Accepted Manuscript Title: Characterization of an exopolysaccharide produced by Lactobacillus plantarum YW11 isolated from Tibet Kefir Author: Ji Wang Xiao Zhao Zheng Tian Yawei Yang Zhennai Yang PII: DOI: Reference:
S0144-8617(15)00186-1 http://dx.doi.org/doi:10.1016/j.carbpol.2015.03.003 CARP 9743
To appear in: Received date: Revised date: Accepted date:
12-9-2014 27-2-2015 2-3-2015
Please cite this article as: Wang, J., Zhao, X., Tian, Z., Yang, Y., and Yang, Z.,Characterization of an exopolysaccharide produced by Lactobacillus plantarum YW11 isolated from Tibet Kefir, Carbohydrate Polymers (2015), http://dx.doi.org/10.1016/j.carbpol.2015.03.003 This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.
Highlights (for review)
1. Physicochemical, rheological properties, structural characterization and
microstructure of the EPS from L. plantarum are reported.
10 11 12
at acidic pHs.
4. Unlike reported before, the EPS from L. plantarum YW11 shows a
3. The EPS has higher viscosity in skim milk, at lower temperature, or
highly branched and porous structure.
5. The EPS has high thermal stability.
for exploitation in food industry.
2. The EPS produced by L. plantarum YW11 is a promising candidate
Ac ce p
Page 1 of 37
Characterization of an exopolysaccharide produced by
Lactobacillus plantarum YW11 isolated from Tibet Kefir
14 15 16 17 18
Ji Wanga,b, Xiao Zhaoa, Zheng Tiana, Yawei Yanga, Zhennai Yanga,b,* a. Beijing Laboratory of Food Quality and Safety, Beijing Technology and Business University, Beijing, China;1 b. School of Biological and Agricultural Engineering, Jilin University, Changchun, China
Abstract: An exopolysaccharide (EPS)-producing strain YW11 isolated from Tibet
Kefir was identified as Lactobacillus plantarum, and the strain was shown to produce
90 mg L-1 of EPS when grown in a semi-defined medium. The molecular mass of the
EPS was 1.1 × 105 Da. The EPS was composed of glucose and galactose in a molar
ratio of 2.71:1, with possible presence of N-acetylated sugar residues in the
polysaccharide as confirmed by NMR spectroscopy. Rheological studies showed that
the EPS had higher viscosity in skim milk, at lower temperature, or at acidic pHs. The
viscous nature of the EPS was confirmed by observation with scanning electron
microscopy that demonstrated a highly branched and porous structure of the
Ac ce p
Keywords: Exopolysaccharide; Lactobacillus plantarum; Rheology; Structural
28 29 30
polysaccharide. The atomic force microscopy of the EPS further revealed presence of many spherical lumps, facilitating binding with water in aqueous solution. The EPS had a higher degradation temperature (287.7 °C), suggesting high thermal stability of the EPS.
*Corresponding author. Beijing Laboratory of Food Quality and Safety, Beijing Technology and Business University, No. 11 Fu-Cheng Road, Hai-Dian District, Beijing, 100048 China. Tel.: +86 10 68984870; Fax: +86 10 68985456. Email: [email protected]
Page 2 of 37
Chemical compounds studied in this article
Glucose (PubChem CID: 79025); Fructose (PubChem CID: 5984); Rhamnose
(PubChem CID: 25310); Glycerol (PubChem CID: 753); Glucuronic acid (PubChem
CID: 444791); Sodium nitrate (PubChem CID: 24268); Sodium chloride (PubChem
CID: 5234); Calcium chloride (PubChem CID: 21226093); Trifluoroacetic acid
(PubChem CID: 6422); Phenol (PubChem CID: 996); Sulfuric acid (PubChem CID:
41 1. Introduction
Exopolysaccharides (EPSs) are extracellular biopolymers that are produced during
the metabolic process of microorganisms such as bacteria, fungi, and blue-green algae
(Amjres et al., 2014). The EPSs could be either covalently associated with the cell
47 48 49
Ac ce p
surface forming a capsule, or be loosely attached, or totally secreted into the surrounding environment during the cell growth (Yang et al., 2010). Among the wide variety of EPS-producing microorganisms, lactic acid bacteria (LAB) are generally regarded as safe due to their long history of safe use in human consumption (Nikolic
et al., 2012). In recent years, owing to the unique physicochemical properties and
biological activities, the EPSs produced by LAB have been used in a wide range of
applications including food products, pharmaceuticals, bioflocculants, bioemulsifiers
and chemical products (Liu et al., 2010; Ye, Liu, Wang, Wang, & Zhang, 2012; Ismail 3
Page 3 of 37
& Nampoothiri, 2010). In the fermented food industry, LAB EPSs are usually used as
natural alternatives to commercial stabilizers because of their viscosifying, stabilizing,
emulsifying or gelling properties for improving the rheology, texture and mouth-feel
of the fermented products including yoghurt and cheese (Ahmed, Wang, Anjum,
Ahmad, & Khan, 2013). In addition, extensive research has revealed that some EPSs
produced by LAB may be correlated with promoting human health and preventing
diseases due to their pharmacological activities, such as immunostimulatory,
immunomodulatory, antitumor, antibiofilm, antioxidant and cholesterol lowering
activities (Liu, Chu, Chou, & Yu, 2011; Kanmani et al., 2011; Li et al., 2014a;
Lindström, Holst, Nilsson, Öste, & Andersson, 2012). Since bacterial EPS are generally
of protective nature, production of EPS may increase the resistance of the bacteria
against unfavorable environmental factors, e.g. resistance to high acidity and bile salts
(de los Reyes-Gavilán et al., 2011).
68 69 70
Ac ce p
In the last decade, a large number of EPS-producing LAB have been isolated from
a variety of fermented foods such as yoghurt, cheese, sausages, kefir, wine and sauerkraut. These LAB strains belong to the species of Streptococcus, Lactobacillus, Pediococcus, Lactococcus and Bifidobacterium (Ramchandran & Shah, 2010; Zhang
et al., 2013; Song, Jeong, Cha, & Baik, 2013; Costa et al., 2010; Prasanna, Bell,
Grandison, & Charalampopoulos, 2012). Among EPS-producing LAB, Lactobacillus
plantarum is famous for its potential probiotic properties and has received
considerable attention in recent years (Zhang et al., 2013; Wang et al., 2014; Wang et 4
Page 4 of 37
al., 2010). It has been shown that the EPS yield, monosaccharide composition and
structure are greatly dependent on the producing microorganisms, their culture
conditions and media compositions (Salazar et al., 2009). L. plantarum C88, an
isolate from fermented dairy tofu, is able to synthesize a high molecular mass capsular
polysaccharide (1.15 × 106 Da) composed of galactose and glucose (1:2) when grown
in a semi-defined medium (Zhang et al., 2013). L. plantarum 70810 isolated from
Chinese Paocai could produce two released EPSs with average molecular weights of
204.6 and 202.8 kDa, and composed of glucose, mannose and galactose with molar
ratios of 18.21:78.76:3.03 and 12.92:30.89:56.19, respectively (Li et al., 2014b).
Besides, L. plantarum KF5 isolated from Tibet Kefir was reported to compose of
mannose, glucose and galactose in an approximate ratio of 1:4.99:6.90 (Wang et al.,
88 89 90 91
EPSs produced by L. plantarum will be increasingly applied in the fermented
Ac ce p
products due to their safe, healthy and desired physicochemical properties in the future, and thus more research is required on the physicochemical, structural and rheological parameters of the EPS. However, so far most of the reports about EPS are focused on the functional characteristics of the EPS-producing L. plantarum strains,
Therefore, in the present study, the EPS from L. plantarum YW11 was prepared and
characterized by gas chromatography–mass spectrometer (GC–MS), nuclear magnetic
resonance (NMR), scanning electron microscopy (SEM), atomic force micrograph
(AFM), thermogram analysis (TGA) and differential scanning calorimeter (DSC) in 5
Page 5 of 37
order to evaluate its potential application in food industry.
2. Materials and methods
2.1. Bacterial strain and media
The EPS-producing strain YW11 used in this study was isolated from Kefir grains
collected from Tibet. The strain was maintained as frozen (−80 °C) stocks in MRS
broth (de Man et al. 1960) supplemented with 20% (v/v) glycerol. The sterile
semi-defined medium (SDM) broth contained (per 1 L): 10 g of bactocasitone (Difco),
5 g of yeast nitrogen base (Difco), 2 g of ammonium citrate, 5 g of sodium acetate,
0.1 g of MgSO4·7H2O, 0.05 g of MnSO4, 2 g of K2HPO4, 20 g of glucose and 1.0 mL
of Tween 80, adjusted to pH 6.6 with 1 M acetic acid (Kimmel & Roberts, 1998).
2.2. Identification of strain YW11
Strain YW11 was primarily identified based on Gram reaction, catalase tests and
cell morphology. Then, the strain was further identified to the species level by API 50
110 111 112
Ac ce p
CHL test (bio-Mérieux, France) and 16S rDNA sequencing analysis. The primers with the
provided by China Agricultural University (Beijing, China). The nucleotide sequences
were compared with standard strains for the sequence similarity through BLAST
Page 6 of 37
2.3. Analysis of bacterial growth and EPS production Strain YW11 was inoculated in 2000-mL Erlenmeyer flasks containing 1000 mL of
SDM broth as mentioned above and incubated at 37 °C. Samples (50 mL) were
withdrawn at different time intervals from 0 to 56 h. The pH was determined with a
pH meter (FE20, Mettler Toledo, Switzerland). The cell viability was determined by
dilution plating with MRS agar medium incubated at 37 °C for 48 h. EPS yield
(expressed as mg L-1) was estimated by phenol-sulfuric acid method using glucose as
a standard (DuBois, Gilles, Hamilton, Rebers, & Smith, 1956).
2.4. Isolation and purification of EPS
The EPS was isolated by using the method of Zhang et al. (2013). The crude EPS
solution (20 mg mL-1, 5mL) was fractionated with an anion exchange chromatography
on a DEAE-cellulose column (26 mm × 40 mm), eluted with deionized water, 0.2 and
0.5 M NaCl solution at a flow rate of 1 mL min-1. Every 5 mL of elution was collected
131 132 133 134
Ac ce p
automatically and the carbohydrate content was determined by phenol-sulfuric acid method. Peak fractions containing polysaccharides were pooled, dialyzed and lyophilized. Further purification of the EPS was performed by a Sepharose CL-6B column (25 mm × 50 mm) and eluted with 0.9% (w/v) NaCl at a flow rate of 0.5 mL min-1. The polysaccharide fractions were detected, pooled, dialyzed and lyophilized.
The purity of the final purified EPS sample was checked. The total carbohydrate
content of the sample was determined by the phenol-sulfuric acid method using
glucose as standard. The total protein content of the sample was determined by the 7
Page 7 of 37
method of binding of Coomassie Brilliant Blue G-250 to protein, using bovine serum
albumin as a standard (Bradford, 1976). The uronic acid content was determined by
the Dische method, using glucuronic acid as standard (Dische, 1947).
The moisture content of the sample was measured by the method of Vijayendra,
Palanivel, Mahadevamma, and Tharanathan (2008). The EPS taken in a dish, which
was previously dried and weighed, was placed along with its lid in an oven
maintained at 105 °C for 5 h and cooled in a desiccator. Drying was repeated till a
constant weight was obtained and the percentage of moisture content was calculated.
2.5. Molecular mass determination of EPS
The molecular mass of the purified EPS was measured by gel-permeation
chromatography (GPC). The GPC system consisted of a Shodex SB-806m-HQ 13 μm,
300 × 8.0 mm column, connected with a SB-G 10 μm, 50 × 6.0 mm guard column.
The EPS were detected using a refractive index detector (RI) (Optilab Wyatt, USA)
152 153 154
Ac ce p
and a multi angle laser-light scattering detector (MALLS) (DAWN HELEOS-II Wyatt, USA), at an internal temperature of 40 °C. The column was eluted with 0.1 M NaNO3 solution at a flow rate of 0.5 mL min-1, and the injection volume of sample was 200
μL, and dn/dc of 0.146 as a refractive index increment was used for polysaccharides
solution (Ai et al. 2008). Data processing was performed with Wyatt Astra software
(Version 184.108.40.206, Wyatt Technology, USA).
2.6. Determination of monosaccharide composition of EPS
Five milligrams of the purified EPS were hydrolyzed with 2 mL (2 M) 8
Page 8 of 37
trifluoroacetic acid (TFA) at 120 °C for 2 h. The hydrolysates were then subjected to
GC-MS analysis. GC-MS was performed on an Agilent 7890A GC fitted with a flame
ionization detector (FID) and a DB-WAX column (30 m length × 0.25 mm inner
diameter × 0.25 µm film thickness; J & W Scientiﬁc, Folsom, CA). The operating
conditions were determined according to Li et al. (2014a). Sugar identification was
done by comparison with standard rhamnose, arabinose, galactose, glucose, mannose
2.7. UV-vis and FTIR spectral analysis of EPS
Ultraviolet-visible (UV-vis) spectroscopy analyses of the EPS were conducted on
UV-vis spectrophotometer (U-3900, Hitachi Ltd., Japan). The EPS solution was
prepared by suspending the sample in distilled water for UV-vis measurement in the
wave-length range of 190-550 nm.
172 173 174 175
The major structural groups of the purified EPS were detected using Fourier
Ac ce p
transform infrared (FTIR) spectroscopy, and the spectrum of the EPS was obtained using a KBr method. The polysaccharide samples were pressed into KBr pellets at sample: KBr ratio 1:100. The FTIR spectra were recorded on a Bruker Tensor 27 instrument (Germany) in the region of 4000-400 cm−1. FTIR spectrum was
determined in transmission mode and the number of scans was 32. The infrared
spectral resolution was 4 cm-1.
2.8. Nuclear magnetic resonance (NMR) spectroscopy analysis of EPS
NMR spectrum of the EPS from strain YW11 were obtained using a Bruker 9
Page 9 of 37
AVANCE 600 MHz spectrometer (Bruker Group, Fällanden, Switzerland) operated at
27 °C with a 5 mm inverse probe. Prior to analysis, samples were exchanged twice in
D2O (99.9 at % D, Cambridge Isotope Laboratories, Inc., Andover, MA, USA) with
intermediate lyophilization, and then dissolved in D2O at concentrations of 5 mg/mL
(for 1 H NMR) and 40 mg/mL (for 13 C NMR). Chemicals shifts (δ) were expressed in
parts per million (ppm). The 2D
decoupling during acquisition of the 1 H FID and used to assign signals.
2.9. Rheological properties of EPS
H–13 C HSQC experiment was recorded with
The rheological behavior of the EPS solutions was carried out in a Brookfield
DV-III ultra programmable rheometer (Brookfield Engineering Laboratories Inc.,
Stoughton, Massachusetts, USA) with a SC4-18 spindle that rotated in chamber
equipped with temperature control system. The lyophilized EPS sample was dissolved
in 0.1 M CaCl2, 0.1 M NaCl solutions, 11% (w/v) skim milk and distilled water at a
194 195 196
Ac ce p
concentration of 2 mg mL-1, respectively. To investigate the effect of pH on the
viscosity of EPS, the pH of EPS solution was adjusted at levels of 4.0, 6.0 and 7.0 by 1 N HCl and NaOH solutions. To investigate the effect of temperature on the viscosity of EPS, the EPS solution was exposed to different temperatures (25 °C, 35 °C and 45
°C). The rheological behavior of the EPS was studied by measuring viscosity as a
function of shear rate from 10 to 300 s−1.
Scanning electron microscopy (SEM) analysis of EPS
The lyophilized samples of the purified EPS (5 mg) were fixed to the SEM stubs 10
Page 10 of 37
with double sided tape, then coated with a layer of gold, ∼10 nm thick. The samples
were observed in a scanning electron microscope (S-4800, Hitachi Ltd., Japan) at an
accelerating voltage of 3.0 kV.
Atomic force micrograph (AFM) analysis of EPS
A stock solution (1 mg mL-1) was prepared by adding the purified EPS into distilled
H2O. The aqueous solution was stirred for about 1 h at 40 °C in a sealed bottle under
N2 stream so that EPS dissolved completely. After cooling to room temperature, the
solution was continuously diluted to the final concentration of 0.01 mg mL-1. About 5
μL of diluted EPS solution was dropped on the surface of a mica sample carrier,
allowed to dry at a room temperature. The AFM images were obtained using a
Dimension® Icon instrument (Bruker Instruments Co., Germany) in tapping mode.
Ac ce p
Thermogram analysis (TGA)
of 100 mL min-1.
213 214 215
The pyrolysis and combustion were carried out in TA SDT-Q600 thermal analyzer
operating at atmospheric pressure. The purified sample of EPS (3 mg) was placed in an Al2O3 crucible and heated at a linear heating rate of 10 °C min-1 over a temperature
range of 25–800 °C. The experiments were performed in air atmosphere at a flow rate
Differential scanning calorimeter (DSC)
The thermal property of the EPS was analyzed using a DSC (DSC Model Q 100, 11
Page 11 of 37
TA instruments). The purified EPS sample (5 mg) was placed in an aluminum pan,
which was sealed and analyzed using empty pan as a reference. The melting point and
enthalpy changes were determined by increasing the heating rate at 10 °C min-1 from
10 to 400 °C (Wang et al., 2010).
3. Results and discussion
3.1. Identification of strain YW11
Strain YW11 was primarily identified as L. plantarum by API 50 CHL test. Further
identification by 16S rRNA sequencing confirmed that the sequence of strain YW11
(Acc. No. KM265361) was identical to those of L. plantarum lp-15 (Acc. No.
FJ763580), L. plantarum LW4 (Acc. No. KJ779096) and L. plantarum TW57-4 (Acc.
No. KJ026699). Therefore, the strain was designated as L. plantarum YW11.
3.2. Bacterial growth and EPS production
233 234 235 236
Change of pH of the medium and EPS production by L. plantarum YW11 during
Ac ce p
growth are shown in Fig. 1. L. plantarum YW11 exhibited a fast growth with a rapid decrease in pH of the medium during the first 8 h of incubation, and the viable cell count reached a maximum of 9.77 log cfu mL-1 at 16 h with pH about 4.0. In the late
stationary phase of growth (after 32 h), the viable count decreased gradually to about
8.60 log cfu mL-1, with a slight decrease in pH to about 3.90 at 56 h. EPS production
by strain YW11 increased rapidly during the initial phase of growth, and continued to
increase to 90 mg L-1 at 56 h. It seemed that the EPS was not degraded during the late
stationary phase of L. plantarum YW11 although the bacterial growth decreased, as 12
Page 12 of 37
previously reported with other EPS-producing L. plantarum strain (Yang et al., 2010).
However, EPS production by L. plantarum C88, L. plantarum 70810, and L.
helveticus MB2-1 decreased after prolonged incubation, probably due to presence of
glycohydrolases in the culture that catalyzed the degradation of polysaccharides
(Zhang et al., 2013; Wang et al., 2014; Li et al., 2014a).
3.3. Isolation and purification of EPS
The crude EPS obtained by ethanol precipitation of the culture supernatant of L.
plantarum YW11 was first separated by anion-exchange chromatography of
DEAE-52. Fractions corresponding to the major peak eluted with 0.2 M NaCl were
found to contain polysaccharides. These fractions that contain the EPS, being an
acidic polysaccharide as it was eluted with NaCl (Gan, Ma, Jiang, Xu, & Zeng, 2011),
were further purified by Sepharose CL-6B gel permeation chromatography. A single
elution peak was generated, and the corresponding fractions were collected and
255 256 257 258
Ac ce p
lyophilized to obtain the purified form of the EPS that was used for the following physicochemical characterization. Analysis of the purified EPS sample showed that it contained 92.35 ± 2.38% of carbohydrate, 2.52 ± 0.12% of moisture, 1.38 ± 0.25% of protein, and 1.56 ± 0.09% of uronic acids. 3.4. Molecular mass and monosaccharide composition of EPS
The molecular mass of the EPS of L. plantarum YW11 was determined by
GPC-MALLS-RI (Fig. 2). The chromatogram of the EPS appeared as a single
symmetrical narrow peak, confirming the homogeneity of the purified EPS sample. 13
Page 13 of 37
The molecular mass of the EPS was determined to be 1.1 × 105 Da (error 3.2%),
which was similar to that (169.6 kDa) of the acidic EPS from L. plantarum 70810
(Wang et al., 2014), lower than that (1.15 × 106 Da) of the neutral EPS of L.
plantarum C88 (Zhang et al., 2013), but higher than that (4.4 × 104 Da) of the EPS of
L. plantarum EP56 (Tallon, Bressollier, & Urdaci, 2003). The polydispersity index of
the EPS of L. plantarum YW11 was determined to be 1.2 (error 2.5%), which was
generally low, indicating no large aggregates formed upon dispersion of this EPS in
aqueous solution. The polydispersity index, as a measure of the width of molecular
mass distribution, was thought to be important due to the relevance and significant
influence of molecular mass distribution on the functional properties of EPS (Zheng et
GC-MS analysis of the monosaccharide composition of the EPS of L. plantarum
YW11 showed that the EPS was composed of glucose and galactose in a molar ratio
276 277 278
Ac ce p
of 2.71:1. Previously, the EPS from L. plantarum C88 was shown to contain glucose
and galactose, but in different molar ratio (Zhang et al., 2013). Other L. plantarum strains, e.g. both EP56 and KF5 produced EPSs consisted of glucose and galactose, and additionally N-acetylgalactosamine (Tallon et al., 2003) and mannose (Wang et
al., 2010), respectively. The monosaccharide composition of EPS produced by LAB
can be affected by the type of strains, culture conditions and medium compositions
(Wang et al., 2014).
3.5. Spectra analysis of the EPS functional groups 14
Page 14 of 37
The UV-vis spectrum of the purified EPS from strain YW11 showed no absorption
at 260 nm or 280 nm, indicating no nucleic acid present in the EPS sample. The FTIR
spectrum (Fig. 3) of the EPS indicated that the polysaccharide contained a significant
number of hydroxyl groups as it displayed a broad and intense stretching peak around
3408 cm-1 (Wang et al., 2010). The stretching band around 2933 cm-1 was due to C–H
stretching vibration (Melo, Feitosa, Freitas, & de Paula, 2002). The absorptions at
1726 cm-1 and 1646 cm-1 were due to the stretch vibration of C=O bond (Ye et al.,
2014), and a peak at 1550 cm-1 could be assigned to N–H bending of amides II of
protein (Lin et al., 2005). The absorption at 1384 cm-1 was possibly due to symmetric
CH3 bending (Pan & Mei, 2010). The bands within the 900–1150 cm-1 region were
attributed to the vibration of C-O-C bond (Ye et al., 2014).
3.6. NMR analysis of EPS
The 1 H NMR and 13 C NMR spectroscopy of the EPS from strain YW11 are shown
297 298 299
Ac ce p
in Fig. 4. Three major chemical shift signals in the anomeric region (δ 4.8–5.5 ppm) were found at δ 5.22, 5.07 and 4.97 ppm in 1 H NMR (Fig. 4A). This indicates that the
EPS from strain YW11 mainly consists of three monosaccharide residues. Based on their chemical shift and the value of the coupling constant for anomeric signals in the
carbons regions (95–110 ppm) and ring carbons (50–85 ppm) regions. Based on the
data reported previously (Li et al., 2014; Wang at al., 2014), the three main signals in
H NMR spectrum, three signals were corresponds to C-1 of α-type configurations. 13
C NMR spectrum (Fig. 4B) of EPS from strain YW11 included anomeric
Page 15 of 37
the anomeric carbons regions at δ 104.58, 104.05 and 101.21 ppm were corresponded
to the α-linkages. Although no aminosugars were detected by monosaccharide
analysis of the EPS as described above, the
signals between 50-60 ppm, and several signals at 22.85, 174.82, 175.30 ppm that
were probably from C-2 and N-acetyl group of the aminosugar, respectively,
indicating possible presence of this sugar in the EPS of strain YW11.
C NMR spectrum demonstrated two
The single-bond correlations between the protons and the corresponding carbons
obtained from 2D 1 H–13 C HSQC spectrum (Fig. 4C) of the EPS from strain YW11
in D2O demonstrated three crosspeaks in the anomeric region resulted from cross link
of the C-1 signals at δ 104.58, 104.05 and 101.21 ppm to the proton signals at δ 5.22,
5.07 and 4.97 ppm, respectively, confirming presence of three monosaccharide
residues in the repeating unit of the polysaccharide. Detailed chemical structure of the
EPS from strain YW11 needs to be further studied.
318 319 320
Ac ce p
3.7. Rheological properties of EPS The rheological behavior of the EPS from strain YW11 was studied in water, skim
milk, different salt solutions (NaCl and CaCl2), and at different pHs (4, 6 and 7) and temperatures (25 °C, 35 °C and 45 °C) (Fig. 5). All the EPS solutions showed a shear
thinning behavior, a decrease of viscosity with increasing shear rates that was caused
mainly by breakdown of structural units in the EPS by hydrodynamic forces generated
during shear (Kavita, Singh, Mishra, & Jha, 2014). This property was considered to be
important for yielding desired sensory properties such as mouth-feel and flavor 16
Page 16 of 37
release properties, as well as some processing operations such as stirring, pouring,
pumping, spray drying (Zhou et al., 2014). The EPS in skim milk was shown to be
more viscous than in water over the whole shear rate range (Fig. 5A). However, no
obvious difference in viscosity was observed with the EPS in 0.1 M NaCl solution or
in 0.1 M CaCl2 solution at all the shear rates tested (Fig. 5B). The higher viscosity of
the EPS solution at acidic pHs (4, 6) than at a neutral pH (7.0) (Fig. 5C) would be
beneficial to the use of this EPS-producing strain to improve texture of fermented
milk that usually has pH values between 4.0 and 4.5. Ahmed, Wang, Anjum, Ahmad,
and Khan (2013) also reported that an EPS produced by L. kefiranofaciens ZW3 was
more viscous at acidic pHs than alkaline pHs. Studies on the effect of temperature
(Fig. 5D) showed that increasing temperature from 25 °C to 35 °C did not affect the
viscosity of the EPS solution over the whole shear rate range, but further increasing to
45 °C resulted in decreased viscosity. Similarly, an acidic EPS produced by
339 340 341
Ac ce p
Streptococcus phocae PI80 had decreased viscosity with increasing temperature from 25 °C to 45 °C (Kanmani et al. 2011). This decrease in viscosity of EPS solution might be attributed to the decreased interactions between the EPS molecules when temperature was increased, thus leading to a loosened polymer structure. Therefore,
the rheological behavior of the EPS produced by L. plantarum YW11 as described
above would make it particular suitable as a potential stabilizer used in dairy foods.
3.8. Scanning electron microscopic (SEM) analysis of EPS
The microstructures of the EPS from strain YW11 and xanthan gum as a reference 17
Page 17 of 37
material are represented by SEM (Fig. 6). The EPS showed a relatively stable three
dimensional structure that appeared to be a porous web (Fig. 6A), whereas the
xanthan gum showed a dispersive structure with irregular lumps of different size (Fig.
6C). At a higher magnification (Fig. 6B and 6D), additional details of the
microstructure of the EPS and xanthan gum were visible. The EPS had a smooth and
glittering surface, but xanthan gum presented a coarse surface. Similar porous
structures were also reported with the purified EPSs of Bifidobacterium longum subsp.
infantis CCUG 52486 and Bifidobacterium infantis NCIMB 702205 (Prasanna, Bell,
Grandisona, & Charalampopoulos, 2012). However, Wang et al. (2010) observed a
sheet-like compact morphology of the EPS from L. plantarum KF5. The highly
branched and porous structure of the EPS from strain YW11 as observed in this study
may facilitate its application in foods to improve the physical properties, e.g. viscosity,
water holding capacity of the product.
360 361 362
Ac ce p
3.9. Atomic force micrograph (AFM) analysis of EPS In recent years exopolysaccharide has been studied extensively by using atomic
force microscopy that provides a powerful tool to characterize the morphological features of polymers (Admed et al., 2013). Images of the strain YW11 EPS deposited
from 0.01 mg mL-1 aqueous solution were obtained by AFM (Fig. 7). There was
presence of many spherical lumps with the height ranged from 3.0 nm to 13.5 nm.
Some regions formed fibrous network, whereas other regions were relatively sparse,
suggesting that the structure of the EPS might be tangled networks, which was in 18
Page 18 of 37
agreement with its fibrous morphology in dry solid form. This structural property of
the EPS was also observed earlier with another viscous polysaccharide from Mesona
blumes gum that had irregular worm-shaped structure (Feng, Gu, Jin, & Zhuang,
2008). The viscous nature of the EPS from strain YW11 described above might also
be caused by its capability of binding with water in the aqueous solution.
Thermogravimetric analysis (TGA)
The thermogravimetric analysis (TGA) of the EPS from L. plantarum YW11 was
carried out dynamically between weight loss vs temperatures. As shown in Fig. 8A,
the EPS showed an initial weight loss of approximately 11.64% between about 25 and
100 °C. This initial weight loss may be associated with the loss of moisture (Wang et
al., 2010), suggesting high content of carboxyl groups in the EPS molecules. Kumar,
Joo, Choi, Koo, and Chang (2004) reported that the high level of carboxyl group in
the EPS increased the degradation of the first phase as carboxyl groups were bound to
381 382 383
Ac ce p
more water molecules. A degradation temperature (Td) of 287.7 °C and a large weight loss of approximately 69.93 % could be observed from the differential thermogravimetric mass loss spectrum (Fig. 8B) and thermogravimetric mass loss spectrum (Fig. 8A) for strain YW11 EPS between about 200 and 300 °C. Then the
weight loss gradually decreased to leave a final residue of ca. 16.07% of the total EPS.
Ahmed, Wang, Anjum, Ahmad, and Khan (2013) and Wang et al. (2010) showed that
the degradation temperatures of ZW3 EPS, KF5 EPS, xanthan and locust gum were
299.62 °C, 278.53 °C, 282.65 °C and 278.46 °C, respectively. The degradation 19
Page 19 of 37
temperature of the EPS from strain YW11 in the present study was within the range of
the KF5 EPS and ZW3 EPS, but slightly higher than xanthan gum and locust gum.
The different thermal stability and degradation behavior of these EPSs probably
attributed to their different carbohydrate compositions. Due to the relatively higher
degradation temperature of the EPS from strain YW11, it would be safe for use of this
EPS in dairy industry where in most of the processes temperature seldom overpasses
Commercial application of an EPS is crucially dependent on its thermal and
rheological behavior. DSC analysis of the EPS from strain YW11 showed an
exothermic peak with heat flow from 10 °C to 400 °C (Fig. 9). The melting point of
the EPS exothermic peak started at about 143.6 °C, and the enthalpy change (△H)
Ac ce p
phocae PI80 endothermic peak started at 120.09 °C and the enthalpy change was
about 404.6 J g-1. Thus the EPS from strain YW11 showed a different thermal
behaviour from these EPSs.
400 401 402
needed to melt 1 g of EPS was 217.8 J. Different results were reported previously for different EPSs. Wang et al. (2010) showed that the melting point and enthalpy change of KF5 EPS isolated from L. plantarum KF5 were 88.35 °C and 133.5 J g-1,
respectively. Kanmani et al (2011) reported that the melting point of the EPS from S.
Page 20 of 37
Conclusion In this study, an EPS-producing strain YW11 isolated from Tibet Kefir was
identified as L. plantarum. The EPS from YW11 was composed of glucose and
galactose in a molar ratio of 2.71:1, with possible presence of N-acetylated sugar
residues in the polysaccharide. The EPS had a molecular mass of 1.1 × 105 Da. It
exhibited a shear thinning behavior under different conditions and had higher
viscosity in skim milk, at lower temperature and acidic pHs. The SEM images of the
EPS demonstrated a porous structure that was highly branched, while the AFM
images of the EPS revealed presence of many spherical lumps. Furthermore, the EPS
had a higher degradation temperature. These characteristics of the EPS produced by L.
plantarum YW11 would make it a promising candidate for its exploitation in food
421 422 423
Ac ce p
This work was financially supported by The Key Project of the Educational
Committee of Beijing City (KZ201310011011), Natural Science Foundation of China (31371804), and National Public Beneﬁt Research (Agriculture) Foundation (201303085).
Ai, L., Zhang, H., Guo, B., Chen, W., Wu, Z., & Wu, Y. (2008). Preparation, partial
characterization and bioactivity of exopolysaccharides from Lactobacillus casei
LC2W. Carbohydrate Polymers, 74, 353‒357. 21
Page 21 of 37
Amjres, H., Béjar, V., Quesada, E., Carranza, D., Abrini, J., Sinquin, C., et al. (2014).
Characterization of haloglycan, an exopolysaccharide produced by Halomonas
stenophila HK30. International Journal of Biological Macromolecules, 72, 117‒
Ahmed, Z., Wang, Y., Anjum, N., Ahmad, A., & Khan, S. T. (2013). Characterization
of exopolysaccharide produced by Lactobacillus kefiranofaciens ZW3 isolated
from Tibet kefire – Part II. Food Hydrocolloids, 30, 343–350.
Bradford, M. M. (1976). A rapid and sensitive method for the quantification of
microgram quantities of protein utilizing the principle of protein–dye binding.
Analytical Biochemistry, 72, 248–252.
Costa, N. E., Hannon, J. A., Guinee, T. P., Auty, M. A. E., McSweeney, P. L. H., &
Beresford, T. P. (2010). Effect of exopolysaccharide produced by isogenic strains
of Lactococcus lactis on half-fat Cheddar cheese. Journal of Dairy Science, 93,
de los Reyes-Gavilán, C. G., Suárez, A., Fernández-García, M., Margolles, A.,
Gueimonde, M., & Ruas-Madiedo, P. (2011). Adhesion of bile-adapted
Bifidobacterium strains to the HT29-MTX cell line is modified after sequential
446 447 448 449 450
Ac ce p
gastrointestinal challenge simulated in vitro using human gastric and duodenal
juices. Research in Microbiology, 162, 514–519.
de Man, J. C., Rogosa, M., & Sharpe, M. E. (1960). A medium for the cultivation of lactobacilli. Journal of Applied Bacteriology, 23, 130–135.
Dische, Z. (1947). A new specific colour reaction of hexuronic acid. The Journal of Biological Chemistry, 167, 189–198.
DuBois, M., Gilles, K. A., Hamilton, J. K., Rebers, P. A., & Smith, F. (1956).
Colorimetric method for determination of sugars and related substances.
Analytical Chemistry, 28, 350–356.
Feng, T., Gu, Z., Jin, Z., & Zhuang, H. (2008). Isolation and characterization of an 22
Page 22 of 37
acidic polysaccharide from Mesona Blumesgum. Carbohydrate Polymers, 71,
159–169. Gan, D., Ma, L., Jiang, C., Xu, R., & Zeng, X. (2011). Production, preliminary
characterization and antitumor activity in vitro of polysaccharides from the
mycelium of Pholiota dinghuensis Bi. Carbohydrate Polymers, 84, 997–1003.
Ismail, B., & Nampoothiri, K. M. (2010). Production, purification and structural
characterization of an exopolysaccharide produced by a probiotic Lactobacillus
plantarum MTCC 9510. Archives of Microbiology, 192, 1049–1057.
Kanmani, P., Kumar, R. S., Yuvaraj, N., Paari, K. A., Pattukumar, V., & Arul, V.
(2011). Production and purification of a novel exopolysaccharide from lactic acid
bacterium Streptococcus phocae PI80 and its functional characteristics activity in
vitro. Bioresource Technology, 102, 4827–4833.
Kavita, K., Singh, V. K., Mishra, A., & Jha, B. (2014). Characterisation and
antibiofilm activity of extracellular polymeric substances from Oceanobacillus
iheyensis. Carbohydrate Polymers, 101, 29–35.
Kimmel, S. A., & Roberts, R. F. (1998). Development of a growth medium suitable
for exopolysaccharide production by Lactobacillus delbrueckii ssp. bulgaricus
RR. International Journal of Food Microbiology, 40, 87–92.
474 475 476 477 478
Ac ce p
Kumar, C. G., Joo, H. S., Choi, J. W., Koo, Y. M., & Chang, C. S. (2004). Purification and characterization of an extracellular polysaccharide from haloalkalophilic Bacillus sp. I-450. Enzyme and Microbial Technology, 34, 673–681.
Liu, C., Chu, F., Chou, C., & Yu, R. (2011). Antiproliferative and anticytotoxic effects of cell fractions and exopolysaccharides from Lactobacillus casei 01. Mutation Research, 721, 157–162.
Li, C., Li, W., Chen, X., Feng, M., Rui, X., Jiang, M., et al. (2014b). Microbiological,
physicochemical and rheological properties of fermented soymilk produced with
exopolysaccharide (EPS) producing lactic acid bacteria strains. LWT-Food 23
Page 23 of 37
Science and Technology, 57, 477–485. Lindström, C., Holst, O., Nilsson, L., Öste, R., & Andersson, K. E. (2012). Effects of
Pediococcus parvulus 2.6 and its exopolysaccharide on plasma cholesterol levels
and inflammatory markers in mice. AMB Express, 2, 66–69.
Lin, M. S., Al-Holy, M., Chang, S. S., Huang, Y., Cavinato, A. G., Kang, D. H., et al.
(2005). Rapid discrimination of Alicyclobacillus strains in apple juice by Fourier
transform infrared spectroscopy. International Journal of Food Microbiology,
Liu, C., Lu, J., Lu, L., Liu, Y., Wang, F., & Xiao, M. (2010). Isolation, structural
characterization and immunological activity of an exopolysaccharide produced
by Bacillus licheniformis 8–37-0–1. Bioresource Technology, 101, 5528–5533.
Li, W., Ji, J., Chen, X., Jiang, M., Rui, X., & Dong, M. (2014a). Structural elucidation
and antioxidant activities of exopolysaccharides from Lactobacillus helveticus
MB2-1. Carbohydrate Polymers, 102, 351–359.
Li, Wei, Ji, J., Tang, W., Rui, X., Chen, X., Jiang, M., & Dong, M. (2014).
Characterization of an antiproliferative exopolysaccharide (LHEPS-2) from
Lactobacillus helveticus MB2-1. Carbohydrate Polymers, 105, 334–340.
500 501 502 503 504 505 506
London, L. E. E., Price, N. P. J., Ryan, P., Wang, L., Auty, M. A. E., Fitzgerald, G. F.,
Ac ce p
et al. (2014). Characterization of a bovine isolate Lactobacillus mucosae DPC 6426 which produces an exopolysaccharide composed predominantly of mannose residues. Journal of Applied Microbiology, 117, 2, 509–517.
Melo, M. R. S., Feitosa, J. P. A., Freitas, A. L. P., & de Paula, R. C. M. (2002). Isolation and characterization of soluble sulfated polysaccharide from the red seaweed Gracilaria cornea. Carbohydrate Polymers, 49, 491–498. Nikolic, M., López, P., Strahinic, I., Suárez, A., Kojic, M., Fernández-García, M., et al.
Lactobacillus paraplantarum BGCG11 and its non-EPS producing derivative 24
Page 24 of 37
strains as potential probiotics. International Journal of Food Microbiology, 158,
Pan, D., & Mei, X. (2010). Antioxidant activity of an exopolysaccharide purified from Lactococcus lactis subsp. lactis 12. Carbohydrate Polymers, 80, 908–914.
Prasanna, P. H. P., Bell, A., Grandison, A. S., & Charalampopoulos, D. (2012).
Emulsifying, rheological and physicochemical properties of exopolysaccharide
produced by Bifidobacterium longum subsp. infantis CCUG 52486 and
Bifidobacterium infantis NCIMB 702205. Carbohydrate Polymers, 90, 533–540.
Ramchandran, L., & Shah, N. P. (2010). Characterization of functional, biochemical
and textural properties of synbiotic low-fat fermented milks during refrigerated
storage. LWT-Food Science and Technology, 5, 819–827.
Salazar, N., Prieto, A., Leal, J. A., Mayo, B., Bada-Gancedo, J. C., de los
Reyes-Gavilán, C. G., et al. (2009). Production of exopolysaccharides by
Lactobacillusand Bifidobacterium strains of human origin, and metabolic
activity of the producing bacteria in milk. Journal of Dairy Science, 92,
Sandal, I., Inzana, T., Molinaro, A., De Castro, C., Shao, J., Apicella, M., et al. (2011).
Identification, structure, and characterization of an exopolysaccharide produced
527 528 529 530 531
Ac ce p
by Histophilus somni during biofilm formation. BMC Microbiology, 11, 186.
Shang, N., Xu, R., & Li, P. (2013). Structure characterization of an exopolysaccharide produced by Bifidobacterium animalis RH. Carbohydrate Polymers, 91, 128–134.
Song, Y. R., Jeong, D. Y., Cha, Y. S., & Baik, S. H. (2013). Exopolysaccharide
produced by Pediococcus acidilactici M76 isolated from the Korean traditional
rice wine, Makgeolli. Journal of Microbiology and Biotechnology, 23, 681–688.
Page 25 of 37
Tallon, R., Bressollier, P., & Urdaci, M. C. (2003). Isolation and characterization of
two exopolysaccharides produced by Lactobacillus plantarum EP56, Research in
Microbiology, 154, 705–712. Vijayendra, S. V. N., Palanivel, G., Mahadevamma, S., & Tharanathan, R. N. (2008).
Physico-chemical characterization of an exopolysaccharide produced by a
nonropy strain of Leuconostoc sp. CFR 2181 isolated from dahi, an Indian
traditional lactic fermented milk product. Carbohydrate Polymers, 72, 300–307.
Wang, K., Li, W., Rui, X., Chen, X., Jiang, M., & Dong, M. (2014). Characterization
of a novel exopolysaccharide with antitumor activity from Lactobacillus
plantarum 70810. International Journal of Biological Macromolecules, 63,
Wang, Y., Li, C., Liu, P., Ahmed, Z., Xiao, P., & Bai, X. (2010). Physical
characterization of exopolysaccharide produced by Lactobacillus plantarum KF5
isolated from Tibet Kefir. Carbohydrate Polymers, 82, 895–903.
Yang, Z., Li, S., Zhang, X., Zeng, X., Li, D., Zhao, Y., et al. (2010). Capsular and
slime-polysaccharide production by Lactobacillus rhamnosus JAAS8 isolated
from Chinese sauerkraut: Potential application in fermented milk products.
Journal of Bioscience and Bioengineering, 110, 53–57.
553 554 555 556 557
Ac ce p
Ye, S., Liu, F., Wang, J., Wang, H., & Zhang, M. (2012). Antioxidant activities of an exopolysaccharide isolated and purified from marine Pseudomonas PF-6. Carbohydrate Polymers, 87, 764–770.
Ye, S., Zhang, M., Yang, H., Wang, H., Xiao, S., Liu, Y., et al. (2014). Biosorption of Cu2+, Pb2+ and Cr6+ by a novel exopolysaccharide from Arthrobacter ps-5. Carbohydrate Polymers, 101, 50–56.
Zhang, L., Liu, C., Li, D., Zhao, Y., Zhang, X., & Zeng, X. (2013). Antioxidant
activity of an exopolysaccharide isolated from Lactobacillus plantarum C88.
International Journal of Biological Macromolecules, 54, 270–275. 26
Page 26 of 37
Zheng, J., Wang, J., Shi, C., Mao, D., He, P., & Xu, C. (2014). Characterization and
antioxidant activity for exopolysaccharide from submerged culture of Boletus
aereus. Process Biochemistry, 49, 1047–1053. Zhou, F., Wu, Z., Chen, C., Han, J., Ai, L., & Guo, B. (2014). Exopolysaccharides
produced by Rhizobium radiobacter S10 in whey and their rheological properties.
Food Hydrocolloids, 36, 362–368.
570 Figure captions
Figure 1. Kinetics of growth and EPS production by L. plantarum YW11 in a
semi-defined medium at 37 °C showing changes of the bacterial cell counts, EPS
yield and pH of the medium during the incubation. Each value represents the average
of triplicate measurements.
Figure 2. GPC-MALLS-RI chromatogram of EPS from L. plantarum YW11.
Ac ce p
purified EPS from L. plantarum YW11 recorded on Bruker AVANCE 600 MHz
spectrometer in D2O.
Figure 5. Rheological behavior of the EPS from L. plantarum YW11 in distilled
577 578 579
Figure 3. UV spectrum of the purified EPS from L. plantarum YW11 in the range of
190-550 nm (A); FTIR spectrum of the purified EPS in the range of 400-4000 cm-1 (B).
Figure 4. The
H NMR (A), 13 C (B) and
H–13 C HSQC (C) NMR spectra of the
Page 27 of 37
water and skim milk (11%, w/v) (A), in 0.1 M NaCl and CaCl2 (B), at different pH
values (4, 6 and 7) (C), and at different temperatures (25, 35 and 45 °C) (D).
Figure 6. Scanning electron micrograph images of the purified EPS from L.
plantarum YW11 (A: 1000 ×; B: 5000 ×) as compared to a reference material xanthan
gum (C: 1000 ×; D: 5000 ×).
Figure 7. Atomic force microscopy images of the purified EPS from L. plantarum
YW11 (A) planar, (B) cubic.
Figure 8. Thermogravimetric mass loss spectrum (A, left hand ordinate ordinate axis),
and differential thermogravimetric mass loss spectrum (B, right hand ordinate
ordinate axis) of the purified EPS from L. plantarum YW11.
Figure 9. DSC thermogram of the purified EPS from L. plantarum YW11.
Ac ce p
Page 28 of 37
Page 29 of 37
Page 30 of 37
Page 31 of 37
Ac ce p
Page 32 of 37
Page 33 of 37
Page 34 of 37
Page 35 of 37
Page 36 of 37
Page 37 of 37