INTRODUCTION:

Colors are used to hide defects of food products and improve its superficial features and also to meet consumer demands¹. For this, the emergence of inorganic pigments as food colors which reported to be very harmful to human being. There is a need for substitution of the synthetic colors used in the food industry with natural colors because of the general toxicity of synthetic colors, making them unaccepted for human consumption².

Various studies have confirmed the use of permitted and non permitted colors or its overconsumption causes hyperactivity, urticaria, dermatitis, asthma, nasal congestion, abdominal pain, inhibition of the immune system, attention deficit disorders, eczema, liver and kidney damage, thyroid tumors, nausea, and proved to be carcinogenic³. As a result, there has been a worldwide demand for natural food colorants development⁴. The benefits of using natural colors are many as they are eco-friendly, unsophisticated and harmonized with nature, safe for body contact, obtained from renewable sources and also their preparation requires a minimum probability of chemical reactions. Generally natural colors are not harmful⁵. The demand for food color in global market in 2000 was 2400 MT which increased to 3000 MT by the year 2005 and further to increase to 8000 MT by the year 2010 and is expected to increase to 15000 MT by the year 2015. The expenditure in natural food color market across the globe has reached to US $ 1 billion and is constantly growing as there is demand for natural food colors in substitution of synthetic food colors⁶. Moreover natural colors are very unstable and easily degraded at the time of food processing. As compared to natural colors, synthetic colors show numerous advantages such as high consistency to light, oxygen and pH, color uniformity, low pollution, relatively less production costs, etc. Natural color degradation occurs during extraction, purification, processing and storage of the pigments. That’s why synthetic food colors are used alternative of natural colors in many food products¹. Anthocyanin are a subclass of flavonoids⁷. Anthocyanins are blue, red, violet and purple coloration in most species of plants². Anthocyanins are very unstable and easily liable to degradation. The stability of anthocyanins is affected by factors like pH, temperature, UV, presence of enzymes, structure and concentration of the anthocyanins⁷. One of the important characteristics of anthocyanins is the change in coloration with the varying pH of the environment. The color and stability of an anthocyanin in solution is more dependent on pH. Anthocyanins are most stable and highly colored at low pH values and slowly lose color as the pH increases, becoming nearly colorless between pH 4.0 and 5.0. This color loss is reversible and will return upon acidification. This limits the use of anthocyanins as a food colorant to products with low pH values. Anthocyanins primarily exist in their colorless forms in neutral to slightly acidic pH. Anthocyanins are more stable in acidic rather than in neutral or alkaline solutions. Current studies using purified anthocyanins have attestable benefits include protection against liver injuries, improvement of eyesight, strong anti-inflammatory and antimicrobial activities, significant reduction of blood pressure, inhibition of mutations caused by mutagens from cooked food and suppression of human cancer cell proliferation. As like other phenolic compounds, these extracts are dynamic scavengers of free radicals, although they can also act as prooxidants. Because of their multiple physiological actions, the consumption of anthocyanins by human beings may play a major role in preventing diabetes, cancer, cardiovascular and neurological disease².

The stability of anthocyanin color can be enhanced by copigmentation phenomenon where the anthocyanin molecule reacts with other natural plant substances through weak interactions forming an enhanced and stabilized color. Copigmentation is a solution phenomenon in which pigments and copigment molecules forms molecular complexes⁸. These causes the pigments to exhibit high color intensity than would be expected from their original⁷. Anthocyanins, like other natural colors, possess low stability. Their degradation can occurs during extraction, purification, processing and storage of the pigments². Copigmentation occurs when a colorless molecule with a planar, pi-electron rich moiety interacts with the planar anthocyanin chromophore forming a colored complex. This interaction prevents water attack on the flavylium cation. The complexation of a copigment with an anthocyanins produce a hyperchromic shift and a bathochromic shift. The hyperchromic shift indicated increase in color intensity while the bathochromic shift indicates shift of the wavelength of maximum absorbance. A number of different substances has been found to act as effective copigments. The most common copigments are flavonoids, ppolyphenolic compounds, alkaloids, amino acids and organic acids⁹. Hibiscus sabdariffa is a tropicalplant which belongs to the Malvaceae fam¹⁰. Its typical red calyx are rich in anthocyanins, minerals, pectin. The major anthocyanins reported are delphinidin-3-glucoside delphinidin-3-sambubioside, and cyanidin-3-sambubioside chiefly responsible for their color and antioxidant properties. The present study was undertaken with an aim to purify anthocyanins from Hibiscus saddariffa and an attempt has been made to improve its stability by copigmentation¹¹. The present study highlights studies on the stabilization of anthocyanins from Hibiscus sabdarriffa.

MATERIALS AND METHODS:

Raw material:

The dried calyces of Hibiscus sabdariffa was used as source of plant materiel investigated in the present study. This materiel was procured from APMC market, Vashi, Mumbai.

Authentication:

Plant material was identified by Gurunanak Khalsa College, Botany Dept, Wadala, Mumbai. The specimen number is 1030955.

Extraction:

100 g of Hibiscus sabdariffa calyx were Soxhlet extracted in 200 ml of distilled water acidified with 0.1 N HCl. The extract was filtered using Whatman filter paper. After vacuum evaporation of the water in rotary evaporator dry extract was obtained. The dry extract was added with 200 ml of distilled water and the aqueous extract was poured on gel XAD7 to eliminate sugars and chlorophyll pigments. The water obtained after filtration was discarded. 100 ml of ethanol added over the gel XAD7 and the ethanolic filtrate obtained was evaporated to dryness with rotary evaporator. The dried extract obtained represents purified extract which was used to for further analysis¹².

Qualitative phytochemical screening:

The presence of phytochemicals was determined by standard phytochemical methods. The purified extract was analysed for presence of alkaloids, polyphenols, flavonoids, tannins, quinones, saponins, tannins, terpenes and sterols¹³.

Confirmative Test for Anthocyanin:

2M NaOH: 1ml of extract solution was added with 2ml of NaOH, and the results were observed¹⁴.

Determination of Total Anthocyanin Content (TAC) by pH differential method:

A spectrophotometric method that involves measurement of absorbance at pH 1.0 and 4.5 The buffer solutions at pH 1.0 (0.025 M potassium chloride) and at pH 4.5 (0.4 M sodium acetate) were prepared. 0.1 gm of Anthocyanin extract was diluted with the buffer to 10 ml. Absorbance was measured at 520 and 700 nm with UV-VIS spectroscopy.

Total anthocyanins content (mg/L) was expressed as cyaniding-3-glucoside according to the following equation and was converted to mg anthocyanin/100 g sample.

^TAC = A x MW x DF x 10³

^{-- -------------------------------------------}

ɛ x1

A = (A520nm – A 700nm) pH 1.0 – (A520nm – A700nm) pH 4.5

MW (molecular weight) = 449.2 g/mol for cyanidin-3-glucoside

DF = dilution factor

ɛ = is the molar absorptivity of cyanidin-3-glucoside (26,900)

1 = cell path length in (1cm)

10³ = factor for conversion from g to mg¹⁵.

Effect of pH:

For studying effect of pH, anthocyanins were diluted with distilled water. The effect of pH variation on the stability of anthocyanins was studied on a wide range of pH values viz; 1.5, 2.5, 3.5, 4.5, 5.5, 6.5, 7.5, 8.5, 9.5, 10.5. pH of solution was adjusted using 0.1 N HCl and 0.1 N NaOH. Absorption spectra were recorded with UV–visible spectrophotometry¹⁶.

Copigmentation studies:

Purified Hibiscus sabdariffa anthocyanin was diluted with distilled water and was then mixed with phenolic acids like gallic acid, tannic acid, feullic acid and ellagic acids and with flavonoids like rutin and quercetin in different concentrations between anthocyanins and copigments viz. 1:0, 1:2, 1:4, and 1:6 at different pH values. Solutions were further sonicated to dissolve copigments. The result was characterized with UV-visible spectrophotometry. Those pigment: copigment ratio will show higher absorption will be used for further study¹⁷.

Effect of temperature:

For thermal stability colorant solution with and without copigment (10 ml) in glass tubes with screw caps were placed in a water bath at 60^oC for 2 hours. Absorbance of the anthocyanin solutions with and without copigment were recorded with UV–visible spectrophotometry¹⁸.

RESULT AND DISCUSSION:

The Hibiscus sabdariffa anthocyanins was extracted in acidic media. Adding acid to the extraction medium i.e. lower pH value had a great effect in stabilizing anthocyanins, and increasing extraction efficiency. The yield of extract was 1560 mg/100gm. Preliminary phytochemical screening revealed the presence of flavonoids and phenolic compounds. The extract was again confirmed for the presence of anthocyanin with 2M NaOH. On addition of NaOH, the initial red color changed blue to green. The total anthocyanin content was found to be 452.53 mg/100 gm. Based on our results, stability of anthocyanin pigment is greatly affected by pH. The results demonstrate evidently that increasing in pH causes greater destruction to anthocyanin in samples as indicated by UV-vis absorption spectra (Fig.1). Hibiscus sabdariffa anthocyanins show higher stability in acidic pH. With increasing pH value of the extracts above 3.5, the colour greatly faded and almost appeared colorless. Highest absorption was observed at pH 2.5 indicated the possibility of using the extract as food colorants in acidic food products. Copigmentation involves hydrophobic π–π molecular interaction, through a vertical stacking, between a planar anthocyanin structure and another planar colorless molecule which results in an enhancement generally an improvement in the original color of the pigment containing solution. The color produced by the anthocyanins in these copigmentation complexes can be many folds that of the original, and the actual enhancement depends primarily on the nature of the pigment, pH, temperature and the ratio of pigment to copigment. The steric conformation of anthocyanin-co-pigment complexes, i.e. the sandwich-type stacking of the phenolic acid moiety superposing the planar anthocyanin molecule, widely prevents or hinders the deteriorative nucleophilic attack of water. The result of the addition of copigments at increasing concentration showed that the copigmentation is solely dependent on copigment concentration. This leads to bathochromic and hyperchromic shifts that confirms copigmentation of anthocyanin in the solutions. Copigmentation study explains that, copigmentation of Hibiscus sabdariffa anthocyanins with all phenolic acids and flavonoids shown hyperchromic shift as illustrated. (Fig.2,3,4,5,6). In ferulic acid copigmentation at pH 2.5, both hyperchromic and bathochromic shift were observed (Fig.8). So, ferulic acid copigmented anthocyanins at pH 2.5 was proceeded to study effect of of higher temperature (60°C) on Hibiscus sabdariffa anthocyanins and results were compared.Temperature is an important factor that affects anthocyanin stability. There is speedy destruction of anthocyanin with increasing temperature. Anthocyanins degrade at higher temperature which is evident by the decrease in absorbance. Fig. 11 reveals that, high temperature and heating period (2 hr) affect stability of anthocyanins. Copigmented anthocyanins with ferulic acid shown higher absorbance and increase in heat stability as compared to natural anthocyanins. Hence, results suggests that, intermolecular copigmentation of Hibiscus sabdariffa anthocyanins with ferulic acid shown significant heat stability.

Fig.1 : Absorption spectra of Hibiscus sabdariffa at diff. pH

Fig.2 : Absorption spectra of Hibiscus sabdariffa -gallic acid at pH 2.5

Fig.3: Absorption spectra of Hibiscus sabdariffa ellagicacid at pH 2.5

Fig.4: Absorption spectra of Hibiscus sabdariffa –tannic acid at pH 2.5

Fig.5: Absorption spectra of Hibiscus sabdariffa –rutin at pH 2.

Fig.6: Absorption spectra of Hibiscus sabdariffa –Quercetin at pH 2.5

Fig.7: Absorption spectra of Hibiscus sabdariffa –ferulic acid at pH 1.5

Fig.8: Absorption spectra of Hibiscus sabdariffa –ferulic acid at pH 2.5

Fig.9: Absorption spectra of Hibiscus sabdariffa –ferulic acid at pH 3.5

Fig.10: Absorption spectra of Hibiscus sabdariffa –ferulic acid at pH 4.5

Fig. 11: Heat stability of Hibiscus sabdariffa natural anthocyanin and ferulic acid copigmented anthocyanin at 60°C

CONCLUSION:

Application of new technologies in the food industry requires untraditional processing of foodstuffs with the aim to improve their quality, durability, storage, nutritional value and visual attraction. The result of present study shows that ferulic acid in 1: 6 concentration has significant ability to stabilize the color of anthocyanins of Hibiscus sabdariffa. Ferulic acid copigmented anthocyanins showed better heat stability as compared to natural anthocyanins. This clearly indicates the formation of a copigmentation complex between ferulic acid and anthocyanins which was demonstrated by hyperchromic and bathocromic shifts. The present study shows that, the process of copigmentation between Hibiscus sabdariffa anthocyanins and the ferulic acid included in the present study is depends on chemical structure of copigments, its concentration, pH of the medium and temperature. Thus industrial processes which use to plan this phenomenon for the improvements of color of their products should take into account strong influence of above factors in all stages of product cycle viz; processing, commercialization, storage etc.

ACKNOWLEDGEMENT:

Authors are thankful to Dr. S. S. Bhujbal, H.O.D, Dept. of Pharmacognosy, Dr. DYPIPSR, Pimpri, Pune, Maharashtra, for providing inspiration and motivational support.

CONFLICT OF INTEREST:

Authors do not have any conflict of interest.

REFERENCES:

1. Rezaei M. Assessment of Synthetic Dyes in Food Stuffs Produced in Confectioneries and Restaurants in Arak. Iran. J. of Synthetic Thrita. 2015, 4(1): 1-5. http://dx.doi.org/ 10.18869/IAHS.3.2.61

2. Eliana FO, Paulo CS, and Milton CC. Stability of anthocyanin in spinach vine (Basella rubra) fruits. Cien. Inv. Agr. 2007; 34(2): 115-120.

3. Naik P. Food colourants and health issues: are we aware?.Current Science. 2014; 106(2): 25-30.

4. Lauro GJ. A primer on natural colors. Am. Assoc. Cereal Chem 1991; 36(11) : 949-953.

5. Mohamad MF, Nasir SN, Sarmadi MR. Degradation kinetics and color of anthocyanins in aqueous extracts of butterfly pea. Asian Journal of Food and Agro industry. 2011. 4(5):306-315.

6. Parmar M, Gupta U. Biocolors: The New Generation Additives. Int. J. Curr. Microbiol. App. Sci. 2015; 4(7): 688-694.

7. Ghasemifar E, Saeidian S, Setareh P. Stability of seedless red grape anthocyanin under the effect of Chlorogenic acid copigment, heating and UV irradiation. Journal of Novel Applied Sciences. 2013; 2(11):594-597.

8. Boulton R. The Copigmentation of Anthocyanins and Its Role in the Color of Red Wine: A Critical Review. Am. J. Enol. Vitic. 2001; 52 (2) : 67-87.

9. Abyari M, Hedari R, Jamei R. The effect of Heating, UV Irradiation and pH on Stability of Siahe Sardasht Grape Anthocyanin-copigment Complex. Journal of Biological Sciences. 2006; 6(4):638-645.

10. Mahadevan N, Shivali K, Pradeep K. Hibiscus sabdariffa Linn.–An overview. Natural Product Radiance. 2009; 8(1): 77-83.

11. Puro K, Sunjuktar R, Samir S, Ghatak S. Medicinal Uses of Roselle Plant (Hibiscus sabdariffa L.): A Mini Review. Indian Journal of Hill Farming. 2016; 27(1): 81-90.

12. Obouayeba A, Diabate S, Joseph D. Phytochemical and antioxidant activity of Roselle. Research J. of Pharmaceutical, biological and Chemical sciences.2014; 5(2):1453-1465.

13. Harborne, J.B. Phytochemical methods: A Guide to Modern techniques of plants Analysis. Chapman and Hall London., U.K. 1998; pp : 64.

14. Rakkimuthu R, Palmurugan S, Shanmugapriya A. Effect of temperature, light, ph on the stability of anthocyanin pigments in cocculus hirsutus fruits. International Journal of Multidisciplinary Research and Modern Education. 2016; 2(2): 91-96.

15. Rasha K, Mohamed AY, Nagwa MH, Rasmy FM, Abu-Salem. Extraction of Anthocyanin Pigments from Hibiscus sabdariffa L. and Evaluation of their Antioxidant Activity. Middle East Journal of Applied Sciences. 2016; 6(4) : 856-866.

16. Kuntapas K, Singh K, Phetkao S. Effect of pH and anthocyanin concentration on color and antioxidant activity of Clitoria ternatea. J. of Food and Applied Bioscience. 2014; 2(1) :31-46.

17. Munawaroh H, Fadillah G, Saputri, Hanif Q, Hidayat R, Wahyuningsih S. The co-pigmentation of anthocyanin isolated from mangosteen pericarp (Garcinia Mangostana L.) as Natural Dye for Dye Sensitized Solar Cells (DSSC). J. of Materials Science and Engineering. 2016; 107: 1-6.

18. Elham G, Shahriar S, Setareh P. Stability of Seedless Red grape anthocyanin under the effect of Chlorogenic acid copigment, heating and UV irradiation. Journal of Novel Applied Sciences. 2013; 2(11) : 595-597.

Received on 24.01.2019 Modified on 23.02.2019

Research J. Pharm. and Tech. 2019; 12(6):2949-2954.

DOI: 10.5958/0974-360X.2019.00496.7