Evaluating the Effects of Anthocyanin Structure and the Role of Metal Ions on the Blue Color Evolution of Anthocyanins in Varied Ph Environments

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Evaluating the Effects of Anthocyanin Structure and the Role of Metal Ions on the Blue Color Evolution of Anthocyanins in Varied pH Environments

DISSERTATION

Presented in Partial Fulfillment of the Requirements for the Degree Doctor of Philosophy in the Graduate School of The Ohio State University

Gregory T Sigurdson, B.S.

Graduate Program in Food Science and Technology

The Ohio State University

2016

Dissertation Committee:

M. Mónica Giusti, Ph.D., Advisor

Lynn Knipe, Ph.D.

Luis Rodriguez-Saona, Ph.D.

Christopher Simons, Ph.D.

Copyrighted by

Gregory Thomas Sigurdson

2016

Abstract

Due to consumer demand and possible health concerns, the use of synthetic food colorants has been decreasing. Synthetic red and yellow colorants are most commonly used in the food industry; however they are considered easier to substitute with alternatives from nature due to their prevalence and various chemical natures.

Alternatives for synthetic blue colorants are more limited due to low prevalence in nature, poor stability, or inability to match the color characteristics of the synthetic dyes.

Anthocyanins are a class of natural pigments responsible for red, purple, and blue colors of edible produce with potential health benefits. In acidic conditions common to many food products, anthocyanins appear in red-purple structural forms limiting their use as blue colorants. However, self-association, co-pigmentation, and metal chelation have been suggested to expand their color expression to purple and blue in acidic pH. Metal ions displace hydrogen ions from B-ring hydroxyl groups, transforming red flavyliums to blue quinonoidal bases. The effects of the anthocyanin substitution pattern and role of various metal ions on their color evolution have not been fully elucidated in the wide pH range common to many foods. The objective of these studies was to replicate these naturally occurring reactions and evaluate the anthocyanin response to metal ions to expand the useful color and pH range of anthocyanins and to better understand the mechanisms of blue color expression.

Acylated and nonacylated derivatives of cyanidin and delphinidin were combined with various metal ions (Mg2+, Al3+, Cr3+, Fe3+, and Ga3+) in factorial excesses to anthocyanin concentration (as large as 2000× [anthocyanin]) in pH 3-8. Phenolic acids were also evaluated as copigments on anthocyanin-metal chelates. The effects on anthocyanin color expression were evaluated by UV-Visible spectrophotometry (380-700 nm) and CIE-L*a*b* colorimetry (transmission, 10º observer angle, D65 illuminant).

Anthocyanin responses to metal ion chelation were dependent on all factors tested. In all pH, anthocyanins exhibited bathochromic and hyperchromic shifts becoming more purple and blue. Largest bathochromic shifts occurred in pH 6, and largest hyperchromic shifts occurred in pH 5. The ratio of metal ion to anthocyanin necessary to induce the largest bathochromic shifts decreased as pH was increased indicating competition for the binding sites by hydrogen ions.

Metal chelation resulted in larger bathochromic shifts on anthocyanins with more

B-ring hydroxyl groups (delphinidin > cyanidin). Anthocyanins lacking acylation typically underwent much larger bathochromic shifts; however, the λmax of acylated anthocyanins with metals was larger in all cases, resulting in more blue color expression.

Anthocyanins bearing diacylation expressed the most blue hues in the widest pH range followed by aromatically monoacylated anthocyanin derivatives. Hydroxycinnamic acid derivatives were shown to act as poor intermolecular copigments of anthocyanin-metal chelates when compared to their effects when covalently attached to the chromophore; they resulted in almost negligible changes to the color of the anthocyanin-chelates.

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Bivalent Mg2+ showed no obvious effect on anthocyanin color. Generally, bathochromic shifts on anthocyanin were greatest with more electron rich metal ions:

Fe3+ ≈ Ga3+ > Al3+ > Cr3+.

Metal chelation by anthocyanins resulted in blue and purple hues similar to currently used synthetic colorants in pH environments where these colors are not typically expressed, further indicating potential to act as alternatives.

Dedication

I dedicate the work of this dissertation to Grandpa, Geoffrey M. Sigurdson, who entered

the realm of the bravest warriors before completion of it.

Acknowledgments

I must first express my most sincere gratitude to my graduate advisor Dr. M.

Mónica Giusti. I began my graduate program under slightly unusual circumstances, and she presented almost no reservation in allowing me to study under guidance. Through her hard work, research funding was secured that not only allowed me to investigate a topic that become very passionate to me but also provided me with ways to grow that many other graduate students do not have the opportunities to experience. Undoubtedly,

Mónica is an expert in her field; and I was gifted the opportunity to absorb as much knowledge as I possibly could from her. The education I received from Mónica was not limited to anthocyanins or color chemistry; she was a great mentor in professional and personal growth, professional relationship development, and understanding the nuances of an academic career. Her passion for our field of study and encouragement throughout my graduate career was evident and often inspired me to strive to learn and undertake more. The gifts Mónica imparted to me will not be forgotten.

I must also thank the members of my advisory committee Dr. Luis Rodriguez-

Saona, Dr. Lynn Knipe, and Dr. Christopher Simons. I am grateful for the time they each have given me providing great advice and also conducting the various necessary components of a graduate level degree. The guidance they provided me is applicable in a variety of settings, and I will carry it with me. The challenges they gave me allowed me to progress in manners I otherwise would not have. Dr. Rodriguez-Saona was especially vi important in my academic growth, having been a professor for what I consider as some of the most formative and foundational courses in food science. He is not only a great advisor but also a great professor and instructor.

I am also indebted to all my peers from the Giusti lab, some who left before me and some who are to follow. Each has played a unique in my education at The Ohio State

University. From them I have learned a wide range of scientific knowledge; experienced unique cultures, foods, and languages; and gained several friends. I am especially grateful to Dr. Neda Ahmadiani, who not only took the time to train me to work in the lab but also to become an efficient lab manager. She was a great source of knowledge and is a great friend and mother. I must thank Alex Westfall who helped me care for the lab during her time here; she is an excellent friend and researcher and can always be counted on to keep things “Moo-ving” along. I am grateful for all my lab mates Jacob Farr, Allison Atnip, Fei

Lao, Peipei Tang, Andrew Barry, Yucheng Zhou, Yingfang Li, Kai Zhang, Xiaoyi Zhu and

Gonzalo Miyagusuku-Cruzado and hope the best for them all.

I would like to thank Dr. Rebecca Robbins and Dr. Thomas Collins from Mars

Chocolate. They have provided continuous support of my work and valuable advice and suggestions that not only improved the quality of my work but also extended my understanding and education. Through their efforts and Mars Chocolate NA, I was provided funding to earn my graduate degree. I am exceedingly grateful for the opportunities they provided me.

I must also acknowledge my family whose pride and encouragement has helped me to continue the course of this program. It has been a long and challenging journey during which my family has been very understanding of my absences and neglect. I am eternally grateful for all the efforts and sacrifices my mother LaDonna Roys has done for me and for the achievement of this

vii degree. I recall times in elementary school when my father Gregory A. Sigurdson would make the extra effort to help me with my homework with my advanced classes while suffering from lack of sleep from working two jobs. These memories have encouraged me to continue my studies even under the most stressful and tiresome of conditions. I can never thank my parents LaDonna Roys,

Gregory Sigurdson and Lynn Sigurdson for their contributions and encouragement during this long journey. I have striven to bring pride to my family and will continue to do so.

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Vita

January 1987 ...... Houston, TX

May 2005 ...... Spencerville High School

December 2011 ...... B.S. Food Science and Technology, The

Ohio State University

2012 to present ...... Graduate Research Associate, Department

of Food Science and Technology, The Ohio

State University

Publications

Sigurdson, G. T.; Robbins, R. J.; Collins, T. M.; Giusti, M. M. (2016). Evaluating the

Role of Metal Ions in the Bathochromic and Hyperchromic Responses of Cyanidin

Derivatives in Acidic and Alkaline pH. Food Chemistry, 208, 1, 26-34.

Sigurdson, G. T. & Giusti, M. M. (2014). Bathochromic and Hyperchromic Effects of

Aluminum Salt Complexation by Anthocyanins from Edible Sources for Blue Color

Development. Journal of Agricultural and Food Chemistry, 62, 6955–6965.

Collins, T.; Robbins, R.; Giusti, M. M.; & Sigurdson, G. T. Colorant Compositions and

Methods of Use Therein. Provisional patent application. Filed: June 30, 2015.

Giusti, M. M. & Sigurdson, G. T. Anthocyanin-Metallo Complexation for Blue and

Purple Colorants for Food Application. Provisional patent application. Filed: September

13, 2014.

Fields of Study

Major Field: Food Science and Technology

Table of Contents

Abstract ...... ii

Dedication ...... v

Acknowledgments...... vi

Vita ...... ix

Table of Contents…………………………………………………………………………xi

List of Tables ...... xvixviii

List of Figures ...... xx

List of Abbreviations…………………………………………………………………..xxiv

Chapter 1: Introduction ...... 1

Chapter 2: Review of Literature ...... 5

2.1: Importance of Color of Foods………………………………………………...5

2.2: Definition of Color……………………………………………………………9

2.3: Measurement of Color………………………………………………………10

2.4: Color Communication Systems……………………………………………..12

2.4.1: CIE-XYZ Tristimulus Systems……………………………………12

2.4.2: CIE-L*a*b* Systems……………………………………………...13

2.5: Food Colorants………………………………………………………………15

2.5.1: Certified Colors……………………………………………………16

2.5.2: Colorants Exempt from Certification……………………………..18

2.5.3: Trends in Coloring of Foods………………………………………21

2.6: Blue Colorants………………………………………………………………23

2.6.1: Synthetic Blue Colorants………………………………………….25

2.6.2: Blue Colorants Derived from Natural Sources……………………26

2.7: Anthocyanins………………………………………………………………..30

2.7.1: Structural Components……………………………………………30

2.7.2: Potential Health Benefits………………………………………….33

2.7.3: Anthocyanins as Food Colorants………………………………….35

2.7.4: Mechanisms to Produce Blue Colors……………………………...38

2.7.4.1: Derivatization of Anthocyanins…………………………38

2.7.4.2: Anthocyanin Self-Association…………………………..39

2.7.4.3: Copigmentation of Anthocyanins……………………….40

2.7.4.4: Metal Chelation and Complexation……………………..43

Chapter 3: Bathochromic and Hyperchromic Effects of Aluminum Chelation by

Anthocyanins from Edible Sources for Blue Color Development……………………….51

3.1: Abstract……………………………………………………………………...51

3.2: Keywords……………………………………………………………………52

3.3: Introduction………………………………………………………………….52

3.4: Materials & Methods………………………………………………………..55

3.4.1: Materials…………………………………………………………..55

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3.4.2: Methods…………………………………………………………...56

3.4.2.1: Extraction and Purification of Anthocyanins…………....56

3.4.2.2: Anthocyanin Purification - Solid Phase Extraction……..57

3.4.2.3: Monomeric Anthocyanin Quantitation………………….57

3.4.2.4: High Pressure Liquid Chromatography (HPLC) Evaluation

of Anthocyanins………………………………………………….58

3.4.2.5: Evaluating Anthocyanin Structures, Concentrations, and

Salt Ratios in Al3+ Complex Formation………………………….60

3.4.2.6: Evaluating Color Stability over time of Al3+ Complexed

Anthocyanins…………………………………………………….60

3.4.2.7: Evaluating the Effect of pH on Al3+ Complex

Formation...... 61

3.4.2.8: Spectrophotometry of Solutions with UV-Visible

Transmission……………………………………………………..61

3.4.2.9: CIE-Lab Color of Solutions by Transmission…………..61

3.4.2.10: Statistical Evaluation of Data………………………….61

3.5: Results and Discussion……………………………………………………...62

3.5.1: Evaluating Results from Reverse Phase HPLC-MS………………62

3.5.2: Evaluating Anthocyanin Structure, Concentrations, and Salt Ratios

in Al3+ Complex Formation……………………………………………...64

3.5.3: Evaluating Color Stability over time (28 days) of Al3+ Complexed

Anthocyanins…………………………………………………………….70

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3.5.4 Evaluating Effects of pH in Al3+ Complex Formation…………….74

3.6: Conclusion…………………………………………………………………..81

Chapter 4: Spectral and Colorimetric Characteristics of Metal Chelates of Acylated

Cyanidin Derivatives…………………………………………………………………….83

4.1: Abstract……………………………………………………………………...83

4.2: Keywords……………………………………………………………………83

4.3: Introduction………………………………………………………………….84

4.4: Materials & Methods………………………………………………………..87

4.4.1: Materials…………………………………………………………..87

4.4.2: Methods…………………………………………………………...88

4.4.2.1: Anthocyanin Extraction…………………………………88

4.4.2.2: Anthocyanin Purification - Solid Phase Extraction……..88

4.4.2.3: Anthocyanin Isolation - High Pressure Liquid

Chromatography (HPLC)………………………………………..88

4.4.2.4: Isolated Anthocyanin Purity…………………………….89

4.4.2.5: Monomeric Anthocyanin Quantitation………………….90

4.4.2.6: Sample Preparation……………………………………...90

4.4.2.7: Spectrophotometry of Solutions by UV-Visible

Transmission……………………………………………………..91

4.4.2.8: Colorimetry of Solutions………………………………..91

4.4.2.9: Statistical Evaluation of Data…………………………...92

4.5: Results and Discussion……………………………………………………...92

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4.5.1: Spectral & Colorimetric Response………………………………..92

4.5.2: Stability……………………………………………………………99

4.6: Conclusion…………………………………………………………………104

Chapter 5: Evaluation of Hydroxycinnamic Acid Derivatives as Inter- and Intra- molecular Copigments of Cyanidin Derivatives and their Metal Chelates……………..106

5.1: Abstract………………………………………………………………….....106

5.2: Keywords….…………………………………………………………….....106

5.3: Introduction………………………………………………………………...107

5.4: Materials & Methods………………………………………………………110

5.4.1: Materials…………………………………………………………110

5.4.2: Methods………………………………………………………….111

5.4.2.1: Anthocyanin Extraction and Purification……………...111

5.4.2.2: Anthocyanin Isolation …………………………………111

5.4.2.3: Isolated Anthocyanin Purity…………………………...112

5.4.2.4: Monomeric Anthocyanin Quantitation………………...113

5.4.2.5: Sample Preparation…………………………………….114

5.4.2.6: Visible Spectrophotometry of Solutions ………………114

5.4.2.7: Colorimetry of Solutions………………………………115

5.4.2.8: Statistical Evaluation of Data …………………………115

5.5: Results and Discussion…………………………………………………….115

5.5.1 Hydroxycinnamic Acid Derivatives as ACN Copigments……….115

5.5.2 Hydroxycinnamic Acid Derivatives as ACN Copigments with the

Presence of Metal Ions…………………………………………………121

5.6 Conclusions…………………………………………………………………126

Chapter 6: Evaluating the Role of Metal Ions in the Bathochromic and Hyperchromic

Responses of Cyanidin Derivatives in Acidic and Alkaline pH………………………..128

6.1: Abstract…………………………………………………………………….128

6.2: Keywords………………………………………………………………….129

6.3: Introduction………………………………………………………………..129

6.4: Materials & Methods………………………………………………………132

6.4.1: Materials……………….………………………………………...132

6.4.2: Methods………………………………………………………….133

6.4.2.1: Anthocyanin Extraction………………………………..133

6.4.2.2: Anthocyanin Purification – Solid Phase Extraction…...133

6.4.2.3: Monomeric Anthocyanin Quantitation ………………..134

6.4.2.4: High Pressure Liquid Chromatography (HPLC) -

Anthocyanin Purity Evaluation…………………………………135

6.4.2.5: Sample Preparation…………………………………….135

6.4.2.6: Spectrophotometry of Solutions by UV-Visible

Transmission……………………………………………………136

6.4.2.7: Statistical Evaluation of Data………………………….136

6.5: Results and Discussion…………………………………………….136

6.5.1: Role of pH on Anthocyanin-Metal Chelation……………137

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6.5.1.1: Role of pH on Bathochromic Responses………137

6.5.1.2: Role of pH on Hyperchromic Responses………138

6.5.1.3: Role of pH on ACN-Mn+ ratios………………...139

6.5.2 Role of Specific Metal Ions in Anthocyanin-Metal

Chelation………………………………………………………..141

6.5.2.1: Magnesium……………………………………..143

6.5.2.2: Aluminum……………………………………...147

6.5.2.3: Chromium……………………………………...149

6.5.2.4: Iron……………………………………………..150

6.5.2.5: Gallium………………………………………...153

6.6: Conclusions………………………………………………………...154

Chapter 7: Concluding Remarks………………………………………………………..157

Bibliography……………………………………………………………………………159

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List of Tables

Table 2.1: List of certified FD&C dyes, their color characterization, and use (Frick,

2003; Sharma, McKone, & Markow, 2011) ...... 17

Table 2.2: List and classification of major colorants exempt from certification, their color characterization, and use (21 CFR 70.3)…………………………………………………19

Table 3.1: Display of anthocyanin, associated molecular weight, and molar absorptivity used for monomeric quantitation of anthocyanins in food samples evaluated…………..58

Table 3.2: Effects of anthocyanin concentration at pH 3 on color shifts, with λmax and

CIE-Lab color characteristics of anthocyanins complexed with AlCl3 100× anthocyanin concentration. In parenthesis () are the standard deviations, n=3………………………..65

Table 3.3: Effects of acidic pH change on color shifts, λmax and CIE-Lab color characteristics of 50 µM anthocyanins (0×) and complexed with AlCl3 100× anthocyanin concentration. (standard deviations), n=3. a values estimated due to low absorbance…..78

Table 4.1: Average and (standard deviation) bathochromic shifts (nm) of Cy derivatives

(50 µM) treated with factorial excess of Al3+ or Fe3+ (1× [ACN] in pH 6 and 0.5× [ACN] in pH 7) to [ACN], n = 3…………………………………………………………………97

Table 4.2: CIE-L*a*b* values average and (standard deviation) of Cy derivatives (50

µM) treated with factorial excess of Al3+ or Fe3+ (1× [ACN] in pH 6 and 0.5× [ACN] in pH 7) to [ACN], n = 3……………………………………………………………………98

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Table 4.3: Average and (standard deviation) of half-life (t1/2 expressed as hr) and rate constants (k) of Cy derivatives (50 µM) in pH 7 treated with factorial excesses of Al3+ or

Fe3+ (0×, 0.5×, 5× [ACN]) during dark storage at 21-25 ºC, calculated by 1st order kinetics by decrease in absorbance of λmax of to, n = 3…………………………………101

3+ Table 5.1: λmax (nm) of samples without Al treatment and bathochromic shifts (nm, compared to CtG) of Cy-3-diglucoside-5-glucoside (CtG) treated with different levels of hydroxycinnamic acid derivatives (0-50× [ACN]), monoacylated (CtgMa) derivatives, and diacylated (CtgDa) derivatives and with specified factorial excess of Al3+ (0-100×

[ACN]) in pH 5-8……………………………………………………………………….116

Table 5.2: CIE-L*C*hº average values (standard deviation) of Cy-3-diglucoside-5- glucoside: CtG derivatives (50 µM) treated with factorial excess of hydroxycinammic acid derivatives (0-50× [ACN]), monoacylated CtgMa derivatives, and diacylated CtgDa derivatives in pH 5-8……………………………………………………………………119

Table 5.3: CIE-L*C*hº average values (standard deviation) of Cy-3-diglucoside-5- glucoside: CtG derivatives (50 µM) treated with factorial excess of hydroxycinammic acid derivatives (0-50× [ACN]), monoacylated CtgMa derivatives, and diacylated CtgDa derivatives with specified factorial excess of Al3+ (0-100× [ACN]) in pH 5-8………...125

Table 6.1: Largest bathochromic shifta expressed in nm (with Mn+ factorial value to

[ACN] inducing shift) of red cabbage and chokeberry ACN induced by different Mn+ in pH 3-8…………………………………………………………………………………..145

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List of Figures

Figure 2.1: Standard observer curves (10 °) demonstrating red (x), blue (z), and green (y) cone sensitivity and the visible spectrum and spectral power distribution curves of three standard CIE illuminants (1964) (CIE 2016)...………………………………………..…10

Figure 2.2: Equations to convert spectral data into CIE-XYZ and CIE-L*a*b* color spaces (Wrolstad & Smith, 2010)………………………………………………………..13

Figure 2.3: CIE-L*a*b* color space (Artec Testnology, 2016)……………………...…15

Figure 2.4: Visible absorbance (380-700 nm) and solution of FD&C Blue No. 1 in

H2O………………………………………………………………………………………24

Figure 2.6: Basic structure of a flavonoid and the most common anthocyanins found in nature………………………………………………………..……………………………31

Figure 2.7: Representation of different color expression of major anthocyanin aglycones adapted from Ananga, Georgiev, Ochieng, Phills, & Tsolova, 2013..…………………..32

Figure 2.8: Major anthocyanin structural forms and color expression due to pH dependence adapted from Houbiers, Lima, Maçanita, & Santos, 1998….………………36

Figure 2.9: Proposed stacking conformations for stabilization mechanisms of acylated anthocyanins adapted from Jackman & Smith, 1996………..…………………………..42

Figure 2.10: Color expression of butterfly pea flower (Clitoria ternatea) anthocyanins in solutions pH 3-8………………………………………………………………………….43

Figure 2.11: Proposed mechanism of metal chelation and structural conversion of anthocyanins adapted from Schreiber, Swink, & Godsey, 2010.………………………..44

Figure 2.12: Cyanidin derivatives (chokeberry anthocyanins) treated with factorial excesses of Al3+ (0-100× [ACN]) in pH 4……………………………………………….47

Figure 2.13: Schematic of the organization of the metalloanthocyanins, adapted from

Yoshida, Mori, & Kondo, 2009………………………………………………………….50

Figure 3.1: Al3+ chelation by cyanidin (B ring exhibiting catechol moiety) or delphinidin-3-p-coumaroyl-rutinoside-5-glucoside (B ring exhibiting pyrogallol moiety), where R1: H or R1: OH, respectively…………………………………………………….53

Figure 3.2: 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, and (G): black carrot with detection at 520 nm showings main pigments identified………………………………………………………………....63

Figure 3.3: Visible absorbance (400-700 nm) of 25, 50, & 100 μM concentrations of

American eggplant anthocyanins treated with factorial increases of AlCl3 (0-1000×) over anthocyanin concentration at pH 3………………………………………………………67

Figure 3.3: Visible absorbance (400-700 nm) of 25, 50, & 100 μM concentrations of

American eggplant anthocyanins treated with factorial increases of AlCl3 (0-1000×) over anthocyanin concentration at pH 3………………………………………………………68

Figure 3.5: 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-

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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)………………………………………………………………………………..72

Figure 3.6: pH change of 50 μM anthocyanins sample solutions initiated by treatment with AlCl3 (0-2000×) in 1 M sodium acetate buffers pH 3, 4, 5, & 6…………………...74

Figure 3.7: 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……………………………………………………..76

Figure 4.1: Identification (and abbreviations) of structure and substitution patterns of isolated ACN……………………………………………………………………………..93

Figure 4.2: λmax (nm) of Cy derivatives in pH 6 (left) or pH 7 (right), treated with factorial excess of Al3+ (0-5×) to [ACN] (50 µM), n = 3………………………………..94

Figure 4.3: Plot of ln(Absorbance at λmax of to) vs. time (hr) of CtGC and CtGSC in pH 7 treated with factorial excesses of Al3+ (0-5× [ACN]), n = 3……………………………..98

Figure 5.1: HPLC chromatogram (detection at 520 nm) of red cabbage anthocyanins illustrating major isolated cyanidin derivatives………………………………………...113

Figure 5.2: λmax (left) and absorbance (right) of Cy-3-diglucoside-5-glucoside (CtG) treated with factorial excesses (0-50× [ACN]) of hydroxycinnamic acid derivatives in pH

5-8, n = 3………………………………………………………………………………..117

Figure 5.3: λmax (left) and absorbance (right) of Cy-3-diglucoside-5-glucoside (CtG) treated with factorial excesses of hydroxycinnamic acid derivatives (0-50× [ACN]),

xxii monoacylated CtgMa derivatives, and diacylated CtgDa derivatives with factorial excess of Al3+ (0-100× [ACN]) in pH 5, n = 3…………………………………………………122

Figure 6.1: HPLC chromatograms and identification of chokeberry and red cabbage

ACN after purification, detection at 520 nm and at 260-700 nm. Cy: cyanidin; soph: sophoroside; glu: glucoside; gal: galactoside; ara: arabinoside………………………...134

Figure 6.2: Bathochromic shift (nm) and hyperchromic shift with addition of equimolar proportions of Fe3+ to red cabbage ACN solutions (pH 3-8) and visible absorbance spectra of red cabbage ACN treated with equimolar proportions of Mn+ (Mg2+, Al3+, Cr3+,

Fe3+, or Ga3+), pH 6……………………………………………………………………..138

Figure 6.3: λmax (nm) and absorbance of red cabbage ACN treated with factorial excess of Al3+ to [ACN] (0-100×), pH 3-8……………………………………………………..140

Figure 6.4: λmax (nm) of chokeberry and red cabbage ACN treated with factorial excess of Mn+ (Mg2+, Al3+, Cr3+, Fe3+, or Ga3+) to [ACN] (0-500×), pH 5…………………….144

Figure 6.5: Absorbance of chokeberry and red cabbage ACN treated with factorial excess of Mn+ (Mg2+, Al3+, Cr3+, Fe3+, or Ga3+) to [ACN] (0-500×), pH 5……………..146

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List of Abbreviations  A: absorbance

 ACN: anthocyanin(s)

 acyl: acylated. Could include: sinapic, ferulic, coumaric, or caffeic acids, etc

 ANOVA: analysis of variance

 arab: arabinoside

 CIE: Commission Internationale de l’Eclairage or International Commission on

Illumination

 CFR: Code of Federal Regulations

 CGM: cyanidin-3-malonyl-glucoside

 CtG: cyanidin-3-diglucoside-5-glucoside (cyanidin-trigylcosylated)

 CtGC: Cy-3-(p-coumaroyl)-diglucoside-5-glucoside

 CtGDa: diacylated Cy derivatives (diacylated fraction)

 CtGF: Cy-3-(feruloyl)-diglucoside-5-glucoside

 CtGMa: monoacylated Cy derivatives (monoacylated fraction)

 CtGS: Cy-3-(sinapoyl)-diglucoside-5-glucoside

 CtGSC: Cy-3-(p-coumaroyl)- (sinapoyl)-diglucoside-5-glucoside

 CtGSF: Cy-3-(feruloyl)-(sinapoyl)-diglucoside-5-glucoside

 CtGSS: Cy-3-(sinapoyl)-(sinapoyl)-diglucoside-5-glucoside

 Cy: cyanidin

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 DF: dilution factor

 Dp: delphinidin

 ESI: electrospray ionization

 EU: European Union

 FDA: U.S. Food and Drug Administration

 fer: ferulic acid

 gal: galactoside

 glurut: glucosyl-rutinoside

 GRAS: Generally Recognized as Safe

 HPLC-MS: high pressure liquid chromatography- mass spectrometry

 hr: hour(s)

 min: minute(s)

 Mn+: metal ion(s)

 MW: molecular weight

 nm: nanometer

 p-cou: para-coumaric acid

 rut: rutinoside

 sop: sophoroside

 TRIS: Tris(hydroxymethyl)aminomethane

 xyl: xyloside

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Chapter 1: Introduction

Color is an integral component of a food product relating to its quality and overall market success. The color of a food product can provide key information regarding its flavor, safety, nutritional value, and several other factors. The food industry uses coloring additives for several purposes in foods including standardization of color of raw ingredients, creation of a color identity for otherwise colorless foods, and account for color loss during processing or storage.

Synthetic colorants were primarily used to color food in the 20th century due to their low cost, inherent chemical stability, and high tinctorial strength. However, consumer demand has been driving the replacement of artificial colorants with those derived from natural sources. The push for use of naturally derived colorants can be related to consumer demand for more “natural” products, possible links of synthetic colorants to hyperactivity in children, and allergenicity to sensitive populations. Although currently used synthetic colorants have long records of safety evaluations and strict regulations, the food industry is seeking alternatives to these colorants to meet the changing market demands.

Warm hued colorants including synthetic reds and yellows have traditionally accounted for the highest amount of food coloring additives but are also considered easier to replicate with pigments derived from natural sources. They can be derived from many edible materials, are prevalent in nature, and have diverse chemical compositions that 1 lend to use in many environments and processing conditions. Although blue colors are not necessarily uncommon in nature, blue pigments are more limited. The blue colors encountered in most animals are actually result of reflection of blue light due to the physical nanoscale characteristics rather than presence of blue pigments. Most naturally occurring blue pigments are encountered in plant systems and some microbiological species; however, their use in foods is typically limited. Typical limitations of naturally derived blue pigments as food colorants include poor heat and/or light stability, low water solubility, sensitivity to pH, or inability to match the color characteristics of synthetic blue colorants.

Anthocyanins are a class of naturally occurring pigments known for imparting a variety of hues in nature ranging from orange to reds, purples, blues, and greens, in some cases. They are commonly found a variety of edible produce and are permitted for use to color food products as fruit or vegetable juice or as grape skin extract. Anthocyanins have shown promising potential to act as alternatives for a variety of synthetic food colorants; however, in the pH environments common to many food products, anthocyanins appear in red-purple structural forms thus limiting their use a blue food colorants. Certain chemical mechanisms found to occur in natural systems may provide methods to increase the blue color expression range of anthocyanins including self-association, molecular copigmentation, and metal chelation. Of these, metal chelation has been considered the most important factor in blue hue expression in acidic pH, which results in conversion of red flavylium anthocyanins to blue quinonoidal structural forms. Although anthocyanins bearing a minimum of an o-dihydroxy system on the B ring have known for several years

2 to chelate metal ions, the exploration of these types of pigments as potential blue food coloring additives is very recent and several factors still remain to be explored.

The work of this dissertation first addressed the role of the substitution pattern of the B ring of the anthocyanin with comparison to acylated counterparts. It is known that the general “blue-ness” of anthocyanins is comparatively increased as the aglycone is increasingly substituted on the B ring. For example the anthocyanin delphinidin, bearing

3 hydroxyl groups on the B ring is comparatively bluer than cyanidin, which only exhibits 2 B ring hydroxyl groups. It should be noted that in acidic pH, however, neither of these anthocyanins appear blue. Chapter 3 compares metal chelation by acylated and nonacylated counterparts of delphinidin and cyanidin for blue color expression and also begins investigation of the role of metal chelation on the stability of anthocyanin pigments.

The findings of chapter 3 seemed to reveal not only B ring structure but also acylation as an important factor in blue color expression of anthocyanin-metal chelates.

Previous works have shown the covalent attachment of different acids to anthocyanins to impact the color expression of the pigments in solution through a mechanism known as copigmentation. This phenomenon occurs when an essentially colorless molecule interacts with an anthocyanin resulting in changes in the visible light absorbance spectrum of the pigment. In acidic pH, it has generally been observed with acylation that the λmax is shifted bathochromically meaning the pigments absorbs wavelengths of less energy (more red) while reflecting or transmitting higher energy photons (more blue).

Interestingly, this effect is enhanced with increasing number of acyl attachments to the

3 pigment. Chapter 4 investigates the role of the amount (monoacylation vs diacylation) and type of acylation (aliphatic vs aromatic) on not only the spectral and colorimetric responses of metal chelation of identical chromophores for blue colorations but also the stability of these pigments.

Copigmentation has also been shown to occur between anthocyanins and free molecules in solution. The molecules are thought to interact through hydrophobic interactions or Π-Π stacking. Unlike intramolecular copigmentation, as occurs for acylated anthocyanins, intermolecular copigmentation (between free molecules) is dependent on many factors such as the molecular structures, their concentrations, and their ratios. The findings of chapter 4 suggested an importance of the acylating moieties on the color expression of the anthocyanin-metal chelates; therefore, chapter 5 investigates the potential role of these acyl moieties as free intermolecular copigments for blue anthocyanin-metal chelates.

The work of previous chapters focused primarily on the effects of the anthocyanin structures on the blue color expression of associated metal chelates. Chapter 6 systemically investigated the role of the environment (pH) and the electronic organization of the metallic cation on the blue color expression of anthocyanin chelates. Food systems can vary dramatically in pH, although most often in the acidic range. It is also well known that anthocyanins adopt various structural resonant forms as response to protonation and deprotonation as a function of pH and express different colors.

Therefore, it should be evaluated as a critical aspect of the blue color appearance of anthocyanin chelates. Different metal ions have also been shown to induce different

4 bathochromic shifts; however conditions and environments varied between the studies making comparison difficult, demonstrating need to further elucidate.

Chapter 2: Review of Literature

2.1 Importance of Color of Foods

Human reaction to foods is considered instinctual, having been ingrained through millennia of learned habits. Historically when searching for foods, humans had to recognize safe and quality foods which could often be related to color (Burrows, 2009).

In modern times, color continues to be part of the first sensory experience related to many food products making it a critical component to a product’s overall success.

The sensory and quality expectations consumers associate with food products are likely to be based partially on the history humans have experienced with different foods.

Certain colors can be associated with certain maturity and ripeness of some fruits and vegetables (Zampini, Sanabria, Phillips, & Spence, 2007). Much of the currently commonly consumed fresh produce changes colors from green to varying shades of yellow-orange and to deep red-purple as it matures. In these green phases, much of this produce contains minimal sugar and relatively higher acidity levels correlating with sourness. However, as red and purple colors develop, fruits and vegetables tend to gain more sugar and volatile compounds which many people then associate with specific fruit/vegetable notes and increased sweetness. Blue colors in nature tend to be relatively limited compared to the more common greens, yellow, and reds. Historically, this atypical color may have been associated with fruits and vegetables that may have some toxicity, such as the blue pigments of some mushrooms (Newsome, Culver, & Breemen, 6

2014). It is also likely that blue colors may have become associated with the spoilage of foods, for many molds and fungi exhibit blue-green colors when growing. The growth of microbial contaminants are known to affect the safety, color, flavor, and texture of many food products, likely leading to the avoidance of that color, historically. Expectations of foods based on colors are also culturally related. The specific color and flavor combinations will vary somewhat between cultures but also based on the products marketed to these specific regions. For those of the U.S., blue has likely become associated with raspberry flavor due to its inception and mass marketing in commonly consumed children food products in the latter half of the 20th century.

Relating more specifically to consumer expectations of food products, color has been found to impact not only flavor perception but also its intensity, the overall sweetness, and even saltiness (Clydesdale, 1993; Zampini et al., 2007). Colors have been found to correlated strongly with flavor perception. The perceived or expected flavor of colored foods was found to often correlate with the color of the food to which the flavor was being matched. For example, orange and yellow colored foods were perceived to be orange and lemon flavored products, respectively (Zampini et al., 2007). While red dyed foods were often thought to berry or cherry flavored (Zampini et al., 2007). Interestingly, color has also played a part in the perception of some of the more basic components of taste. Even sweetness and sourness were thought to be correlated with qualities of similarly colored fruits and vegetables. In sucrose solutions dyed yellow, panelists were unable to detect sweetness until high concentrations were reached when compared to solutions dyed red or green; this suggests the color yellow is not often associated with

7 sweetness, perhaps because it is less common in yellow fruits (lemons) and vegetables

(squash) (Clydesdale, 1993). In colored solutions, a significantly higher concentration of acid was necessary to achieve sourness compared to colorless counterpart, suggesting a higher tolerance of acid due to implied sweetness from ripened fruits (Clydesdale, 1993).

Colors in foods can affect several perceived properties relating not only to sensory aspects such as flavor and sweetness but also with characteristics such as safety or general quality. Colors have been linked to health for centuries, in a variety of manners dependent on culture (Singh, 2006). Although only loosely related to food characteristics, certain colors have been thought to relate to certain health characteristics, such as a relationship of the color red to energy and liver health (Singh, 2006). These perceptions may play a role in individual or cultural preferences for foods of certain colors. In medieval times, food colorants were difficult to source and therefore reserved for the upper classes, leading to the belief that food color indicated both nutritional value and medicinal power. Deeply red colored foods were considered to produce rich, full blood; and golden colors promoted divine healing (Burrows, 2009). The psychological effects of environmental colors have also been utilized in the development of restaurant designs. In fast food restaurants, red and yellow colors are often used to gain consumer attention and stimulate appetite. While in more formal restaurants, cool colors such as blue are often utilized to calm and relax customers to extend their stays and indulge in multiple course meals (Singh, 2006).

2.2 Definition of Color

Color is defined as the sensation experienced by an individual when electromagnetic radiation within the visible spectrum falls upon the retina of the eye

(Wrolstad & Smith, 2010). The electromagnetic spectrum includes radiation of various amounts energy from low energy radio waves to high energy gamma ray; visible light is a small portion of the electromagnetic spectrum of comparative middle energy.

Electromagnetic radiation is considered vibrations of magnetic and electric fields that travels in streams of mass-less particles which then travel in wave-like patterns (Brown,

LeMay (Jr.), Bursten, & Murphy, 2006). The distance from one point on a wave to the same point on an adjacent wave (wavelength) differentiates the different classes of electromagnetic radiation; visible light ranges wavelengths of 380-770 nm falling between ultra-violet and infra-red radiation (Wrolstad & Smith, 2010).

Color is not inherent to the object but to the observer. For color to exist three main requirements must be met; these include presence of a colored object, energy within the visible spectrum, and an observer (Wrolstad & Smith, 2010). Visible light strikes the object and can be absorbed, reflected, and/or scattered in varying amounts or degrees.

Selective absorption or reflectance of certain wavelengths of light by the object as well as the amount and type reaching the retina results in the color of an object (Wrolstad &

Smith, 2010). In general, the wavelengths of light absorbed by an object results in color observation of those associated with the complimentary color. For example, absorbance of wavelengths of red light usually results in observation blue-green colors. This pattern is general and not perfect.

2.3 Measurement of Color

Due its importance in food and several other products, the need to effectively measure and communicate color is essential. Colorimetry is the science of color measurement, capable of defining color in mathematical units (Wrolstad & Smith, 2010).

In measuring color, one must also account for the three requirements of color. Current systems were developed heavily from work conducted by the Commission Internationale de l’Eclairage (CIE or International Commission on Illumination).

To account for variability between unique observers, the CIE developed standard observer curves to correspond to human sensitivity to red, blue, and green light. The human eye contains cells that respond to red, blue and green light; therefore the standard observer curves were intended to correspond to human anatomy. To develop these

10 observer curves, people with normal color vision matched the color of single wavelengths of light by mixing different amounts of three primary lights (red, blue, and green)

(Wrolstad & Smith, 2010). The results of these studies were then averaged providing standardized values representing the human response to colors, Figure 2.1. Originally the curves were developed at field of view of 2 degrees, primarily utilizing the fovea which is the retinal area of greatest acuity (Wrolstad & Smith, 2010). However in 1964, the curves were optimized for a field of view of 10 degrees as the cells responsible for color observation spread beyond the fovea (Wrolstad & Smith, 2010).

Source of lighting also plays an important in the observation of the color of an object and must therefore also be controlled in measuring and communicating color.

Metamerism is a phenomenon in which colors appear to match under 1 source of light but then differ when viewed under a different light source. Different light sources produce different spectral distributions of light, Figure 2.1. The spectral distribution of incandescent light (tungsten lamps or Illuminant A) shows a predominance of long wavelength light with less short wavelength light, producing a warmer, redder light

(Wrolstad & Smith, 2010). In contrast, fluorescent lighting (Illuminant F) produces spectral distributions that are generally higher in shorter wavelengths leading to more blue like light, Figure 2.1. The spectral distribution of daylight is represented by the

Illuminants C and D, with D illuminants accounting for UV wavelength absorption and also spectral differences due to temperature (Wrolstad & Smith, 2010). The inclusion of these factors in D illuminants has made them the industry standards in color measurements.

In communicating color data, the object must also be represented. The physical properties of the object determine the mode of measurement and how the sample is to be presented to the instrument. The color of transparent objects is generally analyzed by measurement of the spectral distribution of the light that passes through the object; while opaque objects are analyzed by measurement of the light reflected from the object. The amount (area) of sample analyzed for reflectance is important as well as the length through which light must pass through a transparent sample for color evaluation.

2.4 Color Communication Systems

A color space is a specific organization of colors that can be an arbitrary classification or a mathematically structured system. The development of standardized observer curves and illuminant spectral distributions allowed for conversion of spectral curves of objects to numerical (colorimetric) color spaces.

2.4.1 CIE-XYZ Tristimulus Systems

The concept for tristimulus color communication systems was derived from the 3- component theory of color vision, in which humans only possess receptors for 3 colors and all others are combinations of these colors (Konica Minolta Inc., 2007). Therefore, the development of the standard observer curves was a critical basis of this system.

Spectral data was converted into 3 numerical values: X, Y, and Z according to the formulas of Figure 2.2. These values were useful to numerical define a color, but they were difficult to visualize (Konica Minolta Inc., 2007). To improve this, the X, Y, and Z values were further reduced into the Yxy system which allowed for 2-dimensional plotting of hue (x,y) independent of lightness (Y), Figure 2.2 (Konica Minolta Inc., 2007;

Wrolstad & Smith, 2010). This improved visualization of the color; however there was not equivalent visual spacing of the color. Wavelength distribution in the chromaticity diagram was over-spaced in the green region, being unequal and smaller for red or blue

(Wrolstad & Smith, 2010). Additionally, the same numerical difference in color did not equate to the same visual difference between all colors (Wrolstad & Smith, 2010).

Figure 2.2: Equations to convert spectral data into CIE-XYZ and CIE-L*a*b* color spaces (Wrolstad & Smith, 2010)

2.4.2 CIE-L*a*b*, CIE-L*c*h* Systems

Further color space systems have been developed after the CIE-XYZ system to help alleviate some of the disadvantages of this system. Empirical approaches were used to reduce the unequal color distributions and also improve the visualization of the color from numerical values (Wrolstad & Smith, 2010). These systems were developed with application of the opponent theory of color perception, which states that color is interpreted by the human visual systems by processing signals in antagonistic manners. Four unique hues are thought to exist: red, green, yellow and blue, and all other hues can be described as mixtures of these four hues (D’Zmura, 1991). These distinct colors are furthermore thought to exist in complimentary pairs (red-green, yellow-blue, and black-white) and do not co-exist in singular time points (D’Zmura, 1991). The more commonly used color systems employ these concepts in developing each color space.

If lightness (L*) is considered a 3rd dimension to color (z-axis), the other 2 dimensions (x- & y-axes) would be a* (positive being red, negative being green) and b*

(positive being yellow, negative being blue), Figure 2.3 (Wrolstad & Smith, 2010). The most current system developed by the CIE is a modification of previous similar systems; the equations to convert spectral data to colorimetric data can be found in Figure 2.2. The system also implemented calculation of hue angle (0-360°), Figure 2.3 and chroma

(saturation of color) to better visual numerical values as colors. Hue is considered by humans to be the most critical component of color (Wrolstad & Smith, 2010); therefore a standardized numerical value for it is useful in communicating color.

Figure 2.3: CIE-L*a*b* color space (Artec Testnology, 2016)

2.5 Food Colorants

In addition to the naturally present colors of food, additives to impart colors may also be used in foods for several purposes. According the U.S. Food and Drug

Administration (FDA), a color additive is “any material that is a dye, pigment, or other substance made by a process of synthesis or similar artifice, or extracted, isolated, or otherwise derived, with or without intermediate or final change of identity, from a vegetable, animal, mineral, or other source and that, when added or applied to a food, drug, or cosmetic or to the human body or any part thereof, is capable of imparting a color thereto (21 CFR 70.3).” The purposes of colorants added to foods vary depending on the products but typically include enhancement and correction of already present colors, standardization of colors of raw materials, giving a color identity to otherwise colorless foods, such as sodas or confections, and also accounting for color loss during storage or processing (Newsome et al., 2014; Potera, 2010).

Food colorants are individually regulated apart from other food additives by the

FDA under the Code of Federal Regulations (CFR) title 21, parts 70-74, 80, and 82.

Unlike other food additives, colorants cannot receive status as Generally Recognized as

Safe (GRAS) because they serve no other purpose in the product than to impart color.

Although some GRAS food additives may impart coloration to foods, but there use as

GRAS additives requires they serve purposes other than imparting colors. Food ingredients (such as fruits, vegetables, and spices) which contribute natural color to the product in which they are added are not considered food colorants unless they are deliberately added for the purposes of coloring the product, such as turmeric to impart yellow color to margarine. As such, colorants permitted in foods and other products must undergo evaluations to prove safety, FDA petition, and subsequent approval (21 CFR

70.3). Current FDA regulations permit 2 major classifications of food colorants: certified and exempt from certification.

2.5.1 Certified Colorants

The classification of food colorants as certified or exempt from certification more or less distinguishes whether the FDA views the colorants as synthetic or natural

(Burrows, 2009). Certified colorants are regulated under the CFR Title 21, part 74.

Current permitted certified pigments are organic molecules that have been chemically synthesized; certification requires that each batch of the colorant produced has evaluated for sufficient identity and purity levels (Downham & Collins, 2000). The FDA currently permits 9 certified (synthetic) colorants in foods; however the use of 2 of these is extremely limited. Table 2.1 lists the currently FDA approved certified food colorants

16 and some physical properties. Certified food colorants must be identified by name on products’ ingredient statements, such FD&C Blue No. 1 or shortened as Blue 1 (Frick,

2003). In Europe 17 synthetic pigments are permitted and are labeled by E-number on packaging. (Downham & Collins, 2000).

Table 2.1: List of certified FD&C dyes, their color characterization, and use (Frick,

2003; Sharma et al., 2011)

Common λ FD&C No. E No. Color max Use in U.S. Name (nm) Erythrosine Red No. 3 127 Bright Pink 527 General Allura Red Red No. 40 129 Red 500 General Tartrazine Yellow No. 5 102 Yellow 422 General Sunset Yellow Yellow No .6 110 Orange 480 General Fast Green Green No. 3 NA Turquoise 625 General Brilliant Blue Blue No. 1 133 Sky Blue 630 General Indigotine Blue No. 2 132 Royal Blue 610 General Yellow- Citrus Red Citrus Red No. 2 NA NA Citrus Peel Orange Sausage Orange B Orange B NA Orange NA Casing * NA: not available

Certified colorants are permitted as 2 forms in foods being dyes and lakes. Dyes are water soluble while lakes are virtually insoluble (Frick, 2003). To produce lake colorants, dyes are precipitated onto an aluminum hydrate substrate and made into fine powders. Due to their insolubility, lakes function in products with low solution volume

(high fat systems, such as icings), in products where color migration is problematic, or in dry powdered products (Downham & Collins, 2000). The hue of lakes is dependent on the concentration of dye as well as the particle size of the final product (Downham & 17

Collins, 2000). Of the 7 synthetic dyes approved for general food usage, only 6 of these are permitted in foods as lakes. The lake of Red No. 3 has been excluded from food use due to a carcinogenic response found in rats, although the dye remains legal (Sharma et al., 2011).

2.5.2 Colorants Exempt from Certification

According to current U.S. regulations, colorants that do not have to undergo batch certification may also be added to foods. Despite lack of certification requirements, these colorants still must adhere to specified identity and purity standards as outlined in the

CFR Title 21, part 73. These colorants exempt from certification are generally considered to include “natural” pigments. The FDA has not yet established a legal definition of the term ”natural” which has led to some overall confusion for the food industry as well as consumers; and the term “natural” is not permitted to describe a color additive. Color may be considered natural when it is an inherent property of the food; however once added to color a food product, it is no longer considered natural. Colorants exempt from certification include a variety of type of pigments derived from plant, animal, or mineral sources and also synthesized compounds, despite the thought that colorants exempt from certification are natural. Iron oxides and titanium are naturally occurring minerals that are legal to impart color to foods; however, U.S. regulations require that they synthesized in order to achieve sufficient levels of purity (21 CFR 70.3).

Table 2.2: List and classification of major colorants exempt from certification, their color characterization, and use (21 CFR 70.3)

Classification Additive Source Color Use in U.S. Non- Grape skin Concord grapes beverage extract Red - purple - Anthocyanin food blue Fruit juice Berries General Vegetable juice Carrot, cabbage, etc General Betalain Beet powder Beet (Beta vulgaris) Pink - red General Caramel Caramel Heating of sugars Brown General Carminic Cochineal Dactylopius coccus Orange - red General Acid extract Annatto Bixa orellana L. General Synthesized, Bacterium (Paracoccus Astaxanthin Fish feed carotinifaciens), Yeast (Phaffia rhodozyma) β-carotene Natural (carrots) or Synthesized General Fish feed & Canthaxanthin Synthesized chicken skin Carotenoid Carrot oil Carrot Yellow - General Corn endosperm orange - red Yellow corn Chicken feed oil Paprika Paprika (Capsicum annum L.) General (oleoresin) Saffron Stigma (Crocus sativus L.) General Aztec Marigold (Tagetes Tagetes Chicken feed erecta L.) Tomato lycopene Tomato (Solanum lycopersicum) General Citrus-based Sodium copper Chlorophyll Alfalfa (Medicago sativa) Green dry beverage chlorophyllin mixes Turmeric Curcuminoid Rhizome (Curcuma longa L.) Yellow General (oleoresin) Yellow, Sausage Iron oxide Synthesized orange, red, casing, pet brown, black feed Mineral General, Mica pearlescent Synthesized Pearlescent restricted Titanium dioxide Synthesized White General General, Phycocyanin Spirulina extract Algae (Arthrospira platensis) Blue-green restricted Vitamin Riboflavin Microbial fermentation Yellow General

As observed from Table 2.2, colorants exempt from certification are derived from several sources and can impart a variety of colors to foods. They are capable of imparting 19 several hues dependent on several parameters of the food product and regulatory status of the colorant. Although not all pigments are directly taken from natural sources, many are chemically modified or synthesized to be identical to naturally occurring pigments. These types of pigments typically carry more limitations when used to color foods compared to synthetic colorants (or certified colorants).

In general, naturally derived pigments used as food colorants are typically more expensive than synthetic counterparts. This can be related to the need for large amounts of raw materials to obtain sufficient quantities of pigments, but also, higher costs are associated with them due to the need for higher usage levels to achieve comparable color intensities to synthetic pigments (Rodriguez-Amaya, 2016; Wrolstad & Culver, 2012).

Due to the source of pigment, many naturally sourced colorants can also impart undesirable flavors and aromas to products. For example, red pigments sourced from red radishes, red cabbages, or red beets may also carry varying amounts of flavor contributing compounds from these vegetables which could then be imparted to the food product. For some synthetic colorants, especially blues, natural options are often unable to match the hue or other color characteristics and require additional work to source and optimize natural alternatives (Wrolstad & Culver, 2012). Naturally derived pigments as food colorants also face several additional challenges compared to synthesized pigments primarily related to lower stability. Most are more sensitive to heat, light, oxygen resulting in color loss or alterations in hue (Lakshmi, 2014; Rodriguez-Amaya, 2016;

Wrolstad & Culver, 2012). Others may be sensitive to environmental matrix conditions

20 such pH, protein, presence of metal ions, or other organic compounds, such as ascorbic acid.

2.5.3 Trends in Coloring of Foods

Despite the comparative limitations of naturally derived pigments, thorough safety evaluations of all food pigments, and strict regulations, the market of food colorants has been shifting substantially toward the replacement of synthetic colorants with naturally sourced alternatives. In 2013, the value of naturally sourced colorants had surpassed that of synthetic colorants (Leatherhead Food Research and Mintel, 2013). Of newly developed products launches, the use of natural colorants has outweighed that of synthetic colorants by 2:1 globally (Leatherhead Food Research and Mintel, 2013). This trend is likely to continue with large companies such as Mars, Inc. and Nestlé having announced plans to remove artificial colorants from their products within the next few years (Mars Incorporated, 2016; Nestlé USA, 2015).

The replacement of artificial dyes with naturally sourced alternatives is being driven by several factors. Consumer demand for “natural” products and ingredients is likely one of the largest contributing factors for the food industry. In a survey of survey grocery shoppers, 55% of ranked the attribute “All natural (no artificial preservatives, additives, colors, sweeteners)” as somewhat to very important (Browne, 2012).

Consumer desire for “natural” food products has been a driving force changes in formulations using less artificial additives or use of more generally known food substances as sources of the food additives, including colorants. Under current regulations, anthocyanins as food colorants may be added to foods primarily in the form

21 of fruit or vegetable juices (21 CFR 73); the ingredient statement bears this same information and is likely familiar to the common consumer.

Other factors leading to the replacement of artificial food dyes with naturally derived counterparts may be related to potential health and safety concerns. Food colorants, certified or exempt, typically are mixtures of the coloring compound and also other constituents. These contaminants may be derived from the starting material or be developed during production. Some of the impurities may include possible carcinogens; however they would not be present in levels to be of concern (Kobylewski & Jacobson,

2010). Possible allergenicity has been a consumer concern with some certified colorants; however evidence is inconclusive and may be only be true for sensitive individuals.

Originally proposed in the 1970’s, the possible link between hyperactivity in children and artificial colorants was further evaluated with release of the “Southampton

Study”(Feingold, 1975; McCann et al., 2007). Children of 2 age groups were fed two solutions of sucrose, blends of Azo type dyes (ex: FD&C Red No. 40, Yellow No. 5, and

Yellow No. 6), and preservatives (sodium benzoate); they were then evaluated for hyperactive symptoms over a 7 day period. Some changes in behavior were observed; however, findings between both age groups and both solutions were inconsistent and only weakly significant (McCann et al., 2007). Due to the mix of pigments and preservatives, it is unclear as to which component or whether the mixture of components was responsible for the small changes in behavior. For some of these reasons the FDA and EU have further reviewed the safety of food colorants. The EU has mandated that labels of food containing the synthetic colorants used in the “Southampton Study” bear warning

22 labels indicating the product may have an adverse effect on activity and attention in children (FDA; CFSAN, 2011). The FDA has concluded that causality has not been proven, and warning labels on foods are unnecessary (FDA; CFSAN, 2011).

The most commonly used colorants in foods are the red and yellow hued pigments. The synthetic red and yellow pigments (FD&C Red No. 40, Yellow No. 5 and

Yellow No. 6) used to account for 90% of all the food colorants used in the industry

(Potera, 2010). The replacement of these synthetic dyes with naturally derived alternatives is considered simpler due to ease of matching hues and the wide variety of options (Newsome et al., 2014). Naturally occurring pigments such anthocyanin, carotenoid, and betalain derivatives are abundant in nature and can impart a wide array of hues in typical food matrices including yellows, oranges, reds, and pinks. These different chemical properties of these different pigments also allow for application in both hydrophilic and lipophilic environments. Naturally occurring pigments are increasingly becoming more popular for their potential health promoting effects in addition to their color expression. Regardless of the comparative low usage levels of blue food colorants, there is considerable industrial demand for naturally derived alternatives for them due to the limited number of sources and inability to match the color characteristics of synthetic dyes.

2.6 Blue Colorants

A blue pigment can be defined as a molecule that adsorbs red light (600 nm region) and appears blue by ordinary color perception, Figure 2.4 (Newsome et al., 2014).

1.2

0.8

Absorbance 0.4

0 380 430 480 530 580 630 680 Wavelength (nm)

Figure 2.4: Visible absorbance (380-700 nm) and solution of FD&C Blue No. 1 in H2O

Most molecules absorb UV energy causing promotion of σ electrons to higher energy σ* or π* orbitals. Fewer molecules exhibit lower energy transitions π → π* or n

→ π* from visible light absorbance, resulting in color expression; these molecules typically contain double bond conjugation (Newsome et al., 2014). Linear conjugation alone does not result in absorbance of lower energy red light, as demonstrated by carotenoid pigments appearing yellow-orange-red. In addition to double bond conjugation, blue pigments typically contain aromatic ring systems, heteroatoms (−NH2,

−OH, =O, −Cl, etc.), and/or ionic charges (Newsome et al., 2014). Transition metal complexes can also result in blue color expression as their d-orbital electrons allow for lower energy electron transitions.

2.6.1 Synthetic Blue Colorants

Blue No. 1 Blue No. 2

Figure 2.5: Chemical structure of FD&C certified blue colorants (Blue No. 1 and Blue

No. 2), (Minioti, Sakellariou, & Thomaidis, 2007)

Currently, two synthetic colorants are permitted in foods as certified colorants,

Table 2.1, and have a long history of use in foods. These include brilliant blue (FD&C

Blue No.1) and indigotine (FD&C Blue No. 2). Together they totaled use of 7.7% of all certified food colorants in foods in 2007, with Blue No. 1 accounting for more than half of this total (Sharma et al., 2011). Brilliant blue (FD&C Blue No. 1) has a molecular weight 792.86 and is water soluble. It is classified as a triphenylmethane compound

(Ahmadiani, 2012). Having a λmax ≈ 630 nm, this dye expresses a brilliant blue color

(with slight green tones) that is widely accepted and used in the food industry for a variety of products. Indigotine (FD&C Blue No. 2) has a slightly lower λmax ≈ 610 nm, so it absorbs lesser amounts of red light, which is observed in its comparatively greyer blue color expression. This dye is commonly used to dye fabrics and is typically responsible for the color of blue jeans. Indigotine is classified as an indigoid compound and has a

25 molecular weight of 466.36 (Sharma et al., 2011). Both are available and permitted in foods as both lakes and dyes.

2.6.2 Blue Colorants Derived from Natural Sources

Blue colors are not necessarily uncommon in nature; however blue pigments are relatively uncommon compared to other pigments, due to the more complex chemical structural requirements to absorb red light (Newsome et al., 2014). Additionally many of the blue colors observed in nature are not due to presence of pigments but rather are result of physical nanoscale structuring that causes reflectance of blue light. The blue colors of most animals, such as butterfly wings or bird feather, are formed by this mechanism; while blue color of plants and microbes are due to presence of pigments

(Newsome et al., 2014).

One of the oldest known colorants to mankind includes indigo (an indole alkaloid or indigoid compound), chemically similar to FD&C Blue No. 2. Although being a naturally sourced alternative to indigotine, this pigment is not currently legal use in foods in the U.S. It has a long historical use for dyeing textiles and use in cosmetics (Jespersen,

Strømdahl, Olsen, & Skibsted, 2005). The pigment was primarily sourced from indigo plants (Indigoera sp.) or woad (Isatis tinctoria), which contain the precursors of the pigment. Indoxyl, a glycoside of indicant or the ester of isatin B which undergo hydrolyzation and oxidation to eventual produce the pigmented indigo and indigotin

(Jespersen et al., 2005). Indigo has very poor solubility and low stability to light exposure. It has a λmax ≈ 605 - 610 nm, resulting in a grey-blue color expression, making

26 it unable to replicate the color characteristics of the more commonly used Blue No. 1.

These characteristics further limit its application in foods.

Blue iridoid derivatives also have a long historical use and therefore a relatively accepted safety record. Similar to the indigo, the pigmented molecule is formed from uncolored precursors sourced from plants of the Rubiaceae family, including gardenias or the huito fruit (Wu, Ford, & Horn, 2009). The colorless iridoid glycosides geniposide and gardenoside are released by juicing the plants which are then hydrolyzed by β- glucosidase releasing genipin and glucose (Jespersen et al., 2005). Blue color is the result of condensation of genipin with amino acids; attachment of different amino acids will result in expression of different blue hues (Wu et al., 2009). These pigments, again, have

λmax ≈ 600 nm, indicating color to be more similar to FD&C Blue No. 2 (Ahmadiani,

2012). Iridoids sourced from gardenia flowers are not legal in the US or EU but are legal in Japan and have been used in China and Korea (Jespersen et al., 2005; Newsome et al.,

2014). When derived from huito and other fruit combinations, the colorant is offered a juice for color according to CFR (Wild Flavors and Specialty Ingredients, 2016).

The options for blue colorants derived from natural sources were recently expanded in the US with the approval of Spirulina extract as a colorant exempt from certification (21 CFR 73). Spirulina extract is sourced from the cyanobacteria Arthrospira platensis and contains the protein complex phycocyanin (21 CFR 73). This compound is the pigment responsible for the blue-green coloration of the extract, and its chemical structure has not yet been fully elucidated (Jespersen et al., 2005). The λmax of the pigment was found to vary from 616 – 620 nm, responding to change in pH, suggesting

27 its color characteristics to be more similar to FD&C Blue No. 1 (Jespersen et al., 2005).

The pigment has also been found to be unstable with light exposure and extremely sensitive to heat, losing color at temperatures ≥ 45 °C (Jespersen et al., 2005). With its bright blue color, this pigment has shown potential in some products including ice creams, confections, and products processed with thermal treatments.

The blue-green coloration of the marine diatom Haslea ostrearia has recently been further evaluated. The bivalve produces the water soluble pigment marennine, which adheres to the gills of the oyster causing a desirable greening (Gastineau et al.,

2014). The structure of the pigment is still unknown, and its color was found to be responsive to pH, being blue in acidic conditions and greener in alkaline pH (Pouvreau et al., 2006). Before use in foods, the pigment must first undergo toxicological evaluations and through the regulatory petition process. The oyster, source of pigment, has been consumed for centuries which may suggest low toxicity (Gastineau et al., 2014).

Tetrapyrrole pigments are known to create almost all colors of the visible spectrum, and some of which express blue hues (Newsome et al., 2014). In the animal kingdom, biliverdin is one of the few blue pigments and is responsible for blue color of bruises (Newsome et al., 2014). Recently, tetrapyrrole compounds were also found to be responsible for the blue-green discoloration of processed garlic (Block, 2010). The formation of these blue heterocyclic pyrrole compounds involves several steps. General reactions include the cleavage of γ-gluatmyl groups, action of allinase on isoalliin, rearrangement to thial-thial S-oxide, and condensation of thiocarbonyls with amino acids

(Block, 2010). There is an inverse relationship between the increasing concentration of

28 pigments and decrease in thiosulfinate compounds, which should also decrease the amount of allium flavors (Block, 2010). As many as 8 pyrolle compounds have been identified as blue-green pigments; all were tri- or tetrapyrolles (Block, 2010). The blue pyrolle pigment bactobilin have also been identified. It is produced by bacteria isolates by

Clostridium tetanopmorphum and Propionibacterium shermanii (Newsome et al., 2014); the latter is used in the production of Swiss cheese. Further investigations will be required if these pigments are to be considered for food use.

Some other blue pigments found in nature include azulene compounds, found to occur in some mushroom and plant species. Guaiazulene is currently approved for use in cosmetics, but the use of these compounds in foods tend to be fairly limited due to their lipophilicity, low molar absorptivity, and low stability (Newsome et al., 2014).

Trichotomine is a bis(indole) alkaloid that naturally occurs in the fruits Clerodendron trichotomum, a flowering shrub native to Asia (Koda, Ichi, Odake, Furuta, & Sekiya,

1992). The pigment has λmax of 618 and 630 nm in methanol, but may be limited for food use due to presence in low levels in the plant (Koda et al., 1992). It is thought to be suitable for food coloring due to its color, stability, and safety (Koda et al., 1992); however, regulatory approval must first be obtained. Additional alternatives for synthetic blue food colorants may be found in anthocyanins (ACN) which are the phenolic compounds responsible for many of red, purple, and blue colors of most commonly consumed fruits and vegetables.

2.7 Anthocyanins

Anthocyanins compose the largest group of water soluble naturally occurring pigments, with more than > 700 unique structures having been identified (Andersen &

Jordheim, 2014). These phenolic compounds are responsible for a wide range colors in plants. The word anthocyanin was derived from the Greek terms anthos and kyanos respectively meaning flower and blue (Schwartz, von Elbe, & Giusti, 2008). In plants, these pigments function to protect against harmful UV irradiation, attract animals for pollination and seed dispersion, may sequester toxic heavy metals in vivo, and perhaps provide antimicrobial activities (Hale et al., 2002; He & Giusti, 2010)

2.7.1 Structural Components

Anthocyanins are a subgroup of flavonoid compounds which are characterized by sharing C6C3C6 carbon skeletal backbone, Figure 2.6; they are differentiated from other flavonoids by the presence of 2 carbon double bonds in the C ring (Ge et al., 2015; Schwartz et al., 2008). This double bond also imparts a positive charge to the molecule in acidic pH (Ge et al., 2015). The basic structure of anthocyanins is 2-phenylbenzopyrylium of flavylium salt, with varying degrees of substitution

(Schwartz et al., 2008). Differentiation between different anthocyanidins, anthocyanins lacking glycosylation, depends on the amount and type of substitution patterns on the B ring and unique substitutions generally on C3, C5, C6, and C7 of the molecule (Andersen

& Jordheim, 2006).

Figure 2.6 demonstrates the structural composition of the most commonly encountered anthocyanins in edible produce. The 6 most common anthocyanins differ

30 primarily in amount of hydroxylation or methoxylation, but more than 30 unique aglycones (anthocyanidins) have been identified (Andersen & Jordheim, 2006). These seemingly small molecular differences impact the color expression of the pigments; generally increasing amount of substitution results in bathochromic shifts on the absorption spectra, resulting in slightly bluer colors, Figure 2.7. The most prevalent anthocyanidin in nature has been found to be Cyanidin (Cy), which bears 2 hydroxyl groups on the B ring (Andersen & Jordheim, 2006). The other hydroxylated anthocyanidins delphinidin and pelargonidin follow in abundance (Andersen & Jordheim,

2006).

Figure 2.6: Basic structure of a flavonoid and the most common anthocyanins found in nature

Figure 2.7: Representation of different color expression of major anthocyanin aglycones adapted from Ananga, Georgiev, Ochieng, Phills, & Tsolova, 2013

Glycosylation, attachment of sugar moieties to the anthocyanidin through O- linkages, improves the stability of the pigment and also increases its water solubility (He

& Giusti, 2010). Aglycones of anthocyanins are rarely found in nature due to the inherent instability of the molecules. Those reported include some uncommon aglycones lacking an oxygen attachment to C3 (3-deoxyanthocyanidins) are reported to occur in nature without glycosylation or may be hydrolysis products formed during pigment extraction

(Andersen & Jordheim, 2006; Schwartz et al., 2008). The stability of the molecule is increased by allowing for formation of intramolecular H-bonding within the anthocyanin

(He & Giusti, 2010). Typically site of glycosylation is C3 of the aglycone followed by C5; less common attachments are known to occur at C7, C3', C4', and C5' (Schwartz et al.,

2008). A variety of sugars have been reported as attachments to anthocyanins from monosaccharides to trisaccharides; tetrasaccharide glycosylation has not been reported

(Andersen & Jordheim, 2006). Glucose and rhamnose are the most common glycosylating moieties, and other common saccharides include galactose, arabinose, xylose, rutinose, sambubiose, and more (He & Giusti, 2010). Glycosylation typically

32 results on small bathochromic shifts on the aglycone, creating more pink-red colors compared to more orange-red colors.

Anthocyanins can also experience further molecular substitution in nature with attachment of acyl (acid) moieties to glycosyl residues; more than 65% of reported anthocyanins are acylated (Andersen & Jordheim, 2006). These structural modifications further add to the diversity of anthocyanin compoundss due to nature, number and linkages positions of the acyl moieties. Acylating groups can be esterified to glycosyl residues at all carbon position, but they are most commonly found bound to C3 of the aglycone (Schwartz et al., 2008). Both aromatic and aliphatic acids have been reported as acyl moieties of anthocyanins. The most common aromatic acid attachments are hydroxycinnamic acid derivatives: p-coumaric > caffeic > ferulic > sinapic > etc.

(Andersen & Jordheim, 2006). Aliphatic acylation with malonic acid is also frequently encountered; and acetic, malic, oxalic, succinic, and tartaric acids are also reported to be found as acyl groups to anthocyanins in more restricted distributions (Andersen &

Jordheim, 2006). Acylation of anthocyanins has been found to further impact color expression of the pigments and also to increase the overall stability of the molecule in solution.

2.7.2 Potential Health Benefits

Interest in dietary anthocyanins and other polyphenolic compounds has been growing in the food industry not only as food colorants but also for potential added health benefits. Epidemiological studies have shown the occurrence of some chronic diseases to have a reverse correlative relationship with diets high in polyphenolic compounds; these

33 have gained further support through in vitro and in vivo evaluations (He & Giusti, 2010;

Wallace & Giusti, 2014).

Anthocyanins are presumed to have occurred in plants before man and therefore to have generally always contributed to part of the diet making it unlikely that anthocyanins have derogatory health implications (Bridle & Timberlake, 1997). Further works have also shown them to be non-toxic and non-mutagenic (Bridle & Timberlake,

1997). In a 2 generational study with rats, dietary anthocyanins were found to have a no observed effect level (NOEL) of 225 mg/kg, which indicates very low toxicity and also can be calculated to an acceptable daily intake (ADI) for humans of 2.5 mg/kg (Clifford,

2000). Human dietary exposure to anthocyanins have been estimated to be as high as 1 mg/kg bw/day for adults and 2 mg/kg bw/day for children, below the ADI (Efsa, 2013).

Related to the phenolic structural components of these compounds, anthocyanins exhibit strong antioxidant activates that likely are related to their potential health benefits (He &

Giusti, 2010). Studies have shown their properties to include protection against liver injuries, reduction of blood pressure, eyesight improvement, some anti-inflammatory and antimicrobial activities, inhibition of mutagens, and cytostatic and cytotoxic activities of human cancer cells (Konczak & Zhang, 2004). Some further health promoting properties include cardiovascular disease prevention, obesity control, and diabetes alleviation (He &

Giusti, 2010). Use of anthocyanins as naturally derived food colorants can not only impart a wide spectrum of colors, improve consumer perception of food products, but also add health benefits beyond standard caloric needs.

2.7.3 Anthocyanins as Food Colorants

Due to the wide array of colors of many fruits and vegetables for which anthocyanins are responsible, they are well known as potential food coloring additives. Their use as food coloring additives does have limitations due to the chemical nature of the pigments. The stability and color expression of anthocyanins is dependent on several factors including pH; acylation and glycosylation; and light, heat and oxygen exposure. Of these factors, pH has been considered the most important (Mazza & Brouillard, 1987). In solution, anthocyanins exist in a structural equilibrium of different resonant species which express different colors as response to the concentration of protons available, Figure 2.8.

Anthocyanins are most stable in an acidic environment in the cationic flavylium form.

Most food and beverage products are slightly to moderately acidic in pH, and in these conditions, anthocyanins typically appear in red-purple structural equilibria. The natural chemistry of anthocyanins (pH dependent structurally equilibria) and stability allows for a wide selection of red-type hues in foods.

Figure 2.8: Major anthocyanin structural forms and color expression due to pH dependence adapted from Houbiers, Lima, Maçanita, & Santos, 1998

Due to the formation of colorless structural forms occurring in moderately neutral pH (3-6), some anthocyanins may also be limited as food colorants in some applications.

Many products have a pH in this range, such as yogurts, and in these conditions, anthocyanins typically have low color expression and stability. To potentially combat this, large amounts of coloring extracts may be necessary which could then impart undesirable flavors from the original plant source to the product. Selection of more chemically complex anthocyanins could also be effective in maintaining color expression by anthocyanins in these conditions. Acylated anthocyanins, those pigments bearing acid moieties, generally exhibit increased stability compared to counterparts lacking acylation

(M. M. Giusti & Wrolstad, 2003). Acylation is also believed to increase the stability of

36 anthocyanins to processing, storage, and other environmental factors (M. M. Giusti &

Wrolstad, 2003). The stacking of the components of the molecule are thought to add stability to molecule and its color by protecting the C ring from hydration by water molecules and avoiding loss of conjugated double bonds.

Acylated anthocyanins from food sources are more often encountered in vegetables while simpler glycosylated anthocyanins are most frequently abundant in fruits. Some sources abundant in acylated anthocyanins that have also been used in industrial food applications include red radishes, red cabbage, black carrots, some grapes varieties, and purple sweet potatoes (M. M. Giusti & Wrolstad, 2003). Acylated pelargonidin derivatives from red radish or red fleshed potatoes are of particular interest as potential alternatives for FD&C Red No. 40 in some applications (such as maraschino cherries or juice systems) (M. M. Giusti & Wrolstad, 2003). Under these conditions (pH

≈ 3.5), the anthocyanin extracts closely resembled the color characteristics of the synthetic red dye (M. M. Giusti & Wrolstad, 2003). In some more challenging matrices, such as yogurt (pH 4.2 – 4.5), extracts of red radish or carrot provided desirable red hues while red cabbage anthocyanins provided a purple hue similar to the color of blueberries

(M. M. Giusti & Wrolstad, 2003). Despite the wide variety of hues more typically anthocyanins can produce in nature, most do not reproduce blue colors in the acidic conditions typical to most food products or provide the necessary color characteristics to act as alternatives to synthetic blue food colorants.

2.7.4 Mechanisms to Produce Blue Colors

Anthocyanins are the basis for many of the blue colors observed in edible fruits, vegetables, and flowers. The anthocyanin as a monomer never exhibits blue color in weakly acidic conditions but expresses purple hues until rapidly losing color (Yoshida et al., 2009). The mechanisms for blue color expression by anthocyanins can be found in nature and can be replicated in order to investigate the pigments as potential alternatives for synthetic blue food colorants. Further derivatization of anthocyanins can lead to larger, water soluble pigments that extend pH range of the blue color expression of these types of pigments. In addition to molecular modification and pH adjustment (increase to alkaline conditions), mechanisms such as anthocyanin self-association, molecular copigmentation, and metal chelation have been proposed to result in alterations of the visible light absorbance by the pigments and possible blue color expression in acidic pH

(Yoshida et al., 2009).

2.7.4.1 Derivatization of Anthocyanins

Anthocyanins can undergo chemical transformation during storage in solution resulting in pyranoanthocyanins and their related polymers found to occur in some products such as red wines occurring during and after the aging process. These anthocyanin derivatives were originally thought to be mediated by acetaldehyde and to result in condensation between anthocyanins and flavanols (Mateus, Oliveira, Haettich-

Motta, & de Freitas, 2004). Several other compounds have since been identified to play a role in production of pyranoanthocyanins, which as monomers exhibit a more orange-red color than their anthocyanin predecessors (Mateus et al., 2004). The derivatives

38 exhibiting blue hues in acidic pH are typically larger molecules in which anthocyanins and flavanols are linked by vinyl bridges (Mateus et al., 2004). These molecules, termed portisins, are thought to be obtained through reaction of pyranoanthocyanins with other compounds that could include flavanols (catechins) or other phenolic compounds followed by decarboxylation, dehydration, and oxidation (Mateus et al., 2004). These final steps are thought to yield the blue coloration by production of extended conjugated

π electron systems. Recently, dimers of pyranoanthocyanins have been discovered in port wine that have been found to produce blue colors in very acidic pH (Oliveira et al.,

2010). One group of the pigments was shown to be stable in pH 2, expressing a λmax of

730 nm and turquoise blue; the other group showed λmax of 680 nm, strongly indicative of blue color expression (Oliveira et al., 2010). Several challenges would first have to be overcome in considering the use of these pigments as food colorants. The low levels found to occur in wines would be a limitation, the safety of the pigments would have to evaluated, and regulatory approval must also be achieve.

2.7.4.2 Anthocyanin Self-Association

The idea of self-association of anthocyanins was proposed partially as another means to account for the large variability of hues expressed by anthocyanins, especially in floral systems. The theory also served the purpose of further explaining stabilization mechanisms of the pigments (Yoshida et al., 2009). It is known that anthocyanins are more stable in concentrated solutions rather than in comparatively dilute solutions likely due to molecular stacking occurring by hydrophobic interactions between aromatic rings, the same mechanisms driving molecular copigmentation (Yoshida et al., 2009).

Self-association of anthocyanins is thought to play an important role in the colors of young wines and has been furthered investigated in model systems to better understand the color evolution of associated anthocyanins. The association of the pigments was confirmed to occur as evidenced by evolution of the color of the solutions (González-

Manzano, Santos-Buelga, Dueñas, Rivas-Gonzalo, & Escribano-Bailón, 2008). Largest impacts on color were found to occur as decreases in L* values and increases in chroma, equating to intensification of the observed color (González-Manzano et al., 2008). Hue angle, however, was found to increase and approach more red-orange like colors rather than purple-blue (González-Manzano et al., 2008). Broadening of the peak of the absorbance spectra of isolated anthocyanins was observed in other studies, accompanied by a hypsochromic shift or decrease in λmax (Yoshida et al., 2009). Self-association, therefore, is likely not the mechanism of blue color expression by anthocyanins.

2.7.4.3 Copigmentation of Anthocyanins

Molecular copigmentation is another means to further explain the large color variation of anthocyanins found in nature; its chemistry is similar to self-association.

Copigmentation occurs when essentially colorless molecules (free or covalently bound to the chromophore) are thought to stack on the planar nucleus of the anthocyanin, interacting through hydrophobic interactions or Π-Π stacking. This stacking results in changes in visible light absorbance by the pigment and therefore alterations its observed color (Di Meo, Sancho Garcia, Dangles, & Trouillas, 2012; Malien-Aubert, Dangles, &

Amiot, 2001). Copigmentation may occur intermolecularly, between free molecules in

40 solutions and anthocyanins, and also intramolecularly, within the same molecule between acyl moieties and the chromophore.

The effects of intermolecular copigmentation depend on several factors including molecular structures, individual reactant concentrations, and their ratios (Gómez-Míguez,

González-Manzano, Teresa Escribano-Bailón, Heredia, & Santos-Buelga, 2006).

Hydroxycinnamic acid derivatives evaluated as intermolecular copigments were generally observed to have small effects on the spectral properties of anthocyanins, inducing bathochromic shifts of up to 18 nm (Dimitrić Marković, Petranović, & Baranac,

2000, 2005). The responses were dose dependent; bathochromic shifts increased as concentration of copigments was increased while anthocyanin concentration was constant

(Dimitrić Marković et al., 2000, 2005). It was also observed the larger and more heavily substituted hydroxycinnamic acids induced larger bathochromic responses: sinapic > ferulic > caffeic acids (Dimitrić Marković et al., 2000, 2005). Copigmentation with additional phenolic compounds on malvidin3-glucoside also found the structure of the copigmenting molecule to affect not only the degree of bathochromic shift but also the overall stability of the complex. Flavan-3-ols (such as catechins) were least effective as copigments but monomeric flavanols (myricitrin and quercitrin) acted as strongest copigments in this study (Gómez-Míguez et al., 2006). Intermolecular copigmentation was also found to be a function of time. Hydroxycinnamic acids showed measureable effects much sooner than catechins or procyanidin B2 which required 2-3 weeks in the parameters of this study (Gómez-Míguez et al., 2006).

Figure 2.9: Proposed stacking conformations for stabilization mechanisms of acylated anthocyanins adapted from Jackman & Smith, 1996

Intramolecular copigmentation is considered a more efficient mechanism than intermolecular copigmentation for color adjustment of anthocyanins. The participating components (the anthocyanin and its acyl moiety) are better spaced for interaction by the covalent linkages between the acyl and glucosyl moieties, Figure 2.9 (Malien-Aubert et al., 2001). In the same conditions, increasing degree of acylation of the same structures on the same chromophore (pelargonidin-3-sophoroside-5-glucoside) was found to lead to increasing λmax of the molecules (Dangles, Saito, & Brouillard, 1993a). Acylation with hydroxycinnamic acid derivatives was found to reduce the rate and extent of hydration of the chromophore resulting in both increasing molecular and color stability in wider environmental conditions (Dangles, Saito, & Brouillard, 1993b). Intramolecular copigmentation by acylation has been thought to help anthocyanins produce more blue like colors in lower pH than nonacylated counterparts, such as the pigments derived from the butterfly pea flower (Clitoria ternatea), Figure 2.10 (Abdullah, Lee, & Lee, 2010).

Figure 2.10: Color expression of butterfly pea flower (Clitoria ternatea) anthocyanins in solutions pH 3-8

The anthocyanins of this flower possess a relatively uncommon acylation pattern occurring on the B ring of the molecule; they have been shown to express blue hues in moderately acidic pH 5-7 (Abdullah et al., 2010). More commonly encountered acylation patterns on C3 of the anthocyanin, such as those found in red cabbage or purple sweet potato, have been found to express blue colors similar to FD&C Blue No. 2 in pH 7-8

(Ahmadiani, 2012).

2.7.4.4 Metal Chelation and Complexation

Historically, the interaction between anthocyanins and metals were investigated primarily to better understand the pigmentation of floral systems (Buchweitz, Carle, &

Kammerer, 2012). The color of hydrangea flowers has been known to shift from pink-red hues to purple-blue with the addition of aluminum to acidic soil systems. The low pH of the soil allows for solubilization of the metallic cation which then can be transported through root system of the plant where it is then localized in anthocyanin vacuoles resulting in color change (Schreiber et al., 2010). It has long been known that

43 anthocyanins bearing at least 2 free hydroxyl groups on the B ring could be qualitatively recognized by development of blue like hues with reaction of iron(III) salts (Bayer,

Egeter, Fink, Nether, & Wegmann, 1966).

For anthocyanins to be capable of metallic cation chelation and express purple- blue colors, the molecule must bear at least 2 free hydroxyl groups on the B ring. As the cation approaches the anthocyanins, it acts in competition of the hydrogen ions for the binding sites. The metal ion displaces these hydrogen ions resulting in conversion of the flavylium cation (red molecule) to quinonoidal bases (blue molecules) (Schreiber et al.,

2010); Figure 2.11 demonstrates this mechanism. This structural conversion is also thought to be accompanied by association with another anthocyanin; the stacking of the molecules is coordinated by the metal ion and helps to stabilize the complex (Schreiber et al., 2010). Various metal ions have been found to induce bathochromic shifts on anthocyanins, but the effects are limited to cations with valency > 1+ (Bayer et al., 1966).

Some include aluminum3+, iron3+, tin4+, titanium3+, and chromium3+ (Bayer and others

1966).

Figure 2.11: Proposed mechanism of metal ion (Mn+) chelation and structural conversion of anthocyanins adapted from Schreiber et al., 2010 44

In the cases such as for some juices or canned fruits, metal chelation by anthocyanins resulting in undesirable color change has been considered a defect.

Cyanidin and delphinidin derivatives from different berries were evaluated for color alterations in the presence of some metallic cations (ferrous iron, ferric iron, and tin) encountered in canning materials. In this study, treatment with iron resulted in brown colors; however, tin was shown to lead to blue color expression by these anthocyanins

(Pyysalo & Kuusi, 1973). Other blue chelates have been shown to form in pH 4-7 with

2+ metals ions of tin(IV), titanium(III), chromium(III), and UO2 (Bayer et al., 1966). In methanol and ethanol at pH 3, large bathochromic shifts on cyanidin derivatives were induced by chelation of lead (25 nm), chromium (152 nm), or cadmium (139 nm)

(Ukwueze, Nwadinigwe, Okoye, & Okoye, 2009). These cations are trace metals in naturally occurring systems and unlikely to be used extensively in food products for some safety concerns, these reactions further illustrate the potential to develop blue colors by anthocyanins in acidic pH.

The chelation of aluminum by anthocyanins has been more extensively studied due to its relationship with the blue coloration of hydrangea floral systems. This cation may have better potential food application as aluminum sulfate is considered a GRAS food additive (21 CFR 182.1125). Relatively stable complexes between delphinidin and aluminum have been found to form in acidified ethanol in a wide pH range (Schreiber et al., 2010). In these systems, color expression of delphinidin started red becoming violet and then blue as pH and anthocyanin concentration were maintained but the aluminum content was increased (Schreiber et al., 2010). Aluminum complexes with synthesized

45 anthocyanin (cyanidin) analogues have been shown to develop in aqueous models, pH 2-

5, resulting in violet colorations (Dangles, Elhabiri, & Brouillard, 1994a). Chelation of aluminum allowed for not only bathochromic shift, an increase in λmax, but also a hyperchromic shift, an increase absorbance at the λmax, Figure 2.12. This indicates not only a shift from red hues to more purple-blue colors but also an intensification of the color so less may be necessary to color products. Complexation of aluminum by synthetic or natural anthocyanins stabilizes the blue quinoidal base against oxygen induced degradation (Moncada et al., 2003). Although not known to occur in nature, an anthocyanin bearing a catechol moiety on the A ring was found to also chelate aluminum and express blue colors (Moncada et al., 2003). Aluminum chelation has allowed for development of some anthocyanin based blue colorants; however, treatment with low proportions of ferric iron resulted in larger bathochromic shifts which may allow for more blue color development (Buchweitz, Carle, et al., 2012).

Figure 2.12: Cyanidin derivatives (chokeberry anthocyanins) treated with factorial excesses of Al3+ (0-100× [ACN]) in pH 4

Other recent works have focused on the role of anthocyanin structure and buffer selection in the production of anthocyanin metal complexes in pectin stabilized systems

(Buchweitz, Nagel, Carle, & Kammerer, 2012). Delphinidin derivatives, having 3 free hydroxyl groups on the B ring, treated with ferric iron showed greatest bathochromic shifts and most blue like hues in the widest pH range compared to cyandin and petunidin glucosides (Buchweitz, Carle, et al., 2012). Similarly the anthocyanin metal chelates with the highest stability during storage over 17 days were those with highest degree of B ring substitution: delphinidin > cyanidin > petunidin (Buchweitz, Carle, et al., 2012). The media of study was also shown to be an important factor in the evaluation of metal 47 chelation by anthocyanins. Buffers systems in aqueous solutions are important to control for potential reduction in pH by addition of the metallic cations; however, some commonly used buffering agents such as phosphates and citrates are also known to chelate metal ions. In solutions with anthocyanins, these pH buffering compounds were found to preferentially bind metal ions or even sequester the ions from already formed anthocyanin-metal complexes leaving the pigments in their flavylium forms (Buchweitz,

Carle, et al., 2012).

Ferric iron chelates of some food safe anthocyanins have also been investigated as blue coloring agents in polysaccharide and gelatin based gels. Anthocyanin sources included various cyanidin derivatives from elderberry (nonacylated), purple carrot

(acylated), and red cabbage (acylated). Gentian blue hues were developed with ferric iron chelation by red cabbage anthocyanins; however, cobalt and more violet blue hues were obtained with purple carrot and elderberry anthocyanins (Buchweitz, Brauch, Carle, &

Kammerer, 2013a). The gelling agents were also found to impact blue color expression of the chelates; generally the polysaccharide blend (agar-agar and amidated pectin) displayed more red like qualities than the gelatin based systems. In addition to the more blue like qualities of the gelatin systems, they also showed high color stability

(Buchweitz et al., 2013a). The stability of the colorant also varied depending on the chemical composition of the anthocyanins; the highest stability was found for the purple carrot based colorants, which contain some acylated anthocyanins (Buchweitz et al.,

2013a). The inclusion of dairy in these gels was found to negatively impact blue color formation resulting primarily in pink and violet hues; this was likely effect of lactic acid

48 preferentially chelating metal ions instead of the anthocyanin (Buchweitz et al., 2013a).

The results of this study suggest application of anthocyanin metal chelates may prove successful as alternatives for synthetic blue food colorants in certain applications and matrices.

A specific subclass of metal coordinated anthocyanin complexes includes the metalloanthocyanins; these macromolecular complexes have been shown to play an important role in the blue coloration of multiple floral systems. Metalloanthocyanins are composed of fixed stoichiometric ratios of anthocyanins, flavones, and metallic cations of

6, 6, 2, respectively (Yoshida et al., 2009). Currently, 5 metalloanthocyanins have been identified and are responsible for violet and blue colors of different floral systems

(Yoshida et al., 2009). The same anthocyanin structural requirements are necessary to form metalloanthocyanins, bearing 2 neighboring hydroxyl groups on the B ring. The known metalloanthocyanins consist of either delphinidin, cyanidin, or petunidin and

Mg2+, Al3+, or Fe3+ as metallic cations (Yoshida et al., 2009). The 2 metal ions are centralized within the macromolecule and the anthocyanins and flavones are self- associated in a chiral, left-handed manner caused by the D-glucsoyl moieties (Yoshida et al., 2009). The anti-clockwise chiral stacking of these molecules produces a characteristic exiton type Cotton effect when evaluated by circular dichroism, Figure 2.13 provides a schematic of the organization of metalloanthocyanins (Yoshida, Kitahara, Ito, & Kondo,

2006). Metalloanthocyanins based on petunidin anthocyanins have not yet been found to produce blue colors, but those based on cyanidin and delphinidin do (Yoshida et al.,

2009). Cyanidin derivaed metalloanthocyanins require ferric iron ions as components to yield blue hues (Yoshida et al., 2009).

Figure 2.13: Schematic of the organization of the metalloanthocyanins, adapted from

Yoshida et al., 2009

One of the oldest known metalloanthocyanins, commelinin, is derived from the

Asiatic dayflower and was prized as blue pigment for printing and art (Yoshida et al.,

2009). The pigment has been shown to produce stable blue colors in very acidic pH as low 2.4 in concentrated solutions; however when diluted, the macromolecule easily dissociated and lost its blue hues (Takeda, Fujii, & Iida, 1984; Yoshida et al., 2009).

Stabilization of the pigment would very likely be necessary for food application as well as further evaluations to consider the pigment as an alternative to synthetic blue food colorants. The need to prove its safety and petition for regulatory approval would likely

50 also be required as the colorant may not necessarily be considered a fruit or vegetable juice.

Chapter 3: Bathochromic and Hyperchromic Effects of Aluminum Chelation by Anthocyanins from Edible Sources for Blue Color Development1

3.1 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 metallo-anthocyanins. 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 while 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.

______

1 Gregory T Sigurdson & M Mónica Giusti Department of Food Science and Technology, The Ohio State University. 2015 Fyffe Rd. Columbus, OH 43210 Published in the Journal of Agriculture and Food Chemistry 2014, 62:6955-6965

3.2 Keywords

Metallo-anthocyanin, 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.

3.3 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 (Potera, 2010). 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 (Potera, 2010). The

European Union has already enforced the use of warning labels indicating that synthetic colorants may cause hyperactivity in children, and the Food and Drug Administration

(FDA) has recommended the safety of synthetic dyes be reviewed (FDA; CFSAN (Food and Drug Administration; Center for Food Safety and Applied Nutrition)., 2011). 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

53 effects, making them viable alternatives for synthetic colorants. Although blue dyes are less used than other hues, there are currently few natural options (Potera, 2010). Those with commercial feasibility have been derived from blue gardenias, huito fruit, and

Spirulina spp., which was only approved for confectionary uses in September 2013

(Buchweitz, Carle, et al., 2012). Food application data of these colorants is relatively scarce (Buchweitz, Carle, et al., 2012). In acidic conditions common to food products, anthocyanins typically appear in red or purple molecular forms. However, self- association, co-pigmentation, and metal complexes can result in acid-stable, blue colorations, like those found in flowers (Yoshida et al., 2009). Most metal-anthocyanin interactions have been studied for better understanding of plant and floral pigmentation

(Buchweitz, Carle, et al., 2012).

For anthocyanins to undergo metal complexation, more than one free hydroxyl group must be present on the B ring (Bayer et al., 1966; Pyysalo & Kuusi, 1973; Yoshida et al., 2009). Figure 3.1 shows catechol and pyrogallol moieties on the B ring of cyanidin

54 and delphinidin, respectively. Multivalent metal ions act in competition of the hydrogen ions attached to these rings inducing their loss, transforming the flavylium cation to a quinoidal base (Fig. 3.1) (Bayer et al., 1966; Schreiber et al., 2010). Simultaneously, a stacking association with another anthocyanin flavylium ion occurs with the transformed molecule forming a metal coordinated complex (Schreiber et al., 2010). Various metal ions are known to induce this effect, all being multivalent (Bayer et al., 1966). Al3+ anthocyanin complexes in ethanol have been reported to be relatively stable in pH 2-5 in reactions with delphinidin (Schreiber et al., 2010). Cyanidin and delphinidin glucosides from berries had previously been found to develop blue like hues with tin chelation while iron chelates were more brown (Pyysalo & Kuusi, 1973). 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 (Buchweitz, Carle, et al., 2012). Unlike previous findings, ferric ion treated delphinidin-3-glucoside showed blue hues and further bathochromic shifts than cyanidin and petunidin glucosides. These samples also showed highest stability over time

(Buchweitz, Carle, et al., 2012). Environment also can affect 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 (Buchweitz, Carle, et al., 2012).

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 (Becaria, Campbell, & Bondy, 2002;

Nayak, 2002). However, no reports of dietary aluminum toxicity were found in literature in healthy individuals (Flaten, Alfrey, Birchall, & Yokel, 1996). Absorption from anthocyanin-aluminum colorants is likely low, based on anthocyanin chelating effects and aluminum’s low bioavailability in water (0.3%) (Flaten et al., 1996; Nayak, 2002).

The FDA considers aluminum sulfate as GRAS (generally recognized as safe) as a food additive with no usage limit, and aluminum salts may comprise 2% of the final product in synthetic lake type dyes (21 CFR 182.B.51, 21 CFR 182.1125).

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 to 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 pH range 3-6 to better understand its role in anthocyanin-metal chelation. Color stability was monitored by a 28 day study of delphinidin and acylated cyanidin chelates in different storage conditions.

3.4 Materials & Methods

3.4.1 Materials

American eggplant (Solanum melongena L), Japanese eggplant (Solanum melongena L), red raspberry (Rubus idaeus), and red cabbage (Brassica oleracea var.

56 capitata f. rubra) were purchased from a local grocery store in Columbus, OH, U.S.A.

Frozen whole black currants (Ribes nigrum) was purchased from CropPharms LLC.

(Staatsburg, NY, U.S.A.). A commercial black carrot (Daucus carota spp. sativus) anthocyanin powder. A commercial chokeberry (Aronia melanocarpa) juice concentrate were also obtained from Artemis International (Fort Wayne, IN, U.S.A.). Aluminum chloride hexahydrate, USP 97.0-101.0 % grade, was purchased and obtained from Sigma-

Aldrich Co. (St. Louis, MO, U.S.A.). ACS grade sodium acetate anhydrous, hydrochloric acid 6N (certified 5.95-6.05), trifluoroacetic acid, and sodium hydroxide N/10 (0.0995-

0.1005) were purchased from Fisher Scientific (Fair Lawn, NJ, U.S.A.) 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 number AL9925) and indigotine or

FD&C blue No. 2 (certification number AL2889), each had a color purity of 92%, were obtained from Noveon Hilton Davis, Inc. (Cincinnati, OH, U.S.A.). FD&C blue no. 1 was diluted to 2.14 × 10-4 M with distilled water while FD&C blue no. 2 was diluted to 6.35 ×

10-4 M.

3.4.2 Methods

3.4.2.1 Extraction and Purification of Anthocyanins

Extraction of anthocyanins from plant materials followed procedures described by

Rodríguez-Saona & Wrolstad, 2001. 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

57 acetone to reduce the brown discoloration, likely due to polyphenoloxidase, that occurred during extraction. After mixing the filtrate with 1-2 volumes of chloroform, phase separation occurred overnight at 4 °C, except eggplant samples being limited to 4 hours.

The chloroform layer containing lipophilic compounds was appropriately discarded; knowing 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.

3.4.2.2 Anthocyanin Purification – Solid Phase Extraction

Crude anthocyanin extracts were purified by loading activated Waters Sep-pak®

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.

3.4.2.3 Monomeric Anthocyanin Quantitation

Monomeric anthocyanins were quantitated by the pH differential method, described by Giusti & Wrolstad, 2001. Briefly, anthocyanins were quantified by measuring the absorbance at 520 nm and 700 nm at pH 1 and pH 4.5 for each sample.

Based on the difference of these absorbances, the concentration of monomeric anthocyanins was determined to use for dilution to known concentrations.

Table 3.1: Display of anthocyanin, associated molecular weight, and molar absorptivity used for monomeric quantitation of anthocyanins in food samples evaluated

Molecular Molar Sample Anthocyanin Weight Absorptivity American delphinidin-3-rutinoside 647.0 26,900a eggplant Japanese delphinidin-3-(p-coumaroylrutinoside)-5- 955.26 26,900a eggplant glucoside Black cyanidin-3-glucoside 484.8 26,900a currant Red cyanidin-3-glucorutinoside 757.0 26,900a raspberry Chokeberry cyanidin-3-glucoside 484.8 26,900a Red cyanidin-3- diglucoside-5-glucoside 773.0 30,175 cabbage Black cyanidin-3- 919.25 26,900a carrot xylosyl(feruloylglucosyl)galactoside a Molar absorptivity of cyanidin-3-glucoside, used for monomeric quantitation

(Giusti & Wrolstad, 2001)

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 molar absorptivity. Table 3.1 shows the values used for each sample’s quantitation.

3.4.2.4 High Pressure Liquid Chromatography (HPLC) Evaluation of Anthocyanins

Reverse phase HPLC was utilized to identify anthocyanins, with comparison to literature. Samples were analyzed using a HPLC (Shimadzu, Columbia, Maryland,

U.S.A.) system equipped with LC-20AD pumps and a SIL-20AC autosampler coupled to a LCMS-2010 Mass Spectrometer (Shimadzu, Columbia, Maryland, U.S.A.) and a SPD-

M20A Photodiode Array (Shimadzu, Columbia, Maryland, U.S.A.) detectors. LCMS

Solution Software (Version 3, Shimadzu, Columbia, Maryland, U.S.A.) was used to view results. Separation of anthocyanins was achieved on a reverse-phase Symmetry C18 column with 5 micron particle size and 4.6 x 150 mm column size (Waters Corp.

Taunton, MA, U.S.A. or Wexford, Ireland) and a 4.6 x 22 mm Symmetry 2 micro guard column (Waters Corp. MA, U.S.A.) was used. Samples were filtered through

Phenomenex® PhenexTM RC 0.2 µm, 15 mm membrane syringe filter (Torrance, CA,

U.S.A.). 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 LCMS grade water and B: LCMS acetonitrile (Fisher Scientific Inc, Fair lawn, NJ, U.S.A.). 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 to 70 min. Spectral data was 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 ion-tunnel mass spectrometer equipped with electrospray ionization (ESI) interface (Shimadzu, Columbia,

Maryland, U.S.A.). Mass spectrometry was performed under positive ion mode. Data was 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).

3.4.2.5 Evaluating Anthocyanin Structures, Concentrations, and Salt Ratios in Al3+

Complex Formation

To solutions of acidified water at pH 3 anthocyanin samples were added at three concentrations, based on monomeric quantitation: 25 μM, 50 μM, 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 M, 0.2288 M, and 0.4576 M. Salt solutions were added to anthocyanin solutions beginning at equal M 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.

3.4.2.6 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 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 conditions: dark storage at refrigeration temperatures (2-4 °C), dark storage at ambient temperatures (18-25 °C), and full light exposure from natural daylight ~8 hours and fluorescent lamps from above at ambient temperatures (18-25 °C).

3.4.2.7 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).

3.4.2.8 Spectrophotometry of Solutions with UV-Visible Transmission

Each sample from each phase of study was subjected to visible spectrophotometry and colorimetry. After 15 min 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 (Shimadzu, Columbia, Maryland,

U.S.A.). Spectrograms were generated using UV Probe software, version 2.21

(Shimadzu, Columbia, Maryland, U.S.A.) associated with this spectrophotometer model.

3.4.2.9 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 Labs, Reston, VA,

U.S.A.). The equipment was set for total transmittance, illuminant D65, and a 10° observer angle for all liquid samples.

3.4.2.10 Statistical Evaluation of Data

The mean and standard deviation of data replicates was calculated using

Microsoft Office Excel 2010 (Office 14.0, Microsoft. Redmond, WA, U.S.A.). Data from

62 evaluation of anthocyanin structures, concentrations, and salt ratios in Al3+ complex formation was conducted by 1-way analysis of variance (ANOVA) (2-tailed, α = 0.05) and Student’s paired t-test (2-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 were 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 1-way ANOVA (2-tailed, α = 0.05). All other figures were also generated using this software.

3.5 Results and Discussion

3.5.1 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 Fig. 3.2. The American eggplant showed delphinidin-3-rutinoside as the primary anthocyanin as indicated from its HPLC chromatogram, mass spectrometry, and by comparison with literature (Azuma et al., 2008). Black currant chromatograms showed profiles similar to previous findings; delphinidin-3-rutinoside and cyanidin-3-rutinoside, each representing ~40% of the anthocyanin profile (Slimestad & Solheim, 2002). Japanese and non-Japanese type eggplants share delphinidin as the primary anthocyanidin; however, the primary Japanese

63 eggplant anthocyanin was acylated, delphinidin-3-(p-coumaroylrutinoside)-5-glucoside

(or nasunin) (Fig. 3.2) (Azuma et al., 2008).

Figure 3.2: Reverse phase HPLC chromatograms of food sample with detection at 520 nm showings main pigments identified 64

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 cyanidin-3- galactoside, representing ~65% of the anthocyanin profile (Oszmianski & Sapis, 1988).

The chromatogram of red raspberries of this study showed 4 major pigments; most prevalent were cyanidin-3-sopohoroside and cyanidin-3-glucorutinoside (Wrolstad &

Boyles, 1993). 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-3-sopohoroside-5-glucoside (McDougall,

Fyffe, Dobson, & Stewart, 2007). Black carrot chromatograms showed 5 major peaks, resembling the profile of the “Deep Purple” variety (Montilla, Arzaba, Hillebrand, &

Winterhalter, 2011). The major pigment of this variety was cyanidin-3- xylosyl(feruloylglucosyl)galactoside, agreeing with literature (Montilla et al., 2011).

3.5.2 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). It 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, or acylation. Each anthocyanin demonstrated

65 both bathochromic and hyperchromic shifts when exposed to the metal ion. Comparing core structures, delphinidin containing samples showed furthest bathochromic shifts and become most blue like at pH 3 (Table 3.2). Glycosylated delphinidin showed λmax bathochromic shifts of 50 nm and 55 nm for American eggplant and black currant, respectively. Cyanidin samples showed lower bathochromic shifts ranging 26-39 nm.

This data indicates that increasing the number of free hydroxyl groups on the aglycone core structure leads to further bathochromic shifts, agreeing with previous studies comparing petunidin, cyanidin, and delphinidin (Buchweitz, Carle, et al., 2012).

Table 3.2: Effects of anthocyanin concentration at pH 3 on color shifts, with λmax and

CIE-Lab color characteristics of anthocyanins (ACN) complexed with AlCl3 100× anthocyanin concentration. In parenthesis () are the standard deviations, n=3.

Color Characteristics [ACN] Sample Δ (μM) L a* b* chroma hue hue American 25 89.3 (1.2) 2.7 (0.4) -5.8 (0.5) 6.3 (0.7) 295.2 (1.5) 279 50 86.1 (0.8) 3.6 (1.2) -10.1 (3.2) 10.7 (3.4) 289.4 (0.5) 284 eggplant 100 80.3 (1.5) 4.4 (1.6) -10.6 (2.1) 11.5 (2.5) 291.9 (4.0) 264 Black 25 89.9 (1.5) 5.0 (0.7) -5.0 (0.9) 7.1 (0.8) 314.7 (7.0) 299 50 81.1 (1.7) 11.1 (1.4 ) -13.3 (3.1) 17.4 (3.2) 310.2 (4.1) 293 currant 100 71.7 (1.0) 17.3 (0.9) -19.3 (1.0) 25.9 (1.5) 311.9 (0.0) 293 Red 25 89.8 (0.1) 7.9 (1.0) -5.0 (0.3) 9.2 (1.2) 329.1 (0.9) 320 50 84.6 (2.6) 14.1 (2.4) -9.7 (1.9) 17.1 (3.1) 325.6 (1.2) 317 raspberry 100 74.3 (0.8) 27.0 (1.6) -16.1 (1.7) 31.3 (2.2) 329.3 (1.1) 318 25 88.1 (1.0) 9.0 (0.6) -4.6 (0.7) 10.2 (0.8) 332.8 (2.8) 318 Chokeberry 50 83.3 (1.4) 13.2 (1.0) -8.6 (1.5) 15.8 (1.7) 327.1 (2.5) 311 100 73.5 (0.3) 22.7 (0.9) -15.2 (0.5) 25.6 (1.4) 325.8 (0.1) 310 25 89.1 (0.9) 4.8 (0.9) -7.7 (0.8) 9.1 (0.9) 301.6 (4.7) -44 Red cabbage 50 81.0 (0.6) 9.0 (1.9) -16.9 (1.2) 19.2 (1.9) 297.8 (3.4) -47 100 70.1 (3.6) 15.1 (3.5) -28.2 (4.0) 30.4 (5.4) 297.9 (2.1) -48 Purple 25 90.2 (0.8) 6.8 (0.2) -5.0 (0.6) 8.4 (0.3) 323.6 (4.0) -36 50 83.7 (0.8) 12.5 (0.1) -11.9 (1.0) 17.3 (0.7) 316.4 (2.6) -42 carrot 100 72.8 (0.3) 22.6 (0.9) -22.4 (0.9) 29.0 (0.4) 315.2 (2.1) -45

Amount and type of common glycosylation does not seem to affect anthocyanin ability to chelate with metal ions but may mildly affect final solution color. When comparing the two glycosylated cyanidin samples, red raspberry anthocyanins demonstrated lower λmax than chokeberry anthocyanins until achieving high levels of

3+ Al ; Table 3.2 shows λmax of red raspberry anthocyanins to be about 10 nm less than 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 3 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 (Giusti & Wrolstad, 2003). 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

3.2). Red cabbage anthocyanins also exhibited more blue-like colors, having a hue angle around ~298; while black carrot anthocyanin Al3+ chelates exhibited hue angles ~316.

Based on this data and the overall longer sugar group chain glycosylation of black carrot anthocyanins suggests the molecular stacking arrangement of acylated black carrot chelates may not overlap as much as red cabbage chelates thus showing lower λmax and less blue hues.

Figure 3.3: Visible absorbance (400-700 nm) of 25, 50, & 100 μM concentrations of

American eggplant anthocyanins treated with factorial increases of AlCl3 (0-1000×) over anthocyanin concentration at pH 3

Regardless of anthocyanin concentration (25-100 µM), all samples chelated Al3+ showing bathochromic and hyperchromic shifts. However, the effects of increasing salt 68 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 (Figs. 3.3 and 3.4), until reaching optimum salt ratios where the same λmax and hue angle 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 1-way ANOVA, indicating mean λmax from each AlCl3 treatment to be different than anthocyanins from the same sample.

Figure 3.4: Visible absorbance (400-700 nm) of 25, 50, & 100 μM concentrations of red cabbage anthocyanins treated with factorial increases of AlCl3 (0-1000×) over anthocyanin concentration at pH 3

Similar results were obtained from 1-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 69 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 (Slimestad & Solheim, 2002). The hue reflects this anthocyanin profile, falling between the complex hues of either primarily delphinidin or cyanidin.

Regarding the atypically high absorbance increases of black currant anthocyanins, responsibility could be attributed to intermolecular copigmentation between the 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 concentrations were found to be between 100-500× anthocyanin concentration, where treated anthocyanins showed the highest intensity of absorbance as well as reaching the furthest bathochromic shift, Figs.

3.3 and 3.4. 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 for these 2 salt concentration ratios on American eggplant, black currant, and red cabbage anthocyanins were found not to differ having p-values greater than 0.06. The other samples were found not to differ with AlCl3 concentrations 500× and

1000× anthocyanin content. Same results from t-tests were obtained for 50 µM anthocyanin samples. For 100 µM anthocyanin samples, only chokeberry and red

70 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. Large excess of salt seemed to initiate some reversion of the color intensification achieved at lower ratios (Figs. 3.3 and 3.4); 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 or less, except 25 µM red raspberry anthocyanin samples, which was still the same at 90%. From Student’s t-test, p-values of 0.03 or less were found for all 50 µM 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. Based on the largest absorbance and λmax observed in these samples, optimal AlCl3 concentration was found to be 100-500× anthocyanin concentration.

3.5.3 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 et al., 2008). However, all three

71 of the proposed pathways involve transformation of the flavylium cation to other forms more prevalent in higher pH which then decompose into degradation products (Schwartz et al., 2008). It has been proposed that chelation of metals by anthocyanins stabilizes the quinonoidal base form as well as protect it from nucleophilic attack by complexation coordination (Bayer et al., 1966; Buchweitz, Carle, et al., 2012). Metal chelation by anthocyanins has been shown to increase stability during storage as well as with thermal treatment (Buchweitz, Carle, et al., 2012; Tachibana, Kimura, & Ohno, 2014). Ferric chelates of cyanidin-3-glucoside showed improved stability to thermal treatment of 60 °C over 80 minutes, with intensity retention of ~70% (Tachibana et al., 2014). 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 (Buchweitz, Carle, et al., 2012).

Figure 3.5: 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)

This study compared the stability of aluminum chelates of delphinidin-3- rutinoside (American eggplant) to that of acylated cyanidin glycosides (red cabbage), as literature is limited about stability of acylated anthocyanin metal chelates. Highest stability for both samples was found in cold, dark storage as expected based on ideal conditions for anthocyanin stability. For delphinidin chelates absorbance decreases were

73 limited to ~0.1 over 28 days while almost no change was noted for acylated cyanidin samples (Fig. 3.5). Much larger decreases in absorbance were noted for samples stored in ambient temperatures. Although those samples receiving light treatment showed slightly larger decreases; 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 (Buchweitz, Carle, et al., 2012). Acylated cyanidin chelates showed more linear degradation patterns (Fig. 3.5). These samples also showed improved color stability, exhibiting lower changes in absorbance over time, likely result of 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 decrease of absorbance noted with high excesses of AlCl3, large amount of salt also was found to diminish stability. From 1-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 based on 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.

3.5.4 Evaluating Effects of pH in Al3+ Complex Formation

Figure 3.6: pH change of 50 μM anthocyanins sample solutions initiated by treatment with AlCl3 (0-2000×) in 1 M sodium acetate buffers pH 3, 4, 5, & 6

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 starting each pH levels tested (pH 3-6), all anthocyanins exhibited roughly equal decreases in pH in 1 M acetate buffer solutions (Fig. 3.6). By 1-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

75 from both aglycones. It is suggested that acidic pH may not limit ability of these anthocyanins to chelate metals, despite the form 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 exceeding AlCl3 salt concentrations 100× anthocyanin content (Fig. 3.6), so color observations were due to complexation rather pH changes.

Even in 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 compete (Buchweitz, Carle, et al., 2012). At low pH where anthocyanins tend to appear in mostly red flavylium structures, those blue anthocyanin-metal complexes compete giving final solution a purple color. Conversely, in more neutral pH where anthocyanins appear in colorless carbinol or yellow chalcone forms, the ultimate solution color is blue or blue-green with metal complex competition.

Figure 3.7: 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

All samples, pH 3-6, followed the same general trend for Al3+ chelation depicted

(Fig. 3.7) 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 77 shifts at low pH, until high levels of AlCl3 2000× anthocyanin concentration. The reverse was noted when beginning at 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 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 non-

3+ acylated counterparts (Table 3.3). Despite similar λmax of Al complexed anthocyanins, hue angles differed with changes in pH, shown in Tab1e 3.3.

Table 3.3: Effects of acidic pH change on color shifts, λmax and CIE-Lab color characteristics of 50 µM anthocyanins (0×) and a complexed with AlCl3 100× anthocyanin concentration. (standard deviations), n=3. values estimated due to low absorbance pH λmax (nm) L a* b* chroma hue Sample 0× 100× 0× 100× 0× 100× 0× 100× 0× 100× 0× 100× 0× 100× 3.08 2.99 523 568 92.5 88.0 6.5 4.6 0.9 -7.1 6.6 8.5 7.9 302.8 (0.11) (0.10) (0) (1) (0.1) (0.3) (0.4) (0.3) (0.1) (0.8) (0.4) (0.8) (1.1) (1.6) 4.00 3.96 523 575 94.1 87.7 2.4 1.7 1.2 -8.2 2.7 8.3 25.7 281.7 American (0.04) (0.05) (4) (1) (0.2) (0.1) (0.0) (0.0) (0.1) (0.5) (0.1) (0.5) (1.9) (0.6) eggplant 5.01 4.97 509a 578 94.5 88.2 0.9 0.2 1.2 -6.2 1.6 6.2 52.7 272.0 (0.01) (0.02) (2) (0.4) (0.1) (0.1) (0.1) (0.2) (0.2) (0.2) (0.2) (1.6) (0.6) 6.00 5.79 509a 579 93.9 90.0 0.4 -0.2 1.2 -2.2 1.3 2.2 73.2 263.9 (0.03) (0.02) (3) (0.6) (0.8) (0.6) (0.1) (0.2) (0.7) (0.4) (0.7) (2.5) (6.5) 3.07 2.98 527 569 92.7 87.6 6.5 3.5 -0.7 -8.3 6.5 9.0 353.6 292.7 (0.10) (0.10) (0) (1) (0.1) (0.4) (0.6) (0.2) (0.4) (1.0) (0.6) (1.0) (2.5) (1.1) 4.01 3.97 529 578 93.9 87.2 2.0 -0.7 0.5 -7.9 2.1 8.0 8.1 264.8 Japanese (0.05) (0.05) (1) (1) (0.3) (0.3) (0.1) (0.1) (0.2) (0.5) (0.1) (0.5) (16.4) (0.6) a 78 eggplant 5.01 4.97 534 581 94.2 87.5 0.5 -2.6 1.1 -6.3 1.2 6.8 63.5 247.2

(0.03) (0.03) (2) (0.3) (1.0) (0.1) (0.3) (0.1) (0.9) (0.1) (1.0) (2.5) (1.1) 6.03 5.80 540a 587 93.2 88.3 -0.6 -2.9 1.4 -2.2 1.6 3.6 115.3 216.2 (0.02) (0.01) (1) (0.6) (1.2) (0.2) (0.5) (0.3) (1.0) (0.2) (0.9) (10.6) (9.8) 3.07 2.98 513 534 92.1 89.0 8.3 9.3 1.8 -3.4 8.5 9.9 12.6 339.9 (0.09) (0.10) (1) (0) (0.6) (1.0) (1.0) (1.1) (0.3) (0.4) (1.0) (1.2) (0.5) (1.0) 4.01 3.97 511 558 93.8 88.6 3.6 5.9 1.1 -6.0 3.8 8.4 17.6 314.3 (0.05) (0.05) (2) (1) (0.3) (0.2) (0.0) (0.1) (0.1) (0.2) (0.0) (0.2) (1.3) (0.7) Chokeberry 5.01 4.97 508a 564 94.3 88.9 1.8 3.3 0.8 -4.7 2.0 5.7 25.1 305.6 (0.02) (0.03) (1) (0.2) (0.2) (0.0) (0.0) (0.2) (0.1) (0.1) (1.0) (4.8) (0.5) 6.04 5.81 528a 568 93.8 89.6 1.0 1.4 0.5 -2.1 1.1 2.5 25.9 302.9 (0.02) (0.02) (1) (0.2) (0.3) (0.1) (0.1) (0.2) (0.2) (0.2) (0.3) (7.2) (1.4) 3.07 2.98 532 551 90.7 87.7 11.6 7.6 -3.3 -8.8 12.1 11.6 344.0 310.9 (0.10) (0.10) (1) (0) (0.4) (0.3) (0.3) (0.3) (0.1) (0.1) (0.3) (0.1) (0.4) (1.2) 4.01 3.97 539 572 91.7 87.1 7.0 2.8 -3.3 -10.6 7.8 10.9 334.5 284.6 Red (0.05) (0.05) (1) (0) (0.2) (0.2) (0.0) (0.0) (0.1) (0.2) (0.0) (0.2) (1.0) (0.5) cabbage 5.01 4.97 549 579 92.5 87.5 3.2 0.5 -3.0 -9.9 4.4 9.9 317.2 272.7 (0.03) (0.03) (2) (1) (0.2) (0.0) (0.1) (0.0) (0.2) (0.3) (0.1) (0.3) (2.4) (0.1) 6.04 5.81 555 589 92.3 88.4 0.1 -1.3 -3.3 -8.3 3.3 8.4 271.8 260.9 (0.02) (0.02) (2) (2) (0.5) (0.2) (0.0) (0.0) (0.3) (0.2) (0.3) (0.2) (0.7) (0.2) 79

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

3.3). Other samples showed more purple-pink hues. Despite delphinidin having the pyrogallol moiety and exhibiting furthest 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 though hue angles differed by about 10.

When increasing 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 most pink; while acylated cyanidin from red cabbage exhibited similar hues to 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

3+ blue color upon complexing with Al . Hue of this sample treated with AlCl3 100× anthocyanin concentration was 265 and remained constant until reaching AlCl3 1,000× and 2,000× becoming 269 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 230-263. Optimal pH for delphinidin-3-glucoside-ferric chelates was found to be 4.5 based on formation and stabilization of the chelates (Buchweitz,

Carle, et al., 2012). Generally absorbance of complexed solutions decreased as pH increased, except comparing pH 3 and 4 where absorbance increased by about 0.1 except chokeberry anthocyanins which was lower. Decreases in absorbance were highest comparing pH 5 and pH 6, ranging 0.9-0.25. From color and intensity evaluation, ideal pH for blue color formation of anthocyanin-Al chelates likely lies in pH 4-5.

Similar trends were noted with starting pH 6; the highest amount of blue hues were 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 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

81 colorant FD&C blue No. 1 expressed a λmax of 630 while blue No. 2 showed λmax of 610 nm, each of which easily correlated to their observed blue colors, in agreement with previous findings (Ahmadiani, 2012). Despite the fact that these metallo-anthocyanin complexes had much lower λmax, some were able to produce similar blue hues.

Unlike the findings from previous studies of anthocyanin-ferric chelates, precipitation of metallo-anthocyanins was not observed until reaching neutral or basic pH

(Buchweitz, Carle, et al., 2012). At pH 5, ferric anthocyanins were found to precipitate within 2 hours of ion introduction leaving the supernatant colorless after 24 hours

(Buchweitz, Carle, et al., 2012). Data was 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.

3.6 Conclusion

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 was 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 hues at

82 pH ≥ 2.5 when complexation occurred at pH 4. Non-acylated 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.

Chapter 4: Spectral and Colorimetric Characteristics of Metal Chelates of Acylated

Cyanidin Derivatives

4.1 Abstract

Colorants derived from nature are increasingly popular due to consumer demand.

Anthocyanins are a class of naturally occurring pigments that produce red-purple-blue hues in nature, especially when interacting with metal ions and co-pigments. The role of various acylations of cyanidin (Cy) derivatives on color expression and stability of Al3+ and Fe3+ chelates in pH 6-7 were evaluated by spectrophotometry (380-700 nm) and colorimetry (CIE-L*a*b*) during dark, ambient storage (48 hr). Increased substitution generally increased λmax of Cy chelates: malonic acid monoacylation < triglycosylated Cy

< Cy monoacylated with hydroxycinnamic acids < diacylated Cy. Patterns were similar regarding bathochromic shifts. Acyl moieties of diacylated Cy with smaller substitution patterns resulted in greater λmax, and no pattern emerged for monoacylated cyanidin.

Pigment stability was improved with increasing proportions of metal ions and acylation.

Stability followed that diacylated cyanidin (p-coumaric-sinapic > ferulic-sinapic > sinapic-sinapic) > monoacylated (malonic ≈ sinapic > ferulic > p-coumaric).

4.2 Keywords

Anthocyanin, anthocyanin-metal chelate, metalloanthocyanin, intramolecular copigmentation, Brassica oleracea var. capitata f. rubra, Zea mays L.

4.3 Introduction

Color additives in food products play an important role in the success of many products. Current regulations allow for use of color additives to enhance and correct already present colors, standardize colors of raw materials, create a color identity to otherwise colorless foods, such as sodas or confections, and also account for color loss during storage or processing (Newsome, Culver, & Breemen, 2014; Potera, 2010).

Consumer demand and possible health concerns are driving the replacement of synthetic food colorants with naturally derived alternatives (Potera, 2010; Sigurdson & Giusti,

2014). Naturally derived options for red and yellow colorants, from pigment classes such as betalains, carotenoids, and anthocyanins (ACN), are considered more prevalent and simpler to reproduce than blue shades (Newsome et al., 2014). ACN, phycocyanin, and iridoid derivatives are pigments sourced from nature with potential as blue food colorants

(Buchweitz, Carle, et al., 2012). Limitations of their use in foods include regulatory restrictions, limited stability or inability to match the hues of currently used synthetic dyes (Buchweitz, Carle, et al., 2012; Newsome et al., 2014; Sigurdson & Giusti, 2014).

Anthocyanins (ACN) are flavonoid compounds responsible for many red, purple, and blue colors of fruits, vegetables, and flowers that undergo structural changes depending on environment and substitution pattern. Deprotonation occurs with increase in pH, resulting in the formation of purple-blue quinonoidal forms (Newsome et al.,

2014). In the acidic conditions of most food products, ACN exist in red-purple structural forms (Sigurdson & Giusti, 2014). Mechanisms like self-association, co-pigmentation,

85 and metal chelation can result in blue colorations in acidic conditions, found in some flowers (Yoshida et al., 2009).

Currently, co-pigmentation is thought to be one of the most efficient color stabilization processes of anthocyanins in plant organs, consisting of the interaction of the

ACN chromophore with essentially colorless co-pigmenting molecules (Dangles et al.,

1993b). The mechanism of co-pigmentation is believed to be based on hydrophobic interactions or Π-Π stacking between the planar ACN chromophore and other phenolic compounds (Di Meo et al., 2012). Intermolecular co-pigmentation occurs between separate molecules. Intramolecular co-pigmentation occurs within the same molecule between the ACN chromophore and acylating attachments. It has been that with increasing degree of acylation of the same chemical structure, increasing bathochromic shifts were observed (Dangles et al., 1993a). Intramolecular co-pigmentation is thought to help to produce more blue hued colors at lower pH than ACN lacking acylation, such as the acylated delphinidin (Dp) derivatives found in the butterfly pea flower (Abdullah et al., 2010).

Other mechanisms for blue color development by ACN include metal chelation, which occurs with aglycones bearing vicinal dihydroxy groups on the B ring (Bayer,

Egeter, Fink, Nether, & Wegmann, 1966; Buchweitz et al., 2012; Schreiber et al., 2010).

ACN-metal complexes are stabilized by intramolecular co-pigmentation by acylation, displacing ACN hydration equilibria (Elhabiri, Figueiredo, Toki, Saito, & Brouillard,

1997). Metal ions induce deprotonation of the ACN B-ring, transforming flavylium cations to the blue quinonoidal bases at lower pH’s (Schreiber et al., 2010). Acylated

ACN, from red cabbage or purple carrot extracts, were found to express larger λmax (most blue color) with metal chelation than non-acylated ACN (Buchweitz et al., 2013a;

Buchweitz, Brauch, Carle, & Kammerer, 2013b; Sigurdson & Giusti, 2014). The ACN of red cabbage and purple carrot are predominantly derivatives of the same chromophore

(cyanidin) bearing acylation of similar hydroxycinnamic acid derivatives but differ in the sugar moieties to which they are attached, therefore changing the spatial geometry of the molecule and effect of co-pigmentation (Li et al., 2012).

In addition to affecting the ACN color expression, metal chelation has also been found to increase the stability of the pigments during storage or with heat treatment, following first order degradation kinetics (Sigurdson & Giusti, 2014; Tachibana,

Kimura, & Ohno, 2014, Buchweitz et al., 2013a, 2013b). Stability of the chelates was found to be affected by several factors including ACN structure, pH, buffer system, metal ion, and ratio of ACN:metal ion (Buchweitz et al., 2013a, 2013b; Buchweitz, Carle, et al.,

2012; Buchweitz, Nagel, et al., 2012; Sigurdson & Giusti, 2014; Tachibana et al., 2014).

Rate of degradation generally decreased with increasing metal contents but lost efficiency with large excesses (Sigurdson & Giusti, 2014), and certain metal ions can promote ACN degradation (Kuusi, Pyysalo, & Pippuri, 1977; Pyysalo & Kuusi, 1973). Chelates of ACN extracts containing acylated derivatives generally showed higher stability than those extracts lacking acylated ACN (Buchweitz et al., 2013b).

Knowing differing patterns of acylation affect the color expression and stability of

ACN, it can be hypothesized that the structure and amount of acylation plays a role in color evolution and stability of ACN-M+ chelates. The objective of this study was to

87 systematically evaluate the role of the degree and structure of acylation, as intramolecular co-pigments, on the spectral responses and stability of ACN-M+ complexes. The findings of this study will provide additional insight on the mechanisms of ACN stabilization while expanding the blue color expression range of ACN based pigments.

4.4 Materials & Methods

4.4.1 Materials

Anthocyanin extracts were derived from fresh red cabbage (Brassica oleracea var. capitata f. rubra) purchased from a local grocery store in Columbus, OH or from purple corn powder (Zea mays L.) (AgroIndustrial, Peru). The predominant ACN of both plant materials were Cy derivatives. However, red cabbage ACN shared the same glycosylation pattern with varying degrees of aromatic acid acylation (Ahmadiani,

Robbins, Collins, & Giusti, 2014). The ACN isolated from purple corn exhibited aliphatic acylation.

Lab grade aluminum sulfate hydrate and reagent grade ferric chloride hexahydrate were purchased from Fisher Scientific (Fair Lawn, NJ).

Tris(hydroxymethyl)aminomethane, 99%) was obtained from Alfa Aesar (Ward Hill,

MA). ACS grade sodium acetate anhydrous, hydrochloric acid (certified ACS Plus), and sodium hydroxide N/10 (0.0995-0.1005) were purchased from Fisher Scientific (Fair

Lawn, NJ) as were all other standard ACS and HPLC grade reagents.

4.4.2 Methods

4.4.2.1 Anthocyanin Extraction

Extraction of ACN from plant materials followed procedures described by

Rodriguez-Saona and Wrolstad with acidified acetone and isolation by phase partition with water and chloroform (Rodríguez-Saona & Wrolstad, 2001).

4.4.2.2 Anthocyanin Purification – Solid Phase Extraction

Crude ACN extracts were purified by loading onto activated Waters Sep-pak®

4.4.2.3 Anthocyanin Isolation – High Pressure Liquid Chromatography (HPLC)

The Cy derivatives were isolated by separation with semi-preparative HPLC with manual collection of desired peaks. Reverse phase HPLC was conducted with a HPLC system (Shimadzu, Columbia, Maryland, U.S.A.) equipped with LC-6AD pumps, CBM-

20A communication module, SIL-20A HT autosampler, CTO-20A column oven, and

SPD-M20A Photodiode Array detector. LCMS Solution Software (Version 3, Shimadzu,

Columbia, Maryland, U.S.A.) was used to analyze results.

Separation of anthocyanins was achieved on a Luna reverse-phase pentafluorophenyl (PFP2) column with 5 µm particle size and 100Å pore size in 250 x

21.2 mm column size (Phenomenex®, Torrance, CA, U.S.A. Prior to injection, samples

89 were filtered through Phenomenex® PhenexTM RC 0.45 µm, 15 mm membrane syringe filter (Torrance, CA, U.S.A.).

Flow rate was set to 10.0 mL/min with a run time of 30 min. A binary gradient was used with solvents A: 4.5% formic acid in HPLC grade water and B: HPLC acetonitrile. Gradient began at 15% B and increased to 30% B from 0-30 min. Elution of

ACN was monitored at 520 nm and desired peaks were manually collected. Residual acetonitrile was removed from eluted peaks in a rotary evaporator at 37 °C under vacuum. The isolated pigments were again subjected to solid phase extraction, as described above without the ethyl acetate washes, to remove high concentrations of formic acid from respective solutions. Pigments were stored in acidified water until further analysis.

4.4.2.4 Isolated Anthocyanin Purity

The ACN isolates were verified for purity by reverse phase HPLC conducted with a similar system (Shimadzu, Columbia, Maryland, U.S.A.), differing only by pumps: LC-

20AD. An analytical column was used in this phase: Kinetix reverse-phase pentafluorophenyl (PFP2) column with 2.6 µm particle size and 100Å pore size in 100 x

4.6 mm column size (Phenomenex®, Torrance, CA, U.S.A. Flow rate was set to 0.6 mL/min with a run time of 50 min. A binary gradient was used with solvents A: 4.5% formic acid in HPLC grade water and B: HPLC acetonitrile. Gradient began at 0% B, increased 0-10% from 0-1 min, 10-30% B from 1-46 min, and 30-40% B from 46-50 min. Spectral data was monitored 270-700 nm with elution of ACN monitored at 520 nm.

ACN were isolated to a purity described as ≥ 88% of the total absorbance when

90 monitoring absorbance from 270-700 nm. Lowest level of purity was 88.4% for Cy-3-

(sinapoyl)-diglucoside-5-glucoside and was highest at 96.4% for Cy-3-(feruloyl)-

(sinapoyl)-diglucoside-5-glucoside.

4.4.2.5 Monomeric Anthocyanin Quantitation

ACN were quantitated by the pH differential method, described by Giusti and

Wrolstad (2001). Molar absorptivities of Cy derivatives from red cabbage were calculated by Ahmadiani, Robbins, Collins, & Giusti, 2015. The concentration of cyanidin-3-malonyl-glucoside (CGM), isolated from purple corn, was expressed as equivalents of cyanidin-3-glucoside. The concentration of monomeric ACN was calculated based on absorbance differences at their respective λmax and 700 nm in pH 1 and pH 4.5.

4.4.2.6 Sample Preparation

Isolated ACN were diluted to 50 µM concentrations in 0.5 M buffers of either sodium acetate for pH 6 or TRIS for pH 7, depending on pH. The pH of samples was monitored with a S220 SevenCompact™ pH/Ion meter (Mettler Toledo Inc., Columbus,

OH). Individual metal salts of Al3+ or Fe3+ were dissolved in distilled water to achieve concentrations of 0.6 M and 0.06 M. These salt solutions were then added to anthocyanin solutions beginning at equal M concentrations and then in factors to an excess of 0.5, 1,

5× [ACN]. Control samples without added salt were maintained at each pH. All samples were equilibrated for 45 min at room temperature in the dark prior to analysis. Three replicates were evaluated for each treatment.

Samples in pH 7 were also evaluated for stability due to predominance of all samples expressing blue hues. After equilibration, the samples were analyzed spectrophotometrically as a 0 hr time point. Samples were stored in sealed glass vials in the dark at 21-25 °C for 48 hours. Aliquots were removed from the vials for measurement at 4, 8, 24, and 48 hours. The ACN-metal chelates of this study were also found to follow

1st order degradation kinetics during the first 24 hours after which changes were negligible (Buchweitz et al., 2013a, 2013b). As absorbance at the λmax is correlated with observation of the desired color, linear regressions were prepared from the natural logarithm of the absorbance (at the λmax of the original time point) during the time points of the study, following the formula ln[At] = -kt + ln[Ao]. Linear regressions showed coefficient of determination values (R2 values) ranging 0.80 – 0.99, with ~80% being ≥

0.90. The half-life (t1/2) was calculated for each of the samples as t1/2 = ln2/k.

4.4.2.7 Spectrophotometry of Solutions by UV-Visible Transmission

After equilibration, 300 µL of each sample was evaluated by visible transmittance

(380-700 nm, 1 nm intervals) spectrophotometry in poly-D-lysine coated polystyrene 96 well plates using a SpectraMax 190 Microplate Reader (Molecular Devices, Sunnyvale,

CA).

4.4.2.8 Colorimetry of Solutions

The collected spectral data (380-700 nm, 5 nm intervals) was used to calculate colorimetric data, expressed in the CIE-L*a*b* communication system. The data was calculated using the CIE standard equations, D65 illuminant spectral distribution, and 10° observer angle functions. ((CIE), 2015; Kheng, 2002; Konica Minolta Inc., 2007)

4.4.2.9 Statistical Evaluation of Data

Figures and data means and standard deviations were produced using Microsoft

Office Excel 2010 (Office 14.0, Microsoft. Redmond, WA). The λmax, bathochromic shifts, and absorbance of the different metal ratio treated individual ACN were evaluated by 1-way analysis of variance (ANOVA) (2-tailed, α = 0.05) and Fisher’s least significant difference (α = 0.05) using Minitab 16 (Minitab Inc., State College, PA). For the specific

ACN-metal ratios, λmax, bathochromic shifts, and colorimetric characteristics (L*, a*, and b* values) of the different isolated ACN were compared by 1-way analysis of variance

(ANOVA) (2-tailed, α = 0.05) and Fisher’s least significant difference (α = 0.05).

Stability was evaluated by 1-way ANOVA (2-tailed, α = 0.05) and Fisher’s least significant difference (α = 0.05) comparing % Δ absorbance (at to λmax), reaction rate (k) from regression plots, and t½ of the different ACN-metal ratios of the same Cy derivative and also between the different isolated Cy receiving the same metal ratio treatment. The reaction rates (k) from regression plots of all Cy derivatives and all treatment ratios of the same metal were compared by two-way ANOVA (2-tailed, α = 0.05).

4.5 Results and Discussion

4.5.1 Spectral & Colorimetric Responses

The ACN of this study were derivatives of Cy aglycones. The Cy derivative from purple corn (CGM, cyanidin-3-malonyl-glucoside) was monoglucosylated at C3 while the ACN from red cabbage exhibited a different glycosylation pattern. These all shared the same glycosylation pattern bearing a glucoside attachment at C5 glucoside and a diglucoside attachment at C3 (CtG, cyanidin triglycosylated), Figure 4.1.

Figure 4.1: Identification (and abbreviations) of structure and substitution patterns of isolated ACN

The difference in these glycosylation patterns may have played a role in different color observation, as it has been found that increasing degree of glycosylation on the same chromophore induced bathochromic shifts (Stintzing, Stintzing, Carle, Frei, &

Wrolstad, 2002). The active chromophore of ACN molecules consists of the intact aglycone which exhibits a unique planar π-conjugated system. Structural distortions of the aglycone (such as stretching, bending, or torsion that could be induced by additional molecular substitution) modify π-delocalization of the system and affects the pigment’s interaction with light and therefore its color expression (Malcıoğlu, Calzolari, Gebauer, 94

Varsano, & Baroni, 2011). Glycosylation of the chomophore alters the absorbance spectra of the molecule due to constraining the dynamics of the molecule, a geometrical effect (Malcıoğlu et al., 2011). Acylation has generally been considered to impact color expression of ACN more strongly. The purple corn ACN bore aliphatic acylation

(cyanidin-3-malonyl-glucoside or CGM), and the red cabbage derivatives varied in type and degree of hydroxycinammic acid attachments, Figure 4.1. Aliphatic acylation

(malonic acid) on C3 of ACN has been found to participate in ACN-M+ chelation by aiding in deprotonation of the aglycone at C7 (Elhabiri et al., 1997).

Figure 4.2: λmax (nm) of Cy derivatives in pH 6 (left) or pH 7 (right), treated with factorial excess of Al3+ (0-5×) to [ACN] (50 µM), n = 3

The phenolic acid attachments of the Cy derivatives from red cabbage included p- coumaric acid, ferulic acid, and sinapic acid; in the diacylated derivatives, one attachment included sinapic acid while the other varied between the three aforementioned acids. The differences in the substitution patterns of the untreated ACN impacted the spectral and colorimetric properties of the pigments, as observed by the differences λmax of the

95 untreated Cy derivatives in Figure 4.2. Generally in the pH used for this study, the λmax of the aromatic diacylated Cy > aromatic monoacylated Cy > triglycosylated Cy >/≈

aliphatic monoacylated Cy in the pH levels used in this study. In pH 6, λmax of the ACN increased from 544 nm ± 6 (CGM) ≈ 541 ± 3 (CtG) < 548 ± CtGS < 551 ± 1 (CtGF) <

555 ± 1 (CtGSC) < 561 ± 0 (CtGC) < 562 ± 1 (CtGSF) << 591 ± 1 (CtGSS). No pattern on λmax was observed when comparing the monoacylated and diacylated counterparts sharing the same phenolic acids. For example, CtGSS (diacylated with sinapic acid) without metal ion treatment showed the largest λmax was; however CtGS (monoacylated with sinapic acid) exhibited the lowest λmax of the aromatically monoacylated ACN.

Interestingly, monoacylated CtGC showed higher λmax than diacylated CtGSC in pH 6 but this was not true in pH 7.

All ACN evaluated in this study responded to the presence of Al3+ or Fe3+, observable with changes in absorbance spectra, observed as increase in λmax in Figure 4.2, or changes in colorimetric values. In solution at pH 6, all ACN experienced bathochromic shifts induced by Al3+ or Fe3+. The magnitude of the shift was also found to depend on the substitution pattern of the ACN. Interestingly, CtG lacking acylation responded to metal chelation strongly and showed larger bathochromic shifts than monoacylated counterparts although the λmax of metal treated CtG was less than those acylated counterparts, agreeing with our preliminary work. CtGSS bearing 2 sinapoyl moieties showed the smallest shift in λmax, as observable in Table 4.1. CtGSC, diacylated with sinapic acid and p-coumaric acid, experienced the largest bathochromic response; while its monoacylated counterpart, CtGC with p-coumaric acid, demonstrated the

96 smallest response of the triglycosylated Cy derivatives. The large impact of metal ion chelation on CtGSC may be explained partially by co-pigmentation of the acyl moieties with the chromophore and possibly by allowing for additional ion chelation by the p- coumaroyl moiety. Coumaric acid lacks methoxyl substitutions on its phenyl group that ferulic and sinapic acids exhibit, and metallic cations may be more prone to bind to the hydroxyl group due to less steric hindrance. As CtGC showed small responses to metal chelation, it can be proposed that weaker interaction occurs between the chromophore and the acyl group compared to the more substituted ferulic or sinapic acids. This trend was similar to the effects of the acyl groups as intermolecular co-pigments, in which bathochromic shifts were found to be greatest with sinapic acid > ferulic acid > caffeic acid (Dimitrić Marković et al., 2000, 2005).

The λmax of metal treated ACN did not follow the same pattern as the spectral responses, such that the ACN with most substituted acylating moieties did not exhibit the largest

λmax. The aliphatic acylated Cy derivative (CGM) exhibited the smallest λmax, indicating that aromatic acylation and increasing glycosylation played a stronger role in copigmentation of ACN-metal chelates. The λmax was slightly larger for the aromatic monoacylated ACN with Al3+, Figure 4.2, where CtGF (ferulic acylation) > CtGC (p- coumaric acylation) > CtGS (sinapic acylation). Diacylated Cy derivatives showed the largest λmax in which CtGSC > CtGSF > CtGSS. Interestingly, the pattern differs between the λmax of the monoacylated Cy and their diacylated Cy derivative counterparts, while the λmax of CtG was always lower.

Table 4.1: Average and (standard deviation) bathochromic shifts (nm) of Cy derivatives

(50 µM) treated with factorial excess of Al3+ or Fe3+ (1× [ACN] in pH 6 and 0.5× [ACN] in pH 7) to [ACN], n = 3

pH 6 pH 7 ACN Al3+ Fe3+ Al3+ Fe3+ CGM 18.3 (6.0) 31.7 (5.7) 11.0 (1.7) 15.3 (1.5) CtG 35.0 (2.6) 38.0 (3.5) -0.3 (0.6) 6.7 (1.2) CtGC 19.0 (1.0) 22.0 (1.0) 0.7 (1.2) 0.7(1.2) CtGF 31.7 (0.6) 43.0 (1.0) 0.7 (1.2) 6.3 (2.5) CtGS 30.7 (4.2) 51.3 (1.2) 0.3 (2.1) 5.0 (4.0) CtGSC 55.0 (1.0) 82.7 (1.2) -1.0 (1.0) 4.3 (1.5) CtGSF 47.0 (1.0) 64.3 (2.3) 0.0 (0.0) 12.7 (1.5) CtGSS 4.3 (0.6) 6.3 (2.6) 0.0 (0.0) 9.7 (1.5)

In pH 7, the spectral responses of these Cy derivatives to metal ion chelation were more difficult to characterize. Bathochromic shifts were large in pH 6, Table 4.1.

However in neutral pH, the observed changes in λmax were small and even hypsochromic in some cases, especially with increasing metal ion concentrations, Figure 4.3. In comparison, the responses of ACN to Fe3+ were slightly larger than to Al3+ in agreement with our previous findings, Table 4.1. The minimal bathochromic responses of the ACN suggest that the ACN may have achieved the largest λmax possible for these chemical structures. It has been found that ACN typically demonstrated their largest λmax in pH 7-8 and then showed hypsochromic responses as pH was further increased (Cabrita, Fossen,

& Andersen, 2000; Fossen, Cabrita, & Andersen, 1998). Although the changes in λmax of

ACN caused by metal chelation were small or negative, alterations in the visible absorption spectra of the pigments occurred and resulted in change in color expression of the pigments in solution.

Figure 4.3: Plot of ln(Absorbance at λmax of to) vs. time (hr) of CtGC and CtGSC in pH 7 treated with factorial excesses of Al3+ (0-5× [ACN]), n = 3

Table 4.2: CIE-L*a*b* values average and (standard deviation) of Cy derivatives (50

µM) treated with factorial excess of Al3+ or Fe3+ (1× [ACN] in pH 6 and 0.5× [ACN] in pH 7) to [ACN], n = 3

pH 6 pH 7 ACN Al3+ Fe3+ Al3+ Fe3+ L* a* b* L* a* b* L* a* b* L* a* b* 82.4 6.8 -11.2 73.9 0.6 ± -14.1 69.4 1.0 -16.4 68.7 -2.7 -12.2 CGM (0.4) (0.2) (0.4) (0.3) (0.0) (0.5) (1.1) (0.3) (0.9) (0.7) (0.1) (0.4) 94.7 1.2 -4.3 89.9 -1.7 -5.0 80.4 -6.2 -22.5 82.0 -7.5 -15.1 CtG (0.3) (0.1) (0.1) (0.2) (0.0) (0.1) (0.0) (0.10 (0.2) (0.5) (0.2) (0.6) 94.4 0.4 -4.5 89.8 -2.0 -5.8 82.5 -9.1 -19.1 79.6 -8.7 -18.8 CtGC (0.2) (0.1) (0.2) (0.5) (0.2) (0.2) (0.7) (0.2) (0.5) (0.8) (0.1) (0.7) 94.6 0.08 -5.3 88.7 -3.5 -6.9 82.5 -8.7 -19.8 80.8 -9.2 -17.2 CtGF (0.3) (0.1) (0.3) (0.9) (0.2) (0.5) (1.2) (0.3) (1.3) (1.3) (0.3) (1.4) 80.8 1.8 -19.4 76.2 -6.0 18.4 74.6 -10.4 -27.1 73.5 -11.6 -22.6 CtGS (0.6) (0.0) (0.1) (0.7) (0.1) (0.3) (0.6) (0.1) (0.4) (0.4) (0.3) (0.6) 64.3 -22.1 -31.2 64.9 -23.2 -23.5 65.2 -35.3 -32.4 64.1 -32.0 -28.5 CtGSC (1.0) (0.3) (0.7) (0.8) (1.4) (0.5) (0.7) (0.2) (0.5) (0.9) (0.1) (0.5) 30.9 -13.0 -39.6 31.3 -14.1 -29.7 71.6 -30.1 -29.3 71.8 -30.1 -28.9 CtGSF (0.0) (0.3) (0.5) (1.2) (1.0) (1.0) (0.8) (0.9) (1.1) (1.7) (1.2) (1.7) 68.6 -9.6 -33.9 70.0 -14.0 -26.5 67.7 -25.6 -34.3 68.1 -26.2 -26.9 CtGSS (1.6) (0.4) (1.6) (1.2) (0.6) (0.7) (0.7) (0.3) (1.0) (1.1) (0.5) (1.1)

In addition to the bathochromic shift of the ACN when chelating metal ions, the visible absorbance spectrum typically showed an increase in absorbance from 380-480 nm. Absorbance in this region typically corresponds to the observation of yellow hues, while the λmax of these ACN typically correspond to blue hued colors. Table 4.2 shows the colorimetric values of the isolated ACN with treatment of Al3+ or Fe3+. In agreement with the spectral absorbance values, metal treated ACN solutions became darker, bluer, and less red. As observable in Table 4.2, the acylating pattern also played a role in the color expression of the ACN-metal chelates. CGM and CtG showed the most red-purple color, having the largest positive a* values or smallest negative a* values. Interestingly, the L* value was smaller for CGM than for the ACN monoacylated with hydroxycinnamic acid derivatives, but the colors exhibited by these pigments were comparatively bluer. In pH 6 and 7, the b* values for the monoacylated derivatives were smallest for CtGS < CtGF < CtGC and were similar for a* values, suggesting stronger bluing effects with increasing substitution of the phenolic acid. In all cases, the diacylated

Cy samples showed the lowest L*, a*, and b* values. The smallest (or most negative) a* values (most green) occurred for CtGSC < CtGSF < CtGSS while no clear pattern was apparent for the b* values. However, negative a* and b* values were observed for all diacylated Cy derivatives, showing blue hues.

4.5.2 Stability

In this study, blue hued colors were developed by some ACN-metal chelates in pH 6; however all chelates expressed variations of blue hues in pH 7. The λmax and absorbance of CtG was lower than those of the other CtG derivative chelates; therefore, it

100 expressed a less intense blue color. Additionally, ferric iron chelates of acylated Cy derivatives from purple carrots have been found to have greater stability and larger half- live values than chelates of nonacylated Cy derivatives from elderberry in pectin systems stored at 20 ºC (Buchweitz et al., 2013b). For these reasons, stability of only the acylated

ACN chelates in pH 7 was evaluated. Significant differences were found between the different Cy derivatives, the treatment ratios, and the two metal ions used.

Metal chelation by ACN was found to decrease the rate of degradation of the pigments and extend their color expression, as found previously (Sigurdson & Giusti,

2014; Tachibana et al., 2014). The kinetics of degradation of the ACN chelates were found to follow first order kinetics agreeing with previous studies (Buchweitz et al.,

2013a, 2013b). Figure 4.3 shows the degradation pattern of CtGC and CtGSC with Al3+ treatment during 24 hours of ambient storage, demonstrating adherence to first order kinetics and also representing the patterns of degradation of all samples. It can also be observed in Table 4.3 that with each factorial increase of metal ion concentration, the rate of ACN degradation decreased significantly expressed by increased half-lives. However, it important to note that certain metal ions or large excesses of them can promote ACN degradation (Kuusi et al., 1977; Pyysalo & Kuusi, 1973; Sigurdson & Giusti, 2014), but findings of this study suggest ratios of ACN:metal ion can be optimized to promote stability of these types of pigments.

101

Table 4.3: Average and (standard deviation) of half-life (t1/2 expressed as hr) and rate constants (k) of Cy derivatives (50 µM) in pH 7 treated with factorial excesses of Al3+ or

Fe3+ (0×, 0.5×, 5× [ACN]) during dark storage at 21-25 ºC, calculated by 1st order kinetics by decrease in absorbance of λmax of to, n = 3

t1/2 (hr) Rate constant (k) Al3+ Fe3+ Al3+ Fe3+ ACN 0× 0.5× 5× 0.5× 5× 0× 0.5× 5× 0.5× 5× 18.7 18.9 62.4 23.7 113.4 -0.037 -0.037 -0.013 -0.029 -0.006 CGM (0.6) (1.2) (32.7) (0.6) (1.5) (0.001) (0.002) (0.001) (0.002) (0.000) 8.1 8.5 22.5 12.6 29.4 -0.086 -0.082 -0.032 -0.055 -0.024 CtGC (1.3) (0.5) (5.4) (0.8) (4.7) (0.013) (0.004) (0.007) (0.003) (0.004) 9.9 12.6 52.9 16.6 30.4 -0.071 -0.055 -0.014 -0.042 -0.023 CtGF (1.4) (0.6) (14.0) (1.7) (5.1) (0.010) (0.003) (0.004) (0.005) (0.004) 14.8 20.1 31.9 20.4 33.9 -0.047 -0.035 -0.022 -0.034 -0.021 CtGS (0.8) (0.9) (3.60 (0.5) (3.4) (0.002) (0.002) (0.002) (0.001) (0.002) 58.8 103.4 149.1 102.1 64.8 -0.012 -0.007 -0.005 -0.007 -0.011 CtGSC (4.0) (24.3) (33.4) (26.5) (11.5) (0.001) (0.002) (0.001) (0.002) (0.002) 34.2 34.6 40.2 105.6 57.0 -0.021 -0.021 -0.017 -0.008 -0.012 CtGSF (6.9) (6.8) (4.7) (54.5) (4.5) (0.004) (0.004) (0.002) (0.003) (0.001) 33.1 33.2 41.0 97.7 53.1 -0.021 -0.021 0.017 -0.007 -0.014 CtGSS (0.1) (0.5) (1.2) (2.6) (11.2) (0.000) (0.000) (0.000) (0.000) (0.003)

The structural composition of the acylating groups was also found to impact the stability of the pigments. As it is well established that acylation plays an important role in the stabilization of ACN pigments, these findings were expected. From linear regression models, the half-life of the ACN-metal chelates were calculated, Table 4.3. Interestingly

ACN-metal chelates showed a larger half-life than most of the other monoacylated peaks despite bearing aliphatic acylation. Generally, ACN acylated with malonic acids are considered to be more labile to hydrolysis than those acylated with hydroxycinnamic acid derivatives (Rodriguez-Saona, Giusti, & Wrolstad, 1999), so this finding was unexpected.

102

Acylation of ACN with malonic acid is thought to participate in metal chelation by aiding in deprotonation of the aglycone at C7 and by forming hydrogen bonds with hydroxyl groups of the chromophore (Elhabiri et al., 1997). This stabilization likely played a role in the increased stability and perhaps by interfering with hydration of the ACN by blocking the chromophore or by attraction of water molecules by the carbonyl groups of the acid. It was found that the other monoacylated chelates showed increasing half-lives as substitution of the hydroxycinnamic acid attachments increased, CtGS > CtGF >

CtGC. These findings are further supported by reports that increasing methoxylation of acylating groups improved the thermal stability of ACN (Patras, Brunton, O’Donnell, &

Tiwari, 2010).

In all cases, the diacylated derivatives showed greater stability and larger half- lives than the monoacylated Cy derivatives, as expected. For example, the diacylated

ACN derivatives found in red radish were more stable than the monoacylated ACN from red potato (M. M. Giusti & Wrolstad, 2003; Rodriguez-Saona et al., 1999). Essentially in opposite order of their monoacylated counterparts, the diacylated ACN with more heavily substituted acylating groups were found to have comparatively shorter half-lives: CtGSC

> CtGSF > CtGSS. In opposition to their co-pigmenting effects, the methoxyl attachments of ferulic and sinapic acids perhaps reduced the strength of interaction between the acylating groups and the chromophore by reducing the planar characteristics of the co-pigments, allowing for increased hydration of the chromophore. Similar to the findings of this study, diacylated ACN derivatives of red cabbage were found to be more stable than monoacylated ACN in simulated gastrointestinal digestion (McDougall et al.,

103

2007). More interestingly CtGSF (diacylated with ferulic and sinapic acids) was found to be more stable than CtGSS (diacylated with sinapic acids), in agreement with the stability findings of this study (McDougall et al., 2007). Additionally, antioxidant activity for hydroxycinnamic acids follows that sinapic > caffeic >> ferulic > p-coumaric acids

(McDougall et al., 2007), suggesting higher reactivity and less stability for these molecules or those bearing them.

The calculations for half-lives were based on changes in absorption at the λmax of time = 0 hr, and during the course of the study, the λmax were not completely static. From

Table 4.3, it can be observed that the differing metal ions affected the stability of the

ACN chelates in addition to the structure of the ACN. Most Fe3+ ACN chelates showed slightly lower absorbance values than the Al3+ chelates, but Fe3+ chelates appeared to have larger half-lives than chelates of Al3+. These findings could be explained by the fact that Fe3+ in solution showed a yellow color causing some retention of absorbance while

ACN degraded, and Al3+ in solution expressed no color. It is also important to note that colors solutions of the Fe3+ chelates were much more brown-gray than those observed for the Al3+ chelates. The absorbance spectrum of the chelates before and during storage reflected these findings and showed comparatively much broader peaks for Fe3+ chelates indicating less of a pure color expression. Previously, stability of the ACN-metal chelates was found to be higher with Fe2+ ions > Al3+ > Fe3+ (Tachibana et al., 2014), in partial agreement with the findings of this study. Despite the larger half-lives of the ACN-Fe3+ complexes, in terms of absorbance, the colors of the ACN-Al3+ chelates changed much less and were comparatively less brown. This suggests Fe3+ may have induced increased

104 rates of oxidation or degradation of ACN pigments resulting in brown hues, perhaps due to the oxidative-reductive properties of the ion.

4.6 Conclusion

The spectral responses and stability of these ACN to metal chelation were result of several factors including the ACN acylation pattern, environment (pH), and the specific metallic cation. Both metal ions of this study induced changes on the color expression of the ACN, but responses were typically larger for chelation of Fe3+ compared to Al3+. In pH 6 where bathochromic shifts were largest, they were greater for diacylated Cy (except CtGSS) than for the monoacylated or nonacylated Cy derivatives.

The small and sometimes even negative bathochromic shifts in pH 7 were difficult to generalize but did result in ACN color changes, becoming bluer or sometimes greener.

The λmax was greatest for metal treated diacylated Cy; therefore, they expressed the most blue hues in the widest range of conditions. Cy acylated with malonic acid only expressed purple hues, but the other monoacylated Cy and triglycosylated Cy samples were comparatively bluer.

Metal chelation by ACN also improved the color stability of the pigments. The complexation induced hyperchromic shifts in absorbance creating more intense colors to be observed. The rates of degradation were observed to sequentially decrease with each increase in metal concentration. Diacylated Cy derivatives were found to have higher stability than the monoacylated ACN. Interestingly acylation with malonic acid seemed to improve the stability of the Cy derivative showing a larger half-life than 2

105 monoacylated Cy derivatives. Of the monoacylated ACN, increasing substitution of the acylating moiety improved stability while having an opposite effect on diacylated Cy.

Metal chelation of these acylated Cy derivatives allowed for the development of differently hued blue colors in lower pH than ACN would typically express these hues.

With appropriate degree and structure of acylation, metal chelation can allow for selection for the desired blue tone hues and also increased stability for ACN based colorants. The chelation of metallic cations by ACN may prove useful in food application by enhancing the stability of these types of pigments.

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Chapter 5: Evaluation of Hydroxycinnamic Acid Derivatives as Inter- and Intra-

molecular Copigments of Cyanidin Derivatives and their Metal Chelates

5.1 Abstract

Anthocyanins are versatile naturally occurring pigments that produce red-purple- blue hues in nature, especially when interacting with metal ions and copigments. The ability to recreate the variety of blue hues of these pigments is not fully understood. To further elucidate, hydroxycinnamic acids were evaluated as inter- (0-50× [ACN]) and intra-molecular copigments of cyanidin derivatives and their Al3+ chelates by spectrophotometry (380-700 nm) and colorimetry (CIE-L*c*hº) in pH 5-8. With no Al3+,

λmax and absorbance was greatest with cyanidin with diacylation > monoacylation > increasing concentration of intermolecular copigments. Al3+ chelation increased absorbance 2-42× and λmax ≳ 40 nm (pH 5-6) compared to copigmentation alone.

Intermolecular copigmentation weakly impacted the color characteristics (ΔE < 5) of cyanidin; acylation resulted in ΔE 5-15. Triglycosylated cyanidin expressed blue color in pH 7-8, indicating glycosylation pattern to alter the chromophore’s planar characteristics.

Metal chelation with intramolecular copigmentation resulted in the most blue hues.

5.2 Keywords

Anthocyanin; anthocyanin-metal chelate; copigmentation; Brassica oleracea var. capitata f. rubra 107

5.3 Introduction

The replacement of synthetic food colorants with naturally derived alternatives has been steadily increasing, being driven by consumer demand (Wrolstad & Culver,

2012). Red and yellow colorants derived from natural sources are considered more prevalent and simpler to reproduce from pigment classes that can include betalains, carotenoids, and anthocyanins (ACN) (Newsome et al., 2014). Some naturally derived blue pigments with potential food use include anthocyanins (ACN), phycocyanins, and iridoid-derivatives (Buchweitz, Carle, et al., 2012; Sigurdson & Giusti, 2014).

ACN are a subset of polyphenolic compounds responsible for many of the red, purple, and blue colors of fruits, vegetables, and flowers. These flavonoid compounds undergo structural changes, depending on environmental pH and substitution pattern, resulting in changes in color observation. The deprotonated quinonoidal forms typically occurring in alkaline pH appear in purple and blue hues (Newsome et al., 2014), but in acidic conditions common to many food products, ACN appear in red-purple structural forms (Sigurdson & Giusti, 2014). Through mechanisms observed in some floral systems such as self-association, molecularly copigmentation, and metal chelation, ACN can adopt arrangements that appear blue even in acidic conditions (Yoshida et al., 2009).

Copigmentation results as changes in the visible absorbance spectrum of the ACN when an essentially colorless molecule stacks on the planar nucleus of the ACN inducing a color change (Malien-Aubert et al., 2001). The mechanism of interaction is thought to be result of hydrophobic interactions or Π-Π stacking (Di Meo et al., 2012). The magnitude of copigmentation depends on many factors including molecular structures,

108 individual reactant concentrations, and their ratios (Gómez-Míguez et al., 2006).

Copigmentation can occur intermolecularly, between 2 unbound molecules and also intramolecularly, within the same molecule. The hydroxycinnamic acids that act as acylating groups of red cabbage ACN have been shown to act as intermolecular copigments, exhibiting increasing bathochromic shifts as the acids were increasingly methoxylation (sinapic > ferulic > caffeic) (Dimitrić Marković et al., 2000, 2005).

Intramolecular copigmentation is considered more efficient as the participating molecules are spaced for interaction by the covalent linkages between the glycosylating and acylating moieties (Malien-Aubert et al., 2001). Increasing amount of acylation on the same chromophore was found to lead to increased bathochromic shifts (Dangles et al.,

1993a).

ACN can also express blue colors by metal chelation; however, bathochromic shifts are only induced on ACN bearing catechol or pyrogallol moieties on the B ring

(Bayer, Egeter, Fink, Nether, & Wegmann, 1966; Buchweitz et al., 2012; Schreiber et al.,

2010). Metal ions displace hydrogen ions from the hydryoxyl groups, inducing conversion of flavylium cations (red) to quinonoidal bases (blue) (Schreiber et al., 2010).

Metal chelation has been found to occur with both acylated and nonacylated ACN

(Buchweitz, Carle, et al., 2012; Sigurdson & Giusti, 2014). However, acylated ACN extracts were found to express comparatively larger λmax and more blue colors

(Buchweitz et al., 2013a, 2013b; Sigurdson & Giusti, 2014). ACN of red cabbage and purple carrot share the same major aglycones and similar acylating moieties, but the

109 sugar moieties to which their acylating groups are attached differ. Therefore, different blue hues were expressed by chelates of these pigment (Buchweitz et al., 2013a, 2013b).

Metal ions have also been found to coordinate macromolecular complexes that express blue colors. Referred to as metalloanthocyanins, these pigments are composed of fixed stoichiometric ratios of ACN, flavones, and metal ions (6:6:2) (Yoshida et al.,

2009). The metal ions are centralized, coordinating both conversion of the ACN to anionic forms and also the chiral association of ACN and flavones in respective anti- clockwise and clockwise manners (Yoshida et al., 2006, 2009). Metal complexes with

ACN have also been found to occur with copigments other than flavones. The blue coloration of hydrangeas is result of “fuzzy metal complexes,” non-stoichiometric metal complex pigments stabilized by copigments (Yoshida et al., 2009). The composition of this pigment was determined to be consist of Dp-3-glucoside, Al3+ ions, 5-O- caffeoylquinic acid , and/or 5-O-p-coumaroylquinic acid (Kondo, Toyama-Kato, &

Yoshida, 2005). Additional blue pigments have been found to be result of combination of

ACN, metal ions, and copigments including flavanols, quercetin, and chlorogenic acid

(Jurd & Asen, 1966; Yoshida et al., 2006).

Copigmentation is thought to play an important role in the formation of blue colors by ACN-metal chelates, especially intramolecular copigmentation as in ACN extracts of red cabbage and purple carrot (Buchweitz et al., 2013a, 2013b; Sigurdson &

Giusti, 2014). From these findings, it can be hypothesized that hydroxycinnamic acid derivatives, the acylating moieties of red cabbage ACN, could play a factor in blue color expression of ACN-metal chelates as intermolecular copigments. The objective of this

110 study was to evaluate the role of the hydroxycinnamic acid derivatives, found on red cabbage ACN, as both intermolecular and intramolecular copigments of Cy derivatives and of Cy-metal chelates to better understand the mechanism of blue color expression.

The findings of this study will expand the current body of knowledge for formation of

ACN-metal based pigments.

5.4 Materials & Methods

5.4.1 Materials

ACN extracts were prepared from fresh red cabbage (Brassica oleracea var. capitata f. rubra) purchased from a grocery store in Columbus, OH. The predominant

ACN of red cabbage were Cy derivatives, sharing the same glycosylation pattern with varying degrees of aromatic acid acylation (Ahmadiani et al., 2014). Hydroxycinnamic acid derivatives for this study included p-coumaric acid, ferulic acid, and sinapic acid. P- coumaric acid was purchased from MP Biomedicals, Inc. (Solon, OH), and ferulic acid

(99%) and sinapic acid (≥98%) were obtained from Sigma-Aldrich, Co. (St. Louis, MO).

Sources of metal ions included lab grade aluminum sulfate hydrate and reagent grade ferric chloride hexahydrate, obtained from Fisher Scientific (Fair Lawn, NJ).

Buffer systems were prepared from tris(hydroxymethyl)aminomethane, 99% (Alfa Aesar,

Ward Hill, MA) and ACS grade sodium acetate anhydrous (Fisher Scientific, Fair Lawn,

NJ). Hydrochloric acid (certified ACS Plus), sodium hydroxide N/10 (0.0995-0.1005) and all other standard ACS and HPLC grade reagents were purchased Fisher Scientific

(Fair Lawn, NJ).

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5.4.2. Methods

5.4.2.1 Anthocyanin Extraction and Purification

ACN were extracted from the red cabbage plant materials with acidified acetone and isolated by phase partition with water and chloroform, following procedures described by Rodríguez-Saona & Wrolstad, 2001. These crude ACN extracts were purified by loading onto activated Waters Sep-pak® C18 (Ireland) cartridges and washed with 0.01% HCl acidified water, ethyl acetate, and recovered with 0.01% HCl acidulated methanol. Pigments were stored in acidified water until further treatment.

5.4.2.2 Anthocyanin Isolation

For evaluation of hydroxycinnamic derivatives as intermolecular copigments of the ACN-metal chelates, ACN of red cabbage were treated with potassium hydroxide to saponify the ester bonds between the acylating and glycosylating moieties, according to procedures of Giusti & Wrolstad, 1994. This Cy triglycosylated derivative (Cy-3- diglucoside-5-glucoside: CtG) was further isolated by semi-preparative high performance liquid chromatography (HPLC). In order to investigate the phenolic acids as intramolecular copigments, the major monoacylated Cy derivatives (CtGMa) and major diacylated Cy derivatives (CtGDa) were collected as respective fractions through use of semi-preparative HPLC.

Reverse phase HPLC was conducted with a HPLC system (Shimadzu, Columbia,

Maryland, U.S.A.) equipped with LC-6AD pumps, CBM-20A communication module,

SIL-20A HT autosampler, CTO-20A column oven, and SPD-M20A Photodiode Array detector. LCMS Solution Software (Version 3, Shimadzu, Columbia, Maryland, U.S.A.)

112 was used to analyze results. To achieve separation of ACN, a Luna reverse-phase pentafluorophenyl (PFP2) column with 5 µm particle size and 100Å pore size in 250 x

21.2 mm column size (Phenomenex®, Torrance, CA, U.S.A) was used with a binary gradient consisting of solvents A: 4.5% formic acid in HPLC water and B: HPLC acetonitrile. Gradient began at 15% B and increased to 30% B from 0-30 min with a flow rate of 10.0 mL/min. Elution of ACN was monitored at 520 nm, and desired peaks were manually collected. After collection, the isolated or fractionated pigments were subjected to solid phase extraction, as described above, and stored in acidified water until further analysis.

5.4.2.3 Isolated Anthocyanin Purity

The purity of the isolated ACN was verified by analytical reverse phase HPLC with a similar system (Shimadzu, Columbia, Maryland, U.S.A.) using different pumps:

LC-20AD. The analytical column was a Kinetix reverse-phase pentafluorophenyl (PFP2) column with 2.6 µm particle size and 100Å pore size in 100 x 4.6 mm column size

(Phenomenex®, Torrance, CA, U.S.A. Flow rate was set to 0.6 mL/min with a run time of 46 min. The same binary system was used for this method, but conditions differed. The gradient began at 0% B, then increased 0-10% from 0-1 min, and followed 10-30% B from 1-46 min. Spectral data was monitored 270-700 nm with elution of ACN monitored at 520 nm. ACN were isolated to a purity representing ≥ 87% of the total absorbance from 270-700 nm.

113

5.4.2.4 Monomeric Anthocyanin Quantitation

ACN were quantitated by the pH differential method, described by Giusti and

Wrolstad (2001). The concentration of monomeric ACN was calculated based on absorbance differences at the λmax for CtG, λ520 for CtGMa, λ530 for CtGDa, and at 700 nm in pH 1 and pH 4.5. The % area under the curve, from analytical HPLC, at the respective λ of detection was used to calculate the concentration of each of the main anthocyanin pigments in each fraction by multiplying this percent by the measured absorbance according the spectrophotometric monomeric anthocyanin quantitation method. These were then summed to calculate the monomeric ACN of each fraction.

Figure 5.1: HPLC chromatogram (detection at 520 nm) of red cabbage anthocyanins illustrating major isolated cyanidin derivatives

All ACN were expressed as the equivalents as follows, based on molecular weight and molar absorptivity. CtG was expressed as Cy-3-diglucoside-5-glucoside equivalents.

ACN content of CtGMa was expressed as the sum of Cy-3-(p-coumaroyl)-diglucoside-5- glucoside, Cy-3-(feruloyl)-diglucoside-5-glucoside, Cy-3-(sinapoyl)-diglucoside-5- glucoside; and ACN concentration CtGDa was expressed as the sum of Cy-3-(p- coumaroyl)-(sinapoyl)-diglucoside-5-glucoside, Cy-3-(feruloyl)-(sinapoyl)-diglucoside- 114

5-glucoside, and Cy-3-(sinapoyl)-(sinapoyl)-diglucoside-5-glucoside, with minor ACN being equivalents of Cy-3-diglucoside-5-glucoside. Figure 5.1 shows the HPLC chromatogram identifying the isolated peaks and fractions. Molar absorptivities of these

Cy derivatives from red cabbage were determined by Ahmadiani, Robbins, Collins, &

Giusti, 2015.

5.4.2.5 Sample Preparation

Isolated ACN were diluted to 50 µM concentrations in 0.5 M buffers of either sodium acetate for pH 5-6 or 1M TRIS for pH 7-8. For evaluation of the phenolic acids as intermolecular copigments, ACN extracts and the hydroxycinnamic acid solutions were diluted together with appropriate buffer solutions for ACN:acyl ratios of 1:0, 1:1, 1:10,

1:25, and 1:50. Individual phenolic acids were dissolved in the same buffer solutions reflective of pH in concentrations of approximately 1 g/L on the day of use. The pH of samples was monitored with a Mettler Toledo Inc. S220 SevenCompact™ pH/Ion meter

(Columbus, OH). Salts of Al3+ were dissolved in distilled water in concentrations of 0.6

M and 0.06 M. These salt solutions were then added to anthocyanin solutions beginning at equal M concentrations and then in factors to an excess of 0.5, 1, 5, 10, 50, 100×

[ACN]. All samples were equilibrated for 45 min at room temperature in the dark prior to analysis. Three replicates were evaluated for each treatment.

5.4.2.6 Visible Spectrophotometry of Solutions

Samples were evaluated by visible transmittance (380-700 nm) spectrophotometry using 1 cm plastic cuvettes in a Shimadzu UV-2450 UV-Visible spectrophotometer

(Shimadzu, Columbia, Maryland, U.S.A.).

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5.4.2.7 Colorimetry of Solutions

Aliquots of samples were transferred to 2 mm path length plastic cells and read for CIE-L*, chroma*, and hue angle* using a Hunter ColorQuest XE (Hunter Labs,

Reston, VA, U.S.A.). The settings for measurement included total transmittance, D65 illuminant, and a 10° observer angle. The mathematical differences of the colors between

CtG and treated samples were calculated according to the CIE-L*a*b* Delta E2000

(ΔE2000) equation (Mokrzycki & Tatol, 2012).

5.4.2.8 Statistical Evaluation of Data

Figures and data means with associated standard deviations were produced using

Microsoft Office Excel 2010 (Office 14.0, Microsoft. Redmond, WA). The λmax, bathochromic shifts, and absorbance of the different metal ratio treated individual ACN were evaluated by 1-way analysis of variance (ANOVA) (2-tailed, α = 0.05) and Fisher’s least significant difference (LSD) (α = 0.05) using Minitab 16 (Minitab Inc., State

College, PA). For the specific ACN-metal ratios, λmax, bathochromic shifts, and colorimetric characteristics (L*, c*, and hº values) of the different isolated ACN were compared by 1-way analysis of variance (ANOVA) (2-tailed, α = 0.05) and Fisher’s least significant difference (α = 0.05).

5.5 Results and Discussion

5.5.1 Hydroxycinnamic Acid Derivatives as ACN Copigments

Previous work from this laboratory has shown hydroxycinnamic acid derivatives as acylating moieties to improve the blue color expression of Cy derivatives and their associated metal chelates. It is also known that intermolecular copigmentation can induce

116 bathochromic shifts on ACN. Therefore it can be hypothesized that hydroxycinnamic acid derivatives may promote additional bathochromic shifts on nonacylated Cy anthocyanins resulting in additional blue color expression.

[ACN]) in pH 5-8

pH 5 pH 6 pH 7 pH 8 Sample λ λ λ λ λ max λ max λ max λ max max Al3+100× max Al3+10× max Al3+0.5× max Al3+0.5× CtG 527.7 39.7 540.7 35.0 591.3 -6.0 594.3 -5.3 1× 521.7 43.7 540.5 40.7 592.7 -4.3 596.7 -3.7 10× 524.0 43.0 555.0 41.0 593.0 -3.3 597.0 -4.3 25× 528.7 43.0 545.3 40.7 591.0 -2.0 597.0 -3.0 50× 528.3 45.7 568.7 40.7 590.3 -1.0 597.0 -1.7 CtGMa 539.3 49.0 549.7 40.0 597.3 1.3 605.7 2.3 CtGDa 548.0 58.0 575.0 50.7 598.0 10.7 612.0 13.0

117

Figure 5.2: λmax (left) and absorbance (right) of Cy-3-diglucoside-5-glucoside (CtG) treated with factorial excesses (0-50× [ACN]) of hydroxycinnamic acid derivatives in pH

5-8, n = 3

Despite aiding in anthocyanin blue hue as bound copigments, the free hydroxycinnamic acid molecules in solution were found to act as poor copigments especially in low concentrations. The λmax of treated samples were found to differ significantly, p-value < 0.05. In general, the greatest λmax was found for ACN samples treated with the highest concentration of hydroxycinnamic acids, table 5.1 and figure 5.2.

The range of the bathochromic responses was generally small, as can be observed by differences in λmax in table 5.1 and figure 5.2. Bathochromic Responses varied from as little as -1.0 nm (pH 7) to 2.7 nm (pH 8), 6.6 nm (pH 5), and as much as 28.2 nm (pH 6).

Significant differences were observed in pH 6-7; however, none were found to occur in pH 5 or 8. Despite the large range of λmax in pH 6, little change in color was observed due to the lack of distinctive peak or λmax and low absorbance < 0.07. The small responses found to occur in pH 5-6 were consistent with previous findings, in which the effects of

118 copigmentation of ACN by sinapic acid were low compared to pH 3.6, the optimal pH level (Dimitrić Marković et al., 2005). No clear trend emerged regarding the bathochromic shifts on ACN with inclusion of these phenolic acids as intermolecular copigments.

The most observable effect of hydroxycinnamic acids on ACN absorbance resulted in hyperchromic shifts. All samples differed significantly by inclusion of the acids and also with the increasing concentration of them. Trends were very clear in that hyperchromic responses were largest with the highest concentration of phenolic acids

50× > 25× > 10× ≈ 1× ≈ 0×; figure 5.2 demonstrates general increase in absorbance with increasing concentration of the copigments. With concentrations of hydroxycinnamic acids 50× [ACN], the effects of intermolecular copigmentation induced increases of absorbance by 46%, 168%, 38%, and 113% in pH 5, 6, 7, and 8, respectively. Some larger hyperchromic responses were observed in pH 7-8 and may have been due in part to the fact that these phenolic acids expressed an observable yellow coloration not seen in acidic pH.

With the covalent attachment of these phenolic acids to the chromophore CtG, greater bathochromic and hyperchromic shifts observed compared to those resulting from the unbound acids. These findings were consist with previous studies which suggested the participating molecules are spaced for interaction by the covalent linkages (Malien-

Aubert et al., 2001). Comparatively, absorbance was increased by 100-200% for monoacylated CtGMa, and CtGDa showed a range of 276-3196% (pH dependent) greater than the absorbance of CtG. The λmax of the nonacylated Cy derivative was increased

119 from 6.0 nm (pH 7) to as much as 17.7 nm (pH 5) with monoacylation and ranged 6.7

(pH 7) to 34.3 (pH 6) nm with diacylation. The bathochromic responses were always greatest for diacylated (CtGDa) samples and were generally followed by monoacylated

(CtGMa) samples. An exception occurred in pH 6 in which bathochromic responses were greater for intermolecular copigment ratios of 25× and 50× [ACN]. These findings were also consistent with previous studies (Dangles et al., 1993a).

pH 5 pH 6 pH 7 pH 8 Sample L* C* hº L* C* hº L* C* hº L* C* hº 89.5 0.8 109.5 95.9 0.7 62.2 92.4 3.7 256.0 87.0 3.5 206.3 CtG (0.1) (0.0) (1.3) (0.0) (0.0) (0.5) (0.0) (0.0) (0.2) (0.0) (0.0) (2.0) 96.5 0.4 17.6 96.4 0.5 5.5 92.5 4.4 257.4 93.4 3.0 213.1 1× (0.0) (0.0) (0.5) (0.0) (0.0) (1.6) (0.0) (0.0) (0.2) (0.1) (0.0) (3.8) 96.4 0.6 10.8 96.2 0.4 29.9 93.0 3.2 250.1 92.8 3.3 192.9 10× (0.0) (0.0) (0.6) (0.0) (0.0) (1.2) (0.0) (0.1) (0.2) (0.1) (0.1) (0.8) 96.0 0.8 352.4 96.2 0.5 21.1 88.9 7.9 249.7 89.4 7.5 209.6 25× (0.0) (0.0) (0.8) (0.0) (0.0) (1.0) (0.1) (0.1) (0.2) (0.0) (0.0) (0.3) 96.1 0.8 21.5 95.6 0.4 33.8 91.1 4.5 261.7 88.9 6.9 205.4 50× (0.0) (0.0) (1.3) (0.1) (0.0) (7.9) (0.0) (0.0) (0.4) (0.1) (0.1) (0.0) 96.1 1.2 335.2 96.2 0.8 326.0 89.3 9.5 262.3 87.2 14.8 235.5 CtGMa (0.0) (0.0) (0.3) (0.0) (0.0) (1.3) (0.1) (0.1) (0.1) (0.0) (0.1) (0.0) 86.2 15.7 317.2 86.8 11.2 287.3 82.5 17.7 251.4 81.2 23.1 231.1 CtGDa (0.1) (0.2) (0.2) (0.0) (0.2) (2.0) (0.1) (0.1) (0.3) (0.1) (0.0) (0.1)

In the low acid conditions of this study, intermolecular copigmentation of CtG with hydroxycinnamic acid derivatives was not found to impact the color expression of

ACN strongly. This can be observed by the small changes (and generally decreases) in

120 chroma, table 5.2. Although the hue angle did vary considerably, especially in pH 5, this was likely result again of low color intensity expression of ACN due to the structural conformations these pigments form in low acid conditions. The effects of intermolecular copigmentation, as acylation, were much more obvious of the color expression of ACN.

In pH 5-6, hue angle proceeded to increase from more red values (0-90º) to more purple values (270-360º), with the diacylated samples showing the most purple-blue colorations.

Of special interest, CtG was found to express blue colorations in neutral and alkaline pH (hue angle 256º and 206º, respectively) despite lack of acylation or added copigments, table 5.2. Acylation has traditionally been considered to play a larger role in the blue color expression of ACN; further supported by previous work from this lab found that Cy-3-glucoside, even with added metal ions, expressed dark purple-brown colors in similar pH conditions. Distortions of the planar aspects of the chromophore

(including stretching, bending, or torsion) alter the pigment’s π-delocalization and therefore affecting its interaction with light and color expression (Malcıoğlu et al., 2011).

Glycosylation of the aglycone alters the absorbance spectra of the molecule due to constraining the dynamics of the molecule (Malcıoğlu et al., 2011). It can also be observed that hue angle of CtG generally decreased, approaching more green hues, with increasing concentration of phenolic acids in solution in pH 7-8. This was likely result due to blue color expression of the ACN in these pH in combination with the yellow color the hydroxycinnamic acids delivered. Acylation of the pigments typically resulted in hyperchromic effects, observable also as decreases in L* value and increases in C* value indicating more intense colorations. In low acidity, diacylation resulted in more

121 blue like color expression than other samples. For trained observers, a just noticeable difference in color could be detected with a ΔE of 2.4; however, noticeable color differences for untrained observers occurs with ΔE values of 3.5-5 (Mokrzycki & Tatol,

2012). In all pH, intermolecular copigment resulted in ΔE less than 5 indicating little color difference and would likely not be observed by the untrained eye. However, covalent attachment of the hydroxycinnamic acid derivatives typically resulted in ΔE greater than 5, illustrating the more pronounced color differences between these samples.

Diacylation resulted in the largest ΔE values, > 11, and therefore, the greatest difference in color expression compared to CtG.

5.5.2 Hydroxycinnamic Acid Derivatives as ACN Copigments with the Presence of

Metal Ions

The induction of bathochromic and hyperchromic shifts on the visible absorbance spectrum of ACN by metal chelation has been well illustrated (Buchweitz et al., 2013b;

Sigurdson & Giusti, 2014). Similar findings were observed from this study, in that all samples showed the same types of responses to the presence of Al3+, observed as increases in λmax and absorbance demonstrated in figure 5.3. The λmax of treated samples were found to differ significantly again by ANOVA, p-value < 0.05. In all pH levels

3+ tested and with all Al ratios, the λmax of CtGDa was greatest followed by CtGMa, table

5.1. In most other cases, hydroxycinnamic acid derivatives in concentrations of 50×

[CtG] showed the next greatest λmax, followed by 25×, similar to findings regarding copigmentation without metals discussed earlier. The λmax of all these samples was substantially less than acylated CtG samples, table 5.1. With evaluation by Fisher’s LSD,

122 the remaining sample treatments with phenolic acids (1× and 10× [ACN]) were found not to significantly differ from the λmax of CtG.

Figure 5.3: λmax (left) and absorbance (right) of Cy-3-diglucoside-5-glucoside (CtG) treated with factorial excesses of hydroxycinnamic acid derivatives (0-50× [ACN]), monoacylated CtgMa derivatives, and diacylated CtgDa derivatives with factorial excess of Al3+ (0-100× [ACN]) in pH 5, n = 3

These findings indicate that these types of chemical structures do not strongly interact with metal ions and are only slightly further coordinated to ACN molecules by them. Similar results were found regarding the structure of the blue hydrangea pigment composed of Dp-3-glucoside, quinic acid substituted hydroxycinnamic acids, and Al3+ ions (Kondo et al., 2005). Without the attachment of quinic acid to caffeic or coumaric acid, blue coloration by these pigments was not observed, indicating the role of the derivative in interaction with the metal ions and coordination to the ACN molecules

(Kondo et al., 2005). It is believed that the Al3+ ions interacts between the vicinal hydroxyl groups of the ACN B-ring and 1-hydroxy, 1-carboxylic acid, and the carbonyl

123 residue in the ester at 5-position of 2 and/or 3 of these chlorogenic like acids (Kondo et al., 2005).

In comparison to the bathochromic shifts induced on CtG by intermolecular copigmentation by hydroxycinnamic acids, those caused by Al3+ chelation were very large, table 5.1. Consistent with previous findings, the largest bathochromic effects were observed in slightly acidic pH 5-6. Generally shifts in λmax in pH 7-8 were hypsochromic, concurring with prior findings from this laboratory. The factorial excesses of Al3+ to

[ACN] to obtain largest bathochromic shifts increased as pH decreased (Sigurdson &

Giusti, 2014), indicating why data from selected factors were presented. Within each factorial ratio of Al3+ to [ACN], the bathochromic shifts induced were found to significantly differ according to ANOVA. Comparing the bathochromic shifts with untreated CtG as point of reference, those experienced by CtGDa were greatest in all Al3+ ratios and in all pH and were generally followed by CtGMa. Similarly with presence of

Al3+, bathochromic shifts were largest with highest concentration of hydroxycinnamic acids in solution and generally decreased in magnitude as concentration of intermolecular copigments decreased. An interesting observation was found when comparing the metal induced bathochromic shifts to the ACN respective control without added Al3+, rather than comparing them to CtG. In these cases, the change in λmax of the acylated Cy derivatives was always less than that of the non-acylated Cy samples. This further implicates the role covalent attachment of aromatic acids to ACN in the degree of bathochromic shift, or effectively intramolecular copigmentation.

124

The hyperchromic effects on ACN sample absorbances induced by Al3+ chelation observed in this study followed the same general trends as described previously regarding the role of molecular copigmentation, figure 5.3. Again, the absorbances of all treatments

(metal and molecule ratios) were found to differ significantly by ANOVA and Fisher’s

LSD. It was also observed that the absorbance of CtG was increased with increasing concentrations of hydroxycinnamic acids and fixed concentrations of Al3+, implicating some interaction between the pigment, copigmenting molecule and metal ion, figure 5.3.

For example, in pH 5 with Al3+ 100× [ACN], absorbance of CtG was increased by 739% with no copigmenting molecules to 1,437%, 1,403%, 1,772%, 1,932% with hydroxycinnnamic acids 1×, 10×, 25×, and 50× [ACN], respectively. When the copigments were bound to the chromphore, absorbance was increased 2,373% with monoacylation and 6,479% with diacylation. These same trends were observed in all pH.

Consistent with the hyperchromic effects of copigmentation without metal ions, largest increases in absorbance were observed in pH 5 > 6 > 8 > 7; although the hyperchromic shifts induced by Al3+ chelation were substantially larger.

125

pH 5, Al3+ 100× pH 6, Al3+ 10× pH 7, Al3+ 0.5× pH 8, Al3+ 0.5× Sample L* C* hº L* C* hº L* C* hº L* C* hº 87.9 1.6 297.3 93.6 2.0 289.8 90.2 6.7 253.5 86.4 4.7 197.7 CtG (0.0) (0.0) (0.3) (0.0) (0.0) (0.2) (0.0) (0.1) (0.7) (0.0) (0.0) (0.2) 91.8 6.1 299.0 94.0 2.6 288.1 90.7 6.8 252.1 93.1 3.7 197.9 1× (0.0) (0.0) (0.0) (0.0) (0.0) (0.1) (0.0) (0.0) (0.3) (0.0) (0.0) (0.3) 91.9 5.6 298.9 94.0 2.0 285.1 91.6 4.9 246.2 92.6 3.9 187.3 10× (0.0) (0.1) (0.0) (0.0) (0.0) (0.2) (0.0) (0.0) (0.1) (0.0) (0.0) (0.5) 90.8 6.9 298.0 93.5 2.6 285.6 89.2 7.7 245.2 89.4 7.5 209.6 25× (0.0) (0.0) (0.1) (0.0) (0.0) (0.2) (0.1) (0.1) (0.2) (0.0) (0.0) (0.3) 90.2 6.9 297.0 94.0 1.4 286.2 88.3 8.4 241.5 88.2 8.7 198.2 50× (0.0) (0.1) (0.0) (0.0) (0.0) (0.2) (0.1) (0.1) (0.1) (0.1) (0.0) (0.3) 89.2 9.8 286.1 92.9 4.4 272.6 86.9 14.0 245.3 86.8 15.7 233.2 CtGMa (0.0) (0.0) (0.0) (0.0) (0.0) (0.1) (0.2) (0.1) (2.5) (0.1) (0.0) (0.4) 78.1 22.5 270.1 81.0 20.1 256.4 83.0 23.3 225.8 83.0 25.1 219.9 CtGDa (0.0) (0.0) (0.0) (0.1) (0.2) (0.1) (0.0) (0.0) (0.1) (0.0) (0.0) (0.0)

Metal chelation by these Cy derivatives resulted in the expression of more blue like hues than caused by molecular copigmentation. Table 5.3 shows hue angles < 300º for each of the Al3+ treated samples in all pH, approaching variants of purple and blues colors. Even with the addition of metal ions, acylation of CtG was found to result in comparatively bluer colors than those with addition of intermolecular copigments.

Similar to samples without added Al3+, the hue angles of CtG with added copigments were approaching green colors (~ 200º hue angle) in pH 8, hue angles less than their counterparts without Al3+. This same observation was found to occur in all the pH levels of this study, tables 5.2 and 5.3, suggesting slightly increased interaction between the

ACN and copigments due to presence the metal ions. Just noticeable color differences for

126 untrained observers may possibly be observed when comparing the colors of CtG derivatives treated with equal concentrations of Al3+ and hydroxycinnamic acid derivatives in pH 5 and 8, as evidenced by ΔE values of 4-5. However in pH 6-7, ΔE values were < 1. With Al3+ chelation, diacylation resulted in ΔE 14-15 in all pH compared to CtG treated with same proportion of Al; while monoacylation resulted in ΔE

6-9, except in pH 6 were ΔE was 2.4. As expected, increasing degree of acylation resulted in the largest color differences and most blue hues.

5.6 Conclusions

Hydroxycinnamic acid derivatives were found to act as intermolecular and intramolecular copigments of ACN and ACN-metal chelates, with the color expression of the pigments to depend on several factors. As intermolecular copigments, with or without added metal ions, hydroxycinnamic acids did not strongly alter the color expression of

ACN based pigments. Their main effects resulted in hyperchromic shifts on the pigments absorption spectra, increasing absorbance with increasing concentrations of the copigments. Interestingly, the acids were found to express yellow colorations in neutral and alkaline pH which impacted the color of the solution containing ACN pigments. As intramolecular copigments, bathochromic and hyperchromic shifts on ACN were much more obvious and allow for more blue color expression in a wider range of environmental conditions.

The interactions of the anthocyanins with metal ions were much more strongly evident, resulting in much larger bathochromic and hyperchromic shifts than the effects of molecular owed for increased blue color expression. The data of this study also

127 demonstrated metal ions to interact with hydroxycinammic acid derivatives free in solution, likely coordinating them to ACN pigments. This was suggested by a decrease in hue angle of about 10 degrees in all pH levels tested when comparing similar sample treatments with and without metal ion treatment. With metal ions, hydroxycinammic acids covalently bound to the ACN resulted in blue colorations in a wider pH range with a much higher intensity.

Application of metal ions has again shown potential to allow for increased color intensity expression by ACN and also allow for expression of more blue hues in a wider range of environmental conditions. This work of this study also provides additional insight in the mechanism of ACN metal chelation, as well as the effects of different phenolic compounds acting as copigments.

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Chapter 6: Evaluating the Role of Metal Ions in the Bathochromic and

Hyperchromic Responses of Cyanidin Derivatives in Acidic and Alkaline pH1

6.1 Abstract

In many food products, colorants derived from natural sources are increasingly popular due to consumer demand. Anthocyanins are one class of versatile and abundant naturally occurring chromophores that produce different hues in nature, especially with metal ions and other copigments assisting. The effects of chelation of metal ions (Mg2+,

Al3+, Cr3+, Fe3+, and Ga3+) in factorial excesses to anthocyanin concentration (0-500×) on the spectral characteristics (380-700 nm) of cyanidin and acylated cyanidin derivatives were evaluated to better understand the color evolution of anthocyanin-metal chelates in pH 3-8. In all pH, anthocyanins exhibited bathochromic and hyperchromic shifts. Largest bathochromic shifts most often occurred in pH 6; while largest hyperchromic shifts occurred in pH 5. Divalent Mg2+ showed no observable effect on anthocyanin color while trivalent metal ions caused bathochromic shifts and hue changes. Generally, bathochromic shifts on anthocyanins were greatest with more electron rich metal ions

(Fe3+ ≈ Ga3+ > Al3+ > Cr3+).

______

1 Gregory T Sigurdson, Rebecca J Robbins, Thomas M. Collins & M Mónica Giusti Department of Food Science and Technology, The Ohio State University. 2015 Fyffe Rd. Columbus, OH 43210 Published in Food Chemistry 2016, 208, 1:26-34

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6.2 Keywords

Anthocyanin; anthocyanin-metal chelate; metalloanthocyanin; Brassica oleracea var. capitata f. rubra; Aronia melanocarpa

6.3 Introduction

Color additives in food may serve several purposes, including enhancement and correction of already present colors, standardization of colors of raw materials, giving a color identity to otherwise colorless foods, such as sodas or confections, and also accounting for color loss during storage or processing (Newsome et al., 2014; Potera,

2010). Although all food coloring additives have undergone safety evaluations and are closely regulated, the use of artificial colorants has become less desirable as consumers are requesting naturally sourced options. Additionally, the European Union (EU) has mandated that foods containing the synthetic colorants used in the “Southampton Study” bear warning labels indicating the dyes may cause hyperactivity in children, furthering the demand for natural options (FDA; CFSAN, 2011). Therefore, the development of food colorants from natural sources has become relevant for food companies to remain competitive in a changing market or in the international marketspace.

Naturally derived options for red and yellow colorants are more prevalent than those for blue and can include pigments such as betalains, carotenoids, and anthocyanins in some cases. The limited options for blue pigments are also considered a limitation in the development of green and purple hues in foods. Some blue pigments that have shown potential for commercial application include phycocyanins (from Spirulina spp.) and iridoid-derivatives (derived from the huito fruit and gardenia flowers) (Buchweitz, Carle,

130 et al., 2012). However, use of these pigments has been limited due to poor stability, regulatory restrictions, or inability to match the hues of currently used synthetic dyes

(Buchweitz, Carle, et al., 2012). Additional alternatives for synthetic blue colorants may be found in anthocyanins (ACN) which can produce some blue hues found in nature.

Therefore the exploration of the color expression of anthocyanins is timely and relevant.

ACN are a class of polyphenolic compounds found in nature that are responsible for many red, blue, and purple colors observed in many fruits, vegetables, and flowers.

Also known for their potential health benefits, they show potential to act as alternatives for synthetic food colorants with added benefits (He & Giusti, 2010). ACN exist in a dynamic equilibria of chemical species whose ratios vary depending on pH, becoming bluer as pH increases. In acidic pH common to many food products, ACN appear in red- purple chemical species. However, self-association, co-pigmentation, and metal chelation can result in blue colorations even in acidic conditions, like those found in flowers

(Yoshida et al., 2009).

For ACN to chelate metal ions (M+) and demonstrate a color change, the chromophore must bear at least 2 free hydroxyl groups on the B ring (Bayer et al., 1966;

Buchweitz, Carle, et al., 2012; Schreiber et al., 2010). Multivalent Mn+ act in competition of hydrogen ions, transforming flavylium cations (red) to the quinoidal base anions (blue)

(Schreiber et al., 2010). It is thought a stacking arrangement then occurs between the transformed ACN with additional ACN molecules, stabilizing a metal coordinated complex (Schreiber et al., 2010). Unlike blue metalloanthocyanins, which are fixed stoichiometric ratios of ACN:flavones:Mn+ of 6:6:2, these ACN-Mn+ chelate complexes

131 seem to exists under equilibria with varying degrees of ACN association from single molecules to complexes of 3 molecules (Sever & Wilker, 2004; Xu, 2013; Yoshida,

Kitahara, Ito, & Kondo, 2006; Yoshida et al., 2009). Recent works have evaluated the effects of ACN structure, regarding B ring and acylation, in acidic pH with trivalent metal ions (Fe3+ and Al3+), finding ACN B rings with 3 free hydroxyl groups to exhibit the largest bathochromic shifts or those with acylation to show greater λmax and most blue hues (Buchweitz, Carle, et al., 2012; Sigurdson & Giusti, 2014).

Many Mn+ have been found to induce bathochromic shifts on the visible absorption of ACN, but all are multivalent (Bayer et al., 1966; Buchweitz, Carle, et al.,

2012; Pyysalo & Kuusi, 1973; Schreiber et al., 2010; Sever & Wilker, 2004; Sigurdson &

Giusti, 2014; Smyk, Pliszka, & Drabent, 2008; Xu, 2013; Yoshida et al., 2006). Some include but are not limited to Mg2+, Al3+, Ti2+, Fe3+, and Cu2+. The degree of the induced bathochromic shift has been found to vary depending on the M+, suggesting the electron density and their organization within specific orbitals of the ion play important roles in the interaction between ACN and M+. In methanol, cyanidin (Cy) aglycones were found to experience bathochromic shifts of 25nm and 152nm in the presence of Pb2+ and Cr3+, respectively (Ukwueze et al., 2009). ACN have also been shown to develop blue colors in the presence of Al3+ and Fe3+, with ACN-Fe3+ chelates experiencing larger bathochromic shifts (Buchweitz, Carle, et al., 2012; Sigurdson & Giusti, 2014). Conversely, the color

ACN from berries was found to evolve into brown hues with Fe2+ and Fe3+ ions (Pyysalo

& Kuusi, 1973).

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To expand the current body of knowledge regarding chelation by ACN, the aim of this study was to explore the response of Cy derivatives in presence of Mn+ ions of increasing size and number of d-orbitals and evaluate their spectral and color responses.

The selection of Mn+ included Mg2+, Al3+, Cr3+, Fe3+, and Ga3+, of which Cr3+ and Ga3+are not approved for food use but reflected varying electron configurations which was valuable for this study. In addition to the multiple ions, this work also systematically evaluated the effects of increasing proportions of Mn+ on known ACN concentrations in pH 3-8, which have been shown to alter the stoichiometric ratios of ligand to Mn+ and

n+ also the λmax of the observed complexes. The ability of ACN to chelate M in alkaline pH was not found to have been evaluated previously.

6.4 Materials & Methods

6.4.1 Materials

Anthocyanin rich extracts of red cabbage (Brassica oleracea var. capitata f. rubra) chokeberry (Aronia melanocarpa) were prepared from respective fresh products and commercial juice concentrate purchased from a local grocery store in Columbus, OH.

The predominant ACN of each is a Cy derivative, having 2 free hydroxyl groups on the B ring; however, red cabbage ACN showed varying degrees of aromatic acid acylation that was not found in chokeberry ACN (Sigurdson & Giusti, 2014).

Lab grade aluminum sulfate hydrate, reagent grade ferric chloride hexahydrate, and reagent grade magnesium chloride hexahydrate were purchased from Fisher

Scientific (Fair Lawn, NJ). Chromium chloride hexahydrate, purity ≥ 98.0%, and

133 anhydrous gallium chloride, purity ≥ 99.999%, were purchased from Sigma-Aldrich Co.

(St. Louis, MO).

Tris(hydroxymethyl)aminomethane, 99%) was obtained from Alfa Aesar (Ward

Hill, MA). ACS grade sodium acetate anhydrous, hydrochloric acid (certified ACS Plus), and sodium hydroxide N/10 (0.0995-0.1005) were purchased Fisher Scientific (Fair

Lawn, NJ) as were all other standard ACS and HPLC grade reagents.

6.4.2 Methods

6.4.2.1. Anthocyanin Extraction

Extraction of ACN from plant materials followed procedures described by

Rodríguez-Saona & Wrolstad, 2001. Plant materials were powdered, treated with 0.01%

HCl acidified acetone and filtered. The filtrate was added to separatory funnel with 1-2 volumes chloroform and stored overnight at 4 °C. The chloroform phase was appropriately discarded; knowing chloroform is a mild carcinogen. The ACN rich aqueous layer was retained.

6.4.2.2 Anthocyanin Purification – Solid Phase Extraction

Crude ACN extracts were purified by loading onto activated Waters Sep-pak®

C18 (Ireland) cartridges. Pigments were washed with 0.01% HCl acidified water, ethyl acetate, and recovered with 0.01% HCl acidulated methanol, which was removed in a rotary evaporator at 37 °C under vacuum. Treatment with water removed polar compounds such as sugars and acids, and ethyl acetate washes removed many other polyphenolic compounds that could potentially act as copigments, especially in the presence of Mn+. Pigments were stored in acidified water until further analysis.

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6.4.2.3 Monomeric Anthocyanin Quantitation

Monomeric ACN were quantitated by the pH differential method, described by

Giusti & Wrolstad, 2001. Briefly, anthocyanins were quantified by measuring the absorbance at 520 nm and 700 nm at pH 1 and pH 4.5 for each sample. Based on absorbance differences, the concentration of monomeric anthocyanins was summarized for dilution to known concentrations. Red cabbage ACN were expressed as cyanidin-3- diglucoside-p-coumaroyl-5-glucoside equivalents (Ahmadiani et al., 2015), and chokeberry ACN were expressed as equivalents of cyanidin-3-glucoside using (M. Giusti

& Wrolstad, 2001). Figure 6.1 shows the identity of the major ACN of each extract.

Figure 6.1: HPLC chromatograms and identification of chokeberry and red cabbage

ACN after purification, detection at 520 nm and at 260-700 nm. Cy: cyanidin; soph: sophoroside; glu: glucoside; gal: galactoside; ara: arabinoside

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6.4.2.4 High Pressure Liquid Chromatography (HPLC) - Anthocyanin Purity

Evaluation

Purified ACN extracts were evaluated for integrity and purity by reverse phase

HPLC. The system was composed of LC-20AD pumps, CBM-20A communication module, SIL-20A HT autosampler, CTO-20A column oven, and SPD-M20A Photodiode

Array detector (Shimadzu, Columbia, Maryland, U.S.A.). LCMS Solution Software

(Version 3, Shimadzu, Columbia, Maryland, U.S.A.) was used to analyze results.

Separation of the pigments was achieved with a Kinetix reverse-phase pentafluorophenyl (PFP2) column with 2.6 µm particle size and 100Å pore size in 100 x

4.6 mm column size (Phenomenex®, Torrance, CA, U.S.A. Flow rate was set to 0.6 mL/min with a run time of 46 min. A binary gradient was used with solvents A: 4.5% formic acid in HPLC grade water and B: HPLC acetonitrile; it began at 0% B, increased

0-10% from 0-1 min, 10-30% B from 1-46 min. Spectral data was monitored 260-700 nm for non-ACN impurities with elution of ACN monitored at 520 nm, Figure 6.1 demonstrates the low levels on non-ACN contaminants.

6.4.2.5 Sample Preparation

ACN of chokeberry or red cabbage were diluted to 50 µM concentrations in 0.5

M buffers of either sodium acetate (pH 3-6) or (TRIS) (pH 7-8), depending on pH. The pH of samples was monitored with a Mettler Toledo Inc. S220 SevenCompact™ pH/Ion meter (Columbus, OH). Individual metal salts were diluted in distilled water to achieve concentrations of 0.6 M and 0.06 M. These salt solutions were then added to anthocyanin solutions beginning at equal M concentrations and then in factors to an excess of 500×

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[ACN]. High molarity buffers were used to minimize decrease in pH associated with addition of high metal excesses, and pH was monitored throughout the course of analysis.

Control samples without added salt were maintained at each pH. All samples were equilibrated for 45 min. at room temperature in the dark prior to further analysis. Three replicates were evaluated for each sample.

6.4.2.6 Spectrophotometry of Solutions by UV-Visible Transmission

After equilibration, 300 µL of each sample was evaluated by visible transmittance

(380-700 nm) spectrophotometry in 96 well plates using a Molecular Devices

SpectraMax 190 Microplate Reader (Sunnyvale, CA). Spectrograms were generated using Microsoft Office Excel 2010 (Office 14.0, Microsoft. Redmond, WA).

6.4.2.7 Statistical Evaluation of Data

Figures as well as means and standard deviations of data replicates were produced using Microsoft Office Excel 2010 (Office 14.0, Microsoft. Redmond, WA). Evaluation of λmax anthocyanin-metal treatments and respective bathochromic shifts was conducted by 1-way analysis of variance (ANOVA) (2-tailed, α = 0.05) and Fisher’s least significant difference (α = 0.05) of λmax as well as associated absorbances using Minitab 16

(Minitab Inc., State College, PA).

6.5 Results and Discussion

ACN extracts were purified by solid phase extraction, as described previously.

Ethyl acetate was a critical solvent as part of these procedures to remove a majority of non-ACN phenolic compounds. These types of compound are known to act as copigments of ACN, modifying their color. Additionally, some other phenolic

137 compounds are known to complex with Mn+ resulting in color expression of these otherwise colorless molecules (Mellican, Li, Mehansho, & Nielsen, 2003). To minimize these potential effects, ACN extracts were semi-purified to achieve low levels of non-

ACN contaminants, Figure 6.1.

6.5.1 Role of pH on Anthocyanin-Metal Chelation

6.5.1.1 Role of pH on Bathochromic Responses

The acylated and non-acylated Cy derivatives evaluated in this study experienced bathochromic and hyperchromic shifts in the presence of most of these Mn+, Figure 6.2, agreeing with previous studies. In trends similar to ACN in solution, pH was found to play an important role in the color expression of the pigment chelates and also the amount of the bathochromic shift and even the ACN:Mn+ ratios. Formation of blue ACN-

Mn+ chelates has previously been found to occur in a limited slightly acidic pH range of

4-6 with low Mn+ levels (Bayer et al., 1966; Buchweitz et al., 2013a; Buchweitz, Carle, et al., 2012; Buchweitz, Nagel, et al., 2012), but previous work from this laboratory has shown chelation to also occur in pH < 4 with large excess of Mn+ (Sigurdson & Giusti,

2014). In support of previous findings, Mn+ chelation appeared to occur most easily in weakly acidic pH 4-6, with the largest bathochromic responses occurring in pH 6 (Figure

6.2). Interestingly, increases in λmax were also found to occur in pH 7-8 but in lesser magnitudes, likely due to predominance of colored basic structural forms of ACN in these pH.

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Fe3+, or Ga3+), pH 6

6.5.1.2 Role of pH on Hyperchromic Responses

Schreiber et al., 2010, have proposed that after conversion of the red flavylium cation to the blue quinoidal base, the Mn+ also coordinates self-association of ACN molecules. Based on the absorbance increases at constant pH with addition of metal ions

(Figure 6.2), this concept has gained further supporting evidence. At neutral and alkaline pH, where bathochromic shifts were minimal, increases in absorbance were still observed

(Figure 6.2), suggesting chromophore association. Self-association of ACN has been found to increase absorbance in a positive non-linear fashion, deviating from the

Lambert-Beer Law (González-Manzano et al., 2008). In studies evaluating Mn+ chelation by catechols, complexes were found to exists in ligand:Mn+ ratios of 1:1 in pH < 5, ratios of 2:1 in pH 6-7, and 3:1 in increasing pH, which may also suggest increased number of

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ACN association with increasing pH in the presence of Mn+ (Sever & Wilker, 2004; Xu,

2013). ACN are thought to self-associate primarily through hydrophobic interactions of the aromatic nuclei of the chromophores, or Π-Π interactions, which help to protect the pigments from hydration by water molecules and increase stability (González-Manzano et al., 2008). It can be suggested that a similar mechanism, coordinated by M+, functions to increase absorbance and stability of ACN-Mn+ chelates.

6.5.1.3 Role of pH on ACN-Mn+ ratios

Environmental pH was also found to affect the proportion of Mn+ needed to achieve ACN color evolution. To achieve bathochromic shifts, the necessary proportion of tested Mn+ to [ACN] decreased as pH increased (Figure 6.3). For example, the amount of necessary Al3+ to obtain largest bathochromic shifts followed an almost logarithmic pattern, decreasing roughly by a factor of 10 for each unit increase of pH, partially observable in Figure 6.3. This finding supports the idea that hydrogen ions compete against the Mn+ for the B ring hydroxyl binding sites (Dangles, Elhabiri, & Brouillard,

n+ 1994b). Very small increases in λmax occurred in pH 3 despite the high of M content used in this study, but previous work showed highest λmax of ACN in pH 3 to occur with

Al3+ excess of 500-1,000× [ACN] (Sigurdson & Giusti, 2014). In pH 4, the largest bathochromic shift was observed with Al3+ content 100-500× [ACN], with [Al3+] 10-50×

[ACN] in pH 5, and with [Al]3+ 1-5× [ACN] in pH 6, following almost logarithmic patterns (Figure 6.3). Not all ACN chelates completely adhered to this pattern due to reactivity of each specific M+, observable in Figure 6.2; further discussion will follow related to the role of the specific Mn+ used in this study.

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Figure 6.3: λmax (nm) and absorbance of red cabbage ACN treated with factorial excess of Al3+ to [ACN] (0-100×), pH 3-8

Similar trends were observable when comparing the absorbance data, which increased in samples with Mn+ treatment (Figure 6.3). Generally, the largest hyperchromic effect was found to occur in pH 5 for most M+. This finding was likely due to conversion of the typical colorless carbinol pseudobase structures to colored forms by

Mn+ treatment. Large excesses of Mn+ were found to have a comparative hypochromic effect on absorbance compared to treatments with lower [Mn+]. In acidic pH, this was found to occur after exceeding 200× [ACN], but in neutral or alkaline pH, hypochromic effects were observed with ≥ 5× [ACN]. A rise in absorbance occurred after 50× [ACN] in pH 7-8; this was attributed to the decrease in solution pH due to the proton like effects of excessive Mn+ concentration and the low buffering capacity of TRIS. Addition of all metals resulted in pH decreases of ≤ 0.15 until factors exceeded 100× [ACN] in acidic pH or until factors were ≥ 50× [ACN] in alkaline pH. The overall trend of decreases in absorbance with high Mn+ concentration were likely due to aggregation and precipitation 141 of ACN-Mn+ chelates which occurred more readily in alkaline pH. Similar observations have been found in the solubility of catechol-Mn+ chelates, suggesting overall net neutrality for the complexes in pH ≥ 7 (Sever & Wilker, 2004).

6.5.2 Role of Specific Metal Ions in Anthocyanin-Metal Chelation

In aqueous solution, metals do not necessarily exist as charged ions as water molecules tend to associated around the charge helping to neutralize it (Mason, 2013).

This fact may also play a role in the amount of Mn+ necessary to induce to bathochromic shifts on ACN in aqueous solutions, as the interference of water molecules would be negligible in alcoholic systems. The occurrence of metal chelation by anthocyanins can be inferred by application of the Crystal Field Theory. This theory assumes that the central Mn+ is regarded as positively charged point surrounded by sets of negatively charged points, representing ligands (Mason, 2013). The electrostatic field (or “crystal field”) generated by the ligands repel the electrons of the M+, raising the energy of the d orbitals; and with the ligands arranged in specific geometries, the relative energy of the d orbitals will reorganize to conserve overall energy (Mason, 2013). The organization of electrons in the reacting orbitals plays a critical role in the formation and stabilization of

M+-ligand complexes.

The d orbitals surrounding the nucleus occupy different regions in space. The orbitals –dxy, -dyz, and -dxz (termed collectively as T2g orbitals) exhibit their dominant electron density off the main axes while the –dx2-y2 and –dz2 (Eg orbitals) have their dominant electron density on the main axes (McCleverty, 1990). With a series of ligands equidistantly located in space, approaching the Mn+ along the x, y, and z axes, the

142 electrons located on the axes (Eg orbitals) would interact directly with the ligands while those off the axes will not (McCleverty, 1990). Under this situation, the Eg and T2g d orbitals no longer share equal amounts of energy due to increased energy from interaction of Eg orbital electrons with the negatively charged ligands. These energy differences in the presence of external ion fields have been termed orbital splitting, but overall, the total energy of the d orbitals remains unchanged (McCleverty, 1990). Therefore, an increase in the energy of the Eg orbitals results in a decrease of the energy of the T2g d orbitals.

Orbital splitting results in deviation from Hund’s Rule (McCleverty, 1990) which states electrons will not pair in an orbital until all orbitals of the same energy contain 1 electron.

This results in different electron configurations; low spin state in which the minimum amount of unpaired electrons in the different orbitals exists. High spin state occurs when adhering to Hund’s Rule, and unpaired electrons exist in the maximum amount throughout the orbitals. Orbital splitting impacts various properties of Mn+ including ionic radii, ionic stability, and importantly, the overall system energy before and after reactions

(McCleverty, 1990). Thus, these effects have important implications in determining degree of reactivity and stability of the ligand-Mn+ complexes and should likely affect the interactions between ACN and different Mn+.

Using quantum theory of atoms in molecules, complexes of Cy with Al3+ or Mg2+ were found to be stabilized by ion-dipole electrostatic interactions rather than by electron pair sharing (Kunsági-Máté, Ortmann, Kollár, Szabó, & Nikfardjam, 2008). The covalent character of these bonds was increased by replacement of Mg2+ with Al3+ and also replacement of water molecules surrounding the Mn+ by the ACN ligands (Kunsági-Máté

143 et al., 2008). These data suggest that increasing electron number and charge of the Mn+ will lead to increased ACN-Mn+ interaction and stability, although a smaller ionic radius creates greater electrostatic attraction, resulting in a more stable complex. Mn+ are known to not only coordinate macromolecular complexes of ACN and copigments resulting in color evolution of ACN but also result in color expression of typically otherwise colorless compounds, as in the case of gallic acid, caffeic acid, and chlorogenic acids in the presence Fe2+ (Andjelković et al., 2006; Yoshida et al., 2009).

6.5.2.1 Magnesium

The only divalent metal cation used in this study was Mg2+, which is an essential

Mn+ to life and commonly found bound to ACN in plant systems (Kunsági-Máté et al.,

2008). In addition to being divalent, also lacks any electrons in d orbitals. In general, the ability to form complexes is generally thought to require the presence of unfilled d or f orbitals (McCleverty, 1990). Some work has found that ACN preferentially bind to Mg2+ and Fe3+, while flavones prefer Ca2+ (Kunsági-Máté et al., 2008). Mg2+ has also been found to be an important in the formation of some metalloanthocyanins. The structures of five metalloanthocyanins have been determined, and Mg2+ was found to play role in the stereochemical configurations of all of them (Yoshida et al., 2009).

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Figure 6.4: λmax (nm) of chokeberry and red cabbage ACN treated with factorial excess of Mn+ (Mg2+, Al3+, Cr3+, Fe3+, or Ga3+) to [ACN] (0-500×), pH 5

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Table 6.1: Largest bathochromic shifta expressed in nm (with Mn+ factorial value to

[ACN] inducing shift) of red cabbage and chokeberry ACN induced by different Mn+ in pH 3-8

Red Cabbage pH Mg2+ Al3+ Cr3+ Fe3+ Ga3+ 3 -0.1 15.5 (500×) 3.9 (500×) 18.9 (10×) 38.9 (50-500×) 4 -0.9 34.5 (200×) 10.8 (500×) 34.1 (10×) 36.1 (5-200×) 5 -2.2 28.5 (50-200×) 30.5 (500×) 45.1 (1×) 35.5 (1×) 6 -1.8 42.2 (1×) 38.5 (50×) 60.2 (1×) 45.9 (1×) 7 -1.5 7.8 (1×) 3.1 (10×) 20.5 (1×) 9.5 (1×) 8 -2.3 5.3 (1×) 0.7 (1×) 11.7 (1×) 2.7 (1×) Chokeberry pH Mg2+ Al3+ Cr3+ Fe3+ Ga3+ 3 1.5 22.2 (500×) 8.8 (500×) b 44.9 (≥100×) 4 1.8 51.3 (500×) 29.7 (500×) 60.0 (10×) 47.3 (100-200×) 5 2.3 67.7 (10-100×) 73.0 (500×) 76.0 (5×) 70.0 (5×) 6 b 83.9 (5-10×) 82.6 (50×) b 82.9 (1×) 7 b 82.8 (10×) 57.5 (50×) 102.9 (5×) 77.8 (5×) 8 b 68.1 (5×) b b 68.8 (5×)

a Calculated against mean of all untreated samples (n = 15) b Unable to calculate due to lack of λmax or precipitation

In general, exposing ACN to Mg2+ had almost negligible effects on the on the

ACN spectra and expressed color. This finding was in agreement with the theory that complex formation generally requires unfilled d or f orbitals (McCleverty, 1990). In acidic pH 3-5, the λmax of red cabbage ACN was not found to differ significantly with

2+ Mg treatment; however, the λmax was significantly different in pH 6-8, according to

ANOVA. In each of these pH, the λmax only differed by 3.7 nm or less, Figure 6.4 shows the typical response of red cabbage and chokeberry ACN in pH 5. Chokeberry ACN in pH 6-8 were difficult to analyze spectrophotometrically due to lack of distinctive peaks,

146 but in lower pH, significant differences in λmax were found for samples in pH 4 and 5.

Again, the bathochromic shifts were small, magnitudes ≤ 3.4 nm, when compared to the respective control treatment for this sample set. The bathochromic shifts were significantly reduced when calculated against the mean of the controls from each Mn+ treatment (Table 6.1). These small changes in λmax did not visually impact the observed color.

Figure 6.5: Absorbance of chokeberry and red cabbage ACN treated with factorial excess of Mn+ (Mg2+, Al3+, Cr3+, Fe3+, or Ga3+) to [ACN] (0-500×), pH 5

Trends were more observable regarding the absorbance of ACN with Mg2+ added, but the changes were of small magnitude (≤ 0.05 AU). The difference in the means of absorbance were generally considered non-significant except pH 4 fors chokeberry ACN

147 and pH 7 for red cabbage ACN. Interestingly, a hyperchromic shift was observed for both

ACN samples in all pH when Mg2+ content was 5× that of [ACN]; however, the effect disappeared with increasing Mg2+ concentration, Figure 6.5. Although Mg2+ seemed unable to convert ACN cationic forms (red) into anionic forms (blue), this hyperchromic response suggested that Mg2+ played a role in organizing ACN into supramolecular assembly similar to the role Mg2+ plays in metalloanthocyanin formation (Yoshida et al.,

2009).

6.5.2.2 Aluminum

Al3+ ions have long been known to induce bathochromic shifts on ACN bearing B ring catechol or pyrogallol moieties, important in blue color development of flowers such as hydrangea. In metalloanthocyanins, Al3+ was found to be able to induce for blue color development with delphinidin (Dp), bearing a pyrogallol moiety on the B ring, but insufficient to lead to blue colors with Cy derivatives (Yoshida et al., 2009). Previous studies have also shown ACN-Al3+ based blue colors to develop alcoholic and aqueous based systems in vitro (Buchweitz, Carle, et al., 2012; Schreiber et al., 2010; Sigurdson &

Giusti, 2014). In calculations, Al3+ was determined to displace Mg2+ in ACN-Mg2+ complexes (Cy based) and form more stable complexes (Estévez, Otero, & Mosquera,

2011). This Mn+ has also even been evaluated as a tool to quantify Cy in ACN extracts from edible sources (Mason, 2013). Similar to Mg2+, Al3+ also lacks electrons in d orbitals but is trivalent when ionized.

Results of this study were found to be similar to previous studies; bathochromic

3+ shifts were induced on ACN from both samples with added Al . In all pH, the λmax of the

148

ACN of both sources were found to be significantly different (p-values = 0.000) with each metal ratio treatment. For non-acylated Cy samples, the largest bathochromic shifts ranged in magnitude of 22.2 - 83.9 nm, dependent on pH; while, acylated Cy samples showed bathochromic shifts of 5.3 - 42.2 nm (Table 6.1). Although the bathochromic

3+ shifts of non-acylated Cy were greater, the λmax of Al treated acylated Cy was greater indicating the development of bluer colors, especially in higher pH, Figure 6.4.

Treatment of ACN with Al3+ also had profound effects on their visible light absorbance. In acidic pH, hyperchromic shifts occurred until Al3+ content was 200×

[ACN]. When exceeding this proportion, hypochromic effects followed; but the absorbance was still higher that than untreated ACN. These decreases in absorbance could be related to decreased solubility of the ACN-Mn+ complexes which resulted in precipitation of some of the complexes. In neutral or alkaline pH, hyperchromic effects were found to be highest with much lower Al3+ levels; they were largest in pH 7 and 8 with Al3+ 10× and 1× [ACN], respectively, and followed by absorbance decreases.

Hyperchromic shifts, also dependent on environment, varied from as little as 0.7% and up to 181.9% times the absorbance of the untreated acylated Cy derivatives (0.6% - 181.2% for Cy from chokeberry). The large increases in absorbance, occurring pH 5-6, could be attributed to conversion of colorless forms of ACN to those which absorb and reflect visible light as well as the Mn+ induced association of ACN molecules.

149

6.5.2.3. Chromium

Chromium is a trace mineral essential for humans with an unclear role (Agency for Toxic Substances and Disease Registry, 2012). It is also well documented that heavy metals, including chromium, can be toxic to humans, animals, and plants. They have the ability to disturb RNA and DNA synthesis and also induce cell ultrastructural changes

(Glińska et al., 2007). ACN known for their potential health benefitting properties, may aid in sequestering these Mn+ and reducing potential toxic effects (Glińska et al., 2007).

However, limited amounts of literature regarding the interaction of ACN with Cr3+ or

Cr6+ was found. When roots of Allium cepa L. were exposed to solutions of heavy metals including Cr3+ with and without ACN rich extracts from red cabbage, the toxic effects of

Cr3+ were found to be reduced when ACN were present (Glińska et al., 2007). This could be due to the chelating properties of ACN and also their antioxidative properties. Cr3+ is a

n+ 3+ trivalent M having electrons in d orbitals. Ionized, Cr outer electrons exist in the T2g orbitals, having dominant electron density off the main axes, which would suggest a lower reactivity. Additionally, Cr3+ is typically considered inert as possible reactions would require electron rearrangement and promotion from T2g d orbital electrons to Eg d orbitals with a high energy expenditure.

The addition of Cr3+ to the ACN of this study was also found to induce bathochromic and hyperchromic shifts in their absorption spectra. In all pH, the mean

λmax of the ACN of both sources were found to be significantly different from one another with each metal ratio treatment. For non-acylated Cy samples, the bathochromic shifts were in magnitudes of 8.8 – 82.6 nm, changing with pH; while, acylated Cy samples

150 showed bathochromic shifts of 0.7 – 38.5 nm. Similar trends occurred with Cr3+ such that the bathochromic shifts of non-acylated Cy were greater, but the λmax of acylated Cy were greater. Although the bathochromic shifts induced on ACN were greater than those that occurring from Al3+ treatment, the amount of Cr3+ necessary was obviously greater,

3+ 3+ Figure 6.4. The larger λmax of the ACN was achieved with Cr when compared to Al producing comparatively bluer colors, which was expected due to the effects of d orbital electrons. The need for a higher Cr3+ content was supported by the theory that ionized

Cr3+ has a d orbital electron configuration that lends itself to decreased reactivity.

Typical to Mn+ chelation by ACN, Cr3+ was found to cause hyperchromic shifts on their absorption spectra. In trends similar to the ACN bathochromic responses, the largest hyperchromic shifts occurred with large Cr3+ content (500× [ACN] for all pH, except pH 7) (Figure 6.5). The hypochromic effects observed with increasing Al3+ were not observed for Cr3+, except in pH 7 when exceeding 100× [ACN]. The largest hyperchromic shifts, again dependent on environment, varied from as little as 18.7% (pH

3) to as large as 319.2% (pH 6) times the absorbance of the red cabbage ACN (5.1% -

333.3% for chokeberry ACN), Figure 6.5. These substantially larger hyperchromic shifts are likely due to the same mechanisms involved with Al3+ chelation, but may also be related to the fact that Cr3+ exhibits a blue-green coloration in solution. Of the Mn+ used in this study, only Cr3+ and Fe3+ exhibited visible colors in solution.

6.5.2.4 Iron

Similar to Al3+, the effects of iron addition to ACN were found to be better characterized in the Literature, especially in its ability to produce some ACN based blue

151 colorations. Conflicting findings regarding the role of Fe2+ were found. In some studies

Fe2+ was not found to form any colored complexes with ACN; however, later findings showed Fe2+ to induce larger bathochromic shifts than Fe3+ (Pyysalo & Kuusi, 1973;

Tachibana et al., 2014). Pigment stability was found to be improved with Fe3+ chelation when compared to Fe2+ chelation (Tachibana et al., 2014). The ability of Fe3+ to form purple and blue ACN chelates in acidic pH was found to be better characterized and more consistent. Fe3+ can induce bathochromic shifts on ACN absorption spectra as well as encourage pigment aggregation and precipitation (Buchweitz et al., 2013a, 2013b;

Buchweitz, Carle, et al., 2012; Kuusi et al., 1977; Yoshida et al., 2006). The role of Fe3+ in blue flower formation has gained further evidence in identifying the vacuolar transporter protein responsible for carrying Fe3+ into ACN vacuoles; tulips lacking the gene responsible for production of this protein were found unable to develop blue petals

(Momonoi et al., 2009). In metalloanthocyanins, Fe3+ is known to play an important role in many of the known pigment macromolecules (Yoshida et al., 2009). Those metalloanthocyanins with Cy chromophores require Fe3+ to produce blue hues, while those based on Dp chromophores can develop blue colors with even Mg2+ ions (Yoshida et al., 2006). The enhanced bathochromic shifts are likely due to electron configuration of

Fe3+ cations. As an ion, the outer electrons of Fe3+ are thought to primarily exist in a high spin configuration, having 1 electron in each of the 5 d orbitals. This allows for better overlap with ligands, forming bonds easily and strongly.

As expected, ACN formed bonds with Fe3+ causing bathochromic and hyperchromic shifts in absorbance, and the mean λmax of the ACN of both sources

152 differed significantly with each metal ratio treatment. Some of the largest bathochromic shifts with Mn+ treatments were observed with Fe3+ application. Bathochromic shifts were in magnitudes of 2.0 – 121.4 nm, for non-acylated Cy; while, acylated Cy samples showed bathochromic shifts of 11.7 – 60.2 nm, Table 6.1. Compared to the previous Mn+ discussed, Fe3+ was able to produce large bathochromic shifts in lower concentrations.

Largest effects in pH 3 occurred with Fe3+ 50× [ACN] compared to 100-500× levels needed for Al3+ or Cr3+, Figure 6.4. The large bathochromic shifts observed are likely due to the unique electron arrangement of Fe3+ that encourages strong bonding and the many absorption bands typically of this electron configuration. Of interest, concentrations of

Fe3+ > 50× [ACN] led to development of brown colors expected with ACN degradation.

This degradation may be attributable to the reductive-oxidative capabilities of Fe2/3+, employing the ACN as antioxidants for form conversion. It has also been postulated that

Mn+ promote oxidation (of flavonols) through reactive oxygen species formation (Makris

& Rossiter, 2000). Being that Fe3+ expresses a yellow-orange-brown color in solution, the visible absorption of spectra of ACN treated with Fe3+ began to express some of the characteristics typically of Fe3+, especially in high ratios. A characteristic increase in absorbance in the region of 380-480 nm was observed, Figure 6.2, which correlated with observed color of the Fe3+ solution. This observation potentially played a role in the brown color observation of ACN with high Fe3+ proportions.

The effects on absorbance intensity were more difficult to characterize due to the brown color formation of ACN with Fe3+ concentration > 50× [ACN]. Ignoring Fe3+ concentration > 50× [ACN], hyperchromic shifts ranged from 2.6% (pH 3) to 102.2%

153

(pH 6) times the absorbance of the untreated red cabbage ACN. The hypochromic responses with excess Mn+ content found with other Mn+ treatments were not observed with these samples (Figure 6.5), likely related to discoloration noted with the higher proportions of Fe3+. Interestingly, the hyperchromic responses of ACN with Fe3+ chelation were less than those observed occurring with either Al3+ or Cr3+.

6.5.2.5 Gallium

Gallium was the largest M+, in terms of ionic weight, used in this study; however, its ionic radius is very similar to that of Fe3+. Ga3+ has been found to displace Fe3+ in biological systems (Kaneko, Thoendel, Olakanmi, Britigan, & Singh, 2007). This capability has been employed to inhibit growth of pathogenic bacteria by action of decreasing bacterial uptake of Fe3+ and by interfering with Fe3+ signaling (Kaneko et al.,

2007). Ga3+ also differs from Fe3+ in its electron configuration, having filled d orbitals and lacking electrons in the outer s and p orbitals. This higher electron density could be assumed to promote larger bathochromic and hyperchromic shifts on ACN, and little could be found in Literature regarding the interaction of ACN with Ga3+. In a study

3+ 3+ n+ evaluating Al or Ga complexation by ACN, pKa values for ACN to the M binding were calculated; and in most cases, the values were lower for ACN-Ga3+ complexes

(Elhabiri et al., 1997). This suggests that Ga3+ can bind more strongly to the ACN and compete for the binding site against hydrogen ions more efficiently than Al3+ (Elhabiri et al., 1997).

Treatment of ACN with Ga3+ resulted again in bathochromic and hyperchromic shifts on the ACN of both sources. Similar to Fe3+ treatment, Ga3+ also resulted in some

154 of the largest bathochromic shifts. For non-acylated Cy bathochromic shifts ranged 44.9 –

82.9 nm and 2.7 – 45.9 nm for acylated Cy samples. Compared to the other M+, lower levels of Ga3+ were needed to induce the largest bathochromic shift in acidic pH. Largest effects in pH 3 occurred Ga3+ 10× (for chokeberry ACN) - 50× (for red cabbage ACN)

[ACN] compared to ≥ 100× levels of Al3+ or Cr3+. Interestingly, the largest bathochromic

3+ shifts for ACN in pH 3 were induced by Ga . These findings further support the pKa binding values calculated for ACN to Al3+ or Ga3+ that indicated ACN binding to Ga3+ can occur at lower pH (Figure 6.4 or Table 6.1).31 These large bathochromic shifts were likely result of the filled d orbitals of Ga3+, enhancing UV-visible light absorption.

Regarding the intensity of absorbance, hyperchromic shifts ranged from 5.8% (pH

8) to 179.0% (pH 5) times the absorbance of the untreated red cabbage ACN and 70.7%

(pH3) to 414.5% (pH 6) times higher for chokeberry ACN, Figure 6.5. Hypochromic effects with high Mn+ excesses were not easily classified. Generally, hyperchromic shifts occurred with most Ga3+ additions; however, precipitation was noted to occur with high

Ga3+ concentrations, making it difficult to generalize findings.

6.6 Conclusions

In general, ACN able to chelate these Mn+ experienced bathochromic and hyperchromic shifts in their absorbance spectrum. The spectral responses were result to several factors including the ACN structure (acylation), environment (pH), and also the atomic organization of the metallic cations. ACN lacking acylation typically underwent much larger bathochromic shifts compared to those with acylation; however, the λmax of the acylated ACN with Mn+ was larger in all cases. As expected with ACN chemistry, the

155 environmental pH played a role in the expressed color of the solutions. Additionally, the ratio of ACN:Mn+ to induce the largest bathochromic shift decreased as pH was increased, further indicating competition for the binding sites existed between the Mn+ and protons. Large excesses of Mn+ (> 100× [ACN]) in very acidic environments (pH ≤

3) were essential for blue color development by ACN which may prove impractical for food application unless additional stabilization methods of the pigments are developed to combat the competition of hydrogen ions for catechol or pyrogallol moieties of the chromophore.

The specific Mn+ evaluated in this study were found to impact the observed colors of ACN chelates. Divalent Mg2+ was found to impact ACN minimally, inducing very small bathochromic and hyperchromic shifts that did not impact the color of the observed

ACN. However, the trivalent Mn+ induced significant changes on the color and spectral characteristics of ACN. It was found that generally with increasing electron density, the degree of bathochromic shift increased. The largest bathochromic shifts results generally resulted from Fe3+ ≈ Ga3+ > Al3+ > Cr3+ >> Mg2+; although there were exceptions in different pH. Due to the electron configuration of Cr3+ and thus its low reactivity, high concentrations of this Mn+ were required to induce effects on ACN. The color of the ACN chelates were found to be stable during the course of analysis, and subsequent studies regarding stability will be conducted to further evaluate the suitability of these types as food colorants. Previous works have shown Mn+ to extend the color expression of ACN pigments over time (Tachibana et al., 2014). However, high concentrations of Fe3+ did result in discoloration of ACN, perhaps as result of the reducing and oxidative

156 capabilities of this cation. Precipitation of ACN-Mn+ chelates was noted in this study, as in some previous studies was found to occur but also found to be function of the specific

Mn+, concentration of the Mn+, the environmental pH, and time.

The bathochromic shifts induced on the ACN allowed for the development of colors similar to the currently used synthetic blue food colorants in lower pH than ACN would typically express these hues. Mn+ chelation also allowed for development of many purple shades and some rather dark, almost black, pigments that could expand the useful colorant range of ACN. Additionally, selection of specific Mn+ can allow for specificity of the desired ACN hue by controlling the amount of bathochromic shift as response to unique electron configuration of each Mn+. Chelation of Mn+ by ACN also expanded the color expression range of ACN in more diverse pH environments, such as pH 5 or 6, where these pigments typically express little to no color.

157

Chapter 7: Concluding Remarks

Anthocyanins bearing ortho-dihydroxyl groups on the B ring were found to undergo bathochromic and hyperchromic shifts in absorbance when undergoing metal chelation. Several factors impacted the extent of evolution of the color expression of the pigments relating to the structure of the aglycone, type and degree of substitution of the aglycone, presence of molecules capable of interacting with anthocyanins, environmental conditions such as pH, and also the electronic configuration of the metal ion.

With metal ion chelation, the largest bathochromic shifts occurred with aglycones bearing the highest degree of substitution on the B ring, being greater for delphinidin than cyanidin. Although the bathochromic shifts experienced by acylated anthocyanins were less than those experienced by nonacylated counterparts, the λmax of the metal chelates of acylated anthocyanins was greater in all cases. With and without metal ions present, cyanidin derivatives with aromatic diacylation always showed a larger than aromatically monoacylated cyanidin which was always greater aliphatic acylated or nonacylated cyanidin. The aromatic acyl moieties as free compounds were found to minimally impact the color expression of anthocyanins or related metal chelates, especially when compared to the effects observed by covalent attachment. In general, the most electron rich metal ions, like ferric iron or gallium, provided the largest bathochromic shifts and largest λmax.

Overall, metal chelation by anthocyanins allowed for development of many blues hues that show potential to act as alternatives for synthetic food colorants, even in acidic

158 pH where most anthocyanins would otherwise appear in red hues. Through selectivity of anthocyanin structure and metal ions, hues could be optimized for specific applications.

159

Bibliography

(CIE), C. I. de l’Eclairage. (2015). Selected Colorimetric Tables. Vienna, Austria. Retrieved from http://www.cie.co.at/index.php/LEFTMENUE/index.php?i_ca_id=298

Abdullah, R., Lee, P. M., & Lee, K. H. (2010). Multiple color and pH stability of floral anthocyanin extract: Clitoria ternatea. CSSR 2010 - 2010 International Conference on Science and Social Research, (09), 254–258.

Agency for Toxic Substances and Disease Registry (ATSDR). (2012). Chromium - ToxFAQs TM. ToxFAQs, (Iii), 1–2.

Ahmadiani, N. (2012). Anthocyanin Based Blue Colorants. The Ohio State University.

Ahmadiani, N., Robbins, R. J., Collins, T. M., & Giusti, M. M. (2014). Anthocyanins contents, profiles, and color characteristics of red cabbage extracts from different cultivars and maturity stages. Journal of Agricultural and Food Chemistry, 62(30), 7524–7531.

Ahmadiani, N., Robbins, R. J., Collins, T. M., & Monica Giusti, M. (2015). Molar Absorptivity (ε) and Spectral Characteristics of Cyanidin-Based Anthocyanins from Red Cabbage. Food Chemistry.

Ananga, A., Georgiev, V., Ochieng, J., Phills, B., & Tsolova, V. (2013). Production of Anthocyanins in Grape Cell Cultures : A Potential Source of Raw Material for Pharmaceutical , Food , and Cosmetic Industries. In D. Puljuha & B. Sladonja (Eds.), The Mediterranean Genetic Code - Grapevine and Olive (1st ed., pp. 247– 288). Rijeka, Croatia: InTech.

Andersen, Ø. M., & Jordheim, M. (2006). The Anthocyanins. In Ø. M. Andersen & K. R. Markham (Eds.), Flavonoids: Chemistry, Biochemistry, and Applications (pp. 472– 537). Boca Raton, FL: CRC Press (Taylor & Francis Group).

160

Andersen, Ø. M., & Jordheim, M. (2014). Basic Anthocyanin Chemistry and Dietary Sources. In T. C. Wallace & M. M. Giusti (Eds.), Anthocyanins in Health and Disease (1st ed., pp. 13–90). Boca Raton, FL: CRC Press (Taylor & Francis Group).

Andjelković, M., Van Camp, J., De Meulenaer, B., Depaemelaere, G., Socaciu, C., Verloo, M., & Verhe, R. (2006). Iron-chelation properties of phenolic acids bearing catechol and galloyl groups. Food Chemistry, 98(1), 23–31.

Azuma, K., Ohyama, A., Ippoushi, K., Ichiyanagi, T., Takeuchi, A., Saito, T., & Fukuoka, H. (2008). Structures and antioxidant activity of anthocyanins in many accessions of eggplant and its related species. Journal of Agricultural and Food Chemistry, 56(21), 10154–10159.

Bayer, E., Egeter, H., Fink, A., Nether, K., & Wegmann, K. (1966). Complex Formation and Flower Colors. Angewandte Chemie International Edition in English, 5(9), 791– 798.

Becaria, A., Campbell, A., & Bondy, S. C. (2002). Aluminum as a toxicant. Toxicology and Industrial Health, 18(7), 309–320.

Block, E. (2010). Garlic and Other Alliums, The Lore and the Science (1st ed.). Cambridge, United Kingdom: The Royal Society of Chemistry.

Bridle, P., & Timberlake, C. F. (1997). Anthocyanins as natural food colours—selected aspects. Food Chemistry, 58(1-2), 103–109.

Brown, T. L., LeMay (Jr.), H. E., Bursten, B. E., & Murphy, C. J. (2006). Chemistry, The Central Science. (N. Folchetti, J. Challice, P. Draper, & E. Al., Eds.) (10th ed.). Upper Sadle River, NJ: Pearson Education, Inc.

Browne, D. (2012). Breakfast cereal – US. Mintel Industry Report. London, UK.

Buchweitz, M., Brauch, J., Carle, R., & Kammerer, D. R. (2013a). Application of ferric anthocyanin chelates as natural blue food colorants in polysaccharide and gelatin based gels. Food Research International, 51(1), 274–282.

Buchweitz, M., Brauch, J., Carle, R., & Kammerer, D. R. (2013b). Colour and stability 161

assessment of blue ferric anthocyanin chelates in liquid pectin-stabilised model systems. Food Chemistry, 138(2-3), 2026–2035.

Buchweitz, M., Carle, R., & Kammerer, D. R. (2012). Bathochromic and stabilising effects of sugar beet pectin and an isolated pectic fraction on anthocyanins exhibiting pyrogallol and catechol moieties. Food Chemistry, 135(4), 3010–3019.

Buchweitz, M., Nagel, A., Carle, R., & Kammerer, D. R. (2012). Characterisation of sugar beet pectin fractions providing enhanced stability of anthocyanin-based natural blue food colourants. Food Chemistry, 132(4), 1971–1979.

Burrows, A. (2009). Palette of our palates: A brief history of food coloring and its regulation. Comprehensive Reviews in Food Science and Food Safety, 8(4), 394– 408.

Cabrita, L., Fossen, T., & Andersen, Ø. M. (2000). Colour and stability of the six common anthocyanidin 3-glucosides in aqueous solutions. Food Chemistry, 68(1), 101–107.

Clifford, M. N. (2000). Review Anthocyanins – nature , occurrence and dietary burden. Journal of the Science of Food and Agriculture, 80, 1063–1072.

Clydesdale, F. M. (1993). Color as a factor in food choice. Critical Reviews in Food Science and Nutrition, 33(1), 83–101.

D’Zmura, M. (1991). Color in visual search. Vision Research, 31(6), 951–966.

Dangles, O., Elhabiri, M., & Brouillard, R. (1994a). Kinetic and thermodynamic investigation of the aluminium-anthocyanin complexation in aqueous solution. Journal of the Chemical Society, Perkin Transactions 2, (12), 2587.

Dangles, O., Elhabiri, M., & Brouillard, R. (1994b). Kinetic and Thermodynamic Investigation of the Aluminium-Anthocyanin Complexation in Aqueous Solution. Journal of the Chemical Society, Perkin Transactions 2, (12), 2587–2596.

Dangles, O., Saito, N., & Brouillard, R. (1993a). Anthocyanin intramolecular copigment effect. Phytochemistry, 34(1), 119–124. 162

Dangles, O., Saito, N., & Brouillard, R. (1993b). Kinetic and Thermodynamic Control of Flavylium Hydration in the Pelagonidin-Cinnamic Acid Complexation. Origin of the Extraordinary Flower Color Diversity of Pharbitis nil. Journal of the American Chemical Society, 115(27), 3125–3132.

Di Meo, F., Sancho Garcia, J. C., Dangles, O., & Trouillas, P. (2012). Highlights on anthocyanin pigmentation and copigmentation: A matter of flavonoid ??-stacking complexation to be described by DFT-D. Journal of Chemical Theory and Computation, 8(6), 2034–2043.

Dimitrić Marković, J. M., Petranović, N. a., & Baranac, J. M. (2000). A spectrophotometric study of the copigmentation of Malvin with caffeic and ferulic acids. Journal of Agricultural and Food Chemistry, 48(11), 5530–5536.

Dimitrić Marković, J. M., Petranović, N. a., & Baranac, J. M. (2005). The copigmentation effect of sinapic acid on malvin: A spectroscopic investigation on colour enhancement. Journal of Photochemistry and Photobiology B: Biology, 78(3), 223–228.

Downham, A., & Collins, P. (2000). Colouring our foods in the last and next millennium. International Journal of Food Science and Technology, 35(1), 5–22.

Efsa. (2013). Scientific Opinion on the re-evaluation of anthocyanins ( E 163 ) as a food, 11(4), 1–51.

Elhabiri, M., Figueiredo, P., Toki, K., Saito, N., & Brouillard, R. (1997). Anthocyanin– aluminium and –gallium complexes in aqueous solution. Journal of the Chemical Society, Perkin Transactions 2, (2), 355–362.

Estévez, L., Otero, N., & Mosquera, R. a. (2011). Molecular structure of cyanidin metal complexes: Al(III) versus Mg(II). Theoretical Chemistry Accounts, 128(4), 485– 495.

FDA; CFSAN (Food and Drug Administration; Center for Food Safety and Applied Nutrition). (2011). Background Document 1. In Background Document for the Food Advisory Committee: Certified Color additives in Food and Possible Association with Attention Deficit Hyperactivity Disorder in Children (pp. 1–13).

163

Feingold, B. F. (1975). Hyperkinesis and Learning Disabilities Linked to Artificial Food Flavors and Colors. American Journal of Nursin, 75(5), 797–803.

Flaten, T. P., Alfrey, A. C., Birchall, J. D., & Yokel, R. A. (1996). Status and Future Concerns of Clinical and Environmental Aluminum Toxicology. Journal of Toxicology and Environmental Health, 48(6), 527–542.

Fossen, T., Cabrita, L., & Andersen, O. M. (1998). Colour and stability of pure anthocyanins influenced by pH including the alkaline region. Food Chemistry, 63(4), 435–440.

Frick, D. (2003). The coloration of food. Review of Progress in Coloration and Related Topics, 33(1), 15–32.

Gastineau, R., Turcotte, F., Pouvreau, J. B., Morancais, M., Fleurence, J., Windarto, E., … Mouget, J. L. (2014). Marennine, promising blue pigments from a widespread Haslea diatom species complex. Marine Drugs, 12(6), 3161–3189.

Ge, X., Timrov, I., Binnie, S., Biancardi, A., Calzolari, A., & Baroni, S. (2015). Accurate and Inexpensive Prediction of the Color Optical Properties of Anthocyanins in Solution. The Journal of Physical Chemistry A, 119(16), 3816–3822. Retrieved from http://pubs.acs.org/doi/abs/10.1021/acs.jpca.5b01272

Giusti, M. M., & Wrolstad, R. E. (1994). Separation and Characterization of Anthocyanins by HPLC. In R. E. Wrolstad, T. E. Acree, H. An, E. A. Decker, M. H. Penner, D. S. Reid, … P. Sporns (Eds.), Current Protocols in Food Analytical Chemistry (1st ed., pp. F1.3.1–F1.3.13). New York, NY: John Wiley & Sons, Inc.

Giusti, M. M., & Wrolstad, R. E. (2003). Acylated anthocyanins from edible sources and their applications in food systems. Biochemical Engineering Journal, 14(3), 217– 225.

Giusti, M., & Wrolstad, R. E. (2001). Characterization and Measurement of Anthocyanins by UV-Visible Spectroscopy. In R. E. Wrolstad, T. E. Acree, H. An, E. A. Decker, M. H. Penner, D. S. Reid, … P. Sporns (Eds.), Current Protocols in Food Analytical Chemistry (1st ed., Vol. 1, pp. F1.2.1–F1.2.13). New York, NY: John Wiley & Sons, Inc.

164

Glińska, S., Bartczak, M., Oleksiak, S., Wolska, A., Gabara, B., Posmyk, M., & Janas, K. (2007). Effects of anthocyanin-rich extract from red cabbage leaves on meristematic cells of Allium cepa L. roots treated with heavy metals. Ecotoxicology and Environmental Safety, 68(3), 343–350.

Gómez-Míguez, M., González-Manzano, S., Teresa Escribano-Bailón, M., Heredia, F. J., & Santos-Buelga, C. (2006). Influence of different phenolic copigments on the color of malvidin 3-glucoside. Journal of Agricultural and Food Chemistry, 54(15), 5422–5429.

González-Manzano, S., Santos-Buelga, C., Dueñas, M., Rivas-Gonzalo, J. C., & Escribano-Bailón, T. (2008). Colour implications of self-association processes of wine anthocyanins. European Food Research and Technology, 226(3), 483–490.

Hale, K. L., Tufan, H. a., Pickering, I. J., George, G. N., Terry, N., Pilon, M., & Pilon- Smits, E. a. H. (2002). Anthocyanins facilitate tungsten accumulation in Brassica. Physiologia Plantarum, 116(3), 351–358.

He, J., & Giusti, M. M. (2010). Anthocyanins: Natural Colorants with Health-Promoting Properties. Annual Review of Food Science and Technology - (New in 2010), 1(1), 163–187.

Houbiers, C., Lima, J. C., Maçanita, A. L., & Santos, H. (1998). Color Stabilization of Malvidin 3-Glucoside : Self-Aggregation of the Flavylium Cation and Copigmentation with the Z -Chalcone Form. J. Phys. Chem. B, 5647(97), 3578– 3585.

Jackman, R., & Smith, J. (1996). Anthocyanins and Betalains. In G. Hendry & J. Houghton (Eds.), Natural Food Colorants (2nd ed.). Great Britain: Blackie Academic & Professional.

Jespersen, L., Strømdahl, L. D., Olsen, K., & Skibsted, L. H. (2005). Heat and light stability of three natural blue colorants for use in confectionery and beverages. European Food Research and Technology, 220(3-4), 261–266.

Jurd, L., & Asen, S. (1966). The formation of metal and “co-pigment” complexes of cyanidin 3-glucoside. Phytochemistry, 5(6), 1263–1271.

165

Kaneko, Y., Thoendel, M., Olakanmi, O., Britigan, B. E., & Singh, P. K. (2007). The transition metal gallium disrupts Pseudomonas aeruginosa iron metabolism and has antimicrobial and antibiofilm activity. Journal of Clinical Investigation, 117(4), 877–888.

Kheng, L. W. (2002). Color Spaces and Color-Difference Equations. Color Research and Application, 24, 186–198.

Kobylewski, S., & Jacobson, M. F. (2010). Food Dyes: A Rainbow of Risks, (5), 58.

Koda, T., Ichi, T., Odake, K., Furuta, H., & Sekiya, J. (1992). Blue Pigment Formation by Clerodendron trichotomum callus. Bioscience, Biotechnology and Biochemistry, 56(12), 2020–2022.

Konczak, I., & Zhang, W. (2004). Anthocyanins — More Than Nature ’ s Colours. Journal of Biomedicine and Biotechnology, 5, 239–240.

Kondo, T., Toyama-Kato, Y., & Yoshida, K. (2005). Essential structure of co-pigment for blue sepal-color development of hydrangea. Tetrahedron Letters, 46(39), 6645– 6649.

Konica Minolta Inc. (2007). Precise color comunication. Minolta Co., 1–62.

Kunsági-Máté, S., Ortmann, E., Kollár, L., Szabó, K., & Nikfardjam, M. P. (2008). Effect of ferrous and ferric ions on copigmentation in model solutions. Journal of Molecular Structure, 891(1-3), 471–474.

Kuusi, T., Pyysalo, H., & Pippuri, A. (1977). The effect of iron, tin, aluminium, and chromium on fading, discoloration, and precipitation in berry and red beet juices. Zeitschrift Für Lebensmittel-Untersuchung Und -Forschung, 163(3), 196–202.

Lakshmi, G. C. (2014). Food Coloring: The Natural Way. Research Journal of Chemical Sciences, 4(2), 87–96.

Leatherhead Food Research and Mintel. (2013). New research reveals natural colours overtake artificial/synthetic colours for first time. Retrieved March 3, 2016, from https://www.leatherheadfood.com/new-research-reveals-natural-colours-overtake- 166

artificial-synthetic-colours-for-first-time

Li, H., Deng, Z., Zhu, H., Hu, C., Liu, R., Young, J. C., & Tsao, R. (2012). Highly pigmented vegetables: Anthocyanin compositions and their role in antioxidant activities. Food Research International, 46(1), 250–259.

Makris, D. P., & Rossiter, J. T. (2000). Heat-induced, metal-catalyzed oxidative degradation of quercetin and rutin (quercetin 3-O-rhamnosylglucoside) in aqueous model systems. Journal of Agricultural and Food Chemistry, 48(9), 3830–3838.

Malcıoğlu, O. B., Calzolari, A., Gebauer, R., Varsano, D., & Baroni, S. (2011). Dielectric and Thermal Effects on the Optical Properties of Natural Dyes: A Case Study on Solvated Cyanin. Journal of the American Chemical Society, 133(39), 15425– 15433.

Malien-Aubert, C., Dangles, O., & Amiot, M. J. (2001). Color stability of commercial anthocyanin-based extracts in relation to the phenolic composition. Protective effects by intra- and intermolecular copigmentation. Journal of Agricultural and Food Chemistry, 49(1), 170–176.

Mars Incorporated. (2016). Mars, Incorporated to remove all artificial colors from its human food portfolio. Retrieved March 2, 2016, from http://www.mars.com/global/press-center/press-list/news- releases.aspx?SiteId=94&Id=6984

Mason, R. P. (2013). Chemical thermodynamics and metal ( loid ) complexation in natural waters. In R. P. Mason (Ed.), Trace Metals in Aquatic Systems (1st ed., pp. 49–123). Chichester, West Sussex, U.K.: Blackwell Publishing (John Wiley & Sons, Ltd.).

Mateus, N., Oliveira, J., Haettich-Motta, M., & de Freitas, V. (2004). New Family of Bluish Pyranoanthocyanins. Journal of Biomedicine and Biotechnology, 2004(5), 299–305.

Mazza, G., & Brouillard, R. (1987). Recent developments in the stabilization of anthocyanins in food products. Food Chemistry, 25(3), 207–225.

167

McCann, D., Barrett, A., Cooper, A., Crumpler, D., Dalen, L., Grimshaw, K., … Stevenson, J. (2007). Food additives and hyperactive behaviour in 3-year-old and 8/9-year-old children in the community: a randomised, double-blinded, placebo- controlled trial. Lancet, 370(9598), 1560–1567.

McCleverty, J. (1990). Metals in Solution. In Chemistry of the First-Row Transition Metals (p. 90). New York, NY: Oxford University Press, Inc.

McDougall, G. J., Fyffe, S., Dobson, P., & Stewart, D. (2007). Anthocyanins from red cabbage--stability to simulated gastrointestinal digestion. Phytochemistry, 68(9), 1285–1294.

Mellican, R. I., Li, J., Mehansho, H., & Nielsen, S. S. (2003). The role of iron and the factors affecting off-color development of polyphenols. Journal of Agricultural and Food Chemistry, 51(8), 2304–2316.

Minioti, K. S., Sakellariou, C. F., & Thomaidis, N. S. (2007). Determination of 13 synthetic food colorants in water-soluble foods by reversed-phase high-performance liquid chromatography coupled with diode-array detector, 583, 103–110.

Mokrzycki, W. S., & Tatol, M. (2012). Colour difference ∆ E - A survey. Machine Graphic and Vision, 1–28.

Momonoi, K., Yoshida, K., Mano, S., Takahashi, H., Nakamori, C., Shoji, K., … Nishimura, M. (2009). A vacuolar iron transporter in tulip, TgVit1, is responsible for blue coloration in petal cells through iron accumulation. Plant Journal, 59(3), 437–447.

Moncada, M. C., Moura, S., Melo, M. J., Roque, A., Lodeiro, C., & Pina, F. (2003). Complexation of aluminum(III) by anthocyanins and synthetic flavylium salts: A source for blue and purple color. Inorganica Chimica Acta, 356, 51–61.

Montilla, E. C., Arzaba, M. R., Hillebrand, S., & Winterhalter, P. (2011). Anthocyanin composition of black carrot (Daucus carota ssp. sativus var. atrorubens Alef.) Cultivars antonina, beta sweet, deep purple, and purple haze. Journal of Agricultural and Food Chemistry, 59(7), 3385–3390.

168

Nayak, P. (2002). Aluminum: impacts and disease. Environmental Research, 89(2), 101– 115.

Nestlé USA. (2015). Nestlé USA Commits to Removing Artificial Flavors and FDA- Certified Colors from All Nestlé Chocolate Candy by the End of 2015. Retrieved March 2, 2016, from http://www.nestleusa.com/media/pressreleases/nestl%C3%A9- usa-commits-to-removing-artificial-flavors-and-fda-certified-colors-from-all- nestl%C3%A9-chocolate-candy-by-the-end-of-20

Newsome, A. G., Culver, C. a, & Breemen, R. B. Van. (2014). Nature’s Palette : The Search for Natural Blue Colorants. Journal of Agricultural and Food Chemistry, 62, 6498–6511.

Oliveira, J., Azevedo, J., Silva, A. M. S., Teixeira, N., Cruz, L., Mateus, N., & De Freitas, V. (2010). Pyranoanthocyanin dimers: A new family of turquoise blue anthocyanin-derived pigments found in port wine. Journal of Agricultural and Food Chemistry, 58(8), 5154–5159.

Oszmianski, J. a N., & Sapis, J. C. (1988). Anthocyanins in Fruits of Aronia Melanocarpa. Journal of Food Science, 53(4), 1241–1242.

Patras, A., Brunton, N. P., O’Donnell, C., & Tiwari, B. K. (2010). Effect of thermal processing on anthocyanin stability in foods; mechanisms and kinetics of degradation. Trends in Food Science & Technology, 21(1), 3–11.

Potera, C. (2010). The Artificial Food Dye Blues. Environmental Health Perspectives, 118(10), 428.

Pouvreau, J. B., Morançais, M., Massé, G., Rosa, P., Robert, J. M., Fleurence, J., & Pondaven, P. (2006). Purification of the blue-green pigment “marennine” from the marine tychopelagic diatom Haslea ostrearia (Gaillon/Bory) Simonsen. Journal of Applied Phycology, 18(6), 769–781.

Pyysalo, H., & Kuusi, T. (1973). The role of iron and tin in discoloration of berry and red beet juices. Zeitschrift Für Lebensmittel-Untersuchung Und -Forschung, 153(4), 224–233.

169

Rodriguez-Amaya, D. B. (2016). Natural food pigments and colorants. Current Opinion in Food Science, 7, 20–26.

Rodriguez-Saona, L. . E., Giusti, M. M., & Wrolstad, R. E. (1999). Color and Pigment Stability of Red Radish and Red-Fleshed Potato Anthocyanins in. Journal of Food Science, 64(3), 451–456.

Rodríguez-Saona, L. E., & Wrolstad, R. E. (2001). Extraction, Isolation, and Purifification of Anthocyanins. (R. E. Wrolstad, T. E. Acree, H. An, E. A. Decker, M. H. Penner, D. S. Reid, … P. Sporns, Eds.)Current Protocols in Food Analytical Chemistry (1st ed.). New York, NY: John Wiley & Sons, Inc.

Schreiber, H. D., Swink, A. M., & Godsey, T. D. (2010). The chemical mechanism for Al3+ complexing with delphinidin: A model for the bluing of hydrangea sepals. Journal of Inorganic Biochemistry, 104(7), 732–739.

Schwartz, S. J., von Elbe, J. H., & Giusti, M. M. (2008). Colorants. In S. Damodaran, K. L. Parkin, & O. R. Fennema (Eds.), Fennema’s Food Chemistry (4th ed., pp. 571– 639). Boca Raton, FL: CRC Press (Taylor & Francis Group).

Sever, M. J., & Wilker, J. J. (2004). Visible absorption spectra of metal – catecholate and metal – tironate complexes, 1061–1072.

Sharma, V., McKone, H. T., & Markow, P. G. (2011). A global perspective on the history, use, and identification of synthetic food dyes. Journal of Chemical Education, 88(1), 24–28.

Sigurdson, G. T., & Giusti, M. M. (2014). Bathochromic and Hyperchromic E ff ects of Aluminum Salt Complexation by Anthocyanins from Edible Sources for Blue Color Development. Journal of Agricultural and Food Chemistry, 62(29), 6955–6965.

Singh, S. (2006). Impact of color on marketing. Management Decision, 44(6), 783–789.

Slimestad, R., & Solheim, H. (2002). Anthocyanins from black currants (Ribes nigrum L.). Journal of Agricultural and Food Chemistry, 50(11), 3228–3231.

Smyk, B., Pliszka, B., & Drabent, R. (2008). Interaction between Cyanidin 3-glucoside 170

and Cu(II) ions. Food Chemistry, 107(4), 1616–1622.

Stintzing, F. C., Stintzing, A. S., Carle, R., Frei, B., & Wrolstad, R. E. (2002). Color and antioxidant properties of cyanidin-based anthocyanin pigments. Journal of Agricultural and Food Chemistry, 50(21), 6172–6181.

Tachibana, N., Kimura, Y., & Ohno, T. (2014). Examination of molecular mechanism for the enhanced thermal stability of anthocyanins by metal cations and polysaccharides. Food Chemistry, 143, 452–458.

Takeda, K., Fujii, T., & Iida, M. (1984). Magnesium in the blue pigment complex commelinin. Phytochemistry, 23(4), 879–881.

Ukwueze, N. N., Nwadinigwe, C. a, Okoye, C. O. B., & Okoye, F. B. C. (2009). Potentials of 3, 3, 4, 5, 7-pentahydroxyflavylium of Hibiscus rosa-sinensis L . ( Malvaceae ) flowers as ligand in the quantitative determination of Pb , Cd and Cr. International Journal of Physical Sciences, 4(2), 58–62.

Wallace, T. C., & Giusti, M. M. (Eds.). (2014). Anthocyanins in Health and Disease (1st ed.). Boca Raton, FL: CRC Press (Taylor & Francis Group).

Wild Flavors and Specialty Ingredients. (2016). Acid-Stable Blue. Erlanger, KY. Retrieved from https://www.wildflavors.com/NA- EN/assets/File/colors_library/ColorsfromNatureAcidStableBlue.pdf

Wrolstad, R. E., & Boyles, M. J. (1993). Anthocyanin composition of red raspberry juice: Influences of cultivar, processing, and environmental factors. Journal of Food Science, 58(5), 1135–1141.

Wrolstad, R. E., & Culver, C. A. (2012). Alternatives to Those Artificial FD&C Food Colorants. Annual Review of Food Science and Technology, 3, 59–77.

Wrolstad, R. E., & Smith, D. E. (2010). Color Analysis. In S. S. Nielsen (Ed.), Food Analysis (4th ed., pp. 573–586). New York, NY: Springer.

Wu, S., Ford, C., & Horn, G. (2009). Stable Natural Color Process, Products, and Use Thereof. United States. 171

Xu, Z. (2013). Mechanics of metal-catecholate complexes: the roles of coordination state and metal types. Scientific Reports, 3, 2914.

Yoshida, K., Kitahara, S., Ito, D., & Kondo, T. (2006). Ferric ions involved in the flower color development of the Himalayan blue poppy, Meconopsis grandis. Phytochemistry, 67(10), 992–998.

Yoshida, K., Mori, M., & Kondo, T. (2009). Blue flower color development by anthocyanins: from chemical structure to cell physiology. Natural Product Reports, 26(7), 884–915.

Zampini, M., Sanabria, D., Phillips, N., & Spence, C. (2007). The multisensory perception of flavor: Assessing the influence of color cues on flavor discrimination responses. Food Quality and Preference, 18(7), 975–984.

172