower susceptibility to syneresis than nonropy strains (Wacher-Rodarte et al., 1993; Rawson and Marshall, 1997). Some LAB (e.g., Lactobacillus casei CG11 and Lactococcus lactis spp. cremoris Ropy352) can express both ropy and mucoid phenotypes depending on the culturing conditions (Cerning et al., 1994; Dierksen et al., 1997). In Lc. lactis spp. cremoris Ropy352, it has been shown that this ability is due to the production of 2 EPS with different chemical compositions (Knoshaug et al., 2000).

View larger version (84K): [in this window] [in a new window]   Figure 2. Macroscopic appearance of the "ropy" strand formed by the cellular mass of a commercial EPS-producing LAB strain growing on the surface of de Man, Rogosa, and Sharpe (MRS) agar plates.

Capsule formation can be found among nonropy and ropy strains. Strep. thermophilus OR901, a strain isolated from commercial yogurt and able to form ropy strands from the cell mass (Ariga et al., 1992), appeared encapsulated using light microscopy with Indian ink staining. This was due to the production of 2 EPS with the same monosaccharide composition and branching linkage, but with different molar mass. The EPS with the higher molar mass appeared in the supernatant of Strep. thermophilus OR901 cultures and was assumed to affect the physical (texture and viscosity) properties of yogurt. The EPS with the lower molar mass was produced in smaller amounts and was thought to be located in the capsule, given that it was only released from the cell surface after sonication. Using confocal scanning laser microscopy (CSLM), Hassan et al. (1995) reported the occurrence of Strep. thermophilus and Lb. delbrueckii spp. bulgaricus capsular EPS-producing strains, with one of these (Lb. delbrueckii spp. bulgaricus RR) also being ropy. The authors detected differences in the aggregation pattern of caseins during milk fermentation between ropy and nonropy capsule-forming Lb. delbrueckii spp. bulgaricus cultures, which also affected the rheological properties of yogurts (Hassan et al., 2002a). The CSLM technique was used to visualize the microstructure of milks fermented with different EPS-producing and nonproducing Lc. lactis spp. cremoris strains. In this case, 2 differential dyes (rhodamine B and acridine orange) were used to distinguish the microorganisms (stained in green) from the casein network (stained in yellow) (Ruas-Madiedo and Zoon, 2003). Recently, a CSLM method was developed for direct visualization of EPS in the fully hydrated dairy product (Hassan et al., 2002). This method involves the staining of EPS with concanavalin A 488 or with wheat germ agglutinin labeled with Alexa fluor 488, and was used to detect EPS produced by several Lc. lactis, Strep. thermophilus, and Lb. delbrueckii spp. bulgaricus strains within the protein network pores of fermented milks. The same staining method was successfully applied for the detection of EPS in Feta cheese made with the EPS-producing strain Strep. thermophilus 3855 (Hassan et al., 2002b.). Finally, the microstructure of cheese and fermented milks containing EPS-producing strains has been recently studied using cryoscanning electron microscopy (Hassan et al., 2003, 2004).

	SCREENING OF EPS-PRODUCING LAB STRAINS

TOP ABSTRACT INTRODUCTION DETECTION OF EPS-PRODUCING... SCREENING OF EPS-PRODUCING LAB... EPS ISOLATION METHODS QUANTIFICATION OF EPS YIELD PRIMARY AND 3-D STRUCTURE... EPS MOLECULAR PARAMETERS RELATED... ACKNOWLEDGEMENTS REFERENCES

 
Today, the increasing demand by consumers for novel dairy products motivates the food industry to better understand the effects of EPS on existing products and to search for new EPS-producing strains with different properties. The isolation of new LAB strains from different dairy and nondairy natural environments makes necessary the use of specific methods for detecting the desired phenotype, i.e., the production of ropy EPS. In the past decade, few articles reporting the screening of EPS-producing LAB from traditional fermented foods have been published. The liquid EPS selection medium (ESM; 90 g of skim milk, 3.5 g of yeast extract, 3.5 g of peptone, and 10 g of glucose per liter) was used by Van den Berg et al. (1993) for selection of EPS-producing LAB from sour dough, sausages, table olives, cheeses, and other dairy products. After incubation at 30°C for 24 h on ESM, the "ropiness" of the culture was determined by its resistance to flow through graduated pipettes according to the method described by Vedamuthu and Neville (1986). Results obtained showed that only 30 out of 607 LAB strains tested (4.9%) were EPS producers. Using a slightly modified ESM containing 50 g/L glucose, Ludbrook et al. (1997) isolated 11 LAB strains from nondairy fermented foods which were able to produce EPS. Four EPS-positive LAB strains from 25 isolates (16%) were found in Nigerian fermented foods using the same modified ESM (Sanni et al., 2002). Furthermore, the use of several carbohydrates (used separately) for the screening, would improve the detection of EPS-producing strains. In that way, Van Geel-Schutten et al. (1998) screened several Lactobacillus strains of different origins (fermented food, gastrointestinal tract of LAB animals, and human dental plaque) for EPS production in de Man, Rogosa, and Sharpe (MRS) medium supplemented with high concentrations (100 g/L) of different sugars: glucose, fructose, maltose, raffinose, sucrose, galactose, or lactose. One hundred eighty-two strains were tested in these media and, after 3 d of incubation at 37°C, the cultures were precipitated with cold ethanol. The resulting pellets were dried at 55°C and the EPS fraction was determined by measuring the total carbohydrate content with the phenol/sulfuric acid method (Dubois et al., 1956). Sixty strains produced EPS and 17 of them rendered more than 100 mg/L, with the sucrose medium being the best for detecting the EPS phenotype. The authors attributed the higher percentage of positive isolates (33%) to the high content of sugar used in the media. Different sugars were used for the screening of EPS-producing LAB from traditional Thai fermented foods (Smitinont et al., 1999). In this case, MRS agar plates containing 20 g/L of glucose, fructose, sucrose, or lactose were streaked with the LAB isolates and incubated at 30°C for 2 to 3 d. Seven out of 104 isolates (6.7%) produced slimy colonies on agar media containing sucrose, and EPS production was confirmed in liquid medium by isolation of the EPS fraction. Two of the 7 EPS-producing strains were identified as Pediococcus pentosaceus (named AP-1 and AP-3), which produced 6.0 and 2.5 g/L of EPS, respectively.

The studies noted above indicated that the carbon source added to the screening media plays an important role in the detection of the EPS phenotype in LAB; the total amount of polysaccharide produced seems to be strongly influenced by the sugar available in the medium. However, no unique sugar gives the best results because in each case the most suitable carbohydrate is largely dependent on the strain tested. Cerning et al. (1994) showed that the production of EPS by Lb. casei CG11 in basal medium containing glucose was higher than in the same medium with lactose or galactose. On the contrary, Mozzi et al. (1999) found that the production of EPS by Lb. casei CRL87 was 1.7-fold higher in galactose than in glucose. In this case, the differences were correlated with variations in activity of the enzymes involved in the synthesis of the sugar nucleotides acting as precursors of the repeating units that build the HePS. Lactococcus lactis spp. cremoris B40 produced higher amounts of EPS in glucose than in fructose (Looijesteijn et al., 1999). This finding was explained by the low activity of the enzyme fructosebi-phosphatase, which catalyzes the conversion of fructose-1,6-diphosphate into fructose-6-phosphate, an essential step in the biosynthesis of sugar nucleotides from fructose but not from glucose. Other differences in EPS production related to the carbon source of the medium have been attributed to the presence of different sugar transport systems in the LAB strains (Chervaux et al., 2000), if the entry of mono- and disaccharides into the cell is the initial step of EPS synthesis. Both sugar transport and the synthesis of sugar-nucleotide precursors are regulated processes that can be under catabolite repression (Boels et al., 2001a; Laws et al., 2001). Studies on the molecular organization of eps gene clusters and enzymes involved in EPS synthesis and polymerization, as well as the factors regulating these processes, will contribute to our knowledge of the influence of the carbon source on EPS production.

	EPS ISOLATION METHODS

TOP ABSTRACT INTRODUCTION DETECTION OF EPS-PRODUCING... SCREENING OF EPS-PRODUCING LAB... EPS ISOLATION METHODS QUANTIFICATION OF EPS YIELD PRIMARY AND 3-D STRUCTURE... EPS MOLECULAR PARAMETERS RELATED... ACKNOWLEDGEMENTS REFERENCES

 
Different culture media have been used to study the qualitative and quantitative production of EPS and to determine the influence of nutrients in the growth of producing strains and the biosynthesis and genetics of these biopolymers in LAB. The media used most often are skim milk, whey, and whey-based media. It is well established that the culture conditions and the composition (not only the carbon source) of the culture media influence the EPS yield and the molecular characteristics of the biopolymers. Therefore, the choice of an adequate EPS production medium is of great importance, given that some of their components could interfere with the EPS analysis. In this sense, Kimmel and Roberts (1998) showed that yeast extract, beef extract, and proteose-peptone accounted for 94% of the total background EPS-equivalent in MRS broth used to grow the EPS-producing strain Lb. delbrueckii spp. bulgaricus RR. Recently, Vaningelgem et al. (2004) indicated that the glucomannans present in yeast extract and peptone are the carbohydrate-polymer material that interferes with the quantification of EPS produced in complex culture media. Semidefined media, eliminating components that interfere with EPS quantification (Kimmel and Roberts, 1998), and chemically defined media containing a carbon source, amino acids, vitamins, nucleic acid bases, and mineral salts have been developed to facilitate EPS analysis (Degeest et al., 2001b).

In general, the complexity of the method used for the isolation and purification of EPS (Table 2) will depend on the composition of the culture medium used for its production. The simplest procedure involves dialysis against water of the cultured medium (after cell removal by centrifugation), followed by lyophilization. This technique was used to isolate EPS from some Lc. lactis spp. cremoris strains grown in chemically defined media (Marshall et al., 1995; van Kranenburg et al., 1997). Ethanol precipitation may be used to concentrate the EPS before dialysis for the isolation of EPS from thermophilic (yogurt starter: Lb. delbrueckii spp. bulgaricus and Strep. thermophilus) and mesophilic (lactococci and lactobacilli) LAB strains (Van Geel-Schutten et al., 1999; Petry et al., 2000; Torino et al., 2000b; Dal Bello et al., 2001; Degeest et al., 2001a; Rimada and Abraham, 2001; Ricciardi et al., 2002). Where culture medium has increased in complexity, additional purification steps have become necessary to reduce the protein content and other components in the final EPS preparation. For EPS obtained from media with high protein content (e.g., skim milk), 3 approaches are commonly used: precipitation with variable amounts of TCA (final concentration ranging from 4 to 14%), digestion with proteases, or a combination of both. With regard to the first approach, the single protein precipitation with 12% TCA followed by dialysis and lyophilization of the supernatant, was used to isolate the EPS produced by LAB in yogurts in which one of the starter strains synthesized EPS (van Marle and Zoon, 1995). The same procedure was used to isolate EPS produced by 4 Lc. lactis spp. cremoris strains in milk (Ruas-Madiedo et al., 2002b). The coefficient of variation of EPS production (100 x standard deviation/mean) in these 2 studies ranged between 10 and 30% for yogurt starters, and between 5 and 20% for Lc. lactis spp. cremoris strains, respectively. The most common procedure used for isolation from complex media involves TCA precipitation and protein removal by centrifugation, followed by concentration of the EPS by ethanol precipitation (García-Garibay and Marshall, 1991; Cerning et al., 1994; Grobben et al., 1995; Dupont et al., 2000; Frengova et al., 2000; Knoshaug et al., 2000; Pham et al., 2000; Marshall et al., 2001a,b; Van Calsteren et al., 2002; Harding et al., 2003). The yields obtained by this method showed a maximum coefficient of variation between duplicates of 5 to 10%. Less frequently, EPS concentration-precipitation after TCA treatment has been performed using acetone instead of ethanol (Stingele et al., 1996; Lemoine et al., 1997; De Vuyst et al., 1998), or using ethanol and acetone (Faber et al., 1998, 2001b). With regard to the second approach, the isolation of EPS from milk cultures involves the digestion of milk proteins with pronase E (protease type XIV) from Streptomyces griseus, which has a wide range of substrate specificity. It was used to isolate the EPS produced by thermophilic yogurt starters (Cerning et al., 1986, 1988; Mozzi et al., 1995; Bouzar et al., 1996, 1997) or mesophilic strains (Cerning et al., 1992; Mozzi et al., 1996; Torino et al., 2000a, 2001) as well as by a Bifidobacterium longum strain (Abbad-Andaloussi et al., 1995). After heat inactivation of the pronase and a concentration step by UF or evaporation, the EPS fraction was precipitated with ethanol. The results showed a coefficient of variation of approximately 10% between duplicates. Finally, a combination of TCA precipitation and protease digestion has been used with yogurt cultures (Gancel and Novel, 1994; Petry et al., 2003).

Recently, Rimada and Abraham (2003) showed that the method used to isolate the EPS fraction exerted a strong influence on the final amount of EPS obtained. Different methodologies were analyzed to determine the EPS recovery of kefir-grain bacteria in milk and whey-base media. The methods involved 1 or 2 steps of ethanol precipitation, 1 step of ethanol precipitation followed by dialysis, direct dialysis with membranes of different cut-off (1000, 6000, and 12,000 Da), and TCA precipitation. It seems that the heat treatment of the samples as a first step of EPS isolation is critical for complete recovery of the polymer. The methods involving ethanol precipitation rendered similar final amounts of EPS, although a single step of ethanol precipitation did not entirely eliminate the residual lactose. The TCA precipitation step reduced (by about 50%) the amount of EPS recovered, given that the biopolymer coprecipitates with the proteins. However, this is the procedure of choice when full physicochemical EPS characterization is required because the resulting EPS fraction has fewer impurities.

	QUANTIFICATION OF EPS YIELD

TOP ABSTRACT INTRODUCTION DETECTION OF EPS-PRODUCING... SCREENING OF EPS-PRODUCING LAB... EPS ISOLATION METHODS QUANTIFICATION OF EPS YIELD PRIMARY AND 3-D STRUCTURE... EPS MOLECULAR PARAMETERS RELATED... ACKNOWLEDGEMENTS REFERENCES

 
After the isolation steps, a lyophilized powder is obtained, its weight being the simplest indication of the EPS yield (De Vuyst et al., 1998; Van Geel-Schutten et al., 1999; Frengova et al., 2000; Degeest et al., 2001a). The EPS production can also be expressed as the equivalent milligrams of dextran per milliliter. This unit was defined by García-Garibay and Marshall (1991) as the amount of polymer that produces the same turbidity at 720 nm as one milligram of dextran (molar mass: 2 x 10³ kDa) under the same conditions of measurement. However, values obtained by these methods are strongly influenced by the degree of purity of the isolated EPS fraction, because proteins and nonEPS carbohydrates (when present) are also quantified. A colorimetric procedure for the determination of sugar and related compounds is the phenol-sulfuric method described by Dubois et al. (1956). Simple sugars, oligosaccharides, polysaccharides, and their derivatives (methyl ethers) render an orange-yellow color when treated with phenol and concentrated sulfuric acid. This method has been widely used as an indication of the EPS yield obtained by different isolation procedures (Cerning et al., 1986, 1988, 1992, 1994; Nakajima et al., 1990, 1992; Gruter et al., 1992, 1993; Gancel and Novel, 1994; Abbad-Andaloussi et al., 1995; Grobben et al., 1995; Marshall et al., 1995; Mozzi et al., 1995, 1996; Roberts et al., 1995; Bouzar et al., 1996, 1997; Stingele et al., 1996; Lemoine et al., 1997; Urashima et al., 1999; Dupont et al., 2000; Frengova et al., 2000; Higashimura et al., 2000; Petry et al., 2000, 2003; Bergmaier et al., 2001; Torino et al., 2001; Macedo et al., 2002). However, the phenol-sulfuric method quantifies the total carbohydrates present in the EPS fraction, including low molecular weight carbohydrates that may be present. Ruijssenaars et al. (2000) determined more precisely the amount of EPS by subtracting the reducing sugar fraction (RS) estimated by the dinitrosalicylic acid method (Miller, 1959) from the total sugar (TS) fraction measured by the phenol-sulfuric method (EPS = TS – RS). Another colorimetric procedure used by several authors for the quantification of sugars (Van den Berg et al., 1995; Levander et al., 2001; Rimada and Abraham, 2001) involves the use of the anthrone reagent (Morris, 1948).

Although colorimetric quantification methods are, in principle, the cheapest and simplest ones, the most accurate techniques for the quantification of the EPS content of EPS isolated fractions are those in which the analytical method is coupled to a separation method. Thus, gel permeation chromatography in an HPLC system allows the separation of the EPS polymers by size exclusion. The simultaneous detection of the EPS molecules by refractive index (RI) can be used to quantify the EPS production in the corresponding EPS elution peak (Faber et al., 1998, 2001b; Ruas-Madiedo et al., 2002b). An additional online ultraviolet detector can provide information on the presence of protein in the sample (Tuinier et al., 1999b; Ruas-Madiedo et al., 2002b).

Finally, a method for the simultaneous quantification of lactic acid, lactose, and EPS yield directly in culture media (without an isolation procedure) has been developed recently using near infrared spectroscopy (NIRS) (Macedo et al., 2002). Results obtained by this method showed a good coefficient of correlation (R²) with reference methods (quantification of lactose and lactic acid by HPLC, and quantification of the EPS fraction isolated by UF with the phenol-sulfuric method). The R² for lactic acid, lactose, and EPS were 99, 99, and 91%, respectively, which suggests that NIRS could be useful for rapid monitoring of EPS and lactic acid production during fermentations.

Due to the diversity of quantification methods existing it is difficult to establish typical EPS yields for specific LAB species. In general, HoPS are produced in larger quantities than HePS (Table 3). Lactobacillus reuteri LB121 can produce 10 g/L of 2 glucan- and fructan-type, homopolymers (Van Geel-Schutten et al., 1999). The reported yields of HePS range from 50 to 350 mg/L for Strep. thermophilus, 60 to 150 mg/L for Lb. delbrueckii spp. bulgaricus, 25 to 600 mg/L for Lc. lactis spp. cremoris, and from 50 to 60 mg/L for Lb. casei (Cerning, 1995).

	PRIMARY AND 3-D STRUCTURE OF EPS

TOP ABSTRACT INTRODUCTION DETECTION OF EPS-PRODUCING... SCREENING OF EPS-PRODUCING LAB... EPS ISOLATION METHODS QUANTIFICATION OF EPS YIELD PRIMARY AND 3-D STRUCTURE... EPS MOLECULAR PARAMETERS RELATED... ACKNOWLEDGEMENTS REFERENCES

The qualitative monosaccharide composition of an EPS has been analyzed in the past by TLC (Cerning et al., 1986, 1988, 1994; Gruter et al., 1992, 1993; Lemoine et al., 1997). However, this method has low discriminatory power and has been largely surpassed by more reliable liquid and gas chromatographic techniques.

The qualitative and quantitative determination of EPS monosaccharides by HPLC involves the separation of monosaccharides by anion-exchange columns and detection by RI. Isocratic separations at 35 to 70°C using 2.5 to 5 mM sulfuric acid as eluent were used to analyze the monomer composition of EPS produced by Lactobacillus rhamnosus (Van Calsteren et al., 2002), Lb. delbrueckii spp. bulgaricus (Grobben et al., 1995), and Strep. thermophilus strains (De Vuyst et al., 1998). Nonetheless, Torino et al. (2000a) used isocratic elution with distilled water to determine the monosaccharide constituents of the EPS produced by Lactobacillus helveticus ATCC 15807. Another technique used for the identification and quantification of mono- and oligosaccharides resulting from the partial hydrolysis of EPS is the high-performance anion-exchange chromatography pulse amperometric detection (HPAEC-PAD) (Gruter et al., 1992, 1993; Lemoine et al., 1997; Levander et al., 2001). Both HPLC-RI and HPAEC-PAD are liquid chromatographic methods that differ in sensitivity and accuracy (Cataldi et al., 2000). Neither of them requires preliminary derivatization or other chemical treatment of samples. The retention mechanism of HPLC anion-exchange columns is based on the formation of weak complexes between the monosaccharides and ions (higher affinity to the immobilized ions in the matrix means longer retention times). The main disadvantages of this method are the low resolving power and the use of a nonselective and low sensitivity RI detection system. In the HPAEC system, the columns have polymer-base matrices characterized by high chemical stability and selectivity (determined by the mobile phase used). Separation is made at high pH with strong alkaline solutions and detection is achieved by monitoring the change in the electric current due to the oxidation of the saccharides in the surface of the gold working electrodes located in the pulse amperometric detector. The procedure is very sensitive and selective for the analysis of sugar compounds in their native state.

The most extensively used technology for the analysis of the monomer composition of EPS isolated from LAB is gas chromatography/mass spectrometry (GCMS). After acid hydrolysis of EPS molecules, generally with trifluoroacetic acid, the resulting monosaccharides are derivatized to alditol acetates (Blakeney et al., 1983) or trimethylsilylated glycosides (Gerwig et al., 1978, 1979), which are then separated by gas chromatography using different gas carriers (He, N₂, H₂), columns, and temperature programs. The identification and quantification of monosaccharides can be carried out with a flame ionization detector or a mass spectrometer, using inositol as an internal standard. The monomeric composition of several EPS produced by different LAB strains such as Lc. lactis spp. cremoris, Strep. thermophilus, Lb. delbrueckii spp. bulgaricus, Lb. helveticus, Lactobacillus sake, Lb. reuteri, Lb. rhamnosus, and Bifidobacterium longum was determined using the GCMS method (Nakajima et al., 1990; Marshall et al., 1995, 2001a; Roberts et al., 1995; Van den Berg et al., 1995; Bouzar et al., 1996, 1997; Lemoine et al., 1997; Van Geel-Schutten et al., 1999; Petry et al., 2000, 2003; Yang et al., 2000; Ricciardi et al., 2002; Lipinski et al., 2003). The sugar linkage analysis is achieved by methylation of EPS, hydrolysis followed by reduction with sodium borodeuteride, and further acetylation. The partially methylated deuterated alditol acetates obtained can be identified by GCMS (Faber et al., 1998, 2001a,Faber et al., b; van Casteren et al., 1998, 2000a, van Casteren et al., b; Harding et al., 2003).

Finally, to obtain the 3-D structure of an EPS molecule, both the ring size (pyranose/furanose) of the monosaccharide residues and the relative orientations of the adjacent monosaccharides have to be determined. Nuclear magnetic resonance is the technique most often used to study the conformation of molecules in solution and allows elucidation of the type of glycosidic linkages and the structure of the repeating units that build the EPS molecules. Before nuclear magnetic resonance analysis, the EPS sample must be exchanged in D₂O (deuteration) during a procedure repeated several times that may involve intermediate lyophilization steps. To date, the structure of several EPS synthesized by Strep. thermophilus (Doco et al., 1990; Lemoine et al., 1997; Urashima et al., 1999; Faber et al., 1998, 2001b, 2002; Marshall et al., 2001a), Lc. lactis spp. cremoris (Gruter et al., 1992; Nakajima et al., 1992; Yang et al., 1999; van Casteren et al., 1998, 2000a, van Casteren et al., b), different Lactobacillus species (Gruter et al., 1993; Van Geel-Schutten et al., 1999; Faber et al., 2001a; Staaf et al., 2000; Yang et al., 2000; Van Calsteren et al., 2002; Harding et al., 2003; Lipinski et al., 2003) and the strain Bifidobacterium adolescentis M101-4 (Hosono et al., 1997) has been determined by nuclear magnetic resonance. For a more detailed compilation of solved EPS structures, refer to the reviews included in Table 1.

	EPS MOLECULAR PARAMETERS RELATED TO THE VISCOSITY INTENSIFYING ABILITY: MOLAR MASS AND RADIUS OF GYRATION

TOP ABSTRACT INTRODUCTION DETECTION OF EPS-PRODUCING... SCREENING OF EPS-PRODUCING LAB... EPS ISOLATION METHODS QUANTIFICATION OF EPS YIELD PRIMARY AND 3-D STRUCTURE... EPS MOLECULAR PARAMETERS RELATED... ACKNOWLEDGEMENTS REFERENCES

 
The ability to confer viscosity of EPS, in aqueous solutions or in fermented products, is largely (but not exclusively) determined by the molecular parameters of the biopolymer, i.e., the molar mass (M) and the radius of gyration (Rg) of the molecule (Tuinier et al., 2001; Ruas-Madiedo et al., 2002b). Briefly, Rg could be considered as a measure of the polymer size and molar mass as an indication of its length. The intrinsic viscosity ([]₀) of an EPS molecule is the parameter that determines its viscosity-intensifying ability and can be calculated from M and Rg according to the equation formulated by Tuinier et al. (1999a):

The molar mass of an EPS can be determined by gel permeation chromatography in an HPLC or fast protein liquid chromatography (FPLC) system. In HPLC, detection is achieved using RI by comparing the EPS peak retention time with known dextran standards of different molar mass (Marshall et al., 1995, 2001a; Mozzi et al., 1996; Torino et al., 2000a, 2001; Van Calsteren et al., 2002; Harding et al., 2003). Less frequently, other standards such as pullulan have been used (Nakajima et al., 1990; Urashima et al., 1999). In fast protein liquid chromatography, the EPS concentration can be determined in the collected fractions by the phenol-sulfuric method (Stingele et al., 1996; Lemoine et al., 1997). The molar mass and Rg of the EPS can be simultaneously determined by gel permeation chromatography using a multiangle laser light scattering detector coupled online with the RI detector in the HPLC system (Van den Berg et al., 1995; Faber et al., 1998; Tuinier et al., 1999b; Higashimura et al., 2000; Ruas-Madiedo et al., 2002b; Petry et al., 2003). The molar mass and Rg of the EPS produced by LAB varied according to strains and, as previously indicated, depended on the polymer type. The molar mass values described in literature for HoPS ranged from 2.7 x 10⁶ to 2.2 x 10⁷ Da for strains of Streptococcus mutans, and from 1.5 x 10⁵ to 3.5 x 10⁶ Da for strains of Lb. reuteri. In general, HePS are smaller and some molar mass values of different LAB species ranged from 4.0 x 10⁴ to 9.0 x 10⁶ Da for Strep. thermophilus strains, 1 x 10⁵ to 2 x 10⁶ Da for Lc. lactis spp. cremoris strains, 2.5 x 10⁴ to 1.4 x 10⁶ Da for Lb. rhamnosus strains, and 3.5 x 10⁵ Da for the strain Bifidobacterium longum BB-79 (Roberts et al., 1995).

In summary, full characterization of an EPS requires information on microbial EPS phenotype, amount of EPS produced, sugar composition, structure, and molecular parameters related to the viscosity intensifying properties. Techniques most often used for the analysis of monomer composition, molar mass, radius of gyration, and structure are those employing gas and liquid chromatography with different detection methods, and nuclear magnetic resonance. It seems evident that the wide variety of culture media and methods used to isolate and quantify EPS has a strong influence on the yield results, which makes it difficult to compare values obtained by different authors. Few studies compare the degree of purity achieved in the EPS fraction after isolation by the different methods currently available. This fact is of great importance given that isolation procedures that render a high purity EPS fraction are desirable, or even necessary, for specific studies on chemical composition, structure, and physical characterization of EPS molecules. In spite of this, it is interesting to note that the development of procedures and analytical methods summarized in this review permitted, in recent years, the isolation and characterization of a large number of EPS produced by LAB strains from different natural ecological niches. The challenge for future research on EPS produced by LAB is to elucidate the structure-function relationship. The influence of the amount, chemical composition, chain length, and molecular size of EPS molecules on the physical properties of dairy fermented products is known. Now it is necessary to correlate a specific EPS structure with specific values of texture and viscosity. Moreover, the EPS molecular characteristics that might play a role in the beneficial effects for human health are still not well understood. However, it is expected that the presence of certain glycosidic linkages and their accessibility to glycolytic enzymes will be important for the putative prebiotic effect of EPS. The elucidation of the key parameters involved in EPS functionality would allow the development of EPS engineering strategies for specific food applications.

	ACKNOWLEDGEMENTS

TOP ABSTRACT INTRODUCTION DETECTION OF EPS-PRODUCING... SCREENING OF EPS-PRODUCING LAB... EPS ISOLATION METHODS QUANTIFICATION OF EPS YIELD PRIMARY AND 3-D STRUCTURE... EPS MOLECULAR PARAMETERS RELATED... ACKNOWLEDGEMENTS REFERENCES

 
The authors are grateful to A. Margolles at IPLA, and P. Lamosa and C. Sánchez at ITQB (Instituto de Tecnología Química e Biológica, Oeiras, Portugal) for the critical reading of this manuscript. A. M. Hernández (IPLA) is acknowledged for her technical advice. P. Ruas-Madiedo was the recipient of an I3P postdoctoral research contract granted by CSIC (Spanish Ministry of Science and Technology) and FEDER funds (European Union). This work was financed by FEDER funds and the Spanish "Plan Nacional de I+D+I" (projects AGL 2001-2296 and AGL 2004-06088-CO2-01/ALI), and by a grant from the "Fundación Príncipe de Asturias" ("Beca Grande Covián").

Received for publication May 26, 2004. Accepted for publication November 30, 2004.

	REFERENCES

TOP ABSTRACT INTRODUCTION DETECTION OF EPS-PRODUCING... SCREENING OF EPS-PRODUCING LAB... EPS ISOLATION METHODS QUANTIFICATION OF EPS YIELD PRIMARY AND 3-D STRUCTURE... EPS MOLECULAR PARAMETERS RELATED... ACKNOWLEDGEMENTS REFERENCES

 


Abbad-Andaloussi, S., H. Talbaoui, R. Marczak, and R. Bonaly. 1995. Isolation and characterization of exocellular polysaccharides produced by Bifidobacterium longum. Appl. Microbiol. Biotechnol. 43:995–1000.[Medline]

Ariga, H., T. Urashima, E. Michihata, M. Ito, N. Morizono, T. Kimura, and S. Takahashi. 1992. Extracellular polysaccharide from encapsulated Streptococcus salivarius subsp. thermophilus OR 901 isolated from commercial yogurt. J. Food Sci. 57:625–628.

Bergmaier, D., C. Lacroix, M. G. Macedo, and P. Champagne. 2001. New method for exopolysaccharide determination in culture broth using stirred ultrafiltration cells. Appl. Microbiol. Biotechnol. 57:401–406.[Medline]

Blakeney, A. B., P. J. Harris, R. J. Henry, and B. A. Stone. 1983. A simple and rapid preparation of alditol acetates for monosaccharide analysis. Carbohydr. Res. 113:291–299.

Boels, I. C., A. Ramos, M. Kleerebezem, and W. M. de Vos. 2001a. Functional analysis of the Lactococcus lactis galU and galE genes and their impact on sugar nucleotide and exopolysaccharide biosynthesis. Appl. Environ. Microbiol. 67:3033–3040.[Abstract/Free Full Text]

Boels, I. C., R. van Kranenburg, J. Hugenholtz, M. Kleerebezem, and W. M. de Vos. 2001b. Sugar catabolism and its impact on the biosynthesis and engineering of exopolysaccharide production in lactic acid bacteria. Int. Dairy J. 11:723–732.

Bouzar, F., J. Cerning, and M. Desmazeaud. 1996. Exopolysaccharide production in milk by Lactobacillus delbrueckii spp. bulgaricus CNRZ 1187 and by two colonial variants. J. Dairy Sci. 79:205–211.[Abstract]

Bouzar, F., J. Cerning, and M. Desmazeaud. 1997. Exopolysaccharide production and texture-promoting abilities of mixed-strain starter cultures in yogurt production. J. Dairy Sci. 80:2310–2317.[Abstract]

Broadbent, J. R., D. J. McMahon, D. L. Welker, C. J. Oberg, and S. Moineau. 2003. Biochemistry, genetics and applications of exopolysaccharide production in Streptococcus thermophilus: A review. J. Dairy Sci. 86:407–423.[Abstract/Free Full Text]

Cataldi, T. R. I., C. Campa, and G. E. De Benedetto. 2000. Carbohydrate analysis by high-performance anion-exchange chromatography with pulsed amperometric detection: The potential is still growing. Fresenius J. Anal. Chem. 368:739–758.[Medline]

Cerning, J. 1990. Exocellular polysaccharides produced by lactic acid bacteria. FEMS Microbiol. Rev. 87:113–130.

Cerning, J. 1995. Production of exopolysaccharides by lactic acid bacteria and dairy propionibacteria. Lait 75:463–472.

Cerning, J., C. Bouillanne, M. J. Desmazeaud, and M. Landon. 1986. Isolation and characterization of exocellular polysaccharide produced by Lactobacillus bulgaricus. Biotechnol. Lett. 8:625–628.

Cerning, J., C. Bouillanne, M. J. Desmazeaud, and M. Landon. 1988. Exocellular polysaccharide production by Streptococcus thermophilus. Biotechnol. Lett. 10:255–260.

Cerning, J., C. Bouillanne, M. Landon, and M. Desmazeaud. 1992. Isolation and characterization of exopolysaccharides from slime-forming mesophilic lactic acid bacteria. J. Dairy Sci. 75:692–699.[Abstract]

Cerning, J., C. M. G. C. Renard, J. F. Thibault, C. Bouillanne, M. Landon, M. Desmazeaud, and L. Topisirovic. 1994. Carbon source requirements for exopolysaccharide production by Lactobacillus casei CG11 and partial structure analysis of the polymer. Appl. Environ. Microbiol. 60:3914–3919.[Abstract/Free Full Text]

Chabot, S., H. L. Yu, L. De Léséleuc, D. Cloutier, M. R. van Calsteren, M. Lessard, D. Roy, M. Lacroix, and D. Oth. 2001. Exopolysaccharide from Lactobacillus rhamnosus RW-9595M stimulate TNF, IL-6 and IL-12 in human and mouse cultured inmunocompetent cells, and IFN-g in mouse splenocytes. Lait 81:683–697.

Chervaux, C., S. D. Ehrlich, and E. Maguin. 2000. Physiological study of Lactobacillus delbrueckii subsp. bulgaricus strains in a novel chemically defined medium. Appl. Environ. Microbiol. 66:5306–5311.[Abstract/Free Full Text]

Dal Bello, F. D., J. Walter, C. Hertel, and W. P. Hammes. 2001. In vitro study of prebiotic properties of levan-type exopolysaccharides from lactobacilli and non-digestible carbohydrates using denaturing gradient gel electrophoresis. Syst. Appl. Microbiol. 24:232–237.[Medline]

Degeest, B., B. Janssens, and L. De Vuyst. 2001a. Exopolysaccharide (EPS) biosynthesis by Lactobacillus sakei 0–1: Production kinetics, enzyme activities and EPS yields. J. Appl. Microbiol. 91:470–477.[Medline]

Degeest, B., F. Vaningelgem, and L. De Vuyst. 2001b. Microbial physiology, fermentation kinetics and process engineering of heteropolysaccharides production by lactic acid bacteria. Int. Dairy J. 11:747–758.

De Vuyst, L., and B. Degeest. 1999. Heteropolysaccharides from lactic acid bacteria. FEMS Microbiol. Rev. 23:153–177.[Medline]

De Vuyst, L., F. De Vin, F. Vaningelgem, and B. Degeest. 2001. Recent developments in the biosynthesis and applications of heteropolysaccharides from lactic acid bacteria. Int. Dairy J. 11:687–708.

De Vuyst, L., F. Vanderveken, S. van den Ven, and B. Degeest. 1998. Production by and isolation of exopolysaccharides from Streptococcus thermophilus grown in a milk medium and evidence for their growth-associated biosynthesis. J. Appl. Microbiol. 84:1059–1068.[Medline]

Dierksen, K. P., W. E. Sandine, and J. E. Trempy. 1997. Expression of ropy and mucoid phenotypes in Lactococcus lactis. J. Dairy Sci. 80:1528–1536.[Abstract]

Doco, T., J. M. Wieruszeski, B. Fournet, D. Carcano, P. Ramos, and A. Loones. 1990. Structure of an exocellular polysaccharide produced by Streptococcus thermophilus. Carbohydr. Res. 198:313–321.[Medline]

Duboc, P., and B. Mollet. 2001. Applications of exopolysaccharides in dairy industry. Int. Dairy J. 11:759–768.

Dubois, M., K. A. Gilles, J. K. Hamilton, P. A. Rebers, and F. Smith. 1956. Colorimetric method for determination of sugar and related substances. Anal. Chem. 28:350–356.

Dupont, I., D. Roy, and G. Lapointe. 2000. Comparison of exopolysaccharide production by strains of Lactobacillus rhamnosus and Lactobacillus paracasei grown in chemically defined medium and milk. J. Industr. Microbiol. Biotechnol. 24:251–255.

Faber, E. J., J. P. Kamerling, and J. F. G. Vliegenthart. 2001a. Structure of the extracellular polysaccharide produced by Lactobacillus delbrueckii subsp. bulgaricus 291. Carbohydr. Res. 331:183–194.[Medline]

Faber, E. J., M. J. van den Haak, J. P. Kamerling, and J. F. G. Vliegenthart. 2001b. Structure of the exopolysaccharide produced by Streptococcus thermophilus S3. Carbohydr. Res. 331:173–182.[Medline]

Faber, E. J., D. J. van Haaster, J. P. Kamerling, and J. F. G. Vliegenthart. 2002. Characterization of the exopolysaccharide produced by Streptococcus thermophilus 8S containing an open chain nononic acid. Eur. J. Biochem. 269:5590–5598.[Medline]

Faber, E. J., P. Zoon, J. P. Kamerling, and J. F. G. Vliegenthart. 1998. The exopolysaccharides produced by Streptococcus thermophilus Rs and Sts have the same repeating unit but differ in viscosity of their milk cultures. Carbohydr. Res. 310:269–276.[Medline]

Frengova, G. I., E. D. Simova, D. M. Beshkova, and Z. I. Simov. 2000. Production and monomer composition of exopolysaccharides by yogurt starter cultures. Can. J. Microbiol. 46:1123–1127.[Medline]

Gancel, F., and G. Novel. 1994. Exopolysaccharide production by Streptococcus salivarius ssp. thermophilus cultures. 1. Conditions of production. J. Dairy Sci. 77:685–688.[Abstract]

Gancel, F., G. Novel, D. Carcano, A. Loones, and P. Ramos. 1989. Selection process for bacterial clones producing exopolysaccharide(s) or muc (plus), by adding cationic colourant to culture medium of bacterial strain. Sodima Union Corp., assignee. French patent FR2632968-A.

García-Garibay, M., and V. M. Marshall. 1991. Polymer production by Lactobacillus delbrueckii bulgaricus. J. Appl. Bacteriol. 70:325–328.

Germond, J. E., M. Delley, N. D’Amico, and S. J. F. Vincent. 2001. Heterologous expression and characterization of the exopolysac-charide from Streptococcus thermophilus Sfi39. Eur. J. Biochem. 268:5149–5156.[Medline]

Gerwig, G. J., J. P. Kamerling, and J. F. G. Vliegenthart. 1978. Determination of the D and L configuration of neutral monosaccharides by high resolution capillary GLC. Carbohydr. Res. 62:349–357.

Gerwig, G. J., J. P. Kamerling, and J. F. G. Vliegenthart. 1979. Determination of the absolute configuration of monosaccharides in complex carbohydrates by capillary GLC. Carbohydr. Res. 77:1–7.

Grobben, G. J., J. Sikkema, M. R. Smith, and J. A. M. de Bont. 1995. Production of extracellular polysaccharides by Lactobacillus delbrueckii ssp. bulgaricus NCFB2772 grown in a chemically defined medium. J. Appl. Bacteriol. 79:103–107.

Gruter, M., B. R. Leeflang, J. Kuiper, P. Kamerling, and F. G. Vliegenthart. 1992. Structure of the exopolysaccharide produced by Lactococcus lactis subspecies cremoris H414 grown in a defined medium or skimmed milk. Carbohydr. Res. 231:273–291.[Medline]

Gruter, M., B. R. Leeflang, J. Kuiper, P. Kamerling, and F. G. Vliegenthart. 1993. Structural characterisation of the exopolysaccharide produced by Lactobacillus delbrueckii subspecies bulgaricus rr grown in skimmed milk. Carbohydr. Res. 239:209–226.[Medline]

Harding, L. P., V. M. Marshall, M. Elvin, Y. Gu, and A. P. Laws. 2003. Structural characterisation of a perdeuteriomethylated exo-polysaccharide by NMR spectroscopy: Characterisation of the novel exopolysaccharide produced by Lactobacillus delbrueckii subsp. bulgaricus EU23. Carbohydr. Res. 338:61–67.[Medline]

Hassan, A. N., M. Corredig, and J. F. Frank. 2002a. Capsule formation by nonropy starter cultures affects the viscoelastic properties of yogurt during structure formation. J. Dairy Sci. 85:716–720.[Abstract]

Hassan, A. N., M. Corredig, J. F. Frank, and M. Elsoda. 2004. Microstructure and rheology of an acid-coagulated cheese (Karish) made with an exopolysaccharide-producing Streptococcus thermophilus strain and its exopolysaccharide non-producing genetic variant. J. Dairy Res. 71:116–120.[Medline]

Hassan, A. N., J. F. Frank, and M. Elsoda. 2003. Observation of bacterial exopolysaccharide in dairy products using cryo-scanning electron microscopy. Int. Dairy J. 13:755–762.

Hassan, A. N., J. F. Frank, M. A. Farmer, K. A. Schmidt, and S. I. Shalabi. 1995. Observation of encapsulated lactic acid bacteria using confocal scanning laser microscopy. J. Dairy Sci. 78:2624–2628.[Abstract]

Hassan, A. N., J. F. Frank, and K. B. Qvist. 2002b. Direct observation of bacterial exopolysaccharides in dairy products using confocal scanning laser microscopy. J. Dairy Sci. 85:1705–1708.[Abstract/Free Full Text]

Higashimura, M., B. W. Mulder-Bosman, R. Reich, T. Iwasaki, and G. W. Robjin. 2000. Solution properties of Viilian, the exopolysaccharide from Lactococcus lactis subsp. cremoris SBT 0495. Biopolymers 54:143–158.[Medline]

Hosono, A., J. Lee, A. Ametani, M. Natsume, M. Hirayama, T. Adachi, and S. Kaminogawa. 1997. Characterization of a water-soluble polysaccharide fraction with inmunopotentiating activity from Bifidobacterium adolescentis M101-4. Biosci. Biotechnol. Biochem. 61:312–316.

Jolly, L., and F. Stingele. 2001. Molecular organization and functionality of exopolysaccharide gene clusters in lactic acid bacteria. Int. Dairy J. 11:733–746.

Jolly, L., S. J. F. Vincent, P. Duboc, and J. R. Neeser. 2002. Exploiting exopolysaccharides from lactic acid bacteria. Antonie Van Leeuwenhoek 82:367–374.[Medline]

Kimmel, S. A., and R. F. Roberts. 1998. Development of a growth medium suitable for exopolysaccharide production by Lactobacillus delbrueckii ssp. bulgaricus RR. Int. J. Food Microbiol. 40:87–92.[Medline]

Kitazawa, H., T. Harata, J. Uemura, T. Saito, T. Kaneko, and T. Itoh. 1998. Phosphate group requirement for mitogenic activation of lymphocytes by an extracellular phosphopolysaccharide from Lactobacillus delbrueckii spp. bulgaricus. Int. J. Food Microbiol. 40:169–175.[Medline]

Kleerebezem, M., R. van Kranenburg, R. Tuinier, I. C. Boels, P. Zoon, E. Looijesteijn, J. Hugenholtz, and W. M. de Vos. 1999. Exopolysaccharides produced by Lactococcus lactis: From genetic engineering to improved rheological properties? Antonie Van Leeuwenhoek 76:357–365.[Medline]

Knoshaug, E. P., J. A. Ahlgren, and J. E. Trempy. 2000. Growth associated exopolysaccharide expression in Lactococcus lactis subspecies cremoris Ropy352. J. Dairy Sci. 83:633–640.[Abstract]

Korakli, M., M. G. Gänzle, and R. F. Vogel. 2002. Metabolism by bifidobacteria and lactic acid bacteria of polysaccharides from wheat and rye, and exopolysaccharides produced by Lactobacillus sanfranciscensis. J. Appl. Microbiol. 92:958–965.[Medline]

Korakli, M., A. Rossmann, M. G. Gänzle, and R. F. Vogel. 2001. Sucrose metabolism and exopolysaccharide production in wheat and rye sourdoughs by Lactobacillus sanfranciscensis. J. Agric. Food Chem. 49:5194–5200.[Medline]

Laws, A., Y. Gu, and V. Marshall. 2001. Biosynthesis, characterization, and design of bacterial exopolysaccharides from lactic acid bacteria. Biotechnol. Adv. 19:597–625.[Medline]

Laws, A. P., and V. M. Marshall. 2001. The relevance of exopolysaccharides to the rheological properties in milk fermented with ropy strains of lactic acid bacteria. Int. Dairy J. 11:709–722.

Lemoine, J., F. Chirat, J. M. Wieruszeski, G. Strecker, N. Favre, and J. R. Neeser. 1997. Structural characterization of the exocellular polysaccharides produced by Streptococcus thermophilus Sfi39 and Sfi12. Appl. Environ. Microbiol. 63:3512–3518.[Abstract]

Levander, F., M. Svensson, and P. Ra °dström. 2001. Small-scale analysis of exopolysaccharides from Streptococcus thermophilus grown in a semi-defined medium. BMC Microbiology, 1: 23. Online. Available: http://www.biomedcentral.com/1471-2180/1/23. Accessed Sept. 26, 2001.[Medline]

Lipinski, T., Ch. Jones, X. Lemercinier, A. Korzeniowska-Kowal, M. Strus, J. Rybka, A. Gamian, and P. B. Heczko. 2003. Structural analysis of the Lactobacillus rhamnosus strain KL37C exopolysaccharide. Carbohydr. Res. 338:605–609.[Medline]