1 Altering Ph, Temperature and Cofactors to Increase the Formation of the More Stable Anthocyanin Derived Pyranoanthocyanin Thes

Altering pH, temperature and cofactors to increase the formation of the more stable anthocyanin derived pyranoanthocyanin

Thesis

Presented in Partial Fulfillment of the Requirements for the Degree Master of Science in

the Graduate School of The Ohio State University

Megan E. Hoehn

Graduate Program in Food Science and Technology

The Ohio State University

2019

Thesis Committee

Dr. M. Mónica Giusti, Advisor

Dr. Luis Rodriguez-Saona

Dr. Christopher Simons

Copyrighted by

Megan E. Hoehn

2019

Abstract

Anthocyanins are responsible for the blue, red and purple hues of many fruits and vegetables. Consumer concern over synthetic colorants has grown and anthocyanins have been looked to as natural alternatives. Anthocyanins are subject to degradation by pH, temperature and bleaching restricting their use as food colorants. Pyranoanthocyanins are a class of compounds derived from the reaction of anthocyanins and a cofactor that possess an additional ring enhancing the stability of the compound. Pyranoanthocyanins increased stability makes them appealing for food use. However, pyranoanthocyanins form at low yields limiting their potential as food colorants. The objective of this study was to determine the environmental conditions, pH, temperature and cofactor, that favor the formation of pyranoanthocyanins. Cyanidin-3-glucoside and cyanidin-3-rutinoside were obtained from blackberry and mixed with pyruvic acid in buffers ranging from pH 1 to 5. At pH 3.0 samples were stored at 5°C, 15°C and 25°C. To determine the effect of cofactor, anthocyanins were mixed with caffeic acid or pyruvic acid at different anthocyanin:cofactor molar ratios ranging from 1:1 to 1:200 in pH 3.0 solution and stored at 25°C. HPLC-PDA and UV-Visible spectrophotometry were used to monitor anthocyanin and pyranoanthocyanin contents and spectral characteristics.

Pyranoanthocyanins formed most rapidly near the pKH of the anthocyanin molecule and as time went on an optimum range of formation between pH 2.8 and 3.5

ii was observed with pyranoanthocyanin formation yields at ~24%. It was expected that the acidic environment would allow for the flavylium cation to predominate but should be low enough to prevent pyruvic acid polymerization. At 25˚C pyranoanthocyanins were forming faster and at higher yields (~25% in 21 days). As the temperature decreased pyranoanthocyanin formation slowed and formation yields were lower (~23.5% in 28 days at 15°C). However, as temperature increased anthocyanin degradation increased.

Increased temperature added more energy to the system likely encouraging pyranoanthocyanin formation. Caffeic acid (pyranoanthocyanin formation yield ~39%) was more effective at forming pyranoanthocyanins than pyruvic acid (pyranoanthocyanin formation yield ~8%) as indicated by higher pyranoanthocyanin formation. The caffeic acid samples had higher absorbance values at the end of the study indicating better pigment retention. The co-pigmentation effect of caffeic acid and the additional stability caffeic acid provided to the carbenium ion intermediate likely encouraged pyranoanthocyanin formation with caffeic acid. Temperature, pH and cofactor were important parameters for pyranoanthocyanin formation and controlling them may help increase pyranoanthocyanin yields for potential industrial use.

iii

Acknowledgments

I would like to acknowledge and thank my advisor Dr. M. Mónica Giusti for her continued guidance and support. Without her constant enthusiasm I would not be where I am at today. When I joined Mónica’s lab as an undergraduate I knew instantly she was the type of professor I wanted to work under in graduate school. She is kind, caring and passionate about teaching and research. Her passion inspired me to find a project that I was passionate about and took pride in, and her encouragement helped motivate me to complete it. She is a renowned researcher who is respected in the scientific community, and it has been an honor working in her lab.

My committee members Dr. Luis Rodriguez-Saona and Dr. Christopher Simons are both inspiring researchers and teachers. I have taken classes taught by Dr. Rodriguez and Dr. Simons. Both individuals are dedicated to their student’s success. They help solve problems and teach students how to critically think to solve their own problems.

Their unwavering support has allowed for my success as both an undergraduate and graduate student.

I would also like to acknowledge my lab members. They have been there to cheer me up when I am down and to congratulate me when I succeed. They have made this journey special and memorable and I am thankful to be able to call them my friends. A special thank goes out to Dr. Gregory Sigurdson and Dr. Jacob Farr. They have been vital

iv teachers in the laboratory. Both are incredible individuals and I am grateful to have them as mentors.

My family deserves acknowledgment. My mother, Elaine, father, Gary, siblings and Adam have encouraged and supported me. They have taught me how to set goals and work to achieve them. My family are my biggest supports and I would not be where I am at today without them. Their love and never-ending support has been extremely valuable in getting to where I am at today.

Lastly, I would like to acknowledge and thank both The Ohio State University and the Food Science Department at Ohio State. Both have provided me with moral and financial support. This university and department have taught me so much in the last five years and I am thankful I decided to study at The Ohio State University.

Vita

December 5, 1995 ...... Born – Lima, Ohio

May 2017 ...... B.S. Food Science and Technology,

The Ohio State University

August 2017 ...... University Fellow, The Ohio State

University

August 2018 to Present ...... Graduate Teaching Associate, Department

of Food Science and Technology, The Ohio

State University

Fields of Study

Major Field: Food Science and Technology

Table of Contents

Abstract ...... ii

Acknowledgments...... iv

Vita ...... vi

List of Tables ...... x

List of Figures ...... xi

Chapter 1. Introduction ...... 1

Chapter 2. Literature Review ...... 4

2.1 Color ...... 4

2.1.1 Importance of Food Color ...... 4

2.1.2 Definition of Color ...... 5

2.1.3 Measuring Color ...... 7

2.2 Food Colorants ...... 9

2.2.1 Current Regulations ...... 9

2.2.2 Certified Colorants ...... 10

2.2.3 Colorants Exempt from Certification ...... 11 vii

2.3 Pigments Obtained from Plants ...... 11

2.3.1 Chlorophyll ...... 11

2.3.2 Carotenoids ...... 12

2.3.3 Betalains ...... 13

2.3.4 Anthocyanins ...... 13

2.4 Pyranoanthocyanins ...... 22

2.4.1 Early Reports of Pyranoanthocyanins ...... 22

2.4.2 Chemical Structure ...... 23

2.4.3 Potential Mechanism of Formation ...... 26

2.4.4 Enhanced Stability ...... 27

2.4.5 Colorimetric Properties ...... 28

2.4.6 Potential use in Food ...... 30

Chapter 3. The effects of pH and temperature on formation of pyranoanthocyanins from cyanidin-3-glucoside and cyanidin-3-rutinoside over time ...... 32

3.1 Abstract ...... 32

3.2 Introduction ...... 33

3.3 Materials & Methods ...... 35

3.3.1 Materials ...... 35

3.3.2 Methods...... 36

viii

3.4 Results and Discussions ...... 40

3.4.1 Effect of pH on Pyranoanthocyanin Formation ...... 40

3.4.2 Effect of Temperature on Pyranoanthocyanin Formation ...... 48

3.5 Conclusion ...... 52

Chapter 4. Comparison of ratios of anthocyanin to cofactor that lead to higher formation of pyranoanthocyanins from cyanidin-3-glucoside and cyanidin-3-rutinoside ...... 53

4.1 Abstract ...... 53

4.2 Introduction ...... 54

4.3 Materials and Methods ...... 57

4.3.1 Materials ...... 57

4.3.2 Methods...... 57

4.4 Results & Discussion ...... 64

4.4.1 Cofactor Pre-Screening ...... 64

4.4.2 Pyranoanthocyanin Formation ...... 65

4.4.3 Spectral Properties ...... 71

4.4.4 Color Properties ...... 73

4.5 Conclusion ...... 76

Chapter 5. Conclusion and Future Work ...... 77

Bibliography ...... 79

List of Tables

Table 3.1. The color properties, based on CIE – L*a*b* and CIE – L*C*abhab color systems, on day 0 and day 30 for pH sample between pH 1.0 and 5.0. Stored at 25°C in the dark. Represented as the average (n=3) and the associated standard deviation in parenthesis...... 47

Table 3.2. The color properties, based on CIE – L*a*b* and CIE – L*C*abhab color systems, on day 0 and day 28 for 5°C, 15°C and 25°C. Stored in the dark at pH 3.0.

Represented as the average (n=3) and the associated standard deviation in parenthesis. . 52

Table 4.1. Colorimetric data for all samples on day 0 and day 42. Data is present as average (n=3) and standard deviation in parenthesis ...... 74

List of Figures

Figure 2.1. The three required components, a light source, object and observer, required to experience color and the visible spectrum (Modified from Konica Minolta Inc., 2007;

National Weather Services, 2010)...... 6

Figure 2.2. An example of the Munsell color system diagram with a value of 5, chroma of 6 and a purple-blue hue (Worlstad & Smith, 2010)...... 8

Figure 2.3. Chemical structure of β-carotene, which is the most abundant natural carotene and can be found in plants, algae, fungi, bacteria and animals (Britton et al.,

2004)...... 13

Figure 2.4. The anthocyanin aglycone and the six main anthocyanidins found in nature

(Giusti & Wrolstad, 2003)...... 14

Figure 2.5. pH dependent conversions on mono-glycosylated anthocyanins when in aqueous solution (He & Giusti, 2010)...... 17

Figure 2.6. The general structure of pyranoanthocyanins that are derived from Malvidin-

3-glucoside and found in red wines. R1=H, COOH, CH3, (vinyl)phenols, (vinyl)flavanols

(De Freitas & Mateus, 2011)...... 24

Figure 2.7. The chemical structure of vitisin A that forms from the reaction of malvidin-

3-glucoside and pyruvic acid. Vitisin A forms naturally in red wines (He et al, 2012). .. 25

Figure 2.8. Nucleophilic addition of carbonyl compounds to Malvidin-3-glucoside to form pyranoanthocyanins with the additional D-ring (from Quaglieri, Jourdes, Waffo- teguo, & Teissedre, 2017)...... 26

Figure 2.9. Absorbance spectrum of malvidin-3-glucoside and carboxypyrano-malvidin-

3-glucoside (Vitisin A) at various pH levels in aqueous solution (Extracted from He et al.,

2010)...... 29

Figure 3.1. Formation of a carboxypyranoanthocyanin by nucleophilic cycloaddition of cyanidin-3-glucoside and pyruvic acid to form 5-Carboxypyranocyanidin-3-glucoside

(Modified from De Freitas & Mateus, 2011)...... 34

Figure 3.2. Pyranoanthocyanin formation yields (Eq 3.1) at pH values between 1.0 and

5.0 for day 3, 7, 14, 21 and 30. Incubated at 25°C in the dark. Points are the averages

(n=3) and the error bars are the associated standard deviations...... 41

Figure 3.3. The percentage of total pigment remaining on days 3, 14 and 30 in relation to day 0 (Eq 3.2) at 12 different pH values. All samples incubated at 25°C in the dark. The averages (n=3) are represented the associated standard deviations are error bars...... 43

Figure 3.4. (A) The lambda maximum at each pH value and 2 control samples (B) and the associated absorbance at the lambda maximum for day 0 and day 30. Samples incubated at 25°C in the dark. Averages are presented (n=3) and the associated standard deviations are represented as error bars ...... 44

Figure 3.5. (A) Chromatogram on day 0 at pH 3.0 and 25°C (B) Chromatogram on day

28 at pH 3.0 and 25°C (C) Spectra from 280nm to 800nm obtained from the uHPLC. The solid line is cyanidin-3-glucoside (C3G) spectra and the dashed line is the spectra of the

xii pyranoanthocyanins (PACNs) derived from cyanidin-3-glucoside (C3G) and cyanidin-3- rutinoside (C3R)...... 46

Figure 3.6 (A) Percentage of pyranoanthocyanin formation yield (Eq. 3.1) and (B) percentage of the total pigment remaining (Eq 3.2) in relation to day 0 at 5°C, 15°C and

25°C in pH 3.0 buffer. Information obtained from chromatograms at 500-520nm. All points are averages (n=3) and error bars are the associated standard deviations...... 49

Figure 3.7. (A) The lambda maximum for each temperature and the (B) associated absorbance at the lambda maximum for day 0 and day 28. Samples incubated in the dark at pH 3.0. Averages are presented (n=3) and the associated standard deviations are represented as error bars ...... 51

Figure 4.1. Formation of a carboxypyranoanthocyanin, 5-carboxypyranocyanidin-3- glucoside, by the reaction of the anthocyanin, cyanidin-3-glucoside, and pyruvic acid

(Modified from Farr & Giusti, 2018)...... 55

Figure 4.2. Pyranoanthocyanin formation and anthocyanin content were monitored with

HPLC. Chromatograms from day 0 and day 42 for CA1:30 and PA1:200 are shown. C3G

= cyanidin-3-glucoside, C3R = cyanidin-3-rutinoside, PACN = Pyranoanthocyanins derived from cyanidin-3-glucoside and cyanidin-3-rutinoside ...... 66

Figure 4.3. (A) Percent pyranoanthocyanin formation yield, Eq 4.1 (B) percent pyranoanthocyanin pigment to total pigment, Eq 4.2. On day 7, 28 and 42, samples are the average (n=3) and the error bars are the associated standard deviations ...... 67

xiii

Figure 4.4. The total amount of pigment remaining at tn in comparison to the total pigment on day 0, Eq 4.3. The average is presented (n=3) and the associated standard deviations are represented as error bars...... 69

Figure 4.5. The percentage of cofactor remaining throughout the experiment. The data points are the average (n=3) and the standard deviations are the error bars. Pyruvic acid remaining was calculated with Eq 4.4. and caffeic acid was calculated with Eq 4.5...... 70

Figure 4.6. (A) The lambda maximum of the control, PA1:200 and CA1:30 samples on day 0, 14, 28 and 42 of the experiment and the (B) associated absorbance at the lambda maximum. The chart shows the number values of the lambda maximum and associated absorbance at the lambda maximum. The average is presented (n=3) with the standard deviations as the error bars or in parentheses...... 72

xiv

Chapter 1. Introduction

Individuals use color to evaluate food products everyday (Clydesdale, 1993). The color of a product provides information regarding the foods flavor, safety, texture and nutritional value (Sigurdson, Tang, & Giusti, 2017). For example, individuals tend to avoid brown fruits and vegetables in the store, because brown is indicative of spoilage and decay. Food companies typically use synthetic dyes to color food or enhance the color of products, because color is such an important attribute to food. Synthetic colorants are often used, because they are stable under a wide range of conditions, are low in cost and have high tinctorial strength (Cevallos-Casals, & Cisneros-Zevallos,

2004). However, there has recently been a demand for naturally derived colorants, because consumers want more natural products and have health concerns over synthetic dyes.

The increasing demand for naturally derived colorants has led food manufactures to explore pigments derived from natural sources. Anthocyanins are pigments with vibrant red, blue and purple hues derived from fruit and vegetable sources (Andersen &

Jordhein, 2014). However, these attractive pigments are subject to degradation by pH, temperature, light, high water activities and bleaching reactions (Cevallos-Casals, &

Cisneros-Zevallos, 2004). To help stabilize anthocyanins co-pigmentation and metal

1 complexation have been studied, and more recently pigments derived from anthocyanins have been gaining attention.

Pyranoanthocyanins were first found in aged wine systems when anthocyanins reacted with yeast metabolites, such as pyruvic acid, to form pyranoanthocyanins

(Somers, 1971; Fulcrand, Cameira dos Santos, Sarni-Manchado, Cheynier, & Favre-

Bonvin, 1996; Bakker, & Timberlake, 1997). Pyranoanthocyanins have an additional fourth ring between C-4 and the C5-hydroxyl group of the anthocyanin molecule (He et al., 2012). This additional ring is likely attributed to the enhanced stability of the pyranoanthocyanin over the anthocyanin counterpart. Pyranoanthocyanins are more resilient to pH changes, water addition and bleaching reactions (Oliveira, Mateus, Silva,

& De Freitas, 2009; Oliveira et al., 2006). However, pyranoanthocyanin formation takes time and pyranoanthocyanins form at low rates limiting their application as potential food colorants (Benito, Morata, Palomero, González, & Suárez-Lepe, 2011; Morata, Calderón,

González, Gómez-Cordovés, & Suárez, 2007). If pyranoanthocyanins are to be used as food colorants their formation yields must be increased.

It was hypothesized that the pH of the solution, the incubation temperature, and the type and quantity of cofactor used could all be altered to help increase pyranoanthocyanin formation yields. The objective of this study was to enhance pyranoanthocyanin formation yields by investigating the effects of incubation temperature, pH and cofactor.

This study first investigated the effect of pH and temperature on pyranoanthocyanin formation. It is known that pH affects the equilibrium state of the

2 anthocyanin molecule which is a required reaction partner for pyranoanthocyanin formation. Additionally, it is likely the flavylium cation form, with electrophilic character, is required for pyranoanthocyanin formation. pH can also affect polymerization of pyruvic acid, which was used as a cofactor in this study. Therefore, it was hypothesized that pH would affect pyranoanthocyanin formation. Temperature is known to increase reaction rates and cause anthocyanin degradation, so it was predicted the incubation temperature would likely affect pyranoanthocyanin formation.

This study also investigated the effect of cofactor type and cofactor quantity on pyranoanthocyanin formation. Pyranoanthocyanins form by the cycloaddition of a cofactor with a nucleophilic vinyl group and anthocyanin molecule (De Freitas &

Mateus, 2011). Studies have shown many different compounds including, hydroxycinnamic acids, vinylphenols, pyruvic acid, acetaldehyde and acetone are all viable cofactors for pyranoanthocyanin formation (Rentzsch, Schwarz, & Winterhalter,

2007). These cofactors all vary in size and chemical structure. Additionally, some cofactors, such as caffeic acid, are known to act as co-pigments with anthocyanins

(Castañeda-Ovando, Pacheco-Hernández, Páez-Hernández, Rodríguez, & Galán-Vidal,

2009). It was hypothesized that the cofactor type and quantity would affect pyranoanthocyanin formation.

Chapter 2. Literature Review

2.1 Color

2.1.1 Importance of Food Color

Individuals learn early in life what acceptable colors are and reactions to color are ingrained. Individuals will purchase one product over another based on the color of the product and will not even noticed their bias to color (Clydesdale, 1993). Color is important in all types of consumer goods including food and beverage products.

Color, texture and flavor are the three main quality attributes of food and beverages. Many experts believe that color may be the most important of the three, because color is the first sensory experience of a food product and an opinion is developed (Burrow, 2009). If the product is not visually appealing the consumer will not purchase the product, so they will never experience the texture and flavor of the product.

Individuals use color to analyze food products before consuming the product, and the color of the product can influence other sensory characteristics.

Color is used to determine the flavor, safety, texture and nutritional value of a product (Sigurdson, Tang, & Giusti, 2017). Early humans used color to identify and avoid toxic and spoiled objects. Individuals still use color to identify food safety concerns. For example, color is used to determine if meat is cooked or raw. People avoid brown fruit and vegetables in the store, because the brown color is associated with

4 spoilage and decay. Humans also use color to predict how a product will taste (Burrow,

2009). Green colored fruits are associated with a sour and bitter flavor, and as the fruit turns to a deep purple or red consumers know the fruit will be sweet. Individuals use color to analyze products without realizing it, making color an important attribute of food.

Color also affects the perceived flavor of products. The effect of color on flavor was shown in a study that demonstrated how red, green and yellow affected the threshold concentrations of sweet, sour, salty and bitter (Maga, 1974). A higher concentration of sucrose was required to reach the sweetness threshold in a yellow solution compared to the other colors indicating individuals did not associate yellow with sweetness. Yellow and green colored solutions decreased the sourness sensitivity indicating individuals associated yellow and green with sourness (Maga, 1974). Sour fruits such as lemons and limes are yellow and green in color creating an association between yellow and green with sourness. It has been demonstrated that color has a significant impact on basic taste.

Color is an important attribute in foods and beverages, because color is used to analyze and predict how a food will taste and if the food is safe. For a product to sell the color must be appealing to the consumer, which is why color communicate systems have been developed to quantitatively define color of products (Wrolstad & Smith, 2017).

2.1.2 Definition of Color

Seeing color is a visual experience that requires a colored object, an observer and a light source in the visible region (Figure 2.1). If one of the three factors are missing color cannot be perceived (Wrolstad & Smith, 2017). Light is separated into a spectrum

5 and different colors have different spectra. The visible spectrum is arranged as red

(longest wavelengths) > orange > yellow > green > blue > indigo > violet (shortest wavelengths). Color is perceived when an object reflects light in the visible spectrum of

380 to 770 nm, and the wavelengths stimulate the retina of the eye (Konica Minolta Inc.,

2007).

Figure 2.1. The three required components, a light source, object and observer, required to experience color and the visible spectrum (Modified from Konica Minolta Inc., 2007; National Weather Services, 2010). The brain interprets the character of light coming off of an object to determine color as seen by the eye. Visible light enters the eye through the cornea and is focused on the retina. Inside of the retina are cones and rods, which are both responsible for vision.

Cones are responsible for daylight and color vision, and contain specific red, green and blue protein receptors allowing us to see and interpret color. Rods are sensitive to low- intensity light and do not discriminate between colors. The optic nerve is responsible for sending signals to the brain where vision and color occur. Based on the “Color Opponent

Theory” the red, green and blue signals are transformed into one brightness signal. The brightness signal includes a lightness and darkness coder, blue-yellow and red-green hue signals (Wrolstad & Smith, 2017).

2.1.3 Measuring Color

Humans can perceive up to 10,000,000 different colors but have limited color memory. The colors people see are based on perception and are subjective. Therefore, an objective measure of color is needed to ensure products meet specifications (Worlstad &

Smith, 2010). Color spaces and instruments to measure color have been developed to interpret and quantify color.

All color systems address color based on hue, value and chroma or saturation, because color is three-dimensional (Worlstad & Smith, 2010). Hue is what we think of when we think of color (red, green, blue, purple, ect..). Value is the lightness and darkness, from black to white on a 0 to 100 scale respectively. Chroma is the transition from gray to pure chromatic color, or how dull or vivid the color is (McGuire, 1992).

Multiple color systems exist based on hue, value and chroma. In 1905 the Munsell

Color system was developed based on visual perception (Figure 2.2). The Munsell Color

System uses colored chips of various hue, lightness and saturation levels to compare

7 colored objects too. The Munsell Book of Colors can be purchased, and comparisons of colors can be made with the book (Konica Minolta Inc., 2007).

Figure 2.2. An example of the Munsell color system diagram with a value of 5, chroma of 6 and a purple-blue hue (Worlstad & Smith, 2010). The other main color systems are based on quantitative values produced by instruments. The CIE (Commission Internationale de l’Eclairage) is the international organization focused on color and color measurement. Two of the main color systems are named with the CIE, CIE XYZ and CIE L*a*b*. The Hunter CIE L*a*b* is another popular method used to measure color (McGuire, 1992; Worlstad & Smith, 2010). The tristimulus XYZ color system was developed in 1931 and is used to define color in a numerical fashion. The L*a*b* color space was developed in 1976 and explains color in a uniform way based on visual differences (Konica Minolta Inc., 2007).

When using color systems instrument settings must be predetermined to ensure reproducible results. The three main CIE illuminants are illuminant A (incandescent 8 light), illuminant C (overcast daylight) and illuminant D65 (average daylight). The most common observer angle is 10, because it best correlates with visual assessment. A 2 observer angle is commonly used as well. When setting observer functions and illuminants, transmission and reflectance curves can be translated into three numerical values used to describe color. These values are described as L for lightness coordinate, a as the red (+) to green (-) coordinate and b as the yellow (+) to blue (-) coordinate

(Worlstad & Smith, 2010). Color systems are important to quantify the color of specific products to ensure products remain in color specifications.

2.2 Food Colorants

Companies want to color their products to ensure their products make a good first impression on the consumer. The Federal Food, Drug and Cosmetic Act indicates that color additives are subject to FDA approval before use in food, drugs, cosmetics and medical devices that contact bodies of people or animals for a significant period of time

(Food and Drug Administration [FDA], 2018). This section will give a review of colorants used in foods and beverages.

2.2.1 Current Regulations

Food colorant regulations vary across the world and there is no standard international system (Sigurdson et al., 2017). In the United States color additives are regulated by the

US Food and Drug Administration (FDA). A color additive is defined as, “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

9 applied to a food, drug, or cosmetic or to the human body or any part thereof, is capable

(alone or through a reaction with another substance) of imparting a color thereto” (Code of Federal Regulations [CFR], 2018a). However, color additives are not treated the same as food additives. Color additives cannot obtain Generally Recognized As Safe (GRAS) exemption, but instead are subjected to premarket approval requirements unless the substance is used solely for a purpose other than coloring (FDA, 2017a).

The color additives regulations can be found in Title 21 of the Code of Federal

Regulations (CFR), parts 70-82. In the CFR all approved color additives are listed with the additives chemical specifications, uses and restrictions, labeling requirements and requirements for certification. Listed color additives fall into one of two categories described by the FDA: color additives subjected to FDA’s certification process

(21CFR74 subpart A-D) and color additives exempt from the certification process

(21CFR73 subpart A-D) (CFR, 2018b).

2.2.2 Certified Colorants

Colorants that are thought of as synthetic are subjected to batch certification. Each batch of colorants produced must be tested by the FDA (CFR, 2018b). In the batch certification process the physical appearance of the color additive is evaluated and the color additive is chemically analyzed for purity, moisture, residual salts, unreacted intermediates, colored impurities, other impurities, lead, arsenic and mercury (FDA,

2017b; CFR, 2018b). Certified colorants are typically called FD&C colorants. FD&C colorants can be in the form of dyes and lakes. Dyes are water soluble pigments, and lakes are used to color oils and fats by dispersion (FDA, 2017b; CFR, 2018b).

2.2.3 Colorants Exempt from Certification

Colorants that are thought to be safe without the batch certification process fall under the category of colorants exempt from certification (FDA, 2017b). Colorants that are exempt from certification are generally considered natural. However, there is no legal definition for natural causing confusion for the industry and consumers. If a colorant is exempt from certification it typically comes from natural sources such as plants, animals, and minerals, but a synthesized pigment that is identical to a pigment found in nature is classified as a colorant exempt from certification. Examples of colorants exempt from certification include: annatto extract, dehydrated beets, caramel, beta-carotene and grape skin extract (FDA, 2017b; CFR, 2018b)

2.3 Pigments Obtained from Plants

2.3.1 Chlorophyll

Chlorophylls are the green pigments present in most green plants, algae and photosynthetic bacteria. Chlorophyll is present in many green vegetables, such as green beans and cucumbers (Schwartz, Cooperstone, Cichon, Von Elbe, & Giusti, 2017).

Chlorophylls are cyclic tetrapyrroles with a Mg2+ atom in the center bound with nitrogen in the porphyrin ring. Chlorophyll a and b are the predominant chlorophylls in plants and vegetables. Chlorophyll a contains a methyl group at C-3 and chlorophyll b contains a formyl group at C-3. Chlorophylls contain a phytol group making chlorophylls lipid soluble (Jackson, 1976). A loss of the vibrant green color during processing is associated with chlorophyll degradation. When the chlorophyll contains Mg2+ in the tetrapyrrole the compound exhibits a green color. However, when the compound is exposed to heat

11 or acid the magnesium can be lost and the compound becomes olive green in color

(Schwartz et al., 2017). For example, when comparing the color of a fresh green bean to a canned green bean the fresh green bean is a more vibrant green and the canned green bean is more of an olive green. Typically, zinc will be added to canned vegetables prior to processing, so the zinc can replace the magnesium. The zinc complex is more stable than the magnesium complex to acid and heat (Schwartz et al., 2017).

2.3.2 Carotenoids

Carotenoids are responsible for the yellow, orange and red colors of some vegetables, fruits and plant life. Carotenoids are heat sensitive and lipophilic. Over 700 carotenoids have been identified in nature and roughly 60 exist in human food (Britton,

Liaaen-Jensen, & Pfander, 2004). Carotenoids are present in all photosynthetic organisms and are the pigments responsible for the yellow and orange leaves in the fall.

Additionally, carotenoids are present in carrots (α-carotene), shrimp (astaxanthin), red peppers (capsanthin), tomatoes (lycopene), leafy greens (violaxanthin) and many other human foods (Schwartz et al., 2017). There are two main classes of carotenoids, based on chemical structure, including hydrocarbon carotenes and oxygenated xanthophylls

(Britton et al., 2004).

Figure 2.3. Chemical structure of β-carotene, which is the most abundant natural carotene and can be found in plants, algae, fungi, bacteria and animals (Britton et al., 2004). 2.3.3 Betalains

Betalains are nitrogen-containing pigments consisting of two structural groups, betacyanins and betaxanthins. Betacyanins are red/violet in color and have a lambda maximum around 535-538 nm. Betaxanthins are yellow/orange in color and have a lambda maximum around 460-477 nm. Roughly 55 betalain structures have been identified, and betalins in red beets are the most studied. Betalins are water soluble and are stable from pH 4.0 to 7.0. When betalins are heated or subjected to acidic solutions the betanin is hydrolyzed and changes from a red to yellow color. When betalins are exposed to oxygen, oxidative degradation of the pigment occurs leading to pigment lose

(Schwartz et al., 2017).

2.3.4 Anthocyanins

Anthocyanins make up the largest group of water-soluble, naturally occurring pigments with more than 700 anthocyanin structures (He & Giusti, 2010; Andersen &

Jordheim, 2014). Anthocyanins are responsible for the red, blue and purple colors of many fruits and vegetables (Andersen & Jordheim, 2014). It is believed that anthocyanins are essential to plant survival as the vibrant colors attract insects and animals, which are

13 essential for seed dispersal and pollination, and protect the plant tissue against UV damage (He & Giusti, 2010).

Chemical Structure

Anthocyanins are polyphenolic compounds that belong to the flavonoid group

(Sigurdson et al., 2017). Anthocyanins have a characteristic aglycone structure composed of a C-6 (A ring) – C-3 (C ring) – C-6 (B ring) carbon backbone (He & Giusti, 2010).

When the anthocyanin aglycone has no sugar attachments the molecule is known as an anthocyanidin. There are 27 naturally occurring anthocyanidins, but roughly 90% are derived from the 6 anthocyanidins shown in Figure 2.4. The anthocyanidin backbones differ by what is attached to the 3` and 5` positions (Schwartz et al., 2017).

Figure 2.4. The anthocyanin aglycone and the six main anthocyanidins found in nature (Giusti & Wrolstad, 2003).

The anthocyanidin is highly unstable, because the 3-hydroxyl group destabilizes the chromophore. Therefore, anthocyanidins are almost always glycosylated and known as anthocyanins. Glycosylation helps to increase water solubility, polarity and stability

(He & Giusti, 2010). Mono-glycosylated anthocyanins typically have sugars attached at

C-3, and di-glycosylated anthocyanins typically have sugars attached at C-3 and C-5.

Although less common anthocyanins can also have sugars attached at C-7, -3`, -4` and/or

-5` (Schwartz et al., 2017). Glucose and rhamnose are the most common sugars attached to the aglycone, but many other sugars such as galactose, arabinose, xylose, rutinose, and sambubiose can also be attached (He & Giusti, 2010). Monosaccharide to trisaccharide’s have been reported as anthocyanidin attachments, but a tetrasaccharide has not been reported. The most abundant anthocyanin found is nature is cyanidin-3-glucoside

(Andersen & Jordheim, 2006).

In addition to sugars anthocyanins can also have acids attached. Roughly 65% of all plant anthocyanins identified are acylated (Andersen & Jordheim, 2014).

Anthocyanins can be acylated with the following aromatic acids, p-coumaric, caffeic, ferulic, sinapic, gallic and p-hydroxybenzoic acids. The following aliphatic acids can also be attached to anthocyanins, malonic, acetic, malic, succinic and oxalic acid. Typically, acylation occurs on the sugar in the C-3 position of the anthocyanin (Schwartz et al.,

2017). Acylation generally increases the overall stability of the anthocyanin in solution making acylated anthocyanins appealing for food use. Common sources of acylated anthocyanins include radishes, red potatoes, red cabbage, black carrots and purple sweet potatoes (Giusti & Wrolstad, 2003).

Anthocyanins structure changes reversibly as pH changes making them unique among flavonoids, as shown in Figure 2.5 (He & Giusti, 2010). The anthocyanin molecule exists in four main forms depending on the pH (Figure 2.5). At acidic pH values (pH 1 to 2) the flavylium cation is predominate and leads to purple and red colors.

Two colorless species, carbinol pseudobases and chalcones, begin to predominate from pH 3 to 6. Around pH 6 and above the quinoidal blue species begins to form. However, when the pH of the solution reaches pH 7 the anthocyanins begin to degrade (Castañeda-

Ovando et al., 2009; He & Giusti, 2010). The various equilibrium states of the anthocyanins lead to color changes as pH changes. Additionally, anthocyanins are relatively unstable at high pH values, and a pH as low as 4.0 promotes anthocyanin degradation (Castañeda-Ovando et al., 2009).

Figure 2.5. pH dependent conversions on mono-glycosylated anthocyanins when in aqueous solution (He & Giusti, 2010). Anthocyanin Stability

Although anthocyanins provide vibrant colors in fruits and vegetables their stability is limited. Anthocyanins undergo many degradative reactions. Anthocyanin stability is decreased as pH increases, and anthocyanins are most stable from pH 1-3

(Cabrita, Fossen, & Andersen, 2000). As temperature and light exposure increase anthocyanin degradation increases (Eiro & Heinonen, 2002). Enzymes such as polyphenoloxidase, peroxidase and glycosidase actively cause anthocyanin degradation.

Increased water activity will promote degradation (Wrolstad, Durst, & Lee, 2005).

Anthocyanins are subject to bleaching by sulfites, ascorbic acid and hydrogen peroxide

(Farr & Giusti, 2018; Wrolstad et al., 2005). Anthocyanins will also condense with phenolics and become polymeric pigments, and acetaldehyde accelerates condensation

(Wrolstad et al., 2005).

Although anthocyanins have limited stability there are ways to help stabilize the anthocyanin pigments. The structure of the anthocyanin molecule determines how rapidly the pigment will degrade. Anthocyanins with more sugars attached are typically more stable, and acylated anthocyanins are even more stable (Giusti & Wrolstad, 2003).

Anthocyanins can also undergo co-pigmentation where the anthocyanin forms a complex with a colorless compound to stabilize and possibility shift the color of the original anthocyanin (Castañeda-Ovando et al., 2009). Anthocyanins can also undergo metal complexation to help stabilize the pigment. Additionally, some anthocyanin derived compounds, such as the pyranoanthocyanin, can help to stabilize the anthocyanin molecule (Castañeda-Ovando et al., 2009).

Food Colorant Trends

Changes and trends in the food industry occur constantly, and it is important to stay up to date and understand these trends to better understand the consumer. Consumers currently want more “natural” products leading to an increased demand for “natural” colorants.

Many large companies, such as Kraft and General Mills, are in the process of removing synthetic colorants from their products and replacing them with natural

18 colorants to meet consumers desires. Dunkin’, formally Dunkin’ Donuts, announced in

2018 that all artificial dyes have been replaced with natural color alternatives in their donuts (Golden, 2018). By removing artificial dyes and using natural colorants brands can attract consumers who want natural ingredients. After Dunkin’ had replaced artificial dyes in their donuts with colors from plant sources, individuals were asked if they would be more likely to purchase donuts from Dunkin’. And 52% of the consumers indicated they would be more interest in purchasing donuts from Dunkin’ (Reed, 2018). The desire for a clean label and “better-for-you” foods is encouraging the use of colorants derived from nature over artificial colorants (Reed, 2018).

An additional reason for the replacement of FD&C colors with natural alternatives is the concern of potential negative health effects associated with synthetic colorants. The fear of potential negative effects grew in 2007 when a South Hampton Study was published linking hyperactivity in 3-year-old and 8/9-year-old children to artificial colors

(McCann et al., 2007). The study consisted of two groups, 3-year-old and 8/9-year-old children, and children were given a drink with sodium benzoate and synthetic colorants or a placebo. Increased hyperactivity was associated with the children who consumed the sodium benzoate and artificial colorant mixture (McCann et al., 2007). After the study was published consumers started to associate synthetic colorants to negative health effects even though the European Food Safety Authority concluded in their review, “In the context of the overall weight, . . . the considerable uncertainties, such as the lack of consistency and relative weakness of the effect and the absence of information on the clinical significance of the behavioral changes observed, the Panel concludes that the

19 findings of the study cannot be used as a basis for altering the ADI of the respective food colours or sodium benzoate” (European Food Safety Authority, 2008, Conclusion, para.

4). The South Hampton Study raised considerable concern over the safety of synthetic colorants even though the study was deemed invalid. Consumer concern over the potential side effects of artificial colorants has continued to grow, so the demand for natural alternatives has increased.

Overall, there is a current trend for foods with clean labels, and to have a clean label,

FD&C colors need to be replaced with naturally derived colorants. Health concerns associated with artificial colorants and potential health benefits associated with colorants derived from nature have encouraged the use of naturally derived colorants. With a demand for colorants derived from nature food manufactures are replacing synthetic colorants with natural alternatives.

Potential Health Benefits of Anthocyanins

When we think of traditional sayings such as, “an apple a day keeps the doctor away” or interesting associations such as the French Paradox we see the effects anthocyanins may have on the body. Epidemiological studies have shown an association between foods high in polyphenols, such as fruits, vegetables and wines, and a decrease in chronic disease (He & Giusti, 2010). Anthocyanins and their health promoting properties are gaining more and more attention by researchers, the industry and consumers.

There is an interest in the potential health benefits of anthocyanins, because of anthocyanins ability to function as antioxidants. Antioxidants protect the body against the

20 damaging effects of free radicals and some chronic diseases that occur during aging.

Antioxidants help to mitigate oxidative stress, which may help protect the body against neurodegenerative diseases. Anthocyanins and phenolics in fruits and vegetables may have an anticancer effect, because of anthocyanins ability to down-regulate cyclooxygenase-2 expression and enzyme activity leading to antiproliferation of some cancer cell lines. Anthocyanins antioxidant activity may inhibit oxidation of LDL and function as anti-inflammatory agents both of which help to protect against cardiovascular diseases (Nile & Park, 2014).

However, a major question that exist is whether anthocyanins are bioavailable in the body. There are more than 700 types of anthocyanins, and anthocyanins can vary in the degree of polymerization, unsaturation, glycosylation, acylation, hydroxylation and methoxylation (Andersen & Jordheim, 2014). The variation in anthocyanins chemical structure makes anthocyanins difficult to study regarding bioavailability (Milbury, Cao,

Prior, & Blumberg, 2002). Anthocyanins have been found in the glycosylated form in human plasma after consumption potentially indicating bioavailability. However, the information on how the anthocyanins are absorbed, the quantity absorbed, and the metabolites formed is limited (Milbury et al, 2002). To understand the health benefits of anthocyanins in the body the bioavailability of anthocyanins in the human body must be better understood.

2.4 Pyranoanthocyanins

2.4.1 Early Reports of Pyranoanthocyanins

Anthocyanins are responsible for the vibrant colors of many fruits, vegetables, flowers and young red wines. Anthocyanins are relatively unstable and subject to color change as pH changes, but in old red wines a vibrant red orange color still exist (Oliveira et al., 2009). Somers predicted that the grape anthocyanins responsible for the red color in young red wines are displaced and transformed over time to more stable pigments

(1971). Somers showed these new pigments are more stable to pH changes in solution and more resistant to decolorization by sulphur dioxide (Somers, 1971). It was likely that pyranoanthocyanins were a part of the pigments Somers investigated.

In the late 1990s and early 2000s pyranoanthocyanins were gaining attention as newly discovered, more stable red/orange pigments. The pigments appeared to be malvidin anthocyanins containing a C3H2O2 group that linked carbon 4 and the 5- hydroxyl group of the anthocyanin forming an additional ring (Fulcrand et al., 1996;

Fulcrand, Benabdeljalil, Rigaud, Cheynier, & Moutounet, 1997; Bakker et al., 1997;

Bakker, & Timberlake, 1997). This new class of compounds appeared to be resistant to sulfur dioxide bleaching and more stable at higher pH values compared to its anthocyanin counterpart (Bakker & Timberlake, 1997). The increased stability of pyranoanthocyanins grabbed the attention of researchers and industries in search of stable naturally derived pigments.

Pyranoanthocyanins were first found in wines, and early research was performed in wine systems. Since, pyranoanthocyanins have been identified in fruit juices (Rein,

Ollilainen, Vahermo, Yli-Kauhaluoma, & Heinonen, 2005) and red onions, Allium cepa, with carboxypyranocyanidin as the pyranoanthocyanin (Fossen & Andersen, 2003).

Strawberries, Fragaria ananassa, have been found to contain trace amounts of 5- carboxypyranopelargonidin (Andersen, Fossen, Torskangerpoll, Fossen, & Hauge, 2004).

Studies have shown that pyranoanthocyanins are both derived from food sources, such as wine and fruit juices, and found naturally in strawberries and onions.

2.4.2 Chemical Structure

The signifying feature of the pyranoanthocyanin is the presence of a fourth ring

(D) between the C-4 and C5-hydroxyl group of the anthocyanin molecule (Figure 2.6).

This additional ring creates a second heteroaromatic ring compared to a free anthocyanin which has one heteroaromatic ring (He et al., 2012). Pyranoanthocyanins differ between each other based on the anthocyanin type and the attachment at C-10 of the new pyrano ring (Oliveira et al., 2009). The most studied pyranoanthocyanins are the pyranoanthocyanins that form from the reaction of the anthocyanin and yeast metabolites produced during fermentation, including pyruvic acid, acetoacetic acid, and acetaldehyde

(De Freitas & Mateus, 2011).

Figure 2.6. The general structure of pyranoanthocyanins that are derived from Malvidin- 3-glucoside and found in red wines. R1=H, COOH, CH3, (vinyl)phenols, (vinyl)flavanols (De Freitas & Mateus, 2011).

Carboxypyranoanthocyanins are the pigments formed by the reaction of an anthocyanin and pyruvic acid. They have a carboxyl group attached to the D ring of the pyranoanthocyanin. Carboxypyranoanthocyanins are one of the most studied pyranoanthocyanins (Fulcrand et al., 1997; Bakker & Timberlake, 1997), because of their prevalence in wine systems as pyruvic acid is produced by yeast and lactic acid bacteria

(Morata et al., 2007). The carboxypyranoanthocyanins derived from Malvidin-3- glucoside, the main wine anthocyanin, are specifically referred to as vitisin A (Figure 2.7)

(Fulcrand et al., 1997). Vitisin A is the major anthocyanin derived pigment found in port red wine after one year of aging, showing the importance of vitisin A in wine color

(Mateus & De Freitas, 2001).

Figure 2.7. The chemical structure of vitisin A that forms from the reaction of malvidin- 3-glucoside and pyruvic acid. Vitisin A forms naturally in red wines (He et al, 2012).

Vitisin B is another major class of pyranoanthocyanins. Vitisin B differs from

Vitisin A, because Vitisin B lacks the carboxyl group at C-10 on the D ring (De Freitas &

Mateus, 2011). Vitisin B is formed by the reaction of acetaldehyde and malvidin-3- glucoside. Acetaldehyde is produced and released by yeast during the fermentation process (Morata et al., 2007).

Pinotins are another group of pyranoanthocyanins. Pinotins form when hydroxycinnamic acids or 4-vinylphenols react with free anthocyanins to form hydroxyphenyl-pyranoanthocyanins. In red wines the main hydroxycinnamic acids present are p-coumaric, caffeic, ferulic and sinapic acids. However, unlike vitisins, hydroxycinnamic acids accumulate after alcohol fermentation. The most rapidly synthesized pinotin, Pinotin A (pyranomalvidin-3-glucoside-catechol), was observed after 2.5 to 4 years of aging. After 2.5 years there are very little free anthocyanins for the caffeic acid, or other hydroxycinnamic acids, to react with (He et al., 2012).

2.4.3 Potential Mechanism of Formation

Anthocyanins in red wines degrade and react to form more complex molecules such as pyranoanthocyanins. Pyranoanthocyanins are predicted to be cycloaddition products, resulting from SN2 nucleophilic substitution at the C-4 position of the anthocyanin moiety (Figure 2.8). This leads to the cyclization and formation of the additional D-ring between the -OH group at C-5 and C-4 (Quaglieri, Jourdes, Waffo- teguo, & Teissedre, 2017). The formation of pyranoanthocyanins requires three important factors, an anthocyanin, a reaction partner and time.

Figure 2.8. Nucleophilic addition of carbonyl compounds to Malvidin-3-glucoside to form pyranoanthocyanins with the additional D-ring (from Quaglieri, Jourdes, Waffo- teguo, & Teissedre, 2017).

Pyranoanthocyanins are derived from anthocyanins, so an anthocyanin source must be present. The formation of pyranoanthocyanins is related to the reactivity of the anthocyanin molecule. Under aqueous conditions, such as wine conditions, anthocyanins

26 react readily and easily to form molecules such as pyranoanthocyanins (De Freitas &

Mateus, 2011; Quaglieri et al, 2017). Anthocyanins are slightly electrophilic in the flavylium cation form (Dangles & Fenger, 2018), so it is predicted the anthocyanin needs to be in the flavylium cation form to form pyranoanthocyanins.

A reaction partner must also be present. Some common partners include hydroxycinnamic acids, vinylphenols, vinylflavanols, pyruvic acid, acetaldehyde, acetone, and further enolizable molecules (Rentzsch et al., 2007). The reaction partners, or cofactors, are typically carbonyl compounds able to undergo keto-enol tautomerization. The cofactor must be in the enol tautomer to react with the anthocyanin to form the pyranoanthocyanin (De Freitas & Mateus, 2011).

The third factor is time. Naturally pyranoanthocyanins form slowly in wines and require time. However, recent research is showing that accelerated conditions can be used to decrease the required time (Rentzsch et al., 2007)

2.4.4 Enhanced Stability

As discussed above the pyranoanthocyanin has an additional D ring. The D ring is oxygenated, which leads to a more dynamic equilibrium. The more dynamic equilibrium state may help to stabilize the pyranoanthocyanin over a wide pH range (He et al., 2012).

This new ring also creates a steric hinderance effect. The steric effect helps to protect the pyranoanthocyanin molecule from nucleophilic addition of water. If water is added to the molecule the colorless hemiketal forms, so the D ring is likely protecting the chromophore (Oliveira et al., 2009). Pyranoanthocyanins are also more resistant to bisulfite bleaching than the anthocyanin. This additional stability can again be attributed

27 to the additional D ring. When anthocyanins undergo bisulfite bleaching the anionic bisulfite is added to the anthocyanin molecule at C-2 or the preferred position of attack,

C-4. The D-ring of the pyranoanthocyanin is attached at C-4 blocking the bisulfite addition. Pyranoanthocyanins may undergo slight color loss in the present of bisulfite, because the less favorable C-2 position is still open for attack (Oliveira et al, 2006). Farr and Giusti found that 5-Carboxypyranocyanidin-3-galactoside had a significantly smaller color shift (three-fold less) than cyanidin-3-galactoside in the presence of ascorbic acid

(2018). The ability of the pyranoanthocyanin to maintain color can likely be attributed to the protection the additional D-ring provides.

2.4.5 Colorimetric Properties

The additional ring pyranoanthocyanins possess, in addition to the three anthocyanin rings, changes the spectral and colorimetric properties of the pyranoanthocyanin from the anthocyanin. Pyranoanthocyanins are hypsochromically shifted in comparison to the parent anthocyanin. For example, cyanidin-3-glucoside has a

λmax of 509 nm at pH 2.0 while 5-Carboxypyranocyanidin-3-glucoside has a λmax of 503 nm at pH 2.0 (Oliveira et al., 2006). The shift in the lambda maximum is associated with a color change from a red to a red/orange color (De Freitas & Mateus, 2011)

Many studies have looked at the spectral and colorimetric changes that occur when pyranoanthocyanins are formed from anthocyanins. Oliveria and others showed that at pH 1 and pH 2 pyranoanthocyanins had lower chroma, meaning the color was less saturated, and higher lightness values in comparison to the anthocyanin counterparts

(2006). However, the reverse was true at pH 5.0 and pH 7.0. At pH 5.0 and 7.0 the

28 pyranoanthocyanins had an increase in chroma and lightness in comparison to the anthocyanin counterparts (Oliveira et al., 2006). Another study showed that the pyranoMV-flavanols were relatively stable from pH 2 to 6 in aqueous solution, while malvidin-3-glucoside was almost colorless at pH 3.6 and above (He, Carvalho, Mateus, &

De Freitas, 2010). These studies showed that at low pH levels, pH 1 to pH 3, the pyranoanthocyanins may not exhibit as much color as the anthocyanin counterparts, but as the pH increases the pyranoanthocyanins remain stable and exhibit more color than the anthocyanins.

Figure 2.9 shows the spectrum of malvidin-3-glucoside and 5- carboxypyranomalvidin-3-glucoside at various pH levels. Figure 2.9 clearly shows the improved stability of the pyranoanthocyanin over the anthocyanin as pH increases (He et al., 2010).

Figure 2.9. Absorbance spectrum of malvidin-3-glucoside and carboxypyrano-malvidin- 3-glucoside (Vitisin A) at various pH levels in aqueous solution (Extracted from He et al., 2010). It is interesting that even at low pH levels the anthocyanins exhibit more color than the pyranoanthocyanins. Based on basic resonance rules it would be predicted that

29 the molecule with the more conjugated π-electrons, more conjugated chromophore, would have a longer wavelength and higher absorbance. However, this is not observed with pyranoanthocyanins (Carvalho, Oliveira, de Freitas, Mateus, & Melo, 2010). A recent theoretical study proposed that the rotamerization of the B-ring leads to less electron conjugation in the pyranoanthocyanin than in the anthocyanin (Carvalho et al.,

2010). The decrease in conjugation may potentially describe the spectral characteristics of the pyranoanthocyanin.

2.4.6 Potential use in Food

As discussed above food colorants fall into one of two categories. Colorants that are subjected to batch certification and colorants exempt from certification (CFR, 2018b).

Colorants that are exempt from certification typically come from natural sources, such as plant, animals and minerals. However, a synthesized pigment that is identical to a pigment found in nature can also be classified as exempt from certification (Sigurdson et al., 2017). For example, β-Carotene is synthesized to be identical in structure to the β-

Carotene found in carrots, so both the synthesized and natural source of β-Carotene are colorants exempt from certification (Sigurdson et al., 2017).

This leads to our discussion on the possibility of pyranoanthocyanins as food colorants. Pyranoanthocyanins are stable over a wide pH range making them applicable for food use. Pyranoanthocyanins form naturally over time in wines and fruit juices

(Bakker & Timberlake, 1997; Rein et al., 2005). However, formation of pyranoanthocyanins takes time and pyranoanthocyanins form at low yields.

Pyranoanthocyanins, if separated from wine, would be extremely expensive to use as

30 food colorants. If the process of formation for pyranoanthocyanins could be optimized outside of the wine system, and the chemical structure would be identical to the pigments found in wine, could these pigments receive exempt from certification status by the

FDA? To my knowledge, pyranoanthocyanins have not been brought to the attention of the FDA as a food colorant, likely because the process of forming pyranoanthocyanins has not been optimized.

Epidemiological data suggest pyranoanthocyanins are likely safe to consume.

Pyranoanthocyanins have been consumed for hundreds of years in wine, and no known issues have been found with moderate consumption of wine. A study by Zhu and others showed that methylpyranoanthocyanins (from red grapes) had a protective effect against

H2O2 induced MRC-5 cell damage (2015). Additionally, when the methylpyranoanthocyanin pigments were fed to mice at high levels no toxicity was found after macro and microscopic assessments, supporting the conclusion that pyranoanthocyanins are non-toxic (Zhu et al., 2015). To my knowledge there is no current evidence that pyranoanthocyanins have a toxic effect, but more research should be carried out to verify the non-toxic effect of pyranoanthocyanins.

Efficient production of pyranoanthocyanins could make them economically feasible to use as food colorants, and if produced under juice processing condition could classify them as colorants exempt from certification making them potential food colorants with increased stability.

Chapter 3. The effects of pH and temperature on formation of pyranoanthocyanins from cyanidin-3-glucoside and cyanidin-3-rutinoside over time

3.1 Abstract

Anthocyanins are vibrant colorants from plant sources, but their limited stability restricts use as food colorants. Pyranoanthocyanins are anthocyanin derived pigments with enhanced stability that are formed at low rates. The objective of this study was to evaluate the effects of pH and temperature on pyranoanthocyanin formation. To determine the effect of pH cyanidin-3-glucoside and cyanidin-3-rutinoside (500 µM) were mixed with pyruvic acid at 200X [anthocyanin]. Anthocyanins and pyruvic acid were brought to 12 different pH values between pH 1.0 - 5.0 and stored at 25°C in the dark. Samples in pH 3.0 buffer were stored at 5°C, 15°C and 25°C in the dark.

Anthocyanin and pyranoanthocyanin contents were monitored with HPLC-PDA and UV-

Visible spectrophotometry. Pyranoanthocyanins were formed at the greatest rate in pH

3.1 (yield ~18% in 7 days), and by day 21 pH 2.8, 3.1 and 3.5 all showed similar yields

(yields ~24% in 21 days) with no significant difference between them, suggesting a maximum amount of formation under the conditions of this study. The highest pyranoanthocyanin formation yields were obtained with incubation at 25°C (yield ~25% in 21 days); reducing temperature slowed formation (yield ~23.5% in 28 days at 15°C).

This study showed that pH and temperature had a significant effect on pyranoanthocyanin formation and could be controlled to increase pyranoanthocyanin formation yields. 32

3.2 Introduction

Synthetic colorants are commonly used to ensure food and food products have vibrant and appealing colors (Burrows, 2009). However, as consumer suspicions of synthetic colorants arise and the demand for natural alternatives grows, there is increasing demand for naturally derived colorants. Anthocyanins are responsible for the blue, red, and purple colors in many fruits and vegetables and have been studied as naturally derived food colorants (Andersen & Jordheim, 2014). Anthocyanins are polyphenolic, water-soluble compounds comprised of more than 700 types known to exist in nature (Sigurdson et al., 2017; Andersen & Jordheim, 2014). However, anthocyanin application is restricted by their limited stability. Anthocyanins are labile compounds subject to color changes and degradation by pH, temperature, light and bleaching reactions (Cabrita et al., 2000; Eiro & Heinonen, 2002; Wrolstad et al., 2005).

Anthocyanins are the main pigments originally in wine systems, but wines can retain vibrant colors for many years. It was predicted and has since been proven that the grape anthocyanins present in young red wines are displaced and transformed into more stable pigments over time (Somers, 1971; Oliveira et al., 2009). Pyranoanthocyanins are one such class of compounds first found in aged red wine, that exhibit enhanced stability compared to their anthocyanin precursors. Pyranoanthocyanins have an additional ring between carbon 4 and the C5-hydroxyl of the anthocyanin molecule which likely contributes to their enhanced stability against pH, temperature, and bleaching (Figure 3.1)

(Fulcrand et al., 1997; Bakker et al., 1997; Bakker, & Timberlake, 1997). The increased

33 stability of pyranoanthocyanins in comparison to anthocyanins makes the pigments appealing as potential food colorants.

Figure 3.1. Formation of a carboxypyranoanthocyanin by nucleophilic cycloaddition of cyanidin-3-glucoside and pyruvic acid to form 5-Carboxypyranocyanidin-3-glucoside (Modified from De Freitas & Mateus, 2011). Pyranoanthocyanin formation studies are typically conducted in model wine systems with pH values near 3.7 to 4.0 and temperatures near 10°C to 20°C (Somers,

1971; Morata et al., 2007). Under these conditions formation generally takes time and formation occurs at low yields. For example, in a model wine system at pH 3.7 at 10°C vitisin A (pyranoanthocyanin from malvidin-3-glucoside and pyruvic acid) formation reached 18.48%, but required 232 days of incubation (Romero & Bakker, 2000). At 32°C maximum formation of vitisin A only reached 3.14% in 31 days (Romero & Bakker,

2000). In a wine system near pH 4.0 stored at 15°C vitisin A formation yields reached

2.03% in 10 weeks (Morata et al., 2007). The low formation yields, and long incubation times limits the application of pyranoanthocyanins.

The pH of the solution greatly affects the stability and equilibrium form of the anthocyanin molecule, which is a precursor in pyranoanthocyanin formation (Figure 3.1).

At acidic pH values (pH < 2) the flavylium cation form predominates. As the pH 34 increases (pH 3 to 6) the carbinol pseudobase and chalcone begin to predominate. If the pH is near neutral the quinonoidal base predominates. As the chemical equilibria of the anthocyanin changes with pH, it was predicted that pH would influence pyranoanthocyanin formation.

Increased temperature accelerates reaction rates, because the increased temperature adds energy to the system increasing collision rates thus increasing reactant interaction (Brown, LeMay, Bursten, Murphy, & Woodward, 2012). However, high temperatures can cause degradation of anthocyanins (Andersen, & Jordheim, 2006). The effect of temperature on a reaction system likely influences pyranoanthocyanin formation. Temperature and pH impact anthocyanin stability and equilibrium state, so it was predicted pH and temperature would affect pyranoanthocyanin formation. The objective of this study was to determine the effect of pH and temperature on pyranoanthocyanin formation in order to increase pyranoanthocyanin yields.

3.3 Materials & Methods

3.3.1 Materials

Blackberries (Rubus sp., SUNBELLE, Los Reyes, Mexico) were purchased from a local grocery store in Columbus, OH, U.S. Lab grade pyruvic acid was purchased from

Sigma Aldrich (St. Louis, Missouri, U.S.). Lab grade acetaldehyde, ACS grade acetone and HPLC grade chloroform were purchased from Fisher Scientific (Hampton, New

Hampshire, U.S.). HPLC-MS grade acetonitrile and HPLC-MS grade water were obtained from Fisher Scientific (Hampton, New Hampshire, U.S.). HPLC grade formic acid was purchased from Sigma Aldrich (St. Louis, Missouri, U.S.).

3.3.2 Methods

Anthocyanin Extraction and Preparation

Fresh blackberries were used as the starting material. Extraction procedures were conducted according to procedures described by Rodriguez-Saona & Wrolstad (2001).

Blackberry anthocyanins are predominantly cyanidin-3-glucoside (C3G) and cyanidin-3- rutinoside (C3R) with small amounts of cyanidin-3-malonylglucoside and cyanidin-3- dioxalyglucoside (Fan-Chiang, & Wrolstad, 2005). The blackberry extract was saponified with 10% KOH for 10 minutes and acidified with 2N HCL and purified by a C18-resin

(Waters Corporation, Milford, Massachusetts, U.S.) using solid phase extraction (SPE)

(Rodriguez-Saona & Worlstad 2001). The sample was washed with two volumes of

0.01% HCl water and three volumes of ethyl acetate. The sample was eluted with methanol. The residual methanol was evaporated with a rotary evaporator (BUCHI

Rotavapor Collegiate Evaporation System, BUCHI Corporation, New Castle, Delaware,

U.S.) under vacuum at 35˚C. The purified pigment was resolubilized in 0.01% HCl water.

The final anthocyanin composition was cyanidin-3-glucoside (93.0%) and cyanidin-3- rutinoside (7.0%), according to results by HPLC monitoring described below.

Anthocyanin and Pyranoanthocyanin Monitoring

A uHPLC (Shimadzu Nexera-I LC-2040C, Columbia, Maryland, U.S.) was used to monitor changes in anthocyanin composition and pyranoanthocyanin formation. A flow rate of 0.4 mL/min was used, and the column oven was set to 60˚C. The injection volume for all samples was 5.0 µL. A Pinnacle DB (Restek Corporation, Bellefonte, PA,

U.S.) C18 column (1.9 µM particle size, 50 x 2.1 mm length) was used. Both the control

36 and treatment samples were ran with the following gradient of Solvent A (4.5% formic acid water) and Solvent B (100% Acetonitrile): 3% B from 0.00 seconds to 1 minute, ramped to 4.3% B by 7.00 minutes, ramped to 40% B by 7.50 minutes, held at 40% B until 8 minutes, decreased to 3% B by 8.50 minutes, and held at 3% B until 10 minutes.

All samples were analyzed by uHPLC during 1 month of storage in the dark. A standard curve was created using cyanidin-3-glucoside (Sigma Aldrich, St. Louis, Missouri, U.S.).

The identities of all peaks were verified using mass spectrometry. The tandem mass spectrometer (LCMS 8040, Shimadzu, Columbia, MD) was coupled with the uHPLC system described above. The nebulizing gas flow was 1.5 L/min, desolvation line temperature was 230˚C, heat block temperature was 200˚C, and the drying gas flow was

15 L/min. A Q1 positive ion scan from 100-1000 m/z was ran to obtain the m/z of intact structures. The secondary collision had an energy of -35.0 eV (argon gas). Precursor ion scans of the aglycones of cyanidin (287 m/z) and of pyranoanthocyanins formed from cyanidin derivatives and pyruvic acid (355 m/z) were also included. (Giusti, Rodriguez-

Saona, Griffin, & Wrolstad, 1999; Farr et al., 2018).

UV-Vis Spectrophotometry and Colorimetry

The absorbance spectra of all samples were collected over a month storage using a micro-well plate reader (SpectraMax 190, Molecular Devices, Sunnyvale, California,

U.S.). Sample aliquots of 50 µL were loaded into a 96-well plate. Samples were evaluated from 380 nm to 720 nm in 1 nm intervals. Distilled water was used as a blank.

Colorimetric date was obtained using a software, ColorBySpectra, that converted the spectral data into colorimetric data (Farr & Giusti, 2017). Color data was obtained

37 with regular transmission, 1964 standard equations, D65 illuminant, and 10˚ observer angle. Color values were reported using CIE-L*a*b* and CIE-L*C*abhab color systems.

Temperature Sample Preparation

The anthocyanin concentration was determined using a standard curve of cyanidin-3-glucoside (Sigma Aldrich, St. Louis, Missouri, U.S.) at an AUC of 500-520 nm. The anthocyanins were diluted in buffer solutions to obtain final concentrations of

500 µM. The buffer was a 0.2 M citrate buffer (citric acid and sodium citrate dihydrate) adjusted to pH 3.0 with HCl and NaOH. A 0.2 M buffer was used to better control the effect temperature had on pH. A stock solution of pyruvic acid was made using pyruvic acid and the citrate buffer. Pyruvic acid was added at a 200X molar ratio of anthocyanin to pyruvic acid (100 mM). 2 mL of all samples were made, and the samples were filtered through a 0.2 µM membrane into 2 mL HPLC vials. Control samples with no pyruvic acid and the treatment samples were stored in the dark for 28 days in 5˚C, 15˚C, and 25˚C incubators. All samples were prepared and ran in triplicate.

pH Sample Preparation

The anthocyanins were diluted with a 0.1 M phosphate buffer to obtain a concentration of 500 µM for all samples. The phosphate buffers were adjusted to pH 1.0,

2.0, 2.5, 3.0, 3.5, 4.0 and 5.0, prior to dilution. A pyruvic acid stock solution was made using pyruvic acid and HPLC water. Pyruvic acid was added to obtain 200X the anthocyanin concentration on a molar ratio (100 mM). The pH values were evaluated with a S220 SevenCompact pH meter (Mettler Toledo, Columbus, OH, U.S.). The pH of the control samples remained within ± 0.05 of the initial pH throughout the study. When

38 pyruvic acid was added to the samples, the pH of the solutions shifted, and treatment samples had pH values of 1.0, 1.8, 2.0, 2.3, 2.6, 2.8, 3.1, 3.5, 3.6, 3.8, 4.1 and 5.0. All pH values varied by less than 0.05 throughout the study. All samples were filtered through a

0.2 µM membrane into 2 mL HPLC vials and stored in the dark for 30 days at 25˚C. All samples were prepared and ran in triplicate.

Equations Used

Various equations were used to determine the percent pyranoanthocyanin formation yields in relation to day 0 and the percent of total pigment remaining.

Eq 3.1.

퐶푎푙푐푢푙푎푡𝑖표푛 표푓 푃푒푟푐푒푛푡 푃푦푟푎푛표푎푛푡ℎ표푐푦푎푛𝑖푛 퐹표푟푚푎푡𝑖표푛 푌𝑖푒푙푑

(AUC C3G PACN + C3R PACN t ) = 500−520nm n ∗ 100 (AUC500−520nm C3G + C3R at Day 0)

Eq 3.2.

퐶푎푙푐푢푙푎푡𝑖표푛 표푓 푃푒푟푐푒푛푡 표푓 푃𝑖푔푚푒푛푡 푅푒푚푎𝑖푛𝑖푛푔

(AUC C3G + C3R + C3G PACN + C3R PACN at t ) = 500−520nm n ∗ 100 (AUC500−520nm C3G + C3R + C3G PACN + C3R PACN at t0)

AUC=Area Under the Curve, C3G=cyanidin-3-glucoside, C3R=cyanidin-3-rutinoside,

PACN=Pyranoanthocyanins formed from the anthocyanin and pyruvic acid

Statistical Analysis

The data was analyzed to determine the statistical significance of the research findings. Figures and tables were created with Microsoft Office 2019 Excel (Redmond,

WA, U.S.). Microsoft Excel was used to find averages and standard deviations. One-way

ANOVAs (2-tailed, α = 0.05) were ran to compare the individual samples to each other

39 on specific days for HPLC, spectra, and color data. Two-way ANOVAs (2-tailed, α =

0.05) were run to compare a single sample across multiple days for HPLC, spectra, and color data. Tukey Post-Hoc tests were used for individual comparisons. Minitab

(Software Version 18) was used for all statistical analysis (State College, PA, U.S.).

3.4 Results and Discussions

3.4.1 Effect of pH on Pyranoanthocyanin Formation

Pyranoanthocyanin Formation

Pyranoanthocyanin formation yields were calculated in relation to the anthocyanin starting concentration on day 0. As shown in Figure 3.2 pyranoanthocyanins were formed at all pH values (data obtained using Eq 3.1), but at different yields. On day

7, a significantly greater pyranoanthocyanin formation yield (18.3%) was observed at pH

3.1 than any other pH value. On day 14, samples at pH 3.1 had the numerically largest pyranoanthocyanin formation yield at 23.0%, but this yield was not significantly larger than those of samples at pH 2.8 and pH 3.5. On day 21, samples at pH 2.8 (24.3%), 3.1

(23.9%) and 3.5 (23.2%) had the highest pyranoanthocyanin formation yields and were not significantly different from each other. Additionally, on day 30 pH 2.8 (23.9%), 3.1

(23.3%) and 3.5 (22.6%) were not significantly different from each other and were not significantly different from day 21 yields. On day 30 pH 2.6 reached similar yields to pH

2.8, 3.1 and 3.5 at 23.0%. It appeared that a maximum yield of ~24% under the conditions of this study, as suggested by lack of significant difference between day 21 and 30. Reduced reaction time is preferable to obtain more pyranoanthocyanins in less time and to avoid anthocyanin and pyranoanthocyanin degradation and polymerization

40 further decreasing pyranoanthocyanin yields (Rein et al., 2005). Importantly, the pyranoanthocyanin formation yields in this study were larger and formed faster than previously recorded yields. For example, one study reported similar yields at 18.48%

Vitisin A formation, but incubation took 232 days (Romero & Bakker, 2000) and another reported only 2.03% Vitisin A formation over 10 weeks (Morata et al., 2007).

Figure 3.2. Pyranoanthocyanin formation yields (Eq 3.1) at pH values between 1.0 and 5.0 for day 3, 7, 14, 21 and 30. Incubated at 25°C in the dark. Points are the averages (n=3) and the error bars are the associated standard deviations. In the first 7 days of the experiment pH 3.1 was the pH of maximum pyranoanthocyanin formation. Between days 14 – 21, formation began to slow and maximum yields were observed between pH 2.8 and pH 3.5 (Figure 3.2). By day 30 maximum yields were observed between pH 2.6 and pH 3.5. The reported pKH of cyanidin-3-glucoside is around 3.01, and the pKH of blackberry has been reported as 3.07

(Stintzing, Stintzing, Carle, Frei, & Wrolstad, 2002), and both are near the maximum of 41 pyranoanthocyanin formation at pH of 3.1. Pyranoanthocyanins form by nucleophilic substitution at the C-4 position of the anthocyanin molecule (Quaglieri et al., 2017). The anthocyanin flavylium cation has electrophilic character and the vinly group of the cofactor (pyruvic acid in the enol form) has nucleophilic character allowing for the formation of pyranoanthocyanins (Dangles & Fenger, 2018). Pyranoanthocyanins likely form most efficiently when anthocyanins are predominantly in the flavylium cation form, because of the electrophilic character of the anthocyanin equilibrium state (Dangles &

Fenger, 2018; Quaglieri, et al., 2017). However, if the only factor affecting formation was the need for the flavylium cation then optimum formation would have been near pH

1.0. This indicated pyruvic acid form and stability likely influenced formation. Pyruvic acid is known to undergo acid polymerization. As the pH decreases polymerization increases, so at low pH levels pyruvic acid polymerizes and becomes unavailable to react with the anthocyanin (Müller & Baumberger, 1939; Hazen, & Deamer, 2007). In addition, at more acidic pH values the keto form is favored over the enol form, so larger pH values encourage the enol equilibrium form (Müller & Baumberger, 1939). Therefore, it was predicted that the optimum pH in this study depended on more than one factor. The medium needed be acidic enough for the flavylium cation to be present in high quantities, but basic enough to prevent acid-catalyzed polymerization of pyruvic acid and basic enough to promote enolization.

In the first 7 days of the experiment it was clear that pyranoanthocyanins were forming most rapidly at pH 3.1, indicating pH 3.1 was likely the preferred pH of formation in this study. As the study continued the non-significant difference for

42 pyranoanthocyanin formation yields (Eq. 3.1) for pH 2.8, 3.1 and 3.5 showed surrounding pH values produced similar pyranoanthocyanin yields but at a slower rate of formation.

Between days 14 and 21, there was a maximum formation between pH 2.8 and 3.5, and this pH range was slightly more acidic than traditional wine pH (pH 3.7 - 4.0) at which most pyranoanthocyanin formation studies have been carried out (Somers, 1971; Morata et al., 2007). Between pH 2.8 and 3.5, the flavylium cation was likely at higher concentrations in solution and near the pKH of the anthocyanins. As the pH increased above 3.5 the colorless, neutral, hemiketal form began to predominate. As pH increased the anthocyanin molecules became more unstable, and began degrading, decreasing the number of anthocyanins present for the pyruvic acid to react with (Figure 3.3).

Spectrophotometric and Colorimetric Properties

Samples showed a hypsochromic shift in the lambda maximum (λmax) excluding pH 1.0 (significant λmax increase) and 1.8 (not significant λmax increase), Figure 3.4. As shown in Figure 3.4 the control sample at pH 2.0 followed a similar trend to treatment samples at pH 1.0 and 1.8 likely indicating the bathochromic shift at pH 1.0 and 1.8 occurred from anthocyanin reactions. Anthocyanins can undergo self-association and polymerization potentially leading to the bathochromic shifts (Brownmiller, Howard, &

Prior, 2008; Fernandes, Brás, Mateus, & De Freitas, 2015). The degree of the hypsochromic shifts for the other samples (pH 2.0, 2.3, 2.5, 2.8, 3.1, 3.5, 3.7, 3.8, 4.1,

5.0) was likely attributed to pyranoanthocyanin formation and pigment degradation.

As shown in Figure 3.5 C pyranoanthocyanins are hypsochromic shifted in the lambda maximum compared to their anthocyanin counter parts (De Freitas & Mateus,

2011; Oliveira et al., 2006). However, even with lesser amounts of pyranoanthocyanins formed at high pH values large hypsochromic shifts were still identified. Figure 3.3 showed as pH increased a greater amount of pigment was degraded, so the large hypsochromic shifts at high pH values were likely attributed to pigment degradation.

Shifts in the lambda maximum (λmax) between day 0 and day 30 for samples with high pyranoanthocyanin formation, pH 2.8 (Δ8nm), 3.1 (Δ10nm) and 3.5 (Δ13 nm), can likely be attributed to both pyranoanthocyanin formation and pigment degradation. Samples at all pH values had a decrease in absorbance at the λmax, consistent with loss of chromophores. Figure 3.4 shows the pH 3.5 control sample had a significantly lower absorbance than the pH 3.5 treatment sample indicating the pyranoanthocyanins that formed may have helped protect the samples pigments.

Figure 3.5. (A) Chromatogram on day 0 at pH 3.0 and 25°C (B) Chromatogram on day 28 at pH 3.0 and 25°C (C) Spectra from 280nm to 800nm obtained from the uHPLC. The solid line is cyanidin-3-glucoside (C3G) spectra and the dashed line is the spectra of the pyranoanthocyanins (PACNs) derived from cyanidin-3-glucoside (C3G) and cyanidin-3- rutinoside (C3R). The color properties of all samples, on day 0 and day 30, are shown in Table 3.1.

All samples showed an increase in lightness besides those at pH 1.0 and pH 5.0. At pH

1.0 anthocyanins are relatively stable; as shown in Figure 3.3, sample at pH 1.0 had significantly more pigment remaining than any other sample which may have helped stabilize the L* value. At pH 5.0 the sample had a large L* value on day 0 and day 30, because the colorless hemiketal form of the anthocyanin predominates at pH 5.0. All samples showed a decrease in a*, meaning the color lost some redness. The b* decreased for all samples with pH values bellow 3.1 indicating samples became less yellow and with pH values greater than 3.1 the b* increased indicating samples became more yellow.

The samples started out with a vibrant red color. As pyranoanthocyanins formed the samples became more orange in color, and as pigments degraded the samples became

46 browner. The changes in a* and b* were likely attributed to both pyranoanthocyanin formation and pigment degradation (Rein et al., 2005; Wrolstad et al., 2005).

The chroma (C*ab) decreased between day 0 and day 30 for samples at all pH values excluding pH 5.0. At pH 5.0 the colorless hemiketal predominated explaining the 47 low initial chroma reading, and the formation of degradation products throughout the study may have increased the chroma. For other samples, chromophores were lost as pigments degraded, and the chroma decreased (He et al., 2012). The hue angle (hab) describes the shade of the color; for example, a hab of 0˚ is a red color and a hab of 90˚ is a yellow color. At pH 1.0, which had a pyranoanthocyanin formation yield of 6.3%, the hue angle decreased from 24.1° on day 0 to 18.0° on day 30. At pH 3.1, which had a pyranoanthocyanin formation yield of 23.9%, the hab increased from 19.3° (day 0) to

30.0° (day 30). Pyranoanthocyanins are more orange in color (De Freitas & Mateus,

2011; Oliveira et al., 2006) which likely attributed to the increase in hab (approaching

90°) for samples with large pyranoanthocyanin formation yields. Additionally, as anthocyanins degrade the color changes from a red to a brown/yellow color. Therefore, the degradation of the anthocyanins could also have contributed to the shift in hue angle

(Wrolstad, Durst, & Lee, 2005).

3.4.2 Effect of Temperature on Pyranoanthocyanin Formation

Pyranoanthocyanin Formation

At all storage temperatures, pyranoanthocyanins were formed; however, there was a significant difference between pyranoanthocyanin formation yields for 5˚C, 15˚C, and

25˚C at each time point. On day 28, storage at 25˚C yielded the highest pyranoanthocyanin formation at 25.4%, followed by 15°C at 23.7% and 5°C at 10.0% formation (Figure 3.6 A). It was expected that as the temperature increased pyranoanthocyanin formation would increase. Collision theory explains that reactant partners must collide with a minimum energy (activation energy) to form a product. As

48 the temperature of the system increases, molecules gain energy, increasing the number of collisions thus increasing the reaction rate (Brown et al., 2012). However, increased temperature also increased anthocyanin degradation (Eiro & Heinonen, 2002; Cabrita et al., 2000) as shown in Figure 3.6 B. At 25°C 32.1% of the total pigments remained (Eq.

3.2) on day 28 and at 15°C 50.0% of the total pigments remained on day 28. Although storage at 25°C yielded significantly more pyranoanthocyanin formation at all time points, it also led to significantly more pigment degradation at all time points.

Figure 3.6 (A) Percentage of pyranoanthocyanin formation yield (Eq. 3.1) and (B) percentage of the total pigment remaining (Eq 3.2) in relation to day 0 at 5°C, 15°C and 25°C in pH 3.0 buffer. Information obtained from chromatograms at 500-520nm. All points are averages (n=3) and error bars are the associated standard deviations.

There was no significant difference between pyranoanthocyanin formation yields

(Eq. 3.1) on day 21 and day 28 when incubated at 25°C. However, a significant difference existed between the percentage of pigment remaining (Eq. 3.2) between day 21 and day 28 for 25ºC. To efficiently form a compound, high yields should be obtained in the shortest amount of time. Therefore, 21 days was the preferred incubation time at 25°C under the conditions of this study because pyranoanthocyanin formation was the same at

28 days but pigment degradation was lower on day 21.

Spectrophotometric and Colorimetric Properties

Spectral properties were recorded at each time point. As shown in Figure 3.7 all samples had a decrease in the lambda maximum (λmax) and a decrease in absorbance. The decrease in λmax and absorbance were not significant at 5°C, because the low temperature likely helped stabilize the pigment. Pyranoanthocyanins had a hypsochromic shift in the

λmax in respect to the anthocyanin counterpart (Figure 3.5C), aligning with literature (Farr et al., 2018). Therefore, the hypsochromic shift in the λmax between day 0 and day 28 was partially attributed to pyranoanthocyanin formation. In addition, pigments were degrading likely contributing to the hypsochromic shift. Chromophores were lost as pigments degraded describing the decrease in absorbance over the 28 days.

Color analyses were conducted and recorded at each time point. The color properties in Table 3.2 followed similar trends to the color properties in Table 3.1. L* increased, a* decreased, C*ab decreased and hab increased. There was no significant difference in b* between day 0 and day 28 at any temperature. These variations can be described by both pyranoanthocyanin formation and pigment degradation, because pyranoanthocyanins are more orange in color compared to their anthocyanin counterpart and degraded pigments are more brown in color (Rein et al., 2005; Wrolstad et al., 2005).

Table 3.2. The color properties, based on CIE – L*a*b* and CIE – L*C*abhab color systems, on day 0 and day 28 for 5°C, 15°C and 25°C. Stored in the dark at pH 3.0. Represented as the average (n=3) and the associated standard deviation in parenthesis.

3.5 Conclusion

This study focused on increasing pyranoanthocyanin yields by investigating effects of pH and incubation temperature. pH 3.1, which was near the pKH, had the highest pyranoanthocyanin formation yields on day 7. Day 21 had an optimum pH range, pH 2.8 to 3.5, for pyranoanthocyanin formation yields (yields ~24%), and day 21 was preferred over day 30. It was hypothesized that the pH of greatest formation depended on the pH being acidic enough for the flavylium cation form to predominated, but basic enough to discourage pyruvic acid polymerization and encourage enol formation.

Therefore, the pH range of greatest formation was more acidic than typical wine pH but not too acidic. 25°C had the highest pyranoanthocyanin formation yields by day 21

(yields ~25%). As temperature decreased pyranoanthocyanin formation took more time and yields decreased, because the energy in the system decreased.

Chapter 4. Comparison of ratios of anthocyanin to cofactor that lead to higher formation of pyranoanthocyanins from cyanidin-3-glucoside and cyanidin-3- rutinoside

4.1 Abstract

Pyranoanthocyanins are gaining attention as potential food colorants because they are more stable than their anthocyanin precursors. Anthocyanins react over time with cofactors to form pyranoanthocyanins, but yields are low limiting pyranoanthocyanin application. The objective of this study was to determine the efficiency of different concentrations of cofactors on pyranoanthocyanin formation. Cyanidin-3-glucoside and cyanidin-3-rutinoside (150 µM) were mixed with pH 3.0 buffer and caffeic acid (CA) at

1:1, 1:10 and 1:30 or pyruvic acid (PA) at 1:10, 1:30, 1:100 and 1:200 on an anthocyanin

: cofactor molar ratio. Samples were stored at 25°C in the dark for 42 days. HPLC with

PDA detector was used to monitor anthocyanin, pyranoanthocyanin, and pyruvic and caffeic acid content. A multi-well plate reader was used to monitor spectral properties during storage. Caffeic acid formed pyranoanthocyanins more efficiently than pyruvic acid. In comparison to day 0, CA1:30 had a pyranoanthocyanin formation yield of 39.1% and PA1:200 had a pyranoanthocyanin formation yield of 8.0% on Day 42. Of the total pigments on day 42 CA1:30 was 73.9% pyranoanthocyanins and PA1:200 was 20.4% pyranoanthocyanins. After storage, there was a hypsochromic shift for PA1:200 (2 nm),

CA1:10 (2 nm), and CA1:30 (7 nm). The cofactor concertation decreased between 2.0%

(CA1:1) and 24.7% (PA1:100) from day 0 to day 42.

4.2 Introduction

The increasing demand for naturally derived colorants has led to the investigation and use of anthocyanins as alternatives to the synthetic lakes and dyes typically used. A major limitation of anthocyanins as food colorants is the limited stability of anthocyanins.

Anthocyanins are subject to degradation and color changes by pH, temperature, light, enzymes, oxygen, sulfites, ascorbic acid, hydrogen peroxide, and high-water activities

(Cabrita et al., 2000; Eiro & Heinonen, 2002; Wrolstad et al., 2005; Farr & Giusti, 2018).

To help stabilize anthocyanins, co-pigmentation and metal complexation have been studied, and more recently anthocyanin derivatives have been gaining attention as potential stable naturally derived colorants (Quaglieri et al., 2017; Farr & Giusti, 2018).

One such type of derivatives includes pyranoanthocyanins which were only recently identified (Fulcrand et al., 1997; Bakker et al., 1997; Bakker, & Timberlake,

1997; Oliveira et al., 2014). The structures of pyranoanthocyanins include an additional ring between carbon 4 and the C5-hydroxyl group of the anthocyanin, Figure 4.1. The additional ring (D ring) of the pyranoanthocyanin likely helps stabilize the pigment over a wider pH range and prevents the nucleophilic addition of water to the anthocyanin (He et al., 2012). The new D ring also protects the chromophore against sulfite, ascorbic acid, and hydrogen peroxide bleaching by blocking the preferred position of attack at carbon 4

(Oliveira et al., 2006; Farr & Giusti, 2018; Wrolstad et al., 2005). The enhanced stability of the pyranoanthocyanin has grabbed the attention of many researchers.

Pyranoanthocyanins were first found in wine systems (Somers, 1971; Fulcrand et al.,

1997; Bakker et al., 1997; Bakker & Timberlake, 1997), so many studies have been conducted in wine systems. Under these conditions pyranoanthocyanins typically form slowly, limiting the application of pyranoanthocyanins as colorants.

Figure 4.1. Formation of a carboxypyranoanthocyanin, 5-carboxypyranocyanidin-3- glucoside, by the reaction of the anthocyanin, cyanidin-3-glucoside, and pyruvic acid (Modified from Farr & Giusti, 2018).

Pyranoanthocyanins form by the cycloaddition of a cofactor with a vinyl group to an anthocyanin molecule (Quaglieri et al., 2017; De Freitas & Mateus, 2011).

Hydroxycinnamic acids, vinylphenols, vinylflavanols, pyruvic acid, acetaldehyde, and acetone are all viable cofactors (Rentzsch et al., 2007). Studies have used yeast strands to create the necessary cofactors to form pyranoanthocyanins, but formation takes time and yields are typically low. For example, hydroxycinnamate decarboxylase yeast,

Saccharomyces cerevisiae and Pichia guillermondii, were used to create 4-vinylphenol from coumaric acid to form vinylphenolic pyranoanthocyanins (Benito, Morata,

Palomero, González, & Suárez-Lepe, 2011). Total formation of vinylphenolic 55 pyranoanthocyanins was around 6% after 12 days in comparison to total anthocyanin content on day 0 (Benito et al., 2011).

It was thought that anthocyanin-vinylphenol adducts (Pinotins) were only formed when a cinnamic acid, such as p-coumaric, sinapic, ferulic, or caffeic acid, was decarboxylated during the fermentation process to form the respective 4-vinylphenol which reacted with the anthocyanin. However, recent research has suggested that anthocyanins can react with hydroxycinnamic acids to form pyranoanthocyanins if the hydroxycinnamic acids have electron-donating substituents (Schwarz et al., 2003). The reaction of 4-vinylphenols and anthocyanins in wines takes years and are typically only found in wines that have been aged for over 2.5 years (Schwarz et al., 2003; He et al.,

2012). However, in model wine systems it has been shown that malvidin-3-glucoside and hydroxycinnamic acids can react quickly (Fulcrand et al., 1996; Schwarz et al., 2003).

Enolizable cofactors can be used to form pyranoanthocyanins. However, formation is generally at low rates. The addition of pyruvic acid to wine systems lead to

2.03% pyranoanthocyanin formation and acetaldehyde lead to 1.35% pyranoanthocyanin formation after 10 weeks of incubation (Morata et al., 2007). With low formation rates it is not economically feasible to consider use of pyranoanthocyanins as food colorants. It was hypothesized that the type of cofactor and the concentration of that cofactor would affect pyranoanthocyanin formation. The objective of this study was to determine the efficiency and the effect of caffeic acid and pyruvic acid on pyranoanthocyanin formation from cyanidin-3-glucoside and cyanidin-3-rutinoside.

4.3 Materials and Methods

4.3.1 Materials

Blackberries (SUNBELLE, Los Reyes, Mexico) of the Rubus sp. were purchased from a local grocery store in Columbus, OH, U.S.. Lab grade pyruvic acid, lab grade caffeic acid and HPLC grade formic acid were purchased from Sigma Aldrich (St. Louis,

Missouri, U.S.). Lab grade acetaldehyde, ACS grade acetone and HPLC grade chloroform were purchased from Fisher Scientific (Hampton, New Hampshire, U.S.).

HPLC-MS grade acetonitrile and HPLC-MS grade water were obtained from Fisher

Scientific (Hampton, New Hampshire, U.S.).

4.3.2 Methods

Anthocyanin Extraction and Preparation

Anthocyanins were extracted from fresh blackberries following procedures according to Rodriguez-Saona & Worlstad (2001). The anthocyanin profile of blackberry is predominantly cyanidin-3-glucoside (C3G) and cyanidin-3-rutinoside (C3R) with small amounts of cyanidin-3-malonylglucoside and cyanidin-3-dioxalyglucoside (Fan-Chiang,

& Wrolstad, 2005). The blackberry extract was saponified to remove the acylation by treatment with 10% KOH for 10 minutes followed by acidification with 2N HCL. After saponification the anthocyanin material underwent Solid Phase Extraction (SPE)

(Rodriguez-Saona & Worlstad, 2001) with a C18 resin (Waters Corporation, Milford,

Massachusetts, U.S.). The sample was washed with two volumes of 0.01% HCl water and three volumes of ethyl acetate and eluted with 0.01% HCl methanol. The residual methanol was evaporated with a rotary evaporator (BUCHI Rotavapor Collegiate

Evaporation System, BUCHI Corporation, New Castle, Delaware, U.S.) under vacuum at

35˚C. The purified pigment was resolubilized in 0.01% HCl water. The final anthocyanin composition was cyanidin-3-glucoside (92.8%) and cyanidin-3-rutinoside (7.2%).

Anthocyanin and Pyranoanthocyanin Monitoring

A uHPLC (Shimadzu Nexera-i LC-2040C, Columbia, Maryland, U.S.) was used to monitor changes in anthocyanin composition and formation of pyranoanthocyanins. A flow rate of 0.4 mL/min was used, and the column oven was set to 60˚C. A Pinnacle DB

(Restek Corporation, Bellefonte, PA, U.S.) C18 column (1.9 µM particle size, 50 x 2.1 mm length) was used. The injection volume was 7 µL. Different solvent gradients were used depending on the cofactor used.

For the samples with pyruvic acid the following gradient of solvent A (4.5% formic acid water) and solvent B (100% acetonitrile) was used: 0-3% B in 3 minutes, 3-

4.3% B from 3-10 minutes, 4.3-40% B from 10-11 minutes, 40% B from 11-12 minutes,

40-0% B by 12.5 minutes, and held at 0% B until 14 minutes. A standard curve for cyanidin-3-glucoside (Sigma Aldrich, St. Louis, Missouri, U.S.) was created using the methodology above.

For the samples with caffeic acid the following gradient of solvent A (4.5% formic acid water) and solvent B (100% acetonitrile) was used: 0-21% B from 0-14 minutes, 21-40% B by 14.5 minutes, 40% B until 16 minutes, 40-0% B by 16.5 minutes, and held at 0% B until 18.5 minutes. A standard curve for anthocyanin quantification, with cyanidin-3-glucoside (Sigma Aldrich, St. Louis, Missouri, U.S.), was created using the methodology above.

The identity of all peaks were verified using mass spectrometry. The mass spectrometry system (LCMS 8040, Shimadzu, Columbia, MD, U.S.) was coupled with the uHPLC system described above. The nebulizing gas flow was 1.5 L/min, desolvation line temperature was 230˚C, heat block temperature was 200˚C, and the drying gas flow was 15 L/min. A Q1 positive ion scan from 100-1000 m/z was ran to get the intact structure m/z. The secondary collision had an energy of -35.0 eV (argon gas). Parent compounds were monitored using precursor ion scanning for m/z of 287 for the cyanidin aglycone, m/z of 355 for the pyranoanthocyanin aglycone from pyruvic acid

(carboxypyranocyanidin), and m/z of 419 for the pyranoanthocyanin aglycone from caffeic acid (pyranocyanidin-catechol) (Giusti et al., 1999; Farr et al., 2018; Vallverdú-

Queralt et al., 2016).

Pyruvic Acid and Caffeic Acid Monitoring

Pyruvic acid concentration was monitored by reverse phase HPLC (Shimadzu,

Maryland, U.S.) with a PDA detector. The HPLC instrument was comprised of LC-20AD pumps, CMB-20A communication system, SIL-20A HT autosampler, CTO-20A column oven, and SPD-M20A Photodiode Array Detector (Shimadzu, Maryland, U.S.). A Prevail

Organic Acid Column was used with a 5 µm particle size and 150 x 4.6 mm length

(Hichrom, Berkshire, U.K.). The mobile phase was HPLC water acidified to pH 2.2 with sulfuric acid at a flow rate of 0.8 mL/min for 30 minutes. Pyruvic acid was monitored at

220 nm. The pyruvic acid concentration in PA1:1 was not monitored due to the detection limit of the instrument. A standard curve for pyruvic acid (Sigma Aldrich, St. Louis,

Missouri, U.S.) was created with the methodology above.

Caffeic acid concentration in samples was monitored with the uHPLC described above. The oven was set to 60°C and the flow rate was 0.4 mL/min. The following gradient of solvent A (4.5% formic acid water) and solvent B (100% acetonitrile) was used: 0% B at 0.00 seconds, ramped to 4.5% B by 3 minutes, ramped to 40% B by 5 minutes, held at 40% B until 7 minutes, decreased to 0% B by 9 minutes, held at 0% B until 13 minutes. The chromatograms were monitored at 320 nm for caffeic acid quantification. The caffeic acid concentration was monitored using a caffeic acid standard curve created with the methodology above.

The organic acid concentration, anthocyanin profile, and pyranoanthocyanin formation were monitored on day 0, 3, 7, 14, 21, 28, 35 and 42.

UV-vis Spectrophotometry and Colorimetry

The absorbance spectra of all samples were collected on Day 0, 3, 7, 14, 21, 28,

35 and 42 using a multi-well plate reader - accuMultiskan GO 1510-04369C (Fisher

Scientific, Waltham, Maryland, U.S.). Sample aliquots of 110 µL were loaded onto a 96- well plate. Samples were evaluated from 380 nm to 720 nm in 1 nm intervals. Distilled water was used as a blank.

Colorimetric data were obtained by converting the spectral data into colorimetric data using the software, ColorBySpectra (Farr & Giusti, 2017). Color data was obtained using the 1964 CIE equations, D65 illuminant spectral distribution, and 10° observer angle functions. Color values were reported using CIE-L*a*b* and CIE-L*C*abhab color communication systems.

Pre-Screening

Anthocyanin concentration was determined using a standard curve of cyanidin-3- glucoside (Sigma Aldrich, St. Louis, Missouri, U.S.) at an AUC of 500-520nm.

Saponified blackberry anthocyanins, cyanidin-3-glucoside and cyanidin-3-rutinsode, were brought to a concentration of 500 µM using a 0.1 M citrate buffer adjusted to pH

3.0 with HCl and NaOH. Potassium sorbate (0.1% w/v) and sodium benzoate (0.1% w/v) were added to the citrate buffer to function as preservatives. Pyruvic acid, acetaldehyde, and acetone were added to obtain a 200X molar ratio of anthocyanin : cofactor (100 mM). Control samples contained anthocyanin and buffer but no cofactor. All samples were filtered through a 0.2 µm membrane into 2 mL HPLC vials. HPLC vials were stored in a 15°C incubator for 28 days. All samples were created and ran in triplicate

The uHPLC described above was used to monitor anthocyanin and pyranoanthocyanin content. The oven was set to 60°C, a flow rate of 0.4 mL/min was used, and the injection volume was 5 µL. A Pinnacle DB (Restek Corporation,

Bellefonte, PA, U.S.) C18 column (1.9 µM particle size, 50 x 2.1 mm length) was used.

The following gradient of solvent A (4.5% formic acid water) and solvent B (100% acetonitrile) was used: held at 0%B from 0.00 min to 1.00 minute, ramped to 4.3% B by

7.00 minutes, ramped to 40% B by 7.50 minutes, held at 40% B until 8.00 minutes, decreased to 3% B by 8.50 minutes, held at 3% B until 10.00 minutes. Readings were taken of all samples on day 0, 3, 7, 14, 21 and 28.

A SpectraMax 190 micro-well plate reader (Molecular Devices, Sunnyvale,

California, U.S.) was used to collect spectral and absorbance data. Sample aliquots of 50

µL were loaded onto a 96-well plate. Samples were evaluated from 380 nm to 720 nm in

1 nm intervals. Distilled water was used as a blank. Colorimetric date was obtained using a software, ColorBySpectra, that converted the spectral data into color data (Farr &

Giusti, 2017). Color data was obtained using the 1964 CIE equations, D65 illuminant spectral distribution, and 10° observer angle functions. Color values were reported using

CIE-L*a*b* and CIE-L*C*abhab color communication systems. Readings were taken of all samples on day 0, 3, 7, 14, 21 and 28.

Sample Preparation

The anthocyanin concentration was determined using a standard curve of cyanidin-3-glucoside (Sigma Aldrich, St. Louis, Missouri, U.S.) at an AUC of 500-

520nm. The anthocyanins were then diluted to obtain a 150 µM concentration (cyanidin-

3-glucoside equivalents) using a 0.1 M citrate buffer (citric acid and sodium citrate dihydrate) adjust to pH 3.0 with HCl and NaOH. Control samples were prepared that contained anthocyanins in buffer with no cofactor. Caffeic acid and pyruvic acid were dissolved in citrate buffer to form stock solutions. Buffer, caffeic acid stock, and anthocyanins were combined to form ratios of 1:1 (CA1:1), 1:10 (CA1:10) and 1:30

(CA1:30) of anthocyanin : caffeic acid molar ratios. Similarly, buffer, pyruvic acid stock, and anthocyanins were combined to form ratios of 1:1 (PA1:1), 1:30 (PA1:30), 1:100

(PA1:100) and 1:200 (PA1:200) of anthocyanin : pyruvic acid molar ratios. All samples were filtered through a 0.2 µm membrane into 2 mL HPLC vials. Vials were stored in a

25˚C incubator (Fisher Scientific, Waltham, Maryland, U.S.) for the duration of the study. All samples were prepared and analyzed in triplicate.

Equations Used

Various equations were used to determine pyranoanthocyanin yields in relation to day 0, percent pyranoanthocyanin pigment in comparison to total pigment, and total pigment degradation.

Eq 4.1.

퐶푎푙푐푢푙푎푡𝑖표푛 표푓 푃푒푟푐푒푛푡 푃푦푟푎푛표푎푛푡ℎ표푐푦푎푛𝑖푛 퐹표푟푚푎푡𝑖표푛 푌𝑖푒푙푑

(AUC C3G PACN + C3R PACN t ) = 500−520nm n ∗ 100 (AUC500−520nm C3G + C3R at Day 0)

Eq 4.2.

퐶푎푙푐푢푙푎푡𝑖표푛 표푓 푃푒푟푐푒푛푡 푃푦푟푎푛표푎푛푡ℎ표푐푦푎푛𝑖푛 퐹표푟푚푎푡𝑖표푛 푡표 푇표푡푎푙 푃𝑖푔푚푒푛푡

(AUC C3G PACN + C3R PACN at t ) = 500−520nm n ∗ 100 (AUC500−520nm C3G + C3R + C3G PACN + C3R PACN at tn)

Eq 4.3.

퐶푎푙푐푢푙푎푡𝑖표푛 표푓 푇표푡푎푙 푃푒푟푐푒푛푡 표푓 푃𝑖푔푚푒푛푡 푅푒푚푎𝑖푛𝑖푛푔

(AUC C3G + C3R + C3G PACN + C3R PACN at t ) = 500−520nm n ∗ 100 (AUC500−520nm C3G + C3R + C3G PACN + C3R PACN at t0)

Eq 4.4

퐶푎푙푐푢푙푎푡𝑖표푛 표푓 푃푒푟푐푒푛푡 푃푦푟푢푣𝑖푐 퐴푐𝑖푑 푅푒푚푎𝑖푛𝑖푛푔

(AUC Pyruvic Acid at t ) = 220nm n ∗ 100 (AUC220nmPyruvic Acid at t0)

Eq 4.5

퐶푎푙푐푢푙푎푡𝑖표푛 표푓 푃푒푟푐푒푛푡 퐶푎푓푓푒𝑖푐 퐴푐𝑖푑 푅푒푚푎𝑖푛𝑖푛푔

(AUC Caffeic Acid at t ) = 320nm n ∗ 100 (AUC320nmCaffeic Acid at t0)

AUC=Area Under the Curve, C3G=cyanidin-3-glucoside, C3R=cyanidin-3-rutinoside,

PACN=Pyranoanthocyanins from C3G and C3R.

Statistical Analysis

The data was analyzed to determine the statistical significance of the research findings. Figures and tables were created with Microsoft Office 2019 Excel (Redmond,

WA, U.S.). Microsoft Excel was used to find averages and standard deviations. One-way

ANOVAs were run to compare the individual samples to each other on specific days for

HPLC, spectra, and color data (2-tailed, α=0.05). Two-way ANOVAs were run to compare a single sample across multiple days for HPLC, spectra, and color data (2-tailed,

α=0.05). Tukey Post-Hoc tests were used for individual comparisons. Minitab 18 was used for all ANOVA analysis (State College, PA, U.S.).

4.4 Results & Discussion

4.4.1 Cofactor Pre-Screening

A preliminary study was ran using 500 µM anthocyanins and 200X the cofactor on an anthocyanin : cofactor molar ratio. Acetaldehyde, pyruvic acid, and acetone were used as cofactors and saponified blackberry was the anthocyanin source. No pyranoanthocyanins were found after 28 days when acetone was used as the cofactor, although other studies have reported small amounts of methyl-pyranoanthocyanin formation when using a smaller dose of acetone (De Freitas & Mateus, 2011; Lu & Foo,

2001). When acetaldehyde was used as a cofactor no identifiable monomeric- pyranoanthocyanins were found; however, it was predicted that the high dosage of acetaldehyde induced rapid polymerization of anthocyanins and pyranoanthocyanins that

64 may have formed as reported in some previous works (Morata et al., 2007).

Pyranoanthocyanins did form when pyruvic acid was used as the cofactor. Previous studies performed in our laboratory showed promising results of caffeic acid as a potential cofactor (Zhu and Giusti, 2018). The current study used various concentrations of pyruvic and caffeic acid to increase pyranoanthocyanin formation.

4.4.2 Pyranoanthocyanin Formation

Pyranoanthocyanin formation was monitored using HPLC (Figure 4.2).

Pyranoanthocyanin yields throughout this paper were expressed as pyranoanthocyanins derived from cyanidin-3-glucoside and cyanidin-3-rutinsodie, because the pyranoanthocyanin peaks for cyanidin-3-glucoside and cyanidin-3-rutinoside coeluted.

Mass spectroscopy was used to identify the coeluting peaks. When pyruvic acid was used as a cofactor the pyranoanthocyanins derived from cyanidin-3-glucoside had a MS/MS transition of 517 m/z to 355 m/z and cyanidin-3-rutinoside had a MS/MS transition of

663 m/z to 355 m/z (Farr et al., 2018). When caffeic acid was used as a cofactor the pyranoanthocyanins derived from cyanidin-3-glucoside had a mass transition of 581 m/z to 419 m/z and cyanidin-3-rutinoside had a MS/MS transition of 727 m/z to 419 m/z

(Vallverdú-Queralt et al., 2016). Based off the starting proportions and intensity data obtained from the MS/MS it was estimated that roughly 90% of the pyranoanthocyanins were derived from cyanidin-3-glucoside and 10% were derived from cyanidin-3- rutinoside.

Figure 4.2. Pyranoanthocyanin formation and anthocyanin content were monitored with HPLC. Chromatograms from day 0 and day 42 for CA1:30 and PA1:200 are shown. C3G = cyanidin-3-glucoside, C3R = cyanidin-3-rutinoside, PACN = Pyranoanthocyanins derived from cyanidin-3-glucoside and cyanidin-3-rutinoside

All samples formed pyranoanthocyanins over a 42 day time period (Figure 4.3).

On day 42 caffeic acid had 1.3% (CA1:1), 14.2% (CA1:10) and 39.1% (CA1:30) pyranoanthocyanin formation yields (Eq 4.1). On day 42 pyruvic acid had 0.1% (PA1:1),

1.4% (PA1:30), 4.5% (PA1:100) and 8.0% (PA1:200) pyranoanthocyanin formation yields (Figure 4.3A). As shown in Figure 4.3A when increasing the concentration of caffeic and pyruvic acid the pyranoanthocyanin formation yields increased. Collision theory explain how chemicals must collide in order to react and more collisions result in greater reaction rates. Therefore, as the concentration of pyruvic acid and caffeic acid increased, the number of collisions between anthocyanins and cofactors increased

66 yielding more products (Brown et al., 2012). The data showed that formation of pyranoanthocyanins was affected by the cofactor concentration.

Low yields of pyranoanthocyanins were obtained under the conditions of this study for pyruvic acid samples. PA1:200 only had 8.0% pyranoanthocyanin formation yields, unlike our previous study which had yields of ~25%. As the concentrations of both the anthocyanins (500 µM to 150 µM) and the pyruvic acid (100 mM to 30 mM)

67 decreased, the pyranoanthocyanin formation decreased. Reaction rate likely depended on both anthocyanin and cofactor concentrations as a decrease in pyranoanthocyanin formation was observed when the anthocyanin concentration was decreased but an increase in pyranoanthocyanin formation was observed when the cofactor concentration increased. However, the degradation of anthocyanins and pyranoanthocyanins throughout the study limited the ability to create accurate rate-law graphs to verify if the reaction was second order (Wrolstad et al., 2005; Brown et al., 2012).

Figure 4.3A and 4.3B clearly demonstrated caffeic acid was more effective at forming pyranoanthocyanins than pyruvic acid. Pigments of sample CA1:30 were composed of 73.9% pyranoanthocyanins on day 42, and pigments of PA1:200 were composed of 20.4% pyranoanthocyanins on day 42 (Eq 4.2). Caffeic acid has been reported to have a co-pigmentation effect with anthocyanins. Co-pigmentation occurs when pigments interact with colorless compounds, generally by π-π stacking, to form an association that enhances the pigments color intensity. This intermolecular interaction may have helped stabilize the anthocyanin molecule, reducing degradation, and bring the reaction partners closer to each other accelerating the reaction (Castañeda-Ovando et al.,

2009). Additionally, Schwarz and others examined the reaction of anthocyanins with cinnamic acids (2003). They predicted that during pyranoanthocyanin formation an electron deficient carbenium intermediate was formed. The intermediate was likely stabilized by the electron donating, aromatic ring of the caffeic acid. The additional stability the aromatic ring provided likely facilitated pyranoanthocyanin formation

(Schwarz et al., 2003). Therefore, the intermolecular interaction and stabilizing effect of

68 the aromatic ring on caffeic acid may have helped increase pyranoanthocyanin formation compared to pyruvic acid.

4.4. The anthocyanins used in this experiment, cyanidin-3-glucoside and cyanidin-3- rutinoside, were simple anthocyanins with no acyl attachments limiting stability

(Wrolstad et al., 2005). CA1:30 had significantly more pigment left on day 42 (52.9%) than any other sample and was comprised of significantly more pyranoanthocyanins

(73.9%) (Figure 4.3B). The decreased pigment degradation for CA1:30 was likely attributed to the increased protection the additional D-ring of pyranoanthocyanins provide

(He et al., 2012; Oliveira et al. , 2014).

98.0%, CA1:10 had 96.8% and CA1:30 had 95.6% of the initial concentration of caffeic acid remaining on day 42 (Figure 4.5). The more pyranoanthocyanins that formed the more caffeic acid was used indicating the amount of caffeic acid used may have been correlated with pyranoanthocyanin formation. A smaller percentage of caffeic acid was used in comparison to pyruvic acid. As shown in Figure 4.5 PA1:100 and PA1:200 had

~75-80% of the cofactor remaining on day 42 (largest decrease). A clear trend in the cofactor remaining was not observed for pyruvic acid. Pyruvic acid is subjected to acid polymerization and decomposition when not in the pure form, so the large decreases in pyruvic acid may have been attributed to the decomposition and polymerization of pyruvic acid (Hazen, & Deamer, 2007; Müller & Baumberger, 1939). Caffeic acid had more pyranoanthocyanin formation and more cofactor remaining than the pyruvic acid samples indicating caffeic acid was more efficient at forming pyranoanthocyanins.

4.4.3 Spectral Properties

Visible absorbance spectra of the samples were recorded at each time point. The lambda maximum (λmax) increased for the control sample, likely due to polymerization,

Figure 4.6A. There was also an increase in the λmax for PA1:1, PA1:30, PA1:100, and

CA1:1. It was expected that when pyranoanthocyanins began to form there would be a hypsochromic shift in the λmax, which was seen for PA1:200, CA1:10 and CA1:30. The hypsochromic shifts in λmax were likely attributed to pyranoanthocyanin formation. For example, CA1:30 had the largest yield of pyranoanthocyanins and showed the largest decrease in λmax of 7 nm. Samples that had low yields of pyranoanthocyanins, such as

PA1:1 and CA1:1, followed trends similar to the control sample. The hypothesis that the more pyranoanthocyanins formed the larger the hypsochromic shift aligned with a study performed by Farr, Sigurdson and Giusti (2018).

The absorbance values in Figure 4.6 showed the absorbance at the λmax. All samples showed a decrease in absorbance throughout the study. Pigments were likely degrading, as shown in Figure 4.4, leading to a decrease in absorbance as chromophores degraded (Giusti, & Worlstad, 2005; Fossen, Andersen, & Cabrita, 2000). On day 42

CA1:30 had a significantly larger absorbance, 0.84, than any other sample. CA1:30 was made up of 73.9% pyranoanthocyanins on day 42. The large amount of 72 pyranoanthocyanins and co-pigmentation may have helped stabilize and protect the samples chromophores (He et al., 2012; Castañeda-Ovando et al., 2009).

4.4.4 Color Properties

As shown in Table 4.1 all samples had an increase in L* during storage, indicating the lightening of color. The control sample had an L* value of 72.1 on day 0 and 75.2 on day 42. While the CA1:30 sample hand an L* of 68.3 on day 0 and increased to 72.6 by day 42. The L* value of CA1:30, which contained 73.9% pyranoanthocyanins, had a similar L* value on Day 42 as the control sample did on day 0. The lower initial L* value of CA1:30 was likely attributed to anthocyanin co-pigmentation with caffeic acid, darkening the color (Castañeda-Ovando et al., 2009).

Table 3.1. Colorimetric data for all samples on day 0 and day 42. Data is present as average (n=3) and standard deviation in parenthesis

Chroma is the color intensity or how saturated the color is. Typically, as pigments degrade chroma decreases, because the intensity of the color decreases as degradation occurs. All samples had a decrease in C*ab. Table 4.1 shows on day 42 CA1:1 had a C*ab of 33.7, CA1:10 had a C*ab of 36.6 and CA1:30 had a C*ab of 42.7. It is likely that the trend of increasing chroma as caffeic acid concentration increased was attributed to both

74 the co-pigmentation effect of the caffeic acid and the increased concentration of pyranoanthocyanins (Castañeda-Ovando et al., 2009; He et al., 2012).

Table 4.1 displayed the hue angle of the samples, representative of the shade of the color; a hue angle of 0˚ is a red color and a hue angle of 90˚ is a yellow color. As the concentration of pyruvic acid increased the change in hab between day 0 and day 42 decreased. The smaller decreases in hab as pyruvic acid concentration increased may have been attributed to pyranoanthocyanin formation. As the concentration of caffeic acid increased an increase in hab was observed; CA1:30 had the greatest increase in hue angle from 10.6° (day 0) to 22.4° (day 42). CA1:30 shifted from red to more of an orange color.

Pyranoanthocyanins are orange in color which explains why CA1:30, which was predominantly pyranoanthocyanins on day 42, had an increase in hue angle.

As demonstrated by the change in L*, C*ab , hab all samples had a color change between day 0 and day 42. ΔE compares the color changes between two different samples and tells how different two colors are from each other. An average consumer would notice a ΔE around 5.0 while an expert would notice a ΔE around 3.0 (Wojciech & Tato,

2011). All samples had a ΔE around 15 when comparing the color on day 0 to the color on day 42. Therefore, all samples changed over the 42 day time period to an extent that an observed would notice the difference. Pyranoanthocyanins are more orange in color and as pigments degrade the color becomes more brown, so a color change was expected for all samples. CA1:1 had significantly more pigment degradation than CA1:30 and

CA1:30 had significantly higher pyranoanthocyanin yields than CA1:1, so since both had

75 similar ΔE it showed the color change was likely attributed to both pigment degradation and pyranoanthocyanin formation.

4.5 Conclusion

The type and concentration of cofactor had a significant effect on pyranoanthocyanin formation. Acetaldehyde and acetone showed no pyranoanthocyanin formation under the conditions of the preliminary study. Caffeic acid was much more efficient at forming pyranoanthocyanins than pyruvic acid. CA1:30 had a pyranoanthocyanin formation yield of 39.1% on day 42 (Eq 4.1) and PA1:200 had a pyranoanthocyanin formation yield of 8.0% on day 42. CA1:30 had the largest hypsochromic shift from day 0 to day 42 likely indicative of pyranoanthocyanin formation. It was predicted that co-pigmentation and the additional stability caffeic acid likely provided to reaction intermediates encouraged pyranoanthocyanin formation with caffeic acid. Caffeic acid samples also had a higher percentage of cofactor remaining at the end of the study than pyruvic acid samples, because pyruvic acid was likely undergoing polymerization. The study showed that caffeic acid was more effective at forming pyranoanthocyanins and increasing the concentration of cofactor increased pyranoanthocyanin yields. Cofactor plays a major role in pyranoanthocyanin formation, so more cofactors should be tested to increase pyranoanthocyanin yields.

Chapter 5. Conclusion and Future Work

Color remains an important quality parameter in food products, and the demand for colorants derived from nature brings many challenges to the food industry. This study provides information on how to more effectively form pyranoanthocyanins which are more stable than the anthocyanin counterpart. It was found that the pH of most rapid pyranoanthocyanin formation was near the pKH of the anthocyanin molecule. The anthocyanin has electrophilic character when in the flavylium state, so the pH needed to be acidic but not so acidic to accelerate pyruvic acid polymerization. The temperature of greatest pyranoanthocyanin formation was 25°C, and at 25°C an incubation time of 21 days was preferred over 30 days. Increased temperature helped increase the energy of the system and speed up pyranoanthocyanin formation. Caffeic acid was found to be the preferred cofactor in this study. Caffeic acid is a hydroxycinnamic acid, and the additional ring likely acted as a co-pigment bringing the reaction partners closer together and stabilizing the carbenium reaction intermediate. Additionally, as the concentration of the cofactor increased the pyranoanthocyanin formation yields increased.

Pyranoanthocyanin formation was generally associated with a hypsochromic shift in the lambda maximum.

Expanding off the current work would help to further increase pyranoanthocyanin yields. A major question left unanswered is the effect of anthocyanin concentration on

77 pyranoanthocyanin formation. A study in which the cofactor concentration is stabilized and the concentration of anthocyanins was changing would give insight into how concentrated the starting pigment should be to increase pyranoanthocyanin formation yields. Another interesting route the study could be expanded on is the use of different anthocyanins. This study used mainly cyanidin-3-glucoside with small amounts of cyanidin-3-rutinoside. Cyanidin-3-glucoside is a relatively unstable anthocyanin. It would be interesting to see the effect of an acyl attachment on pyranoanthocyanin formation. An acyl attachment would help to stabilize the anthocyanin molecule, but may also increase steric hinderance. Many questions remain unanswered, but this study has helped to increase pyranoanthocyanin formation yields by controlling pH, temperature and cofactors.

Bibliography

Andersen, Ø. M., Fossen, T., Torskangerpoll, K., Fossen, A., & Hauge, U. (2004).

Anthocyanin from strawberry (Fragaria ananassa) with the novel aglycone, 5-

carboxypyranopelargonidin. Phytochemistry, 65(4), 405–410.

https://doi.org/10.1016/j.phytochem.2003.10.014

Andersen, Ø. M., & Jordheim, M. (2006). The Anthocyanins. In Ø. M. Andersen & K. R.

Markham (Eds.), Flavonoids: Chemistry, Biochemistry, and Applications (pp. 472–

537). Boca Raton, FL: CRC Press (Taylor & Francis Group).

Andersen, Ø. M., & Jordheim, M. (2014). Basic Anthocyanin Chemistry and Dietary

Sources. In T. C. Wallace & M. M. Giusti (Eds.), Anthocyanins in Health and

Disease (1st ed., pp. 13–89). Boca Raton, FL: CRC Press (Taylor & Francis Group)

Bakker, J., Bridle, P., Honda, T., Kuwano, H., Saito, N., Terahara, N., & Timberlake, C.

F. (1997). Identification of an anthocyanin occurring in some red wines.

Phytochemistry, 44(7), 1375–1382. https://doi.org/10.1016/S0031-9422(96)00707-8

Bakker, J., & Timberlake, C. F. (1997). Isolation, Identification, and Characterization of

New Color-Stable Anthocyanins Occurring in Some Red Wines. Journal of

Agricultural and Food Chemistry, 45(1), 35–43. https://doi.org/10.1021/jf960252c

Benito, S., Morata, A., Palomero, F., González, M. C., & Suárez-Lepe, J. A. (2011).

Formation of vinylphenolic pyranoanthocyanins by Saccharomyces cerevisiae

and Pichia guillermondii in red wines produced following different fermentation

strategies. Food Chemistry, 124(1), 15–23.

https://doi.org/10.1016/j.foodchem.2010.05.096

Britton, G., Liaaen-Jensen, S., & Pfander, H. (Eds.). (2004). Carotenoids Handbook.

Compiled by Mercadante, A. Z., & Egeland, E. S.. Basel, Switzerland: Birkhauser

Verlag. Retrieved from https://link.springer.com/content/pdf/10.1007%2F978-3-

0348-7836-4.pdf

Brown, T. E., LeMay, E. H., Bursten, B. E., Murphy, C., Woodward, P. (2012). Chapter

14: Chemical Kinetics. In Jaworski, A., Zalesky, J., Haugen, T., Hart, J., Neumann,

J., Pierce, L., McDonald, C., Cheselka, G (Eds.) Chemistry: The Central Science,

Custom Edition for The Ohio State University (pp. 556-609. Boston, MA, USA:

Pearson Learning Solutions.

Brownmiller, C., Howard, L. R., & Prior, R. L. (2008). Processing and storage effects on

monomeric anthocyanins, percent polymeric color, and antioxidant capacity of

processed blueberry products. Journal of Food Science, 73(5), 72-79.

https://doi.org/10.1021/jf071994g

Burrows, A. (2009). Palette of our palates: A brief history of food coloring and its

regulation. Comprehensive Reviews in Food Science and Food Safety, 8(4), 394–

408. https://doi.org/10.1111/j.1541-4337.2009.00089.x

Cabrita, L., Fossen, T. & Andersen, Ø.M. (2000). Colour and stability of the six common

anthocyanidin 3-glucosides in aqueous solutions. Food Chemistry, 68(1), pp.101–

107.

Carvalho, A. R. F., Oliveira, J., de Freitas, V., Mateus, N., & Melo, A. (2010). A

theoretical interpretation of the color of two classes of pyranoanthocyanins. Journal

of Molecular Structure: THEOCHEM, 948(1–3), 61–64.

https://doi.org/10.1016/j.theochem.2010.02.020

Castañeda-Ovando, A., Pacheco-Hernández, M. de L., Páez-Hernández, M. E.,

Rodríguez, J. A., & Galán-Vidal, C. A. (2009). Chemical studies of anthocyanins: A

review. Food Chemistry, 113(4), 859–871.

https://doi.org/10.1016/j.foodchem.2008.09.001

Cevallos-Casals, A., & Cisneros-Zevallos, L. (2004). Stability of anthocyanin-based

aqueous extracts of Andean purple corn and red-fleshed sweet potato compared to

synthetic and natural colorants. Food Chemistry, 86, 69–77.

https://doi.org/10.1016/j.foodchem.2003.08.011

Clydesdale, F. M. (1993). Color as a factor in food choice. Critical Reviews in Food

Science and Nutrition, 33(1), 83–101. https://doi.org/10.1080/10408399309527614 81

Code of Federal Regulations (CFR). 2018a. 21CFR70.3: Color Additives, General

Provisions. Washington, DC: US Food and Drug Administration. Retrieved from

https://www.accessdata.fda.gov/scripts/cdrh/cfdocs/cfcfr/CFRSearch.cfm?fr=70.3

Code of Federal Regulations (CFR). 2018b. 21CFR70-82: Food and Drugs (Color

Additives). Washington, DC: US Food and Drug Administration. Retrieved from

https://www.accessdata.fda.gov/scripts/cdrh/cfdocs/cfcfr/cfrsearch.cfm

Dangles, O., & Fenger, J.-A. (2018). The Chemical Reactivity of Anthocyanins and Its

Consequences in Food Science and Nutrition. Molecules, 23(8), 1970.

https://doi.org/10.3390/molecules23081970

De Freitas, V., & Mateus, N. (2011). Formation of pyranoanthocyanins in red wines: A

new and diverse class of anthocyanin derivatives. Analytical and Bioanalytical

Chemistry, 401(5), 1467–1477. https://doi.org/10.1007/s00216-010-4479-9

Eiro, M. J., & Heinonen, M. (2002). Anthocyanin color behavior and stability during

storage: Effect of intermolecular copigmentation. Journal of Agricultural and Food

Chemistry, 50(25), 7461–7466. https://doi.org/10.1021/jf0258306

European Food Safety Authority. (2008). Assessment of the results of the study by

McCann et al. (2007) on the effect of some colours and sodium benzoate on

children’s behaviour: Scientific Opinion of the Panel on Food Additives,

Flavourings, Processing Aids and Food Contact Materials. The EFSA Journal 660:1-

54. https://doi/epdf/10.2903/j.efsa.2008.660 82

Fan-Chiang, H.J., Wrolstad, R.E. (2005). Anthocyanin Pigment Composition of

Blackberries. Journal of Food Science, 70(3), 198-202.

Farr, J. E., & Giusti, M. M. (2017). ColorBySpectra. Available online:

https://buckeyevault.com/products/colorbyspectra (accessed on 27 September

2018).

Farr, J., & Giusti, M. M. (2018). Investigating the Interaction of Ascorbic Acid with

Anthocyanins and Pyranoanthocyanins. Molecules, 23(4), 744.

https://doi.org/10.3390/molecules23040744

Farr, J. E., Sigurdson, G. T., & Giusti, M. M. (2018). Influence of cyanidin glycosylation

patterns on carboxypyranoanthocyanin formation. Food Chemistry, 259, 261–269.

https://doi.org/10.1016/j.foodchem.2018.03.117

Fernandes, A., Brás, N. F., Mateus, N., & De Freitas, V. (2015). A study of anthocyanin

self-association by NMR spectroscopy. New Journal of Chemistry, 39(4), 2602–

2611. https://doi.org/10.1039/c4nj02339k

Food and Drug Administration. 2017a. Color Additives: FDA’s Regulatory Process and

Historical Perspectives. Retrieved from

https://www.fda.gov/ForIndustry/ColorAdditives/RegulatoryProcessHistoricalPerspe

ctives/

Food and Drug Administration. 2017b. Color Certification FAQs. Retrieved from

https://www.fda.gov/ForIndustry/ColorAdditives/ColorCertification/ucm510897.ht

m#batch

Food and Drug Administration (FDA). 2018. Color Additives. Retrieved from

https://www.fda.gov/forindustry/coloradditives/default.htm

Fossen, T., & Andersen, Ø. M. (2003). Anthocyanins from red onion, Allium cepa, with

novel aglycone. Phytochemistry, 62(8), 1217–1220. https://doi.org/10.1016/S0031-

9422(02)00746-X

Fossen, T., Andersen, Ø. M., & Cabrita, L. (2000). Colour and stability of the six

common anthocyanidin 3-glucosides in aqueous solutions. Food Chemistry, 68(1),

101–107. https://doi.org/10.1016/S0308-8146(99)00170-3

Fulcrand, H., Benabdeljalil, C., Rigaud, J., Cheynier, V., & Moutounet, M. (1997). A

new class of wine pigments generated by the reaction between pyruvic acid and

grape anthocyanins. Phytochemistry, 47(7), 1401-1407.

Fulcrand, H., Cameira dos Santos, P. J., Sarni-Manchado, P., Cheynier, V., Favre-

Bonvin, J. (1996). Structure of new anthocyanin- derived wine pigments. Journal of

the Chemical Society, Perkin Transactions 1, 735-739.

Giusti, M. M., Rodriguez-Saona, L. E., Griffin, D., & Wrolstad, R. E. (1999).

Electrospray and Tandem Mass Spectroscopy As Tools for Anthocyanin

Characterization. Journal of Agricultural and Food Chemistry, 47, 4657-4664.

https://doi.org/10.1021/jf981242

Giusti, M. M., & Wrolstad, R.E. (2003). Acylated anthocyanins from edible sources and

their application in food systems. Biochemical Engineering Journal, 14, pp. 217-

225.

Giusti, M. M., & Worlstad, R. E. (2005). Characterization and Measurement of

Anthocyanins by UV-visible spectroscopy. In R. E. Wrolstad, T. E. Acree, E. A.

Decker, M. H. Penner, D. S. Reid, S. J. Schwartz, C. F. Shoemaker, D. Smith, P.

Sporns (Eds). Handbook of Food Analytical Chemistry (pp. F1.2.1-F1.2.13).

Hoboken, NJ, USA, NY: John Wiley &Sons

Golden, R. (2018). Dunkin’ Donuts Removes Artificial Dyes from All Donuts. Dunkin

Newsroom. Retrieved from https://news.dunkindonuts.com/blog/dunkin-donuts-

removes-artificial-dyes-from-all-donuts.

Hazen, R. M., & Deamer, D. W. (2007). Hydrothermal reactions of pyruvic acid:

Synthesis, selection, and self-assembly of amphiphilic molecules. Origins of Life

and Evolution of the Biosphere, 37(2), 143–152. https://doi.org/10.1007/s11084-

006-9027-4

He, F., Liang, N. N., Mu, L., Pan, Q. H., Wang, J., Reeves, M. J., & Duan, C. Q. (2012).

Anthocyanins and their variation in red wines II. Anthocyanin derived pigments and

their color evolution. Molecules, 17(2), 1483–1519.

https://doi.org/10.3390/molecules17021483

He, J., & Giusti, M. M. (2010). Anthocyanins : Natural Colorants with Health-Promoting

Properties. Annual Review of Food Science and Technology, 1, 163-187.

https://doi.org/10.1146/annurev.food.080708.100754

Jackson, A. (1976). Structure, properties and distribution of chlorophyll. In T.W.

Goodwin (Eds.), Chemistry and Biochemistry of Plant Pigments (pp. 1-63). New

York: Academic Press.

Konica Minolta Inc. (2007). Precise color communication. Minolta Co., 1-62

Lu, Y., & Foo, L. Y. (2001). Unusual anthocyanin reaction with acetone leading to

pyranoanthocyanin formation. Tetrahedron Letters, 42(7), 1371–1373.

https://doi.org/10.1016/S0040-4039(00)02246-2

Maga, J. A. (1974). Influence of color on tastes thresholds. Chemical Senses, 1.1(1764),

115–119.

Mateus N., & De Freitas, V. (2001). Evolution and stability of anthocyanin-derived

pigments during port wine aging. Journal of Agriculture and Food Chemistry, 49,

5217-5222.

McCann, D., Barrett, A., Cooper, A., Crumpler, D., Dalen, L., Grimshaw, K., …

Stevenson, J. (2007). Food additives and hyperactive behavior in 3-year-old and 8/9- 86

year-old children in the community: a randomized, double-blinded, placebo-

controlled trial. Lancet, 370(9598), 1560–1567. https://doi.org/10.1016/S0140-

6736(07)61306-3

McGuire, R. G. (1992). Reporting of objective color measurements. HortScience, 27(12),

1254–1255.

Milbury, P. E., Cao, G., Prior, R. L., & Blumberg, J. (2002). Bioavailability of elderberry

anthocyanins. Mechanisms of Ageing and Development, 123(8), 997–1006.

https://doi.org/10.1016/S0047-6374(01)00383-9

Morata, A., Calderón, F., González, M. C., Gómez-Cordovés, M. C., & Suárez, J. A.

(2007). Formation of the highly stable pyranoanthocyanins (vitisins A and B) in red

wines by the addition of pyruvic acid and acetaldehyde. Food Chemistry, 100(3),

1144–1152. https://doi.org/10.1016/j.foodchem.2005.11.024

Müller, O. H., & Baumberger, J. P. (1939). The Keto-Enol Tautomerism of Pyruvate Ion

Studied Polarographically. Journal of the American Chemical Society, 61(3), 590–

596. https://doi.org/10.1021/ja01872a016

National Weather Service. (2010). The Color of Clouds. Retrieved from

https://www.weather.gov/jetstream/color/

Nile, S. H., & Park, S. W.. (2014). Edible berries: Bioactive components and their effect

on human health. Nutrition, 30(2), 134-144.

https://doi.org/10.1016/j.nut.2013.04.007

Oliveira, J., Fernandes, V., Miranda, C., Santos-Buelga, C., Silva, A., De Freitas, V., &

Mateus, N. (2006). Color properties of four cyanidin-pyruvic acid adducts. Journal

of Agricultural and Food Chemistry, 54(18), 6894–6903.

https://doi.org/10.1021/jf061085b

Oliveira, J., Mateus, N., Silva, A. M. S., & De Freitas, V. (2009). Equilibrium forms of

vitisin B pigments in an aqueous system studied by NMR and visible spectroscopy.

Journal of Physical Chemistry B, 113(32), 11352–11358.

https://doi.org/10.1021/jp904776k

Quaglieri, C., Jourdes, M., Waffo-teguo, P., & Teissedre, P. (2017). Trends in Food

Science & Technology Updated knowledge about pyranoanthocyanins : Impact of

oxygen on their contents , and contribution in the winemaking process to overall

wine color. Trends in Food Science & Technology, 67, 139–149.

https://doi.org/10.1016/j.tifs.2017.07.005

Reed, M. (2018, January). New Year, New Colors – Dunkin Brands 2018

Announcement. Retrieved August 24, 2018, from https://sensientfoodcolors.com/en-

us/market-trends/new-year-new-colors-dunkin-brands-2018-announcement/

Rein, M. J., Ollilainen, V., Vahermo, M., Yli-Kauhaluoma, J., & Heinonen, M. (2005).

Identification of novel pyranoanthocyanins in berry juices. European Food Research

and Technology, 220(3–4), 239–244. https://doi.org/10.1007/s00217-004-1063-6

Rentzsch, M., Schwarz, M., & Winterhalter, P. (2007). Pyranoanthocyanins - an

overview on structures, occurrence, and pathways of formation. Trends in Food

Science and Technology, 18(10), 526–534. https://doi.org/10.1016/j.tifs.2007.04.014

Rodriguez-Saona, L. E., & Worlstad, R. E. (2001). Extraction, Isolation, and Purification

of Anthocyanins. In R. E. Worlstad, T. E. Acree, H. An, E. A. Decker, M. H.

Penner, D. S. Reid, & R. Sporns (Eds.). Current protocols in food analytical

chemistry (pp. F1.1-F1.11). Hoboken, NJ, USA, NY: John Wiley &Sons.

Romero, C., & Bakker, J. (2000). Effect of acetaldehyde and several acids on the

formation of vitisin A in model wine anthocyanin and colour evolution.

International Journal of Food Science and Technology, 35(1), 129–140.

https://doi.org/10.1046/j.1365-2621.2000.00372.x

Schwartz, S.J., Cooperstone J.L., Cichon M.J., Von Elbe, J.H., & Giusti, M.M. (2017).

Colorants. In S. Damodaran, & K. L. Parkin (Eds.), Fennema’s Food Chemistry

(5th ed., pp. 682– 752). Boca Raton, FL: CRC Press (Taylor & Francis Group).

Schwarz, M., Wabnitz, T. C., & Winterhalter, P. (2003). Pathway leading to the

formation of anthocyanin-vinylphenol adducts and related pigments in red wines.

Journal of Agricultural and Food Chemistry, 51(12), 3682–3687.

https://doi.org/10.1021/jf0340963

Sigurdson, G. T., Tang, P., & Giusti, M. M. (2017). Natural Colorants: Food Colorants

from Natural Sources. Annual Review of Food Science and Technology, 8(1), 261–

280. https://doi.org/10.1146/annurev-food-030216-025923

Somers, T. C. (1971). The polymeric nature of wine pigments. Phytochemistry, 10, 2175-

2186.

Stintzing, F. C., Stintzing, A. S., Carle, R., Frei, B., & Wrolstad, R. E. (2002). Color and

antioxidant properties of cyanidin-based anthocyanin pigments. Journal of

Agricultural and Food Chemistry, 50(21), 6172–6181.

https://doi.org/10.1021/jf0204811

Vallverdú-Queralt, A., Biler, M., Meudec, E., Guernevé, C. L., Vernhet, A., Mazauric, J.

P., … Dangles, O. (2016). P-hydroxyphenyl-pyranoanthocyanins: An experimental

and theoretical investigation of their acid—base properties and molecular

interactions. International Journal of Molecular Sciences, 17(11), 1–19.

https://doi.org/10.3390/ijms17111842

Wojciech, M., & Tato M. I. (2011). Color difference delta E – A Survey Colour

Difference ∆E – A survey. Machine Graphics and Vision 20(4): 383-411.

Wrolstad, R. E., Durst, R. W., & Lee, J. (2005). Tracking color and pigment changes in

anthocyanin products. Trends in Food Science and Technology, 16(9), 423–428.

https://doi.org/10.1016/j.tifs.2005.03.019

Wrolstad, R.E., & Smith, D.E. (2017). Color Analysis. In S.S. Nielsen (Ed.), Food

Analysis (5th ed., pp. 545-555). New York, NY: Springer.

Zhu, X., & Giusti, M.M. (2018). Pyranoanthocyanin Formation using Cyanidin

Derivatives and Caffeic Acid. Institute of Food Technologist Book of Abstracts.

Zhu, Z., Wu, N., Kuang, M., Lamikanra, O., Liu, G., Li, S., & He, J. (2015). Preparation

and toxicological evaluation of methyl pyranoanthocyanin. Food and Chemical

Toxicology, 83, 125–132. https://doi.org/10.1016/j.fct.201