Simultaneous determination of ten major phenolic acids in sugarcane by a reversed phase HPLC method

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1 Simultaneous determination of ten major phenolic acids in sugarcane by a reversed phase HPLC method Gleichzeitige Bestimmung 10 bedeutender Phenolsäuren in Zuckerrohr mittels RP-HPLC Zhen-Gang Zhao, Li-Cai Zhu, Shu-Juan Yu, Xiong Fu, Xin-an Zeng A high-performance liquid chromatographic method was applied to the determination of ten phenolic acids in sugarcane. The ten phenolic acids were simultaneously analyzed with a Waters XTerra RP 18 column (150 mm 4.6 mm ID, particle size 5.0 µm) by gradient elution using acetonitrile/h 2 O (2% acetic acid) as the mobile phase at 28 C. The flow rate was 1 ml/min, and detection wavelength was set at 280 nm. The recovery of the method was in the range of %, and the relative standard deviation (RSD) was less than 4% for all the compounds. This assay was successfully applied to the determination of ten major phenolic acids in cane juice, fresh leaves and bagasse. The results indicated that the developed HPLC assay could be readily utilized as a suitable method for the determination of the phenolic acids in sugarcane processing. Keywords: RP-HPLC (reversed phase high performance liquid chromatography), phenolic acids, sugarcane, bagasse Zur quantitativen Bestimmung von Phenolsäuren in Zuckerrohr wurde ein HPLC-Verfahren entwickelt. Die 10 Phenolsäuren wurden gleichzeitig analysiert bei 28 C mit Hilfe einer Waters-XTerra-RP 18 -Säule (4.6 mm innerer Durchmesser, 150 mm Länge, Partikelgröße 5.0 µm) mit einer Gradientenelution, bei der ein Acetonitril-Wasser-Gemisch (2 % Essigsäure) als mobile Phase verwendet wurde. Die Flussrate betrug 1 ml/min, gemessen wurde bei einer Wellenlänge von 280 nm. Die Wiederfindungsrate bei diesem Verfahren lag im Bereich 85 bis 109 % und die relative Standardabweichung war <4 % für alle Säuren. Das HPLC-Verfahren wurde erfolgreich zur Bestimmung von 10 bedeutenden Phenolsäuren in Rohrsaft, Zuckerrohr und Bagasse angewandt und kann entsprechend als eine geeignete Methode bei der Zuckerrohrverarbeitung eingesetzt werden. Stichwörter: RP-HPLC (Umkehrphasen-Hochleistungsflüssigkeitschromatographie), Phenolsäuren, Zuckerrohr, Bagasse 1 Introduction Polyphenols including flavonoids and phenolic acids are products of the secondary metabolism of plants. In plants, polyphenols may act as phytoalexins, antifeedants, attractants for pollinators, contributors to plant pigmentation, and protective agents against UV light [1]. In addition, polyphenols exhibit many biological properties such as anticarcinogenic, anti-ulcer, antithrombotic, anti-inflammatory, immune modulating, anti-allergic, antimicrobial, vasodilatory, and antioxidant activities [2]. Polyphenols have become an intense focus of research interest because of their perceived health-beneficial effects. Phenolic acids, low molecular mass phenolics, are derivatives of benzoic and cinnamic acids. They are widely distributed in sugarcane and are color precursors in cane sugar manufacture [3, 4]. Phenolic acids have significant biological and pharmacological properties, some of which were shown to be effective in preventing cancer [5]. They contribute to color formation of juice when sugar cane is crushed, and are also involved in the changes that take place during the processing of sugarcane for the production of raw sugar [6, 7]. Thus, there is need to characterize phenolic compounds present in cane juice. In the present paper, the simultaneous quantification of the ten major phenolic acids (gallic acid, chlorogenic acid, vanillic acid, caffeic acid, p-coumatic acid, ferulic acid, sinapic acid,

2 protocatechuic acid, p-hydroxybenzoic acid, and syringic acid) in sugarcane is intend to report by reversed phase liquid chromatography. Although the presence of these compounds in sugar cane is known, a method for their simultaneous and direct quantification is not available. 2 Materials and methods 2.1 Materials and standards The sugarcane variety Y00318 was obtained from a farm in ZhangJiang, Guang Dong province (China). Gallic acid, chlorogenic acid, vanillic acid, caffeic acid, p-coumaric acid, ferulic acid, sinapic acid, protocatechuic acid, p-hydroxybenzoic acid, and syringic acid were purchased from Sigma-Aldrich (St. Louis, MO, USA). HPLC-grade solvents, methanol, acetonitrile and acetic acid were purchased from Dikma Technology Inc. (Lampoc, CA, USA). Analytical-grade ethyl acetate was purchased from Beijing Beihua Fine Chemicals Co. Ltd. (Beijing, China). For HPLC analysis, ultra-pure water from a Milli-Q system (Millipore, Bedford, MA, USA) was used. 2.2 Apparatus and chromatographic conditions A Waters 600E HPLC system (Waters Technologies, Milford, MA, USA) consisting of a quaternary pump, an on-line degasser, a column temperature controller, a Waters 2487 dual λ detector and Waters Empower2 software. Chromatographic separations were performed on a Waters XTerra RP 18 (150 mm 4.6 mm ID, particle size 5.0 μm) including XTerra RP 18 guard column (3.9 mm 20 mm, particle size 5.0 μm). The sample injection volume was 20 μl. The detection wavelength was set at 280 nm, the flow rate was 1.0 ml/min and the column temperature was maintained at 28 C. The mobile phase was a mixture of acetic acid water (2:98, v/v) (solvent A) and acetic acid acetonitrile water (2:20:78, v/v) (solvent B). The following gradient was used:0~45 min 95% A, 5% B; 45~70 min, from 95% A, 5% B to 10% A,90% B, linear gradient; 70~75 min, 10% A, 90% B. Prior to HPLC analysis, all samples were filtered through a 0.45 μm syringe filter. 2.3 Preparation of standard solutions Stock solutions of g/ml phenolic acids were prepared by dissolving appropriate amount of gallic acid, chlorogenic acid, vanillic acid, caffeic acid, p-coumaric acid, ferulic acid, sinapic acid, protocatechuic acid, p-hydroxybenzoic acid, and syringic acid in methanol, respectively. Standard solutions were prepared by further dilution of stock solutions with methanol. The HPLC mobile phase was prepared fresh daily, filtered through a 0.45 μm membrane filter, and then degassed before injecting into the column. All the stock and standard solutions were stored at 20 C. 2.4 Sample preparation To prepare the juice extracts, 10 ml of fresh sugarcane juice was adjusted to ph 2.0 using 2 mol/l HCl and subsequently extracted 2

3 four times using 40 ml of ethyl acetate each time. The clear organic extracts were combined and dried over anhydrous sodium sulfate, and then the extract was filtered and evaporated to dryness in a rotary vacuum evaporator at a temperature not higher than 40 C. The residue was dissolved in 2 ml of methanol. Finally, the resulting solution was filtered through a syringe filter of 0.45 μm pore size and was ready for chromatographic analysis. 1 g each of fresh sugarcane leaves and dried sugarcane bagasse was extracted in 70% ethanol (3 100 ml, 2 h each), and the supernatants were obtained by centrifugation at 3000 min 1 for 15 min and concentrated by flash evaporation, the ph value was adjusted to 2.0 with 2 mol/l HCl. Phenolic acids were separated by ethyl acetate phase separation (4 50 ml) and the pooled fractions were treated with anhydrous sodium sulfate, filtered and evaporated to dryness in a rotary vacuum evaporator at a temperature not higher than 40 C, the residue was dissolved in 2 ml of methanol. Finally, the resulting solution was filtered through a syringe filter of 0.45 μm pore size and was ready for chromatographic analysis. 3 Results and discussion 3.1 Method development and optimization The standard solutions were scanned at from 200 to 400 nm, using a UV-2102 PC UV-visible spectrophotometer (Unico, China) while the spectra were recorded. The UV spectrum of the phenolic standards exhibited maximum absorptions at about 280 and 320 nm. Detection of phenols is usually achieved at λ = 280 nm even though this is not the maximum wavelength for all the phenolic acids. However, detection at 280 nm may be a compromise because most phenolic acids absorb considerably at this wavelength and other compounds interfere only slightly. In addition, the baseline shifts of the chromatograms did not increase considerably. Consequently, 280 nm was selected as the appropriate wavelength for detection of the phenolic acids. A good separation condition should satisfy the need that the analyzed peaks have baseline separation with adjacent peaks within a short analysis time as far as possible. The effectiveness of HPLC separation was tested using the standard solution of ten phenolic acids including gallic acid, chlorogenic acid, vanillic acid, caffeic acid, p-coumaric acid, ferulic acid, sinapic acid, protocatechuic acid, p-hydroxybenzoic acid, and syringic acid. To obtain chromatograms with a good separation, mobile phase, percentage organic modifier, column temperature and flow rate were, respectively investigated. Initially, the more straightforward isocratic elution condition was investigated. The elution involving the use of methanol/acetonitrile (1:1, v/v): H 2 O/acetic acid (99:1, v/v) at (15:85, v/v) at flow rate of 1.0 ml/min was tested. It was found that chlorogenic acid and caffeic acid, ferulic acid and sinapic acid were not separated, although there was a good separation between the other phenolic acids. Various mixtures of water and methanol were used as the mobile phase but separation was not satisfactory. Reversed-phase liquid chromatography using acetonitrile gradient under acidic mobile phase conditions is a common practice in the separation of complex samples. The results indicated that acetonitrile/water/acetic acid has a better resolution of the peaks than methanol/water/acetic acid in separation, and that a chromatographic gradient system composed of acetonitrile and water, when adding acetic acid, could sharpen peak shapes and improve analytical sensitivity and resolu- 3

4 tion for the HPLC analysis of phenolic acids. It was also found that the concentration of acetic acid was crucial for the simultaneous baseline separation of chlorogenic acid and caffeic acid. Acetic acid could achieve better separation for phenolic acids because it reduces the ionization of phenol, phenolic hydroxyl and carboxyl groups. Therefore, a mobile phase containing of 2% (v/v) acetic acid in acetonitrile/water was selected. The temperature of the column and the flow-rate of the mobile phase, which might affect the separation, were also tested. Raising the column temperature served to decrease viscosity of the mobile phase, and the optimal column temperature was found to be 28 C. At higher temperatures some resolution problems arose, especially between chlorogenic acid and caffeic acid, while at lower temperatures no particular improvement was observed. It also increases the retention times. The most suitable flow rate was found to be at 1 ml/min. The target components in the chromatographic profile of samples were identified by comparing the retention times and the characteristic of the UV spectra of these peaks with those presented in the chromatogram of the mixture standard solution. The chromatograms of pure standards and cane juice samples using the improved and adopted chromatographic conditions are presented in Figures 1 and 2. Fig. 1: HPLC-UV chromatograms of ten phenolic acids standards Fig. 2: HPLC-UV chromatograms of ten phenolic acids in cane juice 3.2 Calibration curves and the limits of detection Under the optimum conditions described above, the results obtained with the proposed method are summarized in Table 1. The external standard method was used to obtain the regression equations. In the regression equation y = ax + b, x represents the concentration of the standard compounds (μg/ml), y is the peak area, a is the intercept of the straight line with y-axis and b is the slope of the line. The r in Table 1 represents the correlation coefficient of the equation. All the standard compounds showed good linearity (r > ) in the range of 2.23~55.00 μg/ml. Sensitivity was investigated by determining the lower limit of detection based on the peak-to-peak noise of the baseline and on a minimal value of the signal-to-noise ratio of 3. The limit of detection (LOD), for all compounds, was in the range of ~ μg/ml. Table Recovery and precision Recovery tests were carried out to further investigate the accuracy of the method by adding three concentration levels (low, medium and high) of the mixed standard solutions to known amounts of samples. The resultant samples were then extracted and analyzed by the described method. The results were calculated with the value detected versus added amounts. The recoveries of the method were in the range of %, with RSD (relative standard deviation) less than 4% as shown in Table 2. From the results of precision test and recovery test, it was known that the method manifested good precision and accuracy. Accuracy and precision were evaluated by adding known escalating amounts of each standard to a solution of a known concentration whose analysis was replicated four times. The precision was 4

5 expressed as RSD and the accuracy as the amount found. Table Sample analysis The samples (Table 3) were prepared as described in section 2.4. A volume of 20 μl of the filtered solution of each sample was injected into the instrument. Peaks in the chromatograms were identified by comparing the retention times and on-line UV spectra with those of the standards. The content of each analysis was calculated from the corresponding calibration curve. Table 3 4 Conclusion A reversed-phase HPLC method has been developed for the simultaneous determination of gallic acid, chlorogenic acid, vanillic acid, caffeic acid, p-coumaric acid, ferulic acid, sinapic acid, protocatechuic acid, p-hydroxybenzoic acid, and syringic acid in sugarcane. This method was shown to be simple, rapid and precise. In this study, the phenolic acid contents of cane juice, fresh leaves and bagasse were determined by the optimized method, respectively. Gallic acid, coumaric acid, ferulic acid, caffeic acid, chlorogenic acid were the main phenolic acids identified in sugarcane. This method could be readily utilized as a suitable method for the determination of the phenolic acids in sugarcane processing. Consequently, further research can be carried out on the relationship between phenolic acids and color formation. Acknowledgements Authors thank Dr. Chung Chi Chou for his keen interest in the work and constant encouragement. The authors also thank the Guangzhou Sugarcane Industry Research Institute for supplying cane of commercial varieties for use in the experiments. This study was supported by grants from the National Natural Science Foundation of China (serial number ). References 1 Naczk, M.; Shahidi, F. (2006): Phenolics in cereals, fruits and vegetables: Occurrence, extraction and analysis. Journal of Pharmaceutical and Biomedical Analysis 41 (5), Wollgast, J.; Anklam, E. (2000): Review on polyphenols in Theobroma cacao: changes in composition during the manufacture of chocolate and methodology for identification and quantification. Food Research International 33 (6), Goodacre, B.C.; Coombs, J. (1978): Formation of color in cane juice by enzyme-catalyzed reactions. Part II. Distribution of enzyme and color precursors. Int. Sugar J. 80 (959), Paton, N.H. (1992): Sugar cane phenolics and first expressed juice color. Part I. Determination of chlorogenic acid and other phenolics in sugar cane by HPLC. Int. Sugar J. 94 (1121), , McCann, M.J., et al. (2007): Anti-cancer properties of phenolics from apple waste on colon carcinogenesis in vitro. Food and Chemical Toxicology 45 (7), Paton, N.H.; Duong, M. (1992): Sugar cane phenolics and first expressed juice color. Part III. Role of chlorogenic acid and flavonoids in enzymic browning of cane juice. Int. Sugar J. 94 (1124), Bucheli, C.S.; Robinson, S.P. (1994): Contribution of Enzymic Browning to Color in Sugarcane Juice. Journal of Agricultural and Food Chemistry 42 (2),

6 Table 1: Retention times, regression equations, and detection limits (LOD) of ten phenolic acids Phenolic acids Retention time) Test range Standard curve r LOD in min in μg/ml in μg/ml Gallic acid ~51.00 y = 50,900x 60, Protocatechuic acid ~53.00 y = 28,600x 2, p-hydroxybenzoic acid ~50.00 y = 36,400x 8, Vanillic acid ~50.00 y = 39,100x 5, Chlorogenic acid ~44.50 y = 11,400x + 2, Caffeic acid ~51.00 y = 37,900x + 5, Syringic acid ~54.50 y = 5,860x + 1, Coumaric acid ~53.50 y = 96,600x 65, Ferulic acid ~51.00 y = 54,600x 4, Sinapic acid ~51.50 y = 29,400x 4, y peak area; x concentration of analyte (μg/ml); LOD (limit of detection): signal-to-noise ratio = 3 Table 2: Analytical results of recoveries (n = 4) Phenolic acid Added Recovery RSD in μg/ml in % in % Gallic acid Protocatechuic acid p-hydroxybenzoic acid Vanillic acid Chlorogenic acid Caffeic acid Syringic acid Coumaric acid Ferulic acid Sinapic acid Table 3: Content of ten phenolic acids in sugarcane (data are means ± standard deviation (n = 3) determined by HPLC analysis) Phenolic acids Cane juice* Fresh leaves Bagasse in mg/l in mg/kg in mg/kg Gallic acid.15 ± ± ± 1.25 Protocatechuic acid 0.89 ± ± ± 1.51 p-hydroxybenzoic acid 0.86 ± ± ± 1.27 Vanillic acid 0.97 ± ± ± 1.05 Chlorogenic acid 0.81 ± ± ± 1.17 Caffeic acid 2.26 ± ± ± 1.38 Syringic acid 0.47 ± ± ± 0.73 Coumaric acid 0.93 ± ± ± 2.55 Ferulic acid.13 ± ± ± 4.30 Sinapic acid 0.42 ± ± ± 0.39 * Cane juice refractometric dry substance content = 16.5%. 6

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