Web of Science (Emerging Sources Citation Index), ISC

Document Type : Original Research Article


Department of Chemistry, College of Science for Woman, University of Baghdad, Baghdad, Iraq


A reversed-phase high-performance liquid chromatography method was used for quantitative tannic acid in (rosemary, anise and cinnamon) with isocratic mode. the column which has been used is C18 (250 mm×4.6 mm i.d.;5 µm) at room temperature, with a mobile phase composed of methanol /water which the best ratio for separation tannic acid was (90:10 v: v%) at (pH 6) and the optimum flow rate was 1.5 mL/min, the run time (5 min) at a detection wavelength 277 nm at room temperature. The linearity of calibration curve was (0.1-60 µg mL-1). The value detection of limit and quantification were calculated to be 0.0037 and 0.0122 µg mL-1 respectively. The precision and accuracy were (98.18-100.095%). The recoveries provided by the method were ranged between (-2.57-0.09).

Graphical Abstract

Extraction and determination of tannic acid in rosemary, anise, and cinnamon by reversal phase RP-HPLC


Main Subjects


Medicinal herbs are really essential to health due to the availability of phytocompounds. Alkaloids, glycosides, tannins, flavonoids, and phenolic compounds have become the most useful of these components[1]. Phenolic chemicals are fundamental for plants' hydroxyl radicals recycling and antioxidant effects; they have a wide range of biological activities, which are mostly ascribed to the antioxidant properties in reducing oxidative stress, inhibiting damage, and ion exchange transitional metals. Hydrogen peroxide, hydroxyl radical, nitric oxide, peroxynitrite, and singlet oxygen are all reactive oxygen species (ROS) [2]. Tannic acid (TA) is a natural source for phenolic molecule found in a variety of fruits and vegetables. Because of the amount of TA in tea, fruits, and beer, it can have a significant impact on their flavor; it is a crucial characteristic to consider when reviewing and ensuring the quality of these goods. In the pharmaceutical sector, tannic acid is utilized as an ingredient in products that are used to treat burns and diarrhea [3]. Tannic acid is a food product that is always used. Based on the type of food to which it is put, the safe amount ranges between 10 to 400 µg [4]. Tannic reduced the growth of several fungi, yeasts, bacteria, and viruses [5]. The Antioxidant effect for tannic acid is due to its relatively hydrophobic “center” and hydrophilic ‘‘crust”. Tannic acid and other polyphenol compounds can be found in a variety of beverages and foods. The foods rich in polyphenol, and plants have inhibitory and protective effects on a number of diseases such as cardiovascular disorders, which could be connected least in part to polyphenols antioxidant activity. Tannic acid has been shown to reduce skin, lung, and stomach cancers caused by polycyclic aromatic hydrocarbon carcinogenic. Tannic acid and some other polyphenols have been shown to have anticancer, antioxidant, and anti-carcinogenic properties. The tannic acid as an antioxidant mechanism, for example, behaves as individual productivity in the condition of copper ions, causing DNA damage, or with an antioxidant, reducing hydroxyl radical generation [4]. It is frequently used as in the healthcare, as a purification for wine and beer in the beverage industry and can be used in leather tanning, and ink compounding [6]. Tannic acid, a type of polyphenol, has a structure with several phenolic hydroxyl groups, giving it great physical and chemical properties as well as amazing pharmacological and biological activity [1]. The chemical formula for tannic acid is C76 H52O46, (M.wt = 1701) [7]. Figure 1 shows structure of the tannic acid. Tannic acid is a water soluble polyphenol found in a wide range of plants [5].

There are several methods for determination of Tannic acid and they differ according to the samples taken and according to the equipment used for the determination, including high-performance liquid chromatography [8-10], HPLC-mass spectroscopy [11], gas chromatography (GC) [12], Gas-Mass spectroscopy [13,14], flow injection analysis [15-17], and pencil graphite electrode[3,18]. The purpose of this study is to extract and determine the method of estimating tannic acid in some herbal plants for instance Iraqi rosemary, cinnamon, and anise through HPLC and UV detection.



A (Shimadzu, UV spectrophotometer U.V-1800, Japan), A (Shimadzue, LC-2010 A, Japan) HPLC instrument with UV-Vis detector, Digital balance, PH meter, (WTW, Ph 720, Germany), Hot plate with Magnetic Stirrer, were purchased.

Reagent and solution

Tannic acid (98%) was provided from (BDH), water deionized. acetic acid (99.5%) from (BDH), potassium hydroxide (≥86%) from (Sigma-Aldrich). All solvents (Methanol and water) for HPLC were used from (CHEM-LAB).

Preparation of stock solution

Tannic acid stock solution 1000 µg mL-1 was prepared by dissolving 0.1 gram in methanol and filled to the mark in a 100 mL volumetric flask and diluted. A series of solution ranged (0.1-60 µg mL-1) were prepared to study calibration curve.

Preparation 1% of acetic acid

Acetic acid was prepared by diluting 1 mL of concentrated CH3COOH in water, then completed to 100 mL of volumetric flask and diluted.

Preparation 0.5 M of KOH

Prepared 0.5 M of potassium hydroxide was by dissolving 1.4 grams from KOH in water, then completed to 50 mL volumetric flask of water.

Plant material and extraction

Anise was purchased from the local market and has been ground in a mill, 25 grams sample was mixed with 500 mL boiling water by magnetic stirrer for 15 min. Then, the extract was filtered by Whatman No.1 paper. The filtered sample was frozen and the extract was placed in a plastic bottle and then stored at -20 °C until used [19].

Dried cinnamon sticks were purchased from the local market and grounded by mill, 30 grams, sample was mixed with 500 mL boiling water by magnetic stirrer for 30 min. Aqueous extract was double-filtered using Whatman paper No.1. After that, aqueous extract solution transferred to a bottle and freeze until use at (-20 °C) [20,21].

Rosemary, Iraqi rosemary, was purchased from the plantation and its leaves were dried, outside at the ambient temperature (25–30 °C), then inside at a temperature of not more than 40±2 °C. The dried leaves were mechanically cut off and crushed into powder. The extracts were made in a standard manner by boiling 8 grams of dried leaves in 100 mL of distilled water for 5 minutes and then letting them infuse for 10 minutes. The extracts were then chilled and filtered before being used [22].

Results and discussion

Selecting wave length of tannic acid

Prepared stock solution 10 µg mL-1 of tannic acid was prepared by dissolving in methanol and scanned from 190-400 nm to choose λmax for tannic acid. 277 nm was detected the best absorbing for tannic acid at concentration of 50 µg mL-1. As shown in Figure 2.

Optimization of the experimental conditions for a separation of tannic acid


Effect ratio of mobile phase on elution for tannic acid


Mobile phase composition was methanol: water in different ratios and pump mode was isocratic flow at 1 mL/min at room temperature, the wavelength was detected at 277 nm. Although the ratios of the mobile phase gave a short retention time, they showed a noticeable tailing and irregular and broad peaks that contain a shoulder. The ratio was chosen (90: 10 MET: W v: v%) because of the highest area and height, and lower tailing. Table 1 shows the best ratio.

Effect of pH on retention time elution of tannic acid

To observe the effect of pH change on retention time for separation and determination tannic acid, (TA) was carried out using isocratic elution with mobile phase mobile phase (methanol/water 90:10 v/v %) with 1.0 mL/min flow rate at various pH ranged from (3-7) by using drops from 1% acetic acid. The study shows effect of changing pH on the resolution and response. The best pH 6 has a good peak shape and the highest peak area. The results are shown in Table 2.

The effect of flow rate on elution of tannic acid

Under the ideal conditions discussed earlier, A 20 µl of 50 µg mL-1 tannic acid solution was injected into an HPLC device at various flow rates ranging (0.25- 1.75) mL/min. 1.5 mL/min was chosen because it gave good response, short retention time and the highest peak height and acceptable capacity. The results are shown in Table 3.

The Effect of volume injection on separation of tannic acid

Under the earlier optimizations selection of tannic acid, 50 µg mL-1 was injected into the HPLC system in various volume ranging from (5 to 20 µL). It was found from the maximum response for injected 20 µL because of the high area and height as shown in Table 4.

The best chromatographic conditions

The previous conditions were studied to determine the best parameters, which are the mobile phase (methanol/ water 90:10 v: v%), at PH 6, the flow rate was 1.5 mL/min and the injection volume was 20 µL, detected at 277 nm wavelength at room temperature. Figure 3 shows chromatogram for tannic acid at optimum condition.

Calibration curve

Separation of calibration plots for tannic acid were constructed by plotting the peak area and peak height against respective concentration by various concentrations of tannic acid (0.1-100 µg mL-1) which were prepared from the standard solution and injected n=3 in the HPLC device under previously optimized conditions. The linearity was ranged from (o.1-60 µg mL-1) as shown in Figure 4. The regression equation, correlation coefficient, limits of detection (LOD) and limits of quantification (LOQ) are given in Table 5.   

Precision and accuracy

Under the recommended procedure, the accuracy and precision of obtained results for study of tannic acid were evaluated. Three replicated analyses were carried out at three concentrations (10, 20, 40) µg mL-1 injected into an HPLC device. The data in Table 6 shows the results which indicate a good accuracy and precision of the proposed method at the studied concentration levels.

Application of method

The proposed method was successfully applied for the determination of rosemary, cinnamon and anise after extracted as mentioned previously; 1 mL of the extract was taken and diluted to 46 mL of distilled water for anise and rosemary [19]. As for cinnamon, 1 mL was diluted to 23 mL of distilled water [21,23-25]. Then, the diluted extract was purified with a 0.45 filter, and was injected into a device under the same conditions as those approved as shown in Table 7.

Tannic acid extraction from herbs was added to four different concentrations (5, 10, 20, and 30 µg mL-1). Tannic acid analytical values are extract from spiked extracted herbs. The procedure was accurate, with recovery results ranging from (90.39-100.22) and RSD percent ranging from (0.269-2.577) as shown in Table 8.


The proposed method is simple, rapid, and economical and was conducted using new conditions for the determination of tannic acid by high-performance liquid chromatography, with column (250mm×4.6 mm,5µm) C18 at a flow rate of 1.5 mL/min and retention time for tannic acid was 1.6 at pH 6. These conditions were successfully applied to the extracts of the herbs, which are rosemary, anise, and cinnamon, and showed good estimated amounts. This method also showed good performance, selectivity, linearity, repeatability, accuracy, and specificity, in addition to good LOD and LOQ.


The writers extend their thanks to the staff of the College of Science for Women/ Department of Chemistry, for their support and provision of requirements to complete this work.


How to cite this article: Halla Talib Salih, Ameera Hassan Hamed*. Extraction and determination of tannic acid in rosemary, anise, and cinnamon by reversal phase RP-HPLC. Eurasian Chemical Communications, 2022, 4(1), 94-102. Link: http://www.echemcom.com/article_142343.html


Copyright © 2022 by SPC (Sami Publishing Company) + is an open access article distributed under the Creative Commons Attribution License(CC BY)  license  (https://creativecommons.org/licenses/by/4.0/), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.


[1] R.K. Sangeetha, R. Soundarya, T. Sudhakaran, T.K. Ravi, Sch. Res. J., 2020, 12, 1-18. [Pdf], [Google Scholar], [Publisher]
[2] T.F. De Brum, M. Zadra, M. Piana, A.A. Boligon, J.K. Fröhlich, R.B. De Freitas, S.T. Stefanello, A.L.F. Froeder, B.V. Belke, L.T. Nunes, R. Da Sjlva Jesus, M.M. Machado, J.B. Teixeira da Rocha, F.A.A. Soares, M.L. Athayde, Molecules, 2013, 18, 8342-8357. [crossref], [Google Scholar], [Publisher]
[3] D.L. Vu, B. Ertek, Y. Dilgin, L. Červenka, Czech J. Food Sci., 2015, 33, 72-76. [crossref], [Google Scholar], [Publisher]
[4] İ. Gülçin, Z. Huyut, M. Elmastaş, H.Y. Aboul-Enein,   Arab. J. Chem., 2010, 3, 43-53. [crossref], [Google Scholar], [Publisher]
[5] T.P. Durgawale, P.P. Durgawale, C.C. Khanwelkar, Der Pharm. Lett., 2016, 8, 123-126. [crossref], [Google Scholar], [Publisher]
[6] D.J. Turley, M.T. Kelly, M.R. Smyth, J. Chromatogr. A, 1990, 513, 263-269. [crossref], [Google Scholar], [Publisher]
[7] A. Chaplin, Stain Technol., 1985, 60, 219-231. [crossref], [Google Scholar], [Publisher]
[8] J. Zhu, J. Ng, L. Filippich, J. Chromatogr. B Biomed. Sci. Appl., 1992, 577, 77-85. [crossref], [Google Scholar], [Publisher]
[9] G. Belleau, M. Dadic, J. Am. Soc. Brew. Chem., 1979, 37, 175-179. [crossref], [Google Scholar], [Publisher]
[10] M.J. Rodrigues, V. Neves, A. Martins, A.P. Rauter, N.R. Neng, J.M.F. Nogueira, J. Varela, L. Barreira, L. Custódio, Food Chem., 2016, 200, 322-329. [crossref], [Google Scholar], [Publisher]
[11] J.P. Salminen, V. Ossipov, J. Loponen, E. Haukioja, K. Pihlaja, J. Chromatogr. A, 1999, 864, 283-291. [crossref], [Google Scholar], [Publisher]
[12] P.J. Hernes, J.I. Hedges, Anal. Chem., 2000, 72, 5115-5124. [crossref], [Google Scholar], [Publisher]
[13] O.A. Adebo, E. Kayitesi, F. Tugizimana, P.B. Njobeh, Food Res. Int., 2019, 121, 326-335. [crossref], [Google Scholar], [Publisher]
[14] P. Viñas, N. Campillo, Polyphenols in Plants, ed: Elsevier, 2019, 285-316. [crossref], [Google Scholar], [Publisher]
[15] Q. Langjin, Z. Jie, Y. Hongzong, S. Yanhua, Chinese J. Anal. Chem., 2000, 7. [crossref], [Google Scholar], [Publisher]
[16] H. Cui, Q. Li, R. Meng, H. Zhao, C. He, Anal. Chim. Acta, 1998, 362, 151-155. [crossref], [Google Scholar], [Publisher]
[17] Y.G. Sun, H. Cui, Y.H. Li, H.Z. Zhao, X.Q. Lin, 2000, 33, 2281-2291. [crossref], [Google Scholar], [Publisher]
[18] S.T. Palisoc, E.J.F. Cansino, I.M.O. Dy, C.F.A. Razal, K.C.N. Reyes, L.R. Racines, M.T. Natividad, Sens. Bio-Sens. Res., 2020, 28, 100326. [crossref], [Google Scholar], [Publisher]
[19] İ. Gülçın, M. Oktay, E. Kıreçcı, Ö.İ. Küfrevıoǧlu, Food Chem., 2003, 83, 371-382. [crossref], [Google Scholar], [Publisher]
[20] I. Gulcin, R. Kaya, A.C. Goren, H. Akincioglu, M. Topal, Z. Bingol, K.C. Cakmak, S. B. Ozturk Sarikaya, L. Durmaz, S. Alwasel, Int. J. Food Prop., 2019, 22, 1511-1526. [crossref], [Google Scholar], [Publisher]
[21] P. Kalin, İ. Gülçin, A.C. Gören, Rec. Nat. Prod., 2015, 9, 496-502. [crossref], [Google Scholar], [Publisher]
[22] M. Haloui, L. Louedec, J.B. Michel, B. Lyoussi, J. Ethnopharmacol., 2000, 71, 465-472. [crossref], [Google Scholar], [Publisher]
[23] E. Bursal, E. Köksal, İ. Gülçin, G. Bilsel, A.C. Gören, Food Res. Int., 2013, 51, 66-74. [crossref], [Google Scholar], [Publisher]
[24] I. Gulcin, A.Z. Tel, E. Kirecci, Int. J. Food Prop., 2008, 11, 450-471. [crossref], [Google Scholar], [Publisher]
[25] I. Gulcin, E. Kirecci, E. Akkemik, F. Topal, O. Hisar, Turk. J. Biol., 2010, 34, 175-188. [crossref], [Google Scholar], [Publisher]
[26] X.Y. Chai, S.L. Li, P. Li, J. Chromatogr. A, 2005, 1070, 43-48. [crossref], [Google Scholar], [Publisher]
[27] F.Z. Saltan, H.S. Canbay, A. Üvez, M. Konak, E.İ. Armutak, FABAD J. Pharm. Sci, 2019, 44, 197-203. [crossref], [Google Scholar], [Publisher]  
[28] N.Y. Sariouml, E. Çakir1, M. Kivanccedil, M. Tunccedil, Afr. J. Microbiol. Res., 2011, 5, 158-163. [crossref], [Google Scholar], [Publisher]  
[29] A. Sharma, M. Bhot, J. Varghese, N. Chandra, Asian J. Chem., 2013, 25, 9097-9100. [crossref], [Google Scholar], [Publisher]  
[30] Y. Nakamura, S. Tsuji, Y. Tonogai, J. Agric. Food Chem., 2003, 51, 331-339. [crossref], [Google Scholar], [Publisher]  
[31] C. Møller, S.H. Hansen, C. Cornett, Phytochem Anal., 2009, 20, 231-239. [crossref], [Google Scholar], [Publisher]