Raman and FTIR spectroscopic study of ...

Food Chemistry 114 (2009) 1091–1098

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Food Chemistry journal homepage: www.elsevier.com/locate/foodchem

Analytical Methods

Raman and FTIR spectroscopic study of carboxymethylated non-starch polysaccharides Sze-Nga Yuen a, Siu-Mei Choi a, David Lee Phillips b, Ching-Yung Ma a,* a b

Food Science Laboratory, School of Biological Sciences, The University of Hong Kong, Pokfulam Road, Hong Kong Department of Chemistry, The University of Hong Kong, Hong Kong

a r t i c l e

i n f o

Article history: Received 7 July 2008 Received in revised form 24 September 2008 Accepted 27 October 2008

Keywords: Carboxymethylation Cellulose Guar gum Locust bean gum Xanthan gum Raman spectroscopy Fourier transform infrared spectroscopy

a b s t r a c t Four types of non-starch polysaccharides including cellulose, guar gum, locust bean gum and xanthan gum were modified by carboxymethylation. The degree of substitution (DS) was determined by a colorimetric method. Raman and Fourier transform infrared spectra of the native and modified polysaccharides were acquired and analysed. Representative marker bands, with intensities and/or integrated areas affected by carboxymethylation, were selected at 1607 cm1 (Raman) and 1315 cm1 or 1605 cm1 (IR), attributed to C@O carbonyl stretching vibration. The ratios of intensities (or areas) of the marker bands to that of an internal standard band, corresponding to the skeletal configuration and linkages (850–950 cm1), were plotted against DS. Linear fits were obtained with high correlation coefficients, r > 0.96 (p < 0.01), suggesting a strong correlation between the spectroscopic data and DS determined by wet chemistry. Some structural changes were also observed from the spectral data. Ó 2008 Elsevier Ltd. All rights reserved.

1. Introduction Non-starch polysaccharides (NSPs) can be chemically-modified to alter their physicochemical properties and to improve specific functional properties. Carboxymethylation (CM) is a well-known etherification process for polysaccharides, in which the hydroxyl groups are etherified with carboxymethyl groups. Carboxymethylated cellulose (CMC) exhibits improved solubility, and has been used in the food, cosmetic, detergent, paper, mining, pharmaceutical and textile industries (Togrul & Arslan, 2003; Verraest, Peters, Batelaan, & Bekkum, 1995). CM guar gum, locust bean gum, and xanthan gum have been used in transdermal drug-delivery systems (Murthy, Hiremath, & Paranjothy, 2004), as alkali-resistant printing thickeners (Deuel & Neukom, 1954), and as a thickener in the paper and textile industry (Schweiger, 1966a), respectively. Apart from the types of substitution, the degree of polymerisation and degree of substitution (DS) are factors influencing the properties of the derivatives (Phillips & Williams, 2000). In most cases, valuable functional properties can only be obtained on the partially substituted, rather than the fully substituted polysaccharides, since more severe synthetic reaction conditions may lead to undesirable reactions such as oxidation or depolymerisation (Yalpani, 1988). The solubility and swell ability of CMC were * Corresponding author. Tel.: +852 2299 0318; fax: +852 2858 3477. E-mail address: [email protected] (C.-Y. Ma). 0308-8146/$ - see front matter Ó 2008 Elsevier Ltd. All rights reserved. doi:10.1016/j.foodchem.2008.10.053

shown to be dependent on DS (Togrul & Arslan, 2003). Hence, DS needs to be monitored, to optimise the properties of modified polysaccharides for specific manufacturing processes. DS is normally determined by traditional wet chemistry methods. The measurement of DS in carboxymethylated samples can be carried out by a colorimetric procedure, based on the amount of glycolic acid produced from the cleavage of carboxymethylated products (Welcher, 1975). However, the procedure, similar to many wet chemistry methods, is destructive to samples, time-consuming, involves the utilisation of harmful chemical reagents, requires a relatively large sample size, and is susceptible to interference from residual impurities. Hence, it is not suitable to be used as a routine monitoring method in the polysaccharide industry. Alternative techniques for determining DS become attractive. Differential scanning calorimetry has been shown to have potential to monitor DS in CM chitin (Kittur, Prashanth, Sankar, & Tharanathan, 2002). 13 C Nuclear magnetic resonance spectroscopy and liquid chromatography have been used to determine DS in CM inulin (Verraest et al., 1995). However, these methods are still destructive to the samples and not amenable for routine quality control purposes. Raman spectroscopy was applied by Phillips, Xing, Liu, Chong, and Corke (1999), to determine the degree of acetylation and succinylation in starches. Raman spectroscopic methods have also been successfully adopted in the quantitative analysis of chemicallymodified food proteins (Wong, Phillips, & Ma, 2007; Zhao, Ma, Yuen, & Phillips, 2004). However, for derivatives of NSPs, Raman

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spectroscopy has only been used to identify new or substituted groups and to monitor reaction kinetics (Zhbankov, 1966). Fourier transform infrared (FTIR) spectroscopy has been used as a quantitative tool in some native polysaccharides, such as pectin (Engelsen & Norgaard, 1996), and in some modified carbohydrates, including starch (Klaushofer, Berghofer, & Diesner, 1976) and cellulose (Zhbankov, 1966). Nevertheless, IR spectroscopy has been used only for the identification of specific functional groups, and has not been widely used to determine the extent of modification in chemically-modified NSPs. Although FTIR spectrometers are more commonly available and less costly than Raman spectrometers, the Raman technique has the distinct advantages of not requiring sample preparation, and is non-destructive to the samples. IR spectroscopy is usually limited to dry or non-aqueous samples due to the strong water signal (Li-Chan, 1996). Some sample treatments, such as dissolution of samples in D2O or making into KBr pellets, are required for traditional IR spectroscopy without attenuated total reflectance detection, whereas solid samples can be directly tested by Raman spectroscopy. In this study, four types of NSPs, cellulose, guar gum, LBG and xanthan gum, were modified by carboxymethylation. An analytical technique based on Raman and FTIR spectroscopic methods was developed with the aim of replacing traditional wet chemistry methods. Raman and FTIR spectral data were also analysed, to provide some structural information on the modified molecules.

Cellulose was carboxymethylated, according to the method of Green (1963). Carboxymethylation of galactomannans was carried out, according to Yueh and Schilling (1972). Carboxymethylated xanthan gum was prepared, according to the method of Schweiger (1966b).

microscopic slides. Raman shift was calibrated by a silicon slide at 520 cm1. The spectra (400–4000 cm1 or 400–2000 cm1) were collected at room temperature in the dark under the following conditions: 100 mW of laser power, 2 cm1 resolution, 30 s exposure time, 10–20 scans. Spectral data were baseline corrected using the GRAMS/32 AI Software (Galactic Industries Corporation, Salem, NH). The sugar backbone band in the range of 870–890 cm1 was used as an internal standard for normalisation. Raman spectra were collected in triplicate and plotted as relative intensity (arbitrary units) against Raman shift in wavenumber (cm1). For IR analysis, samples were prepared in the form of potassium bromide pellets containing 10–15 mg polysaccharide samples and 200 mg of potassium bromide, which were mixed and ground in an agate mortar. The pellet method has an advantage over the D2O method, requiring a smaller sample size (Bociek & Welti, 1975). Infrared spectra (400–2000 or 4000 cm1) were recorded with an FTIR Excalibur spectrometer (Bio-Rad Laboratories, Cambridge, MA). A total of 32 scans for each sample were taken at room temperature at a resolution of 4 cm1. Infrared measurement was carried out in the transmission mode, in which the infrared beam directly passes through the sample, and spectral data were then converted from transmittance into absorbance units. Deconvolution of the infrared spectra was performed using Merlin Software, version 1.2 (Bio-Rad Laboratories, Inc.), according to Byler and Susi (1986). The half-bandwidth used for deconvolution was 10 cm1, and the enhancement factor was 2.0. Quantitative estimation of secondary structure components was performed using Gaussian peaks and curve fitting models, according to Byler and Susi (1986). The spectral data were baseline corrected and normalised against the sugar backbone band in the region of 850–950 cm1 using GRAMS/32 AI software. FTIR spectra were plotted as relative intensity (in arbitrary units) against wavenumber (in cm1). All results were performed in triplicate and reproducible data with standard deviation less than 10% were obtained. Raman and FTIR marker bands, with relative intensities or band areas changing with increases in DS (measured by colorimetric method), were selected. Calibration curves were constructed by plotting the ratios of relative intensities (or areas) of specific marker bands to that of an internal standard band against DS. Raman data were input into a Lorentzian curve fitting program and some broad bands containing multi-components were deconvoluted before analysis.

2.3. Determination of degree of substitution by colorimetric method

3. Results and discussion

A colorimetric method, initially used for sodium carboxymethylcellulose (Welcher, 1975), was employed for all carboxymethylated NSPs in this study. The weight of glycolic acid was determined from a calibration curve by plotting the corrected absorbance (A540–A700) against the amount of glycolic acid (mg). According to Eyler, Klug, and Diephuis (1947),

3.1. Determination of degree of substitution by wet chemistry method

2. Materials and methods 2.1. Materials

a-Cellulose, guar gum, locust bean gum (LBG) and xanthan gum were obtained from Sigma Chemical Co. (St. Louis, MO). All chemicals used in this study were of analytical grade. 2.2. Preparation of carboxymethylated NSPs

DS ¼

162G 76 80G

where G is the weight of glycolic acid (mg/mg of sample), 76 is the molecular weight of glycolic acid, 162 is the molecular weight of an average sugar unit and 80 is the net increase in weight of each unit of sodium carboxymethylate group substituted. 2.4. Vibrational spectroscopic methods Raman spectra of native and modified NSP samples were recorded, using a Renishaw-Raman Imaging Microscope (System 1000), equipped with a 514 nm argon ion laser source (Spectra Physics, Mountain View, CA). Solid samples were prepared on

In each sugar unit, the three hydroxyl groups available for carboxymethylation are the two secondary hydroxyl groups at C2 and C3 and the primary group at C6. Under the influence of environment and conformation of the molecules, these three hydroxyls have different reactivity. The DS in carboxymethylated cellulose (CMC), guar gum (CMG), locust bean gum (CML) and xanthan (CMX) are shown in Table 1. 3.2. Determination of degree of substitution by vibrational spectroscopic methods The Raman and IR spectra (4000–400 cm1) of native cellulose, guar gum, LBG and xanthan gum are presented in Fig. 1. Tentative assignments of major Raman bands according to reported data (Colthup, Daly, & Wiberley, 1990; Galat, 1980; Malfait, Van Dael, & Van Cauwelaert, 1989; Mathlouthi & Koenig, 1986; Zhbankov, Andrianov, & Marchewka, 1997) are shown, while those of IR bands were adapted from others (Bociek & Welti, 1975; Colthup et al.,

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Table 1 Degree of substitution (DS) and Raman area (or IR intensity) ratio in carboxymethylated cellulose (CMC), locust bean gum (CML), guar gum (CMG) and xanthan gum (CMX). Carboxymethylated NSP samples

Sodium chloro acetate/polysaccharide (g/g)

DSa

Raman band area ratioa

IR band intensity ratioa Marker bands 1315 cm1

1605 cm1

CMC Control (A) Sample 1 (B) Sample 2 (C) Sample 3 (D) Sample 4 (E) Sample 5 (F)

0.0 0.6 0.6 1.3 1.6 2.0

0.000 0.210 ± 0.010 0.472 ± 0.003 0.679 ± 0.001 0.946 ± 0.008 1.03 ± 0.020

A1607/A888 0.000 0.757 ± 0.247 1.97 ± 0.358 2.54 ± 0.370 3.55 ± 0.001 4.30 ± 0.225

I1317/I897 0.724+0.010 1.09 ± 0.054 1.70 ± 0.101 2.03 ± 0.033 2.65 ± 0.053 2.73 ± 0.080

CML Control (A) Sample 1 (B) Sample 2 (C) Sample 3 (D) Sample 4 (E) Sample 5 (F)

0.0 0.3 0.5 0.7 1.2 1.8

0.000 0.256 ± 0.013 0.442 ± 0.070 0.525 ± 0.030 0.855 ± 0.056 1.55 ± 0.098

A1607/A868 0.000 1.12 ± 0.001 1.22 ± 0.142 1.54 ± 0.285 3.84 ± 0.002 6.24 ± 0.133

I1315/I872 0.104 ± 0.074 0.615 ± 0.023 1.20 ± 0.029 2.68 ± 0.416 7.50 ± 0.010 10.3 ± 0.933

CMG Control (A) Sample 1 (B) Sample 2 (C) Sample 3 (D) Sample 4 (E)

0.0 0.6 0.8 1.0 1.7

0.000 0.296 ± 0.008 0.435 ± 0.030 0.610 ± 0.019 1.13 ± 0.083

A1607/A880 0.000 1.62 ± 0.281 2.16 ± 0.378 6.26 ± 0.319 9.10 ± 0.652

I1610/I870 2.17 ± 0.229 4.87 ± 0.387 6.32 ± 0.162 7.99 ± 0.881 10.3 ± 0.751

CMX Control (A) Sample 1 (B) Sample 2 (C) Sample 3 (D) Sample 4 (E)

0.0 0.6 0.9 1.1 1.3

0.021 ± 0.006 0.403 ± 0.020 0.566 ± 0.070 0.595 ± 0.001 0.971 ± 0.031

A1607/A887 6.10 ± 0.854 10.2 ± 0.937 13.3 ± 0.010 14.0 ± 0.031 17.4 ± 0.004

I1603/I895 11.0 ± 0.023 25.7 ± 1.30 33.9 ± 2.73 40.5 ± 0.746 57.4 ± 4.54

a

Means of triplicate determinations ± standard deviations.

1990; Galat, 1980; Mathlouthi & Koenig, 1986; Xiao, Weng, & Zhang, 2002; Zhbankov, 1966; Zhbankov et al., 1997). Specific vibrations can be found in similar frequency regions in IR and Raman spectra, since both methods provide information on molecular vibrations. The Raman and IR band in the region of 850–950 cm1, corresponding to the skeleton mode of the anomeric skeletal configuration (a or b conformers) and glycosidic linkages, was used as an internal standard band, due to its insensitivity to carboxymethylation, i.e., the degree of anomerisation is constant (Malfait et al., 1989). The band intensities were normalised relative to this internal standard, thus eliminating problems associated with the potassium bromide method, e.g., influences of concentration and particle size on the height of the IR bands (Klaushofer et al., 1976). Since all four polysaccharides have backbone chains composed of b-glycosidic linkage, a Raman band at 870– 890 cm1 and an IR band at 870–900 cm1 appeared. Raman bands were observed at 888, 880, 868 and 887 cm1 (IR: 897, 870, 872 and 895 cm1) for cellulose, guar gum, LBG and xanthan gum, respectively. They were ascribed to C–C or C–O vibrations coupled with the C–H mode of the anomeric carbon of b-conformers (Malfait et al., 1989). It should be noted that for the internal standard Raman band of CMC and CMX, deconvolution was carried out before peak area measurement since the peak overlapped with the 907 cm1 band in higher DS samples. A new broad band at 1607 cm1 was observed in the Raman spectra of CMG (Fig. 2). This band was also observed in CMC and CML, and in native xanthan (data not shown). With increases in the level of carboxymethylation, the size of this band was increased. The stretching vibration of C@O can be distinguished as two types, according to the location of the bands, a strong band at 1550–1900 cm1 region attributed to carbonyl compounds, and a band at 1740–1800 cm1 ascribed to carboxylic acid

(Colthup et al., 1990). Since sodium chloroacetate was used as the modifying agent instead of chloroacetic acid, the resulting carboxymethylate species in the four polysaccharides were in sodium salt (–COONa+) rather than in the acid form (–COOH), giving a new broad band at 1607 cm1. Although the bands at 907, 1310 and 1402 cm1, and the shoulder band at 1253 cm1, attributed to the carboxymethyl groups, also showed progressive increases after carboxymethylation, they were not suitable marker bands since they were attributed to other vibrations such as C–H stretching, anomeric skeleton and C–O–H stretching and bending, whereas the 1607 cm1 band was attributed solely to the substituted carboxymethylate species. There were increases in three other IR peaks at 1605, 1426 and 1315 cm1 in all the CM polysaccharides, such as CMG (Fig. 3) (data not shown for CMC, CML and CMX). These three bands were attributed to C@O stretching in COO ions, and were in agreement with those bands found in carboxymethylated starch (Klaushofer et al., 1976) and CMC (Zhbankov, 1966). Only slight intensity increases in the 1263 cm1 band, attributed to C–O–C stretching, were shown in CMC and CMG. Comparatively, the extents of increase in these IR bands were higher than those of the corresponding Raman bands, except the 1263 cm1 transition. Since IR spectroscopy detects vibrations caused by a change in the electrical dipole moment of molecules, whereas Raman spectroscopy is sensitive to those caused by the electrical polarisability changes, asymmetric COO polar groups give a much stronger band in IR spectra, while symmetric non-polar C–O–C groups are more prominent in Raman spectra. The Raman marker band was easily selected at 1607 cm1 due to a lack of interference from vibrations in the native polysaccharides. Substantial increase in the intensity of this Raman band was caused only by the introduction of new carboxymethylate groups. However, a water band appeared in the IR region of 1645–1635 cm1, and would overlap with the C@O stretching vibration at 1605 cm1 ascribed to newly

880

1327 1262

919

1414 -1

1607 cm C=O stretch

anomeric region

E

Relative Intensity

C-H bending, wagging

Relative Intensity

O-H stretching

a

C-O stretching; C-O-H stretching, bending, deformation


C-H stretching

1094

X

L

D

C

G

B

C

4000

A 3500

3000

2500 1500

1000

500

-1

Raman shift (cm )

600

400

-1

Fig. 2. Raman spectra of native and carboxymethylated guar gum. A: Native guar gum; B–E: carboxymethylated guar gum with increasing degree of substitution.

Relative Intensity

L

G

870

1263

1427

E

X

1310

1610

anomeric region

C-H bending, wagging; Alcoholic C-O C-O stretching; C-O-H stretching, bending, deformation

Water; C=O stretching in xanthan

O-H stretching

C-H stretching

Raman Shift (cm )

b

Relative Intensity

2000 1800 1600 1400 1200 1000 800

D

C C 4000

3500

3000

2500

2000

1500

1000

500

-1

Wavenumber (cm ) Fig. 1. (a) Raman spectra (4000–400 cm1) and (b) FTIR spectra (4000–400 cm1) of native polysaccharides: cellulose (C), guar gum (G), locust bean gum (L) and xanthan gum (X).

substituted carbonyl groups in carboxymethylate (–COONa+) species. Intensity of the 1426 and 1315 cm1 bands was influenced to various extents by other vibrations, such as C–H stretching and anomeric skeleton, while the 1236 cm1 band was affected by C–O–H vibrations. Thus, it was difficult to find an IR band attributed only to the CM groups. Nevertheless, the feasibility of using the 1315 cm1 and 1605 cm1 vibrations as marker bands for the construction of calibration curves was demonstrated in the present study.

B

A

2000 1800 1600 1400 1200 1000

800

600

400

-1

Wavenumber (cm ) Fig. 3. FTIR spectra of native and carboxymethylated guar gum: A: Native guar gum; B–E: carboxymethylated guar gum with increasing degree of substitution.

Table 1 lists the relative areas of the 1607 cm1 Raman band, and relative intensities of 1315 or 1605 cm1 IR marker band normalised to that of the internal band for different polysaccharide


samples, at varying DS measured by the wet chemistry method. Since the 1607 cm1 Raman band was quite broad, the integrated area was used to more accurately quantify the changes in the amount of C@O in the samples. Plots of intensity (or area) ratios of the Raman and IR marker bands against DS were drawn, and linear regression lines (y = mx + b; where y is the ratio of the marker band to the internal standard band, x is the DS, m is the slope and b is the y-intercept) were obtained (Tables 2 and 3). The intensity (or area) of these marker bands showed a strong linear relationship with DS, as the correlation coefficients (r) of all calibration curves were all larger than 0.96 (p < 0.01). For the 1607 cm1 Raman marker band, the slopes (m) of the area calibration curves were 4.002, 8.625, 4.171, and 12.47 for cellulose, guar gum, LBG and xanthan gum, respectively (Table 2). Zero intensity for the Raman area was almost within the uncertainty of the linear regression y-intercept (b = 0.0365 ± 0.132, 0.432 ± 0.840 and 0.194 ± 0.271 for area ratio in CMC, CMG and CML respectively), except in CMX (b = 5.864 ± 0.590 for area ratio). This confirms that the 1607 cm1 band is mainly attributed to carboxymethylation of hydroxyl groups with little contribution from native polysaccharides. For xanthan gum, the 1607 cm1 marker Raman band has a noticeable background from a near coincident native Raman band, which is mainly due to the C@O group vibrations of acetate, glucouronate and pyruvate in the parent sample. Nevertheless, the 1607 cm1 Raman band still provides a convenient marker band, since a good calibration curve has been obtained for CMX. When all four sets of data were pooled and plotted, the resulting correlation coefficient (r) was markedly decreased to 0.44 (p < 0.05). Since the four polysaccharides have different chemical structures, the C@O groups incorporated into them by CM would experience different molecular environments.

Table 2 Linear regression parameters from the Raman calibration curvesa,b. NSP samples

m

b

r

p

n

Cellulose Guar gum LBG Xanthan gum Pooled datac

4.00 ± 0.197 8.63 ± 1.36 4.17 ± 0.347 12.5 ± 0.987 5.48 ± 2.44

0.0365 ± 0.132 0.432 ± 0.840 0.194 ± 0.271 5.86 ± 0.590 1.89 ± 1.65

0.995 0.965 0.986 0.991 0.448