Effect of Extraction Temperature on the Diffusion ... - Springer Link

Biotechnology and Bioprocess Engineering 19: 369-377 (2014) DOI 10.1007/s12257-013-0733-2

RESEARCH PAPER

Effect of Extraction Temperature on the Diffusion Coefficient of Polysaccharides from Spirulina and the Optimal Separation Method Ratana Chaiklahan, Nattayaporn Chirasuwan, Panya Triratana, Suvit Tia, and Boosya Bunnag

Received: 13 November 2013 / Revised: 9 February 2014 / Accepted: 15 February 2014 © The Korean Society for Biotechnology and Bioengineering and Springer 2014

Abstract The extraction temperature had a significant impact on the concentration of polysaccharides derived from solid-liquid extraction of Spirulina. The polysaccharide concentration was significantly higher when the extraction was performed at 90°C than when it was performed at 80, 70, and 50°C. This result is related to the diffusion coefficients of the polysaccharides, which increased from 1.07 × 10-12 at 50°C to 3.02 × 10-12 m2/sec at 90°C. Using the Arrhenius equation, the pre-exponential factor (D0) and the activation energy (Ea) for Spirulina polysaccharide extraction were calculated as 7.958 × 10-9 m2/sec and 24.0 kJ/mol, respectively. Among the methods used for the separation of Spirulina polysaccharides, cetyltrimethylammonium bromide (CTAB, method I) and organic solvent (ethanol, in methods II and III) provided similar yields of polysaccharides. However, the separation of polysaccharides using an ultrafiltration (UF) process (method III) and ethanol precipitation was superior to separation via CTAB or vacuum rotary evaporation (method II). The use of a membrane with a molecular weight cut-off (MWCO) of 30 kDa and an area of 0.01 m2 at a feed pressure of 103 kPa with a mean permeate flux of 39.3 L/m2/h and a retention rate of 95% was optimal for the UF process. The Ratana Chaiklahan*, Nattayaporn Chirasuwan, Panya Triratana, Boosya Bunnag Pilot Plant Development and Training Institute, King Mongkut’s University of Technology Thonburi, Bang Khun Thian, Bangkok 101-50, Thailand Tel: +66-2470-7483; Fax: +66-2452-3455 E-mail: [email protected]/[email protected] Suvit Tia Department of Chemical Engineering, King Mongkut’s University of Technology Thonburi, Thung Khru, Bangkok 101-40, Thailand Boosya Bunnag School of Bioresources and Technology, King Mongkut’s University of Technology Thonburi, Bang Khun Thian, Bangkok 101-50, Thailand

addition of two volumes (v/v) of ethanol, which gave a total polysaccharide content of approximately 4% dry weight, was found to be most suitable for polysaccharide precipitation. The results of a Sepharose 6B column separation showed that the molecular weights of the polysaccharides in fractions I and II were 212 and 12.6 kDa, respectively. Keywords: polysaccharide, Spirulina, temperature, diffusion coefficients, ultrafiltration

1. Introduction Polysaccharides are used world-wide as food and cosmetic ingredients and in nutraceutical and pharmacological applications. Currently, polysaccharides are produced from plants, seaweed, mushrooms and microalgae such as Chlorella and the cyanobacterium Spirulina and support economically important global industries [1,2] Spirulina can produce a large number of valuable compounds, such as phycocyanin, carotenoids and lipids, which contain essential fatty acids (linoleic acid and gamma-linolenic acid). Spirulina also contain 15 ~ 20% carbohydrate by dry weight, the majority of which is polysaccharide [3,4]. Polysaccharides from Spirulina have a variety of biological activities, including antioxidant, antiherpes simplex virus type 1 (HSV-1) and immunostimulatory activities, as well as inhibitory effects on corneal neovascularization and anticoagulant activity mediated by heparin cofactor II [5-8]. Hot water extracts of Spirulina consist of sulfated polysaccharides called calcium spirulan [9]. Sulfated polysaccharides consist of rhamnose, 3-Omethylrhamnose (acofriose), 2,3-di-O-methylrhamnose, 3O-methylxylose, uronic acids and sulfate. The sulfate

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Biotechnology and Bioprocess Engineering 19: 369-377 (2014)

groups are substituted at the C-2 or C-4 position of 1,3linked rhamnose and at the C-4 position of 1,2-linked 3-Omethylrhamnose (acofriose) [10,11]. Sulfate groups in polysaccharides have been found to be important for biological activities [5,11]. Extraction, separation and purification processes all play important roles in optimizing the yield of polysaccharides. The solid-liquid extraction process involves the use of a solvent to dissolve a soluble fraction from an insoluble, permeable solid [12] and is affected by the transfer of solutes from inside the solid matrix to the solvent. During extraction, the concentration of the solute inside the solid varies, leading to variable conditions that affect the rate, which can be expressed in terms of (mass of solute leached)/unit time or as the change in solute concentration in the solid/unit time (dc/dt or dx/dt) [13]. The extraction equilibrium is described by the equilibrium distribution constant or partition coefficient, which is a function of the concentration at equilibrium in the extract and in the residual material. This relationship can be described as follows [12]: K = Ceq / Cdm

(1)

where K is the equilibrium distribution constant, Ceq is the concentration of a given compound in the solvent at equilibrium and Cdm is the concentration of a given compound in the dry pomace at equilibrium. For larger values of K, more of a given compound will dissolve in the solvent. In addition, the amount of a compound that will dissolve is a function of the characteristics of both the solvent used and the temperature of the solvent [12,14]. Temperature affects the equilibrium and the mass transfer rate of the extraction process [12]. An increase in the working temperature favours extraction, enhancing both the solubility of the solute and the diffusion coefficient (D) as established by the Einstein equation: D ∝ (T/η)

(2)

where T is the absolute temperature and η is the dynamic viscosity coefficient [14,15]. Abdel-Kader [16] reported that the D value for ascorbic acid increased from 0.94 × 10-8 m2/sec at 50°C to 1.94 × 10-8 m2/sec at 90°C. In addition to the extraction process, many previously reported methods have been used to separate polysaccharides, including precipitating the polysaccharides in the supernatant by cetyltrimethyl ammonium bromide (CTAB) [17-19], decreasing the volume of the polysaccharide supernatant using a rotary evaporator or heat followed by precipitation by ethanol [2,20-22], and using ultrafiltration (UF) to separate and purify the polysaccharides. Sheng et al. [23] reported that the separation of polysaccharides from Chlorella pyrenoidosa by a membrane with a molecular

weight cut-off (MWCO) of 30 kDa produced the highest yield. In addition, Challouf et al. [24] and Majdoub et al. [8] used membranes with MWCOs of 30 and 100 kDa, respectively, to separate and purify exopolysaccharides from the culture medium of Arthrospira (Spirulina) platensis. According to a previous study [25], the results showed that the extraction temperature plays an important role in determining the yield of polysaccharides from Spirulina. Therefore, this work sought to investigate the influence of the extraction temperature on the diffusion coefficient of polysaccharides and to determine suitable methods for separating the polysaccharides that could be applied to large-scale production.

2. Materials and Methods 2.1. Materials and extraction conditions Dried Spirulina biomass was provided by Nathong Spirulina Group, Chachoengsao, Thailand. It was extracted three times for 20 min with ethanol at a ratio of 1:5 (w/v) at 60°C [26]. The defatted/depigmented residue was collected by centrifugation and dried. To determine the effect of the extraction temperature on the diffusion coefficient, polysaccharide extractions were carried out under the conditions stipulated by Chaiklahan et al. [25]. Briefly, the extractions were performed at four different temperatures (50, 70, 80, and 90°C) using water as the solvent at a solid-to-liquid ratio of 1:35. The extraction time ranged from 0 to 360 min. After centrifugation at 4,800 × g for 10 min, the supernatant containing the polysaccharides was obtained. 2.2. Determination of the effective diffusion coefficient of the polysaccharides Many reports have shown that polysaccharide extraction from plants and algae is generally controlled by internal diffusion, and solutions of Fick’s second law, given in Eq. (3), were used to determine the diffusion coefficient of the polysaccharide at each temperature. ∂ C/∂ t = D (∂ 2C/∂ r2)

(3)

In general, the dimensionless extract concentration of the solute is defined as the ratio of (C − Ceq)/(C0 − Ceq) [14,27], where C0 and Ceq are the initial and equilibrium concentrations (mg/mL), respectively, of the total sugars at each temperature. The following are the initial and boundary conditions: C = C0 C = C or Cinterface ∂ C/∂ r = 0

of particle

for t = 0 and 0 < r < R, for t > 0 and r = R, and for t > 0 and r = 0

Effect of Extraction Temperature on the Diffusion Coefficient of Polysaccharides from Spirulina …

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The analytical solution for the liquid diffusion equation is represented by the following equations of Crank [27]: ∞

(C − Ceq)/(C0 − Ceq) = (6/π 2) Σn=1((1/n2)exp(−n2π 2Dt/r2)) (4) ln [(C − Ceq)/(C0 − Ceq)] = ln (6/π 2) − π2Dt/r2

(5)

ln [(C − Ceq)/(C0 − Ceq)] = a − bt

(6)

When the logarithm ln (C − Ceq)/(C0 − Ceq) is plotted against time, a straight line should be obtained, and the diffusivity or diffusion coefficient of the polysaccharide can be accessed from its slope b (π 2D/r2). 2.3. Determination of a suitable method for separating the polysaccharide The optimum conditions for polysaccharide extraction from Spirulina are reported by Chaiklahan et al. to comprise a solid-to-liquid ratio of 1:45 (w/v), an extraction temperature of 90°C and an extraction time of 120 min [25]. Thus, the polysaccharide extraction in this study was performed under these conditions in order to determine a suitable method for separating the polysaccharides. After extraction and centrifugation at 4,800 × g for 10 min, the polysaccharide solution was collected. Then, the polysaccharides were separated using three different methods, according to the schematic diagram in Fig. 4. In method I, the polysaccharides (1.2 L) were precipitated by the addition of 1% cetyltrimethyl ammonium bromide (CTAB) solution to a final concentration of 15% (v/v). Then, the precipitate was collected by centrifugation and washed stepwise with saturated sodium acetate in 95% ethanol and absolute ethanol. In method II, the polysaccharide solution (1.2 L) was concentrated in a vacuum rotary evaporator at 55°C whereas in method III, it was concentrated using a UF with a MWCO of 30 kDa [23,24]. When the volume of the polysaccharide solution had decreased to approximately one-fifth of its original volume, ethanol (3 volumes) [28] was added to the concentrated polysaccharide solutions from methods II and III, and precipitation was allowed to proceed overnight at 4°C. The precipitates were collected by centrifugation (10,000 × g, 10 min) and lyophilized. The optimal concentration of ethanol for precipitating the polysaccharides was also determined. After the polysaccharide solution was concentrated to approximately one-fifth of its original volume by UF, ethanol was added to different final concentrations, and the precipitates were separated by centrifugation and lyophilized. The yield of the crude polysaccharide extract (as a percentage of dry weight) was calculated using the equation given below:

Fig. 1. Polysaccharide concentration (A), ln (C − Ceq)/(C0 − Ceq), and extraction time (B) at different temperatures.

Yield (% dry weight) = [weight of polysaccharides / dried sample weight] × 100 (7) 2.4. Separation of polysaccharides by ultra filters with different cut-offs To determine the optimal conditions for the membrane concentration process, the effects of MWCO, effective filtration area and feed pressure were investigated. Crude polysaccharide extract (1.2 L) was fed through membranes of 10, 30, 50, 70, and 100 kDa MWCO (Minimate Tangential Flow Filtration Capsules; Pall Corporation, USA) with areas of 0.005 m2 at a 69 kPa feed pressure to determine the optimum MWCO. After the optimum MWCO was selected, two effective filtration areas (0.005 and 0.01 m2) and two feed pressures (69 and 103 kPa) were tested. The yield, mean permeate flux and retention rate were used as evaluation indexes. The mean permeate flux was defined as the permeate volume flowing through a membrane in one unit of time and filtration area. The retention rate of the polysaccharides was determined using

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the following equation: Retention rate (%) = 100 [1 − total sugar in permeate / total sugar in retentate] (8) 2.5. Analysis A modified method of Dubois et al. [29] was used to determine the total sugar content. Each sample was analyzed by the phenol-sulfuric acid reaction using distilled water and glucose as blank and standard solutions, respectively, and the absorbance was subsequently determined at 490 nm. The average molecular weight of the polysaccharides was estimated using a Sepharose 6B gel filtration column eluted with 0.01 M sodium acetate buffer containing 0.025 M sodium chloride at a flow rate of 0.7 mL/min; fractions of 2 mL were collected. The column was calibrated with standard dextrans (5, 12, 80, 270, and 670 kDa). 2.6. Statistical analysis The results are expressed as the mean ± the standard deviation (SD) of triplicate data sets from independent experiments. Statistical analysis was performed using the analysis of variance (ANOVA) test with a confidence level of 95% (P < 0.05).

to 3.02 × 10-12 m2/sec when the temperature increased from 50 to 90°C, and this had an impact on the polysaccharide concentrations. Similar experiments were conducted by Mulet et al. [30], who reported that the D values for sugar extraction from carob pods were 8.7 × 10-11 and 1.57 × 10-10 m2/sec at extraction medium temperatures of 20 and 50°C, respectively. These results are consistent with the study by Pinelo et al. [31], who studied mass transfer during the continuous solid-liquid extraction of antioxidants from grape byproducts and reported that at higher slope values, higher values of phenolic yields and antioxidant activities were obtained. The diffusivity may be improved by increasing the temperature, which can increase the internal energy of the molecules and the solubilities of the substances in the liquid phase as well as reduce the dynamic viscosity of the solvent [14,32]. The dependence of the effective diffusivity on temperature follows a first-order rate process generally described by the Arrhenius equation, D = D0 exp (−Ea/RT)

(9)

where D0 is the pre-exponential factor (m2/sec), Ea is the activation energy (kJ/mol), R is the gas constant (8.3145 × 10-3 kJ/mol/K), and T is the absolute temperature (K). From Eq. (9), D0 and Ea can be estimated as follows:

3. Results and Discussion ln D = (−Ea/R) (1/T) + ln D0 3.1. Effect of extraction temperature Fig. 1A shows that the polysaccharide concentrations in the liquid increased significantly when the extraction temperature increased. An increase in the extraction temperature resulted in an aqueous solution of lower viscosity and minimized mass-transfer resistance; therefore, the amount of polysaccharide extracted at a high temperature was significantly greater than that obtained at a lower temperature. Graphs of ln (C − Ceq)/(C0 − Ceq) versus the extraction time at temperatures of 50, 70, 80, and 90°C are shown in Fig. 1B. The slopes (π 2D/r2) of the regression lines and the D values for the corresponding temperatures are shown in Table 1, and they indicate that the diffusivities of the polysaccharides were affected by temperature. The polysaccharide diffusion coefficients increased from 1.07 × 10-12

(10)

By plotting the effective diffusivity of the polysaccharide as a function of the absolute value of each temperature, as shown in Fig. 2, the D0 value in the Arrhenius equation was found to be 7.958 × 10-9 m2/sec, and the Ea was found to be 24.0 kJ/mol (R2 = 0.9755). Arroqui et al. [33] reported that the D0 and Ea values were 8.63 × 10-6 m2/sec and 24.89 kJ/mol for distilled water. Moreover, an Ea of 29.1 kJ/mol for glucose diffusion in potatoes was reported by Abdel-Kader [16]. To estimate the goodness-of-fit of the models to Eq. (5), Fig. 3 was used to demonstrate that the predicted polysaccharide concentration in the liquid from the model exhibited a good fit to the experimental data. Therefore, this model is acceptable for representing polysaccharide

Table 1. Diffusivities of polysaccharides at each temperature Temperature (oC) 50 70 80 90 r = approximate 0.250 mm.

Ceq (mg/mL) 0.64 0.93 1.93 2.35

Intercept (a) − 0.1278 − 0.2662 − 0.3073 − 0.2367

Slope (b) − 0.0102 − 0.0165 − 0.0195 − 0.0286

D (m2/sec) 1.07 × 10−12 1.74 × 10−12 2.06 × 10−12 3.02 × 10−12


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Fig. 3. Experimental (symbols) and predicted values (lines) from diffusion model. Fig. 2. The relationship between the effective diffusivity and the absolute temperature.

extraction and can be further applied to large-scale production. 3.2. Efficient method for separating polysaccharides After the hot water polysaccharide extraction and removal of the residue by centrifugation, the polysaccharides were isolated using three methods, as shown in Fig. 4. The

results show that the yields of polysaccharides from the three methods were not significantly different (P < 0.05) (Table 2). To select the most suitable method for scale-up, the advantages and disadvantages of each method were considered. The polysaccharide separation process used in method I was shorter than those used in methods II and III; however, the use of cetyltrimethyl ammonium bromide is not safe for polysaccharides that are intended for use as a

Fig. 4. Schematic diagram of extraction and separation of polysaccharides.

Table 2. Advantages, disadvantages, and yields of separated polysaccharide from each method Methods

Yield (%)

Advantage

Disadvantage

Method I

4.87 ± 1.11

The shortest processing time.

Use chemical substances, no good for health and unable for recycling.

Method II

4.31 ± 1.48

Use edible ethanol and recyclable.

Longer time is required in evaporation process.

Method III

4.26 ± 0.14

Use edible ethanol and recyclable. UF process can adapt to continuous process and simplify for large scale production.

Operating time of UF process was longer than method I, but shorter than method II.

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health food. Moreover, the precipitation of polysaccharides with ethanol may be suitable for large-scale production because ethanol can be recycled. However, to increase the concentration of polysaccharides before precipitation with ethanol, an evaporation or UF step is needed. The use of a vacuum rotary evaporator requires precise control because frothing of the solution can occur if the temperature is set at 55°C under low pressure. The pressure must be gradually reduced to avoid frothing, resulting in a long operating time. The UF used in method III is carried out at ambient temperature and is simpler to achieve than the procedure in method II. Moreover, the use of a membrane has advantages in terms of low energy input [34], and this method can be applied to large-scale production. Therefore, method III was considered the most suitable method for separating polysaccharides. 3.3. Suitable membrane MWCO and optimization of the process Membranes with cut-offs of 10 ~ 100 kDa were tested to identify the optimal filter. Fig. 5A shows that the loss of polysaccharides in the permeate increased when the MWCO increased. Although there were no significant differences among the membranes with cut-offs of 10 ~ 50 kDa (P < 0.05) in terms of the polysaccharide yield and retention rate, there were significant differences in the mean permeate flux, which was approximately 15.7, 19.4, and 22.0 L/m2/h for the 10, 30, and 50 kDa cut-offs, respectively (Fig. 5A and Table 3). The yield, yield loss, retention rate and mean permeate flux were used as criteria for selecting the most suitable membrane cut-off. The polysaccharide yield and retention rate after separation using a membrane with a 30 kDa cut-off were slightly higher than those for a membrane with a 50 kDa cut-off. The yield loss for the 30 kDa cut-off was significantly lower than that for the 50 kDa cut-off, although the mean permeate flux was lower

Fig. 5. Yield of polysaccharides at different membrane cut-offs (A) and at different concentrations of ethanol (B).

than that for the 50 kDa cut-off. Therefore, the membrane with the 30 kDa cut-off was selected for further study to determine the effects of filtration area and feed pressure on polysaccharide extraction through a UF membrane. When the effective filtration area of the membrane with a 30 kDa cut-off was increased from 0.005 to 0.01 m2, a

Table 3. Performances of UF with different cut-offs at 69 kPa feed pressure MWCO (kDa) 10 30 50 70 100

Reduction of water (%) 87 87 87 87 87

Mean permeate flux (L/m2/h) 15.7 ± 0.1 19.4 ± 0.4 22.0 ± 0.6 24.1 ± 0.7 29.1 ± 0.1

Retention rate (%) 95.14 ± 1.33 95.40 ± 0.73 93.78 ± 1.49 91.46 ± 2.31 90.22 ± 4.10

Table 4. Performances of UF with 30 kDa cut-off Effective filtration area (m2) 0.005 0.01 0.01

Feed pressure (kPa) 69 69 103

Mean permeate flux (L/m2/h) 19.3 ± 1.6 30.4 ± 2.9 39.3 ± 6.2

Retention rate (%) 95.00 ± 1.65 95.50 ± 0.99 95.33 ± 0.67


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significant increase in the mean permeate flux from 19.3 to 30.4 L/m2/h (Table 4) was achieved. Similarly, an increase in the feed pressure of the polysaccharide solution through the membrane with a 30 kDa cut-off enhanced the mean permeate flux. Sheng et al. [23] also reported that the membrane flux increased linearly with operating pressure. However, care should be taken when using a high feed pressure because high pressure can induce the formation of a cake or gel on the membrane surface, resulting in a decrease in the permeate flux [35]. At an effective filtration area of 0.01 m2, the mean permeate flux at a feed pressure of 103 kPa was significantly higher than that at a feed pressure of 69 kPa, resulting in a decrease in operating time. Therefore, feeding the polysaccharide extract through a membrane with a 30 kDa cut-off and an area of 0.01 m2 at 103 kPa was selected as the optimal set of operating conditions for the UF process. 3.4. Precipitation of polysaccharides with ethanol Once UF was determined to be the most suitable method for separating and concentrating the polysaccharides, the optimal concentration of ethanol for precipitating the polysaccharides was investigated. When different concentrations of ethanol were added to precipitate the polysaccharides, the polysaccharide yield increased as the volume of ethanol increased, reaching a maximum of approximately 4% dry weight at 2 volumes of ethanol (Fig. 5B). A study conducted by Zha et al. [20] also found that the polysaccharide content increased with increasing final ethanol concentration, and a peak value of approximately 0.118 mg/mL was obtained when the final ethanol concentration reached 80%. Because the polysaccharide yield at 2 volumes of ethanol (4.31%) was significantly higher than that at 1.5 volumes of ethanol (3.63%) and was not different from that at 3 volumes of ethanol (4.46%), 2 volumes of ethanol was considered the optimal concentration for polysaccharide precipitation. Moreover, analysis using an ebulliometer showed that the supernatant (discarded) obtained after crude polysaccharide removal contained approximately 60% alcohol, which can feasibly be recycled. 3.5. Molecular weights of the polysaccharides The molecular weights of polysaccharides depend on the methods of extraction and separation. Majdoub et al. [8] reported that the average molecular weight of the polysaccharides from Arthrospira (Spirulina) platensis was 199 kDa and that it contained sulfate corresponding to 20% of the dry weight, whereas Pugh et al. [7] reported that the molecular weight of polysaccharides from Spirulina platensis was estimated to be above 10 million Da. In addition, in U. S. Patent 5,585,365, the molecular weight of antiviral polysaccharides that were extracted with hot water from

Fig. 6. Standard curve of dextrans for a Sepharose 6B column (A) and elution profile of polysaccharide by a Sepharose 6B column (B).

Spirulina species was between 250 and 300 kDa [36]. Chaiklahan et al. [25] reported that the optimal temperature for polysaccharide extraction from Spirulina was 90°C and that extracts obtained at this temperature showed a high antioxidant capacity. Therefore, the molecular weight (MW) of the polysaccharide component that was extracted at 90°C, separated using a membrane with a 30 kDa cut-off, precipitated with 2 volumes of ethanol and dried was estimated using a Sepharose 6B gel filtration column. The calculation of the molecular weight corresponds to Kav, which is defined as: Kav = (Ve − V0) / (Vc − V0)

(11)

where Ve is the elution volume, V0 is the column void volume, and Vc is the geometric column volume. When the calibration curve of Kav was plotted against the log MW of standard dextrans, the equation of the standard curve was log MW = (Kav − 0.8172) / (− 0.1074) (Fig. 6A). Purification of the polysaccharides using a Sepharose 6B column separated the extract into fractions I and II, as in Fig. 6B. The molecular weights of the purified polysaccharides were estimated to be approximately 212 and 12.6 kDa for fractions I and II, respectively.

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4. Conclusion The extraction temperature plays an important role in determining the amount of polysaccharide extracted, which is related to the increase in the D value that occurs when the extraction temperature is increased. Our results demonstrated that the separation of polysaccharides using a UF process and ethanol precipitation is superior to that using processes involving CTAB or vacuum rotary evaporation. Suitable conditions for the UF process involved the use of a membrane with a cut-off of 30 kDa and an area of 0.01 m2 at a feed pressure of 103 kPa. Purification of the polysaccharides using a Sepharose 6B column led to fractions I and II, which had MWs of approximately 212 and 12.6 kDa, respectively.

Acknowledgement This work was supported by King Mongkut’s University of Technology Thonburi, Bangkok, Thailand.

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