Bathochromic and Hyperchromic Effects of Aluminum Salt

Article pubs.acs.org/JAFC

Bathochromic and Hyperchromic Effects of Aluminum Salt Complexation by Anthocyanins from Edible Sources for Blue Color Development Gregory T. Sigurdson Department of Food Science and Technology, The Ohio State University, 2015 Fyffe Court Road, Columbus, Ohio 43210-1007, United States

M. Monica Giusti* Department of Food Science and Technology, The Ohio State University, 110 Parker Food Science and Technology Building, 2015 Fyffe Court Road, Columbus, Ohio 43210-1007, United States ABSTRACT: Use of artificial food colorants has declined due to health concerns and consumer demand, making natural alternatives a high demand. The effects of Al3+ salt on food source anthocyanins were evaluated with the objective to better understand blue color development of metalloanthocyanins. This is one of the first known studies to evaluate the effects of food source anthocyanin structures, including acylation, with chelation of aluminum. Cyanidin and delphinidin derivatives from different plants were treated with factorial excess of Al3+ in pH 3−6 and evaluated by spectrophotometry and colorimetry over 28 days. Anthocyanin concentration, salt ratio, and pH determined final color and intensity. Pyrogallol moieties on delphinidin showed furthest bathochromic shifts, whereas acylation promoted higher chroma. Blue color developed at lower pH when acylated anthocyanins reacted with Al3+; hue ∼270 occurred with acylated delphinidin at pH ≥2.5. Highest chelate stability was found with AlCl3100−500× anthocyanin concentration. This investigation showed anthocyanin−metal chelation can produce a variety of intense violet to blue colors under acidic pH with potential for food use. KEYWORDS: metalloanthocyanin, anthocyanin−metal complex, metal−chelate complex, anthocyanin, Al3+, natural blue pigments, Solanum melongena L, Rubus idaeus, Brassica oleracea var. capitata f. rubra, Ribes nigrum, Daucus carota spp. sativus, Aronia melanocarpa

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INTRODUCTION Color relates consumer perception to quality and flavor of food products, making it a key to a product’s overall success. Food colorants have several uses in food including color enhancement, giving a color identity to colorless foods, such as margarine, and accounting for color loss during storage.1 The use of artificial colorants has become less desirable due to health concerns and consumer demand for natural products. Artificial food colorants are increasingly becoming linked with allergies, with potential cancer development, and with hyperactivity problems in children.1 The European Union (EU) has already enforced the use of warning labels indicating that synthetic colorants may cause hyperactivity in children, and the U.S. Food and Drug Administration (FDA) has recommended the safety of synthetic dyes be reviewed.2 The development of natural alternatives for food colorants has become a very current and important topic for food safety and for food companies to remain competitive in an international market. Frequently responsible for reds, blues, and purples seen in nature, anthocyanins are a class of naturally derived food pigments that may also impart beneficial health effects, making them viable alternatives for synthetic colorants. Although blue dyes are less used than other hues, there are currently few natural options.1 Those with commercial feasibility have been derived from blue gardenias, huito fruit, and Spirulina spp., © XXXX American Chemical Society

which was approved only for confectionary uses in September 2013.3 Food application data of these colorants are relatively scarce.4 In acidic conditions common to food products, anthocyanins typically appear in red or purple molecular forms. However, self-association, copigmentation, and metal complexes can result in acid-stable, blue colorations, like those found in flowers.5 Most metal−anthocyanin interactions have been studied for better understanding of plant and floral pigmentation.4 For anthocyanins to undergo metal complexation, more than one free hydroxyl group must be present on the B ring.4,6,7 Figure 1 shows catechol and pyrogallol moieties on the B ring of cyanidin and delphinidin, respectively. Multivalent metal ions act in competition with the hydrogen ions attached to these rings, inducing their loss and transforming the flavylium cation to a quinoidal base (Figure 1).6 Simultaneously, a stacking association with another anthocyanin flavylium ion occurs with the transformed molecule, forming a metal-coordinated complex.6 Various metal ions are known to induce this effect, Special Issue: International Workshop on Anthocyanins (IWA2013) Received: November 14, 2013 Revised: February 11, 2014 Accepted: February 18, 2014

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dx.doi.org/10.1021/jf405145r | J. Agric. Food Chem. XXXX, XXX, XXX−XXX

Journal of Agricultural and Food Chemistry

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Figure 1. Al3+ chelation by cyanidin (B ring exhibiting catechol moiety) or delphinidin-3-p-coumaroyl-rutinoside-5-glucoside (B ring exhibiting pyrogallol moiety), where R1 is H or R1 is OH, respectively.

all being multivalent.7 Al3+ anthocyanin complexes in ethanol have been reported to be relatively stable at pH 2−5 in reactions with delphinidin.6 Cyanidin and delphinidin glucosides from berries had previously been found to develop bluelike hues with tin chelation, whereas iron chelates were more brown.8 More recent publications emphasized the effects of B ring structure, acidic range pH, and buffer selection in anthocyanin complexation with trivalent metal ions (Fe3+ and Al3+) in pectin and pectic fractions.4 Unlike previous findings, ferric ion treated delphinidin-3-glucoside showed blue hues and farther bathochromic shifts than cyanidin and petunidin glucosides. These samples also showed highest stability over time.4 The environment also can affect the stability of anthocyanin−metal chelates. In citrate and phosphate buffers, color shifts were not found to occur; however, metal chelation by anthocyanins seemed relatively unhindered in other buffers such as acetate and succinic acid.4 Aluminum has traditionally been considered innocuous to humans, but controversy has risen with links to neurotoxicity, dialysis encephalopathy, microcytic anemia, osteomalacia, and Alzheimer’s disease.9,10 However, no reports of dietary aluminum toxicity were found in the literature in healthy individuals.11 The FDA considers aluminum sulfate as generally recognized as safe (GRAS) as a food additive with no usage limit, and aluminum salts may comprise 2% of the final product in synthetic lake type dyes.12,13 Absorption from anthocyanin− aluminum colorants is likely low, based on anthocyanin chelating effects and aluminum’s low bioavailability in water (0.3%).9,11,14 With still limited knowledge about the conditions supporting anthocyanin−metal chelation, the aim of this study was to explore factors conducive to blue color formation in acidic conditions typical of food products. This is one of the first known studies to evaluate the effects of food source anthocyanin structures, including acylated and nonacylated counterparts of delphinidin and cyanidin, with chelation of aluminum in varied conditions. The effects of differing anthocyanin concentrations and increasing concentrations of aluminum salt were evaluated to determine ideal conditions for complexation leading to most intense bathochromic and hyperchromic shifts in absorbance. These conditions were also carried out in the pH range 3−6 to better understand the role of pH in anthocyanin−metal chelation. Color stability was monitored by a 28 day study of delphinidin and acylated cyanidin chelates in different storage conditions.

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local grocery store in Columbus, OH, USA. Frozen whole black currants (Ribes nigrum) were purchased from CropPharms LLC (Staatsburg, NY, USA). A commercial black carrot (Daucus carota spp. sativus) anthocyanin powder was provided from D. D. Williamson (Louisville, KY, USA), and chokeberry (Aronia melanocarpa) juice concentrate was obtained from Artemis International (Fort Wayne, IN, USA). Aluminum chloride hexahydrate, USP 97.0−101.0% grade, was purchased and obtained from Sigma-Aldrich Co. (St. Louis, MO, USA). ACS grade sodium acetate anhydrous, 6 N hydrochloric acid (certified 5.95−6.05), trifluoroacetic acid, and sodium hydroxide N/10 (0.0995−0.1005) were purchased from Fisher Scientific (Fair Lawn, NJ, USA) as were all other standard ACS and HPLC grade reagents. For comparative purposes, powdered forms of brilliant blue or FD&C Blue No. 1 (certification no. AL9925) and indigotine or FD&C Blue No. 2 (certification no. AL2889), each having a color purity of 92%, were obtained from Noveon Hilton Davis, Inc. (Cincinnati, OH, USA). FD&C Blue No. 1 was diluted to 2.14 × 10−4 M with distilled water, whereas FD&C Blue No. 2 was diluted to 6.35 × 10−4 M. Methods. Extraction and Purification of Anthocyanins. Extraction of anthocyanins from plant materials followed procedures described by Rodriguez-Saona and Wrolstad.15 Whole plant samples (only peels from eggplant samples) were powdered with liquid nitrogen and treated with 0.01% HCl acidified 70% aqueous acetone before filtration, with repetition until plant material was discolored. Eggplant peels were instead treated with 3.0% trifluoroacetic acid acidified 70% aqueous acetone to reduce the brown discoloration, likely due to polyphenol oxidase, that occurred during extraction. After the filtrate was mixed with 1−2 volumes of chloroform, phase separation occurred overnight at 4 °C, except eggplant samples being limited to 4 h. The chloroform layer containing lipophilic compounds was appropriately discarded because chloroform is a mild carcinogen. Care should be taken to avoid introduction of bases to chloroform waste to avoid undue explosions. Aqueous layers were dried in a rotary evaporator at ∼37 °C under vacuum. Anthocyanin Purification: Solid Phase Extraction. Crude anthocyanin extracts were purified by loading activated Waters Seppak C18 cartridges with samples. Loaded cartridges were washed with water acidified to 0.01% HCl to remove organic sugars and acids and then with ethyl acetate to remove phenolics. Anthocyanin pigments were recovered with 0.01% HCl acidulated methanol, which was removed in a rotary evaporator at 37 °C under vacuum. Pigments were stored in acidified water until further analysis. Monomeric Anthocyanin Quantitation. Monomeric anthocyanins were quantitated by the pH differential method, described by Giusti and Wrolstad.16 Briefly, anthocyanins were quantified by measuring the absorbance at 520 and 700 nm at pH 1 and 4.5 for each sample. On the basis of the difference of these absorbances, the concentration of monomeric anthocyanins of the solutions was determined. This information was then used to make dilution to known concentrations. Monomeric anthocyanin pigments were quantitated using the formula ACN (mg/L) = (A × MW × DF × 1000)/(ε × 1), with 1 cm path length. A was the absorbance, based on (Aλ520 − Aλ700)pH 1 − (Aλ520 − Aλ700)pH 4.5. MW was the molecular weight of the sample’s predominant anthocyanin, DF was the dilution factor used, and ε was

MATERIALS AND METHODS

Materials. American eggplant (Solanum melongena L), Japanese eggplant (Solanum melongena L), red raspberry (Rubus idaeus), and red cabbage (Brassica oleracea var. capitata f. rubra) were purchased from a B



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the molar absorptivity. Table 1 shows the values used for each sample’s quantitation.

2000× anthocyanin concentrations. Three replicates were evaluated for each sample in each storage condition, representing different product storages. Samples were stored for 28 days in each condition: dark storage at refrigeration temperatures (2−4 °C), dark storage at ambient temperatures (18−25 °C), and full light exposure from natural daylight ∼8 h and fluorescent lamps from above at ambient temperatures (18−25 °C). (c) Evaluating the Effect of pH on Al3+ Complex Formation. Sodium acetate was used to prepare 1 M buffered solutions at pH 3, 4, 5, and 6. Anthocyanin samples from American eggplant, Japanese eggplant, chokeberry, and red cabbage were diluted to 50 μM solutions in each buffer. AlCl3 solutions were added to achieve factorial excesses of 0×, 1×, 100×, 500×, 1000×, and 2000× over anthocyanin concentration. The pH of these and all samples was measured using a Mettler Toledo International Inc. S220 SevenCompact pH/ion meter (Schwerzenbach, Switzerland). Spectrophotometry of Solutions with UV−Visible Transmission. Each sample from each phase of study was subjected to visible spectrophotometry and colorimetry. After 15 min of equilibration after salt addition, each sample was evaluated by visible transmittance (400−700 nm) spectrophotometry using 1 cm plastic cuvettes in a Shimadzu UV-2450 UV−visible spectrophotometer. Spectrograms were generated using UV Probe software, version 2.21 (Shimadzu) associated with this spectrophotometer model. CIE-Lab Color of Solutions by Transmission. Samples were transferred to 2 mm path length plastic cells and read for CIE-Lab, chroma, and hue angle using a Hunter ColorQuest XE (Hunter Laboratories, Reston, VA, USA). The equipment was set for total transmittance, illuminant D65, and a 10° observer angle for all liquid samples. Statistical Evaluation of Data. The mean and standard deviation of data replicates were calculated using Microsoft Office Excel 2010 (Office 14.0, Microsoft, Redmond, WA, USA). Data from evaluation of anthocyanin structures, concentrations, and salt ratios in Al3+ complex formation were conducted by one-way analysis of variance (ANOVA) (two-tailed, α = 0.05) and Student’s paired t test (two-tailed, α = 0.05) of λmax and associated absorbance comparing different anthocyanin concentration and increasing AlCl3 concentration (0−1000× anthocyanin concentration) using Microsoft Office Excel 2010. Stability over time of American eggplant and red cabbage anthocyanin−aluminum chelates was evaluated by the same testing on the λmax and associated hue from the different salt treatments in differing storage conditions. Finally, the pH decreases likely caused by Al3+ chelation were also evaluated by one-way ANOVA (two-tailed, α = 0.05). All other figures were also generated using this software.

Table 1. Display of Anthocyanin, Associated Molecular Weight, and Molar Absorptivity Used for Monomeric Quantitation of Anthocyanins in Food Samples Evaluated sample American eggplant Japanese eggplant black currant red raspberry chokeberry red cabbage black carrot

anthocyanin

MW

molar absorptivity

delphinidin-3-rutinoside

647.0

26900a

delphinidin-3-(pcoumaroylrutinoside)-5-glucoside cyanidin-3-glucoside cyanidin-3-glucorutinoside cyanidin-3-glucoside cyanidin-3- diglucoside-5-glucoside cyanidin-3-xylosyl(feruloylglucosyl) galactoside

955.26

26900a

484.8 757.0 484.8 773.0 919.25

26900a 26900a 26900a 30175 26900a

a

Molar absorptivity of cyanidin-3-glucoside, used for monomeric quantitation.16

High-Pressure Liquid Chromatography (HPLC) Evaluation of Anthocyanins. Reverse phase HPLC was utilized to identify anthocyanins, with comparison to the literature. Samples were analyzed using a HPLC (Shimadzu, Columbia, MD, USA) system equipped with LC-20AD pumps and an SIL-20AC autosampler coupled to an LCMS-2010 mass spectrometer (Shimadzu) and an SPD-M20A photodiode array (PDA; Shimadzu) detector. LCMS Solution software (version 3, Shimadzu) was used to view results. Separation of anthocyanins was achieved on a reverse-phase Symmetry C18 column with 5 μm particle size and 4.6 × 150 mm column size (Waters Corp., Taunton, MA, USA), and a 4.6 × 22 mm Symmetry 2 micro guard column (Waters Corp., Bedford, MA, USA) was used. Samples were filtered through Phenomenex Phenex RC 0.2 μm, 15 mm membrane syringe filter (Torrance, CA, USA). The flow rate was set to 0.8 mL/min with a run time of 70 min, and PDA detection for 55 min. The solvents were phase A, 4.5% formic acid in LC-MS grade water, and phase B, LC-MS acetonitrile (Fisher Scientific Inc.). A binary gradient was used for solvent B: 0−35 min for 0−25% B, 35− 40 min for 25−50% B, 40−45 min for 50−100% B, and returning B to 0% 50−55 min and maintained until 70 min. Spectral data were obtained from 250 to 700 nm, and elution of anthocyanins was monitored at 520 nm. About 0.2 mL/min flow was diverted to a single-quadrupole iontunnel mass spectrometer equipped with an electrospray ionization (ESI) interface (Shimadzu). Mass spectrometry was performed under positive ion mode. Data were monitored using total ion scan (SCAN) (from m/z 200−1200) and selected ion monitoring at m/z 287 (cyanidin) and m/z 303 (delphinidin). Formation of Complexes with Aluminum and Metal Salts in Acidulated Aqueous Solutions. (a) Evaluating Anthocyanin Structures, Concentrations, and Salt Ratios in Al3+ Complex Formation. To solutions of acidified water at pH 3 were added anthocyanin samples at three concentrations, based on monomeric quantitation: 25, 50, and 100 μM. Anthocyanin sources for this phase included American eggplant, black currant, red raspberry, chokeberry, red cabbage, and black carrot. AlCl3 aqueous solutions were prepared to 0.057, 0.2288, and 0.4576 M. Salt solutions were added to anthocyanin solutions beginning at equal molar concentrations, then in factors of 10× to 100× molarity of anthocyanin content, then 500× and 1000×. Control samples for each concentration were maintained without salt addition for comparison. Three replicates were evaluated for each sample. (b) Evaluating Color Stability over Time of Al3+ Complexed Anthocyanins. Anthocyanins from American eggplant and red cabbage were diluted in acidified water at pH 3 to 50 μM concentration. AlCl3 salt solutions were added to anthocyanin solutions in factorial excesses of 0×, 1×, 100×, 500×, 1000×, and

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RESULTS AND DISCUSSION Evaluating Results from Reverse Phase HPLC-MS. Food samples used in this study represented two anthocyanin aglycone structures: delphinidin and cyanidin. American eggplant and black currant were chosen to represent glycosylated delphinidin. HPLC chromatograms are presented in Figure 2. The American eggplant showed delphinidin-3rutinoside as the primary anthocyanin as indicated from its HPLC chromatogram and mass spectrometry and in agreement with previous literature.17 Black currant chromatograms showed profiles similar to previous findings: delphinidin-3rutinoside and cyanidin-3-rutinoside, each representing ∼40% of the anthocyanin profile.18 Japanese and non-Japanese type eggplants share delphinidin as the primary anthocyanidin; however, the primary Japanese eggplant anthocyanin was acylated, delphinidin-3-(p-coumaroylrutinoside)-5-glucoside (or nasunin) (Figure 2).17 Sources and structures were more varied for cyanidin derivatives, being more prevalent in edible produce. Red raspberry and chokeberry were used as sources for glycosylated cyanidin. The main pigment in chokeberry was found to be C



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or acylation. Each anthocyanin demonstrated both bathochromic and hyperchromic shifts when exposed to the metal ion. By comparison of B ring core structures, delphinidincontaining samples showed the farthest bathochromic shifts and become most blue-like at pH 3 (Table 2). Glycosylated delphinidin showed λmax bathochromic shifts of 50 and 55 nm for American eggplant and black currant, respectively. Cyanidin samples showed lower bathochromic shifts ranging from 26 to 39 nm. These data indicate that increasing the number of free hydroxyl groups on the aglycone core structure leads to farther bathochromic shifts, agreeing with previous studies comparing petunidin, cyanidin, and delphinidin.3 Amount and type of common glycosylation do not seem to affect anthocyanin ability to chelate with metal ions but may mildly affect final solution color. When the two glycosylated cyanidin samples were compared, red raspberry anthocyanins demonstrated lower λmax than chokeberry anthocyanins until high levels of Al3+ were achieved; Table 2 shows λmax values of red raspberry anthocyanins to be about 10 nm less than those of chokeberry anthocyanins with AlCl3 100× anthocyanin content. However, both samples showed λmax of 551−553 nm with AlCl3 500× anthocyanin content, with the exception of the chokeberry anthocyanins in 100 μM concentrations, which shifted to 567 nm but with constant final hue across all three anthocyanin concentrations. This occurrence did not follow the trend of other anthocyanin samples or concentrations. Perhaps due to the high anthocyanin content and high proportion of monoglycosylated cyanidin, additional intermolecular copigmentation could have occurred, leading to the shift in λmax. This postulate does not explain why other anthocyanin samples did not exhibit the same trend, unless their individual complexes had already formed or were hindered related to additional structural components. With acylation, cyanidin samples continued showing intense bathochromic and hyperchromic shifts in absorbance. The acylated samples did not seem to show larger hyperchromic shifts than nonacylated counterparts but did exhibit more intense chroma. The overall higher absorbances could be due to the proposed molecular stacking mechanisms of acylated anthocyanins, in which the acid groups are overlaid the anthocyanins.23 This would increase the amount of electron density in that area, leading to intensification of signals found in spectrophotometry. Although the Al3+ complex colors of the two acylated cyanidin samples followed similar trends of absorbance increases, acylated cyanidin samples from red cabbage showed larger absorbance increases than acylated cyanidin pigments from black carrot (Table 2). Red cabbage anthocyanins also exhibited more blue-like colors, having a hue angle around ∼298, whereas black carrot anthocyanin Al3+ chelates exhibited hue angles at ∼316. These data suggest that the long sugar chain of black carrot anthocyanins may interfere with the molecular stacking arrangement, resulting in lower λmax and fewer and less intense blue hues, as compared to red cabbage chelates. Regardless of anthocyanin concentration (25−100 μM), all samples chelated Al3+, showing bathochromic and hyperchromic shifts. However, the effects of increasing salt concentration (0−1000×) were less in lower anthocyanin concentration samples. At lower concentrations with the same factorial excesses of salt, samples demonstrated lesser bathochromic and hyperchromic shifts in absorbance compared to higher anthocyanin concentrations (Figures 3 and 4), until reaching optimum salt ratios where the same λmax and hue angle

Figure 2. Reverse phase HPLC chromatograms of food sample anthocyanins: (A) American eggplant; (B) Japanese eggplant; (C) black currant; (D) chokeberry; (E) red raspberry; (F) red cabbage; (G) black carrot, with detection at 520 nm showing the main pigments identified.

cyanidin-3-galactoside, representing ∼65% of the anthocyanin profile.19 The chromatogram of red raspberries of this study showed four major pigments; most prevalent were cyanidin-3sopohoroside and cyanidin-3-glucorutinoside.20 Two sources of acylated cyanidin were also included in this study: red cabbage and black carrot. Several peaks for red cabbage were found, primarily various types and amounts of acylated of cyanidin-3sopohoroside-5-glucoside.21 Black carrot chromatograms showed five major peaks, resembling the profile of the ‘Deep Purple’ variety.22 The major pigment of this variety was cyanidin-3-xylosyl(feruloylglucosyl)galactoside, agreeing with the literature.22 Evaluating Anthocyanin Structure, Concentrations, and Salt Ratios in Al3+ Complex Formation. In this portion of the study, samples were limited to sources of glycosylated delphinidin (American eggplant and black currant), glycosylated cyanidin (red raspberry and chokeberry), and acylated cyanidin (red cabbage and black carrot). This was intended to determine what role anthocyanin structure, concentration, and salt ratio played in metal complexation to induce bathochromic shifts toward blue colors. All tested anthocyanins showed complexation to occur with aluminum, regardless of aglycon structure, type of glycosylation, D



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Table 2. Effects of Anthocyanin Concentration at pH 3 on Color Shifts, with λmax and CIE-Lab Color Characteristics of Anthocyanins Complexed with AlCl3 100× Anthocyanin Concentrationa color characteristics

a

sample

[ACN] (μM)

L

a*

b*

chroma

hue

Δ hue

American eggplant

25 50 100

89.3 (1.2) 86.1 (0.8) 80.3 (1.5)

2.7 (0.4) 3.6 (1.2) 4.4 (1.6)

−5.8 (0.5) −10.1 (3.2) −10.6 (2.1)

6.3 (0.7) 10.7 (3.4) 11.5 (2.5)

295.2 (1.5) 289.4 (0.5) 291.9 (4.0)

279 284 264

black currant

25 50 100

89.9 (1.5) 81.1 (1.7) 71.7 (1.0)

5.0 (0.7) 11.1 (1.4) 17.3 (0.9)

−5.0 (0.9) −13.3 (3.1) −19.3 (1.0)

7.1 (0.8) 17.4 (3.2) 25.9 (1.5)

314.7 (7.0) 310.2 (4.1) 311.9 (0.0)

299 293 293

red raspberry

25 50 100

89.8 (0.1) 84.6 (2.6) 74.3 (0.8)

7.9 (1.0) 14.1 (2.4) 27.0 (1.6)

−5.0 (0.3) −9.7 (1.9) −16.1 (1.7)

9.2 (1.2) 17.1 (3.1) 31.3 (2.2)

329.1 (0.9) 325.6 (1.2) 329.3 (1.1)

320 317 318

chokeberry

25 50 100

88.1 (1.0) 83.3 (1.4) 73.5 (0.3)

9.0 (0.6) 13.2 (1.0) 22.7 (0.9)

−4.6 (0.7) −8.6 (1.5) −15.2 (0.5)

10.2 (0.8) 15.8 (1.7) 25.6 (1.4)

332.8 (2.8) 327.1 (2.5) 325.8 (0.1)

318 311 310

red cabbage

25 50 100

89.1 (0.9) 81.0 (0.6) 70.1 (3.6)

4.8 (0.9) 9.0 (1.9) 15.1 (3.5)

−7.7 (0.8) −16.9 (1.2) −28.2 (4.0)

9.1 (0.9) 19.2 (1.9) 30.4 (5.4)

301.6 (4.7) 297.8 (3.4) 297.9 (2.1)

−44 −47 −48

purple carrot

25 50 100

90.2 (0.8) 83.7 (0.8) 72.8 (0.3)

6.8 (0.2) 12.5 (0.1) 22.6 (0.9)

−5.0 (0.6) −11.9 (1.0) −22.4 (0.9)

8.4 (0.3) 17.3 (0.7) 29.0 (0.4)

323.6 (4.0) 316.4 (2.6) 315.2 (2.1)

−36 −42 −45

In parentheses are the standard deviations, n = 3.

Figure 3. Visible absorbance (400−700 nm) of 25, 50, and 100 μM concentrations of American eggplant anthocyanins treated with factorial increases of AlCl3 (0−1000×) over anthocyanin concentration at pH 3.

occurred. From the same anthocyanin source, the λmax means resulting from each combination of salt treatment (0−1000×) and anthocyanin concentration were compared. p values of 6.64 × 10−8 or less were obtained from one-way ANOVA, indicating mean λmax from each AlCl3 treatment to be different from anthocyanins from the same sample. Similar results were obtained from one-way ANOVA on absorbance of the same samples, giving p values of 1.82 × 10−6 or less, with the exception of 25 μM red raspberry anthocyanins. ANOVA resulted in p value of 0.05, for which

the null hypothesis was still rejectable at 90% confidence. Typically, the increase in absorbance continued to rise with increasing anthocyanin content. Largest absorbance increases were found to occur with anthocyanins from black currants, which are roughly equal ratios of glycosylated delphinidin and cyanidin.18 The hue reflects this anthocyanin profile, falling between the complex hues of either primarily delphinidin or cyanidin. With regard to the atypically high absorbance increases of black currant anthocyanins, responsibility could be attributed to intermolecular copigmentation between the E



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Figure 4. Visible absorbance (400−700 nm) of 25, 50, and 100 μM concentrations of red cabbage anthocyanins treated with factorial increases of AlCl3 (0−1000×) over anthocyanin concentration at pH 3.

Figure 5. Changes in absorbance at λmax of 50 μM anthocyanin solutions at pH 3 over 28 days of AlCl3 (0−1000×) treated (A) delphinidin (American eggplant anthocyanins) with dark storage at 4 °C, (B) delphinidin with dark storage at ambient temperatures (19−25 °C), (C) delphinidin with light storage at ambient temperatures (19−25 °C), and (D) acylated cyanidin (red cabbage anthocyanins) with light storage at ambient temperatures (19−25 °C).

concentrations were found to be between 100× and 500× anthocyanin concentration, at which treated anthocyanins showed the highest intensity of absorbance and reached the farthest bathochromic shift (Figures 3 and 4). Comparing the λmax means from AlCl3 concentrations 100× and 500× anthocyanin content of 25 μM by Student’s paired t test, p values of 0.02−0.09 were obtained. The λmax values for these

roughly equal ratios of cyanidin and delphinidin not found in other samples. In addition to anthocyanin structure and concentration, AlCl3 salt ratios were investigated to determine optimal ratios for preferred color development. Anthocyanin chelation with Al3+ was apparent essentially immediately with ion introduction to the system, causing visible color changes. Ideal AlCl3 salt F



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they did not differ significantly from dark storage samples, having p values of 0.1 and 0.5 from Student’s paired t test for AlCl3 levels 100× and 500× anthocyanin content. This indicates anthocyanin−metal complexes are more liable to heat treatment rather than light exposure. Degradation of delphinidin−Al chelates seems related to concentration with rate decreasing with time, in agreement with previous studies.3 Acylated cyanidin chelates showed more linear degradation patterns (Figure 5). These samples also showed improved color stability, exhibiting lower changes in absorbance over time, likely resulting from the proposed molecular folding and stacking acylated anthocyanins can undergo protecting the chromophore. Having used different ratios of salt to anthocyanin, it was possible to estimate concentrations for optimal anthocyanin chelate stability. Similarly to the decrease of absorbance noted with high excesses of AlCl3, a large amount of salt also was found to diminish stability. From one-way ANOVA, all chelated sample means for λmax were found to be different after 28 days, giving p values of ≤0.002 for each storage treatment. With salt 100−500× anthocyanin content, highest stability was noted on the basis of absorbance and hue angle. Those samples that received light and heat abuse showed AlCl3 100× to be slightly more stable than 500× based on hues, which differed by p value ≤0.02 from Student’s paired t test. At this anthocyanin salt ratio (100×) perhaps the largest number of stabilized and protected complexes occurs. Evaluating Effects of pH in Al3+ Complex Formation. Four samples were used in this phase of the study, American eggplant (glycosylated delphinidin), Japanese eggplant (acylated delphinidin), chokeberry (glycosylated cyanidin), and red cabbage (acylated cyanidin), to understand the effects of starting pH on final exhibited color. Decreases of overall solution pH were found to occur likely due to the release of hydrogen ions when anthocyanins chelate metals. At each of the starting pH levels tested (pH 3−6), all anthocyanins exhibited roughly equal decreases in pH in 1 M acetate buffer solutions (Figure 6). By one-way ANOVA, the pH decreases across each sample at starting pH levels were found not to differ by p values ≥0.23. These equivalent decreases in pH indicate the same concentration of H+ ions being released from both aglycones. It is suggested that acidic pH may not limit the ability of these anthocyanins to chelate metals, despite the form

two salt concentration ratios on American eggplant, black currant, and red cabbage anthocyanins were found not to differ, having p values >0.06. The other samples were found not to differ with AlCl3 concentrations 500× and 1000× anthocyanin content. The same results from t tests were obtained for 50 μM anthocyanin samples. For 100 μM anthocyanin samples, only chokeberry and red cabbage anthocyanin samples were similar with AlCl3 concentrations 100−500×, and the remainder was similar with AlCl3 concentrations 500−1000×. In addition to λmax, absorbance was evaluated by ANOVA and Student’s paired t test for optimal salt ratios. A large excess of salt seemed to initiate some reversion of the color intensification achieved at lower ratios (Figures 3 and 4); a decrease in complex intensity can be noted at salt 1000× anthocyanin concentration in all anthocyanin concentrations (25−100 μM). This could be due to metal-induced pigment degradation or complex formation hindrance caused by the high salt concentration. By ANOVA, the absorbances of all anthocyanin concentrations with salt treatment were found different with p values of ≤0.005, except 25 μM red raspberry anthocyanin samples, which were still the same at 90%. From Student’s t test, p values of ≤0.03 were found for all 50 and 100 μM anthocyanin samples with AlCl3 concentrations 500× and 1000× anthocyanin content, except 100 μM chokeberry (p value = 0.09), supporting the observation that absorbance was decreasing with high AlCl3 excesses of 1000×. Some slight decreases in absorbance were noted in 100 μM anthocyanins from American eggplant in the salt range of 30−90×, which did not follow general trends shown by other concentrations. On the basis of the largest absorbance and λmax observed in these samples, optimal AlCl3 concentration was found to be 100− 500× anthocyanin concentration. Evaluating Color Stability over Time (28 Days) of Al3+Complexed Anthocyanins. The mechanism for thermal degradation of anthocyanins has not been fully elucidated and therefore is still not understood (Schwartz and others, 2008). However, all three of the proposed pathways involve transformation of the flavylium cation to other forms more prevalent in higher pH, which then decompose into degradation products.24 It has been proposed that chelation of metals by anthocyanins stabilizes the quinonoidal base form as well as protects it from nucleophilic attack by complexation coordination.3,5 Metal chelation by anthocyanins has been shown to increase stability during storage as well as with thermal treatment.3,25 Ferric chelates of cyanidin-3-glucoside showed improved stability to thermal treatment of 60 °C over 80 min, with intensity retention of ∼70%.25 All chelates exhibited first-order kinetics of degradation, but those with delphinidin-3-glucoside in sugar beet pectin showed highest stability compared to glucosides of cyanidin and petunidin when stored at 20 ± 2 °C for 18−66 days.3 This study compared the stability of aluminum chelates of delphinidin-3-rutinoside (American eggplant) to that of acylated cyanidin glycosides (red cabbage), as the literature is limited about the stability of acylated anthocyanin−metal chelates. Highest stability for both samples was found in cold, dark storage as expected on the basis of ideal conditions for anthocyanin stability. For delphinidin chelates absorbance decreases were limited to ∼0.1 over 28 days, whereas almost no change was noted for acylated cyanidin samples (Figure 5). Much larger decreases in absorbance were noted for samples stored in ambient temperatures. Although those samples receiving light treatment showed slightly larger decreases,

Figure 6. pH change of 50 μM anthocyanin sample solutions initiated by treatment with AlCl3 (0−2000×) in 1 M sodium acetate buffers, pH 3, 4, 5, and 6. G



Article

Figure 7. Visible (400−700 nm) absorbance of 50 μM acylated delphinidin (Japanese eggplant anthocyanins) treated with AlCl3 0−2000× anthocyanin concentration at (A) pH 3, (B) pH 4, (C) pH 5, and (D) pH 6.

but were generally lower for samples starting at pH 3. Delphinidin samples showed greatest increases in λmax versus cyanidin anthocyanins; interestingly, acylated anthocyanins tended to display lower increases in λmax than nonacylated counterparts (Table 3). Despite similar λmax values of Al3+ complexed anthocyanins, hue angles differed with changes in pH, as shown in Tab1e 3. Starting at pH 3, no samples tested showed vivid blue hues. Acylated delphinidin (Japanese eggplant) demonstrated most blue-like hues, having a hue angle of 293 (Table 3). Other samples showed more purple-pink hues. Despite delphinidin having the pyrogallol moiety and exhibiting farthest bathochromic shifts upon Al3+ complexation, competition of H+ ions from the acidic pH likely inhibited formation of pure blue colors. With inclusion of acylation on the molecule, the bathochromic shift was found to be no greater; however, there was a higher initial and terminating λmax with metal chelation. Glycosylated cyanidin samples appeared much pinker than acylated counterparts upon complexation with Al3+, which visually resembled delphinidin samples, although hue angles differed by about 10. When increase in the starting pH to 4, anthocyanin−metal chelates began to approach blue hues. Results followed the same general trend for lower pH. Salt-treated glycosylated cyanidin samples were mostly pink, whereas acylated cyanidin from red cabbage exhibited similar hues to those of delphinidin. Both exhibited more blue-like color than at pH 3 with hues around 280−285; acylated cyanidin exhibited higher, more purple hue. Most interestingly, delphinidin acylated with coumaric acid (Japanese eggplant) showed pure blue color upon complexing with Al3+. Hue of this sample treated with AlCl3 100× anthocyanin concentration was 265 and remained constant until reaching AlCl3 1000× and 2000×, becoming 269

equilibrium shifts. Addition of AlCl3 to aqueous solutions can also cause pH decreases due to its ability to hydrolyze in water; however, using high-molarity buffer solutions helped to reduce these effects. In 1 M acetate buffers, pH decreases were minimized until after AlCl3 salt concentrations exceeded 100× anthocyanin content (Figure 6), so color observations were due to complexation rather pH changes. Even at more neutral pH, where anthocyanins tend to exhibit minimal color intensity, all Al3+-treated samples showed bathochromic and hyperchromic shifts in visible spectrophotometry, becoming bluer than respective untreated anthocyanins. In agreement with previous studies, it was concluded that the predominant anthocyanin structure at said pH acts as the foundation against which the color of the anthocyanin complexes competes.3 At low pH, where anthocyanins tend to appear in mostly red flavylium structures, those blue anthocyanin−metal complexes compete, giving the final solution a purple color. Conversely, at more neutral pH, where anthocyanins appear in colorless carbinol or yellow chalcone forms, the ultimate solution color is blue or bluegreen with metal complex competition. All samples, pH 3−6, followed the same general trend for Al3+ chelation depicted (Figure 7) for acylated delphindin (Japanese eggplant samples), showing hyperchromic shifts. In higher pH solutions, the absorbance of complexed solutions was not as intense as their respective counterparts in lower pH. Samples experienced only bathochromic shifts at low pH, until high levels of AlCl3 2000× anthocyanin concentration. The reverse was noted when the starting point was pH 6. With low salt concentrations (equimolar conditions), bathochromic shifts were first observed but became hypsochromic with increasing salt content. Across the tested pH range, the final λmax terminated at similar wavelengths, within 10 nm, for pH 4−6 H


I

3.07 4.01 5.01 6.04

3.07 4.01 5.01 6.04

chokeberry

red cabbage

(0.10) (0.05) (0.03) (0.02)

(0.09) (0.05) (0.02) (0.02)

(0.10) (0.05) (0.03) (0.02)

(0.11) (0.04) (0.01) (0.03)

0×

pH

2.98 3.97 4.97 5.81

2.98 3.97 4.97 5.81

2.98 3.97 4.97 5.80

2.99 3.96 4.97 5.79

(0.10) (0.05) (0.03) (0.02)

(0.10) (0.05) (0.03) (0.02)

(0.10) (0.05) (0.03) (0.01)

(0.10) (0.05) (0.02) (0.02)

100×

532 539 549 555

(1) (1) (2) (2)

513 (1) 511 (2) 508b 528b

527 (0) 529 (1) 534b 540b

523 (0) 523 (4) 509b 509b

551 572 579 589

534 558 564 568

569 578 581 587

568 575 578 579

(0) (0) (1) (2)

(0) (1) (1) (1)

(1) (1) (2) (1)

(1) (1) (2) (3)

100×

λmax (nm) 0×

90.7 91.7 92.5 92.3

92.1 93.8 94.3 93.8

92.7 93.9 94.2 93.2

92.5 94.1 94.5 93.9

(0.4) (0.2) (0.2) (0.5)

(0.6) (0.3) (0.2) (0.2)

(0.1) (0.3) (0.3) (0.6)

(0.1) (0.2) (0.4) (0.6)

0×

L

87.7 87.1 87.5 88.4

89.0 88.6 88.9 89.6

87.6 87.2 87.5 88.3

88.0 87.7 88.2 90.0

(0.3) (0.2) (0.0) (0.2)

(1.0) (0.2) (0.2) (0.3)

(0.4) (0.3) (1.0) (1.2)

(0.3) (0.1) (0.1) (0.8)

100×

11.6 7.0 3.2 0.1

8.3 3.6 1.8 1.0

6.5 2.0 0.5 −0.6

6.5 2.4 0.9 0.4

(0.3) (0.0) (0.1) (0.0)

(1.0) (0.0) (0.0) (0.1)

(0.6) (0.1) (0.1) (0.2)

(0.4) (0.0) (0.1) (0.6)

0×

a*

7.6 2.8 0.5 −1.3

9.3 5.9 3.3 1.4

3.5 −0.7 −2.6 −2.9

4.6 1.7 0.2 −0.2

(0.3) (0.0) (0.0) (0.0)

(1.1) (0.1) (0.0) (0.1)

(0.2) (0.1) (0.3) (0.5)

(0.3) (0.0) (0.1) (0.1)

100×

(0.3) (0.1) (0.2) (0.2) (0.1) (0.1) (0.2) (0.3)

−3.3 −3.3 −3.0 −3.3

(0.4) (0.2) (0.1) (0.3)

−0.7 0.5 1.1 1.4 1.8 1.1 0.8 0.5

(0.1) (0.1) (0.2) (0.2)

0.9 1.2 1.2 1.2

0×

b*

(0.1) (0.2) (0.3) (0.2)

(0.4) (0.2) (0.1) (0.2)

−3.4 −6.0 −4.7 −2.1 −8.8 −10.6 −9.9 −8.3

(1.0) (0.5) (0.9) (1.0)

(0.8) (0.5) (0.2) (0.7)

−8.3 −7.9 −6.3 −2.2

−7.1 −8.2 −6.2 −2.2

100×

In parentheses are the standard deviations, n = 3. bValues had to be estimated from spectrum due to low absorbance of untreated anthocyanins.

3.07 4.01 5.01 6.03

Japanese eggplant

a

3.08 4.00 5.01 6.00

American eggplant

sample

12.1 7.8 4.4 3.3

8.5 3.8 2.0 1.1

6.5 2.1 1.2 1.6

6.6 2.7 1.6 1.3

(0.3) (0.0) (0.1) (0.3)

(1.0) (0.0) (0.1) (0.2)

(0.6) (0.1) (0.1) (0.2)

11.6 10.9 9.9 8.4

9.9 8.4 5.7 2.5

9.0 8.0 6.8 3.6

(0.1) (0.2) (0.3) (0.2)

(1.2) (0.2) (1.0) (0.3)

(1.0) (0.5) (1.0) (0.9)

(0.8) (0.5) (0.2) (0.7)

100× 8.5 8.3 6.2 2.2

chroma (0.4) (0.1) (0.2) (0.4)

0×

344.0 334.5 317.2 271.8

12.6 17.6 25.1 25.9

353.6 8.1 63.5 115.3

7.9 25.7 52.7 73.2

hue

(0.4) (1.0) (2.4) (0.7)

(0.5) (1.3) (4.8) (7.2)

(2.5) (16.4) (2.5) (10.6)

(1.1) (1.9) (1.6) (2.5)

0×

310.9 284.6 272.7 260.9

339.9 314.3 305.6 302.9

292.7 264.8 247.2 216.2

302.8 281.7 272.0 263.9

(1.2) (0.5) (0.1) (0.2)

(1.0) (0.7) (0.5) (1.4)

(1.1) (0.6) (1.1) (9.8)

(1.6) (0.6) (0.6) (6.5)

100×

Table 3. Effects of Acidic pH Change on Color Shifts, with λmax and CIE-Lab Color Characteristics of 50 μM Anthocyanins (0×) and 50 μM Anthocyanins Complexed with AlCl3 100× Anthocyanin Concentrationa

Journal of Agricultural and Food Chemistry Article



Article

hues at pH ≥2.5 when complexation occurred at pH 4. Nonacylated delphinidin−Al chelates did not express blue color until pH 5, and the same was noted for acylated cyanidin, in that red cabbage anthocyanins expressed blue hues at pH 5. Without acylation, cyanidin did not develop blue colors. This observation may be due to an effect of molecular folding occurring with acylated anthocyanins that increases electron density and resonance. With optimal anthocyanin−salt ratios, blue colors developed from metal−anthocyanins could be viable alternatives for synthetic blue colorants in food systems.

and 274, respectively. As noted before, these high salt concentrations were also correlated with decreases in pH and increased H+ competition against metal ions for the hydroxyl groups of the B ring. Beginning at pH 5, more samples began to exhibit visibly blue colors. Delphinidin anthocyanins from American eggplant and acylated cyanidin from red cabbage showed hues of 272 with AlCl3 concentrations 100× anthocyanin concentration. This hue was maintained until reaching AlCl3 2000×, where color reverted to higher hues. Salt-treated cyanidin samples formed unique purple-pink hues of 305−308, with hue increasing as salt concentration increased. Acylated delphinidin samples exhibited a wider range of hues than other samples from 230 to 263. The optimal pH for delphinidin-3-glucoside− ferric chelates was found to be 4.5 on the basis of formation and stabilization of the chelates.3 Generally, absorbance of complexed solutions decreased as pH increased, except by comparing pH 3 and 4, where absorbance increased by about 0.1 except chokeberry anthocyanins, which were lower. Decreases in absorbance were highest by comparing pH 5 and 6, ranging from 0.9 to 0.25. From color and intensity evaluation, the ideal pH for blue color formation of anthocyanin−Al chelates likely lies in pH 4−5 range. Similar trends were noted with starting pH 6; the highest amount of blue hues was exhibited from the most number of samples tested. Delphinidin and acylated cyanidin showed hues around 260, becoming more blue with the increasing pH. Cyanidin from chokeberry still did not exhibit blue colors at this pH, having hues around 300. It seems cyanidin, without acylation, lacks the necessary electron density and resonance to form blue colors in acidic pH with Al3+. Acylated delphinidin exhibited a range of hues upon chelation with Al3+, 216−260, depending on salt concentration and pH. These hues were found to be the most similar to the synthetic dyes FD&C Blue No. 1 and FD&C Blue No. 2, which were found to exhibit hues of 224.3 and 241.5, respectively. The colorant FD&C Blue No. 1 expressed a λmax of 630, whereas Blue No. 2 showed λmax of 610 nm, each of which easily correlated to their observed blue colors, in agreement with previous findings.26 Despite the fact that these metalloanthocyanin complexes had much lower λmax values, some were able to produce similar blue hues. Unlike the findings from previous studies of anthocyanin− ferric chelates, precipitation of metalloanthocyanins was not observed until neutral or basic pH had been reached.3 At pH 5, ferric anthocyanins were found to precipitate within 2 h of ion introduction, leaving the supernatant colorless after 24 h.3 Data were not included, but anthocyanins were found to chelate to Al3+ in basic pH testing. However, when exposed to pH ≥7, the complexes almost immediately precipitated, leaving the supernatant mildly colored or colorless depending on salt concentration. Precipitates showed colors similar to their respective solution colors at pH 6. Experiments revealed anthocyanins were able to interact with metals in wide concentration ranges, but salt ratio and pH determined final color and strength. Highest stability and color expression were found when catechol- or pyrogallol-bearing anthocyanins were mixed with AlCl3 at ratios of 100−500× anthocyanin concentration. Anthocyanin color stability was also found to improve with metal chelation, exhibiting intense color for increased time. Three free hydroxyl groups on B rings of anthocyanins and acylation were found to further λmax and blue color formation. Acylation proved critical to blue color development at low pH; acylated delphinidin exhibited blue

■

AUTHOR INFORMATION

Corresponding Author

*(M.M.G.) Phone: (614) 247-8016. Fax: (614) 292-0218. Email: [email protected]. Notes

The authors declare no competing financial interest.

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ABBREVIATIONS USED

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REFERENCES

EU, European Union; FDA, U.S. Food and Drug Administration; GRAS, generally recognized as safe; HPLC-MS, highpressure liquid chromatography−mass spectrometry; ESI, electrospray ionization; ANOVA, analysis of variance; ACN, anthocyanin; A, absorbance; MW, molecular weight; DF, dilution factor; Dpn, delphinidin; Cy, cyanidin; rut, rutinoside; p-cou, para-coumaric acid; gal, galactoside; arab, arabinoside; sop, sophoroside; acyl, acylated (could include sinapic, ferulic, coumaric, or caffeic acids); glurut, glucosyl-rutinoside; xyl, xyloside; fer, ferulic acid

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Bathochromic and Hyperchromic Effects of Aluminum Salt

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