RED CABBAGE ANTHOCYANINS: HORTICULTURAL AND CHEMICAL FACTORS
AFFECTING COLOR CHARACTERISTICS OF CRUDE EXTRACTS, SELECT
PIGMENT MIXTURES, AND ISOLATED PIGMENTS
DISSERTATION
Presented in Partial Fulfillment of the Requirements for the Degree Doctor of Philosophy in the Graduate School of The Ohio State University
By
Neda Ahmadiani, M.Sc.
Graduate Program in Food Science and Technology
The Ohio State University
2015
Dissertation Committee:
Dr. M. Monica Giusti, Advisor
Dr. John Litchfield
Dr. Esperanza J. Carcache de Blanco
Dr. Lynn Knipe
© Copyrighted
by
Neda Ahmadiani
2015
All rights reserved
ABSTRACT
The food industry is looking for colorants from natural sources with increased stability, specific color properties, and pure color tones. Red cabbage (Brassica oleracea L.) is an excellent source of food colorant. Red cabbage anthocyanin (RCA) extract contains Cy anthocyanins in various non, mono, and diacylated forms capable of forming various shades of colors from red to blue. The overall objective of this study was to evaluate Red Cabbage
Anthocyanins (RCA), as potential alternatives to red and blue synthetic dyes. The specific objectives were to evaluated RCA colors, content and profiles in various cultivars and maturity stages plants, to develop SPE methods for isolation of the more stable diacylated pigments with different coloring and stability properties, and to determine the molar absorptivity (ε) of different RCA Cy-derivatives.
Anthocyanin concentrations ranged from 1,111 to 1,780 mg Cy3G/100gr DM and did not increase when plants stayed on the ground for longer time. Cultivar and maturation affected pigment profile. Some varieties accumulated ≥30% of di-acylated pigments, and proportions of mono-acylated pigments decreased with time. Extracts from selected varieties at first maturation stage produced colors similar (λmax 520nm and ∆E 6.1-8.8) to FD&C Red No.3 at pH
3.5. Extracts from cabbages with higher proportion of di-acylated pigments at second maturation stage produced color similar (λmax ≃610nm) to FD&C Blue No.2 at pH 7.
Simple SPE methods were developed to segregate diacylated anthocyanins with higher stabilities and better blue coloring properties. When SCX cartridge was used, washing step with
ii buffer pH6+20% MeOH followed by eluting step using buffer pH8+70% MeOH was effective in segregating the target diacylated pigments; however, some anthocyanins permanently retained by the resin. When using the C18 cartridge, washing with 32% MeOH or
18.5% EtOH and eluting with 100% alcohol respectively resulted in high (> 93%) recovery of diacylated pigments. The C18 MeOH combination was the most effective for isolating the target diacylated RCA and produced a better blue color with increased stability.
Knowing the molar absorptivity (ε) of anthocyanins helps with more accurate concentration measurements and provides information about the color strength. The ε of different Cy-based anthocyanins in acidified MeOH and buffer pH1 ranged between 16,000-
30,000 and 13,000-26,000 L/mol.cm, respectively. Higher ε values were obtained for non- acylated pigments, and decreased with increased acylation. Most pigments showed higher ε at pH8 than pH2, and lowest ε between pH 4-6. There were bathochromic shifts (81-105nm) from pH 1-8 and hypsochromic shifts from pH 8-9 (2-19nm). Position and number of acylating groups were critical factors that affected the ε and the spectral behaviors.
Cultivar selection and maturation affected color and stability of red cabbage extracts at different pH values due to their different anthocyanin pigment profiles. The di-acylated pigments with higher stabilities and pure blue color tones were successfully segregated using a
C18 SPE fractionation method. Matrix and the molecular structure of the red cabbage Cy-based anthocyanin affected their ε and spectral characteristics.
Overall, RCA showed potential as suitable alternatives to red and blue synthetic dyes, depending on the application (pH) and the acylations present and can be fractionated to produce better red and blue hues similar to commercial colorants. This study provides a valuable contribution to the use of RCA as colorants for foods.
iii
In dedication to
Marçal and Adrià for bringing joy and happiness to my life
iv ACKNOWLEDGEMENTS
First and foremost, I would like to express the deepest appreciation to my advisor, M.
Monica Giusti for her valuable guidance and advice. I can't thank her enough for the tremendous support and help she provided me. I felt motivated and encouraged throughout the past five years working for her. She inspired me greatly to work on this project. Fulfillment of this research would have been impossible without her willingness to share her extensive knowledge about anthocyanins.
I would also like to thank my committee members Dr. John Litchfieldand, Dr.
Esperanza J. Carcache de Blanco, and Dr. Lynn Knipe for their advice and help with my research, my candidacy exam, and my dissertation. I would also like to take this opportunity to truly thank Greg Sigurdson for sharing with me a CIE-L*a*b* color conversion software that he put extensive effort to improve and being such a great lab manager for our lab. Likewise, I would like to thank my other current lab mates Fei Lao, Peipei Tang, Alexandra Westfall,
Yachen Zhang, and Jacob Farr for sharing their findings in anthocyanin field with me and their continued moral support. I also wish to thank my former lab mate Dr. Allison Atnip, Dr. Kom
Kamonpatana, and Steven Simmons who provided training and assistance for me to learn about anthocyanin purification and analysis techniques using HPLC-MS.
I am deeply thankful to the entire Food Science & Technology department faculty for providing me with the foundation of knowledge necessary for me to achieve my academic goals. Also I am grateful to the staffs in this department who facilitate my study and research.
v Additionally, I would like to express my sincere thanks and gratitude to D. Rebecca J.
Robbins and Dr. Thomas M. Collins from Mars Chocolate for their continuous and valuable feedbacks and suggestion about my project.
I would like to specially thank Mars Chocolate NA for their financial support of this project, and for providing samples to carry out the experiments. I will never be able to thank them enough for providing me with the opportunity to pursue a graduate degree in Food
Science.
Finally, an honorable mention goes to my family, especially my mother, and friends for their never ending love and support despite being far away. Without them the completion of this project would be difficult for me.
vi VITA
August 18, 1978…………………………………………………………...Born, Kermanshah, Iran
2001…………………………………………..B.S. (Audiology), Shahid Beheshti University, Iran
2012………...…………M.Sc. (Food Science and Technology), The Ohio State University, U.S.A
2012-2015………………..OSU Food Science and Technology, The Ohio State University, U.S.A
PUBLICATION
1. Ahmadiani, N; Robbins, R; Collins, T; Giusti, MM. 2014. Anthocyanins contents, profiles and color characteristics of red cabbage extracts from different cultivars and maturity stages. J of Agric and Food Chem 62(30):7524-31. 2. Giusti, MM; Ahmadiani. N; Tang, P; Ottinger, M.A.. Isoflavone and Flavonoid Supplemented Eggs in Health. In: Watson, R. R. and De Meester, F., editors. Handbook of Eggs in Human Function. Wageningen Academic Publishers. P. 333-364. 3. Robbins, RJ; Collins, TM; Giusti, MM; Ahmadiani, N, inventors. 2013. Natural blue anthocyanin-containing colorants from red cabbage. U.S. Patent WO2014152478 A2. 4. Robbins, RJ; Collins, TM; Giusti, MM; Ahmadiani, N, inventors. 2013. Method of Isolating Blue Anthocyanin Fractions. U.S. Patent WO2014152417 A2. 5. Robbins, RJ; Collins, TM; Giusti, MM; Ahmadiani, N, inventors. 2013. Natural Blue Anthocyanin-Containing Colorants. Filed Feb 2013
FIELDS OF STUDY
Major Field: Food Science and Technology
vii
TABLE OF CONTENTS
ABSTRACT ...... ii ACKNOWLEDGEMENTS ...... v VITA ...... vii TABLE OF CONTENTS ...... viii LIST OF TABLES ...... xii LIST OF FIGURES ...... xiv LIST OF ABBREVIATIONS...... xvii CHAPTER 1: INTRODUCTION ...... 1 CHAPTER 2: REVIEW OF LITERATURE ...... 4 2.1. COLOR IN FOOD ...... 4 2.1.1. History of color use in food ...... 5 2.1.2. What are food colorants? ...... 6 2.2. CERTIFIED COLORANTS ...... 7 2.2.1. Types of synthetic certified colorants ...... 8 2.2.2. Reasons for use ...... 10 2.2.3. Limitation of use ...... 11 2.3. COLORS EXEMPT FROM CERTIFICATION ...... 12 2.3.1. Naturally occurring pigments for food use ...... 13 2.3.2. Future trends in natural colors ...... 17 2.4. BLUE COLORANTS ...... 17 2.4.1. Blue color perception ...... 17 2.4.2. Synthetic FD&C blue colorant ...... 19 2.4.3. Alternatives for blue colorants from natural sources ...... 20 2.5. ANTHOCYANINS ...... 22
viii 2.5.1. Chemical structure ...... 23 2.5.2. Health benefits ...... 24 2.5.3. Anthocyanins as food colorants ...... 25 2.5.4. Chemical characterization: ...... 27 2.5.5. Factors affecting the color and blue color formation of anthocyanins ...... 32 2.6. RED CABBAGE ...... 40 2.6.1. Taxonomy ...... 40 2.6.2. Growth location ...... 41 2.6.3. Nutritional composition ...... 41 2.6.4. Production for processing purposes ...... 42 2.6.5. Red cabbage color ...... 42 2.7. POTENTIAL FACTORS AFFECTING ANTHOCYANINS ACCUMULATIONS IN PLANTS ...... 43 2.7.1. Light ...... 43 2.7.2. Nutrients ...... 44 2.7.3. Maturity ...... 45 CHAPTER 3: ANTHOCYANINS CONTENTS, PROFILES AND COLOR CHARACTERISTICS OF RED CABBAGE EXTRACTS FROM DIFFERENT CULTIVARS AND MATURITY STAGES ...... 46 3.1. ABSTRACT ...... 46 3.3. MATERIALS AND METHODS ...... 48 3.3.1. Plant Materials ...... 48 3.3.2. Extraction and Purification ...... 49 3.3.3. Anthocyanins content ...... 50 3.3.4. Alkaline hydrolysis of anthocyanins ...... 50 3.3.5. Chromatographic analysis ...... 51 3.3.6. Color and spectrophotometric analyses ...... 51 3.3.7. Statistical analysis ...... 52 3.4. RESULTS AND DISCUSSIONS ...... 53 3.4.1. HPLC-PDA-MS and identification of major pigments ...... 53 3.4.2. Anthocyanin content and proportion of the pigments ...... 55 3.4.3. Color characteristics and stability ...... 60 3.4.3.1. Color in acidic pH ...... 60 3.5. CONCLUSION ...... 66
ix 3.6. ACKNOWLEDGMENT...... 66 CHAPTER 4: SOLID PHASE FRACTIONATION TECHNIQUES FOR SEGREGATION OF RED CABBAGE ANTHOCYANINS WITH DIFFERENT COLORING AND STABILITY PROPERTIES , ...... 67 4.1. ABSTRACT ...... 67 4.2. INTRODUCTION ...... 68 4.3. MATERIALS AND METHODS ...... 70 4.3.1. Sample preparations ...... 70 4.3.2. SPE fractionation ...... 71 4.3.3. HPLC-MS and chromatographic analysis of the fractions ...... 73 4.3.4. Calculation of R530 recovery and anthocyanin yield ...... 74 4.3.5. Measurement of spectra and degradation of PA, R520 and R530 pigments... 74 4.3.6. Conversion of spectral data to color ...... 75 4.3.7. Color analysis of the fractions ...... 76 4.3.8. Statistical analysis ...... 77 4.4. RESULTS AND DISCUSSIONS ...... 77 4.4.1. Spectra and color analysis ...... 77 4.4.2. Pigments degradations ...... 81 4.4.3. SCX-buffer fractionation of RCA ...... 82 4.4.4. C18-MeOH and EtOH fractionation of RCA ...... 86 4.4.5. Color analysis after fractionation ...... 90 4.5. CONCLUSIONS ...... 92 4.6. ACKNOWLEDGMENT...... 93 CHAPTER 5: MOLAR ABSORPTIVITY (ε) AND SPECTRAL CHARACTERISTICS OF CYANIDIN-BASED ANTHOCYANINS FROM RED CABBAGE, ...... 94 5.1. ABSTRACT ...... 94 5.2. INTRODUCTION ...... 95 5.3. MATERIALS AND METHODS ...... 97 5.3.1. Plant materials and anthocyanins extraction ...... 97 5.3.2. Anthocyanins semi-purification ...... 97 5.3.3. Chromatographic analysis ...... 98 5.3.4. Measurement of ε and Spectral Characteristics ...... 99 5.3.5. Stability comparison between mono and di-acylated Cy- derivatives ...... 100 5.3.6. Statistical analysis ...... 101
x 5.4. RESULTS AND DISCUSSIONS ...... 101 5.4.1. Major Cy based anthocyanins from red cabbage ...... 101 5.4.2.Molar absorptivity and spectral characteristics in MeOH and buffer pH1 .... 103 5.4.3. Molar absorptivity, spectral characteristics, and color in buffers pH 1-9 ..... 108 5.4.4. Stability comparison between mono and di-acylated Cy- derivatives ...... 111 5.5. CONCLUSIONS ...... 113 5.6. ACKNOWLEDGMENT...... 113 OVERALL CONCLUSION ...... 114 BIBLIOGRAPHY ...... 115 APPENDIX A: BUFFERS PREPARATION ...... 128
xi LIST OF TABLES
Table Page
Table 2.1: List of certified FD&C dyes, their color characterization, and use (Sharma et al. 2011; Frick 2003)...... 8 Table 2.2: Important property differences and use of dyes and lakes (Dziezak 1987)...... 10 Table 2.3: Stability of FD&C approved colorants under different environmental conditions (Pintea 2007; Francis 2002)...... 12 Table 2.4: Major types of natural food pigments found among the list of colorants exempt from certification; their major source, properties, and use in foods (Simpson et al. 2012; Chapman 2011; 21CFR73)...... 15 Table 2.5: Color characteristics of two different anthocyanidins (Pg and Cy) and their glycosides with different concentrations in acidic pH values (Giusti and Wrolstad 1996; Torskangerpoll and Andersen 2005)...... 35 Table 2.6: Effect of the addition of one sugar molecule to the Cy anthocyanins on its color characteristics at selected pH values (Torskangerpoll and Andersen 2005)...... 35 Table 2.7: Effect of the concentration of the Cy3-(2-glcglc)-5-glc on the color characteristics at different pH values extracted or modified (Torskangerpoll and Andersen 2005) ...... 36 Table 2.8: Effect of the addition of an acyl group to a glycosylated Cy anthocyanin on the color characteristics of the solution at different pH values (Torskangerpoll and Andersen 2005)...... 37 Table 3.1: The PDA absorbance and the MS data for the RCA. 287 m/z was the major fragment + in all 8 peaks. RT: Retention time; λvis: λ vis-max; λacyl: λ of acylation; M : Mass ion...... 54 Table 3.2: Anthocyanin contents (total monomeric anthocyanin) and percentage of major pigments (% total peak area at 510-540 nm) in seven red cabbage cultivars in two different harvesting times. Different letters in the same column indicate significant differences (p < 0.05); DM: dry matter; FM: fresh matter; non-acylated pigments:
xii peak 1; mono-acylated pigments: peak 2, 3, 4, and 5; di-acylated pigments: peak 6, 7, and 8 (See Table 3.1 for peak identity)...... 58 ◦ Table 3.3: C.I.E. L*a*b*, Chroma (c*), Hue (h ), and λmax of seven red cabbage cultivars extracts at two different harvesting time at pH 3.5 and their comparison with two FD&C
synthetic red dyes. HT: Harvesting time; ∆E1: color difference with FD&C Red No. 3;
∆E2: color difference with FD&C Red No. 40...... 61 Table 3.4: λmax of the red cabbage extracts at two different harvesting times measured after 30min and 6hr refrigeration storage in buffer pH 7compared to FD&C synthetic Blue No. 2, and stabilities (% degradation) during this time. HT: Harvesting time. %Degradation= 100-((absorbance at the λmax after 6 hr/ absorbance at the λmax after 30 min) x100)...... 64 Table 3.5: C.I.E. L*a*b*, Chroma (c*), and Hue (h◦) of seven red cabbage extracts at two different harvesting time after 6 hr refrigeration storage at pH 7 compared to FD&C Blue No. 2. HT: Harvesting time; ∆E: color difference with FD&C Blue No. 2...... 65 Table 4.1: Solvents used in each fractionation method using SCX (S1-S5) and C18 (C1-C6) resins...... 73 Table 4.2: The MS data of the RCA pigments (for the HPLC chromatogram refer to Figure 2). Cy-3diG-5G: Cyaniding-3-diglucoside-5-glucoside. M+: Mass ion...... 78 Table 4.3: Color and spectral characteristics of non-, mono- , and diacylated Cy-3diG-5G from red cabbage at different pH values and their color difference with synthetic red and blue FD&C colorants. Spectral data obtained after 15 min and 8 h at room temperature storage. For the pigments identities refer to Table 4.2...... 80 Table 4.4: Proportion of R530 pigment, their recovery, and yield of anthocyanins in each fraction obtained using method C1-C6 (Table 4.2). %R530 (R530 peaks proportions to the total anthocyanin peaks) obtained from the HPLC chromatograms. For the chromatogram of each fraction refer to Figure 4.5, 4.6, and 4.7...... 90 Table 4.5: Color and spectral characteristics of fractions collected using method C4 compared to the RCA in pH2 and 8...... 91 Table 5.1: ε (L cm-1 mol-1) of the Cy derivatives from red cabbage 1hr after dissolved in acidified MeOH and buffer pH 1(25°C)...... 105
Table 5.2: Molar absorptivity (ε) (L cm-1 mol-1) and maximum absorbance (λmax) of the of the Cy derivatives from red cabbage 1hr after dissolved in pH 1-9 (25°C). Final concentrations were 1.02 x10-04mol/L and 7.50 x 10-05mol/L for peak 1-2 and 3-8, respectively. For peak identity and retention times refer to Figure 5.1...... 108
xiii LIST OF FIGURES
Figure Page
Figure 2.1: Chemical structures of four major categories of synthetic dyes...... 9 Figure 2.2: Blue color perception for a blue pigment and the properties of a red light absorbing molecule using the visible absorbance spectrum of FD&C Blue No.1 solution (λmax= 630nm) as an example (Newsome et al. 2014)...... 19 Figure 2.3: Basic structure of the six major anthocyanins found frequently in the nature (He and Giusti 2010)...... 23 Figure 2.4: Example of color variation of anthocyanins. red radish: RR, red cabbage: RC, purple sweet potato: PSP, blueberry: BB, elderberry: EB...... 26 Figure 2.5: Structural changes in anthocyanins with change in the pH (Stintzing et al. 2002; Wrolstad et al. 2001; Fleschhut et al. 2006)...... 33 Figure 2.6: λmax of six common anthocyanidin-3-glucosids (with different 3' and 5' substitutions) at different pH values 1h after dissolution in buffers at room temperature (Cabrita et al. 2000)...... 34 Figure 2.7: Intramolecular stacking of diacylated anthocyanins above and below the flavylium molecule...... 38 Figure 2.8: Intermolecular copigmentation of anthocyanins: selfassociation (top), copigmentation with a cinnamic acid (bottom)...... 39 Figure 3.1: HPLC chromatograms of two representative red cabbage anthocyanin extracts: second harvested Cairo (top) and first harvested Integro (bottom) red cabbage at 510- 540 nm: refer to Table 1 for major peaks identification. HPLC conditions: solvent A, 4.5% formic acid in LCMS grade water; B, LCMS acetonitrile; gradient: 0-50 min, 0- 30% B. The major peaks (1-8) were found in all other red cabbage samples...... 54 Figure 3.2: Correlation between the first two principal components and the variables, as well as, score plot with respect to cultivors and harvesting time for seven different red cabbage cultivars. a: total monomeric anthocyanin; b: di-acylated pigments; c: mono- acylated pigments; d: non-acylated pigments. Az: Azurro; Ba: Bandolero, Bu:
xiv Buscaro; Ca: Cairo; In: Integro; Ko: Kosaro; Pr: Primero; Symbols in black and gray represent harvest times after 13 or 21 weeks respectively...... 56 Figure 3.3: Change in λmax for the 2nd harvested Buscaro, Bandolero, and Primero at pH 7 over 72 hr refrigeration. Most color changes happened during the first 6hr of storage. ... 62 Figure 4.1: Molecular structure of Cy-3diG-5G, and, percent degradations of non, mono, and diacylated RCA in pH 1-8 after 8 hr. PA:R1=H, R2= H; R520:; R530: R1= acyl, R2= acyl. Acyl groups were cinnamic acids (p-Coumaric, Ferulic, and Sinapic). For the pigments identities refer to Table 4.2. No significant difference was observed up to pH 3 (p > 0.05). % degradation= 100-((absorbance at λmax after 8 hr/absorbance at λmax after 15min) x 100)...... 82 Figure 4.2: Chromatographic separation of different fractions collected using the SCX cartridge and buffers (methods S1 and S2). For the solvent used to isolate each fraction refer to Table 4.1. Identification of each individual peak is shown in Table 4.2. Samples were conditioned with pure buffers prior to collecting each fraction...... 84 Figure 4.3: HPLC chromatograms of the fractionations obtained using SCX cartridge and buffer pH8 and different percentage of MeOH. For the solvent used to isolate each fraction refer to Table 4.1. Sample was conditioned with pure buffer pH8 before collecting the S3-F1 fraction...... 85 Figure 4.4: HPLC chromatograms of method S4 and S5 using SCX cartridge when the conditioning steps using pure buffers were skipped. For the solvent used to isolate each fraction refer to Table 4.1...... 86 Figure 4.5: Chromatographic separation of the three fractions collected using C18 cartridge and MeOH (left) and EtOH (right) in various percentage (method C1 and C2). For the solvent used to isolate each fraction refer to Table 4.1. Conditioning step using pure buffer before collecting each fraction was skipped...... 87 Figure 4.6: HPLC chromatograms of the two fractions collected using C18 cartridge and MeOH in various percentages (method C3 and C4). For the solvent used to isolate each fraction refer to Table 4.1...... 88 Figure 4.7: HPLC chromatograms of the two fractionations obtained using C18 cartridge and EtOH in various percentages (method C5 and C6). For the solvent used to isolate each fraction refer to Table 4.1...... 89 Figure 4.8: Absorbance spectra of the C4-MF1, C4-MF2, and RCA at pH 8 compared to FD&C Blue No. 2 synthetic dye...... 92
xv Figure 5.1: HPLC chromatogram and identity of the major RCA. The major fragment in all 8 peaks was 287 m/z corresponding to Cy3diG-5-G. RT: Retention time; M+: Mass ion (Ahmadiani et al. 2014) ...... 103 Figure 5.2: Normalized UV-vis spectra of a: Cy3diG-5-G (PK1), Cy3diG-5-G+p-Coumaric (PK3), Cy3diG5-G+Ferulic (PK4), Cy3diG-5-G+Ferulic&Ferulic (PK6), and Cy3diG-5-G+Sinapic&Ferulic (PK7); b: Cy3diG-5-G+Sinapic (PK2 and PK5) and Cy3diG-5-G+Sinapic&Sinapic (PK8); c: pure p-Coumaric, Ferulic, and Sinapic acids in MeOH (0.1% HCl)...... 107 Figure 5.3: Color changes of the major Cy based pigments from red cabbage 1hr after dissolved (25°C) in buffer pH 1-9...... 110 Figure 5.4: Visible spectra of a: Cy3diG-5-G (PK1); b: Cy3diG-5-G+Sinapic (PK2); c: Cy3diG- 5-G+Sinapic (PK5); and d: Cy3diG-5-G+Sinapic&Sinapic (PK8) in buffers pH 2, pH 5, pH 7, and pH 9...... 111 Figure 5.5: Spectral degradation comparison of PK5 (6.33x10-04 mol/L) and PK8 (4.86x10-4 mol/L) in pH 1-8 (25°C) stored in dark for 8hr (results were significant at p-value <0.05)...... 112
xvi LIST OF ABBREVIATIONS
ADI Acceptable Daily Intake
AOAC Association of Analytical Communities
Az Azurro
Ba Bandolero
Bu Buscaro
C* Chroma
Ca Cairo
CFR Code of Federal Regulation
Cy cyanidin
Cy3diG-5-G Cyanidin-3-diglucoside-5-glucoside
DM Dry matter
Dp delphinidin
EtOH Ethanol
FDA Food and Drug Administration
FM Fresh matter
GRAS Generally Recognized as Safe h Hours h◦ Hue
HPLC High Pressure Liquid Chromatography
xvii In Integro
Ko Kosaro
L* Lightness
MeOH Methanol
MS Mas Spectroscopy
Mv malvidin
Pg Pelargonidin
Pn peonidin
Pr Primero
Pt petunidin
RCA Red Cabbage Anthocyanins
USDA Department of Agriculture
ε Molar absorptivity or Absorption coefficient
λmax Absorption Maxima
xviii CHAPTER 1: INTRODUCTION
Consumers are decreasing their use of food containing artificial colorants due to possible link between the consumption of certain dyes to behavioral problems in children and allergenicity. In the U.S., color additives regulated under the decision of the Food and Drug
Administration (FDA). Currently, the European Union is requiring foods that contain certain synthetic colorants to carry a label that warns about the possible adverse effect of these dyes; however, such warning labeling is not required in the U.S. Although, evaluating the safety of the synthetic dyes by the FDA is still under investigation, and, the food industry is proactively looking for coloring alternatives from natural sources. In addition to their visual appeal, interest in natural colorants has increased due to their functionality and potential health benefits.
Anthocyanin pigments are found in a wide variety of edible fruits and vegetables and have numerous potential health benefits. Because of their water solubility and capability to form a wide shade of color from orange to red to blue and green, they are good alternatives for coloring foods. Moreover, they are permitted in the U.S. as food colorant in the category of fruit and vegetable juice or grape skin extract. Currently anthocyanins are being used as alternatives to many synthetic dyes in many food and beverages. Their use however, is limited mainly due to their relatively low stability, pH sensitivity, formulation, possible undesirable contribution to the odor and flavor of the food, challenges with obtaining a target pure shade (free from other mixed color tones) of color, and their low color strength compared to FD&C dyes.
1 Red cabbage (Brassica oleracea L.) has high content and high potential yield per unit area of anthocyanins. Red cabbage anthocyanin extract is often used by the food industry as an alternative for synthetic food colorants because of two main reasons: their good stabilities to light and processing temperature and being able to exhibit a wide spectrum of color based on the pH of the media. This extract is a rich source of cyanidin (Cy) based anthocyanins (Cy3diG-
5-G) with various mono and di-acylating groups. Type and acylation of anthocyanins are two important factors that determine their color characteristics at certain pH values. Acylation of anthocyanis is known to cause bathochromic shift (bluing effect) in the color. It also influences the antioxidant properties and stabilities. Di-acylated anthocyanins are known to have higher stability and higher antioxidant activity compared to the other non and mono acylated ones.
Initially we were interested in the effect of red cabbage anthocyanin profiles and their effect on the overall color characteristics. There are several known intrinsic and extrinsic factors which affect anthocyanin content and composition in plants. Plants cultivar and maturation time are among the essential factors that can influence the phytochemical content, among which anthocyanins. In chapter 3, we evaluated the anthocyanin pigment contents and profiles from 7 red cabbage cultivars at two maturity stages (8 weeks apart) and evaluated their color characteristics and behavior under acidic and neutral pH. Knowing the anthocyanin composition of red cabbage cultivar and maturation time would help us to select the cultivar and maturation that could provide the desired color characteristics for a specific application.
In chapter 4, our research aim was to segregate the diacylated RCA with higher stabilities and more pure blue color tones using simple SPE fractionation techniques. Solid phase extraction (SPE) is a simple and affordable separation technique widely used for isolation and concentration of compounds. It has also been used for isolation of anthocyanins. We explored two different cartridges with SCX (Strong Cation Exchange) and C18 sorbents. The
SCX cartridge was used with combination of different buffers (pH 6-8) and MeOH (20-70%);
2 and, the C18 cartridge was used with acidified (0.01% HCl) MeOH (30-100%) and EtOH (10-
100%). With this study we were able to develop a simple, effective, and economic fractionation method to isolate the diacylated RCA pigments with similar coloring properties.
Finally in chapter 5, we assess the color strength and characteristics of different red cabbage Cy-derivatives in different environments by evaluating the molar absorptivity (ε) and spectral behaviors of the major pigments in acidified MeOH and buffers pH 1-9. There is need for newly obtained information about the ε value of the anthocyanins in the literature because these data help with more accurate concentration measurement and provides information about the tinctorial strength of the pigments.
3 CHAPTER 2: REVIEW OF LITERATURE
2.1. COLOR IN FOOD
Color is an important sensory attribute of food which makes it more appealing, attractive, and appetizing for the consumers. Color has a great influence on flavor perception and often perceived prior to aroma. If a food doesn’t look appealing to the eye, it may never be tasted (Walford 1980). In addition, color of the food enables us to identify the product such as candy flavor. Food processing often is one of the main reasons for color and flavor loss, so, addition of color additive plays an important role in the acceptability of the food product after processing (Walford 1980; Frick 2003). There are also other reasons for food coloration such as when foods have no natural color of their own; or, their natural color varies with season and geographic region (Frick 2003).
Food coloration often is very challenging compared to other color application and it requires a great knowledge in food science, engineering, marketing, and law, along with artistic skills (Frick 2003). There are several criteria that need to be considered when coloring our food.
Essentially, the food scientist needs to have sufficient information about the existing coloring compounds (natural and synthetics), their properties, chemical compositions, applications, limitations, and regulatory status. In addition, since foods are very complex matrix, knowledge
4 of the other food ingredients and their interactions among themselves and with the coloring compound is very essential because these interactions may cause discoloration. These interactions are often more prone to happen during processing, consequently, excessive knowledge of chemistry and food processing is also very important for proper selection of food coloring agent (Frick 2003). Finally, being aware of consumers’ demands and marketing trends are very critical for the proper selection of a coloring agent for a specific food product.
2.1.1. History of color use in food
Hundreds of years have passed since the first time that humans started coloring their
food. The addition of colorants to food has been traced back to 1500 BC when candy makers
added natural color extract and wine to their products in Egypt. Naturally derived colorants such
as saffron were normally used to color foods by mid-19th century (Meggos 1995; Downham and
Collins 2000).
Following the industrial revolution and the development of the food industry the
practice of adding color to the “processed food” became more common. During the same era (in
1856), the first synthetic organic dye was discovered by William Henry Perkin called mauve.
The food industry offered a vast array of synthetic colors by the end of the 19th century. At the
time, food colorants were used for decorative purpose, and, sometimes, unfortunately, to cover
low quality food products. Mineral and metal based compounds were the main coloring agents
used to disguise low quality foods; some striking examples include red lead (Pb3O4) and
vermillion (HgS) used to color confectionery and cheese products (Meggos 1995; Downham
and Collins 2000). The uses of some of the unsafe chemicals for food coloring purpose caused
food poisoning and even death back in 1860. By the late 1800, the use of unauthorized colorants
in various food products such as ketchup, mustard, jellies, and wine spread through the U.S. and
5 Europe. Although colors from plants, animals, and minerals were still available, bulk synthesis of chemically derived colorants from toxic materials such as aniline (petroleum based prototypical aromatic amine) became common in 1900s. That is mainly because chemically synthesized colorants were easier to produce, less expensive, had better coloring properties, and didn’t contribute to the flavor of the food products. Increased use of these colorants, however, started to raise safety concerns and so did the stricter regulation (Downham and Collins 2000).
Certification of color in the U.S. was originated in 1906 by the Pure Food and Drug Act as a non-mandatory practice regulated by the U.S. Department of Agriculture (USDA) to acknowledge that each batch of color synthesized conformed to the specifications (Dziezak
1987). This process, however, became mandatory as a result of the Federal Food, Drug, and
Cosmetic Act of 1938. The responsibility of color regulation also was transferred to the Federal
Food and Drug Administration (FDA) at the same time. As a result of 1960 Color Additive
Amendment, a list of those color additives that needed to be certified and the ones exempt from certification was provided (Dziezak 1987; Pintea 2007). In 1993, the Redbook of Colorants was published by the FDA which states the toxicological principles for safety assessments of color additives used in foods (Pintea 2007).
2.1.2. What are food colorants?
According to the 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 (alone or through
reaction with another substance) of imparting a color thereto.” (21CFR70.3)
6 Food colorants are overseen by the FDA Code of Federal Regulation (CFR) 21, under parts 70, 71, 73, 74, 80, and 82. There is no Generally Recognized as Safe (GRAS) exemption to the definition of a color additive because FDA states that a substance that imparts color is a color additive and is subjected to premarket approval otherwise it is used only for a purpose other than coloring. However, a GRAS ingredient can be used to color food only if it is also listed as approved colorant ; for example, ferrous lactate, used for the coloring of ripe olives (21
CFR184.1311 and 21 CFR73.165). There are two major classifications for color additives:
“certified colors” and “colors exempt from certification” (Barrows 2003).
2.2. CERTIFIED COLORANTS
Certified colors are made through chemical synthesis to a high degree of purity, and, they are often in the form of powder, granules, dry blends, and wet-dry blends (Walford 1980).
This type of color additives are currently synthetic organic dyes, lakes, or pigments that are synthesized mainly from raw materials obtained from petroleum. They are, however, referred to as coal-tar colors because of their traditional origins (Barrows 2003). In the U.S., the process of the synthesis of these color additives has to be disclosed to the FDA on the confidential basis.
FDA is in charge of regulating this type of colorants by publication of permitted list
(21CFR.74), checking the purity, and sometimes by setting the level of use of permitted colorant in certain food products. FDA checks each batch of color from the starting materials throughout the processing for compliance with their identity and purity specifications of the final products by applying common analytical tests and assay, as well as, the use of chromatographic techniques (Walford 1980; Barrows 2003).
Certified food colors must be identified by their name on the food’s label. The entire legal name, however, can be shortened, for instance, FD&C Blue No.1 may be labeled as Blue
7 1. It also needs to be specified if the lake form of the color is used, for example Blue 1 lake
(Frick 2003).
2.2.1. Types of synthetic certified colorants
There are currently 9 color additives approved for use in food in the U.S. with two
having specific application (21CFR74). Table 2.1 contains the list of the certified color
additives approved by the FDA and their specifications.
Table 2.1: List of certified FD&C dyes, their color characterization, and use (Sharma et al. 2011; Frick 2003).
Absorption Absorption Dye Category Color Maxima, λmax Coefficient, ε Use (nm) (Lg-1cm-1) Blue 1 Triphenylmethane Sky blue 630 169 Food generally Blue 2 Indigo Royal blue 610 51.5 Food generally Green 3 Triphenylmethane Turquoise 625 165 Food generally Yellow 5 Azo Yellow 422 57.7 Food generally Yellow 6 Azo Orange 480 54.0 Food generally Red 3 Xanthene Bright pink 527 114 Food generally Red 40 Azo Red 500 58.0 Food generally Orange B Azo Orange NA NA Sausage casing Citrus Red 2 Azo Orange to NA NA Orange skins yellow NA: Not Available
According to their chemical structure these dyes can be classified as azo,
triphenylmethane, xanthene, and indigoid dyes (Figure 2.1). Table 2.1 also shows which dye
belongs to which of these four categories. Azo dyes are the largest and least stable among the
four that characterized by one or more azo bonds (-N=N-). Triphenylmethane dyes are known
8 by their three aromatic rings that attached to a central carbon atom which provides high stability and high tinctorial strength. There is only one approved xanthene dye (FD&C Red No.3), and, it has poor light stability with outstanding heat stability; it is, however, insoluble in a low-pH system. Finally, indigoid dyes has Indigo Carmine (FD&C Blue No.2) as its sole representative of its chemical class which is a sulphonated form of indigo (Frick 2003; Barrows 2003).
Figure 2.1: Chemical structures of four major categories of synthetic dyes.
Food color additives can also be classified into dyes and lakes, based on how they are made, their solubility, and the types of foods they are suited for. Dyes are water soluble whereas lakes are the insoluble form of the dye used in foods with fats and oils and little moisture through dispersion. Lakes are also made by reaction of colors with aluminum cation or
9 aluminum hydroxide (Barrows 2003). Table 2.2 shows some major differences between dyes
and lakes.
Table 2.2: Important property differences and use of dyes and lakes (Dziezak 1987).
Properties Lakes Dyes Solubility Insoluble in most solvents Soluble in water, propylene glycol, and glycerin Method of coloring Dispersion Dissolved Usage range 0.1-0.3% 0.01-0.03% Stability Light Better Good Heat Better Good Coloring strength Not proportional to pure dye Directly proportional to pure content dye content Hue Varies with pure dye content Constant Appearance in end product Opaque Transparent Cost More expensive Less expensive Suitable products for use Sugar coating, cake mixes, solid Hard candy, Striped candies, fats and waxes, chewing gum icings, sugar coating, cake mixes, carbonated beverages, dry mix products, chewing gum
2.2.2. Reasons for use
Synthetic dyes are usually the easiest form of food colorants to use in food products due
to their bright, stable, inexpensive, consistent, and relatively trouble free characteristics. Only
very small amount of these types of colorants are adequate to create high tinctorial power for
particularly all applications. The use level ranges usually from 20 or 30 ppm to 300ppm in most
foods (Walford 1980). These types of color additives can be used in a wide variety of
application which results in minimal inventory requirements for the users (Frick 2003). In
addition, they are free from bacteriological problem; don’t contribute to the flavor, and have
10 low cost. Beverages, confections, extrude foods, backed goods, dairy products, and pet foods are typical application for synthetic colorants (Walford 1980).
2.2.3. Limitation of use
In recent years, consumers are increasingly avoiding food containing synthetic dyes.
This is due to decreasing trend in consumption of food containing artificial additives, and, awareness of published studies linking the consumption of certain dyes to behavioral problems in children (Schab and Trinh 2004). Since 2010, the European Union has required foods that contain certain synthetic colorants (azo-dyes) to carry a label that warns consumers that the food
“may have an adverse effect on activity and attention in children”. This was mainly due to a study published in 2007 by a group of researchers at Southampton University on the effect of a combination of some synthetic food colorants and sodium benzoate (a preservative) on childhood behavior. According to this study and other similar study there is a possible link between the consumption of certain dyes and increased hyperactivity in children (Chapman
2011; McCann et al. 2007; Schab and Trinh 2004). In the U.S., the FDA set the Food Advisory
Committee panel up in 2011 to review the scientific evidence on artificial dyes and hyperactivity (FDA 2011). The panel, however, found limited evidence that artificial food colorings contribute to hyperactivity; since then, the FDA evaluated the exposure to FD&C dyes in the U.S. population and concluded that they are below the intake level defined by FDA.
However, this seems to be still ongoing investigation (FDA 2015).
Stability is also another reason that limits the use of these types of colorants. Although certified synthetic color additives are known to have good stability, under certain condition, they can lose colors, precipitate, or react with other components in food such as proteins resulting in color fade. Factors that could influence the stability of these pigments are pH,
11 temperature, light, reducing agents, and the presence of other additives or metals. Table 2.3
shows the stability of the major certified colorant under various conditions. These colorants are
usually unstable in the presence of oxidizing and reducing agents, due to a lot of unsaturated
bonds. They also fade in the presence of metal ions such as Zn, Cu, and Fe in acidic and
alkaline environment especially in the presence of heat (Pintea 2007).
Table 2.3: Stability of FD&C approved colorants under different environmental conditions (Pintea 2007; Francis 2002).
pH Stability after Ascorbic Sulfur Dye Light * Heat * Alkalies * 1 Week† Acid * Dioxide * 3 5 7 8 Blue 1 4 5 4 4 4 sf vsf vsf vsf Blue 2 1 4 3 2 2 af af cf fc Green 3 4 4 4 4 2 sf vsf vsf sf Yellow 5 5 4 2 4 5 naf naf naf naf Yellow 6 4 3 2 4 5 naf naf naf naf Red 3 2 5 2 5 5 ins ins naf naf Red 40 5 3 2 2 4 naf naf naf naf *1= very poor, 5=good †sf= slight fade, vsf= very slight fade, af= appreciable fade, cf= considerable fade, fc= complete fade, naf= no appreciable fading, ins= insoluble.
2.3. COLORS EXEMPT FROM CERTIFICATION
Color additives exempt from certification do not need to be certified; but, their identity
and purity specifications must be followed by the manufacturing and consuming companies. In
the U.S., they include colorants derived from plant or mineral origins and man-made colorants
originating from natural sources through processing or biotechnology. An example would be
caramel color obtained by heat processing of sugar. An exemption to that are cochineal extract 12 and its lake, which is derived from Dactylopius coccus costa (Coccus cacti L.) insect, and
Spirulina extract which is derived from the dried biomass of Arthrospira platensis (A. platensis) cyano bacteria (Socaciu 2007; Barrows 2003).
According to CFR 21, part 73, exempt colors are permitted for food use; however, the permission is limited to certain food matrices. For example grape skin extract is restricted to be used only in beverages. When this type of colorant used, the label needs to state that color added, colored with X, or X color to indicate the type of colorant (21CFR73; Socaciu 2007).
New food colorants, even if they are derived from natural sources, have to be approved by the
FDA as color additives prior to their use in foods unless they can be classified as “fruit juice” or
“vegetable juice” (Newsome et al. 2014).
2.3.1. Naturally occurring pigments for food use
Various pigmenting compounds in nature impart coloration and function in living organisms. These colors include various shades of red, purple, orange, brown, yellow, green, and blue. Application of these pigments into the food could please the consumers who associate the quality and freshness of food with colors (Chapman 2011). These pigments can be found in plants, insects, and microorganisms; and, they can display various shades of red, pink, orange, yellow, green, brown, black, and blue in them. Color, however, is not the only reason for the presence of these pigments; they also carry out different properties and functionalities in the living organisms. They can act as attractants or mating signals for mate attraction, camouflage against predators, participate in metabolic processes (e.g. chlorophylls in energy generation), serve as antioxidant, and have protective effects against sunlight and radiation. Consequently, apart form their visual appeal, functionality and potential health benefits; these compounds have additional advantages (Simpson et al. 2012).
13 Table 2.4 shows the major pigments suitable for food applications that are found in the current exempt color additives. There are, however, other compounds that have coloring properties but they are not usually used for the purpose of coloring food products (Simpson et al. 2012). An example is tannins that can have yellow or brown color. They are mainly found in tea and coffee, as well as, in other foods such as pomegranates, persimmons, cranberries, etc.
Quinones are other compounds that can also have pale yellow to dark brown or black pigments.
There are also minor pigments such as riboflavin (vitamin B2) that has a yellow green color and can be extracted from nature or chemically synthesized. It is used to fortify and/or color foods like dairy products, cereal, and dessert mixes. It is water soluble and heat stable; however, it is sensitive to light. Riboflavin is listed as an exempt color additive; but, it is not extensively used as a food colorant perhaps due to its high cost (Simpson et al. 2012; Chapman 2011).
14 Table 2.4: Major types of natural food pigments found among the list of colorants exempt from certification; their major source, properties, and use in foods (Simpson et al. 2012; Chapman 2011; 21CFR73).
Color additives Source Main use in food Color Major properties Pigment Pigment
Annatto seed, Bixa Annatto extract Mainly cheese Fat soluble, heat and ph stable, orellana L. Yellow sensitive to oxidation especially β-Carotene carrot and palm oil Foods and beverages and when exposed to light, Tomato lycopene Orange encapsulated forms can be water Tomato pulp Foods and beverages
Carotenoids extract soluble
Prepared by saponification and Sodium copper replacement of magnesium by Alfalfa (Medicago sativa) Used for citrus-based dry
1 chlorophyllin Green copper, water soluble, moderate
5 beverage mixes heat and light stability,
Chlorophylls precipitate in acidic conditions
Grape skin extract Grape remaining after Carbonated and alcoholic
(enocianina) pressed for juice or wine beverages Water soluble, pH sensitive, Red/pink Stable during short periods of to purple moderate heating, Fade over time Fresh edible blue berry, to blue on exposure to light
Anthocyanins raspberry, balckberry, red Fruit and vegetable radish, red cabbage, Foods and beverages juice purple potato, black carrot, etc To be continued
15
Table 2.4 continued
Color additives Source Main use in food Color Major properties
Pigment Pigment
Water soluble, Sensitive to heat Dehydrated beets Red beets Foods and beverages Pink/red and light, Prone to oxidation, pH (beet powder)
Betalains stable
Confections, frostings, ice cream and frozen desserts, dessert coatings and toppings, Arthrospira platensis cyano beverage, yogurts, custards, Water soluble, Sensitive to heat Spirulina extract Blue-green 1 bacteria puddings, cottage cheese, and light, Stable at pH 4-7
6 gelatin, breadcrumbs, and Phycocyanins ready-to-eat cereals (excluding
extruded cereals)
Cochineal (Dactylopius Deep Water-soluble, Stable to heat and Cochineal extract; used in alcoholic beverages coccus costa (Coccus cacti orange to light, Resistant to oxidation, Not acid carmine and processed meat products
L.)) red kosher
Carminic Carminic
Ground rhizome of food products such as curry, Lemon Oil soluble, Tends to fade when Turmeric
Curcuma longa L. soups, and confectionery yellow exposed to light, Heat stable Curcumin
16 2.3.2. Future trends in natural colors
From the current trend, it can be anticipated that the use of natural food colorants relative to the synthetic ones will keep increasing. The use of additives in foods would also be expected to decrease unless they are used for safety reasons (e.g. nitrites in bacon). This would influence the search for alternatives from natural food colorant sources (MacDougall 2002).
This could be done by finding novel sources, modification of existing natural pigments, isolation of certain pigments through selective isolation process; or, by selectively breeding or genetically modifying existing food crops to have higher (or certain) colorant contents.
2.4. BLUE COLORANTS
2.4.1. Blue color perception
Color is observed through the selective absorption of visible wave length of light.
Ordinary color perception happens when an object absorbs part of the radiation (from a certain light source) and reflects the remaining photons. The photons that are reflected then enter the human eye; this results in stimulation of the retina which then recognizes the objects color by the brain. When a pigment absorbs red light, it thereby appears blue; and, if it absorb green light, it then appears purple (red and blue combination) (Wrolstad and Smith 2010; Newsome et al. 2014). There are also other less common processes for color perception such as luminescent
(e.g. fluorescence) and phosphorescence which are due to emitted (not absorbed) photons.
Diffraction and scattering of light are also other ways to achieve color perception (Newsome et al. 2014).
17 “Blue pigment is defined as an organic molecule that absorbs red light (600 nm region) and
appears blue by ordinary color perception” (Newsome et al. 2014).
The process of light interaction with molecules can be explained by the electronic absorption theory. According to this theory, when the energy of an incident photon matches an energy gap (∆E= hν) between molecular orbitals such as σ, n, or π, the photon is then absorbed by an electron that is moved to the next higher energy orbital. Ultraviolet photons are absorbed by σ → σ* and σ → π* transition. For the visible light to absorb (color appearance), however, lower energy π → π* and n→ π* transitions are required. Conjugation of π-bond systems can lower (∆E= 3.1-2.5 eV) the π → π* energy gap transitions; this results in blue light absorption and red, orange, and yellow color appearance. For blue color appearance (red light absorption)
(600-700nm, ∆E= 2.1-1.8 eV) linear π-bond conjugation alone is not enough; and, it also requires aromatic ring systems, heteroatoms (-NH2, -OH, =O, -Cl, etc), and ionic charges
(Figure 2.2). In the case of organometallic and transitional metal complexes, their d-orbitals afford many lower energy electronic transitions which sometimes results in bright blue color
(Newsome et al. 2014).
18 FD&C Blue No.1
Blue color perception Full spectrum of visible light hv
2 Requirement for red light absorption: 1.5 • π bond system • Aromatic rings 1 • Heteroatoms
Absorbance • Ionic charge 0.5 • Metal ions
0
560 400 420 440 460 480 500 520 540 580 600 620 640 660 680 700 π (HUMO)→π* (LUMO) Wavelength (nm)
Figure 2.2: Blue color perception for a blue pigment and the properties of a red light absorbing molecule using the visible absorbance spectrum of FD&C Blue No.1 solution (λmax= 630nm) as an example (Newsome et al. 2014).
2.4.2. Synthetic FD&C blue colorant
Brilliant blue (FD&C Blue No.1) and Indigotine (FD&C Blue No.2) are two FDA approved synthetic blue colorants that are currently being used by the food industry. Both of these dyes are general-purpose color additives that can be used in food products alone or in combination with other colorants. The molecular structures and some of their color properties are shown in Figure 2.1 and Table 2.1 respectively.
Brilliant blue (FD&C Blue No.1) is normally supplied as the water-soluble sodium salt.
The powder is a reddish-blue shade, and, its aluminum lake is a bright blue powder. Blue No.1 dye is shown to have a good stability in various environmental conditions (Table 2.3). The color
19 is fairly compatible with most food components. It is not degrade by the microflora in the gastro-intestinal tract which results in a strange coloration of feces, causeing concerns for some consumers. Blue No.1 has low level (<10%) of absorption into the body, and, low degree of toxicity when consumed within the Acceptable Daily Intake (ADI) of 12.5 mg/kg body weight/day (Parkinson and Brown 1981; Emerton 2008).
Indigotine (FD&C Blue No.2) is also supplied as the sodium salt, and, the powder is dark blue hue. This dye has the poorest stability among all the other certified color additives
(Table 2.3) and has very poor compatibility with many food compounds such as sugars
(sucrose, dextrose, and glucose). The ADI of this color is 5mg/kg/ body weight/day, and, there is no toxicological concern with it (Emerton 2008; Pintea 2007).
2.4.3. Alternatives for blue colorants from natural sources
Finding alternatives for blue colorants from natural sources is extremely difficult.
Although blue colors are not unusual in nature, replicating this color in foods is very challenging. The primary concern with finding a naturally derived alternative for blue colorants is matching their shade to the synthetic blue colorants (e.g. FD&C blue no.1). There are also other fundamental concerns such as pH sensitivity, heat and light stability, solubility, interaction with other food matrices’, and etc. Additionally, gaining their regulatory approval to be used in food is another big challenge for the current blue color alternatives from natural sources
(Newsome et al. 2014).
One alternative is a purplish blue pigment (Huito) obtained from fruit of Genipa
Americana tree found mainly in the tropical and part of the subtropical regions of Latin
America. This plant has been used traditionally as a source of blue and black colorants in South
America; and, the colorant is still used for coloring cloth and pottery. There is not, however,
20 enough information about the color characteristics of this colorant, but, it is reported to have good heat stability (> 90◦ C). Huito has also limited approval for use in the U.S. (Gruenwald and
Galizia 2005; Newsome et al. 2014; Echeverry et al. 2011).
Other alternative is Gardenia blue from the fruit of Gardenia jasminoides plant, which is an evergreen shrub originated from southern China and Japan. The main compound of this fruit which is responsible for blue coloration is called genipin (a yellow colorant). This compound can be transformed to blue pigments through a simple modification. These blue pigments are stable after 10 h at temperatures of 60-90 ◦C. These blue pigments are not influenced by pH, and, have visible absorption maximum (λmax) of 596nm. Gardenia blue has limited approval for use in the U.S. (Gruenwald and Galizia 2005; Newsome et al. 2014).
Trichotomine glycosides are also a novel source of blue color which can be extracted from the skin of Japanese Kusagi berry (Clerodendron trichotomum). They are classified as bis- indole alkaloid pigment and used in Japan to color blue textile dyes. The color of this pigment
-1 -1 can have λmax of 658nm in chloroform solution with a large ε value of 70000 M cm .
Trichotomine Glycosides, however, require a significant investment for their regulatory approval (Newsome et al. 2013 and 2014).
Natural indigo is more prominent in the natural blue color market. It is used as a textile dye, but, it also has application in the food and cosmetic industries. It grows all over the world in various climates. Mexico and South India are the two main countries with established commercial indigo fields. The two major species of this plant with high content of blue indigo pigments are Indigofera tinctoria, and Indigofera suffruticosa. The leaves are the major site for the variety of blue pigments. Several steps of solubilization and reduction are required for the extraction of the pigments. As far as the coloring properties of the blue pigments, they can produce a blue color with the λmax of 604nm; however, they are reported to have low stability, especially in beverages (Jespersen et al. 2005; Gruenwald and Galizia 2005).
21 Spirulina extract is a more recently FDA approved blue colorant in the U.S. for coloring candy and chewing gum. Spirulina, derived from the dried biomass of Arthrospira platensis (A. platensis) cyano bacteria, is a phycocyanins (blue protein complex from blue-green algae of
Spirulina platensis). Phycocyanins can have the λmax of 616-620nm; however, it is sensitive to pH, light, and heat. (Dufosse et al. 2005; 21CFR73).
Anthocyanins are also capable of producing blue color. The production of blue color by anthocyanins, however, depends on multiple factors such as pH of the media, molecular structure of the pigment(s), and compounds attached to the anthocyanin molecule(s) (Giusti and
Wrolstad 2003). Stabilization of the blue color produced by anthocyanins is also another important issue which depends on the interaction of anthocyanins within their molecules and with other compounds in the matrix such as other phenolics or metal ions (Castaneda-Ovando et al. 2009). More explanations about the coloring properties of this group of pigments and their potential for formation of blur color will be presented in section 2.5.5.
2.5. ANTHOCYANINS
Anthocyanins are the most important vascular pigments in plants. The word anthocyanin comes from the combination of two Greek words: anthos= flower and
Kianos=blue. Anthocyanin pigments have use as natural water soluble colorants (Castaneda-
Ovando et al. 2009). They are found in flowers, fruits, and vegetables. The main properties of anthocyanins are listed in Table 2.4.
22 2.5.1. Chemical structure
Anthocyanins are flavonoids compounds with a C6-C3-C6 configuration. They are essentially made of two aromatic rings (A and B) that are joined by a 3 carbon heterocyclic ring
(C). The molecular structures of anthocyanins are shown in Figure 2.3. The basic structures of anthocyanins are called anthocyanidins. Anthocyanindins (or aglycons) have no sugar moiety attached to them. Pelargonidin (Pg), Cyanidin (Cy), delphinidin (Dp), peonidin (Pn), petunidin
(Pt), and malvidin (Mv) are the are the major anthocyanidins because their glycosylated forms occur frequently in the nature (Ignat et al. 2011; Harborne 1989). The differences between the six are due to the numbers and the type (hydroxyl or methoxyl groups) of attachments on the B ring.
Anthocyanin Abv R1 R2 Pelargonidin Pg H H Cyanidin Cy OH H
Peonidin Pn OCH3 H Malvidin Mv OCH3 OCH3 Petunidin Pt OCH3 OH Delphinidin Dp OH OH
Figure 2.3: Basic structure of the six major anthocyanins found frequently in the nature (He and Giusti 2010).
23 A sugar molecule can be attached to carbon 3, 5, 7, 3’, 4’ and 5’ in order of occurrence.
Generally, position 3 and 5 are the more common site of this attachment. D-glucose, L- rhamnose, D-galactose, D-xylose and D-arabinose are the most prevalent sugar moieties in the anthocyanins (Francis and Markakis 1989; Da Costa et al. 2000).
Anthocyanins can also have aromatic phenolic acids substituents such as benzoic acid
(e.g. gallic or p-hydroxybenzoic acids) or cinnamic acid (e.g. p-coumaric, caffeic, ferulic acids) derivatives; or, aliphatic organic acid attachments such as malonic, acetic, malic, succinic or oxalic acids. The acyl substituents are frequently attached to C3 sugar or the hydroxyl group of the sugars at C6 or C4 (less common). Due to the varieties in numbers, types, and position of the different sugar or acyl substituents, there exist complex molecular structures of anthocyanins (Goto and Kondo 1991; Giusti and Wrolstad 2003).
2.5.2. Health benefits
Anthocyanins are shown to have numerous potential health benefits such as reducing the risk of chronic disease such as cardiovascular disease and cancers (Ghiselli et al. 1998; He and Giusti 2010; Fernanda Nunez and Magnuson 2013), prevention of obesity and type 2- diabetes (Jayaprakasam et al. 2005), enhancement of sight and acuteness (Tsuda 2012), and anti-allergic and antimicrobial activities (Heins et al. 2001; He and Giusti 2011). Most of the suggested health benefits of anthocyanins are, however, related to their antioxidant an anti- inflammatory activities (Kong et al. 2003).
Anthocyanins are shown to be good antioxidant compounds due to their effective free radical scavenging properties (Wrolstad 2004). Phenolic structures of anthocyanins are the main reason for their antioxidant activity. Anthocyanins have the ability to scavenge reactive oxygen species (ROS) (such as superoxide, singlet oxygen, and peroxide, hydrogen peroxide, and
24 hydroxyl radical). ROS are carcinogenic and can induce severe loss of physiological function of cells and cancer (Wang and Jiao 2000).
Anthocyanins also demonstrated some anti-inflammatory activities. Chronic inflammation has shown to increases the risk of cancer. Chronic inflammation results in production of pro-inflammatory cytokines from inflamed cells that can damage DNA and induce tumorigenesis (Balkwill and Mantovani 2012). Anthocyanins have been reported to change activities of multiple enzymes associated with inflammation. In vitro, Cy derivatives from tart cherries inhibited cyclo-oxygenase-1 and -2 (COX-1 and -2) activities that are responsible for inflamation (Seeram et al. 2006). In vivo studies also have shown anti- inflammatory activity of anthocyanins. Hogan et al. (2010) demonstrated that an inflammatory indicative protein (C-reactive protein ) decreased in rats fed with Norton grape pomace extract
(156.9 mg/g as Cy-3-glu equiv) compared to that in animals fed standard diet.
2.5.3. Anthocyanins as food colorants
Anthocyanins are good alternatives for coloring foods, and, from a regulatory standpoint, as mentioned in section 2.4.1, they are permitted in the U.S. as food colorant in the category of fruit (21CFR73.250), vegetable (21CFR73.260), or grape skin extract
(21CFR73.170). They can form wide shades of orange, red, pink, yellow, purple, blue, green, and gray (Figure 2.4). Also, it is often simple to incorporate them into aqueous food system due to their water solubility (Brouillard 1983; Mazza and Miniati 1993; Brouillard 1982). Currently anthocyanins are being used in many food and beverages as alternatives to many synthetic dyes.
Their use however, is limited due to their relatively low stability to processing, storage conditions, formulation, and possible undesirable contribution to the odor and flavor of the food
(Giusti and Wrolstad 2003; Bąkowska-Barczak 2005).
25
Figure 2.4: Example of color variation of anthocyanins. red radish: RR, red cabbage: RC, purple sweet potato: PSP, blueberry: BB, elderberry: EB.
Stability of anthocyanins depends on many factors such as the molecular structures, pH, presence of oxygen and other compounds, light, and processing condition (Frank et al. 2012;
Bąkowska-Barczak 2005; Mazza and Brouillard 1990; Brouillard 1982). With respect to the molecular structure, for instance, acylation of anthocyanin are shown to be an important factor that influences their stability especially in high heat and medium to high pH range that the anthocyanins are least stable. It has been shown that increasing the number of acylation increase the stability of anthocyanins (Mazza and Brouillard 1987; Giusti and Wrolstad 2003).
Anthocyanins are also shown to be more stable in acidic media and have lower stability in the medium and alkaline pH, so, modification of the food acidity (if doesn’t impact the color and flavor greatly) could help with increasing the stability (Brouillard 1982; Mazza and Miniati
1993). Presence of light, oxygen and some compounds in food (such as ascorbic acid) are also shown to decrease the stability of some anthocyanins (Inami et al. 1996; West and Mauer 2013;
Robert et al. 2010; Bąkowska-Barczak 2005), whereas, other compounds (such as other phenolics) have shown to increase the stability of these pigments upon their presence (Robert et al. 2010; Bąkowska-Barczak 2005; Giusti and Wrolstad 2003). Anthocyanis are heat-labile and would degrade when they are exposed to prolong and extensive heating (Dyrby et al. 2001;
Fracassetti et al. 2013). Modification of processing conditions such as application of non-
26 thermal processing could also help with the stability of anthocyanins. Additionally, there are other strategies (such as encapsulation) that can be taken in order to increase the stability of anthocyanins and make them better alternatives for coloring foods and beverages (Robert et al.
2010).
Removal of undesirable odors from some anthocyanin containing fruits and vegetable sources has also been made possible. Sapers (1982) was successful in preparing an odor-free colorant from red cabbage without affecting the stability and functionality of the colorant by using a polymeric adsorbent (Amberlite XAD-7). The use of resin adsorbent was also successful in removing the undesirable odor from red radish juice (WU et al. 2011).
2.5.4. Chemical characterization:
2.5.4.1. Extraction
Anthocyanins are usually extracted by solvent extraction. The plant materials are initially needed to be grounded, dried, or lyophilized, or, only soaked in a subsequent amount of solvent extract. Since these compounds are polar molecules, the most commonly used solvents for their extractions include EtOH, MeOH, or acetone. Because anthocyanins have good stability in acidified environments, acidified MeOH or EtOH are usually used for their extractions. Although MeOH extractions of anthocyanins are reported to be more efficient,
EtOH is being used in the food industry as the extracting solvent due to MeOH toxicity
(Castañeda-Ovando et al. 2009). There is also a recent report that shows higher proportion of water as an extracting solvent being more effective than other harsher chemical for anthocyanin extraction of Cylon gooseberry (Bochi et al. 2014). The use of polar solvents, however, results in co-extraction of other polar compounds such as sugars, organic acids, and proteins which requires subsequent purification steps (Castaneda-Ovando et al. 2009).
27 2.5.4.2. Quantitation
Accurate measurement of anthocyanins content is important because of two major reasons. Anthocyanin concentration affects color characteristics, and, if used as neutraceutical, knowing the concentration help with a prediction of some of its biological activities such as anti-oxidant activity.
Quantification of anthocyanins is complicated due to the inference from other compounds. Other compounds could include melanoidin pigments or other anthocyanins
(polymeric forms) in a mixture. There are several ways to measure the concentration of anthocyanins; however, pH differential method by spectrophotometry and High Pressure Liquid
Chromatography (HPLC) are two major anthocyanins quantification methods that are commonly used by researchers and the food industry (Wrolstad et al. 2001; Giusti and Jing
2007; Lee et al. 2005).
The pH differential method is an official AOAC method (2005.02) which determines total monomeric anthocyanin content. The principle behind this method is related to the response of monomeric anthocyanin pigments to the pH change by reversible change in their color (Figure 2.5). Total anthocyanin pigments were traditionally determined by measuring the absorption at a single band (between 490 to 550nm) when the pigments were dissolved in acidified media. The downside to this method is that possible presence of degradation products of anthocyanins and melanoidin pigments in the solution cause interference with this method since they also have absorption in the same region. For the pH differential method the sample is dissolved in two different buffers (pH 1 and 4.5), one at a time, and each time the absorbance is measured at the wavelength of maximum absorption (λmax). The total anthocyanin content is measured by the equation bellow. Generally all the pigments exist in the colored (oxonium) form at pH 1; however, only monomeric anthocyanin pigments form the colorless (hemi-ketal) form at pH 4.5 (Wrolstad et al. 2001).
28 Monomeric anthocyanin count (mg/liter)= (A x MW x DF x 1000) / (ε x l)
Where
A= Difference in absorbance between the pH 1 and 4.5 solutions; MW= molecular weight; DF= dilution factor; ε= molar absorptivity; l= pathlength
It is important to select an appropriate molar absorptivity for this measurement based on the major anthocyanin present in a sample. If the ε is not known, the pigment content can also be reported as cyanidin-3-glucoside because of the frequency of its abundance.
The molar absorptivity values of many anthocyanin pigments have not yet been determined; also, there is lack of consistency for the same pigment that has been reported by different investigators. This is due to several reasons such as difficulties in isolation, purification, and preparation of crystalline anthocyanins in sufficient quantities. Factors that affect the molar absorptivity of anthocyanins are the structure and the solvent that they are dissolved in (Giusti et al. 1999).
The HPLC method is another more advanced method that can determine an absolute anthocyanin amount in a sample. This method requires pure anthocyanin standards which are used in the external standard method. It also requires establishment of the identity of the anthocyanin pigments present in a sample prior to their quantifications (Giusti and Jing 2007).
2.5.4.3. Solid phase extraction (SPE)
SPE is a rapid and sensitive technique that uses different types of cartridges with a variety of sorbents. One of the advantages of SPE is that sample preparation and concentration can be done in one step (Andersen and Markham 2006). C18 and Sephadex are two commonly used cartridges for purification and concentration of anthocyanins. Hydroxyl groups of anthocyanins strongly bond into these sorbents and subsequent amounts of these compounds can be separated from other compounds by increasing the polarity with different solvents 29 (Castañeda-Ovando et al. 2009). The majority of interfering sugars and some acids can be eliminated from crude anthocyanin extracts with several washes of acidified water using a C18
SPE cartridge. If higher purification is desired to isolate a specific anthocyanin pigment from a mixture, however, preparatory HPLC is a more powerful technique that can be used.
2.5.4.4. High Pressure Liquid Chromatography (HPLC)
HPLC is one of the most widely used techniques for the separation of anthocyanins for qualitative and quantitative analysis (Andersen 2006). Historically, anthocyanins were separated and analyzed using paper or thin-layer chromatography. HPLC, in comparison, has the advantages of greater resolution, shorter analysis time, and easier quantitation. Reversed phase
HPLC is commonly used for anthocyanins separation which separates them based on their polarities. The order of elution for anthocyanin agylcons depends on the number and type
(methoxy and/or hydroxyl) of attachments on the B-ring (Dp < Cy < Pt < Pg < Pn < Mv).
Anthocyanin glycosides are normally eluted before the aglycones. Numbers of sugars increase the polarity of the compound and (most of the time) decrease the elution time (Triglycosides <
Diglycosides < Monoglycoside). Elution times for hexose glycosides are shorter than pentose glycosides. Substitution of a sugar on C3 carbon of anthocyanins with aromatic and aliphatic acids, on the other hand, decreases the polarity which results in longer retention times
(Wrolstad and Smith 2010; Andersen and Markham 2006).
Two frequently used columns for HPLC analysis of anthocyanins are Silica C18 (for simpler anthocyanins) and Polymeric C18 (more complex anthocyanins) because of their ability to retain these compounds through hydrophobic interactions. Newer generations of columns can also have addition of phenyl phases which can additionally provide aromatic (π-π) interactions.
This aromatic interaction helps with improving the columns’ performances for separation of aromatic compounds such as anthocyanins (Koerner et al. 2010). Acidified (pH<2) water is 30 used often as an initial mobile phase to ensure that the anthocyanins are in oxonium form; this also helps with their UV detection. Gradient methods are used for analyses by decreasing the gradient of the acidified water (more polar phase) and increasing amount of acetonitrile or
MeOH (Andersen and Markham 2006; Wrolstad and Smith 2010).
HPLC can also be used for preparatory purposes to isolate a pure anthocyanin compound from a mixture. Preparative HPLC requires a large column (internal diameter 8-20 mm) that helps with separation and isolation of individual pigments from a large amount of concentrated anthocyanins containing samples. Ordinarily, a separation method needs to be optimized using analytical HPLC prior to transposition to a semipreparative scale to avoid extra cost of solvents (Andersen and Markham 2006).
2.5.4.5. Mass Spectroscopy (MS)
MS is a powerful identification tool used widely in combination with HPLC for analysis of anthocyanins. The MS principle is based on ionization of compounds to generate charged molecules or their fragments and measuring their mass to charge ratio (Sparkman and
Boggess 2001). MS is one of the most sensitive methods for molecular analysis of compounds.
Excellent selectivity is also achieved using this technique due to its high capability of mass separation (Andersen 2006; Wrolstad et al. 2001).
Two powerful techniques that are often used for the characterization of anthocyanins with MS are electrospray (ESMS) and tandem mass spectroscopy (MSMS) (Giusti and
Wrolstad 1996; Wu and Prior 2005b; Ella Missang et al. 2003; Wu and Prior 2005a). Other ionization techniques, however, such as Atmospheric-pressure Chemical Ionization (APCI),
Fast Atom Bombardment (FAB), Matrix-assisted-laser-desorption-ionization (MALDI), and
Matrix-assisted-laser-desorption-ionization-time-of-flight (MALDI-Tof) have also been used
(Zhang et al. 2004; Saito et al. 2002; Wang and Sporns 1999; Castañeda-Ovando et al. 2009). 31 Although anthocyanins are thermally labile, nonvolatile, and polar compounds, they still can be ionized using soft ionization because they possess positive charge in an acid environment, and, the soft ionization methods such as ESI cause very low fragmentation (Castañeda-Ovando et al.
2009). Upon the entry of pure anthocyanin into the mass spectrometer, a distinct mass unit will be generated that is very useful in identification of the pigment. There is usually very little interference from other compounds such as polyphenols (at appropriate voltage) due to the anthocyanins positive charge. In addition to intact anthocyanins, small amount of the aglycon is also generated during ionization in most cases due to the cleavage of the glycosidic pigments
(Wrolstad et al. 2001).
Tandem MS-MS, in addition, can provide clear fragmentation patterns to analyze anthocyanin aglycone and sugar moiety characterization (Ignat et al. 2011). During the mass analysis, individual molecules are selected by the first quadrupole and fragmented in the collision cell using an inert gas such as argon. Second quadrupole will then assist with detection of the fragments. One of the advantages of using MS-MS for the analysis of anthocyanins is that they help with distinguishing between 3- and 3-5-glycosidic substitutions (Wrolstad et al. 2001).
2.5.5. Factors affecting the color and blue color formation of anthocyanins
2.5.5.1. pH
pH of the media is essential in color variations and structural changes of the anthocyanins (Mazza and Brouillard 1987). Flavylium cation, quinoidal base, hemiketal, and chalcone are the four well known anthocyanin structures that are found in equilibrium in any aqueous solution. A change in pH of the media can shift this equilibrium to different directions and increase or decrease the dominance of certain forms. Figure 2.5 shows these structures and their dominance based on the pH. Flavylium cation (red color) is the predominant form at pH 1.
32 When the pH increases, anthocyanins are often become colorless (at pH 4-5), and, the carbinol pseudobase are mostly present. Blue and purple color forms often at neutral to alkaline pH of 6-
8 where the quinoidal base species are the dominant species in the solution; and, pH above 8 pyrylium ring will open up resulting in the formation of Chalcone form (Giusti and Wrolstad
2001; Fleschhut et al. 2006). However, not all the anthocyanins behave the same way, and, this color formation with pH could somewhat change based on the different molecular structure of anthocyanins.
Figure 2.5: Structural changes in anthocyanins with change in the pH (Stintzing et al. 2002; Wrolstad et al. 2001; Fleschhut et al. 2006).
2.5.5.2. B-ring substitution
Anthocyanidin molecular structure is very important in the color behavior of the pigment in different media. Presence and number of hydroxyl and methoxyl substitutions on the 33 B-ring influence their absorbance spectra. Higher amounts of hydroxylation and methoxylation on position 3’ and 5’ causes a bathochromic shift (a shift of λmax toward longer wavelength) which results in a bluer hue (Mazza and Miniati 1993; Cabrita et al. 2000; Giusti and Wrolstad
2003). Figure 2.6 shows the influences of various 3’ and 5’ substitutions of anthocyanins on the
λmax of the spectra at different pH values. As shown in this figure, in pH 1-8.1, glycosylated
MV, Pt, and Dp having OH and/or OCH3 substitutions at position 3’ and 5’had larger λmax than
Pg, Cy, and Pn with only one hydroxyl or methoxyl substitution on those positions.
610 Pg3glc (3':H , 5':H) Cy3glc (3':OH , 5':H) 590 Pn3glc (3':OCH3 , 5':H) Dp3glc (3':OH , 5':OH) 570 Pt3glc (3':OCH3 , 5':OH)
Mv3glc (3':OCH3 , 5':OCH3)
550 nm
530
510
490 1 2 3 4 5 6 7 8 9 pH
Figure 2.6: λmax of six common anthocyanidin-3-glucosids (with different 3' and 5' substitutions) at different pH values 1h after dissolution in buffers at room temperature (Cabrita et al. 2000).
34 2.5.5.3. Glycosylation
There is no clear effect with addition of glucose molecule to the anthocyanins. Giusti et al. (1999) showed that by addition of a glucose molecule to the Pg backbone the hue of the color changed from red to more orange red color at pH 1. Addition of two more glucose molecules to Cy 3-glc, on the other hand, was shown to decrease the hue angle to a more red color at both pH 1 and 3. Bathochromic effect and formation of blue color with addition of the sugar molecules for Cy anthocyanin is more noticeable at neutral to alkaline solutions (pH 7.2 and 8.0) (Table 2.5 and 2.6) (Torskangerpoll and Andersen 2005).
Table 2.5: Color characteristics of two different anthocyanidins (Pg and Cy) and their glycosides with different concentrations in acidic pH values (Giusti and Wrolstad 1996; Torskangerpoll and Andersen 2005). pH Conc. L* C* h◦ Anthocyanin (mM) Pg 1.0 0.0154 90.0 16.7 22.7 Pg 3-glc 1.0 0.0176 90.3 17.6 44.0 Pg 3-(2-glcglc)-5-glc 1.0 0.0185 90.0 20.3 41.0 Pg 3-(2-glcglc)-5-glc 1.0 0.0298 81.8 56.0 53.5 Cy 3-glc 1.1 0.05 81.1 45.9 20.8 Cy 3-glc 1.1 0.15 63.7 81.9 42.4 Cy 3-(2-glcglc)-5-glc 1.1 0.05 81.5 46.7 12.8 Cy 3-(2-glcglc)-5-glc 1.1 0.15 69.5 78.6 38.0
Table 2. 6: Effect of the addition of one sugar molecule to the Cy anthocyanins on its color characteristics at selected pH values (Torskangerpoll and Andersen 2005).
Anthocyanin pH L* C* h◦ Cy 3-glc 3.0 63.0 70.2 25.6 Cy 3-(2-glcglc)-5-glc 3.0 76.9 52.9 4.4 Cy 3-glc 6.0 39.9 32.5 312.9 Cy 3-(2-glcglc)-5-glc 6.0 89.2 13.1 331.1 Cy 3-glc 7.2 32.2 21.9 313.0 Cy 3-(2-glcglc)-5-glc 7.2 29.1 73.4 303.9 Cy 3-glc 8.0 38.7 47.2 335.9 Cy 3-(2-glcglc)-5-glc 8.0 35.6 81.2 325.8 35 2.5.5.4. Concentration
Concentration of anthocyanins also is shown to influence their color properties.
Rationally, by increasing the anthocyanins concentration the lightness decreases and the chroma of the color increases. Torskangerpoll and Anderson (2005) showed that bathochromic shift happened for Cy3-(2-glcglc)-5-glc when the concentration decreased especially around neutral pH values (Table 2.7)
Table 2.7: Effect of the concentration of the Cy3-(2-glcglc)-5-glc on the color characteristics at different pH values extracted or modified (Torskangerpoll and Andersen 2005)
Conc. pH L* C* h◦ (mM) 0.05 3.0 90.5 23.1 375.2 0.15 3.0 76.9 52.9 4.4 0.05 6.0 96.6 4.0 334.0 0.15 6.0 89.2 13.1 331.1 0.05 7.2 69.3 34.7 262.3 0.15 7.2 29.1 73.4 303.9 0.05 8.0 67.1 52.8 319.1 0.15 8.0 35.6 81.2 325.8
2.5.5.5. Acylation
Acylation of anthocyanins with aromatic acids shows a clear bathochromic effect on the absorbance spectra of the anthocyanins. Cinnamic acid acylation of Cy and Pg anthocyanins shifts the λmax to the higher values and decreases the hue angle to purple-blue color (Stintzing et al. 2002; Giusti et al. 1999; Torskangerpoll and Andersen 2005). Effect of the acylation of
Cy with sinapic acid at different pH values is shown as an example in Table 2.8. As shown in this table, in the presented pH values, the hue angle is shifting toward more purple to blue color
36 with the addition of acylating group. The reason for this shift can be explained by a phenomenon called interamolecular co-pigmentation (section 2.5.5.6.).
Table 2.8: Effect of the addition of an acyl group to a glycosylated Cy anthocyanin on the color characteristics of the solution at different pH values (Torskangerpoll and Andersen 2005).
Anthocyanin pH L* C* h◦ Cy 3-(2-glcglc)-5-glc 3.0 76.9 52.9 4.4 Cy 3-(2-(2-singlc)-6-singlc)-5-glc 3.0 48.1 80.6 354.8 Cy 3-(2-glcglc)-5-glc 6.0 89.2 13.1 331.1 Cy 3-(2-(2-singlc)-6-singlc)-5-glc 6.0 35.4 75.8 317.0 Cy 3-(2-glcglc)-5-glc 7.2 29.1 73.4 303.9 Cy 3-(2-(2-singlc)-6-singlc)-5-glc 7.2 57.3 46.6 262.6 Cy 3-(2-glcglc)-5-glc 8.0 35.6 81.2 325.8 Cy 3-(2-(2-singlc)-6-singlc)-5-glc 8.0 28.2 56.1 282.6
2.5.5.6 Co-pigmentation
Co-pigmentation was observed for the first time by Willstatter and Zollinger (1916) when a grape anthocyanin pigment changed to bluer red color in the presence of tannin or gallic acid in the acidic condition. It was however, suggested for the first time in 1931 by Robinson and Robinson when they also observed that the extracted anthocyanins from fuchsia flower change their color to blue-red in the presence of tannin (Robinson and Robinson 1931). It was then defined as bluing of anthocyanins by substances such as flavones. Co-pigment can offer an explanation for a lot of color variation that is observed in flowers in pH range where anthocyanins are colorless (Asen et al. 1972).
There are two major types of co-pigmentation: intra-molecular and inter-molecular co- pigmentation. Interamolecular copigmentation could happen in anthocyanins with multiple
37 aromatic acylation when the acyl residues engage in intra molecular stacking with the anthocyanin (flavylium or quinonoidal) chromophores through hydrophobic interaction (Figure
2.7). This would often results in the folding of the molecule and formation of a “sandwich type” complex (Giusti and Wrolstad 2003).
Figure 2.7: Intramolecular stacking of diacylated anthocyanins above and below the flavylium molecule.
Intermolecular copigmentation (Figure 2.8) is also defined as loose association of certain copigmenting compounds with anthocyanins which results in color modification and stabilities of the later. These compounds could be anthocyanins themselves which in this case this interaction is called self-association (Figure 2.8); or colorless (or mildly colored) compounds such as flavonoids, alkaloids, polysaccharides, amino acids, organic acids, nucleotids, and metal ions (Castaneda-Ovando et al. 2009; Asen et al. 1972).
38
Figure 2.8: Intermolecular copigmentation of anthocyanins: selfassociation (top), copigmentation with a cinnamic acid (bottom).
Copigmentation of anthocyanins generally results in charge-transfer between the anthocyanin chromophore and the copigmenting compound(s), and, this could cause bathochromic and hyperchromic (more intense color) effects (Yoshida et al. 2009; Malien-
Aubert et al. 2001). One possible reason for a bathochromic shift is that the copigmenting molecule interacts with the anthocyanin pigment in its first excited state (electronic transition in the visible region) more intensely than with this pigment in its ground-state. Copigmentation of anthocyanins also helps in the stability of the color since the stacking of the co-pigmenting molecule on the planar nuclei of the anthocyanins prevents the nucleophilic attack of water at 39 the position 2 of the C (pyrylium) ring and partially suppresses the formation of hemiketal and chalcone forms (Malien-Aubert et al. 2001).
2.5.5.7. Metal complexation
The effects of metals in color stabilization of blue flower petals containing anthocyanin pigments have been studied for more than three decades (Yoshida et al. 2009). Anthocyanins with two hydroxyl substitutions on their B-ring, such as Cy, Dp, and Pt, are able to form metal- anthocyanin complex (Boulton 2001). Al, Fe, Cu, Sn, Mg, and Mo are among the metal ions that have shown some abilities to form complex with anthocyanin molecules. Stabilization of anthocyanins blue color could be due to the ability of metals to prevent the oxidation of the blue quinoidal base. Despite the low interest in the food industry, the addition of metal ion to food matrices for stabilizing anthocyanin colors could be a viable alternative for the production and stabilization of the blue color; particularly if the metal ions do not impose any health threat or when they are even part of the essential minerals in diet (Hale and others 2001).
2.6. RED CABBAGE
2.6.1. Taxonomy
Red Cabbage belongs to the family of Brassicaceae (common name: cruciferae). The genus is Brassica (common name: cabbage), and the species is oleracea. Kale, cauliflower, broccoli, brussels sprouts, and kohlrabi are other members of Brassica oleracea family.
Horticultural selection within the Brassica oleracea has led to increase in other conspecific
(belong to the same species) taxon of this species; among which is Ver. capitata L. group which
40 includes red cabbage. Depending on color and shape of the heads; however, other intraspecific taxa of red cabbage also exist (Yamaguchi 1983; Christman 2000; Diederichsen 2001).
2.6.2. Growth location
Red cabbage is an ancient crop originated from the Mediterranean area, the Near East and Europe. Generally, cabbage is grown in cool weather. In Europe, central Europe and the
Netherlands are the most suitable location for red cabbage growth. In the U.S., it can be grown in most places except for Puerto Rico, majority of areas in Alaska and Hawaii and small region in California (Christman 2000; Diederichsen 2001).
2.6.3. Nutritional composition
The caloric energy of red cabbage is low (27 kcal/100g). The edible portion of the plant
(100 g) is composed of: 92% water, carbohydrate (6.1 g), anthocyanin pigment (1.6 g), protein
(1.4 g), fiber (1 g), and fat (0.3 g). There are also K (206 mg), Ca (51 mg), P (42 mg), Na (11 mg), and Fe (0.5 mg) in 100g of fresh plant. Vitamin C (57 mg/100 g) is the major vitamin in the fresh red cabbage which is even higher than common cabbage (47 mg/100 g); the second major vitamin is vitamin A (40 IU/100g). IU stands for the International unit of measurement of a compound based on its biological activity. Other vitamins are rather found in small amounts
(0.03-0.3 mg/100g) in this crop. These amounts might, however, change based on the red cabbage variety and growth condition (Yuan 2009).
41 2.6.4. Production for processing purposes
Important factors for processing red cabbage are color, firmness, trimming and freedom from diseases and damage caused by bursting, freezing, disease, birds, insects, or mechanical and other means. Cabbages for processing are preferred to be larger (5-10 kg) than those for fresh market and are usually mechanically harvested. Generally processing cabbage yield is approximately 5.1- 7.1 tha-1 (Yamaguchi 1983; Maynard 1997). To be marketed as source of colorant from natural source variety selection seems to be the most important factor since different red cabbage variety has shown to have various anthocyanin pigment contents
(Piccaglia et al. 2002; Ahmadiani et al. 2014).
2.6.5. Red cabbage color
Color is the most important aspect for red cabbage. Anthocyanins are the major pigment responsible for the coloration of red cabbage. The major anthocyanin in red cabbage is
Cyanidin-3-diglucoside-5-glucoside (Cy3diG-5-G) found in various acylated forms (Dyrby et al. 2001). There are both di-acylated and mono-acylated anthocyanins in red cabbage.
Aacylation of anthocyanins in this plant are mainly with aromatic acids such as p-coumaric, sinapic, and ferulic (McDougall et al. 2007). Red cabbage is known to have high content of anthocyanins (10-18 g/kg DM) with good stabilities to light and processing temperature (Dyrby et al. 2001; Ahmadiani et al. 2014; Piccaglia et al. 2002). Although anthocyanins concentration in this plant is a little lower than that of some commercially grown berries, the potential yield of fresh material per unit area of cabbage is still higher than that of most small fruits (Piccaglia et al 2002).
42 2.7. POTENTIAL FACTORS AFFECTING ANTHOCYANINS ACCUMULATIONS IN PLANTS
Many intrinsic and extrinsic factors control the anthocyanin accumulation in the plant.
Selecting the right variety (an intrinsic factor) seems to be the most important factor for anthocyanin yield since the enzymatic pathway for anthocyanin production could vary in different varieties (Ahmadiani et al. 2014; Piccaglia et al. 2002). Light, nutrients, maturity, temperature, irrigation, pests, and pH of the soil are among the extrinsic factors which are proven to influence anthocyanin accumulation in various anthocyanin containing plants
(Chalker-Scott 1999). The effects of some of these factors have been discussed below.
2.7.1. Light
Light induce the production of anthocyanins in plants. Light influences several photomorphogenic (the use of light to control structural development in the plant) reactions.
The photomorphogenic phenomenon depends on photoreceptors. Photoreceptors are chemical pigments capable of absorbing specific wavelengths of light. There are 4 different photoreceptors in plants: phytochromes, cryptochromes, UV-B photoreceptors, and protochlorophyllides. Photoreceptors alter the expression of the anthocyanin regulatory and structural genes to induce the accumulation of anthocyanins. Phytochrome and the blue/UV light photoreceptor are two important photoreceptors for anthocyanin production (Khare and
Guruprasad 1993). Several researches have been conducted on various anthocyanin containing plants to evaluate the effect of light. Tsormpatsidis et.al (2008) studied the anthocyanin production in Lollo Rosso lettuce. Plastics were used to block the wavelength across the UV region. The results showed that the anthocyanin production was greatest when both UVA and
UVB were present. Beggs and Wellmann (1985), also, studied the formation of anthocyanin in
43 Zea mays varieties in the presence of UV-B, blue, red, and far-red light. The plants responded to the lights differently. UV-A was more effective than UV-B; and, continuous red light was the least effective in production of anthocyanins. Based on their results, combination and duration of red light and UV, however, seemed to be influential in one variety.
Intensity of the light could also be important in the production of anthocyanins in plants; however, too much radiation in the UVB range inhibits anthocyanin synthesis which could be due to DNA damage (Beggs and Wellmann 1985). Takeda (1988) studied the effect of light on anthocyanin synthesis in carrot cells. In this study, carrot cells were placed two different media at different concentration, and irradiated by white fluorescent lamp with different light intensity. The results showed that anthocyanin production increased by increasing the intensities of the light. These results shows that the anthocyanin content of red cabbage could also change due to the light intensity and the time that the plant is growing.
2.7.2. Nutrients
Optimization of nutrients is an important factor in production of anthocyanins in red cabbage. According to “Carbon-Nutrient Balance” hypothesis, “High nutrient availability leads to an increase in plant growth and development, and decreased allocation of resources towards the production of expendable secondary metabolites such as the phenolic antioxidants” (Jifon
2012). Piccaglia et.al (2002) investigated the effect of some nutrients in anthocyanin content in red cabbage. They found out, however, that P and K had a slight effect in the way that the highest rates of both compounds had a negative effect on the anthocyanin concentration in the cabbage heads. Studying the influence of N, though, is more complicated since N deficiency decrease the plant biomass, and anthocyanins accumulation requires biomass (Piccaglia et al.
2002).
44 2.7.3. Maturity
Maturity stage is also important in the anthocyanin concentration of anthocyanins.
Anthocyanin content of the plant changes due to its maturity stage. In plants such as apple, it has been shown that early in the season (during plant cell division), the concentration is relatively high; and, it gradually decreases during growth; however, it begins to increase again near maturation (Awad et al. 2001). Consequently, red cabbage harvesting time also influences the anthocyanin accumulation and color of the plants.
45 CHAPTER 3: ANTHOCYANINS CONTENTS, PROFILES AND COLOR
CHARACTERISTICS OF RED CABBAGE EXTRACTS FROM DIFFERENT
CULTIVARS AND MATURITY STAGES 1,2
3.1. ABSTRACT
Red cabbage (Brassica oleracea L.) is an excellent source of food colorant. This study aimed to evaluate the anthocyanin pigment contents and profiles from 7 red cabbage cultivars at two maturity stages (8 weeks apart) and evaluate their color characteristics and behavior under acidic and neutral pH. Anthocyanin concentrations ranged from 1,111 to 1,780 mg Cy3G/100gr
DM and did not increase with time. Cultivar and maturation affected pigment profile. Some varieties accumulated ≥30% of di-acylated pigments, and proportions of mono-acylated pigments decreased with time. Extracts from selected varieties at first harvesting time produced colors similar (λmax 520nm and ∆E6.1-8.8) to FD&C Red No.3 at pH 3.5. At pH 7, extracts from the second harvest with higher proportion of di-acylation produced λmax ≃610nm, similar to FD&C Blue No.2. Cultivar selection and maturation affected color and stability of red cabbage extracts at different pH values.
1 Neda Ahmadiania, Rebecca J. Robbinsb, Thomas M. Collinsb, M. Monica Giustia* a Department of Food Science and Technology, The Ohio State University, 2015 Fyffe Road, Columbus, OH 43210, USA,b Analytical and Applied Sciences Group, Mars Inc., 800 High Street, Hackettstown, New Jersey 07840 2 Published in Journal of Agriculture and Food Chemistry 62(30)7524-7531.
46
Keywords: Anthocyanins, Red cabbage (Brassica oleracea L.), Cultivar, Harvesting time, Color.
3.2. INTRODUCTION
Anthocyanins are water-soluble pigments with potential application in coloring of different food products (Giusti et al. 2008; Hendry 1992). Colorants made of these pigments are currently manufactured for food use from horticultural crops and processing wastes (Wrolstad and Smith 2010). Fruit and vegetable juice containing anthocyanins such as concentrated red cabbage, black carrot, purple sweet potato, radish, bilberry, elderberry are being used as approved food color additives in most countries (Socaciu 2007; Hendry 1992). In addition, anthocyanins are proven to be good antioxidant compounds due to their effective free radical scavenging properties and have shown numerous potential health benefits in vitro and vivo studies (Wrolstad 2004; Kong et al. 2003; Heins et al. 2001; He and Giusti 2010; Ghosh and
Konishi 2007).
Red cabbage (Brassica oleracea L.) is an edible source with high content and high potential yield per unit area of anthocyanins (Piccaglia et al. 2002). Red cabbage anthocyanin extract is known to have considerable amount of mono- or di-acylated Cy anthocyanins
(McDougall et al. 2007; Wiczkowski et al. 2013). Type and acylation of anthocyanins are two important factors that determine their color characteristics at certain pH values (Mazza and
Miniati 1993; Cabrita et al. 2000; Giusti and Wrolstad 2003; Stintzing et al. 2002; Giusti et al.
1999). Due to its anthocyanin composition, red cabbage anthocyanin extracts can exhibit a wide spectrum of color, ranging from orange to red, to purple and blue based upon the pH of the environment (Walkowiak-Tomczak and Czapski 2007). Acylation of anthocyanins also
47
influence their antioxidant properties and stability in the food matrix (Tamura and Yamagami
1994; Dyrby et al. 2001; Giusti and Wrolstad 2003). Di-acylated anthocyanins are linked to higher antioxidant activity compared to the other non and mono acylated ones (Wiczkowski et al. 2013). Anthocyanin pigments with higher number of acylation have also shown good stabilitiy to light and processing temperatures (Dyrby et al. 2001).
There are several known intrinsic and extrinsic factors which affect anthocyanin content and composition in plants (Chalker-Scott 1999; Connor et al. 2002; Awad et al. 2001). Plant cultivars and maturation time are among the essential factors that can influence the phytochemical content, among which anthocyanins (Solomon et al. 2006; Fawole and Opara
2013; Josuttis et al. 2013). The objectives of this study were to evaluate the anthocyanins content and profile from different red cabbage cultivars at two maturity stages and evaluate their color characteristics and behavior under acidic and neutral pH. Knowing the anthocyanin composition of red cabbage cultivar and maturation time would help us to select the cultivar and maturation that could provide the desired characteristics for a specific application.
3.3. MATERIALS AND METHODS
3.3.1. Plant Materials
Seven red cabbage cultivars: Cairo, Kosaro, Integro, Buscaro, Azurro, Primero and
Bandolero (three heads from each cultivar), at two maturity stages (harvested 13 and 21 weeks after transplanting) were donated by Bejo Seeds Inc. (New York, USA). Cabbages were grown side-by-side during the summer season. Samples were shipped immediately after harvest and refrigerated until analyzed (within a week). The water content in each sample was determined
48
by placing 4-5 g of sample in a mechanical convection incubator (Precision Scientific, Buffalo,
NY) at 37 °C for 2 days to dry.
3.3.2. Extraction and Purification
Cabbage heads were sliced and ≃30 g were frozen with liquid nitrogen and kept frozen until analyzed the following day. The frozen materials were ground using a stainless steel
WARING Commercial Blender (New Hartford, CT) coupled with a 0.95L container. The acetone/ chloroform extraction procedure was adopted from Giusti and Wrolstad (1996). Frozen plant powder was mixed with 30ml acetone. The mixture was filtered through a Whatman #1 filter (Whatman Inc., Florham, NJ), and, the residual cake was washed with 70% aqueous acetone acidified with 0.1% formic acid (≃ 250ml) until the powder was white and the filtrate was clear. The filtrates were combined, transferred to a separatory funnel and mixed with 1 volume of chloroform. The phases were allowed to separate for 4-5 hr. The aqueous phase was collected and the residual acetone was evaporated using a Büchi rotavapor (Brinkmann
Instruments, Inc, Westbury, NY). The aqueous extract was purified using a Sep-Pak C18 cartridge (Waters Corp., Milford, MA). The cartridge was activated with MeOH, washed with acidified water before loading the sample. The cartridge was further washed with acidified water (0.1% formic acid) and the anthocyanins were recovered with acidified MeOH (0.1% formic acid). The MeOH was removed using the rotavapor; and, the volume was taken to 25 ml with acidified water (0.1% formic acid).
49
3.3.3. Anthocyanins content
The total monomeric anthocyanin content was determined by using the pH differential method according to Giusti and Wrolstad (29). The extract was diluted using pH 1 (0.025M potassium chloride) and pH 4.5 (0.4 M sodium acetate) buffers with a dilution factor of 100.
The solutions were allowed to equilibrate for 15 min in the dark. Absorbance was read on 1 cm pathlength cuvettes at 520 nm and 700 nm using a Shimadzu UV-Visible Spectrophotometer
(Shimadzu, Columbia, MD). The total monomeric anthocyanin was calculated based on the dry matter (DM) and fresh matter (FM) and reported as mg cyanidin-3-glucoside (Cy3G)/ 100g of sample using the following equation:
Total monomeric anthocyanin (mg/L)= [((A520 – A700) pH1 – (A520 – A700) pH4.5) x DF x 1000 X
MW]/(ɛ x P)
Where DF is the dilution factor, MW is the molecular weight (449.2 for Cy3G), ɛ is the molar absorptivity coefficient (26900 cm-1 mg-1 for Cy3G) and P is the cuvette pathlength.
3.3.4. Alkaline hydrolysis of anthocyanins
Alkaline hydrolysis (saponification) was adopted from Giusti and Wrolstad (1996).
Purified red cabbage anthocyanin extract (10 ml) was mixed with 10 ml of 10% KOH in a caped test tube and set aside for 15 min at RT in dark. The solution was neutralized using 2N
HCl untill the color had turned pink. The neutralized sample was then purified using a Sep-Pak
C18 cartridge (Waters Corp., Milford, MA) and prepared for HPLC analysis.
50
3.3.5. Chromatographic analysis
A Shimadzu Prominence® reverse phase high pressure LCMS coupled to a SPD-M20-
A photo-diode array and a single-quadrupole electrospray ionization (ESI) Mass
Spectrophotometer (Shimadzu Scientific, Inc., Columbia, MD). For data analysis, a LCMS
Solution software was used (Shimadzu Scientific, Inc., Columbia, MD). The column was a 100 x 4.5 mm Kinetex PFP 2.6 μm (Phenomenex Inc, CA, USA). The solvents were phase A, 4.5% formic acid in LCMS grade water; B, LCMS acetonitrile (Fisher Scientific Inc, Fair lawn, NJ); gradient: 0-50 min, 0-30% B. Injection volume was 20µl. Spectral data was obtained from 250 to 700 nm and elution of anthocyanins was monitored at 510-540 nm. Peak areas at this region were then integrated and normalized. The proportion of the total peak area of each individual anthocyanin was calculated and reported as percentage of total peak area at 510-540 nm.
For MS analyses, a 0.2 mL/min volume was diverted into the MS and ionized under positive ion condition using an electrospray probe. Data was monitored using total ion scan
(SCAN) (from m/z 200-1200) and selected ion monitoring at m/z 271 (Pg), m/z 287 (Cy), m/z
301 (Pn), m/z 303 (Dp), m/z 317 (Pt), and m/z 331 (Mv).
3.3.6. Color and spectrophotometric analyses
A ColorQuest XE colorimeter (HunterLab, Hunter Associates Laboratories Inc.,
Reston, VA) was used to measure the color characteristics (Hunter CIE LCh) of the samples.
The equipment was set for transmittance with specular included, and D65/10⁰ was used for the measurements. Samples were placed in a 1 cm pathlength plastic cuvette and C.I.E. L*, a*, b*, chroma (c*), and hue angle (h◦) were measured.
51
To measure the color in acidic condition, the extract was mixed (in triplicate) with distilled (DI) water (1:20 v/v). The pH of the solutions were measured after 30 min equilibration and was 3.5. To measure the color in neutral condition, the extract was diluted with water (1:2 v/v, in triplicate). The diluted solutions were then mixed with 0.1M potassium phosphate buffer pH 7 with dilution factor (DF) of 20. The maximum absorbances (λmax) in the visible range of the solutions neutral solutions were recorded using a Shimadzu UV-Visible Spectrophotometer
2450 (Shimadzu, Columbia, MD).
FD&C Red No. 3, FD&C No. 40 and FD&C Blue No. 2 (Noveon Hilton Davis, Inc,
OH) were also dissolved in DI water to concentrations that most closely matched the lightness
(L*) and chroma (c*) of the samples.
3.3.7. Statistical analysis
Seven different cultivars at two maturity stages (3 heads each) for a total of 42 samples were analyzed using Principal Component Analyses (PCA). PCA for total monomeric anthocyanin, non-acylated, mono-acylated, and di-acylated pigments (based on the proportional peak areas shown in Table 3.2) were performed. Autoscaling was used to normalize each variable before the analysis.
To compare the anthocyanin contents and profile, normality of the variables were first checked using Kolmogorov-Smirnov test (α=0.05). Analysis of variance (ANOVA) was then used to analyze the total monomeric anthocyanin and the proportion of each group of pigments separately using the following model: Yijk=µ+vi+tj+vtij+αk + εijk, where Y was the individual variable, µ was the grand mean, vi was the cultivar effect, tj was the harvesting time effect, vtij
52
was the interaction between the main factors, αk was the heads effect defined as random factor, and εijk was the random error of the model. When a significant difference was obtained (P-value
< 0.05), the Tukey means comparison test was used to compare each pair of means.
All statistical analyses were done based on at least three independent replicate samples from each individual head. Results were analyzed by Minitab 16 statistical software (Minitab
Inc).
3.4. RESULTS AND DISCUSSIONS
3.4.1. HPLC-PDA-MS and identification of major pigments
According to the HPLC-PDA data obtained at 510-540nm, up to 23 peaks were observed. In previous studies, up to 36 different anthocyanins have been detected in red cabbage
(Arapitsas et al. 2008; Wu and Prior 2005b; Charron et al. 2007). Figure 3.1 shows examples of anthocyanin profiles (Cairo and Integro extracts) obtained by HPLC-PDA. The eight major peaks, representing ≃ 90% of the total anthocyanins and common to all 7 cultivars at both maturity stages were selected for further identification and analyses. The ʎmax at the UV and
Vis ranges, molecular ions and fragments along with tentative identification of each peak are presented in Table 3.1. The pigments were identified using HPLC-PDA and HPLC-MS and compared with data reported in the literature (Wiczkowski et al. 2013; McDougall et al. 2007).
53
Max Plot - 510 - 540nm 1 5 8
2 ) 3 4 7 6 15 17 12 13 19 AmU 10 14 18 9 11 16 20 21 22 23
3 5
4 1 6 8
Absorbance ( Absorbance 2 7
0 10 20 30 40 50 Time (min)
Figure 3.1: HPLC chromatograms of two representative red cabbage anthocyanin extracts: second harvested Cairo (top) and first harvested Integro (bottom) red cabbage at 510-540 nm: refer to Table 1 for major peaks identification. HPLC conditions: solvent A, 4.5% formic acid in LCMS grade water; B, LCMS acetonitrile; gradient: 0-50 min, 0-30% B. The major peaks (1-8) were found in all other red cabbage samples.
Table 3.1: The PDA absorbance and the MS data for the RCA. 287 m/z was the major fragment + in all 8 peaks. RT: Retention time; λvis: λ vis-max; λacyl: λ of acylation; M : Mass ion.
Peak# RT (min) λvis (nm) λacyl M+ Identification (nm) 1 12.13 513 - 773, (287) Cy3diG-5-G 2 16.95 528 334 979, (287) Cy3diG-5-G+Sinapic* 3 27.41 523 313 919, (287) Cy3diG-5-G+p-Coumaric 4 28.33 523 326 949, (287) Cy3diG5-G+Ferulic 5 28.85 524 329 979, (287) Cy3diG5G+Sinapic 6 30.53 536 319 1125, (287) Cy3diG-5-G+Sinapic& p-Coumaric 7 31.55 536 330 1155, (287) Cy3diG-5-G+Sinapic&Ferulic 8 32.31 536 331 1185, (287) Cy3diG-5-G+Sinapic&Sinapic * Tentative identification.
54
As shown in Table 3.1, 287 m/z was the fragment in all 8 anthocyanins indicating that
Cy derivatives were the major aglycon as reported in previous studies (Wu and Prior 2005b;
Wiczkowski et al. 2013; Park et al. 2014; Scalzo et al. 2008; Sun et al. 2013). All the pigments were nonacylated, monoacylated and di-acylated derivatives of Cy3diG-5-G, which also confirmed by saponification (results are not shown). The acylating groups were aromatic acids: sinapic, ferulic, and p-coumaric acids (Table 3.1).
3.4.2. Anthocyanin content and proportion of the pigments
3.4.2.1. Principal component analysis of red cabbage anthocyanin extracts
The analysis of the objects (i.e. red cabbage cultivars at two maturity stages) is performed visually using the scores plot (Figure 3.2) where the objects are represented in function of the principal components (PCs). As showed in Figure 3.2, the PC1 correlated positively with mono-acylated pigments as opposed to the di-acylated pigments. PC2, on the other hand, was more affected by the non-acylated pigments and the total monomeric anthocyanin. The PC1 and PC2 extracted 83.7% and 12.8% of the total variances, respectively.
According to this analysis, the samples were clearly separated diagonally based on their maturation time. With maturation the percentage of di-acylation increased for most samples, so for most of the second harvested samples PC1 is negative. Also, since the early mature samples
(week 13) had slightly higher amount of total monomeric anthocyanin, PC2 tend to be more positive (Figure 3.2).
PCA is a proper way to obtain relevant information from the original variables into fewer new latent variables (PCs). According to our results, this analysis was helpful in classification of the samples at two maturity stages. Most samples with different maturity times
55
were separated based on the first two principle components. Azurro and Primero cultivars, however, were an exception since their pigment profiles were not significantly changed with maturation.
Figure 3.2: Correlation between the first two principal components and the variables, as well as, score plot with respect to cultivors and harvesting time for seven different red cabbage cultivars. a: total monomeric anthocyanin; b: di-acylated pigments; c: mono-acylated pigments; d: non-acylated pigments. Az: Azurro; Ba: Bandolero, Bu: Buscaro; Ca: Cairo; In: Integro; Ko: Kosaro; Pr: Primero; Symbols in black and gray represent harvest times after 13 or 21 weeks respectively.
3.4.2.2. Anthocyanin content
The average anthocyanin contents for the 13 and 21-week harvested plants were ≃1,442 and 1,269 mg Cy3G/100g DM respectively; and, these values for the fresh matters were ≃150 and 145 mg Cy3G/100g FM, respectively. Piccaglia et al. (2002) also reported the anthocyanin
56
content of three red cabbage cultivars in Italy to be more than 1000 mg/100g DM (Piccaglia et al. 2002). The anthocyanin content for the fresh weight red cabbage reported by Linus Pauling
Institute (Oregon State University) data base and Wu et al. (2006) were, however, 25 and
322±40.8 mg/100g FW, respectively. According to our findings, the cultivar made a difference in anthocyanin contents at both maturity stages.
As shown in Table 3.2, Buscaro and Integro, harvested after 13 weeks, had the highest anthocyanin content. Anthocyanin contents, however, did not change significantly from the first to the second harvest except for Buscaro (DM) cultivar that showed a lower anthocyanin content when plants were left longer on the ground (Table 3.2). Accumulation of anthocyanins can be explained by developmental factors which could be different in different varieties (Awad and others 2001).
3.4.2.2. Proportion of major pigments
Although red cabbage is known to have more than 20 different anthocyanins, relative proportions of 8 of them (Figure 3.1 and Table 3.1), represented ≃90% of the total anthocyanins, were measured and compared among cultivars and maturation stages as they are more likely to impact color and stabilities of the extract. The pigments were also grouped into non-acylated, mono-, and di-acylated.
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Table 3.2: Anthocyanin contents (total monomeric anthocyanin) and percentage of major pigments (% total peak area at 510-540 nm) in seven red cabbage cultivars in two different harvesting times. Different letters in the same column indicate significant differences (p < 0.05); DM: dry matter; FM: fresh matter; non-acylated pigments: peak 1; mono-acylated pigments: peak 2, 3, 4, and 5; di-acylated pigments: peak 6, 7, and 8 (See Table 3.1 for peak identity).
Total monomeric Harvest anthocyanin Mono- Non-acylated Di-acylated Cultivar Time (mg Cy3G/100g) acylated pigments (%) pigments (%) (week) pigments (%) DM FM
13 1111 e 109 gh 21.24 bc 67.63 a 4.56 g Primero 21 1026 e 104 h 26.84 a 63.71 ab 4.09 g
13 1660 ab 185 a 18.17 cde 65.22 ab 7.23 fg Integro 21 1637 ab 188 a 25.59 ab 53.32 cde 12.28 ef
13 1392 bcd 144 bcdef 15.85 def 69.33 a 9.19 fg Azurro 21 1217 de 137 cdef 19.33 cde 65.01 ab 9.21 fg
13 1247 cde 128 efgh 15.51 ef 55.43 cd 11.42 f Kosaro 21 1001 e 115 fgh 20.25 cd 46.74 ef 19.41 cd bcd bcde cde bc ef 13 1389 153 20.08 58.23 12.87 Cairo 21 1256 cde 168 ab 28.5 a 43.96 fg 17.74 de abc abcd f def bc 13 1517 165 13.1 48.67 24.54 Bandolero 21 1512 abc 165 abcd 19.07 cde 36.75 gh 29.1 b a abcd f cde bcd 13 1780 170 13.36 52.1 23.59 Buscaro 21 1236 de 137 defg 17.04 cdef 35.48 h 35.45 a
According to our results, non-acylated pigments represented an average of ≃19.6%
(±4.5) of the major pigments (Table 3.2). Charron et al. also found the percentage of non- acylated pigment in red cabbage to be 21.3% (Charron et al. 2007). Among the major pigments identified, the amount of mono and di-acylated anthocyanins were in average 54.4 (±12.6), and
15.8 % (±8.7), respectively (Table 3.2). Red cabbage has been identified as a highly acylated anthocyanin source according to previous research (Charron et al. 2007; Wu and Prior 2005b).
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Cultivars had a significant (p-value < 0.05) impact on pigment profile. The proportions of mono-acylated and di-acylated pigments were the major differences found among the cultivars. Primero and Azurro had the highest proportions of mono-acylated pigments and the lowest proportions of di-acylated pigments; whereas, Buscaro and Bandolero varieties had the lowest proportions of mono-acyltaed pigments and highest proportion of di-acylated pigments.
The effect of maturity on pigment profile was largely dependent on the cultivar for mono- acylated and di-acylated pigments (p-value <0.05); this interaction was not observed for the non-acylated pigments.
As shown in Table 3.2, the proportion of non-acylated pigments increased significantly in the 21-week mature plants except for Azurro and Buscaro cultivars that did not show a significant increase. More mature Primero, Cairo, and Integro had highest amount of non- acylated pigments. The proportions of mono-acylated anthocyanins were, however, decreased with longer maturation time, except for Primero and Azurro cultivars that did not show a significant decrease in mono-acylated pigments. The proportion of mono-acylated pigments was highest (≃67%) for cultivars Primero, Integro, Azurro first harvest as compared to the other cultivars (≃54%) harvested at the same time (Table 3.2). The proportion of di-acylated pigments varied remarkably among cultivars. They increased significantly in Kosaro and
Buscaro due to maturation. Buscaro and Bandolero, harvested after 21 weeks, had the highest
(≃30%) proportion of di-acylated pigments, while Primero (≃4%) had the lowest at both maturity stages (Table 3.2).
Since the plants were grown side by side, the main reason for the profile differences could arise from intrinsic factors such as plants genetics and enzymes and their activities throughout the maturation process which would influence the anthocyanin synthesis within the plant. Variation in anthocyanins structures can be correlated with alteration of single genes,
59
which influence the enzymatic step of anthocyanin synthesis pathways (Holton and Cornish
1995). The reason for accumulation of certain anthocyanins in certain cultivars could be due to the differences in the activities of different genes. Also, during the maturation, the activity of the genes controlling the synthesis of mono-acylated pigments could have slowed down, whereas, those responsible for the synthesis of non-acylated anthocyanins, or, for the addition of a second acyl group, may have remained active.
3.4.3. Color characteristics and stability
3.4.3.1. Color in acidic pH
Table 3.3 shows the color characteristics of solutions colored with RCA extract from the different cultivars at pH 3.5, and, their color comparisons to synthetic colorants FD&C Red
No. 3 and No. 40. The lightness of the samples was approximately between 60-77 (Table 3.3).
The synthetic colorant solutions were also adjusted to have a similar lightness. Extracts from all seven cultivars produced a deep pink color at pH 3.5 with a hue angle close to 350◦. Chroma of the samples were higher for solutions prepared with Bandolero and Buscaro extracts as opposed to Primero. The wavelength of maximum absorbances (λmax) of the samples were similar to Red
No. 3 (≃520nm) except for late harvested Bandolero and Buscaro cultivars (λmax ≃530nm).
Generally, samples showed more similar color characteristics to FD&C Red No. 3 than FD&C
Red No. 40 synthetic dyes (∆E1<∆E2). Under acidic pH, maturation of the plants did not seem to significantly affect the color of the solutions. Early harvested Azurro, Kosaro, Cairo, and
Integro, at the tested concentrations, were the samples that produced colors most similar to
FD&C Red No. 3 with similar λmax and ∆E between 6.1 to 8.8 (Table 3.3).
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◦ Table 3.3: C.I.E. L*a*b*, Chroma (c*), Hue (h ), and λmax of seven red cabbage cultivars extracts at two different harvesting time at pH 3.5 and their comparison with two FD&C synthetic red dyes. HT: Harvesting time; ∆E1: color difference with FD&C Red No. 3; ∆E2: color difference with FD&C Red No. 40.
HT λmax ◦ Cultivar (week) L* a* b* c* h (nm) ∆E1 ∆E2 Primero 13 76.2±0.8 45.3±0.6 -3.5±0.7 47.4±3.5 359.2±4.4 520±1.2 15.9±2 18±0.7 21 77.1±2 42.1±4 -2.7±0.7 42.2±4 356.3±0.9 520±1.4 18.5±4.2 17.9±1 Integro 13 67.4±2.6 59.5±1.7 -0.5±0.3 59.6±4.6 357.5±2.6 520±0.8 8.8±3.1 23.4±1.3 21 66.2±1.8 61±2.5 -1.9±1.2 61±2.5 358.2±1.2 520±1.1 9.3±2.2 24.8±1.4 Azurro 13 69.6±3 56.1±1 -2.8±2 56.6±5.3 357.7±2.7 520±0.6 6.8±2.9 20.7±2
6
1 21 71.2±2.9 53.2±4.7 -2.9±1.4 53.3±4.6 356.7±1.9 520±0.5 9.1±2.2 20.2±1.4
Kosaro 13 69±3.2 57±3.5 -5.9±1.2 57.3±5 354.1±2.6 520±0.3 6.1±2.7 23.7±3.7 21 69.5±1.5 55.5±2.4 -7.1±0.9 56±2.4 352.7±1 520±0.8 7±0.8 24.8±1.6 Cairo 13 68.9±3.5 57.1±1.6 -4.5±0.6 55.9±3.6 357±1.4 520±1.4 6.6±3.2 23±1.3 21 64.9±1.5 63±2 -4.3±0.4 63.2±2 356.1±0.5 520±0.6 10.1±2 28.1±1.6 Bandolero 13 62.5±1.6 67.3±4.3 -4.7±1.3 68.4±1.5 356.8±4.4 520±0.8 14.8±3 32±1.5 21 60.7±1.8 68.2±2.1 -4.8±1.5 68.4±2 356±1.4 530±1.2 15.9±2.7 33.6±1.3 Buscaro 13 62.9±4.1 66.9±2.3 -3.7±1 64.5±1.2 357.9±1.4 520±0.9 12.8±3 29.9±4.4 21 61.7±4.2 66±4.5 -6.4±2.9 66.4±4.2 354.4±2.8 530±1.5 14.2±5.7 33±3 FD&C Red No. 3 74.2 59.99 -6.1 60.3 354.19 520 FD&C Red No. 40 75.64 45.11 14.48 47.38 17.79 500
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3.4.3.1. Stability in neutral pH
The spectral characteristics of the samples at pH 7 were monitored over 72 hr refrigerated storage. Figure 3.3 shows the changes in λmax for three representative samples.
Solutions colored with Buscaro and Bandolero red cabbage extracts showed the highest variations in the λmax while solutions colored with Primero extract showed the least changes over the 72 hr. Most of the changes observed happened during the first 5-6 hr (Figure 3.3).
615
610
605 Bandolero
λmax 600 Buscaro Primero 595
590 0.5 1 3 6 10 72
Time (hr)
Figure 3.3: Change in λmax for the 2nd harvested Buscaro, Bandolero, and Primero at pH 7 over 72 hr refrigeration. Most color changes happened during the first 6hr of storage.
Table 3.4 shows the λmax of the all 7 varieties 30 min after the extracts were mixed with buffer pH 7 and 6 hr afterward (refrigerated storage). The λmax for samples containing significantly higher amount of di-acylated anthocyanins (Buscaro and Bandolero) at this pH
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seemed to be the highest (≃600nm) compared to the other samples. Torskangerpoll and
Andersen (2005) also investigated the absorbance and color change in Cy 3-(2”-(2’”- sinapoylglucosyl)-6”-sinapoylglucoside)-5-glucoside (a di-acylated pigment isolated from red cabbage) at different pH values and also found that at pH 7.2 the pigment had λmax of ≃605nm
(Torskangerpoll and Andersen 2005). A bathochromic effect of up to 7nm was observed for most samples after 6 hr refrigerated storage. More mature Buscaro and Bandolero varieties produced a λmax (≃610nm) similar to that of FD&C Blue No. 2. For other samples such as
Primero and Integro, however, minute bathochromic effects were observed (Table 3.4).
Anthocyanins tend to have lower stabilities at neutral to alkaline pH values (Fossen et al. 1998; Cabrita et al. 2000). After mixing with buffer pH 7 followed by 6 hr of refrigeration, the color degradation was highest (≃53%) and lowest (≃19%) for the second harvested Primero and Buscaro, respectively (Table 3.4). Higher stabilities of the Buscaro and Bandolero cultivars at the tested pH values could be explained by the larger number of di-acylated anthocyanins which have higher stabilities due to the pigments intramolecular and/or intermolecular copigmentation, and self-association reactions (Giusti and Wrolstad 2003). Torskangerpoll and
Andersen (2005) also demonstrated that di-acylation of anthocyainins increased their color stabilities.
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Table 3.4: λmax of the red cabbage extracts at two different harvesting times measured after 30min and 6hr refrigeration storage in buffer pH 7compared to FD&C synthetic Blue No. 2, and stabilities (% degradation) during this time. HT: Harvesting time. %Degradation= 100- ((absorbance at the λmax after 6 hr/ absorbance at the λmax after 30 min) x100).
HT λmax (nm) Cultivar %Degradation (week) 30 min 6 hr 13 591.4±0.6 591.6±0.8 50.1±5.5 Primero 21 591.3±0.4 591.9±1 53.3±8.9 13 591.4±1.1 592.1±0.1 38.9±5.4 Integro 21 598.6±2.3 599.7±2 44±9.9 13 590.8±0.8 593.5±0.5 41.5±4.9 Azurro 21 595.9±0.7 597.8±2.1 49.3±7.9 13 596.3±1 599.9±1.4 31±3.3 Kosaro 21 602±0.3 604.8±0.8 39.5±5.2 13 593.6±0.2 595.3±4.2 42.1±8.4 Cairo 21 599.7±2.4 602.9±0.5 36.7±8.2 13 601.8±1.2 609.4±1.2 27.1±4 Bandolero 21 599.7±2.3 610.1±1.6 22.5±3.8 Buscaro 13 600.1±0.7 606.7±1.9 28.6±2.3 21 601.8±0.4 610.1±1.8 19.1±3.8 FD&C Blue No. 2 610 610 -
The color characteristics of pH 7 buffer solutions colored with the different red cabbage
extracts after 6 hr refrigeration compared to FD&C Blue No. 2 are shown in Table 3.5. The
lightness of the samples ranged between ≃ 60 to 85. The Chromas were higher for Bandolero
and Buscaro cultivars at the tested pH value (Table 3.5). The synthetic FD&C Blue No. 2 was
diluted until the lightness was similar to that of the evaluated samples. ∆E of the samples when
compared to this synthetic dye was between 17.6 and 26.4 (Table 3.5). The color difference
with FD&C Blue No. 2 at the tested concentration for the late harvested Kosaro sample was the
smallest; however, the λmax for the second harvested Buscaro and Bandolero cultivars were
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closer to this value for FD&C Blue No. 2 (Table 3.4 and 3.5). Acylation of anthocyanins with
aromatic acids (e.g. cinnamic acid) has proven to increase the λmax and shift the hue angle to
purple color under acidic conditions (Giusti et al. 1999; Stintzing et al. 2002). As shown in
Table 3.2, late harvested Buscaro and Bandolero cultivars, with the highest percentage of di-
acylated pigments also exhibited the highest λmax under neutral conditions.
Table 3.5: C.I.E. L*a*b*, Chroma (c*), and Hue (h◦) of seven red cabbage extracts at two different harvesting time after 6 hr refrigeration storage at pH 7 compared to FD&C Blue No. 2. HT: Harvesting time; ∆E: color difference with FD&C Blue No. 2.
HT Cultivar L* a* b* c* h◦ ∆E (week) 13 83.6±0.6 2.6±0.2 -9.9±0.6 10.2±2.4 284.6±1.5 25.7±0.5 Primero 21 84.8±2 2.1±0.6 -8.7±2.2 9±2.2 284.5±5.1 26.4±2 13 70.5±2.9 1.7±2.2 -22.4±2.8 22.5±2.7 274.7±5.8 21.4±1 Integro 21 69.9±3.2 -0.2±0.6 -23.5±2.6 23.5±2.6 269.5±1.3 19.8±0.7 13 77.7±1.5 1.4±0.2 -15.8±1.5 15.9±1.4 275.3±1.3 21.2±1.3 Azurro 21 78.5±3.7 1.2±0.4 -15.2±3.1 15.3±3.1 274.7±1.9 21.5±1.3 13 72.1±1.5 -1.3±0.7 -22.1±1.9 22.1±2.4 266.7±1.3 17.9±2.3 Kosaro 21 71.9±2.5 -1.8±1.8 -22.5±2 22.6±2.2 265.6±4.2 17.6±1.1 13 69.3±4.4 -0.5±1.7 -24.7±4.2 24.7±4.2 269.2±3.5 20.2±1.1 Cairo 21 66.2±3.4 -2.3±0.6 -27.9±3 28±3 265.3±0.8 20.2±1.8 13 60.8±1.8 -4.9±0.7 -31.9±2.7 32.3±4.3 261.2±1.4 22.9±1.2 Bandolero 21 59.6±3.1 -5.7±0.6 -33.2±2.6 33.7±2.5 260.2±1.6 23.9±3.6 Buscaro 13 63.4±3 -3.8±1.5 -29.9±2.9 30.2±2.1 262.9±2.9 21.2±1.6 21 63.4±1.4 -5.4±0.2 -30.6±0.8 31±0.8 260±0.7 20.3±1.4 FD&C Blue No.2 78.06 -17.95 -24.28 30.19 233.52
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3.5. CONCLUSION
In conclusion, anthocyanin content varied among red cabbage cultivars and leaving the cabbages in the ground for additional time did not increase the pigment content. The pigment profiles changed among the cultivars and maturation stages. For most cultivars, the amount of non-acylated and di-acylated pigments increased over maturation time as opposed to the mono- acylated pigments. Cultivars with lower proportions of di-acylated pigments better reproduced the color of FD&C Red No. 3 under acidic conditions, while cultivars with higher proportions of di-acylated pigments better matched the colors of FD&C Blue No. 2 at neutral pH. Pigment profile also affected the color stability at neutral pH. For future studies, however, year to year variability of these cultivars should be investigated.
3.6. ACKNOWLEDGMENT
We are thankful to MARS Chocolate, NJ for providing funding for the project. We also thank Ken McCammon and Bejo Seeds, Inc. for providing the plant materials. Also especial thanks to Marçal Plans Pujolras from UPC (Universitat Politècnica de Catalunya) for his help with data statistical analysis.
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CHAPTER 4: SOLID PHASE FRACTIONATION TECHNIQUES FOR SEGREGATION OF RED CABBAGE ANTHOCYANINS WITH DIFFERENT COLORING AND STABILITY PROPERTIES 1,2
4.1. ABSTRACT
The food industry is looking for colorants from natural sources with increased stability,
specific color properties, and pure color tones. Red cabbage anthocyanin (RCA) extract
contains Cy anthocyanins in various non-, mono-, and diacylated forms capable of forming
various colors from red to blue. Our objective was to develop a simple SPE fractionation
method to segregate diacylated anthocyanins with higher stabilities and better blue coloring
properties. Two different cartridges with SCX (Strong Cation Exchange) and C18 sorbents were
used. In both cases, the resin was activated with MeOH and washed with acidified water (0.01%
HCl). RCA was loaded into the cartridge and washed with acidified water. Anthocyanins were
eluted from the SCX cartridge using a combination of different buffers (pH 6-8) and MeOH
1 Neda Ahmadiania, Rebecca J. Robbinsb, Thomas M. Collinsb, M. Monica Giustia* a Department of Food Science and Technology, The Ohio State University, 2015 Fyffe Road, Columbus, OH 43210, USA,b Analytical and Applied Sciences Group, Mars Inc., 800 High Street, Hackettstown, New Jersey 07840 2 To be published in Journal of Chromatography A.
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(20-70%); and, from the C18 cartridge using acidified (0.01% HCl) MeOH (30-100%) or EtOH
(10-100%). When SCX cartridge was used, washing with
buffer pH 6+20% MeOH followed by elution using buffer pH 8+70% MeOH was effective in segregating the diacylated pigments; however, some anthocyanins were permanently retained by the resin. When using the C18 cartridge, washing with 32%MeOH or 18.5%EtOH and eluting with 100% alcohol, respectively, resulted in high (> 93%) recovery of diacylated pigments. The C18 MeOH combination was the most effective for isolating the target diacylated RCA and produced a better blue color with increased stability.
Keywords: red cabbage, diacylated anthocyanins, fractionation, stable blue color
4.2. INTRODUCTION
The food industry is looking for natural alternatives to synthetic colorants that exert improved stabilities, more pure tone, and certain coloring properties. Anthocyanins are water soluble pigments found in many fruits and vegetables with potential health benefits that are currently being used as alternative to many synthetic dyes. Red cabbage (Brassica oleracea L.) has high anthocyanin content (more than 10 g/kg DM) with low sensitivity to light and heat treatment capable of producing a blue color in proper pH values (Dyrby et al. 2001; Piccaglia et al. 2002a; Ahmadiani et al. 2014).
More than 30 anthocyanin pigments have been detected in RCA extracts (Arapitsas et al. 2008; Wu and Prior 2005a; Charron et al. 2007). The major RCA is Cy-3diG-5G that can be
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found as in the non-, mono-, and diacylated forms; and the acyl groups are mainly aromatic acids such as p-coumaric, sinapic, and ferulic acids (McDougall et al. 2007; Wiczkowski et al.
2013; Wu and Prior 2005b; Park et al. 2014; Scalzo et al. 2008; Sun et al. 2013; Ahmadiani et al. 2014).
Formation of stable blue color by anthocyanins has been associated with the molecular structure of the pigments and the media that the pigments are in. Anthocyanin backbones, as well as, numbers and types of acyl groups are important factors to determine the actual color behaviors at various pH values (Giusti and Wrolstad 2003). In general, additional hydroxylations on the B-ring (Figure 2.3) results in bathochromic shifts which yield a bluer hue
(Mazza and Miniati 1993; Cabrita et al. 2000; Giusti and Wrolstad 2003; Torskangerpoll and
Andersen 2005). Aromatic acylation of anthocyanins has also been shown to have bathochromic effects, as well as, increase the stability of the pigments (Stintzing et al. 2002; Giusti et al.
1999), and the effects are more pronounced with increasing number of aromatic acid acylations
(Giusti et al. 1999; Torskangerpoll and Andersen 2005). Ahmadiani et.al (2014) evaluated RCA extracts from different varieties at two different maturity stages and found that the RCA with higher amount of diacylated pigments better able to produce a blue color in neutral to alkaline conditions, as opposed to, the RCA with higher non and mono acylated pigments. Therefore, in order to obtain an anthocyanin source with high stability and better blue color properties and a more pure color tone, it is desirable to develop a simple, effective, and economical fractionation method to isolate the diacylated RCA pigments with similar coloring properties.
Solid phase extraction (SPE) is a simple and affordable separation technique widely used for sample preparation to segregate selected compounds. The cartridge devices typically consist of a short column (an open syringe barrel) containing a packed silica-based sorbent. The
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sample is loaded (often in a liquid phase) into the column and the target compounds will be bound by the resin (sorbent). Different mobile phases will then be used to wash off undesirable compounds from the solid phase where the target compounds are retained. The target compound will be recovered by elution using a liquid that shows higher affinity for the target compounds that the resin (Poole 2003). SPE (usually C-18) is often used as a method to concentrate anthocyanins extracts by the removal of undesirable compounds such as sugars, small acids and some phenolics, and amino acids from anthocyanin extracts (He and Giusti 2011). Solid phase extraction techniques have also been used to isolate 3,5-diglucosides anthocyanins and their derivatives from the supramolecules (Asada et al. 2014).
The aim of this research was to develop a simple fractionation method based on RCA properties and their interactions with SPE resins and solvents to segregate anthocyanins with higher stabilities and pure blue color properties.
4.3. MATERIALS AND METHODS
4.3.1. Sample preparations
Red cabbages were donated by Bejo Seeds Inc. (Geneva, NY, USA) and the pigments were extracted by acetone and chloroform extraction (Ahmadiani and others 2014). Individual
RCA peaks (Figure 4.2, Table 4.2) were semi-purified (section 4.3.1.1). FD&C Red No. 3 and
40, and FD&C Blue No.1 and 2 (Noveon 146 Hilton Davis, Inc., Cincinnati, OH, USA) were dissolved in distilled (DI) water.
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4.3.1.1. Semi-preparatory HPLC
A Shimadzu Prominence® semi-preparative HPLC coupled with a SPD-M20-A photo- diode array was used to isolate PA. The column was a 250 x 21.20 mm Luna PFP 5 μm
(Phenomenex Inc, CA, USA). The solvents were phase A, 4.5% formic acid in LCMS grade water; B, LCMS acetonitrile (Fisher Scientific Inc, Fair lawn, NJ). Gradient was 0-30 min, 10-
13% B. The flow rate was 12 ml/min and the injection volume was 2000 l. LCMS Solution software (Shimadzu Scientific, Inc., Columbia, MD) was used for spectral analysis. The solvents were evaporated and the sample collected in MeOH (stock). The three pigments in each set (R520 and R530) were mixed in 1:1:1 ration based on their molarities (stock).
4.3.2. SPE fractionation
To begin, an exact aliquot of the red cabbage extract was dissolved in acidified DI water (0.01% HCl) and the monomeric anthocyanins were measured using the pH differential method as described by Wrolstad and Giusti (1999). Two different SPE cartridges with a combination of different solvents, as shown in Table 4.1, were used. The cartridges were Strong
Cation Exchange (SCX) (5g sorbent/ 20cc) (Phenomenex Inc, CA, USA) and C18 (2g sorbent/
12cc) (Waters Corp., Milford, MA).
The cartridge was activated using MeOH, washed with acidified water, and anthocyanins solution was loaded into the cartridge. The mass of the anthocyanins loaded was
1% of the sorbent mass. The loaded sample was then washed again using acidified water
(0.01% HCl).
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For the SCX fractionation methods, the buffers used were potassium phosphate buffers
(0.1M) pH 6, 7, and 8. DI water was used to make the buffer solutions. Each fraction was collected by ∼40ml (two volumes of the cartridge) of the solvent listed in Table 4.1. The final fraction was eluted using enough amount of the solvent. For method S1, S2, and S3 the cartridge was conditioned with one volume (20ml) of buffer (the same as the one used for the collection of each fraction) prior to collecting the fraction. For method S4 and S5 the conditioning step with pure buffer was skipped. Prior to its analysis using HPLC, each fraction was then acidified with a few drops of HCl (6N) and purified using a C18 (Waters Corp.,
Milford, MA) cartridge to eliminate the salts. The cartridge was activated with MeOH and washed with acidified water (0.01% HCl) before loading the sample. The sample was then loaded and washed with 3-4 volumes of acidified water (0.01% HCl) and eluted using acidified
MeOH (0.01%). MeOH was evaporated using a Büchi Rotavapor (Brinkmann Instruments, Inc,
Westbury, NY), and, the sample was collected using acidified water (0.01% HCl) and prepared for HPLC analysis.
For the C18 fractionation technique, the solvents were MeOH and EtOH mixed with DI water (acidified with 0.01% HCl (6N)) used in different percentage for different methods (Table
4.1). Each fraction was collected using enough of the solvent so that no more colored pigments eluted (which were approximately equal to two volumes of the cartridge). To prepare the samples for the HPLC analysis, the MeOH and EtOH in each fraction were evaporated using a
Büchi Rotavapor (Brinkmann Instruments, Inc, Westbury, NY), and, the sample was collected using acidified water (0.01% HCl). The monomeric anthocyanins in each fraction were also measured using the pH differential method.
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Table 4.1: Solvents used in each fractionation method using SCX (S1-S5) and C18 (C1-C6) resins.
Solvent Solvent Solvent 1st 2nd 3rd Method Buffer MeOH Buffer MeoH Buffer MeOH Fraction Fraction Fraction pH (%) pH (%) pH (%) S1 S1-F1 6.0 25 S1-F2 7.0 25 S1-F3 8.0 25 S2* S2-F1 6.0 25 S2-F2 7.0 25 S2-F3 8.0 25 S3 S3-F1 8.0 25 S3-F2 8.0 70 S4 S4-F1 6.0 25 S4-F2 8.0 70 S5 S5-F1 6.0 20 S5-F2 8.0 70 (%) (%) (%) C1 C1-MF1 30 MeOH C1-MF2 40 MeOH C1-MF3 100 MeOH C2 C2-EF1 10 EtOH C2-EF2 20 EtOH C2-EF3 100 EtOH C3 C3-MF1 40 MeOH C3-MF2 100 MeOH C4 C4-MF1 32 MeOH C4-MF2 100 MeOH C5 C5-EF1 17 EtOH C5-EF2 100 EtOH C6 C6-EF1 18.5 EtOH C6-EF2 100 EtOH * Final fraction (S2-F4) was collected using buffer pH8 and 70% MeOH.
4.3.3. HPLC-MS and chromatographic analysis of the fractions
To evaluate the proportion of the pigments in each fraction the fractions were injected
into an HPLC-MS instrument: Shimadzu Prominence® reverse phase analytical HPLC and a
mass spectrophotometer (MS) (Shimadzu Scientific, Inc., Columbia, MD). The MS was coupled
with a single-quadrupole electrospray ionization (ESI system) (Shimadzu Scientific, Inc.,
Columbia, MD). The analytical column was a 100 x 4.5 mm Kinetex PFP 2.6 μm (Phenomenex
Inc, CA, USA). The detector, the solvents, and the software for data analysis were similar to
those described in section 4.3.1.1. The gradient was 0-50 min, 0-30% B, and the injection
volume was 60µl. Spectral data was obtained from 250 to 700 nm. The peaks were integrated at
510-540 nm, and, the area of each individual peak was calculated by the software. The
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proportion of the area of R530 pigments to the total peak area were then calculated and reported as %R530.
Mass Spectroscopy (MS) was used to confirm the identities of the pigments. The flow rate was 0.2 mL/min, and, the positive ion condition was used. Total ion scan (SCAN) (from m/z 200-1200) and selected ion monitoring (at m/z 287) were also applied.
Purity of the isolated pigments was checked by dividing the % Area Under the Curve
(AUC) at 510-540 nm to the max plot (250-700 nm), and, the peaks with purity above 90% were used for further analysis.
4.3.4. Calculation of R530 recovery and anthocyanin yield
The yield of anthocyanins in each fraction was calculated as the percent of total amount of monomeric anthocyanins (mg Cy-3G) that was loaded into the cartridge. The recovery of
R530 pigments were also calculated as the percent of the initial amount of R530 peaks recovered in the sample. The initial amount of the monomeric anthocyanins and the amount of
R530 in the RCA were considered to be 100%.
4.3.5. Measurement of spectra and degradation of PA, R520 and R530 pigments
To measure the spectral characteristics of PA, R520, and R530 pigments, an exact aliquot of the stock solution was pipetted into each well of a 96 well-plate (Greiner Bio-One,
Frickenhausen, Germany), and the plates were allowed to dry in a vacuum chamber for ∼2 hours. To obtain the final concentration of 1x 10-4 mol/L, buffer solutions were added to the dry pigments in each well. The buffers for pH 1, 2-5, and 6-8 were made using KCl (0.25M), citrate
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(0.1M), and phosphate (0.1M) solutions (Appendix I). The dissolved pigments were allowed to equilibrate for 15min in the dark at room temperature. A SpectraMax® 190 UV-Vis Microplate
Spectrophotometer (Molecular Devices, Sunnyvale, CA) was used to record the spectra. The spectra were collected in 1nm increments between 400-700nm wavelengths.
Degradations of the pigments in each pH value were also calculated using the equation below.
% Degradation = 100-((absorbance at λmax after 8hr/ absorbance at λmax after1hr) x100)
4.3.6. Conversion of spectral data to color
To obtain the color components of the samples the color spacing equations in accordance with CIE 1964 supplementary standard colorimetric observer (10 ° observer angle),
5nm intervals (400-700 nm) were used to obtain the X, Y, and Z tristimulus values as shown in
Equation 1 (CIE 2002; Konica Minolta Sensing 2007).
X= 1/N ΣxiSiTi
Y= 1/N ΣyiSiTi Equation 1 Z=1/N ΣziSiTi
Where xi, yi, and zi are the color matching functions for the standard observer, Si is the relative spectral power distribution of an illuminant D65, T is the % transmission of the sample,
* * * and N= ΣyiSi. The X, Y, and Z values were then transformed into the CIE-L a b ColorSpace using the Equation 2 (Kheng 2002; Konica Minolta Sensing 2007).
* L = 116[f(Y/Yr) – 16/116]
* a = 500[f(X/Xr) – f(Y/Yr)] Equation 2
* b = 200[f(Y/Yr) – f(Z/Zr)]
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f(X/Xr) = (X/Xr)1/3 (same formula for f(Y/Yr) & f(Z/Zr), input Y or Z values for X)
Where Xr=94.811, Yr= 100.00, and Zr=107.304, and Xr, Yr, and Zr were normalized to
Y=100.
The L*C*h ColorSpace values were also calculated using Equation 3 (Konica Minolta
Sensing 2007).
* L = 116[f(Y/Yr) – 16/116]
C* = √(푎 ∗)2 + (푏 ∗)2 Equation 3
h = tan-1(b*/a*)
Finally, in order to calculate the color difference (∆E) of the color compared to the
FD&C colorants Equation 4 was used (Kheng 2002).
∆E= [(∆L*)2+(∆a*)2+(∆b*)2]1/2 Equation 4
The ∆L*, ∆a*, and ∆b* values were the difference of L*, a*, and b* values of each sample
(in each pH values) compare to the measured L*, a*, and E* of the FD&C dye solutions.
4.3.7. Color analysis of the fractions
The color components of the two fractions from the most effective method (C4-MF1, and C4-MF2) as well as RCA were measured by A ColorQuest XE colorimeter (HunterLab,
Hunter Associates Laboratories Inc., Reston, VA, USA) using a 1 cm path length plastic cuvette and D65/10°.
The total % transmittance values for the same samples obtained using the colorimeter were then converted to absorbance value using the equation bellow
A=log10(100/%T) Equation 5
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Where A was the absorbance and T was the transmittance.
4.3.8. Statistical analysis
Percent degradation and ∆E (for each of the four groups of comparison) were analyzed using analysis of variance (ANOVA), where pigment and pH were defined as fixed factors, and fixed factor interaction was included in the model.
Recovery in % of R530 pigments was analyzed using one-way ANOVA, where each fraction was considered as an independent level of the fixed factor. Prior to each analysis normality of the variables were checked using the Kolmogorov-Smirnov test (α = 0.05).
In all cases, when a significant difference (P value < 0.05) was obtained, to compare each pair of means, the Tukey means-comparison test was used. Three independent replications were used for the statistical analyses. Statistical analysis was carried out using Minitab 16 statistical software (Minitab Inc.)
4.4. RESULTS AND DISCUSSIONS
4.4.1. Spectra and color analysis
To show the different color properties of the RCA with different molecular structures, the colors of PA (Cy-3diG-5G), R520 (monoacylated Cy-3diG-5G), and R530 (diacylated Cy-
3diG-5G) (Figure 4.1and 4.2, Table 4.2) in different buffer solutions pH 1-8 were analyzed.
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Color behavior of each group of pigment was different at different pH values. All groups of pigment lost their color strength at middle pH range mainly between pH 4 to 6. As shown in Table 4.3, PA had the highest value of chroma (>10) at pH 1, 2, 3, 7 and 8. The color for R520 pigments faded noticeably at pH 5 and 6. R530 pigments, however, maintained some color (chroma > 18) even in medium pH range where the other two groups were almost colorless (Table 4.3).
Table 4.2: The MS data of the RCA pigments (for the HPLC chromatogram refer to Figure 2). Cy-3diG-5G: Cyaniding-3-diglucoside-5-glucoside. M+: Mass ion.
Peak # M+* Identification PA - 773 Cy-3diG-5G 1 919 Cy-3diG-5G+p-Coumaric R520 2 949 Cy-3diG-5G+Ferulic 3 979 Cy-3diG-5G+Sinapic 4 1125 Cy-3diG-5G+Sinapic&p-Coumaric R530 5 1155 Cy-3diG-5G+Sinapic&Ferulic 6 1185 Cy-3diG-5G+Sinapic&Sinapic * 287 m/z was the major fragment in all peaks.
In acidic pH range 1-3, the hue of the color had different shades of red, depending on the type of pigments. At this pH range, PA had more orange shade of red (15> hue> 0.8); whereas, R520 and R530 groups had a rosy pink color with hue angle between 342 and 355
(Table 4.3). When compared to the FD&C red colorants (Red 3 and 40) at this pH range, the colors were generally more similar to Red 3 than Red 40. The lowest color difference was observed for R520 and R530 pigments at pH 1 and 2 (13.3> ∆E >7.7) (Table 4).
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In neutral to alkaline pH range, as expected, the hue turned into a purple-blue color; however, the hues of the blue were different for different type of pigments. For PA at pH 6 a weak purple color with a pink shade (hue= 345) was observed. R520 and R530 pigments, however, had more shade of blue (319>hue>226) (Table 4.3). When they were compared to
FD&C synthetic dyes blue 1 and 2, the color differences were generally lower for blue 2; however, R530 had the lowest (∆E= 18.8) color differences with blue 1 at pH 8. R530 and R520 pigments had lowest color difference with blue 2 especially at pH 7 and 8 (29.4> ∆E >10).
According to these data, amounts of acylation affected the color behavior of the anthocyanins. Di-acylated pigments increased the chroma of the color in the pH range that other non and mono-acylated anthocyanins were essentially colorless. Acylation also caused the hue angle to move counter clockwise toward more violet, purple and blue color in all the evaluated pH values. Torskangerpoll and Andersen (2005) also showed very similar results when they evaluated non- and di-acylated cy based anthocyanins (Cy 3-(2″-glucosylglucoside)-5-glucoside and Cy 3-(2″-(‴-sinapoylglucosyl)-6″-sinapoylglucoside)-5-glucoside ) in pH range between
1.1-10.5. They demonstrated that addition of the aromatic acyl groups to the anthocyanin molecule resulted in more negative hue angle and higher chroma values in the middle pH range
4.1 to 6.8.
According to the spectral data, λmax of the absorbance spectra were varied from 510 nm to 620 nm depending on the pH of the solution and the group of pigments analyzed. The λmax of the spectra were ∼20 nm larger for R530 pigments compared to PA in all the evaluated pH values. The difference between the λmax of the pigments and FD&C Red No.3 were the lowest
(3-5 nm) for R530 pigments at pH 1 and 2 when the color also had good tinctorial strength.
R530 pigments also had lowest λmax difference (10 nm) with blue 1 and 2 at pH 8 (Table 4.3).
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Table 4.3: Color and spectral characteristics of non-, mono- , and diacylated Cy-3diG-5G from red cabbage at different pH values and their color difference with synthetic red and blue FD&C colorants. Spectral data obtained after 15 min and 8 h at room temperature storage. For the pigments identities refer to Table 4.2.
Color Components Spectral Components pH Peak(s) L* C* h◦ ∆E1† ∆E2‡ λmax Degradation(%) PA 77.4±0.9 55.9±0.3 15.3±0.2 22.9±0.2 d 32.4±0.4 a 510 2.1±0.6 1 R520 77.9±0.3 51.2±0.6 355.3±0.7 7.7±0.6a 51.3±0.1c 520 1.8±0.6 R530 76.9±0.7 50.5±1.4 345.3±0.3 13.3±1.5ab NA 530 2.5±0.4 PA 79.8±0.2 50.8±0.5 9.7±0.3 18.6±0.1 cd 41.5±0.7 b 510 2.6±0.2 2 R520 79.2±0.2 49.9±0.5 353.7±0.2 11.5±0.7a 54.1±0.3c 520 0.9±0.3 R530 76.0±0.1 52.2±0.4 345.1±0.1 12.2±0.4ab NA 532 3.7±0.5 PA 88.9±0.1 28.0±0.2 0.8±0.3 35.8±0.2 f NA 510 6.6±0.9 3 R520 85.8±0.2 33.7±0.4 349.4±0.1 29.3±0.5e NA 520 3.0±0.5 R530 77.3±0.4 48.3±0.7 342.9±0.1 16.3±0.7bc NA 535 5.2±0.5
PA 95.8±0.5 8.4±0.0 356.8 ±2.1 NA NA 515 7.5±0.6 4 R520 93.3±0.2 13.7±0.1 342.7±0.1 50.7±0.5h NA 525 5.0±0.4 R530 81.6±0.6 35.4±0.3 335.6±0.4 30.0±1.1e NA 535 0.7±1.0 PA 97.8±0.1 3.1±0.1 353.3 ±6.6 NA NA 525 12.7±1.6 5 R520 96.1±0.0 5.7±0.1 329.9±0.7 NA NA 540 9.4±2.5 R530 85.9±0.5 21.4±0.8 321.3±0.6 45.4±0.7g NA 545 0.4±0.6 PA 97.5±0.0 2.5±0.0 344.6 ±0.7 NA NA 535 17.9±0.2 6 R520 95.4±0.3 4.8±0.1 319.2±0.3 NA NA 545 19.1±2.5 R530 83.3±2.8 18.6±1.7 304.3±2.8 41.6±1.7d 58.8±0.9f 555 7.3±0.12 PA 88.8±0.2 10.3±0.2 320.6 ±0.2 46.8±0.1 e NA 545 74.1±0.9 7 R520 84.2±0.2 15.9±0.3 299.9±0.2 40.7±0.1d 57.1±0.1e 557 68.1±3.1 R530 77.7±0.6 23.9±0.1 276.1±0.6 29.4±0.1b 48.1±0.1c 590 28.6±1.2 PA 54.5±0.4 51.7±0.3 271.7 ±0.4 32.7±0.7 c 51.6±0.6 d 595 89.1±0.1 8 R520 66.6±0.2 40.4±0.2 251.9±0.2 11.0±0.2a 32.1±0.2b 600 64.5±0.1 R530 73.4±0.3 38.0±0.3 226.0±0.3 10.0±0.4a 18.8±0.4a 620 20.6±0.8
Red 3 74.2 60.3 354.2 527 Red 40 63.8 76.5 36.0 500 Blue 1 68.2 54.1 215.7 630 Blue 2 69.2 42.4 237.4 610 † DealtaE1: pH 1-5 color difference with Red 3; pH 6-8 color difference with Blue 2. ‡ Delta E2: pH 1-3 color difference with Red 40; pH 6-8 color difference with Blue 1. Degradation (%) = 100-((absorbance at λmax (8h)/ absorbance at λmax (15min)) x100). NA= Not Applicable, color intensity was low (<10) to be quantified, or ∆E was higher than 60. Letter superscripts compare the ∆E against the target synthetic color. Different letters indicate significant differences (p < 0.05).
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4.4.2. Pigments degradations
Pigments degradations demonstrated by percentage loss in the absorbance spectra (at
λmax) were also monitored during 8 h storage at room temperature. Figure 4.1 shows the degradation of the analyzed pigments in pH 1-8 range. As expected, all the evaluated anthocyanins were more stable in acidic pH with degradation of ∼2%. In the neutral to alkaline pH values of 5-8, the R530 pigment showed a significant stability over PA and R520 groups. In pH 7 and 8, the absorbance of R530 decreased only ∼24%, as opposed to PA and R520 pigments with 80 and 60% loss in the absorbance spectra, respectively. The R530 pigments had also the lowest rate of degradation at pH 4 and 5 (Figure 4.1 and Table 4.3).
Addition of one acylating group to the Cy-3diG-5G increased the maximum absorbance spectra by 5 to 20 nm and resulted in a bathochromic effect. It has also been documented in the literature that the addition of acyl group to the anthocyanin molecule results in bathochromic shifts (Giusti et al. 1999; Torskangerpoll and Andersen 2005).
Diacylation significantly increased the stability of the pigments 3-4 times in the neutral to alkaline pH values where the anthcyanins are more susceptible to color degradations. It also significantly influenced the stability in the middle range of pH (4-5). Torskangepoll and
Anderson (2005) demonstrated that the diacylated cy based anthocyanins (compared to the non acylated ones) had an excellent color stabilities in pH 4.1 and 5.1during 98 days in refrigeration storage condition.
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100
i 90 B 80 h 3 g 70 g 5 2‴ 60 2″ 50 PA 6″ R520
% Degradation % 40 R530 f 30 e e 20 d dc 10 dc dc dba ba b 0 1 2 3 4 5 6 7 8 9 pH
Figure 4.1: Molecular structure of Cy-3diG-5G, and, percent degradations of non, mono, and diacylated RCA in pH 1-8 after 8 hr. PA:R1=H, R2= H; R520:; R530: R1= acyl, R2= acyl. Acyl groups were cinnamic acids (p-Coumaric, Ferulic, and Sinapic). For the pigments identities refer to Table 4.2. No significant difference was observed up to pH 3 (p > 0.05). % degradation= 100-((absorbance at λmax after 8 hr/absorbance at λmax after 15min) x 100).
4.4.3. SCX-buffer fractionation of RCA
The main reason for the application of the SCX (Strong Cation Exchange) cartridge was the ion exchange and non-polar interactions of this sorbent. We hypothesized that anthocyanins based on their pka would acquire different charges at different pH values. At mid pH value of 6 some anthocyanins are positively charged, some have no charge, while others acquire negative
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charge. Since the SCX cartridge interacts with positively charged molecules, when anthocyanins are loaded into the cartridge using pH 6 solvent with low percentage of MeOH can elute the anthocyanins that have no charge or negative charge in that pH range. Then, elute the R530 pigments eluted by increasing the pH and the percentage of MeOH. Because there are not enough data on the pka properties of RCA, several different combination of solvents were studied.
According to our observations, using the SCX cartridge in combination with only
MeOH as solvent did not result in elution of the pigments; however, the same cartridge with combination of mixture of buffer and MeOH resulted in the elution of anthocyanins. When the cartridge conditioned with pure buffer (pH6), using the same buffer containing 25% MeOH afterward was able to elute mostly more polar early eluting peaks and R520 pigments (S1-F1 and S2-F1). Buffer pH 7 could elute both R520 and R530 pigments (S1-F2 and S2-F2) (Figure
4.2). When only buffer pH 8 (with 25% MeOH) used, there were some losses in R530 pigments
(Figure 4.2). When buffer pH 8 (with 25% MeOH) was used (Figure 4.3), it resulted in elution of early eluting peaks and R530 pigments (S3-F1). This suggested that the pH of the solvent was important in the anthocyanin ionic interaction with the SCX sorbent. This could be because
R530 pigments still possessed a positive charge at pH6 and that they remained bonded to the
SCX sorbent. When the pH of the solvent increased, this could result in the R530 pigments to lose their charge or acquire a negative charge, so, they could elute from the cartridge. As shown in Figure 4.2 (right), buffer pH 8 combined with 70% MeOH (S2-F4) was the most effective solvent for the elution of R530 pigments after the more initial and R520 pigments already eluted using the combination of other solutions. Therefore, for further experimentations only pH 6 and pH 8 was used. Also, because some loss in the pigments were observed due to the conditioning
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steps using pure buffers, this step was also skipped for the next round of the experimentation
(S4 and S5).
R530 RCA R520 6 S2-F1 1 2 3 5 4 PA R530
S1-F1 S2-F2
)
AmU
540 nm nm 540 -
S1-F2 S2-F3
Max Plot Max Plot 510 Absorbance (
S2-F4 S1-F3
0 5 10 15 20 25 30 35 0 5 10 15 20 25 30 35 40 Time (min)
Figure 4.2: Chromatographic separation of different fractions collected using the SCX cartridge and buffers (methods S1 and S2). For the solvent used to isolate each fraction refer to Table 4.1. Identification of each individual peak is shown in Table 4.2. Samples were conditioned with pure buffers prior to collecting each fraction.
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R530
RCA
)
AmU 540 nm nm 540
- S3-F1 Max Plot Max Plot 510
Absorbance ( S3-F2
0 5 10 15 20 25 30 35 40 Time (min)
Figure 4.3: HPLC chromatograms of the fractionations obtained using SCX cartridge and buffer pH8 and different percentage of MeOH. For the solvent used to isolate each fraction refer to Table 4.1. Sample was conditioned with pure buffer pH8 before collecting the S3-F1 fraction.
When the conditioning step was eliminated (S4 method), there were some loss in R530 pigments in S4-F1 fraction. In order to minimize the loss the polarity of the amount of MeOH was reduced to 20% as shown in Figure 4.4 (S5). The S5 method was the most effective method in eliminating the initial pigments and concentration of R530 pigments. Although the S5 method was able to elute mainly the R530 pigment in the final fraction, according to the HPLC chromatograms (fraction S5-F1 Figure 4.4) it appeared that some R520 permanently retained on the sorbent.
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R530 R530
RCA RCA
) AmU
540 nm nm 540 S4-F1 S5-F1 -
Max Plot Max Plot 510 S4-F2 S5-F2 Absorbance (
0 5 10 15 20 25 30 35 0 5 10 15 20 25 30 35 40 Time (min)
Figure 4.4: HPLC chromatograms of method S4 and S5 using SCX cartridge when the conditioning steps using pure buffers were skipped. For the solvent used to isolate each fraction refer to Table 4.1.
Utilization of the SCX cartridge in combination with buffers pH 6 (20% MeOH) and buffer pH 8 (70% MeOH) was effective in fractionation of the R530 pigments. However, there are some disadvantages using this method such as cost of the cartridge, difficulties dissolving the buffer and MeOH, loss of the pigments, and low efficiency due to the need for extensive final clean-up step to eliminate salt using C18 cartridge.
4.4.4. C18-MeOH and EtOH fractionation of RCA
The C18 cartridge was used to fractionate the pigments based on their polarity. MeOH and EtOH in water with different percentage were used as solvents (Table 4.1). Initially, three
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fractions were collected as shown in Figure 4.5 using C1 (C18 and MeOH) and C2 (C18 and
EtOH) methods (Figure 5.5). 30% MeOH and 10% EtOH were not able to elute the pigments before R530 pigments. Increasing their amount to 40% MeOH and 20% EtOH, however, resulted in elution of R520 and some R530 pigments. There was ∼30-40% loss of R530 pigments in the second fractions (C1-MF2 and C2-MF2) (Table 4.4).
R530 R530 RCA RCA
C1-MF1 C2-EF1
)
AmU 540 nm nm 540
- C1-MF2 C2-EF2 Max Plot Max Plot 510
Absorbance( C1-MF3 C2-EF3
0 5 10 15 20 25 30 35 0 5 10 15 20 25 30 35 40 Time (min)
Figure4.5: Chromatographic separation of the three fractions collected using C18 cartridge and MeOH (left) and EtOH (right) in various percentage (method C1 and C2). For the solvent used to isolate each fraction refer to Table 4.1. Conditioning step using pure buffer before collecting each fraction was skipped.
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In order to avoid the extra step and achieve a better concentration of R530 pigments, the fractionation steps were reduced to only two as shown for method C3-C6. In the case of fractionation with MeOH (Figure 4.6), 40% MeOH (C3-MF1) was used to elute the initial more polar pigments as well as R520 pigments; however, this resulted in 35.4% loss of R530 in C3-
MF1 fraction (Table 4.4). Consequently, the MeOH (%) was reduced to 32% which resulted in the elution of initial anthocyanin pigments and the R520 group (C4-MF1). This method was shown to be the most efficient in the collection of R530 pigments (98%) in the final fraction
(C4-MF2); there was also only a 1-2% of total anthocyanins loss using this method (Table 4.4).
Two-step fractionation methods (C3 and C4) using MeOH improved the anthocyanin yield compared to the three-step method (C1).
R530 R530
RCA RCA
) AmU
540 nm nm 540 C3-MF1 C4-MF1 -
C3-MF2 C4-MF2
Max Plot Max Plot 510 Absorbance(
0 5 10 15 20 25 30 35 0 5 10 15 20 25 30 35 40 Time (min)
Figure 4.6: HPLC chromatograms of the two fractions collected using C18 cartridge and MeOH in various percentages (method C3 and C4). For the solvent used to isolate each fraction refer to Table 4.1.
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Two-step fractionation was also applied for C5 and C6 methods using EtOH as the solvent. As shown in Figure 4.7, using 17% EtOH was effective in eliminating part of R520 pigments (C5-EF1), the remaining pigments, however, eluted in the final fraction (C5-EF2).
When slightly more EtOH was used (18.5%) the initial fraction (C6-EF1) contained increased amounts of R520 pigments. Although more than 80% of R530 (Table 4.4) was recovered in the final fraction (C6-EF2), there were also some R520 left which did not elute completely in the previous fraction. Two-step fractionation with EtOH also resulted in a high (>98%) anthocyanin yield (Table 4.4).
R530 R530
RCA RCA
) AmU
540 nm nm 540 C5-EF1 C6-EF1 -
C5-EF2 C6-EF2
Max Plot Max Plot 510 Absorbance(
0 5 10 15 20 25 30 35 0 5 10 15 20 25 30 35 40 Time (min)
Figure 4.7: HPLC chromatograms of the two fractionations obtained using C18 cartridge and EtOH in various percentages (method C5 and C6). For the solvent used to isolate each fraction refer to Table 4.1.
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Table 4.4: Proportion of R530 pigment, their recovery, and yield of anthocyanins in each fraction obtained using method C1-C6 (Table 4.2). %R530 (R530 peaks proportions to the total anthocyanin peaks) obtained from the HPLC chromatograms. For the chromatogram of each fraction refer to Figure 4.5, 4.6, and 4.7.
R530 recovered Method Fraction % R530 Anthocyanin yield (%) (%)* C1-MF1 0.5±0.5 0.2±0.2d 10.1±1.1 c C1 C1-MF2 21.5±0.4 43.0±5.2 55.1±5.0 b C1-MF3 90.2±1.4 56.8±5.2 17.4±1.8 C2-EF1 0.6±0.1 0.2±0.0d 10.6±0.2 c C2 C2-EF2 24.9±1.4 34.6±3.8 54.9±3.9 C2-EF3 81.0±0.7 65.2±3.8b 32.2±4.4 C3-MF1 15.3±0.4 35.4±1.9c 58.3±2.0 C3 C3-MF2 76.6±3.4 64.6±2.8b 40.5±2.1 C4-MF1 0.9±0.8 1.3±1.3d 44.2±12.7 C4 C4-MF2 80.2±1.2 98.7±1.3a 54.8±9.1 C5-EF1 0.3±0.0 0.2±0.1d 24.8±1.3 C5 C5-EF2 46.0±0.4 97.9±0.1a 65.1±1.5 C6-EF1 4.7±1.5 7.1±3.1d 50.1±9.8 C6 C6-EF2 61.4±2.1 92.9±3.1a 48.9±5.9 * Different letters indicate significant difference (p < 0.05).
Over all, the final fractions in method C4, C5, and C6 had significantly high amount of
R530 pigments. However, the C5 method resulted in the presence of R520 pigments in the final
fraction, and, method C6 had some loss of R530 pigments in the first fraction (C6-EF1).
4.4.5. Color analysis after fractionation
The colors of the fractions from the most effective method (C4) as well as RCA in two
different pH values (2 and 8) were analyzed and compared to Red 3 and Blue 2 synthetic dyes
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(Table 4.5). The absorbance spectra of the same samples in pH 8 also compared to Blue 2
(Figure 4.8). At pH 2, the first fraction (C4-MF1) collected using 32% MeOH which contained
mostly mono acylated (R520) pigments had the most color similarity to Red 3 (∆E= 11.6, hue
angle=1.17, and λmax= 525). At pH 8, however, the last eluting fraction containing mostly R530
pigments (C4-MF2) had the closest color properties to Blue 2 colorant (∆E= 6.6, hue
angle=235, and λmax= 620). These results were similar to the color analysis of the purified peaks
(section 4.4.1) in which the pure R530 pigments has similar color characteristics to the synthetic
Blue 2.
Table 4.5: Color and spectral characteristics of fractions collected using method C4 compared to the RCA in pH2 and 8. pH Sample L* C* h◦ ∆E† λmax RCA 72.6±0.5 52.0±0.9 353.8±0.2 20.1±0.7 530 2 C4-MF1 70.6±0.9 56.8±1.5 1.17±0.8 11.6±1.7 525 C4-MF2 72.1±4.9 48.2±8.3 347.4±2.6 28.2±4.2 535
RCA 75.1±3.2 25.6±3.3 251.5±2.4 14.9±1.6 614 8 C4-MF1 70.3±2.9 29.2±2.8 257.9±1.5 15.7±0.8 599 C4-MF2 69.2±1.9 32.7±1.3 235.5±1.5 6.9±0.8 620
Red 3 70.1 66.1 7.6 527 Blue 2 74.1 36.9 235.8 610 † pH 2 color difference with FD&C Red No. 3; pH 8 color difference with FD&C Blue No. 2.
As shown in Figure 4.8, the absorbance spectra of this fraction was sharper and more
similar to Blue 2 spectra as compared to RCA and C4-MF1 that had wider spectra at the same
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pH. A narrower spectral peak is normally associated with purer color tone and better color quality.
0.9
0.8
0.7 Blue 2 C4-MF2 0.6 C4-MF1 0.5 RCA
0.4 Absorbance 0.3
0.2
0.1
0 400 450 500 550 600 650 700 nm
Figure 4.8: Absorbance spectra of the C4-MF1, C4-MF2, and RCA at pH 8 compared to FD&C Blue No. 2 synthetic dye.
4.5. CONCLUSIONS
SPE fractionation techniques using strong cation exchange (SCX) or non-polar interaction (C18) resins were effective for the segregation of di-acylated RCA. Using C18 resin the most effective fractionation method used elution using 32 and 100% MeOH. The first
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fraction eluted using 32% MeOH, contained mainly R520 pigments, and had color properties more similar to FD&C Red No. 3 under acidic conditions. Whereas, the color of the fraction eluted using 100% MeOH, contained mostly di-acylated pigments, better matched the colors of
FD&C Blue No. 2 at alkaline pH values. C18 SPE fractionation of the RCA showed to be quite effective, convenient, and practical for isolating the target diacylated pigments with high stabilities, more pure tone, and better blue coloring properties. For practical use, however, the developed method would need to be optimized for scaled-up.
4.6. ACKNOWLEDGMENT
We are thankful to MARS Chocolate, NJ for providing funding for the project.
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CHAPTER 5: MOLAR ABSORPTIVITY (ε) AND SPECTRAL CHARACTERISTICS OF CYANIDIN-BASED ANTHOCYANINS FROM RED CABBAGE1,2
5.1. ABSTRACT
Red cabbage extract contains mono and di-acylated cyanidin (Cy) anthocyanins and is often used as food colorants. Our objectives were to determine the molar absorptivity (ε) of different red cabbage Cy-derivatives and to evaluate their spectral behaviors in acidified MeOH and buffers pH1-9. Major RCA were isolated using a semi-preparatory HPLC, dried and weighed. Pigments were dissolved in MeOH and diluted with either MeOH (0.1% HCl) or buffers to obtain final concentrations between 5x10-5- 1x10-3 mol/L. Spectra were recorded and
ε calculated using Lambert-Beer’s law. The ε in acidified MeOH and buffer pH1 ranged between 16,000-30,000 and 13,000-26,000 L/mol.cm, respectively. Most pigments showed higher ε in pH8 than pH2, and lowest ε between pH4-6. There were bathochromic shifts (81-
105nm) from pH1-8 and hypsochromic shifts from pH8-9 (2-19nm). Anthocyanins molecular structures and the media were important variables which greatly influenced their ε and spectral behaviors.
1 Neda Ahmadiania, Rebecca J. Robbinsb, Thomas M. Collinsb, M. Monica Giustia* a Department of Food Science and Technology, The Ohio State University, 2015 Fyffe Road, Columbus, OH 43210, USA,b Analytical and Applied Sciences Group, Mars Inc., 800 High Street, Hackettstown, New Jersey 07840 2 Submitted in Food Chemistry
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Keywords: Red Cabbage, Cyanidin derivatives, Molar absorptivity (ε), Spectral characteristics
5.2. INTRODUCTION
Anthocyanins are plant pigments with numerous potential preventative and therapeutic health benefits (Fernanda Nunez 2013; He and Giusti 2010). They are known to have antioxidant activities and linked with protection against cardiovascular disease, diabetes, cancers, vision problems, neurodegenerative diseases, aging skin, inflammatory diseases, and other medical conditions (He and Giusti 2010).
Additionally, these water soluble compounds have potential application in coloring of different food products due to their capabilities in creating a wide range of color (Giusti et al.
2008; Hendry 1992). Colorants made of these pigments are currently manufactured for food use from horticultural crops and processing wastes (Wrolstad and Smith 2010). Fruit and vegetable juice containing anthocyanins such as concentrated red cabbage, black carrot, purple sweet potato, radish, bilberry, elderberry are being used as approved food color additives in most countries (Socaciu 2007; Hendry 1992). Due to the growing application of these pigments as food colorants and because their broad color variations depends upon the pH of the product, measurements of anthocyanin contents and their color analysis such as visible spectral characteristics in various solvents and pH values is essential for the food industry and researchers working with this group of pigments.
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True content of anthocyanin pigment in a solvent and pH of choice can be obtained based on the molar absorptivity (ɛ) values of anthocyanins. Molar absorptivity of anthocyanin is a physical property which measures how strongly this pigment absorbs light at a given wavelength when placed in a specific solvent with a known pH value. Molar absorptivity of some anthocyanins in specific solvents have been reported in the literature; however, the data found in the literature are from decades ago and have many discrepancies and inaccuracies due to the limitation of preparing crystalline anthocyanins in a pure form. Along with ε data which provides information about the tinctorial strength of the color, having information about the visible spectra of anthocyanins also provides information about the color characteristics at certain pH values. At a given pH, the wavelength at which anthocyanins have maximum absorbance (λmax) and the total UV-Vis spectral characteristics are a result of the electronic structure of the molecule that will in turn determine the color characteristics.
Red cabbage anthocyanin extract is often used by the food industry as an alternative for synthetic food colorants because of two main reasons: their good stabilities to light and processing temperature and being able to exhibit a wide spectrum of color, ranging from orange through red, to purple and blue based upon the pH of the environment (Walkowiak-Tomczak and Czapski 2007; Dyrby et al. 2001). Red cabbage is a rich source of cyanidin (Cy) based anthocyanins with various mono and di-acylating groups. The objective of this study were to obtain the ε and λmax of major Cy based anthocyanins from red cabbage in MeOH (0.1%HCl) and buffer range pH 1 through 9 and to evaluate the effect of number of acylation on spectral stability of two selected anthocyanin pigments.
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5.3. MATERIALS AND METHODS
5.3.1. Plant materials and anthocyanins extraction
Red Cabbage plants were donated by Bejo Seeds Inc. (New York, USA), variety Cairo.
Samples were refrigerated after they were received and analyzed within a week.
Cabbage heads (≃30 g) were sliced and frozen with liquid nitrogen and kept frozen until analyzed the following day. The frozen plants were ground using a stainless steel blender.
The extraction procedure was adopted from Giusti and Wrolstad (1996). Frozen plant powder was mixed with acetone (30 mL). The mixture was filtered through a Whatman #1 filter
(Whatman Inc., Florham, NJ),and washed with 70% aqueous acidified acetone (0.1% formic acid) (≃ 250 mL) until the powder was white. The filtrates were combined, transferred to a separatory funnel and mixed with 1 volume of chloroform. The phases were allowed to separate for 4-5 hr. The aqueous phase was then collected and the residual acetone was evaporated using a Büchi Rotavapor (Brinkmann Instruments, Inc, Westbury, NY).
5.3.2. Anthocyanins semi-purification
To separate the anthocyanins from sugar and acids, the aqueous extract was semi- purified using a Sep-Pak C18 cartridge (Waters Corp., Milford, MA). The cartridge was activated with MeOH, washed with acidified water (0.01% HCl) before loading the sample.
Prior to semi-preparatory HPLC to improve and ease the pigment isolation, the pigments were concentrated into three different fractions based on their polarity. The fractions recovered contained mainly the early (peak 1 and 2), mid (peak 3-5), and late (peak 6-8) eluting
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pigments (Figure 5.1), named fractions 1, 2, and 3. Solvent conditions used to recover fractions
1, 2 and 3 were 30%, 40%, and 100% acidified MeOH (0.01%) in water respectively. The
MeOH was removed using the rotavapor; and, the concentrated anthocyanins were collected in acidified water (0.01% HCl).
5.3.3. Chromatographic analysis
5.3.3.1. Semi-preparatory HPLC
A Shimadzu Prominence® semi-preparative High Pressure Liquid Chromatograph
(HPLC) coupled to a SPD-M20-A photo-diode array was used to isolate the semi-purified pigments. The column was a 250 x 21.20 mm Luna PFP 5 μm (Phenomenex Inc, CA, USA).
The flow rate was 12 ml/min and the injection volume was 2000 l. The solvents were phase A,
4.5% formic acid in LCMS grade water; B, LCMS acetonitrile (Fisher Scientific Inc, Fair lawn,
NJ).For fraction 1, gradient: 0-15 min, 13% B was used. For fraction 2, gradient: 0-11 min, 18%
B, and 11-35 min, 18- 30% B were used. For fraction 3, gradient: 0-25 min, 20% B was used.
Pigment isolation was done several times, but only the extract with the highest purity level was used for further analysis. LCMS Solution software was used (Shimadzu Scientific, Inc.,
Columbia, MD) for data analysis.
5.3.3.2. Analytical HPLC
A Shimadzu Prominence® reverse phase analytical HPLC and Mass Spectrophotometer
(MS) coupled with a single-quadrupole electrospray ionization (ESI) (Shimadzu Scientific, Inc.,
Columbia, MD) were used to check for the purity of the isolated pigments. The detector and the
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software for data analysis were similar to the those described in section 2.2.1. The column used was a 100 x 4.5 mm Kinetex PFP 2.6 μm (Phenomenex Inc, CA, USA). The solvents were the same as section 5.3.3.1.; and, the gradients were 0-50 min, 0-30% B. Injection volume was
20µl. Spectral data was obtained from 250 to 700 nm and elution of anthocyanins was monitored at 520 nm. Purity of the isolated pigments was checked by dividing the % Area
Under the Curve (AUC) at 510-530 nm to the max plot (250-700 nm), and, the peaks with purity above 90% were used for further analysis.
Mass Spectroscopy (MS) was used to confirm the identities of the pigments. The analysis was done by diverting a 0.2 mL/min volume of solvents into the MS and ionized under positive ion condition using an electrospray probe. Data was monitored using total ion scan
(SCAN) (from m/z 200-1200) and selected ion monitoring at m/z 287 since Cy was the major aglycon anthocyanin in red cabbage.
5.3.4. Measurement of ε and Spectral Characteristics
For both of the analyses described below, purified dried pigments were re-dissolved in
10 mL of 100% HPLC-grade MeOH to make the stock solution. For ε calculations, the
Lambert-Beer’s law and the molecular weight used included the weight of a chloride counter ion and a water molecule of hydration (Giusti et al. 1999).
5.3.4.1. Acidified MeOH and Buffer pH1
To measure the ε and spectral characteristics in acidified MeOH and buffer, an exact aliquot of the stock solution was diluted in MeOH containing 0.1% HCl and buffer pH 1 (0.25M
KCl), by triplicates, to obtain final concentrations of 5x 10-5- 1x 10-3mol/L. 30-40min was
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allowed for equilibrium. Spectra of known dilutions were recorded on a Shimadzu2450 UV spectrophotometer (Shimadzu Scientific, Inc., Columbia, MD) using 1-cm path length quartz cuvette. Spectral data of the p-coumaric, ferulic, and sinapic acids (Sigma-Aldrich, St. Louis,
MO) were also obtained using the same spectrophotometer by dissolving a few mg of the acids in the 0.1% MeOH, so that the UV absorbance would be below 1.0.
5.3.4.2. Buffer Solutions pH 1-9
To measure the ε and spectral characteristics in buffer pH 1-9 (X5), an exact aliquot of the stock solution was pipetted into each well of a 96 well-plate (Greiner Bio-One,
Frickenhausen, Germany). The plates were allowed to dry overnight in dark at room temperature. Buffer solutions were added to the dry pigments in each well to obtain the final concentration of 7.5x 10-5- 1x 10-4 mol/L. The buffers were KCl (0.25M), citrate (0.1M), phosphate (0.1M), and bicarbonate-carbonate (0.1M), for pH 1, 2-5, 6-8, and 9, respectively
(Appendix I). The dissolved pigments were allowed to equilibrate for 1hr in the dark. A
SpectraMax® 190 UV-Vis Microplate Spectrophotometer (Molecular Devices, Sunnyvale, CA) was used. The spectra were collected in 1nm increments between 400-700nm wavelengths. The
ε values for the buffer pH 1 calculated above was used to obtain the accurate path length used by Beer’s law.
5.3.5. Stability comparison between mono and di-acylated Cy- derivatives
To evaluate the effect of number of acylations on the color stabilities, spectra of the isolated pK5 and PK8 were compared to each other. Isolated purified peaks (1.0 x10-4 mol/L)
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were dissolved in the same buffers described in section 2.6.1 pH 1-8 (5 replications of each).
The spectral measurements were also measured the same way described in section 2.6.1 using the UV-Vis Microplate Spectrophotometer, and, the solutions were stored at room temperature
(25°C). The color degradation (% degradation) of the two pigments was calculated by:
% Degradation = 100-((absorbance at λmax after 8hr/ absorbance at λmax after1hr) x100)
5.3.6. Statistical analysis
Two sample t –test was used for statistical analysis for stability comparison of the mono and di-acylated pigments. Microsoft Excel 2010 was used to run the analysis at a 0.05 p-value.
5.4. RESULTS AND DISCUSSIONS
5.4.1. Major Cy based anthocyanins from red cabbage
There were 8 major peaks in red cabbage extract (Figure5.1), labeled as PK1-8.
Ahmadiani et.al. (2014) identified the anthocyanin extracts from 8 different red cabbage varieties including Cairo variety as non-acylated, mono-acylated and di-acylated derivatives of cyaniding-3-diglucoside-5-glucoside in agreement with previously published literature (Wu and
Prior 2005b; Wiczkowski et al. 2013; Park et al. 2014; Scalzo et al. 2008; Sun et al. 2013;
Ahmadiani et al. 2014). The identities of the anthocyanin pigments are shown in Figure 5.1. As shown in this table, the non-acylated pigment eluted first (PK1), followed by mono-acylated
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anthocyanins PK (2-5) and lastly the di-acylated pigments (PK6-8). The major acylating groups were p-coumaric, ferulic, and sinapic acids. The order of elution for the acylating pigments with the same anthocyanin backbone usually depends on the polarity of the attached acylating groups. As shown in Figure 5.1, mono-acylated PK 3,4, and 5 and di-acylated PK 6,7, and 8, followed this law since the order of polarity for the acylating groups is p- coumaric>ferulic>sinapic acid (from the most to the least polar). For PK2, however, having the same mass over charge ratio and with the same acylating group as PK5 (Cy3diG-5-G acylated with one sinapic), the retention time was ∼12min shorter than PK5. This difference in retention time for a similar chemical composition suggests that the difference between PK2 and PK5 is in the position of the acylating group.
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520 nm 1 5 ) 8
AmU 2 4 3 7
6 Absorbance ( Absorbance
0 5 10 15 20 25 30 35 40 45 50 Time (min)
RT Peak M+ Identification (min) 1 12.13 773, (287) Cy3diG-5-G 2 16.95 979, (287) Cy3diG-5-G+Sinapic* 3 27.41 919, (287) Cy3diG-5-G+p-Coumaric 4 28.33 949, (287) Cy3diG5-G+Ferulic 5 28.85 979, (287) Cy3diG5G+Sinapic 6 30.53 1125, (287) Cy3diG-5-G+Sinapic&p-Coumaric 7 31.55 1155, (287) Cy3diG-5-G+Sinapic&Ferulic 8 32.31 1185, (287) Cy3diG-5-G+Sinapic&Sinapic * Tentative identification
Figure 5.1: HPLC chromatogram and identity of the major RCA. The major fragment in all 8 peaks was 287 m/z corresponding to Cy3diG-5-G. RT: Retention time; M+: Mass ion (Ahmadiani et al. 2014)
5.4.2.Molar absorptivity and spectral characteristics in MeOH and buffer pH1
Molar absorptivitiy of the pigments in the acidified MeOH and buffer pH 1 ranged between 16,000 to 30,000 and 13,000 to 26,000 L/mol.cm, respectively (Table 5.1). The highest value belonged to the non-acylated Cy3diG-5-G (PK1) and the lowest was obtained for the Cy
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pigment acylated with one sinapic acid (PK2) in both acidified MeOH and buffer pH1. PK5, despite having the same number and type of acylation as PK2, had the second highest ε in both solutions. Hrazdina et.al. (1977) also reported the ε of some Cy anthocyanins from red cabbage obtained in 1% aqueous HCl using column chromatography techniques on PVC and their values were between 32,000- 38,000 L/mol.cm. Our value for Cy3diG-5-G was slightly closer to the ones calculated based on their data (37,150 L/mol.cm); for the other anthocyanin pigments, however, there were large differences between our values and those reported by the same authors (Hrazdina and others 1977). Chromatographic instrumentations and columns have progressed immensely during the past 30 years, and the differences between the results could be explained by using different and more advanced separation techniques since the presence of other anthocyanins and impurities could result in the inter and intra co-pigmentation which could potentially cause hyperchromic effects (Mazza and Brouillard 1987; Goto and Kondo
1991; Asen et al. 1972) leading to higher ε values. Other possible cause of difference could be the use of less acidic MeOH solution since the acidity of the media greatly influences the color behaviors of anthocyanins (Brouillard 1983; Brouillard and Dubois 1977). Because the concentrations of the pigments were the same in both acidified solutions, we were also able to evaluate the solvent effect. Generally, the solvent in which the anthocyanins are dissolved is important because it has an impact on their quaternary structures, which affects the color of any of the primary, secondary, or tertiary structures of anthocyanins (Bridle and Timberlake 1997).
Anthocyanins had higher λmax and ε in MeOH compared to the aqueous solvent (Table 5.1) resulting in a more violet red to pink color with higher color intensities.
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Table 5.1: ε (L cm-1 mol-1) of the Cy derivatives from red cabbage 1hr after dissolved in acidified MeOH and buffer pH 1(25°C). For peak identity and retention times refer to Figure 5.1.
Weight λmax ε L/(mol.cm) Stock Final Con. Peak# MW Conc. g/L (mol/L) (mg) MeOH* Buffer pH1 MeOH* Buffer pH1
1 22.03 0.88 1.07E-03 826.7 525 512 30247 25541 2 6.42 0.64 4.78E-04 1031.5 528 525 16466 13714 3 5.42 0.54 5.58E-04 972.7 527 523 23147 20259 4 5.40 0.54 5.4E-05 1002.7 527 523 21501 18203 5 4.93 0.49 5.39E-05 1031.5 526 518 26775 21793 6 6.41 0.64 5.45E-05 1178.8 532 531 21293 16869 7 5.00 0.50 4.97E-05 1208.8 528 525 18788 15123 8 7.07 0.71 5.72E-05 1238.8 526 522 15556 12733
*Acidified with 0.1% HCl
Chemical structures also a played role in the spectral variations observed in the
evaluated solvents. The amount of bathochromic shift in the acidified MeOH compared to the
acidified buffer was higher in the absence of acylating group(s). As shown in Table 5.1, this
shift was 13nm for the non-acylated Cy3diG-5-G (PK1). Giusti et.al. (1999), also showed that
non-acylated pigments had the highest solvent effect on spectral characteristics of Pg based
anthocyanins.
In order to compare the spectral shapes of the pigments, the normalized spectra of the
isolated pigments in acidified MeOH are shown in Figure 5.2a and compared to the spectra of
the pure cinnamic acid acylation in the same solution. Both non-acylated and acylated pigments
showed absorbance at 270- 310nm. Absorbance was higher for the non-acylated pigment (PK1)
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in 360- 460 nm. The presence of acylating groups decreases the absorption in the 400-440 nm region for Pg based anthocyanins (Giusti et al. 1999). Acylation resulted in an increase in the absorptivity in 310- 325nm region and, the absorbance maxima in this region corresponded to the absorbance maxima obtained from the pure cinnamic acids (Figure 5.2c). For PK2 and 5 with the same mass and the same building blocks, the spectra were very similar in 285-610nm region (Figure 5.2b); however, PK2 had higher absorptivity in the UV region than PK5. In acidified solutions, anthocyanins normally show a peak at 270 nm (Wrolstad et al. 2001). In the case of PK2, a broad peak was observed in the 260 to 280nm region (Figure 5.2b).
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3
2.5 PK1 (a) 2 PK3 1.5 PK4 1 PK6
0.5 PK7 Absorbance Absorbance 0
-0.5
-1
-1.5 2.5 260 310 360 410 460 510 560 610
2 (b) 1.5 PK2
1 PK5 0.5 PK8 Absorbance Absorbance 0
-0.5
-1
-1.5 260 310 360 410 460 510 560 610 3.5 3 2.5 (c) p-Coumaric 2 Ferulic 1.5 Sinapic 1
0.5 Absorbance 0 -0.5 -1 -1.5 260 310 360 410 460 510 560 610 Wavelength (nm)
Figure 5.2: Normalized UV-vis spectra of a: Cy3diG-5-G (PK1), Cy3diG-5-G+p-Coumaric (PK3), Cy3diG5-G+Ferulic (PK4), Cy3diG-5-G+Ferulic&Ferulic (PK6), and Cy3diG-5- G+Sinapic&Ferulic (PK7); b: Cy3diG-5-G+Sinapic (PK2 and PK5) and Cy3diG-5- G+Sinapic&Sinapic (PK8); c: pure p-Coumaric, Ferulic, and Sinapic acids in MeOH (0.1% HCl).
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5.4.3. Molar absorptivity, spectral characteristics, and color in buffers pH 1-9
The molar absorptivity (ε) of all pigments decreased from pH 1 to pH 6. Pk 1, 3, 4, and
5 had very low absorbencies and faded color at pH 5 and 6. This is consistent with the predominance of the flavylium form at low pH, with formation of the colorless forms as the pH increases. However, diacylated pigments, as well as PK2 did not completely fade in color, with some color remaining in the solution. The molar absorptivity started to increase when the pH approach neutrality, when anthocyanins are expected to go back to their quinonoidal base configuration (Brouillard and Dubois 1977); however, this value started to decrease again at pH
9 (except for PK6) (Table 5.2). For some pigments such as PK7 and PK8 resulted in a noticeably higher ε at pH 8 than pH 1.
Table 5.2: Molar absorptivity (ε) (L cm-1 mol-1) and maximum absorbance (λmax) of the of the Cy derivatives from red cabbage 1hr after dissolved in pH 1-9 (25°C). Final concentrations were 1.02 x10-04mol/L and 7.50 x 10-05mol/L for peak 1-2 and 3-8, respectively. For peak identity and retention times refer to Figure 5.1.
pH Peak 1 2 3 4 5 6 7 8 9 ε 25541 17093 4380 1057 NA NA 3140 22137 8370 1 λmax 511 512 513 516 NA NA 578 594 592 ε 13714 12081 9112 5123 3318 3444 9651 13347 11829 2 λmax 528 528 529 530 539 556 643 633 614 ε 20259 16629 6031 1847 NA NA 4894 17363 14313 3 λmax 521 521 522 524 NA NA 584 601 599 ε 18203 13393 6086 1588 NA NA 6326 16359 13595 4 λmax 522 522 523 523 NA NA 588 603 602 To be continued
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Table 5.2 continued
pH Peak 1 2 3 4 5 6 7 8 9 ε 21793 17801 10804 3965 NA NA 10674 22402 21716 5 λmax 517 517 516 519 NA NA 586 605 603 ε 16869 17342 14322 9340 5845 4718 9887 16573 17851 6 λmax 530 532 534 537 546 554 626 626 617 ε 15123 18003 11282 7577 5788 3836 11078 21187 19411 7 λmax 532 533 537 541 543 548 603 616 609 ε 12733 13459 10795 7075 4921 3570 6303 14849 14071 8 λmax 533 534 536 540 547 551 604 616 608 *NA: Not acquired due to low absorbance.
The actual colors of the pigments at pH 1-9 are shown in Figure 5.3. PK2, 6, 7, and 8
had very similar color behaviors throughout the studied pH range. All of these pigments were
pink at pH 1-4, purple at pH 5- 6, and exhibited a bluer color at pH 7-8. PK3, 4, and 5, on the
other hand, were red at pH 1-3, nearly colorless at pH 4-6, purple at pH 7, and produced bluer
hues at pH 8-9. Finally, PK1 was orange red color at pH 1-3; it then become nearly colorless at
pH4-6, purple color at pH 7, and, it turned into blue color at pH 8-9 (the color change is shown
in the supplementary materials). Figure 5.4, also, compares the spectral characteristics of PK1,
2, 5, and 8 in four different pH values. The spectra were broader at pH 5 and 7, with the lowest
absorbencies at pH 5 for the pigments. The maximum absorbance wavelength shifted toward
the higher wavelength from pH 2 to 9, except for PK2 that the λmax was higher at pH 7 than pH
9. Also, at pH 9 the absorbencies increased at 420nm closer to the UV region.
Table 5.2 also contains λmax of the isolated pigments in buffers pH 1-9. As expected,
there was a bathochromic shift for all the pigments by increasing the pH from 1 to 8, followed
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by a hypsochromic shift from pH 8 to 9 (Giusti et al. 2008). PK1 had the lowest λmax of 511 in pH 1. Generally, PK2, 6, 7, and 8 had higher λmax in the examined pH values compared to PK1,
3, 4, and 5. PK2 compared to the other pigments had the largest λmax of 643 nm and 633nm in pH 7 and 8, respectively. PK6 also had a large maximum absorbance value of 626 nm at both pH 7 and 8. Red cabbage anthocyanin extracts with higher percentage of di-acylated Cy anthocyanins are reported to have larger λmax at a neutral pH of 7 than the ones with larger non and mono-acylated Cy pigment contents (Ahmadiani et al. 2014).
Figure 5.3: Color changes of the major Cy based pigments from red cabbage 1hr after dissolved (25°C) in buffer pH 1-9.
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1.2 0.7
1 a: PK1 pH 2 0.6 b: PK2
pH 5 0.5 0.8 pH 7 0.4 0.6 pH 9 0.3 0.4
Absorbance 0.2
0.2 0.1
0 0 400 450 500 550 600 650 700 400 450 500 550 600 650 700 0.7 Wave length (nm) 0.12 Wave length (nm)
0.6 0.1 c: PK5 d: PK8 0.5 0.08 0.4 0.06 0.3 0.04
0.2 Absorbance 0.1 0.02
0 0 400 450 500 550 600 650 700 400 450 500 550 600 650 700
Wave length (nm) Wave length (nm)
Figure 5.4: Visible spectra of a: Cy3diG-5-G (PK1); b: Cy3diG-5-G+Sinapic (PK2); c: Cy3diG-5-G+Sinapic (PK5); and d: Cy3diG-5-G+Sinapic&Sinapic (PK8) in buffers pH 2, pH 5, pH 7, and pH 9.
5.4.4. Stability comparison between mono and di-acylated Cy- derivatives
Different studies have reported a higher stability of di-acylated anthocyanins as compared to the mono-acylated ones (Giusti and Wrolstad 2003). Having the purified materials we decided to compare the stability of two similar compounds, differing only on the presence or absence of a second acylating group. For this experiment we chose PK5 and PK8. In order to find out how the acylation would influence the spectral stability, the absorbances at λmax for the two pigments were compared to each other after 8 hr storage in room temperature. As shown in
Figure 5.5, degradation was minimal when both pigments were placed in acidified condition
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(pH 1-2); it then started to increase at pH 6, 7, and 8. When the degradation was >10%, at pH 6, there was no significant difference between the amount of the degradation for the two evaluated pigments. The differences, however, were more pronounced in pH 7 and 8 with the mono- acylated pigment being significantly more unstable than the di-acylated one.
Cy3diG5G+Sinapic (PK5)
80 Cy3diG-5-G+Sinapic&Sinapic (PK8)
* 70 5
60 4 * * 50 3 * 40 2 *
% Degradation % 1 30 0 20 1 2 3 4 5
10 * * * 0 1 2 3 4 5 6 7 8 pH
Figure 5.5: Spectral degradation comparison of PK5 (6.33x10-04 mol/L) and PK8 (4.86x10-4 mol/L) in pH 1-8 (25°C) stored in dark for 8hr (results were significant at p-value <0.05).
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5.5. CONCLUSIONS
Molar absorptivities of the red cabbage Cy-based anthocyanins varied with the matrix and the molecular structure of the anthocyanins, and ranged from 30247 for PK1 in acidified
MeOH (0.1% HCl), to 1057 for the same anthocyanin at buffer pH 4. In general, higher ε was obtained with the non-acylated Cy3diG-5-G, and decreased with increased acylation. The pigments generally had higher ε and higher λmax in MeOH than the buffer solution at pH1. The
λmax and the ε of the pigments also changed significantly with pH. For all the evaluated pigments, ε was high in pH 1-2, decreased between pH 3-6, and increased from pH 7-8. For the majorities of the pigments, the ε was higher in pH 8 than pH 2, and, there were 81-105 nm bathochromic shifts in the λmax from pH 2 to pH 8. Generally, pigments with higher molar absorptivity will show higher tinctorial strength when used as colorants.
Another interesting finding was the difference in the ε and spectral behavior of PK2 and
PK5 with the same building blocks (Cy3diG-5-G+Sinapic). PK2 behaved differently compared to the other mono-acylated pigments, having a low molar absorptivity at acidic and alkaline pH values, and a different shade of blue in neutral to alkaline pH 7-9. It, however, behaved more similar to the di-acylated pigments in regards to spectral and color behavior. Therefore, further investigation is required about the spatial configuration of this pigment.
5.6. ACKNOWLEDGMENT
We are thankful to MARS Chocolate, NJ for providing funding for the project.
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OVERALL CONCLUSION
The anthocyanin profiles, which are influenced by the plant cultivars and maturations, affected color and stability of red cabbage extracts at different pH values. Lower proportions of di-acylated pigments better reproduced the color of FD&C Red No. 3 under acidic conditions, whereas, higher proportions of di-acylated resulted in colors similar to FD&C Blue No. 2 at neutral pH.
The developed SPE methods were effective in isolating the target pigments; however, the most effective method with high efficiency was the SPE fractionation technique using a C18 cartridge with 32 and 100% MeOH. This resulted in obtaining an anthocyanin extract from red cabbage with higher stability, more pure tone, and better blue coloring properties.
Our findings showed higher ε for non-acylated pigments, and decreasing ε with increased acylation. Medium and pH were important in the color strength and behavior of anthocyanins. The molar absorptivity and λmax were generally higher in acidified MeOH than in buffer pH 1; and, high ε were found at pH 1-2, decreased at pH 3-6, and increasing again from pH 7-8. Overall, anthocyanin structures were critical factors that affected the ε and spectra.
Overall, RCA showed potential as suitable alternatives to red and blue synthetic dyes, depending on the application (pH) and the acylations present and can be fractionated to produce better red and blue hues similar to commercial colorants.
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APPENDIX A: BUFFERS PREPARATION
Solvents (mL) used to create 100ml of the buffers pH1 (0.25M) and pH2-9 (0.1M).
Trisodium Sodium Sodium Potassium Citric citrate, phosphate, phosphate, Sodium Sodium pH Chloride acid dehydrate dibasic monobasic, bicarbonate carbonate
monohydrate 1* 100.0 2* 82.0 18.0 3 82.0 18.0 4 59.0 41.0 5 35.0 65.0 6 6.2 43.9 7 30.5 19.5 8 47.4 2.7 9* 80.0 20.0 * pH adjustment was done using HCl (6N)
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